Food chemistry, function and analysis. Volume 6, Cereal grain-based functional foods : carbohydrate and phytochemical components 9781788012799, 1788012798, 9781788015325, 1788015320, 9781788011488

Table of contents :
Content: Introduction
Overview of Grain Components and Changes Occurring in Grain Constituents with Different Forms of Processing
Composition and Functionality of Sugars and Oligosaccharides in Cereal Grains
Types and Functionality of Polysaccharides in Cereal Grains
Starch Properties and Modification in Grains and Grain Products
Definition and Analysis of Dietary Fiber in Grain Products
Resistant and Slowly Digested Starch in Grain Products
Functionality of Beta-glucan from Oat and Barley and Its Relation with Human Health
Dietary Arabinoxylans in Grains and Grain Products
Non-digestible Oligosaccharides in Grain Products
Starch-Protein and Starch-Lipid Interactions and Their Effects on the Digestibility of Starch
Types and Distribution of Phenolic Compounds in Grains
Bound Phenolic Constituents as Co-passengers of Dietary Fibre
Anthocyanins, Deoxyanthocuanins and Proanthocyanidins as Dietary Constituents in Grain Products
Interactions Between Grains and the Microbiome
Subject index.

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Published on 06 September 2018 on https://pubs.rsc.org | doi:10.1039/9781788012799-FP001

Cereal Grain-based Functional Foods

Carbohydrate and Phytochemical Components

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Food Chemistry, Function and Analysis

Published on 06 September 2018 on https://pubs.rsc.org | doi:10.1039/9781788012799-FP001

Series editors:

Gary Williamson, University of Leeds, UK Alejandro G. Marangoni, University of Guelph, Canada Juliet A. Gerrard, University of Auckland, New Zealand

Titles in the series:

1: Food Biosensors 2: Sensing Techniques for Food Safety and Quality Control 3: Edible Oil Structuring: Concepts, Methods and Applications 4: Food Irradiation Technologies: Concepts, Applications and Outcomes 5: N  on-extractable Polyphenols and Carotenoids: Importance in Human Nutrition and Health 6: C  ereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components

How to obtain future titles on publication:

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Book Sales Department, Royal Society of Chemistry, Thomas Graham House, Science Park, Milton Road, Cambridge, CB4 0WF, UK Telephone: +44 (0)1223 420066, Fax: +44 (0)1223 420247 Email: [email protected] Visit our website at www.rsc.org/books

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Cereal Grain-based Functional Foods Carbohydrate and Phytochemical Components

Edited by

Trust Beta

University of Manitoba, Canada Email: [email protected] and

Mary Ellen Camire

University of Maine, USA Email: [email protected]

Published on 06 September 2018 on https://pubs.rsc.org | doi:10.1039/9781788012799-FP001

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Food Chemistry, Function and Analysis No. 6 Print ISBN: 978-1-78801-148-8 PDF ISBN: 978-1-78801-279-9 EPUB ISBN: 978-1-78801-532-5 Print ISSN: 2398-0656 Electronic ISSN: 2398-0664 A catalogue record for this book is available from the British Library © The Royal Society of Chemistry 2019 All rights reserved Apart from fair dealing for the purposes of research for non-commercial purposes or for private study, criticism or review, as permitted under the Copyright, Designs and Patents Act 1988 and the Copyright and Related Rights Regulations 2003, this publication may not be reproduced, stored or transmitted, in any form or by any means, without the prior permission in writing of The Royal Society of Chemistry or the copyright owner, or in the case of reproduction in accordance with the terms of licences issued by the Copyright Licensing Agency in the UK, or in accordance with the terms of the licences issued by the appropriate Reproduction Rights Organization outside the UK. Enquiries concerning reproduction outside the terms stated here should be sent to The Royal Society of Chemistry at the address printed on this page. Whilst this material has been produced with all due care, The Royal Society of Chemistry cannot be held responsible or liable for its accuracy and completeness, nor for any consequences arising from any errors or the use of the information contained in this publication. The publication of advertisements does not constitute any endorsement by The Royal Society of Chemistry or Authors of any products advertised. The views and opinions advanced by contributors do not necessarily reflect those of The Royal Society of Chemistry which shall not be liable for any resulting loss or damage arising as a result of reliance upon this material. The Royal Society of Chemistry is a charity, registered in England and Wales, Number 207890, and a company incorporated in England by Royal Charter (Registered No. RC000524), registered office: Burlington House, Piccadilly, London W1J 0BA, UK, Telephone: +44 (0) 207 4378 6556. Visit our website at www.rsc.org/books Printed in the United Kingdom by CPI Group (UK) Ltd, Croydon, CR0 4YY, UK

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Preface Cereals play a part in ameliorating climate change, growing populations, and the increased incidence of diabetes, obesity and cardiovascular disease. Although consumers may view grains as merely sources of carbohydrates, these foods supply crucial nutrients and phytochemicals. We have invited noted cereal scientists from four continents to share their expertise on grain nutraceuticals to stimulate new research and encourage collaborations. Our colleagues have contributed thought-provoking perspectives on grain components and their role in human health. Their efforts are much appreciated. We sincerely thank the staff at the Royal Society of Chemistry for their support and encouragement during the production process. We could not have produced this reference without support from family and friends. Trust Beta and Mary Ellen Camire

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Contents Chapter 1 Introduction  T. Beta

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1.1 Whole Grains as Delivery Packages for Nutrients and Phytochemicals  1.2 Whole Grains and Health Promotion  1.3 Digestion, Bioaccessibility and Bioavailability of Whole Grain Nutrients and Phytochemicals  1.3.1 Genetic Studies  1.3.2 Food Processing  1.3.3 Bioaccessibility and Bioavailability  1.4 Purpose of this Book  1.5 What This Book Does not Set Out to Do  References  Chapter 2 Overview of Grain Components and Changes Occurring in Grain Constituents with Different Forms of Processing  Mary Ellen Camire



2.1 Introduction  2.2 Pre-processing Steps  2.2.1 Cleaning  2.2.2 Tempering and Soaking  2.2.3 Dehulling and Pearling  2.2.4 Milling 

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2.3 Processing with Minimal Heat  2.3.1 Sprouting (Germination)  2.3.2 Fermentation  2.3.3 Pasta and Noodle Production  2.4 Thermal Processing with Water  2.4.1 Nixtamalization  2.4.2 Boiling  2.4.3 Steaming  2.4.4 Baking  2.5 Thermal Processing with Minimal Added Water  2.5.1 Popping and Puffing  2.5.2 Roasting  2.5.3 Microwave Heating  2.5.4 Infrared Heating  2.5.5 Extrusion  2.6 Conclusions  Acknowledgements  References  Chapter 3 Composition and Functionality of Sugars and Oligosaccharides in Cereal Grains  Yongfeng Ai



3.1 Introduction  3.2 Definition, Classification and Determination of Carbohydrates  3.3 Sugars and Oligosaccharides in Mature Cereal Grains  3.4 Changes in the Composition of Sugars and Oligosaccharides During Cereal Kernel Development  3.4.1 Normal Varieties of Cereal Crops  3.4.2 “Sweet” Mutants of Cereal Crops  3.5 Changes in Sugars and Oligosaccharides During Storage and Processing of Cereal Grains  3.5.1 Storage  3.5.2 Reactions Generating Sugars and Oligosaccharides During the Processing of Cereal Grains  3.5.3 Reactions Using Sugars and Oligosaccharides as Substrates During Processing of Cereal Grains  3.6 Conclusions and Future Trends  References 

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Chapter 4 Types and Functionality of Polysaccharides in Cereal Grains  L. Saulnier

4.1 Introduction  4.2 Tissue Organization and Cell Walls in Mature Cereal Grains  4.2.1 Cereal Grain Morphology and Tissue Organization  4.2.2 Cell Wall Composition  4.3 Structure and Properties of Major Polysaccharides from the Starchy Endosperm of Cereal Grains  4.3.1 Arabinoxylans  4.3.2 Mixed-Linked Beta-Glucans  4.3.3 Arabinogalactan Peptides  4.3.4 Interactions of Polymers in Endosperm Cell Walls  4.4 Structure and Properties of Major Polysaccharides from the Outer Layers of Cereal Grains  4.4.1 Heteroxylans  4.4.2 Cellulose  4.4.3 Lignins and Hydroxycinnamic Acids  4.5 Conclusions  Acknowledgements  References  Chapter 5 Starch Properties and Modification in Grains and Grain Products  Harold Corke and Fan Zhu



5.1 What Is Starch and Why Is It so Complicated?  5.2 What Are the Important Properties of Starch and How Are They Measured?  5.2.1 Examining Starch Morphology  5.2.2 Amylose Content (Apparent Amylose)  5.2.3 Viscoamylography  5.2.4 Dynamic Rheology  5.2.5 Gelatinization Temperature  5.2.6 Texture  5.2.7 Retrogradation  5.2.8 Chain Length Distribution  5.2.9 Digestibility and Resistant Starch 

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5.3 Examples of the Impact of Genetic Variations  5.3.1 Wheat: from Udon Noodles to Waxy Starch  5.3.2 More Grains, More Noodles  5.4 Modified Starches  5.5 Interactions of Starch with Polyphenols  5.6 Conclusions  References  Chapter 6 Definition and Analysis of Dietary Fiber in Grain Products  B. V. McCleary, J. Cox, R. Ivory and E. Delaney





6.1 Dietary Fiber as an Important Food Ingredient  6.2 Evolution of the Codex Alimentarius Definition of Dietary Fiber  6.3 Development of a Procedure for the Measurement of Total Dietary Fiber, Including Resistant Starch and Non-digestible Oligosaccharides  6.4 Integrated Procedure for the Measurement of Total Dietary Fiber as Defined by Codex Alimentarius  6.5 Rapid Integrated Procedure for the Measurement of Total Dietary Fiber as Defined by Codex Alimentarius  6.5.1 Preparation of Test Samples  6.5.2 Enzyme Purity  6.5.3 Enzymatic Digestion of Sample  6.5.4 Determination of HMWDF (IDF + SDFP)  6.5.5 Determination of SDFS  6.5.6 Calculations for HMWDF (IDF + SDFP), SDFS and TDF 6.5.7 Safety Considerations  6.6 Addressing Each of the Limitations of the INTDF Procedure  6.6.1 Optimization of PAA and AMG  6.6.2 Choice of HPLC Column  6.6.3 Preparation of Samples for HPLC 

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6.6.4 Recovery of Polysaccharides and Non-digestible Oligosaccharides Under the RINTDF Assay Method  6.6.5 Safety Considerations  6.7 Inter-laboratory Evaluation of the RINTDF Method  6.8 Conclusions  References  Chapter 7 Resistant and Slowly Digested Starch in Grain Products  P. C. Drawbridge and T. Beta



7.1 Introduction  7.2 Types of Resistant Starch  7.3 Starch Synthesis in Grains and Formation of Resistant Starch  7.4 Resistant Starch and Slowly Digestible Starch in Cereal Grains  7.5 Resistant Starch in Processed Grain Products  7.5.1 Commercially Available Resistant Starch for Use in Cereal Grain Foods  7.6 Potential as a Functional Ingredient: Resistant Starch and Health  7.6.1 Effects of Different Types of Resistant Starch on Health  7.6.2 Review of the Differential Benefits Among Types of Resistant Starch  7.7 Conclusions  References  Chapter 8 Functionality of Beta-glucan from Oat and Barley and Its Relation with Human Health  Nancy Ames, Joanne Storsley and Sijo Joseph Thandapilly



8.1 Introduction  8.1.1 Chemical Structure and Occurrence of Beta-glucan  8.1.2 History of Oats and Barley  8.2 Health Benefits of Beta-glucans  8.2.1 Health Claims  8.2.2 Lowering Cholesterol 

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8.2.3 Glycaemic Response  8.2.4 Gut Microbiota  8.2.5 Hypertension  8.3 Physicochemical and Functional Properties  8.3.1 Role of Viscosity  8.3.2 Factors Influencing the Physicochemical Properties of Beta-glucan  8.4 Summary  References  Chapter 9 Dietary Arabinoxylans in Grains and Grain Products  Marta S. Izydorczyk



9.1 Introduction  9.2 Molecular Structure and Physicochemical Properties  9.3 Arabinoxylans in Cereal Grains, Milling Fractions and Cereal-based Products  9.4 Changes in the Content and Properties of Arabinoxylans During Processing  9.5 Extraction of Arabinoxylans  9.6 Arabinoxylans as Prebiotics  9.7 Effects of Arabinoxylans on Glucose Metabolism  9.8 Immunological Effects of Arabinoxylans  9.9 Antioxidant Properties of Arabinoxylans  9.10 Conclusions  References 

Chapter 10 Non-digestible Oligosaccharides in Grain Products  Lovemore Nkhata Malunga and Trust Beta

10.1 Introduction  10.2 Non-digestible Oligosaccharides in Cereal Grains  10.2.1 Cereal Grain Fructans  10.2.2 Raffinose Family Oligosaccharides  10.2.3 Arabinoxylan Oligosaccharides 

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10.3 Potential Health Benefits of Non-digestible Oligosaccharides  10.4 Conclusions  References 

Chapter 11 Starch–Protein and Starch–Lipid Interactions and Their Effects on the Digestibility of Starch  K. G. Duodu and M. N. Emmambux

11.1 Introduction  11.2 Starch–Protein Interactions: Modulatory Effects on Starch Digestibility and Other Functional Properties  11.3 Starch–Lipid Complexes  11.3.1 Chemistry of Amylose–Lipid Complexes  11.3.2 Production of Amylose–Lipid Complexes  11.3.3 Nutritional Impact and Health Benefits of Starch–Lipid Complexes  11.4 Conclusions  References 

Chapter 12 Types and Distribution of Phenolic Compounds in Grains  Victoria Ndolo and Trust Beta

12.1 Introduction  12.2 Types of Phenolic Acids in Grains  12.3 Identification of Phenolic Acids in Grains  12.4 Total Phenolic Content in Whole Grains and Grain Fractions  12.4.1 Major Food Cereals (Wheat, Maize and Rice)  12.4.2 Minor Food Cereals (Barley, Sorghum, Millet, Rye and Oats)  12.5 Composition and Distribution of Phenolic Acids  12.5.1 Whole Grains  12.5.2 Botanical and Milling Fractions  12.6 Conclusions  References 

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Chapter 13 Bound Phenolic Constituents as Co-passengers of Dietary Fibre  Franklin Brian Apea-Bah and Trust Beta

13.1 Introduction  13.2 Chemistry and Biochemistry of Phenolic Compounds Bound to Dietary Fibre in Cereals  13.2.1 Chemistry of Phenolic Acids  13.2.2 Biosynthesis of Phenolic Acids  13.2.3 Dimerization of Hydroxycinnamates  13.3 Sample Preparation, Extraction and Analysis of Phenolic Compounds Bound to Dietary Fibre in Cereals  13.3.1 Sample Preparation and Extraction of Soluble and Insoluble Dietary Fibre  13.3.2 Extraction of Bound Phenolic Acids  13.3.3 Analysis of Phenolic Acids Bound to the Cell Wall in Cereals  13.4 Effects of Processing on Cereal-derived Phenolic Compounds Bound to Dietary Fibre  13.4.1 Removal of Botanical Parts  13.4.2 Solubilization and Leaching  13.4.3 Fermentation  13.4.4 Nixtamalization  13.4.5 Thermal Processing  13.5 Release and Metabolism of Cereal-derived Phenolic Compounds Bound to Dietary Fibre in the Colon  13.6 Beneficial Health Effects of Cereal-derived Phenolic Compounds Bound to Dietary Fibre  13.6.1 Dietary Fibre Antioxidants  13.6.2 In vitro Antioxidant Potential of Cereal Dietary Fibre Antioxidants  13.6.3 In vivo Antioxidant Potential of Cereal Dietary Fibre Antioxidants  13.7 Future Perspectives  References 

Chapter 14 Anthocyanins, Deoxyanthocyanins and Proanthocyanidins as Dietary Constituents in Grain Products Joseph Awika, Leonnard Ojwang and Audrey Girard

14.1 Introduction  14.2 Anthocyanins and 3-deoxyanthocyanins in Cereal Grains 

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14.2.1 Biosynthesis of Anthocyanins and 3-deoxyanthocyanins  14.2.2 Structure and Composition of Anthocyanins in Cereal Grains  14.2.3 Maize Anthocyanins  14.2.4 Anthocyanins in Rice  14.2.5 Anthocyanins in Wheat and Barley  14.2.6 3-Deoxyanthocyanins in Sorghum  14.2.7 Structure of Sorghum 3-deoxyanthocyanins and Basis for Color Stability  14.3 Proanthocyanidins (Condensed Tannins)  14.4 Health Benefits of Cereal Anthocyanins, 3-deoxyanthocyanins and Proanthocyanidins  14.4.1 Anthocyanins and 3-deoxyanthocyanins  14.4.2 Proanthocyanidins (Condensed Tannins)  References 

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Chapter 15 Interactions Between Grains and the Microbiome  S. Brahma and D. J. Rose

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15.1 Introduction  15.2 Grain Components Likely to Interact with the Microbiome  15.2.1 Dietary Fiber  15.2.2 Phenolic Compounds  15.2.3 Other Compounds  15.3 Whole Grain Intervention Studies  15.4 Responders/Non-responders to Whole Grain Interventions  15.5 Increasing Whole Grain–Gut Microbiota Interactions  15.6 Conclusions  References 

Subject Index 

333 334 339 341 342 342 350 350 351 357

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Chapter 1

Introduction T. Beta University of Manitoba, Department of Food and Human Nutritional Sciences, 250 Ellis Building, Winnipeg, Manitoba, Canada R3T 2N2 *E-mail: [email protected]

1.1  W  hole Grains as Delivery Packages for Nutrients and Phytochemicals Whole grains can be viewed as packages of major nutrients (carbohydrates, proteins and lipids), micronutrients (vitamins and minerals) and other phytochemicals. They are heterogeneous structures containing nutrients and phytochemicals distributed unevenly in the pericarp, aleurone, germ and starchy endosperm1–3 (Figures 1.1 and 1.2). Carbohydrates serve as energy sources in the form of digestible starch and simple sugars. The indigestible forms of carbohydrates include high molecular weight polysaccharides and low molecular weight oligosaccharides. More attention can be paid to their enhancement as dietary fibre in the diet given that several countries are witnessing burgeoning levels of obesity. Some dietary fibres already attract health claims, such as beta-glucan from oats and barley. Although indigestible carbohydrates increase the dietary fibre content of foods, they may at the same time serve as functional ingredients and are involved in the texturerelated modification of several foods.

  Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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Figure 1.1  Whole  grain package showing regions dominated by carbohydrates

(sugars, starch and fibre, mainly consisting of non-starch polysaccharides) and non-nutrient phytochemicals. The aleurone layer is part of the outer endosperm, but it is removed along with the outer layers as bran (see details of cytoplasmic organization in Figure 1.2).

The micronutrients in cereal grains include B vitamins (thiamine, riboflavin, niacin, pyridoxine, pantothenic acid and folates) and tocopherols.4 The B vitamins are highly concentrated in the aleurone layer and/or the scutellum part of the germ.5 Whole grains also contribute to the overall mineral content of human diets,6 although the location and/or accumulation of minerals within the grain tissue may influence their dietary availability. Mineral elements include K, Mg, P, Ca, Zn, Fe, Cu, Mn and S.7 Phytochemicals include carotenoids, which are subdivided into hydrocarbons (β-carotene and α-carotene) and their oxygenated derivatives or xanthophylls (lutein, zeaxanthin and β-cryptoxanthin).8 They display provitamin A activity and antioxidant properties. Lutein and zeaxanthin are predominant as minor constituent in cereals,9 whereas β- and α-carotene are found in minimal amounts.10 Lutein and lutein esters make up >90% of the yellow pigment in wheat.11 Phenolic compounds, the main phytochemicals found in whole grains, include phenolic acids, flavonoids, anthocyanidins and phytosterols.12 The phenolic acids present in grains are mostly bound to cell wall polysaccharides as part of dietary fibre. Vitaglione et al.13 asserted that the slow and continuous release of antioxidants bound

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Figure 1.2  Digital  images showing (A) the pericarp, aleurone layer and endosperm

of a cross-section of a barley cereal grain and the concentration of ferulic acid (a major phytochemical) in the cell walls of the aleurone layer; (B) stained wheat aleurone residue with stained starch granules and fragments of empty and filled aleurone cells; and (C) wheat aleurone layer with empty and *filled aleurone cells (the cells are filled with bodies containing protein, phytin and niacin deposits and other B vitamins). Figure 2A reprinted from V. U. Ndolo, T. Beta and R. G. Fulcher, Ferulic acid fluorescence intensity profiles and concentration measured by HPLC in pigmented and non-pigmented cereals, Food Res. Int., 52 (1), 109–118., Copyright 2013, with permission from Elsevier. (B) and (C) reprinted from V. U. Ndolo, R. G. Fulcher and T. Beta, Application of LC–MS–MS to identify niacin in aleurone layers of yellow corn, barley and wheat kernels, J. Cereal Sci., 65, 88–95, Copyright 2015, with permission from Elsevier.

to dietary fibre in the gut determines their nutritional benefits. Ferulic acid is a dominant phenolic antioxidant in grain. This bifunctional molecule, with both carboxylic and phenolic bonding sites, provides a pathway for intra- and intermolecular cross-linking between polysaccharides or between polysaccharides and lignin. This cross-linking is important in the structure of arabinoxylans, for example, causing changes in their solubility in aqueous environments and accessibility for enzymatic hydrolysis by microorganisms in the human colon. Cross-linked phenolic acids change the three-dimensional structure of the polymer, often preventing localized areas from swelling and excluding degradation enzymes.14 Although diferulates are the major cross-linking agents between cell wall biopolymers in cereal dietary fibre,15 sinapate–ferulate cross-products have also been discovered.16

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1.2  Whole Grains and Health Promotion Whole grain cereals deserve more attention as rich sources of bioactive health-promoting compounds17 given their status as a staple part of the human diet relative to fruit and vegetables. The literature indicates that their consumption is associated with a reduced incidence of chronic diseases (such as type 2 diabetes, cardiovascular diseases, obesity and cancer), which are major causes of death and morbidity.18 These protective effects are attributed to a number of bioactive chemical compounds (phytochemicals), which are mostly present in the bran, aleurone and germ fractions of whole grains. Although the protective mechanisms are complex and multifactorial (involving increased satiety, reduced transit time of food in the gut, improved faecal bulking and reduced glycaemic response), antioxidant and anti-inflammatory actions appear to be key factors in the prevention of chronic disease.19 This is because antioxidants prevent free radical damage to proteins (e.g. enzymes and hormones), lipids (e.g. the phospholipid bilayer of cell membranes and low-density lipoproteins), carbohydrates, DNA and other cellular organelles, which leads to inflammation, cell death and, as a consequence, debilitating chronic disease. Research conducted by our group has confirmed that whole grains are excellent sources of biologically active compounds, such as dietary fibre, polyphenols and carotenoids,20,21 which have antioxidant health-promoting properties. The combination of these bioactive compounds could potentially result in overall positive health effects.19

1.3  D  igestion, Bioaccessibility and Bioavailability of Whole Grain Nutrients and Phytochemicals The nutrients and phytochemicals in grain products need to be both bioaccessible and bioavailable on digestion to provide nourishment and for their beneficial health effects to be realized. Bioaccessibility describes the amount of nutrients and other phytochemicals potentially absorbable from the lumen. Bioavailability, in turn, incorporates bioaccessibility, absorption, tissue distribution and bioactivity.22 Exogenous factors affecting bioavailability include the complexity of the grain matrix, the chemical form of the compound of interest, and the structure and amount of co-ingested compounds.23 The amount of phytochemicals released during the digestion of food, and subsequently absorbed into the blood, therefore differs among compounds and also depends on the food matrix and the type of interactions between the phytochemicals and major (carbohydrates, proteins and lipids) or minor nutrients in the food. For instance, ferulic acid is efficiently absorbed when in the free form, such as in beer, but poorly absorbed when bound to arabinoxylans in the cell walls of cereals.24 Ferulic acid and its dehydrodimers (diferulates) are important components of the cereal dietary fibre matrix. Figure 1.3 shows how ferulic acid is involved in grouping together

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Figure 1.3  Possible  linkages between dietary fibre components and hydroxycin-

namic acids. Reprinted from B. B. Buchanan, R. L. Jones and W. Gruissem, Biochemistry and Molecular Biology of Plants, John Wiley and Sons, Hoboken, 2nd edn, 2015, Chapter 2, The Cell Wall, 73.33

non-starch polysaccharides (mainly arabinoxylans) and integrating with lignin through ether or ester linkages. There is, however, little information about the interactions existing among the phytochemicals and also among the phytochemicals and nutrients present in complex food matrices. Even though epidemiological studies indicate that the consumption of whole grains protects against several chronic diseases,25 the mechanisms by which grains confer health benefits are still to be fully established, taking into account issues related to digestibility, bioaccessibility and the bioavailability of nutrients and phytochemicals. There has been a steady growth and expansion of the functional food and nutraceutical market as a result of consumer awareness and demands for

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healthy diet options, as well as advances in food processing and the development of functional foods.26 This provides an opportunity for food scientists and technologists to design and produce new and innovative food products for consumers of all ages using guidance from genetic studies and to contribute to the scientific literature on food processing, bioaccessibility and bioavailability.

1.3.1  Genetic Studies Genetic studies on rice using quantitative trait locus, association and genome-wide mapping techniques have led to the identification of the genes responsible for red and brown pericarp coloration in whole grain rice. These pericarp pigments are related to flavonoids and proanthocyanidins, which impart antioxidant properties to the pigmented whole grain rice.27,28 Because some phytochemicals, such as phenolic compounds, are produced in high amounts in plants in specific environments,27 such information can be used for the selection and breeding of new varieties with enhanced phytochemical contents, via biofortification (genetic engineering), for utilization in the development of novel grain-based functional foods.

1.3.2  Food Processing Several traditional and non-traditional methods (steeping, malting, roasting, fermentation, boiling, baking, frying, nixtamalization, micronization and extrusion) are used to process whole grains into foods. The preparation of food crops into finished food products usually requires a combination of these methods. Our studies have shown that different processing methods affect the total phenolic content, phenolic composition and antioxidant activity of wheat, barley, oat and their mill fractions.1,3,29 Heating wheat bran at 177 °C for 20 minutes did not have a consistent effect on its total phenolic content and antioxidant properties. However, pearling (debranning) appears to improve the roller-milling performance of wheat, concentrating the constituent phenolic compounds in the bran. Although the pearled roller-milled wheat flour may have a relatively low phenolic content, the removed bran can be used as a functional food ingredient. There are, however, gaps in the documentation of the effects of processing methods on the phytochemical composition and health-promoting properties of each whole grain product. These methods – including the extrusion cooking, pressure steaming and canning used in the product development of whole grains – require further investigation to determine the process stability of the free and bound phenolic compounds, vitamins, carotenoids, dietary fibre components and other phytochemicals present in whole grains, as well as the effects of their interactions with each other and with nutrients and non-nutrients on their potential health-promoting properties.

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1.3.3  Bioaccessibility and Bioavailability Research conducted in our laboratory has shown that simulated in vitro gastrointestinal digests from purple wheat bread exhibit antioxidant properties.30 Few other reports are available on the simulated gastrointestinal digestion of whole grain foods and almost all other methods have used static, manually controlled systems for simulated in vitro gastrointestinal digestion. However, the TIM-1 system is an example of a platform using a dynamic, fully automated system that more accurately simulates the physiological processes and conditions in an in vivo upper gastrointestinal system than static systems. It can also be adapted to simulate conditions for young, adult and elderly humans and other mammals.31 Such platforms are useful for studying the stability, release and bioaccessibility of the nutrients and other phytochemicals present in whole grain product matrices after simulated gastrointestinal digestion. Fewer studies have demonstrated the release, bioaccessibility and healthpromoting properties of phytochemical metabolites from grain products after colonic fermentation.32 The metabolites released depend on the food type and the phytochemicals present. Various processing methods affect the constituent phytochemicals in foods differently, leading to digests with different phytochemical compositions. There are gaps in our knowledge of the effects of gastrointestinal digestion on whole grain products and also on the effect of colonic fermentation on the bio-inaccessible phytochemical components in the digests. Although the bioaccessible components will be potentially absorbed from the colon into the systemic circulation, imparting health benefits in the target organs and tissues, the bio-inaccessible portion will remain in the colon for an extended period before excretion in stools. The bio-inaccessible phytochemicals can therefore potentially contribute to the overall health status of the colorectal region of the gut.

1.4  Purpose of this Book This book is intended to be an update on carbohydrates and phytochemical components in whole grain and grain-based foods. Detailed coverage is provided of the composition and functionality of the bulk carbohydrate components (sugars, oligosaccharides and polysaccharides, including starch, beta-glucans and arabinoxylans) and the minor phytochemical components (phenolic compounds, such as phenolic acids, anthocyanins, deoxyanthocyanins and proanthocyanidins) in cereal grains and grain products. Considerable attention is paid to the interactions between the carbohydrate and non-carbohydrate components in grains, including starch–protein and starch–lipid interactions and their effects on the digestibility of starch. The phenolic constituents bound to grain dietary fibre also deserve attention. Chapter 15 presents a review of whole grain–gut microbiota interactions and identifies new areas of research, which may contribute to a better understanding of the underlying mechanisms linked to human health. With the

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Figure 1.4  Functional  foods based on cereal grains: carbohydrate and phytochemical components. DF-PA, phenolic acids linked to dietary fibre; NSP, non-starch polysaccharides; RS, resistant starch; SDS, slowly digestible starch.

exception of the introduction and processing overview, the contents of this book have been divided into carbohydrates and phytochemicals, as shown in Figure 1.4. Chapter 2 provides an overview of the inevitable post-harvest grain processes that transform raw seeds into edible products suitable for human consumption. The processes are classified into those requiring minimal heat and those that are thermal techniques. The latter are further divided into relatively dry and wet processes. Chapter 3 documents the changes in the composition of sugars and oligosaccharides in three different states: (1) during kernel development; (2) at grain maturity; and (3) during storage and processing. During processing, sugars and oligosaccharides can either be generated or used up as substrates. Future research trends are included for these low molecular weight carbohydrates. Chapter 4 focuses on the major polysaccharides (arabinoxylans, mixedlinked beta-glucans, and arabinogalactan peptides) of wheat, rye and barley and their interactions in endosperm cell walls. Heteroxylans are included as polysaccharides of the outer tissues, along with cellulose and lignin and their possible interactions. Chapter 5 introduces starch as a complex entity, the properties of which have been measured using various approaches in an effort to establish structure–function relationships that can then be used to select suitable plant

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varieties or modify the starches. The need to understand starch and starch interactions is highlighted. Chapter 6 considers the definition of dietary fibre and its evolution. Details of the rapid integrated total dietary fibre (RINTDF) method (AOAC Method 2009.01) are provided because it can be applied to all food samples, including whole grains and grain products. Chapter 7 provides a general coverage of resistant starch and slowly digestible starch in grain products, with a focus on resistant starch synthesis in grains, resistant starch formation in processed grain products and the potential of resistant starch as a functional ingredient due to its ease of incorporation into food products. Chapter 8 discusses the functionality of beta-glucan from oat and barley as it relates to health. The health benefits associated with the lowering of cholesterol, blood glucose and blood pressure and the improvement of gut microbial populations are explained, along with their possible mechanisms. The physicochemical properties of beta-glucans, including viscosity, determine their functional properties. Chapter 9 is devoted to arabinoxylans, their molecular structures and physicochemical properties, distribution, and changes in their content and properties in whole grains, milling fractions and grain products. Arabinoxylans can be isolated for further studies of their role as prebiotics, their effect on glucose metabolism, and their immunomodulatory and antioxidant properties. Our need to understand the mechanism by which arabinoxylans exert many health benefits is highlighted. Chapter 10 identifies the major non-digestible oligosaccharides in cereal grains and explains the biosynthesis, structure and distribution of fructans and raffinose family oligosaccharides. This chapter emphasizes the role of non-digestible oligosaccharides in raw and processed products. Arabinoxylan oligosaccharides are also included as potential low-cost prebiotics. Chapter 11 focuses on starch–protein and starch–lipid interactions and their modulatory effects on starch digestibility. The chapter includes the chemistry, production (using classical methods, such as steam jet cooking, extrusion, wet heat processing), nutritional impact and benefits of amylose– lipid complexes. There is potential for the use of starch–lipid complexes as bioactive encapsulants. Chapter 12 is conveniently divided into a discussion of the composition and distribution of phenolic acids in major and minor food cereals as whole grains and as grain fractions. The chapter considers the type of phenolic acids in grains and the methods used for the identification of individual acids. Chapter 13 discusses the bound phenolic constituents as co-passengers of dietary fibre, starting with an overview of the chemistry, biosynthesis, preparation and extraction of phenolic compounds bound to dietary fibre in cereals. A summary is provided on the effects of milling, solubilization and leaching, fermentation, nixtamalization and thermal processing. Their release, metabolism and potential beneficial effects in vitro and in vitro are also considered.

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Chapter 14 covers the biosynthesis and structure of anthocyanins in maize, rice, wheat and barley, and deoxyanthocyanins in sorghum. Proanthocyanidins are considered as relatively rare in cereal grains, except for sorghum, finger millet, barley and rice. A summary of the growing evidence for the potential health benefits of anthocyanins, deoxyanthocyanins and proanthocyanidins is provided. Chapter 15 gives an updated account of whole grain–gut interactions, with special reference to non-digestible carbohydrates (arabinoxylans (AX), beta-glucans, cellulose, fructans and resistant starch) and the phenolic components of cereal grain-based foods, and how increasing the accessibility of carbohydrates to microbiota could be beneficial.

1.5  What This Book Does not Set Out to Do This book is not intended to provide a comprehensive coverage of the interactions among carbohydrate or phenolic components and micronutrients. These interactions are becoming increasingly important due to the prevalence of micronutrient deficiencies in certain segments of the population in both developed and less developed countries. Such interactions merit further investigation and a separate book could be published on this topic.

References 1. T. Beta, S. Nam, J. E. Dexter and H. D. Sapirstein, Cereal Chem., 2005, 82, 390–393. 2. V. U. Ndolo and T. Beta, Food Chem., 2013, 139, 663–671. 3. V. U. Ndolo and T. Beta, Cereal Chem., 2014, 91, 522–530. 4. A. S. Sandberg and R. Ahderinne, HPL method for determination of inositol tri-, teter-, penta- and hexaphosphates in foods and intestinal contents, J. Food Sci., 1986, 51, 547–550. 5. J. Delcour and R. C. Hoseney, Principles of Cereal Science and Technology, AACC International, St Paul, MN, USA, 2010. 6. B. L. O'Dell, A. R. de Boland and R. K. Samuel, J. Agric. Food Chem., 1972, 20, 718–721. 7. S. O. Serna-Saldivar, Cereal Grains: Properties, Processing and Nutritional Attributes, New York, CRC Press, 2010. 8. R. H. Liu, J. Nutr., 2004, 34, 3479S–3485S. 9. M. N. Irakli, V. F. Samanidou and I. N. Papadoyannis, J. Sep. Sci., 2011, 34, 1375–1382. 10. M. Heinonen, V. Ollilainen, E. Linkola, P. Varo and P. Koivistoinen, Cereal Chem., 1989, 66, 270–273. 11. M. Lepage and R. P. A. Sims, Cereal Chem., 1968, 45, 600–604. 12. F. Shahidi and M. Nasck, 1995. Food Phenolics: Sources, Chemistry, Effects, and Application, Technomic Publishing Company, Inc., Lancaster, PA, 1995, 331 pages.

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13. P. Vitaglione, A. Napolitano and V. Fogliano, Cereal dietary fibre: a natural functional ingredient to deliver phenolic compounds into the gut, Trends Food Sci. Technol., 2008, 19, 451–463. 14. R. D. Hatfield, Cell wall polysaccharide interactions and degradability, in Forage Cell Wall Structure and Digestibility, ed. H. G. Jung, D. R. Buxton, R. D. Hatfield and J. Ralph, ASA–CSSA–SSSA, Madison, WI, 1993, pp. 286–314. 15. A. Renger and H. Steinhart, Eur. Food Res. Technol., 2000, 211, 422–428. 16. J. Ralph, M. Bunzel, J. M. Marita, R. D. Hatfield, F. Lu, H. Kim, P. F. Schatz, J. H. Grabber and H. Steinhart, Phytochem. Rev., 2004, 3, 79–96. 17. R. H. Liu, J. Cereal. Sci., 2007, 46, 207–219. 18. PHAC How Healthy Are Canadians? A Trend Analysis of the Health of Canadians from a Healthy Living and Chronic Disease Perspective, Public Health Agency of Canada, Ottawa, ON, Canada, 2016, pp. 1–39. 19. A. Fardet, Nutr. Res. Rev., 2010, 23, 65–134. 20. W. Guo and T. Beta, Food Res. Int., 2013, 51, 518–525. 21. V. U. Ndolo and T. Beta, Food Chem., 2013, 139, 663–671. 22. N. M. Anison, R. van den Berg, R. Havenaar, A. Bast and G. R. M. M. Hainen, J. Cereal. Sci., 2009, 49, 296–300. 23. B. Holst and G. Williamson, Curr. Opin. Biotechnol., 2008, 19, 73–82. 24. C. Manach, G. Williamson, C. Morand, A. Scalbert and C. Rémésy, Am. J. Clin. Nutr., 2005, 81, 230S–242S. 25. J. Slavin, Nutr. Res. Rev., 2004, 17, 99–110. 26. S. K. Basu, J. E. Thomas and S. N. Acharya, Aust. J. Basic .Appl. Sci., 2007, 1, 637–649. 27. Y. Shao, F. Xu, Y. Chen, Y. Huang, T. Beta and J. Bao, Cereal Chem., 2015, 92, 204–210. 28. F. Xu, J. Bao, T.-S. Kim and Y.-J. Park, J. Agric. Food Chem., 2016, 64, 4695–4703. 29. W. D. Li, M. D. Pickard and T. Beta, Food Chem., 2007, 104, 1080–1086. 30. L. Yu, Identification and Antioxidant Properties of Phenolic Compounds during Production of Bread from Purple Wheat Grains and Investigation of Bread Extracts after Simulated Gastrointestinal Digestion, Master of Science (Food Science), University of Manitoba, Winnipeg, Manitoba, Canada, 2014. 31. M. Minekus, The TNO gastro-intestinal model (TIM), in The Impact of Food Bio-Actives on Gut Health - In Vitro and Ex Vivo Models, ed. K. Verhoeckx, P. Cotter, I. López-Expósito, C. Kleiveland, T. Lea, A. Mackie, T. Requena, D. Swiatecka and H. Wichers, Springer, 2015, pp. 37–46. 32. A. Chandrasekara and F. Shahidi, J. Funct. Foods., 2012, 4, 226–237. 33. B. B. Buchanan, R. L. Jones and W. Gruissem, Biochemistry and Molecular Biology of Plants, John Wiley and Sons, Hoboken, 2nd edn, 2015, Chapter 2, The Cell Wall, 73.

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Chapter 2

Overview of Grain Components and Changes Occurring in Grain Constituents with Different Forms of Processing Mary Ellen Camire School of Food and Agriculture, University of Maine, 5735 Hitchner Hall, Orono, Maine, USA *E-mail: [email protected]

2.1  Introduction Cereal grains are an important food source in most cultures and the global food trade allows many types of cereals to be available for consumption. Although many plant foods may be consumed raw, cereals require transformations to become edible. Neanderthals and other early humanoids processed and consumed grass seeds.1 Archeological evidence indicates that sorghum was ground by residents of Mozambique about 100 000 years ago.2 Humans began cultivating wild grasses >10 000 years ago and selected for plants that had non-shattering heads and larger seeds.3 Over the centuries, diverse cultures have created unique cereal-based foods and beverages, but most processes involve some breaking of the exterior bran and gelatinization   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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of the starchy endosperm. The bran protects the grain germ by serving as a barrier to digestion. The crystalline nature of cereal starches is inherently difficult for digestive enzymes to attack, but physical modifications, such as gelatinization, facilitate the activities of mammalian carbohydrases.4 Phenolic acids and other bioactive active components may be bound to cell wall carbohydrates and processing may solubilize these phytochemicals, potentially making them more bioavailable. Following processing, these compounds may simply become more resistant to extraction,5 although some studies have been unable to distinguish inextractability from a reduction or loss of these compounds. The fate of these resistant phytochemicals in the digestive tract is a topic of great interest. Fardet6 reviewed the physicochemical traits of cereals and how they may change with processing and recommended the development of improved models for in vitro digestion to help researchers better understand the interactions of nutrients and phytochemicals within the gastrointestinal tract. The effects of cereal consumption on appetite, satiety and mood are other health effects that processing can modulate. The broad review of processing methods and their effects on grains in this chapter serves as a foundation for subsequent chapters focused on specific categories of beneficial compounds. Although most nations agree that the consumption of whole grains promotes health, defining whole grains for food labeling purposes has been more controversial. The cereal science association AACC International published a definition in 1999 that was further modified by other organizations.7 The European Union HEALTHGRAIN project (FP6-514008) adopted this definition of whole grains:8    Whole grains shall consist of the intact, ground, cracked or flaked kernel after the removal of inedible parts such as the hull and husk. The principal anatomical components – the starchy endosperm, germ and bran – are present in the same relative proportions as they exist in the intact kernel. Small losses of components – that is, less than 2% of the grain/10% of the bran – that occur through processing methods consistent with safety and quality are allowed.    Several nations permit health claims for whole grain foods as a means to encourage consumers to eat whole grains.7,8 The development of convenient, healthful and appealing whole grain products requires an understanding of both grain composition and the effects of processing on the grain. Another health-related cereal topic is the production of gluten-free foods for people with celiac disease and non-celiac gluten sensitivity. The incidence of these conditions has increased.9 Wheat and related grains (barley, triticale and rye) contain gluten. Oats do not naturally contain gluten, but gluten-containing grains may be present as a contaminant.10 As more consumers adopt gluten-free diets, several nutritional deficiencies have been reported.11,12 Gluten-free diets may be low in dietary fiber.12 Therefore gluten-free cereals could play an important part in filling the fiber gap for people who are

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avoiding wheat and gluten. O'Shea et al. summarized the technological issues in designing acceptable gluten-free foods.

2.2  Pre-processing Steps A series of processes may be applied depending on the type of grain and its end use. The ease of removal of the hull differs among species. Although the hulls of most grains can be easily separated during threshing, rice and most barley and oat cultivars have tightly adhering hulls.15 Grains should be dried to reduce the likelihood of microbial contamination, especially molds, which may produce mycotoxins. This step is particularly important in warm, humid climates. Heat- and oxygen-labile compounds, such as phenolic acids, may decrease in quantity during storage.16

2.2.1  Cleaning Harvested grains may arrive at mills mixed with natural products (e.g. stems, husks, stones and insects) or human-made contaminants, such as glass and metal, may be present. Air classification can separate some of these undesirable materials. Magnets can remove most metals. Commercial operations use many types of cleaners, sorters and scoring equipment. Although cleaning steps are important for food safety, they have relatively little impact on nutritional quality.

2.2.2  Tempering and Soaking Grain kernels naturally vary in moisture content, so mills use a process called tempering, which hydrates all the wheat or rye kernels to similar moisture levels, such as 16% for wheat.17 The adjusted moisture level aids the separation of bran from the endosperm during milling. Tempering is a short process because the over-hydration of kernels could stimulate starch-degrading enzymes. Significant changes in nutrients are unlikely during this step. Maize, millet and rice have been reported to lose significant amounts of phytate after soaking for 24 hours, although sorghum did not.18 Lestienne et al.18 concluded that soaking alone did not enhance mineral bioavailability in cereals. Concerns about arsenic levels in rice have stimulated research to reduce the concentration of this toxic element. Yim et al.19 stirred, rinsed and soaked rice to assess the removal of arsenic while retaining nutrients. The process had little effect on thiamine, but decreased dietary fiber in white rice; the opposite trend was found for brown rice.

2.2.3  Dehulling and Pearling Barley and rice are the major cereals requiring hull (husk) removal; other grains have hulls that are more easily removed during threshing.15 The high dietary fiber and mineral content of hulls limit their use as human food.

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For example, rice hulls contain 70% dietary fiber, largely in the form of lignin and cellulose, and silica, which represents 95% of the ash content.15 Rice hulls are removed by abrasive rollers to produce whole grain brown rice and then the bran and aleurone are removed by abrasive pearling to produce white rice. Impact and abrasion dehullers are commonly used to remove barley hulls.20,21 Hull-less barley varieties have been introduced, but hulled types still dominate commercial trade. Whole grain barley has only the hull removed, accounting for 11–12% of the kernel weight, and an additional 6, 11–12 and 20–22% of the kernel weight is abraded for commercial pearled, pot pearled and white barley, respectively. White barley contains no bran, germ or crease.22 Pearl barley is not considered to be a whole grain, but the endosperm is nonetheless rich in healthful beta-glucans.7 As phenolic compounds are concentrated in the outer layers of grains, these phytochemicals have largely been removed in pearled grains, such as barley.16 Giordano and colleagues23 proposed that the pearling of colored wheat might be a method of concentrating anthocyanins for other food uses.

2.2.4  Milling The process of milling reduces the particle size of grains and separates seed fractions with different compositions and physical properties. The bran and aleurone layer are typically rich in water-soluble compounds, whereas the germ is rich in lipid-soluble components. Hemery et al.24 summarized the concentration of functional components in wheat tissues. Tocopherols, tocotrienols, B vitamins, phytic acid and lipids are most likely found in the aleurone and germ; minerals are found in all tissues except the endosperm. In traditional whole grain milling, the entire kernel is crushed by granite or other hard stones, or by steel rollers. In the twentieth century, commercial roller mills separated the different grain fractions (for example, bran, endosperm and germ) and then recombined them on-site (recombination) or at another site (reconstitution).7 The HEALTHGRAIN,8 AACC International7 and other definitions of whole grain permit up to 10% of the outer bran (and not more than 2% of the entire kernel) to be removed because microorganisms, mycotoxins (such as vomitoxin and aflatoxin), heavy metals (arsenic, cadmium, lead and mercury) and agricultural chemicals (pesticides and fungicides) can be concentrated in this outermost layer.25 Lipid oxidation is a concern for traditionally milled flour because native lipids from the germ are intermixed with enzymes and other grain constituents. Stoneground flours may require refrigeration or other methods to slow lipid oxidation via enzymatic and chemical pathways. Phenolic compounds in the germ and bran may be removed during milling16 and thus would be unable to serve as antioxidants in wholegrain flour. A consortium of organizations concluded that milling processes that separate and recombine cereal tissues do not substantially affect the nutritional quality and may enhance shelf life.7

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Roller milling results in a greater retention of carotenoids, such as lutein in yellow wheat.23 Degermination has little effect on the carotenoid concentrations in maize because the germ has a relatively low concentration of these phytochemicals.26,27 For barley, oats and wheat, the removal of the carotenoid-rich germ can significantly reduce the total amount of carotenoids in flour.27 A comparison of a dry degermination process with degermination preceded by tempering found no difference in the total polyphenol, xanthophyll and total dietary fiber concentrations in maize germ and other products, such as flour and grits, but higher total antioxidant levels were detected in the tempered degerminated grits.28 The HEALTHGRAIN project assessed commercial milling procedures to develop recommendations for greater phytonutrient retention. Separation of the nutraceutical-rich aleurone layer can improve the nutritional value of recombined wholegrain flours.29 Dry processing methods to isolate and concentrate specific nutrients and other bioactive compounds in wheat have been proposed.25 Table 2.1 lists the nutrient and phytochemical losses during wheat milling. Wet milling is utilized to separate cereal components, such as starch, protein, oil and dietary fiber. Maize, wheat and rice are commonly wet milled.15 Water-soluble nutrients can be lost in the extraction medium, but the focus of wet milling is ingredient production for which the nutritional composition may not be important. By-products such as corn germ are often used for animal feed. The material and energy costs of wet milling could limit further product and process development.30

2.3  Processing with Minimal Heat 2.3.1  Sprouting (Germination) Germination is a natural process that occurs when seeds are exposed to the appropriate conditions for plant growth. Consumer interest in minimally processed foods is growing. Between 2006 and 2016, new products labeled as “freshly sprouted”, “germinated”, “sprout” or “sprouted” grew more than five-fold.31 Sprouted grains may be consumed directly or further processed by malting or fermentation. The AACC International Board of Directors approved the following definition of sprouted grains in 2008:    Malted or sprouted grains containing all of the original bran, germ, and endosperm shall be considered whole grains as long as sprout growth does not exceed kernel length and nutrient values have not diminished. These grains should be labeled as malted or sprouted whole grain.32    Exposure to warmth, light and moisture induces the release of numerous enzymes that hydrolyze macromolecules.33 Five general steps take place during germination.34 First, the seed absorbs water, signaling production of the hormone gibberellin, which turns on the expression of genes for

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Table 2.1  Nutrient  and phytochemical losses during wheat milling. Grain component

Grain tissue source

Vitamins ●● Thiamine ●● Riboflavin ●● Pyridoxine ●● Folate ●● α-Tocopherol Magnesium Manganese Ferulic acid Sterols Carotenoids Starch Vitamins ●● Niacin ●● Pantothenic acid ●● Pyridoxine ●● Biotin ●● Folate Magnesium Phosphorus Xylans Beta-glucans Ferulic and other phenolic acids Phytic acid Lignans Anthocyanins Carotenoids Arabinoxylans Cellulose Xylans Lignin Phenolic acids Alkylresorcinols

Germ

17

a

Endosperm Aleurone

Bran

a

Data from ref. 15, 25 and 29.

important hydrolytic enzymes. Next, gibberellin migrates from the germ to the aleurone layer; enzymes that hydrolyze starch, nonstarch polysaccharides and protein become activated in the endosperm as enzyme inhibitors are suppressed. The freed sugars, amino acids and lipids then provide fuel for the developing root and shoot in the germ. Singh and Sharma35 reviewed the changes in bioactive components during germination (Table 2.2). The germination of sorghum for malt was limited by the growth of pathogenic bacteria and molds, with risks for the development of mycotoxins. Soaking the seeds in dilute sodium hydroxide, calcium hydroxide or sodium hypochlorite reduced the incidence of these problems.36,37 Donkor et al.37 reported species-specific differences in the proximate analyses of seven grains (barley, brown rice, buckwheat, oat, rye, sorghum and wheat). Methanol extracts of germinated brown rice and oats

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Table 2.2  Nutritional  changes during grain sprouting.

a

Grain component

Change

Starch

Depolymerization into smaller fragments, leading to increase in reducing sugars and oligosaccharides Increased Decreased Increased solubility Release of free amino acids Variable Increased bioavailability Increased folate, niacin, riboflavin, thiamine and vitamin C Mixed response for tocopherols Increased Decreased Decreased Variable changes: species- and process-dependent

Arabinoxylans Beta-glucans Protein Lipids Minerals Vitamins γ-Aminobutyric acid Tannins Phytic acid Phenolic acids a

Data from ref. 33–35 and.37.

had lower α-glucosidase activity than their nongerminated counterparts, but all other grains had higher activity after germination. In the same study, α-amylase significantly increased after germination for all grains. Traditional malt production begins with barley soaking (steeping), followed by germination and then kilning/roasting to stabilize the product and develop flavors and colors. Carvalho et al.38 reviewed the steps in malting and their contributions to the formation of antioxidants and pro-oxidants. Ancient grains such as einkorn, spelt and Kamut have been malted for beer, but differences in cereal composition affect the final composition and quality of the beverages.39 White sorghum steeped in 0.2% calcium hydroxide, germinated, then ground and dried had less flour yield than whole sorghum, but the product was significantly lower in oxalates, phytates and tannins.40 Hydrogen cyanide levels were not affected by malting.

