Vitamins [Reprint 2012 ed.] 9783110859188, 9783110102444

Table of contents :
1 Introduction
2 Vitamin A and Its Provitamins
3 Vitamin D
4 Vitamin E
5 Vitamin K
6 Thiamin, Vitamin B1, Aneurin
7 Vitamin B2: Riboflavin and Its Bioactive Variants
8 Niacin: Nicotinic Acid, Nicotinamide, NAD (P)
9 Vitamin B6
10 Folic Acid and Unconjugated Pteridines
11 Biotin
12 Pantothenic Acid
13 Vitamin B12
14 Vitamin C
15 Literature Supplement
16 Subject Index

Citation preview

Wilhelm Friedrich Vitamins

Cover illustration Vitamin Β, Thiamin hydrochloride With kind permission of Hoffmann-La Roche AG, Grenzach-Wyhlen

Wilhelm Friedrich

Vitamins

w DE

Walter de Gruyter

G Berlin · New York 1988

Author Professor Dr.-Ing. Wilhelm Friedrich Institut für Physiologische Chemie Universität Hamburg 2000 Hamburg 13 FRG

private address Brandheide 2 2000 Hamburg 65 FRG

Library of Congress Cataloging-in-Publication Data Friedrich, W. (Wilhelm), 1913Vitamins. Bibliography: p. Includes index. 1. Vitamins. I. Title. QP771.F77 1988 612'.399 ISBN 0-89925-273-7 (U.S.)

88-3807

CIP-Kurztitelaufnahme der Deutschen Bibliothek Friedrich, Wilhelm: Vitamins / Wilhelm Friedrich. - Berlin ; New York : de Gruyter, 1988 ISBN 3-11-010244-7

Copyright © 1988 by Walter de Gruyter & Co., Genthiner Straße 13, 1000 Berlin 30, FRG Walter de Gruyter, Inc., 200 Saw Mill River Road, Hawthorne, N.Y. 10532 All rights reserved, including those of translation into foreign languages. N o part of this book may be reproduced in any form - by photoprint, microfilm or any other means nor transmitted nor translated into a machine language without written permission from the publisher. The quotation of registered names, trade names, trade marks, etc. in this book does not imply, even in the absence of a specific statement that such names are exempt from laws and regulations protecting trade marks, etc. and therefore free for general use. Cover design: Hansbernd Lindemann, Berlin. - Typesetting: Appi, Wemding. - Printing: Gerike GmbH, Berlin. Binding: Lüderitz & Bauer, Berlin. - Printed in Germany.

Motto

Discovery of the vitamins and their life saving value in the prevention and cure of nutritional deficiency diseases is one of the most important contributions of biochemistry to medicine and human welfare A. Lehninger, Principles of Biochemistry Worth Publishing, New York, 1982

Dedication

To my brothers: Henry, assistant professor at the University of Warsaw, and Richard, teacher at the Gymnasium M. Reya in Warsaw, victims of the Second World War.

Foreword

In the introduction, the author has attempted to convey to the reader some of the basic information on vitamins; the largely tabular format of this chapter should simplify the reading. It is followed by the special chapters on the 13 vitamins. Finally, in the appendix, the author has made an effort to inform the reader of the more important publications, especially those relevant to medicine, which appeared after the corresponding chapters had been written. This list extends to the beginning of May, 1986. The author would not wish to fail to thank the Publisher, Walter de Gruyter, especially Dr. Ing. Rudolf Weber and Dr. Mary Eagleson, for their exemplary cooperation. Thanks to the financial support of the Verband der Chemischen Industrie, it was possible to buy copies of many other references. The author is also grateful to Hoffmann-La Roche AG, Basel and Grenzach, for providing him literature on vitamins, most of it difficult to obtain, for many years. Finally, gratitude is due the libraries of the University of Hamburg for buying literature and to the many donors of reprints from all over the world. Wilhelm Friedrich

Table of Contents

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Introduction Vitamin A and Its Provitamins Vitamin D Vitamin E Vitamin Κ Thiamin, Vitamin B u Aneurin Vitamin B2: Riboflavin and Its Bioactive Variants Niacin: Nicotinic Acid, Nicotinamide, NAD (P) Vitamin B6 Folic Acid and Unconjugated Pteridines Biotin Pantothenic Acid Vitamin B12 Vitamin C Literature Supplement Subject Index

3 63 141 219 285 339 403 475 543 619 753 807 837 929 1003 1021

1 Biochemistry and Physiology of the Vitamins: Introduction

1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.7.1 1.7.2 1.7.3 1.7.4 1.7.5 1.7.6 1.8 1.9 1.10 1.10.1 1.10.2 1.10.2.1 1.10.2.2 1.10.2.3 1.10.2.4 1.10.2.5 1.10.2.6 1.11 1.11.1 1.11.2 1.12 1.13 1.14 1.15 1.16 1.17 1.18 1.19 1.20

Historical Classification, Nomenclature and Structure Physical and Chemical Properties Biosynthesis Commercial Production Vitamin-Containing Preparations and Their Stability Analysis Use of Animals Microbiological Tests Physical and Chemical Methods Enzymatic Methods Evaluation of the Human Vitamin Status Determination of the Biological Activity of Fat-Soluble Vitamins Occurrence, Especially in Foods Enrichment of Foods Vitamin Requirements Recommendations for Human Vitamin Intake Elevated Vitamin Requirements in Human Beings; Vitamin Megadoses as Therapeutics Cancer and Viral Diseases Age Effects of Medication Pregnancy, Delivery Renal Insufficiency Other Situations in Which Vitamin Requirements Are Elevated Absorption, Transport, Distribution, Storage and Excretion Absorption from the Digestive Tract Transport into the Organs and Tissues; Distribution, Storage and Excretion Biological Functions Vitamin Status in Some Population Groups Vitamin Deficiency: Causes and Symptoms Hypervitaminosis, Toxicity Vitamin-Responsive Congenital Protein Defects Synergism and Antagonism of the Vitamins Vitamin Antagonists (Antivitamins) Compounds with Questionable Vitamin Character References

1 Biochemistry and Physiology of the Vitamins: Introduction

This introductory chapter is similar in its outline to the following special chapters, which deal with the individual vitamins. Here, however, the main emphasis is to compare the vitamins and their effects and to indicate interactions among them (synergism, antagonism)1. In this chapter there is more emphasis on tables than on lengthy text. The vitamins are organic, low-molecular weight components of the diet, which act essentially only as catalysts, required by the organism in very small amounts for normal development. (However, in principle, they are not structural components of the body.) They are either not synthesized by the organism, or not in sufficient quantity, and must therefore be ingested with the diet or acquired in some other way. A vitamin deficiency leads to illness which is cured by administration of the vitamin. The above definition distinguishes the vitamins from the essential fatty acids and amino acids, which must be ingested in larger amounts and serve as structural components of the body. However, the distinction is not entirely clear-cut. It is possible that all the vitamins required by human beings for their development are known. Years of experience have shown that patients who are fed entirely parenterally (i. e. receive intravenous doses of known nutritional factors) do not develop deficiency symptoms [Herbert (1980)]. The vitamins are not a chemically uniform group.

1.1 Historical The development of the modem concept of vitamins can be roughly divided into three (broadly overlapping) periods2 [Wagner & Folkers (1964)]: 1) Empirical healing of some diseases by administration of certain foods; 2) experimental induction of dietary diseases in animals; and 3) administration of synthetic diets to discover essential nutritional factors. 1

2

Reviews: Stepp, Kühnau & Schroeder (1957); Wagner & Folkers (1964); Sebrell & Harris (1967-1972); Marks (1975); Willemsen (1977); DeLuca (1978); Goodhart & Shils (1980); Vitamin Compendium (1970, 1980); McCormick & Wright (1970/1971, 1979/1980); Ammon & Dirscherl (1974,1975,1981); Briggs (1981); Bässler & Lang (1981); Isler & Brubacher (1982); Alfin-Slater & Kritchevsky ((1980); Ulimanns Encyklopädie der technischen Chemie (1983); Kirk-Othmer Encyclopedia of Chemical Technology (1984); Briggs (1984); Machlin (1984a); Hanck & Hornig (1985); Marks (1985) Reviews: Wagner & Folkers (1964); Tschesche (1967); Morton (1968); Lieben (1970); György (1976) ; Sharman (1977) ; Snell (1979) ; Rosie (1960) ; Isler & Brubacher (1982); Bechtel (1984)

4

1 Biochemistry and Physiology of the Vitamins

The first phase began many centuries ago, and gradually led to the recognition that night blindness, scurvy, beriberi and rickets are dietary diseases. The ancient Greeks, Romans and Arabs healed night blindness with liver. In the 16th century, scurvy was healed using extracts of spruce needles, and in the 18th century, oranges and lemons were used (J. G. H. Kramer, 1720); the issue of lemon juice to sailors in the English navy was required by law by the beginning of the 19th century. At the end of the 19th century, the Japanese navy realized that beriberi was in some way connected with a diet of rice; the disease was successfully combatted by replacing the rice with barley or by increasing the rations of meat and vegetables. In the same period, it was learned that rickets could be prevented and cured by administration of fish liver oil. The second phase was characterized by the use of experimental animals, in which dietary diseases were deliberately induced and then healed. The animal experiments made possible systematic study of diet-related diseases in human beings and animals. In 1890, C.Eijkman discovered that polished rice, when given to chickens as their main food, caused polyneuritis (degeneration of the peripheral nerves). This disease is similar to human beriberi. Eijkman then demonstrated that non-polished rice, or polished rice + rice bran, when given as feed, healed the polyneuritis. Extracts from the rice bran had the same effect. These experiments led to the postulate that beriberi was caused by the lack of a nutritional substance. Similarly, in 1907, A. Hoist and T. Fröhlich reported experimentally induced scurvy in guinea pigs, and the healing of the condition by the same agents which are effective in human beings. The third phase consists of the search for essential dietary factors. For this, pure nutritional components are fed to experimental animals. As early as 1881, the biochemist N.I.Lunin found that a highly purified diet, consisting only of proteins (e. g. milk protein), carbohydrates and fats, salts and water, is not sufficient for the animal. Lunin postulated the presence (e.g. in fresh milk) of small amounts of unknown substances. W. Stepp showed in 1909 that extracted milk and bread are insufficient as animal feed, because animals fed this way die. The animals survived, however, if the extracts were administered to them separately. Thus around 1900, the idea had been widely accepted that human beings and animals require very small amounts of additional nutritional factors ("accessory growth factors"), the lack of which leads to diseases. F. G. Hopkins reported on this subject in detail in 1912. In the same year, C.Funk coined the term "vitamin". Funk experimented in the area of dietary diseases, and worked on the isolation of the anti-beriberi factor. Because he assumed that this and the other curative factors contain nitrogen, that is, are amines, he named them "vitamines" (vital amines). In 1929, C. Eijkman and F. G. Hopkins shared the Nobel Prize in Medicine for their discoveries in the area of vitamins. Eijkman's discovery that polished rice leads to polyneuritis in birds and beriberi in buman beings prepared the way for the isolation of thiamin. Hopkins showed, by experiments on rats, that milk contains substances which, in very small amounts, make growth possible. In this way he extended the insights of Lunin and confirmed them with more complete experiments. Almost none of the vitamins is a single substance. Instead, there are whole

1.1 Historical

5

Table 1-1. History of the vitamins [Isler & Brubacher (1982)] Vitamin

First isolated

Discovery

Isolation

Structure elucidated

Synthesis

Vitamin A Provitamin A

Fish liver oil Carrot, palm oil Fish liver oil, yeast Wheat germ oil Alfalfa

1909

1931 1831

1931 1930

1947 1950

1918

1932

1936

1959

1922 1929

1936 1939

1938 1939

1938 1939

Rice bran Egg albumin Liver Rice bran Liver, fermentation Liver Liver

1897 1920 1936 (1894) 1934 1926

1926 1933 1935 (1911) 1938 1948

1936 1935 1937 1938 1956

1936 1935 1894 1939 1972

1941 1931

1941 1938

1946 1940

1946 1940

Liver

1931

1935

1942

1943

Adrenal cortex 1912 Lemon

1928

1933

1933

Vitamin D Vitamin E Vitamin Κ Vitamin Vitamin Niacin Vitamin Vitamin

B, B2 B6 B12

Folic acid Pantothenic acid Biotin Vitamin C

families of close chemical relatives which, however, may differ greatly in their biological activity. The substances which we call vitamins must usually be converted within the living cell to their coenzyme or hormone forms in order to become biologically active. For example, vitamin D is hydroxylated, and aquacobalamin is methylated or adenosylated. Vitamin deficiencies were formerly one of the main causes of illness and death. Pellagra, scurvy and beriberi are the best known vitamin deficiency diseases. In the past they decimated whole populations and affected the outcomes of wars and voyages of discovery. Somewhat less well known are the effects of pernicious anemia, rickets and xerophthalmia [Marks (1975)]. An outline of the history of the vitamins is presented in Table 1-1. Today the vitamins are readily available and their mechanisms of action have been thoroughly investigated. Thanks to this development, the vitamin deficiency diseases have become rare in many countries. Some of the sources of vitamins are properly preserved fruit juices, fish liver oil, enriched flour and margarine, the latter enriched with vitamins A, D and E, as well as essential fatty acids. In this way it has become nearly comparable to butter. The vitamins are now produced in large amounts, mostly synthetically. In many countries, more vitamins are consumed than is recommended [Morton (1976)].

6

1 Biochemistry and Physiology of the Vitamins

Table 1-2. Vitamins and provitamins and their most important forms [Isler & Brubacher (1982)] Vitamin group

Most important representative

Important active compounds

Most important commercial forms

Vitamin A

Retinol

Provitamin A

ß-Carotene

Vitamin A acetate, vitamin A palmitate, vitamin A acid ß-Carotene ß-apo-8'-carotenal, ß-apo-8'-carotinic esters

Vitamin D

Cholecalciferol

Vitamin E

α-Tocopherol

Vitamin Κ

Phylloquinone

Retinol, retinal, retinoic acid, 3-dehydroretinol (A2) α-, β- and γ-carotene, ß-apocarotenoids, cryptoxanthine, echinenone Ergocalciferol cholecalciferol α-, β-, γand δ-tocopherol α-, β-, γand δ-tocotrienol Phylloquinone (KO Menaquinone (K2) Menadione (Menaquinone O)

Vitamin Bi

Thiamin

Thiamin and its salts; thiamin diphosphate (cocarboxylase)

Thiamin chloride hydrochloride, thiamin mononitrate thiamin diphosphate (carboxylase)

Vitamin B2

Riboflavin

Niacin

Nicotinamide

Vitamin Be

Pyridoxine

Riboflavin, riboflavin sodium phosphate Nicotinamide, nicotinic acid Pyridoxine hydrochloride, pyridoxal 5'-phosphate (codecarboxylase)

Vitamin Bi2

Cyanocobalamin,

Folic acid

Folic acid

Pantothenic acid

Pantothenic acid

Biotin

d-Biotin

Riboflavin, riboflavin phosphoric acid Nicotinamide, nicotinic acid Pyridoxine, pyridoxal, pyridoxamine, pyridoxal 5'-phosphate (codecarboxylase) Cyanocobalamin, hydroxocobalamin Folic acid, folic acid conjugates Pantothenic acid Pantetheine Panthenol d-Biotin

Vitamin C

Ascorbic acid

Ascorbic acid Dehydroascorbic acid

Ascorbic acid Sodium ascorbate Calcium ascorbate

* More correctly, RRR- and all-rac-α-Tocopherol ** More correctly, RRR- and a/Z-rac-a-Tocopheryl acetate

Vitamin D2, vitamin D3 [d]- and [dl]-aTocopherol* [d]- and [dl]-atocopherol acetate** Vitamin K4 Menadione Menadiol esters

Cyanocobalamin, hydroxocobalamin Folic acid

Calcium pantothenate Sodium pantothenate d-Biotin

1.4 Biosynthesis

7

1.2 Classification, Nomenclature and Structure The vitamins are divided into two groups. One includes the fat-soluble vitamins A, D, E and K, and the other, the water-soluble vitamins thiamin (BO, riboflavin (B2), niacin (B6), folic acid, biotin, pantothenic acid, B12 and C. Because of the differences in their water solubility, these two groups of vitamins differ in some of their basic biological properties. Water-soluble vitamins are poorly stored and intake in excess of the requirement generally leads to their excretion in the urine. Fat-soluble vitamins can be stored, and if they are consumed in excess, hypervitaminoses may result, especially in the cases of vitamins A and D. Before the chemistry of the vitamins was known, the practice was introduced of denoting the vitamins with capital letters. These are still used, usually for the entire group of compounds. Later trivial names were invented, often several for a single substance; many of these are no longer used. Not all of the original isolates were later found to be uniform substances, so that there are gaps in the alphabetical series. The current terminology for the vitamins is shown in Table 1-2; see also "Nomenclature Policy", J. Nutrition 111 (1981) 8. There are many known structural analogs of the vitamins which have no vitamin effect, and are often antagonists of the vitamins. They are often used in animal experiments to generate defined vitamin deficiency states. Sometimes they have clinical uses. Some examples are 4-hydroxycoumarins, antagonists of vitamin K; pyrithiamin and oxythiamin, antagonists of thiamin; isoniazide, antagonist of vitamin B6; methotrexate, antagonist of folic acid; dehydrobiotin, antagonist of biotin.

1.3 Physical and Chemical Properties The vitamins, especially the water-soluble ones, are a very heterogeneous group of substances. The fat-soluble vitamins have at least some common traits in their biosyntheses, in that they are entirely or primarily isoprene derivatives. (Only vitamins E and Κ also contain non-isoprenoid groups in their structure.) The water solubilities of many vitamins are shown in Table 1-3. The structural formulae and the corresponding space-filling models of the vitamins are shown in Fig. 1-1 (pp. 9).

1.4 Biosynthesis The synthesis of the fat-soluble vitamins starts from acetyl-CoA. Isopentenyl diphosphate formed via mevalonic acid condenses with other C-5 units along the usual lines of terpene biosynthesis, via geranyl diphosphate (with 10 C atoms) and farnesyl diphosphate (with 15 C atoms) to geranylgeranyl diphosphate (with 20 C atoms). The path to vitamin D leads via dimerization of farnesylfarnesyl diphosphate to the C-30 hydrocarbon squalene, which is converted to cholesterol, 7-dehydrocholesterol and finally vitamin D. Geranylgeranyl diphosphate is

8

1 Biochemistry and Physiology of the Vitamins

Table 1-3. Solubility of the vitamins in water (in mg/ml) at 25 °C [DeRitter (1982)] Vitamin

Solubility

Thiamin hydrochloride Thiamin mononitrate Riboflavin Riboflavin 5'-phosphate, Na + salt Nicotinamide Pyridoxine hydrochloride Folic acid Biotin Calcium pantothenate Panthenol Cyanocobalamin Ascorbic acid Sodium ascorbate

1000 27 0.066-0.33 43-112 1000 220 0.0016 0.3-0.4 356 infinitely soluble 12.5 333 620

the precursor of vitamins A, E and K. Dimerization of geranylgeranyl diphosphate leads to colorless Carotinoids ; dehydrogenation and cyclization of the ends of the Carotinoid chains produces the carotenes, which are the direct precursors of vitamin A. The biosynthesis of vitamins E and Κ requires the conversion of geranylgeranyl diphosphate to phytyl diphosphate. This is condensed with previously formed benzohydroquinone or naphthohydroquinone components, and then methylated [Isler & Brubacher (1982)]. The biosyntheses of the water-soluble vitamins do not share so many common steps as those of the fat-soluble vitamins (the reader is directed to the corresponding chapters). The animal organism is able to convert pre-synthesized vitamins into the biologically active hormone or coenzyme forms. The evolution of the biosynthesis of vitamin coenzymes and the role of the coenzymes in the origin of life are comprehensively treated in a review by King [King (1980)]. Several vitamins, including thiamin, folic acid, niacin, B6 and B12, are synthesized and excreted by bacteria of the genus Bifidobacterium. These microorganisms make up a large fraction of the human intestinal flora, so their production of vitamins is of interest [Deguchi et al. (1985)].

1.5 Commercial Production All vitamins are now produced commercially. Through improvements in the methods, the prices of vitamins have dropped to a fraction of the 1945 prices. As a result, vitamins are readily available not only for human but for animal consumption. The addition of vitamins accounts for only 0.6-1.0% of the total cost of the feeds [Isler & Brubacher (1982)]. The commercial production of vitamins is primarily by chemical synthesis. Fat-soluble vitamins are also commercially isolated from natural sources, e.g.

1.5 Commercial Production

Vitamin A - Palmitate

Provitamin A

ß-Caroteiie

Vitamin D 3 (Cholecalciferol)

9

10

1 Biochemistry and Physiology of the Vitamins

[dl| - a-Tocopherolacetate

Vitamin K, (Phylloquinone)

^

Ä

• * J F

f;

OH

OH

=

fO °

HO-C-H

CH2OH

L-Ascorbic Acid

1.5 Commercial Production

—CHi—Ν

f

1

%

1

J—CI

l^jLo

C-l^J-HH,

*

H

l *

Thiamin hydrochloride

Vitamin B2

CH,—(CHOH),—CH,«

Riboflavin

^ ^ ^

Pyridoxine hydrochloride

—*

^ ^ ^

12

1 Biochemistry and Physiology of the Vitamins

Vitamin B12

é *

• .·

Cyanocobalamin

-CHi—CH¡—CHi—COOH

φ

*

D-Biotin

Folic Acid

xf

Folic Acid

¿ Q - O - j Ν

TT

MT

% « jm^

Cl

1.5 Commercial Production

13

Nicotinic Acid S

f

NH2

è/ •

è

Nicotinamide

Pantothenic Acid

CHTOH—CtCH3)2—CHOH—CO—NH—CH2—CH2—R

Calcium - D ( + ) - pantothenate Fig. 1-1. Structural formulae and space-filling models of the vitamins (with kind permission of Hoffmann-La Roche AG, D-7889 Grenzach-Wyhlen)

vitamin A from fish liver, vitamin E from soy oil, vitamin Κ from fish meal and vitamin D from liver oil or irradiated yeast [Fryth (1984)]. Microorganisms can be used to produce thiamin, riboflavin, folic acid, pantothenic acid, biotin and vitamin B6. Vitamin B12 is commercially produced only from microorganisms. In the synthesis of ascorbic acid, only the oxidation step from D-sorbitol to L-sorbose is done microbiologically, by Acetobacter suboxydans. Aside from vitamin B12, only riboflavin can be produced by fermentation on a technical scale at a competitive price. The microbial production of ß-carotene is not economical compared to the technical synthesis. However, in Japan other vitamins as well are produced microbiologically [Crueger & Crueger (1984)]. Since some vitamins are also used in their coenzyme forms, efforts have been made to develop methods for extracting these. The preferred approach is to use microorganisms which have been provided chemically synthesized precursors. This method has the advantage over de novo biosynthesis that it bypasses some control mechanisms which inhibit the biosynthesis [Sakai (1980)]. Some microor-

14

1 Biochemistry and Physiology of the Vitamins

Table 1-4. Microbial production of vitamin coenzymes [Sakai (1980)] Microorganism

Coenzyme precursor

Products and Yields

Sorcina lutea Sorcina marginata Saccharomyces logos Saccharomyces marxianus Saccharomyces carlsbergensis Brewer's yeast

FMN, AMP FMN, AMP Nicotinamide, adenine Nicotinamide, adenine Nicotinamide, adenine Pantothenic acid AMP, cysteine Pantothenic acid AMP, cysteine Pantothenic acid, ATP, cysteine Pyridoxine

FAD, 66.5 μg/ml FAD, 38.5 ng/ml NAD, 3.78 mg/g dry cells

Baker's yeast Brevibacterium ammoniagenes Aspergillus oryzae

NAD, 4.08 mg/g dry cells NAD 8.5 mg/g dry cells Coenzyme A, 100 μg/ml Coenzyme A, 128 μg/ml Coenzyme A, 830 μg/ml Pyridoxine phosphate, 7.1 μg/ml Pyridoxal phosphate, 0.9 μg/ml

Table 1-5. Quantitative development of vitamin production [Fryth (1984)] Year

Production in metric tons

1950 1960 1965 1970 1975 1979 1980

1,567 5,018 7,392 10,394 13,586 18,812 19,313

ganisms which carry out these biosyntheses and the coenzyme yields which can be obtained are shown in Table 1-4. The quantitative development of world vitamin production is shown in Table 1-5. The vitamins are used as additives to foods and feeds, as pharmacological preparations and as components of cosmetic products. The use of vitamins in pharmaceutical products is reviewed by DeRitter [DeRitter (1982)].

1.6 Vitamin-Containing Preparations and Their Stability Vitamins belong to a wide variety of chemical classes, and thus display widely varying physical and chemical properties 3 . Each vitamin compound can behave in a variety of ways, depending on the conditions. Properly stored, crystalline 3

Reviews : DeRitter (1982), Kläui (1979), Killeit (1985)

1.6 Vitamin-Containing Preparations and Their Stability

15

pure vitamins normally retain their full biological activity for years. However, during the course of production, storage and preparation of foods, they are exposed to a variety of physical and chemical factors which cause losses. Among the most important of these immediately effective factors are temperature, water content, pH value, oxygen, light, heavy metal ions (iron, copper), interactions with other vitamins and time. A large number of publications on the problem of vitamin stability has appeared; a review of the most important of them was prepared by Killeit [Killeit (1985)]. Some vitamins are relatively stable and do not present serious difficulties during storage and production of vitamin preparations: cholecalciferol, vitamin E acetate, acidic vitamin E succinate, biotin, niacin, pyridoxine and riboflavin. Retinol and its esters, vitamin K, ascorbic acid, folic acid, pantothenic acid, Panthenol and thiamin are relatively labile in pharmaceutical preparations. Double bonds in the molecules of fat-soluble vitamins often cause isomerization in pharmaceutical preparations. In aqueous, acid media, vitamin A undergoes a gradual change from the native all-trans form into a mixture consisting of about 66% alltrans and 33% cis forms, which are less active than the former. For this reason, it has become the practice to produce vitamin A palmitate as an equilibrium mixture of about 66% all-trans and 33% cis forms for incorporation into water-containing preparations ; this mixture, used as a component of multivitamin preparations, loses much less activity over a period of months than pure all-trans preparations. In the case of calciferol, the reversible isomerization in solution leads to an equilibrium mixture of calciferol and precalciferol. Ergocalciferol and cholecalciferol isomerize at about the same rate. Pulverized ergocalciferol isomerizes in the presence of some salts, e.g. CaS0 4 and CaHP0 4 ; the acidity of the salts acts as a catalyst for the isomerization. The isomers which have been identified include 5,6-/ra«s-ergocalciferol, precalciferol, isocalciferol and tachysterol. Oxidation of α-tocopherol with, e.g. nitric acid or ferric chloride, produces a-tocopheryl quinone, then further oxidation products. Under anaerobic conditions, α-tocopherol is stable to heat. Ascorbic acid and pyrogallol protect it from air oxidation. Light decomposes α-tocopherol. Heating thiamin solutions to 75 °C for 60 minutes in the presence of sodium nitrite leads to formation of elemental sulfur, thiochrome and 4-methyl-5-(ß-hydroxyethyl)thiazol. The cleavage of thiamin with sulfite or hydrogensulfite is very rapid at high pH. The thiamin in a hydrogensulfite-containing solution for parenteral infusion is completely destroyed within 24 h. At higher pH values, thiamin is basically unstable [DeRitter (1982)]. Studies on the stability of the active substances in human milk have shown that freezing offers good protection for the vitamins. After storage for three months, there was no significant loss of vitamins [Friend et al. (1983)]. The stability of the vitamins against ionizing radiation has been reviewed [Basson (1983)]. The effect of sterilizing irradiation and of storage time on the content of thiamin, riboflavin and vitamin A in rat feed was studied. As components of foods, most vitamins are very sensitive to radiation. Of the water-soluble vitamins, the most sensitive to sterilizing radiation is thiamin, and the most sensitive fat-soluble vitamin is vitamin A [Hani; sv et al. (1985)]. See also Table 1-6 on the stability of vitamins.

16

1 Biochemistry and Physiology of the Vitamins

Table 1-6. Stability of the vitamins to various effects [DeRitter (1982), Isler & Brubacher (1982), Kläui (1979), Vitamin Compendium (1980), Bässler & Lang (1981)]. L = labile, S = stable Vitamin

Oxygen

UV light Acid

Base

Heavy metal ions

ioo°c

% loss in cooking foods

A

L

L

L

S

L

s

10-30

D

L

L

L

(S)

L

L

slight

E

L

L

S

(S)

L

S without 0 2

50

Κ

(L)

L

S

L

L

S

s

L

L

L

30-50

L

L to sulfite

S

L

L

0-50

L

Max. stability at pH 3.5-4.0

Thiamin L (Β,) Ribofla- S vin (B2)

L, esp. in base

B6

S

L

B,2

s

L

c

S at pH > 3

L in solution with Cu, Fe

S at pH < 9 L

Folic acid

L

L

L at pH < 5

Biotin

S

L

S

L

(S)

s

s

L

L

L

L at pH < 5

(L)

L

Ester more stable than alcohol

about 20

Usually very stable L

L

0-90

Comments

Ester more stable than retinol

20-80

Niacin Ca Panto thenate

Reduction

Max. stability at pH 4.5-5.0 S in solid form

L

L with riboflavin

0-70

Relatively stable

0-30

Usually very stable

0-45

Max. stability at pH 6-7

For use in human medicine, various types of vitamin preparations are available. To produce aqueous dispersions of fat-soluble vitamins, one uses surface-active substances, such as incompletely esterified polyene alcohols. Vitamin A is much better absorbed in vivo when administered as an aqueous dispersion containing a detergent than as a solution in oil. Water-soluble vitamins can be used as aqueous solutions; this is also true of water-soluble forms of fat-soluble vitamins, e.g. menadiol diphosphate tetrasodium salt. Solutions of water-soluble vitamins for injection often contain 1) sodium ascorbate + folic acid + nicotinamide and 2) thiamin hydrochloride + riboflavin 5'-phosphate as sodium salt+Panthenol-I-pyridoxine + cyanocobalamine, separately. Complete injectable multivitamin preparations have also been described, but they do not keep well; the losses affect mainly vitamin A, thiamin, cyanocobalamin and ascorbic acid. Vitamin losses can also occur through adsorption of the infusion solution onto the walls of the container. For oral purposes, both individual vitamins and mul-

1.7 Analysis

17

tivitamin preparations are produced in the form of tablets, emulsions and solutions [DeRitter (1982)].

1.7 Analysis In the early days, practically only microorganisms, animals and even human beings (e.g. pernicious anemia patients in the analysis of B12) were used to determine vitamin activity4. After the vitamins had been purified, physical and chemical methods for their assay became available. The latter methods are now used practically exclusively for pure or highly concentrated substances. In recent years, new and refined analytical methods such as HPLC, microtechniques and automated procedures have been developed. These make it possible to carry out analyses on small blood samples in the clinic. Säuberlich has written a thorough review of the new analytical methods for thiamin, riboflavin, niacin, vitamin B6 and folic acid [Säuberlich (1984)]. The modern methods for vitamin analysis constitute an extensive field; this includes biological and microbiological assays, chromatographic methods, including automated versions, and special techniques for the individual vitamins [Augustin et al. (1985)]. So long as the vitamins had not been purified, their amounts had to be measured in units which corresponded to a certain biological effect. Internationally recognized standard preparations were usually available. Today all vitamins are available in pure form, and their dosages are usually given in units of weight or on a molar basis. 1.7.1 Use of Animals The use of animals to detect a vitamin deficiency disease or to test the curative effect of a vitamin-containing preparation was used by the first vitamin pioneers like Lunin, Eijkman, Holst, Fröhlich and Hopkins (cf. p. 4). Two experimental protocols are used in animal experiments. In the curative protocol, the animals are given a complete diet except for the vitamin to be tested. After a certain time, one observes deficiency symptoms such as reduced growth, skin lesions and the loss of function of certain organs. If the vitamin is now added to the diet, the deficiency symptoms disappear, in so far as they are reversible. In the prophylactic protocol, certain amounts of the vitamin to be tested are added to the diet from the beginning. Both protocols permit quantitative evaluation [Isler & Brubacher (1982)]. 1.7.2 Microbiological Tests The principle of the microbiological test was born at the end of the 19th century and beginning of the 20th, when it was recognized that certain factors are essential for the growth of various fungi [Mackowiak (1979)]. However, it was the clas4

Reviews: Strohecker & Henning (1966), Baker et al. (1980), Granade (1982), Augustin et al. (1985)

18

1 Biochemistry and Physiology of the Vitamins

sic experiments of E.E. Snell (1937-1940) that showed that various microorganisms, especially bacteria, are suitable for the determination of vitamins [Snell (1979)]. Microbiological tests played decisive roles in the first isolation of some vitamins, and are now an important component of clinical diagnostics. Today protozoa and photoflagellata are also used for vitamin determinations. Tetrahymena thermophila (formerly T. pyriformis) requires thiamin, riboflavin, vitamin B6, niacin and pantothenic acid for growth. All these vitamins can be determined using this microorganism (Table 1-7). In these tests, T. thermophila behaves similarly to rats or chicks, in that it is able to utilize conjugates of the vitamins. The vitamins required by Poteriochromonas stipitata (formerly Ochromonas malhamensis), Ochromonas danica and Euglena gracilis include thiamin, biotin and (except for O. danica) vitamin B12 [Augustin et al. (1985)]. The technical procedures for vitamin determinations have been described in detail [Kavanagh & Ragheb (1979), Augustin et al., eds. (1985)]. Some of the microorganisms which can be used in vitamin determinations and the limits of detection attainable with them are listed in Table 1-7. Table 1-7. The more important microbiological methods for vitamin determination and their limits of detection Test microorganism Vitamins and their limits of detection in ng/1 B,

B2

Bö

B,2

Folic Biotin Niacin Pantoacid thenic acid

ATCC No.4

Leuconostoc sp.1 62 62 12500 25 0.5 6250 2 9 Lactobacillus

Ö +

>Λ ν> ν-> ^•Tt·^

ο ο ο ο ο >η ^Ο NO Φ νο

ν> >f> t—

ο ο ο ο ο ο

io io mm

Ο © Ο ^ ^ ^

7->

© Ο Γ^

8 8 r-8 8© v->

s

r" Un

(Ν +

e

ooorjfNíN τ"' (Ν (Ν W"> τ-" τ-"

Ο

Ο +

ο 2-vitamin D3, 276; 1^4^5(OH>3 vitamin D3, 6 [O'Riordan et al. (1982)]. References: 1. Flury & Haldimann (1981); 2. O'Riordan et al. (1982); 3. Mawer (1982); 4. Koshy (1982); 5. Norman (1982); 6. Dokoh et al. (1982); 7. Shinki et al. (1983); 8. Lambert et al. (1982b); 9. Holmes & Kummerow (1983); 10. Bouillon (1983b)

Table 3-9. Average serum concentrations [in nmol/liter] of the most important vitamin D metabolites and their variation through the year [Tjellesen & Christiansen (1983)] Month

24,25(OH)2D

l,25(OH)2D

25(OH)D2

25(OH)D3

25(OH)D

February April June August October December

3.8 5.5 7.2 7.2 4.8 4.6

0.061 0.064 0.071 0.057 0.074 0.072

9.0 4.0 4.5 4.0 7.3 11.8

52.3 67.5 88.3 77.8 67.3 58.5

61.3 71.5 92.8 81.8 74.5 70.3

162

3 Vitamin D

25(OH)-vitamin D2 in 758 healthy Japanese, most of them adults, had the following results [Kobayashi et al. (1983)]: 1.The average concentration of 25(OH)-vitamin D (e.g. 25(OH)-vitamin D2 + 25(OH)-vitamin D3) was 23.8 ng/ml. 2. The average concentration of 25(OH)-vitamin D3 was 23.0 ng/ml (range 3.0-65.3 ng/ml). The plasma level varied with the season, with higher values in summer than in winter; in addition, there was a strong correlation of the plasma level of 25(OH)D3 with the UV content of the sunlight. This shows clearly that the plasma 25(OH)-vitamin D3 is mainly of endogenous origin, and is synthesized from the 7-dehydrocholesterol in the skin. 3.25(OH)-Vitamin D2 was found in only 138 (18.2%) of the subjects, and its average concentration was 4.4 ng/ml (range 1.9-16.5 ng/ml). 4. The average plasma level of 25(OH)-vitamin D3 was higher in the men (26.2 ng/ml) than in the women (19.3 ng/ml); this was attributed to the fact that the men spend more time in the sun than the women. 5. Among subjects in the age range 20-29 years, the 25(OH)-vitamin D3 level was significantly lower than in older subjects. 6. Daily administration of 400 IU vitamin D2 led, after 8 weeks, to a plateau of 11.5 ng 25(OH)-vitamin D 2 /ml. After the dosage was stopped, the level slowly declined with a half-life of 4-5 weeks. The presence of vitamin D2 in human plasma is mainly due to the consumption of multivitamin preparations and enriched foods, which in Japan often contain vitamin D2. The plasma level of 25(OH)-vitamin D is in principle unaffected by pregnancy. Since there is a correlation between the maternal and fetal levels of 25(OH)-vitamin D, it is inferred that the vitamin is transferred across the placenta. Transplacental transfer of calciferol and 25(OH)-vitamin D has been demonstrated in a number of mammalian species. However, the transport protein DBP cannot cross the placenta [Bouillon (1983 b)]. Measurements of the 25(OH)-vitamin D levels in maternal and umbilical cord sera revealed wide variations, which depend on the amount of sunlight and nutritional status. The level of 25(OH)-vitamin D in the newborn infant is 49 to 108% of the maternal serum level. Because of the correlation between the 25(OH)-vitamin D levels of mother and child observed in many studies, it is often inferred that the transport of maternal 25(OH)-vitamin D is a passive or facilitated diffusion [Bruns & Bruns (1983)]. In South Africa, extremely low serum levels (1 ng/ml) of 25(OH)-vitamin D have often been observed in breast-fed children who, like their mothers, received no supplemental vitamin D.The serum levels of the mothers was about 10 ng/ml. In spite of their extremely low serum levels of the vitamin, these children had none of the biochemical symptoms of rickets. On the other hand, some authors have observed rachitic children with 8-20 ng 25(OH)-vitamin D/ml serum; however, since these children had other health problems, these results must be interpreted with care. Rickets is not always the result of a vitamin D deficiency alone; other possible causes are inadequate intake of Ca 2+ or phosphate or genetic disease. It is thought that a serum level of 3-10 ng 25(OH)-vitamin D/ml provides a normal vitamin D metabolism [Holmes & Kummerow (1983)].

