lc-ms in analytical toxicology

LC-MS in analytical toxicology:some practical considerationsLewis Couchman and Phillip E. Morgan*

ABSTRACT: Liquid chromatography, coupled with single-stage or tandem mass spectrometry, is a powerful tool increasinglyused in analytical toxicology. However, the atmospheric pressure ionization processes involved are complex, and subject tointerference from matrix components, for example. Further, the techniques used in sample preparation, chromatography andmass analysis are developing rapidly. An understanding of the advantages and limitations of LC-MS ensures appropriateanalyses are performed, and that reliable results are generated. Consideration should be given to the influence of the samplepreparation and chromatographic conditions on the ionization of the analyte at the mass spectrometer interface. This reviewaims to provide some practical guidance and examples to aid method development for commonly encountered analytes inanalytical toxicology. Copyright © 2010 John Wiley & Sons, Ltd.

Keywords: liquid chromatography; mass spectrometry; analytical toxicology; sample preparation

Introduction

Analytical toxicology is the detection, identification and mea-surement of drugs and other foreign compounds (xenobiotics)and their metabolites in biological and related specimens. Analy-ses tend to fall into (i) emergency and general hospital toxicol-ogy, including ‘poisons screening’ or (ii) more specializedcategories such as forensic toxicology, screening for drugs ofabuse, therapeutic drug monitoring (TDM) and occupational/environmental toxicology. However, there is considerableoverlap between all of these areas. Sample matrices can becomplex, particularly in the case of post-mortem analyses, and ahigh degree of analytical reliability, sensitivity and specificity maybe required (Maurer, 2006, 2007; Flanagan et al., 2007).

Since the first report of an interface between liquid chroma-tography and mass spectrometry a number of interface designs,most importantly that of atmospheric pressure ionization (API),have been developed to improve the efficiency of the ionizationprocess. A better understanding of the physical processesinvolved with analyte ionization means that problems associatedwith co-eluting matrix components (ion suppression andenhancement) can be accounted for and minimized. Whilst gaschromatography–mass spectrometry (GC-MS)—in conjunctionwith detailed GC-MS spectral libraries—remains a very useful toolfor systematic toxicological analysis (STA), non-volatile, polar (e.g.conjugated metabolites), and thermally labile compounds aredifficult or impossible to analyse without lengthy derivatizationprocedures (Marquet and Lachâtre, 1999; Flanagan et al., 2007;Dresen et al., 2010). HPLC with diode-array detection (DAD) pro-vides a means to analyse compounds not suited to GC, but suffersdue to the non-specific nature of UV detection. Certain com-pounds of toxicological relevance also have poor UV absorbance.LC-MS (and LC-tandem MS, LC-MS/MS) may be applied to com-pounds not suited to GC analysis, and spectral libraries now existfor a very wide range of toxicologically relevant compounds(although ionization and fragmentation conditions remain non-standardized). Recent developments in accurate mass measure-

ment have allowed tentative identification of compoundswithout the absolute need for reference materials.

An understanding of the advantages and limitations of MSmethods may help generate reliable quantitative and qualitativedata. Sample collection/pre-treatment procedures and protocols,

* Correspondence to: P. E. Morgan, Toxicology Unit, Department of ClinicalBiochemistry, King’s College Hospital NHS Foundation Trust, Denmark Hill,London SE5 9RS, UK. E-mail: [email protected]

Toxicology Unit, Department of Clinical Biochemistry, King’s College Hospi-tal NHS Foundation Trust, Denmark Hill, London SE5 9RS, UK

Abbreviations used: 6-MAM, 6-monoacetylmorphine; AAFS, The AmericanAcademy of Forensic Sciences; ACN, acetonitrile; APCI, atmospheric pressurechemical ionization; API, atmospheric pressure ionization; APPI, atmosphericpressure photoionization; BEG, benzoylecgonine; CID, collision-induced dis-sociation; CNS, central nervous system; DAD, diode-array detection; DMA,dimethoxyamphetamine; DMANO, dimethylamphetamine N-oxide; DoA,drugs of abuse; DVB, divinylbenzene; EDDP, 2-ethylidene-1,5-dimethyl-3,3-diphenylpyrrolidine; EDTA, ethylene diamine tetra-acetic acid; EMDP,2-ethyl-5-methyl-3,3-diphenyl-1-pyrroline; ESI, electrospray ionization;GC-MS, gas chromatography–mass spectrometry; GHB, gamma hydroxybu-tyrate; GUS, general unknown screening; H-ESI, heated electrosprayionization; HILIC, hydrophilic interaction liquid chromatography; HRMS,high-resolution mass spectrometry; IPA, isopropyl alcohol/2-propanol; ISTD,internal standard; LC-MS, liquid chromatography–mass spectrometry;LC-MS/MS, liquid chromatography–tandem mass spectrometry; LLE, liquid–liquid extraction; LLoQ, lower limit of quantitation; LOD, limit of detection;M3G, morphine-3-glucuronide; M6G, morphine-6-glucuronide; MBDB,methylbenzodioxolylbutanamine; MDA, 3,4-methylenedixoyamphetamine;MDEA/MDE, 3,4-methylenedioxyethylamphetamine; MDMA, 3,4-methylenedioxymetamphetamine; MEPS, micro-extraction by packedsorbent; MTBE, methyl tert-butyl ether; PCP, phencyclidine; PPT, protein pre-cipitation; RAM, restricted access material; RP, reversed-phase; SCX, strongcation-exchange; SOFT, The Society of Forensic Toxicologists; SPE, solid-phase extraction; SRM, selected reaction monitoring; STA, systematic toxi-cological analysis; TDM, therapeutic drug monitoring; TFA, trifluoroaceticacid; THC, tetrahydrocannabinol; THC-COOH, carboxy-tetrahydrocannabinol; TOF, time of flight; TRIS, tris(hydroxymethyl)ami-nomethane; UHPLC, ultra-high-pressure liquid chromatography.

Special Issue: Review Article

Received 30 September 2010, Accepted 4 October 2010 Published online in Wiley Online Library: 10 December 2010

(wileyonlinelibrary.com) DOI 10.1002/bmc.1566

100

Biomed. Chromatogr. 2011; 25: 100–123Copyright © 2010 John Wiley & Sons, Ltd.

and choice of sample preparation and HPLC conditions all influ-ence the final result (Flanagan et al., 2005, 2007; Dinis-Oliveiraet al., 2010). This review highlights some practical points for con-sideration when using LC-MS (or MS/MS) for analysis of the mostcommon biological samples encountered in analytical toxicologylaboratories.

Sample PreparationSample preparation prior to LC-MS analysis aims to reduce matrixeffects via removal of potential interferences, and to get theanalyte into a form amenable to analysis. However, for drugs andlow molecular mass compounds, co-eluting components such asproteins, lipids and salts may cause variability in the efficiency ofanalyte ionization (Bonfiglio et al., 1999; Jemal et al., 2010). Non-volatile components may also cause a reduction in sensitivity andform deposits inside the instrument. The removal of phospholip-ids from plasma/whole blood, compounds known to cause sig-nificant ion suppression in many cases, is the basis for a numberof reports comparing the efficiency of certain sample preparationtechniques (Little et al., 2006; Chambers et al., 2007; Ismaeil et al.,2008; Du and White, 2008; Pucci et al., 2009). As well as sample‘clean-up’, analytes can also be concentrated or diluted duringsample preparation, depending on factors such as samplevolume and the anticipated analyte concentration(s). Poor per-formance may result if sample preparation is overlooked (VanEeckhaut et al., 2009). That said, the superior selectivity of MSdetectors coupled with the versatility of HPLC has prompted thesimplification, miniaturization and greater automation of samplepreparation processes. For STA, the direct injection of urinesamples, usually after filtration or centrifugation and/or dilution(‘dilute and shoot’) has been shown to be useful. Advantagesinclude increased selectivity and lower limits of detection (LODs)in many cases. However, the direct injection strategy is prone tosignificant variations in matrix effects.

The general techniques for the preparation of solid, liquid andgaseous samples for chromatographic analyses have beenreviewed (Smith, 2003; Flanagan et al., 2006; Chen et al., 2008;Nováková and Vlčová, 2009).

General Considerations

The physicochemical properties of the analyte(s), for examplepKa, and octanol–water coefficient (as logP) can help guidetowards an appropriate sample preparation procedure (Flanaganet al., 2007). In liquid samples, for example, manipulation of pH

for analytes possessing ionizable groups often adds selectivity tothe procedure, as well as helping to optimize recovery of ana-lyte(s) from the matrix and ensuring reproducible sample consis-tency (Hendriks et al., 2007). Changes in sample pH may affectthe recovery of other analytes, therefore conditions are often acompromise, particularly when multiple analytes from differentclasses are to be simultaneously investigated. Control ofpH is often through the use of buffer solutions such astris(hydroxymethyl)-aminomethane (TRIS, pH range 7–9), sodiumacetate–acetic acid solutions (pH range 3.5–6), citric acid–citratesolutions (pH range 3–6) and carbonate–bicarbonatesolutions (pH range 9–11). In our experience, solutions of TRIS(2 mol/L) can be used to good effect even at pH 10.6for the extraction of basic drugs from serum or plasma(Flanagan et al., 2001; Morgan et al., 2003). Detailed listsof compounds and their properties are available, for examplehttp://www.sigmaaldrich.com/life-science/core-bioreagents/biological-buffers/learning-center/buffer-reference-center.html(accessed 10 August 2010). If pH control is less of an issue duringsample preparation, simple acidification or alkalinization may beachieved by addition of strongly acidic or alkaline aqueous solu-tions. However, some analytes decompose, undergo structuralrearrangements or react under these conditions, leading to erro-neous results. This is particularly true for certain metabolites,such as N-oxides, but it can also be useful, for example, in theconversion of glucuronidated metabolites to the parent com-pound. Special care must be taken not to introduce non-volatilebuffer salts into the mass spectrometer. Other considerations arelisted in Table 1.

