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Article

Physicochemical Changes and Microbiome Associations during Vermicomposting of Winery Waste

by
Ioanna Karapantzou
1,
Gregoria Mitropoulou
1,
Ioanna Prapa
1,
Dimitra Papanikolaou
1,
Vasileios Charovas
2 and
Yiannis Kourkoutas
1,*
1
Laboratory of Applied Microbiology and Biotechnology, Department of Molecular Biology and Genetics, Democritus University of Thrace, 68100 Alexandroupolis, Greece
2
Evritika Kellaria S.A., 68200 Orestiada, Greece
*
Author to whom correspondence should be addressed.
Sustainability 2023, 15(9), 7484; https://doi.org/10.3390/su15097484
Submission received: 17 March 2023 / Revised: 25 April 2023 / Accepted: 26 April 2023 / Published: 2 May 2023
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Abstract

:
Annually, the wine industry produces high amounts of waste that can be toxic if disposed of without pretreatment. Vermicomposting is an efficient and low-cost method of decomposing organic matter using earthworms under controlled conditions. The organic substrate used in the vermicomposting process affects microbial populations and reflects the dynamics of enzymatic activity, decomposition of organic matter, and nitrogen transformations. However, the microbiome associations during the whole process are still unexplored. Thus, the aim of the present study was to investigate physicochemical, enzymatic, microbial, and microbiome activities during vermicomposting of winery waste. In this vein, a rectangular vermireactor with Eisenia andrei and Eisenia fetida earthworms, loaded with winery waste, was used. At the end of the process, the carbon/nitrogen (C/N) ratio was decreased, the total nitrogen was increased, the pH was neutralized and excess enzymatic activities were recorded. The bacterial and fungal phyla detected by next-generation sequencing analysis identified Armatimonadetes, Bacteriodetes, Candidatus saccharibacteria, Chloroflexi, Cyanobacteria, Planctomycetes, and Proteobacteria and Ascomycota, Basidiomycota, Chytridiomycota, Entomophthoromycota, Glomeromycota, and Mucoromycota, respectively. Physicochemical and microbial changes occurring during vermicomposting of winery waste, along with the microbiome diversity at the beginning and end of the process, may lead to a better understanding of winery-waste biotransformation into effective biofertilizer.

Graphical Abstract

1. Introduction

Wine output reached 279 million hectoliters (Mhl) in 2018 [1]. Winemaking generates 0.3–0.5 kg of wine byproducts per liter, including winter pruning byproducts (vine shoots or vine trimmings and vine leaves). Mature grapes are harvested and transported to wineries, where the destemming process produces additional byproducts known as stalks (or stems) for both red and white wines (and juices). The main solid byproducts of winemaking are vine shoots; grape stalks; grape pomace, which is made up of grape peels and grape seeds in a weight ratio of about 1:1; and wine lees [2,3].
Generally, wine waste has been intended for distillation, landfilling, incineration, and/or land spreading. In particular, grape stalks have been mainly land-spread (76% in Italy, 55% in Spain, and 40% in France), disposed of in landfills (50% in Greece), or destroyed by incineration (36% in France) [2]. Grape pomace has primarily been used as a material for distillation (100% Italy, 90% France, 30–60% Spain) or for land spreading (50% Spain), whereas in Greece it is primarily discarded (67%) or sold as animal fodder (33%) [2]. Finally, in most countries, wine lees are processed further by the distillation method, while grape shoots are land-spread or burned in the field [2].
However, conventional uses of wine waste have been seriously reconsidered. Expensive pretreatments or conditioning steps are needed for wine waste in order for it to be converted to ideal fertilizer because of its low pH, high organic matter content, and high concentrations of macronutrients [3]. Similarly, wine byproducts can inhibit or modify germination properties when used as amendments, and landfilling is strongly discouraged, since wine waste affects soil erosion and decreases groundwater quality because of the high organic-matter losses [4,5]. Grape pomace constitutes a problem as animal feed due to its high polyphenol concentration, which bonds with proteins and produces compounds that are unsuitable for nutritional applications [6]. In addition, with the 2013 Reg. No 1308/2013—European Decree in the matter of wine-waste-disposal rules, distillation of wine byproducts is no longer a remunerative option for wine companies [7]. For these reasons, along with the need to invest in new sustainable and renewable materials, transformation of wine waste to biofertilizers has lately been examined.
In recent years, composting has attracted considerable attention as a sustainable and environmentally friendly alternative for treatment of organic municipal waste. Vermicomposting is the process of decomposing organic matter using earthworms. Thus, the earthworms consume the raw material, which passes through the gut and is then excreted as “worm cast”, rich in nutrients and microflora that continue to decompose the organic material [8,9,10,11]. Raw organic materials that have undergone the metabolic activity of earthworms have enhanced suitability for applications as fertilizing resources. Soil organic matter becomes richer after vermicomposting and plays an important role in sustaining soil fertility. In addition, vermicompost also enhances the physicochemical and biological properties of soil. As an outcome, (i) the soil becomes more resilient to stresses such as drought, diseases, and toxicity; (ii) plant nutrient intake is improved; and (iii) active nutrient cycling is achieved, owing to vigorous microbial activity [8,10,12].
Although a few studies on vermicomposting of agricultural winery waste are available in the literature [13,14], data on the microbiome interactions involved during the process is missing. Hence, the aim of this study was to assess the physicochemical, microbial, and microbiome changes during vermicomposting of winery waste, contributing to a better understanding of winery-waste biotransformation to effective biofertilizer.

