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j.foodchem.2018.08.063

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Food Chemistry 272 (2019) 337–346
Contents lists available at ScienceDirect
Food Chemistry
journal homepage: www.elsevier.com/locate/foodchem
Characterization and storage stability of chlorophylls microencapsulated in
different combination of gum Arabic and maltodextrin
Yu-Ra Kanga, Yun-Kyung Leea, Young Jun Kimb, Yoon Hyuk Changa,
a
b
T
⁎
Department of Food and Nutrition, and Bionanocomposite Research Center, Kyung Hee University, Seoul 02447, South Korea
Department of Food and Biotechnology, Korea University, Sejong 30019, South Korea
A R T I C LE I N FO
A B S T R A C T
Keywords:
Chlorophyll
Microencapsulation
Spray-drying
Microcapsule
Maltodextrin
Gum Arabic
Detailed investigations on the physicochemical and structural characterization of chlorophyll loaded microcapsules and their storage stability have not previously been conducted. Therefore, our objective was to encapsulate unstable chlorophylls using different blends of gum Arabic (GA) and maltodextrin (MD) (GA-MD ratios
of 5:5, 3:7, and 0:10) by spray-drying to improve storage stability of chlorophylls. An increase in concentration
of MD in wall materials was associated with lower moisture content (0.56%), higher encapsulation efficiency
(77.19%), chlorophyll content (46.78 µg/g dry powder), degree of crystallinity, and thermal stability of microcapsules. Furthermore, FTIR, XRD, and DSC analyses confirmed inclusion of chlorophylls within microcapsules. The entrapment of chlorophylls within microcapsules enhanced their storage stability at all temperatures (4, 20, and 40 °C) for ten days; notably, microcapsules coated with MD alone showed the highest
storage stability (94.7–97.5%). In conclusion, microencapsulation of chlorophylls using MD alone was optimal
for enhancing chlorophylls’ storage stability.
1. Introduction
There is a worldwide trend to substitute synthetic colorants with
natural pigments in food products owing to consumers’ concerns about
the harmful effects of synthetic colorants. Chlorophylls are widely
distributed in green fruits and vegetables, and can be used as potential
alternatives to synthetic colorants because they have a brilliant green
color, as well as numerous biological activities. Moreover, chlorophylls
are known to possess therapeutic properties, including anti-oxidant,
anti-inflammatory, anti-bacterial, anti-carcinogenic, deodorizing, and
wound healing activities (Hosikian, Lim, Halim, & Danquah, 2010).
Despite all these health benefits, there are limitations to the commercial-scale application of chlorophylls as natural colorants because they
are very susceptible to environmental stresses, such as oxygen, enzymes, light, high temperature, and acidic or alkaline pH, which result
in degradation and discoloration of chlorophylls (Schoefs, 2002;
Marquez & Sinnecker, 2008).
In recent years, the interest in microencapsulation of susceptible
compounds in a stable wall matrix as a means of protecting functional
compounds from environmental conditions (like oxygen, pH, ionic
strength, and temperature) and improving the bioavailability has increased (Pourashouri et al., 2014). Microcapsules are typically comprised of core (active) and wall (carrier) materials, and there are
⁎
various microencapsulation techniques, such as coacervation, fluidization, lyophilization, extrusion, and spray-drying. Among these techniques, spray-drying is regarded as an economical and effective process in
the food industry and, practically, it is the most commonly used technique used for microencapsulation of natural components (Edris,
Kalemba, Adamiec, & Piątkowski, 2016; Wilkowska, Czyżowska,
Ambroziak, & Adamiec, 2017).
The choice of suitable biopolymers as wall materials is critical to the
success of microencapsulation by spray-drying because the type of wall
material determines the physicochemical and morphological properties
of the produced microcapsules. Moreover, it affects the encapsulation
efficiency, shelf-life, and degree of protection of sensitive core materials. In previous studies, different types of wall materials have been
studied for microencapsulation, including polysaccharides (modified
and hydrolyzed starches, cellulose derivatives, and gums), proteins
(caseinates, whey proteins, and gelatins), and lipids (mono- and diglycerides and stearic acids) (Mahdavi, Jafari, Assadpoor, & Dehnad,
2016). Among the numerous biopolymers, maltodextrin (MD), with
different dextrose equivalents, and gum Arabic (GA) are the most
popular and common wall materials for encapsulating active compounds. GA, the exudate polysaccharides from acacia, has been used as
a wall material for spray-drying microencapsulation due to its ability to
produce a low viscosity solution at high concentrations compared to
Corresponding author.
E-mail address: yhchang@khu.ac.kr (Y.H. Chang).
https://doi.org/10.1016/j.foodchem.2018.08.063
Received 27 March 2018; Received in revised form 6 August 2018; Accepted 16 August 2018
Available online 17 August 2018
0308-8146/ © 2018 Elsevier Ltd. All rights reserved.
Food Chemistry 272 (2019) 337–346
Y.-R. Kang et al.
(GA; Samchun Pure Chemical Co., Seoul, Korea), and Tween® 80
(polyoxyethylene sorbitan monooleate, Junsei Pure Chemical Co.,
Tokyo, Japan) were used as the wall materials and emulsifier, respectively. Potassium bromide (KBr; anhydrous) and petroleum ether were
purchased from Sigma Chemical Co. (St. Louis, MO. USA) and Junsei
Pure Chemical Co., respectively. Isooctane, isopropyl alcohol (isopropanol), ethyl alcohol (95%, denatured), and acetone were obtained
from Daejung Chemicals Co. (Siheung, Korea). All reagents used in the
present study were of analytical grade.
other gums, form a very stable emulsion, and retain volatiles
(McNamee, O'Riorda, & O'Sullivan, 2001; Hosseini, Jafari, Mirzaei,
Asghari, & Akhavan, 2015). However, in recent years, the application of
GA as a wall material has been curtailed by its high price and impurities
(Jafari, Assadpoor, He, & Bhandari, 2008). Thus, researchers are trying
to use GA in combination with other wall materials or completely replace it with suitable and novel biopolymers. On this basis, starch and
its derivatives, such as MD, have been proven to be adequate biopolymers for use with GA. In general, the partially hydrolyzed starch, MD, is
utilized as a secondary wall material in spray-drying microencapsulation, offering several advantages such as comparatively low cost, high
levels of hygroscopicity and solubility, low viscosity at high solid concentrations, film forming ability, and, finally, good protection against
oxidation (Janiszewska-Turak et al., 2017; Moser et al., 2017). However, the greatest limitation of MD is its low emulsifying capacity, and it
is thus generally mixed with other wall materials that can form stable
emulsions, such as GA.
