Saccharomyces cerevisiae Fermentation of Pomegranate Peel By-Product Yields Tannin-Rich Extracts and Potentially Prebiotic Polysaccharides

Abstract

Pomegranate peel, accounting for 35–50% of the fruit weight, is an underutilized agri-food by-product. This study applied, for the first time, fermentation with Saccharomyces cerevisiae as a simple and sustainable strategy to simultaneously obtain tannin-rich extracts and polysaccharide fractions with potential prebiotic activity. Peels from two cultivars, Wonderful and G1, differing in peel thickness, were subjected to three fermentation protocols (air- and not air-exposed) and monitored at 25 °C over 48 and 72 h. HPLC-DAD analysis showed that yeast-inoculated fermentation increased total tannin concentration in dry extracts (up to 70%) without inducing chemical modifications to tannin profiles. As determined by Dynamic Light Scattering, fermentation promoted significant depolymerization of native polysaccharides, while DOSY-¹H-NMR analyses revealed the presence of reduced molecular weight fractions down to 26 kDa. In vitro growth assays confirmed that fermented polysaccharides were more efficiently utilized as a carbon source by Bifidobacterium breve and Lactiplantibacillus plantarum compared to non-fermented controls, likely thanks to polysaccharide depolymerization induced by fermentation. The study demonstrated that air-exposed S. cerevisiae fermentation was an effective process alternative to chemical or enzymatic hydrolysis for modifying pomegranate peel pectin directly within a complex matrix, while simultaneously enhancing tannin recovery. This approach represents a possible sustainable strategy for pomegranate peel valorization into functional ingredients.

1. Introduction

The pomegranate fruit (Punica granatum L.) ranks among the top ten most widely consumed fruits worldwide and has been extensively used since antiquity in traditional medicine across different cultures. In recent years, pomegranate has gained recognition as a “super fruit” due to its high content of bioactive compounds, particularly anthocyanins and ellagitannins [1,2,3,4,5]. Among the fruit components, the juice is especially rich in anthocyanins, whereas the peel contains high levels of ellagitannins, notably α + β-punicalagins [6,7]. During industrial juice production, large quantities of peel are generated as waste, accounting for approximately 35–50% of the total fruit weight. This by-product has attracted increasing interest as a promising raw material for the recovery of high-value compounds through simple, sustainable, and efficient processes [1].

Beyond hydrolysable tannins, pomegranate peel (exocarp and mesocarp) contains 8–12% on dry weight of pectin and pectic polysaccharides (characterized by a reduced percentage of galacturonic acid) [7], which have potential applications in food- and health-related fields [8]. The polysaccharides extracted from the Wonderful cultivar, the most widely cultivated variety globally, are mainly composed of homogalacturonan (HG), and have arabinans and arabinogalactans as side chains. The galacturonic acid content is approximately 56%, with variable degrees of methylation (DM) and acylation (DA), and minor amounts of rhamnogalacturonan [6,7].

While extensive research has focused on the extraction of phenolic compounds from pomegranate peel, methods specifically optimized for pectin recovery remain limited [9]. Notably, extraction protocols, such as decoction, originally developed for phenolic recovery, yield mixed fractions containing both tannins and pectic polysaccharides [1,6]. This compositional complexity may offer a technological advantage for the production of powered, non-hygroscopic dry extracts, without the need for additional carriers such as maltodextrins since pectin perform this function [10].

Microbial fermentation has emerged over the last decade as a sustainable strategy for the valorization of agricultural by-products [9,10]. Fermentation can induce structural and compositional modifications in plant matrices, enhancing their nutritional and functional properties and promoting the release of bioactive compounds [11,12]. Moreover, the consumption of fermented foods has been consistently associated with beneficial health effects [13].

Saccharomyces cerevisiae, often employed for the fermentation processes, was used to treat pomegranate juice for wine production, yielding ethanol concentrations of up to 10% [14,15,16]. In contrast, fermentation of pomegranate peel as a sustainable strategy for exploitation of this by-product remains poorly explored or primarily aimed at ethanol production [11,17] without investigating the fate of other structurally or functionally relevant components. Pomegranate peel fermented by S. cerevisiae produced bioethanol, reaching up to 12.9 g/L yield after a pre-treatment of the peel with hot water (115 °C for 40 min) and addition of specific carbon/nitrogen sources (e.g., glucose, sucrose, meat peptone) [17]. In contrast, the present study was aimed at employing only S. cerevisiae to ferment the dried peel as is, without supplementation of any external nutrients. Few recent works have focused on improving phenolic recovery through solid-state [2] or mixed-culture fermentations [12], reporting increased ellagic acid levels, while largely neglecting the polysaccharide fraction.

Importantly, the effects of yeast-driven fermentation on the structural properties of pomegranate peel pectin have not yet been investigated. In particular, it remains unclear whether fermentation by S. cerevisiae may induce the partial depolymerization of pectin, potentially generating lower-molecular-weight fractions with enhanced prebiotic potential. This aspect is especially relevant in light of previous evidence showing that depolymerized pectin from other agro-industrial by-products, such as apple pomace, exhibit improved prebiotic potential compared to their native counterparts [18]. In vitro studies of colonic fermentations have demonstrated that pomegranate peel powder supports the growth and survival of beneficial gut bacteria, such as Lactobacillus acidophilus [18,19]. Similarly, pectin-derived oligosaccharides, obtained through physical, chemical or enzymatic depolymerization, have been reported to positively modulate gut microbiota composition and activity [20], supporting their potential prebiotic role and their relevance for food applications.

Based on these observations, we hypothesized that the yeast fermentation of pomegranate peel could simultaneously modulate both phenolic and pectic fractions, yielding multifunctional ingredients with complementary health-related properties.

Accordingly, the aim of this study was to evaluate whether the fermentation of pomegranate peel with Saccharomyces cerevisiae enables the sustainable, laboratory-scale production of two high-value functional ingredients: tannin-rich extracts and low-MW pectin with potential prebiotic activity. The novelty of this work lies in the demonstration of a single, yeast-driven fermentation process capable of generating both products simultaneously, thus providing an integrated and efficient strategy for the valorization of an underexploited agro-industrial by-product.

To account for cultivar-dependent differences, two cultivars were fermented: Wonderful and G1, an Italian cultivar selected for its thinner peel. Saccharomices cerevisiae was chosen as it is inexpensive, widely available, and commonly used in domestic preparations. Furthermore, the effects of fermentation with this yeast both on pomegranate tannins and peel polysaccharides have not been previously studied. Three simple fermentation methods were applied at room temperature for up to 72 h. A multimodal analytical approach, combining HPLC-DAD, DOSY ¹H-NMR, and DLS was employed to characterize tannin profiles and monitor pectin depolymerization. Finally, the potential prebiotic activity of low-MW polysaccharides after fermentation was preliminarily evaluated using in vitro assays with beneficial microorganisms, Bifidobacterium breve and Lactobacillus (Lactiplantibacillus) plantarum.

2. Materials and Methods

2.1. Chemicals and Reagents

Food-grade Saccharomyces cerevisiae was obtained from a local supermarket. Solvents for HPLC analyses and D₂O for ¹H-NMR experiments (analytical grade) were purchased from Merck (Saint Louis, MO, USA). Gallic acid (purity ≥ 99%) and ellagic acid (≥95%) were purchased from Extrasynthese S.A. (Lyon, Nord-Genay, France), while punicalagin (≥91%) was purchased from Phytolab (Vestenbergsgreuth, Germany). Maleic acid (98%) was purchased from Merck Life Science S.r.l. (Milano, Italy). Bifidobacterium breve B632 and Lactobacillus plantarum (renamed after 2020 as Lactiplantibacillus plantarum) L12 strains were obtained from the microbial strain collection of the Department of Agro-Environmental Science and Technology, University of Bologna, Italy.

2.2. Fermentation of Pomegranate Samples

The peels of two pomegranate varieties, Wonderful and G1, were collected in 2022. Fermentation experiments were conducted in triplicate, both in the presence of commercial S. cerevisiae (Y) and in its absence (spontaneous fermentation as blank control, B), as outlined in Figure 1.

