Open access peer-reviewed chapter

Antioxidant Compounds from Agro-Industrial Residue

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Beatriz Hernández-Carlos, Norma Francenia Santos-Sánchez, Raúl Salas-Coronado, Claudia Villanueva-Cañongo and Paula Cecilia Guadarrama-Mendoza

Submitted: 26 September 2018 Reviewed: 13 February 2019 Published: 20 March 2019

DOI: 10.5772/intechopen.85184

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Edited by Emad Shalaby

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Agro-industrial residues are a potential source of antioxidant compounds, which in general are phenolic compounds with a large chemical variability. The structure and the complexity of the phenolic compounds (polyphenols) determine their antioxidant capacity, pretreatments, and extraction methods. This chapter gives an overview of the chemical complexity of the phenolic compounds found in extractable and non-extractable fractions of agro-industrial residues, and representative compounds that are present in such residues are shown. Moreover, extraction methods described in this review showed the use of nonconventional technologies and chemical, enzymatic, or thermic treatments, useful to transform non-extractable polyphenols (NEP) to extractable polyphenol (EP) and then apply the EP extraction methods and recover antioxidants.


  • agro-industrial residues
  • total phenol content
  • extractable and non-extractable polyphenols

1. Introduction

The agro-industry produces a huge amount of waste, such as peels and seeds from fruits (juice industry), coffee husks, coffee pulp, spent coffee grounds, cocoa husks, cocoa bean shells, acerola bagasse, soybean expeller, rice straw, wheat straw, and sugar bagasse. Most of these wastes contain value-added substances such as phenol-type compounds, which are important for their antioxidant activity. Phenolic compounds possess an aromatic ring, bearing one or more hydroxyl substituents, and whether they have low or high molecular weights (from one to several aromatic rings), all of them are generally referred to as polyphenols. The chemical complexity of polyphenols and the ease of extraction from vegetal tissues divide them into two main groups. The first group is comprised of low-molecular-weight phenols (LMWP) such as flavonoids, hydroxycinnamic acids, stilbenes, and benzoic acids, which are found in free form or as glycosides ( Figure 1 ) [1, 2, 3]. They are easily extracted by aqueous-organic solvents which is why they are named extractable polyphenols (EP). The second group of compounds are low- or high-molecular-weight polyphenols that include (i) lignans which are phenolic acids or flavonoids associated with the cell wall, such as highly condensed phenylpropanoids [4] and (ii) tannins of high-molecular-weight polyphenols, which can be polymers of phenolic acids and sugars (hydrolysable tannins) or polymers of polyhydroxyflavan-3-ol (condensed tannins or proanthocyanidins) [5]. Due to structural complexity, low solubility, and matrix or vegetal tissue availability, these polyphenols are not easy to extract and therefore are considered non-extractable polyphenols (NEP).

Figure 1.

Extractable (EP) and non-extractable polyphenols (NEP) from ARs.

EP and NEP from agro-industrial residues (ARs) represent sources of value-added compounds with potential uses as ingredients in functional foods [6] or dietary supplements due to their health benefits, including antioxidant activity [7, 8, 9, 10].

Some problems associated with the recovery of antioxidant phenols or polyphenols are the low availability from matrix (NEP), chemical complexity (NEP), low extraction yields (NEP and EP), and the reduction of antioxidant activity during the extraction process (NEP and EP). All of them present challenges to be overcome for the best use of ARs and economic feasibility. In this review, chemical complexity, extraction methods, and antioxidant activity described in the most recent bibliography are presented. Research in the use of agro-industrial waste involves applying nonconventional extraction methods and establishing conditions that prevent the degradation of polyphenols and, consequently, the loss of antioxidant activity.

Phenolic compounds with antioxidant potential in ARs described to date can be grouped into five classes according to the number of carbon atoms in the basic skeleton: C6C1, C6C3, C6C1C6, C6C2C6, and C6C3C6 [11]. There are benzoic acids 16 (C6C1), hydroxycinnamic acids 714 (C6C3), benzophenones 1516 and xanthones 1718 (C6C1C6), stilbenes 1926 (C6C2C6), flavan-3-ols 2730, anthocyanidins 3135, flavonols 3640, flavanones 4145, flavones 4649, isoflavones 5055 and dihydrochalcone 56 (C6C3C6) Figure 2 . Table 1 shows low-molecular-weight phenolic compounds and their occurrence in ARs, mainly pomace, peels, seeds, and fibers from fruits such as acerola (Malpighia) [12], coffee [13], mandarin oranges (Citrus) [14], berries [15], yuzu (Citrus) [16], mangoes [17], apples [3], pears [3], oranges [18], jocote (Spondias purpurea L.) [19], cocoa husks [20], and grapes [21].

Figure 2.

Structures of low-molecular-weight polyphenols obtained from Ars [14, 31, 50, 51, 52, 53].

Polyphenol Example of ARs
Benzoic acids
Protocatechuic acid 1 Mandarin peels [14], grape bagasse [22], spent ground coffee grounds [23], grape pomace [24], sugarcane bagasse [25, 26]
p-Hydroxybenzoic acid 2 Mandarin peels [14]
Vanillic acid 3 Mandarin peels [14]
Gallic acid 4 Mango peels and seeds [17]
Gallic acid derivates theogallin 5 and ellagic acid 6 Orange peel [18], acerola bagasse [12], Vidal grape pomace [27], jocote [19], grape pomace [24]
Hydroxycinnamic acids and derivates
Ferulic acid 7 Coffee pulp [13], grape seed oil press residues [28], Vidal grape pomace [27], pomegranate seeds [29]
Caffeic acid 8 Grape seed oil press residues [28], grape pomace [24]
p-Coumaric acid 9 Grape seed oil press residues [28], mandarin peels [14], acerola bagasse [12], Vidal grape pomace [27]
Sinapic acid 10 Mandarin peels [14], grape pomace [24]
Caftaric acid 11 Grape seed oil press residues [28], grape pomace [24]
Fertaric acid 12 Vidal grape pomace [27], seed oil press residues [28], grape pomace [24]
Coutaric acid 13 Seed oil press residues [28], grape pomace [24]
Caffeoylquinic 14, coumaroylquinic, and feruloylquinic acids Coffee pulp [13], apple fiber [3], Saskatoon berry pomace [15], mandarin peels [14], acerola bagasse [12], green coffee seed residue [30], pear fiber [3], jocote [19]
Maclurin 15, riflophenone 16 Mango peels [17]
Mangiferin 17, isomangiferin 18 Mango peels [17, 31]
Trans-resveratrol 19 Grape pomace [24], grape skin [21], grape cane [32]
Trans-piceid 20, cis-piceid and piceatannol 21 Grape skin [21], grape cane [32]
Hopeaphenol 22 and isohopeaphenol, ampelopsin 23, miyabenol C 24, trans-ε-viniferin 25, r-2-viniferin 26, and ω-viniferin Grape cane [32]
(+)-Catechin 27 Vidal grape pomace [27], cocoa husk [20], pomegranate seeds [29]
(−)-Epicatechin 28 Vidal grape pomace [27], cocoa husk [20], acerola bagasse [12]
Procyanidins B1 29, procyanidins B2 30 Grape skins [21], grape pomace [24], Vidal grape pomace [27], pomegranate seeds [29]
Delphinidin 31 Saskatoon berry pomace [15], blueberry waste [34], grape pomace [24]
Cyanidin 32 Saskatoon berry pomace [15], grape skin [21], blueberry waste [33]
Malvidin 33 Grape skin [21], blueberry waste [33]
Peonidin 34 Grape skin [21], grape pomace [24], blueberry waste [33]
Petunidin 35 Grape pomace [24], blueberry waste [33]
Quercetin 36 and glycosides Saskatoon berry pomace [15], Vidal grape pomace [27], pear fiber [3], grape skin [21], mango peels [31], jocote peels [19], grape pomace [24], lemon pomace [34]
Kaempferol 37 and glycosides Vidal grape pomace [27], grape skin [21], mango peels [31], jocote [19], lemon pomace [35], pomegranate seeds [29]
Isorhamnetin 38, glycosides Apple fiber [3], grape skin [21]
Myricetin 39 glycosides Grape skin [21]
Rhamnetin 40 glycosides Mango peels [27], jocote peels [19]
Naringenin 41 Orange peels [18], lemon pomace [34]
Naringin 42 Orange peels [18], lemon pomace [34], yuzu peels (Citrus junos) [16]
Hesperidin 43 Orange peels [18], lemon pomace [34], yuzu peels (Citrus junos) [16]
Narirutin 44 Orange peels [18]
Hesperetin 45 Orange peels [18], lemon pomace [34]
Diosmetin 46 Orange peels [18], sugarcane bagasse [25, 26]
Tangeritin 47 Orange peels [18]
Luteolin 48 Cocoa bean shells [35]
Tricin 49 Milled rice straw extract [36], sugarcane bagasse [25, 26]
Daidzin 50, glycitin 51, daidzein 52, glycitein 53, genistein 54 Soybean okara [37]
Genistin 55 Soybean okara [37], cherry pomace [38], sugarcane bagasse [26]
Phloretin 56 and glycosides Apple fiber [3], yuzu peels (Citrus junos) [16]

