ISSN 0355-1180

UNIVERSITY OF HELSINKI

Department of Food and Nutrition

EKT Series 1860

DEGRADATION OF OAT LIPIDS: IMPACT ON THEFUNCTIONALITY OF β-GLUCAN RICH INGREDIENTS IN

ACID MILK GELS

Hanna Mahne

Helsinki 2018

ABSTRACTS

HELSINGIN YLIOPISTO ¾ HELSINGFORS UNIVERSITET ¾ UNIVERSITY OF HELSINKI

Tiedekunta/Osasto ¾ Fakultet/Sektion ¾ Faculty

Faculty of Agriculture and ForestryLaitos ¾ Institution ¾ Department

Department of Food and NutritionTekijä ¾ Författare ¾ Author

Hanna MahneTyön nimi ¾ Arbetets titel ¾ Title

Degradation of oat lipids: impact on the functionality of β-glucan rich ingredients in acidmilk gelsOppiaine ¾Läroämne ¾ Subject

Food ChemistryTyön laji ¾ Arbetets art ¾ Level

Master’s thesisAika ¾ Datum ¾ Month and year

October 2018Sivumäärä ¾ Sidoantal ¾ Number of pages

73Tiivistelmä ¾ Referat ¾ Abstract

The present literature was reviewed to understand the recent indications of lipid degradationaffecting β-glucan molecular weight in aqueous solutions, the main topics being oat lipidreactions, oat β-glucan, and oat bran/β-glucan fortified yogurts, as well as the possibleinterplay of these three. A non-heat treated, a heat treated, and a heat treated and defattedoat bran concentrate (OBC) were produced, all with ~20% β-glucan. OBCs were subjectedto a storage trial in dark for 0–12 weeks at 4, 22 and 40 °C. Lipid degradation was followedduring storage by assessing volatile compounds (HS-SPME-GC-MS), neutral lipid classes(NP-HPLC-ELSD) and tocopherol and tocotrienol levels (NP-HPLC-FLD) as well asβ-glucan molecular weights (HP-SEC). Acid milk gels were prepared as a model system foryogurts with added OBCs, and analysed for spontaneous and forced syneresis and texturalchanges. Lipid oxidation was observed in all OBCs, and hydrolysis in the non-heat treatedOBC, but no changes in β-glucan molecular weight were observed in the dry stored OBCs.OBC addition caused structural changes in an acid milk gels, the water retaining propertieswere improved, and the gels were softer and smoother than without added OBCs. Time andtemperature affected OBC lipid oxidation and hydrolysis behaviours. Adding OBCs to gelschanged gel textures compared to both without vs. with OBCs and adding fresh vs. storedOBCs.Avainsanat ¾ Nyckelord ¾ Keywords

Beta-glucan, oat bran concentrate, oat lipids, lipid degradation, yogurts, acid milk gelsSäilytyspaikka ¾ Förvaringsställe ¾ Where deposited

The Digital Repository of University of Helsinki, HeldaMuita tietoja ¾ Övriga uppgifter ¾ Further information

EKT Series 1860

HELSINGIN YLIOPISTO ¾ HELSINGFORS UNIVERSITET ¾ UNIVERSITY OF HELSINKI

Tiedekunta/Osasto ¾ Fakultet/Sektion ¾ Faculty

Maatalous-metsätieteellinen tiedekuntaLaitos ¾ Institution ¾ Department

Elintarvike- ja ravitsemustieteiden osastoTekijä ¾ Författare ¾ Author

Hanna MahneTyön nimi ¾ Arbetets titel ¾ Title

Kauran lipidien hajoamisen vaikutus β-glukaanipitoisten raaka-aineidenfunktionaalisuuteen hapanmaitogeeleissäOppiaine ¾Läroämne ¾ Subject

ElintarvikekemiaTyön laji ¾ Arbetets art ¾ Level

MaisterintutkielmaAika ¾ Datum ¾ Month and year

Lokakuu 2018Sivumäärä ¾ Sidoantal ¾ Number of pages

73

Tiivistelmä ¾ Referat ¾ Abstract

Kirjallisuuskatsaus rakennettiin työkaluksi ymmärtämään viimeaikaisia tutkimustuloksiarasvojen hajoamisen vaikutuksista β-glukaanin molekyylipainoon vesipitoisessamateriaalissa. Kirjallisuuskatsauksen pääaiheet kattoivat kauran lipidien reaktiot, kauranβ-glukaanin ja kaurakuiduilla rikastetut jogurttimallit. Kolme kauralesekonsentraattiavalmistettiin noin 20 %:n β-glukaanipitoisuuksilla: kuumennuskäsittelemätön, kuumennus-käsitelty ekä kuumennuskäsitelty ja rasvauutettu. Lesekonsentraatteja säilytettiin pimeässä0–12 viikkoa 4, 22 ja 40 °C:ssa. Lipidien hajoamista seurattiin määrittämällä muodostuvathaihtuvat yhdisteet (HS-SPME-GC-MS), neutraalilipidiluokat (NP-HPLC-ELSD),tokoferoli- ja tokotrienolipitoisuudet (NP-HPLC-FLD) sekä β-glukaanin molekyylipainot(HP-SEC). Kauralesekonsentraatteja käytettiin jogurtteja mallintavien hapanmaitogeelienainesosina. Geeleistä analysoitiin spontaani ja pakotettu synereesi sekä rakenteellisetmuutokset. Kaikissa kauralesekonsentraateissa havaittiin lipidien hapettumista jakuumennuskäsittelemättömässä kauranäytteessä lipidien hydrolysoitumista. β-glukaaninmolekyylipaino ei kuitenkaan muuttunut kuivissa kauralesekonsentraateissa.Kauralesekonsentraattien käyttäminen hapanmaitogeeleissä aiheutti rakenteellisiamuutoksia. Kauralesekonsentraattien lisäys paransi geelien veden sitomiskykyähuomattavasti ja muutti niiden rakennetta pehmeämmiksi ja sileämmiksi verrattuna ilmanlisättyä kuitua valmistetuttuihin kontrolligeeleihin. Aika ja lämpötila vaikuttivatkauralesekonsentraattien lipidien hapettumiseen ja hydrolysoitumiseen. Kauralese-konsentraattien lisääminen muutti geelien koostumusta ja rakennetta kontrolligeeleihinverrattuna. Erot geelien rakenteissa näkyi myös tuoreen ja säilötyn lesekonsentraatti-lisäysten välillä.Avainsanat ¾ Nyckelord ¾ Keywords

Beetaglukaani, kauralesekonsentraatti, kauran lipidit, lipidien hajoaminen, jogurtit,hapanmaitogeelitSäilytyspaikka ¾ Förvaringsställe ¾ Where deposited

Helsingin yliopiston digitaalinen arkisto, HeldaMuita tietoja ¾ Övriga uppgifter ¾ Further information

EKT-sarja 1860

PREFACE

This Master’s thesis was completed under the Business Finland funded project

OATyourGUT. The experimental work, writing, and instruction were performed as a

collaboration between VTT Technical Research Centre of Finland Ltd (VTT) and University

of Helsinki (UH), from December 2017 to early October 2018. The professor in charge was

Vieno Piironen from UH, and other instructors included Anna-Maija Lampi (UH), and

researchers Natalia Rosa-Sibakov and Martina Lille (VTT). A great big thank you to all my

instructors, for your flexibility and your expertise, we got the puzzle done! I am also grateful

to all the other team members at VTT and UH that helped and contributed to the work. Anni,

you know. Truly invaluable, lunches an all, #graduystävä. Lastly, my fambam and Jaakko.

Thank you for always believing in me, pushing me forward and helping me rest in between.

Hanna Mahne

Helsinki, 12.10.2018

LIST OF ABBREVIATIONS

DAG Diacylglycerol

ELSD Evaporative light scanning detector

FA Fatty acid

FFA Free fatty acid

FLD Fluorescence detector

GDL Glucono delta-lactone

H Non-heat treated OBC sample

HPLC High performance liquid chromatography

HS-SPME-GC-MS Headspace solid phase microextraction gas chromatographycoupled with mass spectrometry

L Heat treated, defatted OBC sample

M Heat treated OBC sample

MUFA Monounsaturated fatty acid

MW Molecular weight

OBC Oat bran concentrate

PUFA Polyunsaturated fatty acid

ROOH (Lipid) hydroperoxide

SC-CO2 Supercritical carbon dioxide

SFA Saturated fatty acid

TA Texture analyser

TAG Triacylglycerol

Tocols Tocopherols and tocotrienols

WHC Water holding capacity

TABLE OF CONTENTS

ABSTRACTS

PREFACE

LIST OF ABBREVIATIONS

1 INTRODUCTION

2 LITERATURE REVIEW 10

2.1 Oat bran, β-glucan, and its enrichment 102.1.1 Oat bran and β-glucan 102.1.2 Dry-fractionation for β-glucan enriched ingredients 13

2.2 Oat lipids 142.2.1 Lipid composition of oat bran 142.2.2 Antioxidants in oat bran 162.2.3 Lipid reactions in oats 18

Chemical oxidationEnzymatic hydrolysis and oxidationLipid reactions during oat processing and storageCo-oxidation of β-glucan

2.3 Yogurts 252.3.1 Milk proteins 252.3.2 Acid milk gels 272.3.3 Fibre fortified yogurts 29

3 EXPERIMENTAL RESEARCH 31

3.1 Background and aims 32

3.2 Overview of the storage trial and yogurt food model experiments 33

3.3 Test materials 343.3.1 Oat bran concentrates (OBCs) 343.3.2 Reagents and reference materials 35

3.4 Methods 363.4.1 Extraction of lipids and analysis of total FA content 363.4.2 Analysis of neutral lipid profiles with NP-HPLC-ELSD 383.4.3 Analysis of tocopherol and tocotrienol contents with NP-HPLC-FLD 393.4.4 Analysis of volatile compounds with HS-SPME-GC-MS 403.4.5 Preparation of acid milk gels 413.4.6 Analysis of spontaneous syneresis and water holding capacity 42

3.4.7 Analysis of gel firmness with TA 43

3.5 Results 433.5.1 OBC characteristics 433.5.2 Lipid degradation 44

Neutral lipidsTocopherols and tocotrienolsVolatile compounds

3.5.3 Properties of acid milk gels 51AcidificationSpontaneous syneresis and water holding capacity (WHC)Firmness of acid milk gels

3.6 Discussion 573.6.1 Lipid degradation behaviour of OBC samples 573.6.2 Effects of OBC addition on acid milk gel functionality 603.6.3 Limitations of study 62

4 CONCLUSIONS 64

REFERENCES 65

APPENDIX 73

Appendix 1. An exemplary graph for the analysis of gel firmness with TA 73

1 INTRODUCTION

It is widely recognized that a typical western diet does not provide enough dietary fibre.

According to the Nordic recommendations, 25–35 g, or approximately 3 g/MJ, of dietary

fibre should be consumed daily (NNR 2012; VRN 2014). Having insufficient fibre intake

coupled with the fact that the West is also plagued with cardiovascular disease and with other

diseases, for example many gastrointestinal disorders, that are all related to insufficient fibre

intake (see e.g. Makki et al. 2018), there seems to be a true need for nutraceuticals and easy-

to-use food products that conveniently add fibre to the Western diet. One interesting

ingredient used for functional foods is oats, and especially its fibre β-glucan.

Oats (Avena sativa) are a resilient crop still suiting Nordic growth conditions (Juhola et al.

2017). Oats have traditionally been used as high-energy feed material, however, the use of

oats as human food has been on a steady rise, especially in the Western world for the past

dozen years. According to the Natural Resources Institute Finland (2017), 92,000 tons of

oats was consumed as food in Finland during crop year 2016–2017, its highest amount for

oats ever recorded. Furthermore, of all cereals exported from Finland last crop year, oats had

the highest export volume of 326,000 tons (LUKE 2017). On their cereal’s balance sheet, it

is seen that a mere 9% of the harvested oats go toward food consumption and most are still

used for feed, indicating there is room to increase the food consumption portion.

The cell wall structures in oat aleurone and endosperm have 3–7% of β-glucan (Cui and

Wood 2000). β-glucan molecules are the main carbohydrates that form the familiar viscous

oat “slime” when preparing oat porridge, and they are also behind the healthy image of oats.

This healthy image has been scientifically supported, as oat β-glucan has three scientifically

proven, EFSA approved health claims: oat β-glucans attribute to (1) maintaining normal

blood cholesterol, (2) lowering blood cholesterol, and (3) reducing post-prandial glycaemic

responses (EFSA 2010, 2011). The beneficial effects are achieved when the daily

consumption of β-glucan reaches 3 grams.

Oats are typically considered as the fattiest cereal, with lipid contents ranging between 2 and

11% in oat grains, depending on the cultivar, growth environment, as well as extraction

method (Zhou et al. 1999a). Oat lipids contain a notable amount of nutritionally interesting

unsaturated fatty acids, such as oleic acid (C18:1) and linoleic acid (C18:2), each comprising

approximately 40% of the fatty acids in oats (Givens et al. 2004). When oats are processed

to flakes or flour, previously secured oat lipids are exposed to the environment, namely pro-

oxidants and oxygen, and it is likely for the lipid autoxidation chain reactions to start,

resulting in loss of lipids and formation of desired and un-desired oxidation products such

as free radicals or volatile compounds (Zhou et al. 1999b). In addition to oxidative reactions,

due to endogenous enzymes, oat lipids also degrade by hydrolysis (Yang et al. 2017). This

means that the possible lipid degradation changes the lipid composition and diminishes the

nutritional status of the oats, whilst forming undesired off-flavours and unpleasant aromas,

to begin with.

The effects of lipid degradation seem to also cause collateral damage, as discovered by Wang

et al. (2016): an oil-in-water emulsion with oxidized lipids induced β-glucan oxidative

degradation, which lead to diminished molecular weight of β-glucan and decreased the

viscosity of the solution. β-glucan amount and molecular weight seem to have significant

roles concerning the physiological actions of β-glucans in humans (Åman et al. 2004;

Andersson and Börjesdotter 2011). The degrading effect of oxidized lipids on β-glucan

might pose a problem in commercial functional foods with oat ingredients high in β-glucan,

as both the food matrix and the oat ingredients in themselves include potentially degradable

lipids. As this interaction between oxidized lipids and β-glucan molecular properties is a

fairly new finding, it is important to research the phenomenon more to understand it fully.

The previous studies have been made in oil-in-water emulsions, which makes it important

to investigate whether the same phenomenon happens in dry oat materials or in aqueous

foods containing oat. Degrading β-glucans in aqueous foods could propose technological

challenges as well as nutritional losses.

The object of this literature review was to gain knowledge and understanding of lipid

degradation in oats, and its impact on the functionality of oat ingredients, especially on the

physicochemical properties of β-glucan within the oat ingredients. The practical work was

designed as a three-month storage trial of β-glucan rich oat ingredients to experiment the

connection between lipid degradation and β-glucan functionality. Firstly, the lipid

degradation state in oat ingredients was analysed by chromatographic means at the

University of Helsinki. Secondly, the stored oat ingredients were used to prepare acid milk

gels, which modelled a yogurt-type semi-solid food, at VTT Technical Research Centre of

Finland. This information on oat lipids and β-glucan was crucial to obtain to be able to create

functional nutraceuticals with β-glucan to aid human health.

