1. Introduction
The agri-food sector generates huge quantities of solid and liquid waste, containing a high organic load. Landfilling or disposal of such waste in an uncontrolled way can cause environmental pollution and economic loss. On the other hand, these residues containing valuable metabolites have an enormous potential for the production of sustainable additives, chemicals, biofuels, and energy [1]. In particular, high added value compounds that are still present in by-products of the food processing industry can be employed as food additives and for the formulation of new functional foods, nutra-/pharmaceuticals, cosmeceuticals, and beauty products, as well as for innovative and more sustainable packaging [2]. Efforts for the systematic reuse of food waste to produce commercial products and energy provided the integrated approach proposed by the biorefinery concept. Figure 1 reports a generic example of the biorefinery solution for food waste valorization.
Agri-food industry by-products comprise several complex biomasses. The waste originated by various branches of the food industry can be divided into two main groups (plant or animal origin) and seven subcategories: cereals, root and tubers, oil crops and pulses, fruits and vegetables, meat products, fish and seafood, and dairy products [3,4]. In agreement with the hierarchical approach proposed by the biorefinery concept [5], these by-products contain valuable compounds that can be exploited in the development and production of new functional ingredients. In particular, interesting bioactive compounds present in food industry residues include proteins, polysaccharides, lipids, polyphenols, carotenoids, tocopherols, and essential oils.
Several extraction techniques have been proposed in the literature for their efficient recovery, from the conventional solid liquid extraction to non-conventional techniques, with the aim of improving the extraction yield of targeted compounds, avoiding deterioration and loss of functionality during processing, and allowing the food grade nature of the final product [6]. These non-conventional techniques include microwave-, ultrasound-, and supercritical fluid-assisted extraction, pressurized liquid extraction, pulsed electric field, and enzyme-assisted extraction techniques. In particular, the use of extraction performed under high-pressure conditions such as supercritical fluids and pressurized liquid extraction offers several advantages for the extraction of bioactive molecules, including short processing times, reduced use of organic solvent, and mild operative conditions able to avoid thermal degradation [7]. These technologies are considered environmentally friendly and are generally able to produce higher yields and high-quality final extracts.
Therefore, this review provides a comprehensive introduction to the literature update on pressure-assisted extraction processes. A critical analysis of the different processes proposed in the literature is reported, with the objective of defining the most suitable process for the extraction of different categories of high added value compounds that can be find in natural waste. Moreover, the possibility of combining different extraction processes in order to complete the valorization of agri-food waste is also analyzed for some case studies.
2. Pressurized Liquid Extraction Processes
In recent years, as a consequence of the deepening of research on biologically active natural substances, many extraction methods have been developed in order to obtain active compounds directly from natural sources. Phenolic compounds, carotenoids, flavonoids, lipids, proteins, polysaccharides, and essential oils are examples of active compounds obtainable from natural sources.
Among the emerging extraction technologies that can enhance the performance of conventional extraction techniques, pressured liquid extraction (PLE) is considered to be environmentally friendly and efficient, and generates a small amount of waste while reducing cost and time [8,9]. The methodology is based on using solvents below their critical point in order to preserve the liquid phase during extraction. Firstly, PLE was introduced as Accelerated Solvent Extraction (ASE) technology by Dionex Corporation [8] in 1995 at the Pittcon Conference. It is also known as pressurized solvent extraction (PSE), enhanced solvent extraction (ESE) or pressurized fluid extraction (PFE) [10,11,12,13].
In the beginning, this technology was used for trace analysis of organic contaminants [8,14,15]. Currently, this technique is mainly applied to the extraction of environmental pollutants in soil, sediments, sewage sludge, and fly ash. However, in recent years, due to the advantages of PLE that include significantly shortened extraction time and low solvent volume required, easy operation, and good performance, this technology is being applied to different fields, including biology, pharmaceuticals, and the food industry [15,16,17,18]. As a result, the number of papers related to pressure-assisted processes in this field have increased significantly (Figure 2).
This growing interest is mainly due to some impressive features of PLE; indeed, it is a process that can be automated, is characterized by reduced extraction time and solvent consumption, and its settings are particularly suitable for compounds that are sensitive to oxygen and light. PLE requires minimal sample pre-treatment, especially for non-fat samples such as homogenization and/or drying [7].
Regarding the mechanism of extraction, PLE combines elevated pressure and temperature with liquid solvents, without exceeding the critical points of the solvent or solvent mixtures, which remains in a liquid state. In this way, it is possible to increase the mass transfer rate from the matrix compounds to the extraction solvent to achieve a fast and efficient solid–liquid extraction. PLE often demonstrates better recovery when compared to other extraction techniques such as microwave-assisted extraction (MAE) or ultrasound-assisted extraction (UAE) [19,20]. Other significant advantages of PLE include the elimination of post-extraction steps, such as filtration and centrifugation, the high level of automation, and the possibility of coupling with different techniques, such as ultrasound [8,21,22].
Regarding the operation mode, there are three possible different modalities for applying the PLE process: static [23,24,25], dynamic [26], and static–dynamic. In the static mode, a fixed volume of extracting solvent is used; in the dynamic mode, the extracting solvent flows continuously through the sample; in the static–dynamic layout, a combination of the two modes is proposed. Figure 3a shows that static pressurized liquid extraction generally consists of the following components: (1) a pump to feed the solvent through the system; (2) an extraction vessel. The extractor consists of an extraction cell (EC) in which the solid matrix is contained, surrounded by a heating system. The layout is completed by (3) a system cooling the fluid from oven temperature to room temperature or even lower; (4) an on/off pressure needle valve located after the cooler to maintain pressure during static extraction; (5) a N2 flushing system, located between the high-pressure pump and the extraction system. The dynamic pressurized liquid extraction system is improved on the basis of the static pressurized extraction system, as shown in Figure 3b: (1) the selector valve connected to the N2 flow is replaced by the inlet valve, because in dynamic mode, there is no need to purge the system; (2) the extraction vessel; (3) a restrictor connected to the outlet of the cooler (in place of the on/off valve) is used to maintain the constant pressure in the system [27].
Other configurations of the pressure-assisted process are also proposed in the literature.
Extractor Naviglio is based on the application of pressure (about 8–9 bar) and decompressions up to atmospheric values, creating a suction effect and a mechanical transport of target compounds from the solid to the solvent phase. The system is composed of two cylinders in which extraction is performed, with the cylinders communicating through a tube on which an electro-valve is intercepted. A pneumatic system working with compressed air moves two pistons to provide a pressure rise during a static stage, during which pressure is held constant for a while; this improves the penetration of the solvent within the solid and a dynamic stage, when the pistons are removed from the equilibrium position and alternately work generating negative pressure gradient between the inside and the outside of the solid matrix and promoting the dragging effect of the extraction [30].This system is designed to work at room or low temperatures, reducing the thermal stress for thermo-sensitive compounds [31].
Moreover, high-pressure and temperature-assisted extraction (HPTE), similar to PLE, exploits the effects of pressure to work at a high temperature for the mass transfer improvements, but a different setup of the extraction cell is employed. Indeed, extraction is carried out in a stainless-steel vessel and a stirrer equipped by impellers provides an excellent and homogeneous mixing action [32]. The extractor can work up to 350 °C and 200 bar, with the two variables monitored by a thermocouple and a pressure gauge, respectively. The thermal sensor is connected to a temperature that maintains the variable at the desired set point value by an external electrical jacket. Before the extraction, an inert atmosphere is created inside the extraction vessel by bubbling inert gases for air removal with the aim of limiting degradation reactions of thermo-sensitive compounds. In HPTE, temperature and pressure are strictly related since the second factor corresponds to the vapor pressure of the solvent at the set point temperature [33].
Manosonication-assisted extraction is an example of pressure-assisted process coupled with other extraction aid, particularly the cavitation effects induced by ultrasounds. The system involves a double-jacket reaction chamber, a sensor, and PID controller for temperature control, while an ultrasonic generator and nitrogen gas controlled by a manometer was supplied to adjust the amount of external pressure applied to the reaction chamber through an inlet fitting [34].
Some authors use a combination of static and dynamic [11,35,36] layout to optimize the extraction process. Both modes have their own advantages and disadvantages. The most significant negative feature of dynamic pressurized extraction is that the solute in the extract is diluted, which requires a concentration step as post-processing. Whereas for the static extraction mode, the extraction efficiency depends on the solute mass transfer balance between the matrix and the extracting solvent [37]. The critical factors are the temperature and time of the extraction. Furthermore, in the static extraction mode, complete extraction might not be achieved due to the limited volume of the extraction fluid.
In general, homemade experimental apparatus is used for the extraction. However, ready-to-use equipment for PLE is available on the market (for example, Parr Instruments Company Illinois (USA), Fluid Management System Inc. Kentucky (USA)).
2.1. Effect of Operative Parameters
In order to improve the extraction efficiency of an active substance, the key parameters that can affect the efficiency of the extraction process must be investigated. For pressurized liquid extraction, the factors that affect the extraction include the type of residue, solvent type, extraction time, pressure, temperature, pH, liquid-to-solid ratio, stirring speed, particle size, and sample moisture [38,39].
Pressure-assisted extractions can employ a wide range of extraction temperatures, usually from room temperature to 200 °C, and pressures up to 200 bar [11]. Temperature is a very important parameter to improve the mass transfer rate of the target compound from the solid matrix to the extraction solvent. Due to the non-compressibility of the liquid (the change in density with pressure is usually insignificant), the effect of pressure does not have a direct effect on the solubility of the analyte in some cases. However, operating at higher pressures can ensure that the solvent remains in a subcritical liquid state, while pressure forces the solvent to penetrate into the matrix pores and facilitates the desorption of the target compound due to a synergistic effect with temperature. The combined effect of high temperature and high pressure provides conditions that are able to significantly increase the solubility and diffusivity of the solute, to perform the rupture of the solute-matrix bond, and to reduce the surface tension and viscosity of the solvent [40,41,42,43].
Solvent selection is a key step in the design of industrial extraction processes. The extraction solvent must be able to dissolve the target solute and minimize the co-extraction of other matrix components. In addition, compatibility with subsequent processing steps (extract purification, target molecule preconcentration, or analytical techniques) and solvent volatility (if extract concentration is required) must also be considered [9,44].
The use of green solvents can have a positive impact on the environmental sustainability of processes; however, extraction yield can be negatively impacted by the selection of solvent less performant compared to the classical solvent. However, when non-conventional processes are performed, operating conditions are able to provide extraction yields and selectivity even higher than conventional processes using conventional solvents. Among the green solvents investigated in the literature, especially for agri-food waste recovery, there are subcritical water and pressurized ethanol aqueous solutions. Since they do not produce residues during the extraction process, they are non-toxic and eco-friendly.
Indeed, as reported by Aliakbarian et al. [45], subcritical water extraction (100 ≤ T ≤ 140 °C and 8 ≤ P ≤ 15 MPa, for 130 min) provided a total flavonoid and polyphenol 12- and 19-fold higher than conventional solid liquid extraction performed with water at 25 °C for 19 h, and with a higher antiradical power. In addition, polyphenol extraction from Teucrium montanum L. carried out by subcritical water gave higher concentrations of total phenolics in comparison to aqueous, methanolic, acetone, ethyl acetate, and petroleum ether extracts obtained by conventional solid/liquid extraction [46]. At the same time, the comparison of HPTE with other extraction techniques, using ethanol as solvent, showed the best performance of the former on the recovery of bioactive compounds from Arthrospira platensis, while solid–liquid extraction provided the worst performance [47].
2.2. PLE of Bioactive Compounds from Waste
Food industry by-products and waste have been proven to be a cheap raw material for the recovery of compounds with commercial value. For example, molecules of interest for the industry are caffeine from spent coffee grounds [33,48], lycopene and carotenoids from tomato waste [49], astaxanthin from crustacean and shrimp waste [50,51], polyphenols from fruit waste and rice [52], and lipids and protein from the waste of olive or microalgae [53,54,55]. These active substances have potential health benefits and find applications as ingredients or additives in the medical field, as well as in the food and cosmetic industries. By enabling the use of green and food-grade solvents, pressure-assisted extraction has been successfully applied to extract active substances (Table 1) such as proteins, polysaccharides (such as lignans), lipids, polyphenolic compounds (carotenoids and tocopherols), and essential oils from food waste [18].
2.2.1. Proteins
The growing world population requires a more sustainable source of protein for food and technological aims. Thus, new sources include insects, fungi, and algae, as well as waste streams from food processing [86]. Conventional methods for the recovery of proteins involve the use of alkali, since the basic environment allows the breakage of disulfide bonds in protein and higher solubility providing high extraction yields, increases their bioavailability and digestibility, but degrades protein quality [87]. In recent years, pressure-assisted extraction has been successfully applied to extract proteins from a variety of food processing residues. Šalplachta et al. [56] optimized pressurized hot water extraction conditions (50 °C, 15 MPa, and 5 min as extraction time) for the recovery of proteins from Sambucus nigra L. branches, demonstrating the highest yields and better reproducibility of the pressure-assisted technique compared to traditional solid–liquid extraction. Zhou et al. [58] used PLE to extract proteins and bioactive compounds from spirulina. Response surface methodology (RSM)-centered composite design (CCD) was used to evaluate and optimize the extraction time (5–15 min), temperature (20–60 °C), and pH (4–10) of the PLE process for extraction (103.4 bar). The results showed that under optimal extraction conditions (10 min, 40 °C, and pH 4), PLE significantly increased the protein yield of the extracts (46.8 ± 3.1%) compared to non-pressurized extraction.
PLE was also applied to produce extracts with antioxidant activity from Sparus aurata side streams. Protein recoveries of 22% (muscle), 33% (head), 78% (viscera), 24% (skin), and 26% (tailfin) were obtained, with values 1.2–4.5-fold higher compared to samples obtained from conventional stirred extraction [59]. Similarly, Fuente et, al [60,61] used the PLE technique to obtain protein extracts with antioxidant capacity from muscle remains, head, viscera, skin, and caudal fins of salmonids. Protein recovery was ≈92% for all samples except viscera, implying a 1.5–4.8-fold increase compared to stirred extraction (≈28%). The effect of pressure in enhancing extraction yields of proteins can be ascribed to the alteration of covalent bonds and to the breakdown of proteins into soluble peptides, as well as to the induced variation of dielectric and physical properties of solvent (such as subcritical water) through the operating conditions enabled by high-pressure-assisted processes [88].
2.2.2. Polysaccharides
Polysaccharides are polymers composed of more than 10 monosaccharides with straight or branched glycosidic bonds [89]. They are characterized by low toxicity, low cost, and high biocompatibility and biodegradability [90]. Natural polysaccharides have several potential applications in food industries due to their properties as stabilizers, emulsifying, thickeners, gelling agents, and film forming [64,90,91,92,93,94]. In addition, they exert bioactivities that can be exploited in medicine, including antioxidant, antitumor, anti-hyperglycemic, and immune regulation activities [95].
