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Homogenization-induced degradation of hydrophobically modified starch determined by asymmetrical flow field-flow fractionation and multi-angle light scattering
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FOOD
HYDROCOLLOIDS Food Hydrocolloids 20 (2006) 1087–1095 www.elsevier.com/locate/foodhyd
Homogenization-induced degradation of hydrophobically modified starch determined by asymmetrical flow field-flow fractionation and multi-angle light scattering Gustaf Modigb, Lars Nilssona,, Bjo¨rn Bergensta˚hla, Karl-Gustav Wahlundb a
Faculty of Engineering LTH, Division of Food Technology, Centre for Chemistry and Chemical Engineering, Lund University, Box 124, S-221 00 Lund, Sweden b Faculty of Engineering LTH, Division of Technical Analytical Chemistry, Centre for Chemistry and Chemical Engineering, Lund University, Box 124, S-221 00 Lund, Sweden Received 10 June 2005; accepted 8 December 2005
Abstract High-pressure homogenization of modified starches is a type of treatment used within the food and pharmaceutical industry for the production of emulsions. Such intense mechanical treatment may, however, influence the size and size distribution of the starch polymer, which may affect the properties of the emulsions. In this work, asymmetrical flow field-flow fractionation (AsFlFFF) connected on-line to a multi-angle light scattering detector (MALS) was used to study the influence of high-pressure homogenization upon molar mass and root-mean-square (r.m.s.) radius of three different octenyl succinate anhydride (OSA) starch samples originating from potato. The homogenization at constant pressure resulted in a significant decrease of both molar mass and r.m.s. radius. The magnitude of the change in molar mass and r.m.s. radius could be correlated to the initial size of the OSA starch and the energy dissipation rate of the homogenization treatment. r 2006 Elsevier Ltd. All rights reserved. Keywords: High pressure; Homogenization; Hydrophobically modified starch; Molar mass; Field-flow fractionation; Multi-angle light scattering
1. Introduction Emulsions can be formed from two immiscible liquids by dispersing one of the phases in the other. This is commonly achieved by applying intense mechanical energy to the system (Walstra, 1983) and can be done with various devices such as stirrers, high-pressure homogenizers, ultrasonication devices, etc. In many of these, such as stirrers and high-pressure homogenizers, turbulent flow plays an important role in the disruption of droplets. In emulsions macromolecules may have different effects (Bergensta˚hl & Claesson, 1997). They are commonly used to provide colloidal stability by adsorption at emulsion droplets to give steric stabilization. Non-adsorbing polymers may create a gel like viscosity of the continuous phase Corresponding author. Tel.: +46 46 222 9670; fax: +46 46 222 9517.
E-mail address: [email protected] (L. Nilsson). 0268-005X/$ - see front matter r 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodhyd.2005.12.011
providing stability. Non-adsorbing polymers may also destabilize emulsions by depletion destabilization. The molar mass distribution of the macromolecule is important in these cases as it affects the adsorption behaviour, the ability to act as a thickener as well as its tendency to induce depletion aggregation. Hydrophobically modified starch, such as octenyl succinate anhydride (OSA) starch (Caldwell & Wurzburg, 1953), is an amphiphilic macromolecule. It offers properties that have many applications within the formation and stabilization of dispersed food systems such as emulsions. Through the chemical modification the starch obtains substituents that show both hydrophobic and anionic properties. The degree of substitution in food applications typically ranges from approximately 0.02 and downwards (Shogren, Viswanathan, Felker, & Gross, 2000). Due to its high molar mass and branched polymer structure hydrophobically modified starch that is adsorbed at the interface
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will give rise to steric stabilization in emulsions and other dispersed systems and it is possible to produce stable emulsions at low OSA starch concentrations (Nilsson, & Bergensta˚hl, 2006; Tesch, Gerhards, & Schubert, 2002). The OSA starch can also act as an associative thickener and can thus be used to change the rheological characteristics of a system (Ortega-Ojeda, Larsson, & Eliasson, 2005). It is known that intense mechanical treatment, such as high-pressure homogenization, can influence the molar mass distribution and functionality of macromolecules. The degradation or disruption of polymers due to mechanical stress has been studied by several authors and some results from these studies are summarized in Table 1. Harrington and Zimm studied the degradation of polystyrene during passage through a piston–cylinder apparatus using viscometry and estimated a critical stress for degradation of the polymer (Harrington & Zimm, 1965). The degradation in this case roughly followed a firstorder rate law and the critical stress depended upon the solvent used, i.e. how stretched the polymer chain was. Buchholz et al. investigated the possibility to narrow the polydispersity index of different high molar mass polymers by injecting the polymers in a capillary under high pressure (Buchholz, Zahn, Kenward, Slater, & Barron, 2004). The authors found a decrease in polydispersity index and average molar mass as analysed by size-exclusion chromatography coupled to multi-angle light scattering and refractive index detectors (SEC–MALS–RI). The phenomenon was interpreted to be of a physical nature and independent of the chemical nature of the polymers investigated. Furthermore, it was suggested that scission occurred when the polymers exceeded a flow-field dependent critical chain length. Studies dealing with the disruption of polysaccharides have been carried out using various methods and types of polysaccharides. Silvestri and Gabrielson (1991) studied
the degradation of gum tragacanth during high-pressure homogenization using a Microfluidizer device. The authors followed the reasoning of Harrington and Zimm (1965) and found that the intrinsic viscosity of samples decreased as the number of passes or pressure increased. Highpressure homogenization using a Microfluidizer device was also used by Kasaai et al. to study fragmentation of chitosan (Kasaai, Charlet, Paquin, & Arul, 2003). The results showed that the extent of degradation increased with the applied chamber pressure and they also showed that the larger polymers were preferentially degraded. Floury et al. studied degradation of methylcellulose during high-pressure homogenization using SEC–MALS–RI and viscometry (Floury, Desrumaux, Axelos, & Legrand, 2002). The authors observed weaker thickening characteristics of the methylcellulose after highpressure homogenization which was coupled to a decrease in molar mass. Since the molar mass of modified starches often belongs in the ultra-high range (i.e. 4107 g/mol) chromatographic fractionation methods like SEC may fail in retrieving the entire distribution of a sample. Application of SEC to such large molecules further involves a high risk of shear degradation due to the fine porous structure of the column packing. Flow field-flow fractionation (FlFFF), however, is due to its open channel geometry and absence of stationary phase, able to size fractionate materials of a very wide size range from 1 nm up to 50 mm (Giddings, 1993). The technique provides very gentle conditions of separation, allowing large fragile molecules to be analysed without degradation (Giddings, 1993). Furthermore, most separations are fast, often requiring less than 10 min. Connecting FlFFF to MALS and RI detectors (FlFFF–MALS–RI) allows determination of molar mass (M) and root-mean-square (r.m.s.) radius as well as their distributions in a size domain where traditional methods are likely to be unsuccessful.
Table 1 Mechanical disruption of various polymers Polymer/processing method
Mw,0 (106 g/mol)
Mw (106g/mol)
P (MPa)
Analysis method
Gum tragacanth/HPHa Methylcellulose/HPH2b Chitosan/HPHc Polystyrene/HPHd Polyethyleneoxide/PLCe Polyacrylamide/PLCe Polydimethylacrylamide/PLCe
0.84 0.30 2.0 10.5 2.7 4.1 4.3
0.41 0.23 1.7 1.5 0.48 1.2 1.9
52 50 21 22 N/A N/A N/A
Viscometry SEC-MALS-RI Viscometry Viscometry SEC-MALS-RI SEC-MALS-RI SEC-MALS-RI
Mw,0 denotes initial weight-average molar mass, Mw denotes weight-average molar mass after processing, and P the pressure used for homogenization. The processing methods are HPH: high-pressure homogenization and PLC: pressure loaded capillary. The analysis methods are viscometry and size exclusion chromatography coupled with multi-angle light scattering and refractive index detection (SEC–MALS–RI). All polymers were dissolved in water except polystyrene, which was dissolved in a-methylnaphtalene. References in the table refer to: a Silvestri and Gabrielson (1991). b Floury et al. (2002). c Kasaai et al. (2003). d Harrington and Zimm (1965). e Buchholz et al. (2004).
