Skip to main content
Journal of Fungi logoLink to Journal of Fungi
. 2021 Nov 4;7(11):939. doi: 10.3390/jof7110939

Advances in the Role of Dark Septate Endophytes in the Plant Resistance to Abiotic and Biotic Stresses

Mila Santos 1,*, Ignacio Cesanelli 1, Fernando Diánez 1, Brenda Sánchez-Montesinos 1, Alejandro Moreno-Gavíra 1
Editors: Nuria Ferrol1, Concepción Azcón Aguilar1
PMCID: PMC8622582  PMID: 34829226

Abstract

Endophytic fungi have been studied in recent decades to understand how they interact with their hosts, the types of relationships they establish, and the potential effects of this interaction. Dark septate endophytes (DSE) are isolated from healthy plants and form melanised structures in the roots, including inter- and intracellular hyphae and microsclerotia, causing low host specificity and covering a wide geographic range. Many studies have revealed beneficial relationships between DSE and their hosts, such as enhanced plant growth, nutrient uptake, and resistance to biotic and abiotic stress. Furthermore, in recent decades, studies have revealed the ability of DSE to mitigate the negative effects of crop diseases, thereby highlighting DSE as potential biocontrol agents of plant diseases (BCAs). Given the importance of these fungi in nature, this article is a review of the role of DSE as BCAs. The findings of increasing numbers of studies on these fungi and their relationships with their plant hosts are also discussed to enable their use as a tool for the integrated management of crop diseases and pests.

Keywords: DSE, endophyte fungi, biological control, abiotic, biotic, stress

1. Introduction

Crop production has always been threatened by biotic and abiotic stress. As a result, different tools have been used, from chemical pesticides to genetically modified organisms (GMO), grafts, and physical barriers, to alleviate their impact. As consumers continue to demand products with no agrochemical residues, such as ecological and organic products, some of the traditional methods of control cannot be employed. Biological pest control is thus a current challenge and microorganisms serve as one of the most important tools to enable this control.

Studies on endophytic fungi have been carried out in the last decades to understand how they interact with their hosts, the types of relationships they establish, and the potential effects of this interaction. The main characteristic of endophyte fungi is that they reside and grow within plant tissues, and sporulate during host senescence [1,2,3].

Two major groups of endophytic fungi were previously identified [4]: the clavicipitaceous endophytes (C-endophytes), also known as class 1 endophytes; and the non-clavicipitaceous endophytes (NC-endophytes), which are divided into class 2, 3, and 4 endophytes. C-endophytes are employed to infect grasses, while NC-endophytes are found in asymptomatic tissues of non-vascular plants, ferns, allies, conifers, and angiosperms [4]. Unlike C-endophytes, NC-endophytes have been found in highly diverse types of ecosystems [5].

As mentioned above, NC-endophytes can be divided into three classes, and exist in several hosts. The first difference among NC-endophytes is that class 3 endophytes present highly localised infections, while class 2 and 4 endophytes can extensively colonise host tissues. Another difference among the classes is the host tissue that they can colonise. Class 2 can colonise the root, shoot, and rhizome; class 3 can only colonise the shoot; and class 4 colonises only the root of their host [4]. In this review, we will focus on dark septate endophytes (DSE), belonging to the class 4 endophytes. DSE are conidial or sterile septate fungal endophytes, usually isolated from healthy plants, that form melanised structures including inter- and intracellular hyphae and microsclerotia in the roots [4]. DSEs do not present any host specificity as they have been associated with 600 plant species, 320 genera, and 114 families [6,7,8], and can live in a facultative manner on different types of organic compounds [9]. However, certain groups of species show certain preferences regarding attachment to their host [7]. DSE exist over a large geographical range [6,8], from South African coastal plains to tropical, temperate, subalpine, alpine, and maritime Antarctic and Arctic zones [6,10,11]. These ecosystems are usually associated with stressful abiotic conditions, such as Arctic or high-altitude ecosystems [10,11], arid or semiarid regions [12,13], drought, high salt content, low fertility, high CO2 concentration or even heavy metal-contaminated soils [14,15,16,17,18,19,20,21]. The presence of DSE in these types of conditions implies that the DSE-host interaction is crucial to plant survival [19]. This ability can be attributed to the development of melanised hyphae and microsclerotia production [22,23]. The general hypothesis suggested by many studies is that the high concentration of this pigment protects plants from free radicals or heavy metal ions owing to its antioxidant effect [24,25]. However, Graber et al. [26] revealed that salt stress tolerance is independent of melanin accumulation. In other types of studies, melanin content was not found to affect processes such as Agrobacterium tumefaciens-mediated transformation efficiency [27]. The colonization of hosts by DSE follows a similar pattern and begins with the appearance of a certain number of superficial hyphae [28]. The hyphae penetrate spaces between adjacent epidermal cells, continuing to the main axis of the root, and passing through the intercellular spaces of the cortical cells [6,29]. Once the hyphae are inside the root, they form masses of packed pigmented fungal cells within the cortical cells, resulting in microsclerotia development [28]. DSE have been characterised in many studies; however, many of the described species do not have a common phylogenetic group [30,31]. Some DSE species have been included in three groups of Heliotiales, while others have been included in Pleosporales, Sordariales or Pezizales. Most DSE belong to the genera Cadophora, Microdochium, Trichocladium and Phialocephala [32]. The Phialocephala genus includes species closely related phylogenetically, are indistinguishable, and are grouped in the complex named Phialocephala fortinii s.l. (sensu lato)–Acephala applanata species complex (PAC). The most frequent DSE in natural forest ecosystems in the Northern hemisphere belong to the PAC [31,33,34,35,36]. Phialocephala fortinii was first described in association with Pinus sylvestris by Wang and Wilcox [37]. Since then, Phialocephala fortinii has been found in many different plant species roots with numerous benefits for the host. Another genus of interest that has been extensively studied is Exophiala, which was investigated by Addy et al. [30,34]. This genus is reported to be an endophyte, and can be found in soil, water, and different hosts [38,39,40,41,42]. Recently, different species of Exophiala were identified as parasites of the eggs of nematodes of species such as Heterodera schachtii [43] or Tylenchulus semipenetrans [44]. The genus Cadophora includes pathogenic plant species associated with roots or wood colonisers [45]. Wang and Wilcox et al. [37] isolated the Cadophora finlandica strain from Pinus sylvestris roots. To date, many other species have been characterised; however, their roles in the interactions with their hosts are mainly unknown [46,47]. Beneficial associations between DSE and plants include nutrient uptake [48,49,50,51,52,53,54], growth promoting effects [54,55,56,57,58,59,60,61,62,63], tolerance to abiotic stress, such as heavy metals [64,65,66,67,68], and tolerance to drought [13,19,21,22,57,58,61,69,70,71,72,73,74,75], salinity [27,76,77], biotic stress [78,79,80,81], and changes in the rhizosphere microbiome [73,81,82], thereby positioning DSE as potential tools for crop production and achieving biological control. According to the map shown in Figure 1, there are few species of DSE responsible for the biological control of diseases.

