1. Introduction
The need for environmentally friendly processing has led to the introduction of green techniques to extract active principles from different plant materials. The current direction aligns with the idea of eco-friendly extraction, which embraces methods aiming to reduce both energy and solvent usage and employ environmentally sustainable solvents [1].
The processing of wood, whether in the timber industry or the pulp and paper industry, generates significant woody by-products, including leaves, stubs, branches, and notably, bark. These residues, due to their richness in biomolecules and bioactive compounds, could be repurposed [2]. The tanning industry stands out as an appealing sector for the utilization of bark acquired as a byproduct. Before the 19th century, natural extracts from wood, barks, leaves, and roots were used in the tanning process of leather [3]. Since then, vegetable tannins have been displaced by chemical tanning agents, such as chromium salts, which are the most used in the worldwide leather manufacturing industry [4]. This is due to the great characteristics they impart to leather products, including physical and mechanical properties and thermal stability [5]. Nevertheless, the large quantities of chrome-containing waste generated during the tanning process raises numerous environmental and health concerns [6]. Considering the detrimental impact of chrome tanning, the current trend is to return to vegetable tannins. The latter are considered eco-friendly, sustainable, and cost-effective. Barks, a by-product of the wood industry, are rich in vegetable tannins (polyphenols), which can be recovered for use in the leather manufacturing industry [7,8,9]. Furthermore, these natural compounds can be easily extracted using green solvents, such as water.
Pedunculate oak (Quercus robur) is a tree belonging to the Euquercus subgenus, section Lepidobalanus. It grows spontaneously in deciduous forests in temperate regions around the world, being native to Europe and Asia [10]. Among its various compounds, oak bark contains hydrolysable and condensed tannins, which are secondary metabolites included in the class of polyphenols [11]. Extracts derived from oak bark exhibit antioxidant [12], antibacterial [13], anti-inflammatory [14], and anticancer [15] effects.
Oak bark, being a lignocellulosic plant material, requires severe treatment conditions to extract the desired compounds [16]. However, subjecting it to such conditions raises the risk of degradation or alteration of the chemical structure of the extracted compounds. An effective technique that could preserve the chemical structure and properties of polyphenols, while allowing for the easy release of targeted compounds from the plant matrix, is ultrasound-assisted extraction (UAE). This method, due to the cavitation phenomenon that occurs in heterogeneous environments, enhances the mass transfer rate [17]. UAE has been demonstrated to be more efficient than conventional methods such as Soxhlet extraction [18] and maceration [19], and it is scalable for industrial applications. Zhang et al. optimized the UAE process for extracting tannins from Acer truncatum seed shells, illustrating their utility for removing cation dyes from water [20]. Duarte et al. employed deep eutectic solvents to obtain polyphenol-enriched extracts from maritime pine residues via UAE. These extracts exhibited significant antioxidant and antimicrobial activities [21]. de Souza Ribeiro et al. optimized the UAE of phenolic compounds from Barbatimao bark, concluding that better yields are achieved compared with conventional methods [22]. Additionally, they evaluated the cytotoxicity and antioxidant cell activity of the extracts, concluding that those obtained via UAE promoted cell viability above 80%, whereas the extracts obtained by the conventional method did not demonstrate that level of activity. Based on these findings, the authors suggest that the extracts could serve as an alternative source of tannins in various fields, including wine, pharmaceuticals, cosmetics, etc.
The aim of this study was to develop a green and efficient pilot-scale extraction technique of polyphenols from oak bark by means of UAE. To achieve this, a modified closed loop Dual-Frequency Reactor (DFR) equipment was employed to ensure mild extraction conditions. Additionally, experiments without ultrasound were carried out to highlight the effectiveness of this technique. The resulting extracts were analyzed physically (determination of extraction yield), chemically (determination of total phenolic content—TPC), microbiologically (determination of antimicrobial activity), and spectroscopically (FTIR analysis). The spectra were further analyzed using principal component and cluster analysis analyses (PCA) to emphasize the influence the extraction conditions have on the molecular profile of the extracts.
The novelty of this study lies in the use of a DFR in a loop system for UAE. The literature only presents multi-frequency ultrasonic reactors using two sonotrodes with different frequencies [23] or a sonotrode in an ultrasound bath [24]. The DFR used in this study has two magnetostrictive transducers with frequencies of 16 and 20 kHz and a diaphragm plate with a diameter of 4 inches. Sound waves from the opposing diaphragm plates superimpose to amplify each other within the dual-frequency processing chamber, which has volumes of 25, 250 or 600 mL. The abnormally high relative pressure amplitude exceeds the capability of a single resonant frequency. The beat frequency rippling effect moves this intense wave throughout the processing cavity for uniform treatment.
Moreover, the loop reactor allows for the treatment of large volumes while ensuring high-intensity ultrasound. By increasing the loading tank capacity, this strategy enables easy scalability of the plant without losing the efficiency of ultrasound treatment.
2. Materials and Methods
2.1. Materials
Oak bark (Quercus robur) was supplied by Hofigal S.A., Bucharest, Romania. The raw material underwent drying in an air-flow heating oven at 40 °C until it reached a constant weight, resulting in a final humidity of 14%. Subsequently, the dried bark was finely milled using an electric grinder and sieved to reach particles smaller than 400 μm. The ground oak bark was then stored in sealed containers at 4–5 °C until it was used in the extraction of polyphenols.