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In the present study, three different OSA starches from potato were examined by asymmetrical flow FFF (AsFlFFF) combined with MALS and RI detection. The aim was to determine if high-pressure homogenization at constant pressure affects the molar mass of OSA starch solutions that have varying initial molar mass. The aim was furthermore to characterize the magnitude of the change in molar mass and r.m.s. radius. 2. Materials and methods 2.1. Solutions of octenyl succinic anhydride starch derivatives Three OSA starch samples, represented as OSA starch I, OSA starch II, and OSA starch III, varying in degree of substitution and expected molar mass were provided by Lyckeby Sta¨rkelsen (Kristianstad, Sweden) and were of native potato origin containing 20% amylose and 80% amylopectin. The degree of substitution was determined with 1H-NMR. The viscosity was measured using Brabender viscometry. The water content was approximately 10% in the samples. OSA starch solutions of 1% (w/v) were prepared by dispersing 1.0 g of OSA starch in a 10 mM phosphate buffer (pH 6.0) containing 20 ppm NaN3, which was then diluted to 100 ml. The OSA starches were then placed in a boiling water bath under stirring for 10 min after which they were left overnight at room temperature. Half of each of the three OSA starch solutions was then homogenized at 15 MPa in a high-pressure lab-scale valve homogenizer, while the other half of the solution was kept nonhomogenized. All homogenizations were carried out at room temperature. The homogenizer was an in-house construction, equipped with a ball valve of stainless steel and has been described in detail previously (Tornberg & Lundh, 1978). Solutions of homogenized and non-homogenized OSA starch were subsequently diluted to 0.025% (w/w) or 0.1% (w/w) in the FFF carrier liquid prior to FFF sample analysis. 2.2. Measurements using AsFlFFF The OSA starch samples were diluted in the FFF carrier liquid once and analysed by AsFlFFF–MALS–RI twice (parallel analysis). The parallel runs were made immediately after each other. The carrier used for AsFlFFF was deionized water (Milli-Q, Millipore, Bedford, MA, USA) containing 10 mM NaNO3 (p.a. grade, Merck, Darmstadt, Germany) together with 0.002% (w/v) NaN3 (Merck). The carrier was filtered through a 0.20 mm pore-size regenerated cellulose filter (ord. no. 18407, Sartorius AG, Goettingen, Germany) to remove particulate impurities. The fractionations were carried out using for the AsFlFFF an Eclipse F System (Wyatt Technology, Santa Barbara, CA, USA) serially connected to a MALS detector Dawn DSP laser photometer (Wyatt Technology) and an
Fig. 1. A schematic drawing of the asymmetrical flow FFF–MALS system with important components indicated. The pump (P) delivers the carrier flow which is degassed by an on-line vacuum degasser (D). The arrows indicate the flow directions during injection (dashed line) and elution (solid line) phase. ‘‘FIN’’ and ‘‘SIN’’ denotes the channel flow and sample flow respectively and ‘‘A’’ the position of the autoinjector. The crossflow (Fc) was adjusted by a software controlled cross-flow regulator denoted ‘‘C’’ The filter ‘‘f1’’ with a pore size of 20 nm ensures that all carrier entering the channel is particle free. The optional post-channel filter (f2) removes large (X2 mm) particulates from the outflow ‘‘Fout’’, before reaching the detectors.
Optilab DSP interferometric refractometer (Wyatt Technology) (Fig. 1). Both detectors used a wavelength of 632.8 nm. In the Eclipse F system, the magnitude of the injection flow and channel flow is regulated by the motordriven needle valves ‘‘N1’’ and ‘‘N2’’, respectively (Fig. 1). The crossflow is controlled using a Liquiflow regulator (Bronkhorst Hightech B.V, Ruurlo, The Netherlands). The needle valves as well as the crossflow-controller ‘‘C’’ are software-controlled and continuously adjusted in accordance with set values, thus allowing the use of programmed fields. The nominal thickness of the channel was 250 mm and the ultrafiltration membrane forming the accumulation wall was made of regenerated cellulose with a cut-off of 10 kD (C010F, Microdyn-Nadir GmbH, Wiesbaden, Germany). An Agilent 1100 Series Isocratic HPLC Pump (Agilent Technologies, Bo¨blingen, Germany) with an inline vacuum degasser delivered the carrier flow to the channel. Between the pump and the channel was placed a Teflon filter holder (an in-house construction) containing a 20 nm pore-sized aluminium oxide filter (Anodisc 25, Whatman, Maidstone, UK). Sample injections were made with an Agilent 1100 Series auto injector (Agilent Technologies) using a 100 ml loop volume resulting in an injected amount of 25 or 100 mg depending on the sample concentration. For the fractionations using a gradient in the crossflow, the injection time was 0.75 min at 0.4 ml/min, followed by a relaxation/focusing time of 1.25 min at a flow rate of 4.0 ml/min. Elution then followed at an outflow rate (Fout), of 1.0 ml/min and with a crossflow rate (Fc) decaying exponentially from 4.0 to 0.0 ml/min with a half-time of 6 min. The exponential gradient was approximated by programming the crossflow to decrease linearly step-wise in
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18 consecutive increments of equal step-length 2.0 min (except for the last step that was 3.0 min), and successively decreasing step-heights in the incremental change of the crossflow rate to fulfil a 6-min half-time. Zero crossflow rate was reached at 47 min retention time. For the fractionations using isocratic crossflow, the injection time was 1.20 min at 0.25 ml/min, followed by a relaxation/ focusing time of 5 min at a flow rate of 1.0 ml/min. Elution then followed at an outflow rate of 1.0 ml/min and with a crossflow rate of 0.2 ml/min. To remove large-sized impurities disturbing the MALS measurements, a small volume filter holder (A355, Upchurch Scientific, Oak Harbor, WA, USA), containing a replaceable 2 mm pore size PEEK frit (A700, Upchurch Scientific), was employed in-line between the channel and the MALS detector. All sample injections and fractionations were carried out at room temperature. 2.3. Data processing and calculations A computer software, ASTRA for Windows 4.73 (Wyatt Technology), was used for data collection and calculation of the molar mass and the r.m.s. radius. The basis for these calculations has been described earlier (Wyatt, 1993). The molar mass and r.m.s. radius were obtained by linear fitting to data in Debye plots according to the Berry method (Berry, 1966) and to guidelines given by Andersson, Wittgren, and Wahlund (2003). In these plots, scattering intensities from at least five different angles were employed. A value of the refractive index increment (dn/dc) of 0.15 ml/g, typical for starch (Aberle, Burchard, Vorwerg, & Radosta, 1994) was used in all calculations and the term containing the second virial coefficient, A2, was assumed to negligible. It was assumed that the low degree of substitution of the starch had no significant influence on the dn/dc. The recovery was calculated from the ratio of the mass eluted from the channel (determined by integration of the RI signal) to the mass injected. 3. Theory 3.1. Asymmetrical flow field-flow fractionation The separation takes place in a thin open channel through which a liquid flow is pumped resulting in a laminar flow with a parabolic velocity profile. The inlet flow is divided into a transverse crossflow, which exits through the semipermeable wall, and an axial flow that transports the sample along the channel (Fig. 2). The separation is induced by the crossflow field, which drives the sample components towards the semipermeable wall. Brownian motion of the sample components will counteract the crossflow-induced drift. This results in different mean distances from the semipermeable wall for sample components that have different Brownian motion characteristics, i.e. different diffusion coefficients, D. Thus, the separation is, in this so-called normal mode (Giddings,
Fig. 2. The principle of asymmetrical flow FFF. The laminar axial carrier flow with a parabolic velocity profile is pumped along the channel. The sample components are driven in the axial direction by the axial flow vector and in the transverse direction by the transverse crossflow vector, which compels them towards the permeable wall. Brownian motion of the sample components will counteract the transverse crossflow-induced drift, resulting in different mean distances from the semipermeable wall for the individual sample components. Consequently, the separation in time is dependent on each sample component’s position above the semipermeable wall, which in turn is determined by the diffusion coefficient of each sample component.
1993), strictly based upon differences in diffusion coefficients, which allows separation of any components that differ in size and shape. If the diffusion coefficient and the flow conditions are known, the retention time, tr, can be predicted from (Wahlund & Giddings, 1987) w2 F c t0 , (1) 6V 0 D where w is the channel thickness, t0 is the void time and V0 is the geometrical volume (void volume) of the channel. Conversely, the diffusion coefficient of a sample component can be calculated from experimental retention times. It should be remembered that Eq. (1) is a simplification, which only applies to rather high retention levels, tr/t0, i.e. 45.7 (Wittgren, Wahlund, Derand, & Wesslen, 1996). Moreover, it represents isocratic conditions where the crossflow rate is constant during the whole separation. If the crossflow rate varies in a gradient run (e.g. a linear decay or an exponential decay) the predictions/calculations of the retention time become more complicated (Williams & Giddings, 1994). The retention time, however, still increases with decreasing diffusion coefficient although not in direct proportion. tr ¼
3.2. Multi-angle light scattering Molar mass and size determination using MALS is well described in the literature and is based on the Rayleigh– Gans–Debye approximation of static light scattering (Wyatt, 1993). The molar mass (M) is derived from Kci 1 1 ¼ þ 2A2 ci (2) PðyÞi M i Ry by measuring the excess Rayleigh scattering, Ry, at different angles in each different slice i of the fractionated sample. The concentration in each slice, ci, is obtained through on-line RI detection. A2 is the second virial coefficient and K is an instrumental coefficient dependent
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on, e.g. the refractive index increment of the sample and the wavelength of the incident light. The particle scattering function P(y) describes the angular dependence of the intensity of the scattered light and depends on the wavelength of the incident light and the r.m.s. radius of the scattering object. Thus, information about the r.m.s. radius can be extracted from the angular dependence of the scattered light. The coupling of MALS to FlFFF has been intensely studied in recent years (Andersson, Wittgren, & Wahlund, 2001; Roessner & Kulicke, 1994; Wittgren & Wahlund, 1997b).