Figure 1.

Flow chart of the experimental plan showing the three fermentation methods applied to powdered dried pomegranate peel from Wonderful (W) and G1 varieties. (Y): fermented samples with the yeast Saccharomyces cerevisiae. (B): fermented samples without yeast; DE, dry extracts. * Dried peel in water, 100 °C for 2 min before the addition of the yeast.

Commercial S. cerevisiae was added to coarsely ground peel samples at a concentration of 25 mg/g of dried peel, corresponding to 2.5 g/L under the applied solid-to-liquid ratio and comparable to yeast amounts commonly added to flour for bread dough preparation [21]. Previous studies have reported the use of higher yeast concentration for pomegranate peel fermentation (30 g/L) [17]. In contrast, the present study used a significantly lower S. cerevisiae amount to better mimic food-related fermentation conditions and to evaluate the modulation of peel fermentation induced by a limited yeast inoculum. A constant matter-to-solvent ratio was applied for all experiments (1 g/10 mL H₂O), and incubation was performed for 48 and 72 h.

Three fermentation experiments were conducted as follows:

• 1N: Anaerobic fermentation with S. cerevisiae. 

Pomegranate peels were fermented with commercial yeast 25 mg/g at 25 °C under anaerobic conditions, with agitation at 380 rpm in an incubator thermo-shaker (PST-60HL Biosan, SIA, Rīga, Latvia) to ensure sample homogeneity. This protocol was designed to evaluate fermentation under anaerobic conditions and the co-presence of the native and variable peel microbiota. The effects observed in yeast-inoculated samples are interpreted as resulting from modulation of the fermentation process induced by S. cerevisiae.

• 2PbN: Blanching followed by anaerobic fermentation with S. cerevisiae. 

Peel samples were pre-treated by blanching in boiling water (100 °C) for 2 min to partially denature endogenous enzymes and reduce the naturally present microbial load (native microbiota). After cooling, samples were inoculated with S. cerevisiae (25 mg/g) and incubated at 25 °C under the same anaerobic conditions of experiment 1N. This approach allowed the assessment of yeast-driven fermentation in a system with minimized interference from endogenous enzymes and native microbiota.

• 3Na: Anaerobic fermentation with S. cerevisiae. 

Peels inoculated with S. cerevisiae (25 mg/g) were incubated at 25 °C under aerobic conditions. Fermentations were conducted in tubes capped with cotton plugs to allow gas exchange while preventing contamination by environmental microorganisms, according to previous literature [22]. This method allowed for the evaluation of the effect of oxygen availability on the fermentation process, considering the ability of S. cerevisiae to perform ethanol fermentation even in the presence of oxygen. This phenomenon, known as the Crabtree effect, which shapes the S. cerevisiae lag phase during the switch between different carbon sources, is widely described in the literature [22,23,24]. During the process, samples were vortexed for 30 s every 8–12 h to facilitate oxygen distribution, sample homogeneity, and uniform yeast activity.

This approach allowed the assessment of yeast-driven fermentation in untreated peel, thereby minimizing the pre-treatment steps prior to fermentation.

For each fermentation method (i.e., 1N, 2PbN, 3Na), six Y and six B samples were produced, allowing for separate analysis of tannins and polysaccharides in triplicate.

Following the completion of the fermentation at 48 and 72 h, all samples were vortexed for 30 s. Ethanol production was determined by q¹H-NMR experiments (see Section 2.6), along with visual observation of gas and foam formation.

2.3. Extraction of Tannins and Polysaccharides

For the analysis of tannins, each sample fermented at 48 and 72 h was centrifuged at 14,000 rpm, and the supernatant was recovered. A 0.5 mL aliquot was diluted twice with water and analyzed by HPLC-DAD-MS. The remaining extract was freeze-dried to obtain the dry extracts (DEs) (Figure 1).

Aiming to maximize the depolimerization process induced by fermentation, the polysaccharides were only extracted from samples obtained after the maximum time of incubation (72 h). Each sample was mixed with 20 mL of water and subjected to a 60 min decoction. After centrifugation at 5000 rpm for 8 min, the solution was recovered and cooled in an ice bath in the presence of 2 volumes of ethanol for 30 min. The precipitated polysaccharides were recovered by centrifugation (5000 rpm for 10 min at 4 °C), washed with 150 mL of 85% acetone with the aid of a 3 min sonication (ultrasonic cleaner at 40 kHz) to remove traces of entrapped ellagitannins, and recovered by another centrifugation step [1,6,10].

A decoction of unfermented peel was performed for 60 min at 100 °C with a water/peel ratio of 1:40 g/mL to recover native polysaccharides [1,6]. The precipitation and washing steps were conducted as described above. Finally, the polysaccharides were dialyzed using a nitrocellulose membrane with a 12–14 kDa cut-off (Medicell Membranes Ltd., Greenwich, London, UK) and lyophilized. The resulting dry samples were used for DLS and ¹H-NMR analyses as well as for glucuronic acid quantification and in vitro assay.

2.4. Analysis of Tannin Content

The supernatant obtained as described in Section 2.3 was used to quantify tannins in dried peel (DP) and the corresponding dry extract (DE). Tannin analysis was performed using the Agilent HP 1260L liquid chromatograph equipped with a DAD detector (Agilent Technologies, Palo Alto, CA, USA) and coupled with a core–shell Kinetex column C18 (100 × 3 mm, 2.6 μm, Phenomenex, Torrance, CA, USA). The solvents used were (A) 0.1% formic acid in water (pH 3.2) and (B) CH₃CN. A multistep linear solvent gradient was applied as follows: 0.1–8 min, 5–25% B; 8–18 min, 25% B; 18–20 min, 25–95% B; 20–26 min, 95% B; 26–28 min, 95–0% B; and 28–32 min, 0–5% B. A flow rate of 0.4 mL/min and 10 min of equilibration time were applied. UV–Vis spectra were recorded from 200 to 500 nm. The Mass Spectrometer Detector was an HP 1260 MSD (G6125B) with an API/electrospray interface (Agilent Technologies, Palo Alto, CA, USA). The following ESI parameters were used: nitrogen flow rate 10.5 L/min, drying gas temperature 350 °C, nebulizer pressure 1811 Torr, and capillary voltage 3500 V. Acquisition was performed in negative ionization mode (150–180 V), and a full scan was performed from 100 to 2000 Da.

According to a previous work [1], tannins were quantified by HPLC-DAD using five-point calibration curves of three external standards. Gallic acid (0.10–1.11 mg/mL in EtOH 80%) at 280 nm (R² = 0.9998) was used to quantify gallic acid. Ellagic acid (0.08–0.24 mg/mL in MeOH) at 350 nm (R² = 0.9994) was used to quantify ellagic acid and its derivatives. Punicalagins (0.1–0.8 mg/mL in MeOH) at 350 nm (R² = 0.999) were used to quantify punicalagins and their derivatives.

2.5. Analyses of Polysaccharides by DLS

According to previous studies [7,25], Dynamic Light Scattering (DLS) measurements of polysaccharides were carried out using a Zsizer Nanoseries ZS90 (Malvern Instrument, Worcestershire, UK) at 25 °C. Samples were prepared in water at 1.25 mg/mL. Measurements included Z-average (Z-Ave, nm) as size distribution by intensity and polydispersity index (PdI), using a 90° scattering angle. Each sample was analyzed in triplicate; for each independent replicate, results were averaged over 14–19 readings. Data was processed using the 60X0 Malvern software package (ver. 7.2).

