ISTANBUL TECHNICAL UNIVERSITY  GRADUATE SCHOOL 

M.Sc. THESIS 

JUNE 2023 

ENCAPSULATION OF AQUEOUS HIBISCUS SABDARIFFA L. EXTRACT IN 

FOOD-GRADE HIGH INTERNAL PHASE PICKERING EMULSIONS 

STABILIZED BY SOY PROTEIN ISOLATE 

Hümeyra ÇAVDAR 

 

Department of Food Engineering 

 

Food Engineering Programme 

 

 

 



 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

  



 

                                                                     

 

 

 

 

 

 

 

  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

ISTANBUL TECHNICAL UNIVERSITY  GRADUATE SCHOOL 

M.Sc. THESIS 

JUNE 2023 

ENCAPSULATION OF AQUEOUS HIBISCUS SABDARIFFA L. EXTRACT IN 

FOOD-GRADE HIGH INTERNAL PHASE PICKERING EMULSIONS 

STABILIZED BY SOY PROTEIN ISOLATE 

Hümeyra ÇAVDAR 

(506211516) 

 

 

Department of Food Engineering 

 

Food Engineering Programme 

 

 

 

Thesis Advisor: Esra ÇAPANOĞLU GÜVEN 

 



 

 

  



 

 

                                  

           

 

 

 

 

 

 

 

 

 

 

 

    

 

 

 

 

 

 

 

 

 

 

 

 

 

              

 

 

 

 

 
HAZİRAN 2023 

İSTANBUL TEKNİK ÜNİVERSİTESİ  LİSANSÜSTÜ EĞİTİM ENSTİTÜSÜ 

SULU HIBISCUS SABDARIFFA L.  EKSTRAKTININ SOYA PROTEİNİ İLE 

STABİLİZE EDİLEN GIDA SINIFI YÜKSEK DAHİLİ FAZLI PICKERING 

EMÜLSİYONLARDA ENKAPSÜLASYONU  

YÜKSEK LİSANS TEZİ 

Hümeyra ÇAVDAR 

 (506211516) 

Gıda Mühendisliği Anabilim Dalı 

 

Gıda Mühendisliği Programı 

 

 

 

 

Tez Danışmanı: Prof. Dr. Esra ÇAPANOĞLU GÜVEN 

 



 

 

 

 

 

 

 

 

 

 

 

 

 

 

   

 



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Thesis Advisor:  Prof. Dr. Esra Çapanoğlu GÜVEN        ............................. 

 Istanbul Technical University 

Jury Members : Assoc. Prof. Dr. Aslı Can KARAÇA      ………………….. 

   Istanbul Technical University 

                                    

                                   Assoc. Prof. Dr. Zeynep Tacer CABA   …………………… 

                                   Bahcesehir University 

    

 

                                                                                                       

Istanbul Technical University 

Hümeyra ÇAVDAR, a M.Sc. student of ITU Graduate School student ID 506211516, 

successfully defended the thesis/dissertation entitled “ENCAPSULATION OF 

AQUEOUS HIBISCUS SABDARIFFA L. EXTRACT IN FOOD-GRADE HIGH 

INTERNAL PHASE PICKERING EMULSIONS STABILIZED BY SOY PROTEIN 

ISOLATE”, which she prepared after fulfilling the requirements specified in the 

associated legislations, before the jury whose signatures are below. 

 

Teslim Tarihi  :16.06.2023  

Savunma Tarihi  :15.06.2023     



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To my beloved family, 

 

 

 



vi 



vii 

FOREWORD 

I would like to extend my heartfelt thanks and appreciation to my supervisor and 

mentor, Prof. Dr. Esra ÇAPANOĞLU GÜVEN, for their invaluable guidance, support, 

and education throughout this journey. Her expertise and mentorship have played a 

crucial role in shaping my scientific and academic development. 

 

I am also grateful to Asst. Prof. Dr. Gülay ÖZKAN, who helped me gain self-

confidence in the laboratory since the beginning of my graduate education, greatly 

enriched my research experience and taught me many of the analyses I performed. 

 

I would like to express my sincere thanks to my office-mates, Elif Feyza AYDAR, 

Zehra MERTDİNÇ, and Fatma Duygu CEYLAN ADRAR, for their support and 

friendship during this time. Their presence has made the journey more enjoyable and 

fulfilling. 

 

Lastly, I would like to convey a special note of appreciation and gratitude to my 

parents, Ayşe and Ali ÇAVDAR, my sister Rümeysa ÇAVDAR, and my husband 

Mehmet DİNÇTÜRK. Their unwavering love, patience, and support have been the 

cornerstone of my education and life. I am deeply grateful for their constant 

encouragement and belief in me. 

 

June 2023        Hümeyra ÇAVDAR 

                (Research Assistant) 

 

 

 

 

 

 



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ix 

TABLE OF CONTENTS 

Pages 

FOREWORD ............................................................................................................ vii 

ABBREVIATIONS ................................................................................................... xi 

SYMBOLS………………………………………………………………………… xiii 

LIST OF TABLE ..................................................................................................... xv 

LIST OF FIGURE ................................................................................................. xvii 

SUMMARY ............................................................................................................. xix 

ÖZET……………………………………………………………………………… xxi 

1. INTRODUCTION .................................................................................................. 1 

1.1 Purpose of Thesis ............................................................................................... 2 

1.2 Literature Review ............................................................................................... 2 

2. MATERIALS AND METHODS ........................................................................ 13 

2.1 Materials ........................................................................................................... 13 

2.2 Preparation of Aqueous Hibiscus Extract (AHE)............................................. 13 

2.3 Preparation of Soy Protein Isolate (SPI) Gel ................................................... 13 

2.4 Preparation of Emulsions ................................................................................. 14 

2.4.1 High internal phase pickering emulsions (HIPPEs) .................................. 14 

2.4.2 High internal phase pickering-double emulsions (HIPP-DEs) ................. 15 

2.5 Storage Stability ............................................................................................... 15 

2.6 Encapsulation Efficiency .................................................................................. 16 

2.7 Determination of Emulsifying Properties ........................................................ 16 

2.8 Particle size, PDI and Zeta Potential ................................................................ 17 

2.9 Creaming Stability ............................................................................................ 17 

2.10 Centrifugation Stability .................................................................................. 17 

2.11 In Vitro Gastrointestinal Digestion ................................................................ 17 

2.12 The Quantification of Polyphenols with HPLC-DAD ................................... 18 

2.13 Spectrophotometric Assays ............................................................................ 19 

2.13.1 Total anthocyanin content (TAC) ........................................................... 19 

2.13.2 Total phenolic content (TPC) .................................................................. 19 

2.13.3 Antioxidant activity ................................................................................. 19 

2.13.4 Bioaccessibility calculations ................................................................... 19 

2.14 Statistical Analysis ......................................................................................... 20 

3. RESULT AND DISCUSSION ............................................................................. 21 

3.1 Storage Stability ............................................................................................... 21 



x 

3.2 Creaming Stability ............................................................................................ 24 

3.3 Centrifugation Stability .................................................................................... 25 

3.4 Determination of emulsifying properties .......................................................... 26 

3.5 Particle size, zeta potential and PDI ................................................................. 29 

3.5.1 Particle size ............................................................................................... 29 

3.5.2 Zeta potential ............................................................................................. 31 

3.5.3 PDI ............................................................................................................ 33 

3.6 Encapsulation Efficiency .................................................................................. 34 

3.7 Effect of In Vitro Gastrointestinal Digestion on TAC, TPC, DPPH and ABTS of 

HIPPEs and HIPP-DEs ........................................................................................... 36 

3.7.1 Total anthocyanin content ......................................................................... 36 

3.7.2 Total phenolic content ............................................................................... 38 

3.7.3 Bioaccessibility of phenolics and anthocyanins ........................................ 41 

3.7.4 Antioxidant activity ................................................................................... 43 

3.7.5 Quantification of Polyphenols with HPLC-DAD ..................................... 45 

4. CONCLUSION AND RECOMMENDATIONS ............................................... 49 

REFERENCES ......................................................................................................... 51 

CURRICULUM VITAE .......................................................................................... 63 

 

 

 



xi 

ABBREVIATIONS 

ABTS : 2,2’ Azinobis (3-Ethylbenzothiazoline-6-Sulfonic Acid) Diammonium Salt 

C3-GE: Cyanidin 3-Glucoside Equivalent 

DPPH : 2,2-Diphenyl-1-picrylhydrazyl 

EAI : Emulsion Activity Index 

ESI : Emulsion Stability Index 

GAE : Gallic Acid Equivalent 

SPI : Soy Protein Isolate  

TAC : Total Anthocyanin Content 

TE : Trolox Equivalent 

TFC: Total Flavonoid Content 

TPC : Total Phenolic Content 

 



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xiii 

SYMBOLS 

g : Gram 

mg : Miligram 

mL : Mililiter 

p : Confidence Level 

pH : Potential of Hydrogen 

 



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xv 

LIST OF TABLES 

Page 

Table 1.1: Nutritional and bioactive compounds in Hibiscus Sabdariffa. ................... 3 

Table 1.2: Encapsulation of bioactives within HIPPEs and DEs stabilized SPI. ...... 10 

Table 2.1: HIPPEs with different formulations. ........................................................ 14 

Table 2.2: HIPP-DEs with different formulations. ................................................... 15 

Table 3.1: EAI and ESI values of HIPPEs. ............................................................... 28 

Table 3.2: EAI and ESI values of HIPP-DEs. ........................................................... 29 

Table 3.3: Total anthocyanin content (mg C-3-GE/ 100 g) of HE and HIPPEs. ...... 38 

Table 3.4: Total anthocyanin content (mg C-3-GE/ 100 g) of HE and HIPP-DEs. .. 38 

Table 3.5: Effect of in vitro digestion on TPC (mg GAE/100 g) of HE and HIPPEs.

 ............................................................................................................................ 40 

Table 3.6: Effect of in vitro digestion on TPC (mg GAE/g) of HE and HIPP-DEs. 41 

Table 3.7: Effects of in vitro digestion on DPPH (mg TE/100 g dw) and ABTS (mg 

TE/100 g dw) of Hibiscus extracts (HE) and HIPPEs. ...................................... 44 

Table 3.8: Effects of in vitro digestion on DPPH (mg TE/100 g dw) and ABTS (mg 

TE/100 g dw) of Hibiscus extracts(HE) and HIPP-DEs. ................................... 45 

Table 3.9: Anthocyanin and phenolic profile of HIPPEs. ......................................... 47 

   



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xvii 

LIST OF FIGURES 

Page 

Figure 3.1: HIPPEs stabilized by only different SPI gel concentrations. ................. 21 

Figure 3.2: HIPPEs stabilized by only different lecithin concentrations. ................. 22 

Figure 3.3: HIPPEs stabilized by different SPI gel and lecithin concentrations....... 23 

Figure 3.4: HIPP-DEs stabilized by different SPI gel and lecithin concentrations .. 23 

Figure 3.5: Changes in creaming index (CI) of HIPPEs during storage ................... 25 

Figure 3.6: Changes in creaming index (CI) of HIPP-DEs during storage............... 25 

Figure 3.7: Degree of phase separation of L4 and L6 after centrifugation ............... 26 

Figure 3.8: HIPP-DEs after centrifugation ............................................................... 26 

Figure 3.9: Change in the particle size and zeta potential between different HIPPEs.