4. Results and discussion 4.1. Effects on molar mass and size by homogenization The molar mass and r.m.s. radius for the homogenized and non-homogenized OSA starches are given in Table 2 as determined by AsFlFFF–MALS–RI. The values presented are the medians of the distributions as these should be less Table 2 Median molar mass (Mm), median r.m.s. radius (rm), solution viscosity, and recovery of the three non-homogenized and homogenized OSA starches, respectivelya,b Sample
Solution viscosity
Mm (g/mol)
rm (nm)
Recovery (%)
OSA OSA OSA OSA OSA OSA
High n/d Low n/d High n/d
3.0 107 1.7 106 7.7 105 6.3 105 3.9 107 2.5 106
135 (1) 44 (5) n/d n/d 157 (4) 33 (3)
87 88 89 88 87 87
I (non-hom.) I (hom.) II (non-hom.) II (hom.) III (non-hom.) III (hom.)
(5) (1) (1) (2) (6) (3)
(1) (3) (1) (2) (2) (1)
Each value represents the average of two measurements. a Values in parentheses represent the percentage relative standard deviations. b Based on the manufacturer’s measurements using Brabender viscometry. 0.4
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influenced by uncertainties at the edges of the distributions than the weight- and z-averages, respectively. The median molar masses were in the ultra-high molar mass range (4107 g/mol) for the non-homogenized OSA starches I and III. These values fall within the range of earlier reports for dissolved native potato starch (BelloPerez, Roger, Baud & Colonna, 1998; Hanselmann, Ehrat & Widmer, 1995; Radosta, Haberer & Vorverg, 2001). The median molar mass was very much lower (o1 106 g/mol) for non-homogenized OSA starch II. After high-pressure homogenization a major reduction in both median molar mass and median r.m.s. radius was observed for the two ultra-high molar mass OSA starches I and III, but not for OSA starch II. Thus, the homogenization caused all three OSA starches to come below the ultrahigh molar mass level and the r.m.s. radii were all o50 nm. The data of Table 2 were based upon AsFlFFF fractionations as illustrated in Figs. 3–5. Due to the similarity in size and molar mass distribution width of the two ultra-high molar mass OSA starches (i.e. I and III) the flow conditions were kept the same for these. The much lower molar mass of OSA starch II made it be chosen to be fractionated under higher crossflow rate conditions in order to obtain an acceptable size resolution. For OSA starches I and III (Figs. 3 and 5 respectively) the LS901 signal was relatively much higher than the RI signal in the latter part of the response curve, than in the earlier part, which is typical for significantly polydisperse samples. This is explained by the higher sensitivity of the light scattering detector for larger sample components. There was further a striking difference in LS901 signal peak height between the homogenized and non-homogenized samples. This is indicative of the size difference, since scattering is directly proportional to molar mass. Moreover, the size reduction after homogenization is also indicated by the decrease in retention time of the LS901 peak maximum, as it is known that the retention time is inversely proportional to the diffusion coefficient (Eq. (1)) when isocratic crossflow is used. Since the same flow
10.0
0.25
9.0
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0.2 7.0 0.1 a 0.0 0
(A)
100 0.10 0.05 c
c t0
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a 0.15
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0.00 0
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8.0
RI signal (V)
0.3
log M
LS 90° signal (V)
b
b 10
t0
5
10
Retention time (min)
Fig. 3. Molar mass and 901 light scattering signal vs. retention time (A) and r.m.s. radius and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch I, and an almost non-visible blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass and r.m.s. radius for homogenized and non-homogenized OSA starch I, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: isocratic crossflow, Fc ¼ 0.2 ml/min, Fout ¼ 1.0 ml/min, t0 ¼ 0.62 min, mass load ¼ 25 mg, carrier: 10 mM NaNO3+0.002% NaN3.
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6.5
0.05
6.0
b a
0.15 RI signal (V)
LS 90o signal (V)
log M
LS 90° signal (V)
0.20
7.0
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0.10
(A)
t0
5
a
c
c 0.00 0
b
0.05
5.0
10
15
0.0 20
0.00 0 t0 (B)
Retention time (min)
5
10
15
20
Retention time (min)
10.0
0.4
9.0
0.3
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0.00
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RI signal (V)
0.20
1000
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100
0.10
(B)
R.m.s. radius
0.5
log M
LS 90° signal (V)
Fig. 4. Molar mass and 901 light scattering signal vs. retention time (A) and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch II, and a blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass for homogenized and non-homogenized OSA starch II, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: crossflow gradient, Fc decaying exponentially from 4.0 to 0.0 ml/min with a half-time of 6 min, Fout ¼ 1.0 ml/min, t0E0.3, mass load ¼ 100 mg, carrier: 10 mM NaNO3+0.002% NaN3.
10 0 t0
5 10 Retention time (min)
Fig. 5. Molar mass and 901 light scattering signal vs. retention time (A) and r.m.s. radius and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch III, and an almost non-visible blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass and r.m.s. radius for homogenized and non-homogenized OSA starch III, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: isocratic crossflow, Fc ¼ 0.2 ml/min, Fout ¼ 1.0 ml/min, t0 ¼ 0.62 min, mass load ¼ 25 mg, carrier: 10 mM NaNO3+0.002% NaN3.
conditions were applied for the non-homogenized OSA starches I and III, the somewhat longer retention time of the latter also shows it to be of a larger size. The trend line for the r.m.s. radius vs. retention time for this sample did, however, show an unexpected negative derivative during the retention time range 0–1.2 min (Fig. 5B). Such apparent deviations from so-called normal mode (Giddings, 1993) behaviour close to the void time may be caused by flow instabilities in the detector cells following the switching from focusing to elution (Litze´n, 1993; Wittgren & Wahlund, 1997a). Therefore, the deviant size data in Fig. 5B during the aforementioned retention time range were excluded from further evaluation. Higher retention conditions (i.e. increased crossflow) could of course have been applied for the homogenized samples in order to better resolve the smallest size components but this was not attempted. The small second peak in the LS901 fractograms (Figs. 3A and 5A) of the homogenized OSA starch I and III samples indicates the presence of ultra-high molar mass
components (likely 4109 g/mol and 4700 nm r.m.s. radius) in very low abundance, possibly remnants of the structure of the granules. No molar mass and size data were available for these peaks due to the too low signal to noise ratio for RI detector signal. For OSA starch II the LS901 and RI fractograms of the homogenized and non-homogenized samples shown in Fig. 4 were nearly identical, which explains the similarities in determined median molar mass (see Table 2). The r.m.s. radius could not be determined due to a very high imprecision of the r.m.s. radii data that scattered the range 10–40 nm. This is close to the lower limit for determination of this parameter. Due to a high RI and MALS signal for the blank, the measurement of molar mass was perturbed during the retention time range 0–2.8 min. No compensation was made for the high blanks and molar mass data were therefore only presented and evaluated for tr42.8 min. For all OSA starch fractionations, molar mass data were only evaluated for an RI detector signal-to-noise ratio
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(S/N)42. Below this level, e.g. at the trailing part of the RI peaks, the molar mass is too unreliable to be reported. The retention times of OSA starch II in Fig. 4 are not directly comparable with those of Figs. 3 and 5 due to the difference in flow conditions in comparison with the ones used for OSA starches I and III. 4.2. Molar mass and size distributions The fractionations illustrated in Figs. 3–5 demonstrate the widths of the molar mass and r.m.s. radius distributions, i.e. the polydisperse character of OSA starch. For the non-homogenized OSA starches I and III the molar mass distribution widths were very similar ranging at least 3 decades (precise enough data from the fronts and tails of the distributions were not obtained) from about 0.8 106 to 2 109 g/mol for OSA starch I, and from 1 106 to 4 109 g/mol for OSA starch III (Figs. 3A and 5A). For the non-homogenized OSA starch II the molar mass distribution width covered only slightly more than one decade, i.e. from 0.8 105 to 3 106 g/mol (Fig. 4A). Likewise, the r.m.s. radius distribution widths were similar for the two non-homogenized OSA starches I and III, i.e. 40–700 and 60–800 nm, respectively (Figs. 3B and 5B). There was a drastic decrease in molar mass and r.m.s. radius upon homogenization of OSA starches I and III (Figs. 3A and 5A) but no change at all for OSA starch II (Fig. 4A). Homogenized OSA starches I and III covered an r.m.s. width of (3–35) 101 and (2–26) 101 nm, respectively (Figs. 3B and 5B). In summary, OSA starches I and III were very similar to each other in molar mass and r.m.s. radius distributions both before and after homogenization whereas OSA starch II was unaffected by homogenization, apparently because its molecular size was much smaller than the others. Indeed, the present determination of the enormous size and polydispersity of these OSA starches illustrates the excellent methodological strength of flow FFF. 4.3. Disruption and shear The results show that high-pressure homogenization of OSA starch solutions affects the molar mass distribution and the r.m.s. radius. The magnitude of the effect seems to depend on the level of initial molar mass and apparent molecular size of the polymer, where a high initial molar mass gives a large reduction in molar mass after homogenization. The resulting molar mass is in a comparable range to those reported by other authors, which are given in Table 1. This change in molar mass could both be due to the breaking of covalent bonds, i.e. intramolecular bonds, or intermolecular non-covalent interactions that would result in the disruption of aggregates. From the results in this study it is impossible to rule out any of these explanations and it is therefore reasonable to assume that both of the suggested events occur. As has been pointed out above it is interesting that the r.m.s. radius becomes
