Changes in the Composition of Olive Pomace after Fermentation: A Preliminary Study

Abstract

Because olive pomace (the main by-product of olive oil production) is phytotoxic, new applications must be investigated to minimize its negative environmental impact. In this work, olive pomace was fermented for 4 and 32 days at room temperature, having in view its valorization as a novel food, thereby creating opportunities for the food industry and addressing a challenge of the olive oil sector. The chemical and microbiological modifications that occurred along the fermentation were followed. The results showed no significant differences (p > 0.05) in total protein between the control and the fermented samples; however, the latter exhibited higher levels of essential amino acids. The major nonessential and essential amino acids were glutamic acid and leucine in all samples. There was a significant increase in the total fat of the 32-day sample and the main fatty acid was oleic acid in all samples. There were considerable reductions in total vitamin E, phenolics, and antioxidant activity values post-fermentation. Hydroxytyrosol replaced oleacein as the main phenolic in the 32-day sample. A sharp increase in total microorganisms occurred (2.20 × 10² to 3.00 × 10⁴–2.01 × 10⁷ colony forming units/mL) but no pathogenic microorganisms were detected. Overall, olive pomace fermentation creates novel products for the food industry with a balanced nutritional composition.

1. Introduction

Food fermentation occurs when the fermentative microorganisms present in the food matrix or in the surrounding environment start growing and initiate enzymatic conversion reactions, creating novel dietary compounds. With the growing emphasis on a healthy diet, consumers are now giving attention to the functional properties of fermented foods. Thus, the food industry has strived to create novel fermented foods with superior flavors, textures, and broader consumer appeal. Secondary metabolites produced during fermentation, particularly, phenolic compounds, have been identified as the major contributors for these functional activities. Despite only a small portion being absorbed by the body, unabsorbed phenolics interact with the gut microbiota, offering various health benefits such as cholesterol reduction, blood sugar and pressure control, fostering the growth of beneficial bacteria while inhibiting harmful bacteria, and inducing the secretion of enzymes that metabolize these compounds into growth-promoting factors [1,2].

Meanwhile, the global human population is growing exponentially, leading to increased challenges in ensuring access to safe and nutritious food. This requires an increase in food production, which compromises natural resources, leading to issues such as soil erosion, loss of biodiversity, and environmental pollution. With approximately one-third of all food produced being wasted, thereby posing a threat to food security, it is recommended to repurpose the by-products from the food industry, which retain high nutritional value because they are a source of various compounds. To address these challenges, valorizing these by-products promotes sustainability within the food chain and the preservation of the environment. This aligns with current European Union guidelines promoting the principles of circular economy and waste utilization [3,4,5,6].

Particularly, the production of olive oil generates phytotoxic by-products such as olive pomace, which pose environmental challenges due to their rich composition in valuable compounds (e.g., sugars, pectins, organic acids, oil, nitrogen, phenolics, and high organic matter). Thus, new solutions for this agri-food waste must be found to minimize its negative environmental impact. In particular, new food alternatives are needed to give response to a much more demanding consumer and to meet sustainability in food production [7,8,9].

The use of fermentation as a strategy to valorize olive pomace was previously emphasized in various works, especially for enzyme production [10,11,12,13,14,15]. Ibrahim et al. focused on using fermented olive pomace as poultry feed, which resulted in a significant decrease in the fat and cholesterol contents in the meat, as well as better stability during storage time by freezing, due to the increase in total phenolics in the groups fed with higher percentages of fermented olive pomace [16].

Other researchers have studied the production of several compounds obtained through olive pomace fermentation, such as ethanol [17], xylitol [18], pigments [19], fatty acids [20], and phenolics [21], particularly, gallic acid [12,22].

Regarding food applications, Goula and Lazarides produced an olive paste spread with olive mill wastewater fermented in a brine for 14 days, before mixing it with olive oil, vinegar, peppers, and herbs [23]; Guermazi and Benincasa produced a spreadable pulp by fermenting olive pomace in a brine for 2.5 weeks [24]; Durante et al. used fermented olive pomace with Saccharomyces cerevisiae for 30 days and then with Leuconostoc mesenteroides for 20 days to enrich a bakery product [25]; Nanis et al. also obtained a paste-type product [7]; Tufariello et al. fermented paté olive cake with the sequential application of S. cerevisiae and L. mesenteroides [26]; and Foti et al. used different microbial strains of Candida boidinii, Wickerhamomyces anomalus, and Lactiplantibacillus plantarum to ferment a paté olive cake [27].

In this work, a fast and long fermentation of olive pomace was carried out for 4 days and 32 days, respectively, aiming its valorization as a novel food, while creating an opportunity for the food industry and helping to minimize a problem of the olive oil sector. A preliminary assessment of several parameters was performed to understand the impact of fermentation on the composition of the resultant olive pomace samples, namely regarding total protein, fat, and phenolics contents; amino acids, fatty acids, vitamin E, and phenolics profiles; antioxidant activity; total count of microorganisms; and pathogenic bacteria evaluation.

2. Materials and Methods

2.1. Raw Material

The olive pomace was generously provided by a Portuguese olive oil producer from Alfândega da Fé (M.C. Rabaçal & Aragão, Lda., Alfândega da Fé, Portugal).

2.2. Fermentation Process

The raw material was mixed with sterile saline solution in a 50:50 ratio (m/V) with a total volume of 2 L. The control was analyzed immediately (no fermentation). The sample was then left to ferment with the microorganisms present in olive pomace, corresponding to the autochthonous microbiota of the product, showing an enrichment and predominance of a type of yeast in which characterization is ongoing. Samples selected for analysis correspond to samples fermented at room temperature for 4 days and 32 days, corresponding to a fast and long fermentation, respectively. Therefore, three samples were studied as follows: the control, the 4-day sample, and the 32-day sample.

