GC/MS ASSAYS

FOR ABUSED

DRUGS IN

BODY FLUIDS

U.S. DEPARTMENT OF HEALTH AND HUMAN SERVICES • Public Health Service • Alchol, Drug Abuse, and Mental Health Administration
GC/MS Assays

for Abused Drugs

in Body Fluids

Rodger L. Foltz, Ph.D.

Center for Human Toxicology

University of Utah

Salt Lake City, Utah 64112

Allison F. Fentiman, Jr., Ph.D.

Ruth B. Foltz

Battelle Columbus Laboratories

Columbus, Ohio 43201

NIDA Research Monograph 32

August 1980

DEPARTMENT OF HEALTH AND HUMAN SERVICES

Public Health Service

Alcohol, Drug Abuse, and Mental Health Administration

National Institute on Drug Abuse

Division of Research

5600 Fishers Lane

Rockville, Maryland 20857

For sale by the Superintendent of Documents, U.S. Government Printing Office

Washington, D.C. 20402
The NIDA Research Monograph series is prepared by the Division of Research

of the National Institute on Drug Abuse. Its primary objective is to provide

critical reviews of research problem areas and techniques, the content of

state-of-the- art conferences, integrative research reviews and significant

original research. Its dual publication emphasis is rapid and targeted

dissemination to the scientific and professional community.

Editorial Advisory Board

Avram Goldstein, M.D.

Addiction Research Foundation

Palo Alto, California

Jerome Jaffe, M.D.

College of Physicians and Surgeons

Columbia University, New York

Reese T. Jones, M.D.

Langley Porter Neuropsychiatric Institute

University of California

San Francisco, California 

William McGlothlin, Ph.D.

Department of Psychology, UCLA

Los Angeles, California

Jack Mendelson, M.D.

Alchol and Drug Abuse Research Center

Harvard Medical School

Mclean Hospital

Belmont, Massachusetts

Helen Nowlis, Ph.D.

Office of Drug Education, DHHS

Washington, D.C.

Lee Robins, Ph.D.

Washington University School of Medicine

St. Louis, Missouri

NIDA Research Monograph series

William Pollin, M.D.

DIRECTOR, NIDA

Marvin Snyder, Ph.D.

DIRECTOR, DIVISION OF RESEARCH, NIDA

Robert C. Petersen, Ph.D.

EDITOR-IN-CHIEF

Eleanor W. Waldrop

MANAGING EDITOR

Parklawn Building, 5600 Fishers Lane, Rockville, Maryland 20857
GC/MS Assays

for Abused Drugs

in Body Fluids
ACKNOWLEDGMENTS

Support for the development and evaluation of the GC/MS

assays described in this monograph and the preparation

of the manuscript was provided by the National Institute

on Drug Abuse, contracts ADM 45-74-140, HSM 42-72-183,

and 271-77-3419 to Battelle Columbus Laboratories, and

contract 271-80-3719 to the Center for Human Toxicology,

and by the Insurance Institute for Highway Safety, project 6762 at the Center for Human Toxicology.

The United States Government does not endorse or

favor any specific commercial product, commodity, or

supplier. Trade or proprietary names and names of

suppliers appearing in this publication are used only

because they are considered essential in the context

of the information presented.

Library of Congress catalog card number 80-600143

DHHS publication number (ADM) 80-1014

Printed 1980

NIDA Research Monographs are indexed in the Index Medicus.

They are selectively included in the coverage of Biosciences

Information Service, Chemical Abstracts, Current Contents,

Psychological Abstracts, and Psychopharmocology abstracts.

i v
Foreword

This monograph presents a collection of methods for the quantitative analysis of several important drugs of abuse by the technique of gas chromatography-mass spectrometry (GC/MS). The material is arranged textbook fashion, by chapters on specific

drugs, to assist researchers in setting up GC/MS assay procedures

in their own laboratories.

These assays are particularly suited to basic studies on the

pharmacology and pharmacokinetics of drugs of abuse. Forensic

toxicology, which sometimes requires the highly sensitive and

specific sample analysis provided by GC/MS, is another important

area of application. Even when an alternate method such as immunoassay is available for such analyses, GC/MS is often the only

means of providing a confirmation assay. The sensitivity and

specificity of GC/MS are especially critical to studies on the

impact of drugs on traffic safety. In addition, demand for legislation relating to drugs and highway safety is increasing, and

enforcement of such laws will necessitate appropriate sample assays.

The unique characteristics of the GC/MS analytical method make it

the only means of determining drug levels under certain circumstances. The inherent complexity of the technique, however, often

dissuades investigators from embarking on studies that involve

its use. At a time when basic questions underlying drug abuse

call for increasingly sophisticated methods of data acquisition,

this monograph should provide the information needed to make

one of these procedures easier to establish and to encourage new

areas of research.

Marvin Snyder, Ph. D.

Director, Division of Research

National Institute on Drug Abuse

 v

Preface

Soon after the establishment of the National Institute on Drug

Abuse in 1974, the development of an analytical program emphasizing gas chromatography/mass spectrometry (GC/MS) was initiated.

A major part of this program was directed at the development,

testing, and eventual publication of GC/MS methods suitable for a

number of abused drugs of interest to this Institute. It was

felt that by making available carefully described quantitative procedures for these drugs, more researchers would be encouraged to

move into several critically important areas of investigation.

A major segment of this development and testing program was carried

out by Battelle (Columbus Laboratories and at the Center for Human

Toxicology at the University of Utah. These GC/MS methods emphasize chemical ionization techniques which frequently offer significant advantages in sensitivity and specificity over techniques of

electron impact ionization.

Each of the techniques described in this monograph has been field

tested by one or more independent laboratories in order to optimize

both the procedure itself and the clarity of its presentation.

This field testing was carried out by Battelle and consisted of

sending the participating laboratories a series of known standard

samples along with the information brochures included herein as

chapters. The participating laboratories then set up the method

in their own facilities and analyzed the spiked samples. Problems

with either the instructional material or the method allowed

Battelle to make many improvements to insure the highest probability

of an inexperienced lab’s success with one of these assays.

This monograph represents the combined efforts of many individuals

and has evolved to its present state over several years. We hope

that it will save research investigators valuable time in establishing quantitative essays of these drugs in their own laboratories.

The basic principles of GC/MS quantitative analysis as presented

here should be useful in setting up procedures for other drugs as

well.

Richard L. Hawks , Ph.D.

Division of Research

vii

Contents

Foreword . . . . . . . . . . . . . . . . . . . . . . . . . . . . v

Marvin Snyder

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . vii

Richard L. Hawks

Chapter 1

Chapter 2

Chapter 3

Chapter 4

Chapter 5

Chapter 6

Chapter 7

Chapter 8

Chapter 9

Chapter 10

Chapter 11

Chapter 12

Chapter 13

Introduction . . . . . . . . . . . . . . . . . . . . 1

Experimental Considerations and Operations Common

to All of the Assays . . . . . . . . . . . . . . . . 5

Phencyclidine(PCP) . . . . . . . . . . . . . . . .25

Methaqualone . . . . . . . . . . . . . . . . . . . .39

Methadone . . . . . . . . . . . . . . . . . . . . .51

-Tetrahydrocannabinol (THC) and Two of Its

Metabolites, 11-Hydroxy- -THC and 11-Nor-9-Carboxy-

-THC . . . . . . . . . . . . . . . . . . . . . . .62

Cocaine and Its Major Metabolite, Benzoylecgonine . 90

Morphine . . . . . . . . , . . . . . . . . . . . . 110

Diazepam and Its Major Metabolite, N-Desmethyldiazepam . . . . . . . . . . . . . . . . . . . . . 128

Amphetamine . . . . . . . . . . . . . . . . . . . 150

Methamphetamine . . . . . . . . . . . . . . . . . 165

2,5-Dimethoxy-4-Methylamphetamine (DOM) . . . . . 179

Mescaline . . . . . . . , . . . . . . . . . . . . 189

i x

Chapter 1

Introduction

The combination of gas chromatography and mass spectrometry (GC/MS)

became widely recognized in the 1960’s as the most sensitive and

versatile tool available for the identification of volatile organic compounds. It was not until the 1970’s, however, and the

development of selected ion monitoring techniques, that the potential of GC/MS for quantitative analysis was generally appreciated.

It now seems certain that the quantitative measurement of specific

organic compounds in complex mixtures will become the major application of GC/MS simply because its sensitivity, accuracy, and versatility are unequalled by any other technique, including the popular immunological assays.

The rapid increase in the use of GC/MS has occurred in spite of

several severe limitations and deterrents. The major limitation

is that for a compound to be analyzed by GC/MS it must have sufficient volatility and thermostability to pass through the gas chromatographic column intact in the vapor state, or be capable of conversion to a derivative which can do so; this requirement prohibits

GC/MS analysis for 80 to 90 percent of known organic compounds.

Fortunately, this is not a serious problem for the toxicologist,

because most drugs and their metabolites are among the structures

which can be analyzed by GC/MS. More serious deterrents to wider

use of GC/MS in toxicological and clinical laboratories are the

expense of GC/MS instrumentation and the relatively high level of

training and skill required to make effective use of these instruments. Although the former is unlikely to be reduced in the future, the latter will undoubtedly become less burdensome as the

reliability of GC/MS instrumentation is improved and its design

simplified.

The purpose of this monograph is to assemble, within a single volume, detailed descriptions of GC/MS procedures for quantitative

measurement of some of the drugs and their metabolites which are

of particular concern to the National Institute on Drug Abuse

(NIDA). The procedures were developed at Battelle Colunbus Laboratories and the Center for Human Toxicology at the University of

Utah, and have been extensively evaluated at both of these laboratories. All of the assays make use of deuterium-labeled internal

standards and packed column gas chromatography combined with chemical ionization mass spectrometry. This combination was chosen for

the following reasons:

Stable isotope labeled analogs constitute nearly ideal

internal standards because their chemical and physical

properties are nearly identical to those of the respective unlabeled compounds.

1
Most analytical laboratories with GC/MS systems are

more familiar with the use of packed columns than

with capillary columns.

Chemical ionization mass spectrometry, using the

technique of selected ion monitoring, is the most

sensitive method of detection available. With chemical ionization most drugs are efficiently ionized,

the sample ion current is concentrated at one or two

m/z values, and the likelihood of interference from

ions generated by column bleed or components of the

biological extract is minimized.

Each drug was originally the subject of a separate analytical

manual. A decision was subsequently made, however, to combine the

manuals into a single volume, with each chapter devoted to a specific drug and its metabolites. As a result of this evolution

of manuscripts, each chapter is relatively self-contained and includes its own list of literature references and its own number

sequence for figures, tables, and structural drawings. For this

reason, anyone who has the tenacity to read the monograph from

cover to cover will find excessive repetition of certain descriptions and explanations. Most readers, however, will not be concerned with all of the drugs included in the monograph, and will

choose to read only selected chapters. Nevertheless, in order to

keep repetition to a minimum, many of the considerations and operations which are common to all of the assays have been removed

from the individual chapters and combined in Chapter 2. We therefore recommend that the reader begin with Chapter 2 and only then

proceed to the chapters which deal with the drugs of particular

interest.

On the premise that an analyst who must determine the concentration

of a drug in biological specimens should know something about the

drug, we have included in each chapter background information which

we believe to be relevant and useful. This background includes:

a brief history of the drug and its uses

a summary of the pharmacology and metabolism of the

drug

a survey of literature procedures for the quantitative

analysis of the drug and its metabolites, with particular emphasis on other GC/MS assays for the drug, or

assays which may be considered as alternatives to GC/

MS analysis

a compilation of physical, chemical and spectroscopic

data on the drug

Although GC/MS instruments are now prominent fixtures in many clinical and toxicological laboratories, too often their use is limited

to the identification of unknowns by matching their electron impact

2
mass spectra to reference spectra. We hope that this monograph

will encourage more analysts to use the power of GC/MS as a quantitative technique. It is important, of course, to recognize when

to use GC/MS for a particular drug analysis and when to use some

other technique. The assays described in this monograph are intended to be used either as reference methods for validation and

quality control of other methods, or as the technique of choice

when other available methods do not offer adequate sensitivity,

specificity or accuracy. The latter situation is often a factor

in forensic toxicology, because the outcome of court litigation may

depend upon the reliability of a drug assay.

GC/MS can also reduce the cost of implementing a new drug assay.

On several occasions we have attempted to use a “less expensive”

technique for a particular drug assay, only to find that the assay

did not have adequate sensitivity or specificity. Each time we

resolved the problem by using GC/MS, but valuable time and money

were expended.

Another reason why GC/MS can often be economic relative to other

chromatographic techniques is that its specificity frequently permits use of simple and rapid extraction procedures. Not only does

a simple extraction procedure lower cost by saving time, but it

also reduces the opportunity for human error. The degree to which

an extraction procedure can be simplified is dependent, however,

upon the level of sensitivity required. For example, tetrahydrocannabinol is psychoactive at blood concentrations of 10 ng/ml,

and less. At these low concentrations a multi-step extraction and

purification procedure is required in order to eliminate interference by endogenous components in the biological medium. In contrast, pharmacologically significant blood concentrations of a drug

such as methaqualone are much higher (> 100 ng/ml); and therefore a

simple, direct extraction of bases without further purification or

concentration is adequate for GC/MS analysis, provided the chromatographic and mass spectrometric conditions are chosen properly.

The exclusive use of chemical ionization (CI) in each of the drug

as says, rather than electron impact (EI) ionization, deserves comment. In qualitative analysis the major advantage of CI is that it

normally generates a prominent peak from which the molecular weight

of a compound can be calculated. In quantitative analysis of drugs

it offers two advantages over EI ionization: 1) the ion current of

the analyte is usually concentrated at one or two compound-characteristic masses which are particularly suitable for selected ion

monitoring, and 2) most drugs and their metabolites can be efficiently ionized by a selective reagent gas such as ammonia. These features usually permit better effective sensitivity with CI than with

EI ionization. Nevertheless, analysts in many laboratories have

been reluctant to use chemical ionization. Part of this reluctance

undoubtedly reflects a lack of familiarity with CI. However, it

is also true that there arc more experimental variables in CI than

in EI ionization, and the task of optimizing and controlling these

variables may discourage some analysts. For this reason we have

included in Chapter 2, and in the discussions of each of the drug

assay procedures, suggestions and guidelines for use of CI.

3
A number of newer chromatographic and mass spectrometric techniques

are in various stages of development and evaluation for analysis

of drugs. Glass capillary columns are rapidly increasing in popularity and will continue to be used in place of packed columns

whenevcr greater separating power and higher sensitivity are needed.

The commercial availability of combined liquid chromatograph/mass

spectrometer (LC/MS) systems makes it feasible to analyze polar

drugs and metabolites directly, without derivatization. The new

technique of MS/MS, in which a mass analyzer is used in place of

a gas chromatograph to separate the components of a mixture prior

to identification or quantitation by mass spectrometry, may ultimately supplant GC/MS where large numbers of samples must be analyzed. Finally, negative ion chemical ionization has been shown

to be capable of dramatically higher sensitivity than positive

ion CI for the GC/MS analysis of compounds which have high electron

affinities. The application of negative ion CI to the analysis of

diazepam and N-desmethyldiazepam is discussed in Chapter 9.

None of these newer techniques are employed in the GC/MS drug assays

in this monograph, primarily because at the time the assays were

developed techniques such as LC/MS, negative ion CI, and MS/MS

were in the early stages of development and were available at only

a few research laboratories. It is now evident, however, that each

of these techniques offers exciting capabilities that will uniquely

satisfy specific analytical needs. It is likely that by the time

this monograph is distributed, many readers will already have had

experience with one or more of the techniques.

The authors gratefully acknowledge the contributions of the many

individuals and institutions that have assisted in the development

of these assays and the preparation of this monograph. First and

foremost, we thank the National Institute on Drug Abuse (NIDA) for

financial support. We especially appreciated the guidance, encouragement, and many helpful discussions with the Project Monitors, Drs.

Richard L. Hawks and Robert E. Willette. Additionally, we would

like to thank the Insurance Institute for Highway Safety for their

financial support which enabled us to further evaluate and refine

several of these assays. Those who contributed to the laboratory

research include Dr. David A. Knowlton, Dr. Pauline A. Clarke, Dr.

Denis C.K. Lin, Dr. Edith G. Leighty, Dr. Bruce A. Petersen, Mr.

Warren E. Bresler, Mr. Bruce J. Hidy, and Dr. Frank W. Crow at

Battelle Columbus Laboratories, and Mr. Dennis M. Chinn and bliss

Cheryl L. Jackson at the Center for Human Toxicology. Others who

helped evaluate the methods include: Dr. Dan S. Pearce, Laboratory

of Criminalistics, Santa Ana, CA; Dr. Jack D. Henion, New York State

College of Veterinary Medicine, Cornell University, Ithaca, NY; Dr.

James Ferguson, Toxicology Department, The Ohio State University,

Columbus Ohio; Dr. Daniel R. DiFeo, Jr., Analytical Chemistry Center,

The University of Texas Health Science Center, Houston, Texas;

and Miss Pamela Ahearn, State of Rhode Island Department of Health,

Providence, Rhode Island.

Finally, special thanks are due to Dr. Bryan S. Finkle at the Center

for Human Toxicology for his invaluable assistance in preparing

and reviewing the monograph manuscript.

4
Chapter 2

Experimental Considerations

and Operations

Common to All of the Assays

Each of the GC/MS assays described in this monograph includes the

following steps:

1. Addition of a deuterium labeled internal standard to the

biological specimen

2. Solvent extraction of the biological specimen

3. GC/MS analysis of the extract using chemical ionizntion

and selected ion monitoring

4. Calculation of the drug concentration in the biological

sample based on the ratio of ion currents generated by

the drug and the internal standard.

This chapter discusses general considerations relevant to each of

these operations, and presents our recommendations regarding the

preparation of calibration graphs and procedures for monitoring

the quality of the analytical data.

Even though all of the assays utilize chemical ionization (CI) , the

procedures can be modified for electron impact (EI) ionization. ‘Ibis

change will usually result in lower sensitivity and specificity because the most abundant ions in the EI mass spectra of these drugs

are generally not as satisfactory for selected ion monitoring as

the abundant ions in the chemical ionization mass spectra (1).

A more rigid requirement of these assays is that the GC/MS system

must be capable of continuously monitoring the ion currents at two

or more selected ion masses. The usefulness of selected ion monitoring (also called “multiple ion detection” and “mass fragmentography”) is now so widely recognized that virtually all GC/MS systems intended for biomedical applications have this capability,

either in the form of an interactive data system or a hardwired

selected ion monitor.

TYPES AND SOURCES OF SPECIMENS

Because the blood concentration of a drug usually gives the best correlation with the pharmacological effect of the drug,

the preferred biological specimens for drug analysis are whole blood,

plasma, or serum. All of the GC/MS assays in this monograph are

5
capable of measuring drug concentrations in blood at least as low

as the minimum level at which the drug exerts a detectable physiological effect.

If a choice of specimen is available, plasma is usually preferred.

The blood should be collected in evacuated tubes containing sodium

fluoride to inhibit bacterial and enzymatic action, and sodium oxalate or heparin to prevent coagulation. The blood should be centrifuged as soon as possible to separate the plasma from the red blood

cells, and the plasma transferred into silylated glass tubes equipped

with Teflon-lined screw caps. If storage or shipment of the plasma

is necessary, it should be frozen and maintained in the frozen state

until the extraction can be performed. Several guaranteed 24-hour

parcel delivery services are now available to most cities in the

United States. These services are well worth the additional cost

for shipment of biological specimens.

The GC/MS assays described here are also applicable to other physiological fluids and tissues, including urine, saliva, cerebrospinal

fluid, gastric fluid, and homogenates from adipose tissue, liver,

kidney, etc. Some specimens, particularly those with a high fat

content, may require additional purification in order to remove interfering endogenous compounds. This can be accomplished in the

case of basic drugs by a “back extraction” procedure in which the

drug is extracted from an organic solvent into aqueous acid. The

acid solution is then made basic and the drug reextracted into an

organic solvent.

Tetrahydrocannabinol (Chapter 6) is unique among the drugs discussed

in this monograph in that it is a highly lipophilic, nonbasic drug,

and therefore difficult to separate from the lipids present in physiological specimens. For this reason, the THC assay is a particularly striking example of the power of GC/MS to selectively measure a specific compound within a complex mixture of similar materials.

INTERNAL STANDARDS

The sensitivity of a GC/MS system depends on a large number of factors, some of which are difficult to control. Consequently, it is

essential to use an internal standard if accurate, quantitative measurements are to be made. An ideal internal standard behaves identically to the analyte throughout the extraction, chromatographic separation, and ionization processes, so that the weight ratio of added internal standard to analyte will remain unaffected by any of

these operations. Stable isotope labeled analogs come the closest

to meeting the characteristics of ideal internal standards. The isotope label exerts a very slight, but usually insignificant effect on

the physical and chemical properties of the compound, yet the higher

mass of the isotope labeled ions of the internal standard permits

them to be separated from the analyte ions by the mass spectrometer.

The major deterrent to the use of stable isotope labeled drugs as internal standards is their limited availability and the high cost of

those that are available. In recognition of this problem, the

6
National Institute on Drug Abuse has contracted the synthesis of

stable isotope labeled analogs of many of the abused drugs and their

metabolites. Milligram quantities of these compounds are available

without charge to authorized investigators engaged in research requiring the quantitative analysis of these drugs. Enquiries regarding the availability of specific isotope labeled drugs and metabolites should be sent to:

National Institute on Drug Abuse

Division of Research/Research Technology Branch

5600 Fishers Lane

Rockville, MD 20857

301-443-5280

Several commercial firms specialize in the synthesis and sale of

stable isotope labeled compounds. These include:

Merck Sharp Dohme Canada Limited

Isotope Division

P.O. Box 899

Pointe Claire-Dorval, Quebec H9R 4P7

Canada

514-697-2823

KOR Isotopes

56 Rogers Street

Cambridge, MA 02142 USA

517-661-8220

Prochem

19, Ox Bow Lane

Summit, NJ 07901 USA

201-273-0440

Chromacol Ltd.

73 a Friern Barnet Lane

London N20 OXT England

368- 7666

Stohler Isotope Chemicals, Inc.

49 Jones Road

Waltham, MA 02154 USA

617-891-1827

Unfortunately, the problems associated with selling controlled substances have discouraged some of the suppliers from any involvement

with this category of drugs. Merck Sharp Dohme Canada synthesizes

and sells stable isotope labeled controlled drugs, but purchasers

in the United States must obtain an American Government Permit to

Import before Merck can ship the product.

Because the market for stable isotope labeled compounds is still

relatively small, their cost is high when purchased from commercial

sources. For this reason laboratories with organic synthesis expertise may elect to synthesize their own isotope labeled internal

7

328-011 0 - 80 - 2
standards. Experimental procedures have been published for preparation of many deuterium labeled drugs and metabolites (2,3).

Several factors arc important to keep in mind when selecting an

isotope labeled compound for use as an internal standard. Most

importantly, the isotope must not undergo exchange under any of

the conditions encountered during the extraction, derivatization,

or chromatography steps of the assay, nor those within the ion

source of the mass spectrometer. The replacement of a deuterium

label by hydrogen within the ion source sometimes occurs unexpectedly under chemical ionization conditions. For example, a drug

labeled with three dcuterium atoms attached to a tertiary amino

methyl group was found to undergo extensive exchange between deuterium and hydrogen atoms when the drug was subjected to methane

chemical ionization (3). Fortunately, the exchange did not occur

when ammonia was used as the reagent gas.

Additional potential problems associated with the use of deuterium

labeled analogs as internal standards are discussed in Millard’s

book entitled Quantitative Mass Spectrometry (5). In particular,

an example is cited in which a prostaglandin was labeled with four

deuterium atoms in apparently stable positions in the molecule, but

after prolonged storage in methanol considerable exchange of hydrogen for deuterium was found to have occurred. Based upon this experience, Millard recommends that stock solutions of deuterium

labeled compounds should be checked at regular intervals for evidence of hydrogen-dcutcrium exchange.

The problem of exchange has prompted some analysts to utilize 13C

or 

15N labels rather than dcuterium. However, deuterium is usually

far easier to incorporate into a molecule than other heavy isotopes

and therefore will undoubtedly continue to be the most frequently

used label for internal standards. The important point is that

the analyst must be aware of the possible problems associated with

the use of deuterium labeled internal standards, and to thoroughly

evaluate each application of a particular internal standard hefore embarking on an extensive series of assays.

A second important point regarding isotope labeling is that the

isotopic variant should have a molecular weight three or more mass

units greater than the unlabeled compound. This is because the

naturally occurring heavy isotope content (primarily 13C) of organic

compounds gives rise to ions of significant intensity at one and

two mass units above each carbon containing ion in the compound’s

mass spectrum. In principle the contributions of these naturally

occurring isotope ions can be calculated and subtracted from the

ion currents resulting from the internal standard, hut in practice

measurement of the intensity of the isotope labeled ion peak is

easier and more accurate if there is no contribution from other ions.

Of course, other types of compounds can bc used as internal standards when a suitable isotope labeled analog is not available.

For example, homologs and other structurally similar compounds have

been used as internal standards and often give satisfactory results

(6-8). Nevertheless, when they can be obtained, stable isotope

8
labeled analogs are the first choice for internal standards of most

practitioners of quantitative mass spectrometry (9).

PREPARATION OF CALIBRATION GRAPHS

The procedure we recommend for preparation of calibration graphs

(“standard curves”) involves addition of a fixed quantity of internal standard to aliquots of an appropriate body fluid containing

known quantities of the analyte. The “spiked standards” are then

extracted, derivatized, and analyzed by exactly the procedure to

be used for unknown samples. The calibration graph is constructed

by plotting the measured ratio of ion current peak heights corresponding to the analyte and the internal standard versus the quantity of analyte added to each sample. Any inaccuracies in measurement

that occur in the preparation of a calibration graph will affect

the accuracy of all subsequent quantitative determinations which

utilize that calibration graph. Consequently, preparation of an

accurate calibration graph is extremely important. The two requirements that are most critical are: 1) the quantity of analyte in

each sample must be accurately known, and 2) exactly the same quantity of internal standard must be added to each sample used to prepare the calibration graph as is added to each of the unknown samples.

As a general rule the sample medium used for preparing a calibration

graph should be as similar as possible to the sample medium to be

analyzed. If the unknowns consist of plasma samples, the biological standards to be used in establishing the calibration graph can

be prepared by adding precise volumes of standard solutions of the

analyte to aliquots of a pool of plasma obtained from a local blood

bank. It is important, of course, to establish that the plasma

pool is completely free of the analyte, or any other compounds which

interfere with the assay.

Ideally, the analyte standard solution and the internal standard

solution should be added in such a way that the biological medium

is not changed and drug compounds are introduced into a molecular

environment which exactly mimics that of the biological specimens

to be analyzed. For this reason it is generally preferable to use

aqueous standard solutions. However, some drugs have very little

solubility in water so that it is difficult to prepare aqueous solutions at the concentrations required. Most drugs are soluble in

methanol, and addition of small quantities of methanolic solutions

to biological media causes relatively minor physical and chemical

alteration. For the sake of uniformity the standard solutions for

all of the drugs and metabolites in this monograph are prepared in

methanol. We have also chosen 100 μl as the volume of the methanolic standard solutions to be added to 1-ml volumes of the biological specimen. This volume of methanolic solution is a compromise

in that it is large enough to be conveniently dispensed with good

accuracy, but is not so large that it causes extensive protein precipitation.

9
As previously stated it is important to add precisely the same

amount of internal standard to each sample to be analyzed, i.e.,

to both standards and unknowns. It is not actually necessary to

know the weight of internal standard added, as long as it is exactly the same for all samples. Nevertheless, the normal practice is

to decide in advance the approximate quantity of internal standard

to be added to each sample, since the ratio of analyte to internal

standard can be measured most accurately when they are present in

similar concentrations. Consequently, the quantity of internal

standard added to a specimen should give a concentration intermediate between the lowest and highest anticipated analyte concentrations, or more precisely, the concentration of internal standard should be such that the lowest anticipated weight ratio of

analyte to internal standard should approximately equal the inverse

of the highest anticipated ratio. Thus, if one desires to measure

concentrations of a drug over the range 1 to 1000 ng/ml, the amount

of internal standard added should give a concentration of approximately 32 ng/ml (1:32 = 32:1000).

Some analysts may favor a lower concentration of internal standard

on the basis that it is more difficult to measure low concentrations

of a drug than high concentrations, and therefore the internal standard concentration should be selected so as to facilitate accurate

measurement of‘ low levels of the drug. On the other hand, relatively large quantities of internal standard are sometimes used in order

for it to act as a “carrier” to minimize losses of analyte due to

adsorption at active sites on the surface of extraction vessels and

within the GC column (7). However, the “carrier” effect is not a

significant factor except at very low analyte concentrations, usually below about 10 ng/ml (8). Based upon these considerations we

have standardized our procedure by specifying the addition of approximately 40 ng of internal standard to each ml of body fluid.

A calibration graph should include measurements at a minimum of

four different concentrations of analyte plus a blank, i.e., a

sample with no added analyte. At least three samples should be prepared and analyzed at each concentration in order to assess the

variance in the measurements at each concentration. The analyte

concentrations used in preparing the calibration graph must include the extremes of concentrations expected to be found in the

unknowns, since linearity must be demonstrated, not assumed. The

remaining points on the graph should correspond to intermediate concentrations.

Since it is important to add the same volume of standard solution

to each body fluid sample in preparing a calibration graph, it is

necessary to prepare a series of standard solutions containing different concentrations of the analyte. A dilution scheme designed

to provide seven analyte concentrations covering the range 1 to

1000 ng/ml is shown in Table 1. To use the scheme, prepare standard stock solutions of the analyte and the internal standard having accurately measured concentrations of approximately 0.10 mg/ml

in methanol as described in the Experimental Procedure sections

within each of the drug chapters. The dilution scheme requires the

10
use of seven 10-ml volumetric flasks, one 1-ml volumetric pipette,

three 2-ml volumetric pipettes, and three 5-ml volumetric pipettes.

TABLE 1. DILUTION SCHEME FOR PREPARING

ANALYTE STANDARD SOLUTIONS

Dilute 1 ml of the 0.10-mg/ml analyte stock

solution to 10 ml to give a 10.0-μg/ml solution.

Dilute 5 ml of the 10.0-μg/ml solution to

10 ml to give a 5.0-μg/ml solution.

Dilute 2 ml of the 5.0-μg/ml solution to

10 ml to give a 1.0-μg/ml solution.

Dilute 5 ml of the 1.0-μg/ml solution to

10 ml to give a 500-ng/ml solution.

Dilute 2 ml of the 500-ng/ml solution to

10 ml to give a 100-ng/ml solution.

Dilute 5 ml of the 100-ng/ml solution to

10 ml to give a 50.0-ng/ml solution.

Dilute 2 ml of the 50.0-ng/ml solution to

10 ml to give a 10.0-ng/ml solution.

Exactly 100 μl of each of these standard solutions is added to 1 ml

of drug-free body fluid along with 100 μl of a 400-ng/ml solution

of the internal standard. Prepare the 400-ng/ml standard solution

by diluting 1 ml of the 0.10-mg/ml stock solution of internal standard to 250 ml with methanol. Each of the body fluid samples will

then contain 40 ng of internal standard and the concentration of

analyte shown in the right hand column in Table 2.

Prepare three or more spiked body fluid samples at each of the listed

concentrations. Extract and analyze each sample as described in the

Experimental Procedure section in the appropriate drug chapter.

Plot the ratio of peak heights of the analyte and the internal standard to the quantity of analyte added to the body fluid sample. A

calibration graph is then obtained by constructing the straight line

which best fits the plotted points. The best fit straight line can

be determined by visual estimation, or more accurately, by the method of “least squares” in which that straight line is drawn which

minimizes the sum of the squares of the vertical deviations from

the line.

Figure 1 is an example of a computer-generated calibration graph.

The significant features of the graph are the slope, the y-intercept, the linear range of the assay, and the variance of the measurements at different analyte concentrations. In this example the

11
FIGURE 1. A CALIBRATION GRAPH FOR ANALYSIS OF TETRAHYDROCANNABINOL IN PLASMA

12
TABLE 2. SCHEME FOR ADDITION OF ANALYTE AND INTERNAL

STANDARD (IS) TO BODY FLUID SAMPLES FOR

PREPARING CALIBRATION GRAPHS

Resulting Analyte

Concentration

Add to 1 ml of Body Fluid

100 μl of the 400- + No analyte

ng/ml IS solution

(ng/ml) 

0

"

"

"

"

"

"

" + 100 μ1 of the 10-ng/ml

analyte solution

1

+ 100 μ1 of the 50-ng/ml

analyte solution

5

+ 100 μ1 of the 100-ng/ml

analyte solution

10

+ 100 μ1 of the 500 ng/ml

analyte solution

50

+ 100 μl of the 1-μg/ml

analyte solution

100

+ 100 μ1 of the 5-μg/ml

analyte solution

500

+ 100 μl of the 10-μg/ml

analyte solution

1000

variance is indicated by vertical bars representing the relative

standard deviation at each concentration measured. The slope and

the y-intercept are the functions used to calculate the quantity

of analyte in an unknown based on the ion current response ratio.

Assays using stable isotope labeled internal standards usually give

calibration lines with positive y-intercepts. The magnitude of the

y-intercept reflects the proportion of internal standard molecules

containing no heavy isotopes (unlabeled molecules). A large y-intercept can significantly limit the sensitivity of the assay.

An important advantage to the use of stable isotope labeled internal standards is that the slope of the calibration line should remain constant as long as the compositions of the standard solutions

remain unchanged and the same volumes of standard solutions are

added to the body fluid samples. Because of the nearly identical

chemical and physical properties of the analyte and its isotope

labeled analog, their weight ratio should not be affected to any

13
significant extent by variations in the extraction, derivatization,

chromatographic and mass spectrometric processes. For this reason

the practice in our laboratories is to determine new calibration

graphs only when new standard solutions arc prepared, or when the

analysis of control specimens containing known concentrations of

the drug shows unacceptable variance from the true values.

If the concentration of a drug in an unknown specimen is found to

be greater than the highest concentration used in preparing the

calibration graph, the specimen should be reanalyzed after dilution

with water or saline (typically 1:10) in order to bring its concentration within the extremes of the calibration graph. The alternative practice of adding a larger quantity of internal standard

to the specimen is less satisfactory because it requires preparation of a new calibration graph.

SAMPLE EXTRACTION

One of the important criteria that guided our selection and evaluation of sample preparation procedures was that they should be as

simple and rapid as possible while still providing adequate recovery

and separation of the analyte from compounds that could interfere

with the GC/MS quantitation. The very high sensitivity and selectivity of GC/MS often permits use of relatively simple extraction

procedures, and we have attempted to fully exploit this feature.

A second major concern was to minimize loss of analyte due to adsorption, evaporation, or decomposition. Even though use of stable

isotope labeled internal standards can compensate for loss of a portion of the analyte through these processes, any loss of analyte

will reduce the sensitivity of the assay. Loss of analyte due to

adsorption on glass surfaces is particularly a problem when the

quantity of analyte is in the low nanogram range. Adsorption can

be minimized by silylation of all glass surfaces with which solutions of the analyte will come in contact. Commercial formulations

are available for liquid-phase silylation of glass surfaces (for

example, Glas-Treet, Regis Chemical Co., Morton Grove, IL 60053).

However, vapor-phase silylation is more convenient and less expensive if large quantities of glassware must be silylated on a regular

basis (10).

All glassware should be thoroughly cleaned before silylation. The

easiest and most thorough method of removing organic contaminants

from glassware is to bake it off in a muffle furnace. Unfortunately, a muffle furnace of sufficient capacity to handle normal laboratory glassware is expensive to purchase and operate. Chromic

acid baths can also be used to remove all traces of organic material

from glassware, but they are messy and are a safety hazard. Consequently, in most laboratories glassware is cleaned by washing with

a detergent solution followed by thorough rinsing with distilled

water. A final rinse with acetone helps to assure removal of any

remaining organic residue (11).

14
It is the practice in our laboratories to use high purity solvents

for preparation of standard solutions and all sample extractions,

even though interference from solvent impurities is not nearly as

much of a problem in GC/MS assays as it is in most other methods

for drug analysis. High purity solvents from several commercial

sources have proven satisfactory. We have not found it necessary

to redistill the solvents.

Some of the drugs discussed in this monograph are quite volatile,

so care must be taken to avoid evaporative loss of the drug during

concentration of the extract. Steps that have been taken to minimize evaporative loss of analyte include:

1. Eliminating the need for concentration of extracts by

using a very small quantity of extraction solvent (see

assays for methaqualone, cocaine, and diazepam).

2. Using highly volatile solvents such as methylene chloride,

hexane, or 1-chlorobutane, which can be removed by heating

at mild temperatures ( 40 to 60°C) under a gentle stream

of air or nitrogen.

3. Adding a small quantity (10 to 20 μl) of a high boiling

liquid to the extract to act as a “keeper solvent” during

evaporative concentration.

Procedures for analysis of volatile basic drugs such as amphetamine

often specify addition of an acid prior to evaporative concentration in order to convert the drug to a nonvolatile salt. In our

experience the use of a keeper solvent such as dimethylformamide

(DMF) is a more effective means of reducing loss due to evaporation

of such drugs (see assays for phencyclidine, methadone, tetrahydrocannabinol, amphetamine, methamphetamine, mescaline, and DOM). However, it is important to establish that the keeper solvent does not

exert adverse effects on the GC/MS assay. For example, we found

that DMF could not be used in combination with trifluoroacetic anhydride for derivatization of amphetamine. Apparently the DMF suppressed removal by evaporation of the trifluoroacetic acid formed in

the derivatization reaction, and subsequent injections of the highly

acidic mixture adversely affected the gas chromatography of the trifluoroacetamide derivative. This problem was avoided by using Nmethyl-bis-(trifluoroacetamide) instead of trifluoroacetic anhydride to prepare the amphetamine trifluoroacetamide derivative (see

assays for amphetamine, methamphetamine, mescaline and DOM).

CALIBRATION AND PERFORMANCE EVALUATION OF THE GC/MS

In spite of substantial improvements in stability and reliability,

gas chromatograph/mass spectrometers remain one of the most attention-requiring of analytical instruments. In addition to having

complex electronic circuitry, the mass spectrometer includes key

components which can only be operated under high vacuum. If leaks

develop in the vacuum system, or if the ionizer, mass analyzer, or

detector become contaminated, the performance of the instrument

15
deteriorates. Yet some contamination is inevitable, since samples

are introduced and effectively decomposed within the instrument’s

vacuum system. Likewise, gas chromatographic columns have a finite

lifetime which often is reduced to only a few weeks when large numbers of crude biological extracts are analyzed. For these reasons

it is imperative to evaluate the total system performance at regular and frequent intervals.

The specific manner in which a GC/MS system is tuned up and calibrated, and its performance evaluated, will vary with different instruments, with the level of performance required for the particular analyses, and according to the personal preferences of the

operator. The procedures described here have proven satisfactory

in our laboratories, and are intended as a guide to assist readers

in establishing a systematic scheme appropriate for their own daily

instrument “check-out.”

A thorough discussion of tune-ups and performance evaluation procedures is beyond the scope of this chapter. Instead, the emphasis

here is on procedures and suggestions which may not be included in

operator and maintenance manuals , but which we have found to be

useful in preparing to perform the types of analyses described in

this monograph.

Performance Evaluation Procedure

Before turning on the ionizer filament and the electron multiplier

voltage to obtain mass spectra it is good practice to check the pressures in the ion source ( < 10-3 mm) and in the analyzer ( < 10-6 mm),

as well as the GC column head pressure, to assure that no major

leaks have occurred. Following this preliminary check, set the GC

column at the maximum temperature to be used for the assay, and observe the electron impact mass spectral background with and without

the helium carrier gas entering the ion source. Some increase in

background will occur when the carrier gas is introduced into the

mass spectrometer. However, a very large increase in backrround is

cause for concern. If the intensities of ions due to air [m/z 28.

(N2

+

), 32 (O2

+

), 40 (Ar+), and 34 (CO2

+

)] become abnormally large;

a leak in the GC system is indicated. The most common locations

of air leaks within the GC system are the injector septum and the

colunm connector fittings. Under methane or ammonia chemical ionization conditions the air peaks will not be seen, and yet an air

leak can significantly affect the instrument sensitivity. Therefore it is important to check for an air leak before switching

to chemical ionization. The appearance of abnormally intense ion

peaks at higher masses suggests that the GC column should be further conditioned or replaced.

Next, introduce methane into the ion source to serve as the CI reagent gas. This is normally done by changing the GC carrier gas

from helium to methane, although some chemical ionization mass

spectrometers require use of a separator in the GC/MS transfer line,

in which case the methane reagent gas is introduced into the ion

source through a separate inlet. Reduce the electron multiplier

16
voltage (to 1.2 KV) and the filament emission current (to 5 0

microamps) to prevent possible damage to the electron multiplier,

and observe the methane CI mass spectrum. It should be similar

to the spectrum shown in Figure 2. The peak at m/z 19 is due to

protonated water and is often quite large, particularly just after the instrument has been down for cleaning and maintenance. The

size of the water peak can usually be reduced by an overnight system bakeout. However, if ammonia is to be used as the reagent gas,

the adsorbed water in the system is unlikely to affect the ionization of sample molecules.

Return the electron multiplier voltage and filament current to their

normal operating valves, and bleed a small quantity (about 10-5mm

partial pressure) of a suitable reference compound such as perfluorotributylamine (often referred to as PFTBA or FC-43) into the

ion source and observe its methane CI mass spectrum. The relative

ion intensities in CI mass spectra are often strongly affected by

the ion source temperature. At a source temperature of approximately 150°C the methane CI mass spectrum of perfluorotributylamine

should look similar to that shown in Figure 3. At higher source

temperatures the high-mass ions tend to decrease in abundance relative to the low-mass ions. Either of the very abundant ions at

m/z 219 and 414 in the methane CI mass spectrum of perfluorotributylamine can be used to optimize source potentials, depending on

which is closest to the analyte and internal standard ions that

will be monitored.

In mass spectrometry, resolution and sensitivity are inversely related. Normally the mass spectrometer is adjusted to give unit resolution, i.e. base-line separation between consecutive integral

mass peaks. However, in quantitative analysis using selected ion

monitoring, a gain in sensitivity can be achieved by decreasing the

mass resolution and thereby increasing the ion current passing

through the mass analyzer. The degree to which the resolution can

be lowered is limited by the specificity required, and since specificity is directly related to resolution, a decrease in resolution

will increase the likelihood of interference from other ions. In

our laboratories the mass spectrometer is normally adjusted to unit

resolution if the ions to be monitored have masses below 300, and

the resolution is reduced by about a factor of 2 when ions above

mass 300 are to be monitored.

After the mass spectral resolution has been set and the ion source

potentials optimized, the system's mass calibration should be checked

and recalibrated if necessary.

If the assay to be performed involves the use of ammonia as reagent

gas, it is best to introduce the ammonia into the ion source through

an inlet line which is adjacent to or concentric with the GC/MS

transfer line. We have found that introducing the ammonia gas

through an inlet into the ion source at a location opposite to the

entering carrier gas gives less satisfactory results.

17
FIGURE 2. METHANE REAGENT GAS IONS (ION SOURCE TEMPERATURE, 150°C; PRESSURE, 1 x 10-3 MM)

18
FIGURE 3. METHANE CI MASS SPECTRUM OF PERFLUOROTRIBUTYLAMINE (QUADRUPOLE MASS ANALYZER. ION

SOURCE TEMPERATURE, 150°C; METHANE PRESSURE,

1 x 10-3 MM; PFTBA PRESSURE, 1 x 10-5 MM)

19
Slowly increase the rate of flow of ammonia into the ion source

while observing the ion intensities at m/z 18 and 35. As the ammonia

flow rate is increased, the intensity of the m/z 18 peak (NH4

+

) should

at first increase and then reach a plateau, at which point a peak

at m/z 35 (NH3•NH4

+

) will appear and begin to increase in intensity.

The optimum ammonia flow rate is the point at which the m/z 18 peak

reaches a maximum and the m/z 35 peak just begins to increase in

height. At this point the methane reagent gas ions will be completely suppressed. The total reagent gas pressure in the ion chamber should be between 0.3 and 1.0 x 10-3

 mm. If a separator is

used, it may be necessary to bleed in methane along with the ammonia to achieve this total pressure. Figure 4 shows an example of

the reagent gas ions observed when methane is used as carrier gas

and ammonia is introduced into the ion source as described.

Set the gas chromatograph and mass spectrometer ion source operating parameters to those that will be used for the assay to be performed. Inject into the GC/MS system a solution containing a specific quantity of ,analyte which is sufficient to yield a good quality mass spectrum. (Usually 100 to 200 ng is sufficient for this

purpose; therefore, it is convenient to inject 1 or 2 μ1 of the 0.1

mg/ml standard stock solution called for in each of the drug assay

experimental procedures). Repetitively scan the mass range from

50 daltons to above the protonated molecule ion of the analyte.

Compare the size and shape of the analyte peak in the total ion

current chromatogram with similar data acquired previously, in order to determine if there has been a significant change in sensitivity or gas chromatographic resolution. Also, compare the acquired

CI mass spectrum of the analyte with the corresponding CI mass spectrum in this monograph.

As a final check of the GC/MS performance, set up the system for

selectcd ion monitoring and inject a standard solution containing

a quantity of analyte close to the lowest level of analyte that the

assay must be capable of measuring. The instrument operator’s manual should provide guidance as to the selected ion monitoring parameters that will provide the highest signal-to-noise ratio and the

best dynamic range. Typically the system will sequentially sample

the ion current, at each mass to be monitored, for 100 msec intervals. It is essential, of course, that the ion currents be measured at the precise center of each of the monitored ion masses.

If the analyte peak signal obtained from the selected ion monitoring

analysis of the standard solution is sufficiently strong relative to

the background noise level to permit reasonably accurate measurement of the peak height (i.e., a signal-to-noise-ratio of at least

3:1), proceed with the analysis of biological extracts.

Since the primary purpose of conducting the foregoing instrument

checkout is to assess the current performance of the system, it is

extremely useful to maintain a record of the data from these evaluations. The ability to compare current performance with data acquired previously under identical experimental conditions will be

a great help in identifying likely causes of any changes that may

occur in instrument performance.

20
FIGURE 4. METHANE-AMMONIA REAGENT GAS IONS (ION SOURCE

TEMPERATURE, 150°C; AMMONIA PRESSURE, 2 x

10-4 MM; METHANE PRESSURE, 8 x 10-4 MM)

21
A common cause of poor sensitivity in GC/MS assays is adsorptive

loss of the analyte within the GC column. Often the sensitivity

improves after 3 number of injections of analyte solution. “Priming” of GC columns in this manner is a common practice in gas chromatography. The effect of priming is most noticeable when a column

was previously. used for analysis of chemically different compounds.

Ideally a separate column should be reserved for each of the different drugs commonly assayed. If this is not practical, columns

used for quantitation of basic compounds should at least not be

interchanged with those used for analysis of acidic compounds.

An inherent danger in column priming is that some of the adsorbed

analyte may be “washed off” the column during subsequent injections,

resulting in an erroneously high value for the concentration of

analyte measured. This potential source of error can be easily

checked by analyzing a sample blank (i.e. a body fluid extract containing the internal standard but none of the analyte) before beginning analysis of sample unknowns.

QUALITY CONTROL AND SOURCES OF ERROR

In addition to determining that the total GC/MS system performance

is satisfactory before beginning the analysis of sample unknowns,

it is important intersperse spiked standards among sample unknowns so that intra-day changes in instrument performance will be

evident and documented. Control charts are effective means of maintaining a continuous record of the accuracy experienced with each

drug assay (12,13).

Probably the largest source of error in GC/MS quantitative drug

assays is that involved in preparing the standard solutions used in

establishing calibration graphs (5). Once the internal standard

is added to the body fluid and becomes equilibrated, there are few

things which will change the weight ratio of the drug to its stable

isotope labeled analog.

Incomplete equilibration of the internal standard in the body fluid

is a potential cause of poor precision. Unfortunately the time and

the conditions required for internal standards to achieve complete

equilibration in terms of protein binding are difficult to determine and undoubtedly vary for each drug and type of specimen. The

assay procedures in this monograph specify equilibration for 15 min

before beginning the extraction, but we have no convincing data upon

which to base this recommendation. Others routinely allow as long

as 12 hr for equilibration (5).

In spite of the high specificity of the selected ion monitoring technique, particularly when ammonia chemical ionization is used, ions

from endogenous compounds will occasionally interfere. Usually a

visual examination of the ion current profile plots will indicate

when an analyte or internal standard peak contains a contribution

from another compound; that is, the peak width will be broader than

22
normal or its retention time will be slightly altered. However,

we have seen a few rare cases where an endogenous compound had an

identical retention time and yielded ions at the same mass as either

the analyte or the internal standard. When this situation occurs

it can easily go undetected and result in erroneous data.

The experimental procedures can be altered in several ways if an

unusually high degree of specificity is required. For example,

the GC/MS analysis can be repeated using a GC column with a different polarity liquid phase, or a high resolution glass capillary

column can be used. Also, prominent fragment ions can be monitored

in addition to the protonated molecule ions. This may require an alteration of the ionization conditions, such as the substitution of

methane for ammonia as the CI reagent gas.

To conclude this chapter, it is appropriate to reemphasize the

remarkable versatility of the gas chromatograph/mass spectrometer

as an analytical instrument. The experienced analyst will become

aware of the full range of capabilities available and can then

choose to modify the procedures in order to best satisfy specific

analytical needs.

REFERENCES

1. R.L. Foltz, D.A. Knowlton, D.C.K. Lin, and A.F. Fentiman, Jr.

Proceedings of the Second International Conference on Stable

Isotopes, Oak Brook, IL, October 20-23, 1975.

2. A.F. Fentiman, Jr. and R.L. Foltz. J. Labelled Compounds,

13, 579 (1977).

3. A.F. Fentiman, Jr. and R.L. Foltz. J. Labelled Compounds,

12, 69 (1976).

4. Personal communication from Dr. Fred Falknier, Pfizer, Central

Research, Groton, CT.

5. B.J. Millard. Quantitative Mass Spectrometry. Heyden, London

(1978).

6. M.G. Lee and B.J. Millard. Biomed. Mass Spectr., 2, 78 (1975).

7. B. Samuelsson, M. Hamberg, and C.C. Sweeley. Anal. Biochem. ,

38, 301 (1970).

