1. Introduction
According to the Food and Agriculture Organization (FAO) of the United Nations, global food production must increase in order to meet the demands of the growing world population. Yet, even under the threat of natural resources depletion and climate change, a considerable percentage of the food produced worldwide is lost or wasted along the food value chain, representing a missed opportunity to improve food security and human nutrition with serious economic, environmental, and social impacts [
1]. Despite international efforts to reduce food wastage and promote circular economy and sovereignty, more investments are needed to encourage these sustainability practices and to show that they can generate multi-level benefits [
2].
Nowadays, food waste biorefineries have been proposed as innovative technological solutions to the growing challenge of waste management, giving their high potential to reduce the environmental burden and produce bio-based chemicals, materials, and energy [
3]. In this way, food waste receives a second chance by entering the value cycle as a renewable biorefinery feedstock to obtain high value-added products. There are studies focused on multi-component cascade biorefinery processes for orange [
4], potato [
5], and pomegranate [
6] peels, among other fruit and vegetable waste. Although a sequential biorefinery process has already been implemented for the valorization of quince (
Cydonia oblonga Mill.) peel and seeds through pectin extraction and subsequent pyrolysis, a process from which bioproducts or biofuels can be obtained [
7], the upcycling of industrial waste from this acidic and astringent fruit through sustainable biorefinery concepts deserves further attention.
Quince is widely used for the production of jams, jellies, marmalade, liqueurs, and other sweet foods. Its edible pulp contains high levels of simple sugars, polysaccharides, and organic acids, while lipids and proteins are found in smaller amounts [
8]. In turn, its peel is particularly rich in fructose, fiber, malic acid, and potassium [
9]. Regarding phenolic compounds, hydroxycinnamic acid derivatives (mainly, 3-caffeoylquinic and 5-caffeoylquinic acids) and polymeric procyanidins (or flavan-3-ols) have been described in whole pitted quince [
10] and its peel [
9]. Both quince pulp and peel extracts have antioxidant activity, and peel extracts can be even more effective in scavenging free radicals and inhibiting the growth of some microorganisms than pulp extracts [
11,
12]. These studies highlighted the bioactive potential of quince peel and its potential to be reintroduced in the value cycle as a source of natural antioxidants, antimicrobials, acidulants, and flavor enhancers for food and beverage formulation, among other applications.
The extraction is a crucial step in the isolation of valuable compounds from quince peel or other food waste, but it must be rethought to achieve the complete valorization of this feedstock. Although most studies on the valorization of agri-food waste use only the filtrate or supernatant from the extraction, the resulting residues should also be used as potential sources of food value ingredients, such as dietary fiber [
9,
13,
14]. Furthermore, since there is no standard extraction procedure due to the wide variety of waste and bioactive molecules it may contain, it is critical to develop targeted extraction processes for particular constituents from specific food waste, thus contributing to efficient utilization and circular bioeconomy. The effects of relevant factors or independent variables on the extraction process can be evaluated using suitable optimization tools, such as response surface methodology (RSM). RSM involves design, modelling, and optimization steps, and allows the development of predictive models that summarize the behavior of the response(s) variable(s) under the tested experimental conditions [
15], allowing one to determine the conditions that maximize the extraction yield, minimizing the degradation of target compounds and making it possible to save solvent and energy.
This study was performed to evaluate the upgrading potential of quince peel for simultaneous production of bioactive extracts (BEs) and dietary fiber concentrates (FCs) for future application as natural food preservatives and fortifiers. For this, the plant material was processed under an RSM-couped 20-run central composite rotatable design (CCRD) and the extraction supernatants (named BEs) were analyzed for phenolic compounds and malic acid, among other water-soluble constituents, while the extraction residues (named FCs) were characterized for their dietary fiber content and color parameters. After determining optimal extraction conditions, the model-predicted outcomes were experimentally validated and the resulting phenolic and malic acid-enriched BEs were characterized for their in vitro antioxidant activity and antimicrobial effects.