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similar for the OSA starches after homogenization. The difference in molar mass between the samples is also somewhat reduced by homogenization. This suggests that there is some limiting size for the OSA starch depending on the processing conditions as has earlier been described by other authors for other polymers (Buchholz et al., 2004; Kasaai et al., 2003; Striegel, 2003). The energy that is released during high-pressure homogenization is usually quite high and is described by the energy dissipation rate e, which can be expressed in W/kg. The energy dissipation rate depends mainly on the homogenization pressure and the volume in which the energy is dissipated. A typical value of e for high-pressure homogenizers is 109 W/kg (Walstra, 1983). During the homogenization the flow conditions will be mostly turbulent and locally it can be assumed that the turbulence is isotropic as this simplifies the description of the turbulence considerably. Isotropic turbulence has been described by Kolmogorov (1949), Levich (1962) and others. An important parameter describing the turbulence is the Kolmogorov scale (Kolmogorov, 1949), which gives the magnitude of the turbulence micro-scale: 3 1=4 n l¼ , (3) e where l is the length scale of the smallest turbulent eddies, u is the kinematic viscosity (m2/s), and e is the energy dissipation rate per unit mass (W/kg). One should, however, remember that this expression is based on scaling laws and thus only gives an estimate of the turbulence micro-scale. In order to cause disruption of a macromolecule the length scale of the turbulent eddies should be smaller than the length scale of the macromolecule, as the eddies then generate a shear field over the macromolecule. This shear field could then ultimately lead to rupture of the macromolecule. If on the other hand, the length scale of the eddies are larger than the macromolecule they would not cause any disruption. In our case, if we assume u ¼ 106 m2/s and e ¼ 109 W/kg then lE200 nm. This value is smaller than the diameter of the OSA starches I and III that were disrupted and larger than II that was not noticeably disrupted. Furthermore, it is also known that macromolecules in turbulent flow become very extended depending on branching and chain flexibility and that scission is likely to occur at the midpoint (Groisman & Steinberg, 2001; Horn & Merrill, 1984). This suggests that the OSA starch is more extended during disruption in the high-pressure homogenizer than during the subsequent analysis with FFF–MALS–RI. 4.4. Molar mass of OSA starch during emulsification In this investigation, we have shown that high-pressure homogenization reduces the molar mass of OSA starch. Thus, it is likely to assume that a similar degradation occurs during a normal emulsification process. During
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emulsification OSA starch is included in the system before the homogenization process to be able to act as an emulsifier in the same moment as the droplets are formed in the homogenizer. A discussion about the role of the molar mass and emulsification properties have rather to be based on the molar mass measured after homogenization than on the molar mass before homogenization. OSA starch typically has quite a high median molar mass, in this investigation, in the range (0.8–40) 106 g/mol. After homogenization the median molar mass still remains high, (0.6–2) 106 g/mol, but it is less extreme. Moreover, for OSA starches I and III the homogenization causes the distribution widths to decrease. 5. Conclusion AsFlFFF–MALS–RI is a unique analysis tool to study the molar mass distribution of OSA starch it being a polydisperse macromolecule with ultra-high molar mass components. High-pressure homogenization causes disruption of OSA starch and the extent of this depends on the initial size of the OSA starch and the energy dissipation rate during processing. The resulting molar mass after high-pressure homogenization is in a comparable range to the results reported by other authors for different polymers (Table 1). A comparison between our results and those given in Table 1 shows that high-pressure homogenized OSA starch is still a high molar mass hydrophobically modified polymer. Acknowledgements This study was financed by the Centre for Amphiphilic Polymers from Renewable Sources (CAP) at Lund University. The support from the Swedish Research Council is acknowledged by one of us (KGW). Lyckeby Sta¨rkelsen, Kristianstad is acknowledged for financial support and for supplying the OSA starch samples. References Aberle, T., Burchard, W., Vorwerg, W., & Radosta, S. (1994). Conformational contributions of amylose and amylopectin to the structural properties of starches from various sources. Starch/Staerke, 46, 329–335. Andersson, M., Wittgren, B., & Wahlund, K.-G. (2001). Ultrahigh molar mass component detected in ethyl hydroxyethyl cellulose by asymmetrical flow field-flow fractionation coupled to multi-angle light scattering. Analytical Chemistry, 73, 4852–4861. Andersson, M., Wittgren, B., & Wahlund, K. G. (2003). Accuracy in multiangle light scattering measurements for molar mass and radius estimations. Model calculations and experiments. Analytical Chemistry, 75, 4279–4291. Bello-Perez, L. A., Roger, P., Baud, B., & Colonna, P. (1998). Macromolecular features of starches determined by aqueous highperformance size exclusion chromatography. Journal of Cereal Science, 27, 267–278. Bergensta˚hl, B., & Claesson, P. (1997). Surface forces in emulsions. In S. E. Friberg, & K. Larsson (Eds.), Food emulsions (pp. 57–109). New York: Marcel Dekker.
Berry, G. C. (1966). Thermodynamic and conformational properties of polystyrene. I. Light scattering studies on dilute solutions of linear polystyrenes. Journal of Chemical Physics, 44, 4550–4564. Buchholz, B. A., Zahn, J. A., Kenward, M., Slater, G. W., & Barron, A. E. (2004). Flow-induced chain scission as a physical route to narrowly distributed high molar mass polymers. Polymer, 45, 1223–1234. Caldwell, C. G., & Wurzburg, O. B. (1953). Polysaccharide derivatives of substituted dicarboxylic acids. US Patent 2-661-349. Floury, J., Desrumaux, A., Axelos, M. A. V., & Legrand, J. (2002). Degradation of methylcellulose during ultra-high pressure homogenisation. Food Hydrocolloids, 16, 47–53. Giddings, J. C. (1993). Field-flow fractionation: Analysis of macromolecular, colloidal, and particulate materials. Science (Washington, DC, United States), 260, 1456–1465. Groisman, A., & Steinberg, V. (2001). Stretching of polymers in a random three-dimensional flow. Physical Review Letters, 86, 934–937. Hanselmann, R., Ehrat, M., & Widmer, H. M. (1995). Sedimentation field flow fractionation combined with multi angle laser light scattering applied for characterization of starch polymers. Starch/Staerke, 47, 345–349. Harrington, R. E., & Zimm, B. H. (1965). Degradation of polymers by controlled hydrodynamic shear. Journal of Physical Chemistry, 69, 161–175. Horn, A. F., & Merrill, E. W. (1984). Midpoint scission of macromolecules in dilute solution in turbulent flow. Nature, 312, 140–141. Kasaai, M. R., Charlet, G., Paquin, P., & Arul, J. (2003). Fragmentation of chitosan by microfluidization process. Innovative Food Science & Emerging Technologies, 4, 403–413. Kolmogorov, A. N. (1949). The breaking of droplets in turbulent flow. Doklady Akademii Nauk SSSR, 66, 825–828. Levich, V. G. (1962). Physicochemical hydrodynamics. Englewood Cliffs: Prentice-Hall. Litze´n, A. (1993). Separation speed, retention, and dispersion in asymmetrical flow field-flow fractionation as functions of channel dimensions and flow rates. Analytical Chemistry, 65, 461–470. Nilsson, L., & Bergensta˚hl, B. (2006). Adsorption of octenyl succinate anhydride starch esters at the oil/water interface. Manuscript. Ortega-Ojeda, F. E., Larsson, H., & Eliasson, A.-C. (2005). Gel formation in mixtures of hydrophobically modified potato and high amylopectin potato starch. Carbohydrate Polymers, 59, 313–327. Radosta, S., Haberer, M., & Vorverg, W. (2001). Molecular characteristics of amylose and starch in dimethyl sulfoxide. Biomacromolecules, 2, 970–978. Roessner, D., & Kulicke, W.-M. (1994). Online coupling of flow field-flow fractionation and multi-angle laser light scattering. Journal of Chromatography, A, 687, 249–258. Shogren, R. L., Viswanathan, A., Felker, F., & Gross, R. A. (2000). Distribution of octenyl succinate groups in octenyl succinic anhydride modified waxy maize starch. Starch/Sta¨rke, 52, 196–204. Silvestri, S., & Gabrielson, G. (1991). Degradation of tragacanth by high shear and turbulent forces during microfluidization. International Journal of Pharmaceutics, 73, 163–169. Striegel, A. M. (2003). Influence of chain architecture on the mechanochemical degradation of macromolecules. Journal of Biochemical and Biophysical Methods, 56, 117–139. Tesch, S., Gerhards, C., & Schubert, H. (2002). Stabilization of emulsions by OSA starches. Journal of Food Engineering, 54, 167–174. Tornberg, E., & Lundh, G. (1978). Functional characterization of protein stabilized emulsions: standardized emulsifying procedure. Journal of Food Science, 43, 1553–1558. Wahlund, K. G., & Giddings, J. C. (1987). Properties of an asymmetrical flow field-flow fractionation channel having one permeable wall. Analytical Chemistry, 59, 1332–1339.
