Nigrospora Classification Essay

1. Introduction

In recent decades, marine organisms focused the attention of researchers for their huge potential in producing bioactive compounds [1,2,3,4]. Among them, the microorganisms gradually took on an important role because they appear to be prolific producers of a wide diversity of secondary metabolites [5,6]. Cultivated in bioreactors, they also represent a sustainable and easily “upscalable” resource, therefore not endangering fragile marine ecosystems. Among these microorganisms, fungi, which had their terrestrial glory days after the discovery of penicillin, are again at the forefront in the search for new marine molecules in view of their rich biodiversity, even in deep-sea niches [7]. Around 70,000 fungal species have already been described worldwide, and among them about 1500 species of marine-derived fungi were mentioned, primarily from coastal ecosystems [8,9]. As 70% of the earth is submerged, Gareth Jones (1998) [10] estimated the total number of marine and marine-derived fungal species to be a minimum of 72,000, indicating that the inherent discovery of new compounds is still in its infancy. Several fungal metabolites from marine origin have already demonstrated their originality and efficacy in different domains. As an example, in the field of therapeuthics, we can mention the two chemically unusual cyclodepsipeptides patented in 2008—scopularides A and B—from the marine-derived Scopulariopsis brevicaulis, demonstrating anticancer activities [11,12], as well as the halimide (plinabuline) from an endophytic Aspergillus sp. CNC-139 isolated from the green algae Halimeda lacrymosa, already achieving phase II clinical tests [13].

Biosynthetically, many extrolites produced by filamentous fungi are polyketides, and several papers report that polyketides seem to dominate marine natural products of fungal origin [14,15]. Polyketides represent an array of often structurally complex natural products including such classes as anthraquinones, hydroxyanthraquinones, naphthalenes, naphthoquinones, flavonoids, macrolides, polyenes, tetracyclines, and tropolones. Many of them have already exhibited either positive or negative effects, as wide as those that are antimicrobial, anticancer, antioxidant, immunomodulatory, cytotoxic, or carcinogenic. This closely concerns the class of anthraquinones whose effects, depending on the nature and amount of compound, can either be beneficial or noxious towards living organisms. These compounds, little studied because of their bad reputation, mainly arising from their benzenic patterns, are, however, worthy of the same attention as other families of fungal compounds, whose members have become pillars of the global pharmacopeia (antibiotics) and are widely used in food or staining industries (azaphilone colorants from Monascus spp. in Asia). Within this review, we investigate the present knowledge of the anthraquinonoid compounds listed to date from marine-derived filamentous fungi′s productions. This overview highlights the molecules identified for the first time and comes along with interesting characteristics: the panel of colors, their known roles in the biology of the organisms, and some specific in vitro biological activities. As the natural products constitute the dynamic element of the present global market, we hope this review can help broadening the horizon towards innovative substances.

2. Anthraquinones from Marine-Derived Fungi

About 700 anthraquinone derivatives were identified in plants, lichens, and fungi; 43 have already been described from fungal cultures [16,17]. Due to their structure, they exhibit interesting chromatic properties and decline a wide range of nuances in colors. Thus, they first presented a great interest in the field of dyeing molecules, highly requested in cosmetics, clothes dyeing and foodstuff industries. From their structures, hydroxyanthraquinone pigments have a relative stability. They also possess good light-fastness properties, which often makes metallization unnecessary. Nevertheless, they can easily form complexes with several metal salts (or cations in general) (aluminium, barium, calcium, copper, palladium, iron) [18,19,20,21,22] and exhibit superior brightness compared to azo-pigments [23,24]. This capacity to form metallic complexes is of a great interest in an industrial context: The complex forms often reduce the solublity in water, enhancing the solvent solubility, without loosing the brightness [20]. In the textile industry, hydroxyanthraquinone are, moreover, considered “reactive dyes,” as they form a covalent bond with the fibers, usually cotton, although they are used to a small extent on wool and nylon. Therefore, they have it it possible to achieve extremely high wash fastness properties by relatively simple dyeing methods. Thereby, the literature abundantly reports the interest for marine organisms with respect to the production of new molecules and, among them, new pigments [25,26]. Besides their coloring properties, anthraquinoid compounds exhibit a wide range of diverse biological activities, sparsely mentioned in the literature. Regarding this aspect, their “Dr Jekyll and Mr Hyde” physiognomy [27] needs to be carefully elucidated in order to examine their potential use in pharmaceutical or alimentary fields, with maximum objectivity.