Plant polysaccharides’ extraction efficiency is often reduced by the presence of cell wall polymers, so extraction aids able to disrupt the cell structure have to be applied to enhance extraction yields. The application of advanced extraction techniques such as PLE shows the great potential of polysaccharides recovery. The better efficiency towards polysaccharides was reported by Dobrinčić et al. [38], who evaluated the effect of MAE and PLE parameters (solvent, temperature, time, and number of cycles) on the polysaccharide yield and chemical composition (total sugars) of Fucus virsoides and Cystoseira barbata. The best PLE parameter was 0.1 M H2SO4 as solvent at 140 °C for two cycles of 15 min. In addition, PLE obtained a significant increase in polysaccharide yield from Fucus virsoides and the polysaccharides obtained using PLE had the highest polydispersity index, fucose, and sulfate content, and the lowest glyoxylate content; however, they had lower antioxidant activity. Herbst et al. [40] studied the yield of extracting valuable compounds from brewer’s grains using PLE under constant pressure with varying temperature, solvent type, and flow rate, and the total phenolic compounds, total flavonoids compounds, antioxidant activity, reducing sugars, and total reducing sugars as indicators, and compared the results with the Soxhlet extract. The results show that PLE can maintain a high extraction rate of reducing sugars and total reducing sugars in the BSG matrix. The application of high pressure can modify the conformation and structure of polysaccharides, changing their biological activities [96,97]. High-pressure extraction was also coupled with ultrasounds, creating hybrid techniques such as high-pressure ultrasonic-assisted extraction [98], which was employed for the recovery of polysaccharides from Hovenia dulcis.
2.2.3. Lipids
Lipids are categorized as either neutral or complex. Neutral lipids include glycerides, wax esters, free fatty acids, free fatty alcohols, carotenoids, sterols, tocopherol and lignin-type antioxidants, and biologically derived hydrocarbons. Complex lipids include phospholipids, glycolipids, ceramides, cerebrosides, and gangliosides.
Many well-known extraction methods have been applied for the extraction of lipids from food waste, including Soxhlet, Bligh and Dyer, and Folch processes, and SFE; more recently, PLE have been successfully developed to enhance lipid extraction. Many applications of PLE methods have been reported, such as the extraction of compounds from plants [28,99,100,101], animals [102,103,104,105], soil [106,107], and algae [108], but few from yeast [109,110]. According to the large potential of lipid production with oleaginous microorganisms, the application of PLE methods to lipid extraction from yeast is a major challenge but with promising perspectives [111,112].
Yongjin et al. [113] used a PLE process to extract microalgal lipids from Isochrysis biomass. Compared with the Soxhlet and Folch methods and PLE using n-hexane, PLE using ethanol exhibits excellent lipid extraction performance. Cescut et al. [114] used PLE to extract total lipids from oleaginous yeast (Rhodotorula glutinis) and compared it with two traditional methods: the Soxhlet method and the improvement of the Bligh and Dyer method. The results showed that the extraction time of PLE was reduced by 10 times, the amount of solvent was reduced by 70%, and the lipid extraction rate was significantly higher than the reference method (Soxhlet method and Bligh and Dyer method). Villanueva-Bermejo et al. [115] used PLE to extract 3-rich oil from chia seeds by using pressurized food grade ethanol at 60 °C. In a short extraction time (10 min), the recovery rate of the seeds was close to 100% [106,116,117,118,119,120,121]. However, since lipids are soluble in non-polar solvents such as hexane, extraction using PLE cannot be performed applying green solvents. For this class of compounds, supercritical fluids extraction is more advantageous.
2.2.4. Polyphenolic Compounds
Among the secondary metabolites produced by plants, phenolic compounds are a relatively abundant category. Generally, phenolic compounds are most often classified into flavonoids and non-flavonoids. Flavonoids are the most abundant plant polyphenols consumed by humans. They are polyphenol compounds with two aromatic rings connected by a three-carbon bridge. For example, more than 800 phenolic compounds have been identified in plants. Chemically, phenolic compounds have at least one aromatic ring and one or more hydroxyl groups [122]. Although phenolic compounds are not necessary for human metabolism, if they are present in the diet, they can improve health and may reduce the risk of serious diseases. In fact, phenolic compounds are widely studied because they can reduce the reactive oxygen species that cause oxidative stress and are related to the prevention and treatment of cardiovascular and chronic degenerative diseases. In addition, phenolic compounds are due to their antibacterial, anti-inflammatory, antidiabetic, and anticancer properties [123,124,125,126,127,128]. At the same time, the antioxidant capacity of phenolic compounds indicates that they can be used as natural additives for functional foods. For example, the phenolic compounds recovered from olive mill wastewater [129] were evaluated as an agent to extend the shelf life of baked products due to their antibacterial properties. In the same way, Basanta et al. [130] extracted phenolic compounds from sugar-exhausted Prunus avium homogenate to be loaded into low-methoxyl-pectin film as an antioxidant barrier for food preservation.
However, the extraction of these compounds is challenging because they may be unstable and their biological activity can be affected by extraction process parameters and external factors such as the presence of oxygen and light. Phenolic compounds have been extracted for decades using traditional methods such as Soxhlet extraction and maceration [131,132,133].
Alternative and more effective extraction processes are needed to overcome these shortcomings and produce higher extraction yields while maintaining the integrity of the compound.
Due to the many advantages of pressure-assisted extraction processes, the research on the extraction of polyphenolic compounds by this technology has become increasingly intense in recent years.
Battistella Lasta et al. [134] recovered phenolic compounds from beetroot (Beta vulgaris L.) residues, leaves, and stems by PLE at a temperature of 40 °C, pressures of 7.5, 10 and 12.5 MPa, and a flow rate of 3 mL min−1. Okiyama et al. [135] demonstrated that PLE was effective in enhancing the extraction rate of flavanols from industrial by-product cocoa shells. Shang et al. [136] optimized extraction conditions for total phenolics and caffeine from waste coffee grounds using the PLE method with water and ethanol.
Considering the thermosensitivity of several polyphenols, the positive effect of pressure in improving the extraction process is of a great interest, allowing good extraction yields even at low temperatures [137] or enabling the use of a very high temperature but for short times [138].
3. Supercritical Fluid Extraction
Supercritical fluid extraction (SFE) is an emerging clean technique that developed in the late 1970s/early 1980s and represents the most studied supercritical carbon dioxide-based technique [139]. Most of the published works focus on the extraction of compounds of interest from plant matrices such as seeds, fruits, or leaves. These studies have mainly been the subject of previous reviews. They are outside the scope of the present review paper, which instead focuses on the extraction of active compounds from food industry waste. The SFE process exploits the advantages of the use of carbon dioxide in the supercritical state: mild critical values of temperature and pressure (Tc = 31.1 °C; Pc = 73.8 bar); total elimination of the use of organic solvents that could remain present in traces in the extract (with a consequent limitation of the environmental impact of the extraction process); the possibility of simply modulating the solvent power of CO2 which changes significantly with pressure and temperature; ease in the scale-up of the process up to the industrial scale [140,141,142,143,144]. It is important to highlight that carbon dioxide in the supercritical state is a “hybrid” between a gas and a liquid; indeed, it has some properties typical of gases (such as high diffusivity, which guarantees a better transport of matter with consequent extraction times drastically reduced than the ones of the classic solvent extraction) and others comparable to those of liquids (such as density, to which the solvent power is directly connected). Moreover, carbon dioxide is non-toxic, non-flammable, available in large quantities, and cheap. The main disadvantage of supercritical carbon dioxide (scCO2)-based techniques is linked to the investment costs (for the design of equipment that operates at high pressures) and operational costs (for pumping CO2). The fundamental elements in an extraction plant are sketched in Figure 4: (a) the carbon dioxide vessel; (b) the pump (with refrigerated head to avoid cavitation phenomena); (c) a heat exchanger to preheat the carbon dioxide; (d) an extraction vessel in which the matrix from which to extract the compounds of interest is loaded; (e) one or more separators at lower pressures than the extractor’s one.
In some cases, due to the low polarity of carbon dioxide, a polar cosolvent can be added to increase the solvent power of the mixture. With this strategy, however, one of the advantages of SFE, which is the complete elimination of organic solvents, is lost.
In general, homemade experimental apparatus is used for the extraction. However, ready-to-use equipment for PLE is available on the market (for example, Extratex and SFE process).
3.1. Effect of the Operating Parameters
The effect of pressure, temperature, average particle size of the matrix, supercritical extraction time, amount of cosolvent, and CO2 flowrate on the extraction yield (g active compounds obtained/g raw material × 100%) is generally considered [17]. Typical values of the operating conditions for the SFE process are pressures in the range 300–500 bar and temperatures in the range 40–80 °C.
3.2. SFE of Bioactive Compounds from Waste
SFE has been widely used recently to reduce the environmental impact of agro-industrial waste. Residues, waste, and by-products generated by the food-growing and -processing industries have proven to be a valuable source of biologically active compounds that can be used in the food, nutraceutical, cosmetic, and pharmaceutical industries. Among the most widespread, there are certainly the waste of the tomato [145], fishery [146], and wine [147] industries; however, an important source of active compounds are also crops and waste from less widespread productions such as cherimoya [148], peel and leaves, or defatted Assaí berries [149].
Industrial tomato by-products, for example, are rich in highly valued compounds such as carotenoids (lycopene, β-carotene), tocopherols, fatty acids, and sitosterols which can be selectively extracted simply modulating the operating conditions. Vági et al. demonstrated that if industrial tomato waste are processed with scCO2 at 460 bar and 80 °C, the extract contained the highest concentration of carotenoids with 90.1% of lycopene, while products rich in tocopherols and phytosterol were obtained above 30 MPa and 40 °C [150].
SFE in some cases has been coupled with other processes to obtain value-added products from food waste. Yu et al. [151], for example, valorized corn, taro, lettuce, and bean sprout using a sequential SFE and thermochemical conversion. The obtained extracts, depending on the vegetal source, were rich in sterols with potential anticancer properties, difatty acids for acne treatment, or unsaturated fatty acids for cholesterol control.
3.2.1. Lipids
Lipids have typically been extracted from fruit, plant, coffee, or seafood processing residues using scCO2 without the need for cosolvents; indeed, lipids are well soluble in scCO2. Fatty acids (FA), common components of complex lipids, can be extracted from red fruits’ seeds and peels or from citrus plants’ seeds. In some papers, the results obtained through supercritical extraction and extraction using organic solvents, such as hexane, have been compared [152,153].
It is well known that red fruits such as raspberry, blueberry, wild strawberry, pomegranate, blackberry, and blackcurrant are very rich in antioxidants, in particular, flavonoids, polyphenols, carotenoids, anthocyanins, and vitamins C and E, which help counteract damage from free radicals, oxidative stress, and cellular degeneration. Waste from the industries that process these fruits (seeds and peels, for example) are rich in FA. Although the total yields in the extract are in some cases higher using hexane as the extracting solvent, the affinity of supercritical CO2 for lipophilic compounds leads to purer extracts, with a higher percentage of fatty acids compared to hexane. For example, Campalani et al. demonstrated that when processing wild strawberry waste, 26.0% of fatty acids were obtained with scCO2 and 1.4% using hexane [152].
Citrus (e.g., mandarin, lemon, lime, orange, tangerine, bergamot, and grapefruit) seeds are commonly considered as agro-industrial waste, but they are a source of by-products given they contain phenolic compounds, limonoids, carotenoids, and tocopherols [154,155]. Rosa et al. [153] compared the extraction of fatty acid and triacylglycerol content by processing mandarin, lemon, and grapefruit seeds with scCO2 extraction and Soxhlet extraction using hexane. Compared to what was observed for the extraction from seeds and peels of red fruits, and also citrus seeds, the total yield was higher for Soxhlet than SFE; the fatty acid profiles obtained through the two extraction techniques were similar but, using SFE, no traces of unwanted solvent were present.
SFE has also been used to extract fatty acids from less common fruit-processing waste worldwide, such as tamarillo (native fruit from Peru, Ecuador, and Colombia), date (cultivated and consumed in African countries), rosehip, and mango. The processing residue from the preparation of tamarillo juices consists of the pericarp and seeds, which represents 24% of the fruit; rosehip seeds constitute about 30% wt of the fruit and is a waste product in the industrial production of herbal tea, fruit juice, jelly, jam, and marmalade; in the case of mango, 35–40% of waste is generated in the form of peels, kernels, and culled fruits [156,157,158].
Depending on the structure of the fruit waste, different SFE optimal operating conditions were identified. For tamarillo, the best results for the extraction of linoleic, oleic, and palmitic acids were obtained at 380 bar and 64 °C [156]; for saturated, monounsaturated, and polyunsaturated fatty acids (with oleic acid as the major constituent) extracted from date stones oil, the optimal conditions were 250 bar and 60 °C [159]; fatty acids were successfully extracted from rosehip seeds at 300 bar and 40 °C [157]; for mango kernel, the highest yield was achieved at a higher pressure (500 bar) and a temperature of 40 °C [160].
In relation to the valorization of agro- and food industrial waste, some research groups are trying to valorize coffee by-products, since coffee is one of the world’s most widely consumed beverages and its processing chain leaves behind large volumes of waste [161]. In this framework, Ahangari and Sargolzaei extracted fatty acids from spent coffee grounds through supercritical fluid, Soxhlet, microwave irradiation, and ultrasonic irradiation extraction [162]. By using SFE, they observed that higher yields were obtained at a higher pressure, lower temperature, and using hexane as a modifier (rather than water or ethanol). All the yields obtained using SFE were higher than those of the other extraction methods using organic solvents except the Soxhlet extraction. Considering the fatty acid compositions, the authors observed that linoleic, palmitic, oleic, and stearic acids were among the most present; in the case of SFE, different operating conditions led to significant differences among several fatty acids, whereas no significant differences were observed in the FA compositions of the extracted oils using organic solvents. FA, above all linoleic and palmitic acids, were extracted from coffee silverskin (the thin layer directly in contact with the coffee bean, which is the only residue from the operation of roasting) by Nasti et al. [163], using both SFE and Soxhlet extraction. From these authors’ findings, it is possible to observe that yields are similar considering the conventional extraction and SFE at the best operating conditions; however, it should be highlighted that when using SFE, the extraction time was drastically reduced and the use of hydrocarbon or chlorinated solvents was avoided.