10

2 LITERATURE REVIEW

2.1 Oat bran, β-glucan, and its enrichment

2.1.1 Oat bran and β-glucan

According to AACC (1989), oat bran is “the food which is produced by grinding clean oat

groats or rolled oats and separating the resulting oat flour by sieving, bolting, and/or other

suitable means into fractions such that the oat bran fraction is not more than 50% of the

original starting material, and has a total β-glucan content of at least 5.5% (dry weight basis)

and a total dietary fibre content of at least 16.0% (dry weight basis), and such that at least

one-third of the total dietary fibre is soluble fibre.” Oat bran is thus a technical term used for

the outer layer milling fraction consisting of the coarse outer parts of the oat kernel: pericarp,

seed coat, aleurone, sub-aleurone, and starchy endosperm. In addition to starch (40–61%),

proteins (10–21%), and fat (8–12%), the bran is rich in minerals (2–4%) and cell wall

polysaccharides such as cellulose, arabinoxylan, and β-glucan (total dietary fibre 11–19%)

(Luhaloo et al. 1998). The main reason for high variations are the effects of oat variety,

agronomic as well as environmental factors (Givens et al. 2004). Additionally, there are

significant amounts of antioxidants present, such as tocopherols and tocotrienols, phytic

acid, avenanthramides and other phenolic compounds (Peterson 2001; Bryngelsson et al.

2002).

β-glucan is a water-soluble non-starch fibre component found in oat aleurone and endosperm

cell wall structures (Figure 1). It counts for approximately 3–11% (Cui and Wood 2000) of

the bran and can be described as a linear glucose homopolymer with β-D-glucopyranosyl

units (Figure 2a). Units are linked with β-(1à4) and β-(1à3) linkages at an approximate

ratio of 7:3. (1à3) linkages stand alone and are usually separated by either two (cellotriose

unit) or three (cellotetraose unit) (1à4) linkages, comprising a polysaccharide group called

mixed linkage β-glucans (Figure 2b).

11

Figure 1. Oat grain and its tissues, cell layers and nutrient distribution (e.g. β-glucan). Scheme by Grundy etal. (Grundy et al. 2018).

Figure 2. Oat β-glucan molecular structures: (a) A basic schematic structure of β-glucans as glucose polymers.(b) Approximate distribution of cellotrioses (one 3G1 and two 4G1s) and cellotetraoses (one 3G1 and three 4G1s)with occasional longer oligomers. 3G1 = (1à3) linkage, 4G1 = (1à4) linkage. Scheme b modified fromLazaridou and Biliaderis (Lazaridou and Biliaderis 2007).

The structure of the β-glucan molecule seems to be a critical aspect when considering its

bioaccessibility as well as its physicochemical properties and functionality, such as solubility

in water, viscosity and gelation properties (Lazaridou and Biliaderis 2007). It is believed that

the ability of β-glucans to form viscous solutions in the intestine helps to maintain normal

blood cholesterol, lower blood cholesterol, as well as reduce post-prandial glycaemic

responses (Regand et al. 2011; Whitehead et al. 2014). The scientifically proven beneficial

12

effect is achieved when the daily consumption of β-glucan reaches 3 grams (EFSA 2010,

2011). To use the EFSA approved health claims, the product must provide 1 g of β-glucan

per portion.

There has been much speculation on the mechanisms by which β-glucan positively affects

human health. In some studies, the desired effects have not been acquired even though the

consumption levels greatly exceeded the required daily amounts (Andersson and

Börjesdotter 2011; Whitehead et al. 2014). Two molecular aspects that seem to be crucial

for β-glucan’s physiological actions are the amount of solubilized β-glucan (in the intestine)

and its molecular weight (MW), both of which need to be sufficiently high (Åman et al.

2004; Andersson and Börjesdotter 2011). Furthermore, structural properties and MW greatly

affect the water solubility, viscosity and gelation properties of oat β-glucans (Cui and Wood

2000). The lack of physiological efficacy is most often connected to the β-glucan MW, and

it seems that medium to high MW β-glucan (530–2210 kDa) would be more efficient in, for

example, actively lowering serum cholesterol levels in humans than low MW β-glucan

(21 kDa) (Wolever et al. 2010). On the other hand, as discussed by Wang et al. (2017), low

MW polysaccharides have been thought to have higher mobility than larger β-glucans, and

thus have a greater antioxidant activity than high MW β-glucans as well as a better ability to

effectively improve the oxidative parameters, for example in the spleen (Błaszczyk et al.

2015). Also, technological properties of β-glucan are dependent on MW, and beverages, for

example, need to obtain low MW β-glucan to avoid aggregation and sedimentation (Sibakov

et al. 2013).

Whitehead et al. (2014) found that the solubility of β-glucans is affected by their

bioaccessibility, in other words by the food matrix. The food matrix may also affect the MW

of β-glucans as the food matrix and its modification (food processing) might induce

degradation of the β-glucan molecules resulting in lowered MW. The effect of commercial

processing and the source and type of extraction of β-glucan are considered in more detail,

for example, in the research of Keogh et al. (2003). Depending on the extraction method and

conditions, solvents, and sample history, the molecular weight of β-glucans can range

anywhere between 65 to over 3000 kDa (Lazaridou and Biliaderis 2007). Thus it should be

noted, as stated in the meta-analysis of Whitehead et al. (2014), that the lack of a

standardized method for β-glucan MW analysis makes it challenging to accomplish a

confident relation between the β-glucan MW and its functionality.

13

Other aspects effecting β-glucan molecular integrity are the presence of enzymes, heat,

mechanical forces, and acids as well as oxidative degradation by ascorbic acid and iron

cations, a pro-oxidant forming hydroxyl radicals that attack β-glucan (Kivelä et al. 2009a,

2009b, 2012; Mäkelä et al. 2015). However, there are differences on degradation behaviour

of β-glucans depending on their sources and levels of residual phytates: oat β-glucan has

been found to have a higher iron binding capacity, quenching and retarding the iron cations

as pro-oxidants in the formation of hydroxyl radicals through the Fenton reaction (Wang et

al. 2017). Therefore, when designing functional foods with β-glucan, it is important to

consider the processes of producing the β-glucan, the food matrix type (solid, semi-solid or

liquid), and the MW of the β-glucan in the final product, to make sure the product is

acceptable and the product’s role as a nutraceutical is not compromised.

2.1.2 Dry-fractionation for β-glucan enriched ingredients

Oat β-glucan can be isolated or enriched by dry milling, wet milling sieving, air classification

and solvent extraction (Tiwari and Cummins 2009). The process usually includes dehulling,

sieving, milling, more sieving and/or air classifying at varying repetitions together with

common heat treatments to remove part of the starch and obtain oat bran and finally β-glucan

enriched ingredients with varying particle sizes (Figure 3). Dry fractionation methods are

often more favoured than wet-fractionation methods, because they are more ecological and

cost-effective as there is no need for drying steps. Additionally, aspects favouring dry-

fractionation include much higher fraction mass yields than in wet-extraction, the ready

usability of all residual fractions, low microbiological risks in dry ingredients, and the fact

that the produced partially purified or enriched oat ingredients are pure enough for most food

products (Sibakov 2014). Dry milling concentration of β-glucan is also nutritionally

interesting, as the oat bran concentrate includes both soluble and insoluble material and thus

has combined beneficial physiological effects of the two dietary fibre classes (Lazaridou and

Biliaderis 2007). Patented dry-fractionation methods to produce oat fractions with high

β-glucan concentrations, have been designed for example by Mälkki et al. (2001). Separation

of oat grain cells can be improved by removing oat lipids with supercritical carbon dioxide

(SC-CO2) (Sibakov et al. 2011; Konak et al. 2014) and it has been found that this type of

processing also improves the foaming properties of oat-derived products or ingredients

(Kaukonen et al. 2011).

14

Figure 3. The structural levels when extracting β-glucan (Grundy et al. 2018).

Because dry-fractionation is a concentration method using only physical and technological

methods to modify β-glucan content, it should be noted that dry-fractioned ingredients are

hybrid ingredients that may contain proteins, lipids, minerals and other possible components,

such as phytic acid. These components will affect the behaviour of the β-glucan rich oat

ingredient in food applications, and should be considered when designing products with oat

bran concentrates.

2.2 Oat lipids

2.2.1 Lipid composition of oat bran

Oil storages in a grain are close to the living tissue, close to lipid degrading enzymes, so that

they can be used during germination (Heneen et al. 2009). Usually the germ of any cereal

grain is considered the main storage place containing these grain’s valuable energy bundles,

lipids. However, relative oat lipid concentration increases from the inner starchy endosperm

to the outer layers of the kernel (Miller and Fulcher 2011). In this way the aleurone layer has

the highest relative lipid concentration within the oat grain (Figure 4), thus making lipids a

significant constituent of oat bran.

15

Figure 4. Oil droplets (bright/yellow) in oat aleurone layer cells visualised by staining with Nile Blue in cryosections of oat grains. Microscopy image by courtesy of VTT Technical Research Centre of Finland Ltd.

Both lipid content and fatty acid composition of oats vary depending on the cultivar,

environment, and agronomic factors, as well as the extraction method of the lipids (Zhou et

al. 1999a; Givens et al. 2004). Luhaloo et al. (1998) explain that crude fat contents in

different oat brans range between 8 and 12%. Fatty acids are chiefly stored as

triacylglycerols (TAGs), and TAGs count for around 80% of all the lipid classes in oats.

TAGs consist of a glycerol and three fatty acids, and in oats, 80% of these fatty acids are

unsaturated. Zhou et al. (1999a) report that the lipid composition of oat brans varies

depending on processing conditions, but in oats in general, around 40% of the fatty acids are

oleic acid (C18:1) and another 40% are linoleic acid (C18:2). The rest of the fatty acids are

palmitic acid (C16:0, 15–20%) with smaller amounts of stearic acid (C18:0), α-linolenic acid

(C18:3), and myristic acid (C14:0). Concerning other lipid classes in oats, around 20% are

polar lipids (phospholipids and glycolipids) and the final percentages mainly sterol esters

(Zhou et al. 1999a). The high amount of unsaturated fatty acids in the oats, including an

essential fatty acid linoleic acid, add to the nutritional value of oats. The high lipid content

is also interesting from a technological aspect. The lipids can have a modifying effect on oat

starch pasting properties, which in turn affect oil stability and instability, and applications as

fat replacers can be made both in food and cosmetic industries (Zhou et al. 1999a).

16

Lipids in oat can be analysed as a whole fraction or as individual lipid classes. The pre-

treatments of the material, the oat fraction and the chosen method, from extraction to

characterisation and quantification, significantly affect the perceived total lipid content.

Most often, the method includes some kind of an extraction step. One of the most popular

extraction solvents used to be hexane, which gives most conservative estimations on the total

lipid content compared to e.g. 85% methanol or ethanol (Zhou et al. 1999a). The more polar

the extraction solvent is, the more it also extracts polar lipids and sterol esters. However, due

to its harmful nature, hexane is now successfully substituted with heptane in lipid analyses,

especially when mixed with more polar solvents, such as 2-propanol (Damerau et al. 2014).

As fatty acids count for most lipids in oats, sometimes it is sufficient enough to add all the

fatty acids together to represent lipid levels in the oat sample. If done so, the lipid content

should be expressed as the total sum of fatty acids, typically mg/g sample or as a percentage.

When extracting and analysing total fatty acids, the process usually includes acid hydrolysis,

saponification and methylation steps to form fatty acid methyl esters detectable by gas

chromatography (GC), as in the AOCS Ce 2c-11 method (Srigley and Mossoba 2017).

Instead of acid hydrolysis, acetone can also be used as an extraction solvent when used with

high enough temperature and pressure with an Accelerated Solvent Extractor (ASE) (Lampi

et al. 2015). Final quantification is most often performed using an internal standard, a fatty

acid not inherently found in the oat lipids.

2.2.2 Antioxidants in oat bran

In addition to lipid classes, the oat bran is also rich in lipid soluble antioxidants. Antioxidants

prevent or inhibit lipid oxidation, and the most abundant fat-soluble antioxidant in the oat

bran is α-tocotrienol (a-T3). Together with α-tocopherol (a-T), β-tocopherol (b-T), and

β-tocotrienol (b-T3), α-tocotrienol is one of four tocopherols and tocotrienols (tocols)

present in the oat bran (Bryngelsson et al. 2002). Tocols are phenolic compounds with

varying amounts and positions of methyl groups within their chromanol ring (Figure 5).

Tocopherols and tocotrienols also differ by the unsaturation level of their phytyl tails.

17

Figure 5. Tocopherol (a) and tocotrienol (b) molecular structures consisting of a chromanol ring and a phytyltail. a, b, g, d = α, β, γ, δ, respectively. T = tocopherol, T3 = tocotrienol.

When oat cell walls are broken during processing, tocols are exposed. This increases the

bioavailability of the antioxidants as well as the chances for their degradation (Bryngelsson

et al. 2002). The radical quenching antioxidant activity is based on the ability of tocols to

donate their phenolic hydrogens to lipid derived free-radicals, making the phenolic

chromanol ring the main location where lipid oxidation is inhibited. As a phenolic hydrogen

is donated to a lipid peroxyl radical, a resonance stabilized chromanoxyl radical is formed

(Kamal-Eldin and Appelqvist 1996). Degradation products of tocols include lactones and

quinones. Depending on the tocol, the environmental factors, as well as the food matrix,

dimers and trimers may also be formed. Because of their ability to act as free radical

scavengers, tocopherol and tocotrienol levels can be used to indirectly follow and assess

lipid oxidation states in oats.

Tocols are usually analysed by chromatographic methods. High performance liquid

chromatography (HPLC) offers the most advantageous methods, and Kamal-Eldin et al.

(2000) note that HPLC methods were fast, simple, sensitive, and selective, superior to GC

methods. The authors continued to explain that although both normal and reversed phase

columns can be used, and although reversed phase (RP) columns offer better stability and

longer durability, the normal phase (NP) columns are better able to separate β- and γ-isomers

of tocopherols and tocotrienols. Due to their heterocyclic nature, tocols can be detected based

on ultraviolet absorbance (UV detector) or fluorescence (fluorescence detector, FLD).

18

Besides tocols, additional oat bran antioxidants include phytic acid, avenanthramides and

other phenolic compounds (Peterson 2001; Bryngelsson et al. 2002). Avenanthramides have

been under vigorous research for the past decade or so, and they are unique for oats.

Avenanthramides are phenolic alkaloids and at least 25 avenanthramide types can be

identified, although most of the avenanthramides consist of three main types (Peterson 2001;

Martínez-Villaluenga and Peñas 2017). They are abundant in the bran fraction, although

measured concentrations depend on processing and analysis methods (Hitayezu et al. 2015).