ARTICLE IN PRESS G. Modig et al. / Food Hydrocolloids 20 (2006) 1087–1095 Walstra, P. (1983). Formation of emulsions. In P. Becher (Ed.), Encyclopedia of emulsion technology, Vol. I (pp. 57–127). New York: Marcel Dekker. Williams, P. S., & Giddings, J. C. (1994). Theory of field-programmed field-flow fractionation with corrections for steric effects. Analytical Chemistry, 66, 4215–4228. Wittgren, B., & Wahlund, K.-G. (1997a). Effects of flow-rates and sample concentration on the molar mass characterization of modified celluloses using asymmetrical flow field-flow fractionation-multi-angle light scattering. Journal of Chromatography, A, 135–149.
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Wittgren, B., & Wahlund, K.-G. (1997b). Fast molecular mass and size characterization of polysaccharides using asymmetrical flow field-flow fractionation-multiangle light scattering. Journal of Chromatography, A, 760, 205–218. Wittgren, B., Wahlund, K.-G., Derand, H., & Wesslen, B. (1996). Aggregation behavior of an amphiphilic graft copolymer in aqueous medium studied by asymmetrical flow field-flow fractionation. Macromolecules, 29, 268–276. Wyatt, P. (1993). Light scattering and the absolute characterization of macromolecules. Analytica Chimica Acta, 272, 1–40.
FOOD
HYDROCOLLOIDS Food Hydrocolloids 20 (2006) 1087–1095 www.elsevier.com/locate/foodhyd
Homogenization-induced degradation of hydrophobically modified starch determined by asymmetrical flow field-flow fractionation and multi-angle light scattering Gustaf Modigb, Lars Nilssona,, Bjo¨rn Bergensta˚hla, Karl-Gustav Wahlundb a
Faculty of Engineering LTH, Division of Food Technology, Centre for Chemistry and Chemical Engineering, Lund University, Box 124, S-221 00 Lund, Sweden b Faculty of Engineering LTH, Division of Technical Analytical Chemistry, Centre for Chemistry and Chemical Engineering, Lund University, Box 124, S-221 00 Lund, Sweden Received 10 June 2005; accepted 8 December 2005
Abstract High-pressure homogenization of modified starches is a type of treatment used within the food and pharmaceutical industry for the production of emulsions. Such intense mechanical treatment may, however, influence the size and size distribution of the starch polymer, which may affect the properties of the emulsions. In this work, asymmetrical flow field-flow fractionation (AsFlFFF) connected on-line to a multi-angle light scattering detector (MALS) was used to study the influence of high-pressure homogenization upon molar mass and root-mean-square (r.m.s.) radius of three different octenyl succinate anhydride (OSA) starch samples originating from potato. The homogenization at constant pressure resulted in a significant decrease of both molar mass and r.m.s. radius. The magnitude of the change in molar mass and r.m.s. radius could be correlated to the initial size of the OSA starch and the energy dissipation rate of the homogenization treatment. r 2006 Elsevier Ltd. All rights reserved. Keywords: High pressure; Homogenization; Hydrophobically modified starch; Molar mass; Field-flow fractionation; Multi-angle light scattering
1. Introduction Emulsions can be formed from two immiscible liquids by dispersing one of the phases in the other. This is commonly achieved by applying intense mechanical energy to the system (Walstra, 1983) and can be done with various devices such as stirrers, high-pressure homogenizers, ultrasonication devices, etc. In many of these, such as stirrers and high-pressure homogenizers, turbulent flow plays an important role in the disruption of droplets. In emulsions macromolecules may have different effects (Bergensta˚hl & Claesson, 1997). They are commonly used to provide colloidal stability by adsorption at emulsion droplets to give steric stabilization. Non-adsorbing polymers may create a gel like viscosity of the continuous phase Corresponding author. Tel.: +46 46 222 9670; fax: +46 46 222 9517.
E-mail address: [email protected] (L. Nilsson). 0268-005X/$ - see front matter r 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodhyd.2005.12.011
providing stability. Non-adsorbing polymers may also destabilize emulsions by depletion destabilization. The molar mass distribution of the macromolecule is important in these cases as it affects the adsorption behaviour, the ability to act as a thickener as well as its tendency to induce depletion aggregation. Hydrophobically modified starch, such as octenyl succinate anhydride (OSA) starch (Caldwell & Wurzburg, 1953), is an amphiphilic macromolecule. It offers properties that have many applications within the formation and stabilization of dispersed food systems such as emulsions. Through the chemical modification the starch obtains substituents that show both hydrophobic and anionic properties. The degree of substitution in food applications typically ranges from approximately 0.02 and downwards (Shogren, Viswanathan, Felker, & Gross, 2000). Due to its high molar mass and branched polymer structure hydrophobically modified starch that is adsorbed at the interface
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will give rise to steric stabilization in emulsions and other dispersed systems and it is possible to produce stable emulsions at low OSA starch concentrations (Nilsson, & Bergensta˚hl, 2006; Tesch, Gerhards, & Schubert, 2002). The OSA starch can also act as an associative thickener and can thus be used to change the rheological characteristics of a system (Ortega-Ojeda, Larsson, & Eliasson, 2005). It is known that intense mechanical treatment, such as high-pressure homogenization, can influence the molar mass distribution and functionality of macromolecules. The degradation or disruption of polymers due to mechanical stress has been studied by several authors and some results from these studies are summarized in Table 1. Harrington and Zimm studied the degradation of polystyrene during passage through a piston–cylinder apparatus using viscometry and estimated a critical stress for degradation of the polymer (Harrington & Zimm, 1965). The degradation in this case roughly followed a firstorder rate law and the critical stress depended upon the solvent used, i.e. how stretched the polymer chain was. Buchholz et al. investigated the possibility to narrow the polydispersity index of different high molar mass polymers by injecting the polymers in a capillary under high pressure (Buchholz, Zahn, Kenward, Slater, & Barron, 2004). The authors found a decrease in polydispersity index and average molar mass as analysed by size-exclusion chromatography coupled to multi-angle light scattering and refractive index detectors (SEC–MALS–RI). The phenomenon was interpreted to be of a physical nature and independent of the chemical nature of the polymers investigated. Furthermore, it was suggested that scission occurred when the polymers exceeded a flow-field dependent critical chain length. Studies dealing with the disruption of polysaccharides have been carried out using various methods and types of polysaccharides. Silvestri and Gabrielson (1991) studied
the degradation of gum tragacanth during high-pressure homogenization using a Microfluidizer device. The authors followed the reasoning of Harrington and Zimm (1965) and found that the intrinsic viscosity of samples decreased as the number of passes or pressure increased. Highpressure homogenization using a Microfluidizer device was also used by Kasaai et al. to study fragmentation of chitosan (Kasaai, Charlet, Paquin, & Arul, 2003). The results showed that the extent of degradation increased with the applied chamber pressure and they also showed that the larger polymers were preferentially degraded. Floury et al. studied degradation of methylcellulose during high-pressure homogenization using SEC–MALS–RI and viscometry (Floury, Desrumaux, Axelos, & Legrand, 2002). The authors observed weaker thickening characteristics of the methylcellulose after highpressure homogenization which was coupled to a decrease in molar mass. Since the molar mass of modified starches often belongs in the ultra-high range (i.e. 4107 g/mol) chromatographic fractionation methods like SEC may fail in retrieving the entire distribution of a sample. Application of SEC to such large molecules further involves a high risk of shear degradation due to the fine porous structure of the column packing. Flow field-flow fractionation (FlFFF), however, is due to its open channel geometry and absence of stationary phase, able to size fractionate materials of a very wide size range from 1 nm up to 50 mm (Giddings, 1993). The technique provides very gentle conditions of separation, allowing large fragile molecules to be analysed without degradation (Giddings, 1993). Furthermore, most separations are fast, often requiring less than 10 min. Connecting FlFFF to MALS and RI detectors (FlFFF–MALS–RI) allows determination of molar mass (M) and root-mean-square (r.m.s.) radius as well as their distributions in a size domain where traditional methods are likely to be unsuccessful.