2. Materials and Methods

2.1. Vermireactor Setup and Sample Collection

The system used was a rectangular (42 L × 42 W × 60 H cm) 4-tray vermireactor (VidaXL, Venlo, The Netherlands), as shown in Figure 1. Briefly, ~1.5 kg of exhausted grape marc, along with ~1 L of worms (Eisenia andrei and Eisenia fetida) and 140 g of coffee grounds, was mixed, homogenized, moistened to reach the appropriate humidity (>75%), divided equally among the trays of the vermireactor, and left in a closed, protected area with a constant temperature between 18 and 22 °C to begin the process. Sample material (~120 g) from the vermicomposting system was collected every 10 and 15 days and for up to 120 days, as previously described [13], and the samples were stored at −20 °C until analysis.

2.2. Chemical Analysis

2.2.1. Moisture Content, pH, Conductivity, and Temperature

Samples were dried at 121 °C for 12 h, and the % of moisture content was determined by calculating the weight loss of the samples. Temperature and conductivity were determined directly with a composting waterproof thermometer made of stainless steel with a temperature measurement range of 0–250 °C and a portable conductivity meter (CON 150 EUTECH, Thermoscientific, Singapore), respectively. Samples were mixed with water (1:10 w/v), and the pH of the supernatant was measured with a pH meter (HANNA, Limassol, Cyprus).

2.2.2. Total C, Total N, C/N Ratio, and O2 Uptake Rate

Total C [15] and total N [16] were determined, as previously described (the lowest limit of detection for C was 3%, and for N, 0.75%). O2 uptake rate was determined directly with a portable oximeter connected to a 1.5 m catheter O2-measuring sensor (HANNA, Limassol, Cyprus).

2.2.3. Macro- and Micronutrients

Carbonate content was determined directly with a calcimeter (HANNA, Cyprus). P, K, Mg, Mn, Zn, Fe, and Cu were determined by atomic absorption spectrophotometry (General Electric photometer, hardware Agrosoil, Bedfordshire, UK) according to EN ISO 9001:2000, as previously described [17]. Of note: the lowest limit of detection was 0.05 ppm.

2.3. Enzymatic Activity

Enzymatic activity was determined colorimetrically. For the determination of dehydrogenase activity, 6 g of compost sample, 0.06 g of CaCO3 (Penta Chemicals, Praha, Czech Republic), 1 mL of 3% aqueous solution of 2,3,5-triphenyl-tetrazolium chloride (TTC) (Applichem, Darmstadt, Germany), and 2.5 mL of demineralized water were mixed and then incubated at 37 °C. After 24 h of incubation, 25 mL of methanol was added and the mixture was filtered twice. The remaining solution was then diluted to 50 mL with methanol. Sample solutions (0.3 mL) were transferred to a 96-well microplate, and absorbance was determined with a SpectraMax® ABS microplate reader (Molecular Devices, San Jose, CA, USA) at 485 nm. Samples with Tris buffer (Sigma Aldrich, UK) and without TTC were used as controls. Deyhodrogenase activity was expressed as μg TPF/g (dm)/24 h and measured with the equation ((S−C) × 100)/(6 × % dm), where S is the mean value of the samples (μg TPF), C is the value of the control (μg TPF), 6 is the initial soil weight (g) and % dm is the percentance of soil dry matter of each sample [18].
Urease activity was determined with the modified Berthelot reaction [19]. Briefly, 5 g of compost was mixed with 2.5 mL (79.9 mM) of urea aqueous solution and incubated at 37 °C for 2 h. After 2 h, 2.5 mL of distilled water and 50 mL of KCL solution (50 mM) were added and the samples were left in a rotary shaker for 30 min. Subsequently, the samples were filtered, and the released ammoniums were extracted and determined based on the reaction of sodium salicylate with NH3 in the presence of sodium dichloroisocyanurate under alkaline conditions. Urease activity was expressed as ammonium (NH+4-N) released per unit sample weight and incubation time [20].
Acid phosphatase and β-galactosidase were measured using the colorimetric pNP-linked substrate approach as described by Jackson et al. [21].