The most common sample preparation techniques currentlyemployed in analytical toxicology are protein precipitation (PPT),liquid–liquid extraction (LLE), and solid-phase extraction (SPE). Inthe following sections, selected applications will be used in orderto highlight the different approaches taken.

Protein Precipitation

The precipitation of proteins from biological fluids is rapid andsimple, and the efficiency of various precipitation reagents hasbeen evaluated (Blanchard, 1981). Chambers et al. (2007) showedthat, for the compounds tested—a range of eight representativepolar and non-polar analytes—recovery was generally good (76–114%) following PPT of plasma. In particular, recovery of polaranalytes included in the test was better than the three LLEmethods used for comparison, and comparable to two of thethree SPE methods investigated. However, matrix effects were

Table 1. Some sample preparation considerations

• Minimize matrix effects as far as practical by (i) removal of endogenous interferences, e.g. phospholipids and (ii) the use ofappropriate internal standard(s).

• Is the chosen method cost effective? Consider the time spent preparing samples, the number of steps involved, and the cost ofreagents and materials.

• Is it possible to automate the procedure for high-thoughput analyses?• Does the method give suitable analyte recovery? Recovery should be reproducible, and independent of analyte concentration.• Evaporation steps should not degrade the analyte(s). The use of an inert gas (e.g. nitrogen) and temperatures as low as

practical are recommended. Additional measures, such as acidification of the eluate prior to evaporation, may be required tominimize loss of amphetamine and related compounds (Mortier et al., 2002).

• Logistical considerations, e.g. fume hood(s), provision of vacuum and compressed air and/or nitrogen, bench space required.• Environmental/health and safety impact. 101

LC-MS in analytical toxicology

Biomed. Chromatogr. 2011; 25: 100–123 View this article online at wileyonlinelibrary.comCopyright © 2010 John Wiley & Sons, Ltd.

considerable, ranging from 47 to 61% suppression of ionization.Moreover, the choice of methanol or acetonitrile as the precipi-tating reagent also affected the abundance of residual phospho-lipids in the supernatant. Although the acetonitrile-treatedsamples contained substantially less phospholipids, the interfer-ence was such that a fully validated method was not deemedviable. However, validated methods have been published(Table 2). Evaporation of the supernatant and reconstitution ofthe residue in mobile phase gave good results in the analysis oflamotrigine and metabolites in plasma (Beck et al., 2006), with noapparent ion-suppression. Dilution of the supernatant prior toinjection is also an option, and can be adjusted according to theexpected analyte concentration. Using this approach, Kirchherrand Kühn-Velten (2006) reported negligible matrix effects for allanalytes with the exception of olanzapine, for which the use ofmatrix-matched calibration solutions was mandatory.

The poor ability of PPT methods to remove certain phospho-lipids from plasma/serum (Chambers et al., 2007; Jemal et al.,2010) means that further treatment of the supernatant, such asSPE or LLE, may be necessary to reduce ion-suppression and toenhance sensitivity (Flanagan et al., 2006). Chilling the precipita-tion reagent prior to use may increase recovery of analytes andimprove reproducibility (Choo et al., 2007). For whole blood andother ‘dirty’ sample matrices, a PPT step is often employed—sometimes with ultrasonication—before the supernatant is sub-jected to SPE (Kristoffersen et al., 2007; Marin et al., 2008;Mercerolle et al., 2008; Chimalakonda et al., 2010; Wu et al., 2010).

Protein precipitation is clearly an attractive sample prepara-tion technique due to its speed, simplicity and the good recov-ery of polar analytes compared with some SPE and LLEprocedures. It is applicable to a range of LC-MS methods rel-evant to toxicology. However, the failure to remove endogenousphospholipids and other potentially interfering compoundsmake it prone to severe matrix effects in the absence of furthertreatment of the supernatant. Moreover, it is non-selective anddifficult to automate, and there may be considerable variationin the effectiveness of precipitation between samples, eventhose of the same matrix.

Liquid–Liquid Extraction

Given appropriate conditions, many analytes readily partitioninto an organic phase from an aqueous sample, the extent ofpartitioning being based on the octanol–water partition coeffi-cients of the analytes. The ideal organic phase is immiscible withthe sample matrix, of low toxicity, volatility and flammability,and efficiently extracts the analytes of interest without alsoextracting endogenous material. Many solvents are used(Table 3), none of which meet the ‘ideal’ criteria listed, and mostof which require specialized storage, handling and disposal pro-cedures. Polar organic solvents such as methyl tert-butyl ether(MTBE) and 1-chlorobutane tend to extract fewer interferencessuch as phospholipids. Depending on the analyte(s), the use ofacidic pH conditions further helps to prevent co-extraction ofthese compounds. Hence, these solvents have been recom-mended as the best single-component solvents in LLE in termsof analyte recovery and extract ‘cleanliness’ (Jemal et al., 2010).However, other solvents may be better suited to particular ana-lytes, matrices or conditions, and should not be excluded (Srini-vas, 2009). Ethyl acetate LLE gave ‘cleaner’ extracts from urinecompared with SPE for analysis of doping agents (Goebel et al.,2004), but caused problems with the extraction of certain

diuretics containing sulfur side chains. This was overcome byusing MTBE instead of ethyl acetate, the caveat being poorrecovery of the other analytes. Mixtures of various solvents canalso prove useful. A rather elaborate extraction solvent (2 mL/sample) consisting of a mixture of dichloromethane (520 mL),dichloroethane (520 mL), heptane (600 mL) and 2-propanol(380 mL) was used to extract benzodiazepines from urine(Glover and Allen, 2010). Various LLE solvent and buffer combi-nations were evaluated for the extraction of 19 antipsychoticdrugs from whole blood (Saar et al., 2009). In that study, TRISbuffer at pH 9.2 (1 mL, 2 mol/L) gave the best extraction effi-ciency regardless of extracting solvent used, as compared withsodium sulfate (1 mL, saturated) and sodium bicarbonate(100 mg). Of the solvents used (8 mL of 1-chlorobutane, ethylacetate or a 50:50 diethyl ether–ethyl acetate mixture),1-chlorobutane gave the highest extraction efficiency for allanalytes, with the exception of sulpiride. Matrix effects werebroadly similar between the tested solvents; however, increasedmatrix effects were observed for olanzapine when extractedusing ethyl acetate. In an evaluation of several extractionmethods for methadone in human plasma, LLE using hexane–isoamyl alcohol was more efficient and less likely to cause ionsuppression than mixed mode SPE, protein precipitation, orcolumn switching arrangements (Souverain et al., 2004).

A mixture of butyl acetate : butanol (9 + 1, v/v) was used for theextraction of amphetamine, metamphetamine and amisulpridefrom serum/plasma (Couchman et al., 2010a), and is applicable toother antipsychotic drugs and amphetamine-related com-pounds. MTBE does not extract phospholipids (Jemal et al., 2010),but can give poor recovery of polar compounds (Chambers et al.,2007). The use of basified solvent and multiple extractions of thesame sample can help improve sensitivity, although the latter issomewhat tedious. MTBE was preferred over toluene and butylacetate for extracting antipsychotics from post-mortem blood(Roman et al., 2008).

A general assumption is that for selective LLE of ionizable ana-lytes, the sample pH should be adjusted to a value at least 2 unitsabove or below the analyte pKa for basic or acidic analytes,respectively. However, in certain circumstances (and indeed inour own experience), enhanced selectivity and good recoveriescan be achieved whilst the analyte is apparently largely ionized(Hendriks et al., 2007).

Liquid–liquid extraction is simple, robust and transferable,and shows good reductions in matrix effects (Guo et al., 2005;Jemal et al., 2010). It may be more suited to urgent analysesthan SPE (Flanagan et al., 2006; Wille and Lambert, 2007).However, it may not be suitable for hydrophilic compounds, andthe formation of emulsions can make it difficult to isolate theextraction solvent. Appropriate treatment of the sample prior toaddition of the solvent can give highly selective extractions insome cases, and direct injection of the extract may also be pos-sible. In most cases, though, there is a need to evaporate thesolvent extract before re-dissolving the residue in mobile phase.This extra step costs time and increases error. Storage, handlingof relatively large volumes, toxicity, disposal and the cost of andneed for high-purity solvents used in LLE are also issues for con-sideration.

Solid-phase Extraction

SPE involves application of the sample onto a bed of material akinto the stationary phase in HPLC. Chemical modification of the

102

L. Couchman and P. E. Morgan

Biomed. Chromatogr. 2011; 25: 100–123View this article online at wileyonlinelibrary.com Copyright © 2010 John Wiley & Sons, Ltd.

Tab

le2.

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103



Tab

le2.

Cont

inue

d

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(s)

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mp

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104



Tab

le3.

Sele

cted

liqui

d–liq

uid

extr

actio

nm

etho

dsin

anal

ytic

alto

xico

logy

LC-M

S

Ana

lyte

(s)

Mat

rixSa

mp

lere

quire

men

tsEx

trac

tion

solv

ent(

s)Re

fere

nce

Tam

oxife

nem

etab

olite

s(6

anal

ytes

)U

rine

3m

LEt

hyla

ceta

teor

MTB

EM

azza

rino

etal

.,20

10D

iure

tics

and

pro

ben

ecid

(18

anal

ytes

)U

rine

2m

LEt

hyla

ceta

te.D

oub

leex

trac

tion

(2p

ortio

nsof

solv

ent)

.LO

D2–

100

ng/m

L

Dev

ente

ret

al.,

2002

Benz

odia

zep

ines

and

met

abol

ites,

zolp

idem

and

zop

iclo

ne(2

8an

alyt

es)

Bloo

d,ur

ine,

hair

250

mL(b

lood

/urin

e)20

mg

(hai

r)1-

Chl

orob

utan

e.La

rge

volu

mes

ofso

lven

tLa

loup

etal

.,20

05

Benz

odiz

epin

es(2

2an

alyt

es).