3.8 Distribution and Storage of Vitamin D

163

3.8.2 Milk Human milk contains relatively little vitamin D. Recent determinations, using a variety of methods, have given the value of 4.2-90 IU/liter (105-2250 ng/liter). Since the daily requirement of an infant is 400 IU vitamin D, it is recommended that mothers' milk be supplemented with vitamin D (400 IU/day). Earlier reports of relatively large amounts of vitamin D sulfate (up to about 25,000 ng/liter) in human milk were not confirmed by more recent measurements; the highest values found were 500 ng/liter (20 IU/liter). Finally, experiments on rats with synthetic vitamin D sulfate showed that the compound has no antirachitic activity [Greer et al. (1982a,b), Makin et al. (1983)]. More exact measurements showed that human milk contains mainly calcidiol, 160-1,200 ng/liter [Holmes & Kummerow (1983)]. According to another report, human milk contains, in addition to calcidiol (320 ng/liter), still smaller amounts of 24,25(OH)2-vitamin D 3 (42 ng/ liter) and l,24,25(OH)3-vitamin D 3 (12 ng/liter) [Koshy (1982)]. On the levels of cholecalciferol and calcidiol in milk and plasma after intravenous injection of cholecalciferol or calcidiol, see Hiridoglou & Knipfel [Hiridoglou&Knipfel (1984)]. On the analysis of vitamin D2, vitamin D3, 25(OH)-vitamin D 2 and 25(OH)-vitamin D3 in human milk, see Hollis (1983). The metabolism and function of vitamin D in lactation are discussed in detail by Toverud (1983). 3.8.3 Liver, Fatty Tissue and Muscle Unlike vitamins Α,Ε and K, vitamin D is not stored in the liver. High concentrations of this vitamin are found only in the livers of some fish. Mammalian liver contains only traces of vitamin D, up to about 1 μg/100g.The primary function of the liver in the economy of vitamin D is the formation of 25(OH)-vitamin D, which is localized mostly in extracellular fluids. Less well understood functions of the liver are the inactivation of vitamin D and the excretion of the break-down products in the bile [Fraser (1983)]. Intracardial application of 50 IU [3H]vitamin D 3 to rachitic chicks leads to a rapid appearance of radioactivity in the liver. A very high level is reached, which then falls by about half after approximately 7 hours. In the rat, too, the liver concentration of [3H]vitamin D3 becomes very high for a short time after application, but the amount in the bones is not much lower. The results of a long-term experiment on human beings is shown in Table 3-10. The subjects were given [3H]vitamin D3, and the levels in various organs were measured 4 to 59 days later. It can be seen that fatty tissue and skeletal muscles are the most important storage organs. Chromatography of the radioactive samples showed that fatty tissue contained mostly unchanged cholecalciferol, while the muscles contained mostly calcidiol [Norman (1979)], p. 199 ff]. In the plasma of the rat, about 95% of the calcidiol is bound to the DBP and about 5% to the lipoproteins; cholecalciferol is distributed 60% to DBP and 40% to lipoproteins. Perfusion experiments on rat liver showed that essentially only lipoprotein-bound vitamin D is taken up in the liver [Silver & Beny (1982)].

164

3 Vitamin D

Table 3-10. Distribution of vitamin D3 in humans, as % of the applied dose [Norman (1979), p. 199 ñ] Organ

Bones Fat Kidneys Liver Skeletal muscle Skin Serum

Days after application of [3H]cholecalciferol 4

22

59

1.8 5.4

0.6 8.6 0.2 1.3 8.6 2.4 3.6

0.8 10.1

-

29.5 10.8 3.3

-

0.3 5.5 1.6 1.5

3.8.4 Birds' Eggs Almost all the calciferol (about 95%) in the egg is found in the egg yolk; about 90% of this is unmodified vitamin. Its transport out of the serum is mediated by one of the two specific serum proteins (p. 156). Of 19 bird species examined, practically only the domesticated fowl (geese, chickens, turkeys and doves), which have extended laying periods, have these two transport proteins. Their concentrations are increased by estradiol. Another protein, phosvitin, is likewise involved in the transport of cholecalciferol from the blood to the egg yolk. Phosvitin is an egg-yolk protein containing 10% phosphate and about 7% Ca 2 + . It is synthesized in the liver, and enters the blood, where it binds to the cholecalciferol-protein complex. The total complex then moves into the developing egg cell. In contrast to the bird embryo, the mammalian fetus receives its vitamin D mainly in the form of 25(OH)-vitamin D, which is transferred across the placenta. The bird embryo controls the metabolism of cholecalciferol autonomously [Fraser & Emtage (1976)].

3.9 Biosynthesis and Metabolism of Vitamin D and Its Localization Before about 1950, practically nothing was known about the metabolism of vitamin D in humans. It was thought that this vitamin was effective as such. However, the remarkably long time between application and observable physiological effect of vitamin D3 was not consistent with this view (A.Carlsson, 1952). After intravenous application of 10 IU vitamin D3, intestinal Ca 2+ transport was not observable for 10-12h, which indicated that the vitamin must undergo conversion. Most of the metabolites of vitamin D 3 which are known today were discovered by using radioactively labelled vitamin D3 in rachitic animals. In spite of the number of these metabolites which have in the meantime been isolated and identified, there seem to be several more which have not been discovered, especially in the bile [Koshy (1982)]. Cholecalciferol is synthesized in the skin from 7-dehydrocholesterol by the action of UV light. Like ergocalciferol, cholecalciferol can also be obtained per

3.9 Biosynthesis and Metabolism of Vitamin D and Its Localization

165

Table 3-11. Organs and enzymes which hydroxylate vitamin D compounds [Norman et al. (1982b)] Organ

Species

Hydroxylase

Liver

Rat Chick Chick Chick Rat Rabbit

25-Hydroxylase 25-Hydroxylase 25-Hydroxylase la-Hydroxylase la-Hydroxylase 1-Hydroxylase 24R-Hydroxylase 1-Hydroxylase 24-Hydroxylase la-Hydroxylase 24R-Hydroxylase 25-Hydroxylase 24R-Hydroxylase 23-Hydroxylase la-Hydroxylase

Kidney

Bones

Human Placenta

Human

Intestine

Chick

Breast

Human

os, i. e. in the food. Before it can exert its known effects in the intestine (promotion of Ca 2+ and phosphate uptake), bones (promotion of bone resorption and mineralization) and other organs (kidneys, parathyroids, muscle, etc.), cholecalciferol must be converted to calcitriol by two hydroxylations, the first in the liver (to calcidiol) and the second in the kidney. The synthesis of calcitriol from calcidiol is stimulated by Ca 2 + deficiency and by increased Ca 2 + requirements (pregnancy, lactation, growth). If the Ca 2 + supply is adequate, the less effective 24,25(OH)2-vitamin D 3 is formed instead [Flury & Haldimann (1981)]. The kidney acts in this case as a classical endocrine gland, in that its synthesis of the two dihydroxylated metabolites is subject to complex hormonal regulation. The following regulators of renal calcidiol hydroxylases are known: parathormone (PTH), calcitonin (CT), prolactin, estrogens, growth hormone, insulin, vitamin D status, serum Ca 2+ and serum phosphate [Flury & Haldimann (1981)]. The kidney is the most important center of vitamin D metabolism and is thus eminently suited to be the site of physiological regulation. One of the regulators is calcitriol, which induces renal 24-hydroxylase and thus the formation of 24,25(OH)2vitamin D 3 [Norman et al. (1982b)]. It can be seen from Table 3-11 that a number of organs other than liver and kidneys have significant amounts of hydroxylases. It is therefore understandable that in patients with no kidneys, a plasma concentration of 10-15 pg calcitriol/ ml was found; this was formed extrarenally. Extrarenal formation may also account for some of the increase in the plasma level of calcitriol in pregnancy and lactation [Norman et al. (1982 b)]. At present, more than 20 chemically well characterized metabolites of vitamin D are known; almost all of these are formed in vivo. The structural formulas of most of the metabolites and their sites of formation are to be seen in Fig. 3-5.

25· increased PTH —· hypophosphatemia —>· rickets [Fraser et al. (1967)]. Some authors consider the primary cause of rickets to be phosphate deficiency, which is primarily the result of defective phosphate transport in the kidney or intestine. Rickets usually only develops, however, when the pre-existing metabolic phosphate deficiency is compounded by a calciferol deficiency (rachitic diseases which are due only to phosphate deficiency are less common). This view is supported by the observation that rats with a simple calciferol deficiency are hypocalcemia but

3.15 Deficiency Symptoms and Therapy

199

not rachitic; the latter condition only appears if the feed contains inadequate amounts of both calciferol and phosphate [Bronner (1976)]. Clinical observations of children have shown that hypocalcemia induced by calciferol deficiency leads to only mild bone disease. However, in children with severe rachitic bone deformation, hypophosphatemia, together with high levels of alkaline phosphatase in the serum, is one of the most obvious symptoms. A phosphate deficiency in childhood can also be nutritionally caused. Because calciferol promotes both Ca 2+ and phosphate transport in the intestine, it is understandable that both groups of mineral deficiency diseases respond to calciferol [Lapatsanis et al. (1976)]

Other causes of rachitis/osteomalacia are impaired intestinal function and a deficiency of bile acids ; these are essential for the absorption of vitamin D [Norman (1982)]. The best therapy for a simple vitamin D deficiency is exposure to the sun or irradiation. Healing can also be achieved, of course, by administration of a vitamin D preparation. Small doses of vitamin D3 or D 2 are recommended, so that several physiological protection mechanisms against intoxication (which accelerate metabolism and excretion) can be avoided. Application of calciferol provides the organism not only with 1,25(0H)2-vitamin D, but 24,25(OH)2vitamin D and other metabolites, which may be important for optimal mineralization [Flury & Haldimann (1981)]. Calciferol treatment (4,000 IU/day) of premature rachitic children leads to normalization. However, calcitriol (e. g. 0.05 μg/ kg body weight and day, intravenously) has also been used successfully. Such children usually also need an increased intake of Ca 2+ and phosphate [Bruns & Bruns (1983)]. If malabsorption is serious, parenteral doses of vitamin D are recommended at first, until the integrity of the intestinal mucosa has been reestablished. It is often possible to achieve success with elevated oral doses of calciferol, but in this case the serum levels of Ca 2+ and 25(OH)-vitamin D must be monitored [Flury & Haldimann (1981), Norman (1982)]. The cases of rachitis/osteomalacia discussed thus far have been associated with inadequate intake of calciferol; the calciferol metabolism is normally intact. However, impairment of the 25-hydroxylase of the liver or nephrosis can also lead to rachitis/osteomalacia, associated with a deficiency of 25(OH)-vitamin D.The following groups of patients are affected : 1. Epilepsy patients who have been treated with phénobarbital and hydantoin preparations. The antiepileptica induce enzymes which accelerate the degradation and biliary loss of vitamin D metabolites, and consequently cause a deficiency of 25(OH)-vitamin D.The affected children absorb too little Ca 2 + , often grow too slowly, have symptoms of rickets and have elevated PTH levels. To prevent rickets in children who are given long-term treatment with antiepileptics, additional doses of vitamin D (up to 4,000 IU/day) are recommended [Flury & Haldimann (1981), Juttman et al. (1982), Koshy (1982), Norman (1982)]. 2. Patients with liver cirrhosis. Reduced formation of 25(OH)-vitamin D due to enzyme impairment reduces the vitamin D reserves. Oral administration of 25(OH)-vitamin D (50-200 μg/day) is recommended [Flury & Haldimann (1981)].

200

3 Vitamin D

3. Nephrotic patients lose 25(OH)-vitamin D in their urine. Additional vitamin D is required [Flury & Haldimann (1981)].

3.15.2 Renal Osteodystrophy Kidney diseases very often lead to skeletal disease (osteodystrophy). Only after the introduction of radioisotopes was it possible to demonstrate that in kidney patients, the intestinal Ca 2+ uptake is usually impaired. We now know that the kidney is the site of formation of l,25(OH)2-vitamin D, which is an essential hormone for intestinal Ca 2 + uptake, and its formation requires intact kidneys [Norman et al. (1982c), p.825-897]. Renal osteodystrophy results from chronic kidney disease of various etiologies. It is a frequent complication in dialysis patients. The more severe the renal insufficiency and the greater the reduction in the glomerular filtration rate, the more frequent and more severe the renal osteodystrophy. The skeleton anomalies include reduction of bone tissue, general demineralization and ostitis fibrosa, associated with bone pains, breaks and deformation, and abnormal posture. In children, growth is impaired. In the sera of untreated patients the results are hypocalcemia, hyperphosphatemia, elevated alkaline phosphatase and elevated PTH concentration. Renal osteodystrophy is more common among children than adults, presumably because of the increased sensitivity of the growing bone to changing levels of vitamin D, phosphate and PTH in chronic kidney disease [Norman (1982)]. On the pathogenesis of renal osteodystrophy, the following is known: Chronic kidney failure leads to hypocalcemia, due to deficient intestinal Ca 2+ resorption and renal phosphate filtration. This leads to hyperphosphatemia and an increase in the secretion of PTH. As the renal insufficiency progresses, the plasma level of l,25(OH)2-vitamin D sinks, whereupon the excess PTH causes a pathological bone resorption [Norman (1982)]. Since PTH is degraded chiefly in the kidneys, its half-life is lengthened in renal insufficiency, which produces an additional increase in the plasma concentration. Ostitis fibrosa, the skeletal symptom of secondary hyperparathyroidism, appears to be the main complaint in the final stages of uremia [Coburn et al. (1982)]. "Dialysis osteomalacia" is a special form of renal osteodystrophy associated with a marked sensitivity to elevated doses of calcitriol. Its cause is probably the toxicity of the aluminum preparations which are administered as phosphate binders. Aluminum complexes are accumulated in the bones and cause bone disease [Coburn et al. (1982), Flury & Haldimann (1981), Goodman et al. (1984), Norman (1982)]. It has been observed that hypocalcemia, hyperphosphatemia and elevated PTH levels are factors which increase the risk that patients with chronic renal insufficiency will develop renal osteodystrophy. Thus it might be assumed that measures which reduce phosphate, administration of Ca 2+ and calcitriol therapy might effectively prevent the bone disease. However, clinical experience in the past few years has shown that this is not the case, and the search continues for further etiological factors and possibilities for therapy [Coburn et al. (1982)].

3.15 Deficiency Symptoms and Therapy

201

There are two main goals in the prophylaxis and treatment of osteodystrophy: 1. Prevention of hyperphosphatemia by administration of phosphate binders, e.g. aluminum preparations (on their side effects, see above), in order to delay the secondary hyperparathyroidism, and 2. Improvement of C a 2 + uptake by oral administration of high doses of C a 2 + or appropriate forms of vitamin D, which are effective in spite of the greatly reduced level of 1-hydroxylase [Fluiy & Haldimann (1981)]. The response of the bones to 1-hydroxylated vitamin D preparations is usually much slower than the rise in C a 2 + uptake, and also depends on the type of bone lesion [Flury & Haldimann (1981)]. This limits the possibilities for therapy [Voigts Table 3-20. Prophylactic and therapeutic effects of some vitamin D forms in patients with chronic renal insufficiency and renal osteodystrophy Vitamin D

Dose/day

Therapeutic effect. Advantages and disadvantages

Calciferol

1-2 mg

Prophylaxis [Fröhling et al. (1982)]

DHT

0.25-1.0 mg

Improved Ca 2+ uptake in dialysis patients [Flury & Haldimann (1981)]

Calcitriol

0.25-1.0 μg

Prophylaxis and therapy of renal osteodystrophy [Ringe (1982)]; no deposit formation, unlike calciferol, calcidiol and DHT [Flury & Haldimann (1981)]; hypercalcemia effect; reduction of PTH and phosphate levels; partial relief of muscle weakness and bone pains [Voigts et al. (1983)].

24,25(OH)2D3

1-10 μg

Stimulation of Ca 2+ uptake (intestine) and bone mineralization; inhibition of bone resorption in nephrectomized rats (calcitriol stimulates bone resorption) [Voigts et al. (1983)]; inhibition of PTH secretion [Norman (1982), Voigts et al. (1983)]; improvement in clinical and pathological pictures, even in patients with high levels of aluminum complexes; these patients did not respond to calcitriol [Ott et al. (1982)].

24,25(0H)2D3 with calcitriol

25(OH)Dj

Improvement of the clinical, biochemical and bone histological picture; partial relief of muscle weakness, reduction of alkaline phosphatase [Evans et al. (1982), Voigts et al. (1983)]. 50-200 μ δ

25(OH)Dj and calcitriol 5,6-transCalcidiol

Improvement in patients with uremia and secondary hyperparathyroidism [Flury & Haldimann (1981), Norman (1982)]; promotion of bone mineralization [Coen et al. (1983)]; hypercalcemia effect [Ofifermann et al. (1982)]. Effective on patients with renal osteodystrophy [Koshy (1982)]

0.05-0.25 mg

Prophylaxis and therapy of renal osteodystrophy [Ringe (1982)]; normalization of the serum Ca 2+ ; the same concentration of calcidiol leads to hypercalcemia; stimulation of intestinal Ca 2+ uptake [Offermann et al. (1982)].

202

3 Vitamin D

et al. (1983)]. Treatment with calcitriol can lead to hypercalcemia, even if the calcitriol level is lower than normal. For example, about 20% of a group of dialysis patients who had been given calcitriol (on the average less than 0.25 μg/day) had serum Ca 2+ levels higher than 11.0 mg/100 ml, even though their serum calcitriol levels were below normal (average 13 pg/ml). Several mechanisms have been suggested for this hypercalcemia [Coburn et al. (1982)]: 1. Impaired Ca 2+ uptake in the bones, perhaps due to the accumulation of trace elements (e.g. Al 3+ ), 2. Impaired renal Ca 2+ excretion, and 3. Localization of the orally administered calcitriol in the intestinal mucosa, where it stimulates Ca 2+ uptake, although the plasma level of calcitriol is subnormal. Hypercalcemia was observed in 73% of dialysis patients after treatment with 0.25 to 0.5 μg calcitriol/day for 1 to 2 years. The hypercalcemia usually becomes apparent within 4 to 8 weeks after the start of the therapy. During long-term therapy, the serum Ca 2+ should therefore be monitored twice a month. If the Ca 2+ concentration exceeds 12-13 mg/100 ml, the medication should be stopped. The serum phosphate level should also be monitored [Voigts et al. (1983)]. The following rule applies: even a slight increase in serum Ca 2+ level due to treatment with a vitamin D preparation leads, in the case of pre-existing hyperphosphatemia, to a critical increase in the Ca-P product, and this in turn causes extraossiary calcification, especially in the vessels and kidneys [Ringe (1982)]. Experiments with 24,25(OH)2-vitamin D 3 have shown that this compound, applied alone or together with calcitriol, has some advantages over pure calcitriol (see Table 3-20.) The Ca 2+ -reducing effect of 24,25(OH)2-vitamin D3 is significant, and it can be expected that this analog, in combination with calcitriol, will be suitable for use with patients with severe ostitis fibrosa, a disease in which the use of calcitriol is limited by hypercalcemia [Hodsman et al. (1983), Voigts et al. (1983)]. 3.15.3 Hypoparathyroidism Hypoparathyroidism, i. e. a lack or reduced production of PTH, can be the result of surgical removal of the parathyroid glands or of a lesion in them. Secretion of PTH is a signal indicating hypocalcemia. Patients with hypoparathyroidism are unable to generate this signal, and thus to convert 25(OH)-vitamin D to l,25(OH)2-vitamin D (see p. 173). As a result, the serum Ca 2 + level continues to drop, and eventually leads to elevated serum phosphate (hyperphosphatemia) [Koshy (1982)]. The deficiency of l,25(OH)2-vitamin D in patients with hypoparathyroidism is the result of the absence of two important stimuli for the 1-hydroxylase: PTH and hypophosphatemia. The therapy for this condition is a compensation for the l,25(OH)2-vitamin D deficit, and consists of administering physiological doses of calcitriol (0.68-2.7 μg/day) or la(OH)-vitamin D 3 (5.0 μg/day) or pharmacological doses of vitamin D (1-3.8 mg/day). Dihydrotachysterol (0.2-0.6 mg/day) together with adequate doses of Ca 2 + is also suitable. The increase in the serum

3.15 Deficiency Symptoms and Therapy

203

Ca 2+ level is independent of the type of vitamin D preparation, mostly due to the stimulation of the intestinal Ca 2+ resorption and less because of mobilization from the bones. Vitamin D therapy and increased Ca 2+ levels cause the phosphate level to drop [Flury & Haldimann (1981), Koshy (1982), Okano et al. (1982), Ringe (1982)]. 3.15.4 Osteoporosis Osteoporosis [Dixon (1983)] is a loss of bone tissue with a retention of the bone structure. The exact cause of osteoporosis is apparently unknown. It affects mainly older people whose sexual hormone levels are decreasing, as well as people who have been treated with steroids, such as glucocorticoids (senile, postmenopause and steroid-induced osteoporosis). Decreasing concentrations of the sexual hormones probably impair vitamin D metabolism and Ca 2+ uptake, so that these functions decrease in old age. Patients with osteoporosis often have relatively low serum levels of l,25(OH)2vitamin D, which is probably due to hormonally caused impairment of the renal hydroxylation of 25(OH)-vitamin D [Koshy (1982)]. Therapeutic success has been obtained by application of calcidiol or calcitriol [Koshy (1982)] and la(OH)-vitamin D3 [Lindholm et al. (1982)]. A 28-day experiment on 489 patients with primary osteoporosis (senile and post-menopausal) showed that la(OH)-vitamin D3 led to an increase in the bone mass. Furthermore, in this experiment a positive correlation was observed between the original serum level of 24,25(OH)2-vitamin D and the healing effect; the higher their 24,25(OH)2vitamin D levels, the better the patients responded to la(OH)-vitamin D3 [Orimo et al. (1982)]. A long-term treatment of patients with osteoporosis with physiological doses of 24,25(OH)2-vitamin D3, however, had no advantages [Reeve et al. (1982)]. 3.15.5 Congenital Impairments of Vitamin D Metabolism A number of hereditary conditions which directly or indirectly influence vitamin D metabolism are known. The best known involve: 1. Tubular phosphate resorption 2. Renal hydroxylation of 25(OH)-vitamin D 3. Cellular receptor proteins for l,25(OH)2-vitamin D and 4. Cellular PTH receptors. 3.15.5.1 Vitamin-D-Resistant Rachitis/Osteomalacia (X-Linked Hypophosphatemic Rickets) Vitamin-D-resistant rickets in childhood or vitamin-D-resistant osteomalacia in adults (also called hypophosphatemia-rachitis/osteomalacia, rachitis renalis, phosphate diabetes) is an X-linked, dominant hereditary disease. It involves: 1. Impairment of the phosphate reabsorption in the proximal kidney tubules, which does not respond to vitamin D (the primary defect is in the proximal tubuli),

204

3 Vitamin D

2. Hypersensitivity to PTH, 3. Impairment of the conversion of 25(OH)-vitamin D to l,25(OH)2-vitamin D [Chan & Wellons (1982)], so that in spite of the hypophosphatemia, which normally stimulates the 1-hydroxylase, too little l,25(OH)2-vitamin D is formed [Flury & Haldimann (1981)] and 4. Impairment of phosphate transport in the intestine [Koshy (1982), Norman (1982)]. It is thought that the tubular phosphate transport and tubular hydroxylation of 25(OH)-vitamin D are controlled by a single mechanism, which is impaired in vitamin-D-resistant .rachitis/osteomalacia [Bonjour et al. (1982)]. Experiments on mice with genetic hypophosphatemia have shown that the renal hydroxylation of 25(OH)-vitamin D functions, in principle, but that primarily 24,25(OH)2-vitamin D is formed [Tenenhouse (1982)]. The disease is accompanied by an increase in the serum alkaline phosphatase. The rachitic skeletal changes usually appear after the first year of life [Ringe (1982)], and growth of the affected children is impaired [Rasmussen et al. (1982)]. The traditional therapy of hypophosphatemic rachitis/osteomalacia consists of administration of phosphate (1.0-4.0 g/day) together with high doses of vitamin D (25,000-50,000 IU). The high doses of vitamin D often led to hypercalcemia and kidney damage [Norman (1982)]. Today the la-hydroxylated forms of vitamin D, namely la(OH)-vitamin D3 and calcitriol, are used, since it is known that the 1 α-hydroxylase is impaired in the affected patients. The dosage is 0 . 2 5 - 3 ^ g / d a y [Flury & Haldimann (1981), Harrelson et al. (1982), Norman (1982), Rasmussen et al. (1982)]. The Fanconi syndrome is associated with several transport defects in the proximal kidney tubules, involving the systems for phosphate, glucose, amino acids and bicarbonate. The bone disease is the result of the renal losses of phosphate (hypophosphatemia) and bicarbonate (metabolic acidosis) and the defective synthesis of l,25(OH)2-vitamin D.Administration of calcitriol or la(OH)-vitamin D 3 and Na + or K + bicarbonate has proved a successful therapy [Norman (1982)]. 3.15.5.2 Congenital Vitamin-D-Dependent Rickets In 1961, V.A. Prader described pseudo-deficiency rickets, a newly discovered congenital mineral metabolic disease. It is a hypocalcemia rickets, which is not responsive to physiological doses of vitamin D, but which does respond to pharmacological doses. Two forms of pseudo-deficiency rickets have been distinguished: type I, the more frequent, is characterized by defective hydroxylation of 25(OH)-vitamin D; and type II, which is less common, is associated with a lack of sensitivity of the target cells to calcitriol [Balsan et al. (1983), Liberman et al. (1982), Visser (1982)]. Congenital Vitamin-D-Dependent

Rickets, Type I

Type I pseudo-deficiency rickets is an autosomal recessive disease caused by a defect of the renal 1-hydroxylase. The clinical picture is one of severe hypocalcemia and hypophosphatemia, associated with secondary hyperparathyroidism

3.15 Deficiency Symptoms and Therapy

205

and rachitic skeletal changes. The serum level of 1,25(0H)2-vitamin D is very low or undetectable [Liberman et al. (1982), Norman (1982)]. The patients (mostly children) respond to physiological doses of calcitriol or la(OH)-vitamin D3 (about 1 μg/day) [Koshy (1982)]. Congenital Vitamin-D-Dependent

Rickets, Type II

Type II pseudo-deficiency rickets, like type I, is an autosomal recessive disease and its symptoms are similar. However, unlike patients with type I disease, type II patients have normal or elevated serum levels of l,25(OH)2-vitamin D and remission is only obtained by generating a still higher plasma level of l,25(OH)2-vitamin D (200-4000 pg/ml) [Liberman et al. (1982)]. The symptoms are rachitic changes in the bones, total baldness, dentai malformation, hypocalcemia and elevated PTH values. The target organs are resistant to calcitriol. No receptor protein for calcitriol was found in the skin fibroblasts of one patient (such receptor proteins are found in the skin of healthy humans and mice) [Holick et al. (1982)]. The observed resistance of the patients to physiological doses of calcitriol suggest that the syndrome is caused by a deficiency of specific receptors for l,25(OH)2-vitamin D.This makes necessary the therapeutic application of massive doses of this metabolite, which exceed the normal dose by five- to one hundred-fold [Norman (1982)]. The skin fibroblasts of patients with pseudo-deficiency rickets type II are used for the study of the mechanism of calcitriol action [Clemens et al. (1983)]. There are a few subgroups of type II pseudo-deficiency rickets: Two subgroups were recently described, each represented by one child. In one child with rickets and baldness, the calcitriol therapy was not successful, and in the other, the same treatment was successful only until its sixth year. Both children displayed hypocalcemia, highly elevated serum PTH and alkaline phosphatase levels, hypophosphatemia and reduced tubular phosphate reabsorption [Balsan et al. (1983)]. Three subgroups were found in other laboratories [Liberman et al. (1982,1983)]. Cell cultures from one patient had normal calcitriol binding in the cytoplasm, but the uptake of calcitriol into the nucleus could not be detected. Similar results were obtained with fibroblasts from the same patient. In another group of patients with the same symptoms, fibroblast cultures were used to show that the cytoplasmic calcitriol receptors were defective [Feldman et al. (1982), Liberman et al. (1983)]. Finally, a further calcitriol-resistant subgroup has been described. In fibroblast cultures from these patients, there was no induction of the 24-hydroxylase [Griffin et al. (1982), Liberman et al. (1983)].

3.15.5.3 Pseudohypoparathyroidism Pseudohypoparathyroidism results from a congenital resistance of the target organs to PTH. Patients with this syndrome secrete enough PTH (unlike patients with hypoparathyroidism, see p. 202) when their serum Ca 2 + drops, but the kidneys and bone tissue do not respond to the hormone. The patients suffer hypocalcemia, although the serum PTH level is high. Oral treatment with calcitriol or la(OH)-vitamin D 3 and Ca 2+ normalizes the serum Ca 2 + level [Koshy (1982)]. The therapeutic effects of several forms of vitamin D have been compared for

206

3 Vitamin D

Table 3-21. Optimal doses (in μg/day) of several forms of vitamin D for patients with pseudohypoparathyroidism (PsH) and postoperative hypoparathyroidism (PoH) [Okano et al. (1982)] Condition

PsH PoH

Form of vitamin D Calcitriol

a(OH)-D 3

DHT

Ergocalciferol

1.5 2.0

2.0 5.0

350 600

3,100 3,800

treatment of patients with pseudohypoparathyroidism (PsH) and postoperative hypoparathyroidism (PoH). The optimal doses are shown in Table 3-21. 3.15.6 Diabetes Mellitus Clinical diabetes mellitus is accompanied by osteopenia (breakdown of the bone tissue). In experimental diabetes, mineral and vitamin D metabolisms are altered. The dominant changes in the diabetic skeletal system are impaired bone and matrix formation and delayed osteoid maturation. These are not accompanied by either hypocalcemia or hypophosphatemia, nor is renal function impaired. Insulin normalizes the condition [Goodman & Hori (1982)]. The formation of l,25(OH)2-vitamin D is usually normal in diabetes patients [S;orensen et al. (1982)]. 3.15.7 Vitamin D Deficiency Symptoms in Animals The reaction of many experimental animals to a vitamin D deficiency has been carefully studied [ Vitamin Compendium (1980)]. General symptoms: Reduced appetite, crippling, impaired growth, weight loss, increased irritability, cramps. Bones: Stiff, painful gait, immobility, hard protrusions on the joints of the limbs and the ribs, bone bending in the limbs, breastbone (chickens) and backbone, increased tendency of the bones to break, soft, pliable bones (domestic animals). Reproduction: Decreased laying of eggs, thin shells, impaired hatching (chickens). 3.15.8 Commercially Available Forms of Pharmaceutical Vitamin D In addition to vitamins D3 and D2, the pharmaceutical industry supplies calcidiol, calcitriol, la(OH)-vitamin D3, dihydrotachysterol (DHT) and 5,6-trans25(OH)-vitamin D3. Tables 3-22 and 3-23 give an overview of the forms of vitamin D and the indications for treatment. 5,6-/raw-25(OH)-Vitamin D3, which does not occur naturally, was introduced especially for the therapy of renal osteodystrophy. la(OH)-Vitamin D3, which is also synthetic, only needs to be hydroxylated at C-25 in the liver in order to be converted to calcitriol, and is thus almost as active as the latter. The classical therapy with vitamin D 3 or D 2 carries the risk of accumulation, because vitamin D is stored in fatty tissue. Even if the therapy is halted because of overdosage, hypercalcemia continues to be manifest, often

3.16 Hypervitaminosis D, Toxicity

207

Table 3-22. Commercial vitamin D preparations [Flury & Haldimann (1981), Norman (1982), Ringe (1982)] Substance

Trade name

Vitamin D 3

Form

Indications

Drops, capsules

Rickets, osteomalacia, (general vitamin D deficiency diseases)

Vitamin D 2

Oldevit, Drisdol, Calciferol

Drops, capsules parenteral forms

As for vitamin D 3

Calcidiol

Calderai, De'drogyl

Drops (5 μg per drop)

Hepatic impairment

Calcitriol

Rocaltrol

Capsules, 0.25 and 0.5 μ β

Renal osteodystrophy, primary and secondary hyperparathyroidism, pseudohypoparathyroidism, hereditary pseudo-deficiency rickets, osteoporosis

la(OH)-Vitamin D 3

1-Alpha

Capsules, 0.25 and 1 μg

In principle, as for calcitriol

DHT

Hytakerol, DHT

Drops, capsules tablets

Renal osteodystrophy, hypoparathyroidism

5,6 -trans25(OH)-D 3

Delakmin

Capsules, 0.05 and 0.125 mg per capsule

Renal osteodystrophy

Table 3-23. Clinical and pharmacological properties of some forms of vitamin D [Flury & Haldimann (1981)] Vitamins D 3 and D 2 Daily dose ^ g ] for rickets 2-10 Daily dose ^ g ] for hypo750-3000 parathyroidism Duration of hypercalcemia 6-18 after cessation [months]

DHT

25(OH)D 3

l,25(OH)2D 3

20-100 250-1000

1-5 50-200

0.1-0.5 0.5-2.0

1-3

4-12

0.5-1.0

for months, and there is a danger of extraossiary calcification. This danger is much less with calcitriol, because its half-life is shorter [Flury & Haldimann (1981), Ringe (1982)].

3.16 Hypervitaminosis D, Toxicity That an overdose of vitamin D is toxic has been known since 1928 (A. F. Hess, J. M. Lewis). Interestingly, vitamin D generated by the action of light on the skin of healthy human beings never has toxic effects [Miller & Hayes (1982)].

208

3 Vitamin D

The minimal serum 25(OH)-vitamin D level required for health in humans is about 3 ng/ml. (Some subjects with no symptoms of osteomalacia have been observed to have serum levels less than 1.7 ng 25(OH)-vitamin D/ml, however.) On the other hand, a serum level of more than 30 ng 25(OH)-vitamin D/ml appears to be toxic. The optimal range of serum levels for 25(OH)-vitamin D is probably 10-30 ng/ml [Holmes & Kummerow (1983)]. Overdose intoxication occurs in most adults after intake of more than 50,000 IU (1.25 mg) calciferol/day [Miller & Hayes (1982)]. This is observed most often in long-term treatment of hypoparathyroidism. Excess calciferol is stored in fatty tissue and the elevated 25(OH)-vitamin D level can persist for months after the treatment has been stopped. DHT is also stored, although its half-life is much shorter than that of calciferol (Table 3-23). Intoxication has been observed with calciferol or DHT with daily therapeutic doses as low as 0.625 or 0.25 mg, respectively [Flury & Haldimann (1981)]. Calcitriol has an even narrower therapeutic range. Patients with Boeck's disease (Morbus Boeck) develop hypercalcemia in response to very low doses of vitamin D or even to exposure to sunlight. This is the result of an increase in l,25(OH)2-vitamin D, whereas 25(OH)-vitamin D remains in the normal range. The concentrations of the l,25(OH)2-vitamin D and calcium can be reduced by treatment with corticosteroids [Flury & Haldimann (1981)].

Ingestion of an overdose of vitamin D is quickly followed by weakness, nausea, loss of appetite, headache, abdominal pains, cramps, vomiting, diarrhea and polyuria. Prompt cessation of the vitamin D therapy leads to normalization [Holmes & Kummerow (1983), Koshy (1982), Norman (1982)]. In general, the result of hypervitaminosis D is the calcification of soft tissues. This can happen in the kidneys, heart, lungs, arteries and practically all tissues. The calcium deposits in the affected tissues can be detected microscopically and chemically. As to the mechanism of the calcinosis, it is assumed that at high serum concentrations, some forms of vitamin D, particularly 25(OH)-vitamin D, penetrate the cell membranes and alter their permeability to Ca 2 + . As a result, Ca 2 + flows into the cell and enters the organelles. The increase of the Ca 2 + concentration in the cytoplasm interferes with normal cellular metabolism [Holmes & Kummerow (1983)]. To avoid hypercalcemia, the serum and urine Ca 2+ levels of the patient should be monitored during any form of vitamin D therapy. The urine Ca 2 + concentration should not be higher than 300 mg/24h. It is further recommended, especially in cases of renal osteodystrophy, that the dose be increased gradually and the reaction of the patient be observed for a time before the next increase is undertaken [Flury & Haldimann (1981)]. Some observations on pregnant women suggest that very high doses of vitamin D (e.g. 100,000 IU/day, prescribed for hypoparathyroidism) can damage the fetus [Miller & Hayes (1982)]. The reaction of animals to an excess of vitamin D is similar to that of human beings. Numerous cases of calcinosis in grazing animals have been traced to the intake of calcinogenic plants. The active components of these plants are glycosides of calcitriol, calcidiol and cholecalciferol [Miller & Hayes (1982)].

3.17 References

209

Some authors warn against any overdose of vitamin D and even consider the officially recommended daily doses (Table 3-19) acceptable only under certain conditions. They argue that sunlight, especially in the summer months, supplies most or all of the body's requirements for vitamin D, and that the "recommendations" should apply only to people w h o are exposed to little sunlight (people with dark skins, inhabitants of northern regions, and the aged, w h o spend most of their time indoors). These critics therefore hold that the enrichment of some foods with vitamin D is questionable and that it should be limited. They point out that many people, especially in the USA, already consume considerable amounts of vitamin D as part of multivitamin preparations and that they use considerably more than an antirachitic dose [Holmes & Kummerow (1983)].

3.17 References Abe, E., Miyaura, C., Sakagami, H., Takeda, M., Konno, K., Yamazaki, T., Yoshiki, S.& Suda, T.(1981) Proc. Natl. Acad. Sa. U.S.A. 78 4990. Abe, E., Miyaura, C., Tanaka, H., Shiina, Y., Kuribayashi, T., Suda, S., Nishii, Y., Deluca, H. F. & Suda, T.(1983) Proc. Natl. Acad. Sci. U.S.A. 80 5583. Agwu, D.E. & Holub, B.J. (1983) Can. J.Physiol. Pharmacol. 61 954. Armbrecht, H. J., Wongsurawat, N., Zenser, T.V. & Davis, B.B. (1983) Arch. Biochem. Biophys. 220 52. Annbrecht, H.J., Wonsurawat, N., Zenser, T.V. & Davis, B.B. (1984) Amer. J.Physiol. 246 E 102.