 ............................................................................................................................ 32 

Figure 3.10: Change in the particle size and zeta potential between different HIPP-

DEs. .................................................................................................................... 33 

Figure 3.11: PDI values of HIPPEs. ......................................................................... 34 

Figure 3.12: PDI values of HIPP-DEs. ..................................................................... 34 

Figure 3.13: EEA and EEP values of HIPPEs .......................................................... 35 

Figure 3.14: EEA and EEP values of HIPPE-DEs.................................................... 36 

Figure 3.15: Bioavailability of phenolics and anthocyanins of HIPPEs. .................. 42 

Figure 3.16: Bioavailability of phenolics and anthocyanins of HIPP-DEs. ............. 43 

Figure 3.17: Change in anthocyanin and phenolic profile of L6S6 during in vitro 

digestion ............................................................................................................. 48 

Figure 3.18: Change in anthocyanin and phenolic profile of L6S6-D during in vitro 

digestion ............................................................................................................. 48 

 

  



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xix 

ENCAPSULATION OF AQUEOUS HIBISCUS SABDARIFFA L. EXTRACT 

IN FOOD-GRADE HIGH INTERNAL PHASE PICKERING EMULSIONS 

STABILIZED BY SOY PROTEIN ISOLATE 

SUMMARY 

Hibiscus Sabdariffa L. is a highly versatile plant that finds applications in various 

industries due to its abundant nutrients, potent bioactive compounds such as phenolics 

and anthocyanins, and natural colour pigments. In addition to its use as a colouring 

agent, its positive health effects, such as antibacterial, antioxidant, anticholesterol and 

prevention of gastrointestinal problems, spread the consumption of the product for 

health purposes. However, the fact that bioactive ingredients are easily affected and 

damaged by many factors limits this product's use in many industries. Therefore, 

integrating Hibiscus Sabdariffa into food products and various formulations becomes 

more complex and limits the methods used. Encapsulation, a preferred method for 

much bioactive protection, appears at this point. Encapsulation comes to fields as a 

technique that allows us to apply in different areas, such as preserving these 

components and controlled release and integrating them into product formulations. 

Emulsification by encapsulation method has gained a solid place in industry and 

literature. High Internal Phase Emulsions are among the leading emulsions with high 

volume encapsulation and high stability against external factors. Similarly, double 

emulsion systems can enhance bioactive components' protection with their nested 

phase structure. The debate over the many adverse effects of preferred emulsifiers and 

stabilizers for emulsion stability, such as human health and environmental pollution, 

has accelerated scientific research into discovering alternative products. Plant proteins 

have gained tremendous interest in recent years as they have the potential to be an 

alternative to conventional emulsifiers and stabilizers. The various functional 

properties and amphiphilic characteristics indicate that these proteins are significantly 

effective in the stability of the emulsion system. Like many other types of plant 

proteins, soy protein is a preferred product due to its easy availability and 

processability. 

In this study, analyses were conducted to identify and evaluate the encapsulation of 

aqueous Hibiscus extract in the High Internal Phase Pickering Emulsions (HIPPE) and 

High Internal Phase Double Emulsions (HIPP-DE) systems stabilized by soy lecithin 

and soy protein isolate. The stability of the emulsions obtained, characteristics and the 

effects of in vitro gastrointestinal digestion on the phenolics and anthocyanins in the 

extract with the presence of soy protein and soy lecithin were investigated. 

The extraction of phenolics and anthocyanins from powdered Hibiscus calyces was 

realized by ultrasonic method, and water was used as a solvent. Emulsions with an 

internal volume of 80% have been created using soy protein isolate (SPI) gel and 

lecithin. A HIPPE resistant to phase separation could not be obtained using soya 

protein isolation gel alone, while emulsions (L4 and L6) containing 4% and 6% 

lecithin and phase separation resistance could be achieved. Combinations of different 

concentrations of lecithin and soy protein isolated gel stabilized other HIPPEs. After 

24 hours of storage, HIPPEs stable against phase separation were obtained by 

homogenizing HIPP-DEs at a volume of 50% with a 6% SPI gel. When emulsions 

stored for 24 hours were observed, SPI gels and lecithin at varying concentrations 

acted as a synergistic mechanism that effectively prevented phase segregation and 

ensured stability in the emulsion system. All HIPP-DEs also showed superior 



xx 

resistance to phase separation. According to the CI results, an increase in the 

concentration of soy lecithin from 2% to 4% in HIPPE containing 6% SPI gel resulted, 

indicating increased stability, leading to a decrease in the CI value on day 1; however, 

no significant difference in CI was observed between HIPP-DEs. EAI and ESI values 

show a statistically significant increase when the SPI concentration increases from 2% 

to 4% in HIPPEs and HIPP-DEs with 4% and 6% lecithin concentrations. Still, 

surprisingly, no significant increase was observed with further increases in SPI. In 

contrast, the particle size of HIPPEs with 4% and 6% lecithin concentrations showed 

a significant decrease as the SPI gel concentration increased from 2% to 6%. In 

HIPPEs, the zeta potential was negative, and the absolute values increased by 

increasing the SPI gel concentration from 2% to 4%, while in L4S4, it was observed 

that the HIPPE had the highest absolute zeta potential (-41.21 ± 1.23 mV). However, 

an increase in SPI gel concentration from 4% to 6% decreased the absolute zeta 

potential value. The increase in the concentration of soy protein isolate (SPI) from 2% 

to 6% resulted in a significant decrease in the PDI values of HIPPE containing 4% and 

6% lecithin. The lowest PDI values were obtained in L4S6 and L6S6 HIPPE with an 

SPI concentration of 6%. The same decline has been observed in HIPP-DEs. 

Different changes have been observed in the stability and characteristics of emulsions, 

as well as in the study of bioactive components and their properties. Encapsulation 

Efficiency (EE) determinations stated that HIPP-DEs produced higher EE than 

HIPPEs due to high SPI gel concentrations. The total anthocyanin content (TAC), total 

phenolic content (TPC), DPPH and ABTS of the resulting aqueous Hibiscus extract 

were analyzed. The TAC (mg Cy-3-GC equivalent/100 g), TPC (mg GAE/100 g), 

DPPH (mg TE/100g) and ABTS (mg TE/100g) values were found as 31.13±1.23, 

2619.01±17.31, 335.12±1.21 and 223.21±2.56 respectively. Significant decreases in 

TPC in Hibiscus extract were observed with in vitro gastrointestinal digestion. In 

HIPPEs prepared with a concentration of 4 % lecithin (L4, L4S2, L4 S4, L4S6), the 

SPI gel concentration increased from 0 % to 4 %, while anthocyanins in the stomach 

environment showed more excellent stability compared to the intestinal environment. 

When the TPC of the digested phenolics of Hibiscus extract was compared to those of 

digestive HIPPEs and HIPP-DEs, higher TPC values were observed in all emulsions. 

Maximum TPC values in the stomach and intestinal digestion reached 709.82 ±2.06 

mg/100g and 1160.71 ±21.01 mg/100g, respectively. In the stomach and intestinal 

phases of in vitro digestion, the rate of phenolic release has been significantly 

influenced by the pH of the environment. According to HPLC results, aqueous 

Hibiscus extract has detected anthocyanins of delphinidin-3-glucoside, cyanidin-3-

glucoside, and cyanidin-3-rutinoside. Four phenolic acids have also been detected in 

the extract and all emulsions, including gallic acid, syringic acid, ferulic acid and 

chlorogenic acid. After gastric digestion, a decrease in the content of chlorogenic acid, 

ferulic acid and gallic acid was observed, while syringic acid increased, showing a 

different tendency. The highest concentrations of each anthocyanins and phenolics 

concentrations were obtained in L6S6-D. 

As a result, changes in SPI gel and lecithin concentrations have been effective in many 

conditions, such as the stability of emulsions, their properties, and the effects of in 

vitro digestion on anthocyanins and phenolics. In addition, according to the results of 

encapsulation efficiency, it can be stated that HIPPEs and HIPP-DEs are effective 

emulsion systems in the encapsulation of Hibiscus extract. 

  



xxi 

SULU HIBISCUS SABDARIFFA L.  EKSTRAKTININ SOYA PROTEİNİ İLE 

STABİLİZE EDİLEN GIDA SINIFI YÜKSEK DAHİLİ FAZLI PICKERING 

EMÜLSİYONLARDA ENKAPSÜLASYONU 

ÖZET 

Hibiscus Sabdariffa L., zengin besin içeriği, fenolik ve antosiyaninler açısından güçlü 

biyoaktif bileşenleri, doğal renk pigmentleri sayesinde birçok endüstride geniş 

kullanım alanı bulmaktadır. Renklendirici olarak kullanımının yanı sıra antibakteriyel 

antioksidan, antikolesterol ve gastrointestinal problemler gibi sağlık üzerindeki olumlu 

etkileri ürünün sağlık amacıyla tüketimini yaygınlaştırmaktadır. Ancak biyoaktif 

bileşenlerin birçok farklı faktörden kolaylıkla etkilenmesi ve zarar görmesi bu ürünün 

birçok endüstride kullanımını sınırlandırmaktadır. Bu nedenle Hibiscus Sabdariffa'nın 

gıda ürünü formülasyonlarına entegrasyonu zorlaşmakta ve kullanılacak yöntemleri 

sınırlandırmaktadır. Birçok biyoaktif koruma önlemi için tercih edilen bir yöntem olan 

enkapsülasyon bu noktada ortaya çıkmaktadır. Enkapsülasyon, yalnızca bu 

bileşenlerin korunması değil, aynı zamanda kontrollü salınım ve ürün tüketimine dahil 

edilmesi gibi farklı alanlarda uygulama sağlayan bir teknik olarak karşımıza 

çıkmaktadır. Emülsifikasyon ile enkapsülasyon yöntemi, hem endüstride hem de 

literatürde sağlam bir yer edinmiştir. Yüksek Dahili Fazlı Emülsiyonlar, yüksek 

hacimli enkapsülasyon, yüksek stabilite ve biyoaktif bileşenleri farklı etkenlere karşı 

koruma özellikleri ile ön plana çıkan emülsiyon türleri arasındadır. Benzer şekilde Çift 

Katlı Emülsiyon sistemleri de iç içe fazlardan oluşan yapıları ile biyoaktif bileşenler 

için yüksek koruma sağlayabilmektedir. Emülsiyon stabilitesi için tercih edilen 

emülgatörler ve stabilizatörlerin insan sağlığı ve çevre kirliliği gibi birçok olumsuz 

etkileri üzerine yapılan tartışmalar alternatif ürünlerin keşfedilmesi üzerine 

gerçekleştirilen bilimsel çalışmaları hızlandırmıştır. Bitkisel proteinler konvansiyonel 

emülgatör ve stabilizatörlere alternatif olma potansiyeline sahip oldukları için son 

yıllarda yoğun ilgi görmektedir. Çeşitli fonksiyonel özellikleri ve amfifilik 

karakterleri, bu proteinlerin emülsiyon sisteminin stabilitesini korumada önemli 

ölçüde etkili olduğunu göstermektedir. Diğer birçok bitkisel protein türü gibi soya 

proteinleri de kolay ulaşılabilirliği ve işlenebilirliği nedeniyle tercih edilen bir üründür. 

Bu çalışmada sulu Hibiskus extraktının soya lesitin ve soya protein izolatı içeren 

Yüksek Dahili Fazlı Pickering Emülsiyonlar (HIPPE) ve Yüksek Dahili Fazlı 

Pickering Çift Emülsiyon (HIPP-DE) sistemlerinde enkapsülasyonu, elde edilen 

emülsiyonların stabilitesi, emülsiyon özellikleri ve in vitro gastrointestinal sindirimin 

soya proteini ve lesitin varlığında ekstrakttaki fenolikler ve antosiyaninler üzerindeki 

etkilerinin ortaya çıkarılması ve değerlendirilmesi üzerine analizler 

gerçekleştirilmiştir.  

