Secreted protein extract analyses present the plant pathogen Alternaria alternata as a suitable industrial enzyme toolbox

Accepted Manuscript Secreted protein extract analyses present the plant pathogen Alternaria alternata as a suitable industrial enzyme toolbox

L. García-Calvo, R.V. Ullán, M. Fernández-Aguado, A.M. García-Lino, R. Balaña-Fouce, C. Barreiro PII: DOI: Reference:

S1874-3919(18)30062-9 https://doi.org/10.1016/j.jprot.2018.02.012 JPROT 3043

To appear in:

Journal of Proteomics

Received date: Revised date: Accepted date:

3 November 2017 1 February 2018 4 February 2018

Please cite this article as: L. García-Calvo, R.V. Ullán, M. Fernández-Aguado, A.M. García-Lino, R. Balaña-Fouce, C. Barreiro , Secreted protein extract analyses present the plant pathogen Alternaria alternata as a suitable industrial enzyme toolbox. The address for the corresponding author was captured as affiliation for all authors. Please check if appropriate. Jprot(2017), https://doi.org/10.1016/j.jprot.2018.02.012

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

ACCEPTED MANUSCRIPT Secreted protein extract analyses present the plant pathogen Alternaria alternata as a suitable industrial enzyme toolbox García-Calvo L1, Ullán RV2, Fernández-Aguado M1, García-Lino AM3, Balaña-Fouce R4, Barreiro C1,5*.

INBIOTEC (Instituto de Biotecnología de León). Avda. Real 1 - Parque Científico de León

PT

1

24006, León, Spain.

mAbxience, Upstream Production. Parque Tecnológico de León, Julia Morros, s/n,

RI

2

3

SC

Armunia, 24009 León, Spain.

Área de Fisiología, Departamento de Ciencias Biomédicas, Universidad de León, Campus

4

NU

de Vegazana s/n, 24071 León, Spain.

Departamento de Ciencias Biomédicas, Universidad de León, Campus de Vegazana s/n,

5

MA

24071 León, Spain.

Departamento de Biología Molecular, Universidad de León, Campus de Ponferrada, Avda.

PT E

D

Astorga s/n, 24401, Ponferrada, Spain.

AC

proteome.

CE

Keywords: Alternaria, Fungi, plant pathogen, feruloyl esterase, lignocellulose, extracellular

*Corresponding author: Dr. Carlos Barreiro. e-mail: [email protected]. Phone: +34 987210308; Fax: +34 987210388. Address: INBIOTEC (Instituto de Biotecnología de León), Parque Científico de León, Avda. Real 1, 24006 León, Spain.

1

ACCEPTED MANUSCRIPT Abstract Lignocellulosic plant biomass is the most abundant carbon source in the planet, which makes it a potential substrate for biorefinery. It consists of polysaccharides and other molecules with applications in pharmaceutical, food and feed, cosmetics, paper and textile industries. The exploitation of these resources requires the hydrolysis of the plant cell wall, which is a complex process. Aiming to discover novel fungal natural isolates with lignocellulolytic

PT

capacities, a screening for feruloyl esterase activity was performed in samples taken from different metal surfaces. An extracellular enzyme extract from the most promising candidate,

RI

the natural isolate Alternaria alternata PDA1, was analyzed. The feruloyl esterase activity of

SC

the enzyme extract was characterized, determining the pH and temperature optima (pH 5.0 and 55-60 ºC, respectively), thermal stability and kinetic parameters, among others. Proteomic analyses derived from two-dimensional gels allowed the identification and

NU

classification of 97 protein spots from the extracellular proteome. Most of the identified proteins belonged to the carbohydrates metabolism group, particularly plant cell wall

MA

degradation. Enzymatic activities of the identified proteins (β-glucosidase, cellobiohydrolase, endoglucanase, β-xylosidase and xylanase) of the extract were also measured. These findings

AC

CE

PT E

D

confirm A. alternata PDA1 as a promising lignocellulolytic enzyme producer.

2

ACCEPTED MANUSCRIPT 1. Introduction The Kingdom Fungi is a complex lineage that comprises 3.5 to 5.1 million species, distributed in most habitats on Earth [1]. Fungi possess a distinctive heterotrophic and absorptive nutrition, which cause them to act as decomposers, mutualists, pathogens and parasites [2]. Due to this versatile enzymatic machinery and secondary metabolite production, they play a key role in society. On the one hand, they are responsible for substantial

PT

economic losses in agriculture and health [3,4], as well as other impacts, such as diseases affecting poultry, cattle and human beings, especially in low-income countries [5–7]. On the

RI

other hand, fungi are profitable microbial cell factories that constitute the most important

SC

group (after Actinobacteria) of pharmaceutical secondary metabolites producers (cholesterol lowering drugs, antibiotics, immunosuppressants, etc.) [8]. Moreover, they are also being employed as heterologous and endogenous enzyme producers. Many filamentous fungi (e.g.

NU

Aspergillus spp., Trichoderma spp.) are natural excellent producers of extracellular enzymes, which makes them interesting candidate hosts for the production of recombinant proteins.

MA

Some of these fungi are capable of secreting very high amounts of proteins and are currently used at an industrial level [9–11].

D

The global industrial enzymes market is a field of increasing economic significance, with an estimated value of 8.18 billion USD in 2015 and a steady annual growth projection for the

PT E

next years [12]. In this regard, proteomic approaches are interesting to study fungal strains with potential industrial uses. In the past years, the number of spectrometry-based proteomic studies of fungi has increased considerably. However, fungal Proteomics is still

CE

underdeveloped and not as thorough for filamentous fungi as it is for other biological systems such as humans and yeast [13]. Greater advances in fungal Proteomics are expected as the

AC

number of sequenced genomes from ascomycetes increases [14]. The evolution of genome sequencing technology has favored the elucidation of several genomes of plant pathogenic fungi, such as Alternaria spp. The Alternaria Genome Database (AGD) is an online resource that includes data from genome sequences of 25 Alternaria spp. species [15]. Alternaria (Ascomycota, Pleosporales) is a ubiquitous fungal genus that includes saprobic, endophytic and pathogenic species. Within this genus, Alternaria alternata is capable of damaging a wide spectrum of plant hosts and may also cause diseases in humans. It can produce metabolites that are toxic to plants, animals and humans, causing upper respiratory tract infections in immunocompromised people and allergic responses [16]. Nevertheless, as an

3

ACCEPTED MANUSCRIPT opportunistic plant pathogen, Alternaria spp. is also an interesting hydrolytic enzyme producer. Among the secreted fungal proteins with positive economic impacts, lignocellulolytic enzymes are of particular interest. These enzymes present applications in the industrial hydrolysis processes for the deconstruction of plant cell walls. Lignocellulosic plant biomass is the most abundant carbon source [17,18] and it constitutes an exploitable alternative to

PT

fossil fuels [19]. The hydrolysis of plant biomass also causes the release of useful molecules: platform chemicals for related commodity compounds and bioactive molecules, such as

RI

hydroxycinnamic acids [18,20]. Among these acids, one of the most relevant is ferulic acid, which displays antioxidant properties and has industrial and medical applications (anti-

SC

inflammatory, antimicrobial, anticarcinogenic, antithrombotic, antiviral) [21]. As a result of the complexity and recalcitrance of the plant cell wall structure, several enzymes are needed

NU

to achieve the release of these useful compounds [17]. Feruloyl esterases (FAEs, EC 3.1.1.73) are auxiliary enzymes in these degradation processes, through the hydrolysis of the ester

MA

bonds that join together hydroxycinnamic acids (e.g. ferulic acid) and lignin-carbohydrate complexes. These cross-links confer stability to the plant cell wall and, thus, its rupture aids the complete cell wall deconstruction [22]. Some of the most used physicochemical

D

pretreatments (hydrothermal, acid, alkali, oxidizing agents) are also capable of breaking these

PT E

bonds and are often combined with lignocellulolytic enzymes (biological treatments) in order to enhance the hydrolysis. However, although physicochemical pretreatments are still widely used due to their higher rates of hydrolysis, biological pretreatments present several

CE

advantages: they are safe, environmentally friendly and less energy consuming [23]. These important advantages emphasize the need for further knowledge about lignocellulolytic

AC

enzymes, such as FAEs. FAEs have applications in fields like biorefinery, textiles, pulp and paper, cosmetics, food, feed and pharmaceutical industries [24–26]. Fungal FAEs have been widely studied throughout the last decades [27]. However, according to the prevalence of putative FAEs on recently described fungal genomes [24], there is still a substantial amount of FAEs to be discovered and further investigated. FAEs are an enzymatic group distributed among simple and complex organisms, existing in different types of ecosystems [28]. Considering the ability of some lignocellulolytic fungi to grow adhered to surfaces [29–31] and the possibility of finding new enzymatic activities in novel environments, the aim of this study was to perform a screening of microorganisms with FAE activity from environmental samples taken from metal surfaces. Although metal is not a lignocellulosic substrate, it 4

ACCEPTED MANUSCRIPT constitutes an unexpected niche for fungal strains capable of producing new lignocellulolytic enzymes with industrial uses. The most promising candidate found after this screening was further studied by means of proteomic approaches.

2. Materials and methods 2.1. Reagents and substrates

PT

Methyl 4-hydroxy-3-methoxy-cinnamate (methyl ferulate, MFA) was purchased from Alfa Aesar (Haverhill, MA, USA). Ethyl 4-hydroxy-3-methoxy-cinnamate (ethyl ferulate), ferulic xylan

from

birchwood,

4-nitrophenyl β-D-xylopyranoside

(pNPX),

3,5-dinitrosalicylic acid

RI

acid,

4-nitrophenyl β-D-cellobioside

(DNS), (pNPC),

SC

4-nitrophenyl β-D-glucopyranoside (pNPG), 4-nitrophenol, carboxymethylcellulose sodium salt (CMC) and D-(+)-gluconic acid δ-lactone were obtained from Sigma-Aldrich. (methyl p-coumarate,

MpCA),

methyl caffeate

(MCA),

NU

Methyl 4-hydroxycinnamate

methyl sinapate (MSA), caffeic acid and sinapic acid were acquired from Carbosynth

MA

(Compton, UK).

Lignocellulosic substrates were obtained from different sources: processed milling oat fiber powder, corn and wheat seeds were purchased from a local supplier; sugar beet pulp (SBP)

D

was kindly provided by Sociedad Cooperativa General Agropecuaria ACOR (Valladolid,

PT E

Spain) as pellets. Substrates were ground until becoming a thick dust (diameter of particles of 1-5 mm).

2.2. Fungal strains and culture conditions

CE

Fungal strains were isolated from metal structures in four different locations: Continental environment (León, Spain), Dry Mediterranean environment (Madrid, Spain), Atlantic Ocean

AC

(Pontevedra, Spain) and Mediterranean Sea (Naples, Italy). Fermentations pipeline for the isolates went as follows: i) culture on solid Potato Dextrose Agar medium (PDA. Sigma-Aldrich) or Power medium [32], at 28 ºC, 5-7 days, until sporulation; ii) scrape and collect conidia from one Petri dish to inoculate a 500 mL non-baffled flask with 100 mL PMMY defined medium (culture volume ratio 1/5) [33], grow at 28 ºC, 250 rpm, 20 h (seed culture); iii) start fermentations inoculating 5% v/v seed culture in 500 mL non-baffled flasks with 100 mL of Lignocellulosic Medium (2.0 g/L NH4NO3; 1.0 g/L K2HPO4; 0.5 g/L MgSO4 7.H20; 0.5 g/L KCl; 1.0 g/L soy peptone; 1.0 g/L D-glucose; 18.7 mg/L FeSO4, pH 5.0)

5

ACCEPTED MANUSCRIPT supplemented with 20 g/L lignocellulosic substrate (sugar beet, wheat, milling oat or corn), grow at 28 ºC, 250 rpm, up to one week. Three biological replicates were used for each strain. 2.3. FAE activity screening FAE activity was analyzed based on the protocol described by Donaghy and McKay [34]. A 1% (w/v) Bacto-agar (BD) sterile solution (50 ºC) was supplemented with 1.5 mL of filtered ethyl ferulate (0.05 g/mL of ethanol) and poured into plates. Triplicate plugs (10 mm

PT

diameter) from PDA or Power medium plates containing sporulated fungal strains were inserted in the screening media and incubated at 28 ºC, 24 h. Plates were stained with

RI

bromocresol green [0.04% (w/v)], 15 min, and washed twice with NaCl 0.9% (w/v), 5 min. Yellow halos against the blue background corresponded to positive strains, due to the pH

SC

decrease after the release of ferulic acid. Agar plates without ethyl ferulate were employed as

NU

controls.

2.4. DNA isolation and identification of selected strains

MA

Genomic DNA extractions were carried out as described by Casqueiro and co-workers [32]. In brief, fungal strains were grown on PMMY medium for 48 h and the mycelium was harvested, recovered by filtration through a 30 µm pore diameter Nytal filter, dried and

D

lyophilized. 500 mg lyophilized mycelium were combined with 1 mL grinding solution and 1

PT E

mL of phenol:chloroform:isoamyl alcohol (25:24:1), mixed, incubated for 30 min at 50 ºC and centrifuged. Genomic DNA was precipitated with ethanol and resuspended in 100 µL TE buffer.

sequencing,

CE

Strains were identified by means of ribosomal DNA (rDNA) and internal transcribed spacer using:

i)

fungus-specific

universal

primers

ITS1

AC

(5′-TCCGTAGGTGAACCTGCGG-3′) and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′), for the non-coding regions between the 18S, 5.8S and 28S rRNA genes; as well as the 5.8S rRNA gene; ii) primers NL-1 (5′-GCATATCAATAAGCGGAGGAAAAG-3′) and NL-4 (5′GGTCCGTGTTTCAAGACGG-3′), for the D1-D2 region of the large subunit rRNA gene [35,36]. For PCR amplification, 7 ng/µL DNA in 25 µL of reaction volume were used, with 0.025 U/µL GoTaq® Flexi DNA Polymerase (Promega). 30 PCR cycles of 94 °C 40 sec, 52 °C 40 sec and 72 °C 1 min, with 10 min initial and final steps, were used. DNA was purified using the Qiagen II Gel Extraction Kit (Qiagen). Species were identified by sequence alignment using the GeneBank, EMBL, DDBJ, PDB and RefSeq databases and the BLASTN tool (http://www.ncbi.nlm.nih.gov/BLAST/). 6

ACCEPTED MANUSCRIPT 2.5. Protein extraction from the culture medium All steps were performed on ice or in a cold room at 4 ºC, to minimize proteolysis. Fermentation broths were filtered through a Nytal 30 µm pore membrane. Dry weight was assessed by drying the recovered mycelium at 80 ºC for one week. Flow-through was centrifuged at 3,220 x g, 4 ºC, 30 min and filtered through a 0.22 μm pore Stericup™ device (Millipore®). The filtrate was precipitated by the addition of ammonium sulfate at a 100%

PT

percentage saturation, according to the EnCorBio ammonium sulfate calculator [37], at 4 ºC, 18 h, with gentle stirring. Precipitated proteins were recovered by centrifugation at

RI

10,400 x g, 4 ºC, 1 h. The resulting pellet was resuspended in acetate buffer 0.02 N pH 5.0 to achieve a 100-fold concentration (v/v), obtaining the samples referred to as “enzyme

SC

extracts”. Enzyme extracts were the samples used in all enzymatic activities analyses, enzyme kinetics and proteomic techniques. Extracts were dialyzed against acetate buffer 0.02

2.6. Quantitative FAE activity assays

NU

pH 5.0, using Amicon ® Ultra-4 Centrifugal Filter Units MWCO 10 kDa.