2.3.2  Fermentation Fermentation is an ancient form of processing and continues to evolve with consumer demand and new technologies. Fermentation preserves foods by lowering the pH, using competitive microbial species or forming ethanol; the microbes may create an improved nutritional profile41 (Table 2.3). Cracked or ground grains are mixed with water and sometimes other ingredients as a first step. Nout42 reviewed the diverse foods and beverages (alcoholic and non-alcoholic) produced from cereals in Africa and Asia. Several types of millet are fermented in Africa to yield beverages and foods.36 Spontaneous or natural fermentation is practiced in many areas. Grain mixtures may be left uncovered to allow inoculation by wild bacteria, usually lactic acid bacteria, molds or yeasts. For example, sourdough bread

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Table 2.3  Improved  nutritional quality in fermented cereals.

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a

Improved digestibility

Increased bioavailability

Starch

Iron and zinc (via reductions in phytate) Phenolic acids (via solubilization)

Protein

Formation of nutrients and other bioactive compounds Thiamine Folate Pyridoxine Riboflavin Ascorbic acid Tocopherols γ-Aminobutyric acid Bioactive peptides

a

Data from ref. 41.

can be fermented by a variety of lactic acid bacteria species and yeasts, but the flora will vary depending on the type of grain, geographical location, method of inoculation and many other factors.43 The process can be accelerated by mixing a small amount of already fermented material in a practice called backslopping. The fermentation time was reduced by 50–75% and the thiamine content was nearly 50% higher in idli (an Indian fermented rice and legume batter) when the mixture was backslopped with 10 or 25% previously fermented idli.44 Phytate and trypsin inhibitors were significantly lower in the backslopped idli, further improving its nutritional value. The B vitamin folate is essential in preventing many health problems, including neural tube birth defects. Lactic acid bacteria have been proposed as a means to increase the folate content of fermented foods, particularly those based on cereals.45 Only the soaking phase of ben-saalga porridge from pearl millet raised folate concentrations; subsequent processing steps led to losses of the vitamin.24 Backslopping did not improve the folate content. In their review of bioactive compounds in fermented seeds, Gan et al.41 summarized the effects of fermentation on the production of γ-aminobutyric acid, phenolic acids and bioactive peptides. Commercial fermentation usually depends on the more controlled addition of selected microorganisms to select for specific fermentation products and to exclude pathogens, toxins and microbes that could reduce product acceptability.46 For example, acetic acid bacteria may contribute undesirable vinegar-like flavor in some food products, but brewers intentionally inoculate these bacterial species for the production of lambic beers.47 De Vuyst et al.48 reviewed the roles of starter cultures and other processing variables in the production of three classes of sourdough fermentation. Fermented beverages are consumed around the world and thus contribute to nutrient intakes. Bamforth49 summarized the nutritional benefits of beer and its components. Brewing styles alter the vitamin content and various treatments after brewing can affect nutritional values. Beer pasteurization

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reduced thiamine, and riboflavin was destroyed by the addition of silica and polyvinylpolypyrrolidone.50 The use of isinglass, bentonite, tannic acids (as a stabilizing agent) and potassium metabisulphite (as an antimicrobial agent) reduced all forms of thiamine and riboflavin in experimental beer.50 Beer and hop polyphenols might alleviate some problems associated with the menopause, but consumers should weigh the benefits of beer versus concerns about alcohol consumption.51 The residue from beer and related fermentations is called brewers' spent grain (BSG). The growth of microbreweries has increased the production of BSG. Although much of the starch and some soluble nutrients are removed during the fermentation and brewing processes, other healthful compounds remain and some are added from the yeast. Ikram et al.52 reviewed the composition of BSG and concluded that the dietary fiber, protein and phenolic acids could be utilized for animal feed or in food for humans. In this author's home town, BSG is used for dairy cattle feed, in dog treats and in pretzels served at a brewpub. Further utilization of this brewing by-product is likely.

2.3.3  Pasta and Noodle Production These products are commonly made from wheat, but other cereals may be used. Durum wheat is the classic basis for pasta. Pasta and noodles are often boiled before eating and the effects of boiling are discussed in Section 2.4.2. Semolina (for pasta) and flour (for noodles) are mixed with water without the inclusion of air and forced through small openings in a pasta machine or extruder to achieve the desired size and shape. Fresh pasta and noodles are consumed immediately, refrigerated for short-term storage or frozen. Most commercial pasta is dried.15 Asian noodles are also fried to create shelf-stable ramen.15 Cubadda et al.53 assessed the mineral content of laboratory-made and commercial pasta samples and found that magnesium and zinc were retained less than other minerals after milling. Cooking the pasta reduced most minerals further. Bran particles interfere with pasta processing. Micronizing whole durum wheat resulted in a higher retention of the total antioxidant capacity and total phenolic acids than was found for traditionally processed durum wheat.54 Figure 2.1 illustrates the changes in phenolic acid content at different steps of pasta preparation.

2.4  Thermal Processing with Water 2.4.1  Nixtamalization The niacin in maize is not very bioavailable and thus reliance on maize as a dietary staple can lead to the niacin deficiency disease pellagra. The early inhabitants of Mexico discovered that cooking and then soaking whole

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Figure 2.1  Changes  in total phenolic compounds as durum wheat is transformed into pasta. Data from ref. 54.

Table 2.4  Compositional  changes in maize due to nixtamalization.a Apparent losses during processing Dietary fiber Phytic acid Thiamine Riboflavin Sodium Anthocyanins

Increased bioavailability

Increased content

Niacin Protein

Calcium Magnesium

a

Data from ref. 60–63.

maize kernels overnight in a calcium hydroxide solution or with wood ash softened the kernels, which could then be stone ground into masa after rinsing off the lime solution. The masa is baked into tortillas.55 This processing improves the bioavailability of essential amino acids and facilitates the conversion of the amino acid tryptophan into niacin.56 Nixtamalization increases the solubility of lower molecular weight proteins, which then aggregate during masa and tortilla production.57 About 25% of the dietary fiber is lost during nixtamalization,58 presumably in the discarded liquid (see Table 2.4). Bressani et al.59 analyzed white and yellow maize as it passed through the stages of home tortilla production in Guatemala. Niacin, riboflavin and thiamine levels decreased by 31–65% from whole corn to masa, with the greatest losses from the germ, not the endosperm. The ash content increased by 0.35–0.40% after soaking in the lime solution for six hours.60 Calcium and magnesium concentrations increased after the lime treatment and subsequent production of tortillas, but sodium and potassium levels decreased.58

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Maize is not a good source of iron or zinc, but maize may be eaten in large enough quantities in some cultures to serve as an important source of these minerals. However, soaking in lime water can increase calcium eight-fold,61 so the advantages of nixtamalization may outweigh the negative nutritional consequences. The phytic acid content decreased by 21.4% from raw maize to tortilla, but residual levels might be sufficient to impair mineral absorption in vivo.61 The water solubility of anthocyanins is a challenge for processors of blue and purple maize. Longer cooking times (25 versus 35 minutes) and higher concentrations of calcium oxide (0.5–0.7% versus 1.0%) caused the greatest loss of anthocyanins in nixtamalized blue–purple corn.62 Nixtamalization reduced the total phenolic compounds in white, yellow, red and blue maize.63 The disposal of the high pH wastewater from nixtamalization is of environmental and economic concern and researchers have explored other techniques to produce masa more sustainably. The substitution of calcium hydroxide with calcium carbonate produced nutritional advantages, such as increased insoluble dietary fiber and resistant starch.64 The ecological nixtamalization process also increased the amount of total dietary fiber and resistant starch and reduced the glycemic index significantly for tamale fillings.65 Extrusion cooking with calcium hydroxide is also being explored as an alternative to traditional nixtamalization. Total carotenoid retention was comparable between tortillas made from extruded yellow corn flour and one produced by the traditional practice.66

2.4.2  Boiling Many cultures cook grains in water. Rinsing refined grains before cooking removes many water-soluble nutrients.67 The cooking water is fully absorbed for products such as rice, oatmeal and farro, or the excess liquid may be discarded. The retention of excess cooking water (beyond that absorbed by the grains) yields soups, gruels and porridges; it assumed that the nutrients that leach into the liquid portion of the food are retained to some degree (Figure 2.2). Rice can concentrate arsenic from soil, but cooking rice in extra water can reduce arsenic concentrations by 40–60% depending on the type of rice.67 Nutrients added to refined rice by enrichment are more easily leached into the cooking water than are the naturally present vitamins and minerals in brown rice.67 There was no difference in the resistant starch content of wheat, rice and barley cooked by boiling and pressure cooking.68 Parboiling, also known as conversion, partially cooks grains. Despite the name of the process, rice is soaked, then steamed and dried for stabilization. Nutrients are leached from the bran and aleurone layer into the endosperm during parboiling, increasing its nutrient content, but some losses of nutrients and phytochemicals occur.69 For example, one study reported that parboiling significantly reduced carotenoids, mostly lutein

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Figure 2.2  Compounds  leached from cereals into cooking water (asterisks indicate compounds not present in all cereals).

and β-carotene, in brown rice, which may be advantageous for consumers who desire white rice.70 Parboiling alters starch crystallization, leading to the slower in vitro digestion of starch in short-grain brown rice.71 Polishing (milling to remove the bran) parboiled rice may improve the digestibility, however. A shorter rice steaming time (20 minutes) resulted in the lowest glycemic index for rice, but 30 minutes of cooking increased the glycemic index.72 Oli et al.73 reviewed the nutritional and physicochemical changes that occur during the production of parboiled rice from a materials science approach.

2.4.3  Steaming Cooking grains by steaming can reduce nutrient losses from leaching. Goufo and Trindade69 summarized the changes in antioxidants in steamed versus boiled rice. Comparison of brown and partially germinated brown rice showed that steaming was more effective at retaining tocotrienols, tocopherols and γ-oryzanol than boiling and frying, which enhanced losses of these bioactive compounds via leaking into the cooking water and thermal degradation, respectively.74 Oats are kilned with steam to inactivate lipases and then rolled and dried to form convenient flakes. These processing steps are targeted at increasing the shelf life, but they also inactivate other enzymes, such as beta-glucanase. Kilning and steaming plus flaking resulted in beta-glucan viscosities that were nine- and 13-fold higher, respectively, than the viscosity of raw oats.75 Although oats are relatively rich in lipids, they also are good sources of antioxidants. Bryngelsson et al.76 reported increases in vanillin and ferulic acid and reductions in tocotrienols, caffeic acid and one avenanthramide during the steaming and flaking of oat groats.

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2.4.4  Baking Many different types of food products are baked. Moisture is lost during baking, but other changes in cereal composition depend on related steps, such as mixing and proving (in the case of yeast-leavened products). The mixing of cereals with other ingredients and water usually incorporates air into the mixture. Enzymes may become active in the presence of adequate moisture. Ktenioudaki et al.77 summarized the enzymatic reactions that occur during mixing. In bread, β-glucanase in wheat can reduce the molecular weight of oat and barley beta-glucan and, in turn, reduce the fiber's ability to lower serum cholesterol when consumed.75 Surface area, baking temperature and other factors influence nutrient retention in baked foods. Carotenoids and folate decrease in bread during baking.77

2.5  Thermal Processing with Minimal Added Water 2.5.1  Popping and Puffing Popping was one of the first means of cooking maize. Maize, sorghum and pearl millet have been the most popular grains to be popped, but interest in local foods has spurred the small-scale popping of other grains. Dry kernels are heated with or without cooking oil and the kernel pericarp serves as a miniature pressure cooker. Water molecules in each kernel vaporize, gelatinizing vitreous starch and causing expansion of the endosperm.78 Cooking kernels in hot sand is practiced in India.79 Brown rice that had been parboiled, dried and then puffed underwent losses of antioxidants, but the processing did not decrease the mineral content.80 Microwave ovens have increased the popularity of popcorn. Although popcorn is considered to be a snack food, popcorn has contributed significantly to intakes of whole grains and dietary fiber in the USA.81 Popping only takes a few minutes, thus minimizing the thermal degradation of nutrients, and no appreciable loss occurs due to leaching into the cooking medium. Red popcorn cooked with or without oil loses anthocyanins, but red and white corn popped in oil shows increased carotenoids and phenolic compounds relative to unpopped kernels.82 Popping does not adversely affect the digestibility of sorghum protein.83,84 The in vitro availability of iron increases, but the availability of zinc, free and bound phenolic compounds and total antioxidant activity decreases after popping sorghum.84 Popped amaranth (Amaranthus cruentus L. and A. hypochondriacus L.) has a similar total phenolic acid and antioxidant activity to the raw seeds, but the total flavonoid content is 72% higher in popped anmaranth.85 Grains are also puffed in “gun” chambers, where seeds are subjected to heat and pressure. Soluble phenolic acids and antioxidant activity increased after einkorn kernels were puffed in a commercial device.86 Puffed and popped grains, particularly those made from ancient grains,

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are finding new uses in snack bars and breakfast cereals. Milling these grains and then extruding them is another common option for new product development.

2.5.2  Roasting Roasting is a process involving high, dry heat. Grains may be roasted for use in coffee blends, such as the Thai beverage oleang88 or as coffee substitutes or adulterants.89 Barley and wheat are commonly roasted for malt, often after sprouting. Both grains can lose significant amounts of thiamine and riboflavin when roasted to darker colors at about 150 °C.90 Roasting has been reported to increase the total phenolic compounds and antioxidant activity.16

2.5.3  Microwave Heating Microwave ovens are common household appliances and cook foods quickly. Food companies use industrial microwaves for multiple purposes, but relatively little has been published on the effects of microwave irradiation on healthful compounds in cereals. The short cooking time suggests that thermal degradation may not be an important factor. Three-minute microwave heating of oatmeal resulted in higher levels of starch than oatmeal microwaved for five minutes or cooked by convection.91

2.5.4  Infrared Heating The heating of foods by infrared radiation is finding more applications in cereal processing. The infrared rays cause common molecular bonds to vibrate and, in turn, generate heat.92 Some researchers have referred to the use of near-infrared or infrared radiation processing of foods as “micronization”. However, micronization may also mean division into fine particles, so the meaning of this term should always be checked when reading research papers. Deepa and Habbar92 reviewed the effects of near-infrared radiation on legumes and cereals. Infrared drying of grains and cereal products can be completed more rapidly and with less product damage than other types of dehydration.93 Maillard reactions can occur at low (90% of peroxidase activity.95 Tryptophan decreased when white and red maize were

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treated with infrared radiation at 140 °C for 40 seconds; infrared processing also significantly reduced the total tocopherols in maize.96 β-Carotene also decreased in treated yellow maize. A promising application for infrared treatment is the inactivation of grain enzymes, particularly those that promote lipid hydrolysis and oxidation. The infrared treatment of oat groats successfully inhibited peroxidase activity, but spraying the grain with water helped to limit browning.97 The optimum retention of all phytochemicals in infrared-stabilized rice bran is challenging. The effective inhibition of lipases must be balanced by the retention of tocopherols, tocotrienols, γ-oryzanol and phenolic compounds. Infrared stabilization of rice bran at 140 °C for 15 minutes resulted in a 20% loss of vitamin E, but improved lipid stability with no change to the γ-oryzanol content.98

2.5.5  Extrusion Extrusion cooking accomplishes several unit operations simultaneously and can produce major changes in food composition and nutrient availability. Unique reactions are possible because macromolecules are conveyed and sheared along the extruder barrel.99 Several extruder parameters can be controlled to guide physicochemical reactions. Alam et al.100 reviewed how these parameters may affect extruded products. Thiamine and ascorbic acid are particularly susceptible to thermal degradation during extrusion.101 The twin-screw extrusion of wheat, barley, rye and oats yielded different changes in bioactive compounds.102 Inositol hexaphosphate, tocopherols, tocotrienols, reduced glutathione and melatonin were lower after extrusion, whereas the amount of phenolic acids increased. When anthocyanin-rich ingredients are mixed with cereals, some of the pigments are retained post-extrusion.103 The growing interest in pigmented grains suggests that these varieties could be targeted for the development of extruded functional foods. Extrusion decreased both free and bound phenolic acids and anthocyanins in refined and whole black rice (Oryza sativa var. Heiyounian), but the free forms of both types of compound increased significantly in extruded rice bran, which is a concentrated source of these natural antioxidants.104 Blue maize lost more than half of its anthocyanins after extrusion and subsequent transformation into tortillas; anthocyanin losses due only to exposure to lime for 20 hours or extrusion were not reported.105

2.6  Conclusions Cereals are an essential part of most people's diet. Grains pass through many steps of processing from farm to table. Few studies track grains through the entire process and laboratory-scale processing may not realistically mimic industrial or home practices. Published research often focuses on a limited number of healthful compounds due to financial and analytical constraints.

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Public–private partnerships may be needed to provide comprehensive research into the changes that can occur in grains throughout commercial and household processing. Subsequent chapters in this book address specific classes of phytochemicals and readers are encouraged to consider how processing might affect those chemicals and their bioavailability.

Acknowledgements Maine Agriculture and Forest Experiment Station external publication #3568. Financial support was provided by the United States Department of Agriculture National Institute of Food and Agriculture Hatch Project Numbers ME0-H-1-00516-13 and ME021804 through the Maine Agricultural & Forest Experiment Station.

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90. B. Hucker, L. Wakeling and F. Vriesekoop, J. Cereal Sci., 2012, 56, 300–306. 91. J. Harasym and R. Oledzki, Nutrients, 2018, 10, 207. 92. C. Deepa and H. U. Hebbar, Food Eng. Rev., 2016, 8, 201–213. 93. M. H. Riadh, S. A. B. Ahmad, M. H. Marhaban and A. C. Soh, Drying Technol., 2015, 33, 322–335. 94. S. Žilić, B. Ataç Mogol, G. Akıllıoğlu, A. Serpen, M. Babić and V. Gökmen, J. Cereal Sci., 2013, 58, 1–7. 95. C. Deepa and H. U. Hebbar, J. Cereal Sci., 2014, 60, 569–575. 96. S. Žilić, V. Šukalović, M. Milašinović, D. Ignjatović-Micić, M. Maksimović and V. Semenčenko, Food Technol. Biotechnol., 2010, 48, 198–206. 97. S. Cenkowski, A. R. Bale, W. E. Muir, N. D. G. White and S. D. Arntfield, Technical Sciences, 2004, 7, 15–26. 98. M. Irakli, F. Kleisiaris, A. Mygdalia and D. Katsantonis, J. Cereal Sci., 2018, 80, 135–142. 99. M. E. Camire, in Advances in Food Extrusion Technology, ed. M. A. Maskan and Aylin, CRC Press, Boca Raton, FL, 2012, pp. 87–102. 100. M. S. Alam, J. Kaur, H. Khaira and K. Gupta, Crit. Rev. Food Sci. Nutr., 2016, 56, 445–473. 101. M. N. Riaz, M. Asif and R. Ali, Crit. Rev. Food Sci. Nutr., 2009, 49, 361–368. 102. H. Zielinski, H. Kozlowska and B. Lewczuk, Innovative Food Sci. Emerging Technol., 2001, 2, 159–169. 103. C. Brennan, M. Brennan, E. Derbyshire and B. K. Tiwari, Trends Food Sci. Technol., 2011, 22, 570–575. 104. H. Ti, R. Zhang, M. Zhang, Z. Wei, J. Chi, Y. Deng and Y. Zhang, Food Chem., 2015, 178, 186–194. 105. J. Aguayo-Rojas, S. Mora-Rochín, E. O. Cuevas-Rodríguez, S. O. Serna-Saldivar, J. A. Gutierrez-Uribe, C. Reyes-Moreno and J. Milán-Carrillo, Plant Foods Hum. Nutr., 2012, 67, 178–185.

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Chapter 3

Composition and Functionality of Sugars and Oligosaccharides in Cereal Grains Yongfeng Ai University of Saskatchewan, Department of Food and Bioproduct Sciences, 51 Campus Drive, Saskatoon, S7N 5A8, Canada *E-mail: [email protected]

3.1  Introduction Cereals are plants that belong to the monocot Gramineae family, including maize (Zea mays L.), wheat [mostly common wheat (Triticum aestivum L.) and durum wheat (Triticum durum Desf.)], rice (Oryza sativa L.), barley (Hordeum vulgare L.), sorghum [Sorghum bicolor (L.) Moench], millet [mostly pearl millet (Pennisetum glaucum (L.) R. Br.)], oats (Avena sativa L.), rye (Secale cereale L.) and triticale (Triticale hexaploide Lart.). Cereals yield edible grains (seeds), which are a major source of nutrients (carbohydrates, proteins and vitamins) and calories for humans, livestock and poultry. Cereal grains are also the primary feedstock for the production of ethanol for alcoholic drinks, biofuel and other industrial uses. Globally, maize, wheat and rice are the most important cereal crops with respect to production and industrial applications. In recent years, pseudocereals—including amaranth (Amaranthus sp.), buckwheat (Fagopyrum esculentum Moench), chia (Salvia hispanica L.) and   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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quinoa (Chenopodium quinoa Willd)—have attracted much research attention, primarily because of their desirable nutritional properties.1,2 Pseudocereals, despite being used in a similar manner to cereal grains in foods, are not members of the Gramineae family. Carbohydrates are the predominant component of cereal grains, the content of which ranges from 66 to 78%.3 Carbohydrates occurring in cereal grains consist of a broad range of organic compounds, which can be further classified into sugars (e.g. glucose, fructose, maltose and sucrose), oligosaccharides (e.g. maltotriose, maltotetraose, raffinose, stachyose and verbascose) and polysaccharides (e.g. starch, cellulose, hemicellulose and β-glucan) based on their degree of polymerization. Compared with polysaccharides, sugars and oligosaccharides have much lower molecular weights and are minor components of mature cereal grains. Sugars and oligosaccharides play vital parts in the development and storage of the seeds of cereal crops. When cereal grains are used to prepare food ingredients and final products, numerous physical, chemical and biological reactions take place to break polysaccharides down into oligosaccharides and sugars, while some reactions convert these minor carbohydrate constituents into other substances. Therefore sugars and oligosaccharides can noticeably influence the functional properties, sensory attributes and nutritional value of food ingredients and products generated from cereal crops. This chapter reviews the analytical techniques available for the separation, identification and quantification of sugars and oligosaccharides in cereal grains. The sugar and oligosaccharide composition of mature cereal grains and how they change during caryopsis development are discussed. The changes occurring to sugars and oligosaccharides during the storage and processing of cereal grains and the resultant impacts on the quality and nutritional value of specific food products are also reviewed. This chapter provides a comprehensive overview of sugars and oligosaccharides in cereal grains and their products, with highlights of existing and future research efforts on these minor carbohydrate components.

3.2  D  efinition, Classification and Determination of Carbohydrates Carbohydrates, as the major component of cereal grains, include a diverse group of compounds. Carbohydrates are defined as polyhydroxy aldehydes (i.e. aldose) or polyhydroxy ketones (i.e. ketose) or a compound that can be derived from them through substitution (e.g. glucose phosphate, amino sugars), reduction (e.g. sugar alcohols), oxidation (e.g. sugar acids) and condensation (e.g. maltose, sucrose, starch, glycogen, chitin).4 Depending on the degree of polymerization (DP), carbohydrates can be classified into sugars (DP = 1–2, e.g. glucose, fructose, maltose and sucrose), oligosaccharides (DP = 3–10, e.g. maltotriose, maltotetraose, raffinose, stachyose and verbascose) and polysaccharides (DP >10, e.g. starch, cellulose, hemicellulose and

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β-glucan). This chapter focuses on the sugars and oligosaccharides present in cereal grains and their products. Some of the sugars—such as glucose, fructose, maltose and lactose—have a free aldehyde or ketone group (i.e. a hemiacetal or hemiketal group not involved in a glycosidic bond) and thus can reduce copper(ii) sulfate in alkaline solution to a red copper(i) oxide precipitate after heating, providing a positive result in Fehling's test.6 These sugars are defined as reducing sugars. By contrast, sucrose is an example of a non-reducing sugar. Because of their low molecular weights and relatively simple structures, sugars and oligosaccharides in cereal grains and their fractions can be extracted using hot water or a hot aqueous solution of ethanol.7,8 Common monosaccharides in the extracts can be quantified using enzymatic or spectrophotometric methods. For example, the concentration of d-glucose can be measured using a glucose oxidase/peroxidase system with a substance (e.g. adrenaline, 4-aminoantipyrine) as an oxygen acceptor for color development.9,10 The absorbance of the developed dye is determined spectrophotometrically and the value can be used to calculate the concentration of d-glucose in the sample. d-Fructose and d-galactose can be quantified using similar methods.11,12 Commercial assay kits are available for these analyses. The quantification of disaccharides (DP = 2, e.g. sucrose and lactose) and oligosaccharides in the extracts can be achieved by degrading the carbohydrates into their monosaccharide constituents using highly purified enzymes, followed by determination of the released monosaccharides.8,10,12 Chromatographic approaches have also been used to identify and quantify sugars and oligosaccharides in cereal grains and their fractions. Paper chromatography and thin-layer chromatography are the earlier methods used for this purpose.13,14 More advanced techniques have been developed for the separation and quantification of carbohydrates in the past few decades, including high-performance liquid chromatography,15,16 high-performance anion-exchange chromatography8,17 and gas chromatography.18,19 The advantages of these methods lie in the improved separation efficiency and quantification accuracy.

3.3  S  ugars and Oligosaccharides in Mature Cereal Grains Sucrose, fructose and glucose are the most important sugars found in mature cereal grains.20,21 Among these sugars, sucrose is present at the highest concentration for all of the cereal grains listed in Table 3.1, whereas fructose and glucose are detected at substantially lower levels. The two most important classes of oligosaccharides are found in cereal grains: (1) galactosyl derivatives of sucrose, such as raffinose, stachyose and verbascose; and (2) fructosyl derivatives of sucrose, such as fructo-oligosaccharides.20 Raffinose is present in trace amounts in sorghum grains and the highest content is found

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Table 3.1  Contents  of sugars and oligosaccharides in the meals of cereal grains.a Content (mg g−1, as-is basis) Sugar

Oligosaccharide

Fructose

Sucrose

Fructosyl (Fructosyl)2 (Fructosyl)3 (Fructosyl)≥4 Data from sucrose sucrose sucroseb sucrose reference (DP = 5) (DP >5) Raffinose Stachyose (DP = 3) (DP = 4)

Wheat Barley Oats Rye

c

LL 0.7 0.01 0.1

1.5 0.8 LL 0.1

7.9 13.6 4.3 7.1

4.7 2.3 0.5 0.8

0.06 0.02 0.3 0.02

2.9 2.6 —d 0.9

1.5 2.0 0.03 1.6

0.5 0.3 — 1.2

3.1 2.3 0.2 12.9

20 20 20 20

Maize (P-3737 hybrid)e Maize (OH 43 inbred) Wheat (Chinese Spring) Rice (Blue Bell) Barley (Himalaya) Barley (Steptoe) Sorghum (WAC 694) Oats (Dal) Rye (Balbo) Triticale (Fas Gro 204)

NAf NA NA NA NA NA NA NA NA NA

NA NA NA NA NA NA NA NA NA NA

14.2 15.0 13.8 5.6 14.2 11.8 8.4 8.8 11.5 8.4

3.1 2.1 7.0 — 7.9 6.3 TR 2.6 7.1 7.2

— — TRg — TR TR TR TR TR TR

NA NA NA NA NA NA NA NA NA NA

NA NA NA NA NA NA NA NA NA NA

NA NA NA NA NA NA NA NA NA NA

NA NA NA NA NA NA NA NA NA NA

21 21 21 21 21 21 21 21 21 21

Cereal grain

a

Glucose

 ugars and oligosaccharides were extracted using an aqueous solution of 80% (v/v) ethanol. S DP = degree of polymerization. c Level too low for reliable estimation. d Not detected. e Variety information is given in parentheses. f Not analyzed in the reference. g Trace amount detected. b

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−1

in barley (Himalaya variety; 7.9 mg g , as-is basis). By contrast, the concentration of stachyose is noticeably smaller and no verbascose can be detected in most cereal grains.7,22 In general, the total concentration of the raffinose family oligosaccharides—including raffinose, stachyose and verbascose—in cereal grains is lower than that in legume seeds.21,22 Therefore there is less concern that the consumption of cereal grains and their products will cause abdominal discomfort, flatulence or even diarrhea in a similar manner to legumes.23 Fructo-oligosaccharides with different DP values have been found in cereal grains and their concentrations are mostly 62%, dry basis), whereas fructan and sugars are minor components, consistent with the data shown in Table 3.1. Similar trends of changes in the contents of sugars, oligosaccharides and starch have been reported during the caryopsis development of durum wheat,34 maize,37 rice38 and barley.39,40 In addition to the reduction in the fructan content, research has shown that the average DP of fructan decreases during the seed maturation of durum wheat, which could be due to the degradation of fructan by enzymes, such as fructan exohydrolases.34

3.4.2  “Sweet” Mutants of Cereal Crops Seeds from normal varieties of cereal crops contain low concentrations of sugars (Table 3.1 and Figure 3.1). However, mutants that possess a substantially higher level of sugars in the kernels also exist. The presence of a large amount of sugars is responsible for a pleasant sweet taste of the kernels and these mutants are described as “sweet” varieties of the crops. Special sweet varieties have been found for maize,41 rice,42 barley,43 sorghum44 and wheat.45 For the sweet varieties, the mutation(s) occurs in the gene(s)

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Figure 3.1  Changes  in (A) the total kernel, dry kernel and moisture weights, (B) the

starch and crude protein contents as a percentage of kernel dry matter and (C) the fructan, sucrose, glucose and fructose contents as a percentage of kernel dry matter as a function of wheat seed (Triticum aestivum L.; Homeros variety) development time. P1, phase 1—cell division and expansion; P2, phase 2—grain-filling; and P3, phase 3—maturation and desiccation. Error bars are the standard deviations of triplicate measurements. Figure 1 has been adapted, with permission from J. Verspreet, S. Hemdane, E. Dornez, S. Cuyvers, A. Pollet, J. A. Delcour and C. M. Courtin, J. Agric. Food Chem., 2013, 61, 9251–9259. Copyright 2013 American Chemical Society.

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that encodes starch biosynthetic enzymes, leading to a deficiency in the enzymes. For example, sugary-1 maize, the most common genotype of “sweet corn (maize)” grown commercially, is deficient in isoamylase debranching enzyme.41 Because this debranching enzyme functions to trim excess branch chains of amylopectin molecules during normal starch biosynthesis, the deficiency in the enzyme causes a diversion from the biosynthesis of starch to the formation of phytoglycogen, a highly branched, water-soluble polysaccharide existing in the form of nanoparticles.41,46 It has also been reported that the sugary-1 phenotype is not expressed until three doses of the sugary-1 gene are present.47 The decrease in the conversion of sucrose to starch leads to the accumulation of sucrose—as well as glucose, fructose and fructan—in the seeds.43,47–49 The total content of sugars in the sweet kernels can be 2–14 times that in the kernels of the corresponding wild type.47,48 Sweet mutants of rice and barley have a genetic background similar to that of sweet corn: sweet rice is lacking two starch debranching enzymes (isoamylase and pullulanase42) and sweet barley is low in isoamylase activity.43 By contrast, one variety of sweet wheat is found to be deficient in granule-bound starch synthase I and starch synthase IIa,45 two important enzymes responsible for the elongation of amylose chain and amylopectin branch chains, respectively. It is crucial to note that the name “sweet sorghum” refers to a variety of sorghum different from these mutants: sweet sorghum accumulates a high level of free sugars in the stalk instead of in the kernels.50 Traditionally, free sugars in the stalk of sweet sorghum are extracted for the manufacture of refined sugar and downstream products. Recent research has focused on the utilization of sweet sorghum as a feedstock for biofuel production.51 The changes in the carbohydrate profiles of the sweet mutants during caryopsis development have been examined. Table 3.2 shows that the endosperm of the sugary-1 maize mutant has considerably higher contents of sucrose, glucose, fructose and sorbitol (a sugar alcohol) than those of the normal variety during seed development.47 The accumulation of soluble sugars in the sugary-1 maize kernels is responsible for an increase in osmotic pressure and thus a significantly greater level of moisture (58–71%) is retained than in the normal variety (34–58%) before maturation and desiccation (i.e. between 20 and 35 days post-pollination). Kernels of sugary-1 maize are commonly harvested before they mature and are consumed as a fresh vegetable or canned food. The amount of sugars present in the kernels determines the sweetness. During the desiccation process, the kernels of sweet mutants of cereal crops lose a larger proportion of their volume due to more moisture loss, resulting in a shrunken and wrinkled phenotype of the sweet kernels.41,49 During caryopsis development, the quantity and content of total sugars in both normal and sugary-1 kernels generally decrease (Table 3.2).47,52 The results, consistent with the changes in the composition of sugars during the development of wheat kernels (Figure 3.1), can be attributed to the conversion of sugars to more complex forms of carbohydrate (e.g. starch and phytoglycogen).

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Table 3.2  Fresh  weight, dry weight and moisture content of kernels, and sugar contents of endosperm in normal and sugary-1 OH43 maize on various days post-pollination (DDP). Adapted with permission from D. C. Doehlert, T. M. Kuo, J. A. Juvik, E. P. Beers and S. H. Duke, J. Am. Soc. Hortic. Sci., 1993, 118, 661–666.d a

Fresh weight (mg per kernel)

a

Dry weight (mg per kernel)

Moisturea (%)

Sucroseb (mg per g dry weight)

Glucoseb (mg per g dry weight)

Fructoseb (mg per g dry weight)

Sorbitolb (mg per g dry weight)

DPP

Normal sugary-1 Normal sugary-1 Normal sugary-1 Normal sugary-1 Normal sugary-1 Normal sugary-1 Normal sugary-1

10 15 20 25 30 35 Mature kernel

80 139 255 295 342 315 223

112** 192** 312** 386** 466** 426** 186**

15 38 108** 156** 192* 207** 202**

16 35 91 142 182 178 169

82 73 58 53 44 34 9

85 82 71** 63* 61** 58* 9

229 51 41 41 19 19 —c

205 121** 124** 94** 43** 70** —

117 4 3 1 0 0 —

104 53** 10 15* 14* 33* —

86 3 3 2 0 0 —

82 33** 5 3 2 6 —

6.6 0 0 0 0 0 —

—

10.6** 8.4** 5.3** 3.3** 3.3** 5.0**

a

Mean of 30 individual kernels. Mean of three extractions, each from kernels of a separate ear. Not available in the reference. d *,** Significant difference between normal and sugary-1 at p = 0.05 and 0.01, respectively. b c

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3.5  C  hanges in Sugars and Oligosaccharides During Storage and Processing of Cereal Grains After the mature seeds have been harvested, they are dried to a moisture content typically 0.2 corresponds to a greater storability of maize seeds.65 It has been suggested that oligosaccharides could stabilize intracellular glasses by enhancing the cytoplasm viscosity and increasing the glass-to-liquid transition temperature, which are likely to retard the deteriorative changes taking place during seed storage.65,67,68

3.5.2  R  eactions Generating Sugars and Oligosaccharides During the Processing of Cereal Grains In the process of transforming cereal grains and their fractions into various food products, treatments are applied to produce more sugars and/or oligosaccharides in the intermediates and end-products. If the generation rate exceeds the depletion rate, the outcome is an increase in the content of sugars and oligosaccharides. During the preparation of bread dough, endogenous amylases in wheat flour, such as α-amylase and β-amylase, can hydrolyze damaged starch to release maltose. Potus et al. found a greater increase in the maltose content of the dough prepared from wheat flour with a larger damaged starch content because damaged starch is susceptible to hydrolysis by the endogenous enzymes.69 The content of maltose in the dough prepared from a patent wheat flour (with the highest damaged starch content, 13.0% on a dry basis) increased 47- and 59-fold after mixing for 5 and 15 min, respectively. A higher mixing rate by alveograph and the addition of exogenous α-amylases led to a further increase in the maltose content. A similar conversion process of damaged starch into maltose by amylases has also been observed in the preparation of sourdough bread with an inoculation of different starter cultures.70,71 It is crucial to note that recent research has shown that some strains of bacteria (e.g. Lactobacillus reuteri, Lactobacillus sanfranciscensis and Weissella spp.) used as starter cultures in sourdough can synthesize a diverse group of oligosaccharides (e.g. gluco- and fructo-oligosaccharides) and exopolysaccharides from sucrose through the action of glycosyltransferases.72–74 The production of these non-digestible substances can potentially enhance the nutritional properties of the sourdough bread because they might be utilized as prebiotics by gut microflora in the gastrointestinal tract of humans.

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Another process that results in substantial increases in the contents of sugars and oligosaccharides during the processing of cereal grains is malting. Allosio-Ouarnier et al. investigated the changes in the carbohydrate profiles of four varieties of barley (Pyramid, Ferment, Alexis and Celinka) during malting for up to five days75 and the data for the representative Celinka variety are shown in Table 3.3. Over the five days of germination, the contents of the examined sugars (arabinose, xylose, glucose, fructose, sucrose, cellobiose and maltose), oligosaccharides from β-glucan (BG3 and BG5) and Table 3.3  Changes  in the contents of sugars (arabinose, xylose, glucose, fructose,

sucrose, laminaribiose, cellobiose and maltose), oligosaccharides from β-glucan (BG3, BG4, BG5 and BG6) and oligosaccharides of glucose (maltotriose, maltotetraose, isomaltose and isomaltotriose) in barley (Celinka variety) during malting for up to five days. Adapted from N. Allosio-Ouarnier, B. Quemener, D. Bertrand and P. Boivin, J. Inst. Brew., 2000, 106, 45–52, copyright ©1999, John Wiley & Sons, Inc. All Rights Reserved.,75 with permission from Wiley.

Malting Variety step Celinka

a

Sugar (mg g−1, dry basis)

Arabinose Xylose Glucose Fructose Sucrose End of 0.047 0.056 1.588 0.815 4.363 steeping Day of germination 1 0.052 0.070 2.388 1.018 3.577 2 0.120 0.105 6.750 2.432 3.843 3 0.200 0.231 13.319 2.919 7.868 4 0.247 0.179 15.353 3.061 11.690 5 0.315 0.412 19.464 3.211 13.988 Sugar or oligosaccharide from β-glucan (mg g−1, dry basis) Laminar- Cellobi- BG3a BG4a BG5a ibiose ose End of 0.066 0.105 0.302 0.030 0.155 steeping Day of germination 1 0.087 0.141 0.335 0.048 0.210 2 0.057 0.341 0.482 0.048 0.260 3 0.078 0.899 0.599 0.033 0.393 4 0.039 1.343 0.692 0 0.488 5 0.068 1.472 0.787 0 0.639 Sugar or oligosaccharide of glucose (mg g−1, dry basis) Maltose Maltotri- MaltoIsomalt- Isomalto­ ose tetraose ose triose End of 2.118 0.210 0.026 0.064 1.173 steeping Day of germination 1 2.699 0.304 0.026 0.099 0.523 2 4.755 0.643 0.032 0.318 0.177 3 6.754 1.204 0.099 0.650 0.095 4 8.006 1.337 0.099 1.387 0.438 5 10.891 2.538 0.106 1.996 0.446

Oligosaccharides with a degree of polymerization of 3, 4, 5 and 6, respectively.

BG6a 0.059 0.141 0.117 0.144 0.059 0.227

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oligosaccharides of glucose (maltotriose and maltotetraose) increased as a result of the enzymatic breakdown of the corresponding carbohydrate molecules with more complex structures. It is noteworthy that the content of isomaltose also increased, which can be attributed to a process of transglucosylation by α-glucosidase. The enzyme does not only catalyze the hydrolysis of α-gluco-oligosaccharides, but also transfers the glucosyl group to the hydroxyl group on the C6 atom of d-glucose.76 However, the contents of other sugars and oligosaccharides (e.g. laminaribiose, BG4, BG6 and isomaltotriose) did not show clear trends of changes during the five-day malting (Table 3.3) and the reasons for this phenomenon are unclear. The activities of the hydrolytic enzymes expressed during malting (e.g. α-amylase, β-amylase, α-glucosidase acting on starch; xylanase, arabinase and β-glucanase acting on cellulose and hemicellulose) can be partly retained after the kilning of malt. When used for beer brewing, the kilned malt is milled, suspended in water and heated to about 65 °C to gelatinize the starch, a step known as mashing.77 During mashing, the remaining endogenous hydrolytic enzymes in the malt, sometimes with the addition of exogenous enzymes (e.g. α-amylase, β-amylase or β-glucanase), can degrade a larger proportion of the complex polysaccharides present into oligosaccharides and sugars, primarily glucose and maltose.78–80 The sugars can be efficiently fermented by yeasts during the brewing process. After the completion of the brewing step, the barley malt residue consists of up to 40% arabinoxylans.81 Efforts have been made to use hydrolytic enzymes (e.g. feruloyl esterases and glycosyl hydrolases) to break down this group of polysaccharides into different arabinoxylo-oligosaccharides, feruloylated oligosaccharides and free ferulic acid.82–84 The released non-digestible oligosaccharides can potentially provide prebiotic functions. The presence of phenolic moieties in the feruloylated oligosaccharides can contribute to antioxidant and antimicrobial properties.85 There is a growing interest in utilizing germination as a clean label approach to modifying the functional and nutritional properties of cereal grains and flours. The biological and chemical changes that occur in cereal grains during the germination process are similar to those in malting. Saman et al. germinated waxy (0.1–0.3% amylose in starch) and normal (28–30% amylose in starch) rice grains at 30 °C under aerobic conditions for up to seven days.86 The concentrations of glucose, maltose and maltotriose increased and reached a maximum on day 3 of germination for both varieties as a result of amylolysis during the germination process. Sugar analysis showed that higher concentrations of sugars are released from the waxy rice than from the normal variety during germination, which may be a result of the greater enzymatic susceptibility of waxy starch due to the absence of amylose.86,87 Other researchers have also observed similar increases in the contents of sugars and reducing sugars in wheat, maize, sorghum, triticale and millet in the initial germination period.88,89 Saman et al.86 showed that the concentrations of sugars gradually decrease from days 4 to 7, which can be attributed to the increased utilization of the sugars by the germinated grains.86 The

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enzymatic degradation of starch, along with other polymers (e.g. protein, cellulose, hemicellulose and β-glucan), can considerably alter the functional properties of the germinated cereal grains and resultant flours. For example, the flours milled from germinated seeds show a significant improvement in their water absorption capacity, oil absorption capacity and emulsifying activity and emulsion stability, but a substantial decrease in their pasting viscosity compared with their non-germinated counterparts.90,91 When germinated brown rice flour is used to replace 30% wheat flour in the preparation of bread, the obtained product is significantly sweeter than the control due to the presence of more sugars.92 Regarding the nutritional value, the partial hydrolysis of starch and other components (e.g. proteins) during germination can markedly enhance the bioavailability of these nutrients and germinated cereal grains and flours can be a good source of essential nutrients and quick energy for infants.93 Malting has therefore been used as a promising method for the preparation of weaning foods.94,95 Extrusion is a continuous processing technology that combines several unit operations—including mixing, kneading, shearing, cooking, structuring and shaping—to transform a variety of raw materials into processed intermediates and end-products. Previous research has shown that extrusion processing can lead to the fragmentation of starch and other polysaccharides.96–98 However, extrusion does not usually generate significant amounts of oligosaccharides or sugars unless a high shear condition is applied.98 The addition of thermostable α-amylase during extrusion processing can efficiently convert starch into malto-oligosaccharides and sugars in barley and maize.99–101 This method has promise as a pretreatment technique to improve the bioavailability of carbohydrates in cereal grains for downstream processing (e.g. fermentation to produce bioethanol).

3.5.3  R  eactions Using Sugars and Oligosaccharides as Substrates During Processing of Cereal Grains Sucrose, fructose and glucose, as the most important sugars present in cereal grains and their fractions (Table 3.1), are readily available for biological conversion and chemical reactions during processing. The sugars occurring in a bread dough can be efficiently used by the incorporated yeasts (mostly Saccharomyces cerevisiae) to primarily produce ethanol and carbon dioxide, along with some minor compounds, such as higher alcohols, short-chain fatty acids and carbonyl compounds.102 Yeast fermentation is a crucial step with respect to leavening and increasing the volume of dough, as well as the aerated structure and aroma of the resultant bread.102,103 Sugars are utilized by yeasts at different rates. Potus et al. showed that sucrose from wheat flour is exhausted by yeasts after hydration and mixing for five minutes,69 which is a result of the efficient hydrolysis of this disaccharide into glucose and fructose by yeast invertase.69,103 In the same study, the content of glucose was shown to decrease at a faster rate than fructose in the dough, suggesting that glucose is the preferred substrate of these two monosaccharides for yeasts.69,103 After glucose and fructose in the dough are exhausted, maltose

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is left as the only sugar available to yeasts. The rate of gas production drops noticeably until the yeasts adapt to ferment maltose.103 Previous studies have also provided evidence that bread yeasts (S. cerevisiae) can efficiently degrade oligosaccharides (e.g. fructan) into simpler forms through the action of invertase.103–105 A positive association has been found between the content of oligosaccharides in wheat flour and the fermentation rate and total amount of gas generated.103 The significant degradation of fructan with larger molecular weights has been observed during the mixing of yeasted dough, and the majority (about 70–80%) of the fructan originally present in the raw whole wheat meal is eliminated after breadmaking.105,106 In another study, 40–62% degradation of fructan was observed for leavened bread baked from a commercial white wheat flour blend, a wholemeal wheat flour blend and a rye flour blend, whereas the unleavened counterparts retained most of the fructan (86–93%).28 The breakdown of fructan by yeasts during the breadmaking process can partly explain the low levels of fructan in leavened bread products from both wheat and rye flours.27,28,105,106 Because fructan can offer potential health benefits as a prebiotic, research has focused on preventing the degradation of this nutrient during breadmaking. The breakdown of fructan can be completely inhibited by using a yeast strain deficient in invertase.105 The nutritional importance of preserving fructan in bread requires further investigation. Traditional sourdough bread production has attracted much research attention in the past few decades because the use of sourdough can greatly improve wheat dough properties, enhance bread texture and flavor, retard the staling process and increase the nutritional value.107–110 However, few bakeries directly utilize sourdough as a leaving agent in bread production on an industrial scale. Instead, starter cultures containing a complex mixture of sourdough-fermenting bacteria and yeasts are commonly used. One of the main research foci in this area is the development of starter cultures with well-defined metabolic properties.111 Among the sourdough-fermenting bacteria, Lactobacillus strains are most commonly found, whereas other species belonging to Leuconostoc, Weissella, Pediococcus, Lactococcus, Enterococcus and Streptococcus genera have also been identified.112 The utilization of sugars and oligosaccharides in the sourdough is an important metabolic feature of starter cultures. Comparative studies have shown that different strains of sourdough-fermenting bacteria have their own patterns of metabolic activities on sugars under the examined conditions.70,71,112,113 Sugars are utilized by starter cultures to support growth and to generate various compounds—such as lactic acid, acetic acid, alcohols, esters and carbonyl compounds—which contribute to the unique aroma and flavor of sourdough and baked bread.114 Some species of sourdough lactobacilli, such as L. reuteri and L. sanfranciscensis, can hydrolyze α-1,6-linked gluco-oligosaccharides.115 Ganzle and Follador116 reviewed the carbohydrate transport systems and activities of the metabolic enzymes responsible for the metabolism of oligosaccharides in various strains of lactobacilli. Another good example of how the utilization of sugars and oligosaccharides can affect product quality is beer production. Sugars and oligosaccharides

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are the key components in the wort used for fermentation to produce beers. Wort is prepared from malt (primarily barley malt) through a series of steps, including milling, mashing, lautering and boiling.117 The sugars in wort can be readily fermented by yeasts to generate ethanol, carbon dioxide and some other substances—such as higher alcohols, esters, aldehydes and vicinal diketones—which contribute to the aroma and flavor of beers.118 Different strains of brewing yeasts are utilized to produce numerous styles of beer. In general, brewing yeasts can be classified into two types based on the flocculation behavior: top fermenting (e.g. ale yeasts) and bottom fermenting (e.g. lager yeasts).119 Brewing yeasts of both types can utilize various sugars—such as glucose, fructose, galactose, sucrose and maltose—with a major difference being that lager yeasts can ferment melibiose, but ale yeasts cannot.119 Lager yeasts have the activity of α-galactosidase to hydrolyze this disaccharide to galactose and glucose.120,121 Most of the sugars in wort are exhausted by yeasts during the brewing process, but some of the malto-oligosaccharides remain in beers. Commercial beer products can contain malto-oligosaccharides (DP 2–7) at a total concentration of 19.3–44.2 g L−1 and these oligosaccharides play crucial parts in the taste, ripeness and overall quality of the products.122 Sugars are also involved in non-enzymatic browning reactions, including the Maillard reaction and caramelization, during food processing. In the early stage of the Maillard reaction, a reducing sugar (e.g. glucose, fructose or maltose) condenses with a compound that has a free amino group (e.g. a free amino acid, the ε-amino group of lysine or the α-amino group of terminal amino acids in proteins) to generate N-substituted glycosylamine. The condensation product undergoes various rearrangement, degradation and other reactions to produce substances such as aldols, N-free polymers, aldimines, ketimines and melanoidins.123 The Maillard reaction can take place during the baking of bread and cakes, the toasting of cereals, the kilning of malt and the extruding of cereal flours to produce breakfast cereals and snacks. By contrast, caramelization occurs in the heating of sugars, particularly sucrose and reducing sugars, in the absence of compounds with a free amino group.124 The final product, caramel, contains a complex mixture of polymeric compounds. Although the optimum conditions for the Maillard reaction and caramelization are different, they may occur simultaneously during food processing and both reactions contribute to the development of aroma, taste and color and can influence the nutritional value of food products.125,126

3.6  Conclusions and Future Trends Cereal grains and their products are an important component of human diets, serving as a major source of nutrients and energy. Despite the fact that sugars and oligosaccharides are minor constituents in cereal grains, tremendous research efforts have been undertaken to advance our understanding of these carbohydrates in kernel development, storage and food processing. This knowledge is of great importance for the preparation of high-quality and nutritious food products from cereal crops. During

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caryopsis development, sugars and fructans are used as precursors for the biosynthesis of starch and other polysaccharides to fill the grains. When mutations occur in the genes encoding starch biosynthetic enzymes, there is a reduction in the conversion of sugars to starch. Consequently, sugars accumulate in the kernels and contribute to a pleasant sweet taste, as in immature sweet corn. The sugar and oligosaccharide composition of the embryo of harvested cereal seeds can affect their storability. During the processing of cereal grains in the food industry, numerous treatments have been utilized to degrade complex polysaccharides into oligosaccharides and sugars of simpler forms, as well as to convert sugars and oligosaccharides into other substances. These changes are important for the generation of food ingredients and final products with desirable functionality, sensory properties and nutritional profiles. One of the main foci of recent research is on increasing the content of non-digestible oligosaccharides in cereal grains and their products. These oligosaccharides are dietary fibers that have potential prebiotic properties and thus an increase in the content can enhance the nutritional value of cereal-based products. Future trends of research on sugars and oligosaccharides in cereal grains and their products include: (1) continued research to advance our understanding of the composition and functionality of these carbohydrate components during kernel development, storage and the processing of minor cereal and pseudocereal crops; (2) the development of new and fast methods to monitor the profiles of these carbohydrate components in these processes; (3) the development of new and efficient technologies to convert complex polysaccharides in cereal-based biomass into sugars for downstream processing (e.g. biofuel production); and (4) an increase in the content of non-digestible oligosaccharides in cereal grains and their products and an advanced and comprehensive understanding of the resultant physiological benefits.