Toz haline getirilen Hibiscus çanaklarından fenolik ve antosiyaninlerin ekstraksiyonu 

ultrasonik yöntemle gerçeleştirilmiş ve çözgen olarak su kullanılmıştır. Soya proteini 

izolat (SPI) jeli ve lesitin kullanılarak %80 iç hacme sahip emülsiyonlar 

oluşturulmuştur. Yalnızca soya proteini izolat jeli kullanılarak faz ayrımına karşı 

dirençli bir HIPPE elde edilemezken, %4 ve %6 lecithin içeren ve faz ayrımına karşı 

dirençli emülsiyonlar (L4 ve L6) elde edilebilmiştir. Diğer HIPPE'ler ise farklı lecithin 

ve soy protein izolat jeli konsantrasyonlarının kombinasyonları ile stabil hale 

getirilmiştir. 24 saat depolama sonrasında faz ayrımı gözlemlenmeyen ve stabil halde 

bulunan HIPPE'ler, %6 SPI jel ile hacimce %50 oranında homojenize edilerek HIPP-

DE'ler elde edilmiştir. 24 saat boyunca depolanan emülsiyonlar gözlemlendiğinde, 



xxii 

HIPPE'lerde değişen konsantrasyonlarda SPI jeli ve lesitinin kullanılması, faz ayrımını 

etkili bir şekilde önleyen ve emülsiyon sisteminde stabilite sağlayan sinerjistik bir 

mekanizma görevi üstlenmiştir. Ayrıca elde edilen tüm HIPP-DE’ler faz ayrımına 

karşı üstün bir direnç göstermiştir. CI sonuçlarına göre, %6 SPI jeli içeren HIPPE'lerde 

soya lesitin konsantrasyonunun %2'den %4'e çıkarılması, 1. günde CI değerinin 

düşmesine yol açarak, stabilitenin arttığını gösterirken, %6 lesitin konsantrasyonu 

artmasına neden olmuştur; ancak, HIPP-DE'ler arasında CI açısından önemli bir fark 

gözlemlenmemiştir. Hem EAI hem de ESI değerleri, %4 ve %6 lesitin 

konsantrasyonlarına sahip HIPPE'lerde ve HIPP-DE'lerde SPI konsantrasyonu %2'den 

%4'e yükseldiğinde istatistiksel olarak anlamlı bir artış sergilerken, şaşırtıcı bir 

şekilde, SPI konsantrasyonu daha da arttığında anlamlı bir artış gözlenmemiştir. Bu 

durumun aksine, %4 ve %6 lesitin konsantrasyonlarına sahip HIPPE'lerin parçacık 

boyutu, SPI jel konsantrasyonu %2'den %6'ya çıktıkça önemli bir azalma sergilemiştir. 

HIPPE'lerde zeta potansiyelinin negatif olduğu ve SPI jel konsantrasyonunun %2'den 

%4'e arttırılmasıyla mutlak değerlerinin arttığı gözlemlenirken, L4S4 ne yüksek 

mutlak zeta potansiyeline (-41,21 ± 1,23 mV) sahip HIPPE olmuştur. Ancak, %4'ten 

%6'ya artan SPI jel konsantrasyonu, mutlak zeta potansiyel değerinde azalmaya 

sebebiyet vermiştir. Soya proteini izolatının (SPI) konsantrasyonunun %2'den %6'ya 

çıkarılması, %4 ve %6 lesitin içeren HIPPE'lerin PDI değerlerinde anlamlı bir 

azalmaya yol açmıştır. En düşük PDI değerleri, SPI konsantrasyonu %6 olduğu olduğu 

L4S6 ve L6S6 HIPPE’lerinde elde edilmiştir. Aynı düşüş HIPP-DE'ler içinde geçerli 

olmuştur. 

Emülsiyon stabilitesi ve karakteristiklerinin yanı sıra emülsiyon sistemlerinin 

biyoaktif bileşenler ve özellikleri açısından incelenmesi ile farklı değişimler ortaya 

konmuştur. Enkapsülayon verimliliği (EE) araştırıldığında görülmüştür ki HIPP-

DE’ler yüksek SPI jel konsantrasyonu sayesinde HIPPE'lere göre daha yüksek EE 

sağlamıştır. Elde edilen sulu Hibiscus ekstraktının toplam antosiyanin miktarı (TAC), 

toplam fenolik miktarı (TPC), DPPH ve ABTS yönünden analiz edilmiştir. TAC (mg 

Cy-3-GC eşdeğeri/100 g), TPC (mg GAE/100 g) , DPPH (mg TE/100 g) and ABTS 

(mg TE/100g ) değerleri sırasıyla 31.13±1.23, 2619.01±17.31, 335.12±1.21 ve 

223.21±2.56 olarak bulunmuştur. In vitro gastrointestinal sindirim ile birlikte Hibiskus 

ekstraktında TPC'de önemli düşüşler gözlemlenmiştir. %4 lesitin konsantrasyonu ile 

hazırlanan HIPPE’lerde (L4, L4S2, L4S4, L4S6) SPI jel konsantrasyonu %0'dan %4'e 

yükselirken, mide sindiriminde antosiyaninler bağırsak ortamına göre daha yüksek 

stabilite göstermiştir. Hibiskus ekstraktının sindirilmiş fenoliklerinin TPC'si 

sindirilmiş HIPPE'lerin ve HIPP-DE'lerinkiyle karşılaştırıldığında, tüm 

emülsiyonlarda daha yüksek TPC değerleri gözlemlenmiştir. Mide ve bağırsak 

sindirimlerinde maksimum TPC değerleri sırasıyla 709.82±2.06 mg / 100g ve 

1160.71±21.01 mg/100 g'a ulaşmıştır. In vitro sindirimin mide ve bağırsak fazları 

enasında fenoliklerin salınım hızı, ortam pH’ından önemli ölçüde etkilenmiştir. HPLC 

sonuçlarına göre, sulu Hibiskus ekstraktında delfinidin-3-glukozit, siyanidin-3-

glukozit, ve siyanidin-3-rutinozit antosiyaninleri tespit edilmiştir. Ayrıca ekstrakt ve 

tüm emülsiyonlarda gallik asit, sirincik asit, ferulik asit, klorojenik asit olmak üzere 

dört fenolik asit tespit edilmiştir. Gastrik sindirimden sonra, klorojenik asit, ferulik asit 

ve gallik asit içeriğinde bir azalma gözlemlenirken, sirincik asit farklı bir eğilim 

göstererek artmıştır. En yüksek konsantrasyon antosiyanin ve fenolik 

konsantrasyonları L6S6-D'de elde edilmiştir.  

Sonuç olarak, SPI jel ve lesitin konsantrasyonlarındaki değişimler, emülsiyonların 

kararlılığı, özellikleri, in vitro sindirimin antosiyaninler ve fenolikler üzerindeki 

etkileri gibi birçok koşulda etkili olmuştur. Ayrıca enkapsülasyon verimliliği 



xxiii 

sonuçlarına göre Hibiskus ekstraktının enkapsülasyonunda HIPPE ve HIPP-DE'nin 

etkili emülsiyon sistemler olduğu söylenebilir. 

 

  



xxiv 

 

 



1 

1.  INTRODUCTION 

Hibiscus Sabdariffa L. finds a wide range of uses in many industries thanks to its rich 

nutritional content and bioactive ingredients rich in phenolic and anthocyanins and the 

natural color pigments obtained. In addition to its use as a colorant, its positive health 

effects including antibacterial antioxidant, anticholesterol and gastrointestinal 

problems make the consumption of the product widespread for this purpose. However, 

the fact that bioactive components are easily affected and damaged by many different 

factors limits the use of this product in many industries. For this reason, the integration 

of Hibiscus Sabdariffa into food product formulations becomes difficult and limits the 

methods to be used. Encapsulation, which is a preferred method for many bioactive 

protection measures, is the sovereignty that emerges at this point. Encapsulation is not 

only the preservation of these measures, but also containment provisions as a technique 

applied in areas such as controlled release and incorporation into product consumption. 

There are many types such as spray drying, freeze drying, coacervation, liposome and 

emulsification. The method of encapsulation by emulsification is technically 

conservative provisions that have gained a solid place in both industry and literature. 

In this method, bioacti production is protected and transported by being trapped in 

water or oil phase, depending on the characteristics of its properties. High Internal 

Phase Emulsions are among the emulsion types that come to the fore thanks to their 

high volume encapsulation, high stability and ability to protect the contents against 

different factors. Similarly, Double Emulsion systems can provide high protection for 

bioactive components with their structure consisting of nested phases.However, 

emulsifiers and stabilizers are used to homogenize these two phases, which do not mix 

with each other as they have a normal emulsion appearance. Lecithin, Tween, Span 

and PGPR are conventionally preferred emulsifiers and stabilizers. However, the 

ongoing discussions on the existence of many negative results such as human health 

and environmental pollution, as well as the use of them, and the studies of scientific 

studies on alternative substances have been accelerated. Plant-based proteins have 

attracted intense interest in recent years because they have the potential to be an 



2 

alternative to conventional emulsifiers and stabilizers. The various functional 

properties and amphiphilic characters feel that these proteins are significantly effective 

in maintaining the stability of the emulsion system. Soy proteins, like many other 

vegetable protein types, are a preferred product due to their easy accessibility and 

processability. 

1.1 Purpose of Thesis 

The aim of this study is to encapsulate aqueous Hibiscus Sabdariffa L. extract in 

lecithin and soy protein based food-grade High Internal Phase Pickering Emulsions 

(HIPPE) and High Internal Phase Pickering Double Emulsions (HIPP-DE) systems, to 

examine the emulsions obtained in terms of emulsion stability and emulsion 

properties, and to reveal the effects of in vitro gastrointestinal digestion on phenolics 

and anthocyanins in the context of bioavailability. Although various studies researched 

the encapsulation of bioactive compounds, no aqueous Hibiscus extract encapsulation 

within food grade HIPPE and HIPP-DE systems have been studies to the best of our 

knowledge.  

1.2 Literature Review 

Hibiscus Sabdariffa L., belonging to the Malvaceae family, has garnered global 

recognition as a medicinal plant in the field of human health (Ismail et al., 2008). 

Commonly known as Roselle, it is referred to by various names such as Kenaf, 

Karkade, and Asam paya, varying across different countries or regions (Ismail et al., 

2008). Widely distributed and cultivated in tropical and subtropical regions, including 

Africa, India, and Sudan, H.Sabdariffa exhibits distinctive characteristics such as 

smooth, cylindrical red stems and green leaves (Da-Costa-Rocha et al., 2014). 

H.Sabdariffa as a valuable ingredient in the food industry, particularly in the 

development of smoothies, beverages, jellies, and confectionery products. Its natural 

reddish color is attributed to the presence of water-soluble color pigments, primarily 

anthocyanins, offering a preferable alternative to synthetic colorants (Amer et al., 

2022). In response to increasing consumer demand for healthier and more natural food 

options, the replacement of artificial colorants with H.Sabdariffa-derived pigments has 

gained significant attention (Ko et al., 2017). From a food science perspective, 

H.Sabdariffa possesses notable nutritional and bioactive compounds that contribute to 



3 

its medicinal properties. The calyces of the plant exhibit a rich composition of 

carbohydrates, proteins, fats, and various phytochemicals (Jabeur et al., 2017). These 

phytochemical constituents, including anthocyanins, flavonoids, and phenolics, have 

been extensively studied for their therapeutic effects in treating diverse diseases 

(Formagio et al., 2015). Notably, H.Sabdariffa demonstrates a wide range of biological 

activities, encompassing antibacterial and antioxidant properties, cancer 

chemoprevention, appetite suppression, lipid metabolism modulation, and beneficial 

effects on hypertension and gastrointestinal issues (Da-Costa-Rocha et al., 2014; Zhen 

et al., 2016). These therapeutic effects are attributed to bioactive compounds present 

in H.Sabdariffa, such as phenolic acids, flavonoids, anthocyanins, and organic acids 

(Herranz-López et al., 2018; Riaz & Chopra, 2018). Table 1.1 represents the nutritional 

and bioactive compounds in Hibiscus Sabdariffa.  

Table 1.1: Nutritional and bioactive compounds in Hibiscus Sabdariffa. 

Compound Bioactive compound Amount Reference 

Carbohydrates  87 ± 1 g/100 g dw Jabeur et al. 

(2017) 

 

Fibers Dietary fiber (total) 33.9 ± 3.56 g/100 g dw 

Proteins  5.5 ± 0.4 g/100 g dw 

Fats  0.59 ± 0.06 g/100 g dw 

Phenolic acid & 

Flavonoids 

Chlorogenic acid 7.55 ± 2.75 g/100 g dw Zhen et al. 

(2016) 

Quercetin 3.2 g/ 100 g dw Alarcón-Alonso 

et al. (2012) Rutin 2.1 g/ 100 g dw 

5-O-Caffeoyl-

shikimic acid 

11.43 ± 0.87 g/100 g dw Piovesana et al. 