LC-M

S-(T

oF)

Urin

e1

mL

Chl

orof

orm

:iso

pro

pyla

lcoh

ol(9

+1)

.LO

D0.

5–3

ng/m

LEl

Sohl

yet

al.,

2006

Risp

erid

one,

9-O

Hris

per

idon

e,b

usip

rone

,zip

rasi

done

,p

erp

hena

zine

,zuc

lop

enth

ixol

,flu

phe

nazi

ne,fl

upen

thix

ol

Post

-mor

tem

blo

od1

gM

TBE.

LOD

bet

ter

than

1mg

/LRo

man

etal

.,20

08

Bup

reno

rphi

ne,n

orb

upre

norp

hine

,na

loxo

nePl

asm

a1

mL

1-ch

loro

but

ane

:ace

toni

trile

(4+

1).

LLO

Q0.

1ng

/mL

Moo

dyet

al.,

2002

Vario

uscl

asse

s:an

tidep

ress

ants

,b

enzo

diaz

epin

es,n

euro

lep

tics,

bet

a-b

lock

ers,

oral

antid

iab

etic

s,b

rain

-dea

thdi

agno

ses

anal

ytes

(140

anal

ytes

)

Plas

ma

500

mLBu

tyla

ceta

te:e

thyl

acet

ate

(1+

1)Re

man

eet

al.,

2010

Ana

bol

icst

eroi

ds(t

etra

hydr

oges

trin

one,

gest

rinon

e,3’

-hyd

roxy

stan

ozol

ol,

17a-

tren

bol

one)

Urin

e5

mL

Die

thyl

ethe

r.LO

D1–

10ng

/mL

Dev

ente

ret

al.,

2006

Am

phe

tam

ine,

met

amp

heta

min

e,M

DA

,MD

MA

,PC

PO

ralfl

uid

1m

LH

exan

e:e

thyl

acet

ate

(1+

1).L

OD

2–10

ng/m

LKa

laet

al.,

2008

Benz

odia

zep

ines

(17

anal

ytes

)U

rine

500

mLD

ichl

orom

etha

ne:d

ichl

oroe

than

e:

hep

tane

:2-p

rop

anol

(52

+52

+60

+38

).LO

D0.

31–2

.5mg

/L

Glo

ver

and

Alle

n,20

10

105



packed bed to provide similar chemistries to the packings used inHPLC means that a large number of different phases are available,in many formats, making SPE a highly versatile technique. Formany SPE materials, allowing the sorbent to become dry risksinactivation of the bonded phase, leading to poor recovery ofanalytes, and to ‘channelling’ of the sample through the packedbed. However, some newer, polymeric sorbents are more resis-tant to this kind of problem, can tolerate relatively ‘dirty’ samplesand are stable over a wider pH range compared with silica-basedmaterials (Yawney et al., 2002). Polymeric sorbents may also beless soluble in certain solvents compared with silica-based mate-rial (Verplaetse and Tytgat, 2010). The presence of some particu-lates is generally tolerated, but the beds can become blocked. Toovercome this, samples can be filtered or centrifuged, after anypH adjustment. Once eluted from the cartridge, the eluate ismost often evaporated and reconstituted. A mobile phase con-taining a high proportion of organic solvent may facilitate thedirect injection of organic sample extracts, saving time andreducing the risk of errors (Couchman et al., 2010a), but is obvi-ously dependant on the initial chromatographic conditionschosen. Direct injection, and dilution of the eluate prior to injec-tion have been reported (Table 4). ‘Generic’ SPE methods havebeen proposed, not necessarily intended for the analysis of everyanalyte, but to provide a set of starting conditions with a reason-able chance of success with minimal adjustment. Schellen et al.(2003) evaluated several SPE materials for extraction of a range ofdrugs from serum/plasma, noting that a divinylbenzene (DVB)material offered the best combination of extraction capacity anddesorption efficiency amongst those tested. A series of SPE sor-bents ranging from non-polar, to mixed-mode, to polymeric,were tested for their performance in the systematic toxicologicalanalysis of a diverse range of drugs (17 analytes) in whole blood(Decaestecker et al., 2003). In this experiment, a C8-modified silicamaterial was found to offer the best overall recovery, followedclosely by a mixed-mode polymeric sorbent. It was reported thatthe C8 material offered the ‘cleanest’ sample extracts. Differencesin analyte ionization efficiencies and concentrations in thesample may be compensated for by adjustment of samplevolumes and/or the sorbent mass used for SPE (Rentsch, 2003;Maralikova and Weinmann, 2004). Silica-based C8 and C18 sor-bents generally offer predictable chemistry and good recovery ofa wide range of analytes from various matrices, often without theuse of extreme pH values. However, they are relatively non-selective, and good recoveries of polar, hydrophilic, and ionizedcompounds may be difficult.

So-called ‘mixed-mode’ materials, in which a combination ofinteractions may be exploited to allow efficient clean-up by usingrelatively harsh wash steps with minimal loss of analyte(s), areincreasingly used for sample preparation. Such methods typicallyinvolve the sequential elution of acidic, neutral and basic com-pounds using solvents at appropriate pH, and this versatility hasled to their increasing popularity. For ‘comprehensive’ analyses,the eluates from different fractions are usually combined andevaporated before reconstitution in an LC-compatible solvent.Otherwise only the fraction containing the analytes of interest iscollected. Control and manipulation of pH is often the key inthese cases. A diverse range of applications in which mixed-modeSPE has been used have been reported (Table 4).

Advantages of SPE include the extraction of relatively hydro-philic compounds such as metabolites of morphine and cocaine(Yawney et al., 2002; Jagerdeo et al., 2008), enhanced selectivityimparted through chemical modification of the particle surface,

ease of automation, high sensitivity and high efficiency.Compared with LLE, SPE is considered to use less solvent, is lesstime-consuming, and gives ‘cleaner’ extracts, especially fromante-mortem blood, plasma or serum (Chambers et al., 2007). Theuse of 96-well SPE plates can increase sample throughput, whilstat the same time reducing sample and solvent volumes (Malletet al., 2003; Ashman et al., 2010).

Limitations of SPE include co-extraction of interfering com-pounds, and poor extraction of some drugs (Yawney et al., 2002;Goebel et al., 2004). The latter was addressed to some extent bySchellen et al. (2003) in using a larger sorbent bed for the extrac-tion of a very polar compound (acetaminophen), and by acidifi-cation of the sample (20 mL phosphoric acid/mL of sample) toimprove recovery of sulfadiazine and sulfamerazine. Anotherproblem may be blockage of the packed bed during sampleapplication. The use of larger SPE cartridges, and/or centrifuga-tion, ultrasonication, protein precipitation or dilution of thesample can be useful in these cases, especially when viscoussamples are encountered (Choo et al., 2007; Saar et al., 2009). Thesensitivity of modern instruments is such that the dilution ofextracts may be routinely required in order to maintain concen-trations within the linear range of the mass spectrometer(Langman et al., 2009).

Batch-to-batch reproducibility of the sorbent bed, althoughless of a problem now compared with a few years ago, is still aconcern (Nováková and Vlčová, 2009) and without automationthe number of samples that can be processed simultaneously isgenerally limited to the number of spaces available on thevacuum manifold, typically 20. Flow-rate through the sorbentbed is difficult to control in most cases, which can lead to variableanalyte recovery. Although the 96-well plate SPE format hasadvantages—low volumes of sample and solvents, small desorp-tion volumes, decreased void volumes, semi-automation—thereare also disadvantages particularly in terms of cost, when not allthe wells are used each time. Micropipette tip-based SPE mayoffer an alternative (Shen et al., 2006).

As well as selectivity, trace enrichment and ease of use com-pared with LLE and PPT, procedures involving SPE are relativelyeasy to automate (Yawney et al., 2002). This can save time andreduce labour costs, and at the same time enhance forensicintegrity (Jagerdeo et al., 2008; Robandt et al., 2009). Instrumentsfor semi-automated or fully automated SPE have been availablefor some years. Despite applications demonstrating robustness,reliability and time-savings (Jourdil et al., 2003; Schellen et al.,2003; Goebel et al., 2004; Robandt et al., 2008, 2009), the capitalcost of the instrumentation required even for a semi-automatedsystem is likely to limit widespread implementation. Moreover,pre-extraction steps such as addition of internal standard,hydrolysis, pre-mixing of the sample with buffer solutions andprotein precipitation, and post-extraction evaporation andreconstitution of the eluate often need to be performed off-line(Jourdil et al., 2003; Maralikova and Weinmann, 2004; Kristoffer-sen et al., 2007; Robandt et al., 2009).

Other Matrices/Preparation Techniques

Direct injection of samples onto size-exclusion HPLC columnshave reported for the analysis of benzodiazepines in diluted urineand plasma (Lee et al., 2003, 2006), and for the measurement ofb-blockers in diluted plasma (Umezawa et al., 2008). Sampleextraction and the need for column switching were eliminated,and good recovery, precision and LODs were reported. Despite

106



Tab

le4.

Sele

cted

solid

-pha

seex

trac

tion

met

hods

inan

alyt

ical

toxi

colo

gyLC

-MS

Ana

lyte

(s)

Mat

rixSa

mp

levo

lum

eSP

Em

ode

and

com

men

tsRe

fere

nce

Qua

tern

ary

amm

oniu

mdr

ugs

and

herb

icid

es(1

1an

alyt

es)

Who

leb

lood

1m

LW

eak

catio

n-ex

chan

geso

rben

t.LO

D3.

