High-Performance Liquid Chromatography and Mass Spectrometry of Porphyrins, Chlorophylls and Bilins (Methods in Chromatography, Vol. 2)

High-Performance Liquid Chromatography and Mass Spectrometry of Porphyrins, Chlorophylls and Bilins METHODS IN CHROMA...

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High-Performance Liquid Chromatography and Mass Spectrometry of Porphyrins, Chlorophylls and Bilins

METHODS IN CHROMATOGRAPHY Series Editor: C. K. Lim (University of London, UK)

Published Vol. 1:

Advances in Liquid Chromatography: 35 Years of Column Liquid Chromatography in Japan edited by T. Hanai & H. Hatano

Vol. 2:

High-Performance Liquid Chromatography and Mass Spectrometry of Porphyrins, Chlorophylls and Bilins by C. K. Lim

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Methods in Chromatography – Vol. 2

High-Performance Liquid Chromatography and Mass Spectrometry of Porphyrins, Chlorophylls and Bilins

Chang Kee Lim University of London, UK

World Scientific NEW JERSEY

•

LONDON

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SINGAPORE

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BEIJING

•

SHANGHAI

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HONG KONG

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TA I P E I

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CHENNAI

Published by World Scientific Publishing Co. Pte. Ltd. 5 Toh Tuck Link, Singapore 596224 USA office: 27 Warren Street, Suite 401-402, Hackensack, NJ 07601 UK office: 57 Shelton Street, Covent Garden, London WC2H 9HE

British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library.

HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY AND MASS SPECTROMETRY OF PORPHYRINS, CHLOROPHYLLS AND BILINS Methods in Chromatography — Vol. 2 Copyright © 2010 by World Scientific Publishing Co. Pte. Ltd. All rights reserved. This book, or parts thereof, may not be reproduced in any form or by any means, electronic or mechanical, including photocopying, recording or any information storage and retrieval system now known or to be invented, without written permission from the Publisher.

For photocopying of material in this volume, please pay a copying fee through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA. In this case permission to photocopy is not required from the publisher.

ISBN-13 978-981-02-3068-5 ISBN-10 981-02-3068-0

Typeset by Stallion Press Email: [email protected]

Printed in Singapore.

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Preface

The porphyrins, chlorophylls, bilins and related tetrapyrroles are referred to as the pigments of life, colours of life and rainbow of life by various groups. They are vital for all living cells and are present in all living organisms. Natural and synthetic tetrapyrroles also have applications in foods, cosmetics, biotechnology, pharmaceuticals, diagnostics and medicine. Methods for their separation and characterisation therefore have a very wide area of applications. High-performance liquid chromatography (HPLC) with fluorescence detection or detection with a variable wavelength uv-visible detector is commonly used for their analysis. More recently, HPLC coupled with mass spectrometry (MS) has significantly improved the sensitivity, accuracy and specificity of tetrapyrrole analysis. At present, HPLC and HPLC/MS techniques have reached a stage for taking stock of their status and books devoted towards this end are lacking. The aim of this monograph is to provide practical HPLC and HPLC/MS protocols, and chromatographic and mass spectrometric reference data for the analysis, identification and characterisation of porphyrins, chlorophylls, bilins and related compounds. Much of the methods described for porphyrins and bile pigments are based on work from our own laboratory. I thank James Rideout, Dennis Wright, Famei Li, Jinli Luo, Rong Guo, Qiang Wang, Hong Cai, Aquib Razzaque, Russell Jones, Abel Gorchein, Francesco de Matteis, Gwyn Lord and Malcolm Danton for their contributions. Methods for chlorophylls and phycobilins are adopted, with suggestions for modifications where necessary, from literature sources described by experts in their particular areas of research. Chang Kee Lim v

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Contents

Preface ...................................................................................... Chapter 1.

1.1. 1.2. 1.3. 1.4.

1.5.

Introduction..................................................................... Nomenclature ................................................................. Biosynthesis of Porphyrins, Haem and Chlorophyll ......... Biosynthesis of Bilins in Animals and Plants .................... 1.4.1. Degradation of haem to bile pigments................ 1.4.2. Biosynthesis of bilins in plants, algae and cyanobacteria .............................................. 1.4.3. Degradation of chlorophyll in senescent higher plants....................................................... Function of Porphyrins and Other Tetrapyrroles ...............

Chapter 2.

2.1. 2.2. 2.3.

Structure, Distribution, Biosynthesis and Function ........................................................

High-Performance Liquid Chromatography of Porphyrins ........................................................

Introduction..................................................................... HPLC of Porphyrin Methyl Esters ..................................... HPLC of Porphyrin Free Acids ......................................... 2.3.1. Stationary phases for reversed–phase HPLC of porphyrins ............................................

vii

v

1 1 2 3 9 9 11 13 15

25 25 26 27 28

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2.3.2.

Mobile phases for reversed–phase HPLC of porphyrins ...................................................... 2.3.3. Procedure for optimisation of porphyrin separation by reversed-phase HPLC.................... 2.3.4. Retention behaviour of porphyrins in reversed–phase HPLC..................................... HPLC of Porphyrinogens...................................................

34 40

Chapter 3. Mass Spectrometry of Porphyrins............................

51

3.1. 3.2.

51

2.4

Introduction..................................................................... Fast Atom Bombardment (FAB) Mass Spectrometry of Porphyrins ................................................................... 3.3. Laser Desorption/Ionisation Time-of-Flight (LDI-TOF) Mass Spectrometry of Porphyrins .................... 3.4 Electrospray Ionisation Mass Spectrometry (ESI-MS) and HPLC/ESI-MS of Porphyrins ....................................... 3.4.1. HPLC/ESI-MS of porphyrins ................................ 3.4.2. ESI-MS/MS fragmentation of porphyrins.............. 3.4.2.1. ESI-MS/MS product ion spectrum and fragmentation pathways of uroporphyrin................... 3.4.3. ESI-MS/MS fragmentation of hydroxyuroporphyrins .................................... 3.4.3.1. ESI-MS/MS product ion spectrum and fragmentation pathways of meso–hydroxyuroporphyrin............ 3.4.3.2. ESI-MS/MS product ion spectrum of hydroxyacetic acid uroporphyrin.... 3.4.3.3. ESI-MS/MS product ion spectrum of α-hydroxypropionic acid uroporphyrin ...................................... 3.4.3.4. ESI-MS/MS product ion spectrum and fragmentation pathways of β-hydroxypropionic acid uroporphyrin ......................................

29 32

52 52 56 58 64

65 65

67 69

71

71

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Contents

3.4.3.5.

3.4.4. 3.4.5. 3.4.6. 3.4.7.

3.4.8. 3.4.9. 3.4.10. 3.4.11.

3.4.12.

Chapter 4.

4.1. 4.2.

ESI-MS/MS product ion spectra and fragmentation pathways of cis- and transhydroxyspirolactoneurochlorins........ 3.4.3.6. ESI-MS/MS product ion spectra and fragmentation pathways of cis- and transdihydroxyurochlorins........................ 3.4.3.7. Characteristic ESI-MS/MS product ions of hydroxyuroporphyrins........... ESI-MS/MS product ion spectra of ketoacid uroporphyrins ................................................... ESI-MS/MS of heptacarboxylic porphyrin.......... ESI-MS/MS of hydroxyheptacarboxylic porphyrins ........................................................ ESI-MS/MS of ketoacid heptacarboxylic porphyrins and formyl heptacarboxylic porphyrin.......................................................... ESI-MS/MS product ion spectrum and fragmentation pattern of coproporphyrin .......... ESI-MS/MS of hydroxymethylcoproporphyrin ..... ESI-MS/MS of β-ketopropionic acid coproporphyrin and formylcoproporphyrin....... ESI-MS/MS product ion spectrum and fragmentation pattern of protoporphyrin ............................................. HPLC/ESI-MS and MS/MS of meso-tetraphenylporphyrin derivatives .............

ix

74

75 80 80 83 86

86 89 93 93

93 97

Porphyrin Profiles in Blood, Urine and Faeces by HPLC and HPLC/ESI-MS................ 107

Introduction..................................................................... Sample Collection and Preparation ................................. 4.2.1. Preparation of urine samples .............................. 4.2.2. Preparation of faecal sample ..............................

107 108 109 110

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4.3.

High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

4.2.3. Extraction of porphyrins in plasma ..................... 4.2.4. Extraction of red blood cell porphyrins............... Porphyrin Excretion Patterns and Enzyme Assays in the Porphyrias .................................................. 4.3.1. Aminolaevulinic acid dehydratase deficiency porphyria (ADP) ................................ 4.3.1.1. Determination of ALA and PBG ......... 4.3.1.2. Determination of erythrocyte Zn-protoporphyrin by HPLC ............... 4.3.2. Acute intermittent porphyria (AIP) ...................... 4.3.3. Congenital erythropoietic porphyria (CEP)................................................... 4.3.4. Porphyria cutanea tarda (PCT) ............................ 4.3.4.1. Determination of uroporphyrinogen decarboxylase activity in erythrocytes by HPLC..................... 4.3.5. Hereditary coproporphyria (HCP) ....................... 4.3.5.1. HPLC assay of coproporphyrinogen oxidase activity in peripheral leucocytes .......................................... 4.3.6. Variegate porphyria (VP)..................................... 4.3.6.1. Determination of protoporphyrinogen oxidase by HPLC............................................. 4.3.7. Erythropoietic protoporphyria (EPP) .................... 4.3.7.1. HPLC assay for ferrochelatase in leucocytes or lymphocytes ............. 4.3.8. Mixed porphyria .................................................

Chapter 5.

5.1. 5.2.

111 112 113 113 113 119 119 123 127

133 135

140 143

144 146 147 149

Isolation and Characterisation of Protoporphyrin Glycoconjugates from Harderian Glands of Rodents by HPLC and HPLC/ESI-MS.................................. 155

Introduction..................................................................... 155 Extraction of Porphyrins from Harderian Glands.............. 159

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Contents

5.3. 5.4.

5.5.

HPLC Separation of Porphyrins from Rat Harderian Gland Extract .................................................. Identification of Protoporphyrin Glycoconjugates from Rat Harderian Glands by Capillary Electrophoresis and HPLC/ESI-MS ................................... 5.4.1. Characterisation of protoporphyrin xyloside isolated from Harderian glands........................... 5.4.2. Characterisation of protoporphyrin monoglucoside isolated from Harderian glands of the rat.................................................. Origin and Function of Protoporphyrin Glycoconjugates in Harderian Glands of Rodents ...........

Chapter 6.

6.1. 6.2. 6.3. 6.4. 6.5.

7.1. 7.2. 7.3. 7.4. 7.5. 7.6.

160

162 166

172 173

HPLC and HPLC/MS of Chlorophyll and Related Compounds ...................................... 177

Introduction..................................................................... Reversed-Phase HPLC of Chlorophyll and Related Compounds in Plant Extracts ........................................... Normal-Phase HPLC of Chlorophyll and Related Compounds ..................................................................... HPLC Separation of Fluorescent and Non–Fluorescent Chlorophyll Catabolites ................................................... HPLC/MS and MS/MS of Chlorophyll and Related Compounds .....................................................................

Chapter 7.

xi

177 177 181 183 185

HPLC and HPLC/MS of Bilins of Animal and Plant Origin................................................... 191

Introduction................................................................... HPLC Separation of Bilirubin IIIα, IXα and XIIIα Isomers .......................................................... HPLC Separation of Bilirubin Conjugates ...................... HPLC of Biliverdins ....................................................... HPLC of Biliviolins ........................................................ HPLC of Urobilinoids ....................................................

191 191 194 195 198 198

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7.7.

7.8. 7.9.

7.10. 7.11.

High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

Analysis and Determination of Bile Pigments in Biological and Clinical Samples by HPLC ................. 7.7.1. HPLC methods for the determination of conjugated and unconjugated bilirubin in serum ............................................. 7.7.1.1. Extraction and determination of conjugated and unconjugated bilirubin in serum following trans-esterification ............................ 7.7.1.2. Solid-phase extraction (SPE) and determination of conjugated and unconjugated bilirubin in serum ........................................... 7.7.1.3. Simultaneous determination of conjugated bilirubin, unconjugated bilirubin and biliprotein or δ-bilirubin by direct injection HPLC................................. HPLC/ESI-MS and MS/MS of Bile Pigments.................... Characterisation of Tetrapyrrole Pigments in Avian Eggshells.......................................................... 7.9.1. Extraction of pigments from eggshells............... HPLC and HPLC/MS of Phycobilins............................... HPLC separation of Phycobiliproteins............................

Chapter 8.

201

202

203

204

207 208 212 213 217 217

Future Directions of HPLC and Mass Spectrometry of Tetrapyrroles............................... 223

Index ......................................................................................... 227

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CHAPTER 1

Structure, Distribution, Biosynthesis, Catabolism and Function

1.1. Introduction Porphyrins are a large class of natural or synthetic pigments having a substituted aromatic macrocyclic ring consisting of four pyrrole residues linked together by four methine bridging groups1 (Fig. 1.1). They are deeply coloured (red or purple), fluorescent compounds with an intense and characteristic absorbance band between 390–425 nm (the Soret band or B band) and two to four much weaker bands (Q bands) between 480–700 nm. The Soret band, with extinction coefficient of 150,000 or more, is often used for the sensitive spectrophotometric detection of porphyrins following separation by high-performance liquid chromatography (HPLC). Porphyrins also have a characteristic fluorescence spectrum. Using an excitation wavelength of 400–405 nm and an emission wavelength of around 600 nm, a much higher intrinsic sensitivity of detection than absorption spectrophotometry can be achieved. A fluorescence detector is therefore preferred for the sensitive detection of porphyrins in chromatographic analysis. Porphyrins are widely distributed in nature. They occur as coloured pigments in the downs of young birds and in higher concentrations in feathers of birds such as Turacos2 (as the copper complex of uroporphyrin III), owls, and bustards.3 The eggshells of birds may also contain porphyrins and/or bile pigments,4,5 usually protoporphyrin IX and biliverdin IXα. The calcareous shells6 and pearls7 of shellfishes often contain porphyrins and the shells of some deepsea bivalves are found to contain high concentrations of pink or red 1

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High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

2

NH HN

NH HN

NH

N H

NH HN

NH HN

N

Pyrrole

Pyrroles

Porphyrinogen

N HN

Porphyrin

Figure 1.1. Structures of pyrrole, porphyrinogen and unsubstituted porphyrin macrocycle.

2

3

1 A

B

4

C

5

8 D 7

NH N

N HN

6

Fischer's numbering system

Figure 1.2.

5 6 7 3 4 2 21 22 8 N 9 1 NH 10 20 19 N HN 11 24 23 12 18 16 14 17 15 13 IUPAC systematic numbering system

The numbering of unsubstituted porphyrin macrocycle.

fluorescent porphyrin deposits, mainly uroporphyrin I. Petroporphyrins8 in coal, oil or shale are formed from dead plants and other photosynthetic organisms by diagenesis deep in the earth over millions of years. It has even been suggested that porphyrin is an ideal biomarker in the search for extraterrestrial life9 because of its presence in virtually all living organisms on Earth. The main physiological significance of porphyrins lies in the pathways of haem10,11 and chlorophyll biosynthesis,12–14 of which they can be considered as intermediary metabolites or oxidised by-products.

1.2. Nomenclature In the conventional Fischer system15,16 of nomenclature, the peripheral positions of the macrocyclic ring are numbered from 1 to 8 (Fig. 1.2). The four pyrrole rings are labelled A, B, C and D and the

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Structure, Distribution, Biosynthesis, Catabolism and Function

3

four methine bridges (the meso-positions) are designated α, β, γ and δ. Trivial names are given to porphyrins of biological and clinical importance and are commonly used.1 They are also used in this book (Table 1.1), unless otherwise stated. In the systematic IUPAC nomenclature17,18 all atoms, including the nitrogen atoms, are numbered. IUPAC system of naming allows a more precise description of a substituent on a carbon or nitrogen atom of the porphyrin macrocycle. Table 1.1 shows the trivial names and structures of some naturally occurring porphyrins.

1.3. Biosynthesis of Porphyrins, Haem and Chlorophyll The first step (Fig. 1.3) is the condensation of glycine with succinyl coenzyme A (CoA), a derivative of succinic acid, to form 5-aminolaevulinic acid (ALA or 5-ALA). The reaction is catalysed by the enzyme 5-aminolaevulinic acid synthase (ALA-S) in the matrix compartment of the mitochondrion. This pathway, called Shemin pathway,19,20 occurs in animals. In plants, the C5 pathway in which ALA is formed from glutamate occurs.21

COOH

COOH

NH2

Glutamate-1-semialdehyde

ALA-S

+ COOH

O SCoA Succiny-CoA

Glycine

H2N

O

5-Aminolevulinic acid

Shemin Pathway (Animals)

Glutamate-1-semialdehyde

aminotransferase Glutamyl-tRNAGlu reductase Glutamyl-tRNAGlu

tRNAGlu ligase

C 5 Pathway (Plants)

COOH

HOOC

NH2

Glutamate

Figure 1.3. The biosynthesis of 5-aminolaevulinic acid (ALA) from glycine and succinyl-CoA (Shemin pathway) and from glutamate (C5) pathway. ALA-S = 5-aminolaevulinic acid synthase.

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High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

Table 1.1. Trivial Names and Structures of some Naturally Occurring Porphyrins. 2

3

1 A

B

4

C

5

8 D

NH N

N HN

6

7

SIDE-CHAIN SUBSTITUENTS Porphyrin

1

2

3

4

5

6

7

8

Uroporphyrin I Uroporphyrin III Heptacarboxylic acid porphyrin I Heptacarboxylic acid porphyrin III (7d) Hexacarboxylic acid porphyrin I (6Iab) Hexacarboxylic acid porphyrin I (6Iac) Hexacarboxylic acid porphyrin III (6ad) Pentacarboxylic acid porphyrin I Pentacarboxylic acid porphyrin III (5abd) Pentacarboxylic acid porphyrin III (5abc) Pentacarboxylic acid porphyrin III (5acd) Pentacarboxylic acid porphyrin III (5bcd) Coproporphyrin I Coproporphyrin III Isocoproporphyrin Tricarboxylic acid porphyrin Protoporphyrin IX Mesoporphyrin IX Deuteroporphyrin IX

Ac Ac Ac Ac Me

Pr Pr Pr Pr Pr

Ac Ac Ac Ac Me

Pr Pr Pr Pr Pr

Ac Ac Ac Ac Ac

Pr Pr Pr Pr Pr

Ac Pr Me Pr Ac

Pr Ac Pr Me Pr

Me

Pr

Ac

Pr

Me

Pr

Ac

Pr

Me

Pr

Ac

Pr

Ac

Pr

Pr

Me

Ac Me

Pr Pr

Me Me

Pr Pr

Me Ac

Pr Pr

Me Pr

Pr Me

Me

Pr

Me

Pr

Me

Pr

Pr

Ac

Me

Pr

Ac

Pr

Me

Pr

Pr

Me

Ac

Pr

Me

Pr

Me

Pr

Pr

Me

Me Me Me Me Me Me Me

Pr Pr Et V V Et H

Me Me Me Me Me Me Me

Pr Pr Pr Pr V Et H

Me Me Ac Me Me Me Me

Pr Pr Pr Pr Pr Pr Pr

Me Pr Pr Pr Pr Pr Pr

Pr Me Me Me Me Me Me

Side-chain abbreviations: Me = methyl, Et = ethyl, Ac = CH2COOH, Pr = CH2CH2COOH, V = vinyl. The letters a,b,c and d denote the positions of the methyl group on rings A, B, C and D, respectively.

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Structure, Distribution, Biosynthesis, Catabolism and Function

COOH

COOH HOOC

COOH O

ALA Dehydratase

O H 2N NH2 5-Aminolevulinic acid (ALA)

Figure 1.4.

5

(ALA-D)

N NH2 H Porphobilinogen (PBG)

The biosynthesis of PBG from two molecules of ALA.

Two molecules of ALA are then condensed with each other (Fig. 1.4) in the soluble part of the cytoplasm to form the monopyrollic precursor, porphobilinogen (PBG). This reaction is catalysed by the enzyme 5-aminolaevulinic acid dehydratase (ALA-D) or porphobilinogen synthase (PBG-S).22 In the next step, four molecules of PBG condense together in a head-to-tail fashion (Fig. 1.5) to yield the symmetrical, linear tetrapyrrole, hydroxymethylbilane (HMB).23,24 This reaction is catalysed by porphobilinogen deaminase (PBG-D), also known as hydroxymethylbilane synthase (HMB-S).25,26 HMB is rearranged and cyclised to yield the asymmetrical uroporphyrinogen III (Fig. 1.5). The reaction is catalysed by uroporphyrinogen III synthase (Urogen III-S).27,28 In the absence of Urogen III-S the unstable HMB, with a half-life of less than 5 minutes at neutral pH, is cyclised spontaneously to the symmetrical and physiologically unimportant uroporphyrinogen I (Fig. 1.5). Uroporphyrinogen III is the common precursor24 in the biosynthesis of haem, sirohaem, the cofactor of sulphite and nitrite reductase, and vitamin B12 (Fig. 1.6). The type isomers I and III denote the arrangements of the four acetic acid groups and four propionic acid groups around the macrocyclic periphery (positions 1–8) of the porphyrins (Table 1.1). There are four possible arrangements and Fischer called these isomers type I, II, III and IV isomers. At this stage of haem biosynthesis, and also in the next three steps, the intermediates are porphyrinogens or hexahydroporphyrins.

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Ac

Pr Ac

Ac Pr COOH HOOC

NH2

N H

Porphobilinogen

PBG-D (HMB-S)

Ac

HO Pr

A

Ac Urogen III-S B

Pr

C

Ac

A

D

NH HN NH HN

B

C

Pr

Pr

Ac

Pr

Uroporphyrinogen III

NH HN

H D Ac

NH HN

Chemical

Pr

Ac

Pr

Hydroxymethylbilane

Ac

Pr

A

D Ac

B

Pr

C

Ac

NH HN NH HN

Pr

Uroporphyrinogen I

Figure 1.5. The biosynthesis of hydroxymethylbilane and uroporphyrinogen III. PBG-D = porphobilinogen deaminase; HMB-S = hydroxymethylbilane synthase; Urogen III-S = uroporphyrinogen III synthase; Ac = CH2COOH; Pr = CH2CH2COOH.

They are colourless, non-fluorescent intermediates in which the pyrrole rings are joined together by methylene rather than methine bridges (Fig. 1.1). The porphyrins are oxidative by-products at these stages and cannot be metabolised themselves. The four acetic acid groups of uroporphyrinogen III are sequentially decarboxylated to methyl groups in a step-wise fashion, starting from ring D through rings A, B and C, to give coproporphyrinogen III (Fig. 1.6).29 The reaction is catalysed by the cytoplasmic enzyme uroporphyringen decarboxylase (Urogen-D).30 Although the clockwise decarboxylation pathway from ring D through rings A, B and C is preferred,31 random decarboxylation has also been observed leading to a complex mixture of isomeric, hepta-, hexa- and penta-carboxylic acid porphyrinogen intermediates.32,33 There are four possible type III hepta-, six type III hexa- and four type III penta-carboxylic acid porphyrinogens.

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Structure, Distribution, Biosynthesis, Catabolism and Function

Ac

Pr Ac

Ac

A

D

NH HN NH HN

Pr

B

Ac

A

Urogen-D C

Ac

Pr Pr

Ac

H3C

Pr

D

NH HN NH HN

H3C

A

Urogen-D C

Pr

Ac

D

H3C

Pr

NH HN NH HN

B

C

Pr

Pr

Ac

Pr

6da

7d

Uroporphyrinogen III

Ac

Pr Pr

B

7

Urogen-D Pr

Precorrin-2

Sirohaem

Vitamin B12

H3C

H3C

A

D Pr

NH HN NH HN

CH3 B

A

Urogen-D C

CH3

Pr H3C

Pr

CH3

Pr

Coproporphyrinogen III

H3C

D

NH HN NH HN

Pr

B

C

Pr

Ac

Pr

5dab

Figure 1.6. The biosynthesis of coproporphyrinogen III by sequential decarboxylation of the four side-chain acetic acid groups of uroporphyrinogen III. Uroporphyrinogen I is similarly decarboxylated to coproporphyrinogen I. Urogen-D = uroporphyrinogen decarboxylase. The letters a, b, c, and d denote the position on which the acetic acid group on ring A, B, C, and D, respectively, has been decarboxylated to a methyl group.

Urogen-D is not specific for uroporphyrinogen III and the symmetrical uroporphyringen I is similarly decarboxylated to coproporphyrinogen I. Coproporphyrinogen III is taken up into the mitochondrion where the remaining steps of haem biosynthesis take place. Coproporphyrinogen III is converted into protoporphyrinogen IX, via 2-vinyl-4,6,7-tricarboxylic acid porphyrinogen, by oxidative decarboxylation of the two propionic acid groups on rings A and B to vinyl groups (Fig. 1.7).10,11 The reaction is catalysed by coproporphyrinogen oxidase (Coprogen-O). This enzyme is highly specific for the type III isomer and will not decarboxylate coproporphyrinogen I which will not be further metabolised. Protoporphyrinogen IX is then oxidised to protoporphyrin IX by protoporphyrinogen oxidase (Protogen-O)10,11 and finally haem is

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H3C

A

CH3

CH3

Pr NH HN

H3C

Pr

B

NH HN Coproporphyrinogen Oxidase

D

H3C

NH HN

Pr

NH HN

(Coprogen-O)

C

CH3

H3C

CH3

Pr

Pr

Pr

Protoporphyrinogen IX

Coproporphyrinogen III

Figure 1.7. The biosynthesis of protoporphyrinogen IX from coproporphyrinogen III.

CH3

CH3

CH3

H3C

H3C

H3C NH

NH HN

N

N Ferrochelatase

Protogen-O N

NH HN CH3

H3C Pr

Pr

Protoporphyrinogen IX

HN

Fe2+ CH3

H3C Pr

N Fe

N

N CH3

H3C

Pr

Protoporphyrin IX

HOOC

COOH

Haem

Figure 1.8. The biosynthesis of protoporphyrin IX and haem. Protogen-O = protoporphyrinogen oxidase.

produced by insertion of ferrous iron into protoporphyrin IX (Fig. 1.8), a step catalysed by the last enzyme of haem biosynthesis, ferrochelatase (FECH).10,11 Note that protoporphyrin IX is the only porphyrin formed in the pathway. An outline of the pathway of haem biosynthesis from glycine to haem is shown in Fig. 1.9. The first enzyme of the pathway, ALA-S, plays a key role in the regulation of haem biosynthesis. Haem, the end product, exercises a negative feedback control on its synthesis,34 by modulating the amount of ALA-S. Protoporphyrin IX is also a precursor in chlorophyll biosynthe12-14 sis via the formation of Mg-protoporphyrin IX followed by monomethyl esterification, methylation, vinyl group reduction and

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NH2

HOOC

ALA-S

+

Glycine

(HMB-S)

N NH2 H

5-Aminolevulenic acid

Porphobilinogen CH3

Fe

Ac

N

A

N

NH HN

D

Ac HOOC

COOH

Haem

Fe2+

Pr

Ac

Ac

A

NH HN

C

Ac

D

Pr

Pr Pr Uroporphyrinogen III

Mitochondria

CH3

CH3

H3C

Pr

H3C

N

N

NH HN

NH HN

Protoporphyrin IX

Ac

Pr

Pr

Protoporphyrinogen IX

Figure 1.9.

Pr NH HN NH HN

CH3

H3C Pr

CH3

H3C

NH HN CH3

H3C Pr

Urogen-D Pr

NH HN

Coprogen-O

HN Pr

C

Uroporphyrinogen I

Pr

NH HN

CH3 Pr

Pr

Ac

CH3

H3C

Protogen-O H3C

B NH HN

Urogen-D

NH

Ac

Ac

Pr Pr

B

C

Chemical

Ac

NH HN

Pr

CH3

H3C

Ferrochelatase

D

Urogen III-S Pr

N

B NH HN

H

HO

Pr Hydroxymethylbilane

Cytoplasm

H3C N

A

PBG-D

O

H2N

SCoA Succinyl-CoA

Ac

ALA-D

COOH

O

Ac

Pr

COOH

COOH

COOH

9

CH3

Pr

Pr

Coproporphyrinogen III

H3C

Pr

Coproporphyrinogen I

The haem biosynthetic pathway.

formation of a fifth ring (ring E) to give protochlorophyllide. In the presence of light protochlorophyllide is reduced to chlorophyllide a, which is esterified by phytyl diphosphate to form chlorophyll a (Fig. 1.10).

1.4. Biosynthesis of Bilins in Animals and Plants 1.4.1. Degradation of haem to bile pigments Bilins is the general term for open-chain tetrapyrroles17,18 derived from names given to bile pigments, the haem degradation products excreted in animal bile. In humans the conversion of haem derived from haemoglobin of effete red cells at the end of their life span to the green bile pigment biliverdin IXα occurs in the reticuloendothelial system by a series of reactions which are catalysed by the microsomal haem oxygenase system.35–37 A small proportion comes from the turnover of other haemoproteins, e.g. cytochrome P450s,

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CH3

CH3

H3C

CH3

H3C NH N

N

N

Mg chelatase Mg2+

HN

CH3

H3C

HOOC

H3C

N

Mg-Protoporphyrin IX methyltransferase

Mg N

N

CH3

H3C

COOH

HOOC

COOH

Mg-Protoporphyrin IX

Protoporphyrin IX

N

N Mg

N

N CH3

H3C

HOOC

COOCH3

Mg-Protoporphyrin IX monomethyl ester Mg-Protoporphyrin monomethyl cyclase

CH3

CH3 H3C

H3C N

Chlorophyll synthetase

Mg H

N

N CH3

H3C HH O

Vinyl reductase N

N

O COOCH3

N Mg

H

N

N

H3C

CH3 N

H

O COOCH3

N

H HOOC

Chlorophyll a

Figure 1.10.

N CH3

H3C

Chlorophyllide a

O

N Mg

H HOOC

Protochlorophyllide reductase H3C CH3 hv

O COOCH3

Protochlorophyllide a

The biosynthesis of chlorophyll a from protoporphyrin IX.

immature erythroid cells and free haem which turnover at a faster rate. Biliverdin IXα is reduced at the C-10 position to bilirubin IXα, the yellow bile pigment (Fig. 1.11). The reaction is catalysed by the cytosolic enzyme NADPH-dependent biliverdin reductase.38,39 Bilirubin forms extensive intra-molecular hydrogen bonds which give it a strongly hydrophobic property.40–42 It is insoluble in aqueous solution at physiological pH and is transported in blood tightly bound to albumin.43 Bilirubin is excreted in the bile into the intestine mainly as the polar bilirubin mono- and di-glucuronide conjugates after esterification of the C-8 and/or C-12 propionic acid side-chains with uridine diphosphate glucuronic acid in the liver. The reaction is catalysed by the microsomal enzyme bilirubin UDP-glucuronosyl transferase (UGT1A1).44,45 Xylose and glucose conjugates have also been detected in small quantities. In the intestine de-conjugation and sequential hydrogenation by intestinal flora results in a series of chromogens which on hydrogenation give a variety of faecal bile pigments with varying degree of double bond conjugation and colour46

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CH3

11

CH3 O O

H3C

haem oxygenase N

N

O2

H3C NH

H 2O

HN

Fe

+ Fe 2+ + CO N

N

N CH3

H3C

HN

NADPH NADP+ H3C

CH3

Biliverdin HOOC

COOH

HOOC

Heme

COOH

biliverdin reductase COOH CH3

H3C

O

2

3

1 N H

4 5

6

CH3 12

8

7 N H

COOH

9 10

11

13

NADPH NADP+

CH3 17

18

16 N 19 N 14 15 H H

O

Bilirubin

Figure 1.11.

The degradation of haem by haem oxygenase system.

ranging from the green-blue biliverdins, violet biliviolins, yellow bilirubins, orange urobilins to the colourless urobilinogens. The structures of these bile pigments are shown in Fig. 1.12.

1.4.2. Biosynthesis of bilins in plants, algae and cyanobacteria Haem oxygenase is present not only in animals but also in plants, algae and cyanobacteria.47–49 The same pathway of enzyme reaction that converts protohaem to biliverdin IXα is observed for all these organisms. From the same universal precursor, plants, algae and cyanobacteria convert biliverdin IXα into phycobilins,47,48 the openchain tetrapyrroles, which include phytochromobilin, phycoerythrobilin, phycocyanobilin, phycobiliviolin and phycourobilin. These, together with chlorophyll a and β-carotene, give the immense variety of colouration seen in algae and cyanobacteria, from blue-green, purple, red and orange to yellow.50

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COOH COOH CH3

H3C 2

3

7

CH3 12

8

13

COOH COOH CH3

17

CH3

H3C

O 1 N 4 6 N 9 16 N 19 O 11 N 14 5 10 15 H H H

O

N H

N H

N H

CH3

N H

N

CH3 N H

O

N H

O

N H

CH3

N H

N H

HOOC

COOH

CH3

H3C

O

COOH CH3 N

CH3 N H

O

i - urobilin

Biliviolin

HOOC

N H

CH3

H3C O

O

N H

HOOC

COOH

CH3

H3C

CH3

Bilirubin

Biliverdin

HOOC

CH3

18

N

H3C

CH3 N H

l - stercobilin

Figure 1.12.

O

O

COOH

CH3 N H

N H

CH3 N H

CH3 N H

O

urobilinogen

Structures of bile pigments.

In higher plants biliverdin IXα is converted to 3(Z )-phytochromobilin by the plastid-localised enzyme phytochromobilin synthase (PΦB synthase) which is a bilin 2,3-reductase (Fig. 1.13). This gives the ethylinene group on ring A essential for covalent linkage to apophytochrome, which occurred after isomerisation of 3(Z)phytochromobilin to 3(E )-phytochromobilin catalysed by a bilin 31,32 cis-trans isomerase.51,52 In red algae biliverdin IXα is first reduced to 15,16-dihydrobiliverdin IXα catalysed by a bilin 15,16-reductase.53 This is then followed by reduction at the 2,3-positions, catalysed by a bilin 2,3-reductase, to give 3(Z)-phycoerythrobilin (Fig. 1.13). It is also believed that the 3(Z)isomer is isomerised to 3(E)-phycoerythrobilin catalysed by a bilin 31,32isomerase.53,54 It has been shown that both the (Z)- and (E)-isomers are eventually converted into phycocyanobilins.47,48,53

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COOH COOH CH3

H3C 2

3

12

8

7

COOH COOH CH3

CH3

17

13

2,3-reductase

18

O 1 N 4 6 N 9 11 N 14 16 N 19 O 5 10 15 H H H

H

CH3

CH3

N H

N H

O

Biliverdin

H 16

N H

N H

15

O

CH3

O

H O

N H

N H

CH3 N

2

(15,16), (18 ,18 )-

O

CH3

CH3

N H

N H

N

3 (Z)-phycocyanobilin

N H

O

N H

CH3

CH3

N H

N H

CH3

CH3

31,32-isomerase O

O

N

N H

O

3 (E)-phycoerythrobilin (15,16), (181,182)-

isomerase CH3

N

N H

isomerase

COOH COOH

COOH COOH H

CH3

COOH COOH H

CH3

3 (Z)-phycoerythrobilin 1

CH3

3 (E)-phytochromobilin

COOH COOH CH3

CH3

N H

15,16-dihydrobiliverdin 2,3-reductase CH3

O

N H

COOH COOH CH3

CH3

N

N

COOH

CH3 N H

CH3

31,32-isomerase

COOH

O

CH3

3 (Z)-phytochromobilin

15,16-reductase

H3C

13

CH3 N H

31,32-isomerase O

H O

CH3

CH3

N H

N H

CH3 N

CH3 N H

O

3 (E)-phycocyanobilin

Figure 1.13. Biosynthetic pathways of bilins in plants, algae and cyanobacteria.

In cyanobacteria, only the 3(Z )-phycocyanobilin isomer is produced55 and the green algae Mesotanium caldariorum is reported to synthesise 3(Z)-phycocyanobilin directly from 3(Z)-phytochromobilin56 (Fig. 1.13).

1.4.3. Degradation of chlorophyll in senescent higher plants The degradation of chlorophylls to colourless nonfluorescent catabolites (NCCs)57–59 is an integral part of leaf senescence and fruit ripening processes. The breakdown pathway begins with de-phytylation of chlorophyll a by chlorophyllase (chlase), followed

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by the removal of Mg by a Mg-dechelating substance or Mgdechelatase to give phaeophobide a. Oxidative ring opening then takes place, catalysed by phaeophobide a oxygenase (PAO), with the conversion of phaeophobide a into the open-chain tetrapyrrole red chlorophyll catabolite (RCC). It has been suggested that chlorophyll b is reduced to chlorophyll a before entering the pathway through PAO.60 RCC is then reduced to a primary fluorescent chlorophyll catabolite (pFCC), catalysed by RCC reductase. Modification of several peripheral side-chains of pFCC occurred and the modified FCCs are transported to the vacuoles61 where, under weakly acidic conditions, they undergo rapid, stereospecific isomerisation to give ubiquitous NCCs (Fig. 1.14). The type of peripheral side-chain modifications within different NCCs are species specific.62,63

CH3 5

3 4 H3C 2 A 1 N

18

H3C O

D

N

CH3 3 4 H3C 2 A 1 NH

8

10

N 14

17 1615 H 132

H

173

7

B N 9 Mg

20

H 19

6

chlorophyllase

11

C 12 CH Mg-dechelation 3

E 13 131

O COOCH3

O

5

6

7

8

B N 9

H3C

10

20

H 19

N

HN

pheophobide a oxygenase

11

14 C 12 CH 18 3 H3C 17 1615 E 13 2 H 13 1 13 O H O COOCH3

D

H H3C O

HO

O CH3 O H NH HN N

HN CH3

H

H

O COOCH3

HO

Pheophobide a

red chlorophyll catabolite (RCC)

Chlorophyll a red chlorophyll catabolite reductase O CH3 O H NH HN

R1 H3C

R2

low pH

NH HN CH3

H3C

O

H3C H

H

O COOR3

HO

nonfluorescent chlorophyll catabolites (NCCs)

H H3C O

O CH3 O H NH HN N

HN CH3

H

H

O COOCH3

HO

primary fluorescent chlorophyll catabolite (pFCC)

Figure 1.14. The pathway of chlorophyll breakdown in higher plants. Sites of peripheral modifications as present in different NCCs are indicated (R1 – R3). R1 = CH2CH3; R2 = H, OH or O-glucosyl; R3 = H or CH3.

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1.5. Functions of Porphyrins and Other Tetrapyrroles The macrocyclic tetrapyrrole structure is ideal for the insertion of metal atoms to form metallo-complexes which are the prosthetic groups in the formation of metalloproteins and metalloenzymes where many essential biochemical processes and bioenergetic reactions of life take place. They are nature’s most important catalysts. Protoporphyrin IX complexes with iron to form the oxygen transport metalloprotein haemoglobin which uses reversible oxygen coordination to iron II for transport of oxygen to organs throughout the body. Myoglobin, found in large amounts in skeletal and cardiac muscles, stores oxygen for use when needed and transports oxygen by diffusion. Other haem containing proteins include the cytochromes, peroxidases, reductases and catalase, which carry out a wide range of important oxidation and reduction reactions vital for all living cells. Sirohaem is the cofactor of sulphite and nitrite reductases. Chlorophylls are magnesium tetrapyrrole complexes which capture and convert absorbed sunlight into usable energy in photosynthesis. Vitamin B12 or cyancobalamin, a cofactor in methyltrasferases, is a cobalt tetrapyrrole complex. Factor F430 is involved in methane formation in certain bacteria, and is a nickel tetraphyrrole complex. Uroporphyrinogen III is the common intermediate to all these cellular tetrapyrrole metal complexes. Bile pigments, especially bilirubin, possess significant antioxidant64–66 and anti-mutagenic properties.67 They are potent free radical scavengers and have been shown to inhibit the mutagenic effects of oxidants and aromatic mutagens such as poly aromatic hydrocarbons and heterocyclic amines. Bilirubin has been hypothesised to have a circadian regulation role in humans.68 The albumin-bound bilirubin resembles phytochromes, which set the biological clock in plants. In higher plants, phytochromobilin, the open-chain tetrapyrrole, is the chromophore of phytochrome which functions as a lightsensing pigment or photoreceptor in plant development.52,69–71 It has the ability to photo-interconvert between red and far-red light absorbing forms by sensing the ambient light conditions. Phytochrome-like

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molecules have also been identified in algae,72 ferns and mosses73 and cyanobacteria.74 Phycobiliproteins50 are a homologous family of phycobilinprotein complexes present in cyanobacteria,50 red algae,50 cryptomonsds,75 and some species of prochlorophytes.76 They are the light-harvesting antennae of these organisms50 with the open-chain tetrapyrrole chromophores covalently linked to protein molecules via cysteine residues. Phycobiliproteins, especially phycocyanin, the blue, lightharvesting pigment in cyanobacteria, rhodophytes and cryptophytes, are water soluble antioxidants with strong fluorescent properties. Phycocyanin has been investigated for potential applications in the food, cosmetic and biotechnology industries, and in diagnostic medicine because of these useful properties.77 Porphyrins and related compounds are excellent photosensitisers78–80 used in photodynamic therapy (PDT) of diseases, including cancer,81,82 dermatological conditions83,84 and wet age-related macular degeneration (AMD).85,86 PDT comprises exogenous administration of a light-absorbing compound (photosensitiser) which can accumulate in a target tissue. Light of wavelength matching its absorption characteristics is directed at the target tissue to photoactivate the sensitiser. This generates free radicals, especially singlet oxygen, at a rate that overwhelms tissue defence and causes cell death. PDT has been investigated as a new anti-microbial strategy87,88 for treating localised infections caused by MRSA and for modulating wound healing. Anti-microbial PDT has also been suggested as a possible method for eliminating pathogenic oral bacteria within the oral cavity.88 Porphyrin photosensitisers have been used for photocatalytic patterning.89 Porphyrins are excited to generate radical species that photocatalytically oxidise, and thereby pattern, chemistries in the local vicinity. The technique, suitable for a wide variety of substrates including proteins and cells, has potential applications in biological and medical sciences.