(2019) 

Quercetin-3-

rutinoside 

7.77 ± 0.28 g/100 g dw 

Anthocyanins Delphinidin-3-

sambubioside 

218.17 ± 12.69 g/100 g 

dw 

Cyanidine-3-

sambubioside 

70.42 ± 5.26 g/100 g dw 

 

The identification and quantification of bioactive compounds play a crucial role in 

understanding the intricate chemistry of plant tissues, which are influenced by a 

diverse array of primary and secondary metabolites. In order to perform accurate 

analytical procedures, it is imperative to prepare a representative sample that reflects 



4 

the real product matrix. Particularly when dealing with complex sample matrices 

containing multiple components, it becomes essential to purify or separate non-target 

compounds from the sample before analysis to prevent interference with the targeted 

compounds (Hapsari & Setyaningsih, 2021). Extraction is commonly employed as a 

sample preparation step to isolate the desired compounds from the sample matrix 

(Bylda et al., 2014). However, the complexity of phenolic compounds and their 

interactions with other constituents in the matrix present challenges in developing an 

optimal extraction method capable of extracting multiple analytes from such a 

complex matrix. Thus, the selection of an appropriate extraction method is pivotal in 

the analytical procedure. Traditional extraction methods like Soxhlet, maceration, 

percolation, and infusion have traditionally been favored for various samples due to 

their simplicity (Hidayat & Wulandari, 2021; López-Bascón & De Castro, 2020). 

Nonetheless, novel extraction methods such as microwave-assisted extraction (MAE), 

supercritical fluid extraction (SFE) and ultrasound-assisted extraction (UAE) have 

emerged in recent years, specifically targeting the extraction of phenolic compounds 

(Mena-García et al., 2019). These advanced techniques have gained popularity due to 

their ability to reduce extraction time and solvent consumption (Jiang et al., 2021). 

Regarding the extraction of bioactive compounds from H. Sabdariffa, UAE, MAE, 

and SFE are the preferred methods (Cassol et al., 2019; Paraíso et al., 2019; Pimentel-

Moral et al., 2019). 

Among the alternative novel extraction techniques, microwave-assisted extraction 

(MAE) operates through a unique heating mechanism that utilizes radiation within the 

frequency range of 300 MHz to 300 GHz (Hapsari & Setyaningsih, 2021). This method 

enhances heat energy transfer via dielectric heating and frictional resistance, resulting 

in efficient extraction (Routray & Orsat, 2012). Extraction time, temperature, solvent 

properties, and the ratio of product to solvent also influence the effectiveness of this 

method. Supercritical fluid extraction (SFE) employs a supercritical solvent above its 

critical point, which exhibits liquid-like properties while maintaining gas-like 

characteristics (Idham et al., 2021). The low viscosity and high diffusivity coefficient 

of supercritical solvents enhance extraction rates by facilitating solvent flow and rapid 

penetration into the matrix, respectively (Da Silva et al., 2016). Consequently, the 

extraction process is accelerated, enabling efficient extraction of targeted compounds 

from the product matrix. ultrasound-assisted extraction (UAE) is the most commonly 

employed method for extracting phenolic compounds from H. Sabdariffa (Hapsari & 



5 

Setyaningsih, 2021). It relies on the application of ultrasonic waves at specific 

frequencies and amplitudes to disrupt cell walls and facilitate the release of compounds 

into the solvent. This phenomenon is achieved through high shear stress generated by 

cavitation bubbles (Lefebvre et al., 2021). Optimal extraction parameters such as pulse 

cycle, ultrasonic power, time, temperature, and solvent characteristics can be adjusted 

based on the specific product to enhance the extraction efficiency (Pollini et al., 2020). 

Each extraction method follows a specific procedure tailored to the type of phenolics 

and sample matrices involved. Typically, the extraction of bioactive compounds from 

H. Sabdariffa utilizes solvents such as methanol, ethanol and distilled water. The 

calyces of Hibiscus sabdariffa are readily available in the commercial market and have 

versatile applications in both medicinal and non-medicinal domains. One notable 

medicinal application involves their use in the preparation of hot and cold beverages. 

Research conducted by Alzweiri et al. (2011) indicates that hot H. sabdariffa tea is 

particularly beneficial for individuals with hypertension, whereas the cold form of the 

beverage is more suitable for those with hypotension. 

In a separate investigation conducted by Halim et al. (2022), the production of jelly 

candy rich in anthocyanins was examined, utilizing red Hibiscus Sabdariffa extract as 

a natural colorant. The jelly candy formulation involved the incorporation of the 

extract alongside corn syrup and gelling ingredients. The addition of Hibiscus 

Sabdariffa extract enhanced the candy's visual appeal due to the presence of 

anthocyanins, which imparted a vibrant coloration. Furthermore, Kim et al. (2019) 

conducted a study involving the production of functional yogurt incorporating 

Hibiscus Sabdariffa. This research revealed the potential use of Hibiscus Sabdariffa 

as a significant ingredient in dairy products, contributing as a coloring agent and 

enhancing the functional properties. The sensory analysis results indicated a positive 

reception. Notably, the authors found it necessary to supplement the yogurt with 

artificial Hibiscus flavor, as the natural flavor from Hibiscus calyces was not 

adequately perceived. Additionally, various studies have investigated the utilization of 

Hibiscus Sabdariffa in diverse food products such as ice cream, pudding, wine, 

chocolates, and fermented beverages (Adadi et al., 2020; Ai et al., 2021; Homayuni 

Rad et al., 2021). 

Although Hibiscus has the potential to be used in different food products, the direct 

utilization of it into food formulations can be challenging due to three main factors. 

Firstly, the high concentrations of bioactives may negatively affect the taste and odor 



6 

of food, affecting its sensory properties (Nejatian et al., 2022). Secondly, the low 

solubility of bioactives in aqueous media and their tendency to bind with other 

components of the food formulation may result in low bioaccessibility in the human 

body (Choi & McClements, 2020). Finally, bioactives may be sensitive to 

environmental and operational conditions, such as light exposure, oxidation, thermal 

processing, and UV irradiation, which can reduce their effectiveness (Ranadheera et 

al., 2016). Encapsulation techniques have been developed as a solution to the 

sensitivity of bioactives when incorporated directly into food formulations. 

Encapsulation involves entrapping bioactives within a carrier or wall material, which 

can be a protein, polysaccharide, fat, or a combination of different materials (Castro-

Enríquez et al., 2020).  

Encapsulation methods play a crucial role in preserving and stabilizing bioactive 

compounds, and several techniques are commonly employed, including spray-drying, 

freeze-drying, coacervation, liposome formation, and emulsification. Each method 

possesses distinct advantages and limitations, and the selection of an appropriate 

technique depends on the specific properties of the bioactive compound and the 

intended application. 

Freeze-drying and spray-drying techniques are particularly effective for solidifying 

and stabilizing heat-sensitive bioactive components such as polyphenols, 

anthocyanins, and probiotics (Jouki et al., 2021; Kanha et al., 2021; Rishabh et al., 

2021; Šturm et al., 2019). Freeze-drying, being a low-temperature process, is widely 

regarded as a reliable method for preserving the bioactive constituents. However, it 

suffers from drawbacks such as being time-consuming and requiring high energy 

input, thereby posing challenges for large-scale production systems (Kandasamy & 

Naveen, 2022). Consequently, there is an increasing demand for alternative methods 

like spray drying, which can produce encapsulated powders with higher efficiency and 

lower cost on an industrial scale (Piñón-Balderrama et al., 2020). 

Coacervation is a process involving the separation of colloidal systems into two 

distinct phases: one phase consists of hydrocolloids or coacervates, while the other 

phase constitutes a dilute continuous phase. Two types of coacervation methods are 

commonly utilized: simple coacervation and complex coacervation. Simple 

coacervates can be generated by gradually introducing ethanol or sodium sulfate to a 

protein solution (Aloys et al., 2016). On the other hand, complex coacervates can be 

formed by associating two oppositely charged phases in an aqueous environment, 



7 

under controlled parameters such as pH, polymer concentration, and ionic strength. 

Notable examples of these distinct phases include combinations of proteins, chitosan 

and anionic polysaccharides, and proteins and polysaccharides (Baiocco et al., 2021; 

Muhoza et al., 2022; Warnakulasuriya & Nickerson, 2018). Emulsions have gained 

widespread recognition as highly effective encapsulation and delivery systems for a 

wide range of bioactive molecules, including lipophilic, hydrophilic, and amphiphilic 

compounds (McClements & Li, 2010). These emulsion systems offer efficient 

encapsulation, contribute to the preservation of the chemical stability of bioactive 

compounds, and enable controlled release (Karim et al., 2022). Notably, emulsion-

encapsulated polyphenols have demonstrated enhanced biological activities compared 

to their free form, particularly during gastrointestinal digestion (Sun et al., 2018). 

The structure of an emulsion consists of two immiscible liquids, typically oil and 

water, with one serving as the dispersed phase and the other as the continuous phase 

(Tadros, 2013). Emulsions can be further classified based on the distribution of the oil 

and water phases. An oil-in-water (O/W) emulsion refers to a system in which oil 

droplets are dispersed within a continuous water phase, as found in milk or mayonnaise 

(Clark, 2013). Conversely, a water-in-oil (W/O) emulsion is formed by dispersing 

water droplets within the oil phase, akin to the composition of butter. The unique 

characteristics of these different phases directly influence the selection of bioactive 

substances suitable for encapsulation. 

Significant advancements in scientific research have resulted in the development of 

diverse emulsion systems specifically designed for the encapsulation and delivery of 

bioactive compounds, thereby unlocking their remarkable health benefits. These 

emulsion systems play a pivotal role in the food industry by serving as encapsulation 

platforms for incorporating bioactives into various food formulations. One prominent 

emulsion system employed for bioactive encapsulation is High Internal Phase 

Emulsion (HIPEs). 

HIPEs, also known as high internal phase emulsions, are characterized by a substantial 

volume of the internal phase, constituting at least 74% (Shi et al., 2020). This higher 

internal volume fraction distinguishes HIPEs from traditional emulsions. Notably, 

HIPEs offer several advantages in terms of encapsulating and preserving bioactive 

compounds. The elevated volume fraction of the internal phase allows for a greater 

loading capacity of bioactives, enabling the encapsulation of higher concentrations of 

active compounds compared to conventional emulsion systems (Díaz-Ruiz et al., 



8 

2021). Depending on the nature of the bioactives to be encapsulated, the internal phase 

of HIPEs can be composed of either water or oil (McClements et al., 2007). Controlled 

release is another crucial attribute of HIPEs, enabling the gradual and sustained release 

of encapsulated bioactives over time (Jin et al., 2017). This controlled release 

mechanism offers potential for targeted and prolonged delivery of bioactive 

compounds (Katouzian & Jafari, 2017). 

HIPEs find extensive applications in various industries, including food, 

pharmaceuticals, and cosmetics (Gao et al., 2021). They have been effectively utilized 

for encapsulating a wide range of bioactive compounds, such as vitamins, essential 

oils, and plant extracts (Chen et al., 2020; Díaz-Ruiz et al., 2021; Feng et al., 2023). In 

addition to HIPEs, double emulsions (DEs) have gained widespread usage for 

bioactive encapsulation. Double emulsions, also known as multiple emulsions or 

W/O/W emulsions, consist of nested emulsion systems (Clegg et al., 2016). The active 

compound is typically enclosed within the internal aqueous phase, which is surrounded 

by an oil phase and an external aqueous phase (Kaimainen et al., 2015). This layered 

structure provides an additional barrier and a controlled environment for the 

encapsulated compounds. By adjusting the properties of the internal and external 

aqueous phases, controlled release of bioactives can be achieved (Giroux et al., 2016).  

Lecithin, Tween, Span, and PGPR are commonly preferred emulsifiers employed to 

stabilize emulsion systems. Lecithin, derived from sources such as soybeans and eggs, 

is a natural emulsifier with both lipophilic and hydrophilic properties (Wang et al., 

2021). Tween and Span are frequently used together as emulsifiers, with Tweens being 

hydrophilic and Spans being lipophilic (Mosca et al., 2013). Polyglycerol 

polyricinoleate (PGPR), a hydrophobic semi-synthetic emulsifier, is commonly 

utilized for emulsion stabilization (Márquez et al., 2010). However, due to increasing 

concerns regarding health and the environment, particularly in the food industry, there 

is a growing reluctance to use synthetic surfactants as emulsifiers. Synthetic 

surfactants have been found to be detrimental to beneficial microorganisms, contribute 

to environmental pollution, and pose risks to human health (Rasheed et al., 2020). 

