Twenty-four bacterial strains, belonging to the microbial collection of the Department of Agricultural Sciences, division of Microbiology of the University of Naples Federico II, were used in this study. These strains, previously isolated from lignocellulosic biomass of A. donax, E. camaldulensis, and P. nigra during biodegradation under natural conditions (Ventorino et al., 2015), were identified as Actinobacteria on the basis of their colony morphology, microscopic features (phase-contrast microscopy, shape, dimension, and presence of spores) and biochemical characteristics (Gram-stains, catalase activity).

Molecular identification was performed by 16S rRNA gene sequencing. Fast DNA SPIN kit for soil (MP Biomedicals, Illkirch Cedex, France) was used to extract and purify the total genomic DNA according to the supplier's recommendations. Approximately 50 ng of DNA was used as template for PCR assays. Synthetic oligonucleotide primers fD1 (5′-AGAGTTTGATCCTGGCTCAG-3′) and rD1 (5′-AAGGAGGTGATCCAGCC-3′) were used to amplify the 16S rRNA gene as reported by Palomba et al. (2011). The PCR conditions were as described by Ventorino et al. (2016a). Amplicons were purified using the QIA quick gel extraction kit (Qiagen S.p.A., Milan, Italy) after visualization by agarose (1.5% wt/vol) gel electrophoresis at 100 V for about 1 h. The DNA sequences were determined and analyzed as previously reported (Pepe et al., 2013a), and were compared to the 16S ribosomal RNA sequences database of GenBank nucleotide data library using the BLAST software at the National Centre of Biotechnology Information website (http://www.ncbi.nlm.nih.gov/Blast.cgi). Multiple nucleotide alignments of the nearly full-length 16S rRNA sequences of bacterial isolates and type strains within each of the defined species were carried out using the ClustalW program from MEGA version 4.0 (Tamura et al., 2007). The nucleotide sequences of the type strains were recovered from the Ribosomal Database Project (RDP—https://rdp.cme.msu.edu). The phylogenetic tree was inferred by the Neighbor-Joining method with the Maximum Composite Likelihood model in MEGA4 program with bootstrap values based on 1000 replications.

Endo and exo-cellulase, cellobiase, xylanase, pectinase, ligninase, peroxidase, and laccase activities were evaluated in all strains by qualitative and semi-quantitative agar spot methods as previously described (Ventorino et al., 2015). Briefly, bacterial cell suspensions (0.5 of McFarland Turbidity Standard corresponding to approximately 1.5 × 10⁸ CFU mL⁻¹) were spotted in triplicate on selective solid media. Cellulase, pectinase and laccase activities were recorded as the “Indices of Relative Enzyme Activity, (I_CMC or I_PEC or I_LAC) = diameter of clearing or halo zone/colony diameter” using carboxymethylcellulose (CMC) agar, Pectin agar or liquid basal medium (LBM) agar (supplemented with 1 g L⁻¹ of 2.2′-azino-bis (3-ethylbenzothiazoline-6-sulphonic acid [ABTS]) media, respectively (Pepe et al., 2013b; Ventorino et al., 2015). Exo-cellulase activity was estimated by inoculating the strains on Avicel agar and observing the development of bacterial colonies after incubation. Cellobiase and xylanase activities were detected by observing a clear zone around the colonies after incubation on cellobiose agar (Ventorino et al., 2015) or Luria-Bertani (LB) agar medium (Oxoid, Milan, Italy) supplemented with 0.05% Remazol brilliant blue-R and a 0.5% solution of sonicated xylan (Sigma-Aldrich, Milan, Italy) as described by Ko et al. (2012). Ligninase activity was assayed using the culture medium guaiacol agar (CGA) supplemented with 2 g L⁻¹ of alkali lignin (Okino et al., 2000) or 2 g L⁻¹ of vegetable biomass powder from A. donax (Ventorino et al., 2015). Finally, LBM agar medium supplemented with 0.1 g L⁻¹ of Azure-B was used to detect peroxidase activity as previously reported (Ventorino et al., 2015).

