Fortified Foods with Vitamins E-Book

• Herausgeber: Wiley-VCH
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 486

Veröffentlichungsjahr: 2011

Beschreibung

Unique in its review of modern analytical approaches to vitamin fortification, this book emphasizes fast, sensitive, and accurate methods, along with assays enabling the detection of various isomers and multiple vitamins.

The expert contributors describe the concepts as well as analytical and assay methods to study fortification, along with applications to create better and safer foods. Taking into considerations regulatory matters, they include data on sampling and extraction methods, and discuss the various pros and cons of each. As a result, readers are able to determine, which type of analytical method is best suited for added vitamins.

A practical guide for food chemists and technologists, as well as analytical laboratories and biochemists.

Leseprobe

Table of Contents

Cover

Table of Contents

Related Titles

Title page

Copyright page

Dedication

Preface

List of Contributors

Part I: Perspectives and General Methodology in Vitamin Analysis

1 Stable Isotope Dilution Assays in Vitamin Analysis – A Review of Principles and Applications

1.1 Principle of Stable Isotope Dilution Assays

1.2 Application of Stable Isotope Dilution Assays to Vitamins

1.3 Outlook

2 Analytical Methods to Assess the Bioavailability of Water-Soluble Vitamins in Food – Exemplified by Folate

2.1 Introduction

2.2 Folate Bioavailability

3 Quantitation of Vitamins Using Microbiological Assays in Microtiter Formats

3.1 Introduction

3.2 Methods and Materials

3.3 Results

3.4 Conclusion

4 Biosensors in Vitamin Analysis of Foods

4.1 Introduction

4.2 Technology

4.3 Surface Plasmon Resonance (SPR)

4.4 Biosensor Assay

4.5 Water-Soluble Vitamin Analysis by Inhibition Protein Binding Assay on Biacore Q

4.6 Validation Considerations

4.7 Conclusions

5 International Perspectives in Vitamin Analysis and Legislation in Vitamin Fortification

5.1 Introduction

5.2 General Requirements for Modern and Future Vitamin Assays

5.3 Fortification with Vitamins – The International Perspective

Part II: Analysis of Water-Soluble Vitamins

6 HPLC Determination of Thiamin in Fortified Foods

6.1 Introduction

6.2 Fortification of Foods with Thiamin

6.3 Analytical Principles

6.4 Extraction Procedures

6.5 Liquid Chromatography Procedures

6.6 Conclusion

7 HPLC Determination of Riboflavin in Fortified Foods

7.1 Introduction

7.2 Materials and Methods

7.3 HPLC Intercomparisons

7.4 Conclusion

8 HPLC–MS Determination of Vitamin C in Fortified Food Products

8.1 Introduction

8.2 Materials and Methods

8.3 Results and Discussion

9 Quantitation of Pantothenic Acid in Fortified Foods by Stable Isotope Dilution Analysis and Method Comparison with a Microbiological Assay

9.1 Introduction

9.2 Materials and Methods

9.3 Results and Discussion

10 Optimization of HPLC Methods for Analyzing Added Folic Acid in Fortified Foods

10.1 Introduction

10.2 Materials and Methods

10.3 Results and Discussion

10.4 Conclusion

11 Studies on New Folates in Fortified Foods and Assessment of Their Bioavailability and Bioactivity

11.1 Introduction

11.2 Materials and Methods

11.3 Results and Discussion

12 Analysis of Vitamin B12 by HPLC

12.1 Introduction

12.2 Sample Preparation

12.3 Separation

12.4 Detection

12.5 Reference Materials for Vitamin B12 Analysis

13 Microbiological Detection of Vitamin B12 and Other Vitamins

13.1 Vitamin B12

13.2 Analysis of Food Vitamin B12 by Bioautography

13.3 Other Vitamins Determined by Microbiological Analysis

14 Multimethod for Water-Soluble Vitamins in Foods by Using LC–MS

14.1 Introduction

14.2 Extraction and Cleanup with Conventional Methods of Analysis

14.3 Liquid Chromatographic Methods for the Analysis of Water-Soluble Vitamins

14.4 LC–MS in Multianalyte Confirmation Analyses

14.5 Electrospray Ionization and Collision-Induced Dissociation

14.6 LC–ESI-MS for Simultaneous Analysis of Water-Soluble Vitamins

14.7 Simultaneous Extraction Procedures

14.8 Conclusions and Future Developments

Part III: Analysis of Fat-Soluble Vitamins

15 Analysis of Carotenoids

15.1 Introduction

15.2 Materials and Methods

15.3 Results and Discussion

16 HPLC Determination of Vitamin E in Fortified Foods

16.1 Introduction

16.2 Fortification of Foods with Vitamin E

16.3 Experimental Procedures Used to Determine Vitamin E in Fortified Foods by HPLC

16.4 Conclusions

17 Determination of Vitamin D by LC–MS/MS

17.1 Introduction

17.2 Materials and Methods

17.3 Results and Discussion

Acknowledgments

18 Quantitation of Vitamin K in Foods

18.1 Introduction

18.2 Biological Role of Vitamin K

18.3 Dietary Vitamin K Sources

18.4 Stability of Vitamin K

18.5 Bioavailability of Vitamin K from Foods

18.6 Dietary Vitamin K Deficiency

18.7 Analytical Methods for the Determination of Vitamin K in Foods

19 Trace Analysis of Carotenoids and Fat-Soluble Vitamins in Some Food Matrices by LC–APCI-MS/MS

19.1 Introduction

19.2 Materials and Methods

19.3 Results and Discussion

19.4 Conclusions

Acknowledgment

Index

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The Editor

Prof. Dr. Michael Rychlik

Bioanalytik Weihenstephan

Technische Universität München

Alte Akademie 10

85354 Freising

Germany

All books published by Wiley-VCH are carefully produced. Nevertheless, authors, editors, and publisher do not warrant the information contained in these books, including this book, to be free of errors. Readers are advised to keep in mind that statements, data, illustrations, procedural details or other items may inadvertently be inaccurate.