2. Materials and Methods
2.1. Plant Material
Quince (Cydonia oblonga Mill.) peels were supplied by local farmers from the Bragança region, Portugal, in October 2020, who grow this fruit for homemade marmalade production. The peel sample containing 74.6 ± 0.2% of water was lyophilized to constant weight, ground with a domestic mill until passing through a 20-mesh sieve, and kept vacuum-packaged at −20 °C until use.
2.2. Experimental Design for Extraction Optimization
The independent variables
X1 (time,
t, 1–119 min),
X2 (temperature,
T, 26–94 °C), and
X3 (EtOH percentage,
S, 0–100%,
v/v) were combined in a CCRD composed of 8 factorial points, 6 axial or star points, and 1 center point replicated six times and investigated using Design-Expert software, 11 (Stat-Ease, Inc., Minneapolis, MN, USA). These independent variables and their range of values were selected based on previous studies [
16,
17]. The CCRD allowed both axial and factorial points to have the same radial distance from the center and, therefore, the same prediction error magnitude. The natural and coded values are presented in
Table 1. The 20 experimental runs were randomized to minimize unexpected variability. For each design point, mean values (
n = 6) were used as observed responses.
2.3. Extraction and Preparation of BEs and FCs
The extraction was performed in a thermostated water bath using sealed vessels to avoid solvent evaporation. Sample weights (1.8 g) were mixed with 30 mL of solvent (0–100% EtOH) and stirred at five levels of time (1–119 min) and temperature (26–94 °C) according to the 20-run design matrix. After extraction, the mixtures were centrifuged at 4000×
g for 10 min and the supernatants (BEs) and solid residues (FCs) were collected (
Figure 1). An aliquot of the supernatants was used to determine the BE yield (%,
w/w) and the remaining portion was concentrated under reduced pressure to remove EtOH and lyophilized for further analysis of phenolic compounds, organic acids, and soluble sugars. The solid residues were oven-dried at 60 °C until constant weight for subsequent determination of FC yield and dietary fiber content and measurement of color parameters.
2.4. Determination of Experimental Responses
2.4.1. Extraction Yields
The BE and FC yields (%, w/w) were determined gravimetrically. For BE, 4 mL of each extraction supernatant (obtained in 2.3) was placed into calcined porcelain crucibles and the solvent was removed at 105 °C for at last 24 h until constant weight. The FC yields were determined by weight difference after oven-drying at 60 °C.
2.4.2. Phenolic Compounds
The BEs were redissolved to 6 mg/mL in 20% EtOH, filtered through 0.22-μm syringe filters, and analysis in a Dionex Ultimate 3000 UPLC system (Thermo Scientific, San Jose, CA, USA) with a diode array detector (DAD, using 280 and 370 nm as preferred wavelengths) and a LTQ XL linear ion trap mass spectrometer (MS, Thermo Finnigan, San Jose, CA, USA) equipped with an electrospray ionization (ESI) source [
18]. Chromatographic separation was made in a Waters Spherisorb S3 ODS-2 C18 column (4.6 mm × 150 mm, 3 µm; Waters, Milford, MA, USA). Compounds were identified by comparison of their retention times and UV-vis and mass spectra with those of available standards and data from the literature. The concentration (mg/g BE) of the identified compounds was calculated by interpolating the peak areas on 7-level calibration curves (
r2 ≥ 0.999) constructed with standards of chlorogenic acid (
y = 168,823
x – 161,172; limit of detection (LOD) = 0.20 µg/mL; limit of quantification (LOQ) = 0.68 µg/mL),
p-coumaric acid (
y = 301,950
x + 6966.7; LOD = 0.68 μg/mL; LOQ = 1.61 μg/mL), catechin (
y = 84,950
x – 23,200; LOD = 0.17 μg/mL; LOQ = 0.68 μg/mL), and quercetin-3-
O-glucoside (
y = 34,843
x – 160,173; LOD = 0.21 µg/mL; LOQ = 0.71 µg/mL) purchased from Extrasynthese (Genay Cedex, France). Compounds were thus expressed in equivalents of their basic constituent or similar compound (i.e., standard used in quantification).
