Abstract
The agriculture and horticulture industries generate many by-products while processing commodities, leading to significant environmental and societal dilemmas. Bioactive compounds obtained from by-products of plants are also known as ingredients in traditional medicines and the food industry due to their potential health benefits for humans as antioxidant, anti-inflammatory, and antimicrobial agents. On the other hand, an extensive array of colorants from natural sources has been scrutinized recently in different sectors to mitigate the negative ecological impacts of synthetic dyes, especially in modern textile manufacturing. Approaching these matters, this paper reviews various chemical compositions of plant secondary metabolites, including flavonoids, tannins, phenolic acids, essential oils, tulipalins, and factors that influence the isolation process. Moreover, the article offers an updated view of the advanced technologies for extraction, focusing on eco-friendly solvents such as water, supercritical carbon dioxide, and deep eutectic solvents. Eventually, a comparison of different extraction methods is highlighted to devise the most appropriate strategy for industries.
Graphical abstract
![](http://media.springernature.com/lw685/springer-static/image/art%3A10.1007%2Fs13399-023-04726-4/MediaObjects/13399_2023_4726_Figa_HTML.png)
Similar content being viewed by others
Explore related subjects
Find the latest articles, discoveries, and news in related topics.Avoid common mistakes on your manuscript.
1 Introduction
With a history spanning millennia, agriculture, including the horticulture industry, has undergone notable development and stands as one of the most paramount fields of human knowledge. Additionally, it has a vital role to support and develop other industries such as cosmetics, pharmaceuticals, and nutraceuticals. Correspondingly, the agriculture industry produces a significant number of residues during the agricultural production process, which can be known as by-products [1]. As a solution, the concept inception of circular bioeconomy is applied to reduce the losses induced by food waste disposal, comprising agricultural by-products. The key point of circular bioeconomy is to diminish environmental—societal and economic—costs, economic competitiveness intensification, and to mitigate poverty and hunger. Moreover, the circular bioeconomy focuses on “waste-to-wealth” concept that creates new technologies, jobs, and livelihoods [2]. Therefore, by-products valorization from plants is one of the crucial biomass to decrease the pollution risks for the environment and, significantly, to enable sustainable development and a generally circular bioeconomy [1, 3]. Those production by-products are traditionally treated as wastes and are typically landfilled or burned, resulting in an observable production of carbon dioxide (CO2) [4]. In this regard, by-products, consisting of roots, flowers, seeds, leaves, buds, fruits, etc. from agriculture and horticulture, are attractive options because of their low cost and biodegradability, besides featuring essential properties [5]. Originating from plants, secondary metabolites are chemical substances that gain diverse beneficial health effects in humans and animals. Some of them are performed as defense compounds against diseases, predators, ultraviolet radiation, parasites, and oxidants, to facilitate the reproductive processes (for example, attractive fragrance and coloring agents) [6]. Bioactive compounds are referred to as secondary plant metabolites that present as natural constituents in plants, including food, which contribute health benefits beyond the fundamental nutrition value of the plants. Some of the bioactive compounds are hydroxybenzoic acids, hydroxycinnamic acids, carotenoids, flavonoids, stilbenes, and lignans [6,7,8]. Several notable attributes of bioactive compounds are antimicrobial, antioxidant, antiviral, anticancer, and antiradical activities [9]. Other applications of bioactive compounds from plants are providing colors for food [10], dyes for textiles [11, 12], dye-sensitized solar cells [13], and fragrance [14].
Excluding the benefits of minimizing the volume of crops residues, the efficient use of by-product resources is proposed to reduce the production costs of natural antimicrobial agents and colorants [15]. Beyond the goal of employing natural compounds and reducing by-product quantities, 92 million tons of textile waste were produced in 2015, and the volume is expected to increase by 62% by 2030 [16]. Many papers have therefore studied the current state of sustainability innovation concerning textile industry. Specifically, researchers have employed frameworks to find novel approaches to remodeling traditional textile methods and processes, which incorporate the basic principles of the circular economy in the textile industry [17,18,19]. These ideas indicate preventive measures to mitigate emissions and effluents from the textile chains by substituting some synthetic compounds with organic natural materials such as bamboo fibers, natural dyes, and less toxic chemical products [20].
Awarding these matters, extracting bioactive compounds from agriculture and horticulture by-products is an essential topic. Particularly, the bioactive substances can be served as natural antimicrobial agents and colorants for the textile industry. To enhance the contribution to the circular economy, innovative technologies including using green solvents will be focused to investigate the possibility of isolating bioactive compounds sustainably. This paper critically considers floral waste such as roses and tulips, vegetable by-products such as tomato and cucumber crop residues, and garden waste like fall foliage. It also conducts and discusses a general outlook, including an order of magnitude estimate of the global potential of horticultural and vegetable production streams. This review paper also points to the bottlenecks that could hinder the implementation of advanced technologies, in which the parameter extraction, solvents used, superiorities, and drawbacks of various isolation methods are comprehensively studied.
2 A general outlook on agricultural and horticultural production streams
This paper analyzes the phenolic-based agricultural by-products in the context of global horticulture and vegetable production. The world’s largest ornamental markets in 2019 and fresh vegetable producers in 2020 are shown in Figs. 1 and 2 respectively. According to the vision of the Association Internationale des Producteurs de l’Horticulture (AIPH) in 2019 [21], China, Japan, and North America (the USA and Canada) were the major mature producer countries for both the production and consumption of the horticultural industry. From a global perspective, Europe will be one of the world’s most prominent actors in horticultural production and consumption growth over the next decade, following the lead of major markets like China, Japan, and North American countries [23]. The global horticultural market was approximately USD 70 billion in 2017 and is expected to reach 100 billion USD by 2027. The Netherlands has been the international flower market leader for years, with a 60% market share. However, it is projected to decline to 35% in 2027, with continued market growth in absolute terms. The market growth decline of the Netherlands is related to the sharp forecast growth rate in Asia at 80%, and in Europe and North America at a relatively modest growth rate of 20% by 2027 [24].
Major producers of ornamentals in the world in 2019 (adapted from [21])
Global production of fresh vegetables in 2020; (a) by region and (b) worldwide (adapted from [22])
Among flowers, roses are still the absolute number one in cut flower production, where red-colored roses represent 30% of the market. About 10 billion rose stems are sold annually worldwide. Otherwise, potted roses (80 million potted plants per year) and garden roses (220 million plants per year) are also important. In the economic overview of roses, the largest rose production areas are India (61,000 hectares) and China (14,348 hectares) [21, 25]. In the Netherlands, rose production has reduced rapidly, decreasing from 850 to 238 hectares in 2016. The reduction in rose production in the Netherlands is more rapid than the overall decrease in total cut flowers [25]. Although the strongest rose production sites are outside Europe, the production of cut roses still shows promise in some countries such as the southern parts of Germany, France, and Italy because of their suitable conditions for rose cultivation [26].
Moreover, the world’s fresh vegetable production (Fig. 3) promptly increased between 2000 and 2019. In Fig. 3, the five main vegetable types accounted for 42–45% of the total amount during the period: tomatoes (16% in 2019); dry onions (9%); cucumbers and gherkins (8%); cabbages (6%); and eggplants (5%). Generally, until 2020, Asia, particularly China and India, where tomatoes, onions, and cucumbers accounted for a considerable volume, produced the largest amount of vegetables [27]. The production volume expansion in recent years therefore boosts biomass availability, opening new opportunities for recovering and expanding the bioproducts field [28], especially its capacity as a good source of phenolic compounds [29].
Fresh vegetable production globally from 2000 to 2019 (adapted from [27])
3 Phytochemical compounds in plants
The main classes of polyphenols in plants include flavonoids, phenolic acids, tannins, stilbenes, and lignans. These polyphenolic compounds are presented in Fig. 4, and some chemical structures are illustrated in Fig. 5. Flavonoids and phenolic acids account for 60% and 30% of the total dietary polyphenols respectively [30]. Overall, phenolics are primarily found in fruit and vegetables, including horticultural and ornamental crops, teas, and cereals. They are often associated with mono- or polysaccharides and functional derivatives such as esters or methyl esters. Phenolics also possess various hydroxylation patterns, and they can appear in acylated and glycosylated forms [31].
The abundant phenolic compounds in some agricultural and horticultural by-products are exhibited in Table 1. Quercetin, phenolic acids, catechins, and tannins are the main compounds in rose plants’ leaves, buds, and flowers [32,33,34,35]. Other chemical constituents include flavonoids, triterpenes, polysaccharides, fatty acids, organic acids, carotenoids, vitamins, and essential oils. Specifically, the phenolic compounds in rose leaves such as ellagitannins (73.69 mg/g of plant dry weight (DW)), gallotannins (21.23 mg/g of plant DW), and hydroxycinnamates (1.47 mg/g of plant DW) were found to be significant. The report also showed that petals accumulated anthocyanins (ANCs), flavonoids (flavonol and dihydroflavonols), and water-soluble polysaccharides, while root samples contained catechin and proanthocyanidins [34].
The fall foliage presents an attractive opportunity to obtain valuable polyphenolic compounds. Manninen et al. [36] studied the collected fall leaves to extract carotenoids and ANCs. The ANC content varied from 0 to 2.8 mg/g of plant DW, with an average of 0.6 mg/g, and carotenoid content ranged from 0 to 83 mg/g, with an average of 26 mg/g. Lisandru et al. [37] studied the extracts from cherry fall leaves. They observed that the carotenoid content was more dominant than the ANC content, and the ANC composition was inversely proportional to carotenoid yield.
From the perspective of agricultural by-products, Silva et al. [38] found phenolic compounds such as gallic acids, chlorogenic acid, rutin, caffeic acid, ferulic acid, and quercetin were abundant in this order in tomato plants comprising stems, leaves, and roots. Another study by Guuntekin et al. [39] reported that the number of phenolic compounds was highest (7.14%) in the extract of tomato stalks compared to other plants. For the most widely grown vegetables in Europe and the USA, around 33 kg of leaf and stem biomass is gathered for every 100 kg of tomato fruit harvested. Each tomato plant produces an average of 0.75 kg of leaf biomass, resulting in a total of 15 tons per hectare [40].
In addition, cucumber (Cucumis sativus L.) stems are unique in that they contain sphingolipid glycosides, which distinguishes them from other field residues [41,42,43]. An equivalent amount (36.6%) of stem and leaf biomass is found in cucumber plants grown in the greenhouse under supplemental light [44]. C. sativus L. leaf extracted with acetone contains alkaloids, glycosides, steroidal terpenes, flavonoids, saponins, reducing sugars, and tannins [45]. The identification of chemical compounds reveals phenols, flavonoids, carotenoids, alkaloids, glycosides, tannins, and terpenes in methanol (MeOH) and ethanol (EtOH) extract from cucumber leaves (C. sativus L.) [46]. Meanwhile, Mallik and Akhter [47] claimed that the EtOH extracts of cucumber leaves and stems showed the absence of flavonoids and reducing sugars, while the presence of alkaloids, glycosides, saponins, sterol, and tannins was found instead. Table 2 lists all the conditions, solvents used, methods, and extracted bioactive compounds from agricultural and horticultural by-products in detail.
3.1 Flavonoids
Flavonoids form one of the major subclasses of polyphenols, representing the most significant group of compounds among over one million metabolites in plants. They consist of two or more aromatic rings, and each ring possesses at least one hydroxyl group and is linked by a three-carbon bridge [52,53,54]. Flavonoids have the most comprehensive color range, from pale yellow to intense violet. Moreover, flavonoids protect against ultraviolet (UV) radiation due to their capacity to absorb the most energetic solar wavelengths, e.g., UV-B and UV-A. This property of flavonoids is the decisive factor for aquatic plants’ evolution and terrestrial existence [55, 56].
Flavonoids primarily have six major subclasses based on their complexity in the chemical structure: flavones, flavanols, isoflavones, flavonols, flavanones, and ANCs [57]. Among these subclasses, ANCs are studied more comprehensively than others, as they are attractive components of the targeted crop residues. ANC originates in flavonol; it has the basic structure of the flavylium ion. ANC is in the form of a glycoside, while an anthocyanidin is known as an aglycone. Anthocyanidins are grouped into 3-hydroxyanthocyanidin, 3-deoxyanthocyanidin, and O-methylated anthocyanidins, whereas ANCs are in the forms of anthocyanidin glycosides and acylated ANCs [58]. Flavonoids are well-known for the red, blue, and purple colors that ANCs produce [36, 58, 59].
ANCs are water-soluble and stored in cell vacuoles. The empirical formula for the flavylium ion of ANC is C15H11O+, with a molecular weight (MW) of 207.24 g/mol. ANCs, glycosylated ANCs, and proanthocyanidins (polymers that produce ANCs when hydrolyzed, or “condensed” tannins) contribute to the color of flowers and fruit, with a range from reddish purple to intense blue. The most common ANCs are cyanidin, pelargonidin, peonidin, petunidin, and malvidin. ANCs are present in higher plants, e.g., in juvenile leaves and senescing plants [58]. For example, in the fall foliage of sugar maple trees (Acer saccharum), the red expression is influenced by foliar nitrogen concentration. Leaves with lower concentrations of nitrogen tend to turn red earlier than those with higher concentrations [60].
3.2 Phenolic acids
Phenolic acids form one-third of phenolic compounds. They are carboxylic acids derived from either benzoic or cinnamic acid. They can be in free and conjugated, and soluble and insoluble states. Hydroxycinnamic acids have nine carbon atoms (C6–C3), but those found in vegetables have seven carbons. Typically, they have a benzene ring, a carboxylic group, and one or more hydroxyl and methoxy groups in the molecule [61]. In contrast, benzoic acids have seven carbon atoms (C6–C1), including gallic and protocatechuic acids [61, 62]. Hydroxycinnamic acids comprise 4-hydroxycinnamic acid, ferulic acid, caffeic acid, and sinapic acid. Hydroxybenzoic derivatives include 4-hydroxybenzoic acid, protocatechuic acid, vanillic acid, gallic acid, and syringic acid [62]. Hydroxybenzoic acids can be found in very low contents in edible plants, while hydroxycinnamic acids are one of the most abundant phenolic compounds. Hydroxycinnamic acids are present in coffee, medicine plants, fresh fruit, and vegetables in free form (rarely) or associated with polysaccharides in the cell wall [63, 64]. For example, ferulic acids are linked to polysaccharides (cellulose, hemicellulose, pectin, and glycoprotein), lignins, and suberin [65].
3.3 Tannins
Tannins are a heterogeneous group and complex phenolic compounds with MWs ranging from intermediate to very high (500–30,000 Dalton (Da)) [66]. Tannins obtain from 12 to 16 phenolic groups and 5 to 7 aromatic rings per 1000 Da. Because of the numerous hydroxyl groups in their structure, it has been demonstrated that tannins possess various biological activities, e.g., antioxidant and antitumor activities [67, 68]. Tannins can conjugate with proteins, carbohydrates, nucleic acid, and alkaloids, either by non-covalent bonds primarily driven by hydrogen bonding and hydrophobic interactions or by covalent binding after polyphenol activation [68, 69].
Tannins fall into two main classes: hydrolyzable tannins and condensed tannins. The other two classes of tannins are complex tannins and phlorotannins. Specifically, hydrolyzable tannins are subdivided into gallotannins and ellagitannins. Gallotannins produce gallic acids and their derivatives from hydrolysis, while ellagitannins produce ellagic acid after hydrolysis [70]. Moreover, condensed tannins have 3 to 8 repetition units and mainly originate in flavan-3-ol and flavan-3,4-diol structures [70, 71]. Hydrolyzable tannins can be found in the fruit, leaves, seeds, and bark of plants of the Leguminosae, Fabaceae, Combretaceae, and Anacardiaceae families [72]. On the other hand, condensed tannins represent 90% of total commercial tannins worldwide. Condensed tannins and their flavonoid precursors are widely distributed in nature, particularly in various Acacia (wattle or mimosa bark extract), Schinopsis (quebracho wood extract), Tsuga (hemlock bark extract), Pinus (bark extract), and Rhus (sumach extract) species [73].
3.4 Essential oils from flowers
The volatile organic compounds are called essential oils, with a molecular weight lower than 300 g/mol. These volatile compounds have a high vapor pressure at atmospheric pressure and room temperature. They belong to various chemical classes, e.g., alcohols, ethers or oxides, aldehydes, ketones, and esters [74]. However, the amounts and types of essential oil are determined by the plant parts e.g., leaves, buds, fruits, flowers, herbs, twigs, bark, wood, roots, and seeds. Another decisive factor is the plant species, as they are derived from terpenes and their oxygenated derivatives such as aromatic terpenoids, aliphatic acid esters, and phenolic compounds [75]. The average extracted oil content in flowers is 0.035% but varies from 0.03 to 0.05%. They typically have a yellow to yellow-green color and characteristic odor [76, 77].
