Characterization of non-solvent precipitated starch using asymmetrical flow field-flow fractionation coupled with multiple detectors

Accepted Manuscript Title: Characterization of non-solvent precipitated starch using asymmetrical flow field-flow fractionation coupled with multiple detectors Authors: Catalina Fuentes, Hisfazilah Saari, Jaeyeong Choi, Seungho Lee, Malin Sj¨oo¨ , Marie Wahlgren, Lars Nilsson PII: DOI: Reference:

S0144-8617(18)31300-6 https://doi.org/10.1016/j.carbpol.2018.10.100 CARP 14232

To appear in: Received date: Revised date: Accepted date:

21-7-2018 11-10-2018 28-10-2018

Please cite this article as: Fuentes C, Saari H, Choi J, Lee S, Sj¨oo¨ M, Wahlgren M, Nilsson L, Characterization of non-solvent precipitated starch using asymmetrical flow field-flow fractionation coupled with multiple detectors, Carbohydrate Polymers (2018), https://doi.org/10.1016/j.carbpol.2018.10.100 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Characterization of non-solvent precipitated starch using asymmetrical flow field-flow fractionation coupled with multiple detectors Catalina Fuentesa,b*, Hisfazilah Saaria, Jaeyeong Choic, Seungho Leec, Malin Sjööa, Marie

a

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Wahlgrena and Lars Nilssona

Food Technology, Engineering and Nutrition, Faculty of Engineering LTH, Lund University, PO

Box 124, S-221 00 Lund, Sweden. b

School of Chemistry, Faculty of Pure and Natural Science, Universidad Mayor de San Andres

Department of Chemistry, Hannam University, Daejeon 34054, Republic of Korea

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c

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(UMSA), PO Box 330, Cota Cota 27 St., La Paz, Bolivia

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*Corresponding author:

Catalina Fuentes, Food Technology, Lund University, P.O. Box 124, SE-22100 Lund, Sweden,

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E-mail to co-authors:

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Fax: +46 46 2224622, cell phone: +46 722889206, E-mail: [email protected]

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E-mail: [email protected] (C. Fuentes) E-mail: [email protected] (H. Saari)

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E-mail: [email protected] (J. Choi)

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E-mail: [email protected] (S. Lee) E-mail: [email protected] (M. Sjöö) E-mail: [email protected] (M. Wahlgren) E-mail: [email protected] (L. Nilsson)

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Highlights Non-solvent precipitated starch (non-SPS) is obtained from waxy maize starch

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Precipitated starch properties after re-dispersion depend on the preparation method

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Precipitated starch has an amorphous structure

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Re-dispersion at room temperature can lead to dissolution of starch precipitates

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Size, molar mass, apparent density, and conformational properties are determined

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Abstract1

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Non-solvent precipitated starch (non-SPS) is a novel component for starch-based emulsions.

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Herein, three non-SPS materials were prepared using ethanol as a precipitant of waxy maize starch

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granules (WMs). The WMs were either untreated (SP) or pre-treated via acid-hydrolysis (AHSP).

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In addition, SP was modified using n-octenyl succinic anhydride (OSA), yielding OSASP. This

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study aimed to investigate the influence of the non-SPS preparation method on the size, molar mass (M), and apparent density (ρapp) of the materials when subjected to different

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dissolution/dispersion procedures using asymmetrical flow field-flow fractionation (AF4). The

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results showed that the molar mass, size, and apparent density depended on the type of non-SPS with a decrease in Mw (1.8 - 9.4 g/mol) and rrms (60 - 148 nm) upon re-dispersion in different

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media in the order: SP > OSASP > AHSP. Moreover, different types of non-SPS materials

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Non-solvent precipitated starch (non-SPS), waxy maize starch granule (WM), n-octenyl succinic anhydride (OSA), asymmetrical flow field-flow fractionation (AF4), differential refractive index (dRI), multiangle light scattering (MALS), dynamic light scattering (DLS), nanoparticle (NP), regenerated cellulose (RC), degree of OSA substitution (DS), bromophenol blue (BPB), bovine serum albumin (BSA), scanning electron microscope (SEM), transmission electron microscopy (TEM).

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displayed different conformational properties and were stable in aqueous solution at room temperature in the investigated time (24 h). Keywords: non-solvent precipitated starch, waxy maize, asymmetrical flow field-flow

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fractionation.

1. Introduction

Starch is the principal carbohydrate reserve in plants as well as a major source of energy in the human diet and animal feed, which makes it suitable for various industrial applications. In

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addition, it often contributes to characteristic properties of food and is added as a functional

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ingredient in many formulated products. In recent years there is a large interest in producing starch

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nanoparticles (NPs), in particular, could have several interesting uses such as emulsifiers for

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Pickering emulsions (Li, Li, Sun & Yang, 2013; Timgren, Rayner, Dejmek, Marku & Sjöö, 2013).