2.4. Microbiological Analysis

Samples (25 g) were blended with sterile ¼ Ringer’s solution (225 mL), followed by decimal serial dilutions and plate counting. The following microbial populations were determined: (a) Total Aerobic Count (TAC) by plating on plate count agar (Condalab, Madrid, Spain) at 30 °C for 72 h; (b) Lactobacillus spp. (Lactic Acid Bacteria, LAB) by plating on acidified MRS agar, pH 5.7 (Condalab, Spain), at 37 °C for 72 h; (c) Enterobacteriaceae by plating on Violet Red Bile Glucose agar (VRBG) (Condalab, Madrid, Spain) at 37 °C for 24 h; (d) coliforms by plating on Violet Red Bile agar (VRBA) (Condalab, Madrid, Spain) at 30 °C for 24 h; (e) Clostridium spp. by plating on tryptose sulfite cycloserine agar (Condalab, Madrid, Spain) at 37 °C for 48 h under anaerobic conditions (Merck Millipore Anaerobic Jar 2.5 L, Merck Millipore 2.5 L Sachets); (f) Escherichia coli by plating on Harlequin™ Tryptone Bile Glucuronide agar (TBX) (LABM, Heywood, UK) at 37 °C for 24 h; (g) Salmonella spp. by plating on xylose lysine deoxycholate agar (VWR International GmbH, Darmstadt, Germany) at 37 °C for 24 h; and (h) yeasts and molds by plating on malt extract agar (Condalab, Spain) at 30 °C for 72 h. Cell levels were expressed as log cfu/g.

2.5. DNA Extraction, PCR Amplification, and 16S and ITS rRNA Sequencing

Samples were collected at the beginning (day 0) and at the end (day 120) of the process. The total DNA was extracted using the NucleoSpin® Soil (MACHEREY-NAGEL GmbH & Co., KG, Düren, Germany), following the manufacturer’s instructions. Next-generation sequencing (NGS) was performed using MiSeq sequencing by MR DNA (http://www.mrdnalab.com (accessed on 3 February 2023), Shallowater, TX, USA). The V1-V3 region of the bacterial 16S rRNA gene was amplified from compost genomic DNA with 27F/519R primers (AGRGTTTGATCMTGGCTCAG/GTNTTACNGCGGCKGCTG). The highly variable Internal Transcribed Spacer (ITS) regions of the fungal ITS1 and ITS2 sequences surrounding the 5.8S coding sequence located between the Small Subunit (SSU) coding sequence and the Large Subunit (LSU) coding sequence of the ribosomal operon was amplified with the primers ITS1/ITS4 (CTTGGTCATTTAGAGGAAGTAA/TCCTCCGCTTATTGATATGC). The archaeal 16S rRNA gene region was amplified with primers Arch2A519F (50-CAGCMGCCGCGGTAA-30) and Arch1017R (50-GGCCATGCACCWCCTCTC-30).
Polymerase Chain Reaction (PCR) amplification was performed using the HotStarTaq Plus Master Mix Kit (Qiagen, Germatown, MD, USA), consisting of 30–35 cycles with the following steps: 95 °C for 5 min, 30 cycles (archaea 35 cycles) of 95 °C for 30 s, 53 °C for 40 s, 72 °C for 1 min, and the final elongation step at 72 °C for 10 min. PCR products were then subjected to electrophoresis in 2% agarose gel to confirm amplification and to determine the relative intensity of bands. Then, the amplicons were purified using Ampure XP beads (Beckman Coulter, Brea, CA, USA).
Samples were subsequently prepared for the Illumina DNA library using MiSeq sequencing, following the manufacturer’s guidelines. Procession of the sequencing data was held using a proprietary analysis pipeline of MR DNA. Operational taxonomic units (OTUs) were defined by clustering at 3% divergence (97% similarity), and the final OTUs were taxonomically classified using BLASTn against a curated database derived from RDPII and NCBI (http://www.ncbi.nlm.nih.gov (accessed on 3 February 2023), http://rdp.cme.msu.edu (accessed on 3 February 2023) and compiled, in each taxonomic level, into both “counts” and “percentage” files. This analysis identified 175 bacterial, 310 fungal and 1 archaeal OTU, and low-abundance (<0.01), rare OTUs were removed. Analysis of raw data in the OTU levels and calculation of a- and b-diversity were performed using the Rhea platform [22].

2.6. Germination Index

Germination index was evaluated according to Paradelo et al. [23]. Specifically, at day 120, 50 mL of aqueous extract (sample/water ratio of 1/10 w/v) from a vermicompost system was used to submerge 50 seeds of Hordeum vulgare L. (barley) for 24 h. After 24 h, 15 seeds were placed on filter paper in a petri dish and 3 mL of the extract was added. The petri dishes were placed at 28 °C for 5 days in the dark. The number of seeds germinated and the length of the roots were recorded, and the Germination Index (GI) was calculated as follows: GI = 100 G/Gc × L/LC (G and L are the germination and radicle growths of the samples, respectively, and GC and LC the germination and radicle growths of the control (distilled water), respectively).

2.7. Statistical Analysis

All experiments were performed at least in triplicate. Data is expressed as average values ± STDEV. Results were analyzed by one-way ANOVA and significant differences were determined by Tukey’s and the Bonferroni post hoc test. ANOVA tables and significance levels (p < 0.05) were calculated with Statistica v.12 software.