8. R. Self. Biomed. Mass Spectr., 6, 315 (1979).

9. M. Claeys, S.P. Markey and W. Maenhaut. Biomed. Mass Spectr.,

4, 122 (1977).

10. D.C. Fenimore, C.M. Davis, J.H. Whitford, and C.A. Harrington.

Anal. Chem., 48, 2289 (1976).

23
11. R.R. Watts, ed. Analysis of Pesticide Residues in Human and

Environmental Samples. U.S. Environmental Protection Agency,

Health Effects Research Laboratory, Environmental Toxicology

Division, Research Triangle Park, NC, Section 3A, p. 1-6 (1980).

12. D.L. Massart, A. Dijkstra, and L. Kaufman, eds. Evaluation and

Optimization of Laboratory Methods and Analytical Procedures.

Elsevier, Amsterdam (1978).

13. H.C. Curtius and M. Roth, eds Clinical Biochemistry, Principles and Methods. Walter de Gruyter, New York, NY, 1, p. 594-

617 (1974).

24
Chapter 3

Phencyclidine (PCP)

Phencyclidine (PCP), 1-(1-phenylcyclohexyl)piperidine (I), was

originally developed in the 1950’s and marketed in 1963 as Sernyl,

a surgical anesthetic. However, human clinical use was soon discontinued because of bizarre adverse side effects, notably postanesthetic delirium. By 1967 the drug was commercially available

for veterinary use only (1).

PCP appeared as an illicit street drug in 1967, but within a year

had fallen out of favor in the drug culture, presumably because

of its unpleasant effects. In spite of the early negative response, street use regained momentum and by 1976 PCP was the fourth

most common cause of hospitalization for drug abuse in San Francisco (2) and the leading cause of inpatient psychiatric admissions in the Washington, D.C. area (3). Until very recently, PCP

sold on the street was rarely represented correctly; either it

was disguised by an esoteric name or it was deliberately misrepresented as THC, mescaline, psilocybin, or some other popular

hallucinogen. It is often mixed with other drugs such as LSD or

amphetamine, and appears in various physical forms including powder, capsules, tablets, or liquid sprayed on plant leaves. Administration among users is by smoking, nasal inhalation, or oral

ingestion (3). “Street” PCP is not generally diverted from legitimate veterinary sources, but is synthesized illegally from

readily available chemicals (5). An intermediate in the synthesis,

1-piperidinocyclohexane carbonitrile (II), is extremely toxic and

is difficult to remove from the final product. Its presence as

a contaminant in imperfectly synthesized PCP has been implicated

in cases of coma and death following PCP ingestion (6,7).

25
PHARMACOLOGY

The pharmacology of PCP is dose-dependent and complex. The drug

acts primarily on the central nervous system as a depressant, but

in a manner distinctly different from that of the sedative-hypnotic

class of depressants. Studies in rats have shown it to be a potent

competitive inhibitor of catecholamine uptake in both the dopaminergic and noradrenergic regions of the brain. This, combined with

its anticholinergic properties, makes PCP an unusual and, therefore, an attractive substance for studies of model psychoses (8).

In its original human surgical doses (0.25 mg/kg body weight given

intravenously), PCP was found to induce unresponsiveness and complete anesthesia although the patients appeared to he awake. Blood

pressure was elevated. With intravenous doses of 0.5-1 mg/kg, agitation and seizure activity appeared. Following even a small oral

“street” dose (commonly 1-6 mg), the initial “high,” during which

the subject is unresponsive but confused and agitated, is followed

by depression, irritability, feelings of isolation, and sometimes

paranoia. Doses of 10-20 mg, possible even by smoking, are sufficient to induce stupor or coma, and the larger amounts possible

with oral ingestion can precipitate fatal epileptic seizures or

hypertensive crises. Blood concentrations of PCP as low as 0.10

μg/ml have been associated with behavioral effects leading to injury

or death, while concentrations greater than 1.0 μg/ml cause coma

in most individuals. Blood concentrations over 2.0-2.5 μg/ml are

generally fatal (9). Recovery from sublethal doses usually occurs

within a few hours when the drug has been smoked or inhaled, but

oral ingestion is characterized by a recovery period of days or

weeks (2). It has been suggested that these prolonged effects

may be due to the lipophilic nature of PCP; in rats high concentrations of the drug were found in adipose tissue long after its disappearance from plasma and urine (10). Also, the analysis of

blood and tissue samples from phencyclidine-related fatalities revealed that in all cases tissue concentrations greatly exceeded

those in blood, and blood levels were similar to those found in the

plasma from casual users (11). Therefore, since PCP concentrates

in tissue, plasma concentrations may not adequately reflect the

severity of ingestion.

Schizophrenic psychosis, as opposed to short-tern intoxication,

can persist for several weeks after the use of PCP, and appears to

be related to personality factors and/or individual drug sensitivity. The drug has been found to be teratogenic (12). While deaths

resulting from true pharmacologic overdoses of PCP have been reported, in many other cases death has resulted from the behavioral

toxicity of the drug; i.e., accidental drowning or violence provoked by aggressive hehavior (13). Clinically significant interactions of PCP with other drugs of abuse have been suggested by

animal experiments. For example, in monkeys PCP strongly potentiates the effects of pentobarbital, including respiratory depression.

PCP has also been found to reinforce drug self-administration in

monkeys, the only hallucinogen which is known to do so, and some

human chronic users report psychological dependence and behavioral

tolerance. However, no evidence for physical dependence has been

reported either in man or animals (14).

26
The pharmacology and toxicology of PCP are reviewed in papers published in 1974 (1.5) and in 1978 (9). Also, a National Institute

on Drug Abuse monograph reviews many aspects of the abuse of PCP,

treatment of PCP intoxications,

(16).

and the pharmacology of the drug

PHARMACOKINETICS AND METABOLISM

Phencyclidine, with a pKa of 8.5, appears to be rapidly metabolized

and excreted. About 60 percent of labeled PCP given intravenously

to monkeys appeared in the urine within 12 hours and 75 percent

within eight days (17). In cases of acute intoxication in humans,

urine concentrations have been used to predict the duration of coma

(2). A comparison of plasma with urinary PCP concentrations in human “overdose” cases showed that urinary concentrations of PCP were

13-19 times the concurrent concentrations in plasma (18). A plasma

elimination half-life of about 11 hours has been reported in a case

of human PCP ingestion (19). It has been shown, however, that PCP

is excreted much more rapidly in acidic urine, and several severely

intoxicated patients have been successfully treated by urine acidification procedures, such as gastric infusion with NH4Cl (20).

In man, PCP is excreted primarily as monohydroxylated derivatives

in the form of conjugates. Two of these metabolites, isolated from

human urine after enzymatic hydrolysis, were identified by mass

spectrometry as 4-phenyl-4-piperidinocyclohexanol (III) and 1-(1-

phenylcyclohexyl)-4-hydroxypiperidine (IV) (21). Both metabolites

were found to exhibit weak pharmacological activity in test animals

(22)

Traces of a N-dealkylated metabolite, 1-phenylcyclohexylamine (V),

have also been detected in the urine of a PCP-intoxicated patient

(23). Dihydroxy metabolites are produced in rats, mice, pigeons,

sheep, and monkeys, and in these animals PCP tends to be less toxic

than in species which have a weaker capacity for hydroxylation (24).

In tests on monkeys, no metabolites were detected in central or

peripheral tissue, although unchanged PCP was present in sciatic

nerve, spinal cord, and brain (17). The detection of PCP metabolites in human blood has not been reported.

27
ANALYTICAL METHODOLOGY

Generally, the urine of patients seen on an emergency basis contains a sufficiently high concentration of unmetabolized PCP

(>0.2 mg/ml) to permit qualitative detection by any of a variety

of analytical methods (25). These have included thin-layer chromatography (TLC), with a sensitivity of 0.2 μg/ml (26) and gas chromatography (GC) with a sensitivity of 2.0 μg/ml (27). GC combined

with mass spectrometry (28,29) and TLC (30) has been used to identify PCP in confiscated street drug samples. However, both clinical work and pharmacokinetic studies have been hampered by the lack

of reliable quantitative methods for analysis of PCP in biological

fluids. Hoffmann-La Roche markets a radio-immunoassay (RIA) kit for

PCP and Syva is field testing a homogeneous enzyme immunoassay

(EMIT) for the drug. To date, however, the sensitivities and specificities of these assays have not been published.

Recently, a method using ultraviolet spectroscopy was reported,

which was able to measure PCP concentrations in urine extracts containing as little as 0.01 mg of the drug by comparison with standard absorbance curves. However, when other basic drugs were present in the sample, GC assay with a flame ionization detector (FID)

was used. Methadone was chosen as the internal standard. The

GC-FID system (Chromosorb W coated with Apiezon L, column temperature 200oC) detected 0.076 μg/ml of PCP added to urine, and measured a range of 0.1-69.4 μg/ml of the drug in samples from hospital admissions (31).

For rapid analysis of urine for PCP in a clinical setting, temperature-programmed (150- 230°) GC-FID on a column of 5 percent SE-30

plus 1 percent Carbowax 20M on Chromosorb WHP has been suggested.

A metabolite, 1- (1-phenylcyclohexyl)-4-hydroxypiperdine (IV) is

also observable by this technique, to confirm the presence of PCP

even when the drug has been extensively metabolized (32).

A GC-FID method using mepivicaine as the internal standard and an

OV-1 or OV-17 column at 180°C, was applied to blood and tissue samples as well as to urine. Blood concentrations in fatal overdose

cases ranged from 0.5 to 5.0 μg/ml, and in liver tissue from the

same individuals, from 5.0 to 36.0 μg/g (33). Human plasma PCP

concentrations in clinical cases, which covered a range of 0.09-

0.22 μg/ml, were measured by a similar GC-FID technique (3.8 percent UCW-98 on Gas Chrom Q at 200°C column temperature) using

ketamine as the internal standard (19). Improved sensitivity and

selectivity were achieved in a GC method by using a nitrogenphosphorus detector (3 percent OV-17, 165°C isothermal). Meperidine was used as the internal standard because of its chemical

similarity to PCP. The method gave a linear calibration curve up

to 1 μg/ml and could detect as little as 10 ng/ml of PCP in plasma

(11).

The combination of GC and MS with selected ion monitoring and the

use of an isotope-labeled variant as the internal standard has

been applied to the quantitation of PCP in blood and urine. Since

an initial report of the adaptation of GC/MS to the assay of PCP

28
(21), the technique has been applied to a study of blood concentrations in 26 PCP intoxications where concentrations ranged from

0.007 to 0.250 μg/ml (34), and to studies of serial plasma and

urine samples in an effort to correlate PCP concentrations with

the duration of psychotropic effects (20). GC/MS analysis of PCP

has also been adapted to an automated procedure using probability

based matching, in a computerized GC/MS system (Olfax) utilizing

a reverse-search algorithm for the measurement of PCP levels in

the urine. The method permits analysis of up to eight samples

per hr and has a detection limit of approximately 10 ng/ml of

PCP (35).

For quantitative studies of PCP, it is important to note that phencyclidine undergoes progressive thermal decomposition to 1-phenylcy-

 clohexene as temperatures rise above 150°C. To minimize pyrolysis

during gas chromatograpny, the injection port temperature should be

kept below 200°C (21,25,36).

Solvents which have been used for the extraction of PCP from alkaline biological fluids include hexane, with an overall recovery of

70-85; percent (21); cyclohexane, 95 percent (used only for urine)

(35); petroleum ether, 76 percent from plasma (19); diethyl ether

(34); 1-chlorobutane (33); 95 percent ethanol (for TLC of urine)

(30); and chloroform, for urine only, with 83 percent recovery reported (27,31). Increasing the pH of the urine to 9 reportedly

allows 94 percent recovery of PCP and quantitative recovery of the

nydroxy metabolite with chloroform (32). Extraction from urine by

a reverse phase XAD-2 resin procedure, with quantitative recover)

of PCP and 89 percent of the metabolite, has also been reported.

Back: extraction into aqueous acid is often used in order to separate PCP from neutral and acidic materials, followed by realkalinization and reextraction with an organic solvent.

EXPERIMENTAL PROCEDURE

Because phencyclidine can be psychoactive at low blood concentrations the GC/MS analytical procedure is designed to measure the

drug at plasma concentrations as low as 1 ng/ml. The extraction

procedure consists of addition of deuterium-labeled phencyclidine

and a pH 9.6 buffer to the plasma, followed by extraction with

hexane, back extraction into acid, and finally, alkalinization and

reextraction with hexane. Chemical ionization with methane and

ammonia as reagent gases is used for the GC/MS analysis.

Standards and Reagents

Phencyclidine hydrochloride was purchased from Applied Science

Laboratories, State College, PA 1680l. Phencyclidine-2H5 (1-[1-

(phenyl-2H5) cyclohexyl]piperidine) was synthesized from bromobenzene2H5

 by a published procedure (38). It was found to be 99

percent pure based on GC/MS analysis. Over 99 percent of the molecules contained five deuterium atoms and less than 0.1 percent of

the molecules were undeuterated.

29
A suitable pH 9.6 buffer can be prepared by dissolving 500 grams

of potassium monohydrogen phosphate (K2HPO4) in about 500 ml of

distilled water. When all of the salt is dissolved, allow the

solution to cool to room temperature and add sufficient water to

give exactly 1 liter.

Stock solutions of phencyclidine and phencyclidine-2H5, used for

the determination of calibration graphs (Chapter 2) and the preparation of working standards, are prepared as follows. Weigh into a

100-ml volumetric flask 11.5 mg of phencyclidine hydrochloride or

10 mg of the free base. Dissolve the drug in methanol and bring

the volume to exactly 100 ml with additional methanol. The resulting stock solution will correspond to a concentration of 0.10

mg/ml of phencyclidine calculated as the free base. A series of

working standard solutions can then be prepared by appropriate

dilution of the stock solution as described in Chapter 2.

The stock solution of phencyclidine-2H5 is prepared in the same

manner. For the measurement of phencyclidine in body fluids within

the concentration range of 1 to 1000 ng/ml, addition of approximately 40 ng of the internal standard to each ml of specimen is

satisfactory. This can be conveniently done by adding 100 μ1 of

a 400-ng/ml phencyclidine-2H5 methanolic solution to each ml of

sample. Prepare the 400-ng/ml standard solution bv diluting 1 ml

of the 0.10-mg/ml stock solution of phencyclidine-2H5 to 250 ml

with methanol. Store stock and working standard solutions in

well-stoppered or capped glass vessels under refrigeration.

Extraction

Transfer 1 ml of specimen (whole blood, plasma, urine, or cerebrospinal fluid) to a 20-ml culture tube equipped with a Teflon-lined

screw cap (0.5 to 2 ml of specimen can be used, but it must be measured accurately). Add 100 μ1 of the 400-ng/ml phencyclidine-2H5

internal standard solution to the body fluid and vortex for 10 sec.

Allow the sample to equilibrate for about 15 min and then add 1 ml

of the pH 9.6 buffer and 5 ml of hexane. Cap the tube and check

for a leak around the cap. Gently mix the contents for 30 min

using a motorized rocker or rotator. Centrifuge for 5 min and then

transfer the organic layer (top) to a clean culture tube by means

of a disposable Pasteur pipette. Add 5 ml of 0.2 N H2SO4 and vigorously agitate for about 1 min. Remove and discard the hexane

layer (top). Add a sufficient quantity of 1 N NaOH to the aqueous

layer to raise the pH to 9.5. Add 5 ml of hexane and again agitate

vigorously for about 1 min. Centrifuge and transfer the organic

layer (top) to a silylated concentrator at least 5 ml in volume

and having a conical or nipple-shaped bottom. Add 20 μ1 of dimethylformamide as a “keeper” solvent to minimize evaporative loss of

the phencyclidine. Remove the hexane by evaporation under a gentle stream of nitrogen or filtered air while heating at 40 to 50°C.

When the extract volume has decreased to about 20 μ1, stopper the

tube or cover the top with Parafilm and store at 0°C until the GC/MS

analysis is to be performed. Inmediately prior to the analysis,

allow the tube to warm to room temperature.

30
GC/MS Analysis

The following experimental conditions are satisfactory for GC/MS

analysis of phencyclidine:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-1 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier gas: Methane, 15-20 ml/min

Temperatures: Injector, 200°C

Column, 180°C isothermal

GC/MS transfer line, 200°C

Ion source, 160°C

Under these conditions the phencyclidine and phencyclidine-2H5

should elute at about 3 min as narrow, symmetrical peaks.

before beginning the GC/MS analysis of samples, the performance

of the total GC/MS system should be evaluated and optimized as

described in Chapter 2. The ions to be monitored occur at m/z

244 and 249. They correspond to the protonated molecule ions for

phencyclidine and phencyclidine-2H5, respectively. With the divert

valve in the divert position, inject 2 to 6 μl of the hexane extract into the gas chromatograph. After approximately 1 min,

switch the divert valve so that the entire carrier gas flow enters

the ion source of the mass spectrometer, and begin data acquisition.

When the phencyclidine peaks have eluted, cease data acquisition

and return the divert valve to the divert position.

The quantity of phencyclidine in the plasma sample is determined

by measuring the heights (or areas) of the phencyclidine and phencyclidine-2H5

 peaks in the selected ion current profiles (“ion

chromatograms”) and relating the ratio of peak heights to a calibration graph. Calculate the concentration of phencyclidine in the

specimen by dividing the measured quantity of phencyclidine by

the exact volume of specimen used in the analysis.

Discussion of the Experimental Procedure

During extraction of phencyclidine from biological fluids, the

basic drug can be separated from neutral and acidic materials by

a back extraction technique. The biological sample is made alkaline and extracted with an organic solvent; the phencyclidine is

then back extracted from the organic solvent with an aqueous acid.

However, if chloroform or methylene chloride is used as the organic solvent in these steps and dilute hydrochloric acid is used

for the back extraction, most of the phencyclidine will remain in

the organic phase, together with the neutral components. The problem is avoided by using hexane as the organic solvent and dilute

sulfuric acid for the back extraction. The aqueous acid extract

is subsequently made alkaline and reextracted with hexane. With

this combination of reagents, the overall recovery of phencyclidine from 1 ml of whole blood containing 5 μg of the drug ranges

31
between 70 and 85 percent.

During early work with phencyclidine, it was noticed that GC analyses of the drug typically showed a second peak eluting considerably

before phencyclidine. The second peak was identified as 1-phenylcyclohexene on the basis of its mass spectrum and comparison with

authentic material (Aldrich Chemical Co., Milwaukee, WI). 1-Phenylcyclohexene is a thermal decomposition product of phencyclidine.

Consequently, when phencyclidine is gas chromatographed, the size

of the 1-phenylcyclohexene peak is dependent on the injection port

temperature. Figure 1 shows the flame ionization detector recordings which resulted from gas chromatography of phencyclidine with

injection port temperatures of 300, 250, and 200°C. The 1-phenylcyclohexene peak decreased in size as the injection port temperature was lowered. At an injection port temperature of 200°C the

1-phenylcyclohexene peak amounted to only 0.5 percent of the phencyclidine peak height. At an injection port temperature of 150°C

the 1-phenylcyclohexene peak was still detectable, but it now represented only 0.1 percent of the phencyclidine peak height. In

view of the thermal lability of phencyclidine, it is important to

keep all surfaces which may come in contact with the drug at the

lowest practical temperature. For example, if the GC/MS separator

is heated to 280°C some of the phencyclidine eluting from the gas

chromatographic column will be thermally converted to 1-phenylcyclohexene.

The electron impact (EI), methane chemical ionization (CICH4) , and

methane-ammonia chemical ionization (CICH4-NH3) mass spectra of

phencyclidine are shown in Figure 2. The EI spectrum of phencyclidine (top) snows an abundant ion at m/z 200 resulting from loss

of C3H7 from the cyclohexyl ring. Since the deuterium atoms in

the internal standard are on the aromatic ring, the corresponding

ion in the mass spectrum of the deuterated phencyclidine is shifted

to m/z 205; therefore, the ions at m/z 200 and 205 are satisfactory for selected ion monitoring. In the methane CI mass spectrum

(middle), the M

+

 is more intense than the MH+, suggesting that

charge transfer competes effectively with proton transfer when methane is used as the reagent gas. In the methane-ammonia CI mass

spectrum (bottom) the protonated molecule ions give the most intense peak and fragmentation is virtually absent. Relative responses for prominent ions in the various modes of ionization are

compared in Table 1. The values in the last column of Table 1

indicate that the best sensitivity is achieved with a mixture of

methane and ammonia as the reagent gas. These data were acquired

on two separate Finnigan 3200 GC/MS systems, one optimized for EI

and the other for CI. If maximum sensitivity is important, each

method of ionization should be evaluated on the specific GC/MS

system(s) to be used for the analysis. Sensitivities can vary

significantly from instrument to instrument.

To evaluate the reproducibility of the method, five independent

replicate analyses were performed on 1-ml samples of whole blood

containing 100 ng of phencyclidine and 50 ng of the internal standard. Methane was used as the reagent gas and ions were monitored

at m/z 159 and 164, and at m/z 243 and 248. The relative standard

deviations for the peak height ratios were 2.9 and 2.3, respectively.

32
FIGURE 1. GAS CHROMATOGRAMS SHOWING GENERATION OF 1-

PHENYLCYCLOHEXENE FROM INJECTION OF PHENCYCLIDINE AT HIGH INJECTION PORT TEMPERATURES

33
FIGURE 2. MASS SPECTRA OF PHENCYCLIDINE

34
TABLE 1. RELATIVE RESPONSES FOR PROMINENT IONS IN THE

EI AND CI MASS SPECTRA OF PHENCYCLIDINE

Relative Response

Percent of Total Per Unit Weight

Method m/z Monitored Ion Current of Drug

EI 200 (M+

 -C3H7) 14 59

CI (CH4

) 243 (M+) 24 20

CI (CH4+NH3

) 244 (MH+) 73 100

Table 2 lists phencyclidine concentrations found in various body

fluids from phencyclidine-intoxicated patients. Blood concentrations ranged from 49 ng/ml for a child in a coma, to 2.7 μg/ml

found in an adult who died following an overdose.

TABLE 2. CONCENTRATIONS OF PHENCYCLIDINE IN BODY FLUIDS

FROM PHENCYCLIDINE INTOXICATED PATENTS

Case Number Blood Urine CSF Serum

1

(Adult, fatal)

2.7 μg/ml

2

(Child, coma)

49 ng/ml 910 ng/ml 6.5 ng/ml

3

(Youth, drowsy)

51 ng/ml 870 ng/ml

4

a

171 ng/ml

5

b

73 ng/ml

aThe condition of the patient was described as "serious."

bThe condition of the patient was described as "fairly serious."

35
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR PHENCYCLIDINE

Nomenclature

Chemical Name: 1-(1-Phencyclohexyl)piperidine

Other Names: Phencyclidine, HOG, PCP, CI-395, Angel dust

Empirical Formula: C17H25N

Chemical Abstracts Registry Number: 77-10-1

Trade Names of Hydrochloride: Sernyl, Sernylan

Physical Constants

Appearance: White crystals

Melting Point: 46-46.5°C; hydrochloride, 243-244°C

Boiling Point: 135-137°C (1.0 mm)

Specific Rotation: Optically inactive

Solubility: The hydrochloride is soluble in hater, ethanol and

chloroform.

REFERENCES

1. Unsigned. PCP (Phencyclidine): The New Disillusionogen.

STASH Press, Madison, WI (1975).

2. Unsigned. Emergency Med. , 8 (3), 265 (1976).

3. P.V. Luisada and B.I. Brown. Clin. Toxicol., 9 (4), 539

(1976).

4. G.D. Lundberg, R.C. Gupta, and S.H. Montgomery. Clin. Toxicol.,

9 (4), 503 (1976).

5. A.T. Snulgin and D.E. Mac Lean. Clin. Toxicol., 9 (4), 553

(1976).

6. C. Helisten and A.T. Snulgin. J. Chromatog., 117, 232 (1976).

7. D. Gagne and R.K. Pike. J. Assoc. Off. Anal. Chem., 60, 32

(1977).

8. K.E. Garey and R.G. Heath. Life Sciences., 18, 1105 (1976).

9. R.S. Burns and S.E. Lerner. Clin. Toxicol., 12, 463 (1978).

10. S.H. James and S.H. Schnoll. Clin. Toxicol., 9 (4), 573 (1976).

36
11. D.N. Bailey, R.F. Shaw, and J.J. Guba. J. Anal. Toxicol., 2,

233 (1978).

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

F. Walker. Science, 184, 189, (1974).

R.S. Burns and S.E. Lerner. Clin. Toxicol., 9 (4), 477 (1976).

R.L. Balster and L.D. Chait. Clin. Toxicol., 9 (4), 513 (1976).

J.C. Munch. Bull. Narcotics, 26 (4), 9 (1374).

R.C. Petersen and R.C. Stillman, eds., Phencyclidine (PCP)

Abuse: An Appraisal. National Institute on Drug Abuse

Research Monograph 21, DHEW Pub. (ADM) 78-728. Supt. of Docs.,

U.S. Govt. Print. Off Washington, D.C. (August 1978).

R.E. Ober, G.W. Gwynn, T. Chang, D.A. McCarthy, and A.J.

Glazko. Fed. Proc., 22, pt. 1, 539 (1963).

E.F. Domino and A.L. Wilson. Clin. Pharmacol. Therap., 22,

421 (1977).

J.A. Marshman, M.P. Ramsay, and E.M. Sellers. Toxicol. Appl.

Pharmacol., 35, 129 (1976).

A. h. Done, R. Aronow, J.A. Miceli, and D.C.K. Lin, in B.H.

Rumack and A.R. Temple, eds., Management of the Poisoned

Patient. Science Press, Princeton, NJ, p. 79 (1977).

D.C.K. Lin, A.F. Fentiman, Jr., K.L. Foltz, R.D. Forney, Jr.,

and I. Sunshine. Biomed. Mass Spectr., 2, 206 (1975).

E.F. Domino. Int. Rev. Neurobiol., 6, 303 (1964).

L.K. Wong and K. Biemann. Biomed. Mass Spectr., 2, 204 (1975).

H.B. Hucker. Ann. Rev. Pharmacol., 10, 99 (1970).

C.B. Liden, F.H. Lovejoy, Jr., and C.E. Costello. J. Amer. Med.

Assoc., 234 (5), 513 (1975).

N.C. Jain, R.D. Budd, W.J. Leung, and T.C. Sneath. J. Anal.

Toxicol., 1, 77 (1977).

B.S. Finkle, E.J. Cherry, and D.M. Taylor. J. Chromatogr. Sci.,

9, 393 (1971).

J.-E. Lindgren, C.-G. Hammer, R. Hessling, and B. Holmstedt.

Am. J. Pharm., 141, 86 (1969).

R.C. Shaler, J.M. Parker, and A. Sage. Clin. Toxicol., 7 (3),

307 (1974).

J.K. Brown, L. Shapazian, and G.D. Griffin. J. Chromatogr.,

64, 129 (1972).

37
31.

32.

33.

34.

3.5.

36.

37.

38.

R.C. Gupta, I. Lu, G-L. Oei, and G.D. Lundberg. Clin. Toxicol.,

8 (b), 611 (1975).

P.M. Froehlich and G. Ross. J. Chromatogr., 137, 135 (1977).

P.C. Reynolds. Clin. Toxicol., 9 (4), 547 (1976).

D.S. Pearce. Clin. Chem., 22, 1623 (1976).

W.D. MacLeod, Jr., D.E. Green, and E. Seet. Clin. Toxicol.,

9 (4), 561 (1976).

L.K. Wong and K. Biemann. Clin. Toxicol., 9 (4), 583 (1976).

A. Kalir, H. Edery, Z. Pelah, D. Balderman, and G. Porath.

J. Med. Chem., 12, 473 (1969).

A.F. Fentiman, Jr., and R.L. Foltz. J. Labelled Compounds and

Radiopharmaceuticals, 12, 69 (1976).

38
Chapter 4

Methaqualone

Methaqualone (2-methyl-3-o-tolyl-4(3H) -quinazolinone, I) was first

synthesized in 1951 after the elucidation of the structure of febrifugine, the active principle of the ancient Chinese antimalarial

Ch’ang Shan, stimulated interest in the medicinal chemistry of

quinazoline derivatives (1).

It was introduced in Europe in 1956 and into the United States in

1965, advertised as a nonbarbiturate, nonaddictive sedativehypnotic, and marketed under several trade names including Quaalude,

Sopor, Optimil, Parest, and Somnafac. Within a few years, it had

become the sixth best selling sedative-hypnotic in this country

(2). Methaqualone abuse was quick to follow, and by 1972 had reached

almost epidemic proportions in the United States, although the first

case of fatal poisoning had been reported in Europe at least ten

years earlier (3).

PHARMACOLOGY

Even in therapeutic doses (150-500 mg), methaqualone can produce

a sensual, euphoric state and a relaxed, intimate mood which explains its popularity and abuse potential. However, the drug may

accumulate in the blood following repeated doses and is addictive

after prolonged use; abrupt withdrawal can result in death if not

properly supervised. Methaqualone is particularly dangerous when

consumed with alcohol. Coma and convulsions can result from acute

ingestion of 2 grams of methaqualone and death from 8 grams, but

in the presence of alcohol these consequences can result from ingestion of considerably smaller quantities of the drug (2). Since

1973, illicit use of methaqualone has dropped dramatically, apparently because of tighter control of its manufacture and distribution.

Methaqualone is a lipophilic, acid-soluble, quinazoline derivative

easily prepared from anthranilic acid, o-toludine and an acetyla39

328-011 0 - 80 - 4
ting agent. It has a characteristic ultraviolet absorption spectrum and is volatile enough for gas chromatography. Its pKa, calculated from UV spectral data, is 2.5 (4). Although resistant to

chemical hydrolysis or oxidation, it is rapidly hydroxylated by

hepatic microsomal enzymes to polar metabolites, which are excreted primarily as ß-glucuronic acid conjugates (1). Since metabolites of methaqualone are not present in significant amounts in

blood (<0.2 μg/ml) after therapeutic doses, they probably contribute very little to the overall pharmacological response to methaqualone (5) . However, a study of urinary metabolite patterns in

six human volunteers given single or repeated therapeutic doses of

methaqualone showed a higher urinary level of one metabolite, 2-

methyl-3-(2'-hydroxymethylphenyl)-4(3H)-quinazolinone, in individuals who demonstrated the most pronounced physiological response

(lower tolerance) to the drug. Since the same metabolite has been

found, unconjugated, in the blood of overdose patients, this compound may be pharmacologically active (6).

PHARMACOKINETICS AND METABOLISM

Binding studies by equilibrium dialysis have established that 80

percent of adlministered methaqualone is protein-bound in plasma

over the whole therapeutic range of concentration. Based on this

observation and on the assumption that only unconjugated methaqualone is pharmacologically active, a comparison of the levels of

unbound drug in serum with levels in cerebrospinal fluid (CSF) by

gas chromatographic measurement has indicated a passive, uninhibited

transfer of drug to the CSF. However, drugs which compete for protein binding sites could theoretically affect the transfer (7).

Other workers have also pointed out the susceptibility of the pharmacokinetics of methaqualone to influence by other drugs, especially

drugs able to influence hepatic microsomal oxidation. This is

not merely of academic interest, since methaqualone is used in many

compound formulations for treatment of a wide range of ailments,

particularly in Germany (1). In this connection, a study of the

kinetics of methaqualone in a formulation containing 250 mg methaqualone, 300 mg carbromal, and 0.33 mg benactyzine, found 6-hr

plasma levels of methaqualone to be lower than those usually reported for methaqualone alone. These results may indicate an effect

of the other drugs on methaqualone metabolism, lending support to

the claim that the pharmaceutical formulation, Staurodorm, alleviates methaqualone “hangover” (8).

The most salient features of the results of pharmacokinetic studies

of methaqualone have been the wide variation in the rate of absorption by different subjects and the faster absorption rate for the

hydrochloride salt as compared with the free base (1). Plasma

levels of methaqualone following a single oral dose of 300 mg generally reach a maximum of 1-8 μg/ml within 2 hr. The average plasma

half life has been reported as 2.6 hr for the first, rapid excretory phase (9). A slow excretory phase becomes apparent as much

as 100 hr after ingestion, and the half life of this phase is

variously reported to be from 19 to 50 hr (10,11).

40
Methaqualone is extensively metabolized in the human body so that

very little unchanged drug, usually <5 percent, is excreted in the

urine (12). In fact, the validity of even the small amounts reported has been called into question by the recent identification

of methaqualone-N-oxide in the urine of human subjects given 250 mg

of methaqualone. This metabolite accounted for 5-9 percent of the

dose in 24 hr, and since it easily undergoes thermal conversion to

methaqualone under gas chromatographic conditions, its presence

could result in mistakenly high calculations for free methaqualone

(13).

Metabolic pathways involving formation of dihydroxy and hydroxy

methoxy derivatives of methaqualone exist, but they are minor in

terms of relative concentrations of methaqualone metabolites in

urine. The major pathway for methaqualone is monohydroxylation in

the liver at any one of ten possible sites. However, hydroxylation

is semi-specific in that five of the possible monohydroxy metabolites predominate over the others in human urine; i.e., those with

substitutions at the 3’, 4’, and 6 positions and on the 2 and 2’-

methyl groups (14). The 3-hydroxy metabolite has only recently

been confirmed in human urine (15) . Stillwell and co-workers identified five dihydrodiol and two hydroxydihydrodiol metabolites

of methaqualone in the unhydrolyzed (unconjugated) fraction of human urine and also detected several unidentified triols. They interpreted these finding as evidence for metabolism of methaqualone in

man by way of epoxide intermediates. Mass spectra indicated that

three of the dihydrodiol and both hydroxydihydrodiols were formed

by epoxidation of the tolyl ring, while the mass spectra of the

other two suggested epoxidation of the quinazolinone nucleus. The

authors suggested that of the many urinary metabolites of methaqualone which have now been reported, including seven dihydrodiols, nine phenols, two methoxyhydroxyphenols, diols, and several

triols, all except the 2- and the 2’-hydroxymethyl derivatives

would be likely to involve epoxide intermediates. None of these

epoxides have yet been isolated, however (16).

ANALYTICAL METHODOLOGY

Because of the widespread use and abuse of methaqualone and because

of its interesting metabolism, virtually every currently popular

technique has been applied to the analysis of the drug. Although

methaqualone’s structure lends itself to spectroptotometric assays,

in practice such methods are rarely sensitive enough for the determination of therapeutic levels. However, they have been widely

applied in toxicological studies involving gross over-dosage. For

spectrophotometric assays, hexane extraction, followed by back

extraction with acid to recover methaqualone free from phenolic

impurities, is to be preferred to extraction with a more polar

solvent such as chloroform because the latter yields a mixture of

strongly UV-absorbing materials with UV spectra indistinguishable

from that of methaqualone (1).

A commercial kit for radioimmunoassay (RIA) has been evaluated for

clinical detection of methaqualone in both urine and blood (17,18).

While it proved to be a rapid technique which required no cumbersome

41
sample preparation, it was unable to distinguish between methaqualone and its metabolites, some of which were as reactive as the

parent compound and others less so or not at all. Metabolites present were additive in their effect on the concentration reading.

Standard RIA curves prepared in serum for methaqualone and five

metabolites demonstrated a detection sensitivity of 2 ng/ml, but

because of the contribution of metabolites to the measured levels,

RIA in the form tested would be useful as a screening procedure

only.

For quantitative studies, gas chromatographic (GC) methods have

been developed, some reporting detection levels as low as 0.01

μg/ml for methaqualone in plasma. This degree of sensitivity was

achieved in a method reported by Berry in 1969, which involved extraction of 5.0 ml plasma, after pH adjustment by addition of 1.0

ml 1N NaOH, with 15 ml of hexane followed by gas chromatography on

a column coated with 3 percent cyclohexanedimethyl succinate. Butobarbital was used as an internal standard. An extraction efficiency

of 96 percent was reported for this method (19).

An extraction efficiency of 98-100 percent was reported for a technique in which saturated NaH2PO3

 solution was added to serum in

order to precipitate proteins and lipoproteins and free the drug

from binding sites before extraction with n-hexane. After centrifugation, the n-hexane phase was evaporated under nitrogen at 57°C

and the residue dissolved in acetone containing hendecyl myristate

as an internal standard. Gas chromatography on a 3 percent OV-17

column with nitrogen as the carrier gas provided a sensitivity of

1.0 μg/ml methaqualone (20).

Quantitation of methaqualone and its five most abundant hydroxyl

metabolites in urine was done by GC in a recent study designed to

characterize metabolite patterns as possible indicators of methaqualone ingestion. Methaqualone was extracted with chloroform from

acid-hydrolyzed urine and chromatographed on 3 percent OV-1 columns

at 21O°C. To achieve complete separation, the metabolites were

silylated and chromatographed on high-resolution SE-30 SCOT columns.

Concentrations measured ranged from 0.20 to 1.24 μg/ml methaqualone

and 0.56 to 31.5 μg/ml among the five principal hydroxy metabolites.

Electron impact mass spectra of the methaqualone and the trimethylsilyl ethers of the metabolites, separated by GC on a 3 percent OV17 column at 225°C, were used to confirm identification of the metabolites (14).

Selected ion monitoring has come into wide use as a highly sensitive

and specific method for measurement of methaqualone as well as for

identification and quantitation of its metabolites. An early attempt

to use methaqualone labeled with deuterium on the 2-methyl substituent as internal standard was abandoned because of poor isotopic

purity. However, the 2-ethyl analog of methaqualone added to plasma

before extraction with diethyl ether provided an internal standard

suitable for GC/MS analysis. The mass spectrometer monitored the

ion current at m/z = 250 for methaqualone and at m/z = 264 for the

internal standard. The method permitted measurement of plasma concentrations of the drug as low as 5 ng/ml (10).

42
More recent utilization of GC/MS in methaqualone studies has led to

techniques allowing detection of 0.20 ng/ml methaqualone and its 6-

hydroxy metabolite in urine. This degree of sensitivity was achieved

by rigorous prepurification of the urine extracts by multiple thinlayer chromatography, use of multilabeled internal standards in

which the level of unlabeled compounds was < 0.01 percent, and analysis of the isotopic ratios by field ionization mass spectrometry

which produces abundant molecular ions with little or no fragmentation. Heptadeuterated methaqualone was prepared from commercial

toluene-2H8 which was converted to o-toluidine-2H7 and condensed

with N-acetyl-anthranilic acid. Heptadeuterated 6-hydroxy-methaqualone was similarly prepared by condensation of the labeled o-toluidine with 5-methoxy-anthranilic acid followed by demethylation.

The internal standards were added to enzyme-hydrolyzed urine samples before adsorption on an Amberlite XAD-2 column and elution with

ethyl acetate: chloroform (3: 2, v/v). From the concentrated eluates,

methaqualone and the 6-hydroxy metabolite were isolated by chromatography four times on thin-layer plates with, respectively, benzene:

n-butanol:methanol (85:10:5); ethyl acetate:methanol:NH4OH (85:10:1);

benzene:acetic acid (9:l); and ether. Other hydroxy metabolites

present were apparently eliminated by this procedure. The technique was applied to a study of elimination kinetics of methaqualone

over an 11-day period following ingestion of one 250-mg tablet,

demonstrating its value in long term investigations when the methaqualone concentration is likely to fall to levels undetectable by

conventional methods (10).

EXPERIMENTAL PROCEDURE

Therapeutic doses of methaqualone are relatively high (150-500 mg).

Consequently, a simple, direct extraction of 0.1 ml or more of biological fluid is usually sufficient to provide enough of the drug

to be measured by GC/MS. Two extraction procedures are described

here. Both procedures are relatively simple and rapid. The first

procedure should give slightly better recovery and provide a cleaner

extract, while the second method is even simpler and completely

avoids evaporation of the solvent extract and therefore the possibility of losing small quantities of the drug through evaporation.

In our hands, precision and accuracy were comparable with both

methods.

Standards and Reagents

The methaqualone hydrochloride used in the development of this method was purchased from Applied Science Laboratories, State College,

PA 16801. No impurities were detected by either gas chromatography or mass spectrometry. Methaqualone- 2H4

 [2-methyl- 3-o-to1yl -4

(3H)quinazolinone-5,6,7,8-2H4] was prepared from phthalimide-2H4 (21).

On the basis of GC/MS analysis this material showed a chemical purity of greater than 99 percent, and had the following isotopic composition: 2H4

, 97.8 percent; 2H3, 2.1 percent; 2H1 and 2H0 <0.1

percent.

43
Prepare a pH 9.6 buffer by dissolving 500 grams of Potassium monohydrogen phosphate (K2HP04

) in about 500 ml of distilled water.

Allow the solution to cool to room temperature and then add sufficient distilled water to make exactly 1 liter.

Stock solutions of methaqualone and methaqualone-2H4 are prepared

as follows for the purpose of determining standard curves and preparing working standards. Weigh into a 100-ml volumetric flask

about 10 mg of methaqualone, or about 11.4 mg of methaqualone

hydrochloride, and record its weight to the nearest 0.1 mg. If

the hydrochloride is used, the equivalent weight of free base is

calculated by multiplying the weight of the hydrochloride by 0.874.

Dissolve the measured methaqualone in about 75 ml of methanol and

then add additional methanol to give exactly 100 ml of solution.

‘This solution will be referred to as the “0.10-mg/ml stock solution” although it should be labeled with its actual concentration

based on the exact measured weight of methaqualone. A series of

working standard solutions can then be prepared by appropriate

dilution of the stock solution as described in Chapter 2.

The stock solution of methaqualone-2H4 is prepared in the same

manner. For the measurement of methaqualone in body fluids within

the concentration range 1 to 1000 ng/ml, addition of approximately

40 ng of the internal standard to each ml of body fluid has proven

satisfactory. This can be conveniently done by adding 100 μl of a

400-ng/ml methaqualone-2H4 methanolic solution to each ml of body

fluid. Prepare the 400-ng/ml internal standard solution by diluting 1 ml of the 0.10-mg/ml stock solution of methaqualone- 2H4

 to

250 ml with methanol.

Store stock and working standard solutions in well-stoppered or

capped glass vessels in the dark at less than 0°C.

Extraction—Method 1

Transfer 1 ml of body fluid (whole blood, plasma or urine) to a

20-ml culture tube equipped with a Teflon-lined screw cap (0.5 to

2 ml of specimen can be used, but it must be measured accurately).

Add 100 μl of the 400-ng/ml methaqualone-2H4 solution and vortex

for 10 sec. Allow the sample to equilibrate for about 15 min and

then add 1 ml of K2HPO4 buffer (pH 9.6) and 5 ml of methylene

chloride. Cap the tube and check for a leak around the cap. Gently

mix the contents for 30 min using a motorized rocker or rotator.

Centrifuge for 5 min, remove the upper aqueous layer by aspiration,

and then transfer by means of a disposable Pasteur pipette the methylene chloride extract (bottom layer) to a silylated concentrator

tube (>5 ml volume) having a conical or nipple shaped bottom. Add

20 μl of dimethylformamide to act as a “keeper solvent” to minimize

evaporative loss of the methaqualone. Remove the methylene chloride

by evaporation under a gentle stream of nitrogen or filtered air

while heating at 50 to 60°C. When the volume of extract decreases

to about 20 μl, stopper the tube or cover it with Parafilm and store

at <0°C until the GC/MS analysis is ready to be performed. Allow

the tube to warm to room temperature immediately prior to the analysis.

44
Extraction—Method 2

Transfer 1 ml of body fluid (whole blood, plasma, or urine) to a

5-ml glass stoppered, conical centrifuge tube. Add 100 μ1 of the

400-ng/ml methaqualone-2H4 internal standard solution and vortex

for 10 sec. Allow the sample to equilibrate for 15 min and then

add 1 ml of K2HPO4 buffer (pH 9.6), followed by 100 μl of a solvent

consisting of toluene:heptane:isoamyl alcohol in a volume ratio of

70:20:10. Stopper the tube and vortex for at least 30 sec. Separate the phases by centrifugation at 2000 x G for 15 min. The

organic solvent should form a narrow upper layer from which an aliquot can be removed with a syringe for injection into the GC/MS.

GC/MS Analysis

The experimental conditions for the GC/MS analysis of methaqualone

are as follows:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-1 or OV-17 on 100/120 mesh Gas

Chrom Q (Applied Science Laboratory, State

College, PA 16801)

Carrier gas: Methane, 15-20 ml/min

Reagent gas: Ammonia, introduced into the ion chamber as

described in Chapter 2

Temperatures: Injector, 270°C

Column, 210°C isothermal

GC/MS transfer line, 260°C

Ion source, 160°C

Under these conditions the methaqualone and methaqualone-2H4 should

elute at about 4 min as narrow, symmetrical peaks.

Prior to beginning GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described

in Chapter 2. The ions to be monitored occur at m/z 251 and 255.

They correspond to the protonated molecule ions for methaqualone

and methaqualone-2H4, respectively. With the divert valve in the

divert position, inject the organic extract into the gas chromatograph (2 to 4 μ1 of the extract from Method 1 or 5 to 10 μ1 of the

extract from Method 2). After approximately 1 min switch the divert

valve so that the entire carrier gas flow enters the ion source of

the mass spectrometer and begin data acquisition. When the methaqualone peaks nave eluted, cease data acquisition and return the

divert valve to the divert position.

The quantity of methaqualone in the specimen is determined by measuring the heights (or areas) of the methaqualone and methaqualone2H4

 peaks in the selected ion current profiles (“ion chromatograms”)

and relating the ratio of peak heights to a calibration graph. Calculate the concentration of methaqualone by dividing the measured quantity of methaqualone by the exact volume of speciman extracted.

45
Discussion of the Experimental Procedure

The distribution ratios of methaqualone between aqueous phases over

a range of pH values and various organic solvents were measured in

order to determine the optimum conditions for extraction of the

drug from biological fluids. For this purpose a stock solution of

methaqualone (100 μg/ml) in 0.1N HCl was prepared. A 5-ml aliquot

of the stock solution was adjusted to a specific pH and transferred

to a 25-ml screw-cap tube along with 5 ml of the organic solvent.

The tube was rocked gently for 30 min and then centrifuged. The

optical densities of both the aqueous and organic layers were measured from 350 to 250 nanometers using a Cary 14 recording spectrophotometer. Water or the appropriate solvent was placed in the

reference cell.

The results indicated that methaqualone is extracted by chloroform

with nearly equal efficiency at all pH values between 2 and 12. At

pH 10 chloroform and methylene chloride are almost equally efficient

extraction solvents, while hexane and cyclohexane are less effective. At pH 2 all of the solvents investigated extract a significant amount of methaqualone. Consequently, a back extraction into

acid, which is often used in drug extractions for removal of nonbasic components, would result in loss of an appreciable proportion

of the methaqualone. Based on these data, we recommend that the

pH of the biological fluid be adjusted to 9-10 before extraction

with methylene chloride,

Since methaqualone contains no NH or OH groups, derivatization before gas chromatography is not necessary. The free drug gas chromatographs well on a variety of GC column packings. both 3 percent

OV-1 and 3 percent OV-17 on 100/120 mesh Gas Chrom Q have been used

successfully, but there is a slight preference for the former because a significantly lower column temperature can be employed.

The electron impact (EI) and chemical ionization (CI) mass spectra

of methaqualone are shown in Figure 1. These spectra were obtained

on quadrupole mass spectrometers (22). The relative sensitivities

achieved by monitoring the ion current at the m/z value corresponding

to the most abundant ion generated by each of the ionization processes are shown in Table 1. If maximum sensitivity is desired, CI

with methane and ammonia as reagent gases should be used. This combination gives slightly better sensitivity than that provided by

methane alone as reagent gas, and is also a more selective mode of

ionization. However, since plasma levels following therapeutic

doses of methaqualone are moderately high (~5 μg/ml), sensitivity

is usually not a problem. At higher ion source temperatures

(>160°C) under methane CI conditions fragment ions at m/z 118 become

more abundant. Since the corresponding fragment ion in the methane

CI mass spectrum of the methaqualone-2H4 retains the deuterium

labels, it appears at m/z 122. Consequently, the ions at m/z 118

and 122 can be monitored in addition to the protonated molecule ions

at m/z 251 and 255 to provide confirmational data.

46
FIGURE 1. MASS SPECTRA OF METHAQUALONE

47
TABLE 1. RELATIVE RESPONSES FOR PROMINENT IONS IN THE

EI AND CI MASS SPECTRA OF METHAQUALONE (22)

Percent of Relative Response

Ionization m/z Sample Ion Per Unit Weight

Method Monitored Current of Drug

EI 250 (M+) 4 6

235 (M+ - CH3) 1.5 20

CI (CH4) 251 (MH+) 57 84

Cl (CH4 + NH3) 251 (MH+) 7 4 100

When solutions of methaqualone and cicosane (internal standard) in

methylene chloride were prepared and stored in capped vials at room

temperature, 2°C, and -10°C, gas chromatographic analyses of the

solutions over a period of six weeks showed no significant changes

in the ratio of eicosane and methaqualone peak heights, indicating

that methaqualone is stable under these conditions.

48
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR METHAQUALONE

Nomenclature

Chemical Name: 2-Methyl-3-o-tolyl-4(3H)-quinazolinone

Empirical Formula: C16H14N2O

Chemical Abstracts Registry Number: 72-44-6

Trade Names: Many, including MAOA, MTZ, TR 495, Catendyl, Dormogen,

Fadormir, Hyprol, Melsomin, Mebedorm, Normi-Nox, Omnyl, Parminol,

Parest, (Quaalude, Roulone, Somnafac, Sonal, Somberol, Soxor, Forinol, Tuazolone

Physical Constants

Appearance: White crystalline powder

Melting Points: Free base: 114-116°C

Hydrochloride: 255-265°C

Specific Rotation: Optically inactive

Solubility: The free base is practically insoluble in water; soluble

in ethanol, ether, and chloroform

pKa: 2.54 (4)

REFERENCES

1. S.S. Brown. In L, Reid, ed., Methodological Developments in

Biochemistry, Vol. 5, p. 179. North-Holland, Amsterdam (1976).

2. J.A. Ostrenga. Clin. Toxicol., 4, 607 (1973).

3. S.S. Brown and S. Goenechea. Clin. Pharmacol. Ther., 14, 314

(1973).