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Reportedly, these NPs can be generated via the non-solvent precipitation of dissolved starch (El-

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Sheikh, 2017; Qin, Liu, Jiang, Xiong & Sun, 2016). Previously, the stabilization of emulsions using non-solvent precipitated waxy maize starch granule (WM) , either unmodified or modified

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with n-octenyl succinic anhydride (OSA), was investigated and compared to OSA-modified starch

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granules (Saari, Fuentes, Sjöö, Rayner & Wahlgren, 2017). Generally, there has been a significant interest in producing nanostructures from starch granules,

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since starch is a green and renewable resource. Indeed, various methods to obtain starch nanostructures have been reported (Ali Razavi & Amini, 2016; Le Corre, Bras & Dufresne, 2010; Wang, Truong & Wang, 2003). However, the techniques employed are said to affect the structure of the material, for instance, whether it is crystalline or amorphous (Ali Razavi & Amini, 2016; Kim, Park & Lim, 2015). 3

Starch nanostructures can principally be prepared in three different ways: i) acidic or enzymatic hydrolysis, which yields nanocrystals at temperatures lower than the gelatinization temperature; ii) precipitation from starch solution, which yields either amorphous or crystalline particles; iii)

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mechanical treatment, such as high pressure homogenization, leading to so called nano-colloids (Ali Razavi & Amini, 2016; Le Corre, Bras & Dufresne, 2010).

More specifically, the hydrolysis is usually carried out in a two-stage pattern (Genkina, Kiseleva & Noda, 2009; Gérard, Colonna, Buléon & Planchot, 2002; Utrilla-Coello et al., 2014). In the

relatively faster, first stage, the amorphous part of the starch granule is affected, resulting in an

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increase of the relative crystallinity. In the slower, second stage, both the amorphous and

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crystalline parts are affected

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In the precipitation procedure, starch granules are first dissolved into polymers, after which a nonsolvent (i.e., ethanol) is added, resulting in the precipitation of smaller particles (non-solvent

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precipitated starch, non-SPS). In fact, methods for the preparation of non-SPS, commonly

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described as nanoprecipitation, have been expensively investigated (Chang, Yan, Wang, Ren, Tong & Zhou, 2017; Chin, Pang & Tay, 2011; Qin, Liu, Jiang, Xiong & Sun, 2016; Qiu, Yang,

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Ge, Chang, Xiong & Sun, 2016; Sadeghi, Daniella, Uzun & Kokini, 2017; Tan, Xu, Li, Liu, Song

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& Wang, 2009; Tan, Xu, Liu, Li, Lu & Wang, 2012; Uzun & Kokini, 2014; Wu, Chang, Fu, Ren, Tong & Zhou, 2016).

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Asymmetrical flow field-flow fractionation (AF4) is a versatile separation method that has been proved suitable for the analysis of NPs and large macromolecules such as starch (Choi, Kwen, Kim, Choi & Lee, 2014; Nilsson, 2013; Wahlund, Leeman & Santacruz, 2011). AF4 is an especially powerful technique when connected to detectors that measure for example differential

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refractive index (dRI) and multiangle light scattering (MALS). For instance, AF4-MALS-dRI has been used in the analysis of starch NPs during their production process (Juna, Damm, Kappe & Huber, 2012; Juna & Huber, 2013; Juna, Huber, Hayden, Damm & Kappe, 2014). However,

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currently, little is known about the behavior of starch NPs and non-SPS as a finished product when they are re-dispersed in an aqueous environment.

The aim of the present study was to investigate the influence of preparation methods for non-SPS, e.g., acid hydrolysis and OSA-modification, on the size, molar mass (M), and apparent density (ρapp) when non-SPS is subjected to different dissolution/dispersion procedures. The method

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employed herein was a combination of acid hydrolysis, employed for the pre-treatment of the

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sample, and a precipitation with a non-solvent such as ethanol (Saari, Fuentes, Sjöö, Rayner &

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Wahlgren, 2017). The stability of the re-dispersed non-SPS was monitored by examining the

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changes in size over a time span of 24 h using AF4-MALS-dRI.

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2. Hypothesis

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The preparation method affects the size, molar mass, and apparent density of non-SPS when

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subjected to different dissolution/dispersion procedures. In addition, the dissolution/dispersion

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procedure affects the stability of non-SPS. 3. Material and methods

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3.1. Materials

WM starch was supplied by Lyckeby Culinar AB (Malmö, Sweden); OSA was obtained from Trigon Chemie (Hesse, Germany); dimethyl sulfoxide (DMSO ≥ 99.5%) and HCl were provided by VWR Chemicals (Leuven, Belgium); ethanol (99%) and citric acid were obtained from SigmaAldrich (St Louis, MO, USA); NaOH was purchased from Merck (Frankfurt, Germany); bovine 5

serum albumin (BSA) was purchased from Sigma Aldrich (St. Louis, MO, USA); NaNO3 (A3125) was provided by AppliChem (Darmstadt, Germany and NaN3 (10369) was supplied by BDH (Poole, UK).

