Plant material and soil characteristics
The olive plants (Olea europaea L.) were obtained from Leccino cultivar, by rooting plant cuttings propagation method, to ensure the production of uniform plant material.
Twenty-centimetre-long branch portions were cut from the tip of 1-year-old healthy olive branches; each branch was cut 0.3 cm below a leaf node. The olive cuttings were placed in container deep enough to support the development of new roots, with the cut end buried in premoistened planting-medium by 2.5–3.8 cm. About 2 months after rooting process, the olive rooted cuttings were put in individual 10-cm-diameter nursery pots filled with a mix of washed sand and milled peat (1:1, v:v). Successively, rooted olive cuttings were transplanted into 0.35-L pots filled with the same potting medium, under lightly shaded conditions. In winter 2017, uniform olive plants with a single main stem were transplanted into pots containing 3.6 L of soil amended with increasing concentrations (0, 0.5, 1.0, 2.0%) of two different types of sheep wool residues (SWR), as described below.
The soil used was collected from the topsoil layer (0–30 cm) of an olive orchard at the experimental site of the Italian National Research Council (CNR) located in Follonica (southern Tuscany, “Santa Paolina” experimental farm (Lat. 43° 49´ 3.032" N, Long. 11° 12´ 4.858" E; 12 m a.s.l.). Physical and chemical characteristics of the soil were as follows: sand 75.63%; silt 8.63%; clay 15.74% (texture ranging from clay loam to sandy loam, Soil Survey Division Staff, 2017); organic matter 0.75%; pH 6.68; total nitrogen 0.08%; available phosphorus 9.9 mg/kg; exchangeable potassium 232 mg/kg; electrical conductivity (EC) 0.705 mS/cm, cation exchange capacity (CEC) 5.6 meq/100 g, active carbonates < 0.1%.
Sheep wool residues (SWR) treatments
The SWR were acquired from the wool scouring company Carbon S.r.l., located in Vernio (Tuscany, Italy). Two SWR types, resulting from two different stages of the wool processing chain (regulated by the Commission Regulation—EU 1063/2012) were utilized: (1) white wool residue (WW), obtained from the mechanic beating of scoured wool and consisting of wool fibre and vegetal residues; (2) black wool residue (BW), obtained from the “carbonization” of the scoured and beaten wool with a solution of sulfuric acid. To determine carbon (C), nitrogen (N), hydrogen (H) and sulphur (S) contents, dry samples of wool residues (WW and BW) were analysed using a CHN Elemental Analyzer (Carlo Erba Instruments, mod 1500 series 2). Percentage contents of N, C, H and S were almost similar in WW and BW: N 11.62, C 44.09, H 6.84, S 2.72 and N 12.31, C 41.87, H 6.35, S 5.76, respectively.
To ensure that the culture substrate was physically homogeneous throughout the pots, the soil was prepared using an electric concrete mixer (100-L capacity, 0.4 hp, 23 rpm). Both control soil and soil–wool mixtures had 20 min of mixing inside the mortar. WW and BW soil–wool mixture samples with (0, 0.5, 1 and 2%) (w:w) were prepared, resulting in 7 different culture substrates, with three replicates for each treatment. The experiment was carried out outdoor from April 2017 to February 2020, at the Institute of BioEconomy (IBE) of CNR in Florence (Central Italy, Lat. 43°49′3.032" N and Long. 11°12′4.858′′ E, 40.5 mt. above s.l.). Pots were arranged in a completely randomized distribution, plants were organized with a fixed-position arrangement (North–South exposition) and placed in open field.
Climatic conditions were typical for Mediterranean regions, where, on average, there are 88 days per year with more than 0.1 mm of rainfall, the driest weather is in July (an average of 39.6 mm), while the wettest weather is in November (an average of 111.2 mm). July is the hottest month with a mean temperature of 24 °C, while January is the coldest having an average of 6 °C. No fertilization and agronomic treatments were supplied during all the experimentation time. The plants were well watered through the experiment thanks to a drip irrigation system with 300 mL each day during summer season. This solution allowed the supply of a regular and uniform water quantity, enough to cover the 80–100% of the field capacity.
Collar diameters of olive plants were measured in six subsequent dates and analysed by one-way ANOVA. The occurrence of significant differences among treatments was established performing the LSD post hoc test. The statistical analyses were carried out in IBM SPSS statistics version 24 software (IBM Corporation, Armonk, NY, USA).
From each pot, two soil cores of 3 cm diameter were collected, taking care to collect also the roots of the olive trees. The two soil subsamples were subsequently mixed in order to produce a single sample.
Diversity and composition of soil bacterial communities by PCR-DGGE
DNA extraction from soil samples and PCR amplification
Genomic DNA was extracted from 250 mg soil samples using DNeasy® PowerSoil Kit® (QIAGEN Group, Germantown, MD) according to the manufacturer’s protocol. The extracted DNA was stored at − 20 °C and subsequently used for the analysis of soil bacterial communities. The amplification of the variable region V3–V5 of 16S rDNA was carried out using the primers 341F (5’-CCT ACG GGA GGC AGC AG-3’) and 907R (5’-CCG TCA ATT CCT TTR AG TTT-3’) (Yu and Morrison 2004). At its 5’ end, the primer 341F had an additional 40-nucleotide GC-rich tail (5’-CGC CCG CCG CGC CCC GCG CCC GTC CCG CCG CCC CCG CCC G-3’). Amplification reaction was prepared in a final volume of 50 μL, using 10–20 ng of DNA, 5 μL of Ex Taq Buffer 10X (Takara Biotechnology), 1.25 U of Takara Ex Taq (Takara Biotechnology), 0.2 mM of each dNTP (Takara Biotechnology) and 0.5 μM of each primer (Eurofins). The fragment obtained was about 560 bp long. The reaction was carried out using an iCycler-iQ Multicolor Real-Time PCR Detection System (Bio-Rad) with the following denaturation, amplification and extension procedure: at 95 °C for 1′; at 94 °C for 30′′, at 60 °C for 30′′, at 72 °C for 30′′ (for 35 cycles); at 72 °C for 5′. The presence of amplicons was confirmed by electrophoresis in 1.5% (w/v) agarose gels stained with 20,000X REALSAFE Nucleic Acid Staining Solution (Durviz, s.l., Valencia, Spain). All gels were visualized using UV light and captured as TIFF format files using the UVI 1D v. 16.11a program for the FIRE READER V4 gel documentation system (Uvitec Cambridge, Eppendorf, Milan, Italy).
