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Citation: Dhaouafi, J.; Nedjar, N.;
Jridi, M.; Romdhani, M.; Balti, R.
Extraction of Protein and Bioactive
Compounds from Mediterranean Red
Algae (Sphaerococcus coronopifolius and
Gelidium spinosum) Using Various
Innovative Pretreatment Strategies.
Foods 2024,13, 1362. https://doi.org/
10.3390/foods13091362
Received: 29 March 2024
Revised: 21 April 2024
Accepted: 23 April 2024
Published: 28 April 2024
Copyright: © 2024 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
foods
Article
Extraction of Protein and Bioactive Compounds from
Mediterranean Red Algae (Sphaerococcus coronopifolius and
Gelidium spinosum) Using Various Innovative
Pretreatment Strategies
Jihen Dhaouafi 1,2 , Naima Nedjar 2, Mourad Jridi 1, Montassar Romdhani 1,2 and Rafik Balti 3, *
1Laboratory of Functional Physiology and Valorization of Bioresources, Higher Institute of Biotechnology of
Beja, University of Jendouba, Avenue Habib Bourguiba, BP, 382, Beja 9000, Tunisia;
jihenedhaouefiii@gmail.com (J.D.); jridimourad@gmail.com (M.J.); romdhanimontassar4@gmail.com (M.R.)
2UMR Transfrontalière BioEcoAgro N◦1158, UniversitéLille, INRAE, UniversitéLiège, UPJV, YNCREA,
UniversitéArtois, UniversitéLittoral Côte d’Opale, ICV—Institut Charles Viollette, 59000 Lille, France;
naima.nedjar@univ-lille.fr
3UniversitéParis-Saclay, CentraleSupélec, Laboratoire de Génie des Procédés et Matériaux, Centre Européen
de Biotechnologie et de Bioéconomie (CEBB), 3 rue des Rouges Terres, 51110 Pomacle, France
*Correspondence: rafik.balti@centralesupelec.fr; Tel.: +33-3-5262-0512
Abstract: In this study, the release of proteins and other biomolecules into an aqueous media
from
two red
macroalgae (Sphaerococcus coronopifolius and Gelidium spinosum) was studied using
eight different cell disruption techniques. The contents of carbohydrates, pigments, and phenolic
compounds coextracted with proteins were quantified. In addition, morphological changes at
the cellular level in response to the different pretreatment methods were observed by an optical
microscope. Finally, the antioxidant capacity of obtained protein extracts was evaluated using three
in vitro
tests. For both S. coronopifolius and G. spinosum, ultrasonication for 60 min proved to be
the most effective technique for protein extraction, yielding values of 3.46
±
0.06 mg/g DW and
9.73 ±0.41 mg/g
DW, respectively. Furthermore, the highest total contents of phenolic compounds,
flavonoids, and carbohydrates were also recorded with the same method. However, the highest
pigment contents were found with ultrasonication for 15 min. Interestingly, relatively high antioxidant
activities like radical scavenging activity (31.57–65.16%), reducing power (0.51–1.70, OD at 700 nm),
and ferrous iron-chelating activity (28.76–61.37%) were exerted by the different protein extracts
whatever the pretreatment method applied. This antioxidant potency could be attributed to the
presence of polyphenolic compounds, pigments, and/or other bioactive substances in these extracts.
Among all the used techniques, ultrasonication pretreatment for 60 min appears to be the most
efficient method in terms of destroying the macroalgae cell wall and extracting the molecules of
interest, especially proteins. The protein fractions derived from the two red macroalgae under
these conditions were precipitated with ammonium sulfate, lyophilized, and their molecular weight
distribution was determined using SDS-PAGE. Our results showed that the major protein bands were
observed between 25 kDa and 60 kDa for S. coronopifolius and ranged from 20 kDa to 150 kDa for
G. spinosum. These findings indicated that ultrasonication for 60 min could be sufficient to disrupt
the algae cells for obtaining protein-rich extracts with promising biological properties, especially
antioxidant activity.
Keywords: red macroalgae; Sphaerococcus coronopifolius;Gelidium spinosum; eco-friendly cell disruption
methods; protein extraction; antioxidant activity
1. Introduction
Already struggling with seven billion humans, the planet’s resources are under enor-
mous and unbearable pressure due to population growth and the increase in food pro-
Foods 2024,13, 1362. https://doi.org/10.3390/foods13091362 https://www.mdpi.com/journal/foods
Foods 2024,13, 1362 2 of 24
duction during the past decades. The global demand for protein is expected to escalate,
exacerbating the need for more sustainable production systems to reduce the carbon foot-
print. These trends encourage the search for alternative protein sources, such as plants,
microorganisms, insects, seaweed, etc., to substitute livestock protein-based diets [
1
]. Par-
ticularly, marine macroalgae are significantly rich in protein and also contain a wide range
of nutritional and bioactive compounds, such as polysaccharides, pigments, minerals, vita-
mins, fatty acids, polyphenols, and peptides. These bioactive substances are associated with
several health benefits including antioxidant, antibacterial, antihypertensive, antidiabetic,
immunomodulatory, anti-inflammatory, and antiviral properties [2–5].
For many decades, macroalgae have been used as natural ingredients in traditional
medicine and cosmetic formulations due to their richness in biologically active compo-
nents [
6
]. In addition, macroalgae have long been consumed extensively as fresh human
food around the world, especially in Asian countries [
7
]. Furthermore, macroalgae biomass
has been discovered to be a high-quality protein-rich food, making it a sustainable alterna-
tive protein source to address current global security challenges [
8
]. It is well known that red
macroalgae have high protein levels, which sometimes exceed conventional protein sources
like soybeans, cereals, eggs, and fish [
9
]. Today, the macroalgae protein market is growing
continuously and is projected to reach $1.131 billion by 2027 [
10
]. Therefore, significant
developments are required to efficiently use macroalgae as a sustainable protein supply.
In terms of the profile of proteins and their derivatives, macroalgae contain significant
amounts of enzymes, glycoproteins, lectins, peptides, and amino acids, as well as phyco-
biliproteins, which are the major photosynthetic accessory pigments in cyanobacteria and
red algae [
11
]. The successful generation of bioactive peptides from the enzymatic hydroly-
sis of macroalgae proteins has been reported. Such peptides can produce a wide range of
bioactive effects and can be used as preservatives and functional ingredients to enhance
the sensory characteristics of food matrices. In fact, they are considered safer than some
synthetic additives, as they present with higher biofunctionality and biospecificity to target
cells, and are rarely associated with adverse effects [
12
]. Recently, seaweed fermentation has
demonstrated the potential to generate novel compounds, including bioactive peptides and
polysaccharides, processed phenolic compounds, enzymes, and organic acids. This biologi-
cal process of algal tissues and extracts can be used to create novel food and nutraceutical
products with high bioactivity and sensory qualities [
13
]. Red macroalgal biomass can
generate hydrocolloids, proteins, and other valuable unique biomolecules, and they are in
high demand in the food, cosmetic, medicine, and pharmaceutical industries [14].