2.3. Standards and Reagents

Absolute ethanol was obtained from Fisher Chemical (Loughborough, UK). Anhydrous sodium sulfate (Na₂SO₄), di-sodium hydrogen phosphate (Na₂HPO₄), di-sodium tetraborate decahydrate (Na₂B₂O₇), Folin–Ciocalteu’s reagent, Kjeldahl catalyst tablets (Na₂S₂O₈/CuSO₄), L-norvaline (≥99% purity), potassium hydroxide (KOH), and n-hexane (HPLC-grade) were acquired from Merck (Darmstadt, Germany). Dichloromethane, sand, and sodium hydroxide (NaOH) were purchased from VWR International (Leuven, Belgium). Boric acid (H₃BO₃) and potassium hydroxide (KOH) were attained from Panreac (Barcelona, Spain). Sodium hydroxide (NaOH) and sulfuric acid (H₂SO₄) were bought from Carlo Erba (Val de Reuil, France). Acetonitrile (ACN, HPLC-grade), hydrochloric acid (HCl), methanol (MeOH, HPLC-grade), and sodium azide (NaN₃) were obtained from Riedel-de Haën (Honeywell, NC, USA). 9-fluorenylmethyl chloroformate (FMOC), borate buffer (0.4 N, pH 10.2), and o-phthalaldehyde/3-mercaptopropionic acid (OPA/3-MPA) were acquired from Agilent Technologies (Santa Clara, CA, USA). The Amino Acids Mix Solution (certified reference material, Trace CERT^®, Sigma-Aldrich, Buchs, Switzerland) was purchased from Sigma-Aldrich (Steinheim, Germany). 1,4-dioxane; 2,2-diphenyl-1-picrylhydrazyl radical (DPPH^•, ≥90% purity); 2,4,6-tripyridyltriazine (TPTZ, ≥99% purity); aluminum chloride (AlCl₃); boron trifluoride (BF₃, 14% in methanol); catechin (≥98% purity); ferric chloride; ferrous sulfate (FeSO₄); gallic acid (≥98% purity); glacial acetic acid; petroleum ether; oleuropein, luteolin, and chlorogenic acid standards (purity 99%, HPLC-grade); sodium acetate; sodium carbonate (Na₂CO₃); sodium nitrite (NaNO₂); and Trolox (≥98% purity) were attained from Sigma (St. Louis, MO, USA). The fatty acids standard mixture (FAME 37 Mix, certified reference material, Trace CERT^®) was bought from Supelco (Bellefonte, PA, USA). Vitamin E standards (α-, β-, γ-, δ-tocopherols and α-, β-, γ-, δ-tocotrienols, ≥95% and ≥97% purity, respectively) were obtained from Calbiochem (La Jolla, CA, USA). Tocol (2-methyl-2-(4,8,12-trimethyl tridecyl)-chroman-6-ol, ≥95% purity) was acquired from Matreya Inc. (Pennsylvania, PA, USA). Ultrapure water was attained in a Milli-Q water purification system (Millipore, Bedford, MA, USA).

2.4. Protein Fraction Analysis

2.4.1. Total Protein Content

The total protein content was assessed via the Kjeldahl method (AOAC 928.08) [28], employing a nitrogen conversion factor of 6.25 [29]. Results are in g/100 g of the sample in dry weight.

2.4.2. Amino Acids (AAs) Profile by HPLC-DAD-FLD

The determination of the AAs profiles followed Machado et al. [30], employing a HPLC system from Jasco (Tokyo, Japan) equipped with MD-2015 Plus multiwavelength and FP-2020 Plus fluorescence detectors, and a ZORBAX Eclipse Plus C18 column (kept at 40 °C, 4.6 × 250 mm, 5 μm, Agilent Technologies, USA). KOH (4 M) was used for alkaline hydrolysis (only for tryptophan, 4 h) and HCl (6 M) was used for acid hydrolysis (24 h) in glass tubes that were heated (110 °C) in a SBH130D/3 heating block (Stuart, Stafford, UK). A fraction of the hydrolysates was mixed with L-norvaline (2 mg/mL, internal standard) and exposed to automatic online derivatization with OPA/3-MPA and FMOC in an AS-4150 autosampler. Fluorescence detection was observed at λ_exc = 340 and λ_em = 450 nm (0–26.2 min) for OPA-derivatives and at λ_exc = 266 and λ_em = 305 nm (26.2–40 min) for FMOC-derivatives. OPA-derivatives were observed at λ = 338 nm and FMOC-derivatives at λ = 262 nm. The gradient solvent system was (A) phosphate/borate buffer (10 mM Na₂HPO₄:10 mM Na₂B₂O₇ (pH = 8.2):5 mM NaN₃) and (B) MeOH:ACN:H₂O (45:45:10, V/V/V): 0.85′ 2% of B; 33.4′ 57% of B; 33.5′ 85% of B; 39.3′ 85% of B; 39.4′ 2% of B; and 40′ 2% of B. The flow rate was 1.5 mL/min. The injection volume was 3 μL. Results are in g/100 g of the sample in dry weight.

2.5. Lipid Fraction Analysis

2.5.1. Total Fat Content

The total fat content was assessed via the Soxhlet method (AOAC 991.36) [28]. Results are in g/100 g of the sample in dry weight.

2.5.2. Lipid Fraction Extraction

The extraction of the lipid fractions followed Alves et al. [31] with slight modifications by Melo et al. [4]. Briefly, the solution was homogenized in a Heidolph Multi Reax Vibrating Shaker (VWR, Pennsylvania, PA, USA). Tocol was used as the internal standard (0.1 mg/mL) and n-hexane was used as the extracting solvent. The final extracts were used for analysis of the FAs and vitamin E profiles.

Fatty Acids (FAs) Profile by GC-FID

The derivatization of FAs into fatty acids methyl esters (FAMEs) followed ISO 12966-2017 [32]. FAs profiles were determined in a gas chromatograph (GC-2010 Plus, Shimadzu, Tokyo, Japan) equipped with an automatic sampler, an AOC-20i split/splitless auto-injector (50:1 split ratio, 250 °C), a flame ionization detector (FID, 270 °C), and an Agilent J&W CP-Sil 88 silica capillary column (50 m × 0.25 mm, 0.20 μm, Agilent Technologies, CA, USA). Carrier gas: helium (3.0 mL/min). The applied temperature program was: 50 °C held for 1 min, 8 °C/min to 120 °C held for 3 min, 2 °C/min to 160 °C held for 15 min, and 2 °C/min to 220 °C. Injection volume was 1 μL. The identification of FAMEs was accomplished through retention time comparison with a standard mixture (FAME 37 Mix). Results are in relative percentage (%) of total FAs.