Consequently, there is a need to explore eco-friendly alternatives that promote good 

health to replace these synthetic emulsifiers. These emulsifiers and stabilizers, which 

are preferred in food emulsion systems, can cause many side effects such as abdominal 

pain, cramps and diarrhea from a certain dose. In addition, consumers may have 

negative attitudes towards these ingredients, which are in the food additives class. One 



9 

promising alternative is the use of solid particles as stabilizers in emulsions, known as 

Pickering emulsifiers (PE). PEs offer several advantages over surfactant-stabilized 

emulsions. They can prevent droplet or particle coalescence in the dispersed phase and 

reduce toxicity (Chevalier & Bolzinger, 2013; Wu & Ma, 2016). PEs have the potential 

for increased utilization in the food and pharmaceutical industries (Shi et al., 2016). 

The use of proteins as an emulsifier can both prevent these side effects and consumer 

concerns and enrich the nutritional content of the products. Various solid particles, 

including nanoparticles and microparticles, can be employed to stabilize PEs. 

Pickering particles derived from polysaccharides, proteins, and polyphenols have vast 

areas of application. Among these, proteins, particularly plant-based proteins, have 

gained significant attention in recent years. Proteins offer a wide range of options for 

creating emulsions with diverse characteristics. Protein-based nanoparticles provide 

advantages such as improved emulsion stability, controlled release of bioactive 

compounds, and enhanced texture and mouthfeel in food products. Their amphoteric 

nature, exhibiting both hydrophilic and hydrophobic properties, makes proteins 

particularly suitable for producing stable food-grade Pickering emulsions (Lam et al., 

2014). In summary, proteins serve as versatile and nutritious emulsifiers, enabling the 

development of various emulsion-based food products with enhanced stability and 

functionality. Plant-based proteins such as soy, pea, kafirin, and zein offer a wide range 

of options for Pickering particles and emulsion stabilization (De Folter et al., 2012; 

Liu & Teng, 2016; Shao & Tang, 2016; Xiao & Huang, 2015). Soy protein isolate 

(SPI), a prominent soy protein product commercially available, is produced through a 

conventional alkali extraction-acid precipitation process (Verfaiilie et al., 2023). These 

proteins possess an amphipathic nature, exhibiting both hydrophilic and hydrophobic 

properties, which enables them to diffuse, adsorb, and stabilize the interfaces of oil 

droplets during emulsification (Djuardi et al., 2020; Yan et al., 2021). Table 2.2 

presents the different studies about the encapsulation of bioactives within High 

Internal Phase Pickering Emulsions (HIPPEs) and Double Emulsions (DEs) stabilized 

by soy protein isolate (SPI). 

 

 

 

 



10 

Table 1.2: Encapsulation of bioactives within HIPPEs and DEs stabilized SPI. 

Emulsion Emulsifiers/Stabilizers Bioactive 

compound 

Effect of 

encapsulation 

Reference 

HIPPE Soy protein isolate β-Carotene The retention of 

β-carotene in 

HIPPEs was 

found to be 

74.1%, which is 

significantly 

higher 

compared to 

that in corn oil. 

Wang et 

al., 2022 

HIPPE Soy protein isolate – 

epigallocatechin-3-

gallate covalent 

composite microgel 

particles 

β-carotene After 42 days 

of storage, the 

HIPPEs 

exhibited a 

higher retention 

rate and 

bioaccessibility 

of β-carotene  

Geng et al., 

2022 

HIPPE Soy protein isolate–

dextran complexes 

Quercetin High 

encapsulation 

efficiency for 

(98.19 %) 

Du et al., 

2022 

DE Bacterial cellulose 

nanofibers/soy protein 

isolate/chitosan 

complex 

Curcumin Higher 

bioaccessibility 

of curcumin 

(77.4%) 

compared to 

extract (32.8 % ) 

Shen et al., 

2023 

DE Soybean protein-

polyglutamic acid 

complex 

Nattokinase Increased 

bioaccessibility 

up to 80.69%. 

Li et al., 

2023 

 



11 

Several studies in the literature have provided evidence that the addition of lecithin 

can enhance the stability of emulsions containing soybean protein and whey protein, 

even when the conditions are close to the proteins' isoelectric point (Mantovani et al., 

2013; Scuriatti et al., 2003). Additionally, Comas et al. (2006) observed improved 

stability in emulsions composed of native or denatured soybean protein isolates, 

sunflower oil, and water when lecithin was included. The interaction between proteins 

and lecithin is attributed to hydrophobic interactions, which play a crucial role in 

maintaining the protein-phospholipid interactions. Recent research conducted by Li et 

al. (2014) further supports the notion that lecithin and soybean protein can interact 

through electrostatic and hydrophobic interactions, leading to favorable changes in 

protein conformation and improved emulsifying properties. 

 

  



12 

 



13 

2.  MATERIALS AND METHODS 

2.1 Materials 

Soy protein isolate (90% protein content) was supplied by Vegrano Co., Ltd. (Istanbul, 

Türkiye). Corn oil was purchased from a local supermarket (Istanbul, 

Türkiye). Hibiscus Sabdariffa L. calyces were supplied by TOS-The Organic Spices 

Co., Ltd. (Antalya, Türkiye). Soybean lecithin was purchased from Sigma-Aldrich. All 

the chemicals utilized in this study were analytical grade. 

2.2 Preparation of Aqueous Hibiscus Extract (AHE) 

Extraction procedure was conducted based on the study of Pinela et al. (2019) with 

some modifications. Dried Hibiscus calyces were ground to obtain powder with a 

blender (Kiwi Classic 575, Bad Homburg, Germany) for 30 s at 22,000 g and stored 

at 4°C temperature in the dark, until use. After weighing 1.5 g of powder into falcon 

tubes (50 ml), 50 ml of distilled water was added (Pinela et al., 2019). The prepared 

mixtures were vortexed for 10 s and kept in an ultrasonic bath (USC900TH; VWR, 

Radnor, PA, USA) for 30 minutes. All samples were centrifuged at 2700 x g (4000 

RPM) for 10 minutes at 4 °C (Universal 32R; HettichZentrifugen, Tuttlingen, 

Germany), and the supernatants were collected. Finally, the resulting supernatant was 

filtered using Whatman filter paper No. 4 to obtain the desired extract.  

2.3 Preparation of Soy Protein Isolate (SPI) Gel 

Soy protein isolate (SPI) solutions with protein concentrations of 2%, 4%, and 6% 

were prepared. The solutions were left overnight with stirring using a magnetic stirrer 

and centrifuged to remove non-hydrated proteins (Urbonaite et al., 2015). After 

complete hydration, SPI solutions were heated in a water bath at 95 °C for 15 minutes, 

followed by rapid cooling in an ice bath to reach room temperature (Huang et al., 

2022).  



14 

2.4 Preparation of Emulsions 

2.4.1 High internal phase pickering emulsions (HIPPEs) 

Different emulsions with varying SPI gel and lecithin concentrations were prepared. 

These include only corn oil and pure aqueous Hibiscus extract (control), SPI (2%, 4% 

and 6%) and Hibiscus extract mixture and corn oil (without lecithin), containing 

Hibiscus extract and corn oil (2%, 4% and 4% lecithin concentration) and finally SPI 

(2%, 4% and 6%)-Hibiscus extract and corn oil (with 2%, 4% and 4% lecithin 

concentrations) emulsions (Table 2.1). Initially, 20 mL corn oil mixture was prepared 

at different lecithin concentrations (0%, 2%, 4%, 6%). To ensure efficient mixing of 

the lecithin with the corn oil, the mixture was stirred on a magnetic stirrer at 5000 rpm 

for 10 minutes. 80 mL of SPI gel solutions were mixed with aqueous Hibiscus extract 

using a magnetic stirrer (5000 rpm, 3 min). Mixing continued until a homogeneous 

SPI-Hibiscus mixture is obtained. First, SPI-Hibiscus extract was mixed with lecithin-

free corn oil. Then, SPI-Hibiscus mixture was added to corn oil samples containing 

lecithin (2%, 4%, 6%). Finally, pure Hibiscus extract (without SPI gel) was mixed 

with corn oil phase containing lecithin. After homogenization, the presence/absence 

of W/O emulsions with 80% internal volume fraction was checked. In each 

homogenization process, Ultraturrax (IKA, T18) was run at 12000 rpm for 3 minutes. 

Table 2.1: HIPPEs with different formulations. 

Phase Corn oil (without 

lecithin) 

Corn oil (2% 

lecithin) 

Corn oil (4% 

lecithin) 

Corn oil (6% 

lecithin) 

Hibiscus extract  Control L2 L4 L6 

Hibiscus extract 

+ 2 % SPI gel 

S2 L2S2 L4S2 L6S2 

Hibiscus extract 

+ 4 % SPI gel 

S4 L2S4 L4S4 L6S4 

Hibiscus extract 

+ 6 %  

SPI gel 

S6 L2S6 L4S6 L6S6 



15 

2.4.2 High internal phase pickering-double emulsions (HIPP-DEs) 

Each 50 mL of W/O HIPPE, which did not show phase separation after 24 hours, was 

homogenized with 50 mL 6% SPI gels with the help of Ultraturrax (12000 rpm, 3 min) 

and it was examined whether W/O/W was formed or not. Table 2.2 presents HIPP-Des 

with different formulations  

Table 2.2: HIPP-DEs with different formulations. 

W/O/W HIPP-DEs Formulation 

L4-D L4+6% SPI gel 

L6-D L6+6% SPI gel 

L2S6-D L2S6+6% SPI gel 

L4S2-D L4S2+6% SPI gel 

L4S4-D L4S4+6% SPI gel 

L4S6-D L4S6+6% SPI gel 

L6S2-D L6S2+6% SPI gel 

L6S4-D L6S4+6% SPI gel 

L6S6-D L6S6+6% SPI gel 

2.5 Storage Stability 

Following the preparation of the emulsions, they were promptly stored at a temperature 

of 4 °C for a duration of 24 hours. During this storage period, stability observation was 

conducted to determine whether any phase separation occurred within the emulsions. 



16 

2.6 Encapsulation Efficiency 

The encapsulation efficiency (EE) of the HIPPEs and HIPP-DEs stabilized by lecithin 

and SPI gel was determined based on the study of Du et al. (2022) with slight 

modifications. 1 mL of each emulsions and 5.0 mL of ethanol was vortexed for 30 

seconds. Then centrifugation was applied at 8000 x g and for 10 minutes. The Total 

Anthocyanin Content (TAC) and Total Phenolic Content (TPC) of the supernatant was 

analyzed. The encapsulation efficiency of each emulsion was calculated using Eqs.  

(2.1) and (2.2). 

EEA (%) = 1 −
𝑇𝐴𝐶 𝑜𝑓 𝑠𝑢𝑝𝑒𝑟𝑛𝑎𝑡𝑎𝑛𝑡 

𝑇𝐴𝐶 𝑜𝑓 𝑒𝑚𝑢𝑙𝑠𝑖𝑜𝑛
 (2.1)  

EEP (%) = 1 −
𝑇𝑃𝐶 𝑜𝑓 𝑠𝑢𝑝𝑒𝑟𝑛𝑎𝑡𝑎𝑛𝑡 

𝑇𝑃𝐶 𝑜𝑓 𝑒𝑚𝑢𝑙𝑠𝑖𝑜𝑛
 (2.2)  

2.7 Determination of Emulsifying Properties 

The emulsifying properties of HIPPEs and HIPP-DEs were analyzed according to the 

method of Li et al. (2014). 50 mL of emulsions were sampled immediately after 

preparation and after 10 min. All samples were diluted 100 times using 0.1% sodium 

dodecyl sulfate (SDS). The absorbance of the samples immediately after preparation 

(A0) and after 10 min (A10) using a spectrophotometer (VWR UV-3100PC, VWR 

International, San Francisco, CA). The emulsifying activity index (EAI) and emulsion 

stability index (ESI) were calculated using Eqs. (2.3) and (2.4), respectively. 

ESI (min) = 
A0

A0−A10
 x (T10 − T0) (2.3)  

EAI (m
2

g⁄ ) = 2 x T x 
A0 x N

10000 X  X L X C
 (2.4)  

Where A0 is the absorbance at 0 min; A10 is the absorbance at 10 min; T10 is the time 

at the 10 min; T0 is the time at the 0 min; T equals to 2.303; N is the dilution factor 

(100);  is the proportion of the oil phase (0.2); L is the thickness of the cuvette (1 

cm); C is the concentration of the SPI (g/ml).  