However, the high structural complexity and rigidity of the algal cell wall is a major
obstacle to the efficient extraction of intracellular bioactive ingredients, principally pro-
teins and their derivates [
15
,
16
]. On the other hand, macroalgae proteins are attached to
non-protein components such as polysaccharides (agar, alginates, and carrageenan) and
polyphenols [
17
,
18
], which is considered among the key factors affecting protein extraction
efficiency [
19
]. Red macroalgae cell walls are composed of a combination of cellulose and
cellulose-like polysaccharides, forming the primary barrier to accessing and extracting
algal proteins and other intracellular compounds [20].
To extract the internal components of a macroalgae cell, it is necessary to first perform
a cell disruption operation that will break down the barrier and allow full access, thus
facilitating the release of cellular biomolecules. In recent years, numerous cell disruption
and protein extraction techniques have been investigated to enhance the extraction yield
and functional properties of macroalgae protein extracts [
8
]. Hence, several strategies
for breaking the cell wall of algae have been evaluated to recover different components,
including bead-beating [
21
,
22
], ultrasonication [
23
,
24
], microwave radiation [
25
], enzymatic
hydrolysis [
9
,
26
], cell homogenizing [
27
], and high-pressure cell disruption [
28
]. All these
extraction methods improve the mass transfer rate and increase the availability of protein
and other high value-added components [
29
]. Until now, there is no proper method to
apply to all macroalgae. The extraction approach must be assessed for each species, and the
Foods 2024,13, 1362 3 of 24
pros and cons must be evaluated regarding the biomass composition to ensure the optimal
protocol for obtaining protein-rich fractions.
Importantly, the methodology applied for protein extraction results in the simulta-
neous release of a wide range of bioactive compounds and therefore significantly affects
the chemical composition of the final extract. During the process of protein extraction
from Ulva sp. and Gracilaria sp., valuable phytochemicals such as phenolic compounds
were co-extracted, which increases the nutritional value of the final products [
30
]. In fact,
polyphenols are bioactive metabolites characteristic of marine macroalgae, and which are
very beneficial to human health, mainly as antioxidant agents [31,32].
The objectives of the present study were first to develop an efficient method for
obtaining protein-rich fractions from two red macroalgae Sphaerococcus coronopifolius (Gi-
gartinales, Sphaerococcaceae) and Geledium spinosum (Gelidiales, Rhodymeniophycidae). The
second objective was to quantify the biomolecules co-extracted concomitantly with pro-
teins and evaluate the antioxidant power of the obtained extracts for possible food and
nutraceutical purposes.
2. Materials and Methods
2.1. Chemicals and Reagents
All chemicals and reagents, including sulfuric acid (H
2
SO
4
), boric acid, hydrochloric
acid (HCl), ammonium sulfate (NH
4
)
2
SO
4
, t-butanol, tris-HCl buffer, Bradford reagent,
bovine serum albumin, glucose, Laemmli buffer, glycine, sodium dodecyl sulfate (SDS),
phenol reagent, acetone, gallic acid, Folin–Ciocalteu reagent, sodium carbonate (Na
2
CO
3
),
sodium nitrite (NaNO
2
), sodium hydroxide (NaOH), aluminum chloride (AlCl
3
), quercetin,
ethanol, 1,1-diphenyl-2-picrylhydrazyl (DPPH), 3,4-dihydro-6-hydroxy-2,5,7,8-tetramethyl-
2H-1-benzopyran-2-carboxylic acid (Trolox), butylated hydroxyanisole (BHA), ferric chlo-
ride (FeCl
2
), ferrozine potassium ferricyanide, trichloroacetic acid (TCA), and 3.5 kDa
MWCO dialysis tubing, were obtained from Sigma-Aldrich (Saint-Quentin-Fallavier, France).
2.2. Collection and Preparation of Algal Materials
The S. coronopifolius species was collected in January and February 2019 from Men-
zel Abderrahmane city, Bizerte, located in the North-West of Tunisia (37
◦
13
′
48
′′
N and
9◦51′36′′ E
). Similarly, the G. spinosum species was collected in the same period from the
Monastir region, which is in the East of Tunisia (35
◦
46
′
N and 10
◦
49
′
E). The collected
samples were packed in polyethylene bags and transported to the laboratory within 2 h.
Upon arrival, the red macroalgal biomass was rinsed with tap water to remove epiphytes,
sediment, and potential contaminants. Fresh macroalgae were subsequently dried in a dark
room at ambient temperature for a few weeks. The dried samples were then ground to a
fine powder using a blender (Knife Mill Grindomix GM 200, Retsch, Haan, Germany) and
kept in airtight glass jars.
2.3. Determination of Crude Protein
The organic nitrogen content was quantified using the Kjeldahl procedure [
33
]. A
total of 0.5 g of dried macroalgae was digested in a Kjeldaltherm
®
block digestion unit
(Gerhardt, Königswinter, Germany) in 15 mL of concentrated H
2
SO
4
and one tablet (2.5 g)
of Kjeldahl catalyst for 1 h. Digestion was completed on the production of a clear solution.
Steam distillation after Kjeldahl digestion was carried out in a Vapodest
®
33 unit (Gerhardt,
Germany). The distillate was collected in an Erlenmeyer flask containing 15 mL of 4%
(v/v) aqueous boric acid solution brought to a final volume of 50 mL. The titration was
performed manually using a standard HCl solution (0.1 M). Nitrogen content, given in g of
nitrogen per 100 g of the sample, was calculated using the following numerical equation
(Equation (1)):
Nitrogen content(%) = 1.4007 ×Vsam ple (mL)−Vblank (mL)×0.1
Wei ght o f sample (g)(1)
Foods 2024,13, 1362 4 of 24
where V
blank
and V
sample
are the volumes of hydrochloric acid consumed during the
titration of the reagent blank and sample, respectively. A factor of 6.25 was used to convert
the nitrogen value to protein [34]. All measurements were performed in triplicate.
2.4. Pretreatment Methods for Red Macroalgae Cell Disruption and Protein Extraction
The different pretreatment techniques applied to red macroalgae biomasses to disrupt
the cell wall and protein extraction are presented in Figure 1. All pretreatments were carried
out in triplicate.
Foods2024,13,xFORPEERREVIEW4of25
wasperformedmanuallyusingastandardHClsolution(0.1M).Nitrogencontent,given
ingofnitrogenper100gofthesample,wascalculatedusingthefollowingnumerical
equation(Equation(1)):
𝑁𝑖𝑡𝑟𝑜𝑔𝑒𝑛 𝑐𝑜𝑛𝑡𝑒𝑛𝑡 %1.4007
.
(1)
whereV
blank
andV
sample
arethevolumesofhydrochloricacidconsumedduringthetitra-
tionofthereagentblankandsample,respectively.Afactorof6.25wasusedtoconvertthe
nitrogenvaluetoprotein[34].Allmeasurementswereperformedintriplicate.
2.4.PretreatmentMethodsforRedMacroalgaeCellDisruptionandProteinExtraction
Thedifferentpretreatmenttechniquesappliedtoredmacroalgaebiomassestodis-
ruptthecellwallandproteinextractionarepresentedinFigure1.Allpretreatmentswere
carriedoutintriplicate.
Figure1.Aschematicofdifferentcelldisruptionandproteinextractionmethodsappliedtotwored
macroalgae(S.coronopifoliusandG.spinosum).