The standard used for determining polyphenols content was gallic acid, purchased from Sigma-Aldrich Co., Bucharest, Romania. Folin–Ciocalteu reagent, sodium carbonate, sodium hydroxide, sodium sulfite, and sodium bisulfite, all of analytical grade, were purchased from Merck, Darmstadt, Germany. The standard extract of tannin, from oak bark, used in the analysis of FTIR spectra was purchased from Silvateam S.p.A, San Michele Mondovi, Italy. The culture media (Mueller-Hinton agar, Sabouraud dextrose agar, MacConkey broth, and casein soya bean digest broth) were purchased from Novachim, Bucharest, Romania. The Escherichia coli and Staphylococcus aureus strains were purchased from Mediclim SRL, Bucharest, Romania.
The standard used for determining polyphenols content was gallic acid, purchased from Sigma-Aldrich Co., Bucharest, Romania. Folin–Ciocalteu reagent, sodium carbonate, sodium hydroxide, sodium sulfite, and sodium bisulfite, all of analytical grade, were purchased from Merck, Darmstadt, Germany. The standard extract of tannin, from oak bark, used in the analysis of FTIR spectra was purchased from Silvateam S.p.A, San Michele Mondovi, Italy. The culture media (Mueller-Hinton agar, Sabouraud dextrose agar, MacConkey broth, and casein soya bean digest broth) were purchased from Novachim, Bucharest, Romania. The Escherichia coli and Staphylococcus aureus strains were purchased from Mediclim SRL, Bucharest, Romania.
2.2. Extraction Procedure
The extraction of polyphenols from oak bark was carried out in two different installations:
A DFR apparatus (Advanced Sonics Processing Systems, Oxford, CT, USA) equipped with a 600 mL batch reactor (Figure 1), with the experiments being performed by introducing ultrasounds into the system (UAE) or without introducing ultrasounds into the system (Control);
A thermostatic water bath, with the experiments being carried out in a glass round-bottom flask connected to a condenser (CONV).
The ultrasound frequencies of DFR are 16 and 20 kHz, and the maximum power of the two generators is approximately 600 W. The equipment was modified to enable a loop extraction process: the apparatus was fitted with a loading tank (5000 mL) for the plant material and solvent. From this tank, the extraction mixture was pumped into the batch reactor, where it was exposed to ultrasound, and then it returned to the loading tank. The extraction mixture was continuously circulated between the ultrasound reactor and the loading tank until the desired extraction time was reached. The raw material was thus intermittently subjected to ultrasound, helping to avoid a potential degradation of the bioactive compounds.
The establishment of the best extraction conditions for polyphenols (tannins) involved three steps (Table 1).
The plant-material-to-solvent ratio was maintained at 1/10 (w/v) for all experiments. The extraction was carried out at a temperature of 50 °C for both UAE and Control. The CONV was conducted for 30 min at a temperature of 70 °C. Due to the loop design, the extraction mixture underwent sonication for 15 s every 60 s of extraction. The Control experiment, conducted to evaluate the efficiency of UAE, was carried out under the same extraction conditions and with the same equipment (DFR) used in the UAE technique, but without introducing ultrasound into the system. After extraction by each method, the mixture was centrifuged for 15 min at 2500 rpm, and the supernatant was stored at 4–5 °C until it was used for characterization.
The objective of these extractions was to obtain extracts with hide-tanning capacity (their tanning ability is the subject of a separate study); thus, both polyphenols (vegetable tannins) and dry extract were quantified. The FTIR analysis and the grouping of spectra by the PCA method allowed for the correlation of the polyphenolic content with the extraction conditions.
The aqueous solution was used as the extraction solvent in order to generate a green tanning agent. The use of very low amounts of sodium hydroxide, sodium sulfite, and bisulfite sought to partially hydrolyze the lignocellulose, allowing polyphenols to be released from the plant matrix [25,26]. Furthermore, research evidence suggests that sodium hydroxide concentrations up to 0.22 and 10% are appropriate for the extraction of hydrolysable [27] and condensable [28] tannins, respectively.
2.3. Determination of Extraction Yield
The extraction yield was determined for all methods used. For this purpose, a portion of the supernatant was weighed and subsequently evaporated using a rotavapor. The dry extract was weighed and expressed as milligrams per gram of dry matter (mg/g DM).
2.4. Total Phenolic Content Determination
With slight adjustments, the TPC was determined colorimetrically following the guidelines outlined in the International Standard ISO 14502-1 [29], as detailed in our previous work [30]. Absorbance was measured at 760 nm. TPC quantification relied on a standard curve which corresponds to 1–5 mg/mL gallic acid solution. The polyphenolic concentration of the extracts was expressed as milligrams of gallic acid equivalents per 1 g of dry matter (mg GAE/g DM). It is important to note that the Folin–Ciocalteu reagent used in this determination is less selective. Consequently, besides tannins, the reagent may be reduced by other polyphenols found in oak bark, as well as non-phenolic compounds. Despite this limitation, the assessment provides a preliminary evaluation of the extracts and can be employed to establish the influence of various parameters on the extraction efficiency.
For TPC and extraction yield determinations, triplicate measurements (n = 3) were performed. The data were reported as the mean value ± SD (standard deviation) and used to evaluate the dissimilarities by univariate one-way ANOVA analysis. Significant statistical variations between the averages of TPC and dry extract amount were identified as reported in our previous work [31]. Differences were considered statistically significant at a p-value below 0.05. The statistical analysis was conducted using XLSTAT Version 2019.1 (Addinsoft, New York, NY, USA).
2.5. FTIR Analysis of the Extracts
Infrared Spectroscopy in Attenuated Total Reflection mode (FTIR-ATR) analyses were carried out using an ALPHA spectrometer (Bruker Optics, Leipzig, Germany) equipped with a Platinum ATR module. The penetration depth, depending on the refractive indices of the ATR crystal and the sample, typically amounts to a few microns (approximately 0.5–3 μm). Spectra were recorded in the 4000–400 cm−1 spectral range with a 4 cm−1 resolution, using 32 scans. Opus software Version 7.0 (Bruker Optics, Leipzig, Germany) was used for the acquisition and elaboration of the spectra, while their deconvolution was performed using PeakFit 4.1 (Jandel Scientific, SanRafael, CA, USA). Multiple peaks of the investigated samples were deconvoluted using the PeakFit asymmetric Gaussian fit function to improve the fit of the asymmetry in the peaks.