17 

2.8 Particle size, PDI and Zeta Potential 

The particle size, PDI and zeta potential of the each emulsions were measured by 

dynamic light scattering device (Nano-ZS, Malvern Instruments, Malvern, UK) based 

on the study of Karaca et al. (2011). 

2.9 Creaming Stability 

The creaming stability analysis was carried out according to the method of Aslan & 

Dogan. (2018) with slight modifications. Creaming index calculation was used to 

determine the tendency of emulsions to separate into two phases, and it involves filling 

a falcon tube with 30 ml of emulsion sample and monitoring it for 14 days at 4°C. The 

height of the clear liquid layer on top of the emulsion (serum) and the total height of 

the emulsion were measured on days 1, 7, and 14. To calculate the creaming index 

value, the height of the serum (Hs) was divided by the total height of the emulsion (Ht) 

and multiplied by 100. The resulting creaming index value can range from 0 to 100 

and indicates the degree of creaming tendency of the emulsion, with higher values 

indicating a greater tendency to separate (Aslan & Dogan, 2018).  

The following formula (2.5) was used to calculate the creaming index value: 

Creaming index =
𝐻𝑆

𝐻𝑡
∗ 100 (2.5)  

2.10 Centrifugation Stability 

The HIPPEs and HIPP-DEs (30 mL) were transferred into 50 mL falcon tubes and 

subjected to centrifugation at 5000 rpm for a duration of 20 minutes (Li et al., 2022). 

The visual appearance of the emulsions after centrifugation was evaluated.  

2.11 In Vitro Gastrointestinal Digestion 

The in vitro gastrointestinal digestion protocol was conducted, making certain 

modifications based on the research conducted by Minekus et al. in 2014. This protocol 

aimed to simulate the process of digestion in the oral, gastric and intestinal digestions. 

During the simulation of oral digestion, each emulsion (5 mL) was mixed with salivary 

juice (4 mL), 0.3 mol/L CaCl2 (25 µL), and distilled water (975 µL). The mixture was 

then incubated for two minutes at 37 °C in a shaking water bath (Memmert SV 1422, 



18 

Memmert GmbH & Co. Nürnberg, Germany). Next, the simulation of gastric digestion 

took place by adding gastric juice (7.5 mL), pepsin (1.6 mL), and CaCl2 (5 µL) to the 

solution. The pH of the mixture was adjusted to 3.0 using 1 mol/L HCl, and distilled 

water was added for volume adjustment. The mixture was incubated once again for 

two hours at 37 °C in a shaking water bath. After simulating gastric digestion, 5 mL 

aliquots were taken from each mixture. The gastric mixture was then combined with 

intestinal juice (8.25 mL), pancreatin (3.75 mL), bile (1.875 mL), and CaCl2 (30 µL). 

The pH of the mixture was adjusted to 7.0 using NaOH, and the total volume of the 

mixture was completed to 30 mL using distilled water. The mixture was incubated for 

two hours in a shaking water bath at 37 °C. Simultaneously, a blank sample containing 

only water instead of emulsion was prepared. Following the incubation period, the 

simulated gastric and intestinal samples were centrifuged for five minutes at 4000 rpm 

and 4 °C (Hettich, Tuttlingen, Germany). The resulting supernatants were stored at -

20 °C until further analysis. 

2.12 The Quantification of Polyphenols with HPLC-DAD 

Polyphenols in the samples were quantified using the method described by Capanoglu 

et al. (2008). Prior to analysis, the samples were passed through a 0.45 μm membrane 

filter (Waters 2996) and then injected into a Waters 2695 HPLC system equipped with 

a PDA detector. The column used was a Supelcosil LC-18 (25 cm x 4.60 mm, 5 μm) 

from Sigma-Aldrich in Steinheim, Germany. Spectral measurements were performed 

at wavelengths of 280 nm, 312 nm, and 360 nm. The eluents used were TFA/MQ water 

(1 mL/L; eluent A) and TFA/acetonitrile (1 mL/L; eluent B). The injection volume 

was 10 mL or the flow rate was set at 1 mL per minute. The gradient elution profile 

was as follows: at 0 minutes, 95% solvent A and 5% solvent B; at 45 minutes, 65% 

solvent A and 35% solvent B; at 47 minutes, 50% solvent A and 50% solvent B; and 

at 54 minutes, the system returned to its initial state. Phenolic acids were quantified 

using authentic standards. The results of each analysis were reported as mg/100 g dry 

sample weight, and each analysis was performed in triplicate. 



19 

2.13 Spectrophotometric Assays 

2.13.1 Total anthocyanin content (TAC) 

The total anthocyanin content (TAC) was determined using the pH-differential method 

as described by Giusti & Wrolstad. (2001). To perform the analysis, 1 mL of the 

extracted solution was transferred into a 10 mL volumetric flask to prepare two 

dilutions of the sample. One dilution was adjusted to volume with potassium chloride 

buffer at pH 1.0, and the other dilution was adjusted to volume with sodium acetate 

buffer at pH 4.5. Both dilutions were allowed to equilibrate for 15 minutes. The 

absorbance of each dilution was then measured at 510 nm and 700 nm, using a blank 

cell filled with distilled water as the reference (Giusti & Wrolstad, 2001). 

2.13.2 Total phenolic content (TPC) 

Total phenolic content (TPC) assay was carried out according to the method of 

Singleton and Rossi (1965). The experiment with Folin–Ciocalteu reagent was 

implemented at 765 nm. Calibration curve was plotted using gallic acid in the range 

of 0.01–0.4 mg/ml and TPC results were reported as mg gallic acid equivalents (GAE) 

per 100 g dw sample.  

2.13.3 Antioxidant activity 

The antioxidant capacities of the infusions were determined by both 2,2-diphenyl-1-

picrylhydrazyl (DPPH) (Molyneux, 2004) and 2,2‘-azinobis (3-ethylbenzothiazoline-

6-sulfonic acid) (ABTS) (Nenadis et al., 2004) assays. Measurements were performed 

at 450 and 517 nm, respectively. Calibration curves were plotted using Trolox in the 

range of 0.01–0.4 mg/ml and results were reported as Trolox equivalents (TE) per 100 

g of dw sample. 

2.13.4 Bioaccessibility calculations 

The bioaccessibility was calculated using following formula; 

 

Bioaccesibility (%) = (BCdigested/BCnon-digested) x100 

 

Where BCdigested was the amount of bioactive compounds (TAC, TPC) recovered in 

the supernatants of the centrifuged final digesta. BCnon-digested was the amount of 

bioactive compounds in non-digested rosehip infusion. 



20 

2.14 Statistical Analysis 

All experiments were conducted with a minimum of three replicates to ensure 

statistical robustness. The error bars displayed on the figures represent the standard 

deviations of the data. The results are presented as the mean value ± the standard 

deviation. For statistical analysis, the SPSS software (version 20.0, SPSS Inc., 

Chicago, IL, USA) was utilized. The processing conditions were evaluated using a 

one-way analysis of variance (ANOVA), followed by a Tukey post hoc test. A 

significance level of p < 0.05 was considered statistically significant. 

 

 

 

 

 

 

 



21 

3.  RESULT AND DISCUSSION 

3.1 Storage Stability 

Storage stability of emulsions were determined based on whether any phase separation 

occurred during 24 h storage at 4 °C. Visual examination revealed phase separation in 

the control emulsion after approximately 2 min at room temperature. Figure 3.1 

exhibits products stabilized by only SPI gel. As can be observed, these products could 

not provide stability for 24 h and even phase separation occurred in about 5 minutes. 

For this reason, it has been observed that a stable emulsion system cannot be obtained 

by using only SPI gel. 

 

Figure 3.1: HIPPEs stabilized by only different SPI gel concentrations. 

Figure 3.2 represents products stabilized by only lecithin. As can be observed, only L2 

could not provide stability for 24 h and even phase separation occurred in about 8 



22 

minutes. However, L4 and L6 presented 24 h stability against phase separation. For 

this reason, it can be stated that increase in lecithin concentration provided enhanced 

stability for emulsions. 

 

Figure 3.2: HIPPEs stabilized by only different lecithin concentrations. 

In the case HIPPEs containing both SPI gel and lecithin, variations in storage stability 

against phase separation were observed. Figure 3.3 shows highly stable HIPPEs 

against phase separation for 24 h and even phase separation could not occured for 45 

h. As it can be stated from here, a synergistic mechanism of action became dominant 

in HIPPEs where SPI gel and lecithin were used at different concentrations and 

significantly prevented phase separation.  



23 

 

Figure 3.3: HIPPEs stabilized by different SPI gel and lecithin concentrations. 

Each 50 mL of stable W/O HIPPE, which did not show phase separation after 24 hours, 

was homogenized with 50 mL 6% SPI gels. Thus, HIPP-DEs can be termed as HIPPEs 

with double emulsions morphology. All HIPP-DEs showed a excellent resistance to 

phase separation. Figure 3.4 demonstrates highly stable HIPP-DEs against phase 

separation for 24 h. 

 

Figure 3.4: HIPP-DEs stabilized by different SPI gel and lecithin concentrations. 

According to these observed results, it can be said that lecithin and soy protein isolate 

present a synergistic mechanism of action. Different studies have indicated that 

lecithin can interact with zein nanoparticles and exhibit synergistic effects to enhance 

emulsion stability (Chuacharoen & Sabliov, 2016; Dai et al., 2016). Taking this into 

consideration, lecithin was incorporated to improve the stability of SPI-stabilized High 

Internal Phase Pickering Emulsions (HIPPEs). Additionally, a two-step emulsification 



24 

process can generate a double emulsion morphology. The findings suggest that lecithin 

plays a role also in forming the HIPP-DE structure, potentially because lecithin is an 

lipophilic surfactant that can stabilize water-in-oil emulsions on its own (Knoth et al., 

2005). 

3.2 Creaming Stability 

The creaming in emulsions is caused by instabilities resulting from the differences in 

densities between the oil and water phases, as well as the gravitational force acting on 

them (Tadros, 2004). Therefore, measuring the amount of cream that separates over a 

given time period is a useful method for evaluating the stability of the emulsion. The 

creaming stability of the emulsions was evaluated by both CI calculation and visual 

observation during storage for 12 h, 1, 5 and 9 days, and the results were presented in 

Figs. 3.5 and 3.6. While the CI value did not show significant difference between 

HIPPEs on the 12th, differences became observable on day 1 (p<.05). The findings of 

this study also indicate that in HIPPEs with 6 % SPI gel concentration, the increase in 

soybean lecithin concentration from 2% to 4 % resulted in a decrease in the creaming 

rate from 24.12 ± 0.45% to 17.79 ± 0.32% (day 1). These results suggest that the 

addition of lecithin enhances the stability of the system, particularly at a concentration 

of 4%. However, further increases in the lecithin concentration up to 6% in HIPPEs 

result in a significant increase in CI. Similar results were also obtained for day 5. ). A 

different result was obtained for HIPP-DEs. Here, the creaming index value did not 

show a significant difference between emulsions (p<.05). The findings of Wang et al. 

(2017) indicate that as the lecithin concentration increases from a certain 

concentration, the creaming index (CI) value also increases, indicating a higher degree 

of instability in the emulsion. The increase in CI is attributed to the sensitivity of the 

adsorbed layer to external forces, as the instability observed can be explained by the 

inability of the interfacial protein layer to withstand surface pressure, resulting in the 

collapse of protein molecules into the serum phase and noticeable creaming of the 

emulsions (Wang et al., 2017). Thus, it can be concluded that the stability of the 

emulsion was improved to a certain extent by the addition of lecithin, and that a 

concentration of 4% was the most effective concentration for enhancing the creaming 

stability of the emulsion. In HIPP-DEs, on day 5 and day 9, CI values did not show a 

statistical difference between emulsions (p<.05). 



25 

 

Figure 3.5: Changes in creaming index (CI) of HIPPEs during storage. 

 

Figure 3.6: Changes in creaming index (CI) of HIPP-DEs during storage. 