Arnaud, C. D. (1983) in D. Bikle, ed., Assay of Calcium Regulating Hormones (Springer, New York) p. 1. Bachelet, M., Lacour, B.& Ulmann, A. (1982) Mineral Electrolyte Metab. 8 261. Baggiolini, E.G., Iacobelli, J. Α., Hennessy, B.M. & Uskokovic, M.R., (1982) J. Amer. Chem. Soc. 104 2945. Balsan, S., Garabedian, M., Liberman, U. Α., Eil, C., Bourdeau, Α., Guillozo, Η., Grimberg, R , Le Deunff, M.J., Lieberherr, M., Guimbaud, P., Broyer, M.& Marx, S.J. (1983) J. Clin. Endocrinol. Metab. 57 803. Bar-Shavit, Z., Teitelbaum, S.L., Reitsma, P., Hall, Α., Pegg, L.E., Trial, J.& Kahn, A. J. (1983) Proc. Natl. Acad. Sa. U.S.A. 80 5907. Bässler, Κ. H. & Lang, Κ. (1981) Vitamine, 2.Auflage (SteinkopfT, Darmstadt). Baxter, L. Α. & DeLuca, Η. F. (1976) J. Biol. Chem. 251 3158. Bengoa, J.M., Bolt, M.J.G. & Rosenberg, I.H. (1983) Gasteroenterology 84 1363, Pt.2. Bhalla, A.K., Amento, E. P., Clemens, T.L., Holick, M. F. & Krane, S.M. (1983) J. Clin. Endocrinol. Metab. 57 1308. Bielecka, L.& Lorenc, R.S. (1983) Post. Biochem. 29 53. Bieri, J.G. & McKenna, M.C. (1981) Amer. J.Clin. Nutr. 34 289. Bikle, D., ed. (1983) Assay of Calcium Regulating Hormones (Springer, New York). Bishop, C.W., Kendrick, N.C. & DeLuca, H.F. (1983) Feder. Proc. 42 Abstr.5199. Bishop, C.W., Kendrick, N.C. & DeLuca, H.F. (1984) J. Biol. Chem. 259 3355. Bonga, S.E. W., Lammers, P.I. & van der Meij, J.C.A. (1983) Cell Tissue Res. 228 117. Bonjour, J.P., Caverzasio, J., Mühlbauer, R., Trechsel, U.& Troehler, U.(1982) in A.W. Norman, ICSchaefer, D.von Herrath & H.G. Grigoleit eds., Vitamin D (de Gruyter, Berlin) p.427. Bouillon, R. (1983 a) J. Steroid. Biochem. 19 921, Part C.

210

3 Vitamin D

Bouillon, R., (1983 b) in M.F. Holick et al., eds. Perinatal Calcium and Phosphorus Metabolism (Elsevier, Amsterdam) p. 291. Bouillon, R.& Van Assche, F. A. (1982) Dev. Pharmacol. Ther. 4 38, Suppl.l. Bouillon, R., Van Baelen, H.& De Moor, P.(1976a) Biochem. J. 159 463. Bouillon, R., Van Baelen, H.& Rombauts, W. (1976 b) Eur. J. Biochem. 66 285. Boyce, R.W. & Weisbrode, S.E. (1983) Lab. Invest. 48 683. Brommage, R., Jarnagin, K., DeLuca, H.F., Yamada, S.& Takeyama, H.(1983) Amer. J.Physiol. 244 E 298. Bronner, F. (1976) Amer. J. Clin. Nutr. 29 1307. Brumbaugh, P.F., Haussler, D.H., Bressler, R.& Haussier, M.R. (1974) Science 183 1089. Bruns, M.E. (1983) in M.F. Holick et al., eds. Perinatal Calcium and Phosphorus Metabolism (Elsevier, Amsterdam) p.183. Bruns, M.E. & Bruns, D.E. (1983) Ann. Clin. Lab. Sei. 13 521. Bruns, M. E. H., Vollmer, S. S., Bruns, D. E. & Overpeck, J. G. (1983) Endocrinology 1131387. Chan, J.C.M. & Wellons, M.D. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds., Vitamin D(de Gruyter, Berlin) p.645. Chandler, J.S., Chandler, S.Κ., Pike, J.W. & Haussler, M.R. (1984) J. Biol. Chem. 259 2214. Chen, T.L., Cone, C.M., Morey-Holton, E.& Feldman, D.(1983) J. Biol. Chem. 258 4350. Christakos, S., Sori, Α., Greenstein, S. M. & Murphy, T. F. (1983) J. Clin. Endocrinol. Metab. 56 686.

Cinti, D.L., Golub, E.E. & Bronner, F. (1976) Biochem. Biophys. Res. Comm. 72 546. Clayton, J., Guilland-Cumming, D.F., Kanis, J.A. & Russell, R.G.G. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.821. Clemens, T. L., Adams, J. S., Horiuchi, N., Gilchrest, Β. Α., Cho, H., Tsuchiya, Y., Matsuo, N., Suda, T.& Holick, M.F. (1983) J. Clin. Endocrinol. Metab. 56 824. Coburn, J.W. & Massry, S.G. (1980) eds., Uses and Actions of 1,25-Dihydroxyvitamin D3 in Uremia (Karger, Basel). Coburn, J.W., Sherrard, D.J., Ott, S.M., Hodsman, A.B., Didomenico, N.C. & Alfrey, A.C. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.827. Coen G. et al. (1983) Mineral Electrolyte Metab. 9 19. Coppenhaver, D. H., Solenne, N. P. & Bowman, Β. Η. (1983) Arch. Biochem. Biophys. 226 218. Dauben, W.G. & Phillips, R.B. (1982) J. Amer. Chem. Soc. 104 5780. Davies, M.& Mawer, E.B. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 523. De Boland, A.R., Albornoz, L.E. & Boland, R.(1983) Calcif. Tissue Int. 35 798. DeLuca, H.F. (1976 a) Ann. Internal Med. 85 367. DeLuca, H.F. (1976b) Amer. J.Clin. Nutr. 29 1258. DeLuca, H.F. (1983) in W.A. Peck, ed. Bone and Mineral Research, Annual 1. (Excerpta Medica, Amsterdam) p. 7. DeLuca, H.F. & Schnoes, H.K. (1976) Annu. Rev. Biochem. 45 632. DeLuca, H.F. & Schnoes, H.K. (1983) Annu. Rev. Biochem. 52 411. Desplan, C., Brehier, Α., Perret, C.& Thomasset, M.(1983) J. Steroid. Biochem. 19 1577. Deyl, Z.& Adam, M.(1983) Biochem. Biophys. Res. Comm. 113 294. Dixon, A.S.J. (1983) ed., Osteoporosis (Acad. Press, London). Dokoh, S., Llach, F.& Haussler, M.R. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.743. Dueland, S., Helgerud, P., Pedersen, J.I., Berg, T.& Drevon, C.A. (1983a) Amer. J.Physiol. 245 E 326. Dueland, S., Pederson, J.I., Helgerud, P., & Drevon, C.A. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin O (de Gruyter, Berlin) p.27. Dueland, S., Pedersen, J.I., Helgerud, & Drevon, C.A. (1983b) Amer. J.Physiol., 245 E 463.

3.17 References

211

Edelstein, S.(1974) Vitam. Hormones 32 407. Edelstein, S.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 127. Edelstein, S., Fullmer, C.S. & Wasserman, R.H. (1984) J. Nutrition 114 692. Eisman, J.A. (1983) in M.K. Agarwal, ed., Principles of Recepterology (de Gruyter, Berlin) p.465. Eisman, J.A., Frampton, R.J., Sher, E., Suva, L.J. & Martin, J.T. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.65. Eisman, J. Α., Hamstra, A.J., Kream, B.E. & DeLuca, H.F. (1976) Arch. Biochem. Biophys. 176 235. Emerson, D.L., Galbraith, R M . & Arnaud, P. (1984) Electrophoresis S 22. Endo, H., Kiyoki, M., Kawashima, K.& Ishimoto, S. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.173. Esparza, M.S., Vega, M.& Boland, R.L. (1982) Biochim. Biophys. Acta 719 633. Evans, R.A., Hills, E., Wong, S. Y. P., Dunstan, C.R. & Norman, A.W. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.835. Fairney, A. (1983) in C.H. Gray & V.H.T. James, eds., Hormones in Blood, Vol. 5 (Acad. Press, London) p. 11. Feldman, D., Chen, T., Cone, C., Hirst, M., Shani, S., Benderli, A.& Hochberg, Ζ. (1982) J. Clin. Endocrinol. Metab. 55 1020. Flury, W.& Haldimann, B.(1981) Schweiz. Med. Wochenschr. 111 314. Fong, G.W.K., Johnson, R.N. & Kho, B.T. (1983) J. Assn. Offic. Anal. Chem. 66 939. Frampton, R.J., Omond, S.A. & Eisman, J.A. (1983) Cancer Res. 43 4443. Frankel, T.L., Mason, RS., Hersey, P., Murray, E.& Posen, S.(1983) J. Clin. Endocrinol. Metab. 57 627. Fraser, D.R. (1983) Lancet 1 969. Fraser, D.R. & Emtage, J.S. (1976) Biochem. J. 160 671. Fraser, D.R. & Kodicek, E.(1970) Nature22» 764. Fraser, D., Kooh, S.W. & Scriver, C.R. (1967) Pediat. Res. 1 425. Freake, H.C., Marcocci, C., Iwasaki, J., Stevenson, J.C. & Maclntyre, 1.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D(de Gruyter, Berlin) p.79. Friedlaender, M.M., Kornberg, Z., Wald, H., & Popovtzer, M. M. (1983) Amer. J. Physiol. 244 F 674. Fröhling, P. T., Kokot, F., Vetter, Κ., Kaschube, I., Schmicker, R, Großmann, I. & Lindenau, K.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.841. Goldsmith, R S . (1982) Miner. Electrolyte Metab. 8 289. Goodman, W.G., Henry, D.A., Horst, R., Nudelman, R.K., Alfrey, A.C. & Coburn, J.W. (1984) Kidney Int. 25 (1984) 370. Goodman, W.G. & Hori, M.T. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.405. Gray, R.W. & Napoli, J.L. (1983) J. Biol. Chem. 258 1152. Gray, T.K. & McAdoo, T.(1983) Clin. Chem. 29 196. Greer, F.R., Reeve, L.E., Chesney, R.W. & DeLuca, H.F. (1982a) Pediatrics69 238. Greer, F.R., Reeve, L.E., Chesney, R.W. & DeLuca, H.F. (1982b) Pediatrics70 499. Griffin, J.E., Chandler, J.S., Haussier, M.R. & Zerwekh, J.E. (1982) Clin. Res. 30 524A. Haddad, J.G., Hillman, L.& Rojanasathit, S.(1976) J. Clin. Endocrinol. Metab. 43 86. Haddad, J.G. & Sanger, J.W. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.1173. Haddad, J.G. & Walgate, J.(1976a) J. Clin. Invest. 58 1217. Haddad, J.G. & Walgate, J.(1976b) J. Biol. Chem. 251 4803. Harnden, D., Kumar, R , Holick, M.F. & DeLuca, H.F. (1976) Science 193 493.

212

3 Vitamin D

Harrelson, J.M., Lyles, K.W., Whitesides, P.C. & Drezner, M.K. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 973. Hashimoto, Y., Kawashima, H., Hoshina, K., Takeshita, T. & Ishimoto, S. (1976) Jap. J. Pharmacol. 26 114P, Suppl. Haussler, M.R. & Brickman, A.S. (1982) in F. Bronner & J.W. Coburn, eds., Disorders of Mineral Metabolism, Vol. 2. Calcium Physiology (Acad. Press, New York) p. 359. Hefti, E., Trechsel, U., Fleisch, H.& Bonjour, J.P. (1983) Amer. J.Physiol. 244 E 313. Henry, H.L. (1982) Mineral Electrolyte Metab. 8 179. Henry, H.L., Noland, T.A., Al-Abdaly, F., Cunningham, N.S., Luntao, E.M. & Amdahl, L.D. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.533. Hidiroglou, M.& Knipfel, J.E. (1984) Can. J. Comp. Med. 48 78. Hodsman, A.B., Wong, E.G.C., Sherrard, D.J., Brickman, A.S., Lee, D.B.N., Singer, F.R., Norman, A.W. & Coburn, J.W. (1983) Amer. J. Med. 74 407. Holick, M. F., Adams, J. S., Clemens, T. L., MacLaughlin, C. J., Horiuchi, N., Smith, E., Holick, S.A., Nolan, J.& Hannifan, N.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 1151. Holick, M. F., Gray, T. K. & Anast, C. S. (1983) eds. Perinatal Calcium and Phosphorus metabolism (Elsevier, Amsterdam). Holick, M.F., Semmler, E.J., Schnoes, H.K. & DeLuca, H.F. (1973) Science 180 190. Hollis, B.W. (1983) Anal. Biochem. 131 211. Holmes, R.P. & Kummerow, F. A. (1983) J. Amer. Coll. Nutrition 2 173. Horst, R.L., Reinhardt, Τ. Α., Pramanik, B.C. & Napoli, J.L. (1983) Biochemistry 22 245. Hosomi, J., Hosoi, J., Abe, E., Suda, T.& Kuroki, T. (1983) Endocrinology Mi 1950. Hummer, L., Nilas, L., Tjellesen, L.& Christiansen, C.(1984) Scand. J. Clin. Lab. Invest. 44 163. Hunziker, W., Walters, M.R., Bishop, J.E. & Norman, A.W. (1983) J. Biol. Chem. 258 8642. Ichikawa, Y., Hiwatashi, A.& Nishii, Y. (1983) Comp. Biochem. Physiol. 75b 479. Imawari, M., Kida, K.& Goodman, D.S. (1976) J. Clin. Invest. 58 514. Ishizuka, S., Ishimoto, S.& Orimo, H.(1982) in A.W. Norman, K Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 341. Isler, 0 . & Brubacher, G. (1982) Vitamine I. fettlösliche Vitamine (Thieme, Stuttgart). IUPAC-IUB (1982 a) Nomenclature of Vitamin D. Recommendations 1981. Pure Appi. Chem. 54 1511. IUPAC-IUB (1982 b) Nomenclature of Vitamin D. Recommendations 1981. Eur. J. Biochem. 124 223. Jarnagin, K., Brommage, R., DeLuca, H. F., Yamada, S. & Takeyama, H. (1983) Amer. J. Physiol. 244 E 290. Jones, G., Schnoes, H.K. & DeLuca, H.F. (1976) J. Biol. Chem. 251 24. Jongen, M.J.M., van der Vijgh, W.J.F., Lips, P.& Netelenbos, J.C. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D{de Gruyter, Berlin) p.797. Jongen, M.J.M., Van Ginkel, F.C., van der Vijgh, W.J.F., Kuiper, S., Netelenbos, J.C. & Lips, P. (1984) Clin. Chem. 30 399. Juttman, J.R., Braun, J.J., Visser, T.J. & Birkenhäger, J.C. (1982) in A. W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.989. Kadowaki, S.& Norman, A.W. (1984) J. Clin. Invest. 73 759. Kanis, J. A. (1982) J. Bone Joint Surg. Brit. 64 542. Kanis, J.Α., Guilland-Cumming, D„ Peterson, A.D. & Russell, R.G.G. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 157. Karsenty, G., Ulmann, Α., Lacour, B., Pierandrei, E.& Drüke, T.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.345. Kawashima, H., Hoshina, K., Saitoh, N.& Hashimoto, Y.(1976) Jap. J.Pharmacol. 26 114P, Suppl.

3.17 References

213

Kawashima, H.& Kurokawa, K.(1983) Miner. Electrol. Metab. 9 227. Keenan, M.J. & Holmes, R.P. (1983) Feder. Proc. 42 Abstr.5193. Kiyoki, M., Endo, H., Kawashima, K.& Ishimoto, S.(1982) in A.W. Norman, K. Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 401. Kobayashi, T., Okano, T., Shida, S., Okada, K., Suginohara, T., Nakao, H., Kuroda, E., Kodama, S.& Matsuo, T.(1983) J. Nutr. Sci. Vitaminol. 29 271. Koshy, K.T. (1982) J. Pharm. Sci. 71 137. Kreutter, D., Matsumoto, T., Peckham, R., Zawalich, K., Wen, W.H., Zolock, D.T. & Rasmussen, H.(1983) J. Biol. Chem. 258 4977. Kübler, W. (1980) in H. D. Cremer, D. Hötzel, & J. Kühnau, eds., Biochemie und physiologie der Ernährung (Thieme, Stuttgart, 1980) p.606. Kumar, R(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.635. Kumar, R.(1984) Physiol. Rev. 64 478. Kumar, R., Wiesner, R., Scott, M.& Go, V.L.W. (1982) Amer. J. Physiol. 243 E 370. Lambert, P.W., Fu, I.Y. & Kaetzel, D.M. (1982a) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.35. Lambert, P.W., Stern, P.H., Avioli, R.C., Brackett, N.C., Turner, R.T., Greene, Α., Fu, I.Y. & Bell, N.H. (1982 b) J. Clin. Invest. 69 722. Lapatsanis, P., Makaronis, G., Vretos, C.& Doxiadis, S.(1976) Amer. J. Clin. Nutr. 29 1222. Lawson, D. E. M. (1978) ed., Vitamin D (Acad. Press, London). Leonard, W.J., Strauss, A.W., Go, M.F., Alpers, D.H. & Gordon, J.I. (1984) Eur. J.Biochem. 139 561. Levy, J., Zuili, I., Yankowitz, N.& Shany, S.(1984) J. Endocrinol. 100 265. Lewin, I.G., Papapoulos, S.E., Hendy, G.N., Tomlinson, H.S. & O'Riordan, J.L.H. (1982) Clin. Sei. 62 381. Liang, C.T. & Sacktor, B.(1982) in A.W. Norman, K. Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.437. Liberman, U.A., Eil, C., Holst, P., Rosen, J.F. & Marx, S.J. (1983) J. Clin. Endocrinol. Metab. 57 958. Liberman, U.A., Eil, C.& Marx, S.J. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.73. Life Sciences Research Office (LSRO) Report (1983) Feder. Proc. 42 2658, No. 10. Lindholm, T.S., Nilsson, O.S., Kyhle, B.R. & Lindholm, T.C. (1982) in A.W. Norman, K-Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D(de Gruyter, Berlin) p.929. Lobaugh, B.& Drezner, M.K. (1983) Anal. Biochem. 129 416. Loo, J.C.K. & Brien, R.(1983) Res. Commun. Chem. Pathol. Pharm. 41139. Lu, J., Endres, D., Mueller, J.& Broughton, A.(1982) Clin. Chem. 28 1592. MacLaughlin, J.A., Anderson, R.R. & Holick, M.F. (1982) Science 216 1001. Maierhofer, W.J., Gray, R.W., Cheung, H.S. & Lemann, J.(1983) Kidney Int. 24 555. Makin, H.L.J., Seamark, D.A., & Trafford, D.J.H. (1983) Arch. Dis. Childh. 58 750. Malluche, H.H., Akmal, M., Mayer, E., Norman, A.& Massry, S.G. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D(de Gruyter, Berlin) p.539. Mangelsdorf, D.J., Koeffler, H.P., Donaldson, C.A., Pike, J.W. & Haussler, M . R (1984) J. Cell. Biol. 98 391. Marx, S.J., Liberman, U.A. & Eil, C.(1983) Vitam. Hormones40 235. Matsui, T., Nakao, Y., Kobayashi, N., Kishihara, M., Ishizuka, S.et al. (1984) Int. J. Cancer33193. Mawer, E.B. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D(de Gruyter, Berlin) p.623. Mayer, E., Bishop, J. E., Ohnuma, N.& Norman, A. W. (1983 a) arch. Biochem. Biophys. 224 671. Mayer, E., Reddy, G.S., Chandraratna, R.A.S., Okamura, W.H., Kruse, J . R , Popjak, G., Bishop, J.E. & Norman, A.W. (1983b) Biochemistry22 1798.

214

3 Vitamin D

McCarthy, D.M., San Miguel, J.F., Freake, H.C., Green, P.M., Zola, H., Catovsky, D.& Goldman, J.M. (1983) Leukemia Res. 7 51. McCormick, D.B. & Wright, L.D. (1980) eds., Methods Enzym. 67F 323-542. Melier, Y., Shainkin-Kestenbaum, R., Shany, S., Zuilli, I., Yankowitz, N., Giat, J., Konforti, A. & Torok, G.(1984) Clin. Orthop. Related Res. No.183p.238. Mendelsohn, M., Min, C., Haddad, J., Hahn, T.& Slatopolsky, E.(1976) Clin. Res. 24 582 A. Merke, J., Senst, S.& Ritz, E. (1984) Biochem. Biophys. Res. Comm. 120 199. Miller, D.R. & Hayes, K.C. (1982) in J.N. Hathock, ed. Nutritional Toxicology, Vol.1 (Acad. Press, New York) p. 81. Miyaura, C., Abe, E., Kuribayashi, T., Tanaka, H., Konno, K., Nishii, Y.& Suda, T. (1981) Biochem. Biophys. Res. Comm. 102 937. Moore, P.B. & Dedman, J.R. (1982) Life Sci. 31 2937. Morris, K.M.L. & Levack, V.M. (1982) Life Sci. 30 1255. Morris, J.F. & Peacock, M.(1976) Clin. Chim. Acta 72 383. Morton, R. A. (1977) Trends Biochem. Sci. 2 Ν 75. Muirhead, Ν., Adami, S., Sandler, L.M., Fraser, R.A., Fraher, L., Catto, G.R.D., Edward, N. & O'Riordan, J.L.H. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 187. Mulkins, M. Α., Manolagas, S.C., Deftos, L. J. & Sussman, H. H. (1983) J. Biol. Chem. 258 6219. Muniz, J . F , Wehr, C.T. & Wehr, H.M. (1982) J. Assn. Offic. Analyt. Chem. 65 791. Napoli, J.L. & Horst, R.L. (1983) Biochem. J. 214 261. Napoli, J.L, Pramanik, B.C., Royal, P.M., Reinhardt, T. A. & Horst, R.L. (1983) J. Biol. Chem. 258 9100. Narbaitz, R , Stumpf, W.E, Sar, M , Huang, S.& DeLuca, H. F. (1983) Calcif. Tissue Int. 35 177. Nemanic, M.& Bikle, D.(1983) Feder. Proc.4,1 Abstr.5192. Nemanic, M.K, Whitney, J , Arnaud, S , Herbert, S.& Elias, P.M. (1983) Biochem. Biophys. Res. Comm. 115 444. Nemere, I.& Norman, A. W. (1983) Feder. Proc. 42 Abstr.5195. Noland, T. A. & Henry, H.L. (1982) in A. W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 549. Norman, A.W. (1979) Vitamin D, The Calcium Homeostatic Steroid Hormone (Acad. Press, New York). Norman, A. W, Leathers, V.& Bishop, J.E. (1983) J. Nutr. 113 2505. Norman, A.W, Leathers, V.L., Bishop, J . E , Kadowski, S.& Miller, B.e. (1982a) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin Ζ)(de Gruyter, Berlin) p. 147. Norman, A.W, Roth, J.& Orci, L.(1982b) Endocrine Rev. 3 331. Norman, A.W, Schaefer, K , Grigoleit, H . G , von Herrath, D.& Ritz, E.(1975) eds, Vitamin D (de Gruyter, Berlin). Norman, A. W, Schaefer, K , Coburn, J. W, DeLuca, H. F , Fraser, D , Grigoleit, H. G. & von Herrath, D.(1977) eds, Vitamin D(de Gruyter, Berlin). Norman, A.W, Schaefer, K , von Herrath, D , Grigoleit, H.G, Coburn, J.W, DeLuca, H . F , Mawer, E.B, Suda, T. (1979) eds. Vitamin D (de Gruyter, Berlin) Norman, A.W, Schaefer, K , von Herrath, D , Grigoleit, H.G. (1982c) eds. Vitamin D (de Gruyter, Berlin). Norman, M.E. (1982) Pediatrie Clinics of N.America 29 947. Nutrition Rev. (1984) 42 27. Offermann, G , Kraft, D.& Delling, G.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.885. Ohnuma, N , Kruse, J . R , Popjak, G.& Norman, A.W. (1982) J. Biol. Chem. 257 5097. Okamura, W.H. (1983) Accounts Chem. Res. 16 81. Okano, K , Furukawa, Y , Morii, H.& Fujita, T.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.1061.

3.17 References

215

Orimo, H., Inoue, T., Fujita, T.& Itami, Y.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 1239. O'Riordan, J.L.H., Adami, S., Sandler, L.M., Clemens, T.L. & Fraher, L.J. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 751. Ott, S.M., Coburn, J.W., Maloney, Ν.Α., Alfrey, A.C. & Sherrard, D.J. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 869. Parfitt, A.M., Mathews, C.H.E., Brommage, R., Jarnagin, K.& DeLuca, H.F. (1984) J. Clin. Invest. 73 576. Perry, H.M., Chappel, J.C., Clevinger, B.L., Haddad, J.G. & Teitelbaum, S.L. (1983) Biochem. Biophys. Res. Comm. 112 431. Posner, A.S. & Betts, F.(1975) Accounts Chem. Res. 8 273. Price, P.A., Williamson, M.K. & Baukol, S.A. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 351. Procsal, D. Α., Okamura, W. H. & Norman, A. W. (1976) Amer. J. Clin. Nutr. 29 1271. Prowedini, D.M. et al. (1983) Science2211181. Puzas, J.E., Turner, R.T., Howard, G. A. & Baylink, D.J. (1983) Endocrinology 112 378. Rasmussen, H., Matsumoto, T.& Kreutter, D.(1982a) in A.W. Norman, K. Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin D(de Gruyter, Berlin) p. 1227. Rasmussen, H., Mazur, Α., Pechet, M., Gertner, J., Baron, R.& Anast, C. (1982b) in A.W. Norman, K. Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 1219. Reddy, G.S., Norman, A.W., Willis, D.M., Goltzman, D., Guyda, H., Solomon, S., Philips, D.R., Bishop, J.E. & Mayer, E.(1983) J. Clin. Endocrinol. Metab. 56 363. Reeve, L , Tanaka, Y.& DeLuca, H.F. (1983) J. Biol. Chem. 258 3615. Reeve, J., Tellez, M., Green, J.R., Hesp, R., Elsasser, U., Wotton, R., Hulme, P., Williams, D., Kanis, J.Α., Russell, R.G.G., Mawer, E.B., Meunier, P.J. (1982) Acta Endocrinol. 101 636. Reinhardt, Τ.Α., Horst, R.L., Orf, J.W. & Hollis, B.W. (1984) J. Clin. Endocrinol. Metab. 58 91. Ringe, J.D. (1982) Deutsche Med. Wochenschr. 107 1483. Rudberg, C., Akerstrom, G., Johansson, H., Ljunghall, S., Malmaeus, J.& Wide, L.(1984) Acta Endocrinol. 105 354. Seelig, M.S. (1983) J. Amer. Coll. Nutrition 2 109. Shiina, Y., Abe, E., Miyaura, C., Tanaka, H., Yamada, S., Ohmori, M., Nakayama, K., Takayama, H., Matsunaga, I., Nishii, Y., DeLuca, H.F. & Suda, T.(1983) Arch. Biochem. Biophys. 220 90. Shinki, T. Shiina, Y., Takahashi, N., Tanioka, Y., Koizumi, H.& Suda, T. (1983) Biochem. Biophys. Res. Comm. 114 452. Shinki, T., Takahashi, N., Kawate, N.& Suda, T. (1982) Endocrinology 111 1546. Schultz, T.D., Fox, J., Heath, H.& Kumar, R.(1983) Proc. Natl. Acad. Sci. U.S.A. 80 1746. Silver, J.& Berry, E.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 53. Simpson, R.U., Hamstra, Α., Kendrick, N.C. & DeLuca, H.F. (1983) Biochemistry 22 2586. Simpson, R.U., Wichmann, J.Κ., Paaren, Η.E., Schnoes, H.K. & DeLuca, H.F. (1984) Arch. Biochem. Biophys. 230 21. Slovik, D.M., (1983) in M.P. Cohen & P.P. Foa, eds., Special Topics in Endocrinology and Metabolism, Vol. 5. (Alan R.Liss, New York) 83. Sömjen, D., Kaye, A.M. & Binderman, 1.(1984) FEBS Lett. 167 281. Sömjen, D., Sömjen, G.J., Weisman, Y.& Binderman, 1.(1982) Biochem. J. 204 31. Serensen, Ο. H., Lund, B., Lund, B., Christiansen, J. S., Parving, H. H. & Andersen, A. R. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.43.

216

3 Vitamin D

Spanos, E., Pike, J.W., Haussler, M.R., Colston, K.W., Evans, I.M.A., Goldner, A.M., McCain, T.A. & Maclntyre, 1.(1976) Life Sci. 19 1751. Stancher, B.& Zonta, F. (1983) J. Chromatogr. 256 93. Suda, T., Abe, E., Miyaura, C., Tanaka, H., Shiina, Y., Kuribayashi, T.& Nishii, Y. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H. G. Grigoleit, eds. Vitamin Ζ) (de Gruyter, Berlin) p. 59. Szebenyi, D.M.E., Obendorf, S.K., Jones, A.J.S. & Moffat, K.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.215. Takada, K.(1983) J. Lipid Res. 24 441. Tanaka, H., Abe, E., Miyaura, C., Shiina, Y.& Suda, T. (1983 a) Biochem. Biophys. Res. Comm. 117 86. Tanaka, H., Abe, E., Miyaura, C.& Suda, T.(1983b) Seigaku 55 1323. Tanaka, Y.& DeLuca, H.F. (1973) Arch. Biochem. Biophys. 154 566. Tanaka, Y.& DeLuca, H.F. (1984) Amer. J.Physiol. 246 E 168. Tanaka, Y., DeLuca, H.F., Akaiwa, Α., Morisaki, M.& Ikekawa, N.(1976) Arch. Biochem. Biophys. 177 615. Tanaka, Y., DeLuca, H.F., Kobayashi, Y.& Ikekawa, N.(1984a) Arch. Biochem. Biophys. 229 348. Tanaka, Y., DeLuca, H.F., Satomura, K., Yamaoka, K.& Seino, Y. (1983 b) J. Lab. Clin. Med. 102 1010. Tanaka, Y., Seino, Y., Ishida, M., Yamaoka, K., Yabuuchi, H., Ishida, H., Seino, S., Seino, Y. & Imura, H.(1984c) Acta Endocrinol. 105 528. Taylor, A.N., Gleason, W. Α., & Lankford, G.L. (1984) J. Dental Res. 63 94. Tenenhouse, H.S. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p. 471. Thierry-Palmer, M.& Gray, T.K. (1983) J. Chromatogr. 262 460. Thomasset, M., Desplan, C.& Parkes, 0.(1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.197. Thomasset, M., Desplan, C.& Parkes, 0.(1983) Eur. J. Biochem. 129 519. Tjellesen, L.& Christiansen, C.(1982) in A.W. Norman, ICSchaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.49. Tjellesen, L.& Christiansen, C.(1983) Scand. J. Clin. Lab. Invest. 43 85. Toverud, S.U. (1983) in M.F. Holick et al., eds. Perinatal Calcium and Phosphorus Metabolism (Elsevier, Amsterdam) p.131. Turner, R.T., Howard, G.A., Puzas, J.E., Baylink, D.J. & Knapp, D.R. (1983) Biochemistry 22 1073. Van Os, C.H. & Ghijsen, W.E.J.M. (1982) in A.W. Norman, K.Schaefer, D.von Herrath, H.G. Grigoleit, eds. Vitamin D (de Gruyter, Berlin) p.295. Vesely, D.L. & Juan, D.(1983) Clin. Res. 31 399 A. Vesely, D.L. & Juan, D.(1984) Amer. J.Physiol. 246 E 115, Pl. Visser, H.K.Α., Degenhart, H.J. & Hogenboezem, T.(1982) J. Endocrinol. 94 7Ρ, Suppl. Vitamin Compendium 2. Auflage. (Hoffmann-La Roche, Basel, 1980). Voigts, A.L., Felsenfeld, A.J. & Llach, F.(1983) Arch. Intern. Med. 143 1205. Warner, M.(1983) J. Biol. Chem. 258 11590. Wasserman, R.H. & Fullmer, C.S. (1983) Annu. Rev. Physiol. 45 375. Wasserman, R.H., Henion, J.D., Haussler, mIr. & McCain, T.A. (1976) Science 194 853. Wong, G.(1982) Miner. Electrol. Metab. 8 188. Yamada, S., Ohmori, M., Takayama, H., Takasaki, Y.& Suda, T. (1983) J. Biol. Chem. 258 457. Zagalak, B., Neuheiser, F., Zagalak, M.J., Küster, T., Curtius, H.C., Exner, G.U., Fanconi, S.& Prader Α. (1983) in A. Frigerio, ed., Chromatography and Mass Spectrometry in Biomedical Sdences, 2 (Elsevier, Amsterdam, 1983) p.347. Zerwekh, J.E., McPhaul, J.J., Parker, T.F. & Pak, C.Y.C. (1983) Kidney Int. 23 401.

4 Vitamin E

4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.7.1 4.7.1.1 4.7.1.2 4.7.1.3 4.7.1.4 4.7.1.5 4.7.2 4.7.2.1 4.7.2.2 4.7.3 4.8 4.9 4.9.1 4.9.1.1 4.9.1.2 4.9.1.3 4.9.2 4.9.3 4.9.4 4.9.5 4.9.6 4.9.7 4.9.8 4.9.9 4.9.10 4.9.11 4.9.12 4.9.13 4.9.13.1 4.9.13.2 4.10 4.10.1 4.10.2

Introduction, History Structure, Nomenclature Isolation from Natural Substrates, Purification Physical Chemical Properties Chemical Synthesis Commercial Forms and Applications Analysis General Biological Methods Fetus Resorption Test on the Rat Encephalomalacia Test on Chicks Myopathy Test Liver Storage Test Erythrocyte Hemolysis Test in the Rat Biological Methods : Direct Measurement of Products of Lipid Peroxidation Malonic Dialdehyde Test on the Rat Ethane and Pentane Exhalation Test on the Rat Physical-Chemical Methods Biosynthesis Biochemical Role Aggressive Oxygen Forms in Animal Organisms Some Properties of the Aggressive Forms of Oxygen On the Formation of Aggressive Oxygen in the Animal Cell and the Protective Role of Vitamin E Enzymatic Defense Systems against Agressive Oxygen Molecular Mechanism of the Autooxidation of Multiply Unsaturated Fatty Acids On the Mechanism of the Antiperoxidative Effect of Vitamin E and Other Antioxidants Vitamins E and C as Antioxidants in the Plant On the Synergistic Interaction of Vitamins E and C in the Mammal On the Synergism between Selenium and Vitamin E Vitamin E as a Specific Protection Factor in the Biological Membrane On the Role of Vitamin E in the Erythrocytes and Blood Platelets Vitamin E and Protein Synthesis On the Anticarcinogenic Effect of Vitamin E Vitamin E and the Neuromuscular System Other Effects Structural Specificity of the Biological Effect Relative Biological Activities Relative Antioxidant Activities Occurrence, Including Occurrence in Foods Vitamin E in the Plant (Except in Fruits and Vegetables) Vitamin E in Fruits and Vegetables

218 4.10.3 4.10.4 4.10.5 4.11 4.12 4.12.1 4.12.2 4.12.3 4.12.3.1 4.12.3.2 4.12.3.3 4.12.3.4 4.12.3.5 4.12.3.6 4.12.4 4.13 4.13.1 4.13.2 4.14 4.14.1 4.14.1.1 4.14.1.2 4.14.1.3 4.14.1.4 4.14.2 4.14.2.1 4.14.2.2 4.14.2.3 4.14.3 4.14.4 4.15 4.15.1 4.15.2 4.15.3 4.15.4 4.16 4.16.1 4.16.2 4.16.3 4.16.4 4.16.5 4.17 4.17.1 4.17.2 4.17.3 4.17.4 4.17.5 4.18

4 Vitamin E Vitamin E in Grains and Grain Products Vitamin E in Plant Oils and Animal Fats Vitamin E in Common Foods Stability in Foods Absorption, Transport, Distribution, Storage, Metabolism and Excretion Intestinal Absorption and Uptake into the Blood Transport Mechanisms Distribution in the Body; Storage Whole Body Plasma Milk Erythrocytes, Thrombocytes (Blood Platelets) and Leukocytes Fat Tissue Brain tissue Metabolism and Excretion Requirements Human Requirements Requirements of Animal Organisms Deficiency and Deficient Utilization Causes of Deficiency in Human Beings Malabsorption Diseases of the Hepatobiliary System Hemolytic Anemias Neonates, Premature Infants Deficiency Symptoms in Human Beings Neuromuscular Anomalies Hemolytic Anemia in Premature Infants Retrolental Fibroplasia Deficiency Symptoms in Animals Diagnosis, Determination of Vitamin E Status Congenital Disorders of the Transport and Function of Vitamin E Abetalipoproteinemia Cholestasis Syndrome Cystic Fibrosis Muscular Dystrophies Therapeutic Applications of Vitamin E Patients with Malabsorption Premature Infants with Retrolental Fibroplasia Patients with Cardiovascular Disorders Patients with Chronic Hemolysis Other Therapeutic Effects Toxic Effects Allergies High Doses of Vitamin E in Adults High Doses of Vitamin E in Neonates and Premature Infants Vitamin E as an Antagonist of Vitamin Κ High Doses of Vitamin E in Animals References

4 Vitamin E

4.1 Introduction, History Around 1922, it was discovered that a purified diet containing adequate amounts of proteins, fats, carbohydrates, mineral salts and the vitamins A, B,, C and D (no other vitamins were known at the time) would support growth and well-being of laboratory rats, but not the male and female reproductive functions1. The animals displayed an atrophy of the reproductive organs, and fetal death and resportion were frequent. In 1922, H. M. Evans and K. S. Bishop reported the existence of a new, fat-soluble nutritional factor, first called Factor X, which prevented fetal death. The same authors found that wheat germ and lettuce were good sources of this fat-soluble factor. Three years later (in 1925), Evans wrote: "We have adopted the letter E as the next serial alphabetical designation, the antirachitic vitamin now being known as vitamin D." By about 1927, vitamin E2 was finally recognized as an essential dietary factor required for the establishment and maintenance of fertility in male and female rats (H. M. Evans, G. O. Burr, 1927). In the next 10 to 20 years, efforts were made to find a practical application for the new vitamin in the prevention or cure of reproductive disorders in veterinary and human medicine, but without success. The first thoughts on the role of vitamin E as an antioxidant were published in 1931 (H.S. Olcott, H.A. Mattili). About the same time, encephalomacia in vitamin-E-deficient chicks and "alimentary muscular dystrophy" in vitamin-E-deficient guinea pigs and rabbits were described (A. M. Pappenheimer, M. Goettsch, 1931). In the first 15 years of vitamin E research, there were considerable difficulties, due in large measure to the lack of vitamin E preparations with high activity. The most active preparation was wheat germ oil. There were also no good biological assay systems for the vitamin E levels in natural products. The situation changed abruptly when Evans and coworkers isolated an alcohol with high vitamin E activity from wheat germ oil in 1936. This had the summary formula C29H50O2 and was named by them α-tocopherol (tocos=birth). In the following year they reported the isolation of two further tocopherols from plant oils, which they named β- and γ-tocopherol; they found that these had less biological activity than α-tocopherol. At the same time, H.S. Olcott and O.H. Emerson (1937) discovered that these tocopherols are effective antioxidants, although the antioxi1

2

Historical reviews : Bieri & Farrell (1976); Janiszowska & Pennock (1976); Mason (1977) ; Machlin (1984); Chow (1985) Reviews: Lubin&Machlin(1982); Vitamin Compendium(\9%0)\Isler& Brubacher(1982); Bieri et al. (1983); Porter & Whelan (1983); Herting (1984); Machlin (1984); Chow (1985)

220

4 Vitamin E

dant and biological activities are not parallel. The structure of α-tocopherol was determined by E. Fernholz (1938), and the substance was first synthesized by P. Karrer (1938). Much later than the first three tocopherols, δ-tocopherol was reported (1947); and still later, about 1959, tocotrienols were described. The latter differ from the tocopherols by the presence of three double bonds in their side chains. Although vitamin E has been considered an essential dietary factor for many animals for several decades, it was first recognized as an essential component of human nutrition by the Food and Nutrition Board in 1968. For a long time the vitamin was not believed to have a special function in human beings. Today these doubts have been satisfied and the only remaining uncertainty concerns the daily requirement. There is still great uncertainty concerning the mechanism of action of vitamin E. Many theories have been developed, but final proof is lacking. The role of vitamin E as an essential antioxidant in the protection of highly unsaturated lipids has been generally accepted, but this antioxidant theory is apparently not able to explain all the effects of vitamin E. The long delay in the recognition of vitamin E as an essential nutritional factor for human beings is due mainly to the fact that avitaminosis E as a result of inadequate diet is practically unknown in the well-nourished countries. In addition, it is difficult to create an avitaminosis E in an adult human being because there are large reserves of this vitamin in the tissues and organs. We now know, however, that avitaminosis E can occur as a result of malabsorption, and that it is common among premature infants, who have only minimal reserves of vitamin E. The study of avitaminosis E has been carried out mainly in rapidly growing small animals. For these, there are a number of typical deficiency manifestations, e. g. myopathy and encephalomalacia, which often occur very soon after the vitamin is withdrawn. Most of these symptoms of a vitamin E deficiency have not been observed in human beings to date. One notes at present an increasing interest in research on the role of vitamin E in human metabolism. The role of vitamin E is especially clearly seen in patients with disorders of fat absorption.

