DEVELOPMENT AND APPLICATIONS OF NOVEL SOLVENT … · 1.6 Solvent Extraction 13 1.7 Solid-Phase Extraction 15 1.8 Motivation of the Project 16 Chapter 2 Development of Novel Solvent-Minimized

DEVELOPMENT AND APPLICATIONS OF NOVEL

SOLVENT-MINIMIZED TECHNIQUES IN THE

DETERMINATION OF CHEMICAL WARFARE

AGENTS AND THEIR DEGRADATION PRODUCTS

LEE HOI SIM NANCY

NATIONAL UNIVERSITY OF SINGAPORE

2008

DEVELOPMENT AND APPLICATIONS OF NOVEL

SOLVENT-MINIMIZED TECHNIQUES IN THE

DETERMINATION OF CHEMICAL WARFARE

AGENTS AND THEIR DEGRADATION PRODUCTS

LEE HOI SIM NANCY

(M.Sc.), NUS

A THESIS SUBMITTED

FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

DEPARTMENT OF CHEMISTRY

NATIONAL UNIVERSITY OF SINGAPORE

2008

i

ACKNOWLEDGEMENTS

My most sincere gratitude goes to my bosses at DSO National Laboratories,

Ms. Sng Mui Tiang, Dr. Lee Fook Kay and Assoc. Prof. Lionel Lee Kim Hock, who

gave me the opportunity to pursue a part-time higher degree and provided me with

much encouragement and support throughout the course of my study. I had the

privilege of working under the expert guidance of Prof. Lee Hian Kee and Dr.

Chanbasha Basheer of the Department of Chemistry at the National University of

Singapore and would like to thank them for this enriching learning experience as well

as their friendship over the years. I sincerely appreciate the help and support from

everyone at the Defense Medical and Environmental Research Institute, DSO

National Laboratories, especially from the Organic Synthesis Group for providing the

analytes used in the study and all users of the GC MSD for sharing the use of the

instrument. Even though people come and go, the memories of the exciting times

especially during the Proficiency Tests, with Dr. Ang Kiam Wee, Ms. Chan Shu

Cheng, Mr. Leonard Chay Yew Leong, Dr. Alex Chin Piao, Dr. Chua Guan Leong,

Ms. Chua Hoe Chee, Mr. Willy Foo, Dr. Diana Ho Sook Chiang, Ms. Krystin Kee

Shwu Yee, Ms. Kwa Soo Tin, Mr. Le Tai Quoc, Dr. Lee Fook Kay, Ms. Leow Shee

Yin, Ms. Lim Hui, Mr. Neo Tiong Cheng, Ms. Ong Bee Leng, Ms. Linda Siow Siew

Lin, Ms. Sng Mui Tiang, Ms. Tan Sook Lan, Ms. Tan Yuen Ling, Ms. Jessica Woo

Huizhen, Ms. Veronica Yeo Mui Huang and Ms. Yong Yuk Lin, will remain with me

for a long time to come. Many thanks also to DSO National Laboratories for the co-

sponsorship throughout the course of my study. Lastly, special mention must be made

of my family members, especially Mum and Tony, who showed me much concern

despite being severely neglected while I worked long hours and throughout the

weekends. Thank you.

ii

TABLE OF CONTENTS

Acknowledgements i

Summary vii

List of Abbreviations ix

Chapter 1 Introduction

1.1 The Chemical Weapons Convention 1

1.2 Chemicals Related To The Chemicals Weapons Convention 2

1.3 The Organization for the Prohibition of Chemical Weapons

(OPCW) 8

1.4 The Official OPCW Proficiency Tests 9

1.5 Recommended Operating Procedures 13

1.6 Solvent Extraction 13

1.7 Solid-Phase Extraction 15

1.8 Motivation of the Project 16

Chapter 2 Development of Novel Solvent-Minimized Extraction Techniques

2.1 Solid-Phase Microextraction 17

2.1.1 Sol-gel SPME Fibers 19

2.1.2 Molecularly Imprinted Polymers for SPME Fibers 22

2.1.2.1 Molecular Imprinting 22

2.1.2.2 Sol-Gel Molecularly Imprinted Polymers 28

2.1.2.3 Current Status 30

2.1.3 Development of Novel SPME Coatings 32

2.2 Liquid-Phase Microextraction 34

iii

Chapter 3 Experimental

3.1 Sol-gel MIPs 40

3.1.1 Materials 41

3.1.2 Synthesis of MIPs and NIPs 42

3.1.3 Procedures 44

3.1.3.1 Evaluation of the effect of endcapping 44

3.1.3.2 Evaluation of the effect of elution solvents and

volume 45

3.1.3.3 Evaluation of binding properties 45

3.1.3.4 Comparison with other sample preparation

techniques 46

3.1.4 Instrumental Analysis 48

3.1.5 Synthesis of sol-gel MIP SPME fibers 48

3.2 SPME using PhPPP-coated fibers 49

3.2.1 Chemicals and Reagents 50

3.2.2 Preparation of PhPPP as a coating for SPME 50

3.2.3 Preparation of Stock Solutions and Samples 52

3.2.4 SPME Procedure 52


3.3 HF-LPME 53

3.3.1 Chemicals and Reagents 54

3.3.2 Preparation of Stock Solutions and Samples 55

3.3.3 Typical HF-LPME Procedures 56

3.3.4 Typical SPME Procedures 57


iv

3.4 Health and Safety Aspects 59

Chapter 4 Results and Discussion

4.1 Sol-gel MIPs 60

4.1.1 Effect of endcapping on PMPA-MIP-SPE 60

4.1.2 Evaluation of elution solvents and volume for

PMPA-MIP-SPE 63

4.1.3 Evaluation of binding properties by PMPA-MIP-SPE 64

4.1.4 Comparison of PMPA-MIP-SPE with other sample

preparation techniques 66


TDG-MIP-SPE 67

4.1.6 Evaluation of binding properties by TDG-MIP-SPE 69

4.1.7 Comparison of TDG-MIP-SPE with other sample



TEA-MIP-SPE 71

4.1.9 Evaluation of binding properties by TEA-MIP-SPE 72

4.1.10 Comparison of TEA-MIP-SPE with other sample



3Q-MIP-SPE 75

4.1.12 Evaluation of binding properties by 3Q-MIP-SPE 76

4.1.13 Comparison of 3Q-MIP-SPE with other sample


v

4.1.14 Evaluation of elution solvents for MIP-SPE using a

mixture of MIPs 78

4.1.15 Comparison of MIP-SPE using a mixture of MIPs with

other sample preparation techniques 79

4.1.16 Preparation of sol-gel MIP fibers 81

4.1.17 Conclusion 81

4.2 SPME using PhPPP-coated fibers 83

4.2.1 Optimization of SPME conditions 85

4.2.2 Method validation 88

4.2.3 Comparison with commercial SPME fibers 89

4.2.4 Conclusion 90

4.3 HF-LPME 91

4.3.1 Optimization of parameters for HF-LPME of chemical

agents 91

4.3.2 Method validation for HF-LPME of chemical agents 96

4.3.3 Comparison of HF-LPME of chemical agents with SPME 97

4.3.4 Optimization of parameters for HF-LPME of CWA

degradation products 100

4.3.5 Method validation for HF-LPME of CWA degradation

products 109

4.3.6 Comparison of HF-LPME of CWA degradation products

with SPME 110

4.3.7 Optimization of parameters for HF-LPME of basic

degradation products 111

vi

4.3.8 Method validation for HF-LPME of basic degradation

products 122

4.3.9 Comparison of HF-LPME of basic degradation products

with SPME 123

4.3.10 Analysis of a 20th Official OPCW Proficiency Test water

sample 124

4.3.11 Conclusion 127

Chapter 5 Concluding Remarks 128

Chapter 6 References 131

Appendices

Appendix 1 List of Schedule 1-3 Chemicals 156

Appendix 2 Total Ion Chromatograms from the MIP-SPE Study 160

Appendix 3 Mass Spectra 166

Appendix 4 List of Poster Presentations and Publications 173

vii

SUMMARY

Two approaches towards the development of solvent-minimized

microextraction techniques are presented in this report. The first approach involved an

attempt to develop solid-phase microextraction (SPME) fibers based on molecularly

imprinted polymers (MIP) synthesized via the sol-gel route for the extraction of

degradation products of chemical warfare agents. In the second approach, hollow

fiber-protected liquid-phase microextraction (HF-LPME) was utilized for the

determination of various chemical warfare agents and their degradation products.

Prior to the development of sol-gel MIPs as SPME fiber coatings, sol-gel

MIPs were first synthesized as powder and evaluated as sorbent packings in solid-

phase extraction (SPE) cartridges. A series of MIPs was synthesized using pinacolyl

methylphosphonic acid (PMPA), thiodiglycol (TDG), triethanolamine (TEA) and 3-

quinuclidinol (3Q) as the templates. A non-imprinted polymer (NIP) was also

synthesized, but in the absence of a template. The polymers were evaluated for their

binding properties towards their respective target analytes in aqueous matrices using

SPE. The elution solvent and volume of elution solvent were optimized for each MIP.

The MIP-SPE procedure was compared with other sample preparation procedures,

namely strong anion-exchange (SAX) SPE and strong cation-exchange (SCX) SPE as

well as a direct rotary evaporation procedure for the analysis of a range of analytes in

an aqueous sample containing polyethylene glycol (PEG).

Commercially-available SPME fibers in which the polymer coatings have

been stripped-off or damaged but with an intact fused silica backbone were used for

the preparation of sol-gel MIP SPME fibers. Several attempts to synthesize the sol-gel

MIP SPME fibers did not proceed well as the fiber coatings cracked and flaked off

viii

upon drying. Hence, efforts were focused on the evaluation of a novel SPME coating

based on poly(1-hydroxy-4-dodecyloxy-p-phenylene) polymer (PhPPP).

PhPPP was investigated as a coating for the SPME of Lewisites from aqueous

samples. Several extraction parameters, namely the choice of derivatizing agent, pH,

salting, and extraction time were thoroughly optimized. Upon optimization of the

extraction parameters, the performance of the novel coating was compared against

that of commercially-available SPME coatings.

HF-LPME was investigated for the extraction of various chemical warfare

agents and their degradation products from aqueous samples. Optimization of several

extraction parameters was carried out where the effects of the extraction solvent, the

derivatizing agent, derivatization procedure, the amount of derivatizing agent (for

degradation products), salting, stirring speed and extraction time were thoroughly

investigated. Upon optimization of the extraction parameters, the HF-LPME

technique was compared against SPME. In addition, the applicability of the technique

for a 20th Official OPCW (Organization for the Prohibition of Chemical Weapons)

Proficiency Test sample was demonstrated.

ix

LIST OF ABBREVIATIONS

A Absorptivity

ACN Acetonitrile

APTEOS 3-Aminopropyltriethoxysilane

BA Benzilic acid

BCVAA Bis(2-chlorovinyl)arsonous acid

BDT 1,4-Butanedithiol

BMPA Isobutyl methylphosphonic acid

BSTFA N,O-Bis(trimethylsilyl)trifluoroacetamide

BT Butanethiol

BZ 3-Quinuclidinyl benzilate

CAR/PDMS carboxen/polydimethylsiloxane

CAS Chemical Abstracts Service

CEES 2-Chloroethyl ethyl sulfide

CEPF O-Cyclohexyl ethylphosphonofluoridate

CEPS Chloroethyl phenylsulfide

CMPA Cyclohexyl methylphosphonic acid

CMPF Cyclohexyl methylphosphonofluoridate

CVAA 2-Chlorovinylarsonous acid

CWA Chemical warfare agent

CWC Chemical Weapons Convention

CW/DVB carbowax/divinylbenzene

DBPP O,O-Dibutyl n-propylphosphonate

DCHMP or DCMP O,O-Dicyclohexyl methylphosphonate

DCM Dichloromethane

x

DEDEP or DEDEPA O,O-Diethyl N,N-diethylphosphoramidate

DIMP O,O-Diisopropyl methylphosphonate

DIPAE 2-(N,N-Diisopropylamino)ethanol

DMEP O,O-Dimethyl ethylphosphonate

DMMP O,O-Dimethyl methylphosphonate

DVB/CAR/PDMS divinylbenzene/carboxen/polydimethylsiloxane

ECPP O-Ethyl O-cyclohexyl n-propylphosphonate

EDEA N-Ethyldiethanolamine

EDEPC O-Ethyl N,N-diethylphosphoramidocyanidate

EDT 1,2-Ethanedithiol

EGDMA Ethylene glycol dimethacrylate

EHES Ethyl 2-hydroxyethyl sulfide

EMPA Ethyl methylphosphonic acid

EPA Ethylphosphonic acid

ET Ethanethiol

EtOH Ethanol

GA Tabun or ethyl N,N-dimethylphosphoramidocyanidate

GB Sarin or isopropyl methylphosphonofluoridate

GC MS Gas chromatography-mass spectrometry

GD Soman or pinacolyl methylphosphonofluoridate

GF Cyclohexyl methylphosphonofluoridate

h hour(s)

HD or SM Sulfur mustard or bis(2-chloroethyl) sulfide

HF-LPME Hollow fiber-protected liquid-phase microextraction

HMDS Hexamethyldisilazane

xi

HN1 Bis(2-chloroethyl)ethylamine

HN2 Bis(2-chloroethyl)methylamine

HN3 Tris(2-chloroethyl)amine

HP Hewlett Packard

HPLC High performance liquid chromatography

IMPA Isopropyl methylphosphonic acid

ISTD Internal standard

L Leak

LC MS Liquid chromatography-mass spectrometry

LLE Liquid-liquid extraction

LOD Limit of detection

Log Kow Logarithm of octanol-water partition coefficient

L1 Lewisite 1 or 2-chlorovinyldichloroarsine

L2 Lewisite 2 or bis(2-chlorovinyl)chloroarsine

L3 Lewisite 3 or tris(2-chlorovinyl)arsine

MAA Methacrylic acid

MDEA N-Methyldiethanolamine

min minute(s)

MIP Molecularly imprinted polymer

MPA Methylphosphonic acid

MSD Mass selective detector

MTBSTFA N-(tert.-butyldimethylsilyl)-N-methyltrifluoroacetamide

MW Molecular weight

m/z mass-to-charge ratio

N Non-endcapped NIP

xii

NE Endcapped NIP

NIP Non-imprinted polymer

NMR Nuclear magnetic resonance spectrometry

nPPA n-Propylphosphonic acid

OPCW Organization for the Prohibition of Chemical Weapons

P Non-endcapped PMPA-MIP

PA polyacrylate

PDMS polydimethylsiloxane

PDMS/DVB polydimethylsiloxane/divinylbenzene

PDT 1,3-Propanedithiol

PE Endcapped PMPA-MIP

PEG polyethylene glycol

PhPPP Poly(1-hydroxy-4-dodecyloxy-p-phenylene) polymer

PIPA Propyl isopropylphosphonic acid

pKa Negative logarithm of acid dissociation constant

PMPA Pinacolyl methylphosphonic acid

PPA Propylphosphonic acid

PT Propanethiol

PTMOS Phenyl trimethoxysilane

Q Sesquimustard or 1,2-bis(2-chloroethylthio)ethane

QOH 1,2-Bis(2-hydroxyethylthio)ethane

R Recovery

r2 Squared regression coefficient

ROP Recommended operating procedure

rpm Revolutions per minute

xiii

RSD Relative standard deviation

SAX Strong anion-exchange

SCX Strong cation-exchange

SD Standard deviation

S/N Signal-to-noise ratio

SPE Solid-phase extraction

SPME Solid-phase microextraction

T O-mustard or bis(2-chloroethylthioethyl)ether

TBDMS tert.-butyldimethylsilyl derivative

TDG Thiodiglycol

TDGS Thiodiglycol sulfoxide

TDGSO Thiodiglycol sulfone

TE Triethylamine

TEA Triethanolamine

TEOS Tetraethoxysilane

TFA Trifluoroacetic acid

TMCS Trimethylchlorosilane

TMS Trimethylsilyl derivative

TOH Bis(2-hydroxyethylthioethyl)ether

TPP Tripropyl phosphate

VERIFIN Finnish Institute for Verification of the Chemical Weapons

Convention

VX O-Ethyl S-2-diisopropylaminoethyl methylphosphonothiolate

w/v weight per volume

3Q 3-Quinuclidinol

1

1 INTRODUCTION

1.1 The Chemical Weapons Convention

The Convention on the Prohibition of the Development, Production,

Stockpiling and Use of Chemical Weapons and on Their Destruction, also known as

the Chemical Weapons Convention (CWC), was opened for signature in Paris, France

on 13 January 1993. The Convention had been the subject of nearly twenty years of

negotiation with the aim to finalize an international treaty banning chemical weapons,

and designed to ensure their worldwide elimination.

The CWC entered into force on 29 April 1997. Today, there are 184 State

Parties with an additional 4 Signatory States that have signed the CWC. A State Party

is one that has signed and ratified or acceded to the CWC and for which the initial 30-

day period has passed (the CWC enters into force for a State only 30 days after its

ratification or accession to the treaty) whereas a Signatory State is one that signed the

CWC prior to its entry into force in 1997 but has yet to deposit its instrument of

ratification with the United Nations in New York. Only 7 Non-Signatory States

world-wide have not taken any action on the Convention. They are Angola,

Democratic People's Republic of Korea, Egypt, Iraq, Lebanon, Somalia and Syrian

Arab Republic. Singapore signed on 14 January 1993 and ratified on 21 May 1997 [1-

4].

The Convention is unique because it is the first multilateral treaty to ban an

entire category of weapons of mass destruction and to provide for the international

verification of the destruction of these weapon stockpiles within stipulated deadlines.

The Convention was also negotiated with the active participation of the global

chemical industry, thus ensuring industry's on-going cooperation with the CWC's

industrial verification regime. The Convention mandates the inspection of industrial

2

facilities to ensure that toxic chemicals are used exclusively for purposes not

prohibited by the Convention [2].

For the purpose of implementing the CWC, several terms have been defined as

follows. Chemical Weapons refers to (a) toxic chemicals and their precursors,

except where intended for purposes not prohibited under this Convention, as long as

the types and quantities are consistent with such purposes; (b) munitions and devices,

specifically designed to cause death or other harm through the toxic properties of

those toxic chemicals specified in subparagraph (a), which would be released as a

result of the employment of such munitions and devices; (c) any equipment

specifically designed for use in connection with the employment of munitions and

devices specified in (b). Toxic Chemical refers to any chemical, which through its

chemical action on life processes can cause death, temporary incapacitation, or

permanent harm to humans or animals. This includes all such chemicals, regardless of

their origin or their method of production, and regardless of whether they are

produced in facilities, in munitions or elsewhere. Precursor refers to any chemical

reactant that takes part at any stage in the production, by whatever method, of a toxic

chemical [5].

1.2 Chemicals Related To The Chemicals Weapons Convention

Besides the definitions, toxic chemicals and precursors, which have been

identified for the application of verification measures, are grouped into lists known as

Schedule 1, 2 and 3. The list of chemicals is tabulated in Appendix 1. Schedule 1

chemicals include those that have been or can be easily used as chemical weapons and

which have very limited, if any, uses for peaceful purposes. These chemicals are

subject to very stringent restrictions, including a ceiling on production of one ton per

3

annum per State Party, a ceiling on total possession at any given time of one ton per

State Party, licensing requirements, and restrictions on transfers. These restrictions

apply to the relatively few industrial facilities that use Schedule 1 chemicals. Some

Schedule 1 chemicals are used as ingredients in pharmaceutical preparations or as

diagnostics. The Schedule 1 chemical, saxitoxin, is used as a calibration standard in

monitoring programs for paralytic shellfish poisoning, and is also used in neurological

research. Ricin, another Schedule 1 chemical, has been employed as a biomedical

research tool. Some Schedule 1 chemicals and/or their salts are used in medicine as

anti-neoplastic agents. Other Schedule 1 chemicals are usually produced and used for

protective purposes, such as for testing chemical weapons protective equipment and

chemical agent alarms. Schedule 2 chemicals include those that are precursors to, or

that in some cases can themselves be used as, chemical weapons agents, but have a

number of other commercial uses (such as ingredients in resins, flame-retardants,

additives, inks and dyes, insecticides, herbicides, lubricants and some raw materials

for pharmaceutical products). For example, BZ (3-quinuclidinyl benzilate) is a

neurotoxic chemical listed under Schedule 2, which is also an industrial intermediate

in the manufacture of pharmaceuticals such as clindinium bromide. Thiodiglycol is

both a mustard gas precursor as well as an ingredient in water-based inks, dyes and

some resins. Another example is dimethyl methylphosphonate, a chemical related to

certain nerve agent precursors that is used as a flame retardant in textiles and foamed

plastic products. Schedule 3 chemicals include those that can be used to produce, or

can be used as chemical weapons, but which are widely used for peaceful purposes

(including plastics, resins, mining chemicals, petroleum refining fumigants, paints,

coatings, anti-static agents and lubricants). Among the toxic chemicals listed under

Schedule 3 are phosgene and hydrogen cyanide, which have been used as chemical

4

weapons, but are also utilized in the manufacture of polycarbonate resins and

polyurethane plastics as well as certain agricultural chemicals. Triethanolamine, a

precursor chemical for nitrogen mustard, is found in a variety of detergents (including

shampoos, bubble baths and household cleaners) as well as being used in the

desulfurization of fuel gas streams [2].

Based on their mode of action, that is, the route of penetration and their effect

on the human body, chemical agents are commonly divided into several categories:

nerve, blister, blood and choking agents [6,7]. The nerve agents such as Tabun, Sarin,

Soman, VX, chlorosarin and chlorosoman are listed in Schedule 1. The blister agents,

namely sulfur mustards, nitrogen mustards and Lewisites, are also listed in Schedule

1. The blood agents, for example hydrogen cyanide and cyanogen chloride, are listed

in Schedule 3. Phosgene, is an example of a choking agent and is listed in Schedule 3.

The nerve agents, known as cholinesterase inhibitors, interfere with the central

nervous system by reacting with the enzyme acetylcholinesterase and creating an

excess of acetylcholine which affects the transmission of nerve impulses [8]. The

classical symptoms of nerve agent poisoning includes difficulty in breathing, drooling

and excessive sweating, vomiting, cramps, involuntary defecation and urination,

twitching, jerking and staggering, headache, confusion, drowsiness, convulsion,

coma, dimness of vision and pinpointing of the pupils [9]. Nerve agent poisoning may

be treated with timely administration of antidotes such as atropine and diazepam.

The blister agents cause blistering of the skin and extreme irritation of the eyes

and lungs. They can be very persistent in the environment. These chemicals cause

incapacitation rather than death but can kill in large doses [10]. Some blister agents

like Lewisite and phosgene oxime are immediately painful while mustard agents may

cause little or no pain for as long as several hours after exposure. No effective medical

5

care exists for the treatment of mustard exposure and care is directed towards

relieving the symptoms and preventing infections [8].

The blood agents are substances that block oxygen utilization or uptake from

the blood, causing rapid damage to body tissues [9,11]. Symptoms are irritation of the

eyes and respiratory tract, nausea, vomiting and difficulty in breathing. Death from

poisoning follows quickly after inhalation of a lethal dose. The victim may recover

quickly from a smaller dose without assistance [12].

The choking agents cause physical injury to the lungs through inhalation.

Membranes may swell and lungs become filled with liquid, and in serious cases, the

lack of oxygen causes death [8]. Phosgene and chlorine are classified as choking

agents but in fact have several industrial uses as well.

Besides the above-mentioned major classes of chemical agents, there exist

incapacitating agents such as vomiting, tearing and riot control agents. These are

generally non-lethal agents that cause temporary physical or mental incapacitation

rather than death. BZ is a hallucinating agent that produces similar effects to atropine

such as changes in heart rate, confusion, disorientation, delusions and slurred speech.

Tearing agents cause irritation to the eyes and skin. Some examples are

chloroacetophenone, o-chlorobenzylidene malononitrile and dibenz-(b,f)-1,4-

oxazepine, which are used as riot control agents. Vomiting agents cause nausea and

vomiting and can also induce cough, headache, and nose and throat irritation. The

vomiting agents are typically solids which when heated, vaporize and condense to

form aerosols. Adamsite, an arsenic-containing chemical, is an example of a vomiting

agent [9,10].

The chemical agents are usually not stable and when subjected to natural

degradation in the environment or decontamination, a myriad of degradation products

6

arise through chemical processes such as hydrolysis, oxidation and elimination [13-

18]. In cases where the parent agent no longer exists, verification of the presence of

CWAs would most likely be based on the detection of the corresponding degradation

products. Hence, the analysis of degradation products of CWAs is equally if not more

important than that of the original substances. Table 1-1 lists the chemical agents and

their corresponding degradation products investigated in this study.

Table 1-1. Chemical agents and degradation products investigated in this study.

Chemical Agent Degradation Product(s)

Tabun (GA)

Not investigated

Sarin (GB) Isopropyl methylphosphonic acid (IMPA)

Soman (GD) Pinacolyl methylphosphonic acid (PMPA)

Cyclohexyl methylphosphonofluoridate (GF)

Cyclohexyl methylphosphonic acid (CMPA)

O-Ethyl S-2-diisopropylaminoethyl methyl phosphonothiolate (VX)

Ethyl methylphosphonic acid (EMPA)

2-(N,N-Diisopropylamino)ethanol (DIPAE)

NP

O

O

N

P

O

OF

P

O

OHO

P

O

OOH

P

O

OF

PO

O

SN

P

O

OHO

NOH

P

O

OHO

P

O

OF

7


Sulfur mustard (HD) Thiodiglycol (TDG)

Sesquimustard (Q) 1,2-Bis(2-hydroxyethylthio)ethane (QOH)

O-mustard (T) Bis(2-hydroxyethylthioethyl)ether (TOH)

Bis(2-chloroethyl)ethylamine (HN1)

N-ethyldiethanolamine (EDEA)

Bis(2-chloroethyl)methylamine (HN2)

N-methyldiethanolamine (MDEA)

Tris(2-chloroethyl)amine (HN3) Triethanolamine (TEA)

2-Chlorovinyldichloroarsine (L1) 2-Chlorovinylarsonous acid (CVAA)

Bis(2-chlorovinyl)chloroarsine (L2) Bis(2-chlorovinyl)arsonous acid (BCVAA)

Tris(2-chlorovinyl)arsine (L3)

Not investigated

ClS

Cl SOHOH

SOHS

OH

OS

OHS

OH

NOH OH

NOH OH

N

OH

OH OH

ClS

SCl

ClS

OS

Cl

ClN

Cl

ClN

Cl

ClN

Cl

Cl

AsCl

Cl

ClAs

OH

OH

Cl

As

Cl

Cl ClAs

OH

Cl Cl

AsCl Cl

Cl

8


3-Quinuclidinyl benzilate (BZ)

Benzilic acid (BA)

3-Quinuclidinol (3Q)

1.3 The Organization for the Prohibition of Chemical Weapons (OPCW)

The Chemical Weapons Convention mandated the Organization for the

Prohibition of Chemical Weapons (OPCW), an independent, international

organization based in The Hague, The Netherlands, to achieve the object and purpose

of the Convention, to ensure the implementation of its provisions, including those for

international verification of compliance with it, and to form a forum for consultation

and cooperation among State Parties. Among the numerous roles of the OPCW, a

complex verification regime is in place in order to ensure steps are taken towards

meeting the objectives of the Convention. On-site inspections and data monitoring are

conducted to ensure that activities within State Parties are consistent with the

objectives of the Convention and the contents of declarations submitted to the OPCW.

There are three types of inspections: routine inspections of chemical weapons-related

facilities and chemical industry facilities using certain dual-use chemicals; short-

notice challenge inspections which can be conducted at any location in any State

Party about which another State Party has concerns regarding non-compliance and

finally investigations of alleged use of chemical weapons [19].

During these inspections, sampling and on-site analysis may be undertaken to

check for the absence of undeclared scheduled chemicals. In cases of unresolved

OH

OHO

N

OH

NO

O

OH

9

ambiguities, samples may be sent to an off-site laboratory, subject to the inspected

State Party's agreement [5]. This off-site laboratory will be selected among several

OPCW designated laboratories. The designation of laboratories is determined through

their performance in the Official OPCW Proficiency Tests.

1.4 The Official OPCW Proficiency Tests

The OPCW proficiency testing scheme was set up with the objective to

simulate sample analysis in order to select laboratories that are capable of performing

trace analysis (at parts per million levels) of chemicals scheduled under the CWC

and/or their degradation products in a wide variety of matrices and of providing the

OPCW with a detailed report on the analysis results that contains analytical proof of

the presence of chemicals reported and provides high certainty of the absence of other

chemicals relevant for the implementation of the CWC and does not contain

information on chemicals not relevant to the CWC. Prior to the Official OPCW

Proficiency Tests, there were four international inter-laboratory comparison tests, also

known as round-robin tests, for laboratories to test the effectiveness of their

procedures for the recovery of CWC-related chemicals and their precursors and

degradation products from various sample matrices [20-23]. Thereafter, an additional

inter-laboratory comparison test [24] was conducted to further test the recommended

operating procedures [25] developed at the Finnish Institute for Verification of the

Chemical Weapons Convention (VERIFIN). Before the 1st Official OPCW

Proficiency Test in May 1996, two trial proficiency tests were held to train

laboratories and to establish procedures for the conduct of this first official test [26].

A laboratory may participate in the official proficiency tests as a regular

participant, whereby the laboratory is given fifteen calendar days to analyze the

10

samples and submit an analysis report to the OPCW [27]. Alternatively, a laboratory

may assist in one of two roles, that of the sample preparation laboratory or the

evaluating laboratory.