Some effort has been made to produce microcapsules for chlorophyll and its derivatives by means of spray-drying and to determine
the optimal preparation conditions. Specifically, Porrarud & Pranee
(2010) noted that the use of OSA-modified starch as a carrier for Znpheophytins resulted in the highest chlorophyll content and the longest
half-life, when compared to the use of GA and MD. In the case of
chlorophyllide, powders produced using GA and MD were more adequate for protecting chlorophyllide from environmental conditions
during storage compared to those prepared with soybean protein isolate
(Comunian et al., 2011). Recently, the microencapsulation of green
pigment extracted from alfalfa by freeze-drying and its application in
heated food (jelly and gummy candy) were evaluated by Raei, Yasini
Ardakani, & Daneshi (2017). In that study, the authors used a blend of
agar and gelatin as wall materials and reported the optimized formula
for making microcapsules with the highest encapsulation efficiency.
Furthermore, they reported that during heating, the microencapsulated
green pigment showed a lower degree of change in the color factors (L*,
a*, and b*) than the non-encapsulated green pigment.
However, as far as the present authors are aware, there are no
documented detailed investigations on the physicochemical and structural characterization of microcapsules entrapping natural chlorophylls
and their storage stability. In our previous study (Kang, Park, Jung, &
Chang, 2018), we prepared semi-purified chlorophylls from spinach
and analyzed their structure by Fourier transform infrared (FTIR) and
nuclear magnetic resonance (NMR) spectroscopy to verify the existence
of chlorophylls. Because extracted chlorophylls are easily degraded
during storage and processing, we postulated that microencapsulation
of chlorophylls in a suitable matrix could provide good protection
against adverse environmental conditions. Therefore, the present study
was aimed at the development of chlorophyll loaded microcapsules
using different blends of GA and MD as protective carriers by spraydrying. To characterize and compare the obtained microcapsules, we
evaluated the moisture content, percentage encapsulation efficiency,
chlorophyll content, particle size distribution, visual color value, and
surface properties of chlorophyll loaded microcapsules. Moreover,
characterization of microcapsules was carried out by FTIR spectroscopy, X-ray diffraction (XRD), and differential scanning calorimetry
(DSC), and the storage stability of these species at different storage
temperatures was evaluated.
2.2. Preparation of emulsions
GA and MD, as wall materials, were mixed according to the compositions listed in Table S1 (Supplementary data) and dissolved in
distilled water (30%, w/v) while stirring at 60 °C for 3 h. The solutions
were then cooled to room temperature (23 ± 1 °C) and kept overnight
in a refrigerator at 4 ± 0.5 °C to ensure complete hydration of the
polymer molecules. After hydration, 1.5% (w/v) of the emulsifier,
Tween® 80, was added to the hydrated wall material solutions and
completely dissolved with vigorous stirring at room temperature. The
core material was prepared by dispersing chlorophylls in MCT oil in the
dark to a final concentration of 0.01 g/mL. The oil-in-water (O/W)
emulsions were produced by blending the wall material solution and
core material in a 2:1 (v/v) ratio using a DIAX 600 homogenizer
(Heidolph, Kelheim, Germany) at 20,500 rpm for 3 min.
2.3. Preparation of microcapsules by spray-drying
The freshly prepared emulsions were spray-dried using a laboratory
scale SD-1000 spray-dryer (Eyela, Tokyo, Japan) under the following
operational conditions: inlet air temperature: 145 °C, outlet air temperature: 95 °C, rotary atomizer: 10 × 10 kPa, blower rate: 0.60 m3/
min, and pump speed: 1.50 mL/min. The obtained microcapsules were
stored in sealed conical tubes at −20 °C for further analysis.
2.4. Characterization of emulsions
2.4.1. Emulsion stability
The emulsion stability index (ESI) of each sample of liquid emulsion
was evaluated by the volumetric method (Chang, Shin, & Lee, 1994).
An aliquot of 10 mL of each emulsion was transferred to a measuring
cylinder and kept at room temperature for 24 h. The volume of the
separated water layer in the bottom was measured after 24 h, and the
ESI within a possible range from 0 to 1 was calculated as follows:
ESI = {1−(Vs/Va)} × 100
(1)
where Va represents the volume of added water in the emulsion and Vs
is the volume of the separated bottom layer after the desired storage
period. A value of 0 indicates poor emulsion stability, whereas a value
of 1 indicates high emulsion stability.
2.5. Characterization of microcapsules obtained by spray-drying
2.5.1. Moisture content
The moisture content of each microcapsule was evaluated using the
AOAC method (AOAC 2005). One gram of each microcapsule was
placed on an aluminium pan and dried at 105 °C for 1 h in a drying
oven. For calculation of the moisture content of each microcapsule, the
following equation was used:
2. Material and methods
2.1. Materials
Moisture content(%) = (Wet powder weight - Dried powder weight)
Freeze-dried chlorophylls, as a core material, were prepared according to our previous study (Kang et al., 2018) and kept at −80 °C
until use. Bergabest MCT oil 60/40 (Sternchemie GmbH & Co. KG,
Germany) was used as a solvent to dissolve the chlorophylls. Maltodextrin (MD; DE = 14–20) (Daesang Co., Seoul, Korea), gum Arabic
/Wet powder weight × 100
(2)
2.5.2. Microencapsulation efficiency
The microencapsulation efficiency is defined as the ratio of the core
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Y.-R. Kang et al.