3.5 Carotenoids in fall leaves
ANCs are not the only color-producing compounds present in fall leaves. Additionally, carotenoids contribute to the bright fall colors, as they absorb light in the range of 400–500 nm. Unlike ANCs, carotenoids are synthesized during the growing season to enable or protect photosynthetic light capture [37, 78]. Carotenoids are non-polar compounds belonging to a class of isoprenoid compounds. They are organic molecules of a C40 polyene backbone containing terminal ionone rings at both ends. The conjugated double bonds of carotenoids make them good light absorbers. The absorbed wavelengths depend on the number and nature of double bonds and other attached substituents. Carotenoids are classified into two types: (1) hydrocarbon carotenes, which include α-carotene, β-carotene, γ-carotene, and lycopene; (2) xanthophylls, which contain oxygen and therefore differ from carotenes. Xanthophylls are therefore more polar compounds than carotenes [15, 79,80,81,82]. Carotenoids occur in plants with 0.07–0.2% of the dry mass. They appear primarily as free carotenoids in leaves and as esterified carotenoids in other tissues. They are commonly found in higher plants’ flowers, fruits, leaves, and roots, e.g., carrots, peaches, tomatoes, paprikas, annatto, and pumpkins [15]. Most leaves contain similar carotenoid composition, while flowers possess individually characteristic carotenoid profiles, which rely on each plant species [79].
3.6 Tulipalin from tulip biomass
Tuliposides are the glucose esters of 4-hydroxy-2-methylenebutanoate and 3,4-dihydroxy-2-methylenebutanoate, the major secondary metabolites in tulips (Tulipa gesneriana). Tulipalins are tulipalin A and tulipalin B, which are the lactonized aglycons of tuliposides: 6-tuliposide A 6-O-(4′-hydroxy-2-methylenebutanoyl-D-glucopyranose and 6-tuliposide B 6-O-[(3S)-3,4–dihydroxy-2-methylidenebutanoyl]-D-glucopyranose [83, 84]. In nature, the 6-tuliposide A and B were found in tulip tissues such as petals, pistils, leaves, stems, bulb scales, and roots [85]. The tuliposides are unstable and are readily chemically converted into their corresponding lactone forms, tulipalin A and tulipalin B (Fig. 6) [84]. In general, tulipalins have the common structure α-methylene-γ-butyrolactone, of which tulipalin A is the simplest α-methylene-γ-butyrolactone first extracted from Erythronium americanum for its anticancer properties [49, 86].
4 Bioactive compounds as antimicrobial agents and colorants for textiles
Apart from being known as plant-derived colorants, bioactive components from plants are also acknowledged for their antimicrobial properties [87, 88]. In textiles, the growth of bacteria on fabrics causes undesirable effects such as unpleasant odors, skin infections, fabric stains, discoloration, and material deterioration. Hence, microbial activity prevention from plants becomes necessary to contribute bioproducts for textiles. For example, ANCs have not only benefited from antioxidant and anti-obesity effects, but they also possess antimicrobial activity against a wide range of microorganisms. ANCs extracted from blueberry, bilberry, raspberry, blackcurrant, and strawberry can inhibit the growth of gram-positive and gram-negative bacteria [58, 89,90,91]. Additionally, antimicrobial finishing in clothing can prevent the formation of foul odors due to bacterial processes in sweat. Colorful and antibacterial ANCs from Cinnamomum camphora waste (fall leaves) extract were considered a potential dyeing agent for wool fibers, valorizing their application in biomedical textiles [92].
Food, cosmetics, and textile industries also use microbial carotenoids for coloration purposes. In textile production, carotenoids are investigated as potent antimicrobial agents [78]. Carotenoids were extracted from annatto by different solvents, and their microbial inhibition activities were observed against Escherichia coli, Klebsiella pneumoniae, Pseudomonas aeruginosa, Bacillus subtilis, and Staphylococcus aureus. The leaf extracts displayed high antibacterial activity even in low concentrations, while the fruit extracts obtained a similar effect in higher concentrations [93].
Tannins have an antibacterial effect, which is explained by the ability to pass through the bacterial cell wall up to the internal membrane, interfere with cell metabolism, and destroy the cell [94]. The abundance of tannins in various natural dyes is reported to contribute strong antibacterial effects against common pathogens [95, 96]. Gupta et al. [97] studied the antibacterial effect of tannin-containing natural dyes against bacteria. Besides E. coli and K. pneumonia, Proteus vulgaris is known to convert urea to ammonia, resulting in unpleasant odors. The study claimed that most of the tested substances were more effective than the commercial antimicrobial Fabshield, as 99% of these bacterial colonies were reduced. In another article, Zhang et al. [98] discovered that tannic acid–treated silk also exhibited excellent antibacterial properties, with an inhibition rate of 99%. Tannin with a metal salt is therefore suggested for improving the antibacterial properties of hospital textiles. It could also be applicable in sports or domestic clothing to avoid unpleasant odors [99].
Essential oils also act as antimicrobial compounds and alternatives for treating infectious diseases, e.g., a disease caused by Herpes simplex virus [100, 101]. By inducing disturbances in membrane-embedded proteins, hindering respiration, and altering ion transportation processes, essential oils exhibit significant microbial prevention properties that ultimately cause bacterial death [102]. Sufficient in vitro data also proved that essential oils could be utilized as antibacterial agents to inhibit a wide range of bacterial strains, comprising Listeria monocytogenes, Salmonella typhimurium, E. coli O157:H7, Shigella dysenteriae, S. aureus, etc. [103, 104]. Walentowska and Foksowicz-Flaczyk [105] successfully applied essential oil from thyme. They evaluated its potential to restrain bacterial growth in linen and cotton fabric. The results showed that using the essential oil of thyme as a finishing agent could highly impede the growth of bacteria for linen-cotton type fiber blends.
For tulipalins from tulip biomass, there are seldom studies that use tulipalins as antimicrobial agents for textiles, despite tulipalins and tuliposides showing strong antimicrobial effects [106]. For example, tulipalins can resist the growth of cancer cells [86], and some gram-positive and gram-negative bacteria [83] such as methicillin-resistant S. aureus [107].
5 Extraction methods
The extraction of antimicrobials from natural sources is one of the most essential steps before the treatment of textile to achieve the target antimicrobial activity. In general, the conventional extraction techniques are maceration, Soxhlet extractions, and hydro-distillation [108]. Maceration is an inexpensive and straightforward technique for obtaining essential oil and bioactive compounds. In this method, the material is soaked in a container by immersing the sample in the solvent with or without stirring. This method consumes extensive solvents and time, and the extraction yield is low. Solvent and time consumption can be reduced by using Soxhlet extraction; yet in Soxhlet extraction (automatic continuous extraction), thermo-labile compounds may be degraded during the procedure [109, 110]. On the other hand, hydro-distillation is a traditional technique for extracting essential oils from plants. Organic solvents are not involved in this process and can be performed before dehydration. There are three types of hydro-distillation: water, water and steam, and direct steam distillation. It has three steps: hydro diffusion, hydrolysis, and decomposition by heat. However, volatile components in plants may be lost when high extraction temperatures are employed, which limits their use for thermo-labile compound extraction [108].
Traditionally, solvent extraction is the common method for isolating bioactive compounds, which applies the mechanisms of a solid matrix [111]. The diffusivity and solubility of bioactive compounds from raw materials promote the extraction process [110]. A solvent is chosen for its selectivity and the ability to extract phenolic compounds (Table 3). The solvent affects the constituent elements in plant cells through a chemical or biophysical alteration. For example, MeOH is used to extract lower MW polyphenols. Alternatively, higher MW polyphenols are extracted with aqueous acetone. Water is a suitable solvent for extracting phenolic acids and their glucosides. It provides higher extraction yields than organic solvents such as MeOH, EtOH, or butanol, particularly under ultrasonic treatment [112, 113]. Moreover, flavonols such as quercetin show high solubility in alcohols, as higher isolation yields are obtained with an increasing percentage of EtOH up to 70% v/v [114]. A few common organic solvents can be listed, from least polar to most polar, such as hexane < chloroform < ethyl acetate < acetone < EtOH < MeOH < water [115, 116]. In addition to the organic solvents, some reagents or salts are utilized industrially to isolate bioactive compounds. As an example of tannin extraction, tannin solution in water generally results in an acidic environment; it forms insoluble precipitates due to the tannin self-condensation reaction. On an industrial scale, reagents such as sodium sulfite (Na2SO3), sodium carbonate (Na2CO3), sodium bisulfite (NaHSO3), EtOH, or sodium hydroxide (NaOH) are therefore normally used [71, 117]. The addition of alkali will increase the pH and the incorporation of EtOH will improve extraction efficiency [118]. Similarly, the addition of salts such as Na2SO3 and Na2CO3 increases the extraction yield of tannins and reduces the viscosity of the extracts derived from the tannin self-condensation reactions [119].
Nevertheless, the observed drawbacks of conventional extraction processes are, e.g., longer extraction times, expensive and high-purity solvents, an excessive amount of energy and solvent used, low extraction selectivity, and poor-quality extracts [108, 120]. Various novel technologies have therefore been investigated to overcome these challenges. The non-conventional extraction techniques use green solvents, including pressurized fluids, deep eutectic solvents (DESs), bio-based solvents, some auxiliaries such as microwave, pulse electric field, and ultrasound [112, 121, 122].
5.1 Solvent extraction
Besides techniques based on the use of compressed fluids as extracting agents and methods with advanced auxiliaries, finding appropriate solvents having low toxicity and diminished environmental effects, elevated extraction ability, and biodegradability is important [123]. Ionic liquids (ILs) are known as novel solvents for extracting phenols besides other green solvents such as hot water or conventional organic liquids. ILs are defined as salts that present as liquids at room temperature and have melting points or glass transition temperatures below 100 °C. ILs are better solvents than water, as they have greater solubility power. Depending on their unique chemical properties, ILs gain many merits in destroying the cell walls and capturing the targets by chemical selectivity, thus, improving the efficiency of the extract in a short time. On the other hand, ILs gain some major disadvantages such as a high cost and poor solvent recovery [123,124,125,126]. ILs are classified based on their physical properties, which are named as neutral ILs, acidic ILs, basic ILs, ILs with amphoteric anions, functionalized ILs, protic ILs, chiral ILs, supported ILs, bio-ILs, poly-ILs, and energetic ILs [127]. Because of a variety of ions combination, ILs can be acknowledged as “designer solvents” that alter their physical properties to tackle the challenges of common volatile organic solvents [128]. In solid–liquid extraction, ILs were used to extract flavonoids from Larix decidua tree bark, in which [C4C1im]Br (25% wt) was evaluated to be the best solvent. The results from the antioxidant activity of extracts, energy-saving process, and flavonoid extractability were very positive, confirming their potential in different applications [129]. Sillero et al. [130] developed a new procedure to extract rutin from the leaves of A. theophrasti using IL [C4mim]Br based on ultrasonic- and microwave-assisted extraction. The isolated rutin from that method yielded 5.49 mg/g, increasing 2.01 folds than that of heating reflux extraction with MeOH.
Since 2003, another advanced selection in solvent extraction has been the substitution of traditional solvents for DESs, a novel technology still in development. Principally, DES is usually known as a subclass of ILs because of many similar characteristics, such as low volatility, low flammability, and broad electrochemical and thermal stability windows. However, these materials are considered inexpensive to synthesize and can be produced by biodegradable and non-toxic constituents [131]. DESs contain large and nonsymmetric ions that possess lattice energy and thus have low melting points [134]. Also, DESs can be characterized by a very large depressing freezing point. They are liquid at temperatures lower than 150 °C, and most of them are liquid between room temperature and 70 °C [135]. In general, DESs are obtained by the complexation of a non-toxic quaternary ammonium salt with a metal salt or a naturally derived uncharged hydrogen-bond donor (organic acids, polyalcohols, sugars, amino acids) in a particular molar ratio [121, 123]. At the optimum component ratio, the mixture obtains the deepest eutectic point. When the temperature exceeds the eutectic point, the mixture will be liquid. Otherwise, it will be solid [129]. For example, ammonium salt mixed with urea in the ratio of 2:1 (urea:salt) leads to the freezing point from –38 to 113 °C for the corresponding DES. In addition, the anion of choline-derived salts also influences the freezing points of DESs. The freezing point of DES made of choline-derived salts and urea decreases in the order: F– > NO3– > Cl– > BF4–, proposing a correlation with the hydrogen bonding strength [135]. Because of the affordable prices, biodegradability, pharmaceutically acceptable toxicity, and sustainability, DESs gain significant attraction as alternatives to conventional solvents to extract phenolic compounds [136,137,138,139], including bioactive flavonoids [129,130,131,132,133]. As an example, rutin was extracted from flower buds of Sophora japonica by choline chloride–based DESs. The solvents showed extremely low toxicity and substantial biodegradability [140]. In a different paper, choline chloride–betaine hydrochloride–ethylene glycol formed DES with the mole ratio of 1:1:2, combined 20% v/v water was proposed to extract and separate flavonoids. The study indicated that DESs could be used as green solvents for the extraction of active components in plant matrices [141]. Despite there are many possible combinations between quaternary ammonium salt and hydrogen-bond donors to make DESs, the predictive comprehension of structure–property relationships of DESs is ambiguous. For instance, tunning high viscosity of DESs is not approached entirely; thus, it hinders applications of DESs in many technologies [131]. As a consequence, to reduce the viscosity, increase the mass transfer, and improve the extraction efficiency, water is often used to control the polarity of DESs and the dissolution of the target compounds [142, 143].
Bio-based solvents can be an alternative besides ILs and DESs, which are produced by renewable sources. The raw materials that are considered feedstocks in biorefinery are usually: agricultural, such as waste from production; forestry, and wood processing; processing waste from other industries, and biomass of marine origin, including algae [144]. The manufacture of bio-based solvents consists of the fermentation of vegetable oils, carbohydrates, and the steam distillation of wood [145]. The most common bio-based solvents are 2-methyl tetrahydrofuran, glycerol, ethanol, γ-valerolactone, p-cymene, and D-limonene. Those solvents are produced from feedstocks such as cellulose, hemicellulose, lignin, and starch [123, 146]. Although they originate from renewable feedstocks, it is essential to mention the energy and solvent consumptions, high cost, and waste generation possibilities. Those factors might influence the valorization of those green solvents [147]. EtOH, ethyl acetate, ethyl lactate, and D-limonene were selected to recover fucoxanthin (an oxygenated carotenoid) from P. tricornutum under pressurized conditions. Among them, D-limonene was the most selective bio-based solvent, although it was not able to extract all fucoxanthin present in biomass unless supercritical fluid under continuous conditions was added [148]. Santos and Martins [149] evaluated the influences of bio-based solvents EtOH, acetic acid, and water on the content of ANC, flavonoids, and phenolics from onions (Allium cepa). The extracts from inions exhibited great antioxidant activities in ABTS, DPPH, and FRAP assays, demonstrating efficient antimicrobial activity against gram-positive bacteria. Especially, bioactive compounds isolated by EtOH 60% v/v showed the best performance that phenol, ANC, and flavonoid content were 3.4, 2.5, and 4.45 times higher than the other solvents, respectively.
5.2 Supercritical fluid extraction
A supercritical fluid (SF) is a liquid with a temperature and pressure above its critical point. SF can diffuse like gas through the solid matrix and dissolve materials like a liquid [150]. SF behaves as a single phase and retains the properties of both gas and liquid. Because SF lacks a phase boundary between gas and liquid, it has no surface tension either. The most significant advantage of SF is its tunability. Close to the critical point, even small changes in temperature and pressure have drastic effects on its density. SF can be adjusted to a more gas-like or liquid-like mobile phase, depending on the material’s solubility [151]. The selection of SF is determined by factors such as safety, hazards, energy requirements, and operability [152].
Among SFs, compressed CO2 fluid is used to extract bioactive compounds. A generalized vapor pressure curve of CO2 and its end are presented in Fig. 7. As the nature of supercritical CO2 fluid (scCO2) is between gas and liquid, it has unusual properties compared to other solvents, e.g., strong solubility and a large mass transfer coefficient; it is non-toxic and easily accessed [153, 154]. However, scCO2 has a disadvantage: It is a non-polar solvent. It therefore needs co-solvents or modifiers like EtOH, MeOH, water, or combinations of co-solvents to increase the polarizability of scCO2 and strengthen the solvating power. The different co-solvent interactions strongly enhance solubility. On the other hand, scCO2 can work as a co-solvent if the mixture contains at least 60% EtOH or water. Conversely, it may act as an anti-solvent when the extract is insoluble in scCO2 during the depressurizing step [155, 156].