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the Deutsche Nationalbibliothek

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© 2011 Wiley-VCH Verlag & Co. KGaA,

Boschstr. 12, 69469 Weinheim, Germany

All rights reserved (including those of translation into other languages). No part of this book may be reproduced in any form – by photoprinting, microfilm, or any other means – nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not specifically marked as such, are not to be considered unprotected by law.

ISBN: 978-3-527-33078-2

ePDF ISBN: 978-3-527-63417-0

ePub ISBN: 978-3-527-63416-3

Mobi ISBN: 978-3-527-63418-7

oBook ISBN: 978-3-527-63415-6

Dedicated to my caring wife and daughters

Preface

In the nutrition sciences, evidence is accumulating that vitamins are essential not only for maintaining physiological functions but also to prevent hazards resulting from oxidative stress or disorders of cell division and DNA repair. For example, the vitamers of the folate group are considered to prevent neural tube defects, Alzheimer’s disease, and colon cancer. Therefore, many foods are supplemented with vitamins, and for cereal products fortification with folic acid is mandatory in the USA.

When considering the recommended intakes of vitamins, consumers have to rely on the labeled contents to make their diet meet the recommendations. Therefore, manufacturers and official laboratories are called upon to control vitamin contents accurately. However, the analysis of vitamins is still demanding due to their occurrence in minute amounts and their lability. The trace amounts to be quantified are sometimes too small even for some highly sensitive instrumental analytical methods, so that bioassays are still important. For the determination of vitamins, organisms can be used that do not produce the analyte compounds by themselves, but need them for their growth. Well into the twentieth century, vertebrates such as chicks were used for vitamin K determination, but nowadays only bacteria are applied. Furthermore, the occurrence of vitamins as conjugates and as various isomers showing different bioactivities render vitamin analysis extremely demanding. In particular, fortification with synthetic vitamins often lacking isomeric purity has aggravated this problem.

The aim of this multi-authored book is, therefore, to review modern analytical approaches to verify the content of all relevant vitamins in fortified foods. Emphasis is placed on fast, sensitive, and accurate methods along with assays that permit the detection of various isomers and multiple vitamins. The individual contributions include both up-to-date reviews and unprecedented reports on new methods. The authors were asked to omit lengthy historical discourses and also laboratory details, as these are easily accessible from original publications.

Internationally renowned experts in vitamin research have contributed to this volume. Their disciplines range from medicine, toxicology, and nutrition science to chemistry, food science, and food chemistry. The authors are active in universities, official laboratories, and companies in many countries throughout the world, including China, Germany, Italy, Japan, Spain, Sweden, and the USA.

The book is intended for professionals, and also advanced students, concerned with and active in the areas of food science, nutraceuticals, human health, and biochemistry. They will benefit from this book which clearly describes the concepts and analytical and assay methods to study fortification, and applications to create better and safer foods.

Acknowledgments

This volume was inspired by the food science and nutrition team of the publisher, the support and assistance of which during the whole production of the book is gratefully acknowledged.

Many thanks are due to all the co-contributors, who have spent so much of their precious time writing high-quality chapters. Many chapters were included as last-minute contributions and have therefore contributed significantly to ensuring that this volume is up-to-date.

List of Contributors

Sameh Ahmed

Assiut University

Faculty of Pharmacy

Department of Pharmaceutical Analytical Chemistry

Assiut 71526

Egypt

Irmgard Bitsch

University of Giessen

Institute of Nutritional Science

Wilhelmstrasse 20

35392 Giessen

Germany

Roland Bitsch

Friedrich Schiller University Jena

Institute of Nutrition

D-07743 Jena

Germany

Volker Böhm

Friedrich Schiller University Jena

Institute of Nutrition

07737 Jena

Germany

Fulvia Caretti

Università “La Sapienza” di Roma

Dipartimento di Chimica

Piazzale Aldo Moro 5

00185 Rome

Italy

Bo Chen

Hunan Normal University

College of Chemistry and Chemical Engineering

Changsha 410081

China

Jonathan W. DeVries

Medallion Laboratories

General Mills

9000 Plymouth Avenue North

Minneapolis, MN 55427

USA

Remedios Fernández Fernández

Almeria University

Department of Analytical Chemistry

Group “Analytical Chemistry of Contaminants”

04071 Almeria

Spain

Antonia Garrido Frenich

Almeria University

Department of Analytical Chemistry

Group “Analytical Chemistry of Contaminants”

04071 Almeria

Spain

Alessandra Gentili

Università “La Sapienza” di Roma

Dipartimento di Chimica

Piazzale Aldo Moro 5

00185 Rome

Italy

Jelena Jastrebova

Swedish University of Agricultural Sciences (Uppsala BioCenter)

Department of Food Science

Undervisningsplan 6C

750 07 Uppsala

Sweden

Afaf Kamal-Eldin

Swedish University of Agricultural Sciences (Uppsala BioCenter)