Chlorophyll b (μg/mL) = 20.13 A 646−4.19 A 662
material in microcapsules to that in the original emulsion and was
calculated using the following equation, as presented elsewhere (Ahn
et al., 2008; Omar, Shan, Zou, Song, & Wang, 2009):
where A646 is the absorbance at 646 nm and A662 is the absorbance at
662 nm. Total chlorophyll content was determined by adding content of
chlorophyll a and b, and expressed as µg/g dry powder.
Microencapsulation efficiency (%) = {(Total oil - surface oil)/Total oil}
× 100
(3)
2.5.4. Particle size distribution
The particle size distribution was analyzed using a Malvern
Mastersizer 2000 (Malvern Instruments Ltd., Worcestershire, UK). A
small amount of microcapsules was dispersed in distilled water and the
refractive index was set at 1.330. The particle distribution was monitored during three successive readings. The particle sizes of all microcapsules were expressed as the volume weighted mean diameter
(D43) and span, which are the mean diameter of a sphere with the same
volume and the width of the particle size distribution, respectively.
Span = (d0.9 − d0.1)/d0.5, where d0.1, d0.5, and d0.9 are the diameters at
10%, 50%, and 90% cumulative volume of the particles, respectively.
2.5.2.1. Total oil. The total oil, including both the encapsulated oil and
surface oil, was analyzed using the method described by QuispeCondori, Saldaña, & Temelli (2011) with minor modifications. An
aliquot of 200 mg of each microcapsule was suspended in 10 mL of
85% ethanol. The suspension was mixed gently and microcapsules were
then rinsed off from the tube with 2 × 5 mL of 85% ethanol solution.
The suspension was placed in a Branson 8510 ultrasonication bath
(Ultrasonic Corporation, Danbury, USA) at room temperature for
15 min to destroy the microcapsule membrane. After sonication, the
oil was extracted five times using a total volume of 50 mL of petroleum
ether as follows: each suspension was mixed vigorously with 10 mL of
petroleum ether. After 10 min of contact, phase separation was
observed and the supernatant containing the extracted oil was
transferred to a dried round bottom flask. The above process was
repeated five times. A round bottom flask containing the supernatant
was subsequently placed in a rotary evaporator to remove the organic
solvent under vacuum conditions. After evaporation, round bottom
flask was dried in an oven at 63 °C until constant weight was achieved
(approximately 3 h). The weight of total oil was calculated based on the
difference between the initial weight of round bottom flask and that
containing the extracted oil residue.
2.5.5. Visual color values
The color values of each microcapsule were measured with a
Minolta Chroma Meter CR-400 (Konica Minolta, Tokyo, Japan) using a
D65 illuminant and an observation angle of 2°; the instrument was
calibrated with a standard white tile. The measurements were expressed as the Hunter color values: L*, a*, and b*. L* represents the
lightness, a* redness (+) and greenness (−), and b* yellowness (+) and
blueness (−). The chroma (C*), indicating the color intensity and hue
angle (H*), were also calculated using the following equations:
2.5.2.2. Surface oil. The surface oil, i.e., the non-encapsulated oil, was
analyzed according to the method of Shen & Quek (2014) with minor
modifications. A 200 mg of each microcapsule was accurately weighed,
added to 15 mL of petroleum ether, and then shaken on an NB-101M
orbital shaker (N-Biotek, Bucheon, South Korea) for 10 min at 150 rpm
to extract the surface oil. Microcapsules and the solvent were separated
by filtration through a No. 5C filter paper (Advantec, Tokyo, Japan).
Microcapsule residue was washed three times using a total volume of
15 mL of petroleum ether, and the filtrate containing the extracted oil
was transferred to a dried round bottom flask. A round bottom flask
containing the filtrate was subsequently placed in a rotary evaporator
to remove the organic solvent under vacuum conditions. After
evaporation, round bottom flask was dried in an oven at 63 °C until
constant weight was achieved (approximately 3 h). The weight of
surface oil was calculated based on the difference between the initial
weight of round bottom flask and that containing the extracted oil
residue.
Chroma (C∗) = [a∗2 + b∗2]1/2
(6)
Hue angle (H∗) = tan−1 (b∗/a∗)
(7)
*
The hue angle (H ) value vary from 0° (pure red color), thru 90°
(pure yellow color), 180° (pure green color), to 270° (pure blue color).
Negative values of H* were changed to positive values by adding 180°
so that they could fall in the 90–180° quadrant. The values for microcapsules were measured in triplicate, and the average of the measurements was used for subsequent statistical analysis.
2.5.6. Particle morphology
The morphology of each microcapsule was studied using a LEOSUPRA 55 field emission scanning electron microscope (FE-SEM; Carl
Zeiss, Jena, Germany) operating at 10 kV. Each microcapsule was
placed on FE-SEM stubs using a double-sided adhesive tape and analyzed after Pt sputtering using a Q150RS sputter coater (Quorum, East
Sussex, UK). Representative micrographs were taken for each microcapsule at magnifications of 5000× and 20,000×.
2.5.7. Fourier transform infrared spectroscopy
Fourier transform infrared (FTIR) spectra of chlorophylls, GA, MD,
and microcapsules were acquired using a Perkin-Elmer Spectrum GX
FTIR spectrometer (Perkin-Elmer, Beaconsfield, UK) equipped with a
deuterated triglycine sulfate (DTGS) detector. The microcapsules were
thoroughly mixed with KBr in a ratio of 1 mg microcapsules to 100 mg
KBr and then pressed into a KBr pellet. The pellet was scanned at room
temperature in the spectral range of 4000–400 cm−1. To improve the
signal to noise ratio for each spectrum, 128 interferograms with a
spectral resolution of 4.0 cm−1 were averaged. Background spectra,
which were collected under identical conditions, were subtracted from
the sample spectra automatically.