6–20

.4ng

/mL

Ariffi

nan

dA

nder

son,

2006

Dop

ing

agen

ts(1

03an

alyt

es)

Urin

e50

0mL

Mix

edm

ode

96w

ellp

late

.Dire

ctin

ject

ion

ofel

uate

Bado

udet

al.,

2010

Com

pre

hens

ive

scre

enin

g(3

92<r

eal>

anal

ytes

,plu

s24

5<t

heor

etic

alm

ass>

inlib

rary

)U

rine

1m

LM

ixed

mod

e.A

cidi

can

db

asic

elua

tefr

actio

nsco

mb

ined

Pela

nder

etal

.,20

03

Com

pre

hens

ive

scre

enin

g(8

15ex

act

mas

ses

for

DoA

,the

rap

eutic

drug

s,de

sign

erdr

ugs.

Rete

ntio

nda

tafo

rha

lfof

thes

e)

Vitr

eous

hum

or1

mL

Mix

edm

ode.

As

Pela

nder

etal

.,20

03.L

ittle

sam

ple

left

ifre

-ana

lysi

sre

quire

dPe

land

eret

al.,

2010

Mor

phi

ne,c

odei

ne,e

thyl

mor

phi

negl

ucur

onid

es,

6-ac

etyl

mor

phi

neU

rine

50mL

Mix

edm

ode.

LOD

5–30

ng/m

LSv

enss

onet

al.,

2007

Keta

min

ean

dse

lect

edm

etab

olite

s(3

anal

ytes

)U

rine

4m

LM

ixed

mod

e.LO

D0.

03(k

etam

ine)

and

0.05

(nor

keta

min

e)ng

/mL

Park

inet

al.,

2008

Can

nab

inoi

ds,o

pio

ids

and

stim

ulan

ts(1

3an

alyt

es)

Bloo

d,p

lasm

a,ur

ine

1m

LC

18so

rben

t.LO

Dra

nge

0.2–

4.0

ng/m

LM

aral

ikov

aan

dW

einm

ann,

2004

MD

MA

and

met

abol

ites

(3an

alyt

es)

Urin

e2.

5m

LM

ixed

mod

e.D

irect

inje

ctio

nof

elua

te.

LOD

0.01

5–0.

04mg

/mL

Jenk

ins

etal

.,20

04

Ant

idep

ress

ants

(16

anal

ytes

)O

ralfl

uid

250

mLM

ixed

mod

e.LO

Q5

ng/m

LC

oulte

ret

al.,

2010

Bup

rop

ion,

hydr

oxyb

upro

pio

n,th

reoh

ydro

bup

rop

ion

Post

-mor

tem

blo

odan

dur

ine

100

mLM

ixed

mod

e.LO

D5

mg/L

Mer

cero

lleet

al.,

2008

Mul

tiple

DoA

and

met

abol

ites

(30

anal

ytes

)U

rine

500

mLM

ixed

mod

e.A

llLO

Ds

<3

ng/m

LFe

nget

al.,

2007

Mul

tiple

hallu

cino

gens

,chl

orp

heni

ram

ine,

keta

min

e,rit

alin

icac

id,a

ndm

etab

olite

s(1

4an

alyt

es)

Urin

e50

0mL

Mix

edm

ode.

LOD

rang

e0.

0003

–2.5

ng/m

LFe

rnan

dez

etal

.,20

07

Coc

aine

and

met

abol

ites

(7an

alyt

es)

Urin

e1

mL

Mix

edm

ode.

LOD

0.25

ng/m

Lal

lana

lyte

sLa

ngm

anet

al.,

2009

Benz

oyec

goni

ne,m

-hyd

roxy

ben

zoyl

ecgo

nine

,p-

hydr

oxyb

enzo

ylec

goni

ne,n

orb

enzo

yecg

onin

eU

rine

1m

LC

8so

rben

t.D

irect

inje

ctio

nof

elua

te.L

OD

1.2

ng/m

LRo

ban

dtet

al.,

2008

Mid

azol

am,h

ydro

xym

idaz

olam

,hyd

roxy

mid

azol

amgl

ucur

onid

e,m

orp

hine

,mor

phi

ne-3

-glu

curo

nide

,m

orp

hine

-6-g

lucu

roni

de

Plas

ma

50mL

Mix

edm

ode

96-w

ellf

orm

at.T

wo

sep

arat

em

etho

dsre

quire

d.D

irect

inje

ctio

nof

elua

te.L

LOQ

<10

ng/m

L

Ash

man

etal

.,20

10

Am

phe

tam

ine,

met

amp

heta

min

e,M

DA

,MD

MA

,M

DEA

,ep

hedr

ine,

pse

udoe

phe

drin

e,p

hent

erm

ine,

phe

nyle

thyl

amin

e

Bloo

d1

mL

Mix

edm

ode

Ap

ollo

nio

etal

.,20

06

107



Tab

le4.

Cont

inue

d

Ana

lyte

(s)

Mat

rixSa

mp

levo

lum

eSP

Em

ode

and

com

men

tsRe

fere

nce

Mor

phi

ne,6

-ace

tylm

orp

hine

,am

phe

tam

ine,

met

amp

heta

min

e,M

DA

,MD

MA

,MD

EA,M

BDB,

ben

zoyl

ecgo

nine

,coc

aine

Bloo

d20

0mL

Mix

edm

ode.

LOD

:0.5

ng/m

Lfo

rm

etam

phe

tam

ine,

MD

MA

,ben

zoye

cgon

ine,

coca

ine;

1ng

/mL

for

mor

phi

ne,6

-MA

M,M

DA

,MD

EA,M

BDB

Con

chei

roet

al.,

2006

Flun

itraz

epam

,7-a

min

oflun

itraz

epam

,3-

hydr

oxyfl

unitr

azep

am,

N-d

esm

ethy

lflun

itraz

epam

Plas

ma

and

urin

e1

mL

Mix

edm

ode.

LOD

(urin

e):0

.025

ng/m

Lflu

nitr

azep

aman

d7-

amin

oflun

itraz

epam

;0.

04ng

/mL

N-d

esm

ethy

lflun

itraz

epam

;0.2

ng/m

L3-

hydr

oxyfl

unitr

azep

am

Jour

dile

tal.,

2003

Vario

usdr

ugs

(aci

dic,

bas

ic,p

olar

,non

-pol

ar,

arom

atic

;11

anal

ytes

)Pl

asm

a10

0mL

Div

inyl

ben

zene

sorb

ent.

Dire

ctin

ject

ion

ofel

uate

.LL

OQ

<1

ng/m

Lex

cep

tac

etam

inop

hen

(2ng

/mL)

Sche

llen

etal

.,20

03

Am

phe

tam

ine,

met

amp

heta

min

e,M

DA

,MD

MA

,M

DEA

,DM

A,D

MA

NO

Urin

e1

mL

Mix

edm

ode.

LOD

<2

ng/m

LKi

met

al.,

2008

Benz

odia

zep

ines

(16)

and

met

abol

ites

(5)

Seru

m10

0mL

Mix

edm

ode.

LOD

rang

e0.

3–11

.4ng

/mL

Nak

amur

aet

al.,

2009

Zona

sim

ide,

lam

otrig

ine,

top

iram

ate,

phe

nob

arb

ital,

phe

nyto

in,c

arb

amaz

epin

e,ca

rbam

azep

ine-

10,1

1-di

ol,1

0-hy

drox

ycar

bam

azep

ine,

carb

amaz

epin

e-10

,11-

epox

ide

Plas

ma

100

mLLO

D<

1ng

/mL

exce

pt

for

carb

amaz

epin

e-10

,11-

diol

(9.7

7ng

/mL)

Sub

ram

ania

net

al.,

2008

Alk

aloi

ds—

acon

itine

,hyp

acon

itine

,gel

sem

ine,

race

anis

odam

ine,

stry

chni

ne,b

ruci

ne.H

igh

per

cent

age

orga

nic

(95–

75–4

5%)f

orb

ette

rio

niza

tion/

sens

itivi

ty

Bloo

dan

dur

ine

500

mLM

ixed

mod

e.LO

D0.

1–1

ng/m

Lin

urin

e;0.

1–1.

5ng

/mL

inb

lood

Wu

etal

.,20

10

Coc

aine

,ben

zoyl

ecgo

nine

,ecg

onin

em

ethy

lest

er,

ecgo

nine

,coc

aeth

ylen

eU

rine

1m

LM

ixed

mod

e.D

irect

inje

ctio

nof

elua

teJa

gerd

eoet

al.,

2008

Basi

cdr

ugs

ofab

use

(18

anal

ytes

)Se

rum

100

mLPP

T(M

eOH

:ZnS

O4)

,eva

por

ate

and

reco

nstit

ute

then

onlin

eco

lum

nsw

itchi

ngSP

E.LO

D<

1ng

/mL

Bouz

aset

al.,

2009

108



matrix effects of 11–30% suppression for the b-blockers, exces-sive variability was not observed.

In a comparison of various LLE and SPE procedures for theextraction of antipsychotics from different blood matrices, i.e.ante-mortem, non-decomposed post-mortem, and decom-posed post-mortem blood, considerable variation was observedin terms of both analyte recovery and matrix effects (Saar et al.,2009). However, the variability appeared to be related to thematrix rather than to the method of extraction. Differences havealso been seen in the recovery of PCP from serumand from whole blood (Chimalakonda et al., 2010) after extrac-tion using mixed-mode SPE, and attempts to improve the LODby reducing the volume of solvent used in the final reconstitu-tion step failed, possibly due to incomplete dissolutionof the analyte, and/or concentration of ion-suppressing matrixcomponents.

Oral fluid is an attractive alternative to urine and plasma,however variations in the matrix warrant stringent method vali-dation. Recovery of analytes from oral fluid is often superiorwhen compared with plasma (Sergi et al., 2009). Directinjection of diluted oral fluid has been reported, however thisresulted in shortened HPLC column lifetimes (Allen et al., 2005).SPE and LLE procedures analogous to those used for urineand plasma samples are usually employed. However, theuse of sampling devices can lead to problems with interferencesand other contaminants (Mortier et al., 2002; Allen et al., 2005),even after an extraction procedure. The use of a micro-SPE(m-SPE) procedure (Sergi et al., 2010) reduced matrix effectscompared with PPT, avoiding the need for a ‘clean-up’ gradientafter each injection. A review of analytical procedures for theanalysis of oral fluids for drugs of abuse is available (Samynet al., 2007).