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References 1. Smith KM. General features of the structure and chemistry of porphyrin compounds. In: Porphyrins and Metalloporphyrins (Smith KM, ed); pp. 3–28. Elsevier Scientific Publishing Company, Amsterdam, Oxford, New York, 1975. 2. Nicholas REH and Rimington C. Isolation of unequivocal uroporphyrin III, a further study of turacin. Biochemical Journal 1952; 50: 194–201. 3. With TK. On porphyrins in the feathers of owls and bustards. International Journal of Biochemistry 1978; 9: 893–895. 4. Kilner RM. The evolution of egg colour and patterning in birds. Biological Reviews 2006; 81: 383–406. 5. McGraw KJ. The mechanics of uncommon colors: Pterins, porphyrins, and psittacofulvins. In: Bird Coloration (Hill GE and McGraw KJ, eds); vol. 1: Mechanisms and Measurements; pp. 354–398. Harvard University Press, Harvard, 2006. 6. Kennedy GY. Porphyrins in invertebrates. Annals of New York Academy of Sciences 1975; 244: 662–673. 7. Iwahashi Y and Akamatsu S. Porphyrin pigment in black-lip pearls and its application to pearl identification. Fisheries Science 1994; 60: 69–71. 8. Xu H, Yu D and Que G. Characterization of petroporphyrins in Gudao residue by ultraviolet-visible spectrophotometry and laser desorption ionization/time-of-flight mass spectrometry. Fuel 84(6): 647–652. 9. Suo Z, Avci R, Schweitzer MH and Deliorman M. Porphyrin as an ideal biomarker in the search for extraterrestrial life. Astrobiology 2007; 7(4): 605–615. 10. Shoolingin-Jordan PM and Cheung KM. Biosynthesis of heme. In: Comprehensive Natural Products Chemistry (Barton DHR, Nakanishi K and Meth-Cohn O, eds); vol. 4 (Kelly JW vol. ed): Amino Acids, Peptides, Porphyrins and Alkaloids; pp. 61–107. Elsevier, Amsterdam, 1999. 11. Ajioka RS, Phillips JD and Kushner JP. Biosynthesis of haem in mammals (review). Biochimica et Biophysica Acta 2006; 1763: 723–736. 12. Masuda T. Recent overview of the Mg branch of the tetrapyrrole biosynthesis leading to chlorophylls. Photosynthesis Research 2008; 96(2): 121–143.

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13. Tanaka R and Tanaka A. Tetrapyrrole biosynthesis in higher plants. Annual Review of Plant Biology 2007; 58: 321–346. 14. GomezMaqueo Chew A and Bryant DA. Chlorophyll biosynthesis in bacteria: The origins of structural and functional diversity. Annual Review of Microbiology 2007; 61: 113–129. 15. Fischer H and Orth H. Die Chemie des Pyrrols. Vol. II 1. Akademische Verlagsgesellschaft, Leipzip, 1937. 16. Fischer H and Orth H. Die Chemie des Pyrrols. Vol. II 2. Akademische Verlagsgesellschaft, Leipzip, 1940. 17. Moss GP. IUPAC Nomenclature of tetrapyrroles. Pure and Applied Chemistry 1987; 59: 779–832. 18. Moss GP. Nomenclature of tetrapyrroles: IUPAC-IUB recommendation 1986. European Journal of Biochemistry 1988; 178: 277–328. 19. Wriston Jr JC, Lack L and Shemin D. The mechanism of porphyrin formation. Further evidence on the relationship of the citric acid cycle and porphyrin formation. Journal of Biological Chemistry 1955; 215: 603–611. 20. Shemin D, Russell CS and Abramsky T. The succinate-glycine cycle. I. The mechanism of pyrrole synthesis. Journal of Biological Chemistry 1955; 215: 613–626. 21. Kannangara CG, Andersen RV, Pontoppidan B, Willows R and Wettstein D. Enzymic and mechanistic studies on the conversion of glutamate to 5aminolaevulinate. In: The Biosynthesis of Tetrapyrrole Pigments (Chadwick DJ and Ackrill K, eds); pp. 3–25. Ciba Foundation Symposium 180. John Wiley & Sons, New York, 1994. 22. Jaffe EK. Porphobilinogen synthase, the first source of haem’s asymmetry. Journal of Bioenergetics and Biomembranes 1995; 27(2): 169–179. 23. Battersby AR, Fookes, CJ, Matcham GW and McDonald E. Biosynthesis of the pigments of life: Formation of the macrocycle. Nature 1980; 285(5759): 17–21. 24. Battersby AR. Tetrapyrroles: The pigments of life (review). Natural Product Reports 2000; 17(6): 507–526. 25. Shoolingin-Jordan PM. Structure and mechanism of enzymes involved in the assembly of tetrapyrrole macrocycle (review). Biochemical Society Transactions 1998; 26(3): 326–336.

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26. Jordan PM and Warren MJ. Evidence for a dipyrrolemethane cofactor at the catalytic site of E. coli porphobilinogen deaminase. FEBS Letters 1987; 225: 87–92. 27. Shoolingin-Jordan PM, Warren MJ and Awan SJ. Discovery that the assembly of the dipyrrolemethane cofactor of porphobilinogen deaminase holoenzyme proceeds initially by the reaction of preuroporphyrinogen with the apoenzyme. Biochemical Journal 1996; 316: 373–376. 28. Shoolingin-Jordan PM. Porphobilinogen deaminase and uroporphyrinogen III synthase: Structure, molecular biology, and mechanism. Journal of Bioenergetics and biomembranes 1995; 27(2): 181–195. 29. Jackson AH, Sancovich HA, Ferramola AM, Evan, N, Games, DE, Matlin SA, Elder GH and Smith, SG. Macrocyclic intermediates in the biosynthesis of porphyrins. Philosophical Transactions of Royal Society of London, Series B Biological Sciences 1976; 273(924): 191–206. 30. Whitby FG, Phillips JD, Kushner JP and Hill CP. Crystal structure of human uroporphyrinogen decarboxylase. EMBO Journal 1988; 17: 2463–2471. 31. Luo J and Lim CK. Decarboxylation of uroporphyrinogen III by erythrocyte uroporphyrinogen decarboxylase. Evidence for a random decarboxylation mechanism. Biochemical Journal 1990; 268: 513–515. 32. Lash TD. Action of uroporphyrinogen decarboxylase on uroporphyrinogen-III: A reassessment of the clockwise decarboxylation hypothesis. Biochemical Journal 1991; 278(Pt 3): 901–903. 33. Luo J and Lim CK. Order of uroporphyrinogen III decarboxylation on incubation of porphobilinogen and uroporphyrinogen III with erythrocyte uroporphyrinogen decarboxylase. Biochemical journal 1993; 289: 529–532. 34. Burnham BF. Evidence for a negative feedback system in the control of porphyrin biosynthesis. Biochemical and Biophysical Research Communications 1962; 7: 351–356. 35. Tennen R, Marver HS and Schmid R. The enzymatic conversion of heme to bilirubin by microsomal heme oxygenase. Proceedings of the National Academy of Sciences USA 1968; 61: 748–755. 36. Maines MD. Heme oxygenase: Function, multiplicity, regulatory mechanisms, and clinical applications. FASEB Journal 1988; 2: 2557–2568.

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37. Ortiz de Montellano PR. Heme oxygenase mechanism: Evidence for an electrophilic, ferric peroxide species. Accounts of Chemical Research 1998; 31: 543–549. 38. Schmid R and McDonagh AF. The enzymatic formation of bilirubin. Annals of New York Academy of Sciences 1975; 244: 533–552. 39. Kutty RK and Maines MD. Purification and characterization of biliverdin reductase from rat liver. Journal of Biological Chemistry 1981; 256: 3956–3962. 40. Bonnett R, Davies JE and Hursthouse MB. Structure of bilirubin. Nature 1976; 262: 327–328. 41. Nogales D and Lightner DA. On the structure of bilirubin in solution. 13 C[1H] heteronuclear Overhauser effect NMR analyses in aqueous buffer and organic solvents. Journal of Biological Chemistry 1995; 270: 73–77. 42. Zunszain PA, Ghuman J, McDonagh AF and Curry S. Crystallographic analysis of human serum albumin complexed with 4Z,15E-bilirubinIXα. Journal of Molecular Biology 2008; 381: 394–406. 43. Brodersen. Bilirubin. Solubility and interaction with albumin and phospholipid. Journal of Biological Chemistry 1979; 254: 2364–2369. 44. Kamisako T, Kobayashi Y, Takeuchi K, Ishihara T, Higuchi K, Tanaka Y, Gabazza EC and Adachi Y. Recent advances in bilirubin metabolism research: The molecular mechanism of hepatocyte bilirubin transport and its clinical relevance. Journal of Gastroenterology 2000; 35: 9–664. 45. Basu NK, Kole L and Owen IS. Evidence for phosphorylation requirement for human bilirubin UDP-glucuronosyltransferase (UGT1A1) activity. Biochemical and Biophysical Research Communications 2003; 303(1): 98–104. 46. Stoll MS and Gray CH. The preparation and characterization of bile pigments. Biochemical Journal 1977; 163: 59–101. 47. Beale SI. Biosynthesis of phycobilins. Chemical Reviews 1993; 93: 785–802. 48. Beale SI. Biosynthesis of open-chain tetrapyrroles in plants, algae, and cyanobacteria. Ciba Foundation Symposium 1994; 180: 156–168. 49. Terry MJ, Linley P and Kohchi T. Making light of it: The role of plant haem oxygenases in phytochrome chromophore synthesis (review). Biochemical Society Transactions 2002; 30: 604–609.

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50. Glazer AN. Light guides. Directional energy transfer in a photosynthetic antenna. Journal of Biological Chemistry 1989; 264(1): 1–4. 51. Terry MJ and Lagarias JC. Holophytochrome assembly. Coupled assay for phytochromobilin synthesis in organelle. Journal of Biological Chemistry 1991; 266: 22215–22221. 52. Terry MJ, McDowell MD and Lagarias JC. 3(Z )- and 3(E )-phytochromobilin are intermediates in the biosynthesis of phytochrome chromophore. Journal of Biological Chemistry 1995; 270: 11111–11119. 53. Beale SI and Cornejo J. Biosynthesis of phycobilins. 15,16-dihydrobiliverdin IXα is a partially reduced intermediate in the formation of phycobilins. Journal of Biological Chemistry 1991; 266: 22341–22345. 54. Beale SI and Cornejo J. Biosynthesis of phycobilins. 3(Z )-phycoerythrobilin and 3(Z )-phycocyanobilin are intermediates in the formation of 3(E )-phycocyanobilin from biliverdin IXα. Journal of Biological Chemistry 266: 22333–22340. 55. Cornejo J and Beale SI. Phycobilin biosynthetic reactions in extracts of cyanobacteria. Photosynthesis Research 1997; 51: 223–230. 56. Wu S-H, McDowell MT and Lagarias JC. Phycocyanobilin is the natural precursor of the phytochrome chromophore in the green alga Mesotaenium caldariorum. Journal of Biological Chemistry 1997; 272: 25700–25705. 57. Oberhuber M, Berghold, J, Breuker K, Hörtensteiner S and Kräutler B. Breakdown of chlorophyll: A nonenzymatic reaction accounts for the formation of the colorless “nonfluorescent” chlorophyll catabolites. Proceedings of the National Academy of Sciences USA 2003; 100: 6910–6915. 58. Pruzinská A, Tanner G, Aubry S, Anders I, Moser S, Müller T, Ongania K-H, Kräutler B, Youn, J-Y, Liljegren, SJ and Hörtensteiner S. Chlorophyll breakdown in senescent Arabidopsis leaves. Characterization of chlorophyll catabolites and of chlorophyll catabolic enzymes involved in the degreening reaction. Plant Physiology 2005; 139: 52–63. 59. Kräutler B. Chlorophyll breakdown and chlorophyll catabolites in leaves and fruit. Photochemical & Photobiological Sciences 2008; 7(10): 1114–1120. 60. Hörtensteiner S. Chlorophyll breakdown in higher plants and algae. Cellular and Molecular Life Sciences 1999; 56: 330–347.

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61. Matile P, Ginsburg S, Schellenberg M and Thomas H. Catabolites of chlorophyll in senescing barley leaves are localized in the vacuoles of mesophyll cells. Proceedings of the National Academy of Sciences USA 1988; 85: 9529–9532. 62. Berghold J, Breuker K, Oberhuber M, Hörtensteiner S and Kräutler B. Chlorophyll breakdown in spinach: On the structure of five nonfluorescent chlorophyll catabolites. Photosynthesis Research 2002; 74: 109–119. 63. Berghold J, Eichmüller C, Hörtensteiner S and Kräutler B. Chlorophyll breakdown in tobacco: On the structure of two nonfluorescent chlorophyll catabolites. Chemistry & Biodiversity 2004; 1: 657–668. 64. Kaur H, Hughes MN, Green CJ, Naughton P, Foresti R and Motterlini R. Interaction of bilirubin and biliverdin with reactive nitrogen species. FEBS Letters 2003; 543(1–3): 113–119. 65. Stocker R. Antioxidant activities of bile pigments. Antioxidants and Redox Signaling 2004; 6(5): 841–849. 66. Mancuso C, Pani G and Calabrese V. Bilirubin: An endogenous scavenger of nitric oxide and reactive nitrogen species. Redox Report 2006; 11(5): 207–213. 67. Bulmer AC, Ried K, Blanchfield JT and Wagner K-H. The anti-mutagenic properties of bile pigments (review). Mutation Research 2008; 658: 28–41. 68. Grass F and Kasper S. Humoral phototransduction: Light transportation in the blood, and possible biological effects. Medical Hypotheses 2008; 71(2): 314–317. 69. Terry MJ, Wahleithner JA and Lagarias JC. Biosynthesis of plant photoreceptor phytochrome. Archives of Biochemistry and Biophysics 1993; 306: 1–15. 70. Furuya M. Phytochromes-their molecular species, gene families, and functions. Annual Review of Plant Physiology. Plant Molecular Biology 1993; 44: 617–645. 71. Quail PH, Boylan MT, Parks BM, Short TW, Xu Y and Wagner D. Phytochromes: Photosensory perception and signal transduction. Science 1995; 268: 675–680. 72. Rudiger W and Lopez-Figueroa F. Photoreceptors in algae. Photochemistry and Photobiology 1992; 55: 949–954.

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73. Schneider-Poetsch HAW, Marx S, Kolukisaoglu HU, Hanelt S and Braun B. Phytochrome evolution-phytochrome genes in ferns and mosses. Physiologica Plantarum 1994; 91: 241–250. 74. Yeh KC, Wu SH, Murphy JT and Lararias JC. A cyanobacterial phytochrome two-component light sensory system. Science 1997; 277: 1505–1508. 75. MacColl R, Eisele LE, Dhar M, Ecuyer J-P, Hopkins S, Marrone J, Barnard R, Malak H and Lewitus AJ. Bilin organization in crytomonad biliprotins. Biochemistry 1999; 38: 4097–4105. 76. Hess WR, Partensky F, van der Staay, GWM, Garcia-Fernandez JM, Börner T and Vaulot D. Coexistence of phycoerythrin and a chlorophyll α/β antenna in a marine prokaryote. Proceedings of the National Academy of Sciences USA 1996; 93: 11126–11130. 77. Eriksen NT. Production of phycocyanin-a pigment with applications in biology, biotechnology, food and medicine (mini-review). Applied Microbiology and Biotechnology 2008; 80: 1–14. 78. Bonnett R and Berenbaum M. Porphyrins as photosensitizers. Ciba Foundation Symposium 1989; 261(1): 277–280. 79. Nyman ES and Hynninen PH. Research advances in the use of tetrapyrrolic photosensitizers for photodynamic therapy. Journal of Photochemistry and Photobiology, B, Biology 2004; 73(1–2): 1–28. 80. Gorman SA, Brown SB and Griffiths J. An overview of synthetic approaches to porphyrin, phthalocyanine, and phenothiazine photosensitizers for photodynamic therapy. Journal of Environmental Pathology, Toxicology and Oncology 2006; 25(1–2): 79–108. 81. Sibata CH, Colussi VC, Oleinick NL and Kinsella TJ. Expert Opinion on Pharmacotherapy 2001; 2(6): 917–927. 82. Juzeniene A, Peng Q and Moan J. Milestones in the development of photodynamic therapy and fluorescence diagnosis. Photochemical and Photobiological Sciences 2007; 6(12): 1234–1245. 83. McCormack MA. Photodynamic therapy. Advances in Dermatology 2006; 22: 219–258. 84. McCormack MA. Photodynamic therapy in dermatology: An update on applications and outcomes. Seminars in Cutaneous Medicine and Surgery 2008; 27(1): 52–62.

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85. Brown SB and Mellish KJ. Verteporfin: A milestone in ophthalmology and photodynamic therapy. Expert Opinion on Pharmacotherapy 2001; 2(2): 351–361. 86. Yang YC. Preserving vision with verteporfin photodynamic therapy. Hospital Medicine 2004; 65(1): 39–43. 87. Maisch T, Bosl C, Szeimies RM, Love B and Abels C. Determination of the antibacterial efficacy of a new porphyrin-based photosensitizer against MRSA ex vivo. Photochemical & Photobiological Sciences 2007; 6(5): 545–551. 88. Maisch T. Anti-microbial photodynamic therapy: Useful in the future? Lasers in Medical Science 2007; 22(2): 83–91. 89. Bearinger JP, Stone G, Christian AT, Dugan L, Hiddessen AL, Wu JJ, Wu L, Hamilton J, Stockton C and Hubbell JA. Porphyrin-based photocatalytic lithography. Langmuir 2008; 24: 5179–5184.

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CHAPTER 2

High-Performance Liquid Chromatography of Porphyrins

2.1. Introduction High-performance liquid chromatography (HPLC) has been used by organic chemists for the separation of porphyrins and related tetrapyrroles since its early development1 in the late 1960s. By the early 1970s, with HPLC equipment available commercially, it gained a foothold in clinical chemistry laboratories and was used for the analysis of porphyrins from clinical samples in the diagnosis of porphyrias.2 Recent advances in HPLC column technology has led to the development of high efficiency columns. HPLC is now the technique of choice in porphyrin analysis and separation. The availability of sensitive and specific detectors such as the fluorescence detector and mass spectrometer allows the high resolution achieved by HPLC to be coupled with sensitive and specific detection, thus providing a very powerful method for the analysis and characterisation of porphyrins. The porphyrins may be separated as their methyl ester derivatives or as the underivatised free acids, depending on the applications required. Porphyrin methyl ester separations are usually performed in chemical synthesis where solubility in an organic solvent is important and more convenient, particularly when intermediates or products are isolated for further characterisation by nuclear magnetic resonance (NMR) spetroscopy. Porphyrin free acids separations are preferred when biological and clinical samples are analysed because they are usually extracted into an aqueous HCl solution and the extract can be analysed by reversed-phase HPLC without further manipulation. 25

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2.2. HPLC of Porphyrin Methyl Esters Porphyrin methyl esters are usually separated on silica columns (3–5 µm particle size) with organic solvent mixtures such as ethyl acetate/n-heptane, ethyl acetate/dichloromethane or chloroform as mobile phases. The proportion is adjusted to suit the required separation. For the separation of a complex mixture of porphyrins, gradient elution is employed, for example, from 30% (v/v) ethyl acetate to 90% ethyl acetate in 30 minutes using ethyl acetate and n-heptane as the gradient mixture. If reproducibility is important, as in the analysis of a batch of clinical samples, removal of traces of water in the organic solvent mixture used for elution, by distillation and drying, is important. Water adsorbed onto the silica stationary phase will gradually change its selectivity, and consequently the retention times of compounds. Reversed-phase HPLC is a better technique for the separation of porphyrin methyl esters because much higher resolution can be achieved. The separation of a mixture of porphyrin methyl esters on an octadecylsilica (ODS, C18) column by gradient elution from 70% acetonitrile in water to 100% acetonitrile in 30 minutes is shown in Fig. 2.1. The type I and type III isomers of hexacarboxylic porphyrin, pentacarboxylic porphyrin and coproporphyrin are easily resolved. The separation of uroporphyrin and heptacarboxylic porphyrin isomers, however, was not achieved. Apart from providing better resolution, reversed-phase HPLC requires a shorter solvent re-equilibration time than normal phase separation on silica and is also easier to maintain. Adsorption of polar compounds on silica columns, such as hydroxylated porphyrins and partially hydrolysed porphyrin methyl esters, is not a problem in reversed-phase HPLC. These compounds are eluted faster than the porphyrin methyl esters. Preparative isolation of porphyrin methyl esters by normal phase HPLC is convenient because the compounds can be recovered by simple solvent evaporation. In reversed-phase HPLC using the acetonitrile/water mixture, the isolated compounds may also be easily recovered by freezing the eluate in a freezer. The upper acetonitrile solution containing the porphyrins, which is not frozen, is decanted and recovered by evaporation.

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Figure 2.1. HPLC separation of a standard mixture of porphyrin methyl esters. Column: Hypersil-BDS-C18 (250 mm × 4.6 mm; 5 µm particle size). Linear gradient elution from 70% acetonitrile in water to 100% acetonitrile in 30 min at a flow rate of 1 ml/min. Detection: 404 nm. Peaks: 8 = uroporphyrin; 7 = heptacarboxylic porphyrin; 6I = hexacarboxylic porphyrin I; 6III = hexacarboxylic porphyrin III; 5I = pentacarboxylic porphyrin I; 5III = pentacarboxylic porphyrin III; 4I = coproporphyrin I; 4III = coproporphyrin III; m = mesoporphyrin; p = protoporphyrin.

Porphyrin methyl esters are usually prepared by treating the porphyrins with 5–10% sulphuric acid in methanol followed by solvent extraction. This procedure may lead to partially esterified derivatives, particularly those with a higher number of carboxyl groups like uroporphyrin. Structural modification may also occur, especially hydration of vinyl groups, methylation of hydroxyl groups, and trans-esterification of porphyrin conjugates. These side reactions could present complications in the identification and quantification of the porphyrins.

2.3. HPLC of Porphyrin Free Acids The type and arrangement of the side-chain substituent groups around the porphyrin macrocycle confer varying degrees of hydrophobicity on the porphyrin molecules. These differences in

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relative hydrophobicity are ideal for separation by reversed-phase chromatography.3–7 The porphyrins may be separated by differences in hydrophobic interaction between the side-chain substituents and the hydrocarbonaceous reversed-phase stationary phase surface3–7 by using an aqueous solution or buffer of controlled pH, or by forming hydrophobic ion-pairs with an ion-pairing reagent such as tetrabutylammonium phosphate,8 with suitable organic modifiers for elution.

2.3.1. Stationary phases for reversed-phase HPLC of porphyrins Porphyrins have been successfully chromatographed on silica chemically bonded with hydrocarbons ranging from C1 (trimethylsilyl) to C18 (octadecylsilyl) in chain length. Columns from almost all manufacturers are suitable. The selectivity of the columns, however, may differ from manufacturer to manufacturer, depending on the bonding chemistry used, the % carbon loading and whether they are endcapped or not. It is therefore important to optimise the separation using a test mixture, ideally consisting of uro-(I + III), copro-(I + III) and meso- (or proto-) porphyrins. Porphyrin standard mixtures for chromatography are also available commercially. The least hydrophobic C1-bonded phase is best for the fast separation of all porphyrins derived from the haem biosynthetic pathway, from uroporphyrin to protoporphyrin.9 The separation of a standard mixture of porphyrins on a C1-bonded phase is shown in Fig. 2.2. C18 or ODS columns are more retentive, but provide better resolution for porphyrins with 4 to 8 carboxylic acid groups (i.e., from copro- to uro-porphyrin). ODS columns are also more stable than C1 columns towards aqueous buffers in prolonged operation, and thus have longer column life. The most commonly used column dimension is 250 mm × 4.6 mm i.d., packed with 5 µm particle sized materials. Unless otherwise stated, this is the column dimension used in the separations described in the following sections. Columns of other dimensions, e.g., 100 mm × 2.0 mm packed with 3 µm ODS may also by used. More recently,

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Figure 2.2. HPLC separation of a standard mixture of porphyrins. Column: SAS-Hypersil (C1-bonded phase; 250 mm × 4.6 mm; 5 µm particle size). Solvent A, 10% (v/v) acetonitrile in 1 mol/l ammonium acetate, pH 5.16; solvent B, 10% (v/v) acetonitrile in methanol. Elution: 30 min linear gradient from 0% B to 65% B followed by isocratic elution at 65% B for a further 10 min. Flow rate: 1 ml/min. Detection: fluorescence excitation 400 nm, emission 618 nm. Peaks: 1 = uroporphyrin I; 2 = uroporphyrin III; 3 = heptacarboxylic porphyrin I; 4 = heptacarboxylic porphyrin III; 5 = hexacarboxylic porphyrin I; 6 = pentacarboxylic porphyrin I; 7 = coproporphyrin I; 8 = coproporphyrin III; 9 = deuteroporphyrin; 10 = mesoporphyrin; 11 = protoporphyrin.

2.1 and 1.7 µm particles have become available but have yet to be explored for porphyrin separation, especially in HPLC/MS application.

2.3.2. Mobile phases for reversed-phase HPLC of porphyrins Mobile phases are chosen to achieve optimal separation within convenient retention times. With the increased importance of HPLC/MS in porphyrin analysis, a mobile phase fully compatible with the mass spectrometer has become essential, and this rules out the involatile

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inorganic phosphate buffers commonly used in reversed-phase ionpair chromatography. The ion-pairing reagent also interferes with mass spectrometric detection. The ideal buffer should be sufficiently volatile and able to provide separation for the whole spectrum of porphyrins, including the type-isomers. One such buffer is ammonium acetate-acetic acid buffer,10 which has been studied in detail for the separation of porphyrins. Figure 2.2 is a typical example of reversedphase chromatography of porphyrins using ammonium acetate buffer and acetonitrile/methanol as mobile phase. Acetonitrile is an excellent organic modifier for reversed-phase chromatography of porphyrins. However, it should not exceed 35% (v/v) when used in conjunction with 1 M ammonium acetate buffer. It is immiscible with 1 M ammonium acetate above this proportion.7 Methanol, on the other hand, is completely miscible with 1 M ammonium acetate at all proportions. It is however, an unsuitable organic modifier for the elution of uroporphyrin because of its hydrogenbonding capability.11 Adsorption of layers of methanol onto the reversed-phase stationary phase surface resulted in extensive hydrogenbonding with the carboxylic acid groups of uroporphyrin (Fig. 2.3), leading to total retention. Porphyrins with less than 8 carboxylic acid groups are not affected by this phenomenon. A mixture of acetonitrile and methanol

Figure 2.3. Hydrogen bonding between side-chain carboxylic acid substituents of uroporphyrin and methanol adsorbed on the surface of the C18 stationary phase. A = acetic acid; P = propionic acid.

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as organic modifier overcomes both the miscibility and uroporphyrin elution problems, as shown in Fig. 2.2. Acetonitrile is more hydrophobic than methanol and is preferentially adsorbed onto the stationary phase surface, thus preventing hydrogen-bonding with uroporphyrin. The following gradient mixtures are recommended for the separation of porphyrins by gradient elution: 10% acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16 (solvent A) and 10% acetonitrile in methanol (solvent B) or 9% acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16 (solvent A) and 9% acetonitrile in methanol (solvent B). The system is very flexible and can be easily modified by adjusting the pH, buffer concentration and types and proportions of organic modifiers in the mobile phase to suit all applications. It should be pointed out that when acetonitrile is used as the sole organic modifier with 1 M ammonium acetate, the column must not be washed with acetonitrile alone at the end of the operation because of the problem of immiscibility. The column can be washed and stored in 10% acetonitrile in methanol/water (95:5, v/v). Buffer solutions, including ammonium acetate, contain trace amounts of metallic elements, usually Cu, Fe, Mg and Zn. In small scale preparative isolation of porphyrins, a small quantity of ethylenediaminetetraacetic acid (EDTA) must be added to the collected fractions to prevent metalloporphyrin formation.4 No metalloporphyrins are formed during the HPLC run, but it can be demonstrated that metalloporphyrins are formed in ammonium acetate buffer when left standing for 2 hours.4 Acidic eluents, e.g., 0.05 or 0.1% trifluoroacetic acid (TFA) in acetonitrile, may also be used for the separation of porphyrins, especially for the fast separation of dicarboxylic acid porphyrins such as protoporphyrin and its derivatives, for example, protoporphyrin amino acid- and peptide-conjugates12 and protoporphyrin glycoconjugates.13 These mobile phases, however, are unable to separate uroporphyrin and heptacarboxylic porphyrin I and III isomers, although the resolution of type I and III isomers of hexacarboxylic-, pentacarboxylic- and copro-porphyrins has been achieved.

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2.3.3. Procedure for optimisation of porphyrin separation by reversed-phase HPLC There are three parameters important for controlling the retention and resolution of porphyrins by reversed-phase HPLC. These are the molar concentration of the buffer, pH and the type and proportion of organic modifier used.3–7 The ammonium acetate buffer system was optimised based on these parameters. The efficiency of a conventional ODS column can be improved by inclusion of ammonium acetate in the mobile phase. It is an effective masking agent for residual silanol groups present on these columns. It has been shown that the optimum pH for ammonium ion adsorption, and therefore exerting its maximum masking effect on conventional ODS columns, is around 5.10–5.20. The fully endcapped ODS columns can also benefit from the use of ammonium acetate. This pH range should be used whenever possible. The porphyrins are zwitterions and their state of ionisation is pH-dependent. Complete ionisation of a porphyrin results in no retention, while complete suppression of ionisation may lead to excessive retention. Studies on the separation of copro- and uroporphyrin isomers have shown that the retention and resolution of these porphyrins decreased with increasing pH and no isomer separation is possible at pH above 5.80. The optimum pH for the simultaneous separation of all porphyrins, particularly for porphyrins with 4 to 8 carboxylic acid groups by gradient elution is 5.16. For dicarboxylic porphyrins, especially in isocratic elution, the pH optimum is around 4.6. The separation of haematoporphyrin diastereoisomers and their derivatives shown in Fig. 2.4 is a typical example.14 The molar concentration of ammonium acetate buffer significantly affected column efficiency and porphyrin retention. Increasing the molar concentration decreases the capacity ratio (k′ ) values of all porphyrins while maintaining the desired resolution, especially in isocratic separation. Earlier studies have concluded that 1 M ammonium acetate provides the best results in terms of peak symmetry, resolution and speed of separation. Peak broadening was observed

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Figure 2.4. HPLC separation of haematoporphyrin diastereoisomers and their derivatives. Column: MOS- Hypersil (C8-bonded phase). Mobile phase: methanol/1 M ammonium acetate, pH 4.6 (60:40, v/v). Flow rate: 1 ml/min. Peaks: 1 = (RS + SR )-haematoporphyrin, 2 = (RR + SS )-haematoporphyrin, 3 = (RS + SR )-3-(1-acetoxyethyl)-8-hydroxyethyldeuteroporphyrin, 4 = (RR + SS )-3-(1-acetoxyethyl)-8-hydroxyethyldeuteroporphyrin, 5 = (RS + SR )8-(1-acetoxyethyl)-3-hydroxyethyldeuteroporphyrin, 6 = (RR + SS )-8-(1acetoxyethyl)-3-hydroxyethyldeuteroporphyrin, 7 = 8-(1-hydroxyethyl)-3vinyldeuteroporphyrin, 8 = 3-(1-hydroxyethyl)-8-vinyldeuteroporphyrin, 9 = (RS + SR )-diacetyldeuteroporphyrin, 10 = (RR + SS )-diacetyldeuteroporphyrin, 11 = 8-(1-acetoxyethyl)-3-vinyldeuteroporphyrin, 12 = 3-(1acetoxyethyl)-8-vinyldeuteroporphyrin. The structure of haematoporphyrin is shown.

when 0.5 M ammonium acetate buffer was used in the isocratic separation of uro- and copro-porphyrin isomers. This problem can, however, be minimised by using gradient elution. A higher molar concentration of ammonium acetate is not only more effective in silanol masking but also provides better column protection, allowing acid solution of porphyrins to be injected. Acetonitrile and methanol are the two most commonly used organic modifiers for reversed-phase HPLC of porphyrins. Either acetonitrile alone or a mixture of acetonitrile and methanol can be used, depending on the application. The proportion of organic modifier in

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the buffer should be adjusted for maximum resolution within the shortest possible retention time in both isocratic and gradient elution. Recent advances in HPLC column technology has led to significant improvements in reversed-phase column efficiency, stability and tolerance to a wide range of pH. Columns allowing separation in the pH range of 2–12 are now common. The selectivity of columns from different suppliers, however, may differ significantly and the above procedure should be used to develop and optimise separation on each individual column. Some of these may not require a high molar concentration of ammonium acetate to achieve column efficiency and, consequently, acetonitrile can be used as an organic modifier with or without methanol.

2.3.4. Retention behaviour of porphyrins in reversed-phase HPLC Hydrophobic interaction between the porphyrin side-chain substituents and the non-polar hydrophobic hydrocarbonaceous stationary phase surface, is the main retention mechanism.5 The number and arrangement of the most hydrophobic groups around the rigid macro-cyclic porphyrin molecules determine their relative elution orders. Referring to Figs. 2.1 and 2.2, the following elution order is observed: uroporphyrin, heptacarboxylic porphyrin, hexacarboxylic porphyrin, pentacarboxylic porphyrin, coproporphyrin, deuteroporphyrin, mesoporphyrin and protoporphyrin. The retention increases with rising number of methyl, ethyl and vinyl groups, i.e., hydrophobic groups. The hydrophobicity of the common porphyrin side-chain substituents increases in the order of CH2COOH < CH2CH2COOH < CH3 < CH2CH3 < CH=CH2. The hydrophobic interaction mechanism is best demonstrated by the elution of the 4 type-isomers of coproporphyrin in the order of I, III, IV and II (Fig. 2.5). Coproporphyrin II has two pairs of adjacent CH3 groups, on positions 1,8 and 4,5, respectively (Fig. 2.6). This arrangement provides the largest hydrophobic surface area among the 4 isomers available for interaction. It is therefore the strongest retaining compound. The symmetrical type I isomer has no adjacent CH3 groups. It has the smallest hydrophobic surface area exposed for interaction and

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Figure 2.5. HPLC separation of coproporphyrin isomers. Column: Hypersil-ODS (C18). Mobile phase: 26% acetonitrile in 1 M ammonium acetate/acetic acid buffer, pH 5.16. Flow rate 1 ml/min.

Pr

3

CH3

H3C 1

Pr NH

H3C 1

5

Pr 7 H3C

Pr NH

N HN

N

CH3 3

Pr

CH3

H3C

Pr

Pr

N

Figure 2.6.

NH

HN

5 Pr

4-II

4 CH 3

H3C 1

N

Pr

Pr

Pr

4 CH 3

H3C 1

8

CH3

4-III

Pr

H3C

5

Pr

4-I

NH

HN

N

8

Pr

N

N CH3

N HN

Pr

6 CH3

7 H3C

Pr

4-IV

Structures of coproporphyrin I, II, III, and IV isomers.

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is the fastest eluting isomer. Coproporphyrin III and IV have one pair of adjacent CH3 groups each, on positions 1,8 and 6,7 respectively. Their hydrophobicities are therefore quite similar. However, when the arrangements of all four CH3 groups is taken into consideration, it clearly shown that the distances between the four CH3 groups on coproporphyrin IV are closer than those on coproporphyrin III. The difference in distance is only one bond length, but this is sufficient to impart a slightly stronger hydrophobicity on the type IV isomer for it to retain longer than the type III isomer. This retention mechanism was confirmed by the separation of the tetramethyl ester derivatives of the four type-isomers of coproporphyrin when the same elution order was observed. The elution orders of pentacarboxylic (Fig. 2.7) and hexacarboxylic porphyrin isomers (Fig. 2.9) can be similarly

Figure 2.7. HPLC separation of pentacarboxylic porphyrin isomers. Column: Hypersil-BDS (C18). Mobile phase: acetonitrile/methanol/1 M ammonium acetate–acetic acid buffer, pH 5.16 (4.5:40.5:55, by vol). Flow rate 1 ml/min. See structures in Fig. 2.8 for peak identification.

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High-Performance Liquid Chromatography of Porphyrins

CH3

Pr H3C

H3C

A

B NH N

D

NH

C

CH3 Pr

Pr

5abd

NH N

N

HN Pr

Pr

5abc

Pr

Pr NH

CH3

CH3

CH3

H3C

N

Ac

HN

H3C

5bcd

Pr Pr

H3C

N

N

Pr

5acd

CH3

Pr

NH

H3C

Pr

Pr

Pr

Ac

N HN

N Ac

CH3

Pr Pr

H3C

N HN

Ac

Pr Pr

37

N HN CH3

Pr Pr

Ac

5I

Figure 2.8. Structures of pentacarboxylic porphyrin isomers. 5I = pentacarboxylic porphyrin I; 5abc, 5abd, 5acd, and 5bcd are type-III isomers. The letters a, b, c, and d denote the position of side-chain CH3 substituents on ring A, B, C, and D, respectively.

explained. Their structures are shown in Figs. 2.8 and 2.10, respectively. For heptacarboxylic porphyrins with a single CH3 group (Fig. 2.11) which dominates the interaction and retention, the capacity values (k′ ) are very similar. The separation of the four type-III isomers has not been achieved, although complete resolution of type-I and type-III (as a group) isomers is not a problem (Fig. 2.12). For uroporphyrins (Fig. 2.13) with 8 carboxylic acid groups and no CH3 substituents, the relative hydrophobicity is determined by the arrangement of the propionic acid groups. Uroporphyrin III, with a pair of adjacent propionic acid groups on positions 6,7, is more hydrophobic than the symmetrical type I isomer with no adjacent propionic acid group (Fig. 2.14). Fast separation of uroporphyrin I and III isomers can also be achieved by isocratic elution on a

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Figure 2.9. HPLC separation of hexacarboxylic acid porphyrin isomers. Column: Hypersil-ODS. Eluent: 16% (v/v) acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16. Flow rate: 1 m/min. Peak: 1 = 6-Iac, 2 = 6-IIIac + 6-IIIbd, 3 = 6-IIIcd, 4 = 6-Iab, 5 + 6 = 6-IIIab + 6-IIIbc and 7 = 6-IIIad (see Fig. 2.10 for structures).