2.1. Anthraquinone′s Basic Structure

Anthraquinones represent a class of molecules of the quinone family, based on a structure composed of three benzene rings. The basic structure 9,10-anthracenedione, also called 9,10-dioxoanthracene (formula C14H8O2), includes two ketone groups on the central ring (Figure 1). The diversity of the anthraquinoid compounds relies on the nature and the position of the substituents, replacing the H atoms on the basic structure (R1 to R8), as diverse as: –OH, –CH3, –OCH3, –CH2OH, –CHO, –COOH, or more complex groups. When n hydrogen atoms are replaced by hydroxyl groups, the molecule is called hydroxyanthraquinone (HAQN). From their structure, HAQN derivatives absorb visible light and are colored.

An important characteristic of the anthraquinone compound is their electronic absorption spectra. The strong absorption in the ultraviolet region is due to the presence of chromophore formed by the system of conjugated double bonds. The spectra of anthraquinone are highly complex because of the presence of absorption bands due to the benzenoid transitions, in addition to quinonoid absorptions. The benzenoid bands appear fairly regularly within the range 240–260, with intense absorption at 250 nm and in 320–330 nm, and with medium absorption at 322 nm, whereas the quinonoid bands absorb at 260–290 nm. These areas of selective absorption are characteristic, and the pattern in the ultraviolet region is not seriously affected by substitution. In addition, hydroxyl anthraquinones show an absorption band(s) at 220–240 nm, not shown by the parent compound. In the visible area, an unsubstituted anthraquinone has a weak yellow color, and its electronic absorption spectrum contains a small peak at 405 nm. The presence of substituents in position 1 and 4 induces a significant bathochromic shift, intensifying the color more significantly than the substituents in the 1,5 and 1,8 positions. Thus, with an alcoholic solution of magnesium acetate, 1,2-dioxyderivative is colored in violet; 1,4-dioxyderivative in purple; and 1,8-dioxyderivative in red-orange [28,29,30]. Therefore, fungal anthraquinones range from pale yellow to dark red or brown colors, through to violet.

2.2. Ecology of Marine-Derived Fungal Anthraquinones Producers

Marine ecosystems host a wide biodiversity of filamentous fungi found in free waters, inert organic or inorganic matter. They can also be included as endophytes or pathogens in marine plants, planktons, vertebrates and invertebrates [31]. Their different roles in these environments are still poorly known, although their implications in lignocellulolytic compounds degradation and mineralization of organic matter has been repeatedly demonstrated [7,32,33]. Yet the notion of “marine fungus” is still under debate in the world of mycologists. Fungi are usually recognized as ubiquitous because they inhabit a plethora of ecosystems, from terrestrial milieus to aquatic environments (Figure 2). Marine and marine-derived fungi therefore form an ecological, not a taxonomic, group [34]. From the widely adopted definition of Kohlmeyer et al. (1979) [8], they are divided into two ecotypes:
  • obligate marine fungi (true ones) that grow and sporulate only in seawater. Their spores are able to germinate and form new thalli in salted environment.

  • transitional marine fungi (marine-derived fungi) that come from terrestrial or freshwater media and have undergone physiological adaptation to survive, grow, or reproduce in the marine environment.

In fungi, anthraquinones are produced from different steps or branches of the polyketides pathway. Today, it is clear that, as far as secondary metabolites and a priori anthraquinoid productions are concerned, a great variability appears among species of the same genus, even among strains in the same species. This could undoubtly be related to the capacities a fungus has to develop, in order to face some specific conditions in specific ecosystems. As an illustration, the composition of the quinoid pigment complexes of P. funiculosum strains isolated from various types of soils are quite different when cultivated in the same artificial culture media [35]. That is why, even if the polyketides pathway is mentioned in a strain, not all strains inside the species are anthraquinones producers. In the same way, if the fungal metabolism is able to express anthtraquinones and (simultaneously) to excrete toxins, the presence of these secondary products are highly dependent on external physico-chemical conditions [36].