Department of Food Science

Undervisningsplan 6C

750 07 Uppsala

Sweden

Current address

United-Arab Emirates University

Department of Food Science

Al Ain

United Arab Emirates

Dorit Kern

Landesamt für Verbraucherschutz des Landes Sachsen-Anhalt

Freiimfelderstrasse 66–68

061112 Halle/Saale

Germany

Naoya Kishikawa

Nagasaki University

Graduate School of Biomedical Sciences

Department of Environmental and Pharmaceutical Sciences

1-14 Bunkyo-machi

Nagasaki 852-8521

Japan

Padmanaban G. Krishnan

South Dakota State University

Department of Health and Nutritional Sciences

Wagner Hall 415, Rotunda Lane

Brookings, SD 57007

USA

Naotaka Kuroda

Nagasaki University

Graduate School of Biomedical Sciences

Department of Environmental and Pharmaceutical Sciences

1-14 Bunkyo-machi

Nagasaki 852-8521

Japan

John L. MacDonald

NP Analytical Labs

Checkerboard Square

Saint Louis, MO 63164

USA

José Luis Martínez Vidal

Almeria University

Department of Analytical Chemistry

Group “Analytical Chemistry of Contaminants”

04071 Almeria

Spain

Sabine Mönch

Technische Universität München

Chair of Food Chemistry

Lise-Meitner-Str. 34

85354 Freising

Germany

David R. Nelson

Caravan Ingredients

7905 Quivira Road

Lenexa, KS 66215

USA

Anthony O’Kane

Queens University Belfast

Institute of Agri-Food and Land Use

David Keir Building

Stranmillis Road

Belfast BT 95AG

UK

Kaname Ohyama

Nagasaki University

Graduate School of Biomedical Sciences

Department of Environmental and Pharmaceutical Sciences

1-14 Bunkyo-machi

Nagasaki 852-8521

Japan

Sudheer R. Musukula

South Dakota State University

Department of Health and Nutritional Sciences

Wagner Hall 425, Rotunda Lane

Brookings, SD 57007

USA

Roberto Romero-González

Almeria University

Department of Analytical Chemistry

Group “Analytical Chemistry of Contaminants”

04071 Almeria

Spain

Dora Roth-Meier

Technische Universität München

Animal Nutrition Center of Life and Food Sciences Weihenstephan

Liesel-Beckmann-Str. 1

Alte Akademie 10

85350 Freising

Germany

Michael Rychlik

Technische Universität München

Chair of Analytical Food Chemistry Center of Life and Food Sciences Weihenstephan

Bioanalytik Weihenstephan Research Center for Nutrition and Food Sciences

Alte Akademie 10

85350 Freising

Germany

Sylvia Stengl

R-Biopharm AG

An der neuen Bergstrasse 17

64297 Darmstadt

Germany

Lennart Wahlström

GE Healthcare

Björkgatan 30, 751 84 Uppsala

Sweden

Fumio Watanabe

Tottori University

Faculty of Agriculture

School of Agricultural

Biological and Environmental Sciences

4-101 Koyama-Minami

Tottori 680-8550

Japan

Wolfgang Weber

Institut für Produktqualität GmbH

Teltowkanalstrasse 2

12247 Berlin

Germany

Cornelia M. Witthöft

Swedish University of Agricultural Sciences

Uppsala BioCenter

Department of Food Science

Undervisningsplan 6C

75007 Uppsala

Sweden

Yukinori Yabuta

Tottori University

Faculty of Agriculture

School of Agricultural

Biological and Environmental Sciences

4-101 Koyama-Minami

Tottori 680-8550

Japan

Da-jin Yang

National Institute for Nutrition and Food Safety

Chinese Center for Disease Control and Prevention

Beijing 100021

China

Part I: Perspectives and General Methodology in Vitamin Analysis

1

Stable Isotope Dilution Assays in Vitamin Analysis – A Review of Principles and Applications

Michael Rychlik

1.1 Principle of Stable Isotope Dilution Assays

1.1.1 General Remarks

The past decades have seen an increasing use of compounds labeled with stable isotopes in research. For instance, labeled precursors facilitate metabolism studies as the label can be followed on its way into different metabolites by mass spectrometry. Another application is the labeling of high molecular weight compounds such as proteins to elucidate their three-dimensional structures by modern nuclear resonance spectrometric methods. The third important use of stable isotopes is in trace analysis by stable isotope dilution assays (SIDAs), which is the topic of this review on vitamin quantitation.

The origin of SIDAs can be traced back to the beginning of the twentieth century when Soddy [1] discovered the existence of isotopes and George Hevesy used radioactive isotopes to determine the content of lead in rocks and the solubility of lead salts in water [2]. The conviction that elements are composed of atoms containing identical nuclei was refuted by Aston [3], who detected different atomic species of the noble gas neon by mass spectrometry. This resulted in a new definition of elements, which accordingly comprise mixtures of nuclei showing identical charge but different masses. As the nuclei with identical charges have the same (Greek isos) place (topos) in the periodic system of the elements, Soddy introduced the term “isotopes” [1]. An element has a natural isotopic distribution and there are two types of isotopes, namely the stable and radioactive ones. For example, carbon shows a natural distribution of C-12 (98%), of C-13 (1.1%), and of C-14, the last of which is radioactive and undergoes β-decay with a half-life of 5370 years.

Radioisotopes have the advantages over their stable analogs of their sensitive detectability and the possible use of low degrees of labeling. However, stable isotopes thereafter found their place as analytical tools when Hevesy and Jacobsen used deuterium oxide to quantify the percentage of extracellular liquid [4]. Interestingly, the percentage of deuterated water was determined by measuring the density of the water, as the mass spectrometers at that time showed very low precision.

The term “stable isotope dilution assay” was first introduced in 1940 by Rittenberg and Foster [5], who quantified amino acids in protein hydrolyzates. SIDAs at that time were very tedious, as mass spectrometry required purification of the compounds to be analyzed. Therefore, several chromatographic and recrystallization steps were essential. Finally, coupling of mass spectrometry (MS) to gas chromatography (GC) [6] opened the door to faster and more sensitive methods. In that way, the first modern type of SIDA was performed by Sweeley et al. [7], who quantified glucose by GC–MS after trimethylsilylation and used [2H7]glucose as the internal standard.