2.6. Analyses of Ethanol and Polysaccharides by ¹H-NMR and DOSY

¹H-NMR spectra of the dialyzed polysaccharides were recorded using a 400 MHz Advance 400 spectrometer (Bruker, Bremen, Germany). To monitor the fermentation process, ethanol content was determined via ¹H-NMR obtaining the data in Table S2. Samples were analyzed after centrifugation, recovery of the supernatant, and addition of 10% D₂O. Quantification was based on a calibration curve built with ethanol (99% purity) in the concentration range 0.4–2% (R², 0.9975). The degree of methylation (DM) and acetylation (DA) of polysaccharides were determined according to guidelines and protocols previously described [25,26]. Briefly, approx. 4 mg of dialyzed polysaccharides was precisely weighed into an NMR tube and dissolved in 1 mL of 0.4 M NaOH in D₂O and incubated at room temperature for 2 h. Then, 20 μL of maleic acid (5.2 mg/mL in D₂O) was added as an internal standard. ¹H-NMR spectra were acquired and used to integrate C-methyl signals at 1.80 ppm (acetic acid), 3.22 ppm (methanol), and the internal standard singlet at 6.24 ppm. The number of moles of acetic acid and methanol were calculated using Equation (1).

$$\mathrm{n}={\displaystyle \frac{I_{CH3}}{I_{istd}}}\times {\displaystyle \frac{N_{istd}}{N_P}}\times {\displaystyle \frac{W_{istd}}{{MW}_{istd}}}\times P_{istd}$$ (1)

• n, number of moles of acetic acid or methanol.

• I_istd, integral of the 2 protons of ISTD.

• I_CH3, integral of CH₃ group of methanol or of acetic acid.

• N_P, number of the protons of the methyl group of acetic acid or methanol.

• N_istd, number of protons for ISTD.

• MW_istd, mw of the ISTD (116.1 g/mol).

• W_istd, weight of ISTD.

• P_istd, purity degree of the ISTD.

For galacturonic acid (Gal-Ac) quantification, polysaccharides were hydrolysed in 2.0 M H₂SO₄ in D₂O at 100 °C for 2.5 h. The α-Gal-Ac and β-Gal-Ac anomeric proton signals were integrated at 5.6 ppm and 4.9 ppm, respectively. Equation (1) was applied, with n representing the number of moles of Gal-Ac and NP the number of anomeric protons for α and β forms. Degree of methylation (DM) and acetylation (DA) were calculated according to Müller-Maatsch et al. [27]:

$$\mathrm{D}\mathrm{M}\left(\mathrm{\%}\right)={\displaystyle \frac{\mathrm{m}\mathrm{o}\mathrm{l} \mathrm{o}\mathrm{f} \mathrm{m}\mathrm{e}\mathrm{t}\mathrm{h}\mathrm{a}\mathrm{n}\mathrm{o}\mathrm{l}}{\mathrm{m}\mathrm{o}\mathrm{l} \mathrm{o}\mathrm{f} \mathrm{g}\mathrm{a}\mathrm{l}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{u}\mathrm{r}\mathrm{o}\mathrm{n}\mathrm{i}\mathrm{c} \mathrm{a}\mathrm{c}\mathrm{i}\mathrm{d}}}\times 100$$

$$\mathrm{D}\mathrm{A}\left(\mathrm{\%}\right)={\displaystyle \frac{\mathrm{m}\mathrm{o}\mathrm{l} \mathrm{o}\mathrm{f} \mathrm{a}\mathrm{c}\mathrm{e}\mathrm{t}\mathrm{i}\mathrm{c} \mathrm{a}\mathrm{c}\mathrm{i}\mathrm{d}}{\mathrm{m}\mathrm{o}\mathrm{l} \mathrm{o}\mathrm{f} \mathrm{g}\mathrm{a}\mathrm{l}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{u}\mathrm{r}\mathrm{o}\mathrm{n}\mathrm{i}\mathrm{c} \mathrm{a}\mathrm{c}\mathrm{i}\mathrm{d}}}\times 100$$

The DOSY-¹H-NMR experiments were carried out on a Bruker FT-NMR Avance III HD 600 MHz spectrometer equipped with a CryoProbe BBO H&F 5 mm (Bruker Biospin GmbH Rheinstetten, Karlsruhe, Germany). After loading the sample into the probe, 5 min was required to achieve thermal equilibrium. Afterward, the magnetic field was locked, the probe head was tuned and matched and the sample shimmed. All the experiments were performed at 300 K and non-spinning. The samples were dissolved in D₂O at a concentration of 3 mg/mL and 550 μL of the solution were transferred into a WILMAD^® NMR tube, 5 mm, Ultra-Imperial grade, 7 in. L, 528-PP (Sigma-Aldrich, Milan, Italy). 3-(trimethylsilyl) propionic-2,2,3,3-d4 acid sodium salt (TSP) was added and used for internal referencing. Two-dimensional Diffusion-Ordered Spectroscopy DOSY-¹H-NMR experiments were performed by using the standard Bruker “ledbpgp2s” pulse program using a longitudinal eddy current (LED) bipolar gradient pulse pair and two spoil gradients. Thirty-two gradient steps were used for the diffusion dimension, from 2 to 98% of the linear gradient amplitude, with 65.7 G/cm as the maximum gradient intensity. The diffusion time (d20, Δ) was optimized for each sample and varied between 0.1 and 0.3 s. The pulse field gradient length (P30, δ) was set at 2500 μs. For all the samples, the other acquisition parameters were set as follows: number of scans, 32; gradient strength (gpz6), 100%; LED delay (d21), 5 ms and the total acquisition time 1 h and 16 min. The DOSY spectra were processed using a Bayesian algorithm implemented in Mnova^® 14.1.2 software (Mestrelab Research, Santiago de Compostela, Spain) and the coefficients of diffusion (D) were determined.

The software allowed a semi-quantitative analysis by integrating the diffusion trace (vertical axis of the 2D spectrum and f1 coordinate (https://mestrelab.com/resources-by-product/resources-nmr/dosy.html, last accessed on 30 January 2026). The molecular weights of polysaccharides were calculated by using the linear regression obtained by plotting the Log D versus Log Mw of six standard dextrans with molecular weights from 20 to 500 kDa (purchased from VWR International, Radnor, PA, USA).

2.7. In Vitro Evaluation of Potential Prebiotic Properties

The growth stimulation of the obtained polysaccharides was assessed on two human gut bacterial strains, B. breve B632 and L. plantarum L12. The polysaccharides recovered after 72 h of fermentation of the peels with S. cerevisiae (Y), or without the yeast (B) were used as substrates. The B. breve B632 strain was cultured in TPY medium for 48 h at 37 °C, while L. plantarum L12 was cultured in MRS medium for 48 h at 37 °C, both under anaerobic conditions (Anaerocult A, Merck, Darmstadt, Germany). The growth experiment for B. breve B632 was carried out using a modified TPY medium (m-TPY) with halved amounts of growth-supporting substrates (tryptone, peptone, and yeast extract) and replacing glucose with different dry polysaccharides (Figure 1, Section 2.3) at 0.5% (w/v) as the carbohydrate source. Positive growth control was performed for each condition using m-TPY with 0.5% glucose (w/v) selected according to previous work [6] and a negative control in m-TPY without any carbon source. The L. plantarum L12 strain was cultured using a modified MRS medium (m-MRS) with halved amounts of substrates such as peptone, yeast extract, and meat extract compared to the original medium. The potential prebiotic activity was evaluated by using the different polysaccharide samples as powder at 0.5% (w/v) in m-MRS. The positive growth control was performed using m-MRS with 0.5% glucose, while for the negative control, m-MRS without any carbon source was used. After 0, 12, 24, 30, and 48 h of incubation at 37 °C in an anaerobic atmosphere, the total bacterial count was performed on TPY or MRS agar for B. breve B632 and L. plantarum L12, respectively. The plates were incubated at 37 °C under anaerobic conditions for 48 h and the results expressed as colony-forming units per milliliter (CFU/mL). Each test was performed in triplicate.

2.8. Statistical Evaluation

A three-way ANOVA was conducted on tannin content and extract yield to assess the effects of treatment (Tr), fermentation type (TyF), and fermentation time (ST), including two- and three-way interactions. To find out the significance of the differences among samples data, Fisher’s Least Significant Difference (LSD) post hoc comparison at p < 0.05 was carried out. ANOVA and LSD analyses were carried out using OriginPro 2023b (Northampton, MA, USA). Growth data were analyzed via one-way ANOVA in R software (version 4.3.1,www.r-project.org, last accessed on 30 January 2026) with the significance threshold set at p ≤ 0.05. Homogeneous groups were determined using Tukey’s HSD test.