4. J.J. Zalipsky, D.M. Patel, R.J. Darnowski, and N.H. ReaveyCantwell. J. Pharm. Sci., 65 (3), 460 (1976).

5. A.F. Delong, R.D. Smyth, A. Polk, R.K. Nayak, and N.H. ReaveyCantwell. Arch. Int. Pharmacodyn., 222 (2), 322 (1976).

6. L. Kazyak, J.A. Kelley, J.A. Cella, R.E. Droege, L.R. Hilpert,

and R.C. Permisohn. Clin. Chem., 3, 2001 (1977).

7. J.M. Christensen and S. Holfort. J. Pharm. Pharmacol., 2, 538

(1975).

8. C. White, E. Doyle, L.F. Chasseaud and T. Taylor. Europ. J.

Clin. Pharmacol., 10, 343 (1976).

49
9. R.N. Morris, G.A. Gunderson, S.W. Babcock, and J.F. Zaroslinski.

Clin. Pharmacol. Ther., 13, 719 (1972).

10. G. Alvan, J.E. Lindgren, C. Bogentoft, and O. Ericsson. Europ.

J. Clin. Pharmacol., 6, 187 (1973).

11. J.M. McReynolds, H. d'A. Heck, and M. Anbar. Biomed. Mass

Spectr., 2, 299 (1975).

12. R. Bonnichsen, K. Dimberg, Y. Marde, and K. Ryhage. Clin.

Chim. Acta, 60, 67 (1975)

13. C.N. Reynolds and K. Wilson. Xenobiotica, 6 (2), 113 (1976).

14. R.C. Permisohn, L.K. Hilpert, and L. Kazyak. J. Forensic Sci.,

21 (1), 98 (1976).

15. D. Burnett, C.N. Reynolds, K. Wilson, and J.K. Francis.

Xenobiotica, 6 (2), 125 (1976).

16. W.G. Stillwell, P.A. Gregory, and M.G. Horning.

olism and Disposition, 3 (4), 287 (1975).

Drug Metab17. A.K. Berman, J.P. McGrath, K.C. Permisohn, and J.A. Cella,

Clin. Chem., 21, 1878 (1975).

18. R.O. Bost, C.A. Sutheimer, and I. Sunshine. Clin. Chem., 22,

689 (1976).

19. D.J. Berry. J. Chromatogr., 42, 39 (1969).

20. D. Chin and E. Fastlich. Clin. Chem., 20, 1382 (1974).

21. A.F. Fentiman, Jr. and R.L. Foltz. J. Labelled Compounds and

Radiopharmaceuticals, 12, 69 (1976).

22. K.L. Foltz, D.A. Knowlton, D.C.K. Lin, and A.F. Fentiman, Jr.

Proceedings of the Second International Conference on Stable

Isotopes, Oak Brook, IL, October 20-23, 1975.

50
Chapter 5

Methadone

Studies on the pharmacology of methadone (I) began in 1946 at which

time the compound was found to be a potent narcotic analgesic (1).

Further investigations of the analgesic effects of methadone in man

(2) led to its subsequent introduction for the control of severe

pain. In contrast to morphine, methadone is effective when taken

orally. In 1963 methadone was introduced as an experimental drug

for the treatment of heroin dependence (3). Methadone dosage can

be maintained or gradually decreased in order to overcome physical

dependence. This “detoxification” process is generally conducted

over a prolonged period during which counseling and other therapy

are also employed. Unfortunately, methadone has considerable abuse

potential so that distribution of the drug must be carefully controlled.

METABBOLISM AND PHARMACOKINETICS

The primary metabolic pathway for methadone in man is N-demethylation followed by cyclization to give a pyrrolidine metabolite (II)

(4):

Additional metabolites, resulting from further N-demethylation,

aromatic ring hydroxylation, and degradation of both side chains,

have been identified.

The pharmacokinetics of methadone in man has been extensively

studied (5,6,7). One study (5) compared acute and chronic administration of methadone to postaddict volunteers. Following a

single 15-mg oral dose, the methadone plasma concentration showed

a biexponential decay; the first phase corresponded to a plasma

elimination half-life of approximately 14 hr, and the second phase

a half-life of approximately 55 hr. The methadone plasma concen51
tration measured at 2 hr after administration was 85 ± 10 ng/ml.

During chronic oral administration of the drug, the plasma concentration underwent a single exponential decay with an average halflife of 22 hr. Patients maintained on daily 40-mg doses showed

average methadone plasma concentrations of 182 ± 35 ng/ml, while

those receiving 80-mg doses showed an average daily methadone concentration of 420 ± 97 ng/ml of plasma. Another study (8) found

a plasma elimination half-life of 7.3 hours following intramuscular

administration of 10 mg of methadone. More recently, marked variations in the pharmacokinetics of the drug between individual human

subjects nave been documented among 12 in-hospital methadone maintenance patients, among whom dose to plasma concentration ratios

varied widely from one subject to another, as did the plasma halflife of methadone (7 to 21 hr). Recoveries of the drug from urine

ranged from 23 percent to 52 percent of total intake in 24-hour

samples (9).

Methadone is eliminated from the body primarily as the parent drug

(I) and the pyrrolidine metabolite (II). Following a single dose,

urine was found to contain about half as much of the metabolite as

of the parent drug, expressed as percentage of daily dose in 24-

hour collections, while chronic administration of the drug increased the percentage of the metabolite to twice that of the drug.

The metabolite was always the predominant form in feces (5). The

increased rate of metabolism and excretion during chronic administration suggests that induction of hepatic microsomal enzymes

responsible for drug metabolism may play a significant role in

development of tolerance to the drug. However, it has been shown

in the rat that methadone does not enhance microsomal drug metabolizing enzyme systems (10). Postmortem examination of tissues from

methadone users demonstrated the presence of both the drug and the

metabolite in liver, kidney, spleen, and brain; and especially high

concentrations in the lung (11).

ANALYTICAL METHODOLOGY

Sensitive and specific methods for analysis of methadone in biological fluids are necessary in order to assess more accurately the

pharmacokinetics of the drug, and also to contribute analytical

information for correlation with the clinical status of patients

who ingest the drug in both maintenance and overdose situations.

Thin-layer chromatography (TLC) (12), UV spectrophotometry (13),

fluorimetry (14), biological assay (15), radioisotopes (16), gas

chromatography (GC) (17-19), and gas chromatography/mass spectrometry (20,21) have been used for detection and quantitation of

methadone.

Immunoassay techniques, which are well suited for the analysis of

large batches of samples, are available for detection of methadone in urine, and are highly sensitive. Radioimmunoassay (RIA)

can detect methadone in urine following doses of as little as 2.5

mg/day, and shows promise as a screening technique because propoxyphene and its metabolites do not interfere as they often do in

TLC and GC procedures (22). However, the antibodies used for RIA,

52
hemagglutination inhibition, or the enzyme multiplied immunoassay

(EMIT) do not react with the primary metabolite II at concentrations of 1000 μg/ml or less. Because the concentration of II is

often equal to or greater than the concentration of methadone in

urine, its detection is important at low drug concentrations. Furthermore, none of the immunological methods are quantitative over

the normal range of methadone concentrations found in urine (23).

The most satisfactory methods for quantitation of methadone in biological fluids have been found to involve gas chromatography. The

combination of GC with electron impact mass spectrometry (GC/EI-MS)

has been used for quantitation of methadone in human plasma (20).

A homolog of methadone was used as the internal standard in order

that both unlabeled and deuterium-labeled methadone could be administered and simultaneously measured in plasma. The method involved

equilibration of 4 ml of plasma with the internal standard, adjustment of the pH to 9.5, and extraction with 1-chlorobutane. The GC

column was packed with 1 percent UCW-98 and operated at 190°C. Ions

monitored corresponded to M

+

 - CH3 for the methadone (m/z 294),

methadone-2H3 (m/z 297), and the internal standard (m/z 308). The

sensitivity of the method was limited by the relatively low abundance of the ions monitored. Under electron impact ionization

most of the ion current for methadone is carried by low mass fragment ions which are unsuitable for selected ion monitoring. Nevertheless, the method is reported to be capable of measuring methadone

concentrations as low as 5 ng/ml with a relative standard deviation

of less than 4 percent.

A GC/MS procedure employing chemical ionization has been reported

to be about five times more sensitive than the electron impact ionization method because of greater relative abundance of ions in the

region of the molecular ion in the CI mass spectrum of methadone

(21). Methadone containing five deuterium atoms attached to one

aromatic ring was synthesized for use as the internal standard in

this assay. Extraction from plasma or urine, after addition of the

deuterated standard and adjustment to pH 9, was with ethyl acetate,

followed by back extraction into 0.05 N HCl. Gas chromatography

was done on a column of 10 percent W-98 on Gas Chrom Q at 180°C.

Ion abundance ratios were measured using isobutane chemical ionization in which the protonated molecule ions at m/z 310 for methadone

and 315 for 2H5-methadone were monitored.

Methadone-2H10 as well as the corresponding decadeuterated methadone metabolites have been synthesized for application to metabolic

studies and as internal standards for GC/MS analysis (24). The

labeled compounds have also proven useful in elucidating the electron

impact fragmentation processes for methadone and its metabolites.

A GC method reported for quantitation of methadone using an electron

capture detector (ECD) involved extraction of methadone from plasma

with n-heptane followed by back extraction into an acidic aqueous

phase. The aqueous extract was then oxidized with barium oxide to

convert the methadone to benzophenone in order to obtain a stronger

ECD response. The method was reported to have sensitivity comparable to the GC/EI-MS technique (17).

53
Several methadone assays using GC with conventional flame-ionization detectors (FID) have been published. In one of them, designed

for determination of methadone in the urine of addicts in a methadone maintenance program, 10 ml of urine was adjusted to pH 9.5

and extracted with chloroform- isopropanol (3:1). The GC column

was packed with 3 percent OV-17 and operated at 240°C. 4-Benzylbiphenyl was used as the internal standard. The method can quantitate methadone in urine within the concentration range 3 to 30

μg/ml with a relative standard deviation of 7 percent (18). Another GC method measured methadone concentrations in blood, plasma,

or urine by a procedure consisting of the addition of n-docosane

as the internal standard, extraction of the body fluid (adjusted

to pH 9.5) with 1-chlorobutane, and gas chromatography on a 4- ft

1 percent W-98 column maintained at 165°C. The minimum concentration of methadone that could be measured by this procedure was 15

ng/ml using 3 ml of blood or plasma (19). A similar initial extraction procedure (1-chlorobutane) was used in another GC assay,

but further purification of the extracted methadone was achieved

by back extraction into 0.2 N HCl. The aqueous acid extract was

then made alkaline and extracted with chloroform. A drug (SKF

525-A) with a structure similar to methadone was used as the internal standard. The gas chromatographic analysis employed a 3 percent SE-30 column operated at 200°C. This method can detect as

little as 15 ng/ml of methadone in 4 ml of plasma (8).

The GC-FID technique using SKF 525-A as an internal standard has

been adapted to the simultaneous determination of methadone, the

pyrrolidine metabolite, cocaine, methaqualone, phencyclidine, and

propoxyphene in small (3 ml) volumes of urine, with detection levels as low as 0.5 μg/ml. The drugs are extracted together at basic pH by toluene: isoamyl alcohol:heptane (76:4:20) and converted

to their less volatile hydrochloride salts by addition of 1 percent HCl in methanol; the reported extraction efficiency for methadone and its metabolite is 100 percent. The residue after evaporation is reconstituted in methanol and subjected to GC on a 2 percent W-98/1 percent OV-17 mixed bed column. For optimal accuracy

in this procedure, silylated glassware and a fresh buffer solution

for the internal standard are required (25). A recently reported

improvement in the extraction procedure for methadone increased

the sensitivity of a GC-FID assay to 5 ng methadone per ml in a

15-ml sample of whole blood. The method involved extraction of

methadone and internal standard (2-dimethylamino-4,4-diphenyl-5-

nonanone, specially synthesized for the analysis) with 1-chlorobutane at pH 9.8, back extraction into 0.5M H2SO4

, alkalinization,

and reextraction into chloroform. Sulfuric acid was used instead

of HCl because the sulfate salt of methadone has a better partition

coefficient (1-chlorobutane:water) than the hydrochloride salt at

pH 0 to 2. In some samples, the aqueous phase had to be equilibrated with 2,2,4-trimethylpentane to break the emulsion formed

after equilibration of the H2SO4 and 1-chlorobutane layers. No

evaporation step was required. Recovery of 14C-labeled methadone

from whole blood by this procedure was ~93 percent. Chromatography was done on 1.5 percent OV-101 on Gas Chrom Q with temperature

programmed from 170- 250°C (26) .

54
EXPERIMENTAL PROCEDURE

A relatively simple extraction procedure is used for removing methadone from biological fluids. The procedure consists of addition of

deuterium-labeled methadone to the specimen together with a pH 9.5

buffer, followed by extraction with 1-chlorobutane. Chemical ionization with methane and ammonia as reagent gases is used for the

GC/MS analysis.

Standards and Reagents

d,1-Methadone (6-dimethylamino-4,4-diphenyl-3-heptanone) was purchased as the hydrochloride salt from Regis Chemical Company, Morton

Grove, IL 60053. Analysis by gas chromatography and mass spectrometry indicated the material was better than 99 percent pure. The

methadone-1,1,1-2H3 hydrochloride was obtained from the National

Institute on Drug Abuse. The deuterated methadone was chromatographically pure (> 99 percent) and, based upon its ammonia CI mass

spectrum, contained an isotopic composition of 2H3, 96.9 percent;

2H2

, 1.9 percent; 2H1, 0 percent; and 2H0, 1.2 percent.

A suitable pH 9.6 buffer can be prepared by dissolving 500 grams

of potassium monohydrogen phosphate (K2HPO4) in about 500 ml of

distilled water. When all of the salt is dissolved, allow the solution to cool to room temperature and add sufficient water to give

exactly 1 liter.

Stock solutions of methadone and methadone-2H3 used for the determination of calibration graphs (Chapter 2) and as reference standards,

are prepared as follows. Weigh into a 100-ml volumetric flask 11.2

mg of methadone hydrochloride. Dissolve the methadone hydrochloride in methanol and bring to exactly 100 ml with additional methanol.

The resulting stock solution corresponds to a concentration of 0.10

mg/ml of methadone calculated as the free base. A series of standard solutions can then be prepared by appropriate dilution of the

stock solution as described in Chapter 2.

The stock solution of methadone-2H3 is prepared in the same manner.

For the measurement of methadone in blood or urine within the concentration range 1 to 1000 ng/ml, addition of approximately 40 ng

of the internal standard to each ml of specimen is satisfactory.

This can be conveniently done by adding 100 μ1 of a 400-ng/ml methadone-2H3

 methanolic solution to each ml of specimen. Prepare the

400-ng/ml standard solution by diluting 1 ml of the 0.10-mg/ml stock

solution of methadone2H3 to 250 ml with methanol. Store stock and

standard solutions in the dark in well-stoppered or capped glass

vessels at 0°C or lower. At room temperature, acidic solutions of

methadone are unstable; they undergo approximately 40 percent decomposition within two weeks.

Extraction

Transfer 1 ml of specimen (whole blood, plasma, or urine) to a 20-ml

culture tube equipped with a Teflon-lined screw cap (0.5 to 2 ml of

specimen can be used, but it must be measured accurately). Add 100

μ1 of the 400-ng/ml methadone-2H3 internal standard solution to the

55

328-011 0 - 80 - 5
specimen and vortex for 10 sec. Allow the sample to equilibrate

for about 15 min and then add 1 ml of pH 9.5 buffer and 5 ml of

1-chlorobutane. Cap the tube and check for a leak around the cap.

Gently mix the contents for 30 min using a motorized rocker or rotator. Centrifuge for 5 min and then transfer the 1-chlorobutane

layer (top) to a silylated concentrator tube at least 5 ml in volume and having a conical or nipple-shaped bottom. Add 20 μl of dimethylformamide as a “keeper” solvent to minimize evaporative loss

of the methadone. Remove the 1-chlorobutane by evaporation with a

gentle stream of nitrogen or filtered air while heating at 40-50°C.

When the extract volume has decreased to about 20 μl, stopper the

tube or cover the top with Parafilm and store at less than 0°C

until the GC/MS analysis is to be performed. Immediately prior to

the analysis, allow the tube to warm to room temperature.

GC/MS Analysis

The following experimental conditions are satisfactory for GC/MS

analysis of methadone:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier gas: Methane, 15-20 ml/min

Temperatures: Injector, 260°C

Column, 210°C

GC/MS transfer line, 210°C

Ion source, 160°C

Under these conditions methadone and methadone-2H3 should elute at

between 3 and 5 min as narrow, symmetrical peaks.

Before beginning the GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described

in Chapter 2. The ions to be monitored occur at m/z 310 and 313.

They correspond to the protonated molecule ions for methadone and

methadone 2H3, respectively. With the divert valve in the divert

position, inject 2 to 6 μl of the extract into the gas chromatograph. After approximately 1 min switch the divert valve so that

the entire carrier gas flow enters the ion source of the mass spectrometer, and begin data acquisition. When the methadone peaks have

eluted, cease data acquisition and return the divert valve to the

divert position.

The quantity of methadone in the specimen is determined by measuring

the heights (or areas) of the methadone and the methadone-2H3 peaks

in the selected ion current profiles (“ion chromatograms”) and relating the ratio of peak heights to a previously established calibration graph. Calculate the concentration of methadone in the biological sample by dividing the measured quantity of methadone by

the exact volume of specimen used in the analysis.

56
Discussion of the Experimental Procedure

The choice of 1-chlorobutane as the extraction solvent was based

primarily on the work of Sullivan and Blake (19) who showed it to

be one of the most efficient solvents for removal of methadone from

biological fluids. Furthermore, it extracts minimal amounts of

endogenous materials. The extraction efficiency is pH dependent,

with a maximum recovery of 85 percent when extractions are made at

pH 9.5 to 10.0.

Of the GC column packings evaluated, 3 percent OV-17 on 100/120 mesh

Gas Chrom Q gave the best combination of good resolution and minimum column bleed, as well as a suitable methadone retention time.

The electron impact (EI), methane chemical ionization (CICH4), and

methane-ammonia chemical ionization (CICH-NH3) mass spectra of

methadone are shown in Figure 1. The EI mass spectrum is dominated

by an intense peak at m/z 72 corresponding to the fragment ion

CH3CH=N (CH3) 2

+

. This ion is unsuitable for selected ion monitoring because it occurs at a m/z value where interfering ions are

likely, and even more important, the corresponding fragment ion

from the deuterated methadone does not retain the deuterium label.

None of the higher mass ions in the EI mass spectrum are sufficiently abundant to be satisfactory for selected ion monitoring. With

methane alone as the reagent gas, the protonated-molecule ion at

m/z 310 and the fragment ion at m/z 265 are abundant and are satisfactory for selected ion monitoring. Monitoring an abundant fragment ion in addition to the protonated molecule ion provides additional confidence in the reliability of the quantitative measurements (see Chapter 2). However, addition of ammonia to the methane

reagent gas results in approximately a fivefold increase in the ion

current generated at m/z 310 by a given amount of methadone (Table

1). Also, the contribution of the methadone-2H3 internal standard

to the m/z 310 ion current is slightly less (1.0 percent vs. 1.9

percent) under ammonia CI than under methane CI. Apparently, in

the latter case, some deuteride abstraction occurs. Consequently,

small amounts of methadone can be measured more accurately in the

presence of large amounts of internal standard when ammonia CI is

used. Finally, addition of ammonia to the reagent gas results in

a more selective ionization since nonbasic compounds are not ionized. In our experience, when methane alone was used, interfering ions prevented reliable quantitation of low concentrations of

methadone ( < 10 ng/ml) in Plasma. The interferences were not a problem when a methane-ammonia mixture was used as the reagent gas.

57
FIGURE 1. MASS SPECTRA OF METHADONE

58
TABLE 1. RELATIVE RESPONSES FOR PROMINENT IONS IN

THE EI AND CI MASS SPECTRA OF METHADONE

Ionization m/z

Method Monitored

EI 309 (M+

)

Percent Relative Response

of Sample Per Unit Weight

Ion Current of Drug

< 0.1 < 0.1

CH3 

72 (CH= 54 750

CI (CH4) 310 (MH+) 24 30

CI (CH4 + NH3) 310 (MH+) 76 100

59
PHYSICAL, CHEMICAL AND SPECTROMEIRIC DATA FOR METHADONE

Nomenclature

Chemical Name: d,1-6-Dimethylamino-4,4-diphenyl-3-heptanone

Empirical Formula: C21H27NO

Chemical Abstracts Registry Numbers: d,1-Methadone, 297881

d,1-Methadone hydrochloride,

125564

Trade Names: Amidone hydrochloride, Adanon hydrochloride, Dolophine

hydrochloride, Physeptone, Heptadon hydrochloride

Physical Constants

Melting Points: 1-Methadone: 99-100°C

1-Methadone hydrochloride: 241°C

d,1-Methadone: 78°C

d,1-Methadone hydrochloride: 233-236°C

Refractive Index: d,1-Methadone hydrochloride (n25

D) = 1.5299

Specific Rotation: 1-Methadone

1-Methadone

pKa: 8.25

Solubility of d,l-Methadone HCl: Water 12 g/100 ml

Ethanol 9 g/100 ml

Isopropanol 2.4 g/100 ml

REFERENCES

1.

2.

3.

4.

5.

6.

C.C. Scott and K.K. Chen. J. Pharmacol. Exp. Ther., 87,

63 (1946).

H. Isbell, A.J. Eisenman, A. Wikler, and K. Frank. J. Pharmacol. Exp. Ther., 92, 83 (1948).

V.P. Dole and M. Nyswander. J. Amer. Med. Assoc., 193, 646

(1965).

A.H. Beckett, J.F. Taylor, A.F. Casy, and M.M.A. Hassan.

J. Pharm. Pharmacol., 20, 754 (1968).

K. Verebely, J. Volavka, S. Mulé, and R. Resnick. Clin.

Pharmacol. Ther., 18, 180 (1976).

E. Anggård, L.-M. Gunne, J. Holmstrand, R.E. McMahon, C.-G.

Sandberg, and H.R. Sullivan. Clin. Pharmacol. Ther., 17, 258

(1975).

60
7. M.J. Kreek. New York State J. Med., 73, 2773 (1973).

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

C.E. Inturrisi and K. Verebey. J. Chromatogr., 65, 361 (1972).

A.K. Dhar and H. Kutt, Fed. Proc., 35, 565 (1976).

A.P. Alvares and A. Kappas. J. Lab. & Clin. Med., 79, 439

(1972).

A.E. Robinson and F.M. Williams. J. Pharm. Pharmacol., 23,

353 (1971).

C.M. Davis and D.C. Fenimore. J. Chromatogr., 104, 193 (1975).

H.E. Hamilton, J.E. Wallace, and K. Blum. J. Pharm. Sci., 63,

741 (1974).

E.J. McGonigle. Anal. Chem., 43, 966 (1971).

C.T. Liu and F.L. Adler. J. Immunol., 111, 472 (1973).

A.L. Misra, R. Bloch, N.L. Vadlamani, and S.J. Mulé. J. Pharmacol. Exp. Ther., 188, 34 (1974).

P. Hartvig and B. Näslund. J. Chromatogr., 111, 347 (1975).

J.L. Valentine, P.E. Wiegert, and K.L. Charles. J. Pharm.

Sci., 61, 796 (1972).

H.R. Sullivan and D.A. Blake. Res. Commun. Chem. Pathol.

Pharmacol., 3, 467 (1972).

H.R. Sullivan, F.J. Marshall, R.E. McMahon, E. Anggard,

L.M. Gunne, and J.H. Holmstrand. Biomed. Mass Spectr., 2,

197 (1975).

D.L. Hachey, M.J. Kreek, and D.H. Mattson. J. Pharm. Sci.,

66, 1579 (1977).

D.L. Roerig, R.I.H. Wang, M.M. Mueller, D.L. Lewand, and

S.M. Adams. Clin. Chem., 22, 1915 (1976).

N.C. Jain, W.J. Leung, T.C. Sneath, R.D. Budd, and D. Chinn.

J. Anal. Toxicol., 1, 6 (1977).

G.I. Kang, F.S. Abbott, and R. Burton. Biomed. Mass Spectr.,

6, 179 (1979).

N.C. Jain, D.M. Chinn, T.C. Sneath, and R.D. Budd. J. Anal.

Toxicol., 1, 192 (1977).

R.K. Lynn, R.M. Leger, W.P. Gordon, G.D. Olsen, and N. Gerber.

J. Chromatogr., 131, 329 (1977).

61
Chapter 6

Tetrahydrocannabinol (THC)

and Two of its Metabolites,

11-Hydroxy- THC

and 11-nor-9-Carboxy- THC

Use of the resin of the hemp plant, Cannabis sativa, for healing

and euphoriant purposes is as old as recorded history. From written

references to its use as a surgical anesthetic in the Chinese pharmacopeia of the 15th century B.C. until modern times, the history

and lore of cannabis have covered a wide range of medical applications, some without value, as in leprosy, and others apparently

justified (1). Its use as a hallucinogen and euphoriant has had

a colorful history as well; the term “assassin” was brought to

Europe by returning crusaders in reference to an 11th century

Arabic religious sect whose devotees (hashishin) allegedly used

the drug to induce ecstatic visions of paradise before setting out

on potentially suicidal missions of political murder. Known in

various parts of the world as “ganja,” “bhang,” “kif,” “hashish,”

or “marijuana,” cannabis in its crude form is eaten or smoked.

The potency of the preparations varies widely according to botanical strain and the parts of the plant used (2). Cannabis retains

its biological activity for long periods when stored in ethanol or

in sesame oil, but the activity is reduced by exposure to light

and oxygen, which helps account for variations in results of early

studies concerning its pharmacology and psychotomimetic effects (3,4).

PHARMACOLOGY

The systematic study of the pharmacology of cannabis has evolved

only in the past 12 years, following the total synthesis of 9-

tetrahydrocannabinol (THC*, I) the principal active component of the

plant material (5). Recent pharmacological research has shown the

drug to have effects that are potentially therapeutic, such as reduction of intraocular pressure, bronchodilation, anticonvulsant

action, and retardation of tumor growth (6,7,8). However, the psychotomimetic effects of cannabis, while having some potential for

therapeutic use in themselves (analgesia, antidepressant action,

etc.), have led to widespread abuse with societal and public health

implications. For example, the contribution of cannabis intoxication

to traffic accidents is the subject of intensive study (9), because

*In this chapter the term “cannabis” will be used to refer to plant

preparations containing a mixture of cannabinoids, while “THC” will

represent isolated or synthetic 9-tetrahydrocannabinol, even though

at least one other active form, 8-THC, is present in small amounts

in some extracts.

62
the principal subjective effects of THC are distortions of perception and time, and impairment of short term memory. THC acts on

the septal area of the limbic system of the brain, where structures

controlling emotional behavior are located (10).

The interaction of THC with other drugs is also of practical interest. THC attenuates the morphine-abstinence syndrome precipitated by naloxone, but exacerbates alcohol withdrawal symptoms,

and it prolongs ether anesthesia. Various cannabis components

are known to inhibit liver enzymes, which prevents detoxification

of other drugs (3).

Toxicological studies have indicated carcinogenic potential in

smoked cannabis; smoke condensates contain considerably higher

amounts of polynuclear hydrocarbons than tobacco smoke (11). Cardiovascular effects of THC include hypotension and tachycardia, and

reduction of blood flow to certain areas of the brain has been shown

in test animals. Tolerance to the cardiac and blood pressure effects

as well as to the psychotropic effects can develop (12). No irreversible pathological changes have been shown to result from THC administration; however, there are reports which suggest possible effects

of THC on immune responses, endocrinological function, and cell metabolism. These may have important clinical implications, especially

in view of the reported persistence of THC in the body in the form

of lipid conjugates (13). Because of its high lipid solubility, THC

is slowly eliminated from the body, and it accumulates in tissues.

It has been reported that even minute amounts of THC disrupt cell

metabolism, preventing formation of DNA, RNA, and protein and retarding cell division. There also appears to be an increased incidence of abnormal forms of spennatoza and a reduced sperm count in

male cannabis users (10). The chemistry and pharmacology of THC

have been the subjects of extensive recent reviews (14,15,16).

PHARMACOKINETICS AND METABOLISM

THC has low solubility in water and is poorly absorbed through the

intestinal tract. Inhalation or intravenous administration, however, causes a rapid rise in plasma THC concentrations followed by a

gradual decline according to a biphasic elimination pattern (17).

The physiological “high” corresponds to the time of maximum THC concentration in the blood.

Cannabinoids are readily metabolized in mammals; over 50 cannabinoid metabolites have been identified, with variations among species,

63
There is some evidence for differential metabolism between lung

and liver within the same species (3). Numerous biotransformation pathways for THC are known, including allylic and aliphatic

hydroxylations; oxidations to acids, aldehydes, and ketones; conjugations with glucuronic acid or fatty acids; epoxidation of the

double bond; and reduction of the terpene double bond. These biotransformations can occur singly to produce monosubstituted metabolites, or in combination to form diols, triols, hydroxy acids,

hydroxy ketones, or substituted hexahydrocannabinols (18). In man,

the conversion of THC to biologically inactive 11-nor-9-carboxy9

-THC (II) is a major metabolic pathway. The main initial biotransformation product of THC is its 11-hydroxy derivative (III),

which is equally and perhaps more active than THC, but like its

precursor it is short-lived in the body and rarely appears in urine.

Recent experiments with radiolabeled THC in rats have demonstrated

that III penetrates the blood-brain barrier more readily than does

THC, possibly because of its high affinity for plasma albumin rather

than the lipoprotein to which THC is bound (19).

Very little unchanged THC is found in urine. One study has reported

that 0.005 percent of the administered drug was detectable in the

urine of four human subjects up to 6 hours after a 30-mg dose.

Excretion of metabolites extended over a period of several days

(20) Although the metabolic breakdown of THC is very rapid, the

drug can be detected in plasma for at least 72 hr after a single

intravenous dose, possibly the consequence of a slowly releasing

deep compartment. THC is highly bound to lipoprotein, 97 percent

by one method of estimation, and has a reported pka’ of 10.6 (21).

An effective oral dose of THC in humans is about 50-200 μg/kg body

weight, and the effective dose when smoked is 25-50 μg/kg (22).

ANALYTICAL METHODOLOGY

Because there is great interest in the metabolism and pharmacology

of THC, and because of the forensic implications of its illicit

use, numerous methods have been developed for its detection and

quantitation; however, its low concentration in biological fluids

64
and the potential interference from endogenous lipids have created

difficult analytical problems. Fluorometric techniques, based on

gallium chelate formation or on cannabinoid dansylation, while

only qualitative, have been suitable for some applications such as

detection of THC in the saliva of marijuana smokers (23-27). A

quantitative fluorometric technique has been reported very recently,

in which a known amount of tritiated THC is added to plasma as an

internal standard prior to extraction with methyl acetate:petroleum

ether:ethanol (66: 33:1.5 v/v, efficiency 70 percent). THC and other

phenols are selectively removed from the organic extract by extraction with Claisen’s alkali reagent. The alkaline solution is then

acidified and reextracted with hexane. The THC is derivatized by

treatment with 14C-labeled dansyl chloride before purification by

thin-layer chromatography (TLC) on silanized silica gel plates

(cyclohexane:ethyl acetate, 95:5 v/v). The dansylated THC spot

is located with UV radiation and eluted into a scintillation vial

for counting of 3H and 14C. The detection level for this method

was found to be 2.5 ng/tube, implying a sensitivity of a few ng/ml

of plasma (28).

For periods up to 2 hours after smoking, the saliva of cannabis

users was reported to contain concentrations of THC sufficient for

detection by two-dimensional TLC with mass spectrometric confirmation. In a study which compared plasma and saliva concentrations of THC in the same subjects, the drug was detected in saliva

less than 30 min after smoking and continued to be detectable in

saliva even after it could no longer be detected in plasma (29).

A new derivatizing reagent (2-p-chlorosulfophenyl-3-phenylindone)

which is reported to be more selective than dansyl chloride has

been used in combination with TLC to detect THC and several of its

metabolites at serum concentrations as low as 0.2 ng/ml (30). However, the fluorescent THC derivative is unstable and must be measured within a few minutes after it is formed on the TLC plate.

Some of the first methods employed for THC quantitation in pharmacological and metabolic studies involved radiolabeled tracers

administered to volunteers. Even minute amounts of cannabinoid

products of the metabolism of radiolabeled THC were detectable by

scintillation counting following TLC. However, the administration

of radioactive substances to human subjects has potential health

hazards which limit the use of the technique (31).

Radioimmunoassays (RIA) have been developed for THC. They have

the advantages of comparative speed and convenience for clinical

uses. This is because they are suited to the simultaneous analysis of multiple samples, often without prior purification. Under

favorable conditions, sensitivities of about 2 ng/ml can be achieved.

However, because of the variables inherent in the preparation of

reagents by serological procedures, RIA lacks the precision of

other methods. In attempts to maximize specificity and affinity

of the antibodies, various derivatives of THC have been used as

haptens to be conjugated to carrier protein. These have included

the hemisuccinate ester (32), an azobenzoic acid derivative (33),

65
an N-oxysuccinimidoyl ester (34), a 10-iodo-9-uriedo linked THC

(35), and an 0-carboxymethyl derivative (36). Although the antibodies elicited by these conjugates are highly specific for the

3-ring cannabinoid nucleus, cross-reactions with other cannabinoids, both natural and metabolic, have not been eliminated. This

can be an advantage in the case of urine studies, where unmetabolized THC is virtually absent. The high degree of nonspecific

binding of THC to lipoprotein is a technical problem in RIA, but

0.1 percent Triton X-405 has been found to effectively solubilize

THC in the aqueous assay medium, and an ethanol extraction step

has been shown to permit > 92 percent recovery of THC from plasma

while removing protein material by precipitation (32). Other RIA’s

have been developed for the measurement of THC and the major metabolite II (Carboxy-THC) and used to determine the relative THC

and Carboxy-THC concentrations in urine and plasma of cannabis

smokers (37).

In order to improve the specificity of an RIA method, high performance liquid chromatography (HPLC) has been used to separate THC

from other cannabinoids that may be present in a plasma sample (38).

The HPLC-RIA method is reported to measure THC plasma concentrations as low as 0.1 ng/ml.

Another immunological technique, the EMIT assay (based on the inhibition of enzymes by antihapten antibodies when the enzymes are

covalently linked to the corresponding hapten) has been adapted

for THC and Carboxy-THC in urine, using relate dehydrogenase. The

hapten synthesized for use in the EMIT system elicited an antibody

that reacted strongly with Carboxy-THC, unlike antibodies previously produced with other THC haptens. The antisera were therefore suitable for general screening for THC and its metabolites

(39). The recommended cut-off for reporting detection of the cannabinoids is 20 ng/ml.

Another approach involves the use of gas chromatography (GC) to

separate THC from other components in plasma. The normal sensitivity of GC using flame ionization detectors (FID) is too low to be

of use for in vivo plasma analysis. However, electron capture detectors (ECD) the necessary sensitivity provided the THC is

converted to a suitable derivative. The pentafluoropropionate

derivative is satisfactory for this purpose; it allows detection

at concentrations as low as 2 ng/ml. However, the derivatizing

reactions are relatively unspecific; therefore biochemical interferences compromise selectivity. A tandem-column GC technique

can be used to achieve greater specificity; for example, THC was

detected in blood as the heptafluorobutyrate on a tandem column

GC system using hexahydrocannabinol as the internal standard. The

first column (5 percent SE-30) provided internal cleanup and the

second, a capillary column coated with OV-17, achieved the required resolution. The technique had the advantage of allowing

temperature programming in the first column which increased speed

and efficiency but did not disturb the base-line response of the

electron capture detector. The sensitivity reported for nonbiological samples was 1 pg injected on column (40).

66
In addition to improved detector response, derivatization of THC

can also improve chromatographic behavior by blocking the polar

phenolic group. THC derivatives which have been prepared for this

purpose include the trimethylsilyl, trifluoroacetyl, acetyl, chloroacetyl , and t-butyldimethylsilyl derivatives. The last named

has been shown to exhibit a high degree of stability to moisture

and bases (41).

Cannabinoids have also been converted to their diethyl phosphate

esters and analyzed by gas chromatography using a nitrogen-phosphorus detector (GC-NPD). The developers of this assay recommend

it for routine quantitative analysis of cannabinoids in all toxicological laboratories (42). The procedure utilizes the n-heptyl

homolog of 8-THC as the internal standard and is capable of detection limits of 0.5 ng/ml in blood and 10 ng/g in brain tissue.

For analysis of brain tissue a back extraction into methanolic

base serves to separate the cannabinoids from interfering endogenous lipids.

HPLC has been used for separation of THC from other cannabinoids

and lipids in biological fluids before analysis by GC-ECD. Reverse phase HPLC elution with 47 percent acetonitrile in water, as

well as normal phase HPLC with 25 percent chloroform in heptane,

was found to separate THC from its active 11-hydroxy metabolite

(III). The resolution of 8

- and 9

-THC was achieved with 5 percent tetrahydrofuran in hexane (43). The GC-ECD analysis of pentafluorobenzylated 9

-THC using HPLC purification had a sensitivity

of 1 ng/ml of plasma, while radiochemical analysis of the HPLC

separated fraction had a sensitivity of 0.2 ng/ml plasma. For

quantitation of THC in plasma in phannacokinetic studies, a known

amount of radiolabeled THC was added as an internal standard either

to the plasma or to the heptane extract. This permitted calculation

of extraction and/or HPLC collection efficiencies for the particular biological sample (44).

The technique of plasma chromatography has been applied to the detection of ultratrace amounts of THC in GC effluents. The positive mobility spectrum for THC shows a strong, single peak with a

Ko = 1.06. The peak is attributed to the protonated molecule ion

of THC (45).

GC/MS analysis using the technique of selected ion monitoring currently is the most reliable method for measuring the very low concentrations of THC in plasma after smoking. The first GC/MS method

for quantitation of THC in blood was reported in 1973 (46). The

method involved addition of a deuterated THC analog as an internal

standard and purification of the extract by liquid chromatography

on a Sephadex LH-20 column. The original procedure was later refined to the point that plasma concentrations as low as 0.3 ng/ml

could be measured. Sensitivity was found to be limited by a small

amount of undeuterated THC contaminating the internal standard, so

the best results have been achieved with a 2H7-analog (47).

67
Because of the very small amounts of THC and 11-hydroxy-THC which

appear in human plasma and urine, the major metabolite (CarboxyTHC) is a candidate for analytical studies as a method to identify

cannabis users. The concentration of Carboxy-THC in plasma exceeds

that of THC within 50 min after smoking, and then remains higher

for more than 24 hr. A modification of the above GC/MS assay for

THC has been applied to identification and estimation of the carboxy metabolite. Extraction from plasma or urine was with diethyl

ether (85 percent recovery from plasma, 90 percent from urine),

followed by a two-step derivatization which involves esterification of the carboxyl group with diazomethane and trimethylsilation of the phenolic hydroxyl group. The silylated methyl ester

of the 8-isomer of the Carboxy-THC was used as an external standard because both isomers have similar retention times but different ion intensities in their mass spectra. In this preliminary

study, a sensitivity in the range of a few ng/ml was indicated (48).

A GC/MS assay for THC in the blood of marijuana smokers has been

reported which is based on the chemistry of the phenolic group

common to all cannabinoids and their metabolites; the basis of the

purification technique is the selective extractability of lipidsoluble phenols from hexane by Claisen’s alkali. After the back

extraction into the aqueous caustic, trimethylanilinium hydroxide

was used for on-column methylation of the phenolic group. The

trideuteromethyl ether of THC was easily synthesized for use as

an internal standard, but the fact that it was, of necessity,

added only after the extraction constitutes a disadvantage of

the method as a quantitative technique (49).

The Claisen’s alkali extraction procedure was also used in an analysis of the active metabolite, 11-hydroxy-THC, in plasma (50). In

this case, tetrahexylammonium hydroxide and ethyl iodide were used

to alkylate the phenolic group of the metabolite. Subsequent trimethylsilylation of the aliphatic hydroxyl group provided a derivative with satisfactory chromatographic and mass spectral properties. When the corresponding pentadeuteroethyl ether was used as

the internal standard, a linear calibration graph was achieved over

the range 7 to 120 ng/ml of plasma. The procedure can be used

for simultaneous determination of both THC and the 11-hydroxy metabolite in plasma (51).

GC/MS was adapted to an automated system using a computer-managed

Olfax II spectrometer and a “probability based matching” algorithm,

to measure THC as its trimethylsilyl derivative in extracts of hydrolyzed human urine, and to identify and measure metabolites.

The procedure showed promise as a means of increasing the speed

and convenience of THC assays while reportedly retaining the specificity of selective ion monitoring (52).

In a pharmacokinetic study of THC and 11-hydroxy-THC in human

plasma, the results of GC/MS (both electron impact and chemical

ionization) were compared with those of GC-ECD and TLC-radiolabel

procedures on the same samples. In the CI procedure THC was derivatized with pentafluoropropionic anhydride, and the pentafluoro68
propionate ester of hexahydrocannabinol was used as the internal

standard. Endogenous plasma constituents did not interfere, so

preliminary chromatographic purification was not required. In

the EI procedure Sephadex LH-20 chromatography was used before

GC/MS, but THC was not derivatized. Pharmacokinetic data for human volunteer subjects were in reasonable agreement for the three

techniques (53).

Further refinements of the GC/EI-MS technique for THC have included

use of THC-2H3 as internal standard and extraction of THC from

plasma with pure hexane to avoid the need for preliminary chromatographic cleanup. The more polar cannabinoids remain with the

plasma; therefore hexane extraction would not be suitable when determination of metabolites is necessary. Both the CI and EI methods had detection limits of about 0.5 ng/ml in clinical samples,

although the EI mode (focusing exclusively on m/z 314 and 317 and

requiring no derivatization step) was reported to have a better

defined linearity on calibration curves than the methane CI method (54).

The combination of HPLC with MS has also been reported for analysis of THC in body fluids. A silica gel column was programmed for

gradient elution using heptane and methylene chloride. Fractions

of the eluent were collected and introduced by direct probe into

the mass spectrometer. THC- 2H3 was added to the body fluid and

served as an internal standard, a marker for effluent collection

from the HPLC (detected by an ultraviolet spectrophotometer) , and

as a carrier for the THC. The technique was used to assay THC

concentrations in human blood, breath, saliva, bile, and postmortem brain tissue. It was shown to be reproducible, accurate, and

linear in the range of 2.5-100 ng/ml of plasma. A positive, statistically significant concentration of THC was detected in the

saliva of marijuana smokers in the first hour after smoking (55).

A paper chromatographic system which employs methanol as the mobile

phase conveniently separates cannabinoids from the large amounts

of endogenous lipid material present in chloroform extracts of

blood samples. The region of the paper near the solvent front is

eluted with methylene chloride and the cannabinoids are identified

by GC/MS. If 15 to 25 ml of whole blood are available, concentrations of THC as low as 0.5 ng/ml can be detected. For quantitative analysis 2,3,4,6-tetrachlorophenyl hexyl ether is suggested

as an internal standard. Under the GC/MS conditions used (3 percent OV-1, EI) the tetrachloro ether has a retention time of 0.45

relative to THC and produces fragment ions at the same nominal

masses as several of THC’s prominent fragment ions. The same report points out the potential for pyrolytic conversion of THC to

cannabinol in GC and GC/MS analyses. Thermal decomposition of

THC is minimized by keeping all GC and MS temperatures to a minimum consistent with satisfactory chromatography (56).

Solvents used for extraction of THC from biological samples include hexane at neutral pH (52, 53); hexane + 5 percent isoamyl

alcohol (49); petroleum ether, sometimes containing 0.5 percent

69
isoamyl ether or 1.5 percent isopentanol (47,53,55); heptane containing 1.5 percent isoamyl alcohol (21,40,57); and petroleum ether:

Ethyl acetate (2:l) (29). Recoveries in excess of 90 percent from

plasma have been reported for heptane:isoamyl alcohol (40); chloroform extraction of whole blood gave a THC recovery of 86 percent

(56).

The proceedings of a symposium on the analysis of cannabinoids in

physiological fluids held in 1977 have been published and provide

a good review of recent research in this area (58). Also, the National Institute on Drug Abuse has sponsored two conferences on

the analysis of cannabinoids in physiological specimens. The proceedings of the first conference, held in 1976, were published as

a NIDA Research Monograph (59). The proceedings of the second conference, held in January, 1980, will be published later in the year

(60).

EXPERIMENTAL PROCEDURE

Two methods are described, one for analysis of THC in whole blood

or plasma and another for analysis of the metabolites, 11-hydroxyTHC and 9-carboxy-THC, in blood, plasma and urine. The first method involves extraction of the body fluid with hexane, a nonpolar,

relatively selective extraction solvent. The very lipophilic THC

is efficiently extracted by hexane, but the polar metabolites are

poorly extracted. The second method employs a more polar extraction solvent (hexane:ethyl acetate, 7:l v/v) which removes the metabolites from the body fluid, but unfortunately also extracts

endogenous components which interfere with measurement of low concentrations (< 10 ng/ml) of THC. A glass capillary GC column has

been used to resolve trimethylsilylated THC from endogenous components (61), but efforts to resolve the interferences using packed

GC columns have so far been unsuccessful. This is the reason why

two similar but separate procedures are used for quantitation of

THC and the two metabolites.

Standards and Reagents

All of the cannabinoid standards used in the development of the

method described here were obtained from the Research Triangle

Institute, Research Triangle, NC, 27709, through the National Institute on Drug Abuse. Purity and isotopic composition data are

given below:

9

-Tetrahydrocannabinol (THC)

Purity: 98.7 percent by GC

5’- 2 H 3-

9

-Tetrahydrocannabinol (THC-2H3)

Purity: > 95 percent by GC, 100 percent by TLC

Isotopic composition: 2H3, 96.6 percent; 2H2, 2.6 percent;

2H1

, 0.77 percent; 2H0, 0 percent.

11-Hydroxy- 9-tetrahydrocannabinol (11-OH-THC)

Purity: >98 percent by GC as the bis-(trimethylsilyl) ether,

100 percent by TLC

70
5'-2H3

-11-Hydroxy- 9-tetrahydrocannabinol (11-OH-THC-2H3)

Purity: >95 percent by GC as the bis-(trimethylsilyl) ether,

100 percent by TLC

Isotopic composition: 2H3 95.8 percent; 2H2, 2.4 percent;

2H1

 1.1 percent; 2H0, 0 percent

11-Nor-9-car-boxy- 9-tetrahydrocannabinol (9-Carboxy-THC)

Purity: 97 percent by GC as the bis-(trimethylsilyl) ether,

100 percent by TLC

5'-2H3-11-Nor-9-carboxy- 9-tetrahydrocannabinol (9-Carboxy-THC-2H3)

Purity: 100 percent by GC as the bis-(trimethylsilyl) derivative, 100 percent by TLC

Isotopic composition: 2H3, 95.8 percent;

2H1

, 1.1 percent; 2H0, 0 percent.

2H2

, 2.4 percent;

The pH 2.0 and pH 7.0 buffer solutions were purchased from Fischer

Scientific Company.

N,O-Bis-(trimethylsilyl)-trifluoroacetamide (BSTFA) containing 1

percent trimethylsilyl chloride was purchased from the Pierce Chemical Company, Rockford, IL, 61105. The reagent was stored at -10°C,

but allowed to warm to room temperature before aliquots were removed for derivatization.

Pure 9-THC is a colorless, viscous syrup. It is therefore difficult to transfer and weigh small quantities accurately. The

9

-THC standards described above were obtained as ethanolic solutions of predetermined concentrations, and were subsequently diluted with methanol to give stock solutions with THC concentrations of 0.10 mg/ml. Of course, the concentrations do not have

to be precisely 0.10 mg/ml, but they must be accurately known.

A THC stock solution concentration of 0.10 mg/ml is used in this

experimental description merely as an example.

The 11-OH-THC and 9-carboxy-THC are solids at room temperature.

Because these two metabolites are measured at the same time in

biological specimens, a single stock solution containing 0.10 mg/ml

of each of the metabolites in methanol can be prepared, as well as

a single stock solution containing 0.10 mg/ml of each of the deuterated analogs (11-OH-THC-2H3

 and 9-car-boxy-THC-2H3) in methanol.

These can be prepared by weighing into a 100-ml volumetric flask

10 mg of 11-OH-THC and 10 mg of 9-carboxy-THC, and weighing into

a second 100-ml volumetric flask 10 mg of each of the deuterated

analogs. Add about 75 ml of methanol to each flask. After the

solids have dissolved add additional methanol to the flasks to

bring each solution to exactly 100 ml. A series of working standard solutions of the undeuterated cannabinoids can be prepared

by appropriate dilution of the stock solutions as described in

Chapter 2.

For the measurement of THC or 11-OH-THC and 9-carboxy-THC in body

fluids within the concentration range 1 to 500 ng/ml, addition of

approximately 40 ng/ml of each of the appropriate deuterated in71

328-011 0 - 80 - 6
ternal standards to each ml of specimen is satisfactory. This can

be conveniently done by adding 100 μ1 of a 400-ng/ml methanolic

solution of THC- 2H3 or 11-CM-THC-2H3 and 9-carboxy-THC-2H3 to each

ml of body fluid. Prepare the 400-ng/ml internal standard solutions by diluting 1 ml of the 0.10-mg/ml stock solutions to 250

ml with methanol.

Cannabinoids, particularly THC, are unstable when exposed to light

(4,62), oxygen (4), acid (21), and/or elevated temperatures (62).

Consequently, store stock and standard solutions in full, stoppered

or capped glass vessels in the dark at below 0°C. The concentrations of the stock solutions should be checked at least every three

months. This can be done by addition of a known quantity of an

internal standard such as 4-androsten-3,17-dione to an aliquot

of the stock solution, and then analyzing the solution by GC with

a flame ionization detector. If the concentration of the cannabinoid solution has changed by a detectable amount, a new calibration

graph should be established for that stock solution.

THC has a strong tendency to adhere to glass surfaces (21). This

adsorption can cause poor sensitivity and precision, but can be

minimized by using clean, silanized glassware for the sample extraction (63).