The PCA used for multivariate analysis of FTIR-ATR spectra to explore similarities and hidden patterns among the composition of the extracts depending on the extraction conditions is the function PCA implemented in MATLAB® (MathWorks, Natick, MA, USA—2024a Documentation). The original set of variables is reduced to a smaller set of linear combinations of the former that account for most of the variance. The first principal component accounts for the largest amount of the total variation, the second principal component accounts for the second largest amount of the total variation, and so on. The focus of PCA is to find out the smallest number of components that account for most of the variability in the original data. The former gives the number of clusters into which spectra are classified. Prior to multivariate analysis, the infrared spectra were smoothed (first polynomial and 15 points per window), normalized (standard normal variation—SNV), derived (first-order, second polynomial and 15 points per window), and mean-centered. The PCA technique was used for multivariate analysis of the FTIR-ATR spectra in the range of 1800–800 cm−1 [32,33].
2.6. Antibacterial Activity and Microbial Contamination of the Extracts
A variety of phenotypic and genotypic antimicrobial susceptibility testing (AST) methods are used in clinical laboratories to detect bacterial susceptibility to conventional antibiotics. Since plant-derived compounds are complex mixes of many compounds, it is expected that may not act as expected in the test system. For this reason, only a few AST methods have found application in determining antibacterial activity of natural products [34]. We selected the agar well diffusion assay to screen the extracts for their antimicrobial potency immediately after extraction, without any preparation. This option is determined by the desire to employ the extracts as such in the tanning process.
The agar well diffusion method on Mueller–Hinton agar (MHA) according to EUCAST Guidelines [35] was used. Escherichia coli (E. coli.; ATCC 11229) and Staphylococcus aureus (S. aureus; ATCC 6538) were used as references for the antibacterial activity of oak bark extracts. Both are included in the ESKAPE pathogens (Enterococcus faecium, Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumannii, Pseudomonas aeruginosa, and Enterobacter species), the leading cause of nosocomial infections throughout the world.
Briefly, MHA agar plates were inoculated with bacterial strain under aseptic conditions and wells (diameter = 6 mm) were filled with 50 µL of the test extract, then incubated at 37 °C for 24 h. The antibacterial agent diffuses in the agar medium and inhibits the growth of the microbial strain tested. Subsequently, the inhibition zone diameters were measured as triplicates (n = 3) achieved for the same tannin extract. The initial inoculum for the tested bacteria reached approximately 1.5 × 108 colony forming units (CFU)/mL (turbidity 0.5 McFarland scale). The turbidity of the bacterial suspension was measured at 600 nm. The diameter of the inhibition zone was calculated as follows:
H = (D − d)/2;
H = inhibition zones diameter, mm;
D = total diameter of the disc and the area of inhibition, mm;
d = diameter of the disc, mm.
To assess the microbiological quality of the extracts, microbial enumeration tests for total aerobic microbial counts (TAMC) and total yeast and mold counts (TYMC) in 1 mL of decimal dilutions were performed according to European Pharmacopoeia Ed. 10/20 guidelines. Microbiological assessment of non-sterile products such as extracts is particularly pertinent since microbial contamination can reduce or even eliminate their antimicrobial effect. The viable aerobic mesophile bacteria count was performed by plating 1 mL of decimal dilutions on casein soy agar. Plates were incubated at 30 °C for 2 days. The molds and yeast count were performed by plating 1 mL of decimal dilutions on Sabouraud dextrose agar. Plates were incubated at 25 °C for 3–5 days. Three Petri dishes (triplicates) were used for each medium. The results are displayed as CFU/g of sample.
For qualitative examination, i.e., as well as the presence of E. coli and S. aureus, 10 mL of prepared sample was added to 100 mL of casein soya bean digest broth. The procedure was dependent on the determination of the absence or limited occurrence of specified microorganism that may be detected:
S. aureus—incubation (24 h, 35 °C) and transmission on Mannitol Salt Agar.
E. coli—incubation (24 h, 35 °C) and transmission to MacConkey broth (incubation 48 h, 35 °C), followed by transmission on MacConkey agar.
3. Results and Discussion
3.1. Ultrasound-Assisted Extraction
An efficient strategy to treat large reaction volumes using equipment capable of ensuring high-intensity ultrasound only in small reactors is the use of a loop-reactor-type system. This allows for the scalability of the plant without losing the efficiency of ultrasound treatment [36]. Therefore, UAE was performed using the DFR equipment presented in Figure 1.
The mass transfer rate can increase because of the cavitation phenomenon, which can promote the breakdown of cell walls in heterogeneous mixtures, such as solid–liquid extraction of active principles from plant materials. Specifically, this phenomenon implies the collapse of cavitation bubbles close to the plant material’s surface, accompanied by microjets towards cells’ walls, which will promote the rupture of the plant matrix and an easier release of the targeted compounds [17].