MA

FAE activity was determined by HPLC-UV using MFA as a standard substrate. Reactions containing 10 μL of the enzyme extracts and 190 μL of a 0.05% MFA solution in acetate

D

buffer 0.1 N pH 5.0 were incubated 30 min at 37 ºC and terminated by boiling, 5 min. Samples were centrifuged 5 min at 16,168 x g, filtered through 0.22 µm syringe filters

PT E

(Fisherbrand) and analyzed by HPLC with a Mediterranea Sea 3 µm reversed-phase column (4.6 x 150 mm, Teknokroma), thermostated to 20 ºC. Substrate and product peaks were evaluated according to an isocratic elution method at a flow velocity of 0.8 mL/min,

CE

following two methods: i) 10 µL of sample were injected and analyzed in 55% mobile phase A (0.01% TFA) and 45% mobile phase B (100% methanol), employing Shimadzu equipment;

AC

ii) 20 µL of sample were injected and analyzed in 60% mobile phase A and 40% mobile phase B, employing Agilent equipment. Both methods included a 5 min re-equilibration step between runs. Peaks were identified by retention times and spectral information (Photodiode Array Detector) and quantified (UV-Visible detector operated at 324 nm) interpolating from linear standard curves. One unit of enzyme activity was defined as the amount of enzyme required to release one µmole of product per min under the tested conditions. All measurements were run, at least, in triplicates. 2.6.1. FAE activity characterization of the enzyme extracts Determination of optimal natural lignocellulosic substrate and sampling time 7

ACCEPTED MANUSCRIPT Fermentations were carried out using four different lignocellulosic substrates in the Lignocellulosic Medium: processed milling oat fiber powder, SBP, processed corn powder and processed wheat powder. Broth samples were collected every 24 h, up to one week, and used for FAE activity determination, expressed in units (U) per µL. pH and temperature optima Reactions were performed as described in section 2.6, with modifications. Acetate buffer

PT

0.1 N pH 5.0 was substituted with citrate 0.1 M-phosphate buffer at pH ranging from 3.0 to

acetate buffer 0.1 N pH 5.0, 30 min, from 37 to 70 ºC.

SC

Effect of temperature on FAE activity and stability

RI

7.0, 30 min at 37 ºC. To determine the temperature optimum, reactions were performed on

Twenty µL of enzyme extracts were incubated for 1 h at temperatures from 30 to 90 ºC, and

NU

then cooled to room temperature and used on FAE reactions (final volume 200 µL MFA in acetate buffer 0.1 N pH 5.0) as described in section 2.6, at 50 ºC, 30 min. These FAE

MA

reactions with the enzyme extract after the thermal treatment were terminated by boiling for 5 min.

Substrate specificity and kinetics parameters

D

Substrate specificity was analyzed performing FAE activity reactions on MFA, MCA, MpCA

PT E

and MSA, measuring the released amount of the corresponding hydroxycinnamic acid (respectively: ferulic, caffeic, p-coumaric and sinapic acid). Km and Vmax were determined measuring initial reaction rates for MFA, MCA and MpCA in

CE

acetate buffer 0.1 N pH 5.0, 50 ºC, at concentrations ranging from 0 to 5 mM. Values were plotted by means of the scientific data analysis and graphing software SigmaPlot® 12, and

Excel [38].

AC

verified by a nonlinear least-squares data fitting method, using the Solver add-in in Microsoft

Enzymatic hydrolysis of natural lignocellulosic substrates Reactions were performed at 50 ºC, 175 rpm, 24 h, using 2% (w/v) SBP in acetate buffer 0.1 N pH 5.0 as substrate. Activity was measured by HPLC-UV and expressed as g of ferulic acid released per kg of lignocellulosic substrate, referred to added protein amount of enzyme extract. 2.7. Cellulase and xylanolytic activity assays

8

ACCEPTED MANUSCRIPT β-glucosidase, cellobiohydrolase and β-xylosidase activities were measured based on the release of 4-nitrophenol after hydrolysis of synthetic substrates [39,40] and presented as specific activities expressed in units per gram of total protein in the enzyme extract. Reactions were performed adding 10 µL enzyme extract to 540 µL substrate solution in acetate buffer 0.1 N pH 5.0, incubated at 50 ºC and terminated by the addition of 550 µL 2% Na2CO3. Absorbance was measured at 405 nm and values were compared to those in a linear standard curve for 4-nitrophenol, prepared in triplicates under the same conditions.

PT

Substrate solutions were: i) for β-glucosidase, 4 mM pNPG; ii) for cellobiohydrolase, 4 mM pNPC supplemented with 1 mg/mL D-glucono-1,5-δ-lactone; iii) for β-xylosidase, 5 mM

RI

pNPX.

SC

Endoglucanase and xylanase activity were determined through the release of reducing sugars, measured adding DNS [41,42]. Reactions were: i) for endoglucanase, 10 µL enzyme

NU

extract in a final volume of 500 µL 1% (w/v) carboxymethyl cellulose (CMC) in acetate buffer 0.1 N pH 5.0; ii) for xylanase, 10 µL enzyme extract in a final volume of 500 µL 1%

MA

(w/v) xylan from birchwood (Sigma-Aldrich) in acetate buffer 0.1 N pH 5.0 (xylan stock solution was prepared in acetate buffer 0.1 N pH 5, boiled 30 min and stirred at 25 ºC overnight; then, centrifuged 15 min and volume adjusted with distilled water). Reactions

D

were performed at 50 ºC, followed by the addition of 500 µL of DNS reagent containing

PT E

1.4% DNS, 0.28% phenol, 0.07% sodium sulfite, 28% Na-K-tartrate (Rochelle salt) and 1.4% NaOH. Reaction tubes were heated at 99 °C, 5 min for color development, and cooled down in an ice-water bath, 5 min for color stabilization. Absorbance of the samples was measured

CE

at 540 nm. The amount of released reducing sugars was determined interpolating from a linear standard curve for glucose/xylose prepared in triplicates under the tested conditions.

AC

Values were presented as specific activities expressed in units per gram of total protein in the enzyme extract.

2.8. Saccharification assay 1.25 mg enzyme extract/g substrate were added to conical 50 mL tubes containing 2% SBP powder in acetate buffer 0.1 N pH 5.0. Reactions were performed at 50 ºC, 175 rpm, 24 h. Saccharification was measured by HPLC-UV (Waters equipment), determining the amount of released mono and disaccharides (sucrose, glucose, fructose, arabinose and galactose) according to an isocratic method developed considering previous works [43–47], using a 8µm Transgenomic CARBOSEP COREGEL 87P column (300 x 7,8 mm) thermostated to 90 ºC.

9

ACCEPTED MANUSCRIPT Twenty µL of sample were injected in Milli-Q Water (mobile phase), at a flow rate of 0.8 mL/min. 2.9. Enzyme extracts treatment for 2-DE and protein identification Enzyme extracts were precipitated adding an equal volume of prechilled 20% (w/v) TCA/acetone containing 0.14% (w/v) DTT [48,49], at -20 ºC, overnight. Precipitated proteins after centrifugation (3,220 x g, 10 min, 4 °C) were washed twice with prechilled acetone plus

PT

0.07% (w/v) DTT and once with 80% (v/v) acetone/distilled water. The final pellet was resuspended in sample buffer [8 M urea, 2% (w/v) CHAPS, 0.5% (v/v) IPG buffer 3-11 NL

RI

(GE Healthcare), 100 mM DTT and 0.002% bromophenol blue] at 25 ºC, shaking for 1 h, and subsequently stored at -80 °C. Protein concentration was determined according to the

SC

Bradford method [50].

NU

2.10. Two-dimensional gel analysis

Two-dimensional gel electrophoresis was carried out as described by Jami and co-workers

MA

[33], using three gels obtained from three independent cultures (biological replicates). In brief, 400 µg of soluble proteins in sample buffer were isoelectrofocused on an IPGphor apparatus, by means of 3-10 NL Immobiline™ 18-cm IPG strips (GE Healthcare). Proteins

D

were focused at 20 °C according to the following program: 1 h, 0 V and 12 h, 30 V

PT E

(rehydration); 2 h, 60 V; 1 h, 500 V; 1 h, 1,000 V; 30-min gradient to 8,000 V; up to 7 h, 8000 V until 50 kVh. Prior to the second dimension separation, IPG strips were equilibrated for 15 min at room temperature in equilibration buffer [2.0% (w/v) SDS, 50 mM Tris-HCl pH

CE

8.8, 6 M urea, 30% (v/v) glycerol, 0.002% bromophenol blue], supplemented with 65 mM DTT. A second equilibration step was performed in equilibration buffer supplemented with 4.0% (w/v) iodoacetamide instead of DTT, for 15 min at room temperature. The second

AC

dimension separation was carried out on SDS-PAGE in 12.5% (w/v) polyacrylamide gels in an Ettan Dalt Six apparatus (GE Healthcare). Precision Plus Protein Standards (Bio-Rad) were used as molecular weight markers. Gels were stained following the “blue silver” Colloidal Coomassie staining method [33]. 2.11. Analysis of protein expression Two-dimensional gels were scanned using an ImageScanner III (GE Heathcare), previously calibrated by using a grayscale marker (Eastman Kodak Co.) and the Labscan 5.00 (v1.0.8) software (GE Healthcare). ImageMasterTM 2D Platinum v5.0 software (GE Healthcare) was employed for gel analysis. After automatic spot detection and manual supervision to 10

ACCEPTED MANUSCRIPT eliminate any possible artifacts, such as streaks and background noise, spot normalization was performed, using relative volumes (volume of each spot divided by the total volume of all the spots in the gel) to quantify and compare the gel spots. Three gels obtained from three independent cultures (biological replicates) were analyzed to guarantee representative results. Protein spots were overlapped and matched, employing landmark features. Only those spots present in all three biological replicates were considered for identification.

PT

2.12. Protein identification Protein spots of interest were manually excised from stained gels by biopsy punches, digested

RI

with trypsin following the method of Havlis and co-workers [51] and processed for further analysis as indicated by Jami and co-workers [33]. Samples were analyzed with a 4800

SC

Proteomics Analyzer (MALDI-TOF/TOF, AB Sciex). A 4700 Proteomics analyzer calibration mixture (Cal Mix 5, AB Sciex) was used as external calibration. All MS spectra

NU

were internally calibrated using peptides from the trypsin auto-digestion. The analysis by MALDI-TOF/TOF mass spectrometry produced peptide mass fingerprints (PMF), and the

MA

peptides observed (up to 65 peptides per spot) were collected and represented as a list of monoisotopic molecular weights with a signal to noise (S/N) ratio greater than 20, using the 4000 Series Explorer v3.5.3 software (AB Sciex). All known contaminant ions (trypsin- and

D

keratin-derived peptides) were excluded from later MS/MS analyses. From each MS spectra,

PT E

the 6 most intensive precursors with an S/N greater than 20 were selected for MS/MS analyses with CID (atmospheric gas was used) in 2-kV ion reflector mode and precursor mass windows of ±7 Da. For protein identification, Mascot Generic Files combining MS (PMF)

CE

and tandem MS (MS/MS) spectra were automatically created and used to query a nonredundant protein database using a local license of Mascot v 2.2 from Matrix Science through

AC

the Global Protein Server v 3.6 (Applied Biosystems). Search parameters for peptide mass fingerprints and tandem MS spectra obtained were as follows: i) Uniprot Ascomycota (date 2017.07.03; 6,766,808 sequences, 3,023,177,811 residues); ii) fixed and variable modifications were considered (Cys as S-carbamidomethyl derivative and Met as oxidized methionine); iii) one missed cleavage site was allowed; iv) precursor tolerance was 100 ppm and MS/MS fragment tolerance was 0.3 Da; v) peptide charge: 1+; vi) the algorithm was set to use trypsin as the enzyme. Protein candidates produced by combined peptide mass fingerprinting/tandem MS search were only considered valid when two criteria were met: i) global Mascot score > 83

11

ACCEPTED MANUSCRIPT (significance level of p < 0.05) and ii) sequence coverage > 15%. If the second criterion was not met, they were only accepted when two or more fragmentation peptides were identified.

3. Results 3.1. FAE activity screening and isolates identification Due to the medical and cosmetic applications of ferulic acid, as well as its uses as a food and

PT

feed additive, the screening of new or more efficient feruloyl esterases (FAEs) has acquired industrial significance. Considering the number of putative FAEs and other lignocellulolytic

RI

enzymes present throughout the fungal genomes readily available [52], and the possibility of finding enzymatic activities in novel and unpredicted environments, a screening for FAE

SC

producing microorganisms originating from metal surfaces in different climates was carried out. Twenty fungal isolates able to grow on lignocellulolytic substrates were screened by

NU

means of a FAE activity plate assay (Supplementary File S1). Seven fungal strains degraded ethyl ferulate. FAE activity results were expressed by the enzymatic index (EI) [53]

colony]. Values are shown on Table 1.

MA

[equation: (diameter of the colony + diameter of the degradation halo) / diameter of the

Positive candidates were identified by 18S rDNA and internal transcribed spacer sequencing.

D

All candidates were classified as fungi belonging to phylum Ascomycota, corresponding to

PT E

genera a) Fusarium: isolates PDA(2)2 and PDA(2)4 (F. oxysporum); b) Alternaria: isolates PDA1 and PDA8 (A. alternata); c) Penicillium: isolate PDA(2)10 (P. roseopurpureum), isolate JCOS10C (P. verrucosum) and d) Pleospora: isolate PDA(3)1 (P. herbarum).