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122. P. Mauri, M. Minoggio, P. Simonetti, C. Gardana and P. Pietta, Rapid Commun. Mass Spectrom., 2002, 16, 743–748. 123. S. I. F. S. Martins, W. M. F. Jongen and M. A. J. S. van Boekel, Trends Food Sci. Technol., 2000, 11, 364–373. 124. L. W. Kroh, Food Chem., 1994, 51, 373–379. 125. L. Manzocco, S. Calligaris, D. Mastrocola, M. C. Nicoli and C. R. Lerici, Trends Food Sci. Technol., 2000, 11, 340–346. 126. E. Purlis, J. Food Eng., 2010, 99, 239–249.

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Chapter 4

Types and Functionality of Polysaccharides in Cereal Grains L. Saulnier INRA, UR 1268 Biopolymers, Interactions, Assemblies, Rue de La Géraudière, Nantes, 44316, France *E-mail: [email protected]

4.1  Introduction Cereals are at the core of human nutrition and their incorporation into a wide range of products is of great economic importance. The major components of the grain are starch (60–70% of the grain, 70–80% of flour) and proteins (10–15%), with non-starch polysaccharides (NSPs) derived from the cell walls [also referred to as cell wall polysaccharides (CWPs)] only accounting for about 3–8% of the total. Nevertheless, these components have major technological effects on the end use of cereal grains1–3 and detrimental effects in animal nutrition (especially poultry feeding). Cell walls and their polysaccharides are the main constituents of dietary fibre and have major beneficial effects on human health, including a reduced transit time and reduction of the risk of cardiovascular disease, diabetes and certain cancers.4,5 The cereal grain is a complex organ consisting of several tissues with specific physiological functions. NSPs show a large diversity of composition,   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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structure and organization, both within the grain tissues and among cereal species. Depending on their tissue of origin, NSPs show different functionalities (e.g. solubility, viscosity and gelling properties), which largely explain their impact in food processing and in human and animal nutrition. This chapter first reviews the organization of cereal grain tissues. Our current understanding of the structure and diversity of cell wall polymers in the different tissues of mature cereal grains is then presented, along with their main physicochemical/functional properties. This chapter focuses on arabinoxylans and (1,3)-(1,4)-beta-d-glucans, also known as mixed-linked beta-glucans (MLGs), which are the major polymers in the cell walls of the starchy endosperm of wheat, rye and barley. Although these polymers are not the most abundant CWPs in the grain (they are found in the outer tissues), they have a major effect on the technological properties of grains and affect human and animal nutrition.

4.2  T  issue Organization and Cell Walls in Mature Cereal Grains 4.2.1  Cereal Grain Morphology and Tissue Organization The cereal grain, botanically the caryopsis, is composed of the pericarp (fruit coat), which originates from the ovary walls (maternal tissues) and encloses the seed. In addition to the embryo and the endosperm, the seed consists of the nucellar epidermis, the remnant of the nucellus, a seed coat (testa) and a cuticle. The grain may be enclosed in a husk, which corresponds to modified leaves and consists of empty cells with lignified secondary walls. In wheat, the caryopses are shed from the plant, whereas the husk remains in close contact with the caryopsis in rice, oats and barley. In naked barley, rice and oats, the husk can be dislodged on milling. The grain shape and form are characteristic of individual cereal species. The morphology and weight can vary considerably within species. Wheat, rye, barley, triticale and oats have a crease on the ventral side of the grain (Figure 4.1A). In the early stages of development, most of the volume of the grain is composed of the maternal tissues, but the outer layers undergo progressive differentiation and then degenerate. In the mature grain, the pericarp is reduced to a layer of four or five cells with thick lignified walls; the cuticle developed on the outer epidermis mainly serves to protect the grain. The evolution of these layers has been described in detail in wheat6 and barley,7 but the organization of these layers in mature grains can vary considerably among species. A schematic diagram of wheat grain tissues is presented in Figure 4.1B.8 The endosperm consists of an outermost layer of a highly specialized tissue known as the aleurone layer (about 7% of the dry grain weight) and of the starchy endosperm (80% of the dry grain weight). The endosperm consists of large cells with thin cell walls in which the starch granules are embedded in a storage protein matrix. The aleurone tissues generally consist of one layer of cells with thick cell walls (up to three layers of cells in barley or

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Figure 4.1  (A)  Example of grain morphologies; E: endosperm; eb: embryo; va: vascular trace (adapted by permission from Springer Nature: S. Krishnan and P. Dayanandan, Structural and histochemical studies on grain-filling in the caryopsis of rice (Oryza sativa L.), J. Biosci., 28, 455–469, Copyright © 2003, Springer Nature).163 (B) Tissue organization in wheat grain (adapted from A. Surget and C. Barron, Ind. Cereales, 2005, 145, 3–7).8

rice). The aleurone tissues remain alive at grain maturity and, on imbibition with water, synthesize and secrete the digestive enzymes that solubilize the storage components accumulated in the starchy endosperm to support the embryo's early growth.

4.2.2  Cell Wall Composition The chemical composition and structure of the CWPs in the mature grain vary depending on their tissue of origin and reflect the functions of the different cell types within the grain (Table 4.1). The cell wall can be differentiated

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Table 4.1  Main  polysaccharides in the cell wall of cereal grains.

a

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Endosperm

Wheat, rye, barley, oat, triticale Rice Corn, sorghum

Starchy endosperm

Aleurone layer

Pericarp

Thin, hydrophilic cell walls

Thick, hydrophilic cell Thick, hydrophobic walls cell walls

AXs and MLGs (partly water soluble) Traces of cellulose, heteromannan AGPs AXs, cellulose, pectin AXs, MLGs

AXs and MLGs

HXs, cellulose, lignin

Traces of cellulose, heteromannan n.r. n.r.

HXs, cellulose, lignin HXs, cellulose, lignin

a

AXs, arabinoxylans; MLGs, mixed-linked beta-glucans; AGPs, arabinogalactan peptides; HXs, heteroxylans; n.r., not reported.

into primary and secondary cell walls based on the source plant and stage of cell development. The primary cell wall has many important functions, such as providing structural and mechanical support, protection against pathogens and dehydration, the maintenance and determination of cell shape, and regulation of the cell turgor pressure. The primary cell walls also allow cell expansion and division during growth. This type of cell wall is mostly composed of cellulose and matrix polysaccharides (e.g. pectin and hemicelluloses), which differ in their nature and ratio according to the type of tissue and plant, and as a function of the cell and stage of organ development.9 They are classified into three classes: type I, found in dicotyldons and non-commelinin monocots; type II, found in grass species;10 and type III, typically found in ferns.11 In cereal grains, the primary cell walls are found in the endosperm tissues, where they represent 2–7% of the tissue and are thin and hydrophilic. The cell walls in the endosperm are essentially formed of two polymers, arabinoxylan and MLG, the proportions of which can vary significantly according to the cereal species – for example, rye, wheat and sorghum are rich in arabinoxylan, whereas barley and oats have high levels of MLG. MLG is a specific polymer of the type II primary wall of grasses and is not found in other plants,12,13 whereas arabinoxylan belongs to the xylan family, which is widely distributed among plants,14 although characteristic of the type II primary cell wall of grasses. Xylans are formed of a beta-1,4-xylan backbone carrying various sugars as side-chains. In the grain endosperm, they are characterized by a high amount of single arabinose side-chains relative to the xylans found in other tissues (e.g. stems and leaves) of grasses or in other plants.2,13 Low amounts of other polymers, such as glucomannan (2–7%) and cellulose (2–4%), are consistently reported in the endosperm cell walls in mature grains,6,7,13,15 whereas xyloglucans and pectins are transitorily observed in developing

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7,15

cereal grains, but are hardly detected in mature grains, with the exception of rice endosperm grains.16,17 These features are unique in the plant kingdom because the cell wall of the cotyledon of dicot seeds generally contains high levels of pectin and xyloglucan and a larger amount of cellulose than the cell wall of the endosperm tissue of cereal grains. Arabinogalactan peptides (AGPs) are also found in the endosperm of cereal grains.18 The secondary cell wall is present in highly specialized cells, such as vessel elements and in ground tissues, especially those originating from stems (e.g. sclerenchyma and collenchyma). It is formed after cell expansion and elongation, is terminated by the deposition of additional layers of polysaccharides (mainly cellulose) between the cytoplasmic membrane and the primary wall, and is reinforced by lignification. The tissues that form the outer part of the kernel primarily have a protective role and are typically secondary cell walls. Cell walls in these tissues are thick, hydrophobic and essentially formed of cellulose, xylans highly substituted by a single arabinose, single glucuronic acid, and more complex neutral sugar side-chains,2,13 which are generally referred to as heteroxylans. The walls of the outermost tissues also contain a significant amount of lignin. Another feature of cell walls in cereal grains that is common to grass and type II primary walls is the presence of hydroxycinnamic acids linked to some of the arabinose side-chains of xylans.19,20 These components play an important part in polymer interactions and the properties of the cell wall because they can dimerize under oxidizing conditions or with UV light, acting as bridging agents interconnecting xylan chains together and with lignins.21,22

4.3  S  tructure and Properties of Major Polysaccharides from the Starchy Endosperm of Cereal Grains There is a large variation in the tissue composition of milling fractions, which differ between species. The starchy endosperm tissue is essentially recovered in white flour, whereas the aleurone tissue in wheat is associated with the bran.23 Table 4.2 gives the concentrations of arabinoxylans and MLGs in the major cereal grains.

4.3.1  Arabinoxylans 4.3.1.1 Composition and General Structure The arabinoxylans are major components of the endosperm cell wall of cereal grains and they affect animal and human nutrition and cereal grain processing. The arabinoxylans in wheat, rye and barley have been studied extensively and many reviews describe the structure and properties of arabinoxylans

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Table 4.2  Cell  wall polysaccharide content of the starchy endosperm of cereal

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grains.b

AXa (g per 100 g flour)

MLG (g per 100 g grain)

Mean (range)

Mean (range)

Bread wheat Durum wheat33

1.9 (1.3–2.7) 1.9 (1.7–2.4)

0.7 (0.5–0.9) 0.3 (0.2–0.4)

Rye40 Barley37 Oat45

3.6 (3.1–4.3) 1.7 (1.4–2.0) 1.2 (1.0–1.3)

1.8 (1.7–2.0) 5.6 (4.6–6.5) 5.1 (4.5–5.6)

Rice2

0.5 (0.2–0.6)

n.r.

Corn44

1.0 (0.7–1.4)

n.r.

33

Other polymers Cellulosic glucose and heteromannans reported in low amounts in cell wall material AGPs are present as water-soluble polymers in the flour of cereal grains On isolated cell walls:164 AX (32%), cellulose (36%), pectin (7%) n.r.

a

 LG, mixed-linked beta-glucans; n.r. – not reported. M The arabinoxylan (AX) content was calculated as the sum of arabinose and xylose. Arabinose content was corrected for the presence of arabinogalactans, except in rice and corn.

b

from the flour or endosperm tissues of these cereal grains.1,2,24,25 By contrast, there has been little research into the arabinoxylans in rice, maize and other cereals. The arabinoxylans in the endosperm of cereal grains are found as water-extractable (WE-AX) and water-unextractable (WU-AX) fractions, which are both characterized by a high molecular weight (200–400 kDa).26–30 The proportion of the WE-AX fraction is variable and depends on the cultivar and cereal species, but it generally makes up to 25–30% of the total endosperm arabinoxylans in bread wheat flours (0.3–0.8% of white flour dry matter).27,31–34 Wheat-related species have comparable WE-AX fractions,33 as does barley,35–38 but the WE-AX fraction is higher in rye (1–2% flour dry matter).38–41 The total arabinoxylan content in endosperm tissues is about 2% (1.3–2.8% flour dry matter) in wheat endosperm,33,34,42 slightly lower for barley36,37 and higher for rye (3–4.5% flour dry matter)40 (Table 4.2). The arabinoxylans make up about 1% of the endosperm of oats, rice, sorghum and maize grains, essentially as the WU-AX fraction.43–45 The primary structures of the WE-AX and WU-AX fractions in the endosperm of wheat, rye and barley are closely related.27 They consist only of arabinose and xylose. Arabinose is mainly present as a single α-l-arabinofuranose (Araf) side-chain unit as a monosubstitution on position O3 (mXyl3) or as a disubstitution on positions O2 and O3 (dXyl) of the β(1,4)-d-xylopyranose (Xylp) residues of the backbone (Figure 4.2). Monosubstitution on O2 (mXyl2) is rare in wheat and rye, but represents a significant proportion in barley.35 On average, 50–60% of the Xylp residues of the backbone are

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Figure 4.2  Structural  features of the main cell wall polysaccharides in cereal grains. unsubstituted (uXyl). The arabinose to xylose ratio (A/X) is often used to characterize the structure of arabinoxylans and a typical average value in wheat is 0.55. This parameter has to be used carefully because it is only a rough characterization of the arabinoxylan structure and substitution. For example, a rye arabinoxylan that has a lower A/X ratio than wheat and barley has a higher level of substitution of its backbone due to a higher proportion of mXyl3.25,46 The WU-AX fraction has been studied less, although it represents most of the arabinoxylan in the cell walls of the endosperm. The structure, established after alkaline extraction, is essentially the same as for the WE-AX fraction, with wheat and rye having a slightly higher A/X ratio.24,29,47 The arabinoxylans in the endosperm of rice, sorghum and maize have a highly branched structure. An A/X ratio close to unity43,44,48,49 has been observed and the presence of glucuronic acid, galactose and short sidechains of arabinose has been reported in rice and sorghum.43,48 In wheat, the arabinoxylans of the aleurone layer, although structurally closely related to the starchy endosperm arabinoxylans, are not water-extractable and have a lower A/X ratio (0.4) than the starchy endosperm arabinoxylans.50 The presence of hydroxycinnamic acids linked to arabinoxylan is a common feature of grasses.20 In arabinoxylans from the endosperm of cereal grains, only ferulic acid is esterified on position O5 of the Araƒ residues (Figure 4.3). The amount of ferulic acid linked to arabinoxylan is low and represents 0.2–0.4% of the WE-AX (w/w) and 0.6–0.9% of the WU-AX fraction in wheat.27,50,51 This amount corresponds to about two to four ferulic acid residues for 1000 xylose residues in the WE-AX fraction (this ratio

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Figure 4.3  Ferulic  acid reactivity and cross-linking of feruloylated arabinoxylan. (A) Formation of phenoxy intermediates by free radical generating agents (e.g. the peroxidase/H2O2 system). The radical mechanism gives rise mainly to 8-8-, 8-5-, 8-O-4- and 5-5-coupled dehydrodimers.92 (B) Arabinoxylan–arabinoxylan cross-linking is effected by the radical dimerization of ferulates. Both ferulates and diferulates enter lignification reactions and cross-link arabinoxylan with lignin. For the ease of representation, only the 5,5′-coupled dehydrodimer is shown.

is two to three times higher in the WU-AX fraction). Similar proportions are observed for the WE-AX fraction in rye and barley.27 Dehydrodiferulic acids were also detected in low amounts (10–15 times less than ferulic acid) in the WE-AX fraction of wheat, barley, rye and triticale.27 The amount of dehydrodiferulic acid is four times lower than the ferulic acid content in the WU-AX fraction.50,52,53 In the aleurone layer, arabinoxylans show a lower A/X ratio and are more heavily esterified by ferulic and dehydrodiferulic acids, representing about 3.2 and 0.45% (w/w) of the polymer, respectively.50,51,53

4.3.1.2 Structural Heterogeneity Wheat, rye and barley endosperm arabinoxylans have common structural features and are characterized by a high heterogeneity of arabinose distribution on the xylan backbone. Populations of arabinoxylans showing different A/X ratios were isolated from water-soluble arabinoxylan polymers by various

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means (size-exclusion chromatography, DEAE cellulose chromatography, graded ethanol or ammonium sulfate fractionation).2 As an example, an arabinoxylan population with an A/X ratio ranging from 0.31 to 1.06 was isolated from the wheat WE-AX fraction with an initial A/X ratio of 0.56.28 Linear relationships were found between the A/X ratio and mXyl, dXyl and uXyl in WE-AX fractions obtained by graded fractionation24,28 and originating from different parts of the kernel54 or isolated from different varieties of grain.55 For wheat, a positive correlation was observed between the A/X ratio and dXyl, whereas mXyl and uXyl were negatively correlated. Therefore the A/X ratio in wheat is essentially dependent on the level of dXyl. Similar trends have been observed for the WE-AX fraction in rye and barley, although, in addition to dXyl, mXyl was also positively correlated with the A/X ratio.25 Structural studies on the WE-AX or WU-AX fraction have been carried out on bulk populations isolated from flours. The range of polymer structures observed clearly reflects structural heterogeneity within the endosperm. This heterogeneity was first suggested by structural studies on milling fractions obtained from wheat54 and barley56 and was further confirmed by studying the composition of the cell wall in wheat endosperm by different approaches, including infrared and Raman micro-spectroscopies, nuclear magnetic resonance spectroscopy and enzymatic fingerprinting.57–62 These studies have shown differences in the composition and structure of the cell wall polymers according to the cell position within the endosperm. The most recent studies on wheat and barley using a combination of enzymatic fingerprinting with matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) imaging illustrate the existence of a gradient in the disubstitution of arabinoxylan on a section of wheat grain (Figure 4.4).63,64 In this experiment, arabinoxylan were digested directly on grain sections with an endoxylanase and the degradation products were analysed in situ. Endoxylanases hydrolyse (1,4) linkages between the β-d-xylopyranosyl residues but the actual cleavage of a (1,4) bond depends on the presence of side chains on the xylose residues.65 Therefore the action of the endoxylanase is affected by the structure of the arabinoxylan and the patterns of arabino-xylo-oligosaccharides in turn give a fingerprint that provides information on the structure of the polymers.66,67 This spatial variation explains the chemical heterogeneity observed among arabinoxylans isolated from wheat flours. A structural variation of arabinoxylan is observed among wheat cultivars.34,55,62 This chemical heterogeneity has been demonstrated by calculating A/X ratio or using enzymatic fingerprinting66,68 and has also been illustrated by MALDI-MS imaging.64 Environmental conditions are likely to affect the amount of arabinoxylan in the endosperm of cereal grains, but the structural variation seems to be mainly genetically determined and a quantitative trait locus for the A/X ratio has been identified on the long arm of chromosome 1B in wheat.68–70 The arabinoxylans in the endosperm of rice, sorghum and maize grains show different features with a high level of substitution of the xylan

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Figure 4.4  Heterogeneity  of arabinoxylan structure within the endosperm of wheat grain as shown by enzymatic fingerprinting MALDI-MS imaging.63,64 (A) Average MALDI mass spectrum of mature wheat section after in situ enzymatic hydrolysis by endoxylanase. DP 5 is a monosubstituted xylotetraose with a single arabinofuranosyl residue at position 3 of the second xylopyranosyl residue from the non-reducing end, whereas DP 6 is disubstituted with arabinofuranosyl residue at both positions 2 and 3 of the same xylopyranosyl residue. (B) Plotting the ratio of DP 6 to DP 5 illustrates the variation in the degree of substitution of arabinoxylan within the endosperm. A high DP 6/DP 5 ratio corresponds to highly substituted arabinoxylan.

backbone. Enzymatic fingerprinting with a pure xylanase does not work on the endosperm arabinoxylan from maize or rice, whereas arabino-xylo-oligosaccharides providing information on the structure of arabinoxylan are obtained from all wheat species, barley, rye, triticale and oats. Possible variations of the arabinoxylan structure within the endosperm or among cultivars of maize, rice and sorghum have not been documented.

4.3.1.3 Molecular Weight, Physicochemical and Functional Properties The determination of the molecular weight (Mw) of polysaccharides can be affected by extraction artefacts and by the method of measurement. For example, the determination of the Mw of arabinoxylans based on sizeexclusion chromatography with a calibration of the column using pullulan or dextran only gives access to an apparent Mw (dextran or pullulan equivalent) because calibration polymers have a more flexible conformation in

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solution than arabinoxylan, meaning a lower hydrodynamic volume at the equivalent Mw. The absolute determination of the molecular weight of polymers can be obtained by light-scattering methods, which are now generally coupled with high-performance size-exclusion chromatography. High-performance size-exclusion chromatography coupled with light-scattering detectors allows the easy determination of the weight-average Mw, the number-average molecular weight (Mn) and the radius of gyration (Rg).71 However, light-scattering techniques are sensitive to aggregates, which can lead to an overestimation of the molar mass and discrepancies between results. These methodological limits explain the wide range of weight-average Mw and sometimes very low or extremely high published values for the WE-AX fraction of wheat, barley and rye. The most reliable studies have determined a weight-average Mw in the range of 200 000–400 000, with a relatively high polydispersity index of 1.7–2 (I = Mw/Mn). The rye WE-AX fraction shows a higher range of Mw than the wheat WE-AX fraction.27,72 No simple relation was observed between the Mw of the WE-AX fraction and the structure of the polymers in wheat, barley or rye.27,28,39,73 Determination of the Mw of the WU-AX fraction from the endosperm of cereal grains was carried out following an alkaline extraction (generally using BaOH2) of the polymer and the WU-AX fraction extracted from wheat74 or barley75 endosperm showed a higher Mw and a similar polydispersity index to the WE-AX fraction of the same sample. The viscosity of a polymer solution is directly related to the fundamental molecular properties (molecular conformation, Mw and distribution of the Mw) and concentration of the polymer. In dilute macromolecular solutions, viscosity measurements allow the determination of a limiting viscosity or intrinsic viscosity ([η]) of the macromolecule.71 The relation between the intrinsic viscosity and Mw of macromolecules is usually expressed in the form of the empirical Mark–Houwink equation: [η] = KMwa, where K and a are empirical parameters that depend on the polymer–solvent pair and the temperature, both of which are related to the chain stiffness.71 The value of the exponent a indicates the general conformation of the polymer: 0.5 ≤ a ≤ 1 for flexible linear chains and a >1 for stiff chains (values up to 1.8 are reported for rod-like chain conformations). Large variations in the intrinsic viscosities determined for the WE-AX fraction from the endosperm of cereal grains are observed between studies, in part due to the extraction procedure, which may or may not include the inactivation of endogenous endoxylanases. Variations were clearly observed between cultivars. In wheat, the intrinsic viscosities reported for the WE-AX fraction are in the range 0.2–0.6 m3 kg−1, with an average value of 0.4 m3 kg−1.31,76 The intrinsic viscosity of the rye WE-AX fraction is larger, with a range of variation from 0.4–1 m3 kg−1.27,38,39 The intrinsic viscosities reported for the barley, oats and triticale WE-AX fractions are generally similar to those reported for wheat.27,38 The exponent a of the Mark–Houwink equations has been

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determined for the wheat (a = 0.74) and rye (a = 0.94) WE-AX fractions. This value indicates a typical semi-flexible random coil behaviour similar to that in thickeners such as galactomannan.71,79 This semi-flexible random coil behaviour of the arabinoxylan chain is further confirmed by the value of the hydrodynamic parameter (ν = 0.47) determined from the radius of gyration–molecular weight relationship (Rg ≈ Mwν), which also gives an indication of the chain flexibility (ν = 1 for a fully extended rigid chain, 1/3 for a compact sphere and 0.5 for Gaussian random coil chains).77 Other measurements of chain rigidity, such as the persistence length (Lp) have been determined for wheat arabinoxylan (Lp = 3–8 nm).77,80 Again, this value is similar to that of galactomannans (4.5–9 nm)71,79 and higher than that for flexible 1 → 6-linked polymers such as pullulans (Lp = 1–3 nm)71 and much lower than stiff polysaccharides such as xanthan (Lp = 100–150 nm).71 The range of values obtained for the persistence length of arabinoxylan, together with the hydrodynamic parameter ν or the Mark– Houwink exponent a confirmed that the WE-AX fraction can be described as random coils in solution and not as extended rigid rods, as was often reported in earlier work.1,24 The conformation of the arabinoxylan chain is only slightly affected by the degree of branching of the xylan backbone and the viscosity of the arabinoxylan solution is mainly dependent on changes in the concentration and Mw of the polymer. Early work on arabinoxylan indicated that 95% of the viscosity of a wheat flour water extract was due to the WE-AX fraction and that the average size of the soluble molecules and the amount of water-soluble polysaccharide were specific varietal characteristics.81 The water solubility of a polysaccharide depends on the balance between chain–chain and chain–solvent interactions. Structural factors such as the chain length, the presence of side-chain groups and their distribution modify this balance and the solubility behaviour of the polymers. In general, the presence of side-chains that prevent chain–chain interactions favours water solubility of the polymers. The arabinoxylans in the endosperm of cereal grains are characterized by a high degree of substitution by arabinose sidechains and are therefore soluble in water at neutral pH, either naturally or after alkaline extraction. The water solubility of arabinoxylans in the endosperm of cereal grains is also related to the presence of covalent linkages or diferulic bridges, which cross-link the arabinoxylan chains and render the WU-AX fraction insoluble in water, although they have a higher A/X ratio than the WE-AX fraction. Modification of the distribution of arabinose side-chains affects the solubility of arabinoxylan chains and chain–chain interactions. The removal of arabinose residues by controlled acid hydrolysis82 or with arabinofuranosidase46,83–85 changes the solubility of arabinoxylans and gives rise to the aggregation and precipitation of the polymer. The aggregation is a result of the intermolecular aggregation of unsubstituted regular parts of the xylan chain. This is not simply related to the global Ara/Xyl ratio, but is also

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influenced by the distribution of substituted xylose residues (e.g. mono- or disubstituted with arabinose). Arabinose substitution also plays a part in the water-binding properties of arabinoxylan chains and this was evidenced by making films with the WE-AX fraction that showed different degrees of substitution.86,87 The effective moisture diffusivities (Deff ) of these films were determined at 0–95% relative humidity at 20 °C. Deff was influenced by the water content and the structure of polysaccharides. Higher Deff values were obtained for films made with highly substituted arabinoxylans. The proton dipolar second moments M2 and water T2 relaxation times measured by time domain nuclear magnetic resonance indicated that the highly branched arabinoxylan films showed a higher nanoporosity, favouring water motion within the films.86 In addition, traction tests showed that the strength and extensibility of arabinoxylan films were affected by the arabinose to xylose ratio. These results suggest that the structural variations in arabinoxylans may modulate the hydration properties of the cell walls and contribute to regulation of the water content of the grain.6 The gelation of feruloylated arabinoxylans from wheat endosperm under oxidizing conditions has been recognized for a long time.88 The reaction is catalysed by chemicals generating free radicals, such as ferric chloride or ammonium persulfate,89 or enzymatic systems (e.g. hydrogen peroxide/ peroxidase, linoleic acid/lipoxygenase or laccase/oxygen).89–91 The gelation primarily results from the formation of three-dimensional networks of arabinoxylan chains anchored by dehydrodimers of ferulic acid. The radical mechanism gives rise to different dehydrodimers92 (Figure 4.3) and 8-5′ (normal and benzofuran forms), 8-O-4′, 5-5′ and 8-8′ dehydrodiferulates have been detected in arabinoxylan gels, with the 8-5′ and 8-O-4′ dehydrodiferulates being the most abundant.27,93 The 4-O-8′,5′-5″ dehydrotrimer was also detected in gelled wheat arabinoxylans, but in lower concentrations than the dehydrodimers.94 The gelling reaction is fast, but enzymatically induced arabinoxylan gels show a significant decrease in their strength after a few days of storage, which has been attributed to partial depolymerization due to radical reactions catalysed by the still-active enzymatic system. Thermal inactivation of the enzyme is required to obtain stable gels.94 The structural characteristics of arabinoxylans play a determinant part in their gelation ability. Comparison of arabinoxylans from different sources has shown that a high content of ferulic acid, a high Mw to hydrodynamic volume and low substitution of the xylan backbone favour the formation of strong gels.24,27,95 A positive correlation has been thus established between gel rigidity and the intrinsic viscosity of arabinoxylans. The better gelling ability of poorly substituted arabinose chains is due to easier contacts between the adjacent feruloyl groups of neighbouring chains. The distribution of the feruloyl groups along the xylan chain could also influence the gelling ability.24,95 Model studies have proved that low Mw adducts of tyrosine

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or tyrosine-rich peptides and ferulate can be produced under specific conditions with regard to enzymes or substrates.91 Arabinoxylan/casein adducts have been obtained96 and dehydrodiferulic acid–tyrosine has been isolated from rye and wheat flour doughs.97

4.3.2  Mixed-Linked Beta-Glucans 4.3.2.1 Composition and Structure The MLGs are linear homopolysaccharides composed of β-d-glucopyranose (Glcp) residues linked by (1,3) and (1,4) bonds. MLGs are found in the primary cell walls of monocotyledons, particularly in Poaceae, in some lower plants and fungi, but not in dicotyledons.12 MLGs can be considered to be cellulose chains (4-O-linked β-d-Glcp units, 70%) interrupted by 3-O-linked β-d-Glcp units (30%) (Figure 4.2). These β-(1 → 3) linkages differentiate cereal β-glucan from cellulose and make the molecule more flexible and thus also soluble in water.98,99 The (1,3) linkages occur singly, leading to a structure of predominantly β-(1,3) linked cellotriosyl (DP3) and cellotetraosyl units (DP4).99–101 Cellulose-like sequences containing more than three consecutive β-(1-4)-linked d-Glcp units generally represent 90% carbohydrate. They are divided into classical and non-classical AGPs.115 Classical AGPs contain a glycosylphosphatidyl inositol (GPI) anchor signal in their

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protein sequence, allowing the formation of a GPI tail for anchorage to the outer surface of the plasma membrane. Other AGPs have been found to be soluble within the cell wall. In wheat and other cereal grains, the AGP are soluble in water, but evidence for their localization in the cell wall is currently lacking. Wheat endosperm AGPs have been estimated to be present at levels of 0.27–0.38% of the flour dry weight.32 The wheat grain AGPs consist of arabinogalactan chains attached through hydroxyproline residues116 to a small core polypeptide accounting for 8.0–9.0% (w/w) of the molecule. The peptide component is composed of 15 amino acids encoded by the grain softness protein 1 gene (GSP-1), which does not contain any GPI signal.117 The AGP peptide core includes three highly conserved hydroxyproline residues, each linked to a carbohydrate chain. The peptide amino acid sequence of spelt and durum wheat AGPs shows a high similarity to the wheat AGP sequence, whereas the triticale, rye and barley AGP peptide cores show less similarity.18 The arabinose to galactose ratio of the polysaccharide chain of about 0.7 (w/w) is stable among cereal species.18 The carbohydrate backbone of AGP is formed of a β-(1,3)-d-galactan backbone with β-(1,6)-d-galactan side-chains, which are highly substituted on position O-3 with single α-l-arabinofuranose residue (Figure 4.2).18,116 Glucuronic acid residues were detected at the non-reducing end of some short β-(1,6)-galactan side-chains in wheat AGPs.118

4.3.3.2 Properties The precise Mw of wheat flour AGPs is still unclear and can range from 22 to 70 kDa depending on the method used. Using size-exclusion chromatography, the apparent Mw of AGPs was compared for different cereal species; a value of about 23 500 was estimated for wheat spelt, durum wheat and barley and slightly higher values were reported for triticale (27 500) and rye (33 000).18 The AGPs are highly soluble molecules with a highly branched structure and thus do not contribute to the viscosity of the aqueous solution. The possible technological impact of AGPs during breadmaking was first suggested by showing a change in the properties of dough after the addition of isolated AGPs (2%) to wheat flour.119 AGPs are concentrated in the dough liquor, which is considered to be a good model of the bread dough liquid phase and has an important role in the creation of an alveolar structure (bread volume/ texture). The interfacial properties of the dough liquor suggest that AGPs are involved in the migration of dough liquor compounds to the interface between liquid films and air.120 The influence of AGPs on surface properties is clearly due to their amphiphilic character and they might contribute to the stabilization of gas cells during breadmaking.120

4.3.4  Interactions of Polymers in Endosperm Cell Walls The cell wall of the endosperm of cereal grains mainly consists of arabinoxylans and MLGs. Various observations using electron microscopy have shown a lamellar organization in wheat endosperm cells walls, especially in

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the aleurone layer, which possibly reflects the assembly of the arabinoxylan and MLG polymers.7,121 Although both arabinoxylan and MLG are co-localized in the wall, interactions between the two polymers have not been clearly demonstrated and there is only indirect and limited evidence of possible non-covalent interactions between arabinoxylans and MLGs.122 The cellulose content of endosperm cell walls is low and its interaction with arabinoxylans or MLGs is not likely to play a major part, although MLGs could naturally interact with cellulose through their longer cellulosic blocks. A number of studies have used the WE-AX fraction to look at their interaction with cellulose, mainly because the structure of the WE-AX fraction can be easily tailored (using enzymes, different sources or chemical fractionation), thus providing interesting insights on the structure–property relationships. In general, the size and substitution level influence the adsorption of xylans to cellulose: the more 0–3 and/or 0–2 linked arabinose, the less wheat arabinoxylan is adsorbed onto bacterial cellulose123,124 and the extent and pattern of arabinose substitution modulates the solubility in water as well as adsorption onto microcrystalline cellulose surfaces.85 As a consequence, the coating on cellulose of highly substituted arabinoxylan from wheat endosperm (A/X 0.6–0.7) is low compared with xyloglucans or with oat spelt xylans, which are considered to be linear.2,123,125 Bioassembly models using cellulose, arabinoxylan and MLG confirmed that neither arabinoxylan nor MLG has equivalent interactions with cellulose to either xyloglucan or pectin.124

4.4  S  tructure and Properties of Major Polysaccharides from the Outer Layers of Cereal Grains Maternal tissues from the outer part of the kernel primarily have a protective role. The cell walls in these tissues are thick, hydrophobic and essentially formed of cellulose and complex xylans, but also contain lignin.1 MLGs represent a few per cent of the cell walls of the outer tissues; in wheat they differ from the endosperm cell wall MLGs in having a higher DP 3/DP 4 ratio.61 The outer tissues of cereal grains isolated by the milling industry and referred to as brans are a complex assembly of different tissues and are not comparable between cereals. In wheat, the bran is composed of different tissues containing different arabinoxylan polymers.50,51,53 Table 4.3 shows that the nucellar epidermis is characterized by a low A/X ratio (0.12), whereas the cross cells and pericarp (also known as beeswing bran) are characterized by a high A/X ratio (1.2). Glucuronic acid (or its 4-O-methyl ester) and galactose are also associated with the arabinoxylan of the outermost tissues of cereal grains that are described as heteroxylans (see Figure 4.2). The structure of heteroxylans is further described in the following section and some of the general structural features of cellulose and lignin are provided with an emphasis on their possible interactions with heteroxylans in the cell walls of the outer tissues.

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Table 4.3  Composition  of the outer tissues of wheat grain (data adapted from C.

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Barron, A. Surget and X. Rouau, J. Cereal Sci., 2007, 45, 88–96).53 HXa (A/X ratio) Weight % of the tissue in grain

Aleurone Nucellar epidermis Inner pericarp + testa + nucellar epidermis Outer pericarp

MLGb

Celluloseb

(g per 100 g of tissue)

Phenolic acid (µg per mg of tissue) FA ester

Linked DHD to ester ligninc

6.5 —

24.4 (0.4) 9.3 60.0 (0.1) 4.4

2.5 —

8.0 10.3

0.3 0.6

— 0

3.5

38.8 (0.4) 3.4

12.3

4.8

0.6

5.8

3.7

47.0 (1.1) 1.5

24.3

3.2

2.3

4.5

a

 he heteroxylan (HX) content was calculated as the sum of arabinose and xylose. T The mixed-linked beta-glucan (MLG) and cellulose contents were calculated from glucose determined with pre-hydrolysis in 72% H2SO4 (MLG + cellulose) and without pre-hydrolysis (MLG). c The ferulic acid (FA) and dehydrodimers of ferulic acid (DHD) ester contents were determined after hydrolysis under mild alkaline conditions (35 °C, 2 M NaOH). The FA and DHD contents were also determined after hydrolysis under hot alkaline conditions (170 °C, 4 M NaOH). The (FA + DHD) esters was deduced from (FA + DHD) determined under hot alkaline conditions to calculate the amount linked to lignin. b

4.4.1  Heteroxylans 4.4.1.1 Composition and Structure The highly branched xylans rich in Araƒ found in the outermost part of cereal grains also contain α-d-glucuronic acid (GlcpA) (or its 4-O-methyl ether) and galactopyranose (Galp) (see Figure 4.2). Heteroxylans from cereal bran are derived from the pericarp tissue. These polymers are soluble at neutral pH after alkaline extraction and are characterized by a highly branched and complex structure that is similar in wheat,126–128 rye,47,129,130 maize,131,132 sorghum133 and rice.134 The level of substitution of the xylan backbone is high (80%) with a high proportion of dXyl (25–35% of the total Xylp). The A/X ratio is 0.8–1 and the Araf residues are mainly found as single unit side-chains. GlcpA and Xylp residues are also found as single unit side-chains; in addition, short side-chains constituted by Araf, Xylp and Galp have been isolated from these heteroxylans.135,136 A high level of hydroxycinnamic acid characterizes the outer tissues of the kernel. The amount of ferulic acid linked to arabinoxylans represents 0.9% (w/w) of the heteroxylans in the pericarp from wheat and up to 5% in the pericarp of maize; similar amounts have been observed for the dehydrodimers.44,53 This amount corresponds to about 30 and 60 ferulic acid residues for 1000 Xylp residues in wheat and maize pericarps, respectively.

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Acetic acid, essentially linked to the xylan backbone of heteroxylans, is observed in cereal brans.132,137,138

4.4.1.2 Properties The Mw of heteroxylans extracted from maize bran by alkaline treatment is about 280 000, with a polydispersity index of 2.139 In the case of maize bran, the Mw of heteroxylan is little affected by the extraction conditions (time, range of pH and temperature). Highly substituted arabinoxylans isolated from bran or cereal endosperm are generally soluble in aqueous media and do not tend to form aggregates. By contrast, low-substituted arabinoxylans isolated from the outer part of the grain have a strong tendency to form aggregates in aqueous solution and dimethylsulfoxide is a better solvent than water for arabinoxylans containing a low amount of arabinose substituents.26,140 The chemical modification of xylans or dioxane as a solvent was also used to determine the Mw of water-insoluble arabinoxylans isolated from rye bran; the value determined (62 600) indicated that aggregation occurred in water-based solvents.141 Heteroxylans extracted by alkaline treatment from the bran of wheat, rice or maize, although highly substituted, are readily converted into short chain fatty acids by the enzymes produced by faecal microbiota, whereas the bran fraction containing intact CWPs are hardly fermented under the same conditions.142 This phenomenon is explained by the high level of interconnection with other cell wall components in this tissue (see Section 4.4.3). These interconnections are limited in the cell walls of endosperm tissues that are easily degraded by enzymes (the WE-AX or WU-AX fraction).66 Cereal bran heteroxylans isolated by alkaline treatment are generally fully de-esterified during extraction. However, under mild alkaline conditions (pH 11–12 at 40–50 °C) it is possible to extract heteroxylans from cereal brans at an lower yield while preserving some of the ferulic acid–arabinose ester linkages. Heteroxylans extracted under these mild alkaline conditions can form gels under oxidative conditions in a similar manner to feruloylated arabinoxylans from the endosperm of wheat grains.143

4.4.2  Cellulose Cellulose is the main reinforcing polysaccharide in plant cell walls and represents up to 40% of the wall of the outer tissues of cereal grains. Its structure has been studied in depth as a result of its high abundance in nature, its major impact on the mechanical properties of cells and organs and its importance in the properties of wood, paper, cotton and other plant fibres. Cellulose is a linear homopolysaccharide composed of 1,4-linked β-d-glucopyranose (Glcp) units, where the repeating unit is the disaccharide cellobiose. The β-1,4 linkage favours the formation of intramolecular hydrogen bonds between Glcp residues, stabilizing an

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extended conformation of the glucan backbone and allowing intermolecular associations through hydrogen bonding of the cellulose chains.99,144 These intermolecular associations result in the lateral aggregation of β-1,4-glucan chains into microfibrils, where crystalline regions alternate with amorphous regions. Cellulose microfibrils are responsible for the strength and stiffness of the cell wall. Hemicelluloses are supposed to be associated with cellulose microfibrils through hydrogen bonding. However, in the case of cereal grains, the interaction of cellulose with arabinoxylans or heteroxylans is not straightforward. On the basis of X-ray diffraction studies, xylans are described as extended chains forming twisted ribbon-like strands with a three-fold symmetry.145 This extended conformation is often compared with that of β-1,4linked polysaccharides such as cellulose, but actually the three-fold screw symmetry of xylans differs from the two-fold symmetry of cellulose and does not favour associations.146,147 The (1,4)-β-xylan chain is more flexible than the conformationally constrained (1,4)-β-cellulose.147 This flexibility is generally explained by the fact that only one hydrogen bond is present between adjacent Xylp residues in xylan,148 whereas two hydrogen bonds are present between adjacent Glcp residues in cellulose. The arabinoxylan hydrogen bonding is clearly restricted by the presence of side-chains that restrict chain–chain interactions, especially for the highly substituted arabinoxylan of cereal grains.124,125 Highly substituted arabinoxylan (DS 0.8) isolated from maize pericarp, which has almost no unsubstituted xylose residues, did not bind to cellulose.2

4.4.3  Lignins and Hydroxycinnamic Acids Lignins are complex aromatic polymers resulting from the polymerization of p-coumaroyl, coniferoyl and sinapoyl alcohols. These are referred to as p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units, respectively, when incorporated into the lignin polymer.149 The composition of lignins (characterized by the ratio of G : S : H units) and the distribution of lignin polymers in tissues are different in woody gymnosperms, angiosperms and grasses. Grass lignins also carry p-coumaric and ferulic acids.150 Lignin plays a vital part in plant growth and development by improving water conduction through the xylem tracheary elements, enhancing the strength of fibrous tissues and limiting the spread of pathogens in plant tissues.149 The lignin content of wheat pericarp is in the range 5–6%, whereas the lignin content of maize pericarp is 35% are rare. Amylose forms a helical structure in a gelatinized (cooked) paste. This property enables it to bind small molecules, such as iodine, as inclusion complexes. Iodine binding in starch is the basis for the common standard methods of amylose determination, referred to as the ‘apparent amylose content’ (because some binding to amylopectin may also occur). Another method for the determination of amylose is by the binding of solubilized amylopectin to the lectin concanavalin A.5 The bound complex is precipitated and the remaining amylose is hydrolysed to glucose. The percentage of glucose determined from the amylose fraction is then compared with the total glucose from hydrolysis of the whole starch. This method has a greater physical basis than the iodine-binding methods, but it is complex and difficult to perform as a routine test. For most purposes, for the comparison of starches within the same species (e.g. in a breeding programme for eating quality in rice), the iodine-binding test is adequate. Other amylose determination methods, for large sets of fairly homogeneous samples, include techniques such as near-infrared and Raman spectroscopy.

5.2.3  Viscoamylography Viscoamylography is the measurement of the viscosity of a starch in an aqueous solution mixture under controlled heating and stirring, where the resistance to stirring is measured over time. The Brabender Viscograph instruments, developed since the 1930s by CW Brabender in Germany, are widely used in the baking industry, primarily for the determination of flour quality (degree of endogenous amylase enzymes) and testing the effect of added exogenous enzymes during the formulation of baked products. A similar instrument, designed for the more rapid quality control characterization of amylase starch damage in wheat, was developed in Australia as the Rapid Visco Analyzer (RVA)6 (developed by Newport Scientific, now owned by Perten Instruments). Some of the multiple applications of this instrument in measuring the properties of starch itself (independent of any enzyme effects) are discussed in detail here. A basic RVA curve (viscogram) illustrates the key information that can be gained from viscoamylography. About 10% starch or flour in water (or a suitable test solution) is placed in the RVA canister.

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The canister, with a stirring paddle inserted, is then placed in the heating block of the RVA instrument. Under constant rapid stirring, the starch–water or aqueous solution mixture undergoes heating and cooling, while the starch undergoes various transitions. The temperature profile is usually set to start at 50 °C for a couple of minutes, ramped up to 95 °C, held at 95 °C for a few minutes, cooled back to 50 °C and then maintained at 50 °C until the end of the test. The total duration of the test is usually about 20 minutes, but may be varied according to experimental need. The viscosity of starch granules in water is near-zero until the system is heated past the pasting temperature, which is closely related to the GT. After pasting, the viscosity continues to increase up to a peak, which is the most important indicative parameter of starch quality for many applications. Over time, with continued stirring at 95 °C, there will be an alignment of the starch molecules in solution and a decrease in viscosity, termed breakdown due to the breakdown of the granular structure, and indicated by the holding strength (a high holding strength indicates less breakdown). Few (but some) native starches are resistant to this breakdown with prolonged stirring, but stability (a high holding strength) is typical of many chemically modified starches. If the application requires a starch that is very resistant to prolonged harsh processing (such as a canned soup) while still maintaining viscosity, then a high holding strength is important. With cooling from 95 °C back to 50 °C (and continued stirring), the effect of a lower temperature usually overcomes the effect of prolonged shearing and the viscosity will increase again. The analysis of RVA curves can be used for the determination of suitable starches for an application and for verifying the consistency of starch properties for quality control purposes.

5.2.4  Dynamic Rheology The dynamic rheological analysis of starch–water systems gives information on both storage (G′) and loss (G″) components during the gelatinization and retrogradation processes7 and tan δ (G″/G′) is commonly deduced. This analysis is based on a small deformation and usually requires a small sample (e.g. 200 mg). A commonly used method is dynamic oscillatory analysis. A starch sample (e.g. 20% solid content) is placed on a stress-controlled rheometer plate under dynamic oscillation (e.g. a strain of 2% and a frequency of 1 Hz). The sample is heated from 40 to 90 °C to gelatinize the starch before cooling back to 25 °C to form a starch paste (Figure 5.3). The paste is subjected to a frequency–time sweep to obtain further information on the mechanical properties of the starch system.7 This method provides more fundamental information on the starch system than viscoamylography and is relatively easy to conduct. The price of an adequate stress-controlled rheometer is similar to that of a viscoamylograph such as the RVA. The dynamic rheological analysis of starch systems has been studied less than viscoamylographic analysis and the data obtained from dynamic rheology remains to be further correlated with the quality attributes of specific starch-based food systems.

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Figure 5.3  Development  of G′ of starches from maize mutants during heating cycle of dynamic oscillatory rheology. Reprinted with permission from F. Zhu, E. Bertoft and G. Li, J. Agric. Food Chem., 2016, 64(34), 6539–6545. Copyright (2016) American Chemical Society.7

5.2.5  Gelatinization Temperature Different starches from genotypes of the same species can vary widely in GT – for example, rice may commonly have values in the range 55–80 °C. The GT is slightly, but not strongly, correlated with the amylose content, so both are usually measured. The most basic significance of the GT is as a measure of the ease of cooking; all else being equal, a rice variety with a 55 °C GT will be preferred over one with an 80 °C GT because the energy required to reach the GT and hence for cooking will be substantially less. There are several ways to measure the GT. A hot-stage microscope can be used. By heating starch granules in water and observing them under polarized light, the point of loss of crystalline order (gelatinization) can be determined by the disappearance of the Maltese cross shape typical of the crystalline starch form. This method is not preferred because it is cumbersome and rather subjective. The alkali spreading method is used as an approximate scale for rice when screening many hundreds or thousands of samples in breeding research. In this test, a few grains of rice are immersed in dilute alkali and the degree of dissolution of the grains after 24 hours gives rough categories for the GT. The RVA pasting profile also gives an estimate of the GT, but it is relatively imprecise because of the lag in time between heating and recording the temperature in the heating block and the actual temperature in the starch– liquid system. The gold standard for the measurement of the GT is differential scanning calorimetry, where the heat required to heat starch in water is measured relative to the heat required to heat a standard (typically indium).

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An endothermic transition, such as the loss of crystallinity in starch gelatinization, can be observed in the differential scanning calorimetry8 curve. The GT (actually the gelatinization peak temperature, Tp) is the main parameter recorded, but the onset (To) and completion (Tc) temperatures and the change in enthalpy of gelatinization can also be recorded and have their own uses. As a result of the cost of equipment and the cost per sample, less precise rapid screening methods, such as the alkali spreading test for rice, are still widely used.