The sample preparation laboratory is tasked with formulating the composition

of test samples according to a test scenario, performing stability studies to ensure the

stability of spiking chemicals in the matrices, preparing the test samples as well as

dispatching a set to each of the participating laboratories in addition to two sets each

to the evaluating laboratory and the OPCW Laboratory. Thereafter, the sample

preparation laboratory proceeds to perform stability studies starting on the dispatch

date until the test period for all participants have expired. A sample preparation report

is submitted to the OPCW Laboratory within two weeks after the stability studies

have been completed. In addition, the sample preparation laboratory assists in the

categorization of the test chemicals and participates in the meeting held at the OPCW

Headquarters in The Hague to discuss the preliminary evaluation results with test

participants [28].

On the other hand, the evaluating laboratory is tasked with analyzing the

samples using at least two different analytical techniques, at least one of which must

be a spectrometric technique, to identify the test chemicals. Thereafter, the evaluating

laboratory submits a sample analysis report to the OPCW Laboratory within twenty

eight days upon receipt of the samples. Upon receipt of copies of the test reports from

participating laboratories (whereby pages identifying respective laboratories have

been removed by the OPCW Laboratory), the evaluating laboratory performs a

detailed evaluation of the reports and also assists in the categorization of the test

chemicals. A draft preliminary evaluation report will be sent to the OPCW Laboratory

within twenty eight days upon receipt of the complete set of copies of all participants'

11

reports. After discussion with the OPCW test coordinator on the draft preliminary

evaluation report, a preliminary evaluation report would be submitted to the OPCW

Laboratory within a week. The evaluating laboratory participates in the meeting held

at the OPCW Headquarters in The Hague to discuss the preliminary evaluation results

with test participants. Laboratories are allowed to submit comments on the

preliminary evaluation results in writing to the OPCW Laboratory within fourteen

calendar days following the preliminary evaluation meeting. If there are comments to

the preliminary evaluation results, the evaluating laboratory will conduct a re-

evaluation of the results affected by the comments, make corrections to the report and

submit the final evaluation report to the OPCW Laboratory within one week

following the receipt of the comments [29].

All participating laboratories that take part in an OPCW proficiency test will

be awarded a performance rating according to the extent to which the laboratory

fulfils the performance criteria as shown in Table 1-2. The sample preparation and

evaluating laboratories will be awarded the maximum performance rating of A

provided the test samples meet the required criteria and the evaluation of results is

performed satisfactorily and within set time lines.

Table 1-2. Method of evaluating performance rating [27].

Performance criteria fulfilled

Identification of chemicals

Performance scoring

Performance rating

Yes Laboratory identifies all chemicals

Maximum possible score

A

Yes Laboratory identifies all chemicals except one

Maximum possible score minus two

B

Yes Laboratory identifies more than half of the chemicals

Score between zero and (maximum possible minus two)

C

Yes Laboratory misses more chemicals than it identifies

Negative score D

No - No score Failure

12

The proficiency tests are a means for OPCW to assess laboratories in their

technical competence and for certifying laboratories that are seeking designation or

retention of designation status. To obtain designation, a laboratory should have

established a quality system and have a valid accreditation by an internationally

recognized accreditation body for the analysis of chemical warfare agents and related

compounds in various types of samples. In addition, a laboratory must obtain a rating

of three As or two As and a B in three consecutive proficiency tests in order for

designation. To retain the designation status, a laboratory must participate in the

proficiency tests at least once per calendar year and maintain the rating of three As or

two As and a B. Otherwise, it may be temporarily suspended or withdrawn in cases

where there is a substantial change in its accreditation status or deterioration in

performance in any proficiency test.

Since 1996 to 2007, a total of twenty two official OPCW proficiency tests

have been conducted. As of the 23rd Official OPCW Proficiency Test, there are

twenty designated laboratories worldwide, namely Belgium, China (two laboratories),

Czech Republic, Finland, France, Germany, India (two laboratories), The

Netherlands, Poland, Republic of Korea, the Russian Federation, Singapore, Spain,

Sweden, Switzerland, the United Kingdom and the United States of America (two

laboratories) [30]. Singapore's participation in the official proficiency tests is

undertaken by the Verification Laboratory of DSO National Laboratories. The

laboratory has been actively taking part since the 2nd Official Proficiency Test and

obtained the designation status in 2003 after the 10th, 11th and 12th Official OPCW

Proficiency Tests. The laboratory has assisted as a sample preparation laboratory in

the 14th Official OPCW Proficiency Test and as an evaluating laboratory in the 20th

Official OPCW Proficiency Test.

13

1.5 Recommended Operating Procedures

The Recommended Operating Procedures (ROPs) for Sampling and Analysis

in the Verification of Chemical Disarmament were proposed by VERIFIN and were

subsequently tested and improved upon through the round-robin tests. In all these

tests and in later proficiency tests, these ROPs have been widely and successfully

applied [5].

The ROPs provide instructions on sampling, sample collection, packing and

handling of samples as well as sample preparation, that is, sample treatment of

various matrices, including air samples, soil, wipe (swab of solid surfaces using

cotton buds, filter paper or glass filter beds), active charcoal, aqueous liquid, concrete,

paint, rubber and other polymeric samples followed by analysis using various

instrumental techniques such as gas chromatography (GC), gas chromatography-mass

spectrometry (GC MS), liquid chromatography-mass spectrometry (LC MS) and

nuclear magnetic resonance (NMR) spectrometry. The ROPs on sample treatment are

essential for the determination of analytes of interest in samples since very often some

form of sample treatment is required in order to extract and/or pre-concentrate the

analytes of interest prior to instrumental analysis. The ROPs are largely based on

solvent extraction and solid-phase extraction (SPE).

1.6 Solvent Extraction

Solvent extraction, in this context of sample treatment, is taken to include both

liquid-liquid extraction (LLE) as well as extraction from solid matrices. Solvent

extraction is often used interchangeably with LLE or liquid-liquid distribution, the

term recommended by The International Union of Pure and Applied Chemistry

(IUPAC). LLE involves the distribution of a solute between two immiscible liquid

14

phases in contact with each other. The solute, initially dissolved in only one of the

two liquids, eventually distributes between the two phases when equilibrium is

reached. LLE commonly takes place with an aqueous solution as one phase and an

organic solvent as the other. Solvent extraction can facilitate the isolation of

analyte(s) from the major component (matrix) and/or the separation of the particular

analyte from concomitant trace or minor elements [31]. LLE has been proven to be an

attractive method of concentrating various organic compounds from aqueous matrices

[32]. On the other hand, solvent extraction can also be applied to the extraction of

solutes from solid materials such as soils, sludges and sediments [33].

With regards to the ROPs for sample treatment of matrices for the analysis of

CWC-related chemicals, LLE of aqueous liquid samples is performed using

dichloromethane as the organic solvent in the first and second extractions for neutral

and basic analytes respectively while a strong cation-exchange cartridge is used to

recover polar acidic analytes in the third extraction. For the solvent extraction of soil

samples, dichloromethane is used in the first extraction in order to extract any non-

polar CWC-related chemicals while distilled, deionized water is used in the second

extraction for polar analytes followed by the use of 1% triethylamine in methanol in

the third extraction to recover basic CWC-related chemicals. Acetone or

dichloromethane can be used in the first extraction of concrete and polymeric

samples, followed by distilled, deionized water in the second extraction. For the

solvent extraction of wipe samples, several non-polar organic solvents such as

acetone, ethyl acetate, dichloromethane or deuterated chloroform (for subsequent

NMR analysis) are recommended for the first extraction while polar solvents such as

acetonitrile, methanol or water can be used in the second extraction. Suitable solvents

15

for the solvent extraction of active charcoal samples are acetone, dichloromethane,

carbon disulfide and deuterated chloroform (for subsequent NMR analysis) [5,25].

1.7 Solid-Phase Extraction

SPE is a form of step-wise chromatography designed to extract, partition,

and/or adsorb one or more components from a liquid phase (sample) onto a stationary

phase (sorbent or resin) [34]. Prior to the loading of the sample, the SPE sorbent is

first wetted and conditioned with solvents. The sorbent can be in the form of pre-

packed cartridges, columns or disks onto which analytes of interest are trapped. A

washing step is performed to remove contaminants trapped on the sorbent without

influencing the elution of the analytes of interest. Finally, elution with a suitable

solvent is carried out to recover the analytes of interest [35]. In this way, SPE may

serve to achieve concentration of the analytes, sample clean-up by removal of

interferences as well as sample matrix removal or solvent exchange into a form

compatible with instrumental analysis [36]. With SPE, many of the problems

associated with LLE, such as incomplete phase separations, less-than-quantitative

recoveries, use of expensive, breakable specialty glassware, and disposal of large

quantities of organic solvents, can be prevented. SPE is more efficient than LLE,

yields quantitative extractions that are easy to perform, is rapid, and can be

automated. Solvent use and laboratory time are reduced [37].

SPE is indeed an established sample treatment method [38] and has been

widely utilized in a variety of applications [36,37,39-49]. Besides, SPE has been

shown to be useful in the analysis of CWC-related chemicals in organic [26,50],

aqueous liquid [51-53], soil [54-56] and biological [57-61] matrices. The SPE

cartridges investigated included silica, C18, strong cation-exchange, strong anion-

16

exchange, quaternary amine and hydrophilic-lipophilic balance (HLB) cartridges. In

fact, SPE can be used in place of LLE in the ROPs for sample treatment of aqueous

samples for the analysis of CWC-related chemicals [62].

1.8 Motivation of the Project

The objective of the project is to improve on current extraction techniques for

the analysis of CWC-related chemicals through the development of solvent-

minimized extraction techniques. The current ROPs mainly utilize solvent extraction

which requires large volumes of organic solvents in addition to substantial amounts of

samples, typically 5 ml of aqueous samples or 5 g of soil samples or more.

Furthermore, the entire procedure is time-consuming and labor-intensive. Besides,

solvent extraction may not be specific such that analytes of interest are extracted

together with contaminants and interfering chemicals. To address these issues, two

approaches were undertaken.

The first approach involved an attempt to develop solid-phase microextraction

(SPME) fibers coated with molecularly-imprinted polymers (MIPs) synthesized via

the sol-gel route to address the issues of selectivity and analysis time. To date, there

have been hardly any reports on sol-gel MIP SPME fibers. At the same time, another

novel SPME coating based on a poly(paraphenylene) polymer was investigated. The

second approach utilized hollow fiber-protected liquid-phase microextraction (HF-

LPME) for the determination of various chemical warfare agents and their

degradation products. This approach allows the miniaturization of the extraction

procedure in terms of reducing the amount of sample, extracting solvents and

glassware required during sample treatment as well as shortening the entire sample

treatment process.

17

2 DEVELOPMENT OF NOVEL SOLVENT-MINIMIZED EXTRACTION

TECHNIQUES

2.1 Solid-Phase Microextraction

SPME is a solvent-free sample preparation technique [63]. Since its

introduction [64], SPME has been extensively investigated for a wide range of

applications [65-81]. In SPME, a 1 cm length of fused silica fiber, coated with a

polymer, is installed in a holder that looks like a modified microliter syringe. There is

also a stainless steel needle that the fiber can be withdrawn into to protect it. The

plunger moves the fused silica fiber into and out of the hollow needle. To use the unit,

the analyst draws the fiber into the needle, passes the needle through the septum that

seals the sample vial, and depresses the plunger, exposing the fiber to the sample or

the headspace above the sample. Organic analytes absorb into/adsorb onto the coating

on the fiber. After adsorption equilibrium is attained, usually in 2 to 30 minutes, the

fiber is drawn into the needle, and the needle is withdrawn from the sample vial.

Finally, the needle is introduced into the GC injector, where the adsorbed analytes are

thermally desorbed and delivered to the GC column, or into the SPME/HPLC

interface [82]. The SPME process is represented in Figure 2-1.

Figure 2-1. The extraction and desorption procedures in SPME [82].

18

SPME fibers are commercially available in various polymeric coatings and

thickness for both GC and LC applications. Commercially-available coatings include

polydimethylsiloxane (PDMS), carboxen/polydimethylsiloxane (CAR/PDMS),

polydimethylsiloxane/divinylbenzene (PDMS/DVB), polyacrylate (PA),

carbowax/divinylbenzene (CW/DVB), polyethylene glycol (PEG) and

divinylbenzene/carboxen/polydimethylsiloxane (DVB/CAR/PDMS) [83].

PDMS and PA fibers are absorptive fibers whereby analytes are extracted by

partitioning onto the liquid phase and are retained by the thickness of the coating.

PDMS/DVB, CW/DVB, CAR/PDMS and DVB/CAR/PDMS fibers are adsorptive

fibers which physically trap or chemically react and bond with analytes owing to their

porous nature and high surface area. The relatively new PEG fibers do not contain an

adsorbent polymer and are meant to replace the CW/DVB fibers [84]. In order to

select the best fiber for the target analyte, several parameters such as the molecular

weight and polarity of the analyte, the polarity and extraction mechanism of the fiber,

minimum detection limit and the linear range requirements have to be considered

[85,86].

A variety of commercial SPME coatings have been evaluated for the analysis

of chemical warfare agents and degradation compounds [87-96]. The coatings that

have been evaluated are 100 m PDMS, 85 m PA, 65 m PDMS/DVB, 65 m

CW/DVB and 75 m CAR/PDMS. Besides, a novel phenol-based polymer coating,

consisting of hydrogen bond acidic hexafluorobisphenol groups alternating with

oligo(dimethylsiloxane) segments, was designed for headspace SPME of Sarin [97].

19

2.1.1 Sol-gel SPME Fibers

Besides using commercial SPME fibers, SPME coatings made of silica-based

polymers can also be fabricated through a simple procedure known as the sol-gel

process. The sol-gel process refers to the preparation of ceramic materials by

preparation of a sol, gelation of the sol and removal of the solvent [98]. A sol is a

fluid, colloidal dispersion of solid particles in a liquid phase where the particles are

sufficiently small to remain suspended by Brownian motion. A gel is a solid

consisting of at least two phases wherein a solid phase forms a network that entraps

and immobilizes a liquid phase [99,100]. The sol-gel process involves mild reaction

conditions such that molecules, particularly those which are water soluble, may be

readily introduced within a highly crosslinked porous host without problems of

thermal or chemical decomposition. Materials in various configurations (for example,

films, fibers, monoliths and powders) can be prepared easily. Specific organic

functional groups can be combined with the inorganic precursor to introduce specific

chemical functionalities into the framework and improve molecular selectivity and

specificity. Furthermore, the materials are stable and the sol-gel processing conditions

can be varied to control the porosity and surface area of the resultant material [101].

The starting materials in the preparation of sol-gel materials are typically

inorganic metal salts or metal alkoxides (M(OR)n) such as tetramethoxysilane

(TMOS) or tetraethoxysilane (TEOS). A general sol-gel route is as follows:

Hydrolysis: M(OR)n + xH2O M(OR)n-x(OH)x + xROH

Condensation: M(OR)3(OH) + M(OR)3(OH) (RO)3M-O-M(OR)3 + H2O

or M(OR)3(OH) + M(OR)3(OH) (RO)3M-O-M(OR)2(OH) + ROH

20

At the end, every oxygen is bridging and hence a pure and highly

homogeneous oxide network is obtained [102]. Through the sol-gel process, materials

can be fabricated in many forms, such as thin films, membranes, powders, dense

ceramics and fibers. Useful applications of sol-gel materials include optical materials,

chemical sensors, catalysts, coatings, membranes, electronic materials and

chromatographic supports [103-109].

Several research groups are actively working on the area of sol-gel SPME

fibers because of the advantages of sol-gel SPME fibers over commercially-available

fibers. These advantages include enhanced thermal stability, solvent stability and the

ease with which inorganic or organic components can be incorporated into the

polymer framework. The enhanced thermal and solvent stability arises from the fact

that sol-gel coatings are chemically bonded to the fused silica backbone. In contrast,

most of the coatings of commercial fibers are physically deposited onto the fused

silica surface [110,111]. Another significant advantage is the fact that sol-gel coatings

contain polar moieties like silanol groups, resulting in the ability of seemingly non-

polar coatings such as PDMS or C11-PDMS to extract both polar and non-polar

analytes [112].

The pioneering work of Malik and co-workers demonstrated that sol-gel

PDMS coatings were capable of extracting both polar analytes such as

dimethylphenols, long chain alcohols and anilines as well as nonpolar analytes such

as polycyclic aromatic hydrocarbons (PAHs) and alkanes [113]. In contrast,

conventionally-coated PDMS fibers do not show sufficient selectivity for polar

compounds. The sol-gel PDMS fibers exhibited higher thermal stability compared to

conventionally-coated PDMS fibers. The sol-gel fibers can be routinely used at 320 C

and higher without any signs of bleeding. Enhanced thermal stability of sol-gel-coated

21

fibers allowed the sample carryover problem, often encountered in SPME of polar

solutes with conventional PDMS fibers, to be overcome. Sol-gel coatings possess a

porous structure and reduced coating thickness that provide enhanced extraction and

mass transfer rates in SPME. High-temperature conditioning of sol-gel-coated PDMS

fibers led to consistent improvement in peak area repeatability for SPME-GC

analysis. Peak area relative standard deviation values of <1% was obtained for PAHs

and dimethylphenols on sol-gel PDMS fibers conditioned at 320 C.

Besides conventional PDMS [114-116], PDMS/DVB [117] and PEG [111,

118-119] coatings, novel sol-gel coatings have been developed and evaluated.

Oligomers [120], novel polymers [121-131], fullerol [132], acrylates [133-136],

crown ethers [137-144], calix[4]arenes [145-151], cyclodextrins [152-154], hybrid

organic-inorganic materials [155,156], silica particles [157] and carbon [158,159]

have been incorporated as fiber coatings on SPME fibers. The fibers showed

improved thermal and chemical stability as well as good extraction efficiency of

target analytes.

Zeng and co-workers have published numerous papers of their work on sol-gel

SPME fibers. Of special interest in the analysis of chemical warfare agents and related

compounds is the development and evaluation of sol-gel PDMS/DVB for the

extraction of trimethylphosphate, tributylphosphate and dimethyl methylphosphonate

(a simulant of Sarin) [117] as well as PDMS coatings with acrylate components for

the extraction of 2-chloroethyl ethyl sulfide (CEES), a simulant of HD [134]. The

authors showed that the performance of the sol-gel PDMS/DVB fiber surpassed that

of commercially-available fibers such as 100 m PDMS, PA and PDMS/DVB for the

extraction of the phosphates and phosphonates from air and water samples. In the

other study, a comparison of acrylate components added to the sol mixture was made

22

and it was found that butyl methacrylate (BMA) gave the best results as compared to

methyl acrylate or methyl methacrylate. The sol-gel PDMS/BMA fiber surpassed that

of commercial PA for the extraction of CEES from soil.

2.1.2 Molecularly Imprinted Polymers for SPME Fibers

One drawback of current SPME coatings is that, other than the target analytes,

the fibers may absorb/adsorb other compounds present in the matrix, which would

possibly interfere with the analysis. One way of introducing selectivity to SPME

fibers is through the use of MIPs as fiber coatings.

2.1.2.1 Molecular Imprinting

Molecular imprinting is a technique used for preparing polymers with

synthetic recognition sites having a predetermined selectivity for the analyte(s) of

interest. The imprinted polymer is obtained by arranging polymerizable functional

monomers around a template (target analyte). Complexes are then formed through

molecular interactions between the analyte and the monomer precursors. These

interactions can either be non-covalent bonds, for example, ionic bonds and hydrogen

bonds, or reversible covalent bonds, for example, through boronic esters. Figure 2-2

depicts examples of the various interactions which may be employed in molecular

imprinting [160]. The complexes are assembled in the liquid phase and fixed by cross-

linking polymerization. Removal of the template from the resulting polymer matrix

creates vacant recognition sites that exhibit affinity for the analyte [161]. The concept

of molecular imprinting is illustrated in Figure 2-3.

23

Figure 2-2. Types of binding interactions that can be exploited during templating: (a) -

interaction; (b) hydrophobic or van der Waals interaction; (c) covalent bonds; (d) (transition) metal-ligand binding; (e) hydrogen binding; (f) crown ether -ion interaction; (g) ionic interaction [160].

Figure 2-3. Illustration of the concept of molecular imprinting [162].

Besides the specificity of MIPs, the other attractive features of MIPs are that

they are easy to prepare in different configurations such as block polymers, particles,

films or membranes and fibers, physically and chemically stable and reusable without

loss of the imprinting effect [163-165]. Interest in molecular imprinting technology

has grown at a phenomenal rate in recent years as seen from the number of original

publications (Figure 2-4) and is being extensively investigated for applications in

separations [167-196], sensors [198-202] and in synthesis and catalysis [203-208].

template and monomers complexation polymerisation

template removal recognition

24

Figure 2-4. A graphical representation of the number of publications within the field of molecular imprinting between 1931 and 1997 [166].

Several research groups have reported good results on the analysis of chemical

warfare agents and their degradation products using MIPs. The techniques of

molecular imprinting and sensitized lanthanide luminescence were combined to create

the basis for sensors that can selectively measure pinacolyl methylphosphonic acid

(PMPA), the hydrolysis product of the nerve agent, Soman, in water [209-213]. The

sensor functions by selectively and reversibly binding PMPA to a functionality-

imprinted copolymer possessing a coordinatively-bound luminescent lanthanide ion,

Eu3+. The MIP is formed by cross-linking styrene with divinylbenzene and templated

for PMPA for the detection of PMPA and isopropyl methylphosphonic acid (IMPA),

the hydrolysis products of Soman and Sarin respectively. This is feasible since the

polymer-bound functional end of PMPA is the same for both PMPA and IMPA. The

sensor is made from a fiber optic probe utilizing a luminescent europium complex.

The use of lanthanide ions as spectroscopic probes of structure and content is an

established technique. The narrow excitation and emission peaks of lanthanide

spectra, typically in the order of 0.01-1 nm full width at half maximum, provide for

the sensitive and selective analyses. Lanthanide complexes exhibit long luminescent

25

lifetimes and are intensely luminescent when complexed by appropriate ligands.

Proper ligand choice, used both to immobilize the lanthanide probe and provide the

enhancements needed for trace analysis, has been shown to provide limits of detection

in parts per trillion or lower [214]. The device has been constructed using europium as

the probe ion since its luminescence spectrum is least complex. Detection of the nerve

agent is based upon changes that occur in the spectrum when the hydrolysis product is

coordinated to Eu3+. This is seen from the presence of a peak at 610 nm in the laser-

excited luminescence spectrum of Eu(DMMB)3(NO3)2(PMPA) as compared to that of

Eu(DMMB)3(NO3)3, where the peak is absent. DMMB refers to the ligand, methyl-

3,5-dimethylbenzoate, which can be converted into polymerizable methyl-3,5-

divinylbenzoate, providing an avenue for self-crosslinking. In addition, the sensor was

evaluated for its physical properties such as luminescence properties, lifetime,

response time and pH dependence. The effect of interferences was also investigated.

The combination of molecular imprinting and luminescence detection provides

multiple criteria of selectivity to virtually eliminate the possibilities of false positive

readings. The sensor can be used in detecting the presence of chemical agents or

pollutants near battlefields, in hospitals or military installations, or in community

water supplies. Investigations were further extended to the imprinting of nerve agents

in which the sensors were evaluated against the presence of nerve agents in various

types of water such as tap, reverse osmosis, and deionized water [215].

In a study of MIP-SPE for the determination of degradation products of nerve

agents (Figure 2-5) in human serum [216], the absorptivities of several degradation

products of nerve agents, namely pinacolyl methylphosphonic acid (PMPA,

degradation product of Soman), ethyl methylphosphonic acid (EMPA, degradation

product of VX), isopropyl methylphosphonic acid (IMPA, degradation product of

26

Sarin), cyclohexyl methylphosphonic acid (CMPA, degradation product of GF),

isobutyl methylphosphonic acid (BMPA, degradation product of Russian VX) and

methylphosphonic acid (MPA, the final degradation product of all nerve agents) on

MIPs imprinted with PMPA, EMPA and MPA were investigated. It was shown that

the MIPs showed cross-selectivity as they could recognize not only the print molecule

but also the degradation products of the other nerve agents because the degradation

products of all nerve agents differ only in the alkyl chain of the phosphonate esters.

On the other hand, the non-imprinted polymer (NIP) showed no absorptivity. The NIP

is a polymer synthesized under the same conditions as the MIP but in the absence of

the template. This is believed to have arisen from the imprinting effect. It was

demonstrated that the cross-selectivity of the PMPA-MIP enabled the extraction of

the possible degradation products of all the nerve agents from human serum with

extraction recoveries of up to 90.5%. The interfering components for the capillary

electrophoresis analysis were successfully removed. Detection limits of 0.1 g ml 1

and relative standard deviations of <9% were obtained.

Figure 2-5. The structures of the nerve agent degradation products investigated [216].

In a separate study [217], a PMPA-imprinted MIP (PMPA-MIP) was

investigated for the analysis of EMPA using solid phase extraction. Two acrylate-

based MIPs were prepared using methacrylic acid (MAA) as a monomer and ethylene

P

O

O

OH

P

O

O

OH

P

O

O

OH

P

O

O

OH

P

O

O

OH

P

O

OH

OH

IMPA (Sarin)

PMPA (Soman) EMPA (VX)

CMPA (GF) BMPA (Russian VX)

MPA

27

glycol dimethacrylate (EGDMA) or trimethylolpropane trimethacrylate (TRIM) as

cross-linkers in dichloromethane and acetonitrile respectively. It was found that both

the MIPs as well as the NIP showed absorptivity of the analyte. This was in contrast

to the results of Meng and co-workers [216] where the NIP did not show any

absorptivity of the analytes. In order to achieve a difference in the recovery between

the MIP and the NIP, a washing step using 10 ml of acetonitrile:methanol (95:5) was

performed. With this step, the MIP gave 87% recovery of EMPA whereas the NIP

gave 34% upon elution with water. The MIP was next evaluated for the sample

cleanup of soil spiked with IMPA and CMPA. An Oasis hydrophilic-lipophilic

balance cartridge was selected to be used prior to the MIP cartridge, resulting in

sample cleanup and 95% recovery of the analytes.

A plastic antibody, that is, an MIP for the specific recognition of sulfur

mustard was fabricated using MAA and EGDMA [218]. The uptake of the plastic

antibody was compared against that of the NIP and the imprinting efficiency was

found to be 1.3. The imprinting efficiency is defined as the ratio of the binding ratio

of the MIP to the NIP, where the binding ratio is the ratio of the adsorbed analyte to

that remaining in solution. The plastic antibody did not experience interference from a

stimulant, dichlorodiethyl ether. The same research group further designed an MIP-

based potentiometric sensor for the specific recognition of MPA, the final degradation

product of nerve agents [219]. The sensor was fabricated from MAA and EGDMA

particles, dispersed in 2-nitrophenyloctyl ether followed by embedding in polyvinyl

chloride matrix to form a polymer membrane. The polymer membrane sensor can be

used for the analysis of MPA in natural waters in the presence of interfering

compounds such as phosphoric acids.

28

In a different approach [220], covalent imprinting was carried out first by

reacting MAA and EGDMA with 3,3-dimethylbutan-2-yl-4-vinylphenyl

methylphosphonate, a vinylphenol template carrying functional groups of PMPA.

This was followed by hydrolysis of the template using caesium fluoride. The binding

affinity of the MIP was studied by colorimetric detection methods where the

imprinting efficiency was determined to be 2.4.

Another interesting approach involved the introduction of a strong nucleophile

(hydroxamic acid) into the MIP to efficiently attack the phosphorus-fluoride bond of

organophosphonate nerve agents [221]. The proof-of-concept study showed that the

MIPs synthesized to be specific for Sarin and Soman, led to accelerated hydrolysis of

corresponding p-nitrophenyl substrates as compared to simultaneous hydrolysis in

buffer. Further studies on actual nerve agents are currently underway.

2.1.2.2 Sol-Gel Molecularly Imprinted Polymers

Besides acrylate polymers, sol-gel MIP materials produced from silane

monomers, have also been extensively investigated for applications mainly in

separations and sensors [222-247]. Of particular interest is the work by Marx et al.

since some of the analytes of interest studied were organophosphorus pesticides. In a

study on the detection of organophosphate pesticides using thin film sol-gel MIPs

[248], it was shown that the parathion-imprinted sol-gel films were highly selective

for the template molecule and not for very closely similar analytes such as methyl-

parathion, paraoxon, triethylphosphate and fenitrothion (Figure 2-6). In addition, the

sol-gel MIP exhibited selective binding of the analyte even in an aqueous matrix. The

sol-gel MIP films were further investigated for specific binding of analytes in the gas

phase [249].

29

Figure 2-6. Results showing high selectivity of the parathion-imprinted MIP [248].

Direct comparisons between acrylic-based MIPs and sol-gel MIPs were made

[231]. It was demonstrated that the thin film sol-gel MIPs, imprinted using

propranolol as a template, possess superior properties over the acrylic MIPs in terms

of selectivity for propanolol as compared to the corresponding NIPs. Even though the

sol-gel system had a lower capacity for binding, the non-specific binding was lower

than the acrylic system. In another study [250], acrylic and sol-gel MIPs imprinted for

2-aminopyridine were compared in terms of specificity and selectivity. Specificity

was defined as the success of the imprinting process as seen from the difference in

binding between the MIP and the NIP while selectivity was defined as the efficiency

in binding structural analogues of the template. It was found that the sol-gel MIP

showed a higher degree of specificity over the NIP in a polar solvent as compared to

the acrylic polymers. In terms of selectivity, the selectivity of the polymers for the

template over its structural analogues could be improved upon.