Although they are very similar in their properties, isotopes can be enriched or depleted due to their different masses. Mixing an element or compound showing a natural isotopic distribution with such an isotopically different material (Figure 1.1) results in a smaller proportion of the naturally abundant isotopes in the resulting material – which led to the term “dilution” in SIDA.

Figure 1.1 Origin of the term “dilution” in stable isotope dilution assays: addition of a standard with a different isotopic distribution – the original isotopic distribution of the analyte has been “diluted.”

The principle of SIDA is simply explained in Figure 1.2. After addition of the labeled standard and its equilibration with the analyte, the ratio of the isotopologs remains stable throughout all subsequent analytical steps. This is due to their almost identical chemical and physical properties. A final MS step enables the isotopologs to be differentiated. Consequently, the content of the analyte in the sample can be calculated with the known amount of the internal standard (IS) added at the beginning. In contrast, a structurally different internal standard may be discriminated against and, therefore, cause systematic errors and imprecision. Hence losses of the analyte are completely compensated for by identical losses of the isotopolog, whereas the structurally different IS may show different losses.

Figure 1.2 The ratio of isotopologic analyte and standard remains stable until final mass spectroscopic analysis. For a structurally different IS, however, the ratio between standard and analyte can alter during sample preparation.

As SIDAs require more or less elaborate syntheses of labeled compounds, their development was at first restricted to very few applications, in particular to those fields in which highest sensitivity and accuracy were essential. Therefore, toxicology, clinical chemistry, and environmental analysis were the first disciplines to use SIDAs. Subsequently, these methods were transferred to foods and emerged as reference methods for food compounds such as lignans [8] and steroids [9].

However, more recently the direction of research changed and assays developed for foods have opened up new prospects in toxicology and nutrition research. In addition to vitamins such as pantothenic acid [10] and folates [11], further examples have been centered on mycotoxins such as trichothecenes [12, 13] and patulin [14] and on odorants [15].

1.1.2 Benefits and Limitations of Using an Isotopologic Internal Standard

As detailed before, due its ideal compensation for losses, SIDA is a perfect tool for a series of analytical applications, in particular for trace analyses. The latter often demand tedious clean-up procedures due to matrix interferences, which typically evoke losses of the analyte. The use of structurally different ISs requires additional recovery and spiking experiments, which often result in imprecise data. In all these cases, SIDA offers significant benefits.

In addition to compensation for losses, thus resulting in improved accuracy, the use of an isotopologic standard enhances the specificity of the determination. In addition to the specific MS information on the analyte, the IS is eluted at an almost identical retention time and shows a distinct mass shift. Therefore, the analyte can be unequivocally assigned in the chromatogram from a SIDA showing the coeluting peaks in the respective mass traces (Figure 1.3).

Figure 1.3 MS/MS chromatograms of fortified breakfast cereals containing 7.94 mg per 100 g of pantothenic acid (PA) and 270 µg per 100 g of folic acid (FA). The isotopically labeled internal standards show retention times identical with those their unlabeled isotopologs in the respective traces below.

A further advantage of adding isotopologic material is to enhance sensitivity by the so-called “carrier effect.” Due to adsorption phenomena on glassware or chromatographic columns, a definite amount of the analyte is likely to be lost during sample clean-up. If the total amount of the analyte in an extract is lower than this loss, the compound will no longer be detectable. However, if an isotopologic standard is added in an amount that exceeds this loss, the total sum of standard and analyte is higher than the loss and, therefore, the isotopologs can be detected. Although there are some conflicting opinions on this topic (for a review, see [16]), there are some applications showing a significant enhancement of sensitivity [17]. For vitamins, however, this effect has not yet been demonstrated.

Regarding the major benefits of SIDAs, that is, specificity and ideal compensation for losses, the question arises of whether they have the potential to be “definite methods.” According to a definition by Cali and Reed [18], “a definite method is one that, after exhaustive investigation, produces analytical results that are accurate, that is, free of systematic errors, to the extent required for the intended end-use(s).” This definition holds true especially for primary methods, which are “methods having the highest metrological qualities, for which a complete uncertainty statement can be written down in terms of SI units, and whose results are, therefore, accepted without reference to a standard of the quantity being measured” [19]. A SIDA can be traced back to a gravimetric (i.e., primary) measurement and, therefore, is considered a primary method. In the case of being validated intensively for the absence of systematic errors, it has the potential to be accurate, that is, to produce the “true” value.

A SIDA of vitamins is only possible if a combination of chromatography with MS and a labeled IS are available. However, whereas the former has increasingly become basic instrumentation in most laboratories, the latter is a narrow bottleneck for the wider application of SIDAs. Of all the vitamins discovered to date, about 20 have been synthesized as labeled analogs, but only seven of these are commercially available. Hence the compound aimed at often has to be synthesized. There are two aspects that arise as a hindrance before starting with these syntheses. The first is that analysts may hesitate to perform chemical syntheses due to a lack of experience. Although the envisaged syntheses on the microscale require often intense purifications by chromatography, analysts are normally familiar with the necessary methods such as distillation in vacuo, purification by high-performance liquid chromatography (HPLC) and checking purity and yield by GC–MS. The procedures are completely detailed in the literature and usually the groups having published these syntheses are willing to give advice in case of practical problems.