4.2 Structure, Nomenclature Vitamin E occurs in nature as a group of substances including eight chemically similar compounds which are derivatives of 6-chromanol. Two to four methyl groups are bound to the chromanol ring, and in position 2 there is also a saturated or unsaturated isoprenoid C16 side chain. The eight natural vitamin E compounds fall into two groups, the four tocopherols (α-, β-, γ- and δ-tocopherol) with saturated side chains (Fig. 4-1), and the four tocotrienols (α-, β-, γ- and δtocotrienol) with unsaturated side chains (Fig. 4-2). The basic skeletons of tocopherols and tocotrienols without methyl groups on the aromatic rings do not occur in nature; they are called "tocol" (Fig.4-1) and "tocotrienol" (Fig.4-2), respectively.

4.2 Structure, Nomenclature

1

R2

5,78-Trimethyl

CH 3

CH 3

CH;

5,8-Dimethyl

CH 3 Η

Η

CH 3

CH; CH·

8-Methyl

Η

Η

Tocol

Η

Η

Tocopherol

-Tocol

aß76—

Fig. 4-1. (1971)]

R

7,8-Dimethyl

R;

CHH

Structural formulae of α-, β-, γ- and δ-tocopherol and of tocol. [from Mayer & Isler

^

-Tocotrienol

RI

R2

CH,

R;

Α-

5,7,8-Tri methyl

CH 3

CH 3

CH:

ß-

5,8-Dimethyl

7Δ-

7,8-Dimethyl

CH 3 Η

Η CH 3

CH; CH;

8-Methyl

Η

Η

CH;

Tocotrienol

Η

Η

H

—

221

Fig. 4-2. Structural formulae of α-, β-, γ- and δ-tocotrienol, and of tocotrienol. [from Mayer & Isler (1971)]

(2 R,L'R, 8'R)-a-Tocopherol

Fig. 4-3.

Stereochemistry of RRR-a-tocopherol. [from Mayer & Isler (1971)]

The stereochemistry of the tocopherols with the three asymmetric centers is illustrated in Fig. 4-3, using α-tocopherol as an example [Isler & Brubacher (1982)]. In the tocols, the saturated side chain together with C-atoms 2 , 3 and 4 and the methyl group on C-2 of the chromanol ring corresponds to the carbon skeleton of phytol. This side chain is unsaturated at positions 3', 7' and 11' in the tocotrienols. The individual tocols and tocotrienols differ within the group only with respect to the number and positions of the methyl groups on the aromatic ring.

222

4 Vitamin E

The term vitamin E applies to all derivatives of tocol (Fig. 4-1) and tocotrienol (Fig. 4-2) which have the qualitative biological activity of α-tocopherol. This term is also used in figures of speech such as "vitamin E deficiency", "vitamin E activity", "vitamin E antagonist", etc. The following rules apply: 1. The α-tocopherol with the configuration 2R, 4'R, 8'R (Fig. 4-3), also formerly called d-a-tocopherol, shall now be called RRR-a-tocopherol. 2. The diastereoisomer with the configuration 2S, 4'R, 8'R is called 2-epi-a-tocopherol. 3. A mixture of RRR-a-tocopherol and 2-ep/-a-tocopherol obtained by synthesis from phytol and the corresponding hydroquinone, is called 2-ambo-a-tocopherol (not dl-a-tocopherol). 4. The totally synthetic vitamin E obtained without any control of the stereochemistry is a mixture of eight diastereoisomers; it is called all-rac-α-tocopherol. Esters of the tocopherols and tocotrienols are called tocopheryl esters and tocotrienyl esters [IUPAC-IUB (1982)].

4.3 Isolation from Natural Substrates, Purification The most important natural sources of vitamin E are steam distillates obtained in the refining of food oils from plants. Because the α-tocopherol is usually mixed with the less valuable tocopherols (β-, γ- and δ-tocopherol) and the tocotrienols, these are converted to α-tocopherol by methylation and hydrogénation. Molecular distillation is used to enrich vitamin E [Isler & Brubacher (1982), Herting (1984)]. An example is discussed below. A pre-purified soy oil with 0.19% of a mixture of α-, γ- and δ-tocopherol is subjected to molecular distillation. The tocopherol fraction which distills below 240°C at 0.004 mm Hg is collected. One removes as many contaminants (mostly steroids) as possible by crystallization from acetone at — 10°C. Glycerides are then removed by hydrolysis. The unsaponifiable fraction containing the tocopherols is then subjected to another molecular distillation. A fraction is obtained which contains at least 60% mixed tocopherols. The non-a-tocopherols can be converted to RRR-a-tocopherol by methylation of the aromatic ring [Herting (1984)].

The extraction and purification steps often serve for analysis, e.g. in the determination of the vitamin E contents of foods and feeds. The method usually consists of the following steps [Isler & Brubacher (1982)]: 1. Extraction of the E vitamins with non-polar solvents and simultaneous alkaline saponification of the tocopheryl acetate which is present. One works under an inert atmosphere, with cooling and the addition of antioxidants. 2. The extract is purified by chromatography (mainly HPLC), steam distillation or molecular distillation, and the amount of α-tocopherol is determined, preferably according to Emmerie-Engle (p. 229). Extraction of the E vitamins from foods, e.g. from grain products, and preparation of the sample for HPLC has been described in detail [Cort et al. (1983)].

4.4 Physical Chemical Properties

223

4.4 Physical Chemical Properties The tocopherols are light yellow substances which are oils at room temperature; some of them crystallize at lower temperatures. α-Tocopherol melts at 2.5-3.5°C, and its acetate melts at 26.5-27.5°C. The tocopherols are insoluble in water, and readily soluble in non-polar solvents [Isler & Brubacher (1982)]. The tocopherols are monoethers of a hydroquinone, and as such, are easily oxidized. By contrast, the esters of the tocopherols, such as the acetate and succinate, are much more stable to air oxygen. Peroxides, ozone and potassium permanganate destroy tocopherols. The oxidation is catalysed by light, and is accelerated by unsaturated fatty acids, metal salts and bases. The tocopherols are very resistant to acids, however, and to bases in the absence of O2· α-Tocopherol is converted to a-tocopherylquinone (Fig. 4-4) by FeCl3. Although a-tocopheryl acetate is rather stable in air, it is saponified to the free α-tocopherol by moisture in an alkaline or acidic medium, and is rapidly oxidized in the presence of air, giving rise to a dark color. α-Tocopherol and its acetate are quite resistant to heat [Isler & Brubacher (1982)]. RRR-a-tocopherol has a weak specific optical rotation [a]o = + 0.32 (in ethanol). The UV absorption spectra of all tocopherols and their acetates in ethanol have a maximum between 280 and 300 nm, and a minimum at 250-260 nm [Isler & Brubacher (1982)]. The UV spectrum of α-tocopherol in ethanol has λ™* at 292 nm (Ε) 1 ^ = 75.8) and a λ,,,ίη at 256 nm. Acetylation of the phenolic hydroxyl group shifts λ™χ to 284 nm ( £ 1 ^ = 43.6) and Cin to 255 nm [Hertig (1984)]. The IR, Ή-NMR, 13C-NMR and mass spectra of the tocopherols have been published [Isler & Brubacher (1982)]. α-Tocopherol is localized primarily in the subcellular membranes of the mitochondria, microsomes and chloroplasts of animals and plants. Synthetic membranes have been made to study the nature of the embedding of the tocopherols in the native membranes. It was shown that α-tocopherol is present mainly in non-ionized form if it is present in a micelle at physiological pH. In addition, the chromanol ring of the vitamin E may lie in the phase boundary between the membrane structure and the water [Drummond & Grieser (1985)].

(2 R, WR, 8'/?)-α-Tocopherol FeClj

W . .J

Fig. 4-4. Oxidation of RRR-a-tocopherol by FeClj to a-tocopherylquinone [from Mayer & Isler (1971)]

224

4 Vitamin E

HC OH

HO

HC

Fig. 4-5. Technical synthesis of all-raoα-tocopherol, starting from trimethylhydroquinone and isophytol [from Isler & Brubacher (1982)]

4.5 Chemical Synthesis The oldest synthesis of vitamin E was published almost simultaneously by several research groups in 1938. It is based on the condensation of trimethylhydroquinone with phytol or phytyl halide. Later, synthetic isophytol was used; this is now preferred in the industrial process. The product of isophytol and trimethylhydroquinone is a//-rac-a-tocopherol (Fig. 4-5), a mixture of eight possible diastereoisomers. The synthesis starting from natural phytol leads to 2-ambo-a-tocopherol. The configuration at C-2 is of eminent significance for the biological activity; 2-epj-a-tocopherol has only 21% of the activity of RRR-a-tocopherol. RRR-a-tocopherol and 2-e/w-a-tocopherol can be separated by fractional crystallization, and RRR-a-tocopherol can now be made by direct synthesis. Many other methods, including some for production of labelled tocopherols, have been reported [Herting (1984), Isler & Brubacher (1982)]. An important process for partial chemical synthesis of α-tocopherol is methylation and hydrogénation of mixtures of E vitamins obtained from refining of food oils from plants. In addition to α-tocopherol, these contain β-, γ- and δ-tocopherols, as well as tocotrienols. Many methods of methylation have been developed and tested [Isler & Brubacher (1982)]. A stereoselective synthesis of the vitamin E side chain has been described [Koreeda & Brown (1983), Helmchen & Schmierer (1983)]. In addition, the chemical synthesis of (S)-chromanomethanol, which can be used as a precursor for chemical synthesis of vitamin E, has been worked out [Takabe et al. (1985)].

4.6 Commercial Forms and Applications a-Tocopheryl acetate, the quantitatively most important form of vitamin E produced, is used as a feed and dietary supplement and in medicine. Unesterified vitamin E is used as an antioxidant in food technology (mainly to prevent fats and oils from becoming rancid), and in the production of some pharmaceuticals. In pharmaceutical preparations, the tocopherols stabilize vitamin A and unsaturated lipids [Herting (1984)]. α-Tocopherol and its esters are components of multivitamin preparations. Here crystalline a-tocopheryl hydrogensuccinate is often used. α-Tocopherol or

4.7 Analysis

225

its esters, dispersed in hydrophilic media, are given orally to patients with malabsorption; however, they can also be used for injections. RRR-a-Tocopheryl-polyethylene glycol-succinate has a wax-like consistency; it forms clear aqueous solutions in concentrations up to 20% without the need for a solubilizer. Supplente for human use range from a few mg in multivitamin preparations to 500-1000 mg. There are also salves and suppositories which contain vitamin E [Herting (1984)]. Because symptoms of a vitamin E deficiency are relatively rare in human beings, there has so far been no extensive enrichment of foods with this vitamin. However, most baby foods are supplemented with vitamin E, as are some grain products and feeds for chickens and turkeys (here, however, mostly to prevent rancidity). Animal feeds, especially those for poultry, consume about 40% of the commercial production of vitamin E [Herting (1984)]. For clinical purposes the main product is all-rac-a-tocopheryl acetate; however, oral preparations often contain RRR-a-tocopheryl acetate. In addition to the acetate form the succinate form is also commercially available. The esters are hydrolysed in the intestine before they are absorbed. Aqueous suspensions are produced for patients with impaired intestinal absorption; these are more readily absorbed [Bieri et al. (1983)]. In 1980, the world production of vitamin E (both natural and synthetic) was about 5300 metric tons. The US production in 1980 was 3297 tons. The price of vitamin E in 1982 was $68 (US)/kg for RRR-a-tocopheryl acetate, and $27/kg for all-rac-a-tocopheryl acetate [Herting (1984)].

4.7 Analysis Numerous methods are available for analysis of vitamin E. Biological methods are based on the ability of certain animals to respond especially rapidly and specifically to vitamin E deficiency. There are also biological methods based on the direct measurement of lipid peroxidation in vitamin-E-deficient animals. Finally, there are physical-chemical methods available; chromatography, and especially HPLC, is an essential tool. 4.7.1 General Biological Methods Many animal species react sensitively, and often quite specifically, to dietary vitamin E deficiency. Table 4-1 gives several examples. Administration of vitamin E can prevent or cure the symptoms. Comparison of the effects of known amounts of vitamin E and the test samples with unknown vitamin E activities allows the latter to be estimated [Chow (1985); Buckingham (1985)]. The preparation of the test animals for the analysis is usually time-consuming. There are numerous criteria which can be used to evaluate the biological activities of vitamin E samples [Chow (1985)]. The following are most often used; the test animals are usually rats and chicks:

226

4 Vitamin E

Table 4-1. Pathology of vitamin E deficiency [simplified from Chow (1985)] Affected tissue; observed change

Animal species

Embryonic degeneration (damage to vessel system)

Female rats, sheep, hens, turkeys

Degeneration of male gonads

Male rats, guinea pigs, hamsters, dogs

Encephalomalacia

Chicken

Myopathy

Rabbit, guinea pig, monkey, duck, rat, chick, turkey

Erythrocyte hemolysis

Rat, chicken

Liver necrosis

Rat, pig

1. Resorption gestation test 2. Encephalomalacia test 3. Myopathy test 4. Liver storage test 5. Erythrocyte hemolysis test 4.7.1.1 Fetus Resorption Test on the Rat The classical procedure for determining vitamin E activity is a modification of the fetus resorption test (resorption gestation method). Female rats are fed a vitamin-E-free diet for 9-10 weeks, and are then bred to normal males. The females are then given graduated amounts of the sample to be tested between the 4 th and 8 th days after breeding. Conception and the first half of pregnancy are normal. However, if no vitamin E is given during the first 10-12 days of the pregnancy, the embryos die and are resorbed. The female remains apparently healthy and can be reused repeatedly. If the added test dose of α-tocopherol exceeds a critical amount, the embryos develop normally. The critical dose of α-tocopherol is 0.3-1.0 mg; less than this amount leads to embryonic death [Herting (1984)]. 4.7.1.2 Encephalomalacia Test on Chicks Alimentary encephalomalacia of the chick is a nerve disorder associated with ataxia, characteristic head position and uncoordinated motion. If vitamin E is not obtained, the disease is fatal. The diagnosis can also be made by histopathological examination of the cerebellum [Hakkarainen et al. (1984)]. Due to closure of vessels in this disease, the nerve substance dies. 4.7.1.3 Myopathy Test Vitamin E is essential for normal muscle function. Myopathy (also called muscular dystrophy) is a prominent symptom of vitamin E deficiency in some animals [Bruce et al. (1985)].

4.7 Analysis

227

In chicks, the muscle degeneration is directly determined 3 to 4 weeks after vitamin E is withdrawn, and substituted by the test dose. The muscle degeneration can also be determined indirectly, as it is associated with a large decrease in many muscle enzymes, which can be measured in the plasma. In some animal species, one also observes a pronounced creatinuria [Machlin (1984); Bässler & Lang (1981)]. The pyruvate kinase activity in the plasma of the rat is often used as an index of myopathy [Machlin et al. (1982)]. The activity of the pyruvate kinase in the plasma of vitamin-E-deficient rats is markedly elevated, and the degree of this elevation corresponds to the severity of the myopathy. Electrophoresis of the plasma of rats with vitamin E deficiency has shown that the increase in pyruvate kinase is due to the increase in an isoenzyme of the M type. Because administration of small doses of vitamin E to the rat cures the myopathy and causes the activity of the pyruvate kinase in the plasma to drop, the curative myophathy rat test was developed on the basis of the reduction of the pyruvate kinase activity in the plasma by vitamin E [Chen & Thacker (1985 a)]. 4.7.1.4 Liver Storage Test This test is based on the observation that the liver levels of vitamin E in rats and chicks is linearly correlated with the vitamin E level in the diet. Chicks with vitamin E deficiency are fed test doses for several days, and then the liver level of vitamin E is determined [Machlin (1984)]. 4.7.1.5 Erythrocyte Hemolysis Test in the Rat The erythrocyte hemolysis test is based on the fact that adult rats with adequate vitamin E in their feed have erythrocytes which are largely resistant to chemically induced hemolysis in vitro. Hemolysis is usually induced with dialuric acid and hydrogen peroxide. Administration of a diet free of vitamin E leads to erythrocytes which are sensitive to hemolysis. The normal test is based on administration of various amounts of vitamin E (the standard and the sample to be tested) to the animals during the period of dietary deprivation. The degree of hemolysis of the erythrocytes is then measured in vitro [Herting (1984)]. About 40-44 hours after application of the test dose, dialuric acid is added to the washed erythrocytes, and the percent which have hemolysed is determined [Machlin (1984)]. 4.7.2 Biological Methods: Direct Measurement of Products of Lipid Peroxidation Lipid peroxidation in animals with vitamin E deficiency forms lipid hydroperoxides which can be degraded to a series of products; these include aldehydes (especially malonic dialdehyde), unsaturated aldehydes, ketones, alkanes (mainly ethane and pentane), etc. Several methods for measurement of these products of lipid peroxidation have been reported. Some methods are discussed below, and two methods of eminent significance are the subjects of the following two sections. Decrease in the lipid substrates. These can be measured in vitro by gas chromatography of organelles, cells and tissue preparations.

228

4 Vitamin E

Measurement of lipid hydroperoxides. This is done by chemical methods, such as iodometric titrations, and by enzymatic methods using peroxidases. The methods are still relatively rarely used. Fluorescence analysis of the products of lipid peroxidation. Amino acids react with malonic dialdehyde, a product of lipid peroxidation, to form Schiffs bases which absorb light strongly and fluoresce; this can be used for analytical purposes. Chemiluminescence. Lipid peroxidation in tissue fractions is associated with light emission, which can be measured. The light emitted during lipid peroxidation is probably due mainly to the formation of singlet oxygen 1 0 2 and its return to the ground state (O2). The wavelengths of these emissions are in the long-wave (red and infrared) region [Slater (1984)]. 4.7.2.1 Malonic Dialdehyde Test on the Rat Malonic dialdehyde is a product of lipid peroxidation in vivo. Rats with vitamin E deficiency have an elevated concentration of malonic dialdehyde in their organs and tissues ; this can be conveniently measured with thiobarbituric acid. The level of malonic dialdehyde is also taken as an index of rancidity of fats [Chan (1985), Bird & Draper (1984)]. To determine malonic dialdehyde, one first treats tissue samples with trichloroacetic acid; after centrifugation, the supernatant is heated with thiobarbituric acid, then cooled and the absorption at 532-535 nm is measured. Although this reaction measures malonic dialdehyde, a number of other substances also react with thiobarbituric acid under the above conditions. The interference is probably relatively slight, however, as was shown by control reactions [Slater (1984)]. Methods for determination of malonic dialdehyde (mainly spectrophotometry, fluorometry and HPLC) have been described [Bird & Draper (1984)]. 4.7.2.2 Ethane and Pentane Exhalation Test on the Rat Volatile hydrocarbons like ethane and pentane are products of the peroxidation of multiply unsaturated fatty acids of membrane lipids, and their formation is considered a sensitive index of lipid peroxidation. After application of xenobiotics, increased amounts of alkanes are registered in the exhaled air from animals with vitamin E deficiency. The main site of formation of alkanes (and of malonic dialdehyde) is the liver. Of the two hydrocarbons ethane and pentane, the latter is a less reliable indicator for measurements in vivo, because it can be metabolized in the liver. This is not the case for ethane [Müller & Sies (1984)]. Ethane and pentane are products of the peroxidative degradation of linolenic and linoleic acids, respectively. The formation of these volatile hydrocarbons, which are exhaled in elevated concentrations by vitamin-E-deficient rats, is reduced or eliminated by re-establishment of the normal vitamin E status [Fraser (1985)]. The measurement of the exhaled ethane was first carried out on the mouse in vivo after it was treated with carbon tetrachloride (to induce lipid peroxidation). It was shown that pretreatment of the mouse with vitamin E effectively reduced the formation of ethane [Lawrence & Cohen (1984)].

4.7 Analysis

229

The exhalation of ethane and pentane corresponds to the degree of lipid peroxidation in vivo. Ethane and pentane have already been measured in perfused organs, cell suspensions and tissue homogenates. Tissue slices seem especially suitable here. Total ethane and pentane (TEP) are released from tissues of vitamin-E-deficient rats in the following order: intestine = brain = kidney > liver = lung > heart > testis = diaphragm > skeletal musculature. The ability of halogenated hydrocarbons to release TEP from slices of liver from vitamin-E-deficient rats decreases in the following order: CBrCl3 >CCl4=l,l,2,2-tetrabromoethane = 1,1,2,2-tetrachloroethane > perchloroethylene. CBrCl3 also stimulates the release of TEP from many other organs, which are thus sites of attack by the toxic CBrCl3. Dietary vitamin E reduces the release of TEP from the liver and kidneys. Doses of iron increase TEP excretion from all tested rat organs except the brain [Gavino et al. (1984)].

4.7.3 Physical-Chemical Methods The following methods are used for determination of vitamin E, especially in preparations with simple compositions: 1. UV spectroscopy. It is suitable only for relatively pure, highly concentrated preparations with little foreign material which interferes with UV absorption. The determination of the difference in absorption of α-tocopherol and a-tocopherylquinone (after oxidation) has proven useful. 2. Emmerie-Engel reaction. This is based on the reduction of two Fe(III) ions to Fe(II) ions by one molecule of free α-tocopherol. The Fe(II) is determined colorimetrically at 520 nm as a red complex with 2,2'-dipyridyl. More recently, 4,7-diphenyl-l,10-phenanthroline, which gives a still more intense color, has been used instead of 2,2'-dipyridyl. 3. Polarographic determination of α-tocopherol after its oxidation to a-tocopherylquinone. 4. Fluorescence measurement. Solutions of free (non-esterified) tocopherols and some of their reaction products have an intense fluorescence in the UV range; this can be measured. The method is specific and sensitive. 5. Gas chromatography (GC) and high-pressure liquid chromatography (HPLC). These are excellent methods for separation; they are used preferentially for pharmaceuticals. All vitamin E mixtures can be separated by HPLC. [Isler & Brubacher (1982)]. The methods for analysis of the tocopherols by thin-layer chromatography and HPLC have been critically compared. The unreliability of the Emmerie-Engel reaction (mainly the side reactions with Carotinoids and cholesterol) has also been pointed out [Ruggeri et al. (1984)]. The deterimination of vitamin E in animal tissues, including blood, by thin-layer chromatography, spectrophotometry and spectrofluorimetry have been described [Desai (1984)]. An automated colorimetrie method for determination of α-tocopherol in the serum has also been described [Sloan & Lappin (1982)], as has a microchemical method for determination of picogram amounts of vitamin E in small (about 3 μg) tissue samples [Buntman et al. (1984)]. The HPLC methods developed in recent years make it possible to measure extremely small amounts (down to 1 ng) of individual tocopherols in biological samples, e.g. plasma, and in foods and feeds [Chow (1985)]. Numerous applica-

230

4 Vitamin E

tions of HPLC for the analysis and isolation of vitamins of the E group have been worked out. A few applications are cited below. Determination of tocopherols and tocotrienols in plasma and foods [Piironen et al. (1984a,b)]; simultaneous determination of vitamins E and A, also in human serum [Miller & Yang (1985); Driskell et al. (1982)]; determination of vitamin E in plasma and tissues [Heng et al. (1983); Zaspel & Csallany (1983)]; simultaneous determination of tocopherols and cholesterol in erythrocytes and plasma [Stump et al. (1984)]; determination of vitamins of the E group [Buttriss & Diplock (1984)]; qualitative and quantitative determination of tocopherols and tocotrienols [Arens et al. (1984)].

Finally, the application of chromatographic methods for determination of vitamin E in serum [Turley et al. (1985)] and for simultaneous determination of vitamins E and A, also in serum, have been described [Williams (1985); Catignani & Bieri (1983)].

4.8 Biosynthesis The aromatic ring of vitamin E arises from shikimic acid, which yields homogentisic acid via several intermediates; homogentisic acid is almost entirely incorporated into vitamin E. Starting from homogentisic acid, the biosynthesis can proceed by at least two paths, the tocopherol and the tocotrienol routes. The former is main route; it takes place in green leaves and generally in most plant tissues, and in algae. In it, phytyl diphosphate condenses with homogentisic acid to δtocopherol; the methyl group in position 8 of δ-tocopherol is derived from the acetate residue of the homogentisic acid. Methylation by S-adenosylmethionine leads via (primarily) γ-tocopherol or ß-tocopherol to α-tocopherol. In the tocotrienol route, homogentisic acid condenses with geranylgeranyl diphosphate to δtocotrienol. Methylations with S-adenosylmethionine lead (primarily) to γ-tocotrienol and (partially) to ß-tocotrienol, and from these to α-tocotrienol. The last step of this route, which is observed in latex and elsewhere, α-tocopherol is formed by hydrogénation (Fig. 4-6) [Janiszowska & Pennock (1976); Isler & Brubacher (1982)]. The biosynthesis of the side chain follows the pathway of terpene synthesis up to geranylgeranyl diphosphate. In the tocopherol pathway, which is found in the green leaves and many other organs of the higher plants, and in algae, the C2o terpene chain is first hydrogenated to phytyl diphosphate, and then condensed with homogentisic acid to 2-methyl-6-phytylhydroquinone (also called 6-phytyltoluquinol). Cyclization and two methylations lead via δ-tocopherol to α-tocopherol, as can be seen in Fig. 4-6. In the tocotrienol pathway, the side chain is built in analagous fashion up to geranylgeranyl diphosphate [Isler & Brubacher (1982)]. That the biosynthesis of tocopherols proceeds via 2-methyl-6-phytylhydroquinone in the chloroplasts has been reconfirmed [Marshall et al. (1985)]. Homogentisic acid is formed from 4-hydroxyphenylpyruvate by the action of 4-hydroxyphenylpyruvate dioxygenase [Fiedler et al. (1982)]. The steric mechanism of the decarboxylation of homogentisic acid has been followed using stereospecifically labelled homogentisic acid as a

4.9 Biochemical Role Tocotrienol route

231

Tocopherol route

OH

ÒH Homogentisic acid

Geranylgeranyl P P

H3C-

Phytyl P P

HoC

I

OH V

¡A

OH

I V

173

6-Geranylgeranyltoluquinol

6-Phytyltoluquinol

δ-Tocotrienol

δ-Tocopherol

/3-Tocotrienol

7-CHo

CH,

5-CH3/

y-Tocotrienol

6H

5-CH3

7-CHÍ a-Tocotrienol

/9-Tocopherol

7-Tocopherol

7-CHK

yî-CHj

a-Tocopherol

Fig. 4-6. Probable course of the biosynthesis of tocopherols and tocotrienols [Janiszowska & Pennock (1976)] precursor. The result was that the decarboxylation occurs with stereochemical retention during the biosynthesis [Kriigel et al. (1985)].

It is thought that there are two sites of tocopherol synthesis in the plant cell: the chloroplast, where mainly α-tocopherol is formed, and outside the chloroplasts, where mainly δ- and γ-tocopherol are formed [Janiszowska & Pennock (1976), Isler & Brubacher (1982)]. Most prokaryotes and yeasts form very little or no vitamin E. Among the most interesting microorganisms which form vitamin E is the unicellular phytoflagellate Euglena gracilis Z, and experiments on producing vitamin E in this way have been undertaken [Ruggeri et al. (1985)].

4.9 Biochemical Role Several decades of vitamin E research have shown that it is essential for the maintenance and function of probably all animal cells3. A deficiency of vitamin E leads often or primarily to destabilization of biological membranes [Buttriss & Diplock (1984)]. The exact mechanism of vitamin E action is not yet known. 3

Review: Simon-Schnaß & Koeppe (1983); Simon-Schnaß (1984)

232

4 Vitamin E

However, evidence is accumulating that it interacts with cellular membranes. It is a component of such biological membranes as the inner mitochondrial membrane and the outer membrane of the red blood cell [Chow (1985)]. It has already been generally accepted that the primary role of vitamin E in tissues is the protection of lipids from peroxidation. This effect in vivo has, however, been definitely proven only for fat tissue, and final proof is lacking for other tissues. In addition, there are other cellular systems for defense against autooxidation, especially the glutathione peroxidase system [Chow (1985)]. It is assumed that vitamin E localized in biological membranes inhibits peroxidation of membrane-bound, multiply unsaturated fatty acids. In addition to erythrocyte hemolysis, a vitamin E deficiency causes damage to the ultrastructure and integrity of mitochondrial membranes, endoplasmic reticulum and nuclear membranes. Damage to the ultrastructure of the blood platelets was observed in vitamin-E-deficient rats [Chow (1985)]. The opinion that vitamin E acts as an antioxidant in biological systems is supported by the observation that some symptoms of vitamin E deficiency in animals can be prevented or cured by administration of synthetic antioxidants. In the absence of antioxidants, especially vitamin E, a peroxidation of polyenic acids occurs in the presence of oxygen. This is associated with a radical chain reaction, which is continuously accelerated by self-catalysis. These reasonably well understood primary processes are then followed by less well understood processes in which aldehydes (mainly malonic dialdehyde), ketones, hydroxy acids, ethane and pentane are formed [Bässler & Lang (1981)]. All attempts to explain the effect of vitamin E entirely by its antioxidative properties, i. e. by the ability of this vitamin to react with free radicals and thus to interrupt chain reactions, have so far failed [Isler & Brubacher (1982)]. In addition to its role as antioxidant of membrane lipids, vitamin E is thought by some authors to have a structural role in biological membranes: vitamin E can interact specifically with the arachidonic acid in the biological membranes, and thus modulate the properties of the membranes. According to another hypothesis, vitamin E can modulate the microviscosity of the membrane. These hypotheses are of little significance, because the approximate molar ratio of the unsaturated fatty acids to vitamin E in biomembranes is about 1000:1 [Massey et al. (1982)]. Finally, some authors suspect that vitamin E has, in addition to its role as antioxidant and membrane-stabilizing structural component, a catalytic and regulatory role in intermediary metabolism [Quintanilha et al. (1982)]. There is much evidence for the action of vitamin E in the maintenance of a normal neurological structure and function. Much data has been provided by studies on patients with chronic impairment of fat absorption, and also by many animal experiments [Muller et al. (1985)]. 4.9.1 Aggressive Oxygen Forms in Animal Organisms Several forms of oxygen, most of them aggressive4, act on the cell. The main one is molecular oxygen, 0 2 , itself, which can react directly with lipid radicals, form4

Review: Packer(1984)

4.9 Biochemical Role

233

ing peroxyl radicals ROO. The superoxide anion radical 0 0 2 (usually called the superoxide anion) is formed very frequently, and it leads to the formation of other, often still more aggressive oxygen forms, such as the hydroxyl radical OH, the hydroperoxy radical H 0 2 , and H 2 0 2 . Finally, the very reactive singlet oxygen, should be mentioned. The oxygen metabolites which attack biological membranes are mainly 0 0 2 , OH and H 2 0 2 . Some animal cells contribute actively to the formation of aggressive forms of oxygen within the framework of their antibacterial action. The defense systems against the aggressive forms of oxygen (including the peroxyl radicals and hydroperoxides, ROOH) are vitamin E and some enzymes, especially superoxide dismutase (EC 1.15.1.1), catalase (EC 1.11.1.6) and glutathione peroxidase (EC 1.11.1.9). 4.9.1.1 Some Properties of the Aggressive Forms of Oxygen The reduction of dioxygen, 0 2 , which eventually produces water, involves the step-wise addition of four electrons. The intermediates in this reduction are damaging to the living cell. The one-electron reduction produces the superoxide anion, Oi [Diplock (1983 b)]. The above reduction is the order of the day in both abiotic and biotic systems; it is the most common route for reduction of 0 2 . In Escherichia coli, 0 2 production accounts for 3% of the total respiration. The mammalian liver produces 24 nmol 0 2 per minute and gram [Fridovich (1984)]. The hydroxyl radical, OH, is formed mainly by ionizing radiation in biological systems, and it is active in biological processes. It is very reactive; one of the reactions is the formation of H 2 0 2 by recombination (eq.l): ΟΗ + ΌΗ — H 2 0 2

(1)

It is known that many normal physiological processes like phagocytosis are linked to the formation of OH. Furthermore, the formation of OH is induced by many toxic agents. The most important pathway to formation of OH in biological systems appears to be the following reaction, in which iron participates (eq.2): Fe(III) + reductant (e.g. O2 or vitamin C) —• Fe(II) Fe(II)+H 2 02 — Fe(III)+ OH + OH-

(2)

[Czapski (1984)]. Singlet oxygen, 1 0 2 , is an excited form of molecular oxygen, and it can be converted to the latter by light emission. Because of its high reactivity, is among the more aggressive oxygen species in biological systems. It reacts with olefins to form hydroperoxides and other products. It can be formed during lipid peroxidation, as a result of the cleavage of lipid peroxides [Cadenas & Sies (1984)]. 4.9.1.2 On the Formation of Aggressive Oxygen in the Animal Cell and the Protective Role of Vitamin E In spite of the destructive effect of 0 2 in the living system, there are specialized cells in the blood which, when activated, generate 0 2 as the main product of the reduction of dioxygen. These are the phagocytes, which produce mainly O \ in

234

4 Vitamin E

the "respiratory burst", for the purpose of killing microorganisms. OH and H 2 0 2 are formed secondarily. The three dioxygen products, Oí, OH and H 2 0 2 , are extremely important defense substances against microbial infections [Fridovich (1984)]. These dioxygen products, however, can damage neighboring tissues as well as the cells which generate them; vitamin E is the main protection against this effect [Lafuze et al. (1983); Chow (1985)]. 0 2 is formed by NADPH oxidase, an enzyme of the neutrophilic leukocytes. This enzyme is inactive in resting cells, but it is activated when the cells are stimulated, e.g. by bacteria. The NADPH oxidase is a membrane-bound flavoprotein, and catalyses the reaction (eq.3): NAD(P)H + 202—NAD(P)+ + H++20i

(3)

[Markert et al. (1984)]. 4.9.1.3 Enzymatic Defense Systems against Agressive Oxygen In addition to vitamin E, there are several enzymes which protect against aggressive oxygen, mainly superoxide dismutase, glutathione peroxidase and catalase. Superoxide dismutase (EC 1.15.1.1) controls the level of the superoxide anion 0 2 ; the enzyme catalyses the following disproportionation (eq.4): 2 O2 + 2H+ —• H2O2 + O2

(4)

Superoxide dismutase is found both in the mitochondria (this form contains manganese) and in the cytosol (this form contains copper or zinc in eukaryotes) [Diplock (1983b, 1984)]. Superoxide dismutase offers protection against 0 2 in hydrophilic media, while in hydrophobic media (in the cell membrane) protection is probably offered by α-tocopherol. The correct localization of the α-tocopherol in the membrane is thus an important requirement for maximal protection [Ozawa & Hanaki (1985); Ozawa et al. (1983)]. Glutathione peroxidase (EC 1.11.1.9), a soluble, selenium-containing enzyme, found both in the mitochondria and in the cytoplasm, reduces hydrogen peroxide to water; the reducing equivalents come from glutathione (eq.5): H202 + 2 GSH —>- 2 H20 + GSSG

(5)

The activity of the glutathione peroxidase is dependent on the presence of selenium. The enzyme also catalyses the reduction of a number of lipid hydroperoxides to hydroxy derivatives (eq.6): ROOH + 2 GSH —»· ROH + GSSG + H20

(6)

[Diplock (1983 b, 1984)]. Catalase (EC 1.11.1.6) decomposes hydrogen peroxide to water and 0 2 in the peroxisomes (eq.7): 2 H2O2 —• 2 H2O+O2 (7) Catalase also acts as a peroxidase, using hydrogen donors (AH2) (eq.8): ROOH + AH2

BJC) + ROH + A

[Aebi (1984), Diplock (1984)].