Jiao and Kermanshahi [157] used scCO2 to extract haskap berries with water as a co-solvent. ANCs were successfully extracted employing 45 × 106 Pa, 65 °C at 15–20 min. The data showed that using scCO2 and water as a co-solvent would result in a higher extraction yield than using water or EtOH as a solvent, 52.7% and 38.3% respectively. Studies by B. Wang et al. [158] and Lao et al. [159] also showed that high-pressure CO2–H2O (107 Pa, 60 °C, 15–20 min) generated more colorants from red cabbage and purple sweet potato respectively than traditional extraction methods with water and 70% acidified EtOH. Both studies revealed the feasibility of the scCO2–H2O extraction method as a promising technology in the future because of higher extract yields. Despite this, this technology has yet to be applied in the industry and is not used on a large scale because of the high cost of the equipment, enormous technical investments, and difficult control parameters [122, 124].
5.3 Pressurized hot water extraction
Pressurized hot water extraction (PHWE) or “subcritical” water extraction (SWE) is operated between 100 and 374 °C. Usually, the temperature ranges from above the boiling temperature (100 °C, 105 Pa) to below the critical temperature (374 °C, 221 × 105 Pa). The pressure is therefore adjusted to maintain the liquid state of water during the extraction. PHWE is superior because of the fast extraction time and high content of the collected phenolics compounds [160]. In PHWE, water can be heated up to 200 °C, and the change in the dielectric constant of the water with the temperature makes water act like an organic solvent. For example, the dielectric constant of water at 200 °C is about 36, roughly similar to MeOH [112].
The water-based extraction process has been reported to be appealing because of its ready-for-use and cost-free availability, nontoxicity, non-flammability, and acceptance as an eco-friendly solvent [161]. Both high pressure and temperature will disrupt the surface equilibrium. The elevated temperature can overcome some obstacles such as the solute-matrix interaction caused by van der Waals forces, hydrogen bonding, dipole attraction of the solute molecules, and active sites in the matrix [162]. Compared to SF extraction, pressurized liquid extraction, including water as a fluid, provides an inert environment and protects samples from light and oxygen. Additionally, higher temperatures and pressure increase the extraction capacity. Otherwise, the essential advantage of supercritical fluid extraction is the increased capacity of diffusion by the extraction fluid for accessing these compounds in the matrix [163, 164].
In PHWE, the use of the optimal high temperature is preferred to the highest temperature. The typical extraction pressure is 5 × 106–107 Pa, but the extraction time depends on the operating parameters such as the amount of water used, temperature, and pressure [160]. PHWE at 100–110 °C appears to be a potent alternative to organic solvents in extracting polyphenolic compounds [165]. The tannins were isolated by PHWE from Phyllanthus niruri at a temperature of 100 °C and pressure of 107 Pa. The PHWE achieved the highest overall tannin and corilagin yield with the shortest extraction time compared to the Soxhlet and SF extraction methods [166]. In another study, gallic acids, ellagic acids, and corilagin from Terminalia chebula Retz. PHWE extracted fruits at temperatures (120–220 °C) and waterflow rates (2–4 mL/min) at a pressure of 4 × 106 Pa. The extracts had the highest phenolic compound content with the highest antioxidant activity, revealing the efficiency of PHWE [167]. Chopping or macerating materials together with the homogenization steps can also increase economic feasibility for industrial purposes [168].
5.4 Ultrasound-assisted extraction
Ultrasound-assisted extraction (UAE) uses the cavitation and strong shear force generated by ultrasound with frequencies ranging between 20 kHz and 100 MHz to improve the extraction and shorten the extraction time [169, 170]. This sonication forms microbubbles, oscillates quickly, and eventually collapses strongly when the acoustic pressure is sufficiently high. The collapse occurs near the solid surface to produce high-speed micro-jets of liquid onto the surface. When these jets are associated with shocking waves, they can break solid surfaces. The behavior is based on two contrasting mechanisms. The first describes the high pressure and temperature produced in the bubbles during adiabatic compression. It is accountable for the cleavage of molecular bonds and radical forming. The second involves micro-discharges because of the high electrical field [169, 171, 172].
UAE is an inexpensive and valuable technology that does not require complex equipment. UAE has been broadly utilized in the extraction of compounds from leaves [173], stalks [114], fruit and berries such as strawberries [174], and plant seeds [175]. The main factors during extraction are ultrasound power, temperature, a solvent with low vapor pressure, extraction time, and sonication mode (continuous or pulsed). These elements determine the extracts’ yield and composition. Non-optimized parameters could lead to the degradation of thermo-labile compounds. If the solvent is too viscous, it will increase the wave amplitude because it is necessary to obtain the mechanical vibrations for the cavitation [171, 172, 176]. A temperature range between 20 and 70 °C can increase the extraction yield. The influence of temperature is explained by the increase of the amount of cavitation bubbles, large solid–liquid contact areas, and solvent diffusivity improvement. The effect therefore enhances the desorption and solubility of the interest compounds. However, if the temperature is near the solvent’s boiling point, the extraction yield will decrease due to the thermal instability of biomass which causes the degradation of bioactive compounds [171, 177, 178]. Notably, most of the works in UAE focus on the application of continuous ultrasound field, and few researchers considered pulsating acoustic field. Although there are no essential differences between modes of UAE such as efficiency of extraction and times, pulse mode results in great energy saving compared to continuous UAE, better temperature control, and a reduction in corrosion on the ultrasound probe tip [176, 179,180,181].
UAE coupled with DES was considered an alternative green method to conventional technologies. Thus, combining DESs and UAE has the benefits of both methods, such as a shorter extraction time and lower solvent consumption than conventional solvent extraction [182, 183]. Sukor et al. [184] introduced intensified DES-mediated ultrasound extraction of tannic acids from onion peels. Using DES in extraction gave a higher tannic acid yield than MeOH. The concentration of tannic acids was maximized by adjusting the DES ratio to 1:1, the solid-to-solvent ratio to 1:10, and the duty cycle to 10%. Aslan Türker and Doğan [185] used a mixture of choline chloride with five different components. Among the tested DESs, choline chloride–based DES prepared with citric acid obtained the best solvent. This finding demonstrated that ultrasound could be used as a substituent energy source. Using DESs would also improve the flavonoid yield and maintain the color of the black carrot extract.
5.5 Pulse electric field extraction
The pulse electric field (PEF) method or pulse electric field pre-treatment is known for improving the pressing, drying, extracting time, and diffusing process without using a high temperature [108, 186]. It is an emerging non-thermal green process for isolating phytochemical compounds from plant parts such as leaves, roots, and fruit [152]. In principle, PEF destroys cell membrane structures to improve extraction. An electric potential pass through the membrane of a living cell in the electric field. The molecules are separated according to the charge in the cell membrane, whose functions are based on the dipole nature of the membrane molecules. Afterward, a drastic permeability increase is induced by the repulsion occurrence when the transmembrane potential exceeds 1V [108].
The impact of PEF on bioactive compounds depends on the field strength, specific energy input, pulse number, treatment temperature, and properties of the materials to be treated [187]. PEF treatment at a moderate electric field (from 500 to 1000 V/cm for 10–4–10–2 s) is found to damage the cell membrane of plant tissue with localized heating, resulting in the inactivation of enzymes and microorganisms. Hence, this can prevent the degradation of heat-sensitive compounds [188,189,190]. The industrial batch PEF extraction of polyphenols from grape peels is normally carried out at 50–60 °C for 20 h [191]. Corrales et al. [192] extracted ANCs from grape by-products using different isolation methods. The result indicated that PEF remarkably enhanced the extraction of ANC monoglucosides compared to the number of acylated glucosides extracted. PEF treatment is preferable before the maceration step in the winemaking process to reduce the extraction time. The performance of the ANC content and total extracted polyphenolic amount increased significantly after the maceration process when the electric field strength increased from 5 to 10 kV/cm [193].
5.6 Microwave-assisted extraction
Compared to conventional solid–liquid extraction, microwave-assisted extraction (MAE) gains higher efficient recovery and superior product quality at a low cost. MAE uses microwave energy to generate the molecular movement rotation of liquid under a permanent dipole, which results in the fast heating of both solvent and samples. This technology extends efficiency and reduces extraction time with low solvent consumption and a high level of automation [194, 195]. During microwave heating, the temperature radiation elevates the temperature of solids and solvents, and the subsequent viscosity of the extract is reduced; it thus promotes the solubility of the compounds, e.g., tannins. In addition, microwave radiation breaks the plant cells’ microstructure. This leads to the reduction of the mass transfer resistance of the components and encourages the diffusion of components from the source materials [196, 197]. MAE has recently been applied to extract various plant resources such as purple sweet potato, red cabbage, blackberry, cranberry, and sour cherry. However, the utilization of MAE and UAE technologies should be controlled with caution, as these methods can destroy the structure of flavonoids and tannins by attributing high temperature and excessive vibration to the samples respectively [122, 124].
5.7 Extraction methods for tulipalins
Few studies focus on extracting tuliposides from tulip biomass, despite the studies of their antibacterial activity [49, 107]. Typically, α-methylene-γ-butyrolactones have been prepared by the organic synthetic method [198]. Nevertheless, researchers have used tulip biomass years later to recover α-methylene-γ-butyrolactones with facile synthetic schemes. Tuliposides are considered the storage form of active tulipalins. Despite the conversion of tuliposides to tulipalins occurring non-enzymatically in nature due to their chemical lability, they can be converted spontaneously to tulipalin under neutral to basic conditions in vitro [199, 200].
Generally, the precursors of tulipalins would be converted into their active counterparts under stress with ethylene gas or an enzyme-mediated condition [201]. For example, Kato et al. [49] facilely extracted tulipalins from tulip tissue (petals). The extract was dissolved in cold water affording a tuliposide-containing extract of the tulip flowers. Potassium phosphate (KPi) buffer was then added to the extract. The procedure was then followed by incubation with and without the partially purified tuliposide-converting enzyme from the tulip bulb. The results showed that the yield of tulipalins was significantly higher than the extract incubated without tuliposide-converting enzyme, suggesting the potential applicability of tulipalins production. The study also indicated that using non-polar organic solvents was more effective than solvents with alkenes.
Similarly, Nomura et al. [50] explained a novel method: tulipalin B was directly formed from the precursor glucose ester 6-tuliposide B by an endogenous Pos-converting enzyme. The compounds were suspended in KPi buffer for 2 h at 4 °C. The combination of gum and resin contained contaminated starch granules, which were removed with a cheesecloth and water solution (equilibrated with 1 M NaCl). Nomura and co-workers also adopted coconut shell–derived activated charcoal (Yashi coal) as an absorbent of the hydrophobic impurities for partial purification of the flower extracts. Kerssemakers in 2018 [51] also introduced a method to extract tulipalin A from tulip bulbs by cutting them into smaller pieces to enhance the surface area. Artificial stressing of the bulb using ethylene gas triggered the mechanism, which eventually led to the conversion of tuliposides. Toluene and demi-water at 4 °C were used as solvents. The study also suggested that decreased polarity resulted in a higher affinity for non-polar solvents. As a result, tulipalin A would be more soluble in non-polar solvents such as benzene or toluene.
6 Impact of the extraction parameters
The effectiveness of the extraction process is principally based on the polarity of solvents because it contributes to the selectivity of the partition system. Therefore, the significant impacts of solvents on the extraction process are the extraction yield, composition profiles, and antioxidation activity of the extract [202]. Besides many selections of solvent as mentioned in Sect. 5, other conditions such as solid-to-solvent ratio, time and temperature, pH of the medium, and particle size of the matrix strongly influence the extraction efficiency [110].
6.1 Time, solid-to-solvent ratio, and temperature
Time, solid-to-solvent ratio, and extraction temperature are vital parameters to optimize to decrease the energy cost of the procedures. Many researchers enhanced the solubility of solute and the diffusion coefficient of the extraction process by elevating time and temperature. However, beyond a certain value, the phenolic compounds can be denatured [203]. The extracting time and temperature vary among studies. For example, the thermal stability of tannins is limited, and they therefore last from roughly 2 to 6 h at high temperatures (80–120 °C), and up to 24 h when the temperature is lower (25 °C) [71]. Sood and Gupta [204] reported that total phenol content extracted from pomegranate peel decreased with increased time and temperature. The same result was shown in the study of Silva et al. [205]. It indicated that 46.8 min with 58.2 °C was the most appropriate duration to extract phenols from Inga edulis leaves, which excessive time was not useful to extract more phenol content. If the temperature is elevated sufficiently, the time to reach equilibrium for the extraction of phenol compounds will be shortened. Nevertheless, long extraction time results in partially extracted polyphenols exposed to oxygen and light that were determined to degrade the extracts [206,207,208].
It is reported that the lower solid-to-solvent ratios would lead to higher extraction yields. Yet, too low solid-to-solvent ratios might lead to an excessive amount of solvent and a longer concentration time [110]. Also, at low solid-to-solvent ratios, the differences between the extraction efficiencies are usually inconsiderable. Thus, this parameter is generally optimized at a certain value [124]. In the study of Wang et al. [209], the increase of solvent used enhanced the flavonoid extract until the point 1:30 w/v, and then the yield decreased. Similarly, although several solid-to-solvent ratios from low to high were used to isolate bioactive compounds, the optimum conditions were not at the highest ratio but at a specific value, which was 1:4 w/v among from 1:1 to 1:7 w/v [210], or 1:8 w/v among 1:2 to 1:10 w/v [211].
A temperature range between 20 and 80 °C can promote higher analytic solubility by accelerating both the mass transfer rate and solubility. The viscosity and surface tension of solvents decrease at higher temperatures, which permit solvents to solvate phenolic compounds more efficiently and enhance the extraction rate. In contrast, some phenolic compounds such as ANCs start to degrade at high temperatures (above 70 °C). It is therefore pivotal to have the correct temperature to control the stability of phenolic compounds and reduce the thermal degradation of flavonoids [112, 212, 213]. For example, the thermostability of ANC is in a temperature range of 80–120 °C, and ANC starts to degrade following the first-order kinetics model in a temperature range of 120–180 °C [214, 215]. Using a temperature that is too high (above 150 °C) during the extraction degraded some antioxidant compounds in grape biomass and cultured red-wine berries [216, 217]. Additionally, tannins were extracted from Quercus infectoria (Manjakani) galls at different temperatures (50, 75, and 100 °C). The temperature effects on tannin content and antioxidant activity were thoroughly examined. The results suggested that the highest concentration of tannin and its highest antioxidant activity was achieved when the temperature was at 75 °C [218].
6.2 pH of the medium
Under the same time and temperature, the yield of chemical extraction depends on the types of solvents with varying polarities and pH. Phenolic compounds such as hydroxycinnamic acids and hydroxybenzoic acids are covalently linked to the polysaccharides of the cell walls by ester bonds. In addition, they are linked by ester or ether bonds to form lignin components. In mild alkaline hydrolysis, ester bonds are cleaved, and the ester-bonded phenolic compounds are released. On the other hand, in acidic conditions, alkyl-aryl-ether bonds are cleaved, and the ether-bonded compounds are released. As a result, many extraction methods are operated under mild alkaline or acidic conditions [219]. The polyphenol yield obtained from vine shoots was examined by basifying water with NaOH and EtOH. The results demonstrated that basified water with NaOH produced a higher polyphenol yield. The finding indicated that NaOH and EtOH had opposing action mechanisms, potentially caused by the carboxylic acid group in phenolic compounds. In an alkaline solution, the COOH group becomes COO–, which is soluble in water and not in EtOH [220]. Other studies found that the highest extraction yield of hydroxycinnamic acid glycosides from flaxseeds was obtained in alkaline hydrolysis by NaOH solution [221].
It has also been shown that acetic acid (about 7% v/v) or trifluoroacetic acid (TFA) (about 3% v/v) is more sufficient than mineral acids because mineral acids may cause the loss of the attached acyl group [222]. The use of hydrochloric acid (HCl) in many studies is therefore restricted, as it may break down the acylate groups. To acquire the best yield of flavonoid extraction, weak organic acids like formic acid, acetic acid, citric acid, tartaric acid, and phosphoric acid or low concentrations of strong acids such as 0.5–3.0% of TFA and less than 1.0% of HCl are suggested [112, 223]. Furthermore, sulfured water can be used as an alternative extraction solvent to reduce the use of organic solvents and extraction costs [224]. Ju and Howard [165] demonstrated that sulfured extraction possessed the highest flavonoid yield compared to subcritical water extraction and conventional solvent extraction (60% v/v MeOH at 50 °C for 1 h). Notably, most of the ANC compounds provide stability in acidic conditions and start to degrade at higher pH. Conversely, at higher pH or alkaline conditions, e.g., petanin, the degradation resistance occurs at pH 8 [58, 225].