2.4.1.Control
Onegramofredseaweedpowderwasdispersedin100mLofdistilledwaterfor2h,
andthesupernatantwasthenrecoveredbycentrifugationat6000×gfor20minat4°C.
Thispreparationservedasacontroltocomparewiththeotherpretreatmentmethods.
2.4.2.Ultrasonication(US)
Redmacroalgaesamplesweresubjectedtoultrasonicationpretreatmentusinganul-
trasoniccelldisruptor(OmniSonicRuptor4000,Kennesaw,GA,USA),followingthe
methodologyofSafietal.[35]withslightmodificationsasdescribedbyMaliketal.[36].
Onegramofthemacroalgaepowderwasdissolvedin100mLofdistilledwaterandthen
ultrasonicallytreatedatafrequencyof20kHzandapowerof200Wwithamplitudesset
at90%(withapulsedurationofon-time15sandoff-time15s).Thesolublefractionswere
collectedbycentrifugationat6000×gfor20minat4°C.
2.4.3.ManualGrinding(MG)
Drymacroalgaeweremanuallygroundusingamortarfor10min,andthen1gwas
dispersedin100mLofdistilledwaterfor2h.Thesupernatantwasrecoveredbycentrif-
ugationat6000×gfor20minat4°C.
2.4.4.CombinationofManualGrindingandUltrasonicationfor30min(MG-US30)
Theredmacroalgaepowdersamplesweremanuallygroundseparatelyusingamor-
tarfor10min,andthen1gofeachsamplewasdissolvedin100mLofdistilledwater.The
Figure 1. A schematic of different cell disruption and protein extraction methods applied to two red
macroalgae (S. coronopifolius and G. spinosum).
2.4.1. Control
One gram of red seaweed powder was dispersed in 100 mL of distilled water for 2 h,
and the supernatant was then recovered by centrifugation at 6000
×
gfor 20 min at 4
◦
C.
This preparation served as a control to compare with the other pretreatment methods.
2.4.2. Ultrasonication (US)
Red macroalgae samples were subjected to ultrasonication pretreatment using an
ultrasonic cell disruptor (Omni Sonic Ruptor 4000, Kennesaw, GA, USA), following the
methodology of Safi et al. [
35
] with slight modifications as described by Malik et al. [
36
].
One gram of the macroalgae powder was dissolved in 100 mL of distilled water and then
ultrasonically treated at a frequency of 20 kHz and a power of 200 W with amplitudes set
at 90% (with a pulse duration of on-time 15 s and off-time 15 s). The soluble fractions were
collected by centrifugation at 6000×gfor 20 min at 4 ◦C.
2.4.3. Manual Grinding (MG)
Dry macroalgae were manually ground using a mortar for 10 min, and then 1 g
was dispersed in 100 mL of distilled water for 2 h. The supernatant was recovered by
centrifugation at 6000×gfor 20 min at 4 ◦C.
2.4.4. Combination of Manual Grinding and Ultrasonication for 30 min (MG-US30)
The red macroalgae powder samples were manually ground separately using a mortar
for 10 min, and then 1 g of each sample was dissolved in 100 mL of distilled water. The
samples were placed in a beaker before ultrasound pretreatment. The pretreatment was
performed at a frequency of 20 kHz and a power output of 200 W (amplitudes of 90%) for
30 min (pulse duration of on-time 15 s and off-time 15 s). The mixtures were centrifuged at
6000×gfor 20 min at 4 ◦C and the supernatants were collected for biochemical analyses.
2.4.5. Three-Phase Partitioning (TPP)
The TPP technique was carried out in accordance with the previously published
method of Chia et al. [
37
] with minor modifications. Before combining with salt solution,
Foods 2024,13, 1362 5 of 24
1 wt%
of red macroalgae biomass was first dissolved in deionized water. Then, 5 mL of pure
t-butanol and 5 mL of 30% saturation (NH
4
)
2
SO
4
were added. The mixture was agitated
using a magnetic stirrer at 200 rpm for 1 h and was allowed to separate for
30 min
at room
temperature. The three phases were observed and separated carefully by pipetting them
out from the beaker. The intermediate protein precipitate was dissolved in an appropriate
amount of tris-HCl buffer and analyzed for soluble protein and other biomolecule contents.
2.4.6. Ultrasound-Assisted Three-Phase Partitioning (UATPP)
The UATPP procedure was carried out using an ultrasonic cell disruptor (Omni Sonic
Ruptor 4000, Kennesaw, GA, USA). For the comparison study, the initial parameters of
UATPP such as the working volume, saturation of salt solution, and weight of biomass
were similar to the TPP method. The preparation and mixing procedure of UATPP was
similar to TPP [37]. The ultrasonic treatment of the mixture was operated at 20 kHz and a
power output of 200 W with an amplitude of 90% for 30 min (pulse duration of on-time
15 s
and off-time 15 s). The treated solution was then taken out and allowed to separate for
30 min at room temperature.
2.4.7. Combination of Freeze Drying with Ultrasonication for 30 Min (FD-US30)
Frozen macroalgae pastes (10 g) were directly introduced to a Lyovapor™ L-200
freeze dryer (BÜCHI Labortechnik AG, Flawil, Switzerland). The pressure was reduced to
0.0010 bar
and the temperature was further decreased to
−
80
◦
C and freeze-drying was
conducted under vacuum for 48 h. Then, one gram of each freeze-dried powder was
dissolved in 100 mL of distilled water to ensure optimal sample homogeneity. Finally, the
ultrasound treatment was carried out at 20 kHz for 30 min with a power output of 200 W
(pulse duration of on-time 15 s and off-time 15 s), followed by centrifugation at 6000
×
gfor
20 min at 4 ◦C for the recovery of the soluble phase.
2.4.8. French Press (FP)
The dried macroalgae biomasses were dispersed in distilled water at 10 g/L and
vigorously mixed in a vortex (Vortex 3, IKA, Staufen, Germany) to ensure the homogeneity
of the macroalgal samples. A high-pressure homogenization method using a One-Shot Cell
Disrupter (Constant Systems Ltd., Warwickshire, UK) was applied to the red macroalgae
suspension in one pass at a pressure of 2700 bar [
35
]. The working volume in this study
was fixed at 8 mL. In this process, the macroalgal cells are forced to flow through a very
small orifice under high-pressure conditions, and, as a result, they could be disrupted
by synergistic mechanical effects, such as cavitation, turbulence, and shear stress [
38
].
Water-soluble protein extracts were obtained by centrifugation (6000×g, 20 min, 4 ◦C).
2.4.9. Bead-Beating (BB)
Red macroalgae cells were disrupted with the bead-beating method according to
Suarez Garcia et al. [
39
] with minor modifications. Macroalgae aqueous suspensions
(
10 g/L
) were transferred to MN Bead Tubes Type C containing 1–3 mm corundum beads
(Macherey-Nagel, Düren, Germany). The samples were subjected to intense mixing using a
Fastprep-24 5G
TM
bead beater (MP Biomedicals, Santa Ana, CA, USA) for
three cycles
at
6 m/s
for 60 s each. A cooling phase of 2 min in between cycles was fixed to avoid overheat-
ing and the decomposition of metabolites (total extraction time
≈
10 min,
T = 25 ◦C±2◦C
).