Discussion

Advances in LC and MS technologies, plus economic pressures,mean that sample preparation centres less on the selective andoften lengthy extraction of specific analytes, and more on theremoval—either during sample preparation or during chromato-graphic analysis—of species likely to interfere in analysis. Sor-bents for SPE have been developed to specifically removephospholipids and proteins from biological samples. Monolithicsorbents less prone to blockages are available as disposable tipsor in 96-well plates. Despite the superior results obtained fromSPE compared with PPT, the cost in terms of labour and materialsmay still be difficult to justify (Mallet et al., 2003).

Automated extraction is routine within the pharmaceuticalindustry, but limited in the toxicology laboratory at present. Tur-bulent flow chromatography offers efficient removal of potentialinterferences (Du and White, 2008), and fast analyses from bio-logical fluids when compared with SPE or LLE procedures (Bernaet al., 2004; Zhou et al., 2005; Morgan et al., 2010). Offline handingof the sample is often limited to centrifugation and dilution(Couchman et al., 2010b). Moreover, in the same way that auto-mated SPE systems can be configured for minimum cycle times(Schellen et al., 2003), such systems are easily adapted to ensuremaximal use of detector time through staggered, parallelmethods in which samples are extracted whilst previous extractsare being analysed (‘multiplexing’). In this way, considerablesavings in terms of time and solvent use can be achieved. Incontrast with SPE, the extraction columns are re-usable forseveral hundred injections (Zeng et al., 2004; Chassaing et al.,

2005). However, as with other automated methods, suchequipment is associated with a high capital cost which hinderswidespread uptake. A review by Xu et al. (2007) reveals many‘home-built’ systems for on-line SPE.

The use in LLE of large volumes of toxic and environmentallypolluting solvents has led to the development of manymicro-extraction techniques. Minimal volumes and a smallnumber of steps are typical. Analyte enrichment and recoveriesare often high, but the methods are difficult to automate andgenerally involve a good deal of manipulation. A few—namely restricted access materials (RAM), turbulent flow chro-matography and micro-extraction by packed sorbent (MEPS)—offer promise in terms of automation, solvent consumption, andease of use (Nováková and Vlčová, 2009). Many recent reviewsof this emerging field are available (Pedersen-Bjergaard andRasmussen, 2008; Blomberg, 2009; Cruz-Vera et al., 2009;Kataoka, 2010; Sarafraz-Yazdi and Amiri, 2010; Vuckovic et al.,2010).

More generally, analytes may be lost during hydrolysis ofsamples prior to analysis or extraction (Jourdil et al., 2003), andlosses from the adsorption of analyte onto the walls of samplecontainers should always be checked for (Verplaetse and Tytgat,2010).

Chromatographic ConsiderationsFor LC-MS, the eluent composition corresponding to optimumanalyte resolution does not always equate to that for optimal MSionization of the analytes of interest. Non-volatile buffers/eluentadditives cannot be used, and strong acids such as trifluoroaceticacid (TFA) may cause significant signal suppression in positiveionization mode through ion-pairing of TFA anions with parentions. The effect of eluent composition, additives, and adduct for-mation, on MS ionization has been reviewed extensively (Zhaoet al., 2002; Mortier et al., 2004; Kostiainen and Kauppila,2009; Table 5). The increasing demand for faster chromatographyexacerbates the problem of co-eluting matrix components,since the most severe matrix effects occur early in thechromatographic run.

Types of Column

Ultra-high pressure liquid chromatography (UHPLC) usingcolumns packed with sub-2 mm particles may shorten analysistime whilst retaining or improving chromatographic efficiency(Nguyen et al., 2006), although ultra-high pressure LC pumps arenecessary. A range of chemically modified stationary phases areavailable (Guillarme et al., 2010a). Such systems have been widelyapplied for high-throughput, targeted drug analyses. Eichhorstet al. (2009) report a semi-quantitative targeted screening analy-sis of 40 drugs/metabolites within 5.2 min, and a capacity for 200urine samples per day. Berg et al. (2009) similarly describe thequantitative analysis of a series of opiates and cocaine/cocainemetabolites within 5.7 min. Matrix effects may be reduced whenusing UHPLC compared with HPLC, as interfering matrix compo-nents are more efficiently separated from compounds of interest(Chambers et al., 2007). If UHPLC hardware is not available, super-ficially porous packing materials based on silica particles withnon-porous cores may offer similar gains in efficiency, but atcolumn pressures within the range of standard HPLC pumps(Kirkland et al., 2007; Ali et al., 2010; Fekete et al., 2009). Rust et al.(2010) used a Phenomenex Kinetex (average 2.6 mm total particle

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diameter) column for the separation of 21 benzodiazepines andthree ‘Z-Drugs’ in human hair. Monolithic columns may be usefulalternatives for fast LC, by virtue of low back pressures, allowinghigh flow-rates (Berna et al., 2004; Guillarme et al., 2010a).Pihlainen et al. (2003) used a Chromolith C18 (Merck) monolith toidentify and quantify 14 different compounds, including amphet-amines, benzodiazepines, opiates and steroids, within 5 min.

Temperature control for any LC separation is important forreproducible retention time data. This is especially true whenconsidering screening analyses in which retention time is oftenused as a criterion for compound identification (Rivier, 2003; deZeeuw, 2004). For thermally stable analytes, one may also con-sider high-temperature LC as a means of speeding up separa-tions. Using such an approach, Nguyen et al. (2007) report theseparation of nine doping agents in less than 1 min, using a sub-2 mm column at 90°C.

An increased risk of column blockage associated with sub-2mm packings is reduced by appropriate filtration of all mobilephases, and the use of in-line filters. Microbial growth in aqueoussolutions can be reduced through regular renewal, or preventedby addition of a small amount of organic solvent. The latter alsohelps to reduce air-bubble formation when high-pressure mixingis used during gradient elution. The narrow peaks generated insuch systems requires the MS cycle time/scan time to be suffi-ciently fast to ensure sufficient data points are collected acrosschromatographic peaks.

Stationary Phase Options

The direct analysis of glucuronidated and sulfated urine conju-gates of many drugs avoids the necessity for lengthy, and oftenpoorly reproducible, enzymatic or chemical hydrolysis stepsduring sample preparation (Kaushik et al., 2006). These andother similarly polar analytes often pose problems when usingtraditional reversed-phase chromatography because the highaqueous content needed for adequate retention of these com-pounds may cause ‘de-wetting’ (or phase-collapse). Moreover,for early eluting compounds, matrix effects caused byco-eluting matrix components may be more apparent. Polar-embedded phases, or packings modified to allow 100%

aqueous eluents to be used are one way to overcome this issue.An alternative approach is that of hydrophilic interaction liquidchromatography (HILIC). HILIC phases (either bare silica ormodified to contain a polar group, e.g. amide, cyano, diol orzwitterionic groups; McCalley, 2010) are now available in anumber of particle sizes, including sub-2 mm and superficiallyporous. At low aqueous eluent composition, the formation of awater-rich layer close to the stationary phase facilitates separa-tion through partitioning (hydrophilic interaction) of the ana-lytes between this layer and the organic eluent component(Alpert, 1990; Hemstrom and Irgum, 2006). HILIC was used byQuintela et al. (2010) for the analysis of cocaine and metabolites(including cocaethylene) in hair, Al-Asmari et al. (2010) andTarcomnicu et al. (2010) for the analysis of ethyl glucuronide,and Luiz Costa and Lanaro (2010) in the analysis of GHB inplasma and urine. There are a number of recent reviews andevaluations of HILIC materials (Chauve et al., 2010; Fountainet al., 2010; Jian et al., 2010; McCalley, 2010). Further, improvedMS response was reported when using HILIC compared withreversed-phase LC (Grumbach et al., 2008).

Peak tailing remains a potential problem for basic compoundson reversed-phase systems due to secondary interactions withresidual silanol groups on the silica surface, and possible solu-tions are detailed in a recent review (McCalley, 2010). We haverecently reported the application of a propylsulfonic acid-modified (strong cation-exchange, SCX) HPLC packing material(Couchman et al., 2010a) using 100% methanolic eluent forLC-MS/MS analysis of amphetamine, metamphetamine andamisulpride (Fig. 1). This simple, isocratic system is also suitablefor analysis of a range of basic drugs.

Non-silica based HPLC packings are rarely used in toxicologicalanalyses, although Stephanson et al. (2002) showed applicationof a porous graphitic carbon column (Hypercarb, Thermo Scien-tific) for the analysis of ethyl glucuronide in urine. Kanno et al.(2009) used a thermoresponsive polymeric material—poly(N-isopropylacrylamide)—with a temperature gradient elution, forthe quantitation of five barbiturates in urine.

MS is an achiral detection system. Hence, if chiral separationsare necessary (for example amphetamine stereoisomers to deter-mine pharmaceutical from ‘street’/clandestine amphetamine), LC

Table 5. Summary of LC eluent considerations for LC-MS (and MS/MS) ionization

Eluent composition• Methanol, acetonitrile and aqueous eluents are most often used for both ESI and APCI. APCI is more amenable to non-polar

solvents.• Non-volatile buffers (e.g. phosphates, borates) should be avoided. The most commonly used LC-MS eluent additives are formic

acid, acetic acid, ammonium formate, and ammonium acetate.• Eluent additives (e.g. ammonium, sodium, lithium, chloride, acetate) can produce adducts. This may be exploited, e.g. in the

analysis of immunosuppressants such as sirolimus, which readily form adducts, or for compounds which themselves do notionize readily.

• ESI is incompatible with high concentrations of eluent additives (>10 mmol/L). APCI can be used with much higherconcentrations of additive.