5.2.6  Texture The textural properties of foods are commonly measured with an Instrontype machine, where a suitable probe is pressed into the material and parameters such as the resistance to cutting, surface stickiness and hardness are measured through texture profile analysis (TPA). In principle, TPA with a probe of suitable geometry has the most direct relationship with the sensory acceptability of the texture of a cooked starch matrix (e.g. bread or noodles)9. TPA is widely used and represents an ‘imitative test’ more similar to the action of human teeth on a final cooked product than other tests such as viscoamylography (with RVA), which is a predictive test. The ease of measurement with TPA and the relative consistency of data compared with human sensory panel testing make it popular in research, product development and quality control in the food industry. However, there are serious problems with the typical TPA methodology. The human jaws are rotational, not a fixed plate and a vertically moving probe as in typical TPA. The geometry of human dentition and the action of biting are complex and are generally not imitated to any degree of precision in the TPA probes. The human sensory assessment of texture in the mouth also depends on the action of the tongue, the lubricating effects of saliva, and the rate and nature of particle size reduction during chewing. A simple texture analysis probe is a poor imitation of all these actions. Sample preparation is another issue. AACC International (formerly the American Association of Cereal Chemists) established an Asian Products Technical Panel, charged with the development of standard methods for the analysis of important Asian food materials and products. The development of a standard method for the measurement of texture in noodles proved almost insurmountably difficult. Problems included variations due to the preparation conditions (e.g. the boiling point of water at different altitudes, the use of salt in water, the exact boiling time, handling after cooking and the exact time before texture measurement). An example of the importance of texture to eating quality can be seen with rice. Consumers have very particular local preferences about the textural quality of rice. Within regions (e.g. China) where rice is typically cooked in a rice cooker with just sufficient water, different localities may prefer more or less sticky rice. Within regions (e.g. much of south and west Asia) where rice is boiled in an excess of water before consumption, the rice grains should

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separate after cooking and match local preferences for hardness and other characteristics. Because the preparation of rice by cooking is fairly simple, most of the quality characteristics are genetic attributes of the varieties used. Breeders select for the ‘eating quality’ and ‘cooking quality’ of rice using a series of simple tests, under the assumption that these are meaningfully related to the gold standard for quality evaluation, the human sensory panel, and to ultimate consumer acceptability. Tests for rice quality at the breeding level include many of those described here: the apparent amylose content (by iodine binding), gel consistency (the spread of a 5% starch gel in a tube, indicating the hardness of cooked rice), the GT (alkali spreading test) and instrumental TPA. However, all these tests represent approximations and do not imitate the complexity of human food processing.

5.2.7  Retrogradation Disordered amylose and amylopectin molecules of gelatinized starch hydrophobically interact with each other to form an ordered structure. This process is termed retrogradation. The retrogradation properties of starch are related to a range of quality attributes of starch-based food products, such as bread staling and digestibility.10 A higher degree of retrogradation is correlated with a higher staling rate of bread and a lower digestibility of starch. The short-term retrogradation (initial hours) of starch is mostly caused by the re-ordering and re-association of amylose, whereas the long-term retrogradation is largely attributed to the recrystallization of amylopectin. There are diverse methods to quantify the retrogradation process. Thermal analysis using differential scanning calorimetry is commonly used to monitor long-term retrogradation. Rheological methods, such as pasting and textural analysis, can also be used to indicate the changes in starch properties during storage. Spectroscopic (Fourier transform infrared, near-infrared, Raman and nuclear magnetic resonance spectroscopy), light scattering (wide-angle and small-angle X-ray scattering) and microscopic (e.g. AFM) methods have been used to probe the structural changes in starch during retrogradation.10 The turbidity and syneresis of gelatinized starch systems can also reflect the extent of retrogradation, which is easy to measure.10 Factors affecting the retrogradation of starch include the chemical composition, the amylopectin structure and the presence of other chemical components. Strategies to increase or decrease the retrogradation of starch have been developed for specific applications.10

5.2.8  Chain Length Distribution From a basic research point of view, it is not sufficient to note that samples of similar amylose content have substantial measured differences in other traits, such as their viscoamylograph profiles or GT, or indeed sensory preference profiles for particular populations. It is desirable to characterize the molecular structure of starch – for example, the chain length size distribution

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and branching pattern of amylopectin and the size and branching, if any, of amylose, along with information on the mode of packing in granules. Ideally, such information would be sufficient to predict precisely the physical properties of starch. For decades, scientists have hoped for ‘designer starches’ where a precise knowledge of the genetics and development of the molecular architecture of starch would combine with knowledge on how these translate into texture and other desired traits to match consumer demand. In such a situation, a physical profile of starch could be defined and breeders could rapidly adapt, for example, a high-yielding variety of rice into different lines, each carrying a particular starch profile. This is the ambition, but it is far from being realized and there may be many decades of research remaining to come near to this level of understanding. The measurement of the chain length distribution of amylopectin is carried out by the debranching and size separation of chains and measuring the relative contribution of each size of chain, usually from 6 to 100 glucosyl units (Figure 5.4). A commonly used separation system is high-performance anion-exchange chromatography coupled with pulsed amperometric detection. High-performance size-exclusion chromatography coupled with refractive index detection is also commonly used for the determination of the unit chain length distribution. The periodic distribution of the amylopectin unit chain length suggests that the unit chains are in the form of clusters.11 The branches of amylopectin are concentrated in the internal part of amylopectin, which is the part from the outermost branches to the reducing end of the molecule.12 Structural analysis of the products from the differential hydrolysis of amylopectin using α-amylase from Bacillus amyloliquefaciens showed that the branches are highly concentrated in certain regions.12 The tightly branched segments are referred to as building blocks. These blocks are

Figure 5.4  Unit  chain length distribution of quinoa (Chenopodium quinoa) amy-

lopectin. Reprinted with permission from G. Li and F. Zhu, Molecular structure of quinoa starch, Carbohydr. Polym., 2017, 158, 124–132. Copyright 2016 Elsevier Ltd.13

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mostly made up of a few chains. Blocks of two chains are the most abundant among different sized building blocks. The internal unit chain composition and building block structure of amylopectins from a range of botanical sources have been analyzed.12,13 Variations in the structural properties have been recorded for different samples. The unit chain length distribution of amylopectin and the composition of building blocks have been correlated with the different physicochemical properties of starch.14 For example, a higher number of short unit chains of amylopectin (degree of polymerization < 8) is correlated with lower GTs because these chains are too short to form helical structures. The inter-block chain length of amylopectin largely determines the degree of parallel packing of double helices. A longer inter-block chain length facilitates parallel packing, resulting in higher GTs.14 It becomes evident that the internal unit chain composition of amylopectin is crucial for the properties of starch.

5.2.9  Digestibility and Resistant Starch Staple foods represent the primary starch or energy source for human populations. Many people depend on grain-based staple foods, such as maize, wheat or rice, and others depend on non-grain starch sources, such as potato or sweet potato. The digestibility of starch enables the freeing of glucose molecules and their availability for energy metabolism. Modern diets are often not calorie-limited and, rather than maximize the caloric yield by selecting and processing for high starch digestibility, there is great interest in reducing it. A starch that is not fully digested by humans will have the added advantage of transferring resistant starch to the intestine, where it may have a probiotic effect on the gut microflora and act as dietary fibre to improve intestinal function and increase faecal bulk. Slowly digested starch may be favoured in helping to modulate the rapid release of glucose into the bloodstream. There is, however, some misunderstanding and confusion about starch digestibility. The digestibility of raw starch is low and it is fairly easy to select plant genotypes with high measured levels of resistant starch. However, raw starch is not edible and these high levels of resistant starch usually drop back to a few per cent after gelatinization (cooking). It is difficult to obtain high levels of indigestible starch after cooking by simple varietal selection. Other methods, such as the physical and chemical modification of starch, may be used to reduce digestibility. Food additives, such as plant polyphenols, may also be used to modulate the digestibility of starch (see Section 5.5).

5.3  Examples of the Impact of Genetic Variations 5.3.1  Wheat: from Udon Noodles to Waxy Starch Little attention was paid to the biological variation in the properties of wheat starch until about 30 years ago. Common wheat (bread wheat), Triticum aestivum L., is hexaploid and each of the genomes (A, B and D) has functional

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alleles affecting the starch composition. As a result, a single mutation will not have the dramatic effects on the properties of starch that may be found in diploid species – waxy rice is fairly common, but waxy wheat does not occur naturally. Within the relatively small variations in wheat starch properties observed in breeding programmes, the genetic variation in these properties does not have much effect on baking quality. The 12% or so of protein in flour is far more important than the 85% of starch in determining baking quality. However, starch has long been of interest to millers and bakers because of the effect of pre-harvest sprouting. Mature wheat stands in the field and dries to a suitable moisture content before harvest. A sudden rain before harvest can wet the grain and induce germination-related enzymatic reactions, including the activity of α-amylase enzymes that rapidly hydrolyse starch molecules. The grain often dries quickly after rain and there is no visible damage. Early testing methods (such as the falling number test) can be used at the point of receiving grain from the farmer to determine whether starch damage caused by enzymes has occurred. The original function of the RVA instrument (Section 5.2.3) was to allow the testing of Australian wheat for amylase-related starch damage (Figure 5.5), but it quickly gained recognition as a versatile research instrument for the quality analysis of undamaged starch. The first example of its use to determine the intrinsic properties of starch came about through the demand from Japanese millers for wheat grain suitable for milling into flour for making udon noodles. Udon noodles – made solely of flour, sodium chloride and water – are cut into thicker strands than other types of white noodles and have a soft and elastic texture. The screening of Australian wheat varieties identified lines, such as Eradu, with a suitably high starch peak viscosity (PV) that was highly correlated with its suitability for udon noodles. A later study indicated that Eradu was ‘partially waxy’ – that is, one of the three genomes of the hexaploid had a waxy mutant, but it was buffered by the other two non-waxy alleles in the other genomes (Figure 5.6). True waxy wheat (combining waxy mutants in each of the three hexaploid genomes) has also been developed and gives a product with interesting technological applications in baking and other products.

5.3.2  More Grains, More Noodles Major commercial noodle products (white salted, yellow alkaline, udon and most types of instant noodles) are made from wheat flour and now have fairly clear expectations from the consumer for the preferred texture. As we have seen with udon noodles, the choice of wheat variety (including the starch and protein characteristics) is essential for successful noodle manufacture. There are other types of noodle made from non-wheat grains (such as rice) or non-grain starches (such as sweet potato). For the less developed, but similar, products (e.g. using novel grain sources such as sorghum), there is more active product development and more dependence on the properties

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Figure 5.5  Effect  of amylase on RVA pasting of flour from wheat of different levels of rain damage (pre-harvest sprouting). (A) For different levels of falling number (indicating α-amylase activity) in water; (B) inactivation of α-amylase by silver nitrate. Reprinted with permission from G. B. Crosbie, A. S. Ross, T. Moro and P. C. Chiu, Cereal Chem., 1999, 76(3), 328–334. Copyright AACC International.15

of starch as the primary determinant of texture (in the absence of wheat gluten). As an example, we can look at a study of the diversity in the starch properties of sorghum to illustrate how the systematic selection and testing of starch variants can be used to control the outcome in terms of the textural quality of the noodle product.

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Figure 5.6  Variation  in RVA pasting profiles of wheat illustrating preferred types for Japanese udon noodles. The group with highest Peak Viscosity (PV) are ‘partially waxy’ and may be suitable for udon. Reprinted with permission from K. K. Y. Tsang, MPhil thesis, The University of Hong Kong, 1999, Figure 2 of Page 65. Copyright (1999) University of Hong Kong.16

Figure 5.7  Rapid  Visco Analyser (RVA) pasting profiles of starches from sorghum landraces selected to illustrate genotypes with high, average, and low peak viscosity. Corn (maize) starch pasting profile for comparison. Reprinted with permission from T. Beta, A. B. Obilana and H. Corke, Cereal Chem., 2001, 78(5), 583–589. Copyright AACC International.17

Beta et al.17 studied a set of 95 landraces (genotypes selected for agricultural use by farmers) originating from Zimbabwe (Figure 5.7). All the landraces had a ‘normal’ amylose content, within a fairly narrow range, and did not include mutants for waxy (0% amylose) or extremely high amylose contents.

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Figure 5.8  Genetic  and environmental variation in peak viscosity (PV) of starch

from each genotype at each environment for 10 sorghum genotypes grown in four different environments in Zimbabwe. Reprinted with permission from T. Beta and H. Corke, Genetic and environmental variation in sorghum starch properties, J. Cereal Sci., 34(3), 261–268. Copyright (2001) Academic Press.18

All the landraces exceeded a normal amylose maize control starch in PV, but generally showed the typical cereal grain RVA pattern of a moderate PV, distinct shear thinning and an increase in viscosity on cooling, reflected as setback. Clearly there is a wide variation, with PVs up to 377 RVU in the sorghum starches compared with 244 for maize. The patterns of shear thinning and setback also differed, with some lines (e.g. z-876) showing a fairly stable holding viscosity and others having a more marked decrease. There is another problem that often occurs in the quality management of starch-based foods. Even if you can source a specific variety of the grain being used, there can be substantial environmental effects (weather, soil conditions or agronomic protocols) on the starch pasting properties that translate into variations in product quality. Some varieties are fairly stable for these traits across environments and some are much less stable. To illustrate this phenomenon, the genetic and environmental variations in the properties of starches from 10 sorghum genotypes grown in four different environments in Zimbabwe were studied.18 The mean amylose content and the mean pasting PV of starch from each genotype in each environment were determined (Figure 5.8). The samples were treated with silver nitrate to prevent any effect of potential variations in the α-amylase content on the results. One genotype had a consistently high PV in each environment and some had fairly consistent medium levels of PV. However, there were clearly several genotypes with extreme variations from environment to environment. Such genotypes would be extremely hard to manage in a food processing context. For example, after the development of a sorghum-based noodle, we would expect the optimum sensory quality to come from a particular starch pasting curve. Verification of the pasting parameters of the incoming raw material would be highly useful for future quality control.

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5.4  Modified Starches A limitation to the use of native starches in food products is that they may be insufficiently stable under processing or in storage and may show undesirable effects, such as viscosity breakdown or syneresis. More stable starches can be made by chemical or physical modifications. Chemical modifications include hydroxpropylation for cross-linking or the addition of phosphate. These methods are effective, but they are expensive, typically resulting in a three to five times higher price, and they may meet consumer resistance as non-natural products. Physical modifications are regarded as ‘natural’ and consist of subjecting starch to temperature cycles (heating or cooling) under controlled moisture conditions. Heat–moisture treatments and annealing are typical physical modifications of starch. Modified starches represent a wide area of research and technology and further discussion of them is beyond the scope of this chapter.

5.5  Interactions of Starch with Polyphenols The interactions of starch with other components, such as phytochemicals, in food systems rich in starch may greatly affect the physicochemical properties of the foods.19 Polyphenols are natural antioxidants ubiquitously present in plants and starch-rich plant food systems. Certain types of grains, such as purple maize, are particularly rich in polyphenols. The interactions between starch and polyphenols are non-covalent. The polyphenols tend to form either V-type inclusion complexes or non-V-type complexes through hydrogen bonding. The formation of V-type inclusion complexes of starch with polyphenols remains to be verified. This uncertainty is due to the fact that polyphenols appear to be too bulky for the formation of inclusion complexes.20 Evidence of the formation of non-Vtype complexes is the increased hydrodynamic radius of amylose in the presence of tea polyphenols as studied by high-performance size-exclusion chromatography.21 The functional properties of starch can be altered by the addition of polyphenols.19 The addition of pomegranate peel extract (rich in ellagitannins, ellagic acid, gallagic acid and gallic acid) increased the PV while reducing the hot paste viscosity of wheat starch during pasting.22 The addition increased the gel hardness while decreasing the GTs of the starch. Ferulic acid and catechin decreased the hot paste viscosity, final viscosity and setback viscosity of maize and sorghum starch pastes, but had no influence on the PV of the former (Figure 5.9).23 Even though the two phenolic compounds increased the breakdown viscosity, it was the addition of ferulic acid which had a greater influence on the hot paste viscosity, the final viscosity, breakdown and setback than catechin. The addition of certain types of polyphenol, such as tea polyphenols, could reduce the digestibility of starch through interacting with the starch and/or enzymes. This reduction has important implications for nutritional applications, such as reducing

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Figure 5.9  Effect  of ferulic acid (FA) and catechin (CE) on maize starch pasting

properties. Reprinted with permission from T. Beta and H. Corke, Cereal Chem., 2004, 81(3), 418–422. Copyright AACC International.23

the chance of developing diabetes and obesity. The effect of polyphenols on the physicochemical properties of starch depends on the type and concentration of polyphenols and starch, the processing conditions and the type of properties measured.19

5.6  Conclusions There are many good books and reviews discussing in great detail the technical aspects of starch chemistry, structure and genetics. It is not the function of a short chapter such as this to compete with the depth and complexity of such works. However, in our experience, having taught grain technology for nearly 30 years, there are a few key points that, if understood, greatly facilitate further understanding and study. Many students do not grasp the dual function of viscoamylography measurements (to measure the properties of starch and to measure the activity of enzymes), they do not understand why starch mutants in wheat are observed less than those in maize or they do not grasp why starches of the same species with the same amylose levels can have such drastically different properties. We have tried to discuss basic points such as these, using examples where possible, in the hope that further studies will be made easier. Starch is surprisingly complex and there is still a lot to discover, especially about structure–function relationships and the true

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ability to define the desired starch properties and use first principles to breed suitable plant varieties with these properties. To use starch-based foods to carry bioactive molecules to the consumer, we need to understand starch and starch interactions before we can attain rational and efficient food product development.

References 1. J. L. Jane, T. Kasemsuwan, S. Leas, H. Zobel and J. F. Robyt, Starch/Stärke, 1994, 46, 121–129. 2. NEUROtiker - Own work, Public Domain, via Wikimedia Commons, https://commons.wikimedia.org/wiki/File:Amylose2.svg, accessed March 21, 2018. 3. NEUROtiker - Own work, Public Domain, via Wikimedia Commons, https://commons.wikimedia.org/wiki/File:Amylopektin_Sessel.svg, accessed March 21, 2018. 4. F. Zhu, Crit. Rev. Food Sci. Nutr., 2017, 57, 3127–3144. 5. T. S. Gibson, V. A. Solah and B. V. McCleary, J. Cereal Sci., 1997, 25, 111–119. 6. I. L. Batey in The RVA Handbook, ed. G. B. Crosbie and A. S. Ross, AACC International, St. Paul, MN, 2007, p. 19. 7. F. Zhu, E. Bertoft and G. Li, J. Agric. Food Chem., 2016, 64, 6539–6545. 8. Shimadzu Corporation, https://www.shimadzu.com/an/industry/foodbeverages/e8o1ci00000005nz_2.htm, accessed March 21, 2018. 9. B. McGregor, Bakerpedia. http://bakerpedia.com/analyzing-texture-of-products/, accessed March 21, 2018. 10. S. Wang, C. Li, L. Copeland, Q. Niu and S. Wang, Compr. Rev. Food Sci. Food Saf., 2015, 14, 568–585. 11. S. Hizukuri, Carbohydr. Res., 1986, 147, 342–347. 12. E. Bertoft, Cereal Chem., 2013, 90, 294–311. 13. G. Li and F. Zhu, Carbohydr. Polym., 2017, 158, 124–132. 14. V. Vamadevan and E. Bertoft, Starch/Stärke, 2015, 67, 55–68. 15. G. B. Crosbie, A. S. Ross, T. Moro and P. C. Chiu, Cereal Chem., 1999, 76, 328–334. 16. K. K. Y. Tsang, MPhil thesis, The University of Hong Kong, 1999. 17. T. Beta, A. B. Obilana and H. Corke, Cereal Chem., 2001, 78, 583–589. 18. T. Beta and H. Corke, J. Cereal Sci., 2001, 34, 261–268. 19. F. Zhu, Trends Food Sci. Technol., 2015, 43, 129–143. 20. R. Karunaratne and F. Zhu, Food Chem., 2016, 199, 372–379. 21. Y. Chai, M. Wang and G. Zhang, J. Agric. Food Chem., 2013, 61, 8608–8615. 22. F. Zhu, Y. Z. Cai, M. Sun and H. Corke, Food Chem., 2009, 112, 919–923. 23. T. Beta and H. Corke, Cereal Chem., 2004, 81, 418–422.

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Definition and Analysis of Dietary Fiber in Grain Products B. V. McCleary*, J. Cox, R. Ivory and E. Delaney Megazyme, Bray Business Park, Southern Cross Road, Bray, County Wicklow, Ireland *E-mail: [email protected]

6.1  Dietary Fiber as an Important Food Ingredient The concept of “available” and “unavailable” carbohydrates was brought into human nutrition early in the 19th century by McCance & Lawrence.1 In their research, the main objective was to differentiate the carbohydrates that affected blood glucose levels, i.e. those “available” for digestion and absorption in the small intestine. Methods were developed for analyzing reducing sugars, sucrose and starch in foods as a measure of the available carbohydrates; unavailable carbohydrates were determined as the insoluble residue, corrected for protein and ash. In 1935, Williams & Olmstedt2 developed a more physiologically based method for the measurement of indigestible material that simulated digestion by incubating the food sample with enzymes. The term “dietary fiber” was introduced by Hipsley3 in 1953 to cover the non-digestible constituents of plants that make up the plant cell wall, including cellulose, hemicellulose and lignin. This definition was broadened by Trowell et al. in 1972 4 and refined in 1976 5 to become primarily a   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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physiological definition based on edibility and resistance to digestion in the human small intestine; the definition included indigestible polysaccharides, such as gums, modified celluloses, mucilages and pectin. The work of Williams & Olmstedt2 formed the basis for enzymic–gravimetric methods developed later by Thomas6 and Hellendoorn et al.7 In 1976, Schaller8 introduced α-amylase treatment into the neutral detergent fiber method of Van Soest and Wine9 to resolve the problems experienced when the method was applied to starchy foods or ingredients in which starch was incompletely solubilized (American Association of Cereal Chemists Method 32–20 for Insoluble Dietary Fiber10). The first gravimetric procedures that measured both the insoluble and soluble components of dietary fiber were developed independently by Furda,11,12 Schweizer and Würsch13,14 and Asp & Johansson.15 These researchers, together with DeVries, Prosky and Harland, designed the enzymic–gravimetric method of AOAC International16 (AOAC Method 985.29). This procedure involved enzymatic treatments to remove starch and protein, followed by the alcoholic precipitation of high molecular weight dietary fiber (HMWDF), isolation of the insoluble components and weighing of the residue containing dietary fiber. This weight was then corrected for protein and ash in the residue. The method was later adapted for the measurement of insoluble and soluble fiber (Lee et al.17 AOAC Method 991.43 18). The removal of starch is also an essential first step in enzymic-chemical methods for the measurement of dietary fiber. Precipitation with 78% (v/v) ethanol is used to separate the high molecular weight soluble dietary fiber polysaccharides from low molecular weight sugars and starch hydrolysis products. Southgate19,20 developed a procedure following the principles of Widdowson & McCance21 that allowed a complete, sequential analysis of sugars, starches, non-cellulose polysaccharides, cellulose and lignin. Problems were experienced in the complete removal of starch and in the semi-specific colorimetric reactions for hexoses, pentoses and uronic acids. Schweizer & Würsch13 developed a gas–liquid chromatography method for the analysis of gravimetrically determined soluble and insoluble dietary fiber residues and Theander & Åman22 published the first version of the Uppsala dietary fiber methodology using gas–liquid chromatography for the determination and characterization of soluble and insoluble fractions of dietary fiber. Englyst et al.23–28 subsequently developed various methods for complete removal of starch and for the measurement of non-starch polysaccharides.

6.2  E  volution of the Codex Alimentarius Definition of Dietary Fiber In 1998, the American Association of Cereal Chemists International (AACCI) began a critical review of the current state of dietary fiber science, including consideration of the state of the definition of dietary fiber. After due deliberation, an updated definition of dietary fiber was delivered to the AACC

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Board of Directors for adoption in early 2000 and was subsequently published,29 namely:    Dietary fiber is the edible parts of plants or analogous carbohydrates that are resistant to digestion and absorption in the human small intestine with complete or partial fermentation in the large intestine. Dietary fiber includes polysaccharides, oligosaccharides, lignin, and associated plant substances. Dietary fibers promote beneficial physiological effects including laxation, and/or blood cholesterol attenuation, and/or blood glucose attenuation.    The Food Nutrition Board of the Institute of Medicine of the National Academies (USA)30 defined dietary fiber as:    Dietary fiber consists of nondigestible carbohydrates and lignin that are intrinsic and intact in plants. Added fiber consists of isolated, nondigestible carbohydrates that have beneficial physiological effects in humans. Total fiber is the sum of dietary fiber and added fiber.    Oligosaccharides that fall under the category of “dietary fiber” are those that are normally constituents of a dietary fiber source (e.g. raffinose, stachyose and verbascose in legumes) and the low molecular weight fructans in foods such as Jerusalem artichokes and onions. “Added fiber” consists of isolated or extracted non-digestible carbohydrates that have beneficial physiological effects in humans. These include fibers that have been isolated or extracted using chemical, enzymatic or aqueous steps, or those that have been chemically modified. Resistant starch and animal-derived, non-digestible carbohydrates are included. Isolated, manufactured or synthetic oligosaccharides of three or more degrees of polymerization are included, but non-digestible monosaccharides, disaccharides and sugar alcohols are not included because they fall under “carbohydrates” on the food label. In 2002, the term “added fiber” was modified to “functional fiber.” The definition of dietary fiber that arose from the 27th Session of the Codex Committee on Nutrition and Foods for Special Dietary Uses (ALINORM 06/29/26) in Bonn, Germany on November 21–25, 2005,31 was similar in many respects to that proposed by AACC, namely:    Dietary fiber means carbohydrate polymers with a degree of polymerization (DP) not lower than 3 which are neither digested nor absorbed in the small intestine. A degree of polymerization not lower than 3 is intended to exclude mono- and disaccharides. It is not intended to reflect the average DP of the mixture. Dietary fiber consists of one or more of: edible carbohydrate polymers naturally occurring in the food as consumed; carbohydrate polymers which have been obtained from raw materials by physical, enzymatic or chemical means, and; synthetic carbohydrate polymers.   

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By contrast, an FAO/WHO Expert Consultation on Carbohydrates in Human Nutrition in Geneva, Switzerland on July17–18, 2006 32 (as discussed at the 28th Session of Codex Committee on Nutrition and Food for Special Dietary Uses WHO/FAO Meeting of Carbohydrates Experts, Geneva, Switzerland, July17–18, 2006) Report CRD 19 33 concluded that the definition of dietary fiber should be more closely linked to fruits, vegetables and wholegrain cereals. To achieve this aim, they stated that the definition should include (1) a source element identifying that the dietary fiber is an intrinsic component of these food groups and (2) a chemical element identifying the component to be measured. The following definition was proposed: “Dietary fiber consists of intrinsic cell wall polysaccharides.” In rationalizing this decision and definition, many points were discussed, including their statement that non-starch polysaccharides can be measured specifically and that the term non-starch polysaccharides relates directly to the content of plant cell walls and the other beneficial substances they contain, such as micronutrients and phytochemicals. This definition has not been taken up, probably because there is no allowance for the range of isolated and synthesized dietary fibers that have become available as food ingredients. Based on the recommendation for the endorsement of the Codex Committee on Nutrition and Foods for Special Dietary Uses in November 2008, a definition of dietary fiber was adopted in June 2009 by the Codex Alimentarius Commission.34 The definition lists three categories of carbohydrates that are not hydrolyzed by the endogenous enzymes in the small intestine of humans. However, the definition left the decision concerning the inclusion, or otherwise, of oligosaccharides with degrees of polymerization (DP) in the range 3–9 to the discretion of national authorities and left the “physiological effect(s) of benefit to health” undefined:34    Dietary fiber consists of carbohydrate polymers (a) with ten or more monomeric units, (b) which are not hydrolyzed by the endogenous enzymes in the small intestine of humans and belong to the following categories: edible carbohydrate polymers naturally occurring in the food as consumed; carbohydrate polymers which have been obtained from food raw material by physical, enzymatic or chemical means and which have been shown to have a physiological effect of benefit to health as demonstrated by generally accepted scientific evidence to competent authorities, and; synthetic carbohydrate polymers which have been shown to have a physiological effect of benefit to health as demonstrated by generally accepted scientific evidence to competent authorities.    (a) When derived from a plant origin, dietary fiber may include fractions of lignin and/or other compounds when associated with polysaccharides in the plant cell walls and if these compounds are quantified by the AOAC gravimetric analytical method for dietary fiber analysis: fractions of lignin and the other compounds (proteic fractions, phenolic compounds, waxes,

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saponins, phytates, cutin, phytosterols, etc.) intimately “associated” with plant polysaccharides in the AOAC 991.43 method.    (b) Decision on whether to include carbohydrates of 3 to 9 monomeric units should be left up to national authorities.    As the Codex definition of dietary fiber was likely to become the reference definition in most countries, an urgent need existed for methodology that could service this definition—that is, a method that could measure dietary fiber as defined by AOAC Method 985.29, but with an accurate measure of resistant starch and accurate and reliable measurement of non-digestible oligosaccharides.

6.3  D  evelopment of a Procedure for the Measurement of Total Dietary Fiber, Including Resistant Starch and Non-digestible Oligosaccharides AOAC Methods 985.29 and 991.43 do not quantitatively measure resistant starch and do not measure non-digestible oligosaccharides. Consequently, other methods were developed for the measurement of specific non-digestible oligosaccharides, namely: fructo-oligosaccharides, AOAC Methods 997.08 35 and 999.03;36 Polydextrose, AOAC Method 2000.11;37 Fibersol 2 (resistant maltodextrins), AOAC Method 2001.03;38 and galacto-oligosaccharides, AOAC Method 2001.02.39 A method was also developed for the accurate measurement of resistant starch: AOAC Method 2002.02.40 It was initially thought that a value for total dietary fiber could be obtained simply by adding the values obtained by these specific methods to that obtained with the Prosky method16 (AOAC Method 985.29). However, this is prevented by the fact that the Prosky method also measures a varying percentage of some specific fibers, such as resistant starch, resistant maltodextrins, Polydextrose and fructans, introducing the problem of “double counting” (Figure 6.1). The need for a single integrated procedure for the measurement of all fiber components as defined by Codex Alimentarius was highlighted.

6.4  I ntegrated Procedure for the Measurement of Total Dietary Fiber as Defined by Codex Alimentarius Of all the dietary fiber components, the most difficult to measure is resistant starch. By definition, resistant starch is starch that is not digested during transit through the human small intestine. In vitro analytical procedures must therefore simulate in vivo digestion in the human small intestine. The integrated total dietary fiber (INTDF) method41 (AOAC Method

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Figure 6.1  Components  measured and not measured with AOAC Method 985.29. Reprinted with permission from B. V. McCleary, C. Mills and A. Draga, Qual. Assur. Saf. Crops Foods, 1 (04), 2009, 213–224.

2009.01/2011.25)42–44 was modeled on a widely used resistant starch method (AOAC Method 2002.02) that uses pancreatic α-amylase (PAA) and amyloglucosidase (AMG).40 It was optimized using a set of food and starch samples with defined resistant starch values (based on ileostomy studies). This method has been adopted as a Codex Type I method45 and has also been adopted by several food authorities worldwide. Samples are incubated with PAA/AMG at pH 6.0 and 37 °C for 16 h. After pH adjustment, the reaction is terminated and proteins in the sample are denatured by heating to about 95 °C. Higher molecular weight soluble dietary fiber, i.e. fiber that is precipitated in the presence of 78% v/v aqueous ethanol (SDFP), is recovered along with insoluble dietary fiber (IDF) by filtration. The recovered residue is dried, weighed and analyzed for both ash and residual protein, which are subtracted from the residue weight to give the amount of HMWDF. The alcoholic filtrate, which contains soluble dietary fiber that remains soluble in the presence of 78% aqueous ethanol (SDFS) is concentrated, desalted and analyzed by high-performance liquid chromatography (HPLC). The evaluation of this method over the past seven years has identified various limitations, or steps that could be improved and simplified. These include:    1. The incubation time of 16 hours with PAA plus AMG does not simulate physiological conditions. Based on numerous published studies, a more likely residence time for food in the small intestine is 4 ± 1 hours.46–51 2. Underestimation of phosphate cross-linked starch [resistant starch 4 (RS4), e.g. FiberRite, Fibersym)]. Much higher dietary fiber values for FiberRite and Fibersym are obtained38 using the Prosky total dietary fiber method (AOAC Method 985.29).52

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3. Underestimation of fructo-oligosaccharides with the HPLC column used. Some commercially available fructo-oligosaccharides contain the trisaccharide fructosyl-β-(2-1)-fructosyl-β-(2-1)-fructose (inulinotriose; F3) which, on chromatography on a Waters Sugar-Pak column, elutes at the same point as disaccharides so, by definition, is not included as a fiber component. 4. Resistant maltodextrins are produced on the hydrolysis of non-resistant starch under the incubation conditions of AOAC Method 2009.01. As these oligosaccharides are readily hydrolyzed by a mucosal α-glucosidase preparation from the small intestine of pigs,52 they clearly are not dietary fiber and should not be included. 5. AOAC Method 2009.01 uses sodium azide as a preservative in the buffer to prevent microbial infection over the extended incubation period of 16 h. The use of sodium azide is of concern to analysts. 6. Current procedures for the preparation of the SDFS fraction for HPLC are tedious.

6.5  R  apid Integrated Procedure for the Measurement of Total Dietary Fiber53 as Defined by Codex Alimentarius In updating the INTDF procedure, a major consideration was to perform the incubation with PAA/AMG in a time frame consistent with the residence time of food in the human small intestine. Numerous published studies indicated that the appropriate time is 4 ± 1 hours.46–51 With this in mind, experiments were performed with a range of control samples to determine the enzyme concentrations (PAA and AMG) and incubation conditions (pH, temperature and agitation) that result, after four hours of incubation, in resistant starch values in agreement with those obtained from ileostomy studies. The optimum conditions included incubation with PAA at 4 KU/sample and AMG at 1.7 KU/ sample at 37 °C and pH 6.0. There is considerable flexibility in the method of agitation of the sample during incubation. Similar dietary fiber values were obtained for the control samples independent of whether they were shaken in tubes or incubated in bottles that were either shaken or stirred throughout the incubation period. The major consideration was that samples must be kept in suspension throughout the entire incubation period. The resulting procedure, the rapid integrated total dietary fiber (RINTDF) procedure,53 is described in detail in the following sections.

6.5.1  Preparation of Test Samples Total dietary fiber should be determined on an as-is basis on dried, low-fat or fat-free samples. The samples are homogenized, weighed and then dried overnight in a vacuum oven at 70 °C. The samples are cooled in a desiccator, reweighed and the weight loss due to drying recorded. A 50 g portion of dried sample is dry-milled to pass through a 0.5 mm sieve. It may be desirable

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to freeze-dry samples with a high moisture content (>25%) before milling. If a high fat content (>10%) prevents proper milling, the sample should be defatted with petroleum ether (three times with 25 ml volumes per gram of sample) and dried in a laboratory hood before milling. When mixed diets are analyzed, the fat should always be extracted before determining the total dietary fiber. The weight loss resulting from fat removal is recorded and the final percentage of dietary fiber is corrected for both the moisture and fat removed. All material is transferred to a wide-mouthed plastic jar, sealed and mixed well by shaking and inversion before storage in the presence of a desiccant.

6.5.2  Enzyme Purity To ensure the presence of appropriate enzyme activity and the absence of undesirable enzyme activity when this procedure is used for cereal grains and products, control materials (beta-glucan, pectin, larch galactan, wheat starch, high amylose maize starch and casein) should be run through the entire procedure. Enzymes should be tested on a yearly basis.

6.5.3  Enzymatic Digestion of Sample 6.5.3.1 Blanks Two blanks should be run along with the samples in each assay to measure any contribution from reagents to the residual weights.

6.5.3.2 Samples 1. Accurately weigh about 1 g of sample (correct to the third decimal place) in duplicate into 250 mL Fisherbrand glass bottles. Record the weight. 2. Wet the sample with 1.0 mL of ethanol (EtOH) or industrial methylated spirits (IMS) and then add 35 mL of 50 mM sodium maleate buffer (50 mM, pH 6.0 plus 2 mM CaCl2) and a 7 × 30 mm stirrer bar to each bottle. Place the bottles on a 2 mag Mixdrive 15 magnetic stirrer in a waterbath at 37 °C. Stir the contents at 170 rpm for 10 minutes to equilibrate to 37 °C. Alternatively, transfer the bottles (without the stirrer bar) to a Grant OLS 200 shaking incubation bath (or similar), secure in place with the shaker frame springs and shake at 150 rpm in orbital motion for 10 minutes. 3. Add 5.0 mL of PAA/AMG solution (PAA 4 KU/5 mL and AMG 1.7 KU/5 mL) to each bottle, cap the bottles and incubate the reaction solutions at 37 °C with stirring at 170 rpm for exactly four hours using a magnetic stirrer bar and a 2 mag Mixdrive 15 magnetic stirrer apparatus. Alternatively, incubate in a shaking water-bath maintained at 37 °C at 150 rpm (orbital motion) for exactly four hours. If using an ammonium sulfate

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−1

111 −1

suspension of PAA/AMG [PAA (2 KU mL )/AMG (0.85 KU mL )] (see Section 6.5.7, point 2), gently swirl the suspension before use and add 2.0 mL of this suspension and 3 mL of maleate buffer to each bottle and incubate as described previously. 4. After four hours, remove all sample bottles from the stirring or shaking water-bath and immediately add 3.0 mL of 0.75 M Tris base solution (pH 11.0) to adjust the pH to about 8.2 (7.9–8.4), at which AMG has no activity. Immediately, slightly loosen the caps of the sample bottles, place the bottles in a boiling water-bath (non-shaking; 95–100 °C) and incubate for 20 minutes with occasional agitation (by hand). This inactivates both PAA and AMG. Ensure that the final temperature of the bottle contents is >90 °C using a thermometer. Checking just one bottle is adequate. At the same time, if only one shaker bath is available, increase the temperature of the shaking incubation bath to 60 °C in readiness for the protease incubation step. 5. Remove all sample bottles from the hot water-bath (using appropriate gloves) and cool to 60 ± 1.0 °C. 6. Add 0.1 mL of protease suspension (50 mg mL−1 in 3.2 M ammonium sulfate) with a positive displacement dispenser (the suspension is fairly thick). Incubate at 60 °C for 30 minutes. 7. Adjust the pH by adding 4.0 mL of 2 M acetic acid to each bottle and mix. This gives a final pH of about 4.3. 8. To each sample, add 1 mL of 100 mg mL−1 glycerol internal standard solution (in 0.02% sodium azide).    NOTE: The PAA/AMG mixture, protease suspension, LC retention time standard, glycerol standard solution (100 mg mL−1) and glycerol/glucose standard solution (10 mg mL−1 of each) are available in the Megazyme Rapid Integrated Total Dietary Fiber Kit (K-RINTDF). Celite (G-CELITE) and ion-exchange resins (G-AMBOH and G-AMBH) are also available from Megazyme.

6.5.4  Determination of HMWDF (IDF + SDFP) 1. To each sample, add 207 mL (measured at room temperature) of 95% (v/v) EtOH preheated to 60 °C and mix thoroughly. Allow the precipitate to form at room temperature for at least 60 minutes (overnight precipitation is acceptable). 2. Prepare the bed of Celite as described previously.52 Tare crucible containing Celite to nearest 0.1 mg. Wet and redistribute the bed of Celite in the crucible using 15 mL of 78% (v/v) aqueous EtOH or IMS from a wash bottle (refer to Figure 2 in ref. 52). Apply suction to crucible to draw Celite onto fritted glass as an even mat. Discard these washings. 3. Under vacuum, filter precipitated enzyme digest from step 1 through the crucible. Using a wash bottle with 78% (v/v) aqueous EtOH or IMS, quantitatively transfer all the remaining particles to the crucible and

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wash the residue successively with two 15 mL portions of 78% v/v aqueous EtOH or IMS. Retain the filtrate and washings for the determination of SDFS (Section 6.5.5, step 1. 4. Using a vacuum, wash the residue successively with two 15 mL portions of the 95% (v/v) EtOH or IMS and then acetone into a “waste” Buchner flask. Discard these washings. Draw air through the crucible for at least 2 minutes to ensure all the acetone is removed and then dry the crucibles in an oven. 5. Dry the crucibles containing the residue overnight in an oven at 105 °C. Loosely cover the crucibles with aluminum foil to prevent the loss of sample during drying. 6. Cool the crucibles in a desiccator for about one hour. Weigh the crucible containing the dietary fiber residue and Celite to the nearest 0.1 mg. To obtain the residue mass, subtract the tare weight, i.e. the weight of the dried crucible and Celite. 7. Analyze the residue from one crucible for protein and analyze the residue of the duplicate for ash. Perform protein analysis on the residue using Kjeldahl or combustion methods. Use a factor of 6.25 to calculate the number of grams of protein. For the ash analysis, incinerate the second residue for five hours at 525 °C. Cool in a desiccator and weigh to the nearest 0.1 mg. Subtract the crucible and Celite weight to determine the ash content.

6.5.5  Determination of SDFS Proper deionization of the filtrate is an essential part of obtaining quality chromatographic data on SDFS. Figure 6.2 shows patterns for glycerol and d-glucose in the presence and absence of buffer salts.    1. Recover the filtrate (from Section 6.5.4, step 3) and transfer into a 500 mL measuring cylinder. Adjust the volume to 300 mL with 78% v/v IMS in water, transfer to a 1 L beaker and mix thoroughly. Transfer about 75 mL (about 25%) of this solution to a 500 mL evaporator flask and concentrate with a rotary evaporator to dryness at 50 °C. (Note: it is not essential to quantitatively transfer all the solution because SDFS is determined by the ratio of these peaks on HPLC to that of the glycerol internal standard). 2. Deionize the sample. Dissolve the residue in the evaporator flask in 8 mL of deionized water. Transfer 5 mL of this solution to a 13 mL polypropylene tube containing 1.5 g of Amberlite FPA53 (OH−) resin and 1.5 g of Ambersep 200 (H+) resin. Cap the container and invert the contents regularly over 5 minutes. Alternatively, if the ammonium sulfate suspension of PAA/AMG is used for starch digestion, then use 2 g of Amberlite FPA53 (OH−) resin and 2 g of Ambersep 200 (H+) to ensure the effective removal of most of the salt in the sample. 3. Prepare samples for HPLC analysis. Remove a sample (about 1.5–2.0 mL) of the supernatant solution from the resin slurry with a syringe

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Figure 6.2  Chromatography  on TSKgel G2500PWXL columns of (a) a non-desalted

mixture of glucose and glycerol in RINTDF incubation buffer mixture; (b) the same sample after desalting 5 mL with 1.5 g of Amberlite FPA53 (OH−) and 1.5 g of Ambersep 200 (H+) resins in a polypropylene tube; and (c) sample from part (b) chromatographed on the same TSKgel G2500PWXL columns, but also with desalting pre-columns in place.

and filter through a polyvinylidene fluoride filter, pore size 0.45 µm. Use this solution as the sample extract in Section 6.5.5, step 6. The HPLC patterns for the non-desalted sample, the sample desalted with resin in the tube and a sample of desalted preparation run onto TSKgel G2500PWXL columns through Bio-Rad de-ashing pre-cartridges are shown in Figure 6.2. 4. Determine the response factor for d-glucose. As d-glucose provides an HPLC refractive index response equivalent to the response factor for the non-digestible oligosaccharides that make up SDFS, d-glucose is used to calibrate the HPLC system and the response factor is used to determine the mass of SDFS. Use a 100 µL liquid chromatography syringe to fill the 50 µL injection loop for the internal standard/d-glucose solution (10 mg mL−1 of both d-glucose and glycerol in sodium azide solution, 0.02% w/v). Inject in triplicate. 5. Obtain the values for the peak areas of d-glucose and glycerol from duplicate chromatograms. The ratio of peak area of the d-glucose/peak area of glycerol to the ratio of the mass of d-glucose/mass of glycerol is the response factor. The average response factor for d-glucose is about 0.82 that of glycerol.

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Figure 6.3  Chromatography  on TSKgel G2500PWXL columns of Raftilose P-95 and glycerol showing the SDFS fraction and the point of demarcation between what is included and what is not included.

6. The response factor (Rf) = (PA-Glycerol)/(PA-Glu) × (Wt-Glu)/(Wt-Glycerol), where PA-Glu = peak area of d-glucose and PA-Glycerol = peak area of glycerol (internal standard), Wt-Glu = mass of d-glucose in 1 mL of glucose/glycerol standard solution (10 mg) and Wt-Glycerol = mass of glycerol in 1 mL of glucose/glycerol standard solution (10 mg). 7. Calibrate the area of the chromatogram to be measured for SDFS. Use a 100 µL liquid chromatography syringe to fill the 50 µL injection loop with the retention time standard (maltodextrins, maltose and glycerol each at about 10 mg mL−1). Inject in duplicate. Determine the demarcation point between DP2 and DP3 oligosaccharides (disaccharide maltose versus higher oligosaccharides). 8. Determine peak areas of SDFS (PA-SDFS) and internal standard (PA-IS) in chromatograms of sample extracts. Inject sample extracts (Section 6.5.5, step 3) in the HPLC system. Record the areas of all peaks of DP greater than the DP2/DP3 demarcation point as PA-SDFS (Figure 6.3). Record the peak area of internal standard as PA-IS.

6.5.6  Calculations for HMWDF (IDF + SDFP), SDFS and TDF 6.5.6.1 Determination of HMWDF

Blank (B) (mg) = [(BR1 + BR2)]/2 – PB – PA

where BR1 and BR2 = residue mass (mg) for two duplicate blank determinations and PB and PA = mass (mg) of protein and ash, respectively, determined on the first and second blank residues.

HMWDF (mg/100 g) = [(R1 + R2)/2 – PB – PA – B]/(M1 + M2)/2] × 100 HMWDF (% w/w) = HMWDF (mg/100 g)/1000

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where R1 = residue mass 1 from M1 (mg), R2 = residue mass 2 from M2 (mg), M1 = test portion mass 1 (g), M2 = test portion mass 2 (g), PA = ash mass (mg) from R1 and PB = protein mass (mg) from R2. Calculations can be simplified by using an Excel-based calculator available at https://secure.megazyme.com/Rapid-Integrated-Total-Dietary-Fiber-Assay-Kit.

6.5.6.2 Determination of SDFS

SDFS (mg/100 g) = Rf ×Wt-IS × (PA-SDFS)]/(PA-IS) × 100/M SDFS (% w/w) = SDFS (mg/100 g)/1000

where Rf = the response factor (Section 6.5.5, steps 4–6), Wt-IS = mass of internal standard (mg) contained in 1 mL of internal standard solution (100 mg mL−1) pipetted into the sample before filtration, PA-SDFS = peak area of SDFS, PA-IS = peak area of internal standard and M = the test portion mass M1 or M2 of the sample, the filtrate of which was concentrated and analyzed by HPLC.

6.5.6.3 Determination of TDF

TDF (% w/w) = HMWDF (% w/w) + SDFS (% w/w)

Calculations can be simplified by using an Excel-based calculator available at https://secure.megazyme.com/Rapid-Integrated-Total-Dietary-Fiber-Assay-Kit.

6.5.7  Safety Considerations 1. Sodium azide should only be used after reading the appropriate Safety Data Sheet. Sodium azide must be weighed and dispensed only under a laboratory hood and should never be added to solutions of low pH. Acidification of sodium azide releases a poisonous gas. Use appropriate personal protection gear and work under a laboratory hood. 2. PAA and/or AMG are allergenic to some people. In such cases, the enzymes should be dissolved in water and precipitated with ammonium sulfate by a non-allergic person. All users should avoid contact with these solutions and, if the enzyme, is spilled, it should be thoroughly mopped up with a wet washcloth. Prepare the enzymes as an ammonium sulfate suspension as follows. Engage an analyst who is not allergic to perform this operation. Gradually add 5 g of PAA/AMG powder mixture (PAA 40 KU g−1 plus AMG 17 KU g−1) to 70 mL of cold, distilled water in a 200 mL beaker on a magnetic stirrer in a laboratory fume hood and stir until the enzymes are completely dissolved (about 5 minutes). Add 35 g of granular ammonium sulfate and dissolve by stirring. Adjust the volume to 100 mL with ammonium sulfate

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solution (50 g/100 mL) and store at 4 °C. This preparation contains PAA at 2 KU mL−1 and AMG at 0.85 KU mL−1 and is stable at 4 °C for three months. 3. Caution should be exercised when using a combustion analyzer for the determination of protein in the residue. Celite volatilized from the sample can clog the transfer lines of the unit.

6.6  A  ddressing Each of the Limitations of the INTDF Procedure 6.6.1  Optimization of PAA and AMG To determine the optimum levels of PAA and AMG for starch hydrolysis, incubations were performed as described in Section 6.5 with either a saturating level of AMG and varying levels of PAA, or with saturating levels of PAA and varying levels of AMG, and glucose released on the hydrolysis of non-resistant starch monitored. The results obtained with incubations performed in Fisherbrand bottles with magnetic stirring are shown in Figure 6.4. Similar results were obtained if the bottles were incubated in a shaking water-bath in orbital motion. The time course of hydrolysis of Hylon VII (Figure 6.4a and d), RMS (Figure 6.4b and e) and Fibersym (Figure 6.4c and f) with varying levels of PAA and AMG at pH 6 is shown. PAA is saturating at >1.5 KU per incubation and AMG is saturating at >0.8 KU per incubation. Thus the final incubation conditions chosen for the analysis of samples for dietary fiber involves the suspension of 1 g of sample in 40 mL of maleate buffer (pH 6) and incubation with stirring at 170 rpm and 37 °C with PAA (4 KU) and AMG (1.7 KU) for four hours. Resistant maltodextrins are not formed under these conditions. In fact, the resistant maltodextrins found on the hydrolysis of non-resistant starch under the incubation conditions of the INTDF method (63-α-d-glucosyl-maltotriose and 63,65-di-α-D glucosyl-maltopentaose52) are completely hydrolysed.52 Under these incubation conditions, there is complete hydrolysis of non-resistant starch in wheat starch and near-complete hydrolysis of starch in regular maize starch. The dietary fiber values for most samples analyzed was in close agreement with those obtained with AOAC Method 2009.01, but much higher values were obtained for Fibersym (RS4) and Hylon VII (high amylose maize starch) (Table 6.1).