2.4.10. Mass Extraction Yield Calculation
All extracts obtained were freeze-dried and weighed. In order to assess the extraction
performances of the evaluated cell disruption methods, the mass extraction yield was
calculated and used as an indicator of the effectiveness of biomass pretreatments. Mass
extraction yields (Y) were calculated according to Equation (2):
Y(%) = w eight o f f re eze −dried extract (g)
weight o f red macroalgae powder (g)×100 (2)
Foods 2024,13, 1362 6 of 24
All calculations are conducted on a dry-weight (DW) basis.
2.5. Optical Microscopic Observation
Suspensions of the untreated and treated red macroalgae were analyzed with an
Optika B-190TB light microscope (OPTIKA, Ponteranica, Italy). Digital images were taken
with a 3.1-megapixel digital color microphotography camera C-B3A (OPTIKA, Ponteranica,
Italy). Acquired images were analyzed and processed using the OPTIKA vision lite 2.1
Software (OPTIKA, Ponteranica, Italy).
2.6. Estimation of Total Soluble Protein
The soluble protein concentration was determined using the dye-binding Bradford
assay [
40
]. Briefly, 0.25 mL of the sample was mixed with 2.5 mL of Bradford reagent (1:50
v/v) and incubated at room temperature for 15 min. The absorbance of the samples was
then measured at 595 nm using a UV–VIS spectrophotometer (SpectraMax
®
ABS Plus, San
Jose, CA, USA) and the soluble protein concentration was calculated using the standard
calibration of bovine serum albumin. The measurements were performed in triplicate.
2.7. Protein Profile by SDS-PAGE
Protein-denaturing sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-
PAGE) of each extract was performed according to the Laemmli method [
41
]. Briefly, the
extracted proteins were solubilized in ultrapure water at a concentration of 2 mg/mL and
diluted in Laemmli buffer containing
β
-mercaptoethanol and SDS before heating at 95
◦
C
for 10 min. Then, 25
µ
L samples and 4
µ
L molecular mass marker solutions (Precision
Plus Protein
TM
Standards, Bio-Rad, Marnes-la-Coquette, France) were deposited on an
Any-kD
TM
Mini-Protean
®
TGX Stain-free
TM
gel (Bio-Rad). Protein migration was allowed
to occur for 1 h in a buffer containing Tris-base (25 mM), glycine (0.19 mM), and SDS
(
3.5 mM
) under a constant voltage of 120 V. Thereafter, fluorescence generated by the
reaction between the gel-trihalo compounds and tryptophan residues of the proteins was
revealed after 5 min of activation time with the Gel Doc
TM
XR+ system and Image Lab 6.1.0
software (Bio-Rad).
2.8. Quantification of Co-Extracted Compounds
Following the different pretreatment methods applied, other intracellular molecules
derived from red macroalgae, such as polyphenols, carbohydrates, and pigments, could
be released simultaneously with the proteins. Indeed, the quantification of the content of
these co-extracted compounds in the final protein extract was carried out.
2.8.1. Total Carbohydrate Analysis
Total carbohydrate content was evaluated by the colorimetric method after adding
phenol and sulfuric acid as described by Dubois et al. [
42
]. Typically, 500
µ
L of the sample
was introduced at the bottom of a 15 mL polypropylene falcon tube. Then, 500
µ
L of
phenol solution (50 g/L) was added to 2.5 mL of sulfuric acid (>96%). After 30 min of
incubation at ambient temperature, the absorbance at 485 nm was measured using a UV–
VIS spectrophotometer. The calibration curve was made using D-glucose and each sample
was analyzed in triplicate.
2.8.2. Pigment Analysis
The quantitative estimation of chlorophyll a and chlorophyll b was carried out with
the method of Arnon [
43
], while carotenoids were determined by following Kirk and
Allen [
44
]. Acetone (80%) was used as the extractant solvent and the absorbance of the
extracted solution was measured using a UV–VIS spectrophotometer at the wavelengths of
480, 645, and 663 nm. The chlorophyll and carotenoid contents were calculated using the
following formulas (Equations (3)–(5)) in triplicate for each pretreatment and expressed as
mg/g of DW.
Foods 2024,13, 1362 7 of 24
Chloro phyll a (mg/g DW)=(12.7 −A663 )−(2.69 ×A645)×Fin al volum e o f ex tract (mL)
Weight o f pretreated macroalgae (g)(3)
Chloro phyll b (mg/g DW)=(22.9 −A645 )−(4.86 ×A663)×Final vol ume o f extrac t (mL)
Weight o f pretreated macroalgae (g)(4)
Carotenoid (mg/g DW)=4×A480 ×Final vol ume o f ex tract (mL)
Weight o f pretreated macroalgae (g)(5)
where A
480
, A
645
, and A
663
are the absorbances at wavelengths of 480, 645, and
663 nm
,
respectively. Each sample was analyzed in triplicate.
2.8.3. Quantitative Polyphenol Analysis
Total phenolic content (TPC) was measured with a colorimetric assay using the Folin–
Ciocalteu phenol reagent [
45
] and using gallic acid as the standard phenolic compound.
Briefly, 50
µ
L of the sample was added to 120
µ
L of Folin–Ciocalteu reagent and 2 mL
of distilled water and mixed thoroughly for 5 min. Then, 375
µ
L of 10% (w/v) sodium
carbonate was added and the mixture was allowed to stand for 2 h at room temperature.
The absorbance was measured at 765 nm using a UV–VIS spectrophotometer. TPC was
expressed as mg gallic acid equivalents (GAE)/g DW. The test was carried out in triplicate.
Total flavonoid content (TFC) was determined using the aluminum chloride colorimet-
ric method [
46
] and using quercetin as the condensed flavonoid standard. Briefly, 400
µ
L of
the sample was mixed with 120
µ
L of 5% (w/v) NaNO
2
and 120
µ
L of 10% (w/v) AlCl
3
was
added. After 6 min, 800
µ
L of NaOH (1 M) was added and the absorbance of the mixture
was measured at 510 nm. TFC was expressed as mg quercetin equivalents (QE)/g DW. The
test was carried out in triplicate.
2.9. Antioxidant Assays
The antioxidant activity of obtained extracts was determined by different
in vitro
methods,
such as the DPPH free radical scavenging, ferrous ion-chelating ability, and reducing power
assays. All tests were carried out in triplicate and average values were considered.
2.9.1. DPPH Free Radical Scavenging Assay
DPPH free radical scavenging activity was measured using the method described
by Bersuder et al. [
47
]. Briefly, a 500
µ
L test sample was mixed with 375
µ
L of 99.5%
ethanol and 125
µ
L of 0.02 mM DPPH ethanol solution. This mixture was shaken then kept
in the dark at room temperature for 30 min before measuring its absorbance at
517 nm
.
DPPH radical scavenging activity was calculated according to the following equation
(Equation (6)):
DPPH radical-scavenging activity (%)=1−A517 of sample
A517 of control ×100 (6)
The control was conducted in the same manner, except that distilled water was used
instead of the sample. Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) at
1 mg/mL was used as the standard.