30

The use of sol-gel MIPs for the analysis of chemical warfare agents and

degradation products is in fact fairly limited. The work by Markowitz et al. is of note

where silica particles were surface-imprinted with PMPA, the degradation product of

Soman [251]. The particle surfaces were imprinted during particle formation by

adding PMPA to a microemulsion. The particles were functionalized with the addition

of organotrialkoxysilanes such as N-trimethoxysilylpropyl-N,N,N-trimethyl

ammonium chloride, 2-(trimethoxysilyl ethyl)pyridine or N-(3-triethoxypropyl)-4,5-

dihydroimidazole and the particle size, surface area, adsorption properties and binding

affinity of organophosphate compounds were studied. It was found that surface-

imprinted quaternary amine, 2-ethylpyridine- and dihydroimidazole-functionalized

silicates had a significantly higher degree of specificity for PMPA than for

structurally similar organophosphates. The binding properties were conducted using

2-propanol as a solvent. Another study focused on the recognition of MPA, the final

degradation product of nerve agents, using surface imprinting whereby the surface of

an indium tin oxide electrode was modified with octadecylsiloxane [252]. High

specificity, selectivity, stability and speed were demonstrated for this potentiometric

chemosensor.

2.1.2.3 Current Status

Thus far, reports on the use of MIPs as SPME fiber coatings have been scarce.

The first reports on MIPs as SPME adsorbents were published in 2001. The first study

[253] involved the use of propranolol-MIP particles for use as a capillary column

material for the determination of propranolol in serum samples by in-tube SPME.

Another study [254], which is also the first report on MIPs as SPME fiber coatings,

showed the feasibility of combining the selectivity of MIPs with the simplicity of

31

SPME. Silica fibers were coated with clenbuterol-imprinted polymers which were

subsequently used for extraction of clenbuterol from biological samples.

Bromobuterol (Figure 2-7), which is a structural analogue of the template,

clenbuterol, and is baseline separated in the liquid chromatography system, was used

to study the extraction and desorption characteristics of the MIP-coated fibers.

Selectivity of the extraction with these fibers was evaluated by comparison with fibers

coated with a NIP in order to investigate the non-selective interactions of the MIP.

The NIP was prepared in the same way as the MIP but without the clenbuterol

template. The selectivity of the MIP-coated fiber was shown by extraction yields of

about 75 and <5% respectively, for brombuterol from acetonitrile, using the imprinted

and non-imprinted polymers. Application of the fiber to the extraction of brombuterol

from spiked human urine gave clean extracts and ~45% yield, demonstrating the

suitability of the fibers for the analysis of biological samples.

Figure 2-7. The structures of clenbuterol and its structural analogue, brombuterol [254].

After a lapse of several years, publications on MIPs as SPME fiber coatings

started emerging in the recent couple of years. An interesting method of making MIP

SPME fibers involved filling untreated fused silica capillaries with the MIP followed

by etching of the fused silica with ammonium hydrogen difluoride [255,256]. Upon

optimization of parameters such as fiber thickness, polymerization time, loading,

Cl

NH2

Cl

NH

OH

Br

NH2

Br

NH

OH

clenbuterol brombuterol

32

washing and elution solvents as well as time and temperature of the loading step, the

fibers were used for the extraction of triazines from soil and vegetable samples.

In an alternative method, prometryn-imprinted polymers were coated onto

silica fibers for SPME of triazines in soil and crop samples [257,258]. Further work

by this research group involved the preparation and evaluation of MIP SPME fibers

for the trace analysis of tetracycline antibiotics in chicken feed, chicken muscle and

milk samples [259].

Using another straight-forward preparation method, monolithic codeine-

imprinted SPME fibers were fabricated through the use specially-prepared moulds.

The resulting fibers were 2 cm in length and 0.3 mm in diameter [260]. The method

was also used to fabricate diacetylmorphine-imprinted SPME fibers for the selective

extraction of diacetylmorphine and its structural analogues followed by GC and/or

GC MS analysis [261].

2.1.3 Development of Novel SPME Coatings

The MIP SPME coatings mentioned in the previous section are solely

acrylate-based coatings formed using methacrylic acid as the monomer and ethylene

glycol dimethacrylate or trimethylolpropane trimethacrylate as the cross-linker.

Hence, one of the aims of this project was to combine the fields of sol-gel chemistry,

molecular imprinting and solid-phase microextraction in the development of novel

SPME coatings in the form of sol-gel MIP SPME fibers for the selective extraction of

degradation products of chemical warfare agents from water samples. To the best of

our knowledge, there has only been one report on sol-gel MIP SPME fibers, for the

determination of ascorbic acid [262].

33

At the same time, another novel SPME coating, based on an asymmetrically

substituted polyhydroxylated poly(paraphenylene) polymer, poly(1-hydroxy-4-

dodecyloxy-p-phenylene) (PhPPP) [263], was investigated. The chemical structure of

the functionalized polymer used in the coating of the fiber is shown in Figure 2-8.

Figure 2-8. The chemical structure of PhPPP.

The incorporation of a long alkoxy chain and hydroxyl groups on either side

of the polymer backbone confers amphiphilic properties to the polymer backbone.

The polymer has been characterized [263-266] and investigated as a coating for

polymer-coated hollow fiber microextraction (PC-HFME) in which it functioned as

the adsorbent for the enrichment of organochlorine pesticides in water [267].

In PC-HFME (Figure 2-9), a short length of the hollow fiber membrane is

coated with a selected polymer. The extraction device is then placed into the sample

solution and allowed to tumble freely and continuously throughout the extraction.

Upon completion of extraction, the fiber is removed prior to desorption of the analytes

in a suitable solvent [268-271].

Figure 2-9. Experimental set-up for PC-HFME [267].

OH

C12H25O

n

34

In our investigation of the PhPPP coating for extraction, the polymer was

coated onto recycled commercial SPME fibers in which the polymeric coating had

been removed, leaving the fused silica backbone intact. The performance of this novel

fiber coating for the extraction of Lewisites from water samples was evaluated against

that of commercially-available fibers.

2.2 Liquid-Phase Microextraction

Single-drop microextraction (SDME) and hollow-fiber protected liquid-phase

microextraction (HF-LPME) are relatively new, solvent-minimized microextraction

techniques. In SDME [80,272-275], also referred to as single drop extraction (SDE)

[276-278], liquid-phase microextraction (LPME) [279-284], solvent microextraction

(SME) [285-287], single-drop LPME (SD-LPME) [288] or liquid-liquid-liquid

microextraction (LLLME) [289], a microdrop of solvent is suspended from the tip of

a conventional microsyringe and then immersed in a sample solution in which it is

immiscible or suspended in the headspace above the sample. After sampling, the

microdrop is retracted into the syringe and transferred to the analytical instrument.

The experimental set-up for SDME is depicted in Figure 2-10.

Figure 2-10. Experimental set-up for SDME [80].

35

An evolution from SDME involves the utilization of a polypropylene hollow

fiber membrane to contain the acceptor phase either in a U-shaped or a rod-like

configuration (Figures 2-11 and 2-12). The former technique was also known as

LPME when it was first described [290-296] whereas the latter has been referred to as

hollow fiber-protected LPME (HF-LPME) [297-307]. In the literature, the term,

LPME , has been used to refer to different techniques and configurations and tends to

lead to some confusion. Hence, the term, HF-LPME , will be used throughout the

rest of this text to specifically refer to the use of the hollow fiber membrane in the

rod-like (the so-called Lee-type) configuration. Alternative terminology for two-phase

HF-LPME include liquid-liquid microextraction (LLME) and microporous membrane

liquid-liquid extraction (MMLLE) [277] while three-phase HF-LPME has also been

referred to as liquid-liquid-liquid microextraction (LLLME) [308-315] and supported

liquid membrane (SLM) LLE [316].

Figure 2-11. Experimental set-up for HF-LPME in the U-shaped configuration [290].

Figure 2-12. Experimental set-up for HF-LPME in the rod-like configuration [297].

36

HF-LPME can be carried out as a two-phase or three-phase procedure. In two-

phase HF-LPME, the hollow fiber is affixed to the tip of the syringe needle and

contains an organic solvent for the extraction of analytes of interest from an aqueous

sample. Upon completion of extraction, the organic solvent is withdrawn into the

syringe and injected directly into a GC instrument for analysis [317]. The hollow fiber

is then discarded and thus resolves any issues of carry-over. In contrast, three-phase

HF-LPME involves extraction from an aqueous sample matrix, through an organic

phase, that is immiscible with water, held in the pores of the hollow fiber, and back

into a fresh aqueous phase inside the lumen (channel) of the hollow fiber [311].

Similar to SPME, besides direct immersion into an aqueous sample, headspace

SDME [318-324] and HF-LPME [325-327] can be carried out over a sample. In

addition, improvement to the technique is possible through dynamic HF-LPME, in

which small volumes of the aqueous sample is a repeatedly pulled in and pushed out

of the hollow fiber with the aid of a syringe pump. During withdrawal of the aqueous

sample, a thin film of organic solvent builds up in the hollow fiber and vigorously

extracts analyte from the sample segment, whereas, during sample expulsion, this thin

film recombines with the bulk organic phase in the syringe. During this

recombination, the portion of analyte extracted in the current cycle is trapped in the

bulk organic solvent. After extraction, which includes many repeated cycles, a portion

of the bulk organic solvent is subjected to further chromatographic analysis [275,328-

333]. Further improvements to the techniques involve the use of ionic liquids [334-

339] or binary solvents [340,341] as the extraction solvent.

Both SDME and HF-LPME techniques are simple and fast techniques to carry

out with the use of inexpensive apparatus. In addition, minimal volumes of solvents,

typically 5 to 50 l, are consumed such that these techniques are practically solvent-

37

less. High pre-concentration of analytes and excellent clean-up can be achieved.

However, SDME has its limitations: (a) the direct immersion mode requires careful

and elaborate manual operation due to the problem of drop dislodgement and

instability; (b) since more complex matrices will compromise the stability of the

solvent drop during extraction, an extra filtration step of the sample is usually

necessary; (c) the sensitivity and precision of SDME are not high due to relatively

long extraction time and the slow stirring rates since fast stirring usually results in

drop dissolution and/or dislodgement; and (d) SDME is not yet a routinely applicable

on-line pre-concentration procedure. Even though HF-LPME is also limited by the

unavailability of commercial equipment, with the protection of the hollow fiber

membrane, the technique is far more robust where samples can be stirred or vibrated

vigorously without any loss of the micro-extract. HF-LPME is expected to be an

important sample preparation technique, complementing existing techniques like

LLE, SPE and SPME [273,275,342,343].

Both SDME and HF-LPME have been investigated for the analysis of CWAs

and their degradation products. In the first report on SDME of CWAs and related

compounds [344], three toxic CWAs and six non-toxic markers of organophosphorus

nerve agents were investigated. The structures of the compounds are shown in Figure

2-13. Extraction parameters were optimized, leading to extraction conditions

requiring a combination of dichloromethane and carbon tetrachloride (3:1) as the

extracting solvent, a stirring rate of 300 rpm, addition of 30% of sodium chloride and

an extraction time of 30 min. The optimized SDME procedure was compared against

SPME and LLE for the extraction of compounds E-I (Figure 2-13). SDME was shown

to be superior over LLE and managed to extract all the compounds when SPME failed

to extract Sarin.

38

Figure 2-13. Structures of CWAs investigated in SDME [344].

In another study [345], HF-LPME was investigated for the extraction of

CWAs from water. The structures of the analytes investigated are shown in Figure 2-

14. The optimized extraction conditions were found to be the use of trichloroethylene

as the extracting solvent, a stirring rate of 1000 rpm, an extraction time of 15 min and

a salt concentration of 30% sodium chloride. It was found that HF-LPME is

advantageous over SDME in terms of operation, sensitivity and reproducibility. The

hollow fiber also provide a clean-up capability that does not apply to SDME whose

drop is exposed directly to the sample.

Figure 2-14. Structures of CWAs investigated in HF-LPME [345].

In a subsequent study [346], the successful analysis of degradation products of

CWAs using HF-LPME with in-situ derivatization was demonstrated. The structures

C2H5

39

of the analytes investigated are shown in Figure 2-15. Prior to extraction, basification

of the spiked water with potassium carbonate, addition of propyl bromide and stirring

of the mixture at 100 C for 2 h were carried out. Upon completion of reaction,

extractions were performed by HF-LPME prior to GC MS analysis. The method was

successfully used in the analysis of the 19th Official OPCW Proficiency Test water

samples.

Figure 2-15. Structures of alkylphosphonic acids investigated in HF-LPME [346].

This approach of developing solvent-minimized extraction techniques for the

determination of various CWAs and related compounds by utilizing HF-LPME was

carried out at approximately the same time as that reported by Gupta and co-workers

described above [345,346]. Besides CWAs and acidic degradation products, basic

degradation products were investigated as well. The results have been published as

three separate articles [347-349] and are discussed in Sections 3.3 and 4.3 of this

report.

40

3 EXPERIMENTAL

3.1 Sol-gel MIPs

Prior to the development of sol-gel MIPs as SPME fiber coatings, sol-gel

MIPs were first synthesized as powder and evaluated as sorbent packings in SPE

cartridges. A series of MIPs was synthesized using pinacolyl methylphosphonic acid

(PMPA), thiodiglycol (TDG), triethanolamine (TEA) and 3-quinuclidinol (3Q) as the

templates. A non-imprinted polymer was also synthesized, but in the absence of a

template. Using the sol-gel process, the polymers formed were crushed up into

powder and sieved. The templates were removed from the polymers by Soxhlet

extraction in ethanol. The polymers were then dried under vacuum.

Endcapping of the polymers was carried out by stirring the polymers in an

equimolar mixture of the endcapping reagents, trimethylchlorosilane (TMCS) and

hexamethyldisilazane (HMDS). The effect of endcapping of the polymers was

investigated by comparing the binding properties of the endcapped versus the non-

endcapped NIP and the PMPA-imprinted MIP when they were used as the sorbent in

SPE cartridges.

The various MIPs (PMPA-MIP, TDG-MIP, TEA-MIP and 3Q-MIP) and the

NIP were evaluated for their binding properties towards their respective target

analytes in aqueous matrices using SPE. The elution solvent and volume of elution

solvent were optimized for each MIP. Subsequently, the MIPs and the NIP were

challenged with water samples containing polyethylene glycol (PEG) to test their

binding properties for the analytes of interest in complex environmental samples

whereby the MIP-SPE procedure was compared with other sample preparation

procedures, namely strong anion-exchange (SAX) SPE and strong cation-exchange

(SCX) SPE as well as a direct rotary evaporation procedure for the analysis of a range

41

of analytes in an aqueous sample containing PEG. All experiments were carried out in

triplicate.

3.1.1 Materials

The monomers (Figure 3-1), tetraethoxysilane (TEOS) ( 99%, Fluka, Buchs,

Switzerland), phenyl trimethoxysilane (PTMOS) (98%, Lancaster, Lancashire,

England, UK) and 3-aminopropyltriethoxysilane (APTEOS) (99%, Aldrich,

Milwaukee, WI, USA), were used without further purification. Chemicals,

methylphosphonic acid (MPA) (98%, Aldrich), ethyl methylphosphonate (EMPA)

(98%, Aldrich), thiodiglycol (TDG) (99%, Merck, Darmstadt, Germany), thiodiglycol

sulfoxide (TDGS) (Chem Service, West Chester, PA, USA), thiodiglycol sulfone

(TDGSO) (65% weight % in aqueous solution, Lancaster), methyl diethanolamine

(MDEA) (99+, Aldrich), ethyl diethanolamine (EDEA) (98%, Aldrich),

triethanolamine (TEA) (98%, Aldrich), 3-quinuclidinol (3Q) (>98%, Fluka) and the

internal standard, tripropyl phosphate (TPP) (99%, Aldrich), were commercially

available while isopropyl methylphosphonate (IMPA), pinacolyl methylphosphonate

(PMPA) and cyclohexyl methylphosphonate (CMPA) were synthesized by the

Organic Synthesis Group of DSO National Laboratories. The endcapping reagents

(Figure 3-2) used were trimethylchlorosilane (TMCS) (98%, Aldrich) and

hexamethyldisilazane (HMDS) (99+%, Lancaster). The solvents used included

absolute ethanol (EtOH) ( 99.9%, Merck), tetrahydrofuran (GR, Merck), methanol

(HPLC, J.T. Baker, Phillipsburg, NJ, USA), dichloromethane (DCM) (ultra resi, J.T.

Baker) and acetonitrile (ACN) (HPLC, J.T. Baker). Concentrated hydrochloric acid

(HCl) (min. 37% AR, Riedel-de Haën, Seelze, Germany) was used as a catalyst as

well as for preparing 0.2 M HCl in methanol while trifluoroacetic acid (TFA) (99+%,

42

Aldrich) and triethylamine (TE) (>99%, Merck) were used as solvent modifiers.

Sodium hydroxide (NaOH) (Merck), of 1.0 M concentration, was used to clean the

fused silica fibers. N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA) (99% with 1%

TMCS, Aldrich) was used as a derivatizing agent for GC analysis. Poly(ethylene

glycol) (PEG) (Number average molecular weight, Mn, ca. 200, Aldrich) was used as

a background contaminant for the water matrix. Deionized water was obtained from a

Milli-Q system (Millipore, Bedford, MA, USA). Strong cation-exchange (SCX) and

strong anion-exchange (SAX) cartridges were commercially available as Varian Bond

Elute SCX (100 mg in 10 ml cartridges) and Supelco Supelclean LC-SAX (100 mg in

1 ml cartridges) respectively.

Figure 3-1. Structures of the monomers.

Figure 3-2. Structures of the endcapping reagents.

3.1.2 Synthesis of MIPs and NIPs

First, 120 ml of TEOS, 8 ml of PTMOS and 120 ml of ethanol were mixed.

Next, 4 ml of concentrated HCl, 8 ml of APTEOS and 40 ml of deionized water were

added dropwise in this order. The mixture was stirred at room temperature for 20 h.

Si Cl Si NH

Si

TMCS HMDS

SiO

OO

O Si

O

O

OO

NH2

SiOO

O

TEOS PTMOS APTEOS

43

For the PMPA-MIP, 100 ml portions of this sol were mixed with 10 ml of a 0.1 M

solution of PMPA (180 mg in ethanol). The solution was stirred for an additional 4 h.

The remaining mixture was used as the non-imprinted polymer [350,351]. The

solutions were then left to stand at room temperature until all the solvent evaporated

to yield hard and transparent polymers. The TDG-MIP, TEA-MIP and 3Q-MIP were

similarly synthesized. A 100 ml portion of the sol solution was mixed with 10 ml of a

0.1 M solution of TDG (127 mg in ethanol), 10 ml of 0.1 M solution of TEA (123 mg

in ethanol) and 10 ml of a 0.1 M solution of 3Q (149 mg in ethanol) respectively. The

polymers were crushed up and ground into fine powder using a mortar and pestle and

sieved through a 100 mesh (150 m) Coulter Particle Sizer sifter screen. The

polymers were repeatedly washed with ethanol using Soxhlet extraction. For the

MIPs, washing was stopped when the template could no longer be detected by GC

MS in the wash. The polymers were then dried under vacuum to constant weight.

Endcapping is a procedure in which the surface silanol groups were reacted

with an equimolar mixture of endcapping reagents. Here, TMCS and HMDS were

used [230]. Endcapping has been shown to prevent the non-specific adsorption of

analytes to surface silanol groups to a great extent and results in a higher imprint

factor [236].

In order to compare the effect of endcapping on the properties of both the NIP

and the MIPs, a portion of the various polymers were weighed out for the endcapping

procedure while the rest remained non-endcapped. Typically, 5 g of polymer was

stirred at room temperature for 24 h in an equimolar mixture of 2.5 g of TMCS and

3.75 g of HMDS. Next, the endcapped polymers were washed with anhydrous

tetrahydrofuran, followed by ACN to remove excess reagents [236] and then dried

under vacuum to constant weight.

44

3.1.3 Procedures

3.1.3.1 Evaluation of the effect of endcapping

First, 50 mg of PMPA-MIP or NIP was packed into recycled Varian 10 ml

SCX cartridges which have been emptied of the contents. The polymer powder was

packed between recycled frits in the cartridges. Next, 5 ml of a deionized water

sample containing 10 g ml 1 of MPA, EMPA, IMPA, PMPA and CMPA was

applied to the cartridges. The eluents collected were analyzed for the percentage

absorptivity, ie. the percentage of analyte that is retained on the polymer and is

obtained by subtracting from 100%, the percentage recovered in the eluent upon

loading of the sample. Elution with 5

2 ml of 1% TE in water, rotary evaporation to

dryness of the eluents and reconstitution in ACN, followed by derivatization using

BSTFA, gave the percentage recovery of the analytes from the polymers. TPP was

used as an internal standard (ISTD). The MIP-SPE procedure is shown in Figure 3-3.

Figure 3-3. Procedure for MIP-SPE.

Pack MIP or NIP intorecycled SCX cartridges.

Load sample. Rotary evaporate eluent to dryness.Reconstitute with 700 l of ACN.

Add 100 l of 100 ppm TPP (ISTD).Withdraw 400 l and add 100 l ofBSTFA. Heat at 60oC for 30 min.

Elute analytes withelution solvent.

Rotary evaporate eluent to dryness.Reconstitute with 700 l of ACN.


GC-MS analysis(% Recovery)

GC-MS analysis(% Absorptivity)

45

3.1.3.2 Evaluation of the effect of elution solvents and volume

The procedure for the packing of the SPE cartridges is as described in the

previous section. In the experiments, 5 ml of a deionized water sample containing 10

g ml 1 of PMPA was applied to the cartridges. For TDG-MIP SPE, 500 mg of TDG-

MIP or NIP were used. In the experiments, 0.5 ml of a deionized water sample

containing 10 g ml 1 of TDG was applied to the cartridges. For TEA-MIP SPE, 500

mg of TEA-MIP or NIP were used. In the experiments, 0.5 ml of a deionized water

sample containing 10 g ml 1 of TEA was applied to the cartridges. For 3Q-MIP SPE,

200 mg of 3Q-MIP or NIP were used. In the experiments, 0.5 ml of a deionized water

sample containing 100 g ml 1 of 3Q was applied to the cartridges.

In order to select the elution solvent that gives the highest recovery of the

analyte from the MIPs, 5

2 ml of ethanol, 1% TFA in water and 1% TE in water

were used separately as the elution solvent. In addition, the volume of elution solvent

required was investigated. Volumes of 1

1 ml, 1

2 ml, 3

2 ml, 4

2 ml, 5

2 ml

and 10

2 ml of elution solvent (giving a total volume of 1, 2, 6, 8, 10 and 20 ml

respectively) were applied to the cartridges.

3.1.3.3 Evaluation of binding properties

In order to evaluate the binding properties of the PMPA-MIP, 5 ml of a

deionized water sample containing 10 g ml 1 of MPA, EMPA, IMPA, PMPA and

CMPA was applied to the cartridges. For the TDG-MIP, 0.5 ml of a deionized water

sample containing 10 g ml 1 of TDG, TDGS and TDGSO was applied to the

cartridges. For the TEA-MIP, 0.5 ml of a deionized water sample containing 10 g

ml 1 of MDEA, EDEA and TEA was applied to the cartridges. For the 3Q-MIP, 0.5

46

ml of a deionized water sample containing 100 g ml 1 of 3Q was applied to the

cartridges. In these experiments, the NIPs were evaluated alongside the MIPs.

The eluents collected were analyzed for the absorptivity. Elution with the

optimal volume of the selected elution solvent, rotary evaporation to dryness of the

eluents and reconstitution in ACN, followed by derivatization using BSTFA, gave the

recovery of the analytes from the polymers. In addition, the polymers were further

challenged with a tap water sample containing the analytes as well as 500 g ml 1 of

PEG as a background contaminant. For such samples, the SPE procedure included a

washing step of 5 ml of water (Figure 3-4).

Figure 3-4. Procedure for MIP-SPE for samples with PEG.

3.1.3.4 Comparison with other sample preparation techniques

The MIP-SPE procedures were compared against other sample preparation

techniques using commercially-available SPE cartridges as well as with a procedure

Pack MIP or NIP intorecycled SCX cartridges.

Load sample.Rotary evaporate eluent to dryness.Reconstitute with 700 l of ACN.


Elute analytes withelution solvent.




GC-MS analysis(% Absorptivity)

Wash with 5 ml of water.GC-MS analysis

(% Leak)

47

involving no sample clean-up. To evaluate the PMPA-MIP, the sample consisted of

10 g ml 1 EMPA, IMPA, MPA, PMPA and CMPA with 500 g ml 1 PEG in tap

water. For the TDG-MIP, 0.5 ml of 10 g ml 1 TDG, TDGS and TDGSO with 500 g

ml 1 PEG in tap water was loaded onto the cartridges. For the TEA-MIP, 0.5 ml of 10

g ml 1 MDEA, EDEA and TEA with 500 g ml 1 PEG in tap water was loaded onto

the cartridges. For the 3Q-MIP, 0.5 ml of 100 g ml 1 3Q with 500 g ml 1 PEG in

tap water was loaded onto the cartridges. Figures 3-5 and 3-6 illustrate the SCX and

SAX SPE procedures respectively. For the procedure without sample clean-up, the

water sample was directly rotary evaporated to dryness, followed by reconstitution

with ACN and derivatization with BSTFA.

Figure 3-5. Procedure for SCX SPE. Figure 3-6. Procedure for SAX SPE.

Lastly, the respective MIPs (PMPA-MIP, TDG-MIP, TEA-MIP and 3Q-MIP)

were mixed in the ratio of 1:10:10:4 respectively and used for the extraction of the

Check pH of solution to ensurepH <\= 7

Load sample.

Condition SCX cartridge:a) 3 x 1 ml of MeOH

b) 3 x 1 ml of deionised water

Yes

No

Repeat CationExchange procedurewith another fresh

SCX cartridge


Add 50 l of 100 ppm TPP (ISTD) and100 l of BSTFA. Heat at 60oC for 30 min.


Wash with 1 ml deionised waterand 1 ml of MeOH.

Load sample.

Condition SAX cartridge:a) 2 x 1 ml of MeOH

b) 2 x 1 ml of deionised water


Add 50 l of 100 ppm TPP (ISTD) and100 l of BSTFA. Heat at 60oC for 30 min.


Elute analytes with 1 ml of0.2M HCl in MeOH.

48

four main template molecules from an aqueous sample. A 500 mg mixture of the

MIPs and corresponding NIP were used for the evaluation of the three elution

solvents, namely ethanol, 1% TFA in water and 1% TE in water. Next, 0.5 ml of a tap

water sample containing 10 g ml 1 PMPA, TDG and TEA and 100 g ml 1 3Q with

500 g ml 1 PEG as a matrix interference was loaded onto the MIP-SPE cartridges.

Elution with 5 × 2 ml of solvents was carried out. This procedure was similarly

compared against the SCX, SAX SPE procedures and the direct rotary evaporation

procedure.

3.1.4 Instrumental Analysis

GC MS analyses were performed on a HP6890 GC/5973 MSD system

(Agilent Technologies, San Jose, CA, USA). Separation was carried out on a HP-5MS

column, 30 m

0.25 mm internal diameter, 0.25 m film thickness, together with a 3

m precolumn. Splitless injections were performed. The temperature program used

was: 60 C, held for 2 min, ramped at 10 C min 1 to 220 C, then 20 C min 1 to

280 C, held for 4 min. The carrier gas was helium at 35 cm s 1. The split/splitless

injector was maintained at 200 C while the transfer line was maintained at 280 C.

The MS source and quadrupole were maintained at 230 and 150 C, respectively. All

analyses were performed in GC MS full scan mode over the range of m/z 40 550 at a

scan rate of 1.49 scans s 1.

3.1.5 Synthesis of sol-gel MIP SPME fibers

Commercially-available SPME fibers (Supelco, Bellefonte, PA, USA) in

which the polymer coatings have been stripped-off or damaged but with an intact

fused silica backbone were used for the preparation of sol-gel fibers. Any remaining

49

coating on the fibers was removed by dipping the fibers in DCM, after which the

coating could be stripped off easily. Prior to coating, the fused silica fibers were

dipped in 1 M NaOH for 1 h to expose the maximum number of silanol groups on the

surface, rinsed with deionized water, dipped in 0.1 M HCl for 30 min to neutralize

excess NaOH, rinsed with deionized water and dried at room temperature in a

desiccator.

To prepare the sol solution, 750 l of TEOS, 50 l of PTMOS and 750 l of

ethanol were mixed. Then, 25 l of concentrated HCl, 50 l of APTEOS and 250 l

of deionized water were added dropwise in this order. The mixture was stirred at

room temperature for 2 h. After this, 1 ml of this sol was mixed with 100 l of a 0.1

M solution of PMPA. The remaining mixture was used as the NIP coating. The

solutions were stirred for 24 h. A fused silica fiber was repeatedly dipped into the sol

solution such that a coating was formed on the fiber. The sol-gel fibers were dried at

room temperature in a desiccator.

3.2 SPME using PhPPP-coated fibers

PhPPP was investigated as a novel coating for the SPME of Lewisites from

aqueous samples. Several extraction parameters, namely the choice of derivatizing

agent, pH, salting, and extraction time were thoroughly optimized. Upon optimization

of the extraction parameters, the performance of the novel coating was compared

against that of commercially-available SPME coatings. All experiments were carried

out in triplicate, unless stated otherwise.