Avenanthramide synthesis is enzyme regulated and these polyphenols are believed to be a

part of a plant defence mechanism (Peterson 2001). Avenanthramides are believed to greatly

add to the health benefits of oats. It is said that these polyphenols exhibit anti-inflammatory,

antiproliferative, and anti-itching activity, which may have bioactive effects alongside

β-glucans, such as contributing to protecting against coronary heart disease and colon cancer,

as well as calming skin irritations (Meydani 2009). As it is with tocols, avenanthramides

might degrade during heat and processing of oats (Bryngelsson et al. 2002), which could be

compensated by the possible release of bound avenanthramides during storage at high

temperatures (Kaur et al. 2018).

2.2.3 Lipid reactions in oats

Chemical oxidation

Lipid oxidation means predominantly the deterioration of double bond containing

unsaturated fatty acids (Schaich et al. 2013). Some of the main factors in affecting the overall

oxidation rate of lipids include the reaction environment, that is e.g. temperature,

the presence of inhibitors and catalysts, and the nature of the compounds involved (Kim and

Min 2008). As oats are high in unsaturated fatty acids, they are susceptible to lipid

degradation (Zhou et al. 1999a).

Chemical lipid oxidation can be described as a series of autocatalytic reactions (Reische et

al. 2008). In this autoxidation process, a free radical chain reaction is generated and

maintained as long as there are substrates and radicals present (Schaich et al. 2013). The

commonly accepted theory has been the free radical chain reaction theory (Ingold 1969;

Schaich et al. 2013). According to this theory, autoxidation can be divided into three main

phases: the initiation, propagation and termination stages (Ingold 1969). Together their

kinetic behaviour can be describes as a sigmoidal curve, where the initiation phase seems

19

long and slow, until the exponential propagation phase begins. The termination phase can be

seen as the final plateau of the curve, and slowing down of reactions.

Because normal oxygen is a triplet according to its energy state, an external initiator or a

catalyst is needed for the autoxidation to begin. Spin states of the reacting molecule parts

need to be corresponding for a reaction to occur, which is why the common triplet oxygen

cannot react spontaneously with unsaturated fatty acids that have singlet state double bonds.

Therefore, the start of the chain reaction, the initiation, most often requires an initiator or a

catalyst that can remove an electron and creates a radical molecule of one of the compounds.

Some of the most typical initiators include preformed or primary radicals, pro-oxidative

metals, enzymes, hemes, and pigments, like chlorophylls. Another considerable group of

initiators includes more physical factors, such as UV-radiation, visible light, and heat, which

all add energy to the lipid system (Shahidi and Wanasundara 2008). These often work

indirectly, for example visible light does not have the required energy to produce radicals

but it can transfer energy and affect oxidation through photosensitizers such as chlorophylls

(Schaich et al. 2013). After a radical has been formed, the radical reacts with a bisallylic

hydrogen of an unsaturated fatty acid and a lipid alkyl radical R• is formed, kick-starting the

free radical chain reaction (Ingold 1969, Kim and Min 2008; Schaich et al. 2013).

Propagation can be described as “the heart of lipid oxidation” (Ingold 1969). In

propagation, oxygen spontaneously converts the alkyl radical to a more reactive peroxyl

radical (ROO•). To quenc its instability, the lipid peroxyl radical will attack an adjacent fatty

acid, abstract a hydrogen, and thus turn into a lipid hydroperoxide (ROOH). In tandem,

another lipid alkyl radical R• is produced. As this process is looped, autocatalysis of lipid

oxidation is enabled (Shahidi and Wanasundara 2008; Schaich et al. 2013). Lipid oxidation

may be extended by excited oxygen atoms, singlet oxygens 1O2. Singlet oxygens are not

radicals, but they are able to react with non-radical, singlet state double bonds and form

hydroperoxides without the intermediate peroxyl radical, because of their high-energy state,

leading to chain branching and supercharging lipid oxidation even more (Kim and Min

2008).

Autocatalytic reaction chains are stopped only if (1) there are no more substrates (fatty acids)

or (2) two free radicals react together to form a more stable non-radical secondary product

(Schaich et al. 2013). Quenching of the radical reaction chains does happen throughout

autoxidation (Schaich et al. 2013). However, Schaich et al. (2013) continue to remark that

20

when the rate of quenching radicals is greater than the rate by which new free radical chains

are produced, the oxidation rate can and will slow down into the termination phase.

Termination has a cumulative effect as individual radicals react together and produce more

stable and non-radical products (Ingold 1969). For an overview on the classical chain

reactions during initiation, propagation, and termination, please refer to Figure 6.

Figure 6. The commonly accepted understanding of how the free radical chain reactions of lipids advance.R•=lipid alkyl radical, ROO•=peroxyl radical, ROOH=hydroperoxide, RR/ROOR=non-radical products.Scheme by Sahahidi and Wanasundara (2008).

The apparent loss of substrates (fatty acids) and possible deterioration on nutritional quality

are not the only changes the oil system undergoes. The increase of the primary oxidation

product, hydroperoxide, contributes to most of the changes in the oil phase. Hydroperoxides

are closely involved in the formation of various secondary products, since hydroperoxides

are transitory in nature and decompose further to other products (Shahidi and Wanasundara

2008). Hydroperoxides are colourless and odourless as such but the secondary products are

often associated with changes in flavour properties because of their highly volatile and

odour-active nature (Kim and Min 2008).

The secondary products are also usually product specific, which stems from the differences

of the fatty acids. Different fatty acids have different activation energies needed to start the

free radical chain reactions (Kim and Min 2008). Based on activation energies, reactions are

most probable in places where bond-energies are at their lowest. As explained by Kerr

(1966), fatty acids have these low bond-energy domains at their allylic C-H bonds.

Furthermore, Cosgrove et al. (1987) showed that the oxidizability of polyunsaturated fatty

acids (PUFA) is linearly dependent on the number of doubly allylic positions, which results

21

in easier abstractions of hydrogens in between double bonds: When a hydrogen is abstracted,

the alkyl is left with an imbalance of electrons. The pentadiene structure compensates the

unevenness by resonance stabilization. This resonance spreads the electrons on a larger area

and exposes the now electron deficient outside positions, which can result in further oxygen

addition and hydroperoxide formation (Schaich et al. 2013). In this manner, PUFA are more

susceptible to oxidation than monounsaturated fatty acids (MUFA) (Schaich et al. 2013).

Depending on the structure of the fatty acid, the creation of these enhanced hydrogen-targets

can enable chain branching and the formation of specific secondary products (Hyde and

Verdin 1968).

Alongside hydrogen abstraction, there is a plethora of other competing chemical reactions

producing secondary lipid oxidation products. These mechanisms include isomerization,

β-scission, and scission, cyclization, recombination and addition, producing more radicals

or polymers, aldehydes, alkanes, oxo compounds, epoxides, and ketones, to name a few

(Schaich et al. 2013). As oats are abundant in linoleic acid, one of the most typical

autoxidation products in oats is the volatile aldehyde hexanal (Zhou et al. 1999a; Lehtinen

and Laakso 2004). However, there are both primary oxidation products (fatty acid

hydroperoxides) as well as other secondary products, such as the aforementioned hexanal.

Enzymatic hydrolysis and oxidation

Lipids are a source of energy for the potentially growing oat seed, and lipid degrading

enzymes in oat are meant to release this energy for germination of the seed (Lascorz and

Drapron 1987). Oat lipids are close to the enzyme production sites that are related to

germination and mobilization (Heneen et al. 2009). In a developing grain, oil bodies from

the embryonic axis and the aleurone layer fuse and accumulate into the endosperm (Heneen

et al. 2008). During germination, enzymes synthetized and secreted from the aleurone and

scutellum tissues mobilize lipid reserves (TAGs) as transportable nutrients (FFAs), which

results in oil droplets accumulating close to the aleurone and scutellar epithelium (Lenova

et al. 2010). For this reason, it is likely for endogenous lipid degrading enzymes to be present

in oat fractions, especially milling fractions containing the aleurone layer.

The main lipid degrading enzymes in oats are lipase, lipoxygenase (LOX), and peroxidase

(POX), and at least lipase is known to reside close to the outer layers of the kernel, or the

bran (Ekstrand et al. 1992). Lipase hydrolyses the ester bound fatty acids from

triacylglycerols (TAGs) first partially into di- and monoacylglycerols, and then soon into

22

free fatty acids (FFAs), which are highly susceptible to autoxidation (Yang et al. 2017).

Accordingly, hydrolysis of oat lipids can be followed by the formation of FFA, which can

start immediately after the cell wall structures are broken in milling, promoting oxidation.

LOX oxidizes fatty acids with pentadiene structures and produces hydroperoxides. This

activity is held to be responsible especially for volatile secondary products contributing to

off-flavours, such as the volatile aldehyde, hexanal and 2-pentylfuran (Molteberg et al. 1996;

Heiniö et al. 2002). LOX activity has also potential to co-oxidate lipid soluble vitamins (e.g.

tocols) (Robinson et al. 1995), which prohibits the vitamins’ antioxidant activity and further

promotes oxidation. However, LOX activity in oats is found to be low, leaving the oxidation

of oat lipids to the non-enzymatic autoxidation (Yang et al. 2017). POX activity produces

fatty acid epoxides and hydroxy fatty acids by transferring oxygen intra- and

intermolecularly from fatty acid hydroperoxides to unsaturated fatty acids (Piazza et al.

2003). These non-volatile compounds are reported to contribute to the bitter taste of stale

oats (Biermann et al. 1980). To prevent enzymatic lipid degradation, enzymes are typically

inactivated by hydrothermal processes.

Lipid reactions during oat processing and storage

The first stage where unwanted lipid hydrolysis can occur, is during dehulling, as there is

the risk of kernel breakage (Doehlert and McMullen 2001). However, unless there is

excessive moisture, this is considered a minor problem with low correlations between kernel

breakage and lipid hydrolysis (Peltonen-Sainio et al. 2004). In milling and fractionation, cell

wall structures are broken, and both lipids and antioxidants are exposed form the protection

of the structures and rendering them to the effects of hydrolysis (if enzymes have not been

inactivated) and oxidation. As the oat bran is high in trace minerals, they too are exposed,

providing initiation sites for autoxidation. If fractionation is done in wet conditions, there is

a risk for lipolytic enzyme activity. However, this kind of enzyme activity can be suppressed

by using, for example, alkaline water (Liukkonen et al. 1993).

To inactivate enzymes, hydrothermal processing (steaming and kilning) is typically applied

to oat kernels. However, heat and added humidity induce degradation of antioxidants, such

as tocols (Bryngelsson et al. 2002), which in turn can promote autoxidative reactions. Heat

treatment has also been found to promote lipid oxidation: e.g. the more sufficient lipase

inactivation is, the more lipid oxidation related products are formed and observed (Lehtinen

et al. 2003). This is especially emphasised in long-term storage, which was the main focus

of the study by Head et al. (2011), in which commercially heated and superheated steam

23

treated oat groats were stored for up to 26 weeks. In their study, the hexanal amounts

increased substantially: more for commercially treated and less for superheated steam treated

oats. However, there were no changes in the colour or content of free fatty acids, indicating

little or no Maillard reaction or hydrolysis in either of the differently processed oat groats.

Extrusion has also been presented as a viable alternative to conventional heat treatments and

oat processing (Lampi et al. 2015). Parker et al. (2000) extruded oat flours in different

conditions and they concluded that extruded debranned oat flour had the highest amounts of

lipid oxidation related alkanals, 2-alkenals, and 2,4-alkedienals. Also, when the extrusion

temperature was at its highest and moisture content at its lowest, the levels of Maillard

reaction products were elevated, such as pyrazines, pyrroles, furans, and compounds

containing sulphur.

Processing also affects the flavour of oats. Initially the formed (volatile) flavour compounds

are considered positive and characteristic to oats. These can include raw (hay-like), oaty

(normal for oats) and toasted notes (derived from Maillard reaction) (Klensporf and Jeleń

2008). However, possible enzymatic hydrolysis and chemical oxidation products can soon

be experienced as bitter and rancid off-flavours, respectively. Hexanal is a typical volatile

lipid oxidation product that gives a grassy off-flavour and has a particularly low aroma

threshold (Heiniö et al. 2002). In fact, hexanal formation is one of the most abundant of the

prominent features of oat lipid oxidation. Interestingly though, nitrogen containing

compounds, such as amino acids, dipeptides, and polyamines, have been found to have

hexanal quenching abilities (Zhou and Decker 1999), which could add to the findings of

Lehto et al. (2003), who found aldehyde dehydrogenase type activity in oats when hexanal

was absorbed into the oat matrix and oxidized to hexanoic acid.

When oat products are stored for a prolonged time, both lipid hydrolysis and oxidation

reactions have time to branch out, yielding more and more degraded lipid-derived products

with high risk of co-oxidation (Schaich et al. 2013). Oat is a hybrid ingredient containing

other components such as proteins, fibres, starch and antioxidants in addition to its lipids.

These compounds will experience degradative reactions, such as co-oxidation, alongside

lipids during storage (Rakić et al. 2014). It is not always clear whether lipid reactions and

certain reactions of these other compounds have a cause and effect relationship. However, it

is very likely that the free radicals from lipid oxidation promote and cause the oxidation of

adjacent compounds (Schaich et al. 2013). Additionally, it is also very important to consider

what type of food is in question. An oat containing emulsion with high water activity and

24

lots of interfacial area will behave very differently compared to dry oat flour in storage. Even

though fine oat flour will have a lot of interfacial area with air, an emulsion will have

technological challenges (emulsion stability (Rousseau 2000)) and chemical challenges

(Laguerre et al. 2009). Time seems to be the one and most important factor in lipid stability

in storage. To slow down and control the inevitable degradation reactions during this time,

light, packaging materials, and storage conditions (temperature, moisture, and oxygen) have

the most considerable effects on lipid stability in stored oat products (Lehtinen and Laakso

2004; Kaur et al. 2018).

In conclusion, hydrolysis of oat lipids can be prevented by inactivating lipolytic oat enzymes

whereas chemical oxidation of oat products is most effectively prevented by controlling

storage temperature, light, and the nature of the container. A rancid oil that has gone through

all three stages of autoxidation is problematic in many ways. Not only can there be toxic

compounds such as epoxides present, but the sensory qualities of the oil suffer, resulting in

changes at least in odour and flavours, as discussed previously. The chemical changes in the

stored oat rise also technological and economical challenges as the functionality of the oat

product might change.

Co-oxidation of β-glucan

It was recently discovered by Wang et al. (2016) that in an emulsion, barley β-glucan

degradation was induced when oxidized lipids were present. The breakdown of β-glucan

molecules was observed as a significant decrease in both emulsion viscosity and β-glucan

molecular weight (MW). These findings raised a lot of questions, as the interactions of

oxidized lipids and β-glucan molecules have not yet been vigorously researched.