CE

A. alternata PDA1 (Fig. 1), isolated from dry Mediterranean climate (Madrid, Spain), showed the highest FAE activity values on plate screening (Table 1). Moreover, it was easily

AC

cultivable, relatively fast growing and capable of sporulating in conditions easily achievable in a laboratory environment. These characteristics made A. alternata PDA1 the most promising candidate and, thus, it was selected for further studies. 3.2. Quantitative FAE activity assays In order to confirm the FAE activity of A. alternata PDA1 and select fermentation conditions, four natural lignocellulosic substrates were assayed: sugar beet pulp powder (medium M-SBP), processed milling oat fiber powder (medium M-OAT), processed wheat powder (medium M-WHEAT) and processed corn powder (medium M-CORN). Fermentation samples were taken every 24 h and FAE activity was determined through the release of 12

ACCEPTED MANUSCRIPT ferulic acid from hydrolysis of MFA (Fig. 2.A). M-OAT and M-SBP were the best substrate candidates. Although FAE activity was higher when A. alternata PDA1 was grown on milling oat, this substrate was discarded due to its suitability to be readily exploited in the food industry, for human consumption. Since the main interest of the circular economy and “green” bio-based industries is to take advantage of waste materials and low-value byproducts, SBP was selected for subsequent assays.

PT

3.2.1 FAE activity characterization It is common that commercial biomass degrading enzyme cocktails contain a mixture of

RI

hydrolytic enzymes, rather than only one purified enzyme. The presence of a variety of enzymes facilitates the processes that lead to the deconstruction of plant cell walls. Because

SC

of this and the fact that this work aims to elucidate the secreted enzymatic machinery of the studied strain, all assays were performed using enzyme extracts obtained from total

NU

fermentation broth. Enzyme extracts were recovered by filtering and precipitating fermentation broth with ammonium sulfate. This precipitation method was chosen based on

MA

several qualities of this inorganic salt: i) relatively inexpensive and affordable; ii) stabilizes protein structure, which can aid in the preservation of the enzymatic activity after precipitation; iii) effective in the process of protein precipitation and iv) much higher

D

solubility than any of the phosphate salts [54]. In order to determine the optimal sampling

PT E

time, FAE activity of the enzyme extracts compared to dry weight was assessed. According to Fig. 2.B, FAE activity of the enzyme extracts increased with fermentation time, reaching a maximum at 96 h. A. alternata PDA1 hyphal growth reached the end of the linear phase

CE

around 48 h and dry weight decrease happened from this time point, which could be attributable to biomass loss due to hyphal lysis and fragmentation [49]. This information

AC

supports 96 h as the optimal sampling time, as it was the time point with higher FAE activity and at longer fermentation times cell lysis was evident, both by the macroscopic culture morphology (data not shown) and by dry weight values. Cell lysis processes should be avoided. Firstly, because the present work focuses on the characterization of the extracellular enzymes produced by A. alternata PDA1, not those that are intracellular and thus, in principle, more difficult to recover and use from an industrial point of view. Secondly, because the presence of proteases can affect the stability of the enzyme extracts and cell lysis events result in the release of intracellular proteases, which add to the secreted proteindegrading enzymes already in the extract. In this case, considering the low representation of

13

ACCEPTED MANUSCRIPT intracellular proteins, cell lysis seemed not to be an important event at the optimal sampling time [49]. pH and temperature optima In order to characterize the enzyme extracts recovered from A. alternata PDA1 grown on SBP, FAE activity was determined at 37 ºC, at pH ranging from 3.0 to 7.0 (Fig. 3.A). The pH optimum was pH 5.0. The enzyme extracts showed activity at a considerably restricted pH

PT

range (pH 4.0 to 6.0). One remarkable finding was that increasing the pH over 5.0 resulted in multiple chromatographic peaks different to those of ferulic acid. These additional peaks can

RI

be considered a result of the degradation of MFA or ferulic acid to other by-products (see Supplementary File S2.A). Thus, pH values over 5.0 should be avoided. The behavior of the

SC

enzyme extract at pH over 5.0 was verified on a natural lignocellulosic material, performing reactions on 20 mg/mL SBP powder (24 h, 50 ºC, 125 rpm). As shown in Supplementary

NU

File S2.II, the appearance of unidentified peaks supported 5.0 as the optimal pH value. Subsequently, the temperature optimum was evaluated at temperatures from 37 to 70 ºC

MA

(Fig. 3.B). Enzyme extracts were active at a broad temperature range (FAE activity was detected at all tested temperatures) and the optimal temperature was 55-60 ºC.

D

Effect of temperature on FAE activity and stability

PT E

Enzymes intended for industrial uses are required to be stable against varying temperature, pH and the presence of certain chemicals [55]. According to this, thermal stability from 30 to 90 ºC was analyzed. Enzyme extracts were submitted to a thermal treatment for an hour,

CE

before using them in FAE activity reactions at optimal conditions (see above). As shown in Fig. 3.C, temperatures above 40 ºC affected the stability of the enzyme extract. Thermal

AC

treatments higher than 50 ºC decreased the FAE activity to values less than half of the maximum. Taking into account these data, 50 ºC was selected as the standard temperature for other lignocellulolytic assays. A decrease in thermal stability as temperature rises, even at temperature values closer to the temperature optimum, has been previously reported in other FAE characterization studies [56–59]. Substrate specificity and kinetic parameters Specific activities were measured with four synthetic substrates: MFA (used as the standard substrate throughout the other quantitative assays), MpCA, MCA and MSA. The behavior of FAEs against these four substrates has traditionally been employed to classify them. The enzyme extract recovered from A. alternata PDA1 fermentations showed higher specific 14

ACCEPTED MANUSCRIPT activity towards MpCA, followed by MCA and MFA (Table 2). The specific activity towards MSA was considerably lower. This pattern corresponds to those feruloyl esterases classified as type B, according to the classical functional classification [60]. Type B feruloyl esterases act preferentially on the phenolic moiety of those substrates containing one (p-coumaric) or two (caffeic acid) hydroxyl substitutions. However, hydrolytic rates are affected when acting on substrates with a methoxy group, such as MFA, and they are not active against MSA

PT

[24,60]. The classification as an extract mainly containing type B feruloyl esterases complies with the fact that this type is preferentially secreted when culturing microorganisms on SBP

RI

[60].

The Km and Vmax values for MFA, MpCA and MCA were determined in acetate buffer 0.1 N

SC

pH 5.0, at 50 ºC. Solutions containing MFA, MpCA and MCA at concentrations ranging from 0 to 5 mM were used as substrates and released free hydroxycinnamic acids (ferulic,

NU

coumaric and caffeic acid, respectively) were determined. Values showed normal Michaelis-Menten kinetics for all substrates (Fig. 4). As shown on Table 2, the Km values

MA

indicated that the enzyme extract possessed a higher affinity for MCA, followed by MFA and MpCA.

D

Enzymatic hydrolysis of natural lignocellulosic substrates FAE activity of the enzyme extracts was characterized by the above-mentioned assays, when

PT E

acting on synthetic substrates (ethyl ferulate, MFA, MpCA, MCA, MSA). However, the aim of a lignocelullolytic enzyme extract with commercial applications is to collaborate on the hydrolysis of natural lignocellulosic substrates and derivatives [61,62]. Accordingly,

CE

reactions were carried out on SBP, adding increasing amounts of enzyme extract to determine the optimal ratio. Even at low concentrations, the enzyme extract was able to release ferulic

AC

acid from SBP, up to approximately 3 g ferulic acid per kg of SBP (≈0.3%). Within the range of 0 mg to 1.25 g of enzyme extract per kg of substrate, the relative values of ferulic acid released per amount of substrate increased considerably. However, the release of ferulic acid was not improved at higher amounts of enzyme extracts. These values support 1.25 g of total enzyme extract protein per kg of SBP as the optimal amount (Supplementary File S3). 3.3. Proteomic analyses of the enzyme extract Proteomic approaches were used to identify and enlarge the information about the range of extracellular proteins present in the A. alternata PDA1 enzyme extract, when grown on SBP. Supernatant samples were collected at the optimal sampling time (96 h cultivation) and 15

ACCEPTED MANUSCRIPT precipitated using ammonium sulfate, in order to recover enzyme extracts. A subsequent precipitation was carried out with TCA/acetone to remove the high content in salts and to concentrate samples. The secreted protein extract 2-DE reference map (Fig. 5) was obtained, with 112 protein spots, which were present in the three biological replicates (less than 18% variability in number of spots/gel). Spot normalization was performed and relative volumes of each protein spot were determined, depending on their intensity. Out of the 112 protein

PT

spots, 109 showed percentages of the total spot volume below 2% (Fig. 6.A). 97 of the 112 protein spots were successfully identified by peptide mass fingerprinting and tandem MS, obtaining 125 identifications (Table 3). The identification of several proteins from one spot in

RI

the reference map is a finding previously reported in other studies [49]. From the 125

SC

identified proteins, 98 corresponded to protein species (isoforms) of 17 different proteins. The remaining 27 protein spots were identified as unique protein species (Supplementary

NU

File S4.A).

The presence of putative signal peptides was analyzed by means of the SignalP software [63–

MA

65]. SignalP performs a predictive analysis of the presence and location of signal peptide cleavage sites in amino acid sequences. The method incorporates a prediction of cleavage sites and a signal peptide/non-signal peptide prediction. For this analysis, sequences derived

D

from protein identification after peptide mass fingerprinting and tandem MS were used. Out

PT E

of 44 distinct proteins, 38 were predicted to have classical signal peptide sequences. The remaining 6 proteins were analyzed through the SecretomeP software [66], which predicts the presence of non-classical secretion signals. Three of them appeared to have a non-classical

CE

secretory targeting signal. Non-classical secretory pathways are important in some organisms belonging to kingdom Fungi, such as some fungal plant pathogens [67] and the yeast

AC

Saccharomyces cerevisiae, where, according to Stein and co-workers, a majority of the proteins in the secretome did not contain a N-terminal signal sequence [68]. A functional annotation of the identified proteins in the enzyme extract was performed by using the COG (Clusters of Orthologous Groups of proteins) system, which also includes eukaryote organisms in its updated version (KOG) [69]. Of all identified proteins, 50% were assigned to COG categories, due to the fact that the COG system is slightly incomplete for eukaryotic organisms when compared to prokaryotes (711 prokaryotic genomes versus 7 eukaryotic genomes). Those proteins not included in COGs were submitted to analysis of conserved domains through the NCBI's Conserved Domain Database (CDD) and the InterPro

16

ACCEPTED MANUSCRIPT database [70,71]. Overall, out of the 44 different proteins identified, 38 were successfully classified into nine of the COG functional categories (Fig. 6.B). Carbohydrate metabolism and transport proteins When grown on natural complex vegetal substrates, fungal species produce and secrete enzymes such as cellulases, hemicellulases (xylanolytic enzymes), amylases and FAEs, among others [72–74]. The majority of the different identified proteins were classified as

PT

enzymes connected with carbohydrate metabolism (Supplementary File S4.B). The protein spot with the highest relative volume in the reference map (12.73%), spot 50, was identified

RI

as an endopolygalacturonase (EPG). EPGs are the best-known enzymes among the pectinases, which are CAZymes (carbohydrate-active enzymes) capable of aiding in the

SC

processes of degradation of pectins (heteropolysaccharides from the plant cell wall). Further putative pectinases were found: i) hydrolases, such as other endopolygalacturonases

NU

(EC 3.2.1.15), exopolygalacturonases (EC 3.2.1.67), α-L-arabinofuranosidases (EC 3.2.1.55; identified from several spots, including spots 62 and 74, which were the second and third

MA

with higher relative volume in the reference map, respectively), endoarabinases (EC 3.2.1.99) and arabinogalactanases (EC 3.2.1.89); ii) lyases, such as pectate lyases (EC 4.2.2.2) and rhamnogalacturonan endolyases (EC 4.2.2.23). These findings denote that the presence of

D

putative pectinases in the secretome was remarkable (Supplementary File S4.B). Commercial

PT E

acidic pectinases are mainly extracted from fungal sources and they have applications in the fruit juice and wine making industries, maceration of plant tissue, or saccharification of biomass [75]. These enzymes have also been reported as virulence factors in fungal

CE

phytopathogens, since they aid in the penetration of the plant tissues [76]. In addition to pectinases, other CAZymes were identified. One of the identified proteins

AC

showed homology with a sucrose-6-phosphate hydrolase (EC 3.1.3.24). Sucrose is the most important molecule to distribute photosynthetically-derived carbon from source to sink tissues in higher plants [77]. Sucrose-6-phosphate hydrolases have been identified in some fungal species, where the glycoside hydrolase activity has importance in their invasive growth and symbiotic plant-fungus associations [78,79]. β-glucosidases (EC 3.2.1.21), identified from spots 1, 2 and 3, are one of the three main enzymatic groups required to degrade

cellulose,

along

with

endoglucanases

(EG,

EC 3.2.1.4)

and

exoglucanases/cellobiohydrolases (EC 3.2.1.91). They have an important industrial role in the last steps of the enzymatic conversion of cellulose to glucose monomers and currently constitute one of the major bottlenecks in the biofuels and platform chemicals industries [80]. 17

ACCEPTED MANUSCRIPT A glycosyl hydrolase family 43 protein was identified from spot 37, involved in the pectins and hemicelluloses degradation processes. Another protein connected to hemicellulase activity is an endo-1,4-β-xylanase (EC 3.2.1.8, spot 95), which possesses industrial applications in fields such as animal feed, food, beverages, pharmaceutical, chemical, textiles, cellulose pulp and paper industries [81], and several identified β-D-xylosidases (EC 3.2.1.37). Spots 8 and 9 were identified as a glycoside hydrolase family 31 protein. The major

PT

CAZy activity in this group are α-glucosidases (EC 3.2.1.20), which are crucial hydrolases in the amylolytic metabolic pathway represented in most organisms, especially in those dependent on plant starch as an energy source [82]. Cutinases (EC 3.1.1.74; spot 94) are

RI

serine esterases capable of hydrolyzing the ester bonds of cutin, an insoluble lipid polyester,

SC

which is the main constituent of the plant cuticle, a protective coating that prevents plant dehydration and protects it against the foliar infection of plant pathogens. As a consequence,

NU

the presence of cutinases is related to the virulence of phytopathogens [83]. Several cutinases from Alternaria spp. have been described in the literature [84,85]. Finally, four endoglucanases were identified (spots 24, 30, 87 and 92), which could be involved in

MA

Alternaria spp. plant infection processes, as described by Eshel and co-workers [86]. Post-translational modification, protein turnover, and chaperones