Undoubtedly, among the food waste containing fatty acids, the fishing industry also has to be considered. In fact, during processing, up to 50% of the body weight of the fish may remain as waste. The oil content of fish viscera is an important source of polyunsaturated fatty acids that have been recovered through SFE from the viscera of Indian mackerel [164] (Rastrelliger kanagurta) or of yellowtail fish [165] (Seriola quinqueradiata). When viscera were taken from Indian mackerel, the highest recovery was obtained using ethanol as a cosolvent and a pressure of 350 bar and temperature of 60 °C, with a yield close to the one obtained with Soxhlet extraction [164]. Considering the extraction from yellowtail fish, Soxhlet extraction with ethanol has a higher yield than SFE (56% vs. 41%), but omega-3 fatty acids extracted by SFE (at 300 bar and 40 °C) have a better oxidative quality than conventionally extracted oils [165]. Special consideration has to be made for carotenoids, since several applications of SFE can be found for the recovery of these natural bioactives.
3.2.2. Carotenoids
Carotenoids are a group of phytochemicals that are responsible for the yellow, orange, and red colors of foods and vegetables [166]. They can be classified in two broad sub-categories: carotenes (such as α-carotene, β-carotene, and lycopene) and xanthophylls (such as lutein, astaxanthin, and cryptoxanthin). From the chemical point of view, xanthophylls are carotenes’ oxygenated derivatives. Indeed, they differ for the presence of oxygen atoms in the structure: carotenes are purely hydrocarbons (for example, α-carotene, β-carotene, and lycopene formula is C40H56), whereas xanthophylls are formed by carbon, hydrogen, and oxygen (lutein formula is C40H56O2, astaxanthin is C40H52O4, and cryptoxanthin is C40H56O).
Although PLE has been used to extract carotenoids, the non-polar nature of some of these components makes SFE using scCO2 the most suitable extraction technique for this category of compounds [17]. Supercritical carbon dioxide has been used alone or with low quantities of ethanol as cosolvent to extract those substances from different sources, such as tomato, vegetables, or seafood waste. Table 2 provides an overview of studies focused on the supercritical carbon dioxide extraction of carotenoids from food waste. The source from which the active principle has been extracted, the eventual amount of cosolvent added to scCO2, the operating conditions in terms of pressure, temperature and extraction time, and the recovery of the carotenoid are reported. The recovery values reported in the table are the ratios between the dry weight of the carotenoid in the extracts and the dry weight of the carotenoid contained in the original sample.
It can be noted that for the extraction of this class of compounds, in some cases CO2 alone is used; in most cases, the addition of a cosolvent is required, which in all cases is ethanol. The amount of ethanol (when present) varies from 5 to 15.5%. Carotenes are extracted in general at 300–350 bar (only in one case has a pressure of 450 bar been adopted to extract lycopene [170]), whereas in the case of xanthophylls, a wider range of pressures can be observed (astaxanthyn is extracted from crustacean waste at 200 bar [50] and focoxanthin is extracted from brown seaweed waste at 400 bar) [172]. From the perspective of operating temperatures, carotenes are extracted at higher temperatures (from 55 to 80 °C) than are xanthophylls (where the range of 40–59 °C is sufficient). The recovery values are generally high, confirming that sc-CO2 extraction is the most appropriate technique for carotenoids.
4. Combined Extractions
Pressurized Liquid Extraction (PLE) and Supercritical Fluid Extraction (SFE) are environmentally friendly advanced techniques enabling the use of non-toxic and/or GRAS solvents such as water, ethanol, or carbon dioxide for the extraction and enhancement of target compounds yield [173,174]. These green technologies are now highly encouraged due to recent increased attention to the environmental impact of processes.
The supercritical fluid extraction of polar compounds, in general, is conducted using an organic solvent as the modifier (frequently ethanol). For example, bioactive compounds are extracted from mandarin peel [175] using scCO2+ 5 and 10% ethanol, from papaya [176] seeds using scCO2+ 2–8% ethanol, from walnut husk [177] using scCO2+ 20% ethanol, or from olive residues [178] (leaves, pomace and prunings) using scCO2+ 60% ethanol. The results in terms of total phenolic compounds and antioxidant activity were, in some cases, compared to the ones obtained through conventional extraction: when phenolic antioxidants were extracted from papaya agro-industrial waste, the yield in the ethanolic extract was slightly higher than the one in the scCO2+ ethanol extract, but the latter had higher antioxidant activity in vegetable edible oil [176]. In the case of walnut green husk, SFE gave far more encouraging yields than those obtained using water, ethanol, or methanol as an extracting solvent [177]. Jafarian Asl et al. [179] compared the extraction of phytosterols and tocopherols from rapeseed oil waste by using scCO2+ ethanol and conventional Soxhlet extraction. At the optimized operating conditions, which for SFE are a temperature of 40 °C, a pressure of 350 bar (for phytosterols) and 400 bar (for tocopherol), and 5 wt% ethanol as cosolvent, SFE resulted in efficiencies that were three times higher than the ones obtained with the modified Soxhlet extraction technique. Considering that a cosolvent is needed, the benefit of using scCO2 is reduced. For this reason, PLE is preferred for the extraction of these compounds. Indeed, because sc-CO2 preferentially extracts non-polar compounds, the residue from supercritical fluid extraction (SFE) can be used as feedstock for a second extraction stage based on PLE (or other pressurized-assisted extraction processes) to complete the extraction of polar molecules. Furthermore, the future direction of food waste valorization is directed towards the combination of these innovative processes in a sequential stage extraction [16,180]. This approach was evaluated by de Aguiar et al. [76], who, from an economical point of view, investigated the possibility of coupling PLE and SFE for the recovery of phytonutrients from biquinho pepper, using supercritical carbon dioxide and ethanol 75% aqueous solution. They estimated that producing biquinho pepper-derived products was economically viable and a combined and sequential process led to a lower manufacturing cost than the commercialization price of the extracts. Similarly, Viganò et al. [181] compared two process alternatives to obtain four extract fractions from passion fruit bagasse. The first, which showed economic promise, involved a sequential multi-stage process comprising three steps of supercritical fluid extraction (SFE) and one step of pressurized liquid extraction (PLE), while the second option included the production of one extract from passion fruit rinds by single-stage PLE process. The efficiency of the biorefining process, based on consecutive supercritical carbon dioxide and pressurized liquid extractions, was also proposed in the work of Tamkutė et al. [182]. The authors used cranberry pomace as a raw material for the recovery linolenic, linoleic acids, and tocopherol rich extract by SFE and ethanol-soluble and water-soluble antioxidants from a sequential PLE extraction using ethanol and water, respectively. The feasibility and efficiency of using a sequential process with green extraction methods based on GRAS solvents from the Amazonian munguba seed was also investigated, showing a free-solvent oil mainly composed of palmitic acid can be firstly recovered by SFE, followed by a phenolic-rich fraction obtained from the defatted cake using ethanol by PLE [101]. The same approach was evaluated for several biomasses [173,183,184], indicating the increasing research interest towards these integrated green-based processes. Nevertheless, different combinations of these high-pressure-assisted processes can be taken into account, to exploit the advantages of supercritical carbon dioxide-based processes to provide solvent-free extracts. This was proposed in the work of Villanueva Bermejoet al. [185], where extract obtained by PLE with ethyl lactate was further processed by supercritical carbon dioxide antisolvent technique for the solvent removal, or extract fractionation [140].
Although PLE and SFE are considered “green extraction technologies”, a technology needs to be evaluated economically and be environmentally friendly before it can be industrialized. A tentative economic evaluation and comparison of supercritical fluid and pressurized liquid extraction analyzing single-stage and continuous-stage processes for phytonutrient extraction from red pepper is reported by Aguiar et al. [76]. It was observed that PLE is more economical for the same yield of target compound. Similarly, this result is consistent with the economic evaluation of the extraction of paclitaxel from passion fruit bagasse by Ferreira et al. [186]. The SFE process produces greater economical consumption. According to some research results, the PLE method is the most environmentally friendly, while the SFE method contributes the most to the environmental impact category. This is because in an extraction scenario, carbon dioxide needs to reach a high pressure to convert it into a supercritical fluid. The LCA results indicate that the equipment used to pump/compress and cool the CO2 stream is responsible for these energy loads, making it a decisive factor in the high environmental loads created by the SFE process. This technology can reach an economic convenience at a large scale of production (for example, industrial coffee decaffeination using SCF). To reduce energy consumption, the operation in a closed loop with total recirculation of CO2 can be used in an isobaric condition in order to avoid the decompression, cooling, and recompression of CO2 [187].
5. Conclusions
High-pressure-assisted extraction is a powerful tool for the extraction of different categories’ compounds. In particular, high-pressure liquid extraction and supercritical fluid-assisted extraction can be complementary to obtaining the complete exploitation of a food waste material before disposal. These can lead to a double effect: on the one side, the possibility of reducing food waste and on the other side, the overall environmental impact reduction.
Thanks to the high operative pressures that characterize both the PLE and SFE processes, the main advantages of these innovative techniques compared to the traditional extraction processes are a high extraction yield, better selectivity, reduced use of toxic solvent (and reduced solvent residue in the final extract), and a better preservation of sensible compounds in a short time. However, the main limitations of the high pressure operation is that on the one hand it can improve the extraction but on the other hand it causes an increase in the overall plant cost, when compared to conventional extraction methods. In addition, costs related to the security assurance of the process have to be considered and this is also related to the high operative pressure. The variable costs can be considered as practically the same as for conventional extraction; in particular, in the SFE, the cost of CO2 is not considered in the option of the closed-loop operation and in the PLE, the solvents can also be completely recovered after the extraction. The general increase in the investment cost related to the high-pressure operation can be balanced from the other side given that in these processes a product with high purity can be obtained, reducing the need for post-processing operations.
Beside the several mentioned advantages over conventional extraction processes, these technologies have mostly been applied only at a lab scale. For this reason, research should be directed towards proving the validity of these high-pressure-assisted processes in larger-scale production in order to promote their industrialization.
Conceptualization, R.C. and I.D.M.; methodology, R.C. and I.D.M.; investigation, J.L.; resources, P.P.; data curation, J.L.; writing—review and editing, R.C., M.P. and I.D.M.; supervision, M.P.; project administration, P.P. All authors have read and agreed to the published version of the manuscript.
This research received no external funding.
Not applicable.
Not applicable.
Not applicable.
The authors declare no conflict of interest.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Enlarge this image.Figure 2. Number of research articles on the main active substances extracted by PLE from 2010 to 2021 (ScienceDirect).
![View Image - Figure 3. PLE system adapted from [27,28,29]. (a) Static mode; (b) Dynamic mode. SR, solvent reservoir; FI, flow indicator; PI, pressure indicator; SV, selecting valve; EC, extraction cell; C, cooler system; PV, pressure valve; INV, inlet needle valve; OV, outlet needle valve; R, restrictor.](https://media.proquest.com/media/hms/THUM/6/trJUM?_a=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%3D%3D&_s=rATeWA8MkpybEY%2B32VxTSq6%2BpqE%3D "View Image - Figure 3. PLE system adapted from [27,28,29]. (a) Static mode; (b) Dynamic mode. SR, solvent reservoir; FI, flow indicator; PI, pressure indicator; SV, selecting valve; EC, extraction cell; C, cooler system; PV, pressure valve; INV, inlet needle valve; OV, outlet needle valve; R, restrictor.")
Enlarge this image.Figure 3. PLE system adapted from [27,28,29]. (a) Static mode; (b) Dynamic mode. SR, solvent reservoir; FI, flow indicator; PI, pressure indicator; SV, selecting valve; EC, extraction cell; C, cooler system; PV, pressure valve; INV, inlet needle valve; OV, outlet needle valve; R, restrictor.
Example for extraction of biocompounds with pressurized extraction. EtOH = ethanol; PWE = Pressurized water extraction; PHWE = Pressurized hot water extraction; SWE = Subcritical water extraction; AS = Agitation speed; S/L = solid to liquid ratio; HPTE = high- pressure high temperature extraction.
| Compounds | Source | Extraction Solvent | Operating Conditions | Sample Weight Treated | Extraction Method | Ref |
|---|---|---|---|---|---|---|
| Proteins | Sambucus nigra L. branches | water | 50 °C and 15 MPa |
4 g | PHWE | [ |
| Proteins | Wheat bran | water | 80 °C, pH 9.3, and 30 bar |
15 g | SWE | [ |
| Proteins | Spirulina | water | 40 C, pH 4, and 103.4 bar |
500 mg | PLE | [ |
| Proteins | Sea |
water | 1500 psi; |
6 g | PLE | [ |
| Proteins | Pomegranate peel (Punica granatum L.) | 70% (v/v) EtOH | 120 °C, 1500 psi static extraction time, 3 min; extraction time, 12 min | 10 g | PLE | [ |
| Proteins | Saccharina japonica | water (1.13–2.40%) | 140 °C and 50 bar for 5 min | 10 g | PLE | [ |
| Fucoidan | Saccharina japonica | NaOH (4.15%) | 140 °C and 50 bar for 5 min | 2 g | PLE | [ |
| L. barbarum |
Fruit of Lycium barbarum | water | 100 °C and 1500 psi |
250 mg | PLE | [ |
| Fucoidan | Saccharina |
water | 127.01 °C, 80 bar, S/L ratio of 0.04 g mL−1, AS of 300 rpm, |
<20 g | SWE | [ |
| β-glucan |
Tuber
|
water and 100% EtOH | 180 °C and 16.7 MPa |
500 mg | PLE | [ |
| Polysaccharides | Fucus virsoides and Cystoseira barbata | water and 0.1 M H2SO4 | 140 °C and 1500 psi |
1 g | PLE | [ |
| Polysaccharides | Sagittaria sagittifolia L. | water | 170 °C, L/S of 7 mL/g for 16 min | 1 g | SWE | [ |
| Polysaccharides | Lentinus edodes | water | 150 °C for 15 min | 1 g | SWE | [ |
| Gallic acid | Cardoon leaf | 100% EtOH | room temperature, 9 bar, static phase 2 min; dynamic phase 2 min | 40 g | Naviglio® method | [ |
| Gallic acid | Cowpea | 50% (v/v) EtOH | 170 °C, 10.34 MPa for 10 min | 5 g | PLE | [ |
| Gallic acid |
Fucus
|
58.65% (v/v) EtOH | 137.18 °C for 4.68 min | 1 g | PLE | [ |
| Gallic acid | Grape marc of Croatina cultivar and olive pomace | 75% (v/v) EtOH | 180 °C for 90 min | 5 g | HPTE | [ |
| Gallic acid | Spent coffee (Coffea arabica L.) | water | 110–190 °C; time: 15–75 min; solid-to-liquid ratio: 1:10–1:70, w/v | 4 g | SWE | [ |
| Gallic acid | Grape skins | methanol | 150 °C for 270 min | <10 g | HPTE | [ |
| Gallic acid | Biquinho pepper | 75% wt ethanol | inlet and outlet temperatures of 190 °C and 88 °C, respectively, 10 MPa | <5 kg | PLE | [ |
| Gallic acid | Red wine pomace | 37.5%: 37.5%: 25% water: EtOH: CO2 | 80 °C, 25 solvent to DW ratio (kg/kg DW), 100 bar for 30 min | <5 kg | PLE | [ |
| Polyphenolic compounds | Solanum stenotomun Peel | 80% (v/v) of ethanol in water acidified to pH 2.6 with acetic acid | 65 °C, 100 bar | 10 g/min | PLE | [ |
| Polyphenolic compounds | Saccharina japonica | NaOH (1.54–3.64%) | 140 °C and 50 bar for 5 min | 10 g | PLE | [ |
| Polyphenolic compounds | Corn Silage | 10% (v/v) EtOH | 180 °C, L/S |
<10 g | HPTE | [ |
| Caffeic acid | Spent coffee (Coffea arabica L.) | 50% (v/v) EtOH | 150 °C | 4 g | HPTE | [ |
| Caffeic acid | Spent coffee (Coffea canephora) | 54% (v/v) EtOH | L/S ratio of 10 mL/g, 150 °C for 60 min | <10 g | HPTE | [ |
| Polyphenolic compounds | Lotus seedpod | water | 140 °C, L/S ratio of 70 mL/g for 20 min | <10 g | SWE | [ |
| Polyphenolic compounds | Agave americana leaves | water | 150 °C and 10 bar for 240 min | <10 g | HPTE | [ |
| Polyphenolic compounds | Grape skins and defatted grape seeds | water | 80–120 °C,10 MPa, 2–5 mL/min, for 2 h | 65 g | SWE | [ |
| Polyphenolic compounds | Silybum marianum | EtOH | over half an hour | <10 g | Navi-glio® method | [ |
| Polyphenolic compounds | Pomegranate peels | water | 40 °C and 102.1 atm |
10 g | PWE | [ |
Extraction of carotenoids by supercritical carbon dioxide. EtOH = ethanol; P = operating pressure; T = operating temperature; t = extraction time.