6.6.2  Choice of HPLC Column In the development of AOAC Method 2009.01,41 a Waters Sugar-Pak HPLC column was used for the analysis of SDFS with D-sorbitol as the internal standard. It was subsequently found that the fructo-oligosaccharides produced by an acidic or enzymatic hydrolysis of inulin are underestimated. One of the hydrolysis products, the trisaccharide fructosyl-β-(2-1)-fructosyl-β-(2-1)-fructose (inulinotriose; F3), elutes from the Waters Sugar-Pak HPLC column at the same point as the disaccharides maltose, sucrose and lactose and thus is

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Figure 6.4  Effect  of the concentration of PAA and of AMG on the extent of hydrolysis of (a, d) Hylon VII, (b, e) RMS and (c, f) Fibersym in Fisherbrand bottles with magnetic stirring at 170 rpm at 37 °C and pH 6.0. Samples (a–c) were incubated with 1.7 KU AMG and varying levels of PAA. Samples (d–f) were incubated with 6 KU of PAA and varying levels of AMG. Samples were analyzed for free d-glucose. Reproduced with permission from B. V. McCleary, N. Sloane, A. Draga and L. Lazewska, Measurement of Total Dietary Fiber Using AOAC Method 2009.01 (AACC International Approved Method 32–45.01): Evaluation and Updates, Cereal Chem., 2013, 90, 396– 414, © AACC International and Wiley-VCH Verlag GmbH & Co. KgaA.

not included in the dietary fiber value as, by definition, only oligosaccharides of DP3 or greater are counted. This problem has been resolved by performing HPLC on TSKgel G2500PWXL columns,38 which gives a clear delineation between F3 and disaccharides (Figure 6.3). Glycerol is the internal standard of choice with these columns. Thus glycerol must be excluded from the enzyme preparations used in the assay. PAA and AMG are used as a stable powder mixture, which is dissolved immediately before use, or, alternatively, are prepared as a stabilized ammonium sulfate suspension (see Section 6.5.7, step 2). Protease is used as an ammonium sulfate suspension.

6.6.3  Preparation of Samples for HPLC Oligosaccharides in the SDFS fraction are separated on TSKgel G2500PWXL columns and quantitated by refractive index detection. Salt in the sample must be removed. Traditionally, this has been performed by percolation

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Table 6.1  HMWDF,  SDFS and TDF values obtained for a range of samples using the

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integrated total dietary fiber (INTDF) and rapid integrated total dietary fiber (RINTDF) methodsa,.b

AOAC Method 2009.01 (INTDF method)

Sample Wholemeal bread Oat bran Weetabix Kellogg all bran Wholewheat pasta Semi-ripe banana Sweet corn (tinned) Garden peas (tinned) Broccoli Carrots Fibersym Hylon VII Polydextrose

HMWDF (% w/w)

SDFS (% w/w)

RINTDF method

TDF (HMWDF + SDFS) HMWDF (% w/w) (% w/w)

SDFS (% w/w)

TDF (HMWDF + SDFS) (% w/w)

12.4

1.8

14.2

12.0

1.5

13.5

18.9 9.8 26.6

0.6 2.8 3.9

19.5 12.6 30.5

19.9 10.1 28.1

1.4 1.4 3.6

21.3 11.6 31.7

9.9

2.8

12.7

10.1

2.2

12.3

30.2

0.9

31.1

30.2

1.4

31.6

12.7

0.4

13.1

12.4

0.5

12.9

29.1

1.4

30.5

29.1

2.2

31.3

28.1 21.8 28.6 48.6 1.5

0.4 0.6 1.1 0.5 83.3

28.5 22.4 29.7 49.3 84.8

29.7 22.2 59.2 58.8 1.1

0.6 1.2 1.0 0.0 83.3

30.3 23.4 60.2 58.8 84.4

a

 MWDF, high molecular weight dietary fiber; SDFS, soluble dietary fiber that remains soluble H in the presence of 78% aqueous ethanol; TDF, total dietary fiber. HMWDF determined gravimetrically; SDFS determined by HPLC using TSK gel G2500PWXL columns; and TDF calculated as the sum of HMWDF and SDFS.

b

of the sample through a column of a mixture of cation and anion resins,38,41 a tedious and time-consuming operation. This can be greatly simplified by using an in-line Bio-Rad deionizing pre-column. These cation and anion deionizing cartridges are costly and only 15–20 samples can be analyzed before the resins are exhausted. The most effective use of these deionizing cartridges can be achieved by the pre-removal of the bulk of the salt in the sample with cation and anion resins in a tube.54 This pre-removal of the bulk of the salt extends the useful life of the Bio-Rad deionizing columns by 10–15 fold, significantly reducing costs and decreasing the sample preparation time. The effect of this process of sample desalting is clearly shown in Figure 6.2, where a mixture of glycerol and d-glucose in the buffers used in the RINTDF procedure were chromatographed through the TSKgel G2500PWXL columns (1) before desalting, (2) after desalting with resins and (3) with the desalted sample applied with the Bio-Rad deionizing pre-columns in place. This process removes all the salt from the sample, allowing the accurate determination of SDFS in the applied samples.

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6.6.4  R  ecovery of Polysaccharides and Non-digestible Oligosaccharides Under the RINTDF Assay Method The purity and activity of the enzymes used in the RINTDF method were checked using a range of polysaccharides and casein. The dietary fiber values obtained for β-glucan, pectin, larch galactan and wheat arabinoxylan are similar to those obtained using AOAC Methods 2009.01 and 991.43,53 and wheat starch and casein are completely hydrolyzed, demonstrating the purity and activity of the enzymes used. With the RINTDF method, a higher value was obtained for the HMWDF fraction of Hylon VII (58.8%) than was obtained with the AOAC Method 2009.01 (49.3%). Dietary fiber values obtained for a range of samples are shown in Table 6.1. Of particular note is the much higher value obtained for Fibersym with the RINTDF method than with AOAC Method 2009.01. This value is still considerably lower than the value reported using AOAC Method 985.29 (about 86% w/w). The value of dietary fiber obtained for Fibersym with AOAC Method 985.29 is critically dependent on the incubation temperature. In our laboratory, the dietary fiber value obtained for a Fibersym sample using the AOAC Method 985.29 with α-amylase incubation at about 100 °C resulted in a high dietary fiber value of about 78% w/w. However, if the incubations are performed at 95 °C, the measured dietary fiber value decreases dramatically to about 43% w/w and at 90 °C to 33% (Figure 6.5). This clearly shows that determination of the dietary fiber value of Fibersym using AOAC Method 985.29 is extremely method-dependent and uses a method that has little relationship to physiological conditions in the human small intestine. This dramatic decrease in the hydrolysis of Fibersym with an increase in temperature is most likely related to the significant rate of inactivation of the thermostable α-amylase as incubation temperatures increase from 80 to 100 °C. In the RINTDF method, incubations are performed with pancreatic α-amylase at 37 °C and pH 6.0, consistent with the physiological conditions in the human small intestine. In the RINTDF method, AMG is added to the incubation mixture to hydrolyze maltose to remove its inhibitory action on PAA. As AMG does have some action on starch, it is conceivable that the extent of hydrolysis of Fibersym by PAA might be inflated by the direct action of the AMG on the starch, rather than the removal of the inhibitory effect of maltose. To check this, AMG in the incubation mixture was replaced by an α-glucosidase from Bacillus stearothermophilis (Megazyme cat. no. E-TSAGL), which readily hydrolyzes maltose, but has no action on soluble starch or on the terminal 1,6-α-linked glucosyl residues in maltosaccharides (e.g. as in panose) (Table 6.2). The degree of hydrolysis of Fibersym by PAA in the presence of 10 KU of α-glucosidase is similar to that with PAA plus AMG (Figure 6.6), indicating that the major effect of the AMG in the RINTDF procedure is the removal of the maltose inhibition of the PAA. The amount of B. stearothermophilis α-glucosidase used in the incubation (10 KU) is much higher than the level of AMG used (1.7 KU). This is to account for the difference in stability of the two enzymes under these incubation conditions. AMG is completely stable

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Figure 6.5  Hydrolysis  of wheat starch and Fibersym under conditions exactly as

described for AOAC Method 985.29, but with incubations performed at 70, 80, 90, 95 and 100 °C. Incubations with protease and AMG were exactly as described in AOAC Method 985.29. A sample of the incubation solution (10 mL) was removed and immediately filtered. A sample (1 mL) was added to 25 mL of distilled water and mixed. An aliquot (0.1 mL) of this was incubated with 30 U of AMG at 40 °C for 15 minutes and measured for glucose using a glucose oxidase/peroxidase reagent. Non-resistant starch was calculated from glucose value and the dietary fiber value (resistant starch) was determined by subtracting the non-resistant starch from the total sample weight. The remaining α-amylase activity in the incubation mixtures was determined using the Ceralpha method.

with no loss of activity over the four hour incubation period. In contrast, B. stearothermophilis α-glucosidase lost 95% of activity after just 2 hours. Dietary fiber values obtained using AOAC Method 2009.01 and the RINTDF method for a range of non-digestible oligosaccharides are shown in Table 6.3.53 For all oligosaccharides except isomalto-oligosaccharides and fructo-oligosaccharides, the values obtained with the two methods are similar, even though much higher levels of both PAA and AMG are used in the RINTDF method. This again demonstrates the high purity of the enzymes used in the procedures. For isomalto-oligosaccharides, a dietary fiber value of 10.8% was obtained with the RINTDF method. This value is similar to that obtained with AOAC Methods 985.29 and 991.43 (about 8%). In separate research55 on the hydrolysis of various oligosaccharides by a preparation of mucosal α-glucosidases from the small intestine of pigs, completely hydrolysis of isomalto-oligosaccharides was observed. The higher value obtained for fructo-oligosaccharides using the RINTDF method compared to AOAC method 2009.01 reflects accurate measurement of inulinotriose by the RINTDF method.

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Table 6.2  Relative  rates of hydrolysis of maltodextrins and starch by Bacillus stearo-

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thermophilis α-glucosidase.

Relative rate of hydrolysisa

Substrate

Structure

Maltose Maltotriose Maltotetraose

1,4-α-d-glucosyl-d-glucose 1,4-α-d-glucosyl-1,4-α-d-glucosyl-d-glucose 1,4-α-d-glucosyl-1,4-α-d-glucosyl-1,4-α-dglucosyl-d-glucose 1,4-α-d-glucosyl-1,4-α-d-glucosyl-1,6-α-dglucosyl-d-glucose Maltotriosyl-1,6-α-d-maltotriose

56.9

1,6-α-d-glucosyl-d-glucose 1,6-α-d-glucosyl-1,6-α-d-glucosyl-d-glucose 1,6-α-d-glucosyl-1,4-α-d-glucosyl-d-glucose 1,6-α-d-glucosyl-d-maltotriose

0.03 0.01 0.30 0.03

Polymer consisting of 1,4-linked and 1,6-linked d-glucose

0.002

Isopanose Maltotriosyl 1,6-α-d-maltotriose Isomaltose Isomaltotriose Panose 63-α-d-glucosyl maltotriose Soluble starch

100 97.6 94.9 63.1

a

An aliquot (0.2 mL) of appropriately diluted enzyme was added to 0.5 mL of malto-oligosaccharide (10 mM) or 0.5 mL of starch (10 mg mL−1) in 100 mM sodium phosphate buffer (pH 6.5) pre-equilibrated at 40 °C. Tubes were incubated for 0, 5, 10 and 15 minutes and the reaction was terminated by placing the tube into a boiling water-bath for 2 minutes. The released glucose was measured with a glucose oxidase/peroxidase reagent. Glucose values for maltose and isomaltose were divided by two to allow for the fact that two molecules of glucose are released for each hydrolytic cleavage.

Figure 6.6  Hydrolysis  of Fibersym under the incubation conditions of the RINTDF

method: (a) with PAA and AMG as described in the standard method; (b) with PAA as described in the standard method, but with no AMG; and (c) with PAA as described in the standard method, but with B. stearothermophilis α-glucosidase (10 KU) in place of AMG. Aliquots of this solution were removed and analyzed. Non-resistant starch was calculated from glucose value.

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Table 6.3  Recovery  of oligosaccharides of DP ≥3 in original samples and on incuba-

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tion of the samples according to AOAC 2009.01 and the rapid integrated total dietary fiber (RINTDF) method.a Recovery of oligosaccharides of DP > 3 as a percentage of total carbohydrate in the sample (% w/w)

Sample Neosugars Raftilose P95 Polydextrose Fibersol 2 Galacto-oligosaccharides Xylo-oligosaccharides Raffinose Isomalto-oligosaccharides AdvantaFiber

Original AOAC method oligosaccharides 2009.01b (INTDF) RINTDF methodb 93.0 89.0 84.3 88.5 76.0 78.0 99.0 65.4

92.9 76.2 85.1 83.4 70.6 78.6 99.0 29.0

92.8 88.2 82.5 82.4 72.0 76.2 98.0 10.8

a

 alculated from HPLC patterns as areas under the peaks for oligosaccharides of DP ≥ 3 (see C Figure 6.3) as a percentage of combined area for all peaks from the sample. b Reproduced in part from B. V. McCleary, N. Sloane and A. Draga, Starch, 67 (9-10), 860–883 with permission from John Wiley and Sons, © 2015 The Authors Starch Published by WileyVCH Verlag GmbH & Co. KGaA.54

6.6.5  Safety Considerations Sodium azide is an effective preservative, but there are concerns over its routine use in analytical laboratories. In the original integrated dietary fiber procedures (AOAC Method 2009.01/2011.25), incubations are performed for 16 hour at 37 °C in maleate buffer at pH 6.0. To prevent microbial contamination of the incubation mixture over this time period, sodium azide (0.02% w/v) was included in the buffer. With the RINTDF procedure, the incubation time is reduced to four hours, so the potential problems associated with microbial contamination are considerably decreased. This was verified by performing incubations under the standard RINTDF procedure (no sodium azide in the buffer) with glucose (1 g) in the incubation mixture. No decrease in determined glucose was observed in incubation mixtures held up to eight hours, indicating that microbial contamination was not a problem. However, in routine use it is suggested that buffer should either be freshly prepared or sterilized before use by heating to about 90 °C in a boiling water-bath or in a microwave oven.

6.7  I nter-laboratory Evaluation of the RINTDF Method The RINTDF method has been subjected to inter-laboratory evaluation under the auspices of the International Association of Cereal Science and Technology and the AACCI according to the protocols of AOAC International. A report on the study has been submitted to the three associations. Thirteen

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Table 6.4  Inter-laboratory  study results for total dietary fiber in foods using the

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rapid integrated total dietary fiber (RINTDF) method. Statistical evaluation according to AOAC International statistics format.a

Parameter

A&D

B&F

C&J

E&H G&N I&M

K&O L&P

No. of laboratories/ analysts Mean percentage sr sR RSDr (%) RSDR (%)

12

13

12

12

13

13

12

13

60.62 0.74 4.67 1.22 7.70

23.70 0.67 0.99 2.81 4.17

29.37 0.36 0.78 1.22 2.64

6.79 0.29 0.91 4.32 13.38

16.15 0.39 0.85 2.41 5.29

19.28 0.29 1.74 1.51 9.01

21.09 0.43 0.57 2.05 2.72

10.76 0.68 0.86 6.34 8.02

a

 amples: A & D = Fibersym; B & F = kidney beans (canned, washed and lyophilized); C & J = S bran cereal; E & H = defatted cookies containing fructo-oligosaccharides; G & N = oat bran; I & M = defatted cookies containing polydextrose and RS2; K & O = dark rye crispbread; L & P = wholemeal bread. sr, within-laboratory variability; RSDr, within-laboratory relative variability; sR, between-laboratory variability; RSDR, between-laboratory relative variability.

collaborating laboratories were involved and they analyzed 16 samples provided as eight blind duplicates. The results of the study are shown in Table 6.4. In total, only four sets of data from the 104 sets submitted were statistically excluded as outliers. The dietary fiber content of the eight test pairs ranged from 6.79 to 60.6%. Total dietary fiber was calculated as the sum of HMWDF (IDF plus SDFP) and SDFS. For total dietary fiber, the within-laboratory variability ranged from 0.29 to 0.74 and the between-laboratory variability ranged from 0.57 to 4.67. The within-laboratory relative variability ranged from 1.22 to 6.34% and the between-laboratory relative variability ranged from 2.64 to 13.38%. In previously adopted methods, the between-laboratory variability ranged from 0.04 to 9.49 and the between-laboratory relative variability from 1.58 to 66.25. The collaborating laboratories were: Megazyme, Bray, County Wicklow, Ireland; Medallion Laboratories/General Mills, Golden Valley, MN, USA; Agriculture and Agri-Food Canada/Agriculture et Agroalimentaires Canada, University of Manitoba – Winnipeg, Manitoba, Canada; Grain Growers Limited, PO Box 7, North Ryde, NSW, Australia; Sanitarium Development and Innovation Analytical Department, Cooranbong, NSW, Australia; CRDS Tienen, Central Department Research, Development and Services, Tienen, Belgium; Finnish Food Safety Authority Evira, Chemistry and Toxicology Research Unit, Helsinki, Finland; Matsutani Chemical Company, Itami City, Hyogo, Japan; NEOTRON SPA, Stradello Aggazzotti, Modena, Italy; Eurofins Food Testing Netherlands BV, Heerenveen, Netherlands;11 Kellogg Company, Battle Creek, MI, USA; Japanese Food Research Laboratories, Japan; Nestlé, Food Science and Technology Carbohydrates; and Nestlé Research Centre, Lausanne, Switzerland. The results of this study have been submitted to the technical committees of International Association of Cereal Science and Technology, AACC International and AOAC International and are currently under review.

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6.8  Conclusions The RINTDF method is an improvement on AOAC Method 2009.01 and is suitable for the measurement of total dietary fiber as defined by Codex Alimentarius. The method simulates physiological conditions in the human small intestine and all the problems identified with AOAC Method 2009.01 (the INTDF method) have been resolved. The method is robust, reliable and reproducible, as demonstrated by the results of the inter-laboratory study. It can be applied to all food samples and to the full range of dietary fiber components available at this point in time.

References 1. R. A. McCance and R. D. Lawrence, Medical Research Council Special Report Series no.135, H.M. Stationery Office, London, 1929. 2. R. D. Williams and W. Olmstedt, J. Biol. Chem., 1935, 108, 653–666. 3. E. Hipsley, Br. Med. J., 1953, 2, 420–422. 4. H. C. Trowell, Atherosclerosis, 1972, 16, 138–140. 5. H. C. Trowell, D. A. T. Southgate, T. M. S. Wolever, A. R. Leeds, M. A. Gassull and D. J. A. Jenkins, Lancet, 1976, 1, 967. 6. B. Thomas, Getreide, Mehl Brot, 1972, 26, 158–165, 168–169. 7. E. M. Hellendoorn, M. G. Noordhoff and J. Slagman, J. Sci. Food Agric., 1975, 26, 1461–1468. 8. D. A. Schaller, Fd. Prodn. Dev., 1976, 11, 70–72. 9. P. Van Soest and R. H. Wine, J. Assoc. Off. Agric. Chem., 1967, 50, 50–55. 10. AACC International Approved Methods of Analysis, Method 32–20.01, AACCI, St. Paul MN, 11th edn, 2000. 11. I. Furda, Cereal Foods World, 1977, 22, 252–254. 12. I. Furda, in The Analysis of Dietary Fiber in Food, ed. W. P. T. James and O. Theander, Marcel Dekker, New York, 1981, pp. 163–172. 13. T. F. Schweizer and P. Würsch, J. Sci. Food Agric., 1979, 30, 613–619. 14. T. F. Schweizer and P. Würsch, in The Analysis of Dietary Fiber in Food, ed. W. P. T. James and O. Theander, Marcel Dekker, New York, 1981, pp. 203–216. 15. N.-G. Asp and C.-G. Johansson, in The Analysis of Dietary Fiber in Food, ed. W. P. T. James and O. Theander, Marcel Dekker, New York, 1981, pp. 173–179. 16. L. Prosky, N.-G. Asp, I. Furda, J. W. DeVries, T. F. Schweizer and B. F. Harland, J. AOAC Int., 1985, 68, 677–679. 17. L. D. Lee, L. Prosky and J. W. DeVries, J. AOAC Int., 1992, 75, 395–416. 18. AOAC International 2007, Methods of Analysis, AOAC Intl., Gaithersburg, MD, 18th edn, 2007. 19. D. A. T. Southgate, J. Sci. Food Agric., 1969, 20, 331–335. 20. D. A. T. Southgate, in The Analysis of Dietary Fiber in Food, ed. W. P. T. James and O. Theander, Marcel Dekker, New York, 1981, pp. 1–19. 21. E. M. Widdowson and R. A. McCance, Biochem. J., 1935, 29, 151–156. 22. O. Theander and P. Åman, Swed. J. Agric. Res., 1979, 9, 97–106.

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23. D. A. T. Southgate, G. H. Hudson and H. N. Englyst, J. Sci. Food Agric., 1978, 29, 979–988. 24. H. N. Englyst, in The Analysis of Dietary Fiber in Food, ed. W. P. T. James and O. Theander, Marcel Dekker, New York, 1981, pp. 173–189. 25. H. N. Englyst and J. H. Cummings, Analyst, 1984, 109, 937–942. 26. H. N. Englyst and G. Hudson, Food Chem., 1987, 24, 63–76. 27. H. N. Englyst, S. M. Kingman and J. H. Cummings, Eur. J. Clin. Nutr., 1992, 46, S33–S39. 28. M. E. Quigley and H. N. Englyst, Analyst, 1992, 117, 1715–1718. 29. Anon, The definition of dietary fiber, Cereal Foods World, 2001, 46, 112–126. 30. Institute of Medicine, Dietary Reference Intakes: Energy, Carbohydrates, Fiber, Fat, Fatty Acids, Cholesterol, Protein and Amino Acids, National Academies Press, Washington, DC, 2002. 31. Codex Committee on Nutrition and Foods for Special Dietary Uses (CCNFSD; ALINORM 06/29/26), 27th Session, Bonn, Germany, 21–25th November, 2005. 32. WHO/FAO Meeting of Carbohydrates Experts, Report CRD 19, Geneva, 17– 18th July, 2006. 33. Codex Committee on Nutrition and Foods for Special Dietary Uses 28th Session, Chiang Mai, Thailand, 30th October–3rd November, 2006. 34. Guidelines on nutrition labelling CAC/GL 2-1985 as last amended 2010, Codex Alimentarius, FAO, Joint FAO/WHO Food Standards Programme, Secretariat of the Codex Alimentarius Commission, Rome, 2010. 35. H. Hoebregs, J. AOAC Int., 1979, 80, 1029–1037. 36. B. V. McCleary, A. Murphy and D. C. Mugford, J. AOAC Int., 2000, 83, 356–364. 37. S. A. S. Craig, J. F. Holden and M. J. Khaled, J. AOAC Int., 2000, 83, 1006–1012. 38. D. T. Gordon and K. Okuma, J. AOAC Int., 2002, 85, 435–444. 39. J. De Slegte, J. AOAC Int., 2002, 85, 417–423. 40. B. V. McCleary, M. McNally and P. Rossiter, J. AOAC Int., 2002, 85, 1103–1111. 41. B. V. McCleary, Anal. Bioanal. Chem., 2007, 389, 291–308. 42. B. V. McCleary, J. W. DeVries, J. I. Rader, G. Cohen, L. Prosky, D. C. Mugford and K. Okuma, J. AOAC Int., 2010, 93, 221–233. 43. B. V. McCleary, J. W. DeVries, J. I. Rader, G. Cohen, L. Prosky, D. C. Mugford and K. Okuma, J. AOAC Int., 2012, 95, 824–844. 44. B. V. McCleary, J. W. DeVries, I. L. Rader, G. Cohen, L. Prosky, D. C. Mugford, M. Champ and K. Okuma, Cereal Foods World, 2011, 56, 238–247. 45. Codex Committee on Methods of Analysis and Sampling, CRD16 from the Thirty-third Session of the Codex Committee on Methods of Analysis and Sampling, FAO, Rome, 2012, available at: ftp://ftp.fao.org/codex/meetings/CCMAS/CCMAS33/CRD/ma33_CRD16e.pdf. 46. N. W. Read, J. Cammack, C. Edwards, A. M. Holgate, P. A. Cann and C. Brown, Gut, 1982, 23, 824–828.

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47. K. P. Geboes, A. Luypaerts, P. Rutgeerts and K. Verbeke, Aliment. Pharmacol. Ther., 2003, 18, 721–729. 48. B. Geypens, R. Bennink, M. Peeters, P. Evenepole, L. Mortelmans, B. Maes, Y. Ghoos and P. Rutgeerts, J. Nucl. Med., 1999, 40, 1451–1455. 49. R. Sadik, H. Abrahamsson, E. Bjornsson, A. Gunnarsdottir and P.-O. Stotzer, Scand. J. Gastroenterol., 2003, 38, 1039–1044. 50. P. O. Stotzer and H. Abrahamsson, Neurogastroenterol. Motil., 2000, 12, 415–419. 51. N. Zarate, S. D. Mohammed, E. O'Shaughnessy, M. Newell, E. Yazaki, N. S. Williams, P. J. Lunniss, J. R. Semler and S. M. Scott, Am. J. Physiol. Gastrointest. Liver Physiol., 2010, 299, G1276–G1286. 52. B. V. McCleary, N. Sloane, A. Draga and L. Lazewska, Cereal Chem., 2013, 90, 396–414. 53. B. V. McCleary, N. Sloane and A. Draga, Starch, 2015, 67, 860–883. 54. Anon, Rapid Integrated Total Dietary Fiber Test Kit Booklet, Megazyme K-rintdf 09/17, 2017. 55. K. Tanabe, S. Nakamura and T. Oku, Food Chem., 2014, 151, 539–546.

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Chapter 7

Resistant and Slowly Digested Starch in Grain Products P. C. Drawbridge* and T. Beta University of Manitoba, Department of Food and Human Nutritional Sciences, 35 Chancellors Circle, Winnipeg, Manitoba, R3T 2N2, Canada *E-mail: [email protected]

7.1  Introduction Starch is the glucose reserve for metabolism in plants. Starch is composed of two water-insoluble homoglucans – amylose and amylopectin – and the proportion and detailed structure of these polymers influence the digestibility of starch. The digestibility of starch varies from rapidly digestible, slowly digestible to non-digestible. The term resistant starch (RS) was first coined by H. Englyst, H.S. Wiggins and J.H. Cummings in their 1982 paper ‘Determination of the non-starch polysaccharides in plant foods by gas–liquid chromatography of constituent sugars as alditol acetates’. Although the determination of RS in foodstuffs was not their primary objective, this paper is highly cited as the first documented measurement of RS. RS was defined as ‘retrograded starch that cannot be hydrolysed by α-amylase and pullulanase, due to food processing’. This definition has evolved as more precise methods of RS analysis have overcome the limitations of the earlier technique. The European FLAIR Concerted Action on Resistant Starch defined RS as ‘the sum of starch and products of starch degradation not absorbed in the small   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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intestine of healthy individuals’. A fifth type of RS (RS5) is excluded from the current definition by Codex Alimentarius (2004):    Resistant starch (RS) is defined as the fraction of starch not absorbed in the small intestine. It consists of physically enclosed starch (RS1), certain types of raw starch granules (RS2) and retrograded amylose (RS3). Modified starches used as food additives may also be partially resistant (RS4).2    RS5 includes amylose–lipid complexes that have been found to be resistant to digestion by enzymes in the small intestine and may even be formed in vivo.3 The amount of RS in grains is related to the amylose content, whereas the slowly digestible starch (SDS) content is primarily associated with amylopectin.4,5 The differences in digestibility of starches suggest that they provide distinct health benefits. Rapidly digestible starch (RDS) provides energy and induces variable post-prandial glycaemia.5 Unlike glycaemic carbohydrates, RS does not induce a glycaemic response, behaves as a form of dietary fibre5 and passes undigested to the large bowel, where it is fermented, yielding beneficial short-chain fatty acids (SCFAs). Consequently, the behaviour and health effects of RS differ from those of traditional fibres. The latter consist mainly of non-starch polysaccharides, which, in general, are non-fermentable. Manipulation of the amylose to amylopectin ratio in cereal grains facilitates the production of starches with novel functional properties and improved health benefits.6 The beneficial health effects of RS result from the SCFAs produced by its fermentation by gut microbiota. The interest in RS is prominently due to the production of SCFAs in the bowel. However, RS not only contributes to large bowel health through fermentation products, but helps manage glycaemic control and obesity when replacing RDS in the diet. SDS induces a slow, prolonged glycaemic response, thereby providing glycaemic control and avoiding the detrimental effects of sustained high blood sugar.5 Starch-rich foods derived from cereals are inaccurately viewed negatively by many consumers, undermining their contribution to human health.6 Fibre and other phytochemicals based on non-starch polysaccharides are more commonly associated with the health benefits of whole grains, although SDS and RS also contribute to the ‘whole grain package’7 of protective agents, thereby necessitating their extensive investigation. A brief description of the five types of RS is provided here along with their sources, formation and the factors affecting their content in cereal grains and grain products. In addition, the benefits of SDS and RS to human health are addressed and the potential for the incorporation of RS into grain-based functional foods.

7.2  Types of Resistant Starch RS1 are physically inaccessible, unavailable starches located in the plant cells of unmilled or partially milled grains. The intact or partially intact hull (and sometimes adjacent outer pericarp layers) resulting from partial

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milling provides a physical barrier and prevents digestive enzymes from acting on the starch. RS2 are native granular starches with structures that reduce their susceptibility to enzymes.8 Their highly dense and partially crystalline structures hinder digestion by enzymes.9 Uncooked starch-containing foods (e.g. raw bananas, legumes and potatoes) and high-amylose corn starches serve as sources of RS2. The company Ingredion Inc. markets an RS2 product called Hi-Maize® made from high-amylose maize. RS3 are retrograded starches formed after heat–moisture treatments of starch gelatinization and cooling. RS3 may be present in cooked and cooled foods, such as corn starch, potatoes and canned peas and beans.9,10 Novelose®, a source of RS3, is marketed in Europe. The X-ray diffraction pattern of RS3 shows a crystalline B-type structure. The crystalline structure consists of short linear segments of organized α-1 → 4-glucans.9 If gelatinized RS is stored at high temperatures, an A-type crystalline structure may be achieved. RS4 are chemically modified starches that have been made resistant to enzymes through oxidation, esterification, etherification, transglycosidation, cross-linking or by adding substituents (e.g. hydroxypropyl). For example, introducing glycosidic bonds via heat treatment that are neither α-1 → 4 or α-1 → 6 reduces the accessibility to amylolytic enzymes.9 An example of a marketed RS4 is Fibersym® flour. Strict regulations limit the amount of chemicals used to produce RS4 and the USA and Japan are currently the only countries permitting RS4 ingredients in foods. RS5 are amylose–lipid complexes. Starch can interact with lipids, fatty acids, alcohols, amylose and long chains of amylopectin.11 On binding lipids, amylases are unable to break down starch due to physical barriers. Another potential type of RS is resistant maltodextrins, such as the commercially available Fibersol-2® and Nutriose®. The starch molecules in resistant maltodextrins have been rearranged to become soluble and indigestible.12 Resistant maltodextrins are currently not included among the different types of RS.

7.3  S  tarch Synthesis in Grains and Formation of Resistant Starch Starch synthesis begins when sucrose is produced via photosynthesis in the leaves. Sucrose is transported to the starch-storing endosperm where it is converted to ADP-glucose, which is the substrate for the starch synthase enzymes. In cereal grains, ADP-glucose is produced primarily in the cytosol by cytosolic ADP-glucose pyrophosphorylase, although its synthesis also occurs in the amyloplast. The amyloplast is the organelle for starch synthesis and storage. In the cytosol, sucrose is first converted to UDP-glucose and then to glucose-1-phosphate. Glucose-1-phosphate is converted to ADP-glucose, which, in turn, enters the amyloplast through a transporter. Starch synthesis occurs through the action of ADP-glucose pyrophosphorylase on

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ADP-glucose. Alternatively, some of the glucose-1-phosphate in the cytosol may be converted to glucose-6-phosphate rather than ADP-glucose directly. Glucose-6-phosphate is capable of entering the amyloplast, where ADP-glucose is produced and converted into starch. Thus both the cytosol and amyloplast serve as sites of ADP-glucose production. Other enzymes are also involved in starch synthesis. Several isoforms of starch synthase and starch-branching enzyme (SBE), as well as starch-debranching enzyme, have unique roles in the biosynthesis of starch. Starch synthases add a glucosyl unit to the non-reducing end of a glucose chain, where each unit is joined by an α-1-4 linkage. Several isoforms of starch synthase may coexist in the endosperm, including granule-bound starch synthase I, starch synthase I, starch synthase II and starch synthase III. In addition to starch synthase, SBEs cleave and transfer a linear glucose chain to a glucose residue through an α-1-6 linkage, thereby forming a branch. Much of the branching complexity of amylopectin molecules is attributed to the presence of multiple interacting isoforms of starch synthase and SBEs.13 The debranching enzymes isoamylase and pullulanase are also important for starch synthesis. However, the precise mechanisms and roles are unclear. Isoamylase cleaves α-1-6 linkages and may play an important part in facilitating the proper branching of amylopectin. In brief, two main hypotheses explain the role of isoamylase as (1) directly trimming ‘pre-amylopectin’ to an organized amylopectin form, which can be incorporated onto the granule surface,or (2) indirectly influencing starch synthesis by degrading soluble glucans that would otherwise be acted upon by starch synthase and SBEs and accumulate as phytoglycogen. In the latter, isoamylase prevents soluble glucans from hindering the activity of enzymes and altering the rate of starch synthesis.13 In summary, granule-bound starch synthase I is required for amylose synthesis, whereas starch synthase I, starch synthase II, starch synthase III and the debranching enzymes isoamylase and pullulanase are involved in the synthesis of amylopectin.14 Amylose and amylopectin assemble to form a starch granule consisting of alternating layers of crystalline and amorphous regions. The formation of the semi-crystalline granular structure generally requires amylopectin and occurs primarily as a physical, self-assembling process. However, a study by Carciofi et al.15 showed that amylopectin is not required for the crystallinity and integrity of the starch granule. The simultaneous suppression of genes encoding all SBEs resulted in starch granules with the unique thermal properties and crystallinity of amylose-only barley. However, amylopectin may still have an important role in rapid starch remobilization because the amylose-only barley exhibited a slower initial growth of shoots.15 The specific roles of enzymes involved in starch synthesis – particularly the starch-debranching enzymes, isoamylase and pullulanase – are nebulous. Also, the process of procurement of the substrate ADP-glucose for starch synthesis in grasses differs from the pathway described here for cereals.13 The

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reasons for the divergent pathways between grasses and cereals are unknown. Current evidence suggests that the flux of carbon into the endosperm is controlled by modulating the activity of ADP-glucose pyrophosphorylase,13,16 but other factors may be involved. Given that starch synthesis is a complex and poorly understood phenomenon, there is no clear answer to what differences in starch synthesis lead to RS, SDS and RDS. The exact mechanism of formation of RS in grains is yet to be fully understood. The starch properties that may contribute to varying degrees of starch digestibility include the ratio of amylose to amylopectin, the granular shape and the crystalline lattice structure. Rice mutants containing higher levels of RS and showing strongly aggregated and mostly round granules following cooking are associated with slower digestibility.17 Short chains with a degree of polymerization of 8–12 are negatively correlated with the digestibility of rice starch. Starch structure therefore appears to be a key factor influencing digestibility of starch. Plant starches can be classified according to the X-ray diffraction pattern given by their crystalline lattices, namely types A, B and C.18 A-type starches are found in cereal endosperms, whereas B and C type starches are found in tubers and pea embryos, respectively. Types B and C starches are more resistant to digestion by amylase than type A starches.19,20 The shape of the granules also affects their resistance to digestion, with more irregular morphologies being more resistant.20 The physiochemical properties, starch granule morphology, starch weight distribution, activities of enzymes involved in starch biosynthesis and their interactions during the development of grain kernels may all have an impact on the RS content of grains.19 A study19 on three rice mutants with different RS contents, including high-RS mutants containing 9.04 and 4.5% RS and a low-RS mutant containing 0.8% RS, elucidated the effect that enzymes, grain development, granular structure, amylose content and amylopectin structure on the RS content of rice kernels. During the first five days following flowering, the high-RS mutants developed more slowly than the low-RS mutants. Grain development affects the formation of RS because several processes throughout grain filling affect the formation of starch granules. The RS content in rice increased with grain maturation and was positively correlated with the molecular mass distribution. Lower activities of ADP-glucose pyrophosphorylase and SBE, and higher activity of starch synthase and starch-debranching enzyme were observed in the high-RS rice. The differing levels of enzyme activities compared with common rice may explain the formation of small irregular starch granules with large spaces among them.19 The formation of RS and SDS in barley is also highly dependent on SBEs. In a novel study,21 which produced a high yield of an amylose-only starch, the genes encoding the synthases SBI, SBIIa and SBIIb were suppressed and the resultant granules were irregularly shaped, with multiple small elongated lobes. The granule size is also related to the RS content of barley. In

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high-RS barley varieties, a large amount of small granules maize > barley > rice > triticale > oats > wheat. Information regarding the digestibility of starch in millet is limited. Based on the results of Srichuwong et al.,30 millet is more digestible than sorghum, but less digestible than maize. Current published values of the RS content of grains are highly variable and are dependent on the grain cultivar and the method of measurement. Researchers from CSIRO in Australia have investigated the development of wheat31 and barley32 cultivars with a low GI and a high-RS content.33 Further work in the area of genetically modified cereal grain lines – including maize,34 rice,35 wheat36 and barley37,38 – has led to the commercially available BARLEYmax. The CSIRO group has also examined the nutritional value and health benefits of novel high-amylose barley. High-amylose barley has a lower GI than normal barley39,40 and can cause a significant increase in starch and SCFAs in the large bowel of rats41 and a significant lowering of plasma cholesterol in pigs.42 Many high-amylose mutants have been developed through the inhibition or mutation of genes encoding SBEs, particularly those coding for inactivation or deficiencies in SBEIIa and SBEIIb activities.27 The novel technology targeting induced local lesions in genomes (TILLING) can detect mutations in target genes and can be used to modify the composition of starch.6 For example, TILLING technology has helped to obtain double mutants, with mutations in two SBEs, which increased the amylose content of wheat by 22% and the RS content by 115%.43 Details on the progress made in the production of high-amylose cereal grain crops have been reviewed by Yu et al.44 These novel cereal grains contain a greater proportion of amylose than common varieties, with as much as 30%–88.2% amylose.5,27 High-amylose mutants are ideal candidates for the production of foods with high-RS contents because resistant starch is formed through the gelatinization and retrogradation of amylose during processing.33

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7.5  Resistant Starch in Processed Grain Products Resistant starch is also present in some processed foods and an estimation of dietary intakes is important. Some researchers have estimated the daily RS intake of a given population based on their typical diet. In Sweden, the estimated daily intake of RS is 3.2 g, most of which is contributed from bread and potato products.45 The RS intakes of 10 different European countries ranged from 3.2 to 5.7 g per day.46 The average intake in Italy has been determined to be 8.5 g per day.47 In Australia, the estimated daily intake ranges from 3.4 to 9.4 g per day.48 The intake in similar countries such as Canada and the USA is probably low.49 The estimated RS intake in the Chinese diet is 14.9 g per day based on the evaluation of the RS content in 121 common foods in the diet, as well as dietary surveys.50 The CSIRO Division of Human Nutrition recommends a daily intake of at least 20 g RS to receive some of the bowel-related benefits.51 The RS intake varies greatly among individual people based on their dietary habits. Resistant starch is also produced commercially and added to processed foods to achieve the desired textures and viscoelastic properties, as well as to increase their fibre content. A survey of an Australian population was conducted by CSIRO in 2010 to determine community engagement and the health awareness of fibre and RS.52 The ultimate goal of the survey was to identify whether the promotion of RS as a dietary fibre would successfully engage the population in increasing their RS consumption, especially through products enriched with RS. The survey found a high level of recognition of the importance of dietary fibre for good health, especially among the women surveyed.52 The survey also showed that people are more accepting towards the idea of RS delivery through its incorporation into healthy staple foods rather than indulgences. It was concluded that there is an adequate level of awareness about the importance of fibre consumption and health in the Australian population. This finding suggests that messages on the health effects of RS will be successfully received by the public through the promotion of foods containing RS.52 Grain products such as bread and muffins are an easy medium in which to incorporate extra fibre, while substituting RDS with RS. Definitive health claims are still pending for RS as a prebiotic and an agent for improving satiety and controlling obesity; however, a health claim for RS has been approved by the European Food Safety Authority (EFSA). The approved health claim relates to RS consumption and the reduction of post-prandial glycaemic responses, with the wording ‘RS can help keep your blood sugar levels balanced after a meal’ or ‘RS may help prevent blood sugar “highs” after a meal’.53 The scientific evidence evaluated for this claim is specific to high-amylose maize RS2, which is viewed by EFSA as being a sufficiently characterized food constituent. However, EFSA acknowledges that all types of RS should reduce post-prandial glycaemic responses when replacing digestible starch in baked foods. High carbohydrate baked foods can only

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bear the claim if at least 14% of the total starch is RS as a replacement for digestible starch.53 Following EFSA's approved health claim, there has been a growing number of commercially available RS for food companies to incorporate into their products.

7.5.1  C  ommercially Available Resistant Starch for Use in Cereal Grain Foods Resistant starch has been added to a range of grain products, including muffins,54–56 spaghetti,57 biscuits,58–60 bread56,60 and tortillas.61 There are several commercially available forms of RS. Table 7.1 highlights some of the most commonly used types of commercial RS available for incorporation into foods. Most of the commercially available RS are currently RS2 – that is, high-amylose maize RS, with the predominant product being Hi-Maize® by Ingredion Inc. Novelose® is a source of RS3 marketed in Europe. An example of a marketed RS4 is Fibersym® flour. Strict regulations limit the amount of chemicals used to produce RS4 and the USA and Japan are the only two countries that currently allow RS4 in foods. In general, the addition of RS has a positive to no effect on the viscoelastic, rheological and thermal properties, and sensory acceptability of foods. The lack of deleterious effects makes it an excellent candidate for functional foods with a high-RS content. The effect of RS on viscoelastic properties has been investigated in wheat flour bread dough,62 spaghetti57 and extruded breakfast cereals.63 RS has a minimal effect on farinograph water absorption: adding 1% RS to wheat flour increases water absorption by 0.17%, less than the effect of adding other fibres, such as locust bean gum and wheat bran, which increase water absorption by 4.27 and 0.45%, respectively.62 Adding RS to wheat flour decreases the farinograph arrival time and increases the dough development time, but has no effect on the farinograph dough stability. Compared with locust bean gum and wheat bran fibres, added RS has the least influence on the farinographic parameters of wheat flour. Table 7.1  Commercial  resistant starch (RS) and associated production companies. RS type, including resistant maltodextrins (RM) and the source are given.

Commercial RS

Company

RS type

Source

HylonVII Hi-Maize® 260 Novelose® 330 CrystaLean® Fibersym® FiberRite® Fibersol-2®

Ingredion Ingredion Ingredion Opta Food Ingredients MGP Ingredients Inc. MGP Ingredients Inc. ADM/Matsutani Chemical Industry Co., Ltd Roquette Roquette

RS2 RS2 RS3 RS3 RS4 RS4 RM

High-amylose corn starch High-amylose corn starch High-amylose corn starch High-amylose corn starch Wheat starch Wheat starch Corn starch

RM RM

Wheat starch Corn starch

Nutriose® FB Nutriose® FM

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The viscoelastic properties of cooked spaghetti noodles containing 5 and 15% RS and bran have also been compared.57 Bran-containing spaghetti was firmer than RS-containing and control spaghetti and had a lower relaxation time. Measurements of viscoelasticity, including stress relaxation and creep recovery tests, showed little difference in the viscoelastic behaviour between the spaghetti containing 5 and 15% RS, with the RS-containing samples being more similar to the control than the bran-containing spaghetti.57 The maximum resistance to extension and strain in pan bread dough containing added RS and the enzyme transglutaminase was increased, as well as the starch gelatinization temperature.64 Certain enzymes probably mitigate the negative protein dilution effect caused by the addition of RS. Resistant starch has been incorporated into extruded breakfast cereals with minimal impact on product quality. The addition of RS (5) with yeast invertase. After 10 minutes, 50, 40, 20 and 10% of the trisaccharides, tetrasaccharides, pentasaccharides and higher DP (>5) fructans, respectively, were hydrolyzed. The inclusion of inulin-type fructans to bread confirmed that lower DP (4000 mg kg−1) and in bran (about 4000 mg kg−1) and shorts (about 3000 mg kg−1).41 In corn, the TPC was highest in high-carotenoid maize [320 mg GAE (100 g)−1], followed by yellow corn, blue corn, white corn and red corn [244 mg GAE (100 g)−1].29 The TPC was significantly reduced during lime cooking in the production of masa and tortilla, although the TPC in yellow corn grown in Mexico ranged from 249 to 1158 mg GAE (100 g)−1 for old varieties and 624 to 1168 mg GAE (100 g)−1 for new varieties.42 The TPC of both free and bound phenolic acids in eight varieties whole grain rice (Oryza sativa L.) of different bran colours were analysed.43 A significantly higher content of free phenolic acids was found in rice with red (6.97) and purple bran (5.47 mg GAE g−1) compared to white, light brown and brown bran (0.44 to 0.82 mg GAE g−1) and to other cereals (corn, wheat and barley). By contrast, bound phenolic acids were very low (0.46–0.95 mg GAE g−1) across the varieties. The levels of free phenolic acids in IITA119 (red bran) reported by Min et al.43 were within the range reported in dark coloured rice bran from Thailand, China and Sri Lanka (79–691 mg GAE 100g−1)44 and in red, white and black rice reported by Shao et al.16 Similarly, free, conjugated and bound phenolic acids were extracted from seven pigmented (Japonica) and 14 non-pigmented (Indica) subspecies. The TPC was four-fold higher in pigmented (4246 mg FAE kg−1) than in non-pigmented genotypes (1073 mg FAE kg−1).45 In pigmented varieties, free and bound phenolic acids ranged from 255 to 4146 and 538 to 1071 mg FAE kg−1, free phenolic acids from 388 to 901 mg FAE kg−1 and bound forms from 306 to 705 mg FAE kg−1 in non-pigmented rice varieties. De Mira et al.45 attributed this variation to pericarp colour, concurring with Goffman and Bergman.46 The TPC was four times higher in pigmented rice (4246 mg FAE kg−1) than in non-pigmented rice (1073 mg FAE kg−1). The high concentration was also due to high levels of free and conjugated (extractable) phenolic acids, which accounted for 81% of the TPC in pigmented varieties. These wide variations were attributed to differences in the extraction solvents, methods of determination (spectrophotometry versus HPLC), the standard used in the quantification (ferulic acid versus gallic acid) and the pericarp colour. Min et al.43 used acetone–water–acetic acid (70 : 29.5 : 0.05 v/v/v) to extract free phenolic acids, whereas most other researchers used 80% chilled acetone or acidified methanol or ethanol. Goufo et al.47 used spectrophotometry and HPLC to determine free and bound phenolic acids in Ariete white and brown rice from the Mediterranean. Spectrophotometry determined a higher concentration of free and bound phenolic acids in white (341.80 ± 59.98 and 422.15 ± 59.74 mg kg−1) and brown (227.38 ± 75.29 and 184. 48 ± 19.74 mg kg−1) rice than the free (4.64 ± 0.52 and 11.56 ± 2.63 mg kg−1) and bound (63.86 ± 0.80 and 132.67 ± 75.29 mg kg−1) levels determined using HPLC. No significant difference was observed between the subspecies indica and japonica within the pigmented and non-pigmented varieties.45

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12.4.2  M  inor Food Cereals (Barley, Sorghum, Millet, Rye and Oats) A limited number of studies have looked at the TPC of minor cereals, despite their role and use in product development. In four dehulled barley varieties, the total phenolic acid content ranged from 334 to 461 mg GAE (100 grain g)−1.48 Shen et al.35 reported a higher TPC of 171 mg GAE g−1 in black highland barley than in previous studies.32,42 Chandrasekara and Shahidi49 found a lower TPC of free, esterified, etherified and insoluble bound fractions in millet grains in the ranges 0.55–16.2, 0.25–2.02, 0.32–3.94 and 3.2–81.6 µmol FAE g−1 defatted meal, respectively. Coarse fractions of three non-waxy and one waxy barley obtained using air classification had 1.2–1.3 times higher concentrations of free and bound phenolic compounds than wholemeal flour.28 Wholegrain buckwheat contains two to five times more phenolic compounds than oats and barley, most of which are present in the free form and are distributed throughout the entire grain, unlike in other cereals.26 F3-3 and F3-4, representing the phenolic-rich fractions of the 16 milling fractions, had a 30-fold higher TPC than those of the FS-1 and FS-2 buckwheat fractions.26

12.5  C  omposition and Distribution of Phenolic Acids 12.5.1  Whole Grains Several studies have reported the phenolic acid profiles of whole grains of wheat,13,20,23,50–51 barley,14,28,31,35,52 corn,53–56 wheat, barley and corn,23 and rice.16,43,45,47,57 Whole grain phenolic acids include p-hydroxybenzoic, protocatechuic, vanillic, gallic and syringic acids and the hydroxycinnamates caffeic, p-coumaric, ferulic and sinapic acids. Ferulic acid dominates in most cereals, including wild rice, wheat, barley, millet, sorghum and maize.38,58–59 The effects of growing area, crop year and gene–environment interactions on the phenolic acids in grains have also been reported.13,45,47,57

12.5.1.1 Major Food Cereals (Wheat, Maize and Rice) The composition and distribution of phenolic acids in whole grain maize, wheat, barley and oats determined using HPLC and LC–MS/MS on alkaline hydrolysates are shown in Table 12.3. Ferulic, p-coumaric, iso-ferulic, syringic, sinapic and vanillic acids were found in all grains, but p-hydroxybenzoic was only observed in yellow corn.23 The phenolic acid content differed significantly (p < 0.05) among cereal types and within cereal varieties, with yellow corn having higher levels than other grains. Even higher concentrations of free and soluble conjugated ferulic acid and lower contents of bound phenolic acids and ferulic acid were found in nixtamalized maize,29 indicating the effect of the lime cooking process. Ferulic, p-coumaric,

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Table 12.3  Phenolic  acid composition and content [total (soluble + bound) (µg g−1, dw) and boundc] in whole grains of major cereals.a,b p-OHA

VA

Wheat (n = 4)

nd

Wheat varieties (n = 3) Wheat (methanol–acetone extraction)d Wheat (acid hydrolysis)d Wheat (alkaline hydrolysis)d Corn Yellow corn (n = 3) Inbred lines (n = 4)e Hybrids (n = 2)e Four genotypes of corn (typical + mutant)f Rice White rice

23–26

Cereal type

GA

PRCA

SRA

p-CA

FA

SA

IsoFA

Reference

70–90

21–29

44–57

600–750

46–59

21–34

9–25

9–10

19–26

449–535

36–69

Ndolo and Beta, 2014 Li & Beta 2015

CA

3.7 ± 0.9

Trace

13.5 ± 0.7 1.07. ±0.3 Trace 13–28

77.9 ± 4.1 Trace

1.2 ± 0.11

181–229

5.08–10.6 3.25–14.71 2.3–25.7

0.11 ± 0.03 0.34 ± 0.07

Arranz et al. 2010

Trace

Arranz et al. 2010 Arranz et al. 2010

Trace

Trace 101–140

12.3–24.48

204–278

1578–1977 117–150

31–67

0.94–1.60 19–21

Ndolo & Beta 2014 Bily et al. 2003

1.05–1.16 19–22 98–211 1552–2969

Bilyet al 2003 Li et al. 2007

8.09 ± 0.03

35–57

119–199

15–36

Light purple rice

2–8

14–71

65–219

11–34

Black rice

8.0–20

6.0–29

26–215

6–45

Zhang et al. 2015 Zhang et al. 2015 Zhang et al. 2015

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White rice (n = 7)g Red rice (n = 4)g Black rice (n = 7)g White rice (n = 6)c Light-purple rice (n = 5)c Black rice (n = 6)c a

0.06–0.36 nd

0.10–0.27 0.1–1.46

0.07–0.12

4.58–8

17–32

0.22–0.53

0.31–0.86

0.12–0.57 0.11–4.44

0.08–0.15

2.61–9.43 18–34

0.38–0.73

0.08–0.17

2.0–6.0

23–44

0.35–1.37

0–8

35–57

119–199

4–15

14–71

65–218

4–20

6–29

26–215

0.56–2.14 Pang et al. 2016 0.73–1.01 Pang et al. 2016 0.53–0.99 Pang et al. 2016 15–36 Zhang et al. 2015 11–34 Zhang et al. 2015 6–45 Zhang et al. 2015

 A, gallic acid; PRCA, protocatechuic; p-OHA, p-hydroxybenzoic acid VA, vanillic acid; CA, caffeic acid; SRA, syringic acid; p-CA, p-coumaric acid; FA, trans-ferulic acid; SA, G sinapic acid; ISOFA, iso-ferulic acid. nd, not detectable. c Reported in bound form only. d mg (100 g−1) fw (fresh weight). e mg g−1. f mg kg−1. g mg (100 g−1). b

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iso-ferulic and syringic acids were found in decreasing order from corn followed by barley, wheat and then oats. The second predominant phenolic acid in yellow corn was p-coumaric acid, whereas it was vanillic acid in wheat and barley,23 A much lower content of p-coumaric acid in yellow corn was reported (0.25 µg g−1)54 compared with 1160 µg g−1 for a pioneer hybrid (P3905)39 and an average of 242 µg g−1 for yellow corn (P1395XR and P1508HR) varieties. Yellow corn contained two to five times more total phenolic acids than barley, wheat and oats. Sosulski et al.9 found that the amount of phenolic acid in yellow corn flour was three times higher than that in wheat, oats and rice. Only ferulic and vanillic acids were significantly different (p < 0.05) among the wheat varieties; purple wheat had the highest content of sinapic and syringic acids.23 The phenolic acid composition of four corn varieties (typical-1, typical-2, waxy and high amylose) included p-hydroxybenzoic, vanillic, caffeic, p-coumaric, ferulic and o-coumaric acids.56 The phenolic acids extracted from wheat flour were dependent on the solvent and the method of extraction used.13 Of the five phenolic acids (p-hydroxybenzoic, p-coumaric, ferulic, sinapic and protocatechuic) quantified, three (p-hydroxybenzoic, ferulic and caffeic) were predominant in methanol–acetone, alkali hydrolysis and acid hydrolysis, respectively. In another study, eight out of 10 acids (gallic, p-hydroxybenzoic acid, vanillic, caffeic, syringic, p-coumaric, ferulic and sinapic acids) were determined in both their free and bound forms in varying concentrations in durum and bread wheat18 (Table 12.1). The levels of phenolic acids reported by Arranz et al.13 and Irakli et al.18 were much lower than those found in the wheat varieties studied by Ndolo and Beta.23 The distribution of antioxidant compounds in white and brown grains of the Mediterranean rice variety Ariete was examined and 10 phenolic acids were identified.47 These phenolic compounds included benzoates (gallic, protocatechuic, p-hydroxybenzoic, vanillic and syringic acids) and cinnamates (chlorogenic, caffeic, p-coumaric, sinapic and ferulic acid acids). All 10 phenolic acids were identified in the free form and only chlorogenic, p-coumaric and ferulic acids were found in the bound form, with the latter two being higher in the bound form than in the free form. The same phenolic acids, except gallic acid, were found in brown, white and germinated brown rice from Japan.6 Two additional acids linked to sugar moieties (feruloylsucrose and sinapoylsucrose) were present. The levels of the nine phenolic acids in white and brown varieties were within the same range, despite the different growing locations (Table 12.3). By contrast, seven phenolic acids (gallic, vanillic, syringic, chlorogenic, caffeic, p-coumaric and ferulic acids) were detected in both free and bound form in black rice from China.22 Black rice, known as Forbidden Rice, has a rich history in China and has been earmarked for its phenolic acids, flavonoids and anthocyanins, which enhance antioxidant activity. However, it is low yielding and new breeds have been developed by breeding the white

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(H32B) and black (Yunanheixiannuo) forms. Comparing the phenolic compounds among the 15 offspring samples from cross-breeds between black and white rice, it was observed that p-coumaric, sinapic, iso-ferulic and vanillic acids were present in the insoluble bound fraction, further confirming the presence of bound forms attached to cell walls within the grains.34 The crossbreed (light purple) had an intermediate amount of all the four monomeric acids and vanillic acid, which was not detected in the parent (white), but was quantified in the cross-breeds, indicating an improvement in the phenolic profiles through the cross-breeding of rice.