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3.2. Preparation of precipitated starch NPs Three different types of non-SPS NPs were prepared from WM, as shown in Table 1. The starting WM starch sample was used for comparative purposes in the analyses. Table 1. Overview of the starch samples prepared in this study

Sample ID AHSP

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Non-SPS of acid-hydrolyzed WM

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Samples

SP

Non-SPS of WM

OSASP

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OSA-modified non-SPS of WM

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The SP sample (non-SPS of WM) was prepared according to a previously reported approach

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(Saari, Fuentes, Sjöö, Rayner & Wahlgren, 2017), which can be briefly summarized as follows. WM starch was dispersed in Milli-Q water at a concentration of 8 mg/mL, and autoclaved at 140

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°C for 20 min in the presence of N2 using a temperature-controlled laboratory autoclave (Roth Model II, 200 mL, Karlsruhe, Germany). In order to precipitate the starch, ethanol was added in a

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volume ratio of 1:1, and the solution was mixed. The precipitated particles were collected by

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centrifugation at 2000 rpm for 10 min, and all batches were pooled. The collected particles were left in the fumehood overnight for complete ethanol removal. The precipitated starch samples were then frozen at -20 °C and freeze-dried in a Hetosicc freeze-dryer (Heto, Birkerød, Denmark) over a period of 4 - 5 d.

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The AHSP sample (non-SPS of acid-hydrolyzed WM) was prepared in a similar way, following an acid pre-treatment. The acid-pretreatment was performed according to a previously reported procedure (Wang, Truong & Wang, 2003) with slight modifications. First, 50 g of WM starch was

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dispersed in 250 mL HCl (1 M) with continuous stirring (200 rpm) at 40 °C for 120 min; the dispersion was adjusted to pH 5.5 using NaOH (1 M). The suspension was then washed three times using Milli-Q water and subsequently centrifuged at 3500 rpm for 10 min. The precipitated solids were air-dried at room temperature for 3 d. Finally, the AHSP sample was obtained by non-solvent precipitation of the acid-hydrolyzed WM, as described above.

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In order to prepare the OSASP sample (OSA-modified non-SPS of WM), a portion of wet SP was

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chemically treated with OSA. The OSA-modification was performed using 3% (w/w) OSA, based

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on the dry weight of SP. The procedure used for the OSA-modification has been previously

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reported (Saari, Heravifar, Rayner, Wahlgren & Sjöö, 2015) and can be briefly summarized as

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follows. Starch was dispersed with continuous stirring in 1.5 times (w/w) more pure water, based

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on the weight of dry starch and the pH of the solution was adjusted to 7.4 - 7.8 using HCl (1 M) or NaOH (1 M). OSA was added in four portions in 15-min intervals, during which the pH was

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maintained constant at 7.6 using automatic titration of NaOH (1 M). The OSA-modification was

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considered to be completed when the pH was stable for at least 15 min. After that, the suspension was centrifuged at 3500 rpm for 10 min, the supernatant was discarded, and the precipitate was

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washed with pure water, citric acid (pH 4.5 - 5.0), and then once again with pure water. The starch sample was freeze-dried as described above. Determination of the degree of OSA substitution (DS) was carried out in duplicate. First, 2.5 g of OSASP and a control sample (SP) were weighed and wetted with a few drops of ethanol, and 25 mL of HCl (1 M) was added. The suspension was then stirred for 30 min and centrifuged at 3500 7

rpm for 10 min. The supernatant was discarded and the residue was washed once with ethanol (25 mL) and twice with pure water (2 × 25 mL). After that, the residue was dissolved in 150 mL pure water, heated in a bath at 90 °C for 10 min, and then rapidly cooled to 25 °C in an ice bath. The

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suspension was titrated while stirring with NaOH (0.1 M) until the pH became 8.2. The volume of NaOH employed in the titration was then used to calculate the mass-based DS, according to the following equation: 𝐷𝑆 (%) =

(𝑉𝑠𝑎𝑚𝑝𝑙𝑒 − 𝑉𝑐𝑜𝑛𝑡𝑟𝑜𝑙)·𝑀·210 𝑚

· 100

(1)

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where V is the volume of NaOH used in the titration, M is the molar concentration of NaOH, 210

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is the molar mass (g/mol) of OSA, and m is the mass of the dry sample. The DS of the OSASP

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sample prepared in this study was determined to be 0.02.

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3.3. Chemical analyses and thermal properties of the starch samples

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The moisture contents of the samples were determined using a moisture analyzer (MAC 110/WH,

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Radwag, Radom, Poland), while the total starch content was determined enzymatically using total starch analysis (Megazyme amyloglucosidase/α-amylase, Megazyme International, Bray, Ireland)

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by subtracting the free glucose value.

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The total protein content was determined by measuring the nitrogen content using an elemental analyzer Flash EA 1112 N (Thermo Fisher Scientific, Waltham, MA, USA). The sample (25–32

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mg) was combusted in a sealed furnace and the nitrogen content was determined through thermal conductivity detection using aspartic acid as a standard. A nitrogen to protein factor of 6.25 was used in the calculations. Each analysis was performed in triplicate.

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The thermal properties were determined using differential scanning calorimetry (DSC 6200, Seiko Instruments Inc., Tokyo, Japan). The sample (20 mg) was weighed into a small tube and Milli-Q water (60 µL) was added (1:3 w/v ratio). The solution was then left for 1 h to equilibrate, after

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which approximately 8 mg of the mixture was weighed into an aluminum pan with lid (TA Instruments, Newcastle, Delaware, USA) using a microbalance (CAHN Instruments, C-30, Samo tronic Microbalance, Paramount, California, USA). The pan was hermetically sealed and analyses were performed over a temperature range of 10 - 160 °C with a scanning rate of 10 °C/min using an empty sealed pan as a reference. Thermo-gravimetric curves were recorded on an Exstar 6000

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Thermal Analyses System equipped with a Muse Standard Analysis software (version 6.4, SII

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3.4. Scanning electron microscope (SEM)

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Nano Technology Inc., Chiba, Japan).