3.3 Centrifugation Stability 

The HIPPEs and HIPP-DEs (30 mL) were transferred into 50 mL falcon tubes and 

subjected to centrifugation at 5000 rpm for a duration of 20 minutes (Li et al., 2022). 

The visual appearance of the emulsions after centrifugation was evaluated. As can be 

clearly observed from Figure 3.7 and Figure 3.8, phase separation was observed only 

in L4 and L6 in HIPPEs. No phase separation was observed in HIPP-DEs. It can be 

said that the reason for this situation is that emulsions using only lecithin cannot resist 

centrifugation, and SPI gel plays an important role in providing emulsion stability. 

0

5

10

15

20

25

30

35

40

45

18 hours day 1 day 5 day 9

C
re

a
m

in
g

 I
n

d
ex

 (
%

)

Storage time

L4 L6 L2S6 L4S2 L4S4 L4S6 L6S2 L6S4 L6S6

0

2

4

6

8

10

12

14

16

18

18 hours day 1 day 5 day 9

C
re

a
m

in
g

 I
n

d
ex

 (
%

)

Storage time

L4-D L6-D L2S6-D L4S2-D L4S4-D L4S6-D L6S2-D L6S4-D L6S6-D

a a a

a
a

a
b b

bb b b

a
a

a a

v

a a

b b
b

b b b

a
a a

a
a

a
a

a a a
a

a

a 
a a 

a a 
a 

a 
a a a 

a a 

b b b 

b b 
b b b b b b 

a 
a a a a a 

c c 
c 

b 
c c 

c 



26 

 

Figure 3.7: Degree of phase separation of L4 and L6 after centrifugation. 

 

Figure 3.8: HIPP-DEs after centrifugation. 

3.4 Determination of emulsifying properties 

The large interfacial area between the water and oil phases results in poor 

thermodynamic stability (Abdulredha et al., 2022). Therefore, flocculation, 

coalescence, and phase separation can occur in the structure of emulsions (Jin et al., 

2021). The ability of a protein to adhere to the oil/water interface is determined by its 

emulsifying property (Liu & Wang, 2021). Determining the emulsifying properties of 

proteins that localize themselves at the oil/water interface in order to reduce the 

interfacial tension (Baldino et al., 2023) is necessary to achieve the desired emulsion 

stability. Also, the effects of commonly used emulsifiers on the specific emulsion 

systems is a subject that needs to be examined. In this study, the emulsification activity 



27 

index (EAI) and emulsification stability index (ESI) were measured to determine 

emulsifying property. The ESI measures the turbidity of an emulsion, whereas the EAI 

indicates the interfacial area (m2) that is stabilized for each weight (g) unit of a protein 

(Cha et al., 2019). Soy protein isolate (SPI), an important plant-based protein, 

possesses amphiphilic properties that allow its use as an emulsifier in food products 

(Deng, 2021; Liao et al., 2021). Lecithin, mixture of phospholipids, 

phosphatidylcholine,  ethanolamine,  and  inositol, is one of the  emulsifiers   that   

actively   reduces   the   surface tension in emulsion production (Wiyani et al., 2016).  

This study aimed to evaluate the EAI and ESI of HIPPEs and HIPP-DEs 

stabilized by SPI and soy lecithin. The emulsions were shown to have different EAI 

and ESI values were depicted in Table 3.1. In HIPPEs containing 4% and 6% lecithin, 

EAI and ESI values increased statistically when the SPI concentration increased from 

2% to 4% (p<.05). The same observation was also achieved for HIPP-DEs (Table 3.2). 

In the study of Pirestani et al. (2017), an increase in EAI and ESI values with increasing 

SPI concentration was attributed to the adsorption of SPI on the o/w interface can 

stabilize the system through viscoelastic film formation around oil droplets. Similarly, 

Guan et al. (2020) showed that the increase in EAI and ESI values was related to the 

emulsifying capabilities of SPI, which makes oil droplets well dispersed in the 

emulsion system. Interestingly, no statistically significant increase occurred when the 

SPI concentration increased from 4% to 6% (p<.05). This phenomenon may be due to 

the protein concentration at the interface exceeding the maximum load and the excess 

protein left from the interface flocculating by forming hydrophobic interaction, which 

results in a constant EAI value (Guan et al., 2020). In addition, each HIPP-DE has 

higher EAI and ESI values than its HIPPEs. The lowest EAI and ESI values were 

observed for L2S6 and L4 emulsions in HIPPEs and for L2S6-D and L4 emulsions in 

HIPPDEs. However, L6 and L6-D showed higher EAI and ESI values than these 

emulsions (p<.05). 

The alterations in EAI and ESI as a function of different lecithin concentrations 

are presented. The findings revealed that as the concentration of lecithin increased 

from 2% to 4%, both EAI and ESI increased gradually and attained their peak values 

at a lecithin concentration of 4%. However, both EAI and ESI decreased slightly at 

higher concentration of lecithin (6%) (p<.05). This indicated that an intermediate level 

of lecithin addition led to significant improvement in emulsifying properties. In the 

study of Zou et al. (2020), this change was attributed to the localization of the majority 



28 

of phospholipid molecules on the aqueous side of the oil-water interface, resulting in 

the formation of protein-lecithin complexes that exhibit enhanced emulsifying 

properties. The elevation in EAI might be due to the increased exposure of 

hydrophobic groups which strengthened the interactions between proteins and lipids 

(Zou et al., 2020). In addition, the decrease in EAI and ESI values with the increase in 

lecithin concentration from 4% to 6% was explained by the decrease in surface 

hydrophobicity. 

Table 3.1: EAI and ESI values of HIPPEs. 

HIPPEs EAI (m2/g) ESI (min) 

L4 49.21±0.56f 39.12±0.45e 

L6 70.19±0.27e 30.29±1.09e 

L2S6 77.12±0.65d 36.46±1.34d 

L4S2 85.21±0.34b 45.11±3.67b 

L4S4 89.92±0.97a 49.83±0.34a 

L4S6 89.96±1.17a 49.96±0.34a 

L6S2 80.12±1.21c 39.12±2.34c 

L6S4 86.12±0.10b 46.14±1.34b 

L6S6 86.18±0.03b 46.10±1.18b 

*The results are represented as mean ± standard deviation (n=3). 
*Different lowercase letters indicate significant differences between different emulsions (p <0.05) 

 

 

 



29 

Table 3.2: EAI and ESI values of HIPP-DEs. 

HIPP-DEs EAI (m2/g) ESI (min) 

L4-D 79.11±0.12e 45.21±0.64e 

L6-D 80.29±0.78e 45.68±1.34e 

L2S6-D 85.22±0.64d 57.12±1.24d 

L4S2-D 95.01±1.14b 60.09±0.04b 

L4S4-D 99.12±1.21a 69.28±2.13a 

L4S6-D 99.16±3.45a 69.92±3.21a 

L6S2-D 90.09±9.98c 60.62±1.67c 

L6S4-D 95.45±1.24b 66.69±1.64b 

L6S6-D 95.56±1.35b 67.09±0.03b 

*The results are represented as mean ± standard deviation (n=3). 
*Different lowercase letters indicate significant differences between different emulsions (p <0.05) 

3.5 Particle size, zeta potential and PDI  

3.5.1 Particle size 

The particle size characteristics of multiple emulsions are depicted in Figure 3.9 and 

Figure 3.10. In emulsions containing 6% SPI gel, the particle size value exhibited a 

statistically significant decrease as the lecithin concentration increased from 2% to 4%. 



30 

The smallest aggregates were observed at the lecithin concentration of 4%. However, 

when the lecithin concentration was increased from 4% to 6%, the particle size value 

increased at 2% and 4% SPI gel concentrations (p<.05). According to Sun et al. (2018), 

as the lecithin concentration increased, the size of whey protein aggregates also 

increased. According to the study by Teng et al. (2020), the particle size of the 

emulsion initially decreased with increasing lecithin concentration up to a certain 

point. However, further increase in lecithin concentration resulted in an increase in 

particle size. This can be explained by the displacement solubilisation principle, where 

excessive lecithin concentration caused the whey protein at the oil-water interface to 

be replaced by lecithin, leading to an increase in the lecithin content in the aqueous 

phase (Martinez et al., 2011). This reduced the amount of protein at the interface, 

resulting in larger particle sizes in the emulsion. The interaction between lecithin and 

whey protein, causing partial unfolding of the protein, was suggested as one possible 

reason for the observed increase in particle size (Teng et al., 2020). Additionally, the 

formation of complexes between lecithin and protein could also contribute to the 

increase in particle size. On the other hand, the decrease in particle size with increasing 

lecithin concentrations was attributed to the formation of a reverse micelle or vesicle-

like structure between protein and lecithin, which led to a reduction in particle size 

(Dai et al., 2016). Similar trends in particle size changes with increasing lecithin 

concentrations were observed in studies conducted by Wang et al. (2017) and Zou et 

al. (2020).  

The particle size of HIPPEs with 4% and 6% lecithin concentrations exhibited a 

significant reduction as the SPI gel concentration increased from 2% to 6%. 

Specifically, the HIPPE with the lowest particle size (400.03±2.23 nm) was observed 

in the L6S6 (6% lecithin and 6% SPI gel) emulsion. This trend was also observed in 

HIPP-DEs, where the lowest particle size(352±3.32 nm) was obtained in L6S6-D (6% 

lecithin and 6% SPI gel). In the study by Sirovec et al. (2022), it was observed that the 

addition of pea protein isolate in the emulsification process of oregano and rosemary 

extracts caused a significant decrease in particle size, with increasing concentrations 

of pea protein isolate showing a stronger effect. Sohn et al. (2017) explained that 

increasing the emulsifier concentration leads to improved coverage of oil droplets at 

the interface, which ultimately leads to a reduction in the size of emulsion droplets. 
 



31 

3.5.2 Zeta potential 

Figure 3.9 and 3.10 illustrate the variation in zeta potential among different emulsions. 

In general, emulsions with higher zeta potential values exhibit enhanced stability due 

to the increased repulsion between droplets, which is a result of electrostatic forces. 

This repulsion prevents droplet coalescence (Zu et al., 2019). Zeta potential serves as 

a quantitative measure of the  surface charge density of proteins and acts as an indicator 

of potential stability of the emulsion system (Li et al., 2020). A higher absolute value 

of negative zeta potential indicates stronger electrostatic repulsion between droplets, 

leading to improved emulsion stability (Tamnak et al., 2016). In HIPPEs, the zeta 

potential was found to be negative and their absolute values increased with increasing 

SPI gel concentration from 2 % to 4 % (at the 4 % lecithin concentration). The highest 

absolute value of the zeta potential (-41.21 ± 1.23 mV) was observed for L4S4. In the 

study of Kwaamba et al. (2015), the increase in the absolute value of zeta potential 

was attributed to changes in the secondary structure of whey protein induced by soy 

lecithin, causing the protein structure to become more extended. Conformational 

changes in protein, resulting in the exposure of negatively charged whey protein 

residues and a decrease in zeta potential (Kwaambwa, Maikokera, & Nermark, 2015). 

As a result, this could reduce the exposure of positively charged groups and increase 

the negative charge of the protein. However, from 4 % to 6 % SPI gel concentration, 

absolute zeta potential value started to decrease. It was attributed that, an increase in 

protein content from a certain limit can lead to the formation of protein aggregates that 

replaced the protein at the oil-water interface, resulting in decreased emulsion stability 

(Li et al., 2018). Other studies have also shown that high protein concentrations can 

cause protein molecules to aggregate, making it difficult for water molecules to 

penetrate them and leading to decreased stability of the emulsions (Berton‐Carabin et 

al., 2014). Also, when the lecithin concentration increased from 4 % to 6 %, decrease 

in absolute zeta potential values was observed for L6S2, L6S4 and L6S6. From a 

certain concentration, the interaction between lecithin and SPI results in electrostatic 

and hydrophobic interactions, which significantly affects the functional properties of 

SPI. The results of the research also suggest that in HIPPEs that contain 6% SPI gel 

the absolute ζ-potential increased from 30.88 ± 1.12 to 35.7 ± 0.98 as the lecithin 

concentration increased from 2% to 4%, with this change being significant (P < 0.05) 

at a concentration of 4%. This decrease can be attributed to the unfolding of proteins 



32 

caused by lecithin binding, which facilitates the exposure of negatively charged groups 

in SPI, thus reducing the zeta-potential (Zou et al., 2020). In the literature, there are 

some studies which states that the lecithin addition increases the absolute zeta-

potential of casein and whey protein (García-Moreno et al., 2017; Wang et al., 2017). 