12.5.1.2 Minor Food Cereals (Barley, Sorghum, Millet, Oats and Rye) The phenolic acid composition of minor food cereals has mainly been studied in barley, sorghum, millet and, to a lesser extent, buckwheat, rye and oats (Table 12.4). In 30 barley varieties, which included a combination of two- and six-rowed, hulled or hull-less, regular or waxy varieties, three benzoates (p-hydroxybenzoic, vanillic and protocatechuic acids) and cinnamates (p-coumaric, caffeic, ferulic and chlorogenic acids) were identified.60 Hot water extracted a trace amount of protocatechuic acid and the highest amount of chlorogenic acid in most barley varieties, whereas acid hydrolysis released all seven acids. By contrast, a combination of acid and α-amylase hydrolysis released significant amounts of caffeic acid, whereas the sequential acid, α-amylase and cellulose extraction gave the highest yields of phenolic acids. Similarly, Holtekjølen et al.14 analysed bound phenolic acids in different barley varieties (two- or six-rowed, hulled and hull-less types, of normal or waxy and high amylose starch) from breeding programmes in Norway and Canada and observed a wide variation in the total phenolic acid content, which ranged from 604 to 1346 µg g−1. The wide variation was influenced by the presence or lack of hulls. p-Coumaric acid was six times higher in hulled (114–268 µg g−1) than in hull-less (15–38 µg g−1) types. Ferulic acid was the dominant acid, in agreement with recent findings.35,52,57 The ferulic and p-coumaric acid contents were 19.14 and 14.59 mg GAE g−1, respectively, in black highland barley (Hordeum vulgare L. var.) extracted using alkaline hydrolysis.35 Three diferulic acids (diFA) 5-5′ DFA, 8-O-4 DFA and 8-5′ DFA were also quantified. Hernanz et al.31 identified three monomeric acids (caffeic, p-coumaric and ferulic acids) and diFA in eight barley grain varieties. Among the diFA, 8-O-4 DFA had highest concentrations (73–118 µg g−1) followed by 8-5 DFA, benzofuran and 5-5′ DFA; 8-5 DFA had the lowest concentrations (10– 23 µg g−1). In non-pigmented and pigmented barley, ferulic acid (731 µg g−1) was predominant, followed by vanillic acid (92 µg g−1), p-coumaric, syringic, sinapic and iso-ferulic acid.23 The phenolic acid composition of six barley whole grain and milling fractions of commercial value, which included three Canadian (AC Parkhill, two-rowed hulled normal; CDC Ratan, two-rowed

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Table 12.4  Phenolic  acid composition and content [total = (soluble + bound) (µg g−1, dw) and boundc] in whole grains of minor cereals.a,b Cereal type

GA

PRCA

VA

0.76–9.86

Dehulled barley (n = 4)d Barley (n = 2) Two rows, normal hulled (n = 2) Six rows, normal hulled (n = 2) Two rows, waxy hull-less (n = 2) Six rows, regular-hulled (n = 8)c Two rows, regular hulled (n = 5)c Six-rows, regular-hull-less (n = 3)c Two rows, regular-hull-less (n = 9)c Two rows, waxy hull less (n = 5)c

p-OHA

nd

75–108

0–2

2–4

6–12

0–2

1–4

2–3

1–2

CA

ChlA

2–4

24–49

SRA

p-CA

FA

SA

IsoFA Reference Zhou et al. 2015

0.94–1.05 48–68 61–66

53–96

657–805

40–61

38–40 Ndolo & Beta, 2014 Gamel & Abdel 2012

0–6

0–2

56–91

232–421

4–10

0–3

0.7–1

41–69

124–466

Gamel & Abdel 2012

2–7

2–3

0

7–34

360–466

Gamel & Abdel 2012

71–91 (80)d

30–42 (35)

10–78 (34)

36–68 (51)

Yu et al. 2001

68–81 (76)

35–50 (41)

24–82 (47)

19–46 (46)

Yu et al. 2001

75–87 (80)

35–40 (37)

32–73 (52)

30–62 (46)

Yu et al. 2001

67–101 (87)

9–33 (19)

32–127 (52)

10–68 (30)

Yu et al. 2001

80–102 (90)

11–29 (23)

44–62 (55)

63–66 (64)

Yu et al. 2001

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Sorghum Ryee

x

x

Oats (n − 1)

1

4.24

46.2 ± 50.1 ± 1.5 367 ± 7.5 57 ± 1.93 0.89 1.3 18.8

98.1

324.4

nd

nd

4.5

45

Kodof

1.8

70.5

31.2

Fingerf

5

119.8

6.3

Foxtailf

4.5

11.8

21.8

Prosof

6.2

72

Littlef

2.1

Pearlf

5.4

Little millet (soluble) Little millet (bound)

5.82 ± 0.18

a

2.8

12.3

Indaf-15

2.54 ± 0.07

x 12.1

7

3.6

60.9

x 1054

139

141.1

802

2209.5

53.2

15.9

25.1

41.4

358.4

0.8

118.7

38.3

18.4

942.7

856.5

16.2

126

168.6

339.2

19

1235.2

444.6

18.9

48.8

32.6

162.4

30.9

23.4

1085.2

355.3

55.4

1.6

47.9

16

30.4

6.3

91.9

812.3

12.8

2.71 ± 0.16 70.36 ± 1.21

129.31 ± 15.07 121.02 ± 2.63

62.24 ± 1.04 2.59 ± 0.07

6.2

4.05 ± 0.13 35.69 ± 1.02 78.33 ± 29.53 ± ± 1.95 0.68 1.32 ± 0.03 23.51 ± 0.69 1.29 ± 0.04 2.53 ± 0.08 ±

nd

Kang et al. 2016 Pihlava et al. 2015 Ndolo & Beta, 2014 Subba Rao & Muralikrishna,2002 Chandrasekara & Shahidi 2013 Chandrasekara & Shahidi 2013 Chandrasekara & Shahidi 2013 Chandrasekara & Shahidi 2013 Chandrasekara & Shahidi 2013 Chandrasekara & Shahidi 2013 Pradeep & Sreema 2017 Pradeep & Sreema 2017

 A, gallic acid; PRCA, protocatechuic; p-OHA, p-hydroxybenzoic acid VA, vanillic acid; CA, caffeic acid; ChlA, chlorogenic acid; SRA, syringic acid; p-CA, p-coumaric acid; FA, G trans-ferulic acid; SA, sinapic acid; IsoFA, iso-ferulic acid. nd, not detectable. c Bound phenolic acid. d mg (100 g−1). e mg kg−1. f total = soluble (free, esterified and etherified) and insoluble bound. b

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hull-less waxy; and Celebrity, six-rowed hulled normal) and three Egyptian (Giza 127, two-rowed hulled normal; Giza 125, six-rowed hulled normal; and Giza 131, two-rowed hull-less waxy) cultivars52 showed major acids (ferulic and p-coumaric) accounting for 73–95 and 2–24%, of the whole grain and pearled fractions, respectively. The Egyptian variety had significantly higher levels than the Canadian cultivars, especially cultivars Giza 125 and 127. In spring wheat, the genotype and environment were also reported to influence the TPC, phenolic acids and antioxidant capacity,61 which is also likely to be the case with barley. In millet grains, both hydroxybenzoates (gallic, protocatechuic, p-hydroxybenzoic, gentistic and vanillic and syringic acids) and hydroxycinnamates (chlorogenic, caffeic, trans-cinnamic, p-coumaric, sinapic, trans-ferulic and cis-ferulic acids) have been reported.49,62–63 The latter were more abundant in the insoluble bound (468–3687 µg g−1) than soluble (45–682 µg g−1) fraction, whereas the converse was seen with benzoates.49,62,80 For example, the benzoic acids in soluble and insoluble bound extracts of defatted meals ranged from 103 to 269 and 32 to 215 µg g−1, respectively. Insoluble bound fractions of Kodo, Foxtail, Proso, little and pearl millets had significantly higher (p < 0.05) levels than the free and soluble conjugated (esterified and etherified) fractions, except for two finger millets (Local and Lava), which had a higher content of free and soluble conjugated than bound fractions.49 Phenolic acids are present in the free form, accounting for 71% of the total, with protocatechuic acid as the major free acid [45 mg (100 g)−1]. Other acids include gallic, caffeic and vanillic acids. Chandrasekara and Shahidi49 studied the phenolic compounds and antioxidant activity of flour from different fractions of whole grain (Table 12.4) and identified 10 benzoic and 15 cinnamic acid forms and their derivatives. Millets also contain dimers and trimers of ferulates, which have a high antioxidant activity compared with their monomers.47 The four diFAs have been identified as 8.5' Benzo DiFA, 8.5' DiFA, 5,5' DiFA and 8,5-Aryl DiFA. Sugar moieties such as protocatechualdehyde, p-hydroxy benzaldehyde and methyl esters of vanillic acid are also present. The major phenolic acids in sorghum include caffeic, p-coumaric, sinapic and ferulic acids,7 as reported by Chiremba et al.36 In sorghum, 77% of the phenolic acids exist in bound form.64 N'Dri et al.17 reported that sorghum, fonio and millet contain more than ten-fold of their phenolic acids in bound form. Hence sorghum has more bound phenolic acids than all other cereals. Protocatechuic, caffeic, ferulic and gallic acids were identified among the 75 phenolic compounds in three types of sorghum (brown, red and white), illustrating that this cereal is a rich source of diverse phenolic compounds, some of which were reported for the first time.65 Another study also identified four phenolic acids, different from those reported by Kang et al.,65 including chlorogenic, protocatechuic, p-coumaric and ferulic acids.66 Buckwheat contains two to five times more phenolic compounds, including phenolic acids, than oats and barley, most of which are present in the free form.33

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12.5.2  Botanical and Milling Fractions 12.5.2.1 Separation of Grain Fractions Using Mechanical and Manual Methods To understand the distribution of phenolic acids in grains, it is important to have an overview of the grain structure. The grain consist of the starchy endosperm, germ and bran. The bran is made up of multi-layer peripheral tissues consisting of the outer pericarp (beeswing bran), the inner pericarp (consisting of cross-cells and tube cells), the seed coat (testa), the hyaline layer (nucellar epidermis) and the aleurone layer. The proportions of these anatomical parts vary widely in different types of grains67,68 (Table 12.5). The distribution of phenolic acids can be measured in situ in grains and in separated grain layers or milling fractions. For in situ analyses, microspectrofluorimetry69,70 and fluorescence microscopy have been used to study the distribution of phenolic acids in the aleurone layer and across the grain.71,72 Ferulic acid is often selected because of its natural ability to auto-fluoresce under UV light.69 Ndolo et al. used a fluorescence microscope to not only visualize the location and distribution of ferulic acid, but also to predict the concentrations using fluorescence intensity profiles (Figure 12.3). The use of fluorescence intensity profiles clearly showed how the concentration of ferulic acid decreases from the outer towards the inner layers of the grain, with the lowest levels in the innermost part of the endosperm. There were significant positive correlations between the fluorescence intensity and the concentration of ferulic acid determined by HPLC.72 Grain botanical or milled fractions may be obtained using various techniques. Mechanical separation, also known as dry fractionation, includes Table 12.5  Percentage  contribution of different parts of cereal grain to whole grain. Adapted from Serna-Saldivar,1 Kent and Evers99 and Ti et al.89 Bran99

Cereal Corn Corn99 Wheat Wheat99 Barley Barley99 Oats Rice89 a

Dent Flint Hard Durum Hulled Hull-less

na, not applicable

Pericarp

Aleurone

Endosperm

Germ

6.0 6.5

2.8 2.2

8.2 12 2.9 3.3

6.7 naa 81.4–84.1 4.8 5.5

78 79.6 79.7–82.0 82 86.4

12 2–3

naa 4–6

12 11.7 11.7–15.2 3.6 1.6 2.5–3.6 3.0 3.4 3.4 3.7 2–3

(Pericarp + testa + aleurone)

5.0–6.5 14.1–15.9 8.8 12 6–9

76.2 87.6 87.6 84 90

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Figure 12.3  Fluorescence  intensity profiles corresponding to line 1 in the raw dig-

ital images showing the distribution of ferulic acid from the outer layers (pericarp and aleurone) into the endosperm: (A, A1) MSUD8006 wheat; (B, B1) USP1395XR yellow corn; and (C, C1) non-pigmented barley. Scale bar = 50 µm. Reprinted from Food Res. Int., 52 (1), V. U. Ndolo, T. Beta and R. G. Fulcher, Ferulic acid fluorescence intensity profiles and concentration measured by HPLC in pigmented and non-pigmented cereals, 109–118., Copyright 2013, with permission from Elsevier.

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pearling, debranning, dehulling, abrasive milling, roller milling, sieving and air classification techniques and is used to obtain milling fractions with different characteristics.28,41,73–76,89–92 Beta et al.41 used incremental pearling and roller milling to obtain 15 fractions designated as break flours (B1–B4, S1), reduction flours (M1–M6, Q1), shorts, bran and bran flour from eight Canadian wheats. Micronization and air classification were used to separate the whole meal produced after the dehulling and pin-milling of hulled barley into two-milled fractions: a coarse fraction (40%) and a fine fraction (60%).28 Eight barley grain varieties, including five high-quality and three low-quality varieties (Sunrise, Iranis and Boira) were mechanically fractionated into three fractions: F1, consisting mainly of the husk and outer layers; F2, intermediate; and F3, consisting mainly of the endosperm.31 Gamel et al.52 used a Satake abrasive mill to pearl barley into the hull, outer layer (F1), middle pearling (F2) and endosperm (F3). Variations in the characteristics of fractions obtained by different debranning processes are summarized in a review by Hemery et al.75 In addition, Hemery et al.76 showed that fractions obtained by various mechanical separation techniques consisted of blends of different grain outer layers, which affect their chemical composition. Blandino et al.77 analysed the distribution of bioactive compounds in maize fractions (germ, maize meal, maize flour and animal feed flour) obtained through two industrial dry-milling processes, dry degermination and tempering–degermination. The method of degermination influenced the bioactive contents in milled fractions and these were unevenly distributed.77 Similarly, Giordano et al.73 compared bioactive compounds in roller-milled and pearled fractions in conventional red and white wheats and pigmented wheat (yellow, purple and blue types) and suggested that the selection of an appropriate fractionation process to produce flours rich in bioactive compounds is crucial. Despite these challenges, the distribution of phenolic acids has mainly been determined in mechanically separated grain fractions from wheat74,78,79 barley,31,52,74 corn25,81 and sorghum.7,36,82 Only a few researchers have used hand-dissected fractions.23,75,83 The manual separation of whole grains yields pure samples suitable for compositional analysis.84 Ndolo and Beta23 manually separated botanical fractions of wheat, barley and corn (Figure 12.4) to study the composition and distribution of phenolic acids in diverse grains. Understanding the distribution of phytochemicals in grains is crucial in guiding the separation of grain fractions rich in these components.85

12.5.2.2 Variation in the Composition and Concentration of Phenolic Acids in Grain Fractions or Milling Fractions The phenolic acid composition and concentration in grain fractions are shown in Tables 12.6 and 12.7. The concentrations differed significantly (p < 0.05) among the fractions of different grains and varieties. In some instances, even the distribution of phenolic acids within grain fractions

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Figure 12.4  Raw  hand-dissected fraction (pericarp, aleurone, endosperm and germ) and fluorescence digital images of (A) purple barley, (B) Dasca corn and (C) Ambassador wheat. Reprinted from V. U. Ndolo and T. Beta, Cereal Chem., 2014, 91 (5), 522–530, with permission from © AACC International and John Wiley and Sons.

varied widely.13,23,25 Free, soluble and insoluble hydroxycinnamic acids were determined by HPLC with diode array detection in selected cereal grain fractions with different characteristics. The identified hydroxycinnamates were ferulic, diferulic, sinapic, p-coumaric and benzoic acid derivatives (p-OH benzoic acid, vanillic acid and vanillin).25 The highest levels of these compounds were present in wheat bran, fermented wheat bran and rye bran, whereas reduced wheat flour and rye flour had the lowest concentrations. Ferulic acid (trans + cis) accounted for 50–80% of the total acids and was the most abundant in wheat middling, air-separated wheat, middling-reduced wheat flour, fresh wheat germ, wheat bran and fermented wheat bran. Insoluble ferulates were highest in wheat bran [about 400 mg (100 g)−1] and rye bran [about 230 mg (100 g)−1], with the same fractions also having high insoluble diferulates [28.7 and 20.1 mg total DFA (100 g)−1], respectively. Fermentation increased the level of diferulates released from wheat bran up to 39.9 mg total DFA 100 g−1. 12.5.2.2.1  Major Food Cereals (Wheat, Maize and Rice).  Eight phenolic acids (vanillic, p-coumaric, ferulic, sinapic, p-hydroxybenzoic, caffeic, syringic and iso-ferulic acids) were identified in manually separated grain fractions: the pericarp (outer and inner pericarp, testa and nucellar epidermis), aleurone, endosperm and germ from four wheat varieties and three yellow

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Table 12.6  Distribution,  composition and content (µg g−1) of phenolic acids in grain fractions of wheat and corn.a,b,c Cereal type/ fraction

GA

PRCA

p-OHBA

VA

SRA

CA

p-CA

FA

SA

Wheat (n = 4) Pericarp

nd

nd

0–56

Aleurone layer

nd

nd

27–65 (47)

Germ

nd

nd

Endosperm

nd

nd

126–190 (152) nf

Wheat flourd

37.9 ± 0.9

Wheat brand AACC wheat bran Red wheat bran White wheat bran Commercial wheat Red wheat bran White wheat bran

164–486 (260) 123–178 (142) 173–251 (213) nf

211–272 (234) 112–117 (113) 83–133 (108) nf

9.9 ± 0.7

nf

126–192 (166) 64–85 (77) 120–185 (158) nf 66–213 (139) nf 19–21 (20) 98.6 ± 11.9 Trace

804.6 ± 19.8

432.09 ± 22.1

3194–3815 (3177) 3172–3616 (3365) 490–1054 (823) 107–118 (111)

60.1 ± 1.9

39.2 ± 0.62

82.6 ± 0.19 125 ± 0.34 0.80 ± 0.00 35.0 ± 0.03 1918 ± 1.59

24.6 ± 0.04

45.4 ± 0.53 71.5 ± 0.16

28.3 ± 0.96

78.0 ± 0.35 182 ± 0.23 1.78 ± 0.00 46.6 ± 0.22 2020 ± 0.94

45.5 ± 0.61

80.8 ± 1.48 145 ± 0.16

38.1 ± 0.05 1376 ± 4.00

38.8 ± 0.03 1992 ± 5.81

173–353 (200) 108–337 (245) 222–656 (403) nf Trace

IsoFA

Data from references

Ndolo & Beta, 2014 102–160 Ndolo & Beta, (139) 2014 110–187 Ndolo & Beta, (162) 2014 124g* Ndolo & Beta, 2014 nf Ndolo & Beta, 2014 Arranz &Calixto, 2010 Arranz &Calixto, 2010 Kim et al. 2006 Kim et al. 2006 Kim et al. 2006 Kim et al. 2006 Kim et al. 2006 Kim et al. 2006 (continued)

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Table 12.6  (continued) Cereal type/ fraction

GA

PRCA

p-OHBA

VA

SRA

CA

p-CA

FA

IsoFA

399–520 (465)

Giordano et al. 2017 Giordano et al. 2017 Giordano et al. 2017 Ndolo & Beta 2014 134–395 Ndolo & Beta (237) 2014

Five wheat varieties Roller-milled bran Refined flour Three yellow corn varieties Pericarp

nd

276–395 (338)

Aleurone layer

152–261 (226)

3700–5101 337–416 (4243) (274)

Germ

113–239 (183)

169–216 (190)

Endosperm Four hard cultivars of maize Bran Flour Four soft cultivars of maize Bran Flour Four specialty maize genotype (F + B)

889–1716 (1371)

nd nd

58–66 (63) nd

2023–3179 14 300– (2555) 22 437 (18 400) 325–492 3944–5652 (412) (4836)

Data from references

SA

1391– 88–136 2282 (106) (1956) 128–173 (146)

293–357 (249) 27–40 (33)

825–919 (863) 152–182 (169)

232–488 (302) 47–83 (65.2)

2740–3471 (3214) 83–129 (108)

88–175 (133) 22–54 (43)

1973–2742 47–69 (2198) (61) 63–113 (81)

89–123 (112)

Ndolo & Beta 2014 Ndolo & Beta 2014 Ndolo & Beta 2014 Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Das & Singh 2016

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Pericarp QPM

499 ± 24.9e

1795

Baby corn

nf

nf

Popcorn

176

2221

Sweet corn

nf

2052 ± 25

QPM

27

537

Baby corn

51

594

Popcorn

35.2 ± 2.4

313

Sweet corn

nf

427

QPM

10.9 ± 0.26

6.8 ± 0.3

Baby corn

nf

Popcorn

nf

0.6 ± 0.005 f 378 ± 8.5

Sweet corn

nf

nf

Germ

Endosperm

a

Das & Singh 2016 86 327 3314 nf 842 ± 36e Das & Singh 2016 296 212 4081 nf 2472 ± Das & Singh 0.8 2016 119 516 ± 13 3594 ± 57.4 376 560 ± Das & Singh 12.2 2016 103 326 4259 413 ± 19 139 Das & Singh 2016 Das & Singh 2016 6.5 17 352 897 ± 57 61 Das & Singh 2016 53.3 ± 1 54.1 ± 3.6 560 411 386 ± 9.6 Das & Singh 2016 46 36 912 171.5 ± 27.9 ± Das & Singh 7.1 1.8 2016 13 67 874 321 827 Das & Singh 2016 Das & Singh 2016 0.66 ± 0.06 0.5 28 nf 1.2 ± Das & Singh 0.05 2016 0.15 ± 0.01f nf 1.52 ± 0.02 206.2 nf Das & Singh 2016 9 10.8 ± 0.18 166 0.4 ± 0.0 f nf Das & Singh 2016 0.51 ± 0.03f 1.5 ± 0.04f 856 244 ± 7 nf Das & Singh 2016

 A, gallic acid; PRCA, protocatechuic acid; 4-OHBA, 4-hydroxybenzoic acid; VA, vanillic acid; CA, caffeic acid; SRA, syringic acid; p-CA, p-coumaric acid; FA, trans-ferulic acid; G SA, sinapic acid;; IsoFA, iso-ferulic acid; QPM, quality protein maize. All values are a total of free + bound forms. c nd, not determined; nf = not found. d Found only in one sample. e bound form only. f free form only. g Only found in the germ of purple wheat. b

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Table 12.7  Distribution,  composition and content (µg g−1) of phenolic acids in grain fractions of rice.a,b Cereal type/ fraction White rice bran (n = 7)c Red rice bran (n = 4)c Black rice bran (n = 7)c White riced Rice brand

GA

PRCA

p-OHA

0.37–3.95

nd–nd

0.63–1.25 0.70–1.22

0.36–0.83 24.28–70.74 118.34–172.52 2.22–5.45 0.19–6.42

Pang et al. 2018

1.25–3.05

nd–2.69

0.57–1.64 0.73–27.84

0.36–0.64 12.36–42.40 61.89–141.03

1.92–5.04 1.88–5.43

Pang et al. 2018

0.32–0.67 9.69–28.52

98.03–146.34

3.43–9.50 1.37–3.33

Pang et al. 2018

VA

CA

6.36–10.58 1.57–10.78 0.68–1.94 17.58–45.37

SRA

p-CA

FA

SA

IsoFA

ChlA

0.22

1.04

0.43

0.23

0.25

0.18

8.3

57

0.33

0.96

3.47

13.2

6.04

3.1

3.49

1.53

223.04

891.33

1.59

3.14

Brown 0.4 riced Rice huskd 1.88

1.43

1.06

0.41

5.14

0.25

29.24

104.22

1.11

0.98

2.13

1.45

0.94

2.44

1.98

807.29

362.2

3.48

0.8

Indica rice (n = 5) Bran/ 16.9–27.1 embryo Endo2.0–3.2 sperm Black rice Rice bran 42.4 Polished rove

2.3

Reference

Goufo et al. 2015 Goufo et al. 2015 Goufo et al. 2015 Goufo et al. 2015

Ti et al. 2014a 6.3–14.1 Ti et al. 2014a

24.4–39.3

37.9–49.7 44.9–64.9 374.9–403.1 1146.1–1416.7

7.2–9.3

7.3–22.9

4.5–11.3 nd

9.3–18.5

69.1–77.7

918.7

49.6

190.9

231.9

1773.8

7706.8

320.2

1

7.5

8.4

82.5

119.7

Ti et al. 2015 Ti et al. 2015

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Brown rice 17.3

299.2

9.9

42.6

34.3

263.2

1588.3

Japonica brown rice Pericarp

Trace

16.1

28.4

10.1

Trace

22

950

2238.2

87.4

14.8

Aleurone layer Embryo

4.4

12.5

33

12.8

Trace

23.2

848

2732.8

124.5

25.7

1.3

4.2

19.2

2.6

9.7

28.8

408.7

983.4

51.1

23

EndoTrace sperm Indica brown rice

Trace

2

Trace

nd

1.5

20.5

128.6

6.3

11.5

Pericarp

1.2

6.7

13.4

15.9

1.7

22.9

939.3

2244.4

119.2

23.6

Aleurone layer Embryo

1.3

1.8

12.1

3.3

2.3

18.3

414.4

1325.3

81.7

17.9

Trace

7.3

6.2

Trace

3

13.5

144.9

350.1

25.2

38.9

Trace

Trace

1.6

nd

Trace

1.2

11.5

104.8

6.9

10.7

Endosperm a

Ti et al. 2015 Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b Ti et al. 2014b

 A, gallic acid; PRCA, protocatechuic; p-OHA, p-hydroxybenzoic acid VA, vanillic acid; CA, caffeic acid; ChlA, chlorogenic acid; SRA, syringic acid; p-CA, p-coumaric acid; FA, G trans-ferulic acid; SA, sinapic acid; IsoFA, iso-ferulic acid; ChlA, chlorogenic acid. b nd, not determined. c All values are a total of free and bound except: bound form only (mg per 100 g). d mg kg−1.

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23

maize, five of which were present in all the fractions (Table 12.6). The grain fractions, excluding the endosperm, had increased levels of phenolic acids compared with the whole grain, further confirming the uneven distribution across the grain kernel and in accordance with Butsat and Siriamornpun30 and Chiremba et al.36 In addition to p-coumaric and ferulic acids, other phenolic acids were significantly high in the pericarp, aleurone and germ of wheat and yellow corn (Table 12.6). As observed with other cereals, ferulic and p-coumaric acids were negligible in the endosperm fractions.23 Chiremba et al.36 identified minimal levels of p-coumaric and ferulic acids in flour fractions and much lower levels of p-coumaric, ferulic and sinapic acids in the bran of both hard and soft cultivars compared with those reported by Ndolo and Beta,23 although they both used HPLC and LC–MS/ MS techniques. However, both ferulic and p-coumaric acids were higher in the bran fractions of hard compared with soft varieties (Table 12.6), suggesting a relationship between maize hardness and the phenolic acid content in bran.36 Iso-ferulic acid, which is rarely reported in cereals, was present in the pericarp and aleurone layer of wheat and yellow maize varieties. Dehydrodimers of ferulic acids, 8-5′ DFA, 5-5′ DFA and 8-O-4 DFA were also identified in the pericarp, aleurone layer and germ fractions of these grains. Das and Singh81 also found iso-ferulic acid in the bound fraction of the germ and pericarp of specialty maize genotypes. Free and bound phenolic acids in the germ, pericarp and endosperm of Indian specialty maize (Zea mays L.) genotypes, quality protein maize (QPM), baby corn, popcorn and sweet corn have also been studied.81 The total free and bound ferulic acid content varied significantly, with the highest and lowest levels in the pericarp and endosperm, respectively, in typical and specialty corn (Table 12.6). Free and bound ferulic acid varied from QPM (5 and 95%), baby corn (23 and 77%), popcorn (4 and 96%) and sweet corn (17 and 83%), with QPM showing the lowest (3694 ± 139 µg g−1) and sweet corn presenting the highest (5989 ± 200 µg g−1). These values are three- to five-fold lower than the total ferulic content reported by Ndolo and Beta23 (19 221–29 190 µg g−1), but higher than those reported by Adom and Liu5 (1749 µg g−1) for yellow maize, by de la Parra et al.29 for various coloured maize (1030–1530 µg g−1) and by Chiremba et al.36 (2198–3214 µg g−1) for white dent maize. Improved acid hydrolysis on wheat bran and wheat flour released phenolic acids identified as hydroxybenzoic, protocatechuic, caffeic, cinnamic, sinapic and ferulic acids and similar to those extracted by methanol–acetone solvent mixtures and alkali hydrolysis, except for gallic and vanillic acids, which were present only in the latter method.13 The type of extraction solvent influenced the type and content of phenolic acid found in the flour and bran. Hydroxybenzoic (41.5 ± 12.3 and 65.8 ± 6.5 µg g−1) and ferulic (0.2 ± 0.002 and 219.3 ± 5.1 µg g−1) acids, respectively, were predominant in wheat flour and bran extracted by methanol–acetone. In the acid hydrolysate, the predominant phenolic acids were caffeic acid in wheat flour and protocatechuic

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acid in wheat bran, although these two phenolic acids were undetectable in methanol–acetone extracts. The free and bound phenolic acids of the roller-milled and pearled fractions of five wheat varieties (Triticum aestivum L.) (PR22R58, red-grained; white bear, white-grained; Bona Vita, yellow-grained; Rosso, purple-grained and Skorpion, blue-grained) clearly showed an uneven distribution in the kernels.73 Only ferulic acid was identified in the refined flour as free and bound forms. Free phenolic acids and phenolic acids bound to the cell wall were highest in the outermost layers of the kernel (0–15%) and decreased towards the inner layers (25–100%) in the pearled fractions, concurring with what was observed by drawing a line across the grain using the fluorescence intensity.72 For the individual phenolic acids, the highest content was found in the first two pearling fractions (accounting for 0–10% of the kernel weight), which partly represented the bran in the roller-milled fractions. Similar distribution patterns of phenolic acids have been reported in the pearled fractions of conventional wheat varieties.41,86 Brewer et al.87 studied the effect of particle size during extraction and found that both free and bound phenolic acids were higher in wheat bran fine particles (0.64 and 6.72 mg FAE g−1) than in coarse (0.45 and 4.73 mg FAE g−1) and medium (0.37 and 5.19 mg FAE g−1) particles. Rice kernels have a hard, protective husk, which is removed by dehulling6 to yield brown rice. The latter is further processed through milling to separate the bran and embryo (germ) from the endosperm, commonly referred to as polished rice. The distribution of phenolic acids in rice has been studied16,22,47,88–91 and Table 12.7 shows the types and concentration found in botanical fractions. Goufo et al.47 examined the distribution of antioxidant compounds in milling fractions (husk, bran, brown rice and white rice) of the Mediterranean rice variety Ariete and identified 10 phenolic acids at 280 nm (gallic, protocatechuic, p-hydroxybenzoic, vanillic and syringic acids) and at 320 nm (chlorogenic, caffeic, p-coumaric, sinapic and ferulic acids). All were present in the free form in all milling fractions, but some were undetectable in the bound form. No benzoate was detected in the bound form in white rice, but p-hydroxybenzoic acid was detectable in brown rice. The predominant acids were ferulic acid followed by p-coumaric acid in all fractions, with rice bran with husks having the highest ferulic acid (891.33 mg g−1) and p-coumaric acid (807.29 mg g−1) content (Table 12.7). Similarly, seven individual phenolic acids (gallic, protocatechuic, chlorogenic, caffeic, syringic, coumaric and ferulic acids) were detected in the bran, embryo and endosperm of indica rice varieties cultivated in southern China,89 but p-hydroxybenzoic, vanillic and sinapic acids were not detectable (Table 12.7). In another study, seven phenolic acids were identified in the milled fractions of black rice,88 six (gallic, chlorogenic, caffeic, syringic, p-coumaric and ferulic acids) of which were similar to those reported by Ti et al.89 Vanillic acid was detected in fractions of black rice not of the indica varieties. All seven were detected in the free form in both fractions; however, those in the bound form were all

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undetectable in the endosperm, whereas only three (protocatechuic, chlorogenic and caffeic acids) were undetectable in the bran/embryo fraction.89 The bran and husk contained the most phenolic acids, especially p-coumaric and ferulic acids (Table 12.7) observed in brown and white rice.41 However, the ferulic acid levels reported by Goufo et al.47 were much lower than those reported by Ti et al.,89 where the total ferulic acid content of the bran/embryo fraction ranged from 1146 to 1416 µg g−1 in indica varieties. In agreement with Goufo et al.,47 ten phenolic acids were identified in four tissue fractions (pericarp, aleurone layer, embryo and endosperm) of japonica and indica whole brown rice.22 Iso-ferulic acid was also detected in the former. There were significant differences (p < 0.05) in the free and bound phenolic contents among the four fractions (Table 12.7). Ferulic and p-coumaric acids were the most abundant in the pericarp of japonica (2238 and 950 µg g−1) and indica (2244 and 939 µg g−1) varieties, mainly in the bound form, and averaging 2204 and 945 µg g−1, respectively (Table 12.7). The aleurone layer had second highest content, followed by the germ. Similar to wheat, yellow maize and barley, the aleurone layer also had a high content of ferulic (2733 and 1325 µg g−1) and p-coumaric (848 and 414 µg g−1) acids, mainly in the bound form. The endosperm had little or only trace amounts of most of the phenolic acids identified. The low levels of bound compounds, especially p-coumaric and ferulic acids, in black, red and white rice varieties confirm that rice, unlike maize and wheat, has a greater amount of free (soluble) than bound phenolic acids. 12.5.2.2.2  Minor Food Cereals (Barley, Sorghum, Millet, Rye and Buckwheat).  While studying the phenolic acids and antioxidant properties of pearling fractions of four barley varieties, Gamel and Abdel-Aal52 identified benzoates (protocatechuic, p-hydroxybenzoic, vanillic and syringic acids) and cinnamates (caffeic, p-coumaric and ferulic acids) in most fractions (Table 12.8). Protocatechuic, syringic and caffeic acids were undetectable in fraction 3 (the endosperm). Ferulic acid was the predominant acid, with the highest levels in the hull and fraction 1 (outer layers: pericarp and testa), followed by fraction 2 (middle pearling parts: of aleurone, testa and germ) and lowest in fraction 3 (the endosperm). The ferulic acid content in fraction 1 and the hull ranged from 616 to 623 and 897.5 to 905.7 µg g−1, respectively, in two-rowed hulled normal barley and from 424 to 601 and 681 to 1018 µg g−1, respectively, in six-rowed hulled normal barley varieties. The hulls also had exceptionally high contents of p-coumaric and syringic acids (Table 12.8). A high concentration of phenolic acids in outer layers has been reported in the husk of Thai rice30 and first and second pearling fractions of wheat grains.41 Ferulic and p-coumaric acids accounted for 43–95 and 2–55% of the total phenolic acids, respectively. Similarly, F1, which was composed of the husk + outer layers, had higher concentrations of ferulic, p-coumaric and ferulic acid dehydrodimers, which accounted for 78–82, 78–86 and 79–87%, respectively. F2 (the intermediate fraction) had lower levels of these

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Cereal type/ fraction PRCA Barley (n = 2) Pericarp Aleurone layer Germ Endosperm

 

p-OHBA

28–107 (72) 192–394 (293) 19–36 (28) 117–122 (119) 40–43 (41) 141–209 (175)

Waxy barley (US) (n = 1) CF (Coarse fraction) WM (Wholemeal) FF (Fine fraction) Non-waxy barley (n = 3) CF (Coarse fraction)

SRA

CA

p-CA

FA

SA

IsoFA

Reference

92–166 (129) 39–47 (43)

nd 64–118 (91)

131–330 (230) 64–73 (68)

44–57 (50)

nd

91–146 (118) 25–29 (27)

1918–3698 (2807) 1008–1651 (1329) 558–571 (565) 116–123 (119)

103–146 (125) 73–153 (113) nd

69–153 (111) 39–133 (87) 175–850 (513)

Ndolo &Beta 2014 Ndolo &Beta 2014 Ndolo &Beta 2014 Ndolo &Beta 2014

14.3

44.7

449.6

1.3

35.2

379.8

0.24

4.58

130.2

1.6–13.7

61–74

459–549

1.2–1.5

50–63

360–470

0.73–0.99

36–49

245–298

GómezCaravaca et al. 2015 GómezCaravaca et al. 2015 GómezCaravaca et al. 2015 GómezCaravaca et al. 2015 GómezCaravaca et al. 2015 GómezCaravaca et al. 2015 (continued)

263

WM (Wholemeal) FF (Fine fraction)

VA

Types and Distribution of Phenolic Compounds in Grains

Table 12.8  Distribution,  composition and content (µg g−1) of phenolic acids in grain fractions of minor cereals (barley).a,b,c

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Cereal type/ fraction PRCA

p-OHBA

264

Table 12.8  (continued) VA

SRA

CA

p-CA

FA

Barley (n = 2) nd/–4.1 7.1–9.1 F1 (Outer layerpericarp/ husk & testa) F2 (Middle nd–nd 0.7–10.0 pearlingpart of aleurone, testa, germ/ intermediate fraction) F3 (Endond. nd 2.2–nd sperm) H (Hull)

4.1–3.6 9.7–17.9

SA

IsoFA

Reference Ndolo & Beta 2014 Gamel & Abdel-Aal 2012

15.0–17.6

4.1–5.7

7.2–26.1

15.7–22.7

616–623

2.0–14.4

nd–0.8

2.0–6.6

12.5–72.0

138.5–154.3

Gamel & Abdel-Aal 2012

1.5–nd

nd–nd

nd–0.88

10.7–16.5

29.8–108.5

19.5–40.5

16.3–21.7

5.5–16.3

805.2–938.3 897.5–905.7

Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012

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1.3–2.5

19–24

91.6–98.6

424–601

F2

1.6–1.8 2.1–3.7

8.1–8.3

0.9–nd

3.6–25.7

22.6–88.7

237–518

F3

nd

1.2–nd

nd

nd

2.6–16.8

25.8–79.5

H

2.1–4.3 6.3–7.0

24.5–25.1

12.6–15.7

1.3–10.0

786.0–1271.9 681–1018

Two row, hull-less waxy barley CDC (n = 2) F1 3.3–6.5 3.5–6.6 15.0–20.5

2.6–2.7

17.6–54.6

14.8–55.9

867–567

F2

2.4–2.7 3.85–4.4

8.4–9.5

nd–1.2

13.2–14.3

7.8–28.8

511.5–383.7

F3

nd

nd–5.4

nd

nd–3.6

1.2–6.8

19.7–99.7

0.2–nd

nd–0.6

Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012 Gamel & Abdel-Aal 2012

Types and Distribution of Phenolic Compounds in Grains

Six row, hull normal barley (n = 2) F1 2.5–3.3 2.6–4.8 10.3–18.6

a

 RCA, protocatechuic acid; 4-OHBA, 4-hydroxybenzoic acid; VA, vanillic acid; SRA, syringic acid; CA, caffeic acid; p-CA, p-coumaric acid; FA, trans-ferulic P acid; SA, sinapic acid; IsoFA, iso-ferulic acid. b nd, not determined. c Numbers in parentheses are mean values.

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compounds compared with F1, but higher levels than F3 (the endosperm), which had the lowest levels, contributing only 1.2–1.9% ferulic acid, 0.9– 1.7% p-coumaric acid and caffeic > p-coumaric > ferulic acid isomers > sinapic acid. There was significant variation in the individual and total phenolic acids in the different pearling fractions. The concentrations of the five acids decreased significantly with the sequential removal of the outer layers. The first three pearling fractions represented about 20% surface removal and accounted for 60% of the total phenolic acids. The total phenolic acids were

Figure 12.5  Changes  in individual and total phenolic acid contents in (A) pearling

fines and (B) pearled kernels of white sorghum as the level of surface removal increased. Error bars represent standard deviation. Reprinted from J. Funct. Foods, 5 (4), D. L. Luthria and K. Liu, Localization of phenolic acids and antioxidant activity in sorghum kernels, 1751–1760, Copyright 2013, with permission from Elsevier.

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−1

four times lower in intact seed (1.4 mg g ) than in the first surface layer (5.50 mg g−1). Moraes et al.82 concurred with previous findings that phenolic acids were concentrated in sorghum bran on examining the correlation among polysaccharides, phenolic compounds, antioxidant activity and glycaemic index in sorghum samples of bran, decorticated sorghum flour and wholegrain flour. The extrusion processing of sorghum bran increased the levels of caffeic, p-coumaric, ferulic and sinapic acids at 180 °C and 20% moisture.94 Ferulic acid, the most abundant, increased by 2.7-fold in extruded bran (EB) compared with in non-extruded bran (NE). Extrusion has been reported to increase the total phenolic acids in other cereals, including barley, rice, oat and wheat, and at different temperatures.88,95 In the four fractions (dehulled, pearled, hull and bran) of foxtail and little millet, eight phenolic acids (gallic, p-hydroxybenzoic, vanillic, syringic, caffeic, chlorogenic, p-coumaric, ferulic and sinapic acids) were identified (Table 12.9).96 The hull had the highest concentration of p-coumaric and ferulic acid than the bran. The bran had the highest content of caffeic and sinapic acid. In foxtail millet hull, p-coumaric and ferulic acid were 21- and 5-fold higher than those in the little millet hull. In rye bran, unlike most bran fractions, sinapic was the second highest followed by p-coumaric while coarse rye fractions had 3-fold higher content of ferulic acid than rye bran97 (Table 12.9). In tartary buckwheat bran (Fagopyrum tartaricum (L.) Gaerth), caffeic acid hexose, chlorogenic and protocatechuic acids and catechin-glucoside were only detected in the bound phenolic fraction.98

12.6  Conclusions The health benefits of whole grain products have been associated with several phytochemicals, including phenolic acids. Phenolic acids, which exist in both free and bound forms, are classified into derivatives of hydroxybenzoic and hydroxycinnamic acids. The composition of these phenolic acids is similar in most cereals. They include gallic, protocatechuic, p-hydroxybenzoic, vanillic, syringic, salicylic, caffeic, trans-cinnamic, p-coumaric, ferulic and sinapic acids, with a few exceptions. Ferulic acid is the most abundant phenolic acid in all cereal grains, followed by p-coumaric acid depending on the type of the grain. Ferulic acid dimers and trimers have also been reported in maize, barley, wheat, millet and sorghum, with 8-O-4′ DFA and 8-5′ DFA as the most common dehydrodimers. However, some phenolic acids are present in specific cereals, including chlorogenic and protocatechuic acid, which are present in some rice varieties. Iso-ferulic acid has been reported in maize, wheat, barley and rice grains. The phenolic acid profiles of whole grains or grain fractions and the amounts reported in published work are influenced by the type of the grain, the genotype, the growing location, gene–environment interactions, the milling or botanical fraction and the type of extraction solvent. Wheat, maize, barley and sorghum tend to have a higher content of bound rather than soluble free forms, whereas rice has high levels of free compared with bound forms. In millet, hydroxybenzoates are higher in the

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Cereal type/fraction Soft sorghum bran (n = 4) Soft sorghum flour (n = 4) Hard sorghum bran (n = 4) Hard sorghum flour (n = 4) Sorghum bran (NEd) Sorghum bran (EBe) Foxtail millet (S + B f) Dehulled

GA

P-OHBA

VA

SRA

CA

ChlA

11–46

p-CA

FA

SA

Reference

103–175

2675–3532

51.5–78.6

140–198

169–205

8.7 ± 0.2 17.3 ± 0.2

19.8 ± 0.2 53.9 ± 1.6

3.4 ± 0.1 7.7 ± 0.2

Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Chiremba et al. 2012 Lopez et al. 2016 Lopez et al. 2016

83–136

14.9 19.9 4.32 ± 0.10

0.48 ± 0.00

2.63 ± 0.04

51.28 ± 3.62

14.58 ± 0.49

8.36 ± 0.23

25.09 ± 0.50

44.12 ± 1.55

Pearled

3.69 ± 0.10

0.28 ± 0.00

2.17 ± 0.06

17.45 ± 0.87

2.62 ± 0.02

1.50 ± 0.03

4.21 ± 0.07

28.25 ± 0.49

Hull

29.48 ± 3.02

5.97 ± 0.26

57.45 ± 1.67

152.52 ± 3.13

42.79 ± 1.39

6478.32 2420.21 ± 94.15 ± 56.15

227.08 ± 5.81

Bran

30.81 ± 1.81

11.42 ± 0.20

67.58 ± 2.32

335.89 ± 12.54

37.46 ± 1.03

595.81 1363.22 ± 15.60 ± 33.04

127.65 ± 1.69

268

Table 12.9  Distribution,  composition and content (µg g−1) of phenolic acids in grain fractions of minor cereals (sorghum, millet, rye).a,b

Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Chapter 12

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4.09 ± 0.08

4.88 ± 0.11

6.76 ± 0.13

95.4 ± 2.09

21.9 ± 0.84

5.0 ± 0.24

27.65 ± 0.70

53.18 ± 0.98

Pearled

1.36 ± 0.03

4.12 ± 0.10

3.15 ± 0.05

60.71 ± 1.61

0.93 ± 0.02

3.86 ± 0.07

17.43 ± 0.33

27.44 ± 0.29

Hull

38.41 ± 1.05

7.26 ± 0.19

30.96 ± 0.80

84.37 ± 1.98

95.35 ± 1.87

308.69 ± 7.63

456.19 ± 12.25

48.47 ± 0.86

Bran

64.99 ± 1.53

13.35 ± 0.26

73.97 ± 1.54

395.61 ± 3.60

127.32 ± 3.86

222.26 ± 4.79

824.2 ± 20.69

119.74 ± 2.19

Rye coarse fractionc

9.6

17.9

3.4

15.7

68.7

1309.6

178.3

Rye endospermc

2

6.4

1

3.6

29.9

233.8

39.7

Rye fine fractionc

4.6

6.4

2.8

11.1

42.5

492.7

108.3

Rye bran fractionc

17.9

19.2

4.5

24.1

125.2

3123.5

315.2

Rye bran R7c

22.1

19.6

5.6

27.2

141.7

2487.2

320.3

Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Pradeep & Sreerama, 2017 Pihlava et al. 2015 Pihlava et al. 2015 Pihlava et al. 2015 Pihlava et al. 2015 Pihlava et al. 2015

Types and Distribution of Phenolic Compounds in Grains

Little millet (S + B) Dehulled

a

 A, gallic acid; VA, vanillic acid; SRA, syringic acid; CA, caffeic acid; ChlA, chlorogenic acid; p-CA, p-coumaric acid; FA, trans-ferulic acid; SA, sinapic acid. G All values are total of free + bound phenolic acids. c mg kg−1 d NE, non-extruded bran. e EB, extruded bran. f S+B, soluble + bound phenolics.. b

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free rather than the bound forms, whereas hydroxycinnamates are higher in the bound rather than the free forms. In contrast with the composition, which is generally similar, the concentrations of phenolic acids in cereals vary widely across the different types of grains. Phenolic acids are unevenly distributed across the grain and are concentrated in the outer grain layers in all types of cereals. Thus the outer grain fractions, the bran and aleurone layers, are a more concentrated source of phenolic acids than the whole grain. The endosperm, as in refined flour, is essentially devoid or contains only trace amounts of phenolic acids.

