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Analytical Profiles of Drug Substances Excipients and Related Methodology Volume 31
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Analytical Profiles of Drug Substances Volume 18 1989
35107892 Analytical Profiles of Drug Substances and Excipients Vol 22 1993 ISBN 0122608224 9780122608223
Analytical Profiles -Volume 19
35578009 Profiles of Drug Substances Excipients and Related Methodology Volume 32 2005
Analytical Profiles of Drug Substances and Excipients Vol 27
Analytical Profiles of Drug Substances Volume 10
Pharmaceutical Coating Technol
Vol 16 Analytical Profiles of Drug Substan(BookSee.org) v 16
Pharmaceutical Excipients-boolket for Pharmacy Students
European Pharmacopoeia 8.0
European Pharmacopoeia 8.2 - 2014 - Index
Affiliations of Editor and Contributors ix
Profiles of Drug Substances
1. Acetylcholine Chloride:
1.1 Physical Profile 3
1.2 Analytical Profile 21
Abdullah A. Al-Badr and Humeida A. El-Obeid
2. Benazepril Hydrochloride: Comprehensive Profile 117
F. Belal, H.H. Abdine and A.A. Al-Badr
3. Ciprofloxacin:
3.1 Physical Profile 163
3.2 Analytical Profile 179
3.3 Drug Metabolism and Pharmacokinetic Profile 209
Mohammed A. Al-Omar
4. Dipyridamole: Comprehensive Profile 215
A. Khalil, F. Belal and Abdullah A. Al-Badr
5. Mefenamic Acid: Analytical Profile 281
Hadi Poerwono, Retno Widyowati, Hajime Kubo,
Kimio Higashiyama and Gunawan Indrayanto
6. Nimodipine
Related Methodology Review Articles
7. Evaluation of the Particle Size Distribution of
Pharmaceutical Solids 379
Harry G. Brittain
Cumulative Index 421
The comprehensive profiling of drug substances and pharmaceutical
excipients as to their physical and analytical characteristics remains at the
core of pharmaceutical development. As a result, the compilation and
publication of comprehensive summaries of physical and chemical data,
analytical methods, routes of compound preparation, degration path-
ways, uses and applications, etc., has always been a vital function to
both academia and industry. Historically, the Profiles series has always
provided this type of information, and has striven to publish the highest
quality available.
As the science of pharmaceutics continues to grow and mature, the need
for information similarly expands along many new fronts and the
appropriate vehicles where investigators may find the information they
need must develop accordingly. As a result, the content of the Profiles
series has expanded so that chapters will therefore fall into one or more of
the following main categories:
1. Physical characterization of a drug substance or excipient
2. Analytical methods for a drug substance or excipient
3. Detailed discussions of the clinical uses, pharmacology, phar-
macokinetics, safety, or toxicity of a drug substance or excipient
4. Reviews of methodology useful for the characterization of drug
substances or excipients
The expansion in scope of the Profiles series first appeared in Volume 30,
and continues in the present volume. Several of the chapters in the
current volume are comprehensive in nature, but others are more
specialized. The volume also contains a methodology review article on
particle size distribution. It is anticipated that future volumes of
the Profiles series will contain similar methodology reviews, as well as
other types of review articles that summarize the current state in a
particular field of pharmaceutics. As always, I welcome communications
from anyone in the pharmaceutical community who might want to
provide an opinion or a contribution.
Editor, Profiles of Drug Substances,
Excipients, and Related Methodology
hbrittain@earthlink.net
AFFILIATIONS OF EDITOR AND CONTRIBUTORS
H.H. Abdine: Department of Pharmaceutical Chemistry, College of
Pharmacy, King Saud University, P.O. Box 2457, Riyadh-11451,
Abdullah A. Al-Badr: Department of Pharmaceutical Chemistry, College
of Pharmacy, King Saud University, P.O. Box 2457, Riyadh-11451,
Mohammed A. Al-Omar: Department of Pharmaceutical Chemistry,
College of Pharmacy, King Saud University, P.O. Box 2457, Riyadh-
11451, Kingdom of Saudi Arabia
F. Belal: Department of Pharmaceutical Chemistry, College of Phar-
macy, King Saud University, P.O. Box 2457, Riyadh-11451, Kingdom of
Harry G. Brittain: Center for Pharmaceutical Physics, 10 Charles Road,
Milford, NJ 08848, USA
Humeida A. El-Obeid: Department of Pharmaceutical Chemistry, College
Kimio Higashiyama: Institute of Medicinal Chemistry Hoshi University,
4-41, Ebara 2-chome, Shinagawa-ku, Tokyo 142-8501, Japan
Gunawan Indrayanto: Faculty of Pharmacy, Airlangga, University Jl.
Dharmawangsa Dalam, Surabaya 60286, Indonesia
A. Khalil: Department of Pharmaceutical Chemistry, College of
Hajime Kubo: Institute of Medicinal Chemistry Hoshi University, 4-41,
Ebara 2-chome, Shinagawa-ku, Tokyo 142-8501, Japan
Hadi Poerwono: Faculty of Pharmacy, Airlangga University, Jl.
Retno Widyowati: Faculty of Pharmacy, Airlangga University, Jl.
Profiles of Drug Substances
Acetylcholine Chloride:
Physical Profile
College of Pharmacy, King Saud University
P.O. Box 2457, Riyadh-11451
PROFILES OF DRUG SUBSTANCES, 3 Copyright ß 2004 Elsevier Inc.
EXCIPIENTS, AND RELATED All rights reserved
METHODOLOGY – VOLUME 31
4 A.A. AL-BADR AND H.A. EL-OBEID
1. Description . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
1.1 Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
1.1.1 Systematic chemical names. . . . . . . . . . . . . 4
1.1.2 Nonproprietary names . . . . . . . . . . . . . . . . 5
1.1.3 Propietary names . . . . . . . . . . . . . . . . . . . 5
1.2 Formulae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5
1.2.1 Empirical formula, molecular weight
and CAS numbers . . . . . . . . . . . . . . . . . . . 5
1.2.2 Structural formula. . . . . . . . . . . . . . . . . . . 6
1.3 Elemental analysis. . . . . . . . . . . . . . . . . . . . . . . . . 6
1.3.1 Acetylcholine . . . . . . . . . . . . . . . . . . . . . . 6
1.3.2 Acetylcholine hydroxide. . . . . . . . . . . . . . . 6
1.3.3 Acetylcholine chloride . . . . . . . . . . . . . . . . 6
1.3.4 Acetylcholine bromide . . . . . . . . . . . . . . . . 6
1.4 Appearance and color . . . . . . . . . . . . . . . . . . . . . . 6
2. Physical Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . 6
2.1 pH. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
2.2 Solubility characteristics . . . . . . . . . . . . . . . . . . . . 6
2.3 Crystallographic properties . . . . . . . . . . . . . . . . . . 7
2.3.1 Crystal data . . . . . . . . . . . . . . . . . . . . . . . 7
2.3.2 X-Ray powder diffraction . . . . . . . . . . . . . 8
2.4 Thermal methods of analysis . . . . . . . . . . . . . . . . . 9
2.4.1 Melting behavior. . . . . . . . . . . . . . . . . . . . 9
2.4.2 Differential scanning calorimetry . . . . . . . 10
2.5 Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
2.5.1 UV/VIS spectroscopy . . . . . . . . . . . . . . . 10
2.5.2 Vibrational spectroscopy . . . . . . . . . . . . . 10
2.5.3 Nuclear magnetic resonance spectroscopy . 11
2.6 Mass spectrometry . . . . . . . . . . . . . . . . . . . . . . . 14
3. Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
4. Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17
5. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17
1. DESCRIPTION
1.1 Nomenclature
1.1.1 Systematic chemical names [1, 2]
Ammonium, 2-(acetyloxy)-N,N,N-trimethylethyl, chloride.
PHYSICAL PROFILE 5
Ethanaminium, 2-(acetyloxy)-N,N,N-trimethyl, chloride.
(2-Acetyloxyethyl)-trimethylammonium chloride.
(2-Acetoxyethyl)-trimethylammonium chloride.
O-Acetyl-choline chloride.
2-(Acetyloxy)-N,N,N-trimethylethanaminium chloride.
(2-Acetoxyethyl)-trimethylammonium hydroxide.
1.1.2 Nonproprietary names
Acetylcholine, Acetylcholine chloride, Chlorune D, Acetylcholinium
chloratum, Acetylcholin, Acetylcholini chloridum [1–3].
1.1.3 Proprietary names
Miochol, Alcon, Acecoline, Arterocholine, Ovisot, Covochol,
CooperVision, Coopervision, Acetylcholin ophthalmologic, Pragmaline,
Tonocholin B, Miovision, Covochol [1–3].
1.2 Formulae
1.2.1 Empirical formula, molecular weight and CAS numbers [1]
1.2.1.1 Acetylcholine
C7H16NO2 þ 146.21 [51843]
1.2.1.2 Acetylcholine hydroxide
C7H17NO3 163.2 —
1.2.1.3 Acetylcholine chloride
C7H16ClNO2 181.68 [60311]
1.2.1.4 Acetylcholine bromide
C7H16BrNO2 226.14 [66239]
1.2.2 Structural formula
O CH3
H3C C O CH2 CH2 N CH3
1.3 Elemental analysis
1.3.1 Acetylcholine
C 57.51%, H 11.03%, N 9.58% O 21.88%.
1.3.2 Acetylcholine hydroxide
C 51.51%, H 10.50%, N 8.58%, O 29.41%.
1.3.3 Acetylcholine chloride
C 46.28%, H 8.88%, Cl 19.52%, N 7.71%, O 17.61% (1).
1.3.4 Acetylcholine bromide
C 37.18%, H 7.13%, Br 35.34%, N 6.20%, O 14.15%.
1.4 Appearance and color
Acetylcholine chloride is obtained as white or off-white hygroscopic
crystals, or as a crystalline powder. The salt is odorless, or nearly
odorless, and is a very deliquescent powder. Acetylcholine bromide is
obtained as deliquescent crystals, or as a white crystalline powder. The
substance is hydrolyzed by hot water and alkali [1–5].
2.1 pH
Following reconstitution, commercially available injectable formulations
are characterized by a pH range of 5–7.8 [4].
2.2 Solubility characteristics
Acetylcholine chloride is freely soluble in water, alcohol, propylene
glycol, and chloroform, and is practically insoluble in ether. The
solubility of 500 mg is complete in 5 mL alcohol, resulting in a colorless
solution. Solutions of acetylcholine chloride in water are unstable.
Likewise, acetylcholine bromide is very soluble in cold water; decom-
posed by hot water or alkalis, soluble in alcohol, and practically insoluble
in ether [3–5].
2.3 Crystallographic properties
2.3.1 Crystal data
The crystal structures of acetylcholine bromide and acetylcholine iodide
have both been investigated by Sorum [6, 7]. The structure was solved in
the P21 space group, with two crystallographically different molecules in
the asymmetric unit. The crystal structure of acetylcholine chloride was
reported by Allen [8].
Acetylcholine chloride is very hygroscopic and extremely soluble, giving a
solution of high viscosity. For this reason, it is difficult to prepare crystals
appropriate for X-ray analysis. At 25 C, a saturated solution contains
approximately 0.962 g/mL of solute, and has a density of 1.19 g/mL.
When a solution, nearly saturated at room temperature, is cooled to
approximately 10 C, the anhydrous salt crystallizes slowly as clusters of
needles. There was no evidence of hydrate formation. Single crystals,
mounted and sealed in glass capillaries were examined about the needle
axis [a] by oscillation and Weissenberg methods using filtered copper
radiation, and by the precession method using molybdenum radiation.
Provided that the sealing was satisfactory these mounted crystals
remained unchanged for a period of several weeks.
The crystal data obtained were as follows:
Acetylcholine chloride (CH3)3NCH2CH2OCOCH3. Cl
Orthorhombic Formula Weight: 181.5
a ¼ 6.28 Å Unit cell volume: 952 Å3
b ¼ 9.93 Å
c ¼ 15.26 Å
Space group: P 212121.
Systematic extinctions observed:
h00 for h ¼ 2n 0k0 for k ¼ 2n 00l for l ¼ 2n
Density: Observed 1.202 g/mL; Calculated 1.27 g/mL
Molecules per unit cell: 4
The density of the crystal was estimated by flotation in benzene/
chloroform mixtures. It is recognized that the value is not very accurate
and is almost certainly lower than the true value because of the difficulty
of freezing the crystal completely from the viscous mother liquor.
However, the method is sufficient to confirm the number of molecules per
unit cell [8].
2.3.2 X-Ray powder diffraction
The X-ray powder diffraction pattern of acetylcholine chloride was
obtained using a Philips PW-1710 diffractometer system with a single
crystal monochromator and using copper Kradiation. The powder
pattern is shown in Figure 1. The scattering angles, d-spacings, and
relative intensities were automatically obtained for a pure sample of
acetylcholine chloride, and are shown in Table 1.
Canepa [9] suggested that one of the two molecular forms of acetylcho-
line bromide reported by Sorum [6, 7] was incorrect. Dunitz [10], on the
other hand, observed some unusual systematic absences and suggested
that the crystal used was a twinned P21/c rather than a single P21
specimen. In response to these observations, Svinning and Sorum [11]
refined the structure of acetylcholine bromide based on previous
photographic intensities corrected for twinning by Canepa, Pauling and
2- Theta -Scale
Scattering Angle (degrees 2-θ)
Figure 1. X-ray powder diffraction pattern of acetylcholine chloride.
Crystallographic Data from the X-ray Powder Diffraction
Pattern of Acetylcholine Chloride
Scattering d-Spacing Relative Scattering d-Spacing Relative
Angle (Å) intensity Angle (Å) intensity
degrees (%) degrees (%)
7.054 12.5219 6.67 11.606 7.6184 6.69
14.592 6.0653 100.00 14.881 5.9484 12.13
16.435 5.3890 37.06 17.497 5.0643 68.38
24.906 3.5721 8.25 27.755 3.2115 5.05
Sorum [12] to a final R of 0.10. Thus, the reinvestigation of the structure
of acetylcholine bromide, [C7H16O2N] þ Br, with X-ray diffraction
intensities collected from two untwinned crystals showed that the crystals
are monoclinic, and are characterized by a space group of P21/n, with
a ¼ 10.966 (4), b ¼ 13.729 (7), c ¼ 7.159 (4) Å, ¼ 108.18 (7) , and
Z ¼ 4. The structure was refined by full-matrix least squares calculations
using 1730 observed reflections, and anisotropic temperature factors for
all non-hydrogen atoms. The final R was found to be 0.041. Atomic
coordinates, thermal parameters, bond lengths, and angles were
compared with those from a previous work on acetylcholine derivatives.
2.4 Thermal methods of analysis
2.4.1 Melting behavior
Acetylcholine chloride, when dried earlier at 110 C in a capillary tube for
one hour, melts between 149 and 152 C [5]. Acetylcholine bromide melts
at 143 C [3].
10 A.A. AL-BADR AND H.A. EL-OBEID
222.42°C
−1 545.8J/g
42.49J/g
Heat Flow (W/g)
0 100 200 300 400 500
Figure 2. Differential scanning calorimetry thermogram of acetylcholine chloride.
2.4.2 Differential scanning calorimetry
Differential scanning calorimetry of acetylcholine chloride was performed
using a Dupont DSC Model TA 9900 thermal analyzer system, interfaced
to a Dupont data collection system. Figure 2 shows the DSC
thermogram, which was recorded from 50 to 350 C.
2.5 Spectroscopy
2.5.1 UV/VIS spectroscopy
Owing to its lack of chromophores, acetylcholine chloride does not
exhibit any absorption bands in the ultraviolet or visible region.
2.5.2 Vibrational spectroscopy
The infrared spectrum of the substance that is shown in Figure 3 was
recorded on a Pye Unicam SP infrared spectrophotometer (KBr pellet
method). The major observed bands have been correlated with the
functional groups, and these are 2970 and 2920 cm1 (CH3 and CH2
stretching), 1755 cm1 (C¼O stretching) and 1465 cm1 (CH deforma-
tion). Clarke reported the principal peaks for acetylcholine bromide at
wavenumbers of 949, 1089, 869, 1231, 1000, 1740 cm1 (KBr pellet) [3].
PHYSICAL PROFILE 11
4000 3000 2000 1500 1000 600
Energy (cm−1)
Figure 3. Infrared absorption spectrum of acetylcholine chloride.
Srivastava et al. carried out a complete study of the vibrational spectrum
of acetylglycine, and evaluated its relation to the spectrum of acetylcho-
line [13]. A normal coordinate analysis of acetylglycine was carried out
using the Wilson's GF matrix method. Vibrational frequencies were
assigned, and the infrared spectra of acetylglycine and acetylcholine
compared. Conformation-sensitive modes of acetylcholine were identi-
fied, and a transferable Urey-Bradley force field was also obtained.
2.5.3 Nuclear magnetic resonance spectroscopy
2.5.3.1 1H-NMR spectra
The 1H-NMR spectrum of acetylcholine chloride in CDCl3 was recorded
on a Varian XL200 NMR spectrometer using TMS as the internal
reference. The simple proton (Figure 4) and HOMCOR (Figure 8)
spectra were used to determine the exact chemical shifts and proton
coupling. The HETCOR spectrum (Figure 9) was used to assign the
protons to their respective carbons in the molecule. The resulting proton
chemical shifts are assigned as listed in Table 2. 1H-NMR studies [14] on
acetylcholine were used in determining its conformation.
2.5.3.2 13C-NMR spectra
The 13C-NMR spectrum of acetylcholine chloride was recorded in
CDCl3, using TMS as the internal reference, on a Varian XL 200 NMR
12 10 8 6 4.143 4 2 0 PPM
Figure 4. 1H-NMR spectrum of acetylcholine chloride in CDCl3.
H-NMR Chemical Shift Assignments for
Acetylcholine Chloride
Chemical shifts Number Multiplicity Assignment
(ppm, relative of protons
to TMS)
2.13 3 singlet
H3C C O CH2 CH
4.14 2 multiplet
N CH2
spectrophotometer. Figures 5, 6, and 7 show the S2PUL pulse sequence,
and the APT and DEPT spectra, respectively. The assigned chemical
shifts are listed in Table 3.
180 160 140 120 100 80 60 40 20 PPM 0
Chemical Shift (ppm)
Figure 5. C-NMR spectrum of acetylcholine chloride in CDCl3.
160 140 120 100 80 60 40 20 PPM 0
Figure 6. C-NMR spectrum (APT) of acetylcholine chloride in CDCl3.
Figure 7. C-NMR spectrum (DEPT) of acetylcholine chloride in CDCl3.
The 13C chemical shifts of acetylcholine chloride were assigned by Murari
and Baumann [15]. The results of their 13C-NMR studies [16] on
acetylcholine chloride were used for analyzing the conformational
isomers of the molecule.
2.6 Mass Spectrometry
The mass spectrum of acetylcholine chloride was obtained utilizing a
Shimadzu PQ-5000 mass spectrometer, with the parent ion being collided
with helium carrier gas. The mass spectrum is shown in Figure 10, and
Table 4 shows the mass fragmentation pattern. Clarke lists the principal
mass spectral peaks at m/z 58, 43, 57, 149, 71, 42, 41, 55 [3].
Mass spectrometry was used in combination with chromatographic
methods for the analysis of acetylcholine in biological systems, with the
mass spectral peak at m/z 58 being most frequently used for detection.
Quantitation of acetylcholine and its related compounds was successfully
performed using GC/MS [17–26] and HPLC/MS [27–29].
3. STABILITY
Aqueous solutions of acetylcholine chloride are found to be unstable.
Such solutions are decomposed by heat, and are incompatible with alkalis
C-NMR Chemical Shift Assignments for
Chemical shift Carbon Assignments
(ppm, relative to TMS)
Found Reported [15]
21.07 21.3 1
170.12 173.8 2
5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0
Figure 8. HOMCOR-NMR spectrum of acetylcholine chloride in CDCl3.
Figure 9. HETCOR-NMR spectrum of acetylcholine chloride in CDCl3.
Figure 10. Mass spectrum of acetylcholine chloride.
and acids. Some reports revealed that the solution is stable for 2–3 h
following reconstitution, after which breakdown products are observed.
The reconstituted solution should be used immediately, and any unused
portion discarded. The unconstituted two-chambered vials of acetylcho-
line chloride for injection should be stored in air-tight containers at
15–30 C, and protected from freezing [1, 4, 5, 30].
Assignments for the Fragmentation Pattern Observed in the
Mass Spectrum of Acetycholine Chloride
m/z Relative Fragment
H3C N CH CH2 H3C N CH CH2
CH3 CH3
H3C N CH2
43 10 H3C–C:O
4. ACKNOWLEDGMENTS
The authors wish to thank Mr. Tanvir A. Butt, Department of
Pharmaceutical Chemistry, College of Pharmacy, King Saud
University, Riyadh, Saudi Arabia, for typing this profile.
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Society, S. Budavari, ed., Zurich, p. 8 (1987).
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30. Product Information: MiocholÕ , Acetylcholine, Cooper, Vision
Pharmaceuticals, San German, Puerto Rico (1995).
Analytical Profile
PROFILES OF DRUG SUBSTANCES, 21 Copyright ß 2004 Elsevier Inc.
1. General Methods of Analysis . . . . . . . . . . . . . . . . . . . . . . 23
1.1 Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . 23
1.2 Reviews on methods of analysis . . . . . . . . . . . . . . 24
1.3 Assay methods . . . . . . . . . . . . . . . . . . . . . . . . . . 25
1.3.1 Method 1: Percentage of acetyl. . . . . . . . . 25
1.3.2 Method 2: Percentage of chlorine . . . . . . . 25
2. Titrimetric Methods of Analysis . . . . . . . . . . . . . . . . . . . . 25
2.1 Acidimetric methods . . . . . . . . . . . . . . . . . . . . . . 25
2.2 Potentiometric methods. . . . . . . . . . . . . . . . . . . . 26
3. Electrochemical Methods of Analysis . . . . . . . . . . . . . . . . 26
3.1 Conductimetric methods . . . . . . . . . . . . . . . . . . . 26
3.2 Polarographic methods . . . . . . . . . . . . . . . . . . . . 27
3.3 Voltammetric methods . . . . . . . . . . . . . . . . . . . . 27
3.4 Coulometric flow titration methods . . . . . . . . . . . 28
3.5 Flow injection method . . . . . . . . . . . . . . . . . . . . 66
3.6 Amperometric methods . . . . . . . . . . . . . . . . . . . . 66
4. Spectrophotometric Methods of Analysis . . . . . . . . . . . . . . 67
4.1 Infrared spectrophotometric method. . . . . . . . . . . 67
4.2 Photometric methods . . . . . . . . . . . . . . . . . . . . . 67
4.3 Mass spectrometric method . . . . . . . . . . . . . . . . . 68
4.4 Colorimetric methods . . . . . . . . . . . . . . . . . . . . . 68
4.5 Fluorimetric methods . . . . . . . . . . . . . . . . . . . . . 70
4.6 Chemiluminescence methods . . . . . . . . . . . . . . . . 72
5. Biological Assay Methods of Analysis. . . . . . . . . . . . . . . . 74
6. Chromatographic Methods of Analysis . . . . . . . . . . . . . . . 75
6.1 Thin layer chromatography (TLC) . . . . . . . . . . . . 75
6.2 Gas chromatography (GC) . . . . . . . . . . . . . . . . . 75
6.3 High performance liquid chromatography
(HPLC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. 79
6.4 Liquid chromatography (LC). . . . . . . . . . . . . . .. 80
6.5 Cation-exchange liquid chromatography . . . . . . .. 97
6.6 High performance liquid chromatography–mass
spectrometry (HPLC/MS) . . . . . . . . . . . . . . . . .. 97
6.7 Gas chromatography–mass spectrometry
(GC/MS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98
6.8 Capillary electrophoresis . . . . . . . . . . . . . . . . . . . 101
6.9 Radioassay methods . . . . . . . . . . . . . . . . . . . . . . 102
7. Other Methods of Analysis . . . . . . . . . . . . . . . . . . . . . . . 105
7.1 Ion-selective method . . . . . . . . . . . . . . . . . . . . . . 105
ACETYLCHOLINE CHLORIDE: ANALYTICAL PROFILE 23
7.2 Microwave methods . . . . . . . . . . . . . . . . . . . . . . 105
7.3 Microdialysis methods. . . . . . . . . . . . . . . . . . . . . 105
8. Acknowledgment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105
9. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106
1. GENERAL METHODS OF ANALYSIS
1.1 Identification
Vizi et al. described a simple and sensitive method for the identification
and assay of acetylcholine [1]. A modification of the acetylcholine
assay technique for guinea pig ileum has been combined with either
minivolume gel filtration or high pressure liquid chromatography
separation of the samples. In addition, labeled acetylcholine (14C-ACh)
was eluted with unlabeled acetylcholine, or with samples expected to
contain acetylcholine, and their elution profiles were compared by
bioassay combined with radioassay of eluted fractions. When the elution
of both biological activity and label occurred in the same fraction, it
was concluded that the substance assayed on guinea pig ileum
was acetylcholine. This method was found to be sensitive to 0.5 ng
(1.8 pmol) of acetylcholine, and its reproducibility was within 5%.
Wiegrebe and Vibig reported a method for the identification of choline
and choline salts [2]. A critique of the methods of identification of
acetylcholine chloride and choline chloride according to the Swiss
Pharmacopoeia VI and of choline chloride according to the German
Pharmacopoeia 8 was given and alternative procedures suggested. As
alternatives, the pyrolytic degradation of acetylcholine to acetaldehyde,
which can be identified by the Simon-Awe method, and of choline to
formaldehyde followed by a reaction with chromotropic acid to give
a deep violet color, are suggested.
Mynka et al. used infrared spectrophotometry for the identification
and determination of acetylcholine [3]. Beer's law was obeyed from 1 to
10 mg/mL of acetylcholine, and the coefficient of variation was 2.7%.
Zhu et al. used an in vivo microdialysis and a reversed phase ion-pair
liquid chromatography-tandem mass spectrometry method for the
determination and identification of acetylcholine in rat brain [4].
A microdialysis probe was surgically implanted in the striatum of a rat,
and this was perfused with Ringer's solution at a flow rate of 2 mL/min.
The resulting microdialysate samples were mixed (9:1) with acetyl-b-
methylcholine (internal standard, 1 ng/mL), and a 950 mL portion was
analyzed on a Discovery C18 column (50 cm  2.1 mm i.d.). The method
used 20 mM ammonium acetate/20 mM heptafluorobutyric acid in
methanol/water (1:9, pH 3.2) as the mobile phase at 0.4 mL/min, and
the analyte and internal standard were detected by tandem ion-trap
mass spectrometry. The calibration graph was linear up to 500 fg/mL of
acetylcholine, and the on-column detection limit was 200 fg acetylcholine.
A sensitive and specific bioautographic method for the identification of
choline and its derivatives, was developed by Lewin and Marcus, utilizing
a Neurospora crassa mutant [5]. As little as 0.03 g of choline chloride
is detectable. The applicability of the method for detection of choline
derivatives in lipid hydrolyzate is discussed. A method for determining
the rate of diffusion of compounds from paper chromatograms into
bioautograph agar is described.
Unger and Ryan used a secondary-ion mass spectrometry method for the
identification of choline and its esters [6]. Secondary-ion mass spectro-
metry provides the mass spectra of choline and its esters, from which
molecular weights and structures may be readily obtained. Structure-
spectral correlations were developed using analogies from other forms of
mass spectrometry. Spectra can be recorded on metal or paper substrates,
and individual components can be identified even in complex samples.
1.2 Reviews on methods of analysis
The recent developments in the determination of acetylcholine and
choline, the advantages and the disadvantages of a variety of analytical
methods used in the analysis, were reviewed and discussed by Maruyama
et al. [7]. Hanin published an overview for the methods used for the
analysis and measurement of acetylcholine [8]. Shimada et al. presented
a review, including the applications of some reactors in the analysis of
choline and acetylcholine. The immobilized enzyme reactors were used
for detection systems in high performance liquid chromatography [9].
Tsai reviewed the separation methods used in the determination of
choline and acetylcholine [10]. This review surveyed the array of anal-
ytical techniques that have been adopted for the measurement of
acetylcholine or its main precursor/metabolite (choline), ranging from
simple (bioassay, radio enzymatic assay, gas chromatography–flame
ionization detection, gas chromatography–mass spectrometry, high
performance liquid chromatography–electrochemical detection, high
performance liquid chromatography–mass spectrometry) to the
sophisticated (combination of microimmobilized enzymatic reactor,
microbore high performance liquid chromatography, and modified
electrode technology for the detection of ultralow levels), with particular
emphasis on the state of the art techniques.
1.3 Assay methods
United State Pharmacopeia XX describes the following assay methods
for acetylcholine as a reagent [11]:
1.3.1 Method 1: Percentage of acetyl
Weigh accurately about 400 mg of analyte, previously dried at 105 C for
3 h, and dissolve in 15 mL of water in a glass-stoppered conical flask. Add
40 mL of 0.1 N-sodium hydroxide VS, and heat on a steam bath for
30 min. Insert the stopper, allow to cool, add phenolphthalein TS, and
titrate the excess alkali with 0.1 N sulfuric acid VS. Determine the exact
normality of the 0.1 N sodium hydroxide by titrating an aliquot of
40.0 mL after it has been treated in the same manner as in the test. Each
milliliter of 0.1 N sodium hydroxide is equivalent to 4.30 mg of CH3CO.
Between 23.2 and 24% of CH3CO is found.
1.3.2 Method 2: Percentage of chlorine
Weigh accurately about 400 mg of acetylcholine chloride, previously
dried at 105 C for 3 h, and dissolve in 50 mL of water in a glass stoppered
125 mL flask. Add with agitation 30.0 mL of 0.1 N silver nitrate VS, then
add 5 mL of nitric acid and 5 mL of nitrobenzene and shake well.
Thereafter, add 2 mL of ferric ammonium sulfate TS, and titrate the
excess silver nitrate with 0.1 ammonium thiocyanate VS. Each milliliter
of 0.1 N silver nitrate is equivalent to 3.545 mg of Cl. Between 19.3 and
19.8% of Cl is found.
2. TITRIMETRIC METHODS OF ANALYSIS
2.1 Acidimetric methods
Szakacs-Pinter and Perl-Molnar described a new and simple procedure
for the determination of choline and its derivative. The method is based
on the acidimetric determination of the volatile base produced by the
K2S2O8 oxidation of choline in an alkaline medium [12]. The procedure
was carried out by distilling the trimethylamine formed continuously
during the reaction. Optimum conditions for the determination are: 0.5
M alkali, 15 min distillation time, and a 10 fold excess K2S2O8. Choline
or acetylcholine could be determined in 0.005–0.1 N solution of
pharmaceutical preparations and animal feed.
2.2 Potentiometric methods
Kakutani et al. described an ion-transfer voltammetry and potentiometry
method for the determination of acetylcholine with the interface between
polymer–nitrobenzene gel and water [13]. The PVC-nitrobenzene gel
electrode was prepared as described by Osakai et al. [14]. The transfer of
acetylcholine ions across the interface between the gel electrode and water
was studied by cyclic voltammetry, potential-step chronoamperometry,
and potentiometry. The interface between the two immiscible electrolyte
solutions acted as the ion-selective electrode surface for the determination
of acetylcholine ions.
Baum reported a potentiometric determination of acetylcholine activity
using an organic-cation-selective electrode [15]. The performance of
a liquid membrane electrode selective for acetylcholine (Corning No.
476.200) was investigated. Measurements of the potential difference
at various concentration of acetylcholine were made against a calomel
reference-electrode.
Nakazawa and Tanaka described a potentiometric titration of salts of
organic bases in non-aqueous solvents with the addition of bismuth
nitrate [16]. The base halide or hydrohalide (0.7 mequiv.) is dissolved in
40 mL of anhydrous acetic acid, and then 40 mL of 1,4-dioxane and
2.5 mL of 5% Bi(NO3)3 solution in acetic acid are added. The Bi(NO3)3
prevents interference by the halide. The solution is titrated to a
potentiometric endpoint with 0.1 M HClO4 (a blank titration is also
carried out). Results of purity assays of acetylcholine chloride and other
compounds were tabulated, and it was found that the coefficient of
variation was 0.18%.
3. ELECTROCHEMICAL METHODS OF ANALYSIS
3.1 Conductimetric methods
An immobilized enzyme flow injection conductimetric system was used by
Godinho et al. for the determination of acetylcholine [17]. Aliquots
of acetylcholine standard solutions were injected into a carrier stream of
0.1 M phosphate buffer of pH 7.4 at 1.19 mL/min, and passed through a
polyethylene tube (3.5 cm  3 mm i.d.) containing acetylcholineesterase
immobilized on glass beads. The solution was merged with a stream of 2 M
H2SO4 at 1.19 mL/min, and passed through a PTFE membrane diffusion
cell. The acetic acid that diffused through the membrane was carried by a
stream of water to a conductimetric cell. The calibration graph was linear
from 10 mM to 1 mM acetylcholine, and the method was applied to the
analysis of acetylcholine in a pharmaceutical formulation.
3.2 Polarographic methods
A quantitative method was reported by Maslova for the determination of
acetylcholine in biological tissues by polarographic analysis utilizing a
rotating platinum electrode [18]. The principle of the method is based
upon a polarographic analysis of the iron ions which remain after the
formation of a specific Fe–acetylhydroxamic acid complex. Using this
method, it was shown that 1 mL of peripheral blood of a healthy adult
man contained 6.6 ng of acetylcholine.
3.3 Voltammetric methods
The ion transfer voltammetry and potentiometry of acetylcholine with the
interface between polymer-nitrobenzene gel and water was reported by
Kakutani et al. [13]. The polyvinyl chloride-nitrobenzene gel electrode was
prepared by Osakai et al. [14], and the transfer of acetylcholine ions across
the interface between the gel electrode and water was studied by cyclic
voltammetry, potential step chronoamperometry, and potentiometry. The
interface between the two immiscible electrolyte solutions acted as the ion
selective electrode surface for the analysis of acetylcholine ions.
Matsue et al. [19] recommended an electrochemical determination
method using Nafion coated electrodes for the determination of
electroinactive medicines. Acetylcholine was determined voltammetri-
cally with the use of a Nafion 117-coated vitreous carbon working
electrode and a saturated calomel electrode. The method is based on
competition between (ferrocenylmethyl) trimethylammonium ion and the
drug cations for entry into the Nafion layer. The presence of acetylcho-
line caused a decrease in the peak current for (ferrocenylmethyl)
trimethylammonium ion, from which the acetylcholine concentration
is determined. The method was described to have permitted the
determination of 1 mM–1 mM of acetylcholine.
Osakai et al. used a microcomputer-controlled system for the ion transfer
voltammetry procedure [20]. The system used is based on a NEC PC-9801
microcomputer, which was designed by using a polarizable oil–water
interface as an ion-selective electrode surface. The system was applied to
the determination of acetylcholine ion by cyclic, differential pulse, and
normal pulse voltammetry at the PVC–nitrobenzene gel electrode. The
amperometric measurement was carried out with voltage pulses of short
durations and constant amplitude.
Hale et al. reported the use of an enzyme-modified carbon paste for the
determination of acetylcholine [21]. The sensor was constructed from
a carbon paste electrode containing acetylcholineesterase and
choline oxidase, and the electron transfer mediator tetrathiafulvalene.
The electrode was used for the cyclic voltammetric determination of
acetylcholine in 0.1 M phosphate buffer at þ 200 mV versus saturated
calomel electrode. Tetrathiafulvalene efficiently re-oxidized the reduced
flavin adenine dinucleotide centers of choline oxidase. The calibration
graph was linear up to 400 mM acetylcholine, and the detection limit was
0.5 mM.
Kawagoe et al. described an enzyme-modified organic conducting salt
microelectrode, comprised of a carbon fiber electrode (20 mM), having a
recessed tip into which crystals of the conducting salt tetrathiafulvalene-
tetracyanoquinodimethane were galvanostatically deposited, followed by
glucose oxidase cross-linked with glutaraldehyde for the detection of
glucose [22]. For the detection of acetylcholine, acetylcholineesterase and
choline oxidase were co-immobilized in the recess instead of glucose
oxidase. The micrographic and cyclic voltammetric characterization of
the recessed electrode was described. The acetylcholine microelectrode
responded to 5 mM or 800 mM acetylcholine within four seconds (Table 1).
By using the electrolysis at the interface between two immiscible
electrolyte solutions, Marecek and Samec determined acetylcholine by
a differential pulse stripping voltammetric method [23].
3.4 Coulometric flow titration methods
Gyurcsanyi et al. studied the determination of acetylcholine after
enzymatic hydrolysis by triangle-programmed coulometric flow titration
[24]. A sample containing acetylcholine was pumped through an enzyme
reactor containing acetylcholineesterase immobilized on controlled-pore
glass. The resulting acetic acid was titrated against KOH with continuous
ACETYLCHOLINE CHLORIDE: ANALYTICAL PROFILE
Biosensors for the Detection and Analysis of Acetylcholine
Biological system Biosensor design Results and remarks Reference
Acetylcholineesterase Carbon paste electrode containing The calibration graph was linear up to [21]
and choline oxidase AChE and ChO and the electron 400 mM ACh and the detection limit
transfer mediator tetrathiafulva- was 0.5 mM. The tetrathiafulvalene-
lene. The electrode was applied to based sensor operated efficiently at
the cyclic voltammetric determina- low applied potential (0 to  100 mV
tion of ACh in 0.1 M phosphate versus SCE).
buffer at þ 200 mV versus SCE.
Acetylcholineesterase Carbon-fiber electrode having a No interference from uric acid. The [22]
and choline oxidase recessed tip into which crystals of electrode responded to 5 mM or
the conducting salts tetrathiafulva- 800 mM ACh within 4 s. Kinetic
lenetetracyanoquinodimethane studies of the biosensor were
were galvanostatically deposited reported.
followed by AChE and ChO
which were co-immobilized in the
recess.
(continued )
Table 1 (continued)
Acetylcholineesterase Glass microelectrode with AChE Nernstian response. Detection limit is [63]
A.A. AL-BADR AND H.A. EL-OBEID
(AChE) immobilized on active surface. pH-dependent. At pH 8.5 detection
limit was 10 mM. In cerebrospinal
fluids it was 0.1 mM.
Choline oxidase and Enzymes immobilized on a nylon Rectilinear response in the range of [64]
acetylcholineesterase net attached to H2O2-selective 1–10 mM. Response time 1–2 min.
amperometric sensor. ChO is used Interferences occur from ascorbic
for choline and AChE and ChO acid, primary amines, and most
for acetylcholine. seriously from betaine aldehyde.
Nicotinic receptor Receptor was fixed into a Sensitive to ACh, H þ , Na þ and K þ . [65]
from Torpedo cross-linked poly(vinylbutyral) Steady state reached after 2 min.
californica membrane covering the gate of an Initial rate of change of the voltage
ISFET which was mounted in a was a rectilinear function of log
sample cell with a reference ISFET ACh concentration was a from 0.1
and Ag–AgCl reference electrode. to 10 mM. When receptor was immo-
bilized in the membrane by using
lecithin an amplified response was
obtained which was due to Na þ
flux across receptor channel.
Acetylcholine receptor Receptor protein noncovalently Response was concentration depen- [66]
from electric organ bound on the surface of a planar dent and specific for ACh and
of Torpedo sp. interdigitated capacitative sensor. inhibited by ( þ )-tubocurarine,
amantidine and -neurotoxin.
Acetylcholineesterase Anal Abst., 1988, 50, 6J174 Sensor Determination of ACh and Ch in rats [67]
previously described. brain extracts. Rectiliner calibration
graphs were obtained for 12–190 mM
acetylcholine. Response times, 2 min.
Recoveries were 93–105% of ACh.
Used for determination of ACh in
ophthalmic preparations.
Acetylcholineesterase Sensor was constructed by attaching Response (amount of H2O2 produced) [68]
and choline oxidase 4 mm2 of film, mounted between was rectilinear for 1–660 mM ACh.
Nephrophan membrane (25 mm) Electrode was stable for 20 days or
to a platinum electrode using a 700 determinations.
manual polarograph or a flow
through system.
Acetylcholineesterase Enzymes were entrapped in a photo- The range of application was 20– [69]
and choline oxidase cross-linkable polymer. Detection 750 mM for ACh. Response time was
of H2O2 liberated from the enzyme 2–4 min. The immobilized enzyme
reaction. membranes were very stable, and
stored in a dry state.
Acetylcholineesterase pH sensitive ISFET (based on a Suitable for determining 0.1–10 mM [70]
silicon-SiO2-Ta2O5) by coating its ACh. A response of 61 mV per
tantalum pentoxide layer with a decade of the substrate
gelatin membrane containing concentration at pH 7.5 was
chloineesterase immobilized with obtained.
use of glutaraldehyde. The sensor
was operated versus silver–silver
chloride in saturated KCl.
Acetylcholineesterase Enzymes were co-immobilized on The response time was 2 min. [71]
and choline oxidase chemically preactivated immun- Detection limit was 50 nM and the
dyne polyamide membrane (thick- response was rectilinear up to
ness 120 mm, size cut-off 3 mm). 20 mM.
Applying 20 mL of 1% ChO solu-
tion in 0.1 M-phosphate buffer (pH
6). Assay was based on electroche-
mical detection of the generated
H 2O 2.
Acetylcholineesterase Enzyme was covalently immobilized The response time was 3–10 min for [72]
on a bovine serum albumin- 0.1–10 mM ACh. pH change of 6–8
modified H þ -selective coated wire had little effect. Coefficient of varia-
electrode. The sensor was used in tion was 5.7 and 5.8%.
5 mM phosphate buffer pH 7 at
Acetylcholineesterase A nereistoxin sensor is prepared Good linearity was observed. Used [73]
from a combination of AChE, for determination of ACh and
ChO and an O electrode. phosphate.
Acetylcholineesterase Carbon fiber electrode (7 mm) Calibration graph was rectilinear [74]
and choline oxidase dipped into poly(vinyl alcohol)- from 0.1 to 1 mM ACh. Detection
quaternized stilbazole-ChO-AChE limit was 0.05 mM, of ACh and
solution, drying in air and exposing detection time is 5 s.
to fluorescent room light for 1 h.
Acetylcholineesterase Electrode was developed by A kinetically controlled bioenzyme [75]
and choline oxidase co-immobilization of AChE with sensor was also used at a low activity
ChO–H2O2. Disposable sensors of ChE for determining inhibitors.
constructed by co-immobilization
in a gelatin membrane on Pt elec-
trode or by immobilizing AChE
in polyurethane on a thick-film
metallized Pt electrode.
Acetylcholineesterase Prepared by mounting a carbon The calibration graph was rectilinear [76]
and choline oxidase fiber (200 mm diam) in a glass capil- from 0.2 to 1 mM of ACh. The
lary with silver paste and epoxy response time was 0.8 min.
resin, electrochemically pretreating
the electrode from 0 to 1.2 V for
15 min, and dipping the electrode
in 11% PVA-Styryl pyridinium
solution containing AChE and
ChO.
Acetylcholineesterase Enzyme membrane in H2O was The best results were obtained at pH 8 [77]
and choline oxidase treated with 11% solution of PVA- and at 30 . The calibration graph
SbQ (polyvinyl alcohol) with styryl was rectilinear for 5 mM ACh. The
pyridinium groups. Mixture was storage stability of the dry mem-
spread on a cellulose nitrate mem- brane was excellent.
brane and air dried. The membrane
was exposed to UV radiation for
3 h and stored at 4 C. The enzyme
membrane was fixed with a Pt elec-
trode. Sample was dissolved in
phosphate buffer and measured.
Acetylcholineesterase Immobilization of horseradish Response time of 30 s and maintained [78]
and choline oxidase peroxidase in the redox polymer; its sensitivity for 24 h at low sub-
poly (4-vinylpyridine-chlorobis- strate concentration. Responses
(2,20 -bipyridyl) osmium cross- were rectilinear up to 10 mM H2O2
linked by means of polyoxyethylene for 100 mM choline with detection
400 diglycidyl ether on polished limit of 10 nM and 1 mM.
vitreous carbon electrodes.
Acetylcholineesterase Carbon-fiber micro-electrode for Calibration graph for ACh and Ch [79]
and choline oxidase determination of ACh and choline, was rectilinear between 0.1 and
mounted in a glass capillary tube. 3 mM. Interference from ascorbic
Enzyme was immobilized on the acid was not observed in the range
surface of the carbon fiber with 0.1 to 0.3 mM.
albumin. Electrode was dip-coated
with Nafion. Evaluation of selec-
tivity and dynamic range, at a
fixed potential of 1.2 V versus
Ag–AgCl.
Enzymes Enzymes immobilized in a polymer Detection limits were 0.1 and 1 mM [80]
solution. Acetylcholine and choline of Ch and ACh, respectively.
are determined at the mM level with Calibration graphs were rectilinear
a precision of 3.5–7%. for 0.1–100 and 1–100 mM of choline
and acetylcholine. Storage stability
of the sensing element is satisfactory.
Response time was 3–10 min.
Acetylcholineesterase Coating a polished vitreous-carbon Calibration graphs were rectilinear [81]
and choline oxidase electrode with 5 mL of a 0.5% solu- for 1–13 mM-choline (sensitivity
tion of choline oxidase which was 48 nA/mM), 0.04–0.06 mM-arseno-
dried by heating. Residue was choline (0.44 mA/mM) and up to
coated with 5 mL of a 1:10 mixture 5 mM acetylcholine. Detection limits
of Nafion and water. The dried were 50 mM, 10 mM and 43 mM,
electrode (ChO activity 0.382 iu/ respectively. There was no inter-
cm2) was used at 0.25 versus SCE ference from anionic and easily
for the amperometric determina- oxidized serum constituents.
tion of choline in 0.1 M phosphate
buffer of pH 6.5 containing 1 mM
K3Fe(CN)6 as redox mediator.
ACh sensor, 0.5 mL of a 0.1% solu-
tion of AChE to choline oxidase
coating and heat drying.
Acetylcholineesterase Carbon fiber was mounted in a glass Calibration graph was linear from [82]
and choline oxidase capillary with Ag paste and epoxy 0.05 to 3 mM of acetylcholine chlo-
resin, washed. Electrochemical pre- ride. A steady-state response was
treatment at 1.2 V dipped into an reached in 2 min. Calibration graph
aqueous solution of ChO and BSA for purified AChE was linear up to
containing polyvinyl alcohol/styryl 50 iu/mL. Results agreed with the
pyridinium polymer, exposed to spectrophotometric method. Sensor
fluorescent light (300–460 nm), was stable for 1.5 months on storage
dip-coated in 5% Nafion solution. at 5 C.
Measurements for AChE were
made at 30 and 1.2 V in 0.1 M-
2-(N-morpholino) ethanesulfonic
acid buffer of pH 7.1 containing
ACh, chloride.
Acetylcholineesterase Co-immobilized enzyme (AChE and Linear response in the range [83]
and choline oxidase ChO) on 7 mm diameter carbon- 0.1–1 mM of ACh. Response
fiber electrode entrapped with time was 5 s.
polyvinyl alcohol quaternized stil-
bazole. Sensor was used as an
amperometric detector for ACh.
Acetylcholineesterase Immobilized by cross linking with Detection limit for ACh and Ch were [84]
and choline oxidase glutaraldehyde vapor on the 20 and 10 pmol, respectively. Sensor
exposed end (diameter 0.2 mm) of was used successfully in the FIA and
a silica-sleeved Pt electrode. for detection in HPLC.
Acetylcholineesterase, Immobilized enzyme on to the Detection limits for ACh was [85]
urease, glucose sensor chip by corsslinking 0.07 mM with corresponding
oxidase and butyryl with glutaraldehyde and BSA. sensitivity of 5.6  0.2 mS/mM.
chloinesterase. Conductivity changes, produced by The device could be also used
the enzyme-catalyzed hydrolysis of for apparent Michaelis
ACh were measured for the constant determination.
analysis.
Acetylcholineesterase Miniaturized multichannel transduc- Optimum pH range for the sensor was [86]
tor with planar Au electrode which 7–9. The calibration graph was
was first covered with a choline- linear from 0.02–10 mm ACh and
selective liquid membrane made detection limit was 5 mM. Response
from 66% PVC-polyvinyl acetate time was 3–5 min. Sensor was sui-
(PVA), 33% 2-nitrophenyl octyl table for determination of ACh in
ether plasticizer and 1% ion-pair biological fluids.
choline phosphotungstate. A sec-
ond layer of 2% AChE in the
PVA-polyethylene dispersion was
spread on the top. The electrode
was used as working electrode
versus Ag/AgCl for potentiometric
measurement of Ch and ACh in
0.1 M Tris buffer at 7.4.
Acetylcholineesterase Co-immobilizing AChE and ChO on The microsensor exhibited a linear [87]
and choline oxidase to a Pt disc microelectrode (200 response for acetylcholine and
mM diameter) with glutaraldehyde choline for 0.05–103 pmol.
vapor. Sensor was used in flow-
injection and LC system at a
potential of 0.6 V versus Ag/AgCl/
KCl (saturated) with mobile phase
containing 0.1 M-phosphate buffer
(pH 8) (for FIA) and 0.05 M phos-
phate buffer at pH 7.5 containing
0.03 mM SDS and 3 mM
tetramethyl ammonium chloride
(for LC). The LC separations were
carried out on an ODS-5 column
(25 cm  0.5 mm i.d.).
Acetylcholineesterase A Cross-linkable polymer, poly The microsensors if they can be suc- [88]
and choline oxidase (vinyl pyridine) derivatized at the cessfully used in vivo will provide
N atoms with a combination of valuable information for brain
iron-linking and redox functional- diseases (Parkinsion's and
ities was used to immobilize the Alzheimer's).
enzymes and to shuttle electrons.
Enzymes were deposited with the
polymer and deposited onto C elec-
trode. For peroxide selectivity over
ascorbate is achieved by incorpora-
tion of Nafion.
Acetylcholineesterase A rotating graphite-disc electrode Calibration graphs were linear from [89]
and choline oxidase was polished, defatted, cleaned 0.6–10 mM substrate. RSD were
and oxidized by immersion in 5% (n ¼ 10). The detection limit
10% HNO3/2.5%, K2Cr2O7 at was 10 mM acetylthiocholine,
2.2 V versus SCE for 10 s. electrode response time was 15 s.
AChE was covalently immobilized
on to the electrode using a
standard method. Measurements
were made in 0.1 M-universal
buffer of pH 7 at 0.8 V versus
SCE.
Acetylcholineesterase A 350 mM diameter coherent The response time of the pH sensor [90]
imaging fiber coated on the distal was 2 s for a 0.5 unit increase in
surface with a planar layer of ana- pH. The biosensor had a detection
lyte-sensitive polymer that was thin limit of 35 mM ACh and a linear
enough not to affect the fiber's response in the range 0.1 mM.
imaging capabilities. It was applied
to a pH sensor array and an ACh
biosensor array (each contain 6000
optical sensor). Fibers were coated
with an immobilized layer of poly
(hydroxyethylmethacrylate)-N-
flurosceinylacrylamide and AChE-
fluorescein isothiocyanate isomer
poly (acryloamide-co-N-acryl
oxysuccinimide), respectively.
Acetylcholineesterase 300 mL 0.1 M phosphate buffer (pH Calibration graphs were linear upto [91]
and choline oxidase 6.5) containing 16 mg BSA and 0.09 and 0.08 mM Ch and ACh,
1 mg each of ChO and AChE were respectively. Detection limits were
mixed with 30 mL 25% glutaral- 0.1 mM of both Ch and ACh.
dehyde diluted 10 fold with phos- Response time was 1 s for both Ch
phate buffer. The solution was and ACh. The use of the sensor as
used to coat the surface of a Pt detector for HPLC analysis for both
electrode. This enzyme electrode Ch and ACh was demonstrated.
was used for the amperometric
measurement of ACh and Ch.
Acetylcholineesterase Au foil was treated with cystamine Electrical communication between the [92]
and choline oxidase to produce a base layer of ami- enzyme and the electrode is achieved
nothiolate units, was derivatized either by the use of ferrocenecar-
by reaction of the amino group and boxylic acid as mediator in the
disodium-4,40 -diisothiocyanato- assay buffer or by immobilization
trans-stilbene-2,20 -disulfonate. of [(ferrocenyl methyl)amino] hexa-
Enzymes were immobilized at the noic acid on the enzyme layer.
isothiocyanate group via
thiourea link. The bifunctional sen-
sor for ACh was prepared by step-
wise immobilization of four layers
of the enzyme ChO and three
layers of AChE. Choline generated
was detected amperometircally
with the use of 2,6-dichloro-
phenolindophenol as a mediator in
Nicotinic Receptor, isolated from electro- The response of the biosensor reached [93]
acetylcholine phorus electricus, was incorporated equilibrium within 5 s for ACh and
receptor into a polymeric film formed in situ remained stable up to 20 min.
on an electronic transductor con- Detection limits  25 ng in a 50 mL
taining two terminal 1.5 cm  1.5 sample (i.e., 0.5 mg/mL).
cm interdigitated gold electrode.
Acetylcholineesterase Coated wire enzyme electrode was Three different methods for immo- [94]
and choline oxidase prepared by coating a Ag wire bilizing the enzyme on the coated
with a homogenized solution of electrode are described with use of
CH2Cl2 containing 0.28 gm of AChE. The model electrode was
polyvinyl bytyral, 0.15 gm of di-n- used to determine ACh from 0.1 to
amylphthalate, 10 mg of tridodecy- 10 mM. Response time was 3–6 min.
lamine and 2 mg of sodium
tetraphenylborate.
Acetylcholineesterase Enzyme membrane electrode was Electrode response was linear for sub- [95]
prepared galvanostatically on a strate over the range 0.1–1 mM. The
stationary Pt and vitreous C disc RSD of the AChE electrode in the
electrode at a current density of determination of 0.1, 0.5 and
0.5 mA/cm2 for 45 s in a solution 1.0 mM ACh were 5.3, 5.0, and
containing 0.4 mL of enzyme solu- 4.8%, respectively.
tion, 0.02 of 25% glutaraldehyde,
0.02 mL ethanol and 0.02 mL of
HCl.
Acetylcholineesterase Pt microelectrode, precoated with The calibration graph was linear from [96]
and choline oxidase cellulose acetate, dipped in a buf- 0.5 mM to 100 mM. Response time
fered solution (pH 6.89) of choline was 15–20 s. The optimum pH was
oxidase or AChE containing glu- 9–10.5. Electrode was stable over
taldehyde as cross linking agent. 41 days.
Acetylcholineesterase Enzyme immobilized over tetra- An applied potential of þ 100 mV [97]
and choline oxidase thiafulvalene tetracyanoquinodi- versus SCE (Pt-wire auxiliary
methane crystals packed into a electrode) and a carrier flow rate of
cavity at the tip of a carbon-fiber 1 mL/min. The Ch and ACh biosen-
electrode. The immobilization sors exhibited linear response upto
matrix consisted of dialdehyde 100 mM and 50 mM, respectively.
starch/glutaraldehyde, and the sen- Response times were 8.2 s.
sor was covered with an outer
Nafion membrane. The ampero-
metric performance of the sensor
was studied with the use of FIA
Acetylcholineesterase Screen-printed electrodes were pre- Responses to H2O2 at þ 700 mV [98]
pared by disposition of a Ag con- versus SCE were linear for 5–100
ductive pad, a C pad and an mM and the RSD was 6.7%.
insulating layer on a polyester Acetylcholine determination was
film substrate. A graphite-based performed by adding AChE to the
ink mixed with 0.5% Ru on activ- electrolyte. Best results were
ated C was used to print the work- achieved using borate buffer of pH 9
ing surface and the electrodes were containing 0.1 M KCl.
heated at 110 C for 10 min.
Acetylcholineesterase Packing a thin layer of a glass bead The calibration graph was linear for [99]
mixture comprising AChE cova- 2.5–25 mM acetylcholine and RDS
lently immobilized on preactivated were <2%; the response was
isothiocyanate glass and thymol reversible.
blue indicator bound on aminopro-
pyl glass on the tip of a bifurcated
optical fiber sensor head. The sen-
sor was incorporated into a flow
cell and used to detect carbamate
and organophosphate pesticides.
Acetylcholineesterase A dual syringe pump, one contain- The H2O2 generated in the first com- [100]
and choline oxidase ing choline solution in 50 mM partment by the enzymatic reaction
phosphate buffer of pH 8 in the of choline was consumed in the sec-
presence of 0.5 mM EDTA. The ond compartment. The unreacted
other containing the sample solu- acetylcholine present in the sample
tion consisting of a mixture of cho- reacted in the third compartment
line and acetylcholine in the same and the generated H2O2 was
buffer solution. The two solutions detected by an osmium-gel-HRP
were pumped via a microdialysis modified vitreous C electrode set at
unit into an enzymatic reactor con- 0 V (versus Ag/AgCl). The calibra-
taining three separate compart- tion graph was linear up to 1 mM
ments filled with ChO, catalase acetylcholine.
and AChE/ChO mixture, respec-
tively.
Acetylcholineesterase A glassy C electrode surface was The detection limit was 4.8 nM, com- [101]
and choline oxidase modified with osmium poly (vinyl- parable to that obtained by LC with
pyridine) redox polymer contain- electrochemical detection combined
ing horseradish peroxidase (Os- with an enzyme reactor. In the
gel-HRP) and then coated with a online measurement of acetylcholine
co-immobilized layer of AChE standard solutions, the sensitivity of
and ChO. A 22 mL pre-reactor, in the biosensor was 43.7 nA/mM at a
which ChO and catalase were flow rate of 16 mL/min.
immobilized on beads in series,
was used to remove choline. The
variation in extracellular concen-
tration of ACh released from rat
hippocampal tissue culture by elec-
trical stimulation was observed
continuously with the online bio-
sensor combined with a micro-
capillary sampling probe.
Measurement of ACh and Ch was
carried out by using a split disc C
film dual electrode.
Acetylcholineesterase A 3 mm diameter, Pt disc electrode, The amperometric response of the [102]
either polished or black, was sensor was measured with a three-
immersed for 20 min in a solution electrode cell at 0.6 V versus Ag/
of avidin (100 mg/mL). After wash- AgCl in 0.1 M phosphate buffer of
ing with PBS solution of pH 7.4, pH 6.8. The response was linear
the electrode was immersed for 20 from 1 mM to 1 mm of acetylcholine.
min in a biotin-labeled ChO solu-
tion (100 mg/mL). After treatment
10 times, the surface of the sensor
was further modified with ChE in
the same manner.
Acetylcholineesterase A stock solution of 0.52 mg/mL of A linear calibration plot was obtained [103]
the pesticide trichlorophen in in the range of 10 ppb–10 ppm with
10 mM phosphate buffer of pH a lowest detection limit of 0.1 ppb.
7.5 was diluted with buffer to var-
ious concentrations. The obtained
solutions were then analyzed using
an ACh biosensor based on the
inhibition effect of trichlorophen
on the function AChE which pro-
motes the hydrolysis of the natural
neurotransmitter, acetylcholine.
The sensor was fabricated by
immobilizing AChE onto the sur-
face of an antimony disc electrode,
which was then used in conjunction
with a double junction Ag/AgCl
(0.1 M-KCl) reference electrode
with a 0.1 M lithium acetate salt
bridge.
Acetylcholineesterase The detector consisted of two Pt Rectilinear calibration graphs for [104]
and choline oxidase electrodes (6 mm  3 mm) sand- 10–100 mM choline and
wiched between Perspex sheets acetylcholine were obtained.
and separated by a 1 mm thick
sheet of silicone rubber, and the
carrier stream (0.5 mL/min) was
0.1 M phosphate buffer (pH 8.2).
AChE and ChO were immobilized
by glutraldehyde cross-linking to
controlled-pore glass and packed
into columns (3 cm  2.5 mm)
that were operated at 25 C.
Nicotinic acetylcho- The biosensors were constructed The detection range was 0.1–10 mM [105]
line receptor with poly (vinylbutyral) mem- acetylcholine.
branes incorporating nicotinic
acetylcholine receptor for the
determination of ACh.
Acetylcholineesterase Biosensors were fabricated from BLM containing AChE was used for [106]
filter-supported solventless bilayer the analysis of acetylcholine follow-
lipid membrane (BLM) and used ing injection of the substrate into a
for the analysis of the substrates 0.1 M KCl/10 mM HEPES carrier
of hydrolytic enzymes in a flow- electrolyte. The enzymatic reaction
through system. The codeposition at the membrane surface caused
of lipid (dipalmitoyl-phosphatidic changes in the electrostatic fields
acid) and protein solutions to and phase structure of the BLM,
form a BLM on a microporous resulting in ion current transients;
glass fiber or polycarbonate ultra- the magnitude of these were linearly
filtration membrane disc was related to the substrate concentra-
described. Enzyme was immobi- tion down to the mM level.
lized on the membrane by incor- The RSD were 5%.
poration of protein solution
into the lipid matrix at the
air-electrolyte interface before
BLM formation.
Acetylcholineesterase The aqueous fermentation Acetylcholine iodide was studied in [107]
solution containing cholineesterase 15 mM NaCl and 13 mM borate
(90–100 mg/mL) was added to buffer of pH 9–10. Detection limit
nitro-cellulose in butyl acetate for acetylcholine was 40 mM.
(30 mg/mL). From 1 to 5 mL of Sensitivity losses was  40% over
the mixture was spotted onto 20 days.
a small area of FET and dried
in the air. The FET was
treated with 5% of glutaraldehyde
for 5–10 min, washed with
10 mM borate buffer (pH 9.18)
for 15 min, dried and stored
at 4 C.
Acetylcholineesterase The ISE biosensors were fabricated A thin-film composed of 40% plyur- [108]
and choline oxidase from membranes containing 1.2% ethane/59.5% bis(1-butylpentyl)-
of 2,6-didodecyl b-cyclodextrin/ adipate/4.8% 2,3,6-triethyl-b-
65.6% O-nitrophenyloctyl ether/ cyclodextrin/0.6% TKB was cast
0.4% sodium tetrabis-3,5-kis-(tri- over the enzyme layer. Acetylcholine
fluoromethyl)phenyl borate (TKB) was determined at 0.18 V versus
and 32.8% PVC or polyurethane. Ag/AgCl. Detection limit was at the
The biosenser was prepared by sub-pM level. Interference with
coating a screen-printed C working other electroactive compounds was
electrode with 1,10 -bis-(methoxy- minimal.
methyl) ferrocene followed by
application of a horseradish perox-
idase/AChE/ChO solution.
