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original scientific paper
ISSN 1330-9862
doi: 10.17113/ftb.56.03.18.5449
Effect of Lactobacillus plantarum Fermentation on the Surface and
Functional Properties of Pea Protein-Enriched Flour
Burcu Çabuk, Andrea K.
Stone, Darren R. Korber,
Takuji Tanaka and
Michael T. Nickerson*
Department of Food and Bioproduct
Sciences, University of Saskatchewan,
51 Campus Drive, Saskatoon, SK, S7N
5A8, Canada
Received: 30 July 2017
Accepted: 13 July 2018
SUMMARY
The effect of Lactobacillus plantarum fermentation on the functional and physicochemical properties of pea protein-enriched flour (PPF) was investigated. Over the course of the
fermentation the extent of hydrolysis increased continuously until reaching a maximum
degree of hydrolysis of 13.5 % after 11 h. The resulting fermented flour was then adjusted
to either pH=4 or 7 prior to measuring the surface and functional attributes as a function
of fermentation time. At pH=4 surface charge, as measured by zeta potential, initially increased from +14 to +27 mV after 1 h of fermentation, and then decreased to +10 mV after
11 h; whereas at pH=7 the charge gradually increased from –37 to –27 mV over the entire
fermentation time. Surface hydrophobicity significantly increased at pH=4 as a function
of fermentation time, whereas at pH=7 fermentation induced only a slight decrease in
PPF surface hydrophobicity. Foam capacity was highest at pH=4 using PPF fermented for
5 h whereas foam stability was low at both pH values for all samples. Emulsifying activity
sharply decreased after 5 h of fermentation at pH=4. Emulsion stability improved at pH=7
after 5 h of fermentation as compared to the control. Oil-holding capacity improved from
1.8 g/g at time 0 to 3.5 g/g by the end of 11 h of fermentation, whereas water hydration capacity decreased after 5 h, then increased after 9 h of fermentation. These results indicate
that the fermentation of PPF can modify its properties, which can lead towards its utilization as a functional food ingredient.
Key words: pea protein-enriched flour, fermentation, functional properties, lactic acid bacteria
INTRODUCTION
*Corresponding author:
Phone: +13069665030;
Fax: +13069668898;
E-mail: Michael.Nickerson@usask.ca
ORCID IDs: 0000-0001-8836-6047
(Çabuk), 0000-0002-0918-1966 (Stone),
0000-0001-6350-209X (Korber),
0000-0003-3304-5830 (Tanaka),
0000-0002-9040-5639 (Nickerson)
Dry edible pea (Pisum sativum) is widely consumed around the globe as a healthy food.
Peas are rich in proteins, carbohydrates, fibre, and vitamins/minerals, and are low in fat (1).
The protein content of pea is higher than many other staple foods, thus pea is considered a
good protein source. Pea protein is rich in lysine, but deficient in the thiol-containing amino
acids (cysteine and methionine) (2,3). As such, it is commonly recommended that one consume pulses such as pea along with cereal grains in order to obtain a complete essential amino acid profile (4), especially in places where animal protein is limited and expensive due to
issues of food insecurity. While peas are a good protein source, they also contain secondary
metabolites considered to be anti-nutritional factors, such as enzyme inhibitors (trypsin and
chymotrypsin inhibitors), oxalates, phytates, oligosaccharides, phenolics, tannins and lectins,
that inhibit protein digestion or mineral absorption when consumed (5).
Peas are usually eaten whole or split, milled into flour (21–24 % protein), air classified into
either a protein-enriched flour (30–50 % protein) or a protein concentrate (50–80 % protein),
or wet processed into a protein isolate (>80 % protein) (6). Each particular fraction may be
incorporated into different products, applications or sectors. Animal-derived proteins from
milk (casein, whey) and egg (ovalbumin), along with plant sources such as wheat (gluten)
and soy (glycinin, conglycinin) dominate the protein ingredient market (7). However, consumers are demanding an increase in alternative protein sources due to the allergen content
of the current protein products. Accordingly, a rise in the demand for pea protein products
July-September 2018 | Vol. 56 | No. 3 411
B. ÇABUK et al. Fermented Pea Protein Functionality and Surface Properties
has occurred since they tend to have low allergenicity. In addition, pea protein products have other market benefits, including not being genetically modified, low in cost, nutritious and
functional (8,9). As the demand for pea increases, so too does
the need for greater variations and choice in pea protein ingredients; processing, such as infrared heating, germination,
enzymatic modification and fermentation can alter the ingredients (9). Fermentation is of great interest to industry as it is a
non-thermal process whose technology acts to partially unravel
the protein conformation to expose buried reactive amino acids and improve its digestibility (10). The use of bacteria or fungi to produce proteases is essential to fermentation processes,
as they not only initiate partial protein unfolding but also facilitate the release of low-molecular-mass peptides with potential
bioactive properties (11-13). Fermentation also acts to reduce
the content of anti-nutritional factors within pulse ingredients,
and help improve mineral absorption through the production
of organic acids which form soluble complexes with minerals,
rendering the minerals unavailable to react with phytates and
become insoluble (14-16).
Fermented pulses are consumed primarily in Asia, Africa and
Europe, with less uptake in North America. However, limited research efforts have been reported on the effects of fermentation conditions on the functional properties of commercial pea
protein ingredients, such as a protein-enriched flour. The overall
goal of the present study is to examine the impact of fermentation of pea protein-enriched flour by Lactobacillus plantarum on
the protein surface and functional properties at pH=4 (indicative
of an acidic food, and near pea protein isoelectric point; pI) and
pH=7 (indicative of the neutralization process in wet processing,
and away from pea protein pI) in order to diversify the pea ingredient line for greater market integration. Fermentation studies commonly use L. plantarum due to its generally recognized
as safe (GRAS) status, growth ability and its ubiquitous nature in
fermented food products (17). We chose it for this study because
of its growth requirements, i.e. the required fermentation conditions do not affect the pea protein quality.