The morphology of AHNP sample was investigated using scanning electron microscopy

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(SEM).The sample was coated with Au/Pb (~10 nm) using a BALTZERS SCD 004 sputter coater

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and inspected using a JSM-6700F scanning electron microscope, (JEOL inc, Japan) operated at 10

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kV using the lower SE detector (LEI).

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3.5. Cryogenic transmission electron microscopy (Cryo-TEM) Micrographs were taken of the AHSP sample at a concentration of 10 mg/mL. The sample was

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dissolved in Milli-Q water and vitrified using an automatic plunge freezer (Leica EM GP). 4 µl drop was placed on a lacy carbon-coated copper grid (Ted Pella) and subsequently plunged into liquid ethane at -183 °C to allow vitrification. The prepared grids were stored in in liquid nitrogen until use. A cryo transfer tomography holder (Fischione Model 2550) was used to transfer the specimen into the electron microscope (JEM 2200FS), equipped with an in-column energy filter 9

(Omega filter). The acceleration voltage was 200kV and zero-loss images were digitally recorded with a camera (TVIPS F416) using serial EM under low dose conditions with a 13 eV energy selecting slit in place.

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3.6. Sample preparation for AF4 analysis In order to perform AF4, the starch samples were prepared in three different ways: i) dissolution at 100 °C in DMSO, ii) dissolution at 100 °C in an aqueous solution (e.g., AF4 carrier liquid), and iii) dispersion at room temperature in an aqueous solution (e.g., AF4 carrier liquid).

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The samples in DMSO were prepared as follows: 100 mg of the respective sample was weighed

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into a glass vial (26 × 76 mm, 25 mL capacity) and 3 mL DMSO was added. The sample vial was

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capped and then heated at 100 °C for 1 h while it was being stirred. Finally, the sample was diluted

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with the AF4 carrier liquid at 100 °C to reach a final concentration of 0.25 mg/mL.

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The aqueous solution samples were prepared at a concentration of 1 mg/mL, heated to 100 °C

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(boiling) for 1 h with continuous stirring, and diluted with the AF4 carrier liquid to a final

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concentration of 0.25 mg/mL.

The samples at room temperature were obtained by dispersing the sample in aqueous solution at a

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concentration of 1 mg/mL for 1 h with continuous stirring and then diluting the mixture with the AF4 carrier liquid to a final concentration of 0.25 mg/mL. The AF4 carrier liquid was composed of

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10 mM NaNO3 and 0.02% NaN3 dissolved in Milli-Q water. 3.7. AF4-MALS-dRI with online Dynamic light scattering (DLS)

The samples were separated with an AF4 system (Eclipse 3+, Wyatt Technology, Dernbach, Germany) and characterized using online MALS (Dawn Heleos II, Wyatt Technology) and dRI 10

detector (Wyatt Technology). Both apparatuses operated at a wavelength of 658 nm. An Agilent 1100 series isocratic pump (Agilent Technologies, Waldbronn, Germany) with an in-line vacuum degasser and an Agilent 1100 series autosampler delivered the carrier flow and handled the sample

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injection into the AF4 channel. Between the pump and channel, a polyvinylidene fluoride membrane with a 100 nm pore size (Millipore Corp., Bedford, MA, USA) was placed to ensure that the particle-free carrier liquid entered the channel.

The AF4 channel (Wyatt Technology, Dernbach, Germany) was short with a trapezoidal geometry (tip-to-tip length of 17.4 cm and inlet and outlet widths of 2.17 and 0.37 cm, respectively) and a

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nominal thickness of 350 µm. The ultrafiltration membrane forming the accumulation wall was a

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regenerated cellulose (RC) with a nominal cut-off of 10 kDa (Merck Millipore, Bedford, MA,

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USA). Experimental determination of the channel thickness and validation of the performance of

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the AF4 system were performed with bromophenol blue (BPB) and bovine serum albumin (BSA)

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(Sigma Aldrich, St. Louis, MO, USA) solutions (1 mg/mL, w/v), respectively, according to a

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previously reported procedure (Nilsson, 2013), using an add MATLAB-based software (FFFHydRad 2.2) (Håkansson, Magnusson, Bergenståhl & Nilsson, 2012; Magnusson, Håkansson,

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Janiak, Bergenståhl & Nilsson, 2012). The actual channel thickness was experimentally

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determined to be 266 µm.