Acetylcholineesterase A glassy carbon electrode (GCE) Significant enhancement in the per- [109]
and choline oxidase modified by electrodepositing sub formance of these biosensors was
mm Pt-black particles on the sur- achieved. The chronoamperometric
face. ACh and Ch. micro biosensor response of 1 m biosensor array was
arrays were fabricated based on linear from 29 to 1200 mM with
immobilization of AChEChO or detection limit of 8.7 mM
ChO by cross linking with gentar- acetylcholine.
aldehyde on Pt-black GCE.
Acetylcholineesterase An acrylohydrazide-methacrylo- Potential measurements were made [110]
hydrazide pre-polymer was with a Radiometer PHM 65 pH
prepared and an aqueous 30% meter and recorder with use of
solution was added to a solution of HEPES buffer containing MgCl2,
AChE in phosphate buffer contain- NaCl, and gelatin as sample buffer
ing gelatin. The enzyme-copolymer and acetylcholine as substrate. The
solution was applied to the tip of a detection limits for nicotine and the
glass electrode, dried and cross- organophosphorus compound were
linked. Other sensors based on given.
pH-sensitive PdPdO and IrIrO2
electrodes were also prepared with
the enzyme and copolymer.
Biological tissues The electrode was constructed from The calibration graph of potential [111]
a slice of pig muscle (0.9–1 mm versus concentration was rectilinear
thick), stuck over the flat end of a from 20 to 600 mg/mL of acetylcho-
glass pH electrode and covered line in 1 mM. KCl at 30 C. There
with a nylon net fixed with a was no interference from a wide
rubber ring. range of biological compounds.
Recovery was 98.2% and the coeffi-
cient of variation was less than 8%
(n ¼ 8).
Nicotinic receptor A silicon-based light addressable The sensor could be applied to deter- [112]
ligands potentiometric sensor was applied mine agonists and antagonists.
to detect biotinylated -bungaro- Allows the detection of nicotinic
toxin bound to carboxy receptor ligands with a light address-
fluorescein-nicotinic acetylcholine able potentiometric sensor. After the
receptor conjugate. This conjugate addition of streptavidin solution, the
(1–20 ng) was incubated for mixture was filtered through the bio-
1 min with 20 nM-biotinylated tinylated nitrocellulose capture
-bungarotoxin in assay membrane.
buffer solution containing
150 mM NaCl, 0.1% bovine
serum albumin and 0.05%
Triton X-100 in 10 mM
Na2H-NaH2PO4 (pH 7).
Acetylcholineesterase Bilayer lipid membranes were pre- Enzymatically generated hydronium [113]
pared by adding a solution of egg ion causes transient current due to
phosphatidylcholine and dipalmi- alteration of the electrostatic field
toyl phosphatidic acid dropwise by the ionization of dipalmitoyl
into the surface of aqueous 0.1 M phosphatidic acid. The response
KCl/10 mM HEPES, near the delay time was directly related to
Saran Wrap partition of a two the substrate concentration where
compartment plexiglass cell. A acetylcholine can be determined
portion of AChE solution in from 1 mM upto mM level.
10 mM Tris hydrochloride buffer
solution of pH 7.4 was applied.
The electrolyte level was momentar-
ily dropped below the orifice and
raised to form a membrane. The
membranes were used as trans-
ducers for the reaction of AChE
with ACh. An external voltage
(25 mV) was applied across the
membrane between two Ag/AgCl
reference electrodes.
Acetylcholineesterase Immobilization of AChE on the sur- The acetylcholine concentration was [114]
face of a glass electrode is achieved found from a calibration graph
by coating the electrode with a which was rectilinear from 0.1 to
AChE-bovine serum albumin mix- 100 mM. The detection limit was
ture and gelatin, and successive 0.01 mM and the response time
treatment with a solution of glutar- was 1 to 3 min. The electrode is
aldehyde and a mixture of glycine intended for the application to
and lysine. The electrode was biological materials.
applied in the determination of
ACh in phosphate buffer pH 8 at
Acetylcholineesterase Sensors were developed consisting of Free choline is measured with a simi- [115]
and choline oxidase a Clarke oxygen electrode modified lar electrode with only choline oxi-
by superposition (over the polypro- dase immobilized on the membrane.
pylene membrane of the electrode) Calibration graphs were rectilinear
of a dialysis membrane on which upto 90 mM choline and upto
AChE and ChO were immobilized. 120 mM acetylcholine. Results were
Both formats are based on the con- presented for the analysis of tissue
version of the substrate to choline extracts and pharmaceutical
which is oxidized in the presence of formulations.
choline oxidase causing a reduction
in pO2 which is detected by the
electrode.
Acetylcholineesterase Amniotic fluid was mixed at Rapid and sensitive discriminating [116]
and choline oxidase 27 C with phosphate buffer determination of AChE activity in
(pH 8) and the current was amniotic fluid with choline sensor.
measured with an electrode The calibration graphs were recti-
consisting of an immobilized linear and the coefficient of varia-
choline oxidase membrane at tion was 2.5%.
650 mV versus Ag/AgCl.
After reaching the steady
state, acetylcholine or
butyrylcholine was added
and the current was
measured.
Acetylcholineesterase Microdialysates were obtained from Detects the basal acetylcholine in rat [117]
and choline oxidase the brain of rats by fitting a can- brain microdialyzate. The detection
nula directly into the striatum and limit obtained by this method was
perfusing with Ringer's solution 10 fmol for acetylcholine. The
(2 mL/min) for 5 h. Fractions were average basal striatal concentration
automatically collected and ana- of acetylcholine was 31 fmol/5 mL
lyzed by liquid chromatography. dialysate.
The eluate was passed through a
10 mM F 8903 enzyme reactor col-
umn containing AChE and ChO
and the H2O2 produced from
ACh by the enzymatic reactions
was detected at vitreous C
electrode operated at þ 100 mV
versus Ag/AgCl.
Acetylcholineesterase Acetylcholineesterase was immobi- Average recovery was 99.5% with [118]
lized on acetyl-cellulose activated RSD of 2.18%. The static and
by NaIO4 to give an enzyme mem- dynamic linear response
brane which was bound to a pH ranges were from 240 to 1300 and
sensing membrane containing 4,40 - 300–1700 mg/mL acetylcholine,
bis-(N,N-didecylaminomethyl) respectively. Optimum measurement
azobenzene of an original medium was 1 mM KCl.
Acetylcholineesterase The sample was diluted with water Enzymatic determinations with rotat- [119]
and choline oxidase and the resulting solution was ing bioreaction and continuous flow
pumped into a flow-through cell stopped flow processing for choline
containing a rotating (1064 rpm) esters in pharmaceuticals. The initial
Teflon disc with AChE and ChO rate of response was measured and
co-immobilized on the top surface used for quantitation of succinylcho-
of the disc. Measurements were line and acetylcholine. The calibra-
made by stopping the flow for 60 s. tion graph was linear from 0.1 to
The H2O2 produced from the enzy- 1.6 mM for acetylcholine.
matic reaction was detected amper-
ometrically at a Pt ring working
electrode located above the
rotating Teflon disc and
held at þ 0.6 V versus Ag/AgCl.
Acetylcholineesterase The surface photovoltage technique The detection limits of the substrates [120]
was applied to the fabrication of a were 9  107 M, 2.7  106 and
biosensor based on immobilized 4.1  106 for butyrylcholine
cholineesterase. The type of the iodide, acetylcholine iodide and
cholineesterase used depends on the acetylcholine chloride, respectively.
type of substrate. On the surface of The activity of the cholinesterase
the silicon wafer the cholineesterase was inhibited by the presence of
layers were immobilized by using alkaloids such as physostigmine and
3-aminopropyl triethoxysilane and neostigmine.
glutaraldehyde.
Characteristics of the sensor were
studied in phosphate buffered
saline containing 15 mM NaCl and
1 mM phosphate buffer (pH 7).
Acetylcholineesterase, Different nonconducting polymers The procedure of immobilization was [121]
choline oxidase and were synthesized on the surface of fast, simple with better long-term
butyrylcholine a Platinum electrode to assemble stability and higher loading of the
esterase fast-response and sensitive enzyme. Studies on reproducibility,
amperometric biosensor for acetyl- life time of immobilized enzyme,
choline choline and butyrylcholine pH, temperature, interference,
based on ChO, AChE or butyryl- buffer, response time, storage and
choline esterase co-immobilized operational time of the biosensor
by crosslinking with BSA and were carried out.
glutaraldehyde. Also, the
electropolymerization conditions
such as scan rate and time were
optimized for each monomer
in order to obtain permselectivity,
the highest rejection of interference
and stability.
Acetylcholineesterase The detector fabrication of glutaral- A liquid chromatography detector [122]
and choline oxidase dehyde co-crosslinking of AChE based on a fast response and sensi-
and ChO with bovine serum albu- tive bienzyme amperometric biosen-
min on the Pt working electrode of sor for ACh and Ch was described.
a conventional thin layer electro- Linear responses were observed over
chemical flow cell. A mobile at least four decades and the abso-
phase of phosphate buffer 0.1 M; lute detection limit were 12 and 27
pH 6.5 containing 5 mM-sodium fmol injected for choline and acetyl-
hexane sulfonate and 10 mM-tetra- choline, respectively.
methylammonium phosphate was
used in the ion-pair, reversed
phase liquid chromatography.
Acetylcholineesterase The electrochemical sensor for acet- The typical response curves for the [123]
and choline oxidase ylcholine/acetylthio choline and detection of choline using mediators
choline/thiocholine are developed and choline oxidase-modified elec-
using two enzyme reactors: AChE trodes below 0.24V versus silver/
encapsulated organically modified silver chloride in 0.1 M Tris–HCl
sol-gel glass and choline oxidase buffer pH 8 are reported.
immobilized within mediator Comparative analytical performance
[tetracyanoquinodimethane, on the mediated electrochemical
tetrathiafulvalene and dimethylfer- responses of the biosensors is
rocene]-modified graphite discussed.
paste electrode. The AChE
immobilized into organically
modified sol-gel glass behaves
as the reactor for enzymatic
hydrolysis of ACh into
choline whereas mediator and
choline oxidase modified
paste electrode are used for
the detection of choline
through mediated mechanism.
Horseradish A three enzyme layered assembly on Acetylcholine was sensed by the tri- [124]
peroxidase (HPP), Au electrodes or Au quartz crystal, component enzyme layered electrode
choline oxidase and consisting of HRP, ChO and using Faradic independence spectro-
acetylcholineesterase AChE is used to sense ACh by scopy, Cyclic Voltammetry and
the HRP-mediated oxidation of Microgravimetric quartz crystal
3,30 ,5,50 -tetramethyl benzidine (1) microbalance transduction methods.
by H2O2 and the formation of the Acetylcholine concentrations
insoluble product (2) on the respec- correspond to 1  105 M are easily
tive transducers. Acetylcholine is sensed by the different transducers.
hydrolyzed by AChE to choline
that is oxidized by ChO and O2 to
yield the respective betaine and
H2O2. The amount of generated
H2O2 and the resulting insoluble
product on the transducer corre-
lates with the concentration of
acetylcholine in the samples.
monitoring of the pH. The end point was detected using a derivative
potential time response curve, and differentiation and smoothing were
carried out using the Stavitzky-Golay algorithm. The experiment was
carried out in a triangle-programmed coulometric flow titration system.
Calibration graphs were established and the detection limit was 80 mM
for acetylcholine.
3.5 Flow injection method
Yao et al. reported a flow injection analytical system for the simultaneous
determination of acetylcholine and choline that made use of immobilized
enzyme reactors and enzyme electrodes [25]. Acetylcholineesterase–
choline oxidase and choline oxidase were separately immobilized by
reaction with glutaraldehyde onto alkylamino-bonded silica, and
incorporated in parallel as the enzyme reactors in a flow injection
system. The sample containing acetylcholine and choline in 0.1 M
phosphate buffer (pH 8.3) carrier solution was injected into the system.
The flow was split to pass through the two reactors, recombined, and
mixed with 0.3 mM K4Fe(CN)6 reagent solution before reaching a
peroxidase immobilized electrode. Because each channel had a different
residence time, two peaks were obtained for choline and total acetylcho-
line and choline. Response was linear for 5 mM–0.5 mM choline, and for
5 mM–1 mM acetylcholine plus choline. The detection limits were 0.4 mM
for choline and 2 mM for acetylcholine.
3.6 Amperometric methods
Gunaratna and Wilson reported the optimization of the multi-enzyme
flow reactors for the determination of acetylcholine [26]. After separation
of acetylcholine and choline by HPLC (Nucleosil C18), the analytes were
eluted into a flow reactor containing immobilized acetylcholineesterase
(which hydrolyzed acetylcholine to choline). Choline oxidase then is used
to convert choline into H2O2, which is then determined amperome-
trically. Of several immobilization techniques investigated, immobiliza-
tion through the avidin–biotin linkage gave the highest activity of the
immobilized enzyme, leading to improved detection limits. Better
sensitivity and detection limits were obtained with both enzymes
immobilized together on the same support through the avidin–biotin
linkage than with separate immobilization in two columns. The post
column system was applied to the determination of acetylcholine and
choline in brain tissue extracts, and linear calibration graphs were
obtained from 0.1 to 500 pmol.
Yao et al. described an online amperometric assay method for the
determination of acetylcholine using microdialysis probes and immobi-
lized enzyme reactors [27]. The probe consisted of a fused silica tube
(350 mm i.d.), in which another two fused-silica tubes (75 mm i.d.) were
inserted to serve as inlet and outlet. A regenerated cellulose dialysis fiber
(200 mm i.d.) was fixed to the interior of the first tube. A stream (2 mL/
min) containing 147 mM sodium chloride, 4 mM potassium chloride,
1.2 mM calcium chloride, and 1 mM magnesium chloride was pumped
through the microdialysis probe, which was immersed in the analyte
solution. The dialysate was merged with the buffer stream (0.1 M
phosphate buffer of pH 7 or 0.1 M borate buffer pH 8) and the mixed
stream was pumped through a PTFE reactor coil (20 mm  0.5 mm i.d.)
packed with controlled pore glass immobilized with a mixture of
acetylcholineesterase and choline oxidase, for the determination of
acetylcholine. Choline was removed from the dialysate. The resulting
H2O2 was detected amperometrically using Pt disc electrode modified
by coating with a film of poly(1,2-diaminobenzene). The detection limit
was 1.5 mM for acetylcholine.
4. SPECTROPHOTOMETRIC METHODS OF ANALYSIS
4.1 Infrared spectrophotometric method
Mynka et al. reported a method for the infrared spectrophotometric
identification and the determination of acetylcholine in the presence of
other esters in pharmaceuticals [3]. Acetylcholine was determined at the
maximum absorbance of 1755 cm1. Beer's law was obeyed in the
concentration range 1–10 mg/mL, and the relative standard deviation was
less than 2.7%. Results obtained were reported to be reliable and
reproducible.
4.2 Photometric methods
Eksborg and Persson reported a photometric method for the determina-
tion of acetylcholine in rat brain after selective isolation by ion-pair
extraction and microcolumn separation [28]. The same authors also
reported a photometric method for the determination of acetylcholine
and choline in brain and urine samples after selective isolation by ion-pair
column chromatography [29].
Barcuchova et al. reported an extractive photometric method for the
determination of choline and its derivatives that made use of
bromothymol blue [30]. The sample, containing the iodide of the choline
derivative, was treated with a solution of bromothymol blue in pH 6.5
phosphate buffer medium. The aqueous solution was extracted with
chloroform, and the absorbance of the extract was measured at 416 nm
against a reagent blank.
4.3 Mass spectrometric method
Suzuki and Haug compared the use of magnetic switching selected ion
monitoring with conventional voltage switching selected ion monitoring
for applicability to acetylcholine and choline analyses in brain samples
[31]. Deuterated and nondeuterated acetylcholine and choline derivatives
were analyzed by mass spectroscopy with magnetic and voltage switching
selected ion monitoring of dimethylmethyleneimmonium ions, m/z 58, 60
and 64. With both monitoring methods, a maxima peak was obtained
at 400 ms/ion, but the smallest variation was obtained at 100 ms/ion.
The method is reported to be an alternative method to voltage switching
selected ion monitoring.
Lehmann et al. determined choline and acetylcholine levels in distinct
rat-brain regions using a stable-isotope dilution and field-desorption
mass spectrometry method [32]. Tissue samples (20 ng) were homo-
genized at 0 C for 30 min with 4% HClO4 (1 mL), including inter-
nal standards of 2H2 choline and 2H9 acetylcholine (10–11 mL).
Each homogenate was centrifuged for 20 min, and 350 mL of the
supernatant solution was transferred to a Reacti-vial with 1.5 mL of
aqueous 0.5 MNa2HPO4 and 1 mL of dipicrylamine in CH2Cl2. After
stirring the mixture for 20 min, the upper layer was discarded and 1 mL of
carbonate buffer solution (pH 9) was added. The lower phase (700 mL
portion) was transferred to another vial, the solvent removed by a
stream of nitrogen, and the residue dissolved in 50 mL of methanol.
Aliquots (5 mL) were subjected to cyclic scans at m/z ¼ 104 and 106 for
choline and at m/z ¼ 148 and 155 acetylcholine (and the deuteron-
analogues). The signal was accumulated through the use of multi-channel
analyses.
4.4 Colorimetric methods
Tsubouchi reported a colorimetric procedure for the analysis of acetyl-
choline [33]. About 2–10 mL of a test solution containing acetylcholine is
extracted into 1,2-dichloroethane in the presence of the ethanolic reagent
(2 mL at 5 mM) and pH 10 0.1 M Na2B4O7–0.2 M Na2HPO4 buffer
(5 mL) in a total volume of 25 mL. The absorptivity of the extract was
measured at 615 nm.
Barletta and Ward described a procedure for the spectrophotometric
analysis of acetylcholine levels in plasma [34]. 0.3 g of K2HPO4 and 0.3 g
of Na2CO3 are added to a sample of plasma (0.5 mL) containing 4–20 ng
of acetylcholine. To this solution is added 0.08% bromophenol blue
solution in 30% K2HPO4 solution (0.5 mL), and the resultant extracted
with 1,2-dichloroethane containing 4% isoamyl alcohol. One then
measures the absorptivity of the organic phase at 600 nm. The
absorbance derived from a similar treatment of a sample of plasma
taken before the administration of angiotensin II is measured. The
change in the concentration of acetylcholine in turtle blood following
the administration of angiotensin II is measured by a modification of the
method of Mitchell and Clark [35].
Das and Abdulla reported a modified method for estimation of
acetylcholine [36]. Acetylcholine reacts with hydroxylamine in alkaline
medium to give hydroxamic acid, which produces a red–violet color when
reacted with FeCl3. Absorbance measurements are made at 540 nm. The
method can be applied to saliva samples stored at 0 C for as long as 72 h.
Abernethy et al. reported an enzymatic method for erythrocyte
acetylcholineesterase [37]. Haemolysate 50 mL was mixed with 67 mM
phosphate buffer of pH 7.4 (2 mL) and 21.5 mM acetylcholine (50 mL),
and the mixture incubated for 20 min at 37 C. A color reagent (500 mL)
containing 4 mM phenol, 0.8 mM aminoantipyrine, peroxidase (5 iUL),
choline oxidase (4 L iU/L), and 0.86 mM physostigmine plus the buffer
was added and, after 20 min the absorbance was measured at 500 nm.
Sakai et al. reported a novel flow injection method for the selective
spectrophotometric determination of acetylcholine using thermochro-
mism of ion associates [38]. Samples (0.14 mL) containing acetylcholine
were injected into a flow injection system with a buffered (pH 11) carrier
stream and a reagent stream (10 mM tetrabromo-phenolphthalein ethyl
ester in dichloroethane) at 0.8 mL/min. The temperature of the flow cell
was 45 C which reduced interference and improved recovery, and the
detection was at 610 nm.
Sakai et al. reported the use of batchwise and flow-injection methods for
the thermo-spectrophotometric determination of acetylcholine and cho-
line with tetrabromophenolphthalein ethyl ester [39]. An aqueous sample
solution was injected into a carrier stream of 0.3 M KH2PO4/0.1 M
sodium borate (adjusted to pH 11 with 1 M sodium hydroxide), which
merged with the extracted solution of tetrabromophenolphthalein ethyl
ester in dichloromethane. The extract was separated with the use of a
PTFE porous membrane and passed through a micro-flow cell at 45 C
before its absorbance was measured at 610 nm.
Takeuchi et al. published a mechanized assay of serum cholinesterase by
specific colorimetric detection of the released acid [40]. The cholinesterase
reaction was carried out on a thermostatted rack at 30 C with a reaction
mixture of serum (10 mL), 50 mM barbitone–HCl assay buffer (pH 8.2;
140 mL), and 12.5 mM acetylcholine solution (50 mL). The solutions were
prepared by programmed needle actions, and a sample blank was also
prepared. The reaction was stopped after 9.7 min by injection of the
mixture into a flow injection analysis system to determine the quantity of
acetic acid formed. The carrier stream (water, at 0.5 mL/min) was
merged with a stream (0.5 mL/min) of 20 mM 2-nitrophenylhydrazine
hydrochloride in 0.2 M HCl and a stream (0.5 mL/min) of 50 mM 1-ethyl-
3-(3-dimethyl-aminopropyl) carbodiimide hydrochloride in ethanol con-
taining 4% of pyridine. The sample was injected into this mixture (pH
4.5), passed through a reaction coil (10 m  0.5 mm) at 60 C, 1.5 M
NaOH was added, and, after passing through a second reaction coil
(1 m  0.5 mm) at 60 C, the absorbance was measured at 540 nm.
4.5 Fluorimetric methods
O'Neill and Sakamoto reported an enzymatic fluorimetric method for the
determination of acetylcholine in biological extracts [41]. Nanomolar
amounts of acetylcholine were determined in perchloric acid extracts of
biological materials (brain tissues) by use of a system containing
acetylcholineesterase, acetyl CoA synthetase, maleate dehydrogenase,
and citrate synthase. The production of NADH2 was stoichiometrically
related to the amount of acetylcholine in the system, and was followed
fluorimetrically. Interfering fluorescent substances in the brain extracts
were removed with acid-washed Florisil.
Sutoo et al. developed a high sensitivity and high linearity fluorescence
microphotometry system for distribution analysis of neurotransmitter
and related substances in the small brain regions [42]. The method makes
use of the fluorescence intensity of not more than 10,000 points in animal
brain slices, which are immunohistochemically and histochemically
stained.
Ricny et al. determined both acetylcholine and choline by flow-injection
with immobilized enzymes and fluorimetric and luminescence detection
[43]. The analytes are injected into a continuous stream of a medium
flowing through a sequence of enzyme reactors containing acetylcho-
lineesterase, choline oxidase, and peroxidase. Additional reactors with
choline oxidase and catalase are used to remove endogenous choline from
tissue extracts in which the contents of acetylcholine is to be measured.
Reaction products are detected fluorimetrically or luminometrically. The
limits of sensitivity are about 10 pmol/sample with luminometric and
0.2 pmol/sample with fluorimetric detection.
A fluorimetric assay method for the determination of acetylcholine with
picomole sensitivity was reported by MacDonald [44]. The method is
based on the hydrolysis of acetylcholine to choline and acetate, catalyzed
in the presence of acetylcholineesterase, oxidation of choline to betaine,
and H2O2 in the presence of choline oxidase, and oxidation of
4-hydroxyphenylacetic acid by H2O2 to a fluorescent product, catalyzed
by peroxidase. The interference in the analysis of brain homogenates
was discussed.
Roulier et al. reported a sensitive and specific method for the
measurement of choline and hydrogen peroxide in sea-water [45].
Choline was oxidized by choline oxidase to produce betaine and H2O2.
The latter was used with horse-radish peroxidase to oxidize hydro-
xyphenyl-propionic acid to produce a fluorescent diphenol end product.
The resulting fluorescence at 410 nm (excitation at 320 nm) was
proportional to the amount of H2O2, and could thus be used to measure
the amount of choline present in the sample. Only 2-dimethyl
aminoethanol interfered. The method was optimized, and used to
determine 0–45 nM choline in coastal sea-water.
Brennan et al. used a method to detect the reaction of acetylcholineesterase
with acetylcholine [46]. The method was based on the use of a monolayer,
consisting of fatty acids having C16 chain lengths, which were covalently
attached to quartz wafers and which contained a small amount of
nitrobenzoxadiazole dipalmitoyl phosphatidylethanolamine (NBD-PE)
(partitioned from water into the membrane). The enzyme–substrate
reaction produced a decrease in fluorescence intensity from the monolayer,
and the detection system was sensitive to the changes in bulk concentration
of as small as 0.1 mM, with a limit of detection of 2 mM of acetylcholine.
The mechanism of transduction of the enzymatic reaction was investigated
using spectrofluorimetric methods and fluorescence microscopy.
A kinetic-fluorimetric method for the determination of choline and
acetylcholine by oxidation with cerium (IV) was reported by Lunar et al.
[47]. To sample solutions containing 0.017–1.0 mM choline and/or
acetylcholine were successively added 6 M H2SO4 (5 mL) and 7.1 mM
Ce(IV) solution (0.35 mL), the mixture diluted to 10 mL with water, and
the solution heated to 80 C for 2 min. A portion of the solution was
transferred to a cell maintained at 20  0.1 C, and after 1 min the Ce(III)
fluorescence intensity was measured at 360 nm (excitation at 260 nm) as a
function of time.
4.6 Chemiluminescence methods
Ternaux and Chamoin described an enhanced chemiluminescence assay
method for the determination of acetylcholine [48]. Reaction medium was
prepared by mixing 250 iu/mL of choline oxidase (100 mL), 2 mg/mL of
horse-radish peroxidase (50 mL) and 10–120 mM luminol in 100 mL of
0.1 M Tris buffer (pH 8.6), or 100 mL of 10–100 mM 7-dimethylamino-
naphthalene-1,2-dicarbonic acid hydrazide, for 10 min in 5 mL of 0.1 M
sodium phosphate buffer (pH 8.6). Aqueous 0.325–80 pmol of acetylcho-
lineesterase (50 mL) purified on a Sephadex G50 coarse column was mixed
with 450 mL of reaction mixture, and the chemiluminescence was
measured at 27 C.
Luterotti and Maysinger reported a chemiluminescence method for the
determination of acetylcholine and choline-related substances in phar-
maceutical preparation by dot-blot [49]. The sample (1 mL) was applied as
a dot to a Hybond-N nylon membrane (Amersham), and 1 mL of a
mixture comprising 50 mL of 0.1 M phosphate buffer at pH 8.6, 10 mL of
choline oxidase (25 iu/mL), 2 mL of horse-radish peroxidase solution
(5 mg/mL), and 50 mL of ECL western blotting reagent mixture
(Amersham, 1:1) was added. Hyperfilm ECL chemiluminescence film
(Amersham) was immediately exposed to the luminescing dots for 20 min,
and then the film was developed, washed with water, fixed and washed
again before densitometric evaluation. Acetylcholine was hydrolyzed by
AChE before the analysis. This method can be used also to determine the
sum total of choline and acetylcholine.
Fan and Zhang determined acetylcholine and choline in rat brain tissue
by a fluorescence immunoassay method, making use of immobilized
enzymes and chemiluminescence detection [50]. Tissue was homogenized
with a 10 fold volume of 0.6 M HClO4, the homogenates were kept on ice
for 30 min, and then centrifuged at 2000 G for 20 min. The pellets were
discarded and the supernatants neutralized with 2 M K2CO3 solution (pH
4.2). Portions of the neutralized supernatant were injected into a carrier
stream, consisting of 0.05 M Tris hydrochloride buffer (0.6 mL/min) in a
flow injection manifold, and the stream passed through two reactors
containing acetylcholineesterase and choline oxidase immobilized on
glass beads (30  2 mm i.d.). The resultant H2O2-containing analyte
stream was merged with streams of 1 mM luminol in 0.1 M NaOH and
25 mM CoCl2 (both at 2.5 mL/min) and the chemiluminescence intensity
signal measured.
Hasebe et al. described a simultaneous flow-injection method for the
determination of acetylcholine and choline based on luminol chemilumi-
nescence in a micellar system with online dialysis [51]. The method was
based on the determination of H2O2 produced from acetylcholine and
choline by enzymatic reaction. A water carrier, a 0.02 M disodium
hydrogen phosphate/NaOH buffer (pH 11), a 0.02 M phosphate buffer
(pH 7), a 1 mM Co(II) solution in 0.01 M HCl (the catalyst for the
chemiluminescence reaction), and a mixed solution of 0.5 mM luminol,
0.1 M sodium hydrogen carbonate, and 0.18% SDS in borate buffer
solution (pH 11), were pumped by the three double-plunger pumps. An
800 mL sample was injected into the carrier, dialyzed to remove protein,
and passed through an ion-exchange column to remove other interfering
substances. Choline was converted to H2O2 at another immobilized
enzyme column. The reactions were performed at 37 C. Hydrogen
peroxide produced from choline was passed through a delay coil before
the chemiluminescence of the acetylcholine and choline reaction products
was measured from 350 to 650 nm. This method determines down to
1 mM at a sampling rate of six samples per hour.
Emteborg et al. developed a peroxyoxalate chemiluminescence method
for the determination of acetylcholine in aqueous solution [52]. The
method was used for the enzymatic detection of the generated H2O2 in
FIA and LC separation modes. The analyte stream (at 0.5–0.6 mL/min)
was merged with the reagent stream (1 mM 1,10 -oxalyl diimidazole) in
acetonitrile, and the flow was passed through the detector cell where
the chemiluminescence was detected using a photomultiplier tube. The
chemiluminescence signal was independent of the composition of the
analyte stream over the pH range 6–7. The detection method was applied
to a FIA system for choline and acetylcholine determination, based on
LC separation, and followed by a choline oxidase acetylcholineesterase
enzyme reactor. Choline and acetylcholine were separated using a cation
exchanger with 5 mM tetramethylammonium nitrate in 30 mM phosphate
buffer at pH 7 as the mobile phase. The linear range for both analytes was
50 nM (detection limit) to 10 mM. The FIA system was employed to
determine choline and acetylcholine in diluted urine.
Wetherell and French determined both acetylcholine and choline by
a chemiluminescence method [53]. The optimum conditions for this
determination were determined by using the method of Israel and
Lesbats, in which acetylcholine was first hydrolyzed to choline by
acetylcholineesterase and choline oxidase. The reaction products were
then further reacted by the oxidation of choline to hydrogen peroxide and
betaine, which in the presence of peroxidase oxidizes luminol to
aminophthalate with the emission of light [54]. Linear standard curves
were obtained for acetylcholine and choline in the picomole range. Since
the enzymes differ in their pH optima, a compromise must be made, so
the use of 0.1 M glycine buffer (pH 9.2) is recommended for light-
sensitivity determinations (1–20 pmol) of acetylcholine and choline.
Israel and Lesbats reported a chemiluminescence method for the
determination of acetylcholine, and continuous detection of its release
from torpedo electric organ synapses and synaptosomes [55]. Birman
described a new chemiluminescence assay method for the determination
of acetylcholineesterase activity with the natural substrate [56].
The method involved monitoring the increase in light emission produced
by accumulation of choline, or measuring the amount of choline
A comparison between the chemiluminescence and the radioenzymatic
assay methods that are used for the measurement of acetylcholine
released from a rat phrenic nerve hemidiaphragm preparation was
reported by Ehler et al. [57]. The comparison demonstrated quantitative
equivalency and limits of detection for different analytes as 2 pmol.
5. BIOLOGICAL ASSAY METHODS OF ANALYSIS
A sensitive biological assay method for the assay of acetylcholine was
used by Vapaatalo et al. [58]. Superfused hamster stomach strip was used
in the bioassay of acetylcholine. The preparation proved very stable and
had no spontaneous movement, and the sensitivity to acetylcholine was
in the range of 10–12 g/mL. Satyanarayana et al. reported the use of a
bioassay method for the analysis of acetylcholine using the arterial blood
pressure of cats [59].
Karube and Yokoyama presented an overview on the developments in
the biosensor technology [60]. The overview describes the use of micro-
machining fabrication techniques for the construction of detection units
for FIA, electrochemical flow cells and chemiluminescence detectors.
Acetylcholine microsensors using carbon fiber electrodes and glutamate
microsensors for neuroscience were discussed.
Aizawa presented an overview on the principles and applications of the
electrochemical and optical biosensors [61]. The current development in
the biocatalytic and bioaffinity bensensor and the applications of these
sensors were given. The optical enzyme sensor for acetylcholine was
based on use of an optical pH fiber with thin polyaniline film.
Mascini and Guilbault presented a review covering the methods used for
immobilization of the enzymes, enzyme electrodes, and sensors for
choline and acetylcholine [62]. A number of reports have appeared in the
literature [21, 22, 63–124] describing biosensors used for the detection
and the analysis of acetylcholine. The important characteristics of some
of these reports are summarized in Table 1.
6. CHROMATOGRAPHIC METHODS OF ANALYSIS
6.1 Thin layer chromatography (TLC)
Acetylcholine was separated by Paul and Ebrahimian from adrenaline
and serotonin [125]. The thin layer chromatographic behavior of
acetylcholine and other quaternary ammonium halides were studied on
silica gel by Iorio et al. [126].
6.2 Gas chromatography (GC)
Acetylchloline has been separated and analyzed by many gas chromato-
graphic systems. Table 2 summarizes some of these systems [127–130].
The important characteristics of other gas chromatography systems
[131–140] will be outlined in the following paragraphs.
Budai et al. described a GC method for the isolation and determination
of choline and choline esters from Krebs-Ringer solution [130]. Samples
containing 5–2000 pmol of choline and choline esters were added to 2 mL
of Krebs-Ringer solution containing 120 mM NaCl, 4.7 mM KCl, 25 mM
NaHCO3, 2.6 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, and 10 mM
glucose and gassed with 5% carbon dioxide in oxygen. Glucose was
Gas Chromatographic Systems
Column Detector Temperature Carrier gas Sample and Reference
and [flow rate] remarks
6 ft  0.25 inch of 1% Flame 180 Nitrogen Acetylcholine and [127]
phenyl diethanolamine ionization [45 mL/min] related compounds
succinate on Polypak 1 micro estimation
(80–120 mesh).
1.8  2 mm of Porapak Ni electron- 145–170 Helium or Serum and urine, [128]
Q (80–100 mesh), capture nitrogen 10 fmol can
Chromosorb 101 [20 mL/min] be detected
(80–100 mesh) or
Chromosorb 105
2 m  3 mm of 0.4% Flame — Nitrogen Tests in urine [129]
Carbowax 1500 on ionization by head-space
graphite (60–80 mesh). GLC
A glass silane-treated N sensitive 170 Nitrogen Krebs-Ringer [130]
column 1.8 m  2 mm [40 mL/min] solution
packed with Pennwalt
223 amine packing
removed by chromatography on a column (2 cm  6 mm) of Amberlite
CG-50 (100–200 mesh, H þ form), from which choline and its esters were
eluted with 2 mL of 0.25 M HCl. The eluent was freeze-dried, acetonitrile
(1 mL) was added, and the solution was centrifuged (15 min; 3000 G).
The supernatant solution was evaporated, choline was converted into
propionylcholine by reaction (10 min) with propionyl chloride (5 mL), and
the dried sample containing butyrylcholine (internal standard)
was demethylated (80 C, 30 min) with 200 mL of 50 mM sodium
benzenethiolate and 25 mM benzenethiol in ethyl methyl ketone. After
cooling, 0.5 M citric acid (30 mL) and pentane (300 mL) were added, the
aqueous layer washed with pentane (2  300 mL), and then mixed with
ethyl ether (200 mL) and 2.5 M ammonium citrate in aqueous 7.5 M
ammonia (30 mL). The ether layer was evaporated (40 C) to 25% of
its volume, and a portion (5 mL) was analyzed by gas chromatography
(see Table 2).
Jenden et al. reported the use of a GC method for the estimation of choline
esters in rat brains [131]. Their earlier gas liquid chromatographic method
[132], which was used for determining acetylcholine as the volatile
dimethylaminoethyl acetate, has been applied successfully to the study of
acetylcholine in rat brain and integrated with a mass spectrometric system.
More sensitivity and more efficient recovery were obtained. Green and
Szilagyi measured acetylcholine by pyrolysis gas chromatographic method
[133]. Acetylcholine was extracted from the tissue with acetonitrile
containing propionylcholine as an internal standard, and after centrifuga-
tion the acetonitrile was removed by shaking with toluene–diethyl ether
mixture. A solution of KI–I2 was added to the aqueous solution to
precipitate the quaternary ammonium compound. After being dissolved in
aqueous acetonitrile, the precipitate was passed through an ion-exchange
resin to convert the periodides of the quaternary compounds to chlorides,
which were simultaneously pyrolyzed and analyzed by GC.
Karlen et al. analyzed acetylcholine and choline by an ion pair extraction
and gas phase method [134]. The gas chromatographic estimation of the
isolated acetylcholine and choline was carried out after demethylation
either with benzenethiolate or by controlled pyrolysis. Acetylcholine was
quantitated by flame ionization detection at the nmol level. Mass
fragment analysis was employed for the determination of acetylcholine in
pmol amounts.
Takahashi and Mouharo developed a pyrolysis apparatus which can be
easily connected, and subsequently disconnected, from a conventional
gas chromatograph for the simultaneous determination of acetylcholine
and choline in brain tissue [135]. Rat brain samples were mixed with
propionyl chloride and butyryl choline and heated in the pyrolyzing
system. Thereafter, the propionylated sample was analyzed by its passage
through the attached gas chromatograph. Three peaks with increasing
retention time were observed, corresponding to acetyl-, propionyl-, and
butyryl-choline.
Maruyama et al. reported the use of a simple pyrolysis gas chromato-
graphic method for the determination of choline and acetylcholine in
brain tissue [136]. Schmidt and Speth reported a simultaneous analysis of
choline and acetylcholine levels in rat brain tissue by a pyrolysis gas
chromatographic method [137]. Kosh et al. reported an improved gas
chromatographic method for the analysis of acetylcholine and choline
in rat brain tissue [138]. Mikes et al. used a syringe micro-pyrolyzer for
the gas chromatographic determination of acetylcholine, choline and
other quaternary ammonium salts [139].
Gilberstadt and Russell determined picomolar quantities of acetylcholine
and choline in a physiological salt solution [140]. Carbon-14 labeled
choline and acetylcholine standards in Krebs–Ringer solution were
subject to chromatography on columns (12 mm  8 mm) packed with
Bio-Sil A (200–400 mesh), and it was found that 95–98% of choline and
acetylcholine were retained. Of the bound choline, 84–97% was eluted
in 1.5 mL of 0.075 M HCl, and then 95–98% of the bound acetylcholine
was eluted in 1.5 mL of 0.03 M HCl in 10% butan-2-one.
Findeis and Farwell described a gas chromatographic method for the
determination of acetylcholine and other biogenic quaternary ammonium
compounds on fused-silica capillary columns with a splitless injector as a
chemical reactor-pyrolysis chamber [141]. A Hewlett Packard model 5880
A chromatograph was used with a split/splitless injector, fused silica
column (20 m  0.32 mm) of DB-1 or (30 m  0.25 mm) of DB-1 or DB-
1701 and flame ionization detection. The column was heated at 35 C for
10 min, and then the temperature was programmed to 200 C at 7.5 C/
min. N-Demethylation was achieved at a concentration of 4.3 mg/mL–
1.5 mg/mL by pyrolysis, or by coinjection with methanolic 0.5%
ethanolamine. Studies were also made using gas chromatography.
Cranmer used a gas chromatographic method for the estimation of the
acetylcholine levels in brain tissue by gas chromatography [142].
Acetylcholine was extracted from frozen rat brain with ethanolic
formic acid. The extract was evaporated to dryness at 50 C under
reduced pressure, and the residue was heated at 95 C with 1 M NaOH.
The resulting hydrolyzate was neutralized with 1 M HCl, diluted with
water, and analyzed for its acetic acid content by gas chromatography.
The glass column (6 ft  0.25 in.) was packed with Porapak Q-S (40–50
mesh) and operated at 165 C, using nitrogen (45 mL/min) as the carrier
gas and a hydrogen flame ionization detector. Recovery ranged from 93
to 100% and the method was reported to be simple and rapid.
6.3 High performance liquid chromatography (HPLC)
The application of immobilized enzyme reactors in HPLC was reported
by Shimada et al. [9] and by Shi and Du [143]. The latter authors cited
the reactors used for the determination of choline and acetylcholine in
their review.
Duan et al. reported the use of a rapid and simple method for the
determination of acetylcholine and choline in mouse brain by high
performance liquid chromatography, making use of an enzyme-loaded
post column and an electrochemical detector [144]. Perchloric acid
extracts of small brain tissue were injected onto the HPLC system with no
prior clean-up procedure. Detection limits for both compounds were
1 pmol, and this method was successfully applied to the measurement of
acetylcholine in discrete brain areas of the mouse.
Tao et al. described an HPLC method for the determination of
acetylcholine in a pharmaceutical preparation [145]. Utilizing reverse
phase ion pairing, acetylcholine was determined in lyophilized ophthal-
mic preparations. Analysis of degraded commercial samples showed the
utility of the method in quantitation, being stability indicating, and useful
in separating acetylcholine from choline.
Rhodes et al. correlated steroid sulfatase inhibition, DU-14
[P-O-(sulfamoyl)-N-tetra-decanoyltyramine] with changes in blood
levels of dihydroepiandrosterone sulfate and dihydroepiandrosterone,
acetylcholine levels in the hippocampus, and the blockade of scopolamine
amnesia in rats [146]. Rats were administered DU-14 by intraperitoneal
injection (30 mg/kg, for 14 days before memory testing) using a
passive avoidance paradigm. A separate group of animals, following
pretreatment with DU-14, were anesthetized and a microdialysis probe
surgically implanted into the hippocampus. Microdialysis samples were
collected and analyzed for acetylcholine via HPLC with electrochemical
detection. Hippocampal acetylcholine increased by almost a factor of
Other high performance liquid chromatography methods reported for the
determination of acetylcholine are summarized (together with their
conditions) in Table 3 [4, 117, 147–193].
6.4 Liquid chromatography (LC)
Niwa et al. determined acetylcholine and choline with platinum-black
ultramicroarray electrodes using liquid chromatography with a post-
column enzyme reactor [194]. An array of platinum-black microelec-
trodes was utilized for the indirect detection of acetylcholine and choline
following microbore liquid chromatography with post-column enzymatic
reaction. The liquid chromatography system consisted of a Unijet
acetylcholine analytical column (53 cm  1 mm i.d.) coupled with Unijet
ACh/Ch immobilized enzyme column and an electrochemical detection
cell equipped with the working electrode, a Ag/AgCl reference electrode,
and a stainless steel auxiliary electrode. The working electrode was
prepared by a plating method and consisted of Pt-black particles
(0.1–0.3 mm) deposited onto an Au film electrode. Each Pt-black particle
operated as an ultramicroelectrode. The detector was operated at
þ 0.5 V, and the injection volume was 5 mL. The mobile phase
(eluted at a rate of 120 mL/min) was 50 mM Na2HSO4 buffer at pH 8
containing 0.5 mM EDTA and 5 mL/L 1% Kathon CG reagent.
Detection limits of 5.7 and 6 fmol were achieved for acetylcholine and
choline respectively.
Mayer determined acetylcholine and choline by enzyme-mediated
liquid chromatography with electrochemical detection [195]. The two
compounds were separated by passing the eluted fractions through a
post-column reactor containing immobilized Acetylcholineesterase and
choline oxidase. In the presence of either compound, the dissolved
oxygen was converted into hydrogen peroxide, which was detected
amperometrically at a platinum electrode. This method was used to
determine choline in rat brain homogenates.
The acetylcholine levels in rat brain regions were determined by
Ikarashi et al. [196]. The method used was an application of FRIT
fast-atom-bombardment liquid chromatography-mass spectrometry.
This technique was used to determine acetylcholine levels in seven
regions of rat brain by monitoring the intact molecular cations of
High Performance Liquid Chromatography Systems
Column Mobile phase and Detection Sample and remarks Reference
[flow rate]
Discovery C18 column 20 mM ammonium Tandem Rat brain [4]
(15 cm  2.1 mm i.d.) acetate/20 mM hepta- ion-trap MS
fluoro butyric acid in
methanol/water (1:9;
pH 3.2) [0.4 mL/min].
MF-8904 cation 50 mM Na2HPO4 con- Vitreous C elec- Detection of basal acetyl- [117]
exchange (53 cm  taining Kathon CG trode operated choline in rat brain
1 mm i.d.) (10 mm) (5 mL/L) of pH 8 at þ 100 mV microdialysate
[140 mL/min]. versus
Ag/AgCl.
15 cm of Bio-Sil ODS-5S 0.01 M sodium acetate- Electrochemical Neuronal tissues. [147]
fitted with an ODS-5 0.02 M citric acid (pH 5) Reversed-phase HPLC.
guard column containing 4.5 mg/L of
sodium octyl sulfate
and was 1.2 mM in
tetramethyl ammonium
chloride.
Reversed-phase packing 20 mM Tris acetate (pH Electrochemical Tissue extracts, AChE [148]
post column reactor 7)–1 mM-tetramethyl- and ChO are absorbed
3 cm  2.1 mm of ammonium chloride- and immobilized on
Aquapore AX 300. 2 mM octanesulfonate the post column
containing 2% of
acetonitrile.
Hamilton PRP-1 poly- 20 mM Tris-acetate (pH Electrochemical Rat or mouse brain or [149]
styrene cartridge 7) 200 mM in octano- salivary gland tissues
(10 cm  4.6 mm) or sulfonate and contain-
column of Ultrasphere ing 2.5% of
ODS. acetonitrile [2 mL/min].
Chemcosorb C-18 (3 mm) — Amperometric Rat brain tissues; [150]
determines 10 pmol
25 cm  4 mm of ODS- 10 mM Sodium acetate Amperometric Use of reactor containing [151]
120 T. (pH 5)–1 mM tetra- ChO and AChE on
methylammonium LiChrosorb-NH2
chloride–10 ppm
[1 mL/min]
10 cm  4.6 mm of 10 mM Sodium acetate, Electrochemical Brain tissues, passed [152]
reversed phase material sodium octyl sulfate through a column con-
(30 mg/L and 2.7% of taining ChO and AChE
acetonitrile, adjusted to
pH 5 with citric acid
[0.8 mL/min].
Radial-Pak mBondapak Butanol–methanol– UV at 254 nm Soya bean leaves. [153]
C18 cartridge. acetic acid–water Detection limit 1 nmol
(4:2:1:43) containing
0.15 mM-1-phenethyl-
2-picolinium bromide
[3 mL/min].
PRP-1styrene-divinyl- Water–acetonitrile– UV at 254 nm Limit of detection 2 mg [154]
benzene (10 mm) propanol (90:7:3)
containing 0.45 M-
KH2PO4 and 0.1 mM-
hexadecyltrimethyl
ammonium bromide
Develosil Ph-5 and Potter et al., Electrochemical Rat brain limit of [155]
Develsil Ph (Anal. Abstr., 1984, 46, detection 5 pmol.
(15–30 mm) guard 9D167).
20 cm  4 mm of styrene Aqueous 2 mM Electrical con- Choline in plants. RP [156]
divinyl benzene hexansulfonic ductivity. ion-pair HPLC.
copolymer acid-acetonitrile.
25 cm  4.6 mm of (A) Acetonitrile:water: Liquid scintilla- Biological material, [157]
Microsorb-silica ethanol:acetic acid: tion counting. choline metabolite
0.83 M sodium acetate
(800:127:68:2:3) at pH
3.6. (B) (400:400:68:53:
7.5 cm  4 mm of 0.2 M phosphate buffer Electrochemical Brain tissue. [158]
Nucleosil 5 SA and a (pH 7.1) and 20 mM Eluate passed through a
guard (7.5 cm  2.1 tetramethyl ammonium column of ChO and
mm) cation exchange. [0.8 mL/min]. AChE.
15 cm  4.6 mm of Sodium acetate (1.36 g/L) Electrochemical Rat brain tissues. Eluate [159]
Nucleosil C18 at 37 . Na2EDTA (3.72 g/L) passed through the
sodium octyl sulfate AChE and ChO
(25 mg/L)–tetramethyl column.
(1.2 mM)–Water (pH 5)
7.5 cm  4.6 mm of Ikarashi et al, Electrochemical Brain tissue [160]
Chemosorb 70 DS-L (Anal. Abstr., 1985,
300 (7 mm) 322, 191).
15 cm  4 mm Nucleosil 100 mM sodium phos- Electrochemical Biological tissues [161]
5 SA phate of pH 7.6, 7 mM
in tetramethyl
ammonium perchlorate
[700 mL/min].
12.5 cm  4 mm of 10 mM phthalic acid, 1.2 Chemi- Eluate was passed [162]
Yanapak ODS 120 T. mM triethylamine and luminescence through a column of
76 mM sodium octane- AChE and ChO.
sulfonate adjusted to
pH 5 with KOH solu-
tion [0.4 mL/min].
15 cm  4.6 mm of 50 mM sodium Electrochemical Brain tissue. Recoveries [163]
PLRP-S (7 mm) and an phosphate (pH 8.3) with a were 108–110%.
immobilized enzyme containing 1 mM tetra- platinum
column (BAS Japan). methyl ammonium electrode at
chloride and 70 mM þ 500 mV
sodium octane-1-sulfo- versus
nate [1 mL/min]. AgAgCl.
10 cm  2 mm of Acetone–aqueous buffer Electrochemical Spinal cord of rat. Eluate [164]
Spherisorb S5 C6 solution (71:929) passed through reactor
containing 50 mM of AChE and ChO
NaH2PO4, 15 mM
chloride and 0.3 mM
sodium dodecyl sulfate
adjusted to pH 7.7
with NaOH
25 cm  4.6 mm of 0.1 M KH2PO4 contain- Electrochemical Brain tissues. Post [165]
Gynkotek Hypersil ing 500–700 mg/L of column enzymatic
ODS (5 mm) tetramethyl ammonium reaction.
chloride at pH 7.
7.5 cm  2.1 mm of 50 mM potassium phos- Chemi- Serum and urine. [166]
Nucleosil 5 SA. phate buffer (pH 7.4) luminescence Detection limit 0.7 and
containing 20 mM 0.5 pmol for ACh and
tetramethyl ammonium Ch.
nitrate [0.5 mL/min]
5 cm  4.6 mm of 10 mM Trizma acetate Electrochemical Neuronal tissues. [167]
RLRPS (5 mm) buffer (pH 7.8)–140 mM Detection limit 0.5 and
octanesulfonic acid– 1 pmol for Ch and
1 mM tetraethyl ammo- ACh.
nium chloride
10 cm  3 mm of 0.2 M Potassium phos- Electrochemical Spinal cord of rats. [168]
Chromspher 5 C18 phate (pH 8) [0.6 mL/ Eluate passed through a
loaded with sodium min]. reactor of AChE and
dodecyl sulfate and a ChO.
pre-column and a
guard column.
100 mm  3.0 mm 0.2 M KH2PO4 buffer Electrochemical Blood plasma and red [169]
reversed phase and 0.005 M KCl at cells in humans and in
chromsphere S C18 pH 8. mice.
with lauryl sulfate
Chemosorb ACh-II — Coulometric Rat brain. Detection [170]
reversed phase. limit was 10–20 pmol
for Ch and ACh.
15 cm  6 mm of Shodex 0.1 M sodium phosphate Amperometric Human cereprospinal [171]
RSpak DE613 medium buffer (pH 8.3) contain- fluid. Post-column
polarity methacrylate ing 1.2 mM tetramethyl enzymatic reaction with
gel. ammonium chloride AChE and ChO.
and 300 mg/L of
sodium decan-1-
sulfonate [1 mL/min].
Eicompak AC-GEL 0.07 M-Phosphate buffer Electrochemical Mouse brain tissues. An [172]
containing 60 ppm of immobilized enzyme
Na2 EDTA, 0.065% of column with ChO and
tetramethyl ammonium AChE was used.
chloride and 0.3 of
sodium octane-
sulfonate
10 cm  3.1 mm of 0.2 M phosphate–0.1 Electrochemical Human cerebrospinal [173]
Nucleosil 5 C18 with mM EDTA (pH 7.9) fluid. An immobilized
guard column containing 0.2 g/L of enzyme was used.
(1 cm  2.1 mm) sodium of octane-1-
sulfonate and 0.4 mM
bromide.
3 or 6 cm  2 mm of 3:47 Acetone-aqueous Electrochemical Both AChE and ChO [174]
Dynosphere PD-051-R buffer (pH 7.5) were immobilized in
(5 mm) post column reactor.
An ODS guard column 0.1 phosphate buffer (pH Electrochemical Cerebrospinal fluids of [175]
(5 mm  4 mm). An 8) containing 65 ppm of rats
AC Gel (10 mm) tetramethyl ammonium
separation column chloride and 300 ppm
(15 cm  6 mm), of sodium octane-
CA trap and sulfonate [1 mL/min].
AC-Enzympak column
10 cm  7 mm of 0.05 M KH2PO4 contain- Electrochemical Mice brain tissues. Post [176]
Spherisorb ODS 2 ing tetramethyl column reactor contain-
(3 mm) ammonium chloride ing ChO and AChE was
and sodium octanesul- used.
fonate (pH 7) [1.2 mL/
min]
15 cm  5 mm of 0.1 M phosphate buffer Electrochemical Human plasma. Eluent [177]
Eicom-pak AC-GEL (pH 8.3) containing 1.2 passed through an
mM tetramethyl ammo- immobilized enzyme
nium chloride and 300 column.
ppm of sodium decan-
sulfonate [1 mL/min]
3 to 25 cm  4 to 4.6 mm 100 mM phosphate buf- — Best results were [178]
of cation exchange fer pH 7.6 containing 7 obtained with Hamilton
columns. Hamilton mM tetramethyl ammo- column
PRP-X 200 column. nium perchlorate [0.5 to
1.5 mL/min]
10 cm  3.2 mm of Tris buffer solution Electrochemical Serum. Detection limit [179]
C18 (3 mm) (pH 7.5) containing was 3 pmol for choline.
acetic acid, tetramethyl
ammonium chloride,
sodium octylsulfate,
NaN3, EDTA, and
6 cm  4 mm of 0.05 M phosphate– Electrochemical Rat brain [180]
polymeric 1.0 mM Na2 EDTA-
styrene-based packing 0.4 mM sodium
1-octansulfonate of pH
8.4 [0.8 mL/min].
15 cm  3 mm of 0.1 M phosphate buffer Electrochemical Erythrocytes extract [181]
Li Chrosorb RP18 (pH 7.4) containing 0.1 from heart or brain
(25 mm) 25  4.1 mm mM EDTA and 4 mM tissue. Reversed phase
of Hamilton PRPX 200 tetramethyl ammonium column provided better
(10 mm). 25 cm  4 mm perchlorate with the use pH stability and longer
of Supelcosil LC8 of a reactor cartridge working life.
(5 mm) and a (3 cm  2.1) of
LiChroCART cartridge immobilized Cho and
12.5 cm  2 mm of AChE [0.5 mL/min].
Superspher 60
RP-Select B).
10 cm  1 mm of 0.2 M phosphate buffer Electrochemical Brain tissues microdia- [182]
Aminex A-9 solution (pH 8) con- lysis or cerebrospinal
taining 5 mM NaCl fluids.
and 0.1% Kathon CG
[60 mL/min].
15 cm  1 mm of 0.1 M sodium phosphate Electrochemical Plasma. Detects anti [183]
Spheron Micro-DEAE buffer (pH 7.4) contain- AChE compound with
300. ing 0.1 mM acetylcho- post column reaction
line and 0.1 mM
EDTA [60 mL/min].
Polymeric reversed phase 100 mM sodium phos- Electrochemical Microdialysis in rat stria- [184]
column (15 cm  3 mm phate of pH 8 contain- using an ESA tum. Analysis in the
i.d.) of ACH-3 (5 mm) ing 0.5 mM tetramethyl model 5040 presence and absence
with an ACH-3-G ammonium chloride, solid state cell of esterase inhibitors.
guard cartridge. 0.005% Reagent MB with a Kel-F
(ESA) and 2 mM octa- or PEEK
nesulfonic acid [0.35 Pt-target work-
mL/min]. ing electrode at
300 mV versus
a Pd reference
Analytical column and Sodium phosphate buffer Electrochemical Human plasma and peri- [185]
post column immobi- pH 8.5 containing at þ 0.5 V toneal dialysis effluent.
lized enzyme reactor Kathon CG (Rohm and versus
supplied as part of a Haas, PA, USA), Ag/AgCl.
ACh/Ch assay kit. [1 mL/min].
(15 cm  1.7 mm i.d.) of 0.2% trifluoroacetic acid Flow fast atom Brain tissues of rodents. [186]
SFPAC-ODSO5-S15 in water containing 1% bombardment Detection limit was 5
(5 mm). glycerol, [40 mL/min for mass and 2 for Ch and ACh.
5 min. then 11 mL/min]. spectrometry
10 mL loop injector on 28 mM Na2HPO4–0.5% Electrochemical Brain dialysis of rats. An [187]
a 10 mm bioanalytical Kathon (pH 8.5 [1 mL/ at 0.5 V versus immobilized-enzyme
system ACh column min]. AgCl. reactor was used.
(53 cm  1 mm i.d.)
A BASJ ion-exchange 0.05 Sodium phosphate Electrochemical Rat frontal cortex [188]
microbore column buffer of pH 8.5 con- at horseradish dialysate samples
(45 cm  1 mm i.d.) taining 0.1 mM EDTA peroxidase
[60 mL/min]. osmium redox
polymer-
vitreous C
electrodes at 0
mV versus
Allteck C-18 (25 cm  Aqueous 10 mM ammo- Electrospray Rat cell culture system. [189]
1 mm i.d.) (5 mm). nium acetate–2 mM ionisation MS Detection limit 10 pM
pyridine–230 mM detection at of acetylcholine.
sodium octanesulfonate m/z 146.
of pH 5 [10 mL/min].
(1) A Stipstik column 29 mM NaH2PO4–22 Electrochemical ACh in microdialysis [190]
(53 cm  1 mm i.d.) mM sodium acetate–5 detection samples.
with ChO/AChE reac- mM Kathon of pH 8.5.
tor and guard columns.
(2) A polymeric reversed 50 mM Na2HPO4–0.3 Amperometric ACh in microdialysis [190]
phase column mM sodium octane-1- samples.
(12 cm  2 mm i.d.) sulfonate–1 mM Na2-
EDTA of pH 8.5.
A polymer gel column 50 mM Na2HPO4 of Electrochemical- ACh and Ch in [191]
(10 cm  4 mm i.d.) pH 8 containing 0.5% ly using Pt micro-dialysates
Kathon CG [0.8 mL/ electrode at from rat brain.
min]. þ 0.5 V versus
Bioanalytical systems 50 mM phosphate buffer Amperometric Acetylcholine and related [192]
acetylcholine (pH 8.4) containing metabolite, detection
separation column 1 mM EDTA and limit are 25 pM.
(6 cm  4 mm i.d.) with 0.4 mM sodium-1-octa-
post column nesulfonate
(5 cm  4 mm i.d.) [0.8 mL/min].
containing immobilized
AChE and ChO.
Crosslinked polystyrene 0.05 M phosphate buffer Amperometric Determines ACh and Ch [193]
column (3 mm) (pH 8.4) containing at þ 0.5 V in biological samples.
(6 cm  4 mm i.d.) with 1 mM EDTA and 0.4 (versus
a precolumn (1 cm  mM sodium 1-octane- Ag/AgCl)
4 mm i.d.) by a slurry sulfonate [0.7 mL/min].
packing with acetone.
acetylcholine and an internal deuterated standard. Results were
compared with other methods, such as pyrolysis GC–MS and LC with
electrochemical detection.
Ikarashi et al. reported the development of a liquid chromatography
multiple electrochemical detector (LCMC) and its application in
neuroscience [197]. The system developed consisted of four parallel
liquid chromatographic systems equipped with multiple electrochemical
detectors, and the fourth was used for the assay of acetylcholine with a
detection limit of 0.1–0.4 pmol.
6.5 Cation-exchange liquid chromatography
Stein reported the separation of choline and acetylcholine by a cation-
exchange chromatographic method [198]. Four columns were compared
by Salamoun et al. for the separation of choline and acetylcholine by
cation exchange liquid chromatography [178]. The mobile phase used was
0.1 M phosphate buffer (pH 7.4), containing 0.1 mM EDTA and 4 mM
tetramethylammonium perchlorate, and was eluted at a rate of 0.5 mL/
min. The system made use of a reactor cartridge (3 cm  2.1 mm)
containing immobilized choline oxidase and acetylcholineesterase, and
electrochemical detection at 0.45 V. The columns were (15 cm  3 mm
i.d.) LiChrosorb RP-18 (5 mm), (25 cm  4.1 mm i.d.) Hamilton PRP-X
200 (10 mm), (25 cm  4 mm i.d.) Supelcosil LC-8 (5 mm), and a
LiChroCART cartridge (12.5 cm  2 mm i.d.) containing Supersphere
60 RP-Select B. Appropriate guard columns were used as well. The use of
reversed phase columns provided better pH stability and a longer
working life than the normal silica-based cation exchangers.
spectrometry (HPLC/MS)
Ishimaru et al. reported the use of high performance liquid chromato-
graphy combined with continuous flow fast atom bombardment mass
spectrometry for the simultaneous determination of choline and
acetylcholine in rodent brain regions [186]. Brain tissues were homo-
genized in 2 mL of acetonitrile containing 50 mg magnesium sulfate and
10 nmol each of the d4 analogs of choline and acetylcholine (as internal
standards). The mixture was centrifuged, and the solution extracted with
two 2 mL aliquots of hexane. The acetonitrile was evaporated under
nitrogen, and the residue was dissolved in 0.2 mL of the mobile phase.
A 20 mL portion of the resulting solution was analyzed by HPLC (see
Table 3 for more details).