50

3.2.1 Chemicals and Reagents

The analytes, 2-chlorovinyldichloroarsine (L1), bis(2-chlorovinyl)chloroarsine

(L2) and tris(2-chlorovinyl)arsine (L3), were synthesized by the Organic Synthesis

Group of DSO National Laboratories and shown to be 95%, 88% and 92% pure by

NMR analysis respectively.

The solvents used included hexane, dichloromethane and acetonitrile (99.8%,

J.T. Baker). Sodium chloride (NaCl) (Merck) and sodium sulfate (Na2SO4) (J.T.

Baker) were used to investigate the effect of ionic strength of the sample during

extraction. Hydrochloric acid (0.1 M, Merck) and ammonium hydroxide (28-30 wt %

solution of NH3 in water, Acros Organics, Geel, Belgium) were used for adjustment

of sample pH. The derivatizing agents (Figure 3-7) used were ethanethiol (ET),

propanethiol (PT), butanethiol (BT) (97%, Fluka), 1,2-ethanedithiol (EDT) (98%,

Fluka), 1,3-propanedithiol (PDT) (99%, Aldrich) and 1,4-butanedithiol (BDT) (90%,

Fluka,). Deionized water was obtained from a Milli-Q system.

Figure 3-7. Structures of the mono- and dithiol derivatizing agents.

3.2.2 Preparation of PhPPP as a coating for SPME

PhPPP was synthesized as summarized in the following scheme [263]:

(i) Br2 in glacial acetic acid, 80%; (ii) NaOH, CH3(CH2)11Br, 45-50 C, 10 h, 65%;

(iii) K2CO3, C6H5CH2Br, 50 C, 10 h, 90%; (iv) n-butyllithium, tetrahydrofuran,

SHSH

SH

SHSH SHSH

SHSH

ET PT BT

EDT PDT BDT

51

78 C, triisopropyl borate, room temperature, 10 h, 70%; (v) 2 M K2CO3, toluene, 3.0

mol % Pd(PPh3)4, reflux, 3 days, (vi) H2, 10% Pd/C,

chloroform/ethanol/tetrahydrofuran.

Figure 3-8. Synthetic scheme of PhPPP [263].

Commercially-available SPME fibers with polymer coatings that were

stripped off but with the fused silica backbone intact were used for coating with

PhPPP. Thin films of the polymers on bare fused silica fibers were prepared by drop-

casting from 0.5 mg ml 1 polymer in chloroform solution under ambient conditions,

without any air flow or temperature control aids. SEM images were taken with a

JEOL JSM 6700 scanning electron microscopy (SEM) and the thickness of the fiber

was measured to be approximately 7 µm.

OH

HO

OH

HO

Br Br BrBr

OR

HO

Br Br

OR

BnO

(ii)

B(OH)2(HO)2B

OR

BnO

(i) (iii)

(iv)

1 2 3 4

5

BrBr

OR

BnOn

OBn

RO

OBn

RO

(v)

(vi)

n

OH

RO

OH

RO

R=C12H25

6

52

3.2.3 Preparation of Stock Solutions and Samples

A stock solution of a mixture of the analytes at a concentration of 1 mg ml 1

was prepared in acetonitrile and stored at 20 C. Aqueous samples (3 ml) were

freshly prepared by spiking deionized water with the analytes at a concentration of 0.5

µg ml 1. Quantitation of the analytes was done by external calibration where a series

of standard solutions was obtained by dilution of the stock solution with

dichloromethane, addition of the respective derivatizing agents and analysis by GC to

obtain linear calibration plots for each analyte based on the total ion chromatographic

peak area.

3.2.4 SPME Procedure

Prior to use, the network fiber was conditioned at 220 C for 30 min in the GC

injection port while the commercially-available fibers from Supelco, 30 m and 100

m PDMS, 65 m PDMS/DVB, 75 m CAR/PDMS, 85 m PA, and 65 m

CW/DVB, were conditioned according to the manufacturer's recommendations [83].

SPME analyses were performed by direct immersion of the fiber to a water sample

spiked with the analytes. Prior to extraction, 1 l of derivatizing agent was added. The

sample was stirred at the maximum rate during the extraction. After extraction, the

fiber was desorbed in the heated injection port of the GC for 5 min.


Optimization experiments were performed with a HP6890 GC, equipped with

a flame ionization detector (FID) (Agilent Technologies). Separation was carried out

on a HP-5 column, 30 m

0.25 mm internal diameter, 0.25 m film thickness.

Splitless injections were performed. The temperature program used was: 60 C, held

53

for 2 min, ramped at 10 C min 1 to 205 C, then 30 C min 1 to 280 C and held for 4

min. The carrier gas was helium at 35 cm s 1. The split/splitless injector was

maintained at 250 C while the detector was maintained at 280 C.

The limits of detection (LODs) for the analytes were estimated at S/N = 3

under GC MS full scan conditions. GC MS analyses were performed in electron

ionization mode (70 eV) on a HP6890 GC/5973 MSD system (Agilent Technologies).

Separation was carried out on a HP-5MS column, 30 m

0.25 mm internal diameter,

0.25 m film thickness, together with a 2 m precolumn. The split/splitless injector

was maintained at 250 C while the transfer line was maintained at 280 C. The

temperature program was identical to that used for the GC FID. The MS source and

quadrupole were maintained at 230 C and 150 C respectively. Analyses were

performed in GC MS full scan mode over the range of m/z 40-400 at a scan rate of

1.36 scans s 1.

3.3 HF-LPME

HF-LPME was investigated for the extraction of various chemical warfare

agents and degradation products from aqueous samples. Optimization of several

extraction parameters was carried out where the effects of the extraction solvent, the

derivatizing agent and derivatization procedure and the amount of derivatizing agent

(for degradation products), salting, stirring speed and extraction time were thoroughly

optimized. Upon optimization of the extraction parameters, the HF-LPME technique

was compared against SPME. In addition, the applicability of the technique for a 20th

Official OPCW Proficiency Test sample was demonstrated. All experiments were

carried out in triplicate, unless stated otherwise.

54

3.3.1 Chemicals and Reagents

The chemical warfare agents, isopropyl methylphosphonofluoridate (Sarin,

GB), pinacolyl methylphosphonofluoridate (Soman, GD), ethyl N,N-

dimethylphosphoramidocyanidate (Tabun, GA), bis(2-chloroethyl)sulfide (Sulfur

mustard, HD) and O-ethyl-S-[(2-(diisopropylamino)ethyl] methylphosphonothiolate

(VX), were synthesized by the Organic Synthesis Group of DSO National

Laboratories. Ethyl methylphosphonic acid (EMPA) (98%), ethyl 2-hydroxyethyl

sulfide (EHES) (97%), methylphosphonic acid (MPA) (98%) and n-propylphosphonic

acid (nPPA) (95%) were purchased from Aldrich, thiodiglycol (TDG) (99%) and

benzilic acid (BA) (98%) were bought from Merck and 1,2-bis(2-

hydroxyethylthio)ethane (QOH) (98%) was supplied by Acros. Isopropyl

methylphosphonic acid (IMPA), pinacolyl methylphosphonic acid (PMPA) and bis(2-

hydroxyethylthioethyl)ether (TOH) were synthesized by the Organic Synthesis Group

of DSO National Laboratories. The basic degradation products, 2-(N,N-

diisopropylamino)ethanol (DIPAE) (98%) and 3-quinuclidinol (3Q) (98%) were

bought from Fluka while N-methyldiethanolamine (MDEA) (99%), N-

ethyldiethanolamine (EDEA) (98%) and triethanolamine (TEA) (98%), were from

Aldrich.

The solvents used included chloroform (CHCl3) (99.9%) and trichloroethylene

(C2HCl3) (99%) from Aldrich, carbon tetrachloride (CCl4) (99.9%) and toluene

(99.9%) from Merck, tetrachloroethylene (C2Cl4) (99.7%), dichloromethane (CH2Cl2,

DCM) (99.8%) and acetonitrile (ACN) (99.8%) from J.T. Baker. Deionized water was

obtained from a Milli-Q system.

Sodium chloride (NaCl) from Merck and sodium sulfate (Na2SO4) from J.T.

Baker were used to investigate the effect of ionic strength of the sample on the

55

extraction. Solutions of hydrochloric acid (Aldrich) and sodium hydroxide (Acros

Organics) were used to adjust the pH of the samples. N,O-

Bis(trimethylsilyl)trifluoroacetamide (BSTFA) (99% with 1% TMCS) and N-(tert.-

butyldimethylsilyl)-N-methyltrifluoroacetamide (MTBSTFA) (97% with 1% tert.-

butyldimethylchlorosilane) purchased from Aldrich were used as derivatizing agents.

Figure 3-9. Structures of the silylating agents used in this work.

3.3.2 Preparation of Stock Solutions and Samples

A stock solution of a mixture of the CWAs at a concentration of 1 mg ml 1

was prepared in acetonitrile and stored at 20 C. Aqueous samples (3 ml) were

freshly prepared by spiking deionized water with CWAs at a concentration of 0.5 g

ml 1 of each while slurry samples were prepared by mixing 60 mg of soil (total

organic content 30 g kg 1) with 3 ml of deionized water prior to spiking CWAs at a

concentration of 0.5 g ml 1 (sample pH 6.5). Extraction of the samples was

performed after 20 min upon preparation. Quantitation of the analytes was done by

external calibration where a series of standard solutions was obtained by dilution of

the stock solution with dichloromethane and analysis by GC MS to obtain linear

calibration plots for each analyte based on the total ion chromatographic peak area.

A stock solution of a mixture of the analytes at a concentration of 2 mg ml 1

for EMPA, IMPA, BA, QOH and TOH, 1 mg ml 1 for EHES and PMPA and 20 mg

Si

OCF3

Si

NNCF3

Si

O

BSTFA MTBSTFA

56

ml 1 for MPA, nPPA and TDG was prepared in acetonitrile and stored at 20 C.

Additionally, a stock solution of a mixture of the analytes at a concentration of 10 mg

ml 1 for 3Q, MDEA, EDEA and TEA and 1 mg ml 1 for DIPAE was prepared in

acetonitrile and stored at 20 C. Based on the results of preliminary experiments the

analytes were prepared at varying concentrations as the extraction efficiency varies

significantly among the analytes. If all the analytes were spiked at a particular

concentration, analytes that were not easily extracted would not be detected while

those that were easily extracted would give saturated signals. Aqueous samples (3 ml)

were freshly prepared by spiking deionized water with the stock solution of the

mixture of analytes. Quantitation of the analytes was done by external calibration

where a series of standard solutions was obtained by dilution of the stock solution

with dichloromethane, addition of BSTFA or MTBSTFA respectively, heating at

60 C for 30 min and analysis by GC MS to obtain linear calibration plots for each

analyte based on the total ion chromatographic peak area.

3.3.3 Typical HF-LPME Procedures

The HF-LPME device consisted of a 10 l microsyringe with 0.63 mm outer

diameter cone-tip needle (SGE, Sydney, Australia) and a 1.2 cm Accurel Q3/2

polypropylene hollow fiber membrane (Membrana, Wuppertal, Germany) with the

dimensions, 600 m inner diameter, 200 m wall thickness and 0.2 m pore size,

attached to the needle tip.

For the extraction of the chemical agents, 5 l of solvent was drawn into the

syringe before a hollow fiber was affixed onto the tip of the syringe needle. The fiber

was immersed in solvent for three seconds to immobilize the solvent in the pores of

the hollow fiber prior to extraction. Upon immersion of the hollow fiber into the

57

sample solution, the syringe plunger was depressed to fill the hollow fiber with

solvent. The sample was stirred during the extraction with a Heidolph MR 3001K

(Kelheim, Germany) magnetic stirrer. After extraction, the analyte-enriched solvent

was drawn back into the syringe and the hollow fiber was discarded. A volume of 1 l

of solvent was injected into the GC MS. In a typical extraction of the degradation

products, a solvent-derivatizing agent mixture was first prepared by mixing

chloroform and MTBSTFA in equal amounts. The procedure is similar to that

described previously. The extraction of the basic degradation products follows that for

the degradation products except that 1 l of extract was injected into the GC MS

together with another microliter of derivatizing agent.

3.3.4 Typical SPME Procedures

The optimized extraction conditions for direct immersion SPME of the

chemical warfare agents were based on a previously-reported procedure [88].

Extractions were performed using a 65 m PDMS/DVB fiber. Prior to use, the fiber

was conditioned at 250 C for 30 min in the GC injection port. The fiber was directly

exposed to a sample spiked with the analytes with 40% (w/v) NaCl. The sample was

stirred at the maximum rate during the 30 min extraction. After extraction, the fiber

was desorbed in the heated injection port of the GC MS for 5 min. Direct immersion

SPME analysis of the water and slurry samples was performed using the same

extraction conditions.

The optimized extraction conditions for SPME of the degradation products

were based on a previously-reported procedure [90]. Extractions of the acidic

degradations were performed using a variety of fibers, namely 100 m PDMS, 65 m

PDMS/DVB, 75 m CAR/PDMS, 65 m CW/DVB and 85 m PA (Supelco) while

58

for SPME of the basic degradation products, the 30 m PDMS was used in place of

the 100 m PDMS fiber and in addition, the newly-introduced 60 m PEG fiber was

evaluated as well. Prior to use, the fibers were conditioned in the GC injection port

according to the manufacturer s recommendations [83]. The extraction procedure

involves first exposing a fiber to the MTBSTFA headspace for 5 min before directly

inserting into a sample spiked with the analytes. The sample was saturated with salt

and stirred during the entire extraction process. After extraction, the fiber was

exposed to the MTBSTFA headspace again for 15 min prior to desorption in the GC

injection port for 5 min.


GC MS analyses were performed in electron ionization mode (70 eV) with a

HP6890 GC/5973 MSD system. Separation was carried out on a HP-5MS column, 30

m×0.25 mm internal diameter, 0.25 m film thickness, together with a 2 m

precolumn. Splitless injections were performed. The carrier gas was helium at 35 cm

s 1. The split/splitless injector was maintained at 250 C while the transfer line was

maintained at 280 C. The MS source and quadrupole were maintained at 230 and

150 C, respectively. For the analysis of the chemical agents, the oven temperature

program used was: 40 C, held for 2 min, ramped at 10 C min 1 to 205 C, then 30 C

min 1 to 280 C and held for 4 min. All analyses were performed in GC MS full scan

mode over the range of m/z 40 400 at a scan rate of 1.36 scans s 1. The oven

temperature program used with BSTFA derivatization was: 60 C, held for 2 min,

ramped at 10 C min 1 to 205 C, 25 C min 1 to 280 C and held for 4 min while with

MTBSTFA derivatization it was: 60 C, held for 2 min, ramped at 10 C min 1 to

59

280 C and held for 4 min. All analyses were performed in GC MS full scan mode

over the range of m/z 40 550 at a scan rate of 1.49 scans s 1.

3.4 Health and Safety Aspects

Caution: extreme care was taken for synthesis of these compounds, as they are

highly toxic. Trained professionals should prepare, handle and use these compounds

in a fume hood equipped with an alkaline scrubber system and adopt proper protective

measures; and international treaties and legal issues when synthesizing or working

with these agents should be adhered to.

60

4 RESULTS AND DISCUSSION

4.1 Sol-gel MIPs

4.1.1 Effect of endcapping on PMPA-MIP-SPE

Preliminary experiments on the binding properties of the PMPA-MIP and the

NIP showed that the NIP did not demonstrate zero absorptivity of the analytes. It was

reported in the study on MIP SPE of nerve agent degradation products using acrylate-

based MIPs that the NIP could not absorb any of the analytes [216]. Hence, an

attempt was made to endcap the surface silanol groups of the sol-gel MIPs and NIPs

synthesized in order to reduce the non-specific adsorption of analytes to the surface

silanol groups to a certain extent and achieve a higher imprint factor [236].

Figure 4-1 shows the structures of the analytes of interest. These analytes are

of particular interest as they are the degradation products of a group of chemical

warfare agents known as the nerve agents. The phosphonic acids, EMPA, IMPA,

PMPA and CMPA, are the degradation products of VX, GA, GD and GF respectively

while MPA is the final degradation product of the phosphonic acids. The difference in

binding properties between the non-endcapped NIP (N) and the endcapped NIP (NE)

as well as that between the non-endcapped PMPA-MIP (P) and endcapped PMPA-

MIP (PE) for selected phosphonic acids, is shown in Figures 4-2 and 4-3 respectively.

Figure 4-1. Structures of the phosphonic acids investigated.

P

O

OHOH

P

O

OHO

P

O

OHO

P

O

OOH

P

O

OHO

MPA EMPA IMPA PMPA CMPA

61

Figure 4-2. Comparison of % absorptivity and recovery between non-endcapped and endcapped NIPs.

The percentage absorptivity (%A) is defined as the percentage of analyte that

is retained on the polymer and is obtained by subtracting from 100%, the percentage

recovered in the eluent upon loading of the sample. The percentage recovery (%R) is

defined as the percentage recovered in the eluent upon elution with elution solvent.

It is expected that the endcapping procedure would reduce non-specific

interactions between the analytes and the polymer. The % absorptivities of the

analytes were observed to be reduced by more than half except for MPA. MPA is the

most polar analyte and shows strong adsorption to the polymer. With endcapping,

recovery of the analytes from the NIP was negligible.

Ideally, the NIP should not show absorptivity of the analytes. The results

imply that the endcapping procedure may be insufficient to completely endcap all the

silanol groups on the polymer and hence eliminate all the non-specific interactions

between the analytes and the polymer such that the NIP shows zero absorptivity. The

endcapping procedure can either be repeated several times or the amounts of

endcapping reagents can be increased. However, as discussed below, endcapping of

the PMPA-MIP adversely affected the binding properties of the MIP.

0

20

40

60

80

100

EMPA IMPA MPA PMPA CMPA

%

%A (N) %R (N) %A (NE) %R (NE)

62

Figure 4-3. Comparison of % absorptivity and recovery between non-endcapped and endcapped PMPA-MIPs.

The non-endcapped PMPA-MIP shows 100% absorptivity for the analytes in

water as none of the analytes was detected in the eluent upon loading of the sample.

On the other hand, the endcapped PMPA-MIP does not show 100% absorptivity. This

observation can be attributed to two factors. Firstly, the process of endcapping has

reduced the non-specific interactions between the PMPA-MIP and the analytes to a

certain extent, hence the endcapped PMPA-MIP is expected to show lower

absorptivity of the analytes. Secondly, there exists competition between the analyte

and the water molecules in the aqueous matrix for binding sites on the polymer. This

would further reduce the absorptivity of the polymer for the analytes.

The recoveries of the analytes with 1% TE in water from the non-endcapped

and endcapped PMPA-MIPs were not quantitative. This indicates relatively strong

binding of the analytes with the polymer matrix such that even a strongly basic eluent

like 1% TE in water was unable to disrupt most of the analyte-polymer interactions. It

was observed that endcapping adversely affected the recovery of the analytes from the

endcapped MIP. Since the non-endcapped MIP showed better binding properties as

compared to the endcapped MIP, the non-endcapped PMPA-MIP and the other non-

endcapped MIPs were used, together with non-endcapped NIPs, for further

experiments.

0

20

40

60

80

100


%

%A (P) %R (P) %A (PE) %R (PE)

63

4.1.2 Evaluation of elution solvents and volume for PMPA-MIP-SPE

Three elution solvents were evaluated for their ability to recover the template,

PMPA, from the PMPA-MIP. Ethanol [249] was chosen as it successfully removed

the templates from the MIPs during MIP synthesis. The two other elution solvents,

1% TFA in water and 1% TE in water, were evaluated since TFA and TE are common

modifiers added to elution solvents used in molecular imprinting studies [352,353].

Although ethanol was effective in the removal of template from the polymer

by Soxhlet extraction, it could not be used as an elution solvent as it was not able to

elute PMPA from the PMPA-MIP. The elution solvents, 1% TFA in water (pH 0) and

1% TE in water (pH 11) were able to elute PMPA from the MIP, however, 1% TE in

water was more effective as it recovered a higher percentage of the analyte loaded.

Hence, 1% TE in water was chosen as the elution solvent in subsequent experiments.

Figure 4-4. Comparison of % recovery of PMPA with the various elution solvents.

Next, the volume of elution solvent required was investigated. Volumes of 1

1 ml, 1

2 ml, 3

2 ml, 4

2 ml, 5

2 ml and 10

2 ml of 1% TE in water (giving a

total volume of 1, 2, 6, 8, 10 and 20 ml respectively) were applied to the cartridges. A

comparison of percentage recovery using the various volumes of elution solvent is

presented in Figure 4-5. As seen from the graph, the optimum volume of elution

solvent was 10 ml. Increasing the volume of elution solvent to 20 ml did not increase

0

5

10

15

20

25

EtOH 1% TFA in H2O 1% TE in H2O

Elution solvent

% R

eco

very

of

PM

PA

64

the % recovery of the analyte significantly. Hence 5 × 2 ml of 1% TE in water was

used in subsequent experiments.

Figure 4-5. Comparison of % recovery of PMPA with the various elution volumes.

4.1.3 Evaluation of binding properties by PMPA-MIP-SPE

In order to study the binding properties of the PMPA-MIP as well as the NIP,

the polymers were challenged with a water matrix spiked with the phosphonic acids, a

range of analytes of similar structure to the template.

Figure 4-6. Comparison of % absorptivity and recovery of PMPA-MIP and NIP.

The % absorptivity and recovery of the PMPA-MIP and the NIP for the

analytes from deionized water are compared in Figure 4-6. It can be seen that the

PMPA-MIP shows 100% absorptivity for all the analytes while the NIP shows 100%

absorptivity for only MPA and particularly low absorptivity for PMPA.

0

10

20

30

0 5 10 15 20 25

Elution volume

% R

eco

very

of

PM

PA

0

20

40

60

80

100


%

%A (MIP) %R (MIP) %A (NIP) %R (NIP)

65

The study by Marx et. al. [249] showed that there was low cross-selectivity of

the parathion-imprinted films for other similar organophosphate pesticides and that

the non-imprinted films showed relatively lower binding. In this case, the PMPA-MIP

was able to quantitatively bind with all the phosphonates. This may not be a

disadvantage when it is necessary for the extraction of a range of unknown analytes of

interest from an environmental sample. Ideally, the NIP should not show binding of

the analytes. However, due to non-specific interactions of the analytes with the silanol

groups on the polymer, the NIP was able to bind to the analytes to a certain extent.

This problem was also encountered by other workers investigating the same polymer

system but with lisinopril dihydrate as the template [238]. The PMPA-MIP showed

similar recoveries of the analytes as compared to the NIP, except for the template

molecule PMPA, where the ratio in % recoveries of the PMPA-MIP to that of the NIP

was 5:1, the highest imprinting efficiency obtained among all the analytes.

Next, the polymers were challenged with a typical water matrix, one of most

common sample types prepared for analysis during the proficiency tests organized by

the OPCW. The analytes were spiked at a concentration of 10 g ml 1, together with

500 g ml 1 of PEG as a background contaminant, into tap water from the laboratory.

The % absorptivity, leak and recovery of the PMPA-MIP and the NIP of the analytes

from this water matrix are compared in Figure 4-7. The % leak (%L) is defined as the

% of analytes found in the wash during the washing step.

It was found that the presence of PEG and the use of tap water as the matrix

did not adversely affect the binding properties of the MIP and NIP as compared to

when deionized water was used as the matrix. The MIP showed a slight decrease in %

recovery for IMPA, PMPA and CMPA and an increase in % recovery for MPA. The

NIP showed a slight decrease in % absorptivity for EMPA, IMPA and PMPA and a

66

decrease in % recovery for all the analytes except for an MPA, for which there was an

increase. On comparison of the MIP and the NIP, it is observed that the ratio of the %

recovery of the MIP to that of the NIP for PMPA was 8:1, the highest among the

range of analytes.

The polymers did not retain much of the PEG present in the sample as PEG

was found in the eluent upon loading of the sample. The purpose of the additional

washing step was to remove more PEG from the polymers. The analytes did not elute

from the MIP during the washing step, whereas for the NIP, the analytes, except for

MPA, were detected in the wash eluent. The final eluent contained very insignificant

amounts of PEG, showing that the MIP-SPE procedure is effective in the sample

clean-up of a water matrix containing PEG as a background contaminant.

Figure 4-7. Comparison of % absorptivity, leak and recovery of PMPA-MIP and NIP for PEG samples.

4.1.4 Comparison of PMPA-MIP-SPE with other sample preparation

techniques

The PMPA-MIP-SPE procedure was compared against other SPE techniques,

namely SCX and SAX as well as a procedure without sample preparation. SCX and

SAX SPE cartridges are used for the removal of cations and PEG in water samples

respectively. These are usually present as background contaminants which may

0

20

40

60

80

100


%

%A (MIP) %L (MIP) %R (MIP) %A (NIP) %L (NIP) %R (NIP)

67

interfere with the analysis. Figure 4-8 shows the % recovery of the analytes using the

various procedures.

Figure 4-8. Comparison of % recovery of the phosphonic acids using the various procedures.

Of the four procedures, only the MIP-SPE and SAX SPE procedures were

capable of sample clean-up such that PEG was detected in insignificant amounts in

the final eluent. PEG was still present in the SCX SPE eluent as well as the sample

which was directly evaporated to dryness. Even though the % recovery of the analytes

from the MIP-SPE procedure cannot surpass that of the SAX SPE procedure, it has

been shown to be effective in the sample clean-up of aqueous samples containing

PEG for the analysis of phosphonates. The total ion chromatograms of the various

procedures are compiled in Appendix 2. The chromatograms were obtained by

reconstitution with DCM instead of ACN as ACN gives split peaks with the HP-5MS

column whereas DCM does not. This is due to the use of ACN, a more polar solvent

as compared to DCM, on the relatively non-polar HP-5MS column.

4.1.5 Evaluation of elution solvents and volume for TDG-MIP-SPE

Preliminary experiments with TDG-MIP showed that the polymer does not

give 100% absorptivity of TDG as TDG was detected in the eluent upon loading the

0

20

40

60

80

100


% R

eco

very

MIP-SPE Direct SCX SAX

68

sample. Hence, to improve on the % absorptivity, a larger amount of polymer was

used (500 mg) together with a smaller volume of sample (0.5 ml) as compared to

those used for PMPA-MIP-SPE. Figure 4-9 shows the % recovery of TDG from the

three elution solvents evaluated. EtOH and 1% TE in water gave comparable

recoveries while 1% TFA in water was unable to recover TDG from the MIP. The

highly acidic elution solvent promoted the interaction between the silanol groups and

TDG such that TDG could not be eluted from the MIP. Since 1% TE gave the highest

% recovery, it was used in subsequent experiments.

Figure 4-9. Comparison of % recovery of TDG with the various elution solvents.

Figure 4-10. Comparison of % recovery of TDG with the various elution volumes.

0

20

40

60

80

100


Elution solvent

% R

eco

very

of

TD

G

0

20

40

60

80

100

0 5 10 15 20 25

Volume of elution solvent (ml)

% R

eco

very

of

TD

G

69

From the graph, the optimum volume of elution solvent was 10 ml. A decrease

in % recovery was observed upon increasing the volume of elution solvent to 20 ml.

Hence 5 × 2 ml of 1% TE in water was used for subsequent experiments.

4.1.6 Evaluation of binding properties by TDG-MIP-SPE

Figure 4-11. Structures of the degradation products of HD.

Figure 4-11 shows the structures of the range of analytes used in the

evaluation of the binding properties of the TDG-MIP and the NIP. TDG is the

degradation product of the blister agent, HD, while TDGS and TDGSO are oxidation

products of TDG.

The % absorptivity and recovery of the TDG-MIP and the NIP for the analytes

from deionized water are compared in Figure 4-12. The TDG-MIP shows 100%

absorptivity only for TDGS while the NIP shows no recovery for TDGS. The %

absorptivity and recovery of the NIP are generally lower than that of the MIP.

Figure 4-12. Comparison of % absorptivity and recovery of TDG-MIP and NIP.

0

20

40

60

80

100

TDG TDGS TDGSO

%


SOHOH

SOHOH

O

SOHOH

O

OTDG TDGS TDGSO

70

Next, the polymers were challenged with tap water samples containing the

range of analytes with 500 g ml 1 of PEG. An attempt to include a washing step to

remove PEG in the TDG-MIP-SPE procedure resulted in the loss of the analytes as

the analytes were eluted during the washing step. Hence, the TDG-MIP-SPE

procedure was carried out according to Figure 3-3, that is, without a washing step.

This ensured reasonable recovery while compromising sample clean-up.

Figure 4-13. Comparison of % absorptivity and recovery of TDG-MIP and NIP for PEG samples.

The % absorptivity and recovery of the TDG-MIP of the analytes from PEG

samples was comparable with that from deionized water sample. The % absorptivity

of the analytes of the NIP showed a decrease in the presence of PEG. The % recovery,

however, remained unaffected. Again, it was observed that the % absorptivity and

recovery of the NIP are generally lower than that of the MIP.

4.1.7 Comparison of TDG-MIP-SPE with other sample preparation techniques

The TDG-MIP-SPE procedure was compared against other SPE techniques,

namely SCX and SAX as well as a procedure without sample preparation. Figure 4-14

shows the % recovery of the analytes using the various procedures.

0

20

40

60

80

100

TDG TDGS TDGSO

%


71

Figure 4-14. Comparison of % recovery of the mustard degradation products using the various procedures.