2.5.3. Chlorophyll content
Chlorophyll content in each microcapsule was determined using the
method explained by Lee, Ganesan, Baharin, & Kwak (2015) with some
modifications. A 0.2 g of each microcapsule was dissolved in 5 mL of
distilled water, followed by vigorous vortexing for 2 min. Thereafter,
40 mL of isooctane:isopropanol solution (3:1, v/v) was added and
mixed vigorously on a vortex mixer for 1 min. The mixtures were
centrifuged at 1000 rpm for 10 min and the supernatants containing the
extracted core materials were transferred to round bottom flasks. The
above process was repeated twice and the supernatants collected in
round bottom flasks were evaporated by using an NN series rotary
evaporator (Eyela, Tokyo, Japan) under vacuum conditions. The extracted core materials containing chlorophylls were dissolved in 2 mL of
acetone. The absorbances at 646 and 664 nm were measured, and the
chlorophyll content in each microcapsule was calculated using the
following equation (Madeira, Ferreira, de Varennes, & Vieira, 2003):
Chlorophyll a (μg/mL) = 11.24 A 662−2.04 A 646
(5)
2.5.8. X-ray diffraction
The crystallinity of chlorophylls, GA, MD, and microcapsules was
evaluated by using powder X-ray diffraction (XRD). An X-ray powder
diffraction instrument, PANalytical X'Pert PRO X-ray diffractometer
(PANalytical, Almeo, Netherlands), was used for phase analysis by
employing Cu-Kα radiation (λ = 1.5406 Å) and a step size of 0.02°. The
(4)
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Y.-R. Kang et al.
Table 1
Emulsion stability index (ESI) of emulsions and moisture content, surface oil, total oil content, microencapsulation efficiency, and chlorophyll content of microcapsules prepared using different blends of wall materials.1)
Samples
GA:MD (5:5)2)
GA:MD (3:7)
GA:MD (0:10)
Emulsions
Microcapsules
ESI
Moisture content (%)
Surface oil (g/g dry
powder)
Total oil (g/g dry
powder)
Encapsulation efficiency
(%)
Chlorophyll contents (ug/g dry
powder)
0.97 ± 0.00b
0.97 ± 0.00b
1.00 ± 0.00a
2.34 ± 0.18a
1.21 ± 0.10b
0.56 ± 0.10c
0.28 ± 0.03a
0.17 ± 0.01b
0.14 ± 0.01b
0.676 ± 0.01a
0.666 ± 0.03a
0.660 ± 0.06a
62.32 ± 0.39c
72.91 ± 1.15b
77.19 ± 1.96a
34.77 ± 0.12c
35.70 ± 0.03b
46.78 ± 0.14a
1)
Each value represents the mean of triplicate experiments ± standard deviation. Mean values in a row with different letters are significantly different
(p < 0.05).
2)
GA and MD indicate gum Arabic and maltodextrin, respectively. GA:MD (5:5), GA:MD (3:7), and GA:MD (0:10) indicate chlorophyll loaded microcapsules
prepared using GA-MD ratios of 5:5, 3:7, and 0:10.
determine significant differences amongst the samples. The means were
compared by using Fisher’s least significant difference (LSD) procedure.
All experiments were replicated at least three times for each treatment,
and the mean ± standard deviation was reported.
samples were analyzed at 2θ angles from 5° to 50°.
2.5.9. Differential scanning calorimetry
Differential scanning calorimetry (DSC) analyses were carried out
for chlorophylls, GA, MD, and microcapsules using a DSC Q2000 (TA
Instruments Inc., New Castle, Delaware, USA) instrument calibrated
with indium. Each sample (20 mg) was placed onto a standard aluminum pan and heated from 25 to 350 °C at a heating rate of 10 °C/min
under constant nitrogen purging at a flow rate of 50 mL/min. An empty
sealed aluminum pan was used as a reference.
3. Results and discussion
3.1. Moisture content
The moisture content is a critical microcapsule property, and is
associated with the stickiness, flowability, water activity, drying efficiency, microbial growth, and oxidation of bioactive agents.
Furthermore, the moisture content of microcapsule can affect the storage stability because the wall material changes from the glassy state to
an amorphous rubbery state at higher moisture levels, resulting in release and degradation of core material during storage (Velasco,
Dobarganes, & Márquez-Ruiz, 2003).
The moisture content of spray-dried microcapsules prepared using
different blends of wall materials (GA or MD) are listed in Table 1. The
present study revealed that the type of wall material influenced
moisture content; the moisture content of microcapsules significantly
decreased with an increase in MD concentrations. These findings are
consistent with those obtained by Premi & Sharma (2017), who reported a relationship between higher amounts of MD in wall material
and a decrease in the moisture content of particles.
The results of the present study can be explained as follows. Firstly,
the difference in the moisture content observed for the different samples is associated with the chemical structures of GA and MD. In comparison with MD, GA has a higher number of hydrophilic groups that
can bind water molecules, and thus, an increasing percentage of GA
decreases the moisture content of particles (Tonon, Brabet, Pallet, Brat,
& Hubinger, 2009; Mohd Nawi, Muhamad, & Mohd Marsin, 2015).
Secondly, according to Premi & Sharma (2017), a rise in the proportion
of GA in wall matrix resulted in higher viscosity of the emulsion, which
then slowed diffusion of the water molecules during spray-drying. Thus,
greater amounts of GA led to an increase in the moisture content of
microcapsules. Thirdly, the presence of surface oil can lower the evaporation of moisture by acting as a vapor obstruction (Calderón-Oliver,
Pedroza-Islas, Escalona-Buendía, Pedraza-Chaverri, & Ponce-Alquicira,
2017). Microcapsules coated with GA-MD in a ratio of 5:5 had the
highest amount of surface oil among all microcapsules, as evidenced in
Table 1 and Fig. 1, and this may contribute to the greater amount of
moisture in these particles.
Consequently, the higher moisture content of microcapsule coated
with GA-MD in a ratio of 5:5 may be due to the higher levels and higher
water binding ability of GA in the feed emulsion and the presence of
greater quantities of surface oil on spray-dried particles.