Thus, a high diversity of molecules is now expected from unexplored marine-derived fungi, which are considered promising novel sources of chemical diversity. The potential of marine-derived microorganisms to produce unique and original molecules could therefore come from specific metabolic or genetic adaptations appearing to meet very specific combinations of physico-chemical parameters (high osmotic pressure, low O2 penetration, low temperature, limited light access, high pressure, or regular tidal ebbs and flows) [37]. Indeed, the two marine ecotypes lead to particular behaviors and consecutively to specific products, compared to the terrestrial congeners: either the challenge of facing unusual living conditions (exogenous fungi) or the use of specific procedures naturally adapted to the marine niches (i.e., indigenous micromycetes, naturally selected for aquatic environments). This skill is, for instance, exemplified by marine macroorganisms′ fungal endophytes as corals or sponges. For now, the highest diversity of marine-derived fungi seems to be found in tropical regions, mainly in tropical mangroves, which are extensively studied because of their high richness in organic matters. Obviously, these biotopes seem favorable to the development of a high diversity of heterotrophic microorganisms based on the diversity of organic and inorganic substrates [8,38].

The questions on the effect of interactions between organisms on microbes extrolites is amply fueled in the case of a very producive lichen′s symbioses. A lichen is a composite organism that emerges from algae or cyanobacteria (or both) living with filaments of a fungus in a mutually beneficial (symbiotic) relationship. About 20,000 lichen species are known in the world, and there are approximately 700 species known from coastal rocks and urbanized shores [39]. Most work on aquatic lichens was done in temperate areas, as, in the tropics, lichens are less developed on costal rocks. One interesting skill is that lichen associations are primarily terrestrial but require alternate wetting and drying regimes for their survival. In marine environments, these circumstances occur principally in tidal zones on coastal rocks, subject to varying water levels and different degrees of inundation. Another feature is that the whole combined life form has properties that are very different from properties of its component organisms alone. Thus, tropical stream margins are promising biota for species and therefore compounds that are new to science.

2.3. Structural Diversity and Colors of Anthraquinoid Extrolites from Marine-Derived Fungi

2.3.1. Present Knowledge about Anthraquinonoid Compounds from Fungi

Today′s knowledge indicates that a large part of compounds identified in terrestrial fungi can often be isolated from the same species living in marine environments. For instance, catenarin, emodin, erythroglaucin, physcion, questin, and rubrocristin or physcion anthrone are produced by marine-derived Aspergillus and/or Eurotium species, as well as by their terrestrial counterparts. According to Bick et al. and Fain et al. [40,41], the most widespread anthraquinones in fungi are 1,8-dihydroxy and 1,5,8 or 1,6,8-trihydroxy anthraquinone derivatives. They appear either as simple forms, as glycosides, or other complexes attached through an O- or C-bond in the side chain, which can enhance the water solubility. Some dimeric structures (formed through C–C bonds) are also produced from fungi, (e.g., alterporriols, skyrin, rubroskyrin, luteoskyrin, icterinoidin, rubellin, rufoolivacin, etc.). Some dimers may contain not only monomeric anthraquinones but also naphthoquinones and other products of polyketide synthesis. According to Fujitake et al. [42,43] and Suzuki et al. [44], the dimeric anthraquinone 5,5′-biphyscion (named hinakurin), chrysotalunin, (−)-7,7′-biphyscion, microcarpin, chrysophanol, and physcion are predominant in soil, but they seem rare in organisms. If hinakurin, chrysotalunin, and (−)-7,7′-biphyscion have not been found in fungi yet, it is now clear that chrysophanol and physcion are frequent fungal productions, and that fungi are able to synthesize dimers. These organisms, widely represented in soil, may have a transient appearance of monomeric and dimeric anthraquinones in telluric biotopes, certainly evolving to complex (humic) polymers. However, these statements rely on decades of terrestrial studies. The increase in knowledge on marine and marine-derived anthraquinones from fungi will certainly elucidate this aspect.