The second psychologic hindrance that prevents analysts from synthesizing labeled mycotoxins is the price of labeled educts. However, this is not a convincing argument, as can be explained by the following example: 1 g of a labeled educt may cost around US$1000 and the yield of a multi-step synthesis may be only 1%, both of which are realistic figures. Then, the price for 10 mg of the labeled product is $1000. However, as less than 1 µg of the labeled standard is required for a SIDA, 10 mg of the standard enables at least 10 000 analyses to be performed. Hence the material cost for using a labeled standard is just $0.10 per sample, which is negligible compared with the cost of labor and equipment.

1.1.3 Prerequisites for Isotopologic Standards

As outlined before, SIDA is based on an isotopolog ratio that remains stable during all analytical steps. Therefore, a stable labeling step is essential for an IS. As carbon–carbon and carbon–nitrogen bonds are very unlikely to be cleaved, 13C and 15N labels are considered to be very stable. In contrast, losses of 18O or 2H at labile positions can occur. On the one hand, 18O in carboxyl moieties can be exchanged in acidic or basic solutions. On the other, deuterium is susceptible to so-called protium–deuterium exchange if it is activated by adjacent carbonyl groups or aromatic systems.

Moreover, an isotopolog ratio may be altered by isotope effects (IEs), that is, small differences in physical or chemical properties of the isotopologs. IEs are due to different energy contents that are caused by the mass differences of the isotopes. The lowest energy level of a molecule, namely the zero point energy, is given by the ground-state vibration of the bonds at absolute zero, where the population in excited vibrational levels is negligible. As frequencies and energies of vibrations are proportional to the encountered masses, heavier isotopologs possess a lower energy content, resulting in higher energies of bond dissociation. IEs are mainly observable in the case of hydrogen, as the mass difference between 1H and 2H is proportionally much higher than between 13C and 12C or 15N and 14N. In the case of chemical reactions involving C–H bond rupture, IEs result in monodeuterated isotopologs reacting up to seven times more slowly then their light analogues. For labeling with 13C, these effects are several hundred times smaller. However, the kinetic IE of 13C is a valuable diagnostic tool in enzyme studies, as 12C isotopologs show lower reaction rates than their 13C counterparts. This discrimination is commonly measured as the δ13C value of the products and is dependent on the transitions states of the encountered enzymatic reactions. Therefore, the measurement of δ13C values allows the characterization of the encountered enzymes and the determination of the products’ origin. For instance, the authenticity of alcoholic beverages [20] or valuable spices such as vanilla [21] can be verified.

The primary IE, which affects direct bond cleavage between the label and adjoining atoms, has to be distinguished from the secondary IE, which has an influence on the bonds between unlabeled atoms. A primary IE would cause either loss of labeling or, in MS, would result in different intensities of the fragments bearing the label. Therefore, all derivatizations or MS fragmentations including a primary IE have to be avoided. Although much less pronounced, a secondary IE, however, cannot be excluded in these reactions, but has not yet been reported in the literature.

In contrast to chemical IEs, a physical IE is more often observable. The latter particularly affects chromatographic behavior and results in different retention times of the isotopologs. In particular for multiple labelings with 2H, the IE may evoke even baseline separation, as shown in the case of eightfold deuterated β-carotene [22], which was clearly separated from its unlabeled isotopolog upon HPLC with ultraviolet (UV) detection (Figure 1.4) In most cases, the heavier isotopologs are eluted earlier than their light analogs, which is unexpected, as the heavier isotopologs should have higher boiling temperatures and, therefore, should be eluted later in GC. This behavior therefore is referred to as an inverse IE.

Figure 1.4 Baseline separation of eightfold deuterated β-carotene from its unlabeled isotopolog using HPLC–UV.

Reproduced with permission from [22].

In order to prevent chromatographic separations of the isotopologs during clean-up and thus changes in the isotopolog ratio, isotope effects have to be minimized either by choosing labelings with 13C or 15N or by introducing only the necessary number of 2H atoms.

For unequivocal quantification, the standard has to be distinguishable from the analyte by MS. This requires the presence of the mass increment introduced by the labels either in the molecular ion or in its fragments. Therefore, a loss of the label prior to detection has to be avoided. However, quantitation by liquid chromatography (LC) coupled with tandem mass spectrometry (MS/MS) is feasible even if the label is lost during the monitored mass transition, as the precursor ions are still differentiated.

A further problem in isotopolog differentiation may arise from spectral overlaps between the standard and analyte. In case of the analyte, the natural abundance of isotopes, in particular of 13C, 18O and 34S, results in isotope clusters of each fragment showing not only the nominal mass mA, but also to a lesser extent mA + 1, mA + 2, or even higher masses. In particular, 13C in compounds consisting of a higher number of carbons causes a significant abundance of m + 1 and m + 2 due to the relatively high natural abundance. To avoid overlap of the signals of those natural isotopologs with mS of the standard, the mass difference therefore has to be sufficient. For vitamins, in which the number of carbon atoms often exceeds 10, generally a mass increment of at least three units is required. However, the number of labels, especially of deuterium, should not be too high, in order to minimize the mentioned chromatographic isotope effect.

In contrast, signals of the labeled material contributing to those of the analyte may also result in spectral overlaps. This may be due to the low isotopic purity of labeled educts or to inadequate labeling during synthetic steps, thus leading to incomplete labels in the standard.

If a spectral overlap cannot be avoided, calculation procedures have been developed that still permit quantification to be achieved. However, these procedures are more complicated the greater the overlap is.

Another important requirement for accurate quantification is complete equilibration between the analyte and standard in the sample to be extracted. As the labeled standard usually is not contained in the sample matrix after its addition, it will likely be recovered to a high extent during the extraction procedure. For the analyte, this might not be true, as it can be trapped in compartments of the matrix and is less extractable by the solvent. Therefore, sufficient time for equilibration of the standard and analyte in all parts of the sample has to be allowed to assure that the analyte and standard show the same concentration ratio in all compartments as far as possible.