Oxidative degradation of β-glucans can happen in food processing and in aqueous food

models (Kivelä et al. 2009a, b, 2012). And as pointed out by Wang (2018), there are

indications of lipid co-oxidation of polysaccharides, where, for example, β-cyclodextrin was

degraded because of linoleate autoxidation and corn starch molecular characteristics

changed because of methyl linoleate. Even so, literature on β-glucan co-oxidation with lipids

is scarce. What can be said is that lipid oxidation induced degradation of β-glucan is most

probably linked to the termination phase of lipid oxidation, as is the case with many other

non-lipid molecules, such as proteins and carbohydrates are co-oxidated (Schaich et al.

2013). Wang (2018) suggests that although lipids are hydrophobic, lipid oxidation products

25

can affect the surrounding molecules through the interface, making lipid oxidation a source

of radicals, which would then degrade β-glucans.

2.3 Yogurts

2.3.1 Milk proteins

Milk is a complex colloidal system of globular fat (mainly TAGs) and proteins in a water-

based solution meant for its primary natural function: nutrition to the young mammals

(Figure 7). Bovine milk contains, on average 87% water, 3.3% protein, 4% fat together with

carbohydrates, vitamins and minerals (Walstra et al. 2006).

Figure 7. Schematic shapes of structural elements in milk illustrating the elements’ size differences, viewed ata magnification of 50 000. Fat globules and casein micelles float in a cloudy or opalscent liquid called serum.Image obtained from Walstra et al. (2006).

Milk proteins are typically divided into two types of proteins. The proteins are grouped based

on their solubility and structure/folding properties into caseins and whey proteins: (1)

Caseins (αS1-, αS2-, β-, and κ-casein) constitute around 80% of total nitrogen in milks.

Caseins have some helical and β-structures in their hydrophobic domains. As their primary

structure is rich in well-conserved prolines, caseins do not have a well-defined tertiary

structure. Consequently, caseins have an open rheomorphic conformation and are

outstandingly heat-stable (Holt and Sawyer 1993). Caseins are also characterised by their

precipitation properties at their isoelectric point (pI) at around pH 4.6. αS1-, αS2-, and β-

caseins are highly phosphorylated, and thus bind calcium cations (Ca2+) effectively making

them precipitate in the presence of Ca2+. κ-caseins, on the other hand, have their C-terminals

glycosylated, do not bind Ca2+ cations and are soluble in the presence of these cations

26

abundant in milk. As the glycosylated part of the C-terminal is a very hydrophilic domain,

and the N-terminals of the κ-caseins are highly hydrophobic, κ-caseins are a

characteristically amphiphilic group of proteins.

Milk is naturally rich in calcium phosphate, which is why caseins organize themselves as

association colloids by hydrophobic interactions and calcium phosphate bridges. These

association groups are often referred to as casein micelles (Figure 8). They vary in sizes (50–

600 nm), are hydrated, stable and polydispersed (Marchin et al. 2007). Because of the

amphiphilic properties of κ-caseins (protruding peptide chains in Figure 8), κ-caseins form

the negatively net charged outer layer of the micelles and are able to associate with the more

hydrophilic serum phase around the micelles (Dalgleish 2011). The inner parts of the

micelles hold calcium-sensitive α- and β-caseins protected. According to the review done by

Dalgleish (2011) caseins are thought to be held tightly together by colloidal calcium

phosphate, yet, micelle-micelle interactions are prevented by the “hairy layer”, the outer

layer of κ-caseins which makes the surface rough. However, there is some proof that the

interior parts of the micelles can be accessed through pores in the outer κ-casein layer

(Marchin et al. 2007; Knudsen and Skibsted 2010; Dalgleish 2011).

Figure 8. Cross-section of a casein micelle model where caseins are thought to be a somewhat organizedassociation colloid with smaller nanoclusters and submicelles making up the inner parts of the casein micelle.Image obtained from Walstra et al. (2006).

After the precipitation of caseins at their isoelectric point, (2) whey proteins are left,

counting up to 15% of the total nitrogen content in milk. Whey proteins are globular proteins

with compact and well-defined secondary and tertiary structures. 50% of whey proteins are

27

β-lactoglobulins (BLG, pI=5.2) and another 20% of the whey proteins are α-lactalbumin

(ALA, pI=4.8) (Haginaka 2000; Chen et al. 2016). Whey proteins have cysteines in their

primary structure which enable them to form intramolecular disulphide bonds. BLG has one

free cysteine that is hidden in its β-barrel structure (Carter and Ho 1994). When the

conformation of the protein changes and the tertiary structure is opened, the free cysteine

domain becomes exposed and reactive (Brownlow et al. 1997; Considine et al. 2007).

2.3.2 Acid milk gels

Traditionally yogurts are prepared using lactic acid bacteria (LAB) to ferment milk lactose

to lactic acid, which slowly acidifies the milk. Nonetheless, to provide control and

reproducibility for both food industry and research purposes, milk can be acidified by adding

an acidifier, such as glucono-δ-lactone (GDL), which gradually hydrolyses into gluconic

acid when in contact with water (Lucey et al. 1998; Lee and Lucey 2004; Marchin et al.

2007; Martin et al. 2009; Liu et al. 2017). When using GDL, the pH drop is similarly gradual

as with LAB fermentation. However, there are always structural differences between GDL

acidified and LAB fermented yogurts, as the initial acidification rate is faster with GDL

(Lucey et al. 1998; Martin et al. 2009). An example of the GDL induced protein network is

presented in Figure 9.

Figure 9. Confocal laser scanning micrographs of acidified acid milk gels at (a) 1 min, (b) 30 min, (c) 60 min,and (d) 120 min after the addition of glucono-δ-lactone, with the target pH of 4.6. Protein particles (bright)were labelled with Rhodamine B. Scale = 25 μm. Images obtained from Auty (2011).

28

Yogurts and acid milk gels can be described as a coarse network of linked protein clusters

and strands (Kaláb et al. 1983). To obtain this protein network, the proteins need to be

modified both with heat and the pH. The heat treatment partially unfolds caseins causing

changes in secondary structures and significant losses of the tertiary structures. This in turn,

causes hidden domains to be exposed which induces further conformational changes in the

proteins that are vital for the gel to form. Whey proteins are also partially unfolded: the heat

treatment (>70 °C) causes dissociation and aggregation of BLG proteins as well as thiol-

disulphide exchange reactions (Considine et al. 2007; Auty 2011). The free thiols will react

with other whey proteins and κ-caseins on the surfaces of micelles, which enables the

formation of micelle-bound and soluble κ-casein/whey protein complexes in milk.

Despite the role of the heat treatment, yogurts are most importantly acidified milk products.

The actual gelation happens by means of acidification even though the heat treatment is

important for the success of the gel. As the pH decreases and the matrix acidifies, micellar

calcium phosphate gradually dissolves and becomes fully soluble around pH 5.2 (Law and

Leaver 1998). This changes the charge-balance within the milk causing the negative surface

charges of the micelles to be balanced. The charge-balancing inflicts the outer hairy layer of

the micelles to collapse, bringing about a loss of steric and entropic effects between micelles

(Roefs and Van Vliet 1990; De Kruif 1997) as well as the increase of electrostatic repulsion

and weak physical interactions (hydrophobic and van der Waals attractions). Moreover, as

the concentration of dissolved Ca2+ increases and the pH approaches 4.6 (the pI of caseins),

the now exposed α- and β-caseins are precipitated. Thus, the pH drop increases micelle-

micelle interactions, which progressively creates a three-dimensional protein network

(Lucey and Singh 1997; Lucey et al. 1998).

The temperature of the heat treatment and the acidity of the milk highly affect the degree of

whey protein denaturation and the distribution of the bound and soluble complexes, thus

effecting how extensive the aggregation formation will be (Lucey and Singh 1997). The lack

of this extensity can be visibly seen on and around the gel as spontaneous syneresis, a

phenomenon where the protein gel network shrinks, becomes firmer, and releases some of

its serum phase (Ercili-Cura et al. 2013). Syneresis is commonly thought as a product defect

and is pronounced in low-fat products and when acidification time is reduced (see e.g.

Aryana and Olson 2017; Kumar et al. 2018).

29

2.3.3 Fibre fortified yogurts

The dietary fibre intake is usually low in the Western diet (Sonnenburg and Sonnenburg

2014). Even though it has been a controversial topic, it is known that dietary fibre has a

significant role in maintaining good human health (see e.g. Bingham et al. 2003; Makki et

al. 2018). To increase dietary fibre intakes, different dietary fibres have been added too

foodstuff as functional ingredients (Dhingra et al. 2012). As yogurts and other cultured dairy

products are expected to continue in their popularity among consumers, adding dietary fibres

to yogurts is an interesting and a convenient way of intervention (Gahruie et al. 2015; Aryana

and Olson 2017).

Dietary fibres used in yogurt fortification follow the definition of dietary fibres, where the

fibres are more often insoluble than soluble in water, are resistant to digestion and absorption

in the small intestine, and are usually polysaccharides, such as cellulose, hemicellulose and

pectins, and non-starch polysaccharides such as arabinoxylans (DeVries et al. 2001). These

fibres are usually extracted from the cell walls of fruits, vegetables, or cereals (Dello Staffolo

et al. 2004).

According to Dello Staffolo et al. (2004), the type of fibre and the storage time of yogurts

significantly affect rheological parameters of the yogurt. The authors continue to explain

that even though different fibres may or may not affect (spontaneous) syneresis and pH, they

do usually affect textural properties and sensory acceptability of yogurts: at their best, added

(dietary) fibres might add to the overall acceptability of fortified yogurt products in relation

to colour, flavour, and texture, and thus establish a functional food with commercial

applications. Some common (dietary) fibres used with yogurts include apple, inulin, wheat,

and bamboo fibres (Dello Staffolo et al. 2004), pineapple peel fibre (Sah et al. 2016), and

oat fibres (Fernández-García et al. 1998; Tomic et al. 2017). Adding oats to yogurts has been

associated with improved body and texture of plain yogurts (Fernández-García et al. 1998).

As the health benefits of oats have been contributed to its β-glucan fibres, it is feasible to not

only add parts of oats, but specifically increase the amount of oat β-glucan in yogurts.

The effect of adding β-glucans to yogurts have generally been associated with improved

water retaining properties observed by increased water holding capacity (WHC) and

decreased spontaneous syneresis (Table 1). β-glucans can be sourced from oats (Andersson

and Börjesdotter 2011) as well as form barley (Regand et al. 2011), spent brewer’s yeast

30

Table 1. Research on yogurts and acid milk gels with added β-glucan.

Reference Yogurt type β-glucan type Fibre inproduct Effects on yogurt quality and acceptability

Brennan andTudorica(2008)

Stirred yogurt Barley β-glucan (86%,Glucagel™)

β-glucan0.5–2.5%,

0.5% β-glucan was enough to improve serum retention and rheological properties. Overallimproved acceptability. Could be used as a fat-replacer.

Lazaridou etal. (2008)

Set acid skim milkgel

Barley β-glucans: 250, 210,180, 140, 70, and 40 kDa 0.5–2%

Weakened protein network, which could be compensated by high viscosity or a secondary gelnetwork with specific β-glucans. Low concentration increased and high concentrationsreduced spontaneous syneresis. Can influence textural attributes to form less cohesive butfirmer, more elastic, adhesive, and gummy products.

Sahan et al.(2008) Non-fat yogurt β-glucan composite (5.5%

soluble β-glucan) 0–1%Gel firmness and WHC1 were not influenced. Viscosity increased and spontaneous syneresisdecreased. In storage, firmness and WHC1 decreased while viscosity and syneresis furtherincreased. Lower levels of β-glucan composite gave satisfactory sensory results.

Elizaquívelet al. (2011)

Yogurt, reconstitutedsemi-skimmed milk

(1,3)-β-D-glucan producedby incorporated LAB2

strains [Pediococccusparvulus (strains 2.6 andCUPV22) and Lactobacillussuebicus CUPV221]

NA3 Did not significantly affect physical and rheological properties

Lazaridou etal. (2014)

Set yogurt,reconstituted milk

Oat β-glucan isolate,high solubility, 117 kDa 1.4%

Resulted in a liquid-like structure: a transient gel structure at an early stage due to phaseseparation of proteins and β-glucan, later retardation of protein aggregation and acidificationkinetics. β-glucan enhanced viability of probiotics in cold storage.

Rinaldi et al.(2015)

Non-fat yogurt,reconstituted milk

Barley β-glucan (75.6%,Glucagel™) 0.4%

Improved viscosity and delayed decrease of viscosity at high shear rate. Observations ofpossible phase separation might explain fast protein digestion for β-glucan enriched yogurts.

Fu et al.(2018) Set yogurt Linear (1,3)-β-D-glucan

(94%, Salecan) 0.5% (w/v)Stabilized product and fortified microstructure: improved anti-shear ability, temperaturestability, and water holding capacity, stabilized fat granules, and anchored casein micelles dueto an additional three-dimensional network.

Raikos et al.(2018)

Stirred skim yogurt,reconstituted milk

Spent brewer’s yeast(Saccharomyces cerevisiae)β-glucan, (Yestimun)

0.2–0.8%Reduced fermentations time, formed small spherical β-glucan clusters within the matrix. Didnot change syneresis, viscosity, colour, or acidity, but did increase hardness and adhesiveness.High concentrations showed adverse flavour and aftertaste, otherwise a potential thickener.

Vanegas-Azuero andGutiérrez(2018)

Yogurt, reconstitutedmilk

Ganoderma lucidumβ-glucan (Ganogen) andsacha inchi seeds(Plukenetia volubilis L.)

0-1.5%β-glucan and

4% sachainchi seeds

Did not affect fermentation kinetics. Significantly increased nutritional value. Increase ofβ-glucan content increased WHC and decreased syneresis. Firmness, consistency,cohesiveness and viscosity decreased. Sensorial acceptance >70%.

1 = Water holding capacity, 2 = Lactic acid bacteria, 3 = Not analysed

31

(Raikos et al. 2018), or mushrooms, such as Ganoderma lucidum (Vanegas-Azuero and

Gutiérrez 2018). The added β-glucan concentrations commonly range from 0.5 to 2%, which

is usually enough to get the 1 g β-glucan per portion required for the use of the health claims

(EFSA 2010, 2011).

Molecular aspects, such as molecular weight, ratio of linkages, and concentration highly

affect β-glucan functionality and physicochemical properties in yogurts (Lazaridou and

Biliaderis 2007; Lazaridou et al. 2008). It is believed that the added polysaccharides induce

phase separation of proteins and β-glucan and thus disturb the protein structure and break

the cohesiveness of the gel (Lazaridou et al. 2014; Rinaldi et al. 2015). However, β-glucans

may compensate for their interference by forming an alternative, water entrapping secondary

gel structure within the protein network, with long polysaccharides among casein micelles

or small spherical clusters within the yogurt matrix (Mårtensson et al. 2001; Fu et al. 2018;

Raikos et al. 2018). This secondary three-dimensional network increases the product’s

viscosity and adhesiveness (Sahan et al. 2008). The extent to which the β-glucans improve

or degenerate the yogurt structure seems to largely be dependent on the source of origin,

extraction and purification method, which in turn determine the molecular attributes of the

β-glucan in use (Lazaridou and Biliaderis 2007; Zhu et al. 2016).