2.3.2. Nature and Colors of Compounds from Marine-Derived Fungi

Most anthraquinoid compounds of natural origin have complex structures with several functional substituent groups. In nature and/or cultures, a wide range of hues appears, from pale yellow to dark brown, through to orange, red, or violet pigmentations. Anthraquinoid compounds generaly color sexual stages or resistance forms (ascomata, spores, conidia, etc.) but sometimes also impregnate mycelium or are excreted in the growth environment. Thus, the structural localization and the colors of these secondary metabolites seem to highly depend on the fungal species and may vary with the amount of compound produced in relation with the environmental conditions.

Genera and Species

Many genera producing anthraquinones have been isolated from marine environments, either from water, sediments, or decaying plants or from living organisms such as invertebrates, plants (endophytes), and algae. To date, strains in the genera—Alternaria, Aspergillus, Eurotium, Fusarium, Halorosellinia, Microsphaeropsis, Monodictys, Nigrospora, Paecilomyces, Penicillium, Phomopsis, and Stemphylium—have been clearly mentioned as marine-derived anthraquinones producers.

Some of the common terrestrial genera—Aspergillus, Eurotium, Alternaria, Penicillium and Fusarium—have been extensively investigated concerning their secondary metabolite′s productions, and the anthraquinoid molecules produced were reviewed by Velmurugan et al. [45], Caro et al. [16], and Gessler et al. [17

1. Introduction

Vascular plants species [1,2], aquatic plants and algae [3,4], mosses and ferns [5,6] examined to date are found to be hosts for endophytic bacteria and fungi [7]. Endophytic microorganisms have been isolated from different parts of plant-like scale primordia, meristem and resin ducts [8,9], leaf segments with midrib and roots, stem, bark, leaf blade, petiole [10], buds [11], and seeds [12]. Successful endophytic colonization is dependent on many factors including plant tissue type, plant genotype, the microbial taxon and strain type, and biotic and abiotic environmental conditions. Fungal endophytes aid plants to withstand and tolerate unfavorable environmental conditions [13,14] and also promote plant growth [15,16]. These inhabitants can produce the same or similar secondary metabolites [17,18,19,20] as their host and play vital roles in vivo such as signaling, defense, and regulation of the symbiosis [21]. Mainly investigations are based on their use as biochemical tools and the end products are to be used in pharmaceutics, industry, and agriculture.

Artemisia is a plant highly evaluated for medicinal and biopesticide traits. A survey of the literature shows this plant genus to be in the hot spot among researchers with over 11,200 publications in Scopus library. Even though Artemisia is a large plant genus with species producing a variety of interesting and active compounds, its endophytic communities are under investigated. The identification of the fungal endophytes in Artemisia spp. is made mainly based on morphological characterization and molecular analysis using nuclear ribosomal DNA sequences, including both the internal transcribed spacers and the 5.8S gene region. To the best of our knowledge, there have been only four studies which investigate the phylogenetic analysis of the Artemisia spp. fungal endophytes [22,23,24,25]. In terms of diversity, the studies are also scarce but interesting facts are brought to light in terms of diversity and plant colonization. For instance, Yuan et al., 2011 [26] performed a comparative study related to infection frequency between cultivated plants and wild plants of Artemisia annua. The results revealed slightly higher infection frequency of the endophytic fungi in cultivated roots (20.9%) than in native roots (16.7%). Further, authors described that the naturally regenerated roots harbored richer fungal genotypes, which supports the hypothesis that wild plant species are predisposed to host rich and novel mycoflora [27]. It is worth mentioning that Qian et al., 2014 [27] reported the presence of Rhodotorula sp. and Fusarium sp. in Artemisia argyi for the first time. The endophytic fungi associated with Artemisia nilagirica were investigated and one strain of Pythium intermedium (Oomycota) and one strain of Rhizopus oryzae (Mucoromycota) were isolated among the majority clade of Ascomycota [28]. Huang et al., 2009 [24] classified 108 fungal isolates obtained from three medicinal plant species Artemisia capillaris, Artemisia indica and Artemisia lactiflora using morphological identification and among the three plant hosts, the highest endophytic colonization rate occurred in Artemisia capillaris, which exhibited highest fungal diversity. Five fungal isolates belonging to Aureobasidium pullulans, Ephelis, Pestalotiopsis, and Pleosporaceae, were only recovered from Artemisia capillaris. Xylaria species was reported to be dominant endophytic fungi in Artemisia indica. Seven Artemisia species were sampled in two locations (Qichun and Wuhan in China) and 21 fungal endophytic species belonging to: Diaporthe, Colletotrichum, Nigrospora, Botryosphaeria, Aspergillus, Penicillium, Neofusicoccum, Cercospora, Rhizoctonia, Alternaria, and Curvularia were found [23]. The highest incidences of colonization frequency per plant host revealed Nigrospora sphaerica in Artemisia sp., Nigrospora oryzae in Artemisia argyi, Alternaria alternata in Artemisia subulata and Artemisia tangutica and Botryosphaeria dothidea in Artemisia lavandulifolia. The authors report for the first time Nigrospora, Neofusicoccum and Curvularia species in Artemisia spp.