• Water-rich eluent may not allow for the most efficient ionization. This is a problem for early-eluting analytes in reversed phaseLC. HILIC may provide an alternative separation mechanism.

• Post-column addition of organic solvents may improve ionization efficiency (Rentsch, 2003).• Eluent pH may be manipulated to promote ionization in the eluent and hence improve ESI signal intensity.Eluent flow-rate• APCI is more amenable to high flow-rates (>1 mL/min) than ESI. ESI can give increased MS signal intensity at lower flow-rates

(0.1 mL/min or less, e.g. in capillary LC).

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separation (using chiral LC columns, Kasprzyrk-Horden et al.,2010) or off-line stereoselective derivatization (Holler et al., 2005;Guillarme et al., 2010b) can be used.

Mass SpectrometryWhen developing an LC-MS method, an important considerationis which type of atmospheric pressure ionization (API) to employ.This decision should be made based upon (i) the structure/physiochemical properties of the analyte, (ii) the LC mobile phasecomposition (and flow-rate) at the expected time of analyteelution, (iii) knowledge of any drug metabolites which may ormay not be chromatographically resolved from the analyte, and(iv) the sample preparation technique used. For the most part,the choice of ionization type will be between electrospray ioniza-

tion (ESI) and atmospheric pressure chemical ionization (APCI).Some modern instruments are supplied with dual-ionizationsources, with the ability to rapidly switch between ESI and APCIduring a chromatographic run (Waters ESCi; Gallagher et al.,2003) or even carry out both processes simultaneously (AgilentMultimode Source, Shimadzu DUIS 2010). Use of atmosphericpressure photoionization (APPI), as described by Robb et al.(2000), has not been widely reported for specific toxicologicalanalyses, though is of potential use for very non-polar com-pounds in conjunction with very low flow-rate LC separations,which cannot be efficiently ionized by APCI.

For quantitative analyses, analyte ionization must be efficientand robust. Source conditions (temperatures, source gas flow-rates and voltages) which ensure complete desolvation reducethe risk of solvent cluster formation which can occur for some

Figure 1. (a) Amphetamine, metamphetamine and amisulpride in serum after liquid–liquid extraction into butyl acetate : butanol (9 + 1, v/v); 20 mL ofthe extract was injected. HPLC: Waters Spherisorb S5SCX; 40 mmol/L ammonium acetate in methanol, pH* 6.0; flow-rate 0.5 mL/min. Detection: TSQQuantum Access (Thermo Fisher Scientific). Results: amphetamine, 2 mg/L; metamphetamine, 197 mg/L; amisulpride, 242 mg/L. (b) Amphetamine andrelated phenethylamines. HPLC conditions: as above.

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analytes. Efforts to minimize, and/or correct for, suppression orenhancement of ionization as matrix components elute from theLC column must always be made. During ‘tuning’ of the MS(Table 6), conditions should mimic, as closely as possible, thoseexpected at the time of analyte elution, including concentrationsof mobile phase additives (which may form adducts with theanalytes), mobile phase composition and flow-rate. For multi-analyte methods, tuning is often a compromise, and should betargeted towards the analyte of lowest concentration and/or ion-ization efficiency.

Electrospray Ionization

Owing to the production of ions with multiple-charges, electro-spray ionization (ESI) is useful for protein analysis and has beenapplied to the analysis of bioactive peptides (such as insulin andinsulin-like growth factor) used in doping (Thomas et al., 2010a).In the field of toxicology, for analysis of smaller molecules(around 100–1000 Da), ESI is the most widely applied ionizationtechnique. Indeed, many laboratories carrying out generalunknown screening (GUS) or systematic toxicological analyses(STAs) utilize positive mode ESI in order to identify as manycompounds as possible. It is suitable for analysis of highly polarcompounds which are ionized in solution, such as glucuronideor sulfate conjugates in urine. However, reversed-phaseLC generally results in early elution of highly polar compoundsat correspondingly high aqueous mobile phase composition,which may result in reduced desolvation (hence ionization)efficiency. To overcome this, some groups advocate post-column addition of organic solvent to improve desolvation ofearly-eluting compounds (Janda et al., 2002; Rentsch, 2003). InHILIC mode, polar compounds are eluted at high organiccompositions and thus ionize more efficiently (Nguyen andSchug, 2008).

The use of high mobile phase electrolyte concentrationsshould be avoided when using ESI, due to the risk of suppressionof ionization and contamination of the source. As a general rule,concentrations of volatile ionic modifiers should remain below10 mmol/L (Kostiainen and Kauppila, 2009).

An important advantage of ESI is that signal intensity (as afunction of the signal-to-noise ratio) is dependent not on the

absolute amount of analyte entering the source, but the concen-tration of the analyte (i.e. the concentration of analyte in theinjected volume) and the eluent flow-rate (Polettini, 2006;Watson and Sparkman, 2007). Because of this, ESI is more appli-cable, and indeed gives increased sensitivity for some applica-tions when LC separations are scaled down from 4.6 mm i.d.columns to 2.1 mm i.d. analytical columns, or from conventionalHPLC to UHPLC. Nano- and capillary-LC applications are alsoreported for the analysis of toxicologically relevant compounds.Murphy et al. (2007) report the use of capillary LC for the analysisof nicotine and cotinine in plasma of smokers. The importance ofcomplete desolvation means that ESI is generally limited to flow-rates of less than 1 mL/min. Flow-splitting is often used at higherflow-rates, since this has no effect on signal for a given concen-tration. Matrix effects are also reduced using this approach(Kloepfer et al., 2005). Heated-ESI (H-ESI) is now a further optionfrom some MS vendors, which provides improved desolvationand, therefore, capacity for increased flow-rates through additionof a heated vapourizer. The thermal stability of analytes should beconsidered when using H-ESI.

Atmospheric Pressure Chemical Ionization

With atmospheric pressure chemical ionization (APCI), ioniza-tion occurs in the gaseous phase, making this type of ionizationinherently more efficient than ESI for non-polar (hydrophobic)analytes, such as steroids, which do not readily form ions insolution (Maurer, 2007). As APCI rarely produces ions with mul-tiple charges, the achievable mass range often equates to thatof the operational range of the instrument. Since APCIrequires use of a heated vapourizer, thermally labile compoundsmay decompose in the ionization source, nullifying thebenefit of ambient or sub-ambient separation achievedwith LC. Indeed, certain compounds give rise to thermallylabile N-oxide metabolites, which decompose in-sourceback to parent compound when using APCI (Peiris et al., 2004).In quantitative analyses where an N-oxide metabolite ispresent but not chromatographically resolved from theparent compound, this can lead to over-estimation of theconcentration of the parent compound (Morgan et al., 2010;Fig. 2). Similarly, some glucuronide metabolites break down in

Table 6. Considerations for analyte optimization for LC-MS (and MS/MS)

• Optimize MS response for all analytes by infusion in eluent (including additives) rather than a different solvent.• For gradient elutions, predict (or test if possible) mobile phase composition at the appropriate elution time, and use this

mixture for analyte tuning.• Always check for presence of adducts—commonly occurring adducts include [M + Na]+ and [M + NH4]+. Check glassware and

liquid handling apparatus as sources of possible adduct formation.• Check for common in-source fragmentation (especially with APCI or H-ESI)—e.g. [M + H - H2O]+.• When optimizing analytes, whenever possible, separately infuse metabolites under the same conditions to check for in-source

conversion back to parent compound, e.g. for N-oxides and S-oxides.• In MS/MS avoid using commonly occurring fragments, such as dehydration or demethylation. Also try to avoid non-specific low

m/z fragments.• Use multiple SRM transitions whenever possible—this can help differentiate isobaric interferences, e.g. naloxone and

6-monoacetylmorphine.• For multi-analyte methods, optimize the method for the analyte for which the highest sensitivity is needed, or which ionizes

the least efficiently.

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source to form protonated aglycone pseudomolecular ions(Polettini, 2006).

Further Considerations

For most toxicological analyses using common LC mobile phasesand columns, both ESI and APCI could be used interchangeably.With regards to quantitation, Beyer et al. (2007) demonstratedvery similar accuracy and precision data when ESI was comparedwith APCI for the quantitation of a series of nine toxic alkaloids.Under the chromatographic conditions used, ESI achieved lowerlimits of detection than APCI.

Despite optimization of LC and MS source conditions, someanalytes will remain poorly ionized. In this situation, the use ofchemical derivatization to generate species more amenable toionization (for instance by addition of a proton-accepting moiety)is an option (Cech and Enke, 2001). Thieme et al. (2008) showedthat improved sensitivity could be achieved for the analysis ofbuprenorphine and norbuprenorphine in plasma with formationof an N-methylpyridinium derivative. Derivatization often servesto increase the signal-to-noise ratio for analytes of lower molecu-lar weight by increasing the observed m/z of the molecular ionspecies, to higher m/z values, where there is less interference.

Matrix Effects

The effects of biological matrix components, such as proteins,lipids (notably phospholipids), sugars and salts, as well as otherdrugs/metabolites (including commonly encountered over-the-counter medications) and mobile phase components/samplepreparation reagents on MS signal intensity are extensivelyreported and reviewed in the literature (Matuszewski et al., 1998;Fu et al., 1998; Annesley 2003; Souverain et al., 2004; Taylor, 2005;Matuszewski, 2006; Leverence et al., 2007; Ismaeil et al., 2008;Srinivas, 2009; Chambers, 2009; van Eeckhaut et al., 2009;Capiello et al., 2010; Gosetti et al., 2010; Marchi et al., 2010;Vogeser and Seger, 2010). Matrix effects may serve to increase(ion enhancement) or decrease (ion suppression) the MS signal,and can have profound effects on assay precision and accuracy inquantitative work. Concentrations of these non-detected inter-fering matrix compounds are often unknown, and highly variablebetween different samples/matrices. This degree of uncertainty isparticularly true for the complex matrices of post-mortem foren-sic samples and has not been extensively studied in alternativematrices such as hair. Especially when considering fully quantita-tive analyses, it is therefore essential (and indeed has become amandatory requirement when validating an bio-assay according

Figure 2. Clozapine, norclozapine and clozapine N-oxide by TurboFlow-LC-MS/MS (0.50 mg/L each analyte in newborn calf serum, 10 mL directlyinjected onto TurboFlow column). Columns: TurboFlow, 50 ¥ 0.5 mm Cyclone; analytical, ACE C18, 50 ¥ 2.1 mm (3 mm average particle size).