32

3 EXPERIMENTAL RESEARCH

3.1 Background and aims

Health effects of β-glucan are believed to be linked with their physicochemical properties,

such as viscosity, which is linked to the concentration and molecular weight of β-glucan

(Regand et al. 2011). Recent research has shown that oxidized lipids may induce the

degradation of β-glucan molecules in aqueous food systems where β-glucan and lipids

co-exist. Wang et al. (2017) designed an oil-in-water emulsion system that contained

β-glucan and oxidized lipids, which they stored for 1 week at room temperature. During this

time, both viscosity of the emulsion, and molecular weight of β-glucan were reported to have

decreased.

This phenomenon needed to be further investigated, which was done as a part of a Business

Finland funded project, OATyourGUT. To further investigate the impact of water, fat, and

the food matrix on β-glucan properties during storage, a yogurt type food model was

designed. Oat β-glucan was to be added to these acid milk gels, with the aim to later link the

molecular level changes of β-glucan to human physiological behaviour and consumption of

oats.

The aim of the experimental part of this Master’s thesis work was to evaluate the impact of

hydrolysis and oxidation of intrinsic oat bran lipids on the functionality of β-glucan in oat

bran concentrate (OBC) ingredients and in acid milk gels (yogurt-type food model) during

storage. The hypothesis was that as the oat ingredients are stored and lipid oxidation is

induced, the β-glucan molecules would be affected and their molecular weight would

decrease, which would then be observed as changes in yogurt texture and stability. The

research questions to be answered were: (1) Does lipid degradation (hydrolysis and

oxidation) affect β-glucan functionality in dry oat ingredients and/or in yogurt models?

(2) Do different oat bran concentrate (OBC) ingredients cause differences in yogurt texture

and stability? This Master’s thesis was conducted as a collaboration between VTT Technical

Research Centre of Finland (VTT) and University of Helsinki (UH) during spring and

summer of 2018.

33

3.2 Overview of the storage trial and yogurt food model experiments

Three types of OBCs with approximately 20% of β-glucan were prepared with different

processing methods in order to induce three levels of lipid degradation. OBC with HIGH

level of lipid degradation (H) was prepared from non-heat treated oat bran; OBC with

MEDIUM lipid degradation (M) was prepared from heat treated oat bran; OBC with LOW

lipid degradation (L) was prepared from heat treated and defatted oat bran. To investigate

the effect of degraded lipids to the degradation and molecular integrity of β-glucan, a storage

trial was conducted: dry OBCs were stored in glass bottles for 0, 1, 2, 4, 8, and 12 weeks at

4, 22, and 40 °C. Separate glass vials with 1 g of sample were prepared and stored for volatile

compound analysis. Lipid degradation states were determined by analysing and following

formed volatile compounds, changes in tocopherol and tocotrienol concentrations, as well

as neutral lipid profiles at UH. At VTT, stored OBCs were used as fibre ingredients in

preparing acid milk gels at each time point of OBC storage test. Chilled acid milk gels were

analysed for spontaneous syneresis, water holding capacity, and texture (deformation test)

the following day from incubation. Spontaneous syneresis and deformation tests were

repeated after refrigerating (4 °C) a set of aliquots for a week, as depicted in Figure 10.

Figure 10. An overview of the experimental research. The experiment was started with lipid analyses (A), afterwhich acid milk gels were prepared (B).

34

3.3 Test materials

3.3.1 Oat bran concentrates (OBCs)

To produce the H sample, oat kernels were dehulled (no heat treatment), milled twice with

a Hosokawa Alpine 100UPZ pin disc mill (17800 rpm) and air-classified using a Minisplit

Air Classifier (British Rema Manufacturing Company Ltd., UK) with classifier wheel speed

of 2900 rpm and air flow of 220 m3/h to separate the bran into the coarse fraction. The coarse

fraction was further milled (2x17800 rpm, 100UPZ pin disc mill) and air classified

(Minisplit Air Classifier, 4500 rpm, 220 m3/h). Powdered concentrate was collected into the

coarse fraction (hereafter referred to as oat bran concentrate or OBC). For M sample, regular

oat bran produced from heat treated oat kernels was milled twice with a pin disc mill

(17800 rpm) and air-classified (Minisplit air Classifier, 3500 rpm, 220 m3/h) to obtain the

OBC in the coarse fraction. For L sample, regular oat bran from heat treated oat kernels was

defatted with supercritical carbon dioxide (SC-CO2) using a Nova Swiss extraction vessel

(Nova Werke AG, Effretikon, Switzerland) with a compressor Chematur Ecoplanning

(Chematur Engineering Ltd., Pori, Finland). Extraction pressure varied between 295 and

305 bar and the temperature in extraction container was 4 °C and in separation container

48 °C. The main points of producing oat bran concentrates with 20% β-glucan can be found

in Figure 11. Additionally, team members at VTT and UH have provided analyses of the

chemical characteristics of the OBC powders, such as β-glucan content, β-glucan molecular

weight (MW), lipase activity and moisture content. β-glucan content was analysed using

Megazyme’s Mixed linkage beta-glucan assay kit (K-BGLU 02/17; McCleary & Codd

1991), β-glucan molecular weight using a high performance size exclusion chromatography

(HP-SEC) method (Suortti 1993), lipase activity according to Yang et al. (2017), and

moisture content by evaporating water from the sample in a 105 °C oven overnight and

counting the moisture content as a percentage based on loss of sample weight. β-glucan

molecular weight was also analysed from fresh and stored (7 days at 4 °C) acid milk gels

with fresh M OBC and M OBC stored at 40 °C for 12 weeks.

35

Figure 11. Enrichment of β-glucan from fresh oat bran. For H and M oat bran concentrates (OBCs), millingand air-classification were performed twice (A and B), before subjecting the H and M OBCs to a storage trial(C). Oat bran for L OBC was milled and air-classified only once (A), since defatting (SC-CO2) the first OBC(β-glucan 15%) was enough to reach target β-glucan concentration before storage (C). Target β-glucanconcentration for all OBCs was 20%.

3.3.2 Reagents and reference materials

All solvents were of HPLC grade and chemicals were of pro-analysis grade and were

purchased from Merck (Darmstadt, Germany), Riedel-de Häen (Seelze, Germany), Sigma-

Aldrich (St. Louis, MI, USA), and Fisher Scientific (Loughborough, UK). Water was

purified by Milli-Q system (Millipore Corp., Bedford, MA, USA) and distilled water was

used for acid milk gel analyses. In acid milk gel preparation, glucono-δ-lactone (GDL)

(Sigma Chemicals, St. Louis, MO, USA) was used as an acidifier, and sodium azide 0.01 %

was prepared as an anti-microbial agent.

To identify and quantify fatty acid methyl esters, GLC-63 mixture (Nu-Check Prep, Inc.,

Elysian, MN, USA) and an internal standard C19:0 methyl ester were used, respectively.

Methylation reagent BF3-CH3OH was purchased from Sigma-Aldrich (St. Louis, MI, USA).

To identify neutral lipid classes, TLC-reference standard 18-6 A (Nu-Check Prep, Inc.,

Elysian, MN, USA) was used. To quantify neutral lipids, calibration mixtures were prepared

in heptane and consisted of tripalmitin for TAGs (>99%, Sigma-Aldrich, St. Louis, MI,

USA), dipalmitin for DAGs (>99%, Nu-Chek Prep, Inc., Elysian, MN, USA) and oleic acid

for FFAs (>99%, Nu-Chek Prep, Inc., Elysian, MN, USA). To identify tocols, tocopherol

reference substances DL-α-, β-, γ-, and δ-Tocopherol (Art. no 15496, stored in a freezer)

were obtained from Merck (Darmstadt, Germany).

To ensure method stabilities, the effectiveness of the ASE extraction and the accurate levels

of tocols and neutral lipids (NP-HPLC methods), in-house references were used and stored

36

in the freezer as follows: NP-HPLC methods, commercial whole wheat flour (Sunnuntai,

Raisio Ltd., Finland), and additionally for tocol analysis, commercial rapeseed oil (used as

an in-house reference in the Food Chemistry research group (UH) since 2011). For the

volatile compounds, an oat bran concentrate with excessive lipid degradation (obtained from

the grain research group (UH)).

3.4 Methods

The oat ingredients (β-glucan enriched OBCs) were stored in different temperatures up to

12 weeks, to study heat-induced lipid degradation (oxidation and hydrolysis). Lipid

degradation was analysed chemically by following three indicators: increase in specific

volatile compounds, tocopherol and tocotrienol degradation and the formation of free fatty

acid. Figure 12 presents an overview of the lipid degradation experiments.

Figure 12. A summary of the experimental design and chosen methods for lipid degradation analysis. a-T3 =α-tocotrienol, FFA = free fatty acids.

3.4.1 Extraction of lipids and analysis of total FA content

Lipid contents and fatty acid compositions of the OBC samples were determined by a gas

chromatographic (GC) method used for fatty acid methyl ester analysis (Lampi et al. 2015).

Lipids were first extracted from OBC samples with acetone (≥99.8% Sigma-Aldrich, St.

Louis, MI, USA) using an accelerated solvent extraction method (ASE) (Dionex ASE 200

Accelerated Solvent Extraction system, Thermo Fisher Scientific). The specifics of the

ASE method were: temperature 100 °C, pressure 1000 psi, heating time 5 min, two times

static cycle time 10 min, flush volume 60%, nitrogen gas purge time 60 s, and rinsing time

1 min. For the extraction, approximately 1 g of the sample was precisely weighed into the

ASE-extraction cell together with an equal amount of drying agent (Sand, General purpose

37

grade, Thermo Fisher Scientific). The cell was then filled with the drying agent, and before

closing the extraction cell, 1 ml of ethanol (≥99.5%, Altia, Rajamäki, Finland) was added on

top of the sand for better extraction of lipids. Lipid extracts were moved to 10 ml KIMAX

Test Tubes with Teflon Liner Caps. 5 mg of C19:0 in heptane was added in each extract as

an internal standard (ISTD). The extracts were transferred quantitatively to 50–100 ml

round-bottom flasks with ethanol, which also aided in evaporating trace water. The acetone

and ethanol were evaporated in a rotary evaporator at ≤37 °C, and redissolved in

3:2 heptane:isopropanol (heptane ≥98%, Merck, isopropanol ≥99.9%, Riedel-de Hän).

For methylation of the extracted lipids, the solvent was first evaporated at ≤40 °C under a

nitrogen flow. Then 1 ml of 0.5 M NaOH in methanol was added to each test tube, and

carefully closed tubes were placed in boiling hot water for 5 minutes. Once the tubes cooled

down, 2 ml of methylation reagent BF3-CH3OH (~10% in methanol, Sigma-Aldrich, St.

Louis, MI, USA) was added and the tubes were heated in boiling hot water for another

5 minutes. 5 ml of heptane and 2 ml of supersaturated NaCl solution were added to cooled-

down extracts, after which the extracts were agitated vigorously for approximately 1 minute.

Using glass Pasteur-pipettes, the upper heptane layers were transferred to clean KIMAX

tubes with some drying agent (NasSO4) at the bottom. After a minimum of 30 minutes,

heptane extracts were loaded to vials for GC analysis as described by Soupas et al. (2005)

and Damerau et al. (2014): The equipment used was an Agilent 6850 Network GC system

coupled with a flame ionization detector (FID). The column in use was a fused-silica

capillary column (Omegawax 250: 30m x 0.25 mm, 0.25 μm, Supelco, Bellefonte, PA,

USA). Helium acted as the carrier gas at a flow of 1 ml/min. The oven temperature program

started at 160 °C and was held there for 1 min, after which the temperature was raised

4 °C/min to 240 °C, and was finally held there for 5 min. The detector temperature was

250 °C, with detector gases helium and air. Injector temperature was 240 °C, and injection

was performed with a split ratio of 1:28 (split injection technique). Agilent ChemStation was

used to operate the instrument, sampling and in result handling.

The identification of fatty acid methyl esters was done based on a standard GLC-63 mixture

and the quantification was conducted using C19:0 methyl ester. Total lipid contents were

calculated as sums of the major fatty acid methyl esters, i.e. C14:0, C16:0, C18:0,

C18:1 (n-9), C18:1 (n-7), C18:2 (n-6), C18:3 (n-3), and C20:1 (n-9), and reported as mg/g

sample on fresh weight basis. Experiments were performed as triplicates. Coefficients of

variation for fatty acid methyl esters were 4 % or lower.

38

3.4.2 Analysis of neutral lipid profiles with NP-HPLC-ELSD

Lipids were extracted from 1.0 g of sample with acetone as described in Section 3.4.1

Extraction of lipids and analysis of total FA content, except no ISTD was used and the

redissolving in 10 ml was performed with HPLC-grade heptane only. After redissolving, the

extracts were further diluted 1:10 in heptane and placed in HPLC vials. To analyse neutral

lipid profiles of the oat ingredients, a method described by Lampi et al. (2015) was used: a

normal-phase HPLC (Agilent 1200 HPLC system, Agilent Technologies, Santa Clara, CA,

USA) with an evaporative light scanning detector (ELSD) (Waters 2420 ELSD, Waters®,

Milford, MA, USA) was used to analyse neutral lipid classes (triacyl, diacyl and monoacyl

glycerols and free fatty acids; TAGs, DAGs, MAGs, FFAs, respectively). A LiChrosorb diol

column (5 μm, 3 x 100 mm, VDS optilab Chromatographie Technik GmbH, Berlin,

Germany) was used to separate neutral lipid classes. A linear gradient elution was a mixture

of heptane and 0.1% acetic acid and an increasing proportion of isopropanol (from 0.09%

isopropanol from 0–5 min, to 3% from 5–15 min, at 3% from 15–35 min, and back to 0.09%

from 35–40 min) with a flow rate of 0.5 ml/min at 25 °C. The stabilisation time between runs

was 10 minute. The ELSD was set to drift tube temperature 60 °C, nebuliser to 42 °C and

the gain to 10. Filtered air was used for nebulisation at a flow rate of 1.4 l/min (i.e. 20 psi).

Quantification was performed by using an external standard method. Calibration mixtures

were prepared in heptane and consisted of tripalmitin, dipalmitin, and oleic acid, as

explained in Section 3.3.2. Calibration mixtures were analysed both at the beginning and the

end of each sequence at six amounts in the following ranges: 100–4000 ng/injection for

TAGs and FFAs, and 100–5000 ng/injection for DAGs. Calibration curves were computed

with a second order regression separately for the three acylglycerol classes and oleic acid.

The regression coefficients for calibration curves were ≥ 0.95. Each OBC lipid extract was

injected in two volumes, 5.0 and 50.0 μl, to meet the calibration ranges. Experiments were

performed as triplicates and the results were reported as mg/g sample on fresh weight basis.