| Carotenoid | Source | Sample Weight Treated | EtOH % | P Bar | T |
t min | Recovery |
Ref |
|---|---|---|---|---|---|---|---|---|
| α-carotene | Apricot peels | 5 g | 15.5 | 350 | 59 | 30 | 97.9 | [ |
| β-carotene | Tomato paste waste | 60 g | 5 | 300 | 65 | 120 | 50 | [ |
| β-carotene | Tomato processing waste | 40–50 g | / | 300 | 80 | / | 88 | [ |
| β-carotene | Sweet potato peels | 5 g | 15.5 | 350 | 59 | 30 | 99.8 | [ |
| β-carotene | Tomato peels | 5 g | 15.5 | 350 | 59 | 30 | 96.9 | [ |
| β-carotene | Apricot peels | 5 g | 15.5 | 350 | 59 | 30 | 99 | [ |
| β-carotene | Pumpkin peels | 5 g | 15.5 | 350 | 59 | 30 | 88.2 | [ |
| β-carotene | Peach peels | 5 g | 15.5 | 350 | 59 | 30 | 99.2 | [ |
| lycopene | Tomato paste waste | 60 g | 5 | 300 | 55 | 120 | 54 | [ |
| lycopene | Tomato processing waste | 40–50 g | / | 300 | 80 | / | 80 | [ |
| lycopene | Tomato processing waste | 2 kg | / | 450 | 70 | 420 | 75 | [ |
| lycopene | Tomato peels | 5 g | 15.5 | 350 | 59 | 30 | 92.5 | [ |
| lycopene | Apricot peels | 5 g | 15.5 | 350 | 59 | 30 | 83.2 | [ |
| lutein | Spinach by-products | 25 g | 10 | 390 | 56 | 215 | 72 | [ |
| lutein | Sweet potato peels | 5 g | 15.5 | 350 | 59 | 30 | 99.8 | [ |
| lutein | Peach peels | 5 g | 15.5 | 350 | 59 | 30 | 75.3 | [ |
| chlorophyll | Spinach by-products | 25 g | 10 | 390 | 56 | 215 | 50 | [ |
| astaxanthin | Crustacean waste | 25 g | 13 | 200 | 40 | 180 | 62 | [ |
| astaxanthin | Shrimp waste | 60 g | / | 250 | 45 | 120 | 47 | [ |
| fucoxanthin | Brown seaweed waste | 56 g | / | 400 | 40 | 180 | 80 | [ |
References
1. Beltrán-Ramírez, F.; Orona-Tamayo, D.; Cornejo-Corona, I.; González-Cervantes, J.L.N.; de Jesús Esparza-Claudio, J.; Quintana-Rodríguez, E. Agro-Industrial Waste Revalorization: The Growing Biorefinery. Biomass Bioenergy-Recent Trends Future Challenges; Abomohra, A. IntechOpen: London, UK, 2019; pp. 83-103. [DOI: https://dx.doi.org/10.5772/intechopen.83569]
2. Fierascu, R.C.; Fierascu, I.; Avramescu, S.M.; Sieniawska, E. Recovery of Natural Antioxidants from Agro-Industrial Side Streams through Advanced Extraction Techniques. Molecules; 2019; 24, 4212. [DOI: https://dx.doi.org/10.3390/molecules24234212] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31757027]
3. Pereira, D.T.V.; Zabot, G.L.; Reyes, F.G.R.; Iglesias, A.H.; Martínez, J. Integration of pressurized liquids and ultrasound in the extraction of bioactive compounds from passion fruit rinds: Impact on phenolic yield, extraction kinetics and technical-economic evaluation. Innov. Food Sci. Emerg. Technol.; 2021; 67, 102549. [DOI: https://dx.doi.org/10.1016/j.ifset.2020.102549]
4. Morini, S. Global Food Losses and Food Waste—Extent, Causes and Prevention; Food and Agriculture Organization of the United Nations: Roma, Italy, 2011; 37.
5. Pettinato, M. Spent Coffee Grounds Valorization by Green and Innovative Extraction Technologies: Process Optimization and Product Stabilization for Industrial Purposes; University of Genoa: Genova, Italy, 2019.
6. Belwal, T.; Chemat, F.; Venskutonis, P.R.; Cravotto, G.; Jaiswal, D.K.; Bhatt, I.D.; Devkota, H.P.; Luo, Z. Recent advances in scaling-up of non-conventional extraction techniques: Learning from successes and failures. TrAC Trends Anal. Chem.; 2020; 127, 115895. [DOI: https://dx.doi.org/10.1016/j.trac.2020.115895]
7. Khaw, K.Y.; Parat, M.O.; Shaw, P.N.; Falconer, J.R. Solvent Supercritical Fluid Technologies to Extract Bioactive Compounds from Natural Sources: A Review. Molecules; 2017; 22, 1186. [DOI: https://dx.doi.org/10.3390/molecules22071186]
8. Andreu, V.; Picó, Y. Pressurized liquid extraction of organic contaminants in environmental and food samples. TrAC Trends Anal. Chem.; 2019; 118, pp. 709-721. [DOI: https://dx.doi.org/10.1016/j.trac.2019.06.038]
9. Carabias-Martinez, R.; Rodriguez-Gonzalo, E.; Revilla-Ruiz, P.; Hernandez-Mendez, J. Pressurized liquid extraction in the analysis of food and biological samples. J. Chromatogr. A; 2005; 1089, pp. 1-17. [DOI: https://dx.doi.org/10.1016/j.chroma.2005.06.072]
10. Dobrincic, A.; Balbino, S.; Zoric, Z.; Pedisic, S.; Bursac Kovacevic, D.; Elez Garofulic, I.; Dragovic-Uzelac, V. Advanced Technologies for the Extraction of Marine Brown Algal Polysaccharides. Mar. Drugs; 2020; 18, 168. [DOI: https://dx.doi.org/10.3390/md18030168]
11. Mustafa, A.; Turner, C. Pressurized liquid extraction as a green approach in food and herbal plants extraction: A review. Anal. Chim. Acta; 2011; 703, pp. 8-18. [DOI: https://dx.doi.org/10.1016/j.aca.2011.07.018]
12. Richter, B.E.; Jones, B.A.; Ezzell, J.L.; Porter, N.L. Accelerated Solvent Extraction: A Technique for Sample Preparation. Angew. Chem.; 1996; 68, pp. 1033-1039. [DOI: https://dx.doi.org/10.1021/ac9508199]
13. Hawthorne, S.B.; Grabanski, C.B.; Martin, E.; Miller, D.J. Comparisons of Soxhlet extraction, pressurized liquid extraction, supercritical fluid extraction and subcritical water extraction for environmental solids: Recovery, selectivity and effects on sample matrix. J. Chromatogr. A; 2000; 892, pp. 421-433. [DOI: https://dx.doi.org/10.1016/S0021-9673(00)00091-1]
14. Schantz, M.M. Pressurized liquid extraction in environmental analysis. Anal. Bioanal. Chem.; 2006; 386, pp. 1043-1047. [DOI: https://dx.doi.org/10.1007/s00216-006-0648-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16896622]
15. Aguilar, L.; Williams, E.S.; Brooks, B.W.; Usenko, S. Development and application of a novel method for high-throughput determination of PCDD/Fs and PCBs in sediments. Environ. Toxicol. Chem.; 2014; 33, pp. 1529-1536. [DOI: https://dx.doi.org/10.1002/etc.2579] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24648260]
16. Santos, P.H.; Kammers, J.C.; Silva, A.P.; Oliveira, J.V.; Hense, H. Antioxidant and antibacterial compounds from feijoa leaf extracts obtained by pressurized liquid extraction and supercritical fluid extraction. Food Chem.; 2021; 344, 128620. [DOI: https://dx.doi.org/10.1016/j.foodchem.2020.128620] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33223301]
17. Herrero, M.; del Pilar Sánchez-Camargo, A.; Cifuentes, A.; Ibáñez, E. Plants, seaweeds, microalgae and food by-products as natural sources of functional ingredients obtained using pressurized liquid extraction and supercritical fluid extraction. TrAC Trends Anal. Chem.; 2015; 71, pp. 26-38. [DOI: https://dx.doi.org/10.1016/j.trac.2015.01.018]
18. Ruiz-Domínguez, M.C.; Fuentes, J.L.; Mendiola, J.A.; Cerezal-Mezquita, P.; Morales, J.; Vílchez, C.; Ibáñez, E. Bioprospecting of cyanobacterium in Chilean coastal desert, Geitlerinema sp. molecular identification and pressurized liquid extraction of bioactive compounds. Food Bioprod. Process.; 2021; 128, pp. 227-239. [DOI: https://dx.doi.org/10.1016/j.fbp.2021.06.001]
19. Gfrerer, M.; Chen, S.; Lankmayr, E.P.; Quan, X.; Yang, F. Comparison of different extraction techniques for the determination of chlorinated pesticides in animal feed. Anal. Bioanal. Chem.; 2004; 378, pp. 1861-1867. [DOI: https://dx.doi.org/10.1007/s00216-004-2492-6]
20. Celeiro, M.; Llompart, M.; Lamas, J.P.; Lores, M.; Garcia-Jares, C.; Dagnac, T. Determination of fungicides in white grape bagasse by pressurized liquid extraction and gas chromatography tandem mass spectrometry. J. Chromatogr. A; 2014; 1343, pp. 18-25. [DOI: https://dx.doi.org/10.1016/j.chroma.2014.03.057]
21. Sumere, B.R.; de Souza, M.C.; Dos Santos, M.P.; Bezerra, R.M.N.; da Cunha, D.T.; Martinez, J.; Rostagno, M.A. Combining pressurized liquids with ultrasound to improve the extraction of phenolic compounds from pomegranate peel (Punica granatum L.). Ultrason. Sonochem.; 2018; 48, pp. 151-162. [DOI: https://dx.doi.org/10.1016/j.ultsonch.2018.05.028]
22. Santos, M.P.; Souza, M.C.; Sumere, B.R.; da Silva, L.C.; Cunha, D.T.; Bezerra, R.M.N.; Rostagno, M.A. Extraction of bioactive compounds from pomegranate peel (Punica granatum L.) with pressurized liquids assisted by ultrasound combined with an expansion gas. Ultrason. Sonochem.; 2019; 54, pp. 11-17. [DOI: https://dx.doi.org/10.1016/j.ultsonch.2019.02.021]
23. Rostagno, M.A.; Palma, M.; Barroso, C.G. Pressurized liquid extraction of isoflavones from soybeans. Anal. Chim. Acta; 2004; 522, pp. 169-177. [DOI: https://dx.doi.org/10.1016/j.aca.2004.05.078]
24. Jiang, Y.; Li, P.; Li, S.P.; Wang, Y.T.; Tu, P.F. Optimization of pressurized liquid extraction of five major flavanoids from Lysimachia clethroide. J. Pharm. Biomed. Anal.; 2007; 43, pp. 341-345. [DOI: https://dx.doi.org/10.1016/j.jpba.2006.06.030] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16935451]
25. Barriada-Pereira, M.; Gonzalez-Castro, M.J.; Muniategui-Lorenzo, S.; Lopez-Mahia, P.; Prada-Rodriguez, D.; Fernandez-Fernandez, E. Comparison of pressurized liquid extraction and microwave assisted extraction for the determination of organochlorine pesticides in vegetables. Talanta; 2007; 71, pp. 1345-1351. [DOI: https://dx.doi.org/10.1016/j.talanta.2006.07.012] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19071456]
26. Murakami, T.; Kawasaki, T.; Takemura, A.; Fukutsu, N.; Kishi, N.; Kusu, F. Identification of degradation products in loxoprofen sodium adhesive tapes by liquid chromatography-mass spectrometry and dynamic pressurized liquid extraction-solid-phase extraction coupled to liquid chromatography-nuclear magnetic resonance spectroscopy. J. Chromatogr. A; 2008; 1208, pp. 164-174. [DOI: https://dx.doi.org/10.1016/j.chroma.2008.08.076] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18778828]
27. Luque-García, J.L.; Luque de Castro, M.D. Comparison of the static, dynamic and static-dynamic pressurised liquid extraction modes for the removal of nitrated polycyclic aromatic hydrocarbons from soil with on-line filtration-preconcentration. J. Chromatogr. A; 2003; 1010, pp. 129-140. [DOI: https://dx.doi.org/10.1016/S0021-9673(03)01106-3]
28. Pitipanapong, J.; Chitprasert, S.; Goto, M.; Jiratchariyakul, W.; Sasaki, M.; Shotipruk, A. New approach for extraction of charantin from Momordica charantia with pressurized liquid extraction. Sep. Purif. Technol.; 2007; 52, pp. 416-422. [DOI: https://dx.doi.org/10.1016/j.seppur.2005.11.037]
29. Wicker, A.P.; Carlton, D.D., Jr.; Tanaka, K.; Nishimura, M.; Chen, V.; Ogura, T.; Hedgepeth, W.; Schug, K.A. On-line supercritical fluid extraction-supercritical fluid chromatography-mass spectrometry of polycyclic aromatic hydrocarbons in soil. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci.; 2018; 1086, pp. 82-88. [DOI: https://dx.doi.org/10.1016/j.jchromb.2018.04.014]
30. Naviglio, D. Naviglio’s Principle and Presentation of an Innovative Solid–Liquid Extraction Technology: Extractor Naviglio®. Anal. Lett.; 2003; 36, pp. 1647-1659. [DOI: https://dx.doi.org/10.1081/AL-120021555]
31. Naviglio, D.; Pizzolongo, F.; Ferrara, L.; Aragòn, A.; Santini, A. Extraction of pure lycopene from industrial tomato by-products in water using a new high-pressure process. J. Sci. Food Agric.; 2008; 88, pp. 2414-2420. [DOI: https://dx.doi.org/10.1002/jsfa.3334]
32. Aliakbarian, B.; Casazza, A.A.; Perego, P. Valorization of olive oil solid waste using high pressure–high temperature reactor. Food Chem.; 2011; 128, pp. 704-710. [DOI: https://dx.doi.org/10.1016/j.foodchem.2011.03.092]
33. Pettinato, M.; Trucillo, P.; Campardelli, R.; Perego, P.; Reverchon, E. Bioactives extraction from spent coffee grounds and liposome encapsulation by a combination of green technologies. Chem. Eng. Process.-Process Intensif.; 2020; 151, 107911. [DOI: https://dx.doi.org/10.1016/j.cep.2020.107911]
34. Hu, W.; Chen, S.; Wu, D.; Zhu, K.; Ye, X. Manosonication assisted extraction and characterization of pectin from different citrus peel wastes. Food Hydrocoll.; 2021; 121, 106952. [DOI: https://dx.doi.org/10.1016/j.foodhyd.2021.106952]
35. Chienthavorn, O.; Insuan, W. Superheated Water Extraction of Lime Peel: A Comparison with Conventional Methods. Anal. Lett.; 2004; 37, pp. 2393-2409. [DOI: https://dx.doi.org/10.1081/AL-200028189]
36. Ding, W.-H.; Fann, J.C.H. Determination of linear alkylbenzenesulfonates in sediments using pressurized liquid extraction and ion-pair derivatization gas chromatography-mass spectrometry. Anal. Chim. Acta; 2000; 408, pp. 291-297. [DOI: https://dx.doi.org/10.1016/S0003-2670(99)00870-3]