The first objective of the research was to highlight the effectiveness of ultrasounds. Thus, experiments under the same conditions as UAE were carried out but without subjecting the system to ultrasounds (Control). Since the extraction time can impact the content of the extracted compounds, its influence on the extraction efficiency was also studied, for both UAE and Control. Until an equilibrium is formed between the constituents inside the plant cells and those previously dissolved by the solvent, the concentration of the extracted components increases over time [37]. Nevertheless, the bioactive compounds may oxidize or undergo conformational changes with extended extraction times [38]. Figure 2 depicts the results of the influence of extraction time and procedure on TPC. It can be noticed that although the TPC values are similar at short times (5 min) for both methods (UAE and Control), these increase by 32% in the UAE experiment lasting 30 min. By UAE, the highest increase in the TPC value occurs between 5 and 10 min, after which it increases slowly up to 30 min. However, ANOVA analysis indicated that TPC increases significantly with increases in the extraction time from 20 to 30 min. Although the best results were achieved for 30 min of extraction, approximately 95% of the polyphenols’ content was achieved in only 10 min. This fact is very useful if a continuous operation of the installation is desired because a short residence time will allow for a higher flow rate of the obtained extract.
In the solid–liquid extraction process, temperature must also be considered. Solubilization of high-molecular-weight compounds during soda pulping is facilitated by higher temperatures. High temperatures, however, could promote recondensation of small molecules, resulting in the formation of high-molecular-weight materials. The solubility of tannin particles dispersed in the continuous phase may be reduced by the self-condensation process at high extraction temperatures, resulting in precipitation [39]. Furthermore, some specific or sub-groups of polyphenols have been acknowledged for their thermolability. If the extraction is carried out at elevated temperatures and for extended times, oxidation and condensation processes may accelerate, potentially causing a decrease in the polyphenolic content [40]. Nevertheless, the structure of the raw material must be considered. Woody materials, which present a lignocellulosic structure, require severe extraction conditions, such as high temperatures, to allow for the penetration of solvent into the plant matrix and solubilize the targeted compounds [16]. Thus, an optimization of the extraction process parameters is required to ensure sufficiently high energetic conditions to release polyphenols from the rigid structure of lignocellulose at a reduced temperature and lower the extraction time to not degrade polyphenols.
Considering the aspects presented above, a conventional extraction at a temperature of 70 °C was performed. It can be noticed, in Figure 2, that although a higher TPC value is achieved for 70 °C (CONV) compared with the Control performed at 50 °C, UAE leads to better results, as confirmed by ANOVA analysis (p < 0.05). The choice of 50 °C for UAE is based on some limitations of the cavitation phenomenon. It has been established that ultrasounds exhibit greater effectiveness at lower temperatures [39]. Rising the temperature can lead to the formation of a higher quantity of bubbles, with their collapse being less violent. Consequently, the enhancement of mass transfer through cavitation is diminished.
The extraction yield, expressed as the amount of dry extract for all methods (UAE, Control, and CONV) is shown in Table 2. It is notable that the quantity of the dry extract follows the same behavior as the TPC, achieving the highest value for the UAE method (p < 0.05) and showing a slight increase in the extraction yield after 10 min. Demidova et al. [41] determined the extractives in oak bark by microwave-assisted extraction using a mixture of water and ethanol at different extraction times (5 to 100 min). Similar with the present work, this study showed that short extraction times (15 min) are required to achieve a high amount of extractives. Demidova et al. reported a maximum yield of 37 mg/g DM, which is less than what was obtained in this study using UAE. Moreover, the addition of sodium hydroxide in the water–ethanol mixture led to lower yields (23 mg/g DM). These differences can be due to both the extraction method and the solvent used. Furthermore, oak age could also influence the bioactive compounds’ content.
The TPC/dry extract ratio falls somewhat with increasing extraction time, indicating that polyphenols are better extracted at the start of the extraction, when the mass driving force is higher; the latter diminishes since the polyphenols’ concentration in oak bark decreases, while it increases in the liquid phase. It can also be observed that the values obtained using UAE are slightly higher than those obtained in the absence of ultrasound (Control).
The extraction of polyphenols from oak bark was performed using a sodium hydroxide solution, as well. This solvent facilitates the hydrolysis and removal of lignin, resulting in an easier release of polyphenols from the plant matrix [25]. Additionally, it increases the solubility of polyphenols due to alkalinity [26]. However, the concentration of sodium hydroxide in water can affect the extraction efficiency. The results for different concentrations of sodium hydroxide in water are shown in Figure 3, indicating that the highest TPC value is achieved for a concentration of 0.5% sodium hydroxide. The TPC increase is 2.1 times higher compared with the extraction performed using water. Dedrie et al. [42] achieved similar results when extracting polyphenols from oak bark by Soxhlet extraction using a mixture of water and ethanol: TPC ranged from 49 to 63 mg/g DM based on the age of the oak.
The extraction yield, expressed as the amount of dry extract, for all sodium hydroxide concentrations used is reported in Table 3. The amount of dry extract increases as the concentration of sodium hydroxide increases, whereas the TPC/dry extract ratio drops, most likely due to increased lignin extraction.
The extracts obtained in this study are intended for hide tanning. Their tanning ability is the subject of a separate study. The extraction performed using sodium hydroxide solution leads to high pH extracts, which could complicate the tanning process. Thus, extractions in water were also carried out. Aqueous solutions of sodium sulfite (0.25%) and bisulfite (0.25%) were also used to verify if they enhance the extraction process. The water and sulfite and bisulfite extracts were concentrated by mixing the extract obtained after the first extraction with a fresh portion of oak bark and subjecting it to a second extraction under the same conditions as the first one. The results are reported in Figure 4 and Table 4. As shown in Figure 4, the addition of sodium sulfite and bisulfite resulted in higher TPC values compared to water (first extract), as confirmed by ANOVA analysis (p < 0.05). The second extract, in the presence of sodium sulfite and bisulfite, showed a significant increase (p < 0.05) compared with sodium hydroxide solution used as a solvent. On the other hand, as depicted in Table 4, the quantity of dry extract is higher (p < 0.05) for the extraction carried out using 0.5% sodium hydroxide.