Acevedo et al. quantified acetylcholine in a cell culture system by semi-
micro high performance liquid chromatography and electrospray
ionization mass spectrometry [189]. Rat cells were washed with
Dullecco's PBS solution (containing 10 mM Physostigmine), and incu-
bated in 2.5 mL 10 mM Physostigmine/50 mM-KCl in Dullecco's PBS for
10 min. Portions (2.5 mL) were mixed with 500 pM N,N,N-trimethyl-
d9-acetylcholine (internal standard), and were applied to a C-18 SPE
column. After washing with water, acetylcholine and choline were eluted
with aqueous 50% acetic acid. The first three fractions (500 mL) were
combined, frozen on dry ice, and lyophilized. After reconstitution in
50 mL of aqueous 10 mM ammonium acetate–2 mM pyridine–230 mM
sodium octanesulfonate (pH 5), 5 mL portions were analyzed on a 5 mm
Alltech C-18 column (25 cm  1 mm i.d.). Electrospray ionization with
MS detection was used at m/z 104 and 146 for choline and acetylcholine,
respectively.
6.7 Gas chromatography–mass spectrometry (GC/MS)
Jenden reported the use of gas chromatography–mass spectrometry
method to measure tissue levels and the turnover of acetylcholine [199].
The systems that catalyze and hydrolyze acetylcholine were inactivated
rapidly by freezing the brain tissue in nitrogen, pulverizing, and extracting
in 15% 1 N formic acid in acetone. After addition of the internal standard,
the homogenate was centrifuged to obtain a supernatant containing the
bound forms of acetylcholine. Acetylcholine was esterified with propionyl
chloride, dried in a stream of nitrogen, demethylated, heated to 80 C for
45 min, and then partitioned between aqueous acid and pentane. The
tertiary amines were removed by extraction with CH3Cl, and the
acetylcholine injected into a gas chromatograph. Quantitative determina-
tion was achieved by mass spectrometry using 2H-labeled compounds as
tracers and internal standards.
Polak and Molenaar described a method for the determination of
acetylcholine from brain tissue by pyrolysis-gas chromatography-mass
spectrometry [200]. The deuterium-labeled acetyl-choline is pyrolytically
demethylated with sodium benzenethiolate, followed by quantitative GC–
MS analysis. In this method, care must be taken so that the samples do not
contain appreciable amounts of choline since exchange of deuterium-
labeled groups between acetylcholine and choline during pyrolysis may
yield erroneous results. The same authors have also reported a method for
the determination of acetylcholine by slow pyrolysis combined with mass
fragment analysis on a packed capillary column [201].
Mikes reported the use of a new injection micropyrolysis apparatus for
acetylcholine and choline determination by gas chromatography–mass
spectrometry [202].
Singh and Drewes described an improved method for the analysis of
acetylcholine and choline in canine brain and blood samples by capillary
gas chromatography–mass spectrometry [203]. Frozen samples were
mixed with butyryl choline (internal standard) and extracted. The GC–
MS system was equipped with an electron-impact ionizer and a 15 m
capillary column coated with a bonded methyl-silicone phase (HP-SE 54).
The column was operated at 60 C for 3 min, and then temperature
programmed (at 25 C/min) to 200 C. The ion at m/z 58 was selected for
monitoring, and the separation of the three compounds was achieved in
less than 5 min.
Liberato et al. reported the separation and quantification of acetycholine
and choline by thermospray liquid chromatography–mass spectrometry
[204]. Choline and acetylcholine were extracted from mouse brain
homogenates, in the presence of 2H9-choline and 2H4-acetylcholine, and
the extract was analyzed directly by HPLC–MS on a column
(15 cm  4.6 mm) of Ultrasphere-I.P. ODS. The mobile phase was
0.1 M ammonium acetate–2 mM pyridine containing octanesulfonic
acid (50 mg/L, pH 5), and was eluted at a flow rate of 1.25 mL/min.
The column was coupled to a quadrupole mass spectrometer by the
thermospray interface, and selected ions were monitored at m/z 104, 113,
146 and 150 for choline, [2H9] choline, acetylcholine and [2H4]
acetylcholine, respectively.
Watanabe et al. used a pyrolysis gas chromatography–mass spectro-
metric method for the determination of acetylcholine in human blood
[205]. Acetylcholine was extracted by an HClO4 treatment of the whole
blood in the presence of eserine to inhibit cholineesterase activity, and
was then determined by pyrolysis GC–MS with butyrylcholine as the
internal standard. Pyrolysis was carried out for 7.5 s at 300 C, and the
product was analyzed on a glass column (2 m  2 mm) of 5% of DDTS
and 5% of OV-101 on Gas Chrom Q (100–200 mesh) at 115 C. Helium
was used as the carrier gas (25 mL/min), and detection was effected using
MS at 28 eV with an ion source temperature of 205 C. The ion at m/z ¼
58 was monitored for both acetylcholine and the internal standard.
Pomfret et al. measured both choline and choline metabolite concentra-
tions using HPLC and GC–MS [206]. Tissues were extracted with
100 A.A. AL-BADR AND H.A. EL-OBEID
chloroform/methanol/water, and the separated phases were evaporated.
The organic phase residue was dissolved in chloroform/methanol (1:1),
and a portion was applied on a silica gel for TLC analysis with
chloroform/methanol–water (65:30:4) as the mobile phase. Zones
corresponding to phosphatidylcholine and lysophosphatidylcholine
were removed and hydrolyzed in methanolic HCl to liberate choline.
The aqueous phase residue was dissolved in methanol/water (10:1), and
the hydrophilic metabolites were separated by HPLC on a column
(8.3 cm  4.6 mm) of Pecosphere-3 CSi (5 mm) with a Supelguard LCSi
guard column (2 cm  4.6 mm) and acetonitrile–ethanol–acetic acid–1 M
ammonium acetate–water–0.1 M sodium phosphate (800:68:2:3:127:10
changing to 400:68:44:88:400:10) as the mobile phase (flow rate of
1.5 mL/min). Fractions corresponding to betaine, acetylcholine, phos-
phocholine, glycerophosphocholine, and choline were separated and
hydrolyzed as before. Choline was determined by GC–MS of each
fraction, and metabolites could be detected down to the 200 pmol level in
10 mg samples.
Terry et al. reported the synthesis of novel selenium-containing
choline and acetylcholine analogues and their quantitation in biological
tissues, using a pyrolysis-gas chromatography–mass spectrometry assay
method [207]. Biological tissues were homogenized in formic acid-
acetonitrile, the homogenate was centrifuged, and the supernatant
solution was extracted with 2 mM dipicrylamine in CH2Cl2 and TAPS
buffer (pH 9.2). After centrifugation, the CH2Cl2 layer was evaporated
under nitrogen, silver p-toluenesufonate in acetonitrile and propionyl
chloride were added, and the solution was set aside for 10 min at room
temperature. The mixture was evaporated to dryness under nitrogen, the
residue was dissolved in sodium acetate buffer (pH 4), and the solution
was washed with butan-2-one before evaporation to dryness at 50 C
under nitrogen. The residue was dissolved in aqueous 90% acetonitrile,
and this was pyrolyzed for 10 s at 325 C in a quartz tube. The pyrolysis
products were trapped in a cold trap for three min before analysis by GC
on a column (30 m  0.25 mm) coated with Stabilwax (0.5 mm) with
helium as the carrier gas and temperature programming from 30 C to
200 C (held for 2.5 min) or 30 C/min and 68 eV EIMS with selected ion
monitoring at m/z 122 and 125. Calibration graphs were linear from
20 pmol to 20 nmol of selenium choline propionyl ester and acetyl-
selenium choline.
Patterson and Kosh described a method for the simultaneous
quantitation of arecoline, acetylcholine, and choline in mouse brain
ACETYLCHOLINE CHLORIDE: ANALYTICAL PROFILE 101
tissues using gas chromatography–electron-impact mass spectrometry
[208]. Mouse brain tissues were homogenized in 1 M formic acid –
acetonitrile (17:3), washed with ethyl ether, and centrifuged for two
min. Arecoline, acetylcholine, and choline in the supernatant were
extracted as ion pairs with 5 mM sodium tetraphenylboron in heptan-
3-one. After centrifugation, the organic layer was evaporated to dryness
under nitrogen at 60 C, and the dried residue was mixed with 5 mM
silver p-toluene sulfonate in acetonitrile. Propionyl chloride was added,
and after 5 min the mixture was evaporated to dryness. The residue was
mixed with sodium thiophenoxide solution and incubated at 80 C for
30 min. The cooled solution was mixed with 0.5 mL citric acid and
washed with ethyl ether and pentane. The solvents were evaporated,
chloroform was added to the solution in an ice bath, which was then
followed by 2 M ammonium citrate–7.5 M ammonium hydroxide
buffer. Following centrifugation, the chloroform layer was analyzed
by gas chromatography–mass spectrometry. Hexadeuterated arecoline
and non-deuterated acetylcholine and choline were used as internal
standards. The analysis was performed on a column (30 m  0.25 mm)
of Stabilwax with ramped temperature programming for 50 C (held for
0.75 min) to 200 C (held for 3.5 min) at 35 C/min and then to 220 C at
25 C/min. Electron impact MS was carried at 68 eV with SIMS
monitoring of m/z ¼ 140 and 146 for arecoline and m/z ¼ 58 and 64
for acetylcholine and choline. The detection limit was 25 pmol for all
the three compounds, and the calibration curves were linear from 0.025
to 50 nmol.
6.8 Capillary electrophoresis
Kappes et al. evaluated the potentiometric detection of acetylcholine and
other neurotransmitters through capillary electrophoresis [209].
Experiments were performed on an in-house capillary electrophoresis
instrument that made use of detection at a platinum wire, dip-coated in
3.4% potassium tetrakis (4-chlorophenyl) borate/64.4% o-nitrohenyl
octyl ether/32.2% PVC in THF. The results were compared to those
obtained using capillary electrophoresis with amperometric detection at a
graphite electrode. Samples prepared in the capillary electrophoresis
buffer were electrokinetically injected (7 s at 5 kV) into an untreated fused
silica capillary (88 cm  25 mm i.d.) and separated with 20 mM tartaric
acid adjusted to pH 3 with MgO as the running buffer. The system used
an applied potential of 30 kV, and detection versus the capillary
electrophoresis ground electrode.
6.9 Radioassay methods
The enzymatic radioassay method for the analysis of acetylcholine
and choline in brain tissue has been reported by Reid et al.
[210]. The method describes the determination of nanogram amounts
of acetylcholine and choline in as little as 10 mg of brain tissue,
involves isolation of acetylcholine by high-voltage paper electrophoresis,
alkaline hydrolysis of acetylcholine to choline, and conversion of
this into [32P]-phosphoryl choline in the presence of choline kinase
and [g32P] ATP. The labeled derivative is isolated by column chroma-
tography on Bio-Rad AG1-X8 resin, using Tris buffer solution as
the eluent. Cerenkov radiation from 32P is counted (at 33% efficiency)
in a liquid scintillation spectrometer. The amount of phosphorylcholine
is proportional to the amount of choline over the range of 0.08–
8.25 nmol.
Bluth et al. described a simplified radio-enzymatic assay method for the
determination of acetylcholine and choline in discrete structures of rat
brain [211].
Gilberstadt and Russell reported the determination of picomole
quantities of acetylcholine and choline in physiologic salt solutions
[140]. Carbon-14 labeled choline and acetylcholine standards in Krebs–
Ringer solution were subjected to chromatography. The unlabeled choline
and acetylcholine were assayed radioenzymatically by using [g32P] ATP,
choline kinase and acetylcholineesterase. The calibration graphs were
rectilinear in the range 10–3000 pmol acetylcholine and 30–3000 pmol
of choline.
Dick et al. used a radio gas chromatography method for monitoring the in-
vivo labeling of postmortem [3H] choline production [212]. After
homogenization in 1 M formic acid–tetrahydrofuran (3:17) and deriva-
tization with butyryl chloride, brain tissues from rats injected with [3H]
choline were assayed for choline, acetylcholine, and their 3H-analogues.
The analysis was performed using GC on a column of Chromosorb 750,
coated with 10% OV-17 and 10% Triton X-100. The method used
temperature programming for 130–150 C, helium as the carrier gas (flow
rate of 40 mL/min), and N-P detection. For subsequent scintillation
counting, 20% of the column effluent was trapped on p-terphenyl, coated
as described above. Propionylcholine and [3H]-hexanoylcholine were used
as internal standards. Detection limits for [3H]-choline and [3H]-
acetylcholine were 150 and 730 fmol, respectively.
A multiple enzyme and multi-substrate cycling system was described by
O'Neill and Hurschak for the assay of choline acetyltransferase [213].
Assay tubes containing sodium [14C] acetate (13 mCi/mmol) and (final
concentration) 0.2 M Tris–HCl, 5 mM choline chloride, 0.1 mM EDTA,
0.05 mM dithiothreitol, 7 mM potassium acetate, 20 mM MgCl2,
0.2 mM eserine sulfate, 0.36 mM ADP, 300 mM KCl, 20 mM-
phosphoenol pyruvate, 0.1 mM-coenzyme A, 0.20% of bovine serum
albumin, pyruvate kinase, phosphate acetyltransferase, and acetate
kinase were centrifuged (1000 G at 3–4 C for 10 min), and kept at 37 C
for 20 min. Brain homogenate was added, and the mixtures were
centrifuged and incubated at 37 C for 1–60 min. The reaction was
stopped by placing the tubes in ice-water. A 10 mM-acetylcholine
solution was added, and the mixture was applied to a column of Ag1-
X8. Elution was with water, and the eluates were collected in phials
containing Chaikoff scintillation solution and assayed by liquid
scintillation counting.
Cooper reported a simple method to determine the released acetylcho-
line in the presence of choline [214]. The use of this method obviates
the necessity both for preventing the re-uptake of choline by
synaptosomes during determination of released acetylcholine, and for
subsequently separating radiolabeled acetylcholine from radiolabeled
choline. To the sample or standard solution were added [3H]-choline
(5 nmol), choline oxidase (10–150 min), and Krebs-Ringer bicarbonate
buffer solution (pH 7.4) to yield a final volume of 0.2 mL. The mixture
was incubated at 37 C for 5 miU, and then 1 mL of 1% tetraphe-
nylborate solution in butyronitrile was added. The mixture was vortex-
mixed for 30 s, and then centrifuged. A 0.5 mL portion of organic phase
was added to 5 mL of Opti-Fluor scintillator for the measurement of
radioactivity.
Eckernas and Aquilonius described a simple radioenzymatic procedure
for the determination of choline and acetylcholine in brain regions of rats
sacrificed by microwave irradiation [215]. The methods are based on
acetylation (in phosphate buffer) of free choline with [14C]-acetylcholine
using purified choline acetyltransferase. After ion-pair extraction of
[14C]-acetylcholine with tertiary phenyl borate contained in toluene-based
scintillation cocktail, the radioactivity was measured.
Spector et al. developed a specific radioimmunoassay method for the
determination of acetylcholine [216]. The described method is suitable
for determining 2–137 pmol of acetylcholine; the radioligand used is
H-acetylcholine, and the hapten is 2-(trimethyl-amino)ethyl-6-amino-
hexanoate hydrobromide toluene-p-sulfonate, which is conjugated to
bovine serum albumin to yield the antigen. The antiserum is raised in
rabbits and is used at a dilution of 1:1000 in phosphate-buffered saline
containing bovine g-globulin and eserine (as a cholinesterase inhibitor).
After incubation of the mixture for 1 or 16 h at 4 C, the bound antigen is
precipitated by adding ammonium sulfate solution, centrifuged off, and
dissolved in water for scintillation counting. Cross-reactivity of the
antiserum with choline is only 0.1%.
Saelens et al. determined acetylcholine and choline by an enzymatic assay
method [217]. This assay method has been made more practical, primarily
by preparing choline acetyltransferase in large quantities in advance.
The preparation of the enzyme from mouse brain is described. The
acetylcholine and choline were separated from brain-tissue extracts by
electrophoresis at 500 V for one hour in a formic acid-acetic acid-water
buffer solution, with tetraethylammonium bromide being used as a
marker. The analyte bands were located through staining with iodine
vapor. The acetylcholine was eluted and hydrolyzed, and the resulting
choline was reacetylated with [acetyl-14C]-coenzyme A in the presence of
choline acetyltransferase. The [acetyl-14C]-acetylcholine was then counted
using a liquid scintillation system. The choline was also separately eluted
from the electropherogram and determined by the same technique.
Higher values for acetylcholine and choline were obtained after
sacrificing mice with liquid nitrogen than after other methods of sacrifice.
The brain tissue was extracted with formic acid-acetone.
Ladinsky and Consolo used an enzymatic radioassay method for the
determination of acetylcholine and choline [218]. The method was based
on the electrophoretic separation of acetylcholine and choline, hydrolysis
of acetylcholine to form choline, and acetylation of the choline with
labeled AcCoA and choline acetyltransferase. The labeled acetylcholine
formed was isolated and quantitated. The method was sensitive and
specific, and permitted the routine handling of a large number of samples
in a single experiment. The standard curves were linear up to at least
42.5 ng (0.4 nmol) choline and 45 ng (0.3 nmol) acetylcholine. The lower
limit of sensitivity was 2 ng, and the recovery of acetylcholine was 95%
when carried through the entire procedure.
Saelens et al. measured choline, acetylcholine, and their metabolites by
combined enzymatic and radiometric techniques [219]. Choline and
acetylcholine were isolated from tissue extracts by a low voltage
electrophoresis and assayed by transfer of the Ac-14C group of acetyl-14C-
coenzyme A to choline by choline-acetyltransferase. The lower limit of
sensitivity for this method was 1011 mol of choline or acetylcholine.
When choline-methyl-3H was infused intravenously into rats, the specific
activity of choline and acetylcholine increased curvilinearly in the cortex,
midbrain, and brainstem. The metabolism of choline involved in
acetylcholine synthesis and the metabolism of acetylcholine itself were
extremely rapid.
7. OTHER METHODS OF ANALYSIS
7.1 Ion-selective method
Jaramillo et al. published an article regarding some ion-selective micro-
electrodes used in the analysis of acetylcholine and choline [220].
7.2 Microwave methods
Szepesy et al. used a microwave apparatus in the determination of
acetylcholine and choline in the central nervous system samples [221].
Schmidt et al. reported the use of a microwave radiation method for the
determination of acetylcholine in the rat brain [222].
7.3 Microdialysis
Kehr et al. described microsurgical methods for the simultaneous
implantation of a microdialysis probe and intraventricular injection
cannula via their respective guide cannulas into the mouse brain [223].
Basal and stimulated release of acetylcholine was determined in the
ventral hippocampus of freely moving mice. No significant differences in
basal potassium-stimulated or scopolamine-induced extracellular ace-
tylcholine levels were observed in 4-month-old wide-type (WT) mice, or
the mice with a loss of function mutation of the galanin gene (KO). In the
aged, 10-month-old animals, the basal extracellular acetylcholine levels
were significantly reduced in both WT and KO groups.
8. ACKNOWLEDGMENT
The authors wish to thank Mr. Tanvir A. Butt, Department of Phar-
maceutical Chemistry, College of Pharmacy, King Saud University,
Riyadh, Saudi Arabia, for typing the manuscript of this profile.
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Benazepril Hydrochloride:
Comprehensive Profile
F. Belal, H.H. Abdine and Abdullah A. Al-Badr
PROFILES OF DRUG SUBSTANCES, 117 Copyright ß 2004 Elsevier Inc.
118 F. BELAL, H.H. ABDINE AND A.A. AL-BADR
1. Description . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119
1.1 Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . 119
1.1.1 Systematic chemical name . . . . . . . . . . . . 119
1.1.2 Nonproprietary names . . . . . . . . . . . . . . . 120
1.1.3 Proprietary names . . . . . . . . . . . . . . . . . . 120
1.2 Formulae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120
1.2.1 Empirical formula, molecular
weight, CAS number . . . . . . . . . . . . . . . . 120
1.2.2 Structural formula. . . . . . . . . . . . . . . . . . 120
1.3 Elemental analysis. . . . . . . . . . . . . . . . . . . . . . . . 120
1.4 Appearance . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121
1.5 Uses and application . . . . . . . . . . . . . . . . . . . . . . 121
2. Method of preparation . . . . . . . . . . . . . . . . . . . . . . . . . . 121
3. Physical characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . 123
3.1 Solubility characteristics . . . . . . . . . . . . . . . . . . . 123
3.2 Optical activity . . . . . . . . . . . . . . . . . . . . . . . . . . 123
3.3 X-Ray powder diffraction pattern. . . . . . . . . . . . . 123
3.4 Thermal methods of analysis . . . . . . . . . . . . . . . . 123
3.4.1 Melting behavior. . . . . . . . . . . . . . . . . . . 123
3.4.2 Differential scanning calorimetry . . . . . . . 123
3.5 Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . 124
3.5.1 UV/VIS spectroscopy . . . . . . . . . . . . . . . 124
3.5.2 Vibrational spectroscopy . . . . . . . . . . . . . 124
3.5.3 Nuclear magnetic resonance spectrometry . 124
3.6 Mass spectrometry . . . . . . . . . . . . . . . . . . . . . . . 128
4. Methods of Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129
4.1 Compendial methods. . . . . . . . . . . . . . . . . . . . . . 129
4.2 Statistically compared methods. . . . . . . . . . . . . . . 130
4.3 Potentiometric methods . . . . . . . . . . . . . . . . . . . . 131
4.4 Spectrophotometric methods . . . . . . . . . . . . . . . . 132
4.5 Colorimetric method . . . . . . . . . . . . . . . . . . . . . . 143
4.6 Voltammetric methods. . . . . . . . . . . . . . . . . . . . . 147
4.7 Chromatographic methods of analysis. . . . . . . . . . 147
4.7.1 Thin-Layer chromatographic methods. . . . 147
4.7.2 Gas chromatographic–mass
spectrometric methods (GC–MS) . . . . . . . 148
4.7.3 High performance liquid
chromatography . . . . . . . . . . . . . . . . . . . 150
4.7.4 Capillary electrophoresis methods . . . . . . . 156
BENAZEPRIL HYDROCHLORIDE: COMPREHENSIVE PROFILE 119
5. Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158
6. Absorption and Pharmacokinetics . . . . . . . . . . . . . . . . . . . 158
6.1 Absorption, metabolism and fate . . . . . . . . . . . . . 158
6.2 Excretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158
7. Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159
1.1.1 Systematic chemical names [1–3]
(3S)-3-[[(1S)-1-Carbethoxy-3-phenylpropyl]amino]-2,3,4,5-tetrahydro-2-
oxo-1H-1-benzazepine-1-acetic acid, hydrochloride.
[S-(R*,R*)]-3-[[1-(Ethoxycarbonyl)-3-phenylpropyl]amino]-2,3,4,5-tetra-
hydro-2-oxo-1H-1-benzazepine-1-acetic acid, hydrochloride.
{(3S)-3-[(1S)-1-Ethoxycarbonyl-3-phenylpropylamino]-2,3,4,5-tetrahy-
dro-2-oxo-1H-1-benzazepin-1-yl}acetic acid, hydrochloride.
(3S)-1-(Carboxymethyl)-[[(1S)-1-(ethoxycarbonyl)-3-phenylpropyl]-
amino]-2,3,4,5-tetrahydro-1H-[1]benzazepin-2-one, hydrochloride.
1-Carboxymethyl-3-[1-ethoxycarbonyl-3-phenyl-(1S)-propylamino]-
2,3,4,5-tetrahydro-1H-1(3S)-benzazepin-2-one, hydrochloride.
(3S)-3-[[(1S)-1-Carboxy-3-phenylpropyl]amino]-2,3,4,5-tetrahydro-2-
oxo-1H-1-benzazepin-1-acetic acid-3-ethyl ester, hydrochloride.
3-{[1-Ethoxycarbonyl-3-phenyl-(1S)-propyl]amino}-2,3,4,5-tetrahydro-2-
oxo-1H-(3S)-benzazepine-1-acetic acid, hydrochloride.
3-[1-(Ethoxycarbonyl)-3-phenyl-(1S)-propylamino]-2,3,4,5-tetrahydro-2-
oxo-1H-1-(3S)-benzazepine-1-acetic acid, hydrochloride.
1H-1-Benzazepine-1-acetic acid, [S-(R*,R*)]-3-[[1-(ethoxycarbonyl)-3-
phenyl-propyl]amino]-2,3,4,5-tetrahydro-2-oxo-, monohydrochloride.
1H-1-Benzazepine-1-acetic acid, 3-[[1-(ethoxycarbonyl)-3-phenylpropyl]-
amino]-2,3,4,5-tetrahydro-2-oxo-, [S-(R*,R*)].
1.1.2 Nonproprietary names [1–4]
Benazepril, Benazepril hydrochloride, CGS-14824 A.
Benazeprilat, CGS-14831.
1.1.3 Proprietary names [1–4]
Briem, Cibacen, Cibacene, Fortekor, Labopal, Lotensin, Tensanil and
Zinadril, Cibace.
1.2.1 Empirical formula, molecular weight, CAS number [1]
Benazepril C24H28N2O5 424.00 [86541755]
Benazepril HCl C24H28N2O5  HCl 460.50 [86541744]
Benazeprilat C22H24N2O5 396.00 [86541788]
COOC2H5 COOH
N H H N H H
CH2COOH CH2COOH
Benazepril Benazeprilat
1.3 Elemental analysis [1]
The calculated elemental compositions are as follows:
Benazepril Benazepril HCl Benazeprilat
Carbon 67.91% 62.54% 66.67%
Hydrogen 06.65% 06.34% 06.06%
Chlorine — 07.68% —
Nitrogen 06.60% 06.07% 07.07%
Oxygen 18.84% 17.37% 20.20%
Benazepril hydrochloride is obtained as a white to off-white crystalline
powder [3].
1.5 Uses and application
Benazepril hydrochloride, an angiotensin-converting enzyme (ACE)
inhibitor, is used in the treatment of hypertension and heart failure.
The family of ACE inhibitors inhibits the angiotensin-converting enzyme,
which is involved in the conversion of angiotensin I to angiotensin II.
Angiotensin II stimulates the synthesis and secretion of aldosterone and
raises blood pressure via a potent direct vasoconstrictor effect. ACE is
identical to bradykininase or kininase II, and ACE inhibitors may reduce
the degradation of bradykinin [2].
Benazepril hydrochloride, being a prodrug of the diacid benazeprilat, is
rapidly absorbed and converted to the active ACE inhibitor, benazepri-
lat. The drug has a half-life of 10–11 h. It is approved for treating
essential hypertension and may be effective for treating congestive heart
failure [2].
Benazepril owes its activity to benazeprilat, to which it is converted after
oral administration. The drug is given by mouth as the hydrochloride
salt, but doses are usually expressed in terms of the free base. In the
treatment of hypertension, the usual initial dose is 10 mg once daily. An
initial dose of 5 mg once daily is suggested for patients with renal
impairment, or who are receiving a diuretic. The usual maintenance dose
is 20–40 mg daily, which may be given in two divided doses if control is
inadequate with a single dose. In the treatment of heart failure, the usual
initial dose is 2.5 mg once daily, adjusted according to response to a
maximum dose of 20 mg daily [2].
2. METHOD OF PREPARATION
Watthey and Watthey et al. [5–7] reported the synthesis of benazepril
using the following procedure (which is illustrated in Scheme 1).
Chlorination of benzazepin-2-one (1) with phosphorous pentachloride in
xylene gave dichlorolactam (2). Compound (2) was reduced to the mono-
chlorolactam (3) by catalytic dehydrogenation (over Pd/C). Treatment of
(3) with sodium azide in dimethyl sulfoxide at 80 C gave azide (4).
H Cl
PCl5 Pd/C
H N Cl
Cl N3
NaN3 BrCH2COOC2H5
N H N H
N3 NH2 NaOH
Pd/C hydrolysis
N H H
CH2COOCH2CH5 CH2COOCH2CH5
NH2 O O
CH2 CH2 C C O C2H5
O (ethyl benzyl pyruvate)
CH2COONa
N CH2 CH2
N H COOCH2CH3
CH2COOH
Scheme 1. Synthesis of benazepril, according to the method of Watthey and
Watthey et al. [5–7].
Alkylation of (4) with ethyl bromoacetate was effected under phase-
transfer conditions with powdered potassium hydroxide and tetrabutyl
ammonium bromide in tetrahydrofuran to yield compound (5), and the
azide function in (5) was then reduced by catalytic hydrogenation (over
Pd/C) to give (6). The aminoester (6) was resolved into its enantiomers via
the tartrate salt. The negatively rotating enantiomer was hydrolyzed to
the sodium salt of the carboxylic acid (7), and this material was
condensed with ethyl benzyl pyruvate in the presence of sodium
cyanoborohydride. Two isomers were obtained in a ratio of 70:30, and
the major isomer (8, benazepril) was obtained in its pure form by
recrystallization.
3.1 Solubility characteristics [3]
The solubility of benazepril hydrochloride is greater than 1 g in 10 mL of
water, ethanol, and methanol.
3.2 Optical activity [1]
The angle observed for the optical rotation of benazepril (C ¼ 1.2% in
ethanol) is  159 , and the corresponding angle of rotation for benazepril
hydrochloride (C ¼ 0.9 in ethanol) is 141.0 and for benazeprilat
(C ¼ 1 in 3% aqueous ammonium hydroxide) is 200.5 .
3.3 X-Ray powder diffraction pattern
The X-ray powder diffraction pattern of benazepril hydrochloride was
obtained using a Simons XRD-5000 diffractometer, and is shown in
Figure 1. A summary of the observed scattering angles (degrees 2-), the
interplanar d-spacings (Å), and the relative intensities (%) of the
observed peaks of benazepril hydrochloride is found in Table 1.
3.4.1 Melting behavior [1]
Benazepril melts at about 148–149 C, benazepril hydrochloride melts at
about 189–190 C, and benazeprilat melts at about 270–272 C.
The differential scanning calorimetry (DSC) thermogram of benazepril
hydrochloride was obtained using a DuPont TA-9900 thermal analyzer
attached to a DuPont Data unit. The thermogram shown in Figure 2 was
obtained at a heating rate of 10 C/min, and was run from 100 to 220 C.
The compound was found to melt at 190.0 C.
2-Theta-Scale (Res. Cent., Sci. Coll., KSU, POB-2455, Riadh-11451, KSA) 08-May-2003 00:52
Figure 1. X-ray powder diffraction pattern of benazepril hydrochloride.
The ultraviolet spectrum of benazepril hydrochloride (obtained at a
concentration of 0.1 mg/mL in water), which is shown in Figure 3, was
recorded using a Shimadzu model 1601 PC ultraviolet–visible spectro-
meter. The drug exhibited only one maximum at 238.5 nm, A[1%,
1 cm] ¼ 198.7, and molar absorptivity ¼ 9160.
The infrared absorption spectrum of benazepril hydrochloride was
obtained in a KBr pellet using a Perkin-Elmer infrared spectro-
photometer. The infrared spectrum is shown in Figure 4, and the
principal peaks were detected at 3435, 2980, 2680, 2467, 1739, 1674, 1524,
1390, 1211, 1003, 754, 702 cm1. The assignments of the major infrared
absorption bands of benazepril hydrochloride are shown in Table 2.
3.5.3 Nuclear magnetic resonance spectrometry
3.5.3.1 1H-NMR spectrum
The proton NMR spectra of benazepril hydrochloride were obtained
using a Bruker Advance System operating at 300, 400, or 500 MHz.
Scattering Angles, d-Spacings, and Peak Relative Intensities from the X-Ray
Powder Diffraction Pattern of Benazepril Hydrochloride
(degrees 2-) (%) (degrees 2-) (%)
6.641 13.2986 53.15 25.244 3.5250 54.16
9.991 8.8460 22.53 25.466 3.4947 38.30
15.335 5.7733 20.66 29.796 2.9960 4.70
20.163 4.4005 100.00 33.818 2.6484 6.00
23.156 3.8380 9.86 43.967 2.05577 4.29
24.369 3.6496 15.88
Total Area / Partial Area
−0.5 178
Figure 2. Differential scanning calorimetry thermogram of benazepril hydrochloride.
220.0 240.0 260.0 280.0 300.0
Wavelength (nm)
Figure 3. Ultraviolet absorption spectrum of benazepril hydrochloride (0.1 mg/mL,
aqueous solution).
%r
4000 3000 2000 1000 600
Wavenumber (cm−1)
Figure 4. Infrared absorption spectrum of benazepril hydrochloride (KBr pellet).
Standard Bruker Software was used to execute the recording of DEPT,
COSY, and HETCOR spectra. The sample was dissolved in D2O, and all
resonance bands were referenced to the tetramethylsilane (TMS) internal
standard. The 1H-NMR spectra of benazepril hydrochloride are shown
in Figures 5–9. The 1H-NMR spectral assignments for benazepril
hydrochloride are provided in Table 3.
3.5.3.2 13C-NMR spectrum
The carbon-13 NMR spectra of benazepril hydrochloride were obtained
using a Bruker Advance Instrument operating at 75, 100, or 125 MHz.
The sample was dissolved in D2O, and tetramethylsilane (TMS) was
added to function as the internal standard. The 13C-NMR spectra are
shown in Figures 10–13, and the gradient HMQC NMR spectra are
shown in Figures 14 and 15. The assignments for the observed resonance
bands associated with the various carbons are provided in Table 4.
The mass spectrum of benazepril hydrochloride was obtained using a
Shimadzu PQ-5000 mass spectrometer. The parent ion was collided with
Assignments for the Infrared Absorption
Bands of Benazepril Hydrochloride
Frequency (cm1) Assignments
3435 Acid O–H stretching
2980 N–H stretching
2680 C–H stretching
2467 C–N stretching
1739 Acid C¼O stretching
1674 C¼C stretching
1524 –CH2 bending
1211 C–C stretching
1003 C–O stretching
754 Di-substituted benzene
702 Mono-substituted benzene
helium as the carrier gas. Figure 16 shows the detailed mass fragmentation
pattern, and Table 5 shows the mass fragmentation pattern of the
substance.
4. METHODS OF ANALYSIS
4.1 Compendial methods
Benazepril hydrochloride is not yet a compendial article in any of the
major pharmacopoeias, so no official tests have been certified as of yet.
ppm 12 10 8 6 4 2
Figure 5. The H-NMR spectrum of benazepril hydrochloride in D2O.
4.2 Statistically compared methods
Wade et al. reported the use of a novel statistical approach for the
comparison of analytical methods to measure angiotensin converting
enzyme [peptidyl-dipeptidase A] activity, and to measure enalaprilat and
benazeprilat [8]. Two methods were used to measure peptidyl-dipeptidase
A, namely hippuryl histidyl leucine (HHL-method) [9], and inhibitor
binding assay (IBA method) [10]. Three methods were used to measure
enalaprilat, namely a radioimmunoassay (RIA) method [11], the HHL
method, and the IBA method. Three methods were used to measure
benazeprilat (then active metabolite of benazepril) in human plasma,
namely gas chromatography–mass spectrometry (GC–MS method) [12],
the HHL method, and the IBA method, and were statistically compared.
First, the methods were compared by the paired t test or analysis of
variance, depending on whether two or three different methods were
under comparison. Secondly, the squared coefficients of variation of the
ppm 4.6 4.4 4.2 4.0 3.8
Figure 6. Expanded 1H-NMR spectrum of benazepril hydrochloride in D2O.
methods were compared by the Jackknife technique. No statistically
significant difference was found between the two assays used to measure
peptidyl-dipeptidase A. Differences were found between the three
methods used to measure enalaprilat, although no obvious reason was
determined for this. Significant differences were also found between the
techniques used to determine benazeprilat; with these being attributed to
the presence of metabolites interfering in the nonspecific assay methods.
Khalil and El-Aliem constructed a coated-wire benazepril-selective
electrode based on incorporation of the benazepril-tetraphenyl borate
ion pair in a poly (vinyl chloride) coating membrane [13]. The influence of
membrane composition, temperature, pH of the test solution, and foreign
ions on the electrode performance was investigated. At 25 C, the
electrode showed a Nernstian response over a benazepril concentration
ppm 2.5 2.0 1.5
range of 1.26  105 to 0.58  102 M, at 25 C and was found to be very
selective, precise, and usable over the pH range of 2.5–9.2. The standard
electrode potentials were determined at 20, 25, 30, 35, 40 and 45 C, and
used to calculate the isothermal temperature coefficient of the electrode.
Temperatures higher than 45 C were found to seriously affect the
electrode performance. The electrode was successfully used for the
potentiometric determination of benazepril hydrochloride both in pure
solutions and in pharmaceutical preparations.
4.4 Spectrophotometric methods
Erk and Onur used two new spectrophotometric methods for the
simultaneous determination of benazepril hydrochloride and hydro-
chlorothiazide in their binary mixtures [14]. In a derivative spectro-
photometric method, dA/d values were measured at 260.7 and 239.8 nm
for benazepril hydrochloride and hydrochlorothiazide, respectively, for
ppm 6 4 2
Figure 8. COSY 1H-NMR spectrum of benazepril hydrochloride in D2O.
their combination solutions. The relative standard deviation of the
method was found to be 0.77% for benazepril hydrochloride and 0.28%
for hydrochlorothiazide. In an absorbance ratio method, quantification
of benazepril hydrochloride and hydrochlorothiazide was performed
using the absorbances measured at 264, 258.3, and 236.4 nm in their
combination solutions. The relative standard deviations of the method
were found to be 1.26% and 0.36% for benazepril hydrochloride and
hydrochlorothiazide, respectively. These two methods have been
successfully applied to tablets containing only these drugs.
Panderi used a second-order derivative spectrophotometric method for
the simultaneous determination of benazepril hydrochloride and hydro-
chlorothiazide in tablets [15]. The determination of benazepril hydro-
chloride in the presence of hydrochlorothiazide was achieved by
ppm 3.5 3.0 2.5 2.0
measuring the second order derivative signals at 253.6 and 282.6 nm,
while the second-order derivative signal at 282.6 nm was measured for the
determination of hydrochlorothiazide. The linear dynamic ranges were
14.80–33.80 mg/mL for benazepril hydrochloride and 18.5–42.2 mg/mL for
hydrochlorothiazide. The correlation coefficients for the calibration
graphs exceeded 0.9998, the precision (% RSD; n ¼ 5) was better than
1.43%, and the accuracy was satisfactory (error less than 0.99%). The
detection limits were found to be 2.46 and 1.57 mg/ml for benazepril HCl
and hydrochlorothiazide, respectively. The method was applied to the
quality control of commercial tablets, and proved to be suitable for rapid
and reliable analysis.
El-Yazbi et al. presented three spectrophotometric methods for
determination of benazepril hydrochloride in its single and multi-
component dosage forms [16]. The methods are (i) derivative
Assignments for the Proton Nuclear Magnetic Resonance
Spectrum of Benazepril Hydrochloride
12 H 17 18 19
N 13 CH2 CH2
H 14 15 16
COOCH2CH3
Chemical Shift Number Multiplicity* Assignment
(ppm, relative of Protons (proton at
to TMS) carbon number)
1.04 3 s (7 Hz) 16
2.08–3.03 8 m (5, 6, 17, 18)
3.73–3.76 1 t (6.5 Hz) 13
3.79–3.82 1 q (8.5 Hz) 4
3.93–4.04 2 m 15
4.31–4.54 2 dd 2
7.09–7.26 9 m (9–12, 20–24)
*s: singlet; t: triplet; q: quartet; m: multiplet; dd: double doublet.
spectrophotometry to resolve matrix interferences, (ii) reaction with
bromocresol green (BCG), and (iii) reaction with 3-methylbenzothialo-
zone hydrazone (MBTH) after alkaline hydrolysis. Powdered tablets
(coating removed for the MBTH method) equivalent to 10 mg of
benazepril hydrochloride were mixed with 25 mL of water (10 mL MBTH
method), shaken for 45 min, filtered, treated and analyzed. For method
(i), derivative spectra were measured at 240 and 244 nm. For method (ii),
ppm 200 150 100 50 13.503
Figure 10. C-NMR spectrum of benazepril hydrochloride in D2O.
the color produced upon reaction with 4 mL 0.1% BCG at pH 2.2 was
measured at 412 nm (" ¼ 18523). For method (iii), after alkaline
hydrolysis, the color produced on reaction with 1 mL each of MBTH
(0.2% in 0.1 M HCl), FeCl3 (1% in 0.1 M HCl) and ethanol was
measured at 593 nm (" ¼ 11059). The color of the BCG and MBTH
products was found to be stable for 20 min. Beer's law was obeyed from
8–24, 4–24 and 4–40 mg/mL using methods (i), (ii) and (iii), respectively,
with intraday RSD values (n ¼ 5) being less than 2%. Recoveries were in
the range of 98.1–101.8%. The BCG approach was found to be a general
method, derivative spectrophotometry was used for benazepril hydro-
chloride in tablet forms (single or in combination with hydrochlorothia-
zide), and the MBTH method was applied to specific determinations of
the degradation product of benazepril hydrochloride.
A derivative UV spectrophotometric method was used by Bonazzi et al.
for the determination of benazepril hydrochloride and other angiotensin
converting enzyme inhibitors in their pharmaceutical dosage forms [17].
ALL PROTONATED
ppm 200 175 150 125 100 75 50 25 0
Figure 11. DEPT 45 C-NMR spectrum of benazepril hydrochloride in D2O.
The drug was determined by second derivative spectrophotometry using
a Uvidec 610 double beam spectrophotometer, and the absorbance was
measured at 259.2 nm. Beer's law was obeyed in range of 10–30 mg/mL.
Banoglu et al. reported the use of rapid, simple, and reliable spectro-
scopic methods (absorbance ratio method and Vierordt's method) for the
quantitative determination of benazepril hydrochloride and hydrochloro-
thiazide in the dissolution testing of these two drugs in commercial tablets
[18]. A 249 nm wavelength was chosen as the isobestic point in the
absorbance ratio method, and the absorbance ratios A236/A249 nm for
benazepril and A269/A249 nm for hydrochlorothiazide were used for
calculation of the regression equation. For the Vierordt' method,
A[ 1%,1 cm] values were obtained at 236 and 269 nm for both drugs and
were used for their quantitative analyses.
CH ONLY
ppm 200 150 100 50
Erk presented a ratio spectra derivative spectrophotometric method
and a Vierordt's method for the simultaneous determination of hydro-
chlorothiazide in binary mixtures with benazepril hydrochloride,
triametrene, and cilazapril [19]. A mass of powder equivalent to one
tablet was dissolved in 1:1 methanol/0.1 M Hydrochloric acid (solvent A),
shaken for 30 min, filtered and the residue was washed with 3  10 mL of
solvent A. Combined solutions were diluted to 100 mL with solvent A,
and then diluted with methanol for analysis by (i) Vierordt's method, and
(ii) ratio spectra derivative spectrophotometry. For method (i), the
absorbance of the solution was measured at 271.7, 238.1, 234.1, and
210.7 nm for hydrochlorothiazide, benazepril, triametrine, and cilazapril,
respectively, and the corresponding concentrations were calculated using
simultaneous equations. For method (ii), the absorption spectra of
solutions containing different amounts of hydrochlorothiazide in a
DEPT 135
CH, CH3 UP
CH2 DOWN
ppm 200 150 100 50 0
Figure 13. DEPT 135 C-NMR spectrum of benazepril hydrochloride in D2O.
binary mixture with benazepril hydrochloride, triametrine, or cilazapril
were recorded within 200–350 nm ( ¼ 1 nm), and the first derivative
ratio spectra obtained. The contents of hydrochlorothiazide with
benazepril hydrochloride, with triametrine and with cilazapril were
determined from signals at 269.5 and 238.8 nm, 260.7 and 284.4 nm, and
273.7 and 236.8 nm, respectively. For both methods, Beer's law was
obeyed over the range of 8–36 mg/mL of benazepril hydrochloride.
Relative standard deviation (n ¼ 5) were 0.3–1.8%, and recoveries were
98.6–99.9%.
El-Gindy et al. presented different spectrophotometric methods for the
chlorothiazide in pharmaceutical tablets [20]. The first method depends
gradient HMQC
123.2 100
ppm ppm
Figure 14. Gradient HMQC NMR spectrum of benazepril hydrochloride in D2O.
on second-derivative ultraviolet spectrophotometry with zero crossing
and peak-to-base measurements. The second derivative amplitude at
214.8 and 227.4 nm were selected for the assay of benazepril
hydrochloride and hydrochlorothiazide, respectively. The second
method depends on the second derivative of the ratio spectra by
measurement of the amplitude at 241.2 and 273.2 nm for benazepril
hydrochloride and hydrochlorothiazide, respectively. Chemometric
methods, classical least squares, and principal component regression
were applied to analyze the mixture. Both the chemometric methods were
applied to the zero and first order spectra of the mixture. These methods
were successfully applied to the determination of the two drugs in
laboratory prepared mixtures and in commercial tablets.
Dinc reported that three chemometric techniques, classical least
squares (CLS), inverse least square (ILS) and principal component
regression (PCR), could be applied to the spectrophotometric analysis
of two pharmaceutical tablet formulations containing hydrochlorothia-
zide and benazepril hydrochloride in the presence of their overlapping
spectra [21]. A training set was randomly prepared by using the
different composition mixtures containing 0–22 mg/mL of hydrochloro-
thiazide and 0–36mg/mL of benazepril hydrochloride in 0.1 M sodium
hydroxide. The chemometric calibrations were constructed by using the
prepared training set and its measurement of the absorbance values at
24 points in the spectrum from 220 to 350 nm. The obtained
chemometric calibrations were used for the prediction of hydrochloro-
thiazide and benazepril hydrochloride levels in the samples. The
validation of the three techniques was realized by analyzing the
synthetic mixtures of the two drugs. The CLS, ILS, and PCR
techniques are suitable for the chemometric determination of benazepril
Assignments for the Carbon-13 Nuclear Magnetic
Resonance Spectrum of Benazepril Hydrochloride
4 CH 24 23
O COOCH2CH3
Chemical Shift Assignment Chemical Shift Assignment
(ppm relative (Carbon (ppm relative (Carbon
to TMS) number) to TMS) number)
172.4 1 127.9 12
168.2 14 64.1 2
139.9 7 59.0 4
139.4 8 57.8 13
129.2 20,24 31.3 17
129.1 10 30.6 18
50 100 150 200 250 300 350 400 450
Figure 16. Mass spectrum of benazepril hydrochloride.
hydrochloride and hydrochlorothiazide in synthetic mixtures and in
tablet formulations.
4.5 Colorimetric method
Belal et al. developed a simple and sensitive spectrophotometric method
for the determination of benazepril hydrochloride in tablets [22]. The
method is based on the reaction of the drug with potassium permanga-
nate in the presence of sodium hydroxide to produce a bluish-green
colored species measurable at 609.4 nm. The absorbance-concentration
plot is linear over the range 1–8 mg/mL, with a limit of detection of 0.1
mg/mL (2.17  107 M). The molar absorptivity was 4.07  104 L/mol/
cm, and the Beer's Law correlation coefficient (n ¼ 6) was 0.9991. The
different experimental parameters affecting the development and stability
of the color were studied carefully, and optimized. This method was
applied successfully to the determination of benazepril hydrochloride in
its dosage forms. The percentage recoveries  SD (n ¼ 9) were
99.79%  1.4 and 100.5  1.48 for tablets containing 10 and 20 mg,
Mass Spectral Fragmentation Pattern for Benazepril HCl
Formula Structure
424 2.5% C24H28N2O5
H +
O COOC2H5
395 1.1% C22H23N2O5 M þ –CH2CH3
378 7.7% C22H22N2O4 M þ –CH3CH2OH
351 100% C21H23N2O3
N+
CH C O
COOH
219 4.81% C12H13NO3
206 9.6% C12H16NO2
190 43% C12H14O2
CH3CH2 O C
146 26.5% C10H10O
O C CH CH
144 26.2% C10H10N
132 24.1% C9H10N
130 24.6% C9H8N
117 28.5% C9H9
HC HC H2C
91 54.5% C7H7
77 7.2% C6H5
42 14.31% C2H2O
H2C C O
respectively. The results obtained were in good agreement with those
obtained using a reference spectrophotometric method. The method
could also be applied to the determination of benazepril in the presence of
co-formulated hydrochlorothiazide..
Belal et al. developed a simple and highly sensitive voltammetric method
for the determination of benazepril and ramipril [23]. The compounds
were treated with nitrous acid, and the cathodic current produced by the
resulting nitroso derivatives was measured. The voltammetric behavior
was studied by adopting direct current (DCt), differential pulse (DPP),
and alternating current (ACt) polarography. Both compounds produced
well-defined, diffusion-controlled cathodic waves over the whole pH
range when studied in Britton-Robinson buffer (BPb). At pH 3 and 5, the
values of diffusion-current constants (Id), were 5.9  0.4 and 6.66  0.61
for benazepril and ramipril, respectively. The current concentration plots
for benazepril were linear over the range of 1.5–40 and 0.1–30 mg/mL in
the DCt and DPP modes, respectively. The limits of detection (S/N ¼ 2)
were 0.015 mg/mL (about 3.25  108 M) and 0.012 mg/mL (about
2.88  108 M) for benazepril and ramipril, respectively (when using the
DPP mode). When applied to the determination of both compounds in
dosage forms, the results obtained using this method were in good
agreement with those obtained using reference methods. The method was
applied to the determination of benazepril in spiked human urine and
plasma. When using the DPP mode, the percentage recoveries were
96.2  1.21 and 95.7  1.61, respectively.
4.7 Chromatographic methods of analysis
4.7.1 Thin-Layer chromatographic methods
Hassib et al. developed a new, simple, sensitive, and fast thin layer
chromatographic scanning densitometric method for the simultaneous
determination of benazepril hydrochloride and hydrochlorothiazide
using ethyl acetate–methanol–ammonia (85:20:10 v/v) as the developing
system [24]. The Rf values were 0.33 and 0.68 for benazepril
hydrochloride and hydrochlorothiazide, respectively. The minimum
detection limit obtained was 0.12 mg/spot and 0.24 mg/spot for benazepril
hydrochloride and hydrochlorothiazide, respectively. The mean percen-
tage recoveries were 100.04  1.02 and 99.31  1.009 for benazepril
hydrochloride and hydrochlorothiazide, respectively. The method could
be successfully used for the determination of pure, laboratory made
mixtures, and pharmaceutical dosage forms. The results obtained were
compared with those obtained by spectrophotometric methods.
El-Gindy et al. described a high performance thin layer chromatographic
(HPTLC) method for the simultaneous determination of benazepril
hydrochloride and hydrochlorothiazide in binary mixtures [25]. The
method is based on the HPTLC separation of the two drugs, followed
by densitometric measurement of their spots at 238 and 275 nm for
benazepril hydrochloride and hydrochlorothiazide, respectively. The
separation was carried out on Merck HPTLC aluminum sheets coated
with silica gel 60 F254, using ethyl acetate–methanol–chloroform (10:3:2
v/v) as mobile phase. A second order polynomial equation was used for
the regression line in the range 2–20 and 2.5–25 mg/spot for benazepril
hydrochloride and hydrochlorothiazide, respectively. The method was
successfully applied to the determination of the two drugs in laboratory
prepared mixtures and in commercial tablets. No chromatographic
interference from tablet excipients was found.
4.7.2 Gas chromatographic–mass spectrometric methods (GC–MS)
Sioufi et al. determined benazepril and its active metabolite, benazeprilat,
in plasma and urine by capillary gas chromatography with mass-selective
detection [12]. Plasma or urine was extracted with Amberlite XAD-2
or Extrelut 1, respectively, and then derivatized with diazomethane.
A portion (2 mL) of the final derivative solution in toluene was analyzed on
a fused silica column (12.5 m  0.2 mm) coated with cross-linked
methylsilicone, operated at 210 C (held for 0.5 min) to 290 C at
50 C/min, with helium as the carrier gas. Detection was by selected-ion
monitoring at m/z ¼ 365 for the methyl ester derivatives of benazepril and
its free acid analogue, benazeprilat and at m/z ¼ 370 for the methyl ester
derivatives of deuterated benazepril and benazeprilat (internal standards).
Detection limits were 4.5 and 6.5 nM for benazepril and benazeprilat,
respectively, in plasma. The corresponding values in urine were 11 and 53
nM. The calibration graphs were linear from 5.4 to 217 and 10.9 to 542.5
nM of benazepril in plasma and urine, respectively, and the corresponding
values for benazeprilat were 6.6–259.6 and 52.9–1310.4 nM
Sereda et al. developed an electron-capture gas chromatographic
detection method for the detection of benazepril and other angiotensin-
converting enzyme (peptidyl-dipeptidase) inhibitors [26]. Sample
solutions (0.001–0.1 mL) containing the inhibitor or metabolite were
vertex-mixed at 70 C for two hours with 10 mg of potassium acetate, 3
mL of acetone, 0.1 mL of 0.1% ramiprilat (the internal standard)
solution, and 0.1 mL of alpha-bromopentafluorotoluene. A 0.25 mL
portion of the cooled solution was evaporated to dryness at 100 C, the
residue was dissolved in 0.25 mL of acetone, and a 0.5 mL portion of the
resulting solution was injected onto a glass column (30 m  0.252 mm)
coated with methyl silicone and operated at 295 C. Helium was used as
the carrier gas (25 psi), and the method employed a 63Ni electron capture
detector. Benazepril and eight ACE inhibitors were well separated, and
calibration graphs were linear over the range of 0.3–15.8 ng of drug on
the column.
Pommier et al. determined benazepril and its active metabolite
(benazeprilat) in human plasma using capillary gas chromatography-
mass-selective detection, with their respective labeled internal standards
[27]. The method was developed and validated according to international
regulatory requirements. After addition of the internal standards, the
compounds were extracted from plasma by solid-phase extraction using
an automated 96-well plate technology. After elution, the compounds
were converted into their methyl ester derivatives by means of a safe and
stable diazomethane derivative. The methyl ester derivatives were
determined by gas chromatography using a mass-selective detector at
m/z ¼ 365 for benazepril and benazeprilat, and at m/z ¼ 370 for the
internal standards. Intra- and interday accuracy and precision were found
to be suitable over a range of concentrations between 2.5 and 1000 ng/mL.
Kaiser et al. developed and validated a specific and sensitive gas
chromatographic–mass spectrometric method for the simultaneous
quantification of unchanged benazepril and its active metabolite (the
dicarboxylic acid) in plasma and urine [28]. The 2H5-labeled analogues of
benazepril and its metabolite were used as internal standards. The
compounds were isolated from plasma and urine under acidic conditions
using XAD-2 resin or Extrelut 1 columns. Following derivatization
with diazomethane, the samples were analyzed by packed-column gas
chromatography–electron impact mass spectrometry with selected ion
monitoring. The analysis of spiked plasma and urine samples demon-
strated the good accuracy and precision for the method, which was found
to be suitable for use in pharmacokinetic and bioavailability studies
with benazepril hydrochloride in humans.
Maurer et al. developed a gas chromatographic–mass spectrometric
screening procedure for the detection of benazepril and other
angiotensin-converting enzyme inhibitors, their metabolites, and angio-
tensin II receptor antagonists in urine as part of systematic toxicologic
analysis procedure for acidic drugs and poisons after extractive
methylation [29]. The part of the phase-transfer catalyst remaining in
the organic phase was removed by solid-phase extraction in a diol phase.
The compounds were separated by capillary gas chromatography, and
identified by computerized mass spectrum in the full scan mode. Using
mass chromatography with ion detection at m/z of 157, 160, 172, 192,
204, 220, 234, 248, 249 and 262, the possible presence of the ACE
inhibitors, their metabolites, and angiotensin II antagonist could be
indicated. The identity of positive signals in these mass chromatograms
was confirmed by comparison of the peak underlying full mass spectra
with reference spectra recorded during this study. This method allowed
detection of therapeutic concentrations of benazepril and five other ACE
inhibitors, their metabolites (or both), as well as the therapeutic
concentration of the angiotensin II antagonist (valsartan) in human
urine samples. The overall recoveries ranged between 80 and 88%, with
coefficient of variation of less than 10%, and the limit of detection of at
least 10 ng/mL (signal-to-noise ratio 3) in the full-scan mode.
4.7.3 High performance liquid chromatography
Panderi and Parissi-Poulou developed a microbore liquid chromato-
graphic method for the simultaneous determination of benazepril
hydrochloride and hydrochlorothiazide in pharmaceutical dosage forms
[30]. The use of a BDS C-18 microbore analytical column was found to
result in substantial reduction in solvent consumption and in increased
sensitivity. The mobile phase consisted of a mixture of 25 mM sodium
dihydrogen phosphate buffer (pH 4.8) and acetonitrile (11:9 v/v), pumped
at a flow rate of 0.4 mL/min. Detection was effected at 250 nm using an
ultraviolet absorbance detector. The intra- and inter-day relative
standard deviation values were less than 1.25% (n ¼ 5), while the
relative percentage error was less than 0.9% (n ¼ 5). The detection limits
obtained according to the IUPAC definition were 0.88 and 0.58 mg/mL
for benazepril hydrochloride and hydrochlorothiazide, respectively. The
method was applied to the quality control of commercial tablets and
content uniformity test, and proved to be suitable for rapid and reliable
Radhakrishna et al. developed a gradient liquid chromatographic method
for the determination and purity evaluation of benazepril hydrochloride
in bulk and pharmaceutical dosage forms [31]. The method is simple,
rapid, and selective. 5-Methyl-2-nitrophenol was used as the internal
standard. The method is linear over the range of 50–800 mg. The precision
of the inter- and intra-day assay variation of benazepril hydrochloride is
below 1.6% relative standard deviation. The accuracy, determined as
relative mean-error (RME) for the intra-day assay, is within  2%. The
method is stability indicating, and is useful in the quality control of bulk
manufacturing and also in pharmaceutical formulations.
Manna et al. described a rapid and accurate reversed-phase ion-paring
liquid chromatographic method, with ultraviolet detection, for the
simultaneous assay of benazepril hydrochloride, fosinopril sodium,
ramipril and hydrochlorothiazide [32]. The separation was achieved on
a LC-8 (12.5 cm  4 mm, 5 mm particle size) column. The mobile phase
consisted of 20 mM sodium heptansulfonate (pH 2.5) and methanol (8:17
v/v). Validation of the method showed it to be precise, accurate, and
linear over the concentration range of analysis, with a limit of detection
of 2 ng for benazepril. The method was applied to the analysis of three
different binary commercial formulations.
El-Gindy et al. described a liquid chromatographic method for the
chlorothiazide [25]. The liquid chromatography was based on the
separation of the two drugs on a reversed phase ODS column at ambient
temperature, using a mobile phase consisting of acetonitrile and water
(7:13) and adjusting to pH 3.3 with acetic acid. Quantitation was
achieved with ultraviolet detection at 240 nm based on peak area, with
linear calibration curves at concentration ranges of 10–60 and 12.5–75
mg/mL for benazepril hydrochloride and hydrochlorothiazide,
respectively. The method was successfully applied to the determination
of the two drugs in laboratory prepared mixtures and in commercial
tablets. No chromatographic interference from tablet excipients was
Dappen et al. studied the performance of a commercial (Polarmonitor,
IBZ, Hannover, Germany) and a laboratory designed polarimetric
detector in conjunction with a HPLC method based on a Nucleosil 100
RP-18 column, an ultraviolet detector, and aqueous 58% methanol as the
mobile phase [33]. The polarimetric signal was proportional to the excess
concentration of one enantiomer in a mixture of two, and measurement
of the total concentration of both enantiomers using the ultraviolet
detector then permitted the enantiomer ratio to be calculated. The
unexpectedly strong polarimetric signals produced by some compounds
(such as 1-phenylethyl alcohol) were attributed to RI-related effects
generated by the eluting peaks. One example of a pharmaceutical
application of polarimetric detection was the detection of the (R,R)-
enantiomer impurity of benazepril (which itself is a pure (S,S)-
enantiomer), with a detection limit of better than 0.5%. Another
application was the determination of the diastereomeric ratio (not
separable on a conventional reversed-phase column) in the preparation of
10-EDAM WS. This chromatographic system used a 5 mm Neocleosil
100 RP-18 reversed phase, packed into a 12.5 cm  4.6 mm column, and
was eluted using Veible buffer–water–methanol (17:25:58). The Vieble
buffer is made of 90 mM potassium chloride and 10 mM hydrochloric
acid at pH 2.06. The flow rate was 0.25 mL/min and temperature was
Bonazzi et al. used an HPLC method and a second derivative ultraviolet
spectroscopy method for the analysis of benazepril and other angio-
tensen-converting enzyme inhibitors [17]. For HPLC, 20 mL sample
solutions containing the drug and an internal standard dissolved in 1:1
acetonitrile/20 mM sodium heptanesulfonate (pH 2.5) were used. HPLC
was performed on a 5 mm Hypersil ODS column (25 cm  4.5 mm) with a
mobile phase mixture consisting of (A) 20 mM sodium heptanesulfonate
(pH 2.5) and (B) 19:1 acetonitrile–tetrahydrofuran, eluted at a flow rate
of 1 mL/min, and with detection at 215 nm. The A/B mixture used was
52:48 for benazepril. A low pH of 2.5 was essential to avoid peak splitting
and band broadening.
Cirilli and La-Torre used a reversed-phase high performance liquid
chromatographic method, on a chiral alpha-acid glycoprotein (AGP)
column, for the stereoselective analysis of benazepril and its stereo-
isomers [34]. A 10 mL sample of benazepril, its enantiomer, two
diastereomers, and a chiral active metabolite (benazeprilat) in ethanol
(24 mg/mL) was injected onto a stainless steel chiral AGP based column
(10 cm  4 mm) or a Kromasil C-8 (5 mm) column (25 cm  4.6 mm).
Analytes were eluted (0.9 mL/min) with 0.01–0.06M potassium dihydro-
gen phosphate buffer (pH 3–7) containing various proportions of
an organic modifier, and detected at 240 nm. A baseline separation
of the five compounds was achieved in less than 35 min on an AGP
column with 0.06 M-phosphate buffer (pH 4) and ethanol (17:3), or
propan-2-ol (12.33:1). Detection limits (calculated as three times the
signal-to-noise), when using propan-2-ol modifier, were 1.5 ng for
benazepril, 5 ng for its enantiomer, the two diastereomers, and 0.6 ng
for benazeprilat.
Gumieniczek and Przyborowski determined benazepril and cilazapril in
pharmaceuticals by a high performance liquid chromatographic method
[35]. Ground tablets equivalent to 25 mg of benazepril or cilazapril were
extracted with methanol. Portions (20 mL) of the extract were analyzed by
HPLC on a 10 mm LiChrosorb RP-18 column (25 cm  4 mm) with
phosphate buffer (pH 2.4)/acetonitrile (7:3) as the mobile phase (eluted at
1 mL/min), and detection at 211 nm. Quantitation was performed using
enalapril maleate (4 mg/mL in methanol) as the internal standard.