37. Herrera, M.C.; Prados-Rosales, R.C.; Luque-García, J.L.; Luque de Castro, M.D. Static–dynamic pressurized hot water extraction coupled to on-line filtration–solid-phase extraction–high-performance liquid chromatography–post-column derivatization–fluorescence detection for the analysis of N-methylcarbamates in foods. Anal. Chim. Acta; 2002; 463, pp. 189-197. [DOI: https://dx.doi.org/10.1016/S0003-2670(02)00427-0]
38. Dobrincic, A.; Pedisic, S.; Zoric, Z.; Jurin, M.; Roje, M.; Coz-Rakovac, R.; Dragovic-Uzelac, V. Microwave Assisted Extraction and Pressurized Liquid Extraction of Sulfated Polysaccharides from Fucus virsoides and Cystoseira barbata. Foods; 2021; 10, 1481. [DOI: https://dx.doi.org/10.3390/foods10071481]
39. Vazquez-Roig, P.; Picó, Y. Pressurized liquid extraction of organic contaminants in environmental and food samples. TrAC Trends Anal. Chem.; 2015; 71, pp. 55-64. [DOI: https://dx.doi.org/10.1016/j.trac.2015.04.014]
40. Herbst, G.; Hamerski, F.; Errico, M.; Corazza, M.L. Pressurized liquid extraction of brewer’s spent grain: Kinetics and crude extracts characterization. J. Ind. Eng. Chem.; 2021; 102, pp. 370-383. [DOI: https://dx.doi.org/10.1016/j.jiec.2021.07.020]
41. Pronyk, C.; Mazza, G. Design and scale-up of pressurized fluid extractors for food and bioproducts. J. Food Eng.; 2009; 95, pp. 215-226. [DOI: https://dx.doi.org/10.1016/j.jfoodeng.2009.06.002]
42. Osorio-Tobón, J.F.; Carvalho, P.I.N.; Rostagno, M.A.; Petenate, A.J.; Meireles, M.A.A. Extraction of curcuminoids from deflavored turmeric (Curcuma longa L.) using pressurized liquids: Process integration and economic evaluation. J. Supercrit. Fluids; 2014; 95, pp. 167-174. [DOI: https://dx.doi.org/10.1016/j.supflu.2014.08.012]
43. Nathia-Neves, G.; Tarone, A.G.; Tosi, M.M.; Marostica Junior, M.R.; Meireles, M.A.A. Extraction of bioactive compounds from genipap (Genipa americana L.) by pressurized ethanol: Iridoids, phenolic content and antioxidant activity. Food Res. Int.; 2017; 102, pp. 595-604. [DOI: https://dx.doi.org/10.1016/j.foodres.2017.09.041]
44. Chen, Y.; Zhou, S.; Wang, Y.; Li, L. Screening solvents to extract phenol from aqueous solutions by the COSMO-SAC model and extraction process simulation. Fluid Phase Equilibria; 2017; 451, pp. 12-24. [DOI: https://dx.doi.org/10.1016/j.fluid.2017.08.007]
45. Aliakbarian, B.; Fathi, A.; Perego, P.; Dehghani, F. Extraction of antioxidants from winery wastes using subcritical water. J. Supercrit. Fluids; 2012; 65, pp. 18-24. [DOI: https://dx.doi.org/10.1016/j.supflu.2012.02.022]
46. Nastić, N.; Švarc-Gajić, J.; Delerue-Matos, C.; Morais, S.; Barroso, M.F.; Moreira, M.M. Subcritical water extraction of antioxidants from mountain germander (Teucrium montanum L.). J. Supercrit. Fluids; 2018; 138, pp. 200-206. [DOI: https://dx.doi.org/10.1016/j.supflu.2018.04.019]
47. da Silva, M.F.; Casazza, A.A.; Ferrari, P.F.; Aliakbarian, B.; Converti, A.; Bezerra, R.P.; Porto, A.L.F.; Perego, P. Recovery of phenolic compounds of food concern from Arthrospira platensis by green extraction techniques. Algal Res.; 2017; 25, pp. 391-401. [DOI: https://dx.doi.org/10.1016/j.algal.2017.05.027]
48. Pettinato, M.; Casazza, A.A.; Ferrari, P.F.; Palombo, D.; Perego, P. Eco-sustainable recovery of antioxidants from spent coffee grounds by microwave-assisted extraction: Process optimization, kinetic modeling and biological validation. Food Bioprod. Process.; 2019; 114, pp. 31-42. [DOI: https://dx.doi.org/10.1016/j.fbp.2018.11.004]
49. Li, J.; Pettinato, M.; Casazza, A.A.; Perego, P. A Comprehensive Optimization of Ultrasound-Assisted Extraction for Lycopene Recovery from Tomato Waste and Encapsulation by Spray Drying. Processes; 2022; 10, 308. [DOI: https://dx.doi.org/10.3390/pr10020308]
50. Nunes, A.N.; Roda, A.; Gouveia, L.F.; Fernández, N.; Bronze, M.R.; Matias, A.A. Astaxanthin Extraction from Marine Crustacean Waste Streams: An Integrate Approach between Microwaves and Supercritical Fluids. ACS Sustain. Chem. Eng.; 2021; 9, pp. 3050-3059. [DOI: https://dx.doi.org/10.1021/acssuschemeng.0c06534]
51. Roy, V.C.; Getachew, A.T.; Cho, Y.-J.; Park, J.-S.; Chun, B.-S. Recovery and bio-potentialities of astaxanthin-rich oil from shrimp (Penaeus monodon) waste and mackerel (Scomberomous niphonius) skin using concurrent supercritical CO2 extraction. J. Supercrit. Fluids; 2020; 159, 104773. [DOI: https://dx.doi.org/10.1016/j.supflu.2020.104773]
52. Ferrari, P.F.; Pettinato, M.; Casazza, A.A.; De Negri Atanasio, G.; Palombo, D.; Perego, P. Polyphenols from Nerone Gold 26/6, a new pigmented rice, via non-conventional extractions: Antioxidant properties and biological validation. J. Chem. Technol. Biotechnol.; 2021; 96, pp. 1691-1699. [DOI: https://dx.doi.org/10.1002/jctb.6694]
53. Spennati, E.; Mirizadeh, S.; Casazza, A.A.; Solisio, C.; Converti, A. Chlorella vulgaris and Arthrospira platensis growth in a continuous membrane photobioreactor using industrial winery wastewater. Algal Res.; 2021; 60, 102519. [DOI: https://dx.doi.org/10.1016/j.algal.2021.102519]
54. Aliakbarian, B.; Casazza, A.A.; Montoya, E.J.O.; Convert, A. Valorisation of Olive Oil Solid Wastes: Valuable Compounds Recovery Using High Pressure- High Temperature. J. Biotechnol.; 2010; 150, 332. [DOI: https://dx.doi.org/10.1016/j.jbiotec.2010.09.341]
55. Casazza, A.A.; Pettinato, M.; Perego, P. Polyphenols from apple skins: A study on microwave-assisted extraction optimization and exhausted solid characterization. Sep. Purif. Technol.; 2020; 240, 116640. [DOI: https://dx.doi.org/10.1016/j.seppur.2020.116640]
56. Šalplachta, J.; Hohnová, B. Pressurized hot water extraction of proteins from Sambucus nigra L. branches. Ind. Crops Prod.; 2017; 108, pp. 312-315. [DOI: https://dx.doi.org/10.1016/j.indcrop.2017.06.053]
57. Celiktas, M.S.; Kirsch, C.; Smirnova, I. Cascade processing of wheat bran through a biorefinery approach. Energy Convers. Manag.; 2014; 84, pp. 633-639. [DOI: https://dx.doi.org/10.1016/j.enconman.2014.04.039]
58. Zhou, J.; Wang, M.; Carrillo, C.; Zhu, Z.; Brncic, M.; Berrada, H.; Barba, F.J. Impact of Pressurized Liquid Extraction and pH on Protein Yield, Changes in Molecular Size Distribution and Antioxidant Compounds Recovery from Spirulina. Foods; 2021; 10, 2153. [DOI: https://dx.doi.org/10.3390/foods10092153] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34574263]
59. de la Fuente, B.; Pallares, N.; Berrada, H.; Barba, F.J. Development of Antioxidant Protein Extracts from Gilthead Sea Bream (Sparus aurata) Side Streams Assisted by Pressurized Liquid Extraction (PLE). Mar. Drugs; 2021; 19, 199. [DOI: https://dx.doi.org/10.3390/md19040199]
60. de la Fuente, B.; Pallares, N.; Barba, F.J.; Berrada, H. An Integrated Approach for the Valorization of Sea Bass (Dicentrarchus labrax) Side Streams: Evaluation of Contaminants and Development of Antioxidant Protein Extracts by Pressurized Liquid Extraction. Foods; 2021; 10, 546. [DOI: https://dx.doi.org/10.3390/foods10030546] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33800768]
61. de la Fuente, B.; Pallarés, N.; Berrada, H.; Barba, F.J. Salmon (Salmo salar) Side Streams as a Bioresource to Obtain Potential Antioxidant Peptides after Applying Pressurized Liquid Extraction (PLE). Mar. Drugs; 2021; 19, 323. [DOI: https://dx.doi.org/10.3390/md19060323]
62. Hernández-Corroto, E.; Plaza, M.; Marina, M.L.; García, M.C. Sustainable extraction of proteins and bioactive substances from pomegranate peel (Punica granatum L.) using pressurized liquids and deep eutectic solvents. Innov. Food Sci. Emerg. Technol.; 2020; 60, 102314. [DOI: https://dx.doi.org/10.1016/j.ifset.2020.102314]
63. Saravana, P.S.; Cho, Y.J.; Park, Y.B.; Woo, H.C.; Chun, B.S. Structural, antioxidant, and emulsifying activities of fucoidan from Saccharina japonica using pressurized liquid extraction. Carbohydr. Polym.; 2016; 153, pp. 518-525. [DOI: https://dx.doi.org/10.1016/j.carbpol.2016.08.014] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27561524]
64. Hao, W.; Wang, S.F.; Zhao, J.; Li, S.P. Effects of extraction methods on immunology activity and chemical profiles of Lycium barbarum polysaccharides. J. Pharm. Biomed. Anal.; 2020; 185, 113219. [DOI: https://dx.doi.org/10.1016/j.jpba.2020.113219] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32145536]
65. Chen, X.J.; Guo, B.L.; Li, S.P.; Zhang, Q.W.; Tu, P.F.; Wang, Y.T. Simultaneous determination of 15 flavonoids in Epimedium using pressurized liquid extraction and high-performance liquid chromatography. J. Chromatogr. A; 2007; 1163, pp. 96-104. [DOI: https://dx.doi.org/10.1016/j.chroma.2007.06.020] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17606269]
66. Saravana, P.S.; Tilahun, A.; Gerenew, C.; Tri, V.D.; Kim, N.H.; Kim, G.-D.; Woo, H.-C.; Chun, B.-S. Subcritical water extraction of fucoidan from Saccharina japonica: Optimization, characterization and biological studies. J. Appl. Phycol.; 2017; 30, pp. 579-590. [DOI: https://dx.doi.org/10.1007/s10811-017-1245-9]
67. Tejedor-Calvo, E.; Morales, D.; Marco, P.; Sanchez, S.; Garcia-Barreda, S.; Smiderle, F.R.; Iacomini, M.; Villalva, M.; Santoyo, S.; Soler-Rivas, C. Screening of bioactive compounds in truffles and evaluation of pressurized liquid extractions (PLE) to obtain fractions with biological activities. Food Res. Int.; 2020; 132, 109054. [DOI: https://dx.doi.org/10.1016/j.foodres.2020.109054] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32331654]
68. Zhang, J.; Wen, C.; Chen, M.; Gu, J.; Zhou, J.; Duan, Y.; Zhang, H.; Ma, H. Antioxidant activities of Sagittaria sagittifolia L. polysaccharides with subcritical water extraction. Int. J. Biol. Macromol.; 2019; 134, pp. 172-179. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2019.05.047]
69. Zhang, J.; Wen, C.; Gu, J.; Ji, C.; Duan, Y.; Zhang, H. Effects of subcritical water extraction microenvironment on the structure and biological activities of polysaccharides from Lentinus edodes. Int. J. Biol. Macromol.; 2019; 123, pp. 1002-1011. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2018.11.194]
70. Mirpoor, S.F.; Varriale, S.; Porta, R.; Naviglio, D.; Spennato, M.; Gardossi, L.; Giosafatto, C.V.L.; Pezzella, C. A biorefinery approach for the conversion of Cynara cardunculus biomass to active films. Food Hydrocoll.; 2021; 122, 107099. [DOI: https://dx.doi.org/10.1016/j.foodhyd.2021.107099]
71. Avanza, M.V.; Álvarez-Rivera, G.; Cifuentes, A.; Mendiola, J.A.; Ibáñez, E. Phytochemical and Functional Characterization of Phenolic Compounds from Cowpea (Vigna unguiculata (L.) Walp.) Obtained by Green Extraction Technologies. Agronomy; 2021; 11, 162. [DOI: https://dx.doi.org/10.3390/agronomy11010162]
72. Sumampouw, G.A.; Jacobsen, C.; Getachew, A.T. Optimization of phenolic antioxidants extraction from Fucus vesiculosus by pressurized liquid extraction. J. Appl. Phycol.; 2021; 33, pp. 1195-1207. [DOI: https://dx.doi.org/10.1007/s10811-020-02362-3]
73. Paini, M.; Casazza, A.A.; Aliakbarian, B.; Perego, P.; Binello, A.; Cravotto, G. Influence of ethanol/water ratio in ultrasound and high-pressure/high-temperature phenolic compound extraction from agri-food waste. Int. J. Food Sci. Technol.; 2016; 51, pp. 349-358. [DOI: https://dx.doi.org/10.1111/ijfs.12956]
74. Xu, H.; Wang, W.; Liu, X.; Yuan, F.; Gao, Y. Antioxidative phenolics obtained from spent coffee grounds (Coffea arabica L.) by subcritical water extraction. Ind. Crops Prod.; 2015; 76, pp. 946-954. [DOI: https://dx.doi.org/10.1016/j.indcrop.2015.07.054]
75. Casazza, A.A.; Aliakbarian, B.; Sannita, E.; Perego, P. High-pressure high-temperature extraction of phenolic compounds from grape skins. Int. J. Food Sci. Technol.; 2012; 47, pp. 399-405. [DOI: https://dx.doi.org/10.1111/j.1365-2621.2011.02853.x]
76. de Aguiar, A.C.; Osorio-Tobón, J.F.; Viganó, J.; Martínez, J. Economic evaluation of supercritical fluid and pressurized liquid extraction to obtain phytonutrients from biquinho pepper: Analysis of single and sequential-stage processes. J. Supercrit. Fluids; 2020; 165, 104935. [DOI: https://dx.doi.org/10.1016/j.supflu.2020.104935]