6.3 The particle size of the matrix
The ideal temperature and the particle size of the samples are the essential factors that affect the yield during extraction. A smaller size prefers higher mass transfer facilitation. The smaller the particle size, the shorter path of the solvent to penetrate the solid matrices [226]. The reduction of the particle diameter is associated with the increased surface area. It therefore increases the contact area between the solvent and the samples [227]. Phenolics can be extracted from fresh, dried, and frozen plant samples. Pre-treated plant samples are usually air-dried and/or freeze-dried because freeze-drying generally retains more phenolics than air-drying [112, 228].
Gião et al. [229] proposed that a particle size of 0.2 mm was sufficient to assure a reasonable extraction rate by conducting conventional coffee mill grinding. Moreover, the highest yield of gallic acid (0.15 mg/g DW) was achieved when reducing the particle size to 125 µm. Particles smaller than 125 µm would not be used for the extraction since they tend to float [226]. Bucić-Kojić et al. [230] discovered that the extraction rate of polyphenol compounds from grape seed was extended when the particles were smaller. In this manner, a smaller particle size class (160–125 µm) induces a higher temperature impact on the extraction extent than a bigger particle size class (630 µm). This observation was supported by the highest value of activation energy (Ea) from the model of the solid–liquid extraction kinetics of total polyphenols.
Nevertheless, downsizing material steps such as grinding and milling consumes intense energy, correlated to the device type, dimensioning, and operating mode. Although the particle size range of biomass (ca. 100 µm) is limited at the laboratory scale due to the small powder amount, it cannot be neglected at the industrial scale. Some issues regarding the size handling process are health risks and explosion hazards. It must therefore be considered as soon as the process design in the scale-up procedures is built, as it influences technological options and induces the implementation of safety procedures and risk management [231].
7 Separation and purification methods
During the extraction process, impurities like sugars, oligosaccharides, proteins, and organic acids should be removed to increase the purity of the target molecules. Purification is a tedious, time-consuming, and expensive procedure. Usually, the protocols are optimized according to the targeted phenolic compounds [122]. Centrifugation for 15–20 min at 20,000–25,000 rpm may be applied to separate the crude plant matrix from the extract [232]. In principle, polyphenols can also be separated based on their MW, with the particles with an MW of 3000 Da to 100 kilodaltons (kDa). It can be removed by gel filtration, and higher MW compounds can be separated by regular ultrafiltration membranes [233].
In addition, the solvents used to extract polyphenols can be evaporated in rotary evaporation under vacuum conditions. The separation of phenolic compounds can also be accomplished using solvent separation methods (liquid–liquid phase separation) followed by chromatographic methods [232]. Until now, the common purification methods for polyphenols include column chromatography, membrane separation, high-speed counter-current chromatography, and high-efficiency preparative liquid chromatography [122]. In particular, few papers work with novel techniques to isolate tulipalin A and B, yet it remains feasible to separate tulipalins by high-efficiency preparative liquid chromatography [106] or by solid–liquid separation by vacuum filtration and rotary evaporation [51].
7.1 Column chromatography
Column chromatography is the conventional method for separating impurities and purifying polyphenols. It is also utilized for active molecule isolation and metabolite extraction from a broad variety of samples. Column chromatography is based on the principle of the various distribution coefficients of phenolic compounds between the solid and mobile phases [122, 234]. In principle, compounds are adsorbed, and the elution is based on the differential adsorption of a substance by adsorbents. In column chromatography, stationary phases can be silica, alumina, calcium phosphate, calcium carbonate, starch, etc. With different polarities, compounds are adsorbed for different reasons and desorbed at compatible solvent polarity. In the elution process, fractions get desorbed and pass through the column when solvents are added [235]. Column chromatography is classified into three classes: gas chromatography (GC), liquid chromatography (LC), and supercritical fluid chromatography (SFC). As the titles imply, the mobile phases in the three methods are gas, liquid, and supercritical fluid chromatography, respectively. As illustrated in Table 4, many chromatography methods are categorized into the first two classes. Notably, LC can be performed either in a column or planar surface. On the other hand, GC and SFC are limited to column procedures, so the column walls enclose the mobile phase [236]. The identification of components separated by column chromatography requires additional analytical methods to confirm the fractions. The crude extracts (complex chemical structures) then can be analyzed by many techniques such as thin layer chromatography (TLC), high-performance liquid chromatography, and GC to distinguish and quantify the desired compounds [234].
The typical column for separations is a reverse phase (RP) C18 column. EtOH, MeOH, acetone, and water are commonly used as eluting a solvent. For example, in the case of size exclusion column chromatography, MeOH is more commonly used than EtOH to elute non-tannin compounds, whereas the acetone–water mixture is preferred to the MeOH–water mixture when polymeric procyanidins are the targeted compounds [112]. The adsorption mechanism of flavonoids on the resin is unclear, as multiple interactions occur at the solid–liquid phase, e.g., hydrophobic, dipole–dipole, and hydrogen bonding. Besides, the rates and equilibria of adsorption vary significantly in physical and chemical properties like surface area, pore radius, porosity, solvent, pH, and temperature. Although column chromatography is the most common purification method for bioactive compounds, it has been abandoned because it is too labor-intensive in large-scale production [112, 122].
7.2 Membrane separation
Membrane separation is an efficient method for purifying and concentrating desirable compounds. The semipermeable membrane allows smaller molecules to pass through and retain larger molecules. The membrane selectively restricts the transportation of various chemicals, including solids, liquids, and gases. The membrane separation is based on the pore size of the applied membrane and can be separated into microfiltration, ultrafiltration, nanofiltration, and reverse osmosis [110, 237]. Membrane filtration is operated in two different ways: dead-end and cross-flow modes. The dead-end operation forces all the feed transportation through the membrane, causing build-up and subsequent performance loss; it therefore requires frequent cleaning. In the cross-flow operation, the flow is parallel to the membrane surface, with the inlet stream passing through the membrane in a specific position [237].
In the food and beverage industries, membrane separation operation is based on the particle or molecular size and is driven by pressure. Other membrane technologies such as osmotic distillation, membrane distillation, and pervaporation have been studied recently for the same application area. In particular, ultrafiltration and nanofiltration have gained great interest in separating and purifying bioactive compounds from plant extracts and among agro-food-based industries [238,239,240,241,242]. The membrane process has been used industrially for agro-food by-products such as olive mill wastewater. For example, Karakulski, Kozlowski, and Morawski [243] introduced ultrafiltration technology to decrease the chemical oxygen consumption for the direct sewer discharge of waste.
Table 5 shows the characteristics of polyphenols in terms of their MW. Nanofiltration has a pore size in the range of 0.5–1 nm, and the primary range of the MW cut-off (MWCO) is between 200 and 1000 Da. Ultrafiltration consists of membranes, with the MWCO in the range of 1–300 kDa and a 0.01 µm pore size [242]. Single membrane filtration does not purify the extracts effectively. A sequence of microfiltration, ultrafiltration, and nanofiltration was applied to isolate a bioactive compound from an olive leaf extract. Microfiltration followed by ultrafiltration separated the impurities more prominently than 5 kDa, whereas the nanofiltration recovered the antibacterial and antioxidant polyphenol compounds [244].
7.3 High-performance liquid chromatography
High-performance liquid chromatography (HPLC) is based on the different distribution coefficients of solute molecules in stationary (pack material of column) and mobile (eluent) phases. Depending on the chemical structure of the samples, the molecules are retarded while passing the stationary phase in the column under high pressure. The intermolecular interactions between the sample molecules and packing material determine the retention time. Thus, the separation process is achieved [246].
There are two types of phase systems used in HPLC: normal phase and reserved phase. In the normal phase, the stationary is a polar adsorbent such as silica. These bonded phases contribute to better elution profiles because they are moderately polar and have a more uniform surface. The mobile phase in this mode includes a non-polar solvent such as hexane that added a more polar modifier, or methylene chloride to control solvent strength and selectivity [247]. On the other hand, HPLC uses mostly reserved phase (RP) as an analytical technique for separating complex mixtures in the chemical, pharmaceutical, and biotechnological industries. RP-HPLC utilizes a non-polar stationary phase and a polar aqueous mobile phase. The mobile phase is usually water mixed with at least one of the water-miscible organic solvents such as acetonitrile, MeOH, EtOH, tetrahydrofuran, and isopropanol, which act as modifiers. The solutes are separated using their hydrophobicity, to which a more hydrophobic solute will be retained on the column longer than a less hydrophobic one [247,248,249]. The selection of the mobile phase compositions, consisting of solvents, buffers, and other additives, will control the interaction degree between the solutes and the stationary phase. For example, acetonitrile is the most popular selection as a modifier due to its low viscosity, which decreases the back pressure, low cut-off wavelength for UV detection, adequate large elution strength, reduced reactivity, and ability to dissolve a wide range of solutes. As alternatives, MeOH or EtOH can be used as a solvent in the mobile phase because they are less toxic, less expensive, and have higher polarity compared to acetonitrile [249]. The introduction of RP columns gains significantly improved HPLC separation for different classes of phenolic compounds, and RP C18 columns are exclusively utilized, followed by C8. The reasons for these references are the relatively high organic content for solute interaction and sufficient chemical stability of the working pH range compared to shorter alkyl chain phases [249, 250]. HPLC is divided into two types: analytical or preparative scale. Analytical HPLC is used for identification, detection, and small-scale preparation. Meanwhile, preparative HPLC is employed to separate and prepare samples [251]. Hence, pure compounds can be obtained by high-performance preparative liquid chromatography. This technique is based on the different physicochemical properties of each component in the crude extract, including ANCs, proanthocyanidins, hydrolyzable tannins, flavonols, flavan-3-ols, flavanones, flavones, and phenolic acids in different plant and food samples [112]. However, the HPLC method has limitations such as with the complex matrixes. An initial preconcentration and purification of the polyphenols from the complex matrixes are therefore crucial before the analysis by HPLC [250].
Also, the chromatographic behavior of the HPLC column will be altered and increased operating pressure under intensive use by the strongly retained materials such as lipids, or hydrophobic proteins. To solve this problem, washing the column with one or a series of strong solvents is essential. Lipids, for example, can be removed by using non-polar solvents such as MeOH, acetonitrile, and tetrahydrofuran. There is no solvents or series of solvent used to strip or remove proteins from the stationary phase. Yet, several solvents, such as acetic acid, (1% v/v in water) or aqueous sodium chloride, sodium phosphate, or sodium sulfate (1–2M) can be applied as eluent or solubilizing agents that strip all adsorbed proteins in HPLC column [252]. In reversed phase column, when a mobile phase does not use buffers or salts, the column can be washed with eluent by the same solvents as that of the mobile phase, only with higher organic solvent concentration [253]. In the case of buffered aqueous mobile phase, using strong solvents immediately is not recommended. A sudden change in the high content of organic solvent might lead to buffer precipitation in the HPLC flowing system and cause plugged frits, plugged connecting tubing, pump seal failure, a scratched piston, or injection valve rotor faults. Therefore, a buffer-free mobile phase, eluent containing the same ratio of water and organic solvent, is used to flush the column then strong solvents are applied to pass through the column [254]. Under the normal phase column, the column is cleaned by using a solution having the same solvents as that of the mobile phase, but in a higher concentration. When polar components adsorb in the column, isopropanol or similar solvents are used to flush them [253].
7.4 High-speed counter-current chromatography
High-speed counter-current chromatography (HSCCC) is an alternative to liquid chromatography. It is a liquid–liquid extraction without the support of any solid that eliminates the irreversible adsorption of compounds onto the solid support matrix. HSCCC is considered a potent candidate for large-scale production, with high recovery and high efficiency, which can improve the loading capacity of samples. The impurities of the desired fraction would be removed, and the yield of purified polyphenols increases after purification. HSCCC can be operated in various phases and modes: normal phases, reserve phase, pH zone refining, dual mode, gradient mode, and recycling mode [255, 256]. According to Degenhardt et al. [257], methyl tert-butyl ether (MTBE)/n-butanol/acetonitrile/water 2:2:1:5 v/v/v/v acidified with 0.1% TFA was employed as eluent to isolate ANCs from red cabbage, black currant, black chokeberry, and roselle. In addition, HSCCC can be applied to separate monomeric catechins, their galloylated esters, and caffeine from tea and other polyphenols from food like procyanidins, phenolic acids, and flavonol glycosides using MTBE/acetonitrile/0.1% aqueous TFA (2:2:3, v/v/v) [258].
8 Challenges and prospects in bioactive compounds extraction
This review provides an adequate understanding of different advanced extraction methodologies used to separate/extract bioactive compounds widely solicited as colorants and antibacterial agents. As a result, Table 6 provides a summary of the different extraction techniques of phytochemicals from plants, presenting details on the conditions, merits, and demerits of each method. In terms of natural products extraction, particularly bioactive compounds from innovative technologies, six principles have been proposed: (1) selection of varieties and use of the plant resources; (2) use of alternative solvents, mainly water or agro-solvents; (3) decrease of energy consumption by advanced extraction technologies; (4) produce co-products instead of wastes, consisting of bio-and agro-refining industry; (5) decrease unit operations and favor safe, robust, and controlled process; (6) target for non-denatured and biodegradable extracts in the absence of contaminants. When choosing a suitable extraction technology, one should follow those principles to connect with the biorefinery concept [259]. Indeed, the use of plant-derived colorants and antimicrobial agents has been perceived as having health-promoting benefits and coloring qualities. However, in the textile field, the major problem of its natural colorants and antimicrobial agents encountered is inadequate availability because of the matters in the collection, prices, compounds’ characteristics, and especially standardized methods of their production [260]. By applying the biorefinery concept, the evaluations of extraction efficiency, economic feasibility, and environmental perspectives are crucial to designing an appropriate extraction process for eco-textile products. Several extraction strategies have been conducted over the years, concerning properties such as sources of matrix, chemical structures, solubility, and chemical properties of both raw materials and solvents used, as well as the conditions during the procedure such as pH, temperature, and extraction time [108].
The manuscript also demonstrates that the utilization of green solvent with auxiliaries under pressurized conditions is the key factor in designing a novel extraction process. Under the advancement of technology, combinations like ultrasound-assisted DES extraction are more effective than a single method. The supplement of enzymes, including cellulose and pectinase for the extraction process such as ultrasound processing, is broadly used to recover ANCs from grape skin and raspberry wine residues. From another perspective, solvent utilization is under investigation to eliminate hazards using non-toxic chemicals and improve the yield of bioactive compounds, and energy consumption. For instance, DESs are promising alternatives to organic solvents [273]. Likewise, the addition of ultrasound-assisted technology allows the tannin extraction process to save 34% energy and 34% water, improving the feasibility of scaling up the extraction process in the industry [274]. As a highlight of solvent selection, the employment of Hansen Solubility Parameters is known as a tool to investigate the solubility parameter theory, to narrow the choice of the most sufficient green solvents to isolate bioactive compounds [275, 276].
Therefore, these advanced techniques encourage the idea that it may be possible to develop chemicals with improved safety profiles, utilizing safe solvents auxiliaries, design for energy efficiency, and degradation prevention [108]. However, scaling up the extraction methods for industry also faces many demerits and complexities such as limited recovery, the degradation of phenolic compounds, equipment cost, and maintenance feasibility. Importantly, energy and solvent consumption can be mentioned as essential subjects [277, 278]. For instance, in the SF extraction process, many factors such as extractors, reactors, CO2 reservoirs, heat exchangers, piping, valves, and operations automation to reduce manpower costs need to be appropriately considered. These units add significant costs, increasing the total investment cost of building a pilot or industrial-scale SF extraction. Moreover, the cleaning section should receive attention due to its time consumption in large-scale operations [279]. Accordingly, the need for scaling up should be integrated with the application of environmental assessment tools (life cycle assessment, etc.) and economic studies to propose safer and cleaner alternatives to more ordinary processes [259]. Consequently, more data from relevant materials and resources to emissions and wastes can be collected to study the potency of different bioactive compound extraction processes for diverse applications [280].
9 Conclusion
Innovative extraction method trends have been evolving. This paper critically reviews the extraction technologies, separation methods, and influential parameters of bioactive compound extraction. The review indicates the potential of novel extraction techniques for utilizing greener solvents in shorter times and less energy, such as scCO2 and PHWE. However, the advances in isolating phenolic compounds are restricted due to the necessity of more optimization efforts between the laboratory and industrial scales. Advanced extraction technologies can be advantageous in time, energy, and product yield; yet the overall efficiency, notably the equipment and maintenance costs, are not as affordable as conventional techniques. Each extraction method can only sufficiently extract some of the valuable compounds in plants. The most appropriate methodology is chosen based on the plant matrix and the targeted compounds. Strategies should therefore be determined to achieve higher efficiency and profitability. The initial step is to select the appropriate plant matrix according to local availability. In addition, extraction conditions such as time, temperature, pressure, reactor types, and the quality of raw materials should be optimized during methodological conduction. Numerous opportunities can expand natural raw feedstocks and their utilization as textile finishing agents and colorants, as well as other valuable applications in the future.
