2. MATERIALS AND METHODS
2.7 Storage of phenolic powder and encapsulated phenolic powder
Phenolic powders (EPP and PEPP) and encapsulated phenolic powders (prepared with 8% MD, 2% GA and EPP; 8% MD, 2% GA and PEPP) were used in the evaluation of storage stability. Samples were stored in two desiccators with different relative humidities (RH) at room temperature (21±2.0 °C). Saturated aqueous solutions of potassium carbonate and potassium chloride were used to obtain 43% and 85% RH, respectively (Greenspan, 1977). Before placing the sample, each desiccator was kept closed overnight to achieve equilibrium. Storage stability analyses were performed once in 5 days for samples stored at 85% RH and once in 10 days for samples stored at 43% RH. The parameters analyzed were total phenolic content, total antioxidant activity and hygroscopicity.
27 2.8 Physical analysis
2.8.1 Particle size analysis
Particle size analysis was performed for extracted concentrates, emulsions and encapsulated phenolic powders by the laser light scattering method using Mastersizer 2000 (Malvern Instruments, Worcestershire, UK). The mean diameter of the particles was expressed as Sauter mean diameter (D[32]) and calculated with the following formula:
𝐷[32] = 𝑛𝑖𝑑𝑖3 𝑛𝑖𝑑𝑖2 (3)
where, ni stands for the frequency of occurrence of particles in size class i, and di for mean diameter (µm) of these particles.
Span of the particle size distributions was calculated with the following formula:
𝑆𝑝𝑎𝑛 = 𝑑 𝑣,90 −𝑑(𝑣,10)
𝑑(𝑣,50) (4)
where, d(v,10), d(v,50), and d(v,90) are the diameters at 10%, 50%, and 90%
cumulative volume, respectively (Elversson et al., 2003). The instrument also reported the specific surface area (m2/g) of the particles.
2.8.2 Color analysis
Color analysis was performed for phenolic powders, capsules, crumb and crust of cakes.
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2.8.2.1 Color measurement of phenolic powder and encapsulated samples
Color determination was performed in CIE (L*, a*, b*) color space by using a UV-2450 UV-Vis spectrophotometer (Shimadzu Co., Kyoto, Japan). It was measured as reflected color from the surface of capsules or powders. Illuminant type C (2° standard observer) was used in this analysis. Mean of 3 determinations was calculated for L*
(darkness/whiteness), a* (greenness/redness) and b* (blueness/yellowness) parameters of each sample. The difference in the color (∆E*) with the reference sample (barium sulfate) was calculated for each sample using the following formula:
∆𝐸∗= ∆𝐿∗2+ ∆𝑎∗2+ ∆𝑏∗2 (5)
where, ∆L*, ∆a* and ∆b* are the differences between the color values of reference and sample.
2.8.2.2 Determination of color of crumb and crust of cakes
Color reflected from the crumb and crust of the cake was measured separately in CIE (L*, a*, b*) color space by using Minolta color reader (CR-10; Japan). The instrument was standardized each time by a white sample. Duplicate readings were performed from different position and mean value of ∆E* calculated by the instrument was recorded. Color measurement was performed only for cakes which contained sugar.
29 2.8.3 Specific volume of cake
Specific volume of cakes which contained sugar was determined by the rape seed displacement method (AACC, 1988). Density of rape seed was determined by filling a glass container with known volume (Vcontainer) and weight (Wcontainer) with seeds through tapping and smoothing the surface with ruler until the constant weight was reached.