3. Results and Discussion

3.1. Moisture Content, pH, Conductivity, and Temperature

Moisture content, pH, conductivity, and temperature measurements were carried out every 10 days. The moisture content of the system was maintained at 78–86% for the well-being of the earthworms, as humidity between 75 and 90% was required [24,25,26]. The pH was slightly alkaline on the first day (7.91), while from the 40th day of operation, it varied from weakly acidic to neutral values (6.5–7.01), as shown in Figure 2, in agreement with other studies [27,28]. This change in pH enhances the potential use of the product as a soil conditioner, since crops respond more favorably when soil pH ranges from a weakly acidic to a weakly alkaline level [19]. Electrical conductivity (Figure 3) was significantly (p < 0.05) increased from 2.13 mmhos/cm to 3.03 mmhos/cm from day 0 to day 20 and then decreased significantly (p < 0.05) to 1.24 mmhos/cm at day 120, in agreement with previous studies [13,25,29]. Electrical conductivity depends on the type of composting material and is related to the concentrations of ions in materials. The decrease in electrical conductivity was associated with the stabilization of the product produced [30]. Composting with earthworms is a mesophilic process, and the temperature of the system ranged from 18 to 22 °C. The final product of vermicomposting has to be ripe to minimize the toxicity of the initial substrate of the winery waste [31]. A possible explanation for the increases in electrical conductivity and pH during the first 20 days is production of salts, ammonium, and inorganic ions during the biotransformation of nonavailable nutrients into absorbable forms [30,32].

3.2. Total C, Total N, C/N Ratio, and O2 Concentration

In agreement with previous studies [13,27], total C increased significantly (p < 0.05), from 37.65% of the dry weight to 54.03%, in the first 15 days of system operation, and a decrease from the 15th day to day 120, with a final value of 21.36% (p < 0.05), was recorded. No significant (p > 0.05) variation in total N between day 0 (1.55%) and day 15 (2.1%) was noted. However, a significant (p < 0.05) decrease between the 15th (2.10%) and the 45th day (1.14%) was reported, probably due to the digestion of the waste by the earthworms. On the 45th day, 1.5 kg of waste (grape marc) was further added to the system, and the percentage of total N increased significantly (p > 0.05) on the 120th day (1.95%). The C/N ratio decreased significantly (p < 0.05), from 24.37 on the day of system initiation to 10.90 on day 120 (Figure 4). During vermicomposting, the respiratory activity of microorganisms and earthworms plays a significant role in organic carbon decrease, along with an increase in total nitrogen through mucus and nitrogenous excretion by the worms and with the presence of nitrifying and denitrifying microbes in the intestinal guts of earthworms [12,33,34,35]. The C/N ratio is used as a factor of stability and maturity of vermicompost, and its reduction is an important indicator of rapid mineralization and decomposition of the initial raw material [24,36,37,38]. According to Bhat et al. [38], a C:N ratio < 20 confirms organic waste mineralization, which indicates compost maturity. However, a C:N ratio < 12 is also preferred for agricultural purposes [38,39]. Ιn our study, the C/N ratio was 10.90 at the end of the process, indicating the maturity and suitability of the final product as an organic biofertilizer.
O2 uptake rate refers to the biological activity of a material and is an indicator of stability in the final product. Specifically, it estimates, through carbonaceous oxygen demand, readily biodegradable organic matter still present in composting material [29,40]. As reported previously, the oxygen uptake rate was significantly (p < 0.05) higher (7.9 g O2/kg) at the beginning of the vermicomposting process, when biodegradable organic matter was in high amounts, but significantly (p < 0.05) lower (7.05 g O2/kg) compared to mature and stable vermicompost (Figure 5) [29,40,41].

3.3. Macro- and Micronutrients

Important nutrients and heavy metals in the produced product were also determined (Table 1). In the first 15 days, significant (p < 0.05) increases in the Ca, Mg, and K concentrations were observed. On day 120, the Ca, Mg, and K levels were significantly (p < 0.05) decreased compared to day 15, while the decrease in Fe was not significant (p > 0.05). No significant (p > 0.05) changes were noticed in the levels of P. According to the literature, both increases and decreases in metals have been observed in earthworm composting products from different types of waste [34,42,43]. The levels of Mn and Cu remained within the limits of European legislation [44]. The passage of organic matter from the guts of earthworms results in the release of nutrients in available forms due to the earthworm gut-enzyme activities, such as phosphatases, glycosidases, ureases, minohydrolases, proteases, etc. [26,42,45].

3.4. Enzymatic Activity

Enzymatic activity is a useful tool to provide information about the potential of compost to perform biochemical reactions and is related to compost characterization and maturity [46]. The activity of four soil enzymes that participate in the metabolic process during organic-matter decomposition was studied. Determination of dehydrogenase activity is related to the metabolic status of microbial feedstock [47]. The release of bound phosphorus into bioavailable forms is linked to acid phosphatase activity [48]. Urease degrades complex nitrogenous organic matter into simpler nitrogenous forms, and β-glycosidase contributes to the carbon cycle by decomposing cellobiose into glucose molecules [46]. On the first 15–45 days, the activity of all enzymes increased significantly (p < 0.05) in the presence of high amounts of biodegradable organic matter and decreased as mature and stable compost was produced, which is in agreement with the literature [47,49,50,51,52]. The results of the enzymatic activity indicated that during the first 15–45 days, the earthworms and the microbes in their excreta exhibited considerable metabolic action (Table 2) [26,52,53].