3. Results and Discussion

3.1. Fermentation Trials on Pomegranate Peels with and Without Addition of S. cerevisiae

Due to the limited amount of data available in the literature, the rationale of this study was to explore the main experimental variables affecting tannin profile and polysaccharide structure after fermentation with Saccharomyces cerevisiae, generating experimental data useful to guide future targeted experimental designs aimed at optimizing the peel fermentation process. Two pomegranate peels with markedly different thicknesses were selected to compare the effects of fermentation on distinct peel morphologies. Peel thickness was considered a functionally relevant parameter, as it reflects differences in the relative contribution of external tissues that vary in both structural polysaccharide composition and hydrolysable tannin content [28]. The inoculum level was selected to minimize yeast addition while remaining within the range commonly used for dough fermentation in bread production [21]. Ethanol production was used as a marker to determine the maximum fermentation time. Preliminary experiments showed minimal ethanol formation at 24 h, while extending fermentation to 96 h did not increase ethanol production compared to 72 h (Table S2). Therefore, 48 h and 72 h were selected as the sampling points, although fermentation times of five days have been often adopted in the literature [2,29,30]. To maximize yeast-driven polysaccharide depolymerization, samples for polysaccharide analysis were collected only after the maximum fermentation time (72 h). Blanching was applied prior to inoculation to partially denature endogenous enzymes and reduce the native microbial load, allowing assessment of the yeast-driven fermentation in a complex system while minimizing interference. Comparison of aerobic versus anaerobic conditions (1N vs. 3Na) allowed evaluation of the effect of oxygen availability on the fermentation process. Finally, to propose simple and low-cost procedures, no nutrients were added to the peel during fermentation.

3.2. Tannin Content in Dry Peel (DP) and Dry Extracts (DEs)

The supernatants recovered from the three fermentation methods mainly contained tannins, as these are highly water-soluble molecules, whereas polysaccharides were not co-extracted, as their recovery requires hot water extraction (Figure 1). Tannin profiles at 48 and 72 h showed excellent reproducibility both in B and Y samples (Figure S1). For each variety, the similarity of tannin profiles between B and Y samples, regardless of fermentation time, blanching pre-treatment (2PbN), or oxygen availability (1N vs. 3Na), indicated that S. cerevisiae did not alter the tannin composition in both the varieties.

The dry extract (DE) obtained by freeze-drying the fermentation supernatants yielded between 28% and 50% relative to dry peel (DP) (Figure 2). For all fermentation methods and in both varieties, the yields were significantly lower in samples inoculated with S. cerevisiae compared to the corresponding blank samples, likely due to the microbial consumption of peel components such as free sugars, mineral salts, and proteins. Notably, only dry extracts inoculated with S. cerevisiae resulted in manageable powders, whereas the corresponding blank extracts were highly hygroscopic, consistent with their higher free sugars content. Furthermore, ethanol and other organic acids were detected exclusively in the samples with S. cerevisiae, as confirmed by ¹H-NMR spectra.

Figure 2.

Yields of the supernatant collected after the fermentation processes (see Figure 1); data are a mean of triplicates expressed as % of dry extracts (DEs) on dried peel (DP). W, Wonderful variety (blue); G1, G1 variety (pink). Bar charts represent the variation as a function of type of treatment, fermentation type and fermentation time. Different letters indicate significant differences at p < 0.05.

Despite the lower extraction yields of DEs from yeast-treated samples, a significant increase in total phenolic content was obtained (Figure 3). This effect is attributed to a simple concentration phenomenon rather than to enhanced tannin extraction. In fact, the increased consumption of non-phenolic soluble components of the peel during yeast fermentation reduced the co-extraction of interfering molecules, thereby increasing the relative phenolic content in the dry extracts.

Figure 3.

Phenolic content expressed as mg/g on dried extract (DE) in Wonderful (blue–light blue) and G1 (orange–pink) varieties. Bar charts represent the variation in phenols in samples as a function of treatment, fermentation type and fermentation time. Data are expressed as average ± SD of triplicates. For each molecule, including total phenols, different letters in a graphic indicate significant differences at p < 0.05.

Analysis of individual phenolic compounds confirmed this interpretation (Figure 3; Table S1). When expressed per gram of DP, gallic acid, ellagic acid, punicalins, and punicalagins showed little or no difference between blank and yeast-treated samples, indicating that tannin recovery from the peel was largely unaffected by fermentation (Figure S2, Table S1). Conversely, when expressed per gram of DE, all major phenolics were significantly higher in yeast-inoculated samples, particularly in the Wonderful variety, reflecting the reduced DE yield (Figure 3, Table S1). Importantly, this increase reflects a higher relative concentration of phenolic compounds in the dry extracts, compared to an increase in their absolute recovery from the peel. Gallic and ellagic acids were minor phenolic compounds in the dry extracts, with maximum concentrations of approximately 30–50 mg/g in the Wonderful variety and slightly lower levels in the G1 variety. Punicalagins were the main phenolic compounds, with the highest amounts observed in the yeast-fermented samples from Wonderful, ranging from approximately 300 to 500 mg/g DE (Figure 3).

A blanching-dependent effect was observed only in the Wonderful variety, where the 2PbN samples exhibited lower punicalin and higher punicalagin levels, suggesting the partial denaturation of hydrolytic enzymes such as peel esterase. Since this conversion occurred in both B and Y samples, it is likely driven by peel enzymes rather than by S. cerevisiae. In contrast, the G1 variety displayed a different pattern, suggesting lower esterase activity in its peel.

The total tannin content expressed on DP was similar across fermentation methods within each variety, with only small statistically significant differences in some cases (Figure 3). Overall, total tannin content in the peel ranged 110–180 mg/g DP for Wonderful and 140–180 mg/g DP for G1, with maximum recovery at 72 h. In contrast, total phenols in DEs were approximately 50% higher in yeast-inoculated samples than the corresponding blanks at both timepoints, resulting in extracts containing up to 60–70% phenols for Wonderful and 50–70% for G1.

Taken together, these results demonstrate that fermentation with S. cerevisiae does not degrade or modify native pomegranate tannins but significantly enhances their concentration in the final dry extract. From an application perspective, a simple 2–3-day yeast fermentation can represent an effective strategy to increase the quality and commercial value of pomegranate peel extracts.

3.3. Chemical and Biological Properties of Polysaccharides

Decoction was applied as an effective method for recovering the polysaccharides after fermentation, as they are usually extracted using hot water, and then isolated by ethanol addition [1,8]. To evaluate if fermentation with S. cerevisiae affected polysaccharide molecular weight, the longest fermentation time (72 h), expected to maximize potential structural modification, was chosen for polysaccharides collection (Figure 1).

3.3.1. Extraction Yields

As a first step, the yields (%) of precipitated polysaccharides from Y and B samples were compared (Table 1).

Table 1.

Polysaccharides recovered after 72 h of fermentation of samples from Wonderful and G1 varieties. For each sample, triplicates were pooled to obtain a sufficient amount of polysaccharides. Percentage yields are expressed on dry peel (DP). Y, samples with S. cerevisiae; B, blank samples.

Samples	Polysaccharides Yield (% on DP)
	G1	Wonderful
B-1N	6.9	8.4
Y-1N	7.2	6.2
B-2PbN	10.5	8.3
Y-2PbN	8.3	6.3
B-3Na	8.3	10
Y-3Na	6.7	7.1

In the Wonderful variety, the highest polysaccharide yields were consistently observed in blank samples for all fermentation methods (8.3–10% on DP), while yeast-fermented samples showed lower recoveries (6.2–7.1%). A similar trend was observed in the G1 variety, except for sample Y-1N, which showed a slightly higher yield than its corresponding blank (B-1N). The polysaccharide content reported in Table 1, often expressed in the literature as pectin percentage, was consistent with values previously reported for the Wonderful variety, which ranged from 8 to 12% on a dry weight basis [7,8,31].