MATERIALS AND METHODS
Materials
Parrheim Foods (Saskatoon, SK, Canada) kindly donated the air-classified pea protein-enriched flour (PPF). 8-Anilino-1-naphthalenesulfonic acid (ANS) and 5,5’-dithiobis-2-nitrobenzoic acid (DNTB) were products of Sigma-Aldrich
(Oakville, ON, Canada), MRS broth was a product of Oxoid (Nepean, ON, Canada), whereas all other chemicals used were of
reagent grade and purchased through Fisher Scientific (Ottawa, ON, Canada). A Millipore Milli-Q™ water purification system (Millipore Corp., Etobicoke, ON, Canada) produced the
water used in this stidy. Canola oil (Great Value™, Wal-Mart
Canada Corp., Mississauga, ON, Canada) was purchased from
a local supermarket. Lactobacillus plantarum NRRL B-4496 was
obtained from the Agricultural Research Service Culture Collection, USDA (Peoria, IL, USA).
412
Fermentation
A growth curve for Lactobacillus plantarum NRRL B-4496
was initially prepared within MRS broth (Oxoid) for approx.
24 h at 32 °C under anaerobic conditions. For fermentation
experiments cell cultivation lasted until the late exponential
phase of growth (approx. 10 h), followed by centrifugation
(10 000×g, 20 min, 4 °C; centrifuge model 5810R; Eppendorf,
Mississauga, ON, Canada), and then washing twice with peptone solution. The resulting pellet was used as the inoculum
for fermentation. Lactobacillus plantarum was added to a 25 %
(m/V) PPF solution (400 mL) in an Erlenmeyer flask at a content of 7 log CFU per g PPF, which was then incubated under
anaerobic conditions at 32 °C for 11 h. Enumeration of L. plantarum was carried out by plating onto MRS medium (Oxoid)
at 37 °C for 48 h under anaerobic conditions. Anaerobic conditions were maintained by placing the experiments within a
rectangular jar with Anaerogen anaerobic gas generating kit
(Thermo Scientific, Waltham, MA, USA). Aliquots (60 mL) were
taken at 0, 1, 5, 9 and 11 h of fermentation and then freezedried for 48 h using a freeze dryer (Labconco, Freezone 12,
Kansas City, MO, USA). All dried samples were then ground
using a coffee grinder (model 80365; Hamilton Beach Custom Grind™, Glen Allen, VA, USA). Fermentation experiments
were run in triplicate, yielding three separate fermented PPF
for each time point.
Composition
Moisture content was determined gravimetrically by calculating the mass loss after drying 2 g sample in an oven
(APT.line™ ED, BINDER GmbH, Tuttlingen, Germany) at 130 °C
for 1 h, according to AOAC method 925.10 (18). Protein content (N/%·6.25) was determined according to AOAC method
920.87 (19) using a Kjeldahl micro digestor (model 6030000;
Labconco) and distillation unit (Rapid Still I; Labconco). Ash
content was determined according to AOAC method 923.03
(20) in which 3 g sample was placed overnight in a muffle furnace (Isotemp®; Thermo Fisher Scientific, Waltham, MA, USA)
at 550 °C and expressed as the difference of the mass of the
sample before and after heating. Crude lipid was determined
gravimetrically after solvent extraction with ether according
to AOAC method 920.39 (21) using a Labconco Goldfisch fat
extractor. Proximate analysis was reported on a dry mass basis. To determine the pH, 15 g of PPF fermentation medium
at each time interval (0, 1, 5, 9, and 11 h) were transferred to a
25-mL beaker and the pH was measured under stirring conditions using a pH meter (B10P Benchtop Meter; VWR, Mississauga, ON, Canada) and magnetic stirrer plate (RO 5; IKA
Works Inc., Wilmington, NC, USA).
Degree of hydrolysis
The degree of hydrolysis (DH/%) of fermented PPF samples was calculated according to Adler-Nissen (22) using the
following formulae:
July-September 2018 | Vol. 56 | No. 3
Food Technol. Biotechnol. 56 (3) 411-420 (2018)
/1/
and
/2/
where Yh is the yield of hydrolysis equivalents (of α-amino
groups formed during the hydrolysis reaction; or α-NH2-Gly
equivalent), ct is the millimolar concentration of α-NH2-Gly
equivalent (measured using a glycine standard curve) obtained from the trypsin-catalyzed protein hydrolysis reactions, cc is the millimolar concentration of α-NH2-Gly equivalent from the non-trypsin treated PPF (control), ctot is the
millimolar concentration of α-NH2-Gly equivalent from the
total PPF hydrolysis, and DF is a dilution factor.
Surface properties
Surface charge
The surface charge (or zeta potential) of each fermented
sample was determined according to Can Karaca et al. (23). In
brief, 0.05 % (by mass) of protein was dissolved in Milli-Q water (Millipore Corp.) and adjusted to pH=4.0 or 7.0 with 0.5 M
NaOH or HCl. The solution was then stirred overnight at 500
rpm using a magnetic stirrer plate (RO 5; IKA Works Inc.) at
room temperature (21–23 °C). The electrophoretic mobility
was measured using a Zetasizer Nano (Malvern Instruments,
Westborough, MA, USA). The zeta-potential (ζ/mV) was determined from the electrophoretic mobility (μE) using Henry’s
equation, as follows:
/3/
where η is the dispersion viscosity, ε is the permittivity, and
ƒ(κα) is a function related to the ratio of particle radius (α) and
the Debye length (κ).