3.5. Microbial Populations

Microbial populations were determined at regular intervals during the process. Significant increases in the cellular levels of all microbial species except Lactobacillus spp. were observed on day 15, and on day 120, all cell loads decreased significantly (p < 0.05). In previous studies, a rapid change in microbial activity in earthworm compost products had been reported in the first 15 days from the beginning of the process [27]. TAC levels increased significantly (p < 0.05), from 6.01 log cfu/g on day 0 to 8.73 log cfu/g on day 15; remained at similar levels until day 45 (8.18 log cfu/g); and decreased significantly (p < 0.05) as the produced product was stabilized. Clostridium spp. levels decreased significantly (p < 0.05) on day 45 compared to day 15, and no significant changes were noted at day 120 compared to day 45, which is in agreement with a previous study, in which no changes at the levels of specific microbial species were observed during earthworm composting [54]. The population of lactobacilli on day 0 reached up to 5.95 log cfu/g due to the high percentage of stems in the composting mixture. However, they were not detected on day 75, which is in agreement with a corresponding study [14], where 16S rRNA high-throughput sequencing showed that the percentage of lactic acid bacteria decreased from 40% at the beginning to 8% on the seventh day and remained at 1% for the rest of the process. According to Regulation (EU) 2019/1009 of the European Parliament and of the Council [44], composting products should have an absence of Salmonella spp. in a 25 g sample, and E. coli cell levels should be below 3 log cfu/g. In all samples, no Salmonella sp. was detected, while regarding E. coli populations, a significant (p < 0.05) increase in cell count was observed on day 15 compared to day 0, from 0 to 6.45 log cfu/g. However, at day 120, the levels decreased (p < 0.05) <3 log cfu/g, fully harmonizing with the European regulation. In agreement with Roubalová et al. [55], the levels of coliforms were undetectable on day 0, but on day 15, they increased (p < 0.05) to values of 7.65 logcfu/g, and they decreased (p < 0.05) on day 120. Yeast/fungal counts increased significantly (p < 0.05), from 6.01 log cfu/g to 7.34 log cfu/g, from day 0 to day 15, and decreased significantly (p < 0.05) on day 120, to 5.33 log cfu/g (Figure 6).
Earthworms’ gut microbiome consists of “nitrogen-fixing” and “decomposer microbes” that are excreted along with nutrients in their vermicasts [12]. Additionally, earthworms enhance microbial activities, leading to increased levels of soil microbial numbers and biomass [12]. In our study, all microbial levels increased during the first 15 days of the system operation, as the substrate for the earthworms was full of organic material. After decomposition of the initial substrate, the final product’s microbial flora changed. The presence of Clostridium spp. may be beneficial in vermicomposting, as according to the literature, most Clostridium spp. are saprophytes and are not involved in a disease process [56,57]. The gut of Eisenia fetida is colonized by various anaerobic N2-fixing bacteria belonging to the Clostridiaceae family, such as Clostridium butyricum, Clostridium beijerinckii, and Clostridium paraputrificum, which are well-known as nitrogen fixers and cellulose digesters [58,59,60,61].