3.3.2. DLS Analysis

Polysaccharides extracted from unfermented peel (W-Dec and G1-Dec) were used as reference samples to compare the molecular weight of native pomegranate pectin with those obtained after fermentation. Dynamic light scattering (DLS) was applied as a preliminary screening technique for estimating the hydrodynamic size distribution of polysaccharides in water and detecting the presence of populations with different molecular dimensions [25]. It should be noted that DLS does not provide absolute molecular weight determination, particularly for heterogeneous, non-spherical polymers such as pectin. The measured hydrodynamic diameter is influenced by molecular conformation, aggregation phenomena, and the presence of minor high-molecular-size species, which can disproportionately affect scattering intensity. Therefore, DLS data were interpreted qualitatively as indicative of relative size changes rather than exact molecular weights. DLS profiles revealed that all samples were characterized by one predominant polysaccharide population (Pk1), accounting for 89–100% of the total scattering intensity (Table 2). In both varieties, polysaccharides extracted from fermented samples obtained using the 1N and 3Na methods, regardless of yeast inoculation, displayed significantly smaller Pk1 sizes compared to the corresponding unfermented references. Hydrodynamic diameters ranged from 282 to 361 nm for Wonderful and from 329 to 461 nm for G1, suggesting a reduction in polysaccharide molecular dimensions following fermentation. This trend is consistent with previous observations reporting the fermentation-induced depolymerization of pectic polysaccharides [18].

Table 2.

Results from DLS analyses of polysaccharides obtained as shown in Figure 1. PdI, polydispersity index; D, diameter measurement; Z-Pk1 average particle size of the main peak; Pk1, Pk2, populations with relative molecular size (kDa); Dec, polysaccharides from decoction of unfermented peel. nd, not detected.

Samples	PdI	Z-Pk1	Pk1	Pk 2	Pk1	Pk 2
		D (nm)	D (nm)	D (nm)	Area %	Area %
W-B-1N	0.24 ± 0.02	342 ± 42	441 ± 58	nd	100	nd
W-Y-1N	0.24 ± 0.01	361 ± 19	463 ± 31	91 ± 1	99 ± 1	0.7 ± 0.2
W-B-2PbN	0.22 ± 0.02	749 ± 24	1017 ± 53	215 ± 14	90 ± 4	10.0 ± 3.5
W-Y-2PbN	0.26 ± 0.01	491 ± 14	563 ± 66	110 ± 11	89 ± 1	10.9 ± 1.2
W-B-3Na	0.23 ± 0.01	314 ± 10	390 ± 13	nd	99 ± 1	0.4 ± 0.5
W-Y-3Na	0.22 ± 0.01	282 ± 10	343 ± 13	nd	99 ± 1	0.8 ± 0.9
W-Dec	0.27 ± 0.03	716 ± 151	1059 ± 186	186 ± 12	88 ± 12	12.2 ± 12.3
G1-B-1N	0.26 ± 0.08	329 ± 8	390 ± 25	798 ± 25	99 ± 1	0.3 ± 0.5
G1-Y-1N	0.27 ± 0.02	395 ± 5	529 ± 37	nd	100	nd
G1-B-2PbN	0.27 ± 0.02	660 ± 29	943 ± 87	115 ± 11	96 ± 6	4.5 ± 5.7
G1-Y-2PbN	0.26 ± 0.01	408 ± 10	498 ± 24	nd	100	7.8 ± 1.1
G1-B-3Na	0.29 ± 0.02	461 ± 71	526 ± 126	nd	100	nd
G1-Y-3Na	0.33 ± 0.03	369 ± 7	525 ± 35	nd	100	nd
G1-Dec	0.27 ± 0.04	652 ± 1	908 ± 7	81 ± 15	97 ± 4	2.8 ± 3.9

An exception was observed for blank samples subjected to blanching prior to fermentation (W-B-2PbN and G1-B-2PbN), which exhibited Pk1 sizes comparable to those of the unfermented reference samples. This finding suggests that the short blanching treatment partially inactivated endogenous polysaccharide-hydrolysing enzymes, thereby limiting molecular size reduction during subsequent incubation. In contrast, yeast-inoculated samples subjected to the same blanching pre-treatment (W-Y-2PbN and G1-Y-2PbN) displayed significantly smaller Pk1 sizes (563 nm and 498 nm, respectively), indicating that S. cerevisiae fermentation counteracted this effect.

These results suggest that yeast-associated or endogenous peel enzymatic activity contributed to polysaccharide depolymerization.

3.3.3. ¹H-NMR Analysis

To characterize the polysaccharides collected after fermentation, ¹H-NMR analyses were performed after chemical hydrolysis to determine the content of galacturonic acid (%) as well as the degree of acylation (DA) and methylation (DM) (Table 3).

Table 3.

Results from q¹H-NMR analyses of dialyzed polysaccharides obtained after 72 h of fermentation. Values are expressed as percentage on dried sample. DM, degree of methylation; DA, degree of acylation.

Samples	CH3OH	CH3COOH	Gal. Ac.	DM	DA
W-B-1N	5.89 ± 0.06	2.54 ± 0.11	39.65 ± 1.20	77.96	38.91
W-Y-1N	5.64 ± 0.37	2.45 ± 0.24	50.71 ± 1.10	67.48	29.34
W-B-2PbN	6.05 ± 0.26	2.72 ± 0.02	46.57 ± 0.92	78.78	35.47
W-Y-2PbN	6.22 ± 0.28	2.86 ± 0.17	50.27 ± 0.95	75.08	34.56
W-B-3Na	6.14 ± 0.26	2.95 ± 0.07	44.77 ± 0.88	69.05	39.98
W-Y-3Na	5.83 ± 0.62	2.63 ± 0.16	52.10 ± 0.78	67.91	30.82
W-Dec	5.45 ± 0.15	2.61 ± 0.03	39.77 ± 1.25	74.29	33.72
G1-B-1N	5.66 ± 0.02	2.10 ± 0.25	45.29 ± 1.40	86.66	38.11
G1-Y-1N	6.01 ± 0.60	2.88 ± 0.09	46.73 ± 0.45	75.34	35.86
G1-B-2PbN	6.38 ± 0.08	2.37 ± 0.42	55.36 ± 0.89	69.90	37.55
G1-Y-2PbN	5.77 ± 0.21	2.46 ± 0.16	55.89 ± 0.98	64.52	37.21
G1-B-3Na	6.46 ± 0.06	1.97 ± 0.27	50.97 ± 1.20	64.78	35.95
G1-Y-3Na	5.44 ± 0.38	2.65 ± 0.11	44.83 ± 0.87	73.66	37.31
G1-Dec	5.92 ± 0.15	2.49 ± 0.21	44.95 ± 0.72	79.88	37.76

¹H-NMR spectra of non-dialyzed polysaccharides from fermented samples showed significantly stronger signals in the typical monosaccharide region (3.2–4.0 ppm) in blank samples compared to yeast-fermented ones, confirming monosaccharide consumption in the presence of the yeast. Signals of citric acid (doublets at 2.5 and 2.6 ppm) were found in all fermented samples, independently of yeast addition and fermentation time, but exclusively in the peel of Wonderful variety. In all samples, weak signals around 1.1 ppm, attributed to methyl groups of rhamnose, indicated the absence of rhamnogalacturonans. Similarly, low-intensity signals at 1.9–2.0 ppm suggested a limited number of acetyl groups bound to the polysaccharides. Furthermore, DM and DA percentages (Table 3) did not show trends between samples fermented with or without S. cerevisiae. The high similarity between B and Y samples was confirmed by the similar MeOH% and CH₃COOH% values determined after acid hydrolysis. The differences observed in DA and DM values, as well as for galacturonic acid, seemed independent on fermentation, since no correlation was found between values and fermentation applied. Overall, neither fermentation nor yeast inoculation appeared to significantly affect galacturonic acid content, DM, or DA, indicating preservation of the primary chemical features of pomegranate pectin.