Surface hydrophobicity
Surface hydrophobicity of fermented samples was determined with 8-anilino-1-naphthalenesulfonic acid (ANS) fluorescent probe (Sigma-Aldrich) using the modified method of
Kato and Nakai (24). Briefly, a 0.025 % (by mass) protein solution was prepared in Milli-Q water (Millipore Corp.), adjusted
to pH=4 or 7.0 using 0.5 M NaOH or HCl, and stirred overnight
at 500 rpm using a magnetic stirrer (RO 5; IKA Works Inc.) at
room temperature. The stock solution was then diluted to final protein mass fractions of 0.005, 0.010, 0.015, and 0.020 %. A
20-µL aliquot of 8 mM ANS solution (in Milli-Q water, pH=4 or
7) was added to 1.6 mL of each protein mass fraction, vortexed
for 10 s and kept in the dark for 5 min. The fluorescence intensity was measured with a FluoroMax-4 spectrofluorometer
(Horiba Jobin Yvon Inc., Edison, NJ, USA) using excitation and
emission wavelengths of 390 and 470 nm, respectively, and a
slit width of 1 nm. Sample blanks were prepared by adding
20 µL of Milli-Q water (pH=4 or 7) to the protein solutions instead of the ANS probe (Sigma-Aldrich). The initial slope of
the plot of the fluorescence intensity (protein solution with
probe minus the same protein solution with water) vs. protein
mass fraction was calculated using linear regression analysis
and used as an index of the surface hydrophobicity. All intensity data were arbitrarily divided by 10 000 prior to statistical
analysis and graphing.
Functional properties
Emulsifying properties
The emulsifying activity (EA) and emulsion stability (ES)
were determined according to Yasumatsu et al. (25). In brief,
1 g of fermented PPF was suspended in 14.3 mL of Milli-Q water (Millipore Corp.) and adjusted to pH=4.0 or 7.0 with either
0.5 M NaOH or HCl. The solution was stirred for 30 min using
a mechanical stirrer (500 rpm) (RO 5; IKA Works Inc.) at room
temperature (21–23 °C). Then, 14.3 mL of canola oil were added, followed by homogenization using an Omni macro-homogenizer (Omni International, Marietta, GA, USA), equipped
with a 20-mm saw tooth probe, at speed 4 (approx. 7200 rpm)
for 1 min. A 10-mL aliquot of the emulsion was poured into
two 15-mL centrifuge tubes and centrifuged at 1300×g for 5
min (model 5810R; Eppendorf, Mississauga, ON, Canada). The
emulsifying activity was determined as follows:
/4/
where h0 is the total height of the emulsion layer prior to centrifugation and h1 is the total height of the emulsion layer after centrifugation. Emulsion stability was determined by preparing the emulsion as previously described, then heating it
at 80 °C for 30 min using a water bath. The emulsion was then
cooled to room temperature using tap water over a 30-minute
period. A 10-mL aliquot of the cooled to room temperature
emulsion was then placed into two 15-mL centrifuge tubes
and centrifuged at 1300×g for 5 min. Emulsion stability was
determined as follows:
/5/
where EAH is the emulsifying activity of the heated emulsion
and EA of the unheated emulsion.
Foaming properties
The foam capacity (FC) and foam stability (FS) were determined according to Liu et al. (26). In brief, 1 g of fermented PPF
was dispersed within 25 mL of Milli-Q water (Millipore Corp.)
and adjusted to pH=4.0 or 7.0 with either 0.5 M NaOH or HCl,
and stirred for 30 min on a mechanical stirrer (500 rpm; RO 5;
IKA Works Inc.) at room temperature (21–23 °C). A 15-mL aliquot was then transferred into a 400-mL beaker for foaming using an Omni Macro Homogenizer (Omni International),
July-September 2018 | Vol. 56 | No. 3 413
B. ÇABUK et al. Fermented Pea Protein Functionality and Surface Properties
equipped with a 20-mm saw tooth probe, at speed 4 (aprox.
7200 rpm) for 5 min. The resulting foamed sample was transferred to a 100-mL graduated cylinder and the foam volume
measured at time 0 and after 30 min. FC and FS were determined as follows:
/6/
and
/7/
where VF0 is the volume of the foam at 0 min, Vsample is the initial volume of sample used (15 mL), and VF30 is the foam volume after 30 min.
Nitrogen solubility index
Fermented PPF samples (1 g) were suspended in 25 mL of
Milli-Q water (Millipore Corp.) and adjusted to pH=4.0 or 7.0
with either 0.5 M NaOH or HCl at room temperature (21–23 °C)
and stirred on a mechanical stirrer (RO 5; IKA Works Inc.) at 500
rpm for 30 min. The suspension was centrifuged at 3070×g for
10 min (model 5804R; Eppendorf). The nitrogen solubility index (in %) was determined by dividing the nitrogen measured
in the supernatant by the original amount in the fermented
samples, multiplied by 100. Nitrogen mass fraction within the
fermented PPF and the supernatant after extraction were determined using AOAC method 920.87 (19).
Water hydration capacity and oil-holding capacity
Water hydration capacity (WHC) and oil-holding capacity
(OHC) values for fermented PPF samples were determined according to Stone et al. (27) with slight modifications. In brief, 10
mL of canola oil (or water, pH=4.0 or 7.0 for WHC) were added to
1 g of PPF in a 50-mL centrifuge tube. The mixture was vortexed
for 10 s every 5 min for 30 min, followed by centrifugation at
11 180×g for 15 min (model 5804R; Eppendorf). OHC or WHC
values were determined as the mass change in fermented protein samples after decanting (wet protein PPF mass minus dry
PPF mass) relative to the dry PPF mass (1 g).
Statistical analysis
All data is reported as the mean value±standard deviation
of PPF derived from triplicate fermentation batches (N=3). A
one-way ANOVA with a Tukey’s test was used to detect statistical differences in response to fermentation time within
compositional, degree of hydrolysis and OHC data. A two-way ANOVA was used to test for significant differences between the main effects of fermentation time and pH, along
with their interaction for all surface and functional (except
for OHC) properties tested. All statistical analyses were performed with Systat v. 10 software (28).