To ensure the performance of the equipment and the stability of the HPLC method, a mixture

of oleyls (TLC-reference standard 18-6 A) was analysed with each HPLC sequence. Oleyls’

retention times remained stable throughout the experiment. To ensure the stability of the

whole method, starting from lipid extraction, a whole wheat flour (Sunnuntai, Raisio Ltd.,

Finland) was used as an in-house reference. Neutral lipid amounts in the in-house reference

stayed stable throughout the study: TAGs remained at 6.37 ± 0.45 mg/g sample, FFAs at

39

4.32 ± 0.26 mg/g sample, and DAGs at 0.78 ± 0.35 mg/g sample, with a total of

11.47 ± 0.51 mg/g sample.

3.4.3 Analysis of tocopherol and tocotrienol contents with NP-HPLC-FLD

Tocopherol and tocotrienol contents were analysed with a normal phase high performance

liquid chromatography method with a fluorescence detector (NP-HPLC-FLD) as described

by Schwartz et al. (2008): Lipids were extracted from 1.0 g of sample with acetone as

described in Section 3.4.1, except no ISTD was used and the redissolving in 10 ml was

performed with HPLC-grade heptane only. Approximately 1 ml of lipid extracts were then

filtered (Acrodisc GHP, 0.54 μm, 13 mm) into HPLC vials, ready to be analysed. To ensure

the performance of the HPLC equipment, rapeseed oil (50 mg in 10 ml heptane) was used

as an in-house reference.

Standard stock solutions were prepared from tocopherol reference substances DL-α-, β-, γ-,

and δ-tocopherol (Art. no 15496, Merck, Darmstadt, Germany, stored in a freezer) to a

concentration of approximately 500 mg/l in ethanol (AAS) and stored at -20 °C. Suitable

amount of each tocopherol were combined and diluted with heptane to create a reference

mixture with 1.5-2 mg/l of each tocopherol. Exact tocopherol concentrations were confirmed

spectrophotometrically (Ryynänen et al. 2004) using reported absorption coefficients for

each vitamer in ethanol E1% at a length of 1 cm. Tocotrienol vitamers were quantified with

each respective tocopherol (Ryynänen et al. 2004).

The HPLC-system consisted of a Waters 515 HPLC pump (Waters Corporation, Milford,

MA, USA), a Waters 717 plus Autosampler, a Guard-Pak Inserts Resolve Silica precolumn

(Waters), a Varian Inertsil 5 SI column (250 x 4.6 mm, 5 μm particle size, Varian Inc., Palo

Alto, CA, USA) and a Waters scanning fluorescence detector model 474. The system

software was Millenium 32 software (Version 4.00, Copyright® 2001 Waters Corporation).

Separation of tocols took 25 minutes, they were separated isocratically using a mobile phase

of 3% 1,4-dioxane and 97% heptane (v/v) with a flow rate of 2 ml/min, and detected

fluorometrically (λex: 290 nm, λem: 325 nm) (Kamal-Eldin et al. 2000). Dissolved air in the

mobile phase was removed first by sonication before analysis and by an online-helium

degasser. The column oven was set at 30 °C.

40

Tocopherols were identified by comparing retention times to those of the standards. The

elution order of the tocopherols and tocotrienols was α-tocopherol first, which was followed

by α-tocotrienol, β-tocopherol and β-tocotrienol, having the polar column retain more polar

molecules for a longer time. Quantification was performed by using external calibration

functions that were established by combining 2x7 calibration points obtained by injecting

tocopherol standard mixture at different volumes (2–100 μl) at the beginning and the end of

each sequence. Quantification of each tocotrienol was done by using the corresponding

tocopherol calibration functions (Ryynänen et al. 2004), and reported as μg/g sample on

fresh weight basis. Experiments were performed as triplicates. Regression coefficients for

the calibration curves were all >0.99.

To ensure the stability of the whole method, starting from lipid extraction, a commercial

whole wheat flour (Sunnuntai, Raisio Ltd., Finland) was used as an in-house reference.

Tocopherol and tocotrienol amounts in the in-house reference stayed stable throughout the

study: a-T remained at 6.95 ± 0.32 μg/g, a-T3 at 3.11 ± 0.12 μg/g, b-T at 4.37 ± 0.11 μg/g,

and b-T3 at 17.45 ± 0.47 μg/g. The tocopherol amounts in the rape seed oil remained within

the standard deviations of the in-house data since 2011.

3.4.4 Analysis of volatile compounds with HS-SPME-GC-MS

Six volatiles typical for lipid oxidation were followed, and changes in concentrations

recorded. The volatiles were, in retention order, 2-ethylfuran, 1-pentanol, hexanal,

2-pentlyfuran, benzaldehyde, and nonanal. Volatiles were measured with a headspace solid

phase microextraction gas chromatography method coupled with a mass spectrometer

detector (HS-SPME-GC-MS) (GC Perkin Elmer, USA) using 1 g (0.99-1.01 g) of the dry

OBC samples in each 20 ml amber headspace vial in triplicates for each time point. Tightly

sealed vials were stored at different temperatures (4, 22, 40 °C) for varying amounts of time

(0–12 weeks) to induce an array of lipid degradation levels. Volatile compounds were

analysed using an HS-SPME injector (combiPAL, CTC Analytics, Zwingen, Switzerland)

with a CAR/PDMS-fibre (85 μm film thickness; Supelco, Bellefonte, PA, USA) together

with a GC (HP 6890 series, Agilent Technologies Inc.), including a capillary column SPB-

624 (30 m x 0.25 mm i.d., 1.4 μm film thickness; Supelco, Bellefonte, PA, USA), coupled

to an MS detector (Agilent 5973 Network, Agilent Technologies Inc.). MSD Chemstation

(Agilent Technologies Inc.) was used for data processing.

41

The analysis was performed as reported earlier by Damerau et al. (2014): Samples were

incubated and extracted for 20 min at 50 °C with an agitation speed of 250 rpm to equilibrate

the volatile compounds between the solid and gas phase. Volatile compounds were identified

by matching their total ion mass spectra (m/z from 50 to 300) with the database Wiley 7N

(Wiley Registry TM of Mass Spectral Data, 7th Edition, Hoboken, NJ, USA) and by

comparing retention times and mass spectra to previously acquired data on oat volatile

compounds.

To ensure the stability of the HS-SPME-GC-MS method, the peak areas of five volatiles

were chosen and followed from an in-house-reference (freezer-stored OBC) in each

sequence, however, the freezer-stored OBC was old and unstable, and did not perform in a

repeatable manner. To monitor the performance of the equipment, air and water checks were

performed daily, and tune evaluation monthly. Experiments were performed as triplicates

and the results are presented as peak areas. The variation coefficients of the peak areas

ranged between 2.5 and 10% within a sequence.

3.4.5 Preparation of acid milk gels

Figure 13. The stages of preparing acid milk gels with added β-glucan (oat bran concentrates).

The preparation and acidification of the acid milk gels was performed as described by

Ercili-Cura et al. (2013) and Rosa-Sibakov et al. (2016) (overview in Figure 13).

Acidified milk gels were prepared in 1000 ml glass bottles with sealable caps to contain

9.57% (w/w) milk solids, 3.3% protein (w/w) and 1 g of β-glucan per portion (portion chosen

was 150 g of yogurt) to achieve the level of β-glucan specified in the health claims (EFSA

2010, 2011) and minimize the added dry matter. This meant an addition of 2.8–3.6% (w/w)

42

of OBC ranging with the assessed β-glucan levels of each sample. The OBC samples, each

with different levels of lipid degradation (stored in varying conditions: 4, 22, or 40 °C for

0–12 weeks), were mixed with distilled water by constant magnetic stirring (190 rpm) at

50 °C for 30 minutes. Skimmed milk powder (34.5% protein, 53% lactose, 8% ash, 3.8%

moisture, and 0.5% fat) (Valio Ltd., Helsinki, Finland) was reconstituted by adding the milk

powder to the OBC-water mixture, and mixed for another 30 minutes by constant magnetic

stirring (190 rpm) at 50 °C. The new mixture (reconstituted skim milk + OBC) was heat

treated by heating the sample ≥86 °C for 5 minutes and cooled down in ice to 40 °C. Sodium

azide (0.01% (w/w)) was added to an aliquot to prevent microbial growth in gels that were

to be stored for 1 week. The total volume of each acid milk gel sample prepared was 500 ml.

The acidification was started by adding 1.3% (w/w) glucono-δ-lactone (GDL) (Sigma

Chemicals, St. Louis, MO, USA) at a constant dosage to the mixture at 40 °C. The mixtures

were then stirred for 10 minutes with magnetic stirring (250 rpm) until GDL was totally

dissolved. Control samples were prepared in a similar manner without the added β-glucan.

The gels were then divided into 40 or 20 g aliquots in tightly sealed plastic cups (ø 3.5 cm)

or sealable 50 ml centrifuge tubes, respectively. Incubation was performed at 40 °C for

approximately 4.5 hours or until the pH reached 4.5–4.6. The acidification process and

changes in pH were recorded during incubation at 10 minute intervals. Each acid milk batch

produced four replicate samples for fresh gels, four replicate samples for stored gels, and

three replicate samples for water holding capacity.

3.4.6 Analysis of spontaneous syneresis and water holding capacity

The water retention ability of an acid milk gel depends on the microstructure of the gel that,

in turn, depends on the acidification temperature and pH combination (Ercili-Cura et al.

2013). Water retention can be assessed by measuring spontaneous syneresis, commonly

thought to be a product defect among consumers, and water holding capacity (WHC) of the

gels. Measuring spontaneous syneresis provides insight into the acceptability of a

prospective yogurt-type product.

After the gels (40 g) had reached the targeted pH (4.5-4.6) and had cooled down in the fridge

overnight, the gels were placed at room temperature for 10 min to condition. The water

released on top of the gel was poured out carefully and weighed. Spontaneous syneresis

43

analyses were performed in quadruplicates, and expressed as the percentage of the amount

of water released on top of the gel in relation to the initial weight of the gel.

For the WHC analysis, the formation of acid milk gels was performed in 50 ml centrifuge

tubes as previously described by Ercili-Cura et al. (2013): 20 g of sample was acidified in

the centrifuge tubes at 40 °C until pH arrived at 4.5–4.6. The gels were centrifuged at 3000g

for 15 minutes at 4 °C. Supernatant was poured out carefully and weighed. WHC analyses

were performed in triplicates, and expressed as the percentage of the weight of the pellet in

relation to the initial weight of the gel.

3.4.7 Analysis of gel firmness with TA

The firmness of the acid milk gels was analysed as previously reported by Ercili Cura et al.

(2009) and Rosa-Sibakov et al. (2016). A large deformation test was done using a

TA.XTPlus Texture Analyser (TA, Stable Microsystems Ltd., Godalming, UK) equipped

with a 5 kg load cell. The samples (40 g) were incubated (40 °C) in plastic cups (ø 3.5 cm)

until pH was 4.5–4.6, after which the gels were refrigerated overnight or for 1 week. Before

the deformation test, the chilled gels were left at room temperature for 10 minutes. Gel

deformation was done by penetrating the gel with a ø 25 mm cylinder probe at a constant

speed of 1.0 mm/s to a distance of 75% of the sample height. Deformation tests were

performed on three to four individual gels for each gel batch. The area under the force vs.

distance curve (up to 15 mm) was determined and taken as a measure of gel firmness. The

reported values are the means of two acid milk gel batches.

3.5 Results

3.5.1 OBC characteristics

The three OBCs contained 18–23% β-glucan (Table 2). OBC samples showed differences in

lipid degradation levels ranging from no degradation to higher levels of degradation. H OBC,

which was not heat treated, had the highest content of total fatty acids, 9.0±1.2%. M OBC,

which was heat treated, was not far off, and had nearly as much fatty acids as H OBC,

7.8±1.0%. L OBC was defatted, and had 1.3±0.2% of fatty acids left. H OBC was the only

oat ingredient to exhibit lipase activity. The MW of β-glucan in all OBCs did not change and

44

remained at >1000 kDa during the storage time. The MW of β-glucan remained >1000 kDa

also in fresh and stored (7 days at 4 °C) acid milk gels with fresh M OBC and M OBC stored

at 40 °C for 12 weeks.

Table 2. β-glucan, fat, and moisture contents of differently processed oat bran concentrates (OBCs)and their lipase activities.

Sample β-glucan(%, dm)

Total FA2

(%)Moisture(%)

Lipase activity(μmol/min/g)

Non-heat treated(H OBC) 18.1±0.4 9.0±1.2 5.7 0.9±0.05

Heat treated(M OBC) 23.2±0.3 7.8±1.0 4.7 0

Heat treated and defatted(L OBC) 19.9±0.2 1.3±0.2 5.2 NA1

1NA = not analysed, 2FA = fatty acid

3.5.2 Lipid degradation

Neutral lipids

The FFAs were eluted as two peaks, which were added together for the total FFA count.

H OBC was the only sample of the three OBCs, where FFAs were formed. When stored at

4 °C, FFAs started to form only after 4 weeks, whereas at the higher temperatures, FFAs

could be seen already at 2 weeks. The FFA amounts increased with time at all the storage

temperatures: at the slowest rate at 4 °C, steadily at 22 °C, and at the fastest rate at 40 °C

(Figure 14). The highest amount of FFAs formed after 12 weeks at 40 °C was

3.8 mg/g sample, which was over 2.5 times more than for the OBC stored at 4 °C (1.5 mg/g).

The total amount of neutral lipids decreased at a similar rate at all storage temperatures

(Figure 15). The share of TAGs to the amount of total neutral lipids was the greatest for

samples stored at 4 °C (91.6%), and the lowest for samples stored at 40 °C (83.7%). As in

compensation, there were more DAGs and FFAs at higher storage temperatures.

45

Figure 14. Free fatty acids (FFA) were formed during a storage trial of a non-heat treated oat bran concentrate(H OBC) with a β-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. H OBC was stored at 4, 22, and40 °C for up to 12 weeks. N = 3.

Figure 15. Neutral lipid classes (triacyl glycerols, diacyl glycerols, and free fatty acids; TAG, DAG, FFA,respectively) were observed in a storage trial of a non-heat treated oat bran concentrate (H OBC) with aβ-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. H OBC was stored at (a) 4 °C, (b) 22 °C, and(c) 40 °C for up to 12 weeks. N=3.

M OBC had less neutral lipids to start with, 49.8±0.8 mg/g sample, when compared to

H OBC’s 58.4±0.3 mg/g sample. A trend of slight decrease of TAGs, increase of DAGs, and

46

decrease of total neutral lipids is visible as time and storage temperature increase (Figure 16).

Fresh L OBC had only possible traces of neutral lipids (data not shown), and thus neutral

lipid classes were not followed during the storage trial.

Figure 16. Neutral lipid classes (triacyl glycerols, diacyl glycerols, and free fatty acids; TAG, DAG, FFA,respectively) were observed in a storage trial of a heat treated oat bran concentrate (M OBC) with a β-glucancontent of 23.2±0.3% and a fat content of 7.8±1.0%. M OBC was stored at (a) 4 °C, (b) 22 °C, and (c) 40 °Cfor up to 12 weeks. N=3.