Figure 1.

Figure 1

Network map of the co-occurrence matrix for the 547 documents published in DSE research. VOSviewer software (version 1.6.15, Leiden University, The Netherlands) was used to map the frequency of keyword co-occurrence networks. Differences in font size imply differences in relevance. The different colours refer to the groups or clusters formed.

2. Nutrient Uptake and Plant Growth Promotion

Nutrient uptake and plant growth are two parameters that are positively correlated. Most studies on DSE–plant interactions could not reveal positive or negative effects [83]; however, some DSE can increase nutritional availability for plants, enabling generally higher growth rates in plants. The effect of the interaction between DSE and the host plant apparently depends on different factors, such as plant and fungal genotype as well as soil fertility [84,85,86,87,88,89]. The enhancement of nutrient absorption by plants has been revealed by numerous researchers. In fact, Xu et al. [52] revealed that the inoculation of maize plants with E. Pisciphila H93 enhanced phosphorus absorption by the host plant. This DSE–plant interaction seemed to occur independently of the DSE-induced genes involved in phosphorus absorption [52]. In addition, Vergara et al. [50] reported that DSE stimulated nutrient uptake in rice plants, increasing the accumulation of N, P, K, Mg, Fe, Ca, and Zn in the aerial parts of rice plants. According to He et al. [69], inoculation with DSE increased the content of N and P in plants but decreased the content of organic material and P in the rhizosphere [14,57]. Similarly, Haselwandter and Read [90] reported increased P content in Carex plants associated with different species of DSE. Recently, Wu et al. [54] revealed increased expression levels of genes related to metabolic processes and genetic information processes. Moreover, responses to environmental signals were found to be enriched under DSE colonization.

The ability of DSE to colonise and have beneficial effects on plants can also be dependent on the nutrient’s nature in the soil or substrate [89,91,92]. For instance, Yakti et al. [53] identified different biostimulant effects exerted by Periconia macrospinosa and Cadophora sp., increasing the root and shoot biomass of tomato plants when organic and inorganic nutrient sources were supplied. When organic nutrient sources were applied, only Periconia macrospinosa increased shoot and root biomass in tomato plants. However, both DSEs promoted shoot growth when cultivated with inorganic fertilisers.

Mutualistic symbiosis relationships between DSE and crops have been investigated. Such relationships promote nutrient uptake and plant growth. Andrade-Linares et al. [62] reported the biostimulant effect of three DSEs (DSE 48, DSE49 and Leptodontidium orchidicola) on tomato plants; however, the effects were only observed in young plants, with none of the DSE strains affecting plants after 22 weeks of cultivation. On the other hand, Usuki et al. [89] revealed that Heteroconium chaetospira forms a mutualistic symbiosis with Chinese cabbage, facilitating the supply of nitrogen in exchange for carbohydrates, resulting in increased plant growth.

Interactions between DSE and other microorganisms have been revealed, mainly with arbuscular mycorrhizal fungi (AMF). In many cases, the interactions may involve a stimulation of DSE fungal development [81] or a reduction in the possible pathogenic effect of DSE on the plant [93,94]. Li et al. [95] assayed the co-occurrence of both AMF and DSE (P. fortinii) in epidermal cells of hair roots. Based on their results, the use of Finnish peat moss as a symbiotic fungi inoculum could enable the establishment of symbiotic fungal colonization and promote rooting and vegetative growth in rabbiteye blueberry cuttings. In addition, the observed positive effect was greater when the colonisation rates of DSE and AMF were considerably higher. Recently, Guo et al. [96] revealed that DSE and mycorrhizal infection rate significantly improved the bacterial diversity and fruit yield of blueberry rhizosphere soil. P. fortinii has been reported to be a plant growth promoter in Asparagus officinalis, Brassica oleracea, and Spinacia oleracea [6,59,97,98,99,100].

Other processes also promote plant growth, such as the production of different secondary metabolites, volatile organic compounds, and phytohormones by the hyphae of DSE [101,102].

3. Abiotic Stress

According to numerous studies, DSE can contribute to the capacity of plants to tolerate abiotic stress, such as salinity, drought, and heavy metal contamination. Different mechanisms have been recognised to be responsible for such protection, such as melanin content in DSE; however, these mechanisms are still unclear, as they often depend on different DSE–host interactions.

Farias et al. [103] revealed that the inoculation of DSE, Sordariomycetes sp-B’2 and Melanconiella elegans-21W2, in cowpea plants induced tolerance to salt stress. The inoculated plants had higher values for leaf area and shoot and root dry mass than control plants at the same level of water salinity. N and P leaf content, photosynthesis and stomatal conductance were also higher in the inoculated plants. Root colonization by Piriformospora indica was found to increase plant growth and attenuate NaCl-induced lipid peroxidation, metabolic heat efflux, and fatty acid desaturation in leaves of the salt-sensitive barley [104]. P. indica significantly elevated the amount of ascorbic acid and increased the activities of antioxidant enzymes in barley roots under salt stress conditions. Other authors revealed that P. indica colonization promoted Arabidopsis growth under salt stress conditions, which might be caused by modulation of the expression levels of the major Na+ and K+ ion channels [105].

Hou et al. [72] revealed that the symbiotic effect of DSE on Artemisia ordosica depended on the DSE species and the salt concentration of the medium. Accordingly, Alternaria chlamydosporigena was found to promote the accumulation of total biomass and enhance superoxide dismutase (SOD) activity; Paraphoma chrysanthemicola promoted the accumulation of root biomass and increased indoleacetic acid (IAA) contents; and Bipolaris sorokiniana enhanced SOD activity and glutathione (GSH) and IAA contents, with changes in the rhizosphere microbiome depending on the salt concentration. Conversely, Gonçalves et al. [106] found that the beneficial effect of the mutualistic interaction between Salicornia and Stemphylium sp. decreased as the salt concentration increased.