Additionally, the electrostatic attraction between negatively charged phosphate groups 

in lecithin and positively charged amine groups in SPI also contributes to the reduction 

in zeta-potential (Surh et al., 2006). HIPP-DEs, as shown in Figure ??, exhibited higher 

absolute zeta potential values compared to HIPPEs. Among the HIPP-DEs, L4-D and 

L6-D had lower absolute zeta potential values compared to the other samples. 

However, there were no statistically significant difference observed among the 

remaining HIPP-DEs. It should be noted that HIPP-DEs have a higher concentration 

of soy protein isolates compared to HIPPEs. This higher concentration of soy protein 

isolates may have contributed to the increase in negative surface charge, thus resulting 

in the significant difference in zeta potential values. 

 

Figure 3.9: Change in the particle size and zeta potential between different HIPPEs. 

 

 

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33 

 

Figure 3.10: Change in the particle size and zeta potential between different HIPP-

DEs. 

3.5.3 PDI 

The graph in Figure 3.11 illustrates how the PDI varies among emulsions. Increasing 

the concentration of soy protein isolate (SPI) from 2% to 6% led to significant 

decreases in the polydispersity index (PDI) of HIPPEs containing 4 % and 6 % lecithin. 

Lowest PDI values were observed when the SPI concentration was 6%. Same decrease 

was also observed HIPP-DEs (Figure 3.12). In the study of Teng et al. (2020), results 

were consistent. SPI functions as an emulsifier by reducing tension at the interface 

between oil and water, preventing oil droplets from aggregating, and increasing the 

thickness of the interfacial layer (Hebishy et al., 2015). This leads to increased 

emulsion stability. At a certain concentration, the interaction between lecithin and SPI 

induces electrostatic and hydrophobic interactions, which significantly affect both the 

emulsion properties and functional properties of proteins (Anal et al., 2019). 

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L4-D L6-D L2S6-D L4S2-D L4S4-D L4S6-D L6S2-D L6S4-D L6S6-D

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34 

 

Figure 3.11: PDI values of HIPPEs. 

 

Figure 3.12: PDI values of HIPP-DEs. 

3.6 Encapsulation Efficiency 

The Figure 3.13 shows EEA and EEP values of different HIPPEs. As it is shown, L4, 

L6 and L2S6 could not present significant difference between EEA and EEP values 

(p<.05).  The L6S6 had the highest (88.9, 83.1) EEA (%) and EEP (%) values, 

respectively. In HIPPEs containing 4 % lecithin, a statistically significant increase is 

observed in the EEA and EEP values of with increasing SPI gel concentration from 

L4S2 to L4S6. The same situation was observed in emulsions containing 6% lecithin. 

In emulsions containing 4% and 6 % SPI gel concentration, lecithin concentration 

0

1

L4 L6 L2S6 L4S2 L4S4 L4S6 L6S2 L6S4 L6S6

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a 

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35 

increased from 2% to 4%, while EEA and EEP values showed a statistically significant 

increase (p<.05). No significant increase was observed as the concentration of lecithin 

increased from 4% to 6%. Additionally, there is a obvious difference between the 

encapsulation efficiencies of phenolics and anthocyanins. EEA values are higher than 

EEP values. Similar observations were achieved in the case of HIPP-DEs. In the study 

of Zahed et al. (2023) similarly, EEA values were higher than EEP values in the system 

containing maltodextrin and soy protein isolate. Higher EEA values were associated 

with the difference in electrical charge between anthocyanins and phenolic compounds 

(Zahed et al., 2023).  

Figure 3.13: EEA and EEP values of HIPPEs. 

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L4 L6 L2S6 L4S2 L4S4 L4S6 L6S2 L6S4 L6S6

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Figure 3.14: EEA and EEP values of HIPPE-DEs. 

3.7 Effect of In Vitro Gastrointestinal Digestion on TAC, TPC, DPPH and ABTS 

of HIPPEs and HIPP-DEs 

The potential application of aqueous Hibiscus extract in functional foods holds 

significant promise due to biological activity of anthocyanins and phenolics. 

Enhancing the stability of anthocyanins and phenolics in the digestive system is crucial 

to enable efficient utilization of them. Therefore, in vitro simulated digestion was 

applied to assess the protective effect of HIPPEs and HIPP-DEs on bioactive 

compounds. As a control group, aqueous Hibiscus extract was used.  

3.7.1 Total anthocyanin content 

Table 3.3 illustrates the variations in TAC of aqueous Hibiscus extract (AHE) and 

various HIPPEs throughout the process of in vitro digestion. To compare the stabilities,  

in vitro digestion protocol was conducted.  For undigested products, all HIPPEs 

presented lower TAC values compared to the AHE. This decrease can be related with 

the entrapment of the anthocyanins in emulsion system and possible interactions 

between soy proteins and anthocyanins. The hydroxyl groups of anthocyanins, 

particularly in Hibiscus, enable them to engage with proteins and peptides in the 

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37 

emulsion (Chickhoune et al., 2017). In a study by Yuksel et al. (2010), it was observed 

that flavonoids, including anthocyanins, possess a strong affinity towards the 

hydrophobic regions located on the surface of proteins. These flavonoids have a 

propensity to bind with these hydrophobic regions on proteins, which can be attributed 

to the hydrophobic nature of flavonoids and the exposed hydrophobic areas on proteins 

(Yuksel et al., 2010). For nonencapsulated AHE, the anthocyanins remained stable and 

TAC remained nearly constant during gastric digestion. In HIPPEs containing 4% 

lecithin (L4, L4S2, L4S4, L4S6), the SPI gel concentration increased from 0% to 4%, 

while the amount of increase in TAC value in gastric digestion decreased in percent. 

However, when the SPI gel concentration increased from 4% to 6%, no statistically 

significant difference was observed in the TAC value during gastric digestion (p<.05). 

In HIPPEs containing 6% lecithin (L4, L4S2, L4S4, L4S6) also provided similar 

changes during gastric digestion. In the study of Betz & Kulozik. (2011), similar 

observation was achieved during encapsulation of bilberry extract within whey protein 

gel. The results showed that increasing the protein concentration from 15% to 20% 

decreased the release rate of anthocyanin during gastric digestion (Betz & Kulozik, 

2011). However, increasing the protein concentration from 20% to 25% (w/w) did not 

significantly affect the release rate. It was suggested that changing the protein 

concentration from a certain concentration do not have a substantial effect on the gel 

network's internal structure to form a complete diffusion barrier (Betz & Kulozik, 

2011). It can be observed that the non-encapsulated aqueous Hibiscus extract remained 

quite stable when exposed to the simulated gastric environment, with no significant 

changes observed in its TAC (p>0.05). Anthocyanins, when present in an acidic 

environment, exist in the flavylium cation form, which imparts a red color (Cooper-

Driver, 2001). These compounds are known to exhibit relatively good stability under 

such conditions (Jackman et al., 1987). Also, TAC values decreased significantly 

during intestinal digestion. According to Lang et al. (2020), anthocyanins are relatively 

stable during simulated gastric digestion. Therefore, the preservation and protection of 

anthocyanins should be focused on the intestinal digestion process. The alkaline 

environment of the intestine is the main factor causing anthocyanin degradation and 

limiting their bioaccessibility (Lang et al., 2020). 

 

 



38 

Table 3.3: Total anthocyanin content (mg C-3-GE/ 100 g) of HE and HIPPEs. 

Emulsion Undigested 

Gastric 

digestion 

Intestinal 

digestion 

HE 31.13±1.23aA 30.09±0.78aA 12.03±0.47cB 

L4 21.07±0.29bA 19.37±0,64dA 15.21±0.24aB 

L6 20.06±0.05bcA 19.47±2,0cdA 15.22±0.17abB 

L2S6 19.21±0.23cA 19.04±4,26dA 13.06±1.31bB 

L4S2 16.12±0.12dA 15.19±0.27abcA 12.96±0.05bB 

L4S4 15.43±0.42dA 15.24±0.04abA 12.72±0.27bcB 

L4S6 15.01±0.01dA 15.88±0.01abA 12.73±0.19bB 

L6S2 16.03±0.05dA 15.15±0.04bcA 12.94±0.04bB 

L6S4 15.43±0.21dA 15.65±0.04deA 12.75±0.04bA 

L6S6 15.13±0.15dA 15.92±0.05aA 13±0.28bcB 

*The results are represented as mean ± standard deviation (n=3). 
*Different lowercase letters indicate significant differences between different emulsions (p <0.05) 
*Different uppercase letters indicate significant differences between digestion phases. 

Table 3.4: Total anthocyanin content (mg C-3-GE/ 100 g) of HE and HIPP-DEs. 

Emulsion Undigested 

Gastric 

digestion 

Intestinal 

digestion 

HE 31.13±1.21aA 30.92±0.76aA 12.03±0.47aB 

L4 11.23±0.25bA 10.23±0.05dB 9.54±0.46bC 

L6 11.04±0.11bA 10.62±0.26cdB 9.55±0.04bC 

L2S6 10.32±0.28bA 9.91±0.03bcdB 9.13±0.23bC 

L4S2 9.88±0.09cA 9.06±0.05bcB 9.01±0.38cC 

L4S4 9.01±0.12cdA 9.17±0.02bcB 8.35±0.28cC 

L4S6 8.76±0.05cdeA 8.46±0.43bB 8.14±0.54cC 

L6S2 9.72±0.17cdeA 9.08±0.03bcB 8.94±0.37cC 

L6S4 9.04±0.04deA 9.17±0.02bB 8.41±0.12cC 

L6S6 8.51±0.49eA 8.49±0.11bB 8.12±0.25cC 

*The results are represented as mean ± standard deviation (n=3). 
*Different lowercase letters indicate significant differences between different emulsions (p <0.05) 
*Different uppercase letters indicate significant differences between digestion phases. 

3.7.2 Total phenolic content 

H. Sabdariffa polyphenols have potential use in functional foods due to their biological 

activity. However, in the case of bioaccessibility, improving their stability in the 



39 

gastrointestinal system is crucial. For this reason, in vitro gastrointestinal digestion 

was used to assess the protective effect of lecithin and SPI gel based HIPPEs and HIPP-

DEs on polyphenols. Aqueous H. Sabdariffa was used as a control group for 

comparison. Aqueous Hibiscus extracts had higher TPC values, concerning their 

emulsions, before in vitro gastrointestinal digestion. Sirovec et al. (2022) conducted a 

study where they compared the total phenolic content (TPC) values of rosemary and 

oregano extracts with protein-based emulsion systems. The findings of this study 

indicated that differences in TPC values between emulsions were appeared. These 

variations in TPC values of emulsified extracts could be attributed to the presence of 

certain compounds like ascorbic acid and proteins, which can react with the Folin-

Ciocalteu reagent and potentially interfere with the accurate determination of TPC 

using the Folin-Ciocalteu assay (Bastola et al., 2017). In the context of HIPPEs with 

the same lecithin concentration, it was observed that an increase in the concentration 

of SPI gel resulted in higher TPC values. This finding aligns with the results obtained 

in the study conducted by Sirovec et al. (2022), where a similar observation was made. 

Specifically, an increase in the concentration of commercial pea protein was found to 

be associated with an increase in the TPC value for emulsified rosemary extracts. From 

Table 3.5, significant decreases in TPC were observed in the Hibiscus extract after in 

vitro digestion. 407.59±6.48 and 927.6±19.3 of TPC of Hibiscus extract remained in 

the gastric and intestinal phase, respectively, representing approximately 15% and 

35% of the initial TPC. Similar significant losses in TPC during in vitro 

gastrointestinal digestion have been reported in other studies, such as elderberry 

(approximately 80% loss), grape pomace (over 50% reduction), Goji berries (only 30% 

TPC remaining in digests and berry pomace (approximately 72% reduction) (Jara-

Palacios et al., 2018; Pinto et al., 2017; Rocchetti et al., 2018; Xiong et al., 2020). 

Comparing the TPC of the digested phenolics of Hibiscus extract with that of t