77. Croxatto Vega, G.; Sohn, J.; Voogt, J.; Birkved, M.; Olsen, S.I.; Nilsson, A.E. Insights from combining techno-economic and life cycle assessment—A case study of polyphenol extraction from red wine pomace. Resour. Conserv. Recycl.; 2021; 167, 105318. [DOI: https://dx.doi.org/10.1016/j.resconrec.2020.105318]
78. Cardoso, L.C.; Serrano, C.M.; Quintero, E.T.; Lopez, C.P.; Antezana, R.M.; Martinez de la Ossa, E.J. High pressure extraction of antioxidants from Solanum stenotomun peel. Molecules; 2013; 18, pp. 3137-3151. [DOI: https://dx.doi.org/10.3390/molecules18033137]
79. Kuzmanovića, M.; Tišmaa, M.; Bucić-Kojića, A.; Casazzab, A.A.; Marco Painib, B.A.; Perego, P. High–Pressure and Temperature Extraction of Phenolic Compounds from Corn Silage. Chem. Eng. Traansactions; 2015; 43, pp. 133-138. [DOI: https://dx.doi.org/10.3303/CET1543023]
80. Silva, M.F.d.; Pettinato, M.; Casazza, A.A.; Maciel, M.I.S.; Perego, P. Design and evaluation of non-conventional extraction for bioactive compounds recovery from spent coffee (Coffea arabica L.) grounds. Chem. Eng. Res. Des.; 2022; 177, pp. 418-430. [DOI: https://dx.doi.org/10.1016/j.cherd.2021.11.011]
81. Yan, Z.; Zhang, H.; Dzah, C.S.; Zhang, J.; Diao, C.; Ma, H.; Duan, Y. Subcritical water extraction, identification, antioxidant and antiproliferative activity of polyphenols from lotus seedpod. Sep. Purif. Technol.; 2020; 236, 116217. [DOI: https://dx.doi.org/10.1016/j.seppur.2019.116217]
82. Ben Hamissa, A.M.; Seffen, M.; Aliakbarian, B.; Casazza, A.A.; Perego, P.; Converti, A. Phenolics extraction from Agave americana (L.) leaves using high-temperature, high-pressure reactor. Food Bioprod. Process.; 2012; 90, pp. 17-21. [DOI: https://dx.doi.org/10.1016/j.fbp.2010.11.008]
83. Duba, K.S.; Casazza, A.A.; Mohamed, H.B.; Perego, P.; Fiori, L. Extraction of polyphenols from grape skins and defatted grape seeds using subcritical water: Experiments and modeling. Food Bioprod. Process.; 2015; 94, pp. 29-38. [DOI: https://dx.doi.org/10.1016/j.fbp.2015.01.001]
84. De Marco, A.; Luongo, G.; Di Marino, C.; De Tommaso, G.; Di Fabio, G.; Zarrelli, A. Silymarin from Silybum marianum by Naviglio’s extractor: A new and very efficient approach. Nat. Prod. Res.; 2021; 35, pp. 2621-2627. [DOI: https://dx.doi.org/10.1080/14786419.2019.1687474] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31691591]
85. Çam, M.; Hışıl, Y. Pressurised water extraction of polyphenols from pomegranate peels. Food Chem.; 2010; 123, pp. 878-885. [DOI: https://dx.doi.org/10.1016/j.foodchem.2010.05.011]
86. Pojić, M.; Mišan, A.; Tiwari, B. Eco-innovative technologies for extraction of proteins for human consumption from renewable protein sources of plant origin. Trends Food Sci. Technol.; 2018; 75, pp. 93-104. [DOI: https://dx.doi.org/10.1016/j.tifs.2018.03.010]
87. Kumar, M.; Tomar, M.; Potkule, J.; Verma, R.; Punia, S.; Mahapatra, A.; Belwal, T.; Dahuja, A.; Joshi, S.; Berwal, M.K. et al. Advances in the plant protein extraction: Mechanism and recommendations. Food Hydrocoll.; 2021; 115, 106595. [DOI: https://dx.doi.org/10.1016/j.foodhyd.2021.106595]
88. Contreras, M.D.M.; Lama-Munoz, A.; Manuel Gutierrez-Perez, J.; Espinola, F.; Moya, M.; Castro, E. Protein extraction from agri-food residues for integration in biorefinery: Potential techniques and current status. Bioresour. Technol.; 2019; 280, pp. 459-477. [DOI: https://dx.doi.org/10.1016/j.biortech.2019.02.040] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30777702]
89. Shi, L. Bioactivities, isolation and purification methods of polysaccharides from natural products: A review. Int. J. Biol. Macromol.; 2016; 92, pp. 37-48. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2016.06.100] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27377457]
90. Shao, P.; Feng, J.; Sun, P.; Xiang, N.; Lu, B.; Qiu, D. Recent advances in improving stability of food emulsion by plant polysaccharides. Food Res. Int.; 2020; 137, 109376. [DOI: https://dx.doi.org/10.1016/j.foodres.2020.109376]
91. Liu, Y.; Sun, Y.; Huang, G. Preparation and antioxidant activities of important traditional plant polysaccharides. Int. J. Biol. Macromol.; 2018; 111, pp. 780-786. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2018.01.086]
92. Ying, M.; Yu, Q.; Zheng, B.; Wang, H.; Wang, J.; Chen, S.; Nie, S.; Xie, M. Cultured Cordyceps sinensis polysaccharides modulate intestinal mucosal immunity and gut microbiota in cyclophosphamide-treated mice. Carbohydr. Polym.; 2020; 235, 115957. [DOI: https://dx.doi.org/10.1016/j.carbpol.2020.115957]
93. Sun, J.; Zhou, B.; Tang, C.; Gou, Y.; Chen, H.; Wang, Y.; Jin, C.; Liu, J.; Niu, F.; Kan, J. et al. Characterization, antioxidant activity and hepatoprotective effect of purple sweetpotato polysaccharides. Int. J. Biol. Macromol.; 2018; 115, pp. 69-76. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2018.04.033]
94. Yan, T.; Nian, T.; Liao, Z.; Xiao, F.; Wu, B.; Bi, K.; He, B.; Jia, Y. Antidepressant effects of a polysaccharide from okra (Abelmoschus esculentus (L) Moench) by anti-inflammation and rebalancing the gut microbiota. Int. J. Biol. Macromol.; 2020; 144, pp. 427-440. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2019.12.138] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31862370]
95. Mohammed, A.S.A.; Naveed, M.; Jost, N. Polysaccharides; Classification, Chemical Properties, and Future Perspective Applications in Fields of Pharmacology and Biological Medicine (A Review of Current Applications and Upcoming Potentialities). J. Polym. Environ.; 2021; 29, pp. 2359-2371. [DOI: https://dx.doi.org/10.1007/s10924-021-02052-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33526994]
96. Huang, H.; Huang, G. Extraction, separation, modification, structural characterization, and antioxidant activity of plant polysaccharides. Chem. Biol. Drug Des.; 2020; 96, pp. 1209-1222. [DOI: https://dx.doi.org/10.1111/cbdd.13794]
97. Leong, Y.K.; Yang, F.C.; Chang, J.S. Extraction of polysaccharides from edible mushrooms: Emerging technologies and recent advances. Carbohydr. Polym.; 2021; 251, 117006. [DOI: https://dx.doi.org/10.1016/j.carbpol.2020.117006]
98. Yang, B.; Wu, Q.; Luo, Y.; Yang, Q.; Wei, X.; Kan, J. High-pressure ultrasonic-assisted extraction of polysaccharides from Hovenia dulcis: Extraction, structure, antioxidant activity and hypoglycemic. Int. J. Biol. Macromol.; 2019; 137, pp. 676-687. [DOI: https://dx.doi.org/10.1016/j.ijbiomac.2019.07.034] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31284006]
99. Luthria, D.; Biswas, R.; Natarajan, S. Comparison of extraction solvents and techniques used for the assay of isoflavones from soybean. Food Chem.; 2007; 105, pp. 325-333. [DOI: https://dx.doi.org/10.1016/j.foodchem.2006.11.047]
100. Xie, J.J.; Lu, J.; Qian, Z.M.; Yu, Y.; Duan, J.A.; Li, S.P. Optimization and comparison of five methods for extraction of coniferyl ferulate from Angelica sinensis. Molecules; 2009; 14, pp. 555-565. [DOI: https://dx.doi.org/10.3390/molecules14010555] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19169202]
101. Teixeira, G.L.; Maciel, L.G.; Mazzutti, S.; Barbi, R.C.T.; Ribani, R.H.; Ferreira, S.R.S.; Block, J.M. Sequential green extractions based on supercritical carbon dioxide and pressurized ethanol for the recovery of lipids and phenolics from Pachira aquatica seeds. J. Clean. Prod.; 2021; 306, 127223. [DOI: https://dx.doi.org/10.1016/j.jclepro.2021.127223]
102. Berrada, H.; Borrull, F.; Font, G.; Marce, R.M. Determination of macrolide antibiotics in meat and fish using pressurized liquid extraction and liquid chromatography-mass spectrometry. J. Chromatogr. A; 2008; 1208, pp. 83-89. [DOI: https://dx.doi.org/10.1016/j.chroma.2008.08.107]
103. Khan, M.R.; Busquets, R.; Santos, F.J.; Puignou, L. New method for the analysis of heterocyclic amines in meat extracts using pressurised liquid extraction and liquid chromatography-tandem mass spectrometry. J. Chromatogr. A; 2008; 1194, pp. 155-160. [DOI: https://dx.doi.org/10.1016/j.chroma.2008.04.058]
104. Bosellia, E.; Velazcoa, V.; Cabonib, M.F.; Lerckera, G. Pressurized liquid extraction of lipids for the determination of oxysterols in egg-containing food. J. Chromatogr. A; 2001; 917, pp. 239-244. [DOI: https://dx.doi.org/10.1016/S0021-9673(01)00688-4]
105. Lund, M.; Duedahl-Olesen, L.; Christensen, J.H. Extraction of polycyclic aromatic hydrocarbons from smoked fish using pressurized liquid extraction with integrated fat removal. Talanta; 2009; 79, pp. 10-15. [DOI: https://dx.doi.org/10.1016/j.talanta.2009.02.048] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19376336]
106. Catchpole, O.; Moreno, T.; Montañes, F.; Tallon, S. Perspectives on processing of high value lipids using supercritical fluids. J. Supercrit. Fluids; 2018; 134, pp. 260-268. [DOI: https://dx.doi.org/10.1016/j.supflu.2017.12.001]
107. Villagrasa, M.; Guillamon, M.; Navarro, A.; Eljarrat, E.; Barcelo, D. Development of a pressurized liquid extraction-solid-phase extraction followed by liquid chromatography-electrospray ionization tandem mass spectrometry method for the quantitative determination of benzoxazolinones and their degradation products in agricultural soil. J. Chromatogr. A; 2008; 1179, pp. 190-197. [DOI: https://dx.doi.org/10.1016/j.chroma.2007.11.087] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18096177]
108. Onofrejova, L.; Vasickova, J.; Klejdus, B.; Stratil, P.; Misurcova, L.; Kracmar, S.; Kopecky, J.; Vacek, J. Bioactive phenols in algae: The application of pressurized-liquid and solid-phase extraction techniques. J. Pharm. Biomed. Anal.; 2010; 51, pp. 464-470. [DOI: https://dx.doi.org/10.1016/j.jpba.2009.03.027]
109. Andujar-Ortiz, I.; Pozo-Bayon, M.A.; Garcia-Ruiz, A.; Moreno-Arribas, M.V. Role of specific components from commercial inactive dry yeast winemaking preparations on the growth of wine lactic acid bacteria. J. Agric. Food Chem.; 2010; 58, pp. 8392-8399. [DOI: https://dx.doi.org/10.1021/jf101132t]
110. Gómez-Ariza, J.L.; Caro de la Torre, M.A.; Giráldez, I.; Morales, E. Speciation analysis of selenium compounds in yeasts using pressurised liquid extraction and liquid chromatography–microwave-assisted digestion–hydride generation–atomic fluorescence spectrometry. Anal. Chim. Acta; 2004; 524, pp. 305-314. [DOI: https://dx.doi.org/10.1016/j.aca.2004.02.044]
111. Bligh, E.G.; Dyer, W.J. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol.; 1959; 37, pp. 911-917. [DOI: https://dx.doi.org/10.1139/y59-099]
112. Folch, J.; Lees, M.; Stanley, G.H.S. A Simple Method for the Isolation and Purification of Total Lipides from Animal Tissues. J. Biol. Chem.; 1957; 226, pp. 497-509. [DOI: https://dx.doi.org/10.1016/S0021-9258(18)64849-5]
113. He, Y.; Huang, Z.; Zhong, C.; Guo, Z.; Chen, B. Pressurized liquid extraction with ethanol as a green and efficient technology to lipid extraction of Isochrysis biomass. Bioresour. Technol.; 2019; 293, 122049. [DOI: https://dx.doi.org/10.1016/j.biortech.2019.122049]
114. Cescut, J.; Severac, E.; Molina-Jouve, C.; Uribelarrea, J.L. Optimizing pressurized liquid extraction of microbial lipids using the response surface method. J. Chromatogr. A; 2011; 1218, pp. 373-379. [DOI: https://dx.doi.org/10.1016/j.chroma.2010.12.003] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21185025]
115. Villanueva-Bermejo, D.; Calvo, M.V.; Castro-Gomez, P.; Fornari, T.; Fontecha, J. Production of omega 3-rich oils from underutilized chia seeds. Comparison between supercritical fluid and pressurized liquid extraction methods. Food Res. Int.; 2019; 115, pp. 400-407. [DOI: https://dx.doi.org/10.1016/j.foodres.2018.10.085] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30599958]
116. Chemat, F.; Abert-Vian, M.; Fabiano-Tixier, A.S.; Strube, J.; Uhlenbrock, L.; Gunjevic, V.; Cravotto, G. Green extraction of natural products. Origins, current status, and future challenges. TrAC Trends Anal. Chem.; 2019; 118, pp. 248-263. [DOI: https://dx.doi.org/10.1016/j.trac.2019.05.037]
117. Mazzutti, S.; Pedrosa, R.C.; Salvador Ferreira, S.R. Green processes in Foodomics. Supercritical Fluid Extraction of Bioactives. Comprehensive Foodomics; Elsevier: Sergipe, Brazil, 2021; pp. 725-743.