Extraction of THC from Plasma or Whole Blood

Transfer a measured quantity (normally 1 ml) of plasma or whole

blood to a 20-ml culture tube equipped with a Teflon-lined screw

cap. Add 100 μl of a 400-ng/ml methanol solution of the THC-2H3

internal standard and vortex for 10 sec. Allow the sample to equilibrate for 15 min. Add 1 ml of pH 7.0 buffer solution and 6 ml of

hexane. Cap the tube tightly and gently mix the contents for 30

min using an automated tube rocker or rotator. Centrifuge for 5

min and then carefully transfer just the hexane (top) layer to a

second culture tube using a disposable Pasteur pipette. The remaining plasma residue can be discarded. The hexane solution is

now washed successively with dilute alkali and dilute acid to remove strongly acidic and basic compounds. Add 2.5 ml of 0.1 N

NaOH solution to the hexane extract. After capping the tube, rotate or rock for 30 min and then centrifuge. Using another disposable pipette, remove the aqueous (bottom) layer and discard.

Add 2.5 ml of 0.1 N HCl solution to the hexane extract. Tap the

tube gently to allow gas bubbles formed by the acid-base reaction

to escape. Again, cap the tube tightly, rock or rotate it for 30

min, and centrifuge. Carefully transfer just the hexane (top)

layer to a 15-ml glass tube with a tapered or cone-shaped bottom.

Concentrate the extract to near dryness at a temperature of 35-

40°C under a gentle stream of nitrogen. The extract residue is

now transferred to a conical 1-ml glass vial equipped with a Teflon-lined screw cap in the following manner. Add 0.5 ml of hexane

to the tube containing the residue and vortex the tube for 10 sec.

Using a disposable pipette, transfer the hexane solution to the

1-ml glass vial. Repeat the last step using an additional 0.5

ml of hexane to transfer any remaining extract residue to the 1-ml

72
vial. Add 10 μ1 of dimethylformamide to the hexane solution to

act as a “keeper” to minimize evaporative loss of the THC during

the final evaporation and to serve as a solvent for the trimethylsilylation. Slowly concentrate the hexane solution by heating the

glass vial in a heating block maintained at 40°C. When all of the

hexane has evaporated (approximately 20 min), add 15 μl of bis-

(trimethylsilyl) trifluoroacetamide containing 1 percent trimethylsilyl chloride to the vial and cap tightly. Vortex the vial and

then heat in an oven at 75°C for 1 hr.

for GC/MS analysis, but it can be

The sample. is now ready

stored at -10°C indefinitely.

GC/MS Analysis of THC-TMS

The experimental conditions for the GC/MS analysis of THC are as

follows:

GC column:

Carrier gas:

Reagent gas:

Temperatures:

1.8 m x 2 mm I.D. glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Methane, 15-20 ml/min

Ammonia introduced into the ion chamber as

described in Chapter 2

Injector, 280°C

Column, 190 to 310°C at 20 °/min

GC/MS transfer line, 280°C

Ion source, 160°C

Under these conditions the THC-TMS should elute from the gas chromatograph at between 2 and 5 min.

Prior to beginning the GC/MS analysis of samples, the performance

of the total GC/MS system should be evaluated and optimized as

described in Chapter 2. The ions to be monitored are the protonated molecule ions for THC-TMS (m/z 387) and THC-2H3

-TMS (m/z

390). With the divert valve in the divert position, inject the

organic extract into the gas chromatograph (2 to 8 μ1). After

approximately 1 min, switch the divert valve so that the entire

carrier gas flow enters the ion source of the mass spectrometer,

and begin data acquisition. When the THC-TMS peaks have eluted,

cease data acquisition and return the divert valve to the divert

position.

The quantity of THC in the biological specimen sample is determined by measuring the heights (or areas) of the THC-TMS and THC2H3

-TMS peaks in the selected ion current profiles (“ion chromatograms”) and relating the ratio of peak heights to a calibration

graph. Calculate the body fluid concentration of THC by dividing

the measured quantity of THC by the exact volume of specimen analyzed.

73
Extraction of Metabolites from Blood, Plasma, or Urine

To 1 ml of specimen (blood, plasma, or urine) in a 20-ml culture

tube equipped with a Teflon-lined screw cap, add 100 μ1 of a methanol solution containing 40 ng each of the internal standards,

11-OH-THC-2H3 and 9-carboxy-THC-2H3 (concentration, 400 ng/ml).

Vortex the tube for 10 sec and allow the sample to equilibrate

for 15 min. Add 1 ml of pH 2 buffer and 4 ml of a hexane:ethyl

acetate solvent (7:1 v/v). Screw the cap on the tube tightly and

check for leaks. Slowly rotate or rock the tube for 15 min. Loosen

the cap and then centrifuge. Using a disposable Pasteur pipette,

transfer the organic layer (top) to a second culture tube and discard the aqueous plasma layer. Add 2 ml of 0.2 N H2SO4

 to the organic extract. Screw on the cap, tighten, and again check for

leaks. Repeat the rocking (15 min) and centrifugation (5 min).

Transfer the organic layer (top) to a centrifuge tube, at least

10 ml in volume. Be careful not to transfer any of the aqueous

(lower) layer to the centrifuge tube. Discard the aqueous layer.

Concentrate the extract to near dryness by heating to 35-40°C under

a gentle stream of nitrogen or filtered air. Wash down the sides

of the tube with 0.5 ml of methylene chloride. Vortex the tube

and transfer the extract to a 1-ml glass vial which has a coneshaped bottom and a Teflon-lined screw cap. Again wash down the

sides of the centrifuge tube with another 0.5 ml of methylene

chloride and transfer the solvent wash to the same 1-ml glass vial.

Add 10 μl of dimethylformamide to the hexane solution to act as a

“keeper” to minimize evaporative loss of the metabolites during

the final evaporation and to serve as a solvent for the trimethylsilylation. Evaporate the combined solvents by heating to 30-40°C

under a gentle stream of nitrogen or filtered air until all of the

methylene chloride is gone (about 20 min). Add 50 μ1 of bis-(trimethylsilyl) trifluoroacetamide containing 1 percent trimethylsilyl

chloride to the vial; cap tightly and heat at 90°C for 1 hr. The

sample is now ready for GC/MS analysis. If necessary it can be

stored in a freezer until it is to be analyzed; however, be sure

to allow the vial to warm to room temperature before removing the

cap to withdraw material for injection into the GC/MS. The trimethylsilylated 9-carboxy-THC is particularly vulnerable to hydrolysis.

Consequently, exposure of the derivatized extract to air should

be kept to a minimum, and prolonged storage should be avoided.

GC/MS Analysis of the 11-OH-THC-TMS and 9-Carboxy-THC-TMS

The experimental conditions for GC/MS analysis of the derivatized

THC metabolites are as follows:

GC column: 1.8 m x 2 mm I.D. glass column packed with

3 percent OV-17 or 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier gas: Methane, 15-20 ml/min

Reagent gas: Ammonia, introduced into the ion chamber as

described in Chapter 2

7 4
Temperatures: Injector, 280°C

Column, 190 to 310°C at 20°/min

GC/MS transfer line, 280°C

Ion source, 160°C

Under these conditions the 11-OH-THC-TMS should elute from the GC

at about 3 min and the 9-carboxy-THC-TMS at about 4.5 min.

The ions to be monitored are the protonated molecule ions for 11-

OH-THC-TMS (m/z 475), 11-OH-THC-2H3-TMS (m/z 478), 9-carboxy-THCTMS (m/z 489), and 9-carboxy-THC-2H3

-TMS (m/z 491). With the divert valve in the divert position inject 2 to 8 μl of the trimethylsilylated extract into the GC/MS. After approximately 1 min switch

the divert valve so that the entire carrier gas flow enters the

ion source. The best sensitivity is achieved by first monitoring

just the ion currents at m/z 475 and 478 until the 11-OH-THC-TMS

has eluted from the GC; then monitor the ion currents at m/z 489

and 491 until the 9-carboxy-THC-TMS has eluted. If the GC/MS data

system does not permit rapid changes of the monitored ion masses,

all four ion masses must be monitored for the entire time required

for elution of both the metabolites. When the 9-carboxy metabolite

has eluted, cease data acquisition and return the divert valve to

the divert position.

The quantities of 11-hydroxy and 9-carboxy metabolites in the biological sample are determined by measuring the heights (or areas)

of the derivatized metabolites and their internal standards in the

selected ion current profiles, and relating the ratios of peak

heights to calibration graphs (Chapter 2). Calculate the concentrations of the 11-hydroxy and 9-carboxy metabolites in the specimen

by dividing the measured quantities of these compounds by the exact

volume of specimen extracted.

Discussion of the Experimental Procedures

The method described above for quantitation of THC in blood and

plasma was developed and refined over a period of four years. During this time over 2500 plasma samples, submitted by various cannabinoid researchers, were analyzed. One out of every six analytical samples corresponded to a control plasma to which a known

amount of THC was added. The measured concentrations of THC in

the spiked plasma samples are summarized in Table 1.

75
TABLE 1. ACCURACY DATA FOR MEASUREMENT OF THC

ADDED TO PLASMA.

Prepared THC Mean Measured

Concentration Concentration ± S.D.

(ng/ml) (ng/ml)

0.92 0.88 ± 0.39

1.84 1.73 ± 0.51

Number of

Samples

Analyzed

15

18

4.60 4.75 ± 0.89 24

9.20 9.32 ± 0.72 22

18.4 18.85 ± 0.45 22

46.0 47.21 ± 1.83 2 1

92.0 91.97 ± 1.92 23

184.0 176.71 ± 1.74 13

Figure 1 shows a representative calibration graph covering the concentration range 0.5 to 184 ng/ml. The correlation coefficient

for the graph is 0.9999.

Most of the THC analyses done at Battelle between September 1977

and September 1979 were performed on a Finnigan 4000 GC/MS coupled

to an INCOS data system. With the system operating normally, as

little as 20 pg of derivatized THC injected into the GC/MS gave

a response sufficient to permit measurement with good precision.

Usually, only 10 to 20 percent of a derivatized extract was injected at one time. Consequently, for a 1-ml plasma sample, concentrations of THC as low as 0.2 ng/ml could be measured. One

technician was able to extract and prepare a set of 36 plasma samples for analysis in a period of 8 hr. An additional 8 hr was required to perform the GC/MS analyses.

The advantages of using CI mass spectrometry with methane and ammonia as reagent gases are evident in Figure 2 and Table 2. Figure 2 compares the EI mass spectrum of the trimethylsilyl derivative of THC with the corresponding CI mass spectra using either

methane or the methane-ammonia combination as reagent gas. In the

methane-ammonia CI mass spectrum almost all of the THC ion current

is concentrated at a single ion mass, corresponding to the protonated molecule ion (m/z 387). The sensitivities achieved by selected ion monitoring analysis of specific quantities of THC are

compared in the fourth column in Table 2. Use of methane and am76
FIGURE 1. CALIBRATION GRAPH FOR ANALYSIS OF THC IN

PLASMA

77
FIGURE 2. MASS SPECTRA OF TRIMETHYLSILYLATED THC

78
monia gives the largest signal per unit weight of THC. Furthermore, the virtual absence of fragment ions under methane-ammonia

chemical ionization means that there is less chance of interference by ions from other components of the extract.

TABLE 2. RELATIVE RESPONSES FOR MAJOR IONS IN THE EI

AND CI MASS SPECTRA OF THE TRIMETHYLSILYL

ETHER OF THC

Ionization

Method

EI

Percent Relative Response

m/z of Samples Per Unit Weight

Monitored Ion Current of THC

386 (M+) 7 50

CI (CH4) 387 (MH+

) 24 20

CI (CH4 + NH3) 387 (MH+

) 67 100

The efficient protonation of THC-TMS under methane-ammonia chemical ionization conditions was unexpected, since the THC molecule

does not contain a basic nitrogen atom. However, apparently in

the gas phase THC and its TMS derivative have high proton affinities. The same is true of the TMS derivatives of the two metabolites, as indicated by the abundant protonated molecule ions

observed in their methane-ammonia CI mass spectra (Figure 3).

The efficiency of ionization and the selectivity of methane-ammonia chemical ionization are major reasons why plasma concentrations of THC as low as 0.5 ng/ml can be measured without employing, a more extensive extraction and sample purification procedure.

However, it is necessary to heat the GC column to temperatures

well above the elution temperature of THC-TMS in order to clear

the column of endogenous compounds, such as cholesterol. If this

is not done, interfering peaks will appear in the ion current profiles generated by subsequent injections. Maintenance of the column at 280-300°C overnight with normal carrier gas flow will also

help to retain good column performance.

If the procedures outlined above are followed, a GC column should

give satisfactory performance for at least several hundred analyses. However, the performance of the column should be evaluated

each day before the analysis of specimens is begun. The resolution of the GC column can be evaluated by injecting a solution

which contains an equal mixture of 8

-THC-TMS and 9

-THC-TMS (~50

μg/ml in hexane). Under the stated GC conditions a good quality

1.8-m x 2-mm packed OV-17 column should give a chromatogram showing at least a detectable valley between the peaks corresponding

to the two cannabinoid derivatives. The sensitivity of the GC/MS

can be evaluated by injecting 1 μl of a standard solution contain79
FIGURE 3. METHANE-AMMONIA CI MASS SPECTRA OF TRIMETHYLSILYL DERIVATIVES OF THC (TOP), 11-

OH-THC (MIDDLE), AND 9-CARBOXY-THC (BOTTOM)

80
ing 1 ng of THC-TMS. The peak in the m/z 387 ion current profile

which corresponds to the THC-TMS should be symmetrical and have a

signal-to-noise ratio of greater than 50:1.

The procedure described above for analysis of the THC metabolites

has not been used as extensively as the THC method, and therefore

it is not as thoroughly validated. A similar procedure, in which

the 9-carboxy-THC was first converted to its methyl ester by treatment with ethereal diazomethane and then trimethylsilylated, was

employed at Battelle for the analysis of approximately 300 plasma

samples. Figure 4 shows representative calibration graphs acquired

at the time these assays were conducted. Table 3 lists the assay

precision for the analysis of plasma samples to which a known concentration of each of the metabolites was added.

TABLE 3. PRECISION DATA FOR MEASUREMENT OF

THC METABOLITES ADDED TO PLASMA

Metabolite

Concentration

(ng/ml)

1

11-OH-THC

RSD

(%)

7

9-Carboxy-THC

RSD

(%)

31

5 4 7

10 2 2

50 1 3

100 3 2

It has been shown recently that up to 80 percent of the 9-carboxyTHC in urine from marijuana smokers is present in the form of a

glucuronide conjugate (64). The glucuronic acid residue appears

to be attached to the metabolite by an ester linkage with the 9-

carboxyl group of the metabolite. The conjugate is not cleaved

by conventional treatment with ß-glucuronidase and sulphatase, but

can be cleaved by alkaline hydrolysis (65). The recommended conditions for hydrolysis are to add the biological specimen to an

equal volume of 1 M sodium hydroxide in methanol and heat on a

steam bath (100°C) for 12 minutes. Acidify the mixture to pH 2

by addition of 1 ml of 1 N sulfuric acid and 1 ml of pH 2 buffer.

The total 9-carboxy-THC concentration can then be determined by

extraction with hexane:ethyl acetate, derivatization, and GC/MS

analysis as previously described.

The analysis of THC in decomposed blood or tissues presents special problems. The main difficulty is that the efficiency of extraction of THC from decomposed blood with hexane is poor and erratic. Even deuterated THC added to decomposed blood as an inter81
FIGURE 4. CALIBRATION GRAPH FOR ANALYSIS OF 11-OH-THC

AND 9-CARBOXY-THC AS THEIR TRIMETHYLSILYL

DERIVATIVES

82
nal standard is difficult to extract efficiently with hexane. Apparently the THC binds to biochemical components in the decomposed

blood in a manner that inhibits its removal by conventional extraction procedures. A procedure was recently reported for the

quantitative determination of THC in cadaver blood (66). The method involves extraction of 3 ml of blood with a total of 50 ml

of acetone, evaporation of the acetone, extraction of the residue

with petroleum ether, and then chromatography on Sephadex LH-20

followed by GC/MS analysis. A simpler procedure was developed

at Battelle for extraction of THC from hemolyzed and decomposed

blood (67); it gave acceptable quantitative results for concentrations of THC down to 1 ng/ml for most batches of hemolyzed

blood. However, occasionally large variations in extraction efficiency did occur, indicating that the method needs further refinement. The procedure consists of addition of the internal standard to 1 ml of hemolyzed blood followed by 1 ml of 3 N H2SO4

 and

1 ml of acetonitrile. The mixture is sonicated and the resulting

protein precipitate removed after centrifugation. The supernatant

is then neutralized with 3 N NaOH, and 1 ml of pH 7.0 buffer is

added. The aqueous solution is extracted twice with 6 ml of hexane. The remaining steps, which include successive washes of the

hexane extract with dilute alkali and dilute acid followed by concentration and derivatization, are identical to the procedure for

extraction of THC from normal plasma.

83
PHYSICAL, CHEMICAL AND SPECTROMETEIC DATA

FOR 9-TETRAHYDROCANNABINOL

Nomenclature

Chemical Name: 6aR-trans-6a,7,8,10a-Tetrahydro-6,6,9-trimethyl3-pentyl-6H-dibenzo[b,d]pyran-1-ol

Empirical Formula: C21H30O2

Chemical Registry Number: 1972-08-3

Physical Constants

Appearance: Colorless viscous syrup

Boiling Point: 200°C at 0.02 mm

Specific Rotation: = -150.5° (c = 0.53 in CHCl3)

Solubility: Very soluble in hexane, chloroform and ethanol. Solubility in water, 3 mg/liter

pKa: 10.6

UV Absorption: max (ethanol) = 283, 276 nm (log E 3.21, 3.20)

PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR

11-HYDROXY- 9-TETRAHYDROCANNABINOL

Nomenclature

Chemical Name: 6aR-trans-6a,7,8,10a-Tetrahydro-1-hydroxy-6,6-

dimethyl-3-pentyl-6H-dibenzo[b,d]pyran-9-methanol

Empirical Formula: C21H30O3

Chemical Registry Number: 36557-05-8

Physical Constants

Appearance: White, amorphous solid

Melting Point: 139-140°C

Solubility: Soluble in ethanol, acetone. Sparingly soluble in

benzene, chloroform. Nearly insoluble in water,

petroleum ether

UV Absorption: max (ethanol) = 276, 282 nm

84
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR

11-NOR-9-CARBOXY- 9

-TETRAHYDROCANNABINOL

Nomenclature

Chemical Name: 6aR-trans-6a,7,8,10a-Tetrahydro-1-hydroxy-6,6-

dimethyl-3-pentyl-6H-dibenzo[b,d]pyran-9-carboxylic acid

Empirical Formula: C21H28O4

Chemical Registry Number: 56354-06-4

Physical Constants

Appearance: White, crystalline solid

Melting Point: 210-213°C

Solubility: Soluble in ethanol, acetone. Sparingly soluble in

chloroform, ethyl ether. Nearly insoluble in water,

benzene, petroleum ether

REFERENCES

1.

2.

3.

4.

5.

6.

7.

8.

9.

S. Cohen in R.C. Petersen, ed., Marihuana Research Findings:

1976. National Institute on Drug Abuse Research Monograph 14,

DHEW Pub. (ADM) 77-501. Supt. of Docs., U.S. Govt. Print.

Off., Washington, D.C., ch. 9 (July 1977).

A. Sperling. J. Chromatogr. Sci., 10, 268 (1972).

R. Karler in R.C. Petersen, ed., Marihuana Research Findings:

1976, ch. 2. See reference 1.

J.W. Fairbairn, J.A. Liebmann, and M.G. Rowan. J. Pharm.

Pharmacol., 28, 1 (1976).

R. Mechoulam and Y. Gaoni. J. Am. Chem. Soc., 87, 3273 (1965).

A.J. Singer, ed. “Marihuana: Chemistry, Pharmacology, and

Patterns of Social Use”, Ann. N.Y. Acad. Sci., 191, 1-269

(1971).

R. Mechoulam, ed. Marijuana, Academic Press, New York and

London (1973).

W.D.M. Paton and J. Crown. Cannabis and its Derivatives,

Oxford Univ. Press, London (1972).

R.E. Willette, ed., Drugs and Driving. National Institute on

Drug Abuse Research Monograph 11, DHEW Pub. (AIM) 77-432.

Supt. of Docs., U.S. Govt. Print. Off., Washington, D.C.

(March 1977).

85
10. G. Nahas. U.N. Bull. Narcotics, 29, 13 (1977).

11. M.L. Lee, M. Novotny, and K.D. Bartle. Anal. Chem., 48,

405 (1976).

12. R. Karler in R.C. Petersen, ed., Marihuana Research Findings:

1976, ch. 3. See reference 1.

13. E.G. Leighty, A.F. Fentiman, Jr., and R.L. Foltz. Res. Commun.

Chem. Pathol. Pharmacol., 14, 13 (1976).

14. R. Mechoulam, W.K. McCallum, and S. Burstein. Chem. Rev., 76,

75 (1976).

15. G.G. Nahas, W.D.M. Paton, and J.E. Idanpaan-Heikkila, eds.,

Marihuana: Chemistry, Biochemistry, and Cellular Effects.

Springer-Verlag, New York (1976).

16. M.C. Braude and S. Szara, eds., Pharmacology of Marihuana,

Vol. 1 2. Raven Press, New York (1976).

17. M.E. Wall. Ann. N.Y. Acad. Sci., 191, 23 (1971).

18. D.J. Harvey, B.R. Martin, and W.D.M. Paton. J. Pharm. Pharmacol., 29, 495 (1977).

19. J. Schou, L.D. Prockop, G. Dahlstrom, and C. Rohde. Acta

Pharmacol. et Toxicol., 41, 33 (1977).

20. S.L. Kanter, L.E. Hollister, F. Moore, and D.E. Green. Clin.

Chem., 20, 860 (1974).

21. E.R. Garrett and C.A. Hunt. J. Pharm. Sci., 63, 1056 (1974).

22. R. Mechoulam. Science, 168, 1159 (1970).

23. R. Boudon. Euro. J. Toxicol., 9, 11 (1976).

24. M. Frei-Hausler and R.W. Frei. J. Chromatogr., 79, 209 (1973).

25. M. Frei-Hausler and R.W. Frei. J. Chromatogr., 84, 214 (1973).

26. A.P. Melikian and I.S. Forrest. J. Pharm. Sci., 62, 1025 (1973).

27. H. Bowd, D.H. Swann, B. Towell, and J.H. Turnbull, in J.B. Birks,

ed., Excited States Biol. Mol. Proc. Int. Conf. 1974. Wiley,

Chichester, England, p. 128 (1976).

28. J.M. Schermann, H. Hoellinger, N.-H.-Nam, R. Bourdon, and E.

Fournier. Clin. Chem. Acta, 79, 401 (1977).

29. W.W. Just, N. Filipovic, and G. Werner. J. Chromatogr., 96,

189 (1974).

86
30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

J.A. Vinson, D.D. Patel, and A.H. Patel. Anal. Chem., 49,

163 (1977).

L. Lemberger, in Garattini F. Hawking, A. Goldin, and I.J.

Kopin, eds., Advances in Pharmacology and Chemotherapy. Academic Press, New York and London, 10, p. 221 (1972).

J.D. Teale, E.J. Forman, L.J. King, E.M. Piall, and V. Marks.

J. Pharm. Pharmacol., 27, 465 (1975).

J.D. Grant, S.J. Gross, P. Lomax, and R. Wong. Nature New

Biol., 236, 216 (1972).

C.E. Cook, M.L. Hawes, E.W. Amerson, C.G. Pitt, and D. Williams

in R.E. Willette, ed., Cannabinoid Assays in Humans. National

Institute on Drug Abuse Research Monograph 7, DHEW Pub. (ADM)

76-339. Supt. of Docs., U.S. Govt. Print. Off., Washington,

D.C., p. 15 (May 1976).

P.T. Tsui, K.A. Kelly, M.M. Ponpipom, M. Strahilevitz, and

A.H. Sehon. Can. J. Biochem., 52, 252 (1974).

A.R. Chase, P.R. Kelley, A. Taunton-Rigby, R.T. Jones, and

T. Harwood in R.E. Willette, ed., Cannabinoid Assays in Humans, p. 1. See reference 34.

S.J. Gross and J.R. Soares in R.E. Willette, ed., Cannabinoid

Assays in Humans, p. 10. See reference 34.

P.L. Williams, A.C. Moffat, and L.J. King. J. Chromatogr.,

155, 273 (1978).

G.L. Rowley, T.H. Armstrong, C.P. Crowl, W.M. Eimstad, W.M. Hu,

J.K. Kam, R. Rogers, R.C. Ronald, K.E. Rubenstein, B.G. Sheldon,

and E.F. Ullman in R.E. Willette, ed., Cannabinoid Assays in

Humans, p. 28. See reference 34.

D.C. Fenimore, R.R. Freeman, and P.R. Loy. Anal. Chem., 45,

2331 (1973).

E.E. Knaus, R.T. Coutts, and C.W. Kazakoff. J. Chromatogr.

Sci., 14, 525 (1976).

N.K. McCallum and E.R. Cairns. J. Anal. Toxicol., 2, 89 (1978).

E.R. Garrett and C.A. Hunt in R.E. Willette, ed., Cannabinoid

Assays in Humans, p. 28. See reference 34.

E.R. Garrett and C.A. Hunt. J. Pharm. Sci., 66, 20 (1977).

F.W. Karasek, D.E. Karasek, and S.H. Kim. J. Chromatogr.,

105, 345 (1975).

87
46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

S. Agurell, B. Gustafsson, B. Holmstedt, K. Leander, J.-E.

Lindgren, I. Nilsson, F. Sandberg, and M. Asberg. J. Pharm.

Pharmacol., 25, 554 (1973).

A. Ohlsson, J.-E. Lindgren, K. Leander, and S. Agurell in R.

E. Willette, ed., Cannabinoid Assays in Humans, p. 48. See

reference 34.

M. Nordqvist, J.-E. Lindgren, and S. Agurell in R.E. Willette,

ed., Cannabinoid Assays in Humans, p. 64. See reference 34.

J.M. Rosenfeld, B. Bowins, J. Roberts, J. Perkins, and A.S.

Macpherson. Anal. Chem., 46, 2232 (1974).

J.M. Rosenfeld and V.Y. Taguchi. Anal. Chem., 48, 726 (1976).

J.M. Rosenfeld. Anal. Letters, 10, 917 (1977).

D.E. Green in R.E. Willette, ed., Cannabinoid Assays in Humans,

p. 70. See reference 34.

M.E. Wall, T.M. Harvey, J.T. Bursey, D.R. Brine, and D. Rosenthal in R.E. Willette, ed., Cannabinoid Assays in Humans, p.

107. See reference 34.

D. Rosenthal, T.M. Harvey, J.T. Bursey, D.R. Brine, and M.E.

Wall. Biomed. Mass Spectr., 5, 312 (1978).

J.L. Valentine, P.J. Bryant, P.L. Gutshall, O.H.M. Gan, E.D.

Thompson, and H.C. Hiu in R.E. Willette, ed., Cannabinoid

Assays in Humans, p. 96. See reference 34. Also, J. Pharm.

Sci., 66, 1263 (1977).

J.N. Pirl, V.M. Papa, and J.J. Spikes. J. Anal. Toxicol., 3,

129 (1979).

L. Lemberger, S.D. Silberstein, J. Axelrod, and I.J. Kopin.

Science, 170, 1320 (1970).

J.A. Vinson, ed., Cannabinoid Analysis in Physiological Fluids,

ACS Symposium Series 98. American Chemical Society, Washington,

D.C. (1979).

R.E. Willette, ed., Cannabinoid Assays in Humans. National

Institute on Drug Abuse Research Monograph 7, DHEW Pub. (ADM)

76-339. Supt. of Docs., U.S. Govt. Print. Off., Washington,

D.C. (May 1976).

R.L. Hawks, ed., The Analysis of Cannabinoids in Biological

Fluids. National Institute on Drug Abuse Resarch Monograph

Series. In preparation.

R.L. Foltz and B.J. Hidy in R.L. Hawks, ed., The Analysis of

Cannabinoids in Biological Fluids. See reference 60.

88
62. C.M. Bonuccelli. J. Pharm. Sci., 68, 262 (1979).

63. D.C. Fenimore, C.M. Davis, J.H. Whitford, and C.A. Harrington.

Anal. Chem., 48, 2289 (1976).

64. A.C. Moffat in R.L. Hawks, ed., The Analysis of Cannabinoids

in Biological Fluids. See reference 60.

65. P.L. Williams, A.C. Moffat, and L.J. King. J. Chromatogr.,

186, 595 (1979).

66. D. Rosenthal and D. Brine. J. Forensic Sci., 24, 282 (1979).

67. B.J. Hidy and R.L. Foltz. Final Report on Quantification of

Cannabinoids in Body Fluids. National Institute on Drug Abuse,

Contract No. HSM-42-72-183, January 1980.

89
Chapter 7

Cocaine and Its Major Metabolite,

Benzoylecgonine

Cocaine (3-benzoyloxy-8-methyl-8-azabicyclo[3.2.1]octane-2-carboxylic

acid methyl ester, I) is an alkaloid obtained from the leaves of

Erythroxylon coca or by synthesis from ecgonine. Although cocaine

hydrochloride has limited medical use as a topical anesthetic, it

is widely used as a stimulant by drug abusers. The major geographical sources of illicit cocaine are Peru, Bolivia, and Ecuador. The

plant from which it comes is largely restricted to the Andes mountain regions of these countries. In these areas the plant leaves

have been chewed by the native population for many centuries for

religious purposes and as an adjunct to work, much as coffee is

consumed as a stimulant in our culture (1) . However, when the psychoactive ingredients of the leaf are isolated and injected or inhaled intranasally in concentrated form, the pharmacological effects

can be quite different from those which occur in the traditional

pattern of cocaine use among Andean Indians. The leaves contain only

about 1 percent cocaine, and the oral route appears to delay absorption and decrease toxicity.

Cocaine was first isolated from coca leaves in 1860, and its therapeutic possibilities were quickly recognized by some physicians,

especially in Europe. By 1890 it was incorporated into a variety

of patent nostrums and “tonics” such as Coca Cola, although since

1910 the latter has not contained any cocaine (2). The dangerous

qualities of cocaine soon became apparent, and following its inclusion in a U.S. narcotics law in 1914, cocaine went “underground,”

to be used primarily in the Bohemian subculture of artists and musicians. It reemerged about 1970 as the “status” drug among middleclass drug abusers in this country, who commonly refer to it as

“coke” or “snow” (3). The reason for its popularity is the nature

of the stimulation it produces; i.e., feelings of heightened sexuality, psychic energy, and self-confidence which are of short duration (a few minutes) but very intense.

90
Cocaine is one of the most powerful natural central nervous system stimulants known (4). It blocks reuptake of norepinephrine

into nerve endings, acting first on the brain cortex to increase

motor activity. Overdose causes anxiety, depression, and confusion, and can eventually lead to convulsions and respiratory arrest. Direct cardiotoxicity can cause cardiac arrest, and even

small doses increase pulse rate and blood pressure. Ventricular

fibrillation and sudden death may result, which sets cocaine apart

from other local anesthetics, such as lidocaine and procaine, which

tend to desensitize the myocardium to fatal arrhythmia. The fatal

oral dose of cocaine is stated to be 1.2 g, but as little as 20 mg

applied to mucous membranes can cause death, and fatal anaphylactic

shock has been reported with use of even lower doses (3). A recent

dose-response study of cocaine in human volunteers showed that

100 mg administered intranasally or 10 mg given intravenously were

sufficient to induce significant subjective and cardiovascular

effects (5). Continued nasal inhalation, commonly called “snorting,” can lead to necrosis of the nasal mucosa.

Although true physical dependence and tolerance are not proven

for cocaine, a very strong psychological dependence can develop.

The cocaine habit is said to be the most seductive, rapidly injurious, and difficult to eradicate of all drug habits (4). Frequent

use of large doses may lead to weight loss, insomnia, and anxiety

(2) With continued high-dose use, a true toxic psychosis may

result as with amphetamine, causing symptoms of paranoia and violent

behavior (3).

PHARMACOKINETICS AND METABOLISM

Cocaine is metabolized rapidly, principally by de-esterification

[hydrolysis by plasma esterases (6) and liver enzymes (7)] and Ndemethylation. These processes result in the formation of benzoylecgonine (II), ecgonine (III), ecgonine methyl ester, and several

N-demethylated products. Of these metabolites, only norcocaine (IV)

has shown biological activity equivalent to cocaine in the inhibition of 3H-norepinephrine uptake by rat brain synaptosomes (8). A

radioisotope study with two human volunteers demonstrated that Ndemethylation does take place in man after oral administration of

cocaine (9).

91
Benzoylecgonine and ecgonine are highly polar and therefore they

do not cross the blood-brain barrier in pharmacologically-significant concentrations. However, norcocaine has been found in the

brains of monkeys given repeated doses of cocaine (8), and the

N-demethylation of cocaine has been shown to occur in the brain of

rats (10). The lipophilic nature of cocaine and norcocaine helps

to account for their penetration across the blood-brain barrier.

In animal studies which compared the disposition and pharmacokinetics of radiolabeled cocaine in acute versus chronically treated

rats, chronic treatment led to consistently higher concentrations

of cocaine being sequestered in fat depots. Although cocaine disappeared rapidly from brain in acute treatment, measurable amounts

persisted in brain and other tissues of chronically treated rats

long after the disappearance of cocaine from plasma (11). In dogs,

however, brain-to-plasma ratios of cocaine were lower in chronically

than in acutely treated animals 2 to 4 hr after injection, and unmetabolized cocaine did not persist in the CNS (12). Radiolabel

studies in rats have also shown localization of cocaine or its

metabolites in other tissues, including kidney, lung, spleen, testes,

intestine, muscle, liver, and heart (13).

In man less than 20 percent, and often only 1 percent, of the parent

drug is excreted in urine; the amount excreted is related to dose

and urine acidity. Cocaine is also found in the bile. The principal urinary metabolites, II (25-54 percent of dose) and III, are

highly water soluble and are largely excreted within 24-36 hr after

administration of cocaine (14). A time-course study of the urinary

excretion of benzoylecgonine following intranasal administration

of cocaine in human subjects showed the appearance of the metabolite in 1-4 hr and a peak urine concentration at 10-12 hr (15).

In another study, peak urine concentrations of benzoylecgonine (up

to 75 μg/ml) occurred 1-12 hr after nasal inhalation of clinical

doses of cocaine (1.5 mg/kg) and diminished slowly over several

days; it was still detectable by radioimmunoassay at 144 hr in one

patient. Maximum urine concentrations of unchanged cocaine (up to

24 μg/ml) occurred within 2 hr of intranasal administration and

diminished rapidly. Benzoylecgonine/cocaine ratios in urine were

too varied among individuals to be of use in predicting cocaine

concentrations from benzoylecgonine data or vice versa (16).

Following an intravenous dose of 100 mg of cocaine hydrochloride to

three human subjects, cocaine concentrations in plasma peaked at

5 min and then declined over 5-6 hr, with a distributional halflife in the plasma of 20-40 min, corresponding to the time-course

of psychologic effects. The mean biologic half-life of cocaine in

plasma was reported to be 2.8 hr. In the same study, benzoylecgonine

was detected in plasma 5 min after intravenous administration of

cocaine. Plasma concentrations of the metabolite increased rapidly

in the first hour, remained stable for 2-4 hr, and then declined

slowly. After 24 hr, the benzoylecgonine was still detectable.

The persistence of benzoylecgonine in plasma suggested that it was

strongly bound to plasma proteins (17). The pharmacokinetic measurements were comparable to those reported for patients receiving

cocaine in the form of a topical anesthetic, and indicated rapid

92
absorption of cocaine through mucous membranes. Plasma concentrations of cocaine in another human study increased rapidly for

15-20 min after intranasal application of 1.5 mg/kg, peaked at

15-60 min, and then decreased gradually for 3-5 hr. Maximum plasma

concentrations ranged from 120 ng/ml to 474 ng/ml (18,19). In

cases of acute intoxication, blood concentrations of cocaine as

low as 200 ng/ml have been associated with fatalities, although

concentrations up to 9.0 μg/ml are more commonly reported in cocaine deaths when no other drugs were present (20).

ANALYTICAL METHODOLOGY

Because of its rapid metabolism, detection and quantitation of cocaine in body fluids is difficult. In addition to the low concentrations likely to be encountered in blood and in urine (21), the

assay of cocaine presents technical problems caused by hydrolysis

of the drug by plasma enzymes and under conditions of storage and

laboratory manipulation. Even when stored in a frozen state, 40

percent of the cocaine in plasma was shown to be lost within 5

days, and 60 percent within a month; at room temperature 45 percent of the cocaine in plasma was lost within 1 hr. However,

addition of sodium fluoride (2.5 mg/ml plasma) to the plasma to

inhibit enzymatic hydrolysis prevented any appreciable loss of

cocaine in plasma maintained at room temperature for 2 hr (22).

Under strongly basic (pH > 10) or acidic (pH< 5) conditions, chemical hydrolysis of cocaine to benzoylecgonine and ecgonine is accelerated. For these reasons, measurement of the cocaine metabolites,

particularly benzoylecgonine, has been emphasized in the development of assay procedures. However, the metabolites of cocaine are

very polar and also amphoteric; they are not readily extractable

into the organic solvents normally used for drug screening, and

therefore specialized methods must be employed for their recovery

from biological materials,

Colorimetric techniques, which are at best semiquantitative, have

been used to determine cocaine in urine (23) and other biological

fluids. An early study on the distribution and metabolism of cocaine in dogs and rabbits was based on such a method; it involved

extraction with chloroform, passage of the extract through a sucrose column, and addition of bromcresol purple. Optical densities of the resulting yellow solutions were determined on a spectrophotometer at 410 nm, and a sensitivity of 0.5-1.0 μg/ml in

urine, blood, and tissue homogenates was reported (24,25). Spectrophotometric methods, however, are neither specific nor sufficiently sensitive for use in most cases of cocaine analysis.

Thin-layer chromatography (TLC), commonly used for routine identification of abused drugs in urine, requires special solvent systems

for separation as well as for extraction of cocaine metabolites.

In one TLC method, cocaine and its metabolites were extracted

from urine with 25 percent ethanol in chloroform at pH 10-11 and

chromatographed on silica gel plates developed in methanol:ammonia

(99:1) and sprayed with an iodoplatinate reagent. The solvent system separated ecgonine from benzoylecgonine and cocaine, but to

93
separate the latter two compounds from each other it was necessary to use ethyl acetate:methanol:ammonia (85:10:10) (26). Another TLC assay involved extraction with a mixture of chloroform:

isopropanol:dichloroethane (8:1:3) at pH 8.5. The sensitivity

for benzoylecgonine analyzed by this procedure was 3-5 μg/ml (27).

More recently, a TLC method using a double solvent system on silica gel sheets involved a butylation step after extraction of cocaine and its metabolites from urine with chloroform:ethanol

(3:2) at pH 8-9; it provided a sensitivity of 3 μg/ml (28). The

sensitivity limits for TLC of cocaine and benzoylecgonine in urine

were subsequently lowered to 0.1 and 0.25 μg/ml, respectively, by

spraying the developed plates with Dragendorff’s reagent followed

by an overspray with dilute H2SO4 and exposure to iodine vapors

(29). Another refinement of TLC detection procedures involved

preliminary absorption of the compounds onto paper loaded with

SA-2 cation exchange resin. Extraction from the resin was made

with chloroform: isopropanol: dichloroethane (4.5:0.9:4.5); sensitivity for cocaine and its metabolites was 1 μg/ml. in 20 ml of

urine. If other drugs were to be assayed in the same urine sample, cocaine and the other drugs were first extracted from the paper at pH 10 and chromatographed in a two-stage solvent system.

Benzoylecgonine was extracted from the aqueous buffer phase after

acidification and readjustment of the pH to 8.1-8.4. The sensitivity limit for benzoylecgonine was 2 μg/ml. Alternatively, benzoylecgonine alone was directly extracted from 5 ml of urine with the

chloroform: isopropanol: dichloroethane solvent and was detected at

a sensitivity limit of 0.5 μg/ml by TLC using Dragendorff’s reagent

and an iodine-potassium iodide reagent (30).

The enzyme-multipled immunoassay technique (EMIT) is widely used

in urine screening for cocaine abuse, and has a detection limit of

1 μg/ml of benzoylecgonine. No extraction is required, and the

assay takes less than 2 min to perform. It does not detect cocaine

nor is it applicable to plasma (31). A commercially available radioimmunoassay has been developed based on a 125I-labeled benzoylecgonine derivative (32). The RIA is now widely used for screening

urine samples for benzoylecgonine and can also be used for quantitative measurement of “benzoylecgonine equivalents” which include

cocaine and other tropane alkaloids which cross-react with the

antibody. The assay will detect benzoylecgonine at urine concentrations as low as 2 ng/ml, but 100 ng/ml has been recommended as

the “cutoff” concentration for positives in routine screening. An

evaluation of the RIA method has been published in which it is compared with GC, TLC and EMIT methods for analysis of benzoylecgonine

in urine (33).

Quantitative analysis of cocaine in biological fluids has relied

primarily on the development of gas chromatographic methods (GC).

Cocaine can be gas chromatographed without derivatization, but its

metabolites cannot. The carboxylic acid groups of II and III must

be esterified with some reagent such as diazomethane, dimethylformamide dimethylacetal, N,N-dicyclohexyl carbodimide, or n-butanolboron trifluoride. Esterification with 2-chloroethanol in the pre94
sence of boron trichloride has the advantage of introducing a halogenated substituent which renders the molecule suitable for electron-capture detection (34). A solvent extraction procedure can

be used to separate benzoylecgonine and ecgonine from basic and

neutral components of biological extracts prior to esterification

and GC analysis of the metabolites (35).

Gas Chromatography (GC) with flame-ionization detection (FID) has

often been used for detection of cocaine and metabolites, especially in urine. In one of the first studies of human cocaine metabolism, GC-FID with benzhexol as an internal standard was used for

measurement of cocaine in urine. Chromatography was on a 3 percent OV-17 column at 185°C, and a linear response was obtained

over a range of 0.5-15.0 μg/ml. Benzoylecgonine was assayed in

this system by continuous extraction with chloroform containing 5-

-cholestane as an internal standard; the extracted benzoylecgonine was methylated to cocaine for gas chromatography (36,37). GCFID on a 2.5 percent SE- 30 column at 200°C was reported as a method

capable of detecting 1 μg/ml of benzoylecgonine in urine. The benzoylecgonine was salted out of the urine into 95 percent ethanol,

purified by TLC, eluted in water, and methylated before gas chromatography . The overall recovery was only 30-40 percent, but an efficiency of 90-100 percent was reported for the initial ethanol extraction (38). Cocaine can be mistaken for pentazocine, levorphenol, or methaqualone on the OV-17 columns commonly used for drug

screening; for this reason, an on-column reaction of cocaine with

trimethyl anilinium hydroxide, which yields an unidentified product with a different retention time, has been suggested as a confirmatory step in the detection of cocaine in urine (39).

For concurrent determination of 9 CNS-active drugs in human plasma,

including cocaine, GC-FID analysis on a 3 percent OV-1 column followed

extraction with benzene-isopropanol at pH 10. Recovery of cocaine

was about 90 percent, and sensitivity limits corresponded to plasma

drug concentrations of 0.05 μg/ml. The technique had the advantage

of comparative simplicity for routine purposes (40). Another procedure designed to determine both cocaine and benzoylecgonine in

urine involved methylation of the metabolite, converting it to cocaine in 72 percent yield before GC-FID analysis on a 3 percent OV17 column. Butylanthraquinone was used as an internal standard.

The extraction technique (chloroform:ethanol, 4:1) gave a recovery

of 93 percent for cocaine and 65 percent for benzoylecgonine at pH

5.5-9.5. The sensitivity reported was 0.20 μg/ml in a 5-ml specimen, which was adequate for the measurement of urinary drug and

metabolite concentrations of postsurgical patients up to 24 hr after

topical cocaine anesthesia (250 mg applied to nasal mucosa) (41).

Similarly, simultaneous determination of cocaine and its metabolites

in urine, using gas chromatography on columns coated with 3 percent

SP-2250-DA or 3 percent SE-30, has been reported. In this case extraction was into chloroform:ethanol (3:1), followed by propylation

of benzoylecgonine using an organic base and 1-iodopropane. Recovery of 99 percent of the cocaine and 80 percent of the benzoylecgonine, combined with 98 percent conversion of the metabolite to its

95
n-propyl ester, provided detection limits of 0.2 μg/ml in 5 ml of

urine. Back extraction and purification were required, and also

synthesis of the n-pentyl ester of benzoylecgonine for use as an

internal standard. The procedure measured concentrations ranging

from 0.2 to 18 μg/ml of benzoylecgonine in samples of urine from

drug screening programs, and demonstrated concentrations of less

than 0.3 μg/ml of cocaine and 8-136 μg/ml of benzoylecgonine in

pooled 8-hour urine samples from postsurgical patients following

cocaine anesthesia (42).

A method reported for simultaneous analysis of cocaine and benzoylecgonine in blood, bile, urine or tissue involved an extractive

alkylation step to convert benzoylecgonine to its ethyl ester which

was then readily separated from cocaine by gas chromatography (3 percent OV-17, 225°C). n-Propyl benzoylecgonine was used as the internal standard (43).

The sensitivity of GC analysis for cocaine can be increased up to

100-fold by use of a nitrogen-phosphorus detector. With this type

of detector sub-nanogram quantities of the drug can be measured according to a reported procedure which involved the use of the propyl

ester of benzoylecgonine as an internal standard, back-extraction

into 0.1 H2SO4, and chromatography on an OV-17 column at 255°C (21).

Another application of GC with nitrogen-phosphorus detection for

the measurement of cocaine in blood involved an ethylmorphine internal standard, extraction with benzene:isopropanol (9:1) , and

chromatography on a 3 percent SP-2100-DB column. Following topical

application of cocaine to nasal mucosa, as little as 20 ng of cocaine

was detected in 1 ml of whole blood by this method (44).

An electron-capture detector has also been used for the measurement

of benzoylecgonine in blood. The procedure involved extraction of

the metabolite and an internal standard (chlorproethazine) from plasma

with chloroform: ethanol (4:1) at pH 9.5. The dried extract was alkylated with pentafluorobenzyl bromide and the derivative partitioned

into benzene to eliminate more polar interfering substances, then

reextracted into acid. Sensitivity for the method was 5 ng of benzoylecgonine per ml of plasma. However, the presence of a background

contaminant peak in urine decreased the sensitivity of the procedure

to 50 ng/ml for urine analysis (17).

Measurement of 20-30 ng/ml of cocaine in urine, without the necessity of evaporative concentration procedures, was achieved in an

adaptation of the GC-ECD technique which used an 0-acylated derivative of reduced cocaine (heptafluorobutyric anhydride or pentafluoropropionic anhydride added to the cyclohexane extract following reduction with LiA1H4

) (45). A very similar procedure was subsequently

used to measure cocaine in plasma as well as urine (46).

High performance liquid chromatography (HPLC) on a reverse phase

column was used for separation and measurement of cocaine and benzoylecgonine extracted from urine. A detection limit of 0.1 μg/ml

of urine was achieved by monitoring the ultraviolet absorption of

the HPLC effluent at 200 nm. The chromatography procedure also per96
mitted the separation and detection of norcocaine and benzoylecgonine; however, debenzoylated metabolites such as ecgonine could

not be detected because of insufficient UV absorption, even at 200

nm (47).

The high specificity and sensitivity of gas chromatography/mass spectrometry (GC/MS) has been applied to the analysis of cocaine and

its metabolites in biological fluids (48-50). Concentrations of

cocaine and norcocaine in human urine were determined by a procedure which involved initial extraction at pH 8.5 into cyclohexane

followed by back-extraction into 0.1 N HCl, and after adjustment

to basic pH with 1 N NH4OH, reextraction with cyclohexane. Before

GC/MS analysis the norcocaine was converted to its N-trifluoroacetamide derivative. Deuterium labeled analogs were used as internal standards. Cocaine-2H3

 was prepared by treatment of benzoylecgonine with dimethylformamide dimethyl-2H6

 acetal (Pierce, Rockford, Illinois), and norcocaine-2H3

 was obtained by N-demethylation of the deuterated cocaine, The assay measured 2 ng/ml of cocaine and norcocaine with a precision of 5 percent (48). A similar

assay was developed by the same research group for quantitation of

cocaine and benzoylecgonine. Deuterium-labeled internal standards

were again used, except the deuterated cocaine was prepared by alkylation of norcocaine with C

2H31 and benzoylecgonine-2H3

 was generated

from the labeled cocaine. Chloroform: isopropanol (4:1) was used to

extract the benzoylecgonine from urine after adjustment of the pH

to 9 with 0.1 N NH4OH. The extracted benzoylecgonine was then converted to its ethyl ester by treatment with diazoethane generated

by addition of N-ethyl-N-nitro-N-nitrosoguanidine to a biphasic

solution of KOH and ether. Electron impact ionization gave molecular ions of moderate abundance for cocaine (m/z 303) and benzoylecgonine ethyl ester (m/z 317) which were monitored together with

the molecular ions for the deuterated internal standards. When all

systems were working well, an assay sensitivity of approximately

2 ng/ml of cocaine and 5 ng/ml of benzoylecgonine could be achieved

(49) 

In another application of GC/MS to the assay of cocaine metabolites

in urine, benzoylecgonine was simultaneously extracted from urine

and converted to its ethyl ester by a 10-min extractive-alkylation

procedure. The butyl ester of benzoylecgonine served as the internal

standard. Quantitation was accomplished by GC/MS using probabilitybased matching (SO).

HPLC is promising as a means of purifying polar metabolites extracted

from body fluids; advantages are extraction selectivity and adaptability to automation (51). Other reported extraction procedures vary

with the analytical method and the purpose of the assay. Cocaine is

easily extracted from alkaline aqueous media into organic solvents,

including cyclonexane (12,45), ether (36,37), 2 percent isoamyl

alcohol in n-heptane (17,18,21), benzene:isopropanol 9:1 (recovery

85-90 percent) (40,44), and chloroform (recovery 85-100 percent)

(24). The recovery of cocaine is best at pH 9.5 (93 percent from

chloroform:ethanol). It drops to 76 percent at pH 10.5 because of

hydrolysis (52).

97
Benzoylecgonine can be removed from the aqueous phase by exhaustive

continuous extraction with chloroform; by single extraction with

chloroform:isopropanol:1,2-dichloroethane (4.5:0.9:4.5) (30) or

chlorofornnethanol (4:1, recovery 65-80 percent) (17,41,42); salted

out together with other polar metabolites into alcohol from a K2CO3

solution or a KH2PO4-K2HPO4 solution (38); or extracted into methanol from lyophilized urine (90-100 percent recovery) (38). The

lower pH possible with salting out procedures avoids saponification

of benzoylecgonine. Adsorption on nonionic (e.g., XAD-2) resins,

followed by elution with chloroform: isopropanol (3:1), is recommended for extraction of some cocaine metabolites from tissue homogenates, but ecgonine and norecgonine are not adsorbed by the resin

(35).