Bulk density (ρseeds) of rape seeds was found to be 693.2 g/cm3. Then, weights of cakes (Wcake) were measured. Finally, cakes were placed in the container and the remaining part was filled with the seed. The container was taped and the surface was smoothed with a ruler. Total weight (Wtotal) was recorded when no change in consecutive weight measurements was observed. Weight of seeds (Wseeds) required for filling of the container was calculated from the following equation:
𝑊𝑠𝑒𝑒𝑑𝑠 = 𝑊𝑡𝑜𝑡𝑎𝑙 − 𝑊𝑐𝑎𝑘𝑒 − 𝑊𝑐𝑜𝑛𝑡𝑎𝑖𝑛𝑒𝑟 (6)
Volume of rape seeds (Vseeds) was calculated from the following equation:
𝑉𝑠𝑒𝑒𝑑𝑠 = 𝑊𝜌𝑠𝑒𝑒𝑑𝑠
𝑠𝑒𝑒𝑑𝑠 (7) Finally, volume of cakes was determined using equation (8):
𝑉𝑐𝑎𝑘𝑒 = 𝑉𝑐𝑜𝑛𝑡𝑎𝑖𝑛𝑒𝑟 − 𝑉𝑠𝑒𝑒𝑑𝑠 (8)
Specific volume (SVcake) was calculated from the following equation:
𝑆𝑉𝑐𝑎𝑘𝑒 =𝑊𝑉𝑐𝑎𝑘𝑒
𝑐𝑎𝑘𝑒 (9) All parameters were in the SI units.
30 2.8.4 Crumb texture
Crumb hardness (N), chewiness (N) and gumminess (N) of cakes with sugar samples were measured by using texture analyzer (TAPlus; Lloyd Instruments, Bognor Regis, UK). Fresh cakes (1 h after baking) with cubic shape (25 mm each side) without crust were compressed for 25% of original thickness by 50 N load cell equipped with cylindrical probe (10 mm in diameter) at a speed of 55 mm/min. Each measurement was duplicated and mean of a selected parameter was calculated.
2.8.5 Hygroscopicity
Hygroscopicity assay was performed in the desiccators containing saturated potassium carbonate and potassium chloride solutions having RH of 43% and 85%, respectively, according to the method proposed by Cai & Corke (2000), with some modifications.
Each sample was weighed (up to 1 g) in the aluminum plate. Hygroscopicity was determined gravimetrically after no change in the mass of samples was observed. Each test was duplicated. Before samples were placed in desiccators weights of aluminum plates (Wplate) and total weights (W0 total) were measured. When no change of total weight was observed in measurements during storage, final total weight (Wfinal total) was measured. Amount of water absorbed per g of capsule or powder was determined by the following equation:
𝐻𝑦𝑔𝑟𝑜𝑠𝑐𝑜𝑝𝑖𝑐𝑖𝑡𝑦 =𝑊𝑓𝑖𝑛𝑎𝑙 𝑡𝑜𝑡𝑎𝑙 −𝑊0 𝑡𝑜𝑡𝑎𝑙
𝑊0 𝑡𝑜𝑡𝑎𝑙 −𝑊𝑝𝑙𝑎𝑡𝑒 (10)
31 2.9 Chemical analysis
2.9.1 Reducing sugar content
It was stated in section 2.6 that high reducing sugar content in the product may interfere with the total phenol reagent. Therefore, it was required to determine the reducing sugar content of the phenolic powders. The method described in the book of Cemeroğlu (2007) with some modifications was used. Fehling-I solution was prepared by dissolving 69.3 g CuSO4.5H2O in distilled water. Then, final volume was brought to 1 liter by addition of distilled water. Fehling-II solution was prepared by dissolving 346 g of potassium sodium tartrate tertrahydrate and 100 g of KOH in distilled water.
Distilled water was added until the final volume was 1 liter. Standardization was performed by using standard glucose solution, which was prepared by dissolving 0.5 g of glucose in 100 mL of distilled water. Firstly, 10 mL of standard glucose solution was mixed with 10 mL of Fehling solution (1:1 solution of Fehling-I and Fehling-II) and heated to its boiling point. After 2 min of boiling, few drops of methylene blue indicator solution were added. After 2 min of boiling, mixture was titrated with standard glucose solution. The amount of glucose required to titrate 10 mL of Fehling solution was determined in this step of standardization.