Table 1.

Polyphenols described in agro-industrial residues.


2. Extractable polyphenols (EP)

The applied methodologies in the use of ARs to obtain EP depend on residue type and polyphenol stability. For example, acerola bagasse contains water [39], and if the extraction procedure is not done quickly, the residue will need to be dried to avoid microbiological contamination without affecting polyphenol stability. On the other hand, the probable water content of cocoa husk is low, and therefore, the polyphenols’ extraction procedures are direct because it is a solid residue. Therefore, drying and extraction technologies or methodologies are necessary to obtain suitable yields of EP with proven antioxidant activity. The description of the drying and extraction methodologies of the EP will focus on the work with anthocyanidins because they are unstable compounds and the conditions of drying or extraction for anthocyanins are important to avoid their decomposition.

2.1 Waste drying

Valorization studies showed acerola bagasse (Malpighia emarginata DC) is a good source of antioxidants due to total phenol content (TPC), which ranges from 0.44 to 10.82 g gallic eq/100 g dm (dry matter) [39, 40, 41], while the contents of anthocyanins ascorbic acid and proanthocyanidins are 1.002 ± 0.014, 1.002 ± 0.014, and 0.7985 ± 0.0213 g/100 g dw, respectively, and the antioxidant activity evaluation by DPPH• method showed 113.7 ± 0.4 μmol trolox/g dw. However, this residue contains water and therefore must be dried for efficient handling. The drying of the residue has been carried out by hot air (60–80 °C, 4–6 m/s). This procedure showed moderate retention of phenolic (26–31%), anthocyanins (23–36%), and proanthocyanidins (21%) compounds [41]. Better results were obtained using a roto-aerated dryer (115 °C, 2.25 m/s) with a pretreatment of the sample (sprayed with ethanol), and total phenol compounds (TPC) increased 104.6% with respect to fresh residue [42]. A more recent drying method for acerola bagasse is dehydration in a thick-layer dryer, where drying was done at low temperatures (31.7 °C, 230 min, 0.4 m/s or 60 °C, 159.3 min, 0.4 m/s), which were enough to obtain a dry residue with TPC values similar to those obtained in fresh residue (2352.4 ± 57.23 mg gallic acid/100 g dm) [42]. Drying studies of other ARs are described for grape and olive waste using thin-layer drying (air temperature 20–110 °C) [43, 44], and sustainable drying strategies such as the use of solar dryers have been described [45].

2.2 ARs extraction

Generally, EP extraction procedures are done using mixtures of water-organic solvents and assisted by microwave (MAE), heating, and ultrasound (UAE). In recent years, pressurized liquid extraction (PLE) and supercritical fluid extraction (SFE) have been applied, which could be better options because polyphenols are not exposed to severe conditions that promote degradation reactions. In addition, temperature control is a common method to assist the extraction procedures. For example, anthocyanidins, procyanidins, and flavonols were obtained from grape skins using MAE and UAE at 50 ± 5 °C. This work also demonstrated that the yields obtained by MAE with UAE were improved up to 40% (86.39–121.18 mg/100 g dm) [21]. Acid conditions have also been tested to improve the extraction of anthocyanins (anthocyanidin glycosides) from grape peels separated from red grape pomace from vintages 2001 to 2002. The extraction was made in two steps. First, the residue was macerated (2 h) with methanol/HCl 0.1 (v/v) with oxygen reduced in the mixture to dissolve polar polyphenols. Then extraction with an organic solvent was done to recover less polar polyphenols. Anthocyanin yields found were better (5967–131, 868 mg/kg dm) [24] than those obtained from grape skins from fresh fruits (1211.8 mg/kg dm) [21] due to previous thermic and enzymatic treatments during wine production, which helped release anthocyanins.

Effects of temperature, time, and solvent concentration on polyphenol extraction from grape marc showed better extraction yields with the increase of water (30–50%) in the ethanol-water mixtures, maintaining the temperature at 60 °C for shorter periods (<8 h) to avoid polyphenol degradation [46]. Similar ethanol-water mixtures (40.4 and 55.4%) were used to extract the major components of grape cane: trans-resveratrol 19, trans-ε-viniferin 25, and ferulic acid 7 ( Figure 2 ). However, a higher temperature (84 °C) was necessary to obtain the highest antioxidant activities (260.8 and 1378.7 μmol TE/g TEAC and ORAC methods) [47]. Anthocyanins extraction from grape skins, stems, and seeds was effective at 70 °C with ethanol-water mixtures, (1:1) and assisted with pulsed electric fields (PEF) (9 kV, 15 s), ultrasound (35 kHz, 70 °C, 1 h), and high hydrostatic pressurization (600 MPa, 70°C, 1 h). The extraction yields for PEF were 81 and 25% higher than those obtained by ultrasound and hydrostatic pressurization [48]. A combination of methods has also been used, such as the consecutive application of UAE (4 min, 80 °C, 20 kHz, 80 W) and SFE (8 MPa, 40 °C, CO2/ethanol) for polyphenols extraction from grape marc. The ultrasound treatment increased mass transfer and accelerated access of solvent (CO2/ethanol) to vegetal tissues, and TPC was increased 27% (2736 ± 11 mg to 3493 ± 61 GAE/100 g dw) [49]. SFE with different conditions (90% CO2, 5% ethanol, 5% water, 20 MPa, 40 °C) was more efficient than pressurized liquid extraction (PLE) in experiments with blueberry waste. Anthocyanidin yields were 808 ± 0.1 mg/100 g and 248 ± 0.2 mg/100 g for SFE and PLE (20 MPa, 50% water, pH 2, 40 °C) extractions, respectively [33]. PLE with increase of pressure and temperature and change of solvent (65 °C, 10 MPa, 75% ethanol) was shown to be more useful in the extraction of gallic acid 4 together with flavanones 4145 from orange peels. The TPC obtained was 14.9 ± 0.7 mg GAE/g dm [18].