Two recently-published books review information on the history,

chemistry, pharmacology, analysis, and sociological aspects of cocaine (1,53).

EXPERIMENTAL PROCEDURE

Because cocaine is rapidly metabolized it is often desirable to analyze for both the parent drug and its major metabolite, benzoylecgonine. Therefore, two procedures are described; both are modifications of published methods developed at the Center for Human Toxicology, University of Utah (54). Procedure 1 consists of a simple,

direct extraction which permits rapid measurement of cocaine concentrations only. Procedure 2 takes longer, but permits simultaneous

measurement of both cocaine and benzoylecgonine concentrations.

The latter procedure consists of extraction with chloroform: isopropanol from a sodium chloride-saturated solution of the body fluid,

derivatization of the extracted benzoylecgonine by heating with N,Ndimethylformide di-n-propyl acetal, sample purification by backextraction of the basic drug and the derivatized metabolite into

acid, and finally adjustment to basic pH and reextraction into an

organic solvent prior to quantitation by (GC/MS.

Cocaine is rapidly hydrolyzed by plasma esterases to ecgonine methyl

ester (55), so it is important to inhibit these enzymes as soon as

blood samples are collected for subsequent cocaine analysis (56).

This can be done by collecting the sample in a tube containing sodium fluoride, such as the grey stoppered 7-ml Venoject tubes (Kimble-Terumo, Inc., Elkton, MD 21921) which are supplied with 17.5 mg

of solid sodium fluoride and 14 mg of potassium oxalate. An additional 50 μl of a saturated sodium fluoride solution is added for

each ml of blood. The sample is then centrifuged and the plasma

transferred to a Teflon-lined, screw cap culture tube and frozen

until analysis.

Standards and Reagents

Cocaine hydrochloride and benzoylecgonine of high purity can be purchased from Applied Science laboratories, State College, PA 16801.

Cocaine hydrochloride labeled with 3 deuterium atoms on the N-methyl

group (cocaine-2H3 HCl), and benzoylecgonine labeled with deuterium

98
on the benzoyl group (benzoylecgonine-2H3

) were obtained from the

National Institute on Drug Abuse. On the basis of mass spectral

analysis the cocaine-2H3 had an isotopic composition of: 2H3, 96

percent; 2H2, 1 percent; 2H1, 1 percent; and 2H0, 0 percent. The

isotopic composition of the benzoylecgonine-2H3 was:

2H5

, 7 percent;

2H4

, 31 percent; 2H3, 40 percent; 2H2, 21 percent; 2H1

 and 1H1, 0

percent.

Prepare a pH 9.6 buffer by dissolving 500 grams of potassium monohydrogen phosphate (K2HP04

) in about 500 ml of distilled water.

Allow the solution to cool to room temperature and then add sufficient distilled water to make exactly one liter.

N,N-Dimethylformamide di-n-propyl acetal was purchased from the

Aldrich Chemical Company, Milwaukee, WI 53233.

Stock solutions of cocaine and benzoylecgonine are prepared as follows for the purpose of determining calibration graphs and preparing

working standards. Weigh into a 100-ml volumetric flask about 11.2

mg of cocaine hydrochloride (equivalent to 10 mg of free base) and

10 mg of benzoylecgonine, and record the weights to the nearest 0.1

mg. Dissolve the measured drug and metabolite in about 75 ml of

methanol and then add additional methanol to give exactly 100 ml

of solution. This solution will be referred to as the “0.10 mg/ml

stock solution” although it should be labeled with its actual concentration based upon the exact measured weights of cocaine and benzoylecgonine. A series of working standard solutions can then be

prepared by appropriate dilution of the stock solution as described

in Chapter 2.

The stock solution of cocaine-2H3 and benzoylecgonine-2H3 is prepared in the same manner. For the measurement of drug and metabolite in body fluids within the concentration range 1 to 1000 ng/ml,

addition of approximately 40 ng of the internal standards to each

ml of body fluid is satisfactory. This can be conveniently done by

adding 100 μl of a methanolic solution containing 400 ng/ml of both

cocaine-2H3 and benzoylecgonine-2H3 to each ml of body fluid. Prepare the 400-ng/ml internal standard solution by diluting 1 ml of

the 0.10-mg/ml stock solution of cocaine-2H3 and benzoylecgonine2H3

 to 250 ml with methanol.

Store stock and working standard solutions in well-stoppered or capped glass vessels in the dark at less than 0°C.

Extraction - Method 1 (Cocaine only)

Transfer 1 ml of whole blood, plasma or urine to a 5-ml glass-stoppered, conical centrifuge tube. Add 100 μl of the internal standard

solution containing cocaine-2H3 and benzoylecgonine-2H3 at concentrations of 400 ng/ml. Vortex the mixture for 10 sec and then allow

it to equilibrate for 15 min. Add 1 ml of K2HPO4

 buffer (pH 9.6)

followed by 100 μl of a solvent consisting of toluene:heptane:isoamyl alcohol in a volume ratio of 70:20:10. Immediately stopper

the tube and vortex for at least 30 sec. Centrifuge the mixture at

2000 x G for 15 min. The organic solvent should form a narrow upper

99
layer from which an aliquot can be removed with a syringe for injection into the GC/MS. If a distinct solvent layer is not obtained, add an additional 100 μl of organic solvent and recentrifuge.

Extraction - Method 2 (Cocaine and Benzoylecgonine)

Transfer 1 ml of whole blood, plasma or urine to a 20-ml culture

tube equipped with a Teflon-lined screw cap (0.5 to 2 ml of sample can be used, but it must be measured accurately,). Add 100 μl

of the internal standard solution containing cocaine-2H3 and benzoylecgonine-2H3

 at concentrations of 400 ng/ml. Vortex the mixture for 10 sec and allow it to equilibrate for 15 min. Check the

pH. If it is not 7.0 ± 0.5, add 1 ml of a pH 7 buffer. Saturate

the mixture with sodium chloride (about 0.5 g) to improve the extraction efficiency of benzoylecgonine. Add 8 ml of a solvent mixture of chloroform:isopropanol (9:1 v/v). Cap the tube and check

for a leak around the cap. Gently mix the contents for 30 min using

a motorized rocker or rotator. Centrifuge for 5 min, remove the

upper aqueous layer by aspiration, and transfer the lower organic

layer into a 12-ml conical, glass-stoppered centrifuge tube. If

particulate matter is present in the organic extract, it should be

removed by filtering the organic extract through silylated glass

wool. It is important that the transferred organic extract be absolutely dry and devoid of any of the lipid interface that forms

between the phases. Remove the solvent by evaporation with a gentle stream of nitrogen or filtered air and heat at not greater than

70°C. All of the volatile solvents must be removed! Add 50 μl of

dimethylformamide and 50 μl of N,N-dimethylformamide di-n-propyl

acetal. Reflux the mixture for 30 sec over an open flame (see Discussion of the Experimental Procedure). Allow the mixture to cool

to room temperature and then add 1 ml of 0.5 N H2SO4. Vortex mix

for about 10 sec. Add 3 ml of a solvent system consisting of toluene:heptane:isoamyl alcohol in a volume ratio of 70:20:10. Vortex

for 30 sec and then centrifuge. Remove the upper organic phase by

aspiration. Neutralize the aqueous acid phase by adding 1 ml of

0.5 N NaOH and 1 ml of the K2HPO4 buffer. Vortex mix and check that

the pH is between 9 and 10. If it is not, adjust by adding a few

drops of 0.5 N NaOH or 0.5 N H2SO4, as appropriate. Immediately

extract the cocaine and benzoylecgonine n-propyl ester from the

basic aqueous solution by adding 3 ml of the toluene:heptane:isoamyl alcohol (70:20:10) solvent, vortex for 30 sec, and centrifuge.

Transfer by means of a disposable Pasteur pipette the upper organic

layer to a silylated, 12-ml concentrator tube having a conical or

nipple-shaped bottom. Remove the solvent by evaporation under a

gentle stream of nitrogen or filtered air and at a temperature not

greater than 70°C. Reconstitute the extract residue in 30 μl of

chloroform for GC/MS analysis.

GC/MS Analysis

The experimental conditions for the GC/MS analysis of cocaine and

benzoylecgonine n-propyl ester are as follows:

100
GC column:

Carrier gas:

Reagent gas:

Temperatures

1.8 m x 2mm (I.D.) glass column packed with

3 percent OV-1 on Gas Chrom Q 100/120 mesh

(Applied Science Laboratory, ‘State College,

PA 16801)

Methane, 15-20 ml/min

Ammonia, introduced into the ion chamber as

described in Chapter 2

Injector, 280°C

Column, 230°C

GC/MS transfer line, 280°C

Ion source, 160°C

Under these conditions the cocaine should elute from the gas chromatograph at between 2 and 4 min, and the benzoylecgonine n-propyl

ester should elute at between 3 and 5 min.

Before beginning the GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored are the protonated

molecule ions for cocaine (m/z 304), cocaine-2H3 (m/z 307), benzoylecgonine n-propyl ester (m/z 332), and benzoylecgonine-2H3

 npropyl ester (m/z 335). With the divert valve in the divert position, inject into the GC/MS 2 to 6 μl of the organic extract from

either procedure 1 or 2. After approximately 1 min switch the divert valve so that the entire carrier gas flow enters the ion source

of the mass spectrometer and begin data acquisition. If the extract

from procedure 1 is being analyzed, only the cocaine ion masses (m/z

304 and 307) are monitored. For analysis of extracts from procedure

2 the best sensitivity is achieved by first monitoring only the cocaine ion masses until the cocaine has eluted; then the benzoylecgonine n-propyl ester ion masses are monitored. If the GC/MS data

system does not permit rapid changes of the monitored ion masses, all

four ion masses must be monitored during the elution of the drug

and the derivatized metabolite. Normally, the latter procedure will

result in only a small sacrifice in sensitivity. When the benzoylecgonine n-propyl ester has eluted, cease data acquisition and return the divert valve to the divert position. The quantities of cocaine and benzoylecgonine in the body fluid are determined by measuring the peak heights (or areas) of the drug, the derivatized metabolite, and their internal standards in the selected ion current profiles, and relating the ratios of peak heights to calibration graphs

(Chapter 2). Calculate the body fluid concentrations of cocaine and

benzoylecgonine by dividing the measured quantities of these compounds by the exact volume of body fluid extracted.

Discussion of the Experimental Procedure

Benzoylecgonine must be converted to a volatile derivative before it

can be gas chromatographed. Various methods of derivatization have

been used, including methylation to cocaine (36,37,41), propylation

by treatment with 1-iodopropane (42), alkylation with pentafluoro101
benzyl bromide (17), and ethylation by treatment with diazoethane

(38) or by extractive alkylation with iodoethane (43,49). The

method of derivatization described here is simple and efficient if

the procedure is followed precisely. The reaction of N,N-dimethylformamide di-n-propyl acetal with benzoylecgonine is rapid and quantitative if the reaction mixture is heated to a sufficiently high

temperature (~150°C). Temperatures of this magnitude are conveniently achieved by repeatedly passing the bottom of the reaction

tube slowly through the tip of a small, open flame until the reaction mixture just begins to boil. Evolution of a white vapor will

occur. If all of the volatile solvents (chloroform and isopropanol)

have not been removed prior to the addition of the derivatizing

agent, the reaction mixture will boil at a lower temperature and

the reaction may not go to completion.

Other N,N-dimethylformamide acetals have been tried, but the di-npropyl acetal was chosen because it gives a benzoylecgonine ester

which has a particularly suitable retention time under the gas chromatographic conditions specified. The N,N-dimethylformamide di-npropyl acetal was purchased from the Aldrich Chemical Company, Milwaukee, WI 53233. Some difficulty was encountered with impurities

in samples of this reagent purchased from a different supplier.

Figure 1 compares the electron impact (EI) and the chemical ionization (CI) mass spectra of cocaine. The corresponding mass spectra

of the propyl ester of benzoylecgonine are similar except that the

abundant ions in the CI mass spectra are shifted 28 daltons higher.

The CI mass spectrum of cocaine obtained using methane and ammonia

as reagent gases shows only one abundant ion, which corresponds to

the protonated molecule ion (m/z 304). The protonated molecule ion

peak in the methane CI mass spectrum has a relative intensity of

about 50 percent of the base peak at m/z 182, which corresponds to a

fragment ion resulting from loss of benzoic acid from the protonated

molecule ion. The EI mass spectrum has a moderately abundant ion

at m/z 182 which could be used for selected ion monitoring analysis

of cocaine, However the analogous ion (m/z 210) in the EI mass

spectrum of the propyl ester of benzoylecgonine would not be a

satisfactory ion to monitor if the deuterium atoms in the internal

standard are located on the benzoyl group because they would be

lost during formation of this fragment ion.

If just the protonated molecule ions are monitored, the methane-ammonia CI gives about a threefold improvement in sensitivity over

methane CI. The methane-ammonia CI also gives the highest selectivity. On the other hand, when methane alone is used as the reagent

gas, the ion masses corresponding to both the protonated molecule

ion and the MH+-PhCO2H fragment ion can be monitored. This will provide additional confidence in the quantitative results because any

co-eluting extract component is unlikely to contribute equally to

the ion currents at both of the ion masses monitored.

Figures 2 and 3 show representative calibration graphs acquired using

extraction procedures 1 and 2, respectively. Table 1 gives accuracy

and precision data obtained using both procedures.

102
FIGURE 1. MASS SPECTRA OF COCAINE

103
FIGURE 1. CALIBRATION GRAPH FOR ANALYSIS OF COCAINE

IN PLASMA BY PROCEDURE 1

FIGURE 3. CALIBRATION GRAPH FOR ANALYSIS OF

IN PLASMA BY PROCEDURE 2

COCAINE

104
TABLE 1. ACCURACY AND PRECISION DATA

Body fluid concentrations of cocaine and benzoylecgonine sometimes

exceed 1 μg/ml. When this occurs the sample should be diluted by

a measured amount with distilled water so that the resulting concentration falls within the range of 0 to 1000 ng/ml. The diluted

sample can then be analyzed as described.

Cocaine, and to a lesser extent benzoylecgonine, can undergo enzymatic and/or chemical hydrolysis (54). The rate of chemical hydrolysis

is greatly accelerated at alkaline pH. For example, at pH 9.8, 40

percent of the cocaine in a 3 μM solution was converted to benzoylecgonine in 1 hr at 37°C (54). Also, benzoylecgonine in methanol

can undergo reestcrification to cocaine, particularly if acid is

present. For these reasons, all body fluid samples to be analyzed

for cocaine should be stabilized with sodium fluoride and stored in

a frozen state. Aqueous solutions of cocaine and benzoylecgonine

and methanolic solutions of benzoylecgonine should be stored under

refrigeration and for no longer than a few days.

105
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR COCAINE

Nomenclature

Chemical Name: 3-Benzoyloxy-8-methyl-8-azabicyclo[3.2.1]octane-2-

carboxylic acid methyl ester

Empirical Formula: C17H21N O4

Chemical Registry Number: 50-36-2

Physical Constants

Appearance: White crystalline powder

Melting Point: 98°C

Boiling Point: 187-188°C @ 0.1 mm

Specific Rotation: = -16° (c = 4 in chloroform)

Solubility: 1 g of cocaine dissolves in 600 ml of water, 6.5 ml

of ethanol, 0.7 ml of chloroform, and 3.5 ml of diethyl

ether.

pKa: 8.70

UV Absorption: max = 231 nm (log = 4.2) and 274 nm (log = 3.0)

PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR BENZOYLECGONINE

Nomenclature

Chemical Name: 3-Benzoyloxy-8-methyl-8-azabicyclo[3.2.1]octane-2-

carboxylic acid

Empirical Formula: C16H19NO4

Chemical Registry Number: 519-09-5

Physical Constants

Appearance: White crystalline powder

Melting Points: 195°C with decomposition, the tetrahydrate melts

at 86°C

Specific Rotation: = -45° (c = 3 in ethanol)

Solubility: Very soluble in hot water, soluble in ethanol, insoluble

in ether

UV Absorption: max = 234 nm and 275 nm (in 0.1 N H2S04

)

106
REFERENCES

1.

2.

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

R.C. Petersen and R.C. Stillman, eds., Cocaine: 1977. National Institute on Drug Abuse Research Monograph 13, DHEW

Pub. (ADM) 77-432. Supt. of Docs., U.S. Govt. Print. Off.,

Washington, D.C. (May 1977).

S. Cohen. J. Amer. Med. Assoc., 231, 74 (1975).

G.R. Gay, C.W. Sheppard, D.S. Inaba, and J.A. Newmeyer. Int.

J. Addictions, 8, 1027 (1973).

T. Harwood. Drug Enforcement, 1, 20 (Spring 1974).

R.B. Resnick, R.S. Kestenbaum, and L.K. Schwartz. Science,

195, 696 (1977).

D. Taylor, V.S. Estevez, L.F. Englert, and B.T. Ho. Res.

Commun. Chem. Pathol. Pharmacol., 14, 249 (1976).

E.G. Leighty and A.F. Fentiman, Jr. Res. Commun. Chem. Pathol.

Pharmacol., 8, 65 (1974).

R.L. Hawks, I.J. Kopin, R.W. Colburn, and N.B. Thoa. Life

Sciences, 15, 2189 (1974).

T. Inaba, D.J. Stewart,

23, 547 (1978).

A.L. Misra, P.K. Nayak,

Mulé. Experientia, 30,

P.K. Nayak, A.L. Misra,

Ther., 196, 556 (1976).

A.L. Misra, M.N. Patel,

and W. Kalow. Clin. Pharmacol. Ther.,

M.N. Patel, N.L. Vadlamani, and S.J.

1312 (1974).

and S.J. Mulé. J. Pharmacol. Exp.

V.R. Alluri, S.J. Mulé, and P.K. Nayak. Xenobiotica, 6, 537 (1976).

A.L. Misra in S.J. Mulé, ed., Cocaine: Chemical, Biological,

Clinical, Social, and Treatment Aspects. CRC Press, Cleveland, OH, p. 73 (1976).

R. Byck and C. Van Dyke in R.C. Petersen and R.C. Stillman,

eds., Cocaine: 1977, ch. V. See reference 1.

C.Van Dyke, R. Byck, P.G. Barash, and P. Jatlow. Clin.

Chem:, 23, 241 (1977).

H.E. Hamilton, J.E. Wallace, E.L. Shimek, Jr., P.Land, S.C.

Harris, and J.G. Christensen. J. Forensic Sci., 22, 697

(1977).

M.J. Kogan, K.G. Verebey, A.C. DePace, R.B. Resnick, and S.

J. Mulé. Anal. Chem., 49, 1965 (1977).

107
18. R. Byck, P. Jatlow, P. Barash, and C. Van Dyke. Psychopharmacol. Bull., 12, 47 (1976).

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

C. Van Dyke, P. Barash, P. Jatlow, and R. Byck. Science,

191, 859 (1976).

B.S. Finkle and K.L. McCloskey in R.C. Petersen and R.C. Stillman, eds., Cocaine: 1977, ch. VIII. See reference 1.

P.I. Jatlow and D.N. Bailey. Clin. Chem., 21, 946 (1975).

J.I. Javaid, H. Dekirmenjian, J.M. Davis, and C.R. Schuster

J. Chromatogr., 152, 105 (1978).

H. Bronisz, B. Wysocka, and W. Rusiecki. Disserationes Pharm.,

15, 343 (1963).

L.A. Woods, J. Cochin, E.J. Fornefeld, F.G. McMahon, and M.H.

Seevers. J. Pharmacol. Exp. Ther., 101, 188 (1951).

L.A. Woods, F.G. McMahon, and M.H. Seevers. J. Pharmacol.

Exp. Ther., 101, 200 (1951).

S.J. Mulé. J. Chromatogr., 55, 255 (1971).

N.N. Valanju, M.M. Baden, S.N. Valanju, D. Mulligan, and S.K.

Verma. J. Chromatogr., 81, 170 (1973).

M.L. Bastos, D. Jukofsky, and S.J. Mulé. J. Chromatogr., 89,

335 (1974).

J.E. Wallace, H.E. Hamilton, H. Schwertner, D.E. King, J.L.

McNay, and K. Blum. J. Chromatogr., 114, 433 (1975).

K.K. Kaistha and R. Tadrus. J. Chromatogr., 135, 385 (1977).

Syva Corp. Palo Alto, CA 94304.

Roche Diagnostics. Nutley, NJ 07110.

S.J. Mulé, D. Jukofsky, M. Kogan, A. DePace, and K. Verebey.

Clin. Chem., 23, 796 (1977).

J.M. Moore. Clin. Chem., 21, 1538 (1975).

M.L. Bastos and D.B. Hoffman in S.J. Mulé, ed., Cocaine:

Chemical, Biological, Clinical, Social, and Treament Aspects, p. 35. See reference 13.

F. Fish and W.D.C. Wilson. J. Chromatogr., 40, 164 (1969).

F. Fish and W.D.C. Wilson. J. Pharm. Pharmacol., 21 135S

(1969).

108
39.

40.

38. S. Koontz, D. Besemer, N. Mackey, and R. Phillips. J. Chromatogr., 85, 75 (1973).

R.H. Hammer, J.L. Templeton, and H.L. Panzik. J. Pharm. Sci.,

63, 1963 (1974).

F. Medzihradsky and P.J. Dahlstrom. Pharmacol. Res. Commun.,

7, 55 (1975).

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

J.E. Wallace, H.E. Hamilton, D.E. King, D.J. Bason, H.A.

Schwertner, and S.C. Harris. Anal. Chem., 48, 34 (1976).

D.L. von Minden and N.A. D'Amato. Anal. Chem., 49, 1974 (1977).

J.C. Valentour, V. Aggarwal, M.P. McGee, and S.W. Goza.

J. Anal. Toxicol., 2, 134 (1978).

B.H. Dvorchik, S.H. Miller, and W.P. Graham. J. Chromatogr.,

135, 141 (1977).

J.W. Blake, R.S. Ray, J.S. Noonan, and P.W. Murdick. Anal.

Chem., 46, 288 (1974).

J.I. Javaid, H. Dekirmenjian, E.G. Brunngraber, and J.M.

Davis. J. Chromatogr., 110, 141 (1975).

P.I. Jatlow, C. Van Dyke, P. Bar-ash, and R. Byck. J. Chromatogr., 152, 115 (1978).

S.P. Jindal, T. Lutz, and P. Vestergaard. Biomed. Mass

Spectr., 5, 658 (1978).

S.P. Jindal and P. Vestergaard. J. Pharm. Sci., 67, 811 (1978).

J.E. Graas and E. Watson. J. Anal. Toxicol., 2, 80 (1978).

A.P. Graffeo, D.C.K. Lin, and R.L. Foltz. J. Chromatogr.,

126, 717 (1976).

S.J. Mulé, M.L. Bastos, D. Jukofsky, and E. Saffer. J. Chromatogr., 63, 289 (1971).

S.J. Mulé, ed., Cocaine: Chemical, Biological, Clinical,

Social, and Treatment Aspects. See reference 13.

D.M. Chinn, D.J. Crouch, M.A. Peat, B.S. Finkle, and T.A.

Jennison. J. Anal. Toxicol., 4, 37 (1980).

D.J. Stewart, T. Inaba, B.K. Tang and W. Kalow. Life Sciences,

20, 1557 (1977).

P. Jatlow in S.J. Mul, ed., Cocaine: Chemical, Biological,

Clinical, Social, and Treatment Aspects, p. 61. See reference 13.

109
Chapter 8

Morphine

Archaeological and historical evidence indicates that opium has

been used as an analgesic since at least the third century B.C.

and probably as long ago as the late Bronze Age in Europe. Morphine, a pentacyclic alkaloid and the chief active component of

opium, was isolated in 1805 by a German pharmacist, although the

correct chemical structure (I) was first proposed in 1923. Total

synthesis was achieved in 1952 (1). In addition to analgesia, morphine causes side effects such as constipation, lethargy and nausea, and a great many derivatives have been synthesized in efforts

to alter or antagonize some of morphine’s pharmacological effects (2).

Heroin (3,6-diacetylmorphine, II) was introduced in 1898. It was

a more effective analgesic than morphine, but it soon became apparent that heroin was more toxic and produced physical dependence

and tolerance more rapidly than morphine. Heroin abuse increased

in the period between the World Wars (1920 to 1935) and again after

1968. By 1974 the U.S. Bureau of Narcotics estimated that there

were over 400,000 heroin addicts in the United States (1). In

spite of voluminous published data reflecting thousands of animal

and human experiments over the past 40 years, the mechanism of

heroin and morphine dependence is still not understood.

Codeine (III), a widely used methyl ether of morphine, has milder

analgesic and respiratory depressant effects, and is an effective

cough suppressant. It is demethylated in vivo and as much as 17

percent of a given dose may appear in the urine as morphine (3).

The tertiary amino group accounts for the basic properties of the

morphine molecule. Natural morphine and its salts are strongly

levorotary. The d-isomer has been synthesized but has no significant pharmacological activity (1).

110
PHARMACOKINETICS AND METABOLISM

Because of the comparative absence of cardiovascular effects, morphine has been revived as a major adjunct to anesthesia, stimulating kinetic studies in human subjects (4). The maximum pharmacological effect of morphine occurs within minutes after intravenous

injection, corresponding to the maximum measured concentration of

the free drug in plasma (5) . The clearance of morphine from blood

is rapid, with about 80 percent of an administered dose excreted

in urine within 8 hr although traces can be detected 72-100 hr

after administration, especially in addicts. The long retention

time is usually attributed to protein binding, but there is also

the likelihood of reabsorption of morphine from the intestinal

mucosa after hydrolysis of conjugates transported from the liver

in the bile (1). After intravenous injection, a plasma half-life

of 2 to 3 hr and a clearance time of 10 to 44 hr have been determined by radioimmunoassay (RIA), but the RIA technique does not

distinguish between the free and conjugated forms of the drug

(4,5). Following intravenous administration of 10-mg doses to

adult volunteers, the serum concentration of morphine ranged from

0.04 to 0.10 μg/ml after 2 hr and fell to 0.002 to 0.007 μg/ml

after 12 hr (5). Morphine is rapidly absorbed after intramuscular

or subcutaneous injection with a maximum pharmacological effect 60

to 90 min after administration, again corresponding to the highest

plasma drug concentrations (6).

Probably because of its high pKa (9.85), morphine is poorly absorbed

from the stomach. The maximum plasma concentration of free morphine

after oral administration in comparison experiments was less than

20 percent of that obtained by intravenous administration of the

same dose. Furthermore, the concentration of conjugated drug in

serum 1 hr after oral administration was 16 times higher than the

free morphine concentration, suggesting a high hepatic extraction

ratio (5). In cases of drug overdose, the minimum lethal concentration of morphine has been reported as 0.2 μg/ml in blood or 0.2-

0.4 μg/g in muscle. For addicts, the figures are higher. In one

case, an addict who had died of other causes had a blood morphine

concentration of 2.8 μg/ml (7). Urine concentrations show no correlation with lethality (7). It has been suggested, but is unsubstantiated, that in some acute deaths from heroin overdose, toxic

adulterants such as quinine may be primary agents in the lethal

reaction (8). Acute respiratory depression has also been suggested

as a likely mechanism of heroin overdose deaths, especially in

cases where the drug is present in combination with other CNS depressants such as alcohol (9).

Morphine possesses no functional groups which readily undergo hydrolysis. Heroin, however, is rapidly hydrolyzed. Less polar than

morphine, it has high lipid and membrane solubility which could

account for rapid absorption and passage through the blood-brain

barrier. The major pharmacologic effects of heroin, however, are

due to 6-monoacetyl-morphine (6-MAM) and morphine; the half-life of

heroin itself in blood is less than 20 min, and the principal product clearing the blood is 6-MAM. Its presence in plasma can be

considered presumptive evidence for heroin use (10). Because 6-MAM

111
is converted to morphine in the liver, the analysis of human urine

after heroin administration yields primarily morphine in the same

ratio of free to conjugated form as that seen after parenteral administration of morphine, i.e., about 1.9 (1). However, the presence

of 6-MAM and its 3-glucuronide in human urine has been reported

following intravenous heroin administration, in amounts representing less than 5 percent of the dose (11). In a study of heroin

kinetics in man, the urinary excretion half-life of free morphine

was found to be 1.3 hr; of 6-MAM, 1.3 hr; of conjugates, 2.8 hr;

and of total normorphine, 2.7 hr (12).

The metabolism of morphine in man is summarized in Table 1. Synthetic reactions are responsible for detoxification by converting

morphine or its metabolites, including normorphine, to readily

excreted water-soluble compounds such as 3- and 6-glucuronides and

3-sulfate, by means of microsomal enzyme systems in liver, brain,

kidney, and intestine. Quantitatively minor synthetic reactions

include methylation of normorphine and 3-0-methylation of morphine to codeine (1). The 0-methyl transferase activity which produces codeine increases in addicts, a fact which suggests altered

metabolism in prolonged opiate use and a possible marker for abuse

(13). Norcodeine, a major metabolite of codeine in man, has also

been identified in the urine of some heroin addicts, in quantities

proportional to the formation of codeine from morphine (14).

The maximal reported recovery of morphine and biotransformation

products is less than 85 percent of the dose, with excretion linearly related to urine output (15). Although small amounts of morphine

and metabolites are eliminated in feces and saliva, the metabolic

fate of 15 to 20 percent of administered drug remains unknown.

ANALYTICAL METHODOLOGY

Thin-layer chromatography (TLC) has been the most commonly employed

method for qualitative (screening) detection of morphine; it is rapid, sensitive, and economical. Technical improvements in solvent

systems, plates, sheets, and sprays have appeared regularly in the

recent literature (16,17). For example, picogram quantities of

morphine in urine or brain homogenates were detected on polyamide

plates by prior treatment of the n-butanol extracts with dansyl

chloride to fonn a highly fluorescent morphine derivative which could

be visualized on the dried plates under ultraviolet light (18).

Spectrofluorometric methods are based on the conversion of morphine

to a fluorophore, excitation and measurement of emission at specific

wavelengths. Most methods employ the oxidation of morphine to pseudomorphine by potassium ferricyanide in weakly alkaline solution,

and achieve detection levels as low as 0.1 μg/ml (19) . Fluorescence

can also be achieved without potassium ferricyanide by treating the

extract (chloroform: isopropanol, 3:1) with concentrated H2SO4, then

concentrated NH40H, and autoclaving for 15 min at 120°C. The latter

procedure can be automated for application to large numbers of samples with detection levels of 0.2 μg/ml (20). As little as 10 ng

of morphine in plasma was detected in a modified method which precipitated plasma proteins before extraction, used only siliconized

112
glassware, and decreased the reaction volume to 40 μ1 in order to

increase sensitivity. Readings were made on a filter fluorometer

(21). Because the decay of pseudomorphine fluorescence is rapid,

fluorometry has largely been limited to qualitative use. However,

coupling the photofluorimeter to a computer system, which reads

reaction kinetics within 3 milliseconds of mixing, has shown promise as a quantitative technique (22).

A quantitative spectrofluorometric method, using high performance

liquid chromatography of the oxidized urine extract with an internal standard of dihydromorphine, has been developed (23). The mixture of hydrolyzed urine and dihydromorphine was extracted with

chloroform: isopropanol (9:1) and mixed with potassium ferricyanide

immediately before injection onto a porous silica column coupled

to a fluorimeter set at ex 320 nm and em 436 nm. The solvent

system contained methanol: 2 N NH4OH:1 N NH4NO3 (30: 20:10). There

was a linear relationship between the amount of morphine in the

original solution and the ratio of the emission peak height of the

mixed dimer (pseudomorphine + pseudodihydromorphine) formed during

oxidation to that of the fluorescent dihydromorphine standard. This

allowed calibration suitable for quantitative analysis with a sensitivity of 0.01 μg/ml. The technique has the advantage of being

highly specific for morphine. Interference by other drugs, particularly normorphine, N-allyl normorphine, dihydromorphine, and 6-

monoacetyl morphine is a potential problem in other fluorometric

procedures because of emission spectra identical to pseudomorphine.

In these instances fluorometry must be combined with a chromatographic separation (19).

Immunoassays developed for morphine include the free radical assay

technique (FRAT) which employs spin labeling with nitroxide-labeled

drugs and is sensitive to 0.5 μg/ml of morphine; the enzyme multiplied immunoassay technique (EMIT) in which morphine is conjugated

to lysozyme and the combination of the labeled drug with antibodies

to morphine renders the enzyme inactive; radioimmunoassay (RIA);

and hemagglutination inhibition (HI) in which the presence of morphine in test material prevents agglutination of morphine-coated

red blood cells by an antimorphine antibody. A major advantage of

immunoassays is that biofluids can be tested directly, without sample extraction or concentration. They can also be extremely sensitive, detecting as little as 0.03 μg/ml morphine (RIA and HI), and

fast (2 to 4 min for EMIT and FRAT) (17). However, immunoassays

are subject to cross-reactivity with drugs of similar molecular

structure. Morphine antibodies have been shown to react with codeine, dextromethorphan, pethidine, dihydromorphinone, propoxyphene,

diamorphine, and 6-MAM (24,25). Attempts to reduce cross- reactions

have produced antiserums raised against various haptens: an antiserum to a conjugate of 6-succinyl morphine with bovine serum albumin reacted only slightly with morphine glucuronide and was used to

measure free morphine in serum of patients receiving oral doses,

at a detection level of 0.3 ng/ml (24). Other haptens tried have

included morphine-3-glucuronide and oxymorphone-6-carboxy-methoxime

(26). Haptens are usually attached to the carrier at one of the

carbon atoms of morphine, but an antiserum capable of distinguishing morphine from codeine was obtained by conjugating carrier pro113
tein to the nitrogen atom of nomorphine. The antibody produced

did not react with codeine or with the 3-0-monoglucuronide of morphine, and detected 15 pg of morphine in the presence of codeine

(27). An antiserum specific for codeine has also been reported,

prepared from a conjugate using N-butyroylnorcodeine as hapten (28).

While immunoassays, especially RIA, can be adapted to quantitative

studies by preparation of calibration graphs, the lack of absolute

specificity remains a problem. Nevertheless, a solid-phase RIA,

which involves the absorption of antibody to polymeric surfaces,

has been adapted for morphine assay using an antibody to morphine6-hemisuccinate purified by affinity chromatography. The assay is

reported to respond to as little as 0.5 ng of morphine per ml of

serum (29).

Gas chromatography (GC) can be used for screening biofluids, but

because of the relatively long time required to complete a GC analysis, it is more often used for confirmation of TLC or immunoassay

results, and for quantitative studies in pharmacokinetics and metabolism. GC systems equipped with flame ionization detectors (GCFID) are sensitive enough to detect 15 ng of morphine/ml. Conversion of the morphine extracted from bile, urine, blood, or liver to

the less polar diacetyl derivative (heroin) can increase sensitivity

of GC-FID to 1 ng (17). The polarity of morphine, which causes

adsorption in the GC column, can also be altered by silylation. A

derivatization procedure using bis-(trimethylsilyl)trifluoroacetamide (BSTFA) plus 1 percent trimethylchlorosilane as both solvent

and silylating agent is reported to give 100 percent conversion of

morphine to its bis-(trimethylsilyl) ether within 20 min at room

temperature (30). Bis-(0-trimethylsilyl) morphine can also be formed

quantitatively and reproducibly by simultaneous injection of trimethylsilylimidazole and the drug solution onto the gas chromatographic

column (31).

A quantitative GC method developed for heroin and 6-monoacetyl morphine in plasma utilized an alkali-flame detector (nitrogen/phosphorus detector). Plasma samples at pH 9.0 containing ethylmorphine as internal standard were extracted with benzene, and the

dried extracts were chromatographed as their acetylated derivatives

after treatment with trifluoroacetic anhydride-benzene (1:5). The

alkali-flame detector permitted quantitation of drug concentrations

as low as 100 ng/ml, with detection of 20 ng/ml. Extraction was

done quickly after venipuncture to avoid in vitro enzymatic hydrolysis of the compounds (32).

Electron-capture detection (GC-ECD) has been used to quantitate

morphine after conversion to halogen-containing derivatives (33-37).

In one procedure nalorphine was added as an internal standard, and

the morphine and nalorphine were trifluoroacetylated following extraction from 1-2 ml of serum or plasma with 10 percent isobutanol in

chloroform, back extraction with 0.5 N HCl, and reextraction with

10 percent isopropanol in ethyl acetate. With a 63Ni detector a

sensitivity limit of 25 ng/ml in serum was reported (33). A refinement of the ECD technique, using a 3H-ECD, detected 0.5 ng/ml morphine in plasma extracts and 0.1 ng morphine in 30 mg of brain tissue from rats. The samples were extracted with toluene:butanol

114
(9:1) at pH 8.9, back extracted into 0.1 N H2SO4

, and reextracted

with toluene:butanol. Morphine and the internal standard (nalorphine) were derivatized with pentafluoropropionic anhydride (34).

Both morphine and codeine have been quantitated by derivatization

with pentafluoropropionic anhydride and analysis by GC-ECD using

a 

63Ni detector. Detection limits were 2 pg for morphine and 20 pg

for codeine in test solutions (35).

A GC-ECD method which can assay as little as 0.1 ng/ml of morphine

in human urine has been reported (36). Morphine was converted to

its heptafluorobutyryl derivative for GC separation on 3 percent

OV-17 after extraction with ethyl acetate of 2 ml of alkalinized

urine. Codeine was used as an internal standard. The sensitivity

limit reported for this procedure was 100 pg morphine/ml of urine.

Fluorinated derivatives were also prepared for GC-ECD quantitation

studies of morphine, codeine, and 6-MAM present as contaminants in

illicit heroin (37). Heptafluorobutyric anhydride (HFBA) yielded

suitable derivatives within a 5-min reaction time, and the derivatives were extracted from an acetonitrile-sodium bicarbonate solution into petroleum ether before chromatography on 3 percent OV-17

on Gas Chrom Q. Morphine, codeine, and 6-MAM in heroin were readily

measured at levels as low as 0.001, 0.01, 0.01 percent, respectively.

The minimum detectable quantity for each compound was ~20 pg for

morphine, 80 pg for codeine, and 100 pg for 6-MAM. However, when

procaine was present as an adulterant, it was highly reactive with

HFBA and caused significant interference with the chromatograms

of the heptafluorobutyryl derivatives.

GC coupled with mass spectrometry (GC/MS) offers the best method

available for the unequivocal identification of morphine and its

metabolites in biological materials. It can serve as a check on

results of less sensitive and specific assays, as well as being

a tool for positive confirmation of structures. The technique of

selected ion monitoring has been applied to quantitation of morphine and related compounds in a variety of body fluids (10,38-43).

Morphine was measured in urine at concentrations as low as 5 ng/ml

using GC/MS with chemical ionization (42). Morphine labeled with

three deuterium atoms on the N-methyl group served as the internal

standard. After adjustment of the urine to pH 8.5 it was extracted

with chloroform: isopropanol (4:1) and the extract was treated with

bis-(trimethylsilyl) acetamide. The resulting bis-(trimethylsilyl)

morphine was separated and measured by chromatography on a 3 percent OV-17 column maintained at 230°C; monitored ions corresponded

to two prominent fragment ions [(M-CH3)

+

 and MH+-HOSi(CH3)3] in

the methane chemical ionization mass spectra of the derivatized

drug and the deuterated internal standard.

Morphine and codeine were measured simultaneously in blood by a

GC/MS procedure involving addition of a deuterated internal standard to the blood, acid hydrolysis of conjugates, a series of extractions to separate the bases from other components of the extract, derivatization with trifluoroacetic anhydride, and measurement of the bis-(trifluoroacetyl) morphine and mono-(trifluoroacetyl) codeine using gas chromatography (3 percent OV-17, 230°C)

and electron impact mass spectrometry (43). The assay was applied

115
to the analysis of 916 blood samples from individuals arrested for

suspicion of opiate intoxication. The average concentration of

morphine was 0.29 μg/ml. For most samples the codeine concentration was approximately 5 to 10 percent of the morphine concentration. The lower limit for acceptable quantitative measurement was

considered to be 20 ng/ml.

In an application of GC/MS to a study of the relationship between

brain levels of morphine and analgesia in rats, the internal standard was again trideuterated morphine and the drug was chromatographed as its trifluoroacetyl derivative (40). Sensitivity was

such that the low concentrations of morphine present in brain following therapeutic doses could be measured easily, with a lower

quantitation limit of 8 ng/g of tissue.

Free and bound morphine, as well as free 6-MAM have been quantitated in rabbit blood by GC/MS (10). Trideuterated morphine and

6-MAM-2H3 were added to 0.2 ml plasma before precipitation of proteins by acetone. The supernatant was evaporated to dryness under

nitrogen and the residue taken up in 2.5 ml of 0.1 N HCl and purified by two extractions with benzene. The aqueous phase was adjusted to pH 8.5, extracted with chloroform:isoamyl alcohol (3:1),

and evaporated to dryness before derivatization with trifluoroacetic anhydride. The derivatives were taken up in chloroform for GC/

MS analysis on a 6 ft x 2 mm silanized column packed with 1 percent

OV-1 on Gas Chrom Q, using helium as carrier gas and a column temperature of 230°C. With electron-impact ionization the highest ion

intensities were associated with fragments at m/z 364 for the protio compound and 367 for the deuterio compound. As an assay for

free and bound heroin metabolites, the method represented a 100-

fold improvement over GC-FID methods for these compounds, with a

sensitivity of 1 ng/ml.

Extractive alkylation with pentafluorobenzyl (PFB) bromide, using

tetrabutylammonium (TBA) as counter ion and ethyl acetate as solvent, has been used to quantitate morphine in plasma at concentrations as low as 5 ng/ml (41). With this procedure morphine was extracted from aqueous solutions with 98 percent efficiency. Alkylation of the phenolic hydroxyl group occurred simultaneously. The

PFB derivative of morphine could then be analyzed directly by GC/

MS. However, better results were obtained if the PFB morphine was

further derivatized by treatment with trifluoroacetic anhydride.

The resulting PFB-TFA derivative was chromatographed on a 2 percent

OV-17 column maintained at 245°C. Quantitation was achieved by

measuring the ion currents at m/z 380 and 383, corresponding to prominent fragment ions in the electron impact mass spectra of the PFBTFA-morphine and its trideuterated analog.

Morphine concentrations determined by spectrofluorometry in blood and

brain were approximately one-half those obtained by GC/MS (44). The

difference may result from quenching interferences in the fluorometric assay and the greater sensitivity of the GC/MS method. Hydrolysis of blood samples increased the apparent concentration of morphine

by about 260 percent, while the corresponding increase in brain

samples was only about 25 percent. Presumably, the increase is

caused by hydrolysis of the glucuronide conjugate of morphine.

116
Charge-exchange ionization using 10 percent nitric oxide in nitrogen as reagent gas can offer an advantage over CI or EI ionization

for the analysis of morphine and heroin in that the charge-exchange

mass spectra of these compounds show less fragmentation (45). Field

ionization also causes little or no fragmentation of morphine-type

molecules and can be used to determine trace amounts of drugs in

biological media by isotope dilution analysis with multilabeled

diluents such as morphine-2H5 and codeine-2H8 (46,47).

Quantitation of total morphine in biological fluids requires hydrolysis to free the drug from conjugates. Enzymatic hydrolysis

has generally proven to be less efficient than acid hydrolysis of

morphine glucuronides because of variations in potency among enzyme

preparations. The most efficient enzymatic recovery found in a

comparative study was 64 percent. Critical parameters included

choice of buffer system, pH, temperature, and enzyme concentration.

High-temperature (autoclaving or refluxing) acid hydrolysis does

not require such critical control of conditions. The same study

reported a yield of 93 percent of available morphine when the conjugate solution was autoclaved at 125°C for 30 min using 8-15 percent hydrochloric acid (48). However, hydrolysis conditions vary

widely in the literature, and some authors have reported complete

hydrolysis of morphine-3-glucuronide in 30 min at 100°C by 4 N

HCl (10).

Extraction procedures for morphine also vary, but because of its

phenolic hydroxyl and tertiary amine the optimum pH for extraction

is 8.5-10 (49). Chloroform is most often used as the organic solvent, usually with the addition of isopropanol (7,15,23), n-butanol

(19,21), or isoamyl alcohol (10) in a ratio of either 9:1 or 3:1.

Recoveries of 60-70 percent have been reported for chloroform:

isopropanol (3:1) (7). However, toluene:butanol (9:1) (34), ethyl

acetate:isobutanol (9:1) (50), ethyl acetate + 10 percent isopropanol (40), benzene (32), ether (51), and 1,2-dichloroethane + 30

percent isopropanol (12) are among other solvents reported. A

morphine recovery of 85-100 percent from urine has been claimed

for a system involving adsorption on a XAD-2 resin column and elution with chloroform: isopropanol (17).

A study (52) of methods of recovering morphine from biological samples prior to analysis resulted in the following conclusions: (1)

acid hydrolysis of morphine conjugates is preferable to enzymatic

hydrolysis unless other acid-labile drugs are to be analyzed, (2)

chloroform:isopropyl alcohol (4:1) is the extraction solvent of

choice because it removes most of the morphine from the biological

sample in a single extraction and it is readily volatile, (3) sodium

bicarbonate and ammonium carbonate are the preferred “salting out”

agents, and (4) the ratio of biological sample to solvent should be

about 1:4.

A review of the analysis of morphine by gas phase methods, including

procedures for extraction from urine and blood for various biomedical applications and quantitation, was published in 1977 (53).

117
EXPERIMENTAL PROCEDURE

The experimental procedure involves a single extraction of a buffered and salt-saturated solution of the biological fluid with a

solvent system consisting of toluene:heptane: isoamyl alcohol (70:

20:10 v/v). ‘The extract is then concentrated and derivatized by

treatment with trifluoroacetic anhydride. Quantitation is achieved

by selected ion monitoring using chemical ionization with methane

and ammonia as reagent gases.

Standards and Reagents

The morphine used in the development of this procedure was purchased

from Applied Science Laboratories, State College, PA 16801. No

impurities were detected by either gas chromatography or mass spectrometry. Morphine labeled with 3 deuterium atoms on the N-methyl

group (morphine-2H3) was prepared by a published procedure (54,55).

Mass spectral analysis of the morphine-2H3 showed that 94 percent

of the molecules contained 3 deuterium atoms, while 0.5 percent

contained no deuterium atoms. Gas chromatography indicated a chemical purity of greater than 99 percent.

Prepare a pH 9.6 buffer by dissolving 500 grams of potassium monohydrogen phosphate (K2HPO4

) in about 500 ml of distilled water. Allow the solution to cool to room temperature and then add sufficient

distilled water to make exactly one liter.

Stock solutions of morphine and morphine-2H3 used to determine calibration graphs and prepare working standards are prepared as follows.

Weigh into a 100-ml volumetric flask about 10 mg of morphine and

record its weight to the nearest 0.1 mg. Dissolve the measured

morphine in about 75 ml of methanol and then add additional methanol

to make exactly 100 ml of solution. In the following paragraphs

this solution will be referred to as the “0.10-mg/ml stock solution”

although it should be labeled with its actual concentration based

on the exact measured weight of morphine. A series of standard solutions can be prepared by appropriate dilution of this stock solution

as described in Chapter 2.

The stock solution of morphine-2H3 is prepared in the same manner.

For the measurement of morphine in body fluids within the concentration range 1 to 500 ng/ml, addition of approximately 40 ng of the

internal standard to each ml of specimen is satisfactory. This can

be conveniently done by adding 100 μl of a 400-ng/ml morphine-2H3

methanolic solution to each ml of specimen. Prepare the 400-ng/ml

internal standard solution by diluting 1 ml of the 0.10-mg/ml stock

solution of morphine-2H3 to 250 ml with methanol.

Store stock and standard solutions in well-stoppered or capped glass

vessels in the dark at less than 0°C.

118

328-011 0 - 80 - 7
Extraction

Transfer 1 ml of the specimen (plasma, serum, whole blood, or urine)

into a 5-ml glass stoppered, conical centrifuge tube. Add 100 μl

of the 400-ng/ml morphine-2H3 internal standard solution and vortex for about 10 sec. Allow the sample to equilibrate for 15 min,

and then add about 0.5 g of solid sodium chloride followed by 1 ml

of the K2HPO4 buffer (pH 9.6). Vortex the mixture for about 10 sec.

Add 1 ml of a solvent system consisting of toluene:heptane:isoamyl

alcohol in a volume ratio of 70:20:10. Stopper the tube and vortex mix for at least 30 sec. Separate the phases by centrifugation

for 10 min. Transfer most of the organic layer (top) to a 12-ml

glass stoppered, conical centrifuge tube. Be careful not to transfer any of the aqueous (bottom) layer. Evaporate the organic extract just to dryness by heating to 70°C under a stream of filtered

air. Stopper the tube and store under refrigeration until shortly

before the GC/MS analysis is ready to be performed. For derivatization, allow the tube to warm to room temperature and then add

200 μl of chloroform plus 100 μl of trifluoroacetic anhydride.

Cap the tube and heat at 70°C for 10 min. Allow the tube to cool

at room temperature for 10 min and then evaporate to dryness by

heating at 40°C under a gentle stream of dry nitrogen or dry air.

The derivatized extract is subject to hydrolytic decomposition upon

prolonged exposure to laboratory air, so it is important to reconstitute the residue immediately in about 30 μl of chloroform and

make the injection into the GC/MS.

GC/MS Analysis

Recommended experimental conditions for the GC/MS analysis of the

trifluoroacetyl derivative of morphine are as follows:

GC column 1.8 m x 2 nun (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier gas: Methane, 15-20 ml/min

Reagent gas: Ammonia, introduced into the ion chamber as

described. in Chapter 2.

Temperatures: Injector, 270°C

Column, 225°C isothermal

GC/MS transfer iine, 260oC

Ion source, 160°C

Under these conditions the bis-(trifluoroacetyl) morphine should

elute at between 2 and 4 min.

Before beginning the GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored occur at m/z 478 and 481.

They correspond to the protonated molecule ions for bis-(trifluoro119

328-011 0 - 80 - 9
acetyl) morphine and the 2H3

-analog, respectively. With the divert valve in the divert position, inject 2 to 6 μl of the chloroform solution of the extract into the gas chromatograph. After

approximately 1 min, switch the divert valve so that the entire

carrier gas flow enters the ion source of the mass spectrometer

and begin data acquisition. When the morphine derivative has eluted, cease data acquisition and return the divert valve to the divert position.