Hypersil-BDS column with 9% acetonitrile in 1 M ammonium acetate-acetic acid buffer, pH 5.5, as eluent (Fig. 2.15). For metalloporphyrins, the insertion of a metal ion which completely occupies the centre of the porphyrin hole alters the electronic environment around the central nitrogen atoms of the porphyrins (Fig. 2.16). The retention is then also influenced by the ability of the species of the inserted metal ion, to accept axial ligands from the mobile phase (Fig. 2.16), in addition to hydrophobic interaction of the side-chain substituents with the stationary phase surface.6 The separation of the Co(III), Fe(III), Zn(II) and Cu(II) complexes of meso- and proto-porphyrins is shown in Fig. 2.17. An elution order of Co, Fe, Zn and Cu complexes is observed for both meso- and protoporphyrins. This is consistent with the fact that Co(III) and Fe(III) complexes are particularly good in accepting axial ligands in solution and may add two extra ligands; Zn(II) complex can add one extra ligand, while further coordination of the Cu(II) complex is only

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CH3

Pr H3C

B

A

NH

D

Ac

N

Pr

Pr NH

C

Pr

CH3

Ac

N

N

HN Pr

Ac

6-Iab

Pr

Pr NH

Ac

CH3

Ac

H3C

N

Pr

HN

H3C

6-IIIcd

Pr Pr

H3C

N

Pr

Pr

6-IIIbd CH3

NH

N Ac

Pr

Pr

Pr NH

H3C

6-IIIbc

Ac

Ac

N HN

N

HN

Pr

Pr

Pr Pr

Ac

N

Pr

Ac Pr

CH3

NH

HN

6-IIIad

Pr Pr

N

H3C

6-IIIac

Ac

N

N

Pr

CH3

NH

HN CH3

6-IIIab Pr

NH

Ac

Pr

Pr

Pr

H3C

N

N Ac

Ac

Pr

H3C

N HN

Ac

Pr

39

N HN CH3

Pr Pr

Ac

6-Iac

Figure 2.10. Structures of hexacarboxylic porphyrin isomers. 6Iab and 6Iac are type-I and 6ab, 6ac, 6ad, 6bc, 6bd, and 6cd are type-III isomers. The letters a, b, c, and d denote the position of CH3 groups.

possible under special conditions. The addition of polar axial ligands, e.g., OH−, decreases the hydrophobicity and therefore in k′ values. Protoporphyrin is more hydrophobic than mesoporphyrin. Znprotoporphyrin and Cu-protoporphyrin are also more hydrophobic and have longer retention times than Zn-mesporphyrin and Cu-mesoporphyrin. However, for the Co and Fe complexes, the reverse is observed with the protoporphyrin complexes eluting before the mesoporphyrin complexes. This is attributed to a decrease in electron

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High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

H3C

Ac

A

D

NH

Pr

B

Pr

C

Ac

NH

7-IIIa

Pr

NH N

7-IIIb

7-IIId

N HN

N

Pr

Pr

7-IIIc Pr

HN

Pr

CH3 Pr

H3C

N

Ac

HN

Ac

NH

H3C

N

Ac

Pr

Pr Pr

Ac

N Ac

Ac

Pr

NH

Ac

Pr

Pr

Ac

N HN

N

Ac

Pr Pr

Ac

N HN

N

CH3

Pr

Ac

Pr

Ac

Pr Pr

Ac

7-I

Figure 2.11. Structures of heptacarboxylic porphyrin isomers. 7I = heptacarboxylic porphyrin I; 7a, 7b 7c, and 7d are type-III isomers. The letters a, b, c, and d denote the position of CH3 groups.

density at the ring nitrogens due to the vinyl groups of protoporphyrin which is reflected in the chelated metals, leading to an increased affinity for the donor electrons of extra ligands and therefore a decrease in hydrophobicity in comparison to the mesoporphyrin complexes. Understanding the retention behaviour is useful for the preliminary assignment of possible side-chain substituent groups on the porphyrin ring.

2.4. HPLC of Porphyrinogens Porphyrinogens are usually not separated by HPLC probably because they are relatively unstable to oxidation in solution. However, they are stable for at least an hour when kept on ice under N2 in the dark.

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Figure 2.12. HPLC separation of heptacarboxylic porphyrin I and III isomers. Column: Hypersil-BDS. Eluent: 15% acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16. Flow rate 1 ml/min.

Ac

Pr

Pr

Ac NH N

Ac

Ac

N

NH

HN

N Ac

Pr Ac

Pr

HN Ac

Pr

Pr

8-II

8-I

Figure 2.13.

Ac

A

N

Ac

Ac

Pr

Pr

Pr

Ac

D

B NH N

N HN

Pr

Pr

Pr Pr

Ac

Ac NH

C Pr

Ac

N

N HN Pr

Pr Ac

8-III

Ac

8-IV

Structures of uroporphyrin isomers.

Earlier studies have shown that isomers of porphyrinogens are much better separated than the corresponding porphyrins.15,16 The separation of porphyrinogens can therefore be used to identify and to confirm the structures of isomers, especially in situations

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Figure 2.14. HPLC separation of uroporphyrin isomers. Column: HypersilODS (C18). Eluent: 13% acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16. Flow rate 1 ml/min.

where complete separation of porphyrin isomers is difficult to achieve. Porphyrinogens are usually prepared by reduction of porphyrins with sodium amalgam (typically 3% w/w) under N2 as follows: 1. Dissolve porphyrin in 3 mM ammonium hydroxide or 10 mM KOH in a stopper-test tube. 2. Add 3% sodium amalgam and flush solution with N2. 3. Shake the mixture vigorously until a clear, colourless solution showing no fluorescence under an UV lamp is obtained. 4. Flush solution with N2 and keep on ice in the dark. Figure 2.18 shows the separation of the four type-isomers of coproporphyrinogen. The elution order of I, II, III, IV is different from

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Figure 2.15. HPLC separation of uroporphyrin I and III isomers on HypersilBDS (C18) with 9% acetonitrile in 1 M ammonium acetate–acetic acid buffer, pH 5.16 (a) and pH 5.5 (b), as mobile phase.

CH3

CH3

CH3

H3C

H3C

H3C NH

N

N

M2+ N

HN

N CH3

H3C

HOOC

COOH

N

N

L

M

HOOC

N

N L N

N CH3

H3C

L M

COOH

CH3

H3C

HOOC

COOH

Figure 2.16. Formation of the equatorial coordination groups and axial coordination groups of a metalloporphyrin. M = Fe, Co, Cu, Zn; L = axial ligand, e.g., OH− or CH3COO−.

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Figure 2.17. HPLC separation of dicarboxylic porphyrins and metalloporphyrins. Column: Hypersil-SAS (C1). Eluent : solvent A, methanol; solvent B, 1 M ammonium acetate-acetic acid buffer (pH 4.6). Elution: 0 to 6 min, 62% A; 6.1 to 13 min, 70% A; 13.1 to 30 min, 75% A. Peaks: 1 = Co-protoporphyrin, 2 = Co-mesoporphyrin, 3 = haemin, 4 = mesohaem, 5 = deuteroporphyrin, 6 = Zn-mesoporphyrin, 7 = Zn-protoporphyrin, 8 = mesoporphyrin, and 9 = protoporphyrin.

that observed for the corresponding coproporphyrins (see Fig. 2.5). While hydrophobic interaction is still expected to be the main retention mechanism, it is complicated by the fact that porphyrinogens are much more flexible molecules than the rigid porphyrin structures. The small CH3 substituents in each isomer may therefore be subjected to varying degrees of steric hindrance or shielding by the larger propionic acid groups, depending on the conformation adopted by the molecule. This alters the expected hydrophobic area available for interaction with the stationary phase surface, making prediction of elution order based on the arrangement of CH3 groups around the molecule difficult. The conformation of porphyrinogens under HPLC conditions is not known. The separation of pentacarboxylic porphyrinogen is shown in Fig. 2.19. The elution order of 5I, 5bcd, 5acd, 5abc, 5abd is again

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Figure 2.18. HPLC separation of coproporphyrinogen isomers on a HypersilODS column with 25% acetonitrile in 1 M ammonium acetate–acetic acid buffer (pH 5.16) containing 0.27 mM EDTA as eluent.

Figure 2.19. HPLC separation of pentacarboxylic porphyrinogen isomers on a Hypersil-ODS column with 40% methanol in 1 M ammonium acetate– acetic acid buffer (pH 5.16) containing 0.27 mM EDTA as eluent.

different from that of the corresponding pentacarboxylic porphyrins (see Fig. 2.7). Improved resolutions of porphyrinogens over porphyrins are much more significant in the separation of the type-III hexa- and,

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Figure 2.20. HPLC separation of hexacarboxylic porphyrinogen isomers on a Hypersil-ODS column with acetonitrile/methanol/1 M ammonium acetate–acetic acid buffer, pH 5.16 (8:12:80, v/v/v) containing 0.27 mM EDTA as eluent.

especially, the hepta-carboxylic porphyrinogens as shown in Figs. 2.20 and 2.21, respectively. Complete separations of the six type-III hexacarboxylic porphyrinogen isomers and the four type-III heptacarboxylic porphyrinogen isomers have been achieved. Similar separations of porphyrins have not been achieved. The improved separations is again due to the steric effect, with different degree of shielding of the methyl group or groups allowing each isomer to interact differently with the hydrophobic stationary phase surface thus achieving separation. Figure 2.22 shows the separation of uroporphyrinogen isomers. The optimum pH for the separation is 4.6. At pH above 5.0 separation of the isomers II, III, and IV was lost, although resolution of the natural I and III isomers was maintained. Suppression of ionisation of the carboxylic acid groups at lower pH increases the hydrophobicity of these relatively polar compounds, thus allowing better retention and more effective separation. Methanol, which causes excessive retention of uroporphyrin on an ODS column because of extensive hydrogen bonding with the carboxylic acid groups (see Fig. 2.3), can be used as

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Figure 2.21. HPLC separation of heptacarboxylic acid porphyrinogen isomers. Column: Asahipak ODP-50 (150 mm × 4.6 mm; 5 µm particle size). Eluent: acetonitrile/methanol/1 M ammonium acetate–acetic acid buffer, pH 5.16 (7:3:90, v/v/v) containing 0.27 mM EDTA.

the organic modifier for the separation of uroporphyrinogens. The flexible uroporphyrinogens are probably able to form intramolecular hydrogen bonds, thus minimising the possibility of intermolecular hydrogen bonding with the adsorbed methanol. This could also explain the reversal of elution order of that observed for the corresponding uroporphyrin isomers (see Figs. 2.14 and 2.15). Porphyrinogens are non-fluorescent, colourless compounds with weak UV absorption at the 220 nm region. Although UV detection can be used, they are best detected electrochemically by the oxidation mode because of the ease of oxidation of these compounds. The electrochemical reaction is shown in Fig. 2.23. Assuming complete oxidation to porphyrin, the porphyrinogen

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Figure 2.22. HPLC separation of uroporphyrinogen isomers on a HypersilODS column with 4% acetonitrile in 1 M ammonium acetate–acetic acid buffer (pH 4.6) containing 0.27 mM EDTA as eluent.

NH HN

- 6H NH HN

Porphyrinogen

Figure 2.23. detection.

NH

N

+6e N

HN

Porphyrin

Electrochemical oxidation of porphyrinogens in amperometric

loses six protons and generates six electrons. Porphyrins are also electro-active but there is no advantage in detecting porphyrins electrochemically since they can be more sensitively detected by a fluorescence detector.

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References 1. Woodward RB. The total synthesis of vitamin B12. Pure and Applied Chemistry 1973; 33: 145–177. 2. Gray CH, Lim CK and Nicholson DC. The differentiation of the porphyrias by means of high pressure liquid chromatography. Clinica Chimica Acta 1977; 77: 167–178. 3. Wright DJ, Rideout JM and Lim CK. High-performance liquid chromatography of coproporphyrin isomers. Biochemical Journal 1983; 209: 553–555. 4. Lim CK, Rideout JM and Wright DJ. Separation of porphyrin isomers by high-performance liquid chromatography. Biochemical Journal 1983; 211: 435–438. 5. Lim CK, Rideout JM and Wright DJ. High-performance liquid chromatography of naturally occurring 8-, 7-, 6-, 5- and 4-carboxylic porphyrin isomers. Journal of Chromatography 1983; 282: 629–641. 6. Lim CK, Rideout JM and Peters TJ. High-performance liquid chromatography of dicarboxylic porphyrins and metalloporphyrins: Retention behaviour and biomedical applications. Journal of Chromatography 1984; 317: 333–341. 7. Lim CK, Li F and Peters TJ. High-performance liquid chromatography of porphyrins (review). Journal of Chromatography 1988; 429: 123–153. 8. Meyer HD, Vogt W and Jacob K. Improved separation and detection of free porphyrins by high-performance liquid chromatography. Journal of Chromatography 1984; 290: 207–213. 9. Lim CK and Peters TJ. Urine and faecal porphyrin profiles by reversedphase high-performance liquid chromatography in the porphyrias. Clinica Chimica Acta 1984; 139: 55–63. 10. Lim CK and Peters TJ. Ammonium acetate: A general purpose buffer for the clinical applications of HPLC. Journal of Chromatography 1984; 316: 397–406. 11. Rideout JM, Wright DJ and Lim CK. High-performance liquid chromatography of uroporphyrin isomers. Journal of Liquid Chromatography 1983; 6: 383–394. 12. Razzaque MA, Lord GA and Lim CK. Amino acid and peptide conjugates of protoporphyrin: Preparation and analysis by high-performance

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13.

14.

15. 16.

High-Performance LC and MS of Porphyrins, Chlorophylls and Bilins

liquid chromatography, high-performance liquid chromatography/ electrospray ionization mass spectrometry and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Communications in Mass Spectrometry 2002; 16: 1675–1679. Lim CK, Razzaque MA, Luo J and Farmer PB. Isolation and characterization of protoporphyrin glycoconjugates from rat Harderian gland by HPLC, CE and HPLC/ESI-MS. Biochemical Journal 2000; 347: 757–761. Meijers JCM, Lim CK, Lawson AM and Peters TJ. Analysis of tumourlocalizing haematoporphyrin derivative by high-performance liquid chromatography and fast-atom bomdardment mass spectrometry. Journal of Chromatography 1986; 352: 231–239. Li F, Lim CK and Peters TJ. HPLC of porphyrinogens with electrochemical detection. Chromatographia 1987; 24: 421–422. Lim CK, Li F and Peters TJ. High-performance liquid chromatography of type-III heptacarboxylic porphyrinogen isomers. Biochemical Journal 1987; 247: 229–232.

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CHAPTER 3

Mass Spectrometry of Porphyrins

3.1. Introduction Mass spectrometry (MS) of porphyrins began with the introduction of the ‘direct’ insertion probes in the mid-1960s. Before then, MS of involatile compounds, including porphyrins, was very difficult or virtually impossible. The availability of fast atom bombardment (FAB)MS in the early 1980s, followed by atmospheric pressure chemical ionisation1 (APCI), electrospray ionisation2 (ESI), and matrix-assisted laser desorption/ionization3,4 mass spectrometry (MALDI)-MS in the late 1980s, dramatically increases the capabilities of MS in porphyrin analysis. APCI, ESI and the relatively new atmospheric pressure photoionisation5,6 (APPI) are atmospheric pressure ionisation (API) techniques well suited for coupling to liquid chromatographic techniques for online separation, detection and tandem MS/MS fragmentation analysis. Modern ESI mass spectrometers are extremely sensitive, requiring only minute quantities of materials for analysis, and can provide not only accurate mass measurements and elemental compositions of compounds, but also structural information. ESI is the MS technique of choice for the analysis porphyrins in biological and clinical samples. Time-of-flight (TOF), quadrupole and ion-trap mass analysers are used with the various ion sources, either singly or in combination, such as a Q-TOF instrument by coupling a quadrupole and a TOF analyser, to improve resolution and sensitivity. The Q-TOF tandem MS/MS instrument allows exact mass measurement on product and precursor ions in tandem MS/MS experiment, greatly improving analytical capability. 51

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Mass analysers measure mass (m) to charge (z) ratio (m/z). In most ionisation processes for molecules below 1000 Da, predominantly singly charged ions are produced, where z = 1 and m/z = mass.

3.2. Fast Atom Bombardment (FAB) Mass Spectrometry of Porphyrins FAB or liquid secondary ionisation or liquid surface ionisation mass spectrometry (LSIMS), although largely superseded by MALDI and ESI, is still useful for the rapid screening of porphyrin mixtures7 in chemical synthesis or in biological extracts where quantity and purity are not important considerations. In FAB or LSIMS the sample is dissolved in a liquid matrix with a low vapour pressure, e.g., glycerol, thioglyerol or 3-nitrobenzyl alcohol. An aliquot of 1–3 µ l is then placed on a small metal target at the end of a probe for insertion into the mass spectrometer. The liquid surface is then bombarded with a beam of high kinetic energy Ar or Xe atoms or Cs ions where molecules are desorbed, enter the gas phase and ionise, forming protonated or de-protonated ions, respectively for positive and negative mode MS. The FAB-MS of a standard mixture of porphyrin methyl esters is shown in Fig. 3.1. The efficiency of ionisation and consequently the sensitivity of detection decrease with increasing number of carboxylic acid substituents. LSIMS and FAB-MS may also be used for the analysis of porphyrins following high-performance thin layer chromatographic (HPTLC) separation on aluminium-backed plates. The bands are excised together with the aluminium backing and the strips are attached to the probe tip using an electro-conducting adhesive. Extraction solvent and liquid matrix are added to the surface of the TLC strips prior to MS.

3.3. Laser Desorption/Ionisation Time-of-Flight (LDI-TOF) Mass Spectrometry of Porphyrins In MALDI-MS the sample is dissolved in the matrix, e.g., 2,5-dihydroxybenzoic acid and sinapinic acid, and allowed to dry and crystallise on a stainless steel target disk. The target disk is then inserted

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Figure 3.1. FAB-MS of a standard mixture of porphyrin methyl esters (see Table 3.2 for m/z value of compounds).

into the mass spectrometer and the surface bombarded with a beam of photons from a pulsed laser beam of appropriate wavelength. Molecules are desorbed and ionised from the surface with little fragmentation or multiple charge formation. MALDI-MS is much more sensitive and has a larger analysable mass range than LSIMS. MALDIMS usually uses a time-of-flight (TOF) mass analyser as in MALDITOF MS. Porphyrins readily absorb UV and visible radiation. They could therefore be desorbed and ionised following excitation by photons of the correct wavelength without the need for added matrix. LDITOF MS rather than MALDI-TOF MS is therefore used in porphyrin analysis.8 LDI-TOF MS is particularly suitable for the fast screening and profiling of porphyrin esters. The porphyrin esters, being soluble in volatile organic solvents such as dichloromethane, can be easily applied onto the target plate and rapidly dried, thus

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Figure 3.2.

LDI-TOF MS of a standard mixture of porphyrin methyl esters.

allowing multi-loading of samples onto a 15-sample target plate. Analysis of 15 samples can be achieved within 10–15 minutes. It has been used for the rapid screening and identification of porphyrins in urine samples obtained from patients with suspected porphyries.8 Figure 3.2 shows the LDI-TOF MS profiles of a standard mixture of porphyrin methyl esters. Again, the sensitivity of detection decreases with increasing number of carboxylic acid groups. Figures 3.3 and 3.4 are those from urine of patients with congenital erythropoietic porphyria (CEP) and porphyria cutanea tarda (PCT), respectively. Less than one minute is required per sample compared to conventional HPLC separation of porphyrin methyl esters which takes up to 30 minutes to complete an analysis. The profiles obtained by LDI-TOF MS are similar to those by HPLC analysis.

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Figure 3.3. LDI-TOF MS of the porphyrin methyl esters isolated from the urine of a patient with CEP with characteristic elevated levels of uroporphyrin I (m/z 943) and coproporphyrin I (m/z 711).

Figure 3.4. LDI-TOF MS of the porphyrin methyl esters isolated from the urine of a patient with PCT showing increased excretion of uroporphyrin (m/z 943), heptacarboxylic porphyrin (m/z 885) and to a lesser extent, hexacarboxylic porphyrin (m/z 827), pentacarboxylic porphyrin (m/z 769) and coproporphyrin (m/z 711).

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Figure 3.5. MALDI-TOF mass spectrum of protoporphyrin-di(cys-gly-tyrgly-pro-lys-lys-lys-arg-lys-val-gly-gly) conjugate. The [M + H]+ ion is at m/z 3316.

MALDI-TOF MS is particularly useful for the analysis of peptide conjugates of protoporphyrin.9 The MALDI-TOF mass spectrum of protoporphyrin — di(cys-gly-tyr-pro-lys-lys-lys-arg-lys-val-gly-gly) conjugate with the [M + H]+ ion at m/z 3316 is shown in Fig. 3.5. The compound was synthesised by reacting protoporphyrinogen with the peptide followed by oxidation to porphyrin. The conjugation is by addition of the peptide to the two vinyl groups via the terminal cysteine group.

3.4. Electrospray Ionisation Mass Spectrometry (ESI-MS) and HPLC/ESI-MS of Porphyrins ESI involves the generation of a fine spray of ionised droplets by applying a high voltage of between 3–5 kilovolts (relative to a counter electrode) to the tip of the outlet of a capillary carrying a

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stream of liquid at atmospheric pressure. The liquid stream may be directly infused sample solution or HPLC eluent. This creates a high electric potential which causes nebulisation and the production of a fine mist of charged droplets. Nitrogen gas is used to enhance nebulisation and also to de-solvate the droplets. The de-solvated ions are propelled into the high vacuum of the mass analyser through a small opening guided by electrical potential difference. ESI is a soft ionisation technique which causes little or no fragmentation, although ‘insource’ fragmentation can be induced by raising the sample cone voltage. ESI is efficient for the ionisation of polar compounds and compatible with solvents used in liquid chromatography, especially reversed-phase HPLC. HPLC/ESI-MS is therefore the ideal technique for the separation, detection and characterisation of porphyrin free acids as well as their ester derivatives. For the less polar porphyrin derivatives, e.g., chlorophyll related compounds (see Chapter 6), APCI may also be used.1 APCI utilises a heated nebuliser to evaporate solvents and is able to accept a flow rate of 1 ml/min commonly used in HPLC. Flow splitting is necessary for ESI-MS except when capillary or microbore HPLC columns are used with flow rates in the 10 to 100 µ l ranges. APCI is less prone to ion suppression and has a wider dynamic concentration range than ESI. However, APCI can produce unexpected fragmentation and is unsuitable for thermally labile compounds. Ionisation in APPI is by proton absorption and electron ejection from a molecule to form a charged molecular ion in the case of nonpolar compounds, or by formation of protonated molecules in the presence of water or protic solvents as with ESI. The technique, in conjunction with Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS), has been used for the direct identification of vanadyl porphyrins and sulphur-containing vanadyl porphyrins in a petroleum asphaltene.10 APPI provides soft ionisation of the porphyrins, while the ultra-high mass resolving power of FTICR-MS was used for positive identification of elemental formulae of the entire family of vanadyl porphyrins and sulphur-containing vanadyl porphyrins in the complex petroleum asphaltene matrix.

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The zwitterionic naturally occurring porphyrins can be analysed by either negative or positive mode MS. The negative mode is generally less sensitive. The pH range of most HPLC mobile phases also favours positive ion MS.

3.4.1. HPLC/ESI-MS of porphyrins Mass spectrometry is a very sensitive and specific technique for the detection and identification of porphyrins and tandem MS/MS analysis allows structural elucidation. The fragmentation patterns and product ion spectra of porphyrin type-isomers obtained by tandem MS/MS, however, are very similar and this makes their identification difficult. This problem is solved by coupling high resolution HPLC to mass spectrometry11-15 (HPLC/MS) in which the porphyrins, including their isomers, are first separated by HPLC (see Chapter 2) and then analysed by MS. HPLC/MS systems incapable of isomer separation are less useful since the same profile can be obtained much more easily and rapidly by LSIMS or LDI-TOF MS analysis. Tandem MS/MS analyses are also possible with LSIMS and LDI-TOF MS. However, the product ion spectra of the type-isomers are very similar and hence could not be differentiated. It is therefore of prime importance that the HPLC system for use with mass spectrometry is capable of high resolution. HPLC separation is also important for minimising or eliminating matrix effects when biological samples are analysed. Endogenous and exogenous interfering components and contaminants in the matrix can suppress the ionisation of the analyte and cause differences in response between the analyte in sample and in standard solution, leading to difficulties in quantitative analysis and compound identification. Matrix effects and interferences and contaminants encountered in modern mass spectrometry have been reviewed.16 A typical HPLC/ESI-MS chromatogram of a mixture of naturally occurring porphyrins12 is shown in Fig. 3.6. The HPLC/MS operation conditions are summarised in Table 3.1 and the m/z ratios of the protonated porphyrin molecules, [M + H]+, are shown in Table 3.2.

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Figure 3.6. HPLC/ESI-MS chromatogram of a mixture of naturally occurring porphyrins. HPLC/MS conditions are listed in Table 3.1. The [M + H]+ ions are shown for uroporphyrin (m/z 831), heptacarboxylic porphyrin (m/z 787), hexacarboxylic porphyrin (m/z 743), pentacarboxylic porphyrin (m/z 699) and coproporphyrin (m/z 655). The type-I isomer eluted before the type-III isomer for all porphyrins. Pentacarboxylic porphyrin isomers were eluted in the order of 5I, 5IIIbcd, 5IIIabc, 5IIIacd and 5IIIabd.

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Table 3.1.

HPLC/ESI-MS and MS/MS Conditions for Porphyrins.

HPLC Column: Gemini C18 (250 mm × 4.6 mm i.d., 5 µ m particle size, from Phenomenex, Macclesfield, UK). Gradient solvent mixtures: solvent A, 9% (v/v) acetonitrile in 1 M ammonium acetate-acetic acid buffer, pH 5.16; solvent B, 9% (v/v) acetonitrile in methanol. Elution: linear gradient from 10% solvent B (90% solvent A) to 90% solvent B (10% solvent A) in 50 min., isocratic elution at 90% B from 50 to 60 min. Flow rate: 1 ml/min. The flow leaving the HPLC column was split in the ratio of 1:9 such that 100 µ l/min entered the mass spectrometer. ESI-MS and MS/MS Mass spectrometer: Q-TOF Micro orthogonal acceleration electrospray ionisation time-of-flight (Micromass, Waters, Manchester, UK). Capillary voltage: 3.50 kV; cone voltage: 90 V; source temperature: 110°C; nebulising gas (N2) flow rate: 50 L/h; desolvation gas (N2) flow rate: 350 L/h with a temperature of 350°C. Data acquisition: continuum data in positive ion mode; mass range: 100-900 Da; scan rate: 3 spectra/s. MS/MS: collision energy: 35 eV; collision gas: argon.

Table 3.2. The m/z Ratios of Protonated Porphyrin Molecules [M + H]+. Porphyrin Uroporphyrin Heptacarboxylic porphyrin Hexacarboxylic porphyrin Pentacarboxylic porphyrin Coproporphyrin 2-Vinyl tricarboxylic porphyrin Mesoporphyrin Protoporphyrin

Free acid m/z

Methyl ester

831 787 743 699 655 609 567 563

943 885 827 769 711 651 595 591

The efficiency of the reversed-phase HPLC system in porphyrin separation is clearly demonstrated by the complete resolution of the five pentacarboxylic porphyrin isomers under gradient elution condition. The separation of hydroxyuroporphyrin derivatives14 (Fig. 3.7) is another example of a high-resolution HPLC/ESI-MS system for the

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Figure 3.7. HPLC/ESI MS chromatogram of uroporphyrin I (m/z 831) and hydroxyuroporphyrin I derivatives formed by photochemical oxidation of uroporphyrinogen I. Peaks: 1, meso-hydroxyuroporphyrin I; 2, α-hydroxypropionic acid uroporphyrin I; 3, β-hydroxypropionic acid uroporphyrin I; 4, hydroxyacetic acid uroporphyrin I; 5, trans-hydroxyspirolactoneurochlorin I; 6, cis-hydroxyspirolactoneurochlorin I; 7, trans-dihydroxyurochlorin I; 8, cis-dihydroxyurochlorin I. Column, Hypersil-BDS; elution, linear gradient from 100% A (0% B) to 90% B (10% A) in 60 minutes. Solvent A was 9% acetonitrile in 1 M ammonium acetate, pH 5.16 and solvent B was 9% acetonitrile in methanol.

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separation and identification of isomeric porphyrin derivatives. Six monohydroxy- and two dihydroxy-uroporphyrin derivatives (Fig. 3.8) have been identified.14 The separation was carried out on a Hypersil BDS-C18 column by gradient elution from 100% A (0% B) to 90% B (10% A) in 60 minutes. Solvent A was 9% acetonitrile in 1 M ammonium acetate, pH 5.16 and solvent B was 9% acetonitrile in methanol. Ammonium acetate buffer at 1 M concentration does not cause significant problems for the MS ion source. Under gradient elution condition it is gradually diluted and by the end of the gradient little remains in the system. It is possible that a column with different selectivity may not need 1 M ammonium acetate buffer to achieve similar separation. It is impossible to investigate all reversed-phase HPLC columns for HPLC/MS of porphyrins. However, reversed-phase columns from major manufacturers/suppliers such as Phenomenex, Thermo Hypersil-Keystone and Waters all show better column efficiency when 1 M ammonium acetate buffer was used. A conventional HPLC column of 250 mm × 4.6 mm packed with 5 µ m particle sized material is usually used in HPLC/ESI-MS of porphyrins. This necessitates splitting the 1 ml/min flow leaving the column in the ratio of 1:9 such that the flow enters the mass spectrometer at 100 µ l/min. Using a column of 100 mm × 1 mm packed with 3 µ m material and eluting at 100 µ l/min, no flow splitting is necessary. ESI-MS sources are concentration sensitive and response to changes in analyte concentration. A smaller diameter column shows higher sensitivity when coupled with ESI-MS, since the analyte eluting in a smaller volume of eluent is more concentrated. A further advantage of using smaller diameter columns is less contaminant enters the mass spectrometer and the ionisation source remains clean longer. When a new column is used for the first time, it is advisable to thoroughly wash and condition the column before coupling to the mass spectrometer. New columns often contain varying amount of surfactant impurities which can contaminate the ionisation source and interfere with MS analysis.

Ac

Pr

COOH

N

NH

HN

Ac

Pr

Ac

Pr

O

Ac

Ac

N

HN

N Ac

Ac

Pr

5

Pr

O

HO

HN

N Ac

Ac

Pr

6

Ac

OH Pr

N

NH

Pr

Pr

Ac Ac

N

NH

Pr

OH O

4

Ac

N

Pr

Ac

3

N Ac

Ac

Pr

7

Pr

N

NH

HN

Pr

OH OH

HN

Pr

Ac Ac

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Ac

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HN

N Pr

Pr

Ac

N

Mass Spectrometry of Porphyrins

HO

HN Ac

2

Ac

Pr

Ac NH

Pr

1

NH

N

Ac

Pr

Pr

Pr

HN

N Ac

COOH

N

NH

COOH

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HO Pr

OH

Ac

HO

OH

Ac

Pr

Ac

N

NH

Ac

Pr

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Pr

Pr

8

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Figure 3.8. Chemical structures of hydroxyuroporphyrin derivatives. 1, meso-hydroxyuroporphyrin I; 2, α-hydroxypropionic acid uroporphyrin I; 3, β-hydroxypropionic acid uroporphyrin I; 4, hydroxyacetic acid uroporphyrin I; 5, transhydroxyspirolactoneurochlorin I; 6, cis-hydroxyspirolactoneurochlorin I; 7, trans-dihydroxyurochlorin I; 8, cis-dihydroxyurochlorin.

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3.4.2. ESI-MS/MS fragmentation of porphyrins Tandem MS/MS analysis provides structural information of compounds. This is carried out by coupling two mass analysers separated by a collision cell. The first analyser selects the ion of interest, the precursor ion, which is then passed into the collision cell pressurised with an inert collision gas, usually argon. Dissociation of the precursor ion into product ions is induced by collision with the argon atoms in the cell. This process is termed collision induced dissociation (CID). The product ions are then analysed in the second analyser to give a product ion spectrum of the original precursor ion. Porphyrins show little or no fragmentation without CID except for derivatives with labile side-chain substituents which may be fragmented in the ion source at a higher cone voltage. The porphyrin macrocycle itself is stable to fragmentation. The fragmentation is therefore centred on the side-chain substituent groups on the porphyrin macrocycle. For porphyrins derived from the haem biosynthetic pathway, this is dominated by the acetic and/or propionic acid substituents. The three most prominent fragmentation pathways, shown in Fig. 3.9, are: 1, benzylic cleavage with the loss of HCOOH (46 mass units) from a protonated . molecule; 2, benzylic cleavage with the loss of a CH2COOH (59 mass unit) radical from a propionic acid substituent; 3, loss of H2O (18 mass unit) from a protonated molecule. The tandem MS/MS fragmentation of the porphyrins can be obtained during the HPLC run. The relative intensity of the product

H3C - CH3 + H (14)

NH N

N HN

H H O C C C OH H H - CH2COOH (59) H O C C OH + H+ H - H2O (18) - HCOOH (46)

Figure 3.9. Major tandem MS/MS fragmentation pathways of porphyrin side-chain substituents.

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ions may differ depending on the type and pH of the mobile phase used, although the pattern should remain the same. The collision energy used also influences the ion intensity and fragmentation. The product ion spectra shown in the following sections are obtained from on-line HPLC/ESI-MS/MS experiments using the conditions described in Table 3.1.

3.4.2.1. ESI-MS/MS product ion spectrum and fragmentation pathways of uroporphyrin The product ion spectrum of uroporphyrin I is shown in Fig. 3.10. Uroporphyrin has four acetic acid and four propionic acid sidechain substituents. Fragmentation of the [M + H]+ precursor ion at m/z 831 follows the expected pathways shown in Fig. 3.7. The product ion at m/z 785 (831 − 46) corresponds to benzylic cleavage and the loss of a HCOOH group form a side-chain acetic acid groups (Fig. 3.11). Benzylic cleavage of a propionic acid substituent by the loss of a . CH2COOH radical gave the peak at m/z 772 (831 − 59). The ion at m/z 727 (831 − 46 − 59 + 1) resulted from the succes. sive loss of a HCOOH group and a CH2COOH radical followed by back protonation of the product ion. The elimination of H2O from the precursor ion gave the peak at m/z 813 (831 − 18), which was further fragmented by the loss of a HCOOH group to give the ion at m/z 767. It must be emphasised that obtaining a product ion spectrum during a HPLC run may be subjected to interference occasionally by coeluting compounds with the same m/z ratio. The most widely encountered interfering compound is poly (ethylene glycol) which is often present in sample extracts.17

3.4.3. ESI-MS/MS fragmentation of hydroxyuroporphyrins Uroporphyrin can be mono-oxygenated to give hydroxyuroporphyrin derivatives. This can take place at the meso-position or on one of the sidechain carboxylic acid substituents.14,18 In general, the fragmentation

813 772

714

669 681

655 663

0 600

610

620

630

640

650

660

670

680

690

699

709

700

710

739

758

743

723

720

730

740

750

760

770

780

790

800

m/z

Figure 3.10.

ESI-MS/MS product ion spectrum of uroporphyrin I.

810

820

830

840

m/z 850

Relative Intensity %

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727

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831

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66

100

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Ac

Pr Ac

Pr NH N

N HN

Pr Ac

CH2

- CH2COOH

N

N

Ac

HN

N

CH2COOH +H]+

Pr Ac

m/ z 831

N

+H

HN CH2+

Ac

- H2O

m/ z 785

N

N

CH2CO+ Pr

Ac

Pr

Pr

HN

m/ z 813

CH3

H2C

- HCOOH

Pr

Figure 3.11.

CH2+ m/ z 727

Pr NH

HN

Ac

Pr

Ac

Ac

N

N

Pr

Ac

Pr

Pr NH

- CH2COOH

Pr

Pr

Ac

Pr NH

- HCOOH

Ac

Pr

Ac

Pr Pr

NH

Ac +H]+

m/ z 772

Ac

Pr Ac

67

N

N

N

HN CH2CO+

Pr Ac

m/ z 767

Pr

Proposed ESI-MS/MS fragmentation pathways of uroporphyrin I.

of hydroxyuroporphyrins12,14 follows the same pathways of uroporphyrin. However, they also show individual characteristics which allow for differential identification of the positional isomers.

3.4.3.1. ESI-MS/MS product ion spectrum and fragmentation pathways of meso-hydroxyuroporphyrin The product ion spectrum of the [M + H]+ ion of meso-hydroxyuroporphyrin I at m/z 847 is shown in Fig. 3.12. The fragmentation pathway is typical for an uroporphyrin derivative, being dominated by benzylic cleavages of the side-chain acetic acid and propionic acid substituents. However, the presence of an OH group at the mesoposition alters the electronic environment around the porphyrin macrocycle and consequently influences the fragmentation pattern of the molecule.

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ESI-MS/MS product ion spectrum of meso-hydroxyuroporphyrin I. Figure 3.12.

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The ion at m/z 801 corresponds to the benzylic cleavage and the loss of a HCOOH group from the acetic acid group of a pro. tonated molecule (847 − 46). Further loss of a CH2COOH radical from this ion with simultaneous back protonation gave the peak at m/z 743 (801 − 59 + 1). An ion of significant intensity at m/z 802, . corresponding to the loss of a COOH radical, was also observed (847 − 45). This appears to be a characteristic feature of mesohydroxyuroporphyrin. With an OH group at the meso-position, which can also be protonated, elimination of a water molecule from the precursor ion is enhanced, giving a prominent peak at m/z 829 (847 − 18). The loss of H2O from a protonated porphyrin molecule is normally a relatively minor pathway. The radical ion at m/z 802 can also eliminate H2O to give the peak at m/z 784 (802 − 18) which was then protonated to give the ion at m/z 785. The ion at m/z 773 appears to be derived from the loss of a CO group from the peak at m/z 801 (801 − 28). This is possible because of a tautomeric oxophlorin structure of the meso-hydroxy form. Elimination of CO would obviously lead to ring opening.

3.4.3.2. ESI-MS/MS product ion spectrum of hydroxyacetic acid uroporphyrin Figure 3.13 shows the product ion spectrum of the [M + H]+ ion of hydroxyacetic acid uroporphyrin I at m/z 847. The most characteristic fragmentation feature of this compound is the ease with which the . entire side-chain hydroxyacetic acid ( CHOHCOOH) group was cleaved to give an ion at m/z 772 (847 − 75). The loss of a HCOOH group gave the ion at m/z 801 (847 − 46) . which was further fragmented by losing a side-chain CHOHCOOH radical, followed by addition of a proton to give the base peak at m/z 727 (801 − 75 + 1). Elimination of H2O from the ions at m/z 847 and 801 gave the ions at m/z 829 and 783, respectively. These are minor pathways of fragmentation.

x1.5

713 743

667 681 684

655

0 600

610

620

630

640

650

660

670

680

708 699

690

700

710

783

720

730

740

829

755

737

750

760

770

780

790

800

810

820

830

840

m/z

Figure 3.13.

ESI-MS/MS product ion spectrum of hydroxyacetic acid uroporphyrin I.

850

m/z 860

Relative Intensity %

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847

FA

727

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70

100

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3.4.3.3. ESI-MS/MS product ion spectrum of α-hydroxypropionic acid uroporphyrin The product ion spectrum of α-hydroxypropionic acid uroporphyrin I is shown in Fig. 3.14. The loss of a HCOOH group from a protonated acetic acid substituent gave the peak at m/z 801. . This was further fragmented by losing a COOH radical from the α-hydroxypropionic acid group to give the most intense ion at m/z 756. This ion was able to pick up a proton to give the peak at m/z 757. . The loss of a side-chain CH2CH2COOH radical by the precursor ion gave the peak at m/z 774 (847 − 73), while the loss of the same radical by the ion at m/z 801 gave the peak at m/z 728 (801 − 73). Similarly, the ion at m/z 757 could fragment by losing a propionic acid radical to give the peak at m/z 684 (757 − 73).

3.4.3.4. ESI-MS/MS product ion spectrum and fragmentation pathways of β-hydroxypropionic acid uroporphyrin The product ion spectrum of β-hydroxypropionic acid uroporphyrin I is the most complex among the hydroxyuroporphyrins and is shown in Fig. 3.15. . The combined loss of a HCOOH group and a CH2COOH radical from the precursor ion at m/z 847 with simultaneous back protonation (+H) of the resulting product ion gave an intense base peak at m/z 743 (847 − 46 − 59 + 1). The ease with which this pathway took place is evidenced by the lack of an ion at m/z 801 (847 − 46) common to all other hydroxyuroporphyrin derivatives. This ion must have been further fragmented rapidly by the loss of . a CH2COOH radical, probably from the β-hydroxypropionic acid group. The loss of an acetic acid group from the β-hydroxypropionic acid group by the precursor ion gave the peak at m/z 787 (847 − 60) while the loss of the entire side-chain β-hydroxypropionic acid group

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ESI-MS/MS product ion spectrum of α-hydroxypropionic acid uroporphyrin I. Figure 3.14.