Ștefănescu et al. performed UAE of polyphenols from oak bark using water as the solvent, achieving a TPC of 267 mg GAE/g DM [2]. However, the significant difference between the present study and that conducted by Ștefănescu et al. is the extraction yield. They achieved only 2.72%, while in the present study, the yield ranges between 7.34% and 19.65%. The higher amount of dry extract could result in better tanning properties. These differences can be attributed to the ultrasound equipment used for the extraction, as Ștefănescu et al. used an ultrasonic bath. The added value of the present work compared to that of Ștefănescu et al. is the use of a DFR in a loop system, which can be easily scaled-up, providing a green and facile technology to process plant materials on an industrial scale.
3.2. FTIR-ATR Spectroscopy
The FTIR-ATR spectra (as used in PCA analysis) of UAE extracts obtained using water, aqueous solution of sodium sulfite and bisulfite, and increasing sodium hydroxide concentrations in water are illustrated in Figure 5. These spectra were compared to the spectrum of a commercial oak tannin, also shown in Figure 5. The most significant changes occur between 1800 and 800 cm−1, where the primary absorption bands for oak tannins are highlighted in black, the lignin absorption bands in brown, and the sulfite absorption bands in green. With increasing sodium hydroxide concentration, tannin-specific bands (1720, 1510, 1450, 1315, 1180 and 1045 cm−1) decrease in intensity, while lignin-specific bands (1600 cm−1, 1370–1380 cm−1) increase in strength, which is consistent with the TPC and dry extract values. Table 5 shows the ATR-FTIR absorption bands found in the literature.
Figure 6 illustrates the ATR-FTIR spectra of extracts obtained by UAE using increasing sodium hydroxide concentrations. At the lowest sodium hydroxide concentration (0.1%), the absorption spectrum is extremely similar to that of water extract: all absorption bands characteristic of oak tannins are present. When the concentration of sodium hydroxide increases to 0.2% and 0.5% several bands either disappear (1180 cm−1) or are attenuated (1720, 1450, 1045 cm−1), while the characteristic bands of lignin (1600, 1370–1380 cm−1) become stronger. Research data indicate the existence of an optimal concentration of sodium hydroxide for the extraction of tannins. Thus, for the extraction of hydrolysable tannins from bark of Coriaria nepalensis, the optimal concentration of sodium hydroxide was reported to be 0.22% [27], and for the extraction of condensable tannins from Pomaceous harrows, the optimal concentration of sodium hydroxide was 10% [28].
PCA analysis of ATR-FTIR spectra of the UAE and Control extracts indicates the presence of four groups, corresponding to the four principal components which explain 97.65% of the variance (Figure 7):
Group 1 contains the commercial tannin sample and the two water extracts in which sulfite and bisulfite were added (extraction time of 30 min.);
Group 2 (quite close to the commercial sample) consists of water extracts and those obtained using 0.1% and 0.2% sodium hydroxide (extraction time of 30 min.);
Group 4 includes the extracts obtained using 0.2% sodium hydroxide with or without ultrasounds, over a period of time from 5 to 30 min;
Group 3 contains only the UAE extract obtained using 0.5% sodium hydroxide (extraction time of 30 min.).
Figure 8, Figure 9 and Figure 10 illustrate the ATR-FTIR spectra of the four groups identified by PCA analysis. Even though the PCA analysis considers the general appearance of the spectra, the groups obtained share common characteristics based on the specific absorption bands presented in Table 5. The analysis of these groups allows for the following observations:
The most coherent group is Group 4 (Figure 10), which gathers the spectra of extracts obtained using 0.2% sodium hydroxide, independent of the extraction method. All these spectra exhibit absorption bands specific to oak tannins plus lignin absorption bands. Although the changes in composition between the Group 4 extracts are minor, the amount of extractables collected varies substantially (Table 2).
Figure 9, where the spectra of water and sodium hydroxide extracts are compared, illustrates how the molecular profile of the extracts changes with the increase in sodium hydroxide concentration. When using water or dilute sodium hydroxide aqueous solutions (0.1% and 0.2%) as a solvent, the FTIR spectra are quite similar, indicating the characteristic bands of tannins. However, when the sodium hydroxide concentration was increased to 0.5%, a significant increase in the lignin specific bands was observed, which results in a different grouping of this sample.
The spectra of extracts obtained by adding sulfite and bisulfite to water are rather like the commercial extract spectrum, except for the sulphonic group signals (Figure 8).
3.3. Microbiological Tests
Leather garments and footwear, which are notoriously susceptible to microbial attack, can act as vectors for microorganisms such as pathogenic bacteria and molds when in contact with the human body. This often results in unpleasant odors, skin infections, product spoilage and allergic reactions, necessitating the development of antimicrobial clothing and leather goods [46]. Plant extracts, which are valuable sources of bioactive compounds, mainly polyphenols, play an important role as a new strategy to inhibit the growth and activity of many microorganisms, including clinically important bacteria and fungi. There is an extensive body of supporting evidence for the potent antibacterial and antifungal activities of polyphenols from various oak species, including those extracted from Quercus robur mature oak bark [2,13]. However, the different solvents used for extraction and the different methods used to evaluate the antimicrobial activity make it difficult to compare the results. In addition, it was recently shown that phenolic and antioxidant compound accumulation of Quercus robur bark diverges based on tree genotype, phenology and extraction method [12]. In this complex context, to confirm the antibacterial activity of UAE extracts, the simple agar well diffusion method on MHA assays was performed in the presence of Escherichia coli (E. coli) and Staphylococcus aureus (S. aureus) bacteria, as reported in the Materials and Methods section. E. coli is a group of Gram-negative bacteria, with large diversity, from virulent to highly pathogenic, with the formation of virulent hybrid microorganisms. This ability also contributes to the development of antibiotic resistance [47]. Its capacity to thrive in both the presence and absence of oxygen [48] can affect a wide range of eukaryotic cellular processes, including cell signaling, ion secretion, protein synthesis, mitosis, cytoskeletal function, and mitochondrial function [49]. S. aureus is a potentially harmful Gram-positive bacterium that grows rapidly and abundantly in aerobic environments, causing diseases, including skin infections. Before the discovery of penicillin to treat S. aureus infections in the early 1940s, the fatality rate for such infections was approximately 80% [50].