2.4.3. Organic Acids
The BEs were redissolved to 6 mg/mL in metaphosphoric acid, filtered through 0.2-μm syringe filters, with analysis in an ultra-fast liquid chromatography (UFLC) system (Shimadzu 20A series, Kyoto, Japan) coupled to a photodiode array (PDA) detector as previously described [
19]. Chromatographic separation was achieved in reverse phase on a C18 column (250 mm × 4.6 mm, 5 µm; Phenomenex, Torrance, CA, USA). Detection was done on PDA at 215 and 245 nm. The detected compounds were identified by chromatographic comparisons with standards and quantified (g/100 g BE) by interpolating the peak areas in calibration curves (
r2 ≥ 0.994) constructed with oxalic acid (
y = 9 × 10
6x + 377.946; LOD = 12.55 µg/mL; LOQ = 41.82 µg/mL), quinic acid (
y = 612.327
x + 16.563; LOD = 24.18 µg/mL; LOQ = 80.61 µg/mL) and malic acid (
y = 912.441
x + 92.665; LOD = 35.76 µg/mL; LOQ = 119.18 µg/mL) standards acquired from Sigma-Aldrich (Saint Louis, MO, USA).
2.4.4. Soluble Sugars
The BEs were redissolved to 6 mg/mL in distilled water, filtered through 0.2-µm syringe filters, and analyzed in by HPLC with refraction index detection as previously described [
20]. Chromatographic separation was achieved with a Eurospher 100-5 NH
2 column (4.6 × 250 mm, 5 mm, Knauer, Berlin, Germany), using acetonitrile/water 70:30 (
v/v) as mobile phase. Soluble sugars were identified by chromatographic comparisons with standards and quantified (g/100 g BE) by interpolating the peak areas in calibration curves (
r2 ≥ 0.999) constructed with fructose (
y = 1.04
x; LOD = 0.05 mg/mL; LOQ = 0.18 mg/mL), glucose (
y = 0.935
x; LOD = 0.08 mg/mL; LOQ = 0.25 mg/mL), and sucrose (
y = 0.977
x; LOD = 0.06 mg/mL; LOQ = 0.21 mg/mL) standards acquired from Sigma-Aldrich (St. Louis, MO, USA).
2.4.5. Dietary Fiber
The dietary fiber content of the FCs was determined by an enzymatic-gravimetric method (AOAC 985.29) [
21]. Briefly, the FCs (250 mg) were gelatinized with heat-stable α-amylase (pH 6.0, 95 °C water bath for 15 min) and then enzymatically digested with protease (pH 7.5, 60 °C water bath for 30 min) and amyloglucosidase (pH 4.5, 60 °C water bath for 30 min) to remove protein and starch (kit from Sigma-Aldrich, Saint Louis, MO, USA). After overnight EtOH precipitation followed by filtration, the residues were successively washed with 78% EtOH, 95% EtOH, and acetone, and then oven-dried at 105 °C and weighed. The protein (AOAC 920.152) and ash (AOAC 940.26) contents were determined by macro-Kjeldahl nitrogen analysis (N × 6.25) or incineration at 525 °C, respectively, for subsequent calculation of the dietary fiber content (g/100 g FC).
2.4.6. Color Parameters
The FC color was measured with a colorimeter (model CR-400; Konica Minolta Sensing Inc., Japan) calibrated with a standard white tile. The parameters
L* (lightness, from
(0) black to
(100) white),
a* (chromaticity from
(–) green to
(+) red), and
b* (chromaticity from
(–) blue to
(+) yellow) were measured as previously described by the authors [
22]. For visual representation, the CIELAB color values were converted to RGB (red, green, blue) color.