To prepare solutions of phenolic powders, 0.5 g of EPP and 0.5 g of PEPP were dissolved separately in 10 mL of distilled water. Then, 0.5 mL of each solution was added to 9.5 mL of distilled water in order to dilute the solutions. Each solution was gently mixed. Then, 10 mL of diluted solution was added to 10 mL of Fehling solution and the same procedure described in the standardization was repeated. The amount of reducing sugar in phenolic powders was determined by reducing sugar content of added standard solution, by multiplying with dilution factor and dividing by 10 mL. Results were reported as % (w/w).
32 2.9.2 Total phenolic content
Total phenolic content (TPC) of EPP, PEPP, capsules and cakes without sugar was determined by modified Folin-Ciocalteu method (Beretta et al., 2005). Two solutions were used in this assay. First solution contained 10% (v/v) of Folin-Ciocalteu’s phenol reagent and 90% of distilled water. Second solution was composed of 7.5 % (w/v) of sodium carbonate dissolved in distilled water. Extraction of polyphenols from the samples was performed with ethanol:water:acetic acid solvent (50:42:8) (Saenz et al.
2009). One hundred milligrams of EPP, PEPP or capsule was added to 1 mL of solvent and agitated by using vortex (ZX3; VELP Scientifica, Usmate, MB, Italy). For the analysis of TPC during storage, dispersions were ultrasonicated (80 W, 50% pulse) for 2 min. Extraction of phenolic compounds from the crumb of cake was performed with some modifications. Firstly, 20 mL of solvent were added to 10 g of crumb. Then, the crumb was crushed manually using glass rod to guarantee efficient ultrasonication.
Solution containing crumb particles was then ultrasonicated in two cycles each for 1 min and with 160 W adjusted power. After the first cycle, dispersion was manually agitated. After the ultrasonication, 7.5 mL of dispersion were centrifuged at 10,000 rpm for 2 min and liquid part was carefully collected. Polyphenolic extracts of EPP, PEPP, capsules and cakes were then filtered through 0.45-µm filter. The dilution rates varied for each sample and they were applied to ensure that phenolic content of the diluted sample fits the calibration curve. Calibration curve for this assay was prepared by using gallic acid as a standard and had a range of 0-100 mg GAE/mL (APPENDIX A1, Fig.A.1).
Samples for the spectrophotometric analysis were prepared in two steps. Firstly, 2.5 mL of 10% Folin-Ciocalteu’s phenol reagent was mixed with 0.5 mL of diluted sample.
Mixture was stored for 5 min in dark. Then, 2 mL of sodium carbonate solution were added to the mixture and stored for 1 h at room temperature in dark place. Finally, samples were analyzed spectrophotometrically at 760 nm. Phenolic content (TPCstd) of
33
the diluted samples was found from the standard curve. TPC of the EPP, PEPP, capsules and cakes was found from following equation:
𝑇𝑃𝐶 = 𝑇𝑃𝐶𝑠𝑡𝑑𝑥𝑊𝑉𝑠𝑜𝑙𝑣𝑒𝑛𝑡
𝑠𝑎𝑚𝑝𝑙𝑒 𝑥 𝐷𝐹 (11)
where, Vsolvent is volume of solvent used in the extraction of phenolic compounds, DF is dilution factor and Wsample is weight of the sample.
Total phenolic content of EPP and PEPP was expressed in mg GAE/ g dry weight.
Total phenolic content loss during storage was expressed in % and was calculated by the following equation:
𝑇𝑃𝐶 𝑙𝑜𝑠𝑠 % =𝑇𝑃𝐶𝑡=0𝑇𝑃𝐶−𝑇𝑃𝐶𝑡=𝑡
𝑡=0 𝑥100 (12) where, TPCt=0 is the initial TPC and TPCt=t is TPC at any time period during storage.