The results of agar well diffusion assays are reported in Table 6 and Table 7. Regarding the measured diameter of the inhibition zone in mm, only results over 7 mm were considered as viable [13]. All assayed extracts show antibacterial activity against E coli and S. aureus and this activity varied depending on the bacteria species and solvent type used in UAE. Depending on the species of bacteria, E. coli exhibited higher sensitivity to sodium hydroxide extracts than S. aureus, whereas small differences were observed for aqueous and sulfitated extracts’ antibacterial inhibitory potential. The solvent has a clear effectiveness on the antibacterial activity, with the sulfitated extracts exhibiting the highest antibacterial activity against both E. coli and S. aureus, while the aqueous extract has the lowest activity. The highest antibacterial activity of the sulfitated extracts could be partially attributed to synergy, as it is known that sodium bisulfite has antibacterial activity against both E. coli and S. aureus [51]. Taking into account the molecular profile of the extracts from their FTIR-ATR spectra, as well as the TPC of the extracts, we can infer a positive correlation between the tannin concentration and the diameter of inhibition zone for the two groups of extracts, namely, tannin-rich extracts (sulfitated, aqueous and sodium hydroxide 0.1%) and lignin-rich extracts (sodium hydroxide 0.2% and 0.5%). This behavior is more evident against E. coli, while the inhibitory efficacy against S. aureus seems not to depend on phenolic concentration.
The results of the mesophilic bacteria and fungi growth test under aerobic conditions (European Pharmacopoeia Ed. 10/20) are listed in Table 8. All extracts comply with the acceptance criteria for pharmaceuticals according to the recommended specifications given by both the U.S. and European Pharmacopoeia. For instance, the TAMC should be under 103 CFU/g and the TYMC should not exceed 102 CFU/g (European Pharmacopeia Acceptance Criteria for Microbiological Quality of Non-Sterile Dosage Forms). In addition, E. coli must be absent. As shown in Table 8, no significant colonies were found after incubation: all values obtained are well below the set limits. This test also confirms the total absence of E. coli and S. aureus bacteria.
4. Conclusions
In this study, ultrasounds were employed for the extraction of polyphenols from oak bark residues. Due to the extracts being rich in tannins, a class of polyphenols, this approach could be of considerable interest in the tanning industry. In order to treat larger volumes of the extraction mixture, modified DFR equipment allowing for a loop-reactor-type system was used. By increasing the loading tank capacity, this procedure allows for easy scalability of the plant without losing the efficiency of the ultrasound treatment. Thus, this strategy provides a green and facile technique for processing vegetable materials at an industrial scale.
UAE proved to be more efficient than the conventional extraction (Control) at 30 min. The TPC value increased by 14.4% and the dry extract by 6.3% in the case of using a 0.2% sodium hydroxide solution as solvent. This indicates that UAE results in a much lower energy consumption, making this approach a greener and more efficient technique compared with conventional extraction procedures. Several possible solvents were tested, including water, sodium hydroxide solutions and a mixture of sulfite and bisulfite solution. For a single extraction, the 0.5% sodium hydroxide solution gave the best results, both for TPC and dry extract.
The antibacterial activity tests indicated that the extracts effectively inhibited the growth of E. coli and S. aureus bacteria. Additionally, the extracts showed no contamination with E. coli and S. aureus, and the mold and yeast count were well below the limits allowed for non-sterile pharmaceuticals. E. coli exhibited higher sensitivity to NaOH extracts than S. aureus, but there were no significant differences between the antibacterial inhibitory potential of aqueous and sulfitated extracts against these bacteria. The most effective antibacterial activity of the sulfitated extracts could be partially related to synergy, as it is known that sodium bisulfite exhibits antibacterial activity against both E. coli and S. aureus. For the extracts grouped according to PCA, a positive correlation between the tannin concentration and the diameter of inhibition zone against E.coli was observed.
The PCA analysis of the spectra of the extracts made it possible to classify them into four groups. In the same group as the spectrum of the commercial extract were the spectra obtained when using a mixture of sulfite and bisulfite solution as solvent, and in the second group were the spectra obtained when using water or diluted sodium hydroxide solutions. When using the 0.2% sodium hydroxide solution, the samples obtained at different times, with or without the use of ultrasounds in the DFR equipment, are part of the same group. Only the spectrum obtained when using the 0.5% sodium hydroxide solution, which shows absorption bands specific to lignin, is not grouped.
Thus, to obtain high amounts of polyphenols and dry extract, UAE using the 0.5% sodium hydroxide solution as the solvent seems to be the best choice. Although the FTIR analysis showed a significant increase in the lignin-specific bands, those specific to tannins are also present, implying that the 0.5% sodium hydroxide solution could be the best choice for the tanning process. However, further studies are required to test the ability of the extracts as tanning agents.