Drought is another abiotic stress where the positive effect of DSE was revealed. The interaction between DSE and their hosts is apparently stronger under water stress conditions. Accordingly, the interaction between P. indica and water stress was found to activate signal oxidation during stress to prevent cell damage and assist in the maintenance of the osmotic pressure in artichoke (Cynara scolymus). Thus, higher water stress levels in the host plant might be attributed to the increased content of phenolics and flavonoids [107]. On the other hand, Li et al. [75] found that Phialophora sp. and Leptosphaeria sp. improved the root biomass, total biomass, nutrient concentration, and antioxidant enzyme activities of host plants under water deficit conditions. The same results were found in cases of colonisation by the DSE fungal group known as PAC (Phialocephala fortinii s.l.–Acephala applanata species complex) [35], which increased under water stress conditions. Landold et al. [100] suggested that the PAC could protect oak roots against desiccation and improve the drought resistance of Q. pubescens due to the more extensive formation of waterproof microsclerotia. Stroheker et al. [63] also proposed the above notion in their assessment of P. abies seedlings. Zhang et al. [108] revealed that sorghum plants inoculated with Exophiala pisciphila showed significantly higher growth and dry matter content than non-inoculated plants, alleviating the negative effects of drought. In addition, plants inoculated with Exophiala pisciphila had higher photosynthetic and leaf transpiration rates. Liu et al. [70] found that Acrocalymma vagum could establish a symbiotic relationship with Ormosia hosiei, allowing the hosts to improve their defence against drought stress, reduce water loss, and maintain normal physiological activity. However, the researchers found that inoculated plants had higher contents of chlorophyll a, chlorophyll b, and carotenoids, with increases of 13%, 28%, and 35%, respectively. Inoculation was also found to significantly enhance net photosynthetic rate, stomatal conductance, and transpiration rates. Morphology differences were also found in leaves, with inoculated plants displaying greater leaf length, area, dry weight, and thicker leaves. According to Panke-Buisse et al. [109], reduced numbers of DSE are associated with a greater drought tolerance. However, there are numerous DSE genera found in plants, such as Paraconiothyrium, Phialophora, Darksidea, Knufia, Leptosphaeria and Embellisia, that lead to drought resistance in other plant species [57,110].

The important role of DSE in heavy metal buffering and tolerance in plants is well documented. Metal resistance to Cd and Zn was described to result from DSE and plant interaction [111,112,113,114,115,116,117]. Zhu et al. [118] revealed that two strains of Phialophora mustea improved the tolerance of metal stress and promoted plant growth and dry weight in tomato plants. Similarly, P. fortinii could promote the growth and survival of Miscanthus sinensis in mine sites under Al stress [119]. This tolerance was based on the reduction in metal contents in roots and shoots and the limited translocation from roots to shoots in inoculated plants compared with non-inoculated controls. DSE stimulated the activities of antioxidant enzymes, superoxide dismutase (SOD), and peroxidase (POD), mitigating the membrane lipid peroxidation damage caused by excessive metal ions. In another study, Shen et al. [120] assessed how maize responds to Cd stress following Exophiala pisciphila inoculation and found that the DSE alleviated the toxic effect of Cd excess and promoted plant growth. Furthermore, Cd accumulation in the root cell walls was higher in inoculated plants; thus, inoculated plants showed a decrease in Cd translocation from roots to shoots. Shadmani et al. [113] determined the effect of Cd uptake and growth on barley plants inoculated with Microdochium bolleyi under three Cd concentrations. Based on their results, root and shoot growth and dry root and shoot biomass were significantly higher in inoculated plants than control plants under all Cd concentration conditions. The study also revealed that Cd concentration in shoots did not increase under higher levels of Cd in the soil due to an efficient regulation system that minimised the heavy metal concentration in the aerial parts of the plant. Similar results were obtained by Yung et al. [121] when Leptodontidium sp. and Phialophora mustea were employed to inoculate Noccaea caerulescens plants, which contributed to the phytoextraction of heavy metals in polluted soils and increased plant growth.

Ban et al. [122] sought to determine the effect of different Pb concentrations on maize growth and photosynthesis efficiency when plants were inoculated with Gaeumannomyces cylindrosporus. Based on their results, inoculated plants had higher height, basal diameter, root length and total biomass, and had more efficient photosynthesis parameters than control plants. Similar to other cases, the results revealed that inoculated plants accumulated more Pb than control plants, and this accumulation was restricted to the roots. This finding suggests that in the inoculated plants, the translocation patterns were changed. Consequently, Pb accumulation in the roots alleviated the aerial parts of the plants.

The use of DSE fungi in phytoremediation can help to minimise the effect of high TE-polluted soils on plants [123]. Previously, Berthelot et al. [124] identified different DSE species isolated from the roots of poplar trees that could produce VOCs and phytohormones, and promote the development of Betula pendula, Eucalyptus globulus and Lolium perenne. Similarly, Exserohilum pedicellatum, Ophiosphaerella sp. and Alternaria alternata displayed technological potential for application in phytoremediation processes owing to their ability to degrade oil [125]. DSE can also contribute to a reduction in the damage caused by global warming in different ecosystems, individually or in combination with other microorganisms [126].

4. Biotic Stress: Biological Control

4.1. Mechanisms of Action

Several researchers evaluated the role of DSE as biocontrol agents of plant diseases, reporting positive results via in vitro and in vivo assays. Fungal strains can establish different types of interactions, such as competition for space, nutrients (Figure 2), or antibiosis, or the development of plant resistance to phytopathogens. Secondary metabolites, such as siderophores or acid compounds, can mobilise nutrients in favour of one strain, thereby limiting the nutritional resources to the other. Secondary metabolites can also reduce growth on the other fungus, generating an antibiosis response [127]. DSE-induced resistance to plant diseases has also been described [128].

Figure 2.

Figure 2

(A). Microsclerotia formed by DSE Rutstroemia calopus. (B). Inhibitory effect of DSE R. calopus against F. solani on PDA medium. Effect of salt stress (C): PDA amended with 5 g·L−1 of NaCl; (D): PDA amended with 20 g·L−1 of NaCl) on microbial antagonism of R. calopus against Sclerotinia sclerotiorum. High salt concentrations significantly reduced the number of microsclerotia and melanin accumulation. No reduction in the antagonistic effect of DSE was observed.

