1. Introduction
The consumption of raw materials such as biomass, fossil fuels, metals, and minerals is expected to double over the next 40 years on a global scale [
1]. Likewise, the waste generated each year is estimated to increase by 70% by 2050 [
2]. With that in mind, the European Commission has recently enforced the Circular Economy Action Plan [
3], one of the main components of the overall European Green Deal, the European new agenda for sustainable growth [
4]. The Green Deal is implementing an EU coordinated strategy to become a climate-neutral, resource-efficient, and competitive economy. Achieving climate neutrality by 2050 will require a decisive effort in supporting the scaling up of circular economy from front-runners to the mainstream economic players; in this way, a decoupling of economic growth from resource use will be encouraged, and the long-term competitiveness of the EU will also be fostered.
Small and medium-sized enterprises are the leading actors of this ecological shifting by adopting advanced and innovative “green” technologies and putting into practice the appropriate know-how, got from Academia, for achieving the replacement of hazardous substances from the production processes. This approach can be applied in almost all manufacturing sectors. The agro-industrial field is of great interest because of the huge amount of residues and by-products that are generated. They could be converted into high value compounds and products by means biorefineries based on environmentally friendly processes [
5,
6]. On the other hand, the effective deployment of integrated biorefinery plants requires the setting up of reliable processing units combined with advanced eco-friendly and economically profitable value chains [
7]. Increasing use of disposable, underused, and residual biomass could increasingly supply the feedstock requirements as expressed by forthcoming developments of new integrated biorefinery systems.
In this context, globe artichoke (
Cynara cardunculus L. subsp.
scolymus (L.) Hayek) represents a very intriguing feedstock for the Mediterranean Area. It is a perennial plant belonging to the Asteraceae family and originating from the Mediterranean region [
8]. It is traditionally cultivated as a polyannual crop through vegetative propagation. Nevertheless, the length of the crop cycle negatively influences yields and the quality of the heads. This led artichoke growers to take an interest in the development of new seed-propagated cultivars for annual crops [
9,
10,
11]. The world production of artichoke accounts for about 1678 Ktons, and Italy is the largest producer with 390 Ktons of artichokes in the world, followed by Egypt (324 Ktons), Spain (208 Ktons), and Perù (155 Ktons) [
12]. Heads (flowers) and stems just below them constitute the edible part of the plant. They are characterized by a high content of bioactive compounds, including phenols, inulin, fibers, and minerals that make them really attractive for market [
13,
14,
15]. The industrial artichoke processing generates a huge amount of waste biomass (80–85%) unsuitable for human consumption that is composed of bracts and stems cut during the harvesting process [
14,
16]. However, according to Gominho et al. [
8], it represents only a small part (15–30% dw) of the entire biomass, depending on genotype, climate, soil, and culture conditions. The remaining part (70–85% dw) is composed by residual leaves, stalks and roots that remain available in the field when cultivated as an annual crop and are usually disposed of as solid waste or left in the field without any further valorization.
Interestingly, the phenolic composition of these residues have been proved to be similar to the edible parts of the plant [
17,
18,
19]. Moreover, according to Zuorro et al. [
16], the phenolic content is higher than other phenols source such as carrot peels, grape pomace, and spent coffee grounds. In addition, an interesting amount of inulin in artichoke roots (6–21% dw) was recently reported by Castellino et al. [
9].
Phenols and inulin from residual biomass of artichoke are excellent candidates for the biorefinery process having many therapeutic properties and biotechnological applications that could contribute in increasing the economic value of residual biomass. The artichoke phenols have been identified since ancient times for their beneficial effects and therapeutic actions including the promotion of blood circulation, mobilization of energy reserves, induction of choleresis, inhibition of cholesterol biosynthesis and low-density lipoprotein (LDL) cholesterol oxidation. A significant antibacterial, antifungal, and antioxidant, as well as strong hepatoprotective effects, are also recognised [
15,
20,
21]. Inulin is a natural storage polysaccharide widely distributed in plants. It is a water-soluble fiber that consists of a mixture of oligo- and polysaccharides of β(2→ 1) linked D-fructose units with a terminal glucose residue, which are classified as fructans [
22]. Inulin-type fructans are considered as prebiotic compounds, i.e., it is indigestible by humans, but it stimulates the growth and activity of specific microorganisms, including
Lactobacilli and
Bifidobacteria in the colon [
23,
24,
25].