D

Six different proteases were classified within this category. Three different proteins with

PT E

peptidase S53 domains (spots 3-6, 8-10, 12, 13 and 85) were identified as serine proteases (EC 3.4.21) belonging to the subtilase family. Subtilases are an extensively represented group of broad-spectrum proteases with important presence in saprophytic fungi, where they

CE

execute a crucial role in nutrient acquisition [87]. Nevertheless, some authors suggest that subtilases are also related to host invasion. This hypothesis would be supported by the fact

AC

that plants have developed mechanisms to produce protease inhibitors as a response to pathogen infection [88]. Three other identified proteins in the A. alternata PDA1 enzyme extract were involved in proteolytic processes: two proteins identified as aspartic proteases (spots 46, 51, 52, 57 and 64) and a deuterolysin metalloprotease (spot 96). The first two belonged to the group of the aspartic-type endopeptidases (EC 3.4.23), which are also known as acidic proteases and have implications in nutrient supply. Aspartic peptidases are employed in industrial fields such as food, beverages, medical and pharmaceutical [89]. The last identified proteolytic enzyme was a metalloprotease. This type of enzyme has recently been isolated from Alternaria solani and characterized [90]. Lipid metabolism 18

ACCEPTED MANUSCRIPT Four distinct hydrolases capable of acting on the ester bonds of carboxylic acids and derived structures were identified from spots 1-4, 12, 18 and 77. Lipases play an ecological role in the colonization or the use of plants as a carbon source by phytopathogens and plant-associated fungi, allowing a starting degradation of the waxes and cuticle, which makes the plant cell wall more accessible [91]. Unassigned proteins

PT

Among the proteins that were not assigned to COGs, spot 97 was identified as the major allergen Alt a 1 subunit from A. alternata. Allergen Alt a 1 from Alternaria spp. is a protein

RI

with a unique structure of a dimeric β-barrel, which may define a new fungal protein family whose function is unknown [16]. Nevertheless, some studies have provided information that

SC

might support a hypothetical role of this allergen in spore germination and a relation to virulence and pathogenicity in fungal plant pathogens [92]. A study carried out by Skóra and

NU

co-workers indicated that environmental strains of Alternaria spp. produced higher levels of this allergen than a strain obtained from a culture collection, when growing them on a

MA

medium containing lignocellulosic components [93], which would explain its presence in the secretome of the natural isolate A. alternata PDA1.

D

3.4. Cellulase and xylanolytic activity assays

PT E

Considering the significant presence of lignocellulolytic enzymes in the analyzed A. alternata PDA1 secretome (63.16% of the identified proteins) and its putative action as a plant pathogen degrading plant tissues, cellulase and xylanolytic activities were analyzed.

CE

Reactions were performed at 50 ºC in acetate buffer 0.1 N pH 5.0, in order to characterize the range of synergistic hydrolytic activities that occur simultaneously at conditions were the

AC

enzyme extract displayed FAE activity. Cellulase activity assays The hydrolysis of cellulose to glucose monomers requires three main groups of extracellular enzymes: i) endoglucanases; ii) exoglucanases (cellodextrinases, EC 3.2.1.74, and cellobiohydrolases) and iii) β-glucosidases. Cellobiohydrolase (CBH) and β-glucosidase activities were spectrophotometrically determined by measuring the release of 4-nitrophenol after the bond cleavage in a chromogenic substrate: pNPC and pNPG, respectively. In CBH reactions, the chromogenic substrate was supplemented with D-glucono-1,5-δ-lactone (a potent inhibitor of β-glucosidases, not affecting exoglucohydrolases) to ensure that CBH activity was not 19

ACCEPTED MANUSCRIPT masked by β-glucosidase. A. alternata PDA1 enzyme extracts showed both cellobiohydrolase and β-glucosidase specific activities, although the latter was comparatively higher (Table 4). Endoglucanase activity was measured by means of the 3,5-dinitrosalicylic acid (DNS) assay (IUPAC commission on biotechnology recommended method) [94]. Enzyme extracts showed endoglucanase activity (Table 4), which is consistent with the identification of two endoglucanases in the 2-DE reference map (Supplementary File S4.B).

PT

Xylanolytic activity assays Xylan is the most abundant molecule among the hemicelluloses in secondary plant cell walls

RI

and it is degraded by the synergistic action of several enzymatic groups [endo-β-l,4-xylanase

SC

(endoxylanase), β-xylosidases, etc] [95].

β-xylosidase activity was measured by means of a spectrophotometric quantitative assay. The

NU

enzyme extracts displayed β-xylosidase activity (Table 4), whereas endoxylanase activity was determined by measuring the release of reducing sugars. Although the DNS assay for the determination of xylanase activity is still widely employed in laboratories around the world, it

MA

has to be taken into account that, according to some authors, it produces results with a 3-8 fold overestimation when compared to other assays, such as the Nelson-Somogyi assay [94].

D

According to these assays, the enzyme extract displayed both β-xylosidase and endoxylanase

PT E

activities (Table 4). 3.5. Saccharification assays

Biotechnological strategies designed to use cellulosic material for feed or fuel depend on the

CE

ability of the proposed enzymatic systems to efficiently degrade the components of the plant cell wall to simple sugars [96]. In order to obtain further information on the ability or

pretreated

AC

A. alternata PDA1 enzyme extracts to catalyze the release of simple sugars from noncomplex

vegetal

substrates,

saccharification

assays

were

performed.

Saccharification was measured determining the amount of released mono and disaccharides (sucrose, glucose, fructose, arabinose and galactose) by HPLC-UV. All detected monosaccharides were not present in the control reactions performed without adding enzyme extract, which confirms that the obtained values are not attributable to a spontaneous release after incubation. The addition of enzyme extract successfully released limited amounts of glucose, fructose and arabinose (Table 5). However, neither sucrose nor galactose were detected.

4. Discussion 20

ACCEPTED MANUSCRIPT Photosynthetic organisms such as plants directly or indirectly provide most of the energy that is available to living organisms. Moreover, their cellular structures enclose valuable compounds, such as ferulic acid, with multiple economic applications. A successful industrial exploitation of this energy and chemicals source requires the hydrolysis of the plant cell wall. However, plant cell walls are dynamic structures that vary to a great extent between plant species, tissues, cells, primary or secondary wall and even inside of the same layer [97]. This

PT

heterogeneity requires the combination of physicochemical and enzymatic treatments. The enzymatic degradation of plant biomass is currently a challenge still in development, with

RI

major bottlenecks such as the degradation of lignocellulose to glucose [74]. Fungal microorganisms and their enzymatic machinery are crucial in lignocellulosic

SC

industries. Some ascomycetes with potential industrial applications in biomass degradation are genera Aspergillus [98], Penicillium [99], Trichoderma and Neurospora [100]. In this

NU

regard, fungal plant pathogens are of special interest as their mechanism of infection usually implies the production and secretion of lignocellulolytic enzymes to penetrate the host

MA

tissues. This is the case of Alternaria, a genus mainly containing saprobic fungi, which also encompasses some phytopathogenic species. Being a ubiquitous genus, it is possible to encounter species in very different ecosystems, associated with substrates including plant and

D

animal products and tissues, soil, atmosphere and aquatic environments [101]. Alternaria spp.

PT E

has also been isolated from less rich environments, such as building materials and carwash effluents [102]. This ubiquity explains why it was found on the metal surfaces analyzed in the present work and, also, the fact that two of the seven isolates with FAE activity were

CE

identified as Alternaria spp. Their opportunistic phytopathogenic character is related to their potential as lignocellulolytic enzyme producers shown in the present paper. Previous works

AC

have investigated this potential on genus Alternaria. Xiao and co-workers studied a full enzyme complement originating from an A. alternata strain isolated from Canadian wood log, with promising results on the release of ferulic acid from wheat and tritricale brans [103]. Alternaria showed lignocellulolytic activity on leaf litter from a subtropical forest containing mostly biomass from Quercus variabilis [104] and on synthetic substrates, when grown on pretreated delignified sugarcane bagasse, soybean bran, pectin and xylan [105]. The results shown in the present paper enlarge the knowledge about the Alternaria spp. lignocellulolytic machinery, identifying new CAZymes from a natural isolate and confirming this genus as an interesting candidate for industrial applications.

21

ACCEPTED MANUSCRIPT Regarding the industrial applications of Alternaria, further studies should be carried out in order to ensure that A. alternata PDA1 is a safe host for upscaled protein production. Firstly, being a potential human pathogen, it should be demonstrated that the strain is not capable of producing diseases on healthy individuals [106]. Alternaria spp. can produce different toxins, namely tenuazonic acid (tetramic acid derivative); alternariol, alternariol monomethyl ether, altenuene (dibenzopyrone derivatives) and the altertoxins (perylene derivatives) [107].

PT

However, toxigenicity is a trace that is common to other fungi currently being used as hosts for enzyme and metabolite production, such as Fusarium spp., Aspergillus spp. or Penicillium spp, and it is unlikely that these toxins are present in the processed enzyme extract.

RI

Alternaria spp. can also produce allergens. Another approach to avoid the potential

SC

pathogenicity, toxigenicity and allergenicity is the use of molecular biology techniques to express particular enzymes in a well-characterized production strain with a history of safe

NU

use, or to obtain genetically engineered strains that are safer [106]. The natural isolate A. alternata PDA1 was classified as genus Alternaria section Alternata,

MA

according to its macroscopical and microscopical features [101] (see Fig. 1). However, taxonomic classifications based solely on morphological features are not advisable for this genus, as it exhibits a substantial morphological plasticity depending on culture conditions,

D

substrate, temperature, light and humidity, even within the same culture [108]. Thus, rDNA

PT E

sequencing approaches were applied in order to verify the identification. Nucleotide BLAST searches returned results of high percentages of identity (99-100%) with regions of several Alternaria species belonging to the Alternata section, according to the classification proposed

CE

by Woudenberg and co-workers [101]. The complexity in the identification inside genus Alternaria is a common trait to several previous studies, particularly between the small-

AC

spored Alternaria species (A. tenuissima, A. alternata, among others) [108–110]. In this regard, some authors suggest that several misidentifications of Alternaria species might have occurred [111]. The identification in the present work was tentatively determined as A. alternata, as it is the representative species in section Alternata. Sugar beet pulp (SBP) is a complex natural lignocellulosic material whose content in ferulic acid makes it a good substrate for the utilization of fungal FAEs. Its low lignin and high sugar content makes it an appropriate raw material for the biofuels industry [112] and it has been successfully degraded to a great extent with the use of enzymatic preparations [113]. Sakamoto and co-workers reported the extraction of ferulic acid from SBP, using an enzyme mixture from a P. chrysogenum strain grown on a medium including this natural substrate 22

ACCEPTED MANUSCRIPT [114]. At least two FAEs were induced when cultivating Aspergillus niger on SBP, hypothetically due to its high content of hemicellulosic materials, which may act as strong FAE inducers [115]. Apart from high levels of ferulic acid (which make it a suitable substrate for the production of “natural” vanillin), SBP also contains other exploitable polysaccharides such as galacturonic acid, arabinose and rhamnose, which can be extracted through the use of enzymatic cocktails [116]. The utilization of wheat and corn flour as a FAE inducer in the

PT

present work did not produce as promising results as the use of SBP, possibly on account of their differential polysaccharide composition (a higher starch and lower hemicelluloses content) [117,118]. Considering the average composition values for SBP [119,120] and the

RI

values from a study that used the same SBP as the present work (ferulic acid content of

SC

0.6-1.1%) [121], A. alternata PDA1 enzyme extract successfully released a 27-50% of the total ferulic acid, in 24 h. Enzyme mixtures such as the one analyzed in the present study

NU

have shown higher competence in the release of ferulic acid from natural substrates than purified FAEs, possibly because of the concurrence of other lignocellulolytic enzymes that

MA

act synergistically to open the plant cell wall structure to the action of FAEs [120]. Several FAEs have been purified and characterized in the last decades. FAEs show activity at pH values ranging from 3.0 to 10.0 and temperatures from 20 to 75 ºC, although the majority

D

are active at pH 4.0-7.0 and temperatures lower than 50 ºC [24]. The studied A. alternata

PT E

PDA1 enzyme extract showed a pH optimum of 5.0, which coincides with other fungal FAEs analyzed in previous works [24]. The additional peaks that appeared in the HPLC analyses of the degradation of substrates at pH greater than 5.0 might be a consequence of the activity of

CE

other enzymes present in the heterogeneous enzyme extract, or a phenomenon derived of an alkaline hydrolysis of the substrate as pH in the reaction medium rises, with the liberation of

AC

side products [122]. The temperature optimum was 55-60 ºC, which is slightly higher than the majority of characterized fungal FAEs and is a potentially exploitable feature from an industrial point of view. In general, FAEs possess a limited thermostability at high temperatures [24], which was the reason why the reaction temperature was conservatively established at the lowest temperature close to the optimal range, 50 ºC. In general, Vmax values for A. alternata PDA1 enzyme extract were considerably lower than those determined in previous studies for other FAEs [123]. Km values for MFA, MCA and MpCA for the enzyme extract were lower than those of a FAE-II from F. oxysporum, which indicates a higher affinity for those substrates [124]. Recombinant Fae-1 from Neurospora crassa, showed Km values of approximately half the Km value for MFA in the analyzed 23

ACCEPTED MANUSCRIPT enzyme extract, and 20-fold lower for MCA and MpCA, which denotes a very high affinity for MCA and MpCA [125]. Xiao and co-workers determined specific FAE activities for cultures of a natural isolate of A. alternata (strain FC007) on wheat and triticale brans, finding activity values 5-fold, 2.8-fold and 7.3-fold higher for MFA, MpCA and MCA, respectively [103]. In summary, type B FAEs from other fungal species showed lower Km values than A. alternata PDA1 [123], which implies that the affinity towards MFA, MpCA

PT

and MCA of the analyzed enzyme extract is not as high as previously studied enzyme preparations.

RI

The kinetic parameters determined for MFA, MCA, MpCA and MSA corresponded to those of an enzyme belonging to the type B FAE group, according to the classical classification

SC

[60]. This finding can be explained by the inducing substrate utilized in this study: SBP is known to be the preferential natural substrate for the production of type B FAEs [22]. Even

NU

though this classical ABCD classification was useful initially, Dilokpimol and co-workers suggested an updated classification that includes 13 subfamilies and takes into account

MA

phylogenetic analyses. Thus, different FAEs seem to have derived from tannases, acetyl xylan esterases, lipases, choline esterases, xylanases and α-l-rhamnosidases, which might explain the differences in the specificity towards distinct hydroxycinnamic acids [24].