Conceptualization, I.C. and E.B.; methodology, I.Q., P.C., R.-R.C. and I.P.; software, I.Q., V.L. and I.P.; validation, I.C., V.L. and E.B.; formal analysis, I.Q., V.F., P.C., E.-G.D., R.-R.C. and I.P.; investigation, I.Q., V.F., P.C., E.-G.D., R.-R.C. and I.P.; resources, I.C. and E.B.; data curation, I.Q., V.L. and I.P.; writing—original draft preparation, I.Q. and I.P.; writing—review and editing, I.C., V.L., E.B. and I.P.; visualization, I.Q., N.D.I. and I.P.; supervision, I.C. and E.B.; project administration, I.C., E.B. and N.D.I.; funding acquisition, N.D.I. All authors have read and agreed to the published version of the manuscript.
The original contributions presented in the study are included in the article; further inquiries can be directed to the corresponding author.
This work was supported by a grant of the Ministry of Research, Innovation and Digitization, CNCS/CCCDI—UEFISCDI, project number PED 639/2022, within PNCDI III. Author Ferrara V. thanks the Programma Operativo Nazionale, Ricerca e Innovazione 2014−2020 (CCI 2014IT16M2OP005), Fondo Sociale Europeo, Azione I.1 “Dottorati Innovativi con caratterizzazione Industriale - CUP D43D20002240006, and the University of Salerno and the Ministry of University and Research in Italy for their financial support.
The authors Quaratesi, I.; Calinescu, I.; Lavric, V.; Badea, E.; Chipurici, P.; Dumbravă, E.-G.; Constantinescu, R.-R.; Ignat, N.D.; Popa, I. declare no conflicts of interest. The author Ferrara V. was pursuing a joint PhD between the University of Salerno (Italy) and the company Bi-QEM Specialities S.p.A. The PhD scholarship was fully funded by a grant from the Ministry of University and Research in Italy. Ferrara V. declares that the research was conducted in the absence of any commercial or financial relationships with Bi-QEM S.p.A. that could be construed as a potential conflict of interest.
Footnotes
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Enlarge this image.Figure 1. Scheme of the equipment for the ultrasound-assisted extraction (UAE) of polyphenols from oak bark. (a) Dual-Frequency Reactor (DFR); (b) loop reactor (1—batch reactor fixed with sanitary flange by the two ultrasound transducers; 2 and 3—ultrasound transducers of different frequencies, cooled with water; 4—diaphragm plate; 5—dual-Frequency processing cavity; 6—pump; 7—loading tank).
Enlarge this image.Figure 2. Influence of extraction time and type of method on the total phenolic content (TPC) from oak bark extracts (solvent—0.2% NaOH). The different letters (a–e) within graph show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test.
Enlarge this image.Figure 3. Influence of sodium hydroxide concentration on the TPC from oak bark extracts (extraction time—30 min). The different letters (a–d) within graph show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test.
Enlarge this image.Figure 4. Influence of solvent type and number of extractions on the TPC from oak bark extracts (extraction time—30 min; water-1 and water-2—first and second extract, respectively, performed with water as solvent; sulfite-1 and sulfite-2—first and second extract, respectively, carried out with an aqueous solution of sodium sulfite and bisulfite). The different letters (a–e) within graph show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test.
Enlarge this image.Figure 5. ATR-FTIR spectra of extracts obtained by UAE using different solvents compared to ATR-FTIR spectrum of a commercial sample in the 1800–800 cm−1 range.
Enlarge this image.Figure 6. ATR-FTIR spectra of extracts obtained by UAE using increasing sodium hydroxide concentrations in the 1800–800 cm−1 range.
Enlarge this image.Figure 7. Clusters found after principal component analysis (PCA) of the spectra of the 15 tannin extracts (the percent variance explained by the principal four components is 97.65).
Enlarge this image.Figure 10. ATR-FTIR spectra of the extracts grouping in Group 4 (solvent—0.2% sodium hydroxide).
Steps for establishing the best extraction conditions for polyphenols (tannins).
| First step |
|
| Second step |
|
| Third step |
|
Influence of extraction time and type of method on the extraction yield (solvent—0.2% NaOH). The different letters (a–h) within table show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test. Control—control experiment, CONV—conventional extraction method.
| Time, min | 5 | 10 | 20 | 30 | |
|---|---|---|---|---|---|
| UAE | Dry extract, mg/g DM | 99.0 ± 0.87 g | 129.0 ± 1.57 c | 137.7 ± 2.38 b | 146.7 ± 1.56 a |
| TPC/Dry extract | 0.47 | 0.48 | 0.46 | 0.45 | |
| Control | Dry extract, mg/g DM | 94.6 ± 0.86 h | 113.1 ± 1.45 f | 120.3 ± 2.73 e | 122.0 ± 3.18 d,e |
| TPC/Dry extract | 0.50 | 0.43 | 0.41 | 0.42 | |
| CONV | Dry extract, mg/g DM | - | - | - | 138.0 ± 2.43 b |
| TPC/Dry extract | - | - | - | 0.41 | |
Influence of sodium hydroxide concentration on the extraction yield (extraction time—30 min). The different letters (a–d) within table show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test.
| Solvent | Water | Sodium Hydroxide Concentration (%) | ||
|---|---|---|---|---|
| 0.1 | 0.2 | 0.5 | ||
| Dry extract, mg/g DM | 73.4 ± 1.79 d | 111.8 ± 0.76 c | 146.7 ± 1.56 b | 196.5 ± 3.24 a |
| TPC/Dry extract | 0.47 | 0.57 | 0.45 | 0.37 |
Influence of solvent type and number of extractions on the extraction yield (extraction time—30 min). The different letters (a–e) within table show the significant difference between groups analyzed by ANOVA (p < 0.05) and Duncan’s new multiple comparison test.