Interactions between DSE-hosts and biological control mechanisms have been reported in many studies. For instance, Tellenbach et al. [78] studied the mechanism employed by Phialocephala strains to control Phytophthora species. The P. europaea strain significantly reduced Phytophthora plurivora growth compared with the other strains evaluated. During the evaluation of metabolite production, four compounds were found: sclerin, sclerolide, sclerotinin A, and sclerotinin B. Iron is an essential micronutrient for every organism. Bartholdy et al. [129] tested the capacity of Phialocephala fortinii strains to excrete siderophores into the nutrient medium for the mobilization and uptake of nutrients, limiting the resource for other fungi. In the assay, the researchers identified ferricrocin, ferrirubin, and ferrichrome C as the main types of excretions. Siderophore production is influenced by several factors. For example, the researchers measured the highest siderophore concentrations at low pH values (4–4.5), whereas the highest levels of pH tended to inhibit siderophore excretion. The initial Fe concentration also affected siderophore synthesis. Although some initial Fe concentration is required for growth, high iron concentration inhibited siderophore synthesis. Thus, optimal levels depend on each individual organism.

Su et al. [130] found that Harpophora oryzae could control Magnaporthe oryzae in rice plants by inducing the accumulation of H2O2, an important type of ROS, in the site of infection; this effect was proportional to the stage of development of the DSE. In addition, the presence of ROS in the non-infected cells adjacent to the primary infection could be observed, inducing resistance to secondary infections. When foliar infection was analysed, it was clear that unlike susceptible control, plants inoculated with H. oryzae limited the lesions of M. oryzae to localised spots, controlling the damage, and indicated that H. oryzae induced systemic resistance to the disease. Deng et al. [79] studied how Phialocephala bamuru and Rhizoctonia solani affect physiological indices in Pinus Sylvestris var. mongolica. By comparing P. bamuru and control seedlings inoculated with R. solani, the researchers found that β–1,3-Glucanase and chitinase activity, proline, and soluble protein contents in seedlings from treatment via DSE inoculation increased by 18.41%, 92.54%, 101.27%, and 30.48%, respectively, in comparison to the control. In the study, the survival rate significantly increased from 33% with control treatment to 80% with P. bamuru treatment.

Understanding the mechanisms of action of the different DSE that can control diseases, as well as the different interactions that occur between DSE, the host and other microorganisms present in the rhizosphere of the plant, is of great importance for its potential use in agriculture. It is important to unravel not only the advantages, but also the disadvantages, such as potential risk of toxicity to the environment or human health. From a commercial perspective, the high microsclerotia formation (Figure 2A) rate of DSE can lead to higher viability of the future formulation. In addition, it is important to understand whether any additional requirements are necessary to increase the benefit of DSE in the plant, or if, on the contrary, the nutrient solution and/or water requirements can be adjusted without any loss in production.

4.2. Biological Control Assays

The role of DSE in the biological control of plant diseases has been less studied than their effects as biostimulants under abiotic stress conditions. Nevertheless, several studies have reported that DSE fungi can be used as biocontrol agents for pathogenic fungi (Table 1). The beneficial effects of their application in plants include changes in root architecture, disease alleviation, and improved plant growth, among other benefits.

Table 1.

In vitro and in vivo biological control tests carried out using DSE.

DSE Phytopathogen Host References
Cadophora sp. Fusarium oxysporum f. sp. meloni Cucumis melo [131]
Verticillium dahliae In vitro/tomato [132]
Rhizoctonia solani In vitro/tomato [132]
Pythium aphanidermatum In vitro/tomato [132]
Heterobasidion annosum In vitro [127]
Cladophialophora chaetospira Fusarium oxysporum Brassica pekinensis [133]
Cladosporium cladosporioides Bursaphelenchus xylophilus Pinus tabulaeformis [134,135]
Fragaria vesca [133]
Cryptosporiopsis ericae Heterobasidion parviporum In vitro [136]
Cryptosporiopsis sp. Phytophtora pini, Heterobasidion parviporum In vitro [137]
Botrytis cinerea
Exophiala pisciphila Fusarium oxysporum Brassica pekinensis [133]
Fusarium oxysporum f. sp. fragariae Fragaria vesca [134]
Tylenchulus semipenetrans Screening/in vitro [44]
Exophiala salmonis Bursaphelenchus xylophilus Pinus tabulaeformis [134]
Exophiala sp. Fusarium oxysporum Brassica pekinensis [133]
Fusarium oxysporum f. sp. fragariae Fragaria vesca [133]
Heterodera schachti Beta vulgaris [43]
Ophiostoma ulmi Ulmus [138]
Gaeumannomyces cylindrosporus Bursaphelenchus xylophilus Pinus spp. [135]
Bursaphelenchus xylophilus Pinus tabulaeformis [134]
Harpophora oryzae Magnaporthe oryzae Rice [130]
Heteroconium chaetospira Verticillium longisporum, Plasmodiophora brassicae Brassica pekinensis [80,139]
Pseudomonas syringae pv. Macricola, A. brassicae [128]
Leptodontidium orchidicola Verticillium dahliae Tomato [62]
Leptodontidium sp Pythium intermedium, Phytophthora citricola, H. annosum In vitro [127]
Meliniomyces variabilis Verticillium longisporum Brassica pekinensis [80,140]
Paraphoma chrysanthemicola Bursaphelenchus xylophilus Pinus spp. [135]
Pinus tabulaeformis [134]
Phialocephala bamuru Rhizoctonia solani Pinus silvestris [79]
Phialocephala sphareoides Heterobasidion parviporum In vitro/Picea abies [134]
Phialocephala europaea Phytophthora citrícola In vitro [127]
Phialocephala fortinii Verticillium longisporum Brassica pekinensis [80]
Fusarium oxysporum f. sp. asparagi Asparagus officinalis [141]
PAC Heterobasidion parviporum In vitro [136]
Phialophora mustea Phytophtora citricola, Pythium intermedium H. annosum In vitro [127]
Bursaphelenchus xylophilus Pinus tabulaeformis [134]
Phialocephala subalpina Phytophthora plurivora In vitro [132]
Elongisporangium undulatum In vitro [132]
Veronaeopsis simplex Fusarium oxysporum In vitro [142]

Vascular diseases caused by Fusarium and Verticillium are controlled using different DSE species. Khastini et al. [131] studied the control of Fusarium wilt by applying Cadophora sp. to melon seedlings in petri dish and field tests. The results revealed disease alleviation of 40–65% in both types of tests, with greater hyphal colonisation in the roots when the amino acid valine was added to the medium.