Vitamin E Profile by HPLC-DAD-FLD

Vitamin E profiles were determined in an HPLC system (Jasco, Tokyo, Japan) composed of a multiwavelength diode array detector (DAD, MD-2015), a fluorescence detector (FLD, FP-2020, λ_exc = 290 and λ_em = 330 nm) and a Supelcosil^TM LC-SI normal phase column (75 mm × 3.0 mm, 3.0 μm, Supelco, Bellefonte, PA, USA). The mobile phase was 1.5% 1,4-dioxane in n-hexane (V/V). The flow rate was 0.7 mL/min. The injection volume was 20 μL. α-, β-, γ-, δ-tocopherols and α-, β-, γ-, δ-tocotrienols identification was accomplished based on their UV spectra and through retention time comparison with individual standards. Results are in mg/100 g of the sample in dry weight.

2.6. Antioxidant Fraction Analysis

2.6.1. Antioxidant Fraction Extraction

After optimization, the extraction was performed with 200 mg of each sample mixed with 30 mL of water/ethanol (1:1) in a Heidolph Multi Reax Vibrating Shaker (VWR, Pennsylvania, USA) at 2000 rpm for 30 min, in triplicate. A reextraction was performed with 20 mL of solvent for 15 min. After centrifugation, the supernatants of both extractions were combined, filtered, and stored at −20 °C prior to analysis of the total phenolics and antioxidant activity assays.

2.6.2. Total Phenolics Content (TPC) and Antioxidant Activity Assays by Spectrophotometric Methods

TPC, ferric reducing antioxidant power (FRAP), and 2,2-diphenyl-1-picrylhydrazyl radical scavenging ability (DPPH^•-SA) assays determination followed Ferreira et al. [6]. The calibration curves were prepared with gallic acid (5–100 mg/mL; R² = 0.999), ferrous sulphate (25–500 μmol/L; R² = 0.999), and Trolox (5.62–175.34 mg/L; R² = 0.998), respectively. The absorbance was measured in a Synergy HT GENS5 microplate reader (BioTek Instruments, Vermont, USA) at λ = 765, 595, and 525 nm, respectively. Results are in g of gallic acid equivalents (GAE), mmol of ferrous sulphate equivalents (FSE), and g of Trolox equivalents (TE) per 100 g of the sample in dry weight, respectively.

2.7. Phenolic Compounds Profile by HPLC-DAD-MS-ESI⁺

2.7.1. Sample Preparation

To prepare the sample, 0.2 g of the lyophilized sample with 2 mL of methanol acidified with 1% HCl (37%) were vortexed (1 min), followed by ultrasonic treatment (30 min, room temperature) and centrifugation (10,000 rpm, 10 min). The supernatant, rich in the extracted polyphenols, was filtered (nylon filter, pore size 0.45 μm) and 20 μL was injected in the HPLC system.

2.7.2. Chromatographic Conditions

A HP-1200 liquid chromatograph equipped with a quaternary pump, autosampler, a DAD detector, and an MS-6110 single quadrupole API-electrospray detector (Agilent-Technologies, USA) was used for analysis. The positive ionization mode was applied to detect the phenolic compounds; different fragmentor, in the range 50–100 V, were applied. Column: Kinetex XB-C18 (5 μm; 4.5 × 150 mm, Phenomenex, USA). For the mobile phase, (A) water acidified by acetic acid 0.1%; and (B) ACN acidified by acetic acid 0.1%. The multistep linear gradient was 5% of B for 2 min; 5–90% of B in 20 min, hold for 4 min at 90% of B, then 6 min to arrive at 5% of B. The total time of analysis was 30 min. The flow rate was 0.5 mL/min. The oven temperature was 25 °C. Mass spectrometric detection of positively charged ions was performed employing the Scan mode. The experimental conditions were gas temperature 350 °C, nitrogen flow 7 L/min, nebulizer pressure 35 psi, capillary voltage 3000 V, fragmentor 100 V, and m/z 120–1500. Figure 1 presents the chromatograms obtained at λ = 280 and 340 nm. The Agilent ChemStation software (v. B.02.01-SR2) was employed for data acquisition.

2.7.3. Quantification

The tyrosol content was calculated with a 5-point calibration curve of oleuropein (y = 9.6467x − 2.0489, R² = 0.9978, linearity range: 10–50 μg/mL). The flavones content was calculated with a 5-point calibration curve of luteolin (y = 68.857x − 25.113, R² = 0.9972, linearity range: 1–100 μg/mL). The hydroxycinnamic acid and cyclitol content was calculated with a 5-point calibration curve of chlorogenic acid (y = 22.585x − 36.728, R² = 0.9937, linearity range: 10–50 μg/mL). Results are in mg/g of the sample.

2.8. Microbiological Analysis

2.8.1. Total Count of Microorganisms

Each sample was accompanied through 32 days. In order to analyze the microbiota population, a total of 1 g of each sample was diluted in 9 mL of sterile saline solution, making dilutions until 10⁻⁸, then 100 μL of each were spread in the Plate Count Agar (PCA, Liofilchem, Italy) and incubated at 37 °C for 48 h. Results are in colony forming units (CFU)/mL.

2.8.2. Pathogenic Bacteria Evaluation

Molecular methodology, through PCR assay, was performed aiming the verification of pathogenic species in the microbiota of the samples. The original sample was submitted to an enrichment step using the following two broths: Tryptic Soy Broth (TSB, Liofilchem, Roseto degli Abruzzi, Italy) and Rappaport-Vassiliadis Soya Peptone Broth (RVS, Liofilchem, Roseto degli Abruzzi, Italy). Each broth’s DNA was extracted by centrifuging 1 mL of it, discarding the liquid, adding saline solution (0.85%), heating the suspension to 100 °C for 15 min, and then centrifuging it again to separate the DNA from the lysed cells. The extracted DNA was analyzed using the PCR method with the primers presented at

Table 1. List of primers used to check the presence of putative pathogens.