Polyphenols different from anthocyanins, such as phenolic acids (147.4–492.7 g/kg), gallic acid 4 (93.1–353.7 mg/kg), and flavonoids (2.52–13.5 mg/kg), were obtained with acidified methanol from residues of grape (Vitis vinifera L.) seed oil production [28], while protocatechuic acid 1 and chlorogenic acid 14 were extracted with hot water (92 ± 3 °C, 2 min) [13]. Other solvents used for EP extraction are acetone/water mixtures (80%, 3 h stirring) to obtain flavonol 37, 40 glycosides and xanthones 1718 from lyophilized mango peels ( Table 1 and Figure 2 ) [31]. In general, EP extraction is made with water-organic solvents and assisted by conventional and nonconventional methods. However, when extraction conditions are severe as when heating above 70 °C, acid extraction or ultrasound exposure for long periods of time, then these conditions are also useful for obtaining NEP, where the objective is promoting the breakdown of chemical bonds.


3. Non-extractable polyphenols (NEP)

Non-extractable polyphenols (tannins and lignins) are low- or high-molecular-weight compounds associated with vegetal tissue macromolecules; therefore, they are retained in the residue matrix during the extraction process. Depending on the monomeric structures and chemical reactivity of tannins, these are grouped in condensed and hydrolysable tannins. Condensed tannins are polyhydroxyflavan-3-ols oligomers and polymers linked by carbon-carbon bonds between flavanol units. These are also known as proanthocyanidins because the butanol/HCl/heat treatment produces a red anthocyanidin [54]. Hydrolysable tannins are multiple esters of gallic acid with glucose and products of oxidative reactions, and they can be soluble (EP) or non-soluble (NEP) ( Figure 3 ) [55]. Lignin is a phenylpropanoid (C6C3) polyphenol where the monomeric units are p-coumaryl 71, coniferyl, and sinapyl. Occurrence of monomeric units in lignin varies according to the taxonomic origin of the ARs, e.g., gymnosperms or angiosperms [56].

Figure 3.

Structures of condensed tannins (proanthocyanidins), hydrolysable tannins (gallotannins), and lignin (5-hydroxyguaiacyl residue) [55, 57].

Non-extractable polyphenols are common in almost all ARs and represent significant polyphenol percentages of total phenol content (TPC) in vegetal tissues. For example, NEP quantification in peels from apple, banana, kiwi, mandarin, mango, nectarine, orange, pear, and watermelon showed that, of the total phenols found, 7–82% correspond to NEP [58]. Currently research in appropriate methodologies for the extraction of NEP is a priority topic because of the economic advantages of the use of ARs. Some examples of research on the best conditions for the extraction of NEP from ARs are those for the use of cocoa by-products, which involve extractions assisted by ultrasound [20, 35], thermic treatment [59, 60], hydrodynamic cavitation [35], pressurized liquids [50], pulsed electric field [61], subcritical water hydrolysis [62], and solid fermentation [63]. There are also reports on detailed chemical studies of the NEP structure thanks to modern analytical instrumentation, such as liquid chromatography (LC) coupled to matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS), electrospray/ionization time-of-flight mass spectrometry (ESI-TOF-MS), LC × LC coupled to tandem mass spectrometry, pyrolysis/gas chromatography/mass spectrometry (Py/GC/MS), and nuclear magnetic resonance (NMR), which has been key to making detailed chemical studies of high-molecular-weight polyphenols [36, 64, 65, 66]. In the following paragraphs, some examples of chemical studies of NEP are presented to give an overview of the structural complexity that exists in them.

3.1 Tannins

Studies in pomegranate by-products led to the identification of several polyphenols. EP was extracted with acetone 70% (ultrasound-assisted 20 min, 30 °C), and NEP was previously subjected to basic hydrolysis of insoluble residues before extraction under the same conditions. Ellagic acid 6 and monogalloyl-hexoside 57 were the main compounds, in addition to ellagic acid derivates, valoneic acid bilactone I and II 58, punicalagin 59 and isomers, trigalloylglucopiranose I and II, and granatin, among 34 hydrolysable tannins described in Figure 4 . Moreover, condensed tannins such as procyanidin dimers 30 and gallocatechin (or catechin) hexoside were described. Antioxidant activity for polyphenol (EP and NEP) fractions was 0.12–3.58 mmol TE/g dm (DPPH) and 5.34–208.73 μmol TE/g dm (ABTS radical-scavenging activity) [66]. Oligomeric proanthocyanidins such as dimer, trimer, tetramer, pentamer, and hexamer units (MW 600, 889, 1177, 1465, and 1753 amu) were identified in coffee pulp by MALDI-TOF-MS analysis. Extraction of these tannins was made with acid aqueous acetone [67]. Acetone/water also was used to extract anthocyanidins from litchi pericarp (Litchi chinensis), after its incubation with Aspergillus awamori. Total extractable tannin recovery was increased up to 59%, and ESI-TOF-MS analysis revealed A. awamori degraded B-type condensed tannin and showed low capacity to degrade the condensed tannin A-type. Differences between both compound types are C4-C8 couple C2-O-C7´ linkages for A-type tannin, while C4-C8 linkages are found in B-type tannin ( Figure 5 ) [68].

Figure 4.

Hydrolysable tannins identified in pomegranate by-products [55, 69].

Figure 5.

Condensed A- and B-type condensed tannins. A-type C4-C8 couple C2-O-C7 linkage and B-type C4-C8 linkage [68].

3.2 Lignin

Studies of chemical composition of NEP corresponding to lignin were made for that recovered from black liquor (BL) and milled rice straw extract (RSE). BL is produced from basic hydrolysis/heat of rice straw (straw alkali oxygen cooking). Lignin and phenolics from rice straw were obtained by Soxhlet extraction (ethanol/benzene 1:2, v/v, 8 h). Detailed studies by infrared and nuclear magnetic resonance spectroscopy showed a number of residues from rice straw lignin, including β-O-4’ethers 60, β-O-4’ethers with acylated γ-OH 61, phenylcoumarans 62, resinols 63, dibenzodioxocyns 64, α,β-diaryl ethers 65, tricin units 66, Cα-oxidized guaiacyl unit 67, syringyl units 68, Cα-oxidized syringyl 69, p-hydroxyphenyl units 70, p-coumaroyl 71, guaiacyl units 72, feruloyl 73, cinnamyl alcohol end-groups 74, and cinnamyl aldehyde end-groups 75 ( Figure 6 ). Antioxidant evaluation (DPPH• and ABTS• methods) showed lignin from BL had a better radical-scavenging ability than RSE, which was due to the release of p-hydroxyphenyl units by rice straw alkali oxygen cooking. This process caused the destruction of tricin [70]. Lignin and phenolics could also be extracted from milled rice straw by hydrothermal treatment (210 °C) to release cellulosic components, delignification (ethanol/water 60.5%, 130 °C) to fractionate the lignocellulosic biomass and separation (nanofiltration cutoff 280 Da) to isolate the polyphenols [71].

Figure 6.

Residues identified in lignin from rice straw [70].