(8)

4.9 Biochemical Role

235

4.9.2 Molecular Mechanism of the Autooxidation of Multiply Unsaturated Fatty Acids Lipid-containing foods, even when they are stored at low temperatures, can become rancid. This is a sign that they react with atmospheric oxygen. The process is called autooxidation. It has been investigated in a number of substances; the studies showed that it is a chain reaction in which free radicals are involved. If the organic oxidizable substance is RH (e. g. linoleic acid) and the radical formed from it is R·, the autooxidation consists of the following reactions (eq.9-12): Initiation RH —>• R + H Propagation R + 0 2 — R O O ROO + RH — ROOH + R Termination ROO + ROO—>· molecular products

(9) (10) (11) (12)

The initiation (eq.9) consists of the production of the radical R . The reaction can be initiated by, for example, heat, light or ionizing radiation. In the presence of oxygen, the radical R· is rapidly converted to the peroxyl radical ROO (eq. 10). The peroxyl radical attacks the organic substance, RH, in a much slower reaction which forms hydroperoxide, ROOH, and regenerates the radical R· (eq.ll); this radical adds 0 2 and thus continues the chain reaction. Many RH molecules are oxidized before the chain is terminated, for example by the reaction of two peroxyl radicals with one another (eq.12). Starting from a single free radical R, a large number of hydroperoxides ROOH can be formed. The hydroperoxides can in turn be converted to free radicals by abstraction of hydrogen, and thus add to their own number; so that the autooxidation proceeds exponentially. This process, which was described in eq.9-12, is illustrated in Fig.4-7 [Burton & Ingold (1983), Fraser (1985)]. The most common cause of initiation of the chain (eq.9) is the degradation of hydroperoxides, ROOH, which were formed previously by oxidation. Hydroperoxides are easily cleaved by light and by certain metal ions (eq. 13-15):

M

RH

/pH

m0H

bSH»0"0''

Fig. 4-7. Schematic representation of the autooxidation of multiply unsaturated fatty acids (e.g. linoleic acid). These are symbolized by RH, their hydroperoxides by ROOH and their peroxyl radicals by ROO. Vitamin E interrupts the radical chain reaction at 1 and hydroperoxidedestroying antioxidants act at 2 [from Fraser (1985)]

236

4 Vitamin E ROOH

hv

RO + OH

(13)

ROOH + Fe(II) — RO + OH" + Fe(III) +

ROOH + Fe(III) — ROO + H + Fe(II)

(14) (15)

The reactions of eq.14 and 15 form a catalytic cycle, so that very small traces of iron (or copper or cobalt) can initiate many chain reactions. One class of preventive antioxidants consists of chelating agents, which complex the heavy metals and thus make them ineffective [Burton & Ingold (1983)]. Aerobic organisms are exposed to much greater oxidative stress than, for example, foods in cold storage. In spite of this, they do not become rancid as long as they are alive. Living organisms have mechanisms which protect them from autooxidation. The greatest protection is probably required by the multiply unsaturated fatty acids of the membrane lipids. Autooxidation of the lipids of the biological membranes destroys the latter, and interrupts many life functions [Burton & Ingold (1983)]. Many toxic agents can be activated in the cell, forming free radicals which initiate the damaging lipid peroxidation. Light is an important initiator of free radicals, and of the peroxidation of polyenic acids, especially in the lens of the eye [Slater (1984); Varma et al. (1982)]. It is known that plants which contain peroxidizable lipids are able to form tocopherols, and in amounts which are about directly proportional to the concentration of polyene lipids. That antioxidants are necessary to protect nutritional fats from becoming rancid seems obvious. However, for a long time it was not so obvious that the same or related lipids, after they had become components of animal cells, need such protection [Horwitt (1976)]. 4.9.3 On the Mechanism of the Antiperoxidative Effect of Vitamin E and Other Antioxidants There are two basic types of antioxidants which can protect multiply unsaturated lipids from peroxidation. The preventive or primary antioxidants reduce the speed of initiation of autooxidation, usually by conversion of the hydroperoxides ROOH to the corresponding alcohols ROH. The chain interrupting or secondary antioxidants generally act by addition of a hydrogen atom to the peroxyl radical; these are mainly phenolic substances (ArOH) and include vitamin E (eq.16): ROO + ArOH — ROOH + ArO

(16)

The resulting ArO radical (for example the tocopheryl radical) is normally too unreactive to continue the chain reaction. However, it can react rapidly with a peroxyl radical, forming molecular products (eq.17): ROO + ArO —>· molecular products

(17)

In the end effect, every molecule of the antioxidant ArOH (and thus vitamin E) can capture two peroxyl radicals and interrupt two reaction chains [Burton & Ingold (1983); Lambelet & Löliger (1984); McCay (1985)]. To protect its products, nature uses both preventive and chain-interrupting antioxidants. Glutathione peroxidase, which can reduce a number of organic hy-

4.9 Biochemical Role

237

droperoxides, is among the best known preventive antioxidants; catalase is another (eq.6 and 8) [Burton & Ingold (1983)]. The biological chain-interrupting (secondary) antioxidants can be divided into two groups, depending on where they destroy radicals: in the aqueous or the lipid phase. The most important radical in the aqueous phase, which must be eliminated, is the superoxide ion, O2. It is not very reactive itself, but it forms the highly reactive hydroperoxyl radical (eq.18): O2 + H + ^ HOO

(18)

At physiological pH values, the above equilibrium (eq.18) favors the superoxide ion by about two orders of magnitude. The latter is decomposed by the enzyme superoxide dismutase (eq.4), while HOO· can be converted into H 2 0 2 by ascorbic acid [Burton & Ingold (1983)]. In the lipid phase of biological systems, vitamin E is probably practically the only chain-interrupting antioxidant. The individual tocopherols differ in their effectiveness; the relative effectiveness in vitro of α-, β-, γ- and δ-tocopherol are, respectively, 2.4, 1.7, 1.6 and 0.65. The corresponding values for two synthetic phenolic antioxidants are 2.1 and 0.21. α-Tocopherol is more efficient in vitro than any commercial phenolic antioxidant. Its effect in vivo is also very great [Burton & Ingold (1983)]. 4.9.4 Vitamins E and C as Antioxidants in the Plant The distribution of the tocopherols in plant tissue indicates that they have special functions. Three possible functions are discussed: 1. Protection against oxidants 2. Membrane stabilization 3. Initiation of flowering. In the plant, vitamin E probably protects mainly the lipid depots from peroxidation, as in animal cells, by interrupting radical chain reactions. A correlation was found between the amounts of unsaturated fatty acids and tocopherols in plant oils. In the chloroplasts α-tocopherol could protect sensitive unsaturated membrane lipids from photooxidation. It was also shown that α-tocopherol can induce flowering in some plants [Janiszowska & Pennock (1976)]. Numerous observations indicate that some herbicides, e.g. the bipyridylium salts, induce peroxidation of the biomembranes in higher plants and algae. There are several antioxidative systems which offer protection against damaging peroxidation of lipids, including a number of enzymes, low-molecular-weight substances like glutathione and ß-carotene, and above all, vitamins E and C [Finckh & Kunert (1985)]. Vitamins E and C protect the plant against peroxidation by aerial pesticides and against some herbicides. In one experiment, vitamin E prevented membrane damage in an alga and a higher plant treated with herbicide. Experiments have shown that the peroxidation of membranes depends directly on two factors, namely the absolute concentration and the relative amounts of vitamins E and C in the plant. Young transplants, which contain substantial amounts of both vi-

238

4 Vitamin E

tamins, and in which the ratio of vitamin C to vitamin E by weight is between 10:1 and 15:1, are largely protected against the phytotoxic effect of the herbicide. The cell damage is higher in experiments on plants with a much lower or much higher ratio of vitamin C to vitamin E.The cell damage is also greater when the total level of the two antioxidants is lower. Thus vitamins C and E are a very effective protection against peroxidative cell damage in the plant. The vitamin C level in the plant is generally higher than the vitamin E level. Vitamin C probably forms a reservoir of antioxidative potential for the purpose of regenerating vitamin E during peroxidative stress. Plants are badly damaged by peroxidation if the ratio of the two vitamins is 1:1 or less. This is the case, for example, in old beech leaves, and it is associated with an increase in peroxidative processes [Kunert & Ederer (1985); Finckh & Kunert (1985)]. Experiments on beech leaves and spruce needles showed that the vitamin E level increased with age, while the level of vitamin C did not increase. The ratio of the concentration of vitamin C to vitamin E thus decreases in the course of aging. The decrease in the ratio is accompanied by an increase in lipid peroxidation, as predicted by the rule; this increase was confirmed by measurement of the malonic dialdehyde [Kunert & Ederer (1985)].

4.9.5 On the Synergistic Interaction of Vitamins E and C in the Mammal In 1968 for the first time it was suggested that ascorbic acid is capable of reducing tocopheroxyl radicals which are formed in vivo by the destruction of free radicals during metabolic processes. A single tocopherol molecule is able to trap many radicals. A direct reaction between vitamin E radicals and ascorbate was demonstrated in vitro. It is still unclear what the mechanism of the interaction between vitamins E and C may be in vivo, since the first is probably localized in the membrane, and the second only in the aqueous phase. The reaction is apparently made possible by the localization of the phenolic group of vitamin E in the region of the phase boundary. It is thought that tocopherol is oxidized to the tocopheroxyl radical in the reaction with the peroxyl radical, ROO, and that the tocopheroxyl radical is then reduced to vitamin E by the subsequent reaction with vitamin C [McCay (1985)]. Several experiments in vitro have confirmed that vitamin E traps peroxyl radicals and that the resulting tocopheroxyl radical reacts with vitamin C to form vitamin E.Thus vitamins E and C act synergistically in the inhibition of lipid peroxidation [Niki et al. (1984); Burton et al. (1983 a); Summerfield & Tappel (1984)]. 4.9.6 On the Synergism between Selenium and Vitamin E Synergistic relations between vitamin E and selenium have long been known, liiere are a number of vitamin E deficiency symptoms which can be cured by doses of selenium, including atrophied testes in the hamster, dog and rooster; liver necrosis in the rat and pig; decrease in plasma proteins in chicken and turkey; exudative diathesis in the chicken; and kidney degeneration in the rat, monkey and mink [Bässler & Lang (1981)].

4.9 Biochemical Role

239

We now know that selenium is a component of the enzyme glutathione peroxidase, which takes part in the protection of membrane lipids against peroxidation. Selenium is thus not a synergist of vitamin E as such. 4.9.7 Vitamin E as a Specific Protection Factor in the Biological Membrane The peroxidation of lipids by free radicals is a permanent biological process, which when uncontrolled leads to damage of cellular and intracellular membranes, including the enzymes and other macromolecules bound to them. Vitamin E, an effective lipid antioxidant, prevents this destruction by reacting with peroxyl radicals, ROO, and interrupting the chain reaction [Buttriss & Diplock (1984); Diplock (1984)]. The side chain of vitamin E is involved in this function; it probably interacts with the acyl chains of the multiply unsaturated phospholipids, especially with the arachidonyl residue. This interaction might serve mainly to anchor the vitamin correctly in the membrane [Buttriss & Diplock (1984); Leb et al. (1985); Niki et al. (1985)]. Some authors suspect a specific effect of α-tocopherol on the architecture of the biomembrane other than its antioxidant function. This might occur via complex formation with arachidonyl residues or protection against phospholipases, which would lead to stabilization of the biomembrane [Erin et al. (1983, 1984, 1985)]. The small amount of vitamin E in the membrane weighs against a stabilizing role; in the erythrocyte membrane, the molar ratio of vitamin E/arachidonic acid is about 1:500 to 1:1000 [Burton et al. (1983 a); Diplock (1983 b)]. The chemical nature of the side chain contributes significantly to the reactivity of the vitamin E in vitro. It probably effects the correct association of the vitamin E with the membrane. The exchange of the side chain for a methyl residue in the vitamin E molecule leads to inactivity in vivo, but retains the full activity in vitro [McCay (1985)]. Studies on microsomes from various tissues of the rat, mouse and rabbit revealed a correlation between vitamin E content and resistance to lipid peroxidation in vitro. Analogous observations showed that vitamin E deficiency led to an increase in peroxidation in microsomes and mitochondria in the rat liver [Borrello et al. (1985)]. These observations are taken as direct evidence that vitamin E has a membrane-protecting role. 4.9.8 On the Role of Vitamin E in the Erythrocytes and Blood Platelets Because the erythrocytes are involved in biological gas exchange, they are relatively vulnerable to oxidative stress from oxidizing components of the air. Normally the erythrocytes are able to cope with most interfering effects. If they are not able to cope, their membranes are damaged. They have a number of enzymatic and non-enzymatic systems to protect them against oxidative damage. Vitamin E is a major antioxidative system in erythrocytes; vitamin-E-deficient erythrocytes are more sensitive to hemolysis than normal ones. On the other hand, no structural differences were found in the membrane proteins of normal and vitamin-E-deficient erythrocytes. However, biochemical experiments showed that a decreased intake of vitamin E increased the tendency of erythrocyte mem-

240

4 Vitamin E

branes to form disulfide bridges between proteins by an oxidative process [Chow (1985)]. Eiythrocytes from adults and children with reduced vitamin E intake and with low vitamin E serum levels are relatively sensitive to oxidants in vitro, and the lifetime of these erythrocytes is shortened. In premature infants the lifetime of erythrocytes is shorter than in normal infants. This is at least partly due to the vitamin E deficiency in premature infants [Chow (1985)]. Hemoglobin catalyses lipid peroxidation by converting lipid hydroperoxides to free radicals. Autooxidation of oxyhemoglobin to methemoglobin simultaneously forms the superoxide anion O2. The latter can react with hydrogen peroxide in the erythrocytes, forming hydroxyl radicals and singlet oxygen, which attack unsaturated membrane lipids [Chow (1985)]. These observations underline the protective function of vitamin E. The blood platelets have a relatively high level of multiply unsaturated fatty acids in their membrane phospholipids, which must be protected from oxidation by vitamin E. Vitamin E also protects the membrane from phospholipases. Finally, vitamin E inhibits platelet aggregation, which is an oxygen-consuming process. Patients with impaired intestinal absorption, e.g. with cystic fibrosis or abetalipoproteinemia, who do not receive increased doses of vitamin E show an intensified platelet aggregation, and sometimes a shortening of platelet lifetime [Chow (1985)]. Many authors have investigated the question whether there are lipid-soluble substances in human blood other than vitamin E which, like the tocopherols, are able to trap radicals and break off chain reactions. Their experiments have shown that it is most probably not the case, and that there are no significant amounts of another chain-interrupting antioxidant in human plasma and erythrocytes [Burton & Ingold (1983); Burton et al. (1983 a,b); Burton, Joyce & Ingold (1982); Chow (1985); McCay (1985)].

4.9.9 Vitamin E and Protein Synthesis Changes in the activities of a number of enzymes have been observed in the tissues of vitamin-E-deficient animals, and it has been postulated on this basis that vitamin E might have a specific regulatory role in protein synthesis. Of the many enzymes whose activities change during vitamin E deficiency, muscle creatine kinase and liver xanthine oxidase display the greatest changes. A vitamin E deficiency markedly increases the turnover rate of the creatine kinase and the rate of de-novo synthesis of xanthine oxidase in animal experiments [Chow (1985)]. It is considered a proven fact thàt in the case of xanthine oxidase and creatine kinase, the elevated enzyme levels in animals with vitamin E deficiencies are due to increased synthesis of the enzymes. This observation led to the suggestion that vitamin E acts as a repressor of the synthesis of some enzymes [Machlin (1984)]. On the other hand, an elevated plasma level of many enzymes is a sure index of tissue loss, because the enzymes in question diffuse out of the damaged cells into the plasma. The plasma concentrations of many enzymes have been found to be altered in animals with vitamin deficiencies. Rats with vitamin E deficiency were found to have elevated plasma activities of pyru-

4.9 Biochemical Role

241

vate kinase, and this elevation is used as a sensitive index of myopathy in vitamin E deficiency [Chow (1985)].

4.9.10 On the Anticarcinogenic Effect of Vitamin E Some experimental treatments of cancer patients with vitamin E have been described. The first efforts were made with patients with cervical tumors, but without significant success. 2-amòo-a-tocopheryl acetate had an inhibitory effect on a murine neuroblastoma. The application of the acid succinate of vitamin E caused irreversible growth inhibition in some melanoma cells in culture. In addition, vitamin E succinate had some inhibitory effect on a human neuroblastoma transplanted into mice [Helson et al. (1983)]. Vitamin E inhibited the growth of some experimental tumors in hamsters and mice. Clinical experiments on human epithelial tumors of the ovary, however, showed no significant effect of vitamin E on the development of the disease [Heinonen et al. (1985); Helson et al. (1983)]. The treatment of breast cancer with vitamin E is in the discussion stage [London et al. (1985 a)]. A dietary deficiency of selenium increases the risk of cancer; this effect is enhanced by inadequate intake of vitamin E [Salonen et al. (1985 b)]. Vitamin E and selenium inhibit mutagenesis in vitro [Chow & Gairola (1984)]. Lipid extracts from rat hepatomas had distinctly more vitamin E than lipid extracts from normal rat liver, which might be related to a relatively low consumption of vitamin E in the tumor tissue. Microsomal tumor membranes appear generally to have a limited lipid peroxidation; however, they also have a relatively low level of multiply unsaturated fatty acids [Burton et al. (1983 a); Borrello et al. (1985)]. These observations show that vitamin E has a certain anti-cancer effect in animal experiments, but not in human beings. There appear to be close connections between vitamin E deficiency, formation of malonic dialdehyde as product of lipid peroxidation and DNA damage, which can lead to tumor formation. Malonic dialdehyde, a highly reactive product of lipid peroxidation, reacts with the amino groups of proteins, and with RNA and DNA. Reaction with malonyldialdehyde probably leads to cross-linking of the DNA. The compound is mutagenic in bacteria, Drosophila and human fibroblasts in culture. Correspondingly, animals whose diets promote peroxide formation have a higher frequency of tumors, and the template activity of their DNA is reduced. The frequency of tumors is increased in rats whose feed contains malonic dialdehyde. The pathogenic effect of lipid peroxidation, which leads to massive formation of malonic dialdehyde, is potentiated by vitamin E deficiency, irradiation and many drugs. As indicated earlier, it is linked to DNA damage. The formation of malonic dialdehyde and DNA damage are often linked to ageing in animals [Summerfield & Tappel (1984)]. An opinion on the role of vitamins E and C in the etiology of human cancer has been published [Brightsee (1984)]. 4.9.11 Vitamin E and the Neuromuscular System The frequency of neurological and muscular disorders in vitamin E deficiency conditions of varying etiology, and the (usually attainable) reversibility of these

242

4 Vitamin E

ε o. ε

2

3 S

ε 2 α ε 60 ο 2 3υ < Ζ

ω ε « •>

υ

Ö Ν « S ™ Ο

Βυ < D οû •ο

ω ce

S

υ "3 «3 S

3

α> L_ "2t-H Λ J2 S Ä Ο s 2 g ^ £ -o e -δ 1) — ^ ω ILë h >„ > g ω s Ή ' Μ ε ° α , aï ο ° & ce β 5 Ήu e χ .. 3 ω S s « u es Κ α ^ Ε C -D. S «υ > ρ ? sM< - . 2 Β £ ; g < 3 Ο Ό < «> S S GD ^ TO

cd ε 9 g> "53 o ιΟ. O α

β 2 υ υ

ce — X) c e 12 J= o cυ ÈΒ 2α> Ο » Ν 43 § &

tQ C ε ce

o •s ca .Βa Γ-3Ö S *

>> υ β β Ä fl> U -o

Ό

2 υ .S

Η

υ 'δ

υ> ta-ri co S CK

4.9

Biochemical

R o l e

243

s « OO

4>

2 C-

'S «

5 . 3

S 3

Ή G ο

ω

e U eo υ

•S 2 3 - g m 3 υ o g

υ ί§

'S

£

ε -a « (O ~ ω

S

ε

-

ce

.2 t:

Q

3

.S

u

»

s

ε - s 3 J3 a O oo s oo o (A ce >> f C Ό υ ce t» Χι Λ C Jw3 ce 1 g ce {Λ a. 3 3 o Xi ε

>>

o)

S

.g

C o

£O

•3 ed w 'S O •tS

Λ -ι ÍN τ-·

5

· ·

«

£

w

OO

_* ce

Ά

ce ~υ

ceε

&!· S? 5Λ «2

J

τ-

* à

g

ο ce

ε 60

g

00

s

S

σ.

R C?

ο M

!

0 «

( o §

_ ce

' 1 C O

-S o) 'S

3 o C

4g

ε

s

Ν υ

« Ζ

ce >

§

ο

00

·;

2 W

.s

fi? ΟΟ

ΓC τ-ι Ο ' CA " Ü "Μ Ζ « ΓΛ

00 C

Λ

'•§ Sí °ο °. κ 42 Τ-Η a w

υ

a 1

E β -t—1 > .H β o h rt υ

¿ δc .a o c u ·ο

£ 2 o

s

§ ω

«

Ίί

·β g

á » υ c

e § ·ς >

ε .2 C JO

Ο

CA

I ce u S

o

0\

^

σ\

'S « «

Χ t 2 ? ON

-d

C

3 υ 'C

fe S 3

ts >

^ ο£ ο

s

Ο α

ω«

73 !

fN

? "

. •

J

1 ? ® oo ¡ >

0\

e? o ts ^ ¡ce o e λ .S c .3 λ

c 00 S 3

« S bû

•S c >"

3

o S

.3

? ·· •(j· ^ oo

.

C ·

0 0

s s

i u

.. —; ^ ce c i T „ oo ^ «t. o . U w ce υ . S 'S « 3 —H O 2 « τ™ ce « λ • • H . u 2 m M

SMa S

Q""

ω

«

ce

" S S β J f ^

c

x

· 0 0

»

C

'S

244

4 Vitamin E

Table 4-3. Neurological symptoms in patients with untreated abetalipoproteinemia and other conditions of severe chronic malabsorption [Muller et al. (1983b)] Symptom

Abetalipoproteinemia

Other disorders of fat absorption

Areflexia Cerebellar ataxia Impaired sense of position Retinal disease Impaired perception of vibrations Ophthalmoplegia Muscle weakness Nystagmus Loss of sense of touch and pain

+ + + + +

+ + + + + + + + +

++ ++ ++ ++ +

+ + + + + +

+ + + + +

++ ++

+ + + , Very common (>75%); + + , common (50-75%); + , less common (20-30%) (+), rare (- chorismic acid — 15%). Most of those affected were children, youth, pregnant women and the old [Becker & Bitsch (1979), Lonsdale & Shamberger (1980), Reinken et al. (1979)]. The transketolase activity is measured in hemolysed erythrocytes or hemolysed whole blood by measurement of sedoheptulose 7-phosphate formation [Boni et al. (1980), Davis & Icke (1983)].

6.16 Thiamin Deficiency in Human Beings and Animals

385

The diagnostic value of the transketolase activity and of the TDP effect were tested on a large number of healthy controls and patients. The transketolase activities were lowest in diabetics, and were probably due to a lack of apoenzyme. Patients with polyneuritis also had low activities. Anemic patients scored similarly to the controls, but pernicious anemia patients had decidedly high values, probably due to a high level of apoenzyme [Kjosen & Seim (1977), Säuberlich (1984)]. The following observations are also significant: The transketolase activity may be elevated in severe illness [Leevy (1980)]. Animals after long-term lack of thiamin have subnormal apotransketolase levels [Reinken et al. (1979), Nutrition Rev. 35 (1977) 185]. The transketolase activity is often reduced in alcoholics, but addition of TDP does not increase it, so that here the T D P effect does not correlate with the thiamin status, because it is too low [Davis & Icke (1983), Somogyi et al. (1980)]. Measurements of the transketolase and the T D P effect in erythrocytes of 1028 healthy German children showed a distinct dependence on age: the transketolase activity drops temporarily in the second year of life, and then increases continuously until the 6th year; after another four years, it declines steadily [Reinken et al. (1979)]. The stimulation of the erythrocyte transketolase by addition of T D P depends on pH [Tate et al. (1984)].

To determine the stability of the transketolase, whole blood and washed erythrocytes from normal persons and from aged patients (who had abnormal diets) were stored 14 days at 4°C and at - 2 0 ° C . Both the ETK activity and the TDP effect decreased with time at both temperatures. Interestingly, at — 20 °C, the TDP effect in the samples from the aged patients declined significantly within 4 days, while this did not occur with the samples from normal persons until after 14 days [Puxty et al. (1985)]. Measurements of the ETK activity in 24 children who had died of SIDS revealed about the same values as were found in measurements on children who had died of other causes. This result seems to exclude thiamin deficiency as a cause of death in the SIDS [Peterson et al. (1981)]; however, cf. p. 370. 8 variants of the transketolase in human erythrocytes have been discovered. These differ distinctly in their isoelectric points. There are at least two classes of transketolase from human fibroblasts, which differ in their Km values for TDP. These differences are probably of genetic origin. They are important in connection with the determination of thiamin status by the ETK method, because the method is based on the assumption that the affinity of erythrocyte transketolase for TDP is constant. However, this assumption is apparently negated by the above results [Kaczmarek & Nixon (1983)]. 6.16.3.2 Measurement of the Thiamin Level in the Blood There has been much discussion of the possibility of determining thiamin status by determining the thiamin level in whole blood or its components. It is assumed that there is a correlation between the blood and tissue levels of thiamin, although this is not known for certain with human beings. The blood contains only about 0.8% of the total thiamin in the body, and the thiamin level in blood is thus very low (6-12 μg/100 ml). Because most of the thiamin in blood is in the cells, the serum contains much less. The methods used to determine it must therefore be very sensitive. O. danica has proven suitable for measurement of thiamin in whole blood and its components [Gubler (1984)]. In the past few years, very effective chromatographic methods have been described, most of which make use of HPLC and fluorimetry. They are able to determine thiamin and its esters in

386

6 Thiamin, Vitamin B l t Aneurin

the blood components very precisely [Warnock (1982), Kimura et al. (1982 a), Kimura & Itokawa (1983), Bontemps et al. (1984), Floridi et al. (1984), Sauberlich (1984), Weber & Kewitz (1985), Kimura & Itokawa (1985)]. 6.16.3.3 Measurement of Thiamin Excretion in the Urine Thiamin excretion in urine was formerly the most common method of determining thiamin status, using either the thiochrome method or a microbiological test. A good correlation was found between the degree of thiamin deficiency and the decrease in thiamin excretion, or between the daily thiamin intake and thiamin excretion. It is useful, but not very practical, to measure the 24 hour excretion. Instead, the 4 hour excretion is usually taken, and the thiamin is related to the simultaneously excreted creatinine [Gubler (1984)]. The ratio of urinary thiamin to creatinine is normally proportional to the consumption of thiamin [Gregory & Kirk (1978)]. The normal thiamin excretion by adults is more than 66 μg/g creatinine, and values less than 27 μg/g creatinine indicate thiamin deficiency [Herman et al. (1976)]. HPLC is often used to measure thiamin in the urine. Thiamin coming off the column is converted to thiochrome and measured fluorimetrically [Säuberlich (1984)]. Riboflavin can be measured at the same time [Mansourian et al. (1982)]. 6.16.4 Therapy of the Thiamin Deficiency Diseases In mild adult beriberi, one administers 10 mg thiamin 3 χ daily, orally. Severe cases are treated with 25 mg thiamin 2 χ daily, intravenously. Children with mild beriberi should be given 5 mg thiamin/day orally; in severe cases, 10 mg thiamin 2 χ daily, intravenously. In the case of ambulant patients who have recovered from beriberi heart disease, it is recommended that the treatment be continued until they are completely healed, to avoid sudden death. Fulminant beriberi heart disease requires, in addition to the other therapy, 100 mg thiamin hydrochloride (intravenously); in this case, thiamin alone is not sufficient. Patients with Wernicke encephalopathy should receive intravenous doses of 25 to 50 mg thiamin/ day [Herman et al. (1976)]. High doses of thiamin led to normalization of the symptoms in a 24-year-old, overweight woman with Wernicke encephalopathy which had been caused by gastric folding [MacLean (1982)]. Very high doses of thiamin (300 mg/day for several weeks) were therapeutically effective for patients with Kearns-Sayre syndrome (progressive external ophthalmoplegia, or paralysis of the eye muscles). This disease is associated with impaired metabolism of pyruvate and lactate [Lou (1981)]. In the treatment of alcoholics, thiamin is first applied parenterally. Allithiamins (p. 389) may be applied orally [Leevy (1982)]. The administration of 50 mg thiamin/day produced normalization of the TDP effect within 10 days, but the transketolase activity and the thiamin level rose insignificantly [Waldenlind et al. (1981)]. Liver diseases, even those which last for a relatively short time (4-20 days) can lead to thiamin deficiency. Patients with liver damage should therefore be treated with high doses of thiamin, e.g. 100 mg thiamin hydrochloride 2 χ daily [Laba-

6.17 Congenital Impairments of Thiamin-Dependent Metabolism

387

darios et al. (1977)] or 200 mg thiamin hydrochloride/day, for one week [Roosouw et al. (1978)]. Experiments on calves have shown that thiamin (100 mg/day, subcutaneously) will largely protect them from poisoning with lead acetate (5 mg/day, for 20 days). Administration of the lead acetate alone led to the death of almost half the animals. It is assumed that thiamin will generally protect against lead, which is present in considerable concentration in the air, in humans as well as animals [Bratton et al. (1981)].

6.17 Congenital Impairments of Thiamin-Dependent Metabolism 6.17.1 Maple Syrup Urine Disease, Branched-Chain Disease The cause of maple syrup urine disease9 is a congenital10 lack of the enzyme branched-chain 2-oxoacid dehydrogenase (EC 1.2.4.4). The disease is autosomal recessive. The metabolism of leucine, isoleucine and valine are affected. These amino acids and the corresponding 2-oxoacids (which cannot be degraded by oxidative decarboxylation) are found in very high concentrations in the serum and urine. The urine of the patients characteristically smells like maple syrup. The disease affects 1 of 250,000 neonates. There are several variants of the disease, which differ in their prognoses. In the severe form, there is essentially no activity of the branched-chain 2-oxoacid decarboxylase in the leukocytes and fibroblasts; the mild form is almost free of the typical symptoms, although the usual biochemical anomalies can be detected. The classic form of the disease does not respond to administration of thiamin, but there are rare thiamin-responsive variants in which very high doses of thiamin lead to improvement. It is thought that in these cases, the thiamin stabilizes the enzyme. The severe form of the disease is coupled with mental retardation, lethargia and vomiting, and is fatal early in life. If branched-chain amino acids in the diet are restricted early enough, the classic manifestations can be somewhat alleviated [Auerbach & DiGeorge (1973), Herman et al. (1976), Elsas & Danner (1982), Davis & Icke (1983), Bartlett (1983), Heffelfinger et al. (1984)]. A one-year-old child with maple syrup urine disease was treated with high oral doses of thiamin (100 mg/day, later 300 mg/day). After about three weeks a partial improvement was achieved, in that the frequency of acute crises became less [Gray et al. (1982)]. The examination of another patient with thiamin-responsive maple syrup urine disease revealed a reduced affinity of the branched-chain 2-oxoacid decarboxylase for TDP and for 2-oxoisovalerate [Chuang et al. (1982)]. A few other children with maple syrup urine disease were recently discovered [Wendel et al. (1983)].

9 10

Reviews : Naughten et al. (1985) General review: Duran & Wadman (1985)

388

6 Thiamin, Vitamin Β,, Aneurin

6.17.2 Congenital Lactate Acidosis This is a group of several congenital defects which are collectively called lactate acidosis. The disease has been observed primarily in children. The clinical symptoms are lactic acidosis, pyruvic acidosis, neurological impairment and delayed growth. The most important causes are probably defects in the pyruvate dehydrogenase complex, or in the substituent enzymes of this complex. Patients with a lack of pyruvate decarboxylase (one of the substituent enzymes) have been treated with high doses of thiamin [Bartlett (1983), Koike & Koike (1982)]. 6.17.3 Leigh Syndrome The Leigh syndrome, also called subacute necrotizing encephalomyelopathy, was first described in 1951. By 1976,100 cases of the disease were known. It is a rapidly fatal disease of infancy and early childhood. It is associated with weakness, anorexia (lack of appetite), irregular eye motions, difficulties with speech and cessation of development. The affected children appear to develop normally at first, but the clinical symptoms then appear very rapidly. Autopsies revealed necroses in the brain stem and spinal cord. The biochemical changes included elevated levels of pyruvate, 2-oxoglutarate and lactate in the blood, which suggests impaired oxidative decarboxylation of the 2-oxoacids. The disease is familial, and is thought to have autosomal recessive inheritance. The neuropathological and clinical symptoms in patients with Leigh syndrome are similar to those in patients with Wernicke encephalopathy. Post mortem, patients with Leigh syndrome were found to have a reduced level of thiamin triphosphate (TTP) in the brainstem. Some authors found an inhibitor of the phosphorylation of TDP to TTP in the urine, blood and CSF of the patients. The successful treatment of patients with Leigh syndrome with very high oral doses (up to 2.0 g/day) of thiamin has been reported. The most useful forms were thiamin propyldisulfide and thiamin tetrahydrofurfuryldisulfide, which enter the CSF more readily than thiamin [Davis & Icke (1983), Herman et al. (1976), Cooper & Pincus (1973)]. 6.17.4 Thiamin-Responsive Megaloblastic Anemia In 1969, an eleven-year-old child with megaloblastic anemia, deafness and diabetes mellitus was first observed. The anemia was resistant to vitamin B12, folic acid and vitamin B6, but it did respond to oral doses of 20 mg thiamin/day 11 . Another case of this disease appeared in 1978. Two more cases were then found, a pair of twins. The inheritance is thought to be autosomal recessive [Haworth et al. (1982)]. Examination of two patients with thiamin-responsive megaloblastic anemia indicated that the thiamin level in the blood, the erythrocyte transketolase and the pyruvate dehydrogenase and 2-oxoglutarate dehydrogenase complexes of the leukocytes were normal. Administration of 50 mg thiamin/day led to a rapid increase in the reticulocytes and hemoglobin [Bartlett (1983)]. It is thought that the 11

Review: Nutrition Rev.38 (1980) 374

6.18 Modified Forms of Thiamin

389

cause of the disease is impaired thiamin transport through the biological membranes of some organs and tissues, including the enterocytes and the red blood cells [Poggi et al. (1984)]. In all, at least five cases of thiamin-responsive anemia have so far been reported. All the patients developed megaloblastic anemia and diabetes and lost their hearing, and all of them responded to high doses of thiamin, although the thiamin status appeared to be normal from the beginning [Mandel et al. (1984)].

6.18 Modified Forms of Thiamin 6.18.1 Thiamin Analogs with Full or Improved Vitamin Activity Many lipophilic derivatives of thiamin are known which have improved vitamin activity. Some of them have been adopted in therapy and the food industry, due to their ease of absorption. In most of these derivatives, the thiazole ring has been opened, and the sulfur carries a lipophilic substituent, e.g. S-alkyl or S-tetrahydrofurfuryl in the allithiamins (Fig. 6-13). The latter two are probably the most important derivatives ; the individual representatives differ in the type of S-substituent. The best known are thiamin allyldisulfide (Fig.6-13a), thiamin propyldisulfide (Fig.6-13b) and thiamin tetrahydrofurfuryldisulfide (Fig. 6-13 c).

(a) (b)

R=CH2CH = CH;'2 R = CH2CH2CH3

(c) Fig. 6-13. Some of the more important allithiamins [Oka (1984)]. a. Thiamin allyldisulfide; b. Thiamin propyldisulfide; c. Thiamin tetrahydrofurfuryldisulfide

Allithiamins were discovered by M.Fujiwara and H.Watanabe (1952). These authors observed that when extracts of some plants, especially garlic (Allium sativum) were heated with thiamin to 60 °C, the thiamin was modified and no longer gave the typical thiochrome reaction. However, it had the full vitamin activity in tests with animals. It was found that thiamin had been converted to an alkyldisulfide. Such a modification also occurs in the presence of extracts of onion (Allium cepa) and many other plants. Finally, the allithiamins and their modifications were produced synthetically; this group also includes S-acyl derivatives and 0,Sdibenzoylthiamin. All of these compounds are less water-soluble than thiamin. They are rapidly absorbed from the intestine, and appear to bypass the control

390

6 Thiamin, Vitamin B^ Aneurin

mechanisms for absorption. Free thiamin appears in the urine after oral administration. There are indications that thiamin propyldisulfide (Fig. 6-13 b) is better retained in the body and leads to a higher thiamin level in the erythrocytes than the conventional form of the vitamin. Thiamin propyldisulfide enters the erythrocytes after intestinal absorption and is thereafter reduced to free thiamin, in the presence of glutathione. The free thiamin is then slowly released into the blood plasma. It has also been shown that thiamin propyldisulfide readily enters the CSF, the nervous tissue and the eye. S-Acylthiamin is just as readily absorbed from the intestine, but it does not enter the erythrocytes as easily as thiamin propyldisulfide. Probably only thiamin propyldisulfide and thiamin tetrahydrofurfuryldisulfide will be clinically applied [Davis & Icke (1983)]. The formation of allithiamins from thiamin in the presence of plant extracts is due to the reactions of various allicines, which have the structure: R - S=0

I R'-S

where R and R' may be methyl, ethyl, propyl or allyl groups. An enzymatically catalysed transfer of the R'-S- group to the sulfur of thiamin produces allithiamin [Fujiwara (1976)]. Thiamin propyldisulfide (Fig. 6-13 b) is used, among other things, to test the function of the liver. Healthy liver degrades this compound mainly to 2-hydroxypropylmethylsulfone (2-HPMS), which has the structure CH3SO2CH2CHCH3 I OH

2-HPMS is excreted in the urine and measured. The excretion of 2-HPMS decreases in proportion to the severity of the liver damage [Oda et al. (1984)]. Synthetic O, S-dibenzoylthiamin is recommended as a food additive, because of its unusually low water solubility and its stability to heat [Heywood et al. (1985)]. Table 6-6. Some structural modifications of the thiamin molecule and their effects on biological activity of the analogs Structural modification

Biological effect

H or ethyl instead of methyl on C-4

Partial thiamin activity in doves

Elimination or methylation of the 3,5'-CH2 group

Loss of biological activity

Replacement of the 3,5'-CH2 by an ethylene group

60% as much catalytic activity as thiamin

Replacement of N-l' by -CH =

Loss of biological activity

Replacement of CH3-group on C-2' by ethyl or propyl

Bioactivity still present

Replacement of the CH3- on C-2' by H

Loss of bioactivity

Methylation of the NH2- group

Loss of bioactivity

6.18 Modified Forms of Thiamin

391

6.18.2 Thiamin Analogs with Weak or No Vitamin Effects A description of the thiamin analogs will not be given here; only some deviations from the thiamin structure and their effects on the bioactivity are summarized in Table 6-612. 6.18.3 Thiamin Analogs with Antagonistic Effects The biochemical basis of the changes accompanying thiamin deficiency in human beings and animals has been intensively studied. The experiments in vivo and in vitro were made easier or even possible by the use of thiamin antagonists, especially oxythiamin (OH instead of NH 2 in the thiamin molecule) and of pyrithiamin (CH = CH instead of S in the thiamin molecule, see Fig. 6-14 a).