In the AF4 analysis of the samples, the detector flow rate was kept constant at 1 mL/min, while the

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sample injection into the channel was performed at a flow rate of 0.2 mL/min for a total of 6 min. The sample injection volume was in the range of 5 - 80 µl for injected sample mass of approximately 1.25 - 20 µg. In order to ensure that no overloading occurred, the injection mass was optimized so that the retention time was independent of the injection mass. After injection, a 4-min focusing/relaxation was performed, with the focusing flow rate being identical to the initial 11

cross flow rate before the sample elution. The applied crossflow was set to decay exponentially with time in order to avoid excessive retention and to obtain a more uniform selectivity over the entire size distribution (Leeman, Wahlund & Wittgren, 2006). The crossflow decay was

𝑄𝑐 (𝑡) = 𝑄𝑐 (0)𝑒

(−

𝑙𝑛2 𝑡) 𝑡1/2

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programmed according to the following equation:

(2)

where Qc (t) is the crossflow rate as a function of time t after elution starts, Qc (0) is the initial

cross flow rate, and t1/2 is the half-life of the decay. For all samples, the elution started with an

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initial cross flow rate of 2.5 mL/min, which decreased exponentially over time to 0.13 mL/min,

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was flushed without any cross flow for 11 min.

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with t1/2 = 4 min. This rate was then maintained for 22 min. Before the next injection, the channel

Data were recorded using the Astra software (v. 6.1.5.22, Wyatt Technology). The values of M

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and the z-average root-mean-square radius (rrms) were obtained from the 1st order fitting of the

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MALS data at a scattering angle in the range of 60.0°–152.5° using the Berry method (Andersson, Wittgren & Wahlund, 2003; Berry, 1966). In this case, the second virial coefficient (A2 ) was

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neglected and a specific refractive index (dn/dc) of 0.146 mL/g was used for amylopectin in water

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(Brandrup, Immergut & Grulke, 1999). The apparent density (ρapp) was calculated from M and rrms (Nilsson, Leeman, Wahlund & Bergenståhl, 2006), whereas the hydrodynamic radius (rh) was

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calculated using the MATLAB add-on FFFHydRad 2.2. For online DLS measurements, an Eclipse 2 system (Wyatt Technology) was coupled online with a DAWN EOS MALS detector (Wyatt Technology) operating at a wavelength of 690 nm. A MALS detector with a scattering angle of 110.7° was connected to the DLS instrument (DynaPro

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NanoStar, Wyatt Technology) via a glass fiber. In this setup, the channel thickness was determined to be 258 µm. 4. Results and discussion

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4.1. Chemical analysis and thermal transition properties of the non-SPS samples The moisture, total starch, and total protein contents of the three non-SPS samples and WM starch are shown in Table 2. The moisture content was similar in all samples (11 - 13%), while the total starch content varied in the range of 90 - 95%, with the SP sample showing the highest value of about 95%. Notably, the starch content of the reference (WM) was lower than those of the AHSP

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and SP samples. One possible explanation could be that the precipitation procedure removed some

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impurities, resulting in a lower starch content in the WM sample. However, the starch content of

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the OSASP sample was slightly lower than that of the WM sample, which may be due to a

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possible interference of the OSA substituents with the enzymatic degradation to glucose.

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Furthermore, the protein contents of the non-SPS samples were similar to each other, in the range

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of 0.12% - 0.15%. In contrast, the protein content of the WM sample was lower than those of the non-SPS samples, probably because of the removal of impurities during the precipitation

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procedure, as suggested above.

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Table 2. Moisture, total starch, and total protein contents of the starch samples

Sample

Moisture (%) a

Total starch (%) a,b

Total protein (%) a,b

AHSP

13.1 ± 0.7

93.7 ± 0.7

0.13 ± 0.00

SP

12.9 ± 0.9

95.1 ± 0.6

0.12 ± 0.00

OSASP

12.1 ± 1.1

89.9 ± 0.7

0.15 ± 0.01

WM

11.1 ± 1.6

90.9 ± 1.4

0.07 ± 0.03

a

The values include a standard error of the mean (±) based on three replicates.

b

Values reported on dry basis.

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In addition, DSC analysis for WM gave the onset temperature (T0) = 64.1 ± 1 °C, peak temperature (Tp) = 70.0 ± 0.5 °C, conclusion temperature (Tc) = 75.6 ± 1.4 °C, and enthalpy (∆H)

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14.9 ± 2.0 mJ/mg. Since the non-SPS samples did not show any transition in the endothermic curve (no peak was detected), the crystallinity was not measurable. This indicates that the granule disruption was complete and the non-solvent precipitation did not induce any major formation of starch crystals (Das, Sanson, Fava & Kumacheva, 2007; Kim, Lee, Kim, Lim & Lim, 2012). Therefore, all three non-SPS samples (AHSP, SP, and OSASP) had amorphous structures.

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4.2. Effect of the dissolution/dispersion procedure on the molar mass and size of the non-

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SPS samples

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Table 3 shows the weight-average molar mass (Mw), rrms, and sample recoveries obtained from the AF4-MALS-dRI analysis of starch samples dissolved in DMSO. Considering the fact that the

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recovery values were ≥ 95% for all non-SPS samples, we concluded that there was no significant

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sample losses during the AF4 analysis. In contrast, in the case of WM, the recovery was much

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lower (67%), which could be related to the uncertainty in drawing the baseline of the rather weak dRI-signal of WM. In fact, in order to avoid overloading, the injection mass of WM was kept very

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low (1.9 µg) compared to those of the non-SPS samples (about 10 µg), thus making the determination of the mass recovery rather uncertain (Perez-Rea, Bergenståhl & Nilsson, 2015).