Following a general trend towards the recycling biomass, including food waste and agro-industrial residues, several research studies have been performed in order to find new pathways for valorizing the residues of artichoke [
26]. Some examples include the extraction of phenolic compounds [
16,
27,
28], the production of biofuels [
14,
29], the recovery of enzyme peroxidase for wastewater treatment [
30], and the extraction and purification of high-molecular-weight inulin [
9,
31]. In the light of our knowledge, little attention has been paid in sequential and eco-friendly processes that could convert residual artichoke biomass into a plethora of high-value compounds (phenols, inulin, etc.), bioenergy and agricultural application at the same time. Nevertheless, no studies have been performed aiming at designing an integrated biorefinery process for the valorization of different parts of globe artichoke plant residues (heads, leaves, stalks, and roots) when cultivated as an annual crop.
Therefore, this work aimed to investigate a sequential microwave extraction process of phenols and inulin from artichoke crop residues, inspired to green chemistry principles, to convert waste into valuable compounds with a biorefinery approach. Moreover, two different valorization pathways of extracted residual biomass as bioenergy feedstocks and green manure for agriculture application were evaluated.
2. Materials and Methods
2.1. Sampling and Biomass Preparation
Artichoke plants (
C. cardunculus L. subsp.
scolymus (L.) Hayek), Madrigal
® (Nunhems SAS, Beaucouze, France) seed propagated cultivar (hybrid variety), were collected at the end of the productive stage of heads (June 2019, after harvesting for market) from a 4 Ha cultivated field (FIMAGRI Farm) located at Candela in Southern Italy (latitude 41°08′ N, longitude 15°31′ E altitude 499 m above sea level). We randomly selected four sampling squares (3.5 m × 3.5 m) in the field. The number of plants was counted in each sampling unit. Then, three entire plants along the diagonal of each square were collected. Sampled plants (12) were immediately brought to the laboratory where the 4 main components were hand separated: Heads (residual), leaves, stalks and roots. The wet weight was measured for each component. Samples were dried in a ventilated oven at 60 °C according to National Renewable Energy Laboratory (NREL) protocol [
32]. Dried samples were ground in a cutting mill (Pulverisette 15, Fritsch, Idar-Oberstein, Germany) to pass through a 1 mm sieve. The mixed portions of dried tissue were used for chemical characterization and extractions.
2.2. Chemical Characterisation of Biomass
Proximate analysis (moisture, ash, volatile solids, fixed carbon) of raw biomass (heads, leaves stalks and roots) and extracted biomass, after the sequential extraction of phenols and inulin, was performed using a thermogravimetric analysis system (TGA 701, LECO, St. Joseph, MI, USA), following ASTM D7582 method.
Ultimate analysis (C, H, N, S, O) was performed using a CHNS628 elemental analyzer (LECO, St. Joseph, MI, USA) and following the method LECO-ASTM-D 5291. The oxygen content was calculated by difference, including the ash content.
Protein content was calculated by multiplying elemental N concentration by a factor of 6.25, according to standard method AOAC-2016.
The high heating value (HHV) was experimentally determined in the laboratory with an adiabatic bomb calorimeter AC-500 (LECO, St. Joseph, MI, USA) following the standard method CEN/TS 14918:2005.
Elemental analysis (micro- and macro-elements) was performed by digesting 0.25 g dw of the sample in 20 mL of HNO3 in a closed vessel of microwave digester (CEM-Mars6) for 20 min at 220 °C. The metals in the solution were analyzed by inductively coupling plasma optical emission spectroscopy (ICP-OES Agilent 720, Agilent Technologies, Santa Clara, CA, USA), calibrated with external standard (TraceCERT®, Sigma–Aldrich, St. Louis, MO, USA).