The Streptomyces strains were grown on plate count agar (PCA) medium (0.5% tryptone, 0.25% yeast extract, 0.1% glucose, and 1.5% agar) for 14 days at 28°C. The pre-inoculum was prepared by adding one agar plug (4 mm) from a 2-week old plate into 10 mL of liquid medium containing 10 g L⁻¹ carboxymethylcellulose (CMC), 0.5 g L⁻¹ yeast extract, 2.5 g L⁻¹ (NH₄)₂SO₄, 2.7 g L⁻¹ KH₂PO₄, 5.3 g L⁻¹ Na₂HPO₄, 0.2 g L⁻¹ NaCl, 0.2 g L⁻¹ MgSO₄, and 0.05 g L⁻¹ CaCl₂. After 48 h of incubation on a rotary shaker (250 rpm) at 28°C, the spore concentration was determined by using the Bürker-chamber through decimal serial dilutions in isotonic Ringer's solution. Fermentation was carried out in 100 mL plugged Erlenmeyer flasks, each containing 20 mL of the same medium used for the preinoculum. A quantity of 2 × 10⁶ spores mL⁻¹ was inoculated. The flasks were incubated for 10 days at 28°C on a rotary shaker (200 rpm). Samples were withdrawn every 24 h for 10 days and the supernatants after removal of cells by three centrifugations (4629 × g, 30 min each at 4°C) used for measurement of extracellular endo-cellulase activity. The assay was performed by using Azo-CMC (Megazyme, Ireland) as substrate, following supplier's instructions and determined by referring to Trichoderma sp. endo-cellulase standard curve.

The optimum temperature and pH of the cellulase activity of four selected Streptomyces strains was determined using the supernatant of bacterial cultures recovered and concentrated by ultrafiltration with a 10 kDa polyethersulfone membrane (Millipore Corporation, Bedford, MA, USA).

To assess the optimum temperature, the enzymatic activity assay was performed at 40, 45, 50, and 55°C by using Azo-CM-Cellulose (Azo-CMC; Megazyme, Ireland) dissolved in 50 mM Na citrate at pH 5.0 as substrate, following supplier's instructions.

To determine the optimum pH of cellulase activity, the experiments were performed at 40°C in 50 mM Na-citrate buffer, at pH 5.0 and 6.0, using Azo-CMC (Megazyme, Ireland) as substrate dissolved in the above-mentioned buffer, following supplier's instructions.

The reported results correspond to mean values of the three independent experiments each one performed in three replicates.

A. donax biomass, pretreated according to Garbero et al. (2010) and De Bari et al. (2013) was utilized as substrate in biotransformation experiments carried out in capped tubes, on the rotary shaker ThermoMixer C (Eppendorf, Milan, Italy) at 50°C up to 72 h. The reaction mixture contained 50 mM sodium citrate buffer pH 5.0 or 6.0, the enzymatic cocktail and a concentration of pretreated biomass of 5% (w/v) in a total volume of 2.5 mL. The components of the enzymatic mix, Accelerase 1500, Accelerase BG and Accelerase XY, (Mix 1), were obtained from Genencor and were applied at the following loadings, expressed as units per gram of pretreated substrate: 5.4, 145, and 4000, respectively.

Enzymatic extracts endowed with endo-cellulase activity from four selected Streptomyces argenteolus strains were obtained by removal of cells through three subsequent centrifugations (5500 × g, 30 min each at 4°C) and concentration by using the stirred ultrafiltration Amicon® system (Millipore Corporation, Bedford, MA, USA) with a 10 kDa polyethersulfone membrane. These crude extracts were utilized in replacement of the commercial counterpart represented by the Accelerase 1500. Samples were collected at different time intervals (0, 48, and 72 h), cooled on ice, centrifuged at 16,500 × g for 30 min at 4°C and the amount of sugars released were quantified in the recovered supernatants, as described below. The saccharification yields, that are means of three replicates, were expressed as percentages with respect to the sugar content of the pretreated material before the hydrolysis (Garbero et al., 2010; De Bari et al., 2013).

The sugars released upon enzymatic hydrolysis were determined by high-performance liquid chromatography (HPLC) using a system 7Q (Dionex, California, USA), equipped with an anionic exchange column (Carbopac PA-100) and a pulsed electrochemical detector. Glucose and xylose were separated with 16 mM sodium hydroxide at a flow rate of 0.25 mL min⁻¹, and identified by the respective standards. Fucose was used as internal standard.

Protein concentration of crude enzyme preparation was determined by using Bradford reagent of Biorad (München, Germany) following supplier's instructions. Bovin serum albumin (BSA) was used to set up the standard curve.