Extraction of NEP from residues that come from industrial processes has been done using methodologies applied to EP because they were released by the industrial process involved. For example, polyphenolics from sugarcane bagasse were extracted with 95% ethanol (maceration 7d) and p-coumaric acid; tricin 66, luteolin, tricin 7-O-β-glucopiranoside, diosmetin 46, 6-C-glucoside, and protocatehuic acid 1 ( Figure 2 ) were isolated from ethanol extracts (ethanol 95%, 7d, maceration). TPC from extract was 3.0 mg GAE/100 mg dm [25]. Antioxidant activities (ORAC assay) of tricin 49, p-coumaric 9, and protocatechuic 1 acids were 9.76 ± 1.01, 7.22 ± 0.73, and 6.40 ± 0.62 μmol TE/μmol, respectively. Phenols yields were from <0.2 to 2.12 mg/kg dm. Other compounds reported in this residue were genistein 52 (15.22 ± 1.28 mg/g), genistein 53 (trace), and quercetin 36 (9.98 0.40 mg/g DW) [26]. Polyphenol recovery was improved (117.1%) when residue was treated with glacial acetic acid and hydrogen peroxide at 60 °C for 7 h and later subjected to hydrolysis with xylanase (Clostridium thermocellum ATCC 27405). The antioxidant activity was increased 73% (695.8 ± 105.3 μmol trolox eq./L) [72].

Other residue that has undergone industrial processes enough to release polyphenols is the lignin from the ozone, soaking aqueous ammonia pretreatment of wheat. Py/GC/MS analysis showed the presence of 17 phenolic compounds derived from guaiacyl 72, syringyl 68, and p-hydroxyphenyl 70 units in ratios of 65.09, 23.36, and 11.5%, respectively, the main compounds being Phenol 2-methoxy (guaiacol) 76, phenol 2-methoxy-4-vinyl (4-vinylguaiacol) 77, and 2,6-dimethoxy phenol (syringol) 78. The residue was identified as a potential source of antioxidants because it showed 86.9 ± 0.34% of inhibition of DPPH radicals similar to that of commercial BHT 103.3 ± 1% [36].


4. Antioxidant activity (AA) and extraction methods

Antioxidant activity (AA) obtained from AR’s extracts varies according to the residue type and the extraction method. In case of cocoa residues, UAE (methanol/water 1:1 and acetone water 7:3) showed better method than maceration because AA was improved 8.8% (EC50 0.0486 ± 0.0018 mg/mL) in comparison with that obtained from maceration with the same solvents (EC50 0.0533 ± 0.0022 mg/mL), while total phenol content (TPC) was 41.96% higher ( Table 2 ) [20]. Extracts with high TPC (55 mg GAE/g) but low AA values (EC50 8.18 mg/mL) were obtained by hydrothermal treatment (170 °C, 30 min) [60], in comparison with those values for extracts obtained by UAE and maceration. Thermic treatment improves yields of polyphenols because it promotes release of NEP, but polyphenols can loss their antioxidant capacity. A method that significantly improved the TPC without reducing the AA of the extracts is the hydrodynamic cavitation (HC), which was used to assist polyphenols extraction from cocoa bean shells with hexane/ethanol/water mixtures [35]. The authors compared the efficiency of this method with UAE (ethanol/water); AA were similar (EC50 62 ± 3.2 and 66.9 ± 2.4 μg/mL), while TPC was 125 and 197.4 mg GAE/g for extracts obtained by UAE and HC, respectively.

Residue/ref. Extraction method Antioxidant activity TPC/(dry matter)
Cocoa husk [20] UAE 25 kHz, 30 min, MeOH/H2O 1:1, and acetone/water 7:3 EC50 0.0486 ± 0.0018 mg/mL (DPPH) 25.34 ± 1.82 mg GAE/g
Maceration MeOH/H2O 1:1, 2 h stir, acetone/water 7:3 EC50 0.0533 ± 0.0022 mg/mL (DPPH)
Ascorbic acid 0.0243 ± 0.0009 mg/mL
17.85 ± 1.33 mg GAE/g
Cocoa husk [60] Hydrothermal, 170 °C, 30 min EC50 8.18 mg/mL (DPPH) 55 mg GAE/g
Cocoa bean shells [61] Extraction EtOH/water rotatory agitation 25 °C with a pretreatment time of 11.99 μs, number of pulses of 991.28, PEF strength of 1.74 kV cm−1, ethanol 39.15%, 118.54 min 101.1–321.97 μM TE/g (DPPH) 17.88–55.16 mg GAE/g
Cocoa bean shells [35] UAE EtOH/water (70:30), 15 min, 150 W, 19.9 kHz, 40 °C EC50 66.9 ± 2.4 μg/mL and 235.3 ± 8.4 μM TE/g (DPPH) 125 mg GAE/g
Cocoa bean shells [50] PLE EtOH, 10.35 MPa, 90 °C, 30 min 65 ± 2 μM TE/g (DPPH)
84 ± 4 μM TE/g (FRAP)
10 ± 0.3 mg ECE/g
Cocoa bean shells [35] Hydrodynamic cavitation and Hex/EtOH/H2O mixtures (30:49:21) scale-up reactor EC50 62.0 ± 3.1 μg/mL (DPPH) and 256.7 ± 9.9 μM TE/g (DPPH) 197.4 mg GAE/g
Cocoa bean shells [63] Solid state fermentation with Penicillium roqueforti and EtOH/water extraction 81.3% inhibition (DPPH)
23.2 μM ferrous sulfate/g (FRAP)
926.6 ± 61 mg GAE/100 g
Acerola bagasse [12] MeOH 50%, 80 °C, 15 min 405.11 ± 1.83 μM TE/Lg
(Rutin 1473.07 ± 21.39 μM TE/Lg) (ABTS)
Acerola bagasse [41] Water and stirring 30 min 21.7–24.0 μM TE/g (DPPH) 2710.2–3171.9 mg GAE/100 g
Coffee pulp [13] Water, 92 ± 3 °C, 2 min EC50 18–27 μg/mL (ABTS)
82–153 μg/mL (DPPH)
7.61–17.40 mg GAE/L
Coffee husk [23] SFE 200 bar/323.15 K, CO2 + 8% EtOH EC50 630 μg/mL (DPPH)
141 ± 1 μM TE/g (ABTS)
36 ± 1 mg CAE/g
UAE, EtOH EC50 235.1 μg/mL (DPPH)
161 ± 3 μM TE/g (ABTS)
133.3 ± 0.6 mg CAE/g
Spent coffee grounds [23] SFE 200 bar/323.15 K, CO2 + 4% EtOH EC50 516.2 μg/mL (DPPH)
169 ± 3 μM TE/g (ABTS)
57 ± 3 mg CAE/g
Soxhlet extraction, EtOAc EC50 202.23 μg/mL (DPPH)
160.13 ± 13 μM TE/g (ABTS)
182.6 ± 28.2 mg CAE/g
Coffee silver skin [1] Hydroalcoholic solvent (50%) at 40°C, 60 min 326.0 ± 5.7 mg TE/L (DPPH)
1791.9 ± 126.3 mg FSE/L (FRAP)
302.5 ± 7.1 mg GAE/L
Spent coffee grounds [73] Solid state fermentation with Bacillus clausii (37 °C, 39 h), defatted (hexane) and EtOH/water (80:20) extraction (orbital shaker, 30 °C, 50 rpm, 3 h) 17.894 μM TE/100 g (ABTS) 1051 mg GAE/100 g
Increased 36%
Blueberry waste [33] SFE 90% CO2, 5% H2O, 5% EtOH, 20 MPa 1658 ± 160 μM TE/g (DPPH)
199 ± 20 μM TE/g (ABTS)
134 ± 11 mg GAE/g
Wet matter
PLE 40 °C, 20 MPa, 15 min, 5 mL cell, EtOH/water 1:1 1746 ± 71 μM TE/g (DPPH)
66 ± 1 μM TEAC/g (ABTS)
90 ± 2 mg GAE/g
Wet matter
Grape cane [47] Water/EtOH 40.4 and 55.4%, 84 °C 238.6 μM TE/g (ABTS)
1259.6 μM TE/g (ORAC)
8.93 mg resveratrol eq./g
Grape cane [32] Acetone/water 6:4, room temperature 1700–5300 μM TE/g (ORAC) Stilbene total content 2.62–3.30 mg/g
Grape skins [21] MAE, 600 W, 2450 MHz, 50 ± 5 °C, water/EtOH/phosphoric acid 50:50:1 nd Stilbenes 1.5 mg/100 g
UAE, 130 W, 40 kHz, 50 ± 5°C, water/EtOH/phosphoric acid 50:50:1 nd Stilbenes 0.71 mg/100 g
Soybean okara [37] Solid state fermentation with Saccharomyces cerevisiae r. f. bayanus 72 h. Water/MeOH (80%) extraction 24.04 mM TE/g (DPPH)
20.65 mM TE/g (ABTS)
Increase 15%
116 mg GAE/10 g
UAE, ultrasound-assisted extraction; EC50, effective concentration at 50%; PLE, pressurized liquid extraction; PEF, pulsed electric field; SFE, supercritical fluid extraction; TPC, total phenol content; MeOH, methanol; EtOH, ethanol; AcOEt, ethyl acetate; CAE, chlorogenic acid equivalent; GAE, gallic acid equivalent; ABTS, 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt; DPPH, 2,2-diphenyl-1-picrylhydrazyl; ORAC, oxygen radical absorbance capacity; MAE, microwave-assisted extraction; TE, trolox equivalent; FRAP, ferric-reducing antioxidant power; FSE, ferrous sulfate equivalent; nd, not described