Broth	Species	Primer	Sequence	Size (bp)	Ref.
TSB	Enterococcus faecalis	E1 (1551)	5′-ATCAAGTACAGTTAGTCTT-3′	941	[33]
		E2 (1552)	5′-ACGATTCAAAGCTAACTG-3′
	Enterococcus faecium	F1 (1553)	5′-GCAAGGCTTCTTAGAGA-3′	550
		F2 (1554)	5′-CATCGTGTAAGCTAACTTC-3′
	Klebsiella pneumoniae	ntrA F	5′-CATCTCGATCTGCTGGCCAA-3′	90	[34]
		ntrA R	5′-GCGCGGATCCAGCGATTGGA-3′
	Enterobacter cloacae	Atpd F	5′-CGAGAGCCTGUTGCTG-3′	180
		Atpd R	5′-GATTGGCTGACCCAAT-3′
	Citrobacter spp.	16s rRNA F	5′-GCTCAACCTGGGAACTGCATCCGA-3′	529
		16s rRNA R	5′-AGTTCCGGCCTAACCGCTGGCAA-3′
	Escherichia coli	UidA F	5’-CTGGTATCAGCGCGAAGTCT-3′	556
		UidA R	5’-AGCGGGTAGATATCACACTC-3′
	Shiga toxin-producing E. coli	Stx1F	ATAAATCGCCATTCGTTGACTAC	180	[35]
		Stx1R	AGAACGCCCACTGAGATCATC
		Stx2F	GGCACTGTCTGAAACTGCTCC	255
		Stx2R	TCGCCAGTTATCTGACATTCTG
		eaeAF	GACCCGGCACAAGCATAAGC	384
		eaeAR	CCACCTGCAGCAACAAGAGG
		hlyAF	GCATCATCAAGCGTACGTTCC	534
		hlyAR	AATGAGCCAAGCTGGTTAAGCT
RVS	Salmonella spp.	invA 1	5′-ACAGTGCTCGTTTACGACCTGAAT-3′	243	[36]
		invA 2	5′-AGACGACTGGTACTGATCGATAAT-3′

Tryptic Soy Broth (TSB), Rappaport-Vassiliadis Soya Peptone Broth (RVS).

. The genes amplification by PCR were analyzed through electrophoresis using agarose gel 1.5–2% (NZYTech, Lisbon, Portugal) and the running conditions were 200 A, 90–120 V during 30–9 min—depending on whether the PCR was multiplex (testing more than one gene at once) or simplex (testing one gene at once).

2.9. Statistical Analysis

Significant differences between samples’ means (p < 0.05) were assessed by one-way ANOVA and the Tukey post hoc test, employing IBM SPSS Statistics v.26 (IBM Corp., New York, NY, USA). Experiments were performed in triplicate (n = 3).

3. Results

Table 2. Chemical characterization of the protein fraction of the olive pomace samples.

Parameter	Control	4 Days	32 Days
Total protein content	5.75 ± 0.05 a	5.48 ± 0.31 a	5.84 ± 0.59 a
Aspartic acid	0.46 ± 0.02 b	0.49 ± 0.02 a,b	0.52 ± 0.03 a
Glutamic acid	0.58 ± 0.02 b	0.62 ± 0.02 a,b	0.65 ± 0.04 a
Serine	0.24 ± 0.01 b	0.25 ± 0.01 a,b	0.27 ± 0.02 a
Histidine *	0.08 ± 0.01 a	0.09 ± 0.00 a	0.09 ± 0.01 a
Glycine	0.25 ± 0.01 a	0.27 ± 0.01 a	0.27 ± 0.02 a
Threonine *	0.21 ± 0.01 b	0.23 ± 0.01 a,b	0.24 ± 0.01 a
Arginine	0.33 ± 0.01 a	0.34 ± 0.01 a	0.34 ± 0.02 a
Alanine	0.25 ± 0.01 b	0.26 ± 0.01 a,b	0.28 ± 0.02 a
Tyrosine	0.11 ± 0.01 a	0.117 ± 0.004 a	0.12 ± 0.01 a
Valine *	0.25 ± 0.01 b	0.27 ± 0.01 a,b	0.29 ± 0.02 a
Methionine *	0.02 ± 0.00 b	0.03 ± 0.00 a	0.03 ± 0.00 a
Tryptophan *	0.02 ± 0.00 a	0.02 ± 0.00 a	0.02 ± 0.00 a
Phenylalanine *	0.28 ± 0.01 b	0.30 ± 0.01 a,b	0.32 ± 0.02 a
Isoleucine *	0.22 ± 0.01 b	0.24 ± 0.01 a	0.25 ± 0.01 a
Leucine *	0.54 ± 0.02 b	0.58 ± 0.02 a,b	0.61 ± 0.03 a
Lysine *	0.12 ± 0.00 b	0.13 ± 0.00 a	0.12 ± 0.00 b
Hydroxyproline	0.12 ± 0.00 b	0.12 ± 0.00 b	0.13 ± 0.01 a
Proline	0.26 ± 0.01 b	0.28 ± 0.01 a,b	0.29 ± 0.01 a
∑Essential amino acids	1.75 ± 0.06 b	1.89 ± 0.05 a	1.97 ± 0.09 a
∑Non-essential amino acids	2.59 ± 0.09 b	2.76 ± 0.09 a,b	2.87 ± 0.16 a
∑Total amino acids	4.40 ± 0.07 b	4.65 ± 0.14 a,b	4.84 ± 0.25 a

* Essential amino acid. Results are in g/100 g of the sample dry weight (average ± s.d., n = 3). Distinct letters in each line denote significant differences between samples (p < 0.05). Asparagine and glutamine were not identified because, in hydrolysis conditions, they were converted into aspartic and glutamic acids, respectively [30].

presents the chemical characterization of the protein fraction of the olive pomace samples. There were no significant differences (p > 0.05) between the total protein content of the samples (about 5.5–5.8 g/100 g). However, there was a significant increase in the sum of essential AAs in the fermented samples in relation to the control (1.89–1.97 vs. 1.75 g/100 g, respectively). The 32-day sample also showed significant increases in the nonessential and total AAs sums in comparison to the control (2.87 vs. 2.59 and 4.84 vs. 4.40 g/100 g, respectively). The AAs profile revealed that the major nonessential AA was glutamic acid in all samples (0.58–0.65 g/100 g) but the 32-day sample presented a significantly higher amount than the control (Table 2). The main essential AA in all samples was leucine, and the 32-day sample presented also significantly higher (p < 0.05) quantities compared with the control (around 0.6 vs. 0.5 g/100 g, respectively).