CH 3

Fig. 6-14. Structures of the thiamin antagonists pyrithiamin (a) and amprolium (b) [Oka (1984)]

Only the latter affects the central nervous system in vivo; it acts not only as an antagonist of thiamin, but probably also directly on the conductivity and secretions of the neurons [Hirsch & Gibson (1984)]. Both oxythiamin and pyrithiamin cause weight loss and bradycardia in animal experiments, but only pyrithiamin causes polyneuritis. It has been found that pyrithiamin administered parenterally or orally can pass the blood-brain border unhindered and is accumulated in the brain. Oxythiamin cannot pass the blood-brain barrier. Experiments on rabbit vagus in vitro have also indicated that only pyrithiamin, but not oxythiamin, can enter nerve cells from an incubation solution [Cooper & Pincus (1979)]. Finally, amprolium (Fig. 6-14 b) is of considerable interest. This is a prototype of a class of thiamin antagonists which are effective against coccidiosis. They inhibit intestinal thiamin transport by blocking a transport protein which carries thiamin into the cell [Rogers (1982)]. Amprolium also (like pyrithiamin) blocks thiamin transport across the blood-brain barrier in the rat [Greenwood & Pratt (1981)]. Some properties of the three thiamin antagonists discussed above are summarized in Table 6-7. Other thiamin antagonists are treated by Oka and Gubler [Oka (1984), Gubler (1984)]. 12

Review: Rogers (1962), Oka (1984), Gubler(1984)]

392

6 Thiamin, Vitamin B1( Aneurin

Table 6-7. Some properties of pyrithiamin, oxythiamin and amprolium Thiamin Antagonist

Anti-thiamin effect

Pyrithiamin (Fig. 6-14a)

Substrate of thiamin pyrophosphokinase (1). Easily passes the blood-brain barrier; causes neurological symptoms (4). Stored in the brain, mostly as phosphates (1). Blocks thiamin transport into the brain (1). Strongly inhibits pyrophosphorylation of thiamin (2, 3, 4) and thus leads to rapid urinary excretion of thiamin (2). Presumably acts as a competitor of thiamin at the cell membranes in the nervous system (2).

Oxythiamin

Probably inhibits all TDP-dependent reactions (5). Forms oxythiamin diphosphate (OTDP) in the presence of thiamin pyrophosphokinase; OTDP binds to pyruvate decarboxylase and blocks binding of TDP (6). Cannot cross the blood-brain barrier (7).

Amprolium (Fig. 6-14b)

Agent against coccidiosis (8, 9). Inhibits thiamin absorption in the chicken (10). Is not phosphorylated (10).

References: (1) Iwashima et al. (1976); (2) Waldenlind (1978); (3) Yoshioka (1984); (4) Dreyfus (1982); (5) Waldenlind (1979a); (6) Shin et al. (1979); (7) Cooper & Pincus (1979); (8) Rogers (1982); (9) Oka (1984); (10) Lumeng et al. (1979).

6.19 Enzymes and Low-Molecular-Weight Plant Components Which Destroy Thiamin Many natural substrates contain components which destroy thiamin. These include enzymes and also low-molecular-weight substances. The enzymes, called thiaminases, have been thoroughly investigated, but the low-molecular-weight substances are still objects of controversy. There is a thiamin-destroying activity in tea; continuous administration to rats leads to thiamin deficiency and inhibition of acetylcholine synthesis [Ruenwongsa & Pattanavibag (1984)]. In some areas of Thailand, where much tea is consumed, thiamin deficiency is common in the population [Hayakawa et al. (1984)]. 6.19.1 Thiaminases Two thiamin-degrading enzymes are known. Thiaminase I (EC 2.5.1.2) has been found in shellfish, fresh water fish, some sea fish and plants, especially, ferns. Some microorganisms also synthesize thiaminase I.Thiaminase II (EC 3.5.99.2) has been found chiefly in bacteria. In the living cell the thiaminases are normally inactive, but upon homogenization of the cell in water at pH 4-8 they are activated. They may also be excreted into the medium by microorganisms. Thiaminase I catalyses the cleavage of thiamin by an exchange reaction with a nitrogen base or an SH compound. This reaction is a displacement of the methylene group of the

6.19 Enzymes and Low-Molecular-Weight Plant Components

(α)

X = SO3H

(b)

0/ X = -Ni

(c)

X = OH

393

0>

Fig. 6-15. Thiamin cleavage by a. Sulfite b. Thiaminase I c. Thiaminase II [Oka (1984)]. The reaction is an exchange of the thiazole group for the group X

pyrimidine part of the thiamin, which eliminates the thiazole part. Pyridine, quinoline, nicotinic acid, aniline, cysteine, etc. may serve as the displacement base or SH compound. The above reactions follow the same mechanism as the one with sulfite. Pure thiaminase I is only active in the presence of a nitrogen base or an SH compound. Thiaminase II catalyses the simple hydrolysis of thiamin to pyrimidine and thiazole portion (Fig. 6-15) [Murata (1982), Edwin (1979b)]. Not only is thiamin destroyed by the reactions catalysed by thiaminase I, but thiamin analogs may be produced which, e.g. after intestinal absorption, may block thiamin-dependent metabolic reactions [Brent & Bartley (1984)]. Bacterial thiaminase I is an exoenzyme which is bound to the cell surface. It has been obtained from various bacteria, including B. thiaminolyticus, and from the rumen contents and feces of animals with polioencephalomalacia (PEM). The purified bacterial enzyme has a fnolar mass of 67,000 daltons, but the enzyme from PEM-afïlicted animals is much heavier (158,000-540,000 daltons). The thiaminases I from Clostridium sporogenes and B. thiaminolyticus are also not identical. The preferred cosubstrate for thiaminase I from PEM-afïlicted animals is aniline (100%), followed by niacin (23-45%) and pyridoxin (23-41%); histamine and imidazole are nearly as good cosubstrates as niacin. The cause of polioencephalomalacia is believed to be the thiaminase I in the rumen, which cleaves thiamin in the presence of various cosubstrates, and which also forms thiamin analogs which may interfere with thiamin metabolism [Brent & Bartley (1984)].

6.19.2 Low-Molecular-Weight Thermostable Plant Components As early as 1946, non-enzymatic plant substances which inactivate thiamin were observed. What was probably the earliest communication reported that rats whose diets contained 40% fern plants lost weight and displayed symptoms of thiamin deficiency. Thereafter a number of fruits and vegetables were systematically investigated for antithiamin activity, and the highest activity was found in blueberries, red chicory, red currants, red beets, brussels sprouts and red cabbage. The activity was most often found in colored plants. As further experiments showed, the antithiamin activity is bound to phenolic compounds, especially those which contain two adjacent OH groups. Caffeic acid (3,4-dihydroxycinnamic acid) was isolated from ferns and blueberries as one of the antithiamin

394

6 Thiamin, Vitamin B1; Aneurin

factors in these plants [Hilker & Somogyi (1982), Hayakawa et al. (1984)]. The effects of polyphenols on thiamin have been described [Panijpan & Ratanaubolchai (1980), Hayakawa et al. (1984)]. The reaction by which thiamin is inactivated by ortho diphenols, e.g. caffeic acid, has been reinvestigated. Ή-NMR and thin-layer chromatography failed to reveal a change of the thiamin by caffeic acid or other ortho diphenols in aqueous solution at pH 7.8 and 37 °C (the original conditions used). The unmodified thiamin was demonstrated by the thiochrome reaction on the thin-layer chromatograms. This cast doubt on the inactivation of thiamin by ortho diphenols in vitro, which had been previously reported [Horman & Brambilla (1982), Horman et al. (1981)].

6.20 Pharmacology, Toxicology Very high intravenous doses of thiamin are lethal to animals: 125 mg/kg in the mouse, 250 mg/kg in the rat, 300 mg/kg in the rabbit, and 350 mg/kg in the dog. Experiments on dogs indicated that when they were given artificial respiration, they could tolerate much higher doses than under normal conditions; with artificial respiration, a blood level of 36.9 mg/100 ml is not yet lethal in the dog, although 7.2-10.0 mg/100 ml is otherwise lethal. The cause of death is thought to be impairment of the respiratory center [Davis & Icke (1983)]. In one experiment, mice died of suffocation after injection of 8 mg thiamin per mouse [Davis et al. (1982)]. In human beings, parenteral doses of thiamin may have some toxic effects, but only when the recommended intake is exceeded by more than 100-fold (Table 6-5). The symptoms are headaches, convulsions, weakness, paralysis, heart arrythmia and allergic reactions. By contrast, even very high oral doses have been found not to be toxic in human beings, but a maximum of 5 mg/dose is absorbed [Miller & Hayes (1982)]. Some allergic reactions to parenteral administration of thiamin hydrochloride in a 44-year-old man have been reported [Kolz (1980)]. The biosynthesis of vitamin B6 is inhibited in many yeasts by exogenous thiamin (1.5 nmol/ml). Cell growth is not affected, however. (In bacteria, neither growth nor vitamin B6 synthesis is affected by thiamin.) The cellular level of vitamin B6 is elevated, however, in the presence of the thiamin antagonists pyrithiamin or oxythiamin, at concentrations which do not yet affect cell growth [Minami et al. (1982)].

6.21 References Abdel-Hamid, M. E., Barary, M. H., Hassan, E.M. & Elsayed, M. A. (1985) Analyst 110 831. Aksoy, M., Basu, T.K., Brient, J.& Dickerson, J.W.T. (1980) Eur. J. Cancer 16 1041. Ang, C. Y. W. & Moseley, F. A. (1980) J. Agrie. Food Chem. 28 483. Auerbach, V.H. & Di George, A. M. (1973) in: Hommes, F.A. & van den Berg, C.J., eds., Inborn Errors of Metabolism (Academic Press, London) p. 337.

6.21 References

395

Auhagen, E. (1982) Trends Biochem. Sci. 7 225. Bachevalier, J., Joyal, C.& Botez, M. E. (1981) Internat. J. Vitam. Nutr. Res. 51 205. Baker, H.& Frank, 0.(1976) J. Nutr. Sci. Vitaminol. 22 63, Suppl. Barchi, R.L. (1979) Methods Enzymol. 62D 118. Barclay, L.L., Gibson, G.E. & Blass, J.P. (1980) J. Clin. Res. 28 516 Α. Barker, J.N., Jordan, F., Hillman, D.E. & Barlow, 0.(1982) Ann. Ν. Y. Acad. Sci. 378 449. Bartlett, Κ. (1983) Adv. Clin. Chem. 23 141. Bässler, K.H. & Lang, K.(1981) Vitamine 2. Auflage (Steinkopf, Darmstadt). Bayliss, R.M., Brookes, R., McCulloch, J., Kuyl, J.M. & Metz, J.(1984) Internat. J. Vitam. Nutr. Res. 54 161. Becker, D.P. & Bitsch, R.(1979) Biologie in Unserer Zeit 9 187. Beitz, J., Schellenberger, Α., Lasch, J.& Fischer, J.(612) Biochim. Biophys. Acta 612 451. Bergquist, J.E. & Hanson, M. (1983) Exper. Neurol. 79 622. Bisswanger, H.(1980) Biochim. Biophys. Res. Comm. 95 513. Blass, J.P. & Gibson, G.E. (1977) N. Engl. J.Med. 297 1367. Blum, K.U. (1975) Med. Welt 26 119. Boni, L., Kleckens, L.& Hendrikx, A.(1980) J. Nutr. Sei. Vitaminol. 26 507. Bonjour, J.P. (1980) Internat. J. Vitam. Nutr. Res. 50 321. Bontemps, J., Philippe, P., Bettendorff, L., Lombet, J., Dandrifosse, G., Schoffenich, E.& Crommen, J.(1984) J. Chromatogr. 307 283. Brandt, D.R. & Roche, T.E. (1983) Biochemistry 22 2966. Brandt, D.R., Roche, T.E. & Pratt, M.L. (1983) Biochemistry 22 2958. Bratton, G.R., Zmudzki, J., Bell, M.C. & Warnock, L.G. (1981) Toxicol. Appi. Pharmacol. 59 164. Breen, K.J., Buttigieg, R., Iossifidis, S., Lourensz, C.& Wood, Β.(1985) Amer. J.Clin. Nutr. 42 121.

Brent, Β. E. & Bartley, E. E. (1984) J. Anim. Sci. 59 813. Brown, G. M. (1971) Comprehensive Biochemistry 21 1. Brown, G.M. & Williamson, J.M. (1982) Advances Enzymol. 53 345. Butterworth, R.F. (1982) Neurochem. Internat. 4 449. Campbell, C.H. (1984) Lancet 2446. Caster, W.O. & Meadows, J.S. (1980) Internat. J. Vitam. Nutr. Res. 50 125. Chan-Palay, V., Plaitakis, Α., Nicklas, W.& Beri, S. (1977) Brain Res. 138 380. Chen, L.T., Bowen, C.H. & Chen, M.F. (1984) Nutr. Rep. Int. 30 433. Chuang, D.T., Ku, L.S. & Cox, R.P. (1982) Proc. Natl. Acad. Sci. USA 79 3300. Claus, D., Eggers, R., Warecka, K.& Neundörfer, B.(1985) Eur. Arch. Psychiatr. Neurol. Sci. 234 390. Cooper, J.R., Nishino, Κ., Nishino, N.& Piros, K.(1982) Ann. Ν. Υ. Acad. Sci. 378 177. Cooper, J.R. & Pincus, J.H. (1979) Neurochem. Res. 4 223. Cooper, J.R. & Pincus, J.H. (1973) in: Hommes, F. A. & van den Berg, C.J., eds., Inborn Errors of Metabolism (Academic Press, London) p. 119. Corbett, M.D. & Corbett, B.R. (1982) in: Kehl, H., ed., Chemistry and Biology of Hydroxamic Acids (Karger, Basel) p. 160. Crane, S.& Price, J.(1983) J. Nutr. Sci. Vitaminol. 29 381. Cremer, H.D. (1980) in: Cremer, H.D., Hötzel, D.& Kiihnau, J., eds., Ernährungslehre und Diätetik Band /(Thieme, Stuttgart) p. 14. David, S., Estramareix, B., Fischer, J.& Th;eerisod, M.(1981) J. Amer. Chem. Soc. 103 7341. David, S., Estramareix, B., Fischer, J.& Th;eerisod, M.(1982) J. Chem. Soc. Perkin Trans. I 2131. Davis, R.E. & Icke, G.C. (1983) Adv. Clin. Chem. 23 93. Davis, R.E., Icke, G.C. & Hilton, J.M. (1982) Clin. Chim. Acta 123 321. Davis, R.E., Icke, G.C., Thorn, J.& Riley, W.J. (1984) J. Nutr. Sci. Vitaminol. 30 475.

396

6 Thiamin, Vitamin B1; Aneurin

Deutsche Gesellschaft für Ernährung (1981) Empfehlungen fur die Nährstoffzufuhr (Umschau Verlag, Frankfurt/Main). Documenta Geigy (1975) Wissenschaftliche Tabellen (Thieme, Stuttgart). Doerge, D.R., Sakurai, T.& Ingraham, L.L. (1981) Analyt. Biochem. 112 236. Dong, M.H., Green, M.D. & Sauberlich, H.E. (1981) Clin. Biochemistry 14 16. Dreyfus, P.M. (1976) J. Nutr. Sci. Vitaminol. 22 13, Suppl. Dreyfus, P. M. (1982) Ann. N. Y. Acad. Sci. 378 365. Dunbar, W.E. & Stevenson, K.E. (1979) J. Assn. Offic. Anal. Chem. 62 642. Duran, M.& Wadman, S.K. (1985) J. Inher. Metab. Dis. 8 70, Suppl. 1. Dykner, T., Ek, Β., Nyhlin, H.& Wester, P.O. (1985) Acta Med: Scand. 218 129. Echols, R.E., Harris, J.& Miller, R.H. (1980) J. Chromatogr. 193 470. Echols, R.E., Miller, R.H., Winzer, W., Carmen, D.J. & Ireland, Y.R. (1983) J. Chromatogr. 262 257. Eder, L.& Dunant, Y.(1980) J. Neurochem. 35 1278. Eder, L., Dunant, Y.& Loctin, F.(1980) J. Neurochem. 35 1287. Edijala, J. K. (1979) Analyst 104 637. Edwin, E.E. (1979a) Methods Enzymol. 62D 51. Edwin, E.E. (1979b) Methods Enzymol. 62D 113. Eliasson, S.G. & Scarpellini, J.D. (1976) Neurochem. Res. 1191. Elsas, L.J. & Danner, D.J. (1982) Ann. Ν. Y. Acad. Sci. 378 404. Estramareix, B.& Th;eerisod, M.(1984) J. Amer. Chem. Soc. 106 3857. Evered, D.R. & Mallet, C , Life Sä. 32 1355. Floridi, Α., Pupita, M., Palmerini, C.Α., Fini, C.& Fidanza, A.A. (1984) Internat. J. Vitam. Nutr. Res. 54 165. Fujiwara, M.(1976) J. Nutr. Sei. Vitaminol. 22 57, Suppl. Gaitonde, M.K. (1982) Neurochem. Internat. 4 465. Gibson, G., Barclay, L.& Blass, J.(1982) Ann. N. Y. Acad. Sei. 378 382. Gibson, G. E., Ksiezak-Reding, H., Sheu, K.F.R., Mykytyn, V.& Blass, J. P. (1984) Neurochem. Res. 9 803. Gill, G.V., Henry, L.& Reid, H.A. (1980) Brit. J.Nutr. 44 273. Gomez-Moreno, C.& Edmondson, D.E. (1985) Arch. Biochem. Biophys. 239 46. Gounaris, A.D. & Schulman, M.(1980) Analyt. Biochem. 102 145. Gray, R.G.F., Green, A.& Bates, G.R. (1982) J. Med. Genet. 19 65. Greenwood, J.& Pratt, O.E. (1981) J. Physiol. 317 65P. Gregory, J.F. & Kirk, J.R. (1978) J. Agrie. Food Chem. 26 338. Gubler, C.J. (1979) Methods Enzymol. 62D 101. Gubler, C. J. (1984) in : Machlin, L. J., ed., Handbook of Vitamins (Dekker, New York) p. 245. Gubler, C.J., Fleming, G.& Kuby, S.A. (1982) Ann. N. Y. Acad. Sci. 378 459. Gubler, C.J. & Hemming, B.C. (1979) Methods Enzymol. 62D 63. Hakim, A.M. (1984) Ann. Neurol. 16 673. Hartman, G.J., Carlin, J.T., Scheide, J.D. & Ho, C.T. (1984) J. Agrie. Food Chem. 32 1015. Hassan, S. S. M., Iskander, M. L. & Nashed, N. E. (1985) Fresenius' Ζ. Analyt. Chem. 320 584. Haworth, C., Evans, D.I.K., Mitra, J.& Wickramasinghe, S.N. (1982) Brit. J.Haematol. 50 549. Hayakawa, F., Ueda, K.& Murata, Κ. (1984) J. Nutr. Sci. Vitaminol. 30 319, 327. Hayashi, K., Yoshida, S.& Kawasaki, T.(1981) Biochim. Biophys. Acta 641106. Heffelfinger, S.C., Sewell, E.T., Elsas, L.J. & Danner, D.J. (1984) Amer. J. Hum. Genet. 36 802. Henderson, G.B., Kojima, J.M. & Kumar, H.P. (1985) Biochim. Biophys. Acta 813 201. Henderson, G.B. & Zevely, E.M. (1978) J. Bacteriol. 133 1190. Herman, R.H., Stifel, F.B. & Greene, H.L. (1976) in: Dietschy, J.M., ed., Disorders of the Gastrointestinal Tract (Grune & Stratton, New York) 390. Heywood, R„ Wood, J.D. & Majeed, S.K. (1985) Toxicol. Lett. 26 53. Hilker, D.M. & Clifford, A.J. (1982a) Feder. Proc. 41 Abstr.1161.

6.21 References

397

Hilker, D.M. & Clifford, A.J. (1982b) J. Chromatogr. 231 433. Hilker, D.M. & Somogyi, J.C. (1982) Ann. Ν. Y. Acad. Sci. 378 137. Hilvert, D.& Breslow, R.(1984) Bioorg. Chem. 12 206. Hirsch, J. A. & Gibson, G.E. (1984) Biochem. Pharmacol. 33 2325. Hoeller, H , Fecke, M.& Schaller, K.(1977) J. Anim. Sci. 44 158. Hogg, J.L. (1981) Bioorg. Chem. 10 233. Horman, I.& Brambilla, E. (1982) Ann. Ν. Y. Acad. Sci. 378 467. Horman, I., Brambilla, E.& Stalder, R.(1981) Internat. J. Vitam. Nutr. Res. 51 187, 385. Hoyumpa, A.M. (1982) Ann. Ν. Y. Acad. Sci. 378 337. Hoyumpa, A.M., Strickland, R., Sheehan, J. J., Yarborough, G.& Nichols, S.(1982) J. Lab. Clin. Med. 99 701. Hurst, W.J. (1983) Internat. J. Vitam. Nutr. Res. 53 239. Iber, F.L., Blass, J. P., Brin, M.& Leevy, C.M. (1982) Amer. J. Clin. Nutr. 38 1067. Inoue, M., Hirano, H., Sugiyama, K., Ishida, T.& Nakagaki, M.(1982) Biochim. Biophys. Res. Comm. 108 604. Irle, E.& Markowitsch, H.J. (1982) Exper. Brain Res. 48 199. Irle, E.& Markowitsch, H.J. (1983) Behav. Brain Res. 9 277. Ishii, K., Sarai, Κ., Sanemori, H.& Kawasaki, T.(1979a) Analyt. Biochem. 97 191. Ishii, K., Sarai, Κ., Sanemori, H.& Kawasaki, T.(1979b) J. Nutr. Sci. Vitaminol. 25 517. Itokawa, Y (1976) J. Nutr. Sci. Vitaminol. 22 17, Suppl. Itokawa, Y., Kimura, M.& Nishino, K.(1982) Ann. Ν. Y. Acad. Sci. 378 327. Iwashima, A. (1981) Seikagaku 53 444. Iwashima, Α., Nishimura, H.& Nose, Y.(1979) Biochim. Biophys. Acta 557 460. Iwashima, A.& Nose, Y.(1976) J. Bacteriol. 128 855. Iwashima, Α., Wakabayashi, Y.& Nose, Y.(1976) J. Biochem. 79 845. Iwata, H.(1982) Trends Pharmacol. Sci. 3 171. Iwata, H., Matsuda, T., Maeda, S.& Baba, A. (1979) Biochim. Biophys. Acta 583 159. Iwata, H., Yabushita, Y., Doi, T.& Matsuda, T.(1985) Neurochem. Res. 10 779. Jansen, B.C. P. (1972) in: Sebrell, W.H. & Harris, R.S., eds., The Vitamins, Vol.5 (Academic Press) p. 99. Kaczmarek, M.J. & Nixon, RF. (1983) Clin. Chim. Acta 130 349. Karlberg, B.& Thelander, S.(1980) Analyt. Chim. Acta 114 129. Karlson, P. (1980,1984) Biochemie 11. Aufl. 1980; 12. Auf. 1984 (Thieme, Stuttgart). Karlson, P. (1984) Trends Biochem. Sci. 9 536. Kato, N., Higuchi, T., Sakazawa, C., Nishizawa, T., Tani, Y.& Yamada, H.(1982) Biochim. Biophys. Acta 715 143. Katz, D„ Metz, J.& van der Westhuyzen (1985) Amer. J. Clin. Nutr. 42 666. Kawai, C., Wakabayashi, Α., Matsumura, T.& Yui, Y.(1980) Amer. J.Med. 69 383. Kawasaki, T.(1979) Methods Enzymol. 62D 69. Kennedy, C.A. & McCleary, B.V. (1981) Analyst 106 344. Kimura, M., Fujita, T.& Itokawa, Y. (1982 a) Clin. Chem. 28 29. Kimura, M., Fujita, T., Nishida, S.& Itokawa, Y.J. Chromatogr. 188 417. Kimura, M.& Itokawa, Y.(1977) J. Neurochem. 28 389. Kimura, M.& Itokawa, Y.(1981) J. Chromatogr. 211 290. Kimura, M.& Itokawa, Y.(1983) Clin. Chem. 29 2073. Kimura, M.& Itokawa, Y.(1985) J. Chromatogr. 332 181. Kimura, M., Panijpan, B.& Itokawa, Y.(1982b) J. Chromatogr. 245 141,144. Kjesen, B.& Seim, S. H. (1977) Amer. J. Clin. Nutr. 30 1591. Koike, M.& Koike, K.(1982) Ann. Ν. Y. Acad. Sci. 378 225. Kolz, R , Lonsdorf, G.& Burg, G.(1980) Hausarzt 31 657. Koyama, S., Egi, Y., Shikata, H., Yamada, K.& Kawasaki, T.(1985) Biochem. Internat. 11371. Krampitz, L.O. (1969) Annu. Rev. Biochem. 38 213.

398

6 Thiamin, Vitamin B15 Aneurin

Krampitz, L.O. (1982) Ann. Ν. Y. Acad. Sci. 378 Kremer, A.B., Egan, R.M. & Sable, H.Z. (1980) J. Biol. Chem. 255 2405. Kiibler, W.(1980) in: Cremer, H.D., Hötzel, D.& Kühnau, J., eds., Biochemie und Physiologie der Ernährung, Bd. I (Thieme, Stuttgart) p. 606. Kuriyama, M., Yokomine, R., Arima, H., Hamada, R.& Igata, A.(1980) Clin. Chim. Acta 108 (1980) 159. Labadarios, D., Rossouw, J.E., McConnell, J.B., Davis, M.& Williams, R.(1977) Internat. J. Vitam. Nutr. Res. 47 17. Laffï, R., Tossani, N.& Marchi Marchetti, M.(1985) Internat. J. Vitam. Nutr. Res. 55 181. Lazarov, Y., Bohorov, 0 . & Doncheva, A.(1980) Comp. Biochem. Physiol. 66A 145. Leach, F.R., Carraway, C.A.C. (1979) Methods Enzymol. 62D 76. Leevy, C.(1980) Amer. J. Clin. Nutr. 33 172. Leevy, C.M. (1982) Ann. N. Y. Acad. Sei. 278 316. Lonsdale, D.& Shamberger, R. J. (1980) Amer. J. Clin. Nutr. 33 205. Lou, H.C. (1981) Arch. Neurol. 38 469. Lowe, Ρ.Ν., Leeper, F.J. & Perham, R.N. (1983) Biochemistry22 150. Lumeng, L., Edmondson, J.W., Schenker, S.& Li, T. (1979) J. Biol. Chem. 254 7265. MacLean, J.B. (1982) J. Amer. Med. Assoc. 248 1311. Mahmood, S., Dani, H. M. & Mahmood, A. (1984) Amer. J. Clin. Nutr. 40 226. Malathy, P.V. & Adiga, P.R. (1985) J. Biosci. 7 77. Mandel, H., Berant, M., Hazani, A.& Naveh, Y.(1984) N. Engl. J.Med. 311 836. Mansourian, R., Barclay, D.& Dirren, H. (1982) Internat. J. Vitam. Nutr. Res. 52 228. Martin, P.R., Majchrowicz, E., Tamborska, E., Marietta, C., Mukherjee, A.B. & Eckhardt, M. J. (1985) Science 227 1365. Mather, M., Schöpfer, L.M., Massey, V.& Gennis, R.B. (1982) J. Biol. Chem. 257 12887. Matsuda, T.& Cooper, J.R. (1981a) Proc. Natl. Acad. Sci. USA 78 5886. Matsuda, T.& Cooper, J.R. (1981b) Analyt. Biochem. 117 203. Matsunaga, T., Karube, I.& Suzuki, S. (1978) Analyt. Chim. Acta 98 25. McClean, H.E., Dodds, P.M., Stewart, A.W., Beaven, D.W. & Riley, C.G. (1976) N. Zealand Med. J. 84 345. McCormick, D.B. & Wright, L.D., eds., (1979) Methods Enzymol. 62D 51-125. Meilgaard, M.C. (1982) J. Stud. Alcohol 43 427. Miller, D.R. & Hayes, K.C. (1982) in: Hathock, J.N., ed., Nutritional Toxicity (Academic Press, New York) p. 81. Minami, J., Kishi, T.& Kondo, M.(1982) J. Gen. Microbiol. 128 2909. Mitsuda, H., Takii, Y., Iwami, K.& Yasumoto, K.(1975) J. Nutr. Sci. Vitaminol. 21 19. Mitsuda, H., Takii, Y., Iwami, K., Yasumoto, K.& Nakajima, K.(1980) Methods Enzymol. 62D 107. Mitsuda, H., Tanaka, T.& Kawai, F.(1970) J. Vitaminol. 16 263. Molin, W.T. et al. (1980) Plant Physiol. 66 308, 313. Muniyappa, K.& Adiga, P.R. (1979) Biochem. J. 177 887. Muniyappa, K.& Adiga, P.R. (1980) Biochem. J. 186 201. Muniyappa, K.& Adiga, P.R. (1981) Biochem. J. 193 679. Murata, Κ.. (1982) Ann. Ν. Υ. Acad. Sci. 378 146. Murphy, J.V. (1976) J. Nutr. Sci. Vitaminol. 22 69, Suppl. Murphy, J.V., Frerman, F.E. & Hodach, A. (1980) Neurochem. Res. 5 145. Naughten, F.R., Saul, I.P., Roche, G.& Mullins, C.(1985) J. Inher. Metab. Dis. 8 131, Suppl.2. Neumann, C.G., Swendseid, M.E., Jacob, M., Stiehm, E.R. & Dirige, O.V. (1979) Amer. J. Clin. Nutr. 32 99. Nishimune, T.& Hayashi, R.(1980) Experientia 36 916. Nishimura, H., Sempuku, K., Nosaka, K.& Iwashima, A.(1984a) J. Biochem. 96 1289. Nishimura, H., Uehara, Y., Sempuku, K.& Iwashima, A. (1984b) J. Nutr. Sci. Vitaminol. 301.

6.21 References

399

Nishimura, H., Yoshioka, K.& Iwashima, A.(1984c) Analyt. Biochem. 139 373. Nishino, K„ Itokawa, Y., Nishino, N., Piros, K.& Cooper, J.R. (1983) J. Biol. Chem. 258 11871. Nishino, K., Itokawa, Y., Nishino, N., Piros, K.& Cooper, J.R. (1982) Ann. N. Y. Acad. Sci. 378 453. Nishino, Α., Nishino, H.& Iwashima, A. (1980) J. Nutr. Sci. Vitaminol. 26 415. Nutrition Rev. 35 (1977) 185. Nutrition Rev. 37 (1979) 24. Nutrition Rev. 37 (1979) 261. Nutrition Rev. 38 (1980) 374. Nutrition Rev. 40 (1982) 53. Nutrition Rev. 40 (1982) 316. Nutrition Rev. 42 (1984) 328. Oda, R., Okumura, M., Yamano, S.& Yoshimura, H.(1984) Int. J.Clin. Pharmacol. Ther. Toxicol. 22 591. Ogawa, K.& Sakai, M.(1982) Ann. Ν. Y. Acad. Sci. 378 188. Oguchi, F., Okazaki, M., Hobara, R., Hajikano, M., Kato, H.& Sakamoto, K.(1980) Folia Pharm. Japónica 76 553, 567. Ohta, H., Baba, T., Suzuki, Y.& Okada, E.(1984) J. Chromatogr. 284 281. Oka, Y. (1984) Kirk-Othmer Encyclopedia of Chemical Technology, Vol. 24 (Wiley, NY) ρ.125. Panijpan, B.& Detkriangkraikun, P. (1979) Amer. J. Clin. Nutr. 32 723. Panijpan, B., Kimura, M.& Itokawa, Y. (1983) J. Chromatogr. 258 307. Panijpan, B.& Ratansubolchai, K.(1980) Internat. J. Vitam. Nutr. Res. 50 247. Parker, W.D., Haas, R., Stumpf, D.A., Parks, J., Eguren, L.A. & Jackson, C.(1984) Neurology 34 1477. Passia, D., Hilsher, B., Goslar, H. G., Hilscher, W.& Hofmann, N.(1979) Hormone Metab. Res. 11 697. Pattini, C.& Rindi, G.(1980) Internat. J. Vitam. Nutr. Res. 50 10. Pawlik, F., Bischoff, A.& Bitsch, 1.(1977) Acta Neuropathol. 39 211. Penttinen, H.K. (1978) Acta Chem. Scand. B32 609. Penttinen, H.K. (1979) Methods Enzymol. 62D 112. Peterson, D.R., Labbe, R.F., van Belle, G.& Chinn, N.M. (1981) Amer. J. Clin. Nutr. 34 65. Petzold, D.R., Hübner, G., Fischer, G., Neef, H.& Schellenberger, A. (1982) Studia Biophys. 87 15. Pincus, J. H., Solitare, G. R. & Cooper, J. R. (1976) Arch. Neurol. 33 759. Pipkin. J.D. & Stella, V.J. (1978) J. Pharm. Sci. 67 818. Plaitakis, Α., Nicklas, W.J. & Beri, S.(1978) Neurology 28 691. Pletcher, J., Sax, M., Turano, A.& Chang, C.H. (1982) Ann. Ν. Υ. Acad. Sci. 378 454. Poggi, V., Longo, G., DeVizia, B., Andria, G., Rindi, G., Pattini, C.& Cassandre, E. (1984) J. Inher. Metab. Dis. 7 153, Suppl.2. Puxty, J. A.H., Haskew, A. E., Ratcliffe, J.G. & McMurray, J.(1985) Ann. Clin. Biochem. 22 423, part 4. Ranhotra, G., Gelroth, J., Novak, F.& Bohannon, F.(1985) J. Nutrition 115 601. Ratanaubolchai, K.& Panijpan, B.(1979) Clin. Chem. 25 1670. Recny, M.A. & Hager, L.P. (1982) J. Biol. Chem. 257 12878. Reggiani, C., Pattini, C.& Rindi, G (1984) Brain Res. 293 319. Reinken, L., Stolley, H.& Droese, W.(1979) Eur. J.Pediatr. 131 229. Rindi, G.(1978) Boll. Soc. Ital. Biol. Sper. 54 TR5, Suppl. Rindi, G.(1982) Acta Vitaminol. Enzymol. 4 59. Rindi, G., Comincioli, V., Reggiani, C & Pattini, C.(1984) Brain Res. 293 329. Rindi, G.& Ferrari, G.(1977) Experiential 211. Rindi, G., Gastaldi, G., Casirola, D.& Ferrari, G.(1985) IR CS Med. Sci. 13 234. Rindi, G., Pattini, C., Comincioli, V.& Reggiani, C.(1980) Brain Res. 181 369.

400

6 Thiamin, Vitamin Bi, Aneurin

Rindi, G., Patrini, C.& Poloni, M. (1981) Experìentia 37 975. Roche, T. E., Brandt, D.R. & Pratt, M.L. (1982) Ann. N. Y. Acad. Sa. 378 462. Roche, T.E. & Lawlis, V.B. (1982) Ann. N. Y. Acad. Sci. 378 236. Rogers, E. F. (1962) Ann. N. Y. Acad. Sci. 98 412. Rogers, E. F. (1982) Ann. N. Y. Acad. Sci. 378 157. Rose, R.C. (1985) J. Inher. Metab. Dis. 8 13, Suppl.l. Roser, R.L., Andrist, A.H., Harrington, W.H., Naito, H.K. & Lonsdale, D.(1978) J. Chromatogr. 146 43. Rossouw, J.E., Labadarios, D., Krasner, N., Davis, M.& Williams, R.(1978) Scand. J.Gastroent. 13 133. Ruenwongsa, P.& Cooper, J.R. (1977) Biochim. Biophys. Acta 482 64. Ruenwongsa, P.& Pattanavibag, S.(1983) Nutr. Rep. Int. 27 713. Ruenwongsa, P.& Pattanavibag, S.(1984) Life Sci. 34 365. Sable, Η. Ζ. & Gubler, C. J., eds., (1982) "Thiamin" Ann. Ν. Y. Acad. Sci. 378 1-468. Sane, R.T., Doshi, V.J., Jukar, S., Joshi, S.K, Savant, S.V. & Pandit, U.R. (1985) J. Assn. Offic. Anal. Chem. 68 83. Sasser, L.B., Hall, G.G., Bratton, G.R. & Zmudzki, J.(1984) J. Nutrition 114 1816. Säuberlich, Η. E. (1984) Annu. Rev. Nutr. 4 377. Säuberlich, Η.E., Herman, Y.F., Stevens, C.O. & Herman, R.H. (1979) Amer. J.Clin. Nutr. 32 2237. Sauberlich, H.E., Kretsch, M.J., Johnson, H.L. & Nelson, R.A. (1982) in: Beitz, D.C. & Hansen, R. G., eds. Animal Products in Human Nutrition (Academic Press, New York) p. 339. Scheiner, J.M., Araujo, M.M & Deritter, E.(1981) Amer. J.Hosp. Pharm. 38 1911. Schellenberger, A.(1967) Angew. Chemie 79 1050. Schellenberger, A.(1982) Ann. N. Y. Acad. Sei. 378 51. Schrijver, J., Dias, T.& Hommes, F. Α. (1978) Neurochem. Res. 3 699. Schwartau, M., Doehn, M.& Bause, H.(1981) Klin. Wochenschr. 59 1267. Sen, I.& Cooper, J.R. (1976) Neurochem. Res. 1 65. Shin, W., Pletcher, J., Sax, J.& Blank, G.(1979) J. Amer. Chem. Soc. 101 2462. Ski an, D.& Trostler, N.(1977) J. Nutr. 107 353. Soliman, A. G. M. (1981) J. Assn. Offic. Anal. Chem. 64 616. Somogyi, J.C., Kopp, P.M., Filippini, L.& Monnat, Α.(1980) J. Nutr. Sci. Vitaminol. 26 221. Soukop, M.& Calman, K.C. (1978) Brit. J. Cancer 38 180. Spector, R.(1982) Ann. N. Y. Acad. Sci. 378 344. Sümegi, B.& Alkonyi, 1.(1983) Eur. J.Biochem. 136 347. Takahe, M.& Itokawa, Y.(1980) Experientia 36 327. Takabe, M.& Itokawa, Y.(1983) J. Nutr. Sci. Vitaminol. 29 509. Tate, J., Kaczmarek, M.J. & Nixon, P.F. (1984) Clin. Chim. Acta 137 81. Thérisod, M., Fischer, J.& Estramareix, B.(1981) Biochim. Biophys. Res. Comm. 98 374. Thompson, R.H.S. (1983) Trends Biochem. Sci. 8 460. Thornber, E.J., Dunlop, R.H. & Gawthorne, J.M. (1980) J. Neurochem. 35 713. Thurnham, D.I. (1981) Proc. Nutr. Soc. 40 155. Trostler, Ν., Guggenheim, Κ., Havivi, E.& Sklan, D.(1977) Nutr. Metab. 21 294. Ullrich, J.(1982) Ann. Ν. Y. Acad. Sci. 378 287. Ullrich, J., Deus, B.& Holzer, Η. (1968) Internat. J. Vitam. Nutr. Res. 38 273. Ullrich, J., Ostrovsky, Y.M., Eyzaguirre, J.& Hölzer, H.(1970) Vitamins Hormones 28 365. Uray, G.(1982) Monatsh. Chemie 113 1475. Usmanov, R. A. & Kochetov, G. A. (1982) Biochemistry Internat. 5 727. Usmanov, R. Α., Neef, H., Pustynnikov, M.G., Schellenberger, A.& Kochetov, G. A. (1985) Biochemistry Internat. 10 479. Van der Westhuyzen, J., Davis, R.E., Icke, G.C. & Metz, J.(1985) Internat. J. Vitam. Nutr. Res. 55 173.