2.5. Extraction Process Modelling and Statistical Verification of the Models
The response variables considered for the optimization of the extraction process were grouped into three main groups: (i) bioactive extract (BE) and its main compounds (BE yield (%, w/w), Σ phenolic compounds (mg/g BE), Σ phenolic acids (mg/g BE), Σ flavan-3-ols (mg/g BE), and malic acid (g/100 g BE)); (ii)) other water-soluble BE constituents (quinic acid (g/100 g BE), Σ organic acids (g/100 g BE), fructose (g/100 g PE), glucose (g/100 g BE), and Σ soluble sugars (g/100 g BE)); and iii) fiber concentrate (FC) and its properties (FC yield (%, w/w), dietary fiber (g/100 g FC), lightness (L*), redness (a*), and yellowness (b*)).
The response surface models were fitted using the quadratic Equation (1):
where
Y corresponds to the dependent variable,
X define the independent variables,
b0 is the constant coefficient,
b1,
b2, and
b3 are linear term coefficients,
b11,
b22, and
b33 are quadratic term coefficients, and
b12,
b13, and
b23 are interaction term coefficients. Subscripts 1, 2, and 3 in each term stand for time (
t), temperature (
T), and solvent (
S), respectively.
Fitting procedures, coefficient estimates, and statistical analysis were performed using Design-Expert software. The significance of the models and their terms was assessed by an analysis of variance (ANOVA) at a 95% confidence level, and only the significant terms or those necessary for the hierarchy were considered in the fitting procedures. The lack-of-fit test, the coefficients R2 and R²adj and the adequate precision were used to assess the model-fitting adequacy. Design-Expert was also used to generate the 2D and 3D plots.
2.6. Experimental Validation of the Models and Evaluation of Bioactive Properties of BEs Obtained under Optimized Conditions
Enriched quince peel extracts were obtained under the optimal settings obtained for phenolic compounds, malic acid, and both phytoconstituents. The extraction and analysis were caried out as described in the previous sections and the quantitative results were used for experimental validation of the theoretical models, by comparing experimental and model-predicted data. The BE and FC yields and the FC color parameters were also evaluated, as well as the BE antioxidant and antimicrobial activities. These experiments were performed in triplicate and each sample was measured three times (n = 9).
2.6.1. Antioxidant Activity
The ability of the BEs to inhibit oxidative hemolysis and the formation of thiobarbituric acid reactive substances (TBARS) such as the aldehyde malondialdehyde (MDA) was evaluated in vitro as described below. Trolox and the food additives calcium ascorbate (E302) and sodium metabisulfite (E223) were tested as positive controls.
For TBARS, a porcine brain tissue solution (1:2,
w/v; 0.1 mL) was mixed with 0.1 mL of FeSO
4 (10 µM) and 0.1 mL of ascorbic acid (0.1 mM) and incubated with 0.2 mL of BE (0.016–8 mg/mL), E302 (0.01–2.56 mg/mL), E223 (0.01–2.56 mg/mL), or trolox (3.125–100 µg/mL) for 60 min at 37 °C. After adding 0.5 mL of trichloroacetic acid (28%
w/v) and 0.38 mL of thiobarbituric acid (2%,
w/v), the mixtures were incubated at 80 °C for 20 min and the color intensity provided by MDA-TBA adducts was monitored (Specord 200 spectrophotometer, Analytik-Jena, Jena, Germany) at 532 nm. Half-maximal effective concentration (EC
50) values (µg/mL) were calculated as previously described by the authors [
23].
For oxidative hemolysis inhibition, a red blood cell (RBC) solution (2.8%,
v/v; 0.2 mL) prepared in PBS (pH 7.4) was mixed with 0.4 mL of either BE (0.125–4 mg/mL), E302 (0.01–2.56 mg/mL), E223 (0.01–2.56 mg/mL), trolox (7.81–250 µg/mL), PBS (control), or water (baseline) in a 48 well plate. After 10 min pre-incubation at 37 °C with shaking, 0.2 mL of the free radical generator 2,2′-azobis(2-methylpropionamidine) dihydrochloride (AAPH, 160 mM) was added, and the optical density was monitored at 690 nm (Elx800 plate reader, BioTek) over time. Half-maximal inhibitory concentration (IC
50) values (µg/mL) were calculated for Δ
t of 60 and 120 min, as previously described by the authors [
23].