3.6. Next-Generation Sequencing DNA Analysis

Bacteria and fungi play major roles in decomposition of organic matter. The Next-Generation DNA Sequencing analysis revealed significant changes in the microbial communities during the vermicomposting process. Diversity was determined with the Shannon and Simpson indices [62], and significant (p < 0.05) bacterial and fungal diversity were noted during vermicomposting as the Shannon index increased and the Simpson index decreased. In total, seven bacterial and six fungal phyla were detected. Armatimonadetes, Bacteroidetes, Candidatus saccharibacteria, Chloroflexi, Cyanobacteria, Planctomycetes, and Proteobacteria were reported during the process. Proteobacteria were the most abundant at the beginning, representing 95.25% of the total identified sequences. At the end of the process, all bacterial phyla increased significantly (p < 0.05), except for Candidatus saccharibacteria, which increased (p > 0.05), and Proteobacteria, which decreased significantly (p < 0.05). Although Proteobacteria decreased, it remained the third most abundant phylum, while Bacteroidetes was the most abundant, followed by Planctomycetes (Table 3).
The Ascomycota, Basidiomycota, Chytridiomycota, Entomophthoromycota, Glomeromycota, and Mucoromycota fungal phyla were detected, with no significant (p > 0.05) changes at their percentage level, except for Mucoromycota, which increased significantly (p < 0.05) but remained at a low percentage (1.21%). The most predominant fungi phylum was Ascomycota on day 0 and also on day 120 with a percentage level of 88.43–97.47%.
At the genus level, 43 bacterial genera were identified. Acetobacter (90.26%), Anaerolinea (2.04%) Ohtaekwangia (1.84%), Paracoccus (1.18%), Methylomarinum (0.65%), Devosia (0.53%), Brevundimonas (0.30%), and Mesorhizobium (0.28%) were the eight abundant genera at day 0. At day 120, the microbial population changed, and the eight most predominant bacteria were Ohtaekwangia (68.11%), Phycisphaera (10.31%), Anaerolinea (3.93%), Porphyrobacter (2.8%), Synechococcus (2.66%), Algisphaera (2.04%), Amaricoccus (1.27%), and Mesorhizobium (0.59%). Three of the genera detected (Acetobacter, Anaplasma, and Methylomarinym) decreased significantly (p < 0.05), 18 genera out of 43 increased significantly (p < 0.05) (Algisphaera, Amaricoccus, Anathece, Bradyrhizobium, Ensifer, Fimbriimonas, Mesorhizobium, Methyloceanibacter, Methylosinus, Methylovirgula, Novosphingobium, Ohtaekwangia, Phycisphaera, Porphyrobacter, Rhizobium, Sphingobium, Synechococcus, Thermanaerothrix), and the remaining 22 genera underwent no significant (p > 0.05) change (Agrobacterium, Altererythrobacter, Anaerolinea, Asticcacaulis, Blastomonas, Bosea, Brevundimonas, Candidatus saccharimonas, Caulobacter, Cellvibrio, Chryseolinea, Devosia, Dokdonella, Ochrobactrum, Paracoccus, Phenylobacterium, Pseudaminobacter, Reyranella, Shinella, Sphingomonas, Sphingopyxis, Taibaiella) (Figure 7).
Ohtaekwangia belongs to the proteolytic family Cytophagaceae, and it has been described as a nitricifier, able to produce marinoquinolines, which can provide protection to the rhizosphere from pathogens and predators [63,64,65]. According to Wiegand et al. [66], Planctomycetes (Phycisphaera and Algisphaera) plays a crucial role in the carbon and nitrogen cycles, and Phycisphaera has been reported to absorb and hydrolyze heteropolysaccharides excreted by soil microorganisms and degrade wood in marine sediments [67]. Anaerolinea belongs to the family of Anaerolineales, which digests organic waste and can sequester damaging substances to plant growth [68]. Astafyeva et al. [69] studied the most expressed genes in Porphyrobacter, and their results included antagonistic interaction mechanisms, vitamin and fatty-acid biosynthesis, and secretion systems that can promote plant growth. Synechococcus belongs to the phylum of Cyanobacteria, bacteria with the ability to fix elemental nitrogen into more easily assimilable forms [70]. Furthermore, they produce secondary metabolites with growth-promoting effects or that can enhance resistance to plant diseases. Mesorhizobium is a symbiotic N2-fixing soil bacterium that needs a host, and along with the host, it can produce various phytohormones and stimulate plant growth [71].
In total, forty-six fungal genera were detected, and the seven most abundant on the first day of vermicomposting were Pichia (48.53%), Saccharomyces (13.74%), Wickerhamomyces (13.13%), Hydnellum (9.92%), Scedosporium (6.50%), Saccharomycodes (1.84%), and Hygrocybe (0.53%). At the end of the process, Galactomyces (30.82%), Fusarium (26.09%), Scedosporium (12.15%), Microcera (11.48%), Pichia (6.59%), Peziza (6.25%), and Mortierella (1.20%) were detected as the seven most predominant genera (Figure 8).
Aleuria, Amanita, Ambispora, Cryptococcus, Gigaspora, Guehomyces, Hydnellum, Ophiocordyceps, Pichia, Saccharomyces, Saccharomycodes, Torulaspora, Trichosporon, and Wickerhamomyces decreased significantly (p < 0.05), while Arthrographis, Ceratocystis, Chytridium, Cylindrocarpon, Dipodascus, Fusarium, Galactomyces, Humicola, Malbranchea, Mortierella, Peziza, Scedosporium, Scutellinia, and Veluticeps increased significantly (p < 0.05). On the other hand, no significant (p < 0.05) change was observed in Basidiobolus,Batrachochytrium, Cladosporium, Coltricia, Doratomyces, Gibellulopsis, Hygrocybe, Hyphopichia, Kalaharituber, Lobulomyces, Lomentospora, Microcera, Nectria, Ogataea, Parascedosporium, Penicillium, Phialemonium, or Scytalidium.
According to the literature, antagonistic activities against phytopathogenic fungi have been associated with Galactomyces and thus it can be used as a biocontrol agent [72]. Μetagenomic and metataxonomic analyses of grape-marc vermicompost microbes, performed by Losada et al. [73], showed that the second most dominant genus was Fusarium, followed by Scedosporium, but since species of these genera are considered opportunistic pathogens, phytotoxicity of vermicompost products should be evaluated before application in the field. Microcera is an entomopathogenic fungus [74] that infects and kills insects and arthropods and can be used as a biopesticide. Pichia and Mortierella belong to plant-growth-promoting fungi and can improve the physicochemical and biological properties of soil, while Peziza is a fungus that consumes cellulose, lignocellulose, and lignin and is abundant in mature vermicompost [75,76,77].
The archaeal genus Methanotorris, which was detected at the beginning of the process, is made up of strictly anaerobic species [78]. Vermicomposting is an aerobic procedure, and this genus’ abundance was not detected on the final day of the system.