Data availability
Not applicable.
References
Nguyen VT (2017) Recovering bioactive compounds from agricultural wastes. Wiley, Incorporated, New York, United Kingdom
Sharma P, Gaur VK, Sirohi R et al (2021) Sustainable processing of food waste for production of bio-based products for circular bioeconomy. Bioresour Technol 325:124684. https://doi.org/10.1016/j.biortech.2021.124684
Sharma M, Sridhar K, Gupta VK, Dikkala PK (2022) Greener technologies in agri-food wastes valorization for plant pigments: step towards circular economy. Curr Res Green Sustain Chem 5:100340. https://doi.org/10.1016/j.crgsc.2022.100340
Gunarathne DS, Udugama IA, Jayawardena S et al (2019) Resource recovery from bio-based production processes in developing Asia. Sustain Prod Consum 17:196–214. https://doi.org/10.1016/j.spc.2018.11.008
Ortega F, Versino F, López OV, García MA (2022) Biobased composites from agro-industrial wastes and by-products. Emergent Mater 5:873–921. https://doi.org/10.1007/s42247-021-00319-x
Zhao Y, Wu Y, Wang M (2015) Bioactive substances of plant origin. In: Cheung PCK, Mehta BM (eds) Handbook of food chemistry. Springer, Berlin, Heidelberg. https://doi.org/10.1007/978-3-642-36605-5_13
Biesalski H-K, Dragsted LO, Elmadfa I et al (2009) Bioactive compounds: definition and assessment of activity. Nutrition 25:1202–1205. https://doi.org/10.1016/j.nut.2009.04.023
Bernhoft A (2010) Bernhoft A (2010) Bioactive compounds in plants: Benefits and risks for man and animals. In: Proceedings from a Symposium Held in Norwegian Academy of Science and Letters, Oslo, 13-14 November 2008. Novus Forlag
Veneziani G, Novelli E, Esposto S et al (2017) Chapter 11 - Applications of recovered bioactive compounds in food products. In: Galanakis CM (ed) Olive Mill Waste. Academic Press, pp 231–253. https://doi.org/10.1016/B978-0-12-805314-0.00011-X
Delgado-Vargas F, Jiménez AR, Paredes-López O (2000) Natural pigments: carotenoids, anthocyanins, and betalains–characteristics, biosynthesis, processing, and stability. Crit Rev Food Sci Nutr 40:173–289. https://doi.org/10.1080/10408690091189257
Vankar PS, Shukla D (2019) Chapter 1 - Newer natural dyes for various textiles. In: Vankar PS, Shukla D (eds) New trends in natural dyes for textiles. Woodhead Publishing, pp 1–6969. https://doi.org/10.1016/B978-0-08-102686-1.00001-9
Uddin MA, Rahman MM, Haque ANMA et al (2022) Textile colouration with natural colourants: a review. J Clean Prod 349:131489. https://doi.org/10.1016/j.jclepro.2022.131489
Torchani A, Saadaoui S, Gharbi R, Fathallah M (2015) Sensitized solar cells based on natural dyes. Curr Appl Phys 15:307–312. https://doi.org/10.1016/j.cap.2015.01.003
Wang C, Chen L (2005) Aromachology and its application in the textile field. Fibres Text East Eur. https://api.semanticscholar.org/CorpusID:67851456
Brudzyńska P, Sionkowska A, Grisel M (2021) Plant-derived colorants for food, cosmetic and textile industries: a review. Materials 14:3484. https://doi.org/10.3390/ma14133484
Luján-Ornelas C, Güereca LP, Franco-García M-L, Heldeweg M (2020) A life cycle thinking approach to analyse sustainability in the textile industry: a literature review. Sustainability 12:10193. https://doi.org/10.3390/su122310193
Furferi R, Volpe Y, Mantellassi F (2022) Circular economy guidelines for the textile industry. Sustainability 14:11111. https://doi.org/10.3390/su141711111
Harsanto B, Primiana I, Sarasi V, Satyakti Y (2023) Sustainability innovation in the textile industry: a systematic review. Sustainability 15:1549. https://doi.org/10.3390/su15021549
Franco MA (2017) Circular economy at the micro level: a dynamic view of incumbents’ struggles and challenges in the textile industry. J Clean Prod 168:833–845. https://doi.org/10.1016/j.jclepro.2017.09.056
de Oliveira Neto GC, Ferreira Correia JM, Silva PC et al (2019) Cleaner Production in the textile industry and its relationship to sustainable development goals. J Clean Prod 228:1514–1525. https://doi.org/10.1016/j.jclepro.2019.04.334
AIPH (2019) Ornamental horticulture: a growing industry? Int Assoc Horticult Producers (AIPH). https://aiph.org/wp-content/uploads/2020/11/GIIC_AIPH_IVP_Report2019.pdf
Statista (2022) Global fresh vegetable production by region 2020. In: Statista. https://www.statista.com/statistics/264066/global-vegegable-production-by-region/. Accessed 6 Sep 2022
Gabellini S, Scaramuzzi S (2022) Evolving consumption trends, marketing strategies, and governance settings in ornamental horticulture: a grey literature review. Horticulturae 8:234. https://doi.org/10.3390/horticulturae8030234
van Heck E (2021) Technology meets flowers: unlocking the circular and digital economy. Springer International Publishing, Cham
Leus L, Van Laere K, De Riek J, Van Huylenbroeck J (2018) Rose. In: Van Huylenbroeck J (ed) Ornamental Crops. Springer International Publishing, Cham, pp 719–767
Marriott M (2017) Modern (post-1800). In: Reference module in life sciences. https://doi.org/10.1016/B978-0-12-809633-8.05063-9
FAO (2021) World food and agriculture – statistical yearbook 2021. FAO, Rome, Italy. https://doi.org/10.4060/cb4477en
Baskar C, Ramakrishna S, Baskar S et al (2022) Handbook of solid waste management: sustainability through circular economy. Springer Nature Singapore, Singapore. https://doi.org/10.1007/978-981-16-4230-2
Hembrom R (2017) Horticultural waste-a potential source of antioxidants
Nichenametla SN, Taruscio TG, Barney DL, Exon JH (2006) A review of the effects and mechanisms of polyphenolics in cancer. Crit Rev Food Sci Nutr 46:161–183. https://doi.org/10.1080/10408390591000541
Bueno JM, Ramos-Escudero F, Sáez-Plaza P et al (2012) Analysis and antioxidant capacity of anthocyanin pigments. Part I: General considerations concerning polyphenols and flavonoids. Crit Rev Anal Chem 42:102–125. https://doi.org/10.1080/10408347.2011.632312
Bitis L, Sen A, Ozsoy N et al (2017) Flavonoids and biological activities of various extracts from Rosa sempervirens leaves. Biotechnol Biotechnol Equip 31:299–303. https://doi.org/10.1080/13102818.2016.1277956
Cendrowski A, Ścibisz I, Mitek M et al (2017) Profile of the phenolic compounds of Rosa rugosa petals. J Food Qual 2017:e7941347. https://doi.org/10.1155/2017/7941347
Olennikov DN, Chemposov VV, Chirikova NK (2021) Metabolites of prickly rose: chemodiversity and digestive-enzyme-inhibiting potential of Rosa acicularis and the main ellagitannin rugosin D. Plants 10:2525. https://doi.org/10.3390/plants10112525
Wang Y, Zhao Y, Liu X et al (2022) Chemical constituents and pharmacological activities of medicinal plants from Rosa genus. Chin Herb Med 14:187–209. https://doi.org/10.1016/j.chmed.2022.01.005
Manninen M, Vesterinen V-M, Salminen J-P (2020) Chemistry of autumn colors: quantitative spectrophotometric analysis of anthocyanins and carotenoids and qualitative analysis of anthocyanins by ultra-performance liquid chromatography–tandem mass spectrometry. J Chem Educ 97:772–777. https://doi.org/10.1021/acs.jchemed.9b00436
Lisandru T, Bunea A, Füstös A et al (2018) Chromatographic analysis of carotenoids and anthocyanins in sweet cherry autumn leaves used in ornamental landscapes. Stud Univ Babeş-Bolyai Chem 63:143–156. https://doi.org/10.24193/subbchem.2018.2.13
Silva N, Ruiz-Cruz S, Chaidez-Quiroz C et al (2014) Chemical constitution and effect of extracts of tomato plants byproducts on the enteric viral surrogates. Int J Environ Health Res 25:1–13. https://doi.org/10.1080/09603123.2014.938030
Guuntekin E, Uner B, Karakus B (2009) Chemical composition of tomato (Solanum lycopersicum) stalk and suitability in the particleboard production. J Env Biol, pp 731–734
Junker-Frohn LV, Lück M, Schmittgen S et al (2019) Tomato’s green gold: bioeconomy potential of residual tomato leaf biomass as a novel source for the secondary metabolite rutin. ACS Omega 4:19071–19080. https://doi.org/10.1021/acsomega.9b01462
Tang J, Meng X, Liu H et al (2010) Antimicrobial activity of sphingolipids isolated from the stems of cucumber (Cucumis sativus L.). Molecules 15:9288–9297. https://doi.org/10.3390/molecules15129288
Miazek K, Lebecque S, Hamaidia M et al (2016) Sphingolipids: promising lipid-class molecules with potential applications for industry. A review. BASE. https://doi.org/10.25518/1780-4507.13071
Alves E, Dias M, Lopes D et al (2020) Antimicrobial lipids from plants and marine organisms: an overview of the current state-of-the-art and future prospects. Antibiotics 9:441. https://doi.org/10.3390/antibiotics9080441
Hao X, Papadopoulos AP (1999) Effects of supplemental lighting and cover materials on growth, photosynthesis, biomass partitioning, early yield and quality of greenhouse cucumber. Sci Hortic 80:1–18. https://doi.org/10.1016/S0304-4238(98)00217-9
Tuama AA, Mohammed AA (2019) Phytochemical screening and in vitro antibacterial and anticancer activities of the aqueous extract of Cucumis sativus. Saudi J Biol Sci 26:600–604. https://doi.org/10.1016/j.sjbs.2018.07.012
Dick E, Onyemaechi John O, Pierre A et al (2016) Comparative studies of the phytochemistry, proximate analysis, mineral and vitamin compositions of the methanol leaf extracts of Cucumis sativus L. and Daucus carota L. Int J Pharmacol Res 6:282–285. https://doi.org/10.7439/ijpr/16
Mallik J, Akhter R (2012) Phytochemical screening and in-vitro evaluation of reducing power, cytotoxicity and anti-fungal activities of ethanol extracts of Cucumis sativus. Int J Pharm Biol Arch 3:555–560
Panina Y, Fravel DR, Baker CJ, Shcherbakova LA (2007) Biocontrol and plant pathogenic Fusarium oxysporum-induced changes in phenolic compounds in tomato leaves and roots. J Phytopathol 155:475–481. https://doi.org/10.1111/j.1439-0434.2007.01260.x
Kato Y, Yoshida H, Shoji K et al (2009) A facile method for the preparation of α-methylene-γ-butyrolactones from tulip tissues by enzyme-mediated conversion. Tetrahedron Lett 50:4751–4753. https://doi.org/10.1016/j.tetlet.2009.06.018
Nomura T, Hayashi E, Kawakami S et al (2015) Environmentally benign process for the preparation of antimicrobial α-methylene-β-hydroxy-γ-butyrolactone (tulipalin B) from tulip biomass. Biosci Biotechnol Biochem 79:25–35. https://doi.org/10.1080/09168451.2014.946395
Kerssemakers A (2018) A novel approach for the production and isolation of tulipalin-A from bulbous tulip biomass. Wageningen University. https://edepot.wur.nl/444903
Beecher GR (2003) Overview of dietary flavonoids: nomenclature, occurrence and intake. J Nutr 133:3248S-3254S. https://doi.org/10.1093/jn/133.10.3248S
Yonekura-Sakakibara K, Higashi Y, Nakabayashi R (2019) The origin and evolution of plant flavonoid metabolism. Front Plant Sci 10. https://doi.org/10.3389/fpls.2019.00943
Afendi FM, Okada T, Yamazaki M et al (2012) KNApSAcK family databases: integrated metabolite–plant species databases for multifaceted plant research. Plant Cell Physiol 53:e1. https://doi.org/10.1093/pcp/pcr165
Alzand K (2012) Flavonoids: Chemistry, biochemistry and antioxidant activity. J Pharm Res 5(8):4013–4020
Kumar S, Pandey AK (2013) Chemistry and biological activities of flavonoids: an overview. Sci World J 2013:162750. https://doi.org/10.1155/2013/162750
Cui L, Liu X, Tian Y et al (2016) Flavonoids, flavonoid subclasses, and esophageal cancer risk: a meta-analysis of epidemiologic studies. Nutrients 8:350. https://doi.org/10.3390/nu8060350
Khoo HE, Azlan A, Tang ST, Lim SM (2017) Anthocyanidins and anthocyanins: colored pigments as food, pharmaceutical ingredients, and the potential health benefits. Food Nutr Res 61:1361779. https://doi.org/10.1080/16546628.2017.1361779
Zhao D, Tao J (2015) Recent advances on the development and regulation of flower color in ornamental plants. Front Plant Sci 6. https://doi.org/10.3389/fpls.2015.00261
Schaberg PG, Van Den Berg AK, Murakami PF et al (2003) Factors influencing red expression in autumn foliage of sugar maple trees. Tree Physiol 23:325–333. https://doi.org/10.1093/treephys/23.5.325
Giada M de LR (2013) Food phenolic compounds: main classes, sources and their antioxidant power. IntechOpen. https://doi.org/10.5772/51687
Yang K, Zhang L, Liao P et al (2020) Impact of gallic acid on gut health: focus on the gut microbiome, immune response, and mechanisms of action. Front Immunol 11. https://doi.org/10.3389/fpls.2019.00943. https://doi.org/10.3389/fimmu.2020.580208
El-Seedi HR, Taher EA, Sheikh BY et al (2018) Chapter 8 - Hydroxycinnamic acids: natural sources, biosynthesis, possible biological activities, and roles in Islamic medicine. In: Atta-ur-Rahman (ed) Studies in natural products chemistry. Elsevier, pp 269–292. https://doi.org/10.1016/B978-0-444-64068-0.00008-5
Muronetz VI, Barinova K, Kudryavtseva S et al (2020) Natural and synthetic derivatives of hydroxycinnamic acid modulating the pathological transformation of amyloidogenic proteins. Molecules 25:4647. https://doi.org/10.3390/molecules25204647
Mattila P, Pihlava J-M, Hellström J (2005) Contents of phenolic acids, alkyl- and alkenylresorcinols, and avenanthramides in commercial grain products. J Agric Food Chem 53:8290–8295. https://doi.org/10.1021/jf051437z
Serrano J, Puupponen-Pimiä R, Dauer A et al (2009) Tannins: current knowledge of food sources, intake, bioavailability and biological effects. Mol Nutr Food Res 53:S310–S329. https://doi.org/10.1002/mnfr.200900039
Okuda T, Ito H (2011) Tannins of constant structure in medicinal and food plants—hydrolyzable tannins and polyphenols related to tannins. Molecules 16:2191–2217. https://doi.org/10.3390/molecules16032191
Fraga-Corral M, Otero P, Echave J et al (2021) By-products of agri-food industry as tannin-rich sources: a review of tannins’ biological activities and their potential for valorization. Foods Basel Switz 10:137. https://doi.org/10.3390/foods10010137
Le Bourvellec C, Renard CMGC (2019) Interactions between polyphenols and macromolecules: effect of tannin structure. In: Melton L, Shahidi F, Varelis P (eds) Encyclopedia of food chemistry. Academic Press, Oxford, pp 515–521
Arbenz A, Avérous L (2015) Chemical modification of tannins to elaborate aromatic biobased macromolecular architectures. Green Chem 17:2626–2646. https://doi.org/10.1039/C5GC00282F
Corral M, García Oliveira P, González Pereira A et al (2020) Technological application of tannin-based extracts. Molecules 25:614. https://doi.org/10.3390/molecules25030614
Oluwole O, Fernando WB, Lumanlan J et al (2022) Role of phenolic acid, tannins, stilbenes, lignans and flavonoids in human health – a review. Int J Food Sci Technol 57:6326–6335. https://doi.org/10.1111/ijfs.15936
Pizzi A, Mittal KL (2003) Handbook of adhesive technology, revised and expanded, 2nd edn. CRC Press, Boca Raton
Dhifi W, Bellili S, Jazi S et al (2016) Essential oils’ chemical characterization and investigation of some biological activities: a critical review. Med Basel Switz 3:E25. https://doi.org/10.3390/medicines3040025