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ESI-MS/MS product ion spectrum of β-hydroxypropionic acid uroporphyrin I.

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Figure 3.15.

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with simultaneous back protonation gave the peak at m/z 759 (847 − 89 + 1) as shown: P-CHOH-CH2COOH − CH3COOH → P-CHO (m/z 787) . P-CHOH-CH2COOH − CHOH-CH2COOH + H → PH (m/z 759) P = protonated porphyrin molecule The precursor ion can eliminate H2O from the protonated molecule as well as from the β-hydroxypropionic acid substituent to give the ion at m/z 829. Elimination of a H2O molecule by the precursor ion is therefore another major fragmentation pathway, which gave a relatively intense peak at m/z 829 (847 − 18). Loss of H2O from the β-hydroxypropionic acid group can be shown as: P-CHOH-CH2COOH − H2O → P-CH=CH-COOH (m/z 829) . Further fragmentation of this ion by losing a COOH (45 mass units) radical with simultaneous back protonation gave the peaks at m/z 785 (829 − 45 + 1) represented as: . P-CH=CH-COOH − COOH + H → P-CH=CH2 (m/z 785) The m/z 829 peak may also eliminate H2O or HCOOH to give ions at m/z 811 (829 − 18) and m/z 783 (829 − 46), respectively. The ion at m/z 765 was derived from the peak at m/z 811 by losing a HCOOH group (811 − 46 = 765). The peak at m/z 829 may be fragmented by losing a CH3COOH group from the β-hydroxypropionic acid substituent to give the ion at m/z 769 (829 − 60).

3.4.3.5. ESI-MS/MS product ion spectra and fragmentation pathways of cis- and transhydroxyspirolactoneurochlorins Hydroxyspirolactoneurochlorins are formed following exposure of uroporphyrinogen to light or during the auto-oxidation of

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uroporphyrinogen.14,18 They exist as the cis and trans isomers and are most probably derived from an epoxide structure with the oxygen inserted across a β-pyrrole ring which is hydrolysed to a compound with a diol structure followed by facile dehydration and internal cyclisation.18 The product ion spectrum of trans-spirolactoneurochlorin I is shown in Fig. 3.16. The most characteristic feature is the ease with which the acetic acid and spirolactone (-CH2CH2COO-) substituents attached to the reduced pyrrole ring can be cleaved, giving ions at m/z 788 (847 − 59) and 775 (847 − 72), respectively. The ion at m/z 775 was further fragmented by losing a HCOOH group to give the base peak at m/z 729. The loss of a HCOOH group by the precursor ion followed by elimination of H2O gave the peaks at m/z 801 (847 − 46) and m/z 783 (801 − 18), respectively, while the loss of H2O by the precursor ion gave the peak at m/z 829 (847 − 18), which was further fragmented by . losing a CH2COOH radical to give the ion at m/z 770 (829 − 59). The fragmentation pattern of cis-spirolactoneurochlorin I (Fig. 3.17) is similar to that of the trans isomer, although sufficiently different to allow differentiation between them. The loss of the spirolactone group was more prominent by the cis-isomer giving a more intense peak at m/z 775. This ion was further fragmented by losing a HCOOH group to give the ion at m/z 729 (775 − 46) which was protonated to give the base peak at m/z 730. . A further difference is that the loss of a COOH radical was preferred to the loss of a HCOOH group by the cis-isomer, resulting in the reversal in the m/z 801 and 802 peak intensity.

3.4.3.6. ESI-MS/MS product ion spectra and fragmentation pathways of cis- and trans-dihydroxyurochlorins Dihydroxyurochlorins are derived from epoxyurochlorins by hydrolysis.18 The tandem MS/MS product ion spectrum of trans-dihydroxyurochlorin I is shown in Fig. 3.18. Like the hydroxyspirolactoneurochlorins, groups attached to the reduced pyrrole ring are easily fragmented. The precursor ion at m/z 865 thus eliminated an acetic acid group to give the base peak

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ESI-MS/MS product ion spectrum of trans-hydroxyspirolactoneurochlorin I. Figure 3.16.

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ESI-MS/MS product ion spectrum of cis-hydroxyspirolactoneurochlorin I.

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Figure 3.17.

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ESI-MS/MS product ion spectrum of trans-dihydroxyurochlorin I. Figure 3.18.

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at m/z 805 (865 − 60), which then cyclised by eliminating H2O to give the peak at m/z 787 (805 − 18). The precursor ion also cyclised by losing H2O to give a spirolactone structure at m/z 847 (865 − 18) which was further fragmented by losing another molecule of H2O to give the ion at m/z 829 (847 − 18), a HCOOH group to give the peak at m/z 801 (847 − 46), a . CH2COOH radical to give the ion at m/z 788 (847 − 59) or a spirolactone (-CH2CH2COO-) group to give the peak at m/z 775 (847 − 72). The proposed fragmentation pathway is shown in Fig. 3.19. cis-Dihydroxyurochlorin I gave very similar product ion spectrum except the base peak was at m/z 847, probably because this isomer is more easily cyclised to the spirolactone structure by losing H2O.

Pr

Ac

HO

Pr

O

OH

Ac NH

Pr

N

m/ z 865

Pr

N

+ H]+ N

Ac Ac

NH

HN

Pr

- H2O

m/ z 805

Pr

Ac

HO

O

Pr

O

N

N + H]+ Ac

HN

Pr Ac

Ac Ac

Pr

O

O

N + H]+

N

HN

Pr

Pr

Ac Ac

m/ z 847

HN

m/ z 787

HO

NH

- CH2COOH

N

Pr

Pr

Ac NH

O

+ H]+

Ac Ac

NH N

- H2O Ac

O

HN

Pr

Pr

O

Ac

- CH3COOH

+ H]+ N

Pr

OH

Ac

- H2O

Pr

m/ z 788

- (CH2CH2COO) Pr

HO

Ac NH N

+

N

H

Ac Ac

m/ z 775

Ac NH N

HN

Pr Pr

Ac + O

Pr

Ac

O

N HN

Pr

Ac Ac

Pr

m/ z 829

Figure 3.19. Proposed ESI-MS/MS fragmentation pathways of transdihydroxyurochlorin I.

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3.4.3.7. Characteristic ESI-MS/MS product ions of hydroxyuroporphyrins The product ion spectra of the above hydroxyuroporphyrin derivatives show clearly identifiable patterns and characteristic product ions for each derivative. This allows the isomers to be identified by tandem MS/MS analysis. The most characteristic product ions are summarised in Table 3.3.

3.4.4. ESI-MS/MS product ion spectra of ketoacid uroporphyrins Ketoacetic acid uroporphyrin and β-ketopropionic acid uroporphyrin18 are derived from hydroxyacetic acid uroporphyrin and βhydroxypropionic acid uroporphyrin, respectively, by further oxidation. Ketoacid porphyrins, particularly β-ketopropionic acid porphyrins, are able to form stable intramolecular H-bonds (Fig. 3.20) and this gives them the characteristic fragmentation patterns. The product ion spectrum of β-ketopropionic acid uroporphyrin I is shown in Fig. 3.21. The small peak at m/z 813 (845 − 32) is highly characteristic. It represents the loss of O2 which is only possible from an intramolecularly H-bonded β-ketopropionic acid ring structure. Further loss of CO from the ion at m/z 813 gave the peak at m/z 785 (813 − 28). The proposed pathway is shown in Fig. 3.22.

Table 3.3.

Characteristic ESI-MS/MS Product Ions of Hydroxyuroporphyrins. Porphyrin

Meso-Hydroxyuroporphyrin Hydroxyacetic acid uroporphyrin α-Hydroxypropionic acid uroporphyrin β-Hydroxypropionic acid uroporphyrin trans-Hydroxyspirolactoneurochlorin cis-Hydroxyspirolactoneurochlorin trans-Dihydroxyurochlorin cis-Dihydroxyurochlorin

Product ion (m/z) 829, 802, 801, 783, 801, 774, 829, 811, 829, 788, 829, 802, 847, 829, 847 (base

801 (base peak), 773, 743 772, 743, 727 (base peak) 756 (base peak), 728, 684 787, 759, 743 (base peak) 783, 775, 729 (base peak) 783, 775, 730 (base peak) 805 (base peak), 788, 775 peak), 829, 805, 788, 775

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O

H O

COOH

Pr

81

O

O

Pr Pr

Ac

Pr

Ac

N

NH

N

NH HN

N

HN

N

Ac

Pr

Ac

Pr Pr

Ac

Ac

1 Ac

Pr

COOH

N

NH

Ac

Pr Pr

Ac

3

H O

Ac

HN

N

Ac

Pr

O

Ac NH

Pr

2

N

O O

N HN Ac

Pr Pr

Ac

4

Figure 3.20. Chemical structures of ketoacetic acid uroporphyrin I (1 and 2) and β-ketopropionic acid uroporphyrin I (3 and 4).

The second proposed pathway of fragmentation involves the loss of HCOOH from a protonated molecule followed by benzylic cleavage of the side-chain β-ketopropionic acid substituent, giving ions at m/z 799 and 741, respectively (see Fig. 3.22). The peak at m/z 827 (845 − 18) shows the loss of H2O from a protonated molecule, which is further fragmented by losing a HCOOH group to give the ion at m/z 781 (827 − 46). The above fragmentation pathways are also observed for ketoacetic acid uroporphyrin I. The product ion spectrum is shown in Fig. 3.23. However, there are also some characteristic features not present in β-ketopropionic acid uroporphyrin which allow differentiation of the two compounds. These are firstly, the net loss of a CO2 group from the ketoacetic acid substituent resulting from the loss of a COOH group with simultaneous back protonation of the product ion to give the peak at m/z 801

827

683 755 681

695

713

669

753

699

671

654 658

0 650

783

677

663 655

709 675

690

715 717

758

723 746 750

735

801

769 773 764

813 825

797

832

m/z 660

670

680

690

700

710

720

730

740

750

760

770

780

790

800

810

820

830

840

850

m/z

Figure 3.21.

ESI-MS/MS product ion spectrum of β-ketopropionic acid uroporphyrin I.

Relative Intensity %

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Ac

Pr

H O

Ac

Pr

Ac

N

O

N

Ac

NH

- O2

CH2COOH + H] +

HN

CH2COOH + H] +

Ac

O

OH

N HN CH2+

Ac

Pr

m/z 799

Pr

m/ z 785

Ac

Pr

O

.

NH

+H

N

O

N

Ac NH

HN CH2+

Pr Pr

Ac

m/ z 741

Ac

Pr

Ac

- CH2COOH N

CH2COOH + H] +

- HCOOH

Ac

Pr

HN

Ac

m/z 813

- HCOOH

NH

N

N

Pr

Pr

Ac

m/z 845

Pr

NH

-CO

Pr

Pr

Ac

O

N

N

HN

Pr

Ac

Pr

O Ac

NH

83

N

O

N HN CH2+

Pr Pr

Ac

m/z 767

Figure 3.22. Proposed ESI-MS/MS fragmentation pathways of β-ketopropionic acid uroporphyrin I.

(845 − 44); secondly, the benzylic cleavage of a propionic acid substituent to give the ion at m/z 786 (845 − 59) and thirdly, the loss of an entire COCOOH substituent giving the ion at m/z 772 (845 − 73).

3.4.5. ESI-MS/MS of heptacarboxylic porphyrin The fragmentation pathways of heptacarboxylic porphyrin (Fig. 3.24) are essentially similar to those observed for uroporphyrin. The loss of a HCOOH group gave the peak at m/z 741 (787 − 46). The presence of a CH3 substituent is clearly shown by . the loss of a CH3 radical from the ion at m/z 741 to give the peak at m/z 726 (741 − 15), which on protonation gave the ion at m/z 727. The successive loss of two HCOOH groups gave the ion at m/z . 695, while the loss of a HCOOH group and a CH2COOH radical . gave the peak at m/z 682. Elimination of a CH3 radical from the ion at m/z gave the peak at m/z 667, which on protonation gave the peak at m/z 668.

801

827 667

772

681 753

695 625 609

0 600

713

651 636

699 640 646

709

813

721

654 664

677

763

735

691

m/z 610

620

630

640

650

660

670

680

690

700

710

720

730

740

750

760

770

780

790

800

810

820

830

m/z

Figure 3.23.

ESI-MS/MS product ion spectrum of ketoacetic acid uroporphyrin I.

840

850

Relative Intensity %

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FA

845

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84

100

x2

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637

%

741 651 564 577

714

608 617 605 611

721

664

567 523

550 534

698

581 503

517

557

571

587

600

644

660

709

769

674 758

688

0 500

510

520

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Relative Intensity %

682

530

540

550

560

570

580

590

600

610

620

630

640

650

660

670

680

690

700

710

720

730

740

750

760

780

770

780

790

m/z 800

m/z

FA

ESI-MS/MS product ion spectrum of heptacarboxylic porphyrin III.

85

Figure 3.24.

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3.4.6. ESI-MS/MS of hydroxyheptacarboxylic porphyrins Heptacarboxylic porphyrin gave hydroxylated derivatives similar to those of the corresponding uroporphyrin derivatives. In addition, the CH3 substituent can also be hydroxylated to give a hydroxymethyl derivative (Fig. 3.25). Hydroxymethylheptacarboxylic porphyrin is easily distinguished from the other hydroxylated heptacarboxylic porphyrin derivatives by its characteristic fragmentation pattern. The product ion spectrum of hydroxymethylheptacarboxylic porphyrin I is shown in Fig. 3.26. The loss of a CH2O (30 mass units) group from the hydroxymethyl substituent of the precursor ion gave the most characteristic product ion for a hydroxymethylporphyrin structure at m/z 773 (803 − 30). The product ion spectrum of β-hydroxypropionic acid heptacarboxylic porphyrin I is shown in Fig. 3.27 for comparison. No m/z 773 ion was observed for this compound on fragmentation.

3.4.7. ESI-MS/MS of ketoacid heptacarboxylic porphyrins and formyl heptacarboxylic porphyrin The presence of a methyl group can also lead to the formation of a formyl derivative of heptacarboxylic porphyrin in addition to ketoacetic acid- and β-ketopropionic acid-heptacarboxylic porphyrins (Fig. 3.28).

Pr

CH2OH

Ac

Pr NH N

Pr

NH N

HN Ac Pr

1

Pr Pr

N

Pr Ac

CH3

Ac

Ac

Pr

N

N Ac

Ac

CHOHCH2COOH

2

N

NH

HN

Pr

CH3

HN

Pr

CHOHCOOH Ac

Pr

3

Figure 3.25. Chemical structure of 1, hydroxymethylheptacarboxylic porphyrin; 2, β-hydroxypropionic acid heptacarboxylic porphyrin; 3, hydroxyacetic acid heptacarboxylic porphyrin.

x5

x2

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803

726

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739 699

Relative Intensity %

681 639 609

669 622

641

655 715 651

597

741

627

581

709

605

685

649

721

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Mass Spectrometry of Porphyrins

773

757

693 577

647

618

587

767

730

759

631 593

575

583

603 737

677

0 570

580

590

600

610

620

630

640

650

660

670

680

690

700

710

720

730

740

750

760

770

780

790

800

m/z 810

m/z

FA

ESI-MS/MS product ion spectrum of hydroxymethylheptacarboxylic porphyrin I.

87

Figure 3.26.

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ESI-MS/MS product ion spectrum of β-hydroxypropionic acid heptacarboxylic porphyrin I. Figure 3.27.

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O

Pr

Pr

Ac

Pr

N

Ac

HN

N Ac

Ac

Pr

1

N

NH

Pr

Pr Pr

N

NH

CH3

89

Pr NH

HN

Pr

Ac Ac

COCH2COOH

2

CH3

Ac

N

N HN

Pr

COCOOH Ac

3

Pr

Figure 3.28. Chemical structure of 1, formylheptacarboxylic porphyrin; 2, β-ketopropionic acid heptacarboxylic porphyrin; 3, ketoacetic acid heptacarboxylic porphyrin.

For formylheptacarboxylic porphyrin I, the most characteristic product ion is at m/z 773 (Fig. 3.29) derived from the precursor ion (m/z 801) by the loss of a CO group. This ion is absent in the product ion spectrum of β-ketopropionic acid heptacarboxylic porphyrin I (Fig. 3.30), which characteristically eliminates O2 (as seen in all ketoacid uroporphyrin derivatives, see Sec. 3.4.4) to give a peak at m/z 769 (801 − 32). The product ion spectrum of β-ketopropionic acid heptacarboxylic porphyrin I (Fig. 3.30) indicated the presence of ketoacetic acid heptacarboxylic porphyrin I as an impurity due to incomplete HPLC separation of the compounds.

3.4.8. ESI-MS/MS product ion spectrum and fragmentation pattern of coproporphyrin The product ion spectrum of coproporphyrin III is shown in Fig. 3.31. With no acetic acid substituents, fragmentation of the precursor [M + H]+ ion at m/z 655 was dominated by the successive benzylic cleavages of the four propionic acid groups to give product ions at m/z 596 (655 − 59), 537 [655 − (2 × 59)], 478 [655 − (3 × 59)] and 419 [655 − (4 × 59)], respectively (Fig. 3.31). There was no product ion corresponding to the loss of a HCOOH (46 Da) group. This clearly shows that the loss of a HCOOH group can only occur when there is an acetic acid substituent and serves to differentiate an acetic acid from a propionic acid substituent.

Relative Intensity %

669

609 755

709 623 727 605 714

579 565

551

737

641

593

741 618

597

633

691 685

573

663 677

783

723

0 550

560

570

580

590

600

610

620

630

640

650

660

670

680

690

700

710

720

730

740

750

760

770

780

790

m/z

Figure 3.29.

ESI-MS/MS product ion spectrum of formylheptacarboxylic porphyrin I.

800

m/z 810

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ESI-MS/MS of β-ketopropionic acid heptacarboxylic porphyrin. Figure 3.30.

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ESI-MS/MS product ion spectrum of coproporphyrin III. Figure 3.31.

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Peaks at m/z 582 (655 − 73) and 509 [655 − (2 × 73)] could be . derived from the loss of one and two CH2CH2COOH radicals, respectively. However, they are more likely to results from the loss of . a CH3 radical from the ions at m/z 596 and 523, followed by back protonation of the resulting product ions, i.e., 596 − 15 + 1 = 582 and 523 − 15 + 1 = 509, respectively. The ion at m/z 523 was similarly . derived from the peak at m/z 537 by the loss of a CH3 radical followed by protonation (537 − 15 + 1 = 523). This was also observed for the ions at m/z 478 and 419, which fragmented to give peaks at m/z 464 (478 − 15 + 1) and 405 (419 − 15 + 1), respectively. The loss of H2O from a protonated molecule to give the ion at m/z 637 (655 − 18) was observed as a minor pathway.

3.4.9. ESI-MS/MS of hydroxymethylcoproporphyrin The product ion spectrum of hydroxymethylcoproporphyrin III is shown in Fig. 3.32. The loss of CH2O by the precursor ion was clearly seen, giving a prominent product ion at m/z 641 (671 − 30). This most characteristic pathway is similar to that observed for hydroxymethylheptacarboxylic porphyrin.

3.4.10. ESI-MS/MS of β-ketopropionic acid coproporphyrin and formylcoproporphyrin The fragmentation pathway of β-ketopropionic acid coproporphyrin III is similar to that of the corresponding of β-ketopropionic acid uroporphyrin and β-ketopropionic acid heptacarboxylic porphyrin and is characterised by the loss of O2, giving a peak at m/z 637 (Fig. 3.33). For formylcoproporphyrin, there was no loss of O2 but a CO group was eliminated from the formyl substituent.

3.4.11. ESI-MS/MS product ion spectrum and fragmentation pattern of protoporphyrin Figure 3.34 shows the product ion spectrum of protoporphyrin IX. The fragmentation pathway is similar to those observed for coproporphyrin

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ESI-MS/MS product ion spectrum of hydroxymethylcoproporphyrin III. Figure 3.32.

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ESI-MS/MS product ion spectrum of β-ketopropionic acid coproporphyrin III.

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Figure 3.33.

563

504

390

400

410

420

430

440

450

460

470

480

490

500

510

520

530

540

550

m/z Figure 3.34.

ESI-MS/MS product ion spectrum of protoporphyrin IX.

560

570

580

m/z 590

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and is dominated by the successive benzylic cleavages of the two propionic acid substitutes. Thus the precursor ion at m/z 563 is frag. mented by the loss of one and two CH2COOH radicals to give product ions at m/z 504 (563 − 59) and 445 [563 − (2 × 59)], respectively. Again, as for coproporphyrin, the peak at m/z 490 was derived from . the ion at m/z 504 by the loss of a CH3 radical followed by back protonation (504 − 15 + 1 = 490). The ion at m/z 431 was similarly . derived from the loss of a CH3 radical by the ion at m/z 445 followed by protonation. Elimination of H2O from a protonated molecule was also observed, giving the ion at m/z 545 (563 − 18).

3.4.12. HPLC/ESI-MS and MS/MS of mesotetraphenylporphyrin derivatives A wide variety of compounds based on the meso-tetrakistetraphenyporphyrin structure have been synthesised as potential photosensitisers for photodynamic therapy.19–22 5,10,15,20-Tetra(m-hydroxyphenyl)-chlorin and 5,10,15,20-tetra(m-hydroxyphenyl)bacteriochlorin (Fig. 3.35) are examples.19,20 HPLC/ESI-MS and MS/MS have been developed for the separation and analysis of hydroporphyrins of the meso-tetra (hydroxyphenyl) porphyrin series. The separation is best carried out by reversed-phase HPLC with various proportions of acetonitrile in 0.1% trifluoroacetic acid (TFA) as mobile phase.23–25 TFA is believed to be an unfavourable mobile phase component for HPLC/ESI-MS, since it can cause ion suppression by forming neutral ion pairs with protonated basic molecules in positive mode ESI-MS and suppressing the ionisation of acidic compounds in negative ESI-MS. Formic acid is often used as an alternative to TFA in HPLC/ESI-MS. However, TFA provides far superior separation than formic acid for porphyrins and the ion suppression effect is minimal and does not have a significant impact on the sensitivity of detection. Figure 3.36 shows the HPLC separation of β-hydroxy-m-THPC (m/z 697), m-THPC (m/z 681) and m-THPP (m/z 679) in the liver extract of a mouse treated with m-THPC. The separation was carried out on a Hypersil-ODS column with 77% (v/v) acetonitrile in 0.1%

HN

N

N

OH

NH

N

N

HN

N HN OH

OH

HO

HO

m-THPP

m-THPC OH

OH

OH OH

HO

NH N

OH

HO

N

NH

HN

N

HO-m-THPC

NH

HN

OH

HO

OH

HO

N

N

OH

HO

(HO)2-m-THPC

N HN OH

HO

HO-m-THPBC

Figure 3.35. Chemical structures of 5,10,15,20-tetra(m-hydroxyphenyl)porphyrin (m-THPP); 5,10,15,20-tetra(mhydroxyphenyl)chlorin (m-THPC); 5,10,15,20-tetra(m-hydroxyphenyl)bacteriochlorin (m-THPBC); β-hydroxym-THPC (HO-m-THPC); β-dihydroxy-m-THPC [(HO)2-m-THPC], and β-hydroxy-m-THPBC (HO-m-THPBC).

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HO

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OH

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98

OH

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Figure 3.36. HPLC/ESI-MS of β-hydroxy-m-THPC (m/z 697), m-THPC (m/z 681) and m-THPP (m/z 679). Column: Hypersil-ODS; eluent: acetonitrile/0.1% TFA (77:23, v/v).

TFA as the mobile phase. m-THPP and β-hydroxy-m-THPC are oxidation products of m-THPC. The compounds have been characterised by ESI-MS/MS analysis. The most characteristic fragmentation pattern of this group of compounds is the sequential elimination of two of the four phenolic rings. Thus on tandem MS/MS, the [M + H]+ ion of m-THPC at m/z 681 gave product ions at m/z 588 (681 − 93), 495 [681 − (2 × 93)], respectively. The tetrapyrrole macrocycle is relatively stable to fragmentation. The precursor ion of m-THPP (m/z 679) similarly gave characteristic product ions at m/z 586 and 493, respectively. Protonation of the ion at m/z 493 to give a peak at m/z 494 was also observed. For β-hydroxy-m-THPC (m/z 697), elimination of H2O from the precursor ion to give the ion at m/z 679 (697 − 18) is a

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Figure 3.37. HPLC/ESI-MS chromatogram of m-THPC (m/z 681), m-THPBC (m/z 683), β-hydroxy-m-THPC (m/z 697), and β-hydroxy-m-THPBC (m/z 699). Column: Hypersil-BDS column (100 mm × 4.6 mm i.d., 3 µ m particle size); elution: linear gradient from 50% A (0.1% TFA in water): 50% B (0.1% TFA in acetonitrile) to 100% B in 15 min.

characteristic feature. The pathway was then similar to that observed for m-THPP. Figure 3.37 shows the HPLC/ESI-MS chromatogram of m-THPC (m/z 681), m-THPBC (m/z 683), β-hydroxy-m-THPC (m/z 697) and βhydroxy-m-THPBC (m/z 699). The separation was carried out on a Hypersil-BDS column (100 mm × 4.6 mm i.d., 3 µ m particle size) by linear gradient elution from 50% A (0.1% TFA in water): 50% B (0.1% TFA in acetonitrile) to 100% B in 15 min at a flow rate of 1 ml/min. The retention time of m-THPC, m-THPBC, β-hydroxy-m-THPC and βhydroxy-m-THPBC were 12.3, 9.6, 5.4, and 4.6 minutes, respectively.

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ESI-MS/MS product ion spectrum and fragmentation pathway of β-dihydroxy-m-THPC.

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Figure 3.38.

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It is interesting that under these HPLC conditions, m-THPP was eluted at a retention time of about 3 minutes, way before elution of m-THPC. A probable explanation is that at higher mobile phase acidity protonation of the porphyrin nitrogen is enhanced, leading to an increase in polarity and therefore rapid elution. The TFA/acetonitrile system is therefore a very flexible HPLC system for the separation of these compounds by allowing easy manipulation of the mobile phase to suit a particular application. β-Hydroxy derivatives of chlorin and bacteriochlorin are easily characterised by the ease in which they eliminate H2O when subject to tandem MS/MS analysis. Thus β-hydroxy-m-THPC gave a peak at m/z 679 (697 − 18) and β-hydroxy-m-THPBC an ion at m/z 681 (699 − 18), respectively. Chlorin and bacteriochlorin can also form dihydroxy derivatives and they also show characteristic tandem MS/MS fragmentation patterns. The fragmentation of β-dihydroxy-m-THPC shown in Fig. 3.38 is a typical example. The [M + H]+ ion of β-dihydroxy-m-THPC is at m/z 713. This precursor ion eliminated H2O to give the ion at m/z 695 which was then cleaved by losing CO from the reduced pyrrole ring to give the base peak at m/z 667 (see insert in Fig. 3.38). The above HPLC/ESI-MS/MS systems are expected to be suitable for the analysis of related synthetic compounds.

References 1. Hayen H and Karst U. Strategies for the liquid chromatographic-mass spectrometric analysis of non-polar compounds (review). Journal of Chromatography A 2003; 1000: 549–565. 2. Fenn JB, Mann M, Meng CK, Wong SF and Whitehouse CM. Electrospray ionization for mass spectrometry of large biomolecules (review). Science 1989; 246(4926): 64–71. 3. Tanaka K, Waki H, Ido Y, Akita S, Yoshida Y, Yoshida T and Matsuo T. Protein and polymer analyses up to m/z 100000 by laser ionization time-of-flight mass spectrometry. Rapid Communications in Mass Spectrometry 1988; 2(8): 151–153.

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4. Karas M and Hillenkamp F. Laser desorption ionization of proteins with molecular masses exceeding 10000 daltons. Analytical Chemistry 1988; 60: 2299–2301. 5. Raffaelli A and Saba A. Atmospheric pressure photoionization mass spectrometry. Mass Spectrometry Review 2003; 22(5): 318–331. 6. Robb DB and Blades MW. State-of-the-art atmospheric pressure photoionization for LC/MS. Analytica Chimica Acta 2008; 627: 34–49. 7. Luo J, Lamb JH and Lim CK. Analysis of urinary and faecal porphyrin excretion patterns in human porphyrias by fast atom bombardment mass spectrometry. Journal of Pharmaceutical and Biomedical Analysis 1997; 15: 1289–1294. 8. Jones RM, Lamb JH and Lim CK. Urinary porphyrin profiles by laser-desorption ionization time-of-flight mass spectrometry without the use of classical matrices. Rapid Communications in Mass Spectrometry 1995; 9: 921–923. 9. Razzaque MA, Lord GA and Lim CK. Amino acid and peptide conjugates of protoporphyrin: preparation and analysis by high-performance liquid chromatography, high-performance liquid chromatography/electrospray ionization mass spectrometry and matrix-assisted laser desorption/ ionization time-of-flight mass spectrometry. Rapid Communications in Mass Spectrometry 2002; 16: 1675–1679. 10. Qian K, Mennito AS, Edward KE and Ferrughelli DT. Observation of vanadyl porphyrins and sulphur-containing vanadyl porphyrins in a petroleum asphaltene by atmospheric pressure photoionization Fourier transform ion cyclotron resonance mass spectrometry. Rapid Communications in Mass Spectrometry 2008; 22(14): 2153–2160. 11. Danton M and Lim CK. Identification of monovinyl tripropionic acid porphyrins and metabolites from faeces of patients with hereditary coproporphyria by high-performance liquid chromatography/electrospray ionization quadrupole time-of-flight tandem mass spectrometry. Rapid Communications in Mass Spectrometry 2004; 18: 2309–2316. 12. Lim CK, Danton M, Clothier B and Smith AG. Dihydroxy-, hydroxyspirolactone- and dihydroxyspirolactone-urochlorins induced by 2,3,7,8-tetrachlorodibenzo-p-dioxin in liver of mice. Chemical Research in Toxicology 2006; 19: 1660–1667.

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13. Danton M and Lim CK. Porphyrin profiles in blood, urine and faeces by HPLC/electrospray ionization tandem mass spectrometry. Biomedical Chromatography 2006; 20(6–7): 612–621. 14. Danton M and Lim CK. High-performance liquid chromatography/electrospray ionization tandem mass spectrometry of hydroxylated uroporphyrin and urochlorin derivatives formed by photochemical oxidation of uroporphyrinogen I. Biomedical Chromatography 2007; 21(5): 534–545. 15. Silva EM, Domingues P, Tomé JP, Faustino MA, Neves MG, Tomé AC, Dauzonne D, Silva AM Cavaleiro JA, Ferrer-Correia AJ and Domingues MR. Electrospray tandem mass spectrometry of beta-nitroalkenyl mesotetraphenylporphyrins. European Journal of Mass Spectrometry 2008; 14(1): 49–59. 16. Keller BO, Sui J, Young AB and Whittal RM. Interferences and contaminants encountered in modern mass spectrometry. Analytica Chimica Acta 2008; 627: 71–81. 17. Danton M and Lim CK. Porphomethene inhibitor of uroporphyrinogen decarboxylase: analysis by high-performance liquid chromatography/ electrospray ionization tandem mass spectrometry. Biomedical Chromatography 2007; 21(7): 661–663. 18. Lin W and Timkovich R. Oxygenated tetrapyrroles produced from porphyrinogens. Bioorganic Chemistry 1994; 22: 72–94. 19. Bonnett R, White RD, Winfield UJ and Berenbaum MC. Hydroporphyrins of the meso-tetra(hydroxyphenyl)porphyrin series as tumour photosensitizers. Biochemical Journal 1989; 261(1): 277–280. 20. Bonnett R, Nizhnik AN, White SG and Berenbaum MC. Porphyrin sensitizers in tumour phototherapy. Novel sensitizers of the chlorin and bacteriochlorin class with amphiphilic properties. Journal of Photochemistry and Photobiology B. Biology 1990; 6(1–2): 29–37. 21. Serra VV, Domingues MR, Faustino MA, Domingues P, Tomé JP, Neves MG, Tomé AC, Cavaleiro JA and Ferrer-Correia AJ. Electrospray tandem mass spectrometry of new porphyrin amino acid conjugates. Rapid Communications in Mass Spectrometry 2005; 19(18): 2569–2580. 22. Frochot C, Di Stasio B, Vanderesse R, Belgy M-J, Dodeller M, Guillemin F, Viriot M-L and Barberi-Heyob M. Interest of RGD-containing linear or cyclic peptide targeted tetraphenylchlorin as novel photosensitizers for

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selective photodynamic activity. Bioorganic Chemistry 2007; 35: 205–220. 23. Wang Q, Altermatt HJ, Ris H-B, Reynolds, BE, Stewart JCM, Bonnett R and Lim CK. Determination of 5,10,15,20-tetra-(m-hydroxyphenyl)chlorin in tissue by high-performance liquid chromatography. Biomedical Chromatography 1993; 7(3): 155–157. 24. Cai H, Wang Q, Luo J and Lim CK. Study of temoporfin metabolism by HPLC and elecrospray mass spectrometry. Biomedical Chromatography 1999; 13(5): 354–359. 25. Jones RM, Wang Q, Lamb JH, Djelal BD, Bonnett R and Lim CK. Identification of photochemical oxidation products of 5,10,15,20-tetra(m-hydroxyphenyl)chlorin by on-line high-performance liquid chromatography-eectrospray ionization tandem mass spectrometry. Journal of Chromatography A 1996; 722: 257–265.

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CHAPTER 4

Porphyrin Profiles in Blood, Urine and Faeces by HPLC and HPLC/ESI-MS

4.1. Introduction Haem biosynthesis is effectively regulated and controlled by a feedback mechanism so that the amount of haem required for the formation of the various haemoproteins is readily made, with little waste of the intermediates. The haem precursors, ALA, PBG, and porphyrins are therefore accumulated in various tissues and excreted in urine and faeces only in relatively small amounts under normal conditions. ALA and PBG are carried by the circulating blood to the kidney and excreted only in the urine. Normal urinary ALA and PBG levels are <34 µ mol/l and <10 µ mol/l, respectively. They are found in concentrations below the limits of detection in the plasma. Normal human urine contains small amounts of porphyrins (formed by aromatisation of excreted porphyrinogens), predominantly coproporphyrins with approximately 75–80% of the type III isomer. Uroporphyrin and traces of hepta-, hexa- and pentacarboxylicporphyrins are also present. Coproporphyrinogen has been shown to be normally excreted exclusively in the urine and coproporphyrin in the bile and faeces, unless there is damage to the hepatic excretory mechanism. Protoporphyrin is excreted exclusively in the faeces via the biliary route. Faecal protoporphyrin may also be derived from chlorophyll and haemoprotein of dietary origin or from intestinal haemorrhages. 107

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Normal faecal protoporphyrin excretion, which constitutes over 70% of total faecal porphyrins, is therefore variable. Coproporphyrin (approximately 70% type I isomer) is also excreted in faeces via the bile flow into the intestinal tract. Normal human erythrocytes contain approximately 20–60 µ g protoporphyrin and 1–2 µ g coproporphyrin per 100 ml of packed red cells. Normal human plasma contains little or no porphyrins. Enzyme defects in the haem biosynthetic pathway causes genetic diseases known as porphyrias,1–3 which are characterised by reduced activity of specific enzymes and the excessive production, accumulation and excretion of porphyrins and/or porphyrin precursors. Porphyrias due to deficiency in the activity of each enzyme of the haem biosynthetic pathway have been described, except for the first enzyme ALA-synthase. Porphyrias may also be caused by environmental chemicals and toxins that affect an enzyme of the haem biosynthetic pathway. The main types of porphyrias and the sites of enzyme deficiencies are summarised in Table 4.1. Porphyrias are diagnosed by clinical and laboratory investigation.4–6 Each porphyria has characteristic clinical features and patterns of haem precursor overproduction. The analysis of porphyrins in blood, urine and faeces is therefore essential for the biochemical diagnosis and differentiation of the porphyrias. HPLC with spectrophotometric or fluorometric detection is commonly used for the quantitative and profile analysis of porphyrins.7,8 More recently, HPLC/ESI-MS/MS provides additional resolution and specificity for the unequivocal identification of porphyrins, thus significantly improving the diagnostic application of porphyrin profile analysis.9,10

4.2. Sample Collection and Preparation To prevent photo-induced degradation of PBG and porphyrins, all samples must be protected from light. Random urine (10–20 ml) and faeces (2–5 g) are collected without preservative; blood samples are collected with EDTA as anticoagulant. Samples are best analysed fresh but can be stored in a freezer at −20°C until analysis.

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Table 4.1. Summary of Haem Biosynthesis, Sites of Enzyme Deficiencies and Porphyrias. Enzymes

Glycine + Succinyl CoA

Porphyrias

ALA synthase ALA ALA-D deficiency porphyria

ALA dehydratase PBG

Acute intermittent porphyria (AIP)

PBG deaminase (Hydroxymethylbilane synthase) Hydroxymethylbilane Uroporphyrinogen III synthase Uroporphyrinogen III Uroporphyrinogen decarboxylase Coproporphyrinogen III

Congenital erythropoietic porphyria (CEP) Porphyria cutanea tarda (PCT) Familial/ sporadic/toxic Hereditary coproporphyra (HCP)

Coproporphyrinogen oxidase Protoporphyrinogen IX Protoporphyrinogen oxidase

Variegate porphyria (VP) Protoporphyrin IX

Ferrochelatase Haem

Erythropoietic Protoporphyria (EPP)

4.2.1. Preparation of urine samples Fresh, clear urine sample (1 ml) is centrifuged for 1 minute at 9600 × g in a 1.5 ml polypropylene disposable sample cup in a micro-centrifuge and 100 to 500 µ l of the supernatant injected into the HPLC. Urine often forms precipitates on standing or when frozen at −20°C or below. The precipitates, which include calcium phosphate, may contain adsorbed porphyrins and must be re-dissolved before HPLC separation. The sample (5 ml) is vortex-mixed with 0.2 ml of concentrated HCl until the precipitate re-dissolves. The solution is centrifuged at 9600 × g for 1 minute and the supernatant used for analysis.

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For urine containing low concentrations of porphyrins, or when analysis and identification of a minor component is required, solid phase extraction can be used to concentrate the porphyrins before analysis. This is most conveniently carried out on a C18 extraction cartridge widely available from most HPLC packing manufacturers. A typical extraction procedure is as follows: 1. Condition a Bond Elute C18 or C8 cartridge (30 mg or 50 mg) with 2 ml of methanol/acetonitrile (9:1, v/v) followed successively by 2 ml of water and 2 ml of 1 M ammonium acetate — acetic acid buffer (pH 5.16). 2. Mix the urine sample with an equal volume of 1 M ammonium acetate-acetic acid buffer (pH 5.16). 3. Load the urine sample (up to 10 ml) slowly into the cartridge. 4. Wash the cartridge with 2 ml of 1 M ammonium acetate-acetic acid buffer (pH 5.16). 5. Elute porphyrins from the cartridge with 2 ml of methanol/ acetonitrile (9:1, v/v). 6. Evaporate the eluate to dryness and re-dissolve porphyrins in 150 µl of concentrated HCl. 7. Dilute with 350 µl of 1 M ammonium acetate solution just before HPLC analysis.

4.2.2. Preparation of faecal sample The procedure used to extract faecal porphyrins11 for spectrophotometric assay is also suitable for preparing porphyrin solutions for HPLC analysis as follows: 1. Vortex-mix about 50–100 mg of faeces with 1 ml of concentrated HCl to disintegrate the faecal particles and leave to stand for 5 minutes with occasional vortex-mixing. 2. Add 3 ml of peroxide free ether and vortex-mix to give an emulsion. 3. Add 3 ml of water and again vortex-mix.

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4. Centrifuge to give an upper ether layer, a pad of insoluble material at the interface and a lower HCl layer containing porphyrins. 5. Discard the ether layer and withdraw 1 ml of the HCl solution. 6. Centrifuge at 9600 × g for 1 minute to obtain a clear solution for HPLC analysis.