1.1.4 Calibration Procedures

From the intensity ratio of suitable ions measured by MS and a calibration function, the isotopolog ratio can be assessed, which directly allows the amount of analyte present in the sample to be calculated. The relation between isotopolog ratio and intensity ratio has to be elucidated by analyzing the intensity ratios of a series of defined standard–analyte mixtures. If there is no spectral overlap, the calibration function is assumed to be linear. However, typically there are still residues of unlabeled analytes in the labeled material and low intensities of natural isotopologs of the analyte contributing to the signals of the standard. Therefore, the calibration function can be expected to be linear only in a restricted region, as illustrated in Figure 1.5. The outlined function appears under the supposition that the standard may contain 2% of unlabeled material und natural isotopologs may contribute 5% intensity to the signal of the standard. In this case, the calibration function can only be assumed to be a straight line for molar ratios of analyte to standard ranging between 0.2 and 5. With excess of the standard or of the analyte, the function is dominated by the unlabeled residues in the standard or by the natural isotopologs of the analyte, respectively.

Figure 1.5 “Real” calibration function (dashed line) for a SIDA under the supposition that the standard contains 2% of unlabeled material with natural isotopologs contributing 5% intensity to the standard’s signal.

However, for complicated structures or elaborate syntheses, a spectral overlap often cannot be avoided and, therefore, suitable procedures for calculations are required. In general, there are several ways to cope with this problem: First, hyperbolic or polynomial models have been elaborated [23, 24], which approximate the real calibration relation by a mathematical function. As these procedures are rather complicated, several authors have proposed linearization methods, which convert the nonlinear function into a linear function. In a study on odorants, Fay et al. [25] compared four of these methods using [2H1]benzaldehyde as the IS for quantification of benzaldehyde. This assay shows a spectral overlap of about 12% between the analyte and the standard. The authors demonstrated that the method of Colby and McCaman [26] did not yield a linear function. By contrast, both the average mass approach [27] and linearization using isotopic enrichment factors [28] gave straight lines, whose calibration points, however, were spread very inhomogeneously. The only procedure giving a calibration line with homogeneously distributed calibration points was the method of Bush and Trager [29], which notably includes only rather simple calculations.

The third way to cope with a nonlinear calibration function is the “bracketing” approach [30], which requires the measurement of further calibration points lying in the proximity of the intensity ratio measured in the sample. Although this method is widely accepted as the most accurate one, it is very elaborate and, therefore, is only seldom applied.

1.2 Application of Stable Isotope Dilution Assays to Vitamins

1.2.1 Fat-Soluble Vitamins

This group of vitamins is naturally embedded in food lipids. For their analysis, these compounds have to be separated from fat prior to detection. Due to their low volatility, GC applications are scarce. Moreover, the hydrophobicity of these vitamins is responsible for a high retention on common reversed phases in HPLC, which limits their separation on these phases. As a consequence thereof and as LC–MS combinations need aqueous mobile phases, reports on LC–MS detection are also rare. However, with the development of new reversed-phase materials, LC–MS applications have become increasingly popular.

For fat-soluble vitamins, the use of stable isotopologs is restricted to β-carotene, vitamin A, α-tocopherol, vitamin K, and vitamin D. To date, nearly all applications have been directed towards quantitation of these vitamins in blood or studies of their bioavailability. SIDAs in foods have not been reported so far.

1.2.1.1 Vitamin A

For vitamin A, most applications of labeled compounds have been dedicated to the administration of labeled β-carotene and quantification of the generated retinol in blood serum. For this purpose, β-carotene and retinol have been labeled with deuterium [22] or carbon-13 [31]. These compounds have been applied in several studies, and the bioefficacy of β-carotene was found to range between 30 and 40% [32].

1.2.1.2 Vitamin E

Because it functions as an antioxidant in tissues, an important part of tocopherol research is dedicated to studying its reaction with oxygen and to elucidating the respective reaction products. An extensive SIDA for the quantification of α-tocopherol and its oxidation products such as α-tocopherolquinone, α-tocopherolhydroquinone, and several epoxy-α-tocopherolquinones by using their deuterated analogs was presented by Liebler et al. [33]. They oxidized endogenous α-tocopherol in rat liver microsomes with the radical generator azobis(amidino­propane) and quantified the isotopologs by GC–MS after trimethylsilylation.

1.2.1.3 Vitamin D

Cholecalciferol (vitamin D3) and its major metabolites calcidiol and calcitriol were quantified in human blood by two research groups in the 1980s. Whereas Zagalak and Borschberg [34] used [2H8]cholecalciferol along with [2H3]calcitriol as IS, Coldwell et al. [35] applied 2H6-labeled isotopologs of cholecalciferol, calcidiol, calcitriol, and some other metabolites for SIDAs. As a further isotopolog of vitamin D3, [2H7]cholecalciferol was synthesized by Kamao et al. [36] and used in a SIDA for several fat-soluble vitamins in human breast milk.

An actual LC–MS/MS method for vitamin D in mineral tablets and baby food is presented in Chapter 17.

1.2.1.4 Vitamin K

One of the seldom used SIDAs for fat-soluble vitamins has been reported for vitamin K1(20) in plasma using GC–MS [37]. The IS was [2H3]phylloquinone, which was prepared by deuteromethylation of 1,4-naphthoquinone and subsequent coupling with phytol. The suitability of this IS, however, has been called into question for LC–MS, as the deuterium label is at the acidic 2-methyl position and, therefore, is highly susceptible to H–D exchange in aqueous eluents. In contrast, Suhara et al. synthesized 18O2-labeled phylloquinone homologs starting from 2-methyl-1,4-naphthoquinone diacetate in a four-step procedure including an oxidation in the presence of H218O [38]. These compounds were used as ISs in a multiple SIDA for the quantitation of vitamin K along with vitamin K1, K4, and K7 compounds in human breast milk [36]. Further applications of these compounds to foods have not yet been reported.