414
RESULTS AND DISCUSSION
Physicochemical properties
Table 1 shows changes to the proximate composition and
pH of the freeze-dried PPF as a function of fermentation time.
All crude protein, ash, and lipid mass fractions changed significantly during fermentation (p<0.05), presumably due to an
increase in the bacterial biomass present and loss of carbohydrates during the process. In the case of the latter, carbohydrates decreased from approx. 53 to 37 % (based on calculated
difference) during fermentation. Crude fat content increased
from 2.5 to 3.5 % after 11 h of fermentation, most likely due to
the loss of carbohydrates. Literature reports various effects of
fermentation on the crude fat content of pulses. For instance,
solid-state fermentation of chickpea tempeh flour decreased
the fat content from 6.1 % in the raw chickpea flour to 2.6 %
in the tempeh flour (29), whereas cowpea fat content slightly increased after fermentation, from 0.9 to approx. 2 % (30).
In the current study, protein mass fraction increased sharply
from 42.9 to approx. 47 % between 1 and 5 h of fermentation
before leveling off, a result hypothesized to be associated with
the exponential growth of the L. plantarum cells and the loss
of carbohydrates. In contrast, the ash mass fraction increased
steadily from 4.2 to 11.0 % over the 11-hour fermentation time,
again thought to be due to the loss of carbohydrates.
Like other Lactobacillus spp., growth of L. plantarum during
fermentation leads to the production of weak acids and the
release of small peptides from the proteins, resulting in a reduction in pH from pH=7.5 at time 0 to pH=4.3 after 11 h of
fermentation. Chandra-Hioe et al. (31) have previously reported a reduction in pH during fermentation of fermented chickpea and faba bean flour. Because of the pH reduction, the pH
of the starting material and the fermented PPF (taken at different fermentation times) was readjusted to pH=4 (near the
protein pI) and pH=7 (representative of the neutralization process commonly used in commercial wet processing of protein
ingredients) prior to measuring their surface and functional
characteristics. Increase in protein mass fraction during fermentation appears to be system-dependent. Chandra-Hioe
et al. (31) reported no change in the protein content of desi or
kabuli chickpea flour after 16 h of fermentation; however, the
protein content of faba bean flour rose from 23 to 30 % over
the same fermentation period. Reyes-Moreno et al. (29) and
Akubor and Chukwu (32) reported a 22 and 18 % increase in
protein mass fraction of fermented chickpea flour and full fat
African oil bean seed flour, respectively.
Changes to the degree of hydrolysis (DH) of PPF during
fermentation showed a sigmoidal increase with a maximum
value of approx. 13 % after 11 h (Table 1). A one-way analysis of
variance found that changes to the degree of hydrolysis with
fermentation time were significant (p<0.05), where values increased from 0 % at time 0 to 9.7 % after 5 h, 10.6 % after 9 h,
and a further increase to 13.5 % after 11 h. During a preliminary study (data not shown), fermentation up to 48 h did not
yield any further changes to the DH values, possibly because:
July-September 2018 | Vol. 56 | No. 3
Food Technol. Biotechnol. 56 (3) 411-420 (2018)
Table 1. Changes to the composition (on dry mass basis), degree of hydrolysis and pH of pea protein-enriched flour fermented by Lactobacillus
plantarum over an 11-hour time course
t/h
w(crude protein)/%
w(crude ash)/%
w(crude lipid)/%
w(crude CHO)/%
DH/%
pH
0
(40.1±1.2)a
(4.2±1.2)a
(2.5±0.1)a
53.2
-
(7.5±0.0)a
1
(42.9±1.3)a
(5.9±0.9)ab
(2.9±0.4)ab
48.3
(6.1±0.2)a
(7.2±0.0)b
5
b
(46.6±0.7)
(5.9±0.7)
a
b
(6.3±0.0)c
9
c
11
d
(2.6±0.0)
44.9
(9.7±0.5)
(46.4±0.1)
bc
(8.4±1.4)
b
(3.4±0.0)
41.8
(10.6±0.8)
(4.4±0.1)d
(48.1±0.4)
(11.0±0.4)
(3.5±0.0)
37.4
(13.5±0.0)
(4.3±0.0)e
b
b
ab
c
b
DH=degree of hydrolysis, CHO=carbohydrates determined based on the percentage difference between 100 % and the mean values of protein,
ash, and lipid mass fractions. Data represent the mean value±standard deviation (N=3). Data with different superscript letters in the same column
are significantly different (p<0.05)
ζ/mV
The surface charge (zeta potential) and hydrophobicity
of all PPF samples at both pH=4 and 7 are given in Figs. 1a
and 1b, respectively. Overall, the zeta potential (ZP) of the PPF
was positive when adjusted to pH=4 and negative when at
pH=7, as the proteins would be below and above, respectively, the isoelectric point of pea protein (pI~4.6) (Fig. 1a). As determined by a two-way ANOVA for ZP data, both the effects of
fermentation time and pH, and their interaction, were significant (p<0.001). At pH=4 the ZP increased from approx. +14 mV
in the unfermented sample to a maximum of +27 mV in 1-hour
fermented samples, followed by a gradual decline to +10 mV
by 11-hour fermentation. In contrast, at pH=7 the ZP increased
gradually from –37 mV at time 0 to –27 mV at 11 h of fermentation (Fig. 1a). The decreases in charge density at both pH values
may indicate that limited hydrolysis of PPF leads to exposure
a) 60
pH=4
of few numbers of both positively- and negatively-charged
groups
since
changes
in
net
charge
at
both
pH
values
pH=7were
40
low. A decrease in ZP at pH=7 due to an increase in degree of
hydrolysis
was reported by Ghribi et al. (33) where chickpea
20
protein isolate was modified via enzymatic hydrolysis.