2.5.10. Storage stability at different temperatures
To determine the storage stability of encapsulated chlorophylls over
storage time at different temperatures, samples of each microcapsule
(0.1 g) were transferred to glass vials. The glass vials were then hermetically capped and stored at various temperatures (4, 20, 40 °C) in
the MIR-153 incubator (Sanyo, Gunma, Japan) for a period of 10 days
in the dark. The samples were withdrawn at two day intervals to
monitor the degradation of encapsulated chlorophylls. The withdrawn
samples were dispersed into 2.5 mL distilled water, followed by vigorous vortexing for 2 min. Thereafter, 20 mL of isooctane:isopropanol
solution (3:1, v/v) was added and mixed vigorously for 1 min using a
vortex mixer. The samples were centrifuged at 1000 rpm for 10 min,
and the organic solvent phase containing extracted core materials was
collected in round bottom flasks. The above process was repeated twice
and the organic solvent was evaporated using rotary vacuum evaporator. The remaining core materials were dissolved in 2 mL of
acetone for the subsequent spectrophotometric assay for estimation of
chlorophyll content (Eq. (4), Eq. (5)). In all cases, non-encapsulated
chlorophylls (control samples; chlorophylls in MCT oil to a final concentration of 0.01 g/mL) were also stored and analyzed under the same
conditions. For control samples, the chlorophyll content was evaluated
by dissolving them in 2 mL of acetone and measuring absorbance at 646
and 664 nm (Eq. (4), Eq. (5)). The retention percentage, which was
defined as the ratio between the content of chlorophylls that retained in
samples after selected times for 10 days and the original content of
chlorophylls in samples, was used to evaluate the stability of encapsulated chlorophylls and non-encapsulated chlorophylls.
Storage stability(%)
= (Chlorophyll content remaining in control samples or microcapsules)
÷ (Initial chlorophyll content in control samples or microcapsules)
× 100
(8)
2.6. Statistical analysis
All statistical analyses were conducted by using SAS version 9.3
(SAS Institute Inc., Cary, NC, USA). Analysis of variance (ANOVA) was
performed using the general linear models (GLM) procedure in order to
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Y.-R. Kang et al.
Fig. 1. FE-SEM micrographs of different microcapsules of chlorophylls obtained after spray drying with the following wall materials: GA-MD at ratios of 5:5 (A); 3:7
(B); 0:10 (C), magnification 5000× and 20000×. The bold arrows indicate the surface oil for microcapsules coated with GA-MD in ratio of 5:5.
correlated with the emulsion stability index (ESI). The emulsion stability was evaluated using a volumetric method after preparing chlorophyll loaded O/W emulsions. Among the emulsions, emulsion prepared using a GA-MD ratio of 0:10 exhibited the highest ESI value of
1.00, as indicated in Table 1. It is known that GA has good emulsifying
ability and is usually used in the food industry as an emulsifier. However, MD had no capacity for emulsification due to the lack of a surface
binding ability at the oil-water interface. Therefore, it was expected
that the emulsions containing higher amounts of GA should exhibit a
higher ESI. However, the results obtained did not conform to these
expectations. This can be explained as follows. In the emulsion systems
containing Tween® 80 and MD, Tween® 80 can be bound to MD by
inserting the non-polar tails into a helical coil built from the MD chain,
thereby imparting emulsifying ability to MD (Klinkesorn, Sophanodora,
Chinachoti, & McClements, 2004). In the case of GA, Tween® 20 can
detach proteins at the oil droplet surface; thus, the interaction between
the proteins and oil droplet surface was weakened (Courthaudon,
Dickinson, Matsumura, & Clark, 1991). According to Matsumura,
Satake, Egami, & Mori (2000), addition of Tween® 20 reduced the
emulsifying ability of GA, because the peptide moieties of GA were
detached from the oil droplet surface.
3.2. Microencapsulation efficiency and chlorophyll content
The encapsulation efficiency reflects the degree of retention and
protection of core materials embedded within the wall materials. The
surface oil, total oil, encapsulation efficiency, and chlorophyll content
of the microcapsules prepared using GA-MD ratios of 5:5, 3:7, and 0:10
are presented in Table 1.
The microcapsules coated with GA-MD in a ratio of 5:5 showed the
highest surface oil content, followed by those coated with GA-MD in
ratios of 3:7 and 0:10. The surface oil content is an indicator of nonencapsulated oil and has been used as a crucial parameter for determining the quality of microcapsules because the non-encapsulated
oil is prone to oxidation, resulting in off-flavors. The total oil, including
both encapsulated oil and surface oil, for all microcapsules varied, and
no significant differences were observed. The chlorophyll content and
encapsulation efficiency of microcapsules significantly increased with
an increase in MD concentration in wall materials. The greatest amount
of encapsulated chlorophylls in microcapsules coated with MD alone
might be due to the thermal protective ability of MD which was used as
a thermoprotectant during spray-drying and storage.
According to Binsi et al. (2017), the encapsulation efficiency is
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Y.-R. Kang et al.
Table 2
Mean diameter (D43), size distribution (Span), and color values of chlorophyll loaded microcapsules prepared using different blends of wall materials.1)
Samples
GA:MD (5:5)2)
GA:MD (3:7)
GA:MD (0:10)
Particle size distribution
Hunter color values
D43 (um)
Span
L*
a*
b*
C*
H*
9.72 ± 0.12b
7.71 ± 0.02c
11.44 ± 0.04a
2.64 ± 0.00c
2.93 ± 0.00a
2.85 ± 0.01b
21.55 ± 0.01c
25.41 ± 0.00b
26.22 ± 0.01a
−12.10 ± 0.01a
−17.40 ± 0.01c
−16.72 ± 0.02b
13.39 ± 0.01c
20.08 ± 0.01a
19.64 ± 0.00b
18.17 ± 0.01c
26.57 ± 0.04a
25.78 ± 0.01b
128.32 ± 0.04c
130.32 ± 0.03b
131.43 ± 0.04a
1)
Each value represents the mean of triplicate experiments ± standard deviation. Mean values in a row with different letters are significantly different
(p < 0.05).