| Solvent | Water | 0.5% Sodium Hydroxide | Sodium Sulfite and Bisulfite | ||
|---|---|---|---|---|---|
| Number of extractions | 1 | 2 | 1 | 1 | 2 |
| Dry extract, mg/g DM | 73.4 ± 1.79 e | 100.5 ± 1.08 d | 196.5 ± 3.24 a | 142.7 ± 1.27 c | 159.5 ± 3.49 b |
| TPC/Dry extract | 0.47 | 0.58 | 0.37 | 0.44 | 0.46 |
ATR-FTIR absorption bands.
| Bands (cm−1) | Assignment | Identification |
|---|---|---|
| Signals specific for tannins obtained from oak bark [ | ||
| 1720 L | The C=O stretching of esters of hydrolysable tannins, especially derivatives of gallic acid. | Hydrolysable tannins |
| 1605 L | The stretching of the C=C–C aromatic bond. | |
| 1510 S | C=C asymmetric. Aromatic character. | Condensed tannins |
| 1450 L | C–H bending. Aromatic. Asymmetric. C–C stretching. Ring B, C–O stretching. Aromatic. | |
| 1315 L | C–O stretching. Ring gallic/pirogallic. | Hydrolysable tannins |
| 1180 L | Phenyl acetates, C–O bending C–H bending. Aromatic. Symmetric. | Hydrolysable tannins |
| 1045 L | C–O stretching. Aromatic. (Esters) C–H bending. Aromatic. Symmetric. In-plane. | |
| Specific for sulfite extraction | ||
| 1118 | Asymmetric stretching of SO4 group. | NaHSO3 + Na2SO3 [ |
| 971 | Symmetric stretching of SO4 group. | NaHSO3 + Na2SO3 [ |
| Specific for lignin | ||
| 1370–1380 | Phenolic stretch vibration of OH and aliphatic CH deformation in methyl groups. This band is common to lignin. | Lignin [ |
| 1600, 1515, and 1426 | Aromatic skeleton vibrations. | Lignin [ |
| 1462 | The C–H deformation combined with aromatic ring vibration. | Lignin [ |
Antibacterial activity against Escherichia coli (ATCC 11229).
| Extract | Inhibition Zone | Zone of Clearance (mm) |
|---|---|---|
| Sulfite-1 | [Image omitted. Please see PDF.] | H = (52–10)/2 |
| Sulfite-2 | [Image omitted. Please see PDF.] | H = (54–10)/2 |
| 0.1% NaOH | [Image omitted. Please see PDF.] | H = (45–10)/2 |
| 0.2% NaOH | [Image omitted. Please see PDF.] | H = (39–10)/2 |
| 0.5% NaOH | [Image omitted. Please see PDF.] | H = (50–10)/2 |
| Water-1 | [Image omitted. Please see PDF.] | H = (35–10)/2 |
| Water-2 | [Image omitted. Please see PDF.] | H = (38–10)/2 |
Antibacterial activity against Staphylococcus aureus (ATCC 6538).
| Extract | Inhibition Zone | Zone of Clearance (mm) |
|---|---|---|
| Sulfite-1 | [Image omitted. Please see PDF.] | H = (48–10)/2 |
| Sulfite-2 | [Image omitted. Please see PDF.] | H = (50–15)/2 |
| 0.1% NaOH | [Image omitted. Please see PDF.] | H = (35–10)/2 |
| 0.2% NaOH | [Image omitted. Please see PDF.] | H = (31–10)/2 |
| 0.5% NaOH | [Image omitted. Please see PDF.] | H = (33–10)/2 |
| Water-1 | [Image omitted. Please see PDF.] | H = (34–10)/2 |
| Water-2 | [Image omitted. Please see PDF.] | H = (31–10)/2 |
Total aerobic microbial count (TAMC) and total mold and yeast count (TYMC) amounts for oak bark extracts obtained by UAE.
| Extract | TAMC (CFU/g) | TYMC (CFU/g) | E. coli | S. aureus |
|---|---|---|---|---|
| Allowable Limits | ≤103 | ≤102 | Absent | Absent |
| Sulfite-1 | 1.5 × 101 | 3.33 | Absent | Absent |
| Sulfite-2 | 1.8 × 101 | 1.2 × 101 | Absent | Absent |
| 0.1% NaOH | 2.1 × 101 | 1 × 101 | Absent | Absent |
| 0.2% NaOH | 2.4 × 101 | 8.66 | Absent | Absent |
| 0.5% NaOH | 1.9 × 101 | 6.66 | Absent | Absent |
| Water-1 | 3.2 × 101 | 1.6 × 101 | Absent | Absent |
| Water-2 | 2.8 × 101 | 1.2 × 101 | Absent | Absent |
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; Calinescu, Ioan 2
; Lavric, Vasile 2
; Ferrara, Vittoria 3 ; Badea, Elena 4
; Petre Chipurici 2
; Elisa-Gabriela Dumbravă 1 ; Constantinescu, Rodica-Roxana 1 ; Ignat, Nicoleta Daniela 5 ; Popa, Ioana 2
1 Leather and Footwear Research Institute (ICPI) Division, National Research and Development Institute for Textile and Leather (INCDTP), 031215 Bucharest, Romania;
2 Faculty of Chemical Engineering and Biotechnologies, National University of Science and Technology Politehnica Bucharest, 011061 Bucharest, Romania;
3 Department of Chemistry and Biology “A. Zambelli”, University of Salerno, I-84084 Fisciano, Italy;
4 Leather and Footwear Research Institute (ICPI) Division, National Research and Development Institute for Textile and Leather (INCDTP), 031215 Bucharest, Romania;
5 Faculty of Entrepreneurship, Business Engineering and Management, National University of Science and Technology Politehnica Bucharest, 060042 Bucharest, Romania;