Similar equation was used for the calculation of the retention of polyphenols after baking. Retention of TPC (%) was found by subtracting TPC loss (%) from 100%. Due to the presence of antioxidants in margarine and a possibility that other ingredients may affect measurement of TPC of cakes, control cake without sugar was also determined.
This amount was then subtracted from the experimental value of TPC of cakes. In this calculation initial TPC was a theoretical value of polyphenols and TPCt=t was measured TPC of cakes. Weight loss factor which was not studied in this work was taken into consideration for the calculation of theoretical TPC of cakes and was equal to 1.1.
Correction factor due to the presence of the reducing sugars in the phenolic powders was not used. Reducing sugar content of EPP and PEPP was found to be 65.3% and 63.2%, respectively. However, in the analysis of TPC, all samples were diluted and final concentration of reducing sugars in any sample was ranging from 0.52% (w/v) to
34
0.44%. Waterhouse (2002) proposed to use the correction factor for the sweet and semisweet wines having reducing sugar content >2% (w/v) sugar.
2.9.3 Surface phenolic content and encapsulation efficiency of capsules
Surface phenolic content (SPC) of capsules is an important parameter for the evaluation of encapsulation efficiency (EE%). SPC was determined by modified Folin-Ciocalteu method (Beretta et al., 2005). One hundred milligrams of capsules were agitated for 1 min with 1 mL of methanol:ethanol solvent (1:1) and then filtered through 0.45-µm filter. Calibration curve was prepared with gallic acid and ethanol:methanol solvent (APPENDIX A.1, Fig.A.2). Measurement of SPC was performed at the same conditions described in section 2.9.2. SPC was expressed as mg GAE/g of capsules.
Encapsulation efficiency was calculated from the following equation:
𝐸𝐸% =𝑇𝑃𝐶 −𝑆𝑃𝐶𝑇𝑃𝐶 𝑥100 (13)
2.9.4 Total antioxidant activity of phenolic powders, capsules and cakes
Total antioxidant activity (TAA) of capsules, phenolic powders and cakes was measured by (1,1-diphenil-2-picrylhydrazyl) DPPH method (Yen & Duh, 1994) with some modifications. The same diluted samples, described in the section 2.9.2 were used for the TAA determination. One hundred microlitters of diluted samples were added to 25 ppm DPPH solution (DPPH dissolved in methanol) and left to stand in the dark place for 1 h. After that, samples were analyzed spectrophotometrically at 517 nm.
DPPH content corresponding to each sample was determined from the calibration curve (APPENDIXT A.1, Fig.A.3). TAA was calculated from the following formula:
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𝑇𝐴𝐴 = 𝑇𝐴𝐴𝑡=0− 𝑇𝐴𝐴𝑡=1 (14)
where, TAAt=0 is initial DPPH concentration and TAAt=1 h is concentration of DPPH in the sample after 1 h.
Dilution rate, volume of solvent and weight of sample were used in the calculation of TAA of the samples. TAA was expressed as DPPH equivalent/g of sample.
During storage percentage of TAA loss was calculated from the following equation:
𝑇𝐴𝐴 𝑙𝑜𝑠𝑠 % = 𝑇𝐴𝐴𝑖𝑛𝑖𝑡𝑖𝑎𝑙 −𝑇𝐴𝐴𝑡 =𝑡
𝑇𝐴𝐴𝑖𝑛𝑖𝑡𝑖𝑎𝑙 𝑥100 (15)
where, TAAinitial is TAA of capsules before storage and TAAt=t is TAA at any time period during storage.
Similarly with TPC analysis, TAA was also determined for the control cake. The correction factor due to weight loss was equal to 1.1. Retention of TAA after baking was also calculated by subtracting TAA loss (%) (Equation 15) from 100%. TAAinitial
was theoretical TAA of cake and TAAt=t was TAA of cake. TAA of cake was calculated by subtracting TAA of control cake from experimentally determined TAA of cake.