118. Quintin, D.; Garcia-Gomez, P.; Ayuso, M.; Sanmartin, A.M. Active biocompounds to improve food nutritional value. Trends Food Sci. Technol.; 2019; 84, pp. 19-21. [DOI: https://dx.doi.org/10.1016/j.tifs.2018.03.024]
119. Osorio-Tobon, J.F. Recent advances and comparisons of conventional and alternative extraction techniques of phenolic compounds. J. Food Sci. Technol.; 2020; 57, pp. 4299-4315. [DOI: https://dx.doi.org/10.1007/s13197-020-04433-2] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33087945]
120. Temelli, F. Perspectives on supercritical fluid processing of fats and oils. J. Supercrit. Fluids; 2009; 47, pp. 583-590. [DOI: https://dx.doi.org/10.1016/j.supflu.2008.10.014]
121. Wianowska, D.; Gil, M. Critical approach to PLE technique application in the analysis of secondary metabolites in plants. TrAC Trends Anal. Chem.; 2019; 114, pp. 314-325. [DOI: https://dx.doi.org/10.1016/j.trac.2019.03.018]
122. Hogervorst Cvejić, J.; Atanacković Krstonošić, M.; Bursać, M.; Miljić, U. Polyphenols. Nutraceutical and Functional Food Components; Academic Press: Novi Sad, Serbia, 2017; pp. 203-258.
123. Socrier, L.; Quero, A.; Verdu, M.; Song, Y.; Molinie, R.; Mathiron, D.; Pilard, S.; Mesnard, F.; Morandat, S. Flax phenolic compounds as inhibitors of lipid oxidation: Elucidation of their mechanisms of action. Food Chem.; 2019; 274, pp. 651-658. [DOI: https://dx.doi.org/10.1016/j.foodchem.2018.08.126]
124. Yousefian, M.; Shakour, N.; Hosseinzadeh, H.; Hayes, A.W.; Hadizadeh, F.; Karimi, G. The natural phenolic compounds as modulators of NADPH oxidases in hypertension. Phytomedicine; 2019; 55, pp. 200-213. [DOI: https://dx.doi.org/10.1016/j.phymed.2018.08.002]
125. Luna-Guevara, M.L.; Luna-Guevara, J.J.; Hernández-Carranza, P.; Ruíz-Espinosa, H.; Ochoa-Velasco, C.E. Phenolic Compounds: A Good Choice Against Chronic Degenerative Diseases. Stud. Nat. Prod. Chem.; 2018; 59, pp. 79-108.
126. Ullah, M.; Uddin, Z.; Song, Y.H.; Li, Z.P.; Kim, J.Y.; Ban, Y.J.; Park, K.H. Bacterial neuraminidase inhibition by phenolic compounds from Usnea longissima. S. Afr. J. Bot.; 2019; 120, pp. 326-330. [DOI: https://dx.doi.org/10.1016/j.sajb.2018.10.020]
127. Chen, L.; Gnanaraj, C.; Arulselvan, P.; El-Seedi, H.; Teng, H. A review on advanced microencapsulation technology to enhance bioavailability of phenolic compounds: Based on its activity in the treatment of Type 2 Diabetes. Trends Food Sci. Technol.; 2019; 85, pp. 149-162. [DOI: https://dx.doi.org/10.1016/j.tifs.2018.11.026]
128. Martini, S.; Conte, A.; Tagliazucchi, D. Bioaccessibility, bioactivity and cell metabolism of dark chocolate phenolic compounds after in vitro gastro-intestinal digestion. J. Funct. Foods; 2018; 49, pp. 424-436. [DOI: https://dx.doi.org/10.1016/j.jff.2018.09.005]
129. Galanakis, C.M.; Tsatalas, P.; Charalambous, Z.; Galanakis, I.M. Control of microbial growth in bakery products fortified with polyphenols recovered from olive mill wastewater. Environ. Technol. Innov.; 2018; 10, pp. 1-15. [DOI: https://dx.doi.org/10.1016/j.eti.2018.01.006]
130. Basanta, M.F.; Rojas, A.M.; Martinefski, M.R.; Tripodi, V.P.; De’Nobili, M.D.; Fissore, E.N. Cherry (Prunus avium) phenolic compounds for antioxidant preservation at food interfaces. J. Food Eng.; 2018; 239, pp. 15-25. [DOI: https://dx.doi.org/10.1016/j.jfoodeng.2018.06.028]
131. Wong-Paz, J.E.; Muñiz-Márquez, D.B.; Aguilar-Zárate, P.; Ascacio-Valdés, J.A.; Cruz, K.; Reyes-Luna, C.; Rodríguez, R.; Aguilar, C.N. Extraction of Bioactive Phenolic Compounds by Alternative Technologies. Ingredients Extraction by Physicochemical Methods in Food; Academic Press: Cambridge, MA, USA, 2017; pp. 229-252.
132. Caldas, T.W.; Mazza, K.E.L.; Teles, A.S.C.; Mattos, G.N.; Brígida, A.I.S.; Conte-Junior, C.A.; Borguini, R.G.; Godoy, R.L.O.; Cabral, L.M.C.; Tonon, R.V. Phenolic compounds recovery from grape skin using conventional and non-conventional extraction methods. Ind. Crops Prod.; 2018; 111, pp. 86-91. [DOI: https://dx.doi.org/10.1016/j.indcrop.2017.10.012]
133. Alara, O.R.; Abdurahman, N.H. Microwave-assisted extraction of phenolics from Hibiscus sabdariffa calyces: Kinetic modelling and process intensification. Ind. Crops Prod.; 2019; 137, pp. 528-535. [DOI: https://dx.doi.org/10.1016/j.indcrop.2019.05.053]
134. Battistella Lasta, H.F.; Lentz, L.; Gonçalves Rodrigues, L.G.; Mezzomo, N.; Vitali, L.; Salvador Ferreira, S.R. Pressurized liquid extraction applied for the recovery of phenolic compounds from beetroot waste. Biocatal. Agric. Biotechnol.; 2019; 21, 101353. [DOI: https://dx.doi.org/10.1016/j.bcab.2019.101353]
135. Okiyama, D.C.G.; Soares, I.D.; Cuevas, M.S.; Crevelin, E.J.; Moraes, L.A.B.; Melo, M.P.; Oliveira, A.L.; Rodrigues, C.E.C. Pressurized liquid extraction of flavanols and alkaloids from cocoa bean shell using ethanol as solvent. Food Res. Int.; 2018; 114, pp. 20-29. [DOI: https://dx.doi.org/10.1016/j.foodres.2018.07.055]
136. Shang, Y.F.; Xu, J.L.; Lee, W.J.; Um, B.H. Antioxidative polyphenolics obtained from spent coffee grounds by pressurized liquid extraction. S. Afr. J. Bot.; 2017; 109, pp. 75-80. [DOI: https://dx.doi.org/10.1016/j.sajb.2016.12.011]
137. Posadino, A.M.; Biosa, G.; Zayed, H.; Abou-Saleh, H.; Cossu, A.; Nasrallah, G.K.; Giordo, R.; Pagnozzi, D.; Porcu, M.C.; Pretti, L. et al. Protective Effect of Cyclically Pressurized Solid(-)Liquid Extraction Polyphenols from Cagnulari Grape Pomace on Oxidative Endothelial Cell Death. Molecules; 2018; 23, 2105. [DOI: https://dx.doi.org/10.3390/molecules23092105] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30134642]
138. Casazza, A.A.; Aliakbarian, B.; Mantegna, S.; Cravotto, G.; Perego, P. Extraction of phenolics from Vitis vinifera wastes using non-conventional techniques. J. Food Eng.; 2010; 100, pp. 50-55. [DOI: https://dx.doi.org/10.1016/j.jfoodeng.2010.03.026]
139. Hubert, P.; Vitzthum, O.G. Fluid Extraction of Hops, Spices, and Tobacco with Supercritical Gases. Angew. Chem.; 1978; 17, pp. 710-715. [DOI: https://dx.doi.org/10.1002/anie.197807101]
140. Reverchon, E.; De Marco, I. Supercritical fluid extraction and fractionation of natural matter. J. Supercrit. Fluids; 2006; 38, pp. 146-166. [DOI: https://dx.doi.org/10.1016/j.supflu.2006.03.020]
141. Herrero, M.; Cifuentes, A.; Ibanez, E. Sub- and supercritical fluid extraction of functional ingredients from different natural sources: Plants, food-by-products, algae and microalgae: A review. Food Chem.; 2006; 98, pp. 136-148. [DOI: https://dx.doi.org/10.1016/j.foodchem.2005.05.058]
142. Reverchon, E. Supercritical fluid extraction and fractionation of essential oils and related products. J. Supercrit. Fluids; 1997; 10, pp. 1-37. [DOI: https://dx.doi.org/10.1016/S0896-8446(97)00014-4]
143. Lang, Q.; Wai, C.M. Supercritical fluid extraction in herbal and natural product studies—A practical review. Talanta; 2001; 53, pp. 771-782. [DOI: https://dx.doi.org/10.1016/S0039-9140(00)00557-9]
144. Pourmortazavi, S.M.; Hajimirsadeghi, S.S. Supercritical fluid extraction in plant essential and volatile oil analysis. J. Chromatogr. A; 2007; 1163, pp. 2-24. [DOI: https://dx.doi.org/10.1016/j.chroma.2007.06.021] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17624357]
145. Romano, R.; Aiello, A.; Pizzolongo, F.; Rispoli, A.; De Luca, L.; Masi, P. Characterisation of oleoresins extracted from tomato waste by liquid and supercritical carbon dioxide. Int. J. Food Sci. Technol.; 2020; 55, pp. 3334-3342. [DOI: https://dx.doi.org/10.1111/ijfs.14597]
146. Kuvendziev, S.; Lisichkov, K.; Zeković, Z.; Marinkovski, M. Artificial neural network modelling of supercritical fluid CO2 extraction of polyunsaturated fatty acids from common carp (Cyprinus carpio L.) viscera. J. Supercrit. Fluids; 2014; 92, pp. 242-248. [DOI: https://dx.doi.org/10.1016/j.supflu.2014.06.007]
147. Obregón Tinoco, H.; Huayta Socantaype, F.; Cardenas Toro, F. Optimización del Proceso de Extracción por Fluidos Supercríticos en la Obtención de Aceite de Semillas de Uva con el Empleo de la Metodología Taguchi y Superficie de Respuesta. Proceedings of the 16th LACCEI International Multi-Conference for Engineering, Education, and Technology: “Innovation in Education and Inclusion; Lima, Peru, 18–20 July 2018; 268.