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Another reason for the low mass recovery of WM may be the immobilization of the sample at the accumulation wall and the subsequent elution as a release peak when the cross flow is turned off. For the WM sample, the release peak area was about 19% of the injected sample.

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The mean sizes of the non-SPS samples were smaller than that of WM (rrms = 204 nm), increasing in the order of AHSP < OSASP < SP. The Mw values of the non-SPS samples were also lower than that of WM (2.4·108 g/mol), maintaining the same trend as for the mean sizes.

Sample

Mw·108 (g/mol)a

rrms (nm)a

AHSP

0.18

60

SP

0.94

148

OSASP

0.44

98

WM

2.4

204

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Table 3. Results from the AF4-MALS-dRI analysis of starch samples dissolved in DMSO Recovery (%)b 97 95 96 67

Mw is the weight-average molar mass; rrms is the z-average root-mean-square radius.

b

The mass recovery was determined in percentage from the ratio of the mass eluted from

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a

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the separation channel (integration of the dRI signal) to the injected mass (based on the

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analyzed starch content of the sample).

The AF4 fractograms of the three non-SPS samples, i.e., AHSP, SP, and OSASP are presented in

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Fig. 1A, B, and C, respectively. The samples were prepared according to three different

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dissolution/dispersion procedures, as described in section 3.4.

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In a previous study (Perez-Rea, Bergenståhl & Nilsson, 2015), in which starch granule samples were dissolved in DMSO, near complete dissolution (95 ± 3%) was observed. Therefore, it is

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expected that the differences in the AF4 results of sample dissolved in DMSO compared to other dissolution/dispersion procedure would indicate an incomplete dissolution of the sample due to the

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presence of particles or supramolecular aggregates Notably, no significant differences were observed in the AF4 fractograms as well as the molar mass and size distributions between the same sample dissolved in DMSO (blue) and boiling aqueous solution (green), respectively (Fig. 1A - C). This indicates that all three non-SPS samples 15

were well dissolved in either DMSO or boiling aqueous solution. In contrast, considering the samples prepared in aqueous solutions at room temperature (red solid lines in Fig. 1A - C), each non-SPS sample showed a different behavior. More specifically, the AF4-MALS fractogram of the

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AHSP sample prepared in an aqueous solution at room temperature (red solid line, Fig. 1A) indicated the existence of two populations, unlike for the same sample prepared in DMSO (blue solid line) or boiling aqueous solution (green solid line). The elution time (approx. 22 - 34 min) and elution profile of the first population were similar to those of the same sample prepared

differently and with slightly higher M and rrms distributions. In addition, the second population

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(elution time approx. 37 - 45 min), which did not exist in the sample prepared differently, showed

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higher M (about 109 g/mol) and rrms (about 200 nm). In fact, the existence of the second population

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suggests the presence of some undissolved material in AHSP sample in an aqueous solution at

M

room temperature.

D

The AF4-MALS fractogram of the SP sample prepared in aqueous solution at room temperature

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(red solid line, Fig 1B) showed a much different elution profile than the one prepared in DMSO, thus indicating that dissolution was not complete. In particular, the elution profile was narrower

EP

than that of the DMSO. SP sample (blue solid line), mainly because much of the later-eluting material was missing. Due to the same reason, the SP sample in an aqueous solution at room

CC

temperature exhibited lower M and rrms distributions than the same sample prepared differently.

A

By contrast, no significant differences were observed among the three AF4-MALS fractograms of the OSASP samples (Fig. 1C). Moreover, the M and rrms distributions were similar, indicating that OSASP was dissolved well in all three preparation procedures.

16

(A) 20

30

35

40

45 1010

300

250 109

108

150

0,4

SC RI PT

200 rrms (nm)

0,8

100

Molar Mass (g/mol)

Differential refractive index

1,2

25

107

50

0

0,0

20

25

30

35

40

0,0

D

108

TE

150

100

0

(B) 45 109

A

200

50

40

M

250

EP

0,4

35

300

rrms (nm)

0,8

30

25

30

35

40

106 45

Time (min)

A

CC

20

107

Molar mass (g/mol)

Differential refractive index

1,2

25

N

20

U

Time (min)

106 45

17

(C) 20

30

35

40

45

300

109

250 108

150

0,4

107

100

50

0,0

SC RI PT

200 rrms (nm)

0,8

Molar mass (g/mol)

Differential refractive index

1,2

25

106

0 20

25

30

35

40

45

N

U

Time (min)

Fig. 1. AF4 fractograms of the non-SPS samples (AHSP (A), SP (B), and OSASP (C)) dissolved in DMSO (blue), boiling aqueous

A

solution (green), and aqueous solution at room temperature (red). Symbols denote the molar mass (g/mol) (○), root-mean-square

M

radius rrms (nm) (▲), MALS-signal at 90° scattering angle (∙∙∙∙), and dRI signal (▬).