A 0two-way ANOVA for surface hydrophobicity data found
that both the fermentation time and pH, and their interaction,-20
were all significant (p<0.001). In samples at pH=4 surface
hydrophobicity remained constant (approx. 9 arbitrary units,
-40
AU) between 0 and 1 h of fermentation, and then steadily increased to approx. 21 AU in 9-hour samples, at which time it
-60
plateaued
at6pH=7 surface
hydrophobici0 (Fig. 1b).
2 In contrast,
4
8
10
12
ty declined very slightly from
approx.
8
AU
in
the
unfermented
t(fermentation)/h
samples to approx. 7 AU in 11-hour fermented samples (Fig.
1b). It is hypothesized that at pH=4.0, after 1 h of fermentation
hydrolysis of the PPF leads to a partial unraveling of the protein and release of peptides which exposed buried reactive
charged and hydrophobic sites (34). However, after 1 h, fermentation-induced changes to the surface properties reflect
the continued unraveling of the protein structure and the increase in bacterial biomass protein. Since the protein would
be only weakly charged at this pH, conformational entropy
would be greater allowing it to unravel more, as evidenced by
a)
60
b)
pH=4
pH=7
40
20
ζ/mV
Surface properties
an increase in hydrophobicity and a slight decline in charge.
In contrast, for samples at pH=7, sufficient electrostatic repulsion between particles would overshadow the minor changes that would occur in protein conformation (and hence surface charge and hydrophobicity) as the result of fermentation.
0
-20
-40
-60
0
2
4
6
8
10
12
10
12
t(fermentation)/h
b)
Surface hydrophobicity/AU
(i) the low pH of the medium (approx. 4.3) restricted further
cleavage of the proteins, and (ii) no additional proteinase production occurred during the stationary phase of growth (22).
30
25
20
15
10
5
0
0
2
4
6
8
t(fermentation)/h
Fig. 1. Effect of fermentation time and pH on: a) surface charge (ζ),
and b) hydrophobicity of pea protein-enriched flour fermented by
Lactobacillus plantarum. Data represent the mean value±standard
deviation (N=3)
Functional properties
Emulsification
The emulsifying activity (EA) and emulsion stability (ES)
of fermented PPF as a function of fermentation time and pH
are given in Figs. 2a and 2b. A two-way ANOVA found that
July-September 2018 | Vol. 56 | No. 3 415
B. ÇABUK et al. Fermented Pea Protein Functionality and Surface Properties
fermentation time, pH and their associated interaction have a
highly significant effect on both EA and ES (p<0.001). The EA,
a measure of emulsion forming ability, was constant (approx.
45 %) between 0 and 5 h of fermentation, and then declined
significantly to approx. 5–7 % after 9 h of fermentation (Fig.
2a) at pH=4. This decline corresponds to the sharp increase in
hydrophobicity observed at the corresponding pH in 5-hour
fermented samples. The increased hydrophobicity may have
impacted the protein’s ability to migrate to the oil-water interface to lower interfacial tension and facilitate emulsion formation and possibly favour the aggregation of released peptides
and unhydrolyzed proteins (35). Emulsion formation was lower (EA approx. 35 %) at pH=7 compared to pH=4 between fermentation time 0 and 5 h, however, EA remained relatively constant over the 11 h of fermentation at pH=7, possibly due to the
higher charge density and conformational flexibility of peptide
and protein molecules (35). However, the emulsions formed at
pH=4 were inherently less stable than those formed at pH=7,
except after 11 h of fermentation where the ES was similar at
each pH. At pH=4, ES values were relatively constant at approx. 20 % over the entire fermentation time course; whereas,
at pH=7, ES values increased from approx. 37 % at time 0 to approx. 56 % at 5 h, and then declined to approx. 20 % by the end
of fermentation (Fig. 2b). Unfortunately, a clear mechanism underlining emulsion stability could not be reached in this study,
due to several confounding effects associated with changes in
surface characteristics and solubility during the fermentation
time/pH, as well as other factors, not measured as part of this
study such as differences in droplet sizes and rate of creaming
based on Stokes’s law, and the concentration and viscoelasticity of the film formed at the oil-water interface.
constant surface properties at this pH (Fig. 1). Foam stability
at pH=4 was relatively constant at approx. 20 % for samples
up to 5 h of fermentation, but then dropped to 9 % after 9 h
of fermentation. It was hypothesized that longer fermentation
times lead to greater hydrophobicity, which may have inhibited the migration of protein to the air-water interface where
bubbles continually broke and reformed. In contrast, at pH=7,
FS of samples remained relatively constant at approx. 20 %
over the entire fermentation period (Fig. 2d). No large changes in the surface properties of the PPF were evident at pH=7,
leading to the observed constant FS. The foaming properties
of African oil bean seed (Pentaclethra macrophylla) flour (32),
African breadfruit seed (Treculia africana) flour (36), and peanut protein concentrate (37) all improved after fermentation.
Nitrogen solubility indices
The nitrogen solubility indices (NSI) of PPF as a function
of fermentation time and pH are shown in Fig. 2e. The effect
of fermentation time, pH and their associated interaction, all
had a significant effect on NSI (p<0.001). Overall, NSI were relatively low at approx. 8–11 % at pH=4 regardless of the fermentation time; whereas, at pH=7, NSI decreased from approx. 43
to 36 % after 11 h of fermentation (Fig. 2e). Increased nitrogen
solubility at the higher pH is associated with the greater surface charge and amount of electrostatic repulsive forces present relative to pH=4, which is closer to the pI of PPF. The slight
decline in NSI at pH=7 is hypothesized to be attributed to the
increase in biomass protein which was presumed to have lower solubility than the pea protein.
Water hydration capacity
Foaming
The foam capacity (FC) and stability of PPF and fermented
PPF as a function of fermentation time and pH is given in Figs.
2c and 2d. Fermentation time, pH and their associated interaction all had a very significant effect on FC (p<0.001). In contrast, for FS data only the effect of fermentation time and the
interaction of fermentation time with pH were significant
(p<0.001). The foam-forming properties of PPF at pH=4 increased FC from approx. 80 % at time 0 to approx. 90 % FC
after 5 h of fermentation, and thereafter declined to approx.