Table 1 Mechanical disruption of various polymers Polymer/processing method
Mw,0 (106 g/mol)
Mw (106g/mol)
P (MPa)
Analysis method
Gum tragacanth/HPHa Methylcellulose/HPH2b Chitosan/HPHc Polystyrene/HPHd Polyethyleneoxide/PLCe Polyacrylamide/PLCe Polydimethylacrylamide/PLCe
0.84 0.30 2.0 10.5 2.7 4.1 4.3
0.41 0.23 1.7 1.5 0.48 1.2 1.9
52 50 21 22 N/A N/A N/A
Viscometry SEC-MALS-RI Viscometry Viscometry SEC-MALS-RI SEC-MALS-RI SEC-MALS-RI
Mw,0 denotes initial weight-average molar mass, Mw denotes weight-average molar mass after processing, and P the pressure used for homogenization. The processing methods are HPH: high-pressure homogenization and PLC: pressure loaded capillary. The analysis methods are viscometry and size exclusion chromatography coupled with multi-angle light scattering and refractive index detection (SEC–MALS–RI). All polymers were dissolved in water except polystyrene, which was dissolved in a-methylnaphtalene. References in the table refer to: a Silvestri and Gabrielson (1991). b Floury et al. (2002). c Kasaai et al. (2003). d Harrington and Zimm (1965). e Buchholz et al. (2004).
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In the present study, three different OSA starches from potato were examined by asymmetrical flow FFF (AsFlFFF) combined with MALS and RI detection. The aim was to determine if high-pressure homogenization at constant pressure affects the molar mass of OSA starch solutions that have varying initial molar mass. The aim was furthermore to characterize the magnitude of the change in molar mass and r.m.s. radius. 2. Materials and methods 2.1. Solutions of octenyl succinic anhydride starch derivatives Three OSA starch samples, represented as OSA starch I, OSA starch II, and OSA starch III, varying in degree of substitution and expected molar mass were provided by Lyckeby Sta¨rkelsen (Kristianstad, Sweden) and were of native potato origin containing 20% amylose and 80% amylopectin. The degree of substitution was determined with 1H-NMR. The viscosity was measured using Brabender viscometry. The water content was approximately 10% in the samples. OSA starch solutions of 1% (w/v) were prepared by dispersing 1.0 g of OSA starch in a 10 mM phosphate buffer (pH 6.0) containing 20 ppm NaN3, which was then diluted to 100 ml. The OSA starches were then placed in a boiling water bath under stirring for 10 min after which they were left overnight at room temperature. Half of each of the three OSA starch solutions was then homogenized at 15 MPa in a high-pressure lab-scale valve homogenizer, while the other half of the solution was kept nonhomogenized. All homogenizations were carried out at room temperature. The homogenizer was an in-house construction, equipped with a ball valve of stainless steel and has been described in detail previously (Tornberg & Lundh, 1978). Solutions of homogenized and non-homogenized OSA starch were subsequently diluted to 0.025% (w/w) or 0.1% (w/w) in the FFF carrier liquid prior to FFF sample analysis. 2.2. Measurements using AsFlFFF The OSA starch samples were diluted in the FFF carrier liquid once and analysed by AsFlFFF–MALS–RI twice (parallel analysis). The parallel runs were made immediately after each other. The carrier used for AsFlFFF was deionized water (Milli-Q, Millipore, Bedford, MA, USA) containing 10 mM NaNO3 (p.a. grade, Merck, Darmstadt, Germany) together with 0.002% (w/v) NaN3 (Merck). The carrier was filtered through a 0.20 mm pore-size regenerated cellulose filter (ord. no. 18407, Sartorius AG, Goettingen, Germany) to remove particulate impurities. The fractionations were carried out using for the AsFlFFF an Eclipse F System (Wyatt Technology, Santa Barbara, CA, USA) serially connected to a MALS detector Dawn DSP laser photometer (Wyatt Technology) and an
Fig. 1. A schematic drawing of the asymmetrical flow FFF–MALS system with important components indicated. The pump (P) delivers the carrier flow which is degassed by an on-line vacuum degasser (D). The arrows indicate the flow directions during injection (dashed line) and elution (solid line) phase. ‘‘FIN’’ and ‘‘SIN’’ denotes the channel flow and sample flow respectively and ‘‘A’’ the position of the autoinjector. The crossflow (Fc) was adjusted by a software controlled cross-flow regulator denoted ‘‘C’’ The filter ‘‘f1’’ with a pore size of 20 nm ensures that all carrier entering the channel is particle free. The optional post-channel filter (f2) removes large (X2 mm) particulates from the outflow ‘‘Fout’’, before reaching the detectors.
Optilab DSP interferometric refractometer (Wyatt Technology) (Fig. 1). Both detectors used a wavelength of 632.8 nm. In the Eclipse F system, the magnitude of the injection flow and channel flow is regulated by the motordriven needle valves ‘‘N1’’ and ‘‘N2’’, respectively (Fig. 1). The crossflow is controlled using a Liquiflow regulator (Bronkhorst Hightech B.V, Ruurlo, The Netherlands). The needle valves as well as the crossflow-controller ‘‘C’’ are software-controlled and continuously adjusted in accordance with set values, thus allowing the use of programmed fields. The nominal thickness of the channel was 250 mm and the ultrafiltration membrane forming the accumulation wall was made of regenerated cellulose with a cut-off of 10 kD (C010F, Microdyn-Nadir GmbH, Wiesbaden, Germany). An Agilent 1100 Series Isocratic HPLC Pump (Agilent Technologies, Bo¨blingen, Germany) with an inline vacuum degasser delivered the carrier flow to the channel. Between the pump and the channel was placed a Teflon filter holder (an in-house construction) containing a 20 nm pore-sized aluminium oxide filter (Anodisc 25, Whatman, Maidstone, UK). Sample injections were made with an Agilent 1100 Series auto injector (Agilent Technologies) using a 100 ml loop volume resulting in an injected amount of 25 or 100 mg depending on the sample concentration. For the fractionations using a gradient in the crossflow, the injection time was 0.75 min at 0.4 ml/min, followed by a relaxation/focusing time of 1.25 min at a flow rate of 4.0 ml/min. Elution then followed at an outflow rate (Fout), of 1.0 ml/min and with a crossflow rate (Fc) decaying exponentially from 4.0 to 0.0 ml/min with a half-time of 6 min. The exponential gradient was approximated by programming the crossflow to decrease linearly step-wise in
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18 consecutive increments of equal step-length 2.0 min (except for the last step that was 3.0 min), and successively decreasing step-heights in the incremental change of the crossflow rate to fulfil a 6-min half-time. Zero crossflow rate was reached at 47 min retention time. For the fractionations using isocratic crossflow, the injection time was 1.20 min at 0.25 ml/min, followed by a relaxation/ focusing time of 5 min at a flow rate of 1.0 ml/min. Elution then followed at an outflow rate of 1.0 ml/min and with a crossflow rate of 0.2 ml/min. To remove large-sized impurities disturbing the MALS measurements, a small volume filter holder (A355, Upchurch Scientific, Oak Harbor, WA, USA), containing a replaceable 2 mm pore size PEEK frit (A700, Upchurch Scientific), was employed in-line between the channel and the MALS detector. All sample injections and fractionations were carried out at room temperature. 2.3. Data processing and calculations A computer software, ASTRA for Windows 4.73 (Wyatt Technology), was used for data collection and calculation of the molar mass and the r.m.s. radius. The basis for these calculations has been described earlier (Wyatt, 1993). The molar mass and r.m.s. radius were obtained by linear fitting to data in Debye plots according to the Berry method (Berry, 1966) and to guidelines given by Andersson, Wittgren, and Wahlund (2003). In these plots, scattering intensities from at least five different angles were employed. A value of the refractive index increment (dn/dc) of 0.15 ml/g, typical for starch (Aberle, Burchard, Vorwerg, & Radosta, 1994) was used in all calculations and the term containing the second virial coefficient, A2, was assumed to negligible. It was assumed that the low degree of substitution of the starch had no significant influence on the dn/dc. The recovery was calculated from the ratio of the mass eluted from the channel (determined by integration of the RI signal) to the mass injected. 3. Theory 3.1. Asymmetrical flow field-flow fractionation The separation takes place in a thin open channel through which a liquid flow is pumped resulting in a laminar flow with a parabolic velocity profile. The inlet flow is divided into a transverse crossflow, which exits through the semipermeable wall, and an axial flow that transports the sample along the channel (Fig. 2). The separation is induced by the crossflow field, which drives the sample components towards the semipermeable wall. Brownian motion of the sample components will counteract the crossflow-induced drift. This results in different mean distances from the semipermeable wall for sample components that have different Brownian motion characteristics, i.e. different diffusion coefficients, D. Thus, the separation is, in this so-called normal mode (Giddings,
Fig. 2. The principle of asymmetrical flow FFF. The laminar axial carrier flow with a parabolic velocity profile is pumped along the channel. The sample components are driven in the axial direction by the axial flow vector and in the transverse direction by the transverse crossflow vector, which compels them towards the permeable wall. Brownian motion of the sample components will counteract the transverse crossflow-induced drift, resulting in different mean distances from the semipermeable wall for the individual sample components. Consequently, the separation in time is dependent on each sample component’s position above the semipermeable wall, which in turn is determined by the diffusion coefficient of each sample component.