Calibration graphs were linear from 10–50 and 40–200 mg/mL of
benazepril and cilazapril, respectively. Recoveries of the two drugs
ranged from 96.34–102.04% and 103.08–107.96%, respectively. Relative
standard deviations (n ¼ 5) were less than 4.45% and less than 3.5 for
benazepril and cilazapril, respectively.
Hassib et al. developed two chromatographic procedures for the
chlorothiazide in laboratory made mixtures, and in pharmaceutical
dosage forms (Cibadrex tablets) using reversed phase HPLC and TLC
methods [24]. For reversed phase HPLC, a very sensitive, rapid, and
selective method was developed. The linearity ranges were 32–448 ng/
20 mL and 40–560 ng/20 mL for benazepril hydrochloride and hydro-
chlorothiazide, respectively. The corresponding recoveries were 99.38
1.526 and 99.2  1.123. The minimum detection limits were 7 ng/20 mL
for benazepril and 14 ng/20 mL for hydrochlorothiazide. The method
could be successfully applied for the determination of laboratory made
mixtures and for pharmaceutical dosage forms. The results obtained were
compared with those obtained by a spectrophotometric method.
Banoglu et al. reported a comparison of the spectrophotometric
methods (absorbance ratio method and Vierordt's method) with
HPLC for quantitative determination in the dissolution testing of
benazepril hydrochloride and hydrochlorothiazide commercial tablets
[18]. In the HPLC method, simultaneous determination of the two
drugs from the dissolution medium was achieved using a mobile phase
containing phosphate buffer (0.01 M, pH 6.2) and acetonitrile (65:35)
on a Supelcoil LC-18 (4.6  250, 5.6 mm) reversed phase column.
Dissolution tests of commercial tablets were carried out according
to USP XXII method (0.1 M hydrochloric acid at 37  0.5 C, and
paddle rotation of 50 rpm). Comparison of the dissolution data from
the HPLC and the two spectroscopic methods indicated good
correlation between the methods. It was concluded that both the
spectroscopic methods and the HPLC method are suitable for routine
analyses of benazepril hydrochloride and hydrochlorothiazide in
dissolution testing of commercial tablets.
Sreenivas et al. described a validated HPLC method for the assay of
quinapril and benazepril hydrochloride in pharmaceuticals, and the purity
evaluation of quinapril bulk drug substance [36]. Solutions of quinapril
hydrochloride (2.5 mg/mL) and benazepril hydrochloride (1.5 mg/mL)
were prepared by dissolving the compounds in mobile phase A (5:3:2
mixture of 0.05 M potassium dihydrogen phosphate (pH 4)/acetonitrile/
methanol). The internal standard for quinapril hydrochloride was 0.3 mg/
mL benazepril hydrochloride, and the internal standard for benazepril
hydrochloride was 0.3 mg/mL quinapril hydrochloride. Extracts were
prepared by grinding up to twenty Acupril 40 mg tablets and Lotensin
40 mg tablets, and extracting with methanol. Extracts were diluted to 0.2
mg/mL and 0.3 mg/mL for each drug substance. Samples (10 mL) were
analyzed on a 5 mm Hypersil C-18 BDS column (25 cm  4.6 mm) at
50 C, eluted with mobile phase A, and employing detection at 215 nm.
The calibration curve was linear from 0.1 to 0.6 mg/mL of quinapril, and
from 0.15 to 0.75 mg/mL of benazepril hydrochloride. Intra-day precision
and accuracy was evaluated (n ¼ 3) at concentrations of quinapril
hydrochloride and benazepril hydrochloride of 0.15 mg/mL, 0.3 mg/
mL, and 0.45 mg/mL. Relative standard deviations ranged from 0.57 to
0.97% and 0.3 to 0.88% for quinapril hydrochloride and benazepril
hydrochloride, respectively. Interday precision and accuracy were
calculated (n ¼ 3) for three days. Relative standard deviations ranged
from 0.65–0.94% and 0.74–1.19% for quinapril hydrochloride and
benazepril hydrochloride, respectively. Recoveries from tablets ranged
from 96.3 to 98.4% (relative standard deviations 0.32–0.43%) for
quinapril, and 100.4 to 103.5% (relative standard deviations ¼ 0.98–
1.17%) for benazepril hydrochloride. The detection limit was 2.5 mg/mL
for a 10 mL injection of quinapril hydrochloride and benazepril
hydrochloride. The quantitation limit was 10 mg/mL for both quinapril
hydrochloride and benazepril hydrochloride.
Gana et al. developed and validated a reversed-phase high performance
liquid chromatographic method for the kinetic investigation of the
chemical and enzymatic hydrolysis of benazepril hydrochloride [37].
Kinetic studies on the acidic hydrolysis of benazepril hydrochloride were
carried out in 0.1 M hydrochloric acid solution at 50, 53, 58 and 63 C.
Benazepril hydrochloride appeared stable in pH 7.4 phosphate buffer at
37 C, and showed susceptibility to in vitro enzymatic hydrolysis with
porcine liver esterase (PLE) in a pH 7.4 buffered solution at 37 C.
Benazeprilat appeared to be the major degradation product in both the
chemical and enzymatic hydrolysis studies. Statistical evaluation of the
HPLC methods revealed their good linearity and reproducibility.
Relative standard deviation was less than 4.76, while detection limits
for benazepril hydrochloride and benazeprilat were 13  107 and
9  107, respectively. Treatment of kinetic data of the acidic hydrolysis
was carried out by nonlinear regression analysis, and k values were
determined. The kinetic parameters of the enzymatic hydrolysis were
determined by nonlinear regression analysis of the data using the
Michaelis–Menten equation. Chromatographic separation was per-
formed on a reversed-phase BDS C18 column (25 cm  3 mm i.d., 5 mm
particle size) Shandon Scientific Ltd. The mobile phase consisted of 0.020
M disodium hydrogen phosphate adjusted to pH 3 with phosphoric acid
and methanol (40:60, v/v), which was filtered through a 0.20 mm nylon
membrane filter and degassed under vacuum prior to use. A flow rate of
0.35 mL/min with a column inlet pressure of 2000 psi was used in order to
separate the drug and its degradation products.
Graf et al. used an enzymatic method for the determination of
benazeprilat in plasma and urine [38]. For this determination, urine
and plasma samples were diluted 1:20 with phosphate buffer (pH 8.3)
and heated at 75 C for 5 min to inactivate endogenous angiotensin-
converting enzyme (ACE). Plasma samples were then incubated in blank
plasma in 0.25 M disodium hydrogen phosphate/0.75 M sodium
chloride (adjusted to pH 8.3 with 1 M-phosphoric acid) for 12 min
before the addition of hippuryl-L-histidyl-L-leucine as substrate and
further incubation for 80 min. The reaction was stopped with
methanolic phosphoric acid, whereupon acetonitrile and mandelic acid
(internal standard) were added, and the mixture centrifuged. A 15 mL
portion of the supernatant solution was analyzed on a column (25
cm  4.6 mm) of LiChrosorb-NH2 (10 mm) with a mobile phase (2.3
mL/min) consisting of 1:4 acetonitrile/10 mM potassium dihydrogen
phosphate (adjusted to pH 3 with 1 M phosphoric acid). Detection was
effected on the basis of the UV absorbance at 228 nm. Urine was treated
as for plasma with rabbit lung ACE solution in place of blank plasma,
[3H]-hippuryl-glycyl-glycine as the substrate, and 2 M hydrochloric acid
saturated with sodium sulfate to stop the reaction. The released [3H]-
hippuric acid was extracted into a water-immiscible scintillation cocktail
and counted without separation of the phases. Mean recoveries of
benazeprilat were 97.1–104% and 92.9–116% from plasma and urine,
respectively, with a coefficient of variation (n ¼ 8) of 8% for 20 nM
benazeprilat. Detection limits were 10 and 20 nM benazeprilat in urine
and plasma, respectively. The results agreed well with those obtained
by GC–MS.
Cakir et al. reported the use of a high performance liquid chromato-
graphic method and a spectrophotometric method for the analysis of
benazepril hydrochloride and hydrochlorothiazide [39].
Barbato et al. reported the analysis of benazepril and six other
angiotensin converting enzyme inhibitors by a HPLC method [40]. The
influence of different organic modifiers and counter-ions in the mobile
phase at different pH values has been investigated, allowing the
identification of the best experimental conditions for the analysis of the
drug and the other ACE inhibitors. The synergistic effect of triethylamine
and propanol in the eluting phase appeared to be a critical factor in order
to obtain satisfactory peak shapes and resolution. A reversed phase
Spherisorb 5 ODS-2 stainless steel column (25 cm  4.6 mm) (Phase
Separation-Clwid-UK) was employed. The mobile phases were con-
stituted by phosphate buffer at various pH values, and contained
counter ions at different percentages in mixtures with different amounts
of acetonitrile, methanol, tetrahydrofuran, or n-propanol organic
modifiers. Cetyltrimethylammonium bromide (CTAB), sodium dodecyl
sulfate, n-butylamine, and triethylamine were used as counter ions. The
analyses were performed at room temperature. The HPLC was
performed on a Waters liquid chromatographs equipped with a
Model 7125 Rheodyne injection valve and a Model 486 UV detector
set at 240 nm.
4.7.4 Capillary electrophoresis methods
Gotti et al. developed a capillary electrophoresis method for the
determination of benazepril hydrochloride and other angiotensin-
converting enzyme inhibitors in pharmaceutical tablets [41]. Since a
free solution capillary electrophoresis system failed to achieve a complete
separation of closely related compounds (benazepril and others),
alkylsulfonic additives (sodium heptane sulfonate and 10 camphor-
sulfonic acid) were added to the running buffer. This resulted in an
improved separation, suggesting a favorable effect of ion-pairing
interactions between analytes and additives. The separations were carried
out in acidic medium and a systematic investigation of electrophoresis
parameters was made to evaluate the performance of the selected
additives. Under optimized conditions, benazepril and ramipril in their
commercial dosage forms were determined, confirming the applicability
of the developed capillary electrophoresis approach to the analysis of
pharmaceutical samples. The results were compared with those obtained
applying a previously described and validated HPLC method.
Hillaert and Van den Bossche used capillary electrophoresis method for
the separation of benazepril among eight other angiotensin-converting
enzyme inhibitors [42]. The method was investigated with respect to the
pH of the running buffer, the organic modifiers, and plausible
surfactants. The most critical parameter was found to be the pH of the
running buffer. The addition of sodium dodecyl sulfate had a negative
influence on the peak symmetry, and the selectivity was not improved.
The separation of the eight compounds can be performed by means of
two phosphate buffers (each 100 mM) at pH 7 and 6.25, respectively. The
combination is necessary for the selective identification of structurally
related substances because of their similar pKa values.
The same authors also applied capillary electrophoresis to the study of
benazepril hydrochloride and several angiotensin-converting enzyme
inhibitors [43]. Separation of the compounds was performed by means of
two phosphate buffers (each 0.1 M) at pH 7 and 6.25, respectively [42].
Due to the highest selectivity of the first mentioned running buffer, the
same system has been applied for the quantification of benazepril and
other compounds in their corresponding pharmaceutical formulations.
It was found that the possibility of simultaneous identification and
quantification of the active ingredient in the finished products was
especially attractive, and that excipients do not adversely affect the
results. This article deals with the validation of some parameters of the
quantitative analysis, namely linearity, precision, accuracy, and robust-
ness [43].
Hillaert et al. used a statistical experimental design to optimize a capillary
electrophoresis separation method for benazepril and other angiotensin-
converting enzyme inhibitors [44]. Because a free solution capillary
electrophoresis system did not achieve a complete separation of these
compounds in one run, the usefulness of alkylsulfonates as ion-pairing
agents was investigated. After preliminary investigation to determine the
experimental domain and the most important factors, a three-level full-
factorial design was applied to study the impact of pH and molarity of the
ion-pairing agent on the separation. Improved separations were obtained,
suggesting a favorable effect of ion-pairing interactions between analytes
and the additive. However, it remained impossible to separate all of these
in a single run, so a combination of two systems was still necessary to
enable selective identification of all structurally-related substances.
Benazepril is an ester prodrug, whose ester linkage can be cleaved
chemically or enzymatically by hydrolysis or proteins displaying an
esterase-like activity. It should be a good substrate for the porcine liver
esterase, but it can be resistant to acidic hydrolysis. The kinetics of the
acidic and enzymatically catalyzed hydrolysis of benazepril was studied
using a HPLC method [37]. The acid-hydrolysis was studied in 0.1 M HCl
at 50, 53, 58, and 63 C. The kinetics of the degradation (benazeprilat
being the major degradation product) followed first-order kinetics, with
reaction rate constants around 10 h1 and half-lives ranging from 420 to
1400 h. The activation energy was found to be 19.9 kcal/mol. The
enzymatic hydrolysis was carried out at 37 C, in phosphate buffer of pH
7.4. It was found that benazepril remained stable for at least 30 min.
6. ABSORPTION AND PHARMACOKINETICS
6.1 Absorption, metabolism and fate
Following oral administration, at least 37% of a dose of benazepril is
absorbed. Benazepril is almost completely metabolized in the liver to
benazeprilat. Peak plasma concentrations of benazeprilat following an
oral dose of benazepril have been achieved after 1–2 h in the fasting state,
or after 2–4 h in the non-fasting state. Both benazepril and benazeprilat
are found to be about 95% bound to plasma protein [45–48].
Activation of benazepril involves cleavage of the ethoxy carbonyl group
by hepatic esterase [49, 50]. After oral administration, peak plasma
concentrations of benazepril and benazeprilat occur within half an hour
and one to one and a half hours, respectively [49, 51]. Benazepril is
extensively metabolized with less than 1% of dose being excreted in the
urine as unchanged benazepril [49, 52]. The primary urinary metabolites
of benazepril were benazeprilat (accounting for 17% of the oral dose),
and glucuronide conjugates of benazepril and benazeprilat (accounting
for 4% and 8% of the dose).
6.2 Excretion
Benazeprilat, the active metabolite of benazepril, is excreted mainly in the
urine, with only about 11–12% being excreted in the bile. The effective
half-life for accumulation of benazeprilat is 10–11 h following adminis-
tration of multiple doses of benazepril. The elimination of benazeprilat is
slowed in renally impaired individuals. Small amounts of benazepril and
benazeprilat are distributed into breast milk [2].
The main route of elimination of benazeprilat is by way of the kidneys,
although biliary excretion may also occur [49,51]. No clinically significant
metabolic drug interactions with benzepril have yet been described [49].
The authors would like to thank Mr. Tanvir A. Butt for his secretarial
assistance in preparing this manuscript.
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Ciprofloxacin: Physical Profile
164 M.A. AL-OMAR
1. General Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164
1.1.1 Systematic chemical names. . . . . . . . . . . . 164
1.1.4 Synonyms. . . . . . . . . . . . . . . . . . . . . . . . 165
1.2.1 Empirical formula, molecular weight, CAS
number . . . . . . . . . . . . . . . . . . . . . . . . . 165
2. Physical Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . 165
2.1 Solution pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165
2.6.1 Ultraviolet spectroscopy. . . . . . . . . . . . . . 170
2.6.3 Nuclear magnetic resonance
spectrometry . . . . . . . . . . . . . . . . . . . . . . 172
3. Stability and Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . 177
1. GENERAL INFORMATION
1-Cyclopropyl-6-fluoro-1,4-dihydro-4-oxo-7-(1-piperazinyl)-3-
quinolinecarboxylic acid.
CIPROFLOXACIN: PHYSICAL PROFILE 165
Ciprofloxacin
1.1.3 Proprietary names [2, 3, 4]
Aceto, Baycip, Ciflox, Cifluran, Ciloxan, Ciplox, Ciprinol, Cipro,
Ciprobay, Ciproxan, Ciproxin, Flociprin, Septicide, Velmonit,
Xorpic.
1.1.4 Synonyms [3]
Bay-q-3939.
1.2.1 Empirical formula, molecular weight, CAS number [3, 4]
C17H18FN3O3 331.35 [0085721-33-1].
F COOH
C ¼ 61.62% H ¼ 5.48% F ¼ 5.73% N ¼ 12.68% O ¼ 14.49%.
1.4 Appearance [1, 3]
Ciprofloxacin is obtained as a light yellow crystalline powder.
2.1 Solution pH
A 2.5% solution in water has a pH of 3.0–4.5 [3].
Ciprofloxacin is practically insoluble in water, very slightly soluble in
dehydrated alcohol and in dichloromethane, and soluble in dilute acetic
acid [3].
2.3 Optical activity
Since ciprofloxacin has no centers of dissymmetry, it does not exhibit
optical activity.
2.4 X-Ray powder diffraction
The X-ray powder diffraction pattern of ciprofloxacin was obtained using
a Simons XRD-5000 diffractometer, which is shown in Figure 1. Table 1
contains a compilation of values for the observed scattering peaks (in
units of degrees 2-), the interplanar d-spacings (units of Å), and the
relative intensities associated with the powder pattern.
Scattering Angle
(degrees 2-θ)
Figure 1. X-ray powder diffraction pattern of ciprofloxacin.
Crystallographic Data from the X-Ray Powder Diffraction Pattern
of Ciprofloxacin
CIPROFLOXACIN: PHYSICAL PROFILE
(degrees 2h) (%) (degrees 2) (%)
8.185 10.7928 5.94 9.065 9.7475 21.43
19.37 4.5864 43.48 19.810 4.4779 23.85
(degrees 2) (%) (degrees 2) (%)
22.575 3.9354 5.10 23.192 3.8321 13.76
M.A. AL-OMAR
57.924 1.5907 1.70
Ciprofloxacin is found to melt in the range of about 318–320 C [2].
The differential scanning calorimetry (DSC) thermogram of ciprofloxacin
was obtained using a DuPont TA-9900 thermal analyzer interfaced with a
DuPont Data Unit. The thermogram shown in Figure 2 was obtained at
a heating rate of 10 C/min, and was run from 40 to 400 C. The sole
observed thermal event was the melting endotherm which was observed
at 322 C.
2.6.1 Ultraviolet spectroscopy
The UV spectrum of ciprofloxacin dissolved in methanol was recorded
using a Shimadzu ultraviolet–visible Spectrophotometer 1601 PC, and is
Figure 2. Differential scanning calorimetry thermogram of ciprofloxacin.
Figure 3. The ultraviolet absorption spectrum of ciprofloxacin in methanol.
shown in Figure 3. The spectrum of ciprofloxacin was found to exhibit
the following characteristics at the three observed maxima:
Wavelength A[1%, 1 cm] Molar absorptivity
maximum (nm) (L mol1 cm1)
320.6 1.95 129
280 5.76 382
The infrared absorption spectrum of ciprofloxacin was obtained in a KBr
pellet using a Perkin Elmer infrared spectrophotometer. The infrared
spectrum is shown in Figure 4, and the principal peaks were noted at
3490, 3320, 2930, 2840, 1696, 1605, 1480, 1435 cm1. The assignments for
the major infrared absorption bands are summarized in Table 2.
880 960 1040 1120 1200 1280 1360 1440 1520 1600 1680 1760 1840 1920 2000 2080 2160 2240
Peak Energy (cm−1)
Figure 4. The infrared absorption spectrum of ciprofloxacin obtained in a
KBr pellet.
The proton NMR spectrum of ciprofloxacin was obtained using a Bruker
Advance system, operating at 300, 400, and 500 MHz. The sample was
dissolved in D2O, and tetramethylsilane (TMS) was used as the internal
standard. The proton NMR spectrum is shown in Figure 5, and
assignments for the 1H-NMR resonance bands of ciprofloxacin are found
in Table 3.
The carbon-13 NMR spectrum of ciprofloxacin was obtained using a
Bruker Advance system operating at 75, 100, and 125 MHz. Standard
Bruker Software was used to obtain DEPT spectra. The sample was
standard. Assignments for the various carbons of ciprofloxacin are
shown in Table 4.
Assignments for the Infrared Absorption Bands of
3490 O–H stretch
3320 N–H stretch of piperazinyl moiety
2930 Aliphatic C–H stretch
2840 N–C stretch
1696 C¼O stretch of carboxyl group
1605 C¼O stretch of quinoline
1480, 1435 C–N stretch
Figure 5. The H-NMR spectrum of ciprofloxacin in D2O.
Assignments for the observed Resonance Bands in the
H-NMR Spectrum of Ciprofloxacin
1a 1b
HN 4' 2' 1c
5' 1'
6' N 8a N
5 4 3a
Proton Chemical shift Multiplicity Number of
atoms (ppm relative (s: singlet, proton
to TMS) d: doublets) atoms
H-2 8.63 s 1
H-5 7.46 d 1
H-20 ,60 or 30 ,50 3.66 d 2
H-1a 1.22 s 1
H-1b, 1c 1.47 d 2
The electron impact (EI) spectrum of ciprofloxacin is presented in
Figure 6, and was recorded using a Shimadzu PQ-5000 GC-MS
spectrometer. The spectrum shows a mass peak (M.) at m/z 332, and a
base peak at m/z 288 resulting from the loss of the group. All these and
Assignments for the Observed Resonance Bands in the C-NMR Spectrum of
Carbon Chemical shift Assignment at Carbon Chemical Assignment
atoms (ppm relative carbon number atoms shift (ppm at carbon
to TMS) relative number
C-1a, C-1b 36.90 2 C-5 111.26 1
C-1c 43.96 1 C-6 107.14 1
C-2 139.48 1 C-7 145.40 1
C-3 155.60 1 C-20 , C-60 43.96 2
C-3a 169.31 1 C-30 , C-50 47.05 2
C-4a 148.69 1 C-8a 152.26 1
Figure 6. Mass spectrum of ciprofloxacin.
other proposed fragmentation patterns of the drug are presented in
3. STABILITY AND STORAGE [3, 5]
Solutions of ciprofloxacin are light sensitive and should be protected
from light and freezing [3]. When the concentrate formulated for
intravenous injection, or the 1.2 g pharmacy bulk package, is diluted
with 5% dextrose injection or 0.9% sodium chloride injection to a final
concentration of 0.5–2 mg/mL, the resultant solution is stable for upto 14
days when stored at room temperature or when refrigerated at 2–8 C [5].
The commercially available injection for intravenous infusion that
contains 2 mg/mL in 5% dextrose is provided in a plastic container
fabricated from a specially formulated polyvinyl chloride (PVC).
Tammilehto et al. used thin-layer chromatography to study the
degradation of ciprofloxacin hydrochloride solutions after these were
irradiated by a high pressure mercury lamp [6].
Assignments for the Fragmentation Pattern Observed in
the Mass Spectrum of Ciprofloxacin
Mass number Relative Structural
(m/z) intensities assignment
332 45 C17H18FN3O3e þ þ mass peak (M.)
314 55 M–H2O
288 100 M–CO2
245 30 M–(CO2  C2H5N)
1. Remington's: Pharmaceutical Sciences, 20th edn., A.R. Gennaro,
ed., Mack Publishing Co., Pennsylvania, p. 1539 (2000).
2. The Merck Index, 12th edn., S. Budavari, ed., Merck and Co., NJ,
3. Martindale, The Complete Drug Reference, 33rd edn., S.C.
Sweetman, ed., The Pharmaceutical Press, Chicago, p. 182
4. Index Nominum 2000: International Drug Directory, 17th edn.,
Swiss Pharmaceutical Society, Medpharm Scientific Publishers,
Ontario, p. 239 (2000).
5. Drug Information, 95 edn., G.K. McEvoy, ed., American Society
of Health-System Pharmacists, p. 493 (1995).
6. S. Tammilehto, H. Salomies, and K. Torniainen, J. Planar.
Chromatogr. Mod. TLC, 7, 368 (1994).
Ciprofloxacin: Analytical Profile
180 M. A. AL-OMAR
1. Compendial Methods of Analysis . . . . . . . . . . . . . . . . . . . 180
1.1 Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . 180
1.2 Compendial analyses . . . . . . . . . . . . . . . . . . . . . . 181
1.2.1 Appearance of solution . . . . . . . . . . . . . . 181
1.2.2 Fluoroquinolonic acid content . . . . . . . . . 181
1.3 Related substances: . . . . . . . . . . . . . . . . . . . . . . . 181
1.4 Heavy metals . . . . . . . . . . . . . . . . . . . . . . . . . . . 183
1.5 Loss on drying . . . . . . . . . . . . . . . . . . . . . . . . . . 183
1.6 Sulfated ash . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183
1.7 Chlorine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183
1.8 Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184
1.9 Titrimetric assay method for bulk
drug substance . . . . . . . . . . . . . . . . . . . . ...... 184
1.10 Assay method for tablets . . . . . . . . . . . . . ...... 184
1.11 Assay method for intravenous infusion
formulations . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185
1.12 Impurities. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185
1.12.1 Chromatographic purity . . . . . . . . . . . . . 187
2. Reported Methods of Analysis . . . . . . . . . . . . . . . . . . . . . 188
2.1 Titration methods . . . . . . . . . . . . . . . . . . . . . . . . 188
2.3 Chemiluminescence methods . . . . . . . . . . . . . . . . 191
2.4 Electrochemical methods . . . . . . . . . . . . . . . . . . . 192
2.5 Thin-layer chromatography . . . . . . . . . . . . . . . . . 192
2.6 High performance liquid chromatographic
methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193
2.7 Gas chromatography . . . . . . . . . . . . . . . . . . . . . . 202
2.9 Immunoassay methods . . . . . . . . . . . . . . . . . . . . 203
1. COMPENDIAL METHODS OF ANALYSIS
The European Pharmacopoeia [1] recommends the use of infrared
absorption spectrophotometry for the identification of the pure drug
substance. The procedure entails an examination by infrared absorption
CIPROFLOXACIN: ANALYTICAL PROFILE 181
spectrophotometry, and a comparison with the spectrum obtained with
the ciprofloxacin reference standard.
1.2 Compendial analyses
1.2.1 Appearance of solution
Dissolve 0.25 g of the substance to be tested in 0.1 M hydrochloric acid,
and dilute to 20 mL with the same solvent. The solution is clear, and not
more intensely colored than the reference solution GY4.
1.2.2 Fluoroquinolonic acid content
Examine by thin-layer chromatography, using as the coating substance a
suitable silica gel with a fluorescent indicator having an optimal intensity
at 254 nm.
Test solution: Dissolve 50 mg of the substance to be examined in
dilute ammonia R1 and dilute to 5 mL with the same solvent.
Reference solution: Dissolve 10 mg of fluoroquinolonic acid CRS in
a mixture of 0.1 mL of dilute ammonia R1 and 90 mL of water R,
and dilute to 100 mL with water R. Dilute 2 mL of this solution to
10 mL with water R.
Apply 5 mL of each solution to the plate. At the bottom of a
chromatographic tank, place an evaporating dish containing 50 mL of
concentrated ammonia R. Close the tank and expose the plate to the
ammonia vapor for 15 min. Withdraw the plate, transfer to a chromato-
graphic tank, and develop over a path of 15 cm using a mobile phase
mixture consisting of 10 volumes of acetonitrile R, 20 volumes of
concentrated ammonia R, 40 volumes of methanol R, and 40 volumes of
methylene chloride R. Allow the plate to dry in air and examine under 254
nm ultraviolet light. Any spot corresponding to fluoroquinolonic acid in
the chromatogram obtained with the test solution is not more intense than
the spot in the chromatogram obtained with the reference solution (0.2%).
1.3 Related substances [1, 2]
Examine the substance in question by liquid chromatography, using the
following procedure:
Test solution (a): Add 0.2 mL of dilute phosphoric acid R to 25 mg
of the substance to be examined, and dilute to 50.0 mL with the
mobile phase. Treat in an ultrasonic bath until a clear solution is
obtained.
Test solution (b): Dilute 0.1 mL of test solution (a) to 50 mL with
the mobile phase.
Reference solution (a): Dissolve 2.5 mg of ciprofloxacin impurity B
CRS in the mobile phase and dilute to 50.0 mL with the mobile
phase. This solution is also used to prepare the reference solution
(d). Dilute 1.0 mL of this solution to 50 mL with mobile phase.
Reference solution (b): Dissolve 2.5 mg of ciprofloxacin impurity C
CRS in mobile phase and dilute to 50.0 mL with mobile phase. This
solution is also used to prepare reference solution (d). Dilute 1 mL
of this solution to 50 mL with mobile phase.
Reference solution (c): Dissolve 2.5 mg of ciprofloxacin impurity D
CRS in mobile phase and dilute to 50 mL with mobile phase. This
of this solution to 50.0 mL with mobile phase.
Reference solution (d): Mix 0.1 mL of test solution (a), 1 mL of
reference solution (a), 1.0 mL of reference solution (b), 1 mL
of reference solution (c) and dilute to 50 mL with mobile phase.
The chromatographic procedure may be carried out using:
– Chromatographic column: a stainless steel column, 0.25 m long
and 4.6 mm internal diameter, packed with base-deactivated
octadecylsilyl silica for chromatography R (5 mm particles). The
column is maintained at a temperature of 40 C.
– Mobile Phase: a mixture of 13 volumes of acetonitrile R and
87 volumes of 0.025 M phosphoric acid, previously adjusted to
pH 3.0 with triethylamine R. The flow rate is to be about
1.5 mL/min.
– Detector: a spectrophotometer set at a wavelength of 278 nm.
Inject separately 50 mL of test solution (b) and 50 mL of reference
solutions (a), (b), (c), and (d). When the chromatogram is recorded using
the conditions described above, the retention time of ciprofloxacin is
about 9 min. Adjust the sensitivity of the system so that the height of the
ciprofloxacin impurity C peak in the chromatogram obtained with
reference solution (d) is at least 40% of the full scale deflection of the
recorder. The test is not valid unless in the chromatogram obtained with
reference solution (d), the resolution between the peaks corresponding
to ciprofloxacin impurity B and ciprofloxacin impurity C is at least 1.3,
and the resolution between the peaks corresponding to ciprofloxacin
and ciprofloxacin impurity D is at least 3.
Inject separately 50 mL of test solution (a) and 50 mL of reference
solutions (b) and (c), and record the chromatograms for twice the
retention time of ciprofloxacin. In the chromatogram obtained with test
solution (a), the areas of the peaks corresponding to ciprofloxacin
impurity C and ciprofloxacin impurity D are not greater than the
corresponding peaks in the chromatogram obtained with reference
solutions (b) and (c) (i.e., not more than 0.2%). The areas of any other
secondary peaks are not greater than the peak in the chromatogram
obtained with reference solution (b) (i.e., not more than 0.2%). The sum
of the areas of all the peaks, apart from the principal peak, is not greater
than 2.5 times the area of the peak in the chromatogram obtained with
reference solution (b) (i.e., not more than 0.5%). Disregard any peak
whose area is less than 0.25 times the area of the peak in the
chromatogram obtained with reference solution (b).
1.4 Heavy metals [1]
Dissolve 0.5 g of the substance to be tested in dilute acetic acid R, dilute
to 30 mL with the same solvent, and then add 2 mL of water R. The
filtrate complies with limit test E for heavy metals (not more than
20 ppm). The comparison stand is prepared using 5 mL of lead standard
solution (2 ppm Pb) R.
1.5 Loss on drying [1]
The sample does not contain more than 1.0% total volatile content when
1.000 g is dried under vacuum at 120 C.
1.6 Sulfated ash [1]
The sample does not contain more than 0.1% sulfated ash, determined
on 1.0 g.
1.7 Chlorine
The turbidity exhibited by the test solution does not exceed that of the
standard solution (0.02%) [3].
1.8 Sulfate
The turbidity exhibited in the tube containing the test solution
does not exceed that of the tube containing the standard solution
(0.04%) [3].
1.9 Titrimetric assay method for bulk drug substance
Dissolve 0.300 g of the substance under test in 80 mL of anhydrous acetic
acid R. Titrate with 0.1 M perchloric acid, using potentiometric means
to determine the end-point. In the titration, each milliliter of 0.1 M
perchloric acid is equivalent to 33.14 mg of C17H18FN3O3.
1.10 Assay method for tablets [2]
The tablet assay is performed using a liquid chromatographic method,
which requires the following solutions. Weigh and powder 20 tablets. For
solution (1), add a quantity of powdered tablets containing the equivalent
of 2 g of ciprofloxacin to 750 mL of water, mix for 20 min, add sufficient
water to produce 1000 mL, and mix again. Centrifuge a portion of the
resulting suspension, and dilute the clear supernatant liquid with
sufficient water to produce a solution containing the equivalent of
0.05% w/v of ciprofloxacin. Solution (2) contains 0.058% w/v of
ciprofloxacin hydrochloride EPCRS in the mobile phase. Solution (3)
contains 0.025% w/v of ciprofloxacin impurity C EPCRS (the
ethylenediamine compound) in solution (2). For solution (4), dilute 1
volume of solution (3) to 100 volumes with the mobile phase.
The chromatographic procedure may be carried out using a stainless steel
column (25 cm  4.6 mm) packed with a stationary phase C (7 mm)
(Nucleosil C18 is suitable), whose temperature is to be maintained at
40 C. The mobile phase is eluted at a flow rate of 1.5 mL/min, and
consists of a mixture of 13 volumes of acetonitrile and 87 volumes of a
0.245% w/v solution of orthophosphoric acid (the pH of which has been
adjusted to 3.0 with triethylamine). Analytes are detected on the basis
of their UV absorption at 278 nm. The Assay is not valid unless in
the chromatogram obtained with solution (3), the resolution factor
between the peaks due to ciprofloxacin and ciprofloxacin impurity C is
at least 3.
Calculate the content of C17H18FN3O3 in the tested tablets using the
declared content of C17H19ClFN3O3 in ciprofloxacin hydrochloride
EPCRS. Each mg of C17H19ClFN3O3 is equivalent to 0.9010 mg of
C17H18FN3O3. The content of ciprofloxacin should be 95.0–105.0% of
the label claim.
1.11 Assay method for intravenous infusion formulations [2]
Ciprofloxacin intravenous infusion is a sterile solution of ciprofloxacin
lactate (prepared by the interaction of ciprofloxacin and lactic acid) in
sodium chloride intravenous infusion solution.
The assay of an IV formulation is performed using a liquid
chromatographic method, which requires the following solutions. For
solution (1), dilute a quantity of the intravenous infusion with sufficient
mobile phase to produce a solution containing the equivalent of 0.05%
w/v of ciprofloxacin. Solution (2) contains 0.058% w/v of ciprofloxacin
hydrochloride EPCRS in mobile phase. Solution (3) contains 0.025%
w/v of ciprofloxacin impurity C EPCRS (ethylenediamine compound)
in solution (2). For solution (4), dilute 1 volume of solution (3) to 100
volumes with the mobile phase. The chromatographic procedure may be
carried out using a stainless steel column (12.5 cm  4 mm) packed with
stationary phase C (5 mm) (Nucleosil C18 is suitable). The mobile
phase is eluted at a flow rate of 1.5 mL/min, and consists of a mixture
of 13 volumes of acetonitrile and 87 volumes of a 0.245% w/v solution
of orthophosphoric acid (the pH of which has been adjusted to 3.0 with
triethylamine). Analytes are detected on the basis of their UV
absorption at 278 nm. The assay is not valid unless in the
chromatogram obtained with solution (3), the resolution factor between
the peaks due to ciprofloxacin and the ciprofloxacin impurity C is
at least 1.5.
Calculate the content of C17H18FN3O3 in the intravenous infusion
formulation using the declared content of C17H19ClFN3O3 in ciproflox-
acin hydrochloride EPCRS. Each milligram of C17H19ClFN3O3 is
equivalent to 0.9010 mg of C17H18FN3O3. The ciprofloxacin content
should be 95.0–105.0% of the label claim.
1.12 Impurities [1, 3]
The European Pharmacopoeia [1] (Section 1.3) and the US
Pharmacopoeia [3] describe liquid chromatographic methods for the
determination of five related substances, denoted as substances A, B, C,
D, and E. The following structure is used to define substances A and C:
R N
Substance A: R ¼ Cl; systematic name is 7-chloro-1-cyclopropyl-
6-fluoro-4-oxo-1,4-dihydroquinoline-3-carboxylic
acid (fluoroquinolic acid).
Substance C: R ¼ NH–[CH2]2–NH2; systematic name is 7-[(2-
aminoethyl)amino]-1-cyclopropyl-6-fluoro-4-oxo-
1,4-dihydroquinoline-3-carboxylic acid (also
known as the ethylenediamine compound).
The following structure is used to define substances B and E:
R′ R
Substance B: R ¼ COOH and R0 ¼ H; systematic name is 1-cyclo-
propyl-4-oxo-7-(piperazin-1-yl)-1,4-dihydroquino-
line-3-carboxylic acid (also known as the desfluoro
compound).
Substance E: R ¼ H and R0 ¼ F; systematic name is 1-cyclopropyl-
6-fluoro-7-(piperazin-1-yl)quinolin-4(1H)-one (also
known as the decarboxylated compound).
Substance D: 7-Chloro-1-cyclopropyl-4-oxo-6-(piperazin-1-yl)-1,4-
dihydroquinoline-3-carboxylic acid.
Cl N
N COOH
1.12.1 Chromatographic purity [3]
The chromatographic purity of ciprofloxacin is determined using the
following general procedure:
Standard Preparation Solution: Transfer about 25 mg of USP
ciprofloxacin RS, accurately weighed, to a 50-mL volumetric flask.
Add 0.2 mL of 7% phosphoric acid to dissolve, dilute to volume
with mobile phase, and mix.
Resolution Solution: Dissolve a quantity of USP ciprofloxacin
ethylenediamine analog RS in the Standard Preparation Solution,
to obtain a solution containing about 0.5 mg/mL.
Assay Preparation Solution: Transfer about 25 mg of ciprofloxacin,
accurately weighed, to a 50-mL volumetric flask. Add 0.2 mL of 7%
phosphoric acid to dissolve, dilute to volume with mobile phase,
and mix.
Mobile Phase: Prepare a filtered and degassed 87:13 v/v mixture of
0.025 M phosphoric acid (previously adjusted to a pH of 3.0  0.1
with triethylamine) and acetonitrile.
Chromatographic System: The liquid chromatograph is equipped
with a 278-nm detector, and with a 4-mm  25-cm column that
contains packing L1. The column is maintained at a temperature of
40  1 C, and the flow rate is about 1.5 mL/min.
System Suitability: (A) Chromatograph the Resolution Solution,
and record the responses as directed under Procedure. The retention
time for ciprofloxacin is between 6.4 and 10.8 min. Relative
retention times are about 0.7 for ciprofloxacin ethylenediamine
analog, and 1.0 for ciprofloxacin. The resolution between the
ciprofloxacin ethylenediamine analog peak and the ciprofloxacin
peak is not less than 6. (B) Chromatograph the Standard
Preparation Solution. The column efficiency, determined from the
ciprofloxacin peak, is not less than 2500 theoretical plates, the
tailing factor for the ciprofloxacin peak is not more than 4.0, and
the relative standard deviation for replicate injections is not more
than 1.5%.
Procedure: Separately inject equal volumes (about 10 mL) of the
Standard Preparation and the Assay Preparation Solutions into
the chromatograph, record the chromatograms, and measure the
responses for the major peaks. Calculate the quantity (in mg) of
C17H18FN3O3 in the sample of ciprofloxacin taken using the
formula 50 C (rU/rS), in which C is the concentration (units of mg/
mL) of USP ciprofloxacin RS in the Standard Preparation Solution,
and rU and rS are the ciprofloxacin peak responses obtained from
the chromatograms of the Assay Preparation and Standard
Preparation Solutions, respectively.
Calculations: Calculate the percentage of each impurity peak in the
chromatogram obtained from the assay preparation taken using the
formula, 100ri/rt, in which ri is the response of each impurity peak,
and rt is the sum of the responses of all the peaks. Not more than
0.4% of ciprofloxacin ethylenediamine analog and not more than
0.2% of any other individual impurity peak is found, and the sum
of all impurity peaks is not more than 0.7%.
2. REPORTED METHODS OF ANALYSIS
Several methods were reported in the literature for the determination of
ciprofloxacin in pharmaceutical formulations and in biological fluids.
2.1 Titration methods
Four DNA topisomerase (gyrase) inhibitors, including ciprofloxacin
(alone or in mixture with other drugs), were determined by a titration
method [4]. Tablets containing ciprofloxacin were extracted three times
with 30 mL of methanol. A portion of the resulting solution was mixed
with 2 mL water, and that solution was titrated with aqueous
5 mM NaOH, 5 mM tetrabutyl-ammonium hydroxide, or 5 mM AgNO3
by adding 0.1 mL increments at 2 min intervals. The end-point was
detected using the solution conductance.
Zhang et al. determined ciprofloxacin lactate in injectable solutions by
acid dye biphasic titration [5]. A 200 mg dried sample was dissolved in
and diluted with water to a volume of 100 mL, and then a 2 mL aliquot of
the solution was mixed with 10 mL Na2HPO4/citric acid buffer (pH 7)
and 15 mL CHCl3. This solution was titrated with 0.5 mM-bromothymol
blue, with agitation, until a light-blue end-point appeared in the aqueous
phase. Recoveries were 99.9–102%, with a relative standard deviation
(n ¼ 4) of 0.22–0.24.
Kilic et al. selected five different antibiotics, including ciprofloxacin, and
reported a titration method based on the use of tetrabutylammonium
hydroxide as the titrant in a non-aqueous assay method [6]. Ciprofloxacin
was dissolved in water and diluted to 25 mL with pyridine. A 20 mL
portion of the resulting solution was titrated with 0.04 M tetrabutyl-
ammonium hydroxide in methanol/propan-2-ol at 25 C under nitrogen
in a jacketed glass reaction cell. End-point detection was performed with
an Orion 720A digital pH ion-meter, equipped with a combination pH
electrode that contained a saturated solution of anhydrous methanolic
KCl in the reference. The method was applied to pharmaceutical
preparations, and enabled recoveries in the range of 99.95–101.53%,
with a RSD of 0.5–1.09%.
Rizk et al. developed a sensitive and selective derivative UV-
spectrophotometric method for the determination of three fluoroqui-
nolone compounds, including ciprofloxacin, in formulations and spiked
biological fluids [7]. The method depends on the complexation of Cu(II)
with the studied compounds in an aqueous medium. A linear
correlation was established between the amplitude of the peak and
the drug concentration over the range of 35–120 ng/mL. The detection
limit was reported as 1.3 ng/mL. The method was used for the
determination of the ciprofloxacin bulk drug substance and its tablet
formulation, with an overall percentage recovery of 99.22  0.55 to
100.33  1.60.
Ciprofloxacin in both the tablets and blood serum was determined by
a fluorimetric method that was based on the intrinsic luminescence
of complexed terbium(III) [8]. A standard solution of ciprofloxacin
(0.1 mM) was mixed with 2 mL of a 3 mM Tb(III) solution and 0.1 M
sodium acetate/0.1 M acetic acid buffer (pH 6). The mixture was diluted
to 10 mL with water, and after 30 s the fluorescence intensity was
measured at a wavelength of 545 nm (using an excitation wavelength
of 325 nm). The calibration curve was linear from 13 ng/mL to 1 mg/mL
of ciprofloxacin, with a detection limit of 10 ng/mL. The recoveries were
91.3–106.5% in the tablets with an RSD (n ¼ 5) of 0.8%, and
93.9–105.2% in the blood serum.
Navalon et al. developed a method for the determination of trace
amounts of ciprofloxacin, based on solid-phase spectrofluorimetry [9].
The relative fluorescence intensity of ciprofloxacin fixed on Sephadex SP
C-25 gel was measured directly after packing the gel beads in a 1 mm
silica cell, using a solid-phase attachment. The wavelengths of excitation
and emission were 272 and 448 nm, respectively. The linear concentration
range of the compound was 0.3–10 ng/mL, with a RSD of 1.2% (for a
level of 4 ng/mL) and a detection limit of 0.1 ng/mL. This method was
used for the determination of ciprofloxacin in human urine and serum
samples with a recovery of 100% in all cases.
Amin described a simple and sensitive spectrophotometric method for the
determination of three gyrase inhibitors, including ciprofloxacin, in
pharmaceutical formulations [10]. This method is based on the formation
of an ion pair with Sudan III in 40% v/v aqueous acetone. The color of
the dye changes at 566 nm in the presence of ciprofloxacin. Beer's law is
obeyed over the range of 0.4–10.4 mg/mL, and the results showed good
recovery (100  1.7%) with a RSD of 1.08%.
An analytical procedure for the determination of ciprofloxacin in serum
without previous extraction has been developed by Djurdjevic et al. [11].
The determination was carried out at pH 3, using iron(III) nitrate as the
chromogenic agent with the addition of sodium dodecyl sulfate.
Absorbance values were measured at 430 nm. The linearity range was
between 0.5 and 20 mg/mL, with a detection limit of 0.2 mg/mL.
Five drugs used as antibacterial agents, including ciprofloxacin, were
measured spectrophotometrically by Avadhanulu et al. [12]. Cipro-
floxacin was determined in the tablets after extraction with aqueous
50% acetone on the basis of its absorbance at 524 nm. The recovery
and RSD for ciprofloxacin was 99.3–100.2% and 0.3–0.55%,
Abdel-Gawad et al. described a spectrophotometric method for the
determination of ciprofloxacin in its pure form and in its tablet
formulations that made use of charge-transfer complexation reactions
[13]. In this method, either 5 mM 2,3-dichloro-5,6-dicyano-p-benzoqui-
none (I), 5 mM 7,7,8,8-tetracyanoquinodimethane (II), or 5 mM
p-chlorail (III) was added to a methanolic solution of ciprofloxacin.
After 15 min at 25 C for (I), or at 60 C for (II) and (III), the solution was
diluted to 10 mL with acetonitrile, and the absorbance was measured at
460, 843, or 550 nm for (I), (II), and (III), respectively. Beer's law was
obeyed over the concentration range of 5–50 mg/mL of ciprofloxacin, with
a RSD of 1.4% (n ¼ 7).
The effect of pH on the fluorescence of ciprofloxacin and norfloxacin has
been studied, and the ground-state microscopic dissociation constants
were determined [14]. After their acidic solutions (pH 2–4) were allowed
to stand for 5 min at 25 C, the optimum wavelength for fluorescence
determinations was found to be 445 nm (excitation at 280 or 331 nm) of
these compounds.
Ciprofloxacin was determined in tablets and intravenous infusion
solutions by an oxidative spectrophotometric method [15]. A yellow–
orange complex was obtained upon reaction of ciprofloxacin with 5%
ammonium sulfamate, 5 N H2SO4, and 0.1 M cerium(IV) ammonium
sulfate. The complex was extracted into CHCl3, dried with anhydrous
Na2SO4, and measured at 345 nm within up to 4 h. Beer's law was obeyed
over the range of 12–120 mg/mL, and the molar absorptivity was found to
be 5090.
Aly et al. [16] reported a rapid and sensitive chemiluminescence (CL)
method for the determination of three fluoroquinolone derivatives,
including ciprofloxacin, in both pharmaceutical dosage forms and in
biological fluids. The method is based on the CL reaction of the drugs
with tris(2,20 -bipyridyl)ruthenium(II) and cerium(IV) in sulfuric acid
medium. The CL intensity was proportional to the concentration of
ciprofloxacin in solution over the range 0.05–6 mg/mL, and the limit of
detection was reported to be 26 nM.
A flow-injection CL method was developed by Rao et al. for the
determination of fluoroquinolones, including ciprofloxacin, based on the
CL reaction of sulfite with cerium(IV) [17]. The linear range for
ciprofloxacin-CL is 0.4–30 mg/mL, with a RSD of 2.1–2.6% (n ¼ 10).
Ciprofloxacin hydrochloride was determined in capsules and tablets using
5 mM Na2SO3 in a flow-injection chemiluminescence system [18]. The
system was eluted at a flow rate of 6 mL/min, with a carrier/reagent
stream consisting of 0.4 mM CeSO4  2(NH4)2SO4 containing 50 mM
H2SO4. The CL signal was measured in an on-line flow cell.
The calibration graph was linear over the range of 1–20 mg/L of
ciprofloxacin, with a detection limit of 0.27 mg/L and a method recovery
of 96–108% [18].
2.4 Electrochemical methods
The voltammetric behavior of the Ni(II) complexes of three 4-quinolone
antibacterial compounds in tablets, including ciprofloxacin, was studied
using direct current (DCt), differential pulse (DPP) and alternating
current polarography (ACt) [19]. A well-defined, cathodic wave was
detected at pH 5.5 for ciprofloxacin. The current–concentration relation-
ship was found to be linear over the range 2–6.4  105 M and
0.8–5.6  105 M for ciprofloxacin using DCt and DPP modes,
respectively. Limits of detection (S/N ¼ 2) for these species were found
to be about 2  107 M. The average percent recovery for ciprofloxacin
was 99.58  0.72 to 100.50  0.79, and 99.50  0.71 to 100.17  0.29,
when using DCt and DPP, respectively.
Oscillopolarography was used to determine the ciprofloxacin content in
tablets [20]. The method was performed with measurement of the second-
derivative peak height at 1.16 V. The calibration graph for the drug was
found to be linear from 0.985 to 19.7 mM, with recoveries of 95.8–101.4%.
Ciprofloxacin hydrochloride, in pharmaceutical raw material and
capsules, has been analyzed by oscillopolarography [21]. A sample that
was mixed with 2 mL of 0.5 M KH2PO4/KOH buffer (pH 6.9), and
diluted with water, yielded a reductive peak potential at 1.51 V vs. SCE.
The calibration graph was linear from 0.1 to 20 mM, with an average
recovery of 101.1% and a RSD of 1.14%.
2.5 Thin-layer chromatography
Mixtures of antibacterial agents (including ciprofloxacin hydrochloride)
together with p-aminophenol (internal standard) in 0.1 M HCl were
analyzed by thin-layer chromatography (TLC) [22]. The system used a
silica gel GF254 plate, and a mobile phase consisting of chloroform/
methanol/concentrated ammonium hydroxide (15:10:3). The chromato-
gram was examined under a UV lamp at 254 nm, and it was found that
all spots were cleanly separated.
Quinolones (including ciprofloxacin) have been shown to be separable by
TLC using thin layers of Diol-silica adsorbent, and solutions of di-(2-
ethylhexyl)-orthophosphoric acid (HDEHP, an ion-pairing reagent) in
polar solvents as the mobile phase [23]. Retention and selectivity in the
adsorption-ion-association system can be controlled by adjusting the
concentration of HDEHP (typically 5–10%), or by changing the polar
diluents.
Wang et al. reported the simultaneous determination of trace amounts
of ciprofloxacin, pefloxacin, and norfloxacin in serum and urine by
TLC-fluorescence spectrodensitometry [24]. The drug substances were
applied to the silica gel plates, and developed to 9 cm with CHCl3/
methanol/toluene/CH2Cl2/aqueous NH3 (27:46:17:5:5). Fluorescence
spots were observed upon excitation at 254 nm, while the fluorescence
spectrodensitometric measurements were performed at 400 nm (excita-
tion at 278 nm). The calibration graph was linear upto 75 ng/spot for
ciprofloxacin, with an RSD that was less than 8.6%. The recovery was
96–108%.
The degradation of ciprofloxacin hydrochloride solution after irradiation
by a high pressure mercury lamp was studied using TLC [25]. Samples of
solution were taken and analyzed on 10  10 cm silica gel 60F254
HPTLC plates, after applying samples 5 mm apart at opposite edges of
the plate and eluting in a horizontal HPTLC developing chamber. The
plate was developed from opposite edges with acetonitrile/10% ammonia
containing 0.3 M ammonium chloride (13:5). Plates were scanned in
absorbance/reflectance mode at 283 nm. The working range was 10–100
ng/spot, and the RSD values (n ¼ 11) were 2.4 and 1.5% for the 20 and
the 80 ng/spot, respectively.
2.6 High performance liquid chromatographic methods
Numerous high performance liquid chromatography (HPLC) methods
have been reported, and the salient features of these are summarized in
Reported HPLC Methods for Ciprofloxacin
No. Material Column Mobile phase Internal Flow rate Detection Ref.
standard wavelength
1 Raw Waters C18 (5 mm) Phosphate buffer p-Aminobenzoic 1 mL/min 277 nm 22
material 15 cm  4.6 mm 0.98 mg/mL acid
i.d. sodium heptane
M. A. AL-OMAR
sulfonate pH 2.4/
methanol (13:7)
2 Human LiChrospher 60 RP 5% Acetic acid/ Lomefloxacin NA 280 nm 26
plasma (5 mm) methanol/acetoni-
trile (18:1:1)
3 Influenza Purospher RP-18e Acetonitrile/water/ NA 0.6mL/min 280 nm 27
vaccine (5 mm) 12 cm  phosphoric acid
3.5 mm i.d. (85%) (60:340:1)
4 Ointment Water Nova-pak 50 mM citric acid/ NA 1 mL/min 277 nm 28
C18 (10 mm) acetonitrile
15 cm  3.9 mm (82:16) pH 3.5
5 Urine Micro Bondapak- Methanol/ethyl Lomefloxacin 0.7 mL/ 285 nm 29
NH2 (30 cm  3.9 acetate/water/ min
mm i.d.) acetic acid
(50:10:40:1)
CIPROFLOXACIN: ANALYTICAL PROFILE
6 Plasma Kromasil C8 (5 mm) Acetonitrile/20 NA 1 mL/min 278/418 30
and 15 cm  4.6 mm mM- KH2PO4 nm
Mueller i.d. (pH 3.5) (3:97)
-Hinton
7 Dosage Purosher-RP-18e Methanol/20 NA 0.8mL/min 220 nm 31
forms (12.5 cm  4 mm mM-H3PO4 (1:3)
i.d.)
8 Dosage LiChrospher 100 100 mM-tertrabu- NA 1 mL/min 444/491 32
forms RP-18 (5 mm) tylammonium nm
25 cm  4.6 mm hydroxide in ace-
i.d. tonitrile/water
(93:7)/25 mM-
H3PO4 in acetoni-
trile/water (93/7)
standard wave-
9 Plasma YMC pack A-3-2 5% Acetic acid/ Lomefloxacin 1 mL/min 280 nm 33
and (5 mm) 15 cm  acetonitrile/
urine 6 mm) methanol (9:5:5)
10 Human Lichrosorb RP-18 Acetonitrile/metha- Theophylline NA 275 nm 34
serum (5 mm) 25 cm  nol/0.4 M citric
4 mm i.d. acid (1:3:6)
11 Tablets Nacalai Cosmosil Methanolic 3 mM Piroxicam 0.8 mL/ 257 nm 35
and 5C18 MS SDS/40 mM- min
capsules (25 cm  4.6 mm phosphate buffer/
i.d.) acetonitrile
(5:11:4) pH 3.5
12 Human Novapak C18 car- Methanol/acetoni- Pipemidic acid 1 mL/min 278/450 36
aqueous tridge (4 mm) trile/0.4 M citric nm
humor 10 cm  8 mm i.d. acid (3:1:10)
13 Raw LiChrospher 100 0.1 M-tetrabutyl- NA NA 278 nm 37
material C18 (5 mm) ammonium
25 cm  4 mm i.d. bromide in 25
mM H3PO4 of
pH 3.89/acetoni-
trile (93:7)
14 Impuri- RP-C-18 (10 mm) Methanol/water/ NA 1 mL/min 254 nm 38
ties 30 cm  3.5 mm acetic acid
i.d. (840:159:1)
15 Dosage Shimpack CLC- Tetrabutylammo- NA 1 mL/min 280 nm 39
forms ODS (15 cm  nium hydroxide
6 mm i.d.) buffer/acetonitrile
16 Clinical Spheri-5-OD-5A 0.1 M-KH2PO4/ NA 1 mL/min 280 nm 40
speci- (25 cm  4.6 mm) acetonitrile (1:1)
17 Plasma, Micro Bondapak 0.1 M H3PO4-45% Difloxacin 1 mL/min 278/470 41
whole C18 (10 mm) KOH (pH 2.5) nm
blood 30 cm  3.9 mixed 3:1 with
and mm i.d.) acetonitrile
erythro-
18 Human MB C18 radial pak 0.1 M- Quinine 4 mL/min 277/453 42
plasma (10 mm) 10 cm  (NH4)2HPO4 (pH bisulfate nm
8 mm i.d. 2.5)/acetonitrile/
(80:13:7)
19 Tablets Micro Bondapak Acetonitrile/metha- Cephalothin 2 mL/min NA 43
C18 (10 mm) nol/acetic acid/10
30 cm  3.9 mM KH2PO4
mm i.d. (15:12:0.3:73)
20 Dosage Micro Bondapak Methanol/water/ Nalidixic acid 1.8mL/min 254 nm 44
forms C18 (10 mm) triethylamine
30 cm  3.9 (250:250:1) pH 2.4
mm i.d.
21 Serum Phenyl Spherisorb Water/acetonitrile/ NA 1 mL/min 178.6 nm 45
and (3 mm) 10 cm  methanol
pro- 4.6 mm i.d. (83:15:2)
22 Clinical Ultropac Acetonitrile/0.4 M- Pipemidic acid 1 mL/min 340 or 275 46
speci- LiChrosorb RP- citric acid (1:5) nm
mens 18 (10 mm)
25 cm  4 mm)
23 Serum, Micro-Pak MCH Acetonitrile/0.1 M- NA 0.8mL/min NA 47
urine 10 (10 mm) KH2PO4 (pH 2.5;
and 30 cm  4 mm i.d. 19:81)
24 Serum Micro Bondapak 14% Methanol/ 1-(4-fluorophe- 1.7mL/min 278/470 48
and C18 (10 mm) 5%acetonitrile/ nyl)-7-(4- nm
urine 30 cm  3.9 0.3%tetrabutyl- methyl pipera-
mm i.d. ammonium zin-1-yl)
hydroxide (pH 3)
25 Body Poly(styrene-divi- Acetonitrile/metha- NA 0.5mL/min 277 nm 49
fluid nylbenzene) nol/0.02 M
(5 mm) 15 cm  trichloroacetic
4.6 mm i.d. acid
26 Biologi- Spherisorb ODS II 0.025 M H3PO4 NA 2 mL/min 277/445 50
cal (5 mm) 25 cm  (pH 3)/acetoni- nm
fluids 4 mm trile (19:1)
27 Dosage LiChrospher C18 Water/acetonitrile/ NA 1.5 mL/ 280 nm 51
forms (5 mm) 25 cm  triethylamine min
and 4 mm i.d. (80:20:0.6) pH 3
bulk with orthophos-
drug phoric acid
28 Stability LiChrospher C18 Methanol/0.245% NA 1.5 mL/ 278 nm 52
in (5 mm) 25 cm  H3PO4 (3:22) pH min
tablets 4 mm i.d. 3. or aqueous
acetonitrile pH
29 Biologi- Inertsil OSD-2 50% Aqueous Nalidixic acid 0.8 mL/ 320/545 53
cal (5 mm) 25 cm  methanol of min nm
material 4.6 mm i.d. pH2.5
30 Human Nucleosil C18 50 mM KH2PO4 Eprofloxacin 2 mL/min 277 nm 54
serum (3 mm) 3 cm  (pH3)/water/0.1
4.6 mm i.d. M tetrabutylam-
monium bromide
(pH 3)/acetoni-
trile (12:6:1:1)
NA: Not available.
2.7 Gas chromatography
Gas chromatography was used to analyze the residual piperazine of
ciprofloxacin and norfloxacin in pharmaceutical preparations [55]. The
analyte was dissolved in water, and the solution analyzed by GC on
a fused-silica column (30 m  0.3 mm i.d.) coated with 5% cross-linked
ph-Me silicone (3 mm). The temperature program consisted of an increase
to 50 C, holding for 5 min, ramping to 180 C (ramp ¼ 15 C/min), and
finally holding for 10 min at 180 5 C/min. Nitrogen was used as the
carrier gas (flow rate of 1 mL/min), detection was effected using flame
ionization. Piperazine was determined in the two mixtures by mixing for
10 min with cyclohexane, filtering, partitioning the filtrate with water,
heating the aqueous phase at 45 C for 10 min, and then analyzing as
above. The calibration graph was linear for 0.4–10 ppm piperazine, with
a RSD (n ¼ 10) of 2.1% for 2 mg piperazine. The recovery from
ciprofloxacin was 98.1–98.4%.
Six quinolone antibiotics (including ciprofloxacin) were separated and
determined by CE on fused-silica capillaries (57 cm  75 mm i.d.; 50 cm to
detector) at 25 C, with injection on the anode side, an applied voltage of
10 kV, and detection at 280 nm [56]. The buffer was 100 mM HEPES/
acetonitrile (9:1). The calibration graphs were linear from 0.25 to
40 mg/mL and detection limits were approximately 0.25 ng/mL.
Sun and Wu described a capillary electrophoresis method for the
determination of seven fluoroquinolones, including ciprofloxacin, in
pharmaceutical formulations [57]. Pipemidic acid (10 mg) was added as
an internal standard to 25 mg active ingredient dissolved in 100 mL of
methanol. The clear solutions (4.1 nL) were hydrodynamically injected at
60 mbar for 3 s on fused silica capillaries (67 cm  50 mm i.d.; effective
length ¼ 52 cm). The compounds were well separated in less than 8.5 min
when using 65 mM sodium borate/35 mM sodium dihydrogen phos-
phate/60 mM sodium cholate of pH 7.3 in acetonitrile (72:28) as the
running buffer, an applied voltage of þ 27 kV, and detection at 275 nm.
The calibration graphs were linear from 25 to 300 mg/mL of the
fluoroquinolones, with recoveries within 95–105% of their label claims.
A mixture of four fluoroquinolones (including ciprofloxacin hydrochlo-
ride) was determined by high performance capillary electrophoresis using
caffeine as an internal standard [58]. A portion of the solution was
introduced by pressure-differential sampling at 10 cm for 10 s, whereupon
the drugs were separated and determined by high-performance CE. The
method used a fused-silica capillary column (60 cm  0.75 mm i.d.;
effective length 55 cm), operated at an applied voltage of 18 kV, with
50 mM Na2B4O7/NaH2PO4 buffer of pH 8.5 as the running buffer, and
detection at 214 nm. The calibration graph for ciprofloxacin was linear
from 20 to 100 mg/L, and the detection limits and RSD were 0.40–0.48
mg/L and less than 2%, respectively.
Ciprofloxacin and its metabolite desethyleneciprofloxacin were deter-
mined in human plasma using capillary electrophoresis in the presence of
N-(1-naphthyl)ethylenediamine dihdrochloride as the internal standard
[59]. The sample was injected hydrodynamically into the capillary (37 cm
 50 mm i.d.), and the running buffer was 0.1 M H3PO4/0.1 M H3BO3
adjusted to pH 2.3 with triethylamine. The applied voltage was 28 kV,
and laser-induced fluorescence detection at 450 nm (excitation at 325 nm)
was used. The limits of quantitation were 20 and 10 mg/L for cipro-
floxacin and its metabolite (desethyleneciprofloxacin), respectively, with a
RSD (n ¼ 6) being less than 9%.