Table 2.

Antioxidant activity and extraction methods in agro-industrial residues.

In general, SFE (with the optimal conditions) is an extraction method more convenient in order to avoid degradation reactions and therefore reduces the AA of extracts. Blueberry waste extracts were obtained by SFE (90% CO2, 5% H2O, 5% ethanol, 20 MPa) and PLE (40 °C, 20 MPa, 15 min). Both methods showed extracts with similar AA (1658 ± 160 and 1746 ± 71 μM TE/g), but SFE yields extracts with TPC 48% more higher (134 ± 11 mg GAE/g) than those from PLE (90.2 ± 2 mg GAE/g) ( Table 2 ) [33]. However, extracts obtained by UAE (ethanol) and Soxhlet (ethyl acetate) from coffee residues showed better AA values (EC50 235.1 and 202.23 μg/mL) than those observed in the extracts obtained by SFE (200 bar/323.15 K, CO2 + 8 or 4% ethanol) (EC50 630 and 516.2 μg/mL) [23].

Pretreatments as solid-state fermentation before polyphenolic extraction have shown effects on TPC and AA, for example, in spent coffee grounds, an increase in TPC and AA of 36%, and 15% were observed in fermented extracts by Bacillus clausii followed by ethanol/water extraction [73]. The same increase in AA was observed when soybean residues were subjected to solid-state fermentation using Saccharomyces cerevisiae (20.39–24.04 mM TE/g) [37]. However, the use of Penicillium roqueforti in this fermentation type with cocoa shells showed a weak increase of AA from 79.2 to 81.3% (2.6%) and the reduction of TPC from 2120 ± 20 mg GA/100 g to 926.6 ± 61 mg/100 g (56%) [63].

Grape cane residues are rich in stilbene compounds, which can be extracted with mixtures of water/ethanol or acetone/water, and their antioxidant activities depend on stilbene type present in extracts, e.g., quantitative structure-antioxidant activity relationship studies showed structural facts as planar geometry of trans-isomers has direct relation with the AA because polyphenols increase their free radical-stabilizing properties [74]. This may explain why AA is lower in extracts with higher stilbene contents; water/ethanol extracts with 8.93 mg resveratrol eq./g showed an AA of 1259.6 μM TE/g (ORAC), while acetone/water extracts with stilbene total content of 2.62–3.30 mg/g showed from 1700 to 5300 μM TE/g. Therefore, stilbene extraction accelerated by temperature can have consequences on stereoisomer content and therefore on the AA. Extraction of grape skins at 50 °C assisted by MAE and UAE showed better stilbenes contents for MAE (1.5 mg/100 g), but authors did not describe the AA [21]. Heat resistance phenol compounds and with significant AA are those found in residues previously treated to release NEP such as 56, 76-79 [36, 75] ( Figures 2 and 7 ). For example, apple pulp was subjected to reflux with water for 2 h, and the EtOAc extract showed an AA of IC50 10.59 ± 2.77 μg/mL (DPPH), while 5-hydroxymaltol 79, isolated from AcOEt extract, showed an IC50 value of 8.22 ± 1.83 μg/mL, which was 48 times higher than α-tocopherol (IC50 0.17 ± 0.04 μg/mL) [75].

Figure 7.

Heat resistance phenol compounds identified in ARs [36, 75].


5. Conclusions

Most of the bibliography related to the study of waste is focused on the search for conditions for the greater extraction of polyphenols from ARs and evaluating the feasibility of using these residues as a source of antioxidants. To evaluate the extraction efficiency of the proposed methods, the content of total phenols (TCP), the quantification and/or identification of specific polyphenols and determination of AA have been described. Antioxidant activity of polyphenols varies mainly by the temperature, which could promote the compound degradation or only small structural changes, mainly with anthocyanidins and stilbenes. Extraction methods applied to ARs described in this review showed the use of nonconventional technologies such as SFE and LPE for EP extraction while chemical, enzymatic, or thermic hydrolysis has been used to transform NEP to EP to apply the EP extraction methods and recover antioxidants. Moreover, significant contributions to the knowledge of the chemistry of ARs are summarized and representative compounds are shown that cover most types of phenols that exist in the plant kingdom and that are present in such residues. The chemical structures of 79 low-molecular-weight compounds, mainly EP and some examples of tannin and lignin residues, are described. Therefore, the use of ARs to recover polyphenols is growing due to the knowledge of ARs chemistry and to the development of nonconventional extraction methods and more efficient dry methods.



The authors thank Carol Ann Hayenga for her assistance with English language in the preparation of this manuscript. The Technological University of the Mixteca provided support.


Conflict of interest

The authors declare that there are no conflicts of interests regarding the publication of this chapter.