1.2.2 Water-Soluble Vitamins

Because of their low hydrophobicity and low retention on reversed phases, this group of vitamins is very suited to LC–MS in aqueous solvents. Numerous applications of LC–MS have been published, so it is not surprising that some SIDAs have also been developed. In the following sections, such assays for pyridoxin, niacin, folic acid, and pantothenic acid are described.

1.2.2.1 Vitamin B6

Due to the wide variety of pyridoxine vitamers, SIDA applications for this group of vitamins are rare. The only assay reported in the literature dates back to 1985, when Hachey et al. [39] used deuterated analogs of pyridoxine, pyridoxal, pyridoxamine, and pyridoxic acid to quantify these compounds and their phosphates in guinea pig liver, human urine, feces, and goat milk. In the last case, pyridoxal phosphate was found to be the main vitamer contributing almost 70% to total pyridoxine.

1.2.2.2 Niacin

As [2H4]nicotinic acid is commercially available, it has been used as the basis of SIDAs for the niacin group. Following the first application to determine nicotinic acid and six of its metabolites in urine by Li et al. [40], the first SIDA for foods was presented by Goldschmidt and Wolf [41]. However, the latter assay did not measure nicotinamide, which might also contribute to niacin activity in food samples. Nevertheless, the assay showed excellent accuracy for the analysis of certified reference materials such as wheat flour, milk powder, and multivitamin tablets.

1.2.2.3 Ascorbic Acid

Although its importance for the diet is well documented, LC–MS quantitations of ascorbic acid are surprisingly scarce. Examples of the rare reports are a study of degradation products of dehydroascorbic acid and ascorbic acid [42] and the most recent quantitation of vitamin C along with nine other vitamins in multivitamin products by Chen et al. [43].

Moreover, labeled ascorbic acid for LC–MS has not yet been used. The only SIDA for vitamin C was reported in 1988, when Ellerbe et al. applied 13C-labeled ascorbic acid to milk powder analysis by GC–MS of the tert-butyldimethyl derivatives [44].

1.2.2.4 Folic Acid

The first SIDA for folic acid in fortified foods was published by Pawlosky and Flanagan [45], who used commercially available [13C5]folic acid ([13C5]PteGlu) as the IS. One year later, we synthesized fourfold deuterated folic acid along with the most abundant folate monoglutamates for quantifying endogenous food folates and folic acid in fortified foods [46]. Further developments included the use of chicken pancreas in addition to rat plasma for improved deconjugation [47] and the use of 4-morpholinoethanesulfonic acid (MES) for enhancing folate stability [48]. In particular, the need for labeled folate tracers in bioavailability research spurred the generation of new stable isotope-labeled folates. Starting with the first dual label study using [2H2]PteGlu and [2H4]PteGlu by Gregory and Quinlivan [49], early investigations were restricted to the sole measurement of total urinary folate isotopologs by GC–MS [50, 51] and often were hampered by spectral overlap due to insufficient mass increments interfering with naturally occurring isotopologs. For this reason, syntheses of differently labeled folates have been developed, such as [2H4]PteGlu and [13C6]PteGlu labeled in the glutamate moiety and the benzene moiety, respectively [52]. For tracer studies, these folates were suitable for so-called extrinsic labeling, that is, by simple addition to foods. However, as added substances may not behave like the endogenously occurring folates, so-called intrinsically labeled foods were produced by growing spinach in a 15N-labeled environment [53], which generated 15N1–7-labeled 5-methyltetrahydrofolate (5-CH3-H4folate) in the latter vegetable.

A further methodological improvement arose from the use of LC–MS in folate isotopolog analysis, when Wright et al. [54] measured 13C6-labeled, 15N1–7-labeled, and unlabeled 5-CH3-H4folate using [2H2]folic acid as the IS in the single ion monitoring mode. However, the use of a structurally different IS such as [2H2]PteGlu may decrease the accuracy by ion suppression and, moreover, quantitation was hampered by spectral overlap of [15N1–7]-5-CH3-H4folate with, on the one hand, [13C6]-5-CH3-H4folate, and, on the other, unlabeled 5-CH3-H4folate in single-stage LC–MS. This handicap was overcome by Melse-Boonstra et al. [55], who measured 13C6-labeled along with 13C11-labeled 5-CH3-H4folate as tracer isotopologs and simultaneously quantified unlabeled 5-CH3-H4folate by using [13C5]-5-CH3-H4folate as the IS. In the latter study, spectral overlaps of 5-CH3-H4folate isotopologs were avoided by labeling different moieties of the target molecule and their differentiation by LC–MS/MS. However, this investigation was restricted to plasma 5-CH3-H4folate without an application to food samples.

With more 13C-labeled folates now being offered commercially, SIDA for folates in foods is currently being used by various groups throughout the world. Of all folate vitamers, H4folate, 5-CH3-H4folate, 5-formyl-H4folate, 5,10-methenyl-H4folate and folic acid are currently available as 13C5-labeled isotopologs. Whereas SIDA for monoglutamates can be considered a fairly well-established technology, the analysis of endogenous folates is still challenging. In particular, the polyglutamates and their complete conversion to the monoglutamates are still not fully understood and are the target of further research. The most recent approach is the use of [13C5]pteroylheptaglutamate for confirmation and quantitation of the degree of deconjugation. The latter compound has been produced by a Merrifield-like solid phase synthesis [48].