Fourteen quinolone antibacterial compounds (including ciprofloxacin)
were separated by fused-silica column (59 cm  50 mm i.d.; 43 cm to
detector) at 30 kV in less than 8 min and monitored at 260 nm [60]. In this
method, a background electrolyte of pH 7.3 containing 32 mM borate,
39 mM cholate, 8 mM heptanesulfonate, and 18 mM phosphate (as the
sodium salts), modified with 28% acetonitrile, was used as running
buffer.
Ciprofloxacin was separated from its impurities and determined by free-
solution capillary electrophoresis in a fused-silica tube (72 cm  50 mm
i.d.) operated at 20 kV and 30 C with a 50 mM borate buffer acidified
with H3PO4 to pH 1.5 and detection at 272 nm [61]. The linear calibration
range was 5 mg/mL–0.75 mg/mL of ciprofloxacin, and the elution time
was 14.8 min. The limit of determination was 1.6 mMm and the RSD was
4.9% (n ¼ 10). Six impurities were detected.
2.9 Immunoassay methods
A high performance immunoaffinity chromatography (HPIAC) column,
containing covalently bound anti-sarafloxacin antibodies, was developed
to determined fluoroquinolones in bovine serum [62]. In this method,
the HPIAC column was used to capture ciprofloxacin with other
fluoroquinolones, while allowing the remainder of the serum components
to elute to waste. After binding to HPIAC column, the fluoroquinolones
were eluted directly onto a reversed-phase column for final separation of
the compounds prior to their fluorescence detection (observation at
444 nm after excitation at 280 nm). The recovery of the method exceeded
95%, and its RSD was less than 7%.
Snitkoff et al. reported a specific immunoassay for monitoring the levels
of ciprofloxacin in human samples [63]. Serum was incubated at 37 C
with shaking for 2 h, with rabbit anti-ciprofloxacin keyhole limpet
haemocyanin primary antibody in microtiter wells coated with cipro-
floxacin-BSA conjugate. Further incubation with horseradish peroxidase-
labeled goat anti-rabbit IgG was carried out. The samples were shaken in
0.1 M citric buffer (pH 4.5), plates were incubated for 10 min, and then
read at 450 nm. Calibration graphs were linear for 10 pg/mL to 10 ng/mL
of ciprofloxacin, and the within-day RSD was less than 10%.
Holtzapple et al. developed an immunoassay method for determination
of four fluoroquinolone compounds (including ciprofloxacin) in liver
extracts [64]. In this method, an immunoaffinity capture SPE column was
used, that contained anti-sarafloxacin antibodies covalently cross-linked
to protein G. After interfering liver matrix compounds had been washed
away, the bound ciprofloxacin was eluted directly onto the HPLC
column. The HPLC system used a 5 mm Inertsil phenyl column
(15 cm  4.6 mm i.d.), with 0.1 M-glycine hydrochloride/acetonitrile
(17:3) as the mobile phase (eluted at a rate of 0.7 mL/min). Fluorimetric
detection at 444 nm was used after excitation at 280 nm. The recovery
of ciprofloxacin ranged from 85.7 to 93.5%, and the detection limit was
0.47 ng/mL.
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Ciprofloxacin: Drug Metabolism and
Pharmacokinetic Profile
1. Uses, Applications, and Associated History . . . . . . . . . . . . . 210
2. Absorption and Bioavailability. . . . . . . . . . . . . . . . . . . . . . 211
3. Metabolism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211
4. Excretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212
1. USES, APPLICATIONS, AND ASSOCIATED HISTORY
Since 1980, many effective antimicrobial drugs of the synthetic
fluoroquinolone series have been developed for the treatment of bacterial
infections in humans. Ciprofloxacin is one of the most widely used drugs
of this group, and exhibits bactericidal effects by inhibition of DNA
gyrase [1]. The drug is structurally related to other quinolones, including
cinoxacin, enoxacin, lomefloxacin, nalidixic acid, norfloxacin, and
pefloxacin. This family of compounds contains a 4-quinolone nucleus
with a nitrogen at position-1, a carboxyl group at position-3, and
a ketone moiety at position-4. Ciprofloxacin is a fluoroquinolone
carboxylic acid, since it contains a fluorine atom at position-6, and
a carboxylic acid moiety at position-3 of the 4-quinolone nucleus.
Fluorinated quinolones have a wider spectrum of activity and more
potency as compared to non-fluorinated quinolone [2]. The piperazine
group at position-7 of the 4-quinolone nucleus is responsible for the
antipseudomonal activity of ciprofloxacin. The drug also contains a
cyclopropyl group at position-1, which enhances its antimicrobial
Ciprofloxacin is commercially available as the monohydrate phase of its
hydrochloride salt, and has been formulated for oral and ophthalmic
administration. The compound has also been developed as the lactate salt
for use in intravenous administration [3].
Ciprofloxacin is approved for use in the treatment of bone and joint
infections, infectious diarrhea caused by Shigella or Campylobacter,
lower respiratory tract infections, skin infections, and urinary tract
infections. It is the drug of choice for the treatment of infections caused
by Campylobacter jejuni. In addition, it has found off-label use as an
alternative drug for the treatment of gonorrhea, salmonella, and yersinia
DRUG METABOLISM AND PHARMACOKINETIC PROFILE 211
infections [3, 4]. In general, ciprofloxacin is active against susceptible
gram-negative and gram-positive aerobic bacteria, so therefore it should
not be used alone for mixed aerobic–anaerobic bacterial infections [4, 5].
2. ABSORPTION AND BIOAVAILABILITY
All formulations of ciprofloxacin are rapidly and well absorbed from the
GI tract following oral administration, and the compound undergoes
minimal first-pass metabolism [3]. The presence of food in the GI tract
decreases the rate, but not the extent, of drug absorption. Antacids
containing magnesium, aluminum, and/or calcium, decrease the oral
bioavailability of ciprofloxacin [6]. The oral bioavailability of ciproflox-
acin is 50–85%, and peak serum concentrations of the drug are generally
attained within 0.5–2.3 h. Peak serum concentrations, and areas under
serum concentration-time curves (AUC), are slightly higher in geriatric
patients than in younger adults. In adults who receive a single 200 mg
dose of ciprofloxacin by intravenous (IV) injection over 10 min, serum
concentrations of the drug immediately following the injection average
6.3–6.5 mg/mL, and serum concentrations 1 and 2 h later average 0.87
and 0.1 mg/mL, respectively [4].
Ciprofloxacin is widely distributed into body tissues and fluids following
oral and IV administration. Highest concentrations of the drug generally
are found in bile, lung, kidney, liver, gall bladder, uterus, seminal fluid,
prostatic tissue and fluid, tonsils, endometrium, fallopian tubes, and
ovaries [4, 7, 8]. It also distributed into bone, aqueous humor, sputum,
saliva, nasal secretions, skin, muscle, adipose tissue, and cartilage [4, 7, 9].
The apparent volume of distribution of ciprofloxacin is 2–3.5 L/kg, and
the apparent volume of distribution at steady state is 1.7–2.7 L/kg. Only
low concentrations of ciprofloxacin are distributed into cerebrospinal
fluid (CSF). Peak CSF concentrations may be 6–10% of peak serum
concentrations. The drug is moderately bound to serum protein
(16–43%), and crosses the placenta and is produced with the mammary
milk [6, 10].
3. METABOLISM
At least four active metabolites have been identified, namely desethylene-
ciprofloxacin, sulfo-ciprofloxacin, oxo-ciprofloxacin, and N-acetyl-
ciprofloxacin. Oxo-ciprofloxacin appears to be the major urinary
metabolite, and sulfo-ciprofloxacin the primary fecal metabolite [4, 11].
Carolyn et al. have reported two additional ultraviolet-absorbing
metabolites in urine specimens [12]. The drug is partially metabolized in
the liver by modification of the piperazinyl group. Oxo-ciprofloxacin and
N-acetyl-ciprofloxacin microbial activities are comparable to norfloxacin,
and desethylene-ciprofloxacin is comparable to nalidixic acid for certain
organisms. About 40–50% of an oral dose is excreted unchanged in the
urine, and 15% as metabolites. Upto 70% of a parental dose may be
excreted unchanged, and 10% as metabolites within 24 h [13, 14].
Ciprofloxacin and its metabolite desethylene-ciprofloxacin were deter-
N-(1-naphthyl) ethylenediamine dihydrochloride as an internal standard.
Krol et al. [15] have thoroughly investigated the biotransformation of
ciprofloxacin in body fluids using high performance liquid chromato-
graphy (HPLC) and proposed the sequence of metabolic pathways
illustrated in Figure 1.
4. EXCRETION
Ciprofloxacin is eliminated by renal and non-renal mechanisms. The
drug is partially metabolized in the liver by modification of the
piperazinyl group to atleast four metabolites. These metabolites, which
have been identified as desethylene-ciprofloxacin, sulfo-ciprofloxacin,
oxo-ciprofloxacin, and N-acetyl-ciprofloxacin, have microbiological
activities that are less than that of the parent drug, but may be similar
to or greater than that of some other quinolones [4, 8, 9].
Ciprofloxacin and its metabolites are excreted in urine by both
glomerular filtration and by tubular secretion. Following oral adminis-
tration of a single 250-, 500-, or 750-mg dose in adults with normal renal
function, 15–50% of the dose is excreted in urine as unchanged drug, and
10–15% as metabolites within 24 h. 20–40% of the dose is excreted in
feces as the unchanged drug and metabolites within 5 days [4, 5, 16, 17].
Most, but not all, of the unchanged ciprofloxacin in feces appears to
result from biliary excretion [4, 5].
Renal clearance of ciprofloxacin averages 300–479 mL/min in adults
with normal renal function, and the drug is 16–43% bound to serum
protein in vitro. It crosses the placenta and is distributed into the amniotic
fluid in humans. The usual human dosage has not revealed evidence of
harm to the fetus.
HN C N
H o
N N N
F COOH F COOH
Desethylene-ciprofloxacin N-Accetyl-ciprofloxacin
HO3S N HN
N N N N
Sulfo-ciprofloxacin Oxo-ciprofloxacin
Figure 1. Biotransformation of ciprofloxacin in body fluids, as proposed by
Krol et al. [15].
The drug is also distributed into human-milk [4, 5, 10]. In lactating
women who received 3 doses every 12 h of 750 mg of ciprofloxacin, the
concentration of the drug in milk that was obtained 2–4 h after a dose
averaged 2.26–3.79 mg/mL. Concentrations in milk were higher than
concomitant serum concentrations for up to 12 h after a dose [10]. The
plasma half-life is about 3.5–4.5 h, and there is an evidence of modest
accumulation [4].
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Dipyridamole: Comprehensive Profile
Department of Pharmaceutical Chemistry,
P.O. Box 2457, Riyadh-11451, Saudi Arabia
216 A. KHALIL, F. BELAL AND A.A. AL-BADR
1. Description . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217
1.2.1 Empirical formula, molecular weight,
CAS number. . . . . . . . . . . . . . . . . . . . . . 218
1.5 Uses and applications . . . . . . . . . . . . . . . . . . . . . 219
2. Methods of Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . 219
3.1 Ionization constant . . . . . . . . . . . . . . . . . . . . . . . 220
3.3 X-ray powder diffraction pattern . . . . . . . . . . . . . 221
3.5 Ultraviolet spectroscopy . . . . . . . . . . . . . . . . . . . 223
3.6 Vibrational spectroscopy . . . . . . . . . . . . . . . . . . . 224
3.7 Fluorescence spectrum. . . . . . . . . . . . . . . . . . . . . 225
3.8 Nuclear magnetic resonance spectrometry . . . . . . . 226
3.8.1 1H-NMR spectra. . . . . . . . . . . . . . . . . . . 226
3.8.2 13C-NMR spectra . . . . . . . . . . . . . . . . . . 226
4.1 Compendial tests. . . . . . . . . . . . . . . . . . . . . . . . . 230
4.1.1 USP 24 compendial tests . . . . . . . . . . . . . 230
4.1.2 British pharmacopoeia
compendial tests . . . . . . . . . . . . . . . .... 245
4.2 Methods of analysis reported
in the literature . . . . . . . . . . . . . . . . . . . . . . . . . . 250
4.2.1 Identification . . . . . . . . . . . . . . . . . . . . . 250
4.2.2 Titrimetric methods . . . . . . . . . . . . . . . . . 251
4.2.3 Potentiometric methods . . . . . . . . . . . . . . 251
4.2.4 Extraction-gravimetric method . . . . . . . . . 252
4.2.5 Conductimetric method . . . . . . . . . . . . . . 252
DIPYRIDAMOLE: COMPREHENSIVE PROFILE 217
4.2.6 Spectrophotometric methods . . . . . . . . . . 253
4.2.7 Electrochemical methods . . . . . . . . . . . . . 264
4.2.8 Mass spectrometry . . . . . . . . . . . . . . . . . 266
4.2.9 Chromatographic methods . . . . . . . . . . . . 267
5. Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276
6. Pharmacokinetics and Metabolism . . . . . . . . . . . . . . . . . . . 276
7. Acknowledgment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276
2,20 ,200 ,2000 -[(4,8-Di-1-piperidinyl-pyrimido-[5,4-d]-pyrimidine-2,6-
diyl)-dinitrilo]-tetrakisethanol.
2,6-Bis-(diethanolamino)-4,8-dipiperidinopyrimido-[5,4-d]-
pyrimidine.
2,20 ,200 ,2000 -[(4,8-Dipiperidinopyrimidino-[5,4-d]-pyrimidine-2,6-
diyl)-dinitrilo]-tetraethanol.
2,20 ,200 ,2000 -[(4,8-Di-piperidin-1-yl)-pyrimido-[5,4-d]-pyrimidine-
2,6-diyl]-dinitrilo]-tetraethanol.
Ethanol, 2,20 ,200 ,2000 -[(4,8-di-1-piperidinylpyrimido-[5,4-d]-
pyrimidine-2,6-diyl)-dinitrilo]-tetrakis.
2,6-Bis-[di-(2-hydroxyethyl)-amino]-4,8-dipieridinopyrimido-
[5,4-d]-pyrimidine.
2,6-Bis-[di-(2-hydroxyethyl)-amino]-4,8-dipieridinopyrimidino-
Dipyridamolum, Dipyridamole, Dipyridamol, Dipiridamol.
Agredamol, Angipec, Apo-Dipyridamole, Atrombin, Cardiwell,
Cleridium, Cordantin, Coribon, Coronair, Coronamole,
Coronarine, Corosan, Coroxin, Corantyl, Didamol, Diphar,
Dipiridamol, Dipramol, Diprimol, Dipyridamole, Dipyrida,
Dipyridan, Dipyrin, Drisentin, Functiocardon, Kardisentin,
Miosen, Natyl, Novo-Dipiradol, Novodil, Penselin, Perazodin,
Peridamol, Perkod, Persantin, Persantine, Persentic, Piroan,
Prandiol, Protangix, Proxicor, Rombosit, Santhimon, Stinocor,
Tinol, Trancocard, Trodamol, Tromboliz, Trombosentin,
Trombostaz, Vazodi.
C24H40N8O4, 504.63 [58322]
N N OH
HO N N
Carbon 57.12%
Hydrogen 7.99%
Nitrogen 22.21%
Oxygen 12.68%
Dipyridamole is obtained as deep yellow needles from ethyl acetate, or as
an intensely yellow crystalline powder. Its solutions are yellow, and show
strong blue-green fluorescence.
1.5 Uses and applications
Dipyridamole is an adenosine reuptake inhibitor and a phosphodiester-
ase inhibitor with antiplatelet and vasodilating activity. As a result, the
compound is therefore used in thrombo-imbolic disorders. Orally
administered dipyridamole is used in association with orally administered
anticoagulants for the prophylactic treatment of thromboimbolism
following cardiac valve replacement. It is also used as a coronary
vasodilator [5–8].
Dipyridamole exerts its antiplatelet action by several mechanisms [7].
One of these is through the inhibition of phosphodiesterase enzyme in
platelets, resulting in an increase in intraplatelet cyclic AMP and the
consequent potentiation of the platelet inhibiting actions of prostacyclin.
Another is the direct stimulation of the release of this eicosanoid by
vascular endothelium, and the third is the inhibition of cellular uptake
and metabolism of adenosine (thereby increasing its concentration at the
platelet vascular interface).
The antiplatelet/antithrombotic activity of dipyridamole has been
demonstrated in laboratory and in animal models, and has been shown
to inhibit platelet aggregation and vessel-wall thrombogenesis [9–11].
Dipyridamole has been given either alone or with aspirin in the
management of myocardial infarction and stroke. For the secondary
prevention of stroke or transient ischemic attack, the drug may be given
as a modified-release preparation in a dose of 200 mg twice daily.
Dipyridamole administered intravenously results in a marked coronary
vasodilation and is used in stress testing in patients with ischemic heart
disease [5].
2. METHODS OF PREPARATION
A general outline of the procedure for synthesizing dipyridamole is
shown in Scheme 1. Reaction of the pyrimidino pyrimidine-2,4,6,8-
tetraol (1) with a mixture of phosphorous oxychloride and phosphorous
pentachloride gives the tetrachloro derivative (2). The halogens at the
peri positions 4 and 8 are more reactive to substitution than are the
remaining halogen pairs 2 and 6, which are in effect the two positions of
the pyrimidines. Thus, reaction with piperidine at ambient temperature
gives the 4, 8 diamine (3). Subsequent reaction with bis-2-hydroxy
ethylamine under more strenuous conditions gives dipyridamole (4)
[12, 13].
OH Cl
N OH N Cl
N N Piperidine
POCl3/PCl5
N N − 2 HCl
HO N Cl N
CH2CH2OH
H N
N Cl CH2CH2OH
N − 2 HCl
Scheme 1. General outline of the procedure for synthesizing dipyridamole.
3.1 Ionization constant [2, 3]
Dipyridamole exhibits a pKa value of 6.4.
3.2 Solubility characteristics [2, 3]
Dipyridamole is slightly soluble in water, and soluble in dilute acid
having a pH of 3.3 or below. The bulk drug substance therefore dissolves
in dilute solutions of mineral acids. It is also very soluble in methanol,
ethanol, chloroform, but not too soluble in acetone, benzene and ethyl
acetate. Dipyridamole is practically insoluble in ether.
The X-ray powder diffraction pattern of dipyridamole was obtained
using a Simons XRD5000 diffractometer. The XRPD powder pattern
of the drug is shown in Figure 1, and a summary of the crystallographic
data deduced from the powder pattern is shown in Table 1.
−11.13
−4.277
Figure 1. X-ray powder diffraction pattern of dipyridamole.
The X-Ray Powder Diffraction Pattern of Dipyridamole
Scattering angle d-spacing Relative intensity
(degrees 2-) (Å) (100*I/IO)
7.171 12.3173 1.07
7.987 11.0606 100.00
8.851 9.9825 36.13
3.4.1 Melting behavior [2, 3]
Dipyridamole melts in the range of 164–167 C.
The differential scanning calorimetry characterization of dipyridamole
was performed using a Dupont DSC model TA 9900 compiler thermal
analyzer system. The DSC thermogram shown in Figure 2 was recorded
at a heating rate of 10 C/min (under a nitrogen flow) over a temperature
range of 50–450 C. The compound was found to melt at 169.32 C, with
an enthalpy of fusion equal to 4.400 J/g.
3.5 Ultraviolet spectroscopy
The ultraviolet (UV/VIS) absorption spectrum of dipyridamole was
recorded on a Shimadzu model 1604 PC UV/VIS spectrophotometer. The
spectra were recorded at a concentration of 10 mg/mL, with the drug
4.400J/g
Figure 2. Differential scanning calorimetry thermogram of dipyridamole.
Dipyridamole in ethanol(.001%)
292.0, 0.575
Figure 3. Ultraviolet absorption spectrum of dipyridamole.
being dissolved in ethanol, 0.1 M hydrochloric acid, and 0.1 M sodium
hydroxide. The spectrum of dipyridamole in ethanol is shown in Figure 3.
The values calculated for molar absorptivity and A1% 1cm at different
wavelengths in the three media are shown in Table 2.
Literature values reported for the drug dissolved in acidic methanol
were at a wavelength maxima of 230 nm and 285 nm A1% 1cm ¼ 650 a). In
aqueous alkali, the wavelength maximum is observed at 295 nm [2].
3.6 Vibrational spectroscopy
The infrared absorption spectrum of dipyridamole was obtained using a
Perkin-Elmer infrared spectrophotometer, with the drug being pressed in
a KBr pellet. The IR spectrum is shown in Figure 4, and the principal
lines are at 3460 (OH stretch), 2360, 2340, 1597 (C¼C stretch), 1479,
1397, 1337, 1107, 965, 772, and 594 cm1.
Literature values reported for the principal peaks (KBr pellet) are at
wavenumbers 1526, 1214, 1010, 1076, 1041, and 1251 cm1 [2].
Molar Absorptivity and A1%
1cm Values from the Ultraviolet
Absorption Spectrum of Dipyridamole in Different Media
Solvent max (nm) A1%
1cm Molar
(L/Mol/cm)
Ethanol 429 147 7.43  103
409.5 163 8.23  103
292 573 2.89  104
0.1 M NaOH 415.2 162 8.18  103
206.2 2429 1.22  105
0.1 M HCl 399.6 133 6.71  103
3.7 Fluorescence spectrum
The native fluorescence spectrum of dipyridamole was recorded in
ethanol (5 mg/mL) using a Kontron spectrofluorimeter, Model SFM 25 A,
equipped with a 150 W xenon-high pressure lamp and driven by a PC
Pentium-II computer. As shown in Figure 5, the excitation maximum was
found at 297 nm, and the emission maximum was located at 467 nm.
Figure 4. Infrared absorption spectrum of dipyridamole.
3.8 Nuclear magnetic resonance spectrometry
3.8.1 1H-NMR spectra
The 1H-NMR spectrum of dipyridamole was obtained using a Bruker
system operating at 300, 400, or 500 MHz. Standard Bruker software was
used to execute recording of DEPT, COSY, and HETCOR spectra. The
sample was dissolved in CDCl3, and all resonance bands were referenced
to tetramethylsilane (TMS) internal standard. The 1H-NMR spectra are
shown in Figures 6 and 7, while the assignments for the observed
resonance bands are given in Table 3 along with an appropriate atom
numbering system. The 1H-NMR spectrum was also obtained using
a JEOL instrument, using methanol-D3 as the solvent and TMS as the
reference standard. The spectra are shown in Figures 8 and 9, and
assignments for the observed bands are also shown in Table 3. The COSY
H-NMR spectrum is shown in Figure 10.
3.8.2 13C-NMR spectra
The 13C-NMR spectra of dipyridamole dissolved in CDCl3 was obtained
using a Bruker system operated at 75, 100, or 125 MHz. All resonance
Figure 5. The fluorescence absorption spectrum of dipyridamole (EX ¼ excitation
spectrum, and EM ¼ emission spectrum).
bands were referenced to tetramethylsilane (TMS) internal standard.
The 13C-NMR spectrum is shown in Figure 11, while the DEPT 135
C-NMR spectrum is shown in Figure 12. The HMQC NMR spectra are
shown in Figures 13 and 14, and the HMBC NMR spectrum is shown in
Figure 15. Assignments for the observed bands are given in Table 4
together with an appropriate atom numbering system.
The mass spectrum of dipyridamole was obtained using a Shimadzu PQ-
5000 mass spectrometer. The parent ion was collided with helium as the
Figure 6. The H-NMR spectrum of dipyridamole in CDCl3.
Figure 7. Expanded H-NMR spectrum of dipyridamole in CDCl3.
Assignments for the Resonance Bands in the 1H-NMR
Spectrum of Dipyridamole
N CH2CH2OH
3 N 1 N
5 4 CH2CH2OH
HOH2CH2C N
N 1 N 3
Chemical shift Number Multiplicity Assignment
(ppm, relative of protons (m ¼ (proton at
to TMS) multiplet) carbon #)
Bruker* Jeol*
1.64 1.68 12 M 7 and 8
3.68 3.30 4 M 5 (OH)
3.79, 3.8 3.76 8 M 6
3.89 4.08 8 M 4
*NMR Instruments
carrier gas. Figure 16 shows the detailed mass fragmentation pattern, and
Table 5 shows the proposed mass fragmentation pattern. The reported
values of the principal peaks are at m/z values of 504, 473, 429, 505, 221,
474, 84, and 430 [2].
Figure 8. The 1H-NMR spectrum of dipyridamole in deuterated methanol.
4.1 Compendial tests
4.1.1 USP 24 compendial tests [14]
4.1.1.1 Tests for dipyridamole bulk drug substance
Identification. When dipyridamole is tested according to General
Method h197 Ki, the infrared absorption spectrum of the tested
dipyridamole must be equivalent to that of the dipyridamole reference
Melting range. When dipyridamole is tested according to General
Method h741i, the melting range of the tested dipyridamole is between
Figure 9. Expanded 1H-NMR spectrum of dipyridamole in deuterated methanol.
162 C and 168 C, but the range between the beginning and end of the
melting does not exceed 2 C.
Loss on drying. When dipyridamole is dried according to General
Method h731i (i.e., 105 C for 3 h, it loses not more than 0.2% of its
weight.
Chloride content. Dissolve 500 mg of dipyridamole in 5 mL of alcohol
and 2 mL of 2 N nitric acid, and then add 1 mL of silver nitrate TS.
No turbidity or precipitate is produced.
Residue on ignition. When dipyridamole is tested according to
General Method h281i, it contains not more than 0.1% of non-ignitable
residue.
Figure 10. COSY 1H-NMR spectrum of dipyridamole in CDCl3.
Heavy metals. When dipyridamole is tested according to General
Method h231i (Method II), it contains not more than 0.001% heavy
metal content.
Organic volatile impurities. When dipyridamole is tested according to
General Method h467i (Method IV), it meets the requirements.
Chromatographic purity. The purity of dipyridamole is established using
a HPLC method, which is defined by the following characteristics:
Mobile Phase: Dissolve 250 mg of dibasic sodium phosphate in 250
mL of water and adjust with dilute phosphoric acid (1 in 3) to a
Figure 11. C-NMR spectrum of dipyridamole in CDCl3.
pH of 4.6. Add 750 mL of methanol, mix, filter through a 0.5 mm
membrane filter, and degas. Adjustments can be made to the
composition to enable System Suitability to be achieved.
with a 288 nm detector, and a 3.9 mm  30 cm column that contains
packing L1. The flow rate is about 1.5 mL/min.
Test Preparation A: Prepare a solution of dipyridamole in methanol
having a known concentration of 1 mg per mL.
Test Preparation B: Dilute 1 mL of Test preparation A with
methanol to 100 mL, and mix.
Procedure: Inject 10 mL of Test Preparation B into the chromato-
graph by means of a sampling valve, adjusting the operating
parameters so that the response of the main peak (retention time
Figure 12. DEPT-135 C-NMR spectrum of dipyridamole in CDCl3.
about 6.5 min) is about 5% full scale. Inject 10 mL of Test
Preparation A, and run the chromatograph for 10 min. The sum
of responses of all the secondary peaks obtained from Test
Preparation A is not greater than the response of the main peak
obtained from Test Preparation B (i.e., 1.0%).
Assay. Transfer about 450 mg of dipyridamole, accurately weighed, to
a 250 mL beaker, and dissolve in 50 mL of glacial acetic acid. Stir for
30 min, add 75 mL of acetone, and then stir for an additional 15 min.
Titrate with 0.1 N perchloric acid VS to a potentiometric end point
using a glass electrode and a silver–silver chloride reference electrode
system. Perform a blank titration, and make any necessary correction.
Each mL of 0.1 N perchloric acid is equivalent to 50.46 mg of
C24H40N8O4.
Figure 13. The HMQC NMR spectrum of dipyridamole in CDCl3.
4.1.1.2 Tests for dipyridamole in formulations
Identification. Triturate a quantity of finely powdered tablets, contain-
ing the equivalent of about 100 mg of dipyridamole, with 10 mL of 0.1 N
hydrochloric acid, and filter. Add 0.1 N sodium hydroxide to the filtrate
until the solution is basic and a precipitate forms. Heat the mixture on
a steam bath for 1 min, cool, filter, and dry the residue at 105 C for 1 h.
The residue so obtained responds to the identification tests for
dipyridamole.
Dissolution. Dissolution testing of dipyridamole tablets is to be
performed according to the directives of General Method h711i, with
the inclusion of the following method characteristics:
Medium: 900 mL of 0.01 N hydrochloric acid.
Apparatus: Method 2 (paddles), rotation speed of 50 rpm.
Figure 14. The HMQC-NMR spectrum of dipyridamole in CDCl3.
Procedure: Determine the amount of C24H40N8O4 dissolved by
employing UV absorption at the wavelength of maximum absor-
bance at about 282 nm on filtered portions of the solution under test,
suitably diluted with Dissolution Medium, if necessary. The com-
parison is to be made with a Standard solution having a known
concentration of USP Dipyridamole RS in the same medium.
Tolerances: Not less than 70% (Q) of the labeled amount of
C24H40N8O4 is dissolved in 30 min.
Content uniformity. The determination of Content Uniformity is made
following General Method h9057i. The test is conducted by transferring
Figure 15. The HMBC-NMR spectrum of dipyridamole in CDCl3.
1 tablet to a 100 mL volumetric flask, adding 50 mL of 1 N hydrochloride
acid, heating in a steam bath for 5 min, and shaking by mechanical
means for 30 min. The mixture is cooled to room temperature, diluted
with 1 N hydrochloric acid to volume, and mixed. The resulting
mixture is filtered (discarding the first 25 mL of the filtrate), and then
an accurately measured portion of the subsequent filtrate is diluted with 1
N hydrochloric acid to provide a solution containing about 10 mg/mL of
dipyridamole. Concomitantly, the absorbances of this solution and of
a solution of USP Dipyridamole RS in the same medium having a
known concentration of about 10 mg/mL in 1 cm cells at the wavelength
of maximum absorbance (about 282 nm) using 1 N hydrochloric acid
as the blank, are determined. The quantity, in units of mg, of
Assignments for the Resonance Bands in the C-NMR
N 4 5
Chemical shift Assignment of Number
(ppm relative carbon number of carbons
62.86 5 4
52.6 6 4
Figure 16. Mass spectrum of dipyridamole.
C24H40N8O4 in the tablet taken, is calculated by the formula:
TC AU
D AS
where T is the labeled quantity (in mg) of dipyridamole in the tablet, C is
the concentration (in mg/mL) of USP Dipyridamole RS in the Standard
solution, D is the concentration (in mg/mL) of dipyridamole in the
solution from the tablet based upon the labeled quantity per tablet and
the extent of dilution, and AU and AS are the absorbances of the solution
of the tablet and the Standard Solution, respectively.
Assay. The assay value of dipyridamole tablets is established using a
HPLC method, which is defined by the following characteristics:
Mobile phase: Dissolve 250 mg of dibasic sodium phosphate in
250 mL of water, and adjust with dilute phosphoric acid (1 in 3) to
Assignments for the Observed Mass Spectrum Fragments
of Dipyridamole
504 48% C24H40N8O4
N N(CH2CH2OH)2
(HOH2CH2C)2N N
487 4% C24H38N8O3
N N O
N OH
N N O H
385 3% C17H28N8O
296 8% C16H20N6
221 25% C11H17N4O
191 16% C10H15N4
177 8% C9H13N4
99 32% C4H6N2O
HN C N O
84 25% C5H10N
69 23% C3H5N2
H N C N
58 70% C3H8N H
44 100% C 2H 4O
H2C CH OH
43 60% C2H5N
H2C CH CH3
a pH of 4.6. Add 750 mL of methanol, mix, filter through a 0.5 mm
membrane filter, and degas. Make adjustment, if necessary.
Standard preparation: Using an accurately weighed quantity of USP
Dipyridamole RS, prepare a solution in Mobile phase having a
known concentration of about 15 mg/mL.
Assay preparation: Transfer not less than 20 tablets to a 1000 mL
volumetric flask, add 100 mL of water, and sonicate for 15 min.
Add about 750 mL of methanol, and shake by mechanical means
for 30 min. Dilute with methanol to volume, mix, and centrifuge.
Dilute an accurately measured volume (VS, in units of mL) of the
clear supernatant solution quantitatively with Mobile phase to
obtain a solution (VA, in units of mL) containing about 15 mg/mL of
Chromatographic system and system suitability: The liquid chro-
matograph system is equipped with a 288 nm detector and a
3.9 mm  30 cm column that contains packing L1. The flow rate is
about 1.5 mL/min. Chromatograph the standard preparation, and
record the peak responses as directed under Procedure. The column
efficiency determined from the analyte peak is not less than 1000
theoretical plates, the tailing factor for the analyte peak is not more
than 2.0, and the relative standard deviation for replicate injections
is not more than 2.0%.
Standard preparation and the Assay preparation into the chro-
matograph, record the chromatograms, and measure the responses
for the major peaks. Calculate the quantity (in mg) of C24H40N8O4
in the tablet taken by the formula:
VA rU
VS rS
in which C is the concentration (in mg/mL) of USP Dipyridamole
RS in the Standard preparation, VA is the volume (in mL) of the
assay preparation, VS is the volume (in mL) of supernatant solution
taken for the assay preparation, and rU and rS are the peak
responses obtained from the assay preparation and the Standard
preparation, respectively.
4.1.2 British pharmacopoeia compendial tests [4]
Identification.
Test 1: The melting point range of the tested substance is in the
range of 162 C–168 C.
Test 2: Dissolve 10 mg in a mixture of 1 volume of 0.1 M
hydrochloric acid and 9 volumes of methanol R, and dilute to
50 mL with the same mixture of solvents. Dilute 5 mL of
this solution to 100 mL with a mixture of 1 volume of 0.1
M hydrochloric acid and 9 volumes of methanol R. Examine
between 220 nm and 350 nm, where the solution will exhibit
absorption maxima at 232 nm and 284 nm. The ratio of absorbance
measured at the 284 nm maximum to that measured at the 232 nm
maximum is in the range of 1.25–1.45.
Test 3: Examine by infrared absorption spectrophotometry,
comparing with the spectrum obtained with dipyridamole CRS.
Examine the substances as pellets prepared using potassium
bromide R.
Test 4: Dissolve about 5 mg in a mixture of 0.1 mL of nitric
acid R and 2 mL of sulfuric acid R. An intense violet color is
produced.
Related substances. Examine the test article using a liquid chromato-
graphic system that meets the following requirements:
Test solution: Dissolve 10 mg in the mobile phase, and dilute to
20 mL with the mobile phase.
Reference solution (a): Dilute 1 mL of the Test Solution to 20 mL
with the mobile phase, and then dilute 1 mL of this solution to
Reference solution (b): Dissolve 10 mg of diltiazem hydrochloride
CRS in the mobile phase, and dilute to 10 mL with mobile
phase. Dilute 1 mL of this solution to 20 mL with Reference
Solution (a).
The chromatographic system is composed of the following:
1. Column: A stainless steel column 0.25 m long and 4.6 mm
in internal diameter, is packed with octylsilyl silica gel for
chromatography R (5 mm). The temperature of the column is
maintained at 30 C.
2. Mobile Phase: It is prepared by dissolving 0.504 g of potassium
dihydrogen phosphate R in 370 mL of water R, and adjusting
to pH 3 with phosphoric acid R. To this is added 80 mL of
acetonitrile R and 550 mL of methanol R. The mobile phase is
eluted at a flow rate of 1.3 mL/min.
3. Detector: A spectrophotometer set at an analyzing wavelength
of 290 nm.
Inject 20 mL of each solution and continue the chromatography in the test
solution for a length of time that is at least nine times the retention time
of dipyridamole. The test is not valid unless in the chromatogram
obtained with reference solution (b), the resolution between the peaks
corresponding respectively to diltiazem and dipyridamole is at least 2.
In the chromatogram obtained with the test solution, the area of any
peak (apart from the principal peak) is not greater than the area of the
peak in the chromatogram obtained with reference solution (a) (i.e.,
0.5%). The sum of the areas of all the peaks (apart from the principal
peak) is not greater than twice the area of the peak in the chromatogram
obtained with reference solution (a) (i.e., 1%). Disregard any peak with
an area less than 0.1 times that of the peak in the chromatogram obtained
with reference solution (a).
Chloride. To 0.250 g of the substance under test, add 10 mL of water R
and shake strongly. Filter, rinse the filtrate with 5 mL of water R, and
dilute to 15 mL with water R. The combined filtrates comply with the
limit test for chloride (not more than 200 ppm).
Loss on drying. After drying 1.000 g in an oven at 100 C–105 C, the
substance loses not more than 0.5% of volatile substances.
Sulfated ash. After testing 1 g, the substance contains not more than
0.1% sulfated ash.
Assay. Dissolve 0.400 g in 70 mL of methanol R, and titrate with 0.1 M
perchloricacid,determiningtheendpointpotentiometrically. Eachmilliliter
of 0.1 M perchloric acid is equivalent to 50.46 mg of C24H40N8O4.
Storage. Store in a well-closed container, protected from light.
Impurities. 2,20 -[[4,6,8-tri(piperidin-1-yl)-pyrimido-[5,4-d]-pyrimidin-2-
yl]nitrilo] diethanol.
This impurity has the following structure:
2,20 ,200 ,2000 ,20000 ,200000 -[[8-(Piperidin-1-yl)-pyrimido-[5,4-d]-pyrimidine-2,4,6-
triyl]-trinitrilo]-hexaethanol. This impurity has the following structure:
HO OH
2,20 -[[2-Chloro-4,8-di-(piperidin-1-yl)-pyrimido-[5,4-d]-pyrimidin-6-yl]
nitrilo] diethanol. This impurity has the following structure:
Test 1: Shake a quantity of the powdered tablets containing 50 mg
of dipyridamole with 20 mL of chloroform, filter, and evaporate to
dryness. The infrared absorption spectrum of the residue (see
Appendix IIA of the BP) is concordant with the reference spectrum
of dipyridamole (RS 108).
Test 2: To a quantity of powdered tablets containing 10 mg of
dipyridamole, add 50 mL of methanol, warm slightly, shake for 15
min, and allow to cool. Add sufficient methanol to produce 100 mL,
and filter. To 10 mL of the filtrate, add 1 mL of 1 M hydrochloric
acid and sufficient methanol to produce 100 mL. The light
absorption of the resulting solution (see Appendix II B of the BP)
in the range of 220–450 nm exhibits three maxima, namely 230, 285,
and 405 nm.
Related substances. Carry out the method for liquid chromatography
(see Appendix III D of the BP), using the following solutions:
Solution (1): Shake a quantity of powdered tablets containing
50 mg of dipyridamole with 100 mL of mobile phase for 15 min,
filter (Whatman GF/C filter is suitable), and use the filtrate.
Solution (2): Dilute 1 volume of Solution (1) to 200 volumes with
mobile phase.
Solution (3): Dilute 1 volume of a 0.1% w/v solution of diltiazem
hydrochloride EPCRS to 20 mL with Solution (2).
Chromatographic System: It consists of the following:
Column: A stainless steel column (25 cm  4.6 mm), packed
with octylsilyl silica gel for chromatography (5 mm)
(Lichrosorb RP8 5 m is suitable). The column is maintained
at 30 C.
Mobile Phase: Dissolve 0.504 g of potassium dihydrogen
orthophosphate in 370 mL of water and adjust the pH to 3
with orthophosphoric acid. To this, add 80 mL of acetonitrile
and 550 mL of methanol. The mobile phase is eluted at a flow
rate of 1.3 mL/min.
Detector: UV absorbance at a wavelength of 290 nm.
Procedure: Inject 20 mL of each solution. Continue the chromato-
graphy of Solution (1) for at least nine times the retention time of
dipyridamole. This test is not valid unless in the chromatogram
obtained with Solution (3), the resolution factor between the peaks
due to diltiazem and dipyridamole is at least 2.
In the chromatogram obtained with Solution (1), the area of any
secondary peak is not greater than the area of the principal peak
in the chromatogram obtained with Solution (2) (i.e., 0.5%). In
addition, the sum of the areas of any such peaks is not greater than
twice the area of the peak in the chromatogram obtained with
Solution (2) (i.e., 1%). Disregard any peaks with an area less than
0.1 times that of the peak in the chromatogram obtained with
Solution (2) (0.05%).
Assay. To 10 whole tablets, add 300 mL of 1 M hydrochloric acid, heat
at 40 C for 20 min with shaking, allow to cool, and add sufficient 1 M
hydrochloric acid to produce 500 mL. Filter and dilute, if necessary, with
1 M hydrochloric acid to produce a solution containing 0.05% w/v of
Dipyridamole. Dilute 1 volume of this solution to 50 volumes with water,
and measure the absorbance of the resulting solution at the maximum
283 nm (see Appendix II B of the BP). Calculate the content of
C24H40N8O4 from the absorbance of a solution obtained by diluting
1 volume of a 0.05% w/v solution of dipyridamole BPCRS in 1 M
hydrochloric acid to 50 volumes with water.
4.2 Methods of analysis reported in the literature
4.2.1 Identification
The following two color tests have been recommended for the
identification of dipyridamole [2]:
Mandelin's Test: Prepare the reagent by dissolving 0.5 g of
ammonium vanadate in 1.5 mL of water and diluting to 100 mL
with sulfuric acid. When dipyridamole is reacted with this reagent,
the resulting solution has a violet color.
Marquis' Test: Prepare the reagent by mixing 1 volume of
formaldehyde with 9 volumes of sulfuric acid. When dipyridamole
is reacted with this reagent, the resulting solution has an orange
4.2.2 Titrimetric methods
Ganescu et al. used a volumetric and a spectrophotometric method for
the determination of dipyridamole with Reinecke salt analogues [15].
Eight complex salts of dipyridamole, namely dipyridamole H2[Cr(SCN)4
(amine)2]2 (amine ¼ aniline or other aromatic heterocyclic amines), were
prepared and characterized. Dipyridamole in these salts was determined
through oxidation (using KMnO4, KBrO3 or KIO3), through spectro-
photometry, or through complexation (using EDTA). The results were
subjected to statistical analysis.
Ganescu et al. used thiocyanato–Cr(III)-complexes in the titrimetric and
spectrophotometric determination of dipyridamole [16]. For titrimetric
analysis of the drug (1.36–27.2 mg), sample solutions (25–50 mL) were
acidified with hydrochloric acid and treated with excess of a 3% solution
of ammonium dianilinetetrakis (thiocyanato) chromium (III) in aqueous
25% ethanol. The red-violet precipitate was collected and washed,
decomposed with NaOH, and the mixture again acidified. The liberated
thiocyanate was titrated with potassium permanganate, potassium
bromate, or potassium iodate.
Qi et al. studied the effect of water content in perchloric acid on the non-
aqueous potentiometric titration of dipyridamole and other nitrogen-
containing compounds [17]. The USP non-aqueous titrimetric method
specifies a 0.1 N perchloric acid solution, with a water content in the
range of 0.02–0.05%. Experiments were carried out on nitrogen-
containing compounds to see if this water range could be extended.
When using an increased water content, the results showed a maximum
difference of 0.7% when compared with those obtained using the
standard method. Relative standard deviations were less than 0.4%. The
study authors recommended that perchloric acid with a water content in
the range of 0.02–0.5% should be permitted for the titration of nitrogen-
containing compounds.
4.2.3 Potentiometric methods
Hassan and Rizk developed potentiometric dipyridamole sensors based
on lipophilic ion-pair complexes and native ionic polymer membranes
[18]. The sensors are based on the use of the ion-association complexes
of dipyridamole cation with tetraphenylborate and reineckate counter-
anions as ion-exchange sites in plasticized PVC matrices. A plasticized
native polymer (carboxylated polyvinyl chloride) can also be used. These
sensors exhibit linear and near-Nernestian responses for 10 mM–1 mM
dipyridamole, with a cationic slope of 53.4–55.1 mV/decade, and a
working pH range of 2–5. The sensors proved to be useful for deter-
mining dipyridamole in various dosage forms. The average recovery was
reported to be 98.7%, with a standard deviation of 0.6%.
Issa et al. described a potentiometric titration method based on the use of
ion-selective electrodes [19]. The electrode response was Nernestian over
the range of 6.3 mM–0.1 M, when operated in the pH range of 2–7 and at a
temperature range of 2570 C. The response time was reported to
be 20–30 s. The recovery was 97.37–103.35%, with relative standard
deviation of 0.06–0.5%.
4.2.4 Extraction-gravimetric method
Liu et al. determined dipyridamole in tablets using a modified extraction-
gravimetric method combined with a surface acoustic wave resonator
sensor [20]. The powdered sample (200 mg) was mixed with 0.02 M
hydrochloric acid and filtered. A portion of the filtrate was mixed with
0.2 M sodium hydroxide, shaken with chloroform, and a portion (1 mL)
of the organic layer was then applied to a surface acoustic wave resonator
sensor. After the chloroform was evaporated by a nitrogen flow,
frequency shift responses were measured after 3 min. The device com-
prised 62 MHz one-port resonators fabricated on Y-cut 2-propagation
LiNbO3 crystals with aluminum metallization, which were coated with
polyvinylidene fluoride. The sensor was mounted in a measuring chamber
and connected to an oscillator. The calibration graph was linear for
0.065–1.12 mg dipyridamole, and the relative standard deviation (n ¼ 5)
for 0.48 mg was 3.2%. The detection limit was 38 ng, and the mean
recovery was 99.8%.
4.2.5 Conductimetric method
Bahbouh et al. used a conductimetric method for the analysis of
dipyridamole [21]. Ground tablets containing the drug were dissolved in
100 mL of 67% ethanol, and a portion was transferred to a titration cell
equipped with a conductivity cell. The sample solution was diluted to
50 mL with water and titrated with 5 mM phosphotungstic acid. After
each addition of titrant, conductivity measurements were recorded after
a 2 min interval and plotted against the amount of titrant added. The
end point was determined at the curve break. Results obtained by this
method were in good agreement with those obtained by the official
pharmacopoeial methods.
4.2.6 Spectrophotometric methods
4.2.6.1 Infrared spectrometry
Huvenne and Lacroix described a mathematical procedure for the
correlation of band intensities of the Fourier-transform infrared
absorption spectra with those of the corresponding infrared transmission
spectra of compounds in KBr discs [22]. The procedure was applied to
spectra of flunitrazepam, dipyridamole, and lactose that were obtained
through the use of a Nicolet 7199 B FTIR spectrometer with photo-
acoustic detection. When the photoacoustic spectrum of a plant charcoal
was used to correct the spectra for inequalities in the incident light flux
before applying the procedure, the correlated band intensities were
generally consistent with those obtained using infrared transmission
spectra. The procedure may be useful for the direct identification of the
Umapathi et al. determined aspirin and dipyridamole simultaneously in
bulk form, and in dosage forms, by an infrared spectrophotometric
method [23]. Accurately weighed quantities of the drug (250 mg, 375 mg,
500 mg, 625 mg, 750 mg, 875 mg, and 1000 mg) were transferred quan-
titatively to volumetric flasks, dissolved in and diluted to 25 mL with
chloroform. This procedure yielded solutions containing 10–40 mg/mL
and 10–60 mg/mL of aspirin and dipyridamole, respectively. Two other
mixed standard solutions of aspirin and dipyridamole were pre-
pared containing varying concentrations (10–40 mg/mL) of aspirin and
powdered tablets equivalent to 600 mg dipyridamole (aspirin) and 480,
800 or 320 mg aspirin (dipyridamole) dissolved in and diluted to 25 mL
with chloroform. The solutions were filtered, and the infrared spectra
recorded using a calibrated sodium chloride cavity cell (path length
0.5 mm) against a chloroform reference. The absorbance at 920 and
2935 cm1 for aspirin and dipyridamole, respectively, was used for
quantitation.
4.2.6.2 Ultraviolet spectrometry
El-Yazbi et al. reported an application of a derivative-differential
ultraviolet spectrophotometric method for the determination of oxaze-
pam or phenobarbitone in the presence of dipyridamole [24]. Tablets
containing the drugs were powdered and dissolved in ethanol. For
solutions of oxazepam and dipyridamole, two portions of each were
diluted with 0.1 N sulfuric acid and 0.05 M sodium borate, and subjected
to differential spectrophotometry with measurements being made at
283, 292, 298 and 282, and 307, and 296 nm. First derivative (D1)
and second derivative (D2) determinations were made. For solutions of
dipyridamole and phenobarbitone, two portions of each were diluted
with 0.1 N sulfuric acid and 0.1 N sodium hydroxide, and differential
measurement were made at 253 and 309, 238 and 310, and 244 and 315
nm for D1 and D2, respectively. The calibration graphs were linear
over the range of 0.2–1.6 mg %, the recoveries were similar (close to
100%), and the coefficients of variation were in the range of 0.8–1.2%.
Hu et al. described an ultraviolet spectrophotometric method for the
analysis of dipyridamole [25]. A sample (50 mg) of the powdered tablet
was dissolved and diluted to 100 mL with 0.01 M hydrochloric acid.
A 2 mL portion of the solution was diluted to 100 mL with 0.01 M
hydrochloric acid, and the absorbance of the final solution was measured
at 283 nm (or at 403 nm for pure dipyridamole). The calibration graph
was linear for upto 12 or 60 mg/mL, and suitable for high-content
determinations. Results were in good agreement with those obtained by
titration with bromate.
Barary et al. used a derivative spectrophotometric method for the
determination of dipyridamole and carbocromen hydrochloride in the
presence of their oxidative degradation products [26]. Dipyridamole
in 0.1 M hydrochloric acid was determined by first derivative
(D1) spectrophotometry at 240 and 260 nm, and by second derivative
(D2) spectrophotometry at 246 and 268 nm. For both drugs,
0.1 M-hydrochloric acid was used as a blank. Graphs of D1 and D2
were linear for 4–12 mg/mL of dipyridamole. The technique effectively
corrects for the presence of degradation products. This method was used
successfully to determine dipyridamole in persantin tablets.
Khashaba et al. descried a first derivative spectrophotometric method for
the analysis of dipyridamole in the presence of aspirin in a ratio of 1:4.4,
as well as for buphenine hydrochloride in the presence of aescin in a ratio
of 1:20 [27]. The method depends on the measurement of the first
derivative amplitude (peak to zero) at 312 nm and at 250 nm for minor
components dipyridamole and buphenine hydrochloride, respectively.
No interference was observed from the other combined drugs (major
components) at the selected wavelength. The method response obeys
Beer's law in the concentration range of 2–18 mg/mL for dipyridamole,
and 3–38 mg/mL for buphenine hydrochloride. Laboratory synthetic
mixtures of the combined drugs in different ratios were analyzed showing
good and reasonable recoveries. For dipyridamole the reported
recovery range was 97.11  0.09 to 100.17  0.47, and for buphenine
hydrochloride the reported range was 98.23  0.89 to 99.53  0.31.
The method was also used for the assay of both drugs in single
pharmaceutical dosage forms. Small relative standard deviation values
(i.e., less than 2%) were noted, and the results obtained were found to be
in good agreement with those obtained using official methods.
Korany and Haller described a spectrophotometric method for the
determination of dipyridamole and oxazepam in two component
mixtures [28]. The pj method was applied to the simultaneous
determination of the two drugs in two component mixture. Authentic
mixtures and tablets containing the two drugs were assayed by measuring
the absorbances in 0.1 N sulfuric acid and 0.05 M borax solutions, and
then calculating the p2 at 2 wavelength sets {272312 nm (at 8 nm
intervals) and 288328 nm (at 8 nm intervals)}. The concentration is then
determined by solving a pair of simultaneous equations. The results
obtained were reasonably reproducible, exhibiting a coefficient of
variation that was less than 2%. The method was compared with a A
modification of Vierordt's method of two-component analysis. The npj
method was found to yield more accurate and precise results.
4.2.6.3 Colorimetric methods
Sane et al. used a spectrophotometric method for the determination of
dipyridamole and methdilazine hydrochloride from pharmaceutical
formulations [29]. The method uses a simple technique of ion pair
extraction of the two drugs with acid dyes like solochrome cyanine
(SCR), solochrome dark blue (SDB), solochrome black T (SBT), and
chromotrope 2B (C2B) in the presence of an acidic buffer. The colored
complexes of the drugs are extracted into chloroform, and their
absorbances measured at the wavelength of maximum absorption. The
drug was detected at 460, 520, and 520 nm when using SCR, SDB, and
SBT dyes, respectively. The percentage recovery of the SCR, SDB,
and SBT dye products from tablet formulations were 100.42, 99.55, and
98.98%, respectively, and the corresponding standard deviations were
0.122, 0.171, and 0.147. Beer's law was obeyed over the respective ranges
of 4–28, 4–24, and 4–32 mg/mL and the respective coefficients of variation
were 0.95%, 1.18%, and 1.17%.
Mahrous et al. reported the use a spectrophotometric method for the
determination of dipyridamole and other cardiovascular drugs using
p-chloranilic acid [30]. Tablets, or aqueous solutions containing salts of
dipyridamole and the other drugs, were rendered alkaline with aqueous
ammonia and extracted with chloroform (4  20 mL). The extract was
diluted to 100 mL with chloroform, a portion (0.4 to 2 mL) was
evaporated to dryness, and the residue was dissolved in 2 mL of
acetonitrile. 0.2% chloranilic acid in acetonitrile (1 mL) was added,
and the mixture was diluted to 5 mL with acetonitrile. The absorbance
was measured at 522 nm against a reagent blank. Recoveries were in the
range 99–102%, and Beer's law held from 0.1–0.5 mg/mL for
Issa et al. used 2,3-dichloro-5,6-dicyano-p-benzoquinone for the spectro-
photometric determination of dipyridamole and other cardiovascular
drugs [31]. Aqueous sample solutions are extracted with chloroform,
portions of the organic solution are evaporated, and the residues
dissolved in 2 mL of acetonitrile. After that, 2 mL of the cited reagent
(0.2% solution in acetonitrile) was added, and the solutions diluted to
5 mL with acetonitrile. After 15 min, the absorbance was measured at
430 nm. Calibration graphs are linear in the range 10–50 mg/mL of
dipyridamole (molar absorptivity equal to 10,300). The results obtained
using this method compared well with those obtained by the official
Ganescu et al. used a thiocyanto–Cr(III)-complex for the spectro-
photometric determination of dipyridamole (1–15 mg) in 25 mL of
solution [16]. The solution was acidified with 1–2 mL of 1 M hydrochloric
acid, and treated with a slight excess of ammonium diimidazoletetrakis
(thiocyanato) chromium(III). The precipitate was collected and washed,
dissolved in acetone, and diluted to 25 mL with the same solvent. The
absorbance was measured at 540 nm (molar absorptivity equal to 846).
Shoukry et al. used selected chromotropic acid azo dyes to develop a
spectrophotometric method for the determination of dipyridamole and
chlorpheniramine maleate [32]. When using p-chlorophenylazochromo-
tropic and 2-( p-nitrophenylazo)chromotropic acids, the method is
simple, accurate, and highly sensitive for the analysis of the drugs. The
method consists of extracting the ion-pair complexes into methylene
chloride. After reaction with dipyridamole, the p-chlorophenylazochro-
motropic acid product exhibits an absorption maximum at 544 nm, while
the 2-( p-nitrophenylazo)chromotropic acid product exhibits a maximum
at 524 nm. Dipyridamole was determined upto 3.5–3.78 mg/mL using
these reagents. The molar absorptivity and Sandell sensitivity of the
reaction products were calculated. Statistical treatment of the results
reflects that the procedure is precise, accurate, and is easily applied for the
determination of the drugs in pure form and in their pharmaceutical
preparations.
4.2.6.4 Fluorimetric methods
Li described a spectrofluorimetric method for the determination of
dipyridamole in plasma [33]. Three to 5 mL aliquots of chloroform are
used to extract dipyridamole from 0.5 to 1 mL samples of plasma. After
centrifugation, the fluorescence of the extract is measured at 465 nm
(excitation at 415 nm) against a blank. The calibration graph was found
to be linear over the range of 40–800 ng/mL, and the detection limit was
10 ng/mL. The coefficients of variation (n ¼ 5) were 3.43 and 0.97% at
76.31 and 261.02 ng/mL, respectively.
Shao et al. recommended the use of a simultaneous fluorimetric method
for the determination of the dissolution rate of dipyridamole and aspirin
tablets [34]. The powdered tablets (equivalent to weight of one tablet) of
dipyridamole and aspirin were dissolved in simulated digestive fluids
at 37 C, cooled, and diluted to 1 L with simulated digestive juice. The
solution was filtered, and a 1 mL portion of the filtrate was mixed with
0.1 M sodium hydroxide and then set aside at room temperature for 1 h.
The solution was mixed with 8 mL of phosphoric acid buffer (pH 6.8)
and fluorimetrically detected for dipyridamole at 493 nm (excitation at
418 nm). The coefficients of variation for within-day and within 5 days
were 2% for both dipyridamole and aspirin.
Pulgarin et al. reported the use of a direct method for the determination
of dipyridamole in serum [35]. Dipyridamole (50–1000 ng) was added to
2.8 mL of 1 M ammonium chloride/sodium hydroxide buffer solution of
pH 10 and 1.5 mL ethanol, followed by 0.5 mL of serum. The resultant
mixture was diluted to 10 mL with water. Detection was by fluorescence
spectrometry at 495 nm (excitation at 412.2 nm). Calibration graphs were
linear from 5–100 ng/mL dipyridamole, and the detection limit was
2.8 ng/mL. Relative standard deviations were 1.6–14.4% (n ¼ 3) for
repeatability, and 2.5–23.3% (n ¼ 3) for reproducibility. Recoveries were
in the range of 99.1–102.7%. The effect of varying ethanol content, pH,
and temperature was discussed.
Umapathi et al. used spectrofluorimetric methods for the estimation of
aspirin and dipyridamole in pure admixtures and in dosage forms [36].
The methods are described for the determination of the two drugs in
pharmaceutical preparation without prior separation from each other
and the matrix. A finely ground sample of the tablets was extracted
with chloroform. The extract was filtered, and appropriate volumes of
the filtrate were diluted with 1% acetic acid in chloroform for the
determination of aspirin and pure chloroform for the determination of
dipyridamole. Aspirin and dipyridamole were determined at 345 nm
(excitation at 246 nm) and 475 nm (excitation at 420 nm), respectively.
Linear calibration graphs were obtained for 1–12 mg/mL aspirin and
2–12 mg/mL dipyridamole. Recoveries from 60 to 100 mg aspirin samples,
and 75 mg dipyridamole from tablets, were 98.9–99.75 and 98.94–
100.14%, respectively. The relative standard deviations (n ¼ 5) were less
than 0.53%.
Steyn described and made a comparison between two spectrofluorimetric
methods for the determination of dipyridamole in serum [37]. The thin-
layer chromatographic-fluoridensitometric method utilizes 1 mL of
plasma, which is extracted at pH 10 with diethyl ether–dichloromethane
(80:20). The organic phase is evaporated to dryness, reconstituted in
250 mL dichloromethane, and 5 mL aliquots spotted on a silica gel 60
plate. The plate is developed in ethyl acetate–methanol–ammonia
(85:10:5), dried, dipped in a paraffin wax solution, dried, and scanned
at an excitation wavelength of 380 nm. Using a 430 nm cut-off filter, all
emitted light is collected using a photomultiplier tube. Quantitation was
performed by the external standard method, with peak heights being
measured and a calibration graph constructed. For the spectrofluori-
metric method, 1 mL of plasma is extracted at pH 10 with 8 mL of
hexane-isoamyl alcohol (95:5), and the organic phase directly used for the
measurement of the fluorescence response (excitation at 405 nm, and
emission at 495 nm). Quantitation was accomplished by measuring the
fluorescence of standards that were treated as above and constructing
a calibration graph of concentration versus fluorescence intensity.
Concentrations of unknowns were found by interpolation from this
graph. The two methods were found to exhibit good correlation, but the
spectrofluorimetric method proved to be more amenable to the analysis
of a large number of samples.
Murillo-Pulgarin et al. used a non-linear variable-angle synchronous
fluorescence method for the simultaneous determination of dipyridamole
and three other drugs that have potential for abuse by athletes [38]. The
drugs studied (atenolol, propranolol, amiloride, and dipyridamole) were
determined simultaneously at concentrations of 10–400, 6–200, 5.6–280,
and 5–100 ng/mL, respectively. Analyses were performed at 20 C in
aqueous 70% ethanol at pH 7.5, adjusted with trishydroxymethyl
aminomethane buffer solution. A spectrum based on a single scan was
obtained by nonlinear variable-angle synchronous fluorimetry, with
measurement of the absolute values of the first derivative at excitation/
emission wavelengths of 228.8/300, 287.2/340, 366.4/412.8, and 288/487.2
nm for the four drugs, respectively. The detection limits were 5.2, 1.9, 2.1,
and 0.1 ng/mL for the four drugs, respectively, and the relative standard
deviations for the repeatability and reproducibility were 0.8–2.5% and
0.85–11.5%, respectively. When applied to dosage forms of one com-
pound to which the others were added, recoveries were in the range of
99.8–110%.
Gong and Zhang developed a modified b-cyclodextrin based fluorosensor
for dipyridamole [39]. Samples (2 mL) were injected into a carrier stream
(2 mL/min) consisting of 0.1 M-sodium hydroxide–0.1 M sodium dihy-
drogen phosphate buffer of pH 8. The stream passed through a flow cell
(25 mL) packed with silica gel (100–120 mesh) on which 2,6-O-diethyl-b-
cyclodextrin had been immobilized, and the fluorescence of the complex
with dipyridamole was measured at 468 nm (excitation at 414 nm).
Between injection of samples or standard, 2 mL of aqueous 70% methanol
was injected to desorb the dipyridamole from the cell. Calibration graphs
were linear, with a detection limit of 0.8 nM dipyridamole. The relative
standard deviation (n ¼ 10) was 1.4% for 50 nM dipyridamole. No
interference was observed with the 15 compounds tested (sugars, ascorbic
and salicylic acids, trace metals, or starch). Urine samples were analyzed
after mixing 5 mL with 5 mL of the buffer and diluting to 25 mL.
Recoveries of 100–150 nM dipyridamole solutions were in the range
96–101%. Extracts of tablets and dilutions of injection formulations of
dipyridamole gave results that were in agreement with the labeled values
and with those obtained using the 1995 Chinese Pharmacopoeial method.