4.3. Conformation study using the hydrodynamic radii obtained from the AF4 theory and

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D

online DLS

In order to investigate the conformational properties of the non-SPS samples, they were dispersed

EP

in aqueous solutions at room temperature, as they could be useful in the preparation of Pickering emulsions (Saari, Fuentes, Sjöö, Rayner & Wahlgren, 2017). Herein, the conformational parameter

CC

was obtained from the ratio rrms/rh, where rh is the hydrodynamic radius that can be obtained in two ways: AF4 online with DLS or calculations from the AF4 theory. The results obtained from AF4

A

connected to online DLS are shown in the supplementary material (Fig. S1-S3). The results had severe limitations and they showed considerable noise, if the noise is disregarded, the rh appeared to be underestimated compared to AF4 theory (Fig. S4). To a large degree, the rh appeared to be independent on retention time although rrms and M were increasing. This suggests that there is an

18

upper limit of rh that could be determined (although it should be noted that separation in AF4 is not based on rrms or M). The diffusion rate is relatively low for large analytes and the residence time in the detector cell may be insufficient for an adequate data collection, resulting in an

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underestimation of rh as well as a maximum value. The results are in good agreement with previous results found in literature. Reportedly, the upper limit for an adequate determination of rh for starch polymers, using online DLS, is approx. 50 nm. While determination of higher rh required stop-flow (Rolland-Sabaté, Guilois, Jaillais & Colonna, 2011). It has also been shown for other large macromolecules that commonly applied flow rates in AF4-DLS results in underestimated rh

U

while calculation of rh from retention times provided reliable data (Boye, Ennen, Scharfenberg,

N

Appelhans, Nilsson & Lederer, 2015). The limitation was reported to be overcome by a decrease in

A

the flow to the detector, which in turn, led to lower resolution. Furthermore, the concentration in

M

the detector cell is known to strongly influence the determination of rh and the analyte concentration at the detector is required to be > 0.1 mg/mL (Runyon & Williams, 2014). In the

D

current investigation, the concentration at the dRI peak maxima (Fig. 1A - C) was between 0.3 - 1

TE

µg/mL, which is, at least a factor 100 lower than recommended. To increase injected amounts in

EP

order to overcome this limitation would not be possible, as this will rapidly lead to overloading of the separation channel (Perez-Rea, Bergenståhl & Nilsson, 2015). Similarly, it would not be

CC

feasible to decrease the detector flow, to allow for sufficient residence time of the analytes, as this would widen and lower the peaks and lead to even lower concentrations in the DLS detector.

A

Hence, AF4 with online DLS does not seem to be a viable characterization method for the starches investigated. The summary of the conformation results based on rh from AF4 theory is shown in Table 4. For SP and OSASP samples rrms/rh = 1.22 and 1.04 respectively. These values correspond to what is 19

expected from a highly branched macromolecule (Burchard, 1999) and are in good agreement for what has been reported for amylopectin (rrms/rh = 1.02 to 1.29) (Roger, Bello-Perez & Colonna, 1999). For AHSP (first eluting population) rrms/rh = 0.86. It suggest that this sample has a

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conformation that does not agree with what is expected for amylopectin in solution and is clearly different from SP and OSASP samples. Taking into consideration that the pre-treatment with acid hydrolysis could have an impact on the starch structure, thereby making the chain length

distribution narrower compared to native WM, which, in turn, could favor the formation of

spherical objects (Kim, Lee, Kim, Lim & Lim, 2012; Miao, Jiang, Zhang, Jin & Mu, 2011; Saeng-

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on & Aht-Ong, 2017). To further investigate these results, SEM and cryo-TEM were utilized to

N

characterize both the dry AHSP sample and AHSP in aqueous dispersion. In the SEM micrographs

A

(Fig 2. A), the presence of small paricles (d approx. 250 nm) can be clearly seen. However, upon

M

dispersion in an aqueous environment, the cryo-TEM micrographs (Fig 2. B.) does not show such well defined particles. Rather, the sample appears to consist of smaller and less well-defined

D

objects. The conformation from rrms/rh is suggested to be something in between a highly branched

TE

object (rrms/rh ≥ 1.0) and a spherical object with homogeneous mass distribution (rrms/rh = 0.778)

EP

(Burchard, 1999). The result is in reasonable agreement with what is displayed in the cryo-TEM micrograph (Fig. 2. B.). Furthermore, it can be concluded that upon dispersion the AHSP sample

CC

loses its well defined particle-like nature and more resembles an aggregate of polymers. Most likely, this is due to partial dissolution of the particles. It should also be noted that the AHSP

A

sample contains a second, much larger population (rrms approx. 200 nm) present at very low concentration (Fig. 1. A.). These components are larger than the particles observed in SEM and were not visible in the cryo-TEM results (probably due to their very low concentration). Possibly,

20

this second population might consist of either larger particles or aggregates of smaller particles, but not conformational data could be obtained for this population. Table 4. Values of rrms, rh, and conformation parameter obtained from AF4 theory rrms (nm)a

rh (nm)b

AHSP

70

81

SP

116

95

OSASP

103

99

rrms/rh

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Sample

0.86 1.22 1.04

rrms is the z-average root-mean-square radius

b

rh is the z- weighted average hydrodynamic

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TE

D

M

A

N

U

a

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Fig. 2. Micrographs from (A) Scanning electron microscopy (SEM) and (B) Cryogenictransmission electron microscopy (TEM) of AHSP sample showing the morphology in the dry state and in aqueous dispersion respectively.