Of the four sample preparation techniques, only SAX SPE was able to

completely remove PEG from the sample such that PEG could not be observed in the

GC chromatogram. However, none of the compounds were recovered. It was

observed that the compounds were not retained on the cartridge and were eluted upon

loading of the sample onto the cartridge. This implies that TDG, TDGS and TDGSO

were not anionic at the sample pH (pH 6) and were thus not retained on the SAX

cartridge. Even though the TDG-MIP-SPE, SCX and direct rotary evaporation

methods could recover the analytes of interest, PEG could not be totally removed

from the aqueous sample. However, on comparison of the residual amount of PEG, it

is observed that the TDG-MIP-SPE procedure has considerably reduced the amount

of PEG. The total ion chromatograms of the various procedures are compiled in

Appendix 2.

4.1.8 Evaluation of elution solvents and volume for TEA-MIP-SPE

Like the TDG-MIP, the TEA-MIP does not give 100% absorptivity of TEA.

Hence, 500 mg of polymer was used together with 0.5 ml of sample. Figure 4-15

show the % recovery of TEA from the three elution solvents evaluated. All three

0

20

40

60

80

100

TDG TDGS TDGSO

%


72

elution solvents gave comparable recoveries with 1% TE giving the highest %

recovery. Hence, the latter was used in subsequent experiments.

Figure 4-15. Comparison of % recovery of TEA with the various elution solvents.

The volume of elution solvent did not significantly affect the % recovery of

TEA. A volume of 1 ml of 1% TE in water was sufficient for a reasonable recovery of

TEA from the MIP. The % recovery did not improve with the use of 10 or 20 ml of

elution solvent. From the graph, 4

2 ml of 1% TE was determined to be the

optimum volume of elution solvent required.

Figure 4-16. Comparison of % recovery of TEA with the various elution volumes.

4.1.9 Evaluation of binding properties by TEA-MIP-SPE

Figure 4-17 shows the structures of the analytes used in the evaluation of the

binding properties of the TEA-MIP and the NIP. The ethanolamines, EDEA, MDEA

0

10

20

30

40

50


Elution solvent

% R

eco

very

of

TE

A

0

10

20

30

40

50

60

0 5 10 15 20 25

Volume of elution solvent (ml)

% R

eco

very

of

TE

A

73

and TEA, are the degradation products of the nitrogen mustards, HN1, HN2 and HN3

respectively.

Figure 4-17. Structures of the ethanolamines.

The % absorptivity and recovery of the TEA-MIP and the NIP for the analytes

from deionized water are compared in Figure 4-18. Both the TEA-MIP and NIP show

similar % absorptivity and recovery for each of the analytes. The NIP shows slightly

higher % absorptivity of the analytes but lower % recovery as compared to the TEA-

MIP.

Figure 4-19 shows the % absorptivity and recovery of TEA-MIP for PEG

samples. The presence of PEG in the tap water resulted in a slight decrease in the %

absorptivity and recovery as compared to the deionized water sample for both the

TEA-MIP and the NIP. The % absorptivity and recovery of the NIP is generally lower

as compared to that of the TEA-MIP.

Figure 4-18. Comparison of % absorptivity and recovery of TEA-MIP and NIP.

0

20

40

60

80

100

MDEA EDEA TEA

%


NOHOH

NOHOH

OH

NOHOH

MDEA EDEA TEA

74

Figure 4-19. Comparison of % absorptivity and recovery of TEA-MIP and NIP for PEG samples.

4.1.10 Comparison of TEA-MIP-SPE with other sample preparation techniques

The TEA-MIP-SPE procedure was compared against other SPE techniques,



Only the TEA-MIP-SPE and the direct rotary evaporation procedures were

able to recover the ethanolamines from the PEG sample. The SCX and SAX SPE

procedures could not recover the ethanolamines, implying that the ethanolamines are

cationic at the sample pH (pH 6). The cationic ethanolamines were retained on the

SCX cartridge while they eluted upon loading onto the SAX cartridge. Only the SAX

SPE procedure was able to remove PEG from the sample. The total ion

chromatograms are compiled in Appendix 2.

The recovery of the ethanolamines using the TEA-MIP-SPE procedure was

less than half that of the direct rotary evaporation procedure. However, in terms of

sample cleanup, the amount of PEG introduced into the GC system was considerably

less as compared to the direct rotary evaporation procedure.

0

20

40

60

80

100

MDEA EDEA TEA

%


75

Figure 4-20. Comparison of % recovery of the ethanolamines using the various procedures.

4.1.11 Evaluation of elution solvents and volume for 3Q-MIP-SPE

Here, 200 mg of polymer was used together with 0.5 ml of sample. Figure 4-

21 shows the % recovery of 3Q from the three elution solvents evaluated. As 1% TFA

gave the highest % recovery of 3Q, it was used in subsequent experiments.

Figure 4-21. Comparison of the % recovery of 3Q with the various elution solvents.

Figure 4-22. Comparison of the % recovery of 3Q with the various elution volumes.

0

20

40

60

80

100

MDEA EDEA TEA%


0

10

20

30

40

50


Elution solvent

% R

eco

very

of

3Q

0

10

20

30

40

50

0 5 10 15 20 25

Elution volume

% R

eco

very

of

3Q

76

From the graph, the optimum volume of elution solvent was determined to be

10 ml. Increasing the volume to 20 ml led to a decrease in the % recovery. Hence, 5 ×

2 ml of 1% TFA in water was used in subsequent experiments.

4.1.12 Evaluation of binding properties by 3Q-MIP-SPE

Figure 4-23 shows the structure of 3Q which is the degradation product of the

psychogenic agent, BZ. The % absorptivity and recovery of the 3Q-MIP and the NIP

for 3Q from deionized water are compared in Figure 4-24. Both the 3Q-MIP and the

NIP do not show 100% absorptivity of the analyte. The % absorptivity and the

recovery of the NIP are lower as compared to that of the 3Q-MIP.

Figure 4-23. The structure of 3Q.

Figure 4-24. Comparison of the % absorptivity and recovery of 3Q-MIP and NIP.

Figure 4-25. Comparison of the % absorptivity and recovery of 3Q-MIP and NIP for PEG samples.

0

20

40

60

80

100

3Q

%


0

20

40

60

80

100

3Q

%


N

OH

3Q

77

The polymers were further challenged with a tap water sample containing 3Q

with 500 g ml 1 of PEG. Both the 3Q-MIP and NIP show a decrease in %

absorptivity with the tap water sample containing PEG. On the other hand, the %

recovery of the analyte is slightly higher as compared to the deionized water sample.

Here, the ratio of % recovery improved slightly over the deionized water sample.

4.1.13 Comparison of 3Q-MIP-SPE with other sample preparation techniques

The 3Q-MIP-SPE procedure was compared against other SPE techniques,



Figure 4-26. Comparison of % recovery of 3Q using the various procedures.

Only the 3Q-MIP-SPE and the direct rotary evaporation procedures were able

to recover 3Q from the PEG sample. Similar to the experiments with TEA, the SCX

and SAX SPE procedures could not recover 3Q, implying that 3Q is also cationic at

the sample pH (pH 8). The cationic 3Q was retained on the SCX cartridge [23] while

it eluted upon loading onto the SAX cartridge. Only the SAX SPE procedure was able

to remove PEG from the sample (Appendix 2).

The recovery of 3Q using the 3Q-MIP-SPE procedure marginally surpasses

that of the direct rotary evaporation procedure. In addition, in terms of sample

0

20

40

60

80

100

3Q

%


78

cleanup, the amount of PEG introduced into the GC system was considerably less as

compared to the direct rotary evaporation procedure.

4.1.14 Evaluation of elution solvents for MIP-SPE using a mixture of MIPs

Next, MIP-SPE using a mixture of the various MIPs was investigated for the

extraction of the various template molecules from aqueous matrices. From the

evaluation studies, 50 mg of PMPA-MIP, 500 mg of TDG-MIP and TEA-MIP and

200 mg of 3Q were used. Hence the MIPs, PMPA-MIP, TDG-MIP, TEA-MIP and

3Q-MIP, were mixed in the ratio of 1:10:10:4 respectively. The % absorptivity and

recovery using the elution solvents, ethanol, 1% TFA in water and 1% TE in water,

are compared in Figures 4-27 to 4-29.

Figure 4-27. Comparison of % absorptivity and % recovery of the mixture of MIPs and NIP with EtOH.

Figure 4-28. Comparison of % absorptivity and % recovery of the mixture of MIPs and NIP with 1% TFA in water.

0

20

40

60

80

100

3Q PMPA TDG TEA

%


0

20

40

60

80

100

3Q PMPA TDG TEA

%


79

Figure 4-29. Comparison of % absorptivity and % recovery of the mixture of MIPs and NIP with 1% TE in water.

Except for 1% TFA for the recovery of TDG where no recovery was obtained,

all the solvents were able to recover the analytes to varying extents. The highest %

recovery of 3Q and PMPA was obtained with 1% TFA while ethanol gave the highest

% recovery of TDG and TEA. This is in contrast with the results from the evaluation

of elution solvents for each MIP where deionized water samples without PEG were

used. Those results showed that the highest % recovery for 3Q was obtained with 1%

TFA while 1% TE gave the highest recovery for PMPA, TDG and TEA. The presence

of the other analytes, presence of PEG and the use of tap water may have contributed

to this observation. In terms of the biggest ratio of % recovery between the MIP and

the NIP, 1% TFA was the solvent of choice for 3Q, 1% TE for PMPA and ethanol for

TDG and TEA. The % absorptivity and recovery were generally lower for the NIP as

compared to that of the mixture of MIPs.

4.1.15 Comparison of MIP-SPE using a mixture of MIPs with other sample

preparation techniques

The % recovery of the analytes using the mixture of MIPs with the different

elution solvents as well as the different sample preparation procedures are compared

0

20

40

60

80

100

3Q PMPA TDG TEA

%


80

in Figure 4-30. SAX SPE is the only method capable of removing PEG from the

aqueous samples. However, it has its limitations. It is unable to recover the entire

range of analytes as only PMPA was detected. SCX SPE was unable to remove PEG

from the aqueous samples and in addition; it was only able to recover PMPA and

TDG. Except for TDG using TFA as the elution solvent, the MIP-SPE procedure was

able to recover the entire range of analytes. As expected, all the analytes were

detected using the direct rotary evaporation procedure. The total ion chromatograms

of the various procedures are compiled in Appendix 2. The chromatograms were

obtained by reconstitution with DCM instead of ACN, as ACN gave split peaks with

the HP-5MS column whereas DCM did not.

Figure 4-30. Comparison of % recovery using the various procedures.

Both the MIP-SPE and direct rotary evaporation procedures have their

advantages and disadvantages. The direct rotary evaporation procedure is fast and

gives a higher recovery of analytes. However, the procedure was found to introduce

more PEG into the GC column as seen from the great difference in areas under the

PEG peaks when the chromatograms are overlaid. This is important as it is not

0

20

40

60

80

100

3Q PMPA TDG TEA

%

MIP-SPE (EtOH) MIP-SPE (TFA) MIP-SPE (TE)Direct SCX SAX

81

advisable to introduce high concentrations of matrix interference into the GC MS

system as this may not only result in contamination of the column and MS detector

but also in shortening the lifespan of the MSD filament. On the other hand, the MIP-

SPE procedure gives lower recovery, and introduces less PEG into the GC column.

4.1.16 Preparation of sol-gel MIP fibers

Next, the development of sol-gel MIP fibers was attempted. Upon coating the

sol solution onto the fused silica fiber, the coating was dried at room temperature in a

desiccator. The coating solidified to give a shiny crystalline layer, however, it was of

unequal thickness along the length of the fiber. Prior to use, the fiber was conditioned

at 250 C for 1 h in the GC inlet. The heat treatment caused some of the coating to

flake off. Preliminary experiments with the fiber for the extraction of analytes of

interest from a water matrix and subsequent desorption in the heated inlet of the GC

caused further flaking off of the coating. As such, further investigation of sol-gel MIP

SPME was not carried out and efforts were instead focused on the evaluation of the

novel PhPPP coating.

4.1.17 CONCLUSION

The MIPs as well as the NIP were evaluated for their binding properties for a

range of analytes in deionized water samples as well as tap water samples containing

PEG as a background contaminant. The MIPs were not only able to bind to their

respective template but also to structurally similar compounds. This is an advantage

for the analysis of a wide range of similar analytes. The NIPs used in this study

showed some adsorption of the analytes. Ideally, they should not show adsorption of

analytes. However, the absorptivity and recovery were generally lower than that of the

MIPs. Only the PMPA-MIP showed 100% absorptivity of the analytes.

82

Each MIP-SPE procedure was compared with other sample preparation

procedures, namely SAX SPE and SCX SPE as well as a direct rotary evaporation

procedure for the analysis of a range of analytes in an aqueous sample containing

PEG. All the procedures were able to recover the phosphonates. However, only the

PMPA-MIP-SPE and SAX SPE procedures were capable of removing PEG from the

sample. Except for PMPA-MIP-SPE, a problem encountered for the other MIP-SPE

procedures was that analytes were eluted during the washing step. Hence to ensure

recovery of the analytes, the washing step was omitted and this compromised the

capability for sample clean-up. All the procedures except for SAX SPE were able to

recover TDG while only the MIP-SPE and the direct rotary evaporation procedures

were able to recover TEA and 3Q.

The various MIPs were mixed together for the analysis of a range of analytes,

namely the templates used during the polymer synthesis. The MIP-SPE procedures

using ethanol or 1% TE in water as the elution solvent, as well as the direct rotary

evaporation procedure, were able to recover the entire range of analytes. This is in

contrast to the procedures using commercially available SPE cartridges which did not

work for all of the analytes investigated. Although the direct rotary evaporation

procedure gave higher recovery, it was observed that the MIP-SPE procedure

introduces a lower amount of PEG into the GC column.

The development of sol-gel MIP SPME fibers posed a problem as the sol-gel

coatings synthesized using alkoxysilane monomers cracked and flaked off upon

drying at room temperature. Flaking off of the SPME coating was also observed upon

exposure to the heated GC inlet.

83

4.2 SPME using PhPPP-coated fibers

L1 is an organoarsenic agent listed under Schedule 1 of the Chemicals

Weapons Convention. The synthesized agent is actually composed of cis and trans

isomers and contains impurities such as L2 and L3. The structures of the compounds

are shown in Figure 4-31. Similar to sulfur mustard, L1 possesses vesicant properties.

However, in contrast to sulfur mustard, it is known to produce an immediate burning

sensation upon contact with the skin. Several arsenic-containing compounds were

introduced as chemical warfare agents during World War I. Among arsine,

diphenylchloroarsine, phenyldichloroarsine, diphenylaminechloroarsine,

diphenylcyanoarsine, methyldichloroarsine, ethyldichloroarsine and

ethyldibromoarsine, Lewisite was considered as the best arsenical war gas [354].

Unlike other chemical warfare agents, L1 requires derivatization in order to be

amenable to GC MS analysis as direct analysis leads to deterioration of the column

performance and corrosion of the detector [58].

Figure 4-31. Structures of the Lewisites.

Upon contact with water, L1 is rapidly hydrolyzed to 2-chlorovinylarsonous

acid (CVAA) which is as toxic. Subsequently, 2-chlorovinylarsenous oxide (CVAO)

and polymerized chlorovinylarsenous oxide is formed. The hydrolysis pathway is

shown in Figure 4-32. Similarly, L2 is expected to undergo similar hydrolysis to the

corresponding bis(2-chlorovinyl)arsonous acid (BCVAA) [356]. Except in old

abandoned chemical bombs where intact Lewisites 1 and 2 could still be found [357],

AsCl

Cl

ClAs

Cl

Cl ClAs

Cl Cl

Cl

L1 L2 L3

84

the existence of Lewisites in the environment would most likely be indicated by the

presence of CVAA, BCVAA, L3 or other degradation products. The SPME of CVAA

from aqueous and soil samples [89] as well as in urine [92] has previously been

investigated. In these studies, dithiols such as 1,2-ethanedithiol and 1,3-propanedithiol

were used as the derivatizing agents. In this study, we explored the use of the novel

PhPPP fiber for the extraction of the arsenic compounds as well as using monothiols

as derivatizing agents in SPME.

Figure 4-32. Hydrolysis pathway of L1 [355].

In the evaluation of the PhPPP-coated SPME fiber for the analysis of

Lewisites, extraction conditions such as sample pH, ionic strength, derivatizing agent

and extraction time were first optimized. Using the optimal extraction conditions, the

precision, linearity and limit of detection of the method were investigated using

spiked deionized water samples. The performance of the network fiber was compared

Cl OHAs

OH

AsCl

Cl

Cl

ClAs

O

ClAs

O

+ 2 H2O + 2 HCl

CVAA

CVAO

n

L1

+ H2O

85

0

20

40

60

80

0 2 4 6 8

pH

Up

take

(n

g)

L3

L2-PT

L1-PT

against that of a series of commercially available fibers, namely 30 m and 100 m

PDMS, 85 m PA, 65 m PDMS/DVB, 75 m CAR/PDMS and 65 m CW/DVB.

All experiments were performed in triplicate, unless stated otherwise.

4.2.1 Optimization of SPME conditions

Recommended operating procedures for the extraction of Lewisite in aqueous

samples involve the adjustment of the pH of the sample to 2 [62]. Hence, the effect of

sample pH on uptake by the fiber was investigated. The water sample was adjusted to

an acidic pH of 0 and 2, left unadjusted at pH 6 and adjusted to a basic pH of 8.

Figure 4-33 shows the uptake of the analytes with respect to sample pH. It was

found that adjustment of pH to 2 or analysis without pH adjustment did not

significantly affect the uptake. On the other hand, an extremely acidic pH of 0 and a

moderately basic pH of 8 adversely affected the uptake of the analytes. The behavior

of the analytes at extreme acidity is not well-understood and the lower uptake may be

due to the degree of ionization of the analytes while the poor uptake at pH 8 is the

result of decomposition in alkaline media [354]. Hence, in subsequent experiments,

the pH of the sample was not adjusted.

Figure 4-33. Effect of pH on uptake of the Lewisites. Spiking concentration: 0.5 µg ml 1; derivatizing agent: PT; salt concentration: 40% (w/v) NaCl; extraction time: 15 min.

86

Increasing the ionic strength of the aqueous sample by the addition of salt has

been shown to improve the SPME uptake of nerve agents [88]. However, the reverse

is observed for the Lewisites. As seen from Figure 4-34, saturating the sample with

the addition of 40 % (w/v) NaCl or Na2SO4 led to a decrease in uptake of the analytes.

Since the derivatized analytes were relatively non-polar as compared to the nerve

agents, salting was not required for uptake onto the fiber. Hence, further experiments

were conducted without the addition of salt. Omitting the need for adjustment of

sample pH and addition of salt resulted in a simple and straightforward extraction

procedure.

Figure 4-34. Effect of salting on uptake of the Lewisites. Spiking concentration: 0.5 µg ml 1; pH: 6; derivatizing agent: PT; extraction time: 15 min.

L1 can be chromatographed on a new column but leads to rapid deterioration

of the column. L2 can be chromatographed but is better derivatized. L3, on the other

hand, does not require derivatization [358]. The dithiols form cyclic derivatives with

L1 while for L2, a free thiol group remains. In other studies on CVAA, 1,2-

ethanedithiol and 1,3-propanedithiol were commonly used as derivatizing agents

[89,91,92,359]. An interesting fact to note is that derivatization of L1 or CVAA with

any particular thiol leads to the same derivative. The same is true for L2 and BCVAA.

0

40

80

120

L3 L2-PT L1-PT

Up

take

(n

g)

0%

40% Sodium sulfate

40% Sodium chloride

87

0

50

100

150

L3 L2 L1

Up

take

(n

g)

Ethanethiol Propanethiol ButanethiolEthanedithiol Propanedithiol

A series of monothiols and dithiols as derivatizing agents for the analytes was

investigated. The monothiols investigated included ethanethiol, propanethiol and

butanethiol while the dithiols were 1,2-ethanedithiol and 1,3-propanedithiol. The

structures of the derivatives are presented in Figure 4-35. The results are shown in

Figure 4-36. As 1,4-Butanedithiol was not fully soluble in water, it was excluded from

further investigations.

Figure 4-35. Structures of the various derivatives of L1 and L2.

Figure 4-36. Effect of derivatizing agents on uptake of the Lewisites. Spiking concentration: 0.5 µg ml 1; pH: 6; salt concentration: 0%; extraction time: 15 min.

Cl

S

AsS Cl

S

SAs

Cl

S

SAs

Cl

S

AsClCl

S

AsClCl

S

AsCl

Cl

S

AsCl

SH

Cl

S

AsCl

SH

S

S

AsCl

AsS

S

Cl

L1-ET L1-PT L1-BT

L1-EDT L1-PDT

L2-ET L2-PT L2-BT

L2-EDT L2-PDT

88

Since L3 does not require derivatization, its uptake did not vary significantly

with the derivatizing agent used while that of L2 and L1 showed significant variation.

It was obvious that the monothiols were superior over the dithiols as derivatizing

agents for L2 and L1. As our findings indicated that propanethiol gave the highest

uptake of L2 and L1, propanethiol was thus selected as the derivatizing agent for the

analytes in water samples in subsequent experiments.

Extraction time profiles (Figure 4-37) were obtained at extraction times

ranging from 5 min to 60 min. It was observed that equilibrium was achieved for the

analytes at 30 min. Hence, subsequent experiments were conducted using an

extraction time of 30 min.

Figure 4-37. Effect of extraction time on uptake of the analytes. Spiking concentration: 0.5 µg ml 1; pH: 6; derivatizing agent: PT; salt concentration: 0%; extraction time: 15 min.

4.2.2 Method validation

Using the optimal extraction conditions, the precision, linearity and limit of

detection of the method were investigated using spiked deionized water samples. The

results are compiled in Table 4-1.

0

50

100

150

0 15 30 45 60

Extraction Time (min)

Up

take

(n

g)

L3

L2-PT

L1-PT

89

0

5

10

15

20

25

L3 L2-PT L1-PT

Up

take

(n

g)

PhPPP fibre

30 um PDMS

100 um PDMS

PDMS/DVB

CAR/PDMS

PA

CW/DVB

Table 4-1. Quantitative results of SPME of the Lewisites.

Analyte Equationa

r2

% RSD (n = 6)

LODb

(µg l 1)

LODc

(µg l 1)

L3 y = 0.2799x + 2.1886 0.9975

8 0.31 0.17

L2-PT y = 0.4502x

0.5715 0.9997

6 0.19 0.24

L1-PT y = 0.5856x + 3.8997 0.9901

8 0.08 0.10

a linear range 0.001-0.5 µg ml 1

b LODs of SPME using PhPPP fiber at S/N = 3. c LODs of SPME using 100 m PDMS fiber at S/N = 3.

The linearity of SPME calibration plots was investigated over a concentration

range of 0.001-0.5 g ml 1. The analytes exhibited good linearity with good squared

regression coefficients of >0.990. This allowed the quantification of the compounds

by the method of external calibration. The relative standard deviations (RSDs) for six

extractions were below 8% for all the analytes. The limits of detection (LODs) for the

analytes were estimated at S/N = 3 under GC MS full scan conditions. The LODs

ranged between 0.08-0.31 g l 1.

4.2.3 Comparison with commercial SPME fibers

Figure 4-38. Comparison of uptake using the various fibers. Spiking concentration: 10 g l 1; pH: 6; derivatizing agent: PT; salt concentration: 0%; extraction time: 30 min.

90

The performance of the novel PhPPP fiber in the uptake of the analytes in

aqueous samples was compared against that of several commercially available fibers

from Supelco. As seen in Figure 4-38, the CAR/PDMS fiber gave the lowest uptake

of the analytes since it is more suitable for gases and low molecular weight

compounds up to a molecular weight of 225. Furthermore, it is not surprising that the

CW/DVB and PA fibers, suitable for polar compounds, showed poor uptake of the

analytes since they are considered to be relatively non-polar. Our results showed that

the PhPPP fiber performed better than all the commercial fibers in the uptake of

derivatives of CVAA and BCVAA but marginally poorer than the 100 m PDMS and

PDMS/DVB fibers in the uptake of L3. The LODs for the analytes using the PhPPP

fiber and the best-performing fiber, the 100 m PDMS fiber, are compared in Table

4-1. The use of propanethiol instead of propanedithiol as derivatizing agent for

CVAA lowered the LOD of 1 g l 1 obtained in a previous study [89] by an order of

magnitude. The lowest LOD reported for the analysis of CVAA is 7 pg ml 1 in urine

using an automated SPME GC MS system [92].

4.2.4 Conclusion

The novel PhPPP fiber shows great promise as an alternative to commercially

available fibers for the SPME of Lewisites in aqueous samples. PhPPP fibers are easy

to prepare and are significantly less costly than commercial fibers.

91

4.3 HF-LPME

4.3.1 Optimization of parameters for HF-LPME of chemical agents

The chemical warfare agents investigated included the nerve agents, GB, GD,

GA and VX as well as the blister agent, HD. The structures of the analytes are

presented in Figure 4-39. Factors affecting the extraction efficiency such as stirring

speed, sample ionic strength, and extraction time were optimized. All experiments

were performed in triplicate.

Figure 4-39. The chemical warfare agents investigated in this study.

Several commonly-used organic solvents in microextraction such as toluene,

cyclohexane, isooctane and their combinations were initially investigated. However,

higher extraction efficiency was observed with chlorinated solvents and their

combinations [344,345]. Hence, dichloromethane, chloroform, carbon tetrachloride,

trichloroethylene and their combinations were studied. For each solvent, extractions

were performed using deionized water spiked with CWAs, 30% (w/v) NaCl and with

stirring at 700 rpm for 15 min. Figure 4-40 shows the effect of the different solvents

on extraction efficiency.

P

O

OF

P

O

OF

NP

O

O

N

ClS

ClP

O

O

SN

GB GD GA

HD VX

92

Figure 4-40. Effect of extraction solvent on uptake of the CWAs. Salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

Dichloromethane, when used as a single solvent, was not suitable as it

evaporated too easily during the course of the extraction. As seen from Figure 4-40,

chloroform gave the highest uptake for GB and VX and relatively similar uptake for

GD, GA and HD as compared to the other solvents. In other preliminary studies, the

uptake was found to decrease when chloroform was used in combination with other

solvents. There is no observed relationship between the uptake of the CWAs as

related to the polarity indices [360] or octanol/water partition coefficients (log Kow)

[361] of the solvents and the log Kow of the CWAs [15]. The polarity indices of the

solvents as well as the log Kow of the CWAs are compiled in Tables 4-2 and 4-3.

Chloroform, the most polar solvent among the chlorinated solvents investigated, was

able to extract the two analytes with extreme log Kow values, GB and VX. On the

other hand, trichloroethylene, the least polar solvent, showed poor uptake for most of

the analytes. The results are in contrast to those reported previously [345] where

chloroform, among other solvents, was discarded due to problems with non-

0

10

20

30

40

50

GB GD GA HD VX

ChloroformCarbon tetrachlorideDichloromethane-Carbon tetrachloride 1:1Dichloromethane-Carbon tetrachloride 3:1TrichloroethyleneToluene

Up

take

(n

g)

93

compatibility with fiber pores, instability and miscibility with water. Our results show

that the extraction efficiency of the CWAs with chloroform is significantly better than

that with toluene and with trichloroethylene. Several of the solvents were unable to

extract VX. Hence, chloroform was selected as the extraction solvent.

Table 4-2. Data of the solvents. Table 4-3. Data of the CWAs.

Extractions without the addition of salt were performed. However, uptake of

the analytes was poor, particularly for VX which was not recovered. Hence

experiments were performed with addition of salt. Increasing the ionic strength of the

water sample has been shown to decrease the affinities of the chemical agents for the

aqueous matrix and hence enhancing their uptake [88]. The effect of two different

salts, NaCl and Na2SO4, was compared in Figure 4-41. At the same salinity level of

30% (w/v), neither NaCl nor Na2SO4 was the obvious choice. The uptake of VX was

better whereas the uptake of HD was adversely affected with Na2SO4. However, since

NaCl gave higher uptake of most the analytes, further experiments were conducted

with NaCl. Salt concentrations were varied from a low salinity level of 10% (w/v) to a

saturation level of 40% (w/v).

Solvent Polarity Index [360] Chloroform 4.1 Dichloromethane 3.1 Toluene 2.4 Carbon tetrachloride 1.6 Trichloroethylene 1.0

CWA Log Kow

[361]

GB 0.299 GD 1.824 GA 0.384 HD 1.37 VX 2.09

94

0

10

20

30

40

10 20 30 40

Salt concentration (w/v % )

Up

take

(n

g) GB

GDGAHDVX

Figure 4-41. Effect of salt on uptake of the CWAs. Extraction solvent: chloroform; stirring speed: 700 rpm; extraction time: 15 min.

Figure 4-42. Effect of salt concentration on uptake of the CWAs. Extraction solvent: chloroform; stirring speed: 700 rpm; extraction time: 15 min.

Figure 4-42 shows the effect of salt concentration on extraction efficiency. It

was observed that the optimum salt concentration for all the analytes was 30% (w/v).

At higher salinity levels of 33% or higher, the sample solution was saturated such that

the presence of suspended salt particles interfered with the uptake and led to the

0

10

20

30

40

50

GB GD GA HD VX

Up

take

(n

g)

Sodium chloride

Sodium sulfate

95

0

20

40

60

80

300 500 700 900 1100

Stirring Speed (rpm)

Up

take

(n

g)

GBGDGAHDVX

decrease in recoveries observed. The salt concentration was thus maintained at 30%

(w/v) NaCl for subsequent experiments.