6.21 References

401

Van Dort, H. M., van der Linde, L.M., de Rijke, D.(1984) J. Agrie. Food Chem. 32 454. Vimokesant, S., Kunjara, S., Rungruangsak, K., Nakornchai, S. & Panijpan, B. (1982) Ann. N. Y. Acad. Sei. 378 123. Vimokesant, S., Nakornchai, S., Rungruangsak, K., Dhanamitta, S.& Hilker, D.M. (1976) J. Nutr. Sei. Vitaminol. 22 1, Suppl. Vir, S.C. & Love, A. H. G. (1979) Internat. J. Vitam. Nutr. Res. 49 291. Vitamin Compendium (1980) (Hoffmann-La Roche, Basel). Volpe, J.J. & Marasa, J.C. (1978) J. Neuroehem. 30 975. Voskoboyev, A. I. & Ostrovsky, Y.M. (1982) Ann. N. Y. Acad. Sei. 378 161. Wakabayashi, Y., Iwashima, A.& Nose, Y.(1979) Methods Enzymol. 62D 105. Wakabayashi, Α., Yui, Y.& Kawai, C.(1979) Japan. Cireulatio. J. 43 995. Waldenlind, L.(1978) Acta Physiol Scand. 459 1, Suppl. Waldenlind, L. (1979 a) Acta Physiol. Scand. 105 1. Waldenlind, L. (1979b) Nutr. Metab. 23 38. Waldenlind, L., Borg, S.& Vikander, B.(1981) Acta Med. Scand. 209 209. Waldenlind, L., Elfman, L.& Rydquist, B.(1978) Acta Physiol. Scand. 103 154. Walter, W.& Bacher, A. (1977) J. Gen. Microbiol. 103 359. Warnock, L.G. (1982) Analyt. Biochem. 126 394. Warnock, L.G., Prudhomme, C.R. & Wagner, C.(1978) J. Nutrition 108 421. Weber, W.& Kewitz, H.(1985) Eur. J.Clin. Pharmacol. 28 213. Wendel, U., Lombeck, I.& Bremer, H.J. (1983) N. Engl. J.Med. 308 1100. Whelan, W.J. (1976) Trends Biochem. Sci. 1189. White, R.H. (1981) Science 214 797. White, R L . & Spenser, I.D. (1979) Biochem. J. 179 315. White, R.L. & Spenser, I.D. (1982) J. Amer. Chem. Soc. 104 4934. Wick, H., Schweizer, K.& Baumgartner, R.(1977) Agents Actions 1 405. Wielders, J.P.M. & Mink, C.J.K. (1983) J. Chromatogr. 277 145. Wimalasiri, P.& Wills, R.B.H. (1985) J. Chromatogr. 318 412. Witt, E.D. & Goldman-Rakic, P.S. (1983) Ann. Neurol. 13 376, 396. Wood, T.(1981) Biochim. Biophys. Acta 659 233. Wood, B.& Breen. K.J. (1980) Med. J.Australia 1 461. Wuest, H. M. (1962) Ann. N. Y. Acad. Sci. 98 385. Yagi, N.& Itokawa, Y.(1979) J. Nutr. Sci. Vitaminol. 25 281. Yamada, K.& Kawasaki, T.(1980) J. Bacteriol. 141 254. Yamada, K., Kumaoka, H.(1982) Biochemistry Internat. 5 771. Yamada, K.& Kumaoka, H.(1983) J. Nutr. Sci. Vitaminol. 29 389. Yamada, K., Morisaki, M.& Kumaoka, H.(1983) Biochim. Biophys. Acta 756 41. Yamada, K., Yamamoto, M., Hayashiji, M., Tazuya, K.& Kumaoka, H. (1985) Biochem. Int. 10 689. Yamamoto, S., Koyama, S.& Kawasaki, T. (1981) J. Biochem. 89 809. Yoshimura, K., Nishibe, Y., Inoue, Y., Hirono, S., Toyoshima, K.& Minesita, T. (1976) J. Nutr. Sci. Vitaminol. 22 429. Yoshioka, Κ (1984) Biochim. Biophys. Acta 778 201. Zoltewicz, J. Α., Kaufïman, G.& Uray, G.(1982) Ann. Ν. Y. Acad. Sci. 378 7. Zoltewicz, J. Α., Uray, G., Baugh, T.D. & Schultz, Η. (1985) Bioorg. Chem. 13 135.

7 Vitamin B2: Riboflavin and Its Bioactive Variants

7.1 7.2 7.3 7.3.1 7.3.2 7.3.3 7.3.3.1 7.3.3.2 7.3.3.3 7.3.3.4 7.3.4 7.3.5 7.3.6 7.4 7.5 7.5.1 7.5.2 7.5.3 7.6 7.6.1 7.6.1.1 7.6.1.2 7.6.1.3 7.6.1.4 7.6.2 7.6.2.1 7.6.3 7.6.4 7.7 7.8 7.9 7.9.1 7.9.1.1 7.9.1.2 7.9.1.3 7.9.2 7.9.2.1 7.9.3 7.9.4 7.9.5 7.9.5.1 7.9.6 7.9.6.1

Introduction, History Isolation, Purification and Commercial Preparations Nomenclature, Structure, Properties Nomenclature Structure Chemical Reactivity Redox Reactions Oxidized Flavins, Flox Dihydroflavins, FlredH2 Flavin radicals Solubility, Crystallization Spectra Photochemistry Chemical Synthesis Analysis Microbiological Methods Physical-Chemical Methods Other Methods Biosynthesis Mechanism of Biosynthesis in Microorganisms Reductase and Deaminase Reactions Formation of 6,7-Dimethyl-8-ribityllumazine Riboflavin Synthase Reaction Biosynthesis of the Coenzyme Forms of Riboflavin Biosynthesis of the Coenzyme Forms and Their Binding to the Apoenzymes in the Animal Organism Regulation Covalent Linkage of Coenzyme and Apoenzyme Enzymatic Methods for Preparative Production of Flavins Occurrence, Including Occurrence in Foods Stability, Especially in Foods Flavin-Dependent Enzymatic Reactions and Enzymes One-Electron Transferases Bacterial Flavodoxins Electron-Transfer Flavoprotein, ETF (EC 1.3.99.2-3) ETF-Ubiquinone Reductase Dehydrogenases (Pyridine-Nucleotide Dependent) NADPH-Methemoglobin Reductase Dehydrogenases (Not Requiring Pyridine Nucleotides) Pyridine Nucleotide Disulfide Oxidoreductases Reactions of Reduced Flavoproteins with Oxygen Flavoprotein Oxidases (0 2 -»• H 2 0 2 ) Flavoprotein Monooxygenases Bacterial "Aromatic" Hydroxylases

404

7 Vitamin B2: Riboflavin and Its Bioactive Variants

7.9.6.2 Bacterial Luciferases 7.9.7 On the Role of the Flavoproteins in Human Oxygen Metabolism 7.9.8 Riboflavin as Attractant 7.9.9 Isolation and Analysis of the Flavoproteins 7.9.10 Congenital Defects in Flavin-Dependent Enzymatic Reactions 7.10 Absorption, Transport, Distribution, Storage, Metabolism and Excretion in Human Beings and Animals 7.10.1 Intestinal Absorption 7.10.2 Transport, Transport Proteins and Other Riboflavin-Binding Proteins 7.10.2.1 Estrogen-Inducible Riboflavin-Binding Proteins (RFBPs) 7.10.2.2 Riboflavin Transport in the Brain 7.10.3 Distribution, Storage 7.10.3.1 Milk 7.10.3.2 Whole Blood 7.10.3.3 Eye 7.10.3.4 Various Organs 7.10.4 Metabolism 7.10.5 Excretion 7.11 Uptake into the Cell and Metabolism in Microorganisms and Plants 7.12 Riboflavin Deficiency in Human Beings and Animals 7.12.1 Important Causes 7.12.1.1 Insufficient Dietary Intake 7.12.1.2 Alcoholism 7.12.1.3 Oral Contraceptives 7.12.1.4 Phototherapy of New-Born Infants 7.12.2 Deficiency Symptoms in Human Beings 7.12.3 Deficiency Symptoms in Animals 7.12.3.1 General Symptoms 7.12.3.2 Impairment of Enzyme Functions 7.12.4 Determination of Deficiency: Riboflavin Status 7.12.4.1 Measurement of the EGR Activity 7.12.4.2 Measurement of Riboflavin Excretion in the Urine 7.12.4.3 Other Methods 7.12.4.4 Riboflavin Status in Old Age 7.12.4.5 Riboflavin Status and Malaria 7.13 Riboflavin Requirements in Human Beings and Animals 7.14 Therapeutic Application of Riboflavin and Its Analogs 7.15 Some of the More Important Synthetic Analogs of Riboflavin 7.15.1 Riboflavin Analogs with Modified Ribityl Residues 7.15.2 Riboflavin Analogs with a Side Chain Other than Ribityl 7.15.3 1- and 5-Deazaflavins 7.15.4 Flavins Modified in Positions 6,7 and 8 of the Isoalloxazine Ring 7.16 Flavin and Flavoprotein Antagonists 7.17 Toxicology 7.18 References

7 Vitamin B2: Riboflavin and Its Bioactive Variants

7.1 Introduction, History Soon after the isolation of vitamin B, (thiamin) from yeast concentrates, the search began for a "second nutritional factor", whose existence in the same substrate was suspected1. This "second factor" appeared to be a thermostable growth factor which had anti-pellagra activity (it healed the dermatitis in rats). However, it soon became apparent that this "second factor" was not a single substance (in addition to riboflavin, it contained pyridoxine, pantothenic acid and nicotinic acid). Finally, using the rat test and the compound's yellow-green fluorescence as assays, R.Kuhn, P.György & T. Wagner-Jauregg (1933) isolated pure riboflavin2 from yeast, egg white and whey. Its structure was established by R.Kuhn and Wagner-Jauregg in 1933-34, and it was synthesized in 1934-35 by R.Kuhn, F. Weygand & P.Karrer. About the same time, the flavin coenzymes were discovered. In 1932, O.Warburg and W.Christian isolated the "old yellow enzyme" from yeast; its coenzyme was found to be flavin mononucleotide, FMN (H.Theorell, 1934). The synthesis of FMN was accomplished in 1936 by R. Kuhn and H. Rudy. In 1938, O.Warburg discovered flavin adenine dinucleotide, FAD, as the coenzyme of D-amino acid oxidase. Other flavin-dependent enzymes were discovered thereafter at a rapid rate. Most of them were found to contain a firmly bound coenzyme (Ad < 10"6M. However, it could usually be removed by denaturation (heat, methanol or acids), dialysis in the presence of salts, or ammonium sulfate precipitation. In 1955, it was observed for the first time that some tissue flavins can be extracted only after proteolysis, which speaks for a covalent binding of these flavins to their apoproteins. It was found that most of the "insoluble flavin" is associated with the succinate dehydrogenase of the inner mitochondrial membrane. In 1960, E.B. Kearney isolated a small peptide with covalently bound FAD from a trypsin/chymotrypsin hydrolysate of succinate dehydrogenase from bovine heart. Not until 1969 (P. Hemmerich et al.) was it proven that the bonding between FAD and the apo-succinate dehydrogenase involves C-8a. This demonstration depended mainly on spectroscopic methods, including ESR. The reactivity of the C-8a group is indicated, for example, by the fact that its H atoms exchange for deuterium in D 2 0 rather quickly at pH 6.8. Halogens bound to C-8a 1

2

Historical reviews: Wagner & Folkers (1964), Wagner-Jauregg (1972), Hemmerich (1976), Sebrell ((1979), Nielsen, Rauschenbach & Bacher (1983) General reviews: Rivlin (1975), Singer (1976), Yagi & Yamano (1980), Yagi (1975), Hemmerich (1976), Walsh (1980), Hemmerich et al. (1977), Foy & Mbaya (1977), McCormick & Wright (1980), Vitamin Compendium (1980), Merrill et al. (1981), Bässler & Lang (1981), Massey & Williams (1982), Yoneda (1984), Bray et al. (1984), Cooperman & Lopez (1984)

406

7 Vitamin B2: Riboflavin and Its Bioactive Variants

are also very reactive. The immediate binding partner of the FAD was discovered by degrading succinate dehydrogenase in various ways. One of the products was a pentapeptide with the sequence Ser-His(FAD)-Thr-Val-Ala, and then a fragment consisting of flavin and histidine (P. Hemmerich et al., T. P. Singer et al., 1970-72). The binding of the flavin via N-3 of the histidine was then determined by comparison with synthetic histidyl flavins (T. P. Singer et al., 1971). On other types of binding, cf. p. 424. The flavosemiquinones, and thus the first stable flavin radicals, were discovered quite early (R. Kuhn, 1937), but their exact characterization had to wait for about 20 years (H. Beinert, 1956). In addition to riboflavin, FMN and FAD, a number of other biologically active flavins have been found in microorganisms and plants, including factor 420, which has 5-deazaflavin as its chromophore. Synthetic 5-deazariboflavin has a number of very interesting properties. The research in flavins has become very active in the last few years. This is expressed by the large number of international flavin symposia, among other things.

7.2 Isolation, Purification and Commercial Preparations The flavins are light-sensitive. Light attacks the ribityl side chain, forming lumiflavin or lumichrome. Therefore, all operations on flavins must be carried out in the dark or in red light, in order to avoid photochemical degradation reactions. If FAD is treated with acetic acid, this must be done with cooling, because otherwise the molecule is degraded to FMN [Decker & Hamm (1980)]. Many methods for purification of flavins have been described, especially chromatographic methds. Some of these shall be briefly mentioned here: A preparative method for obtaining FMN by column chromatography makes use of DEAEcellulose and silicagel in ethanol/lM triethylammonium hydrogencarbonate, pH 7.5 [Don Johnson et al. (1978)]. The isolation of FAD from Sorcina lutea, cultured in the presence of adenine and riboflavin, consists of the following steps: heating the culture medium to 80°C for 3 min; centrifugation; filtration of the supernatant, which contains 8 g FAD, through a column containing 1.5 1 Florisil; washing the column with 3 1 0.5% acetic acid; elution with phenol/ acetic acid/water, 1:6:6 (by vol.); chromatography of the FAD fraction on Amberlite IRA-401 (CI - form); phenol extraction; transfer of the FAD to the aqueous phase. At this point, the FAD is 75% pure, and the yield is 60%. Very pure FAD can be obtained by chromatography on p-acetoxymercurianiline-agarose [Chibata et al. (1980)]. The extraction, separation and purification of radioactively labelled FAD and FMN from Clostridium kluyveri consists in principle of extraction of the bacterial cells with perchloric acid, adsorption on DEAE-cellulose, elution of FMN with 3mM HCl/15mM LiCl, elution of FAD with 3mM HCl/35mM LiCl, and deionization on Florisil [Decker & Hamm (1980)]. FAD and its analogs were extracted and purified by affinity chromatography on a column of Sepharose 4 Β with covalently bound glucose oxidase and D-amino acid oxidase. The bound enzymes were quite stable; the half-life was about 11 months. However, ion-exchange and gel chromatography have also been used for this purpose. Flavodoxin from Peptostreptococcus elsdenii bound to Sepharose 4 Β is very good for the purification of FMN and its analogs by affinity chromatography [Massey & Mendelsohn (1979)].

7.3 Nomenclature, Structure, Properties

407

Reduced riboflavin is only very slightly soluble. This property is utilized in the extraction and purification of the vitamin. Reduction, e.g. with dithionite, causes it to precipitate; it can then be separated and reoxidized [Cooperman & Lopez (1984)]. Riboflavin is widely used in the pharmaceutical industry and for the enrichment of foods and feeds. The pharmaceutical industry produces tablets for oral administration and solutions for injection [Yoneda (1984)]. Because of the poor solubility of riboflavin (10-13 mg/100 ml water at room temperature), tryptophan and nicotinamide are added to solutions for oral or parenteral application. This increases the solubility. However, the sodium salt of riboflavin 5'-phosphate (FMN), which is quite soluble, is commercially available [Cooperman & Lopez (1984)]. The salt used is the monosodium dihydrate; it is about 200 times more soluble in water than riboflavin, and it is therefore suitable for preparing aqueous solutions without recourse to solubilizing agents (these are necessary for riboflavin) [Yoneda (1984)].

7.3 Nomenclature, Structure, Properties 7.3.1 Nomenclature The chemical name for riboflavin is 7,8-dimethyl-10-(l'-D-ribityl)isoalloxazine (Fig. 7-1 a). The terms "lactoflavin", "ovoflavin", "uroflavin", etc., were used formerly, but now are of historical interest only. There are also two radical forms of riboflavin, the anion (Fig. 7-1 b) and the neutral molecule (Fig. 7-1 c), and dihydroriboflavin (Fig. 7-1 d). The redox forms are named as follows : the fully oxidized form is flavoquinone (Flox); the flavin radical is flavosemiquinone (F1H) and the fully reduced form, flavohydroquinone (FlredH2, dihydroflavin, leucoflavin) [Hemmerich (1976)]. The accepted names of the coenzyme forms of riboflavin are flavin mononucleotide (FMN, Fig. 7-2 a) and flavin adenine dinucleotide (FAD, Fig. 7-2 b). These are not actually correct, since FMN is not a nucleotide, and FAD is not a dinucleotide. Some authors would therefore have preferred the names riboflavin monophosphate and riboflavin adenine diphosphate. Some authors also use the name "riboflavin" for the coenzyme forms of the vitamin, which often makes the interpretation of their publications difficult. Some authors take the name "vitamin B2" as a general term, and the names "riboflavin", "FMN" and "FAD" as specific terms, which seems more practical. In this text, the second practice will be followed where possible. 7.3.2 Structure Riboflavin (Ci7H2oN406, rei. molecular mass, 376.4) is a yellow, tricyclic molecule (Fig. 7-1 a) which is usually phosphorylated in biological systems, and then adenylated. This produces the two coenzyme forms, flavin mononucleotide

408 (a)

7 Vitamin B 2 : Riboflavin and. Its Bioactive Variants CH 2 0H HOCH

Ir

HOCH

Ir Ir

HOCH

Fig. 7-1. Structural formulae of riboflavin in various redox states. a. Oxidized riboflavin, FloX b. Red radical anion, Fl~ c. Blue neutral radical, Fl 1 H d. 1,5-Dihydroriboflavin, FlredH2 R=Ribityl. Formulae taken from Yoneda [Yoneda (1984), p. 108]

(FMN, Fig. 7-2 a) and flavin adenine dinucleotide (FAD, Fig. 7-2 b). Some important commercial forms are sodium riboflavin phosphate and free riboflavin. Riboflavin 5'-phosphate, FMN (456.6 daltons), was first obtained from the "yellow enzyme" by O.Warburg and W.Christian in 1932.The coenzyme was separated from the enzyme by acidification and dialysis. It was first synthesized chemically by R. Kuhn. Flavin adenine dinucleotide, FAD (785.56 daltons) was first extracted from D-amino acid oxidase and identified in 1938 by O. Warburg and W.Christian. The first synthesis was achieved by A.R. Todd (1952) [Yoneda (1984)]. The two ring systems (isoalloxazine and adenine) of FAD are probably arranged one above the other and are nearly coplanar; this is suggested by the Xray analysis of a crystalline 1:1 complex of riboflavin and adenine [Fujii et al. (1977)]. Many flavins, which differ structurally and functionally from riboflavin, FMN and FAD, have been discovered in the organic world. Their physiological roles are not known in every case. Several of these analogs are discussed briefly below. Riboflavin 2',5'-cyclophosphate was extracted from Aspergillus oryzae. Several riboflavinyl 5'-glycosides have been isolated, but their physiological roles are not known. Glycosides of

7.3 Nomenclature, Structure, Properties

409

NH,

(α)

CH20-P-0H

It·

HOCH Ir HOCH Ir HOCH

OH

Ir

H,C H,C

Fig. 7-2. Structural formulae of the coenzyme forms of riboflavin. a. Flavin mononucleotide, FM Ν b. Ravin adenine dinucleotide, FAD riboflavin are also formed easily by irtww-glycosidases in rat liver, plants and microorganisms. Maltose, dextrin, starch and glycogen may be glycosyl donors for riboflavinyl glucoside; lactose for riboflavinyl-galactoside, and sucrose for riboflavinyl fructoside. However, these glycosides are not normally found in significant concentrations [Merrill et al. (1981 b)]. Riboflavin analogs in which the 5'-hydroxyl group is oxidized to an aldehyde (in riboflavinal) or carboxyl group (in riboflavinoic acid) have been isolated from Schizophyllum commune, and are also found in edible Basidiomycetes ; in a Candida species, they stimulate fermentation [Merrill et al. (1981b), Tachibana et al. (1979)]. 5'-Malonylriboflavin has been isolated from young oat plants. The 5'-malonyl ester bond is relatively labile to hydrolysis and photolysis. This novel flavin is probably a blue-light receptor in phototropic organisms; its function would be to convert the light into a signal which orients the growth in the direction of the light. In this case, 5'-malonylriboflavin would have a major role in the plant kingdom [Ghisla et al. (1984)]. 6-Hydroxy-FMN was isolated as the cofactor of the glycolate oxidase from pork liver. 6-Hydroxy-FAD is found in the electron-transfer flavoprotein from Megasphaera elidenti [Merrill et al. (1981 b), Yoneda (1984)]. 8-Hydroxy-FAD was isolated from D-lactate dehydrogenase from M. elsdenii [Walsh (1984), Merrill et al. (1981b)]. Roseoflavin, 405.4 daltons, was isolated from the culture medium of Streptomyces davawensis as fine, red-brown needles. The 8-methyl group of riboflavin is replaced by a dimethylamino group in this compound. Roseoflavin is an antibiotic which is effective against gram positive bacteria [Yoneda et al. (1984)]. Roseoflavin is phosphorylated by the flavokinase of rat liver; it is an antagonist of riboflavin in the rat and is toxic for poultry [Merrill et al. (1981 b)]. R. Wolfe et al. isolated factor 420 (F42o) from methane bacteria in 1975; they identified it as a 7,8-didemethyl-8-hydroxy-5-deazariboflavin derivative. Its side chain is ribityl 5'-phosphate, and this has a phosphodiester bridge to lactyl diglutamate. Factor (F4M) has an absorbance maximum at 420 nm. It participates in methanogenesis, and has several other functions as well. F420 is found in Streptomycetes as well as in methane bacteria [Walsh (1984), Yamazaki et al. (1982), Jacobson & Walsh (1984)]. 5-Deazariboflavin (with CH instead of Ν in position 5) was synthesized as early as 1970. It differs considerably from riboflavin with respect to its redox behavior, in that it, like the pyridine nucleotides, can take part only in 2-electron transfers [Walsh (1984)].

410

7 Vitamin B2: Riboflavin and Its Bioactive Variants

There are structural and functional analogies between flavins and pteridines (see chapter on Folic Acid). Both classes of compound are redox cofactors of enzymes which catalyse hydroxylation reactions. Pteridines and flavins both contain a pyrimidine and a pyrazine ring in condensed form. The structural analogies are a result of the biosynthetic pathways, which are shared in part. Both types of cofactor are formed from GTP, and the first steps of the synthesis are identical (see chapter on Folic Acid). Both pteridines and flavins activate molecular oxygen by forming angular hydroperoxides [Wetzel & Ghisla (1983)]. 7.3.3 Chemical Reactivity The catalytic properties of the flavins are due to the unique combination of benzenoid, pyrimidine and pyrazine rings to the isoalloxazine ring system. The catalytic functions of the flavins are carried out primarily by the positions N-l, C-4a and N-5. These are components of the pyrazine and pyrimidine rings. The benzenoid ring was long thought to be chemically inert, but now it is known that position 8 is very reactive. The methyl group bound at the 8 position exchanges protons in D 2 0, and enters covalent bonds with enzyme proteins. An antibiotic, roseoflavin (cf. p. 5), is a riboflavin analog with -N(CH3)2 at C-8. Position 6 is also reactive. Recently 6-S-cysteinyl-FMN was found as the covalently bound coenzyme of trimethylamine dehydrogenase [Moore et al. (1979), Singer & Mclntyre (1984)]. Riboflavin can be relatively easily esterifled in the ribityl moiety. Many fatty acid esters of this type, including riboflavin 5'-butyrate and riboflavin 5'-palmitate, have been reported [Okuda & Horiguchi (1980 b)]. Riboflavin forms a dark red silver salt. Cu + and Hg 2+ form similar complexes; these are 1:1 adducts. Their color indicates charge transfer between the metal and the flavin [Yoneda (1984)]. When reduced with sodium dithionite (Na 2 S 2 0 4 ), zinc in acid or catalytically activated hydrogen, riboflavin takes up two hydrogen atoms and forms the almost colorless 1,5-dihydroriboflavin (Fig. 7-1 d). This is reoxidized in the presence of air [Yoneda (1984)]. The flavin coenzymes are among the most versatile redox cofactors known to biochemistry. Since flavins can participate in either one- or two-electron redox reactions, they act as switching sites between the two types of electron transfer. When reduced, they can also react with molecular oxygen, due to their ability to exist as radicals [Walsh (1984)]. The structures of oxidized and reduced flavins have been studied in detail by 13 C and 15N NMR. These and other methods were used to show that reduced flavin has a bent ("butterfly") conformation [Moonen et al. (1984b)]. The binding of flavins to apo-flavoproteins is often covalent. It has been an object of intense investigation. X-ray analyses, e. g. of the structures of glutathione reductase and 4-hydroxybenzoate hydroxylase, have given good insight into the nature of the flavin-protein interaction [Merrill et al. (1981b)].

7.3 Nomenclature, Structure, Properties

411

7.3.3.1 Redox Reactions The flavin coenzymes, along with the nicotinamide coenzymes, are among the most important electron acceptors and donors in biological redox systems. The tricyclic flavins, however, are in some ways more versatile catalysts than the monocyclic nicotinamides, a) The nicotinamides always transfer two electrons at a time, and are apparently not able to take part in biological one-electron processes; the flavins can do both, b) Reduced nicotinamides are inert to oxygen, but dihydroflavins can react with 0 2 . Due to these properties, flavins can act, on the one hand, as redox switches between obligate two-electron donors (e.g. NADH, succinate) and obligate one-electron acceptors (iron-sulfur proteins, heme proteins) in cellular redox metabolism. On the other hand, thanks to their ability to react with molecular oxygen, flavins can serve as two types of cofactor: a) in the two-electron reduction of 0 2 to H 2 0 2 , and b) in the reductive four-electron activation and cleavage of 0 2 in the monooxygenase reactions according to Eq.l (X = flavin): X H 2 + 0 2 + substrate-H — X + H 2 0 + substrate - OH

(1)

In the redox process, the flavin molecule changes its shape; it is planar in the oxidized form and folded in the reduced form. If the apoprotein has different affinities for the oxidized and reduced forms of the flavin, this affects the redox potential of the bound flavin [Walsh (1980), Hemmerich (1976)]. The mechanism by which flavins are reduced by dihydropyridines has been thoroughly examined. Non-enzymatic reduction of various flavin analogs by 1-propyl-dihydronicotinamide indicated a hydride transfer (2 e~ transfer). There is also other evidence for the hydride transfer, including the fact that dihydronicotinamide is not able to reduce a flavin radical to the dihydroflavin by a oneelectron transfer [Powell & Bruice (1983)]. 7.3.3.2 Oxidized Flavins, Flox Oxidized flavin, or flavoquinone, is the best known form of the flavins, due to its characteristic light absorption and fluorescence (p. 413), and its stability in the presence of 0 2 . Binding to proteins shifts the absorption bands to longer wavelengths, and it can also quench the fluorescence of the flavin. The flavoquinone is protonated at N-l (pK about 0.0). The yellow flavin chromophore was useful in many kinetic experiments with flavoproteins due to its typical light absorption and fluorescence [Hemmerich (1976), Walsh (1980)]. 7.3.3.3 Dihydroflavins, FlredH2 Dihydroflavins (in contrast to Flox) are colorless, but only in dilute solutions, and do not fluoresce in protein-free solutions. There are prohably only a few biological functions of flavins in which FlredH2 does not participate [Hemmerich (1976)]. Flavins and flavoproteins are reduced with dithionite, borohydrides and hydrogen in the presence of catalysts, as well as by natural substrates. The reduc-

412

7 Vitamin B 2 : Riboflavin and Its Bioactive Variants

.0

H '

H

0

1,5-Dihydroflavin

4a,5-Dihydroflavin

Fig. 7-3. Two forms of dihydroflavins. The substituent in position 10 was left out. [Ghisla (1980), p. 360]

tion with hydrogen produces only 1,5-dihydroflavins. A two-electron reduction of Flox, however, can lead formally to several tautomeric reduced forms. Only two forms appear to be significant in enzymatic catalysis, namely 1,5-dihydroand 4 a,5-dihydroflavins (Fig. 7-3) [Ghisla (1980)]. Reduction of the flavins by thiols has been discussed in detail. Spectrophotometric experiments suggest the formation of thiol-flavin adducts via C-4a of the flavin [Loechler & Hollocher (1980)]. The most important biological acceptors of the electrons from reduced flavins are a) Fe 3+ in cytochromes and adrenodoxines in the flavoprotein one-electron transfer reactions, b) flavin itself, e.g. in electron transfer flavoprotein, and c) molecular oxygen in all flavoprotein oxidases and monooxygenases ; oxygen can react with the flavins in three ways: as a 1 e " acceptor, forming the superoxide radical O2, as a 2 e~ acceptor, forming hydrogen peroxide, and as 4 e~ acceptor in the flavoprotein monooxygenases [Hemmerich (1976)]. N-l of the flavin ring system is the most basic, but not the most nucleophilic atom (the steric hindrance from the substituents at N-10 is responsible for this). The reduced flavin ring system is most often alkylated at N-5, although C-4a is also an important alkylation site. The two sites compete with one another, regardless whether the alkylation reaction is electrophilic or nucleophilic [Hemmerich (1976)]. Free flavohydroquinones undergo a rapid inversion at N-5 and N-10, even when the substituent groups are large. This change in conformation may be the reason that free reduced flavins in principle do not fluoresce in solution. Low temperature ("freezing" of the oscillation) restores the fluorescence [Hemmerich (1976)]. 7.3.3.4 Flavin radicals At least two forms of the flavosemiquinones exist under physiological conditions: binding of one electron to Fl0x leads to the red radical anion Fl~ ( λ ^ 490 nm; Fig. 7-1 b). In a protein-free system, this is protonated, pK ca.8.5, and forms the blue neutral radical 5-HF1 (Xmax 570 nm; Fig.7-1 c). Thermodynamically, this blue radical is moderately stable in protein-free systems. It disproportionates to Flox and FlredH2 via a reactive dimer (eq.2): 2 Η Fl * (HF1)2 ^ FTox+Fl^Hj

(2)

The blue radical 5-HF1 can be stabilized, biochemically by binding a suitable apoprotein, or chemically, by alkylation to 5-RFl· [Hemmerich et al. (1977)]. The

7.3 Nomenclature, Structure, Properties

413

neutral radical form HR· binds a proton (pK ca.3.0); the red cation H2F1+ was crystallized by R. Kuhn in 1937 [Hemmerich (1976)]. The one-electron reduction of riboflavin bound to the protein RFBP, (p. 446) yields first the anionic semiquinone (Fig. 7-1 b), which is then rapidly converted to the stable neutral semiquinone (Fig. 7-1 c) [Klapper & Faraggi (1983)]. The one-electron reduction potentials for riboflavin, FMN and FAD have been determined [Anderson (1983)]. 7.3.4 Solubility, Crystallization Riboflavin (376.4 daltons) is moderately soluble in water (10-13 mg/100 ml at 25-27°C) and in absolute ethanol (4.5 mg/100 ml at 27.5°C). It is insoluble in ether, chloroform or acetone. It is very soluble in dilute basic solutions, but the solutions are unstable. Riboflavin crystallizes out of water, alcohol or pyridine in fine, yellow-orange needles which have a bitter taste. They melt at 278-282°C with decomposition. Polymorphic crystalline forms of riboflavin are known. Aqueous solutions of riboflavin fluoresce with a greenish yellow color; the fluorescence is maximal at pH 3-8, and is quenched by acidifying the solution or making it more basic. The optical activity of riboflavin in neutral and acid solution is [afe0 = +56.5-59.5° (0.5%, dilute hydrochloric acid). Riboflavin cannot be extracted with the usual organic solvents. It can be extracted with CHCI3 as lumiflavin (for analytical purposes) after photochemical cleavage of the ribityl residue. Riboflavin is very well adsorbed from aqueous solution by activated charcoal, but elution is very difficult [Werner, Gani & Fresenius (1980), Yoneda (1984), Foy «fe Mbaya (1977)]. Riboflavin 5'-phosphate (FMN), 456.5 daltons, forms fine yellow crystals which melt at 195°C; [a]g = +44.5° (2%, hydrochloric acid). 3.0 g of the sodium salt dissolves in 100 ml water at 25 °C. The substance is very sensitive to UV light [Yoneda (1984)]. 7.3.5 Spectra In the oxidized state (Flox), flavins and flavoproteins are yellow pigments3 with two typical absorption bands at about 370 and 450 nm. In solution, this chromophore gives rise to fluorescence with a maximum around 520 nm. If it is in a flavoprotein, the emission may be quenched [Ghisla (1980)]. In water, the green fluorescence of riboflavin has the same quantum yield as that of FMN. FAD fluoresces about 10 times less intensely, although the spectral distribution is the same in all these compounds [Visser & Müller (1980)]. Substitution on the ring can drastically change the spectrum. For example, roseoflavin (8-dimethylaminoriboflavin, p.409) has a strong maximum at 350 nm [Shiga et al. (1980)]. The absorption spectrum of FMN is shown in detail in Fig. 7-4. As would be expected, riboflavin has the same maxima [Moore et al. (1979), Shiga et al. (1979), Otto et al. (1981), Grasselli (1973)]. 3

Reviews : Massey & Ghisla (1974) ; Ghisla (1980)

414

7 Vitamin B2: Riboflavin and Its Bioactive Variants

300

400

500

Wavelength (nm)

Fig. 7-4. Absorption spectrum of FMN in water in the oxidized (Flox), reduced (Flred), neutral and anionic states. [Ghisla (1980), p. 360]

Temperature-difference spectra in the UV and visible have been described for flavins and flavoproteins [Müller & Mayhew (1980)]. The electronic spectra of the flavins are affected by hydrogen bonds. It is assumed that the spectral characteristics of many flavin enzymes are at least partly due to such Η-bonds [Yagi (1980)]. Reduced flavins and flavoproteins (FlredH2) have very atypical and variable UV and visible spectra above 300 nm. The sensitivity of reduced flavins to oxygen makes spectroscopy more difficult, but the spectra are especially relevant, due to the biological significance of reduced flavins. 1,5-Dihydroflavins have a rather featureless absorption spectrum (cf. Fig. 7-4), which is affected by temperature, pH and solvent. The 1,5-dihydroflavins do not fluoresce in solution, but when they are incorporated into a glass matrix, they have a pale bluish fluorescence around 500 nm. Fluorescence is also seen at very low temperatures [Ghisla (1980)]. The 13C NMR spectroscopy of reduced flavins has been reported [Van Schagen & Müller (1980)], as have Ή , 13C and M P NMR [Müller et al. (1980)] and resonance Raman [Benecky et al. (1979), Kitagawa et al. (1979), Nishina et al. (1980), Dutta et al. (1980), Bowman & Spiro (1981)] spectroscopy of various flavins and flavoproteins. Spectroscopy and photochemistry of the flavins and flavoproteins (Raman, bioluminescence, and chemical luminescence) have been reviewed in depth [Müller (1981)].

7.3 Nomenclature, Structure, Properties

415

7.3.6 Photochemistry The flavins, especially FMN and riboflavin, are sensitive to light4. This sensitivity is primarily a property of flavins not bound to proteins [ H e m m e r i c h (1976)]. Riboflavin and other isoalloxazines with various substituents at N-10 are bleached by light in 02-free, polar solutions. The changes occur mainly in the aliphatic side chain. The photochemical reactions begin with the intramolecular abstraction of hydrogen, and may be divided into two groups. In the reversible reactions, the carbon skeleton of the side chain is retained, although it is oxidized; in the irreversible reactions, at least part of the carbon skeleton of the side chain is split off, or the isoalloxazine ring is opened. The hydrogen is usually abstracted from the OH-carrying C atom of the side chain which is closest to the ring system. Completely reversible photoreduction of the riboflavin is also observed, chiefly in alcoholic solutions with labile hydrogens. Excited flavin can abstract the hydrogen from the alcohol and form the flavosemiquinone [Getoff et al. (1978)]. The photolysis of the side chain of the riboflavin was first observed by O. Warburg and W.Christian in 1933.In an alkaline medium, it produces lumiflavin (7,8,10-trimethylisoalloxazine, Fig. 7-5 a), and in a neutral medium, lumichrome (7,8-dimethylalloxazine, Fig. 7-5 b). The reaction is complex [Hemmerich et al. (1980)]. (α)

(b)

o

o

Fig. 7-5. Two photolysis products of riboflavin a. Lumiflavin b. Lumichrome Formulae from Yoneda [Yoneda (1984), p. 108]

Lumichrome was characterized and synthesized in 1934 by P. Karrer, and lumiflavin, by R.Kuhn in 1933. Another irradiation product of riboflavin is 7,8-dimethyl-10-formylmethylisoalloxazine [Yoneda (1984)]. Lumiflavin is produced by subjecting a solution of riboflavin in 0.5M NaOH to light for 3h.The solution is then neutralized and extracted with chloroform. The chloroform is evaporated, and the residue is dissolved in water and chromatographed on DEAE-cellulose; 0.5M sodium acetate is used for elution. Lumichrome is produced by irradiating riboflavin in water for lOOh.The irradiated solution is poured directly into the chromatography column and eluted with 0.9% NaCl [Suzuki et al. (1979)].

The application of laser technology to the analysis of the flavins has been described. Laser spectroscopy permits detection of many substances, by fluorescence, even at very low concentrations. The limits of detection for riboflavin are 4

Review: Hellis (1982)

416

7 Vitamin B2: Riboflavin and Its Bioactive Variants

1.0 χ 10 -10 M by conventional methods, and 1.25 χ 10~12M by laser fluorescence. Laser spectroscopy offers similar advantages in the detection of vitamin A and pyridoxine [Richardson (1980)]. Flavins stimulate the photooxidation of a number of biologically important molecules, e.g. amino acids, deoxyribonucleotides, retinol and ascorbate, presumably by a radical mechanism [Heelis et al. (1981)]. Riboflavin is also thought to act as a photoreceptor in plants, animals and microorganisms [Galston (1977)]. Light can cause considerable losses of riboflavin. In a few hours, sunlight destroys up to about 70% of the riboflavin in milk. Phototherapy of newborn infants for hyperbilirubinemia leads to riboflavin deficiency [Merrill et al. (1981 b)].