2)
GA and MD indicate gum Arabic and maltodextrin, respectively. GA:MD (5:5), GA:MD (3:7), and GA:MD (0:10) indicate chlorophyll loaded microcapsules
prepared using GA-MD ratios of 5:5, 3:7, and 0:10.
GA-MD in a ratio of 5:5. The H* values of microcapsules were significantly decreased with a decrease in MD concentration, where a
lower H* value indicates a reduction in the greenness. According to
Tuyen et al. (2010), spray-drying at higher temperature caused a significant loss of red color of Gac fruit owing to thermal destruction of
carotenoid pigments. Therefore, considering the lower L*, −a*, C*, and
H* values for microcapsule covered with GA-MD in a ratio of 5:5, the
chlorophylls in these microcapsule may undergo more thermal degradation than in the others (GA-MD ratios of 3:7 and 0:10) during
spray-drying. These results are consistent with the chlorophyll content
of the respective microcapsules.
Therefore, the higher ESI value for a GA-MD ratio of 0:10 is attributed to the emulsifying ability of MD, which was improved by the
interaction with Tween® 80. Additionally, the greater emulsion stability
obtained for microcapsule coated with a GA-MD ratio of 0:10 may be
associated with its higher encapsulation efficiency.
3.3. Particle size distribution
Fig. S1 (Supplementary data) and Table 2 indicate the particle size
distribution and volume weighted mean diameters (D43) of chlorophyll
loaded microcapsules prepared using different combinations of GA and
MD as wall materials. All microcapsules exhibited a unimodal distribution with span values in the range of approximately 2.64–2.93 and
D43 values ranging from 7.71 to 11.44 µm. In the case of D43, McNamee
et al. (2001) synthesized soy oil loaded microcapsules by spray-drying,
where microcapsules were comprised of different blends of GA, MD (DE
5.5-38), glucose, sucrose, and lactose. They reported that microcapsules
composed of 50% GA and 50% MD (DE 18.5, 28, and 38) as wall materials presented the D43 values in the range of 10.8–15.5 µm. Carneiro,
Tonon, Grosso, & Hubinger (2013) also produced flaxseed oil encapsulated microcapsules using different combinations of wall materials and found a higher value of D43 (approximately 23.03 µm) for
microcapsules prepared using a blend of GA and MD compared to those
of the present samples. The smaller size of the present microcapsules
may be associated with the spray-drying conditions, such as the slower
feed flow rate than that used in foregoing studies. These results are
especially interesting in the case of spray-dried microcapsules, because
the smaller microcapsules can penetrate into the spaces between the
bigger ones, thus occupying less space and decreasing the powder volume. These characteristics are suitable for transportation and packaging purposes.
3.5. Structural characteristics
3.5.1. Particle morphology
Fig. 1 shows the FE-SEM micrographs of microcapsules prepared
using different blends of wall materials (GA and MD) containing
chlorophylls. Most of the microcapsules exhibited a smooth and spherical shape, but some of the microcapsules were observed to have
wrinkles and dimples. According to Comunian et al. (2011), the occurrence of wrinkles and dimples on the surface is due to rapid water
evaporation during spray-drying. Agglomerates were observed for all
microcapsules but comparatively more were observed for those covered
with GA-MD in a ratio of 5:5. The large amount of adhering microcapsules could be attributed to the presence of higher quantities of
surface oil on the microcapsules with GA-MD ratio of 5:5. Because of
the presence of surface oil, the surface characteristic of microcapsules
covered with GA-MD in a ratio of 5:5 could not be accurately observed.
Similar to the present study, Mahdavi et al. (2016) elucidated that
anthocyanin loaded microcapsules obtained from GA-MD as wall materials were smooth but not uniform, showing dents on the surface and
slight agglomeration
3.4. Visual color values
3.5.2. Analysis of Fourier transform infrared spectra
FTIR spectroscopic analysis was performed (Fig. 2A) to understand
the characteristics of the intermolecular interactions between the wall
and core materials.
The spectrum of GA (Fig. 2A-a) presented absorption bands at 3413
(OeH stretching), 2930 (CeH stretching), 1613 (C]O stretching and
NeH bending), 1141, 1070 (C-O stretching), 840, and 773 (OeH deformation vibrations), which agree with previous studies (Williams,
Gold, Holoman, Ehrman, & Wilson, 2006; Banerjee & Chen, 2007;
Tiwari, 2007; Ursescu, Măluţan, & Ciovică, 2009).
The MD spectrum (Fig. 2A–b) showed absorption bands at 3392
(OeH stretching), 2925 (CeH stretching), 1653 (C]O stretching),
1457 (CH2 bending), 1371 (OeH bending), 1158, 1081, 1020 (CeO
stretching and CeOeH bending), 928, 848, 761, 708, and 576 (skeletal
vibrations of the pyranoid ring), which agree with the results reported
by Smrčková et al. (2013), Tabani, Mahyari, Sahragard, Fakhari, &
Shaabani (2015), and Krishnaiah, Sarbatly, & Nithyanandam (2012).
The characteristic bands of chlorophylls appeared at 3455 (OeH
stretching), 2927 (CeH stretching in phytol), 2855 (asymmetric and
The variations of the color values of microcapsules are related to
differences in the structures of microcapsules and the amounts of core
and wall materials. Table 2 illustrates the effects of the different wall
materials on the color characteristics (L*, a*, b*, C*, and H*) of spraydried microcapsules. The L* values of microcapsules were significantly
increased with an increase in MD concentration. Generally, an increase
in the L* value of microcapsules was observed with higher MD and
lower GA concentrations due to the white color of MD (Tuyen, Nguyen,
& Roach, 2010). Comparable results were also obtained for purple
sweet potato and tamarind pulp extract loaded microcapsules produced
by spray-drying (Ahmed, Akter, Lee, & Eun, 2010; Cynthia, Bosco, &
Bhol, 2015).
For the color parameter a*, all the microcapsules showed negative
values, indicating the greenness of all samples. A relatively higher value
of −a* was obtained for the microcapsules coated with GA-MD in ratios
of 3:7 and 0:10, indicating more greenness than that of microcapsule
covered with GA-MD in a ratio of 5:5.