Although agitation of the sample can enhance the extraction and reduce the

time to thermodynamic equilibrium, higher agitation speed may result in drop

dislodgement and solvent loss particularly for SDME, over long extraction times. As

mentioned above, an obvious advantage of HF-LPME over SDME is the fact that

higher stirring speeds can be employed without affecting the solvent drop. The effect

of stirring speed on the extraction efficiency was investigated over a range of 300 rpm

to the maximum of 1250 rpm. Figure 4-43 shows the effect of stirring speed on

extraction efficiency. The uptake of VX was relatively unaffected by stirring speed or

extraction time due to its slow rate of diffusion [88], the extraction efficiencies of the

other analytes increased with increasing stirring speeds. Hence the maximum

allowable stirring speed on the magnetic stirrer, i.e. 1250 rpm, was used in subsequent

experiments.

Figure 4-43. Effect of stirring speed on uptake of the CWAs. Extraction solvent: chloroform; salt concentration: 30% (w/v); extraction time: 15 min.

96

0

25

50

75

100

125

15 30 45 60

Extraction Time (min)

Up

take

(n

g) GB

GD

GA

HD

VX

HF-LPME is an equilibrium-based rather than exhaustive extraction process.

Generally, the equilibrium time is selected as extraction time. However, it is usually

not practical to prolong an extraction unnecessarily for equilibrium to be established.

This is because the longer the extraction, the greater the potential of solvent loss due

to dissolution in the sample solution. Additionally, there are obvious benefits in

conducting more time-efficient extraction. Thus, for quantitative analysis, achieving

equilibrium is not necessary as long as extraction conditions are kept rigorously

constant. Figure 4-44 shows the effect of extraction time on extraction efficiency.

Except for VX which was relatively unaffected by extraction time, the other analytes

showed increasing uptake with increasing extraction times. The extraction time of 30

min was selected since reasonable uptake was achieved in this shorter analysis time.

Figure 4-44. Effect of extraction time on uptake of the CWAs. Extraction solvent: chloroform; salt concentration: 30% (w/v); stirring speed: 1250 rpm.

4.3.2 Method validation for HF-LPME of chemical agents

Using the optimal extraction conditions, the precision, linearity and limit of

detection of the method were investigated using spiked deionized water samples. The

results are shown in Table 4-4. External calibration was performed for all

97

experiments. To evaluate the linearity of the calibration plots, samples were spiked

with CWA over a concentration range of 0.01 1 g ml 1 and then extracted. The GC

peak area counts were plotted against the respective analyte concentrations to

generate calibration curves. The CWAs exhibited good linearity with very good

squared regression coefficients of >0.9950. This allowed the quantification of the

compounds by the method of external calibration. The limits of detection for the

CWAs were estimated at S/N = 3 under GC MS full scan conditions. The LODs

ranged between 0.02 and 0.09 g l 1 and were better than those reported for SDME

and HF-LPME using trichloroethylene [344,345]. Also, these values better the

requirement for the safe level (50-200 g l 1 in 2-3 L) in drinking water contaminated

with CWAs [88]. The RSDs for six extractions were below 10% for all the analytes.

Table 4-4. Quantitative results of HF-LPME of the CWAs.

Analyte

Equationa

r2

% RSD

(n = 6) LODb

LODc

LODd

LODe

LODf

GB

y = 0.1164x + 1.0640

0.9950

5

0.03

0.03

10

75

0.05

GD

y = 0.1840x + 0.0447

0.9999

7

0.09

0.10

-

-

0.05

GA

y = 0.1802x + 0.2885

1.0000

6

0.02

0.02

-

-

0.05

HD

y = 0.1698x + 0.3817

0.9999

9

0.09

0.10

1.0

30

-

VX

y = 0.0375x + 1.9906

0.9977

7

0.09

0.20

-

-

0.5

a linear range 0.01-1 µg ml 1

b LODs (in g l 1) of HF-LPME for water samples at S/N = 3. c LODs (in g l 1) of HF-LPME for slurry samples at S/N = 3. d LODs (in g l 1) of HF-LPME using trichloroethylene as a solvent at S/N = 10 [345]. e LODs (in g l 1) of SDME using dichloromethane:carbon tetrachloride as a solvent at S/N = 10 [344]. f LODs (in g l 1) of SPME using 65 m PDMS/DVB fiber at S/N =3 [88].

4.3.3 Comparison of HF-LPME of chemical agents with SPME

In order to demonstrate the capability of HF-LPME in the extraction of the

CWAs, a relatively simple matrix such as deionized water samples as well as the

much more complex slurry samples (20 mg of soil in 1 ml of water) were employed.

98

Figure 4-45 shows typical total ion chromatograms after HF-LPME extraction of the

CWAs. As compared to the deionized water samples, the slurry samples were meant

to pose a greater challenge to the extraction procedure due to the presence of soil

particles as well as extraneous materials.

Figure 4-45. Total ion chromatograms after HF-LPME extraction of spiked deionized water sample (a) and spiked slurry sample (b) at a concentration of 0.5 µg ml 1. Extraction solvent: chloroform; salt concentration: 30% (w/v); stirring speed: 1250 rpm; extraction time: 30 min.

The extraction efficiency of HF-LPME was also compared with that of direct

immersion-SPME. Previously-reported optimum conditions for direct immersion

SPME were utilized [88]. As seen from Figure 4-46, neither technique demonstrated

6.00 7.00 8.00 9.0010.0011.0012.0013.0014.0015.0016.0017.0018.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

Time-->

Abundance

TIC: WCF2.D

VX

HD

GA

GD

GB

Time (min)

6.00 7.00 8.00 9.0010.0011.0012.0013.0014.0015.0016.0017.0018.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

Time-->

Abundance

TIC: SCF2.D

VX

HDG

A

GD

GB

Time (min)

99

superiority over the other. The HF-LPME technique gave higher uptake for GB, GA

and VX while direct immersion SPME showed higher uptake of GD and HD. The

PDMS/DVB fiber showed higher uptake of GD and HD, over the other CWAs, owing

to their relative hydrophobicity. Except for VX, the uptake of the analytes using HF-

LPME was not affected by the more complex slurry matrix. On the other hand, a

difference between the uptake of the analytes by direct immersion SPME from water

and slurry samples is observed. In fact, VX was not recovered at all from slurry

samples by direct immersion-SPME. This experiment demonstrated an obvious

advantage of HF-LPME over direct immersion SPME where the hollow fiber served

as a filter that prevented the soil particles in the complex slurry matrix from

interfering with the analysis. After each extraction, the hollow fiber can be discarded

and a fresh one used for the next extraction. In contrast, the fiber was observed to

deteriorate with repeated direct immersion SPME from the slurry samples.

Figure 4-46. Comparison of HF-LPME and SPME extraction efficiency for the CWAs. HF-LPME extraction solvent: chloroform; salt concentration: 30% (w/v); stirring speed: 1250 rpm; extraction time: 30 min. SPME salt concentration: 40% (w/v); stirring speed: 1250 rpm; extraction time: 30 min.

0

50

100

150

200

GB GD GA HD VX

Up

take

(n

g)

HF-LPME (water)

HF-LPME (slurry)

SPME (water)

SPME (slurry)

100

4.3.4 Optimization of parameters for HF-LPME of CWA degradation products

Several main degradation products of CWAs were selected, ranging from

those of nerve agents such as GB, GD, VX to blister agents, including HD, 1,2-bis(2-

chloroethylthio)ethane (sesquimustard, Q), bis(2-chloroethylthioethyl)ether (O-

mustard, T), and a psychotomimetic agent, BZ. The actual analytes of interest were

EMPA (from VX), IMPA (from GB), PMPA (from GD), MPA which is the final

degradation product of alkyl methylphosphonic acids, n-propylphosphonic acid

(nPPA), ethyl 2-hydroxyethyl sulfide (EHES), thiodiglycol (TDG) (from HD), 1,2-

bis(2-hydroxyethylthio)ethane (QOH) (from Q), bis(2-hydroxyethylthioethyl)ether

(TOH) (from T), and benzilic acid (BA) (from BZ). The structures of the analytes are

shown in Figure 4-47 while the pKa and log Kow data of the analytes are compiled in

Table 4-5. Factors affecting the extraction efficiency such as extraction solvent, pH,

sample ionic strength, stirring speed, and extraction time were optimized. All

experiments were performed in triplicate.

Figure 4-47. CWA degradation products investigated in this study.

P

O

OHOH

P

O

OHO

P

O

OHO

P

O

OOH

SOHOH

OS

OHS

OH

SOHS

OH

P

O

OHOH

SOH

OH

OHO

MPA

EMPA IMPA PMPA

TDG QOH

TOH

nPPA

EHES

BA

101

Table 4-5. pKa and log Kow data of the CWA degradation products.

Analyte pKa [362,363]

Log Kow

[362]

EMPA 2.32 -0.15 EHES 14.73 0.44 IMPA 2.32 0.27 PMPA 2.30 Not available MPA 2.12a

-0.70 nPPA 3.13 0.28 TDG 14.68 -0.63 BA 3.05a

2.30 QOH 14.64 Not available TOH Not available Not available

a Obtained from the Interactive PhysProp Database [362].

HF-LPME for analytes requiring derivatization prior to GC MS analysis has

been carried out by extracting the analytes of interest first using pure solvent before

drawing up derivatizing agent into the hollow fiber prior to injection [364], or

introducing derivatizing agent into the GC injection port immediately after injection

of the extract [365]. Alternatively, the derivatizing agent is added to the sample for

simultaneous derivatization and extraction of analytes [366,367]. Coupled two-step

derivatizations in which hydroxycarbonyls were first extracted and derivatized with

O-(2,3,4,5-pentafluorobenzyl)-hydroxylamine by HF-LPME followed by single drop

microextraction over the headspace of bis(trimethylsilyl)trifluoroacetamide (BSTFA)

have been reported [368]. Another successful approach for derivatization was to use a

mixture of solvent and derivatizing agent as the extraction medium held within a HF

for the simultaneous clean-up, extraction and derivatization of 11-nor- 9-

tetrahydrocannabinol-9-carboxylic acid from urine [369]. The solvent mixture

consisted of BSTFA and octane in the ratio of 5:1. After an 8-min extraction, 6 l of

the mixture was analyzed by GC MS. Similarly, a HF-LPME technique involving

simultaneous in-fiber silylation for unconjugated anabolic steroids in urine has been

developed [370]. The hollow fiber was first pre-conditioned with dihexyl ether and

102

then filled with a mixture of N-methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA),

ammonium iodide and dithioerythritol as the acceptor phase. The extraction was

performed at 45 C for 30 min. A 2- l portion of the acceptor phase was then drawn

from the fiber and injected directly into the GC MS instrument.

In-situ derivatization HF-LPME of alkylphosphonic acids has been carried

out by basification of the spiked water with potassium carbonate, addition of propyl

bromide and stirring of the mixture at 100 C for 2 h. Upon completion of reaction,

extractions were performed by HF-LPME prior to GC MS analysis [346]. Here, HF-

LPME with in-situ derivatization was investigated for the extraction of degradation

products of CWAs from aqueous samples in which extractions were performed using

a mixture of solvent and derivatizing agent held in the hollow fiber. Although the

derivatizing agent used, N-(tert.-butyldimethylsilyl)-N-methyltrifluoroacetamide

(MTBSTFA), is moisture-sensitive, it is protected within the hollow fiber and allowed

the successful extraction of the analytes of interest prior to GC MS analysis. In

addition, tert.-butyldimethylsilyl (TBDMS) derivatives are known to be 104 times

more stable to hydrolysis than trimethylsilyl (TMS) derivatives [371].

In order to determine an optimum derivatization method, three derivatization

procedures were compared. In the first method, pure solvent (chloroform) was held in

the hollow fiber for the extraction of the analytes. Extractions were performed using

deionized water spiked with the analytes, 30% (w/v) NaCl and with stirring at 700

rpm. After 15 min of extraction, 1 l of MTBSTFA was combined with 1 l of extract

in the syringe for injection. In the second method, a mixture of chloroform and

MTBSTFA in a 1:1 ratio was held in the hollow fiber for extraction, followed by

injection of 1 l of the extract. The third method was essentially identical to the

second except that the 1 l of extract was combined with another 1 l of MTBSTFA

103

prior to injection. It was concluded from our experiments that the second method

using the chloroform/MTBSTFA solvent-derivatizing agent mixture (1:1) without the

additional microliter of derivatizing agent was optimum for extractions. The effect of

the derivatization procedure on the uptake of the analytes is shown in Figure 4-48.

Figure 4-48. Effect of derivatization method on uptake of the degradation products. Spiking concentration: 2 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 1 µg ml 1

for EHES and PMPA and 20 µg ml 1 for MPA, nPPA and TDG; extraction solvent: solvent/MTBSTFA (1:1); pH: 1.5; salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

The experiments on HF-LPME of CWAs, as discussed in Section 4.3.1,

demonstrated that chlorinated solvents were ideal for this class of compounds. Hence,

several chlorinated solvents, namely chloroform, carbon tetrachloride,

trichloroethylene and tetrachloroethylene, were evaluated together with a commonly-

used non-chlorinated solvent, toluene. Each of the solvents was mixed in a 1:1 ratio

with MTBSTFA and 5 l of the solvent-derivatizing agent mixture was held in the

hollow fiber for extraction. From Figure 4-49, it can be seen that none of the solvents

gave the highest extraction for all the analytes. Chloroform was selected for further

experiments as it gave the optimum extraction for most of the analytes.

0

50

100

150

200

EMPA EHES IMPA PMPA MPA nPPA TDG BA QOH TOH

Up

take

(n

g)

Chloroform:MTBSTFA 1:1

Chloroform:MTBSTFA 1:1 + MTBSTFA

Chloroform + MTBSTFA

104

0

300

600

900

1200


Up

take

(n

g)

MTBSTFA

BSTFA

Figure 4-49. Effect of extraction solvent on uptake of the degradation products. Spiking concentration: 2 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 1 µg ml 1

for EHES and PMPA and 20 µg ml 1 for MPA, nPPA and TDG; extraction solvent: solvent/MTBSTFA (1:1); pH: 1.5; salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

Next, two commonly-used silylating derivatizing agents, BSTFA and

MTBSTFA, were compared in order to determine the derivatizing agent of choice.

The results are presented in Figure 4-50. Except for BA where BSTFA gave higher

uptake than MTBSTFA, MTBSTFA is the derivatizing agent of choice for the rest of

the analytes. Subsequent experiments were carried out using MTBSTFA as the

derivatizing agent.

Figure 4-50. Effect of derivatizing agent on uptake of the degradation products. Spiking concentration: 2 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 1 µg ml 1

for EHES and PMPA and 20 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/derivatizing agent (1:1); pH: 1.5; salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

0

50

100

150


Up

take

(ng

)

ChloroformCarbon tetrachlorideTrichloroethyleneTetrachloroethyleneToluene

105

0

50

100

150

200

10 20 30 40 50 60

% MTBSTFA

Up

take

(n

g)

EMPA

EHES

IMPA

PMPA

MPA

nPPA

TDG

BA

QOH

TOH

The amount of MTBSTFA required for derivatization was investigated (Figure

4-51). The use of 100% MTBSTFA was not feasible as the liquid rapidly solidified

upon exposure to water. Solidification was also observed with an MTBSTFA content

of 65% and above. Hence, experiments were conducted using chloroform with an

MTBSTFA content of 10%, 40%, 50% and a maximum of 60% as extracting solvent.

It was found that 50% of MTBSTFA in chloroform, that is, chloroform/MTBSTFA

(1:1), gave optimal results and was hence used in subsequent experiments.

Figure 4-51. Effect of MTBSTFA content on uptake of the degradation products. Spiking concentration: 2 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 1 µg ml 1

for EHES and PMPA and 20 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/MTBSTFA; pH: 1.5; salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

As the analytes possess acidic protons, pH has a significant impact on the

degree of ionization and hence extraction. A comparison of the extraction of the

analytes at an extreme pH of 0, pH 1.5 and pH 4, where the sample pH was

unadjusted, was made. Figure 4-52 clearly shows that pH 0 was optimum for the

extraction of the analytes since they are expected to be fully unionized at this pH.

Hence the pH of the samples was adjusted to 0 in subsequent experiments.

106

0

50

100

150

200

0 2 4

pH

Up

take

(n

g)

EMPAEHESIMPAPMPAMPAnPPATDGBAQOHTOH

Figure 4-52. Effect of pH on uptake of the degradation products. Spiking concentration: 2 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 1 µg ml 1 for EHES and PMPA and 20 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/MTBSTFA (1:1); salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

Increasing the ionic strength of the water sample has been shown to decrease

the affinities of the CWAs for the aqueous matrix, thus enhancing their extraction by

the extractant solvent [88]. The effect of two different salts, NaCl and Na2SO4, was

compared. At the same salinity level of 30% (w/v), NaCl gave higher uptake of most

of the analytes as compared to Na2SO4 (Figure 4-53).

Figure 4-53. Effect of salt on the uptake of the degradation products. Spiking concentration: 0.7 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 0.3 µg ml 1 for EHES and PMPA and 7 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/derivatizing agent (1:1); pH: 1.5; salt concentration: 30% (w/v); stirring speed: 700 rpm; extraction time: 15 min.

0

10

20

30

40


Up

take

(n

g)

Sodium chloride

Sodium sulfate

107

0

20

40

60

20 30 40

% NaCl

Up

take

(ng

)


NaCl concentrations were then varied from a salinity level of 20% (w/v) to a

highly saturated level of 40% (w/v). PMPA and nPPA showed vastly opposite trends

with increasing ionic strength. This can be explained by the fact that nPPA is

relatively more polar than PMPA and its uptake is enhanced by increasing salt

concentration. From Figure 4-54, the optimum salt concentration was determined to

be 33% (w/v). The salt concentration was adjusted to 33% (w/v) NaCl for subsequent

experiments.

Figure 4-54. Effect of salt concentration on the uptake of the degradation products. Spiking concentration: 0.7 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 0.3 µg ml 1 for EHES and PMPA and 7 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/MTBSTFA (1:1); pH: 0; stirring speed: 700 rpm; extraction time: 15 min.

The effect of stirring speed on the extraction efficiency was investigated over

a range of 300 rpm to the maximum of 1250 rpm. As seen from Figure 4-55

extraction of several of the analytes were relatively unaffected by the stirring speed.

However, it was observed that the extraction of PMPA, IMPA and TOH increased

with increasing stirring speed up to 1000 rpm. Increasing the stirring speed led to

increased diffusion of these analytes, which possess branched or long alkyl chains.

108

0

25

50

75

300 500 700 900 1100

Stirring speed (rpm)

Up

take

(n

g)


Hence the stirring speed was set at 1000 rpm in subsequent experiments, since at this

value, most of the analytes were optimally extracted.

Figure 4-55. Effect of stirring on uptake of the degradation products. Spiking concentration: 0.7 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 0.3 µg ml 1 for EHES and PMPA and 7 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/MTBSTFA (1:1); pH: 0; salt concentration: 33% (w/v); extraction time: 15 min.

Figure 4-56. Effect of extraction time on uptake of the degradation products. Spiking concentration: 0.7 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 0.3 µg ml 1 for EHES and PMPA and 7 µg ml 1 for MPA, nPPA and TDG; extraction solvent: chloroform/MTBSTFA (1:1); pH: 0; salt concentration: 33% (w/v); stirring speed: 1000 rpm.

0

50

100

150

200

15 30 45 60

Extraction time (min)

Up

take

(n

g)


109

Figure 4-56 shows the time profile of extraction up to 60 min. Except for

EHES and BA which were relatively unaffected by extraction time, the uptake of the

other analytes increased with increasing extraction times up to 45 min. The extraction

time of 45 min was selected since it was the optimum extraction time for most of the

analytes.

4.3.5 Method validation for HF-LPME of CWA degradation products

Using the optimum extraction conditions, the precision, linearity and LODs of

the method were investigated using spiked water samples. The results are shown in

Table 4-6.

Table 4-6. Quantitative results of HF-LPME of the degradation products.

Analyte Equation r2

% RSD (n = 6)

LODa

(µg l 1) EMPA y = 0.1108x + 0.4105 0.9997 17 0.16 EHES y = 0.0436x + 5.4783 1.0000 9 0.08 IMPA y = 0.1751x + 1.7730 0.9977 12 0.03 PMPA y = 0.4045x + 2.7154 0.9989 13 0.05 MPA y = 0.0022x + 2.4999 0.9940 14 0.02 nPPA y = 0.0187x + 3.5046 0.9995 13 0.01 TDG y = 0.0079x

0.8692 0.9996 11 0.10 BA y = 0.0073x + 0.7016 0.9987 22 0.54 QOH y = 0.1434x

5.6745 0.9934 22 0.46 TOH y = 0.2744x

4.9917 0.9929 20 0.39 a at S/N = 5.

The linearity of HF-LPME calibration plots was investigated over a

concentration range of 0.005 5 g ml 1. The analytes exhibited good linearity with

squared regression coefficients of >0.993. This allowed the quantification of the

compounds by the method of external calibration. The LODs for the analytes were

estimated at S/N = 5 under GC MS full scan conditions by considering the lowest

concentration of each of the analytes that could be detected. The LODs ranged

110

between 0.01 and 0.54 g l 1. The relative standard deviations for six extractions were

between 9 and 22% for the analytes. Due to the need for derivatization, the % RSD

values tend to be larger in general than that of extractions without the need for this

additional step.

4.3.6 Comparison of HF-LPME of CWA degradation products with SPME

In order to compare the performance of HF-LPME with in-situ derivatization

with that of SPME, extractions of the degradation products from deionized water

samples were conducted under the optimized HF-LPME conditions and previously-

reported optimum conditions for SPME [90]. Several commercially available SPME

fibers were used, ranging from non-polar PDMS to polar PA fibers. Figure 4-57

shows the results obtained. It was obvious that HF-LPME with in-situ derivatization

surpassed that of SPME for all of the analytes except for BA. The CW/DVB fiber

gave the highest extraction for BA owing to the polarity as well as porosity of the

CW/DVB coating. Due to its hydrophilic nature as compared to the other analytes,

MPA posed a problem for both extraction techniques as seen from the low recoveries

of both procedures. Besides the simplicity and low cost, an added advantage of HF-

LPME over SPME is that after each extraction, the hollow fiber can be discarded and

a fresh one used for the next extraction. In contrast, the SPME fiber required

additional conditioning steps prior to and after extractions in order to prevent

carryover problems. The limited lifetime of an SPME fiber subjected to direct

immersion extraction is also an unfavorable consideration.

111

Figure 4-57. Comparison of HF-LPME and SPME of the degradation products. Spiking concentration: 1 µg ml 1 for EMPA, IMPA, BA, QOH and TOH, 0.5 µg ml 1

for EHES and PMPA and 10 µg ml 1 for MPA, nPPA and TDG; HF-LPME extraction solvent: chloroform/MTBSTFA (1:1); pH: 0; salt concentration: 33% (w/v); stirring speed: 1000 rpm; extraction time: 45 min. SPME pH: 1.5; salt concentration: 40% (w/v); stirring speed: 1000 rpm; extraction time: 30 min.

4.3.7 Optimization of parameters for HF-LPME of basic degradation products

Several basic degradation products of CWAs were selected, namely those of a

nerve agent (VX), the nitrogen mustards blister agents (HN1, HN2 and HN3) and a

psychotomimetic agent (BZ). The actual analytes of interest were 2-(N,N-

diisopropylamino)ethanol (DIPAE) (from VX), N-methyldiethanolamine (MDEA)

(from HN2), N-ethyldiethanolamine (EDEA) (from HN1), triethanolamine (TEA)

(from HN3) [15] and 3-quinuclidinol (3Q) (from BZ) [372]. The structures of the

analytes are shown in Figure 4-58. The negative logarithm of the acid dissociation

constants (pKa) and logarithms of the octanol-water coefficients (log Kow) of the

analytes are tabulated in Table 4-7.

Figure 4-58. Structures of the basic analytes investigated in this study.

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)HF-LPMEPDMSPDMS/DVBCAR/PDMSCW/DVBPA

NOH OH

N

OH

OH OHN

OH OH

N

OHNOH

DIPAE 3Q

MDEA EDEA TEA

112

Table 4-7. pKa and log Kow data of the basic analytes.

Analyte pKa

[362,363] Log Kow

[362]

DIPAE 10.1 0.88 3Q 9.16a

0.17

MDEA 8.52 -1.50 EDEA 8.75a

-1.01

TEA 7.76 -1.00 a Based on the SPARC On-Line Calculator [363]

Factors affecting the extraction efficiency such as choice of derivatizing agent,

extraction solvent for HF-LPME, pH, sample ionic strength, stirring speed, and

extraction time were optimized. All experiments were performed in triplicate.

In order to determine an optimum derivatization method for HF-LPME, three

derivatization procedures were compared together with the evaluation of MTBSTFA

and BSTFA as the more suitable derivatizing agent. In the first method, pure solvent

(chloroform) was held in the hollow fiber for the extraction of the analytes.

Extractions were performed using deionized water spiked with the analytes, 30%

(w/v) NaCl and with stirring at 1000 rpm. After 15 min of extraction, 1 l of

derivatizing agent was combined with 1 l of extract in the syringe for injection. In

the second method, a mixture of chloroform and derivatizing agent in a 1:1 ratio was

held in the hollow fiber for extraction, followed by injection of 1 l of the extract.

The third method was essentially identical to the second except that the 1 l of extract

was combined with another 1 l of derivatizing agent prior to injection. The results

are shown in Figure 4-59.

113

Figure 4-59. Effect of derivatization method on uptake of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/derivatizing agent (1:1); pH: 10; salt concentration: 30% (w/v) NaCl; stirring speed: 1000 rpm; extraction time: 15 min.

It is interesting to note that in the case of derivatization with MTBSTFA, 3Q

was largely underivatized, and only a small amount of the MTBSTFA derivative was

observed. This problem did not occur with BSTFA derivatization. This observation

can conceivably be attributed to the steric bulk of both 3Q and MTBSTFA that

prevents effective derivatization. Furthermore, the secondary alcohol group of 3Q

could have affected its reactivity towards MTBSTFA. Hence, quantification was

based on the underivatized 3Q instead of the MTBSTFA derivative for all subsequent

experiments that involved MTBSTFA.

The extraction with the chloroform:MTBSTFA mixture followed by another 1

l of MTBSTFA gave the highest uptake of 3Q even though, as mentioned above, it

remained largely underivatized in the presence of MTBSTFA. Separate experiments

were performed to investigate the recovery of 3Q using HF-LPME without

MTBSTFA which resulted in negligible recovery of 3Q. Hence it is speculated that

MTBSTFA probably aided in the uptake of 3Q similar to an ion-pairing mechanism

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Chloroform:MTBSTFA 1:1

Chloroform:MTBSTFA 1:1 + MTBSTFA

Chloroform + BSTFA

Chloroform:BSTFA 1:1

Chloroform:BSTFA 1:1 + BSTFA

114

but eventually failed to fully derivatize the compound. This phenomenon has not been

thoroughly investigated and hence is not yet fully understood. This could be the

subject of future studies. It was concluded from our experiments that the third method

using the chloroform/BSTFA solvent-derivatizing agent mixture (1:1) together with

the additional microliter of derivatizing agent was optimum for HF-LPME as it gave

the highest uptake for most of the analytes, especially TEA which could not be

extracted using the other methods. On the other hand, SPME using derivatization with

BSTFA gave negligible recoveries of the analytes. Hence, SPME experiments were

conducted using MTBSTFA as the derivatizing agent.

The experiments on HF-LPME of CWAs and degradation products, as

discussed in Sections 4.3.1 and 4.3.4, demonstrated that chlorinated solvents were

ideal for this class of compounds. Hence, several chlorinated solvents, namely

chloroform, carbon tetrachloride, trichloroethylene and tetrachloroethylene, were

evaluated together with a commonly-used non-chlorinated solvent, toluene. Each of

the solvents was mixed in a 1:1 ratio with BSTFA and 5 l of the solvent-derivatizing

agent mixture was held in the hollow fiber for extraction. As seen from Figure 4-

60(a), chloroform was the best extraction solvent and was selected for further

experiments. On the other hand, the uptake of the analytes using various fibers was

evaluated. Several fibers gave negligible uptake of the analytes. As seen from Figure

4-60(b), even though the PDMS fiber gave the highest uptake of DIPAE, the most

non-polar analyte, the CAR/PDMS gave the highest uptake for the other analytes and

was thus used in subsequent experiments.

115

Figure 4-60. (a) Effect of extraction solvent on uptake of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); pH: 10; salt concentration: 30% (w/v) NaCl; stirring speed: 1000 rpm; extraction time: 15 min. (b) Effect of SPME fiber on uptake of the basic analytes. Spiking concentration: 5 µg ml 1 for DIPAE and 50 µg ml 1 for 3Q, MDEA, EDEA and TEA; pH: 10; salt concentration: 30% (w/v) Na2SO4; stirring speed: 1000 rpm; extraction time: 15 min.

The amount of BSTFA required for derivatization in HF-LPME was

investigated through experiments using chloroform with a BSTFA content of 25%,

50% and 75% and pure BSTFA as extracting solvent. It was found that 50% of

BSTFA in chloroform, that is, chloroform/BSTFA (1:1), gave optimal results (Figure

0

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)

Chloroform

Carbon tetrachloride

Trichloroethylene

Tetrachloroethylene

Toluene

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3Q DIPAE MDEA EDEA TEA

Up

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(ng

)

PDMS

PDMS/DVB

CAR/PDMS

(b)

(a)

116

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% BSTFA

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(n

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DIPAE

3Q

MDEA

EDEA

TEA

4-61) and was hence used in subsequent experiments. The reason for the decreasing

recoveries at higher amounts of BSTFA could be attributed to the decreasing

solubility of the analytes with decreasing amounts of CHCl3 solvent. The use of

BSTFA means that the poor derivatization of 3Q by MTBSTFA as observed earlier

was no longer a concern in HF-LPME.