4.2.3. Extraction of porphyrins in plasma The most commonly used method for extracting plasma porphyrins is by solvent partition as follows: 1. Add 5 ml of peroxide-free ether/glacial acetic acid (4:1, v/v) to 0.5 ml of plasma in a 15 ml centrifuge tube and vortex-mix for 1 minute. 2. Centrifuge for 10 minutes at 2000 × g and collect the supernatant into a clean test tube. 3. Add 3 ml of 2.7 M HCl and vortex-mix for 30 seconds. 4. Remove the lower acid layer for HPLC analysis. Plasma porphyrins may also be extracted with a 1:1 (v/v) mixture of 20% (w/v) trichloroacetic acid (TCA)/dimethyl sulphoxide (DMSO) as follows:10 1. Add 0.5 ml of freshly prepared 1:1 (v/v) mixture of 20% TCA/DMSO to 0.5 ml of plasma and vortex-mix for 1 minute. The extraction mixture must be pre-mixed and must not be added to the plasma sample separately. 2. Centrifuge for 10 minutes at 2000 × g and collect the supernatant. 3. Dilute the supernatant with an equal volume of 1 M ammonium acetate-acetic acid buffer (pH 5.16) before HPLC analysis. A mixture of methanol/DMSO (4:1, v/v) has also been used to extract plasma porphyrins. Plasma (250 µ l) is vortex-mixed with 1 ml of the extraction mixture and centrifuged. The supernatant is diluted with an equal volume of 1 M ammonium acetate buffer

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just before HPLC analysis. Again, the extraction mixture must be pre-mixed.

4.2.4. Extraction of red blood cell porphyrins Total red blood cell porphyrin can be extracted with diethyl ether/glacial acetic acid12 as follows: 1. Add 0.05 ml of EDTA anti-coagulated whole blood to 0.45 ml of physiological saline with vortex-mixing. 2. Add 5 ml of a mixture of freshly prepared diethyl ether/glacial acetic acid (4:1, v/v) and continue vortex-mixing for a further 30 sec. 3. Centrifuge and transfer supernatant into a fresh tube. 4. Add 3.0 ml of 2.7 M HCl, vortex-mix and centrifuge. Discard the upper ether layer and transfer the lower aqueous acid solution of porphyrins into a clean tube. The porphyrin solution may be used for spectrofluorometric determination12 or HPLC separation. Red blood cells may contain protoporphyrin as well as Znprotoporphyrin. For the extraction of Zn-protoporphyrin a neutral extraction solvent, such as ethanol,13 is essential in order to avoid de-metallation of Zn-protoporphyrin and the extraction of haemin. Heparin or EDTA anti-coagulated whole blood is used and a typical extraction procedure is as follows: 1. Add 150 µ l of distilled water to 50 µ l of whole blood and while vortex-mixing add 1 ml of absolute ethanol. 2. Vortex-mix for 20 seconds and centrifuge at 2000 × g for 10 minutes. 3. Dilute the ethanol extract with an equal volume of 1 M ammonium acetate buffer (pH 5.16) just before injection into the HPLC. Ethanol may be replaced with 1 ml of a mixture of methanol/ DMSO (4:1, v/v) for extraction10 in the above procedure.

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4.3. Porphyrin Excretion Patterns and Enzyme Assays in the Porphyrias 4.3.1. Aminolaevulinic acid dehydratase deficiency porphyria (ADP) ADP is a very rare autosomal dominant disorder14 characterised by marked increased excretion of ALA and coproporphyrin (mainly type III) in the urine. There is also a moderate increase in urinary PBG concentration. The faecal porphyrin excretion is normal. The condition is very similar to lead poisoning with decrease ALA-D activity and increased Zn-protoporphyrin in erythrocytes. They can, however, be distinguished by determining blood lead concentration which is obviously elevated in lead poisoning and by measuring erythrocyte ALA-D activity. ALA-D inhibition has been shown to correlate with lead concentration in whole blood, the logarithm of ALA-D activity decreasing linearly as blood lead concentration increases. The inhibition can be reversed in lead poisoning by incubating the inhibited enzyme with zinc and dithiothreitol (DTT) in vitro but not in ADP. The ratio of ALA-D activity before and after reactivation with zinc and DTT is thus an important indicator of blood lead levels.

4.3.1.1. Determination of ALA and PBG ALA is not usually measured in clinical laboratories. PBG in urine is commonly determined using a commercially available kit from BioRad Laboratories which is based on the method described by Mauzerall and Granick.15 It involves anion-exchange purification of PBG followed by reaction with 4-dimethylaminobenzaldehyde (Ehrlich’s reagent) in acid to give a red colour product with a characteristic absorption maximum at 553 nm and a shoulder at about 540 nm. The kit is provided with reagents, supplies and experimental procedure instructions from the manufacturer. PBG has also been determined by HPLC following derivatisation with Ehrlich’s reagent.16 A polybenzimidazole resin column (2.5 ml resin, 150–250 µ m particle size) from Celanese Corp., Charlotte,

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NC, USA was used to extract PBG from urine for derivatisation. The derivative was separated by reversed-phase HPLC on a Whatman RP ODS-1 column (250 mm × 5.6 mm, 10 µ m particle size) by fast gradient elution from 10% sodium phosphate buffer (10 mmol/l, pH 3.0) in methanol to 100% methanol in 3 min and maintained at 100% methanol for a further 3 min. A stable isotope dilution HPLC/MS/MS method has been described for the sensitive and specific determination of PBG in urine using [2,413 C]PBG as internal standard.17 PBG was extracted from urine using an Oasis HLB extraction cartridge (1 ml, 30 mg of packing, Waters, USA) and separated by reversed-phase HPLC on a Supelcosil LC-18 column (33 mm × 4.6 mm, 3 µ m particle size) for tandem MS/MS determination by the selected-reaction monitoring (SRM) mode. PBG and [2,413 C]PBG were monitored through their own precursor and product ion settings (m/z 227 to 210 and m/z 229 to 212, respectively). The HPLC mobile phase was acetonitrile/1.4 g/l formic acid (30:100, v/v). PBG is not well retained on a reversed-phase column without an ion-pairing agent. Resolution was achieved mainly by mass spectrometry. PBG is well-retained on reversed-phase HPLC columns in the presence of an ion-pairing agent such as 1-heptanesulphonic acid.18 Micellar electrokinetic capillary chromatography (MEKC) has also been employed for the analysis of PBG.19 The separation was carried out in a 72 cm fused-silica capillary (50 cm to the detector) with an inner and outer diameter of 50 and 370 µ m, respectively. PBG was detected at 220 nm, although the charged PBG molecule can also be detected at the 400–420 nm regions. The running voltage and temperature was 20–25 kV and 30°C, respectively. The running buffer was 20 mM NaH2PO4 and 20 mM Na2B4O4 containing 50 mM sodium dodecyl sulphate adjusted to pH 9.5 with 1 M NaOH. The method has been applied for the determination of ALA-D activities in human erythrocytes. Red blood cells are prepared as follows: 1. Centrifuge blood collected in a heparinised tube at 2000 × g for 15 minutes at 4°C. 2. Discard plasma and leucocytes.

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3. Wash erythrocytes with cold 0.15 M NaCl solution and centrifuge at 2000 × g for 10 minutes after washing. 4. Repeat washing and centrifuging as described in step 3. 5. Store washed erythrocytes at −20°C if not use immediately. The procedure for ALA-D assay is as follows: 1. Vortex-mix 10 µl of washed red blood cells with 200 µl of 0.2% (v/v) Triton X-100 in water and 200 µl of 0.2 M potassium phosphate buffer, pH 6.8, containing 50 mM DTT. 2. Add 200 µl of 20 mM ALA solution in water and incubate the mixture for 1 hour at 37°C in the dark in a shaking water bath. 3. Terminate reaction by adding 500 µl of ice-cold 10% (w/v) TCA. 4. Vortex-mix and centrifuge for 5 minutes at 2000 × g. 5. Load supernatant into a C18 Sep-Pak cartridge (1 ml, 30 mg packing) which has been pre-conditioned by washing successively with methanol (2 ml), water (2 ml) and 5 mM sodium acetate buffer, pH 3.5, containing 5 mM octanesulphonic acid (2 ml). 6. Wash cartridge with 1 ml of 5 mM sodium acetate buffer, pH 3.5, containing 5 mM octanesulphonic acid to remove potential interfering impurities. 7. Elute PBG from the cartridge with 1 ml of the MEKC running buffer for separation and determination. Figure 4.1 shows the MEKC electropherogram of PBG in the incubation mixture for the determination of ALA-D in erythrocytes. Zero-time blank (Fig. 4.1a) was carried out by using boiled red cells. PBG concentration was determined from a calibration curve constructed by plotting peak areas against concentrations and enzyme activity was expressed as µM of PBG formed per hour per ml of red blood cells at 37°C. MEKC is not suitable for coupling with mass spectrometry because the running solution contains phosphate and borate buffers which are not compatible with mass spectrometry. An alternative is to use capillary zone electrophoresis/electrospray ionisation tandem mass spectrometry (CZE/ESI-MS/MS),20 which is an extremely sensitive and specific method for the analysis of ALA and PBG. The compounds

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Figure 4.1. MEKC electropherogram for the determination of ALA-D in human erythrocytes. Capillary length: 50 cm; running buffer: 20 mM sodium phosphate and 20 mM sodium borate (pH 9.5) containing 50 mM sodium dodecyl sulphate (SDS); voltage: 20 KV; temperature: 30°C.

were effectively separated on a 70 cm fused-silica capillary (75 µm inner diameter) with a mixture of 50 mM ammonium acetate-acetic acid buffer, pH 5.2, and acetonitrile (90:10, v/v) as the running buffer solution. The extracted ion electropherogram for ALA at m/z 132 and PBG at m/z 227 is shown in Fig. 4.2. The ion for protonated PBG at m/z 227 is relatively labile and easily fragmented to give a dominant ion at m/z 210, as shown in the product ion spectrum (Fig. 4.3). Figure 4.4 shows a postulated pathway for this fragmentation in which the basic nitrogen is readily protonated, ammonia is lost and a stable methylenepyrrolenine ion (m/z 210) is formed. The loss of ammonia from protonated PBG was found to occur easily in the electrospray source, even at relatively low cone voltage. The ion at m/z 210 thus provides the best sensitivity for the detection of PBG in quantitative analysis, especially by monitoring the transition m/z 227 to 210 in multiple reaction monitoring (MRM) acquisitions, as used in the HPLC/ESI-MS/MS method for the determination of PBG in

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Figure 4.2. CZE/ESI-MS electropherogram for ALA (m/z 132) and PBG (m/z 210). Capillary: 70 cm long (75 µm o.d. and 375 µm i.d.); running buffer: 50 mM ammonium acetate-acetic acid buffer, pH 5.2/acetonitrile (90:10, v/v); cone voltage: 30 V; source temperature: 70°C.

Figure 4.3.

Product ion spectrum of protonated PBG at m/z 227.

urine. The proposed structures for the minor fragment ions are shown in Fig. 4.4. The protonated ALA molecule (m/z 132) is relatively stable and showed the best sensitivity under the same conditions used for PBG analysis.

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COOH COOH

HOOC - NH3 N NH2 H + H+ Protonated PBG (m/ z 227)

- HCOOH

- C2H2 + N H

COOH

+ H+ COOH HOOC

HOOC

+ 2H

Methylenepyrrolenine (m/ z 210)

H

+

CH

H2C H

CH

HN

HN

m/ z 186

m/ z 140

- CH3COOH HOOC

CH2+ H

CH HN

m/ z 126

Figure 4.4.

Proposed fragmentation pathway of PBG.

CZE/MS is only available in specialised research laboratories and also requires a skilled operator. In the above method, CZE capillaries were linked to the mass spectrometer via a co-axial probe consisting of a Micromass “Tri-axial” probe, modified by exchanging the probe tip and stainless steel sheath capillary with those as used in a Micromass CE co-axial probe. Samples were injected into the CZE-ESI-MS system by gravity by raising the capillary outlet 15 cm above the probe outlet for 10 seconds. Methanol/aqueous 0.1% formic acid (1:1, v/v) was used as the sheath “make-up” flow solvent at a flow-rate of 6 µl/minute. A voltage of 20 kV was applied to the anodic, injection end of the CZE capillary, with an electrospray voltage of 3.5 kV at the cathodic end, resulting in a net voltage of 16.5 kV for electrophoretic separation. The CZE/ESI-MS data was acquired in continuum mode at a scan rate of 1 spectrum/s, over the range m/z 100–500, using positive ion mode, with a cone voltage of 30 V and a source temperature of 70°C. Product ion spectra were obtained using argon as the collision gas at a pressure of 1.0 × 10−3 mbar with collision potential offset of 20 V. CZE/ESI-MS/MS is applicable to the determination of PBG in urine like HPLC/ESI-MS. The same solid-phase extraction procedure

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using an Oasis HLB column can be used for sample preparation as follows: 1. Condition an Oasis HLB cartridge (1 ml, containing 30 mg packing material) by washing with 1 ml of 1 M acetic acid followed by 1 ml of water. 2. Load 1 ml of freshly acidified urine sample (pH 2.0) into the cartridge. 3. Wash cartridge with 0.5 ml of water. 4. Elute and collect PBG from cartridge with 1 ml of 1 M acetic acid.

4.3.1.2. Determination of erythrocyte Zn-protoporphyrin by HPLC The separation of red blood cell porphyrins in a case of lead exposure showing excessive accumulation of Zn-protoporphyrin is shown in Fig. 4.5a. Mesoporphyrin was used as the internal standard in quantitative analysis. Figure 4.5b shows the red cell porphyrins of a patient with erythropoietic protoporphyria where protoporphyrin predominates. The HPLC conditions were: column, Hypersil SAS (C1bonded phase); eluent, methanol/1 M ammonium acetate buffer, pH 5.16 (80:20, v/v); flow-rate, 1 ml/min; detection, 404 nm.

4.3.2. Acute intermittent porphyria (AIP) AIP is the most prominent acute hepatic porphyria, inherited as an autosomal dominant trait, due to deficient PBG-deaminase or hydroxymethylbilane synthase (HMB-S) activity.1–3 AIP is characterised by the excretion of excess ALA and PBG in the urine. The urinary porphyrin profile during acute attack when PBG concentration is high shows increased levels of uroporphyrins (I and III), coproporphyrins, and to a lesser extent, heptacarboxylic-, hexacarboxylic and penta-carboxylic-porphyrins. Uroporphyrins are mainly formed by non-enzymatic condensation of PBG. Nonenzymatically formed uroporphyrinogens can be decarboxylated to

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Figure 4.5. HPLC separation of red blood cell porphyrins of (a), a patient with erythropoietic protoporphyria and (b), lead poisoning. Column: Hypersil-SAS (C1); eluent: 80% methanol in 1 M ammonium acetate, pH 5.16. Peaks: 1 = Znprotoporphyrin, 2 = mesoporphyrin (internal standard), 3 = protoporphyrin.

coproporphyrinogens by uroporphyrinogen decarboxylase. The urinary porphyrin and occasionally PBG can be normal in asymptomatic adult patients. In childhood AIP carriers, PBG excretion is normal and only about a third of asymptomatic carriers show increased PBG excretion after puberty. Faecal porphyrin excretion is usually normal but may also be increased.21 Determination of erythrocyte HMB-S activity is a convenient first step to the diagnosis of AIP and detection of asymptomatic carriers. A method for the simultaneous determination of HMB-S and uroporphyrinogen III synthase (Urogen III-S) in erythrocytes has been described.22 It is based on the fact that Urogen III-S can be completely deactivated by heating while HMB-S is heat-stable. Thus if red blood cells are heated for one hour at 56°C (or 65°C)23 to deactivate Urogen III-S prior to incubation with PBG, the uroporphyrinogen formed is

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exclusively of the type I isomer derived from the non-enzymatic cyclisation of hydroxymethylbilane (HMB) produced by the HMB-S reaction. This system can therefore be used to generate HMB for incubation with unheated red cells in the coupled-enzyme assay of Urogen III-S. The incubation and procedure for the enzyme assay are as follows: 1. Mix 1.5 g of MgCl2 and 1 ml of Triton X-100 in 1 litre of 50 mMTris/HCl buffer, pH 8.25. This is the incubation medium. 2. Accurately pipette 30 µl of washed red blood cells with a volume adjustable micropipette into 1.40 ml of incubation medium and vortex-mix. 3. Heat the mixture for one hour at 56°C in a water bath in the dark and then cool the mixture down to room temperature. 4. Add 3 µl of unheated red blood cells with a fixed-volume micropipette into the mixture, vortex-mix and pre-incubate for 5 minutes at 37°C. 5. Add 50 µl (83 µg) of PBG substrate, vortex-mix and incubate for 30 minutes at 37°C. 6. Terminate enzyme reaction by adding 1.50 ml of ice-cold 10% (w/v) trichloroacetic acid (TCA) containing 0.5% (w/v) I2, vortexmix and centrifuge at 2000 × g for 15 minutes at 4°C. 7. Collect the clear supernatant for HPLC separation and quantitation of uroporphyrin I and III isomers formed by the enzyme reactions. The total uroporphyrin (I + III isomers) formed is due to the activity of HMB-S in 33 µl of red blood cells, while the uroporphyrin III formed is a result of Urogen III-activity in 3 µl of red blood cells. The enzyme activities were expressed as nmol/l of uroporphyrin/ml red cells/hour and were calculated as follows: HMB-S = (Uro I + Uro III) × 2.983/1000 × 1000/33 × 60/30 Urogen III-S = Uro III × 2.983/1000 × 1000/3 × 60/30 Where Uro I and Uro III are the concentrations (nmol/l) of uroporphyrin I and uroporphyrin III formed, respectively, and 2.983/ 1000 is the dilution factor.

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Figure 4.6. HPLC chromatograms for the simultaneous determination of HMB-S and Urogen III-S in erythrocytes. Uroporphyrin I and III formed in the incubation mixture of (a), a normal subject, (b), a patient with AIP, (c), a patient with VP, (d), a patient with HCP and (e) a patient with CEP. Column: Hypersil-BDS C18 ; eluent: 9% acetonitrile in 1 M ammonium acetate pH 5.16.

Two blanks were analysed in parallel; the first contains enzyme solution without PBG substrate and the second contains PBG and enzyme solution but the reaction was terminated at zero time. Figure 4.6 shows the HPLC chromatograms for the determination of HMB-S and Urogen III-S in a normal control subject and in patients with various types of porphyrias. HMB-S activity in erythrocytes has also been determined using ESI-MS/MS.24 The enzymatically formed uroporphyrin I was extracted into 1-butanol for tandem MS/MS analysis (multiple reaction monitoring, MRM) by monitoring the product ion of uroporphyrin at m/z 727.

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4.3.3. Congenital erythropoietic porphyria (CEP) CEP is a rare autosomal recessive human porphyria due to uroporphyrinogen III synthase defect.1–3 It causes the overproduction and excretion of type I porphyrin isomers, especially uroporphyrin I and coproporphyrin I, in the urine (Fig. 4.7) and coproporphyrin I in the faeces (Fig. 4.7). There is also a marked increased in erythrocyte uroporphyrin I and coproporphyrin I concentrations

Figure 4.7. Urinary and faecal porphyrin profiles in a patient with CEP. (a), urine and (b), faeces. Column: Hypersil-ODS; solvents: 10% acetonitrile in 1 M ammonium acetate, pH 5.16 (A) and 10% acetonitrile in methanol (B); elution: linear gradient from 10% B to 90% B in 30 min followed by isocratic elution at 90% B for a further 10 min. Peaks: 2, 4, 5, 6, 7, and 8 are protoporphyrin, coproporphyrin, penta-, hexa- and hepta-carboxylic porphyrin, and uroporphyrin, respectively; I and III denote type I and type III isomers.

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Figure 4.8. HPLC separation of porphyrins in the plasma of a patient with CEP. HPLC column and conditions are as described in Fig. 4.7 except solvent A was 9% instead of 10% acetonitrile in 1 M ammonium acetate, pH 5.16. Peaks: 1 = uroporphyrin I, 2 = heptacarboxylic porphyrin I, 3 = hexacarboxylic porphyrin I, 4 = pentacarboxylic porphyrin I, 5 = coproporphyrin I.

which could diffuse into, and elevate the plasma porphyrin levels25 (Fig. 4.8). The excessive excretion of uroporphyrin I in urine is accompanied by small quantities of oxygenated derivatives, the hydroxyuroporphyrin I and keto acid uroporphyrin I derivatives. Figure 4.9 shows the separation of meso-hydroxyuroporphyrin I, β-hydroxypropionic acid uroporphyrin I, hydroxyacetic acid uroporphyrin I and hydroxyspirolactone urochlorin I in the urine of a patient with CEP. The compounds showed characteristic tandem ESI-MS/MS fragmentation patterns as described in Chapter 3, Sec. 3.4.4.

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Figure 4.9. HPLC separation of meso-hydroxyuroporphyrin I (peak 1), β-hydroxypropionic acid uroporphyrin I (peak 2), hydroxyacetic acid uroporphyrin I (peak 3), hydroxyspirolactone urochlorin I (peak 4) and uroporphyrin I (peak 5) in the urine of a patient with CEP. Column: Hypersil ODS; solvents: A, 9% acetonitrile in 1 M ammonium acetate, pH 5.16, and B, 10% acetonitrile in methanol; elution program: 0 to 16 min, 0% B (100% A) to 11 % B; 16 to 24 min, isocratic elution at 11% B; 24 to 35 min, isocratic elution at 90% B.

The HPLC/ESI-MS chromatogram of β-ketopropionic acid uroporphyrin I (m/z 845) and ketoacetic acid uroporphyrin I (m/z 845) in the CEP urine is shown in Fig. 4.10. The peaks were characterised by tandem MS/MS analysis with characteristic fragmentation patterns (see Sec. 3.4.4). These compounds are eluted after uroporphyrin (m/z 831) because intramolecular hydrogen-bonding (see Chapter 3, Fig. 3.20) increases their hydrophobicity. β -Ketopropionic acid uroporphyrin I eluted before ketoacetic acid uroporphyrin I. The m/z 847 peaks are hydroxyuroporphyrins.

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Figure 4.10. HPLC/ESI-MS chromatogram of β-ketopropionic acid uroporphyrin I (m/z 845) and ketoacetic acid uroporphyrin I (m/z 845) in the urine of a patient with CEP. Column: Hypersil-BDS C18; solvents: 9% acetonitrile in 1 M ammonium acetate, pH 5.16 (A) and 9% acetonitrile in methanol (B); elution program: linear gradient from 0% B to 90% B in 60 min at a flow rate of 1 ml/min. The flow was split in the ratio 1:9 such that 100 µl/min entered the mass spectrometer.

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Although CEP is normally presented at birth, late-onset CEP presented in adults is also known to occur.26 The characteristic type I series of porphyrins in blood, urine and faeces allows this condition to be differentiated from other human porphyrias. Determination of uroporphyrinogen III synthase activity in erythrocytes22,23 is useful for the diagnosis of heterozygotes and homozygotes with CEP and differentiation from other forms of human porphyrias.

4.3.4. Porphyria cutanea tarda (PCT) PCT is the commonest type of human porphyria1–3,27 caused by partial deficiency in the activity of uroporphyrinogen decarboxylase (Urogen-D). There are three main types: familial, sporadic and toxic PCT induced by exposure to toxic chemicals such as polyhalogenated aromatic hydrocarbons, for example, hexachlorobenzene and 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD). All types show decreased hepatic Urogen-D activity. In familial PCT, which is inherited as an autosomal dominant trait, erythrocyte enzyme activity is also reduced. Sporadic and toxic PCT show normal erythrocyte Urogen-D activity. A very rare form of PCT with homozygous inheritance, hepatoerythropoietic porphyria, has also been described. The faecal porphyrin profile (Fig. 4.11) shows elevation of every porphyrin intermediate between uro- and copro-porphyrin, with significant increase in heptacarboxylic porphyrin III (7d). Pentacarboxylic porphyrin III (5IIIabd) may occasionally also be elevated significantly. Hexacarboxylic porphyrin III (6IIIad) is increased to a lesser extent. The urinary porphyrin profile of PCT (Fig. 4.11) is dominated by uroporphyrin I and III and heptacarboxylic porphyrin III. The heptacarboxylic porphyrin is over 95% type III with the ring D acetic acid group decarboxylated into a methyl group (7d). Varying amounts of mono-oxygenated porphyrins, such as hydroxyand keto acid-porphyrins, may be present in the urine of PCT patients. These are the oxidation products of porphyrinogens, probably derived from a combination of both enzymatic and chemical

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Figure 4.11. HPLC profiles of faecal (a) and urinary (b) porphyrins of a patient with PCT. HPLC conditions and peak identification are as described in Fig. 4.7.

oxidation, and are difficult to identify using spectrophotometric or fluorescence detection. They are often reported as “unknown” porphyrins, for examples, the peaks eluted before uroporphyrin I and between uroporphyrin III and heptacarboxylic porphyrin III in Fig. 4.12. The separation was carried out by an extended, 120-minute gradient elution program. These porphyrins can be positively identified using HPLC/ESI-MS instead of fluorescence detection. Peaks 1 to 5 were shown to be hydroxylated uroporphyrins and peaks 6 and 7 were β-ketopropionic acid uropophyrin and ketoacetic acid uroporphyrin, respectively.

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Figure 4.12. HPLC separation of porphyrins in the urine of a patient with PCT using an extended, 120-min gradient elution program on a Hypersil-ODS column with 9% acetonitrile in 1 M ammonium acetate, pH 5.16, and 10% acetonitrile in methanol as gradient elution solvent mixtures.

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Figure 4.13. HPLC/ESI-MS chromatogram of keto acid uroporphyrins (m/z 845) in the urine of a patient with PCT.

Figures 4.13, 4.14 and 4.15 show the presence of keto acid uroporphyrins (m/z 845), keto acid heptacarboxylic porphyrins (m/z 801) and keto acid pentacarboxylic porphyrins (m/z 713), respectively. Keto acid porphyrins are more hydrophobic than, and are therefore eluted after, the parent compounds because of their ability to form stable intramolecular H-bonded ring structures. They all show the characteristic fragmentation patterns of keto acid porphyrins (see Chapter 3, Sec. 3.4.4). Hydroxy- and keto acid-pentacarboxylic porphyrins are also detected in the faeces of PCT patients, especially when pentacarboxylic porphyrin concentration is significantly elevated. They are probably formed by a combination of enzymatic and non-enzymatic oxidation prior to excretion in the faeces. The detection of these

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Figure 4.14. HPLC/ESI-MS chromatogram of keto acid heptacarboxylic porphyrins (m/z 801) in the urine of a patient with PCT.

porphyrins in urine and faeces may serve as additional evidence of PCT, as with the presence of isocoproporphyrin in faeces. The accumulation of pentacarboxylic porphyrinogen, particularly 5IIIabd porphyrinogen, led to the formation and excretion of

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Figure 4.15. HPLC/ESI-MS chromatogram of keto acid pentacarboxylic porphyrins (m/z 713) in the urine of a patient with PCT.

isocoproporphyrin and related compounds (Fig. 4.16) in the faeces of PCT patients.28 The pentacarboxylic porphyrinogen 5IIIabd is a substrate for coproporphyrinogen oxidase29 and is converted to dehydroisocoproporphyrin by the enzyme. Further metabolism by intestinal flora resulted in a series of isocoproporphyrin derivatives (Fig. 4.16). Isocoproporphyrin is not generally available as a chromatographic standard. It elutes after coproporphyrin III by reversed-phase HPLC in the region where several other coproporphyrin derivatives, notably β-ketopropionic acid coproporphyrins and hydroxyethyltricarboxylic porphyrin, also elute. Furthermore, hydroxyethyl- and keto-isocoproporphyrin are eluted in the area where several ketopropionic acid

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Figure 4.16. Chemical structures of isocoproporphyrin and related metabolites. 1 = dehydroisocoproporphyrin, 2 = isocoproporphyrin, 3 = ketoisocoproporphyrin, 4 = hydroxyisocoproporphyrin, 5 = deethylisocoproporphyrin.

pentacarboxylic porphyrin isomers also elute. Figure 4.17 shows the complexity of porphyrins present in the urine of a patient with PCT eluted in this region. Care should therefore be taken when identifying these compounds by HPLC with fluorometric detection.

4.3.4.1. Determination of uroporphyrinogen decarboxylase activity in erythrocytes by HPLC The procedure, using pentacarboxylic porphyrinogen I as substrate, is as follows: 1. Add 10 µl of washed red blood cells to 100 µl of 0.1 M K2HPO4/ KH2PO4 buffer (pH 6.8) containing 150 µmol/l EDTA, 1 ml/l Triton-X 100 and 20 mmol/l dithiothreitol in a 1.5 ml polythene tube.

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Figure 4.17. HPLC separation of porphyrins in the urine of a patient with PCT showing the complexity of peaks eluted in the pentacarboxylic porphyrin and coproporphyrin region.

2. Pre-incubate the mixture for 5 minutes at 37°C. 3. Add 10 µl (20 µM) of pentacarboxylic porphyrinogen I (prepared by reduction of pentacarboxylic porphyrin I with 3% Na/Hg), flush with N2, cap the tube and incubate mixture for 30 minutes at 37°C in the dark. 4. Terminate reaction by vortex-mix with 500 µl of 10% TCA containing 0.5% (w/v) I2/DMSO (1:1, v/v). 5. Centrifuge at 9600 × g for 3 minutes and collect supernatant for HPLC separation and quantitation of coproporphyrin I formed.

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Figure 4.18. HPLC chromatograms for the determination of uroporphyrinogen decarboxylase activity in human erythrocytes using pentacarboxylic porphyringen I as substrate. Peak 1 = coproporphyrin I formed in the incubation mixture of (a) a normal subject and (b) a patient with familial PCT.

Figure 4.18 shows the HPLC chromatogram for the determination of Urogen-D activity in erythrocytes. Pentacarboxylic porphyrin I from commercial sources may contain small amounts of the type III isomer as well as coproporphyrins as impurities and should be purified by HPLC before use. ESI-MS/MS has also been to determine Urogen-D activity in erythrocytes.30 Since coproporphyrin I can be rapidly separated and detected by HPLC with more than adequate sensitivity, tandem mass spectrometry does not provide any advantage.

4.3.5. Hereditary coproporphyria (HCP) This autosomal dominant porphyria is caused by a partial deficiency of coproporphyrinogen oxidase,1–3 the enzyme which catalyses the

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Figure 4.19. HPLC profiles of urinary (a) and faecal (b) porphyrins of a patient with HCP. HPLC conditions and peak identification are as described in Fig. 4.7.

sequential oxidative decarboxylation of the 2- and 4-propionic acid substituents of coproporphyrinogen III into vinyl groups to give protoporphyrinogen IX. The intermediate is 2-vinyl-4,6,7-tripropionic acid porphyrinogen. The porphyrinogens are oxidised to porphyrins and excreted in the faeces. HCP is therefore characterised by the excessive excretion of coproporphyrin III in the faeces (Fig. 4.19). Urinary coproporphyrin III is also increased (Fig. 4.19). Acute attacks in HCP are very rare. The excretion of excessive coproporphyrin III led to the formation of hydroxycoproporphyrins and β-ketopropionic acid coproporphyrins in the urine and faeces. Figure 4.20 shows the HPLC/ESI-MS chromatogram of coproporphyrin III (m/z 655), hydroxycoproporphyrin III (m/z 671) and four β-ketopropionic acid coproporphyrin III isomers (m/z 669) in the urine of a patient with HCP. The four β-ketopropionic acid coproporphyrin III isomers have virtually identical tandem MS/MS fragmentation patterns with a characteristic fragment

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Figure 4.20. HPLC/ESI-MS chromatogram of coproporphyrin III (m/z 655), hydroxycoproporphyrin III (m/z 671) and four β-ketopropionic acid coproporphyrin III isomers (m/z 669) in the urine of a patient with HCP. HPLC/ESI-MS conditions are as described in Fig. 4.10.

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ion at m/z 637 (669 − 32) showing the loss of O2 from the precursor ion (Fig. 4.21). Again, because of intramolecular hydrogenbonding, β-ketopropionic acid coproporphyrins are eluted after coproporphyrin III. A very rare variant of HCP, homozygous HCP or “harderoporphyria”, in which the defect is believed to be in the conversion of the tricarboxylic porphyrinogen into protoporphyrinogen, has also been identified.31 It is characterised by very low coproporphyrinogen oxidase activity (usually <10%) and the excretion of excess 2-vinyl4,6,7-tripropionic acid porphyrin or “harderoporphyrin” in faeces. The term “harderoporphyrin” was used to name the porphyrin, believed to be 2-vinyl-4,6,7-tripropionic acid porphyrin, isolated from the Harderian glands of rats and other rodents.32,33 It is a misnomer and should no longer be used because it has been shown conclusively that the Harderian glands of rats and other rodents contain virtually no tricarboxylic porphyrin34 (see Chapter 5), and that the porphyrin isolated and characterised as tricarboxylic porphyrin was in fact an artefact generated by the isolation procedure.35 Similarly the term “harderoporphyria” should not be used to describe homozygous HCP. Tricarboxylic porphyrin is not available commercially for use as a chromatographic marker. It is not uncommon to wrongly identify a peak as tricarboxylic porphyrin based on its elution between coproporphyrin and protoporphyrin IX when spectrophotometric or fluorometric detection is used. Uneqivocal identification of tricarboxylic porphyrin requires mass spectrometry. Figure 4.22 shows the HPLC/ESI-MS chromatograms of coproporphyrins (CI and CIII; m/z 655), 2-vinyl-4,6,7-tripropionic acid porphyrin (m/z 609), and protoporphyrin (PP; m/z 563) extracted from the faeces of a patient with HCP. 2-Vinyl-4,6,7-tripropionic acid porphyrin was confirmed by tandem MS/MS analysis and the product ion spectrum is shown in Fig. 4.23. The sequential loss of side-chain CH2COOH radicals by benzylic cleavages typical for porphyrins can be clearly seen. Tricarboxylic porphyrin is elevated not only in the faeces of homozygous HCP but also in all HCP.9 The increased excretion of

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Figure 4.21.

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Figure 4.22. HPLC/ESI-MS chromatograms of coproporphyrin I and III (CI and CIII; m/z 655), 2-vinyl-4,6,7-tripropionic acid porphyrin (m/z 609) and protoporphyrin (PP; m/z 563) extracted from the faeces of a patient with HCP. Column: Hypersil-BDS C18 ; solvents: 9% (v/v) acetonitrile in 1 M ammonium acetate buffer, pH 5.16 (A) and 9% (v/v) acetonitrile in methanol (B); elution: linear gradient from 0% B (100% A) to 90% B in 60 min.

tricarboxylic porphyrin in the faeces of HCP also led to the modification of the vinyl group and formation of ethyl-, hydroxyethyl- and keto-derivatives9 (Fig. 4.24) similar to those observed for the isocoproporphyrin series.

4.3.5.1. HPLC assay of coproporphyrinogen oxidase activity in peripheral leucocytes Peripheral blood leucocytes were isolated by erythrocyte sedimentation and NH4Cl lysis.36 The enzyme assay procedure37 is as follows: 1. Freeze-thaw the leucocyte suspension three times before assay. 2. Add 100 µl of leucocytes to 100 µl of Tris/HCl buffer (pH 7.0) containing 1 mmol/l EDTA.

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ESI-MS/MS product ion spectrum of 2-vinyl-4,6,7-tripropionic acid porphyrin (m/z 609; M + H+).

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Figure 4.23.

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CH3 3

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Figure 4.24. Chemical structures of 2-vinyl-4,6,7-tripropionic acid porphyrin and metabolites detected in the faeces of a patient with HCP.

3. Pre-incubate at 37°C for 5 minutes. 4. Add 50 µl of coproporphyrinogen III substrate (final concentration 1 µmol/l), mix and incubate for 1 hour at 37°C in the dark. 5. Terminate reaction by vortex-mix with 750 µl of methanol/ DMSO (4:1, v/v) containing mesoporphyrin as internal standard. 6. Cool in ice for 15 minutes to assist protein precipitation. 7. Centrifuge at 2100 × g for 10 minutes. 8. Collect the clear supernatant for HPLC separation and quantitation of protoporphyrin formed. Figure 4.25 shows the HPLC chromatogram for the measurement of coproporphyrinogen oxidase activity in human leucocytes. ESI-MS/MS has also been used to determine coproporphyrinogen oxidase activity in lymphocytes.30

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Figure 4.25. HPLC chromatograms for the determination of coproporphyrinogen oxidase activities in human leucocytes. (a), a standard mixture of mesoporphyrin (peak 1, internal standard) and protoporphyrin (peak 2); (b), blank (boiled leucocytes); (c), control subject; (d), a patient with HCP. Column: HypersilODS; eluent: 86% methanol in 1 M ammonium acetate, pH 5.16; flow rate: 1.5 ml/min; detector: fluorescence, excitation 400 nm and emission 618 nm.

4.3.6. Variegate porphyria (VP) VP is an autosomal dominant porphyria caused by partial deficiency in the activity of protoporphyrinogen oxidase.1–3 The accumulation of protoporphyrinogen resulted in the formation of covalent protoporphyrinprotein conjugates (probably via addition of cysteine in protein to vinyl in protoporphyrinogen) in the plasma having a characteristic fluorescence emission maximum38,39 at 624–626 nm when scanned between 550–650 nm with an excitation wavelength of 405 nm. This has been widely used for the diagnosis of VP. The addition reaction of

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Figure 4.26. HPLC profiles of (a) urinary and (b) faecal porphyrins of a patient with VP. HPLC conditions and peak identification are as described in Fig. 4.7.

cysteine, glutathione and cysteine containing polypeptides to the vinyl groups of protoporphyrinogen has been demonstrated.40 Protoporphyrin will not react under the same reaction conditions. The faecal porphyrin profile shows elevated protoporphyrin and to a lesser extent coproporphyrin III in VP (Fig. 4.26). Urinary porphyrin is normal during remission but uroporphyrins (I and III) and coproporphyrins may be increased, together with hepta-, hexa-, and penta-carboxylic porphyrins, during acute attack when PBG level is high as observed for AIP.

4.3.6.1. Determination of protoporphyrinogen oxidase by HPLC For the detection of asymptomatic carriers in family with VP, determination of protoporphyrinogen oxidase activity in lymphocytes

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or leucocytes is necessary.39 The HPLC assay procedure41 is as follows: 1. Prepare protoporphyrinogen solution by shaking a solution of protoporphyrin IX in freshly prepared 0.01 M KOH/ethanol (4:1, v/v) with 3% (w/w) sodium amalgam in a N2 flushed and capped tube vigorously until little or no fluorescence detected. 2. Mix the resulting protoporphyrinogen solution with an equal volume of ice-cold 0.25 M Tris-HCl buffer (pH 8.6) containing 5 mM glutathione, 5 mM EDTA and 1% Tween 20 (w/v), flush with N2, capped and keep on ice in the dark. This is the substrate solution. 3. Add 100 µl of leucocyte suspension (or lymphocytes) to 50 µl of 0.25 M Tris-HCl buffer (pH 8.6) containing 5 mM glutathione, 5 mM EDTA and 1% Tween 20 (w/v). 4. Preincubate for 5 minutes at 37°C in the dark. 5. Add 100 µl of protoporphyrinogen IX substrate solution (approximately 35 µM) and incubate for 10 minutes in the dark without shaking. 6. Terminate reaction by vortex-mix with 1 ml of ice-cold methanol/DMSO (4:1, v/v) containing mesoporphyrin as internal standard. 7. Centrifuge for 10 minutes at 4°C and collect supernatant for HPLC separation and determination of protoporphyrin formed. The reduction of protoporphyrin to protoporphyrinogen is often incomplete and removal of protoporphyrin may be necessary. This can be achieved by passing the substrate solution through a small strong anion exchange solid-phase extraction cartridge (Bond-Elut SAX) which has been pre-conditioned by washing with methanol, water and 0.25 M Tris-HCl buffer (pH 9.0). Protoporphyrin was retained by the column while protoporphyrinogen was unretained and passed straight through the column and then collected.42 Protoporphyrinogen and protoporphyrin are also well-separated42 by reversed-phase HPLC, as shown in (Fig. 4.27). The HPLC chromatogram for the measurement of protoporphyrinogen oxidase activity is shown in Fig. 4.28.