The lowest C* value was observed for microcapsule covered with
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Fig. 2. FTIR (A) and XRD (B) spectra of chlorophylls (Chls), GA (a), MD (b), and microcapsules prepared using GA-MD ratios of 5:5 (c), 3:7 (d), and 0:10 (e).
Reference is made to the FTIR spectrum of chls from our previous study (Kang et al., 2018) and the bold arrows in that spectrum indicate the characteristic peaks of
chlorophylls.
symmetric CH2 and CH3 stretching), 1745 (C-173]O and C-133]O
stretching), 1694 (C-131]O stretching), 1607 (skeletal C]C and C]N
stretching of aromatic system in chlorophyll), and 1286 (C-173-O and
C-133-O stretching) (Fig. 2A-chls). These results are consistent with
those obtained in our previous study (Kang et al., 2018).
For all microcapsules, the characteristic hydroxyl peaks (OeH
stretching) were observed around 3400 cm−1. Compared to the peak
intensities for GA and MD, the intensities of the hydroxyl peaks for all
microcapsules considerably decreased. This implies that the hydroxyl
groups in GA and MD participated in chemical reactions during spraydrying, such as hydrogen bonding and/or esterification between MD
and GA. A peak corresponding to the amine or carbonyl group of GA
(1613 cm−1) appeared in the spectra of microcapsules coated with GAMD in ratios of 5:5 and 3:7, but was not apparent in that of
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confirmed.
Furthermore, the intensity of the XRD peaks for microcapsules
progressively increased with increasing MD content in chlorophyll
loaded microcapsules, where a relatively higher intensity was observed
for microcapsule prepared using MD alone than the other microcapsules. It is well known that the increased sharpness of the diffraction
peaks in XRD profiles indicates an increased degree of crystallinity of
the sample. Accordingly, it was demonstrated that microcapsules
coated with GA-MD in a ratio of 0:10 had more crystalline characteristics than the other microcapsules due to the higher content of MD in
the wall materials of the former.
The crystalline and amorphous state are also related to the physicochemical properties and storage stability of powders. In general,
amorphous materials are more soluble and hygroscopic than crystalline
materials, resulting in water absorption during storage (Nambiar,
Sellamuthu, & Perumal, 2017). Furthermore, it is known that the water
absorption of materials is related to weight gain, collapse of the microstructure, degradation of nutrient, and potential microbiological
instability during storage (Borrmann, Pierucci, Leite, & da Rocha Leão,
2013). Therefore, the XRD results imply that microcapsule prepared
using MD alone may have higher storage stability (Fig. 4) because of its
higher degree of crystallinity compared to the other microcapsules.
microcapsule prepared using MD alone. Furthermore, the spectra of all
microcapsules (Fig. 2A-c, d, e) showed new peaks at 2927, 2855, and
1745 cm−1, which were not detected in the spectra of GA and MD.
Based on the spectra of chlorophylls, the aforementioned peaks indicate
the presence of chlorophylls in all microcapsules.
Consequently, it was confirmed that microencapsulation by spraydrying was successfully performed.
3.5.3. X-ray diffraction analysis
Because the crystallinity of microcapsules is related to their stability, it is important to determine whether the microcapsules have
crystalline or amorphous structures through XRD analysis. Generally,
the presence of diffuse and broad peaks in the XRD profile represents
amorphous structures because amorphous materials are disordered and
thus yield disperse bands. However, crystalline materials yield sharp
and defined peaks due to their well ordered state.
Fig. 2B presents the XRD profiles of chlorophylls, GA, MD, and
chlorophyll loaded microcapsules coated with GA-MD in ratios of 5:5,
3:7, and 0:10. Chlorophylls presented two broad and non-defined peaks
at 2θ = 8.6° and 19.5°, demonstrating an amorphous structure with
minimum crystallinity. GA, MD, and chlorophyll loaded microcapsules
coated with GA-MD in ratios of 5:5, 3:7, and 0:10 also had an amorphous structure with minimum organization, as indicated by the occurrence of broad and diffuse peaks. These results are consistent with
those obtained from several previous studies, which reported that GA
and MD generally had amorphous structures and the spray-drying
process did not influence the crystalline characteristics of the wall
materials (Silva, Coelho, Calado, & Rocha-Leão, 2013; Botrel, de Barros
Fernandes, Borges, & Yoshida, 2014).
The characteristic peaks of chlorophylls disappeared in the XRD
profiles of spray-dried microcapsules. The results provide further evidence that chlorophylls were largely embedded in the wall materials
composed of GA and MD, as revealed by DSC analysis (Fig. 3). In other
words, formation of chlorophyll loaded microcapsules were further
3.5.4. Differential scanning calorimetry analysis
The thermal behavior of chlorophylls and microcapsules coated
with GA-MD in ratios of 5:5, 3:7, and 0:10 evaluated by DSC, is presented in Fig. 3. Chlorophylls showed two endothermic peaks around
133 °C and 163 °C. The endothermic peak at 133 °C may be related to
melting of chlorophylls, which are known to melt at 117–130 °C (Guad,
Surana, Talele, Talele, & Gokhale, 2006). The other endothermic peak
at 163 °C may be due to melting of carotenoids (such as lutein and βcarotene) in the range of approximately 175–200 °C (Sy et al., 2012),
given that we used semi-purified chlorophylls extracted from spinach
by the ‘dioxane method’. Additionally, for microcapsules coated with
GA-MD in ratios of 5:5, 3:7, and 0:10, the DSC thermograms showed
endothermic peaks at 143 °C, 159 °C, and 197 °C, respectively. These
endothermic peak temperatures are associated with the melting point of
each microcapsule. However, the endothermic peak of chlorophylls
completely disappeared in the DSC thermograms of all microcapsules.