DGGE and profile analyses
For DGGE analyses, 20 μL of amplicons were separated in 8% (w/v) polyacrylamide gels with a 36–58% urea–formamide gradient, using the DCode™ Universal Mutation Detection System (Bio-Rad, Milan, Italy). Gels were run at 90 V and 60 °C for 16 h, stained for 30′ in 500 mL of TAE buffer 1X containing 50 μL of Sybr Gold Nucleic Acid Gel Stain (Thermo Fisher Scientific, Italia) and visualized as previously described. The sample V18 was added on each side and in the centre of DGGE gels as DGGE marker.
DGGE profiles were digitally processed with BioNumerics software version 7.6 (Applied Maths, St-Martens-Latem, Belgium) and bacterial community composition was assessed by cluster analysis of DGGE profiles, as reported in Palla et al. (2018). Similarities between DGGE patterns were calculated by determining Pearson’s similarity coefficients for the total number of lane patterns from the DGGE gel using the band-matching tool with an optimization of 1%. The similarity coefficients were then used to generate the dendrogram utilizing the clustering method UPGMA (unweighted pair group method using arithmetic average).
DGGE banding data were used to estimate four different indices treating each band as an individual operational taxonomic unit (OTU). Richness (S) indicates the number of OTUs present in a sample and was determined from the number of fragments. Shannon–Weaver (Hs) and the dominance index of Simpson (D) were calculated using the equations Hs = −Σ(Pi x lnPi) and D = ΣPi2, respectively, where the relative importance of each OTU is Pi = niN−1, and ni is the peak intensity of a band and N is the sum of all peak intensities in a lane. Evenness index (E), which allows the identification of dominant OTUs, was calculated as E = H (lnS)−1.
One-way ANOVA was applied to diversity indices with SWR/soil ratios as the variability factor. The means were compared by the Tukey's test (P < 0.05). Analyses were carried out with the SPSS version 23 software (IBM Corp., Armonk, NY, USA).
DGGE band sequencing
The main bands of DGGE profiles were excised from the gels for sequencing at the Eurofins Genomics MWG Operon (Ebersberg, Germany) as reported in Agnolucci et al. (2019). DNA was extracted by eluting for 3 days in 50 μL UltraPure™ DNase/RNase-Free Distilled Water (Invitrogen) at 4 °C. Two microliters of the supernatant diluted 1:10 were used to re-amplify the V3–V5 region of the DNA, using the 341F primer without the GC clamp. PCR products were than purified with the QIAquick PCR Purification Kit (Qiagen) according to the manufacturer's protocol, quantified and 5’ sequenced by Eurofins Genomics (Ebersberg, Germany). Sequences were analysed using BLAST on the NCBI web (http://blast.ncbi.nlm.nih.gov/Blast.cgi). The related sequences were collected and aligned using MUSCLE (Edgar 2004a, b), and phylogenetic trees were constructed using the Neighbor-Joining method based on Kimura’s 2-parameter model (Kimura 1980) in Mega X software (http://www.megasoftware.net/) with 1000 bootstrap replicates. Sequences were submitted to the European Nucleotide Archive under the accession numbers: from OU745530 to OU745538 (WW) and from OU745381 to OU745389 (BW) (Project: PRJEB47775).
Mycorrhizal inoculum potential of the soil
Mycorrhizal inoculum potential (MIP) bioassay was performed to verify the activity of native AMF occurring in the soil of pot-grown olive plants and was assessed using Cichorium intybus L. as host plant, as described in Turrini et al. (2018). Briefly, soil samples from each pot (50 g soil) were dried, sieved using a 4-mm sieve, and put in 50-mL tubes. For each MIP determination, six replicated tubes were used, filled with 45 mL of soil and sown using the biotest plant. Then, they were put in sun-transparent bags and maintained in a growth chamber at 27 °C under a 16/8 h light/dark daily cycle until harvest. Four days after germination, plants were thinned to three per tube. Each tube was watered as needed. Plants were harvested 28 days after sowing and shoots were excised and discarded. Roots were removed from soil, washed with tap water, then cleared in 10% KOH in a 80 °C water bath for 15′, neutralized in 2% aqueous HCl, and stained with 0.05% Trypan blue in lactic acid. The percentage of AMF colonization was calculated using a dissecting microscope at × 25 or × 40 magnification and the gridline intersect method (Giovannetti and Mosse, 1980).
The percentage of AMF root colonization was determined on 5 g of thoroughly washed olive root samples, after clearing and staining, as described above. Percentages of AM fungal root colonization were assessed on representative root samples from each plant under a dissecting microscope (Wild, Leica, Milano, Italy) at × 25 or × 40 magnification by the gridline intersect method (Giovannetti and Mosse 1980).
Data of MIP and olive root mycorrhizal colonization (after arcsin transformation) were analysed by one-way ANOVA. The occurrence of significant differences among treatments was established performing the Tukey post hoc test. The statistical analyses were carried out in IBM SPSS statistics version 23 software (IBM Corporation, Armonk, NY, USA).