Appositions and thickenings were observed in the cells of the plant cell wall. Similar results were found in a biocontrol assay of Veronaeopsis simplex against Fusarium oxysporum in Chinese cabbage [142], which resulted in 70% disease alleviation. Competition for space or infection sites between V. simplex and F. oxysporum was observed; however, no direct antagonistic activity was demonstrated in vitro. On the other hand, Surono [141] detected 100% disease alleviation caused by Fusarium oxysporum f. sp. asparagi by applying Phialocephala fortinii. Harsonowati et al. [133] reported the control of Fusarium oxysporum f. sp. fragariae by Exophiala sp., Exophiala pisciphila, and Cladophialophora chaetospira, each causing disease alleviation of 62, 85, and 90%, respectively. In addition, DSE promoted plant growth by increasing photosynthetic rates and accelerating flower initiation and fruit formation.

Andrade-Linares et al. [62] reported the control of Verticillium dahliae in tomato plant assays using DSE49 and Leptodontidium orchidicola, which reduced the severity of the disease by 30% under field conditions. Narisawa et al. [80] also evaluated the interaction between Phialocephala fortinii and Heteroconium chaetospira against Verticillium longisporum in Chinese cabbage under field conditions and found a reduction in the disease by 80% and 50%, respectively. Other important pathogens have also been studied. Berthelot et al. [127] evaluated the interaction between DSE and pathogen in dual cultures, and reported that the growth of the pathogen, Heterobasidion annosum, was inhibited by Phialophora mustea and Cadophora sp. Moreover, P. mustea was found to inhibit Phytophthora citricola. On the other hand, Berthelot et al. [127] employed DSE/DSE strains but did not find any negative effects in the interaction. Finally, Tellenbach and Sieber [143] demonstrated that Phialocephala subalpina could reduce disease intensity caused by Elongisporangium undulatum and Phytophthora plurivora in Norway spruce seedlings.

As previously discussed, the parasitism of nematode eggs and juveniles by DSE has been revealed [43,44,134,135]. Gené et al. [44] found that parasitism was related to the presence of eggs or females and to the amount of magnesium or phosphorus in the soil. Disease alleviation by DSE was related to changes in root architecture as well as changes and conservation of the microbiota present in the rhizosphere. However, to our knowledge, very few studies have examined the control of bacterial diseases by DSE [128].

DSE do not always suppress pathogens. For instance, Yakti et al. [132] studied the in vitro interaction between Cadophora sp and three pathogens, namely Verticillium dahliae, Rhizoctonia solani, and Pythium aphanidermatum. Based on their findings, although Cadophora sp. reduced the growth of pathogens, the pathogens also reduced the growth of DSE at a higher percentage. As this behaviour was not observed in plant tests, the in vitro results were not consistent with the in vivo results. Similarly, Martínez-Arias et al. [138] did not obtain good results for control of Ophiostoma ulmi in Ulmus by applying Exophiala sp. However, studies revealed the inhibition of the pathogen, but to the detriment of plant growth [144]. In general, the interactions between DSE and their hosts are not necessarily beneficial. Although DSE do not behave as a pathogen, they can lead to a reduction in plant growth [144].

According to Berthelot et al. [102], studies to date are still at an early stage and thus require more mature validation. New DSE with different functions are being characterised [145], such as the production of anticancer metabolites which, however, showed no biostimulant effect in the tested plant species [146], or biostimulants of growth and production of anticancer compounds in plants [147]. More studies are required to understand the different interactions of DSE within cultivation systems and to incorporate DSE into pest and disease management methods. Compatibility studies with fungicides commonly used in agriculture are also needed [148].

5. Conclusions

Although more information has been obtained concerning the role of DSE in alleviating biotic stress, much work remains to be done before DSE can be successfully introduced into the market. Market trends and legislation for agricultural production systems highlight the need for products that are environmentally friendly and waste-free and are produced under environmentally sustainable parameters. Consequently, workers in the agricultural sector must follow these trends and develop production models that meet these requirements. As part of this adaptation, alternatives to disease control that eliminate the use of chemical tools as much as possible should be discovered and employed. As revealed in this review, DSE can colonise and establish relationships with their hosts, resulting in benefits to plants not only in terms of disease control, but also in terms of growth, nutrient solubilisation and absorption, and tolerance to abiotic stresses, such as salinity and drought. Accordingly, we believe that these types of fungi can become a very important player in new agricultural production methods that will be presented in the short and medium term. Therefore, their behaviour within these methods must be studied. Agronomic studies are necessary to determine how the DSE present in each ecosystem can be incorporated into the current production systems of each region and crop, integrating them as tools for daily use.

Acknowledgments

The present work benefited from the input of the project TRFE-I-2019/020 and was supported by University of Almeria- Bucephalus Seed, S.L.

Author Contributions

Conceptualization, M.S. and I.C.; methodology, I.C., F.D., B.S.-M. and A.M.-G.; software, F.D. and I.C.; validation, M.S.; writing—original draft preparation, I.C. and M.S. writing—review and editing, I.C., M.S. and F.D.; supervision, M.S. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