3.7. Germination Index

After 120 days of vermicomposting, the final product was tested for potential phytotoxicity in barley seeds. Germination and radical growth were measured, and the Germination Index (GI) was calculated. According to Paradelo et al. [23] a GI > 80 is a good indicator at which no phytotoxicity of the tested product is observed. Our study’s vermicompost had no toxic effect on germination of barley seeds, and its GI was calculated to be 119.85, in accordance with previous studies, in which vermicomposts of different initial materials showed no phytotoxicity [79,80,81]. Before applying vermicompost products to soil, it is highly suggested to check potential phytotoxicity to avoid environmental risks. Majlessi et al. [82] indicated that even if the final product is stable (low C/N ratio, neutral pH, low electrical conductivity), there is a possibility that it will remain phytotoxic, and stability, maturity, and phytotoxicity tests are required.

4. Conclusions

Vermicomposting has numerous benefits, including reducing the amount of organic waste, producing high-quality fertilizer, and improving soil health and plant growth. It is also an environmentally friendly and sustainable alternative to manage organic waste. In the present study, the physicochemical, enzymatic, microbial, and microbiome activities during vermicomposting of winery waste were assessed. Excess enzymatic activities during the process, along with the physicochemical results (low C/N, total N increased, neutral pH), indicated that the final product was stable and mature and constituted a rich source of beneficial microorganisms, such as phosphate solubilizers, nitrogen-fixing bacteria, enzyme-producing bacteria, and bacteria that stimulated plant growth. However, more research is still required to fully understand the mechanism of action of winery-waste biotransformation into efficient biofertilizer and verify its effectiveness in real plant cultures.

Author Contributions

Conceptualization, Y.K.; data curation, I.K., G.M. and I.P.; formal analysis, I.K. and G.M.; funding acquisition, V.C. and Y.K.; investigation, I.K., G.M., I.P. and D.P.; methodology, G.M. and I.P.; project administration, V.C. and Y.K.; resources, V.C. and Y.K.; software, D.P.; supervision, Y.K.; validation, G.M., I.P. and D.P.; visualization, Y.K.; writing—original draft, I.K., G.M. and I.P.; writing—review and editing, Y.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research was cofinanced by the European Union and National Resources under the Operational Programme “Eastern Macedonia and Thrace” 2014–2020, AΜΘΡ7-0074893, MIS number: 5076618, “AgroWasteCompost”, and by the “Infrastructure of Microbiome Applications in Food Systems—FOODBIOMES” (MIS 5047291), which was implemented under the Action “Regional Excellence in R&D Infrastructures”, funded by the Operational Programme “Competitiveness, Entrepreneurship and Innovation” (NSRF 2014–2020) and cofinanced by Greece and the European Union (European Regional Development Fund).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

All data is available upon request.

Conflicts of Interest

The authors declare no conflict of interest.