2.9.2. Ferrous Ion-Chelating Ability Assay
The chelating activity on Fe
2+
was determined using the method described by Decker
and Welch [
48
]. An aliquot of 100
µ
L of the sample solution was mixed with 50
µ
L of 2 mM
FeCl
2
and 450
µ
L of distilled water. The mixture was then reacted with 200
µ
L of 5 mM
ferrozine for 10 min at room temperature. The absorbance of the Fe
2+
–ferrozine complex
with red or violet color was read at 562 nm. The control was prepared in the same manner
except that distilled water was used instead of the sample. Butylated hydroxyanisole (BHA)
Foods 2024,13, 1362 8 of 24
tested at 1 mg/mL was used as a reference. Chelating activity was then calculated as
follows (Equation (7)):
Chelating activity (%) = 1−A562 of sample
A562 of control ×100 (7)
2.9.3. Reducing Power Assay
The ability of the samples to reduce iron (III) was determined according to the method
of Yildirim et al. [
49
] with slight modifications. An aliquot of 0.5 mL of each sample was
mixed with 1.25 mL of 0.2 M phosphate buffer (pH 6.6) and 1.25 mL of 1% potassium
ferricyanide. The mixture was incubated at 50
◦
C for 30 min, followed by the addition of
1.25 mL of 10% (w/v) trichloroacetic acid. The mixture was then centrifuged at 3000
×
g
for 10 min. Finally, 1.25 mL of the supernatant was mixed with 1.25 mL of distilled water
and 0.25 mL of 0.1% (w/v) ferric chloride. After a 10 min reaction, the absorbance of the
resulting solution was measured at 700 nm. Increased absorbance of the reaction mixture
indicated increased reducing power. BHA (1 mg/mL) was used as a positive control.
2.10. Statistical Analysis
Each extract was independently produced in triplicate. All the experimental tests
were performed at least in triplicate. Values are expressed as mean
±
standard deviation
(SD). Data analysis was carried out with the GraphPad Prism 9.0 software (GraphPad
Software, Inc., San Diego, CA, USA) using a one-way ANOVA analysis followed by post
hoc Tukey’s honestly significant difference (HSD) tests. Differences were considered
statistically significant at p< 0.05.
3. Results and Discussion
3.1. Total Protein Contents of S. coronopifolius and G. spinosum
The total protein contents of the raw materials were determined using the Kjeldahl
method by converting elemental total nitrogen into protein percentage. The protein
content of S. coronopifolius was 17.62
±
0.20%. Consistent findings were reported by
Patarra et al. [50]
for the same species, recording a protein content of 19.56%. On the other
hand, G. spinosum exhibited a protein content of 22.71
±
0.31%. This value appeared slightly
lower compared to the results presented by Ben Said et al. [
51
], who reported a protein
value of 29% for G. spinosum harvested from the Monastir coasts (Tunisia). Generally, the
content of proteins from macroalgae is strongly influenced by the geographical origin,
species, and season [
52
,
53
]. In fact, higher levels of proteins are reported in winter [
54
].
For example, the protein content of G. spinosum varied between 18 and 29% in April and
January, respectively [
51
]. Also, our results are comparable to those reported for other
red macroalgae species such as Gracilaria edulis (25.3%) [
55
], Palmaria palmata (15.3%) [
56
],
Chondracanthus chamissoi (17.6%) [57], and Pyropia orbicularis (13.6%) [58].
Despite the high nutritional value of algal proteins [
9
], their availability is limited
by the rigidity of the algal membrane. For this reason, our study aimed to evaluate the
proteins released in the aqueous media after different cell disruptions. Compounds such
as polyphenolic compounds, carbohydrates, and pigments are extracted simultaneously
with the proteins during the different treatments. The results are based not only on the
mechanical rigidity of each macroalga cell wall but also on its chemical properties.
3.2. Optical Scanning Microscopy Morphology Observation of Macroalgae Cells
To better interpret the efficacy of each pretreatment, morphological changes of the
S. coronopifolius
and G. spinosum control and pretreated cells were observed using an optical
microscope. The results are displayed in Figure 2.
Foods 2024,13, 1362 9 of 24
Foods2024,13,xFORPEERREVIEW9of25
Despitethehighnutritionalvalueofalgalproteins[9],theiravailabilityislimitedby
therigidityofthealgalmembrane.Forthisreason,ourstudyaimedtoevaluatethepro-
teinsreleasedintheaqueousmediaafterdifferentcelldisruptions.Compoundssuchas
polyphenoliccompounds,carbohydrates,andpigmentsareextractedsimultaneously
withtheproteinsduringthedifferenttreatments.Theresultsarebasednotonlyonthe
mechanicalrigidityofeachmacroalgacellwallbutalsoonitschemicalproperties.
3.2.OpticalScanningMicroscopyMorphologyObservationofMacroalgaeCells
Tobeerinterprettheefficacyofeachpretreatment,morphologicalchangesoftheS.
coronopifoliusandG.spinosumcontrolandpretreatedcellswereobservedusinganoptical
microscope.TheresultsaredisplayedinFigure2.
Figure2.CellmorphologyobservationofS.coronopifolius(1)andG.spinosum(2)under40×magni-
fication:untreatedcells(A),USfor15min(B),USfor30min(C),USfor60min(D),MG(E),MG-
US30(F),TPP(G),UATPP(H),FD-US30(I),BB(J),andFP(K).Scalebar,10mm.
Figure2(1A,2A)showstheimageofalgalcellsofS.coronopifoliusandG.spinosum,
respectively,beforebeingsubjectedtosonication(untreatedcells).RegardingS.coronopi‐
folius,thecellsaresphericalandhaveintactintracellularcompartments(2.00–3.5mmin
Figure 2. Cell morphology observation of S. coronopifolius (1) and G. spinosum (2) under 40
×
magnifi-
cation: untreated cells (A), US for 15 min (B), US for 30 min (C), US for 60 min (D), MG (E), MG-US30
(F), TPP (G), UATPP (H), FD-US30 (I), BB (J), and FP (K). Scale bar, 10 mm.
Figure 2(1A,2A) shows the image of algal cells of S. coronopifolius and G. spinosum,
respectively, before being subjected to sonication (untreated cells). Regarding S. coronopi-
folius, the cells are spherical and have intact intracellular compartments (2.00–3.5 mm in
diameter). In contrast, the untreated cells of G. spinosum have an ellipsoidal shape, with the
intracellular compartments intact within the cell (4.00–10.00 mm in diameter).
Figure 2(1B–K,2B–K) shows the morphology of the cells after different pretreatments,
where the cells were subjected to the cell disintegration treatments. As can be observed
from Figure 2(1B,C,2B,C), the cells treated with US for 30 and 15 min were not disrupted
significantly. As shown in Figure 2(1D,2D), the cells were fragmented after US for 60 min.
This proves that sonication is important for breaking the cell wall to facilitate the release of
protein and other compounds. Acoustic cavitation can effectively destroy the gas vacuoles
that control the floating of algal cells in water [
59
,
60
]. Therefore, sonication should reduce
the suspension of algal cells and accelerate their sedimentation.