1993), strictly based upon differences in diffusion coefficients, which allows separation of any components that differ in size and shape. If the diffusion coefficient and the flow conditions are known, the retention time, tr, can be predicted from (Wahlund & Giddings, 1987) w2 F c t0 , (1) 6V 0 D where w is the channel thickness, t0 is the void time and V0 is the geometrical volume (void volume) of the channel. Conversely, the diffusion coefficient of a sample component can be calculated from experimental retention times. It should be remembered that Eq. (1) is a simplification, which only applies to rather high retention levels, tr/t0, i.e. 45.7 (Wittgren, Wahlund, Derand, & Wesslen, 1996). Moreover, it represents isocratic conditions where the crossflow rate is constant during the whole separation. If the crossflow rate varies in a gradient run (e.g. a linear decay or an exponential decay) the predictions/calculations of the retention time become more complicated (Williams & Giddings, 1994). The retention time, however, still increases with decreasing diffusion coefficient although not in direct proportion. tr ¼
3.2. Multi-angle light scattering Molar mass and size determination using MALS is well described in the literature and is based on the Rayleigh– Gans–Debye approximation of static light scattering (Wyatt, 1993). The molar mass (M) is derived from Kci 1 1 ¼ þ 2A2 ci (2) PðyÞi M i Ry by measuring the excess Rayleigh scattering, Ry, at different angles in each different slice i of the fractionated sample. The concentration in each slice, ci, is obtained through on-line RI detection. A2 is the second virial coefficient and K is an instrumental coefficient dependent
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on, e.g. the refractive index increment of the sample and the wavelength of the incident light. The particle scattering function P(y) describes the angular dependence of the intensity of the scattered light and depends on the wavelength of the incident light and the r.m.s. radius of the scattering object. Thus, information about the r.m.s. radius can be extracted from the angular dependence of the scattered light. The coupling of MALS to FlFFF has been intensely studied in recent years (Andersson, Wittgren, & Wahlund, 2001; Roessner & Kulicke, 1994; Wittgren & Wahlund, 1997b).
4. Results and discussion 4.1. Effects on molar mass and size by homogenization The molar mass and r.m.s. radius for the homogenized and non-homogenized OSA starches are given in Table 2 as determined by AsFlFFF–MALS–RI. The values presented are the medians of the distributions as these should be less Table 2 Median molar mass (Mm), median r.m.s. radius (rm), solution viscosity, and recovery of the three non-homogenized and homogenized OSA starches, respectivelya,b Sample
Solution viscosity
Mm (g/mol)
rm (nm)
Recovery (%)
OSA OSA OSA OSA OSA OSA
High n/d Low n/d High n/d
3.0 107 1.7 106 7.7 105 6.3 105 3.9 107 2.5 106
135 (1) 44 (5) n/d n/d 157 (4) 33 (3)
87 88 89 88 87 87
I (non-hom.) I (hom.) II (non-hom.) II (hom.) III (non-hom.) III (hom.)
(5) (1) (1) (2) (6) (3)
(1) (3) (1) (2) (2) (1)
Each value represents the average of two measurements. a Values in parentheses represent the percentage relative standard deviations. b Based on the manufacturer’s measurements using Brabender viscometry. 0.4
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influenced by uncertainties at the edges of the distributions than the weight- and z-averages, respectively. The median molar masses were in the ultra-high molar mass range (4107 g/mol) for the non-homogenized OSA starches I and III. These values fall within the range of earlier reports for dissolved native potato starch (BelloPerez, Roger, Baud & Colonna, 1998; Hanselmann, Ehrat & Widmer, 1995; Radosta, Haberer & Vorverg, 2001). The median molar mass was very much lower (o1 106 g/mol) for non-homogenized OSA starch II. After high-pressure homogenization a major reduction in both median molar mass and median r.m.s. radius was observed for the two ultra-high molar mass OSA starches I and III, but not for OSA starch II. Thus, the homogenization caused all three OSA starches to come below the ultrahigh molar mass level and the r.m.s. radii were all o50 nm. The data of Table 2 were based upon AsFlFFF fractionations as illustrated in Figs. 3–5. Due to the similarity in size and molar mass distribution width of the two ultra-high molar mass OSA starches (i.e. I and III) the flow conditions were kept the same for these. The much lower molar mass of OSA starch II made it be chosen to be fractionated under higher crossflow rate conditions in order to obtain an acceptable size resolution. For OSA starches I and III (Figs. 3 and 5 respectively) the LS901 signal was relatively much higher than the RI signal in the latter part of the response curve, than in the earlier part, which is typical for significantly polydisperse samples. This is explained by the higher sensitivity of the light scattering detector for larger sample components. There was further a striking difference in LS901 signal peak height between the homogenized and non-homogenized samples. This is indicative of the size difference, since scattering is directly proportional to molar mass. Moreover, the size reduction after homogenization is also indicated by the decrease in retention time of the LS901 peak maximum, as it is known that the retention time is inversely proportional to the diffusion coefficient (Eq. (1)) when isocratic crossflow is used. Since the same flow
10.0
0.25
9.0
0.20
1000
0.2 7.0 0.1 a 0.0 0
(A)
100 0.10 0.05 c
c t0
6.0
a 0.15
5.0 5 10 Retention time (min)
0.00 0
(B)
R.m.s. radius
8.0
RI signal (V)
0.3
log M
LS 90° signal (V)
b
b 10
t0
5
10
Retention time (min)
Fig. 3. Molar mass and 901 light scattering signal vs. retention time (A) and r.m.s. radius and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch I, and an almost non-visible blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass and r.m.s. radius for homogenized and non-homogenized OSA starch I, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: isocratic crossflow, Fc ¼ 0.2 ml/min, Fout ¼ 1.0 ml/min, t0 ¼ 0.62 min, mass load ¼ 25 mg, carrier: 10 mM NaNO3+0.002% NaN3.
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6.5
0.05
6.0
b a
0.15 RI signal (V)
LS 90o signal (V)
log M
LS 90° signal (V)
0.20
7.0
0.10
5.5
0.10
(A)
t0
5
a
c
c 0.00 0
b
0.05
5.0
10
15
0.0 20
0.00 0 t0 (B)
Retention time (min)
5
10
15
20
Retention time (min)
10.0
0.4
9.0
0.3
8.0
0.2
7.0
0.1
6.0
0.05
5.0
0.00
0.0 0
(A)
t0
5 10 Retention time (min)
RI signal (V)
0.20
1000
0.15
100
0.10
(B)
R.m.s. radius
0.5
log M
LS 90° signal (V)
Fig. 4. Molar mass and 901 light scattering signal vs. retention time (A) and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch II, and a blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass for homogenized and non-homogenized OSA starch II, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: crossflow gradient, Fc decaying exponentially from 4.0 to 0.0 ml/min with a half-time of 6 min, Fout ¼ 1.0 ml/min, t0E0.3, mass load ¼ 100 mg, carrier: 10 mM NaNO3+0.002% NaN3.
10 0 t0
5 10 Retention time (min)
Fig. 5. Molar mass and 901 light scattering signal vs. retention time (A) and r.m.s. radius and RI signal vs. retention time (B) for homogenized (a) and non-homogenized (b) OSA starch III, and an almost non-visible blank run (c). The symbols ‘J’ and ‘&’ notate the log molar mass and r.m.s. radius for homogenized and non-homogenized OSA starch III, respectively. Continuous line curves represent the fractograms obtained from the MALS photometer at angle 901 (LS901) and the RI detector, respectively. Fractionation conditions: isocratic crossflow, Fc ¼ 0.2 ml/min, Fout ¼ 1.0 ml/min, t0 ¼ 0.62 min, mass load ¼ 25 mg, carrier: 10 mM NaNO3+0.002% NaN3.