  1. 1. Costa AS, Alves RC, Vinha AF, Barreira SV, Nunes MA, Cunha LM, et al. Optimization of antioxidants extraction from coffee silver skin, a roasting by-product, having in view a sustainable process. Industrial Crops and Products. 2014;53:350-357. DOI: 10.1016/j.indcrop.2014.01.006
  2. 2. Peralbo-Molina A, Priego-Capote F, Luque de Castro MD. Tentative identification of phenolic compounds in olive pomace extracts using liquid chromatography-tandem mass spectrometry with a quadrupole-quadrupole-time-of-flight mass detector. Journal of Agricultural and Food Chemistry. 2012;60:11542-11550. DOI: 10.1021/jf302896m
  3. 3. Delpino-Rius A, Eras J, Vilaró F, Cubero MÁ, Balcells M, Canela-Garayoa R. Characterization of phenolic compounds in processed fibers from the juice industry. Food Chemistry. 2015;172:575-584. DOI: 10.1016/j.foodchem.2014.09.071
  4. 4. Lapierre C. Application of new methods for the investigation of lignin structure. In: Forage Cell Wall Structure and Digestibility, (Forage Cell Walls). Madison, WI: ASA, CSSA, SSSA; 1993. pp. 133-166. DOI: 10.2134/1993.foragecellwall.c6
  5. 5. Pérez-Jiménez J, Saura-Calixto F. Fruit peels as sources of non-extractable polyphenols or macromolecular antioxidants: Analysis and nutritional implications. Food Research International. 2018;111:148-152. DOI: 10.1016/j.foodres.2018.05.023
  6. 6. Martirosyan DM, Singh J. A new definition of functional food by FFC: What makes a new definition unique? Functional Foods in Health and Disease. 2015;5:209-223. DOI: 10.31989/ffhd.v5i6.183
  7. 7. Frémont L. Biological effects of resveratrol. Life Sciences. 2000;66:663-673. DOI: 10.1016/S0024-3205(99)00410-5
  8. 8. Kong JM, Chia LS, Goh NK, Chia TF, Brouillard R. Analysis and biological activities of anthocyanins. Phytochemistry. 2003;64:923-933. DOI: 10.1016/S0031-9422(03)00438-2
  9. 9. Macrae WD, Towers GN. Biological activities of lignans. Phytochemistry. 1984;23:1207-1220. DOI: 10.1016/S0031-9422(00)80428-8
  10. 10. Lee S, Monnappa AK, Mitchell RJ. Biological activities of lignin hydrolysate-related compounds. BMB Reports. 2012;45:265-274. DOI: 10.5483/BMBRep.2012.45.5.265
  11. 11. Mann J, Davison RS, Hobbs JB, Benthorpe DV, Harbone JB. Natural Products. 1st ed. Longman: Addison Wesley; 1995. p. 455
  12. 12. Rezende Marques T, Aparecida Caetano A, Avelar Rodrigues LM, Assaid Simão A, Andrade Machado GH, Duarte Corrêa A. Characterization of phenolic compounds, antioxidant and antibacterial potential the extract of acerola bagasse flour. Acta Scientiarum. Technology. 2017;39:143-148. DOI: 10.4025/actascitechnol.v39i2.28410
  13. 13. Duangjai A, Suphrom N, Wungrath J, Ontawong A, Nuengchamnong N, Yosboonruang A. Comparison of antioxidant, antimicrobial activities and chemical profiles of three coffee (Coffea arabica L.) pulp aqueous extracts. Integrative Medicine Research. 2016;5:324-331. DOI: 10.1016/j.imr.2016.09.001
  14. 14. Ma YQ, Chen JC, Liu DH, Ye XQ. Simultaneous extraction of phenolic compounds of citrus peel extracts: Effect of ultrasound. Ultrasonics Sonochemistry. 2009;16:57-62. DOI: 10.1016/j.ultsonch.2008.04.012
  15. 15. Li R, Hettiarachchy N, Rayaprolu S, Eswaranandam S, Howe B, Davis M, et al. Phenolics and antioxidant activity of Saskatoon berry (Amelanchier alnifolia) pomace extract. Journal of Medicinal Food. 2014;17:384-392. DOI: 10.1089/jmf.2012.0278
  16. 16. Assefa AD, Saini RK, Keum YS. Extraction of antioxidants and flavonoids from yuzu (Citrus junos Sieb ex Tanaka) peels: A response surface methodology study. Journal of Food Measurement and Characterization. 2017;11:364-379. DOI: 10.1007/s11694-016-9405-1
  17. 17. Dorta E, González LMG, Sánchez-Moreno C, de Ancos B. Screening of phenolic compounds in by-product extracts from mangoes (Mangifera indica L.) by HPLC-ESI-QTOF-MS and multivariate analysis for use as a food ingredient. Food Research International. 2014;57:51-60. DOI: 10.1016/j.foodres.2014.01.012
  18. 18. Barrales FM, Silveira BPDPM, Ruviaro AR, Paulino BN, Pastore GM, Martinez J. Recovery of phenolic compounds from citrus by-products using pressurized liquids—An application to orange peel. Food and Bioproducts Processing. 2018;112:9-21. DOI: 10.1016/j.fbp.2018.08.006
  19. 19. Engels C, Gräter D, Esquivel P, Jiménez VM, Gänzle MG, Schieber A. Characterization of phenolic compounds in jocote (Spondias purpurea L.) peels by ultra-high-performance liquid chromatography/electrospray ionization mass spectrometry. Food Research International. 2012;46:557-562. DOI: 10.1016/j.foodres.2011.04.003
  20. 20. Quiroz-Reyes CN, Aguilar-Méndez MA, Ramírez-Ortíz ME, Ronquillo-De Jesús E. Comparative study of ultrasound and maceration techniques for the extraction of polyphenols from cocoa beans (Theobroma cacao L.). Revista Mexicana de Ingeniería Químic. 2013;12:11-18
  21. 21. Crupi P, Dipalmo T, Clodoveo ML, Toci AT, Coletta A. Seedless table grape residues as a source of polyphenols: Comparison and optimization of non-conventional extraction techniques. European Food Research and Technology. 2018;244:1091-1100. DOI: 10.1007/s00217-017-3030-z
  22. 22. Farías-Campomanes AM, Rostagno MA, Meirele MAA. Production of polyphenol extracts from grape bagasse using supercritical fluids: Yield, extract composition and economic evaluation. The Journal of Supercritical Fluids. 2013;77:70-78. DOI: 10.1016/j.supflu.2013.02.006
  23. 23. Andrade KS, Gonçalvez RT, Maraschin M, Ribeiro-do-Valle RM, Martínez J, Ferreira SR. Supercritical fluid extraction from spent coffee grounds and coffee husks: Antioxidant activity and effect of operational variables on extract composition. Talanta. 2012;88:544-552. DOI: 10.1016/j.talanta.2011.11.031
  24. 24. Kammerer D, Claus A, Carle R, Schieber A. Polyphenol screening of pomace from red and white grape varieties (Vitis vinifera L.) by HPLC-DAD-MS/MS. Journal of Agricultural and Food Chemistry. 2004;52:4360-4367. DOI: 10.1021/jf049613b
  25. 25. Zheng R, Su S, Li J, Zhao Z, Wei J, Fu X, et al. Recovery of phenolics from the ethanolic extract of sugarcane (Saccharum officinarum L.) bagasse and evaluation of the antioxidant and antiproliferative activities. Industrial Crops and Products. 2017;107:360-369. DOI: 10.1016/j.indcrop.2017.05.050
  26. 26. Zheng R, Su S, Zhou H, Yan H, Ye J, Zhao Z, et al. Antioxidant/antihyperglycemic activity of phenolics from sugarcane (Saccharum officinarum L.) bagasse and identification by UHPLC-HR-TOFMS. Industrial Crops and Products. 2017;101:104-114. DOI: 10.1016/j.indcrop.2017.03.012