D

The proteomic approaches carried out in this work shed light on the complexity of

PT E

A. alternata PDA1 secretome. Several proteins were successfully identified from the reference map and a vast majority of them were assigned to A. alternata strains, which is coincident with the natural isolate species identification. In general, the majority of them

CE

were extracellular proteins possessing signal peptides and no intracellular proteins suggesting an important concurrence of cell lysis processes were found. Most of the identified proteins

AC

belonged to the carbohydrate metabolism and transport functional group, particularly CAZymes. A considerable number of pectinases and hemicellulases were found, which can be due to the high content of pectins in SBP [118], and its carbohydrate composition in general. Pectinases have been proposed as enzymes involved in the pathogenicity of Alternaria spp. [86] Other enzymes presumably related to phytopathogenicity in Alternaria spp. were identified in the present work, such as endopolygaracturonases [76]. A cutinase was identified from the enzyme extracts displaying FAE activity. Cutinases have previously been reported as proteins that can show FAE activity [126]. The presence of CAZymes in the enzyme extract of A. alternata PDA1 was supported by the cellulase and xylanolytic enzymatic activities found in assays. β-glucosidase and 24

ACCEPTED MANUSCRIPT endoglucanase were of special interest, showing the highest activity values, followed by the xylanolytic activities. On the contrary, cellobiohydrolase activity values were considerably lower. Lee and co-workers evaluated several lignocellulolytic activities in three white rot fungi grown on softwood, finding that their cellobiohydrolase activity was low and it only appeared at long incubation times [127]. Some studies have shown promising increases in the cellobiohydrolase activity values when heterologously expressing these enzymes in fungi

PT

[128,129]. Further insights on the Alternaria spp. genomes and molecular engineering tools for this genus will open the possibility of heterologous expression of cellobiohydrolases,

RI

oriented at the production of more balanced enzyme cocktails.

The enzyme extract studied in the present work contains a complex mixture of hydrolytic

SC

enzymes, as confirmed by proteomic techniques. The enzymatic activity values obtained for endoglucanase, β-glucosidase, cellobiohydrolase, endoxylanase and β-xylosidase may be

NU

compared to those calculated for other microorganisms in previous works. Enzyme cocktails from Trichoderma reseei [130], several fungi isolated from a deep-sea sponge [131] and

MA

edible mushroom fungi [132] appeared to have higher activity values than the enzyme cocktail obtained from A. alternata PDA1. Nevertheless, the relative proportion of β-glucosidase and β-xylosidase activities was remarkably higher in A. alternata PDA1. Song

D

and co-workers [99] compared enzymatic activities of three commercial cellulase

PT E

preparations from Penicillium oxalicum (SP) and T. reesei (ST; Celluclast® 1.5L, SigmaAldrich). A. alternata PDA1 showed similar or higher β-glucosidase and β-xylosidase values than these commercial preparations, which confirms this strain as a promising candidate for

CE

the preparation of industrial cocktails with these enzymatic activities.

5. Conclusions

AC

The recalcitrance of the plant cell wall requires the characterization and utilization of an increasing number of novel lignocellulolytic enzymes with stability and activity conditions that make them suitable for industrial applications. In this regard, the present work provides new insights into the complex extracellular enzymatic machinery of A. alternata PDA1 as a lignocellulolytic fungus, particularly focusing on the degradation of polysaccharides. Enzymatic assays and proteomic approaches point Alternaria spp. out as a promising lignocellulolytic factory, being capable of producing a considerable variety of CAZymes, (including cellulases, xylanolytic enzymes, pectinases, sucrose-6-phosphate hydrolases and cutinases), proteases and lipases. Further knowledge on the characteristics of the range of 25

ACCEPTED MANUSCRIPT enzymes produced by this phytopathogen will open new perspectives and applications, both considering it a source for new industrial enzymes and a microorganism responsible for economical and material losses.

Acknowledgements This work was funded by the ERDF (European Regional Development Fund) and the Spanish Ministry of Economy, Industry and Competitiveness (project ref. no. CTM2012-32026).

PT

Ms. Laura García-Calvo was supported by a PhD grant awarded by the Spanish Ministry of Economy, Industry and Competitiveness (ref. no. BES-2013-064578). The funding sources

RI

had no involvement in the study design; collection, analysis and interpretation of data; in the

SC

writing of the report and in the decision to submit the article for publication. Special thanks go to the BIOCORIN project (“New biocoating for corrosion inhibition in

NU

metal surfaces”, Project ID: 282881, FP7-ENVIRONMENT), as source of the natural isolates tested in this study; ProWood project (“Wood and derivatives protection by novel bio-coating

MA

solutions” Project ID: PCIN-2016-081, ERA-IB 7th Joint Call) and Instituto para la Competitividad Empresarial ICE (former ADE. Project “Biorrefinería enfocada a la valorización de subproductos agroindustriales”). We would also like to thank for their

D

collaboration to all the members of INBIOTEC whose valuable help aided in the completion

AC

CE

PT E

of this work and the University of León (Spain).

26

ACCEPTED MANUSCRIPT References [1]

M. Blackwell, The fungi: 1, 2, 3 ... 5.1 million species?, Am. J. Bot. 98 (2011) 426– 438.

[2]

D.J. McLaughlin, D.S. Hibbett, F. Lutzoni, J.W. Spatafora, R. Vilgalys, The search for the fungal tree of life, Trends Microbiol. 17 (2009) 488–497. N.J. Mitchell, E. Bowers, C. Hurburgh, F. Wu, Potential economic losses to the US

PT

[3]

corn industry from aflatoxin contamination, Food Addit. Contam. Part A. 33 (2016)

[4]

RI

540–550.

T.E. Cleveland, P.F. Dowd, A.E. Desjardins, D. Bhatnagar, P.J. Cotty, United States

SC

Department of Agriculture - Agricultural Research Service research on pre-harvest prevention of mycotoxins and mycotoxigenic fungi in US crops, Pest Manag. Sci. 59

[5]

NU

(2003) 629–642.

J.M. Wagacha, J.W. Muthomi, Mycotoxin problem in Africa: Current status,

Microbiol. 124 (2008) 1–12.

D. Bhatnagar, J. Yu, K.C. Ehrlich, Toxins of filamentous fungi, Chem. Immunol. 81

D

[6]

PT E

(2002) 167–206. [7]

MA

implications to food safety and health and possible management strategies, Int. J. Food

M.C. Fisher, D.A. Henk, C.J. Briggs, J.S. Brownstein, L.C. Madoff, S.L. McCraw, S.J. Gurr, Emerging fungal threats to animal, plant and ecosystem health, Nature. 484

[8]

CE

(2013) 1–18.

C. Barreiro, J.F. Martín, C. García-Estrada, Proteomics shows new faces for the old

15. [9]

AC

penicillin producer Penicillium chrysogenum, J. Biomed. Biotechnol. 2012 (2012) 1–

L. Gifre, A. Arís, À. Bach, E. Garcia-Fruitós, Trends in recombinant protein use in animal production, Microb. Cell Fact. 16 (2017) 40.

[10] O.P. Ward, Production of recombinant proteins by filamentous fungi, Biotechnol. Adv. 30 (2012) 1119–1139. [11] D.B. Archer, Filamentous fungi as microbial cell factories for food use, Curr. Opin. Biotechnol. 11 (2000) 478–483. [12] Manisha, S.K. Yadav, Technological advances and applications of hydrolytic enzymes 27

ACCEPTED MANUSCRIPT for valorization of lignocellulosic biomass, Bioresour. Technol. (2017). [13] R. González-Fernández, E. Prats, J. V. Jorrín-Novo, Proteomics of Plant Pathogenic Fungi, J. Biomed. Biotechnol. 2010 (2010) 1–36. [14] J.M.P.F. de Oliveira, L.H. de Graaff, Proteomics of industrial fungi: trends and insights for biotechnology, Appl. Microbiol. Biotechnol. 89 (2011) 225–237. [15] H.X. Dang, B. Pryor, T. Peever, C.B. Lawrence, The Alternaria genomes database: a

RI

and allergenic species, BMC Genomics. 16 (2015) 239.

PT

comprehensive resource for a fungal genus comprised of saprophytes, plant pathogens,

[16] I. Kustrzeba-Wójcicka, E. Siwak, G. Terlecki, A. Wolańczyk-Mędrala, W. Mędrala,

SC

Alternaria alternata and Its Allergens: a Comprehensive Review, Clin. Rev. Allergy Immunol. 47 (2014) 354–365.

NU

[17] C.P. Kubicek, E.M. Kubicek, Enzymatic deconstruction of plant biomass by fungal enzymes, Curr. Opin. Chem. Biol. 35 (2016) 51–57.

MA

[18] G. Jäger, J. Büchs, Biocatalytic conversion of lignocellulose to platform chemicals, Biotechnol. J. 7 (2012) 1122–1136.

D

[19] F. Cherubini, The biorefinery concept : Using biomass instead of oil for producing

PT E

energy and chemicals, Energy Convers. Manag. 51 (2010) 1412–1421. [20] M. Bilal, M. Asgher, H.M.N. Iqbal, H. Hu, X. Zhang, Biotransformation of lignocellulosic materials into value-added products — A review, Int. J. Biol.

CE

Macromol. 98 (2017) 447–458.

[21] N. Kumar, V. Pruthi, Potential applications of ferulic acid from natural sources,

AC

Biotechnol. Reports. 4 (2014) 86–93. [22] A.E. Fazary, Y.H. Ju, Feruloyl esterases as biotechnological tools: Current and future perspectives, Acta Biochim. Biophys. Sin. (Shanghai). 39 (2007) 811–828. [23] F. Talebnia, D. Karakashev, I. Angelidaki, Production of bioethanol from wheat straw: An overview on pretreatment, hydrolysis and fermentation, Bioresour. Technol. 101 (2010) 4744–4753. [24] A. Dilokpimol, M.R. Mäkelä, M.V. Aguilar-Pontes, I. Benoit-Gelber, K.S. Hildén, R.P. de Vries, Diversity of fungal feruloyl esterases: updated phylogenetic classification, properties, and industrial applications, Biotechnol. Biofuels. 9 (2016) 28

ACCEPTED MANUSCRIPT 231. [25] A.E. Fazary, Y.-H. Ju, The large-scale use of feruloyl esterases in industry, Biotechnol. Mol. Biol. Rev. 3 (2008) 95–110. http://www.academicjournals.org/journal/BMBR/article-abstract/68CC9AE40231. [26] N. Gopalan, L.V. Rodríguez-Duran, G. Saucedo-Castaneda, K.M. Nampoothiri, Review on technological and scientific aspects of feruloyl esterases: A versatile

PT

enzyme for biorefining of biomass, Bioresour. Technol. 193 (2015) 534–544. [27] I. Benoit, E.G.J. Danchin, R.J. Bleichrodt, R.P. De Vries, Biotechnological

RI

applications and potential of fungal feruloyl esterases based on prevalence,

SC

classification and biochemical diversity, Biotechnol. Lett. 30 (2008) 387–396. [28] D.B.R.K.G. Udatha, I. Kouskoumvekaki, L. lsson, G. Panagiotou, The interplay of

NU

descriptor-based computational analysis with pharmacophore modeling builds the basis for a novel classification scheme for feruloyl esterases, Biotechnol. Adv. 29

MA

(2011) 94–110.

[29] G.K. Villena, M. Gutiérrez-Correa, Morphological patterns of Aspergillus niger biofilms and pellets related to lignocellulolytic enzyme productivities, Lett. Appl.

D

Microbiol. 45 (2007) 231–237.

PT E

[30] R. Liu, J. Zhu, Z. Yu, D. Joshi, H. Zhang, W. Lin, M. Yang, Molecular analysis of long-term biofilm formation on PVC and cast iron surfaces in drinking water distribution system, J. Environ. Sci. 26 (2014) 865–874.

CE

[31] M. Gutiérrez-Correa, Y. Ludeña, G. Ramage, G.K. Villena, Recent Advances on Filamentous Fungal Biofilms for Industrial Uses, Appl. Biochem. Biotechnol. 167

AC

(2012) 1235–1253.

[32] J. Casqueiro, O. Bañuelos, S. Gutiérrez, M.J. Hijarrubia, J.F. Martín, Intrachromosomal recombination between direct repeats in Penicillium chrysogenum: Gene conversion and deletion events, Mol. Gen. Genet. 261 (1999) 994–1000. [33] M.-S. Jami, C. Barreiro, C. García-Estrada, J.-F. Martín, Proteome Analysis of the Penicillin Producer Penicillium chrysogenum, Mol. Cell. Proteomics. 9 (2010) 1182– 1198. [34] J.A. Donaghy, A.M. McKay, Novel screening assay for the detection of phenolic acid esterases, World J. Microbiol. Biotechnol. 10 (1994) 41–44. 29

ACCEPTED MANUSCRIPT [35] T.J. White, T.D. Bruns, S.B. Lee, J.W. Taylor, Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics, in: M.A. Innis, D.H. Gelfand, J.J. Sninsky, T.J. White (Eds.), PCR Protoc. A Guid. to Methods Appl., Academic Press, Inc., New York, 1990. [36] X. Wang, Y.-F. Fu, R.-Y. Wang, L. Li, Y.-H. Cao, Y.-Q. Chen, H.-Z. Zhao, Q.-Q. Zhang, J.-Q. Wu, X.-H. Weng, X.-J. Cheng, L.-P. Zhu, Identification of Clinically

PT

Relevant Fungi and Prototheca Species by rRNA Gene Sequencing and Multilocus PCR Coupled with Electrospray Ionization Mass Spectrometry, PLoS One. 9 (2014)

RI

e98110.

[37] P. Wingfield, Protein Precipitation Using Ammonium Sulfate, in: Curr. Protoc. Protein

SC

Sci., John Wiley & Sons, Inc., Hoboken, NJ, USA, 1998: p. A.3F.1-A.3F.8.

NU

[38] G. Kemmer, S. Keller, Nonlinear least-squares data fitting in Excel spreadsheets, Nat. Protoc. 5 (2010) 267–281.

MA

[39] C.R.F. Terrasan, J.M. Guisan, E.C. Carmona, Xylanase and β-xylosidase from Penicillium janczewskii: Purification, characterization and hydrolysis of substrates, Electron. J. Biotechnol. 23 (2016) 54–62.