Footnotes

Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  • 1.Sherwood M., Carroll G. Fungal succession on needles and young twigs of old-growth Douglas fir. Mycologia. 1974;66:499–506. doi: 10.2307/3758493. [DOI] [Google Scholar]
  • 2.Carroll G. Fungal endophytes in stems and leaves–from latent pathogen to mutualistic symbiont. Ecology. 1988;69:2–9. doi: 10.2307/1943154. [DOI] [Google Scholar]
  • 3.Stone J.K., Polishook J.D., White J.R.J. In: Endophytic fungi. In Biodiversity of Fungi: Inventory and Monitoring Methods. Mueller G., Bills G.F., Foster M.S., editors. Elsevier; Burlington, VT, USA: 2004. pp. 241–270. [Google Scholar]
  • 4.Rodriguez R.J., White J.F., Jr., Arnold A.E., Redman R.S. Fungal endophytes: Diversity and functional roles. N. Phytol. 2009;182:314–330. doi: 10.1111/j.1469-8137.2009.02773.x. [DOI] [PubMed] [Google Scholar]
  • 5.Arnold A.E., Lutzoni F. Diversity and host range of foliar fungal endophytes: Are tropical leaves biodiversity hotspots? Ecology. 2007;88:541–549. doi: 10.1890/05-1459. [DOI] [PubMed] [Google Scholar]
  • 6.Jumpponen A., Trappe J.M. Dark septate endophytes: A review of facultative biotrophic root-colonizing fungi. N. Phytol. 1998;140:295–310. doi: 10.1046/j.1469-8137.1998.00265.x. [DOI] [PubMed] [Google Scholar]
  • 7.Han L., Zuo Y., He X., Hou Y., Li M., Li B. Plant identity and soil variables shift the colonisation and species composition of dark septate endophytes associated with medicinal plants in a northern farmland in China. Appl. Soil. Ecol. 2021;167:104042. doi: 10.1016/j.apsoil.2021.104042. [DOI] [Google Scholar]
  • 8.Mandyam K., Jumpponen A. Seeking the elusive function of theroot-colonising dark septate endophytic fungi. Stud. Mycol. 2005;53:173–189. doi: 10.3114/sim.53.1.173. [DOI] [Google Scholar]
  • 9.Day M.J., Currah R.S. Role of selected dark septate endophyte species and other hyphomycetes as saprobes on moss gametophytes. Botany. 2011;89:349–359. doi: 10.1139/b11-023. [DOI] [Google Scholar]
  • 10.Schmidt S.K., Sobieniak-Wiseman L.C., Kageyama S.A., Halloy S.R.P., Schadt C.W. Mycorrhizal and dark-septate fungi in plant roots above 4270 meters elevation in the Andes and Rocky Mountains. Arct. Antarct. Alp. Res. 2008;40:576–583. doi: 10.1657/1523-0430(07-068)[SCHMIDT]2.0.CO;2. [DOI] [Google Scholar]
  • 11.Newsham K.K. A meta-analysis of plant responses to dark septate root endophytes. N. Phytol. 2011;190:783–793. doi: 10.1111/j.1469-8137.2010.03611.x. [DOI] [PubMed] [Google Scholar]
  • 12.Porras-Alfaro A., Herrera J., Sinsabaugh R.L., Odenbach K.J., Lowrey T., Natvig D.O. Novel root fungal consortium associated with a dominant desert grass. Appl. Environ. Microbiol. 2008;74:2805–2813. doi: 10.1128/AEM.02769-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Knapp D.G., Pintye A., Kovács G.M. The dark side is not fastidious–dark septate endophytic fungi of native and invasive plants of semiarid sandy areas. PLoS ONE. 2012;7:e32570. doi: 10.1371/journal.pone.0032570. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Alberton O., Kuyper T.W., Summerbell R.C. Dark septate root endophytic fungi increase growth of Scots pine seedlings under elevated CO2 through enhanced nitrogen use efficiency. Plant Soil. 2010;328:459–470. doi: 10.1007/s11104-009-0125-8. [DOI] [Google Scholar]
  • 15.Sonjak S., Beguiristain T., Leyval C., Regvar M. Temporal temperature gradient gel electrophoresis (TTGE) analysis of arbuscular mycorrhizal fungi associated with selected plants from saline and metal polluted environments. Plant Soil. 2009;314:25–34. doi: 10.1007/s11104-008-9702-5. [DOI] [Google Scholar]
  • 16.Regvar M., Likar M., Piltaver A., Kugonič N., Smith J.E. Fungal community structure under goat willows (Salix caprea L.) growing at metal polluted site: The potential of screening in a model phytostabilisation study. Plant Soil. 2010;330:345–356. doi: 10.1007/s11104-009-0207-7. [DOI] [Google Scholar]
  • 17.Deram A., Languereau F., Haluwyn C.V. Mycorrhizal and endophytic fungal colonization in Arrhenatherum elatius L. roots according to the soil contamination in heavy metals. Soil Sediment Contam. 2011;20:114–127. doi: 10.1080/15320383.2011.528470. [DOI] [Google Scholar]
  • 18.Zhang Y., Li T., Zhao Z.W. Colonization characteristics and composition of dark septate endophytes (DSE) in a lead and zinc slag heap in Southwest China. Soil Sediment. Contam. 2013;22:532–545. doi: 10.1080/15320383.2013.750267. [DOI] [Google Scholar]
  • 19.Knapp D.G., Kovács G.M., Zajta E., Groenewald J.Z., Crous P.W. Dark septate endophytic pleosporalean genera from semiarid areas. Persoonia. 2015;35:87–100. doi: 10.3767/003158515X687669. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Lugo M.A., Reinhart K.O., Menoyo E., Crespo E.M., Urcelay C. Plant functional traits and phylogenetic relatedness explain variation in associations with root fungal endophytes in an extreme arid environment. Mycorrhiza. 2015;25:85–95. doi: 10.1007/s00572-014-0592-5. [DOI] [PubMed] [Google Scholar]
  • 21.Dos Santos S.G., da Silva P.R.A., Garcia A.C., Zilli J.É., Berbara R.L.L. Dark septate endophyte decreases stress on rice plants. Braz. J. Microbiol. 2017;48:333–341. doi: 10.1016/j.bjm.2016.09.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Barrow J.R. Atypical morphology of dark septate fungal root endophytes of Bouteloua in arid southwestern USA rangelands. Mycorrhiza. 2003;13:239–247. doi: 10.1007/s00572-003-0222-0. [DOI] [PubMed] [Google Scholar]
  • 23.McLellan C.A., Turbyville T.J., Wijeratne E.K., Kerschen A., Vierling E., Queitsch C., Gunatilaka A.L. A rhizosphere fungus enhances Arabidopsis thermotolerance through production of an HSP90 inhibitor. Plant Physiol. 2007;145:174–182. doi: 10.1104/pp.107.101808. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Ban Y., Tang M., Chen H., Xu Z., Zhang H., Yang Y. The response of dark septate endophytes (DSE) to heavy metals in pure culture. PLoS ONE. 2012;7:e47968. doi: 10.1371/journal.pone.0047968. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Zhan F., He Y., Zu Y., Li T., Zhao Z. Characterization of melanin isolated from a dark septate endophyte (DSE), Exophiala pisciphila. World J. Microbiol. Biotechnol. 2011;27:2483–2489. doi: 10.1007/s11274-011-0712-8. [DOI] [Google Scholar]