conditions were applied for the non-homogenized OSA starches I and III, the somewhat longer retention time of the latter also shows it to be of a larger size. The trend line for the r.m.s. radius vs. retention time for this sample did, however, show an unexpected negative derivative during the retention time range 0–1.2 min (Fig. 5B). Such apparent deviations from so-called normal mode (Giddings, 1993) behaviour close to the void time may be caused by flow instabilities in the detector cells following the switching from focusing to elution (Litze´n, 1993; Wittgren & Wahlund, 1997a). Therefore, the deviant size data in Fig. 5B during the aforementioned retention time range were excluded from further evaluation. Higher retention conditions (i.e. increased crossflow) could of course have been applied for the homogenized samples in order to better resolve the smallest size components but this was not attempted. The small second peak in the LS901 fractograms (Figs. 3A and 5A) of the homogenized OSA starch I and III samples indicates the presence of ultra-high molar mass
components (likely 4109 g/mol and 4700 nm r.m.s. radius) in very low abundance, possibly remnants of the structure of the granules. No molar mass and size data were available for these peaks due to the too low signal to noise ratio for RI detector signal. For OSA starch II the LS901 and RI fractograms of the homogenized and non-homogenized samples shown in Fig. 4 were nearly identical, which explains the similarities in determined median molar mass (see Table 2). The r.m.s. radius could not be determined due to a very high imprecision of the r.m.s. radii data that scattered the range 10–40 nm. This is close to the lower limit for determination of this parameter. Due to a high RI and MALS signal for the blank, the measurement of molar mass was perturbed during the retention time range 0–2.8 min. No compensation was made for the high blanks and molar mass data were therefore only presented and evaluated for tr42.8 min. For all OSA starch fractionations, molar mass data were only evaluated for an RI detector signal-to-noise ratio
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(S/N)42. Below this level, e.g. at the trailing part of the RI peaks, the molar mass is too unreliable to be reported. The retention times of OSA starch II in Fig. 4 are not directly comparable with those of Figs. 3 and 5 due to the difference in flow conditions in comparison with the ones used for OSA starches I and III. 4.2. Molar mass and size distributions The fractionations illustrated in Figs. 3–5 demonstrate the widths of the molar mass and r.m.s. radius distributions, i.e. the polydisperse character of OSA starch. For the non-homogenized OSA starches I and III the molar mass distribution widths were very similar ranging at least 3 decades (precise enough data from the fronts and tails of the distributions were not obtained) from about 0.8 106 to 2 109 g/mol for OSA starch I, and from 1 106 to 4 109 g/mol for OSA starch III (Figs. 3A and 5A). For the non-homogenized OSA starch II the molar mass distribution width covered only slightly more than one decade, i.e. from 0.8 105 to 3 106 g/mol (Fig. 4A). Likewise, the r.m.s. radius distribution widths were similar for the two non-homogenized OSA starches I and III, i.e. 40–700 and 60–800 nm, respectively (Figs. 3B and 5B). There was a drastic decrease in molar mass and r.m.s. radius upon homogenization of OSA starches I and III (Figs. 3A and 5A) but no change at all for OSA starch II (Fig. 4A). Homogenized OSA starches I and III covered an r.m.s. width of (3–35) 101 and (2–26) 101 nm, respectively (Figs. 3B and 5B). In summary, OSA starches I and III were very similar to each other in molar mass and r.m.s. radius distributions both before and after homogenization whereas OSA starch II was unaffected by homogenization, apparently because its molecular size was much smaller than the others. Indeed, the present determination of the enormous size and polydispersity of these OSA starches illustrates the excellent methodological strength of flow FFF. 4.3. Disruption and shear The results show that high-pressure homogenization of OSA starch solutions affects the molar mass distribution and the r.m.s. radius. The magnitude of the effect seems to depend on the level of initial molar mass and apparent molecular size of the polymer, where a high initial molar mass gives a large reduction in molar mass after homogenization. The resulting molar mass is in a comparable range to those reported by other authors, which are given in Table 1. This change in molar mass could both be due to the breaking of covalent bonds, i.e. intramolecular bonds, or intermolecular non-covalent interactions that would result in the disruption of aggregates. From the results in this study it is impossible to rule out any of these explanations and it is therefore reasonable to assume that both of the suggested events occur. As has been pointed out above it is interesting that the r.m.s. radius becomes
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similar for the OSA starches after homogenization. The difference in molar mass between the samples is also somewhat reduced by homogenization. This suggests that there is some limiting size for the OSA starch depending on the processing conditions as has earlier been described by other authors for other polymers (Buchholz et al., 2004; Kasaai et al., 2003; Striegel, 2003). The energy that is released during high-pressure homogenization is usually quite high and is described by the energy dissipation rate e, which can be expressed in W/kg. The energy dissipation rate depends mainly on the homogenization pressure and the volume in which the energy is dissipated. A typical value of e for high-pressure homogenizers is 109 W/kg (Walstra, 1983). During the homogenization the flow conditions will be mostly turbulent and locally it can be assumed that the turbulence is isotropic as this simplifies the description of the turbulence considerably. Isotropic turbulence has been described by Kolmogorov (1949), Levich (1962) and others. An important parameter describing the turbulence is the Kolmogorov scale (Kolmogorov, 1949), which gives the magnitude of the turbulence micro-scale: 3 1=4 n l¼ , (3) e where l is the length scale of the smallest turbulent eddies, u is the kinematic viscosity (m2/s), and e is the energy dissipation rate per unit mass (W/kg). One should, however, remember that this expression is based on scaling laws and thus only gives an estimate of the turbulence micro-scale. In order to cause disruption of a macromolecule the length scale of the turbulent eddies should be smaller than the length scale of the macromolecule, as the eddies then generate a shear field over the macromolecule. This shear field could then ultimately lead to rupture of the macromolecule. If on the other hand, the length scale of the eddies are larger than the macromolecule they would not cause any disruption. In our case, if we assume u ¼ 106 m2/s and e ¼ 109 W/kg then lE200 nm. This value is smaller than the diameter of the OSA starches I and III that were disrupted and larger than II that was not noticeably disrupted. Furthermore, it is also known that macromolecules in turbulent flow become very extended depending on branching and chain flexibility and that scission is likely to occur at the midpoint (Groisman & Steinberg, 2001; Horn & Merrill, 1984). This suggests that the OSA starch is more extended during disruption in the high-pressure homogenizer than during the subsequent analysis with FFF–MALS–RI. 4.4. Molar mass of OSA starch during emulsification In this investigation, we have shown that high-pressure homogenization reduces the molar mass of OSA starch. Thus, it is likely to assume that a similar degradation occurs during a normal emulsification process. During
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emulsification OSA starch is included in the system before the homogenization process to be able to act as an emulsifier in the same moment as the droplets are formed in the homogenizer. A discussion about the role of the molar mass and emulsification properties have rather to be based on the molar mass measured after homogenization than on the molar mass before homogenization. OSA starch typically has quite a high median molar mass, in this investigation, in the range (0.8–40) 106 g/mol. After homogenization the median molar mass still remains high, (0.6–2) 106 g/mol, but it is less extreme. Moreover, for OSA starches I and III the homogenization causes the distribution widths to decrease. 5. Conclusion AsFlFFF–MALS–RI is a unique analysis tool to study the molar mass distribution of OSA starch it being a polydisperse macromolecule with ultra-high molar mass components. High-pressure homogenization causes disruption of OSA starch and the extent of this depends on the initial size of the OSA starch and the energy dissipation rate during processing. The resulting molar mass after high-pressure homogenization is in a comparable range to the results reported by other authors for different polymers (Table 1). A comparison between our results and those given in Table 1 shows that high-pressure homogenized OSA starch is still a high molar mass hydrophobically modified polymer. Acknowledgements This study was financed by the Centre for Amphiphilic Polymers from Renewable Sources (CAP) at Lund University. The support from the Swedish Research Council is acknowledged by one of us (KGW). Lyckeby Sta¨rkelsen, Kristianstad is acknowledged for financial support and for supplying the OSA starch samples. References Aberle, T., Burchard, W., Vorwerg, W., & Radosta, S. (1994). Conformational contributions of amylose and amylopectin to the structural properties of starches from various sources. Starch/Staerke, 46, 329–335. Andersson, M., Wittgren, B., & Wahlund, K.-G. (2001). Ultrahigh molar mass component detected in ethyl hydroxyethyl cellulose by asymmetrical flow field-flow fractionation coupled to multi-angle light scattering. Analytical Chemistry, 73, 4852–4861. Andersson, M., Wittgren, B., & Wahlund, K. G. (2003). Accuracy in multiangle light scattering measurements for molar mass and radius estimations. Model calculations and experiments. Analytical Chemistry, 75, 4279–4291. Bello-Perez, L. A., Roger, P., Baud, B., & Colonna, P. (1998). Macromolecular features of starches determined by aqueous highperformance size exclusion chromatography. Journal of Cereal Science, 27, 267–278. Bergensta˚hl, B., & Claesson, P. (1997). Surface forces in emulsions. In S. E. Friberg, & K. Larsson (Eds.), Food emulsions (pp. 57–109). New York: Marcel Dekker.
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Wittgren, B., & Wahlund, K.-G. (1997b). Fast molecular mass and size characterization of polysaccharides using asymmetrical flow field-flow fractionation-multiangle light scattering. Journal of Chromatography, A, 760, 205–218. Wittgren, B., Wahlund, K.-G., Derand, H., & Wesslen, B. (1996). Aggregation behavior of an amphiphilic graft copolymer in aqueous medium studied by asymmetrical flow field-flow fractionation. Macromolecules, 29, 268–276. Wyatt, P. (1993). Light scattering and the absolute characterization of macromolecules. Analytica Chimica Acta, 272, 1–40.