Table 3. Chemical characterization of the lipid fraction of the olive pomace samples.

Parameter	Control	4 Days	32 Days
Total fat content	8.43 ± 0.13 b	9.19 ± 0.49 b	11.54 ± 0.83 a
C16:0 Palmitic acid	10.49 ± 0.14 a	10.18 ± 0.13 b	10.22 ± 0.06 a,b
C16:1 Palmitoleic acid	0.45 ± 0.01 a	0.45 ± 0.03 a	0.46 ± 0.03 a
C17:0 Margaric acid	0.06 ± 0.01 b	0.08 ± 0.00 a	0.08 ± 0.01 a
C18:0 Stearic acid	2.88 ± 0.06 a	2.79 ± 0.03 a	2.85 ± 0.06 a
C18:1n9c Oleic acid	74.78 ± 0.16 a	75.40 ± 0.36 a	75.18 ± 0.23 a
C18:2n6c Linoleic acid	10.07 ± 0.25 a	9.44 ± 0.17 b	9.57 ± 0.15 b
C20:0 Arachidic acid	0.25 ± 0.01 b	0.38 ± 0.01 a	0.35 ± 0.01 a
C18:3n3c α-Linolenic acid	0.63 ± 0.01 b	0.66 ± 0.01 b	0.74 ± 0.01 a
C20:1n9c cis-11-Eicosanoic acid	0.17 ± 0.00 b	0.25 ± 0.00 a	0.26 ± 0.01 a
C22:0 Behenic acid	0.14 ± 0.01 b	0.20 ± 0.01 a	0.18 ± 0.00 a
C24:0 Lignoceric acid	0.08 ± 0.01 c	0.17 ± 0.01 a	0.11 ± 0.00 b
∑Saturated fatty acids	13.90 ± 0.08 a	13.80 ± 0.14 a	13.80 ± 0.08 a
∑Monounsaturated fatty acids	75.40 ± 0.14 b	76.11 ± 0.28 a	75.90 ± 0.20 a,b
∑Polyunsaturated fatty acids	10.69 ± 0.22 a	10.10 ± 0.13 b	10.31 ± 0.13 a,b

Total fat results are in g/100 g of the sample dry weight. Fatty acids results are in relative % of total fatty acids. Results are expressed as average ± s.d. (n = 3). Distinct letters in each line denote significant differences between samples (p < 0.05).

presents the chemical characterization of the lipid fraction of olive pomace samples. There was a significant increase in the total fat content (around 3 g/100 g) in the 32-day sample in relation to the control. However, the overall FAs profiles were not altered during fermentation because the same FAs were identified in all samples and the major one was oleic acid (roughly 75%) in all samples. Palmitic and linoleic acids were the following most abundant FAs with values ranging from around 9 to 10% (Table 3). Regarding the statistical analysis, a significant reduction of around 0.3% was verified for palmitic acid in the 4-day sample in comparison to the control. Similarly, the percentage of linoleic acid was decreased with fermentation from 10.07% in the control to 9.44–9.57% in the fermented samples. The minor FAs increased slightly but significantly with fermentation, in particular, margaric, arachidic, cis-11-eicosanoic, and behenic acids (Table 3). The α-linolenic acid percentage increased significantly only in the 32-day sample (Table 3).

Table 4. Chemical characterization of the antioxidant fraction of olive pomace samples.

Parameter	Unit	Control	4-Days	32-Days
α-tocopherol	mg/100 g	4.23 ± 0.39 a	1.46 ± 0.35 b	1.93 ± 0.14 b
β-tocopherol		0.24 ± 0.01 a	0.25 ± 0.01 a	0.22 ± 0.00 b
γ-tocopherol		0.39 ± 0.02 a	0.36 ± 0.02 a	0.37 ± 0.01 a
∑Total vitamin E		4.86 ± 0.41 a	2.06 ± 0.37 b	2.52 ± 0.14 b
Total phenolics content	g GAE/100 g	4.08 ± 0.31 a	3.43 ± 0.40 b	4.11 ± 0.21 a
FRAP	mmol FSE/100 g	41.75 ± 2.64 a	29.27 ± 3.06 c	33.37 ± 1.66 b
DPPH•-SA	g TE/100 g	4.28 ± 0.25 a	3.28 ± 0.35 b	3.53 ± 0.27 b

GAE—gallic acid equivalents, FRAP—ferric reducing antioxidant power, FSE—ferrous sulfate equivalents, DPPH^•-SA—2,2-diphenyl-1-picrylhydrazyl radical scavenging ability, TE—Trolox equivalents. Dry weight results are expressed as average ± s.d. (n = 3). Distinct letters in each line denote significant differences between samples (p < 0.05).

presents the results related to the antioxidant fraction of olive pomace covering polar (phenolics) and apolar (vitamin E) antioxidants. The major vitamin E isomer was α-tocopherol in all samples, δ-tocopherol was not identified neither any of the tocotrienols. The total vitamin E significantly decreased from around 4.9 mg/100 g in the control to 2.0–2.5 mg/100 g in the fermented samples (Table 4).

Similar to vitamin E, it is possible to see a reduction in the TPC from roughly 4.1 g GAE/100 g in the control to 3.4 g GAE/100 g in the 4-day sample. However, that was not verified for the 32-day sample (Table 4). Despite that, in the quantification by HPLC-DAD-MS-ESI⁺, there was a reduction in total phenolics from around 93 in the control to 85–86 mg/g in the fermented samples (

Table 5. Phenolic compounds profile by HPLC-DAD-MS-ESI⁺ of olive pomace samples.