On the other hand, when pretreated with FD-US30 (Figure 2(1I) and Figure 2(2I) for
S. coronopifolius
and G. spinosum, respectively), the majority of the cells were broken, while
some of them remained intact and cells maintained their globular form. Combination
Foods 2024,13, 1362 10 of 24
treatment, such as MG-US30, showed that the cells are similar to the cells observed from
US30 only (Figure 2(1F) and Figure 2(2F) for S. coronopifolius and G. spinosum, respectively).
In the case of BB treatment (Figure 2(1J) and Figure 2(2J) for S. coronopifolius and
G. spinosum, respectively), the cells were partially disrupted with the presence of some
organelles that were liberated, and many cells were quiet in this pretreatment. High-
pressure cell disruption using FP (Figure 2(1K) and Figure 2(2K) for S. coronopifolius and
G. spinosum
, respectively) was relatively efficient for the two red macroalgae; the majority
of cells were broken while some of them remained intact.
3.3. Mass Extraction Yield
The mass recovery yields achieved through various disruption methods are depicted
in Figure 3. The mass extraction yields ranged from 13.47
±
0.003% DW to
60.19 ±0.007%
DW and from 19.86
±
0.07% DW to 47.43
±
0.14% DW for S. coronopifolius and G. spinosum,
respectively. Among all tested techniques, the US 60 min cell disruption appears to be the
best technique (p< 0.05) for both S. coronopifolius (60.19% DW) and G. spinosum (47.44% DW)
compared to the untreated cells (30.53
±
0.07% DW and 37.68
±
0.07% for S. coronopifolius
and G. spinosum, respectively). These findings showed the necessity of effective cell dis-
ruption to maximize the release of intracellular compounds from the two red macroalgae.
Foods2024,13,xFORPEERREVIEW11of25
Figure3.MassextractionyieldsofS.coronopifolius(A)andG.spinosum(B)pretreatedwithdifferent
celldisruptiontechniques.a,b,c,d,e,f,g,handk:differentleersmeansignificantdifferences
betweenextracts(p<0.05).Resultsareexpressedasaverage±standarddeviation(SD)(n=3).US,
ultrasonication;MG,manualgrinding;TPP,three-phasepartitioning;UATPP,ultrasonication-as-
sistedthree-phasepartitioning;MG-US30,manualgrinding+ultrasonicationfor30min;FD-US30,
freezedrying+ultrasonicationfor30min;FP,Frenchpress;andBB,bead-beating.
USpowerdemonstratedasignificanteffect(p<0.05)ontheincreaseoftheextraction
yieldsforS.coronopifoliusandG.spinosum(Figure3),unlikethoseobtainedwithoutultra-
soundpretreatment(0W/L)suchasintheuntreatedcells,TPP,andUATPP.Thisindicated
thepositiveeffectoftheultrasonicationmethodontheextractionofintracellularcom-
pounds.Sonicationdurationisalsoaveryimportantparameterthatdeterminesthetreat-
mentefficiency.ByapplyingalongerextractiontimeusingUSfor60min,themacroalgae
biomassesaresubjectedtoprolongedexposuretoultrasonicwaveswhichgeneratean
implosionofcavitationbubblesinthefluid.Theseconditionsareeasilycapableofdis-
ruptingcellwallsandmembranesandreleasingintracellularcompoundsinanefficacious
andrapidmanner.Intracellularcompoundssuchasproteins,polysaccharides,lipids,vit-
amins,minerals,andantioxidantscantherebybeeffectivelyextractedusingpowerultra-
sonics.Inviewofthesehighyields,USappearstobeasuitableprimaryextractionmethod
forthedisintegrationofcellwalls,therebyfacilitatingthereleaseoftargetcompounds
fromredmacroalgae.
Manyresearchershavebeeninterestedinultrasonicationextractionsinceitprovides
agreaterbiomoleculeextractionyield[61,62].Theextractionyieldofbioactivechemicals
(laminarin,fucose,uronicacid,andphenolics)fromthebrownalgaeA.nodosumisim-
provedbyultrasonication-assistedextraction[26,63].Previousresearchhasshownthat
molecularvibrationsgeneratedbyhigh-powerultrasonicwavescanhelpdisruptchemi-
calinteractionsbetweenmoleculesandfacilitatemolecularmobility[64].Usually,the
compositionofthecellwallmustalsobeconsideredsinceitconstitutesadetermining
factorthatcansignificantlyinfluencetheefficiencyoftheextraction[65].
Figure 3. Mass extraction yields of S. coronopifolius (A) and G. spinosum (B) pretreated with dif-
ferent cell disruption techniques. a–k: different letters mean significant differences between ex-
tracts (p< 0.05). Results are expressed as average
±
standard deviation (SD) (n= 3). US, ultra-
sonication; MG, manual grinding; TPP, three-phase partitioning; UATPP, ultrasonication-assisted
three-phase partitioning; MG-US30, manual grinding + ultrasonication for 30 min; FD-US30, freeze
drying + ultrasonication for 30 min; FP, French press; and BB, bead-beating.
US power demonstrated a significant effect (p< 0.05) on the increase of the extrac-
tion yields for S. coronopifolius and G. spinosum (Figure 3), unlike those obtained without
ultrasound pretreatment (0 W/L) such as in the untreated cells, TPP, and UATPP. This
indicated the positive effect of the ultrasonication method on the extraction of intracellular
Foods 2024,13, 1362 11 of 24
compounds. Sonication duration is also a very important parameter that determines the
treatment efficiency. By applying a longer extraction time using US for 60 min, the macroal-
gae biomasses are subjected to prolonged exposure to ultrasonic waves which generate an
implosion of cavitation bubbles in the fluid. These conditions are easily capable of disrupt-
ing cell walls and membranes and releasing intracellular compounds in an efficacious and
rapid manner. Intracellular compounds such as proteins, polysaccharides, lipids, vitamins,
minerals, and antioxidants can thereby be effectively extracted using power ultrasonics.
In view of these high yields, US appears to be a suitable primary extraction method for
the disintegration of cell walls, thereby facilitating the release of target compounds from
red macroalgae.
Many researchers have been interested in ultrasonication extraction since it provides
a greater biomolecule extraction yield [
61
,
62
]. The extraction yield of bioactive chemicals
(laminarin, fucose, uronic acid, and phenolics) from the brown algae A. nodosum is improved
by ultrasonication-assisted extraction [
26
,
63
]. Previous research has shown that molecular
vibrations generated by high-power ultrasonic waves can help disrupt chemical interactions
between molecules and facilitate molecular mobility [
64
]. Usually, the composition of
the cell wall must also be considered since it constitutes a determining factor that can
significantly influence the efficiency of the extraction [65].
The combined treatment with FD-US30 showed a significantly (p< 0.05) lower yield
(44.69% DW and 39.88% DW for S. coronopifolius and G. spinosum, respectively) than the
US 30 min alone (55.65% DW and 42.14% DW for S. coronopifolius and G. spinosum, respec-
tively). It has been demonstrated that the freeze-drying process, which preserves biological
materials well, makes protein extraction more difficult for particular algae species [
66
].
Furthermore, after freeze-drying, the cells become more aggregated, reducing the contact
surface with the extracting solvent and potentially affecting the cell wall integrity [
67
,
68
].