1.2.2.5 Pantothenic Acid

Along with folic acid, pantothenic acid was one of the first vitamins to be quantified by SIDA based on LC–MS. An extensive description of [13C3,15N]pantothenic acid used as internal standard and recent results are presented in Chapter 9.

1.3 Outlook

Although numerous labeled vitamins are used in clinical chemistry and bioavailability research, they have been only marginally applied in the SIDA of foods (Table 1.1). However, with the advances in LC–MS/MS in vitamin analysis, labeled ISs will gain more importance. In particular for the analysis of multivitamin products, multiparametric SIDAs will be developed, when the labeled standards become available, as can be seen from the past for folic acid and pantothenic acid [56]. In particular for the analysis of reference materials, SIDAs are essential to establish the certified reference contents. In this context, a SIDA for the analysis of seven water-soluble vitamins in an infant/adult nutritional formula has been reported very recently [57].

Table 1.1 Applications of SIDAs to vitamins in foods.

References

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2

Analytical Methods to Assess the Bioavailability of Water-Soluble Vitamins in Food – Exemplified by Folate

Cornelia M. Witthöft

2.1 Introduction

Folate is the generic term for a large group of vitamers (Figure 2.1) participating in one-carbon transfer reactions required for thymidylate and purine biosynthesis and amino acid interconversions. Folate forms differ in the oxidation status of the pteridine moiety being oxidized or reduced as 7,8-dihydro- (H2folate) and 5,6,7,8-tetrahydrofolate (H4folate) and their one-carbon substituent at N-5 or N-10. Native food folates are mainly folyl polyglutamates with up to seven glutamic acid residues [1, 2]. Food composition tables indicate that the folate content in food is usually low, in the region of several micrograms per 100 g of food [3]. Foods are sometimes classified into good and moderate folate sources with folate concentrations ranging from 50 to 100 µg and from 15 to 50 µg per serving, respectively [4].

Figure 2.1 Structure of native food folates and their substituent groups and positions. n, number of glutamates.

Individual native folates are susceptible – to different extents depending on their chemical nature – to oxidation leading to cleavage of the molecule at the C-9–N-10 bond and subsequent loss of their biological activity (summarized in [5, 6]). Some storage and food processing, under both household and industrial conditions, can result in substantial losses of the vitamin activity of dietary folates, as summarized in [2]. For fortification purposes, the synthetic and fully oxidized vitamer folic acid is used for economic reasons (low cost of synthesis) and due to its high stability. In retention trials using food model systems and different food processing methods, folic acid exhibited higher retention and a half-time (t½) up to more than 100-fold compared with native folates [2]. Regarding the analysis of folic acid in fortified foods, for further information see Chapter 10.

Several health benefits are associated with a good folate status, due to which this vitamin has attracted both scientific and public interest in recent years. The role of folate in the prevention of neural tube defects (NTDs), such as spina bifida, is now well established [7–10]. With respect to the prevention of coronary heart disease, folate is assumed to play a key role in risk reduction by lowering serum homocysteine concentration, which has been suggested as an independent risk factor [11, 12]. However, evidence about the health protective role of folate – or the mechanisms behind it – is less consistent with respect to certain cancers, for example, the role of folate status regarding cancer initiation, progression and growth of subclinical cancers, efficacy of antifolate treatment drugs [13–17], and neuro-psychiatric disorders such as dementia and Alzheimer’s disease [18, 19]. As summarized by de Bree et al. [20] in 1997, adults in several European countries had an average daily folate intake from below 200 to around 300 µg, which was in line with recommendations at that time aimed at preventing folate deficiency. However, the authors stated that – with respect to health benefits from a good folate status – the desired dietary intake should be higher than 350 µg per day, and that only a small part of the studied European populations reach that goal. They also pointed out methodological difficulties in the estimation and comparison of micronutrient intake. The observation of suboptimal folate intakes by European populations has also been pointed out in recent studies [21], and this is commonly attributed to the low bioavailability of natural food folates. Data from the US National Health and Nutrition Examination Survey (NHANES) show that before the introduction of mandatory fortification, 20–30 and 35–60% of the population were at risk for low serum and erythrocyte folate concentrations, respectively [22]. In the USA and Canada, the concept of dietary reference intakes was developed for folates and other micronutrients [23] with the aim of including in current concepts the role of nutrients in long-term health, going beyond deficiency diseases. According to new recommendations, based in large part on information about bioavailability, the recommended intake levels were increased to 400–600 µg per day for females of childbearing age and during pregnancy and lactation [23]. A similar development of increased intake recommendations, but to a somewhat lower extent, has been observed in several European countries since 1998. The development of harmonized best practice guidance for a science base for setting micronutrient recommendations is also an aim of the European network of excellence EURRECA [24, 25].

The introduction of nationwide mandatory fortification of cereal-grain products and ready-to-eat cereals in the USA and Canada in 1998 had the particular aim of reducing the incidence of NTD-affected pregnancies. The US folic acid fortification level of 140 µg per 100 g, chosen to increase the intake of women of childbearing age while preventing excessive intake by other groups in the population, was aimed to provide an additional 100 µg of folic acid to the average diet. In several other countries, such as Chile and Costa Rica, which introduced folic acid fortification in staple foods to reduce the number of births complicated by NTD, this public health goal was reached [26, 27]. In the USA, NTD prevalence was reduced by 26% [28]. Furthermore, in most population groups, folate status improved in the post-fortification period so that less than 1 and 5% are at risk for too low serum and erythrocyte folate concentrations, respectively [22]. However, the same report stated that with the current fortification practice, the part of the US population having serum folate concentrations above the normal range of 20 ng ml−1