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to the US Food and Drug Administration guidelines) to evaluateand attempt to minimize the incidence of matrix effects (FDA/CDER, 2001; Peters et al., 2007). Consideration of sample collec-tion and pre-treatment, as well as MS conditions, are essential. Itis acknowledged that with regards MS conditions, ESI is moresusceptible to matrix effects compared with APCI or APPI, andthat ionization in the negative mode is more selective than thepositive mode (LeBeau et al., 2000; Annesley, 2003; Dams et al.,2003b; Schuhmacher et al., 2003; Matuszewski, 2006; Maurer2007; Flanagan et al., 2007; Smith et al., 2007). That said, matrixeffects are known to occur and should always be evaluated forAPCI methods as well (Sangster et al., 2004). Modern instrumentstend to use a spray orthogonal to the entry orifice of the massspectrometer; hence only ionized species enter the MS, meaninghigher flow-rates can be applied without significant increases innoise (Niessen, 2003; de Hoffman and Stroobant, 2007).

Two main methods exist for matrix effect evaluation, both ofwhich are widely cited. The method proposed by Bonfiglio et al.(1999) uses post-column infusion of target analytes to qualita-tively highlight regions of suppression/enhancement in a chro-matogram, whilst that proposed by Matuszewski et al. (2003)suggests a method to quantify the degree of matrix effects forparticular analytes using peak area/height ratios from analysescarried out with and without the presence of matrix compo-nents. Using both of these methods in combination allows formanipulation of chromatographic conditions and evaluation/adaptation of sample pre-treatment procedures to minimizematrix effects. For exogenous compounds, analyte-free matricesobtained from different sources are recommended for thorough

evaluation of matrix effects (as required in the FDA bio-analytical method validation guidelines; FDA/CDER, 2001).Matuszewski (2006) suggested that, for quantitative batchanalyses, quality control samples prepared in different, indepen-dent matrices from the calibration standards should beincluded.

It is well-documented that the use of stable isotope-labelledinternal standards (ISTDs) serves to compensate for matrix effects(Matuszewski, 2006; O’Halloran and Ilett, 2008; Tan et al., 2009;Marchi et al., 2010; Table 7). These isotopes should, in theory,exhibit identical behaviour to the analyte throughout the entireanalytical procedure, including ion-suppression or enhancementeffects. When the cost of these labelled compounds is prohibi-tive, for example in multi-analyte procedures, some groups havereported the use of analogous compounds as ISTDs; however inforensic cases, ingestion of the analogous compounds can neverbe unequivocally ruled out (Maurer, 2006). Moreover, recentreports of revised interest within the pharmaceutical industry indeuterium-substituted drugs may complicate the situation in thefuture. Drug analogues which are not licensed for treatment canusually be sourced from the drug manufacturers and other sup-pliers. Use of a representative ISTD for more than one analyte hasalso been reported (Liang et al., 2003; Remane et al., 2010). Whilstanalogues may compensate for some matrix effects, labelledISTDs have proved more useful and should be used whereverpossible. However, Wang et al. (2007) described an example inwhich a deuterated ISTD (racaemic [2H5]-carvedilol) interacteddifferently to the unlabelled analyte in different matrices. Thisstudy also suggested that ISTDs labelled with 13C, 15N or 17O may

Table 7. Internal standard considerations for LC-MS (and MS/MS)

• Use 13C, 15N or 18O-labelled ISTDs if possible. The ratio of the masses of these isotopes to 12C, 14N and 16O, respectively, is smallerthan that of 2H to 1H, thus they will behave more similarly to the analyte. Also, these labels are often present at sites integral tomolecular structure (e.g. 13C-enriched aromatic rings), and so are less likely than deuterated ISTDs to undergo exchangereactions. Despite this, deuterium-labelled ISTDs are the most commonly used.

• Ideally, use one ISTD for each analyte in multi-analyte methods.• In MS/MS mode, try to use fragments which retain labelled atoms where possible. It it important to know the structures of

labelled ISTDs—this may help identify the structure of fragments (e.g. for clozapine-D8, it can be deduced that fragmentationoccurs as below).

• Always evaluate matrix effects for ISTDs as well as analytes. Check whether the ISTD suppresses/enhances the analyte signaland vice versa.

• If optimizing MS response for stable isotope-labelled ISTDs, always check for the presence of unlabelled/partially labelledimpurities.

• Whilst stable isotope labelled ISTDs may appear expensive, when calculated on a cost-per-test basis, they are often notprohibitively so.

• Stable isotope labelled ISTDs are not available for all compounds. Analogues of the analyte (sometimes available frompharmaceutical manufacturers) may be useful in these instances.

• Deuterated ISTDs may chromatographically separate from their unlabelled analogues, even under achiral LC conditions(Flanagan et al., 2007).114



be better at compensating for matrix effects than deuteriumlabelled ISTDs (although they are often more costly). Likewise,Stovkis et al. (2005) and Lindegardh et al. (2008) emphasize thatisotopically-labelled standards sometimes show different recov-eries and chromatographic retention times relative to theirun-labelled analogues. There is evidence that the ionization ofsome stable isotopically labelled ISTDs is itself suppressed orenhanced by the un-labelled analogues (Liang et al., 2003; Sojoet al., 2003; Maurer, 2005; Remane et al., 2010). It is good practicewhen optimizing MS conditions for a new labelled ISTD to checkfor unlabelled (or partially-labelled) impurities, and to select afragment ion (or ions) retaining at least one isotopically-labelledatom where possible. A further consideration should be thematrix in which the ISTD is used. Addition to plasma of a relativelylarge proportion (20%, v/v) of acetonitrile containing ISTDresulted in a large number of matrix interferences when com-pared with lower proportions (Jemal et al., 2010).

The influence of the solvents used in sample preparation onmatrix effects, and as a source of alkali metals as cations foradduct formation in some assays, has been reported (Annesley,2007; Keller et al., 2008; Napoli, 2009). The volume (Liu et al., 2007)and composition of solvent mixtures (Zhang et al., 2008) used insample preparation may significantly affect the degree of matrixinterference and should be evaluated during method develop-ment. For certain analytes, where sensitivity permits, dilution ofmatrix components as far as possible serves to reduce matrixeffects (Schuhmacher et al., 2003; Kruve et al., 2009).

Mei et al. (2003) highlighted the problems which can becaused by lithium-heparin containing sample collection tubes.Formation of [M + Li]+ adducts may affect both analyte recoveryand chromatographic retention, and will lead to a reducedsignal of the [M + H]+ ion. Similarly, Chin et al. (2004) reportedreduced sensitivity to olanzapine when blood samples were col-lected in tubes containing potassium EDTA or sodium-heparin.The exact mechanism of adduct formation is not fully under-stood; however it has a significant effect on quantitative LC-MS(Medvedovici et al., 2010). Mortier et al. (2004) concluded that amajor concern is the reproducibility of adduct formation, par-ticularly due to the variation in cation (Na+, K+, NH4

+, etc.) con-centrations in biological samples, and thus adduct ions shouldnot ideally be used for quantitation. However, the use of stableisotope-labelled ISTDs was again advocated, as these shouldbehave similarly to the analyte itself with regards to adduct for-mation. However, mobile phase additives (e.g. ammoniumacetate) have been used to generate specific adduct ions forquantitation of certain compounds, notably immunosuppres-sants such as sirolimus and tacrolimus, which readily formadducts (Holt et al., 2000; Bogusz et al., 2007). Li et al. (2002)showed improved assay precision by monitoring and summingthe signals of all adducts found compared with just monitoringa single adduct. However, as noted by Nozaki et al. (2010), thismethod assumes equal ionization efficiency for all adductforms. Sodiated adducts have been shown to produce differentfragment ions than their relative protonated adducts (Nozakiet al., 2010; Medvedovici et al., 2010).

Mass Analysis

Mass analysers can be broadly classified into two groups: (i)scanning mass analysers (which only allow transmission ofsingle m/z at a time—these include quadrupoles, ion-traps andmagnetic sector instruments) and (ii) those which allow trans-

mission of ions of differing m/z simultaneously (such as time-of-flight (TOF) instruments) (de Hoffman and Stroobant, 2007).The choice of mass analyser(s) will largely be decided by theanalytical requirements and, of course, the capital cost of theinstrument.

For targeted, quantitative analyses, for instance in therapeuticdrug monitoring (TDM) and targeted drugs of abuseanalysis, LC-MS/MS is nowadays considered the method ofchoice. Triple quadrupole instruments are well-regardedas the ‘gold-standard’ for such analyses, due largely to theability to perform selective reaction monitoring (SRM)experiments. Rapid electronic control of the quadrupoles allowsfor many SRM experiments to be carried out very quickly,which is of advantage when considering accurate quantitationcoupled with the sharp chromatographic peaks achievablewith fast LC and UHPLC. A vast number of applicationshave been reported using triple quadrupole instruments inSRM mode. Reports of multi-targeted ‘screening’ proceduresusing SRM mode LC-MS/MS are available (Gergov et al.,2003; Nordgren and Beck, 2004; Nordgren et al., 2005; Eichhorstet al., 2009).