Bahbouh et al. used a spectrophotometric method for the determination
of dipyridamole in pure drug and its dosage forms using chromotrope 2B
and phosphotungestic acid [21]. Ground tablets containing 0.1–1 mg
dipyridamole were dissolved in 100 mL of 1 mM hydrochloric acid.
A portion of this solution was transferred to a 50 mL separating funnel,
treated with 1 mL of 0.1 M of hydrochloric acid and 10 mL of 1 mM
chromotrope 2B, and then extracted with two 5 mL aliquots of
chloroform. The separated organic extracts were centrifuged, and the
absorbance was measured at 540 nm against a reagent blank. Beer's law
was obeyed from 5 to 60 mg/mL, and the recoveries were 101.9–102.8%.
Results obtained using this method were in good agreement with those
obtained by the official pharmacopoeial methods.
El-Yazbi et al. developed a spectrophotometric method for the
determination of dipyridamole in ternary mixtures by the derivative
ratio spectrum-zero crossing approach [40]. This method was used for the
resolution of ternary mixtures with overlapping spectra, and is based on
the simultaneous use of the zero-crossing and Salinas method developed
by Berzas-Nivado et al. [41]. The method involves the use of the first
derivative of the ratio spectra (obtained by dividing the absorption
spectrum of the mixture by that of one of its components) and zero-
crossing wavelength measurements. The concentrations of the other
components were obtained from their calibration graphs. The utility
of the method was illustrated by resolution of two ternary mixtures,
one being dipyridamole, aspirin, and salicylic acid, and the other
being dipyridamole, oxazepam, and 2-amino-5-chlorobenzophenone.
The method was found to be suitable for both synthetic mixtures and
commercial dosage forms. The strength of the procedure was in that
it requires no solvent extraction and provides a direct estimate of
degradation products in the presence of the intact drug.
Cao et al. used chemometrics to enhance a spectrofluorimetric resolution
for the determination of dipyridamole in mixture with a amiloride and
propranolol (closely overlapping drug mixtures) [42]. The mixture was
determined in aqueous solution by fluorimetry, with excitation wave-
lengths of 220–230 nm (4 nm intervals) and emission wavelengths of
330–522 nm (4 nm intervals), at a scan rate of 1200 nm/min. A
49  26  9 data array was collected, and treated by a modified parallel
factors analysis algorithm with penalty diagonalization error. Linear
ranges for dipyridamole, amiloride, and propranolol were 20–110,
10150, and 9–100 ng/mL, respectively, and the corresponding correla-
tion coefficients were better than 0.999.
Ruiz-Medina et al. prepared a flow-through optosener with fluorimetric
transduction for the sensitive and selective determination of dipyridamole
in aqueous solutions and biological fluids [43]. The method is based on a
monochannel flow-injection analysis system using Sephadex QAE A-25
resin, placed into a Hellma 176-QS fluorimetric flow-through cell, as an
active sorbing substrate. The native fluorescence of dipyridamole fixed on
the solid sorbent is continuously monitored at wavelengths 305 nm and
490 nm for excitation and emission, respectively. After obtaining the
maximum fluorescence intensity, the eluted solution (potassium dihydro-
gen phosphate/sodium hydroxide buffer solution, c(T) ¼ 0.05 mol/L, pH
6) is allowed to reach the flow cell, the analyte is removed, and the resin
support is regenerated. When a sodium hydroxide (104 mol/L)/sodium
chloride (0.1 mol/L) solution is used as a carrier solution at a flow rate
of 1.56 mL/min, the sensor responds linearly in the measuring range of
10–500 mg/L with a detection limit of 0.94 mg/L and a throughput of
22 samples per hour (300 mL of sample volume). The relative standard
deviation for ten independent determinations (200 mg/L) is less than
0.82%. The method was satisfactorily applied to the determination of
dipyridamole in pharmaceutical preparations and in human plasma.
4.2.6.5 Phosphorimetry methods
Murillo-Pulgarin et al. used a phosphorimetry method for the
determination of dipyridamole in pharmaceutical preparations [44]. Ten
tablets or capsules were powdered and homogenized, and then 0.1 g was
dissolved in 0.1 M sodium dodecyl sulfate. The determination of
dipyridamole was carried out in 26 mM sodium dodecyl sulfate/
15.6 mM thallium nitrate/20 mM sodium sulfite, whose pH was adjusted
to 11.5 by the addition of sodium hydroxide. After 15 min at 20 C, the
phosphorescence was measured at 616 nm (after excitation at 303 nm).
The calibration graph was linear from 100–1600 ng/mL, with a detection
limit of 16.4 ng/mL. Relative standard deviations were in the range of
0.5–7.3%, and sample recoveries were in the range of 95–97%.
Munoz-de-la-Pena et al. described a micellar-stabilized room temperature
phosphorescence method for the stopped-flow determination of dipyr-
idamole in pharmaceutical preparations [45]. The pharmaceutical
preparation was dissolved in and diluted with 0.1 M sodium dodecyl
sulfate. A portion of the resulting solution was mixed with 9 mM sodium
dodecyl sulfate, 1 mM sodium hydroxide, and 15 mM sodium sulfite in
a drive syringe. A second drive syringe was filled with 1 mM sodium
hydroxide, 15 mM sodium sulfite, and 60 mM thallium nitrate. The
contents of the two syringes were mixed in a stopped-flow mixing
chamber, and the phosphorescence intensity was recorded at 616 nm
(after excitation at 303 nm). Dipyridamole was quantified by measuring
the phosphorescence intensity after 10 s (method A), or by measuring the
maximum slope of phosphorescence development over a time interval of
1 s (Method B). The calibration graph was linear for 50–400 ng/mL
dipyridamole for both methods. The detection limits were 21.5 and 37.5
ng/mL with methods A and B, respectively. The relative standard
deviations were 1.4–11.5% for method A and 2.7–21% for method B.
Salinas-Castillo et al. used a new, rapid, sensitive, simple, and selective
phosphorimetric method for the determination of dipyridamole in
pharmaceutical preparation [46]. The phosphorescence signals are a
consequence of intermolecular protection when analytes are exclusively
in the presence of heavy atom salts, and sodium sulfite is used as an
oxygen scavenger to minimize RTP quenching. The determination was
performed in 0.1 mol/L thallium(I) nitrate and 8 mmol/L sodium sulfite
at a measurement temperature of 20 C. The phosphorescence intensity
was measured at 635 nm, after excitation at 305 nm. A linear concentra-
tion range was obtained between 0 and 100 ng/mL, with a detection limit
of 940 ng/L, an analytical sensitivity of 2.5 ng/mL, and a standard
deviation of 2.7% at 60 ng/mL concentration. The method has been
successfully applied to the analysis of dipyridamole in a unique Spanish
commercial formulation containing 100 ng/mL per capsule. The recovery
was 101.6%, with a standard deviation of 6.5%. The method has been
validated using standard addition methodology.
4.2.6.6 Chemiluminescence methods
Nakushima et al. determined dipyridamole on the basis of its
chemiluminescence intensity after injecting into a carrier stream of
triethylamine in acetonitrile (1 mL/min) combined with a stream of
bis-(2,4,6-trichlorophenyl)oxalate and hydrogen peroxide [47].
Sugiura et al. studied the effect of additives of reaction solvents in a
peroxyoxalate chemiluminescence detection system for liquid chromato-
graphy [48]. The method described previously by Imai et al. [49] was
adopted to obtain the reaction profile and optimum conditions for the
chemiluminescent detection with bis[4-nitro-2-(3,6,9-trioxadecyloxycar-
bonyl) phenyl]oxalate and hydrogen peroxide by changing the reaction
coil lengths between the post-column mixer and the detector. The
addition of a phthalate ester to the basic solvent mixture for the
chemiluminogenic reagents, ethyl acetate-acetonitrile (1:1), gave a
considerable reduction in the baseline noise level without decreasing
the fluorescent signal from the test compound, dipyridamole. A mixture
of 0.25 mM bis-[4-nitro-2-(3,6,9-trioxadecyloxycarbonyl) phenyl]oxalate
and 25 mM hydrogen peroxide in ethyl acetate–acetonitrile–0.25%
di(2-ethylhexyl)phtlalate was optimum for detection of amol levels of
fluorescent compounds. The detection limit of dipyridamole was 10 amol.
Imai et al. studied the catalytic effect of imidazole and other bases on the
peroxyoxalate chemiluminescence reaction for HPLC [49]. The peak
height of dipyridamole obtained using the eluent containing buffers was
largest at pH 7, a few times less at pH 6 and pH 5, 100 times less at pH 4,
and a few hundred times less at pH 3. Dipyridamole was separated on the
ODS column, with a detection limit of dipyridamole equal to 40 amol.
Sugiura et al. also developed a chemiluminescence detection system
using bis-[4-nitro-2-(3,6,9-trioxadecyloxycarbonyl) phenyl]oxalate for
the sensitive determination of dipyridamole and the other fluorescent
compounds separated with an acidic mobile phase [50]. Dipyridamole
samples (10 fmol) and dansylated amino acids (10 fmol) were subjected
to HPLC on two systems. System A involved the mixing of the cited
chemiluminescent reagent and an imidazole buffer solution in a coil,
resulting in the generation of active intermediates, which were then
mixed with the column eluate in a two dimensional mixing device.
System B involved the mixing of the chemiluminescent reagent, buffer,
and column eluate directly in a rotating flow-mixing device fitted with
three directional inlets (0.3, 0.3, and 0.5 mm, i.d.) followed by
peroxyoxalate chemiluminescence detection. Detection limits were 10
amol and sub-fmol for dipyridamole and the dansylated amino acids,
Nie et al. used a flow injection chemiluminescence method for the
determination of dipyridamole in Persantin tablets [51]. A flow injection
analysis system equipped with a chemiluminescence detector was used.
Sodium hypochlorite was used as a chemiluminescent substance, and
Triton X-100 was used as an enhancer of the chemiluminescence reaction.
Sample solutions (pH 4–6) was injected into 0.2% Triton X-100 and
treated with 0.5% sodium hypochlorite, and the chemiluminescence
intensity then measured. The calibration graph was linear from 0.04 to
10 mg/mL of Persantin tablets (as its active ingredient dipyridamole), with
a detection limit of 11 ng/mL. Recoveries were quantitative and the
relative standard deviation (n ¼ 3) was 2.7%. The method has been used
to determine Persantin in injection and tablets, and the results obtained
using this method agreed with those obtained by the Chinese
Pharmacopoeial method.
Zheng and Zhang developed a new electrogenerated chemiluminescence
method for the determination of Persantin with flow-injection analysis
[52]. A sample obtained from 20 ground tablets was dissolved in water,
and a 90 mL portion of the solution was injected into a carrier stream
of 8 mM luminol at 4 mL/min in a flow-injection analysis manifold.
The mixed stream was merged with a reagent stream containing
hypobromite (obtained online by galvanostatic electrolysis of 0.15 M
potassium bromide in sodium carbonate/sodium bicarbonate buffer of
pH 11) at 3 mL/min, in an electrolytic flow-cell, at a current of
200 mA. The reduced chemiluminescence due to inhibition caused by
the dipyridamole in Persantin was measured. The calibration graph
was linear from 0.01 to 2 mg of dipyridamole, with a detection limit of
4 mg/L. The relative standard deviation was 2%, and no interference
was observed.
Yang et al. determined dipyridamole by flow-injection analysis method
with chemiluminescence detection [53]. Portions (50 mL) of a standard
solution of dipyridamole were injected into a carrier stream of 2 mM
hydrogen peroxide at 1.3 mL/min in a flow-injection manifold, and
merged with a confluent of two reagent streams (50 mM potassium
permanganate and 2 M sulfuric acid) flowing at the same rate. The
merged stream was mixed in transfer tubing (0.8 mm i.d.), and the
resulting chemiluminescence intensity was measured. The calibration
graph was linear from 0.2 to 80 mg/mL of dipyridamole, with a detection
limit of 58 ng/mL. Most co-existing ions and substances did not interfere.
The method was used in the direct analysis of dipyridamole in both
tablets and injection solutions, with relative standard deviations of
1.2–1.4%.
Pascual et al. studied the effect of antioxidants on the luminal chemi-
luminescence produced by dipyridamole [54]. The method is based on the
use of the reaction between luminal and horse radish peroxidase, and is
proposed for the determination of dipyridamole.
4.2.7 Electrochemical methods
4.2.7.1 Polarography method
Tuncel et al. investigated the polarographic behavior of dipyridamole in a
supporting electrolyte consisting of 0.2 M potassium chloride, 20% (v/v)
ethanol, and buffer [55]. The optimum polarographic conditions were pH
7–8, potential rate of 4 mV/s, drop time of 1 s, and a pressure of 1000
dyne cm2. The effect of concentration on the limiting current was
examined using direct current, superimposed constant amplitude pulse,
superimposed amplitude pulse, and differential pulse polarographic
modes at pH 7.8 and optimum conditions. Differential pulse polaro-
graphy was compared with ultraviolet spectrophotometry for the
determination of dipyridamole in tablets. The methods were similar
in terms of simplicity, reproducibility, and time consumption, but the
differential pulse polarography method was ten times more sensitive and
more specific.
4.2.7.2 Voltammetric methods
Zeng et al. reported the measurement of trace dipyridamole by adsorptive
stripping voltammetry [56]. A solution (10 mL) of dipyridamole in 0.05
M sodium hydroxide containing 10% ethanol was purged with nitrogen
for 4 min before pre-concentration of dipyridamole on a static-Hg-drop
electrode at 1.2 V versus silver/silver chloride. A platinum wire counter
electrode was used. After quiescence for 15 s, the voltammogram was
recorded by applying a linear-sweep scan to 1.6 V at 100 mV/second.
The calibration graph was rectilinear from 0.005 to 1 mM dipyridamole
(pre-concentration for 1 min), and a detection limit of 1 nM was
attainable by pre-concentration for 5 min. For 0.5 mM dipyridamole
and a pre-concentration of 15 s, the relative standard deviation was 1.7%
(n ¼ 12). The method was applied to injection solutions (recoveries
88–92%), tablets (recovery was 105%, and relative standard deviation
of 9.4%), and urine.
Wang et al. determined trace amounts of dipyridamole by stripping
voltammetric using a Nafion modified electrode [57]. Human serum
(0.5 mL) containing dipyridamole was mixed with 0.5 mL water and
1 mL 20% TCA, and centrifuged. A portion (0.1 mL) of the supernatant
was mixed with 10 mL of Britton-Robinson buffer (pH 1.7), and analyzed
by anodic stripping voltammetry using a Nafion-modified vitreous C
working electrode (0.126 cm2). After accumulation for 1 min at 0 V
versus silver/silver chloride (platinum counter electrode) with stirring,
and a 30 s rest period, then a positive scan was initiated at 100 mV/s until
þ 1 V was reached. The calibration graph for dipyridamole was linear for
1–80 nM, and the detection limit was 80 pM for 4 min accumulation.
The relative standard deviation for the determination of 3.78 mg/mL
of dipyridamole in serum was 4.76%, and recoveries were in the range
of 96.7–104%.
Almeida et al. studied the oxidation of dipyridamole in acetonitrile
and ethanol by voltammetric and spectroscopic methods [58]. The
electrochemical oxidation of dipyridamole in acetonitrile and ethanol
at high and low scan speeds was similar, and exhibited two reversible
voltammetric waves corresponding to consecutive one-electron
reactions. The E1/2 values were 120 and 400 mV in acetonitrile,
and 470 and 680 mV in ethanol. The release of one electron
produced a radical cation which was detected by electron spin
resonance spectrometry. The electrochemical oxidation of dipyrida-
mole was accompanied by a decrease in fluorescence intensity at 470
nm, and significant changes in the ultraviolet–visible absorption
spectrum. The oxidation of dipyridamole proceeded more slowly in
ethanol than in acetonitrile.
Yang studied the anodic voltammetry of dipyridamole at a glassy carbon
electrode [59]. Portions of a standard dipyridamole solution were mixed
with 10 mM hydrochloric acid for anodic voltammetry, using a glassy
carbon working electrode and measurement of the anodic oxidation peak
at 0.62 V (versus saturated calomel electrode). The calibration graph was
linear from 0.5 to 10 mg/L of dipyridamole, and over 20 foreign species
did not interfere. The method was directly applied to the analysis of
dipyridamole in tablets and urine, with recoveries of 100.5–101.2% and
a relative standard deviation of 1.5–2.2%.
Ghoneim et al. described a cathodic adsorptive square-wave stripping
voltammetric method for the assay of dipyridamole in human serum
[60]. The rapid and sensitive square-wave voltammetric procedure was
optimized for the determination of dipyridamole after its adsorption
pre-concentration onto a hanging mercury drop electrode. The peak
current of the first of the two peaks developed for this drug in Britton-
Robenson buffer at pH 8 was considered for the analytical study.
An accumulation potential of 1.0 V against Ag/AgCl/KCls, a pulse
amplitude equal to 100 mV, a scan increment of 10 mV, and a
frequency of 120 Hz were found to be the optimal experimental
parameters. Dipyridamole can be determined in the concentration
range of 9 nM–5 mM using accumulation times of 30–300 s. A detection
limit of 40 pM was achieved after a 300 s accumulation time.
Applicability to serum samples was illustrated. The average recoveries
for dipyridamole spiked to serum at 0.25–4.5 mg/mL were 96–102%,
with a standard deviation of 2.9%. A detection limit of 0.06 mg/mL of
serum was obtained.
4.2.8 Mass spectrometry
Rodrigues Filho et al. studied the fragmentation of dipyridamole and
several of its derivatives by electrospray ionization combined with
collisional activated decomposition mass spectrometry in both positive
and negative modes [61]. These compounds produce abundant mono-
charged ions ([M þ H] þ ) under electrospray ionization. Interpretation
of the collisional activated decomposition spectra showed that fragmen-
tation occurs preferentially in the ethanolamine groups attached at C-2,
C-4, C-6, and C-8. 2-Methoxyethanol is eliminated when ethanolamines
are in positions C-2/C-6, and 2-aziridinethanol is eliminated from
C-4/C-8 ethanolamines. The fragmentation schemes were supported by
deuterium labeling experiments and tandem mass spectrometry.
4.2.9 Chromatographic methods
4.2.9.1 Thin-layer chromatography
Strelyuk described a thin-layer chromatographic method for the
qualitative detection of dipyridamole [62]. Data were presented on the
use of various reagents for the detection by solution color change and
precipitate formation in the thin-layer chromatography of dipyridamole.
Thin-layer chromatography was on silica gel–calcium sulfate dihydrate,
with methanol–aqueous ammonia or benzene-dioxane being used as the
mobile phase with detection at 270–330 nm. The Rf values were 0.7–0.75
and 0.5–0.55 for the two solvent systems, and the detection limit was
0.08 mg.
Argekar and Kunjir reported the use of a high performance thin-layer
chromatographic method for the simultaneous determination of aspirin
and dipyridamole in pharmaceutical preparations [63]. Tablets contain-
ing aspirin and dipyridamole were powdered and dissolved with
ultrasonication in methanol-ethanol (1:1). The solution was filtered and
further diluted with methanol. The resulting solution was applied to
aluminum-backed silica gel 60F254 high performance thin-layer chroma-
tographic plates. The plates were developed with the upper layer of ethyl
acetate–ethanol–13.5 M ammonia (5:1:1) to 7 cm, and the absorbance of
the spots was measured by densitometry at 290 nm. Calibration graphs
were linear for 60–340 ng of aspirin and 60–380 ng of dipyridamole, with
detection limits of 25 and 10 ng, respectively, and average recoveries of
99.72 and 97.08%, respectively.
The following three TLC systems [2, 64] were reported for the separation
of dipyridamole:
System 1:
Plate: Silica gel G, 250 mm thick, dipped in, or sprayed with, 0.1M
potassium hydroxide in methanol, and dried.
Mobile phase: Methanol:strong ammonia solution (100:1.5).
Reference compounds: Diazepaim Rf 75, chlorprothexine Rf 56,
codeine Rf 33, and atropine Rf 18.
Analyte Rf value: 68 [2, 64].
Plate: Silica gel G, 250 mm thick, dipped in, or sprayed with, 0.1 M
Mobile phase: Cyclohexane:toluene:diethylamine (75:15:10).
Reference compounds: Dipipanone Rf 66, pethidine, Rf 37,
desipramine Rf 20, codeine Rf 0.6.
Mobile phase: Chloroform:methanol (90:10).
Reference compounds: Meclozine Rf 79, caffeine Rf 58, dipipanone,
Rf 33, desipramine Rf 11.
Analyte Rf value: 37 (acidified iodoplatinate solution, positive)
[2, 64].
4.2.9.2 Gas chromatography
The following gas chromatographic system [2, 65] was recommended for
the separation of dipyridamole:
Column: 2.5% SE-30 on 80100 mesh Chromosorb G (acid washed
and dimethyl dichlorsilane-treated), 2 m  4 mm internal diameter
glass column. It is essential that the support is fully deactivated.
Column temperature: Normally between 100 C and 300 C. As an
approximate guide, the temperature to use is the retention index
divided by 10.
Carrier gas: Nitrogen at 45 mL/min.
Reference compounds: n-alkanes, with an even number of carbon
atoms.
Analyte retention index: 1640 [2, 65].
4.2.9.3 High performance liquid chromatography
The following high performance liquid chromatography (HPLC) system
[2, 66] has been described for the analysis of dipyridamole:
Column: Silica (Spherisorb S5W, 5 mm 12.5 cm  4.9 mm i.d.).
Eluent: A solution containing 1.175 g (0.01 M) of ammonium
perchlorate in 1000 mL of methanol, adjusted to pH 6.7 by the
addition of 1 mL of 0.1 M sodium hydroxide in methanol.
k0 Values: 0.2 [2, 66].
Zhou et al. developed an HPLC method for the determination of
dipyridamole, aspirin, and salicylic acid [67]. The HPLC system consisted
of a Lichrospher 5-C18 (4.6 mm  150 mm) column and a UV detector
operated at 227 nm. The mobile phase (eluted at a flow rate of 1 mL/min)
was composed of disodium hydrogen phosphate solution(methanol
(50:50, pH adjusted to 3 by addition of phosphoric acid). The mean
recovery of dipyridamole and aspirin was 100.9–104.1% and 97.01–97.89,
Chevalier et al. analyzed dipyridamole directly in the blood samples by a
reversed phase high performance chromatographic method with fluori-
metric detection [68]. The HPLC system uses a 15 cm  4.8 mm column
packed with Lichrosorb RP8 (10 mm) and a mobile phase consisting of
40:60 v/v acetonitrile–water. Detection was made on the basis of
fluorimetric detection at 410 nm (excitation at 305 nm). Depending on
the model of the detector used, the detection limit of the method was
between 2  1013 and 2  1015 mole injected. The precision was 5% for
a concentration of 180 ng/mL of dipyridamole, and the method was used
for a biodisposition study of dipyridamole in dogs.
Schmid et al. described a rapid and sensitive high-performance liquid
chromatographic method for the determination of dipyridamole in
human plasma [69]. The column used was a 12.5 cm  4.6 mm filled with
Lichrosorb RP 18, 5 mm. The mobile phase consisted of methanol–
0.2 M Tris HCl buffer (80:20), eluted at a flow rate of 1 mL/min.
Spectrofluorimetric detection at an emission wavelength of 478 nm
(excitation at 415 nm) was used for detection. The results were confirmed
by re-chromatographing the eluate from the column on silica gel G thin-
layer plates, which were developed using a solvent system composed
of toluene–isopropanol–ethanol–ammonia (70:15:15:1) and n-butanol–
methyl ethyl ketone (80:20). The Rf values of dipyridamole for the two
solvent systems are 0.60 and 0.80, respectively. The plates were dried in a
stream of cold air, and then inspected under ultraviolet light at 254 nm.
Wolfram and Bjornsson described a rapid, sensitive, and specific high
performance liquid chromatographic method for the quantitative
analysis of dipyridamole in plasma and whole blood [70]. The method
involves a single extraction on an alkalinized sample with diethyl ether,
followed by evaporation of the organic solvent and ion-pair chromato-
graphy using fluorescence detection. The HPLC system consists of a
Waters model 6000 A high-pressure solvent delivery system (equipped
with a Model U6K injector), and a Waters mBondapak C18 reversed-
phase column (30 cm  39 mm; 10 mm). The mobile phase is a mixture
of methanol–water (65:35), containing 1-heptanesulphonic acid sodium
salt (0.005 M) with 0.1% acetic acid. The mobile phase is eluted at a
flow rate of 2 mL/min with a column input of 170 atm (2500 p.s.i.).
The column is insulated with sponge rubber to minimize ambient
temperature variation. The fluorescence signal was measured with a
Schoeffel Model FS 970 fluorimeter. An excitation wavelength of 285
nm was selected in conjunction with a 470 nm emission filter. The lower
limit of sensitivity for dipyridamole is 1 ng/mL. Concentrations of
dipyridamole between 1 and 500 ng per sample are measured, with an
average coefficient of variation of 4.5% for plasma and 7.4% for whole
Williams et al. used a high performance liquid chromatographic assay
method for dipyridamole monitoring in plasma [71]. The HPLC system
uses a Waters model 6000 A solvent delivery pump equipped with a U6K
injector, a mBondapak C19 column (30 cm  39 mm; 10 mm), and a Model
440 absorbance detector. The signal from the detector was quantified
using a Shimadzu data processor and an Omni-Scribe recorder. A mobile
phase flow rate of 1.5 mL/min was produced by a pressure of
approximately 102 atm (1500 p.s.i.). The mobile phase was 50:50 mixture
of acetonitrile and 0.01 M sodium phosphate in water (adjusted to pH 7).
The absorbance reading of dipyridamole in methanol was made at 280
nm.
Rosenfeld et al. described a high-performance liquid chromatographic
method with ultraviolet detection for the determination of dipyridamole
in plasma [72]. The method is sufficiently sensitive to follow the
concentration of dipyridamole in man for 48 h, and is linear over the
range of concentrations found in man after a standard therapeutic
regimen. The HPLC system used was a Waters model 6000 liquid
chromatograph equipped with a U6K injector and a model 440 dual-
channel filter absorbance detector in conjunction with a Helwett-
Packard 3380 A integrator. The column used was an Altex Ultrasphere
5mm reversed-phase C18 column (25 cm  0.46 mm). The drug was
eluted with 33% acetonitrile in 0.02 M phosphate buffer containing
0.01 M N,N,N,N-tetramethylethylenediamine at pH 2.9. The flow rate
was 2.5 mL/min, which was generated a pressure of approximately 238
bar. The effluent was monitored at 280 nm.
Sane et al. described a HPLC method for the simultaneous estimation of
the combined dipyridamole and aspirin dosage form [73]. Twenty tablets
containing dipyridamole and aspirin were accurately weighed and
powdered. Powder equivalent to 75 mg of dipyridamole was accurately
weighed and 40 mg of pyrimethamine (internal standard) was added. The
contents were first dissolved in 20 mL of acetonitrile, and then in about
40 mL of the mobile phase. The solution was filtered through Whatman
filter paper no. 1. The residue was washed with the mobile phase. The
filtrate and the washings were collected in a 100 mL standard volumetric
flask, and diluted to the mark with mobile phase. A 5 mL portion was
transferred into a 50 mL volumetric flask, and diluted to the mark with
mobile phase. A 10 mL portion of this solution was further diluted to
10 mL, and then 20 ml of this solution was injected onto the HPLC
system. The chromatographic system used a Spectra physics HPLC
equipped with a SP 8810 precision isocratic pump, and a SP 8450 UV–
visible detector with wavelength of detection as 254 nm. The column used
was a Nucleosil C18 (25 cm  4 mm, 10 mm), with a mobile phase
consisting of methanol : acetonitrile : water : triethylamine (55:0:40:0.1;
pH adjusted to 4 using phosphoric acid). The flow rate was 1.5 mL/
min, and the sample injected at ambient temperature using a Rheodyne
7125 (20 mL loop) injector. The standard deviation, coefficient of
variation, and percent recovery for dipyridamole were 0.77, 1.03, and
101.09%, respectively.
Pederson described a specific HPLC method for the determination of
dipyridamole in serum [74]. The HPLC system used was a Waters model
600 liquid chromatograph equipped with a U6K injector, a mBondapak
C18 column (30 cm  39 mm) (10 mm), and a model 440 dual channel filter
absorbance detector in conjunction with a Tarkan W þ W 600 recorder.
The mobile phase was a 75:25 mixture of methanol and a 0.02 M solution
of sodium acetate (adjusted to pH 4 with acetic acid). The solvent flow
rate of 2 mL/min was produced by an applied pressure of approximately
2000 p.s.i. Detection of the analyte was made at the UV absorption
maximum of 280 nm.
Table 6 contains a summary of the conditions of other reported HPLC
methods that has also been reported for the determination of
dipyridamole [49, 75–84].
HPLC Conditions of the Methods Used for the Determination of Dipyridamole Hydrochloride
Column Mobile phase and flow rate Detection Remarks Ref.
A. KHALIL, F. BELAL AND A.A. AL-BADR
15 cm  4 mm of 50 mM imidazole in 50 mM phos- Chemiluminescence Separation of the 49
TSK ODS 80 TM phate buffer of pH 6-acetonitrile drug and benzy-
(5 mm) (1:1) [1 mL/min] and 0.5 mM damine HCl.
bis-[4-nitro-2-(3,6,9-trioxadecy- Detection limit
loxy-carbonyl)-phenyl]oxalate for dipyridamole
(TDPO) in acetonitrile-25 mM- was 40 amol.
H2O2 in ethyl acetate for eluents
of pH 5–7 and a mixture of 0.8
mMTDPO and 4 mM-bis-(2,4-
dinitrophenyl)oxalate for elu-
ents at pH 3–4.
30 cm  3.9 mm 0.1 M Sodium acetate buffer pH Amperometric at Detection and 75
of mBondapak C18 4-methanol (1:3) [1 mL/min] þ 0.70 V vs a determination of
(10 mm) SCE the drug by HPLC
in pharmaceuticals
7 cm  4.6 mm Methanol-0.02 M ammonium 280 nm Analysis of free 76
of Ultrasphere XL acetate buffer pH 5 (13:7) and plasma pro-
ODS (3 mm) [1.5 mL/min]. tein-bound dipyri-
damole by HPLC
DIPYRIDAMOLE: COMPREHENSIVE PROFILE
10 cm  2 mm of Aqueous 20 mM-Na2HPO4 (con- 305 nm and by Rapid isocratic 77
Hypersil ODS taining 50 mM-Na dodecyl fluorimetry at 468 HPLC assay for
(5 mm) sulfate and H3PO4 to pH 2)- nm (excitation at the drug using
acetonitrile (13:12) [0.5 mL/min] 293 nm) a micro-bore col-
umn technique.
Down to 0.05 mg/
mL of the drug
can be detected.
25 cm  4.6 mm of Methanol-aqueous 200 mM 288 nm Stability indicating 78
Phenomenex sodium pentanesulfonate (7:3) analysis used to
ODS 1 (5 mm) containing triethylamine 2 mL/L analyse the raw
operated at 60 C. and adjusted to pH 3 with phos- material and the
phoric acid [1.5 mL/min). capsule formula-
30 cm  3.9 mm 0.1 M sodium acetate buffer of Coulometric at Sensitive isocratic 79
of mBondapak C18 pH 4/methanol (9:11) [1 mL/ þ 0.65 V HPLC for the
(10 mm) min]. assay of the drug
in plasma
TSK gel ODS-80- 5 mM phosphate buffer pH 5- Chemiluminescent Separates the drug 80
TM acetonitrile (3:2) [1 mL/min] reagent. and dansylated
and 5 mM bis [4-nitro-2-(3,6,9- valine. Detection
trioxadecyl oxycarbonyl)phenyl] limit under the
-oxalate TDPO and 50 mM- optimum condi-
acetonitrile [1.3 mL/min] tions is 20 amol
for dipyridamole.
30 cm  3.9 mm Sodium acetate-acetic acid of pH 276 nm Stability indicating 81
of mBondapak C18 5.1  0.1 in methanol (70:130) HPLC method.
(10 mm) [1 mL/min]. Analysis of the
drug in injection
30 cm  3.9 mm Aqueous sodium acetate pH 5– 276 nm Stability indicating 82
of Waters 36% acetic acid–650 mL metha- HPLC method
mBondapak nol [1 mL/min]. for analysis of
C18 (10 mm) the degradation
products in dipyr-
idamole injection.
25 cm  4 mm of 15 mM-NaH2PO4 buffer (pH 7.6) 280 nm and by Purity evaluation 83
LiChrosorb RP-18 containing 60–90% of acetoni- fluorimetry at of the drug by
trile [2 mL/min] 470 nm (excita- HPLC. Three
tion at 285 nm) impurities were
separated from
the drug.
15 cm  4.6 mm of 50 mM imidazole buffer (pH 6)- Chemiluminescence Analysis in rat 84
TSK ODS 80 TM acetonitrile (1:1). Eluate was plasma by HPLC
(5 mm) operated at mixed with 0.25 mM-bis-[(4- of dipyridamole
40 C nitro-2-(3,6,9-trioxadecyloxy- and benzydamine.
carbonyl)phenyl]oxalate and
12.5 mM-H2O2 solution in acet-
onitrile–ethyl acetate (1:1)
The stability of freshly prepared standard solutions of dipyridamole
exposed to daylight showed considerable variation [74]. The rate of
degradation seemed to follow first-order kinetics, and the rate constant
was obviously dependent on the light intensity in the laboratory. The
maximum degradation rate was found in the solutions with TRIS-
buffered water, in which the half-time of dipyridamole varied from about
3–30 h. The corresponding half-time in ethanol was 12–240 h. Samples
prepared with deproteinized serum were stable for several days. No
difference between the reagent tubes was found, and there was no
evidence of adsorption on the glassware or plastic tubes. These findings
strongly indicate that all samples and standards should be protected from
light [74].
6. PHARMACOKINETICS AND METABOLISM
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Mefenamic Acid: Analytical Profile
Hadi Poerwono1, Retno Widyowati1,
Hajime Kubo2, Kimio Higashiyama2
and Gunawan Indrayanto1
Faculty of Pharmacy, Airlangga University
Jl. Dharmawangsa Dalam
Surabaya 60286, Indonesia
Institute of Medicinal Chemistry Hoshi University
4-41, Ebara 2-chome, Shinagawa-ku
282 H. POERWONO ET AL.
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283
2. Compendial Methods of Analysis . . . . . . . . . . . . . . . . . . . . 284
2.1.1 Bulk drug substance . . . . . . . . . . . . . . . . 284
2.1.2 Drug substance in pharmaceutical
preparations . . . . . . . . . . . . . . . . . . . . . . 286
2.2 Physical methods of analysis . . . . . . . . . . . . . . . . 287
2.2.1 Loss on drying . . . . . . . . . . . . . . . . . . . . 287
2.2.2 Residue on ignition . . . . . . . . . . . . . . . . . 288
2.2.3 Sulfated ash . . . . . . . . . . . . . . . . . . . . . . 288
2.2.4 Light absorption . . . . . . . . . . . . . . . . . . . 288
2.3 Impurity analyses . . . . . . . . . . . . . . . . . . . . . . . . 288
2.3.1 Heavy metals . . . . . . . . . . . . . . . . . . . . . 288
2.3.2 Copper. . . . . . . . . . . . . . . . . . . . . . . . . . 288
2.3.3 Chromatographic purity . . . . . . . . . . . . . 289
2.3.4 Related substances . . . . . . . . . . . . . . . . . 289
2.3.5 2,3-Dimethylaniline . . . . . . . . . . . . . . . . . 290
2.4 Assay methods . . . . . . . . . . . . . . . . . . . . . . . . . . 291
2.4.1 Titration methods . . . . . . . . . . . . . . . . . . 291
3. Titrimetric Methods of Analysis . . . . . . . . . . . . . . . . . . . . 291
3.1 Non-aqueous titration methods . . . . . . . . . . . . . . 291
3.2 Potentiometric titration methods . . . . . . . . . . . . . 292
4. Electrochemical Methods of Analysis . . . . . . . . . . . . . . . . . 293
4.1 Polarography . . . . . . . . . . . . . . . . . . . . . . . . . . . 293
4.2 Conductimetry . . . . . . . . . . . . . . . . . . . . . . . . . . 293
5. Spectroscopic Methods of Analysis . . . . . . . . . . . . . . . . . . 293
5.1 Spectrophotometry . . . . . . . . . . . . . . . . . . . . . . . 293
5.2 Colorimetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296
5.3 Fluorimetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298
5.4 Chemiluminescence . . . . . . . . . . . . . . . . . . . . . . . 301
5.5 Proton magnetic resonance spectrometry. . . . . . . . 302
5.7 Atomic absorption spectrometry. . . . . . . . . . . . . . 302
6. Chromatographic Methods of Analysis . . . . . . . . . . . . . . . . 303
6.2 Column chromatography . . . . . . . . . . . . . . . . . . . 304
MEFENAMIC ACID: ANALYTICAL PROFILE 283
6.4 High performance liquid chromatography . . . . . . . 309
6.5 Supercritical fluid chromatography . . . . . . . . . . . . 313
6.7 Capillary isatachophoresis . . . . . . . . . . . . . . . . . . 328
7. Determination in Body Fluids and Tissues. . . . . . . . . . . . . . 329
7.1 Spectroscopic methods. . . . . . . . . . . . . . . . . . . . . 329
7.2 Chromatographic methods . . . . . . . . . . . . . . . . . . 330
Mefenamic acid has the following structure:
HO O
H CH3
The elemental analysis of mefenamic acid calculates as follows:
Nitrogen 5.81%
The empirical formula of mefenamic acid is C15H15NO2, and its formula
weight is calculated to be 241.285.
According to USP 26 [1] and the Indonesian Pharmacopoeia [4],
mefenamic acid is specified to contain not less than 98.0% and not
more than 102.0% of C15H15NO2, calculated on the dried basis. The
substance is to be preserved in tight, light-resistant containers. The
British Pharmacopoeia 2002 [2], European Pharmacopoeia [5] and Indian
Pharmacopoeia [6] describe that mefenamic acid contains not less than
99.0%, and not more than the equivalent of 100.5% of N-2,3-
xylylanthranilic acid, calculated with reference to the dried substance.
The Pharmacopoeia of the People's Republic of China [7] describes
that mefenamic acid is 2-[(2,3-dimethylphenyl)amino]benzoic acid,
and contains not less than 99.0% C15H15NO2, calculated on the dried
basis.
According to USP 26 [1], Indonesian Pharmacopoeia [4], Indian
Pharmacopoeia [6], and Pharmacopoeia of the People's Republic of
China [7], mefenamic acid capsules are specified to contain not less
than 90.0% and not more than 110.0% of the labeled amount of
C15H15NO2. According to the British Pharmacopoeia 2002 [3], mefe-
namic acid capsules and tablets are specified to contain not less than
95.0% and not more than 105.0 percent of the stated amount of the
2.1.1 Bulk drug substance
According to the United States Pharmacopoeia 26 [1] and Indonesian
Pharmacopoeia [4], mefenamic acid is identified by examination using
infrared absorption spectrophotometry (method h197 Ki), comparing
with the spectrum obtained with USP Mefenamic Acid reference
standard (Test A). In addition to the infrared absorption test, the USP
26 [1] includes examination using Liquid Chromatography, where the
retention time of the major peak in the chromatogram of the Assay
preparation corresponds to that in the chromatogram of the Standard
preparation, as obtained in the Assay (Test B). Test B according to the
Indonesia Pharmacopoeia [4] uses a Thin Layer Chromatography test,
where the Rf value of the main spot in the chromatogram of the Test
solution corresponds to that in the chromatogram of the Standard
Identification testing in the British Pharmacopoeia 2002 [2] and
European Pharmacopoeia [5] is arranged in two tiers, classified as the
(a) first identification by test B, and (b) the second identification by tests
A, C and D.
Test A: About 20 mg of the acid sample is dissolved in a mixture
of 1 volume of 1 M hydrochloric acid and 99 volumes of
methanol, and diluted to 100 mL with the same mixture
of solvent. About 5 mL of the solution is diluted to
50 mL in a mixture of 1 volume of 1 M hydrochloric
acid and 99 volumes of methanol. The resulting solution
is examined between 250 nm and 380 nm, whereas the
solution shows two absorption maxima at 279 nm and
350 nm. The ratio of the absorbance measured at the
maximum at 279 nm to that measured at 350 nm is 1.1
to 1.3.
Test B: Examination of the acid sample by infrared absorption
spectrophotometry, comparing with the spectrum obta-
ined with mefenamic acid reference standard. Examine
the substances prepared as KBr pellets. If the spectra
obtained show differences, dissolve the substance to be
examined and the reference substance separately in
alcohol, evaporate to dryness, and record new spectra
using the residue.
Test C: About 25 mg of the acid sample is dissolved in 15 mL of
methylene chloride. The solution is examined in ultravio-
let light at 365 nm to exhibit a strong greenish-yellow
fluorescence. Carefully add 0.5 mL of a saturated solution
of trichloroacetic acid dropwise and examine under
ultraviolet light at 365 nm. The solution does not exhibit
fluorescence.
Test D: About 5 mg of the acid sample is dissolved in 2 mL of
sulfuric acid, to which is added 0.05 mL of potassium
dichromate solution. An intense blue color is produced,
turning rapidly to brownish-green.
Identification testing according to the Indian Pharmacopoeia [6] is
performed by means of three tests (denoted as tests A, B, and C).
Test A: The infrared absorption spectrum is concordant with the
reference spectrum of mefenamic acid or with the
spectrum obtained from mefenamic acid reference stan-
dard. If the spectra are not concordant, dissolve a
sufficient quantity of the substances in the minimum
volume of ethanol (95%), evaporate to dryness, and
prepare new spectra of the residues.
Test B: About 25 mg of the acid sample is dissolved in 15 mL of
chloroform. The solution is examined under ultraviolet
light at 365 nm to exhibit a strong greenish-yellow
Test C: About 5 mg of the acid sample is dissolved in 2 mL of
sulfuric acid, to which is added 0.05 mL of 0.0167 M
potassium dichromate solution. An intense blue color is
produced, which fades rapidly to brownish-green.
According to the Pharmacopoeia of the People's Republic of China [7],
mefenamic acid is identified by the performance of four tests (denoted as
Tests 1, 2, 3, and 4).
Test 1: About 25 mg of the acid sample is dissolved in 15 mL of
light at 254 nm to exhibit a strong green fluorescence.
Test 2: About 5 mg of the acid sample is dissolved in 2 mL of
sulfuric acid, to which is added 0.05 mL of 0.5%
potassium dichromate solution. A deep blue color is
produced which fades immediately to brownish-green.
Test 3: The light absorption of a 20 mg/mL solution in a mixture
of hydrochloric acid (1 mol/L) and methanol (1:99),
exhibits two absorption maxima, at 279 nm and 350 nm.
The absorbance of these is about 0.69 0.74 and
0.59 0.6, respectively.
Test 4: The infrared absorption spectrum is concordant with the
spectrum of mefenamic acid reference standard.
2.1.2 Drug substance in pharmaceutical preparations
Identification test of mefenamic acid in capsule preparations according to
the United States Pharmacopoeia 26 [1] and Indonesian Pharmacopoeia
[4] is performed by the execution of two tests (denoted as Tests A and B)
Test A: A portion of capsule contents (equivalent to about 250 mg
of mefenamic acid) is placed in a 250 mL volumetric flask,
to which is added about 100 mL of a mixture of
chloroform and methanol (3:1) with vigorous shaking. It
is then diluted with a mixture of chloroform and methanol
(3:1) to volume, mixed, and filtered and the filtrate is
obtained. This solution responds to the Thin-Layer
Chromatographic Identification Test h201i when eluted
with a solvent system consisting of a mixture of chloro-
form, ethyl acetate, and glacial acetic acid (75:25:1). The
Ordinary Impurities general test h466i visualization
technique is used.
Test B: Examination using Liquid Chromatography: the retention
time of the major peak in the chromatogram of the Assay
preparation corresponds to that in the chromatogram of
the Standard preparation, obtained as directed in the
Assay.
According to the British Pharmacopoeia 2002 [2] and Indian
Pharmacopoeia [6], mefenamic acid in capsule and tablet preparations
are identified by examination using infrared absorption spectrophotome-
try as the following procedure. Extract a quantity of the capsule contents
(or powdered tablets) containing 0.25 g of mefenamic acid with two 30 mL
quantities of ether. Wash the combined extracts with water, evaporate to
dryness on a water bath, and dry the residue at 105 C. Dissolve a sufficient
quantity in the minimum volume of absolute ethanol, and evaporate to
dryness on a water bath. The infrared absorption spectrum is concordant
with the reference spectrum of mefenamic acid.
According to the Pharmacopoeia of the People's Republic of China [7], the
method of identification testing of mefenamic acid in capsule and tablet
preparations is based on light absorption spectrophotometry. A specific
quantity of the powdered contents of capsules (or powdered tablets),
equivalent to 0.25 g mefenamic acid, is dissolved in a mixture of 10 mL of
0.1 mL/L hydrochloric acid/methanol (1:99), shaken, and filtered. Then
some quantity of the filtrate is diluted with the above-mixed solution to
produce a solution having a concentration of about 20 mg/mL. The absor-
ption spectrum of the solution exhibits maxima at 279 nm and 350 nm.
2.2 Physical methods of analysis
2.2.1 Loss on drying
When the acid sample is dried at 105 C for 4 h according to the USP 26
general procedure h731i, or by the procedure of Indonesian and Indian
Pharmacopoeias, it loses not more than 1.0% of its weight [1, 4, 6].
According to the British and European Pharmacopoeias, as well as the
Pharmacopoeia of the People's Republic of China, it loses not more
than 0.5% determined on 1.000 g by drying at 100–105 C to a constant
weight [2, 5, 7].
2.2.2 Residue on ignition
When examined according to the USP general procedure h281i as well
as the procedure of Indonesian Pharmacopoeia and Pharmacopoeia of
the People's Republic of China, the ash obtained for an acid sample
may not exceed 0.1% [1, 4, 7].
2.2.3 Sulfated ash
When examined according to the British Pharmacopoeia 2002 general
procedure h2.4.14i, as well as the procedure of the European and Indian
Pharmacopoeias, the sulfated ash obtained for an acid sample may not
exceed 0.1%, determined on 1.0 g [2, 5, 6].
2.2.4 Light absorption
The absorbance of a 0.002% w/v solution (in a mixture of 99 volumes of
methanol and 1 volume of 1 M hydrochloric acid) is 0.69–0.74 at the first
maximum of 279 nm, and 0.56–0.60 at the second maximum of 360 nm [6].
2.3 Impurity analyses
2.3.1 Heavy metals
The heavy metal content of mefenamic acid is determined using the USP
26 general procedure h231i, as well as the procedure of Indonesian
Pharmacopoeia [1, 4]. The limit on heavy metals in the substance is
0.002%.
2.3.2 Copper
The copper content of mefenamic acid is determined using atomic
absorption spectrometry, according to the British Pharmacopoeia 2002
general procedure h2.2.23, Method Ii, as well as the procedure of the
European Pharmacopoeia [2,5]. The substance may not contain more
than 10 ppm of Cu.
Indian Pharmacopoeia determines the copper content using a color
intensity method, according to the following procedure. Moisten 1 g of
the substance with sulfuric acid and ignite until all the carbon is removed.
Add 10 mL of 1 M sulfuric acid to the residue and allow to stand for
10 min. Transfer to a separating funnel using 20 mL of water and add 10
mL of a solution containing 20% w/v diammonium hydrogen citrate and
5% w/v of disodium edetate. Add 0.2 mL of thymol blue solution and
neutralize with 5 M ammonia. Add 10 mL of sodium diethyldithiocar-
bamate solution and 15 mL of carbon tetrachloride, shake, and allow to
separate. The yellow color of the carbon tetrachloride layer is not more
intense than that produced by treating 2 mL of copper standard solution
(10 ppm Cu) in the same manner beginning at the directions, ''Transfer
to a separating funnel using 20 mL of water, as described above'' (20
ppm) [6].
According to the Pharmacopoeia of the People's Republic of China, the
above separated carbon tetrachloride layer is measured by the light
absorption spectrophotometry at 435 nm. The absorbance of that
solution is not more than 0.35 [7].
2.3.3 Chromatographic purity
Chromatographic purity according to the USP 26 is determined using
Liquid Chromatography. The impurity content of the substance is not
more than 0.1% for any individual impurity, and not more than 0.5%
for total impurities [1].
2.3.4 Related substances
According to the British Pharmacopoeia 2002 and the European and
Indian Pharmacopoeias, the content of related substances is examined by
thin-layer chromatography using TLC silica gel GF254 plates. A test
solution is prepared by dissolving 0.125 g of the substance to be examined
in a mixture of 1 volume of methanol and 3 volumes of methylene chloride,
and diluting to 5 mL with the same mixture of solvents. Reference solution
(a) is prepared by diluting 1 mL of the test solution to 50 mL with a
mixture of 1 volume of methanol and 3 volumes of methylene chloride,
then diluting 1 mL of the solution to 10 mL with the same mixture of
solvents. Reference solution (b) is prepared by dissolving 5 mg of
flufenamic acid and 5 mg of mefenamic acid reference standard in a
and diluting to 10 mL with the same mixture of solvents. Apply 20 mL of
each solution to the TLC silica gel GF254 plate. Develop over a path of
15 cm using a mixture of 90 volumes of toluene, 25 volumes of dioxane, and
1 volume of glacial acetic acid. Dry the plate in a current of warm air,
and expose the plate to iodine vapor for 5 min, and examine in ultraviolet
light (254 nm). Any spot in the chromatogram obtained with the test
solution, apart from the principal spot, is not more intense than the spot in
the chromatogram obtained with reference solution (a) (0.2%). The test is
not valid unless the chromatogram obtained with reference solution (b)
shows two clearly separated spots [2, 5, 6].
A similar procedure is mentioned in the Pharmacopoeia of the People's
Republic of China that uses a mixture of 3 volumes of isobutanol and
1 volume of concentrated ammonia solution as the mobile phase [7].
2.3.5 2,3-Dimethylaniline
The 2,3-dimetylaniline content is determined by color intensity method,
according to the British Pharmacopoeia 2002 and the European
Pharmacopoeia. Solution (a) is prepared by dissolving 0.250 g of the
substance to be examined in a mixture consisting of 1 volume of
methanol and 3 volumes of methylene chloride, and diluting to 10 mL
with the same mixture of solvents. This solution is used to prepare the test
solution. Solution (b) is prepared by dissolving 50 mg of 2,3-dimethylani-
line in a mixture of 1 volume of methanol and 3 volumes of methylene
chloride, and diluting to 100 mL with the same mixture of solvents. Then
1 mL of the solution is diluted to 100 mL with the same mixture of
solvents. This solution is used to prepare the standard. Using three flat-
bottomed tubes, place 2 mL of solution (a) in the first, 1 mL of solution
(b) in the second, and 1 mL of a mixture of 1 volume of methanol and
3 volumes of methylene chloride in the third. A blank tube is prepared by
making 2 mL of a mixture of 1 volume of methanol and 3 volumes of
methylene chloride. To each tube add 1 mL of a freshly prepared 10 g/L
solution of dimethylamino-benzaldehyde in methanol and 2 mL of glacial
acetic acid. Allow to stand at room temperature for 10 min. The intensity
of the yellow color of the test solution is between that of the blank and
that of the standard (100 ppm) [2, 5].
According to the Indian Pharmacopoeia, 2,3-dimethylaniline is examined
by TLC using a silica gel G plate, and a mixture of 90 volumes of toluene,
25 volumes of dioxane, and 1 volume of 18 M ammonia as the mobile
phase. Apply 40 mL of each of the two solutions in a mixture of 3 volumes
of chloroform and 1 volume of methanol containing (1) 2.5% w/v of the
substance being examined separately to the plate, and (2) 0.00025% w/v
of 2,3-dimethylaniline. After removal of the plate, dry it in a current of
warm air. Spray the dried plate with ethanolic sulfuric acid (20%), heat at
105 C for 30 min, and immediately expose to nitrous fumes in a closed
glass chamber for 15 min (the nitrous fumes may be generated by adding
dilute sulfuric acid dropwise to a solution containing 10% w/v of sodium
nitrite and 3% w/v of potassium iodide). Place the plate in a current of
warm air for 15 min, and spray with a 0.5% w/v solution of N-(1-
naphthyl)ethylenediamine dihydrochloride in ethanol (95%). If neces-
sary, allow to dry, and repeat the spraying. Any spot corresponding
to 2,3-dimethylaniline in the chromatogram obtained with solution (1) is
not more intense than the spot in the chromatogram obtained with
solution (2) [6].
2.4.1 Titration methods
According to the British Pharmacopoeia 2002, the European and Indian
Pharmacopoeias, and the Pharmacopoeia of the People's Republic of
China, the assay for mefenamic acid is performed by a titration method.
About 0.200 g of the substance to be assayed is dissolved with the aid of
ultrasound in 100 mL of warm ethanol that has been previously neutrali-
zed to the phenol red endpoint. About 0.1 mL of phenol red solution is
added and titrated with 0.1 M sodium hydroxide. Each milliliter of 0.1 M
NaOH is equivalent to 24.13 mg of mefenamic acid [2, 5–7].
According to the USP 26 and the Indonesian Pharmacopoeia, the assay
for mefenamic acid is performed by a liquid chromatography method. A
buffer solution is prepared as a 50 mM solution of monobasic ammonium
phosphate, adjusted with 3 M ammonium hydroxide to a pH of 5.0. The
system uses a filtered and degassed mixture of acetonitrile, buffer
solution, and tetrahydrofuran (23:20:7) as the mobile phase. The liquid
chromatograph is equipped with a 254 nm detector and a 4.6 mm
25 cm column that contains packing L1. The flow rate is about 1 mL/min.
The column efficiency is not less than 8200 theoretical plates, the tailing
factor for the analyte peak is not more than 1.6, and the relative standard
deviation for replicate injections is not more than 1.0% [1, 4].
3.1 Non-aqueous titration methods
A quantity of the mixed contents of 20 mefenamic acid capsules
containing 0.5 g of the substance, or a quantity of the powdered tablets
(after weighing and powdering of 20 mefenamic acid tablets) containing
0.5 g of the substance, is dissolved in 100 mL of warm absolute ethanol
previously neutralized to the phenol red endpoint. The resulting solution
of each preparation is titrated with 0.1 M sodium hydroxide using phenol
red solution as the indicator. Each mL of 0.1 M NaOH is equivalent to
24.13 mg of mefenamic acid [2, 6, 7].
Based on the fact that mefenamic acid reacts quantitatively with
N-bromosuccinimide (NBS) in an acidic medium, Hassib et al. [8]
reported the titrimetric determination of mefenamic acid. A solution of
mefenamic acid in acetic acid (0.02% m/V) was prepared. A volume of
this solution containing 1.5–3.5 mg of mefenamic acid was allowed to
react with 40 mL of 0.05 M NBS solution for 10 min in the dark
(temperature of 25  2 C). Potassium iodide solution (10 mL) was added,
and the solution was titrated with 0.01 N sodium thiosulfate to the starch
end-point. A blank determination was carried out. For the determination
of mefenamic acid in the capsules, the contents of 20 PonstanÕ capsules
were mixed thoroughly and an accurately weighed portion of the mixed
powder (nominally containing 25 mg of mefenamic acid) was extracted
with diethyl ether (3  10 mL). The residue remaining after evaporation
of diethyl ether was dissolved in acetic acid, and the procedure was
continued as described for the determination of mefenamic acid.
Issa et al. [9] used various metal ions for the volumetric determination of
mefenamic acid. Mefenamic acid was precipitated from its neutral
alcoholic solution by a standard solution of either silver nitrate,
mercurous acetate, or potassium aluminum sulfate. In the argentimetric
procedure, residual Ag(I) was titrated with standard NH4SCN. With
Hg(OAc)2 or potash alum, the residual metal was determined by adding
EDTA and conducting back titration of excess of EDTA with standard
Pb(NO3)2 using xylenol orange indicator. The applied methods were used
for the determination in bulk drug substance, and in its formulations.
3.2 Potentiometric titration methods
Çakirer et al. [10] reported a potentiometric titration method in
nonaqueous media for the determination of some commonly used
antiinflammatory agents. The direct potentiometric titration of mefe-
namic acid, fenbufen, and ibuprofen, and the indirect potentiometric
titration of diclofenac sodium, were carried out in acetonitrile using
tetrabutylammonium hydroxide solution in 2-propanol/methanol as the
titrant. The titration was performed at 25 C under an atmosphere of
nitrogen. This method gave highly precise recoveries, having a relative
standard deviation of not more than 1.0% for all antiinflammatory
agents studied.
4.1 Polarography
Song et al. [11] developed a polarographic method for the determination
of mefenamic acid in tablets, which was based on rapid nitrosation of
mefenamic acid with sodium nitrite in acetic acid, and subsequent
measurement of the N-nitroso derivative of mefenamic acid by linear-
sweep polarography. The method is simple, sensitive, and specific, and
was characterized by a detection limit of 2  107 mol/L.
4.2 Conductimetry
Aly and Belal reported the conductimetric determination of some non-
steroidal antiinflammatory drugs in their dosage forms [12].
5. SPECTROSCOPIC METHODS OF ANALYSIS
5.1 Spectrophotometry
On the basis of spectral changes induced by changing the solvent medium
from HCl (0.01 M) to NaOH (0.01 M), Hassan and Shaaban [13]
developed a coefficient-difference spectrophotometric method of analysis
for mefenamic acid using the six-points quadratic order of the orthogonal
polynomials. The optimum wavelength range was 326–386 nm, measured
at 12 nm intervals. The mean recovery was 100.1  0.89 % over a
concentration range of 0.6–2.6 mg/100 mL ( p ¼ 0.05). Trials to adapt
single wavelength difference technique as well as the cubic order of the
orthogonal polynomials were also performed; their results were discussed
on a statistical basis. The developed procedures have been applied to the
analysis of some randomly collected market preparations, and the results
proved suitability of the method for its application in routine analysis.
Shinkuma et al. [14] reported a spectrophotometric assay of different
commercial mefenamic acid capsules. The contents of 20 capsules of each
brand were carefully removed and accurately weighed. After finding the
mean weight of the capsule contents, the contents were mixed uniformly
and used as the brand sample. A weighed amount of this powder,
equivalent to about 150 mg of mefenamic acid, was dissolved in 200 mL
of hydrochloric acidmethanol solution (0.84 mL of concentrated
hydrochloric acid/1000 mL methanol). After filtering and appropriate
dilution, the sample was assayed spectrophotometrically at 350 nm using
a double-beam spectrophotometer. Beer–Lambert law was obeyed over
the concentration range used in this work.
Das et al. [15] established a spectrophotometric procedure for the
simultaneous determination of mefenamic acid and paracetamol in
mixtures. Using 0.01 M methanolic hydrochloric acid as a solvent, the
absorbance of the mixture is measured at 248, 279, and 351 nm. The
concentration of each component can be calculated by solving two equat-
ions using two wavelengths, either 248 and 279 nm, or 248 and 351 nm.
The amounts of mefenamic acid and paracetamol were calculated using
either of two sets of equations:
Set A: Mass (mg) of mefenamic acid ¼ 332.2A351
Mass (mg) of paracetamol ¼ 113.04A248  84.4A351
Set B: Mass (mg) of mefenamic acid ¼ 314.8A279  53.0A248
Mass (mg) of paracetamol ¼ 126.6A248  79.9A279
The method was applied for the determination of these drugs in
commercial tablets with recoveries in the ranges of 99.1101.7%
(set A), and 96.7100.75% (set B) for mefenamic acid and
95.999.5% (set A) and 96.499.6% (set B) for paracetamol.
Gangwal and Sharma [16] described a simultaneous spectrophotometric
method for the determination of mefenamic acid and paracetamol in their
combined dosage forms based on the native UV absorbance maxima of
mefenamic acid and paracetamol in 0.02 M NaOH. Mefenamic acid
exhibits two absorption maxima at 285 nm and 333 nm, while paracetamol
has one absorbance maxima at 257 nm. In a separate study, the same
group [17] also reported a spectrophotometric procedure for mefenamic
acid and paracetamol in two component tablet formulations. The method
is based on the two-wavelength method of calculation. The difference in
absorbances at 217 nm and 285 nm was used for determination of
mefenamic acid, and the difference in absorbances at 257 nm and 308.8 nm
was used for the determination of paracetamol. Beer's law is obeyed by
both the drugs within the concentration ranges employed for analysis. The
method has been statistically validated, and was found to be satisfactory.
Parimoo et al. [18] established a UV spectrophotometric method for the
combination preparations. The determination was carried out in two
different solvents, namely methanol and 0.1 N NaOH. The wavelength
maximum for mefenamic acid was found at 284 nm, and 248 nm for
paracetamol. The wavelength maximum for mefenamic acid in NaOH
was found at 219 nm, and 256 nm for paracetamol. There were no
interferences in the analyte estimations.
A direct and simple first derivative spectrophotometric method has been
developed by Toral et al. [19] for the determination of mefenamic acid
and paracetamol in pharmaceutical formulations. A methanolic hydro-
chloric acid solution was used as a solvent for extracting the drugs from
the formulations, and subsequently the samples were directly evaluated
by derivative spectrophotometry. Simultaneous determination of both
the drugs can be carried out using the zero-crossing and the graphical
methods, and the methods do not require simultaneous equations to be
solved. The calibration graphs were linear in the ranges from 1.8  106
to 1.6  104 M of mefenamic acid, and from 4.1  106 to 1.4  104 M
of paracetamol. The ingredients commonly found in commercial
pharmaceutical formulations did not interfere, so the proposed method
was applied to the determination of these drugs in tablets.