3.3.4. DOSY Experiments

2D-DOSY-NMR experiments provide the diffusion coefficients (D) of molecules in solution by applying increasing magnetic field gradient strength to track their translational motions, which depend on the molecular size. DOSY has been applied for studying the molar mass or hydrodynamic volume of polysaccharides and their oligomers. In particular, the study of commercial dextran from Leuconostoc spp demonstrated that DOSY analysis can be used to estimate the size of dextrans in aqueous solution even at high molecular weight [32]. DOSY was also applied to monitor the production of oligomers by enzymatic hydrolysis of ulvan from Ulva lactuca [33].

In this study, DOSY-¹H-NMR experiments were employed to complement DLS analysis and to detect the presence of lower-molecular-weight polysaccharide populations not detectable by DLS. Since the D of a molecule is affected by the physical properties of the surrounding environment such as the viscosity, DOSY experiments were performed on diluted solutions. Furthermore, since high dilution of polysaccharides with large molecular weight is not applicable, the performance, accuracy, and reproducibility of DOSY measurements are defined by the quality of the spectra along with the intensity decay curve. An optimal concentration of 3 mg/mL was identified to provide a satisfactory signal-to-noise ratio across the increasing gradient strength steps. At this concentration, the D was comparable to that obtained from a more diluted solution of the same polysaccharide, confirming the reliability of the determination and excluding viscosity-related effects.

For each experiment, the vertical trace was extracted, the area of each diffusion peak was determined, and the relative abundance percentage was calculated obtaining the distribution shown in Table 4.

Table 4.

Mean molecular weight (kDa) and relative abundance of pomegranate polysaccharides determined by DOSY ¹H-NMR. Population, group of molecules with a homogeneous size; Dec, decoction of unfermented peel. * Out of the calibration curve range.

Populations kDa
Samples	705–690	517–429	385–197	159–91	72–50	38–26	<20
W-B-1N	690 (10%) *	516 (37%)	357 (37%)	109 (13%)	-	-	-
W-Y-1N	700 (25%) *	508 (36%)	264 (17%)	98 (19%)	51 (3%)	-	-
W-B-2PbN	701 (18%) *	-	351 (42%)	94 (7%)	51 (5%)	-	<20 (28%)
W-Y-2PbN	693 (33%) *	501 (67%)	-	-	-	-	-
W-B-3Na	-	-	367 (21%)	-	72 (59%)	-	<20 (20%)
W-Y-3Na	-	-	385 (31%)	159 (40%) 108 (25%)	69 (4%)	-
W-Dec	705 (96%) *	-	-	-	-	-	-
G1-B-1N	-	509 (11%)	266 (13%)	-	70 (13%)	-	<20 (63%)-
G1-Y-1N	-	429 (4%)	265 (59%)	155 (16%)	68 (10%)	-	<20 (11%)
G1-B-2PbN	-	-	278 (9%) 197 (10%)	96 (14%)	-	38 (10%) 26 (10%)	<20 (47%)
G1-Y-2PbN	-	499 (69%)	357 (26%)	-	67 (5%)	-	-
G1-B-3Na	-	500 (20%)	356 (27%)	138 (24%)	69 (11%)	38 (18%)	-
G1-Y-3Na	-	495 (17%)	367 (36%)	91 (17%)	-	34 (4%)	<20 (26%)
G1-Dec	-	517 (39%)	367 (42%)	-	71 (7%)	-	<20 (12%)

DOSY results confirmed that polysaccharides from W-Dec were homogeneous in size, with a molecular weight tentatively estimated at around 700 kDa. In contrast, polysaccharides from G1-Dec were more heterogeneous, with two major populations around 500 and 360 kDa, and additional species below 20 kDa. All fermented samples showed a reduced molecular weight compared to native polysaccharides (W-Dec and G1-Dec). Unlike DLS, DOSY experiments displayed the presence of several additional polysaccharides at reduced molecular weight (below 72 kDa) in almost all fermented samples, confirming the greater performance of DOSY analysis for polysaccharides characterization. This observation confirms that while DLS is suitable as a preliminary screening tool, it fails to detect species below approximately 150 kDa, as confirmed by using dextran standards of known molecular weight.

The polysaccharide patterns resulting from the three fermentation methods was different between the two varieties, possibly due to differences in peel thickness (thicker in Wonderful and thinner in G1) and the different availability of the nutrients for microbial growth. In Wonderful samples, fermentation under air-exposure conditions (3Na) greatly reduced the size of native polysaccharides, yielding 59% in the 72–50 kDa range in blank samples and 65% in the 159–91 kDa range in yeast-fermented samples. In contrast, the highest concentration of polysaccharides below 159 kDa was observed in the G1-B-1N sample (sum 76%), confirming the extensive depolymerization of G1 pectin under these conditions.

For each fermentation method, comparable depolymerization patterns were observed in both B and Y samples. In particular, the highest number of populations below 20 kDa in G1 were found in the three blank samples (Table 4). The photograph provided by the DOSY spectra highlighted the strong reduction in molecular weight of pectin after fermentation, leading to the formation of oligosaccharide-rich extracts.

3.3.5. In Vitro Potential Prebiotic Activity on B. breve and L. plantarum

To investigate the fermentability of polysaccharides from pomegranate peels, the ability of B. breve B632 and L. plantarum L12 strains to use polysaccharides extracted from both fermented and non-fermented pomegranate peel as carbon sources was investigated. Bacterial growth on these substrates was compared with that observed on glucose (used as a positive control, as it is a readily fermentable carbon source) and on a sugar-free medium (used as a negative controls).

Both Bifidobacterium and Lactobacillaceae strains are known to compete with enteric pathogens for nutrients and adhesion sites on the intestinal mucosa, while also stimulating the development of the mucosal immune system [34,35]. Furthermore, B. breve B632 is known for its anti-inflammatory activity and ability to colonize the human gut as well as for the capacity to protect the gut epithelial integrity [36]. Recent studies demonstrated the prebiotic properties of oligosaccharides produced after hydrolysis of pomegranate pectin. Devecioglu et al. [31] showed that enzymatic hydrolysates of pomegranate pectin promoted the growth of Levilactobacillus brevis and enhanced the production of γ-aminobutyric acid [31]. Similarly, Bachari et al. [37] demonstrated that ultrasonic treatment induced partial hydrolysis of pomegranate peel pectin, significantly reducing its molecular weight and promoting the growth of Bifidobacterium longum and Lactobacillus casei.

In vitro assay represents a simplified and preliminary screening model aimed at comparing the relative fermentability of different carbon sources and highlighting differences in bacterial utilization between fermented and non-fermented pomegranate peel polysaccharides. It does not reproduce the complexity of colonic fermentation, including microbial cross-feeding interactions, host responses, or metabolite production. Nevertheless, this approach provides useful initial evidence of improved accessibility and microbial utilization of pomegranate pectin following fermentation-induced depolymerization.

The results of the in vitro growth assays with the B. breve B632 strain are shown in Table 5. In the presence of glucose, a 1 × 10² CFU/mL increase was observed after 12 h compared to time 0, reaching 3.5 ± 0.21 × 10⁸ CFU/mL. Cell counts slightly increased after 24 h, approaching a growth plateau. At 12 h, all other fermented polysaccharide growth substrates showed bacterial concentrations around 10⁷ CFU/mL—lower than that of glucose, but higher than at time 0, and significantly greater than the negative control (no carbon source). Fermented samples also showed higher cell counts than those of the native polysaccharides from non-fermented samples (W-Dec and G1-Dec). After 24 h, notable differences emerged among samples: in both varieties, all samples fermented with S. cerevisiae exhibited higher cell counts (10⁸ CFU/mL) compared to samples fermented without yeast inoculation (10⁷ CFU/mL). Thus, S. cerevisiae-inoculated fermentation samples produced more easily fermented oligosaccharides than spontaneous fermentation with native microbiota (B samples). The highest growth was observed in aerobic fermentation samples (W-Y-3Na and G1-Y-3Na), reaching 10⁹ CFU/mL at 30 h, even exceeding the positive control with glucose. After 48 h, a slight decrease in growth was observed across all samples.