The quantity of free morphine in the biological sample is determined by measuring the heights (or areas) of the bis-(trifluoroacetyl) morphine peak and the 2H3

-analog peak in the ion current

profiles (“ion chromatograms”) and relating the ratio of peak

heights to a calibration graph. Calculate the specimen concentration

of free morphine by dividing the measured quantity of morphine by

the exact volume of specimen extracted.

Discussion of the Experimental Procedure

The procedure described here is relatively rapid, yet it results

in excellent sensitivity and specificity. Morphine concentrations

as low as 1 ng/ml can be measured accurately and interference from

other components of the extract is seldom a problem. If interferences are encountered it may be possible to remove them by incorporating a back-extraction into the method. When a back-extraction

is desirable, follow the previously described procedure to the

point where the toluene:heptane:isoamyl alcohol extract (approximately 1 ml) is transferred into a 12-ml glass stoppered, conical

centrifuge tube. Then add 1 ml of 0.2 N H2SO4. Stopper the tube

and shake or vortex for at least 30 sec. Discard the organic layer

(top) and neutralize the aqueous layer (bottom) by addition of 1 ml

of 0.2 N NaOH. Saturate the solution with sodium chloride (~1 g)

and add 2.0 ml of the K2HPO4 buffer. Confirm that the pH of the

solution is between 8.5 and 9.0, and then extract the morphine with

2 ml of the toluene:heptane:isoamyl alcohol solvent. Finally, proceed with the derivatization and GC/MS analysis as previously described.

Preparation of the bis- (trimethylsilyl) derivative of morphine by

heating the extract with bis-(trimethylsilyl) trifluorocetamide

(BSTFA) at 60°C for 1 hr, is an alternative to formation of the

trifluoroacetyl derivative. Both derivatives have been used with

satisfactory results. Trifluoroacetylation occurs somewhat more

rapidly than the trimethylsilyation. However, the excess trifluoroacetic anhydride should be removed by evaporation prior to GC/MS

analysis to avoid degradation of the GC column. (Injection of excess

BSTFA onto the GC column does not appear to degrade column performance). Figures 1 and 2 show the methane-ammonia CI mass spectra

of bis-(trifluoroacetyl) morphine and bis-(trimethylsilyl) morphine,

respectively. In both spectra the only abundant ions correspond to

the protonated molecule ions. A direct comparison has not been made

of the stability of these two morphine derivatives, but no major

differences have become evident so far.

120
FIGURE 1. METHANE-AMMONIA CI MASS SPECTRUM OF BIS-(TRIFLUOROACETYL) MORPHINE

FIGURE 2. METHANE-AMMONIA CI MASS SPECTRUM OF BIS-(TRIMETHYLSILYL) MORPHINE

121
The procedure described here measures only free morphine in the

body fluid. In urine, most of the excreted morphine is present

as glucuronide and sulfate conjugates. Consequently, in order to

measure the total morphine it is necessary to hydrolyze the conjugated morphine before extraction. Acid hydrolysis can be performed

as follows (43):

Transfer 1 ml of specimen into a 12-ml culture tube. Add 100 μl

of the 400-ng/ml morphine-2H3 internal standard solution and vortex for about 10 sec. Allow the sample to equilibrate for 15 min.

Add 1 ml of 6 N HC1 (prepared by a 1 to 1 dilution of concentrated

hydrochloric acid with distilled water). Vortex the mixture and

then heat in a boiling water bath for 1 hr. After cooling to room

temperature neutralize the mixture by slowly adding 1 ml of 6 N

NaOH. Saturate the solution with sodium chloride to facilitate

extraction of the free morphine. Add 2 ml of the K2HPO4 buffer

and check to see that the pH of the mixture is between 8.5 and 9.0.

Extract the morphine with 1 ml of the toluene:heptane:isoamyl alcohol solvent, and proceed with the derivatization and GC/MS analysis as previously described.

Other opiate narcotics such as codeine and 6-monoacetyl morphine

can be measured simultaneously by the described method if the

corresponding deuterium-labeled analogs are available for use as

internal standards. The protonated molecule ions for trifluoroacetyl codeine and its 2H3

-analog occur at m/z 396 and 399, respectively; in the case of the trifluoroacetyl derivative of 6-monoacetyl morphine and its 2H3

- analog, these ions occur at m/z 4.24

and 427. Figures 3 and 4 show representative calibration graphs for

morphine and codeine in plasma. The graphs are linear for drug

concentrations from 1 to 500 ng/ml. If higher concentrations are

encountered or anticipated, the specimen should be diluted with distilled water in order to reduce the drug concentration so that it

falls within the linear range of the calibration graph.

122
FIGURE 3. CALIBRATION GRAPH FOR ANALYSIS OF MORPHINE

IN PLASMA

FIGURE 4. CALIBRATION GRAPH FOR ANALYSIS OF CODEINE

IN PLASMA

123
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR MORPHINE

Nomenclature

Chemical Name: 7,8-Dihydro-4,5-epoxy-17-methyl-morphinan-3,6-

diol

Empirical Formula: C17H19NO3

Chemical Abstracts Registry Number: 57-27-2

Physical Constants

Melting Points: 254°C with decomposition

Specific Rotation: = -132° (methanol)

Solubility: 1g of morphine dissolves in 5000 ml of water at

room temperature, 1100 ml of boiling water, 210 ml

of ethanol, 1220 ml of chloroform, 6250 ml of diethylether, or 10 ml of boiling methanol.

pKa: 9.85

UV Absorption: max 287 nm (ethanol)

REFERENCES

1. U. Boerner, S. Abbott, and R.L. Roe. Drug Metabolism Reviews,

4, 39 (1975).

2. A.F. Casy in D. Clouet, ed., Narcotic Drugs: Biochemical Pharmacology. Plenum, New York, ch. 1 (1971).

3. G.J. Mannering, A.C. Dixon, E.M. Baker III, and T. Asami. J.

Pharmacol. Exp. Ther., 111, 142 (1954).

4. P.A. Mitenko and P.C. Duke. Clin. Research, 22, 726A (1974).

5. S. Spector and L.S. Vesell. Science, 174, 421 (1971).

6. S.F. Brunk and M. Delle. Clin. Research, 21, 467 (1973).

7. S. Felby, H. Christensen, and A. Lund. Forensic Sci., 3, 77

(1974).

8. D.C. Perry. Clin. Toxicol, 8, 239 (1975).

9. J.R. Monforte. J. Forensic Sci., 22, 718 (1977).

10. W.O.R. Ebbighausen, J.H. Mowat, H. Stearns, and P. Vestergaard. Biomed. Mass Spectr., 1, 305 (1974).

124
11. S.Y. Yeh, R.L. McQuinn, and C.W. Gorodetzky. J. Pharm. Sci.,

66, 201 (1977).

12. S.Y. Yeh, C.W. Gorodetzky and R.L. McQuinn. J. Pharmacol.

Exp. Ther., 196, 249 (1976).

13. U. Boerner and S. Abbott. Experientia, 29, 180 (1973).

14. U. Boerner, K.L. Roe, and C.E. Becker. J. Pharm. Pharmacol.,

26, 393 (1974).

15. S.Y. Yeh. J. Pharmacol. Exp. Ther., 192, 201 (1975).

16. J.F. Taylor in D.Clouet, ed., Narcotic Drugs: Biochemical

Pharmacology. Plenum, New York, ch. 2 (1971).

17. S.J. Mulé. J. Chromatogr. Sci., 12, 245 (1974).

18. H.H. Loh, I.K. Ho, T.M. Cho, and W. Lipscomb. J. Chromatogr.,

76, 505 (1973).

19. H. Kupferberg, A. Burkhalter, and E.L. Way. J. Pharmacol.

Exp. Ther., 145, 247 (1964).

20. S.J. Mulé and P.L. Hushin. Anal. Chem., 43, 708 (1971).

21. A.E. Takemori. Biochem. Pharmacol., 11, 1627 (1968).

22. J.M. Hicks, R.A. Sonoff, and K.J. Koch. Clin. Chem., 20,

901 (1974).

23. I. Jane and J.F. Taylor. J. Chromatogr., 109, 37 (1975).

24. G.W. Aherne, V. Marks, B.A. Morris, E.M. Piall, J.D. Robinson,

and R.G. Twycross. Brit. J. Pharmacol., 54, 228P (1975).

25. D. Sohn, J. Simon, M.A. Hanna, G.V. Ghali, R.A. Tolba, and V.

Melkonian. Anal. Chem., 45, 1498 (1973).

26. M. Koida, M. Takahashi, and H. Kaneto. Japan. J. Pharmacol.,

24, 707 (1974).

27. G. Gintzler, E. Mohacsi, and S. Spector. Europ. J. Pharmacol.,

38, 149 (1976).

28. J.W.A. Findlay, R.F. Butz, and K.M. Welch. Life Sci., 19,

389 (1976).

29. M. Steiner and J.L. Spratt. Clin. Chem., 24, 339 (1978).

30. G.R. Wilkinson and E.L. Way. Biochem. Pharmacol., 18, 1435

(1969).

31. K.E. Rasmussen. J. Chromatogr., 120, 491 (1976).

125
32. D.A. Smith and W.J. Cole. J. Chromatogr., 105, 377 (1975).

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46 .

47.

48.

49.

50.

51.

52.

J.E. Wallace, H.E. Hamilton, K. Blum, and C. Petty. Anal.

Chem., 46, 2107 (1974).

B. Dahlström and L. Paalzow. J. Pharm. Pharmacol., 27, 172

(1975).

M.J. Kogan and M.A. Chedekel. J. Pharm. Pharmacol., 28, 261

(1976).

G. Nicolau, G. Van Lear, B. Kaul, and B. Davidow. Clin. Chem.,

23, 1640 (1977).

J.M. Moore. J. Chromatogr., 147, 327 (1978).

W.O.R. Ebbighausen, J.H. Mowat, and P. Vestergaard. J. Pharm.

Sci., 62, 146 (1973).

W.O.R. Ebbighausen, J.H. Mowat, P. Vestergaard, and N.S. Kline.

Adv. in Biochem. Psychopharmacol., 7, 135 (1973).

P.P. Hipps, M.R. Eveland, E.R. Meyer, W.R. Sherman, and T.J.

Cicero. J. Pharmacol. Exp. Ther., 196, 642 (1976).

W.J. Cole, J. Parkhouse and Y.Y. Yousef. J. Chromatogr. 136,

409 (1977).

P.A. Clarke and R.L. Foltz. Clin. Chem., 20, 465 (1974).

D. Pearce, S. Wiersema, M. Kuo, and C. Emery. Clin. Toxicol.,

14, 161 (1979).

D. Reed. Clin. Toxicol., 14, 169 (1979).

I. Jardine and C. Fenselau. J. Forensic Sci., 20, 373 (1975).

M. .Anbar and W.H. Aberth. Anal. Chem., 46, 59A (1974).

J.A. Lawson, J.I. Degraw, and M. Anbar. J. Heterocyclic Chem.,

13, 593 (1976).

F. Fish and T.S. Hayes. J. Forensic Sci., 19, 676 (1974).

T.N. Higgins and J.D. Taylor. Clin. Biochem., 7, 280 (1974).

G.R. Nakamura and E.L. Way. Anal. Chem., 47, 775 (1975).

F. Fish and W.D.C. Wilson. J. Chromatogr., 40, 164 (1969).

D.B. Predmore, G.D. Christian, and T.A. Loomis. J. Forensic

Sci., 23, 481 (1976).

126
53. J.P.G. Tehnot and K.D. Haegele in D. Glick, ed., Methods of

Biochemical Analysis, Vol. 24. Interscience, John Wiley and

Sons, New York, p. 101 (1977).

54. C. Elison, H.W. Elliott, M. Look, and H. Rapoport. J. Med.

Chem., 6, 237 (1963).

55. H. Rapoport and M. Look. U.S. Patent 2,890,221; Chem. Abstr.,

54, 612f (1960).

127
Chapter 9

Diazepam and its Major Metabolite,

N-Desmethyldiazepam

Diazepam (7-chloro-1,3-dihydro-1-methyl-5-phenyl-2H-1,4-benzodiazepin-2-one, I) was first synthesized in 1959 and introduced clinically in the early 1960’s as an antianxiety agent and a muscle relaxant (1). It can be synthesized by the reaction of 2-methylamino-5-chlorobenzophenone and ethyl glycinate, or by a variety of

other routes (2) . Diazepam has been marketed since 1963 as Valium

and has become the most commonly used prescription drug in the

Western world (1).

The most important clinical effects of the drug are mediated through

the central nervous system (CNS); the limbic system of the brain,

which controls emotions, has been implicated in attempts to locate the site of antianxiety activity. Diazepam has muscle relaxant and strong anticonvulsant properties also, but the sites of

action for these activities have not been identified. Unlike

barbiturates, drugs of the benzodiazepine class do not appear to

produce clinically important hepatic microsomal induction and are

more benign than other sedative-hypnotic drugs with respect to

cardiovascular and respiratory depression (1). Tolerance does develop, however, so that larger doses become necessary to achieve

the desired effects. Recent clinical studies have indicated that

sensitivity to the CNS-depressant effects of diazepam increases

with age (3). Side effects of therapeutic doses (usually 20 mg

daily or less) can include drowsiness, ataxia, disorientation, and

vertigo (4). There is evidence for an association between the ingestion of diazepam in the first trimester of pregnancy and the

incidence of oral clefts in infants (5). However, because of its

relatively wide margin of safety, death due to diazepam overdose

is rare. Significant impairment of psychomotor skills at doses of

10-20 mg was demonstrated in a study related to driving ability,

and this impairment combined with the sleep-inducing effect of the

drug makes its possible contribution to traffic accidents an impor128
tant consideration (6). Diazepam potentiates the depressant effect

of methadone, adding to problems of addict management. Abuse of

diazepam (40-80 mg daily), leading to severe withdrawal symptoms,

has been reported in drug treatment programs (7). These dangers

associated with the misuse of diazepam caused the Food and Drug

Administration to reclassify it as a Schedule IV drug.

METABOLISM AND PHARMACOKINETICS

In man, diazepam undergoes two major metabolic transformations:

N-demethylation and 3-hydroxylation. In the first of these routes,

diazepam is metabolized by liver microsomal enzymes to N-desmethyldiazepam (II) which is then hydroxylated to form oxazepam (III).

The diazepam hydroxylation pathway yields IV, which is subsequently

demethylated to III (3). The major metabolite present in blood is

II, although III and IV are also found, and with chronic administration II will equal or exceed diazepam blood concentrations.

The contribution of N-desmethyldiazepam (II) to the clinical effects

of diazepam in man is not clear (3,8), although the pharmacological

properties of the metabolite are generally considered to be similar

to its precursor; and the anticonvulsant effect, in fact, seems to

require dealkylation (9). Oxazepam exerts anticonvulsant activity

in mice comparable to that of N-desmethyldiazepam. In man, 60 to

75 percent of a dose of diazepam administered orally is excreted

in the urine. The most abundant metabolite in human urine is the

oxazepam-glucuronide, but N-desmethyldiazepam, IV-glucuronide, and

p-hydroxy-diazepam (V) are also found (10). Intact diazepam has

not been reported in more than trace amounts in urine (11).

The increase in plasma concentrations of N-desmethyldiazepam relative to the parent drug following repeated doses of diazepam has

suggested that diazepam might induce its own metabolism (12). However, there is evidence for an alternative explanation; i.e., the

tissues may be saturated with the metabolite following the first

pass hepatic metabolism so that repeated doses result in elevated

plasma concentrations (13). N-desmethyldiazepam has also been reported in human plasma as a metabolite of chlorodiazepoxide (Librium) (14).

Diazepam is highly bound (> 97 percent) to plasma proteins (15)

and is metabolized completely but slowly in man, with a terminal

elimination half-life usually reported to be between 20 and 50 hr,

although it can range up to 90 hr in normal individuals (1,8,16).

The half-life is age-dependent; i.e., up to 5 times longer in elderly persons than in young adults (16). The half-life of desmethyldiazepam is longer, about 50-100 hr (8,17), and it too is

about 97 percent protein-bound. Diazepam is lipid soluble but

only sparingly soluble in water at physiological pH, so that intramuscular absorption is slow and erratic, and oral or intravenous

routes of administration are preferred (15). Because of its long

half-life, diazepam will accumulate in plasma. For example, if

administered three times per day, the concentration may increase

3-fold over that following a single dose. Desmethyldiazepam accumulates more slowly, usually appearing in plasma 24-36 hr after

129
the first dose, and both the parent drug and the metabolite reach

steady state after 5-7 days (18,19). Both diazepam and N-desmethyldiazepam can be present in pharmacologically active concentrations

a week after medication is stopped (19). However, wide variations

in pharmacokinetic profiles are seen between individuals, and no

simple correlation exists between clinical response and plasma levels of diazepam and N-desmethyldiazepam. Nevertheless, a minimum steady state plasma concentration of diazepam of 400 ng/ml

has been reported to be necessary for relief of acute anxiety (20).

During chronic therapy in patients given doses of 4-60 mg diazepam

per day, plasma concentrations are reported to range from 20-200

ng/ml of diazepam and 32-2750 ng/ml of N-desmethyldiazepam. In

the same study, absorption of diazepam and its metabolite by erythrocytes was observed after 11 weeks of administration. Uncon130
jugated forms of diazepam and metabolites appeared in urine following several weeks of medication (21).

After intravenous injection of 20 mg of diazepam, a maximum concentration of 1600 ng/ml was reached in 15 min, while 20 mg taken

orally gave a maximum blood concentration of 490 ng/ml in 30 min

and the maximum after intramuscular administration was 290 ng/ml

in 60 min (22). In other investigations, however, oral administration of 10 mg diazepam resulted in peak plasma concentrations

of 100-200 ng/ml after 1.5 hr (23). ‘The drug was labeled with

tritium for this study, and the decline of radioactivity in the

plasma indicated a rapid and extensive uptake by tissues followed

by a redistribution into blood of both diazepam and N-desmthyldiazepam.

The rate at which diazepam returns to the central compartment from

the deep peripheral compartment has been shown to be the ratecontrolling factor in the elimination of diazepam from the body

and in formation of dealkylated metabolites; only 18 percent of

the drug in the body is present in the central compartment and

available for elimination at any given time (17). In vitro displacement of diazepam from human serum albumin by lauric acid demonstrated that protein binding, as well as tissue distribution, could

be affected by fluctuations in plasma free fatty acids (24). Because human bile contains virtually no diazepam, N-desmethyldiazeam, or their glucuronide conjugates, as shown by a study using

14C-diazepam their long half-lives in plasma cannot be explained

by enterohepatic circulation (10). The presence of liver disease,

however, has been found to decrease the protein binding of diazepam

and to prolong its plasma elimination half-life (16). Chronic

ingestion of alcohol. (25), as well as food intake (26), can also

affect the pharmacokinetics of intravenously administered diazepam.

The entry of free, unbound diazepam and N-desmethyldiazepam into

the cerebrospinal fluid (CSF) appears to be by passive diffusion,

quickly reaching concentrations equivalent to 2-4 percent of the

total plasma concentration (11). After long-term diazepam therapy,

however, some accumulation of the metabolite in the CSF relative to

plasma has been observed which could reflect high concentrations

of the metabolite in brain tissue following repeated administration

of diazepam (9). Desmethyldiazepam is known to accumulate in fetal

brain, heart, liver, and lung tissues when the mother has taken

diazepam (27).

The concentration of diazepam in saliva has been shown to be directly proportional to the concentration of unbound drug in plasma

(saliva/plasma ratio 0.03). This finding is significant in that

it raises the possibility of obtaining a sample by a noninvasive

method for the determination of plasma concentrations (28).

An extensive review of the clinical pharmacokinetics of diazepam

and N-desmethyldiazepam has been published (29).

131
ANALYTICAL METHODOLOGY

A quantitative ultraviolet spectrophotometric assay capable of measuring blood concentrations of diazepam and N-desmethyldiazepam

resulting from doses greater than 50 mg was reported in 1966.

Quantitation was based on measurement of absorbance values at 240

nm. The sensitivity limit of 0.3 μg/ml was sufficient for toxicological purposes (18). A fluorometric screening method capable

of detecting benzodiazepines in blood and urine at concentrations

of 25 ng/ml has been reported more recently, but it does not distinguish between diazepam and its metabolites. The benzodiazepines

present are hydrolyzed to benzophenones and then converted to fluorescent 9-acridanones by heating with lead dioxide (30).

For human metabolic studies using 14C-labeled diazepam, thin-layer

chromatography (TLC) of extracts, using chloroform: acetone (9:1) as

developing solvent, followed by scintillation counting of the appropriate sections of gel, has been used for measurement of diazepam and its metabolites in blood, urine, and bile (10). Recently,

another TLC procedure was developed for use in bioavailability

studies. Carbamazepine was added as an internal standard to buffered serum (pH 8) before extraction with n-hexane containing 10

percent isobutanol. The dried residue was dissolved in chloroform

and spotted on the TLC plate; the developing solvent was ethyl acetate:benzene:ammonia (35:15:0.15). After drying, the plates were

exposed to HCl gas for 30 min and then to radiation at 254 nm. Diazepam, N-desmethyldiazepam, and the internal standard were converted to fluorescent compounds by this procedure, and fluorescence

was enhanced by dipping the plates in a 10 percent solution of paraffin wax in light petroleum. Quantitative fluorimetry ( ex 360 nm,

em 460 nm) was then possible based on calibration graphs relating

peak height ratios to drug concentrations over the range 50-500

ng/ml (31).

Differential pulse polarography based on the reduction peak of the

4,5-azomethine group, while not specific for diazepam among 1,4-

benzodiazepines, can measure this class of compounds in blood at

a sensitivity of 250 ng/ml. The concentration of drugs and metabolites is calculated on the basis of the resultant current of appropriate internal standards. Resolution of particular benzodiazepines can be achieved by incorporation of a TLC step, which also

increases the sensitivity to 50 ng/ml (32).

Currently, the techniques most widely used for analysis of benzodiazepines include high performance liquid chromatography (HPLC),

immunoassays, and gas chromatography with electron capture detection (GC-ECD). HPLC has the advantage that it can be used to analyze for any of the benzodiazepines and their major metabolites.

The immunoassay techniques (RIA, EMIT) offer greater speed and,

when large numbers of samples are involved, lower cost than chromatographic methods; however, they have a very limited dynamic range

and are more suitable for qualitative screening than quantitative

measurement. GC-ECD offers excellent sensitivity and specificity

if a high quality column is used, but is unsatisfactory for the

132
analysis of some benzodiazepines and their metabolites because of

their thermal instability. A more detailed discussion of each of

the techniques follows.

Quantitative high performance liquid chromatography on porous silica, using n-heptane:isopropanol:methanol (40:10:1) with detection

by UV absorption at 232 nm, was developed for routine forensic determinations of diazepam and N-desmethyldiazepam in human blood.

Extraction was made with benzene followed by a back extraction into

acid. When concentrations were greater than 100 ng/ml., 1 ml of

blood was sufficient for quantitation by HPLC, but for lower concentrations (25-100 ng/ml), 2 ml of blood and repetition of each

extraction step was necessary (33).

A reverse-phase HPLC column was used for simultaneous measurement

of diazepam and its pharmacologically active metabolites, N-desmethyldiazepam and oxazepam (34). An internal standard (chlordiazepoxide) was added to 250 μl of serum or plasma together with

a sodium phosphate buffer (pH 11) and the mixture was extracted

with chloroform. The extract was chromatographed on a μ-Bondapak

CN column (Waters Association, Milford, MA) using a solvent system

of acetonitrile and phosphate buffer (1:2). UV detection at 254

nm gave a linear response from 20 to 2000 ng for each drug and

metabolite. Day-to-day precision was reported to be 5 to 10 percent. Two recent reports also describe reverse-phase HPLC procedures for diazepam which were used to confirm measurements obtained

by GC-ECD). Both methods employed UV detection at 254 nm (35,36).

An electrochemical detector operated in a reduction mode served as

the detector for an HPLC assay for diazepam, nitrazepam, and chlordiazepoxide. A mobile phase of methanol:water (60:40) containing

0.05 M ammonium acetate (pH 7.25) was used with a reverse-phase

column. Interference due to reduction of oxygen is a major problem when electrochemical detectors are operated in the reduction

mode. Attempts to remove oxygen from the solvent and sample failed

to completely eliminate this interference. However, diazepam,

which can be measured at a reduction potential of -0.93 v, could

be detected at quantities as low as 3 ng (37).

A radioimmunoassay (RIA) procedure for diazepam in blood extracts

was developed following production of two different antibodies, one

specific for diazepam and the other recognizing both diazepam and

its metabolite, N-desmethyldiazepam. The RIA was capable of detecting 1 ng of the drug in dried ether extracts of whole blood,

and the response was linear to 100 ng (38). In a separate RIA

developed specifically for N-desmethyldiazepam, cross reaction

with diazepam and most other benzodiazepines was less than 1 percent; this assay permitted detection of as little as 3 ng/ml of

the metabolite in only 10 μl of plasma (39). Another immunoassay

procedure, EMIT, has also been reported for benzodiazepines (40),

but it is not specific for diazepam and is used only for urine because of interference by enzymes in plasma.

133
Gas chromatography (GC) has been used extensively in the analysis

of benzodiazepines. The high polarity and low volatility of the

benzodiazepines require use of a thermally-stable liquid phase

and well-deactivated column packings. Because gas chromatography

of these compounds, and particularly N-desmethyldiazepam, is difficult, some of the earlier GC methods involved conversion of the

benzodiazepines to benzophenones by acid hydrolysis prior to extraction and GC analysis (41,42). However, with recent improvements in GC column technology it is now possible to chromatograph

both diazepam and N-desmethyldiazepam without prior chemical transformations (Table 1).

Gas chromatography with flame ionization detection (FID) does not

offer sufficient sensitivity to assay blood concentrations of diazepam after therapeutic dosage. However, because of the presence

of the electronegative chlorine substituent, diazepam and its metabolites are well suited for electron capture detection (ECD) and

many assays using this detector have been published (9,28,35,36,

43-50).

In general, the GC-ECD assays are able to measure diazepam concentrations as low as about 10 ng/ml, although a sensitivity of 1 ng/

ml has been reported for an assay of the drug in human cerebrospinal fluid when a 63Ni detector was used and griscofulvin was added

as an internal standard to the extraction residue just before injection into the chromograph (9). An equally sensitive (1 ng/ml)

method for the assay of N-desmethyldiazepam, suitable for pharmacokinetic studies, uses N-desmethyl tetrazepam as the internal standard and relies on extraction from plasma with diethyl ether and an

acid back extraction for additional purification. A recovery of

at least 70 percent is reported. The dried ether residue is dissolved in hexane:acetone (4:1) and chromatographed on a column containing 1 percent Poly-A103 on Gas Chrom Q with a 63Ni detector.

This degree of sensitivity requires scrupulously clean glassware,

extremely pure reagents, and careful extraction/purification procedures (4 7). ECD assays are time-consuming and are usually unsuitable for rapid analysis of large numbers of clinical samples;

therefore a simplified procedure was developed in which 2 ml of

Na3PO4-saturated distilled water was added to 1 ml of whole blood

to precipitate proteins (final pH = 12.8). After thorough vortex

mixing, the sample was extracted with benzene:methylene chloride

(9:1). The extract was evaporated to dryness, dissolved in 100

μl benzene:acetone:methanol (80:15:10) and analyzed by GC-ECD without further purification. The sensitivity reported for this procedure was 5 ng/ml of blood (43).

In another adaptation of GC-ECD for rapid (< 1 hr) quantitation of

diazepam in 1-ml samples of human blood, methyl clonazepam was

added to serum or plasma as an internal standard prior to extraction with toluene at pH 9, back extraction into acid, and reextraction with toluene. Underivatized diazepam and N-desmethyldiazepam

were analyzed on a 3 percent OV-17 column equipped with 63Ni-ECD

(44) Similarly, sensitivities of 5 ng diazepam and 10 ng N-desmethyldiazepam per ml of plasma have been reported for automated

134
TABLE 1. GAS CHROMATOGRAPHIC ASSAYS FOR DIAZEPAM

AND/OK N-DESMETHYLDIAZEPAM

Extraction

Solvent

Toluene:heptane

(4:1)

Diethyl ether

plus back

extraction

Benzene:

methylene

chloride (9:1)

Toluene plus

back extraction

Benzene 3% OV-17

Toluene:heptane

(4:1) plus 1.6%

isoamyl alcohol

Toluene:heptane:

isoamyl alcohol

(76:20:4)

Hexane:ethyl

acetate (7:3)

Toluene

Toluene:heptane

(9:1)

Diethyl ether

plus back

extraction

Benzene:isoamyl

alcohol

Ethyl acetate

GC

Column

2% OV-17

1%

Poly-A103

3% OV-17

3% OV-17

3% OV-17

3% OV-17 Flunitrazepam ECD 35

3%

SP2250-DB

3% OV-17

3% OV-17

3% OV-17

3% OV-17

3% OV-17

Internal

Standard

Griseofulvin

Desmethyltetrazepam

Methylclonazepam

Methyl

nitrazepam

Medazepam

Prazepam ECD 36

Flurazepam ECD 28

Prazepam ECD 49

Prazepam NPD 51

50

52

53

Ro 7-9957 and

Ro 7-9749

N-C2H7-

diazepam

N-C2H7-

diazepam

135

Detector

ECD

ECD

ECD

ECD

ECD

ECD

ECD

MS

MS

Reference

9

47

43

44

45,48

46

- -
Extraction

Solvent

Diethyl ether

plus back

extraction

G C Internal

Column Standard

1% OV-17 Medazepam

Detector Reference

FID 54

Butyl chloride 2% OV-17 Prazepam FID 55

GC-ECD analysis using benzene for extraction and methylnitrazepam

as internal standard (45). When a single-step extraction method

was combined with GC using a frequency-response ECD, linear sensitivity over the range of 10- 500 ng/ml of serum was achieved (46).

The method permitted results of up to 40 clinical samples to be

reported on the same day.

Another rapid extraction procedure has been applied to simultaneous

quantitation of four benzodiazepines in blood and plasma. One ml

of saturated borate buffer was added to 0.05 to 1.0 ml of body fluid

and the mixture extracted with 1 ml of a solvent system consisting

of toluene:heptane:isoamyl alcohol (76:20:4). After centrifugation,

1.5 μl of the organic phase was removed and injected directly into

the GG-ECD. The method gave adequate sensitivity for measurement

of therapeutic levels of the benzodiazepine drugs and their metabolites (35).

N-Desmethyldiazepam has been measured in plasma by GC-ECD following

administration of dipotassium clorazepate, a drug which undergoes

spontaneous decarboxylation to N-desmethyldiazepam at pH < 4 (48).

A rapid and convenient extraction procedure was used, consisting

of addition of 0.1 ml saturated potassium chloride solution to 0.1

ml of plasma and extraction with 1 ml of benzene containing 25 ng

of methylnitrazepam as the internal standard.

Various methods have been reported for improving the chromatographic characteristics of the benzodiazepines, and particularly Ndesmethyldiazepam. The need for priming the GC column before beginning each day’s analyses is frequently mentioned. Priming

agents that have proven beneficial include: 1) blank plasma extracts (35,50,59), 2) lecithin in ethanol (5 g/l) (49), and 3)

cholesterol in acetone (49). A new liquid phase (SP 2250-DB, Supelco, Inc.) was found to considerably reduce the GC peak tailing

of N-desmethyldiazepam when compared with OV-17 or OV-1 columns

(36).

Because the free N-hydrogen in N-desmethyldiazepam can hydrogen

bond to active sites in the GC column, conversion of the metabolite

to an N-alkyl derivative can result in dramatically improved chromatographic behavior. The N-butyl derivative was formed by treatment of a dimethylacetamide solution of the extracted residue with

tetrabutylammonium hydroxide and 1-iodobutane (49). The N-butylation procedure gave more reliable results than formation of the

136
trimethylsilyl derivative (54). A similar derivatization involving formation of the N-propyl derivative of N-desmethyldiazepam

was combined with use of a nitrogen-phosphorus detector for quantitation of diazepam and its major metabolite in whole blood (51).

Although the FID is not as sensitive as the ECD, it is less easily

contaminated and because of its wider range of linearity it is

often preferred for routine use in pathology and toxicology laboratories. The incorporation of back extraction procedures and internal standards allows diazepam to be measured at serum concentrations as low as 80 ng/ml (54). However, because many neutral

and basic drugs may be present in the final extract, confirmation

of identity by TLC is desirable (55).

The use of mass spectrometry as the detector system for GC separation of drugs in biological fluids is indicated when the drugs

are present in the picogram to low nanogram per ml range. N-C2H3

Diazepam was used as the internal standard in two studies concerned with the quantitation of diazepam in plasma (52) and in

breast milk (53) by GC/MS.

Both diazepam and N-desmethyldiazepam are readily extracted from

biological media by various organic solvents. The extractions are

normally done at alkaline pH. Back extraction into acid has been

employed to achieve additional sample purification. However, some

salts of the drug and its metabolites are difficult to extract

from solvents such as chloroform and methylene chloride.

Diethyl ether is commonly used for extraction of diazepam and Ndesmethyldiazepam from blood, urine, or bile (16,18,38,54,56); at

pH 7 the recovery efficiency is 90 percent. Other solvents reported for diazepam extraction include benzene (> 90 percent from

blood, urine hydrolysates, or tissue homogenates) (10,26,43,45,50);

benzene:methylene chloride (9:1, > 85 percent from blood) (3,32,43);

ethyl benzoate (56); chloroform (90 percent) (58,59); hexane:isoamyl alcohol (98.3:1.7) from blood or gastric fluid (30); 1-chlorobutane followed by chloroform after back extraction into acid

(100 percent from blood or tissue homogenates) (55); and toluene:

heptane:isoamyl alcohol (78: 20:2, 96 percent at pH 9.5 from blood,

homogenates, and urine) (21,35); or toluene containing 1.5 percent

amyl alcohol to prevent absorption onto glass (83 percent from

blood) (44).

The choice of solvent and pH depends in part on the metabolites

to be measured. For example, the hydroxy metabolites have different partition properties from diazepam, and oxazepam is lost at

a pH greater than 10 (21). Because diazepam metabolites occur

mainly in the form of water-soluble conjugates in urine, Amberlite

XAD-2 resin has been recommended for extraction of these compounds

from urine made alkaline to pH 8-9. Absorbed compounds are eluted

with ethyl acetate:methanol:acetic acid (90:10:0.1). The eluate

is evaporated and the residue is taken up in buffer and hydrolyzed

with ß-glucuronidase/arylsulfatase before extraction with ethyl

acetate. A recovery of 95 percent has been reported for this procedure (60).

137
Diazepam is remarkably stable; no loss of concentration was found

to occur in plasma samples stored for 1 year at 20°C, 8 weeks at

4°C, or 20 days at room temperature (61).

Extensive reviews of the analysis of diazepam and other benzodiazepines in biological samples have been published (4.1,62).

EXPERIMENTAL PROCEDURE

Blood concentrations of the active metabolite, N-desmethyldiazepam,

are often comparable to, or greater than that of the parent drug.

Therefore it is important to measure the concentrations of both

compounds. The following procedure involves addition of deuterated analogs of diazepam and its major metabolite to the body fluid

specimen for use as internal standards and a simple direct extraction into a small voulme of organic solvent followed by injection

of an aliquot of the organic extract into the GC/MS. The ratios

of concentrations of the drug and metabolite to their deuterated

analogs are measured by selected ion monitoring using chemical

ionization with methane and ammonia as reagent gases. Because

the method involves only a single extraction, no concentration of

the extract, and no derivatization, it is very rapid; also it should

provide sufficient sensitivity to measure drug and metabolite concentrations at least as low as 5 ng/ml.

Standards and Reagents

The diazepam used in the development of this method was purchased

from Applied Science Laboratories, State College, PA 16801. The

N-desmethyldiazepam was obtained as a gift from Hoffmann-La Roche

Inc., Nutley, N.J. 07110. The pentadeuterated analogs of diazepam

and N-desmethyldiazepam were synthesized starting from bromobenzene2H5

 by published procedures (63). 0n the basis of GC/MS analysis, both deuterated analogs were more than 99 percent pure and

had the following isotopic composition: 2H5, 96.0 percent; 2H4,

3.4 percent; 2H3, 0.4 percent; and 2H2, 

2H1

 and 2H0 < 0.2 percent.

All of the deuterium atoms were located on the phenyl ring of each

structure. The uncorrected melting points for the deuterated compounds were: diazepam-2H5

, 131-132°C; N-desmethyldiazepam-2H5,

215-216°C.

Prepare a pH 9.6 buffer by dissolving 500 grams of potassium monohydrogen phosphate (K2HP04

) in about 500 ml of distilled water.

Allow the solution to cool to room temperature and then add sufficient distilled water to make exactly one liter.

For the purpose of preparing calibration graphs and working standards

stock solutions of diazepam and N-desmethyldiazepam are prepared

as follows. Weigh into a 100-ml volumetric flask about 10 mg of

diazepam and 10 mg of N-desmethyldiazepam, and record the weights

to the nearest 0.1 mg. Dissolve the measured drug and metabolite

in about 75 ml of methanol and then add additional methanol to

givo exactly 100 ml of solution. This solution will be referred

to as the “0.10-mg/ml stock solution” although it should be labeled

138
with its actual concentration based upon the exact measured weights

of diazepam and N-desmethyldiazepam. A series of working standard

solutions can then be prepared by appropriate dilution of the stock

solution as described in Chapter 2.

The stock solution of diazepam-2H5 and N-desmethyldiazepam-2H5 is

prepared in the same manner. For the measurement of drug and metabolite in body fluids within the concentration range 5 to 1000 ng/

ml, addition of approximately 40 ng of the internal standards to

each ml of specimen is satisfactory. This can be conveniently

done by adding 100 μl of a methanolic solution containing 400 ng/

ml of both diazepam2H5 and N-desmethyldiazepam-2H5 to each ml of

specimen. Prepare the 400-ng/ml internal standard solution by diluting 1 ml of the 0.10-mg/ml stock solution of diazepam-2H5

 and

N-desmethyldiazepam-2H5 to 250 ml with methanol.

Store stock and working standard solutions in well-stoppered or

capped glass vessels in the dark at less than 0°C.

Extraction Procedure

Transfer 1 ml of body fluid (whole blood, plasma, or urine) to a

5-ml glass-stoppered, conical centrifuge tube. Add 100 μl of the

internal standard solution containing 400 ng/ml of diazepam-2H5

and N-desmethyldiazepam-2H5 and vortex the mixture for 10 sec. Allow the sample to equilibrate for 15 min and then add 1 ml of the

K2HPO4 buffer (pH 9.6), followed by 100 μl of a solvent consisting

of toluene:heptane:isoamyl alcohol in a volume ratio of 70:20:10.

Stopper the tube and vortex mix for at least 30 sec. Separate the

phases by centrifugation for 10 min. The organic solvent will have

formed a narrow upper layer from which an aliquot can be removed

with a syringe for injection into the GC/MS, If there is not a

clean separation of layers and a clear upper solvent layer, repeat

the centrifugation at a higher speed ( 2000 X G).

GC/MS Analysis

The experimental conditions for the simultaneous GC/MS analysis of

diazepam and N-desmethyldiazepam are as follows:

GC column:

Carrier gas:

Reagent gas:

Temperatures

1 m x 2 mm (I.D.) glass column packed with 3

percent OV-17 on Ultra-bond 20M, 100/120 mesh

(RFR Corp., Hope, RI, 02831)

Methane, 15-20 ml/min

Ammonia, introduced into the ion chamber as

described in Chapter 2

Injector, 290°C

Column, 270°C isothermal

GC/MS transfer line, 280°C

Ion source, 160°C

139

328-011 0 - 80 - 10
Under these conditions the diazepam should elute at 1 to 2 min and

the N-desmethyldiazepam at 2 to 4 min.

Prior to beginning the GC/MS analysis of samples, the performance

of the total GC/MS system should be evaluated and optimized as

described in Chapter 2. The ions to be monitored occur at m/z 285,

290, 271 and 276. They correspond to the protonated molecule ions

for diazepam, diazepam2H5, N-desmethyldiazepam, and N-desmethyldiazepam- 2H5

, respectively. With the divert valve in the divert

position, inject 4 to 10 μl of the organic extract layer into the

GC/MS. After approximately 1 min switch the divert valve so that

the entire carrier gas flow enters the ion source of the mass spectrometer and begin data acquisition. For maximum sensitivity monitor just the diazepam ion masses (m/z 285 and 290) until the dliazepam elutes from the GC column, and then monitor the N- desmethyldiazepam ion masses (m/z 271 and 276) until it elutes. If the

GC/MS data system does not permit rapid changes in the monitored

ion masses, all four ion masses must be monitored during the elution period for the drug and its metabolite. Normally, the latter

procedure will result in only a small sacrifice in sensitivity.

When the N-desmethyldiazepam has eluted, cease data acquisition

and return the divert valve to the divert position. The quantities

of diazepam and N-desmethyldiazepam in the body fluid are determined

by measuring the peak heights (or areas) of the drug, the metabolite and the internal standards in the selected ion current profiles and relating the ratios of peak heights to calibration graphs

(Chapter 2). Calculate the body fluid concentrations of diazepam

and N-desmethyldiazepam by dividing the measured quantities of

these compounds by the exact volume of specimen extracted.

Discussion of the Experimental Procedure

Diazepam and some of the other benzodiazepines and their metabolites are most often analyzed by GC-ECD which can provide very high

sensitivity and specificity. The GC/MS method described here is

capable of comparable sensitivity and even better specificity than

the GC-ECD methods. Figures 1 and 2 compare the methane CI and

the methane-ammnonia CI mass spectra of diazepam and N-desmethyldiazepam. All of the spectra are dominated by abundant protonated molecule ion peaks. For these two compounds either reagent

gas can be used and should provide similar sensitivity, but the

methane-ammonia reagent gas combination will result in a more selective ionization. High selectivity is necessary because the single, direct extraction procedure results in a relatively “dirty”

extract.

Additional specificity can be provided by also monitoring the 37Clisotope peaks (m/z 287, 292, 273, and 278) for each of the protonated molecule ions. The peak height of the 37CI-isotope peak should

be exactly one-third the intensity of the corresponding 35Clisotope peak height for each compound. Any deviation from a

ratio of one-third indicates an interfering compound is contributing to the ion current at one of the monitored masses. Usually,

a visual examination of the ion current profiles will disclose

140
FIGURE 1. CHEMICAL IONIZATION MASS SPECTRA OF DIAZEPAM

141
FIGURE 2. CHEMICAL IONIZATION MASS SPECTRA OF N-DESMETHYLDIAZEPAM

142
which of the peak heights is inaccurate due to the interference;

i.e. the peak will have a shoulder, or will be perceptibly broadened, or will have an abnormal retention time.

If even greater sensitivity is needed, negative ion chemical ionization can be used. Diazepam and N-desmethyldiazepam have high

electron affinities and therefore are efficiently ionized by electron capture under methane chemical ionization conditions. In unpublished work conducted at Battelle Columbus Laboratories, up to

a 30-fold improvement in sensitivity was achieved by switching

from positive ion detection to negative ion detection while operating the mass spectrometer under conventional methane CI conditions. Unfortunately, there are significant problems associated

with quantitative measurements obtained with electron-capture negative ion chemical ionization. One difficulty is illustrated in

Figure 3, which compares the negative ion CI mass spectra of diazepam acquired at ion source temperatures of 80° and 170°C. In

the spectrum acquired at the lower ion source temperature the molecular anion at m/z 284 is the only prominent peak observed, and

the chloride anion at m/z 35 has a relative intensity of only 10

percent. At an ion source temperature of 170°C a dissociative

electron capture process dominates so that the chloride anion becomes the most intense peak in the spectrum, while the molecular

anion peak has virtually vanished (about 2 percent relative intensity). Consequently, in order to use electron-capture negative

ion chemical ionization for quantitative measurement of diazepam

with high sensitivity, it is necessary to maintain a low ion source

temperature (< 100°C) even though the ion source tends to become

contaminated more rapidly at low temperatures. The large temperature effect experienced with negative ion chemical ionization

can also cause poor reproducibility in quantitative measurements,

although the use of deuterated analogs as internal standards largely compensates for changes in the ionization process.

The choice of chromatographic column is critical in the GC/MS analysis of diazepam and N-desmethyldiazepam, particularly when low

nanogram-quantities are injected onto the column. The Ultra-bond

20M column packing supplied by the RFR Corporation is a very inactive surface, well suited for gas chromatography of polar drugs

and metabolites. Both diazepam and N-desmethyldiazepam give narrow, relatively symmetrical peaks when chromatographed under the

conditions specified here.

143
FIGURE 3. NEGATIVE ION METHANE CI MASS SPECTRA OF

DIAZAPEM RECORDED AT DIFFERENT ION SOURCE

TEMPERATURES

144
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR DIAZEPAM

Nomenclature

Chemical Name: 7-Chloro-1,3-dihydro-1-methyl-5-phenyl-2H-1,4-benzodiazepin-2-one

Empirical Formula: C16H13C1N2O

Chemical Abstracts Registry Number: 439-14-5

Trade Names: Ansiolin, Apavrin, Apozepam, Atensine, Atilen, Bialzepam, Calmpose, Ceregulart, Dipam, Eridan, Faustan,

Lembrol, Levium, Morosan, Noan, Pacitran, Paxate,

Paxel, Relanium, Seduxen, Setonil, Stesolid, Stesolin, Tranimul, Valium, Vival, Vivol

Physical Constants

Physical Appearance: Off-white to yellow crystalline powder

Melting Point: 125-126°C

Specific Rotation: optically inactive

pKa: 3.4

Solubility: At room temperature 1 g of diazepam will dissolve in

20 1 of water, 56 ml of diethyl ether, 20 ml of methanol, 4.5 ml of benzene, and less than 2 ml of chloroform.

UV Absorption: max at 242, 285 and 368 nm (in 2N HCl)

PHYSICAL, CHEMICAL AND SPECTROMETRIC

DATA FOR N-DESMETHYLDIAZEPAM

Nomenclature

Chemical Name: 7-chloro-1,3-dihydro-5-phenyl-2H-1,4-benzodiazepin-2-one

Empirical Formula: C15H11C1N2O

Chemical Abstracts Registry Number: 1088-11-5

Physical Constants

Physical Appearance: Off-white to yellow crystalline powder

Melting Point: 216-217°C

145
REFERENCES

1. D.J. Greenblatt and Richard I. Shader. New England J. Med.,

291, 1011 (1974).

2. A. MacDonald, A.F. Michaelis, and B.Z. Senkowski, in K. Florey,

ed., Analytical Profiles of Drug Substances, Vol. 1. Academic

Press, New York, p. 79 (1972).

3. M.M. Reidenberg, M. Levy, H. Warner, C.B. Coutinho, M.A.

Schwartz, G. Yu, and J. Cheripko. Clin. Pharmacol. Ther.,

23, 371 (1978).

4. J.J. Tanja, C.R. Clark, and D.E. Griffies. J. Anal. Toxicol.,

1, 175 (1977).

5. I. Saxén and L. Saxén. Lancet, 2 (7933), 498 (1975).

6. J.F.W. Haffner, J. Morland, J. Setekleiv, C.E. Stromsaether,

A. Danielsen, P.T. Frivik, and F. Dybing. Acta Pharmacol.

Toxicol., 32, 161 (1973).

7. J. Wiersum. J. Amer. Med. Assoc., 227, 79 (1974).

8. L. Hillestad, T. Hansen, and H. Melsom. Clin. Pharmcol.

Ther., 16, 485 (1974).

9. J. Hendel. Acta Pharmacol. Toxicol., 37, 17 (1975).

10. W.A. Mahon, T. Inaba, T. Umeda, E. Tsutsumi, and R. Stone.

Clin. Pharmcol. Ther., 19, 443 (1976).

11. J. Kanto, L. Kangas, and T. Siirtola. Acta Pharmacol. Toxicol.,

36, 328 (1975).

12. R. Sellman, J. Xanto, E. Raijola, and A, Pekkarinen. Acta

Pharmacol. Toxicol., 37, 34.5 (1975).

13. K. Korttila, M.J. Mattila, and M. Linnoila. Acta Pharmacol.

Toxicol., 36, 190 (1975).

14. R. Dixon, M.A. Brooks, E. Postma, M.R. Hackman, S. Spector,

J.D. Moore, and M.A. Schwartz. Clin. Pharmacol. Ther., 20,

450 (1976).

15. U. Klotz, K.H. Antonin, and P. Bieck. Arch. Pharmacol., 287,

Suppl, R90 (1975).

16. U. Klotz, G.R. Avant, A. Hoyumpa, S. Schenker, and G.R. Wilkinson. J. Clin. Inv., 55, 347 (1975).

17. S.A. Kaplan, M.L. Jack, K. Alexander, and R.E. Weinfeld. J.

Pharm. Sci., 62, 1789 (1973).

146
18. J.A.F. de Silva, B.A. Koechlin, and G. Bader. J. Pharm. Sci.,

55, 692 (1966).

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

E. van der Kleijn, P.J. M. Guelen, C.G.W.M. van Wijk, I.

Baars, and T.B. Vree, in G. Levy, ed., Clinical Pharmokinetics: Symposium. Amer. Pharm. Assoc., Acad. Pharmaceutical

Sciences (October 1974), p. 79.

H. Dasberg. Psychother. Psychosom., 24, 113 (1974).

I.A. Zingales. J. Chromatogr., 75, 55 (1973).

L. Hillestad, T. Hansen, H. Melsom, and A. Drivenes. Clin.

Pharmacol. Ther., 16, 479 (1974).

M.A. Schwartz, B.A. Koechlin, E. Postma, S. Palmer, and G.

Krol. J. Pharmacol. Exp. Ther., 149, 423 (1965).

E. Tsutsumi., T. Inaba, W.A. Mahon, and W. Kalow. Biochem.

Pharmacol., 24, 1361 (1975).

R. Sellman, J. Kanto, E. Raijola, and A. Pekkarinen. Acta

Pharmacol. Toxicol., 36, 33 (1975).

M. Linnoila, K. Korttila, and M.J. Mattila. Acta Pharmacol.

Toxicol., 36, 181 (1975).

M. Mandelli, P.L. Morselli, S. Nordio, G. Pardi, N. Principi,

F. Sereni, and G. Tognoni. Clin. Pharmacol. Ther., 17, 564

(1975).