Chouhan S, Sharma K, Guleria S (2017) Antimicrobial activity of some essential oils—present status and future perspectives. Medicines 4:58. https://doi.org/10.3390/medicines4030058
Atanasova T, Kakalova M, Stefanof L et al (2016) Chemical composition of essential oil from Rosa Damascena mill., growing in new region of Bulgaria. Ukr Food J 5:492–498. https://doi.org/10.24263/2304-974X-2016-5-3-8
Zolotilov V, Nevkrytaya N, Zolotilova O et al (2022) The essential-oil-bearing rose collection variability study in terms of biochemical parameters. Agronomy 12:529. https://doi.org/10.3390/agronomy12020529
Fazal-ur-Rehman, Adeel S, Khan Khosa K et al (2019) Revitalization of carotenoid-based natural colorants in applied field: a short review. In: Textiles and clothing. Wiley, Ltd, pp 45–78. https://doi.org/10.1016/B978-0-444-64068-0.00008-5
Zhu C, Bai C, Sanahuja G et al (2010) The regulation of carotenoid pigmentation in flowers. Arch Biochem Biophys 504:132–141. https://doi.org/10.1016/j.abb.2010.07.028
Venil CK, Velmurugan P, Dufossé L et al (2020) Fungal pigments: potential coloring compounds for wide ranging applications in textile dyeing. J Fungi Basel Switz 6:E68. https://doi.org/10.3390/jof6020068
Maghsoudi S, Taghavi Shahraki B, Rabiee N et al (2022) The colorful world of carotenoids: a profound insight on therapeutics and recent trends in nano delivery systems. Crit Rev Food Sci Nutr 62:3658–3697. https://doi.org/10.1080/10408398.2020.1867958
Zaripheh S, Erdman JW (2002) Factors that influence the bioavailablity of xanthophylls. J Nutr 132:531S-534S. https://doi.org/10.1093/jn/132.3.531S
Shoji K, Ubukata M, Momonoi K et al (2005) Anther-specific production of antimicrobial tuliposide B in tulips. J Jpn Soc Hortic Sci 74:469–475. https://doi.org/10.2503/jjshs.74.469
Nomura T, Ogita S, Kato Y (2012) A novel lactone-forming carboxylesterase: molecular identification of a tuliposide A-converting enzyme in tulip1. Plant Physiol 159:565–578. https://doi.org/10.1104/pp.112.195388
Shigetomi K, Kishimoto T, Shoji K, Ubukata M (2008) First total synthesis of 6-tuliposide B. Tetrahedron Asymmetry 19:1444–1449. https://doi.org/10.1016/j.tetasy.2008.05.019
Tzeng C-C, Lee K-H, Wang T-C et al (2000) Synthesis and cytotoxic evaluation of a series of γ-substituted γ-aryloxymethyl-α-methylene-γ-butyrolactones against cancer cells. Pharm Res 17:715–719. https://doi.org/10.1023/A:1007534416561
Yusuf M, Shabbir M, Mohammad F (2017) Natural colorants: historical, processing and sustainable prospects. Nat Prod Bioprospecting 7:123–145. https://doi.org/10.1007/s13659-017-0119-9
Manzoor M, Singh J, Gani A, Noor N (2021) Valorization of natural colors as health-promoting bioactive compounds: phytochemical profile, extraction techniques, and pharmacological perspectives. Food Chem 362:130141. https://doi.org/10.1016/j.foodchem.2021.130141
Abdel-Shafi S, Al-Mohammadi A-R, Sitohy M et al (2019) Antimicrobial activity and chemical constitution of the crude, phenolic-rich extracts of Hibiscus sabdariffa, Brassica oleracea and Beta vulgaris. Molecules 24:4280. https://doi.org/10.3390/molecules24234280
Giusti MM, Wrolstad RE (2003) Acylated anthocyanins from edible sources and their applications in food systems. Biochem Eng J 14:217–225. https://doi.org/10.1016/S1369-703X(02)00221-8
Puupponen-Pimiä R, Nohynek L, Ammann S et al (2008) Enzyme-assisted processing increases antimicrobial and antioxidant activity of bilberry. J Agric Food Chem 56:681–688. https://doi.org/10.1021/jf072001h
Rather LJ, Ali A, Zhou Q et al (2020) Instrumental characterization of merino wool fibers dyed with Cinnamomum camphora waste/fallen leaves extract: an efficient waste management alternative. J Clean Prod 273:123021. https://doi.org/10.1016/j.jclepro.2020.123021
Venugopalan A, Giridhar P (2012) Bacterial growth inhibition potential of annatto plant parts. Asian Pac J Trop Biomed 2:S1879–S1882. https://doi.org/10.1016/S2221-1691(12)60513-9
Kaczmarek B (2020) Tannic acid with antiviral and antibacterial activity as a promising component of biomaterials—a minireview. Materials 13:3224. https://doi.org/10.3390/ma13143224
Singh R, Jain A, Panwar S et al (2005) Antimicrobial activity of some natural dyes. Dyes Pigments 66:99–102. https://doi.org/10.1016/j.dyepig.2004.09.005
Shahid M, Shahid-ul-Islam MF (2013) Recent advancements in natural dye applications: a review. J Clean Prod 53:310–331. https://doi.org/10.1016/j.jclepro.2013.03.031
Gupta D, Khare SK, Laha A (2004) Antimicrobial properties of natural dyes against Gram-negative bacteria. Color Technol 120:167–171. https://doi.org/10.1111/j.1478-4408.2004.tb00224.x
Zhang W, Yang Z-Y, Tang R-C et al (2020) Application of tannic acid and ferrous ion complex as eco-friendly flame retardant and antibacterial agents for silk. J Clean Prod 250:119545. https://doi.org/10.1016/j.jclepro.2019.119545
Ghaheh FS, Nateri AS, Mortazavi SM et al (2012) The effect of mordant salts on antibacterial activity of wool fabric dyed with pomegranate and walnut shell extracts. Color Technol 128:473–478. https://doi.org/10.1111/j.1478-4408.2012.00402.x
Safaei-Ghomi J, Ahd AA (2010) Antimicrobial and antifungal properties of the essential oil and methanol extracts of Eucalyptus largiflorens and Eucalyptus intertexta. Pharmacogn Mag 6:172–175. https://doi.org/10.4103/0973-1296.66930
Astani A, Reichling J, Schnitzler P (2010) Comparative study on the antiviral activity of selected monoterpenes derived from essential oils. Phytother Res 24:673. https://doi.org/10.1002/ptr.2955
Devi KP, Nisha SA, Sakthivel R, Pandian SK (2010) Eugenol (an essential oil of clove) acts as an antibacterial agent against Salmonella typhi by disrupting the cellular membrane. J Ethnopharmacol 130:107–115. https://doi.org/10.1016/j.jep.2010.04.025
Shuaib A, Rohit A, Piyush M (2016) A review article on essential oils. J Med Plants Stud 4:237–240
Fisher K, Phillips C (2008) Potential antimicrobial uses of essential oils in food: is citrus the answer? Trends Food Sci Technol 19:156–164. https://doi.org/10.1016/j.tifs.2007.11.006
Walentowska J, Foksowicz-Flaczyk J (2013) Thyme essential oil for antimicrobial protection of natural textiles. Int Biodeterior Biodegrad 84:407–411. https://doi.org/10.1016/j.ibiod.2012.06.028
Christensen LP, Kristiansen K (1999) Isolation and quantification of tuliposides and tulipalins in tulips (Tulipa) by high-performance liquid chromatography. Contact Dermatitis 40:300–309. https://doi.org/10.1111/j.1600-0536.1999.tb06080.x
Shigetomi K, Shoji K, Mitsuhashi S, Ubukata M (2010) The antibacterial properties of 6-tuliposide B. Synthesis of 6-tuliposide B analogues and structure–activity relationship. Phytochemistry 71:312–324. https://doi.org/10.1016/j.phytochem.2009.10.008
Azmir J, Zaidul ISM, Rahman MM et al (2013) Techniques for extraction of bioactive compounds from plant materials: a review. J Food Eng 117:426–436. https://doi.org/10.1016/j.jfoodeng.2013.01.014
Bucar F, Wube A, Schmid M (2013) Natural product isolation – how to get from biological material to pure compounds. Nat Prod Rep 30:525–545. https://doi.org/10.1039/C3NP20106F
Zhang Q-W, Lin L-G, Ye W-C (2018) Techniques for extraction and isolation of natural products: a comprehensive review. Chin Med 13:20. https://doi.org/10.1186/s13020-018-0177-x
Turturică M, Oancea AM, Râpeanu G, Bahrim G (2015) Anthocyanins: naturally occuring fruit pigments with functional properties. Ann Univ Dunarea Jos Galati Fascicle VI - Food Technol 39:9–24
Dai J, Mumper RJ (2010) Plant phenolics: extraction, analysis and their antioxidant and anticancer properties. Molecules 15:7313–7352. https://doi.org/10.3390/molecules15107313
Oreopoulou A, Tsimogiannis D, Oreopoulou V (2019) Chapter 15 - Extraction of polyphenols from aromatic and medicinal plants: an overview of the methods and the effect of extraction parameters. In: Watson RR (ed) Polyphenols in plants (second edition). Academic Press, pp 243–259
Yang Y, Zhang F (2008) Ultrasound-assisted extraction of rutin and quercetin from Euonymus alatus (Thunb.) Sieb. Ultrason Sonochem 15:308–313. https://doi.org/10.1016/j.ultsonch.2007.05.001
Altemimi A, Lakhssassi N, Baharlouei A et al (2017) Phytochemicals: extraction, isolation, and identification of bioactive compounds from plant extracts. Plants 6:42. https://doi.org/10.3390/plants6040042
Reichardt C (2002) Classification of solvents. In: Solvents and solvent effects in organic chemistry. Wiley, Ltd, pp 57–91. https://doi.org/10.1002/3527601791
Sealy-Fisher VJ, Pizzi A (1992) Increased pine tannins extraction and wood adhesives development by phlobaphenes minimization. Holz Als Roh- Werkst 50:212–220. https://doi.org/10.1007/BF02663290
Aires A, Carvalho R, Saavedra MJ (2016) Valorization of solid wastes from chestnut industry processing: extraction and optimization of polyphenols, tannins and ellagitannins and its potential for adhesives, cosmetic and pharmaceutical industry. Waste Manag 48:457–464. https://doi.org/10.1016/j.wasman.2015.11.019
Panamgama LA (2007) Polyphenolic extracts of Pinus radiata bark and networking mechanisms of additive-accelerated polycondensates. J Appl Polym Sci 103:2487–2493. https://doi.org/10.1002/app.24466
Olalere OA, Gan C-Y, Adeyi O et al (2022) Upscalability and techno-economic perspectives of nonconventional essential oils extraction techniques. Jundishapur J Nat Pharm Prod 17.https://doi.org/10.5812/jjnpp-122792
Gould KS, Markham KR, Smith RH, Goris JJ (2000) Functional role of anthocyanins in the leaves of Quintinia serrata A. Cunn J Exp Bot 51:1107–1115. https://doi.org/10.1093/jexbot/51.347.1107
Tan J, Han Y, Han B et al (2022) Extraction and purification of anthocyanins: a review. J Agric Food Res 8:100306. https://doi.org/10.1016/j.jafr.2022.100306
Hashemi B, Shiri F, Švec F, Nováková L (2022) Green solvents and approaches recently applied for extraction of natural bioactive compounds. TrAC Trends Anal Chem 157:116732. https://doi.org/10.1016/j.trac.2022.116732
de Hoyos-Martínez PL, Merle J, Labidi J, Charrier – El Bouhtoury F (2019) Tannins extraction: a key point for their valorization and cleaner production. J Clean Prod 206:1138–1155. https://doi.org/10.1016/j.jclepro.2018.09.243
Xiao J, Chen G, Li N (2018) Ionic liquid solutions as a green tool for the extraction and isolation of natural products. Mol J Synth Chem Nat Prod Chem 23:1765. https://doi.org/10.3390/molecules23071765
Cvjetko Bubalo M, Radošević K, Radojčić Redovniković I et al (2014) A brief overview of the potential environmental hazards of ionic liquids. Ecotoxicol Environ Saf 99:1–12. https://doi.org/10.1016/j.ecoenv.2013.10.019
Hajipour AR, Rafiee F (2015) Recent progress in ionic liquids and their applications in organic synthesis. Org Prep Proced Int 47:249–308. https://doi.org/10.1080/00304948.2015.1052317
Lim JR, Chua LS, Mustaffa AA (2022) Ionic liquids as green solvent and their applications in bioactive compounds extraction from plants. Process Biochem 122:292–306. https://doi.org/10.1016/j.procbio.2022.10.024
Sillero L, Prado R, Welton T, Labidi J (2021) Energy and environmental analysis of flavonoids extraction from bark using alternative solvents. J Clean Prod 308:127286. https://doi.org/10.1016/j.jclepro.2021.127286
Han D, Row KH (2011) Determination of luteolin and apigenin in celery using ultrasonic-assisted extraction based on aqueous solution of ionic liquid coupled with HPLC quantification. J Sci Food Agric 91:2888–2892. https://doi.org/10.1002/jsfa.4553
Spittle S, Poe D, Doherty B et al (2022) Evolution of microscopic heterogeneity and dynamics in choline chloride-based deep eutectic solvents. Nat Commun 13:219. https://doi.org/10.1038/s41467-021-27842-z
Wei Z-F, Wang X-Q, Peng X et al (2015) Fast and green extraction and separation of main bioactive flavonoids from Radix Scutellariae. Ind Crops Prod 63:175–181. https://doi.org/10.1016/j.indcrop.2014.10.013
Cui Q, Peng X, Yao X-H et al (2015) Deep eutectic solvent-based microwave-assisted extraction of genistin, genistein and apigenin from pigeon pea roots. Sep Purif Technol 150:63–72. https://doi.org/10.1016/j.seppur.2015.06.026
Smith EL, Abbott AP, Ryder KS (2014) Deep eutectic solvents (DESs) and their applications. Chem Rev 114:11060–11082. https://doi.org/10.1021/cr300162p
Zhang Q, Vigier KDO, Royer S, Jérôme F (2012) Deep eutectic solvents: syntheses, properties and applications. Chem Soc Rev 41:7108–7146. https://doi.org/10.1039/C2CS35178A
García A, Rodríguez-Juan E, Rodríguez-Gutiérrez G et al (2016) Extraction of phenolic compounds from virgin olive oil by deep eutectic solvents (DESs). Food Chem 197:554–561. https://doi.org/10.1016/j.foodchem.2015.10.131
Peng X, Duan M-H, Yao X-H et al (2016) Green extraction of five target phenolic acids from Lonicerae japonicae Flos with deep eutectic solvent. Sep Purif Technol 157:249–257. https://doi.org/10.1016/j.seppur.2015.10.065
Dai Y, Witkamp G-J, Verpoorte R, Choi YH (2013) Natural deep eutectic solvents as a new extraction media for phenolic metabolites in Carthamus tinctorius L. Anal Chem 85:6272–6278. https://doi.org/10.1021/ac400432p
Wei Z, Qi X, Li T et al (2015) Application of natural deep eutectic solvents for extraction and determination of phenolics in Cajanus cajan leaves by ultra performance liquid chromatography. Sep Purif Technol 149:237–244. https://doi.org/10.1016/j.seppur.2015.05.015
Zhao B-Y, Xu P, Yang F-X et al (2015) Biocompatible deep eutectic solvents based on choline chloride: characterization and application to the extraction of rutin from Sophora japonica. ACS Sustain Chem Eng 3:2746–2755. https://doi.org/10.1021/acssuschemeng.5b00619
Qi X-L, Peng X, Huang Y-Y et al (2015) Green and efficient extraction of bioactive flavonoids from Equisetum palustre L. by deep eutectic solvents-based negative pressure cavitation method combined with macroporous resin enrichment. Ind Crops Prod 70:142–148. https://doi.org/10.1016/j.indcrop.2015.03.026
Ling JKU, Chan YS, Nandong J (2020) Extraction of antioxidant compounds from the wastes of Mangifera pajang fruit: a comparative study using aqueous ethanol and deep eutectic solvent. SN Appl Sci 2:1365. https://doi.org/10.1007/s42452-020-3153-x
Vilková M, Płotka-Wasylka J, Andruch V (2020) The role of water in deep eutectic solvent-base extraction. J Mol Liq 304:112747. https://doi.org/10.1016/j.molliq.2020.112747
Tobiszewski M (2019) Analytical chemistry with biosolvents. Anal Bioanal Chem 411:4359–4364. https://doi.org/10.1007/s00216-019-01732-2
Vian M, Breil C, Vernes L et al (2017) Green solvents for sample preparation in analytical chemistry. Curr Opin Green Sustain Chem 5:44–48. https://doi.org/10.1016/j.cogsc.2017.03.010
Calvo-Flores FG, Monteagudo-Arrebola MJ, Dobado JA, Isac-García J (2018) Green and bio-based solvents. Top Curr Chem 376:18. https://doi.org/10.1007/s41061-018-0191-6
Gu Y, Jérôme F (2013) Bio-based solvents: an emerging generation of fluids for the design of eco-efficient processes in catalysis and organic chemistry. Chem Soc Rev 42:9550–9570. https://doi.org/10.1039/C3CS60241A