Tocopherols and tocotrienols

H OBC had the highest amounts of tocopherols and tocotrienols (hereafter referred as tocols)

compared to M OBC and L OBCs. The amounts of tocols in fresh H OBC were

7.5±0.08 μg/g sample for a-T, 66.7±0.9 μg/g sample for a-T3, 0.9±0.01 μg/g sample for b-T,

and 8.2±0.2 μg/g sample for b-T3 (Figure 17). All the tocopherols degraded during time, and

more intensively with increased temperature: the final amounts for a-T, b-T, and b-T3 neared

zero at all temperatures, and a-T3 persisted at a range of 36.7±0.3 to 12.7±0.2 μg/g for

samples stored at 4 and 40 °C, respectively.

47

Figure 17. α-tocopherol (alpha), α-tocotrienol (aT3), β-tocopherol (beta), and β-tocotrienol (b3T)concentration levels were observed during a storage trial of a non-heat treated oat bran concentrate (H OBC)with a β-glucan content of 18.1±0.4% and a fat content of 9.0±1.2%. H OBC was stored at (a) 4 °C, (b) 22 °C,and (c) 40 °C for up to 12 weeks. N = 3.

Tocols in M OBC decreased with increased time and temperature (Figure 18) in a similar

manner as in H OBC. Higher temperature also accelerated the degradation of all tocols.

L OBC had only possible traces of tocols (data not shown), and thus were not followed for

during the storage trial.

48

Figure 18. α-tocopherol (alpha), α-tocotrienol (aT3), β-tocopherol (beta), and β-tocotrienol (b3T)concentration levels were observed during a storage trial of a heat treated oat bran concentrate (M OBC) witha β-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. M OBC was stored at (a) 4 °C, (b) 22 °C, and(c) 40 °C for up to 12 weeks. N = 3.

Volatile compounds

H OBC showed resilience towards storage and heat. There were no detectable volatile

compounds typical for lipid oxidation in samples stored at 4 or 22 °C during 12 weeks. Some

enzyme activity related 2-pentylfuran could be detected in samples stored at 40 °C, however,

in a small measure (Figure 19).

49

Figure 19. 2-pentylfuran formation was observed in a storage trial of a non-heat treated oat bran concentrate(H OBC) with a β-glucan content of 18.1±0.4% and a fat content of 9.0±1.2%. H OBC was stored at 40 °C forup to 12 weeks. N = 3.

M OBC portrayed more typical volatile profiles for heat treated and stored oat ingredients.

Autoxidation product hexanal was the dominant volatile compound together with a little

nonanal. Hexanal amounts increased steadily during 8 weeks of storage until it decreased

towards week 12. The higher the storage temperature was, the more there was hexanal

(Figure 20).

Figure 20. Hexanal formation was observed in a storage trial of a heat treated oat bran concentrate (M OBC)with a β-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. M OBC was stored at 4, 22, and 40 °Cfor up to 12 weeks. N = 3.

Similarly, nonanal peaked at 8 weeks, however, the highest amounts of nonanal seemed to

be formed at lower temperatures (Figure 21). The samples stored in 40 °C showed also some

formation of 2-pentylfuran (Figure 22), approximately double the amount found in H OBC

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(Figure 19). All L OBC samples formed traces of hexanal, possibly showing slightly higher

amounts in samples stored at 40 °C.

Figure 21. Nonanal formation was observed in a storage trial of a heat treated oat bran concentrate (M OBC)with a β-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. M OBC was stored at 4, 22, and 40 °Cfor up to 12 weeks. N = 3.

Figure 22. 2-pentylfuran formation was observed in a storage trial of a heat treated oat bran concentrate(M OBC) with a β-glucan content of 23.2±0.3% and a fat content of 7.8±1.0%. M OBC was stored at 4, 22,and 40 °C for up to 12 weeks. N = 3.

As H OBC showed stability to elevated temperatures and time, and produced only little

amounts of volatiles, an extra experiment was performed in order to understand the

phenomena and exclude the possibility of an error in the sampling. Raw materials for H and

M OBC were subjected to a similar storage test as for the OBCs storage conditions being 8

weeks at 40 °C, and then analysed for volatile compounds. For H OBC, milled non-heat

treated oat kernels (H RAW), i.e. material used before obtaining oat bran and before the air-

51

classification procedures (Figure 11) to produce H OBC, was used. For M OBC, the basic

heat treated commercial type OBC (M RAW, β-glucan content ~15%), was used. Some

2-pentylfuran was formed in H RAW at 4 and 8 weeks of storage, which counted for double

the amount H OBC had at its highest at 8 weeks. Additionally H RAW showed signs of

oxidation as there was some formation of hexanal at weeks 4 and 8, although low in amounts

(Figure 23a). M RAW was more stable than M OBC, and showed some hexanal and

2-pentylfuran formation (Figure 23 b). However, it should be noted that e.g. the

concentration of hexanal was minor, four times smaller than the amounts formed in M OBC

stored at 40 °C. The main finding from analysing the RAW materials was their different

profiles with different volatile compounds.

Figure 23. Volatile compounds were observed in a storage trial of (a) a non-heat treated oat flour and (b) heattreated oat bran concentrate (β-glucan content ~15%), both raw materials for oat bran concentrates (H OBCand M OBC). Samples were stored at 40 °C for up to 8 weeks. N = 3.

3.5.3 Properties of acid milk gels

Acidification

Control acid milk gels (without added OBC) had initial pH values around 6.5, and were

acidified to pH 4.6 in 4–5 hours as shown in the acidification curve in Figure 24. When

H OBC was added to milk gels, the acidification time was longer and it would have taken

over 20 h to get close to pH 4.6 (Figure 24). For practical reasons, a pH value of 4.6±1.0

with an acidifying time of 5-6 h was settled upon. M and L OBCs arrived to the target pH in

a shorter time, and after approximately 5 h of acidifying, the pH was 4.5-4.6. The effect of

storage of the OBCs on acidification times was most pronounced in acid milk gels with

H OBC. For example, when using fresh H OBC, it took just over 6 hours for the pH to reach

4.7. However, when H OBC was stored 12 weeks at 40 °C and then used as an ingredient,

in the same amount of time, the pH reached 4.6 (Figure 25, shown with arrows).

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Figure 24. Acidification of acid milk gels without (control) or with fresh oat bran concentrates (OBC). H OBCwas non-heat treated, M OBC heat treated, and L OBC heat treated and defatted.

Figure 25. Acidification of acid milk gels with added non-heat treated oat bran concentrate (H OBC). H OBCwas stored in dark at 40 °C for 0–12 weeks prior to use as an ingredient in the acid milk gels. Storage of oatingredients shortened the acidification time, indicated with arrows.

Spontaneous syneresis and water holding capacity (WHC)

Control acid milk gels (without added OBC) had 9±3% spontaneous syneresis (Figure 26c),

and a water holding capacity of 39±10% (Figure 27c). None of the gels with added OBCs

showed spontaneous syneresis (Figure 26ab). When gels with M and L OBCs were subjected

53

to centrifuging, water retention was a perfect 100% (Figure 27ab). However, the water

holding capacity of gels with fresh and stored H OBCs were not far and ranged between 96%

and 99% (Table 3).

Figure 26. Acid milk gels (40 g) (a, b) with and (c) without added oat bran concentrates (OBCs). The additionof OBCs improved the water retention properties of the gels leading to having gels with no visible spontaneoussyneresis on top of the gels (a, b).

Figure 27. Acid milk gels (20 g) (a, b) with and (c) without added oat bran concentrates (OBCs) were placedin individual 50 ml tubes and centrifuged in 3000g at 4 °C for 15 min. The addition of OBCs substantiallyimproved the water retention properties of the gels: from (c) 39% to (a, b) 96–100%.

Table 3. Water holding capacity (WHC) of acid milk gels (a) without and (b) with added oat fibre. Added oatfibre was a non-heat treated oat bran concentrate (H OBC) with a β-glucan content of 23.2±0.3% and a fatcontent of 7.8±1.0%. H OBC was stored at 4, 22, and 40 °C for up to 12 weeks before being used as aningredient in the acid milk gels. N =3. 1NA = not applicable.

SampleStorage

temperature(°C)

WHC (%)

0 wks 2 wks 4 wks 8 wks 12 wks

a) Control NA1 38.8±10.4 NA1 NA1 NA1 NA1

b) H OBC(non-heattreated)

4 95.8±3.74 98.1±0.17 99.4±0.07 99.3±0.08 98.0±0.54

22 95.8±3.75 96.4±1.12 99.3±0.02 99.3±0.06 98.9±0.15

40 95.8±3.76 98.0±0.19 99.5±0.06 99.7±0.27 97.2±0.23

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Firmness of acid milk gels

Control acid milk gels (without added OBC) were firm but brittle, and would immediately

release serum when deformed. Acid milk gels with OBCs were softer and smoother than

control gels (Figure 28a). Small oat specs could be seen in all OBC gels, however, having

H OBC as an ingredient seemed to produce the grainiest gels. The control sample had the

largest area under the force vs. distance curve, 15.3±1.7 N.mm, indicating a very firm acid

milk gel. Compared to the control, the gels with fresh OBCs were not as firm: 5.1±0.4,

7.9±4.2, and 5.2±1.1 N.mm for H, M and L OBC gels, respectively. All gels with added

fresh OBCs were stored 7 days at 4 °C. After this storage time in the fridge, gels with H OBC

were softer than they were as freshly prepared gels, gels with M OBC were similar to what

they were as freshly prepared gels, and gels with L OBC were harder than what they were

as freshly prepared gels. An example of the force vs. distance curve is provided in Appendix

1.

Figure 28. The firmness of acid milk gels changed depending on the types of oat bran concentrates (OBCs):(a) a fresh gel with fresh OBC, (b) a fresh gel with OBC stored for 12 weeks at 22 °C, and (c) a gel stored at4 °C for 7 days with OBC stored for 12 weeks at 22 °C. Control sample was prepared without added OBC.H OBC was non-heat treated, M OBC heat treated, and L OBC heat treated and defatted.

All H OBC gels were firmer than or at the same firmness level as gels with fresh H OBC

(week 0), regardless of the temperature H OBCs were stored at (Figure 29). Between 2 and

12 weeks of storage of H OBCs, gels with H OBCs stored both at 4 and 40 °C decreased in

firmness. There were no differences between gels of weeks 2 and 12 with H OBCs stored at

22 °C (Figure 29). Gels stored for 7 days at 4 °C were generally firmer than fresh gels (Figure

28bc (H OBC) and Figure 29).

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Figure 29. Firmness of acid milk gels with a non-heat treated oat bran concentrate (H OBC, 18.1±0.4%β-glucan and 9.0±1.2% fatty acids). H OBC was stored at 4, 22, and 40 °C for up to 12 weeks, before used inacid milk gels. A control sample without added fibre was also prepared (on the left). N = 4.

M OBC gels gained firmness after M OBCs were stored 8 weeks at any storage temperature,

only to decrease in firmness with ingredients stored for 12 weeks (Figure 30). Although,

standard deviations were notably high. For gels with M OBC, it was not clear whether storing

gels for 7 days at 4 °C had an increasing or decreasing effect on gel firmness compared to

freshly prepared gels (Figure 28bc (M OBC) and Figure 30).

Figure 30. Firmness of acid milk gels with a non-heat treated oat bran concentrate (M OBC, 23.2±0.3%β-glucan and 7.8±1.0% fatty acids). M OBC was stored at 4, 22, and 40 °C for up to 12 weeks, before used inacid milk gels. A control sample without added fibre was also prepared (on the left). N = 2x4.

L OBC gels were generally the firmest of all the OBC gels, having the largest areas under

the force vs. distance curve. However, behaviour of gel firmness over the weeks varied

depending on the storage temperatures of L OBC: Gels with L OBC stored at 4 °C were the

firmest at week 8, decreasing towards week 12. Gels with L OBC stored at 40 °C grew firmer

week by week, and gels with L OBC stored at 22 °C showed no differences in firmness

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(Figure 31). The effect of the 7 day’s storage of gels at 4 °C was most pronounced in L OBC

gels, especially at weeks 8 and/or 12 at 4 and 22 °C (Figure 28bc (L OBC) and Figure 31).

Although L OBC gels showed increasing likeness to the firmness of the control gel, L OBC

gels continued to be short, smooth and pleasant.

Figure 31. Firmness of acid milk gels with a non-heat treated oat bran concentrate (L OBC, 19.9±0.2%β-glucan and 1.3±0.2% fatty acids). L OBC was stored at 4, 22, and 40 °C for up to 12 weeks, before used inacid milk gels. A control sample without added fibre was also prepared (on the left). N = 2x4.

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3.6 Discussion

3.6.1 Lipid degradation behaviour of OBC samples

To fully understand the effect degraded lipids have on β-glucan functionality, the different

pre-treatments (no heat treatment (H OBC), heat treatment (M OBC), no heat treatment and

defatting (L OBC)) as well as storage temperatures (4, 22, 40 °C) were chosen to create a

range of oat ingredients with different stages of lipid hydrolysis and oxidation. Regarding

different OBCs, it was hypothesised that H OBC would represent a rancid material with both

hydrolysis and oxidation products, M OBC would be a commercial industry-type oat bran

concentrate with indications of lipid oxidation in long-term storage, and due to defatting,

L OBC would have very little, if any, indications of lipid degradation. Lipid extraction

performed on L OBC was efficient and the ingredient behaved according to the hypothesis:

the changes in the lipid phase were not detectable due to low lipid concentrations. This is

why this section will concentrate on the results of H and M OBCs.

Based on previous studies, storage temperature was known to affect lipid degradation

kinetics, and it was hypothesised that 4 °C would slow down and 40 °C would accelerate the

degradation reactions (Kaur et al. 2018). The effect of storage temperature and its impact on

reaction kinetics (Labuza 1971) could be seen in the results of both H and M OBC. The

accelerating effect of higher storage temperatures on the extent of lipid degradation was

emphasized in, for example, the formation of FFA in H OBC (Figure 14) and hexanal in

M OBC (Figure 20), as well as in the degradation of tocols in both samples (Figures 17 and

18). The temperature most probably also provided favourable conditions for enzyme activity,

as samples stored at 40 °C quickly formed FFAs. However, this formation rate did not stay

as high as in the beginning, which could also be attributed to higher temperatures. FFAs

might have reacted further at a faster rate than they were formed, appearing as a slowed-

down formation rate.

H OBC had a higher total FA content than M OBC (Table 2), which probably resulted from

M OBC having a slightly higher β-glucan content. During storage, total neutral lipids

decreased in H OBC (Figure 15). TAGs were degraded to DAGs and FFAs, but as DAGs

and FFAs would react further at a faster rate than TAGs were degraded, the total amount of

neutral lipids would slowly decrease. A higher storage temperature increased the relative

58

portion of DAGs and FFAs in the samples, which is probably related to increased enzyme

activity at 40 °C.