D

[40] C. Li, F. Lin, Y. Li, W. Wei, H. Wang, L. Qin, Z. Zhou, B. Li, F. Wu, Z. Chen, A β-

PT E

glucosidase hyper-production Trichoderma reesei mutant reveals a potential role of cel3D in cellulase production, Microb. Cell Fact. 15 (2016) 151. [41] Z. Xiao, R. Storms, A. Tsang, Microplate-based carboxymethylcellulose assay for

CE

endoglucanase activity, Anal. Biochem. 342 (2005) 176–178. [42] C. Mattéotti, J. Bauwens, C. Brasseur, C. Tarayre, P. Thonart, J. Destain, F. Francis, E.

AC

Haubruge, E. De Pauw, D. Portetelle, M. Vandenbol, Identification and characterization of a new xylanase from Gram-positive bacteria isolated from termite gut (Reticulitermes santonensis), Protein Expr. Purif. 83 (2012) 117–127. [43] S. Gorinstein, R. Moshe, J. Deutsch, F.H. Wolfe, K. Tilis, A. Stiller, I. Flam, Y. Gat, Determination of basic components in white wines by HPLC, FT-IR spectroscopy, and electrophoretic techniques, J. Food Compos. Anal. 5 (1992) 236–245. [44] I.M.P.L.V.. Ferreira, A.M.. Gomes, M.. Ferreira, Determination of sugars, and some other compounds in infant formulae, follow-up milks and human milk by HPLCUV/RI, Carbohydr. Polym. 37 (1998) 225–229. 30

ACCEPTED MANUSCRIPT [45] J.L. Chávez-Servín, A.I. Castellote, M.C. López-Sabater, Analysis of mono- and disaccharides in milk-based formulae by high-performance liquid chromatography with refractive index detection, J. Chromatogr. A. 1043 (2004) 211–215. [46] L.R.V. De Sá, M.A.L. De Oliveira, M.C. Cammarota, A. Matos, V.S. Ferreira-Leitão, Simultaneous analysis of carbohydrates and volatile fatty acids by HPLC for monitoring fermentative biohydrogen production, Int. J. Hydrogen Energy. 36 (2011)

PT

15177–15186. [47] J. Ma, X. Hou, B. Zhang, Y. Wang, L. He, The analysis of carbohydrates in milk

RI

powder by a new “heart-cutting” two-dimensional liquid chromatography method, J.

SC

Pharm. Biomed. Anal. 91 (2014) 24–31.

[48] F.J. Fernández-Acero, I. Jorge, E. Calvo, I. Vallejo, M. Carbú, E. Camafeita, J.A.

NU

López, J.M. Cantoral, J. Jorrín, Two-dimensional electrophoresis protein profile of the phytopathogenic fungus Botrytis cinerea, Proteomics. 6 (2006) S88–S96.

MA

[49] M.-S. Jami, C. García-Estrada, C. Barreiro, A.-A. Cuadrado, Z. Salehi-Najafabadi, J.F. Martín, The Penicillium Chrysogenum Extracellular Proteome. Conversion from a Food-rotting Strain to a Versatile Cell Factory for White Biotechnology, Mol. Cell.

D

Proteomics. 9 (2010) 2729–2744.

PT E

[50] M.M. Bradford, A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72 (1976) 248–254.

CE

[51] J. Havliš, H. Thomas, M. Šebela, A. Shevchenko, Fast-Response Proteomics by Accelerated In-Gel Digestion of Proteins, Anal. Chem. 75 (2003) 1300–1306.

AC

[52] U. Weingart, Y. Lavi, D. Horn, Data mining of enzymes using specific peptides, BMC Bioinformatics. 10 (2009) 446. [53] C.M.P. Braga, P. da S. Delabona, D.J. da S. Lima, D.A.A. Paixão, J.G. da C. Pradella, C.S. Farinas, Addition of feruloyl esterase and xylanase produced on-site improves sugarcane bagasse hydrolysis, Bioresour. Technol. 170 (2014) 316–324. [54] R.R. Burgess, Chapter 20 Protein Precipitation Techniques, in: Methods Enzymol., 1st ed., Elsevier Inc., 2009: pp. 331–342. [55] C.J. Yeoman, Y. Han, D. Dodd, C.M. Schroeder, R.I. Mackie, I.K.O. Cann, Chapter 1 - Thermostable Enzymes as Biocatalysts in the Biofuel Industry, in: Adv. Appl. 31

ACCEPTED MANUSCRIPT Microbiol., 2010: pp. 1–55. [56] S. Kelle, A. Nieter, U. Krings, K. Zelena, D. Linke, R.G. Berger, Heterologous production of a feruloyl esterase from Pleurotus sapidus synthesizing feruloylsaccharide esters, Biotechnol. Appl. Biochem. 63 (2016) 852–862. [57] Y.-S. Lee, J.B. Heo, J.-H. Lee, Y.-L. Choi, A Cold-Adapted Carbohydrate Esterase from the Oil-Degrading Marine Bacterium Microbulbifer thermotolerans DAU221:

PT

Gene Cloning, Purification, and Characterization, J. Microbiol. Biotechnol. 24 (2014) 925–935.

RI

[58] A. Nieter, P. Haase-Aschoff, D. Linke, M. Nimtz, R.G. Berger, A halotolerant type A

SC

feruloyl esterase from Pleurotus eryngii, Fungal Biol. 118 (2014) 348–357. [59] M. Zhou, Z. Huang, B. Zhou, Y. Luo, G. Jia, G. Liu, H. Zhao, X. Chen, Construction

NU

and expression of two-copy engineered yeast of feruloyl esterase, Electron. J. Biotechnol. 18 (2015) 338–342.

MA

[60] V.F. Crepin, C.B. Faulds, I.F. Connerton, Functional classification of the microbial feruloyl esterases, Appl. Microbiol. Biotechnol. 63 (2004) 647–652. [61] C. Escamilla-Alvarado, J.A. Pérez-Pimienta, T. Ponce-Noyola, H.M. Poggi-Varaldo,

D

An overview of the enzyme potential in bioenergy-producing biorefineries, J. Chem.

PT E

Technol. Biotechnol. 92 (2017) 906–924. [62] S.H. Toushik, K.-T. Lee, J.-S. Lee, K.-S. Kim, Functional Applications of Lignocellulolytic Enzymes in the Fruit and Vegetable Processing Industries, J. Food

CE

Sci. 82 (2017) 585–593.

AC

[63] H. Nielsen, Predicting Secretory Proteins with SignalP, in: D. Kihara (Ed.), Springer New York, New York, NY, 2017: pp. 59–73. [64] J. Dyrløv Bendtsen, H. Nielsen, G. von Heijne, S. Brunak, Improved Prediction of Signal Peptides: SignalP 3.0, J. Mol. Biol. 340 (2004) 783–795. [65] T.N. Petersen, S. Brunak, G. von Heijne, H. Nielsen, SignalP 4.0: discriminating signal peptides from transmembrane regions, Nat. Methods. 8 (2011) 785–786. [66] J.D. Bendtsen, L.J. Jensen, N. Blom, G. von Heijne, S. Brunak, Feature-based prediction of non-classical and leaderless protein secretion, Protein Eng. Des. Sel. 17 (2004) 349–356.

32

ACCEPTED MANUSCRIPT [67] T. Liu, T. Song, X. Zhang, H. Yuan, L. Su, W. Li, J. Xu, S. Liu, L. Chen, T. Chen, M. Zhang, L. Gu, B. Zhang, D. Dou, Unconventionally secreted effectors of two filamentous pathogens target plant salicylate biosynthesis, Nat. Commun. 5 (2014) 4686. [68] K.R. Stein, B.J. Giardina, H. Chiang, The Non-classical Pathway is the Major Pathway to Secrete Proteins in Saccharomyces cerevisiae, Clin. Exp. Pharmacol. 4 (2014).

PT

[69] R.L. Tatusov, N.D. Fedorova, J.D. Jackson, A.R. Jacobs, B. Kiryutin, E. V Koonin, D.M. Krylov, R. Mazumder, S.L. Mekhedov, A.N. Nikolskaya, B.S. Rao, S. Smirnov,

RI

A. V Sverdlov, S. Vasudevan, Y.I. Wolf, J.J. Yin, D.A. Natale, The COG database: an

SC

updated version includes eukaryotes, BMC Bioinformatics. 4 (2003) 41. [70] R.D. Finn, T.K. Attwood, P.C. Babbitt, A. Bateman, P. Bork, A.J. Bridge, H.-Y.

NU

Chang, Z. Dosztányi, S. El-Gebali, M. Fraser, J. Gough, D. Haft, G.L. Holliday, H. Huang, X. Huang, I. Letunic, R. Lopez, S. Lu, A. Marchler-Bauer, H. Mi, J. Mistry, D.A. Natale, M. Necci, G. Nuka, C.A. Orengo, Y. Park, S. Pesseat, D. Piovesan, S.C.

MA

Potter, N.D. Rawlings, N. Redaschi, L. Richardson, C. Rivoire, A. Sangrador-Vegas, C. Sigrist, I. Sillitoe, B. Smithers, S. Squizzato, G. Sutton, N. Thanki, P.D. Thomas,

D

S.C.E. Tosatto, C.H. Wu, I. Xenarios, L.-S. Yeh, S.-Y. Young, A.L. Mitchell, InterPro in 2017—beyond protein family and domain annotations, Nucleic Acids Res. 45

PT E

(2017) D190–D199.

[71] A. Marchler-Bauer, Y. Bo, L. Han, J. He, C.J. Lanczycki, S. Lu, F. Chitsaz, M.K.

CE

Derbyshire, R.C. Geer, N.R. Gonzales, M. Gwadz, D.I. Hurwitz, F. Lu, G.H. Marchler, J.S. Song, N. Thanki, Z. Wang, R.A. Yamashita, D. Zhang, C. Zheng, L.Y. Geer, S.H. Bryant, CDD/SPARCLE: functional classification of proteins via subfamily domain

AC

architectures, Nucleic Acids Res. 45 (2017) D200–D203. [72] G. Liu, L. Zhang, X. Wei, G. Zou, Y. Qin, L. Ma, J. Li, H. Zheng, S. Wang, C. Wang, L. Xun, G.-P. Zhao, Z. Zhou, Y. Qu, Genomic and Secretomic Analyses Reveal Unique Features of the Lignocellulolytic Enzyme System of Penicillium decumbens, PLoS One. 8 (2013) e55185. [73] H. Ravalason, S. Grisel, D. Chevret, A. Favel, J.G. Berrin, J.C. Sigoillot, I. HerpoëlGimbert, Fusarium verticillioides secretome as a source of auxiliary enzymes to enhance saccharification of wheat straw, Bioresour. Technol. 114 (2012) 589–596. [74] W.R. de Souza, Microbial Degradation of Lignocellulosic Biomass, in: Sustain. 33

ACCEPTED MANUSCRIPT Degrad. Lignocellul. Biomass - Tech. Appl. Commer., InTech, 2013: pp. 135–152. [75] D.. Kashyap, P.. Vohra, S. Chopra, R. Tewari, Applications of pectinases in the commercial sector: a review, Bioresour. Technol. 77 (2001) 215–227. [76] A. Isshiki, K. Akimitsu, M. Yamamoto, H. Yamamoto, Endopolygalacturonase Is Essential for Citrus Black Rot Caused by Alternaria citri but Not Brown Spot Caused by Alternaria alternata, Mol. Plant-Microbe Interact. 14 (2001) 749–757.

PT

[77] H.-S. Kim, H.-J. Park, S. Heu, J. Jung, Molecular and Functional Characterization of a Unique Sucrose Hydrolase from Xanthomonas axonopodis pv. glycines, J. Bacteriol.

RI

186 (2004) 411–418.

SC

[78] X. Liao, W. Fang, L. Lin, H.-L. Lu, R.J.S. Leger, Metarhizium robertsii Produces an Extracellular Invertase (MrINV) That Plays a Pivotal Role in Rhizospheric Interactions

NU

and Root Colonization, PLoS One. 8 (2013) e78118.

[79] J. Parrent, T.Y. James, R. Vasaitis, A.F. Taylor, Friend or foe? Evolutionary history of

MA

glycoside hydrolase family 32 genes encoding for sucrolytic activity in fungi and its implications for plant-fungal symbioses, BMC Evol. Biol. 9 (2009) 148. [80] A. Sørensen, M. Lübeck, P. Lübeck, B. Ahring, Fungal Beta-Glucosidases: A

D

Bottleneck in Industrial Use of Lignocellulosic Materials, Biomolecules. 3 (2013)

PT E

612–631.

[81] M.L.T.M. Polizeli, A.C.S. Rizzatti, R. Monti, H.F. Terenzi, J.A. Jorge, D.S. Amorim, Xylanases from fungi: properties and industrial applications, Appl. Microbiol.

CE

Biotechnol. 67 (2005) 577–591.

AC

[82] M. Okuyama, W. Saburi, H. Mori, A. Kimura, α-Glucosidases and α-1,4-glucan lyases: structures, functions, and physiological actions, Cell. Mol. Life Sci. 73 (2016) 2727–2751.

[83] S. Chen, L. Su, J. Chen, J. Wu, Cutinase: Characteristics, preparation, and application, Biotechnol. Adv. 31 (2013) 1754–1767. [84] C. Fan, W. Köller, Diversity of cutinases from plant pathogenic fungi: differential and sequential expression of cutinolytic esterases by Alternaria brassicicola, FEMS Microbiol. Lett. 158 (1998) 33–38. [85] K. Koschorreck, D. Liu, C. Kazenwadel, R.D. Schmid, B. Hauer, Heterologous

34

ACCEPTED MANUSCRIPT expression, characterization and site-directed mutagenesis of cutinase CUTAB1 from Alternaria brassicicola, Appl. Microbiol. Biotechnol. 87 (2010) 991–997. [86] D. Eshel, A. Lichter, A. Dinoor, D. Prusky, Characterization of Alternaria alternata glucanase genes expressed during infection of resistant and susceptible persimmon fruits, Mol. Plant Pathol. 3 (2002) 347–358. [87] G. Hu, R.J. St. Leger, A phylogenomic approach to reconstructing the diversification

PT

of serine proteases in fungi, J. Evol. Biol. 17 (2004) 1204–1214.

[88] T.A. Valueva, V. V. Mosolov, Role of inhibitors of proteolytic enzymes in plant

RI

defense against phytopathogenic microorganisms, Biochem. 69 (2004) 1305–1309.

SC

[89] L.W. Theron, B. Divol, Microbial aspartic proteases: current and potential applications in industry, Appl. Microbiol. Biotechnol. 98 (2014) 8853–8868.