Precipitated proteins were dissolved in 1 mL of Tris 300 mM pH 8.0, urea 6 M, EDTA 10 mM, and disulfide bridges were reduced with 1,4-Dithiothreitol (DTT) 10 mM (final concentration) at 37°C for 2 h and then cysteines alkylated by adding iodoacetamide (IAM) 100 mM (final concentration) at room temperature for 30 min in the dark. The protein sample was desalted by size exclusion chromatography on a Sephadex G25M column (GE Healthcare). Protein containing fractions were concentrated by evaporation under vacuum to 100 μL. Enzymatic digestion was then performed by adding 2 μg trypsin in a sample volume of 100 μL. After incubation at 37°C for 16 h, the surnatant was recovered by centrifugation at 45 × g, and the peptide mixture was filtered through a 0.22 μm PVDF membrane (Millipore), concentrated and purified using reverse phase C18 Zip Tip pipette tips (Millipore). Peptides were eluted with 20 μL of a solution made of 50% acetonitrile/0.1% formic acid in MilliQ water and analyzed by LC-MS/MS. LC-MSMS analyses were carried out on a 6520 Accurate-Mass Q-TOF LC/MS System (Agilent Technologies, Palo Alto, CA) equipped with a 1200 HPLC system and a chip cube (Agilent Technologies). After loading 10% of the recovered sample, the peptide mixture was first concentrated and washed in 40 nL enrichment column (Agilent Technologies chip), with 0.1% formic acid in 2% acetonitrile as the eluent. Peptides were then fractionated on a C18 reverse-phase capillary column (Agilent Technologies chip) at a flow rate of 400 nL min⁻¹, with a linear gradient of eluent B (0.1% formic acid in 95% acetonitrile) in A (0.1% formic acid in 2% acetonitrile) from 7 to 80% in 50 min. Data analysis was performed using data-dependent acquisition of one MSscan (mass range from 300 to 1800 m/z) followed by MS/MS scans of the five most abundant ions in each MS scan. MS/MS spectra were measured automatically when the MS signal surpassed the threshold of 50,000 counts. Double and triple charged ions were preferably isolated and fragmented. The acquired MS/MS spectra were transformed in Mascot generic format (.mgf) and used for protein identification in the unreviewed set of protein entries that are present in the NCBInr database for Actinobacteria, with a licensed version of MASCOT software (www.matrixscience.com) version 2.4.0.

Additional MASCOT search parameters were: peptide mass tolerance 10 ppm, fragment mass tolerance 0.6 Da, allowed missed cleavages up to 3, carbamidomethylation of cysteines as fixed modification, oxidation of methionine, and pyro-Glu N-term Q, as variable modifications. Only doubly and triply charged ions were considered. Ions score was −10 log (P), where P is the probability that the observed match is a random event. Individual ion scores >43 indicated identity or extensive homology (P < 0.05). Protein scores were derived from ion scores as a non-probabilistic basis for ranking protein hits (http://www.matrixscience.com/help/interpretation_help.html).

Trypsin, dithiothreitol, iodoacetamide and NH₄HCO₃ were purchased from Sigma-Aldrich. All other reagents and solvents were of the highest purity available from Baker (Mumbai, MH, India).

The 16S rRNA gene sequences obtained from bacterial strains were deposited in the GenBank nucleotide database under accession numbers from KX431234 to KX431257 (http://www.ncbi.nlm.nih.gov).

One-way ANOVA followed by Tukey's HSD post-hoc for pair-wise comparison of means (at P < 0.05) was used to assess the difference in the enzymatic activities of bacterial strains such as I_CMC, I_PEC, and Azo-CMCase as well as in the glucose and xylose yields (%). Statistical analyses were performed using SPSS 19.0 statistical software package (SPSS Inc., Cary, NC, USA).

Twenty-four lignocellulosic-adapted bacterial strains, with potential enzymatic degradation activities on lignocellulosic biomass, were grown on starch casein agar medium and characterized from a phenotypic point of view by analysis of colony and cell morphology, presence of spores, gram reaction, and catalase activity. All strains were Gram and catalase positive and produced a spore-bearing aerial mycelium. They were identified as Actinobacteria and grouped in seven phenotypes on the basis of colony morphology (Figure 1).

As shown in Table 1, the strains were identified by 16SrRNA gene sequencing showing that all strains belonged to the genus Streptomyces (S.). In particular, the most highly represented species was S. argenteolus representing 50% of strains (phenotype A), while eight strains were identified as members of species Streptomyces flavogriseus (synonym S. flavovirens) (phenotype B) and Streptomyces fimicarius (synonyms S. acrimycini, S. baarnensis, S. caviscabies, and S. flavofuscus) (phenotype C). Finally, the other 4 strains belonged to the species Streptomyces drozdowiczii, Streptomyces rubrogriseus, Streptomyces albolongus, and Streptomyces ambofaciens (phenotypes D, E, F, and G).