Peak No.	Retention Time (min)	UV λmax (nm)	[M + H]+ (m/z)	Compound	Subclass	Control	4 Days	32 Days
1	2.91	225	193	Quinic acid	Cyclitol	2.34 ± 0.03 b	2.59 ± 0.04 a	2.54 ± 0.02 a
2	9.17	280	317	Hydroxytyrosol glucoside	Tyrosol	5.59 ± 0.07 a	5.23 ± 0.09 b	0.00 ± 0.00 c
3	9.74	280	155	Hydroxytyrosol	Tyrosol	13.48 ± 0.04 c	17.42 ± 0.03 b	32.83 ± 0.07 a
4	12.07	280	139	Tyrosol	Tyrosol	3.59 ± 0.05 c	3.83 ± 0.03 b	5.71 ± 0.04 a
5	13.55	290	405	Oleoside 11-methyl ester	Tyrosol	3.82 ± 0.02 a	3.37 ± 0.06 b	3.01 ± 0.02 c
6	15.35	330	363	Ligstroside-aglycone	Tyrosol	11.94 ± 0.05 b	13.06 ± 0.03 a	10.27 ± 0.06 c
7	15.76	332	625	Verbascoside	Hydroxycinnamic acid	6.41 ± 0.05 a	4.84 ± 0.02 b	3.81 ± 0.05 c
8	16.02	340	449	Luteolin-7-O-glucoside	Flavone	0.73 ± 0.02 a	0.73 ± 0.03 a	0.26 ± 0.06 b
9	17.42	341	433	Apigenin-7-O-glucoside	Flavone	0.30 ± 0.05 a	0.35 ± 0.02 a	0.11 ± 0.05 b
10	18.27	280	557	Hydroxyoleuropein	Tyrosol	4.72 ± 0.02 a	2.28 ± 0.07 b	0.69 ± 0.09 c
11	18.68	280	379	Oleuropein-aglycone	Tyrosol	4.55 ± 0.04 a	2.18 ± 0.05 b	1.40 ± 0.07 c
12	20.12	295	197	Hydroxytyrosol-acetate (3,4-DHPEA-AC)	Tyrosol	8.43 ± 0.06 a	6.08 ± 0.08 b	5.84 ± 0.03 c
13	21.09	340	287	Luteolin	Flavone	1.01 ± 0.03 c	1.33 ± 0.03 b	1.66 ± 0.09 a
14	22.95	290	321	Oleacein (3,4-DHPEA-EDA)	Tyrosol	26.26 ± 0.05 a	22.76 ± 0.09 b	16.71 ± 0.06 c
Total Phenolics (mg/g)						93.18 a	86.05 b	84.86 c

Results are expressed as average ± s.d. (n = 3). Distinct letters in each line denote significant differences between samples (p < 0.05).

). Nevertheless, in terms of antioxidant activity results, both assays (FRAP and DPPH^•-SA) revealed considerable decreases in their values with both fermentation processes, which is in agreement with the depletion of total vitamin E and phenolics. This decrease was especially notorious in the FRAP value of the 4-day sample (Table 4). This seems to be correlated with the lower TPC value found for this sample, because phenolic compounds are significant contributors to the antioxidant capacity measured by the FRAP assay. Lower TPC usually indicates a lower amount of phenolic antioxidants, which, in turn, results in lower reducing power.

Table 5 presents the DAD and MS data obtained after positive ionization of the olive pomace samples and the content of phenolic compounds. In the present study, the same 14 phenolics from four different subclasses were identified before and after fermentation as follows: a cyclitol–quinic acid [37,38]; 9 tyrosols–hydroxytyrosol (HT) glucoside [37,39], HT [37,40], tyrosol [41,42], oleoside 11-methyl ester [37,43], ligstroside-aglycone [41,44], hydroxyoleuropein [37,45], oleuropein-aglycone [37,46], HT-acetate [47,48], and oleacein [37,49]; a hydroxycinnamic acid–verbascoside [37,50]; and 3 flavones–luteolin-7-O-glucoside [37,51], apigenin-7-O-glucoside [37,52], and luteolin [47,53].

Despite that, several differences in the amounts of the identified phenolics are noticed after fermentation as follows (Table 5): quinic acid increased slightly post-fermentation; HT glucoside exhibited a slight decline after the fast fermentation but was not identified after the long fermentation; HT had a notable increase of nearly 4 mg/g in the fast fermentation and around 19 mg/g in the long fermentation; tyrosol increased slightly in the 4-day sample but it increased about 2 mg/g in the 32-day sample; oleoside 11-methyl ester was reduced by roughly 0.8 mg/g; ligstroside-aglycone increased in the fast fermentation slightly more than 1 mg/g but decreased in the 32-day sample; verbascoside decreased to nearly half; luteolin-7-O-glucoside and apigenin-7-O-glucoside presented similar amounts in the control and 4-day sample but decreased around 0.5 and 0.2 mg/g, respectively, after the long fermentation; hydroxyoleuropein, oleuropein-aglycone, and HT-acetate experienced reductions exceeding 4 mg/g, 3 mg/g, and almost 2.6 mg/g, respectively; and luteolin showed a slight increase of approximately 0.6 mg/g, whereas oleacein decreased by nearly 10 mg/g.

Overall, oleacein stood out as the predominant phenolic in the control. However, the long fermentation induced a notable shift in the profile, with HT emerging as the most abundant phenolic. As previously stated, after fermentation, there was a notable decline in oleacein levels from roughly 26 to 23 and 17 mg/g in the 4-day and 32-day samples, respectively. Conversely, the HT content exhibited an increase during fermentation, rising from approximately 13 to 17 and 33 mg/g in the 4-day and 32-day samples, respectively (Table 5).

The microbiological analysis revealed that the control (no fermentation) presented 2.20 × 10² CFU/mL in the total count of microorganisms. We noticed a sharp increase in this amount with fermentation, especially at 4 days (2.01 × 10⁷ CFU/mL) and also at 32 days (3.00 × 10⁴ CFU/mL). Despite this increase, none of the pathogenic microorganisms (Enterococcus faecalis, Enterococcus faecium, Klebsiella pneumoniae, Enterobacter cloacae, Citrobacter spp., Escherichia coli, Shiga toxin-producing E. coli, and Salmonella spp.) were detected.

4. Discussion

The results revealed that olive pomace fermentation led to noteworthy modifications in the chemical composition of this by-product, particularly in the protein, lipid, and antioxidant profiles.

Concerning the protein fraction, the differences between the results of total protein (determined by the Kjeldahl method and calculated using nitrogen value) and the AAs sum (HPLC-DAD-FLD technique) could be due to the presence of other AAs and derivatives not analyzed with the employed HPLC methodology or to the presence of nonprotein nitrogen [30]. Thus, in future work, the nonprotein nitrogen will be determined to better understand these differences.