NU

[90] M. Chandrasekaran, R. Chandrasekar, S.-C. Chun, M. Sathiyabama, Isolation, characterization and molecular three-dimensional structural predictions of

MA

metalloprotease from a phytopathogenic fungus, Alternaria solani (Ell. and Mart.) Sor., J. Biosci. Bioeng. 122 (2016) 131–139. [91] J. Barriuso, A. Prieto, M. Martínez, Fungal genomes mining to discover novel sterol

PT E

D

esterases and lipases as catalysts, BMC Genomics. 14 (2013) 712. [92] M.F. Gabriel, I. Postigo, C.T. Tomaz, J. Martínez, Alternaria alternata allergens: Markers of exposure, phylogeny and risk of fungi-induced respiratory allergy,

CE

Environ. Int. 89–90 (2016) 71–80. [93] J. Skóra, A. Otlewska, B. Gutarowska, J. Leszczyńska, I. Majak, Ł. Stępień,

AC

Production of the Allergenic Protein Alt a 1 by Alternaria Isolates from Working Environments, Int. J. Environ. Res. Public Health. 12 (2015) 2164–2183. [94] A. V. Gusakov, E.G. Kondratyeva, A.P. Sinitsyn, Comparison of Two Methods for Assaying Reducing Sugars in the Determination of Carbohydrase Activities, Int. J. Anal. Chem. 2011 (2011) 1–4. [95] R. Chávez, P. Bull, J. Eyzaguirre, The xylanolytic enzyme system from the genus Penicillium, J. Biotechnol. 123 (2006) 413–433. [96] R. Jimenez-Flores, G. Fake, J. Carroll, E. Hood, J. Howard, A novel method for evaluating the release of fermentable sugars from cellulosic biomass, Enzyme Microb.

35

ACCEPTED MANUSCRIPT Technol. 47 (2010) 206–211. [97] H. Wei, Q. Xu, L.E. Taylor, J.O. Baker, M.P. Tucker, S.-Y. Ding, Natural paradigms of plant cell wall degradation, Curr. Opin. Biotechnol. 20 (2009) 330–338. [98] R.A. Batista-García, E. Balcázar-López, E. Miranda-Miranda, A. Sánchez-Reyes, L. Cuervo-Soto, D. Aceves-Zamudio, K. Atriztán-Hernández, C. Morales-Herrera, R. Rodríguez-Hernández, J. Folch-Mallol, Characterization of Lignocellulolytic

PT

Activities from a Moderate Halophile Strain of Aspergillus caesiellus Isolated from a Sugarcane Bagasse Fermentation, PLoS One. 9 (2014) e105893.

RI

[99] W. Song, X. Han, Y. Qian, G. Liu, G. Yao, Y. Zhong, Y. Qu, Proteomic analysis of

system, Biotechnol. Biofuels. 9 (2016) 68.

SC

the biomass hydrolytic potentials of Penicillium oxalicum lignocellulolytic enzyme

NU

[100] N.L. Glass, M. Schmoll, J.H.D. Cate, S. Coradetti, Plant Cell Wall Deconstruction by Ascomycete Fungi, Annu. Rev. Microbiol. 67 (2013) 477–498.

MA

[101] J.H.C. Woudenberg, J.Z. Groenewald, M. Binder, P.W. Crous, Alternaria redefined, Stud. Mycol. 75 (2013) 171–212.

[102] T. Sibanda, R. Selvarajan, M. Tekere, H. Nyoni, S. Meddows-Taylor, Potential

D

biotechnological capabilities of cultivable mycobiota from carwash effluents,

PT E

Microbiologyopen. 6 (2017) e00498. [103] Z. Xiao, H. Bergeron, P.C.K. Lau, Alternaria alternata as a new fungal enzyme system for the release of phenolic acids from wheat and triticale brans, Antonie van

CE

Leeuwenhoek, Int. J. Gen. Mol. Microbiol. 101 (2012) 837–844.

AC

[104] J. Hao, X. Tian, F. Song, X. He, Z. Zhang, P. Zhang, Involvement of Lignocellulolytic Enzymes in the Decomposition of Leaf Litter in a Subtropical Forest, J. Eukaryot. Microbiol. 53 (2006) 193–198. [105] D. Robl, P. da Delabona, C. Mergel, J. Rojas, P. dos Costa, I. Pimentel, V. Vicente, J.G. da Cruz Pradella, G. Padilla, The capability of endophytic fungi for production of hemicellulases and related enzymes, BMC Biotechnol. 13 (2013) 94. [106] V. Sewalt, D. Shanahan, L. Gregg, J. La Marta, R. Carrillo, The Generally Recognized as Safe (GRAS) Process for Industrial Microbial Enzymes, Ind. Biotechnol. 12 (2016) 295–302.

36

ACCEPTED MANUSCRIPT [107] A. Logrieco, A. Moretti, M. Solfrizzo, Alternaria toxins and plant diseases: an overview of origin, occurrence and risks, World Mycotoxin J. 2 (2009) 129–140. [108] B.M. Pryor, T.J. Michailides, Morphological, Pathogenic, and Molecular Characterization of Alternaria Isolates Associated with Alternaria Late Blight of Pistachio, Phytopathology. 92 (2002) 406–416. [109] N.P. Kwiatkowski, W.M. Babiker, W.G. Merz, K.C. Carroll, S.X. Zhang, Evaluation

PT

of Nucleic Acid Sequencing of the D1/D2 Region of the Large Subunit of the 28S rDNA and the Internal Transcribed Spacer Region Using SmartGene IDNS Software

RI

for Identification of Filamentous Fungi in a Clinical Laboratory, J. Mol. Diagnostics.

SC

14 (2012) 393–401.

[110] M. Andrew, T.L. Peever, B.M. Pryor, An expanded multilocus phylogeny does not

NU

resolve morphological species within the small-spored Alternaria species complex, Mycologia. 101 (2009) 95–109.

MA

[111] G.S. de Hoog, R. Horré, Molecular taxonomy of the Alternaria and Ulocladium species from humans and their identification in the routine laboratory, Mycoses. 45 (2002) 259–276.

D

[112] S. Kühnel, H.A. Schols, H. Gruppen, Aiming for the complete utilization of sugar-beet

PT E

pulp: Examination of the effects of mild acid and hydrothermal pretreatment followed by enzymatic digestion, Biotechnol. Biofuels. 4 (2011) 14. [113] J. Thibault, X. Rouau, Studies on Enzymic Hydrolysis of Polysaccharides in Sugar

CE

Beet Pulp, Carbohydr. Polym. 13 (1990) 1–16. [114] T. Sakamoto, S. Nishimura, T. Kato, Y. Sunagawa, M. Tsuchiyama, H. Kawasaki,

AC

Efficient Extraction of Ferulic Acid from Sugar Beet Pulp Using the Culture Supernatant of Penicillium chrysogenum, J. Appl. Glycosci. 52 (2005) 115–120. [115] C. Brézillon, P. Kroon, C. Faulds, G.M. Brett, G. Williamson, Novel ferulic acid esterases are induced by growth of Aspergillus niger on sugar-beet pulp, Appl. Microbiol. Biotechnol. 45 (1996) 371–376. [116] E. Bonnin, H. Grangé, L. Lesage-Meessen, M. Asther, J.F. Thibault, Enzymic release of cellobiose from sugar beet pulp, and its use to favour vanillin production in Pycnoporus cinnabarinus from vanillic acid, Carbohydr. Polym. 41 (2000) 143–151. [117] N.W. Jaworski, H.N. Lærke, K.E. Bach Knudsen, H.H. Stein, Carbohydrate 37

ACCEPTED MANUSCRIPT composition and in vitro digestibility of dry matter and nonstarch polysaccharides in corn, sorghum, and wheat and coproducts from these grains, J. Anim. Sci. 93 (2015) 1103–1113. [118] L. Saulnier, J.F. Thibault, Ferulic acid and diferulic acids as components of sugar-beet pectins and maize bran heteroxylans, J. Sci. Food Agric. 79 (1999) 396–402. [119] V. Micard, C.M.G.C. Renard, J.F. Thibault, Enzymatic saccharification of sugar-beet

PT

pulp, Enzyme Microb. Technol. 19 (1996) 162–170.

[120] I. Benoit, D. Navarro, N. Marnet, N. Rakotomanomana, L. Lesage-Meessen, J.-C.

RI

Sigoillot, M. Asther, M. Asther, Feruloyl esterases as a tool for the release of phenolic

SC

compounds from agro-industrial by-products, Carbohydr. Res. 341 (2006) 1820–1827. [121] R. López Alonso, C. Ramírez de Lara, L. Rivero, A.E. Ruiz, C.T. Torres Zapata, R. V

NU

Ullán, Aplicación de una nueva FAE en la liberación químico-enzimática de ácido ferúlico a partir de pulpa de remolacha, Ciencias Agronómicas. 19 (2011) 21–25.

MA

[122] A. Aarabi, M. Mizani, M. Honarvar, H. Faghihian, A. Gerami, Extraction of ferulic acid from sugar beet pulp by alkaline hydrolysis and organic solvent methods, J. Food Meas. Charact. 10 (2016) 42–47.

D

[123] D.W.S. Wong, Feruloyl Esterase: A Key Enzyme in Biomass Degradation, Appl.

PT E

Biochem. Biotechnol. 133 (2006) 87–112. [124] E. Topakas, H. Stamatis, P. Biely, D. Kekos, B.. Macris, P. Christakopoulos, Purification and characterization of a feruloyl esterase from Fusarium oxysporum

CE

catalyzing esterification of phenolic acids in ternary water–organic solvent mixtures, J.

AC

Biotechnol. 102 (2003) 33–44. [125] V.F. Crepin, C.B. Faulds, I.F. Connerton, A non-modular type B feruloyl esterase from Neurospora crassa exhibits concentration-dependent substrate inhibition, Biochem. J. 370 (2003) 417–427. [126] A. Andersen, A. Svendsen, J. Vind, S.. Lassen, C. Hjort, K. Borch, S.. Patkar, Studies on ferulic acid esterase activity in fungal lipases and cutinases, Colloids Surfaces B Biointerfaces. 26 (2002) 47–55. [127] J.-W. Lee, K.-S. Gwak, J.-Y. Park, M.-J. Park, D.-H. Choi, M. Kwon, I.-G. Choi, Biological pretreatment of softwood Pinus densiflora by three white rot fungi, J. Microbiol. 45 (2007) 485–91. 38

ACCEPTED MANUSCRIPT [128] A. Miettinen-Oinonen, M. Paloheimo, R. Lantto, P. Suominen, Enhanced production of cellobiohydrolases in Trichoderma reesei and evaluation of the new preparations in biofinishing of cotton, J. Biotechnol. 116 (2005) 305–317. [129] M. Zoglowek, P.S. Lübeck, B.K. Ahring, M. Lübeck, Heterologous expression of cellobiohydrolases in filamentous fungi – An update on the current challenges, achievements and perspectives, Process Biochem. 50 (2015) 211–220.

PT

[130] T.T. Lai, T.T.H. Pham, K. Adjallé, D. Montplaisir, F. Brouillette, S. Barnabé, Production of Trichoderma Reesei RUT C-30 Lignocellulolytic Enzymes Using Paper

RI

Sludge as Fermentation Substrate: An Approach for On-Site Manufacturing of

SC

Enzymes for Biorefineries, Waste and Biomass Valorization. 8 (2017) 1081–1088. [131] R.A. Batista-García, T. Sutton, S.A. Jackson, O.E. Tovar-Herrera, E. Balcázar-López,

NU

M.D.R. Sánchez-Carbente, A. Sánchez-Reyes, A.D.W. Dobson, J.L. Folch-Mallol, Characterization of lignocellulolytic activities from fungi isolated from the deep-sea

MA

sponge Stelletta normani, PLoS One. 12 (2017) e0173750. [132] J.A. Buswell, Y.J. Cai, S.T. Chang, J.F. Peberdy, S.Y. Fu, H.S. Yu, Lignocellulolytic enzyme profiles of edible mushroom fungi, World J. Microbiol. Biotechnol. 12 (1996)

AC

CE

PT E

D

537–542.

39

ACCEPTED MANUSCRIPT Tables Table 1. Enzymatic Index (EI) of the candidates showing FAE activity. Values are given as

8,21 ± 0,86

JCOS10C

4,13 ± 0,34

PDA8

3,90 ± 0,83

PDA(2)2

3,88 ± 0,42

PDA(2)10

3,61 ± 1,02

PDA(3)1

3,33 ± 0,29

PDA(2)4

3,27 ± 0,37

RI

PDA1

SC

EI

NU

Isolate

PT

means, including standard deviations. The best producer is highlighted in bold.

MA

Table 2. Specific activities and kinetic parameters of the enzyme extract for methyl ferulate, methyl p-coumarate, methyl caffeate and methyl synapate. Values are given as means,

D

including standard deviations.

Specific activity (U/g total

PT E

Substrate

protein)

MFA

MCA

a

AC

MSA

CE

MpCA

Km (mM)

Vmax (U/g total protein)

88.93 ± 0.67

0.43 ± 0.05

104.01 ± 2.38

168.53 ± 1.60

0.47 ± 0.05

218.01 ± 5.28

96.29 ± 3.54

0.36 ± 0.05

155.04 ± 4.32

7.60 ± 0.32

N/Aa

N/Aa

N/A: not applicable

40

ACCEPTED MANUSCRIPT Table 3. Protein spots identified by peptide mass fingerprinting and tandem mass spectrometry present in the enzyme extract from A. alternata PDA1. MOWSE score from Mascot search (Score) and a predictive analysis for the presence of signal peptides with SignalP (Signal pep) and non-classical secretion (Non-classical secretion) are described. COG classes (COG) are also included. Further information about sequence coverage, matched and non-matched peptides, RMS error, MW, pI, and global functional classification

1

A0A177DVF3_ALTAL

2

A0A177DVF3_ALTAL

A0A177E3B6_ALTAL

A0A177E3B6_ALTAL 3

Protein description

a

Score

Uncharacterized protein (β-glucosidase/βxylosidase) Carboxylic ester hydrolase Uncharacterized protein (β-glucosidase/βxylosidase) Carboxylic ester hydrolase

Signal pep

Nonclassical secretion

COG

RI

Entry name (UNIPROT)

267

YES

G

244

YES

I/E

565

YES

G

202

YES

I/E

SC

Prot. No.

PT

can be found in Supplementary File S4.