G.J. DiGregorio, A.J. Piraino, and E. Ruch. Clin. Pharmacol.

Ther., 24, 720 (1978).

M. Mandelli, G. Tognoni, and S. Garattini. Clin. Pharmacokinetics, 3, 72 (1978).

J.C. Valentour, J.R. Monforte, B. Lorenzo, and I. Sunshine.

Clin. Chem., 21, 1976 (1975).

P.J. Van der Merwe and J.M. Steyn. J. Chromatogr., 148,

549 (1978).

M.A. Brooks, L. D'Arconte, M.R. Hackman, and J.A.F.de Silva.

J. Anal. Toxicol., 1, 179 (1977).

A. Bugge. J. Chromatogr., 128, 111 (1976).

J.J. MacKichan, W.J. Jusko, P.K. Duffner, and M.E. Cohen.

Clin. Chem., 25, 856 (1979).

M.A. Peat and L. Kopjak. J. Forensic Sci., 24, 46 (1979).

J.E. Wallace, H.A. Schwertner, and E.L. Shimek, Jr. Clin. Chem.,

25, 1296 (1979).

147
37. W. Lund, M. Hannisdal, and T. Greibrokk. J. Chromatogr.,

173, 249 (1979).

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56. J.F. Steyn and H.K.L. Hundt. J. Chromatogr., 107, 196 (1975).

B. Peskar and S. Spector. J. Pharmacol. Exp. Ther., 186,

167 (1973).

R. Dixon, W. Glover, J. Earley, and M. Delaney. J. Pharm. Sci.,

68, 1471 (1979).

B.H. Haden, K.G. McNeil, N.A. Huber, W.A. Khan, P. Singh, and

R.S. Schneider. Clin. Chem., 22, 1200 (1976).

D.M. Hailey. J. Chromatogr., 98, 527 (1974).

A.K. Dhar and H. Kutt. Clin. Chem., 25, 137 (1979).

J.A.F. de Silva, I. Bekersky, C.V. Puglisi, M.A. Brooks, and

R.E. Weinfeld. Anal. Chem., 48, 10 (1976).

J.L. Ferguson and D. Couri. J. Anal. Toxicol., 1, 171 (1977).

R.E. Weinfeld, H.N. Posmanter, K.C. Khoo, and C.V. Puglisi.

J. Chromatogr., 143, 581 (1977).

M. Linnoila and F. Dorrity, Jr. Acta Pharmacol. Toxicol.,

41, 458 (1977).

A. Viala, J.-P. Cano, J.-P. Reynier, and R. Rispe. J. Chromatogr., 147, 349 (1978).

D. Haidukewych, E.A. Rodin, and R. Davenport. Clin. Chem.,

26, 142 (1980).

J.J. De Gier and B.J. 't Hart. J. Chromatogr., 163, 304 (1979).

D.J. Greenblatt. Clin. Chem., 24, 1838 (1978).

H.H. McCurdy, E.L. Slightom, and J.C. Harrill. J. Anal. Toxicol., 3, 195 (1979).

B.M. Yehia, J.S. Oliver and A. Mackay. Vet. Hum. Toxicol.,

21 (Suppl.), 167 (1979).

M.G. Horning, W.G. Stillwell, J. Nowlin, K. Lertratanangkoon,

D. Carroll, I. Dzidic, R.N. Stillwell, and E.C. Horning. J.

Chromatogr., 91, 413 (1974).

M.S. Greaves. Clin. Chem., 20, 141 (1974).

R.C. Baselt, C.B. Stewart, and S.J. Franch. J. Anal. Toxicol.,

1, 10 (1977).

148
57. A.G. Howard, G. Nickless, and D.M. Hailey. J. Chromatogr.,

90, 325 (1974).

58. G. Banelli and P. Marini. Boll. Chim. Unione Ital. Lab. Prov.,

2, 310 (1976).

59. L.B. Foster and C.S. Frings. Clin. Chem., 16, 177 (1970).

60. H. Sawada, A. Hara, S. Asano, and Y. Matsuaoto. Clin. Chem.,

22, 1596 (1976).

61.

62.

P.J. Howard. J. Pharm. Pharmacol., 30, 136 (1978).

B.J. Gudzinowicz and M.J. Gudzinowicz, Analysis of Drugs and

Metabolites by Gas Chromatography-Mass Spectrometry, Vol. 2:

Hypnotics, Anticonvulsants, and Sedatives. Marcel Dekker,

New York, ch. 2, VII (1977).

63. A.F. Fentiman, Jr. and R.L. Foltz. J. Labelled Compounds and

Radiopharmaceuticals, 13, 579 (1977).

149
Chapter 10

Amphetamine

For many centuries the khat plant has been cultivated in the Middle

East for its central nervous system (CNS) stimulating properties.

A synthetic relative of the khat alkaloids, amphetamine ( - methylphenethylamine, I) was first produced in 1887, but its psychopharmacological properties were not described until 1927 (1). For the

next 30 years, medical use of amphetamine (as a bronchodilator, as

an appetite suppressant, in the treatment of hyperkinesis in children, and for narcolepsy) was extensive (2), but by 1967 the potential for abuse of the drug was causing widespread concern. Regular

oral use of amphetamine can cause paranoid psychosis and a disabling dependence, and intravenous use is even more likely to lead

to these effects. Primarily because of an initial euphoria, especially following intravenous injection, amphetamine has become a major drug of abuse. Appetite suppression, insomnia, sudden changes

in mood, and compulsiveness are the common results of even moderate amphetamine use. Tolerance develops rapidly, so that increasing doses are required to achieve the desired stimulation (3).

Amphetamine is marketed for medical purposes as Dexedrine (d-amphetamine) and as Benzedrine (d,l-amphetamine), or in combination with

a barbiturate (e.g. Dexamyl, d-amphetamine + amobarbital) (4). However, because of the relative ease of manufacture, illicit preparations of amphetamine are common in “street” use. As a result, contaminants are often present. Significant quantities of zinc (1-2

percent) have been found in some illicit samples, as well as polycondensation products whose pharmacological and toxicological properties are unknown (5), although there is evidence that the intensity of euphoria following intravenous injection may depend on contaminants (3). Severe reactions can occur in nontolerant persons

following a dose of 30 mg of amphetamine, while persons who have

developed tolerance can endure doses of 400-500 mg. While true

physical dependence has not been proven, very strong psychological

habituation to amphetamine is common. In its extreme form, the

“speed cycle” of heavy intravenous use for days at a time results

150
in sleep deprivation (hallucinations, etc.), malnutrition, and often

infection (1). For medical purposes, many structural modifications

of amphetamine have been made in efforts to eliminate the undesirable effects of the drug while preserving the crucial structural

feature (the d-phenethylamine skeleton) for its antifatigue, anorexic, or other pharmacological properties. Structurally related

drugs include p-chloroamphetamine, phentermine, fenfluramine, phenmetrazine, and methylphenidate, some of which have also become

drugs of abuse (6).

The mechanism of the CNS stimulant action is thought to be a triggering of the release of catecholamines (epinephrine, dopamine, norepinephrine) and blockage of their reuptake (7). Amphetamine also

exhibits peripheral - and ß-adrenergic activity (1). It has been

shown to increase cerebral blood flow in rats (8), and in man an

increased susceptibility to strokes and cerebral hemorrhages has

been reported. Elevated blood pressure and cardiac arrhythmia are

the common cardiovascular effects of amphetamine (1).

The optical isomers of amphetamine exhibit different pharmacological

effects. With respect to the CNS, the d-isomer is 4-10 times as

potent as the 1-isomer, but in cardiovascular effects the 1-isomer

is more potent (1). More d-amphetamine is retained in heart and

brain tissue, and only the d-isomer leads to the presence of p-hydroxy norephedrine as a metabolite in the brain. It is possible

that this metabolite could function as a false neurotransmitter,

which might account for the development of psychosis in some individuals after prolonged use of large doses of amphetamine (9).

METABOLISM AND PHARMACOKINETICS

The metabolism of amphetamine in man yields at least two biologically active metabolites, p-hydroxyamphetamine (II) and p-hydroxynorephedrine (III), both of which are found in significant levels

in the urine and undoubtedly contribute to the overall pharmacology

of the drug (10). ß-Hydxoxylation is carried out by an extra hepatic enzyme located in sympathetic nerve endings (11).

151
Metabolic N-oxidation of amphetamine has also been reported (12).

The hydroxylamine (IV) is unstable and is quickly converted by

chemical or metabolic processes to the oxime (V) and to phenylacetone (VI).

Based on the report that 30 percent of an administered dose of damphetamine in human subjects was found to be excreted as unchanged

drug, 3 percent as II (free and conjugated), 3 percent as phenylacetone, 20 percent as free and conjugated benzoic acid, and 5 percent as III, deamination seems to be the predominant metabolic

route in man, although p-hydroxylation is the primary route in rats.

Deamination is catalyzed by a liver microsomal enzyme that requires

reduced NADP (9). Deamination is stereoselective; the S-(+)-isomer

of amphetamine is preferentially metabolized (11).

The pKa of amphetamine is 9.77 (13), so it is ionized at physiological pH; nevertheless, it does readily penetrate the blood-brain

barrier. Excretion of the unchanged drug has been shown to fluctuate from 10 percent to 60 percent with changes in urinary pH (14).

A computerized method of predicting excretion curves (and by extension, body concentrations) has been developed (15).

Pharmacokinetic studies in humans have confirmed that the drug has

a high extravascular concentration in the body. In one of these

studies, peak blood concentrations (~35 ng/ml) were reached about

2 hr after ingestion of 10 mg of d-amphetamine sulfate, followed

by an exponential decline over the next 48 hr, with a plasma elimination half-life of 11-13 hr. Mean total urinary recovery of unchanged drug was 45 percent of the ingested dose (16). Another

report indicated an 8 hr plasma half-life for human subjects on

an acid diet, but 22 hr when on an alkaline diet (9). In a study

involving patients with amphetamine psychosis, the intensity of

psychosis correlated with the concentration of basic polar metabolites of the drug in urine but not with the amphetamine concentration in plasma. Amphetamine metabolism in these patients was

followed with the aid of a tritium-labeled analog of the drug. Under metabolic conditions producing acid urine (pH <6.6), renal elimination of unchanged drug (67-73 percent of the dose) was associated with a plasma elimination half-life of 7-14 hr and a rapid

reduction in psychotic symptoms. In patients with alkaline urine,

the plasma half-life of amphetamine was 18-34 hr, psychosis was of

longer duration, and hydroxylated metabolites in urine accounted

152
for 6-13 percent of the dose as compared with 3-8 percent in

acidic urine (17).

In a study using rats as models, inhibition of metabolism increased

the brain concentration of amphetamine by a factor of 5, indicating

that metabolism is more important than renal excretion as the route

of elimination of amphetamine, at least in that species (18). In

human subjects receiving 1500 mg of ascorbic acid daily prior to

amphetamine administration, the 24-hr excretion of a single 30 mg

dose of drug was decreased by an average of 38 percent, regardless

of urinary pH, although the acute pharmacological effects of the

drug were not altered by the ascorbic acid pretreatment. There

was no increase in hydroxylated metabolites to account for the

smaller quantity of amphetamine excreted (19).

A definitive compendium of information on the pharmacology and metabolism of amphetamine was published in 1970 (20), and an extensive

review of amphetamine metabolism in mammals, including man, appeared

in 1976 (11).

ANALYTICAL METHODOLOGY

The development of methods sufficiently sensitive, specific, and

reproducible for the identification and measurement of amphetamine

in biological fluids is important because there is a need to establish therapeutic dosages and plasma concentrations, to understand

tolerance and the stereoisomeric effects, to carry out toxicity

studies, and simply to identify drug users. The concentrations of

amphetamine found in urine, even after small doses, are high enough

to be detected by most methods available to clinical laboratories.

However, plasma concentrations are usually in the 5-100 ng/ml

range. Techniques used have included ultraviolet spectrophotometry

(UV), thin-layer chromatography (TLC), fluorometry, immunoassay,

gas chromatography (GC), and combined gas chromatography/mass spectrometry (GC/MS).

A quantitative UV method, with the advantages of relative speed and

simplicity for screening purposes, was reported in 1968. The amphetamine extract from urine, blood, or tissue homogenate was oxidized with ceric sulfate, using 0.8 N HCl as a catalytic agent, to

form a reaction product with a well-defined UV absorption curve

( max 287 nm, ,min 253 nm in hexane). Sensitivity was 0.5 μg/ml,

but the method was unable to distinguish between amphetamine and

methamphetamine (21).

Until the early 1970’s, fluorometry was the most widely used method

for the quantitative analysis of biogenic amines and amphetamines.

A fluorometric method, based on conversion of amphetamine to its dihydrolutidine derivative, utilized an automated continuous-flow apparatus for extraction of the derivatized drug from urine (22). The

detection limit was 0.15 μg/ml, but 2-phenylethylamine and 2-aminoheptane gave false positive reactions, Fluorometry combined with

TLC is widely used, especially for screening urine specimens for

amphetamine. For example, the drugs in urine can be separated

and concentrated by XAD-2 resin, chromatographed on silica gel in

153
a suitable solvent system, and sprayed with a color reagent. Sequential spraying with ethanolic ninhydrin-phenylacetaldehyde, aqueous sodium phosphate, and then p-dimethylaminobenzaldehyde has been

shown to distinguish amphetamine from other primary amines of forensic and toxicological interest such as mescaline, DOM, phenylpropanolamine, ß-phenethylamine, benzocaine, chlorphentermine , and

procaine. A final spray with chromotropic acid was found to be

necessary to distinguish amphetamine from 3,4-methylenedioxyamphetamine (MDA). A 2 μg detection limit for amphetamine was reported

for this technique (23). A Fluorescamine (4-phenyl- spiro [furan2(H)-1’-phthalen]- 3,3'dione) spray reagent was found to be more

sensitive than ninhydrin, detecting 250 ng of amphetamine in urine

extracts following TLC. Methamphetamine does not react with Fluorescamine (24). The Fluorescamine technique was adapted to in situ

spectrodensitometry of thin-layer chromatograms for quantitation

of amphetamine in urine extracts without elution. A solvent system of ethyl acetate:methanol:water:ammonia (85:13.5:10:0.5) was

used to separate amphetamine from extraneous fluorescent material.

A concentration of 0.25 μg/ml was detectable (25). 4-Chloro-7-

nitrobenzo-2,1,3-oxadiazole (NBD-Cl) has also been used to form a

fluorescent derivative of amphetamine in urine and blood extracts

before separation by TLC, and the fluorescence intensity used as a

quantitative measure of the amphetamine present. Sensitivity is

sufficient for measurement of therapeutic concentrations in blood

(10-100 ng/ml) (26). Although any primary or secondary amine will

yield a fluorescent derivative with NBD-Cl, TLC analysis using a

two-solvent system [e.g., pentane: ethyl ether (1:1) or methanol:

carbon tetrachloride (80: 30)] can be used to separate amphetamine

from compounds likely to yield false results (27).

Some assays developed for amphetamine do not require chromatographic

separation. As little as 10 ng/ml of amphetamine in plasma was detected by an enzyme assay based on the N-methylation of the drug

to form radioactive methamphetamine by means of a rabbit lung Nmethyltransferase and S-adenosyl-L-[methyl-3H] methionine as methyl

donor. Sensitivity, specificity, and reproducibility were achieved

by a combination of several extraction and drying procedures used

prior to liquid scintillation counting (28).

For large numbers of samples or for rapid screening, immunological

techniques are often the method of choice. They do not require

extensive extraction of biological materials or a high degree of

skill in operation. Radioimmunoassay (RIA) depends upon competition

between the unlabeled drug in the sample and a known amount of radiolabeled drug for the binding sites on specific drug antibodies in

rabbit serum. The antibody in some RIA assays for amphetamine binds

equally well with methamphetamine, but the method can detect 30

ng/ml of amphetamine in urine (29). A single RIA takes about 40

min, but several samples can be run concurrently (30). The other

common immunoassay, HIT, is based on the linkage of an enzyme

(lysozyme) to the amphetamine molecule to form the antigen reagent.

Antibody binds the drug and its attached enzyme, inactivating the

lysozyme. Any free amphetamine present in the sample competes with

the drug-enzyme complex for antibody binding sites; while unbound

enzyme is then free to hydrolyze the cell walls of bacteria used as

154
an indicator in the assay. The clearing of the cell walls suspension

(decrease in absorbance) is proportional to the amount of enzyme

label displaced from the antibody and hence to the amount of competing free drug present in the specimen. EMIT cannot presently

be used for blood, and it is subject to interference by lysozyme

naturally present in some urine samples and to false negatives

because of inactivation of enzyme in alkaline urines. However,

it takes only one minute to perform, it is as sensitive and specific as RIA, and it gives semiquantitative results based on calibration graphs (30).

Gas chromatography (GC) has been used at least since 1969 for measurement of amphetamine in urine and blood. Gas chromatography

of extracts, after addition of N,N-dimethylaniline as an internal

standard, on a column coated with 5 percent KOH + 10 percent Carbowax 6000 and using a flame ionization detector (FID), was sensitive enough to detect 0.1 μg/ml of amphetamine in urine (31). A

rapid GC method developed for routine use in urine screening incorporated diethylamine as the internal standard and confirmation by

preparation of a ketone derivative of amphetamine. Detection limits

were 0.01 ng/ml (32). For quantitation of amphetamine in blood by

GC a number of specific steps were recommended. These included

the use of Teflon-coated glassware to minimize absorption, coextraction with a scavenger amine (diethylamine), formation of the nonvolatile HCl salt, preparation of the trifluoroacetamide derivative,

and use of an internal standard (1-adamantanamine HCl). This combination of conditions gave a nearly complete (98 percent) recovery

of amphetamine from blood, and concentrations of 10-100 ng/ml were

measurable (33).

Because of the sensitivity limits of the flame ionization detector,

electron capture detection (ECD) after formation of suitable derivatives offers advantages over FID for GC assay of the very low

concentrations of amphetamine often present in blood samples. The

heptafluorobutyl (34), trichloroacetamide (35), and N-acetyl (36)

derivatives, among others, have been reported for this purpose. Oncolumn acylation of amphetamine with N- (trifluoroacetyl) imidazole,

or with N-(heptafluorobutyryl) imidazole, has also been reported.

The derivatives were formed quantitatively and reproducibly, making the technique suitable for quantitative analysis. The imidazole

reagents have the advantage over acyl chlorides or acid anhydrides

in that no acids are released into the chromatographic system (37).

A comparison study of the ECD response of several fluorinated amphetamine derivatives indicated that a sensitivity of 10 pg for the

pentafluorobenzamide derivative was possible using a 63Ni detector,

but pure compounds and not biological extracts were used in the

study (38). GC-ECD of the trichloroacetate derivative, also using

a 

63Ni detector, was able to measure concentrations of amphetamine

as low as 5 ng/ml in plasma (39). Although EC detectors may improve sensitivity, they can have the disadvantages of excessive

background or of severe quenching due to water and solvent. Preliminary studies of the analysis of amines by GC-ECD, using model

compounds such as phenethylamine and utilizing glass capillary columns, have suggested that the high resolving power of capillary

columns could improve quantitative analysis at the subnanogram level

155

328-011 0 - 80 - 11
of neurotransmitters and related compounds like amphetamine (40).

GC analysis of the metabolite N-hydroxyamphetamine is difficult

because of its instability under normal GC conditions, but the decomposition has been shown to be the result of catalytic activity

of the solid support rather than thermal conditions or oxidation

in the column. Direct GC analysis was found to be possible without

derivatization, by the use of a column containing 0.2 percent Carbowax 20M with glass beads as support material, operated at 100°C

(41).

For selected ion monitoring analysis of drugs and metabolites in

biological fluids preparation of derivatives is often desirable in

order to achieve better gas chromatographic characteristics and

to generate ions in the mass spectrometer which are more suitable

than those generated by the parent drug. Acetylation, trimethylsilylation (TMS), and perfluroacylation give amphetamine derivatives

which show good GC and MS characteristics (39).

Some acetylation procedures call for reaction times of up to several

hours and are therefore unsuitable for rapid analyses. Recently,

a rapid and quantitative acetylation of amphetamine was reported

in which trifluoroacetic anhydride was used in the presence of mercuric trifluoroacetate as a catalyst. The reagent acts on amphetamine in the form of either the free base or salt, and the method

requires no prior extraction of street drug samples to remove adulterants. Application of this derivatization procedure to GC/MS

analysis allowed detection of amphetamine at the picomole level,

within about 10 min total analysis time (42).

For purposes of mass spectral identification of primary amines in

urine samples, isothiocyanate derivatives have been shown to possess

some advantages over TMS derivatives because of a fragmentation pattern which eliminates a CNS-containing radical from the molecular

ion and allows for charge retention on the moiety containing the

phenyl group (43). The isothiocyanate derivative of amphetamine

is claimed to be even more stable than the trifluoroacetamide derivative. By monitoring the molecular ion of the isothiocyanate

derivative a signal-to-noise ratio of 3:1 was achieved for 1 ng

injected on column (44).

Dansylation of amphetamine with 5-dimethylamino-1-naphthalene sulfonyl chloride yields a UV-absorbing derivative which was employed

in an assay for amphetamine in urine and tissue homogenates. Amphetamine- 2H5

 was added to urine (pH 10) before extraction and dansylation. A benzene extract of the dansyl-amphetamine was chromatographed on thin-layer plates developed with benzene:triethylamine (8:1), and the zone identified by UV spectroscopy (365 nm)

was eluted with ethyl acetate, dried, and chromatographed on another plate (petroleum ether:toluene:acetic acid:water, 133:67:170:

30 v/v). The final extract was analyzed by high resolution mass

spectrometry using selected ion monitoring. A linear relationship

between amphetamine concentration and ion current response was established over the range 5 pg to 500 ng, and as little as 0.1 ng/ml

of amphetamine was detectable in urine. However, because the inter156
nal standard was of poor isotopic purity, reliability of the method

suffered at concentrations lower than 2 ng/ml (45).

In a GC/MS procedure involving quantitation of amphetamine as the

isothiocyanate derivative, amphetamine-2H5 and ß-methylphenethylamine were compared as internal standards. The two internal standards were reported to give comparable results (44).

A comparison of an RIA and a GC/MS assay for amphetamine showed

that the GC/MS method was about twice as sensitive and gave better precision than the RIA method, However, taking into consideration factors such as cost, time, and degree of training required to perform the assays, RIA performed in duplicate or triplicate was recommended for routine measurements. The measured concentrations in unknown samples by both techniques showed a good

correlation, lending confidence in the accuracy of both methods

(46).

The optical isomers of amphetamine can be separated by formation

of a N-trifluoroacetyl-L-prolyl derivative and separation of the

diastereoisomers by TLC. For qualitative purposes, such as forensic differentiation of illicit samples (47), the individual isomers

can be measured in urine (48). However, GC/MS, with its high inherent sensitivity, has provided the first means of studying the

stereochemical profile of amphetamine in human plasma and saliva

at ng/ml concentrations following oral administration of 10 mg

of racemic amphetamine. The drug, following extraction, was derivatized by reaction with a chiral agent, N-pentafluorobenzoyl-S-(-)-

prolyl-1-imidazole. The resulting diastereoisomers were easily resolved by capillary GC (5 percent OV-275 coated on Chromosorb W-AW)

and quantitated by chemical ionization mass spectrometry using methane as the reagent gas. Amphetamine-2H2

 served as the internal

standard (49). An analytical procedure developed for simultaneous

determination of the concentrations of amphetamine enantiomers in

rat urine also utilized GC/MS with selected ion monitoring, but

used the N-(S)- -methoxy- -(trifluoromethyl)phenylacetyl derivative

and amphetamine labeled with deuterium on the methyl group as the

internal standard (50).

A general method for extraction of amphetamine from biological materials involves adjustment of the pH to about 10-12 and subsequent

extraction with an organic solvent or elution from ion exchange columns. The latter is not as selective and the resulting extracts show

many extraneous peaks, so the solvent extraction method is preferred

for GC analysis. Solvents used have included chloroform (22) or

chloroform: isopropanol (96: 4) (24); pentane (28); diethyl ether, for

which recoveries of 70-75 percent from blood and 83 percent from

urine have been reported (26,32,51); hexane (89 percent from blood,

97 percent from urine) (21,49), or hexane:isooctane (75:25 v/v) (35);

and benzene (33,52). Amphetamine is about 15 percent bound to plasma

proteins, but both free and bound forms are extracted equally well,

suggesting that losses in recovery are mainly a consequence of the

drug’s high volatility. Mild conditions are therefore essential

for evaporation of the solvent (51).

157
EXPERIMENTAL PROCEDURE

The experimental procedure for extraction, derivatization, and GC/

MS analysis of amphetamine is identical to the procedure described

in Chapter 9 for analysis of methamphetamine except for the internal standard (amphetamine-2H3

 is used instead of methamphetamine2H5

) and the specific ions monitored. Consequently, the concentrations of both drugs can be measured simultaneously by adding deuterated analogs of the two drugs to the biological sample for use

as internal standards. Then during data acquisition, the ion currents at four m/z values are monitored. These are m/z 232, 235,

246, and 251, which correspond to the protonated molecule ions for

the trifluoroacetamide derivatives of amphetamine, amphetamine-2H3,

methamphetamine, and methamphetamine-2H5, respectively.

Standards and Reagents

d-Amphetamine hydrochloride can be purchased from Applied Science

Laboratories, State College, PA 16801, for preparation of standard

solutions. The amphetamine-2H3 hydrochloride used in the development of this procedure was synthesized by a literature procedure

(53). The deuterium-labeled amphetamine had an isotopic purity of

99.9 percent (

2H3

).

For the purpose of preparing calibration graphs and working standards,

stock solutions of amphetamine and amphetamine-2H3 are prepared as

follows. Weigh into a 100-ml volumetric flask about 12.6 mg of

amphetamine hydrochloride and record its weight to the nearest 0.1

mg The equivalent weight of free base is calculated by multiplying the weight of amphetamine hydrochloride by 0.79. Dissolve the

measured amphetamine in about 75 ml of methanol and then add additional methanol to give exactly 100 ml of solution. In the following

paragraphs this solution will be referred to as the “0.10-mg/ml stock

solution” although it should be labeled with its actual concentration based upon the exact measured weight of amphetamine hydrochloride. A series of working standard solutions can be prepared by

appropriate dilution of this stock solution as described in Chapter 2.

The stock solution of amphetamine-2H3 is prepared in the same manner.

For the measurement of amphetamine in body fluids within the concentration range 1 to 500 ng/ml, addition of approximately 40 ng of the

internal standard to each ml of specimen is satisfactory. This can

be conveniently done by adding 100 μl of a 400-ng/ml amphetamine2H3

 methanolic solution to each ml of specimen. Prepare the 400-ng/

ml internal standard solution by diluting 1 ml of the 0.10-mg/ml

stock solution of amphetamine-2H3 to 250 ml with methanol.

Store stock and standard solutions in well-stoppered or capped glass

vessels in the dark at less than 0°C.

Extraction

Transfer 1 ml of the specimen (whole blood, plasma or urine) to a

20-ml culture tube equipped with a Teflon-lined screw cap (0.5 to 2

ml of specimen can be used, but it must be measured accurately).

158
Add 100 μl of the 400-ng/ml amphetamine-2H3

 soiution and vortex

for about 10 sec. Allow the sample to equilibrate for about 15

min and then increase the pH to greater than 10 by adding 1 ml of

K2CO3-saturated distilled water. Extract the basic mixture with

5 ml of 1-chlorobutane by gently mixing the contents for 30 min

using a motorized rocker or rotator. Centrifuge for 5 min and then

transfer the organic phase (top) to another 20-ml culture tube by

means of a disposable Pasteur pipette. Extract the amphetamine

and other organic bases from the organic phase into aqueous acid

by adding 3 ml of 0.2 N H2SO4 and gently mixing for at least 15

min. Centrifuge and remove the top organic phase by aspiration.

Add 0.5 ml of 10 N NaOH to the aqueous phase, mix the contents, and

check the pH to make certain the solution is strongly basic (pH > 10).

Finally, reextract the organic bases from the aqueous alkaline solution by adding 5 ml of methylene chloride and gently mixing for at

least 15 min. Centrifuge for 5 min, aspirate off the upper aqueous

layer, and transfer by means of a disposable Pasteur pipette the

methylene chloride extract (bottom layer) to a silylated concentrator tube having a volume of at least 5 ml, a conical bottom, and a

Teflon-lined screw cap. Add 10 μl of dimethylformamide to act as

a “keeper solvent” to minimize evaporative loss of the amphetamine.

Remove the methylene chloride by evaporation under a gentle stream

of nitrogen or filtered air while heating the tube to no higher

than 40°C. When the volume of extract has decreased to about 10

μ1, add 15 μl of N-methyl bis- (trifluoroacetamide) “MBTFA” (Pierce

Chemical Co., Rockford, IL 61105). Cap the tube and heat it at

70°C for 15 min. The capped tube can be stored under refrigeration until the GC/MS analysis is ready to be performed. Immediately before the analysis, allow the tube to warm to room temperature.

GC/MS Analysis

The experimental conditions for the GC/MS analysis for amphetamine

trifluoroacetamide are as follows:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801).

Carrier and

reagent gas: Methane, 15-20 ml/min

Temperatures: Injector, 260°C

Column, 120°C, isothermal

GC/MS transfer line, 260°C

Ion source, 160°C

Under these conditions, the amphetamine trifluoroacetamide should

elute in 2 to 4 min as a narrow, symmetrical peak.

prior to beginning GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored occur at m/z 232 and

235. They correspond to the protonated molecule ions for ampheta159
tamine trifluoroacetamide and amphetamine-2H3

 trifluoroacetamide,

respectively. With the divert valve in the divert position, inject 2 to 6 μl of the organic extract into the gas chromatograph.

After approximately 1 min, switch the divert valve so that the entire carrier gas flow enters the ion source of the mass spectrometer, and begin data acquisition. When the amphetamine trifluoroacetamide peak has eluted, cease data acquisition and return the

divert valve to the divert position.

The quantity of amphetamine in the specimen is determined by measuring the heights (or areas) or the amphetamine trifluoroacetamide

and amphetamine-2H3 trifluoroacetamide peaks in the selected ion

current profiles (“ion chromatograms”) and relating the ratio of

peak heights to a calibration graph. Calculate the concentration of

amphetamine in the specimen by dividing the measured quantity of

amphetamine by the exact volume of specimen extracted.

Discussion of the Experimental Procedure

The electron impact (EI) and chemical ionization (CI) mass spectra

of amphetamine trifluoroacetamide are shown in Figure 1. The most

abundant ions in the CI mass spectra correspond to the protonated

molecule ion (m/z 232) and the ammonia-attachment ion (m/z 249), depending on whether methane alone or methane and ammonia are used

as reagent gases. Even though the procedure specifies use of methane alone as the reagent gas, the combination of methane and ammonia

is also satisfactory and should provide slightly better sensitivity.

Perhaps most important is the fact that the methane and ammonia combination offers an opportunity to monitor a different pair of ion

masses if interferences are encountered at m/z 232 or 235 when methane alone is used.

An additional option which can be explored if interferences are encountered in the selected ion monitoring measurements, is to generate a derivative different from the trifluoroacetamide. For example, the acetamide derivative could be prepared by substituting

N-acetylimidazole for the N-methyl bis-(trifluoroacetamide). Also,

the pentafluoroacetamide or heptafluorobutyramide derivatives could

be used. The trifluoroacetamide derivative has proven satisfactory,

but the use of other derivatives has not been fully evaluated.

A potential problem associated with concentration of organic extracts

containing amphetamine is the loss of the drug through evaporation

because of its relatively high volatility. For this reason, some

amphetamine assay procedures specify addition of an acid to convert

the drug to a nonvolatile salt prior to concentration of the extract.

In the procedure described here, losses due to evaporation are minimized by the following means. First, methylene chloride is used in

the final extraction because it has a lower boiling point (39°C)

than 1-chlorobutane (78°C). Second, a small quantity of dimethylformamide is added just prior to evaporative concentration to act

as a “keeper solvent.” If these procedures are followed and care

is taken not to allow the concentrated residue to remain at an elevated temperature for an extended period, no more than 20 percent of

the amphetamine should be lost due to evaporation.

160
FIGURE 1. MASS SPECTRA OF THE TRIFLUOROACETAMIDE

DERIVATIVE OF AMPHETAMINE

161
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR AMPHETAMINE

Nomenclature

Chemical Name: -Methylphenethylamine

Empirical Formula: C9H13N

Chemical Abstracts Registry Number: 300-62-9

Trade Names: Actedron, Allodene, Benzedrine, Adipan, Sympatedrine,

Psychoedrine, Isomyn, Isoamyne, Mecodrin, Norephedrone,

Novydrine, Elastonon, Ortedrine, Phenedrine, Profamina,

Propisamine, Sympamine, Simpatedrin

Physical Constants

Appearance: Colorless liquid

Boiling Point: 200-203° (760 mm); 82-85° (13 mm)

Solubility of the free base: Slightly soluble in water; soluble

in acid, alcohol, and ether

pKa: 9.77

Specific Rotation: d- -methylphenethylamine sulfate = +21.8°

(c=2)

REFFRENCES

1.

2.

3.

4.

5.

6.

7.

8.

9.

J.B. Hart and J. Wallace. Clin. Toxicol., 8, 179 (1975).

Unsigned. Brit. Med. J., 1, 476 (1975).

J.C. Kramer. J. Psychedelic Drugs, 2, 8 (1969).

M. Lampe. Drugs: Information for Crisis Treatment. STASH

Press, Beloit, WI, p. 9 (1972).

J.N. Lomonte, W.T. Lowry, and I.C. Stone. J. Forensic Sci., 2,

575 (1976).

J.H. Biel, in E. Costa and S. Garattini, eds., Amphetamines

and Related Compounds. Raven Press, New York, p. 3 1970).

A. Carlsson, in E. Costa and S. Garattini, eds., Amphetamines

and Related Compounds. Raven Press, New York, p. 289 (1970).

C. Carlsson, M. Hagerdal, and B.K. Siesjö. Acta Physiol.

Scand., 94, 128 (1975).

J. Axelrod, in E. Costa and S. Garattini, eds., Amphetamines

and Related Compounds. Raven Press, New York, p. 207 (1970).

162
10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

L.G. Dring, R.L. Smith, and R.T. Williams. J. Pharm.

Pharmacol., 18, 402 (1966).

J. Caldwell. Drug Metab. Revs., 5, 219 (1976).

A.H. Beckett, H.T. Coutts, and F.A. Ogunbona. J. Pharm.

Pharmacol., 25, 708 (1973).

A.H. Beckett and S. Al-Sarraj. J. Pharm. Pharmacol., 25,

328 (1973).

J.W. Gorrod. Life Sci., 13, lviii (1973).

A.H. Beckett, R.N. Boyes, and G.T. Tucker. J. Pharm. Pharmacol.,

20, 277 (1968).

M. Rowland. J. Pharm. Sci., 58, 508 (1969).

E. Änggård, L.-E. Jonsson, A.-L. Hogmark, and L.-M. Gunne.

Clin. Pharmacol. Ther., 14, 870 (1973).

A.K. Cho, B.J. Hodshon, B. Lindeke, and G.T. Miwa. J. Pharm.

Sci., 62, 1491 (1973).

M.P. Kullberg and J.D. Griffith. Fed. Proc., 35, 565 (1976).

E. Costa and S. Garattini, eds., Proc. Int. Symp Amphetamines

and Related Compounds. Raven Press, New York (1970).

J.E. Wallace, J.D. Biggs, and S.L. Ladd. Anal. Chem., 40,

2207 (1968).

T.S. Hayes. Clin. Chem., 19, 390 (1973).

K.J. Bussey and R.C. Backer. Clin. Chem., 2, 302 (1974).

B. Klein, J.E. Sheehan, and E. Grunberg. Clin. Chem., 20,

272 (1974).

J. Sherma, M.F. Dobbins, and J.C. Touchstone. J. Chromatogr.

Sci., 12, 300 (1974).

F. Van Hoof and A. Heyndrickx. Anal. Chem., 46, 286 (1974).

T.J. Hopen, R.C. Briner, H.G. Sadler, and R.L. Smith. J.

Forensic Sci., 21, 842 (1976).

D.S. Kreuz and J. Axelrod. Science, 183, 420 (1974).

A. Castro and H. Malkus. Res. Commun. Chem. Pathol. Pharmacol.,

16, 291 (1977).

R.O. Bost, C.A. Sutheimer, and I. Sunshine. Clin. Chem., 22,

789 (1976).

163
31. K.D.R. Setchell and J.D.H. Cooper. Clin. Chim. Acta, 35,

67 (1971).

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

J.E. O'Brien, W. Zazulak, V. Abbey, and O. Hinsvark. J. Chromatogr. Sci., 10, 336 (1972).

R.B. Bruce and W.R. Maynard, Jr. Anal. Chem., 41, 977 (1969).

J.S. Noonan, P.W. Murdick, and R.S. Ray. J. Pharmacol. Exp.

Ther., 168, 205 (1969).

P.A. Toseland and P.H. Scott. Clin. Chim. Acta, 25, 75 (1969).

G. Brugaard and K.E. Rasmussen. J. Chromatogr., 147, 476

(1978).

G.R. Wilkinson. Anal. Lett., 3, 289 (1970).

E. Änggård, in E. Costa and S. Garattini, eds., Amphetamines

and Related Compounds. Raven Press, New York, p. 191 (1970).

J.J. Franken and M.M.F. Trijbels. J. Chromatogr., 91, 425

(1974).

A.H. Beckett and R. Achari. J. Chromatogr., 135, 200 (1977).

A. Wu. Clin. Toxicol., 8, 225 (1975).

N. Narasimhachari and P. Vouros. Anal. Biochem., 45, 154

(1972).

N. Narasimhachari, R.O. Friedel, F. Schlemmer, and J.M. Davis.

J. Chromatogr., 164, 386 (1979).

T.J. Danielson and A.A. Boulton. Biomed. Mass Spectr., 1, 159

(1974).

K.H. Powers and M.H. Ebert. Biomed. Mass Spectr., 6, 187 (1979).

D. Eskes. J. Chromatog., 117, 442 (1976).

L.-M. Gunne. Biochem. Pharmacol., 16, 863 (1967).

S.B. Matin, S.H. Wan, and J.B. Knight. Biomed. Mass Spectr.,

4, 118 (1977).

J. Gal. Biomed. Mass Spectr., 5, 32 (1978).

D.B. Campbell. J. Pharm. Pharmacol., 21, 129 (1969).

A.K. Cho, B. Lindeke, B.J. Hodshon, and D.J. Jenden. Anal.

Chem., 45, 570 (1973).

B. Lindeke and A.K. Cho. Acta Pharm. Suecica, 9, 363 (1972).

164
Chapter 11

Methamphetamine

The first synthesis of methamphetamine ( d -N, -dimethylphenethylamine, I), a potent central nervous systerm stimulant, was reported

in 1919 (1). Methamphetamine has since been used clinically as an

analeptic to counteract alcohol, barbiturate, or narcotic stupor,

and also during surgery to maintain the blood pressure of patients

under anesthesia (2). Clinical use of the drug has declined in

recent years but methamphetamine continues to be abused. It is

available on the street as Methedrine tablets or as illicit preparations which are often injected intravenously. Like other stimulants of the amphetamine group, methamphetamine produces an initial

state of euphoria. However, this feeling is followed by restlessness, agitation, irritability and sometimes extreme paranoia, as

well as depression and exhaustion as the stimulant effects of the

drug wear off. In consequence there is a strong tendency to continue use in order to maintain the “high” and avoid the unpleasant

after-effects. Tolerance develops rapidly following repeated use

and increasing doses are required to maintain the original level

of stimulation. Physical dependence has not been proven, but the

likelihood of continued use is very high (3). Methamphetamine has

been found in human neonatal tissues, and the fetus rarely survives

to term when the mother is a methamphetamine abuser (4).

For forensic purposes, it may be necessary to use procedures which

can identify the stereoisomers, because 1-methamphetamine can be

found in the urine of persons using at least one of the common, nonprescription inhaler formulations. As in the case of amphetamine,

the isomers differ in their pharmacological activity; 1-methamphetamine has greater sympathomimetic properties while d-methamphetamine

has greater anorexic and stimulant properties (5).

A comparative study in rats showed that the N-methyl group rendered methamphetamine less toxic to liver microsomal enzyme systems

than amphetamine, but in these animals the methylated drug was more

active, and required a smaller dose to achieve the same pharmaco165
logical effect (6). Also in rats, methamphetamine was found to

cause a short-term (less than 6 hr) depletion of norepinephrine

and serotonin in brain and a long-lasting (more than 48 hr) depletion of heart norepinephrine. Amphetamine was detected in these

tissues after methamphetamine administration, as well as the p-hydroxyl derivative of amphetamine and one or more unidentified metabolite(s) represented by radioactivity. The latter, more polar

than the parent drug or amphetamine, persisted at higher concentrations in heart than in brain tissue, paralleling the time course

of norepinephrine depletion. Methamphetamine and amphetamine levels

were lower in heart than in brain and disappeared from heart tissue

within 5 hr (7).

METABOLISM AND PHARMACOKINETICS

A study of 14C-methamphetamine metabolism in man showed that about

90 percent of the 14C was excreted in the urine within four days.

An average of 22 percent of the dose was excreted in urine within

24 hr as unchanged drug, and 15 percent was excreted as 4-hydroxymethamphetamine (II).

Minor metabolites include 4-hydroxyamphetamine (III, 1 percent),

amphetamine (IV, 2-3 percent), 4-hydroxynorephedrine (V, 1-2 percent), norephedrine (VI, 2 percent), hippuric acid (5 percent),

and an acid-labile precursor of benzylmethyl ketone (1 percent).

An average of only 62 percent of the radioactivity ingested appeared

in urine within 24 hr (8). Radioactivity was not monitored in

sweat or saliva, although methamphetamine is known to be excreted

by these routes (9). Although its products account for only 4 percent of the dose in man, ß-hydroxylation is significant because norephedrine and 4-hydroxy norephedrine could act as false neurotransmitters (10). The hydroxylated derivatives of methamphetamine are excreted primarily as glucuronic acid conjugates. In

those metabolites where both parahydroxylation and N-demethylation

have occurred, some evidence indicates that the latter occurs first.

N-demethylation is to some degree stereospecific, occurring more

readily with the d-isomer of methamphetamine than with the l-isomer

(7,11).

166
Alteration of urinary pH has been show to affect the amount of

unchanged methamphetamine excreted in human urine. When urine was

metabolically acidified by ingestion of NH4Cl or alkalinized by ingestion of NaHC03

, 16-hr excretion of the parent drug was much greater in acid than in alkaline urine. Under alkaline urine conditions,

subjective effects of methamphetamine were more prolonged, indicating reabsorption and longer retention in the body (11).

Therapeutic doses of methamphetamine in man are relatively small,

5-10 mg. The drug is a moderately strong organic base (pKa = 9.89)

(11), and therefore blood concentrations are usually low (20-60 ng/

ml) although as much as 550 ng/ml has been reported in overdose

cases .

ANALYTICAL METHODOLOGY

A quantitative method employing thin-layer chromatography (TLC) of

urine extracts on alkaline silica gel plates, followed by spectrophotometric determination of a color reaction in the spots developed on TLC, was used to measure methamphetamine in the urine of

horses. The intensity of the color was directly proportional to

the quantity of drug over the range of 200-2000 μg (12). An improved

sensitivity (10 ng/ml) was achieved by ultraviolet spectrophotometry

of extracts from urine, serum, or tissue homogenates after oxidation with cerium sulfate, but the procedure was relatively nonspecific for methamphetamine (13).

A fluorometric technique, which could in principle be used for quantitation of methamphetamine if the drug were first purified chromatographically, has been used to measure basic amines in urine. The

drugs were converted to highly fluorescent compounds by reaction

with 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole (NBD chloride) (14).

This technique has in fact been used to screen for methamphetamine

in human urine, with specificity determined by the presence of twin

fluorescent spots on TLC, corresponding both in color and RF values

to the NBD derivatives of methamphetamine and its metabolite, amphetamine (15).

Because of the small amounts of methamphetamine found in blood,

improvements in both sensitivity and specificity of assay techniques

have been needed. A GC method using electron-capture detection (ECD)

167
of the heptafluorobutyramide derivative of the drug in blood extracts was sensitive to 20 ng/ml of methamphetamine, the amount

found in human blood 6 hr after a 5-mg oral dose (16). In another

application of GC-ECD in which therapeutic blood concentrations of

methamphetamine were monitored, the trichloroacetamide derivative

was prepared before chromatography on 3 percent OV-1. This derivative was superior in chromatographic characteris tics and more sensitive to ECD than either the heptafluorobutyramide or the pentafluorobenzamide. Blood concentrations over a 24-hr period were

found to range from 10 to 20 ng/ml following a 12.5-mg dose of

methamphetamine, but the method was capable of measuring amounts

as low as 25 pg (17).

Human urinary excretion rates of several amphetamine drugs, including methamphetamine, were followed by a GC procedure with the

addition of phenethylamine as an internal standard and the use of

a flame ionization detector (FID). The drug was chromatographed

as its acetamide derivative on a column packing coated with 8 percent ethylene glycol adipate, but the sensitivity was reported

only in terms of μg base/mg creatinine (18). In a similar method,

blood was first treated with tungstic acid to precipitate proteins,

and N-propylamphetamine was added as an internal standard. The

supernatant was made strongly basic for ether extraction of methamphetamine and the internal standard before conversion to the acetamide derivatives for GC-FID analysis. This method measured 30 ng/

ml methamphetamine in human blood 2 hr after ingestion of a single

10-mg dose (19).

For clinical use, a procedure to measure methamphetamine in urine by

GC-FID was &signed to eliminate the false positives caused by interfering substances and to shorten analysis time to 30 min. First,

the drug was determined as the free base on a 10 percent Apiezon L10 percent KOH column. Extracts found positive with reference to

GC peaks of standard solutions were then chromatographed on 3 percent OV-17, with on-column derivatization with trifluoroacetic anhydride. ß-Phenethylamine was the internal standard in both GC systems. The amount of methamphetamine was measured with an accuracy

of ±7 percent and the sensitivity was 0.1 μg/ml. The procedure

was adaptable to rapid analysis of a large number of samples (20,21).

Although chromatographic techniques have proved to be highly sensitive for methamphetamine, the addition of mass spectrometry is often

desirable for confirmation of its structure, even in routine analysis

such as monitoring in competitive sports. For confirmation purposes of this kind, the trifluoroacetylated derivative of methamphetamine was incorporated into a GC/MS protocol which detected 0.3

μg of methamphetamine/ml of human urine (22). GC/MS had been employed several years earlier as a method of identification, but not

quantitation, of methamphetamine in the urine of participants in

sporting events, with confirmation based on spectra of the N-acetyl

and N-propionyl derivatives (23). Mercuric acetate has been used

as a catalyst for rapid (30 sec at 80°C) quantitative formation of

trifluoroacetamide derivatives of methamphetamine and other phenylalkylamine drugs prior to GC/MS analysis (24).

168
Immunoassay techniques have been used in the screening of body fluids

for methamphetamine and have the advantages of being highly sensitive and requiring no extraction procedure. However, those in present use cannot distinguish between amphetamine and methamphetamine,

and they are subject to interference by other phenylethylamines (21).

Radioimmunoassay (RIA) did prove successful in detecting metabolites

of methamphetamine in rat saliva following injection of the drug.

Neither methamphetamine itself nor its 4-hydroxy derivative was

reactive in the RIA procedure, but the rat saliva gave positive

RIA reactions which resulted from amphetamine, 4-hydroxynorepnedrine, and 4-hydroxyamphetamine, either alone or in combination

(25). A commercial RIA system is available and can be used for

quantitative measurement of phenethylamines in both blood and

urine (26). However, its dynamic range is very limited and consequently it is primarily used as a screening technique with confirmation and quantitation achieved by other methods. The EMIT

technique is widely used for detection of phenethylamines in urine.

More than 70 tests per hr can be conducted with a detection level

of 2 μg/ml of urine (27).

Extraction solvents used for methamphetamine include diethyl ether

(19,22,23), n-hexane (13,17), pentane (16), cyclohexane (28), and

chloroform (14,15,18,21), under pH conditions ranging from 8 to 14.

Back extraction into acid is sometimes employed (21). Blood proteins can be precipitated and acid substances removed by treatment

with tungstic acid prior to extraction of methamphetamine at alkaline pH (17,19). A recovery of 100 percent has been reported for

extractions of the drug from blood with pentane (16) and from urine

with chloroform (18), both after alkalinization with NaOH.

An extensive review of the GC and GC/MS of amphetamines and their

derivatives has been published (29).

EXPERIMENTAL PROCEDURE

The blood concentrations of methamphetamine following normal dosages

are usually in the low ng/ml range. Consequently, the extraction

procedure includes a back extraction purification in order to provide a relatively clean extract, which in turn permits GC/MS measurement of the methamphetamine with very high sensitivity and little likelihood of interferences from endogenous components of the

blood. After addition of the deuterium-labeled internal standard,

the sample is made strongly basic with a saturated potassium carbonate solution and extracted with 1-chlorobutane. The methamphetamine and other organic bases are back extracted into dilute sulfuric acid, which is subsequently made basic with sodium hydroxide

and reextracted with 1-chlorobutane. The organic extract is then

treated with trifluoroacetic anhydride to convert methamphetamine

to its N-trifluoroacetamide derivative prior to concentration by

evaporation. Quantitation is achieved by selected ion monitoring

using chemical ionization with methane as the reagent gas.

Standards and Reagents

The d , 1-methamphetamine hydrochloride used in the development of

this method was acquired from the National Institute on Drug Abuse.

169
It had a melting point range of 170-172°C. No impurities were detected by either gas chromatography or mass spectrometry. d ,1-Methamphetamine-2H5

 hydrochloride was synthesized at Battelle Columbus

Laboratories using a published procedure (30). The location of

the deuterium atoms is indicated in structure VII. On the basis

of mass spectral analysis, its isotopic composition was: 2H5, 94

percent; 2H4, 5.7 percent; 2H3, 0.3 percent; 2H2, 

2H1

 and 2H0 less

than 0.1 percent.

For the purpose of preparing calibration graphs and working standards,

stock solutions of methamphetamine and methamphetamine-2H5 are prepared as follows. Weigh into a 100-ml volumetric flask about 12.4

mg of methamphetamine hydrochloride and record its weight to the

nearest 0.1 mg. The equivalent weight of free base is calculated

by multiplying the weight of methamphetamine hydrochloride by 0.805.