Figure 4-61. Effect of amount of derivatizing agent on uptake of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA; pH: 10; salt concentration: 30% (w/v) NaCl; stirring speed: 1000 rpm; extraction time: 15 min.

As they are basic compounds (Table 4-7), pH has a significant impact on the

degree of ionization and hence extraction of the analytes. A comparison of HF-LPME

of the analytes at an extreme pH of 14, pH 12, pH 10 and pH 7, where the sample pH

was unadjusted, was made. On the other hand, the extraction pH using SPME was

evaluated at pH 8, pH 9, pH 10 and pH 11, which was the maximum acceptable

working pH of the Carboxen/PDMS fiber. Figure 4-62 shows that pH 12 and pH 10

117

was optimum for HF-LPME and SPME respectively. Hence, the pH of the samples

was adjusted accordingly in subsequent experiments.

Figure 4-62. (a) Effect of pH on HF-LPME of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); salt concentration: 30% (w/v) NaCl; stirring speed: 1000 rpm; extraction time: 15 min. (b) Effect of pH on SPME of the basic analytes. Spiking concentration: 5 µg ml 1 for DIPAE and 50 µg ml 1 for 3Q, MDEA, EDEA and TEA; salt concentration: 30% (w/v) Na2SO4; stirring speed: 1000 rpm; extraction time: 15 min.

0

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7 8 9 10 11 12 13 14

pH

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(n

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DIPAE

3Q

MDEA

EDEA

TEA

(a)

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120

160

8 9 10 11

pH

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(n

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3Q

DIPAE

MDEA

EDEA

TEA

(b)

118

Increasing the ionic strength of the water sample has been shown to decrease

the affinities of analytes for the aqueous matrix and hence enhancing their extraction

by the extractant solvent [90]. The effect of two different salts, NaCl and Na2SO4, was

compared. At the same salinity level of 30% (w/v), Na2SO4 gave higher uptake of the

analytes as compared to NaCl for both HF-LPME and SPME (Figure 4-63).

Figure 4-63. (a) Effect of salt on HF-LPME of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); pH 12; salt concentration: 30% (w/v); stirring speed: 1000 rpm; extraction time: 15 min. (b) Effect of salt on SPME of the basic analytes. Spiking concentration: 5 µg ml 1 for DIPAE and 50 µg ml 1 for 3Q, MDEA, EDEA and TEA; pH 10; salt concentration: 30% (w/v); stirring speed: 1000 rpm; extraction time: 15 min.

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Sodium chloride

Sodium sulfate

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Sodium chloride

Sodium sulfate

(a)

(b)

119

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20 25 30 35 40

% Na2SO4

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(n

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DIPAE

3Q

MDEA

EDEA

TEA

(a)

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15 20 25 30 35

% Na2SO4

Up

take

(n

g)

3Q

DIPAE

MDEA

EDEA

TEA

(b)

The concentration of Na2SO4 was then varied from a salinity level of 20%

(w/v) to a highly saturated level of 40% (w/v) for HF-LPME while that of SPME was

varied from 15% (w/v) to 35% (w/v). Extractions without the addition of salt are

expected to yield negligible recoveries of the analytes. Figure 4-64 shows the effect of

salt concentration on extraction efficiency where the optimum salt concentration was

determined to be 30% (w/v). In both cases, increasing salt concentration aided in the

uptake of the polar analytes except for HF-LPME of DIPAE, where increasing salt

concentration led to a decrease in the uptake of the least polar analyte. The salt

concentration was adjusted to 30% (w/v) Na2SO4 for subsequent experiments.

Figure 4-64. (a) Effect of salt concentration on HF-LPME of the basic analytes. Spiking concentration: 0.5 µg ml 1 for DIPAE and 5 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); pH 12; stirring speed: 1000 rpm; extraction time: 15 min. (b) Effect of salt concentration on SPME of the basic analytes. Spiking concentration: 5 µg ml 1 for DIPAE and 50 µg ml 1 for 3Q, MDEA, EDEA and TEA; pH 10; stirring speed: 1000 rpm; extraction time: 15 min.

120

The effect of stirring speed on the extraction efficiency was investigated over

a range from 300 rpm to the maximum of 1250 rpm. As seen from Figure 4-65, the

optimum stirring speed was achieved at 1000 rpm for HF-LPME while the uptake of

DIPAE and 3Q by SPME showed opposite trends with increasing stirring speed. This

observation may be attributed to competition between DIPAE and 3Q, both relatively

bulky molecules, for uptake onto the fiber. Hence the stirring speed was set at 1000

rpm for HF-LPME and at 700 rpm for SPME in subsequent experiments.

Figure 4-65. (a) Effect of stirring on HF-LPME of the basic analytes. Spiking concentration: 0.5 µg ml 1 for DIPAE and 5 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); pH 12; salt concentration: 30% (w/v) Na2SO4; extraction time: 15 min. (b) Effect of stirring on SPME of the basic analytes. Spiking concentration: 5 µg ml 1 for DIPAE and 50 µg ml 1 for 3Q, MDEA, EDEA and TEA; pH 10; salt concentration: 30% (w/v) Na2SO4; extraction time: 15 min.

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(n

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DIPAE

3Q

MDEA

EDEA

TEA

(a)

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Up

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(n

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3Q

DIPAE

MDEA

EDEA

TEA

(b)

121

Figure 4-66 shows the time profile from 10 min to 25 min for HF-LPME and

from 15 min to 60 min for SPME respectively. Extraction times of 20 min for HF-

LPME and 30 min for SPME were selected since these were the optimum extraction

times for virtually all of the analytes.

Figure 4-66. (a) Effect of extraction time on HF-LPME of the basic analytes. Spiking concentration: 0.5 µg ml 1 for DIPAE and 5 µg ml 1 for 3Q, MDEA, EDEA and TEA; extraction solvent: chloroform/BSTFA (1:1); pH 12; salt concentration: 30% (w/v) Na2SO4; stirring speed: 1000 rpm. (b) Effect of extraction time on SPME of the basic analytes. Spiking concentration: 2.5 µg ml 1 for DIPAE and 25 µg ml 1 for 3Q, MDEA, EDEA and TEA; pH 10; salt concentration: 30% (w/v) Na2SO4; stirring speed: 700 rpm.

0

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Up

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(n

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DIPAE

3Q

MDEA

EDEA

TEA

(a)

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Upt

ake

(ng

)

3Q

DIPAE

MDEA

EDEA

TEA

(b)

122

4.3.8 Method validation for HF-LPME of basic degradation products

Using the optimum extraction conditions, the precision, linearity and limit of

detection of both methods were investigated using spiked deionized water samples.

The results are shown in Tables 4-8 and 4-9.

Table 4-8. Quantitative results of HF-LPME of the basic analytes.

Analyte Equation r2

% RSD

(n = 6)

LOD

(S/N = 5) (µg l 1)

DIPAE y = 0.0556x + 2.2385 0.9966 9 0.04 3Q y = 0.0016x

0.1049 0.9996 6 0.22 MDEA y = 0.0009x

1.7186 0.9975 10 0.09 EDEA y = 0.0072x

8.1536 0.9959 9 0.08 TEA y = 0.0001x

0.0649 0.9982 8 0.36

Table 4-9. Quantitative results of SPME of the basic analytes.

Analyte Equation r2

% RSD

(n = 6)

LOD

(S/N = 5) (µg l 1)

DIPAE y = 0.0434x + 45.988 0.9997 14 0.06 3Q y = 0.0022x

1.6675 0.9998 5 0.34 MDEA y = 0.0001x

0.0444 0.9985 22 0.11 EDEA y = 0.0005x

0.7572 0.9946 21 0.19 TEA y = 0.0001x

0.8472 0.9965 6 0.77

The linearity of calibration plots was investigated over a concentration range

of 0.05-25 µg ml 1 for HF-LPME and 0.5-25 µg ml 1 for SPME. The analytes

exhibited good linearity with squared regression coefficients of >0.994 for both

techniques. This allowed the quantification of the compounds by the method of

external calibration. The limits of detection for the analytes were estimated at a S/N

ratio of 5 under GC MS full scan conditions. The LODs ranged between 0.04 and

0.36 µg l 1 for HF-LPME and 0.06 and 0.77 µg l 1 for SPME. The relative standard

deviations for six extractions were between 6 and 10% for LPME and 5 and 22% for

SPME.

123

4.3.9 Comparison of HF-LPME of basic degradation products with SPME

In order to compare the performance of HF-LPME with in-situ derivatization

with that of SPME, extractions of the degradation products from deionized water

samples were conducted under the respective optimized HF-LPME and SPME

conditions. Figure 4-67 shows the results obtained.

Figure 4-67. Comparison of HF-LPME and SPME of the basic analytes. Spiking concentration: 1 µg ml 1 for DIPAE and 10 µg ml 1 for 3Q, MDEA, EDEA and TEA; HF-LPME extraction solvent: chloroform/BSTFA (1:1); pH 12; salt concentration: 30% (w/v) Na2SO4; stirring speed: 1000 rpm; extraction time: 20 min. SPME pH 10; salt concentration: 30% (w/v) Na2SO4; stirring speed: 700 rpm; extraction time: 30 min.

It was obvious that HF-LPME with in-situ derivatization surpassed that of

SPME for all of the analytes. Besides the simplicity and low cost, an added advantage

of HF-LPME over SPME is that after each extraction, the hollow fiber can be

discarded and a fresh one used for the next extraction. In contrast, the SPME fiber

required additional conditioning steps prior to and after extractions in order to prevent

carryover problems. Furthermore, unlike SPME where the fibers are limited to a

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(n

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HF-LPME

SPME

124

working pH range, HF-LPME does not have this constraint. In addition, the limited

lifetime of an SPME fiber subjected to direct immersion extraction is an unfavorable

consideration.

4.3.10 Analysis of a 20th Official OPCW Proficiency Test water sample

The OPCW conducts a biannual proficiency test for chemical verification

laboratories around the world to benchmark their capability. In the 20th Official

OPCW Proficiency Test held in October 2006, two chemicals, namely EMPA and 2-

(N,N-diisopropylamino)ethanol (DIPAE), were spiked into an aqueous waste sample.

The samples were prepared by Centre d Etudes du Bouchet (CEB), France. Using

only 0.3 ml of sample adjusted to pH 0 and topped up to 3 ml with deionized water,

EMPA was successfully detected by HF-LPME with in-situ derivatization using the

optimized extraction conditions discussed in Section 4.3.4. Similarly, after adjusting

the pH to 12 of only 0.3 ml of sample topped up to 3 ml with deionized water, DIPAE

was successfully detected by the same technique using the optimized extraction

conditions described in Section 4.3.7. Figures 4-68 and 4-69 show the respective

chromatograms obtained. The large signal at 11.39 min in Figure 4-68 was confirmed

to be ethyl tert.-butyldimethylsilylmethyl phosphonate (the TBDMS derivative of

EMPA) while this analyte was absent in the blank, as expected. The large signal at

9.38 min in Figure 4-69 was confirmed to be 2-(N,N-diisopropylamino)ethyl

trimethylsilyl ether (the TMS derivative of DIPAE) which was not present in the

blank.

125

Figure 4-68. Total ion chromatograms of HF-LPME under acidic conditions with in-situ derivatization of a 20th Official OPCW Proficiency Test aqueous sample (top) and blank (bottom). HF-LPME extraction solvent: chloroform/MTBSTFA (1:1); pH 0; salt concentration: 33% (w/v); stirring speed: 1000 rpm; extraction time: 45 min.

11.00 11.20 11.40 11.60 11.80 12.00 12.20 12.40 12.60 12.80 13.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

7500000

8000000

8500000

9000000

9500000

1e+07

1.05e+07

1.1e+07

1.15e+07

Time-->

Abundance

TIC: WB_1.D

10.80 11.00 11.20 11.40 11.60 11.80 12.00 12.20 12.40 12.60 12.80 13.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

7500000

8000000

8500000

9000000

9500000

1e+07

1.05e+07

1.1e+07

1.15e+07

Time-->

Abundance

TIC: W_1.D

EMPA-TBDMS

126

Figure 4-69. Total ion chromatograms of HF-LPME under basic conditions with in-situ derivatization of a 20th Official OPCW Proficiency Test aqueous sample (top) and blank (bottom). HF-LPME extraction solvent: chloroform/BSTFA (1:1); pH 12; salt concentration: 30% (w/v) Na2SO4; stirring speed: 1000 rpm; extraction time: 20 min.

8.00 8.20 8.40 8.60 8.80 9.00 9.20 9.40 9.60 9.80 10.00 10.20 10.40 10.60

5000000

1e+07

1.5e+07

2e+07

2.5e+07

3e+07

3.5e+07

4e+07

Time-->

Abundance

TIC: WSI_1.D

DIPAE-TMS

8.00 8.20 8.40 8.60 8.80 9.00 9.20 9.40 9.60 9.80 10.00 10.20 10.40 10.60

5000000

1e+07

1.5e+07

2e+07

2.5e+07

3e+07

3.5e+07

4e+07

Time-->

Abundance

TIC: WBSI_1.D

127

These experiments show that the technique can be applied to a wide range of

analytes of interest, whether they are acidic or basic in nature and only a very small

amount of sample and extraction solvent are required. The technique will continue to

be further tested and evaluated in future OPCW proficiency tests in order to

demonstrate its applicability to the entire range of analytes of interest. It is hoped that

the technique will complement and even substitute liquid-liquid extraction in the

recommended operating procedures for the analysis of CWAs and related compounds.

4.3.11 Conclusion

HF-LPME has been successfully utilized for the analysis of the several CWAs

and degradation products in aqueous samples. For analytes that required

derivatization, even though the derivatizing agents used were moisture-sensitive,

protection afforded by the hollow fiber allowed simultaneous extraction and

derivatization to be carried out followed by direct injection into the GC MS, without

the need for an additional off-line or separate derivatization step which requires

heating of the samples prior to HF-LPME. The procedure is simple, convenient to

perform and only a few microliters of organic solvent and derivatizing agents are

required. In addition, HF-LPME significantly improved the LODs over that of SPME.

The significantly lower cost of the hollow fiber membrane (one bundle of 2600 pieces

of 53.5 cm length costs only ~$200 USD) [297] as compared to commercially-

available SPME fibers (~$120 USD) further enhances its advantages over SPME. The

successful analysis of an OPCW Proficiency Test sample further affirms the

capability of the procedure.

128

5 CONCLUDING REMARKS

Thus far, there have been many reports on sol-gel solid-phase microextraction

(SPME) fibers, sol-gel molecularly imprinted polymers (MIPs), MIP SPME fibers but

very few reports on sol-gel MIP SPME fibers. In an attempt to develop sol-gel MIP

SPME fibers, the synthesized MIPs were first evaluated as sorbent material for solid-

phase extraction (SPE). It was found that the non-imprinted polymers (NIPs) did not

show zero absorptivity of the analytes. Hence, endcapping of the polymers was

carried out to reduce the non-specific adsorption of analytes on the surface silanol

groups of the polymers in order to achieve a high imprint factor. With endcapping, the

non-specific interaction of analytes with the NIPs was reduced but not totally

eliminated. Moreover, endcapping resulted in lower absorptivities and recoveries of

the analytes with the MIPs. Hence, non-endcapped MIPs and NIPs were used in

subsequent experiments. The various MIPs were evaluated for their binding properties

towards their respective templates as well as similar analytes. In addition, the MIP-

SPE procedure was compared against several sample preparation techniques for the

extraction of analytes of interest in samples with polyethylene glycol (PEG) as matrix

interference. Next, the development of sol-gel MIP SPME fibers was attempted.

However, the sol-gel coatings were prone to cracking, and flaked off upon drying at

room temperature. To successfully develop sol-gel MIP SPME fibers, it is expected

that a lot more effort has to be invested in optimizing the composition of the sol-gel

solution used for coating of the fibers. This can be achieved by using computer

modeling for the selection of the best monomers for molecular imprinting [373] prior

to the synthesis of the actual polymer systems.

On the other hand, a simple method of fabricating SPME fibers was achieved

by coating a novel poly(1-hydroxy-4-dodecyloxy-p-phenylene) polymer (PhPPP) as

129

the SPME coating. The novel PhPPP-coated fibers were evaluated for the analysis of

Lewisites after derivatization with thiols. The in-house prepared PhPPP fibers gave

comparable performance as compared to commercially-available SPME fibers. In

addition, they were easy to prepare and were significantly less costly in contrast to the

fairly expensive commercially-available fibers.

The extraction and determination of selected CWAs and degradation products

has been successfully demonstrated using hollow fiber-protected liquid-phase

microextraction (HF-LPME) and gas chromatography-mass spectrometry (GC MS).

In-situ derivatization of degradation products with silylating agents such as BSTFA

and MTBSTFA was possible as a result of the protection of the moisture-sensitive

derivatizing agents afforded by the hollow fiber. In order for HF-LPME to fully

substitute liquid-liquid extraction as a recommended operating procedure in the

analysis of chemical warfare agents and their degradation products, it is essential to

show that HF-LPME is applicable to all the various classes of chemicals. Future work

on HF-LPME will include the investigation of HF-LPME for the determination of

extremely volatile chemicals such as pinacolyl alcohol and chloropicrin as well as for

the determination of Lewisites with in-situ derivatization using mono- or dithiols as

derivatizing agents.

Future Work

A recently-introduced sample preparation technique, dispersive liquid-liquid

microextraction (DLLME), is being extensively investigated for various applications

[374-377] but has yet to be explored for the analysis of chemical warfare agents and

their degradation products. DLLME is a very simple and rapid method for extraction

and pre-concentration of organic compounds from water samples. In this method

130

(Figure 5-1), an appropriate mixture of extraction solvent (8.0 l of C2Cl4) and

disperser solvent (1.0 ml of acetone) is rapidly injected into the aqueous sample (5.0

ml) using a syringe. As a result, a cloudy solution is formed consisting of fine droplets

of extraction solvent dispersed entirely into the aqueous phase. Through

centrifugation, the extraction solvent is collected at the bottom of a conical sample

vial, withdrawn with a microsyringe and injected into a GC. DLLME is characterized

by very short extraction times, mainly due to the large surface area between the

solvent and the aqueous phase. Other advantages are the simplicity of operation, low

cost, high recoveries (82-111% at a spiking level of 5 g l 1) and enrichment factors

(603-1113), offering potential for ultra trace analysis (LODs between 0.007 and 0.030

g l 1, with RSD below 10% at 2 g l 1) [374]. It will be highly worthwhile to

investigate this microextraction technique in comparison with SPME and HF-LPME.

It is proposed that this be explored in a future study.

Figure 5-1. Photography of different steps in DLLME: (a) before injection of mixture of disperser solvent (acetone) and extraction solvent (C2Cl4) into sample solution, (b) starting of injection, (c) end of injection, (d) optical microscopic photography, magnitude 1000 (that shows fine particles of C2Cl4 in cloudy state), (e) after centrifuging and (f) enlarged view of sedimented phase (5.0±0.2 l) [374].

131

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PhysProp Database, North Syracuse, NY, 1999;

http://www.syrres.com/esc/physdemo.htm. (Accessed on 28 March 2008.).

[363] SPARC On-Line Calculator, University of Georgia, Athens, GA, release

w4.2.1374-s4.2.1370, 2008; http://ibmlc2.chem.uga.edu/sparc. (Accessed on

28 March, 2008.).

[364] J. Zhang, H.K. Lee, J. Chromatogr. A 1117 (2006) 31.

[365] C. Basheer, H.K. Lee, J. Chromatogr. A 1057 (2004) 163.

[366] M. Kawaguchi, R. Ito, N. Endo, N. Okanouchi, N. Sakui, K. Saito, H.

Nakazawa, J. Chromatogr. A 1110 (2006) 1.

[367] K.-J. Chia, S.-D. Huang, J. Chromatogr. A 1103 (2006) 158.

[368] P.-S. Chen, S.-D. Huang, J. Chromatogr. A 1118 (2006) 161.

http://www.syrres.com/esc/physdemo.htm

http://ibmlc2.chem.uga.edu/sparc

155

[369] K.E. Kramer, A.R.J. Andrews, J. Chromatogr. B 760 (2001) 27.

[370] A Leinonen, K. Vuorensola L.-M. Lepola T. Kuuranne, T. Kotiaho, R.A.

Ketola, R. Kostiainen, Anal. Chim. Acta 559 (2006) 166.

[371] K. Blau, J.M. Halket, Handbook of Derivatives for Chromatography, Wiley

Chichester, 1993, p. 51.

[372] G.D. Byrd, R.C. Paule, L.C. Sander, L.T. Sniegoski, E. White, H.T. Bausum,

J. Anal. Toxicol. 16 (1992) 182.

[373] D. Pavel, J. Lagowski, C.J. Lepage, Polymer 47 (2006) 8389.

[374] M. Rezaee, Y. Assadi, M.R. Milani Hosseini, E. Aghaee, F. Ahmadi, S.

Berijani, J. Chromatogr. A 1116 (2006) 1. Figure 5-1 reprinted from reference,

Copyright (2006), with permission from Elsevier.

[375] E. Zeini Jahromi, A. Bidari, Y. Assadi, M.R. Milani Hosseini, M.R. Jamali,

Anal. Chim. Acta 585 (2007) 305.

[376] A.S. Yazdi, N. Razavi, S.R. Yazdinejad, Talanta 75 (2008) 1293.

[377] A.P. Birjandi, A. Bidari, F. Rezaei, M.R. Milani Hosseini, Y. Assadi, J.

Chromatogr. A 1193 (2008) 19.

156

Schedule number

Chemical name CAS number Chemical Structure Molecular formula (Molecular Weight)

1.A.01 O-Alkyl (<C10, incl. cycloalkyl) alkyl (Me, Et, n-Pr or i-Pr)-phosphonofluoridates

e.g. Sarin: O-Isopropyl methylphosphonofluoridate (GB) 107-44-8

C4H10FO2P (140)

e.g. Soman: O-Pinacolyl methylphosphonofluoridate (GD)

96-64-0

C7H16FO2P (182)

1.A.02 O-Alkyl (<C10, incl. cycloalkyl) N,N-dialkyl (Me, Et, n-Pr or i-Pr) phosphoramidocyanidates

e.g. Tabun: O-Ethyl N,N-dimethyl phosphoramidocyanidate (GA) 77-81-6

C5H11N2O2P (162)

1.A.03 O-Alkyl (H or <C10, incl. cycloalkyl) S-2-dialkyl(Me, Et, n-Pr or i-Pr)-aminoethyl alkyl(Me, Et, n-Pr or i-Pr) phosphonothiolates and corresponding alkylated or protonated salts

e.g. VX: O-Ethyl S-2-diisopropylaminoethyl methyl phosphonothiolate 50782-69-9

C11H26NO2PS (267)

1.A.04 Sulfur mustards: 2-Chloroethylchloromethylsulfide 2625-76-5

C3H6Cl2S (144)

Mustard gas: Bis(2-chloroethyl)sulfide (HD)

505-60-2

C4H8Cl2S (158)

Bis(2-chloroethylthio)methane

63869-13-6

C5H10Cl2S2 (204)

Sesquimustard: 1,2-Bis(2-chloroethylthio)ethane (Q)

3563-36-8

C6H12Cl2S2 (218)

1,3-Bis(2-chloroethylthio)-n-propane

63905-10-2

C7H14Cl2S2 (232)

1,4-Bis(2-chloroethylthio)-n-butane

142868-93-7

ClS

SCl

C8H16Cl2S2 (246)

1,5-Bis(2-chloroethylthio)-n-pentane

142868-94-8

ClS S

Cl

C9H18Cl2S2 (260)

Bis(2-chloroethylthiomethyl)ether

63918-90-1

C6H12Cl2OS2 (234)

O-Mustard: Bis(2-chloroethylthioethyl)ether (T) 63918-89-8

ClS

OS

Cl

C8H16Cl2OS2 (262)

1.A.05 Lewisites: Lewisite 1: 2-Chlorovinyldichloroarsine (L1) 541-25-3

C2H2AsCl3 (206)

Lewisite 2: Bis(2-chlorovinyl)chloroarsine (L2) 40334-69-8

C4H4AsCl3 (232)

Lewisite 3: Tris(2-chlorovinyl)arsine (L3) 40334-70-1

C6H6AsCl3 (258)

P

O

OF

P

O

OF

NP

O

O

N

PO

O

SN

AsCl

Cl

Cl

Appendix 1 List of Schedule 1-3 Chemicals

ClS Cl

ClS

Cl

ClS S

Cl

ClS

SCl

ClS S

Cl

ClS O S

Cl

As

Cl

Cl Cl

AsCl Cl

Cl

157

Schedule number


1.A.06 Nitrogen mustards: HN1: Bis(2-chloroethyl)ethylamine 538-07-8

C6H13Cl2N (169)

HN2: Bis(2-chloroethyl)methylamine 51-75-2

C5H11Cl2N (155)

HN3: Tris(2-chloroethyl)amine 555-77-1

C6H12Cl3N (203)

1.A.07 Saxitoxin 35523-89-8

C10H17N7O4 (299)

1.A.08 Ricin 9009-86-3

Structure undefined 60 kDa

1.B.09 Alkyl (Me, Et, n-Pr or i-Pr) phosphonyldifluorides

e.g. DF: Methylphosphonyldifluoride

676-99-3 CH3F2OP (100)

1.B.10 O-Alkyl (H or <C10, incl. cycloalkyl) O-2-dialkyl(Me, Et, n-Pr or i-Pr)-aminoethyl alkyl(Me, Et, n-Pr or i-Pr) phosphonites and corresponding alkylated or protonated salts e.g. QL: O-Ethyl O-2-diisopropylaminoethyl methylphosphonite

57856-11-8 C11H26NO2P (235)

1.B.11 Chlorosarin: O-Isopropyl methylphosphonochloridate 1445-76-7

C4H10ClO2P (156)

1.B.12 Chlorosoman: O-Pinacolyl methylphosphonochloridate 7040-57-5

C7H16ClO2P (198)

2.A.01 Amiton: O,O-Diethyl S-[2-(diethylamino)ethyl] phosphorothiolate and corresponding alkylated or protonated salts 78-53-5

C10H24NO3PS (269)

2.A.02 PFIB: 1,1,3,3,3-Pentafluoro-2-(trifluoromethyl)-1-propene 382-21-8

C4F8 (200)

2.A.03 BZ: 3-Quinuclidinyl benzilate 6581-06-2

C21H23NO3 (337)

ClN

Cl

ClN

Cl

ClN

Cl

Cl

P

O

FF

PO O

N

P

O

OCl

P

O

OCl

PO

O

SN

O

F

FF

F

F

FF

F

NO

O

OH

158

Schedule number


2.B.04 Chemicals, except for those listed in Schedule 1, containing a phosphorus atom to which is bonded one methyl, ethyl or propyl (normal or iso) group but not further carbon atoms e.g. Methylphosphonyl dichloride 676-97-1

CH3Cl2OP (132)

Dimethyl methylphosphonate 756-79-6

C3H9O3P (124)

Exemption: Fonofos: O-Ethyl S-phenyl ethylphosphonothiolothionate 944-22-9

C10H15OPS2 (246)

2.B.05 N,N-Dialkyl (Me, Et, n-Pr or i-Pr) phosphoramidic dihalides

2.B.06 Dialkyl (Me, Et, n-Pr or i-Pr) N,N-dialkyl(Me, Et, n-Pr or i-Pr)-phosphoramidates

2.B.07 Arsenic trichloride 7784-34-1

AsCl3 (180)

2.B.08 Benzilic acid: 2,2-Diphenyl-2-hydroxyacetic acid 76-93-7

C14H12O3 (228)

2.B.09 Quinuclidin-3-ol 1619-34-7

C7H13NO (127)

2.B.10 N,N-Dialkyl (Me, Et, n-Pr or i-Pr) aminoethyl-2-chlorides and corresponding protonated salts

2.B.11 N,N-Dialkyl (Me, Et, n-Pr or i-Pr) aminoethane-2-ols and corresponding protonated salts

Exemptions: N,N-Dimethylaminoethanol and corresponding protonated salts 108-01-0

C4H11NO (89)

Exemptions: N,N-Diethylaminoethanol and corresponding protonated salts 100-37-8

C6H15NO (117)

2.B.12 N,N-Dialkyl (Me, Et, n-Pr or i-Pr) aminoethane-2-thiols and corresponding protonated salts

2.B.13 Thiodiglycol: Bis(2-hydroxyethyl)sulfide 111-48-8

C4H10O2S (122)

P

S

SO

P

O

ClCl

P

O

OO

PN

O

XX

R

R

PN

O

OO

R

R R

R

AsCl

Cl

Cl

OH

O

OH

NOH

ClN

R

R

OHN

R

R

OHN

OHN

SHN

R

R

OHS

OH

159

Schedule number


2.B.14 Pinacolyl alcohol: 3,3-Dimethylbutan-2-ol 464-07-3

C6H14O (102)

3.A.01 Phosgene: Carbonyl dichloride 75-44-5

CCl2O (98)

3.A.02 Cyanogen chloride 506-77-4

CClN (61)

3.A.03 Hydrogen cyanide 74-90-8

CHN (27)

3.A.04 Chloropicrin: Trichloronitromethane 76-06-2

CCl3NO2 (163)

3.B.05 Phosphorus oxychloride 10025-87-3

POCl3 (152)

3.B.06 Phosphorus trichloride 7719-12-2

PCl3 (136)

3.B.07 Phosphorus pentachloride 10026-13-8

PCl5 (206)

3.B.08 Trimethyl phosphite 121-45-9

C3H9O3P (124)

3.B.09 Triethyl phosphite 122-52-1

C6H15O3P (166)

3.B.10 Dimethyl phosphite 868-85-9

C2H7O3P (110)

3.B.11 Diethyl phosphite 762-04-9

C4H11O3P (138)

3.B.12 Sulfur monochloride 10025-67-9

S2Cl2 (134)

3.B.13 Sulfur dichloride 10545-99-0

SCl2 (102)

3.B.14 Thionyl chloride 7719-09-7

SOCl2 (118)

3.B.15 Ethyldiethanolamine 139-87-7

C6H15NO2 (133)

3.B.16 Methyldiethanolamine 105-59-9

C5H11NO2 (117)

3.B.17 Triethanolamine 102-71-6

C6H15NO3 (149)

OH

Cl Cl

O

NCl

NH

NO2

Cl

Cl

Cl

PCl

O

ClCl

PCl Cl

Cl

PCl Cl

ClCl

Cl

PO O

O

PO O

O

PO O

O

H

PO O

O

HS

SCl

ClS

ClCl

SClCl

O

OHN

OH

OHN

OH

OHN

OH

OH

160

APPENDIX 2 Total Ion Chromatograms from the MIP-SPE Study

Figure A2-1. Total ion chromatograms of samples upon (a) direct rotary evaporation; (b) SCX; (c) SAX; (d) loading onto PMPA-MIP-SPE; (e) washing of PMPA-MIP-SPE; (f) elution of PMPA-MIP-SPE cartridges.