2.10 Surface morphology of phenolic powders and capsules
The micrographs of the surface of encapsulated phenolic powders, EPP and PEPP were obtained by using scanning electron microscopy (SEM). Dry samples were coated by
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gold/palladium using Hummel VII sputter coating device (Anatech USA, Union City, CA, USA) and analyzed with JSM-6400 electron microscope (Jeol Ltd., Tokyo, Japan) operating at 20 kV. Images were taken at 100x and 500x magnification.
2.11 Sensory analysis of cakes
A 30-member panel consisting of students of Food Engineering Department of METU was used in the sensory evaluation of cake samples. Three types of cakes were analyzed by the panelists. One cake was the control cake (containing no encapsulated phenolic powder) and other two contained encapsulated phenolic powders prepared from a) 8% MD, 2% GA and EPP and b) 8% MD, 2% GA and PEPP. The panelists were asked to give scores for flavor, texture and color of cakes ranging from 1 to 5.
Each cake had a three digit code. Sensory analysis evaluation sheet and data of panelist scores are shown in the APPENDIX A.2.
2.12 Statistical analysis
All experiments if not implied in the method part were replicated twice. Data was analyzed by one-way or two-way analysis of variance (ANOVA) (P≤0.05) using MINITAB software 15 version (Minitab Inc., State College, PA, USA) and SAS software version 9.1 (SAS Institute Inc., NC, USA), respectively. Two-way ANOVA was used for storage experiment which had independent variables of powder type, composition of coating materials and RH. Dependent variables were, EE%, SPC, D[32], span, SSA, TAA, L*, a*, b*, ∆E* (of capsules, powders, crumb and crust), TAA loss (storage and baking), TPC loss (storage and baking), hygroscopicity, hardness, chewiness, gumminess, flavor, and texture.
37 CHAPTER 3
RESULTS AND DISCUSSION
3.1 Effect of degritting on physical and chemical properties of polyphenolic powders and capsules
3.1.1 Effect of degritting on particle size of extracted concentrates
In order to study the effect of centrifugation on particle size distribution of extracted polyphenolic concentrate, three different samples were analyzed (Fig 3.1). It was found that centrifugation had significant effect on particle size distribution of the samples.
Sample had larger particle size prior to centrifugation (P1). Sample spun at 10,000 rpm (P3) contained more particles with smaller size when compared to the sample spun at 5,000 rpm (P2). In other words, more particles with larger size were removed from the concentrate as centrifugation rotational speed was increased. This resulted in the shift of particle size distribution curve to the left (lower diameters) which can be explained by Stoke’s Law. General equation of Stoke’s Law (Leung, 2007) can be expressed as:
𝑉𝑠𝑜 = 𝜌𝑆−𝜌18𝜇𝐿 𝑔𝑑2 (16)
where, Vso is separation velocity, ρS density of solid, ρL density of liquid, g centrifugal acceleration, d diameter of the particle, and µ viscosity of liquid. Subscript “o” in
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separation velocity stands for separation of an individual particle with no interaction with other particles in an ideal dilute solution. In this analysis all parameters were the same for three samples except centrifugal acceleration. As angular velocity increased critical diameter of the particle to remain in the suspended form decreased.
Fig. 3.1 Effect of degritting on particle size distribution of concentrated polyphenolic extracts P1, P2 and P3.
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It can be seen from the Equation (16) that P3 had smaller particles in the suspended form when compared to P2 during centrifugation. Undoubtedly, P1 contained larger suspended particles when compared to P2 and P3, since no centrifugal acceleration was applied on it. Hence, sample spun at the higher angular velocity was comprised of particles with smaller Sauter mean diameter (D[32]) (Table 3.1; Table B.1). On the contrary, span of P3 had the highest value and it was significantly different (p≤0.001) from that of P1 and P2 samples (Table B.2). This can be explained by analyzing its particle size distribution curve (Fig 3.1). Since most of the particles of P3 are smaller this shifted the particle size distribution curve to the left (smaller size), therefore difference between d(v,90) and d(v,10) is high (Equation (4)). Definitely, this difference is lower for P2 and P1. In addition, d(v,50) is the lowest for P3. Removal of the large particles by centrifugation from P3 made it possible to detect smaller particles which could be shadowed during particle size analysis and thus, not present in the results of P1 and P2. Therefore, particle size distribution curve of P3 shifted to the left.