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43. B. Min, L. Gu, A. M. McClung, C. J. Bergman and M.-H. Chen, Free and bound total phenolic concentrations, antioxidant capacities, and profiles of proanthocyanidins and anthocyanins in whole grain rice (Oryza sativa L.) of different bran colours, Food Chem., 2012, 133(3), 715. 44. R. Sompong, S. Siebenhandl-Ehn, G. Linsberger-Martin and E. Berghofer, Physicochemical and antioxidative properties of red and black rice varieties from Thailand, China and Sri Lanka, Food Chem., 2011, 124(1), 132. 45. N. V. M. de Mira, I. L. Massaretto, C. D. S. C. I. Pascual and U. M. L. Marquez, Comparative study of phenolic compounds in different Brazilian rice (Oryza sativa L.) genotypes, J. Food Compos. Anal., 2009, 22(5), 405–409. 46. F. Goffman and C. Bergman, Rice kernel phenolic content and its relationship with antiradical efficiency, J. Sci. Food Agric., 2004, 84(10), 1235. 47. P. Goufo, L. M. Ferreira, H. Trindade and E. A. Rosa, Distribution of antioxidant compounds in the grain of the Mediterranean rice variety ‘Ariete’, CyTA-J. Food, 2015, 13(1), 140. 48. Y. Zhu, T. Li, X. Fu, A. M. Abbasi, B. Zheng and R. H. Liu, Phenolics content, antioxidant and antiproliferative activities of dehulled highland barley (Hordeum vulgare L.), J. Funct. Foods, 2015, 19, 439. 49. A. Chandrasekara and F. Shahidi, Determination of antioxidant activity in free and hydrolyzed fractions of millet grains and characterization of their phenolic profiles by HPLC-DAD-ESI-MS n, J. Funct. Foods, 2011, 3(3), 144. 50. S. Siebenhandl, H. Grausgruber, N. Pellegrini, D. Del Rio, V. Fogliano and R. Pernice, et al., Phytochemical profile of main antioxidants in different fractions of purple and blue wheat, and black barley, J. Agric. Food Chem., 2007, 55(21), 8541. 51. G. Dinelli, I. Marotti, R. Di Silvestro, S. Bosi, V. Bregola and M. Accorsi, et al., Agronomic, nutritional and nutraceutical aspects of durum wheat (Triticum durum Desf.) cultivars under low input agricultural management, Ital. J. Agron., 2013, 8(2), 12. 52. T. Gamel and E.-S. M. Abdel-Aal, Phenolic acids and antioxidant properties of barley wholegrain and pearling fractions, Agric. Food Sci., 2012, 21(2), 118. 53. L. X. Lopez-Martinez, R. M. Oliart-Ros, G. Valerio-Alfaro, C.-H. Lee, K. L. Parkin and H. S. Garcia, Antioxidant activity, phenolic compounds and anthocyanins content of eighteen strains of Mexican maize, LWT-Food Sci. Technol., 2009, 42(6), 1187. 54. Q.-P. Hu and J.-G. Xu, Profiles of carotenoids, anthocyanins, phenolics, and antioxidant activity of selected color waxy corn grains during maturation, J. Agric. Food Chem., 2011, 59(5), 2026. 55. J.-G. Xu, Q.-P. Hu, X.-D. Wang, J.-Y. Luo, Y. Liu and C.-R. Tian, Changes in the main nutrients, phytochemicals, and antioxidant activity in yellow corn grain during maturation, J. Agric. Food Chem., 2010, 58(9), 5751.

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71. R. Fulcher, T. O'brien and J. Lee, Studies on the aleurone layer I. Conventional and fluorescence microscopy of the cell wall with emphasis on phenol-carbohydrate complexes in wheat, Aust. J. Biol. Sci., 1972, 25(1), 23. 72. V. Ndolo, T. Beta and R. Fulcher, Ferulic acid fluorescence intensity profiles and concentration measured by HPLC in pigmented and non-pigmented cereals, Food Res. Int., 2013, 52(1), 109. 73. D. Giordano, M. Locatelli, F. Travaglia, M. Bordiga, A. Reyneri and J. D. Coïsson, et al., Bioactive compound and antioxidant activity distribution in roller-milled and pearled fractions of conventional and pigmented wheat varieties, Food Chem., 2017, 233, 483. 74. G. G. Bellido and T. Beta, Anthocyanin composition and oxygen radical scavenging capacity (ORAC) of milled and pearled purple, black, and common barley, J. Agric. Food Chem., 2009, 57(3), 1022. 75. Y. Hemery, X. Rouau, V. Lullien-Pellerin, C. Barron and J. Abecassis, Dry processes to develop wheat fractions and products with enhanced nutritional quality, J. Cereal Sci., 2007, 46(3), 327. 76. Y. Hemery, U. Holopainen, A.-M. Lampi, P. Lehtinen, T. Nurmi and V. Piironen, et al., Potential of dry fractionation of wheat bran for the development of food ingredients, part II: electrostatic separation of particles, J. Cereal Sci., 2011, 53(1), 9. 77. M. Blandino, M. Alfieri, D. Giordano, F. Vanara and R. Redaelli, Distribution of bioactive compounds in maize fractions obtained in two different types of large scale milling processes, J. Cereal Sci., 2017, 77, 251. 78. C. M. Liyana-Pathirana and F. Shahidi, The antioxidant potential of milling fractions from breadwheat and durum, J. Cereal Sci., 2007, 45(3), 238. 79. C. M. Liyana-Pathirana and F. Shahidi, Importance of insoluble-bound phenolics to antioxidant properties of wheat, J. Agric. Food Chem., 2006, 54(4), 1256. 80. H. Zieliński and H. Kozłowska, Antioxidant activity and total phenolics in selected cereal grains and their different morphological fractions, J. Agric. Food Chem., 2000, 48(6), 2008. 81. A. K. Das and V. Singh, Antioxidative free and bound phenolic constituents in botanical fractions of Indian specialty maize (Zea mays L.) genotypes, Food Chem., 2016, 201, 298. 82. É. A. Moraes, R. da Silva Marineli, S. A. Lenquiste, C. J. Steel, C. B. de Menezes and V. A. V. Queiroz, et al., Sorghum flour fractions: Correlations among polysaccharides, phenolic compounds, antioxidant activity and glycemic index, Food Chem., 2015, 180, 116. 83. C. Barron, A. Surget and X. Rouau, Relative amounts of tissues in mature wheat (Triticum aestivum L.) grain and their carbohydrate and phenolic acid composition, J. Cereal Sci., 2007, 45(1), 88.

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84. F. Brouns, Y. Hemery, R. Price and N. M. Anson, Wheat aleurone: separation, composition, health aspects, and potential food use, Crit. Rev. Food Sci. Nutr., 2012, 52(6), 553. 85. J. S. Sidhu, Y. Kabir and F. G. Huffman, Functional foods from cereal grains, Int. J. Food Prop., 2007, 10(2), 231. 86. V. Sovrani, M. Blandino, V. Scarpino, A. Reyneri, J. D. Coïsson and F. Travaglia, et al., Bioactive compound content, antioxidant activity, deoxynivalenol and heavy metal contamination of pearled wheat fractions, Food Chem., 2012, 135(1), 39. 87. L. R. Brewer, J. Kubola, S. Siriamornpun, T. J. Herald and Y.-C. Shi, Wheat bran particle size influence on phytochemical extractability and antioxidant properties, Food Chem., 2014, 152, 483. 88. H. Ti, R. Zhang, M. Zhang, Z. Wei, J. Chi and Y. Deng, et al., Effect of extrusion on phytochemical profiles in milled fractions of black rice, Food Chem., 2015, 178, 186. 89. H. Ti, Q. Li, R. Zhang, M. Zhang, Y. Deng and Z. Wei, et al., Free and bound phenolic profiles and antioxidant activity of milled fractions of different indica rice varieties cultivated in southern China, Food Chem., 2014, 159, 166. 90. Y. Pang, S. Ahmed, Y. Xu, T. Beta, Z. Zhu and Y. Shao, et al., Bound phenolic compounds and antioxidant properties of whole grain and bran of white, red and black rice, Food Chem., 2018, 240, 212. 91. M. Zaupa, L. Calani, D. Del Rio, F. Brighenti and N. Pellegrini, Characterization of total antioxidant capacity and (poly) phenolic compounds of differently pigmented rice varieties and their changes during domestic cooking, Food Chem., 2015, 187, 338. 92. P. Van Hung, Phenolic compounds of cereals and their antioxidant capacity, Crit. Rev. Food Sci. Nutr., 2016, 56(1), 25. 93. J. M. Awika and L. W. Rooney, Sorghum phytochemicals and their potential impact on human health, Phytochemistry, 2004, 65(9), 1199. 94. N. J. Salazar Lopez, G. Loarca-Piña, R. Campos-Vega, M. Gaytán Martínez, E. Morales Sánchez, J. M. Esquerra-Brauer, G. A. Gonzalez-Aguilar and M. R. Sánchez, The extrusion process as an alternative for improving the biological potential of sorghum bran: Phenolic compounds and antiradical and anti-inflammatory capacity, Evidence-Based Complementary Altern. Med., 2016, 2016, 1. 95. H. Zielinski, H. Kozlowska and B. Lewczuk, Bioactive compounds in the cereal grains before and after hydrothermal processing, Innovative Food Sci. Emerging Technol., 2001, 2(3), 159. 96. P. Pradeep and Y. N. Sreerama, Soluble and bound phenolics of two different millet genera and their milled fractions: Comparative evaluation of antioxidant properties and inhibitory effects on starch hydrolysing enzyme activities, J. Funct. Foods, 2017, 35, 682. 97. J.-M. Pihlava, E. Nordlund, R.-L. Heiniö, V. Hietaniemi, P. Lehtinen and K. Poutanen, Phenolic compounds in wholegrain rye and its fractions, J. Food Compos. Anal., 2015, 38, 89.

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98. F. Li, X. Zhang, S. Zheng, K. Lu, G. Zhao and J. Ming, The composition, antioxidant and antiproliferative capacities of phenolic compounds extracted from tartary buckwheat bran [Fagopyrum tartaricum (L.) Gaerth], J. Funct. Foods, 2016, 22, 145. 99. N. L. Kent and A. D. Evers, Kent's Technology of Cereals, Woodhead Publishing, Cambridge, 4th edn, 1994.

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Chapter 13

Bound Phenolic Constituents as Co-passengers of Dietary Fibre Franklin Brian Apea-Bah*a,b and Trust Betaa a

Department of Food and Human Nutritional Sciences, University of Manitoba, Winnipeg, Manitoba, Canada; bBiotechnology and Nuclear Agriculture Research Institute, Ghana Atomic Energy Commission, P.O. Box LG 80, Accra, Ghana *E-mail: [email protected]

13.1  Introduction Phenolic acids are the most abundant phytochemicals present in the dietary fibre of cereals and consist of hydroxycinnamic and hydroxybenzoic acids. Hydroxycinnamic acids, such as ferulic acid, are usually bound to arabinoxylans in the cell wall of cereals through ester linkages.1 Other hydroxycinnamates, such as p-coumaric acid and sinapic acid, also interact with arabinoxylans, lignins and proteins through covalent associations.1–3 In addition to the hydroxycinnamates, hydroxybenzoates have also been found to be associated with cell wall components, albeit to a lesser extent. Bound phenolic acids contribute to cross-linkages between the arabinoxylan molecules and therefore strengthen the cell wall networks. They provide protection to whole grains against insect pests and microbial attack and therefore play a key part in plant defence.4 Phenolic compounds are all derived from the phenylpropanoid pathway using l-phenylalanine, the end-product of the   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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shikimic acid pathway, as their precursor. l-Phenylalanine therefore serves as a link between the primary metabolites and secondary metabolites in plant systems.5 Most of the beneficial health effects of whole grain cereals are attributed to dietary fibre and phenolic compounds, including bound phenolic acids. However, the most widely studied beneficial health effect of bound phenolic acids is their antioxidant properties. Bound phenolic acids can donate hydrogen atoms to quench free radicals, such as reactive oxygen and nitrogen species, which, when overproduced in the body and not effectively controlled, can react with and damage bioactive macromolecules (e.g. bioactive proteins, lipids and DNA). This damage may lead to diseases related to oxidative stress, such as cardiovascular and coronary heart disease, neurodegenerative disease and various cancers, including colorectal cancer. For cereal grains to be palatable for consumption, they need to be cooked using various processing operations. Some of these processing operations can affect the bound phenolic composition of the grains by disrupting the cell wall structure and promoting the release of bound phenolic acids. The effect of these processing operations is important because they ultimately affect the antioxidant properties of the bound phenolic acids in the resulting food product. Water-insoluble dietary fibre is abundant in whole grain cereals and is covalently associated with bound phenolic acids. The bound phenolic acids are therefore referred to as co-passengers of dietary fibre. They are believed to be transported through the gastrointestinal tract in this form without undergoing digestion until they reach the colon, where they are metabolized by the colonic microbiota and released for absorption into the systemic circulation.

13.2  C  hemistry and Biochemistry of Phenolic Compounds Bound to Dietary Fibre in Cereals 13.2.1  Chemistry of Phenolic Acids The major types of bound phenolic acids in cereals are the hydroxycinnamic acids, with ferulic acid being the most predominant, followed by the diferulic acids, which are formed by the covalent binding of two ferulic acid molecules. Other hydroxycinnamic acids (p-coumaric acid, sinapic acid and caffeic acid) are also present in varying amounts depending on the cereal type and variety (Tables 13.1 and 13.2). Hydroxybenzoic acids (e.g. protocatechuic acid, p-hydroxybenzoic acid, syringic acid and vanillic acid) have also been reported in cereals.6,7 A number of ferulic acid dimers and trimers have been reported in bound forms in cereals, including 8-8′-dehydrodiferulic acid, 8-5′-dehydrodiferulic acid, 8-O-4′-dehydrodiferulic acid, 5-5′-dehydrodiferulic acid and 4-O-5′-dehydrodiferulic acid, as well as 5-5′/8′-O-4″-dehydrotriferulic acid.8,9

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mented cereals.a

Monomeric phenolic acid Sample

Class PCA

CFA

SYA

13.22 71.72 4.58 35.61 22.12 4.11 — 13.91 — 6.07 34.63 72.75 20.14 10.21 — 18.29 — 20.44 443.90 1349.50 31.00 558.90 —

65.90

— —

497.20 —

—

—

188.61 — — — — — — —

— — — — — —

59.80 248.80 —

11.32 14.29

166.90 272.70

450.30 —

31.29 29.37 101.07 81.66 576.52 123.00

27.80 510.50

756.40 1328.4 —

p-COA

200.4

13.86 10.90 45.67 9.97 — — — — — — — — — — — —

Dimeric ferulic acid

FA

SIA

iso-FA 8-5′

5-5′

8-O-4′

Total

2287.56 43.37 186.7 30.79 27.68 46.55 2743.46 1412.16 28.88 27.54 31.46 16.49 35.59 1709.23 3543.82 32.71 155.00 43.35 27.90 55.96 3979.79 3081.57 66.14 75.54 28.94 13.41 37.75 3522.74 8322.16 107.56 47.38 112.30 79.17 140.61 9424.43 1823.10 — — — — — —

Reference 6 6 6 6 6 101

68.20 2324.90 —

—

—

—

—

—

101

5.10 —

—

—

—

—

—

101

136.50 2730.40 —

—

—

—

—

—

101

250.80

—

—

—

—

—

101

—

—

—

—

—

101

31.11 15.17 — — — — — —

16.04 17.14 — — — — — —

9.16 8.52 — — — — — —

18.52 27.80 — — — — — —

2609.48 1041.92 — — — — — —

6 6 102 102 102 102 102 102

—

0.45

332.20 — 22.9

—

425.29 1814.34 70.33 36.74 829.17 37.45 842.61 592.92 — 1139.06 325.50 — 36.70 294.70 — 679.52 1685.04 — 917.64 178.82 — 20.68 638.91 —

Chapter 13

Barley IDF Purple barley Wheat Purple wheat Yellow corn Dent corn pericarpc Dent corn germc Dent corn endospermc Flint corn pericarpc Flint corn germc Flint corn endospermc Red rice Oats Foxtail millet Proso millet Finger milletd Kodo millet Little millet Pearl millet

VNA

c

280

Table 13.1  Composition  of phenolic acids (µg g−1 sample)b derived from insoluble and soluble dietary fibre in pigmented and non-pig-

a

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SDF

— — — — — — — — — — — — —

— — — — — — — — — — — — —

— — — — — — — — — — — — —

— — — — — — — — — — — — —

15.29 21.45 15.03 15.43 31.96 31.94 20.59 54.18 15.86 0.89 14.31 70.31 16.96

146.72 140.06 94.87 99.46 103.59 100.67 107.41 93.14 28.95 12.34 196.18 94.09 60.14

12.00 15.63 5.90 3.99 32.27 26.31 8.64 — — — — — —

13.05 10.39 9.65 13.07 13.10 14.72 8.71 — — — — — —

— — — — — — — — — — — — —

— — — — — — — — — — — — —

— — — — — — — — — — — — —

187.06 187.53 125.45 131.95 180.92 173.64 145.35 — — — — — —

6 6 6 6 6 6 6 102 102 102 102 102 102

I DF, insoluble dietary fibre; SDF, soluble dietary fibre; PCA, protocatechuic acid; VNA, vanillic acid; CFA, caffeic acid; SYA, syringic acid; p-COA, p-coumaric acid; FA, ferulic acid; SIA, sinapic acid; iso-FA, iso-ferulic acid. Values are expressed as the mean of two measurements (standard deviation is not shown). c Not detected. d Values are mean of two varieties. b

Bound Phenolic Constituents as Co-passengers of Dietary Fibre

Barley Purple barley Wheat Purple wheat Yellow corn Red rice Oat Foxtail millet Proso millet Finger milletd Kodo millet Little millet Pearl millet

281

soluble dietary fibre (SDF).a

Sample b

a

TPC

TEAC

DPPH

ORAC

H2O2

FRAP

IDF

2.72–7.11 6.16 8.81 9.09 18.35 42.45 24.23 1.26 42.60 17.57 1.48 15.04 3.97 11.59 2.21 3.52 18.64 9.64 9.14 2.10 6.60–0.85 1.33 0.88 1.05 1.45 1.81 1.13

8.59 — — — — 798.90 457.60 12.5 357.10 416.50 63.70

4.54–11.21 9.45 11.13 12.47 18.72 1667.60 1262.90 21.80 1534.80 1133.50 60.80 18.55 6.77

28.14 — — — — — — — — — —

— — — — 367.60 217.50 39.50 127.00 113.20 21.6

— — — — 27.40 16.10 0.70 27.70 11.80 2.60

2300.00 1.04–2.42 1.03 0.88 0.95 1.00 1.10 0.93

— 15.09

SDF

40.61 11.14 5.90 86.13 10.50 3.02

8.25 2.96 6.70 29.33 —

—

Reference 6 and 103 6 6 6 6 101 101 101 101 101 101 6 6 102 102 102 102 102 102 57 6 and 103 6 6 6 6 6 6

TPC, total phenolic content (mg FAE/g sample); DPPH, 2,2′-diphenyl-1-picrylhydrazyl (µmol TE/g sample); TEAC, trolox equivalent antioxidant capacity (µmol TE/g sample); ORAC, oxygen radical absorbance capacity (µmol TE/g sample); FRAP, ferric-reducing antioxidant power (µmol AAE/g defatted sample); FAE, ferulic acid equivalent; TE, trolox equivalent; AAE, ascorbic acid equivalent. b Range of mean values from different studies. c TPC values converted from µmol FAE/g to mg FAE/g. FRAP values expressed as µmol TE/g sample. d values mean of two varieties.

Chapter 13

Barley Purple barley Wheat Purple wheat Yellow corn Dent corn pericarpc Dent corn germc Dent corn endospermc Flint corn pericarpc Flint corn germc Flint corn endospermc Red rice Oat Foxtail millet Proso millet Finger milletd Kodo millet Little millet Pearl millet Red, non-tannin sorghum Barley Purple barley Wheat Purple wheat Yellow corn Red rice Oats

Class

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Table 13.2  Total  phenolic content and radical scavenging capacities of phenolic extracts derived from insoluble dietary fibre (IDF) and

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Bound Phenolic Constituents as Co-passengers of Dietary Fibre

Figure 13.1  Bound  phenolic acids in cereals. (A) Ferulic acid, (B) p-coumaric acid,

(C) caffeic acid, (D) sinapic acid, (E) protocatechuic acid, (F) vanillic acid, (G) syringic acid, (H) 8,5′-diferulic acid, (I) 8,8′-diferulic acid, (J) 5,5′-diferulic acid, (K) 8-O-4′-diferulic acid, (L) 4-O-5′-diferulic acid and (M) 5-5'/8′-O-4″-triferulic acid. Reproduced from, M. Bunzel, Chemistry and occurrence of hydroxycinnamate oligomers, Phytochem. Rev., 9, 2009, 47–64, Copyright Springer Science + Business Media B.V. 2009. With permission of Springer.

Figure 13.1 shows the structures of some of the bound phenolic acids present in cereals. Most cereals have higher levels of phenolic acids bound to insoluble dietary fibre than to soluble dietary fibre. In comparing the levels of different forms of ferulic acid in whole grain cereals

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10

(maize, wheat, oats and rice), Adom and Liu noted that the free, soluble-conjugated and bound forms existed in the ratio 0.1 : 1 : 100. The bound phenolic acids are covalently bound to the arabinoxylan backbone in hemicellulose through ester linkages between the carboxylic group of the phenolic acid and the hydroxyl groups of the arabinoxylan. Some phenolic acids act as a point for intramolecular and intermolecular crosslinks between the arabinoxylan molecules and contribute to the integrity and mechanical strength of the cell walls. Although ferulic acid and its dimers are the major cross-linking phenolic compounds,11 p-coumaric acid dimers, sinapic acid dehydrodimers and ferulic acid dehydrotrimers also contribute to the cross-linking, thereby strengthening the network of the cell wall polysaccharides.11

13.2.2  Biosynthesis of Phenolic Acids Phenolic acids, whether hydroxycinnamates or hydroxybenzoates, originate from the aromatic amino acid l-phenylalanine through the phenylpropanoid pathway via a series of biosynthetic steps. The details of the biosynthetic cascade, with the associated enzymes, coenzymes and genes coding for the expression of the enzymes, have been reviewed by Fraser and Chapple.12 In the first step, l-phenylalanine is deaminated by the enzyme phenylalanine ammonia lyase to form trans-cinnamic acid. In the second step, trans-cinnamic acid is hydroxylated by the enzyme cinnamate 4-hydroxylase (trans-cinnamate 4-monooxygenase), in the presence of the coenzyme reduced nicotinamide adenine dinucleotide phosphate, to form p-coumaric acid (4-hydroxycinnamic acid). p-Coumaric acid is then converted to its coenzyme A (CoA) thioester, 4-hydroxycinnamoyl-CoA (p-coumaroyl-CoA) in the presence of the enzyme 4-coumarate-CoA ligase. Other reactions following the formation of p-coumaroyl-CoA involve hydroxylation with or without methylation to yield the other hydroxycinnamic (caffeic, ferulic and sinapic) acids. A schematic diagram of the biosynthesis is shown in Figure 13.2. Two biosynthetic pathways have been proposed for hydroxybenzoic acids, with the main pathway involving the loss of an acetate group from the sidechain of the corresponding hydroxycinnamic acids. The alternative pathway, which derives from an intermediate in the shikimic acid pathway, involves a series of enzymatic reactions that convert 3-dehydroshikimic acid to various hydroxybenzoic acids.13,14

13.2.3  Dimerization of Hydroxycinnamates Ferulates, p-coumarates and sinapates are able to react with each other to form dimers through one of two reaction mechanisms: (1) through oxidative or radical-induced coupling reactions; and (2) through photochemical (light)-induced coupling reactions. Phenoxy radicals, for example, can be generated in ferulic acid within the cell wall to form dehydrodiferulate esters. Electron delocalized phenoxy radicals couple at the 4-O-, 8-O-, C5- or

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285

Figure 13.2  Schematic  representation of the phenylpropanoid pathway show-

ing the biosynthesis of hydroxycinnamic acids. PAL, phenylalanine ammonia lyase; C4H, cinnamate 4-hydroxylase; CCL, 4-coumarate-CoA ligase; HCT, hydroxycinnamoyl-CoA shikimate:quinate hydroxycinnamoyl-transferase; C3′H, p-coumaroyl shikimate 3′-hydroxylase; CCoAOMT, caffeoyl-CoA 3-O-methyltransferase; CCR, cinnamoyl-CoA reductase; F5H, ferulate 5-hydroxylase; COMT, caffeic acid/5-hydroxyferulic acid O-methyltransferase; HCALDH, hydroxycinnamaldehyde dehydrogenase; CoA, coenzyme A. Adapted with permission of American Society of Plant Biologists, from Fraser and Chapple;12 permission conveyed through Copyright Clearance Center, Inc.

C8- positions to give rise to 4-O-5′, 8-O-4′, 5-5′, 8-5′ and 8-8′ coupled diferulate esters. The detailed reaction mechanisms and the formation of other oligomers, such as ferulic acid trimers and tetramers, have been reviewed by Bunzel.1 It is worth noting that some of these oligomers have been identified in maize and other cereals.8,9,15,16

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13.3  S  ample Preparation, Extraction and Analysis of Phenolic Compounds Bound to Dietary Fibre in Cereals 13.3.1  S  ample Preparation and Extraction of Soluble and Insoluble Dietary Fibre Several researchers have reported methods for the sample preparation and extraction of phenolic compounds bound to the soluble and insoluble dietary fibre fractions in cereals.6,9 Soluble and insoluble dietary fibre are obtained from the whole grain cereals by the enzymatic hydrolysis of carbohydrates using heat-stable α-amylase and amyloglucosidase, followed by protein hydrolysis using proteases, all at elevated temperatures. The reaction mixture is then centrifuged and both the supernatant and residue are kept. The residue is washed with hot distilled water, 95% aqueous ethanol and 95% aqueous acetone. It is then filtered under vacuum, placed in a fume hood to remove the organic solvent and dried in a vacuum oven at 35–40 °C to obtain the insoluble dietary fibre. The supernatant (which contains the sugars, peptides and amino acids from the enzyme hydrolysis, as well as the free and soluble bound phenolic compounds) is combined with the washings from the residue and precipitated in four volumes of 80% aqueous ethanol (preheated to 60 °C) overnight. The precipitate is placed in a fume hood to remove the organic solvent and then dried as before to give the soluble dietary fibre. The soluble and insoluble dietary fibre may then be milled into fine powders of the desired particle size.

13.3.2  Extraction of Bound Phenolic Acids The phenolic acids covalently bound to cell wall polysaccharides are released either by acid or alkaline hydrolysis. Alkaline hydrolysis (2 or 4 M NaOH), however, is more efficient in breaking the ester and other covalent cross-links between phenolic acids and cell wall polysaccharides in dietary fibre17 and is therefore the preferred method for releasing bound phenolic acids.6,7,11 This step is usually performed under nitrogen gas to prevent the oxidation of the phenolic acids. The phenolate salts formed are acidified to pH 1.5–2 using a strong acid (6 or 12 M HCl) to release the phenolic acids, which are then extracted using an organic solvent immiscible with the aqueous medium, such as ethyl acetate, diethyl ether or equal volumes of the two solvents. The liquid–liquid extraction may be carried out three or four times with about twice the volume of the aqueous medium to ensure as much extraction of the phenolic acids as possible. The extract is then dried, either using a rotary evaporator or under a stream of nitrogen gas, and reconstituted in an appropriate organic solvent (methanol, ethanol or their aqueous derivative) for analysis.

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13.3.3  A  nalysis of Phenolic Acids Bound to the Cell Wall in Cereals Liquid chromatography–mass spectrometry (LC–MS) is the preferred method for the analysis of bound phenolic acids in cereals.6,7,18 This method is preferred because, under suitable operating conditions, the liquid chromatographs separate the non-volatile phenolic compounds based on their relative polarity and adsorption properties on a reversed-phase column, before introducing each compound to a detector. Gas chromatography–mass spectrometry has also been used for the analysis of bound phenolic acids. However, this method requires the derivatization of the non-volatile compounds into the dimethylsilylated volatile forms.9,11,15 A known volume of the reconstituted extract is filtered through a 0.45 or 0.2 µm membrane filter and then injected onto the liquid chromatography column. Gradient elution with a mobile phase system – usually involving purified water acidified with formic, acetic or trifluoroacetic acid as solvent A and acidified alcohol (methanol or ethanol) as solvent B – is used to separate the constituent phenolic acids on a reversed-phase column. Separation is based on the relative interactions between the analytes (phenolic acids) and the mobile phase and stationary phase particles. The nature and strength of the interactions depend on the respective adsorption properties and relative polarities of the analytes. The separated compounds are detected by a UV–visible detector set at a particular wavelength, or a photodiode array detector that can scan the analytes over a wide wavelength range, usually 200–800 nm. Phenolic acids usually show a maximum absorption at 80% of the total anthocyanins; peonidin-3-O-glucoside is also significant, accounting for about 10–15% of the anthocyanins in black rice.

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Figure 14.4  Effect  of increasing lime concentration on visual appearance of different color classes of maize: A, red/blue; B, blue; C, red; and D, purple maize. The left-hand column is 0% lime, 0.5% lime is in the middle and 1.0% lime is in the far right-hand column. Reprinted with permission from A. Collison, L. Yang, L. Dykes, S. Murray and J. M. Awika, J. Agric. Food Chem., 2015, 63 (22), 5528–5538. Copyright (2015) American Chemical Society.19

The anthocyanin pigments in rice are typically located in the bran layer, which is exposed after the rice husk is removed. Because the normal rice polishing process results in a highly concentrated source of these pigments in bran,18 rice appears to be an attractive source of pigments for commercial

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extraction as food colorants. However, a major disadvantage that negatively affects the exploitation of black rice pigments as natural colorants is the fact that the content of the more stable acylated anthocyanins is generally low to non-existent. Nevertheless, pigmented rice types are gaining interest in health food applications. An interesting new development is the transformation of rice to produce anthocyanins in the endosperm through genetic engineering.3 Through transgene stacking, it was possible to produce purple endosperm rice with up to 1000 mg g−1 anthocyanins in the endosperm. This accomplishment is important because the endosperm forms the bulk of cereal grains, especially rice (>90%). Thus such a high level of anthocyanin accumulation in the endosperm implies that the net yield of anthocyanins from cereal grains can be increased several-fold. This type of grain could be used at low levels in food products to provide the health benefits associated with anthocyanins. Also, with the concomitant enhancement of the acylated anthocyanin biosynthetic pathway, rice could be a competitive source of natural food colorants.

14.2.5  Anthocyanins in Wheat and Barley Anthocyanins are present in diverse colored species of barley and wheat, both domesticated and wild. These compounds have been identified in Triticum aestivum (bread wheat), T. durum (pasta wheat) and various species of domesticated and wild einkorn.24–26 Like other cereal grains, the anthocyanins are either localized in the pericarp or aleurone layer of the seed. Values can vary widely depending on the phenotype, but average anthocyanin content of pigmented wheat and barley range between 50 and 600 µg g−1 (Table 14.1), with values that are five to six times higher in the bran fractions.27–29 The profile of anthocyanins in wheat, as well as pigmented barley, are generally similar to those reported for rice, with cyanidin-3-O-glucoside being the most common compound. However, blue wheat appears to contain delphinidin-based anthocyanins (primarily delphinidin-3-O-glucoside and delphinidin-3-O-rutinoside) as the dominant pigments.18 Similarly, blue barley also contains delphinidin-3-O-glucoside and delphinidin-3-O-malonylglucoside as the major anthocyanins.27 This profile is in contrast with maize and rice, where the delphinidin-based pigments have not been identified. With the exception of some barley varieties,27 pigmented wheat and barley are generally low in acylated anthocyanins. The low level of acylated anthocyanins in wheat and barley severely limits the potential application of these pigments as natural food colorants due to their poor stability. Diczházi and Kursinszki27 identified malonated delphinidin and cyanidin glucosides as major components of anthocyanins in three varieties of barley from a Hungarian collection, accounting for about 50% of the total anthocyanins. This finding suggests that specific varieties of barley could potentially be a source of valuable commercial food colorants.

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14.2.6  3-Deoxyanthocyanins in Sorghum The 3-deoxyanthocyanins (Figure 14.3) are a unique feature of sorghum as a cereal grain, or more broadly, as a food crop. Mature sorghum plants do not appear to synthesize anthocyanins in detectable quantities and instead exclusively accumulate the 3-deoxyanthocyanins in various plant tissues, including the seed pericarp. Unlike the anthocyanins ubiquitous in most plants, the 3-deoxyanthocyanin pigments are unsubstituted at the C-3 position. This small structural difference is highly relevant to food processing because it significantly stabilizes the structure of the molecule and makes it less susceptible to nucleophilic attack and unfavorable structural transformations.30–34 The 3-deoxyanthocyanins also show important differences in their bioactive properties compared with their anthocyanin analogs.35,36 For these reasons, the sorghum pigments are of major interest to the food industry as functional natural food colorants. The genetics of biosynthesis of the 3-deoxyanthocyanins is largely known. The yellow seed1 (ys1) gene found in most sorghum varieties controls the biosynthetic pathway that leads to the accumulation of 3-deoxyflavanoid compounds (including 3-deoxyanthocyanins).37 The synthesis of the 3-deoxyanthocyanins in sorghum is controlled by a set of two genes, R and Y. A homozygous recessive yy will produce a white pericarp without any pigment, whereas a recessive rr gene will produce a yellow pericarp with very little 3-deoxyanthocyanin. Dominance at R_ and Y_ will result in a red pericarp;38 these sorghums accumulate significant levels of the pigments. Other interesting traits have also been observed in sorghum pericarp pigmentation—for example, some red sorghum varieties will turn black during grain maturation as a result of exposure to UV light. This trait is due to the enhanced synthesis of the 3-deoxyanthocyanin compounds in the pericarp tissue, apparently an inducible response.39 The genetic basis for the response is not fully understood, but it suggests that these sorghum varieties may be producing these compounds as a means of protecting the seed against UV radiation. The 3-deoxyanthocyanins are also known to be synthesized in sorghum as phytoalexins in response to pathogens or pest invasion.40 Even though the anthocyanin biosynthetic pathway is functional in sorghum, as observed in seedlings grown in sterile environments,41 the enzymes responsible appears to be almost completely suppressed when sorghum is exposed to biotic stress, as is inevitable when plants are grown in a normal environment. Thus the 3-deoxyanthocyanins are the only detectable pigments in mature sorghum plants and grain. Some sorghum landraces native to West Africa accumulate very high levels of the 3-deoxyanthocyanins in their leaf sheath, with values up to 90 mg g−1 reported in some varieties;31,42 these values are 6–15 times higher than those reported in black sorghum bran (4–16 mg g−1 bran).6,43 These sorghums landraces are grown primarily for their pigments and are thus called dye sorghums. The pigments from these sorghums have been used

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for centuries, not only for food coloring (traditional cheese and porridge), but also for diverse applications such as dyeing leather and in traditional medicine.42 These sorghums could, at a minimum, provide a source of genetic material for engineering the enhanced biosynthesis of 3-deoxyanthocyanins in hybrid sorghum tissue for large-scale commercial use. In fact, mutagenesis breeding has been used to induce the over-accumulation of 3-deoxyanthocyanins in the leaves of high biomass sorghum, with promising results.4 Because these non-grain tissues are a much larger part of the plant biomass, there is an opportunity to exploit them as a commercial source of the pigments.

14.2.7  S  tructure of Sorghum 3-deoxyanthocyanins and Basis for Color Stability A unique property of the 3-deoxyanthocyanins is that they exist in nature primarily as aglycones, i.e. without sugar substituents. This characteristic is unusual for most flavonoids (including anthocyanins) or other phenolic compounds found in nature. Most phenolic compounds in nature are conjugated to sugars and organic acids, either as free molecules in the cell vacuole or as components of cell wall polysaccharides. Anthocyanins are especially unstable at near-physiological pH and are thus always conjugated for improved stability. Perhaps, due to their inherent stability, the 3-deoxyanthocyanins do not require conjugation. Alternatively, the fact that these compounds are inducible phytoalexins may make the more reactive aglycone more advantageous to the sorghum plant. The 3-deoxyanthocyanins of sorghum are composed primarily of apigeninidin and luteolinidin and their derivatives (Figure 14.3). Depending on the sorghum variety, the 3-deoxyanthocyanidin aglycones can constitute 75–98% of the extractable sorghum pigments, in sharp contrast to anthocyanins, which are >99% conjugated. The aglycones are commonly O-methyl substituted at position 7 (C7 of the A-ring), with the 7-O-methylapigenindin and 7-O-methylluteolinidin accounting for 20–50% of the aglycones in most pigmented sorghum.35 Because the favored glycosyl substitution position in anthocyanins, C3, is unsubstituted, the 3-deoxyanthocyanins are most commonly glycosylated at position 5 (C5 of the A-ring).35 Di-glycosides and acyl-glycosides of these compounds have also been identified in sorghum leaves and germinating seedlings, respectively.4,44 Condensed flavene dimers of apigeninidin and pyrano-apigeninidin (pyrano-apigeninidin-4-vinylphenol) have also been identified in sorghum leaf sheath (Figure 14.3).31,45 The condensed apigeninidin dimers were shown to be highly resistant to pigment degradation induced by a changing pH and oxidizing agents such as sulfur dioxide, and are thus of significant interest in food processing. The pyrano-apigeninidin is also likely to have improved stability due to the protective effect of cyclic addition at C4 and C5, which protects the reactive C5.32

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Figure 14.5  Structural  transformation of 3-deoxyanthocyanidins and anthocyani-

dins in aqueous solution. Unlike in anthocyanins where equilibrium favors hydration reactions in moderate to mildly acidic environments (Kh > Ka), leading to the formation of colorless chalcone, the 3-deoxyanthocyanins have Ka > Kh at most pH values, hence deprotonation is the favored reaction.30,79

An important consequence of the lack of substitution at C-3 of 3-deoxyanthocyanins is that it results in a region between C-5 and C-4′ (Figures 14.1 and 14.3) that has greater hydrophobicity than their anthocyanin analogs and is less reactive with hydrophilic molecules. This region causes 3-deoxyanthocyanins to be less susceptible to nucleophilic attack and hydration, which is a primary mechanism for the structural transformation of anthocyanins in solution to colorless forms (Figure 14.5). For example, monomeric anthocyanins are almost completely colorless in the pH range 4.0–5.0 due to the near-complete transformation of the flavylium cation into the colorless carbinol pseudobase. The carbinol pseudobase is highly reactive and readily transforms into the open-ring chalcones, which can participate in other degradative reactions during food processing.34 The 3-deoxyanthocyanins, however, resist hydration and retain most of their flavylium cation over a broad acidic pH range. Their favored transformation as the pH increases is deprotonation (Figure 14.5), which leads to reddish blue quinoidal bases, which are more stable than chalcones. Their reduced hydration constant also makes the 3-deoxyanthocyanins a lot more resistant to thermal degradation than anthocyanins because chalcone formation is the primary path for the thermal degradation of these compounds.34 Consequently, the 3-deoxyanthocyanins have a relatively stable color profile over a broad pH range (Figure 14.6) at high temperatures and on exposure to oxidizing agents such as ascorbic acid and sulfites.31–33 This trait is an

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Figure 14.6  Effect  of pH on sorghum 3-deoxyanthocyanin pigment hue compared with anthocyanin pigments from fruits. Color stability and predictable hue over a broad pH range is a distinct advantage of the 3-deoxyanthocyanins. Reprinted from Gluten-Free Ancient Grains, A volume in Woodhead Publishing Series in Food Science, Technology and Nutrition, Joseph M. Awika, Chapter 3 – Sorghum: Its Unique Nutritional and Health-Promoting Attributes, 21–54, Copyright 2017, with permission from Elsevier.5

advantage in providing predictable color hues in foods. An additional advantage of the 3-deoxyanthocyanins is that they provide red hues at near-neutral pH, complementing the anthocyanins, which tend to be blue at neutral pH. As a result of their attractive advantages, and the growing interest in natural ingredients, various food and ingredient companies are currently testing the sorghum 3-deoxyanthocyanins. Two important challenges that limit the use of the 3-deoxyanthocyanins include their relatively poor extractability from plant (especially bran) tissue and their tendency to self-associate in aqueous systems. The pigments are located in cell vacuoles within the pericarp cells, high in the cellulosic cell wall material. Thus disruption of the cell wall is essential to improving their extractability. As a result of their relatively high thermal stability,35 high-temperature systems can probably be exploited to enhance the extractability of these compounds in aqueous systems. However, a recent test using an accelerated solvent extraction system combining high temperatures and pressures did not produce positive results.46 Preliminary work from our group suggests that microwave-assisted extraction is a likely effective alternative with the choice of the right solvent.

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The self-association of 3-deoxyanthocyanins in aqueous systems is largely due to their reduced hydrophilicity relative to anthocyanins. The fact that they exist mainly as aglycones further exacerbates this problem. This problem can most likely be mitigated through encapsulation with hydrocolloids and stabilizers with an appropriate hydrophilic–lipophilic balance. The 3-deoxyanthocyanins can also be readily used in products where suspension in the water phase is not essential, e.g. alcoholic beverages, as well various non-liquid food applications, e.g. snacks, breakfast cereals, and bakery and confectionery products.

14.3  Proanthocyanidins (Condensed Tannins) Although ubiquitous in nature, the proanthocyanidins are relatively rare in cereal grains. Apart from some genotypes of sorghum (Sorghum bicolor), finger millet (Eleusine coracana), barley (Hordeum vulgare) and rice (Oryza sativa), most other cereal grains contain none or inconsequential amounts of these compounds (Table 14.2). Proanthocyanidins are believed to be largely responsible for the red pericarp pigmentation in red wheat (T. aestivum). However, these compounds appear to be tightly bound and are difficult to extract and therefore they have not been well characterized in mature seeds. It thus remains largely unknown which types of proanthocyanidins are present in wheat, their levels and their possible effects on nutrition and health. Even among the cereal grains with significant amounts of detectable tannins, data on their molecular weight profile or structural properties are scarce, with the exception of sorghum, which has been extensively studied. In general, however, cereal grains appear to contain only condensed tannins, with no tannin acid (hydrolysable tannins) detected in any cereal grain. Table 14.2  Cereal  grains with confirmed presence of proanthocyanidins.a Commodity (phenotype)

Estimated content (µg g−1, db) Major anthocyanins present

Sorghum (brown)

890–33 000

Finger millet (brown) Barley

1400–1800b 150–1600

Rice (red and black)

190–325

Wheat (red)

Unknown

a

Mostly high molecular weight B-type proanthocyanidins, mean degree of polymerization 20 Molecular weight profile unknown Heteropolymeric, molecular weight profile largely unknown Procyanidins with A- and B-type linkages, up to decamers reported Positively identified in immature red wheat, unextractable in mature wheat

Proanthocyanidins only present in specific genotypes of these grains. Incomplete quantitative data available.

b

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The most common substituents of the proanthocyanidins are the flavan3-ols, a subclass of flavonoids, with catechin and gallocatechin and their isomers and derivatives (e.g. epicatechin, epigallocatechin gallate) being the most well known. In some plants (e.g. tea and various legumes such as cowpeas), the flavan-3-ols mainly exist as monomers or low molecular weight condensed polymers. However, in cereal grains, the monomeric forms tend to be trivial and the vast majority of these compounds exist as high molecular weight condensed polymers—in sorghum, the mean degree of polymerization is about 20.47 This size is important to nutrition and health because the high molecular weight tannins are known to produce the most significant consequences on the digestion of macronutrients. The proanthocyanidins are formed in the flavonoid biosynthetic pathway and generally polymerize late in the plant maturation cycle as carbo-cations of quinone methides and leucoanthocyanidin derivatives condense with catechin, epicatechin or other flavonoids.48 Depending on the enzymes involved in the biosynthetic pathway, a wide diversity of polymer substituents can exist in terms of interflavan linkages (Figure 14.7), hydroxyl substitution on the B-ring (Figure 14.8), glycosyl substitution and other factors. However, a mystery still remains as to the source of the carbo-cationic extension units of proanthocyanidins. Xie and Dixon49 speculated that polyphenol oxidase might be involved in converting flavan-3-ols into reactive quinoidal intermediates, which can be reduced to carbo-cations through coupled non-enzymatic oxidation. Thus, in addition to the common flavan-3-ols, other 3-deoxyflavans are also involved in the polymeric structure of cereal tannins. For example, glycosylated 3-deoxyflavan polymers (proluteolinidin and proapigeninidin) with flavanones (eriodictyol or naringenin) and their glycosides as the terminal

Figure 14.7  Examples  of proanthocyanidins with B-type C4 → C8 interflavan bonds (left) and A-type (right).

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Figure 14.8  Repeating  units of B-type proanthocyanidins with differences in B-ring hydroxylation.

units have been reported in sorghum and barley.50,51 Also, in addition to the classical 4 → 8 B-type flavan linkages, A-type interflavan linkages (Figure 14.7) have also been reported in sorghum,51 although they seem to be a relatively minor component. The overall body of evidence suggests significant heterogeneity in the structure of cereal grain tannins. In sorghum, in particular, the condensed tannins have been the most widely investigated group of flavonoids. The main reason that they have been such a subject of scientific curiosity is their observed negative impact on the feed value of sorghum. Tannins are well known to bind strongly to proteins, a property used for centuries to convert hide into leather. When sorghum containing significant amounts of tannins is cooked, the tannins complex with the proteins to restrict protein digestibility by as much as 70% or more.52 In addition, the tannins can also potentially, in theory at least, chelate micronutrients, specifically divalent minerals such as iron and zinc, reducing their bioaccessibility. Biological evidence in this regard is, however, equivocal. In the free form (not complexed with other food macronutrients), the tannins can also affect nutrition by directly complexing with and inhibiting digestive enzymes (themselves proteins). However, these effects are strongly dependent on the content of tannins in the grain or food matrix.52,53 For these reasons, most commercially produced sorghum in places such as the USA does not contain tannins. However, in some regions of Africa and South America, tannin sorghums are still produced as a matter of necessity. An important practical benefit of tannins is that they appear to significantly reduce bird pest predation of sorghum grain during maturation. When tannin and non-tannin sorghum are present in the same vicinity, the observed bird preference for the non-tannin sorghums is dramatic. This phenomenon clearly indicates that tannins serve an important protective role against predation. Consequently, in regions where bird pest pressure is a major problem, a significant portion of sorghum grown will be the tannin type. Growing interest in the potential health benefits conferred by the tannins to humans may see a deliberate resurgence in the commercial production of tannin sorghum.

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14.4  H  ealth Benefits of Cereal Anthocyanins, 3-deoxyanthocyanins and Proanthocyanidins Cereal grains are generally low in flavonoid compounds and, consequently, the health benefits associated with their phenolic compounds have often largely been attributed to their high phenolic acid content. However, in cereal grains that contain anthocyanins, 3-deoxyanthocyanins and tannins, these flavonoids constitute a significant proportion, often most, of the grain phenolic compounds. Thus they are expected to play a prominent part in contributing to the health benefits of the intake of these whole grains. Some of the cereal flavonoids, as a result of their structure, can produce significant bioactive properties at low levels, probably due to specific and efficient interactions with relevant target biological receptors.35,54–56 Thus the flavonoid composition of grains is highly relevant to their contribution to health.