Artemisia thuscula is an endemic plant of Canary Islands and community of endophytes housed inside its plant tissues remains unexplored. With the idea of exploring endemic medicinal plants for useful and underexplored fungal endophytes, we strategically pinned down to Artemisia thuscula that has been harboring in western areas of islands i.e., Tenerife and La Palma, for ages. Elements of phylogeny and diversity were framed for the strains obtained from both islands with a case study of Tenerife where diversity was intended to be enhanced by using different nutrient media and stem ages. Questions on host specificity were explored, having one plant species and various collection locations.

2. Materials and Methods

2.1. Plants Sampling

Plants of Artemisia thuscula species were collected from Canary Islands (La Palma and Tenerife). 10 plants specimens were sampled in total. Three plants were sampled from La Palma and seven plants were sampled from Tenerife; GPS coordinates are mentioned in Table 1. In situ, plants were observed for their healthy appearance prior to the sampling, only those individuals that did not show symptoms of attack by pest or disease were selected. From each plant only stems segments were cut, labeled and kept in paper bags inside zip-locked bags at T = 4–5 °C until transported to the laboratory and then processed within 24 h. Identification of the plant species was performed using classical morphological examination. The plants were deposited at the University of La Laguna (ULL) herbarium (TFC).

2.2. Fungal Endophyte Isolation

Surface sterilization method was used to suppress epiphytic microorganisms from the plant [23]. Thus, stem fragments were first washed with sterile water, then immersed in 70% ethanol for 1 min, followed by an immersion in 15% sodium hypochlorite for 1 min, again in 70% ethanol for 1 min and lastly were washed with sterile distilled water. To assure a successful sterilization, fragments were rolled on potato dextrose agar (PDA) medium and drops of last step sterilization water were poured on medium, as a control check for complete sterilization. After this process, plant material was dried on sterile blotting sheet, excised in pieces of 2 cm and cut longitudinally with a sterile scalpel. Segments were placed in PDA (Sigma-Aldrich, St. Louis, MI, USA) Petri plates amended with tetracycline (10 mg L−1). Plates incubated with the plant segments were incubated at 25 °C in the dark for two weeks and observed daily for fungal growth. When fungal outgrowth from the plant tissues occurred observations on emerged fungi were made. Only the fungi with different morphological characteristics were subcultured. Eventually, when an endophyte was acquired in pure culture it was preserved in Czapek medium (Fluka Analytical, Sigma-Aldrich), T = 5 °C and in glycerol (≥99.5, Sigma-Aldrich) 20% in deionized H2O, T = −32 °C and identified. To analyze the fungal diversity, each replicate of the distinct stem fragments was noted. To enhance bioprospection and diversity, variable nutritive media were utilized to incubate stem fragments (with ages less than one year and more than one year) of eight plants from Tenerife. Therefore, V8 tomato juice agar and lignocellulose agar (LCA) [29] media were additionally used. All the reagents were purchased from Sigma-Aldrich, except Agar Agar—GUINAMA (Valencia, Spain) and Potassium chloride—PanReac AppliChem (Barcelona, Spain).

2.3. Fungal Endophyte Collection and Maintenance

Every isolate and its plant origin were dully recorded for calculation of colonization rate from host, counting the same isolate identification only once if it emerges from the same plant segment. After purification of each isolate, it was subjected to microscopical observations followed by molecular analysis to identify at genus and/or species level. Isolates are presently maintained in three types of media: Czapek, T = 5 °C; mineral oil (Sigma-Aldrich), T = 5 °C and glycerol (Sigma-Aldrich) 20% diH2O, T = −80 °C. For short term use, fungal isolates were maintained on PDA, 25 °C.