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Figure 1. (a) Vermireactor system (VidaXL, The Netherlands) and (b) vermicomposting material loaded with earthworms.
Figure 1. (a) Vermireactor system (VidaXL, The Netherlands) and (b) vermicomposting material loaded with earthworms.
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Figure 2. Changes in pH values during vermicomposting of winery waste for 120 days.
Figure 2. Changes in pH values during vermicomposting of winery waste for 120 days.
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Figure 3. Changes in electrical conductivity, expressed as mmhos/cm at 25 °C, during vermicomposting of winery waste for 120 days.
Figure 3. Changes in electrical conductivity, expressed as mmhos/cm at 25 °C, during vermicomposting of winery waste for 120 days.
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Figure 4. Changes in C/N ratio during vermicomposting of winery waste.
Figure 4. Changes in C/N ratio during vermicomposting of winery waste.
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Figure 5. Oxygen uptake rate during vermicomposting, expressed as g O2 g/kg.
Figure 5. Oxygen uptake rate during vermicomposting, expressed as g O2 g/kg.
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Figure 6. Microbial population levels during vermicomposting of winery waste. Levels are expressed as log cfu/g of vermicompost. TAC: Total Aerobic Count, LAB: Lactic Acid Bacteria.
Figure 6. Microbial population levels during vermicomposting of winery waste. Levels are expressed as log cfu/g of vermicompost. TAC: Total Aerobic Count, LAB: Lactic Acid Bacteria.
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Figure 7. Bar plot of the mean relative genus abundances of the predominant bacteria during vermicomposting of winery waste from day 0 to day 120.
Figure 7. Bar plot of the mean relative genus abundances of the predominant bacteria during vermicomposting of winery waste from day 0 to day 120.
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Figure 8. Bar plot of the mean relative genus abundances of the predominant fungi during vermicomposting of winery waste from day 0 to day 120.
Figure 8. Bar plot of the mean relative genus abundances of the predominant fungi during vermicomposting of winery waste from day 0 to day 120.
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Table
Table 1. Concentrations of macro- and micronutrients during vermicomposting of winery waste for 120 days.
Table 1. Concentrations of macro- and micronutrients during vermicomposting of winery waste for 120 days.
NutrientsDay 0Day 15Day 45Day 90Day 120
Total Ca (% Dry Weight)6.71 ± 0.869.77 ± 0.475.21 ± 0.080.95 ± 0.211.86 ± 0.21
Active Ca (% Dry Weight)1.12 ± 0.036.46 ± 0.351.10 ± 0.040.31 ± 0.160.41 ± 0.01
CaCO3 (% Dry Weight)7.83 ± 0.8916.23 ± 0.816.31 ± 0.131.26 ± 0.061.45 ± 0.20
P (ppm)0.28 ± 0.040.41 ± 0.060.21 ± 0.010.10 ± 0.010.24 ± 0.07
K (ppm)4.23 ± 0.047.20 ± 0.143.78 ± 0.041.48 ± 0.040.35 ± 0.14
Mg (ppm)1.60 ± 0.144.88 ± 0.321.42 ± 0.120.78 ± 0.042.48 ± 0.71
Zn (ppm)35.80 ± 9.6247.88 ± 0.4637.25 ± 0.3557.55 ± 3.4655.25 ± 2.47
Mn (ppm)60.86 ± 0.7661.15 ± 1.254.48 ± 1.1062.08 ± 3.4629.90 ± 4.81
Fe (ppm)49.30 ± 1.5631.78 ± 2.345.75 ± 2.0536.35 ± 1.0618.00 ± 0.00
Cu (ppm)11.25 ± 1.0612.08 ± 1.5210.50 ± 1.4113.75 ± 0.3525.5 ± 7.78
Table
Table 2. Enzymatic activity of four different soil enzymes related to biochemical reactions during decomposition of organic matter by earthworms.
Table 2. Enzymatic activity of four different soil enzymes related to biochemical reactions during decomposition of organic matter by earthworms.
Enzymatic ActivityDay 0Day 15Day 45Day 60Day 90Day 120
Dehydrogenase (μg TPF/g dm/ 24 h)5.77 ± 0.926.1 ± 10.164.4 ± 9.342.3 ± 5.725.6 ± 1.227.0 ± 2.1
Acid phosphatase (μmoles pNP/g dm/ h)8.2 ± 1.728.6 ± 11.322.9 ± 6.522.5 ± 4.122.2 ± 4.16.7 ± 0.6
Urease
(μg N/g dm/ 2 h) 		42.1 ± 22.01546.0 ± 138.62579.6 ± 80.32448.2 ± 85.62240.7 ± 160.92505.2 ± 88.4
β-glycosidase (μmoles pNP/g dm/ h) 4.4 ± 0.218.5 ± 8.714.5 ± 0.010.2 ± 0.57.7 ± 0.53.6 ± 0.2
Table
Table 3. Changes in bacteria and fungi phylum abundances (%) during vermicomposting of winery waste, as determined by next-generation sequencing.
Table 3. Changes in bacteria and fungi phylum abundances (%) during vermicomposting of winery waste, as determined by next-generation sequencing.
Bacterial PhylaDay 0Day 120Fungal PhylaDay 0Day 120
Armatimonadetes0.020.06Ascomycota88.4397.70
Bacteroidetes1.9368.44Basidiomycota11.000.62
Candidatus saccharibacteria0.010.05Chytridiomycota0.210.24
Chloroflexi2.215.53Entomophthoromycota0.10.01
Cyanobacteria0.033.05Glomeromycota0.190.23
Planctomycetes0.5712.38Mucoromycota0.081.21
Proteobacteria95.2510.48
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Karapantzou, I.; Mitropoulou, G.; Prapa, I.; Papanikolaou, D.; Charovas, V.; Kourkoutas, Y. Physicochemical Changes and Microbiome Associations during Vermicomposting of Winery Waste. Sustainability 2023, 15, 7484. https://doi.org/10.3390/su15097484

AMA Style

Karapantzou I, Mitropoulou G, Prapa I, Papanikolaou D, Charovas V, Kourkoutas Y. Physicochemical Changes and Microbiome Associations during Vermicomposting of Winery Waste. Sustainability. 2023; 15(9):7484. https://doi.org/10.3390/su15097484

Chicago/Turabian Style

Karapantzou, Ioanna, Gregoria Mitropoulou, Ioanna Prapa, Dimitra Papanikolaou, Vasileios Charovas, and Yiannis Kourkoutas. 2023. "Physicochemical Changes and Microbiome Associations during Vermicomposting of Winery Waste" Sustainability 15, no. 9: 7484. https://doi.org/10.3390/su15097484

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