The obtained results were in line with those of Barbino et al. [
69
] for the two macroalgae
Sargassum vulgare and Chnoospora minima.
The MG-US30 combined pretreatment is gaining importance due to it resulting in higher
yields than US 30 min alone (p< 0.05) (49.84% DW and 43.54% DW for
S. coronopifolius
and
G. spinosum, respectively). For FP pretreatment, the yield was in the order of 39.53% DW
and 45.79% DW for S. coronopifolius and G. spinosum, respectively. On the other hand, the
lowest yields were obtained for the TPP and UATPP methods (13.47% DW and 16.89% DW
for S. coronopifolius and 19.86% DW and 22.07% DW for G. spinosum, respectively) due to the
rigidity of the cell walls of the studied macroalgae species. Aqueous maceration alone has
been shown to generally have a relatively low extraction yield compared to other alternative
liquid extraction systems [37].
3.4. Release of Soluble Protein
Protein extraction yields from algae sources are significantly higher than those from
protein-rich crops such as lupin, soybean, and legumes [
70
]. Therefore, significant develop-
ments are required to efficiently exploit marine macroalgae as an alternative sustainable
protein source. This study compared the efficiency of different protein extraction techniques
from S. coronopifolius and G. spinosum. Figure 4shows the protein contents obtained by
the different pretreatment methods of biomass and untreated cells. The extracted protein
content was very low in untreated cells for S. coronopifolius (0.24 mg/g DW) and it is in the
order of 4.05 mg/g DW for G. spinosum, which confirms the need for efficient cell disruption
to enhance protein release. These results were in line with expectations as the extraction
was carried out in water. However, the osmosis phenomenon was not strongly effective for
red macroalgae, which are known to have rigid cell walls [37].
Foods 2024,13, 1362 12 of 24
Foods2024,13,xFORPEERREVIEW13of25
Figure4.ProteincontentsofS.coronopifolius(A)andG.spinosum(B)extractsusingdifferenttech-
niquesofcelldisruption.a,b,c,d,e,andf:differentleersmeansignificantdifferencesbetween
extracts(p<0.05).Resultsareexpressedasaverage±standarddeviation(SD)(n=3).US,ultrasoni-
cation;MG,manualgrinding;TPP,three-phasepartitioning;UATPP,ultrasonication-assistedthree-
phasepartitioning;MG-US30,manualgrinding+ultrasonicationfor30min;FD-US30,freezedrying
+ultrasonicationfor30min;FP,Frenchpress;andBB,bead-beating.
Forbothmacroalgae,theproteinlevelsafter10minofMGhavenosignificantdiffer-
ence(p>0.05)comparedtountreatedcells.Ontheotherhand,thecomparisonofother
techniquesrevealedthesuperiorityofUS60min,followedbytheFPmethod.Thetotal
quantityofextractableproteinfromS.coronopifoliuswas3.43±0.06mg/gDW.Similarly,
fortheredmacroalgaG.spinosum,theproteinamountwas9.73±0.41mg/gDWwhenUS
wasappliedfor60min.Thiswasfurthersupportedbymicroscopicobservation,which
revealedacompletestructuralalteration(Figure2).
Consideringpreviousdataonthetimerequiredforeffectiveultrasoundpretreatment
[71,72],theevaluationoftheUStechniquewascarriedoutwithdifferentpretreatment
periodsrangingfrom15to60min.OurstudydemonstratedthatUSfor60mincoupled
withapoweroutputof200Wsignificantlyaffectstheconcentrationsofproteinreleased.
SimilarresultswerereportedbyAl-Zuhairetal.[73],illustratinghowultrasoundinflu-
encedthelevelsofextractedproteinfromvariousmicroalgaespeciesincludingChlorella
sp.,Ankistrodesmusbraunii,Pseudochlorococcumsp.,Tetraselmissp.,andNannochloropsissp.
Furthermore,ourresultsareconsistentwiththefindingsofPernetandTremblay[74]who
determinedthatdifferentdisruptionmethods,particularlysonicationtechniques,signifi-
cantlyaffectedtheextractedproteinlevelsfromthemarine-centricdiatom,Chaetoceros
gracilis.Inaddition,FPapplication(600MPafor4min)tofacilitateproteinextractionfrom
tworedalgae(PalmariapalmataandChondruscrispus)wasstudiedbyO’Connoretal.[28].
Indeed,FPtreatmentat400MPafor20minofSoleriachordalisonlyresultedinanincrease
of2.60%(w/w)inproteinyield[75].
Figure 4. Protein contents of S. coronopifolius (A) and G. spinosum (B) extracts using different tech-
niques of cell disruption. a–f: different letters mean significant differences between extracts (p< 0.05).
Results are expressed as average
±
standard deviation (SD) (n= 3). US, ultrasonication; MG, manual
grinding; TPP, three-phase partitioning; UATPP, ultrasonication-assisted three-phase partitioning;
MG-US30, manual grinding + ultrasonication for 30 min; FD-US30, freeze drying + ultrasonication
for 30 min; FP, French press; and BB, bead-beating.
For both macroalgae, the protein levels after 10 min of MG have no significant differ-
ence (p> 0.05) compared to untreated cells. On the other hand, the comparison of other
techniques revealed the superiority of US 60 min, followed by the FP method. The total
quantity of extractable protein from S. coronopifolius was 3.43
±
0.06 mg/g DW. Similarly,
for the red macroalga G. spinosum, the protein amount was 9.73
±
0.41 mg/g DW when US
was applied for 60 min. This was further supported by microscopic observation, which
revealed a complete structural alteration (Figure 2).
Considering previous data on the time required for effective ultrasound pretreat-
ment [
71
,
72
], the evaluation of the US technique was carried out with different pretreatment
periods ranging from 15 to 60 min. Our study demonstrated that US for 60 min coupled
with a power output of 200 W significantly affects the concentrations of protein released.
Similar results were reported by Al-Zuhair et al. [
73
], illustrating how ultrasound influ-
enced the levels of extracted protein from various microalgae species including Chlorella
sp., Ankistrodesmus braunii,Pseudochlorococcum sp., Tetraselmis sp., and Nannochloropsis sp.
Furthermore, our results are consistent with the findings of Pernet and Tremblay [
74
] who
determined that different disruption methods, particularly sonication techniques, signif-
icantly affected the extracted protein levels from the marine-centric diatom, Chaetoceros
gracilis. In addition, FP application (600 MPa for 4 min) to facilitate protein extraction from
two red algae (Palmaria palmata and Chondrus crispus) was studied by O’ Connor et al. [
28
].
Indeed, FP treatment at 400 MPa for 20 min of Soleria chordalis only resulted in an increase
of 2.60% (w/w) in protein yield [75].