70 % after 9 h of fermentation (Fig. 2c). Similar to emulsions,
the foam forming properties of a protein depend on its ability
to migrate to the air-water interface to lower surface tension,
and then realign its hydrophobic groups towards the apolar
phase and the hydrophilic groups towards the polar phase.
At pH=4, an increase in FC was observed as the protein unravels and exposes hydrophobic groups; however, after 5 h
of fermentation, it was presumed that an overabundance of
hydrophobic groups were exposed, reducing the ability of the
protein to migrate to the interface and therefore decreasing
FC. At pH=7, some variability was evident within the fermentation period, but FC values remained relatively constant at approx. 70 % (Fig. 2c), which was most likely due to the relatively
416
The WHC of fermented PPF as a function of fermentation
time and pH is given in Fig. 2f. A two-way ANOVA of WHC data
found only the effect of fermentation time (p<0.001) and its interaction with pH (p<0.01) to be significant, but not the effect
of pH alone (p>0.05). WHC values declined from approx. 1–1.2
g/g at time 0 to 0.8–0.9 g/g after 5 h of fermentation, and then
increased to 1.5–1.6 g/g after 9 h of fermentation at both pH
values, however, which one was greater (pH=4 vs. 7) seemed
to fluctuate a little within this trend (Fig. 2f). For instance, WHC
at pH=4 was slightly greater than WHC at pH=7 at t=0 and 9 h,
whereas WHC at pH=7 was slightly greater than WHC at pH=4
for t=1 and 5 h, with both pH values being similar in magnitude
at t=11 h (Fig. 2f). We hypothesized these findings were associated with protein hydrolysis to a point where the pea proteins
unravel to expose buried hydrophilic sites which then interact
with more water. Similarly, Xiao et al. (38) found that the fermentation of chickpea with Cordyceps militaris SN-18 significantly increased the WHC. In another study, Oloyede et al. (39)
reported that the WHC of Moringa oleifera seed flour increased
during the first 24 h of fermentation and then, after 72 h, it declined. Similarly, Akubor and Chukwu (32) reported that the
water absorption properties of African oil bean (Pentaclethra
macrophylla) seed flour increased by 36 % when fermented.
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Food Technol. Biotechnol. 56 (3) 411-420 (2018)
a) 100
pH=4
pH=7
80
60
80
ES/%
EA/%
b) 100
40
20
60
40
20
0
0
0
2
4
6
8
t(fermentation)/h
10
12
0
4
80
80
60
60
FS/%
d) 100
40
20
8
10
12
8
10
12
10
12
40
20
0
0
0
2
4
6
8
10
12
0
2
4
t(fermentation)/h
6
t(fermentation)/h
e) 100
f) 2.0
80
1.5
60
WHC/(g/g)
NSI/%
6
t(fermentation)/h
c) 100
FC/%
2
40
1.0
0.5
20
0
0.0
0
2
4
6
8
t(fermentation)/h
10
12
0
2
4
6
8
t(fermentation)/h
Fig. 2. Effect of fermentation time and pH on: a) emulsifying activity (EA), b) emulsion stability (ES), c) foam capacity (FC), d) foam stability (FS), e)
nitrogen solubility index (NSI), and f) water hydration capacity (WHC) of pea protein-enriched flour fermented by Lactobacillus plantarum. Data
represent the mean value±standard deviation (N=3)
Oil-holding capacity
Fig. 3 shows the OHC of PPF as a function of fermentation time. Fermentation time had a significant effect on OHC
(p<0.001) as determined by a one-way ANOVA. Overall, OHC
increased from approx. 1.8 g/g at time 0 h to approx. 3.5 g/g after 1 h of fermentation, after which it declined to 2.5 g/g at 9 h
of fermentation and then increased again to 3.5 g/g after 11 h
(Fig. 3). Since we did not adjust the pH during the OHC test, the
pH would have differed depending on the fermentation time
(Table 1). The pH declined from pH=7.5 to 4.3 over the 11 h of
fermentation, which resulted in a decrease in protein surface
charge and solubility as well as an increase in hydrophobicity
to allow for increased protein interactions with oil. As evident
in Fig. 1b, surface hydrophobicity increased greatly at pH=4
relative to pH=7. During fermentation, the proteins become
partially unraveled to expose buried hydrophobic groups that
can bind oil. Fermentation also increased bacterial biomass in
the PPF, which in this case altered OHC in a positive manner.
There are also reports on an increase in OHC in fermented
chickpea and faba bean flours (31). Periago et al. (40) reported
a similar increase in OHC with an increase in degree of hydrolysis (via enzymatic modification) of pea flour.
July-September 2018 | Vol. 56 | No. 3 417
B. ÇABUK et al. Fermented Pea Protein Functionality and Surface Properties
REFERENCES
6
1. Tulbek MC, Lam RSH, Wang Y, Asavajaru P, Lam A. Pea: A sustainable vegetable protein crop. In: Nadathur SR, Wanasundara JPD, Scanlin L, editors. Sustainable protein sources.
Cambridge, MA, USA: Academic Press; 2017. pp. 145–64.
OHC/(g/g)
5
4
3
https://doi.org/10.1016/B978-0-12-802778-3.00009-3
2
2. Leterme P, Monmart T, Baudart E. Amino acid composition
of pea (Pisum sativum) proteins and protein profile of pea
flour. J Sci Food Agric. 1990;53(1):107–10.
1
https://doi.org/10.1002/jsfa.2740530112
0
0
2
4
6
8
10
12
t(fermentation)/h
Fig. 3. Effect of fermentation time on the oil-holding capacity (OHC)
of pea protein-enriched flour fermented by Lactobacillus plantarum.