Dissolve the measured methamphetamine in about 75 ml of methanol

and then add additional methanol to give exactly 100 ml of solution.

In the following paragraphs this solution will be referred to as

the “0.10-mg/ml stock solution” although it should be labeled with

its actual concentration based upon the exact measured weight of

methamphetamine hydrochloride. A series of working standard solutions can be prepared by appropriate dilution of this stock solution as described in Chapter 2.

The stock solution of methamphetamine-2H5 is prepared in the sane

manner. For the measurement of methamphetamine in body fluids within the concentration range 1 to 500 ng/ml, addition of approximately

40 ng of the internal standard to each ml of specimen is satisfactory. This can be conveniently done by adding 100 μl of a 400-ng/ml

methamphetamine-2H5 methanolic solution to each ml of specimen. Prepare the 400-ng/ml internal standard solution by diluting 1 ml of

the 0.10-mg/ml stock solution of methamphetamine-2H5 to 250 ml with

methanol.

Store stock and standard solutions in well-stoppered or capped

glass vessels in the dark at less than 0°C.

Extraction

Transfer 1 ml of body fluid (whole blood, plasma or urine) to a 20-ml

culture tube equipped with a Teflon-lined screw cap (0.5 to 2 ml of

specimen can be used, but it must be measured accurately). Add 100

μl of the 400-ng/ml methamphetamine-2H5 solution and vortex for

170
about 10 sec. Allow the sample to equilibrate for about 15 min and

then increase the pH to greater than 10 by adding 1 ml of K2CO3-

saturated distilled water. Extract the basic mixture with 5 ml of

1-chlorobutane by gently mixing the contents for 30 min using a

motorized rocker or rotator. Centrifuge for 5 min and then transfer the organic phase (top) to another 20-ml culture tube by means

of a disposable Pasteur pipette. Extract the methamphetamine and

other organic bases from the organic phase into aqueous acid by

adding 3 ml of 0.2 N H2SO4 and gently mixing for at least 15 min.

Centrifuge and remove the top organic phase by aspiration. Add

0.5 ml of 10 N NaOH to the aqueous phase, mix the contents, and

check the pH to make certain the solution is strongly basic (pH> 10).

Finally, reextract the organic bases from the aqueous alkaline solution by adding 5 ml of methylene chloride and gently mixing for at

least 15 min. Centrifuge for five minutes, aspirate off the upper

aqueous layer, and transfer by means of a disposable Pasteur pipette

the methylene chloride extract (bottom layer) to a silylated concentrator tube having a volume of at least 5 ml, a conical bottom,

and a Teflon-lined screw cap. Add 10 μl of dimethylformamide to

act as a “keeper solvent” to minimize evaporative loss of the methamphetamine. Remove the methylene chloride by evaporation under a

gentle stream of nitrogen or filtered air while heating the tube to

no higher than 40°C. When the volume of extract has decreased to

about 10 μl, add 15 μl of N-methyl bis-(trifluoroacetamide) ‘MBTFA”

(Pierce Chemical Co., Rockford, IL 61105). Cap the tube and heat

it at 70°C for 15 min. The capped tube can be stored under refrigeration until the GC/MS analysis is ready to be performed. Immediately before the analysis, allow the tube to warm to room temperature .

GC/MS Analysis

The experimental conditions for the GC/MS analysis for methamphetamine trifluoroacetamide are as follows:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier and

reagent gas: Methane, 15-20 ml/min

Temperatures: Injector, 260°C

Column, 120°C, isothermal

GC/MS transfer line, 260°C

Ion source, 160°C

Under these conditions, the methamphetamine trifluoroacetamide should

elute in two to four minutes as a narrow, symmetrical peak.

Prior to beginning GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored occur at m/z 246 and

251. They correspond to the protonated molecule ions for methamphetamine trifluoroacetamide and methamphetamine-2H5

 trifluoroacetamide,

171
respectively. With the divert valve in the divert position, inject

2 to 6 μl of the organic extract into the gas chromatograph. After

approximately 1 min, switch the divert valve so that the entire

carrier gas flow enters the ion source of the mass spectrometer,

and begin data acquisition. When the methamphetamine trifluoroacetamide peak has eluted, cease data acquisition and return the

divert valve to the divert position.

The quantity of methamphetamine in the specimen is determined by

measuring the heights (or areas) or the methamphetamine trifluoroacetamide and methamphetamine-2H5

 trifluoroacetamide peaks in

the selected ion current profiles (“ion chromatograms”) and relating the ratio of peak heights to a calibration graph. Calculate the

concentration of methamphetamine in the specimen by dividing the

measured quantity of methamphetamine by the exact volume of specimen extracted.

Discussion of the Experimental Procedure

The procedure for analysis of amphetamine (Chapter 10) is virtually

identical to this procedure, except for the ions monitored. Consequently, the concentrations of both drugs can be measured simultaneously by adding deuterated analogs of the two drugs to the biological sample for use as internal standards. Then during data

acquisition, the ion currents at four m/z values are monitored.

These are m/z 232, 235, 246, and 251, which correspond to the protonated molecule ions for the trifluoroacetamide derivatives of

amphetamine, amphetamine-2H3, methamphetamine, and methamphetamine2H5

, respectively.

The electron impact (EI) and chemical ionization (CI) mass spectra

of metamphetamine trifluoroacetamide are shown in Figure 1. The

relative sensitivities achieved by monitoring the ion current at

the m/z value corresponding to the most abundant ion generated by

each of the ionization processes are shown in Table 1. In this

case, the highest response was obtained by monitoring the fragment

ion at m/z 154 generated by EI ionization. Nevertheless, CI should

give effective sensitivities at least as high as EI ionization because of the lower background usually experienced in CI. The most

abundant ions in the CI mass spectra correspond to the protonated

molecule ion (m/z 246) and the ammonia-attachment ion (m/z 263),

depending on whether methane alone or methane and ammonia are used

as reagent gases. Even though the procedure specifies use of methane alone as the reagent gas, the combination of methane and ammonia

is also satisfactory and should provide slightly better sensitivity.

Perhaps most important is the fact that the methane and ammonia combination offers an opportunity to monitor a different ion mass if

interferences are encountered at m/z 246 or 251 when methane alone

is used.

An additional option which can be explored if interferences are encountered in the selected ion monitoring measurements, is to generate a derivative different from the trifluoroacetamide. For example, the acetamide derivative could be prepared by substituting

N-acetylimidazole for the N-methyl bis-(trifluoroacetamide). Also

the pentafluoroacetamide or heptafluorobutyramide derivatives could

172
FIGURE 1. MASS SPECTRA OF METHAMPHETAMINE TRIFLUOROACETAMIDE

173
be used. The trifluoroacetamide derivative has proven satisfactory,

but the use of other derivatives has not been fully evaluated.

TABLE 1. RELATIVE RESPONSES FOR PROMINENT IONS IN THE

EI AND CI MASS SPECTRA OF THE TRIFLUOROACETAMIDE

DERIVATIVE OF METHAMPHETAMINE

% of Sample Relative Response Per

Method m/z Monitored Ion Current Unit Weight of Drug

EI 154 18 100

CI (CH4) 264 50 32

CI (CH4 + NH3) 263 72 59

A potential problem associated with concentration of organic extracts containing methamphetamine is the loss of the drug through

evaporation because of its relatively high volatility. For this

reason, some methamphetamine assay procedures specify addition of

an acid to convert the drug to a nonvolatile salt prior to concentration of the extract. In the procedure described here, losses

due to evaporation are minimized by the following means. First,

methylene chloride is used in the final extraction because it has

a lower boiling point (39°C) than 1-chlorobutane (78°C). Second,

the methamphetamine is converted to the less volatile trifluoroacetamide derivative prior to removing the solvent. Finally, a

small quantity of dimethylformamide is added just prior to evaporative concentration to act as a “keeper solvent.”

A comparison study showed that all of the organic solvents investigated except diethyl ether are effective in extracting methamphetamine from 2.5 N sodium hydroxide (Table 2). 1-Chlorobutane was

selected as the solvent to extract methamphetamine from biological

fluids. It represents a compromise of several factors including

extraction efficiency, solvent volatility, cleanliness of the extract, and ease of separation of the organic phase from the biological fluid. Diethyl ether, ethyl acetate, and ethylene chloride

were eliminated from consideration because of their relatively low

partition coefficients (4.3, 25.8, and 41.9, respectively). Chloroform:isopxopanol 9:1 (v/v), chloroform, and methylene chloride all

had high partition coefficients (82.8, > 90, and 78.5, respectively),

but they form the lower layer of an aqueous/organic two-phase system and it is therefore difficult to separate them from body fluids

using a pipette without introducing contamination from the aqueous

phase. 1-Chlorobutane, hexane, and cyclohexane all have relatively

high partition coefficients (70.3, 81.3, and 74.3, respectively)

and all form the upper layer of the aqueous/organic two-phase system. Of these, hexane was found to cause emulsion problems and

174
was eliminated from further consideration. Although 1-chlorobutane and cyclohexane have similar partition coefficients, 1-chlorobutane was selected for methamphetamine extraction based on previous successful experience using this solvent for drug extractions.

At pH 10, the percent of methamphetamine extracted from aqueous

solution reaches a maximum (about 97 percent) and this does not

significantly change at the higher pH region.

TABLE 2. PARTITION COEFFICIENTS FOR METHAMPHETAMINE

BETWEEN 2.5 N SODIUM HYDROXIDE AND

VARIOUS ORGANIC SOLVENTS (a)

Organic Solvent

Diethyl Ether

Ethyl Acetate

Chloroform:Isopropanol

9:1 (v/v)

Methylene Chloride

Chloroform

Ethylene Chloride

1-Chlorobutane

Partition Percent Methamphetamine

Coefficient in Organic Phase

4.3 81.1

25.8 96.3

82.8 98.8

78.5 98.7

90 100

41.9 97.7

70.3 98.6

Hexane 81.3 98.8

Cyclohexane 74.3 98.7

(a) Equal volumes of 2.5 N sodium hydroxide and organic phase

were used to determine the partition coefficient.

175
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR METHAMPHETAMINE

Nomenclature

Chemical Name: d-N, -Dimethylphenethylamine

Empirical Formula: C10H15N

Chemical Abstracts Registry Numbers: d -Methamphetamine: [537-46-2]

 d-Methamphetamine Hydrochloride: [51-57-0]

Trade Names: Adipex, Amphedroxyn, Desfedrin, Desoxyfed, Desoxyn,

Destim, Dexoxal, Doxephrin, Drinalfa, Efroxine,

Gerobit, Hiropon, Isophen, Madrine, Methedrine,

Methylisomyn, Pervitin, Semoxydrine, Soxysympamine,

Syndrox, Tonedron

Physical Constants

Melting Points: d-Methamphetamine Hydrochloride: 173-175°C

Specific Rotation: d-Methamphetamine = + 14° to + 20°(C = 1)

Solubility of d-Methaqhetamine Hydrochloride: 1 g/2 ml Water,

1 g/3 ml Ethanol, 1 g/5 ml Chloroform

Acid Dissociation Constant: d-Methamphetamine Hydrochloride pKa =

9.89

Refractive Index: d-Methamphetamine Hydrochloride = 1.5101

Boiling Point: d-Methamphetamine 212°C

REFERENCES

1. A. Ogata. J. Pharm. Soc. Japan, 451, 751 (1919).

2. T.J. Haley. J. Am. Pharm. Assn., 36, 161 (1947).

3. M. Lampe. Drugs: Information for Crisis Treatment. STASH

Press, Beloit, WI (1972).

4. J.C. Garriott and F.G. Spruill. J. Forensic Sci., 18, 434

(1973).

5. M.D. Solomon and J.A. Wright. Clin. Chem., 23, 1504 (1977).

6. G. Feuer, D.R. Miller, S.D. Cooper, F.A. de la Iglesia, and

G. Lumb. Int. J. Clin. Pharmacol., Ther., and Toxicol., 7,

13 (1973).

176
7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

C.D. Morgan, F. Cattabeni, and E. Costa. J. Pharmacol. Exp.

Ther., 180, 127 (1972).

J. Caldwell, L.G. Dring, and R.T. Williams. Biochem. J.,

129, 11 (1972).

T.B. Vree, T.J.M. Muskens, and J.M. Van Rossum. Arch. Int.

Pharmacodynamie, 199, 311 (1972).

H. Thoenen, A. Hürlimann, K.F. Gey, and W. Haefely. Life Sci.,

5, 1715 (1966).

A.H. Beckett and M. Kowland. J. Pharm. Pharmacol., 17,

suppl.. 109S (1965).

M.S. Karawya, M.A. El-Keiy, S.K. Wahba, and A.R. Kozman.

J. Pharm. Pharmacol., 20, 650 (1968).

J.E. Wallace, J.D. Biggs, and S.L. Ladd. Anal. Chem., 40,

2207 (1968).

J. Monforte, R.J. Bath, and I. Sunshine. Clin. Chem., 18,

1329 (1972).

R.N. Gupta, B.G. Chittim, and P.M. Keane. J. Chromatogr.

Sci., 12, 67 (1974).

R.B. Bruce and W.R. Maynard, Jr. Anal. Chem., 41, 977 (1969).

R.C. Driscoll, F.S. Barr, B.J. Gragg, and G.W. Moore. J.

Pharm. Sci., 60, 1492 (1971).

J.W. Schweitzer and A.J. Friedhoff. Clin. Chem., 16, 786

(1970).

P. Lebish, B.S. Finkle, and J.W. Brackett, Jr. Clin. Chem.,

16, 195 (1970).

N.C. Jain, T.C. Sneath, and R.D. Budd. Clin. Chem., 20,

1460 (1974).

N.C. Jain, R.D. Budd, and T.C. Sneath. Clin. Toxicol., 8,

211 (1975).

K. Kamei, M. Murata, K. Ishii, M. Namekata, and A. Momose.

Chem. Pharm. Bull., 21, 1996 (1973).

A.H. Beckett, G.T. Tucker, and A.C. Moffat. J. Pharm.

Pharmacol., 19, 273 (1967).

A. Wu. Clin. Toxicol., 8, 225 (1975).

G.J. Pigregorio and E.K. Kniaz. Drug and Alcohol Dependence,

1, 377 (1976).

177
26. Abuscreen. Roche Diagnostics, Nutley, NJ 07110.

27. EMIT. Syva, Palo Alto, CA 94304.

28. J.W. Blake, R.S. Ray, J.S. Noonan, and P.W. Murdick. Anal.

Chem., 46, 288 (1974).

29. B.J. Gudzinowicz and M.J. Gudzinowicz. Analysis of Drugs

and Metabolites by Gas Chromatography - Mass Spectrometry,

Vol. 4. Marcel Dekker, Inc., New York (1978).

30. A.F. Fentiman, Jr. and R.L. Foltz. J. Labelled Compounds

and Radiopharmaceuticals, 12, 69 (1976).

178
Chapter 12

2, 5-Dimethoxy-4-

methylamphetamine (DOM)

In the late 1960’s a street drug known to users as “STP” was extensively circulated in the United States as a hallucinogen (1).

The psychotomimetic component of the illicit preparations was identified as 1-(2,5-dimethoxy-4-methylphenyl)-2-aminopropane (I), a

structure more conveniently referred to as 2,5-dimethoxy-4-methylamphetamine or DOM, which had been developed as an experimental

compound by Dow Chemical Company. Early tests of its psychotropic

activity in mice showed behavioral responses very similar to those

evoked by mescaline, but which required only about one-fiftieth of

the dose for an equivalent response (2). Controlled dosage studies

in human volunteers indicated a hallucinogenic potency 100 times

greater than mescaline but only about one-thirtieth that of LSD.

Effects of the drug included elevation of pulse rate, systolic

blood pressure and temperature, as well as nausea, paresthesia,

and tremors. Psychoactivity began with moderate euphoria within

1-2 hr after administration of as little as 2 mg; and doses in

excess of 5 mg always produced marked hallucinogenic effects of

dose-related intensity which subsided within 8 hr (1).

The sensory and hallucinogenic effects of DOM are limited to the

R-(-) enantiomer (3). In vivo metabolism of DOM in rabbits is stereoselective (4), and in rabbit liver preparations S-(+)-DOM-2H6

is metabolized more rapidly than R-(-)-DOM by microsomal enzymes

(5). Several methoxyl and methyl analogs were prepared for a

study of comparative pharmacological activity in mice. These studies showed that DOM was almost as effective as amphetamine in

decreasing pentobarbital-induced sleep time, and that the 4-methyl

group was apparently essential for the psychotomimetic activity

of DOM while the 2-methoxyl group was not (6). DOM has no known

therapeutic value but has been used for research investigations

concerning model psychoses.

179
METABOLISM AND PHARMACOKINETICS

In vivo studies in rats, using radiolabeled tracers, showed that

the major metabolic pathway of DOM in that species is hydroxylation of the 4-methyl group to 1-(2,5-dimethoxy-4-hydrovthylphenyl)-2-aminopropane (II), followed by oxidation of II to 1-(2,

5-dimethoxy-4-carboxyphenyl)-2-aminopropane (III). Nearly 50 percent of the urinary radioactivity was present as II plus its glucuronide and sulfate conjugates. Unchanged DOM accounted for 8

percent of urinary radioactivity and III accounted for 28 percent.

1-(2,5-Dimethoxy-4-methylphenyl)-2-propanone (IV) was detected in

trace amounts. In feces, 83 percent of the radioactivity was due

to III; 6 percent was due to II; only a trace amount was due to

unchanged DOM (7). Identification of the metabolites was based on

comparison of TLC Rf values with those of synthesized standards

(8). The same metabolic pathways for DOM, including the minor one

involving oxidative deamination, have been demonstrated in rabbits

in vivo (4). N-Hydroxylation (V) and O-demethylation reactions

have been reported in rabbit liver microsomal preparations incubated with DOM (9). The O-demethylated derivative of DOM might be

of interest in light of speculation that a bis-O-demethylated metabolite could be oxidized to the corresponding p-hydroquinone

which in turn could cyclize to form 5-hydroxy-2,6-dimethylindole

in a manner analogous to that observed for 6-hydroxydopamine.

The 5-hydroxy-2,6-dimethylindole has been suggested as a possible

cause of hallucinogenic activity (10).

180
In the human study previously mentioned, about 20 percent of the

injected dose appeared in the urine within 24 hr as the unchanged

drug. The peak of urinary excretion, between 3 and 6 hr after administration, corresponded to the peak clinical effects (1). In

rats, excretion of radiolabeled DOM and its metabolites was nearly

complete within 24 hr (7). Plasma concentration of DOM monitored

over a 24-hr period in two 9-kg dogs given 2 mg and 4 mg of DOM

respectively, suggested a bi-phasic elimination process, with an

initial rapid decrease and a slower phase corresponding to a halflife of 1.5 hr (11).

ANALYTICAL METHODOLOGY

Even though DOM has been a recognized drug of abuse for at least

ten years, little effort appears to have been devoted to development of methods for identification and quantitation of the drug in

biological media for forensic or research purposes. In 1968, several authors reported data on the physical and chemical characteristics for DOM including its TLC Rf values and its ultraviolet,

infrared, nuclear magnetic resonance, and mass spectra. These analytical techniques were used for identification of the drug in confiscated illicit preparations (2,12). An ultraviolet spectrophotometric method for determination of DOM following oxidation by

cerium sulfate was reported (13). However, UV and IR spectra were

unsatisfactory for samples containing impurities because of inadequate specificity. Mass spectrometry was suggested as a reliable

means of identification of DOM in confiscated drugs (14).

A fluorometric method has been used for the analysis of unchanged

DOM in the urine of human volunteers during a study of hallucinogenic reactions (1). More recently, a highly sensitive radioimmunoassay (RIA) has been reported in which a radioiodinated derivative of DOM was used as the test antigen. The specificity of

the antibody, raised in rabbits by DOM conjugated to human serum

albumin, was such that structurally related molecules (i.e., amphetamine, mescaline, catecholamine metabolites, or N,N-diethyltryptamine) were several orders of magnitude less effective than

DOM as inhibitors (15).

In vivo metabolic studies in rats measured excretion of tritiumlabeled DOM and its metabolites in urine and in fecal homogenates

by liquid scintillation counting. Metabolites were extracted

with diethyl ether and identified by comparison of TLC Rf values

with those of synthesized standards. The major metabolites were

further characterized by gas chromatography with a flame-ionization detector (GC-FID) on columns of 3 percent OV-1 or 3 percent

SE-30 maintained at 160°C. Quantitation of individual metabolites

was done by scintillation counting of the appropriate spots cut

from paper chromatograms (7). Similarly, concentrations of urinary DOM and metabolites from rabbits receiving carbon-14 labeled

DOM were measured by isotope dilution analysis during a study of

stereoselective metabolism. DOM was extracted from the urine with

diethyl ether and recrystallized for measurement of radioactivity

(4).

181
In vitro studies of stereoselective metabolism have utilized directprobe chemical ionization mass spectrometry with deuterium-labeled

DOM as an internal standard for quantitation of DOM and its metabolites in rabbit liver preparations (5). In other metabolic

studies with rabbit liver microsomes, the technique of selected

ion monitoring was used to measure DOM metabolite formation, with

DOM-2H6 as substrate, and a synthetic N-hydroxylated metabolite

of DOM as the internal standard. The deuteratcd metabolite produced by the enzyme system was identified as its bis-(trifluoroacetyl) derivative by GC/MS. Electron impact mass spectra were

obtained following GC separation on a 2 percent Dexil 300 column

maintained at 160°C. GC separations prior to isobutane chemical

ionization mass spectrometry were done on 3 percent OV-25 or 3

percent OV-1 columns (9).

In published studies which involved extraction of DOM from biological specimens, the extraction solvent has usually been diethyl

ether, with no extraction efficiencies reported (4,7).

EXPERIMENTAL PROCEDURE

The procedure for extraction, derivatization, and GC/MS quantitation of DOM is the same as that described for the analysis of methamphetamine (Chapter 9) and the other phenylalkylamine drugs (i.e.,

amphetamine and mescaline). After addition of the deuterium-labeled

internal standard, the sample is made strongly basic with a saturated potassium carbonate solution and extracted with 1-chlorobutane. The DOM and other organic bases arc back extracted into dilute sulfuric acid, which is subsequently made basic with sodium

hydroxide and reextracted with 1-chlorobutane. The organic extract

is then treated with trifluoroacetic anhydride to convert DOM to

its N-trifluoroacetamide derivative prior to concentration by evaporation. Quantitation is achieved by selected ion monitoring using

chemical ionization with methane as the reagent gas.

Standards and Reagents

The DOM•HCl used in the validation of this procedure was obtained

from the National Institute on Drug Abuse. It had a melting point

of 187-189°C. The DOM•HCl can be purchased from Applied Science

Laboratories, State College, PA 16801. DOM-2H6•HCl was synthesized

at Battelle Columbus Laboratories using a published procedure (15).

On the basis of mass spectral analysis the DOM-2H6 had the following isotopic composition: 2H6

 96.1 percent; 2H5, 3.1 percent; 2H4,

0.6 percent; 2H3, 0.2 percent; 2H2, 

2H1

, and 2H0, <0.1 percent.

For the purpose of preparing calibration graphs and working standards,

stock solutions of DOM and DOM-2H6 are prepared as follows. Weigh

into a 100-ml volumetric flask about 11.7 mg of DOM•HCl and record

its weight to the nearest 0.1 mg. The equivalent weight of free

base is calculated by multiplying the weight of DOM•HCl by 0.85.

Dissolve the measured DOM in about 75 ml of methanol and then add

additional methanol to give exactly 100 ml of solution. In the

following paragraphs this solution will be referred to as the “0.10-

mg/ml stock solution” although it should be labeled with its actual

182
concentration based upon the exact measured weight of DOM hydrochloride. A series of working standard solutions can be prepared by

appropriate dilution of this stock solution as described in Chapter 2.

The stock solution of DOM-2H6 is prepared in the same manner. For

the measurement of DOM in body fluids within the concentration

range 1 to 500 ng/ml, addition of approximately 40 ng of the internal standard to each ml of specimen is satisfactory. This can be

conveniently done by adding 100 μl of a 400-ng/ml DOM-2H6 methanolic solution to each ml of specimen. Prepare the 400-ng/ml internal standard solution by diluting 1 ml of the 0.10-mg/ml stock solution of DOM-2H6

 to 250 ml with methanol.

Store stock and standard solutions in well-stoppered or capped

glass vessels in the dark at less than 0°C.

Extraction

Transfer 1 ml of body fluid (whole blood, plasma or urine) to a

20-ml culture tube equipped with a Teflon-lined screw cap (0.5 to

2 ml of specimen can be used, but it must be measured accurately).

Add 100 μl of the 400-ng/ml DOM2H6 solution and vortex for about

10 sec. Allow the sample to equilibrate for about 15 min and then

increase the pH to greater than 10 by adding 1 ml of K2CO3- Saturated distilled water. Extract the basic mixture with 5 ml of lchlorobutane by gently mixing the contents for 30 min using a motorized rocker or rotator. Centrifuge for 5 min and then transfer

the organic phase (top) to another 20-ml culture tube by means of

a disposable Pasteur pipette. Extract the DOM and other organic

bases from the organic phase into aqueous acid by adding 3 ml of

0.2 N H2SO4 and gently mixing for at least 15 min. Centrifuge and

remove the top organic phase by aspiration. Add 0.5 ml of 10 N

NaOH to the aqueous phase, mix the contents, and check the pH to

make certain the solution is strongly basic (pH > 10). Finally,

rcextract the organic bases from the aqueous alkaline solution by

adding 5 ml of methylene chloride and gently mixing for at least

15 min. Centrifuge for 5 min, aspirate off the upper aqueous layer,

and transfer by means of a disposable Pasteur pipette the methylene

chloride extract (bottom layer) to a silylated concentrator tube

having a volume of at least 5 ml, a conical bottom, and a Teflonlined screw cap. Add 10 μl of dimethylformamide to act as a

“keeper solvent” to minimize evaporative loss of the DOM. Remove

the methylene chloride by evaporation under a gentle stream of

nitrogen or filtered air while heating the tube to no higher than

40°C. When the volume of extract has decreased to about 10 μl,

add 15 μl of N-methyl bis-(trifluoroacetamide) “MBTFA” (Pierce

Chemical Co., Rockford, IL 61105). Cap the tube and heat it to

70°C for 15 min. The capped tube can be stored under refrigeration until the GC/MS analysis is ready to be performed. Immediately before the analysis, allow the tube to warm to room temperature.

183
GC/MS Analysis

The experimental conditions for the GC/MS analysis for DOM trifluoroacetamide are as follows:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science laboratory, State College,

PA 16801)

Carrier and

reagent gas: Methane, 15-20 ml/min

Temperatures: Injector, 260°C

Column, 165°C, isothermal

GC/MS transfer line, 260°C

Ion source, 160°C

Under these conditions, the DOM trifluoroacetamide should elute

at about 4 min as a narrow, symmetrical peak.

Prior to beginning GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored occur at m/z 306

and 312. They correspond to the protonated molecule ions for DOM

trifluoroacetamide and DOM-2H6 trifluoroacetamide, respectively.

With the divert valve in the divert position, inject 2 to 6 μl of

the organic extract into the gas chromatograph. After approximately

1 min, switch the divert valve so that the entire carrier gas flow

enters the ion source of the mass spectrometer, and begin data acquisition. When the DOM trifluoroacetamide peak has eluted, cease

data acquisition and return the divert valve to the divert position.

The quantity of DOM in the specimen is determined by measuring the

heights (or areas) or the DOM. trifluoroacetamide and DOM-2H6 trifluoroacetamide peaks in the selected ion current profiles (“ion

chromatograms”) and relating the ratio of peak heights to a calibration graph. Calculate the concentration of DOM in the specimen

by dividing the measured quantity of DOM by the exact volume of

specimen extracted.

Discussion of the Experimental Procedure

The methane chemical ionization mass spectrum of the trifluoroacetamide derivative of DOM is shown in Figure 1. In addition to the

protonated molecule ion at m/z 306, there is also an abundant fragment ion (m/z 193) which is formed by loss of H2NCOCF3

 from the

protonated molecule ion. These ions are shifted to m/z 312 and

199 in the methane CI mass spectrum of DOM-2H6. Normally only the

ion currents at m/z 306 and 312 are monitored during GC/MS analysis,

but the ion currents at m/z 193 and 199 can also be monitored if

additional specificity is desired. The slope of the calibration graph

obtained by measuring the m/z 193/199 ratio for samples containing

known concentrations of DOM and DOM-2H6 may be slightly different

184
FIGURE 1. METHANE CI MASS SPECTRUM OF THE TRIFLUOROACETAMIDE DERIVATIVE OF DOM

185
from the slope obtained for the m/z 306/312 ratio. However, if

the concentration of DOM calculated from the m/z 193/199, peak

height ratio differs significantly from the concentration based

on the measured m/z 306/312 peak height ratio, it is likely that

an interfering compound is contributing to one or more of the DOM•

TFA peaks in the ion current profiles. When this occurs, usually

a visual examination of the ion current profiles will disclose

which of the peak heights is inaccurate due to the interference;

i.e. the peak will have a shoulder, or will be perceptibly broadened, or will have an abnormal retention time.

186
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR DOM

Nomenclature

Chemical Name: 2,5-Dimethoxy-4-methylamphetamine

Empirical Formula: C12H19NO2

Chemical Abstracts Registry Numbcr: 15588-95-1

Physical Constants

Appearance: White crystals

Melting Point: 61°C

Hydrochloride, 190°C

Specific Rotation: Synthetic material is usually racemic

Solubility: Insoluble in water, soluble in chloroform

W Absorption: m a x = 214, 225 and 288 nm (in dilute HCl)

REFERENCES

1.

2.

3.

4.

5.

6.

7.

8.

9.

10.

S.H. Snyder, L. Faillace, L. Hollister. Science, 158, 669

(1967).

G.F. Phillips and R.J. Mesley. J. Pharm. Pharmacol., 21, 9

(1969).

A.T. Shulgin. J. Pharm. Pharmacol., 25, 271 (1973).

S.B. Matin, P.S. Callery, J.S. Zwieg, A. O'Brien, R. Rapoport,

and N. Castagnoli, Jr. J. Med. Chem., 17, 877 (1974).

K.J. Weinkam, P. Callery, J. Gal, and N. Castagnoli, Jr. 22nd

Ann. Conf. MS and Allied Topics, Philadelphia. Abst., p. 174

May 19-24, 1974.

B.T. Ho, W.M. McIsaac, R. An, L.W. Tansey, K.E. Walker, L.F.

Englert, Jr., and M.B. Noel. J. Med. Chem., 13, 26 (1970).

B.T. Ho, V. Estevez, L.W. Tansey, L.F. Englert, P.J. Creaven,

and W.M. McIsaac. J. Med. Chem., 14, 158 (1971).

B.T. Ho and L.W. Tansey. J. Med. Chem., 14, 156 (1971).

J. Gal, L.D. Gruenke, and N. Castagnoli, Jr. J. Med. Chem.,

18, 683 (1975).

J.S. Zweig and N. Castagnoli, Jr. J. Med. Chem., 17, 747

(1974).

187

328-011 0 - 80 - 13
11. R.J. Martin and T.G. Alexander. J. Assoc. Off. Anal. Chem.,

51, 159 (1968).

12. J.F. Wallace, J.D. Biggs, and S.L. Ladd. Anal. Chem., 40,

2207 (1968).

13. S.W. Bellman. J. Assoc. Off. Anal. Chem., 51, 164 (1968).

14. L.J. Riceberg, H. van Vunakis, and L. Levine. Anal. Biochem.,

60, 551 (1974).

15. A.F. Fentiman, Jr., and R.L. Foltz. J. labelled Compounds

and Radiopharmaceuticals, 12, 69 (1976).

188
Chapter 13

Mescaline

Mescaline (3,4,5-trimethoxyphenylethylamine, I) is the hallucinogenic constituent of the peyote cactus Lophophora williamsii, a

plant which has been employed by Indians in northern Mexico from

the earliest recorded time as an adjunct to religious rites, to

relieve hunger and fatigue, and for treatment of disease (1). In

1889, a tincture of the peyote buttons appeared in the Parke-Davis

catalog as a cardiac and respiratory stimulant for treatment of

angina pectoris and pneumothorax. Mescaline was first isolated

from peyote in 1896 but a complete synthesis was not reported until

1918. During the 1940’s mescaline was one of several hallucinogens

used in research concerning “model” psychoses (1). At present the

use of mescaline, except in a religious context by certain North

American Indians, is primarily as an illicit hallucinogen. However,

street drugs sold as mescaline often contain little or none of the

drug. Instead they may contain amphetamines, DOM, belladonna alkaloids, LSD, or phencyclidine (PCP) as the active ingredient.

Mescaline, unlike most other hallucinogenic drugs which are indole

compounds, is chemically reiated to the hormone neurotransmitters,

epinephrine and norepinephrine. Pharmacological effects vary among

animal species. Physical dependence has not been demonstrated in

man, although repeated doses of mescaline cause tolerance to the

physiological, subjective, and psychological effects of the drug,

within 3 to 6 days. Cross tolerance to these effects has been

demonstrated between mescaline and LSD or psilocybin. Certain drugs,

such as insulin and barbiturates, potentiate the toxicity of mescaline. Mescaline poisoning may cause abdominal cramps, nausea, vomiting, and other symptoms which can be mistaken for acute gastroenteritis, appendicitis, or pancreatitis (2). An hallucinogenic dose of

mescaline is about 500-1500 mg (1). Distribution studies of mescaline-14C in cat brains showed the highest concentration of radioactivity in cortical and subcortical grey matter (3). In vitro experiments with sheep umbilical vasculature (4) and in vivo studies of rat

responses (5) have shown that antagonists of serotonin also antagonize the action of mescaline, supporting the hypothesis that mesca189
line initiates its pharmacological activity via serotonin receptors.

Efforts to synthesize specific antagonists to mescaline have led to

production of the N-allyl and N-n-propyl derivatives of mescaline

which alone have no effect on the swim-maze behavior of mice but

do antagonize the effect of mescaline on mouse behavior. No correlation has been demonstrated, however, between the swim-maze effects of mescaline in mice and psychotomimetic potency in humans

(6) .

METABOLISM AND PHARMACOKINETICS

Mescaline metabolism has been studied in several species, stimulated

by evidence that the hallucinogenic properties of mescaline are actually caused by one or more of its metabolites. This is suggested

because the psychotomimetic effects typically take several hours to

reach their maximum and the hallucinogenic dose is so large. Cases

of “flashbacks” over a period of several months following a single

dose of mescaline have also been reported (7). The major urinary

metabolic product in mammals is 3,4,5-trimethoxyphenylacetic acid

(II) produced by oxidative deamination by way of the corresponding

aldehyde. The aldehyde can also be reduced to 3,4,5-trimethoxyethanol, which has been detected in rat urine and brain, although the

aldehyde intermediate itself has not been isolated (8). The enzyme

involved in deamination differs from monoamine oxidase in that it

is cyanide-sensitive and resistant to octanol, and it has been characterized as a type-II enzyme (diamine oxidase) (9,10). However,

mescaline is not extensively metabolized in man or in laboratory

animals. Metabolic studies with mescaline-14C in human subjects

showed that 55-60 percent of the drug was excreted unchanged in the

urine. Trimethoxyphenylacetic acid accounted for 27-30 percent of

urinary radioactivity, while N-acetyl-ß-(3,4-dimethoxy-5-hydroxyphenyl)

ethylamine (III), in the form of a conjugate, represented 5 percent

and N-acetylmescaline (IV) less than 1 percent. The proportions

varied with time following administration. Five other minor metabolites were only partially characterized (11). Other studies have

shown the presence of 3,4-dimethoxy-5-hydroxyphenethylamine (V) and

the glutamine conjugate of 3,4-dihydroxy-5-methoxyphenylacetic acid

(VI) in human urine following administration of mescaline (7,12).

Compounds I, II, III, and IV were also found in human cerebrospinal

fluid (CSF) after oral administration of mescaline-14C. Mescaline

represented at least 50 percent of the CSF radioactivity, and the

concentration of radioactivity in the CSF appeared to coincide with

the peak of behavioral manifestations. The average biological halflife of radioactivity ingested as mescaline in this study was 6 hr,

with 87 percent of the dose excreted within 24 hr and 92 percent

within 48 hr (11). Neither trimethoxyphenylacetic acid nor N-acetylmescaline, in doses as high as 12 mg/kg, had any physiological or

psychological effects in human subjects, indicating that N-acetylation and deamination are both detoxification pathways for mescaline. The discrepancy in the relative amount of trimethoxyphenylacetic acid and N-acetylmescaline in CSF as compared to urine suggested that in the central nervous system N-acetylation is a more

important pathway (11). N-Acetylmescaline has been identified in

190
the CNS of mice (13) and of rats, in which acetylation appeared to

occur predominantly in the soluble cell fractions (14). 0-Demethylation of mescaline has been reported; however, 0-demethylation can

occur spontaneously under acidic conditions, which casts doubt on

the authenticity of these urinary metabolites (12).

In the continuing effort to identify the mechanism of hallucinogenic

activity of mescaline, rats trained for discriminative responses

were given inhibitors of aldehyde dehydrogenase or amine oxidase

before treatment with mescaline or three of its major rat metabolites. A role in mescaline-induced interoceptive stimuli could not

be established for any of the metabolites (8); however, in mouse

brain, side-chain degradation of mescaline to 3,4,5-trimethoxybenzoic acid has been demonstrated in vivo and in vitro, catalyzed by

enzymes in the nuclear and microsomal fractions of brain tissue (15).

The authors of this report suggest that complete elucidation of the

mechanism of the oxidative degradation of mescaline to 3,4,5-trimethoxybenzoic acid could provide a basis for disclosure of the assumed

relationship between the hallucinogenic properties of mescaline and

its metabolism.

ANALYTICAL METHODOLOGY

Thin-layer chromatography (TLC) and gas chromatography (GC) have

been the methods used in most studies for isolation and identification of mescaline in biological fluids or plant materials (16,17).

For TLC, mescaline extracted from urine can be developed in ethyl

191
acetate:methanol:NH40H (85:5:5) and sprayed with ninhydrin before

irradiation under 350 nm light, or in chloroform:methanol (9:1)

treated with Marquis reagent. The detection limit reported for TLC

is 40 ng/ml. GC conditions recommended for confirmation of mescaline in urine extracts are 3 percent SE-30 on Chromosorb W-AW 100/

200 mesh in a Teflon-lined aluminum column, and an oven temperature

of 150°C (2). A recent study of mescaline metabolism in mouse brain

tissue used mass spectrometry to confirm the identification of 3,4,5-

trimethoxybenzoic acid as its brominated derivative after TLC separation (15). A quantitative radioimmunoassay, which can distinguish between mescaline and its most common urinary metabolite, trimethoxyphenylacetic acid, has been reported. The assay detected as

little as 100 pg of mescaline (18).

A very sensitive GC/MS method for quantitation of mescaline in blood

or urine has been reported (19). Blood plasma was made basic (pH 10)

and extracted with benzene. The extracted mescaline was converted

to its trifluoroacetyl derivative which was measured by GC/MS using

a 2.5 percent OV-1 on Varaport column packing and a column temperature of 195°C. Deuterated mescaline was used as an internal standard. The limit of detection was reported to be 0.5 ng injected on

column.

Mescaline can be separated from its acid metabolites in urine by

extraction of the latter from acid solution with ethyl acetate. Mescaline remains in the aqueous phase and can be quantitatively extracted with benzene after the solution is made strongly basic (11).

Mescaline extraction from urine at pH 8.5 in n-butanol has also been

reported, with a recovery of 65 percent after back-extraction into

0.1N HCl (20). There are also reports of mescaline extraction from

urine buffered at pH 9.2 using chloroform or chloroform:isopropanol

(3:1) (2,3) and with ethylene dichloride:isoamyl alcohol at pH 10

(12)

EXPERIMENTAL PROCEDURE

The procedure for extraction, derivatization, and GC/MS quantitation

of mescaline is the same as that described for the analysis of methamphetamine (Chapter 11) and the other phenylalkylamine drugs (i.e.,

amphetamine and DOM). After addition of the deuterium-labeled internal standard, the sample is made strongly basic with a saturated potassium carbonate solution and extracted with 1-chlorobutane. The

mescaline and other organic bases are back-extracted into dilute sulfuric acid, which is subsequently made basic with sodium hydroxide

and reextracted with 1-chlorobutane. The organic extract is then

treated with trifluoroacetic anhydride to convert mescaline to its

N-trifluoroacetamide derivative prior to concentration by evaporation.

Quantitation is achieved by selected ion monitoring using chemical

ionization with methane as the reagent gas.

Standards and Reagents

The mescaline hydrochloride (m.p. 183-184) and the mescaline-2H3 hydrochloride (m.p. 179-182) used in the validation of this procedure

were obtained from the National Institute on Drug Abuse. No impuri192
ties were detected in either compound by gas chromatography. The

mescaline-2H3 HCl was further examined by thin-layer chromatography

(silica gel plates, 1.5 percent ammonia in methanol, Rf = 0.19)

and found to give a single spot. The three deuterium atoms in mescaline-2H3

 were located on the 4-methoxy group. Its isotopic com2H0

, < 1 percent.

position was: 2H3, 93 percent; 2H2, 3 percent; 2H1, 3 percent;

For the purpose of preparing calibration graphs and working standards

stock solutions of mescaline and mescaline-2H3 are prepared as follows. Weigh into a 100-ml volumetric flask about 11.7 mg of mescaline hydrochloride and record its weight to the nearest 0.1 mg.

The equivalent weight of free base is calculated by multiplying the

weight of mescaline hydrochloride by 0.85. Dissolve the measured

mescaline in about 75 ml of methanol and then add additional methanol to give exactly 100 ml of solution. In the following paragraphs

this solution will be referred to as the “0.10-mg/ml stock solution”

although it should be labeled with its actual concentration based

upon the exact measured weight of mescaline hydrochloride. A series

of working standard solutions can be prepared by appropriate dilution of this stock solution as described in Chapter 2.

The stock solution of mescaline-2H3 is prepared in the same manner.

For the measurement of mescaline in body fluids within the concentration range 1 to 500 ng/ml, addition of approximately 40 ng of

the internal standard to each ml of specimen is satisfactory. This

can be conveniently done by adding 100 μl of a 400-ng/ml mescaline

methanolic solution to each ml of specimen. Prepare the 400-ng/ml

internal standard solution by diluting 1 ml of the 0.10-ng/ml stock

solution of mescaline-2H3 to 250 ml with methanol.

Store stock and standard solutions in well-stoppered or capped glass

vessels in the dark at less than 0°C.

Extraction

Transfer 1 ml of body fluid (whole blood, plasma or urine) to a 20-ml

culture tube equipped with a Teflon-lined screw cap (0.5 to 2 ml of

specimen can be used, but it must be measured accurately). Add 100

μl of the 400-ng/ml mescaline-2H3 solution and vortex for about 10

sec. Allow the sample to equilibrate for about 15 min and then increase the pH to greater than 10 by adding 1 ml of K2CO3

-saturated

distilled water. Extract the basic mixture with 5 ml of 1-chlorobutane by gently mixing the contents for 30 min using a motorized

rocker or rotator. Centrifuge for 5 min and then transfer the organic phase (top) to another 20-ml culture tube by means of a disposable Pasteur pipette. Extract the mescaline and other organic

bases from the organic phase into aqueous acid by adding 3 ml of 0.2

N H2SO4 and gently mixing for at least 15 min. Centrifuge and remove the top organic phase by aspiration. Add 0.5 ml of 10 N Na0H

to the aqueous phase, mix the contents, and check the pH to make

certain the solution is strongly basic (pH > 10). Finally, reextract the organic bases from the aqueous alkaline solution by adding 5 ml of methylene chloride and gently mixing for at least 15 min.

193
Centrifuge for 5 min, aspirate off the upper aqueous layer, and

transfer by means of a disposable Pasteur pipette the methylene

chloride extract (bottom layer) to a silylated concentrator tube

having a volume of at least 5 ml, a conical bottom, and a Teflonlined screw cap. Add 10 μl of dimethylformamide to act as a

“keeper solvent” to minimize evaporative loss of the mescaline.

Remove the methylene chloride by evaporation under a gentle stream

of nitrogen or filtered air while heating the tube to no higher

than 40°C. When the volume of extract has decreased to about 10

μl, add 15 μl of N-methyl bis-(trifluoroacetamide) “MBTFA” (Pierce

Chemical Co., Rockford, IL 61105). Cap the tube and heat it at

70°C for 15 min. The capped tube can be stored under refrigeration until the GC/MS analysis is ready to be performed. Immediately before the analysis, allow the tube to warm to room temperature.

GC/MS Analysis

The experimental conditions for the GC/MS analysis for mescaline

trifluoroacetamide are as follows:

GC column: 1.8 m x 2 mm (I.D.) glass column packed with

3 percent OV-17 on 100/120 mesh Gas Chrom Q

(Applied Science Laboratory, State College,

PA 16801)

Carrier and

reagent gas: Methane, 15-20 ml/min

Temperatures: Injector, 260°C

Column, 160°C, isothermal

GC/MS transfer line, 260°C

Ion source, 160°C

Under these conditions, the mescaline trifluoroacetamide should

elute at about 4 min as a narrow, symmetrical peak.

Prior to beginning GC/MS analysis of samples, the performance of

the total GC/MS system should be evaluated and optimized as described in Chapter 2. The ions to be monitored occur at m/z 308

and 311. They correspond to the protonated molecule ions for mescaline trifluoroacetamide and mescaline-2H3

 trifluoroacetamide,

respectively. With the divert valve in the divert position, inject 2 to 6 μl of the organic extract into the gas chromatograph.

After approximately 1 min, switch the divert valve so that the

entire carrier gas flow enters the ion source of the mass spectrometer, and begin data acquisition. When the mescaline trifluoroacetamide peak has eluted, cease data acquisition and return the

divert valve to the divert position.

The quantity of mescaline in the specimen is determined by measuring the heights (or areas) or the mescaline trifluoroacetamide

and mescaline-2H3 trifluoroacetamide peaks in the selected ion

current profiles (“ion chromatograms”) and relating the ratio

of peak heights to a calibration graph. Calculate the concentration

of mescaline in the specimen by dividing the measured quantity of

mescaline by the exact volume of specimen extracted.

194
Discussion of the Experimental Procedure

The methane chemical ionization mass spectrum of the trifluoroacetamide derivative of mescaline is shown in Figure 1. The only

abundant ion in the mass spectrum is the protonated molecule ion

at m/z 308. The fragment ion formed by loss of H2NCOCF3

 from the

protonated molecule ion is considerably less abundant than it is

in the methane CI mass spectra of the other phenylalkylamine drugs

included in this monograph. This is probably because the amide

nitrogen is attached to a primary carbon, rather than a secondary

carbon as is the case in amphetamine, methamphetamine, and DOM.

Elimination of H2NCOCF3 from a secondary carbon would lead to a

more stable carbonium ion.

195
FIGURE 1. METHANE CI MASS SPECTRUM OF: THE TRIFLUOROACETAMIDE DERIVATIVE OF MESCALINE

196
PHYSICAL, CHEMICAL AND SPECTROMETRIC DATA FOR MESCALINE

Nomenclature

Chemical Name: 3,4,5-Trimethoxybenzenethanamine or

3,4,5-Trimethoxyphenethylamine

Empirical Formula: C11H17NO3

Chemical Abstracts Registry Number: 54-04-6

Physical Constants

Appearance: White crystals

Melting Point: 35-36°C

Hydrochloride, 181°C

Boiling Point: 180° (12mm)

Specific Rotation: Optically inactive

Solubility: Moderately soluble in water; soluble in alcohol,

chloroform, benzene; almost insoluble in ether

and petroleum ether

UV Absorption: max = 268( = 30)

REFERENCES

1.

2.

3.

4.

5.

6.

7.

8.

Staff of Student Association for the Study of Hallucinogens,

Mescaline and Peyote: A Literature Review. STASH Press,

Madison, WI (1974).

D.T. Teitelbaum and D.C. Wingeleth. J. Anal. Toxicol., 1,

36 (1977).

N. Neff, G.V. Rossi, G.D. Chase, and J.L. Rabinowitz. J.

Pharmacol. Exp. Ther., 144, 1 (1964).

D.C. Dyer, R. Benington, and R.D. Morin. Arch. Int. Pharmacodyn. Ther., 217, 197 (1975).

J.C. Winter. Arch. Int. Pharmacodyn. Ther., 214, 250 (1975).

F. DeSantis, Jr. and K.A. Nieforth. J. Pharm. Sci., 65,

1479 (1976).

J. Harley-Mason, A.H. Laird, and J.R. Smythies. Confinia

Neurologica, 18, 152 (1958).

R.G. Browne and B.T. Ho. Pharmacol. Biochem. and Behavior,

3, 109 (1975).

197
9. E.A. Zeller, J. Barsky, E.R. Berman, M.S. Cherkas, and J.R.

Fouts. J. Pharmacol. Exp. Ther., 124, 282 (1958).

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

L.J. Riceberg, M. Simon, H. Van Vunakis, and R.H. Abeles.

Biochem. Pharmacol., 24, 119 (1975).

K.D. Charalampous, K.E. Walker, and J. Kinross-Wright. Psychopharmacologia, 9, 48 (1966).

J. Ratcliff'e and P. Smith. Chem. Ind., p. 925 (July 18, 1959).

N.S. Shah and H.E. Himwich. Neuropharmacol., 10, 547 (1971).

B.T. Ho, S.F. Pong, R.G. Browne, and K.E. Walker. Experientia,

29, 275 (1973).

L. Demisch and N. Seiler. Biochem. Pharmacol., 24, 575 (1975).

G.F. Phillips and J. Gardiner. J. Pharm. Pharmacol., 21,

793 (1969).

J. Lundstrom and S. Agurell. J. Chromatogr., 36, 105 (1968).

L.J. Riceberg, H.Van Vunakis, and L. Levine. Anal. Biochem.

60, 551 (1974).

C. Van Peteghem, M. Van den Heede, and A. Heyndrickx in A.

Frigerio, ed., Recent Dev. Mass Spectr. Biochem. Med., 4th

Volume. Plenum Press, New York, p. 153 (1978).

J.M. Musacchio and M. Goldstein. Biochem. Pharmacol., 16,

963 (1967).

198
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