14.4.1  Anthocyanins and 3-deoxyanthocyanins Anthocyanins are relatively common in fruits and beverages (e.g. red wine) and thus have been the subject of considerable investigations for their bioavailability, pharmacokinetics and bioactive properties. Even though direct evidence on the biological effects of the consumption of anthocyanin-rich cereals is scarce, the composition of anthocyanins in the cereals is similar to those from well-studied fruits and vegetables, thus the evidence from the latter provides useful insights. In general, an intake of anthocyanin-rich foods has been associated with beneficial effects against fat accumulation (obesity), diabetes and cardiovascular disease.57 However, because anthocyanin-rich foods are also commonly rich in other health-promoting compounds, isolating the effect of anthocyanins can often be tricky. Similar to evidence from fruit-derived anthocyanins, cereal anthocyanins have been shown to provide specific health benefits. For example, an anthocyanin-rich extract from purple maize was shown to suppress weight gain induced by a high-fat diet, as well as insulin resistance in mice, partially through action against the enzymes involved in fatty acid and triacylglycerol synthesis.58 The diet was formulated to provide 2 g of cyanidin-3-O-glucoside (C3G)/kg diet and thus provided a good basis for comparing related studies on anthocyanins. The C3G was the dominant anthocyanin in the purple maize extract and is also the dominant anthocyanin in pigmented wheat and rice (Table 14.1), which makes the evidence particularly relevant. A related study using black rice showed that the black rice anthocyanins reduced weight gain and platelet hyperactivity (which can promote platelet aggregation and adhesion) in dyslipidemic rats induced with a high-fat diet.59 Anthocyanins derived from cereal grains are also known to improve cardiovascular health. For example, a controlled study that fed rats isogenic diets of anthocyanin-rich or anthocyanin-free extracts from maize showed

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that the anthocyanin-rich extract significantly protected the heart of rats from ischemia.60 Using an ex vivo model of isolated perfused rat heart, it was further demonstrated that the anthocyanins could induce a state of myocardial resistance through reduced infarct size following regional ischemia and perfusion.60 The cardioprotection was associated with increased myocardial glutathione levels and it was suggested that this implied that the anthocyanins might act by modulating endogenous cardiac antioxidant defenses. It was also found that the maize anthocyanins were absorbed as intact glycosides and acyl-glycosides. This type of study is important because it provides compelling evidence for the direct contribution of dietary anthocyanins to disease prevention. The cardioprotective effects of rice anthocyanins have also been demonstrated.61 Anthocyanins derived from black rice were shown to protect mouse liver from injury induced by carbon tetrachloride by alleviating the release of aminotransferases from hepatocytes and reversing the imbalance of redox homeostasis.62 Using pure anthocyanins, it was shown that C3G, the dominant anthocyanin in black rice, was more active than peonidin-3-O-glucoside (Pn3G), the second most dominant anthocyanin in the rice extract. It is likely that the antioxidant-related mechanism of protection is related to the ability of the anthocyanins to induce endogenous antioxidant system, as reported by Toufektsian et al.60 The evidence further shows the beneficial effects of the dominant anthocyanins in cereal grains. Evidence for the health benefits of the sorghum-derived 3-deoxyanthocyanins is scarce. As a result of their important structural difference from anthocyanins, their biological activity is expected to be different. For example, sorghum-derived 3-deoxyanthocyanidins were found to be strong phase II enzyme inducers, a property that has not been reported for their anthocyanin analogs.35,63 The structure of the 3-deoxyanthocyanins had a major impact on their ability to influence the phase II enzymes, with O-methyl substitution being essential for their activity. Phase II enzymes are important in metabolism and the detoxifying of xenobiotics relevant to cancer prevention. A comparative study of the dominant sorghum 3-deoxyanthocyanidins (luteolinidin and apigeninidin) with their anthocyanidin analogs (cyanidin and pelargonidin) showed that the 3-deoxyanthocyanidins were significantly more effective against HL-60 leukemia and HepG2 cancer cell proliferation in vitro.36 Thus the structural differences may confer unique bioactive properties to 3-deoxyanthocyanins compared with their anthocyanin analogs. Stud­ ies that target the specific biological properties of the 3-deoxyanthocyanins are warranted.

14.4.2  Proanthocyanidins (Condensed Tannins) Condensed tannins have been widely investigated for their potential health benefits due to their widespread intake from fruits and vegetables. These compounds have been associated with a myriad of potential health benefits, including direct antioxidant function, anti-inflammatory properties,

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chemoprevention and protection against cardiovascular disease. The proanthocyanidins can produce their impact on health in three main ways: direct bioactive function; interactions with gut microflora; and interactions with macronutrients. Proanthocyanidin monomers and, to a lesser extent, dimers and trimers, are absorbed in vivo, evidenced by, for example, the presence of flavan-3ols and their glucuronidated, acylated or sulfonated forms in the urine or plasma of rats after the consumption of a proanthocyanidin-rich diet.68 Proanthocyanidins with a degree of polymerization >3 generally survive digestion and reach the colon, where they are fermented by various bacteria into low molecular weight phenolic acids, including hydroxylated phenylacetic, phenylpropionic and phenylvaleric acids. These molecules can then be absorbed and undergo phase II metabolism. For example, benzoic, phenylacetic and phenylproprionic acid metabolites were found in the serum and urine of rats fed sorghum tannins.69 Similarly, phenylpropionic, phenylacetic and benzoic acids and valerolactone derivatives were found in the plasma of rats fed grape seed tannins.68 Even though evidence on the direct bioactive properties of tannins specific to cereal grains is limited, studies suggest that, when present, the tannins may confer unique benefits to cereal grains. For example, a study comparing tannin-containing sorghum with those without tannins showed that the presence of tannins conferred stronger protection against the proliferation of colon cancer cells.63 Sorghum tannins were found to act as strong antioxidants in the gastrointestinal tract even when complexed to food macronutrients and were thus proposed as effective biological free radical sinks.65 Because of their poor absorption in the upper gastrointestinal tract, tannins are far more likely to directly exert bioactivity through their action in the colon and interaction with colon bacteria. For example, Gu et al.69 reported that microbial-derived phenolic acid metabolites of tannins were the predominant phenolic compounds in the systemic circulation of rats after the consumption of tannin sorghum bran. This finding suggests significant action on the tannins by colon bacteria and thus the likely significant interaction of the tannins and their microbial metabolites with the colon epithelium. A study using extruded tannin sorghum showed that when cellulose fiber and starch were substituted with the tannin sorghum, a significant reduction in lipogenesis and epididymal adipose tissue was observed in obese rats fed a high-fat diet.64 The proinflammatory cytokine TNF-α was significantly decreased, whereas the anti-inflammatory cytokine interleukin-10 was significantly increased in the sorghum-fed rats.64 This result suggests that the tannins in sorghum may contribute to weight management in obese patients via their action against lipogenesis and chronic inflammation. The tannins and their phenolic acid metabolites can also exert a modulating effect on the microbiota in the colon. Various studies have shown the selective effect of tannins from dietary sources on the population and

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activity of the colon microbiota. Monomeric flavan-3-ols incubated with fecal bacteria generated valeroactones and phenylpropionic acid and led to an increase in the Clostridium coccoides–Eubacterium rectale group, Bifidobacterium spp. and Escherichia coli, while decreasing Clostridium hystolyticum.70 When proanthocyanidins with a degree of polymerization of 2–3 were added to cultured fecal bacteria, Lactobaccillus spp. and Enterococcus spp. increased and C. hystolyticum decreased.71 Human consumption of high molecular weight cocoa proanthocyanidins for four weeks increased Bifidobacterium spp., Lactobaccillus spp. and Enterococcus spp. and decreased Clostridium spp. in human fecal samples.72 Broadly, Bifidobacterium spp. and Lactobaccillus spp. have health-promoting functions, whereas Clostridium spp. has deleterious effects. Thus it can be concluded that, to some extent, dietary proanthocyanidins can exert prebiotic benefits, thus producing beneficial effects beyond what can be predicted from their direct activity. Sorghum tannins also present an interesting opportunity to modify starch digestion and the calorie impact of foods in general.5 Starch is a major contributor to the calorie intake of humans and is the major component of cereal grains. With the prominent role obesity currently has in chronic diseases globally, strategies to reduce the caloric impact of foods based on starch and cereal grains is likely to produce important impacts on global nutritional health. In addition to the known cross-liking with proteins, evidence indicates that sorghum tannins can complex with starches and make them less digestible or completely non-digestible.2,73–75 As with proteins, the binding of tannins with starch appears to be specific and thus could be manipulated to produce desired properties in starch. High molecular weight fractions of sorghum tannins were found to most effectively complex with starch, whereas monomeric flavonoids such as catechin do not bind with starch,2,76 similar to what is observed with proteins. The amylose component of starch interacts more efficiently with tannins than amylopectin, further suggesting the specificity of starch–tannin complexes. Starch–tannin binding mechanisms appear to involve mainly hydrogen bonding, stabilized by hydrophobic interactions, similar to the mechanisms observed for proteins.2 Thus the reaction conditions can be manipulated to produce a wide range of starch digestion profiles, from starches high in slowly digestible starch to those that contain only resistant starch.73 For example, depending on the degree of starch gelatinization, the tannins can affect the starch digestibility profile in different ways. For instance, in vitro slowly digestiible starch increased two-fold (3×) from 97 to 274 mg g−1 in normal maize starch when tannins were reacted with partially gelatinized starch in a high moisture environment.73 In the same experiment, rapidly digesting starch was reduced by 46%. When starch gelatinization was inhibited during the reaction, however, subsequent starch digestion was almost completely blocked (>90% resistant starch). Thus the tannins provide a new opportunity to ‘naturally’ modify starch to benefit nutrition.

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The beneficial effect of tannins on starch digestion was found to be retained, to some extent, in a food matrix that contained proteins, known to have a much higher affinity for the tannins. Dunn et al.74 found that slowly digestible starch was significantly increased, whereas rapidly digestible starch was decreased in tortillas fortified with tannin sorghum bran relative to those made with non-tannin sorghum or wheat bran. This type of evidence is highly relevant to food processing and human health because it indicates that sorghum tannins can be feasibly used as ingredients to naturally make starch less nutritionally available in food processing. Additional evidence suggests that the tannin interaction with proteins may impart favorable quality attributes as food processing ingredients. For example, Girard et al.47,77 showed that condensed tannins from sorghum interact strongly with wheat gluten proteins to dramatically increase gluten strength and resilience. These interactions primarily involve high molecular weight tannins (degree of polymerization >3) and the high molecular weight glutenin subfractions of gluten proteins. It was also shown that the higher the molecular weight of the tannins, the stronger the interactions with sorghum tannins (mean degree of polymerization 19.5), producing much stronger effects than grapeseed tannins (mean degree of polymerization 8.3). A monomeric flavan-3-ol (catechin) had the opposite effect of weakening gluten due to the expected antioxidant effect. Limited evidence also suggests the sorghum tannins may benefit the blood glucose response in vivo, either through anti-inflammatory function or through interactions with digestive enzymes or starch. Arbex et al.64 reported a significant improvement in post-prandial hyperglycemia in obese mice fed a high-fat diet containing extruded tannin sorghum compared with control mice fed a high-fat diet containing cellulose and starch. A human trial (N = 10) showed that the consumption of a drink based on tannin sorghum 30 minutes prior to the ingestion of a 25 g glucose solution significantly decreased the glycemic response compared with non-tannin sorghum drinks or controls.78 This suggests that the tannins in sorghum may be acting by partially inhibiting the action of membrane glucose transporters. Thus the tannins could be a valuable tool in treating patients with type 2 diabetes and in weight management via various mechanisms.

References 1. J. M. Awika, L. W. Rooney and R. D. Waniska, Food Chem., 2005, 90, 293–301. 2. D. Amoako and J. M. Awika, Curr. Opin. Food Sci., 2016, 8, 14–18. 3. Q. Zhu, S. Yu, D. Zeng, H. Liu, H. Wang, Z. Yang, X. Xie, R. Shen, J. Tan, H. Li, X. Zhao, Q. Zhang, Y. Chen, J. Guo, L. Chen and Y.-G. Liu, Mol. Plant, 2017, 10, 918–929.

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4. C. Petti, R. Kushwaha, M. Tateno, A. E. Harman-Ware, M. Crocker, J. Awika and S. DeBolt, J. Agric. Food Chem., 2014, 62, 1227–1232. 5. J. M. Awika, in Gluten-free Ancient Grains, ed. J. R. N. Taylor and J. Awika, Woodhead Publishing, 2017, ch. 3, pp. 21–54, DOI: 10.1016/ B978-0-08-100866-9.00003-0. 6. J. M. Awika, C. M. McDonough and L. W. Rooney, J. Agric. Food Chem., 2005, 53, 6230–6234. 7. J. B. Harborne, Comparative biochemistry of flavonoids–II: 3-desoxyanthocyanins and their systematic distribution in ferns and gesnerads, Phytochemistry, 1966, 5, 589–600. 8. E. Grotewold, B. J. Drummond, B. Bowen and T. Peterson, Cell, 1994, 76, 543–553. 9. J. B. Harborne, Phytochemistry, 1967, 6, 1643–1651. 10. T. A. Holton and E. C. Cornish, Plant Cell, 1995, 7, 1071–1083. 11. K. Stich and G. Forkmann, Phytochemistry, 1988, 27, 785–789. 12. S.-C. C. Lo, K. De Verdier and R. L. Nicholson, Physiol. Mol. Plant Pathol., 1999, 55, 263–273. 13. C. S. Winefield, D. H. Lewis, E. E. Swinny, H. Zhang, H. S. Arathoon, T. C. Fischer, H. Halbwirth, K. Stich, C. Gosch, G. Forkmann and K. M. Davies, Physiol. Plant., 2005, 124, 419–430. 14. K. Stich and G. Forkmann, Z. Naturforsch., C: J. Biosci., 1988, 43, 311–314. 15. H. Halbwirth, S. Martens, U. Wienand, G. Forkmann and K. Stich, Plant Sci., 2003, 164, 489–495. 16. C.-H. Shih, I. K. Chu, W. K. Yip and C. Lo, Plant Cell Physiol., 2006, 47, 1412–1419. 17. H. Liu, Y. Du, H. Chu, C. H. Shih, Y. W. Wong, M. Wang, I. K. Chu, Y. Tao and C. Lo, Plant Cell Physiol., 2010, 51, 1173–1185. 18. E.-S. M. Abdel-Aal, J. C. Young and I. Rabalski, J. Agric. Food Chem., 2006, 54, 4696–4704. 19. A. Collison, L. Yang, L. Dykes, S. Murray and J. M. Awika, J. Agric. Food Chem., 2015, 63, 5528–5538. 20. M. Paulsmeyer, L. Chatham, T. Becker, M. West, L. West and J. Juvik, J. Agric. Food Chem., 2017, 65, 4341–4350. 21. K. Petroni, R. Pilu and C. Tonelli, Planta, 2014, 240, 901–911. 22. T. Nakayama, H. Suzuki and T. Nishino, J. Mol. Catal. B: Enzym., 2003, 23, 117–132. 23. G. Pereira-Caro, G. Cros, T. Yokota and A. Crozier, J. Agric. Food Chem., 2013, 61, 7976–7986. 24. D. B. M. Ficco, V. De Simone, S. A. Colecchia, I. Pecorella, C. Platani, F. Nigro, F. Finocchiaro, R. Papa and P. De Vita, J. Agric. Food Chem., 2014, 62, 8686–8695. 25. A. Brandolini, A. Hidalgo, S. Gabriele and M. Heun, J. Cereal Sci., 2015, 63, 122–127. 26. F. S. Hosseinian, W. Li and T. Beta, Food Chem., 2008, 109, 916–924. 27. I. Diczházi and L. Kursinszki, Cereal Chem., 2014, 91, 195–200.

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28. D. C. Knievel, E. S. M. Abdel-Aal, I. Rabalski, T. Nakamura and P. Hucl, J. Cereal Sci., 2009, 50, 113–120. 29. G. G. Bellido and T. Beta, J. Agric. Food Chem., 2009, 57, 1022–1028. 30. J. M. Awika, Food Res. Int., 2008, 41, 532–538. 31. B. Geera, L. O. Ojwang and J. M. Awika, J. Food Sci., 2012, 77, C566–C572. 32. L. Ojwang and J. M. Awika, J. Sci. Food Agric., 2008, 88, 1987–1996. 33. L. O. Ojwang and J. M. Awika, J. Agric. Food Chem., 2010, 58, 9077–9082. 34. L. Y. Yang, L. Dykes and J. M. Awika, Food Chem., 2014, 160, 246–254. 35. L. Y. Yang, J. D. Browning and J. M. Awika, J. Agric. Food Chem., 2009, 57, 1797–1804. 36. C. H. Shih, S. O. Siu, R. Ng, E. Wong, L. C. M. Chiu, I. K. Chu and C. Lo, J. Agric. Food Chem., 2007, 55, 254–259. 37. J. Boddu, C. Svabek, F. Ibraheem, A. D. Jones and S. Chopra, Plant Sci., 2005, 169, 542–552. 38. W. L. Rooney, in Sorghum: Origin, History, Technology, and Production, ed. C. W. Smith and R. A. Frederiksen, John Wiley and Sons, New York, 1st edn, 2000, pp. 261–307. 39. L. Dykes, L. M. Seitz, W. L. Rooney and L. W. Rooney, Food Chem., 2009, 116, 313–317. 40. J. D. Hipskind, R. Hanau, B. Leite and R. L. Nicholson, Physiol. Mol. Plant Pathol., 1990, 36, 381–396. 41. S.-C. C. Lo and R. L. Nicholson, Plant Physiol., 1998, 116, 979–989. 42. A. P. P. Kayodé, M. J. R. Nout, A. R. Linnemann, J. D. Hounhouigan, E. Berghofer and S. Siebenhandl-Ehn, J. Agric. Food Chem., 2011, 59, 1178–1184. 43. J. M. Awika, L. W. Rooney and R. D. Waniska, J. Agric. Food Chem., 2004, 52, 4388–4394. 44. J. D. Hipskind, R. Hanau, B. Leite and R. L. Nicholson, Physiol. Mol. Plant Pathol., 1990, 36, 381–396. 45. A. Khalil, R. Baltenweck-Guyot, R. Ocampo-Torres and P. Albrecht, Phytochem. Lett., 2010, 3, 93–95. 46. F. Barros, L. Dykes, J. M. Awika and L. W. Rooney, J. Cereal Sci., 2013, 58, 305–312. 47. A. L. Girard, S. R. Bean, M. Tilley, S. L. Adrianos and J. M. Awika, Food Chem., 2018, 245, 1154–1162. 48. J. Zhao, Y. Pang and R. A. Dixon, Plant Physiol., 2010, 153, 437–443. 49. D.-Y. Xie and R. A. Dixon, Phytochemistry, 2005, 66, 2127–2144. 50. M. J. Brandon, L. Y. Foo, L. J. Porter and P. Meredith, Phytochemistry, 1980, 21, 2953–2957. 51. C. G. Krueger, M. M. Vestling and J. D. Reed, J. Agric. Food Chem., 2003, 51, 538–543. 52. B. N. Mitaru, R. D. Reichert and R. Blair, J. Nutr., 1984, 114, 1787–1796. 53. J. M. Awika and L. W. Rooney, Phytochemistry, 2004, 65, 1199–1221. 54. S. Agah, H. Kim, S. U. Mertens-Talcott and J. M. Awika, Mol. Nutr. Food Res., 2017, 61, DOI: 10.1002/mnfr.201600625.

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55. J. M. Awika, in Advances in Cereal Science: Implications to Food Processing and Health Promotion, American Chemical Society, 2011, ch. 9, vol. 1089, pp. 171–200. 56. L. Yang, K. F. Allred, B. Geera, C. D. Allred and J. M. Awika, Nutr. Cancer, 2012, 64, 419–427. 57. T. Tsuda, Mol. Nutr. Food Res., 2012, 56, 159–170. 58. T. Tsuda, F. Horio, K. Uchida, H. Aoki and T. Osawa, J. Nutr., 2003, 133, 2125–2130. 59. Y. Yang, M. C. Andrews, Y. Hu, D. Wang, Y. Qin, Y. Zhu, H. Ni and W. Ling, J. Agric. Food Chem., 2011, 59, 6759–6764. 60. M.-C. Toufektsian, M. de Lorgeril, N. Nagy, P. Salen, M. B. Donati, L. Giordano, H.-P. Mock, S. Peterek, A. Matros, K. Petroni, R. Pilu, D. Rotilio, C. Tonelli, J. de Leiris, F. Boucher and C. Martin, J. Nutr., 2008, 138, 747–752. 61. M. Xia, W. H. Ling, J. Ma, D. D. Kitts and J. Zawistowski, J. Nutr., 2003, 133, 744–751. 62. F. Hou, R. Zhang, M. Zhang, D. Su, Z. Wei, Y. Deng, Y. Zhang, J. Chi and X. Tang, J. Funct. Foods, 2013, 5, 1705–1713. 63. J. M. Awika, L. Y. Yang, J. D. Browning and A. Faraj, LWT Food Sci. Technol., 2009, 42, 1041–1046. 64. P. M. Arbex, M. E. d. C. Moreira, R. C. L. Toledo, L. de Morais Cardoso, H. M. Pinheiro-Sant'ana, L. d. A. Benjamin, L. Licursi, C. W. P. Carvalho, V. A. V. Queiroz and H. S. D. Martino, J. Funct. Foods, 2018, 42, 346–355. 65. K. M. Riedl and A. E. Hagerman, J. Agric. Food Chem., 2001, 49, 4917–4923. 66. A. E. Hagerman, K. M. Riedl, G. A. Jones, K. N. Sovik, N. T. Ritchard, P. W. Hartzfeld and T. L. Riechel, J. Agric. Food Chem., 1998, 46, 1887–1892. 67. J. M. Awika and K. G. Duodu, J. Funct. Foods, 2017, 38, 686–697. 68. M. Margalef, Z. Pons, F. I. Bravo, B. Muguerza and A. Arola-Arnal, J. Funct. Foods, 2015, 12, 478–488. 69. L. Gu, S. E. House, L. Rooney and R. L. Prior, J. Agric. Food Chem., 2007, 55, 5326–5334. 70. X. Tzounis, J. Vulevic, G. G. Kuhnle, T. George, J. Leonczak, G. R. Gibson, C. Kwik-Uribe and J. P. Spencer, Br. J. Nutr., 2008, 99, 782–792. 71. C. Cueva, F. Sánchez-Patán, M. Monagas, G. E. Walton, G. R. Gibson, P. J. Martín-Álvarez, B. Bartolomé and M. V. Moreno-Arribas, FEMS Microbiol. Ecol., 2013, 83, 792–805. 72. X. Tzounis, A. Rodriguez-Mateos, J. Vulevic, G. R. Gibson, C. Kwik-Uribe and J. P. E. Spencer, Am. J. Clin. Nutr., 2011, 93, 62–72. 73. D. B. Amoako and J. M. Awika, Food Chem., 2016, 208, 10–17. 74. K. L. Dunn, L. Yang, A. Girard, S. Bean and J. M. Awika, J. Agric. Food Chem., 2015, 63, 1234–1241. 75. F. Barros, J. M. Awika and L. W. Rooney, J. Agric. Food Chem., 2012, 60, 11609–11617. 76. F. Barros, J. M. Awika and L. W. Rooney, J. Sci. Food Agric., 2014, 94, 1212–1217.

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77. A. L. Girard, M. E. Castell-Perez, S. R. Bean, S. L. Adrianos and J. M. Awika, J. Agric. Food Chem., 2016, 64, 7348–7356. 78. P. C. Anunciação, L. d. M. Cardoso, V. A. V. Queiroz, C. B. de Menezes, C. W. P. de Carvalho, H. M. Pinheiro-Sant'Ana and R. d. C. G. Alfenas, Eur. J. Nutr., 2018, 57, 251–257. 79. G. Mazza and R. Brouillard, J. Agric. Food Chem., 1987, 35, 422–426.

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Chapter 15

Interactions Between Grains and the Microbiome S. Brahmaa and D. J. Rose*a,b,c a

University of Nebraska-Lincoln, Department of Food Science & Technology, 1901 N. 21st St., Lincoln, NE 68588, USA; bUniversity of Nebraska-Lincoln, Department of Agronomy & Horticulture, 202 Keim Hall, Lincoln, NE 68583, USA; cUniversity of Nebraska-Lincoln, Nebraska Food for Health Center, 1901 N. 21st St., Lincoln, NE 68588, USA *E-mail: [email protected]

15.1  Introduction The human gastrointestinal tract is one of the largest interfaces in the human body between the host and the environment. The microbes that colonize the gastrointestinal tract are termed the gut microbiota and these microorganisms have evolved with the host to form an intricate and mutually beneficial relationship.1 The number of microorganisms populating the gastrointestinal tract has been estimated to be 1013 cells, equivalent to the number of human body cells.2 The adult human microbiota typically includes five dominant commensal phyla—Firmicutes, Bacteroidetes, Proteobacteria, Fusobacterium and Actinobacteria—of which Firmicutes and Bacteroidetes are present in the greatest abundance (>90%).1 The microbiota confers many benefits to the host, such as regulating immune function,3 harvesting energy4 and maintaining gut integrity.5 The potential disruption of the microbial   Food Chemistry, Function and Analysis No. 6 Cereal Grain-based Functional Foods: Carbohydrate and Phytochemical Components Edited by Trust Beta and Mary Ellen Camire © The Royal Society of Chemistry 2019 Published by the Royal Society of Chemistry, www.rsc.org

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composition may lead to dysbiosis, which can promote the development of metabolic diseases.6–8 Although the composition of the microbiota is relatively stable within an individual, both long- and short-term perturbations, such as dietary changes, have been reported to induce both structural and functional changes to the gut microbiota.9–11 Cereal grains are major source of dietary non-digestible food carbohydrates that are potentially available to be fermented by the gut microbiota in the large intestine. The human genome does not encode for enzymes that break down the complex carbohydrates (e.g. cellulose, arabinoxylan, β-glucan and fructans) that make up the dietary fibers in whole grains; however, bacteria are able to use these substrates for energy. Bacterial metabolism of these carbohydrates confers health benefits to the host—for example, by producing beneficial short chain fatty acids (SCFAs) such as acetate, propionate and butyrate.12 Whole grain cereals are also abundant in phytochemicals such as phenolic compounds, flavonoids and anthocyanins, which are considered to have significant health impacts in the prevention of chronic diseases. Several studies have documented the impact of whole grain foods and components of whole grains on human metabolic health and the gut microbiota.7,8,13–21 However, much is still unknown about how the specific components of whole grains interact with the gut microbiota and how they pertain to human health. Findings from whole grain intervention studies are not consistent with respect to shifts in the microbiota and the corresponding host benefits.13,17,19 Complicating matters further, some intervention trials suggest that the composition of the gut microbiota at enrollment into a study is predictive of host benefits in response to whole grains.22–24 Not all non-digestible components in whole grains are available for metabolism by the microbiota25,26 and therefore optimizing the processing methods and grain types to enhance the amount of carbohydrates available for gut microbial fermentation is an area ripe for research.27–33 The purpose of this review is to discuss whole grain– gut microbiota interactions and to identify new areas of research that may contribute to a better understanding of the underlying mechanisms linked to human health.

15.2  G  rain Components Likely to Interact with the Microbiome The most important whole grain components that are likely to interact with the microbiota are dietary fibers and polyphenols. Other non-digestible compounds (e.g. waxes, saponins, phytates, phytosterols, other lipophilic compounds and resistant proteins) may also interact with the gut microbiota,34 but much less is known about the impacts of these compounds on the gut microbiota. The carbohydrate and polyphenol compositions of whole grains vary among grains (Figure 15.1 and Table 15.1).

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Figure 15.1  Typical  non-digestible carbohydrate composition in selected whole grains. Data from ref. 35–38.

Table 15.1  Major  bioactive components of selected whole grains (% dry matter). Data from ref. 39–46.

Component Barley

Corn

Phytic acid Tocols Phenolic acids Phytosterols

Oats

Rice

0.04–0.11 0.16–0.17 0.04–0.12 0.45–0.8 0.005–0.01 0.38–0.47 0.002–0.004 0.4–0.9 0.03–0.07 0.06–0.09 0.04–0.09 Not reported 0.09–0.12 0.01–0.02 0.06–0.07 Not reported Alkylresorci­ 0.003–0.01 Not Not Not nols present present present Avenanthra- Not Not 0.004–0.01 Not mides present present present

Rye

Wheat

0.05–0.15 0.04–0.14 0.004–0.01 0.003–0.01 0.05–0.11 0.03–0.12 0.11–0.14 0.07–0.09 0.08–0.12 0.02–0.07 Not Not present present

15.2.1  Dietary Fiber The concentration of dietary fiber in whole grains depends on many factors and typically ranges from as little as 4% in brown rice to as much as 16% in rye (Figure 15.1). The main dietary fiber components in whole grains are non-starch polysaccharides, which can be classified into poorly fermentable (by the gut microbiota) (e.g. cellulose and water-unextractable arabinoxylans) and readily fermentable (e.g. mixed-linkage β-glucans and water-extractable arabinoxylans).47 Compositional and structural descriptions of the major dietary components that escape digestion in the human small intestine are outlined in the following subsections.

15.2.1.1 Arabinoxylans Arabinoxylans are the major dietary fiber components in grains, comprising roughly 50% of the dietary fiber in all whole grains except for oats and barley, which contain about 30% dietary fiber as arabinoxylans (Figure 15.1).

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These polysaccharides are composed of a linear backbone of β-d-xylopyranosyl (Xylp) residues linked through (1-4) glycosidic bonds. The backbone can contain α-l-arabinofuranosyl (Araf) substitutions at the O-3 and/or O-2 positions on the Xylp residues.48 Some Araf residues contain an ester-linked ferulic acid moiety (see Section 15.2.2) at O-5, which can form oxidative cross-linkages with other arabinoxylan chains and other components of the cell wall.49 Oligosaccharide branches consisting of glucose, arabinose and xylose are also common, as are glucuronic acid residues. Arabinoxylans can be categorized as water-extractable or water-unextractable. Water-extractable arabinoxylans dissolve in aqueous solutions and are present in far lower concentrations than the insoluble water-unextractable arabinoxylans.48 The water-extractable arabinoxylans can be considered “precursors” to the water-unextractable arabinoxylans, which act as the “glue” that holds the plant cell wall together through phenolic cross-linkages and non-covalent bonds. Because water-unextractable arabinoxylans are made unextractable in large part by ester-linked phenolic cross-linkages, a large portion of the water-unextractable arabinoxylans can be made soluble by treatment with alkali. The structure of arabinoxylans can vary among grain types. For example, wheat arabinoxylans contain more O-2 and O-2,3 substituted Xylp residues than rye, which contains more O-3 substituted Xylp residues.49 Rye also contains more unsubstituted Xylp residues more uniformly distributed along the xylan backbone, whereas wheat contains less Xylp residues, which tend to cluster in contiguous groups along the backbone.50 The structure of arabinoxylans also varies among different anatomical parts of the grain. For instance, when water-extractable arabinoxylans were analyzed from wheat bran and the starchy endosperm, the arabinose to xylose ratios (a measure of the degree of branching) as well as the concentrations of arabinoxylans were different in each fraction.48–50 Structural features associated with the degree of branching, molecular weight, the spatial arrangement of arabinoxylans and the ratio of arabinose to xylose in cereals influence their fermentability, which, in turn, could further affect the functionality of the gut microbiota.51,52 For instance, Rose et al.51 found that among the alkali-extracted fractions of corn bran, rice bran and wheat bran, corn arabinoxylans resulted in the highest production of SCFAs. Rice and corn arabinoxylans were hypothesized to degrade by a debranching mechanism due to their regular branching patterns, whereas wheat arabinoxylans were hypothesized to ferment in two stages due to the irregularity of the branches along the Xylp backbone, with the unsubstituted regions fermenting first, followed by the highly branched regions.51 In another study, no difference was reported in the fermentation rate patterns with respect to the molecular mass or arabinose to xylose ratio, although rice and sorghum arabinoxylans were shown to have a simple branched structure associated with more rapid fermentation than the wheat and corn arabinoxylans.52 Other studies have shown the impact of arabinoxylans on the modulation of the gut microbiota by promoting certain probiotic bacteria

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(e.g. Lactobacillus and Bifidobacterium) and enhancing the production of SCFAs.29,53,54 For instance, Damen et al.53 studied the impact in rats of arabinoxylan fractions isolated from wheat bran. The water-unextractable (40% purity), water-extractable (80% purity) and arabinoxylan oligosaccharide (79% purity) fractions, and their combinations, were included in a standardized diet at 5% arabinoxylan for 14 days. The ternary combination of water-extractable, water-unextractable and arabinoxylan oligosaccharides increased butyrate production in the colon, promoted a reduced pH, and limited proteolytic metabolites and Bifidobacterium growth in the colon relative to diets with only the individual arabinoxylan fractions. Truchado et al.54 studied the modulatory effects of two doses of water-extractable, long-chain arabinoxylans (3 and 6 g L−1) given three times per day for three days on the luminal and mucosal microbiota in a human intestinal microbial ecosystem (M-SHIME). They concluded that the higher dose stimulated Bifidobacterium and could be potentially beneficial to human host health. The fermentation of isolated arabinoxylan fractions is very different from that of arabinoxylan in whole grains due to the extensive cross-linkages present in native arabinoxylans.51,55,56 Cross-linking and other factors limit the availability of native arabinoxylans for microbial fermentation, although the extent of fermentation may be altered by various means, such as processing (see Section 15.5).

15.2.1.2 β-Glucans β-Glucans are non-digestible polysaccharides composed of mixed-linkage (1,3) and (1,4)-β-d glucose units with a molecular mass ranging between 50 and 2300 kDa and are present in the greatest amounts in oats and barley.57 The highest content of β-glucans has been reported for barley (2–20 g) and oats (3–8  g) [g (100 g dry weight)−1]. Other cereals (e.g. corn, wheat and rye) also contain β-glucan, but in lower concentrations.57 As with arabinoxylans, the structure and molecular features of β-glucan— such as the ratio of (1,3) to (1,4) linkages, the ratios of cellotriosyl to cellotetraosyl units (DP3/DP4) and the molecular weight—play significant parts in the viscosity, solubility, dispersibility and, consequently, physiological functions, including cholesterol-lowering and glucose-attenuating effects in the gastrointestinal tract.58–60 For instance, in a human feeding trial, Wang et al.60 showed the impact of following four β-glucan-based experimental diets for five weeks on the composition of the gut microbiota of patients with mild hypercholesterolemia. The experimental diets included: (1) a wheatand rice-based control; (2) 3 g day−1 low molecular weight barley β-glucan (288 kDa); (3) 5 g day−1 low molecular weight barley β-glucan (292 kDa); and (4) 3 g day−1 high molecular weight barley β-glucan (1349 kDa). Among the treatment groups, the 3 g day−1 high molecular weight barley β-glucan increased Bacteriodetes and decreased Firmicutes relative to the control diet.

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At the genus level, the high molecular weight barley β-glucan diet increased Bacteroides, decreased Dorea and tended to increase Prevotella. These genera were correlated with changes in the markers for cardiovascular disease. The low molecular weight barley β-glucan treatments did not induce any changes in the composition of the gut microbiota. Other studies have indicated the effectiveness of β-glucan in modulating the composition of the gut microbiota and increasing the production of SCFAs by the microbiota.17,19,61 For instance, Dong et al.17 studied how oat products modulated the gut microbiota and reduced obesity in rats. Rats were fed either a normal chow diet, a high-fat diet or a high-fat diet supplemented with oatmeal, oat flour or oat bran for eight weeks. The diets containing any of the oat products modulated the overall gut microbiota composition by increasing the Bacteriodetes to Firmicutes ratio. Acidobacteria was only detected in the group following treatment with oat products and this was more pronounced in the oat bran group. A significant increase in fecal SCFAs was also noted in the oat products groups relative to the control group. Increases in the abundance of Bacteroidetes and the Bacteriodetes to Firmicutes ratio were also found to be negatively correlated with markers of obesity, dyslipidaemia and inflammation. It was suggested that oat products help in controlling obesity and related metabolic disorders while regulating the composition of the gut microbiota in obese rats. By contrast, Martínez et al.19 reported that rolled whole grain barley (60 g day−1) caused a decrease in the Bacteriodetes to Firmicutes ratio, along with an increase in the abundance of the genus Blautia in a human feeding trial. The bacteria that changed in response to the whole grain barley treatment encode for β-glucanase genes that assist in utilizing the substrate during fermentation.

15.2.1.3 Cellulose Cellulose is an essential component of cereal cell walls and consists of linear chains of (1,4)-linked β-d-glucose units. As a result of the linear structure of β-glucan, cellulose is insoluble in water and can form three-dimensional microfibrillar aggregates that are resistant to digestion by microbial enzymes.62 Robert et al.63 showed that the ability of the microbiota to degrade microcrystalline cellulose was greatest in methanogenic subjects. Methanogenic bacteria from these subjects belonged mostly to the Ruminococcus genus, together with some Enterococcus. A few mouse studies have compared diets containing cellulose with those containing more fermentable fibers and have concluded that diets containing fermentable fibers are more important to gut health than cellulose.64,65 Native cellulose in plant cell walls behaves differently in the gut from purified cellulose.64 For instance, Van Soest65 compared the effects of controlled diets with the addition of cellulose from three sources (cabbage, wheat bran or purified) on the microbial ecology of the healthy volunteers. They reported not only the lowest fermentation for purified cellulose,

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but also that purified cellulose did not induce bacterial fermentation and depressed the breakdown of other cell wall polysaccharides from the diet. The fermentation of purified cellulose showed a lag of 17–20 h, much longer than cellulose from natural sources. Although cellulose was present in the diets along with other carbohydrate polymers, more research needs to be conducted to establish whether native cellulose has unique properties in the gastrointestinal tract.

15.2.1.4 Fructans Fructans are naturally occurring plant oligo- and polysaccharides built on the repeated fructosylation of sucrose.66 Among the grains, rye has the highest fructan levels, ranging from 3.3 to 6.6%, followed by wheat (Figure 15.1). Wheat contains fructans known as graminans, which contain both β-(2,1) and β-(2,6) fructosyl linkages in the same molecule and contain an internal glucose unit instead of a terminal glucose.66,67 Unfortunately, structural information on fructans from other grains is not currently available. Although some studies have shown the prebiotic potential of fructans,68,69 few studies have documented the impact of cereal fructans on gut health.70,71 Belobrajdic et al.70 observed similar SCFA concentrations in the cecum and colon digesta of rats fed diets containing oligofructose, wheat stem fructans or barley grain fructans at the 5% level. Although the number of Bifidobacteria in the cecum increased only for the oligofructose group, there was a significant decrease in the pH of the colonic digesta in the barley grain fructan group. Similar to this finding, another research group studied the impact of the chain length of fructans isolated from wheat stem and barley on the gut microbiota during in vitro fermentation and compared the findings with those for inulin and oligofructose.71 The graminan fructans produced comparable levels of total SCFAs to oligofructose and inulin, suggesting that fructans from such novel sources could have metabolic benefits.

15.2.1.5 Resistant Starch Starch can be divided into three categories: rapidly digestible starch; slowly digestible starch; and resistant starch (RS).72 RS is the fraction of starch that is relevant to gut health because it survives transit to the large intestine. RS can be classified into five categories: RS1 (physically inaccessible); RS2 (granular or native semi-crystalline); RS3 (retrograded or re-crystallized); RS4 (chemically modified); and RS5 (amylose–lipid complexes).73 Studies have shown the health benefits of RS on the composition of gut microbiota.74–76 Upadhyaya et al.75 fed 20 people with signs of metabolic syndrome RS4 (30% v/v in flour) or a control wheat flour for 12 weeks, each in a crossover design. The RS4 group had higher concentrations of fecal SCFAs, such as propionate and butyrate, together with a higher abundance of Bacteroides, Parabacteroides, Oscillospira, Blautia, Ruminococcus, Eubacterium and Christensenella. Significant correlations were reported between changes

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in the composition of the gut microbiota induced by RS4 and increased fecal SCFAs. For instance, acetate and butyrate levels were correlated with changes in Ruminococcus lactaris and Oscillospira species. The total SCFAs were correlated with changes in Methanobrevibacter species and R. lactaris and propionate and iso-butyrate were correlated with Methanobrevibacter species, Eubacterium dolichum, Christensenella minuta and R. lactaris. No significant correlation was noted between changes in the gut microbiota and SCFA production in the control flour intervention. Goldsmith et al.76 studied the impact of whole grain corn flour with RS on the gut microbiota in obese rats for 11 weeks. The study included four diet groups: normal corn starch; whole grain control flour (containing 6.9% RS); isolated RS-rich corn starch (25% RS); and whole grain corn flour (25% RS). The isolated RS-rich corn starch contributed to a higher Bacteroides to Firmicutes ratio than the other diet groups, whereas the high RS whole grain treatment induced higher SCFA production and a lower cecal content pH than isolated RS.

15.2.2  Phenolic Compounds Whole grains are good sources of phenolic compounds, which may act as antioxidants and have anti-inflammatory, antimicrobial and anticarcinogenic effects against degenerative diseases, such as heart disease and cancer.77 Phenolic compounds are secondary metabolites of plants that are involved in defense mechanisms against UV radiation or to protect the plant from pathogens.77 The total phenolic content in grains ranges from 0.04% in oats up to 0.4% in foxtail millets (Table 15.1). Brans have a higher percentage of phenolic compounds than their corresponding whole grains, ranging between 0.42 and 0.45%.78 All phenolic compounds have a phenolic ring and can be classified into different categories as a function of the number of phenol rings they contain and the structural elements that attach these rings to one another.79 Examples of the most common categories of phenolic compounds are phenolic acids, flavonoids, condensed tannins and alkyl resorcinols.79 Phenolic acids are derivatives of benzoic and cinnamic acids and are usually represented by two types: (1) hydrobenzoic acids, such as gallic, vanillic and syringic acids; and (2) hydrocinnamic acids with C6–C3 structures, such as coumaric, caffeic, ferulic and sinapic acids.78 Flavonoids have a typical C6–C3–C6 structure, consisting of two aromatic rings attached by a three-carbon linkage, and include flavonols, flavones, isoflavones, flavanones, anthocyanidins and flavanols (catechins and proanthocyanidins). Flavonoids are mostly found in sorghum, millet, barley, maize, rye, rice and wheat.43 Condensed tannins are polymerized flavanol units that can bind to proteins, carbohydrates and minerals; they are mostly found in grains such as sorghum, barley and red finger millets.43 Lignans are phytoestrogens. The two most commonly found plant lignans are secoisolariciresinol and matairesinol, which are present predominantly in cool seasonal cereal grains, such as barley, oats, rye, triticale and wheat.80 Alkylresorcinols are mostly present in the bran of wheat,

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rye, triticale and barley, but not in maize, oats, millet, rice or sorghum. They are 1,3-dihydroxybenzene derivatives with an odd-numbered n-alkyl sidechain at C-5 on the benzene ring.81 Phenolic acids are the most abundant antioxidants in whole grains and can be present in free and bound forms. The bound phenolic acids are mostly linked to arabinoxylan chains (see Section 15.2.1.1). Grains have higher bound phenolic acids than the free: about 85, 75 and 62% of the total phenolic acids present in corn, wheat and rice, respectively, are in insoluble bound forms.82 Some varieties of barley contain between 54 and 90% bound phenolic acids.83 The release and absorption of free phenolic acids from the food matrix occurs either by direct solubilization in the intestinal fluids under gastrointestinal conditions and/or by the action of digestive enzymes that hydrolyze macronutrients and favor the release of phenolic acids from the food matrix.84 Once absorbed, phenolic compounds may be subjected to biotransformation in the enterocytes and hepatocytes, generating water-soluble conjugate metabolites (e.g. methyl, glucuronide and sulfate derivatives), which are then distributed to the host tissues and ultimately excreted in urine.84 By contrast, the release of bound phenolic acids from the food matrix occurs to only a limited extent. Kroon et al.85 reported that gastric and small intestinal enzymatic treatment released 0.41 and 2.46 nmol of free ferulic acid, respectively, and 6.91 and 4.70 nmol of esterified ferulic acid, which, in total, accounted for only 2.6% of total feruloyl groups in the wheat bran fiber. The majority of bound phenolic acids traverse the small intestine intact, along with dietary fiber, and reach the colon, where they serve as substrates for the gut bacteria.86 Andreasen et al.87 compared the release of free diferulic acids (8-5-diferulic acid, 5-5-diferulic acid, 8-O-4-diferulic acid and 8-5-benzofuran diferulic acid) from wheat and rye bran by human fecal microbiota. They reported that the microbiota could release 36% of 8-5-diferulic acid, 4% of 5-5-diferulic acid, 4% of 8-O-4-diferulic acid and 7% of 8-5-benzofuran diferulic acid during fermentation of the wheat bran matrix. In rye bran, human fecal microbiota were unable to release any of the 8-5-diferulic acid or 5-5-diferulic acid and only small amounts of 8-O-4-diferulic acid and 8-5benzofuran diferulic acid (6 and 3%, respectively). However, the extent of release of bound phenolic acids from the matrix can be altered (usually increased) by non-thermal and thermal processing techniques, such as fermentation processes in food, germination, roasting, extrusion cooking and boiling (see Section 15.5). Once released by the gut bacteria, phenolic compounds are rapidly metabolized through hydrogenation, demethylation, dehydroxylation and decarboxylation. For instance, the first step in the fermentation of methyl ferulate in the colon by the human gut microbiota is the process of demethylation into ferulic acid, followed by several reactions that ultimately yield phenylpropionic acid.88 Only a few bacterial genera (e.g. Escherichia, Bifidobacterium, Lactobacillus, Bacteroides and Eubacterium) have been documented to be able to metabolize phenolic acids.84

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Covalently attached phenolic acids in grains or grain fractions can affect gut health.89,90 Duncan et al.89 reported that wheat bran promoted the enrichment of five key species of bacteria (Butyrivibrio fibrisolvens, Eubacterium rectale, Roseburia faecis and Roseburia intestinalis and Eubacterium siraeum), which were not only known butyrate producers, but also released ferulic acid and thus had a pivotal role in fermenting wheat bran. Yang et al.90 fed a low-fat diet, a high-fat diet and a high-fat diet supplemented with maizederived non-digestible feruloylated oligo- and polysaccharides to mice for eight weeks. Blooms were observed in the gut microbial genera Blautia and Akkermansia in three of the mice fed with feruloylated oligo- and polysaccharides and these shifts were found to be associated with decreased body and adipose tissue weights compared with the mice fed with the control high-fat diet, indicating that the changes observed could depend on the ability of an individual's microbiota to ferment feruloylated oligo- and polysaccharides. However, the underlying interactions between whole grain dietary fibers and the associated phenolic acids remain elusive. The specific effects of dietary phenols on the modulation of gut ecology remains vague and needs to be further investigated.

15.2.3  Other Compounds Whole grains also contain other combinations of minerals and phytochemicals depending on the type of cereal. In addition to phenolic compounds, other examples of phytochemicals are phytosterols and tocols (terpenes and terpenoids), betaine, folate, α- and β-carotene, lutein, β-cryptoxanthin and zeaxanthin and phytates.91 Phytosterols are steroid compounds present in plants and can be classified into sterols and stanols depending on the number of carbon side-chains and the presence or absence of double bonds. Sterols are unsaturated compounds with a double bond in the ring, whereas stanols are saturated compounds because they lack double bonds. Stanols represent only 10% of total dietary phytosterols; sitosterols and campesterols are the most abundant sterols found in plants and human diets.92 Phytic acid, also known as inositol hexaphosphate, is the storage form of phosphorus present in grains and cereals. The concentration of phytate in grains varies among cereals, ranging from 0.5% to 2.0%.93 Some studies have revealed the impact of phytosterols and phytic acid on the gut microbiota.94,95 For instance, Markiewicz et al.94 studied how diet shaped the ability of the microbiota to degrade phytate in an in vitro study using fecal samples from adults following conventional and vegetarian diets and breast-fed infants. Regardless of the diet group, the gram-positive anaerobes and lactobacilli had the lowest ability to degrade phytate, whereas coliforms and proteobacteria–bacteroides cultures showed the highest potential to degrade phytate to intermediate myo-inositol phosphates. It was concluded that a well-balanced cooperation of aerobic and anaerobic bacteria is essential to degrade phytate and a diet rich in phytate could enhance the potential of the microbiota to degrade phytate. Another study by Rasmussen

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et al. showed how plant sterol esters made with fatty acids from soybean oil, beef tallow or purified stearic acid could affect the absorption of cholesterol when fed to male hamsters for four weeks. A control group was also included where the hamsters were fed a diet devoid of sterol esters. The hamsters fed with purified stearic acid and plant sterol esters showed significantly lower cholesterol absorption and reduced concentrations of plasma nonhigh-density lipoprotein cholesterol and liver cholesterol, suggesting that cardioprotective benefits can be achieved by increasing the consumption of stearate-enriched plant sterol esters.

15.3  Whole Grain Intervention Studies Intervention studies have been conducted to understand the impact of whole grain consumption on markers of cardiovascular and metabolic health (Table 15.2). These studies have shown that the consumption of whole grains and their components have been associated with a lower body mass index (BMI), less adipose tissue, and lower levels of obesity, cardiovascular disease and type 2 diabetes, although the findings are not consistent. These studies have also shown differing effects on the composition of the gut microbiota, which could be due to differences in the study design and the types of whole grain foods used. Whole wheat and wheat bran breakfast cereals caused an increase in lactobacilli/enterococci and Bifidobacterium spp.16 Lappi et al.18 examined the differences in the composition of the gut microbiota after the intake of high fiber rye bread and low fiber wheat bread in Finnish adults. They reported a decrease in Bacteroidetes and an increase in Clostridium cluster IV, Collinsella and Atopobium spp. during the 12-week intervention. In another study, human subjects consumed a daily dose of whole grain barley, brown rice or an equal mixture of both whole grain barely and brown rice for 17 weeks in a randomized crossover design.19 There was a decrease in Bacteroidetes and an increase in Firmicutes. Vanegas et al.20 showed that replacing whole grains with refined grains in a six-week randomized trial using healthy human subjects had only a modest impact on the composition of the gut microbiota, accompanied by an increase in Lachnospiraceae, a decrease in Enterobacteriaceae and an increase in fecal acetate and total SCFAs.

15.4  R  esponders/Non-responders to Whole Grain Interventions The prebiotic literature discusses a phenomenon termed “responder/non-responder” and is based on how an individual's microbiota changes in response to dietary prebiotic interventions.22–24 This division among the individuals originated based on whether the microbiota remained stable (unchanged) during the intervention or whether the expected changes, such as an increase in Bifidobacterium, were evident after treatment with a prebiotic.19,22,23,113

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Characteristics of participants

Feeding trial

WG treatment

Study design Non-significant results Significant results

46 healthy adults; WG vs. RG; WG based Bread, rice, pasta, Six week paralsnacks, breakon estimated BMI 20–28; low lel-arm fast cereals, energy needs (e.g., whole grain tortilla, baking 13.7 g fiber day−1 consumers (