Data represent the mean value±standard deviation (N=3)
CONCLUSIONS
Lactobacillus plantarum fermentation of pea protein-enriched flour over the course of 11 h achieved 13.5 % degree
of hydrolysis, a drop in pH and significant changes to the protein surface and functional properties. Crude protein and ash
mass fractions of the PPF increased as fermentation time increased due to the microbial proliferation, as the amount of
biomass increased and the amount of carbohydrates declined.
During fermentation, enzymes cleave part of the protein causing the changes in conformation, exposing the buried hydrophobic sites and resulting in higher hydrophobicity of the PPF,
at pH=4, after longer fermentation times. Depending on the
time, different levels of hydrolysis led to different surface characteristics and functionality of PPF. The final functionality of
the fermented ingredients, like other proteins, can be further
tailored depending on extrinsic factors, such as solution pH.
For instance, fermented PPF exhibited relatively better emulsion stability at pH=7 after 5 h of fermentation and improved
foam capacity at pH=4 after 5 h of fermentation. It is important to note that the final ingredient is a blend of both pea protein and microbial biomass, giving its novel properties, which
is highly dependent on both pH and length of fermentation.
Overall, the fermented PPFs have the potential to be incorporated into food products, such as beverages, sports bars, nutritional supplements, and so on. The use of fermented pea protein may be advantageous or disadvantageous, depending
on the level of hydrolysis and food environment, all of which
could influence its functional attributes. Although not tested
as part of this study, fermented ingredients also tend to have
unique flavour profiles, reduced non-nutritive compounds
and bioactive peptides for enhanced nutrition.
3. Reinkensmeier A, Bußler S, Schlüter O, Rohn S, Rawel HM.
Characterization of individual proteins in pea protein isolates and air classified samples. Food Res Int. 2015;76(Pt. 1):
160–7.
https://doi.org/10.1016/j.foodres.2015.05.009
4. Multari S, Stewart D, Russell WR. Potential of fava bean as future protein supply to partially replace meat intake in the human diet. Compr Rev Food Sci Food Saf. 2015;14(5):511–22.
https://doi.org/10.1111/1541-4337.12146
5. Patterson CA, Curran J, Der T. Effect processing on antinutrient compounds in pulses. Cereal Chem. 2017;94(1):2–10.
https://doi.org/10.1094/CCHEM-05-16-0144-FI
6. Kiosseoglou V, Paraskevopoulou A. Functional and physicochemical properties of pulse proteins. In: Tiwari BK, Gowen A, McKenna B, editors. Pulse foods: Processing, quality
and nutraceutical applications. Oxford, UK: Academic Press;
2011. pp. 57–90.
https://doi.org/10.1016/B978-0-12-382018-1.00003-4
7. Global protein ingredients market size, industry report,
2018–2025. San Francisco, CA, USA: Grand View Research;
2018. Report ID: 978-1-68038-451-2.
8. Hall C, Hillen C, Garden Robinson J. Composition, nutritional value, and health benefits of pulses. Cereal Chem. 2017;
94(1):11–31.
https://doi.org/10.1094/CCHEM-03-16-0069-FI
9. Sozer N, Holopainen-Mantila U, Poutanen K. Traditional and
new food uses of pulses. Cereal Chem. 2017;94(1):66–73.
https://doi.org/10.1094/CCHEM-04-16-0082-FI
10. Shiba K, Negishi Y, Okada K, Nagao S. Chemical changes
during sponge-dough fermentation. Cereal Chem. 1990;
67(4):350–5.
11. Fernandez-Orozco R, Frias J, Muñoz R, Zielinski H, Piskula MK, Kozlowska H, Vidal-Valverde C. Fermentation as a
bio-process to obtain functional soybean flours. J Agric
Food Chem. 2007;55(22):8972–9.
https://doi.org/10.1021/jf071823b
ACKNOWLEDGEMENTS
The Global Institute for Food Security at the University of
Saskatchewan (Saskatoon, SK) and the Saskatchewan Ministry
of Agriculture Development Fund (ADF#: 2014-0283) provided
financial support for this research.
418
12. Lee IH, Hung YH, Chou CC. Solid-state fermentation with
fungi to enhance the antioxidative activity, total phenolic
and anthocyanin contents of black bean. Int J Food Microbiol. 2008;121(2):150–6.
https://doi.org/10.1016/j.ijfoodmicro.2007.09.008
July-September 2018 | Vol. 56 | No. 3
Food Technol. Biotechnol. 56 (3) 411-420 (2018)
13. Tavano OL. Protein hydrolysis using proteases: An important tool for food biotechnology. J Mol Catal B Enzym.
2013;90:1–11.
https://doi.org/10.1016/j.molcatb.2013.01.011
14. Hemalatha S, Platel, K, Srinivasan K. Influence of germination
and fermentation on bioaccessibility of zinc and iron from
food grains. Eur J Clin Nutr. 2006;61:342–8.
https://doi.org/10.1038/sj.ejcn.1602524
complexes. Food Res Int. 2010;43(2):489–95.
https://doi.org/10.1016/j.foodres.2009.07.022
27. Stone AK, Karalash A, Tyler RT, Warkentin TD, Nickerson MT.
Functional attributes of pea protein isolates prepared using
different extraction methods and cultivars. Food Res Int.
2015;76(Pt.1):31–8.
28. Systat v. 10 software, Systat Software Inc., San Jose, CA, USA;
2000.
15. Martín-Cabrejas MA, Sanfiz B, Vidal A, Mollá E, Esteban R,
López-Andréu FJ. Effect of fermentation and autoclaving on
dietary fiber fractions and antinutritional factors of beans
(Phaseolus vulgaris L.). J Agric Food Chem. 2004;52(2):261–
6.
https://doi.org/10.1021/jf034980t
29. Reyes-Moreno C, Cuevas-Rodríguez EO, Milán-Carrillo J,
Cárdenas-Valenzuela OG, Barrón-Hoyos J. Solid state fermentation process for producing chickpea (Cicer arietinum
L) tempeh flour. Physicochemical and nutritional characteristics of the product. J Sci Food Agric. 2004;84(3):271–8.