Dinç et al. [20] developed four new methods for the simultaneous
determination of mefenamic acid and paracetamol in their combination
formulations. In the ratio spectra derivative method, analytical signals
were measured at the wavelengths corresponding to either maxima or
minima for both the drugs in the first derivative spectra of the ratio
spectra obtained by dividing the standard spectrum of one of the two
drugs in a mixture of 0.1 M NaOH and methanol (1:9). In three
chemometric methods {classical least-squares, inverse least-squares, and
principal component regression (PCR)}, the conduct of procedures was
randomly selected by using the different mixture compositions containing
two drugs in a mixture of 0.1 M NaOH and methanol (1:9). The
absorbance data was obtained by the measurement at 13 points in the
wavelength range of 235–355 nm. Chemometric calibration was con-
structed from the absorbance data and the training set for the prediction
of the amount of mefenamic acid and paracetamol in samples. In
the PCR method, the covariance matrix corresponding to the absor-
bance data was calculated for the basis vectors and matrix containing
the new coordinates. The obtained calibration curve was used to
determine the drugs in their mixtures. Linearity range in all the
methods was found over the range of 2–10 mg/mL for mefenamic acid
and 4–20 mg/mL for paracetamol. Mean recoveries were found to be
satisfactory (greater than 99%). The procedures do not require any
separation step, and were successfully applied to a pharmaceutical
formulation (tablet).
5.2 Colorimetry
Deveaux et al. [21] reported that mefenamic acid, being an N-aryl
derivative of anthranilic acid, can be characterized by two color
reactions. The color reactions, negative with N-aryl derivatives of
aminonicotinic acid, are associated with the diphenylamine structure.
For the first color reaction, add to a test tube approximately 0.5 g of
oxalic acid dihydrate and at least 1 mg of the test material. Place the tube
into an oil bath at 180–200 C for 4–5 min. After cooling, dissolve the
residue in 10 mL of 95% ethanol to obtain a stable, intense blue solution
(absorption maximum at 586–590 nm). To use the reaction for capsule
formulations, extract the active ingredient with acetone and filter prior to
the assay. For the second color reaction, add mefenamic acid (100–800
mg) in 1 mL of HOAc–H2SO4 (98:2, v/v), 5 mL of HOAc–HCl (50:50, v/v)
and 1 mL of 0.10% (w/w) aqueous levulose. Heat the mixture for 25 min
at 100 C, and after cooling, measure the absorbance at 597 nm.
Vinnikova [22, 23] described the spectrophotometric determination of
mefenamic acid at 490 nm after conversion to its colored complex with
Fast Red B salt at pH 6.60 (phosphate buffer). The method was applied
for the determination of free and bound mefenamic acid, and found to be
useful for studying the blood plasma protein binding, absorption,
distribution, metabolism, and excretion of mefenamic acid.
Sastry and Rao [24] reported the determination of mefenamic acid and
some NSAIDs in pharmaceuticals by a spectrophotometric method. The
method is based on the formation of a blue complex with Folin-Ciocalteu
reagent, and subsequent measurement of the absorbance at 750–760 nm.
The method was found to be simple, sensitive, accurate, and rapid,
with recoveries of 98.0–102.3% and relative standard deviations of
1.04–1.66%.
Sastry et al. [25] also described a fairly sensitive spectrophotometric
method for the determination of mefenamic acid based on the formation
of a chloroform-soluble, colored ion-association complex between
mefenamic acid and methylene violet at pH 7.6. The absorbance of the
separated chloroform layer was measured at 540 nm. The molar
absorptivity is 2.29  104 L/mol  cm, and the limit of detection is 2–
16 mg mefenamic acid/mL. The method was applied successfully to the
determination of mefenamic acid in bulk samples and in pharmaceutical
preparations, with recoveries of 99.2–99.9%.
Another report by Sastry's group [26] described the determination of
mefenamic acid based on the reaction of the mefenamic acid with p-N,N-
dimethylphenylenediamine in the presence of S2O82or Cr(VI), whereby
an intensely colored product having maximum absorbance at 740 nm is
developed. The molar absorptivity is 4.15  104 L/mol cm. The reaction
is sensitive enough to permit the determination of 0.25–4.0 mg mefenamic
acid/mL, and was successfully applied to the determination of mefenamic
acid in bulk samples and in pharmaceutical preparations, with recoveries
of 99.4–99.8%.
Mahrous et al. [27] described the colorimetric determination of
mefenamic acid with potassium ferricyanide in NaOH medium. The
orange product is measured at 464 nm, and the molar absorptivity is
1.9  103 L/mol cm. The method was applied successfully to the
determination of mefenamic acid in capsules. Garcia et al. [28] reported
a flow injection spectrophotometric method for the determination of
mefenamic acid in bulk samples and pharmaceuticals, also based on the
reaction of mefenamic acid with potassium ferricyanide in NaOH media.
The absorbance of the product obtained was measured at 465 nm, and
the corresponding calibration graph was linear over the range of 1.00–
100 mg/L, with a limit of detection of 0.18 mg/L.
Hassib et al. [29] determined mefenamic acid by ferric hydroxamate
complex formation and measurement of the absorbance of the colored
complex at 530 nm. The method is applicable to mefenamic acid amounts
varying from 0.5 to 74.5 mg/25 mL, and PonstanÕ capsules were
successfully determined using this method.
Khier et al. [30] determined mefenamic acid after complexation with
copper(II) amine sulfate. The complex is extracted with chloroform and
treated with diethyldithiocarbamate solution, whereupon another copper
(II) complex (wavelength maximum of 430 nm) is formed. Beer's law is
followed over the mefenamic acid concentration range of 6–48 mg/mL.
The method was applied successfully to the determination of mefenamic
of 98.0–101.0%.
Abdel-Hay et al. [31] described the determination of mefenamic acid
based on its reaction in ethanolic medium with 2-nitrophenylhydrazine in
the presence of dicyclohexylcarbodiimide to give an acid hydrazine,
which showed intense violet color (maximum absorption at around 550
nm). The effect of reagent concentration (2-nitrophenylhydrazine-HCl,
pyridine, and dicyclohexylcarbodiimide), heating temperature, and
heating time were studied to optimize the reaction conditions. The
method was successfully applied to the determination of mefenamic
acid in pure and dosage forms, with a relative standard deviation less
than 2%.
Bojarowicz and Zommer-Urbanska [32] reported that the ferric
complexes of mefenamic acid (stoichiometries of 1:1, 1:2, 1:3, and 1:4,
depending on the amount of the ligand used) were formed in 90%
aqueous methanol at pH 2.8. All showed absorbance at 540–550 nm.
A simple photometric method in the visible region was described by Babu
et al. [33] for the determination of mefenamic acid in its pharmaceutical
dosage forms. The method is based on the reaction of the drug with
4-aminophenazone and potassium ferricyanide to yield a reddish-green
colored chromophore, which exhibits an absorption maximum at 590 nm.
The chromophore is stable for 40 min, and Beer's law is obeyed over the
concentration range of 0.5–4 mg/mL.
Amin et al. [34] reported the determination of mefenamic acid based on
the formation of the blue complex obtained from the reaction of
methylene blue with mefenamic acid. The method was applied for the
analysis of pharmaceutical preparations containing mefenamic acid, and
found to be simple, precise and reproducible.
The nitrosation conditions of aniline and diphenylamine derivatives
(paracetamol, phenacetin, mefenamic acid, and diclofenac sodium), with
subsequent formation of colored products in alkaline media, were studied
by Shormanov et al. [35]. Suitable procedures for quantitative
determination of analytes in bulk drug substance and tablets were
5.3 Fluorimetry
The influence of sample pretreatment by alkaline saponification and
oxidation by K2Cr2O7 in acetic acid, and the influence of the solvent and
addition of haloacetic acids (e.g., trichloroacetic acid required for
fluorescence), on the fluorescence intensity of mefenamic acid and other
N-arylanthranilic acids was studied by Dell and Kutschbach [36] with
reference to their fluorimetric determination in urine. Hydrolysis is
necessary for determining esters and amides of mefenamic acid. Polar
solvents decrease the fluorescence intensity, and solvents with a negligible
dielectric constant (especially CCl4, Cl2C ¼ CCl2, and decalin) were
required to render the solution fluorescent. Among the haloacetic acids
studied, only acids with pK values less than 1 were found to be suitable
(i.e., Cl3CCO2H and F3CCO2H). Aryl substitution affected the position
of the fluorescence maximum but not the intensity. Anthranilic acid was
found to fluoresce in CCl4–Cl3CCO2H, whereas its meta and para
analogs and Ph2NH were not.
Deveaux et al. [21] explained the fluorescent reaction of mefenamic acid
by the formation of a substituted acridone after dissolving mefenamic
acid in concentrated H2SO4 and heating for 10 min at 100 C. The
acridone exhibits an intense green fluorescence when excited by white
light, and blue when excited by ultraviolet light.
In contrast to the above Deveaux method, Mehta and Schulman [37]
described that the native fluorescence of mefenamic, flufenamic and
meclofenamic acids is more useful for determination of these drugs
compared to the fluorescence of the derivatives-substituted acridones and
benzoxazines of these drugs by treatment with H2SO4 and HCHO,
respectively. The determination of mefenamic acid at trace level by
fluorescence spectrometry was also reported by Miller et al. [38].
Huang and Xu [39] developed a fluorimetric method for analysis of some
acid drugs by using pyronine to form a drug-dye complex. In Na2HPO4
solution, mefenamic acid reacts with pyronine to form a complex which is
soluble in CHCl3. Pyronine fluoresces strongly at an excitation
wavelength of 542 nm, and emits at an emission wavelength of 566 nm.
The intensity of pyronine in the complex is proportional to the amount of
mefenamic acid in the complex. The method sensitivity was found to be
10–50 ng/mL, and good recovery and precision were obtained.
Based on the formation of the complex of the compound with Al(III) in
an ethanolic medium, a sensitive and rapid flow injection fluorimetric
method was proposed by Albero et al. [40] for the determination of
mefenamic acid. The calibration graph resulting from the measurement
of the fluorescence after excitation of the complex with mefenamic acid at
355 nm (emission wavelength of 454 nm) was linear over the range of
0.30–16.1 mg/mL. The method has been applied to the determination of
the substance in pharmaceutical preparations.
Ioannou et al. [41] reported the use of terbium sensitized fluorescence to
develop a sensitive and simple fluorimetric method for the determination
of the anthranilic acid derivative, mefenamic acid. The method makes use
of radiative energy transfer from anthranilate to Tb(III) in alkaline
methanolic solutions. Optimum conditions for the formation of the
anthranilate-Tb(III) complex were investigated. Under optimized condi-
tions, the detection limit was 1.4  108 mol/L, and the range of
application was 2.5  108 to 5.0  105 mol/L. The method was
successfully applied to the determination of mefenamic acid in serum
after extraction of the sample with ethyl acetate, evaporation of the
organic layer under a stream of nitrogen at 40 C, and reconstitution of
the residue with alkaline methanolic terbium solution prior to instru-
mental measurement. The mean recovery from serum samples spiked
with mefenamic acid (3.0  106, 9.0  106, 3.0  105 mol/L) was
101  5%. The within-run precision (RSD) for the method for the two
serum samples varied from 2 to 8%, and the day-to-day precision for two
concentration levels varied from 2 to 13%.
The use of second-derivative synchronous fluorescence spectrometry was
reported by Ruiz et al. [42] to develop a simple, rapid and sensitive
fluorimetric method for the determination of binary mixtures of the
nonsteroidal antiinflammatory drugs flufenamic (FFA), meclofenamic
(MCFA) and mefenamic (MFA) acids in serum and in pharmaceutical
formulations. The method is based on the intrinsic fluorescence of these
compounds in chloroform. A differential wavelength of 105 nm was used
for the resolution of FFA–MFA and MFA–MCFA mixtures, whereas
the FFA–MCFA mixture was determined at a differential wavelength of
40 nm. Serum samples were treated with trichloroacetic acid to remove
the proteins, and the analytes were extracted in chloroform prior to
determination. Pharmaceutical preparations were analyzed without prior
separation steps.
Capitán-Vallvey et al. [43] has developed a spectrofluorimetric method
for the quantitative determination of flufenamic, mefenamic and
meclofenamic acids in mixtures by recording emission fluorescence
spectra between 370 and 550 nm with an excitation wavelength of 352 nm.
The excitation-emission spectra of these compounds are deeply over-
lapped which does not allow their direct determination without
previous separation. The proposed method applies partial least squares
multivariate calibration to the resolution of this mixture using a set of
wavelengths previously selected by Kohonen artificial neural networks.
The linear calibration graphs used to construct the calibration matrix
were selected in the ranges from 0.25 to 1.00 mg/mL for flufenamic and
meclofenamic acids, and from 1.00 to 4.00 mg/mL for mefenamic acid. A
cross-validation procedure was used to select the number of factors. The
selected calibration model has been applied to the determination of these
compounds in the synthetic mixtures and pharmaceutical formulations.
Sabry [44] reported that the fluorescence properties of flufenamic and
mefenamic acids were obtained in aqueous and binary aqueous-organic
solvents with or without a- and b-cyclodextrins (CDs). The spectral
behavior of the model compounds in aqueous micellar systems of ionic
and non-ionic surfactants were investigated. In aqueous acidic solution,
the investigated analytes were better incorporated in CDs (flufenamic
acid) and micelles (flufenamic and mefenamic acids). The luminescence
emission from the analytes were found to be greatly enhanced by
surfactants. The fluorescence of the analyte–surfactant system was
quenched by increasing amounts of organic solvents. The changes in
the dissociation constant of flufenamic and mefenamic acids in solutions
of anionic and nonionic surfactants (sodium dodecyl sulfate and Triton
X-100, respectively) were studied spectrofluorimetrically. The fluores-
cence intensity enhancements in CDs media relative to aqueous solution
ranged from 1.3 to 1.6. Sensitive micellar-enhanced spectrofluorimetric
methods for the determination of flufenamic and mefenamic acids were
developed, with ranges of application down to ng/mL. The linear
dynamic ranges and relative standard deviations were reported. The
methods were applied satisfactorily to the determination of both the
compounds in pharmaceutical preparations with recoveries of 98–101%.
5.4 Chemiluminescence
Aly et al. [45] has investigated a novel chemiluminescent method using
flow injection for the rapid and sensitive determination of flufenamic and
mefenamic acids. The method is based on a tris(2,20 -bipyridyl)ruthenium
(III) chemiluminescence reaction. Ru(bipy)33 þ is chemically generated by
mixing two streams containing solution of tris(2,20 -bipyridyl)ruthenium
(II) and acidic cerium (IV) sulfate. After selecting the best operating
parameters calibration graphs were obtained over the concentration
ranges 0.07–6.0 and 0.05–6.0 mg/mL for flufenamic and mefenamic acids,
respectively. The limits of detection (s/n ¼ 3) were 3.6  109 M
flufenamic acid and 2.1  107 M mefenamic acid. The method was
successfully applied to the determination of these compounds in
pharmaceutical formulations and biological fluids.
5.5 Proton magnetic resonance spectrometry
Mansour et al. [46] developed a simple, rapid efficient quantitative proton
magnetic resonance method for the determination of mefenamic acid in
PonstanÕ capsules. The method is based on a comparison between the
sum integrals of the two methyl singlets of mefenamic acid and that
integral of the sharp singlet of p-methoxybenzylidenemalononitrile
(internal standard). The method gave accurate and reproducible results.
De Kanel et al. [47] described a method for the simultaneous analysis of
14 nonsteroidal antiinflammatory drugs in human serum using negative
electrospray ionization-tandem mass spectrometry. After addition of
internal standard and protein precipitation using acetonitrile, samples
were transferred to autosampler vials for direct analysis without
chromatography. Injection of an air bubble with the sample and a
multiple reaction monitoring method using argon collision-induced
dissociation of analyte (M–H) ions permitted integration of the product
ion peak areas to produce reproducible quantitative data over the range
of concentrations expected in serum during routine use of these drugs.
The method permitted the analysis of 30 samples per hour. Two hundred
and fifty consecutive analyses did not adversely affect instrument
sensitivity.
5.7 Atomic absorption spectrometry
Salem et al. [48] reported simple and accurate methods for the
quantitative determination of flufenamic, mefenamic and tranexamic
acids utilizing precipitation reactions with cobalt, cadmium and
manganese. The acidic drugs were precipitated from their neutral
alcoholic solutions with cobalt sulfate, cadmium nitrate or manganese
chloride standard solutions followed by direct determination of the ions
in the precipitate or indirect determination of the ions in the filtrate by
atomic absorption spectroscopy (AAS). The optimum conditions for
precipitation were carefully studied. The molar ratio of the reactants was
ascertained. Statistical analysis of the results compared to the results of
the official methods revealed equal precision and accuracy. The suggested
procedures were applied for determining flufenamic, mefenamic and
tranexamic acids in pharmaceutical preparations as well and proved
validity.
The similar method using nickel (II) instead of cobalt, cadmium and
manganese was also investigated by Salem et al. [49] for the quantitative
determination of flufenamic, mefenamic and tranexamic acids, furose-
mide, diclofenac sodium and thiaprofenic acid. Statistical analysis of the
results compared to assays used in pharmacopoeias and the Amax
methods revealed equal precision and accuracy. Furthermore, the assays
were also applied for the determination of these drugs in pharmaceutical
Alpdogan and Sungur [50] developed an indirect atomic absorption
spectroscopy method for the determination of mefenamic and flufenamic
acids, and diclofenac sodium, based on the complexation with copper (II)
amine sulfate. The complex was extracted into chloroform, and the
concentrations of substances were determined indirectly by AAS
measurement of copper after re-extraction into 0.3 N nitric acid solution.
The developed method was applied to the assay of the substances in
commercial tablet formulations. The results were statistically compared
with those obtained by HPLC method by t- and F tests at 95%
confidence level. Calculated t and F values were both lower than the
table values.
Khalil et al. [51] described the microquantitative determination of
mefenamic acid based on the reaction of mefenamic acid with a silver
nitrate solution in a neutral alcoholic medium. The formed precipitation
is quantitatively determined directly or indirectly through the silver
content of the precipitation formed or the residual unreacted silver ions in
the filtrate by atomic absorption spectrophotometry. The results obtained
in both the procedures either in their pure form or in their pharmaceutical
formulations are accurate and precise. The stoichiometric relationship of
the reaction was studied using Job's continuous variation method, and it
was found to be (1:1) drug:Ag þ for the mefenamic acid.
The detection of mefenamic acid and its metabolites {N-(2-methyl-3-
carboxyphenyl)anthranilic acid and N-(2-methyl-3-hydroxymethyl-phe-
nyl)anthranilic acid} in urine by thin layer chromatography was reported
by Demetriou and Osborne [52]. Plates were prepared from a slurry of
silica gel G (30 g) in water (65 mL). Layers of thickness 250 m were
prepared and dried at 110 C for 30 min. For the extraction procedure,
urine (10 mL) was treated with sodium hydroxide (10 M, 1 mL) for 20 min
at room temperature. The sample was then acidified with concentrated
hydrochloric acid, and extracted with petroleum ether (b.p. 40–60 C;
10 mL) for 10 min on a mechanical shaker. The organic layer was
separated, and placed in a 10 mL conical tube, then the solvent was
removed. The extract was reconstituted with methanol (100 mL) and a 30-
mL aliquot of the solution was used for spotting, along with 10-mL aliquot
of the reference solution (mefenamic acid in methanol, 1 mg/1 mL). The
solvent system used for the development of the chromatogram was
toluene–acetic acid (9:1) providing Rf value of 0.75 for mefenamic acid.
Sodium nitrite (1% in 1% sulfuric acid) was used as the detection reagent
to give a green color of mefenamic acid.
By et al. [53] described the identification and chromatographic
determination of some undeclared medicinal ingredients of traditional
oriental medicine. The qualitative examination of indomethacin, mefe-
namic acid, diazepam and hydrochlorothiazide was performed on silica
gel using solvent system of ethyl acetate (system A) and ethyl acetate–
methanol (4:1, v/v) (system B). The spots were identified using short-wave
UV light to give Rf values of 0.42 and 0.50 for mefenamic acid in solvent
system A and B, respectively. The utilization of the TLC method was also
reported by Dharmananda [54] for the investigation of imported Chinese
herb products adulterated with drugs.
Argekar and Sawant [55] developed simultaneous determination of
paracetamol and mefenamic acid in tablets by high performance thin
layer chromatography method (HPTLC). The procedure was performed
on silica gel using toluene–acetone–methanol (8:1:1) as the solvent
system. Detection and quantification of the substance was assayed by
densitometry at 263 nm. The linearity ranges were 120–360 ng/mL; and
the RSD were between 0.41 and 0.78%. Limit of detection and limit of
quantification were found to be 1.80 and 5.5 ng/mL, respectively.
Recoveries were in the range of 95.50–103.60%.
6.2 Column chromatography
Adams [56] reported ion-pairs partition chromatography of mefenamic
acid with tetraalkylammonium cations as an effort to develop an
analytical method from the extraction data. Ion pairs of mefenamic
acid with Me4NOH, Et4NOH, Pr4NOH and Bu4NOH were formed and
extracted from aqueous solutions with CHCl3 or mixtures of CHCl3 and
isooctane. The ion-pair extraction constant (KQA) increased by a factor of
 500,000 from the methyl to butyl homologs, and the plot of KQA vs. C
number was linear. The distribution ratio (D) of the ion-pairs between
aqueous and organic phases was affected by solvent polarity, which was
varied by adding up to 60% isooctane to the CHCl3. Chromatography
requires D values <0.01 for retention and >0.06 for elution. For
chromatography, Celite 545 is mixed with Pr4NOH (the best ion-pairing
reagent), placed in a column, covered with an alkaline solution of ion-
paired with excess Pr4NOH and containing more Celite 545. The column
is washed with 33% CHCl3 in isooctane to reduce the distribution ratio
to <0.01, and pure CHCl3 is used to elute the ion-pair, which is collected
in HCl–MeOH to break the ion-pair. Mefenamic acid is then detected by
spectrometry in CHCl3 at 353 nm. The average recovery from standard
solutions was 99.01%, and results with capsule solutions were
Bland et al. [57] reported the determination of mefenamic acid in blood
and urine of the horse. To 2.0 mL of biological fluids were added 1.0 mL
of an aqueous solution of saturated KH2PO4 and 2.0 mL of CH2Cl2. The
drug was extracted into CH2Cl2 phase, transferred to a clean culture
tube, and evaporated to dryness under a stream of nitrogen. To the
residue was added 50 mL of pentafluoropropionic anhydride and 50 mL
of ethyl acetate saturated with ZnCl2 which acts as a Lewis acid. The
tube was capped and placed in a water bath at 60 C for 5 min to
complete the derivatization. Following derivatization, 1.0 mL of
cyclohexane was added to the residue and the solvent was washed
vigorously with 8.0 mL of a saturated sodium tetraborate solution. After
centrifugation, the derivative in the cyclohexane phase was then analyzed
using gas-liquid chromatograph which was a Beckman GC-5, equipped
with a Beckman nonradioactive electron capture detector. The column
was a 4 ft, 2 mm i.d. glass column packed with 3% OV-1 on 80/100 mesh
Chromosorb GHP provided by Supelco, Inc. The instrument was
operated isothermally at an oven temperature of 190 C. The samples
were injected onto the column, at an inlet temperature of 215 C.
Detector compartment temperature was 290 C. The carrier gas (He) flow
was 40 mL/min. The drug derivative eluted in 3 min 43 s under these
conditions. As little as 50 ng/mL of mefenamic acid can be quantitated
with this procedure.
Gas-liquid chromatographic determination of mefenamic acid in human
serum was established by Dusci and Hackett [58]. A 2 mL sample of
serum was acidified with 0.5 mL of 1 M hydrochloric acid and extracted
with 15 mL of CH2Cl2 by shaking for 5 min in a glass tube. The tube was
centrifuged and the aqueous phase aspirated from the surface. The
organic layer was filtered through a solvent-washed Whatman No. 54
filter paper. To a 12 mL aliquot of the solvent was added 50 mL of 1000
mg/L solution of 5-( p-tolyl)-5-phenylhydantoin in ethanol as the internal
standard, and the mixture taken to dryness at 50 C under a stream of
nitrogen. To the residue were added 80 mL N,N-dimethylacetamide, 10 mL
tetramethylammonium hydroxide and 20 mL 1-iodobutane. The contents
of the tube were mixed after each addition and left to stand for 5 min.
The sample was centrifuged and an aliquot of 2.0 mL of the supernatant
injected to a Pye GCD gas chromatograph equipped with a flame
ionization detector. The column was a 1.5 m  4 mm i.d. glass
tube packed with 3% SP 2250 DA on Supelcoport 100–200 mesh. The
instrument settings were as follows; injection temperature 285 C; detector
temperature 290 C; carrier gas flow-rate, 60 mL/min; hydrogen flow-rate,
240 mL/min. Under these conditions the retention time of mefenamic
acid and the internal standard were 1.8 min and 4.1 min, respectively. The
ratio of the peak height of mefenamic acid to that of the internal standard
was linear over a range of 0.05–0.8 mg of the drug.
Singh et al. [59] described a gas chromatographic method for the
simultaneous analysis of flunixin, naproxen, ethacrynic acid, indometha-
cin, phenylbutazone, mefenamic acid and thiosalicylic acid in horse
plasma and urine. For sample preparation from plasma, a 1-mL plasma
sample containing 0.1, 0.5, 1.0, 2, 2.5, 5.0 and 10 mg of various drugs
was mixed with 1 mL of 0.1 M hydrochloric acid and 10 mL of
dichloromethane. The mixture was rotoracked for 10 min and centrifuged
(1500 g) for 15 min, and the aqueous phase was aspirated from the
surface. The organic layer was collected into another tube and dried at
45 C under a stream of nitrogen. The dried residue was derivatized by
mixing with 10 mL of bis(trimethylsilyl)trifluoroacetamide (BSTFA)
before analysis. Mefenamic acid and indomethacin was used as the
internal standard for the quantitative analysis. For sample preparation
from urine, 1–2 mL of urine (control or NaOH-hydrolyzed) were
acidified to pH 3 with saturated phosphate buffer and extracted with
10 mL of dichloromethane. The dichloromethane layer was separated by
centrifugation at 10,000 g, and dried at 45 C under nitrogen. The dried
residue was derivatized by adding 10 mL of BSTFA to the sample, and
1 mL of the derivatized residue was injected into the GC–MS system.
GC–MS analysis was performed by using an Econocap capillary column,
SE-54 (30 m  0.25 mm i.d.). The oven temperature was programmed at
20 C/min from an initial temperature of 150 C to a final temperature of
280 C; the run time was 15 min. The injector temperature was 250 C and
the injection mode was splitless. For selected-ion monitoring (SIM), three
ions were selected for each drug. The recovery of each drug from plasma
was ca. 95%, and the assay demonstrated good precision. The accuracy
of the assay was best at the 10.0 mg/mL level.
Giachetti et al. [60] compared the performance of mass selective detector
(MSD), electron capture detector (ECD) and nitrogen–phosphorus
detector (NPD) of gas chromatography systems in the assay of six non-
steroidal antiinflammatory drugs in the plasma samples. As a practical
test, six NSAIDs (mefenamic, flufenamic, meclofenamic and niflumic
acids, diclofenac and clonixin) added to plasma samples were detected
and quantified. The analyses were carried out after solvent extraction
from an acidic medium and subsequent methylation. The linearity of
response was tested for all the detection systems in the range of
1–25 ng/mL. Precision and accuracy were detected at 1, 5 and 10 ng/mL.
The minimum quantifiable level for the six drugs was about 1 ng/mL with
each of the three detection systems.
Gonzáles et al. [61] described a gas chromatographic–mass spectrometric
(GC–MS) procedure for the detection of seventeen nonsteroidal anti-
inflammatory drugs (including mefenamic acid) in equine plasma and
urine samples. The extraction of the compounds from the biological
matrix was performed at acidic pH (2–3) with diethyl ether. Ethereal
extracts were washed with a saturated solution of sodium hydrogencar-
bonate (urine) or treated with a solid mixture of sodium carbonate and
sodium hydrogencarbonate (plasma). The ethereal extracts were dried
and derivatized by incubation at 60 C with methyl iodide in acetone in
the presence of solid potassium carbonate. Mono- or bismethyl
derivatives of the NSAIDs were obtained. After derivatization kinetic
studies, 90 min was the incubation time finally chosen for screening
purposes for adequate methylation of all the compounds under study.
For individual confirmation analyses, shorter incubation times can be
used. The chromatographic analysis of the derivatives was accomplished
by a Model 5890A Series II gas chromatograph coupled to a Model 5970
electron impact mass-selective detector via a direct capillary interface
(Hewlett-Packard). The column was a 25 m  0.2 mm i.d. fused silica
cross-linked methylsilicone with a 0.11-mm film thickness (Hewlett-
Packard). Helium was used as the carrier gas at 0.65 mL/min. The
injection port and detector temperatures were 280 C. The oven
temperature was increased from 100 to 200 C at 25 C/min and then to
300 C at 15 C/min, with a final hold time of 2.3 min; the total run time
was 13 min. The injection volume was 2 mL and a splitting ratio of 10:1
was used. In general, extraction recoveries ranged from 23.3 to 100% in
plasma and from 37.5 to 83.8% in urine samples. Detection limits from
less than 5 to 25 ng/mL were obtained for both plasma and urine samples
using selected-ion monitoring. The procedure was applied to the
screening and confirmation of NSAIDs in routine doping control of
equine samples.
Maurer et al. [62] developed a gas chromatographic–mass spectrometric
(GC–MS) screening procedure for the detection of NSAIDs in urine as
part of a systematic toxicological analysis procedure for acidic drugs and
poisons after extractive methylation. The compounds were separated by
capillary GC and identified by computerized MS in the full-scan mode.
Using mass chromatography with the ions m/z 119, 135, 139, 152, 165,
229, 244, 266, 272, and 326, the possible presence of NSAIDs and their
metabolites could be indicated. The identity of positive signals in such
mass chromatograms was confirmed by comparison of the peaks
underlying full mass spectra with the reference spectra recorded during
this study. The method allowed the detection of therapeutic concentra-
tions of acemetacin, acetaminophen (paracetamol), acetylsalicylic acid,
diclofenac, diflunisal, etodolac, fenbufen, fenoprofen, flufenamic acid,
flurbiprofen, ibuprofen, indomethacin, kebuzone, ketoprofen, lonazolac,
meclofenamic acid, mefenamic acid, mofebutazone, naproxen, niflumic
acid, phenylbutazone, suxibuzone, tiaprofenic acid, tolfenamic acid, and
tolmetin in urine samples. The overall recoveries of the different NSAIDs
ranged between 50 and 80% with coefficients of variation of less than
15% (n ¼ 5), and the limit of detection of the different NSAIDs were
between 10 and 50 ng/mL (s/n ¼ 3) in full-scan mode. Extractive
methylation has also been used for plasma analysis.
Takeda et al. [63] described a systematic analysis of acid, neutral and
basic drugs in horse plasma by a combination of solid-phase extraction,
nonaqueous partitioning and gas-chromatography–mass spectrometry.
Sample preparation method started by using a 6 mL column packed with
1 g of Bond Elut Certify for the extraction of 4 mL of horse plasma.
Fractionation is performed with 6 mL of CHCl3–Me2CO (8:2) and 5 mL
of 1% TEA–MeOH according to its property. Simple and effective
clean-up based on nonaqueous partitioning is adopted to remove
co-eluted contaminants in both acid and basic fractions. Two kinds of
1-(N,N-diisopropylamino)-n-alkanes are co-injected with the samples
into the GC–MS system for the calculation of the retention index. Total
recoveries of 107 drugs (including mefenamic acid) are examined. This
procedure achieves sufficient recoveries and clean extracts for GC–MS.
This method is able to detect ng/mL drug levels in horse plasma.
Lo et al. [64] reported acidic and neutral drugs screen in blood with
quantitation using capillary gas chromatography–ion flame detector.
Blood previously acidified with aqueous saturated ammonium chloride
solution was extracted with ethyl acetate. The dried extract was subjected
to acetonitrile–hexane partition. The acetonitrile portion was analyzed
for the presence of acidic and neutral drugs by GC–FID (25 m narrow-
bore  0.25 mm i.d. HP-5 column with 0.33 mm film thickness). The
protocol was found to be suitable for both clinical toxicology (including
emergency toxicology) and postmortem toxicology. Quantitation was
facilitated by incorporating calibration blood standards in each assay
batch. The five drugs most commonly encountered in clinical blood
specimens (1150 cases) were: paracetamol (47.4% of the cases),
chlormezanone (6.6%), theophylline (1.74%), naproxen (1.65%) and
mefenamic acid (1.56%).
Arrizabalaga et al. [65] reported the determination of the composition of a
fraudulent drug by coupled chromatographic methods (GC–MS,
GC–FTIR, HPLC–DAD). The Chinese pills, Chuifong Toukuwan, as
traditional herbal medicine claimed to have a natural composition, was
studied by these appropriate methods. The analysis of crude and
derivatized extracts of the pills showed that they comprise a mixture of
synthetic drugs (mefenamic acid, indomethacin, hydrochlorothiazide,
and diazepam) and some herbal compounds. A similar procedure was
also reported by Fraser et al. [66] for the determination of undeclared
prescription drugs in methanol–ether extracts of Black Pearl products, a
type of Chinese herbal preparation generally marketed to American
consumers suffering from arthritis and back pain. They detected the
following prescription drugs: diazepam, diclofenac, hydrochlorothiazide,
indomethacin and mefenamic acid.
As mentioned above, a compendial assay using high performance liquid
chromatography was described by the USP 26 and the Indonesian
Pharmacopoeia [1, 4].
The separation of samples that contain more than 15–20 analytes (n>15–
20) is typically difficult and usually requires gradient solution.
Considering this problem, Dolan et al. [67, 68] reported reversed-phase
liquid chromatographic (RP–LC) separation of complex samples by
optimizing temperature and gradient time. They examined the reversed-
phase liquid chromatographic separation of 24 analytes with 8  n  48
(n ¼ the sum of analytes) as a function of temperature T and gradient
time tG. The required peak capacity was determined for each sample,
after selecting T and tG for optimum selectivity and maximum sample
resolution. A comparison of these results with estimates of the maximum
possible peak capacity in reversed-phase gradient elution was used to
quantify the maximum value of ''n'' for some required sample resolution
(when T and tG have been optimized). These results were also compared
with literature studies of similar isocratic separations as a function of
ternary-solvent mobile phase composition, where the proportions of
methanol, tetrahydrofuran and water were varied simultaneously. This
turn provides information on the relative effectiveness of these two
different method development procedures (optimization of T and tG vs.
%methanol and %tetrahydrofuran) for changing selectivity and achiev-
ing maximum resolution. The chromatographic conditions: 0–100%
acetonitrile in buffer gradient in 40 min, temperature 50 C, 25  0.46 cm
column, 2.0 mL/min. This observation is surprising, since the ability of
either T or tG to vary band spacing and maximize separation is much less
than observed for changes in solvent type (e.g., varying the ratio of
methanol to tetrahydrofuran). An alternative approach is to carry out
two separations with different conditions (T, tG) in each run. The
combination of results from these two runs then allows the total analysis
of the sample, providing that every sample component is adequately
resolved in one run or the other. One of the compounds having the
poorest-resolved peak was mefenamic acid (Rs ¼ 1.7, tG ¼ 103 min,
T ¼ 74 C).
Ahrer et al. [69] developed methods for the determination of drug
residues in water based on the combination of liquid chromatography
(LC) or capillary electrophoresis (CE) with mass spectrometry (MS). A
2 mM ammonium acetate at pH 5.5 and a methanol gradient was used
for the HPLC–MS allowing the separation of a number of drugs such as
paracetamol, clofibric acid, penicillin V, naproxen, bezafibrate, carba-
mazepin, diclofenac, ibuprofen, and mefenamic acid. Apart from the
analytical separation technique, water samples have to be pretreated in
order to get rid of the matrix components and to enrich the analytes; the
usual way to accomplish this aim is to perform a solid-phase extraction
(SPE) step employing suitable stationary phase (reversed-phase materi-
als) and conditions. Using the optimized SPE procedure, a 500 mL
volume of water sample was brought to pH 2 with concentrated
hydrochloric acid and passed through an SPE cartridge (6 mL) packed
with 500 mg Bondesil ODS 40 mm (Varian) conditioned with acetone,
methanol and water, pH 2 (one cartridge volume each). The flow-rate was
adjusted to approximately 10 mL/min. After the cartridges had been
allowed to dry for 30 min, the drugs were eluted using an overall volume
of 2 mL methanol. The extract was brought to dryness in a nitrogen
stream (purity 4.6) and finally redissolved in 50 mL methanol and diluted
with 300 mL water which was then used for injection (100 mL injection
volume). Liquid chromatography separations were performed on a HP
1100 HPLC system equipped with a HP 1050 autosampler (Agilent)
employing a YMC ODS-AM column 250  2 mm i.d. (YMC). Using a
flow-rate of 150 mL/min the optimized mobile phase was a ternary
gradient of a 2 mM ammonium acetate solution adjusted to pH 5.5 with
1 M acetic acid (A), methanol (B) and water (C) in the following form:
0 min: A–B–C (10:15:75); 3 min: A–B–C (10:15:75); 10 min: A–B–C
(10:50:40); 20 min: A–B–C (10:90:0). MS detection was performed on a
quadrupole system HP 5989B (Agilent) equipped with a radiofrequency-
only hexapole (Analytica of Branford) using either a pneumatically
assisted electrospray ionization (ESI) interface HP 59987A (Agilent) or
an atmospheric pressure chemical ionization (APCI) interface (Analytica
of Branford) for the combination with LC instrument. HPLC–MS is a
powerful technique for the determination of drug residues in water
offering both a high selectivity and good detection limits between 0.05
and 1 mg/L in the solution injected; with a sample pretreatment procedure
based on SPE before LC separation, detection limits in the sample of
about 1 ng/L and below can be achieved. The applicability of the
technique could be demonstrated for the analysis of several river
water samples; concentrations between approximately 2 and 130 ng/L
of bezafibrate, carbamazepine, diclofenac, and mefenamic acid were
Ku et al. [70] used HPLC with photodiode array detector (DAD) for the
identification of adulterants in rheumatic and analgesic Chinese
medicine. A study was performed by using a gradient elution with
acetonitrile and 0.1% acetic acid solution on a C-18 column. The method
was applied to simultaneously screen seventeen synthetic drugs adulter-
ated in traditional Chinese medicine within half an hour. The drugs
belonged to six pharmacological categories, namely antipyretic analge-
sics, glucocorticoids, diuretics, CNS stimulants, muscle relaxants
and sedatives. They included acetaminophen, aminopyrine, bucetin,
ethoxybenzamide, indomethacin, ketoprofen, mefenamic acid, phenyl-
butazone, piroxicam, salicylamide, dexamethazone, prednisolone, chlor-
mezanone, chlorzoxazone, hydrochlorothiazide, diazepam, and caffeine.
This method can provide higher resolution and greater efficiency than
thin-layer chromatography for screening adulterated synthetic drugs.
A reversed-phase HPLC post-column ion-pair extraction system was
developed by Kim and Stewart [71, 72] for the analysis of carboxylic acid
drugs and their salts (sodium formate, sodium acetate, 3-bromopropionic
acid, 6-aminocaproic acid, 11-bromoundecanoic acid, 1-heptanesulfonic
acid, p-nitrophenylacetic acid, sodium benzoate, sodium salicylate,
valproic acid, probenecid, naproxen, ketoprofen, ibuprofen, mefenamic
acid, flufenamic acid, and cefuroxime sodium) using a-(3,4-dimethoxy-
phenyl)-40 -trimethylammoniummethylcinnamonitrile methosulfate
(DTM) as a fluorescent ion-pair reagent. The HPLC system is composed
of three pumps; an Alcott Chromatography Model 760 HPLC pump for
the mobile phase, a LDC/Milton Roy mini-pump VS with pulse
dampener (SSI Model LP-21) for the ion-pair reagent, and a Kratos
Spectraflow 400 solvent delivery system with pulse dampener for the
extraction solvent; a Rheodyne Model 7125 injector equipped with a
50 mL sample loop, and two mixing tees (Alltech). The detector was a HP
Model 1046A programmable Fluorescence Detector (Hewlett-Packard)
set at an excitation of 355 nm and an emission of 460 nm (also equipped
with a 408 nm emission cut-off filter). The analytical column was a 5 mm
RP-8 column (Spheri-5, 10 cm  4.6 mm i.d.). The extraction coils were
three dimensional knitted teflon coils (1.8 m  0.3 mm i.d., LDC/Milton
Roy) knitted using a four pin ''Strickliesel''. The mobile phase was
prepared as an absolute methanol–pH 7 aqueous phosphate buffer
mixture (30:70, v/v) and the flow rate was set at 1 mL/min. An ion-pair
reagent concentration of 1.12  104 M (5 mg/100 mL of mobile phase)
was the most appropriate in order to minimize the contribution of
background noise in the system. A 30% n-pentanol in chloroform
was used as the extracting solvent. Sodium salicylate, ketoprofen,
ibuprofen, probenecid and valproic acid were used as model compounds
to evaluate the ion-pair extraction system. The method was applied to
pharmaceutical dosage forms containing ketoprofen and valproic acid.
Other acidic compounds evaluated using the ion-pair reagent showed
that lipophilic acids produced more extractable ion-pairs and higher
sensitivities than hydrophilic acids. Mefenamic acid exhibited the
capacity factor (k0 ) of 7.5039 and the limit of detection of 100 ng and
2 mg/mL (s/n ¼ 2).
Shinozuka et al. [91] developed a sensitive method for the determination
of four anthranilic acid derivatives (diclofenac sodium, aluminium
flufenamate, mefenamic and tolfenamic acids) by HPLC procedure.
The four drugs were converted into methylphthalimide (MPI) derivatives
in a constant yield by reaction with N-chloromethylphthalimide at 60 C
for 30 min. The production of the MPI derivatives were confirmed by
mass spectrometry. The MPI derivatives of the four drugs were separated
by HPLC using a C-18 bonded phase LiChrospher RP-18 column
(250  4 mm i.d.) with acetonitrile–water (80:20, v/v) as mobile phase.
The flow rate was 0.8 mL/min. The UV absorbance was measured at
282 nm. The calibration curves of the MPI derivatives of the drugs were
linear from 1.0 to 5.0 mg/mL. The detection limits of the four drugs were
0.5–5 ng. The extraction procedure for the four anthranilic acid
derivatives added in the plasma and urine was performed by using
Extrelut 1 column. Yields of column extraction of 100 mL of plasma and
urine samples (containing 0.5 mg of anthranilic acid derivatives) with 6 mL
of ethyl acetate were 84–106%.
Other methods for the HPLC analysis of mefenamic acid, as described by
several groups, are presented in Table 1.
6.5 Supercritical fluid chromatography
Jagota and Stewart [102] investigated super critical fluid chromatography
(SFC) for the separation of nonsteroidal antiinflammatory agents.
Chromatography was performed on a Lee Scientific Model 600D
supercritical fluid chromatograph equipped with a pump, oven and
flame-ionization detector and controlled by a Dell computer (ACI 600D,
software version 2.2). SFC was performed on three different stationary
phases: a 5 m  100 mm i.d. SB-methyl-100 (200 mm o.d. and 0.25 mm
flim thickness), 10 m  50 mm i.d. SB-biphenyl-30 and a SB-cyanopro-
pyl-50 (both 195 mm o.d. and 0.25 mm film thickness). Solutions of each
NSAID were prepared by accurately weighing 5 mg of each drug and
dissolving in 5 mL of absolute methanol to give a final concentration of
approximately 1 mg/mL. Chromatographic parameters were pump
program, multilinear pressure program: 7 min hold at an initial pressure
of 100 atm, then 25 atm/min ramp to 250 atm, followed by a 4.0 atm/min
ramp to 290 atm; oven program, isothermal at 130 C; injection type, time
split set at 200 ms, injection ratio approximately 20:1 giving an injection
volume of approximately 25 mL; detector, flame ionization at 375 C;
mobile phase, supercritical fluid chromatography grade carbon dioxide.
Baseline separation of a flufenamic acid, mefenamic acid, fenbufen and
HPLC Methods for Mefenamic Acid
Column Mobile phase Detection Detection Samples/ Ref.
limit matrices
Daisopak ODS column Acetonitrile–acetate buf- UV, 220 nm 11.5–75 Human 73
(SP-120-5-ODS-BP, fer (pH 3.5; 0.1 M)– ng/mL plasma
H. POERWONO ET AL.
250  4.6 mm i.d., methanol (35:40:25,
5 mm) v/v/v), 1 mL/min
Lichrospher 60 RP select 0.01 M Acetic acid pH 3 Photodiode- 0.016 mg/mL Bull plasma 74
B column (250  4 mm in methanol (35:65, array at 240, at 240 nm,
i.d., 5 mm) and v/v), 1 mL/min, 35 C 254, 284 nm 0.027 mg/mL
Lichrospher 60 RP at 254 nm
select B guard column
(4  4 mm i.d., 5 mm)
Discovery RP-Amide Acetonitrile–25 M UV, 230 nm NDa Solution 75
C16, 15 cm  4.6 mm, KH2PO4, pH 3.0
5 mm particles (40:60), 1 mL/min,
Wakosil ODS 5 1.9 g Tetra-pentylammo- UV, 230 nm 2 ng/mL Human urine, 76
C18column nium bromide in 1 L of pharmaceu-
(150  4.6 mm a mixture of acetic tical
i.d., 5 mm) acid–sodium acetate preparations
MEFENAMIC ACID: ANALYTICAL PROFILE
buffer solution, pH 5.0,
and acetonitrile (11:9,
v/v), 1.0 mL/min, 40 C
ODS II Inertsil Acetic acid–acetonitrile Photodiode- 0.05– Plasma 77
(250  4.6 mm, 5 mm) (80:20, 2 min; 36:64, array 64 ng/mL
column connected to 25 min; 80:20, 5 min), 240, 278,
pre-column ODS II 1 mL/min, 40 C 290 nm
Inertsil (8 mm, 5 mm)
Inertsil ODS-2 50 mM Phosphate buf- UV, 230 nm 0.05 mg/mL Human urine 78
(150  4.6 mm, fer–acetonitrile (58:42,
i.d., 5 mm) column v/v) at pH 5.0, 0.9 mL/
min, ambient tempera-
LiChrosorb-RP 18 6 mM Phosphoric acid– UV, 205 nm 0.05 mg/mL Human 79
column (250  4 mm acetonitrile (50:50), plasma
i.d., 7 mm) 2.0 mL/min, 50 C
Hypersil ODS NDa Diode-array NDa Blood 64
(200 mm  2.1 mm detection
i.d.) column
Lichrosorb RP 18 column Acetonitrile–water–0.1 M UV, 280 nm NDa Urine 80
(250  4 mm i.d., 7 mm) phosphoric acid
(55:42:3,v/v),
2.0 mL/min, 45 C
TSK Gel ODS-80 TM 10 mM Ammonium UV, 254 nm 0.05 mg/mL Urine 81
column (250  4.6 mm dihydrogen phosphate–
i.d., 5 mm) methanol (98:2, v/v) at
pH 5.0, 1.0 mL/min,
Nucleosil C18 analytical Methanol–water UV, 280 nm 0.1 mg/mL Rat plasma 82
column (5 mm) (77:23, v/v), 0.8 mL/
min, room temperature
Hypersil SI (100  2.0 Hexane–5% water in iso- Diode-array NDa Equine urine 83
mm i.d., 5 mm) column propanol (98:2, 70:30, UV, 280 nm
0:100, v/v), 0.4 mL/min,
Lichrosorb RP 18 column Methanol–water UV, 230, nm NDa Pharmaceuti- 84
(250  4.6 mm, 7 mm) (30:70,35:65,40:60, cal prepara-
45:55,50:50, tions
55:45,60:40,
65:35,70:30,75:25,80:20,
v/v) containing
1% acetic acid, 1.0
mL/min, room temp.
Vydac stainless-steel 10 mM Phosphoric acid– UV, 280 nm 0.08 mg/ml Human 85
analytical column acetonitrile (40:60, v/v), plasma
(250  4.6 mm i.d.), pH 2.6, 0.9 mL/min,
C18 bonded-phase ambient temperature
silica, 5 mm
Lichrosorb RP 18 0.065 M Ammonium UV, 282 nm 0.7 ng Blood serum, 86
(250  4 mm i.d., acetate–methanol urine,
10 mm) column (25:75, 30:70), 0.8 mL/ tablets, cap-
min, 22 C sules,
suppositor-
ies,
suspensions,
Lichrosorb RP 18 0.05 M Ammonium UV, 285 nm 1 pg Blood serum, 87
mm, 10 mm) column (78:22, 72:28), 0.81 mL/ tablets, sup-
min, 22 C positories,
Cyano column Water–acetonitrile– UV, 290 nm 0.05 mg/mL Human 88
(25  0.4 cm, 10 mm) methanol–17 M acetic plasma
acid (69:15:15:1, v/v)
Wakosil 5 C 18 Acetonitrile–water– UV, 280 nm NDa Pharmaceuti- 89
(150  4.6 mm) triethylamine cal
(60:40:0.1, v/v/v) preparation
adjusted with H3PO3 to
a pH of 3.0, 0.8
mL/min, 40 C
YMC ODS (70  20 mM pH 5.0 UV, 219 nm 0.1 ng/mL Human 90
4.6 mm i.d., 5 mm) Phosphate buffer–acet- serum
onitrile (65:35, v/v)
onitrile (55:45, v/v) 1.0
TSK Gel ODS 80 TM 20 mM pH 6.0 UV, 219 nm 0.1 ng/mL Human 90
(150  4.6 mm i.d., Phosphate buffer–acet- serum
5 mm) onitrile (55:45, v/v)
Supelcosil LC 8 0.05 M Phosphoric acid– Diode-array, 100–500 Plasma, urine 59
(7.5 cm  4.6 mm i.d., acetonitrile (55:45, v/v), 209–402 nm ng/mL
3 mm) 1.0 mL/min
Supelcosil LC 8 0.05 M Phosphoric acid– UV, 235 nm 50–150 ng/mL Plasma, urine 59
(7.5 cm  4.6 mm i.d., acetonitrile (55:45, v/v),
Supelcosil LC 8 0.05 M Phosphoric acid– Fluorescence 10 ng/mL Plasma, urine 59
(7.5 cm  4.6 mm i.d., acetonitrile (55:45, v/v), detection
3 mm) 1.0 mL/min (exc ¼
235 nm, em
¼ 405 nm
10 mm mBondapak Methanol–glacial acetic Photodiode NDa Pharmaceuti- 92
phenyl column acid–water array, cal prepara-
(300  3.9 mm, 10 mm) (85:2:15, v/v), 278 nm tions
1 mL/min
Nova-Pak C 18 column Acetonitrile–THF– Photodiode NDa Pharmaceuti- 92
(150  3.9 mm, 5 mm) water–glacial acetic array, cal prepara-
acid (15:40:45:2, v/v), 278 nm tions
Shimpack CLC-ODS 1000 mL of 85% Electrochemi- 0.4 pg Human 93
(15 cm  6.0 mm) Methanol containing cal detection serum
0.05 M sodium
perchlorat and 5.7 mL
of glacial acetic acid,
Hypersil ODS Acetonitrile–sodium UV, 280 nm 2 mg/mL Plasma/ 94
(25  0.4 cm i.d., 5 mm) acetate buffer pH 4.2 serum
column (60:40, v/v),
2.5 mL/min
LiChrosorb RP 6.5 mM Phosphoric UV, 280 nm 0.1 mg/L Plasma 95
18 column acid–acetonitrile
(250  4 mm i.d., 7 mm) (45:55, v/v),
Octyl coated silica of Acetonitrile–0.05 M UV, 240 nm NDa Black pearl 53
RP-8 Spheri-10 column potassium dihydrogen (black
(250  4.6 mm, 10 mm) phosphate pH 3.3 herbal pill)
(15:85, 50:50, 70:30),
Zorbax C 8 column Methanol–water–acetic UV, 280 nm NDa Human 96
(25 cm  4.6 mm i.d.) acid (70:30:1), 1.4 mL/ serum
min, 35 C albumin
Hypersil ODS 0.1 M Potassium dihy- UV, 240 nm NDa Blood 97
stainless-steel column drogen phosphate–for-
(16 cm  5 mm i.d., mic acid–isopropanol
5 mm) (1000:1:17, v/v),
(16 cm  5 mm mic acid–isopropanol
i.d., 5 mm) (1000:1:176, v/v),
Hypersil ODS 0.1 M Potassium UV, 240 nm NDa Blood 97
stainless-steel column dihydrogen
(16 cm  5 mm phosphate–formic
i.d., 5 mm) acid–isopropanol
(1000:1:540, v/v),
ODS silica stainless-steel Methanol–1% UV, 254 nm NDa Pharmaceuti- 98
column (150  1 mm acetic cal prepara-
i.d., 3 mm) acid, 30 mL/min tions
column (150  1 mm ammonia, 30 mL/min cal prepara-
i.d., 3 mm) tions
Hibar RP 18 column Acetonitrile–0.05 M UV, 254 nm NDa Urine 99
(250  4 mm i.d., 5 mm) phosphate buffer pH
and short pre column 4.5 (from 25–55%
Pherisorb RP 8 acetonitrile),
(30  4 mm i.d., 0.8 mL/min, 35 C
30–40 mm)
Cyanopropylsilane Water–acetonitrile– UV, 254 nm 1 mg/mL Plasma 100
stainless steel column acetic acid (60:30:10,
(30 cm  4 mm i.d., v/v/v), 1.0 mL/min
10 mm)
Zorbax SB-C 8 0.1% TFA in water–acet- NDa NDa Solution 101
(50  2.1 mm, 5 mm) onitrile (60:40),
ND ¼ no data available
indomethacin mixture was achieved on the SB-biphenyl-30 column using
a pressure gradient. A mixture containing flufenamic acid, mefenamic
acid, acetylsalicylic acid, ketoprofen and fenbufen, and another mixture
containing ibuprofen, fenoprofen, naproxen ketoprofen and tolmetin
were well separated on the SB-cyanopropyl-50 column using pressure
gradients. Typical analysis time for a mixture of NSAIDs on the biphenyl
or cyanopropyl column was approximately 20–25 min. Application of the
method using the biphenyl column to the determination of NSAIDs
present in selected dosage forms was performed.
Ku et al. [103] developed a micellar electrokinetic capillary chromato-
graphy (MEKC) to identify adulterants in rheumatic and analgesic
traditional Chinese medicine. This chromatographic technique was
carried out using a buffer of (pH 9.0) 0.02 M sodium tetraborate and
dihydrogenphosphate (containing 0.15 M sodium lauryl sulfate). This
method was applied to simultaneously analyze fourteen synthetic
chemical drugs adulterated in traditional Chinese medicine within 25
min. They represent five pharmacological categories, namley antipyretic
analgesics, glucocorticoids, CNS stimulants, muscle relaxants and
sedatives, and include acetaminophen, aminopyrine, bucetin, diclofenac
sodium, ethoxybenzamide, indomethacin, ketoprofen, mefenamic acid,
phenylbutazone, dexamethazone, prednisolone, chlorzoxazone, diaze-
pam and caffeine. The recovery of synthetic chemical drugs added to
Dwu-Hwo-JIh-Seng-Tang ranged from 92.1 to 102.5%. This method was
used to assess two traditional Chinese medicines sold by the distributors
of Chinese natural drugs collected by the consumer centers of local health
bureaus from October 1991 to June 1992. MEKC was found to be an
efficient and sensitive chromatographic technique to measure adulterants
in rheumatic and analgesic traditional Chinese medicine.
The possibility of separating flufenamic, meclofenamic and mefenamic
acids by capillary electrophoresis using b-cyclodextrin was studied by
Pérez-Ruiz et al. [104]. A capillary electrophoresis P/ACE System 5500
(Beckman Instruments) equipped with a diode-array detector, an
automatic injector, a fluid-cooled cartridge and a System Gold data
station were used in this study. Electrophoresis was performed in a 57
cm  75 mm i.d. (50 cm to the detector) fused-silica capillary tube
(Beckman Instruments). The high voltage power supply was set to 12 kV
(normal polarity, equivalent to a field strength of 210 V/cm) resulting in a
typical current of 90–92 mA. Sample introduction was made at the anode
side using the pressure option (0.5 p.s.i.  33.3 mbar) for 4–10 s. The
detection of analytes was performed at 285 nm. The best approach
involved combining a suitable pH of the carrier electrolyte (pH 12.0) with
the host-guest complexation effects of b-cyclodextrin. A running buffer
consisting of 30 mM phosphate buffer (pH 12.0), 2 mM b-CD and 10%
(v/v) acetonitrile was found to provide a very efficient and stable
electrophoresis system for the analysis of fenamic acids by capillary zone
electrophoresis (CZE). Responses were linear from 0.4 to 40 mg/mL for
the three drugs with detection limits of about 0.3 ng/mL. Intra- and inter-
day precision values of about 1–2% R.S.D. (n ¼ 11) and 3–4% R.S.D.
(n ¼ 30), respectively, were obtained. The method is highly robust and no
breakdowns of the current or capillary blockings were observed for
several weeks. The general applicability of this rapid CZE procedure
(migration less than 12 min) is demonstrated for several practical
samples, including serum, urine, and pharmaceuticals.
Cherkaoui and Veuthey [105] described the simultaneous separation of
nine nonsteroidal antiinflammatory drugs by nonaqueous capillary
electrophoresis (NACE). NACE was found to be a good alternative for
the analysis of pharmaceutical drugs and their metabolites, which are
difficult to separate in aqueous media. CE data were generated in a
Hewlett-Packard CE system equipped with an on-column diode array
detector, an auto sampler and a power supply able to deliver upto 30 kV.
The total capillary length (Composite Metal Services) was 48.5 cm, while
the length of the detector was 40 cm, with a 50 mm internal diameter. An
alignment interface, containing an optical slit matched to the internal
diameter, was used and the detection wavelength was set at 200 nm with a
bandwidth of 10 nm. All experiments were performed in cationic mode
(anode at the inlet and cathode at the outlet). The capillary was
maintained in a thermostat at 20 C, unless otherwise stated. A constant
voltage of 30kV, with an initial ramping of 500 V/s, was applied during
analysis. Sample injection (8 nL injection volume) was achieved using the
pressure mode for 10 s at 25 mbar. The selected electrophoretic medium,
a methanol–acetonitrile (40:60, v/v) mixture containing 20 mM ammo-
nium acetate, yielded the best compromise in terms of analysis time,
selectivity and separation efficiency for the simultaneous separation of
baclofen, indoprofen, ibuprofen, fenoprofen, indomethacin, ketoprofen,
suprofen, diclofenac, and mefenamic acid.
residues in water based on a combination of liquid chromatography (LC)
or capillary electrophoresis (CE) with mass spectrometry (MS). A 20 mM
ammonium acetate solution adjusted to pH 5.1 with acetic acid was
employed as the carrier electrolyte for the separation of clofibric acid,
naproxen, bezafibrate, diclofenac, ibuprofen, and mefenamic acid by CE–
MS. For the determination of the drug residues by CE–MS the water
samples were pretreated by liquid–liquid extraction (LLE) prior to solid-
phase extraction (SPE). A 500 mL volume of water sample was brought
to pH 2 with concentrated hydrochloric acid; after the addition of 50 g
sodium sulfate the sample was extracted twice with 25 mL of a mixture of
hexane–MTBE (methyl tert-butyl ether) (1:1, v/v). The organic phase was
re-extracted twice with 50 mL 2 mM sodium hydroxide solution, brought
to pH 2 with hydrochloric acid and diluted to 250 mL with water, pH 2.
The extract was passed through an SPE cartridge (6 mL) packed with 500
mg Bondesil ODS 40 mm (Varian) conditioned with acetone, methanol,
and water, pH 2 (one cartridge volume each). The flow-rate was adjusted
to approximately 10 mL/min. After the cartridges had been allowed to
dry for 30 min, the drugs were eluted using an overall volume of 2 mL
methanol. The extract was brought to dryness in a nitrogen stream
(purity 4.6) and finally redissolved in 50 mL of a mixture of methanol–
carrier electrolyte (80:20, v/v) prior to injection (5 kPa, 0.3 min). A
Crystal 310 CE instrument (Thermo CE) was employed for CE–MS
experiments. Fused-silica capillaries of 50 mm i.d. were obtained from
Polymicro Technologies; new capillaries with a length of 70 cm were
conditioned by flushing with 0.1 M NaOH followed by water (each for 10
min). Finally, the conditioning procedure was completed by flushing with
the CE carrier electrolyte consisting of 20 mM ammonium acetate, pH
5.1 for 5 min; prior to each run the capillary was flushed with carrier
electrolyte for 3 min. MS detection was performed on a quadrupole
system HP 5989B (Agilent) equipped with a radiofrequency-only
hexapole (Analytica of Branford) using a pneumatically assisted
electrospray ionization interface HP 59987A (Agilent). A CE probe
was used for CE–MS experiments. The sheat-liquid for CE–MS consisted
of 2-propanol–water (80:20, v/v) containing 0.1% (v/v) acetic acid for the
positive detection mode or 0.1% (v/v) triethylamine for the negative
mode and was delivered by a syringe pump (Model 22; Harvard
Apparatus) at a flow rate of 4 mL/min. The drying gas flow-rate was 1.4
L/min at 150 C. CE–MS proved to be a useful alternative to HPLC–MS,
although the sample pretreatment had to be expanded by an LLE prior to
the SPE. Because of rather high standard deviations of the recoveries due
to the combination of three extraction steps, a method based on standard
addition is recommended for quantitation. As expected the detection
limits for the CE–MS method are poorer than the detection limits
obtained by HPLC–MS namely between 27 and 93 mg/L for standard
injections which resulted in detection limits in the samples between 4.8
and 19 ng/L. Nevertheless, CE–MS may be a useful technique for the
confirmation of questionable results obtained by HPLC–MS.
Regarding the work described above, Ahrer and Buchberger [106] noted
that in contrast to the technique of gas chromatography (GC) coupled
with MS, CE–MS has an advantage in that derivatization of nonvolatile
analytes can be avoided. HPLC–MS yields better detection limits, but
CE–MS is a valuable complementary alternative, which can be used
when separation selectivity of HPLC is not satisfactory or when results
from HPLC must be confirmed by a different technique. Therefore, they
developed a nonaqueous capillary electrophoresis (NACE) for the
separation of clofibric acid, penicillin V, diclofenac, mefenamic acid,
bezafibrate, naproxen and ibuprofen using carrier electrolyte consisting
of 20 mM ammonium acetate, 60 mM acetic acid in methanol–
acetonitrile (40:60). Capillary electrophoresis separation was performed
using an HP3D instrument (Agilent Technologies). Mass spectrometry
detection has been done on an HP 5989 MS equipped with an HP
59897A electrospray interface (Agilent Technologies). The sprayer
needle for CE–MS coupling is a grounded stainless steel capillary
with a coaxial CE capillary inside. Between the CE capillary and the
steel capillary is a sheath flow that forms the electrical contact and
provides a stable spray. The CE capillary was 50 mm  70 cm fused
silica. A constant voltage of 30 kV was applied during analysis.