del Pilar S-C, Pleite N, Herrero M et al (2017) New approaches for the selective extraction of bioactive compounds employing bio-based solvents and pressurized green processes. J Supercrit Fluids 128:112–120. https://doi.org/10.1016/j.supflu.2017.05.016
Santos LG, Martins VG (2022) Recovery of phenolic compounds from purple onion peel using bio-based solvents: thermal degradation kinetics and color stability of anthocyanins. J Food Process Preserv 46:e17161. https://doi.org/10.1111/jfpp.17161
Rombaut N, Tixier A-S, Bily A, Chemat F (2014) Green extraction processes of natural products as tools for biorefinery. Biofuels Bioprod Biorefining 8:530–544. https://doi.org/10.1002/bbb.1486
Amran MA, Palaniveloo K, Fauzi R et al (2021) Value-added metabolites from agricultural waste and application of green extraction techniques. Sustainability 13:11432. https://doi.org/10.3390/su132011432
Panja P (2018) Green extraction methods of food polyphenols from vegetable materials. Curr Opin Food Sci 23:173–182. https://doi.org/10.1016/j.cofs.2017.11.012
Nikolai P, Rabiyat B, Aliev A, Abdulagatov I (2019) Supercritical CO2: properties and technological applications - a review. J Therm Sci 28:394–430. https://doi.org/10.1007/s11630-019-1118-4
Kumar M, Dahuja A, Tiwari S et al (2021) Recent trends in extraction of plant bioactives using green technologies: a review. Food Chem 353:129431. https://doi.org/10.1016/j.foodchem.2021.129431
Cvjetko Bubalo M, Vidović S, Radojčić Redovniković I, Jokić S (2015) Green solvents for green technologies. J Chem Technol Biotechnol 90:1631–1639. https://doi.org/10.1002/jctb.4668
Maria V, Cunha B, da Silva M et al (2018) Carbon dioxide use in high-pressure extraction processes Provisional chapter Carbon Dioxide Use in High-Pressure Extraction Processes. https://doi.org/10.5772/intechopen.71151
Jiao G, Kermanshahi A (2018) Extraction of anthocyanins from haskap berry pulp using supercritical carbon dioxide: influence of co-solvent composition and pretreatment. LWT 98:237–244. https://doi.org/10.1016/j.lwt.2018.08.042
Wang B, Yang S, Xu L et al (2022) Evaluation study on extraction of anthocyanins from red cabbage using high pressure CO2 + H2O: a fuzzy logic model and metabolomic analysis. Sustainability 14:1369. https://doi.org/10.3390/su14031369
Lao F, Cheng H, Wang Q et al (2020) Enhanced water extraction with high-pressure carbon dioxide on purple sweet potato pigments: comparison to traditional aqueous and ethanolic extraction. J CO2 Util 40:101188. https://doi.org/10.1016/j.jcou.2020.101188
Castro-Puyana M, Herrero M, Mendiola JA, Ibáñez E (2013) Subcritical water extraction of bioactive components from algae. In: Functional ingredients from algae for foods and nutraceuticals. Elsevier, pp 534–560. https://doi.org/10.1533/9780857098689.3.534
Ameer K, Shahbaz HM, Kwon J-H (2017) Green extraction methods for polyphenols from plant matrices and their byproducts: a review. Compr Rev Food Sci Food Saf 16:295–315. https://doi.org/10.1111/1541-4337.12253
Teo CC, Tan SN, Yong JWH et al (2010) Pressurized hot water extraction (PHWE). J Chromatogr A 1217:2484–2494. https://doi.org/10.1016/j.chroma.2009.12.050
Palma M, Piñeiro Z, Barroso CG (2001) Stability of phenolic compounds during extraction with superheated solvents. J Chromatogr A 921:169–174. https://doi.org/10.1016/S0021-9673(01)00882-2
Piñeiro Z, Palma M, Barroso CG (2006) Determination of trans-resveratrol in grapes by pressurised liquid extraction and fast high-performance liquid chromatography. J Chromatogr A 1110:61–65. https://doi.org/10.1016/j.chroma.2006.01.067
Ju Z, Howard LR (2005) Subcritical water and sulfured water extraction of anthocyanins and other phenolics from dried red grape skin. J Food Sci 70:S270–S276. https://doi.org/10.1111/j.1365-2621.2005.tb07202.x
Markom M, Hasan M, Daud WRW et al (2007) Extraction of hydrolysable tannins from Phyllanthus niruri Linn.: effects of solvents and extraction methods. Sep Purif Technol 52:487–496. https://doi.org/10.1016/j.seppur.2006.06.003
Rangsriwong P, Rangkadilok N, Satayavivad J et al (2009) Subcritical water extraction of polyphenolic compounds from Terminalia chebula Retz. fruits. Sep Purif Technol 66:51–56. https://doi.org/10.1016/j.seppur.2008.11.023
Mohd Jusoh NH, Subki A, Yeap SK et al (2019) Pressurized hot water extraction of hydrosable tannins from Phyllanthus tenellus Roxb. BMC Chem 13:134. https://doi.org/10.1186/s13065-019-0653-0
Ötleş S (2009) Handbook of food analysis instruments. CRC Press, Boca Raton, FL
Belwal T, Huang H, Li L et al (2019) Optimization model for ultrasonic-assisted and scale-up extraction of anthocyanins from Pyrus communis ‘Starkrimson’ fruit peel. Food Chem 297:124993. https://doi.org/10.1016/j.foodchem.2019.124993
Chemat F, Rombaut N, Sicaire A-G et al (2017) Ultrasound assisted extraction of food and natural products. Mechanisms, techniques, combinations, protocols and applications. A review. Ultrason Sonochem 34:540–560. https://doi.org/10.1016/j.ultsonch.2016.06.035
Chaves JO, de Souza MC, da Silva LC et al (2020) Extraction of flavonoids from natural sources using modern techniques. Front Chem 8. https://doi.org/10.3389/fchem.2020.507887
Albu S, Joyce E, Paniwnyk L et al (2004) Potential for the use of ultrasound in the extraction of antioxidants from Rosmarinus officinalis for the food and pharmaceutical industry. Ultrason Sonochem 11:261–265. https://doi.org/10.1016/j.ultsonch.2004.01.015
Herrera MC, Luque de Castro MD (2004) Ultrasound-assisted extraction for the analysis of phenolic compounds in strawberries. Anal Bioanal Chem 379:1106–1112. https://doi.org/10.1007/s00216-004-2684-0
Rostagno MA, Palma M, Barroso CG (2003) Ultrasound-assisted extraction of soy isoflavones. J Chromatogr A 1012:119–128. https://doi.org/10.1016/s0021-9673(03)01184-1
Christou A, Stavrou IJ, Kapnissi-Christodoulou CP (2021) Continuous and pulsed ultrasound-assisted extraction of carob’s antioxidants: processing parameters optimization and identification of polyphenolic composition. Ultrason Sonochem 76:105630. https://doi.org/10.1016/j.ultsonch.2021.105630
Zhang Z-S, Wang L-J, Dong L et al (2008) Ultrasound-assisted extraction of oil from flaxseed. Sep Purif Technol 62:192–198. https://doi.org/10.1016/j.seppur.2008.01.014
Palma M, Barroso CG (2002) Ultrasound-assisted extraction and determination of tartaric and malic acids from grapes and winemaking by-products. Anal Chim Acta 458:119–130. https://doi.org/10.1016/S0003-2670(01)01527-6
Patience NA, Schieppati D, Boffito DC (2021) Continuous and pulsed ultrasound pectin extraction from navel orange peels. Ultrason Sonochem 73:105480. https://doi.org/10.1016/j.ultsonch.2021.105480
Brás T, Paulino AFC, Neves LA et al (2020) Ultrasound assisted extraction of cynaropicrin from Cynara cardunculus leaves: optimization using the response surface methodology and the effect of pulse mode. Ind Crops Prod 150:112395. https://doi.org/10.1016/j.indcrop.2020.112395
Pan Z, Qu W, Ma H et al (2012) Continuous and pulsed ultrasound-assisted extractions of antioxidants from pomegranate peel. Ultrason Sonochem 19:365–372. https://doi.org/10.1016/j.ultsonch.2011.05.015
Xue H, Tan J, Li Q et al (2020) Optimization ultrasound-assisted deep eutectic solvent extraction of anthocyanins from raspberry using response surface methodology coupled with genetic algorithm. Foods 9:1409. https://doi.org/10.3390/foods9101409
Savic Gajic I, Savic I, Boskov I et al (2019) Optimization of ultrasound-assisted extraction of phenolic compounds from black locust (Robiniae pseudoacaciae) flowers and comparison with conventional methods. Antioxidants 8:248. https://doi.org/10.3390/antiox8080248
Sukor NF, Selvam VP, Jusoh R et al (2021) Intensified DES mediated ultrasound extraction of tannic acid from onion peel. J Food Eng 296:110437. https://doi.org/10.1016/j.jfoodeng.2020.110437
Aslan Türker D, Doğan M (2021) Application of deep eutectic solvents as a green and biodegradable media for extraction of anthocyanin from black carrots. LWT 138:110775. https://doi.org/10.1016/j.lwt.2020.110775
Segovia FJ, Luengo E, Corral-Pérez JJ et al (2015) Improvements in the aqueous extraction of polyphenols from borage (Borago officinalis L.) leaves by pulsed electric fields: pulsed electric fields (PEF) applications. Ind Crops Prod 65:390–396. https://doi.org/10.1016/j.indcrop.2014.11.010
Heinz V, Toepfl S, Knorr D (2003) Impact of temperature on lethality and energy efficiency of apple juice pasteurization by pulsed electric fields treatment. Innov Food Sci Emerg Technol 4:167–175. https://doi.org/10.1016/S1466-8564(03)00017-1
Ade-Omowaye BIO, Angersbach A, Taiwo KA, Knorr D (2001) Use of pulsed electric field pre-treatment to improve dehydration characteristics of plant based foods. Trends Food Sci Technol 12:285–295. https://doi.org/10.1016/S0924-2244(01)00095-4
Fincan M, Dejmek P (2002) In situ visualization of the effect of a pulsed electric field on plant tissue. J Food Eng 55:223–230. https://doi.org/10.1016/S0260-8774(02)00079-1
Lebovka NI, Praporscic I, Vorobiev E (2004) Effect of moderate thermal and pulsed electric field treatments on textural properties of carrots, potatoes and apples. Innov Food Sci Emerg Technol 5:9–16. https://doi.org/10.1016/j.ifset.2003.12.001
Medina-Meza IG, Barbosa-Cánovas GV (2015) Assisted extraction of bioactive compounds from plum and grape peels by ultrasonics and pulsed electric fields. J Food Eng 166:268–275. https://doi.org/10.1016/j.jfoodeng.2015.06.012
Corrales M, Toepfl S, Butz P et al (2008) Extraction of anthocyanins from grape by-products assisted by ultrasonics, high hydrostatic pressure or pulsed electric fields: a comparison. Innov Food Sci Emerg Technol 9:85–91. https://doi.org/10.1016/j.ifset.2007.06.002
López N, Puértolas E, Condón S et al (2008) Effects of pulsed electric fields on the extraction of phenolic compounds during the fermentation of must of Tempranillo grapes. Innov Food Sci Emerg Technol 9:477–482. https://doi.org/10.1016/j.ifset.2007.11.001
Sun Y, Liao X, Wang Z et al (2007) Optimization of microwave-assisted extraction of anthocyanins in red raspberries and identification of anthocyanin of extracts using high-performance liquid chromatography – mass spectrometry. Eur Food Res Technol 225:511–523. https://doi.org/10.1007/s00217-006-0447-1
Zheng X, Wang X, Lan Y et al (2009) Application of response surface methodology to optimize microwave-assisted extraction of silymarin from milk thistle seeds. Sep Purif Technol 70:34–40. https://doi.org/10.1016/j.seppur.2009.08.008
Pan Y, He C, Wang H et al (2010) Antioxidant activity of microwave-assisted extract of Buddleia officinalis and its major active component. Food Chem 121:497–502. https://doi.org/10.1016/j.foodchem.2009.12.072
Xue H, Xu H, Wang X et al (2018) Effects of microwave power on extraction kinetic of anthocyanin from blueberry powder considering absorption of microwave energy. J Food Qual 2018:e9680184. https://doi.org/10.1155/2018/9680184
Brzezinski LJ, Rafel S, Leahy JW (1997) The asymmetric Baylis-Hillman reaction as a template in organic synthesis. Tetrahedron 53:16423–16434. https://doi.org/10.1016/S0040-4020(97)01025-9
Beijersbergen JCM, Lemmers CBG (1972) Enzymic and non-enzymic liberation of tulipalin A (α-methylene butyrolactone) in extracts of tulip. Physiol Plant Pathol 2:265–270. https://doi.org/10.1016/0048-4059(72)90009-4
van Rossum MWPC, Alberda M, van der Plas LHW (1998) Tulipaline and tuliposide in cultured explants of tulip bulb scales. Phytochemistry 49:723–729. https://doi.org/10.1016/S0031-9422(98)00199-X
McGovern R, Elmer W (2017) Diseases of tulip. In: McGovern, R., Elmer, W. (eds) Handbook of Florists' Crops Diseases. Handbook of Plant Disease Management. Springer, Cham. https://doi.org/10.1007/978-3-319-32374-9_49-1
Gil-Martín E, Forbes-Hernández T, Romero A et al (2022) Influence of the extraction method on the recovery of bioactive phenolic compounds from food industry by-products. Food Chem 378:131918. https://doi.org/10.1016/j.foodchem.2021.131918
Spigno G, Tramelli L, De Faveri DM (2007) Effects of extraction time, temperature and solvent on concentration and antioxidant activity of grape marc phenolics. J Food Eng 81:200–208. https://doi.org/10.1016/j.jfoodeng.2006.10.021
Sood A, Gupta M (2015) Extraction process optimization for bioactive compounds in pomegranate peel. Food Biosci 12:100–106. https://doi.org/10.1016/j.fbio.2015.09.004
Silva EM, Rogez H, Larondelle Y (2007) Optimization of extraction of phenolics from Inga edulis leaves using response surface methodology. Sep Purif Technol 55:381–387. https://doi.org/10.1016/j.seppur.2007.01.008
Antony A, Farid M (2022) Effect of temperatures on polyphenols during extraction. Appl Sci 12:2107. https://doi.org/10.3390/app12042107
Mahmud I, Mirghani MES, Yusof F, Al-khatib M (2019) Effects of time, temperature, and solvent ratio on the extraction of non-extractable polyphenols with anticancer activity of Barhi date palm kernels extracts using response surface methodology. Preprints 2019, 2019070055. https://doi.org/10.20944/preprints201907.0055.v1
Thoo YY, Ho SK, Liang JY et al (2010) Effects of binary solvent extraction system, extraction time and extraction temperature on phenolic antioxidants and antioxidant capacity from mengkudu (Morinda citrifolia). Food Chem 120:290–295. https://doi.org/10.1016/j.foodchem.2009.09.064
Wang Y, Xi G-S, Zheng Y-C, Miao F-S (2010) Microwave-assisted extraction of flavonoids from Chinese herb Radix Puerariae (Ge Gen). J Med Plants Res 4:304–308. https://doi.org/10.5897/JMPR09.426
Chen F, Sun Y, Zhao G et al (2007) Optimization of ultrasound-assisted extraction of anthocyanins in red raspberries and identification of anthocyanins in extract using high-performance liquid chromatography–mass spectrometry. Ultrason Sonochem 14:767–778. https://doi.org/10.1016/j.ultsonch.2006.12.011
Wu X, Yu X, Jing H (2011) Optimization of phenolic antioxidant extraction from Wuweizi (Schisandra chinensis) pulp using random-centroid optimazation methodology. Int J Mol Sci 12:6255–6266. https://doi.org/10.3390/ijms12096255
Dyrby M, Westergaard N, Stapelfeldt H (2001) Light and heat sensitivity of red cabbage extract in soft drink model systems. Food Chem 72:431–437. https://doi.org/10.1016/S0308-8146(00)00251-X
Rein M (2005) Copigmentation reactions and color stability of berry anthocyanins. Helsingin Yliopisto
Ursu MS, Aprodu I, Milea Ștefania A et al (2020) Thermal degradation kinetics of anthocyanins extracted from purple maize flour extract and the effect of heating on selected biological functionality. Foods 9:1593. https://doi.org/10.3390/foods9111593
Nayak B, Berrios JDJ, Powers JR, Tang J (2011) Thermal degradation of anthocyanins from purple potato (Cv. Purple Majesty) and impact on antioxidant capacity. J Agric Food Chem 59:11040–11049. https://doi.org/10.1021/jf201923a