Foods 2024,13, 1362 13 of 24
The combined treatment of MG-US30 (3.09
±
0.03 mg/g DW and 5.19
±
0.18 mg/g
DW for S. coronopifolius and G. spinosum, respectively) also seems interesting due to the
higher protein concentrations compared to the US for 30 min alone (2.61
±
0.19 mg/g DW
and
6.71 ±0.96 mg/g
DW for S. coronopifolius and G. spinosum, respectively). In addition,
the protein contents recorded with UATPP (1.89
±
0.04 mg/g DW and
6.96 ±0.25 mg/g
DW for
S. coronopifolius
and G. spinosum, respectively) are higher (p< 0.05) than that
obtained with TPP (0.72
±
0.09 mg/g DW and 4.89
±
0.41 mg/g DW for
S. coronopifolius
and
G. spinosum
, respectively). The combination with the ultrasonic treatment, capable of
cracking the cell wall of macroalgae, probably explains the differences in protein contents
between UATPP and TPP. These findings are consistent with those of Chia et al. [
37
]
who reported that UATPP was found to be an improved technique compared to TPP
for the extraction of proteins from Chlorella vulgaris FSP-E. Moreover, the lowest protein
concentrations for both macroalgae were obtained in untreated cells and through MG,
especially for S. coronopifolius.
The ultrasonication method is based on liquid shear forces caused by high-frequency
wave sounds (up to 15–20 kHz). These sound waves form gas bubbles or cavities in
the liquid, which reach a threshold size after a certain number of cycles, collapsing and
releasing significant amounts of energy. Acoustic cavitation also causes cell wall destruction
by increasing local temperatures and producing hydroxyl radicals [
76
]. It has been reported
that ultrasonication with lower frequencies and higher power causes more violent cavitation
reactions [
77
–
79
] and has substantial mechanical effects on solid particles, potentially
enhancing mass transfer during extraction [
80
]. The effect of ultrasound is due to bubble
cavitation, which facilitates the disruption of biological matrices [29].
3.5. Protein Molecular Weights Profile
US for 1 h appears to be the most effective method for extracting the maximum protein
content from both macroalgae. These proteins were precipitated with ammonium sulfate.
Recently, ammonium sulfate (80% w/v) precipitation in combination with dialysis using
a 3.5 kDa MWCO membrane of proteins extracted from macroalgae by sonication (1 h
at 42 Hz) has been demonstrated [
28
]. Ammonium sulfate is the salt of choice due to its
food-grade status, cost effectiveness, exceptional solubility, and its ability to efficiently
stabilize protein structures [81,82].
The molecular weight distribution of the extracted macroalgae soluble proteins by US
60 min from S. coronopifolius and G. spinosum was estimated under denaturing conditions by
SDS-PAGE (Figure 5). In general, the SDS-PAGE profile of protein extracts for the two red
macroalgae samples showed that the protein bands were resolved clearly without having
too much smearing. Some variations in the protein pattern between the two species of red
macroalgae were also observed. Effectively, the major protein bands of S. coronopifolius
were observed between 25 and 60 kDa (Figure 5, lane 2). In contrast, the G. spinosum
protein extract revealed that a majority showed a pattern containing approximately six
discrete bands with molecular weights of 25 to 150 kDa (Figure 5, lane 3). Bands containing
abundant proteins were observed in the two species, but their intensity varied among the
extracts of red macroalgae. Previous research showed that most protein fractions of red
macroalgae visualized by SDS-PAGE ranged between 6.5 and 116 kDa. In addition, band
sizes with low molecular weight (<15 kDa), however, varied, revealing the differences in
types of proteins among various seaweed samples.
The SDS-PAGE profile of aqueous soluble proteins extracted from milled oven-dried
Palmaria palmata has presented a greater number of protein bands ranging in size from
15.5 to 97 kDa with four main protein bands ranging from 14.8 to 55 kDa compared to the
alkaline soluble protein extract [83].
Foods 2024,13, 1362 14 of 24
Foods2024,13,xFORPEERREVIEW15of25
Figure5.Sodiumdodecylsulphatepolyacrylamidegelelectrophoresisprofilesofredmacroalgae
solubleproteinextracts.Lane1:molecularweightstandards.Lane2:watersolubleproteinfromS.
coronopifolius.Lane3:watersolubleproteinfromG.spinosum.
Phycoerythrin,whichisaphotosensitiveredpigmentfromthephycobiliproteinfam-
ilypredominantlypresentinredalgae,iscomposedofthreesubunits(α,β,andγ)with
apparentmolecularweightsof18,20,and30–33kDa,respectively.Thiswater-soluble
chromoproteinisalargeoligomercharacterizedbytheaggregationofitssubunitstoform
abasicunitwithdifferentarrangementslikethecomplexesofαβthatcanhaveamolecu-
larweightofabout38kDa.
3.6.ReleaseofSolubleCarbohydrates
Separatingcarbohydratesfromproteinsproveschallengingduetothesubstantial
presenceofthesepolymerswithinthealgalmatrix,potentiallyleadingtoprotein–poly-
saccharideinteractions.Overcomingthischallengestillremainsamajorissueintheex-
tractionofproteinsfromalgae[84].InthecaseofS.coronopifolius,carbohydratecontents
rangedfrom9.03mg/gDWto235.25mg/gDW(Figure6A).Similarly,G.spinosumextracts
presentedvaluesbetween79.56±1.56mg/gDWand199.18±7.53mg/gDW(Figure6B).
Notably,theuseofUSimprovedtheextractionprocess,yieldingsignificantly(p<0.05)
highercarbohydratecontentsforbothredmacroalgae.Moreover,thedurationofUShad
asignificantinfluence(p<0.05)onthereleaseofcarbohydratescomparedtothecontrol
group.ThelowestcontentswereobservedwithMGforS.coronopifolius(60.43mg/gDW)
andTPPforG.spinosum(74.42±6.47mg/gDW).Inlightofthesefindings,USaloneorin
combinationwithotherbiomasspretreatmenttechniquehasenormouspotentialininter-
cellularcarbohydratesrelease.
Figure 5. Sodium dodecyl sulphate polyacrylamide gel electrophoresis profiles of red macroalgae
soluble protein extracts. Lane 1: molecular weight standards. Lane 2: water soluble protein from
S. coronopifolius. Lane 3: water soluble protein from G. spinosum.
Phycoerythrin, which is a photosensitive red pigment from the phycobiliprotein
family predominantly present in red algae, is composed of three subunits (
α
,
β
, and
γ
)
with apparent molecular weights of 18, 20, and 30–33 kDa, respectively. This water-soluble
chromoprotein is a large oligomer characterized by the aggregation of its subunits to form
a basic unit with different arrangements like the complexes of
αβ
that can have a molecular
weight of about 38 kDa.
3.6. Release of Soluble Carbohydrates
Separating carbohydrates from proteins proves challenging due to the substantial pres-
ence of these polymers within the algal matrix, potentially leading to protein–polysaccharide
interactions. Overcoming this challenge still remains a major issue in the extraction of
proteins from algae [
84
]. In the case of S. coronopifolius, carbohydrate contents ranged
from 9.03 mg/g DW to 235.25 mg/g DW (Figure 6A). Similarly, G. spinosum extracts pre-
sented values between 79.56
±
1.56 mg/g DW and 199.18
±
7.53 mg/g DW (Figure 6B).
Notably, the use of US improved the extraction process, yielding significantly (p< 0.05)
higher carbohydrate contents for both red macroalgae. Moreover, the duration of US had
a significant influence (p< 0.05) on the release of carbohydrates compared to the control
group. The lowest contents were observed with MG for S. coronopifolius (60.43 mg/g DW)
and TPP for G.