7.4 Chemical Synthesis In 1935, R. Kuhn and P. Karrer independently carried out the total synthesis of riboflavin5, and thus proved that its structure is 7,8-dimethyl-10-D-ribitylisoalloxazine. The above syntheses are based on the condensation of 6-D-ribitylamino3,4-xylidine with alloxan in acid solution (Fig. 7-6). Most commercial syntheses are based on the condensation of the appropriately substituted xylol and uracil rings [Yoneda (1984)]. H

ΟγΝ^Ο o· a-NADP + + A D P

(2)

If NADP is heated in aqueous solution at pH 8.4 and 100°C, 2'P-ADP is produced (Eq.3): NADP — 2T-ADP+nicotinamide +

ribose

(3)

2'P-ADPR may be an intermediate in this reaction. 2'P-ADP was originally thought to be an isomer of ATP, from which it could not be separated. It can be differentiated from ATP by means of the luminescence test with firefly luciferase [Bernofsky (1980 d)]. In the presence of 0 2 , NADH is sensitive to light. Photochemically produced singlet oxygen OO2) reacts with NADH to form NADH peroxide, 1-HOO-NADH+ (Eq.4, Fig.8-3) [Bernofsky & Wanda (1982a,b)]: NADH + 1 0 2 + H + — P-t-H0 2 -NADH +

(4)

The most-used method for chemical synthesis of nicotinic acid is based on the oxidation of the corresponding substituted pyridine; 2-methyl-5-ethylpyridine is

8.5 Chemistry

H H

NADH + 'O, + Η δ

485

CONH,

ho: HCOH HCOH I HC 0 I ch 2 or

Fig. 8-3. Reaction of N A D H with photochemically generated singlet oxygen 1C>2 to form 1-HOO-NADH+; R=5'-adenosine diphosphate [from Bernofsky & Wanda (1982b)]

preferred as starting material because of its relatively low price. Nitric acid or air is the oxidizing agent; in this case both substituents are oxidized to carboxyl groups, and decarboxylation at position 2 of the pyridine ring produces nicotinic acid. The carboxylation of pyridine is another feasible method. The old method of oxidation of nicotine with nitric acid is also used sometimes [Offermanns et al. (1984), Hankes (1984)]. For the chemical synthesis of nicotinamide, one can start with aliphatic compounds like ethylene, acetaldehyde or acrolein together with ammonia. The intermediates are 3-methylpyridine, 2-methyl-5-ethylpyridine, nicotinic acid and its nitrile. Alkaline hydrolysis of the nitrile is one of the commercial methods for production of nicotinamide [Offermanns et al. (1984)]. Nicotinamide is also produced commercially by ammonolysis of nicotinic acid esters. Methyl nicotinate, which is commercially available, is autoclaved in the presence of gaseous ammonia to form nicotinamide (200-270°C, high pressure, yield 96-99% [Hankes (1984)]. Nicotinic acid and nicotinamide are produced by chemical synthesis in numerous pharmaceutical factories. The registered countries (excluding the USSR) produce about 22,000 tons per year [Offermanns et al. (1984)]. Nicotinamide riboside can be synthesized from NAD + . One first applies snake venom phosphodiesterase, and separates nicotinamide mononucleotide from 5'-AMP by column chromatography on Dowex-l-formate. The nicotinamide mononucleotide is then cleaved to nicotinamide riboside using a monoesterase [Kasarov & Moat (1980)]. There are two other ways to produce nicotinamide mononucleotide: 1.Cleavage of NAD(P) by acid hydrolysis or NAD pyrophosphatase (isolated, e.g. from potatoes); the NAD pyrophosphatase can be bound to a carrier such as Sepharose. 2. Total synthesis, starting from nicotinamide, ribose and phosphate. This total synthesis has already been worked out by several groups [Jeck & Woenckhaus (1980)]. A chemical/enzymatic synthesis of NAD + has been reported. Nicotinamide mononucleotide (NMN) is produced in a series of reactions, starting from ribose 5-phosphate. The NMN is coupled to AMP, using the enzyme NAD pyrophosphorylase (EC 2.7.7.1) in the presence of ATP. NAD can be converted to NADP by NAD kinase (EC 2.7.1.23) (cf. also p. 488) [Walt et al. (1980,1984)]. X-ray analyses of the structures of NAD and its complexes with Li + or malate dehydrogenase have been reported [Parthasarathy & Fridey (1984)].

486

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P)

In addition, an enzyme-catalysed synthesis of deuterated nicotinamide cofactors [Wong & Whitesides (1983)] and a synthesis of polymer-bound NADP derivatives and their coenzyme activities [Araki et al. (1984)] have been reported.

8.6 Physical Properties Nicotinic acid and nicotinamide are colorless, crystallizable substances. Nicotinic acid is slightly soluble in water (about 1 g/60 ml) and alcohol (about 1 g/100 ml), but it dissolves easily in alkaline solutions. It melts with sublimation at 236-237 °C. Nicotinamide is very soluble in water (about 1 g/ml) and reasonably so in alcohol; melting point 129-132°C; both substances are either insoluble or only very slightly so in ether and chloroform. Nicotinic acid and nicotinamide have similar absorption spectra in water, with an absorption maximum around 262 nm; the extinctions are pH dependent (Table 8-3). The UV spectrum of NAD(P)H has two maxima (Fig. 8-4). One, at 260 nm, is given by the adenine ring, and the other, at 340 nm, is due to the nicotinamide ring. Various wavelengths are used for quantitative measurements, depending on the available source of light (Table 8-4). NAD(P)H fluoresces around 470 nm Table 8-3. Physical properties of nicotinic acid and nicotinamide [Offermanns et al. (1984), Vitamin Compendium (1980)] Properties

Measured values

Rei. molar mass (daltons) Melting point (°C)

Nicotinic acid 123.11 236-237 (Sublimation)

Isoelectric point in water, 25 °C Solubility (g/100 ml) Water, room temperature Water, 38 °C Ethanol, 95%, room temp. Ethanol, 95%, 78 °C Methanol, 62 °C Glycerol Acetone, chloroform Ether

Nicotinamide 122.12

129-132 (stable modification) 105-106 (unstable modification)

pH 3.42

1.6

100.0

2.47 1.0

Easily soluble

7.6 34.5 Practically insoluble Practically insoluble

Soluble Slightly soluble Practically insoluble

Crystallinity

White crystalline powder

Absorption maximum in water

Emax about 262 nm

8.7 Biosynthesis and Metabolism of Niacin

260

300

340

487

380 nm

Wavelength

Fig. 8-4. Absorption spectrum of the nicotinamide nucleotides. The reduced form has a maximum at 340 nm [from Karlson (1984 a)]

when excited with light of either 260 nm or 340 nm wavelength. When the adenine portion is excited, the fluorescence is the result of an energy transfer to the nicotinamide part. The spectra (UV, fluorescence, CD and NMR) are relatively strongly dependent on the temperature, which is associated with changes in conformation [Malcolm (1980)]. The absorption spectrum of NADH shows a slight red shift (about 1%) at high hydrostatic pressure (2 kbar), and a slight decrease in absorption at 339 nm [Jaenicke et al. (1981)]. Table 8-4. (1980)]

Extinction coefficients of NADH at various wavelengths and 25 °C [Malcolm,

Wavelength (nm)

334

340

366

E x l O " 3 (M-'cm- 1 )

6.0

6.16-6.29

3.30

8.7 Biosynthesis and Metabolism of Niacin The biosynthesis de novo of niacin4 (Fig. 8-5) goes via quinolinic acid in all organisms which have so far been studied. Some microorganisms synthesize quinolinic acid from asparagine and a C3 compound which has been identified as dihydroxyacetone phosphate or glyceraldehyde phosphate; the exact biosynthetic route is still open. In addition to the 4

Reviews : Foster & Moat (1980), Gopalan & Rao (1975)

488

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P) Η,Ν 0 2 1 II CH2CHCOH

\

\

Tryptophan

rr

0 II COH

0 II COH

li 0

0 Η,Ν 0 II I II ^CCH2CHCOH

OH

AC MSA

OH

3-OH-Ant

v

3-OH-Ky

0 » ^COH

0 C,

@C

I

"N" "COH

11

COH +

X Ν"TOH

O

O

QA

Asp

QA -PRPP C02+PPr ATP

PRPP

NA

-

-NH3

V

-NARP

PP¡

PPi ATP

PRPP Me-NAm-2-NAm ·

\ 2

- V -

NMN

PPi

k

Y "

- NAAD -

Gln+ATP

•

Glu+AMP+PPj

NAD-^NADP

PP;

Γ> ADPR

Fig. 8-5. Biosynthesis de novo and metabolism of NAD(P). This biosynthesis always goes via quinolinic acid (QA). In some microorganisms, dihydroxyacetone phosphate or glyceraldehyde phosphate serves as quinolinic acid precursors; these C3 bodies supply the C-4, C-5 and C-6 of the quinolinic acid; aspartic acid (Asp) is the source of the remaining atoms of the quinolinic acid. In animals and some microorganisms, and in at least some plants, tryptophan is the quinolinic acid precursor. The most important intermediates are 3-hydroxykynurenine (3-OH-Ky), 3-hydroxyanthranilic acid (3-OH-Ant) and α-amino-ß-carboxymuconic semialdehyde (ACMSA). The biosynthesis of NADP, starting from nicotinamide (NAm), proceeds in bacteria and yeasts via reactions 7 , 1 , 2, 4 and 10, and in mammals via reactions 5, 6 and 10. Reactions 1, 2 and 4 constitute the Preiss-Handler pathway. Some of the important metabolic products are adenosine diphosphate ribose (ADPR), 1-methylnicotinamide, l-methyl-6-pyridone 3-carboxamide and l-methyl-4-pyridone 3-carboxamide. Literature sources: Henderson (1983), McCreanor & Bender (1983) Bender et al. (1982), Liu et al. (1982), Sakai (1980), Kinney et al. (1979), Gopalan & Rao (1975), Arditti & Tarr (1979), Tarr & Arditti (1982), Magboul & Bender (1983), Offermanns et al. (1984)

8.7 Biosynthesis and Metabolism of Niacin

489

asparagine route, there is a pathway to quinolinic acid which starts from tryptophan; this is active in animals [Arditti & Tarr (1979); Kinney et al. (1979); Gopalan & Rao (1975); Tarr & Arditti (1982); Henderson (1983); Offermanns et al. (1984); Cooper et al. (1982)] (Fig. 8-5). E. coli, Mycobacterium tuberculosis, Salmonella typhimurium and many other microorganisms synthesize niacin via aspartic acid [Tarr & Arditti (1982); Cooper et al. (1982); Kinney et al. (1979)]. This is the only route for synthesis of niacin in some bacteria, fungi and plant seeds [Arditti & Tarr (1979), Tarr & Arditti (1982)], maize seedlings and orchids [Tarr & Arditti (1982) Cooper et al. (1982)], and in birds and mammals [Arditti & Tarr (1979), Tarr & Arditti (1982)]. Algae form niacin by more than one route; in some, tryptophan serves as a precursor. Saccharomyces cerevisiae forms niacin under anaerobic conditions via aspartate and glyceraldehyde 3-phosphate, but under aerobic conditions, via

Symbols, abbreviations ACMSA ADPR Asp C3

α-Amino-ß-carboxymuconate semialdehyde Adenosine diphosphate ribose Aspartic acid as donor of C 4 and Ν of quinolinic acid Three-carbon unit of quinolinic acid, donated by dihydroxyacetone phosphate or glyceraldehyde phosphate Gin Glutamine Glu Glutamic acid Me-NAm 1-Methylnicotinamide NA Nicotinic acid NAAD Nicotinic acid adenine dinucleotide NAD Nicotinamide adenine dinucleotide NADP Nicotinamide adenine dinucleotide phosphate NAm Nicotinamide NARP Nicotinic acid ribonucleotide NMN Nicotinamide ribonucleotide 3-OH-Ant 3-Hydroxyanthranilic acid 3-OH-Ky 3-Hydroxykynurenine PPi Pyrophosphate PRPP 5-Phosphoribosyl 1-diphosphate QA Quinolinic acid Enzymes 1 =Nicotinate phosphoribosyltransferase (EC 2.4.2.11) 2 = Deamido-NAD pyrophosphorylase (EC 2.7.7.18) 3 = Quinolinate phosphoribosyltransferase (EC 2.4.2.19) 4 = NAD synthetase (EC 6.3.5.1) 5 = Nicotinamide phosphoribosyltransferase (EC 2.4.2.12) 6 = NAD pyrophosphorylase (EC 2.7.7.1) 7 = Nicotinamide deamidase (EC 3.5.1.19) 8 = NAD(P) + glycohydrolase (EC 3.2.2.5 and 3.2.2.6) 9=Nicotinamide methylase (EC 2.1.1.1) 10=NAD kinase (EC 2.7.1.23) 11 = 1-Methylnicotinamide oxidases (formation of l-methyl-4(6)-pyridone 3-carboxamide 12 = Kynureninase (EC 3.7.1.3)

490

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P)

tryptophan (Fig. 8-5). Thus it is possible for two different biosynthetic pathways for niacin to be present in one organism [Arditti & Tarr (1979)]. Animals and bacteria convert quinolinic acid to nicotinic acid ribonucleotide (NARP) by transfer of phosphoribosyl from 5-phosphoribosyl 1-diphosphate, with loss of C0 2 (Fig. 8-5). Some results indicate that the reaction catalysed by quinolinate phosphoribosyltransferase is the rate-limiting step for the synthesis in the rat [Henderson (1983)]. It is interesting that quinolinic acid, when administered orally or intraperitoneally, is a relatively ineffective precursor for pyridine nucleotides in rats [Henderson & Gross (1979 a)]. Two pathways for the biosynthesis of NAD(P), starting from nicotinamide (NAm) are known. One (Fig. 8-5, reactions 5 and 6) leads directly to NAD via nicotinamide ribonucleotide (NMN) as an intermediate. In the second pathway (Fig. 8-5, reactions 7,1, 2,4), NAm is first deamidated to nicotinic acid (NA), and the latter is converted to the dinucleotide (NAAD) via NARP; amidation (Reaction 4) leads to NAD (Fig. 8-5) [Schuette & Rose (1983)]. The nicotinamide deamidase activity of the animal cells is very slight. In contrast, intestinal bacteria are able to deamidate nicotinamide effectively, and also to amidate nicotinic acid [Offermanns et al. (1984)]. The metabolism of the niacin coenzymes includes the release of nicotinamide by the action of NAD(P) + glycohydrolase, followed by reincorporation into the coenzyme, deamidation to nicotinic acid, or 1-methylation (Fig. 8-5). The most important excretion products of niacin in the urine of mammals are 1-methylnicotinamide and its pyridones. The regulation of the level of NAD(P) probably occurs primarily via hydrolysis of these dinucleotides to nicotinamide (Fig. 8-5) [Nutrition Rev. 42 (1984) 62]. 8.7.1 Niacin Biosynthesis in Mammals and Birds 8.7.1.1 Tryptophan as Niacin Precursor and Regulation of the Tryptophan Metabolism Dietary tryptophan can be converted enzymatically to NAD(P) in the animal body. This conversion is probably regulated by the enzyme quinolinate phosphoribosyltransferase, which is the key enzyme in the metabolism of tryptophan via quinolinic acid [Satyanarayana & Rao (1983)] (Fig. 8-5). The tryptophan route is not very efficient, and in human beings, 60 mg tryptophan yields only 1 mg niacin. However, this ratio does not hold under conditions of dietary niacin deficiency, low tryptophan intake or pregnancy [Offermanns et al. (1984)]. Human beings utilize about 2.8% of their dietary tryptophan for the biosynthesis of niacin [Tarr & Arditti (1982)]. Niacin synthesized by this route can supply up to about two thirds of the human requirement. Pyridoxine deficiency reduces the conversion of tryptophan to niacin [Bausch et al. (1980)]. Some animals make more efficient use of tryptophan as a niacin source than human beings do; the niacin equivalent in birds is 1/45, and in the rat, 1/50. The synthesis of NAD from tryptophan in the animal liver does not go via nicotinic acid or nicotinamide. Nicotinamide formed by degradation of NAD is, however, very efficient-

8.7 Biosynthesis and Metabolism of Niacin

491

ly reconverted to NAD via nicotinamide ribonucleotide (NMN) (Fig. 8-5, Reactions 5, 6). Recycling of the nicotinamide via nicotinic acid (NA) (Preiss-Handler route, Fig. 8-5, Reactions 1, 2, 4) is normally not very significant in the mammal, but at very high doses of nicotinamide, it is used preferentially. Human tissue cells contain little nicotinamide deamidase, to be sure, but nicotinamide can be deamidated in the intestinal tract by enzymes from the microflora [Offermanns et al. (1984)]. Some observations suggest that leucine inhibits the tryptophan-to-NAD metabolic pathway, at a step which comes before 3-hydroxyanthranilic acid [Shin et al. (1982)]. Similar conclusions were drawn from experiments on rats which showed that an excess of leucine in the feed, together with an inadequate supply of niacin and/or tryptophan, produces pellagra. It is thought probable that leucine inhibits tryptophan metabolism at the kynureninase level (Fig. 8-5) [Magboul & Bender (1983)]. Other experiments showed that an addition of 5% leucine to the feed of rats elevated the activity of the hepatic NAD(P) + glycohydrolase, and thus decreased the NAD level. It was also observed, on isolated rat liver cells, that 2-oxoisocaproate, the 2-oxo analog of leucine, as well as other ketone bodies, decrease NAD biosynthesis from both tryptophan and nicotinic acid [Yamada et al. (1979,1983)]. Tryptophan promotes both growth and development in chicks. Addition of 4.8% leucine to the diet has no effect. The results show that leucine does not impair tryptophan metabolism via quinolinic acid in chicks [Penz et al. (1984)]. 8.7.1.2 NAD Biosynthesis from Nicotinic Acid and Nicotinamide In mammals, the biosynthesis of the pyridine nucleotides (Fig. 8-5) and the regulation of their intracellular concentrations are complicated, because there are three precursors, tryptophan, nicotinic acid and nicotinamide. Although each organ and tissue apparently is able to form its own pyridine nucleotides, there is an exchange among the tissues, primarily at the level of nicotinamide. In the rat, nicotinic acid is the most important coenzyme precursor in the liver, kidneys, brain and erythrocytes, but nicotinamide appears to be a better precursor in the testes and ovaries. The liver rapidly converts nicotinate to the amide via the synthesis of NAD by the Preiss-Handler pathway (Fig. 8-5), followed by the action of NAD(P) + glycohydrolase. The nicotinamide is transported into other tissues [Henderson & Gross (1979 a)]. [14C]Nicotinamide injected into the portal vein of the mouse reaches the gastrointestinal tract via the liver. In the gut, it is deamidated to form nicotinic acid. This is reabsorbed and converted to NAD in the liver [Arditti & Tarr (1979)] (Fig. 8-5). The nicotinamide can be converted to NAD via nicotinamide ribonucleotide (NMN) (Fig. 8-5, Reactions 5, 6). 8.7.1.3 Niacin Biosynthesis in the Ungulates Rumen microorganisms form niacin, and this synthesis is normally sufficient for the optimal development of the animal. It has been found, however, that this is not the case for animals subjected to the stress of production. In cattle, for example, a niacin supplement (50-250 ppm) increased the rate of growth and milk

492

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P)

production. Furthermore, niacin stimulated the fermentation and protein formation in the rumen. Rumen bacteria form niacin from tryptophan [Brent & Bartley (1984)]. 8.7.2 Niacin Biosynthesis in Higher Plants and Microorganisms The data on niacin biosynthesis in the higher plants are rather sparse [Arditti & Tarr (1979)]. The studies on maize and orchid seedlings indicated that here the synthesis proceeds via tryptophan, as in animals and some microorganisms, and that aspartic acid is not a precursor of niacin in these plants [Tarr & Arditti (1982), Cooper et al. (1982)]. Escherichia coli and Salmonella typhimurium synthesize NAD(P) de novo, starting from dihydroxyacetone phosphate and aspartate. Furthermore, they can start from nicotinamide, and they can also use nicotinic acid (Fig. 8-5, Reactions 7,1, 2, 4) [Liu et al. (1982)]. In most microorganisms, the first step of the biosynthesis of NAD, starting from nicotinamide, is the deamidation to nicotinic acid and ammonia by nicotinamide deamidase. This is followed by conversion to nicotinic acid ribonucleotide, nicotinic acid adenine dinucleotide, and NAD [Tanigawa et al. (1980), Kinney et al. (1979)] (Fig. 8-5). Trigonellin (1-methylnicotinic acid), which is found in many plants, is thought to be a storage form of nicotinic acid. A conversion of nicotinic acid and nicotinamide to trigonellin in the plant has been demonstrated [Arditti & Tarr (1979)]. 8.7.3 On the Biosynthesis of NAD(P) and Control of the NAD(P) Level NAD and NADP are formed from several precursors, including tryptophan, in animal tissue (Fig. 8-5). In the pathway starting from nicotinamide, nicotinic acid is normally not an intermediate in NAD synthesis. This is apparently only the case when the nicotinamide level is much higher than physiological, and in this case, the nicotinamide deamidase of the intestinal bacteria comes into play. The conversion of nicotinamide to NAD is normally controlled very precisely by the activity of the nicotinamide deamidase, the nicotinamide phosphoribosyltransferase and the nicotinamide methylase [Bender et al. (1982)]. It was shown in cultured liver sections from adult rats that nicotinamide serves in equal measure as precursor for 1-methylnicotinamide and for NAD, if the nicotinamide in the medium is present in physiological concentration (11500 μΜ). After the nicotinamide level has been elevated to 1 mM or more, the formation of 1-methylnicotinamide reaches a plateau, and the rate of NAD synthesis is dramatically increased. This increase rather exactly parallels the elevation of the intracellular nicotinamide pool, which suggests that the intracellular nicotinamide concentration is rate-determining for NAD synthesis. From various experiments, it is concluded that 1-methylnicotinamide is an inhibitor of NAD synthesis from nicotinamide. The toxicity of the nicotinamide is probably due to the fact that its conversion to NAD consumes much cellular ATP. Simultaneous application of 1-methylnicotinamide and nicotinamide at least partly inhibits the elevation of NAD level and loss of ATP [Hoshino et al. (1984)]. There are many

8.7 Biosynthesis and Metabolism of Niacin

493

observations which suggest that 1-methylnicotinamide is involved in the control of hepatocellular DNA synthesis and proliferation [Hoshino et al. (1982 a,b)]. It is suspected that the nicotinamide level in the plasma is regulated by hormonally controlled release of nicotinamide from the "storage NAD" in the liver. The "storage NAD" is probably not in equilibrium with the "functional NAD", which is involved in redox processes. The sharp increase in the hepatic NAD concentration in response to elevated nicotinamide intake is ascribed mainly to an increase in the "storage NAD" [Henderson (1983)]. The NAD metabolism in cultivated brush-border membranes from rat kidneys has been described [Angielski et al. (1982)]. Hydrolysis has an essential role in the control of the NAD(P) levels in animal organs. The microsomal NAD(P) + glycohydrolase cleaves NAD(P) to ADPR and nicotinamide (Fig. 8-5) [McCreanor & Bender (1983)]. Extracellular glycohydrolases in the synaptosomes of rat brain quickly degrade NAD, which serves here as modulator of the synaptic activity [Snell et al. (1984)]. In patients with Fanconi's anemia, there was a reduced activity of the NAD(P) + glycohydrolase in the repair of UV-irradiated DNA [Klocker et al. (1983)]. NAD kinase and ATP are required for the synthesis of NADP from NAD (Fig. 8-5). Nicotinic acid adenine dinucleotide phosphate, NAADP + , can be formed in a base-exchange reaction, starting from NADP + , nicotinic acid and NAD(P) + glycohydrolase (EC 3.2.2.6) from bovine spleen (Eq.5): NADP + + Nicotinic Acid — NAADP + + Nicotinamide

(5)

+

NAD reacts analogously [Bernofsky (1980c)]. The half-life of NAD in E. coli and in the animal cell is 1-2h. About 95% of the NAD synthesized in mammals is consumed in catabolic reactions in the nucleus, and only about 5% is used to maintain the NAD coenzyme pool [Brown et al. (1979)]. NADPH is formed mainly in the pentose phosphate cycle. In rat liver, this cycle is probably controlled mainly by the intracellular NADPH/NADP ratio [Fabregat et al. (1985)]. NAD is obtained from yeast and a number of other microorganisms. With S. cerevisiae, fermentation yields about 1 g NAD/kg dry cells, but with Saccharomyces carlsbergensis, up to 4.2 g NAD/kg dry cells may be obtained [Walt et al. (1984)]. NADP was first obtained in larger quantities by extraction from microbial cells, but later it became the practice to phosphorylate NAD to NADP using NAD kinase (cf. Fig. 8-5 and Eq.6): NAD + ATP — NADP 4- AD Ρ

(6)

It was then found to be more favorable to immobilize bacteria with high NAD kinase activities. A number of modifications of this method have been introduced, which make use of immobilized enzymes and in which the ATP is generated in situ [Miyawaki et al. (1982)]. 8.7.4 Bacterial Degradation of Nicotinic Acid Nicotinic acid is degraded anaerobically by Clostridium barkeri, forming stoichiometric amounts of acetate, propionate, carbon dioxide and ammonia. Vitamin B12 is involved in this degradation (Fig. 8-6) [Kollmann-Koch & Eggerer (1984)].

494

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P) ,C0 2 H

,CO2H

xy

CT Ν

O^N' Η

Vitamin B12 ΗΟ2Ο'

'CO 2 H

HO 2 C'

¿H3

+ H2O H,C Η J

| I

HO2C-C-C-CO2H

H 3 CCH 2 CO 2 H + H 3 CCOCO 2 H

Fig. 8-6. Degradation of nicotinic acid by Clostridium barken [from Kollmann-Koch & Eggerer (1984)] Hydroxylation and partial reduction of nicotinic acid yields 6-oxotetrahydronicotinic acid. Hydrolytic ring cleavage and deamination lead to 2-methyleneghitarate. A vitamin B12-dependent enzymatic conversion and an isomerization then lead to 2,3-dimethylmaleate, which is further degraded to propionate and pyruvate. The microorganism obtains the energy required for its growth from the conversion of pyruvate to acetate, which yields 1 molecule ATP [Pirzer et al. (1979), Kollmann-Koch & Eggerer (1984)]. The first enzyme in the above reaction chain is nicotinic acid hydroxylase. It is a flavoprotein (300,000 daltons) containing iron, sulfur, molybdenum, selenium and a pterin cofactor [Dilworth (1983)]. Some properties of the selenium-containing part of the nicotinic acid hydroxylase from C. barkeri have been described [Dilworth (1982)].

The bacteria Pseudomonas aeruginosa and Serratia marcescens are able to hydroxylate nicotinic acid at position 6, but E. coli and Klebsiella pneumoniae are not able to. This ability can be used to identify P. aeruginosa and S. marcescens in the clinic [Shiraishi et al. (1985)]. 8.7.5 Important Enzymes and Enzymatic Reactions Nicotinate phosphoribosyltransferase (EC 2.4.2.11) and nicotinamide phosphoribosyltransferase (EC 2.4.2.12) are important for the biosynthesis of NAD, starting from nicotinic acid (Preiss-Handler pathway) or from nicotinamide (nicotinamide pathway) (cf. Fig. 8-5). A rapid assay for nicotinamide phosphoribosyltransferase has been developed [Elliott et al. (1980)]. Quinolinate phosphoribosyltransferase (EC 2.4.2.19) catalyses the biosynthesis of nicotinic acid ribonucleotide, starting from quinolinic acid. The reaction takes place in the liver according to Eq. 7 : Quinolinate + PRPP

N A R P + C 0 2 + PP¡

(7)

The activity of the enzyme can be determined by measuring the 1 4 C0 2 which is released in stoichiometric quantity from the a-carboxyl of radiolabeled quinolin-

8.7 Biosynthesis and Metabolism of Niacin

495

ic acid [Iwai & Taguchi (1980)]. The reaction of Eq. 7 also takes place in bacteria [Kinney et al. (1979)]. Quinolinate phosphoribosyltransferase is essential for the synthesis of NAD de novo, both in eukaryotes and in prokaryotes. Unphysiologically high concentrations of 0 2 damage the enzyme, and thus decrease the pyridine nucleotide level. The situation can then be corrected by nicotinic acid, but not by quinolinic acid. Nicotinic acid could therefore be significant in human medicine, if patients must be treated with potentially toxic concentrations of 0 2 [Brown & Song (1980)]. Deamido-NAD pyrophosphorylase (EC 2.7.7.18) and NAD pyrophosphorylase (EC 2.7.7.1) catalyse the reaction of mononucleotides (NARP, NMN) with ATP to form dinucleotides (NAAD, NAD) and PP¡ (cf. Fig. 8-5, Reaction 2 and 6). NAD synthetase (EC 6.3.5.1) catalyses the amidation of NAAD to NAD in a reaction which involves ATP. Glutamine serves as the donor of the amino group (cf. Fig. 8-5, Reaction 4). Nicotinamide deamidase, nicotinamide amidohydrolase (EC 3.5.1.19) is always present in the microsomal fractions of rat and rabbit tissues. It is present in relatively low amounts in the central nervous system. Its presence may be correlated with the synthesis of NAD [Wintzerith et al. (1979)]. The enzyme (230,000 daltons) consists of subunits of 65,000 and 50,000 daltons [Wintzerith et al. (1980)]. The nicotinamide deamidase is very active in tissue homogenates, but in vivo, in animal tissues, it is practically inactive [Henderson (1983)] (cf. Fig. 8-5, Reaction?). NAD(P)+ glycohydrolase (EC 3.2.2.6), which occurs widely in microorganisms and mammals, splits NAD + at the nicotinamide-ribose bond. NAD contains two ß-glycosidic bonds, both of which are available for hydrolytic cleavage. Generally the bond at the nicotinamide residue is cleaved, although Aspergillus niger has the ability to split NAD at the adenine-ribose bond, forming nicotinamide ribose diphosphate ribose (NAmRDPR) and adenine. In this case, a purine nucleosidase is involved in the cleavage of the NAD [Kuwahara (1980)]. The microbial enzymes generally catalyse an irreversible hydrolysis of NAD + . Mammalian enzymes, by contrast, can catalyse the exchange of the nicotinamide part of the NAD for free nicotinamide, or for another pyridine base [Everse et al. (1980)]. It has been suggested that the increase in activity in the NAD(P) + glycohydrolase (EC 3.2.2.6) could be used as an enzymatic indicator of the malignancy of Burkitt's tumor cells [Skala et al. (1982)]. The NAD(P) + glycohydrolase from Neurospora crassa was the first to be studied [Jorge & Terenzi (1984)]. An NAD+ glycohydrolase (about 62,000 daltons) from the inner mitochondrial membrane of rat liver has also been studied. This enzyme has only a little activity with respect to NADP + , and is inhibited by ATP and nicotinamide [Moser et al. (1983)]. The mammalian NAD(P) + glycohydrolase is a membrane-bound enzyme. The highest activity has been found in the microsomal fraction of various tissues. Some workers have reported that the hepatic NAD(P) + glycohydrolase is part of the endoplasmic reticulum, while others found the enzyme in the plasma membrane, the nucleus and the lysosomes. One of the physiological roles of the enzyme may be the control of the intracellular NAD level [Muller et al. (1983)]. Nicotinamide methylase (EC 2.1.1.1) is located mainly in the liver in adult rats.

496

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P)

Fetal liver contains only 2% of the nicotinamide methylase activity found in adult liver. This enzyme catalyses the methylation of nicotinamide to 1-methylnicotinamide, which is excreted in the urine, as are its oxidation products, 1-methyl-6-pyridone 3-carboxamide and l-methyl-4-pyridone 3-carboxamide [Seifert et al. (1984)]. (The structural formulae of the niacin metabolites are given in Fig. 8-1.) Rat liver tissue in which cell division is more frequent than normal, e.g. after partial hepatectomy, produces more nicotinamide methylase than normal. Substances like thioacetamide, which stimulate proliferation of the hepatocytes, have a similar effect. This increase in activity is associated with a considerable drop in the NAD level and a several-fold increase in the rate of synthesis of the 1-methylnicotinamide. It is assumed that nicotinamide methylase and its product, 1-methylnicotinamide, are involved in the control of hepatocyte DNA synthesis and proliferation. 1-Methylnicotinamide decreases the hepatic NAD level by inhibiting the synthesis of NAD from nicotinamide, or by stimulating the microsomal NAD(P) + glycohydrolase [Hoshino et al. (1982 b)]. Two 1-methylnicotinamide oxidases, both of about 150,000 daltons, have been isolated from rat liver. These proteins differ immunologically and with regard to their heat stability and pH optima. They catalyse the conversion of 1-methylnicotinamide to l-methyl-6-pyridone 3-carboxamide and l-methyl-4-pyridone 3-carboxamide. The physiological role of these oxidases is not known [Ohkubo et al. (1983)]. NAD kinase (EC 2.7.1.23) catalyses the reaction of Eq.8: NAD + + ATP — NADP+ + ADP

(8) +

Their activity can be measured fluorimetrically, after reduction of the NADP by glucose 6-phosphate in the presence of glucose 6-phosphate dehydrogenase [Blomquist (1980)]. The NAD kinase activity in rat uterus is stimulated by progesterone, probably by regulation of gene expression and protein synthesis [Cummings & Yochim (1983)]. a-Amino-§-carboxymuconate semialdehyde decarboxylase (picolinate carboxylase, EC 4.1.1.45) is one of the most important regulators of the conversion of tryptophan to nicotinic acid in the rat. An increase in the activity of this enzyme, for example in response to a hormone, reduces the yield of quinolinate and nicotinate [Sanada & Miyazaki (1980), Magboul & Bender (1983)]. Nucleotide pyrophosphatase (EC 3.6.1.9) cleaves the pyrophosphate bond of NAD. Experiments with enzymes from potatoes, the venom of Crotalus adamanteus, Neurospora crassa and sheep liver have been reported [Christ, Coper (1980)]. Nicotinamide N-oxide is formed enzymatically in the microsomes of rat liver [Nomura et al. (1983)]. It can be reduced to nicotinamide in vivo; the reduction is catalysed by the aldehyde oxidase (EC 1.2.3.1) of the liver [Kitamura & Tatsumi (1984)]. Kynureninase (EC 3.7.1.3) is one of the enzymes which regulates tryptophan degradation. This enzyme is probably inhibited by excess leucine in the diet, and this effect may be one of the causes of the pellagrogenic activity of leucine [Magboul & Bender (1983)].

8.7 Biosynthesis and Metabolism of Niacin

497

0

II

Ν CH2-0-®~ ©-0-I HO\|

/oh

Ό' OH

OH

+

Fig. 8-7. Structure formula of NAD showing the site of the high-energy bonds. The bond between the nicotinamide and adenosine diphosphate ribose (ADPR) parts is cleaved by NAD(P) + glycohydrolases [from Hilz (1981)]

8.7.6 NAD(P) + Glycohydrolases and ADP-Ribosylation The ß-nicotinamide-ribose bond in NAD(P) + is a high-energy bond (Fig. 8-7). The ADPR group thus has a high transfer potential, and there is a group of enzymes which make use of this5. As early as 1966, a nuclear enzyme was discovered which polymerizes the ADPR part of NAD + . The bonding is 0-glycosidic, and occurs between the released C-l of the ribose and the C-2' of the adenosine part of another ADPR molecule [Hilz (1981)]. In 1967, the transfer of the ADPR group to macromolecules was observed [Henderson (1983)]. 8.7.6.1 ADPR Transfer to Water and Small-Molecule Bases ("Base Exchange") Enzymes which cleave the nicotinamide-ribose bond in NAD(P) + are frequently (and not quite accurately) called general NAD(P) + glycohydrolases. However, this term applies primarily to enzymes which catalyse the hydrolytic cleavage with release of ADP-ribose and nicotinamide (NAm) according to Eq.9: ADPR-NAm + + H 2 0 — ADPR + NAm + Η +

(9)

This is a matter of transferring the ADPR group to water [Henderson (1983)]. Some NAD(P) + glycohydrolases catalyse the exchange of other bases for nicotinamide, which produces NAD analogs (Eq.10): ADPR-NAm + + Base — ADPR-Base + + NAm

(10)

+

In rat liver, there is an NAD(P) glycohydrolase localized in the plasma membranes of the Kupffer cells [Amar-Costesec et al. (1985)]. 8.7.6.2 Mono-ADPR Transfer to Macromolecules It was observed that some NAD(P) + glycohydrolases have a transglycosidase activity which is characterized by the transfer of mono-ADPR to macromolecules. This ADP-ribosylation occurs according to E q . l l : 5

Reviews: Hilz (1981), Hayasishi & Ueda (1982), Henderson (1983), Vaughan & Moss (1983), Wold & Moldawe (1984), Althaus, Hilz & Shall (1985)].

498

8 Niacin: Nicotinic Acid, Nicotinamide, NAD(P) ADPR-NAm + 4- M —• ADPR-M + NAm + H +

(11)

[Henderson (1983)]. The macromolecule (M) appears usually to be a protein, so that Eq.ll applies mostly to the synthesis of mono-ADPR-proteins. The monoADPR transferases are classified in three groups: 1) Bacteriophage enzymes, 2) Enzymes of animal cells, and 3) Bacterial toxins which act on animal cells by catalysing the ADP-ribosylation of specific cellular proteins [Vaughan & Moss (1983)]. Protein-bound ADPR groups have been found in all eukaryotic tissues examined for them. Usually they are single ADPR groups ("mono(ADPR) groups"), but more rarely they are polymeric (ADPR)n chains. The ADPR transferases of the prokaryotes transfer only single ADPR groups to proteins. Studies of the subcellular distribution indicated that more than 95% of the mono(ADPR) conjugates in rat liver are found in extranuclear compartments [Hilz (1981)]. Many well studied examples of mono-ADP-ribosylations catalysed by bacterial toxins are available. Diptheria toxin is one of the best studied ADPR transferases; it inhibits protein synthesis in HeLa cells in the presence of NAD by ADP-ribosylation of some elongation factors (EF). The activity of cholera toxin is another example of mono-ADP-ribosylation by bacteria; here a monomer is bound to the membrane protein which regulates adenylate cyclase. Cholera toxin stimulates the cyclase activity, probably by ADP-ribosylation of an arginine residue [Henderson (1983)]. Cholera toxin also catalyses the ADP-ribosylation of tubulin [Hawkins & Browning (1982)]. Tubulin is a protein component of the microtubuli, which are responsible for motions within the cell. The ADP-ribosylation catalysed by diptheria and cholera toxins is associated with inversion of the N-glycosidic bond from β to α [Henderson (1983)].

The mono-ADP-ribosylation of nuclear proteins from chicken liver inhibits phosphorylation reactions [Tanigawa et al. (1983)]. 8.7.6.3 Poly-ADPR Transfer to Macromolecules All types of eukaryotes contain enzymes in their nuclei which catalyse the transfer of multiple ADPR residues to proteins (Eq.12): nADPR-NAm + + M —