  • 26.Gaber D.A., Berthelot C., Camehl I., Kovács G.M., Blaudez D., Franken P. Salt Stress Tolerance of Dark Septate Endophytes Is Independent of Melanin Accumulation. Front. Microbiol. 2020;11:562931. doi: 10.3389/fmicb.2020.562931. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Berthelot C., Perrin Y., Leyval C., Blaudez C. Melanization and ageing are not drawbacks for successful agro-transformation of dark septate endophytes. Fungal Biol. 2017;121:652–663. doi: 10.1016/j.funbio.2017.04.004. [DOI] [PubMed] [Google Scholar]
  • 28.Ohki T., Masuya H., Yonezawa M., Usuki F., Narisawa K., Hashiba T. Colonization process of the root endophytic fungus Heteroconium chaetospira in roots of Chinese cabbage. Mycoscience. 2002;43:191–194. doi: 10.1007/S102670200027. [DOI] [Google Scholar]
  • 29.Currah R.S., Tsuneda A., Murakami S. Morphology and ecology of Phialocephala fortinii in roots of Rhododendron brachycarpum. Can. J. Bot. 1993;71:1639–1644. doi: 10.1139/b93-199. [DOI] [Google Scholar]
  • 30.Addy H.D., Piercey M.M., Currah R.S. Microfungal endophytes in roots. Can. J. Bot. 2005;83:1–13. doi: 10.1139/b04-171. [DOI] [Google Scholar]
  • 31.Grünig C.R., Queloz V., Sieber T.N. Structure of Diversity in Dark Septate Endophytes: From Species to Genes. In: Pirttila A.M., Frank A.C., editors. Endophytes of Forest Trees: Biology and Applications, Forestry Sciences 80. Springer Science Business Media B.V.; Berlin/Heidelberg, Germany: 2011. pp. 3–30. [Google Scholar]
  • 32.Grünig C.R., Queloz V., Sieber T.N., Holdenrieder O. Dark septate endophytes (DSE) of the Phialocephala fortinii s.l.–Acephala applanata species complex in tree roots: Classification, population biology and ecology. Botany. 2008;86:1355–1369. doi: 10.1139/B08-108. [DOI] [Google Scholar]
  • 33.Stoyke G., Currah R.S. Endophytic fungi from the mycorrhizae of alpine ericoid plants. Can. J. Bot. 1991;69:347–352. doi: 10.1139/b91-047. [DOI] [Google Scholar]
  • 34.Addy H.D., Hambleton S., Currah R.S. Distribution and molecular characterization of the root endophyte Phialocephala fortinii along an environmental gradient in the boreal forest of Alberta. Mycol. Res. 2000;104:1213–1221. doi: 10.1017/S0953756200002896. [DOI] [Google Scholar]
  • 35.Stroheker S., Dubach V., Vögtli I., Sieber T.N. Investigating Host Preference of Root Endophytes of Three European Tree Species, with a Focus on Members of the Phialocephala fortinii—Acephala applanata Species Complex (PAC) J. Fungi. 2021;7:317. doi: 10.3390/jof7040317. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Brenn N., Menkis A., Grünig C.R., Sieber T.N., Holdenrieder O. Community structure of Phialocephala fortinii s. lat. in European tree nurseries, and assessment of the potential of the seedlings as dissemination vehicles. Mycol. Res. 2008;112:650–662. doi: 10.1016/j.mycres.2007.12.003. [DOI] [PubMed] [Google Scholar]
  • 37.Wang C.J.K., Wilcox H.E. New species of ectendomycorrhizal and pseudomycorrhizal fungi: Phialophora finlandia, Chloridium paucisporum, and Phialocephala fortinii. Mycologia. 1985;77:951–958. doi: 10.1080/00275514.1985.12025185. [DOI] [Google Scholar]
  • 38.Maciá-Vicente J.G., Glynou K., Piepenbring M. A new species of Exophiala associated with roots. Mycol. Prog. 2016;15:1. doi: 10.1007/s11557-016-1161-4. [DOI] [Google Scholar]
  • 39.Najafzadeh M.J., Dolatabadi S., Saradeghi Keisari M., Naseri A., Feng P., de Hoog G.S. Detection and identification of opportunistic Exophiala species using the rolling circle amplification of ribo-somal internal transcribed spacers. J. Microbiol. Methods. 2013;94:338–342. doi: 10.1016/j.mimet.2013.06.026. [DOI] [PubMed] [Google Scholar]
  • 40.Wang Q., Shen S.K., Zhang A.L., Wu C.Y., Wu F.Q., Zhang X.J., Wang Y.H. Isolation and diversity analyses of endophytic fungi from Paris polyphylla var. Yunnanensis. Zhongguo Zhong Yao Za Zhi. 2013;38:3838–3844. [PubMed] [Google Scholar]
  • 41.Piercey M. Master’s Thesis. Department of Biological Sciences, University of Alberta; Edmonton, AB, Canada: 2003. Dark Septate Root Endophytic (DSE) Fungi in Canada: Geographic Distribution and Patterns of Variation. [Google Scholar]
  • 42.Summerbell R.C. Microfungi associated with the mycorrhizal mantle of and adjacent microhabitats within the rhizosphere of black spruce in boreal Canada. Can. J. Bot. 1989;67:1085–1095. doi: 10.1139/b89-142. [DOI] [Google Scholar]
  • 43.Nuaima R.H., Ashrafi S., Maier W., Heuer H. Fungi isolated from cysts of the beet cyst nematode parasitized its eggs and counterbalanced root damages. J. Pest Sci. 2021;94:563–572. doi: 10.1007/s10340-020-01254-2. [DOI] [Google Scholar]
  • 44.Gene J., Verdejo-Lucas S., Stchigel A.M., Sorribas F.J., Guarro J. Microbial parasites associated with Tylenchulus semipenetrans in citrus orchards of Catalonia, Spain. Biocontrol Sci. Technol. 2005;15:721–731. doi: 10.1080/09583150500135941. [DOI] [Google Scholar]
  • 45.Harrington T.C., McNew D.L. Phylogenetic analysis places the Phialophora-like anamorph genus Cadophora in the Helotiales. Mycotaxon. 2003;87:141–151. [Google Scholar]
  • 46.Ellis M.B. More Dematiaceous Hyphomycetes. Commonwealth Mycological Institute; Kew, UK: 1976. [Google Scholar]
  • 47.Currah R.S., Sigler L., Hambleton S. New records and new taxa of fungi from the mycorrhizae of terrestrial orchids of Alberta Canada. Can. J. Bot. 1987;65:2473–2482. doi: 10.1139/b87-336. [DOI] [Google Scholar]