According to Paramera, Konteles, & Karathanos (2011), the absence of
thermal events of a core material in the thermal profile of microcapsules is regarded as evidence of true inclusion. Therefore, the obtained results confirmed that chlorophylls were successfully covered
with the wall materials, forming inclusion complexes. It is also important to point out that the melting points of each microcapsule shifted
to higher temperatures than that of chlorophylls. Particularly, the microcapsule coated with GA-MD in a ratio of 0:10 showed the highest
endothermic peak (197 °C), followed by microcapsules coated with GAMD in ratios of 3:7 and 5:5. Based on the DSC thermograms of MD (data
not shown), it is hypothesized that the higher endothermic peaks of
microcapsules might be related to the higher level of MD, which was
used as a wall material, because the endothermic peak (melting point)
of MD was observed at approximately 209 °C in this study. These results
indicate that thermal stability of chlorophylls was improved by microencapsulation using GA and MD as wall materials. Furthermore, the
optimal thermal stability of chlorophyll loaded microcapsules was
achieved when MD was only used as the wall materials.
3.6. Evaluation of storage stability at different temperatures
It is well known that the storage temperature is a crucial factor for
preserving heat sensitive materials such as nutrients, flavors, and microorganisms. Therefore, the importance of the storage temperature for
the stability of both non-encapsulated and encapsulated chlorophylls
was studied. The chlorophylls dissolved in MCT oil and all microcapsulated chlorophylls were stored at different temperatures (4, 20,
Fig. 3. DSC thermograms of chlorophylls (a), chlorophyll loaded microcapsules
prepared using GA-MD ratios of 5:5 (b), 3:7 (c), and 0:10 (d).
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and 40 °C) for 10 days in the dark, as shown in Fig. 4.
The retention of chlorophylls dissolved in MCT oil was approximately 92.3, 89.0, and 87.3% after 10 days of storage at 4, 20, and
40 °C, respectively. On the other hand, chlorophyll retention in microcapsules at the end of the period of 10 days was 93.9–97.5% at 4 °C,
94.2–96.5% at 20 °C, and 92.1–94.7% at 40 °C, indicating greater retention of encapsulated chlorophylls than non-encapsulated chlorophylls. The foregoing results apparently indicate that the stability of
chlorophylls was affected by the storage temperature, and entrapment
of chlorophylls within microcapsules augmented their storage stability.
Moreover, at the end of storage, microcapsules coated with MD
alone presented the highest storage stability of all the microcapsules,
followed by those coated with GA-MD in ratios of 3:7 and 5:5 at all
storage temperatures. It is proposed that an increase in MD concentration might have resulted in better protection against thermal
degradation of encapsulated chlorophylls. This hypothesis can be explained based on the above mentioned moisture content, XRD, and DSC
analyses of microcapsules. According to Velasco et al. (2003), as the
moisture content increases, a solid-state change from the glassy state to
amorphous state with a higher molecular mobility at a given temperature occurs, and is termed the glass transition temperature, indicating the point at which a solid collapses. In the case of microcapsules, it was reported that partial release of encapsulated materials
was associated with the foregoing physical changes of the solid, namely
changes from the glassy state to amorphous state. Therefore, in our
work, the highest storage stability of microcasule covered with MD
alone might be attributed to the lower molecular mobility of wall materials, induced by the lower moisture content compared to that of the
other microcapsules. The highest storage stability of chlorophylls
coated with MD alone may also be related to the characteristic XRD
profiles of this sample. As illustrated in Section 3.5.3, the amorphous
characteristics of particles lead to greater water absorption, and thus,
lower stability during storage compared to the crystalline particles. In
this sense, the superior storage stability of microcapsule coated with
MD alone may be derived from its higher degree of crystallinity due to
the greater content of MD in wall matrix. Lastly, the DSC thermograms
clearly indicated that an increment in the content of MD in wall matrix
contributed to enhancement of the thermal stability of microcapsules,
as evidenced by the shift in the endothermic peaks towards higher
temperature with increasing amounts of MD in samples (Fig. 3).
Conclusively, the foregoing results demonstrate that all chlorophyll
loaded microcapsules had good storage stability under various temperature conditions in comparison with non encapsulated chlorophylls.
Notably, the microencapsulation of chlorophylls using MD alone was
optimal for reducing chlorophyll degradation and enhancing the storage stability of the chlorophylls. In this regard, it is suggested that
microencapsulation by spray-drying with MD may be applicable for
enhancing the storage stability of chlorophylls or for the control of its
release properties.
4. Conclusions
The results obtained in this study demonstrate that MD alone can be
recommended as a carrier material for adequate microencapsulation of
chlorophylls as this material provides minimum moisture content and
maximum encapsulation efficiency and chlorophyll content, and exhibits better potential for the protection of microencapsulated chlorophylls from degradation. However, the addition of GA to the wall
materials resulted in a higher amount of surface oil on microcapsules
and more thermal degradation of chlorophylls during spray-drying, as
confirmed by SEM observation and the hunter color values, respectively. Generally, due to its low ability to form stable emulsions, MD is
used as a wall material by mixing with other biopolymers with good
emulsifying property. However, the present study demonstrates that
microcapsule coated with MD alone showed better physicochemical
characteristics and stability than microcapsules coated with blends of
Fig. 4. Stability of chlorophyll loaded microcapsules prepared using GA-MD
ratios of 5:5, 3:7, and 0:10 at 4 °C (A), 20 °C (B), and 40 °C (C) over 10 days. The
stability (%) was calculated as follows: relative amount of chlorophyll remaining in microcapsules with respect to the initial amount in microcapsules.
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GA and MD. This is because Tween® 80 was bound to MD, and thus
improved its emulsion forming ability. Although the current findings
indicate that spray-drying using MD as a carrier agent is a good option
for encapsulation of chlorophylls, the pattern of chlorophyll release in
the gastrointestinal tract was not demonstrated. Therefore, further
studies will be needed to evaluate the stability and sustained release of
chlorophylls under simulated gastrointestinal conditions.
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Appendix A. Supplementary data
Supplementary data associated with this article can be found, in the
online version, at https://doi.org/10.1016/j.foodchem.2018.08.063.
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