Table 5.

Evaluation of the potential prebiotic properties of pomegranate peel from Wonderful and G1 cultivars on Bifidobacterium breve B632. Different letters indicate significant difference between the different fermentation samples according to HSD Tukey’s test; ns, ** and ***: effect not significant or significant at p ≤ 0.01 or p ≤ 0.001.

B. breve B632	0 h	12 h	24 h	30 h	48 h
	CFU/mL	CFU/mL	CFU/mL	CFU/mL	CFU/mL
W-Y-3Na	(2.1 ± 0.2) × 105	(3.5 ± 0.2) × 107 b	(9.8 ± 0.2) × 108 a	(2.1 ± 0.3) × 109 a	(8.4 ± 0.3) × 108 a
W-Y-1N	(2.2 ± 0.2) × 105	(2.7 ± 0.2) × 107 b	(6.6 ± 0.2) × 108 a	(8.7 ± 0.2) × 108 c	(2.1 ± 0.3) × 108 e
W-Y-2PbN	(2.1 ± 0.2) × 105	(3.1 ± 0.2) × 107 b	(8.3 ± 0.3) × 108 b	(9.3 ± 0.4) × 108 c	(5.4 ± 0.3) × 108 c
W-B-3Na	(2.4 ± 0.2) × 105	(3.3 ± 0.2) × 107 b	(4.4 ± 0.4) × 107 g	(3.4 ± 0.3) × 108 de	(3.6 ± 0.2) × 107 g
W-B-1N	(2.2 ± 0.2) × 105	(2.5 ± 0.2) × 107 b	(1.8 ± 0.1) × 107 g	(8.6 ± 0.2) × 107 ef	(1.2 ± 0.3) × 107 g
W-B-2PbN	(2.4 ± 0.1) × 105	(2.8 ± 0.1) × 107 b	(2,9 ± 0.2) × 107 g	(1.3 ± 0.3) × 108 ef	(2.5 ± 0.2) × 107 g
G1-Y-3Na	(2.3 ± 0.2) × 105	(3.4 ± 0.2) × 107 b	(7.1 ± 0.3) × 108 c	(1.7 ± 0.2) × 109 b	(6.1 ± 0.3) × 108 b
G1-Y-1N	(2.1 ± 0.1) × 105	(2.9 ± 0.2) × 107 b	(3.4 ± 0.2) × 108 f	(4.9 ± 0.3) × 108 d	(1.3 ± 0.1) × 108 f
G1-Y-2PbN	(2.2 ± 0.2) × 105	(3.0 ± 0.3) × 107 b	(6.6 ± 0.3) × 108 d	(8.8 ± 0.2) × 108 c	(3.2 ± 0.2) × 108 d
G1-B-3Na	(2.6 ± 0.2) × 105	(3.0 ± 0.3) × 107 b	(4.7 ± 0.3) × 107 g	(5.4 ± 0.2) × 108 d	(3.0 ± 0.3) × 107 g
G1-B-1N	(2.4 ± 0.2) × 105	(2.6 ± 0.3) × 107 b	(1.4 ± 0.3) × 107 g	(8.8 ± 0.3) × 107 ef	(1.0 ± 0.5) × 107 g
G1-B-2PbN	(2.4 ± 0.1) × 105	(2.7 ± 0.4) × 107 b	(3.2 ± 0.4) × 107 g	(1.2 ± 0.2) × 108 ef	(1.7 ± 0.3) × 107 g
W-Dec	(2.5 ± 0.2) × 105	(8.4 ± 0.3) × 105 c	(8.4 ± 0.2) × 106 g	(8.2 ± 0.3) × 106 f	(6.7 ± 0.6) × 106 g
G1-Dec	(2.3 ± 0.2) × 105	(7.7 ± 0.2) × 105 c	(6.8 ± 0.1) × 106 g	(7.6 ± 0.2) × 106 f	(4.5 ± 0.3) × 106 g
No Carbon source	(2.1 ± 0.3) × 105	(2.6 ± 0.2) × 105 c	(3.8 ± 0.2) × 106 g	(4.7 ± 0.3) × 106 f	(1.3 ± 0.2) × 106 g
Glucose	(2.2 ± 0.1) × 105	(3.5 ± 0.2) × 108 a	(4.3 ± 0.2) × 108 e	(3.8 ± 0.3) × 108 de	(2.5 ± 0.2) × 108 e
	ns	***	***	**	***

A similar trend was observed for L. plantarum L12 (Table 6). Growth with glucose was rapid after 12 h (8.5 ± 0.15 × 10⁸ CFU/mL), reaching a maximum at 24 h (8.3 ± 0.16 ×10⁹ CFU/mL), followed by a plateau. Similarly to the previous case, all Y samples exhibited higher cell counts compared to B samples. High growth occurred in the W-Y-3Na and G1-Y-3Na samples after 12 h, with cell count of 3.4 ± 0.2 × 10⁸ CFU/mL and 4.3 ± 0.2 × 10⁸ CFU/mL respectively. After 30 h, all S. cerevisiae-fermented samples reached 10⁸ or 10⁹ CFU/mL. Again, the samples with no carbon source and those from unfermented peel (W-Dec and G1-Dec) exhibited a similar growth significantly lower than that of blank samples from fermentation. Although slightly higher growth was observed in G1 samples throughout the experiment, no significant differences were found between the different fermentation conditions. As for the B. breve strain, L. plantarum growth was higher in yeast-fermented samples compared with those undergoing spontaneous fermentation (B samples), with the highest values observed for aerobic 3Na fermentation.

Table 6.

Growth assay test of pomegranate peel from Wonderful and G1 cultivars on Lactobacillus plantarum L12. Different letters indicate significant difference between the different fermentation samples according to HSD Tukey’s test; ns, ** and ***: effect not significant or significant at p ≤ 0.01 or p ≤ 0.001.

L. plantarum L12	0 h	12 h	24 h	30 h	48 h
	CFU/mL	CFU/mL	CFU/mL	CFU/mL	CFU/mL
W-Y-3Na	(1.8 ± 0.1) × 105 a	(3.4 ± 0.2) × 108 c	(4.7 ± 0.3) × 109 d	(4.2 ± 0.3) × 109 c	(2.3 ± 0.2) × 109 c
W-Y-1-N	(1.5 ± 0.2) × 105 a	(1.5 ± 0.2) × 108 e	(3.8 ± 0.2) × 109 e	(3.5 ± 0.2) × 109 d	(1.5 ± 0.4) × 109 b
W-Y-2PbN	(1.6 ± 0.2) × 105 a	(2.4 ± 0.2) × 108 d	(3.4 ± 0.4) × 109 e	(3.4 ± 0.2) × 109 d	(1.9 ± 0.2) × 109 cd
W-B-3Na	(1.7 ± 0.2) × 105 a	(8.4 ± 0.2) × 107 f	(5.4 ± 0.3) × 108 fg	(3.7 ± 0.2) × 108 ef	(2.8 ± 0.3) × 108 e
W-B-1N	(1.4 ± 0.2) × 105 a	(7.5 ± 0.2) × 107 f	(3.5 ± 0.3) × 108 fgh	(1.4 ± 0.2) × 108 ef	(1.8 ± 0.2) × 108 e
W-B-2PbN	(1.7 ± 0.2) × 105 a	(7.8 ± 0.6) × 107 f	(4.6 ± 0.2) × 108 fgh	(1.9 ± 0.2) × 108 ef	(2.1 ± 0.4) × 108 e
G1-Y-3Na	(1.6 ± 0.2) × 105 a	(4.3 ± 0.2) × 108 b	(7.7 ± 0.4) × 109 b	(6.4 ± 0.3) × 109 b	(4.2 ± 0.2) × 109 a
G1-Y-1N	(1.3 ± 0.2) × 105 a	(3.6 ± 0.2) × 108 c	(6.5 ± 0.3) × 109 h	(6.3 ± 0.6) × 109 b	(3.3 ± 0.1) × 109 b
G1-Y-2PbN	(1.7 ± 0.2) × 105 a	(2.3 ± 0.4) × 108 d	(7.1 ± 0.3) × 109 b	(6.8 ± 0.2) × 109 b	(4.4 ± 0.4) × 109 a
G1-B-3Na	(1.5 ± 0.2) × 105 a	(7.8 ± 0.2) × 107 f	(5.8 ± 0.2) × 108 f	(5.1 ± 0.3) × 108 e	(4.6 ± 0.3) × 108 e
G1-B-1N	(1.8 ± 0.2) × 105 a	(4.8 ± 0.2) × 107 fg	(2.6 ± 0.3) × 108 fgh	(2.1 ± 0.3) × 108 ef	(1.8 ± 0.3) × 108 e
G1-B-2PbN	(1.9 ± 0.2) × 105 a	(6.6 ± 0.4) × 107 f	(5.9 ± 0.3) × 108 f	(5.7 ± 0.3) × 108 e	(2.7 ± 0.3) × 108 e
W-Dec	(1.7 ± 0.2) × 105 a	(8.4 ± 0.3) × 105 g	(8.4 ± 0.4) × 106 gh	(8.2 ± 0.2) × 106 f	(6.7 ± 0.1) × 106 e
G1-Dec	(1.6 ± 0.2) × 105 a	(8.9 ± 0.2) × 105 g	(8.9 ± 0.2) × 106 gh	(1.5 ± 0.2) × 107 f	(2.4 ± 0.2) × 107 e
No Carbon source	(1.8 ± 0.2) × 105 a	(6.5 ± 0.1) × 105 g	(5.7 ± 0.2) × 106 b	(8.6 ± 0.3) × 106 f	(1.4 ± 0.3) × 107 e
Glucose	(1.4 ± 0.2) × 105 a	(8.5 ± 0.2) × 108 a	(8.3 ± 0.2) × 109 a	(7.8 ± 0.2) × 109 a	(4.5 ± 0.3) × 108 e
S	ns	***	**	***	***