A0A177D675_ALTAL

Subtilisin-like protein

267

YES

O

A0A177E3B6_ALTAL

232

YES

I/E

150

YES

G

168

YES

O

146

YES

not assigned

137

YES

I/E

5

A0A177D675_ALTAL

Subtilisin-like protein

298

YES

O

6

A0A177D675_ALTAL

264

YES

O

127

YES

G

87

YES

G

320

YES

O

237

NO

209

YES

185

NO

398

YES

G

A0A177D4E0_ALTAL

Subtilisin-like protein Uncharacterized protein (rhamnogalacturonan lyase) Uncharacterized protein (rhamnogalacturonan lyase) Subtilisin-like protein Uncharacterized protein (glycoside hydrolase family 31 protein) Subtilisin-like protein Uncharacterized protein (glycoside hydrolase family 31 protein) Uncharacterized protein (rhamnogalacturonan lyase) Tripeptidyl-peptidase 1

257

YES

O

A0A177DZ08_ALTAL

Sucrose-6-phosphate hydrolase

574

YES

G

A0A177D4E0_ALTAL

Tripeptidyl-peptidase 1

504

YES

O

A0A177DNT5_ALTAL

Carboxylic ester hydrolase Uncharacterized protein (rhamnogalacturonan lyase) Subtilisin-like protein Uncharacterized protein (no putative conserved domains) Uncharacterized protein (ubiquitin 3 binding protein But2 C-terminal domain; cell division protein DedD) Uncharacterized protein (ubiquitin 3 binding protein But2 C-terminal domain; cell division protein DedD) Uncharacterized protein (ubiquitin 3 binding protein But2 C-terminal domain; cell division protein DedD) Carboxylic ester hydrolase Uncharacterized protein (ubiquitin 3 binding protein But2 C-terminal domain; cell division protein

193

YES

I

284

YES

G

214

YES

O

386

YES

not assigned

273

YES

D

137

YES

D

158

YES

D

254

YES

I/E

462

YES

D

A0A177DB39_ALTAL 7

A0A177DB39_ALTAL

8

A0A177D675_ALTAL A0A177DPL2_ALTAL

9

A0A177D675_ALTAL

11 12

13

A0A177DB39_ALTAL

AC

10

CE

A0A177DPL2_ALTAL

MA

A0A177D675_ALTAL A0A177D3A4_ALTAL

D

4

PT E

A0A177DVF3_ALTAL

NU

A0A177E3B6_ALTAL

Carboxylic ester hydrolase Uncharacterized protein (β-glucosidase/βxylosidase) Subtilisin-like protein Uncharacterized protein (no putative conserved domains) Carboxylic ester hydrolase

A0A177DB39_ALTAL A0A177D675_ALTAL

14

A0A177DCA7_ALTAL

15

A0A177D4W2_ALTAL

16

A0A177D4W2_ALTAL

17

A0A177D4W2_ALTAL

18

A0A177DI60_ALTAL

19

A0A177D4W2_ALTAL

41

YES

G O

YES

G

ACCEPTED MANUSCRIPT DedD) 20

A0A177DCA7_ALTAL

21

A0A177DCA7_ALTAL

A0A177D4C3_ALTAL

A0A177D4C3_ALTAL 22

A0A177DCA7_ALTAL

YES

not assigned

145

YES

G

153

YES

not assigned

152

YES

G

169

YES

not assigned

111

YES

G

260

YES

G

94

YES

G

Q9UVP4_ALTAL

112

NO

25

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

957

YES

G

26

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

952

YES

G

27

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

732

YES

G

28

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

545

YES

G

29

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

513

YES

G

30

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

647

YES

G

A0A177DKY7_ALTAL

Glucanase

241

YES

G

31

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

931

YES

G

32

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

997

YES

G

33

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

888

YES

G

34

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

664

YES

G

35

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

849

YES

G

36

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

995

YES

G

37

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

529

YES

G

A0A177DUN7_ALTAL

Glycosyl hydrolase family 43 protein

396

YES

G

38

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

666

YES

G

39

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

877

YES

G

40

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

746

YES

G

41

A0A177DQV8_ALTAL

Phosphoesterase-domain-containing protein

413

YES

M/P

Acid phosphatase

240

YES

M/P

Arabinofuranosidase/B-xylosidase

143

YES

Glycoside hydrolase

370

NO

Arabinofuranosidase/B-xylosidase

414

YES

Ribonuclease T2

215

NO

A0A0L1HZ20_9PLEO A0A177D5S3_ALTAL 42

A0A177DUC6_ALTAL

SC

NU

MA

D

A0A177DE80_ALTAL

PT

24

Sucrose-6-phosphate hydrolase Uncharacterized protein (glycoside hydrolase family 79, β-glucuronidase) Glucanase

PT E

A0A177DZ08_ALTAL

202

RI

A0A177D4C3_ALTAL 23

Uncharacterized protein (no putative conserved domains) Polygalacturonase Uncharacterized protein (no putative conserved domains) Polygalacturonase Uncharacterized protein (no putative conserved domains) Polygalacturonase

NO

G

G NO

M

A0A177D5S3_ALTAL

44

A0A177DVG9_ALTAL

45

A0A177D1Q5_ALTAL

Pectin lyase-like protein

476

YES

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

196

YES

46

A0A177DSP2_ALTAL

Acid protease

612

NO

47

A0A177DXD0_ALTAL

Endopolygalacturonase

100

YES

G

48

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

703

YES

G

49

A0A177DXD0_ALTAL

Endopolygalacturonase

496

YES

G

50

A0A177DXD0_ALTAL

Endopolygalacturonase

610

YES

G

51

A0A177DWS3_ALTAL

Acid protease

591

YES

O

52

A0A177DWS3_ALTAL

Acid protease

774

YES

O

53

A0A177D4F4_ALTAL

Pectin lyase-like protein

142

YES

G

54

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

94

YES

G

55

A0A177DR40_ALTAL

Pectin lyase A

165

YES

G

56

A0A177DR40_ALTAL

Pectin lyase A

494

YES

G

57

A0A177DWS3_ALTAL

Acid protease

616

YES

O

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

100

YES

G

58

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

512

YES

G

59

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

333

YES

G

60

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

86

YES

G

AC

CE

43

42

G YES

J G G

YES

O

ACCEPTED MANUSCRIPT 61

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

577

YES

G

62

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

370

YES

G

63

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

313

YES

G

64

A0A177DWS3_ALTAL

Acid protease

487

YES

O

Endopolygalacturonase

301

YES

G

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

89

YES

G

66

A0A177D4F4_ALTAL

Pectin lyase-like protein

546

YES

G

67

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

586

YES

G

68

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

392

YES

G

69

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

487

YES

G

70

A0A177D5G1_ALTAL

Endopolygalacturonase

325

YES

G

71

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

107

YES

G

72

A0A177DTI8_ALTAL

Arabinogalactan endo-beta-1,4-galactanase

149

YES

G

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

115

YES

G

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

274

YES

G

166

YES

G

373

YES

G

503

YES

G

RI

73

PT

A0A177DXD0_ALTAL 65

Endopolygalacturonase

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

75

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

76

A0A177DTI8_ALTAL

Arabinogalactan endo-beta-1,4-galactanase

220

YES

G

77

A0A177DIB2_ALTAL

261

YES

U/I

78

A0A0L1I1D6_9PLEO

91

YES

not assigned

79

A0A177D4F4_ALTAL

Alpha/beta-hydrolase Uncharacterized protein (no putative conserved domains) Pectin lyase-like protein

514

YES

G

80

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

178

YES

G

81

A0A177D3Z9_ALTAL

Glycoside hydrolase

570

YES

G

82

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

630

YES

G

83

A0A177E3J2_ALTAL

Arabinan endo-1,5-alpha-L-arabinosidase

528

YES

G

84

A0A177E3J2_ALTAL

Arabinan endo-1,5-alpha-L-arabinosidase

560

YES

G

85

A0A177DGQ5_ALTAL

Subtilisin-like serine protease-like protein PR1A

157

YES

O

A0A177E3J2_ALTAL

Arabinan endo-1,5-alpha-L-arabinosidase

153

YES

G

D

MA

NU

SC

A0A177DXD0_ALTAL 74

A0A177DAL9_ALTAL

SGNH hydrolase

130

YES

E

87

A0A177DZE3_ALTAL

Concanavalin A-like lectin/glucanase (Fragment)

304

YES

G

88

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

146

YES

G

89

A0A177D5S3_ALTAL

Arabinofuranosidase/B-xylosidase

658

YES

G

90

A0A177DXD0_ALTAL

Endopolygalacturonase

441

YES

G

Arabinofuranosidase/B-xylosidase

159

YES

G

Arabinofuranosidase/B-xylosidase

877

YES

G

A0A177D5S3_ALTAL

PT E

86

A0A177D5S3_ALTAL

92

A0A177DH30_ALTAL

Endoglucanase A

500

YES

G

93

A0A0L1HTJ6_9PLEO

284

YES

not assigned

255

NO

CE

91

94

A0A177D761_ALTAL

Cell wall protein Uncharacterized protein (no putative conserved domains) Cutinase

150

YES

G

95

A0A177E449_ALTAL

Endo-1,4-beta-xylanase

167

YES

G

96

A0A177D506_ALTAL

Deuterolysin metalloprotease

483

YES

O

97

Q71SP0_ALTAL

Alt a 1

686

YES

not assigned

AC

A0A177D114_ALTAL

a

NO

not assigned

The assignment of the biological or enzymatic function to the hypothetical proteins (based on BLASTP analyses) is shown between parentheses and in italic characters.

43

ACCEPTED MANUSCRIPT Table 4. Cellulase and xylanolytic activities of the enzyme extract. Reactions were performed at 50 ºC, in acetate buffer 0.1 N pH 5.0. One unit corresponds to the amount of enzyme that releases one µmole of product (4-Nitrophenol for cellobiohydrolase, β-glucosidase and β-xylosidase; glucose for endoglucanase; xylose for endoxylanase) per min at the tested conditions. Values are given as means, including standard deviations. Measured activity

Specific activity (U/g total

9.99 ± 0.52

Endoglucanase

547.03 ± 49.29

β-xylosidase

250.98 ± 10.57

Endoxylanase

280.48 ± 16.77

RI

Cellobiohydrolase

SC

1147.07 ± 126.40

NU

β-glucosidase

PT

protein)

MA

Table 5. Amounts of simple sugars released when adding A. alternata PDA1 enzyme extract to a solution of a natural lignocellulosic substrate: sugar beet pulp. Values are given as

Sucrose Glucose

Fructose

CE

Galactose

a

AC

Arabinose

Saccharification power (g/L released)

PT E

Monosaccharide/disaccharide

D

means, including standard deviations.

N/Da 1.57 ± 0.10 N/Da 1.35 ± 0.03 2.18 ± 0.12

N.D: non-detected

44

ACCEPTED MANUSCRIPT Figure captions Fig. 1. Optical microscope images of A. alternata PDA1 conidia. Scale bar = 25 µm.

Fig. 2. A) FAE activity of A. alternata PDA1 (U/L fermentation broth) along fermentation on different natural lignocellulosic substrates. M-SBP: sugar beet pulp powder; M-OAT:

PT

processed oat fiber powder; M-CORN: processed corn powder; M-WHEAT: processed wheat powder. B) FAE activity (U/g total protein in the enzyme extract) and dry weight of the

RI

culture A. alternata PDA1 (g/L fermentation broth), when grown on sugar beet pulp powder

SC

(M-SBP).

NU

Fig. 3. Effect of pH and temperature on FAE activity of A. alternata PDA1 enzyme extract against methyl ferulate. Data expressed as relative FAE values compared to the maximum

MA

(100%). A) FAE activity dependence on pH, at 37 ºC. B) FAE activity dependence on temperature, at pH 5.0. C) Thermal stability. Residual FAE values of the enzyme extract, after incubating for one hour at the indicated temperature and subsequently FAE activity

PT E

D

analysis under optimal conditions.

Fig. 4. Michaelis-Menten kinetics plotted using SigmaPlot® 12, for methyl ferulate, methyl

CE

p-coumarate and methyl caffeate, at the optimal assay conditions. A) Kinetics for methyl

AC

ferulate 13. B) Kinetics for methyl p-coumarate. C) Kinetics for methyl caffeate.

Fig. 5. Reference map of the A. alternata PDA1 partial extracellular proteome. A total of 112 protein spots were visualized in a 3.0-10.0 pH range, 97 of which (numbered in the figure) were successfully identified. Molecular weights are marked on the left and the assigned numbers correspond to those in Supplementary File S4.

Fig. 6. A) Number of protein spots sorted according to their relative volume referred to the total spot volume. Only 13 out of 112 spots showed a relative volume above 1% of the total. 45

ACCEPTED MANUSCRIPT B) COG classification of all the identified proteins. Category identification goes as follows: D (cell cycle control and mitosis), E (amino acid metabolism), G (carbohydrate metabolism and transport), I (lipid metabolism), J (translation), M (cell wall/membrane/envelope biogenesis), O (post-translational modification, protein turnover, chaperone functions), P

AC

CE

PT E

D

MA

NU

SC

RI

PT

(inorganic ion transport and metabolism) and U (intracellular trafficking and secretion).

46

ACCEPTED MANUSCRIPT

Significance Although plant biomass is an abundant material that can be potentially utilized by several industries, the effective hydrolysis of the recalcitrant plant cell wall is not a straightforward process. As this hydrolysis occurs in nature relying almost solely on microbial enzymatic systems, it is reasonable to infer that further studies on lignocellulolytic enzymes will

PT

discover new sustainable industrial solutions. The results included in this paper provide a promising fungal candidate for biotechnological processes to obtain added value from plant

RI

byproducts and analogous substrates. Moreover, the proteomic analysis of the secretome of a

SC

natural isolate of Alternaria sp. grown in the presence of one of the most used vegetal substrates on the biofuels industry (sugar beet pulp) sheds light on the extracellular enzymatic machinery of this fungal plant pathogen, and can be potentially applied to developing new

NU

industrial enzymatic tools. This work is, to our knowledge, the first to analyze in depth the secreted enzyme extract of the plant pathogen Alternaria when grown on a lignocellulosic

MA

substrate, identifying its proteins by means of MALDI-TOF/TOF mass spectrometry and

AC

CE

PT E

D

characterizing its feruloyl esterase, cellulase and xylanolytic activities.

47

ACCEPTED MANUSCRIPT Highlights

1. New enzymatic tools are demanded and required by the biomass degrading industries.

PT

2. Screening for feruloyl esterase (FAE) producing fungi was done on natural isolates.

RI

3. Best candidate, Alternaria alternata PDA1, showed FAE, cellulase and xylanolytic

SC

activities.

NU

4. The reference map of the A. alternata PDA1 protein extract was elaborated.

AC

CE

PT E

D

MA

5. Ninety-seven protein spots were identified and assigned to functional categories.

48

Graphics Abstract

Figure 1

Figure 2

Figure 3

Figure 4

Figure 5

Figure 6