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Figure 4.27. HPLC separation of protoporphyrinogen (peak 1) and protoporphyrin (peak 2). Column: Hypersil-ODS; eluent: 83% methanol in 1 M ammonium acetate, pH 5.16, for 7 min, then 95% methanol in the buffer for a further 10 min; flow rate: 1.5 ml/min.

4.3.7. Erythropoietic protoporphyria (EPP) EPP, also known as erythrohepatic protoporphyria (EHP), is an autosomal dominant porphyria resulting from ferrochelatase (FECH) deficiency.1–3 EPP is usually presented in early childhood and is characterised by excessive accumulation of protoporphyrin in erythrocytes, plasma, liver and skin. The determination of protoporphyrin in red blood cells and in plasma is therefore used to diagnose EPP. In EPP, free protoporphyrin levels always exceed that of Znprotoporphyrin. This distinguishes it from lead poisoning (see Fig. 4.5), iron deficiency anaemia, chronic infection, inflammatory disease or malignancy where Zn-protoporphyrin is the predominant red cell porphyrin. A homozygous, recessive mode of inheritance of the disease has also been reported in a small percentage of patients,43–45 with higher protoporphyrin accumulation in erythrocytes. The urinary porphyrin profile is normal in EPP, except in patients who develop liver failure. Faecal protoporphyrin is elevated

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Figure 4.28. HPLC chromatograms for the measurement of protoporphyrinogen oxidase activity in human leucocytes. (a), Protoporphyrin formed in the incubation mixture of a normal subject; (b) blank incubation with boiled leucocytes. Column: Hypersil-ODS; eluent: 88% methanol in 1 M ammonium acetate, pH 5.16; flow rate: 1.5 ml/min. Peaks: 1 = mesoporphyrin (internal standard), 2 = protoporphyrin.

(Fig. 4.29), but cannot be used to diagnose EPP because the profile may be similar to that of normal subjects on haem-containing diets.

4.3.7.1. HPLC assay for ferrochelatase in leucocytes or lymphocytes Bivalent metal ions such as Fe2+, Zn2+, Cu2+ and Co2+ are all substrates for ferrochelatase (FECH). The enzyme also catalyses the insertion of bivalent metal ions into mesoporphyrin. FECH activity is therefore conveniently measured using Zn2+ and the more stable mesoporphyrin as substrates.46,47 Zn-mesoporphyrin is also a fluorescent

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Figure 4.29. HPLC profiles of (a) faecal and (b) red blood cell porphyrins of a patient with EPP.

compound which can be detected with great sensitivity by a fluorescence detector. The incubation mixture consisted of 100 µl of 0.25 M Tris-HCl buffer (pH 8.0) containing Tween 20 or Triton-X-100 (10 g/l) and palmitic acid (1.75 mM), 50 µl of leucocytes or lymphocytes, and 50 µl of mesoporphyrin (200 µM). The mixture was preincubated for 5 minutes at 37°C. The enzyme reaction was then started by adding 50 µl of 200 µM zinc acetate in water. The incubation was continued for 30 minutes at 37°C. The reaction was terminated by vortex-mixed with 1 ml of methanol/DMSO (4:1, v/v) containing 270 µM of EDTA. The mixture was cooled in ice for 15 minutes to ensure complete protein precipitation before centrifugation at 2100 × g for 10 minutes. The supernatant was analysed by HPLC and the chromatogram is shown in Fig. 4.30.

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Figure 4.30. HPLC chromatograms for the determination of ferrochelatase in human leucocytes. (a) Enzyme incubation mixture and (b) blank incubation with boiled leucocytes. Column: Hypersil-ODS; eluent: 88% methanol in 1 M ammonium acetate, pH 5.16; flow rate: 1.5 ml/min. Peaks: 1 = Zn-deuteroporphyrin (internal standard), 2 = Zn-mesoporphyrin, 3 = mesoporphyrin.

4.3.8. Mixed porphyria The co-existence of PCT and VP in some patients, mainly in South Africa, has been described.48 Enzyme assay combined with detailed HPLC or HPLC/MS analysis of porphyrins should allow the diagnosis of this “dual porphyria”.

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References 1. Kappas A, Sassa S, Galgraith RA and Nordmann Y. The porphyrias. In: The Metabolic and Molecular Bases of Inherited Disease, 7th edn, Scriver CR, Beaudet AL, Sly WS, Vale D (eds). McGraw-Hill: New York, 1995; 2103–2159. 2. Bickers DR and Frank J. The porphyrias. In Fitzpatrick’s Dermatology in General Medicine, 7th edn, Wolff K, Goldsmith LA, Katz SI, Gilchrest BA, Paller AS, Leffell DJ (eds). McGraw-Hill: New York, 2008; part 8, section 24, chapter 132. 3. Nordmann Y and Puy H. Human hereditary hepatic porphyrias. Clinica Chimica Acta 2002; 325: 17–37. 4. Elder GH, Smith SG and Smyth JS. Laboratory investigation of the porphyrias. Annals of Clinical Biochemistry 1990; 27: 395–412. 5. Bonkovsky HL and Barnard GF. Diagnosis of porphyric syndromes: A practical approach in the era of molecular biology. Seminars in Liver Disease 1998; 18: 57–66. 6. Deacon AC and Elder GH. Front line tests for the investigation of suspected porphyria. Journal of Clinical Pathology 2001; 54: 500–507. 7. Lim CK and Peters TJ. Urine and faecal porphyrin profiles by reversedphase high-performance liquid chromatography. Clinica Chimica Acta 1984; 139: 55–63. 8. Kuhnel A, Gross U, Jacob K and Doss MO. Studies on coproporphyrin isomers in urine and feces in the porphyrias. Clinica Chimica Acta 1999; 282: 45–58. 9. Danton M and Lim CK. Identification of monovinyl tripropionic acid porphyrins and metaboites from faeces of patients with hereditary coproporphyria by high-performance liquid chromatography/ electrospray ionization quadrupole time-of-flight tandem mass spectrometry. Rapid Communications in Mass Spectrometry 2004; 18: 2309–2316. 10. Danton M and Lim CK. Porphyrin profiles in blood, urine and faeces by HPLC/electrospray ionization tandem mass spectrometry. Biomedical Chromatography 2006; 20(6–7): 612–621. 11. Lockwood WH, Poulous V, Rossi E and Curnow DH. Rapid procedure for fecal porphyrin assay. Clinical Chemistry 1985; 31: 1163–1167.

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12. Blake D, Poulos V and Rossi R. Diagnosis of porphyria-recommended methods for peripheral laboratories. Clinical Biochemistry Reviews 1992; 13(suppl 1): S1–24. 13. Garden JS, Mitchell DG, Jackson KW and Aldous KM. Improved ethanol extraction procedure for determining zinc protoporphyrin in whole blood. Clinical Chemistry 1977; 23: 264–269. 14. Maruno M, Furuyama K, Akagi R, Horie Y, Meguro K, Gabaczewski L, Chiorazzi N, Doss M, Hassoun A, Mercelis R, Verstraeten L, Harper P, Floderus Y, Thunell S and Sassa S. Highly heterogenous nature of deltaaminolevulinate dehydratase (ALAD) deficiencies in ALAD porphyria. Blood 2001; 97(10): 2972–2978. 15. Mauzerall D and Granick S. The occurrence and determination of δ-aminolaevulinic acid and porphobilinogen in urine. Journal of Biological Chemistry 1956; 219: 435–446. 16. Jamani A, Pudek M and Schreiber WE. Liquid chromatographic assay of urinary porphobilinogen. Clinical Chemistry 1989; 35(3): 471–475. 17. Ford RE, Magera MJ, Kloke KM, Chezick PA, Fauq A and McConnell JP. Quantitative measurement of porphobilinogen in urine by stable-isotope dilution liquid chromatography-tandem mass spectrometry. Clinical Chemistry 2001; 47(9): 1627–1632. 18. Crowne H, Lim CK and Samson D. Determination of 5-aminolaevulinic acid dehydrase activity in erythrocytes by high-performance liquid chromatography. Journal of Chromaography 1981; 223: 421–425. 19. Luo JL, Deka J and Lim CK. Determination of 5-aminolaevulinic acid dehydratase activity in erythrocytes and porphobilinogen in urine by micellar electrokinetic capillary chromatography. Journal of Chromatography A 1996; 722: 353–357. 20. Lord GA, Luo JL and Lim CK. Capillary zone electrophoresis/mass spectrometry of 5-aminolaevulinic acid and porphobilinogen. Rapid Communications in Mass Spectrometry 2000; 14: 314–316. 21. Rossi E. Increased fecal porphyrins in acute intermittent porphyria. Clinical Chemistry 1999; 45: 281–283. 22. Wright DJ and Lim CK. Simultaneous determination of hydroxymethylbilane synthase and uroporphyrinogen III synthase in erythrocytes by high-performance liquid chromatography. Biochemical Journal 1983; 213: 85–88.

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23. Tsai SF, Bishop DF and Denick RJ. Coupled-enzyme and direct assays of uroporphyrinogen III synthase activity in human erythrocytes and cultured lymphoblasts. Enzymatic diagnosis of heterozygotes and homozygotes with congenital erythropoietic porphyria. Analytical Biochemistry 1987; 166(1): 120–133. 24. Wang Y, Scott CR, Gelb MH and Turec ek F. Direct assay of enzymes in heme biosynthesis for the detection of porphyrias by tandem mass spectrometry. Porphobilinogen deaminase. Analytical Chemistry 2008; 80: 2606–2611. 25. Gorchein A, Guo R, Lim CK, Raimundo A, Pullon HWH and Bellingham AJ. Porphyrins in urine, plasma, erythrocytes, bile and faeces in a case of congenital erythropoietic porphyria (Gunther’s Disease) treated with blood transfusion and iron chelation: Lack of benefit from oral charcoal. Biomedical Chromatography 1998; 12: 350–356. 26. Western MJ, Nicholson DC, Lim CK, Clark KG, McDonald A, Henderson MA and Williams R. Congenital erythropoietic porphyria (Gunther’s Disease) presenting in a middle age man. International Journal of Biochemistry 1978; 9: 921–926. 27. Elder GH. Porphyria cutanea tarda. Seminars in Liver Disease 1998; 18(1): 67–75. 28. Elder GH. Differentiation of porphyria cutanea tarda symptomatica from other types of porphyria by measurement of isocoproporphyrin in faeces. Journal of Clinical Pathology 1975; 28: 601–607. 29. Cooper CL, Stob CM, Jones MA and Lash TD. Metabolism of pentacarboxylate porphyrinogens by highly purified human coproporphyrinogen oxidase: Further evidence for the existence of an abnormal pathway of heme biosynthesis. Bioorganic & Medicinal Chemistry 2005; 13: 6244–6251. 30. Wang Y, Gatti P, Sadilek M, Scott CR, Turec ek F and Gelb MH. Direct assay of enzymes in heme biosynthesis for the detection of porphyrias by tandem mass spectrometry. Uroporphyrinogen decarboxylase and coproporphyrinogen III oxidase. Analytical Chemistry 2008; 80: 2599–2605. 31. Lamoril J, Puy H, Whatley SD, Martin C, Woolf JR, Da Silva V, Deybach JC and Elder GH. Characterization of mutations in the CPO gene in British patients demonstrates absence of genotype-phenotype correlation

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33.

34.

35. 36. 37.

38. 39.

40.

41.

42.

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and identifies relationship between hereditary coproporphyria and harderoporphyria. American Journal of Human Genetics 2001; 68: 1130–1138. Kennedy GY. Harderoporphyrin: A new porphyrin from the Harderian glands of the rat. Comparative Biochemistry and Physiology 1970; 36: 21–36. Kennedy GY, Jackson AH, Kenner GW and Suckling CJ. Isolation, structure and synthesis of a tricarboxylic porphyrin from the Harderian glands of the rat. FEBS Letters 1970; 6: 9–12. Lim CK, Razzaque MA, Luo J and Farmer PB. Isolation and characterization of protoporphyrin glycoconjugates from rat Harderian gland by HPLC, capillary electrophoresis and HPLC/electrospray ionization MS. Biochemical Journal 2000; 347: 757–761. Gorchein A and Lim CK. Harderoporphyrin: A misnomer. Biomedical Chromatography 2005; 19(8): 565–569. Cutts JH. Cell separation methods in haematology. Academic Press: New York, London 1970; 49–54. Guo R, Lim CK and Peters TJ. Accurate and specific HPLC assay of coproporphyrinogen III oxidase activity in human peripheral leucocytes. Clinica Chimica Acta 1988; 177: 245–252. Poh-Fitzpatrick MB. Plasma porphyrin fluorescence marker for variegate porphyria. Archives of Dermatology 1980; 116: 543–547. Da Silva V, Simonin S, Deybach JC, Puy H and Nordmann Y. Variegate porphyria: Diagnosis value of fluorometric scanning of plasma porphyrins. Clinica Chimica Acta 1995; 238: 163–168. Razzaque MA, Lord GA and Lim CK. Amino acid and peptide conjugates of protoporphyrin: Preparation and analysis by high-performance liquid chromatography, high-performance liquid chromatography/ electrospray ionization mass spectrometry and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Communications in Mass Spectrometry 2002; 16: 1675–1679. Guo R, Lim CK and Peters TJ. High-performance liquid chromatographic assays for protoporphyrinogen oxidase and ferrochelatase in human leucocytes. Journal of Chromatography 1991; 566: 383–396. Li F, Lim CK and Peters TJ. An HPLC assay for protoporphyrinogen oxidase activity in rat liver. Biochemical Journal 1987; 243: 863–866.

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43. Norris PG, Nunn AV, Hawk JL and Cox TM. Genetic heterogeneity in erythropoietic protoporphyria: A study of the enzymatic defect in nine affected families. Journal of Investigative Dermatology 1990; 9(3): 260–263. 44. Todd DJ. Erythropoietic protoporphyria. British Journal of Dermatology 1994; 131(6): 751–766. 45. Goerz G, Bunselmeyer S, Bolsen K and Schürer NY. Ferrochelatase activities in patients with erythropoietic protoporphyria and their families. British Journal of Dermatology 1996; 134(5): 880–885. 46. Li F, Lim CK and Peters TJ. An HPLC assay for rat liver ferrochelatase. Biomedical Chromatography 1987; 2(4): 164–168. 47. Rossi E, Costin KA and Garcia-Webb P. Ferochelatase activity in human lymphocytes, as quantified by a new high-performance liquid chromatographic method. Clinical Chemistry 1988; 34(12): 2481–2485. 48. Day RS, Blekkenhorst GH and Eales L. Co-existent variegate and symptomatic porphyria in S.A. families. South African Medical Research Council News 1978; 10(11): 1–2.

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CHAPTER 5

Isolation and Characterisation of Protoporphyrin Glycoconjugates from Harderian Glands of Rodents by HPLC and HPLC/ESI-MS

5.1. Introduction The Harderian gland is a bi-lobed alveolar gland located around the posterior half of the eyeball cavity. It is named after Johann Jakob Harder who first reported its presence in the stag in 1694. It is found in all mammals and, although vestiges can still be found, is virtually absent in man. The Harderian gland synthesises and stores large amounts of porphyrins,1 especially in some species of rodents, such as rats, mice, rabbits, hamsters and gerbils. The major porphyrin component in rodents was thought to be protoporphyrin.2–4 The presence of coproporphyrin III has also been reported.5 The function of the Harderian gland and the significance of porphyrins remain unknown. In 1970 a porphyrin was isolated from Harderian glands6 and characterised as a tricarboxylic porphyrin believed to be 2-vinyl4,6,7-tripropionic acid porphyrin.7 The presence of a significant amount of this porphyrin (up to 29% of total porphyrin) in the Harderian gland led to the name “harderoporphyrin” being given to the porphyrin.7 Subsequently, the term “harderoporphyria” was used to describe homozygous hereditary coproporphyria where 2-vinyl4,6,7-tripropionic acid porphyrin is significantly elevated (see 155

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Chapter 4, Sec. 4.3.5). These terms are still being widely used, despite the fact that it is now clear that the Harderian gland contains no tricarboxylic porphyrin.8 The “tricarboxylic” porphyrin isolated was in reality an artefact generated from protoporphyrin by the experimental procedures, probably initiated by hydration of a vinyl group of protoporphyrin to give the hydroxyethyl derivative which was then acetylated to give a acetoxyethyl-vinyl structure.8 The dimethyl ester of the acetoxyethylvinyl derivative has the same molecular weight (650 Da) as the trimethyl ester of 2-vinyl-4,6,7-tricarboxylic porphyrin (Fig. 5.1). The tandem ESI-MS/MS fragmentation of the protonated trimethyl ester of 2-vinyl-4,6,7-tricarboxylic porphyrin at m/z 651 gave product ions at m/z 578, 505 and 432, resulted from the successive benzylic cleavage and the loss of one, two and three CH2COOCH3 groups from the side-chain propionic ester substituents, respectively9 (Fig. 5.2). The dimethyl ester of the acetoxyethyl-vinyl derivative would also be expected to give similar loss of three CH2COOCH3 groups, two from the propionic esters and one from the acetoxyethyl group. This similarity in fragmentation probably contributed to the initial inaccuracy in assignment of structure based on EI mass spectrometry. The main component of porphyrins in the Harderian glands of rodents has been shown conclusively to be protoporphyrin-1-O-acyl β-xyloside10 (Fig. 5.3). Protoporphyrin and protoporphyrin-1-O-acyl β-glucoside are minor components.

3

2 H3C

H3C

1

8

NH

CH3 4

N

N

HN

7

5

CH3 CH2CH2COOCH3

NH N CH3

6

H3COOCH2CH2C

CH2CH2COOCH3

1

H 3C N

CH3 C H OCOCH3

HN CH3

H 3C H3COOCH2CH2C

CH2CH2COOCH3

2

Figure 5.1. Chemical structures of 2-vinyl-4,6,7-tripropionic acid trimethyl ester (1) and 2-vinyl-4-acetoxyethyl-6,7-dipropionic acid porphyrin dimethyl ester (2).

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Figure 5.2. (m/z 651).

N

H3C

N

NH

HN

COOH

Protoporphyrin IX

CO HOH2C

O H

H OH

HN

N CH3

H3C

COOH H

H

OH

H OH

Protoporphyrin-1-acyl beta-xyloside

HN CH3

H3C

O

N

CH2OH O H H OH

CO

COOH

O H

H

OH

Protoporphyrin-1-acyl beta-glucoside

Figure 5.3. Chemical structures protoporphyrin IX, protoporphyrin-1-O-acyl β-xyloside and protoporphyrin-1-Oacyl β-glucoside from Harderian glands of the rat.

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CH3

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CH3

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The isolation, identification and characterisation of the glycoconjugates of protoporphyrin were achieved by a combination of HPLC, HPLC/ESI-MS and capillary electrophoresis.

5.2. Extraction of Porphyrins from Harderian Glands Porphyrins in tissues and other biological materials have traditionally been isolated as methyl esters by solvent extraction following esterification with 5–10% methanolic sulphuric acid, or as the free acid porphyrins with 1.5–3.0 M HCl. These procedures are not suitable for the extraction and isolation of protoporphyrin glycoconjugates. The former trans-methylates porphyrin glycoconjugates into methyl ester derivatives, while the latter hydrolyses the glycoconjugates into porphyrin free acids. Porphyrins can be extracted with ethyl acetate-acetic acid without degradation, but the extract is not suitable for injection into the HPLC system before removal of ethyl acetate by evaporation. Porphyrins isolated for the first time from biological materials should be extracted into a neutral solvent mixture to prevent structural modification when the presence of glycoconjugates or other acid-labile groups is expected. Extraction solvents containing methanol or other alcohols must be avoided. Transmethylation of glycoconjugates may take place even under neutral conditions. The following procedure is recommended for the extraction of porphyrins from Harderian glands: 1. Wash freshly dissected Harderian glands briefly in phosphatebuffered saline and cut into small pieces with a scissors or scalpe. 2. Homogenise 250–300 mg in 1 ml of acetonitrile-dimethyl sulphoxide (4:1, v/v) with a motor-driver plunger in a Teflon-glass or glass-glass homogeniser. 3. Centrifuge at 2000 × g for 10 minutes and collect the supernatant. 4. Repeat homogenisation and centrifugation on the pellets until the supernatant is free of fluorescence under UV light.

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5. Pool the supernatants and either use immediately for analysis or store under nitrogen at −20°C for further analysis. 6. Dilute the supernatant with an equal volume of water before injection into the HPLC column. Injection of undiluted supernatant may lead to peak distortion.

5.3. HPLC Separation of Porphyrins from Rat Harderian Gland Extract All reversed-phase columns are suitable for the separation of porphyrins extracted from Harderian glands and various mobile phase mixtures have been investigated. The methanol/ammonium acetate solvent system for the separation of dicarboxylic porphyrins (see Chapter 2) was initially used in an attempt to isolate 2-vinyl-4,6,7tripropionic acid porphyrin as a standard for the study of hereditary coproporphyria. A typical HPLC chromatogram using 88% methanol in 1 M ammonium acetate, pH 5.16 as eluent on a Hypersil-BDS C18 column is shown in Fig. 5.4. The Harderian gland extract was resolved into three peaks. The longest retaining compound (20 min) was identified as a protoporphyrin by comparison of retention time and by co-injection with authentic protoporphyrin standard. The other two peaks (eluted at 15 and 18 min), believed to be tricarboxylic porphyrins, were isolated by peak collection. It was noticed then that these two compounds were unstable in the methanol/ammonium acetate solution, and on re-chromatography gave an additional peak which was more hydrophobic than and eluted after protoporphyrin. Since protoporphyrin was stable and there are no chemical reasons why the tricarboxylic porphyrin should be less stable than protoporphyrin, it was concluded that the compounds were not tricarboxylic porphyrins and HPLC/ESI-MS was used for further investigation. To prevent the possibility of transmethylation, which was suspected, a mobile phase system without methanol as a component was developed. The system, with a mixture of 75% acetonitrile (v/v) in 0.05% trifluroacetic acid (TFA) as the mobile phase, effectively

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Figure 5.4. HPLC separation of porphyrins extracted from Harderian glands of rat. Column: Hypersil-BDS C18 ; eluent: 88% methanol in 1 M ammonium acetate, pH 5.16. Peaks: protoporphyrin, 20.7 min; protoporphyrin glycoconjugates, 15.0 and 18.2 min.

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resolved the porphyrin components in the rat Harderian gland extract (Fig. 5.5). It was then used to analyse a sample exposed to methanol, the chromatogram of which clearly shows the presence of an additional late eluting peak (Fig. 5.6). Porphyrins from Harderian glands were usually identified by HPLC based on their retention times. Since the reported identification of “harderoporphyrin” in 1970,6,7 it was generally assumed that the porphyrins eluting between the tetracarboxylic coproporphyrin and the dicarboxylic protoporphyrin must be tricarboxylic porphyrins. This has led some to assume that the compounds detected in the Harderian glands are the so-called “harderoporphyrin” and isoharderoporphyrin.11 It is obvious from the above results that unequivocal characterisation requires HPLC/MS/MS analysis.

5.4. Identification of Protoporphyrin Glycoconjugates from Rat Harderian Glands by Capillary Electrophoresis and HPLC/ESI-MS The above solvent system was modified in order to achieve better separation in HPLC/ESI-MS analysis. The elution program was isocratic elution with acetonitrile/0.05% TFA (65:35, v/v) for 30 minutes, followed by linear gradient elution from 65 to 90% acetonitrile in 10 minutes for a total run time of 40 minutes. This system also allows column cleaning between runs, thus minimising matrix effect when used in HPLC/ESI-MS analysis. Figure 5.7 shows the reconstructed HPLC/ESI mass chromatograms of porphyrins in the Harderian gland extract. There were three peaks (1, 2 and 3) showing an m/z of 563 corresponding to protonated protoporphyrin. Peak 3 can be positively identified as protoporphyrin by retention time and characteristic tandem MS/MS fragmentation patterns. Peaks 1 and 2 are obviously product ions generated from peaks 6 (m/z 695) and 8 (m/z 725), respectively in the ion source. Tandem MS/MS analysis of the ions at m/z 695 and 725 (Fig. 5.5) confirmed that they were indeed product ions. Peaks 4 and 5 are isomers of peak 6, while peak 7 is an isomer of peak 8. They showed similar fragmentation characteristics.

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Figure 5.5. HPLC separation of porphyrins extracted from Harderian glands of the rat. Column: Hypersil-BDS C18 ; eluent: 75% (v/v) acetonitrile in 0.05% TFA. Peaks: protoporphyrin, 14.4 min; protoporphyrin glycoconjugates, 7.8, 8.6 and 9.6 min.

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Figure 5.6. HPLC separation of porphyrins extracted from Harderian glands of rat showing the generation of a late eluting peak in the presence of methanol. HPLC conditions as in Fig. 5.4. Peaks: protoporphyrin, 14.3 min; late eluting peak, 27.0 min.

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Figure 5.7. HPLC/ESI mass chromatograms of porphyrins from the rat Harderian gland extract. Column: HypersilBDS C18 . The elution program was isocratic elution with acetonitrile/0.05% TFA (65:35, v/v) for 30 minutes, followed by linear gradient elution from 65 to 90% acetonitrile in 10 minutes for a total run time of 40 minutes.

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5.4.1. Characterisation of protoporphyrin xyloside isolated from Harderian glands The mass difference of 132 Da between the precursor (m/z 695) and product (m/z 563) ions of peak 6 indicates conjugation of a pentose to the protoporphyrin molecule. HPLC isolation and acetylation of this compound with anhydrous pyridine/acetic anhydride gave a product with the [M + H]+ ion at m/z 821, an increase of 126 Da in mass. This corresponds to the addition of three acetyl groups and is consistent with a pentose conjugate. The most likely site of conjugation is via one of the propionic acid groups by forming a β-glycosidic 1-O-acyl conjugate. This was confirmed by treating the glycoconjugate with 2% (w/v) methanolic KOH which transmethylates and converts the conjugate into protoporphyrin monomethyl ester. HPLC/ESI-MS analysis of the protoporphyrin monomethyl ester derivative gave the expected [M + H]+ ion at m/z 577 (Fig. 5.8). The transmethylation procedure is as follows: 1. Isolate protoporphyrin monoxyloside by HPLC. 2. Evaporate the purified fraction to dryness. 3. Dissolve the residue in 100 µl of methanolic KOH (2%, w/v) by vortex-mixing. 4. Add 15 µl of 2.7 M HCl, vortex-mix and inject solution for HPLC/ESI-MS identification of protoporphyrin monomethyl ester. The monosaccharide released by the transmethylation reaction can be characterised by capillary electrophoresis (CE) following reaction with 3-methyl-1-phenyl-2-pyrazolin-5-one (MPP). MPP reacts with monosaccharides to form bis-MPP derivatives which are well resolved by CE and can be used to identify the individual sugar species.12 Using this technique the monosaccharide was identified as xylose and the major protoporphyrin glycoconjugate in the Harderian glands of rats is therefore protoporphyrin-1-O-acyl β-xyloside (Fig. 5.3).

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Figure 5.8. HPLC/ESI-MS chromatogram of the protoporphyrin monomethyl ester derivative (m/z 577) formed by trans-esterification of protoporphyrin monoglycoside (m/z 695).

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The procedure for the identification of monosaccharides by CE is as follows: 1. Isolate sufficient quantity of protoporphyrin monoxyloside by HPLC. 2. Evaporate the combined purified fractions to dryness under reduced pressure. 3. Add 50 µl of MPP reagent (0.5 M in methanol) and 50 µl of 0.3 M NaOH to the residue and vortex-mix. 4. Heat the solution at 70°C for 30 minutes in a screw-capped tube. 5. Cool solution down to room temperature and add 50 µl of 0.3 M HCl. 6. Evaporate the solution to dryness under reduced pressure. 7. Dissolve the residue in 200 µl of water. 8. Add 200 µl of ethyl acetate, vortex-mix and centrifuge. 9. Discard the upper ethyl acetate layer and repeat step 8 two more times. 10. Evaporate the final aqueous layer to dryness using a SpeedVac. 11. Dissolve the residue in 200 µl of water and analyse the monosaccharide-MPP derivative by CE. The CE conditions were fused silica capillary: 72 cm total length (53 cm to detector) and 50 µm in inner diameter; running buffer: disodium tetraborate (100 mM, pH 9.5); running temperature: 30°C; running voltage: 20 kV; injection: vacuum, 3 seconds; detection: UV 245 nm. Figure 5.9 shows the capillary electropherograms of the MPP derivatives of a standard mixture of pentose monosaccharides (a) and xylose (b) monosaccharide released from the protoporphyrin monoxyloside in Harderian gland. The bis-MPP-xylose derivative gave an expected [M + H]+ ion at m/z 481 when analysed by HPLC/ESI-MS (Fig. 5.10). The reaction scheme for the characterisation of protoporphyrin1-O-acyl β-xyloside is summarised in Fig. 5.11.

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Figure 5.9. Capillary electropherograms of the bis-3-methyl-1-phenyl-2pyrazolin-5-one derivatives of a standard mixture of pentose monosaccharides (a), and the monosaccharide released from the protoporphyrin monoglycoside (b) in the Harderian gland extract. CE conditions: fused silica capillary, 72 cm total length (53 cm to detector) and 50 µm in inner diameter; running buffer, di-sodium tetraborate (100 mM, pH 9.5); running temperature, 30°C; running voltage, 20 kV; injection, vacuum, 3 seconds; detection, UV 245 nm.

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Figure 5.10. HPLC/ESI-MS chromatograms of the bis-3-methyl-1-phenyl-2-pyrazolin-5-one (MPP) derivative of the monosaccharides released from the protoporphyrin monoglycosides in the Harderian gland extract.

H3C

N

NH

N

N

HN CH3

H3C

HOH2C

H3COOC

HOH2C

H H

H OH

COOH

O

O H

COOH

CH3

OH

Protoporphyrin monoxyloside (m/z 695)

OH

O

MPP

H

H

H

H OH

Bis-MPP xylose derivative

OH

Xylose Capillary electrophoresis

Figure 5.11. Reaction scheme for the characterisation of protoporphyrin-1-O-acyl β-xyloside extracted from the Harderian glands of the rat. MPP = bis-3-methyl-1-phenyl-2-pyrazolin-5-one.

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5.4.2. Characterisation of protoporphyrin monoglucoside isolated from Harderian glands of the rat The same procedures described for the identification of protoporphyrin monoxyloside were used. The capillary electropherograms of MPP derivatives of a standard mixture of hexose monosaccharides and glucose released from protoporphyrin monoglucoside are shown in Figs. 5.12(a) and 5.12(b), respectively. The HPLC/ESI-MS chromatogram of the bis-MPP-glucose derivative with the [M + H]+ ion at m/z 511 is shown in Fig. 5.10.

Figure 5.12. Capillary electropherograms of bis-3-methyl-1-phenyl-2pyrazolin-5-one (MPP) derivatives of (a), a standard mixture of hexose monosaccharides and (b), glucose released from protoporphyrin monoglucoside.

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5.5. Origin and Function of Protoporphyrin Glycoconjugates in Harderian Glands of Rodents The mechanism of formation of protoporphyrin glycoconjugates in the Harderian gland remains to be elucidated. It is probably an enzymatic reaction similar to the formation of bilirubin mono- and di-glucuronides in the hepatic microsomes catalysed by uridine 5’-diphosphoglucuronyltransferase (UDP-glucuronosyltransferase). Whether such an enzyme system exists in the Harderian gland is not known. The principal bilirubin conjugates are mono- and di-glucuronides, with traces of glucose and xylose conjugates also detected in the bile. Conjugation of the lipophilic unconjugated bilirubin resulted in the formation of water soluble conjugated bilirubin. The function of protoporphyrin glycoconjugates in the Harderian gland may be similar — to improve the solubility of protoporphyrin. Excessive accumulation of insoluble protoporphyrin in the liver of patients with erythrohepatic protoporphyria results in gall stones formation. It may also be responsible for liver failure. No protoporphyrin glucuronide, dixyloside or diglucoside were detected in the Harderian gland. The principal porphyrin in the Harderian glands of rodents was protoporphyrin monoxyloside, with a smaller amount of unconjugated protoporphyrin and traces of protoporphyrin monoglucoside. No other porphyrins, including 2-vinyl-4,6,7-tripropionic acid porphyrin, were detectable by HPLC/ESI-MS. In view of these results, the term “harderoporphyrin” should logically be dropped, and, if a name is desired, protoporphyrin-1-O-acyl β-xyloside should be used. The variant form of hereditary coproporphyria (homozygous coproporphyria) in which there is a high faecal excretion of 2-vinyl-4,6,7-tripropionic acid porphyrin also deserves a better name than the current misnomer “harderoporphyria”. Protoporphyrin xyloside and protoporphyrin glucoside in the Harderian glands of rodents are the only naturally occurring glycoconjugates known so far. The extraction and identification of these

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compounds emphasises the importance of using correct experimental procedures for the characterisation of new porphyrins in nature.

References 1. Derrien E and Turchini J. Sur l’accumulation d’une porphyrine dans la glande de Harder des rongeurs du genre Mus et sur son mode d’excretion. Comptes rendus de la Societe de Biologie 1924; 91: 637–639. 2. Thomas J. Action physiologique des porphyrines à l’état libre. Bulletin dela Societe de Chemie Biologique 1938; 20: 878–891. 3. Towbin EJ, Fanta PE and Hodge HC. The porphyrin of Harder’s gland. Proceedings of the Society for Experimental Biology and Medicine 1945; 60: 228–231. 4. Bittner JJ and Waton CJ. The possible association between porphyrins and cancer in mice. Cancer Research 1946; 6: 337–343. 5. Tomio JM and Grinstein M. Porphyrin biosynthesis 5. Biosynthesis of protoporphyrin IX in Harderian glands. European Journal of Biochemistry 1968; 6: 80–83. 6. Kennedy GY. Harderoporphyrin: A new porphyrin from the Harderian glands of the rat. Comparative Biochemistry and Physiology 1970; 36: 21–36. 7. Kennedy GY, Jackson AH, Kenner GW and Suckling CJ. Isolation, structure and synthesis of a tricarboxylic porphyrin from the Harderian glands of the rat. FEBS Letters 1970; 6: 9–12. 8. Gorchein A and Lim CK. Harderoporphyrin: A misnomer. Biomedical Chromatography 2005; 19(8): 565–569. 9. Danton M and Lim CK. Identification of monovinyl tripropionic acid porphyrins and metabolites from faeces of patients with hereditary coproporphyria by high-performance liquid chromatography/electrospray ionization quadrupole time-of-flight tandem mass spectrometry. Rapid Communications in Mass Spectrometry 2004; 18: 2309–2316. 10. Lim CK, Razzaque MA, Luo J and Farmer PB. Isolation and characterization of protoporphyrin glycoconjugates from rat Harderian gland by HPLC, capillary electrophoresis and HPLC/electrospray ionization MS. Biochemical Journal 2000; 347: 757–761.

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11. Ng JC, Oi L and Moore MR. HPLC measurement of harderoporphyrin in the Harderian glands of rodents as a biomarker for sub-lethal or chronic arsenic exposure. Toxicology Letters 2002; 133(1): 93–101. 12. Honda S. Monosaccharide analysis by capillary electrophoresis. In: Capillary Electrophoresis of Carbohydrates, Thibault P and Honda S (eds). Humana Press: Totowa, New Jersey, 2003; 81–92.

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CHAPTER 6

HPLC and HPLC/MS of Chlorophyll and Related Compounds

6.1. Introduction Chlorophyll and related compounds are significantly less polar or more hydrophobic than the porphyrin molecules discussed in the previous chapters. In general, they are soluble in common organic solvents such as acetonitrile, ethanol and methanol, but the very apolar, fully esterified derivatives may require ethyl acetate, dichloromethane, hexane or other hydrocarbon solvents for solution. They can be separated by normal-phase HPLC on a silica column like the esterified porphyrins, or by reversed-phase HPLC using a high proportion of organic modifier or even totally non-aqueous solvents for elution. The peaks are either detected and identified with a UV-Visible or fluorescence detector, or characterised by MS and tandem MS/MS analysis. The structures of chlorophyll a (Chl a), chlorophyll b (Chl b), their C-132 epimers (Chl a’ and Chl b’, respectively) and some of the biosynthetic intermediates and degradation products which have been separated by HPLC are shown in Fig. 6.1.

6.2. Reversed-Phase HPLC of Chlorophyll and Related Compounds in Plant Extracts A reversed-phase HPLC system using mobile phases similar to those developed for the separation of porphyrins (see Chapter 2) was used to separate chlorophyll-related compounds in plant extracts.1 177

N

N

N H CH3

H3C H

N

N

18

CH3

H3C H

N

H

D

10

N 14

C

11 12

E

132

O

CH3OOC

CH3

H3C 17 1615 E 13 H 132 131 O H O 173 COOCH3

O COOCH3

HOOC

Mg

20

8

B N 9

H

C-132 epimer

O Protochlorophyllide

Chlorophyllide Chlorophyll 3 4 H3C 2 A NH 1

H3C N

N Mg

N

CH3

H3C O

173

H

N

H

O COOCH3

O

6

7

R

8

B N 9

3

H3C 2 A

10

20 19

18

5

D

N

HN 14

1

11

C 12 CH 3

H3C 17 1615 E 13 H 132 131 O H O COOCH 3 173 O

4

NH

5

6

8

10

20

H 19

7

B N 9

N

HN

11 12

D 14 C 18 CH3 H3C 17 1615 E 13 2 H 13 131 O H O COOCH3 HO Pheophobide

Protochlorophyll

Pheophytin

Figure 6.1. Chemical structures of chlorophyll a (R = CH3), chlorophyll b (R = CHO) and related compounds. Chlorophyll a’ and b’ are C-132 epimers of chlorophyll a and b, respectively.

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R

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The method was applied to the analysis of degradation products of chlorophyll during the industrial processing of fruits and vegetables. Depending on the methods and conditions of processing chlorophyll undergoes various degradation reactions, which include demetalation, epimerisation, dephytylation, demethoxycarbonylation and allomerisation. Thus a complex mixture of compounds is formed and an analytical method capable of separating and identifying these compounds is needed. The compounds were separated on a Lichrosphere 100-RP18 column (5 µm partical size, 250 mm × 4.6 mm) by gradient elution. Solvent A was 1 M ammonium acetate/methanol (1:4, v/v) and solvent B was acetone/methanol (1:4, v/v). The linear gradient was run at a flow rate of 1 ml/min from 100% A to 100% B in 15 min and held at 100% B for a further 30 min or 50 min for the elution of highly hydrophobic components if necessary. Table 6.1 shows the retention times of the compounds separated by this system. Chl b and Chl b’ eluted before Chl a and Chl a’, as expected for reversed-phase HPLC. Replacing the CH3 group with a CHO group at position 7 increases the polarity of the b chlorophylls. Chl a eluted before Chl a’ because the -COOCH3 substituent at the C-132 position is not on the same plane of the C-17 phytyl group and is therefore less hindered and thus more polar than Chl a’. Similarly, Chl b eluted before Chl b’. The chlorophyllides, with an unesterified propionic acid group at C-17, are much less hydrophobic than the chlorophylls. They were the first group to elute. Demetallation increases the hydrophobicity of chlorophyllrelated compounds. The pheophobides were therefore more strongly retained than the chlorophyllides. Pheophytins, the demetallated chlorophylls, were the longest retaining compounds. Non-aqueous reversed-phase HPLC using acetone (A), acetonitrile (B) and methanol (C) as gradient solvent mixture2 have also been developed for the separation of chlorophylls and derivatives in plant extract. The separation of chlorophylls and related compounds in Gynostemma pentaphyllum Makino was carried out on a HyPurity C18 column (5 µm particle size, 150 mm × 4.6 mm i.d., from Thermo

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Table 6.1. Retention Time of Chlorophylls Separated by Reversed-Phase HPLC.1 Compound

Retention time (min)

Chlorophyllide b Chlorophyllide a Chlorophyllide a’ Protochlorophyllide a Pheophobide b Pheophobide b’ Pheophobide a Pyropheophorbide b Pheophobide a’ Pyropheophorbide a Hydroxychlorophyll b Chlorophyll b Chlorophyll b’ Hydroxychlorophyll a Methoxychlorophyll a Methoxylactone chlorophyll a Chlorophyll a Chlorophyll a’ Pheophytin b Pheophytin b’ Pheophytin a Pheophytin a’

6.7 13.6 14.3 17.0 18.1 18.7 20.4 20.4 21.1 22.8 25.5 26.6 27.1 27.6 28.2 28.7 29.3 30.1 39.6 41.5 45.6 48.9

Column: Lichrosphere 100-RP-18; solvent A, 1 M NH4Ac/CH3OH (1:4, v/v); B, acetone/CH3OH (1:4, v/v); elution: 100% A to 100% B in 15 min; held at 100% B.