Taken together, these results demonstrate that polysaccharides derived from fermented pomegranate peels support the growth of both B. breve and L. plantarum, with a clear enhancement associated with S. cerevisiae inoculation and oxygen exposure during fermentation. The observed effects were consistent across two bacterial strains and two pomegranate varieties, indicating that controlled fermentation can enhance the health-promoting potential of peel-derived polysaccharides.

The ability of polysaccharides to selectively stimulate the growth of one or a limited number of bacteria in the colon is strongly influenced by their molecular weight. Several studies have demonstrated that pectic oligosaccharides and partially depolymerized pectins are utilized more readily by the gut microbiota than native, high-molecular-weight pectins. DOSY-¹H-NMR analysis of fermented samples, particularly those obtained under aerobic conditions, clearly revealed the presence of variable populations with polysaccharides fractions below 100 kDa. These fractions fall within the molecular weight range considered optimal for prebiotic activity [38].

The different growth patterns observed for B. breve and L. plantarum strains can be explained by their distinct carbohydrate utilization strategies. Bifidobacterium species primarily metabolize low-molecular-weight oligosaccharides, whereas members of Lactobacillaceae are capable of metabolizing larger partially depolymerized polysaccharides through their extracellular or cell-associated enzymes. As a result, B. breve shows greater sensitivity to fermentation-induced depolymerization, while L. plantarum exhibits a broader growth response [39,40].

Unlike commercial pectin, which generally has a high molecular weight and show limited fermentability, the fermented pomegranate peel polysaccharides described in this study display a heterogeneous molecular weight distribution. This pattern resembles that of pectic oligosaccharides and other established prebiotics, including fructooligosaccharides and galactooligosaccharides. These characteristics indicate their potential application as functional ingredients with prebiotic activity.

4. Conclusions

The present study proposes an innovative and sustainable strategy for the valorization of pomegranate peel, a major by-product of juice production, through controlled fermentation aimed at generating high-value functional ingredients for nutraceutical applications. The three investigated fermentation methods demonstrated excellent reproducibility at the laboratory scale and yielded extracts with comparable tannin contents, with only minor differences between yeast-inoculated and non-inoculated conditions.

While fermentation did not alter the chemical structure of tannins, S. cerevisiae inoculation resulted in a marked increase in total phenolic content in dry extracts through the consumption of oligosaccharides and other fermentable substrates of the peel.

The use of Diffusion-Ordered Spectroscopy Proton Nuclear Magnetic Resonance (DOSY-¹H-NMR) and Dynamic Light Scattering (DLS) enabled effective monitoring of fermentation-induced depolymerization in peel polysaccharides. DLS proved to be a valuable and rapid screening method for tracking polysaccharide depolymerization. DOSY-¹H-NMR provided insights into the co-presence of distinct oligomer and polymer populations and revealed significant molecular weight reduction in native pectin after fermentation. Importantly, all the applied yeast-fermentation methods significantly reduced pectin size compared to the native structures accounting for the enhanced in vitro growth of the target gut strains widely recognized as probiotic species.

Although the prebiotic properties of hydrolysed pectin and pectic oligosaccharides are well-documented, their depolymerization is typically achieved through chemical or enzymatic treatments and not by fermentation. Among the tested approaches, aerobic fermentation showed strong potential for future scale-up due to its operational simplicity, low water and energy requirements, and minimal processing steps.

This study demonstrates that controlled fermentation can be an effective dual approach and a low-impact bioprocessing strategy capable of inducing pectin depolymerization directly within a complex agri-food matrix without the need for external hydrolytic treatments. Overall, the study highlights controlled fermentation as a sustainable and economically feasible strategy for converting pomegranate peel into functional ingredients with potential prebiotic activity, thereby supporting circular bioeconomy and waste valorization initiatives.

Abbreviations

The following abbreviations are used in this manuscript:

UAE	ultrasound-assisted extraction
MAE	microwave-assisted extraction
SFE	supercritical fluid extraction
DE	dry extract.
DP	dry peel
HPLC-DAD	high-performance liquid chromatography-diode array detector
DOSY-1H-NMR	Diffusion-Ordered Spectroscopy Proton Nuclear Magnetic Resonance
DLS	Dynamic Light Scattering

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/foods15040605/s1, Figure S1. Profiles at 370 nm of the triplicates of samples from G1 variety after 48 h of fermentation: B, blank sample and Y sample with yeast. Figure S2. Phenolic content expressed as mg/g on dried peel (DP) in Wonderful (blue–light blue) and G1 (orange–pink) varieties. Bar charts represent the variation in phenols in samples as a function of treatment, fermentation type and fermentation time. Data are expressed as average ± SD of triplicates. For each molecule, including total phenols, different letters in a graphic indicate significant differences at p < 0.05. Table S1. Data processing of the main tannins analyzed for Wonderful (A) and G1 (B) varieties. For each variable, results from three-factor ANOVA are reported, where the factors are the treatment (Tr), the type of fermentation (TyF) and the fermentation time (ST). The two-way and three-way interactions are also reported. Table S2. Percentage of ethanol after fermentation by Saccharomyces c, at 48 h and 72 h.

Author Contributions

Conceptualization: M.K. and N.M.; methodology: M.K. and M.B.; software, L.C.; validation: M.K., N.M., M.B. and L.C.; formal analysis: L.C.; investigation, B.Z., S.D., D.B., E.T., E.P., and D.D.G.; resources, N.M., D.B. and D.D.G.; data curation, M.K., L.C., M.B., B.Z. and E.P.; writing—original draft preparation, M.K., L.C., D.B., D.D.G. and N.M.; writing—review and editing: M.K., L.C., D.B., D.D.G. and N.M.; visualization: M.B., E.T., S.D., E.P. and B.Z.; supervision: N.M.; project administration: N.M.; funding acquisition: N.M. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article/Supplementary Material. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

Authors declared no conflicts of interest.

Funding Statement

This research was supported by the Italian Ministry of Education, University and Research (MIUR) and funded by European Union—Next Generation EU (project PRIN2022 MUR, N° 2022X3WZAF). The APC was funded by project PRIN2022 MUR, N° 2022X3WZAF.