  27. 27. Luo L, Cui Y, Zhang S, Li L, Suo H, Sun B. Detailed phenolic composition of Vidal grape pomace by ultrahigh-performance liquid chromatography-tandem mass spectrometry. Journal of Chromatography B. 2017;1068:201-209. DOI: 10.1016/j.jchromb.2017.10.031
  28. 28. Maier T, Schieber A, Kammerer DR, Carle R. Residues of grape (Vitis vinifera L.) seed oil production as a valuable source of phenolic antioxidants. Food Chemistry. 2009;112:551-559. DOI: 10.1016/j.foodchem.2008.06.005
  29. 29. He L, Xu H, Liu X, He W, Yuan F, Hou Z, et al. Identification of phenolic compounds from pomegranate (Punica granatum L.) seed residues and investigation into their antioxidant capacities by HPLC-ABTS+ assay. Food Research International. 2011;44:1161-1167. DOI: 10.1016/j.foodres.2010.05.023
  30. 30. Castro ACCM, Oda FB, Almeida-Cincotto MGJ, Davanço MG, Chiari-Andréo BG, Cicarelli RMB, et al. Green coffee seed residue: A sustainable source of antioxidant compounds. Food Chemistry. 2018;246:48-57. DOI: 10.1016/j.foodchem.2017.10.153
  31. 31. Berardini N, Fezer R, Conrad J, Beifuss U, Carle R, Schieber A. Screening of mango (Mangifera indica L.) cultivars for their contents of flavonol O- and xanthone C-glycosides, anthocyanins, and pectin. Journal of Agricultural and Food Chemistry. 2005;53:1563-1570. DOI: 10.1021/jf0484069
  32. 32. Guerrero RF, Biais B, Richard T, Puertas B, Waffo-Teguo P, Merillon JM, et al. Grapevine cane’s waste is a source of bioactive stilbenes. Industrial Crops and Products. 2016;94:884-892. DOI: 10.1016/j.indcrop.2016.09.055
  33. 33. Paes J, Dotta R, Barbero GF, Martínez J. Extraction of phenolic compounds and anthocyanins from blueberry (Vaccinium myrtillus L.) residues using supercritical CO2 and pressurized liquids. The Journal of Supercritical Fluids. 2014;95:8-16. DOI: 10.1016/j.supflu.2014.07.025
  34. 34. Di Donato P, Taurisano V, Tommonaro G, Pasquale V, Jiménez JMS, de Pascual-Teresa S, et al. Biological properties of polyphenols extracts from agro industry’s wastes. Waste and Biomass Valorization. 2018;9:1567-1578 DOI: 10.1007/s12649-017-9939-4
  35. 35. Grillo G, Boffa L, Binello A, Mantegna S, Cravotto G, Chemat F, et al. Cocoa bean shell waste valorization; extraction from lab to pilot-scale cavitational reactors. Food Research International. 2019;115:200-208. DOI: 10.1016/j.foodres.2018.08.057
  36. 36. Azadfar M, Gao AH, Bule MV, Chen S. Structural characterization of lignin: A potential source of antioxidants guaiacol and 4-vinylguaiacol. International Journal of Biological Macromolecules. 2015;75:58-66. DOI: 10.1016/j.ijbiomac.2014.12.049
  37. 37. Santos VAQ, Nascimento CG, Schimidt CA, Mantovani D, Dekker RF, da Cunha MAA. Solid-state fermentation of soybean okara: Isoflavones biotransformation, antioxidant activity and enhancement of nutritional quality. LWT-Food Science and Technology. 2018;92:509-515. DOI: 10.1016/j.lwt.2018.02.067
  38. 38. Kołodziejczyk K, Sójka M, Abadias M, Viñas I, Guyot S, Baron A. Polyphenol composition, antioxidant capacity, and antimicrobial activity of the extracts obtained from industrial sour cherry pomace. Industrial Crops and Products. 2013;51:279-288. DOI: 10.1016/j.indcrop.2013.09.030
  39. 39. Silva PB, Duarte CR, Barrozo MA. Dehydration of acerola (Malpighia emarginata DC) residue in a new designed rotary dryer: Effect of process variables on main bioactive compounds. Food and Bioproducts Processing. 2016;98:62-70. DOI: 10.1016/j.fbp.2015.12.008
  40. 40. Resende-Marques TR, Corrêa AD, Lino JBDR, Abreu CMPD, Simão AA. Chemical constituents and technological functional properties of acerola (Malpighia emarginata DC.) waste flour. Food Science and Technology. 2013;33:526-531. DOI: 10.1590/S0101-20612013005000085
  41. 41. Nóbrega EM, Oliveira EL, Genovese MI, Correia RT. The impact of hot air drying on the physical-chemical characteristics, bioactive compounds and antioxidant activity of acerola (Malphigia emarginata) residue. Journal of Food Processing and Preservation. 2015;39:131-141. DOI: 10.1111/jfpp.12213
  42. 42. Silva DI, Silva NC, Mendes LG, Barrozo MA. Effects of thick-layer drying on the bioactive compounds of acerola residues. Journal of Food Process Engineering. 2018;41:e12854. DOI: 10.1111/jfpe.12854
  43. 43. Doymaz I, Akgün NA. Study of thin-layer drying of grape wastes. Chemical Engineering Communications. 2009;196:890-900. DOI: 10.1080/00986440802668422
  44. 44. Celma AR, Rojas S, Lopez F, Montero I, Miranda T. Thin-layer drying behavior of sludge of olive oil extraction. Journal of Food Engineering. 2007;80:1261-1271. DOI: 10.1016/j.jfoodeng.2006.09.020
  45. 45. Montero I, Blanco J, Miranda T, Rojas S, Celma AR. Design, construction and performance testing of a solar dryer for agroindustrial by-products. Energy Conversion and Management. 2010;51:1510-1521. DOI: 10.1016/j.enconman.2010.02.009
  46. 46. Spigno G, Tramelli L, De Faveri DM. Effects of extraction time, temperature and solvent on concentration and antioxidant activity of grape marc phenolics. Journal of Food Engineering. 2007;81:200-208. DOI: 10.1016/j.jfoodeng.2006.10.021
  47. 47. Karacabey E, Mazza G. Optimization of antioxidant activity of grape cane extracts using response surface methodology. Food Chemistry. 2010;119:343-348. DOI: 10.1016/j.foodchem.2009.06.029
  48. 48. Corrales M, Toepfl S, Butz P, Knorr D, Tauscher B. Extraction of anthocyanins from grape by-products assisted by ultrasonics, high hydrostatic pressure or pulsed electric fields: A comparison. Innovative Food Science & Emerging Technologies. 2008;9:85-91. DOI: 10.1016/j.ifset.2007.06.002
  49. 49. Da Porto C, Natolino A, Decorti D. The combined extraction of polyphenols from grape marc: Ultrasound assisted extraction followed by supercritical CO2 extraction of ultrasound-raffinate. LWT-Food Science and Technology. 2015;61:98-104. DOI: 10.1016/j.lwt.2014.11.027
  50. 50. Okiyama DC, Soares ID, Cuevas MS, Crevelin EJ, Moraes LA, Melo MP, et al. Pressurized liquid extraction of flavanols and alkaloids from cocoa bean shell using ethanol as solvent. Food Research International. 2018;114:20-29. DOI: 10.1016/j.foodres.2018.07.055
  51. 51. Rivière C, Papastamoulis Y, Fortin PY, Delchier N, Andriamanarivo S, Waffo-Teguo P, et al. New stilbene dimers against amyloid fibril formation. Bioorganic & Medicinal Chemistry Letters. 2010;20:3441-3443. DOI: 10.1016/j.bmcl.2009.09.074