2.4. Morphological Identification

Prior to taxonomic identification, a preliminary classification was made to avoid the selection of identical strains arising from the same plant individual, separating isolates into morphotypes. Observations targeted characteristics related to the colony and medium as: colony shape, texture and colour; exudates, medium colour and growth rate. For the microscopic observations, a strain was inoculated onto a PDA Petri plate and a sterile cover slide was attached at two centimeters. Once the growth of the fungus partially covered the cover slide, the slide was removed, inverted on a slide with cotton blue (for the slightly coloured colonies) and observed under microscope.

2.5. Molecular Identification

Out of several procedures for genomic DNA extraction, the most efficient protocol, although time consuming, was the one described by Shu et al., 2014 [20] to which the following modifications were made. Samples were centrifuged for 15 min at 12,000 rpm; after the chloroform (≥99.5, Sigma-Aldrich) procedure the upper phase was mixed with 10% Sodium acetate (ReagentPlus®, ≥99.0%, Sigma-Aldrich) and 60% Isopropyl alcohol (Aldrich ≥ 97.0%, Sigma-Aldrich), incubated for 10 min at −30 °C and centrifuged (10 min, 12,000 rpm). Finally, the pellet was washed twice with 75% ethanol (before maintained at −20 °C) and centrifuged (10 min, 12,000 rpm). The solvent was removed by evaporation, keeping the sample in the laminar flow cabinet. The purified DNA was suspended in 20 µL TE buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA); all reagents were purchased from Sigma-Aldrich. RNase A was added, and the sample was incubated for 1 hour at room temperature (long-term storage at −32 °C).

The second protocol for DNA extraction involves no purification of DNA but acceptable results were garnered (around 50% samples succeeded). 20 µL of TE buffer was pipetted into a microtube and glass beads (diameter = 0.4–0.6 mm) were added to make up 3/4 of the reagent’s volume. A small quantity of fungal mycelium was added (2–5 mm/2–3 mg) with a needle. Samples were homogenized using FastPrep 24™ 5 G (MP Bio, Santa Ana, California, USA) at 4 m/s, 20 s. Subsequently samples were centrifuged at 13,000 rpm for 1 min and maintained on ice. One µL of the supernatant was used for the PCRs.

The third and fourth protocol involved two genomic DNA extraction kits. First one used was E.Z.N.A. Fungal DNA Kit according to the manufacturer indications (OMEGA bio-tek, Norcross, Georgia, USA) with overall good results (around 80% of the samples succeeded). The second one tested was Fungi/Yeast Genomic DNA Isolation Kit, according to the manufacturer indications (NORGEN Biotek, Thorold, ON, Canada) with overall good results also (approximately 70% of the samples succeeded).

The fourth protocol approaches nucleic acid extraction by application of silica coupled to magnetic particles, which is efficient and automated. Genomic fungal DNA was extracted using Maxwell 16 Mouse Tail DNA purification kit. The Promega kit is designed for automated DNA extraction from tissue samples using the Maxwell™ 16 platform (Promega BioSciences, San Luis Obispo, CA, USA). This protocol was performed at the University Institute of Tropical Diseases and Public Health of the Canary Islands, University of La Laguna.