The nearly full-length gene sequences (>1400 bp) of the Streptomyces bacteria identified were grouped by phylogenetic analysis into seven different clusters, generating a consensus tree. The phylogenetic tree was generated from the distance data using the Neighbor-Joining method with the Maximum Composite Likelihood model in the MEGA4 Program (Figure 2). The nucleotide sequences of related type strains of different species were included in the data set. High bootstrap values were observed and indicated significant branching points in the phylogenetic tree. Strains representative of the dominant species of S. argenteolus were placed in a cluster with bootstrap values higher than 70%. The other species were included in clusters with bootstrap values from 80 to 99%, except for the cluster of strains identified as S. fimicarius (41%, Figure 2).

The Streptomyces strains were assayed for different enzymatic activities specific for the degradation of the lignocellulosic biomass using qualitative and semi-quantitative screening. One hundred percent of the bacterial strains exhibited multiple enzymatic activities (from three to six) as shown in Table 2. Endo-cellulase activity was highlighted in all Streptomyces strains with I_CMC values ranging from 20 to 40. In particular, five strains (S. argenteolus AE58P, S. albolongus AP71X, S. argenteolus AE82P, S. argenteolus AA85A, and S. argenteolus AP51A) showed the highest activity with I_CMC > 30 (P < 0.05). Furthermore, 16 strains showed endo-cellulase activity between 25 ≤ I_CMC ≤ 30, whereas three other strains (S. argenteolus AP73A, S. flavogriseus AA72A, and S. argenteolus AP64C) showed I_CMC <25. Moreover, all bacterial strains showed also exo-cellulase activity, since were able to grow on culture medium containing microcrystalline cellulose (Avicel) as sole carbon source, and 22 Streptomyces strains exhibited pectinase activity with I_PEC ≥ 10 (Table 2). The highest pectinase activity was observed in the strains S. argenteolus AE82P and S. argenteolus AA63A (I_PEC = 28, P < 0.05).

Xylanase activity was detected in approximately 80% of bacterial strains, whereas only eight strains were able to hydrolyze cellobiose.

Finally, the two strains identified as S. argenteolus AP51A and S. flavogriseus AA72A showed a high laccase activity (I_LAC = 20) in LBM agar medium supplemented with ABTS, whereas peroxidase activity was detected only in the strain S. flavogriseus AA62C. Lignin hydrolysis was not detected in any of the strains tested (Table 2).

A further screening of the 24 lignocellulosic-adapted Streptomyces strains was carried out in liquid medium containing the sole carbon source CMC as inducer substrate for endo-1,4-ß-glucanases. In Table 3, the values of the maximum AZO-CMCase activity measured for each strain and the corresponding time of production are reported. This screening confirmed three of the five Streptomyces strains with the highest activity in solid medium (S. argenteolus AE58P, S. argenteolus AE82P, and S. argenteolus AP51A) as the best endo-cellulase producers in liquid medium, whilst S. albolongus AP71X and S. argenteolus AA85A strains proved to be less effective producers than on solid medium. The S. argenteolus AE58P produced the highest endo-1,4-ß-glucanase activity level (0.41 ± 0.05 U mL⁻¹; P < 0.05) after 6 days of fermentation and was chosen as reference. The maximum values of endo-cellulase production for S. argenteolus AP51A, S. argenteolus AE710A, S. argenteolus AE82P, and S. argenteolus AE612P were at least 50% in comparison to the maximum value shown by S. argenteolus AE58P. These strains were classified as high producers. The strains S. argenteolus AP73A, S. fimicarius AA88P, S. flavogriseus AA62C, S. drozdowiczii AA86P, S. argenteolus AP71C, S. argenteolus AE61P, S. flavogriseus AA72A, S. fimicarius AE78P showed a maximum value of activity between 34 and 45 % in comparison to the reference value. The other strains produced an endo-cellulase activity less than 34% of the reference.