A

4.4. Stability of SP and AHSP in an aqueous dispersion during a 24-h dissolution/dispersion process

As discussed above, the AHSP and SP samples dispersed in an aqueous environment at room temperature appeared not to dissolve upon re-dispersion. In order to gain more insight into this phenomenon, the stabilities of the dispersed samples were investigated during a time span of 24 h. 21

AF4-MALS-dRI was used to determine the Mw, rrms, and ρapp values, as well as the sample recovery, and the obtained results were summarized in Table 5. As a result, we found that the samples contained rather high M species, with Mw values ranging from 1.7·107 to 6.7·107 g/mol,

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rrms in the range of 68 - 116 nm, and sample recoveries higher than 85%. The results of the AHSP sample showed very small fluctuations over the full time span. As can be seen from Fig. 2, a slight growth occurred in the later eluting second population until the twelfth hour, after which no further growth was observed. Notably, this population constituted a very small fraction of the entire sample (< 2 mass %).

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Considering the SP sample, although different fractograms were obtained when prepared in an

N

aqueous solution at room temperature and through the other preparation methods, the results

A

seemed stable over the full time span, and the measured Mw and rrms values were in agreement with

M

the previously reported ones (Juna, Williams & Davies, 2011; Naguleswaran, Vasanthan, Hoover, Chen & Bressler, 2014; Qin, Liu, Jiang, Xiong & Sun, 2016).

rrms

ρapp

Recovery

SP sample Mw·107

rrms

ρapp

Recovery

(nm)a

(kg/m3)b

(%)c

(g/mol)a

(nm)a

(kg/m3)b

(%)b

70

16.2

92

6.7

116

8.3

85

69

16.3

91

6.6

114

8.4

86

1.7

68

13.7

91

6.2

110

8.9

86

1.9

69

14.9

88

5.7

111

7.9

86

2.3

70

17.6

87

6.4

111

9.1

86

2.2

6

2.1

12 18

A

24

CC

1

EP

AHSP sample Mw·107 Time (h) (g/mol)a

TE

D

Table 5. Average parameter values obtained from AF4-MALS-dRI analyses of AHSP and SP in aqueous solutions at room temperature over a time span of 24 h

a

Mw is the weight-average molar mass, rrms is the z-average root-mean-square radius

b

ρapp was calculated using z-average rrms and Mw.

c

The sample recovery was determined from the ratio of the mass eluted from the AF4 channel (integration of the dRI

signal) to the injected mass (based on the analyzed starch content of the sample).

22

20

25

30

35

40

45

1,0

300

24thhour

200 12thhour

0,5

6thhour 1sthour

100

0,0 15

20

25

30

35

40

0 45

N

U

Time (min)

rrms (nm)

MALS-signal 90°

18thhour

SC RI PT

15

Fig. 3. AF4-MALS fractogram of AHSP in an aqueous solution at room temperature over a time span of 24 h. The Rayleigh ratio

A

obtained from the MALS-signal at a scattering angle of 90° is indicated by the solid lines and the root-mean-square radius (rrms)

M

distribution vs the elution time (min) is depicted using open circles.

D

5. Conclusion

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In this study, three different types of non-solvent precipitated starch (non-SPS) samples were prepared from waxy maize starch granules, and their properties were examined. As a result, we

EP

found that they differed in rrms and M upon re-dispersion in different media, with a decrease in Mw

CC

in the order SP > OSASP > AHSP. These changes depended on the type of treatment used (with or without an acid hydrolysis) and the post precipitation OSA modification. Notably, all three non-

A

SPS samples showed an amorphous structure and displayed different conformational properties when dispersed in aqueous solutions at room temperature. While OSASP was successfully dissolved when dispersed in an aqueous solution at room temperature, the AHSP and SP aqueous solutions at room temperature had different size distributions than the DMSO or boiling aqueous solutions of these materials. However, AHSP and SP were stable after dispersion in aqueous 23

solution at room temperature in the investigated time span (24 h). These results indicate that the preparation procedure had a strong influence on the size and conformation of the resulting nonSPS. Furthermore, depending on the preparation procedure, these materials were to a large extent

SC RI PT

dissolved when re-dispersed in an aqueous environment. This was especially true for non-SPS materials, which have undergone a more extensive chemical treatment, i.e., acid hydrolysis or

chemical modification by OSA. Nevertheless, in order to obtain non-SPS with a more pronounced

U

nanoparticle character, additional treatments are needed.

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Acknowledgements

A

The present study was supported by the Swedish International Development Agency

M

(SIDA/SAREC) in a collaborative project between the Higher University of San Andres UMSA (Bolivia) and Lund University (Sweden); the National Research Foundation (NRF) of Korea

D

[NRF-2013K2A3A1000086, NRF-2016R1A2B012105]; and the Swedish Foundation for

TE

International Cooperation in Research and Higher Education (STINT).

EP

Authors would like to thank Dorin Vataire and Dr. Claudia Zielke (Lund University) for their help

CC

during the preparation of the OSASP sample and Anna Carnerup (Physical Chemistry – Lund

A

University) for their help during the SEM and Cryo-TEM analysis

24

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