Table 3.1 Influence of angular velocity on purification of dispersion
* Columns having different letters (a, b & c) are significantly different (p ≤ 0.05).
Specific surface area was also significantly different (p≤0.001) for all samples (Table B.3). The reason for this is its indirect proportionality to D[32] (Table 3.1). The highest
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specific surface area was reported for P3. Consequently, it can be understood that filtration process was insufficient for the removal of particles with the diameters in the range of 1-100 µm from the polyphenolic extract, so extra treatment had to be applied.
The origin of these particles which were present in the extract can be organic or inorganic. Soluble and insoluble tissues of pomace with different size and geometry could be extracted during maceration. Drying of the pomace under the sun could lead to contamination with particles from the environment such as mineral crystals or dust suspended in the air. These particles with size under the critical diameter of suspension could remain in the extract.
3.1.2 Effect of degritting of concentrates on particle size distribution of emulsions Two different core materials (EPP and PEPP) were used to prepare emulsions. Core materials were entrapped in two different coating materials including 10% MD and combination of 8% MD and 2% GA. Results of the particle size analysis of the emulsions are given in the Table 3.2.
Table 3.2 Particle size analysis results of emulsions prepared with different coating materials and powder types
* Columns having different letters (a & b) are significantly different (p ≤ 0.05).
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Purification of polyphenolic extracted concentrates had significant (p≤0.001) influence on D[32] (Table B.4), span and specific surface area values of emulsions (Table B.5;
Table B.6). Emulsions prepared with PEPP contained smaller particles when compared to emulsions prepared with EPP. Most of these particles were in the nano range, resulting from the D[32] values of P3. Therefore, degritting was found to be a critical parameter in the preparation of nano-emulsions. Coating materials used in this study had no significant influence (p>0.05) on the particle size distribution of the emulsions.
Similarly, it can be seen on the Fig.3.2 that particle size distribution curves are very similar for emulsions prepared with different coating materials. On the contrary, they appear very different when emulsions containing PEPP were compared with emulsions containing EPP.
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Fig. 3.2 Particle size distribution of micro-emulsions (containing EPP) prepared with 10% MD (solid line) and 8% MD-2% GA (dotted line), nano-emulsions (containing PEPP) prepared with 10% MD (dashed line) and 8% MD-2% GA (dash dotted line)
Similar to extracted polyphenolic concentrates, degritting had significant (p≤0.05) effect on the span values of the emulsions. High span value of P3 resulted in the high span values of the emulsions prepared with PEPP. These emulsions contained large range of particles with different sizes. Most of these particles were in the nano range, and there was continuous and approximately equal percent volume range of large particles (Fig 3.2) which had remained in the concentrate after degritting, and were present in the PEPP. However, particle size distribution appeared more narrow (lower span values) for emulsions prepared with EPP, but still contained large range of
43
particles with different diameters. Jafari et al. (2007a) reported that homogenization in blender and ultrasonication increased span values of the sub-micron emulsions. In addition, since PEPP contained lower amount of impurities, energy density of ultrasonication of total solids in the emulsions was higher than that of the emulsions prepared with EPP, leading to more disruption and formation of smaller particles.
Gordon and Pilosof (2010) reported that ultrasonication for 10 min caused the formation of many small particles. Specific surface areas of the emulsions prepared with PEPP were higher and significantly (p≤0.001) different when compared to emulsions prepared with EPP (Table 3.2), due to the indirect proportionality to Sauter mean diameter.
3.1.3 Effect of degritting of concentrates on particle size distribution of capsules
3.1.3 Effect of degritting of concentrates on particle size distribution of capsules