Sample injection (10 mg/L each) was achieved using the pressure mode
for 18 s at 50 mbar. The capillary was flushed with hexadimethrin
bromide prior to the first use to obtain a permanent reverse of the
display electroosmotic flow (EOF). In contrast to the aqueous CE
method, the separation of basic analytes could not be performed using
these conditions, but the acidic compounds could be detected with
quantitation limits between 10 and 160 mg/L.
6.7 Capillary isatachophoresis
Polásek et al. [107] used anionic capillary isatachophoresis (ITP) with
conductimetric detection for determining selected nonsteroidal antiin-
flammatory and analgesic drugs of the phenamate group, namely
tolfenamic, flufenamic, mefenamic and niflumic acids. Isatachophoresis
analyses were carried out with the use of a computer-controlled EA
100 ITP analyzer (VILLA s.r.o.) operated in the single-column mode.
The analyzer was equipped with a 30-mL sampling valve, a 160
mm  0.3 mm i.d. analytical capillary made of fluorinated ethylene-
propylene (FEP) copolymer and a conductivity detector. Initially the
pKa values (proton lost) of tolfenamic, flufenamic, mefenamic, and
niflumic acids were determined as 5.11, 4.91, 5.39, and 4.31,
respectively, by the UV spectrophotometry in aqueous 50% (w/w)
methanol. The optimized ITP electrolyte system consisted of 10 mM
HCl þ 20 mM imidazole (pH 7.1) as the leading electrolyte and 10
mM 5,50 -diethylbarbituric acid (pH 7.5) as the terminating electrolyte.
The driving and detection currents were 100 mA (for 450 s) and 30 mA,
respectively (a single analysis took about 20 min). Under such
conditions the effective mobilities of tolfenamic, flufenamic, mefenamic
and niflumic acids varied between 23.6 and 24.6 m2 V1 s1 (evaluated
with orotic acid as the mobility standard). The calibration graphs
relating the ITP zone length to the concentration of the analytes were
rectilinear (r ¼ 0.9987–0.9999) in the range 10–100 mg/L of the drug
standard. The RSDs were 0.96–1.55% (n ¼ 6) when determining 50
mg/L of the analytes in pure test solutions. The method has been
applied to assay the phenamates in six commercial mass-produced
pharmaceutical preparations (MobilisinÕ gel and ointment, LysalgoÕ
capsules, NiflurilÕ cream, NiflugelÕ gel, and ClotamÕ capsules).
According to the validation procedure based on the standard addition
technique the recoveries were 98.4–104.3% of the drug and the RSD
values were 1.25–3.32% (n ¼ 6).
7. DETERMINATION IN BODY FLUIDS AND TISSUES
7.1 Spectroscopic methods
Chemiluminescence can be used to determine mefenamic acid in biological
fluids according to the procedure described earlier by Aly [45] (see Section
5.4). For plasma, the extraction process was carried out with ethyl acetate
after acidification with acetic acid. For spiked plasma the calibration
graphs were obtained over the concentration ranges 0.07–6.0 and 0.05–
6.0 mg/mL for flufenamic and mefenamic acids, respectively. The
respective mean percentage recoveries were 99.57  0.56 and
99.17  0.98 for flufenamic and mefenamic acids, respectively. In the
case of urine, the extraction process was carried out with diethyl ether after
acidification with hydrochloric acid. For spiked urine, the chemilumines-
cence intensity (I, mV) was linearly related to flufenamic and mefenamic
acids concentrations over the range 0.5–6.0 mg/mL. The respective mean
percentage recoveries were 99.94  0.41 and 100.10  0.57 for flufenamic
and mefenamic acids, respectively.
The simultaneous analysis of 14 nonsteroidal antiinflammatory drugs in
human serum using negative electrospray ionization-tandem mass
spectrometry without chromatography was described by De Kanel [47]
(see Section 5.6).
7.2 Chromatographic methods
A simple detection of mefenamic acid and its metabolites in urine by thin
layer chromatography was reported earlier by Demetriou and Osborne
[52] (see Section 6.1).
As described in the Section 6.3, the gas chromatography procedure has
been used to determine mefenamic acid in plasma and urine of human
and horse [57–64].
Similarly, the high-performance liquid chromatography is shown to be
the most applied procedure for the determination of mefenamic acid in
biological fluids (see Table 1).
Determination of flufenamic, meclofenamic and mefenamic acids in
human urine and serum samples using capillary zone electrophoresis
(CZE) was described by Pérez-Ruiz [104] (see Section 6.6). Analysis of
drug in urine samples by CZE is always a delicate problem. If urine is
directly injected into the capillary, proteins and the other biomolecules
in the urine matrix adsorb to the wall of the capillary and thus quickly
harm the column's performance. In addition, appropriate conditions
must be found where no other components comigrate with the analytes.
Experiments with different urine dilution ratios have shown that a
10 fold dilution of urine was suitable for the analysis because of
flufenamic, meclofenamic and mefenamic acids were kept free of
adverse matrix effects. The serum was treated with acetonitrile to
separate proteins. After centrifugation (3 min at 1000 g), the liquid
supernatant was treated with 0.1 mL of 0.01 M sodium hydroxide,
filtered through a 0.45 mm filter and diluted with demineralized water to
appropriate volume. In the concentration range studied (3–20 and 3–30
mg/mL for urine and serum samples, respectively), the calibration curves
ware linear and the migration times were reproducible. Separation of
the fenamic acids under the best conditions for human urine and serum
samples gave results with mean recovery in the range of 98.6–102.1%
and 98.0–102.7% for mefenamic acid in urine and serum samples,
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Nimodipine: Physical Profile
1. General Information. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 338
3. Stability and Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . 353
1,4-Dihydro-2,6-dimethyl-4-(3-nitrophenyl)-3,5-pyridinedicarboxylic
acid 2-methoxyethyl 1-methylethyl ester.
2-Methoxyethyl 1,4-dihydro-5-(isopropoxycarbonyl)-2,6-dimethyl-4-(3-
nitrophenyl)-3-pyridinecarboxylate.
NIMODIPINE: PHYSICAL PROFILE 339
Isopropyl 2-methoxyethyl 1,4-dihydro-2,6-dimethyl-4-(3-nitrophenyl)-
3,5-pyridinedicarboxylate.
2,6-Dimethyl-4-(30 -nitrophenyl)-1,4-dihydropyridine-3,5-dicarboxylic
acid-3-b-methoxyethyl, 5-isopropyl diester.
2,6-Dimethyl-4-(m-nitrophenyl)-1,4-dihydropyridine-3,5-dicarboxylic
acid-3-b-methoxy ethyl-5-a-methylethyl diester.
5-(Isopropoxycarbonyl)-3-(2-methoxyethoxycarbonyl)-4-(3-nitrophenyl)-
2,6-dimethyl-1,4-dihydropyridine.
Nimodipine.
Admon, Nimotop, Periplum, Nimodipina, Norton, Noodipinia, Oxigen,
Brainox, Modina, Sobrepina, Trinalion, Brainal, Calnit, Kenesil, Modus,
Remontal, Vasotop, Nivas, Cebrofort, Acival, Eugerial.
Bay-e-9736
C21H26N2O7 418.459 [66085-59-4].
(CH3)2CHO C C OCH2CH2OCH3
C 60.28% H 6.26% N 6.69% O 26.76%.
Nimodipine is obtained as a light yellow to yellow crystalline powder.
The pH of a 1% (w/v) solution is 6–6.8 [3].
Nimodipine is practically insoluble in water, freely soluble in ethyl
acetate, and sparingly soluble in absolute alcohol [3].
The specific rotation of the ( þ )-form is [a]20 þ 7.9 (c ¼ 0.439 in dioxane),
and the specific rotation of the ( )-form is [a]20 7.9 (c ¼ 0.374 in
dioxane) [2]. When tested according to the British Pharmacopoeia, the
angle of optical rotation, determined on solution S, is –0.10 to þ 0.10 .
The X-ray powder diffraction pattern of nimodipine is shown in Figure 1
and was obtained using a Simons XRD-5000 diffractometer. Table 1
shows the values of the scattering angles (units of degrees 2y), the
interplanar d-spacings (Å), and the relative peak intensities.
Nimodipine melts at about 125 C [2].
The differential scanning calorimetry thermogram of nimodipine was
obtained using a DuPont TA-9900 thermal analyzer attached to a
a heating rate of 10 C/min, and was run from 40 to 400 C. The
compound was found to melt at 130.9 C.
Scattering Angle (degrees 2θ)
Figure 1. X-ray powder diffraction pattern of nimodipine.
The UV spectrum of nimodipine (10 mg/mL) in ethanol is shown in
Figure 3, and was recorded using a Shimadzu UV–visible Spectro-
photometer 1601 PC. Nimodipine exhibited two maxima at 355 and 236.6
nm, which are characterized by the following values:
max(nm) A [1%, 1 cm] Molar absorptivity
(L mol1 cm1)
The infrared absorption spectrum of nimodipine shown in Figure 4
was obtained in a KBr pellet, using a Perkin Elmer infrared
spectrophotometer. The principal peaks were observed at 3379, 3271,
3216, 3087, 2977, 1698, 1644, 1576, 1529, 1498, and 1360 cm1.
Assignments for the infrared absorption bands of nimodipine are given
Pattern of Minodipine
(degrees 2y) (%) (degrees 2y) (%)
6.485 13.6180 58.19 10.303 8.5786 0.89
Figure 2. Differential scanning calorimetry thermogram of nimodipine.
Figure 3. The ultraviolet absorption spectrum of nimodipine in ethanol.
Figure 4. The infrared absorption spectrum of nimodipine obtained in a KBr pellet.
Assignments for the Infrared Absorption Bands
of Nimodipine
3379, 3271 N–H stretch
3216, 3087 Aromatic C–H stretch
1698 C¼O stretch
1644, 1576 N–H bending
1529, 1498, 1360 C¼C ring stretch
1360 C–C(¼O)–O stretch of a,b-unsa-
turated ester
The proton NMR spectrum of nimodipine was obtained using a Bruker
Avance Instrument operating at 300, 400, or 500 MHz. Standard Bruker
Software was used to obtain COSY and HETCOR spectra. The sample
was dissolved in D2O, and tetramethylsilane (TMS) was used as the
internal standard. The proton NMR spectra are shown in Figures 5 and
6, the COSY 1H-NMR spectra are shown in Figures 7 and 8, and the
gradient HMQC-NMR spectrum is shown in Figure 9. Assignments for
the 1H-NMR resonance bands of nimodipine are given in Table 3.
The carbon-13 NMR spectrum of nimodipine was obtained using a
Bruker Avance Instrument operating at 75, 100, or 125 MHz. Standard
Figure 5. The 1H-NMR spectrum of nimodipine in DMSO-d6.
Figure 6. Expanded 1H-NMR spectrum of nimodipine in DMSO-d6.
Figure 7. C-NMR spectrum of nimodipine in DMSO-d6.
Figure 8. DEPT 90 1H-NMR spectrum of nimodipine in DMSO-d6.
Assignments of the 1H-NMR Chemical Shifts of Nimodipine
18 NO2
H3C 8 20 19 7 5
CHOOC 10 4 9 COOCH2CH2OCH3
3 H 2
Proton Chemical shift Multiplicity* Number of
atoms (ppm relative proton atoms
H-2, H-3 1.01, 1.02 Two d 6
H-6, H-7 3.43–3.49, m 4
H-8 4.81 q 1
H-13 7.53 Two d 1
H-14 7.63 d 1
H-12 8.01 s 1
N–H 9.01 s 1
Figure 10. DEPT 135 NMR spectrum of nimodipine in DMSO-d6.
dissolved in D2O and tetramethylsilane (TMS) was used as the internal
standard. The carbon-13 NMR spectrum of nimodipine is shown in
Figure 10, and the DEPT-NMR spectra are shown in Figures 11 and 12.
Assignments for the various carbon resonance bands of nimodipine are
The electron impact (EI) spectrum of nimodipine shown in Figure 13
was recorded using a Shimadzu PQ-5000 GC–MS spectrometer. The
spectrum shows a mass peak (M.) at m/z ¼ 418, and a base peak at m/z ¼
296 resulting from the loss of the nitrophenyl group. These and
Figure 11. HSQC NMR spectrum of nimodipine in DMSO-d6.
Figure 12. HMBC NMR spectrum of nimodipine in DMSO-d6.
ssignments of the C-NMR Chemical Shifts of Nimodipine
NIMODIPINE: PHYSICAL PROFILE
Carbon atoms Chemical shift Assignments at Carbon atoms Chemical shift Assignments at
(ppm relative carbon number (ppm relative carbon number
to TMS) to TMS)
C-1 18.13 2 C-12 122.10 1
C-2, C-3 21.42, 21.74 2 C-13 129.46 1
C-5 57.92 1 C-15, C-16 146.07, 146.68 2
C-9, C-10 100.79, 101.52 2 C-19, C-20 165.88, 166.39 2
C-11 120.94 1
Mass Spectrum of Nimodipine
Mass Relative Structural
number (m/z) intensity (%) assignment
418 6.86 C21H26N2O7eþ þ . mass peak (M.)
359 9.8 M–C3H7O
331 5.7 M–C4H7O2
296 100 [M–C6H4NO2] base peak
273 10.8 M–(C3H7O2 þ CO
þ C3H6)
254 68.6 M–(C3H7O þ C2H4O
þ NO2 þ CH3)
227 4.9 M–(C3H7O2 þ CO
þ C3H6 þ NO2)
196 53.9 M–(C3H7O2 þ CO þ C3H6
þ NO2 þ 2CH3 þ H)
122 4.8 M–C15H22NO5
67 5.8 [CH3–C:C–C:O] þ
59 10.8 [CH3OCH2CH2] þ
3. STABILITY AND STORAGE
Solutions of nimodipine are incompatible with some plastics, including
polyvinyl chloride. The solutions are also light-sensitive, and should be
protected from light [3]. The degradation half-lives of nimodipine are 16
and 56 h following exposure of aqueous solution of the drug to UV light
Figure 13. Mass spectrum of nimodipine.
at 360 nm and daylight, respectively [5]. Such solutions should be
protected from freezing, and stored at 15–30 C.
ed., Mack Publishing Co., Pennsylvania, p. 855 (1990).
2. The Merck Index, 12th edn., S. Budavari, ed., Merck and Co., NJ
Sweetman, ed., The Pharmaceutical Press, Chicago, p. 946 (2002).
of Helth-System Pharmacists, p. 1259 (1995).
Nimodipine: Analytical Profile
1. Compendial Methods of Analysis . . . . . . . . . . . ......... 356
1.1 Identification methods for the
drug substance . . . . . . . . . . . . . . . . . . . . . . . . . . 356
1.2 Tests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 356
1.3 Impurities and related substances . . . . . . . . . . . . . 356
2. Methods of Analysis Reported in the Literature. . . . . . . . . . 359
2.5 Liquid chromatography . . . . . . . . . . . . . . . . . . . . 363
2.6 Capillary electrophoresis (CE) . . . . . . . . . . . . . . . 366
1.1 Identification methods for the drug substance
absorption spectrophotometry for the identification of the pure drug.
The procedure is as follows:
Examine by infrared absorption spectrophotometry comparing with the
spectrum obtained with nimodipine CRS. If the spectra obtained in the
solid state show differences, record further spectra using 20 g/L solutions
in methylene chloride R and a 0.2-mm cell.
1.2 Tests [1]
1. Examine by infrared absorption spectrophotometry or compar-
ing with the spectrum obtained with nimodipine CRS.
2. Dissolve 1 g in acetone and dilute to 20 mL with acetone. The
solution should be clear.
1.3 Impurities and related substances [1]
The European Pharmacopoeia [1] described a liquid chromatographic
method for the determination of three related substances, denoted as
NIMODIPINE: ANALYTICAL PROFILE 357
A, B, and C:
Substance A: 2-methoxyethyl 1-methylethyl-2,6-dimethyl-4-(3-nitrophe-
nyl) pyridine-3,5-dicarboxylate.
H3C O O
C C OCH3
CH3 O O
Substance B: In the structure below, R ¼ CH(CH3)2. The systematic
compound name is bis (1-methylethyl)-2,6-dimethyl-4-(3-nitrophe-
nyl),1,4-dihydro-pyridine-3,5-dicarboxylate.
RO OR
Substance C: In the structure above, R ¼ CH2–CH2–OCH3. The
systematic compound name is bis (2-methoxyethyl)-2,6-dimethyl-4-
(3-nitrophenyl)-1,4-dihydro pyridine-3,5-dicarboxylate.
The method requires preparation of the following three solutions.
Test Solution: Dissolve 40 mg of the substance to be examined in 2.5
mL of tetrahydrofuran and dilute to 25 mL with the mobile phase.
Reference Solution (a): Dilute 1 mL of the test solution to 100 mL
with the mobile phase. Dilute 2 mL of the solution to 10 mL with
Reference Solution (b): Dissolve 20 mg of nimodipine impurity A in
2.5 mL of tetrahydrofuran and dilute to 25 mL with the mobile
phase. Dilute 1 mL of the solution to 20 mL with the mobile phase.
Reference Solution (c): Dilute 0.5 mL of the test solution to 20 mL
with the mobile phase.
Reference Solution (d): Mix 1 mL of reference solution (b) and 1 mL
of reference solution C, and dilute to 25 mL with the mobile phase.
The chromatographic procedure is carried out using a stainless steel
column 12.5 cm long and 4.6 mm in internal diameter packed with
octadecylsilyl silica gel (5 mm), which is maintained at 40 C. A mobile
phase consisting of a mixture of 20 volumes of methanol, 20 volumes of
tetrahydrofuran, and 60 volumes of water, is eluted at a flow rate of 2 mL/
min. Detection is effected at 235 nm. When 20 mL of reference solution
(d) is injected, the retention time of impurity A is about 7 min, and
nimodipine elutes at about 8 min. Separately inject 20 mL of the test
solution and 20 mL of reference solution (a). Record the chromatogram
for the test solution for four times longer than the retention time of
nimodipine. In the chromatogram obtained with the test solution, the area
of the peak due to impurity A is not greater than the corresponding peak
in the chromatogram obtained with reference solution (d) (i.e., 0.1%).
None of the peaks (apart from the principal peak and the peak due to
impurity A) has an area greater than the area of the principal peak in the
chromatogram obtained with reference solution (a) (i.e., 0.2%). The sum
of the areas of all peaks (apart from the principal peak) is not greater than
2.5 times the area of the principal peak in the chromatogram obtained
with reference solution (a) (i.e., 0.5%). Disregard any peak due to the
solvent and any peak with an area less than 0.5 times the area of the
principal peak in the chromatogram obtained with reference solution (d).
Loss on drying
The permissible content is not more than 0.5% when determined on a
1.0 g sample.
Sulfated ash
The permissible sulfated ash content is not more than 0.1% when
determined on 1.0 g sample.
Assay method for the bulk drug substance
The European Pharmacopoeia [1] recommends the following assay
method. Dissolve 0.180 g with gentle heating in a mixture of 25 mL of
2-methyl-2-propanol and 25 mL of perchloric acid. Add 0.1 mL of
ferroin, and titrate with 0.1 M ceric sulfate (titrate slowly toward the end
of the titration). Carry out a blank experiment, and correct if necessary.
Each milliliter of 0.1 M ceric sulfate is equivalent to 20.92 mg
Assay method for tablets [1]
The assay method for nimodipine in tablets is performed according to the
following procedure. Weigh and pulverize 20 tablets, and prepare the
following solutions for use in the liquid chromatography method:
Solution (1): Add 50 mL of methanol to a quantity of the powdered
tablets containing 60 mg of nimodipine, and mix with the aid of
ultrasound for 5 min. Then add sufficient methanol to produce
100 mL, centrifuge, and use the supernatant liquid.
Solution (2): Prepare to contain 0.06% w/v of nimodipine BPCRS
in methanol.
Solution (3): Prepare to contain 0.002% w/v of nimodipine BPCRS
and 0.002% w/v of nimodipine impurity ABPCRS in methanol.
The chromatographic system and conditions described under Related
Substances are used. The test is not valid unless in the chromatogram
obtained with solution (3), the resolution factor between the peaks due
to nimodipine and nimodipine impurity A is not less than 1.5, and the
symmetry factor due to nimodipine is not less than 2.
Calculate the content of nimodipine in tablets using the declared content
of C21H36N2O7 in nimodipine BPCRS. The content of nimodipine in
the tablet should be 95–105% of the label claim.
2. METHODS OF ANALYSIS REPORTED
IN THE LITERATURE
Reddy et al. described two spectrophotometric methods for nimodipine
in its dosage forms [2]. The first method involves the use of para
dimethylaminocinnamaldehyde as a colorimetric reagent, and the
measurement of the generated color at 510 nm. The second method is
based on the use of the Folin–Ciocalteu reagent, and a colorimetric
measurement at 640 nm. The Beer's law is obeyed over the ranges of
0.5–4 and 5–20 mg/mL, respectively.
Formation of a Schiff base product of the reduction product of
nimodipine (using the Zn–HCl system) and 4-dimethylaminobenzalde-
hyde formed the basis of a spectrophotometric method for the
determination of nimodipine in tablets [3]. A pink color exhibiting an
absorption maximum at 580 nm was produced and remained stable for
30 min. The percent recovery was 99–101%, with coefficients of variation
of 0.3–0.5.
Reduction of nimodipine to the corresponding aminoderivative (using
Zn–HCl), followed by diazotization and coupling with the Bratton–
Marshall reagent was reported [4]. The resulting color was measured at
550 nm, and the Beer's law was obeyed upto 40 mg/mL. Squella et al.
determined nimodipine by measuring the absorbance of its solution in an
acetate-phosphate buffer (pH 5) at 360 nm. The coefficient of variation
for this method was 2.5% [5]. A similar method based on measuring the
absorbance of nimodipine in ethanol at 237 nm was reported [6]. A linear
relationship was observed between the concentration and the absorbance
over the range of 0.14–6 mg/mL.
Belal et al. developed a fluorimetric method for the determination of
nimodipine [7]. The method involves reduction of nimodipine with Zn–
HCl, and measuring the fluorescence at 425 nm after excitation at 360
nm. The addition of triton X-100 was found to enhance the fluorescence.
The useful concentration range for the fluorescence measurement is from
0.1 to 5.0 mg/mL, with a detection limit of 0.06 mg/mL (1.62  107 M).
The method was applied to the analysis of commercial tablets, and was
further applied to spiked human urine. The percent recovery in the latter
study was 107.1  2.54 (n ¼ 4).
The electrochemical behavior of nimodipine was studied in ammonia
buffer containing 10% (v/v) ethanol [8]. A single-sweep oscillopolaro-
graphic method was then developed for nimodipine in tablets. The
calibration graph (peak current at 0.73 V vs. concentration) was linear
from 0.2 to 70 mM, and the detection limit was 10 mM. The same authors
applied linear sweep voltammetry for the determination of nimodipine in
tablets [9]. A reduction peak at 0.62V vs. the Ag/ACl reference
electrode was obtained over the range 10 nM0.8 mM with a detection
limit of 1 nM. The method involved a preconcentration period of 60 s.
The anodic electrochemical behavior of various 1,4-dihydropyridines
(including nimodipine) was studied in acetate-phosphate buffer (pH 8)
containing 70% (v/v) ethanol [10]. An anodic peak due to a two-electron
oxidation of the dihydropyridine ring to a pyridine derivative was
obtained. Recoveries were 91.7–104%, with a RSD being less than
3%. A cathodic polarographic method for nimodipine in tablets was
developed. The dependence of peak current on concentration in acetate-
phosphate buffer of pH 5 containing 20% ethanol was linear over the
range of 1 mM–1 mM, with a coefficient of variation of 0.2% [5].
Enantiometric mixtures of nimodipine were separated on b-cyclodextrin-
bonded silica gel plates. The plates were developed with light petroleum/
ethyl acetate/methanol, methanol/1% triethylammonium acetate (pH
4.1), or methanol/acetonitrile/1% triethylammonium acetate as mobile
phases. Spots were identified by illumination at 365 nm, or by exposure
to iodine vapor [11].
Trace amounts of nimodipine could be determined using TLC on silica
gel GF254 plates and a mobile phase consisting of chloroform/methanol/
methylene chloride/n-hexane (26:12:10:5). A. TLC-scanner was used to
measure the absorbance at 365 nm (Rf ¼ 0.48). The calibration graph
was linear from 5 ng to 1 mg, with a detection limit of 5 ng [12].
Nimodipine was determined using a HPTLC method in tablets after their
extraction with methanol (paracetamol was used as an internal standard).
The plates were developed with toluene/ethylacetate (1:1), and densito-
metric detection was effected at 240 nm. Calibration graphs were linear
from 25 ng (the detection limit) to 200 ng. Recovery was 96.03%, and the
relative standard deviation was 0.56% [13].
Metabolites of the dihydropyridines (including nimodipine) could be
determined in urine through the use of gas chromatography–mass
spectrometry after extractive methylation [14]. Derivatization was
affected using methyl iodide. The derivative was dissolved in ethyl
acetate, and was injected into a HP capillary (12 m  0.2 mm;
cross-linked methylsilicone film thickness 330 nm) operated with
temperature programming from 100 C and held for 3 min. Helium was
used as a carrier gas (flow rate of 1 mL/min), and 70 eV EIMS were used.
The overall recoveries ranged from 67 to 77%, with relative standard
deviations less than 10%.
Mixtures of dihydropyridine derivatives (including nimodipine) were
qualitatively and quantitatively determined by GC–MS [15]. Methanolic
samples were injected into a HP column (12 m  0.2 mm i.d.), operated
with temperature programming to 280 C at 2 C/min, and then to 300 C
at 10 C/min. Helium was used as the carrier gas (flow rate of 4.5 mL/
min), and 70 eV EMIS was used for detection. The analysis time was
45–60 min. The recoveries were better than 99%, and the relative
standard deviations were 2.11–2.46%.
Nimodipine was determined in tablets by GC after extraction with
chloroform (using nifedipine as an internal standard) [16]. A column
(3 ft  2 mm) of 10% of OV-1 on Chromosorb G (AW), 80–100 mesh
treated with DMCS and operated at 225 C, was used with nitrogen as
the carrier gas. Flame ionization was used as the means of detection. The
calibration graph was linear over the range 0.5–2 ng/mL, and the
coefficient of variation was 2.28.
Nimodipine in plasma was determined by a GC method [17]. Plasma was
treated with 2 M NaOH after addition of the internal standard, and then
extracted with toluene. The column (10 m  0.31 mm) consisted of cross-
linked 5% phenylmethyl silicone, and the method used temperature
programming from 90 C (held for 1 min) to 255 C at a heating rate of
25 C/min. Helium was used as a carrier gas, and the N-P detection mode
was employed. The calibration graph was linear from 2 to 50 ng/mL,
and the detection limit was 0.5 ng/mL.
The external factors influencing nimodipine concentrations during intra-
venous administration were studied using GC–electron-capture detection
[18]. Nimodipine was extracted from plasma and analyzed using a column
(1.8 m  2 mm) packed with 3% of OV-17 on gas-Chrom Q (50–100
mesh). The column was operated at 255 C with nitrogen as the carrier gas
(flow rate of 25 mL/min), and 63 Ni ECD. The calibration graph was
linear for upto 1000 ng/mL, and the limit of detection was 0.5 ng/mL.
Nimodipine could be determined by GC in plasma and cerebrospinal
fluid at concentrations as low as 1 ng/mL [19]. To avoid decomposition
during the GC analysis, nimodipine was first oxidized to its pyridine
derivative. A column (1.8 m  3 mm) of 2% of OV-17 on Anakrom Q
was operated at 250–260 C, with nitrogen as the carrier gas (flow rate of
25–30 mL/min) and ECD detection.
2.5 Liquid chromatography
The concentration–time profiles of nimodipine following intravenous and
oral administration were followed using high-performance liquid
chromatography (HPLC) after a solid phase extraction (SPE).
A column of 3 mm Spherisorb ODS2 was eluted with a mobile phase
(of 0.8 mL/min) consisting of 1 mM phosphate buffer (pH 5)/methanol
(7:13), and UV detection was used. Diazepam was used as the internal
standard, and the calibration graph was linear over the range of 5–200
ng/mL. Enantiometric ratios were determined on a 5 mm Ultron ES-
OVM chiral column using a mobile phase (flow rate of 1 mL/min)
consisting of ethanol/20 mM phosphate buffer (47:153). The data
evaluation entailed using the peak area ratio of the two enantiomers [20].
The stability of nimodipine injectable formulations was studied using
reversed phase HPLC, which made use of methyltestosterone as an
internal standard. A YWG C18 column was eluted with methanol/water
(13:7, v/v), and detection was effected at 238 nm. The calibration graph
was linear over the range of 5.98–299 mM/L [21].
1,4-Dihydropyridines (including nimodipine) were determined in human
plasma using HPLC coupled with electrochemical detection. The
separation was performed using a Supelcosil LC-ABZ-Plus C18 column.
A mobile phase consisting of methanol/H2O (7:3), that contained 2 mM
acetate buffer (pH 5), was eluted at a flow rate of 1 mL/min. The detector
was equipped with a glassy carbon electrode that was operated at 1000
mV vs. an Ag/AgCl reference electrode in the direct current mode. The
total elution time was less than 18 min [22].
Nimodipine, tarcine, and their metabolites were determined by HPLC in
the plasma of patients with Alzheimer's disease [23]. After extraction and
evaporation under nitrogen to dryness, the residue was reconstituted into
acetonitrile/H2O (1:3), and analyzed on a 5-mm Hypersil phenyl column
(25 cm  4.6 mm i.d.) equipped with a Nova Pak phenyl guard column
and operated with acetonitrile/4.4 mM KH2PO4 buffer (39:61) as the
mobile phase (flow rate of 1.5 mL/min). Detection was effected either
fluorimetrically at 330 nm excitation and 360 nm emission, or by its UV
absorbance at 239 nm. Calibration graphs were linear over the range
of 2–500 mg/mL for nimodipine and two of its metabolites, and 4–500
mg/mL for the third metabolite.
Six drugs (including nimodipine) used together in a program of
hypertension therapy were determined simultaneously by HPLC [24].
The tablets were extracted with methanol, and felodipne was added as an
internal standard. A column (25 cm  4 mm i.d.) of Jasco Metaphase
ODS silica (5 mm) was used with a mobile phase (flow rate of 1.5 mL/min)
of 10 mM phosphate buffer of pH 4.5/acetonitrile (1:1). Detection was
effected at 250 nm, and the linear operational range was found to be
25–3200 ng/mL.
Five dihydropyridine enantiomers (including nimodipine) were separated
by HPLC using cyclodextrin as the chiral selector [25]. Columns
containing b-cyclodextrin covalently bonded to 5 mm silica (25 cm
4 mm i.d.), or of (R)- or (S)-naphthyl ethyl carbomyl b-cyclodextrin
covalently bonded to 5 mm silica (25 cm  4.6 mm i.d.), were used. The
mobile phases (flow rate of 0.8 mL/min) were mixtures of ethanol or
acetonitrile, with 0.1% triethylamine, adjusted to pH 5 with acetic acid.
Nimodipine was determined in human plasma using HPLC and
methyltestosterone as an internal standard [26]. A Spherisorb ODS,
10 mm column (25 cm  4.6 mm i.d.) with methanol/H2O/n-butylamine
(65000-35000-1) was used as the mobile phase (flow rate of 1.2 mL/min),
with detection at 238 nm. The calibration graph was linear over the range
of 5–100 ng/mL, the detection limit was 2 ng/mL, and the average
recovery was 92%.
The enantiospecific determination of nimodipine in human plasma could
be established by liquid chromatography–tandem mass-spectroscopy
[27]. The separation was effected using a 8 mm Chiral OJ MOD column
(25 cm  2 mm i.d.), operated at 35 C with ethanol–n-heptane (1:4)
containing 2 mM ammonium acetate as the mobile phase (flow rate
of 300 mL/min) and MS detection. Calibration graphs were linear over
the range of 0.5–75 ng/mL, with a detection limit of 0.25 ng/mL for
each enantiomer.
Five chiral calcium antagonists (including nimodipine) were separated
using b-cyclodextrin as the stationary phase in a Merck Chiradex column
(25 cm  4.6 mm i.d.) with 0.1% triethylamine containing 5–100% of
methanol or acetonitrile (as modifiers) [28]. The solvent was adjusted
to pH 5 with acetic acid, and eluted at a flow rate of 0.8 mL/min.
UV absorbance was used for the detection.
The metabolism of nimodipine in rat liver microsomes was studied by
HPLC [29]. The centrifugate of the reaction mixture was analyzed using
a column (10 cm  4 mm i.d.) of Spherisorb S3 ODS II at 40 C, with a
mobile phase (flow rate of 0.5 mL/min) of 5 mM ammonium acetate
buffer (pH 6.6)/methanol (2:3) and detection at 218 and 238 nm.
Calibration graphs were linear over the range of 0.2–50 mM for
nimodipine, and 0.2–20 mM for the three main metabolites with detection
limits of 30–80 nM.
Zhang et al. described a HPLC method for the simultaneous determina-
tion of six different dihydropyridine calcium antagonists (including
nimodipine) in plasma [30]. After extraction with ethyl ether/n-heptane
(1:1) and centrifugation, the organic phase was heated at 50 C and the
residue dissolved in aqueous 60% acetonitrile (the mobile phase). An
Ultrasphere 5 mm ODS column (25 cm  4.6 mm i.d.) was used as the
means of separation. The mobile phase flow rate was 1 mL/min, and
detection was effected at 238 nm. The calibration graphs were linear over
the range of 50 ng/mL–6.4 mg/mL, with a detection limit of 5 ng/mL.
Qian and Gallo used HPLC to determine nimodipine in monkey plasma
[31]. After extraction with ethyl acetate and evaporating the organic
phase to dryness, the residue was dissolved in the mobile phase (aqueous
65% methanol). A Hypersil ODS column (15 cm  4.6 mm i.d.) was
used. The mobile phase flow rate was 1 mL/min, and detection was
effected at 238 nm. The calibration graph was linear over the range of
0.01–2 mg/mL, and the recovery was 95–106%.
Jiang et al. studied optimization of the mobile phase composition for the
HPLC separation of dihydropyridine drugs, including nimodipine [32].
These workers used a stainless steel column (15 cm  4.6 mm) 10 mm
operated at 30 C. The mobile phase was methanol-acetonitrile-water
(24.8:27:48.2).
Wang et al. studied the stability of nimodipine by HPLC, using
beclomethazone dipropionate as an internal standard [33]. The method
used a stainless steel column (25 cm  4 mm) containing YWG C18H37,
which was eluted at a flow rate of 1 mL/min) with methanol–water–ethyl
ether (35:15:4). Detection was effected at 238 nm. The calibration graph
was linear over the range of 0.03–0.72 mg/mL, the recovery was 100.1%,
and the coefficient of variation was 0.83%.
Jain and Jain reported a HPLC method for the microquantification of
nimodipine in intravenous infusion fluids [34]. A column (5 cm  4.6 mm
i.d.) of Shim-pack FLC-ODS was eluted with aqueous 75% methanol
as the mobile phase (flow rate of 1 mL/min). Detection was effected at
238 nm. The calibration graph was linear over the range of 5–25 mg/mL,
and recoveries were from 98.31 to 100.6%.
HPLC was used to evaluate the enantiomeric resolution of dihydrope-
pidine enantiomers (including nimodipine), using phenylcarbamates of
polysaccharides as a chiral stationary phase [35]. A column (25 cm  4.6
mm) packed with the arylcarbamate derivatives of amylase, cellulose, and
xylem was used. Detection was effected using polarimetry at 435 nm.
Using xylem bis-(3,5-dichlorophenylcarbamate) and a mobile phase (flow
rate of 0.5 mL/min) of 0.1%, diethylamine in hexane-propan-2-ol (17:3)
yielded separation of nimodipine.
Patil et al. utilized supercritical fluid chromatography for the simulta-
neous assay of three dihydropyridines, including nimodipine [36]. These
workers used a 10 mm JASCO-RP C18 column (25 cm  4.6 mm),
operated at 45 C and 13.7 mPa, with 13.04% methanol in CO2 as the
mobile phase (flow rate of 2 mL/min), and detection at 230 nm.
Calibration graphs were linear upto 10 mg/mL, with detection limit of
0.3 mg/mL. The same technique was used by Bhoir et al. for the
separation of seven vasodilating agents [37]. A JASCO-RP-C18 column
(25 cm  4 mm i.d.) operated at 45 C was used with a mobile phase
consisting of 14.2% methanol in CO2 and a pressure of 10.13 mPa, and
detection at 224 nm. The calibration graph was linear over the range of
0.2–10.8 mg/mL.
2.6 Capillary electrophoresis (CE)
Micellar electrokinetic chromatography was used as a fast screening
method for the determination of 1,4-dihydropyridines, and was used for
the separation of six drugs of this group including nimodipine [38].
A fused silica tube (58.5 cm  75 mm i.d., 50 cm to detector) was
operated at 25 kV with detection at 200–236 nm. The optimized buffer
(pH 8.2) contained 20 mM SDS in 50 mM borate buffer modified with
acetonitrile. Linear calibration graphs extended upto 70 mg/mL from
lower limits of 6.7 mg/mL.
The same technique was used by Wang et al. for the separation of five
dihydropyridine calcium channel blockers [39]. The method used a fused
silica capillary (75 cm  75 mm i.d.; 68 cm to detector) operated at 60 C
with an applied voltage of 24 kV, and a running buffer of 40 mM borax/
4 mM SDS at pH 8. Detection at 200 nm was used for quantitation.
Enantioseparation of dihydropyridine derivatives could be accomplished
using capillary electrophoresis by means of neutral and negatively
charged b-cyclodextrin derivatives [40]. The method used a fused silica
capillary (47 cm  75 mm; 40 cm to detector), operated at 25 C. The
running phase was 50 mM phosphate buffer (pH 3) containing
cyclodextrins and organic modifiers, and an applied voltage of 6–20 kV
was applied. Detection was effected at 190–300 nm.
Gilar et al. have also applied CE to the separation of dihydropyridine
calcium antagonists [25]. Their method used an uncoated capillary (76
cm  75 mm i.d.) at 20 kV, a running buffer of 20–25 M phosphate-
borate buffer (pH 9–9.5) containing 1% urea, and detection at 240 nm.
The buffer also contained 10 mM b-cyclodextrin, and varying amounts of
SDS.
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Nimodipine: Drug Metabolism and
1. Uses, Application, and Associated History . . . . . . . . . . . . . 372
1. USES, APPLICATION, AND ASSOCIATED HISTORY
Nimodipine is a dihydropyridine calcium channel blocker derivative
which is used in the treatment of cerebrovascular disorders. Particularly,
it is used as a prophylactic and in the treatment of ischemic neurological
deficits that follow aneurysmal or traumatic subarachnoid hemorrhage
[1]. It may also have an effect on patients with dementia of vascular or
degenerative origin which is most commonly due to Alzheimer's disease
and/or cerebrovascular disease [2, 3]. The drug acts mainly by dilating the
cerebral blood vessels and improving the cerebral blood flow, and it may
also prevent or reverse ischemic damage to the brain by limiting
transcellular calcium influx [4]. It is well-known that calcium has a role in
regulating various brain functions, which links membrane excitation to
subsequent intracellular enzymatic responses. Nimodipine is an isopropyl
calcium channel blocker which can easily cross the blood–brain barrier.
It reduces the number of open calcium channels, and thus decreases the
influx of calcium ions into the neurons [5].
The oral dose for the prophylaxis of neurological deficit is 60 mg every
4 h. Treatment should begin within 4 days of the onset of hemorrhage,
and should continue for 12 days. If cerebral ischemia has already
occurred, the neurological deficit may be treated by intravenous infusion
of 1 mg per hour for 2 h as an initial dose. The starting dose should be
increased to 2 mg per hour provided that no severe decrease in blood
pressure is observed [1].
Nimodipine is rapidly absorbed from the gastrointestinal tract following
oral administration, but undergoes extensive first-pass metabolism in the
liver [1]. The oral bioavailability is reported to be about 13%. Although
the drug administration using capsules could cause one to reach peak
levels faster, the bioavailability was not significantly different from that
of the tablets [6]. The drug is highly bound to plasma protein (greater
than 95%) [1, 7].
Mean cerebrospinal fluid concentrations were estimated to be 0.3  0.2
mg/L in patients whose mean plasma concentrations were 76.9  34
mg/L [8]. This indicates that nimodipine does not apparently bind to
the cerebrospinal fluid proteins. The bioavailability of dihydropyridines
following oral administration was found to increase after food
intake, such as grapefruit juice [9]. After intravenous administration,
nimodipine is distributed rapidly and uniformly. On the other
hand, after oral administration, nimodipine is strongly differentiated,
with high concentrations being found in the liver, kidneys, and fat,
low concentrations in the brain and testes, and the rest in the
plasma [10, 11]. Only small amounts of nimodipine pass through the
placenta [11].
Nimodipine is extensively metabolized in the liver, and undergoes
extensive first-pass metabolism [1]. More than 18 metabolites have been
detected and identified from the biotransformation of nimodipine. The
drug undergoes different reactions before its excretion. These reactions
include, dehydrogenation of the 1,4-dihydropyridine moiety, oxidation of
the two ester groups, and oxidative demethylation which is followed by
carboxylic acid formation via oxidation of the resulting alcohol. In
addition to the oxidation reactions, hydroxylation of the methyl groups
at the pyridine ring, hydroxylation of one methyl group at the isopropyl
ester moiety, reduction of the aromatic nitro group, and glucuronidation
can take place [12]. Nimodipine is mainly metabolized by cytochrome
P450-3A (CYP3A), which is involved in the dehydrogenation and
oxidation reactions [13]. The oxidation of 1,4-dihydropyridines is
proposed to produce a reactive oxygen species that could be harmful
to vulnerable tissues such as the brain [14]. Nimodipine is comple-
tely metabolized, and no unchanged drug was detected in either urine or
feces [12].
Scherling et al. [12] have thoroughly investigated the biotransformation
of nimodipine in rats, dogs, and monkeys, and have proposed the
metabolic pathway scheme shown in Figure 1.
NO2 NO2 NO2
(CH3)2CHOOC COOH (CH )2CHOOC COOCH2CH2OCH3 (CH3)2CHOOC
3 COOCH2CH2OH
H3C N CH3 H3C N CH3 N CH3
H H3C
nimodipine
NO2 NO2
(CH3)2CHOOC COOCH2CH2OH
(CH3)2CHOOC COOH (CH3)2CHOOC COOCH2CH2OCH3
CH3 H3C N CH3
H3C N CH3 H3C N
NH2 NO2
HOOC COOCH2COOH
HOOC COOCH2
(CH3)2CHOOC COOH CH2OH
HOOC COOH
HOCH2 N CH3
H3C N CH2OH
CH2OH
NO2 H COOC COOCH2CH2OH
CH2OH N
H COOC COOH NH2 NO2
COOH HOOC COOH
(CH3)2CHOOC
H3C N CH2OH HOCH2 N CH2OH
Figure 1. Metabolic pathway of nimodipine in rats, dogs, and in monkeys [12].
Nimodipine is excreted in feces via the bile duct, and in urine via the
glomular filtration, as metabolites [1, 10]. Fecal excretion is the major
excretory route (greater than 67%) [11]. The parent compound and its
metabolites were detected in breast milk [9]. The average elimination half-
life for the drug is reported to be 9 h, but the initial decline in plasma is
much more rapid, equivalent to a half-life of 1–2 h [1]. An average of
43% of bile-excreted nimodipine is subject to pronounced enterohepatic
circulation, and 57% is excreted with the feces [15, 16]. Due to this cycle
of excretion/reabsorption, the plasma concentration decreases and the
increases. More than 80 and 19% of the reabsorbed quantity is excreted
via bile and urine, respectively [15, 16]. The excretion routes detected in
the fetus are essentially the same as that of the mother [11].
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C. Wunsche, Arzneimittelforschung, 41, 392 (1991).
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CUMULATIVE INDEX
Bold numerals refer to volume numbers.
Acebutolol, 19, 1 Azintamide, 18, 1
Acetaminophen, 3, 1; 14, 551 Aztreonam, 17, 1
Acetazolamide, 22, 1
Acetohexamide, 1, 1; 2, 573; 21, 1 Bacitracin, 9, 1
Acetylcholine chloride, 31, 3, 21 Baclofen, 14, 527
Acyclovir, 30, 1 Benazepril hydrochloride, 31, 117
Adenosine, 25, 1 Bendroflumethiazide, 5, 1; 6, 597
Allopurinol, 7, 1 Benperidol, 14, 245
Amantadine, 12, 1 Benzocaine, 12, 73
Amikacin sulfate, 12, 37 Benzoic acid, 26, 1
Amiloride hydrochloride, 15, 1 Benzyl benzoate, 10, 55
Aminobenzoic acid, 22, 33 Betamethasone diproprionate, 6, 43
Aminoglutethimide, 15, 35 Bretylium tosylate, 9, 71
Aminophylline, 11, 1 Brinzolamide, 26, 47
Aminosalicylic acid, 10, 1 Bromazepam, 16, 1
Amiodarone, 20, 1 Bromcriptine methanesulfonate, 8, 47
Amitriptyline hydrochloride, 3, 127 Bumetanide, 22, 107
Amobarbital, 19, 27 Bupivacaine, 19, 59
Amodiaquine hydrochloride, 21, 43 Busulphan, 16, 53
Amoxicillin, 7, 19; 23, 1
Amphotericin B, 6, 1; 7, 502 Caffeine, 15, 71
Ampicillin, 2, 1; 4, 518 Calcitriol, 8, 83
Apomorphine hydrochloride, 20, 121 Camphor, 13, 27
Arginine, 27, 1 Captopril, 11, 79
Ascorbic acid, 11, 45 Carbamazepine, 9, 87
Aspartame, 29, 7 Carbenoxolone sodium, 24, 1
Aspirin, 8, 1 Cefaclor, 9, 107
Astemizole, 20, 173 Cefamandole nafate, 9, 125; 10, 729
Atenolol, 13, 1 Cefazolin, 4, 1
Atropine, 14, 325 Cefixime, 25, 39
Azathioprine, 10, 29 Cefotaxime, 11, 139
422 INDEX
Cefoxitin sodium, 11, 169 Cortisone acetate, 26, 167
Ceftazidime, 19, 95 Crospovidone, 24, 87
Ceftriaxone sodium, 30, 21 Cyanocobalamin, 10, 183
Cefuroxime sodium, 20, 209 Cyclandelate, 21, 149
Celiprolol hydrochloride, 20, 237 Cyclizine, 6, 83; 7, 502
Cephalexin, 4, 21 Cyclobenzaprine hydrochloride,
Cephalothin sodium, 1, 319 17, 41
Cephradine, 5, 21 Cycloserine, 1, 53; 18, 567
Chloral hydrate, 2, 85 Cyclosporine, 16, 145
Chlorambucil, 16, 85 Cyclothiazide, 1, 65
Chloramphenicol, 4, 47; 15, 701 Cypropheptadine, 9, 155
Chlordiazepoxide, 1, 15
Chlordiazepoxide hydrochloride, Dapsone, 5, 87
1, 39; 4, 518 Dexamethasone, 2, 163; 4, 519
Chloropheniramine maleate, 7, 43 Diatrizoic acid, 4, 137; 5, 556
Chloroquine, 13, 95 Diazepam, 1, 79; 4, 518
Chloroquine phosphate, 5, 61 Dibenzepin hydrochloride, 9, 181
Chlorothiazide, 18, 33 Dibucaine, 12, 105
Chlorpromazine, 26, 97 Dibucaine hydrochloride, 12, 105
Chlorprothixene, 2, 63 Diclofenac sodium, 19, 123
Chlortetracycline hydrochloride, Didanosine, 22, 185
8, 101 Diethylstilbestrol, 19, 145
Chlorthalidone, 14, 1 Diflunisal, 14, 491
Chlorzoxazone, 16, 119 Digitoxin, 3, 149; 9, 207
Cholecalciferol, 13, 655 Dihydroergotoxine
Cimetidine, 13, 127; 17, 797 methanesulfonate, 7, 81
Ciprofloxacin, 31, 163, 179, 209 Diloxanide furoate, 26, 247
Cisplatin, 14, 77; 15, 796 Diltiazem hydrochloride, 23, 53
Citric Acid, 28, 1 Dioctyl sodium sulfosuccinate,
Clarithromycin, 24, 45 2, 199; 12, 713
Clidinium bromide, 2, 145 Diosgenin, 23, 101
Clindamycin hydrochloride, 10, 75 Diperodon, 6, 99
Clioquinol, 18, 57 Diphenhydramine hydrochloride,
Clofazimine, 18, 91; 21, 75 3, 173
Clomiphene citrate, 25, 85 Diphenoxylate hydrochloride, 7, 149
Clonazepam, 6, 61 Dipivefrin hydrochloride, 22, 229
Clonfibrate, 11, 197 Dipyridamole, 31, 215
Clonidine hydrochloride, 21, 109 Disopyramide phosphate, 13, 183
Clorazepate dipotassium, 4, 91 Disulfiram, 4, 168
Clotrimazole, 11, 225 Dobutamine hydrochloride, 8, 139
Cloxacillin sodium, 4, 113 Dopamine hydrochloride, 11, 257
Clozapine, 22, 145 Dorzolamide hydrochloride, 26, 283;
Cocaine hydrochloride, 15, 151 27, 377
Codeine phosphate, 10, 93 Doxorubicine, 9, 245
Coichicine, 10, 139 Droperidol, 7, 171
Echothiophate iodide, 3, 233 Gadoteridol, 24, 209
Econazole nitrate, 23, 127 Gentamicin sulfate, 9, 295; 10, 731
Edetic Acid (EDTA), 29, 57 Glafenine, 21, 197
Emetine hydrochloride, 10, 289 Glibenclamide, 10, 337
Enalapril maleate, 16, 207 Gluthethimide, 5, 139
Ephedrine hydrochloride, Gramicidin, 8, 179
15, 233 Griseofulvin, 8, 219; 9, 583
Epinephrine, 7, 193 Guaifenesin, 25, 121
Ergonovine maleate, 11, 273 Guanabenz acetate, 15, 319
Ergotamine tartrate, 6, 113 Guar gurn, 24, 243
Erthromycin, 8, 159
Erthromycin estolate, 1, 101; Halcinonide, 8, 251
2, 573 Haloperidol, 9, 341
Estradiol, 15, 283 Halothane, 1, 119; 2, 573; 14, 597
Estradiol valerate, 4, 192 Heparin sodium, 12, 215
Estrone, 12, 135 Heroin, 10, 357
Ethambutol hydrochloride, 7, 231 Hexestrol, 11, 347
Ethynodiol diacetate, 3, 253 Hexetidine, 7, 277
Etodolac, 29, 105 Histamine, 27, 159
Etomidate, 12, 191 Homatropine hydrobromide, 16, 245
Etopside, 18, 121 Hydralazine hydrochloride, 8, 283
Eugenol, 29, 149 Hydrochlorothiazide, 10, 405
Hydrocortisone, 12, 277
Fenoprofen calcium, 6, 161 Hydroflumethaizide, 7, 297
Fenoterol hydrobromide, 27, 33 Hydroxyprogesterone caproate,
Flavoxoate hydrochloride, 28, 77 4, 209
Flecainide, 21, 169 Hydroxyzine dihydrochloride, 7, 319
Fluconazole, 27, 67 Hyoscyamine, 23, 155
Flucytosine, 5, 115
Fludrocortisone acetate, 3, 281 Ibuprofen, 27, 265
Flufenamic acid, 11, 313 Imipramine hydrochloride, 14, 37
Fluorouracil, 2, 221; 18, 599 Impenem, 17, 73
Fluoxetine, 19, 193 Indapamide, 23, 233
Fluoxymesterone, 7, 251 Indinivar sulfate, 26, 319
Fluphenazine decanoate, 9, 275; Indomethacin, 13, 211
10, 730 Iodamide, 15, 337
Fluphenazine enanthate, 2, 245; Iodipamide, 2, 333
4, 524 Iodoxamic acid, 20, 303
Fluphenazine hydrochloride, 2, 263; Iopamidol, 17, 115
4, 519 Iopanoic acid, 14, 181
Flurazepam hydrochloride, 3, 307 Ipratropium bromide, 30, 59
Flutamide, 27, 115 Iproniazid phosphate, 20, 337
Fluvoxamine maleate, 24, 165 Isocarboxazid, 2, 295
Folic acid, 19, 221 Isoniazide, 6, 183
Furosemide, 18, 153 Isopropamide, 2, 315; 12, 721
Isoproterenol, 14, 391 Mercaptopurine, 7, 343
Isosorbide dinitrate, 4, 225; 5, 556 Mesalamine, 25, 209; 27, 379
Isosuprine hydrochloride, 26, 359 Mestranol, 11, 375
Ivermectin, 17, 155 Metformin hydrochloride, 25, 243
Methadone hydrochloride, 3, 365;
Kanamycin sulfate, 6, 259 4, 520; 9, 601
Ketamine, 6, 297 Methaqualone, 4, 245
Ketoprofen, 10, 443 Methimazole, 8, 351
Ketotifen, 13, 239 Methixen hydrochloride, 22, 317
Khellin, 9, 371 Methocarbamol, 23, 377
Methotrexate, 5, 283
Lactic acid, 22, 263 Methoxamine hydrochloride, 20, 399
Lactose, anhydrous, 20, 369 Methoxsalen, 9, 427
Lansoprazole, 28, 117 Methylclothiazide, 5, 307
Leucovorin calcium, 8, 315 Methylphenidate hydrochloride, 10,
Levallorphan tartrate, 2, 339 473
Levarterenol bitartrate, 1, 149; Methyprylon, 2, 363
2, 573; 11, 555 Metipranolol, 19, 367
Levodopa, 5, 189 Metociopramide hydrochloride, 16,
Levothyroxine sodium, 5, 225 327
Lidocaine, 14, 207; 15, 761 Metoprolol tartrate, 12, 325
Lidocaine hydrochloride, 14, 207; Metronidazole, 5, 327
15, 761 Mexiletine hydrochloride, 20, 433
Lincomycin, 23, 275 Minocycline, 6, 323
Lisinopril, 21, 233 Minoxidil, 17, 185
Lithium carbonate, 15, 367 Mitomycin C, 16, 361
Lobeline hydrochloride, 19, 261 Mitoxanthrone hydrochloride, 17,
Lomefloxacin, 23, 327 221
Lomustine, 19, 315 Morphine, 17, 259
Loperamide hydrochloride, 19, 341 Moxalactam disodium, 13, 305
Lorazepam, 9, 397
Lovastatin, 21, 277 Nabilone, 10, 499
Nadolol, 9, 455; 10, 732
Mafenide acetate, 24, 277 Nalidixic acid, 8, 371
Malic Acid, 28, 153 Nalmefene hydrochloride, 24, 351
Maltodextrin, 24, 307 Nalorphine hydrobromide, 18, 195
Mandelic Acid, 29, 179 Naloxone hydrochloride, 14, 453
Maprotiline hydrochloride, 15, 393 Naphazoline hydrochloride, 21, 307
Mebendazole, 16, 291 Naproxen, 21, 345
Mebeverine hydrochloride, 25, 165 Natamycin, 10, 513; 23, 405
Mefenamic acid, 31, 281 Neomycin, 8, 399
Mefloquine hydrochloride, 14, 157 Neostigmine, 16, 403
Melphalan, 13, 265 Nicotinamide, 20, 475
Meperidine hydrochloride, 1, 175 Nifedipine, 18, 221
Meprobamate, 1, 207; 4, 520; 11, 587 Nimesulide, 28, 197
Nimodipine, 31, 337, 355, 371 Phytonadione, 17, 449
Nitrazepam, 9, 487 Pilocarpine, 12, 385
Nitrofurantoin, 5, 345 Piperazine estrone sulfate, 5, 375
Nitroglycerin, 9, 519 Pirenzepine dihydrochloride, 16, 445
Nizatidine, 19, 397 Piroxicam, 15, 509
Norethindrone, 4, 268 Polythiazide, 20, 665
Norfloxacin, 20, 557 Polyvinyl alcohol, 24, 397
Norgestrel, 4, 294 Polyvinylpyrollidone, 22, 555
Nortriptyline hydrochloride, 1, 233; Povidone, 22, 555
2, 573 Povidone-Iodine, 25, 341
Noscapine, 11, 407 Pralidoxine chloride, 17, 533
Nystatin, 6, 341 Praziquantel, 25, 463
Prazosin hydrochloride, 18, 351
Ondansetron hydrochloride, 27, 301 Prednisolone, 21, 415
Ornidazole, 30, 123 Primidone, 2, 409; 17, 749
Oxamniquine, 20, 601 Probenecid, 10, 639
Oxazepam, 3, 441 Pyrazinamide hydrochloride, 4, 333;
Oxyphenbutazone, 13, 333 28, 251
Oxytocin, 10, 563 Procaine hydrochloride, 26, 395
Procarbazine hydrochloride, 5, 403
Pantoprazole, 29, 213 Promethazine hydrochloride, 5, 429
Papaverine hydrochloride, 17, 367 Proparacaine hydrochloride, 6, 423
Particle Size Distribution, 31, 379 Propiomazine hydrochloride, 2, 439
Penicillamine, 10, 601 Propoxyphene hydrochloride, 1, 301;
Penicillin-G, benzothine, 11, 463 4, 520; 6, 598
Penicillin-G, potassium, 15, 427 Propyl paraben, 30, 235
Penicillin-V, 1, 249; 17, 677 Propylthiouracil, 6, 457
Pentazocine, 13, 361 Pseudoephedrine hydrochloride, 8,
Pentoxifylline, 25, 295 489
Pergolide Mesylate, 21, 375 Pyrazinamide, 12, 433
Phenazopyridine hydrochloride, 3, Pyridoxine hydrochloride, 13, 447
465 Pyrimethamine, 12, 463
Phenelzine sulfate, 2, 383
Phenformin hydrochloride, 4, 319; Quinidine sulfate, 12, 483
5, 429 Quinine hydrochloride, 12, 547
Phenobarbital, 7, 359
Phenolphthalein, 20, 627 Ranitidine, 15, 533
Phenoxymethyl penicillin potassium, Reserpine, 4, 384; 5, 557; 13, 737
1, 249 Riboflavin, 19, 429
Phenylbutazone, 11, 483 Rifampin, 5, 467
Phenylephrine hydrochloride, 3, 483 Rutin, 12, 623
Phenylpropanolamine hydrochloride,
12, 357; 13, 767 Saccharin, 13, 487
Phenytoin, 13, 417 Salbutamol, 10, 665
Physostigmine salicylate, 18, 289 Salicylamide, 13, 521
Salicylic acid, 23, 427 Thiabendazole, 16, 611
Scopolamine hydrobromide, Thiamine hydrochloride, 18, 413
19, 477 Thiamphenicol, 22, 461
Secobarbital sodium, 1, 343 Thiopental sodium, 21, 535
Sertraline hydrochloride, 24, 443 Thioridazine, 18, 459
Sertraline lactate, 30, 185 Thioridazine hydrochloride,
Sildenafil citrate, 27, 339 18, 459
Silver sulfadiazine, 13, 553 Thiostrepton, 7, 423
Simvastatin, 22, 359 Thiothixene, 18, 527
Sodium nitroprusside, 6, 487; Ticlopidine hydrochloride, 21, 573
15, 781 Timolol maleate, 16, 641
Solasodine, 24, 487 Titanium dioxide, 21, 659
Sorbitol, 26, 459 Tobramycin, 24, 579
Sotalol, 21, 501 a-Tocopheryl acetate, 3, 111
Spironolactone, 4, 431; 29, 261 Tolazamide, 22, 489
Starch, 24, 523 Tolbutamide, 3, 513; 5, 557;
Streptomycin, 16, 507 13, 719
Strychnine, 15, 563 Tolnaftate, 23, 549
Succinycholine chloride, 10, 691 Tranylcypromine sulfate, 25, 501
Sulfacetarnide, 23, 477 Trazodone hydrochloride, 16, 693
Sulfadiazine, 11, 523 Triamcinolone, 1, 367; 2, 571; 4, 521;
Sulfadoxine, 17, 571 11, 593
Sulfamethazine, 7, 401 Triamcinolone acetonide, 1, 397;
Sulfamethoxazole, 2, 467; 4, 521 2, 571; 4, 521; 7, 501; 11, 615
Sulfasalazine, 5, 515 Triamcinolone diacetate, 1, 423;
Sulfathiazole, 22, 389 11, 651
Sulfisoxazole, 2, 487 Triamcinolone hexacetonide, 6, 579
Sulfoxone sodium, 19, 553 Triamterene, 23, 579
Sulindac, 13, 573 Triclobisonium chloride, 2, 507
Sulphamerazine, 6, 515 Trifluoperazine hydrochloride,
Sulpiride, 17, 607 9, 543
Triflupromazine hydrochloride,
Talc, 23, 517 2, 523; 4, 521; 5, 557
Teniposide, 19, 575 Trimethaphan camsylate, 3, 545
Tenoxicam, 22, 431 Trimethobenzamide hydrochloride,
Terazosin, 20, 693 2, 551
Terbutaline sulfate, 19, 601 Trimethoprim, 7, 445
Terfenadine, 19, 627 Trimipramine maleate, 12, 683
Terpin hydrate, 14, 273 Trioxsalen, 14, 705
Testolactone, 5, 533 Tripelennamine hydrochloride,
Testosterone enanthate, 4, 452 14, 107
Tetracaine hydrochloride, 18, 379 Triprolidine hydrochloride, 8, 509
Tetracycline hydrochloride, Tropicamide, 3, 565
13, 597 Tubocurarine chloride, 7, 477
Theophylline, 4, 466 Tybamate, 4, 494
Valproate sodium, 8, 529 X-Ray Diffraction, 30, 271
Valproic acid, 8, 529 Xylometazoline hydrochloride,
Verapamil, 17, 643 14, 135
Vidarabine, 15, 647
Vinblastine sulfate, 1, 443; 21, 611 Yohimbine, 16, 731
Vincristine sulfate, 1, 463; 22, 517
Vitamin D3, 13, 655 Zidovudine, 20, 729
Zileuton, 25, 535
Warfarin, 14, 423 Zomepirac sodium, 15, 673
X Ray Crystallography
Nuclear Magnetic Resonance Spectroscopy
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