  52. 52. Sharma K, Mahato N, Cho MH, Lee YR. Converting citrus wastes into value-added products: Economic and environmently friendly approaches. Nutrition. 2017;34:29-46. DOI: 10.1016/j.nut.2016.09.006
  53. 53. Choi SY, Ha TY, Ahn JY, Kim SR, Kang KS, Hwang IK, et al. Estrogenic activities of isoflavones and flavones and their structure-activity relationships. Planta Medica. 2008;74:25-32. DOI: 10.1055/s-2007-993760
  54. 54. Schofield P, Mbugua DM, Pell AN. Analysis of condensed tannins: A review. Animal Feed Science and Technology. 2001;91:21-40. DOI: 10.1016/S0377-8401(01)00228-0
  55. 55. Domínguez-Rodríguez G, Marina ML, Plaza M. Strategies for the extraction and analysis of non-extractable polyphenols from plants. Journal of Chromatography A. 2017;1514:1-15. DOI: 10.1016/j.chroma.2017.07.066
  56. 56. Sette M, Wechselberger R, Crestini C. Elucidation of lignin structure by quantitative 2D NMR. Chemistry-A European Journal. 2011;17:9529-9535. DOI: 10.1002/chem.201003045
  57. 57. Ragauskas AJ, Beckham GT, Biddy MJ, Chandra R, Chen F, Davis MK, et al. Lignin valorization: Improving lignin processing in the biorefinery. Science. 2014;344:1246843. DOI: 10.1126/science.1246843
  58. 58. Pérez-Jiménez J, Díaz-Rubio ME, Saura-Calixto F. Non-extractable polyphenols in plant foods: Nature, isolation, and analysis. In: Watson RR, editor. Polyphenols in Plants. Academic Press: Elsevier; 2014. pp. 203-218. DOI: 10.1016/C2011-0-08711-2
  59. 59. Adjin-Tetteh M, Asiedu N, Dodoo-Arhin D, Karam A, Amaniampong PN. Thermochemical conversion and characterization of cocoa pod husks a potential agricultural waste from Ghana. Industrial Crops and Products. 2018;119:304-312. DOI: 10.1016/j.indcrop.2018.02.060
  60. 60. Hernández C, Morales Sillero A, Fernández-Bolaños J, Bermúdez Oria A, Azpeitia Morales A, Rodríguez-Gutiérrez G. Cocoa bean husk: Industrial source of antioxidant phenolic extract. Journal of Science and Food Agriculture. 2019;99:325-333. DOI: 10.1002/jsfa.9191
  61. 61. Barbosa-Pereira L, Guglielmetti A, Zeppa G. Pulsed electric field assisted extraction of bioactive compounds from cocoa bean shell and coffee silverskin. Food and Bioprocess Technology. 2018;11:818-835. DOI: 10.1007/s11947-017-2045-6
  62. 62. Mayanga-Torres PC, Lachos-Perez D, Rezende CA, Prado JM, Ma Z, Tompsett GT, et al. Valorization of coffee industry residues by subcritical water hydrolysis: Recovery of sugars and phenolic compounds. The Journal of Supercritical Fluids. 2017;120:75-85. DOI: 10.1016/j.supflu.2016.10.015
  63. 63. Lessa OA, dos Santos Reis N, Leite SGF, Gutarra MLE, Souza AO, Gualbert SA, et al. Effect of the solid-state fermentation of cocoa shell on the secondary metabolites, antioxidant activity, and fatty acids. Food Science and Biotechnology. 2018;27:107-113. DOI: 10.1007/s10068-017-0196-x
  64. 64. Arapitsas P. Hydrolysable tannin analysis in food. Food Chemistry. 2012;135:1708-1717. DOI: 10.1016/j.foodchem.2012.05.096
  65. 65. Toro-Uribe S, Montero L, López-Giraldo L, Ibáñez E, Herrero M. Characterization of secondary metabolites from green cocoa beans using focusing-modulated comprehensive two-dimensional liquid chromatography coupled to tandem mass spectrometry. Analytica Chimica Acta. 2018;1036:204-213. DOI: 10.1016/j.aca.2018.06.068
  66. 66. Ambigaipalan P, de Camargo AC, Shahidi F. Phenolic compounds of pomegranate byproducts (outer skin, mesocarp, divider membrane) and their antioxidant activities. Journal of Agricultural and Food Chemistry. 2016;64:6584-6604. DOI: 10.1021/acs.jafc.6b02950
  67. 67. Ramirez-Coronel MA, Marnet N, Kolli VK, Roussos S, Guyot S, Augur C. Characterization and estimation of proanthocyanidins and other phenolics in coffee pulp (Coffea arabica) by thiolysis-high-performance liquid chromatography. Journal of Agricultural and Food Chemistry. 2004;52:1344-1349. DOI: 10.1021/jf035208t
  68. 68. Lin S, Li Q, Yang B, Duan X, Zhang M, Shi J, et al. Transformation of litchi pericarp-derived condensed tannin with Aspergillus awamori. International Journal of Molecular Sciences. 2016;17:1067. DOI: 10.3390/ijms17071067
  69. 69. Mämmelä P, Savolainen H, Lindroos L, Kangas J, Vartiainen T. Analysis of oak tannins by liquid chromatography-electrospray ionisation mass spectrometry. Journal of Chromatography A. 2000;891:75-83. DOI: 10.1016/S0021-9673(00)00624-5
  70. 70. Jiang B, Zhang Y, Gu L, Wu W, Zhao H, Jin Y. Structural elucidation and antioxidant activity of lignin isolated from rice straw and alkali-oxygen black liquor. International Journal of Biological Macromolecules. 2018;116:513-519. DOI: 10.1016/j.ijbiomac.2018.05.063
  71. 71. Moniz P, Serralheiro C, Matos CT, Boeriu CG, Frissen AE, Duarte LC, et al. Membrane separation and characterisation of lignin and its derived products obtained by a mild ethanol organosolv treatment of rice straw. Process Biochemistry. 2018;65:136-145. DOI: 10.1016/j.procbio.2017.11.012
  72. 72. Mandelli F, Brenelli LB, Almeida RF, Goldbeck R, Wolf LD, Hoffmam ZB, et al. Simultaneous production of xylooligosaccharides and antioxidant compounds from sugarcane bagasse via enzymatic hydrolysis. Industrial Crops and Products. 2014;52:770-775. DOI: 10.1016/j.indcrop.2013.12.005
  73. 73. Rochín-Medina JJ, Ramírez K, Rangel-Peraza JG, Bustos-Terrones YA. Increase of content and bioactivity of total phenolic compounds from spent coffee grounds through solid state fermentation by Bacillus clausii. Journal of Food Science and Technology. 2018;55:915-923. DOI: 10.1007/s13197-017-2998-5
  74. 74. Mikulski D, Molski M. Quantitative structure-antioxidant activity relationship of trans-resveratrol oligomers, trans-4, 4′-dihydroxystilbene dimer, trans-resveratrol-3-O-glucuronide, glucosides: Trans-piceid, cis-piceid, trans-astringin and trans-resveratrol-4′-O-β-D-glucopyranoside. European Journal of Medicinal Chemistry. 2010;45:2366-2380. DOI: 10.1016/j.ejmech.2010.02.016
  75. 75. Demirci MA, Ipek Y, Gul F, Ozen T, Demirtas I. Extraction, isolation of heat-resistance phenolic compounds, antioxidant properties, characterization and purification of 5-hydroxymaltol from Turkish apple pulps. Food Chemistry. 2018;269:111-117. DOI: 10.1016/j.foodchem.2018.06.147

Written By

Beatriz Hernández-Carlos, Norma Francenia Santos-Sánchez, Raúl Salas-Coronado, Claudia Villanueva-Cañongo and Paula Cecilia Guadarrama-Mendoza

Submitted: 26 September 2018 Reviewed: 13 February 2019 Published: 20 March 2019