In developing quantitative, (multi-)targeted assays, it isimportant to consider the practical limitations of SRM-basedanalysis (Sauvage et al., 2008; Maurer, 2010). When just SRMscans are carried out, the absence of full-scan mass spectraldata provides information relating only to the targeted com-pound(s). With this in mind, full-scan data may be useful inevaluating the effects of co-eluting, non-isobaric matrix compo-nents. Whilst SRM transitions are highly specific, numerousexamples exist of isobaric interference from other compounds,particularly when only a single SRM transition per analyte ismonitored. Allen (2006) reported interference during the analy-sis of tramadol by LC-MS/MS arising from ingestion of the anti-depressant venlafaxine. Nordgren et al. (2005) reported a 35%false positive rate when monitoring 23 analytes using single-transition SRM mode. However, this improved significantly uponaddition of a second SRM transition. The use of multiple transi-tions for a single compound (and the ratio between the signalintensities for each) is, for this reason, commonplace (Kushniret al., 2005; Concheiro et al., 2007; Sauvage et al., 2008). Interfer-ence may also arise from metabolites, or other compoundswhich fragment or thermally degrade in-source to form isobariccompounds (Peiris et al., 2004; Vogeser and Seger, 2010). Wehave reported such an example in the LC-MS/MS analysis ofclozapine and norclozapine. Clozapine N-oxide, a minor plasmametabolite of clozapine, degrades under the APCI sourceconditions back to clozapine itself, making chromatographicresolution of these compounds essential (Morgan et al., 2010).SRM-mode, and the application of product ion ratios, is obvi-ously limited for compounds which do not fragment reproduc-ibly. A well-documented example is that of buprenorphine andnorbuprenorphine. Whilst many laboratories overcome thisissue by monitoring the non-fragmented pseudomolecular ionin quadrupole 3 (Q3), Ceccato et al. (2003) report that, by usingcollision pressure/energy at a level just below that which causescomplete fragmentation of the precursor ion, isobaric interfer-ences could be fragmented, thus increasing the observedsignal-to-noise ratio.

To minimize the risk of interference, multiple transitions(ideally avoiding non-specific transitions such as water loss(es) orlow molecular weight fragments) should always be used wherepossible. For compounds which do not fragment well, or have

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only one major fragment, extra steps should be taken to ensureselectivity during sample preparation and LC separation(Sauvage et al., 2008).

Systematic Toxicological Analysis/GeneralUnknown Screening

In forensic and post-mortem toxicology, systematic toxicologicalanalysis (STA)/general unknown screening (GUS) is the startingpoint from which further, targeted quantitative analyses follows(Flanagan et al., 2007), and aims to sensitively and reliably detectas wide a range of compounds as possible. It is the most crucialstage in forensic and post-mortem toxicological analysis, and tobe as comprehensive as possible is a prerequisite. For many yearsGC-MS, despite the problems associated with larger, non-volatileand thermally labile compounds, was considered the best strat-egy for these analyses. The reproducibility of GC-MS ionization/fragmentation allowed for the development of comprehensivemass spectra libraries for reliable structural identification. The lackof equivalent LC-MS based spectral libraries, which prevents uni-versal adoption of this technique, is due to ‘soft’ ionizationachieved with API (compared with GC-MS ionization), and also thepoor inter-instrument reproducibility of LC-MS fragmentation(Marquet, 2002; Kushnir et al., 2005; Jansen et al. 2005; Maurer andPeters, 2005;Yadav et al., 2008). Some groups however, report thatreproducibility can be achieved with standardization of instru-ment settings (Bristow et al., 2004; Gergov et al., 2004; Hopleyet al., 2008; Oberacher et al., 2009). Early STA methods usingsingle-stage LC-MS (usually quadrupole) instruments, usingin-source collision-induced dissociation (CID) to generate production mass spectra showed some promise, and users began to buildconsiderable ‘in-house’ spectral libraries of their own (Marquetet al., 2000; Hough et al., 2000; Lips et al., 2001; Saint-Marcouxet al., 2003; Venisse et al., 2003). However, LC-MS/MS (triple qua-drupole or hybrid quadrupole-ion trap instruments) withinformation- or data-dependant acquisition is fast becoming con-sidered a better way to perform STAs. Improved instrument scanrates and collision cell functions (for example collisional energyramping and spectra averaging, enhanced product-ion scans, andreduction of space-charging effects) make for more product rich,reproducible mass spectra, which are then applicable to‘in-house’reference libraries/databases (Marquet et al., 2003; Saint-Marcouxet al., 2003; Mueller et al., 2005; Dresen et al., 2006, 2009, 2010;Sauvage et al., 2006; Politi et al., 2007; Liu et al., 2009, 2010; Lynchet al., 2010; Maurer, 2010).

An emerging approach to STA analysis is that of accurate mass(or exact mass, high-resolution) MS. By carrying out full-scan MSexperiments at high mass accuracy (m/z up to 5 decimal places), itis possible to very precisely filter the full-scan data and extractanalyte chromatograms with very low background noise. In thisway, one can distinguish between compounds which have thesame nominal, but different exact masses (e.g. benzoylecgonineand atropine, flecainide and diltiazem). High-resolution MS(HRMS) can be performed, as with quadrupoles and ion-traps, aseither single-stage MS (Gergov et al., 2001; Pelander et al., 2003,2008, 2010; Ojanperä et al., 2005; Polettini et al., 2008; Lee et al.,2009) or as MS/MS (quadrupole-TOF, Q-TOF), allowing HRMSproduct ion scan data, with information-dependant acquisition(Pavlic et al., 2006; Decaestecker et al., 2004; Peters et al., 2010).TOF-HRMS is of particular interest in the application of empiricalformula-based data libraries, with isotope pattern-matching soft-ware, and the potential to screen for unknown compounds (and

identify their metabolites) by knowledge of elemental composi-tion alone, without the absolute need for reference material (Ojan-perä et al., 2006). Exact mass identification of specific metabolites(Liotta et al., 2010) and systematic fragmentation (Q-TOF-MS)approaches (Tyrkko et al., 2010) have shown that even structuralisomers can be distinguished using accurate mass. Further, retro-spective interrogation of full-scan data can be useful to investi-gate the presence of new compounds/metabolites (such as new‘designer drugs’) without re-analysis (Peters et al., 2010). NewerOrbitrap®/Exactive™ technology (ThermoFisher Scientific), alsocapable of HRMS, is finding some toxicologically relevant applica-tions (Thomas et al., 2010b; Weider et al., 2010; Johnson andKozak, 2010) and may be a useful tool for the future.

Where possible, for unequivocal compound identification,library matching should be carried out against databases pro-duced ‘in-house’. New mass analysis technologies should beexploited whenever possible, in order to improve the product-ion spectra data obtained, and thus create a more useful refer-ence library. Mass spectra libraries and compound formulaedatabases from elsewhere may not be completely transferablebetween instruments (certainly with respect to product/fragment ion ratios; Jansen et al., 2005) and library data will notgive any indication of chromatographic retention time, which is auseful criterion for absolute identification (Rivier, 2003; Mara-likova and Weinmann, 2004; de Zeeuw, 2004; Fox et al., 2009).Lynch et al. (2010) highlight the importance of manual interroga-tion of mass spectral data alongside automated library searchingsoftware to avoid false identifications.

Logistical Considerations

Vogeser and Seger (2008) highlighted the considerable logisticalrequirements for installation of LC-MS instrumentation withregards to noise and space. A further important considerationrelating to installation is that of ambient operating temperature.Many instruments are designed to operate optimally within apre-determined temperature range. Exceeding these parametersmay cause analytical problems, notably mass calibration drift.Placing instruments below air-conditioning vents, which mayresult in significant temperature variation throughout the day,should therefore be avoided.

Method Validation and Laboratory Accreditation

The importance of analytical toxicology results necessitates reli-able, accurate and often legally defensible results. A number ofuseful reports, official guidelines and reviews exist on therequirements and practicalities of bioanalytical method valida-tion (Wood, 1999; Shah et al., 2000; FDA/CDER, 2001; VanderHeyden et al., 2001; Thompson et al., 2002; Taverniers et al., 2004;SOFT/AAFS, 2006; Viswanathan et al., 2007; EMEA/CHMP, 2009)and how these guidelines are best applied to clinical and forensicanalytical toxicology (Peters and Maurer, 2002; Peters et al., 2007).The more recent of these guidelines/reviews highlight the needfor matrix effect evaluation with LC-MS methods. For qualitativeanalyses, sensitivity (‘the ability to detect truly positive samplesas positive’), specificity (‘the ability to detect truly negativesamples as negative’) and limit of detection (LOD) should beascertained at the very least, plus ideally investigations intoanalyte recovery, assay precision and robustness (Trullols et al.,2004; Peters et al., 2007). Forensic laboratory accreditation isbased upon international standards (ISO/IEC 17025, 2005) whichinclude requirements for method validation. The Society of

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Forensic Toxicologists (SOFT) and the Toxicology Section of theAmerican Academy of Forensic Sciences (AAFS) have issueduseful guidelines for preparation of a laboratory for accreditationinspection (SOFT/AAFS, 2006).

ConclusionsLC-MS and LC-MS/MS combine the versatility of HPLC with thesensitivity and selectivity of MS detection. This overcomes thelimitations associated with GC-MS when analysing certain polarand non-volatile compounds, many of which are of significancein toxicological investigations, but brings other problems.Advances in interface technology, and a better understanding ofthe complex physiochemical processes occurring during ioniza-tion, mean that LC-MS is becoming commonplace for routine,high-throughput, and specialist toxicological investigations. Withregards to matrix effects, the importance of sample preparationand chromatographic separation cannot be over-emphasized. Anumber of approaches to minimizing matrix effects are available,and should form the basis of thorough method development.

AcknowledgementsThe authors wish to thank Victoria Lay (Department of Chemistry,Loughborough University, UK) and Professor Robert Flanagan(King’s College Hospital, UK) for valuable assistance and sugges-tions, and also Dr Chang Kee Lim for the kind invitation to con-tribute to this Special Issue of Biomedical Chromatography.

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