Hypersil-Keystone). The elution program was from 2% A, 93% B and 5% C to 2% A, 71% B and 27% C in 3 min; 2% A, 64% B and 34% C in 6 min; 2% A, 45% B and 53% C in 9 min; 2% A, 39% B and 59% C in 21 min; 2% A, 24% B and 74% C in 24 min; 20% A, 0% B and 80% C in 26 min; 40% A, 0% B and 60% C in 28 min; 50% A, 0% B and 50% C in 30 min and returned to 2% A, 93% B and 5% C in 35 min. The mobile phase flow rate was 1 ml/min and detection was

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Table 6.2. Retention Time and Capacity Ratio of Chlorophylls2 Separated by Non-Aqueous Reversed-Phase HPLC. Compound

Retention time (min)

Capacity ratio (K′′)

Hydroxychlorophyll b Chlorophyll b Chlorophyll b’ Hydroxychlorophyll a Chlorophyll a Chlorophyll a’ Hydroxypheophytin b Hydroxypheophytin b’ Pheophytin b Pheophytin b’ Hydroxypheophytin a Hydroxypheophytin a’ Pheophytin a Pheophytin a’ Pyrophephytin a

7.17 8.19 8.80 10.21 11.66 12.74 14.82 15.82 17.00 18.71 21.64 23.68 25.80 28.11 31.19

2.83 3.38 3.71 4.46 5.24 5.81 6.93 7.46 8.09 9.00 10.57 11.66 12.80 14.03 15.68

Column: HyPurity C18 (Thermo Hypersil-Keystone); Solvents and elution: gradient elution with acetone (A), acetonitrile (B) and methanol (C).

at 660 nm. The retention times and capacity ratios are shown in Table 6.2.

6.3. Normal-Phase HPLC of Chlorophyll and Related Compounds The separation of Chl a, Chl a’, pheophytin a (Pheo a) and protochlorophyll (PChl) with the 173 carbons esterified with geranylgeranyol (GG), dihydrogeranylgeranyol (DHGG), tetrahydrogeranylgeranyol (THGG) and phytol (Pyt) groups3 has been achieved on a silica column (150 mm × 6 mm Senshupak Silica 2151-N) with hexane/ toluene/methanol (100/4/0.8, by volume) as mobile phase at a flow rate of 1 ml/min. The column temperature was maintained at 4°C. The elution of the four groups of compounds was in the order of pheophytin, protochlorophyll, chlorophyll a’ and chlorophyll a. Within each group, the retention time increased with increasing

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PheophytinGG

R=

CH2O CH3

PheophytinDHGG

R=

CH2O

3 4 H3C 2 A NH 1

5

6

18

PheophytinTHGG

R=

CH2O

H3C 17 O

173

8

10

20

H 19

7

B N 9

N

HN

11 12

14 C 16 15 E 13 2 H 13 131 O H

D

CH3

COOCH3

R Pheophytin a

R=

Figure 6.2.

CH2O

Pheophytin a and related C-173 side-chain derivatives.

number of double bonds in the C173 isoprenoid side-chain. Thus, an elution in the order of Pheo a, Pheo aTHGG , Pheo aDHGG and Pheo aGG (Fig. 6.2) was observed. The elution order was reversed when these compounds were separated by reversed-phase HPLC on a µBondapak C18 column (300 mm × 4 mm, 10 µm particle size) with methano/acetone (90:10, v/v) as mobile phase.4 The protochlorophyll, chlorophyll a’ and chlorophyll a groups of compounds behaved similarly. The retention times and capacity ratios of the compounds separated are listed in Table 6.3. The method was applied to the analysis of possible biosynthetic intermediates of pheophytin a and chlorophyll a’ in greening etiolated barley leaves.3 Normal-phase HPLC has also been used for the separation of chlorophyll a allomerisation products in “wet” methanol.5,6 The separation was carried out on a 250 mm × 4.6 mm silica column (Spherisorb S5 W). The mobile phase was hexane (solvent A) and 2-propanol/methanol (1:1, v/v, solvent B). The elution program was as follows: 0 to 20 min, isocratic elution at 1.5% B; 20 to 35 min, linear gradient elution from 1.5% B to 10% B; 35 to 40 min, back to 1.5% B. The flow rate was 2 ml/min and detection was at 425 nm. Seven compounds have been identified in the allomerisation mixture. They were Mg-purpurin-7 dimethyl phytyl ester and the (R)- and

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Table 6.3. Retention Time and Capacity Ratio of Chlorophylls3 Separated by Normal-Phase HPLC. Compound

Retention time (min)

Capacity ratio (k′′)

Pheophytin a Pheophytin aTHGG Pheophytin aDHGG Pheophytin aGG Protochlorophyll ProtochlorophyllTHGG ProtochlorophyllDHGG ProtochlorophyllGG Chlorophyll a’ Chlorophyll a’THGG Chlorophyll a’DHGG Chlorophyll a’GG Chlorophyll a Chlorophyll aTHGG Chlorophyll aDHGG Chlorophyll aGG

11.3 11.9 12.7 13.3 17.1 18.0 19.1 20.2 23.1 24.6 26.4 27.9 34.2 36.4 39.3 41.5

1.82 1.97 2.17 2.33 3.27 3.50 3.77 4.06 4.78 5.15 5.61 5.97 7.55 8.09 8.84 9.37

Column: Senshupak Silica 2151-N (Senshu Science, Japan); eluent: hexane/toluene/methanol (100:4:0.8, by vol); column temperature: 4°C.

(S )-epimers of 132-hydroxy Chl a, 132-methoxy Chl a and 151methoxylactone Chl a (Fig. 6.3). The elution order was Mg -purpurin-7 dimethyl phytyl ester, 2 13 (S )-hydroxy Chl a’, 151(S )-methoxylactone Chl a’, 151(R)methoxylactone Chl a, 132(R)-methoxy Chl a’ + 132(R)-hydroxy Chl a, and 132(S )-methoxy Chl a.

6.4. HPLC Separation of Fluorescent and Non-Fluorescent Chlorophyll Catabolites Reversed-phase HPLC systems have been developed for the analysis and preparative isolation of chlorophyll catabolites.7–10 Analytical separation was typically performed on a C18 column, e.g., HypersilODS by gradient elution. The solvents were (A), 50 mM potassium phosphate buffer, pH 7.0 and (B), 25 mM potassium phosphate

11 12

16

1

N

H 19

N

10

14 15

Mg

20

CH3

18

13

H3C 17

COOCH3 COOCH3

1

8

11 12

14 C 16 15 E 13 H 132 131 O R

D

132(S)-Hydroxychlorophyll a; R1 = OH, R2 = COOCH3

10

N

1

O

7

B N 9

132(R)-Hydroxychlorophyll a; R1 = COOCH3, R2 = OH CH3

132(S)-Methoxychlorophyll a; R1 = OCH3, R2 = COOCH3 132(R)-Methoxychlorophyll a; R1 = COOCH3, R2 = OCH3

R2 O Phytyl

H3C 2

3 4 5 6

N

1

N

H 19 H3C

N Mg

20 18

17

16

7

8

9

151(S)-Methoxylactone chlorophyll a; R1 = OCH3, R2 = COOCH3

10

N 14 15

H R1

O

CH3

13 151

O R2

11 12

CH3

151(R)-Methoxylactone chlorophyll a; R1 = COOCH3, R2 = OCH3

O

O Phytyl

Figure 6.3.

Chemical structures of chlorophyll a allomers. 1 = Mg-purpurin-7 dimethyl phytyl ester.

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Mg

H O

O

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H3C 17

H3C 2 A

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4

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N

20

CH3 3

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1

N

184

H3C

2

CH3

3 4 5 6

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buffer, pH 7.0/methanol (20:80, v/v). The elution program was: 0 to 60 minutes, linear gradient from 25% B to 75% B; 60 to 65 minutes, 75% B to 100% B; 65 to 70 minutes, isocratic elution at 100% B. The flow rate was 0.5 ml/min. The peaks were detected by sequential monitoring with a photodiode array detector (200–600 nm) and a fluorescence detector (excitation 320 nm, emission 450 nm).

6.5. HPLC/MS and MS/MS of Chlorophyll and Related Compounds For HPLC/MS and tandem MS/MS operations, normal phase, nonaqueous reversed-phase system and reversed-phase HPLC solvent systems are all suitable. Atmospheric pressure chemical ionisation (APCI) is the preferred ionisation technique for HPLC/MS and MS/MS analysis of the relatively hydrophobic chlorophyll-related compounds. A ternary mobile phase system consisted of 0.1 M ammonium acetate (A), methanol (B) and acetone (C) was used to identify chlorophylls in plant extracts.1 The column used was Lichrosorb RP-18. The elution program was: 0 to 15 min, linear gradient from 20% A, 80% B and 0% C to 0% A, 80% B and 20% C; 15 to 50 min, isocratic elution at 0% A, 80% B and 20% C; 50 to 60 min, isocratic elution at 0% A, 45% B and 55% C. The precursor [M + H]+ and main product ions of the compounds analysed are shown in Table 6.4. The mechanism of fragmentation and the characteristic fragmentation pattern of chlorophylls have been studied. The positive ion mass spectra are characterised by the presence of an intense protonated molecule. Thus Chl a and Chl a’ both gave an [M + H]+ ion at m/z 893. Upon MS/MS, a major fragment ion at m/z 615 (893 − 279 + 1), corresponding to the loss of the phytyl chain with simultaneous back transfer of a proton from the leaving group to the carboxylic oxygen of the product ion was observed.11 Another characteristic fragment is the elimination of a CH3COOC20H39 group, presumably from a protonated phytyl ester side-chain, to give the ion at m/z 555.

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Table 6.4. Atmospheric Pressure Chemical Ionization MS and MS/MS of Chlorophylls. Compound

[M + H]+

Chlorophyll a Chlorophyll b Hydroxychlorophyll a Methoxychlorophyll a Methoxylactone chlorophyll a Pheophorbide a Pyropheophorbide a Pyropheophorbide b Pheophytin a Pheophytin b Hydroxy Pheophytin a Pyropheophytin a Pyropheophytin b

893 907 909 923 939 593 549 535 871 885 887 813 827

Main fragment ions 615, 879, 891, 891, 907, 575, 527, 513, 593, 857, 869, 535 799,

555 629, 631, 645, 879, 535, 501 487 533 607, 609,

596 613 613 852, 821, 661, 629 517

547 591

549

Although Chl a, and Chl a’ gave essentially identical fragmentation patterns, the ease of fragmentation and therefore the relative abundance of the fragment ions are different under identical conditions. Chl a’ is more easily fragmented and gave a more intense ion at m/z 615 than Chl a. This is attributed to the greater steric strain resulting from the C-132 COOCH3 and the C-17 phytyl ester groups lying on the same plane of the macrocyclic ring. The above characteristic fragmentation feature was also observed for Chl b and Chl b’ (m/z 907) which were further characterised by the loss of CO from the CHO group on the C-7 position to give an ion at m/z 879 (907 − 28). The pheophytins have similar MS and MS/MS characteristics to the chlorophylls. Pheophytin a and pheophytin a’ gave the [M + H]+ ion at m/z 871 and characteristic product ions at m/z 593 (871 − 278) and 533 (871 − 338), corresponding to the loss of a phytadiene group and the elimination of a CH3COOC20H39 group, respectively. As with the chlorophylls, the more strained pheophytin a’ is more easily fragmented and gave a more intense product ion at m/z 593

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than pheophytin a. This was similarly observed for pheophytins b and b’. APCI-MS and MS/MS studies5,6 of chlorophyll a allomers have shown that facile fragmentations involve the methoxy and hydroxy groups at the C-132 or C-151 chiral centres and the losses of C-132 or C-151 hydroxy or methoxy substituents occur more easily from the (S )-epimer than from the (R)-epimer. This is resulting from the greater steric strain and repulsion between the substituents and the C-17 phytyl ester group on the same plane of the macrocyclic ring. For examples, enhanced loss of the C-132 substituent by 132(S )hydroxy Chl a as water and 132(S )-methoxy Chl a as CH3OH, respectively, was observed. Similarly, 151(S )-methoxylactone Chl a showed more facile loss of CH3OH than its (R )-epimer. This characteristic can be used to differentiate and assign the stereo configuration of the epimers. The most important MS/MS fragment ions of chlorophyll a allomers and their relative intensities are shown in Table 6.5. APCI-MS and MSn have also been applied to the study of products of allomerisation of bacteriochlorophyll a, and bacterioviridin a5,6 and the structural assignment of sedimentary bacteriochlorophyll derivatives.12

Table 6.5.

Most Important Product Ions in MS2 for Chlorophyll a Allomers.4,5

Compound

[M + H]+

Productions

132(R )-Hydroxychlorophyll a 132(S )-Hydroxychlorophyll a 132(R )-Methoxychlorophyll a 132(S )-Methoxychlorophyll a 151(R )-Methoxylactonechlorophyll a 151(S )-Methoxylactonechlorophyll a Mg-Purpurin-7 dimethyl phytyl ester

909 909 923 923 939

819(18), 631(100), 613(74) 819(30), 631(52), 613(100) 891(26), 645(100), 613(79) 891(32), 645(27), 613(100) 907(21), 879(6), 852(24), 821(6), 661(100), 629(29) 907(40), 879(26), 852(28), 821(12), 661(100), 629(70) 911(3), 907(20), 879(3), 852(38), 661(100), 660(64)

939 939

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References 1. Gauthier-Jaques A, Bortlik K, Hau J and Fay LB. Improved method to track chlorophyll degradation. Journal of Agricultural and Food Chemistry 2001; 49: 1117–1122. 2. Huang SC, Hung CF, Wu WB and Chen BH. Determination of chlorophylls and their derivatives in Gynostemma penta phylum Makino by liquid chromatography-mass spectrometry. Journal of Pharmaceutical and Biomedical Analysis 2008; 48(1): 105–112. 3. Nakamura A, Tanaka S and Watanabe T. Normal-phase HPLC separation of possible biosynthetic intermediates of pheophytin a and chlorophyll a. Analytical Sciences 2001; 17: 509–513. 4. Schoch S, Lempert U, Wieschhoff H and Scheer H. High-performance liquid chromatography of tetrapyrrole pigments. Journal of Chromatography 1978; 157: 357–364. 5. Jie C, Walker JS and Keely BJ. Atmospheric pressure chemical ionization normal phase liquid chromatography mass spectrometry and tandem mass spectrometry of chlorophyll a allomers. Rapid Communications in Mass Spectrometry 2002; 16: 473–479. 6. Walker JS, Jie C and Keely BJ. Identification of diastereomeric chlorophyll allomers by atmospheric pressure chemical ionization liquid chromatography/tandem mass spectrometry. Rapid Communications in Mass Spectrometry 2003; 17: 1125–1131. 7. Ginsburg S and Matile P. Identification of catabolites of chlorophyll-porphyrin in senescent rape cotyledons. Plant Physiology 1993; 102: 521–527. 8. Oberhuber M, Berghold, J, Breuker K, Hörtensteiner S and Kräutler B. Breakdown of chlorophyll: A nonenzymatic reaction accounts for the formation of the colorless “nonfluorescent” chlorophyll catabolites. Proceedings of the National Academy of Sciences USA 2003; 100: 6910–6915. 9. Pruzinská A, Tanner G, Aubry S, Anders I, Moser S, Müller T, Ongania K-H, Kräutler B, Youn, J-Y, Liljegren, SJ and Hörtensteiner S. Chlorophyll breakdown in senescent Arabidopsis leaves. Characterization of chlorophyll catabolites and of chlorophyll catabolic enzymes involved in the degreening reaction. Plant Physiology 2005; 139: 52–63.

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10. Kräutler B. Chlorophyll breakdown and chlorophyll catabolites in leaves and fruit. Photochemical & Photobiological Sciences 2008; 7(10): 1114–1120. 11. Van Breemen RB, Canjura FL and Schwartz SJ. Identification of chlorophyll derivatives by mass spectrometry. Journal of Agricultural and Food Chemistry 1991; 39: 1452–1456. 12. Wilson MA, Hodgson DA and Keely BJ. Atmospheric pressure chemical ionization liquid chromatography/multistage mass spectrometry for assignment of sedimentary bacteriochlorophyll. Rapid Communications in Mass Spectrometry 2005; 19: 38–46.

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CHAPTER 7

HPLC and HPLC/MS of Bilins of Animal and Plant Origin

7.1. Introduction HPLC was used for the separation of synthetic bile pigments since the early 1970s. The dimethyl ester derivatives of the pigments were separated on silica columns with mobile phases similar to those used in thin-layer chromatography,1 typically a binary or ternary mixture consisting of methyl acetate, dichloromethane and n-heptane or isooctane. Normal phase HPLC remains a useful technique for the analytical separation and preparative isolation of bile pigments in the chemical laboratory. Reversed-phase HPLC is suitable for the separation of free acid bile pigments2 as well as their dimethyl ester derivatives. The technique is well suited to the analysis of bile pigments from biological extracts2 from both animal and vegetable sources.

α, IXα α 7.2. HPLC Separation of Bilirubin IIIα α Isomers and XIIIα The rubins are yellow compounds with absorption maximum at around 450 nm. Because of extensive intramolecular hydrogen bonding free rubins are relatively hydrophobic and are insoluble in aqueous solution at neutral pH and methanol. They are soluble in alkaline solution as the sodium salt and in organic solvents such as chloroform, dichloromethane and dimethyl sulphoxide (DMSO). The dimethyl esters of rubins are readily soluble in polar organic solvents. 191

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Bilirubin IIIα and XIIIα isomers are easily formed in small quantity in vitro by fission of the central methene bridge followed by recombination of the dipyrroles, especially in the presence of strong acids.3 Commercially available bilirubin IXα contains varying amount of IIIα and XIIIα isomers as impurities (Fig. 7.1). Bilirubin isomers are easily separated by reversed-phase HPLC. The choice of mobile phase depends on the application required. In preparative purification and isolation of a pure isomer, a C18 or ODS column with a mixture of acetonitrile, dimethyl sulphoxide (DMSO) and 0.1 to 0.5 M ammonium acetate (pH 4.6–5.2) as eluent is recommended.4 Figure 7.2 shows the separation of bilirubin IIIα, IXα and XIIIα isomers on a Hypersil-ODS column with a mixture of 0.1 M ammonium acetate (pH 4.6), acetonitrile and DMSO (40:60:60, v/v/v) as mobile phase at a flow rate of 1 ml/min. COOH COOH CH3

H3C O

N H

CH3

N H

CH3

N H

O

N H

1 COOH COOH CH3

H3C O

N H

CH3

N H

CH3

N H

N H

O

2

COOH COOH CH3 CH3 O

N H

CH3

N H

N H

CH3

N H

O

3

Figure 7.1.

Chemical structure of bilirubin IIIα (1), IXα (2) and XIIIα (3).

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Figure 7.2. HPLC separation of bilirubin IIIα, IXα and XIIIα isomers. Column: Hypersil-ODS; eluent: acetonitrile/DMSO/0.1 M ammonium acetate, pH 4.6 (40:60:60, v/v/v).

The presence of DMSO improves the solubility and chromatography of bilirubins. The system is also suitable for the separation of bilirubin methyl esters. The simultaneous separation of bilirubin, bilirubin monomethyl ester and bilirubin dimethyl ester (Fig. 7.3) was achieved by modification of the above mobile phase system to 0.5 M ammonium acetate (pH 4.6)/acetonitrile/DMSO (40:50:50, v/v/v). The retention of these compounds can be precisely controlled by manipulation of buffer concentration, pH and/or organic modifier proportion in the mobile phase.4 Mobile phase pH provides the best parameters for retention control. The retention of bilirubin and bilirubin monomethyl ester decreases with increasing pH, with the former more significantly affected than the latter. Conversely, the retention of bilirubin dimethyl ester increases with increasing pH. This can be explained by the fact that at higher pH the two propionic acid groups of bilirubin are more likely to be ionised with a consequent decrease in hydrophobicity. Without a free acid group, the ionisation state of the pyrrole nitrogens becomes a main factor in bilirubin dimethyl

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Figure 7.3. HPLC separation of bilirubin, bilirubin monomethyl ester, bilirubin dimethyl ester, and their isomers. Column: Hypersil-ODS; eluent: 0.5 M ammonium acetate, pH 4.6/acetonitrile/DMSO (40:50:50, v/v/v). Peaks: 1 = bilirubin XIIIα monomethyl ester; 2 = bilirubin IXα monomethyl ester; 3 = bilirubin IIIα monomethyl ester; 4 = bilirubin XIIIα; 5 = bilirubin IXα; 6 = bilirubin IIIα; 7 = bilirubin XIIIα dimethyl ester; 8 = bilirubin IXα dimethyl ester; 9 = bilirubin IIIα dimethyl ester.

ester. Increasing the pH suppresses nitrogen ionisation, thus increasing the hydrophobicity and, therefore the retention. At pH above 5, the elution order of bilirubin monomethyl ester, bilirubin and bilirubin dimethyl ester observed in Fig. 7.3 was changed to bilirubin, bilirubin monomethyl ester and bilirubin dimethyl ester with a reversal of elution order between bilirubin and bilirubin monomethyl ester. In analytical separation where solubility is less of a problem the separation can be carried out with binary solvent mixture such as acetonitrile and 0.1–0.5 M ammonium acetate at near neutral pH without DMSO (see below).

7.3. HPLC Separation of Bilirubin Conjugates Bilirubin mono- and di-glucuronide conjugates (Fig. 7.4) can be separated by reversed-phase HPLC with 0.1 M ammonium acetate

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OO

195

CH3

H3C NH HN NH HN H3C HOOC HO

O

HO

OH

O

CH3

8

12 C O

C O O

HO

OH

Figure 7.4. conjugates.

NH HN CH3

12 C

CH3

NH HN

NH HN

O

OH OH

H3C

NH HN

O

HO

OO

H3C

HOOC HO

COOH

CH3

OO

H3C

O

COOH O

H3C

8

CH3 O

HOOC

O

C

O

HO

COOH OH OH

Chemical structures of bilirubin mono- and di-glucuronide

buffer adjusted to pH 5.16-acetonitrile-DMSO (85:50:50, v/v/v) as mobile phase4 (Fig. 7.5). The retention and resolution are easily controlled to suit a particular application by adjusting the pH and/or buffer concentration of the mobile phase. Buffer concentrations of between 0.1–0.25 M at pH values of 5.0–6.8 are most suitable.

7.4. HPLC of Biliverdins Verdins are green-blue compounds with absorption maximum at around 666 nm. The dimethyl esters have the absorption maximum in the range of 630–662 nm, depending on the solvent used. The verdins are chemically the most stable bile pigments. Free verdins are soluble in both aqueous acid and alkaline solutions and in methanol

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Figure 7.5. HPLC separation of mono- and di-glucuronide. Column: Hypersil-ODS; eluent: 0.1 M ammonium acetate, pH 5.16/acetonitrile/ DMSO (85:50:50, v/v/v). Peaks: 1 = bilirubin diglucuronide; 2 = bilirubin C-8 monoglucuronide; 3 = bilirubin C-12 monoglucuronide.

but not chloroform or dichloromethane. The methyl esters are soluble in chloroform and hot methanol. The enzymatic oxidative cleavage of haem gives biliverdin IXα as the major product accompanied by the IXβ, IXγ and IXδ isomers (Fig. 7.6) as minor products. The four isomers can also be generated by non-enzymatic coupled oxidation of haemin with reducing agents. In such a reaction, for example by coupled oxidation of haemin with oxygen and ascorbate in aqueous pyridine, cleavage of all four meso-positions of haemin takes place with formation of the four isomers.5,6 The dimethyl esters of the four biliverdin isomers have been separated by reversed-phase HPLC on a Zorbax ODS column (150 × 4.6 mm) maintained at 44°C with an equal volume of acetonitrile, methanol and 0.01 M sodium acetate buffer, pH 3.65 as mobile phase7 at a flow rate of 1 ml/min. The isomers were eluted in the order of IXα, IXβ, IXδ and IXγ.

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CH3

OO

CH3

H3C

H3C

NH HN

N

HN

N

CH3

H 3C

HOOC

197

COOH

HN

O O

NH HN CH3

H3C

HOOC

COOH

1

2

CH3

CH3 H3C

H3C N

NH

HN

O O

NH HN CH3

H 3C

N

NH HN CH3

H3C

OO HOOC

COOH

3

COOH

HOOC

4

Figure 7.6. Chemical structure of biliverdin IXα (1), bilirubin IXβ (2), bilirubin IXγ (3), and bilirubin IXδ (4).

Normal phase HPLC on a Zorbax Sil-850 column (250 × 4.6 mm) with dichloromethane/methanol/water (99.0:0.9:0.1, v/v/v) as mobile phase at a flow rate of 1.5 ml/min was also successfully used to separate the dimethyl esters of biliverdin isomers.8 An elution order of IXβ, IXα, IXγ and IXδ was observed. Reversed-phase ion-pair HPLC with tetrabutylammonium hydroxide as ion-pairing agent was developed for the separation of the four free acid biliverdin isomers.9 The column used was LiChrosorb 5 RP-8 (150 × 4.6 mm). The mobile phase consisted of a mixture of acetonitrile, ethyl acetate, methanol and water containing 10 mM of the ion-pairing agent at pH 4.0. The elution order of the

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isomers was IXα, IXδ, IXβ and IXγ. Hexafluoroacetone and pyridine have also been used as ion-pairing agents for the reversed-phase ion-pair HPLC of biliverdin isomers.10 The isomers were also separated by reversed-phase HPLC on a µBondapak C18 column (300 mm × 4 mm; 10 µm particle size) with methanol/water (78:22, v/v) as eluent.11 The elution order was IXα, IXβ, IXδ and IXγ. The IIIα, IXα and XIIIα isomers of biliverdin could be separated by reversed-phase HPLC with a mixture of acetonitrile and ammonium acetate as mobile phase (see Sec. 7.8 below). This system should therefore also be suitable for the resolution of the IXα, IXβ, IXδ and IXγ isomers.

7.5. HPLC of Biliviolins Violins are violet compounds soluble in most organic solvents with an absorption maximum at around 564–587 nm. The columns and mobile phases used for the separation of bilirubin and derivatives are also applicable to the separation of the violins. The dimethyl esters of biliviolins can be separated by normal phase HPLC on silica with a mixture of methyl acetate and isooctane as mobile phase. Table 7.1 shows the retention time synthetic biliviolin dimethyl esters (Fig. 7.7) separated on a Hypersil (silica) column with methyl acetate/isooctane (135:115, v/v) as mobile phase. For the separation of free acid violins reversed-phase HPLC with acetonitrile/ammonium acetate buffer as mobile phase is recommended.

7.6. HPLC of Urobilinoids Urobilinoids is the name given to a group of orange colour bile pigments which include urobilin with two unsaturated pyrrolinone end-rings (dipyrrolinone), stercobilin with two saturated pyrrolidone end-rings (dipyrrolidone) and half-stercobilin with one saturated pyrrolidone and one unsaturated pyrrolinone end ring (pyrrolidone-pyrrolinone).

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H3COOC

COOCH3

CH3 CH3 O

N H

N H

H3C O

COOCH3

CH3

CH3 N H

3 COOCH3

CH3

CH3

N H

N H

N 5

COOCH3

CH3

CH3

N 7

N H

O

O

O

CH3

H3COOC

N H

H3C

CH3

N

H3COOC

H3C O

1

N H

N H

H

N H

N

H3COOC

N H

H3C O

CH3 CH3

N H

O

O

COOCH3

CH3

CH3

CH3

N

COOCH3

CH3

CH3

N H

CH3 N H

N 4

H3COOC

COOCH3

CH3

CH3

N H

N 6

O

N H

2

H3COOC

N H

H3C O

H3COOC

N H

H 3C

199

O

CH3 N H

N H

O

CH3 N H

O

Figure 7.7. Chemical structure dimethyl ester of mesobiliviolin IIIα (1), mesobiliviolin IXα (2), mesobiliviolin XIIIα (3), A-dihydrobiliviolin IXα (4), D-dihydrobiliviolin IXα (5), A-dihydroisobiliviolin IXα (6), and ethylideneisomesobiliviolin IXα (7).

Table 7.1.

Retention Time and Capacity Ratio of Biliviolin Dimethyl Esters.

Compound

Retention time (min)

Capacity ratio (k′)

Ethylidene isomesobiliviolin IXα A-Dihydrobiliviolin IXα Mesobiliviolin IIIα A-Dihydrobiliviolin IXα Mesobiliviolin IXα D-Dihydrobiliviolin IXα Mesobiliviolin XIIIα

6.2 7.4 9.2 10.5 10.9 11.6 12.7

0.87 1.24 1.78 2.18 2.30 2.51 2.84

Column: Hypersil; eluent: methyl acetate/isooctane (135:115, v/v); flow rate: 1.5 ml/min.

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The synthetic urobilinoids is a mixture of RR + SS and RS + SR diastereoisomers, while the natural i-urobilin is a racemate mixture of two enantiomers, RR and SS. They have been resolved into D- and L-enantiomers. D-urobilin and L-stercobilin occur naturally and have also been isolated from faeces. The free base urobilins are soluble in methanol with an absorption maximum of 450 nm. The HCl salt is soluble in chloroform or dichloromethane with an absorption maximum at 490 nm. The dimethyl esters are soluble in both chloroform and methanol. The dimethyl esters of urobilins (Fig. 7.8) were effectively separated by normal phase HPLC on silica.12 The compounds separated were (SS )-stercobilin, (RR,SS )-half-stercobilin, (RR,SS )-urobilin,

H3C O

N H

H3COOC

COOCH3

CH3

CH3

N H

CH3 O

N H

N

stercobilin dimethyl ester

H3C O

N H

H3COOC

COOCH3

CH3

CH3

N H

CH3 O

N H

N

half-stercobilin dimethyl ester

H3C O

N H

H3COOC

COOCH3

CH3

CH3

N H

N

CH3 N H

O

urobilin dimethyl ester

Figure 7.8. Chemical structures of stercobilin dimethyl ester, half-stercobilin dimethyl ester and urobilin dimethyl ester.

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Figure 7.9. HPLC separation of dimethyl esters of (SS)-stercobilin (1), (RR,SS )-half-stercobilin (2), (RR,SS )-urobilin (3), (RS,S )-half-stercobilin (4), and (RS,SR)-urobilin. Column: Hypersil; eluent: n-heptane/methyl acetate/ methanol containing 1% diethylamine (75:25:2, v/v/v).

(RS,SR)-half-stercobilin and (RS,SR)-urobilin (Fig. 7.9). A Hypersil silica column (250 mm × 4.6 mm) was used with n-heptane/methyl acetate/methanol containing 1% diethylamine (75:25:2, v/v/v) as mobile phase. Column efficiency was improved by the addition of a small quantity of diethylamine.

7.7. Analysis and Determination of Bile Pigments in Biological and Clinical Samples by HPLC The analysis and determination of urinary and faecal bile pigments, unlike the determination and profile analysis of porphyrins, are of limited clinical value. The determination of bilirubin in serum, on the other hand, is of important clinical value for the investigation of familial hyperbilirubinaemia13 and the diagnosis of various forms of pre-hepatic and post-hepatic jaundice in the adult. Jaundice is due to an increased concentration of bilirubin in the blood caused by excessive haemolysis of red blood cells in the reticuloendothelial

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system (haemolytic jaundice) or to obstruction in the biliary passages by stone or tumour (obstructive jaundice). The so-called catarrhal jaundice is believed to be due to obstruction of the bile ducts by catarrh.

7.7.1.

HPLC methods for the determination of conjugated and unconjugated bilirubin in serum

In normal subjects serum bilirubin is almost entirely unconjugated. Unconjugated bilirubin is increased in neonatal jaundice where there is a deficiency of the conjugating enzyme. In haemolytic anaemia or pre-hepatic jaundice serum bilirubin is mainly unconjugated and conjugated bilirubin concentration is often low. However, if increased haemopoisis fails to maintain adequate concentrations of haemoglobin, the anaemia may become sufficiently severe that anoxia causes liver damage and an increase in conjugated bilirubin in serum. In post-hepatic (obstructive) jaundice and also in cholestatic jaundice common in many patients with severe hepatitis, serum bilirubin is almost all conjugated. In hepatitis with less cholestasis unconjugated bilirubin is increased in the serum accompanied by small quantity of conjugated bilirubin. The proportions of conjugated and unconjugated bilirubin are probably of little significance in hepatitis. The determination of conjugated and unconjugated bilirubin in serum is necessary to distinguish Dubin- Johnson and Rotor’s syndrome where the serum contains both conjugated and unconjugated bilirubin, from the pure unconjugated hyperbilirubinaemia of Gilbert’s syndrome due to a defect of transport of unconjugated bilirubin and of the Crigler- Najjar syndrome where there is a defect in the enzymatic conjugation mechanism. Dubin- Johnson syndrome is due to a defect in the excretion of conjugated bilirubin with no impairment of the conjugation mechanism. The abnormality in Rotor’s syndrome is the failure of uptake and storage of unconjugated bilirubin. Bilirubin covalently bound to serum albumin14,15 (biliprotein or δ-bilirubin) is an important fraction of total bilirubin in conditions

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where the hepatic excretion of bilirubin is impaired, as in the DubinJohnson syndrome and in patients with hepatocellular and cholestatic jaundice. It may account for up to 90% of total serum bilirubin. It has been demonstrated that the ratio of conjugated bilirubin/ δ-bilirubin and concentration of conjugated bilirubin are useful as a prognostic index after treatment of percutaneous trans-hepatic cholangiodrainage and liver transplantation.16-18 There are basically three HPLC procedures for the determination of conjugated and unconjugated bilirubin in serum. The first is by trans-esterification of the glycoconjugates into the methyl esters with 2% methanolic KOH followed by solvent extraction and HPLC separation; the second is by solid-phase extraction (SPE) of bilirubin and its glycoconjugates for separation and determination and the third is by direct injection of serum into a suitable HPLC column without sample preparation. Whichever method is used it is important to remember that bilirubin and its derivatives must be handled with care to avoid undesired degradation or structural modification. The samples and standards must be protected from exposure to light and all experiments carried out in dim light in deoxygenated solvents preferably in the presence of an anti-oxidant, e.g., ascorbic acid, and EDTA.

7.7.1.1. Extraction and determination of conjugated and unconjugated bilirubin in serum following trans-esterification The quantitative determination of conjugated and unconjugated bilirubin in serum by trans-esterification of the glycoconjugates into the methyl esters19 has two main advantages. These are effective and quantitative extraction of bilirubin and bilirubin mono- and dimethyl esters into organic solvents and the availability of calibration standards. Bilirubin mono- and dimethyl esters can be prepared in sufficient quantities from commercially available bilirubin as follows: 1. Dissolve bilirubin (100 mg) in dichloromethane (125 ml) with refluxing.

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2. After cooling to room temperature add an excess quantity of a solution of diazomethane in ether. 3. Evaporate the mixture to dryness at 35°C using a rotary evaporator. The above reaction gave a mixture of bilirubin mono- and di-methyl ester which can be purified by preparative thin-layer chromatography on silica or by HPLC. The procedure for the trans-esterification reaction is as follows: 1. Add 30 mg of ascorbic acid and 1 mg of EDTA to 250 µl of serum. 2. Add 3 ml of 2% methanolic KOH (2g of KOH in 100 ml of methanol), vortex-mix for 1 min and leave standing for 1 min. 3. Add 5 ml of dichloromethane and 5 ml of glycine/HCl buffer, pH 2.7 (glycine in 0.4 M HCl) sequentially, shake mixture vigorously and centrifuge at 2000 × g for 5 min. 4. Collect the dichloromethane extract and evaporate to dryness at 30°C. Store at −20°C under nitrogen until analysis by HPLC. Figure 7.10 shows the reversed-phase HPLC separation of conjugated and unconjugated bilirubins in serum after alkaline methanolysis on a Hypersil ODS (C18) column with acetonitrile/DMSO/0.5 M ammonium acetate, pH 4.6 (50:50:40, v/v/v) as eluent.

7.7.1.2. Solid-phase extraction (SPE) and determination of conjugated and unconjugated bilirubin in serum The main advantage of SPE is it allows direct extraction of bilirubin and its glycoconjugates without the needs for derivatisation. The technique is based on the fact that caffeine/sodium benzoate solution is able to release conjugated and unconjugated bilirubin from their protein binding sites, thus allowing them to be extracted onto a reversed-phase cartridge.20,21 A wide range of cartridges can be used, for examples, Bond-Elute C8 and Sep-Pak C18 cartridges. The caffeine/ sodium benzoate solution, similar to that used for the classical Jendrassik-Grof method for the estimation of serum bilirubin, is prepared by adding caffeine (37.5 g), sodium benzoate (57.0 g) and sodium

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Figure 7.10. HPLC separation of conjugated and unconjugated bilirubins following alkaline methanolysis of the plasma of a patient with hyperbilirubinaemia. Column: Hypersil-ODS; eluent: 0.5 M ammonium acetate, pH 4.6/acetonitrile/DMSO (40:50:50, v/v/v).

acetate (94.5 g) to water (500 ml) at 60°C with stirring until completely dissolved. The solution is made up to 1 litre with water after cooling to room temperature. A typical extraction procedure is as follows: 1. Condition a C8 cartridge containing 100–200 mg of packing material with 2 ml of methanol followed by 2 ml of caffeine/ sodium benzoate solution. 2. Mix serum (100–500 µl) with 500 µl of caffeine/sodium benzoate solution and 10 µl of mesobilirubin in DMSO. 3. Load the mixture into the cartridge and collect the eluate. 4. Wash cartridge with 2 ml of caffeine/sodium benzoate solution and collect the eluate. 5. Combine eluate 3 and 4 in a 5 ml volumetric flask, make up to volume and use for determination of covalently bonded bilirubin-albumin (biliprotein or δ-bilirubin) by a modified

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Jendrassik- Grof method. 6. Wash cartridge with water (1 ml) followed by 20% (v/v) methanol in 0.1 M ammonium acetate buffer, pH 5.16 (1 ml) to remove caffeine/sodium benzoate from the cartridge. 7. Elute the adsorbed conjugated and unconjugated bilirubin from the cartridge with 1 ml of acetonitrile/methanol (1:1, v/v) into a 2 ml polypropylene screw top tube and evaporate to dryness under nitrogen at 40°C. 8. Dissolve the residue in 25 µl of DMSO, dilute with 500 µl of HPLC mobile phase and sonicate for 1 min to ensure solution of unconjugated bilirubin. The solution is used for HPLC analysis. The residue may also be stored at −20°C in the dark until HPLC analysis. Mesobilirubin (internal standard) is prepared by hydrogenation of bilirubin in 0.1 M NaOH with palladium on charcoal as catalyst. Covalently bound bilurubin-albumin or biliprotein reference material can be isolated from patients with drug-induced cholestasis by membrane filtration.20,22 Figure 7.11 shows the separation of conjugated and unconjugated bilirubin in the serum extract of a patient with hyperbilirubinaemia (a) and a normal subject (b) on a Hypersil ODS column (5 µm particle size, 250 × 4.6 mm) with 0.5 M ammonium acetate buffer, pH 5.16 (solvent A) and 1:1 mixture of acetonitrile/DMSO (solvent B) as eluent. The elution program, suitable for analysis of a batch of samples using an auto-sampler, is as follows: 1. 2. 3. 4.

0 to 5 min, linear gradient from 55 to 80% B. 5 to 8 min, isocratic elution at 80% B. 8.1 min, reset to 55% B. 13 min, program complete, reset to time 0, automatic injection of second sample and restart elution program.

The separation can also be carried out isocratically using the mobile phase described in Sec. 7.3.

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Figure 7.11. Separation of conjugated and unconjugated bilirubin in plasma from a patient with hyperbilirubinaemia (a) and a normal subject (b). Column: Hypersil-ODS; eluents: 0.5 M ammonium acetate, pH 5.16 (A) and 1:1 (v/v) acetonitrile/DMSO (B); elution: 0 to 5 min, linear gradient from 55 to 80% B; 5 to 8 min, isocratic elution at 80% B. Peaks: 1 = bilirubin diglucuronide; 2 = bilirubin C-8 monoglucuronide; 3 = bilirubin C-12 monoglucuronide; 4 = mesobilirubin; 5 = bilirubin.