Molecular identification of the fungal Dicarya strains was performed using ITS1 (5′-TCCGTAGGTGAACCTGCGG-3′) and ITS4 (5′-TCCTCCGCTTATTGATATGC-3′) primer pair to amplify the 5.8S rDNA and the two internal transcribed spacers ITS1 and ITS2 [30] for the majority of the samples and NL-1 (5′-GCA TAT CAA TAA GCG GAG GAA AAG-3′) and NL-4 (5′-GGT CCG TGT TTC AAG ACG G-3′) primer pair to amplify the 5′ end of 28S rDNA spanning domains D1 and D2) [31]. PCRs were performed in a total volume of 25 µL containing 10 ng genomic DNA, 0.5 µM primer, 200 µM dNTPs, 1X Buffer Taq, 0.0125U of Taq DNA Polymerase. For ITS sequences, PCR cycling parameters were carried out according to Shu et al. 2014 [20] with slight modifications: 94 °C for 2.5 min; 40 cycles of 94 °C for 30 s, 58 °C for 30 s, and 72 °C for 1 min; and a final extension at 72 °C for 10 min. For 28S rDNA domain, the PCR conditions were denaturation for 4 min at 95 °C followed by 45 s at 95 °C and then annealing for 45 s at 58 °C, 1 min at 72 °C, followed by an extension at 72 °C for 5 min. The final step was at 16 °C for 5 min. A total of 40 cycles were performed. All PCR products were detected by agarose gel electrophoresis (110V, 35 min, on 2% agarose gels, 1X TAE Buffer) loading 5 µL PCR product, 1 µL Loading Buffer (6X) and 2 µL SYBR Green I (Sigma-Aldrich; dilution 1:10,000). PCR and electrophoresis reagents were purchased from Sigma-Aldrich. PCR products were purified using GenElute™ PCR Clean-Up Kit (Sigma-Aldrich) and sequenced by Sequencing Services SEGAI (La Laguna, Spain). The sequences were run through the BLASTN search page using Megablast program (National Center for Biotechnology Information; Bethesda MD, USA) where the most identical hits and their accession numbers were obtained. Further, only ITS sequences were used for the phylogenetic analysis, therefore details on 28S sequenced strains are listed in Table 2.

2.6. Phylogenetic Analysis

ITS sequences [i.e., endophytic fungi—Table 3, their most similar hits from GenBank (NCBI, Bethesda MD, USA) and type sequences of the selected taxa] were aligned with the multiple alignment program ClustalW [32] as implemented in Mega 6.0 (Molecular Evolutionary Genetics Analysis) [33] and indels corrected manually to minimize alignment gaps [34]. Designated outgroup was Caloscypha fulgens (GenBank Accession No. DQ491483). After the exclusion of non-overlapping leading/trailing gaps the length of the alignment was 603 bps. Because of the high number of indels, these were recoded as a binary matrix by means of the simple indel coding algorithm [35], appending the fragments to the nucleotide data as additional characters, as implemented in FastGap 1.21 (Department of Biosciences, Aarhus University, Denmark) [36]. This “indel matrix” was used in all Bayesian and maximum likelihood analyses. Formerly, Gblocks program (hosted at was used to eliminate poorly aligned positions and divergent regions [37]. Best-fit models were compared in jModelTest 2 according to Bayesian Information Criterion (BIC) [38]. Best fit according to the BIC criterion model (K80 + G) was selected to reconstruct the Bayesian tree. Bayesian Inference analysis was conducted with MrBayes 3.2.3 (hosted by Mobyle SNAP Workbench, North Carolina State University) [39] and run for 1 × 107 generations with a sampling frequency of 100 generations. Of the resulting trees, the first 25,000 trees were discarded as burn-in and the following 75,001 were used to estimate topology and tree parameters. The percentage number of times a node occurred within these 75,001 was interpreted as the posterior probability of the node [40]. Convergence of the runs was indicated by an average standard deviation of split frequencies between duplicate runs of less than 0.01. The consensus trees were drawn using Treegraph 2 software (Institute for Evolution and Biodiversity, University of Munster, Germany) [41] and edited with Adobe Illustrator CS3 (Adobe Systems Incorporated, San Jose, CA, USA).

2.7. Diversity Analysis

The colonization rate (CR%) was calculated as the total number of stem fragments in a sample (plant/nutritive medium) yielding at least one isolate divided by the total number of stem fragments in that sample. Colonization frequency (CF%) was calculated as the total number of fragments in a sample (plant/location) colonized by a species divided by the total number of fragments plated. For the diversity of endophytic fungi, the Margalef index, Brillouin index, Fisher’s alpha index and Simpson’s dominance index were used. Margalef index [42] measures species richness while Brillouin index combines richness and evenness. The Margalef index was calculated using formula , where S is the number of species and N is the number of individuals in the sample. The Brillouin index [43,44] was calculated using formula: , where N is the total number of individuals, S is the number of taxa and ni is the number of individuals belonging to i species. Fisher’s logarithmic series model [45] is a species-abundant model and describes the relationship between the number of species and the number of individuals of those species. It was calculated using formula , where S is number of taxa, n


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