Mori K, Goto-Yamamoto N, Kitayama M, Hashizume K (2007) Loss of anthocyanins in red-wine grape under high temperature. J Exp Bot 58:1935–1945. https://doi.org/10.1093/jxb/erm055
Vergara-Salinas JR, Bulnes P, Zúñiga MC et al (2013) Effect of pressurized hot water extraction on antioxidants from grape pomace before and after enological fermentation. J Agric Food Chem 61:6929–6936. https://doi.org/10.1021/jf4010143
Iylia Arina MZ, Harisun Y (2019) Effect of extraction temperatures on tannin content and antioxidant activity of Quercus infectoria (Manjakani). Biocatal Agric Biotechnol 19:101104. https://doi.org/10.1016/j.bcab.2019.101104
Max B, Salgado JM, Cortés S, Domínguez JM (2010) Extraction of phenolic acids by alkaline hydrolysis from the solid residue obtained after prehydrolysis of trimming vine shoots. J Agric Food Chem 58:1909–1917. https://doi.org/10.1021/jf903441d
Rajha HN, Boussetta N, Louka N et al (2014) A comparative study of physical pretreatments for the extraction of polyphenols and proteins from vine shoots. Food Res Int 65:462–468. https://doi.org/10.1016/j.foodres.2014.04.024
Corbin C, Fidel T, Leclerc EA et al (2015) Development and validation of an efficient ultrasound assisted extraction of phenolic compounds from flax (Linum usitatissimum L.) seeds. Ultrason Sonochem 26:176–185. https://doi.org/10.1016/j.ultsonch.2015.02.008
Andersen OM, Markham KR (2005) Flavonoids: chemistry, biochemistry and applications. CRC Press, Boca Raton
Alappat B, Alappat J (2020) Anthocyanin pigments: beyond aesthetics. Molecules 25:5500. https://doi.org/10.3390/molecules25235500
Cacace JE, Mazza G (2002) Extraction of anthocyanins and other phenolics from black currants with sulfured water. J Agric Food Chem 50:5939–5946. https://doi.org/10.1021/jf025614x
Tanaka Y, Sasaki N, Ohmiya A (2008) Biosynthesis of plant pigments: anthocyanins, betalains and carotenoids. Plant J 54:733–749. https://doi.org/10.1111/j.1365-313X.2008.03447.x
Yeop A, Sandanasam J, Pan SF et al (2017) The effect of particle size and solvent type on the gallic acid yield obtained from Labisia pumila by ultrasonic extraction. MATEC Web Conf 111:02008. https://doi.org/10.1051/matecconf/201711102008
Li H, Guo H, Luo Q et al (2021) Current extraction, purification, and identification techniques of tea polyphenols: an updated review. Crit Rev Food Sci Nutr 0:1–19. https://doi.org/10.1080/10408398.2021.1995843
Abascal K, Ganora L, Yarnell E (2005) The effect of freeze-drying and its implications for botanical medicine: a review. Phytother Res 19:655–660. https://doi.org/10.1002/ptr.1651
Gião MS, Pereira CI, Fonseca SC et al (2009) Effect of particle size upon the extent of extraction of antioxidant power from the plants Agrimonia eupatoria, Salvia sp. and Satureja montana. Food Chem 117:412–416. https://doi.org/10.1016/j.foodchem.2009.04.020
Bucić-Kojić A, Planinić M, Tomas S et al (2007) Study of solid–liquid extraction kinetics of total polyphenols from grape seeds. J Food Eng 81:236–242. https://doi.org/10.1016/j.jfoodeng.2006.10.027
Mayer-Laigle C, Rajaonarivony RK, Blanc N, Rouau X (2018) Comminution of dry lignocellulosic biomass: part II. Technologies, improvement of milling performances, and security issues. Bioengineering 5:50. https://doi.org/10.3390/bioengineering5030050
Routray W, Orsat V (2013) Preparative extraction and separation of phenolic compounds. In: Ramawat KG, Mérillon J-M (eds) Natural products: phytochemistry, botany and metabolism of alkaloids, phenolics and terpenes. Springer, Berlin, Heidelberg, pp 2013–2045
Garcia AA, Bonen MR, Ramirez-Vick J, Vuppu A (1999) Bioseparation process science. https://api.semanticscholar.org/CorpusID:92612833
Kumari VBC, Patil SM, Ramu R et al (2022) Chapter 5 - Chromatographic techniques: types, principles, and applications. In: Egbuna C, Patrick-Iwuanyanwu KC, Shah MA et al (eds) Analytical techniques in biosciences. Academic Press, pp 73–101
Srivastava N, Singh A, Kumari P, et al. (2021) Chapter 21 - Advances in extraction technologies: isolation and purification of bioactive compounds from biological materials. In: Sinha R p., Häder D-P (eds) Natural bioactive compounds. Academic Press, pp 409–433. https://doi.org/10.1016/B978-0-12-820655-3.00021-5
Skoog DA, Holler FJ, Crouch SR (2017) Principles of instrumental analysis. Cengage Learning
Martín J, Díaz-Montaña EJ, Asuero AG (2018) Recovery of anthocyanins using membrane technologies: a review. Crit Rev Anal Chem 48:143–175. https://doi.org/10.1080/10408347.2017.1411249
Díaz-Reinoso B, Moure A, Domínguez H, Parajó JC (2009) Ultra- and nanofiltration of aqueous extracts from distilled fermented grape pomace. J Food Eng 91:587–593. https://doi.org/10.1016/j.jfoodeng.2008.10.007
Li J, Chase HA (2010) Applications of membrane techniques for purification of natural products. Biotechnol Lett 32:601–608. https://doi.org/10.1007/s10529-009-0199-7
Cissé M, Vaillant F, Pallet D, Dornier M (2011) Selecting ultrafiltration and nanofiltration membranes to concentrate anthocyanins from roselle extract (Hibiscus sabdariffa L.). Food Res Int 44:2607–2614. https://doi.org/10.1016/j.foodres.2011.04.046
Cassano A, Conidi C, Ruby-Figueroa R (2014) Recovery of flavonoids from orange press liquor by an integrated membrane process. Membranes 4:509–524. https://doi.org/10.3390/membranes4030509
Conidi C, Cassano A, Caiazzo F, Drioli E (2017) Separation and purification of phenolic compounds from pomegranate juice by ultrafiltration and nanofiltration membranes. J Food Eng 195:1–13. https://doi.org/10.1016/j.jfoodeng.2016.09.017
Karakulski K, Kozlowski A, Morawski AW (1995) Purification of oily wastewater by ultrafiltration. Sep Technol 5:197–205. https://doi.org/10.1016/0956-9618(95)00123-9
Khemakhem I, Gargouri OD, Dhouib A et al (2017) Oleuropein rich extract from olive leaves by combining microfiltration, ultrafiltration and nanofiltration. Sep Purif Technol 172:310–317. https://doi.org/10.1016/j.seppur.2016.08.003
Galanakis CM (2015) Separation of functional macromolecules and micromolecules: from ultrafiltration to the border of nanofiltration. Trends Food Sci Technol 42:44–63. https://doi.org/10.1016/j.tifs.2014.11.005
KNAUER Wissenschaftliche Geräte (2023) HPLC principles and parameters. In: KNAUER. https://www.knauer.net/en/Systems-Solutions/Analytical-HPLC-UHPLC/HPLC-Basics---principles-and-parameters. Accessed 2 Mar 2023
Reuhs BL (2017) High-performance liquid chromatography. In: Nielsen SS (ed) Food analysis. Springer International Publishing, Cham, pp 213–226
Shabir GA (2010) Development and validation of a reversed-phase HPLC method for the determination of hydroxybenzene in a cream formulation. Indian J Pharm Sci 72:307–311. https://doi.org/10.4103/0250-474X.70475
García-Alvarez-Coque MC, Baeza-Baeza JJ, Ramis-Ramos G (2015) Reversed phase liquid chromatography. In: Analytical separation science. Wiley, Ltd, pp 159–198. https://doi.org/10.1002/9783527678129.assep008
Ignat I, Volf I, Popa VI (2011) A critical review of methods for characterisation of polyphenolic compounds in fruits and vegetables. Food Chem 126:1821–1835. https://doi.org/10.1016/j.foodchem.2010.12.026
Mu T-H, Sun H-N (2019) Chapter 22 - Sweet potato leaf polyphenols: preparation, individual phenolic compound composition and antioxidant activity. In: Watson RR (ed) Polyphenols in plants (second edition). Academic Press, pp 365–380. https://doi.org/10.1016/B978-0-12-813768-0.00022-0
Timothy Wehr C (1984) [6] High-performance liquid chromatography: care of columns. In: Methods in Enzymology. Academic Press, pp 133–154. https://doi.org/10.1016/S0076-6879(84)04087-8
YMC Europe GmbH (2019) YMC classic column. https://ymc.eu/files/imported/publications/282/documents/YMC_Classics_eCatalogue_0123.pdf
Ronald E. Majors (2003) The cleaning and regeneration of reversed-phase HPLC columns. LCGC North America 21
Chen T, Yang X, Wang N et al (2018) Separation of six compounds including two n-butyrophenone isomers and two stibene isomers from Rheum tanguticum Maxim by recycling high speed counter-current chromatography and preparative high-performance liquid chromatography. J Sep Sci 41:3660–3668. https://doi.org/10.1002/jssc.201800411
Xue H, Zhu X, Tan J et al (2021) Counter-current fractionation-assisted bioassay-guided separation of active compound from blueberry and the interaction between the active compound and α-glucosidase. Foods 10:509. https://doi.org/10.3390/foods10030509
Degenhardt A, Knapp H, Winterhalter P (2000) Separation and purification of anthocyanins by high-speed countercurrent chromatography and screening for antioxidant activity. J Agric Food Chem 48:338–343. https://doi.org/10.1021/jf990876t
Yanagida A, Shoji A, Shibusawa Y et al (2006) Analytical separation of tea catechins and food-related polyphenols by high-speed counter-current chromatography. J Chromatogr A 1112:195–201. https://doi.org/10.1016/j.chroma.2005.09.086
Herrero M, Ibañez E (2018) Green extraction processes, biorefineries and sustainability: recovery of high added-value products from natural sources. J Supercrit Fluids 134:252–259. https://doi.org/10.1016/j.supflu.2017.12.002
Samanta AK (2020) Bio-dyes, bio-mordants and bio-finishes: scientific analysis for their application on textiles. In: Chemistry and technology of natural and synthetic dyes and pigments. IntechOpen. https://doi.org/10.5772/intechopen.92601
Breitenbucher JG, Arienti KL, McClure KJ (2001) Scope and limitations of solid-supported liquid−liquid extraction for the high-throughput purification of compound libraries. J Comb Chem 3:528–533. https://doi.org/10.1021/cc010039f
Justino CIL, Duarte K, Freitas AC et al (2014) Chapter 3 - Classical methodologies for preparation of extracts and fractions. In: Rocha-Santos T, Duarte AC (eds) Comprehensive analytical chemistry. Elsevier, pp 35–57. https://doi.org/10.1016/B978-0-444-63359-0.00003-3
Silva S, Costa EM, Calhau C et al (2017) Anthocyanin extraction from plant tissues: a review. Crit Rev Food Sci Nutr 57:3072–3083. https://doi.org/10.1080/10408398.2015.1087963
Suwal S, Marciniak A (2018) Technologies for the extraction, separation and purification of polyphenols – a review. Nepal J Biotechnol 6:74–91. https://doi.org/10.3126/njb.v6i1.22341
Seabra IJ, Braga MEM, Batista MT, de Sousa HC (2010) Effect of solvent (CO2/ethanol/H2O) on the fractionated enhanced solvent extraction of anthocyanins from elderberry pomace. J Supercrit Fluids 2:145–152. https://doi.org/10.1016/j.supflu.2010.05.001
Vatai T, Škerget M, Knez Ž et al (2008) Extraction and formulation of anthocyanin-concentrates from grape residues. J Supercrit Fluids 45:32–36. https://doi.org/10.1016/j.supflu.2007.12.008
Golmohamadi A, Möller G, Powers J, Nindo C (2013) Effect of ultrasound frequency on antioxidant activity, total phenolic and anthocyanin content of red raspberry puree. Ultrason Sonochem 20:1316–1323. https://doi.org/10.1016/j.ultsonch.2013.01.020
Carabias-Martínez R, Rodríguez-Gonzalo E, Revilla-Ruiz P, Hernández-Méndez J (2005) Pressurized liquid extraction in the analysis of food and biological samples. J Chromatogr A 1089:1–17. https://doi.org/10.1016/j.chroma.2005.06.072
Petersson EV, Liu J, Sjöberg PJR et al (2010) Pressurized Hot Water Extraction of anthocyanins from red onion: a study on extraction and degradation rates. Anal Chim Acta 663:27–32. https://doi.org/10.1016/j.aca.2010.01.023
Armenta S, Garrigues S, de la Guardia M (2008) Green analytical chemistry. TrAC Trends Anal Chem 27:497–511. https://doi.org/10.1016/j.trac.2008.05.003
Ranjha MMAN, Kanwal R, Shafique B et al (2021) A critical review on pulsed electric field: a novel technology for the extraction of phytoconstituents. Mol Basel Switz 26:4893. https://doi.org/10.3390/molecules26164893
Nowosad K, Sujka M, Pankiewicz U, Kowalski R (2021) The application of PEF technology in food processing and human nutrition. J Food Sci Technol 58:397–411. https://doi.org/10.1007/s13197-020-04512-4
Xue H, Tan J, Li Q et al (2020) Ultrasound-assisted deep eutectic solvent extraction of anthocyanins from blueberry wine residues: optimization, identification, and HepG2 antitumor activity. Molecules 25:5456. https://doi.org/10.3390/molecules25225456
Faye A, Leung ACW, Guyot-Reeb S et al (2021) Extraction of tannins from yellow birch: enhanced process for water conservation and energy savings. J Clean Prod 299:126807. https://doi.org/10.1016/j.jclepro.2021.126807
Al-Hamimi S, Abellan Mayoral A, Cunico LP, Turner C (2016) Carbon dioxide expanded ethanol extraction: solubility and extraction kinetics of α-pinene and cis-verbenol. Anal Chem 88:4336–4345. https://doi.org/10.1021/acs.analchem.5b04534
Sánchez-Camargo AP, Montero L, Cifuentes A et al (2016) Application of Hansen solubility approach for the subcritical and supercritical selective extraction of phlorotannins from Cystoseira abies-marina. RSC Adv 6:94884–94895. https://doi.org/10.1039/C6RA16862K
Belwal T, Ezzat SM, Rastrelli L et al (2018) A critical analysis of extraction techniques used for botanicals: trends, priorities, industrial uses and optimization strategies. TrAC Trends Anal Chem 100:82–102. https://doi.org/10.1016/j.trac.2017.12.018
Tena N, Asuero AG (2022) Up-to-date analysis of the extraction methods for anthocyanins: principles of the techniques, optimization, technical progress, and industrial application. Antioxidants 11:286. https://doi.org/10.3390/antiox11020286
Perrut M (2000) Supercritical fluid applications: industrial developments and economic issues. Ind Eng Chem Res 39:4531–4535. https://doi.org/10.1021/ie000211c
Kalakul S, Malakul P, Siemanond K, Gani R (2014) Integration of life cycle assessment software with tools for economic and sustainability analyses and process simulation for sustainable process design. J Clean Prod 71:98–109. https://doi.org/10.1016/j.jclepro.2014.01.022
Funding
Open Access funding provided by LUT University (previously Lappeenranta University of Technology (LUT)). The Jane and Aatos Erkko Foundation funded this project.
Author information
Authors and Affiliations
Contributions
Each author performed sufficiently in the work. T.L.N.: writing—original draft and visualization. T.L.N., A.O., S.T.H., A.R., R.R., M.K-M., K.M.: conceptualization, methodology, investigation, writing, editing, reviewing.
Corresponding author
Ethics declarations
Ethical approval
This study complies with ethical standards for scientific research.
Competing interests
The authors declare no competing interests.
Additional information
Publisher's note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Nguyen, T.L., Ora, A., Häkkinen, S.T. et al. Innovative extraction technologies of bioactive compounds from plant by-products for textile colorants and antimicrobial agents. Biomass Conv. Bioref. (2023). https://doi.org/10.1007/s13399-023-04726-4
Received:
Revised:
Accepted:
Published:
DOI: https://doi.org/10.1007/s13399-023-04726-4