16. Mohite BV, Chaudhari GA, Ingale HS, Mahajan VN. Effect
of fermentation and processing on in vitro mineral estimation of selected fermented foods. Int Food Res J.
2013;20(3):1373–7.
17. de Vries MC, Vaughan EE, Kleerebezem M, de Vos WM. Lactobacillus plantarum-survival, functional and potential probiotic properties in the human intestinal tract. Int Dairy J.
2006:16(9);1018–28.
https://doi.org/10.1016/j.idairyj.2005.09.003.
30. Prinyawiwatkul W, Beuchat LR, McWatters KH, Phillips RD.
Fermented cowpea flour: Production and characterization
of selected physico-chemical properties. J Food Process
Preserv. 1996;20(4):265–84.
18. AOAC Official Method 925.10. Solids (total) and moisture
in flour. Gaithersburg, MD, USA: AOAC International; 2005.
19. AOAC Official Method 920.87. Protein (total) in flour. Gaithersburg, MD, USA: AOAC International; 2005.
20. AOAC Official Method 923.03. Ash of flour. Gaithersburg,
MD, USA: AOAC International; 2005.
21. AOAC Official Method 920.39. Fat (crude) or ether extraction
in animal feed. Gaithersburg, MD, USA: AOAC International; 2005.
22. Adler-Nissen J. Determination of the degree of hydrolysis of
food protein hydrolysates by trinitrobenzenesulfonic acid.
J Agric Food Chem. 1979;27(6):1256–62.
https://doi.org/10.1021/jf60226a042
23. Can Karaca A, Low N, Nickerson M. Emulsifying properties
of chickpea, faba bean, lentil and pea proteins produced by
isoelectric precipitation and salt extraction. Food Res Int.
2011;44(9):2742–50.
https://doi.org/10.1016/j.foodres.2011.06.012
24. Kato A, Nakai S. Hydrophobicity determined by a fluorescence probe method and its correlation with surface properties of proteins. Biochim Biophys Acta Protein Struct.
1980;624(1):13–20.
https://doi.org/10.1016/0005-2795(80)90220-2
25. Yasumatsu K, Sawada K, Moritaka S, Misaki M, Toda J, Wada
T, Ishii K. Whipping and emulsifying properties of soybean
products. Agric Biol Chem. 1972;36(5):719–27.
https://doi.org/10.1080/00021369.1972.10860321
26. Liu S, Elmer C, Low NH, Nickerson MT. Effect of pH on the
functional behaviour of pea protein isolate–gum Arabic
https://doi.org/10.1002/jsfa.1637
https://doi.org/10.1111/j.1745-4549.1996.tb00747.x
31. Chandra-Hioe MV, Wong CHM, Arcot J. The potential use of
fermented chickpea and faba bean flour as food ingredients. Plant Foods Hum Nutr. 2016;71(1):90–5.
https://doi.org/10.1007/s11130-016-0532-y
32. Akubor PI, Chukwu JK. Proximate composition and selected
functional properties of fermented and unfermented African oil bean (Pentaclethra macrophylla) seed flour. Plant
Foods Hum Nutr. 1999;54:227–38.
https://doi.org/10.1023/A:1008100930856
33. Ghribi AM, Gafsi IM, Sila A, Blecker C, Danthine S, Attia H, et
al. Effects of enzymatic hydrolysis on conformational and
functional properties of chickpea protein isolate. Food
Chem. 2015;187:322–30.
https://doi.org/10.1016/j.foodchem.2015.04.109
34. Zheng L, Zhao Y, Xiao C, Sun-Waterhouse D, Zhao M, Su G.
Mechanism of the discrepancy in the enzymatic hydrolysis efficiency between defatted peanut flour and peanut
protein isolate by Flavorzyme. Food Chem. 2015;168:100–6.
https://doi.org/10.1016/j.foodchem.2014.07.037
35. Liang HN, Tang CH. pH-dependent emulsifying properties
of pea [Pisum sativum (L.)] proteins. Food Hydrocoll. 2013;
33(2):309–19.
https://doi.org/10.1016/j.foodhyd.2013.04.005
36. Fasasi OS, Eleyinmi, AF, Oyarekua MA. Effect of some traditional processing operations on the functional properties of African breadfruit seed (Treculia africana) flour. LWT
- Food Sci Technol. 2007;40(3):513–9.
https://doi.org/10.1016/j.lwt.2005.11.009
37. Yu J, Ahmedna M, Goktepe I. Peanut protein concentrate:
Production and functional properties as affected by processing. Food Chem. 2007;103(1):121–9.
https://doi.org/10.1016/j.foodchem.2006.08.012
July-September 2018 | Vol. 56 | No. 3 419
B. ÇABUK et al. Fermented Pea Protein Functionality and Surface Properties
38. Xiao Y, Xing G, Rui X, Li W, Chen X, Jiang M, Dong M. Effect of solid-state fermentation with Cordyceps militaris
SN-18 on physicochemical and functional properties of
chickpea (Cicer arietinum L.) flour. LWT – Food Sci Technol.
2015;63(2):1317–24.
https://doi.org/10.1016/j.lwt.2015.04.046
39. Oloyede OO, James S, Ocheme OB, Chinma CE, Akpa VE.
Effects of fermentation time on the functional and pasting
420
properties of defatted Moringa oleifera seed flour. Food Sci
Nutr. 2016;4(1):89–95.
https://doi.org/10.1002/fsn3.262
40. Periago MJ, Vidal ML, Ros G, Rincón F, Martínez C, López
G, et al. Influence of enzymatic treatment on the nutritional and functional properties of pea flour. Food Chem.
1998;63(1):71–8.
https://doi.org/10.1016/S0308-8146(97)00199-4
July-September 2018 | Vol. 56 | No. 3