In order to evaluate the best conditions for long-term storage, the residual enzymatic activity of the best five endo-cellulase producing strains (S. argenteolus AE58P, S. argenteolus AP51A, S. argenteolus AE710A, S. argenteolus AE82P, and S. argenteolus AE612P) were assayed after up to 30 days of storage at 4, −20 and −80°C. The S. argenteolus AE710A retained most of the endo-1,4-ß-glucanase activity after 30 days (60% at 4°C and more than 80% at −20 and −80°C). Moreover, the S. argenteolus AE58P preserved 50–60% enzymatic activity after 30 days at all three tested temperatures, while the S. argenteolus AE82P and S. argenteolus AP51A strains retained 80–100% of activity at −20 and −80°C up to 30 days and lost 50% activity after 6 days at 4°C. By contrast, the tested storage conditions were not suitable for the endo-cellulase(s) produced by the S. argenteolus AE612P (data not shown). Based on these results, this strain was not subjected to further experiments.

The optimal temperature and pH of endo-cellulase activities produced by S. argenteolus AE58P, S. argenteolus AP51A, S. argenteolus AE710A, and S. argenteolus AE82P were also evaluated. In the Na-citrate buffer, the cellulase activity produced by the analyzed Streptomyces strains showed an optimum at same pH value (5.0) (Figure 3). Furthermore, the optimum temperature for all the strains was 50°C (Figure 4), which represents the condition mostly used for the enzymatic hydrolysis of lignocellulosic biomasses.

Enzymatic crude extracts of the four S. argenteolus strains (AE58P, AE710A, AE82P, and AP51A) selected as the best endo-cellulase producers in liquid medium were concentrated by ultrafiltration and adopted to perform saccharification experiments on pretreated A. donax whose macromolecular composition in terms of percentage of glucans, xylans, and Klason lignin, was 36, 20, and 21%, respectively. The different degrees of cellulose and xylan hydrolysis were evaluated by determining the kinetics of glucose and xylose release during a 72 h incubation of the pretreated A. donax biomass in the presence of cellulose degrading enzymes (cellulase and β-glucosidase) and xylan related activities (xylanase and β-xylosidase). To this end the monosaccharide yields were evaluated on samples collected at different time intervals (0, 48, and 72 h). Cellulose and xylan conversion, when conducted for 72 h at 50°C in the presence of the commercial (hemi)cellulolytic activities from Genencor (Accellerase 1500) (see Mix 1 composition under Materials and Methods Section and Table 4), gave glucose and xylose yields of 30.17 and 68.9%, respectively. The replacement of the cellulolytic commercial preparation with the endo-cellulase from S. argenteolus AE58P (Mix 2) allowed glucose yield to reach, after 72 h, almost 76% of that obtained in the presence of the commercial counterpart after the same incubation time (Table 4). Significantly lower glucose yields (%) were obtained after 72 h when cellulases from S. argenteolus AE710A and S. argenteolus AE82P (Mix 3 and 5, Table 4) were utilized instead of the activities from the strain S. argenteolus AE58P. The most limited glucan hydrolysis was reached with the preparation from S. argenteolus AP51A (Mix 4, Table 4) and accounted for 43.6% of the maximum value (22.05%) reached with Mix 2. Moreover, two-fold increase in protein loading for the biocatalysts produced by all strains analyzed (Mix 2A, 3A, 4A, and 5A, Table 4) did not cause additional benefits to cellulose conversion. Also for the xylan hydrolysis the significant highest yields were obtained with the enzymatic extract from S. argenteolus AE58P (Mix 2) that accounted for 74% of the value obtained with the commercial biocatalysts (68.9% reached with Mix 1).

In order to improve the glucose and xylose yields, biotransformation experiments with Mix 1 and 2 were performed at pH 6.0 but the maximum glucose yields obtained after 72 h of incubation (29.50 and 13.88%) were lowered by 13.5 and 44 %, respectively compared to the result obtained at pH 5.0 (data not shown).

Due to the promising results obtained by using the enzymatic crude extracts from S. argenteolus AE58P in the pretreated A. donax saccharification experiments, the proteins putatively responsible for endo-cellulase activity in this strain were tentatively identified by proteomic strategy.

The secreted proteins from the above mentioned strain were precipitated by the addition of ammonium sulfate, recovered and diafiltered in 50 mM Na₂HPO₄, pH 6.

The peptide mixture obtained by enzymatic digestion by trypsin was analyzed by LC-MS/MS and query of the NCBInr database for Actinobacteria with a licensed version of MASCOT software (www.matrixscience.com). The identified proteins are reported in Table 5.

Several proteins were confidently identified in different Streptomyces spp., eight of which belong to the class of Carbohydrate active enzymes. In detail, three proteins were annotated as containing carbohydrate-binding modules (CBM), three annotated as cellulases of family GH5, one as a cellulase of family GH12, and another as a cellulase that is homologous to glycosyl hydrolases of GM109 class.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.