148. Galarce-Bustos, O.; Fernandez-Ponce, M.T.; Montes, A.; Pereyra, C.; Casas, L.; Mantell, C.; Aranda, M. Usage of supercritical fluid techniques to obtain bioactive alkaloid-rich extracts from cherimoya peel and leaves: Extract profiles and their correlation with antioxidant properties and acetylcholinesterase and alpha-glucosidase inhibitory activities. Food Funct.; 2020; 11, pp. 4224-4235. [DOI: https://dx.doi.org/10.1039/D0FO00342E] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32353090]
149. Santana, Á.L.; Albarelli, J.Q.; Santos, D.T.; Souza, R.; Machado, N.T.; Araújo, M.E.; Meireles, M.A.A. Kinetic behavior, mathematical modeling, and economic evaluation of extracts obtained by supercritical fluid extraction from defatted assaí waste. Food Bioprod. Process.; 2018; 107, pp. 25-35. [DOI: https://dx.doi.org/10.1016/j.fbp.2017.10.006]
150. Vági, E.; Simándi, B.; Vásárhelyiné, K.P.; Daood, H.; Kéry, Á.; Doleschall, F.; Nagy, B. Supercritical carbon dioxide extraction of carotenoids, tocopherols and sitosterols from industrial tomato by-products. J. Supercrit. Fluids; 2007; 40, pp. 218-226. [DOI: https://dx.doi.org/10.1016/j.supflu.2006.05.009]
151. Yu, I.K.M.; Attard, T.M.; Chen, S.S.; Tsang, D.C.W.; Hunt, A.J.; Jérôme, F.; Ok, Y.S.; Poon, C.S. Supercritical Carbon Dioxide Extraction of Value-Added Products and Thermochemical Synthesis of Platform Chemicals from Food Waste. ACS Sustain. Chem. Eng.; 2018; 7, pp. 2821-2829. [DOI: https://dx.doi.org/10.1021/acssuschemeng.8b06184]
152. Campalani, C.; Amadio, E.; Zanini, S.; Dall’Acqua, S.; Panozzo, M.; Ferrari, S.; De Nadai, G.; Francescato, S.; Selva, M.; Perosa, A. Supercritical CO2 as a green solvent for the circular economy: Extraction of fatty acids from fruit pomace. J. CO2 Util.; 2020; 41, 101259. [DOI: https://dx.doi.org/10.1016/j.jcou.2020.101259]
153. Rosa, A.; Era, B.; Masala, C.; Nieddu, M.; Scano, P.; Fais, A.; Porcedda, S.; Piras, A. Supercritical CO2 Extraction of Waste Citrus Seeds: Chemical Composition, Nutritional and Biological Properties of Edible Fixed Oils. Eur. J. Lipid Sci. Technol.; 2019; 121, 1800502. [DOI: https://dx.doi.org/10.1002/ejlt.201800502]
154. Castillo-Herrera, G.A.; Farías-Álvarez, L.J.; García-Fajardo, J.A.; Delgado-Saucedo, J.I.; Puebla-Pérez, A.M.; Lugo-Cervantes, E. Bioactive extracts of Citrus aurantifolia swingle seeds obtained by supercritical CO2 and organic solvents comparing its cytotoxic activity against L5178Y leukemia lymphoblasts. J. Supercrit. Fluids; 2015; 101, pp. 81-86. [DOI: https://dx.doi.org/10.1016/j.supflu.2015.02.026]
155. Moulehi, I.; Bourgou, S.; Ourghemmi, I.; Tounsi, M.S. Variety and ripening impact on phenolic composition and antioxidant activity of mandarin (Citrus reticulate Blanco) and bitter orange (Citrus aurantium L.) seeds extracts. Ind. Crops Prod.; 2012; 39, pp. 74-80. [DOI: https://dx.doi.org/10.1016/j.indcrop.2012.02.013]
156. Dorado Achicanoy, D.; Hurtado Benavides, A.; Martinez-Correa, H.A. Study of supercritical CO2 extraction of tamarillo (Cyphomandra Betacea) seed oil containing high added value compounds. Electrophoresis; 2018; 39, pp. 1917-1925. [DOI: https://dx.doi.org/10.1002/elps.201700430]
157. Salgın, U.; Salgın, S.; Ekici, D.D.; UludaĿ, G. Oil recovery in rosehip seeds from food plant waste products using supercritical CO2 extraction. J. Supercrit. Fluids; 2016; 118, pp. 194-202. [DOI: https://dx.doi.org/10.1016/j.supflu.2016.08.011]
158. Larrauri, J.A.; Rupérez, P.; Borroto, B.; Saura-Calixto, F. Mango Peels as a New Tropical Fibre: Preparation and Characterization. Chem. Lwt-Food Sci. Technol.; 1996; 29, pp. 729-733. [DOI: https://dx.doi.org/10.1006/fstl.1996.0113]
159. Louaer, M.; Zermane, A.; Larkeche, O.; Meniai, A.H. Experimental study and optimization of the extraction of Algerian date stones oil (Phoenix dactylifera L.) using supercritical carbon dioxide. J. Food Process Eng.; 2019; 42, 13049. [DOI: https://dx.doi.org/10.1111/jfpe.13049]
160. Kayathi, A.; Chakrabarti, P.P.; Bonfim-Rocha, L.; Cardozo-Filho, L.; Jegatheesan, V. Selective extraction of polar lipids of mango kernel using Supercritical Carbon dioxide (SC-CO2) extraction: Process optimization of extract yield/phosphorous content and economic evaluation. Chemosphere; 2020; 260, 127639. [DOI: https://dx.doi.org/10.1016/j.chemosphere.2020.127639] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32758767]
161. Murthy, P.S.; Madhava Naidu, M. Sustainable management of coffee industry by-products and value addition—A review. Resour. Conserv. Recycl.; 2012; 66, pp. 45-58. [DOI: https://dx.doi.org/10.1016/j.resconrec.2012.06.005]
162. Ahangari, B.; Sargolzaei, J. Extraction of lipids from spent coffee grounds using organic solvents and supercritical carbon dioxide. J. Food Process. Preserv.; 2013; 37, pp. 1014-1021. [DOI: https://dx.doi.org/10.1111/j.1745-4549.2012.00757.x]
163. Nasti, R.; Galeazzi, A.; Marzorati, S.; Zaccheria, F.; Ravasio, N.; Bozzano, G.L.; Manenti, F.; Verotta, L. Valorisation of Coffee Roasting By-Products: Recovery of Silverskin Fat By Supercritical CO2 Extraction. Waste Biomass Valorization; 2021; 12, pp. 6021-6033. [DOI: https://dx.doi.org/10.1007/s12649-021-01435-9]
164. Ferdosh, S.; Sarker, M.Z.I.; Abd Rahman, N.N.N.; Selamat, J.; Karim, M.R.; Razak, T.A.; Abd Kadir, M.O. Fish Oil Recovery from Viscera of Indian Mackerel (Rastrelliger kanagurta) by Supercritical Fluid: An Optimization Approach. J. Chin. Chem. Soc.; 2012; 59, pp. 1421-1429. [DOI: https://dx.doi.org/10.1002/jccs.201100235]
165. Franklin, E.C.; Haq, M.; Roy, V.C.; Park, J.S.; Chun, B.S. Supercritical CO2 extraction and quality comparison of lipids from Yellowtail fish (Seriola quinqueradiata) waste in different conditions. J. Food Process. Preserv.; 2020; 44, e14892. [DOI: https://dx.doi.org/10.1111/jfpp.14892]
166. Rao, A.V.; Rao, L.G. Carotenoids and human health. Pharmacol. Res.; 2007; 55, pp. 207-216. [DOI: https://dx.doi.org/10.1016/j.phrs.2007.01.012]
167. de Andrade Lima, M.; Kestekoglou, I.; Charalampopoulos, D.; Chatzifragkou, A. Supercritical Fluid Extraction of Carotenoids from Vegetable Waste Matrices. Molecules; 2019; 24, 466. [DOI: https://dx.doi.org/10.3390/molecules24030466]
168. Baysal, T.; Ersus, S.; Starmans, D.A.J. Supercritical CO2 Extraction of â-Carotene and Lycopene from Tomato Paste Waste. J. Agric. Food Chem.; 2000; 48, pp. 5507-5511. [DOI: https://dx.doi.org/10.1021/jf000311t] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11087510]
169. Sabio, E.; Lozano, M.; de Espinosa, V.M.; Mendes, R.L.; Pereira, A.P.; Palavra, A.F.; Coelho, J.A. Lycopene and â-Carotene Extraction from Tomato Processing Waste Using Supercritical CO2. Ind. Eng. Chem. Res.; 2003; 42, pp. 6641-6646. [DOI: https://dx.doi.org/10.1021/ie0301233]
170. Longo, C.; Leo, L.; Leone, A. Carotenoids, fatty acid composition and heat stability of supercritical carbon dioxide-extracted-oleoresins. Int. J. Mol. Sci.; 2012; 13, pp. 4233-4254. [DOI: https://dx.doi.org/10.3390/ijms13044233] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22605975]
171. Derrien, M.; Aghabararnejad, M.; Gosselin, A.; Desjardins, Y.; Angers, P.; Boumghar, Y. Optimization of supercritical carbon dioxide extraction of lutein and chlorophyll from spinach by-products using response surface methodology. Lwt; 2018; 93, pp. 79-87. [DOI: https://dx.doi.org/10.1016/j.lwt.2018.03.016]
172. Quitain, A.T.; Kai, T.; Sasaki, M.; Goto, M. Supercritical carbon dioxide extraction of fucoxanthin from Undaria pinnatifida. J. Agric. Food Chem.; 2013; 61, pp. 5792-5797. [DOI: https://dx.doi.org/10.1021/jf400740p]
173. Bendif, H.; Adouni, K.; Miara, M.D.; Baranauskiene, R.; Kraujalis, P.; Venskutonis, P.R.; Nabavi, S.M.; Maggi, F. Essential oils (EOs), pressurized liquid extracts (PLE) and carbon dioxide supercritical fluid extracts (SFE-CO2) from Algerian Thymus munbyanus as valuable sources of antioxidants to be used on an industrial level. Food Chem.; 2018; 260, pp. 289-298. [DOI: https://dx.doi.org/10.1016/j.foodchem.2018.03.108]
174. Gil-Ramírez, A.; Aldars-García, L.; Palanisamy, M.; Jiverdeanu, R.M.; Ruiz-Rodríguez, A.; Marín, F.R.; Reglero, G.; Soler-Rivas, C. Sterol enriched fractions obtained from Agaricus bisporus fruiting bodies and by-products by compressed fluid technologies (PLE and SFE). Innov. Food Sci. Emerg. Technol.; 2013; 18, pp. 101-107. [DOI: https://dx.doi.org/10.1016/j.ifset.2013.01.007]
175. Quispe-Fuentes, I.; Uribe, E.; López, J.; Contreras, D.; Poblete, J. A study of dried mandarin (Clementina orogrande) peel applying supercritical carbon dioxide using co-solvent: Influence on oil extraction, phenolic compounds, and antioxidant activity. J. Food Process. Preserv.; 2021; 46, e16116. [DOI: https://dx.doi.org/10.1111/jfpp.16116]
176. Castro-Vargas, H.I.; Baumann, W.; Ferreira, S.R.S.; Parada-Alfonso, F. Valorization of papaya (Carica papaya L.) agroindustrial waste through the recovery of phenolic antioxidants by supercritical fluid extraction. J. Food Sci. Technol.; 2019; 56, pp. 3055-3066. [DOI: https://dx.doi.org/10.1007/s13197-019-03795-6]
177. Romano, R.; Aiello, A.; Meca, G.; De Luca, L.; Pizzolongo, F.; Masi, P. Recovery of bioactive compounds from walnut (Juglans regia L.) green husk by supercritical carbon dioxide extraction. Int. J. Food Sci. Technol.; 2021; 56, pp. 4658-4668. [DOI: https://dx.doi.org/10.1111/ijfs.15161]
178. Caballero, A.S.; Romero-García, J.M.; Castro, E.; Cardona, C.A. Supercritical fluid extraction for enhancing polyphenolic compounds production from olive waste extracts. J. Chem. Technol. Biotechnol.; 2019; 95, pp. 356-362. [DOI: https://dx.doi.org/10.1002/jctb.5907]
179. Jafarian Asl, P.; Niazmand, R.; Yahyavi, F. Extraction of phytosterols and tocopherols from rapeseed oil waste by supercritical CO2 plus co-solvent: A comparison with conventional solvent extraction. Heliyon; 2020; 6, e03592. [DOI: https://dx.doi.org/10.1016/j.heliyon.2020.e03592] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32258458]
180. Martins, C.M.; Guedes, J.A.C.; de Brito, E.S.; Ferreira, S.R.S. Valorization of tamarind seeds using high-pressure extraction methods to obtain rich fractions in fatty acid and phenolic compounds. J. Supercrit. Fluids; 2022; 183, 105556. [DOI: https://dx.doi.org/10.1016/j.supflu.2022.105556]
181. Viganó, J.; Zabot, G.L.; Martínez, J. Supercritical fluid and pressurized liquid extractions of phytonutrients from passion fruit by-products: Economic evaluation of sequential multi-stage and single-stage processes. J. Supercrit. Fluids; 2017; 122, pp. 88-98. [DOI: https://dx.doi.org/10.1016/j.supflu.2016.12.006]
182. Tamkutė, L.; Liepuoniūtė, R.; Pukalskienė, M.; Venskutonis, P.R. Recovery of valuable lipophilic and polyphenolic fractions from cranberry pomace by consecutive supercritical CO2 and pressurized liquid extraction. J. Supercrit. Fluids; 2020; 159, 104755. [DOI: https://dx.doi.org/10.1016/j.supflu.2020.104755]
183. Mazzutti, S.; Rodrigues, L.G.G.; Mezzomo, N.; Venturi, V.; Ferreira, S.R.S. Integrated green-based processes using supercritical CO2 and pressurized ethanol applied to recover antioxidant compouds from cocoa (Theobroma cacao) bean hulls. J. Supercrit. Fluids; 2018; 135, pp. 52-59. [DOI: https://dx.doi.org/10.1016/j.supflu.2017.12.039]
184. Kraujalienė, V.; Pukalskas, A.; Venskutonis, P.R. Biorefining of goldenrod (Solidago virgaurea L.) leaves by supercritical fluid and pressurized liquid extraction and evaluation of antioxidant properties and main phytochemicals in the fractions and plant material. J. Funct. Foods; 2017; 37, pp. 200-208. [DOI: https://dx.doi.org/10.1016/j.jff.2017.07.049]
185. Villanueva Bermejo, D.; Ibáñez, E.; Reglero, G.; Turner, C.; Fornari, T.; Rodriguez-Meizoso, I. High catechins/low caffeine powder from green tea leaves by pressurized liquid extraction and supercritical antisolvent precipitation. Sep. Purif. Technol.; 2015; 148, pp. 49-56. [DOI: https://dx.doi.org/10.1016/j.seppur.2015.04.037]
186. Ferreira, W.S.; Viganó, J.; Veggi, P.C. Economic assessment of emerging extraction techniques: Hybridization of high-pressure extraction and low-frequency ultrasound to produce piceatannol-rich extract from passion fruit bagasse. Chem. Eng. Process.-Process Intensif.; 2022; 174, 108850. [DOI: https://dx.doi.org/10.1016/j.cep.2022.108850]
187. Trucillo, P.; Campardelli, R.; De Marco, I. Environmental and Sustainability Analysis of a Supercritical Carbon Dioxide-Assisted Process for Pharmaceutical Applications. Processes; 2021; 9, 1788. [DOI: https://dx.doi.org/10.3390/pr9101788]
© 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/). Notwithstanding the ProQuest Terms and Conditions, you may use this content in accordance with the terms of the License.
Details
; Pettinato, Margherita 1
; Campardelli, Roberta 1
; De Marco, Iolanda 2
; Perego, Patrizia 1
1 Department of Civil, Chemical and Environmental Engineering (DICCA), University of Genoa, Via Opera Pia 15, 16145 Genova, GE, Italy;
2 Department of Industrial Engineering, University of Salerno, Via Giovanni Paolo II, 132, 84084 Fisciano, SA, Italy