Structural-carbohydrates from cellulose and hemicelluloses together with ‘‘Klason lignin” were measured using a strong acid hydrolysis, according to National Renewable Energy Laboratory (NREL) method [
33]. Monosaccharides (i.e., glucose, xylose, arabinose, fructose, galactose), uronic sugars (glucuronic and galacturonic) and dehydration products (levulinic acid, hydroxyl methyl furfural, and furfural) were analyzed by HPLC (Agilent 1260, Agilent Technologies, Santa Clara, CA, USA) coupled to a refractive index detector (RID). The analysis was carried out with a Hi-Plex H column at 60 °C. The eluent was ultrapure water (MilliQ
®, Merk Millipore, Burlington, MA, USA) 5 mM H
2SO
4 under a flow rate of 0.7 mL min
−1. The system was calibrated with pure chemicals from Sigma–Aldrich (St. Louis, MO, USA). Thereafter, cellulose and hemicelluloses contents were estimated as follows:
where 1.11 is the conversion factor for glucose-based polymers (glucose) to monomers and 1.13 is the conversion factor for xylose-based polymers (arabinose and xylose) to monomers.
Total Carbohydrates were quantified as the sum of monomeric carbohydrates, uronic sugars, and dehydration products (levulinic acid, hydroxymethylfurfural, and furfural) converted into corresponding hexose and pentose sugars amount as follows:
where 1.55 is the hexose–LA molecular weight ratio, 1.43 is the hexose–HMF molecular weight ratio and 1.56 is the pentose–Furfural molecular weight ratio.
2.3. Microwave-Assisted Extraction (MAE) of Phenols
Microwave extraction was carried out using a microwave reaction system MARS-6 (CEM srl, Cologno Al Serio, Italy) consisted of 12 closed extraction vessels equipped with an infrared temperature sensor. Direct measurement of pressure and temperature (by means optic fiber probe) was performed in the reference vessel. The magnetic stirring was set at 300 rpm. There were two green solvents, water, and ethanol at four different concentrations in water (0, 25, 50, and 75% v/v), used for the extraction. The sample weight to solvent volume ratio was maintained constant at 1:10 (w/v ratio) for all extractive experiments. In a typical extraction process, 1 g dw of biomass was extracted with 10 mL of solvent. We tested three temperature levels (50, 75 and 100 °C) with three different extraction times (5, 10, and 20 min). The microwave frequency used for the extraction was 2450 MHz. At the end of the extraction, the vessels were cooled down to 25 °C using compressed air. Then, the mixture was centrifuged at 4100 rpm for 10 min, and the liquid phase was filtered (0.20 µm). The extracts were then flushed with nitrogen gas and stored in the dark at −40 °C until HPLC analysis of phenolic profile. The wet solid residue (Phenols Extracted Residue—PER) was weighed and stored for sequential inulin extraction. A portion was dried in an oven at 60 °C, weighed and stored for further chemical characterizations.
2.4. Conventional Extraction (CE) of Phenols
With the aim to evaluate the extraction efficiency of MAE, the phenols extraction was performed following the conventional extraction (CE) method shaking the biomass with solvents reported above for MAE (1:10, w/v ratio) for 20 min, 8, 24, and 48 h at room temperature and using an orbital shaker (Stuart, Cole-Parmer srl, Cernusco sul Naviglio, Italy) set at 70 rpm. The following work sequence was the same as described above for MAE.
2.5. Microwave-Assisted Extraction of Inulin
Microwave-assisted extractions (MAE) of inulin from raw and residual biomass (PER) were performed in the same microwave reaction system (MARS-6, CEM srl, Cologno Al Serio, Italy) used for phenol extraction. Sample (0.5 g dw) was transferred to an extraction vessel containing 50 mL of water (MilliQ®, Merk Millipore, Burlington, MA, USA). The operational parameters employed in the MAE apparatus were the following: magnetron power 100%, ramp temperature and extraction time of 5 min, respectively. During operations, both temperature and pressure were monitored. Then, three temperature levels were tested for extractions: 60, 80, and 100 °C. After the extraction, the vessels were cooled down to room temperature, and the mixture was centrifuged at 4100 rpm for 10 min. The liquid phase was filtered (0.45 µm) and analyzed by HPLC for inulin quantification. The extracted inulin was precipitated with EtOH, dried and then analyzed by FT-IR ATR (Attenuated Total Reflectance). The solid residue (Inulin Extracted Residue—IER) was dried in an oven at 60 °C, weight and stored for further chemical characterizations. The inulin extraction efficiency of MAE compared to CE was evaluated.