The presence or absence of a heat treatment expectedly affected lipase activity, and thus, the

presence of hydrolytic products. H OBC was not heat treated and was the only OBC to have

its lipase still active (Table 2), which explains why it was the only OBC ingredient to form

FFAs. However, only little enzyme activity related volatiles, such as 2-pentylfuran, were

found (Figure 19), even in the stored raw material (Figure 23). The heat treatment also

affected the amounts of tocols. α-tocotrienol was the most abundant tocol in both samples,

which is typical for an oat bran ingredient (Bryngelsson et al. 2002). However, H OBC had

significantly higher concentrations of α-tocotrienol compared to M OBC both at the

beginning and at the end of the storage time (Figures 17 and 18). M OBC was heat treated,

which could explain the concentration difference, as tocols are reported to be sensitive to

heat during processing (Bryngelsson et al. 2002).

Quite surprisingly, H OBC did not show indications of lipid oxidation. Even after the

extended storage period, oxidation products, such as hexanal, in H OBC were scarce. The

extraction fibre in use could have affected the extracted volatile compounds, as previous

studies with volatiles related to oat lipid oxidation used DVB/CAR/PDMS-fibres (e.g.

Damerau et al. 2014). However, using a CAR/PDMS-fibre should not have adversely

affected volatile profiles, but either had a potentially neutral or improving effect on the

volatile extraction process (Poinot et al. 2007, Raffo et al. 2015). Furthermore, the same

extraction fibre was used in the analysis of other OBC samples and volatiles were detected

in M OBC, which supports the use of CAR/PDMS-fibre. To be sure, more studies on the

impact of DVB/CAR/PDMS- and CAR/PDMS- fibres on the extraction of oat volatile

compounds should be conducted.

The overall lipid stability of H OBC in storage was still a puzzle, as usually non-heat treated

oats are thought to become rancid quicker because of active enzymes, giving reason to

inactivate the enzymes by heat treating oats (Molteberg et al. 1996). The stability of H OBC

lipids could lie in oxidation kinetics and the presence of antioxidants. The higher

concentrations of α-tocotrienol and slower α-tocotrienol degradation rates, even during 12

weeks’ storage at 40 °C, could tell of high antioxidant activity that could protect lipids from

degradation. On the other hand, tocols also exhibit a proxidative nature, as side reactions in

tocol degradation can be prooxidative (Kamal-Eldin and Appelqvist 1996). There are also

59

other antioxidants in the bran that could possibly affect oxidation kinetics. Purified oat

β-glucan with residual phytic acid is shown to quench and bind iron and scavenge hydroxyl

radicals (Wang et al. 2017), which could participate in protecting the lipids from oxidation

and the formation of both primary and secondary oxidation products. During storage,

β-glucan molecules did not degrade and the molecular weight remained high, over 1000 kDa.

But then again, Wang et al. (2017) explain that in an aqueous food system, β-glucans with

low molecular weight were more efficient in retarding oxidation than with high molecular

weight.

The presence of antioxidants is not a fully satisfactory explanation to the stability of H OBC

lipids. When considering the location of lipids in oat bran, the lipids are mainly located inside

thick cell walls of the aleurone layer (Miller and Fulcher 2011). Compared to other studies

that experimented with non-heat treated oats, the oat samples were wholegrain flours,

meaning the lipids in their oat samples were mainly from the starchy endosperm, where the

cell walls are lighter (Grundy et al. 2018) and the lipids thus more easily accessed and

susceptible to oxidation (Molteberg et al. 1996; Heiniö et al. 2002; Lehtinen et al. 2003).

The lipid degrading enzymes from nearby tissues should be able to mobilize lipid reserves

even through thick cell walls (Lenova et al. 2010), and lipid hydrolysis, although minimal,

should promote lipid oxidation by providing substrates for LOX and autoxidation by the

formation of FFAs (Ekstrand et al. 1992). However, it might just be that in the bran

ingredient, the cell structures were still intact and lipids protected and immobile. Larger

particle sizes have been shown to protect β-glucan from degradation in sourdough bread

(Johansson et al. 2018). The authors visualized, how in coarse oat brans the cell walls were

more intact, which limited the accessibility of β-glucan to degrading enzymes. It might be

possible that in H OBC, the lipid reservois were not accessible to lipid degrading enzymes,

or to free radicals, for that matter, resulting in a more stabile OBC product. The puzzling

fact was that M OBCs were produced using the same method as for H OBC, the only

difference being the heat-treatment, which should not affect oat cell structures. However, for

M OBC, part of the processing was done in the industry, where mere processing scale aspects

might influence the end product.

M OBC showed signs of lipid oxidation as typical lipid autoxidation related volatiles (Zhou

et al. 1999a) were produced (Figures 20 and 21) and tocopherols and tocotrienols were

degraded in the course of time and progress of lipid oxidation (Figure 18). Hexanal is known

to first fill the airspace fairly quickly and then to dissipate and/or be absorbed into the oat

60

matrix (Zhou and Decker 1999; Lehto et al. 2003), which is the most probable explanation,

why hexanal amounts decreased in M OBC after 8 weeks of storage at all storage

temperatures (Figure 20). Tocols are known to quench free radicals by their own expense

(Kamal-Eldin and Appelqvist 1996), which is why with time, when the oxidation reaction

chains advanced and the amount of formed free radicals increased, tocols were consumed

and their amounts deceased. The same degrading effect on antioxidants could be seen in the

amount of total phenolic compounds in the study of Rakic et al. (2014). Even though lipid

degradation was observed in M OBC, it did not cause changes in β-glucan molecular weight

in the dry stored ingredients. It seems that β-glucan needs an aqueous food matrix to undergo

degradation (e.g. Kivelä et al. 2009b).

To verify the compatibility and thus comparability of H and M OBC, it would be beneficial

to analyse the particle sizes of both OBCs. It would also be interesting to measure LOX

activity in H and M OBCs to better understand enzyme activity or inactivity in the OBC

ingredients.

3.6.2 Effects of OBC addition on acid milk gel functionality

OBCs were included in acid milk preparations at dosages aiming to achieve the same amount

of β-glucan in each of the final gel to deliver the 1 g of β-glucan per portion, as specified in

the health claims. Adding OBC to yogurts significantly increased acidification times in a

similar way as in the study of Rosa-Sibakov et al. (2016) using xylans as dietary fibre

enrichment. This effect might be contributed to the amount of soluble components in the

OBC, as they might affect the buffering properties of the solution. In previous studies, adding

oat fibre to yogurt has had different effects on the acidification times of yogurts. In the study

done by Fernández-García et al. (1998), oat fibre did not affect the acidification time, but in

the study done by Raikos et al. (2018), β-glucan from spent brewer’s yeast reduced the

acidification times. However, it must be acknowledged that in both studies the yogurts were

acidified with LAB and the former had also sweeteners affecting culture activity in their

yogurts, which means their yogurts were even more complex systems than GDL acidified

acid milk gels, making it challenging to compare results.

The acid milk gel textures varied a lot with both added OBC type and OBC storage time.

Both acid milk gels and OBCs are complex, heterogeneous systems. Mixing the two together

makes it increasingly difficult to isolate the effect of oxidized lipids on β-glucan

61

functionality, especially as it seems that there were no changes in β-glucan MW. There might

be other interactions affecting the final result. For example, because lipids have been found

to interfere separating oat components as fractions (Sibakov et al. 2011), OBCs might still

have some starch in them. The presence of milk ingredients, such as skim milk powder, have

been found to affect the behaviour of oat starch, including pasting, textural, rheological,

freeze-thaw, and swelling properties, both during and after cooking (Kumar et al. 2018).

Therefore, to get an idea on the effect that β-glucan might have on the gel, it is fitting to

observe the possible changes the gel matrix undergoes with the OBC addition. Some changes

to be followed are the changes in water retaining properties, as they can tell of β-glucan

functionality in the acid milk gel matrix. In this study, adding OBCs to the acid milk gels

improved the water retaining properties significantly. This can be explained with previous

knowledge on the water-binding characteristics of β-glucan molecules, as they are known to

bind water and make viscous solutions (Lazaridou and Biliaderis 2007), even in acid milk

gels (Lazaridou et al. 2008). The effect of the additional fibre network in the gel and

increased viscosity resulted in no spontaneous syneresis and less or no forced syneresis

(WHC) in the gels of the experiment (Figures 26 and 27, Table 3). The effect of storing gels

could be seen as increased firmness. When the protein network rearranges and concentrates,

serum will be released, which increases the gel protein content with an outcome of a firmer

gel (Ercili-Cura et al. 2013). This could explain why control gels were more brittle than gels

with OBC, and why gels, both control gels and gels with OBCs, became denser and firmer

after 7 days’ storage in the fridge (Figures 29, 30, and 31).

The molecular weight of β-glucan also indicates β-glucan functionality. The β-glucan

molecular weight was measured in fresh and stored acid milk gels with M OBC stored 0 and

12 weeks at 40 °C. The molecular weights remained unchanged, over 1000 kDa. However,

the method used supported only molecular weights up to 1000 kDa, meaning it was not sure

if the large β-glucans had actually degraded, but were still over the detection limit. In the

study of Wang et al. (2016), nearly pure β-glucans were added to an emulsion with highly

oxidised lipids, resulting in β-glucan degradation and lowered molecular weight and

viscosity. This is not the same when adding OBC to acid milk gels with mildly degraded

lipids. OBCs are complex systems, with parts of plant tissue possibly intact, and act as the

carriers of β-glucans and lipids. This means that in the OBCs, the β-glucans are not as

accessible as in the emulsion study with nearly pure β-glucan, which already hinders the

possibility of the degradation of β-glucans. Additionally, as it is probable that the β-glucans

in the OBCs create a viscous secondary gel network within the acid milk gel (Lazaridou et

62

al. 2008), both the protein network and the secondary β-glucan network act as steric effects,

hindering the β-glucan degradation even further.

Because the solubility of β-glucans affect their concentration in the food matrix and thus the

technological and functional properties (Lazaridou and Biliaderis 2007), it would have been

of great benefit to measure β-glucan concentrations in the gels after acidification to see how

much β-glucan has been solubilized. The amount of soluble β-glucan in the product could

also affect the product’s nutritive value as a carrier of β-glucan.

3.6.3 Limitations of study

This study had some challenges with repeatability when preparing acid milk gels. The

challenges most probably arose from the properties of the raw material. The OBCs were not

fully soluble, which made it difficult to disperse the material in a repeatable manner. The

agitation was done with magnetic stirring, and speed and length of agitation seemed to have

an impact on the solubility of the material. It was only after the heat treatment, which was

designed for protein denaturation, that OBC sedimentation was somewhat restrained.

Solubility of β-glucan is a general technological challenge in high moisture food systems

(Aktas-Akyildiz et al 2018).

When performing gel penetration tests, the gel temperatures were not constant as laboratory

and fridge temperatures varied despite attempts to control them. Having some deviation in

gel firmness results is considered acceptable, however, the fluctuating gel temperatures

added to repeatability challenges when measuring gel firmness. Low repeatability makes it

difficult to draw conclusions on the effect of storage and lipid degradation on β-glucan

functionality in acid milk gels. This why the gel firmness results should be regarded as

suggestive, and used to support other results describing β-glucan functionality, for example

viscosity.

In order to complete the storage trial and all the experiments in a limited time, not all things

could be taken into account, including slight variations in OBC compositions. Although the

aim of preparing OBC with differences in their lipid degradation states was obtained, the

OBCs most probably had slight compositional variations firstly due to small differences in

β-glucan contents, but also in L OBC due to its defatting procedures. These slight

compositional variations might have affected gel properties. To maintain a constant β-glucan

63

content in the gels, the amount of OBCs in the recipe slightly varied, affecting the dry matter

content of the final product. The milk protein amounts were fixed, so the possible variations

in dry matter contents of acid milk gels were based on added OBCs.

H OBC lipids were expected to degrade rapidly, both with hydrolysis and oxidation.

However, as H OBC did not perform as expected, additional experiments were needed,

rendering the prepared H OBC batch scarce. This lead to making only one batch of acid milk

gels per time point instead of two batches, which could add to the errors both in results and

in interpretation of the results. However, the effect is assumed to be minimal, as the standard

deviations within the batches were small, and the trend of changes in gel firmness over time

were clear.

Concerning β-glucan molecular weight analysis, the reliable detection range of the method

was only up to 1000 kDa. This study would have benefited of a range at least up to 2000 kDa,

as there were indications of high molecular weight β-glucans, which could have halved in

sized without detecting it. Additionally, for volatile analyses, the reference material revealed

to have excessive lipid degradation, as the followed volatile compounds did not stay stable.

Having a clogged CAR/PDMS-fibre was ruled out by changing the fibre in the middle of the

analyses. Not being sure if the instability originates form the instrument or the reference

material might add to errors in the results.

64

4 CONCLUSIONS

A storage trial was designed to better understand the impact of lipid hydrolysis and oxidation

in oat bran concentrates on the functionality of β-glucan in acid milk gels, a yogurt type food

model. In order to design functional foods, it has been of great interest to investigate, how

oat bran concentrates with high β-glucan concentrations (~20%) function in regard to

progressive lipid degradation both in dry oat ingredients and/or in yogurt models. It was also

investigated whether different oat bran concentrates caused differences in yogurt texture and

stability.

Lipid hydrolysis was observed in non-heat treated oat bran concentrates (H OBC, 9% fatty

acids, 18% β-glucan) and lipid oxidation was observed in heat treated oat bran concentrate

(M OBC, 8% fatty acids, 23% β-glucan). However, neither caused changes in β-glucan

molecular weight in the dry stored ingredients. Defatting oat bran concentrates was efficient

and changes in the lipid phase could not be followed (L OBC, 1% fatty acids, 20% β-glucan).

Acid milk gels with added OBCs had excellent water retaining properties and were softer

and smoother than without added OBCs. Changes in gel texture were observed when using

different OBCs and when using them after storage. However, the changes might be attributed

to other factors than molecular weight of β-glucan or products from lipid degradation in the

dry stored ingredient. Assessing OBC particle sizes and the β-glucan molecular weight from

more of the acid gels would be interesting to better understand the changes in β-glucan

molecular weight and functionality in regard to lipid degradation.

There have not been similar storage studies on hybrid ingredients researching the impact of

oxidized lipids on β-glucan. Thus this study provides novel information on the interactions

of lipid degradation (hydrolysis and oxidation) and β-glucan functionality. Even when lipids

were oxidized within the dry oat ingredient, β-glucan molecules did not degrade, which

might be due to having protective cell wall structures intact in the oat bran concentrates.

65

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APPENDIX

Appendix 1. An exemplary graph for the analysis of gel firmness with TA

A deformation test was performed on 40 g aliquots of gel, acidified in plastic cups (ø 3.5 cm), pH 4.6±1.0, by penetrating the gel with a ø 25 mm cylinder probe at a constant speed of1.0 mm/s to a distance of 75% of the sample height. The area under the force vs. distance curve (up to 15 mm) was determined and taken as a measure of gel firmness