7.7.1.3. Simultaneous determination of conjugated bilirubin, unconjugated bilirubin and biliprotein or δ-bilirubin by direct injection HPLC There are two useful methods reported for the simultaneous separation and determination of conjugated bilirubin, unconjugated bilirubin and δ-bilirubin by direct injection of serum into organic polymer-based column packing materials. In the first method,23,24 a Micronex RP-30 column (150 mm × 6 mm i.d.) from Sekisui Chemical Co., Osaka, Japan, was used. The packing material was 8–10 µm porous cross-linked organic polymer gel consisted of 80% poly(ethylene glycol) dimethylacrylate and 20% tetramethylol methane tetraacrylate. The eluents were 5 mM pentanesulphonic acid solution containing 0.1 M of acetic acid (solvent A) and acetonitrile (solvent B). A typical elution program at a flow rate of 1 ml/min is as follows: 1. 0 to 20 min, linear gradient from 100% A (0% B) to 50% A (50% B).

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2. 20 to 27 min, linear gradient from 50% B to 100% B. 3. 27 to 32 min, isocratic elution at 100% B. The elution was in the order of δ-bilirubin, bilirubin diglucuronide, bilirubin monoglucuronide and unconjugated bilirubin. Photobilirubins (the photo-induced derivatives of bilirubin), if present, can also be detected. A Shodex Asahipak GS-320HQ column (300 mm × 7.6 mm i.d.) was used in conjunction with a 50 mm × 7.6 mm i.d. GS-2G 7B grad in series in the second method.24 The columns were from Showadenko, Tokyo, Japan, and contained 6 µm particle size polyvinylalcohol gel. The separation of δ-bilirubin, bilirubin diglucuronide, bilirubin monoglucuronide and unconjugated bilirubin was achieved by isocratic elution at 0.7 ml/min with 30% acetonitrile and 70% phosphate buffer (0.3 M, pH 6.5) containing 1% Brij 35 and 0.08% sodium ascorbate (v/v) as mobile phase. The buffer solution was prepared by dissolving Brij 35 in 0.3 M phosphate buffer adjusted to pH 6.5 and sodium ascorbate was added to the buffer just before use since it is relatively unstable at pH 6.5. HPLC methods for the determination of conjugated bilirubin, unconjugated bilirubin and δ-bilirubin are best used as reference methods for routine clinical chemistry procedures of bilirubin measurement, especially when the results are doubtful or inconsistent.

7.8. HPLC/ESI-MS and MS/MS of Bile Pigments For HPLC/MS analysis DMSO, which is relative involatile, can be ommited from the mobile phase system. The HPLC/MS systems described for porphyrins (see Chapter 3) are also applicable to bile pigments, including systems using ammonium acetate buffer with acetonitrile and methanol, and TFA with acetonitrile. Gradient elution is generally used for the analysis of bile pigments in biological extracts. Figure 7.12 shows the HPLC/ESI-MS chromatograms of biliverdin and bilirubin IIIα, IXα and XIIIα isomers. The separation was carried out on a Hypersil BDS C18 column (250 mm × 4.6 mm, 5 µm particle size)

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Figure 7.12. HPLC/ESI-MS chromatograms of bilirubin XIIIα (19.9 min), IXα (A; 21.0 min), IIIα (22.2 min), and biliverdin XIIIα (17.2 min), IXα (B; 17.6 min) and IIIα (17.9 min) isomers. Column: Hypersil-BDS C18 ; Solvents: 0.5 M ammonium acetate (A) and acetonitrile (B); elution: isocratic elution at 5% B (95% A) for 5 min, then linear gradient elution from 5 to 80% B in 20 min.

with 0.5 M ammonium acetate (solvent A) and acetonitrile (solvent B) as gradient elution solvent mixtures. The elution program was an initial isocratic elution at 5% B (95% A) for 5 minutes, then followed by linear gradient elution from 5% B to 80% B (20% A) in 20 minutes. The total run time was 25 minutes. The system, developed for the simultaneous analysis of bilirubin and the dipyrroles formed by bilirubin degradation,25 is also suitable for the separation of photobilirubins as shown in Fig. 7.13. The initial isocratic elution can be omitted if separation of dipyrroles is not required. The mass spectrum of bilirubin showed the expected [M + H]+ ion at m/z 585. Radical cation of bilirubin at m/z 584 was also observed. Similarly, the mass spectrum of biliverdin gave a [M + H]+ ion at m/z 583 and a radical cation at m/z 582.

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Figure 7.13. HPLC/ESI-MS chromatograms of bilirubins and photobilirubins (16.0, 16.4 and 16.6 min). HPLC/MS conditions are as described in Fig. 7.11.

Tandem ESI-MS/MS of bile pigments is dominated by cleavage of the central tetrapyrrole bridge to give a prominent dipyrrole product ion. Thus protonated bilirubin gave a prominent product ion at m/z 299 resulting from cleavage of the central methane bridge, and protonated biliverdin gave an intense ion at m/z 297. The product ion spectrum of biliverdin (m/z 583) is shown in Fig. 7.14. Cleavage of the protonated biliverdin molecule at the central tetrapyrrole bridge gave the most abundant product ion at m/z 297 and a minor radical cation at m/z 285. The ion at m/z 297 was further fragmented by the loss of CO (28 mass units) to give an intermediate which can either cyclise to give the ion at m/z 269 (297 − 28) or accept two protons to give the structure at m/z 271 (297 − 28 + 2H). The ion at m/z 297 may also lose H2O to give a cyclic structure at m/z 279 (297 − 18) which was further fragmented by losing CO and accepting 2H to give the structure at m/z 253. The proposed fragmentation pathways are shown in Fig. 7.15.

CH3 100

x24

COOH

CH3

CH3

297

N

N H

583

m/z 583 [M + H] +

565

271

279 285

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227

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211

Figure 7.14.

220

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COOH

HOOC CH3

CH3

COOH

CH3

CH3

CH3

+ H]+ O

N H m/ z 297

N

N H

N H

N H

N H

COOH

COOH

CH3

CH3

- CO O

N H

H

N

O

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CH3

CH3

O

+ 2H

CH3

H2N

N

H +

m/ z 271

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CH3

CH3

- CO

CH3

HN

N m/ z 269

CH3

CH3 O

O H +

O

N H

N

+

+ 2H

H2N

m/ z 279 (297 - 18)

N

+

m/ z 253

Figure 7.15. Proposed ESI-MS/MS fragmentation pathways of protonated biliverdin molecule.

Like protoporphyrin, the precursor ion loses H2O to give the peak at m/z 565 (583 − 18) and the sequential loss of two .CH2COOH radicals gave the ions at m/z 524 (583−59) and m/z 465 (524−59), respectively.

7.9. Characterisation of Tetrapyrrole Pigments in Avian Eggshells The origin, evolution and functions of avian eggshell colour and patterning are of interest to evolution biologists and ornithologists.26,27 These pigments can be easily and accurately characterised by the HPLC/ESI-MS/MS technique described for porphyrins and bile pigments. The successful application of the technique, however, is dependent on a suitable extraction method before HPLC/MS analysis. Zn-biliverdin and Zn-protoporphyrin have been reported in the eggshells, methods using methanolic sulphuric acid or hydrochloric

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acid to extract eggshell pigments must therefore be avoided, because they demetallate these compounds.

7.9.1. Extraction of pigments from eggshells An extraction procedure28 using EDTA and acetonitrile-acetic acid or acetonitrile-DMSO, which do not demetallate Zn-protoporphyrin, is described below. 1. Break eggshells into small pieces and place 100–200 mg in a 1.5 ml conical polypropylene Eppendorf centrifuge tube. 2. Add 0.5 ml of an aqueous solution of disodium EDTA (100 mg/ml) adjusted to pH 7.2 with NaOH. 3. Vortex-mix for 1 min and carefully release the stopper of the tube. 4. After cessation of effervescence, centrifuge the tube at 15000 × g for 30 sec. 5. Discard the supernatant from step 4 and repeat steps 2, 3 and 4 three times. 6. Add 1.0 ml of acetonitrile-acetic acid (4:1, v/v) or acetonitrileDMSO (4:1, v/v) to the final pellet and vortex-mix vigorously for 2 min in 30 sec burst. Uncap the tube to allow escape of CO2. 7. Centrifuge at 15000 × g for 2 min. 8. Transfer supernatant from step 7 into a clean tube for HPLC/ ESI-MS and MS/MS or store at 4°C until analysis, generally within 24 hours of preparation. The above method was effective for the extraction of protoporphyrin, Zn-protoporphyrin and biliverdin from eggshells for characterisation by HPLC/ESI-MS/MS. EDTA at neutral pH does not demetallate Zn-protoporphyrin. Zn-biliverdin, however, was demetallated by EDTA and could not be recovered unchanged. The method is also not suitable for quantitative extraction and analysis because complete recovery of pigments would require much longer final extraction with acetonitrile-acetic acid or acetonitrile-DMSO.

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Without prior treatment with EDTA, eggshell pigments could not be extracted with organic solvents such as acetonitrile, methanol and DMSO even on prolonged contact. EDTA demineralises the calcium carbonate matrix of eggshells, thus releasing the pigments for extraction into organic solvents. Figure 7.16 shows the HPLC/ESI-MS chromatogram of the bile pigment and porphyrin extracted from the eggshell of the common nighthawk. The peaks at m/z 583 and m/z 563 are the protonated molecules of biliverdin and protoporphyrin, respectively, and have been confirmed by tandem MS/MS analysis. The separation was carried out on a Hypersil BDS C18 column (250 mm × 4.6 mm, 5 µm particle size). The solvents were 0.05% trifluoroacetic acid in water (solvent A) and acetonitrile (solvent B). The elution program was linear gradient from 10% B to 100% B in 25 minutes, followed by isocratic elution at 100% B for a further 10 min. The eluent from the HPLC was split in the ratio of 9:1, allowing 100 µl/min to enter the mass spectrometer. TFA at 0.05–0.1% does not demetallate Zn-protoporphyrin. It will, however, remove Zn from Zn-biliverdin. An alternative solvent system in which Zn-biliverdin is stable, with 0.5 M ammonium acetate in water (solvent A) and acetonitrile (solvent B) as gradient mixtures, was used for the separation of biliverdin and its Zn complex. The elution program was linear gradient from 20% B to 55% B in 20 minutes. Figure 7.17 demonstrates further the applicability of the method by the detection of a small amount of protoporphyrin in the presence of a large quantity of biliverdin in the blue-green eggshell of the emu. The method has been used to analyse eggshell pigments from other bird species, including captive Japanese quail, American robin, the common cuckoo, the great tit and the sooty tern.28 In all the above avian eggshell types the principal pigments characterised by HPLC/ESI-MS/MS are biliverdin and protoporphyrin. No Zn-protoporphyrin could be detected.28 It is likely that previous reports of the occurrence Zn-protoporphyrin and Zn-biliverdin in avian eggshells were artefacts resulted from sample preparation and

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Figure 7.16. HPLC/ESI-MS chromatograms of biliverdin (m/z 583) and protoporphyrin (m/z 563) extracted from nighthawk eggshell. Column: Hypersil-BDS C18 ; solvents: 0.05% TFA (A) and acetonitrile (B); elution: linear gradient from 10 to 100% B in 25 min, then isocratic elution at 100% B for a further 10 min.

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18.91

33.82 35.26

13.27

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5.00

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17.50

20.00

22.50

25.00

27.50

30.00

32.50

35.00

37.50

40.00

42.50

Time 45.00

Figure 7.17. HPLC/ESI-MS chromatograms of biliverdin (m/z 583) and protoporphyrin (m/z 563) extracted from emu eggshell. HPLC/ESI-MS conditions are as described in Fig. 7.15.

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EMU 400-800 CONE 60V

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extraction procedures. In the presence of Zn ion in aqueous solutions biliverdin forms Zn-complex easily.

7.10. HPLC and HPLC/MS of Phycobilins The phycobilins are structurally similar to the bile pigments described in the previous sections. They were mainly separated by reversed-phase HPLC on C18 columns. Mobile phases developed for the analytical and semi-preparative separation of phycobilins include acetone/ethanol/100 mM formic acid (25:65:10; v/v/v),29 acetone/ 20 mM formic acid (50:50; v/v)30 and ethanol/acetone/water/acetic acid (48:34:17:1; v/v/v/v).31 Linear gradient elution with water and a mixture of acetone/ethanol/water/acetic acid as the gradient solvents, has also been described.32 Gradient elution using 0.05–0.1% TFA/acetonitrile, 0.1–0.5 M ammonium acetate buffer/acetonitrile or 0.1–0.5 M ammonium acetate buffer/acetonitrile/methanol as solvent mixtures is recommended for HPLC/MS analysis. These systems are widely used for porphyrin and bile pigment analysis and should also be suitable for the phycobilins. Phycobilins are usually obtained from phycobiliproteins by the HgCl2-assisted methanolysis procedure.33–35

7.11. HPLC Separation of Phycobiliproteins Phycobiliproteins can by separated by reversed-phase HPLC systems commonly used for the chromatography of peptides and proteins. This is usually performed by gradient elution on a C-4 wide-pore reversed-phase column with 0.005–0.1% TFA and acetonitrile as the gradient solvent mixtures. For example, phycocyanin isolated from the blue-green alga Aphanizomenon flos-aquae was analysed on a Vydac Protein C-4 column (250 mm × 4.6 mm; 5 µm particle size) by linear gradient elution from 20% acetonitrile in a mixture of acetonitrile/0.1% TFA (20:80; v/v) to 100% acetonitrile in 60 minutes, followed by isocratic elution at 100% acetonitrile for a further 10 min.36

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B-phycoerythrin, R-phycoerythrin, allophycocyanin and C-phycocyanins have been separated using similar systems.37,38 There are considerable recent interests in the potential applications of phycocyanin in biotechnology, food and medicine.39

References 1. Stoll MS, Lim CK and Gray CH. The separation of bile pigments by HPLC. In: High Pressure Liquid Chromatography in Clinical Chemistry (Dixon PF, Gray CH, Lim CK and Stoll MS eds); Academic Press, London, New York, San Francisco 1976; pp. 97–108. 2. Heirwegh KP, Fevery J and Blanckaert N. Chromatographic analysis and structure determination of biliverdins and bilirubins. Journal of Chromatography 1989; 496: 1–26. 3. Stoll MS and Gray CH. The preparation and characterization of bile pigments. Biochemical Journal 1977; 163: 59–101. 4. Li F, Lim CK and Peters TJ. Reversed-phase high-performance liquid chromatography of conjugated and unconjugated bilirubins in body fluids. Journal of Chromatography 1986; 353: 19–26. 5. Levin EY. Studies on verdohemochrome. Biochemistry 1966; 5(9): 2845–2852. 6. Bonnett R and McDonagh AF. The meso-reactivity of porphyrins and related compounds. VI. Oxidative cleavage of the haem system. The four isomeric biliverdins of the IX series. Journal of Chemical Society, Perkin 1; 1973; 9: 881–888. 7. Hirota K, Yamamoto S and Itano A. Urinary excretion of isomers of biliverdin after destruction in vivo of haemoproteins and haemin. Biochemical Journal 1985; 229: 477–483. 8. Cole KD and Little GH. Isocratic high-performance liquid chromatography of bile pigments. Journal of Chromatography 1982; 227: 503–509. 9. Rasmussen RD, Yokoyama WH, Blumenthal SG, Bergstrom DE and Ruebner BH. High-performance liquid chromatographic separation and quantification of the four biliverdin dimethyl ester isomers of the IX series. Analytical Biochemistry 1980; 101(1): 66–74. 10. Sakamoto H, Omata Y, Adachi Y, Palmer G and Noguchi M. Separation and identification of the regioisomers of verdoheme by reversed-phase

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12.

13.

14.

15.

16.

17.

18.

19.

20.

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ion-pair high-performance liquid chromatography, and characterization of their complexes with heme oxygenase. Journal of Inorganic Biochemistry 2000; 82(1–4): 113–121. Schoch S, Lempert U, Wieschhoff H and Scheer H. High-performance liquid chromatography of tetrapyrrole pigments. Journal of Chromatography 1978; 157: 357–364. Bull VA, Lim CK and Gray CH. High-performance liquid chromatography of bile pigments: separation and chracterization of the urobilinoids. Journal of Chromatography 1981; 218: 647–652. American Academy of Pediatrics. Practice parameter: Management of hyperbilirubinemia in the healthy term newborn. Pediatrics 1994; 94: 558–565. Weiss JS, Gautam A, Lauff JJ, Sundberg MW, Jatlow P, Boyer JL and Seligson D. The clinical importance of a protein-bound fraction of serum bilirubin in patients with hyperbilirubinemia. New England Journal of Medicine 1983; 309(3): 147–150. McDonagh AF, Palma LA, Lauff JJ and Wu TW. Origin of mammalian biliprotein and rearrangement of bilirubin glucuronides in vivo in the rat. Journal of Clinical Investigation 1984; 74(3): 763–770. Arvan D and Shirey TL. Conjugated bilirubin: A better indicator of impaired hepatobiliary excretion than direct bilirubin. Annals of Clinical and Laboratory Sciences 1985; 15: 252–259. Wu TW, Levy GA, Yiu S, Au JX, Greig PD, Strasberg SM, Ettles M, Abecassis M, Superina RA, Langer B, Blendls LM, Phillips MJ and Taylor BR. Delta and conjugated bilirubin as complementary marker of early rejection in liver-transplant recipients. Clinical Chemistry 1990; 36(1): 9–14. Akamatsu Y, Ohkohchi N, Seya K and Satomi S. Analysis of bilirubin fraction in the bile for early diagnosis of acute rejection in living related liver transplantation. Tohoku Journal of Experimental Medicine 1997; 181: 145–154. Blanckaert N. Analysis of bilirubn mono- and di-conjugates. Determination of their relative amounts in biological samples. Biochemical Journal 1980; 185: 115–128. McKavanagh SM and Billing BH. The use of Bond-Elut for the estimation of serum bile pigments bonded covalently to albumin. Biomedical Chromatography 1987; 2(2): 62–65.

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21. Li F, Lim CK and Peters TJ. Solid phase extraction and direct HPLC separation of bilirubin and its conjugates in plasma. Chromatographia 1987; 24: 637–638. 22. Lauff JJ, Kasper ME, Wu TW and Ambrose RT. Isolation and preliminary characterization of a fraction of bilirubin in serum that is firmly bound to protein. Clinical Chemistry 1982; 28(4 Pt 1): 629–637. 23. Adachi Y, Inufusa H, Yamashita M, Kambe A, Yamazaki K, Sawada Y and Yamamoto T. Clinical application of serum bilirubin fractionation by simplified liquid chromatography. Clinical Chemistry 1988; 34(2): 358–388. 24. Osawa S, Sugo S, Yoshida T, Yamaoka T and Nomura F. An assay for separating and quantifying four bilirubin fractions in untreated human serum using isocratic high-performance liquid chromatography. Clinica Chimica Acta 2006; 366: 146–155. 25. De Matteis F, Lord GA, Lim CK and Pons N. Bilirubin degradation by uncoupled cytochrome P450. Comparison with chemical oxidation system and characterization of the products by high-performance liquid chromatography/electrospray ionization mass spectrometry. Rapid Communications in Mass Spectrometry 2006; 20: 1209–1217. 26. Kilner RM. The evolution of egg colour and patterning in birds. Biological Reviews 2006; 81: 383–406. 27. McGraw KJ. The mechanics of uncommon colors: Pterins, porphyrins, and psittacofulvins. In: Bird coloration (Hill GE, McGraw KJ eds), Vol. 1: Mechanisms and Measurements, pp. 354–398. Harvard University Press: Harvard, MA, 2006. 28. Gorchein A, Lim CK and Cassey P. Extraction and analysis of colourful eggshell pigments using HPLC and HPLC/electrospray ionization tandem mass spectrometry. Biomedical Chromatography 2009; 23(6): 602–606. 29. Terry MJ and Kendrick RE. The aurea and yellow-green-2 mutants of tomato are deficient in phytochrome chromophore synthesis. Journal of Biological Chemistry 1996; 271: 21681–21686. 30. Wu S-H, McDowell MT and Lagarias JC. Phycocyanobilin is the natural precursor of the phytochrome chromophore in the green alga Mesotaenium caldariorum. Journal of Biological Chemistry 1997; 272: 25700–25705.

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31. Terry MJ, McDowell MT and Lagarias JC. (3Z )- and 3(E )-phytochromobilin are intermediates in biosynthesis of the phytochrome chromophore. Journal of Biological Chemistry 1995; 270: 11111–11119. 32. Schluchter WM and Glazer AN. Characterization of cyanobacterial biliverdin reductase. Journal of Biological Chemistry 1997; 272: 13562–13569. 33. Arciero DM, Bryant DA and Glazer AN. In vitro attachment of bilins to apophycocyanin. I. Specific covalent adduct formation at cysteinyl residues involved in phycocyanobilin binding in C-phycocyanin. Journal of Biological Chemistry 1988; 263: 18343–18349. 34. Beale SI and Cornejo J. Biosynthesis of phycobilins. 3(Z )-phycoerythrobilin and 3(Z )-phycocyanobilin are intermediates in the formation of 3(E )-phycocyanobilin from biliverdin IXα. Journal of Biological Chemistry 1991; 266: 22333–22340. 35. Terry MJ, Maines MD and Lagarias JC. Inactivation of phytochrome- and phycobiliprotein-chromophore precursors by rat liver biliverdin reductase. Journal of Biological Chemistry 1993; 268: 26099–26106. 36. Benedetti S, Rinalducci S, Benvenuti F, Francogli S, Pagliarani S, Giorgi L, Micheloni M, D’Amici GM, Zolla L and Canestrari F. Purification and characterization of phycocyanin from the blue-green alga Aphanizomenon flos-aruae. Journal of Chromatography B 2006; 833: 12–18. 37. Swanson RV and Glazer AN. Separation of phycobiliprotein subunits by reverse-phase high-pressure liquid chromatography. Analytical Biochemistry 1990; 188(2): 295–299. 38. Bermejo R, Talavera EM and Alvarez-Pez JM. Chromatographic purification and characterization of B-phycoerythrin from Porphyridium cruentum. Semipreparative high-performance liquid chromatographic separation and characterization of its subunits. Journal of Chromatography A 2001; 917(1–2): 135–145. 39. Eriksen NT. Production of phycocyanin-a pigment with applications in biology, biotechnology, food and medicine. Applied Microbiology and Biotechnology 2008; 80: 1–14.

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CHAPTER 8

Future Directions of HPLC and Mass Spectrometry of Tetrapyrroles

High-performance liquid chromatography and mass spectrometry have become the indispensable techniques for the separation, analysis, and characterisation of the tetrapyrroles. Applications range from diagnosis of diseases, metabolic profiling, identification and characterisation of metabolites in animals and plants, drug discovery, impurity and degradation product analysis in pharmaceuticals, biotechnology, and geochemistry. Over the past 5 years significant advances and developments in both HPLC and mass spectrometry have been achieved. The sensitivity and resolution of mass spectrometers continue to improve and advances in HPLC column technology and instrumentation led to the development of capillary high-performance liquid chromatography1 and ultra performance liquid chromatography2–5 (UPLC) for coupling with mass spectrometry. The Thermo Electron (San Jose, CA, USA) LTQ OrbitrapTM mass spectrometer can achieve a specified resolving power of >60,000 full width at half height (FWHM) using Fourier transforms. High resolution HPLC coupled to high resolution and high sensitivity mass spectrometry will be the technique of choice for the analysis and characterisation of porphyrins, chlorophylls and bilins in the future. It is clear from the previous chapters that sophisticated and complicated mobile phases are not necessary and two simple mobile phase systems are recommended for reversed-phase HPLC and HPLC/ MS of all these tetrapyrroles. These are: 1, ammonium acetate/acetic acid buffer (0.1–1 M, pH 4.6–6.8) with acetonitrile and/or methanol 223

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as organic modifier; 2, TFA (0.01–0.1%) with acetonitrile as the organic modifier. Capillary HPLC, which uses 30–50 µm internal diameter fusedsilica columns eluting at a very slow mobile phase flow rate1 of ~50 nl/min, is not suitable for the separation of porphyrins. At a very slow flow rate porphyrins are completely adsorbed and immobilised onto the silica adsorbance and are therefore not eluted. Although the eluent leaving the HPLC column enters the mass spectrometer via a fused-silica capillary, adsorption of porphyrins onto the relatively short capillary is minimal at the faster flow rate (100 µl/min) in conventional or microbore HPLC/MS operation. The adsorption problem is, however, compounded in capillary HPLC/MS analysis of porphyrins. Adsorption onto fused-silica capillary is compound dependent and porphyrins are particularly prone to adsorption. UPLC uses small particle size (1.7 µm) packing materials to increase the speed, sensitivity and resolution of HPLC analysis.2–5 This requires the HPLC instrument and columns capable of providing and withstanding liquid flow of up to 1034 bar. For most commercial HPLC instruments the maximum operation pressure is around 400 bar. The ACQUITY ultra performance liquid chromatography system (Waters Corp., Milford, USA) has been shown to provide superior speed, resolution and sensitivity of analysis for many compounds.2–5 The technique has great potential and should also be explored for application to the tetrapyrroles. At present, 5 µm particle size packing material is preferred, although 2.1–3.5 µm materials are also available. Columns packed with smaller particles require more thorough and cleaner sample preparation procedures because they are more easily blocked when biological extracts are analysed. It is sometimes difficult to develop an ideal sample preparation method for a compound or group of compounds of interest, for example, the extraction porphyrins in the Harderian glands of rodents (Chapter 5) and the pigments, especially Zn-biliverdin, from the eggshells of birds (Sec. 7.9, Chapter 7). Desorption electrospray ionisation6,7 (DESI) and Direct Analysis in Real Time8 (DART) are two ambient desorption ionisation mass spectrometric techniques9 which could be useful in such a situation. DESI uses a spray of electrospray solvent

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composed of charged droplets as ionisation agent and has been successfully used to ionise compounds in solid samples. DART uses primary and secondary ionisation brought about by metastable atoms of helium or molecules of nitrogen and is a dry technique. When coupled to TOF mass analysers, it permits high-resolution, accurate mass measurement of compounds in gas, liquid and solid samples.10–13 It would be interesting to see whether DART could be successfully used to analyse the pigments on the surface of eggshells and/or the porphyrins in the Harderian glands of rodents. Other potential applications of DART include the identification of petroporphyrins and the direct analysis of tetrapyrroles in cell and tissue samples of both animal and plant origin.

References 1. Ito S, Yoshioka S, Ogata I, Yamashita E, Nagai S, Okumoto T, Ishii K, Ito M, Kaji H, Takao K and Deguchi K. Capillary high-performance liquid chromatography/electrospray ion trap time-of-flight mass spectrometry using a novel nanaflow gradient generator. Journal of Chromatography A 2005; 1090: 178–183. 2. de Villiers A, Lestremau F, Szucs R, Gélébart S, David F and Sandra P. Evaluation of ultra performance liquid chromatography Part I. Possibilities and limitations. Journal of Chromatography A 2006; 1127: 60–69. 3. Guillarme D, Nguyen DTT, Rudaz S and Veuthey J-L. Recent development in liquid chromatography — Impact on qualitative and quantitative performance. Journal of Chromatography A 2007; 1149: 20–29. 4. Guillarme D, Nguyen DTT, Rudaz S and Veuthey J-L. Method transfer for fast liquid chromatography in pharmaceutical analysis: Application to short columns packed with small particle. Part II: Gradient experiments. European Journal of Pharmaceutics and Biopharmaceutics 2008; 68: 430–440. 5. Russo R, Guillarme D, Nguyen DTT, Bicchi C, Rudaz S and Veuthey J-L. Pharmaceutical applications on columns packed with sub-2 microm particles. Journal of Chromatographic Science 2008; 46(3): 199–208. 6. Takats Z, Wiseman JM, Gologan B and Cooks RG. Mass spectrometry sampling under ambient conditions with desorption electrospray ionization. Science 2004; 306(5695): 471–473.

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7. Takats Z, Wiseman JM and Cooks RG. Ambient mass spectrometry using desorption electrospray ionization (DESI): Instrumentation, mechanism and applications in forensics, chemistry, and biology. Journal of Mass Spectrometry 2005; 40(10): 1261–1275. 8. Cody RB, Laramée JA and Durst HD. Versatile new ion source for the analysis of materials in open air under ambient conditions. Analytical Chemistry 2005; 77(8): 2297–2302. 9. Verter A, Nefliu M and Cooke RM. Ambient desorption ionization mass spectrometry. Trends in Analytical Chemistry 2008; 27(4): 284–290. 10. Cody RB. Observation of molecular ions and analysis of nonpolar compounds with the direct analysis in real time ion source. Analytical Chemistry 2009; 81(3): 1101–1107. 11. Yu S, Crawford E, Tice J, Musselman B and Wu J-T. Bioanalysis without sample clean up or chromatography: The evaluation and initial implementation of direct analysis in real time ionization mass spectrometry for quantitation of drugs in biological matrixes. Analytical Chemistry 2009; 81(1): 193–202. 12. Banerjee S, Madhusudanan KP, Chattopadhyay SK, Rahman LU and Khanuja SP. Expression of tropane alkaloids in the hairy root culture of Atropa acuminate substantiated by DART mass spectrometric technique. Biomedical Chromatography 2008; 22(8): 830–834. 13. Yew JY, Cody RB and Kravitz EA. Cuticular hydrocarbon analysis of an awake behaving fly using direct analysis in real time time-of-flight mass spectrometry. Proceedings of the National Academy of Science USA 2008; 105(20): 7135–7140.

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Index

Bile pigments, 1, 9–12, 15, 191, 195, 198, 201, 208, 210, 212, 217 Bilin biosynthesis, 11, 13 Bilirubin, 10, 11, 15, 173, 192–198, 201–210 Bilirubin IIIα, IXα and XIIIα isomers, 191–193 Bilirubin albumin complex, 202 Bilirubin conjugates, 173, 194 Bilirubin monoglucuronide, 208 Bilirubin diglucuronide, 196, 207, 208 Bilirubin UDPglucuronosyltransferase, 10 Biliproteins, 16, 217 Biliverdin, 1, 9, 10–12, 195–198, 208–217, 224 Biliverdin IXα, IXβ, IXγ and IXδ isomers, 196 Biliverdin reductase, 10 Biliviolins, 11, 198

Acute intermittent porphyria (AIP), 109, 119, 120, 122, 144 Allophycocyanin, 218 Aminolaevulinic acid (ALA), 3, 5, 107–109, 113, 115–117, 119 Aminolaevulinic acid dehydratase (ALA-D), 5, 109, 113–116 Aminolaevulinic acid dehydratase deficiency porphyria (ADP), 113 Aminolaevulinic acid synthase (ALA-S), 3, 8 Allomerisation, 179, 182, 187 Allomers, 184, 187 Atmospheric pressure chemical ionisation (APCI), 51, 57, 185, 186 Atmospheric pressure ionisation (API), 51 Atmospheric pressure photoionisation (APPI), 51, 57 Avian eggshell pigments, 212

C5 pathway, 3 Capillary electrophoresis (CE), 118, 159, 162, 166, 168, 169

Bacteriochlorin, 97, 98, 102 Bacteriochlorophyll, 187 227

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Capillary HPLC, 224 Capillary zone electrophoresis (CZE), 115, 118 Capillary zone electrophoresis/mass spectrometry (CZE/MS), 115, 118 Chlorophylls, 13, 15, 179–181, 183, 185, 186, 223 Chlorophyll a allomerisation, 182 Chlorophyll a allomers, 184, 187 Chlorophyll biosynthesis, 2, 8 Chlorophyll catabolites, 183 Chlorophyll degradation, 188 Chlorophyllides, 179 Congenital erythropoitic porphyria (CEP), 54, 55, 109, 122–127 Coproporphyrin, 4, 26, 27, 29, 34–36, 55, 59, 60, 89, 92, 93, 95, 97, 107, 108, 113, 123, 124, 132, 134–140, 144, 155, 162 Coproporphyrin I, II, III, IV isomers, 34, 35 Coproporphyrinogen, 6–8, 42, 45, 107, 109, 132, 135, 136, 138, 140, 142, 143 Coproporphyrinogen I, II, III, IV isomers, 42, 45 Coproporphyrinogen oxidase (Coprogen-O), 7, 109, 132, 135, 138, 140, 142, 143 Coupled oxidation of haem, 196 Crigler-Najjar syndrome, 202

Desorption electrospray ionisation (DESI), 224 Direct Analysis in Real Time (DART), 224, 225 Deuteroporphyrin, 4, 29, 34, 44, 149 Dicarboxylic porphyrins, 32, 44, 160 Dihydrobiliverdin, 12 Dihydrogeranylgeraniol (DHGG), 181 Dihydroxy-mtetra(hydroxyphenyl)chlorin, 98, 101, 102 Dihydroxyurochlorins, 75 Dubin-Johnson syndrome, 202, 203 Eggshell pigment extraction, 213, 214 Electrochemical detection, 50 Electrospray ionisation mass spectrometry (ESI-MS), 56, 57, 60, 62, 97 Epoxyurochlorins, 75 Erythrocyte porphyrin extraction, 112 Erythropoietic protoporphyria (EPP), 109, 119, 120, 146–148 Faecal porphyrin extraction, 110 Faecal porphyrin profiles, 123 Ferrochelatase, 8, 109, 146, 147, 149 Fischer’s numbering system, 2

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Index

Fluorescent chlorophyll catabolites, 183 Formylcoproporphyrin, 93 Formylheptacarboxylic porphyrin, 89, 90 Fourier Transform ion cyclotron resonance mass spectrometry (FTICR-MS), 57 Geranylgeranyol (GG), 181 Glucose identification by CE, 172 Glucoside of protoporphyrin, 156, 173 Glycoconjugates of protoporphyrin, 159 Haematoporphyrin, 32, 33 Haem biosynthesis, 5, 7, 8, 107, 109 Haem cleavage, 196 Half-stercobilin, 198, 200, 201 Harderian gland porphyrins, 155, 159, 162, 224, 225 Harderoporphyrin, 138, 155, 162, 173 Hepatoerythropoietic porphyria, 127 Heptacarboxylic porphyrins, 37, 86, 130, 131 Heptacarboxylic porphyrinogens, 46 Hexacarboxylic porphyrins, 26, 27, 29, 34, 36, 39, 55, 59, 60, 119, 124, 127

229

Hexacarboxylic porphyrinogens, 46 Homozygous coproporphyria, 173 Hydrophobic interaction retention mechanism, 34, 44 Hydroxychlorophyll a, 180, 181, 186, 187 Hydroxycoproporphyrins, 136 Hydroxymethylbilane (HMB), 5, 6, 109, 119, 121 Hydroxymethylbilane synthase (HMB-S), 5, 6, 109, 119–122 Hydroxyheptacarboxylic porphyrins, 86 Hydroxymethylcoproporphyrin, 93, 94 Hydroxypentacarboxylic porphyrins, 130 Hydroxypheophytin a, 181 Hydroxyurochlorins, 75 Hydroxyuroporphyrins, 65, 67, 71, 80, 125 Hyperbilirubinaemia, 201, 202, 205–207 Ion trap, 51 Isocoproporphyrin, 4, 131–133, 140 IUPAC nomenclature, 3 Jaundice, 201–203 Ketoacetic acid uroporphyrin, 80, 81, 84, 125, 126, 128 Ketoacid heptacarboxylic porphyrins, 86

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Ketoacid uroporphyrins, 80 Ketopropionic acid coproporphyrin, 93, 95, 132, 136–139 Ketopropionic acid uroporphyrin, 80–83, 93, 125, 126

Micellar electrokinetic capillary chromatography (MEKC), 114–116 Mixed porphyria, 149 Monosaccharides separation by CE, 169, 172

Laser desorption/ionization timeof-flight (LDI-TOF), 52–56, 58 Lead poisoning, 113, 120, 146 Light harvesting complex, 16 Liquid secondary ion mass spectrometry (LSIMS), 52, 53, 58 Liquid surface ionization mass spectrometry (LSIMS), 52, 53, 58

Non-aqueous reversed-phase HPLC of bile pigments, 191 Non-aqueous reversed-phase HPLC of chlorophylls, 177, 181 Non-fluorescent chlorophyll catabolites, 183 Normal phase HPLC of chlorophylls, 181, 183

Magnesium chelatase, 10 Magnesium purpurin-7 dimethyl phytyl ester, 182–184, 187 Matrix assisted laser desorption/ionization time-of flight (MALDI-TOF), 53, 56 Matrix effect, 58, 162 Mesoporphyrin, 4, 27, 29, 34, 39, 40, 44, 60, 119, 120, 142, 143, 145, 147–149 Metalloporphyrins, 31, 38, 44 Meta-tetra(hydroxyphenyl)chlorin (m-THPC), 97–102 Methoxychlorophyll a, 180, 186, 187 Methoxylactone chlorophyll a, 180, 186

Open-chain tetrapyrroles, 9 Orbitrap mass spectrometer, 223 Oxidative ring opening, 14 Oxygenated derivatives of uroporphyrin, 124 Pentacarboxylic porphyrins, 45, 130, 132 Pentacarboxylic porphyrinogens, 44, 45, 131–135 Peptide conjugates of protoporphyrin, 31 Pheophorbides, 180, 186 Pheophytins, 179, 186, 187 Photochemical oxidation of uroporphyrinogen, 61 Photobilirubins, 208–210 Photodynamic therapy (PDT), 16, 97

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Index

Photosensitisers, 16, 97 Phycobilins, 11, 217 Phycobiliproteins, 16, 217 Phycocyanin, 16, 217, 218 Phycocyanobilin, 11–13 Phycoerythrin, 218 Phycoerythrobilin, 11, 12 Phytochrome, 12, 15 Phytochromobilin, 11–13, 15 Plasma porphyrin extraction, 111 Porphobilinogen (PBG), 5, 6, 107–109, 113–122, 144 Porphobilinogen deaminase (PBGD), 5, 6 Porphyrias, 25, 108, 109, 113, 122, 127 Porphyria cutanea tarda (PCT), 54, 55, 109, 127–135, 149 Protochlorophyll, 181–183 Protoporphyrin, 1, 4, 7, 8, 10, 15, 27–29, 31, 34, 39, 40, 44, 56, 60, 93, 96, 107–109, 112, 119, 120, 123, 138, 140, 142–147, 155, 156, 158–164, 166–173, 212–216 Protoporphyrin-amino acid conjugates, 31 Protoporphyrin-peptide conjugates, 31 Protoporphyrin glucoside, 173 Protoporphyrin glycoconjugates, 31, 155, 159, 161–163, 173, 174 Protoporphyrin monomethyl ester, 166, 167

231

Protoporphyrin xyloside, 166, 173 Protoporphyrinogen oxidase (Protogen-O), 7, 8, 109, 143–145, 147 Pyropheophytin a, 186 Pyrrole, 1, 2, 6, 75, 102, 193 Quadrupole time-of-flight (Q-TOF), 51, 60 Red blood cell porphyrin extraction, 112 Retention behaviour of porphyrins, 34 Rotor’s syndrome, 202 Shemin pathway, 3 Sirohaem, 5, 15 Stercobilin, 198, 200 Transmethylation of glycoconjugates, 159 Tricarboxylic porphyrin, 60, 132, 138, 140, 155, 156, 160, 162 Ultra-performance liquid chromatography (UPLC), 223, 224 Urinary porphyrin extraction, 109, 110 Urobilinoids, 198, 200 Urobilinogen, 11 Uroporphyrin, 1, 2, 4, 26–32, 34, 37, 41–43, 46, 47, 55, 59–63, 65–67, 69–73, 80–84, 86, 89, 93, 107, 121–128

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Uroporphyrin isomers, 32, 41, 42, 47 Uroporphyrinogens, 47, 119 Uroporphyrinogen decarboxylase (Urogen-D), 6, 7, 109, 120, 127, 133, 135 Uroporphyrinogen III synthase (Uro III-S), 5, 6, 109, 120, 123, 127 Vanadyl porphyrins, 57

Variegate porphyria (VP), 109, 122, 143, 144, 149 Violins, 198 Vitamin B12, 5, 15 Xylose identification by CE, 166 Xyloside of protoporphyrin, 166, 173 Zinc-biliverdin, 212–214, 224 Zinc-protoporphyrin, 44, 112, 113, 119, 146, 212–214