The major nonessential AA found in the samples was glutamic acid (Table 2). Glutamine is the body’s most prevalent AA, which plays a pivotal role in intermediary metabolism and is utilized extensively by immune cells. Many elite athletes utilize dietary glutamine supplements to restore immune function post-training [54]. Additionally, the main essential AA found in the samples was leucine (Table 2). This is a branched-chain AA that must be supplied by the diet. It plays numerous health-related metabolic functions such as protein synthesis, tissue regeneration, and metabolism. Moreover, its carbon skeleton can be used to produce ATP [55].

Regarding the lipid fraction, oleic acid was the major FA identified, which is a monounsaturated FA associated with cardiovascular protection [56]. Oleic acid’s mechanisms include improved blood lipid profiles by lowering total cholesterol, low-density lipoprotein, and triglycerides, while increasing high-density lipoprotein and inhibiting enzymes involved in cholesterol synthesis [56]. Also, it lowers blood pressure by affecting membrane lipid composition and G-protein signaling, enhancing vasodilation [56]. Moreover, it improves vascular endothelial function by increasing nitric oxide availability, reducing inflammation, and inhibiting oxidative stress pathways [56]. It also improves insulin sensitivity and stabilizes atherosclerotic plaques, reducing thrombosis risk [56].

Nanis et al. found that all the yeast strains that they identified in the fermented solid residue of olive mill wastewaters were catalase-positive, contributing to the preservation of the fermented product against FAs oxidation [7]. In the present study, we found that the 4-day sample had a significant but small reduction in the sum of polyunsaturated FAs, which can be related to the slight decrease in linoleic acid of around 0.6% in relation to the control (Table 3). However, the 32-day sample did not show significant differences in the FAs sums in comparison to the control (Table 3); therefore, this could probably be a result of the higher metabolic activity of the microorganisms in the long fermentation, helping to protect against FAs oxidation [7].

Other studies have analyzed the FAs profile. Durante et al. [25] identified the same FAs as in the present study (Table 3), except lignoceric acid (C24:0). Tufariello et al. obtained similar FAs sums for both unfermented and fermented samples: about 65%, 20%, and 14% for monounsaturated, saturated, and polyunsaturated FAs sums, respectively [26]. Although we also obtained similar values between the unfermented and fermented samples, these values differ from our results (Table 3).

The main energy sources for bacteria and yeast’s metabolic processes include glucose (a simple sugar), which can be metabolized through glycolysis to produce ATP, or other sugars such as fructose, sucrose, maltose, and lactose, which can be broken down through various metabolic pathways to generate ATP. Some species can also metabolize organic acids like acetic acid, lactic acid, and citric acid to obtain energy. Also, AAs can be broken down through processes like transamination and oxidative deamination to generate ATP. Certain species are capable of metabolizing FAs that undergo β-oxidation to generate acetyl-CoA, which enters the citric acid cycle to produce ATP [57,58].

The microorganisms probably used the available sugars and organic acids present in olive pomace as an energy source, because this by-product is a source of both types of compound [7]. It does not seem like the microorganisms used AAs as an energy source because the AAs sums actually increased after fermentation (Table 2) as mentioned above. Similarly, it does not appear that the microorganisms used FAs as an energy source because there was a significant increase in the total fat content in the 32-day sample and its FAs profile did not present many noteworthy changes post-fermentation (Table 3). In future work, we intend to determine the sugars profile to better understand these findings.

Fathy et al. evaluated the chemical composition of olive pomace after solid-state fermentation with the yeast Kluyveromyces marxianus, having found higher protein and fat contents after fermentation [22]. In the present data, that was only verified in the fat content after the long fermentation process, as previously mentioned.

In terms of antioxidants, the major vitamin E isomer was α-tocopherol in all samples (Table 4), which is the most biologically active form in the organism [4]. α-tocopherol significantly contributes to preventing lipid peroxidation and scavenging lipid peroxyl radicals [9]. There was sharp decrease in total vitamin E post-fermentation (Table 4). This depletion is probably due to its use as a protection against oxidative processes, because although the fermentation occurred in a closed system, there was a residual amount of air in the top of the flasks. Also, the fermentation occurred in the presence of light and vitamin E can also be degraded by this external factor [59]. Unlike the present study, Tufariello et al. found no significant changes in the content of α-tocopherol after the fermentation process [26].

Phenolics can stimulate enzymes secretion (e.g., esterase, gallate decarboxylase, glycosidase, phenolic acid decarboxylase, and tannase) by microorganisms, which, in turn, metabolize challenging phenolics into growth factors. Concurrently, several biotransformation pathways cause the breakdown of phenolics by microorganisms that can enhance their bioavailability in food materials, resulting in a range of components that are more easily digested and absorbed by the body [2].

The differences in the results of TPC (Table 4) and the quantification of total phenolics by HPLC-DAD-MS-ESI⁺ (Table 5) could probably be explained by the higher sensitivity of the HPLC technique in comparison to the spectrophotometric assay with Folin–Ciocalteu’s reagent, which can sometimes overquantify due to the interference of the food matrix, for example [60]. Indeed, the TPC method can be influenced by various factors such as matrix effects, chemical interferences, calibration standard variations, and solvent influences. The electron-transfer reaction fundamental to the Folin–Ciocalteu assay, occurring under basic conditions, renders it susceptible to interference from nonphenolic compounds present in the sample matrix, potentially compromising the accuracy of the analysis [60].

A significant reduction in TPC was also previously reported after fermentation of the solid residue of olive mill wastewaters, which the authors correlated to oxidation, condensation processes, and enzymatic activities of yeasts [7].

In terms of sensory analysis, the overall reduction in total phenolics is interesting because it can help mitigate the astringency and bitterness of olive pomace, potentially enhancing consumer acceptance as previously suggested [24].

Both antioxidant activity assays (FRAP and DPPH^•-SA) revealed considerable decreases in their values after fermentation, which is in agreement with the depletion of total vitamin E and phenolics.

Mahmoud et al. attributed the antioxidant and anticancer bioactivities to several phenolics present in fermented olive pomace by K. marxianus such as carvacrol, thymol, eugenol, caryophyllene oxide, and methyl isopalmitate [61]. Unlike the present study, where we used 50% aqueous ethanol, Mahmoud et al. tested 100% ethanol and found that their fermented sample showed higher antioxidant activities in DPPH^•-SA in comparison to their unfermented counterpart [61].