2.6. Conventional Extraction of Inulin
The conventional extraction of inulin was performed by means hot water diffusion procedure at an average temperature of 85 °C for one hour with a continuous stirring as reported by Zutela and Sambucetti [
34]. The sample (1 g dw) was transferred into 200 mL Pyrex beaker, added with 100 mL of hot water at pH 6–8, and kept at 85 °C with continuous magnetic stirring for 1 h. The mixture was then cooled down to room temperature, and the volume was made up to 100 mL. The solution was filtered (0.45-µm) and characterized by HPLC analysis.
2.7. HPLC Analysis of Phenolic Profile
The extracted phenols were analyzed by means high pressure liquid chromatography (HPLC, 1260 Infinity, Agilent Technologies, Santa Clara, CA, USA) coupled with a Diode Array Detector (DAD). A reversed-phase Zorbax Stable Bond SB-C18 column (250 × 4.6 mm i.d., particle size 5 µm, Agilent Technologies, Santa Clara, CA, USA) with a Zorbax precolumn guard cartridge (10 mm × 4 mm i.d. 5 µm) was used for chromatographic separation at room temperature (25 °C). The injection volume was set at 20 µL.
A gradient binary elution was performed using solvent A (0.1% formic acid in water) and solvent B (0.1% formic acid in acetonitrile/methanol, 60:40
v/
v) at a constant flow of 0.8 mL min
−1. The gradient program reported by Rouphael et al. [
13] was used: 20–30% B (6 min), 30–40% B (10 min), 40–50% B (8 min), 50–90% B (8 min), 90–90% B (3 min), 90–20% B (3 min).
HPLC was calibrated using commercial standards provided by Sigma–Aldrich (St. Louis, Missouri, USA): chlorogenic acid (≥96% purity), caffeic acid (≥97% purity), 1,3-dicaffeoylquinic acid (≥95% purity), ferulic acid (≥95% purity), 1,5-dicaffeoylquinic acid (≥95% purity), 1-caffeoylquinic acid (97% purity), apigenin (95% purity), luteolin (≥95% purity), apigenin-7-O-glucoside (≥95% purity), luteolin 7-O-glucoside (≥95% purity).
Caffeoylquinic derivatives were quantified at 330 nm, while apigenin and luteolin derivatives were quantified at 330 nm and 350 nm, respectively. Phenolic compounds were identified by comparison with commercial standards and available data in the literature [
13,
35]. All data are reported as mg g
−1 of dry matter (dw). Calibration curves and calculation of limit of detection (LOD) and limit of quantification (LOQ) values are reported in
Table S1 (Supplementary Materials).
2.8. HPLC Analysis of Inulin
Inulin was quantified by HPLC analysis following the analytical method reported by Zutela and Sambucetti [
34]. The chromatographic equipment consisted of an Agilent 1260 HPLC (Agilent Technologies, Santa Clara, CA, USA) equipped with a refractive index detector and an Aminex HPX-87C (Bio-Rad Laboratories S.r.l., Segrate, Italy) anion exchange column. The mobile phase was ultrapure water (MilliQ
®, Merk Millipore, Burlington, MA, USA) at 85 °C at a flux rate of 0.6 mL min
−1 and an injection volume of 20 µL. Pure inulin from chicory (Sigma–Aldrich, St. Louis, MO, USA) was used as an external standard for calibration. A calibration range between 0.5 and 5 mg mL
−1 was set for inulin quantification.
2.9. FT-IR ATR Analysis of Inulin
Standards and mixtures were analyzed by Fourier transform infrared attenuated total reflectance (FT-IR ATR) spectroscopy by Perkin Elmer Spectrum TWO FT-IR spectrometer equipment (Waltham, MA, USA) operating at 4 cm−1 resolution with 64 scans per test. Spectra were collected in the transmittance mode from 4000 to 400 cm−1.
2.10. Statistical Analysis
All the experiments were repeated three times. Unless otherwise stated, all data were expressed as mean ± standard deviation (SD). The means of all the parameters were examined for significance by analysis of variance (ANOVA) using the software JMP version 9 (SAS Institute Inc., Cary, NC, USA). When F values showed significance, individual means were compared using Tukey’s honest significant difference (HSD). Significant differences were considered when p < 0.05.