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

Time-->

Abundance

TIC: P5_A.D(d) Absorptivity of

PMPA-MIP-SPE

TP

P

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

Time-->

Abundance

TIC: P5_W.D(e) Washing of

PMPA-MIP-SPE T

PP

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

Time-->

Abundance

TIC: SAX2.D

(c) SAX

IMPA

-TM

S

MP

A-T

MS

PMPA

-TM

S

TP

P

CM

PA-T

MS

EM

PA-T

MS

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

3.2e+07

Time-->

Abundance

TIC: V2.D(a) Direct

IMPA

-TM

S M

PA

-TM

S

PM

PA-T

MS

TP

P

CM

PA-T

MS

EM

PA

-TM

S

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

3.2e+07

Time-->

Abundance

TIC: SCX1.D(b) SCX

IMPA

-TM

S M

PA-T

MS

PMPA

-TM

S

TPP

CM

PA-T

MS

EM

PA-T

MS

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

7500000

8000000

8500000

9000000

9500000

1e+07

Time-->

Abundance

TIC: P5_R.D (f) Recovery of

PMPA-MIP-SPE

IMPA

-TM

S M

PA

-TM

S

PMPA

-TM

S

TPP

CM

PA-T

MS

EM

PA-T

MS

161

Figure A2-2. Total ion chromatograms of samples upon (a) direct rotary evaporation; (b) SCX; (c) SAX; (d) loading onto TDG-MIP-SPE; (e) elution of TDG-MIP-SPE cartridges.

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.000

1000000

2000000

3000000

4000000

5000000

6000000

7000000

8000000

9000000

1e+07

1.1e+07

1.2e+07

1.3e+07

1.4e+07

1.5e+07

1.6e+07

1.7e+07

1.8e+07

Time-->

Abundance

TIC: VAP1.D

TP

P

TD

G-T

MS

TD

GS-T

MS

TD

GS

O-T

MS

(a) Direct

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.000

1000000

2000000

3000000

4000000

5000000

6000000

7000000

8000000

9000000

1e+07

1.1e+07

1.2e+07

1.3e+07

1.4e+07

1.5e+07

1.6e+07

Time-->

Abundance

TIC: SCX3.D

TP

P

TD

G-T

MS

TD

GS

-TM

S

TD

GSO

-TM

S

(b) SCX

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.000

200000

400000

600000

800000

1000000

1200000

1400000

1600000

1800000

2000000

Time-->

Abundance

TIC: SAX4.D

TPP

(c) SAX

6.00 8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

Time-->

Abundance

TIC: T4_A.D

TPP

TD

G-T

MS

TD

GSO

-TM

S

(d) Absorptivity of

TDG-MIP-SPE

6.00 8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

Time-->

Abundance

TIC: T4_R.D

TPP

TD

G-T

MS

TD

GS

-TM

S

TD

GSO

-TM

S

(e) Recovery of

TDG-MIP-SPE

162

Figure A2-3. Total ion chromatograms of samples upon (a) direct rotary evaporation; (b) SCX; (c) SAX; (d) loading onto TEA-MIP-SPE; (e) elution of TEA-MIP-SPE cartridges.

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

3.2e+07

3.4e+07

3.6e+07

Time-->

Abundance

TIC: VAP5.D

TPP

MD

EA

-TM

S

TE

A-T

MS

ED

EA

-TM

S

(a) Direct

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

3.2e+07

Time-->

Abundance

TIC: SCX5.D

TPP

(b) SCX

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

200000

400000

600000

800000

1000000

1200000

1400000

1600000

1800000

2000000

2200000

2400000

2600000

Time-->

Abundance

TIC: SAX5.D

TPP

(c) SAX

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

100000

200000

300000

400000

500000

600000

700000

800000

900000

1000000

1100000

1200000

1300000

1400000

1500000

1600000

1700000

1800000

1900000

2000000

2100000

Time-->

Abundance

TIC: TPEG4_A.D

TPP

MD

EA

-TM

S

TE

A-T

MS

ED

EA

-TM

S

(d) Absorptivity of

TEA-MIP-SPE

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

1000000

2000000

3000000

4000000

5000000

6000000

7000000

8000000

9000000

1e+07

1.1e+07

1.2e+07

1.3e+07

1.4e+07

1.5e+07

1.6e+07

1.7e+07

1.8e+07

1.9e+07

Time-->

Abundance

TIC: TPEG4_R.D

TPP

MD

EA

-TM

S

TE

A-T

MS

ED

EA

-TM

S

(e) Recovery of

TEA-MIP-SPE

163

Figure A2-4. Total ion chromatograms of samples upon (a) direct rotary evaporation; (b) SCX; (c) SAX; (d) loading onto 3Q-MIP-SPE; (e) elution of 3Q-MIP-SPE cartridges.

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

Time-->

Abundance

TIC: VAP5.D

TPP

3Q-T

MS

(a) Direct

(b) SCX

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

Time-->

Abundance

TIC: SCX5.D

TPP

(c) SAX

7.00 8.00 9.00 10.0011.0012.0013.0014.00 15.0016.0017.0018.0019.00

200000

300000

400000

500000

600000

700000

800000

900000

1000000

1100000

1200000

1300000

1400000

Time-->

Abundance

TIC: SAX5.D

TPP

TPP

3Q-T

MS

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00

200000

400000

600000

800000

1000000

1200000

1400000

1600000

1800000

2000000

2200000

2400000

2600000

Time-->

Abundance

TIC: 3Q_DCM_A.D(d) Absorptivity of

3Q-MIP-SPE

TP

P

3Q-T

MS

7.00 8.00 9.00 10.0011.0012.0013.0014.0015.0016.0017.0018.0019.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

Time-->

Abundance

TIC: 3Q_DCM_R.D (e) Recovery of

3Q-MIP-SPE

164

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

3e+07

3.2e+07

3.4e+07

3.6e+07

Time-->

Abundance

TIC: VAP5.D

3Q-T

MS

PMPA

-TM

S

TD

G-T

MS

TP

P

TE

A-T

MS

(a) Direct

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

2000000

4000000

6000000

8000000

1e+07

1.2e+07

1.4e+07

1.6e+07

1.8e+07

2e+07

2.2e+07

2.4e+07

2.6e+07

2.8e+07

Time-->

Abundance

TIC: SCX5.D

PM

PA

-TM

S

TD

G-T

MS

TPP

(b) SCX

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

200000

400000

600000

800000

1000000

1200000

1400000

1600000

1800000

2000000

2200000

2400000

2600000

2800000

Time-->

Abundance

TIC: SAX5.D

PMPA

-TM

S

TPP

(c) SAX

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

Time-->

Abundance

TIC: MET4_A.D

3Q-T

MS

TD

G-T

MS

TPP

TE

A-T

MS

(d) Absorptivity of

MIP-SPE

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

7500000

8000000

8500000

9000000

9500000

1e+07

1.05e+07

1.1e+07

1.15e+07

1.2e+07

1.25e+07

1.3e+07

Time-->

Abundance

TIC: MET4_R.D

3Q-T

MS PM

PA-T

MS

TD

G-T

MS

TPP

TE

A-T

MS

(e) Recovery of MIP-SPE

with EtOH

165

Figure A2-5. Total ion chromatograms of samples upon (a) direct rotary evaporation; (b) SCX; (c) SAX; (d) loading onto MIP-SPE; (e) elution of MIP-SPE cartridges with EtOH; (f) loading onto MIP-SPE; (g) elution of MIP-SPE cartridges with 1% TFA in water; (h) loading onto MIP-SPE; (i) elution of MIP-SPE cartridges with 1% TE in water.

(f) Absorptivity of

MIP-SPE

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

Time-->

Abundance

TIC: MTFA4_A.D

3Q-T

MS

TD

G-T

MS

TPP

TE

A-T

MS

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

1000000

2000000

3000000

4000000

5000000

6000000

7000000

8000000

9000000

1e+07

1.1e+07

1.2e+07

1.3e+07

1.4e+07

1.5e+07

Time-->

Abundance

TIC: MTFA4_R.D

3Q-T

MS

PMP

A-T

MS

TPP

TE

A-T

MS

(g) Recovery of MIP-SPE

with 1% TFA in water

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

500000

1000000

1500000

2000000

2500000

3000000

3500000

4000000

4500000

5000000

5500000

6000000

6500000

7000000

Time-->

Abundance

TIC: MTE4_A.D3Q-T

MS

TD

G-T

MS

TPP

TE

A-T

MS

(h) Absorptivity of

MIP-SPE

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00

1000000

2000000

3000000

4000000

5000000

6000000

7000000

8000000

9000000

1e+07

1.1e+07

1.2e+07

1.3e+07

1.4e+07

Time-->

Abundance

TIC: MTE4_R.D

3Q-T

MS

PM

PA

-TM

S

TD

G-T

MS

TP

P

TE

A-T

MS

(i) Recovery of MIP-SPE with 1% TE in water

166

APPENDIX 3 MASS SPECTRA

Figure A3-1. Mass spectrum of GB (MW: 140) Figure A3-2. Mass spectrum of GD (MW: 182)

Figure A3-3. Mass spectrum of GA (MW: 162) Figure A3-4. Mass spectrum of HD (MW: 158)

Figure A3-5. Mass spectrum of VX (MW: 267) Figure A3-6. Mass spectrum of L3 (MW: 258)

Figure A3-7. Mass spectrum of L2-ET (MW: 258) Figure A3-8. Mass spectrum of L1-ET (MW: 258)

20PPMCWA #81-89 RT: 5.52-5.54 AV: 6 SB: 18 5.34-5.45 , 5.81-5.96 NL: 2.28E5T: + c EI Full ms [40.00-209.00]

40 50 60 70 80 90 100 110 120 130 140 150m/z

0

10

20

30

40

50

60

70

80

90

100

Re

lativ

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99

125

8143

7947 9867 10059 12682


40 60 80 100 120 140 160 180m/z

0

10

20

30

40

50

60

70

80

90

100

Re

lativ

e A

bund

ance

12699

8269

41

5783

4381 85

9855 67 12745 12570 8649 79 100

20PPMCWA #1051 RT: 10.65 AV: 1 SB: 9 9.66-10.35 , 11.06-11.49 NL: 1.52E5T: + c EI Full ms [40.00-165.00]

40 50 60 70 80 90 100 110 120 130 140 150 160 170m/z

0

10

20

30

40

50

60

70

80

90

100

Re

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7043

44

133

162106

117

108

4765 92 135119 1479071 93 161 16376 13655 60


40 50 60 70 80 90 100 110 120 130 140 150 160m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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109

111

15863

160

45 59 736547 9658 123 162989557 12581 113

20PPMCWA #2515-2522 RT: 18.39-18.41 AV: 4 NL: 2.29E5T: + c EI Full ms [40.00-267.00]

40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

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114

72

127

70 79 84 11543 112 13985 1675649 98 25261 144

50ET #1966-1972 RT: 15.49-15.52 AV: 7 NL: 3.72E5T: + c EI Full ms [40.00-264.00]

40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

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ance

136

77

145

110

113147

100

51

17187 135 161115101 148 173

150 19775 1248961 163 25849 1348463 17560 151 262187 201


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

136

145107

258171

113 203

260147

205229

173197119100 23185 12287 16114859

615845 262124 19314994 20775 163 233159 22317563 79 208


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

107 136148

171

197258

229

143173108

165260

23119910959 15061

1358945 17510058 122 20387 19675 16157 26123362 223

P

O

OF

P

O

OF

NP

O

O

N

ClS

Cl

PO

O

SN

AsCl Cl

Cl

Cl

S

AsCl Cl

S

AsS

167

Figure A3-9. Mass spectrum of L2-PT (MW: 272) Figure A3-10. Mass spectrum of L1-PT (MW: 286)

Figure A3-11. Mass spectrum of L2-BT (MW: 286) Figure A3-12. Mass spectrum of L1-BT (MW: 314)

Figure A3-13. Mass spectrum of L1-EDT (MW: 228) Figure A3-14. Mass spectrum of L2-EDT (MW: 290)

Figure A3-15. Mass spectrum of L1-PDT (MW: 242) Figure A3-16. Mass spectrum of L2-PDT (MW: 304)

50PT #2294-2298 RT: 17.22-17.24 AV: 5 NL: 3.74E5T: + c EI Full ms [40.00-278.00]

40 60 80 100 120 140 160 180 200 220 240 260 280m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

43

145

272150

274

107127 203

13685 110 229171

205169

161 23111910184211

17345 185 27659 19794 15951 75 21323361 17576 239 277

50PT #2593-2600 RT: 18.81-18.82 AV: 3 NL: 8.85E5T: + c EI Full ms [40.00-292.00]

40 60 80 100 120 140 160 180 200 220 240 260 280m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

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ance

176

15043

107

286169 211

143116 185 243

165 288139 213

151 24518745 136101 201 22559 1527558 117 2899473 21419376 246

50BT #2502-2507 RT: 18.33-18.34 AV: 3 NL: 6.35E5T: + c EI Full ms [40.00-293.00]

40 60 80 100 120 140 160 180 200 220 240 260 280m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

57

286

145

41288

164147

85107

229141 171110 20316155

23120511947 10084 22558 173135 197 2908959 149 176

19375 232207 25173 291

50BT #2870-2874 RT: 20.27-20.29 AV: 5 NL: 1.72E6T: + c EI Full ms [40.00-320.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300 320m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

204

57164

314

107 169225

41257

130 143 316205171 227201 20655 259139 14547 58 197 224 253110101 31789 183 2282218775 279260

50EDT #2791-2794 RT: 19.85-19.86 AV: 2 SB: 17 19.34-19.69 , 20.41-20.56 NL: 9.06E3T: + c EI Full ms [40.00-281.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

61

121

168145

107 123119 167

147169110 161 19659

10045 1369447 173149 198 22863 9253 85 132 20375 184 23115984

50PDT #2467-2474 RT: 18.14-18.17 AV: 7 NL: 6.44E5T: + c EI Full ms [40.00-248.00]

40 60 80 100 120 140 160 180 200 220 240m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

242

107

149

181

244

106

148165

45

142110 132 1537346 100 20718316858 64 75 112 24678 201121 2091848549 17115693

50PDT #2914-2918 RT: 20.50-20.52 AV: 2 NL: 1.42E5T: + c EI Full ms [40.00-282.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

107

73

106

14541 16747147

133 169109 14975 18113679 1006149 112 203197132 24288 207157 229

Cl

S

AsCl Cl

S

SAs

Cl

S

AsCl Cl

S

SAs

S

S

AsCl

Cl

S

AsCl

SH

AsS

S

ClCl

S

AsCl

SH

50EDT #2212-2218 RT: 16.79-16.82 AV: 7 NL: 7.54E5T: + c EI Full ms [40.00-234.00]

40 60 80 100 120 140 160 180 200 220m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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167

165

107 228

230

200

16945 139110 14258 202100 121 13675 1476046 23257 92 12311885 174 193156 204

168

Figure A3-17. Mass spectrum of EMPA-TMS (MW: 196) Figure A3-18. Mass spectrum of EMPA-TBDMS (MW: 238)

Figure A3-19. Mass spectrum of IMPA-TMS (MW: 210) Figure A3-20. Mass spectrum of IMPA-TBDMS (MW: 252)

Figure A3-21. Mass spectrum of MPA-TMS (MW: 240) Figure A3-22. Mass spectrum of MPA-TBDMS (MW: 324)

Figure A3-23. Mass spectrum of nPPA-TMS (MW: 268) Figure A3-24. Mass spectrum of nPPA-TBDMS (MW: 352)

20_10SI #358-369 RT: 7.76-7.77 AV: 2 NL: 2.29E4T: + c EI Full ms [41.00-318.00]

40 60 80 100 120 140 160 180 200m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

153

75

73

45

77 121 1371025949 154 17291 18156 61 79 15186 107 16998 18751 12384 19615570 14493 17313065 119 167

20PPM_10MT #293-300 RT: 11.46-11.47 AV: 2 NL: 4.04E6T: + c EI Full ms [40.00-310.00]

40 60 80 100 120 140 160 180 200 220 240m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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ance

153

181

75

154155 18212173 77 19545 137 151 1835947 107 2239184 179

20_10SI #445 RT: 8.16 AV: 1 NL: 4.39E4T: + c EI Full ms [40.00-209.00]

40 60 80 100 120 140 160 180 200 220m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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ance

153

7543 151

7745 73 169154

121107 13747 9149 79 19561 15512384 119 1701681067255 9265 139135 197 209179


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

153

15475

19515512173 2371517741 13745 57 19610784 193


40 60 80 100 120 140 160 180 200 220 240m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

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ance

225

7773

133

22645135 147

227105 24059 11547 20912191 137 1951511077261 1818958 123 165 210 22892 241196138

20PPM_10MT #1078 RT: 15.10 AV: 1 NL: 3.69E6T: + c EI Full ms [40.00-326.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300 320m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

267

268

73

269153135 195

212 225 3091477559 105 251 27012157 181 197 213 226154

REFSTD5 #505-508 RT: 10.95-10.98 AV: 2 NL: 2.22E6T: + c EI Full ms [40.00-342.00]

40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

253

226

254225211

195135 25573 147 237

224209121 228 26845 75 149136 24118111759 165105


50 100 150 200 250 300 350m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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295

296

29773

195135 181 337147 253240198 223 29812175 279 33859 18241 165

P

O

OO

Si

P

O

OO

Si

P

O

OO

Si

P

O

OO

Si

P

O

OO

Si

Si

P

O

OO

Si

Si

P

O

OO

Si

Si

P

O

OO

Si

Si

169

Figure A3-25. Mass spectrum of PMPA-TMS (MW: 252) Figure A3-26. Mass spectrum of PMPA-TBDMS (MW: 294)

Figure A3-27. Mass spectrum of CMPA-TMS (MW: 250) Figure A3-28. Mass spectrum of CMPA-TBDMS (MW: 292)

Figure A3-29. Mass spectrum of BA-TMS (MW: 372) Figure A3-30. Mass spectrum of BA-TBDMS (MW: 456)

Figure A3-31. Mass spectrum of EHES-TMS (MW: 178) Figure A3-32. Mass spectrum of EHES-TBDMS (MW: 220)


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

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100

Rel

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153

41

7516915157 121

19573

4313777 15469 196

5545 16813685 91 107 170 19761 179120 135103 163 237198138 181


40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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153

237154

21175 121 182155 19573 151 23841 57 12377 193137 27969 21285 179 196107

20PPM #649-651 RT: 14.10-14.13 AV: 3 NL: 1.50E6T: + c EI Full ms [40.00-600.00]

40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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153

169

1547515173 155 17013712141 776755 84 221107 179167

20PPMPA #1393-1397 RT: 15.97-15.98 AV: 2 NL: 2.90E6T: + c EI Full ms [40.00-310.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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153

15475 211

15573 19541 12155 7767 152137 23521284 179107 196

BASI

#1006 RT: 18.53 AV: 1 NL: 2.95E6T: + c EI Full ms [41.00-361.00]

50 100 150 200 250 300 350 400m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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255

73

256

147165

239 257 32916645 74 240105 148135 330258179 35722511559 207

BAMT

#1345-1349 RT: 22.74-22.75 AV: 2 NL: 2.47E6T: + c EI Full ms [40.00-445.00]

50 100 150 200 250 300 350 400 450m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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ance

297

147

73 371

298 399165

372239166 400

299240 37375 133 167 401105 20959 225 241 26877 374300281 325 441


40 60 80 100 120 140 160 180m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

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73

75

89

16345

11910359

61

47 8858178

43 101 11691877749 72 93 10555 16412062 13386 18070 147135

100_10MT #831-835 RT: 11.60-11.60 AV: 2 NL: 1.21E5T: + c EI Full ms [40.00-208.00]

40 60 80 100 120 140 160 180 200 220m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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89163

75

119574161 73

4547 16491

887756 11710193 121 2057253 85 166135115 149

P

O

OO

Si

P

O

OO

Si

O

O

Si

OSi

O

O

Si

OSi

SO

Si

SO

Si

P

O

OO

Si

P

O

OO

Si

170

Figure A3-33. Mass spectrum of TDG-TMS (MW: 266) Figure A3-34. Mass spectrum of TDG-TBDMS (MW: 350)

Figure A3-35. Mass spectrum of TDGS-TMS (MW: 282) Figure A3-36. Mass spectrum of TDGSO-TMS (MW: 298)

Figure A3-37. Mass spectrum of QOH-TMS (MW: 326) Figure A3-38. Mass spectrum of QOH-TBDMS (MW: 410)

Figure A3-39. Mass spectrum of TOH-TMS (MW: 370) Figure A3-40. Mass spectrum of TOH-TBDMS (MW: 454)


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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73

11675 147

103133

45 59

1171018747 61149 161105 17613566 9349 77 242154120 192106 251165


50 100 150 200 250 300 350m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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147

73

293

87

75 189148119 233

133 29414959101

29516345 66 190103 23413484 21912099 296265165 335

20PPM_T #829-830 RT: 16.33-16.35 AV: 2 NL: 9.30E5T: + c EI Full ms [40.00-600.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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73

117

75 166267

147135116103 26523845 1335966 268104 14887 16747 122 223177 239136 26489 191

20PPM_T #842-844 RT: 16.49-16.52 AV: 3 NL: 8.18E5T: + c EI Full ms [40.00-600.00]

40 60 80 100 120 140 160 180 200 220 240 260 280 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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73

283

117

14775284

28510166 239885945 116 13387 11897 148 193 28647 255240139 167115 13190


50 100 150 200 250 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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73

11775103

45 87 17759 61 130 17686 13447 101 104 149 161 17814749 23621766 210


50 100 150 200 250 300 350 400m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

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219

73

159 220119

75 22110387 16159 133 14845 207 353195179 222 251


50 100 150 200 250 300m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

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73

103116

75

117

45 1771198759 133101 16347 72 86 104 17814991 174


50 100 150 200 250 300 350 400m/z

0

10

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30

40

50

60

70

80

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100

Rel

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219

73

159

22011575

22120787 103 133 16159 8145 134 170 191 222 281 295

SOO

SiSi SOO

SiSi

SOO

SiSi

O

SOO

SiSi

OO

SOS

OSi

Si

SOS

OSi

Si

OS

OS

OSi Si

OS

OS

OSi Si

171

Figure A3-41. Mass spectrum of DIPAE-TMS (MW: 217) Figure A3-42. Mass spectrum of DIPAE-TBDMS (MW: 259)

Figure A3-43. Mass spectrum of MDEA-TMS (MW: 263) Figure A3-44. Mass spectrum of MDEA-TBDMS (MW: 347)

Figure A3-45. Mass spectrum of EDEA-TMS (MW: 277) Figure A3-46. Mass spectrum of EDEA-TBDMS (MW: 361)

Figure A3-47. Mass spectrum of TEA-TMS (MW: 365) Figure A3-48. Mass spectrum of TEA-TBDMS (MW: 491)

20_5NSI #444-449 RT: 9.42-9.43 AV: 3 NL: 1.63E6T: + c EI Full ms [40.00-372.00]

40 60 80 100 120 140 160 180 200 220m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

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ance

114

72

73 11520243 14470 8659 75 160 21749 94 116100 203128 14561 103

20_5NMT #1038-1040 RT: 12.47-12.48 AV: 2 SB: 5 11.78-12.34 , 12.59-13.26 NL: 3.36E6T: + c EI Full ms [40.00-281.00]

40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

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114

72115

7324443 75 14759 70 144 23356 86 259245100 202116 160128


40 60 80 100 120 140 160 180 200 220 240 260m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

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160

73

161147

11716275 2485942 11684 13045 101 14870 86 263144

20_5NMT #2074-2080 RT: 16.85-16.87 AV: 6 NL: 5.95E6T: + c EI Full ms [40.00-351.00]

50 100 150 200 250 300 350m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

202

203

73

204 290147130 3327559 116101 159 29142 84 200 205


40 60 80 100 120 140 160 180 200 220 240 260 280m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

174

73

175

147117 1767559 130 26284 10145 144 24486 14811672 172 277188


50 100 150 200 250 300 350m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

216

217

73

218147130 304 34659 75 11510157 15989 219214


50 100 150 200 250 300 350m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

262

26373

264350130 147117

7559 351101 174 26545 14886

NO

Si NO

Si

NO O

Si Si NO O

Si Si

NO O

Si Si NO O

Si Si

NO O

Si Si

OSi

50PPM_1 #1820 RT: 20.18 AV: 1 SB: 1 20.00-20.13 , 20.23-20.25 NL: 1.25E5T: + c EI Full ms [40.00-537.00]

50 100 150 200 250 300 350 400 450 500m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

346

73

347

348144

57 13075 101 35522141 434281115 295 476186 313172 267 344 401

NO O

Si Si

OSi

172

Figure A3-49. Mass spectrum of 3Q (MW: 127) Figure A3-50. Mass spectrum of 3Q-TMS (MW: 199)

Figure A3-51. Mass spectrum of 3Q-TBDMS (MW: 241)

20_5NMT #460-467 RT: 10.03-10.06 AV: 7 SB: 53 9.52-9.87 , 10.19-10.26 NL: 8.21E4T: + c EI Full ms [40.00-207.00]

40 50 60 70 80 90 100 110 120 130m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

42

127

58

43 98

7082

57 11255 12644 11069 9683 9971

8473 1285467 8681 108947952 65 11345 59 10093

20_5NSI #677-680 RT: 10.90-10.91 AV: 2 SB: 8 10.25-10.81 , 10.97-11.04 NL: 8.94E4T: + c EI Full ms [40.00-207.00]

40 60 80 100 120 140 160 180 200m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

73

19912996

184

198110101 14542

75 15617014270 116 13010844

55 20082 15712759 83 18568 9453 14180 146 158126 20161 172 18692 147 1598752 140


40 60 80 100 120 140 160 180 200 220 240m/z

0

10

20

30

40

50

60

70

80

90

100

Rel

ativ

e A

bund

ance

110184

96

73

18575

241

41 101

8449 141 1561278659 157108 226111

13155 70 81 142 186170 240115 24217114368 21319812694 158 227 243132 172144 214

N

OH N

OSi

N

OSi

173

APPENDIX 4 LIST OF POSTER PRESENTATIONS AND

PUBLICATIONS

Poster Presentations

[1] N.H.S. Lee, M.T. Sng, C. Basheer, H.K. Lee, Sol-Gel Molecularly Imprinted

Polymers for Solid Phase Extraction, 4th Singapore International Symposium

on Protection Against Toxic Substances (4th SISPAT) in association with

Chemical and Biological Medical Treatment Series, 6-10 December 2004,

Shangri-La Hotel, Singapore.

[2] N.H.S. Lee, M.T. Sng, C. Basheer, H.K. Lee, Hollow-Fibre Protected Liquid-

Phase Microextraction for the Analysis of Chemical Warfare Agents and

Degradation Products, Workshop on the Analysis related to Chemical

Weapons to Mark the Tenth Anniversary of the Entry into Force of the

Convention, 5 - 7 September 2007, Gustavelund Conference Hotel, Tuusula,

Finland.

[3] N.H.S. Lee, M.T. Sng, C. Basheer, H.K. Lee, Hollow-Fibre Protected Liquid-

Phase Microextraction for the Determination of Basic Degradation Products of

Chemical Warfare Agents, SICC-5: Singapore International Chemistry

Conference 5 & APCE 2007: 7th Asia-Pacific International Symposium on

Microscale Separation and Analysis, 16-19 December 2007, Singapore

International Convention & Exhibition Centre (Suntec Singapore), Singapore.

Publications

[1] H.S.N. Lee, C. Basheer, H.K. Lee, J. Chromatogr. A 1124 (2006) 91.

[2] H.S.N. Lee, M.T. Sng, C. Basheer, H.K. Lee, J. Chromatogr. A 1148 (2007) 8.

[3] H.S.N. Lee, M.T. Sng, C. Basheer, H.K. Lee, J. Chromatogr. A 1196-1197

(2008) 125.

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