2.6.2. Antimicrobial Activity
The BEs redissolved in 30% EtOH at 20 mg/mL were screened against the Gram-positive bacteria Staphylococcus aureus (ATCC 11632), Bacillus cereus (food isolate), and Listeria monocytogenes (NCTC 7973) and the Gram-negative bacteria Escherichia coli (ATCC 25922), Salmonella enterica subsp. enterica serovar Typhimurium (ATCC 13311), and Enterobacter cloacae (clinical isolate). The micromycetes Aspergillus fumigatus (human isolate), Aspergillus niger (ATCC 6275), Aspergillus versicolor (ATCC 11730), Penicillium funiculosum (ATCC 36839), Penicillium verrucosum var. cyclopium (food isolate), and Trichoderma viride (IAM 5061) were used to assess antifungal activity. The microorganisms were obtained from the Mycological laboratory, Department of Plant Physiology, Institute for Biological Research “Siniša Stanković”, National Institute of the Republic of Serbia, University of Belgrade, Serbia.
The lowest extract concentration (mg/mL) that inhibited the visible microbial growth at the binocular microscope (minimum inhibitory concentration, MIC) and the lowest concentration (mg/mL) required to kill the original inoculum (minimal bactericidal or fungicidal concentrations, MBC and MFC, respectively) were determined by the serial microdilution method and the
p-iodonitrotetrazolium violet (INT) colorimetric assay [
24,
25]. The antibiotics streptomycin and ampicillin, the antifungals ketoconazole and bifonazole, and the food additives sodium benzoate (E211) and potassium metabisulfite (E224) were used as positive controls, while 30% EtOH was the negative control.
2.6.3. Evaluation of Statistical Differences between Extracts’ Bioactivity
The antioxidant activity results were expressed as the mean ± standard deviation and differences among samples were assessed using one-way analysis of variance (ANOVA). Data normality and variance homogeneity were evaluated by the Shapiro–Wilk and Levene’s tests, respectively. Results were compared using the Tukey’s HSD test. Additionally, a Pearson’s correlation was performed to assess possible correlations between antioxidant activity and BE constituents. Statistical tests were performed at a 5% significance level using SPSS® Statistics, Version 28.0 (IBM Crop, Armonk, NY, USA).
4. Conclusions
BEs and FCs were simultaneously obtained from quince peel through a “zero-waste” biorefinery approach. The BEs contained phenolic acids, flavan-3-ols, glycosylated flavonols, malic acid, fructose, and glucose, among other water-soluble constituents. The optimal extraction conditions for BE yield (66.4 min, 28.4 °C, and 42.6% EtOH), phenolic compounds (64.2 min, 88.0 °C, and 0% EtOH) and malic acid (87.7 min, 92.7 °C, and 54.4% EtOH) were associated with response values of 69% (w/w), 10.6 mg/g BE, and 7.9 g/100 g BE, respectively. For FCs, higher yields tended to be associated with lower fiber percentages, and those containing more dietary fiber were darker. The maximum fiber content (67 g/100 g FC) was associated with the use of 92.2 °C and 35.5% EtOH. The conditions foreseen for the extraction of phenolic compounds and malic acid were experimentally validated and the BEs obtained showed ability to inhibit lipid peroxidation and oxidative hemolysis and antimicrobial potential against foodborne bacteria and fungi. Furthermore, these BEs stood out in some aspects compared to synthetic food additives, especially the malic acid-enriched BE. Overall, the bioactive and functional food extracts resulting from the total upcycling of quince peel could be used as natural ingredients for food preservation and fortification. In future studies, it will be interesting to investigate the effect of solid/liquid ratio on extraction yields and assess the stability and preservative and fortifying capacity of the extracts in food matrices, including beverages.