Similar to our study (Table 5), Durante et al. also identified HT, tyrosol, verbascoside, and oleacein after enriching a bakery product with olive pomace paste [25]. Tufariello et al. [26], in addition to those four compounds, also identified HT-acetate, like in our study (Table 5).

Nanis et al. found that the reduction in oleuropein in the fermented solid residue of olive mill wastewaters was due to the enzymatic activity of yeasts, particularly esterase and β-glucosidase [7]. The activation of oleuropein hydrolysis enzymes was also previously suggested [24]. Thus, the reduction in the oleuropein derivatives in the present study (namely hydroxyoleuropein and oleuropein-aglycone) might be due to the metabolic activity of yeasts in the olive pomace and/or enzymatic activity.

Initially, we hypothesized that the significant surge in HT content could be attributed to its secondary metabolite status derived from tyrosine as observed in a study with wine yeasts [27,62]. However, the analysis of tyrosine contents before and after fermentation revealed no significant differences (0.11–0.12 g/100 g, Table 2). Given that oleacein is converted to HT through hydrolysis [63,64] and that we have added saline solution to the olive pomace, this probably facilitated this conversion process, explaining our findings.

The degradation of other phenolics can also explain these results, particularly, oleuropein-aglycone can also be converted into HT by hydrolysis [63,65]. Indeed, the first compound experienced a reduction exceeding 3 mg/g throughout fermentation, so it could also have contributed to the obtained HT’s contents.

In addition, verbascoside is formed with HT, caffeic acid, and the sugar α-L-rhamnopyranosyl-(1→3)-β-D-glucopyranose [66]; therefore, it is possible that verbascoside’s degradation (Table 5) also contributed to the final HT’s contents. Similarly, HT glucoside was not identified after the long fermentation, therefore it was probably converted to HT because it is one of its derivatives. The increase in HT’s amount and the reductions in oleacein and verbascoside contents were corroborated by the results of a previous study that used paté olive cake from two different cultivars [26].

The fermented samples are a considerable source of HT, especially after the long fermentation (Table 5). HT exhibits various health effects through multiple mechanisms of action [67]. HT acts as a potent anti-inflammatory agent by inhibiting inflammatory cytokines expression such as tumor necrosis factor-α and interleukin-1 [67]. HT inhibits the activation of microglial cells and reduces the production of reactive oxygen species induced by inflammatory stimuli [67]. These effects contribute to its neuroprotective role and its potential in mitigating neuroinflammation associated with conditions like Parkinson’s and Alzheimer’s by inhibiting the formation of α-synuclein and β-amyloid fibrils, respectively [67]. In addition, HT is able to inhibit angiogenesis, which is particularly important in combating cancer and cardiovascular diseases in which abnormal angiogenesis plays a crucial role in disease development and progression [67]. HT also exhibits anti-tumoral activity by inducing apoptosis and anti-proliferative effects in cancer cells [67]. HT has been shown to exert insulin-like effects on target cells, contributing to its anti-diabetic effects [67].

In agreement with the results of the microbiological analysis of our study, the authors did not find lactic acid bacteria, staphylococci, pseudomonads, nor Enterobacteriaceae, which is likely due to the high phenolic content and acidic environment that discouraged their growth, indicating the suitability of the food product for human consumption [7]. Yeasts were the predominant microorganisms, especially S. cerevisiae and Pichia membranifaciens, due to the favorable conditions of high salt concentration and low pH [7].

The highest total count of microorganisms observed in the 4-day sample can probably be explained by the high metabolic activity of the microflora present in the olive pomace in the beginning of fermentation. During this period, the amount of available nutrients is still high, allowing a big growth period in the population, probably in the exponential phase of the microbial growth curve. In the 32-day sample, the amount of toxic products starts to accumulate, probably leading to a death phase, explaining why the total microbial count was reduced [68].

Since this was a preliminary assessment of the changes in olive pomace composition post-fermentation, the next line of work will be to investigate a possible probiotic effect due to the increase in the total microbial count. Additionally, we aim to scale up the production of the fermented samples to obtain higher quantities and facilitate further analysis, including sensory evaluations. Through these assessments, we seek to gauge the extent to which tasters perceive the reduction in the phenolic content and, consequently, in bitterness, and whether they approve the final product. Thus, future research will help to further elucidate the nutritional and health-promoting properties of the obtained products.

5. Conclusions

In conclusion, the fermentation of olive pomace led to notable changes in its chemical composition, particularly in the protein, lipid, and antioxidant profiles.

Both fermented samples exhibited increased levels of essential AAs, with the 32-day sample showing significant enhancements in nonessential and total AAs sums. This underscores the potential of fermentation in enhancing the nutritional value of olive pomace.

The lipid fraction showed a significant increase in total fat content in the 32-day sample, while maintaining the beneficial FAs profile dominated by oleic acid.

The fermentation also influenced the antioxidant fraction, resulting in a considerable reduction in total vitamin E and phenolics. However, these reductions may positively impact sensory characteristics, potentially reducing bitterness and improving consumer acceptance.

The phenolic profile exhibited alterations post-fermentation, with notable shifts in the quantities of various compounds, suggesting the metabolic activity of microorganisms in degrading certain phenolics, as well as conversion reactions between phenolics. Interestingly, the increase in HT content, despite no significant changes in tyrosine levels and oleacein’s reduction, suggests its conversion into the first compound.

Microbial analysis revealed a significant increase in total microorganism count post-fermentation, although none of the tested pathogenic microorganisms were detected, indicating the product’s safety.

Overall, these findings highlight the potential of fermentation as a valuable bioprocessing technique for enhancing the nutritional quality and functional properties of olive pomace, thereby adding value to this by-product of olive oil production. At the same time, it reduces the amount of olive pomace, ameliorating a problem of the olive oil sector, while it also gives response to an increasingly competitive sector such as the food industry by meeting consumer demand for novel, functional, and sustainable foodstuffs. Moreover, the valorization of olive pomace through fermentation is aligned with food security, circular economy, and sustainability issues.