  • 48.Caldwell B.A., Jumpponen A., Trappe J.M. Utilization of major detrital substrates by dark-septate root endophytes. Mycologia. 2000;92:230–232. doi: 10.2307/3761555. [DOI] [Google Scholar]
  • 49.Della Monica I.F., Saparrat M.C.N., Godeas A.M., Scervino J.M. The co-existence between DSE and AMF symbionts affects plant P pools through P mineralization and solubilization processes. Fungal Ecol. 2015;17:10–17. doi: 10.1016/j.funeco.2015.04.004. [DOI] [Google Scholar]
  • 50.Vergara C., Araujo K.E.C., Alves L.S., de Souza S.R., Santos L.A., Santa-Catarina C., da Silva K., Pereira G.M.D., Xavier G.R., Zilli J.E. Contribution of dark septate fungi to the nutrient uptake and growth of rice plants. Braz. J. Microbiol. 2018;49:67–78. doi: 10.1016/j.bjm.2017.04.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Vergara C., Araujo K.E.C., Sperandio M.V.L., Santos L.A., Urquiaga S., Zilli J.E. Dark septate endophytic fungi increase the activity of proton pumps, efficiency of 15N recovery from ammonium sulphate, N content, and micronutrient levels in rice plants. Braz. J. Microbiol. 2019;50:825–838. doi: 10.1007/s42770-019-00092-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Xu R., Li T., Shen M., Yang Z.L., Zhao Z.W. Evidence for a Dark Septate Endophyte (Exophiala Pisciphila, H93) Enhancing Phosphorus Absorption by Maize Seedlings. Plant Soil. 2020;452:249–266. doi: 10.1007/s11104-020-04538-9. [DOI] [Google Scholar]
  • 53.Yakti W., Kovács G.M., Vági P., Franken P. Impact of dark septate endophytes on tomato growth and nutrient uptake. Plant Ecol. Divers. 2018;11:637–648. doi: 10.1080/17550874.2019.1610912. [DOI] [Google Scholar]
  • 54.Wu F.L., Qu D.H., Tian W., Wang M.Y., Chen F.Y., Li K.K., Sun Y.D., Su Y.H., Yang L.N., Su H.Y., et al. Transcriptome analysis for understanding the mechanism of dark septate endophyte S16 in promoting the growth and nitrate uptake of sweet Cherry. J. Integr. Agric. 2021;20:1819–1831. doi: 10.1016/S2095-3119(20)63355-X. [DOI] [Google Scholar]
  • 55.Haselwandter K., Read D.J. Fungal associations of roots of dominant and sub-dominant plants in high-alpine vegetation systems with special reference to mycorrhiza. Oecologia. 1980;45:57–62. doi: 10.1007/BF00346707. [DOI] [PubMed] [Google Scholar]
  • 56.Wu L.Q., Lv Y.L., Meng Z.X. The promoting role of an isolate of dark-septate fungus on its host plant Saussurea involucrata Kar et Kir. Mycorrhiza. 2010;20:127–135. doi: 10.1007/s00572-009-0268-8. [DOI] [PubMed] [Google Scholar]
  • 57.Li X., He X.L., Hou L.F., Ren Y., Wang S.J., Su F. Dark septate endophytes isolated from a xerophyte plant promote the growth of Ammopiptanthus mongolicus under drought condition. Sci. Rep. 2018;8:7896. doi: 10.1038/s41598-018-26183-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Zuo Y., Su F., He X., Li M. Colonization by dark septate endophytes improves the growth of Hedysarum scoparium under multiple inoculum levels. Symbiosis. 2020;82:201–214. doi: 10.1007/s13199-020-00713-9. [DOI] [Google Scholar]
  • 59.Surono , Narisawa K. The cellulolytic activity and symbiotic potential of dark septate endophytic fungus Phialocephala fortinii to promote non-mycorrhizal plants growth. IOP Conf. Ser. Earth Environ. Sci. 2021;648:012165. doi: 10.1088/1755-1315/648/1/012165. [DOI] [Google Scholar]
  • 60.Lin L.C., Lin C.Y., Lin W.R., Tung Y.T., Wu J.H. Effects of ericoid Mycorrhizal fungi or dark septate endophytic fungi on the secondary metabolite of Rhododendron pseudochrysanthum (R. Morii) seedlings. Appl. Ecol. Environ. Res. 2021;19:1221–1232. doi: 10.15666/aeer/1902_12211232. [DOI] [Google Scholar]
  • 61.Hesse U., Schoberlein W., Wittenmayer L.F., Orster K., Warnstorff K., Diepenbrock W., Merbach W. Effects of Neotyphodium endophytes on growth, reproduction and drought: Tress tolerance of three Lolium perenne L. genotypes. Grass Forage Sci. 2003;58:407–415. doi: 10.1111/j.1365-2494.2003.00393.x. [DOI] [Google Scholar]
  • 62.Andrade-Linares D.R., Grosch R., Restrepo S., Krumbein A., Franken P. Effects of dark septate endophytes on tomato plant performance. Mycorrhiza. 2011;21:413–422. doi: 10.1007/s00572-010-0351-1. [DOI] [PubMed] [Google Scholar]
  • 63.Stroheker S., Dubach V., Queloz V., Sieber T.N. Resilience of Phialocephala fortinii s.l. e Acephala applanata communities e effects of disturbance and strain introduction. Fungal Ecol. 2018;31:19–28. doi: 10.1016/j.funeco.2017.10.006. [DOI] [Google Scholar]
  • 64.Likar M., Regvar M. Isolates of dark septate endophytes reduce metal uptake and improve physiology of Salix caprea L. Plant Soil. 2013;370:593–604. doi: 10.1007/s11104-013-1656-6. [DOI] [Google Scholar]
  • 65.Xu R.B., Li T., Cui H.L., Wang J.L., Yu X., Ding Y.H., Wang C.J., Yang Z.L., Zhao Z.W. Diversity and characterization of Cd-tolerant dark septate endophytes (DSEs) associated with the roots of Nepal alder (Alnus nepalensis) in a metal mine tailing of southwest China. Appl. Soil Ecol. 2015;93:11–18. doi: 10.1016/j.apsoil.2015.03.013. [DOI] [Google Scholar]
  • 66.Hou L., Yu J., Zhao L., He X. Dark Septate Endophytes Improve the Growth and the Tolerance of Medicago sativa and Ammopiptanthus mongolicus under Cadmium Stress. Front. Microbiol. 2020;10:3061. doi: 10.3389/fmicb.2019.03061. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67.Yu Y., Teng Z., Mou Z., Lv Y., Li T., Chen S., Zhao D., Zhao Z. Melatonin confers heavy metal-induced tolerance by alleviating oxidative stress and reducing the heavy metal accumulation in Exophiala pisciphila, a dark septate endophyte (DSE) BMC Microbiol. 2021;21:40. doi: 10.1186/s12866-021-02098-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 68.Jin H.Q., Liu H.B., Xie Y.Y., Zhang Y.G., Xu Q.Q., Mao L.J., Li X.J., Chen J.C., Lin F.C., Zhang C.L. Effect of the dark septate endophytic fungus Acrocalymma vagum on heavy metal content in tobacco leaves. Symbiosis. 2018;74:89–95. doi: 10.1007/s13199-017-0485-4. [DOI] [Google Scholar]
  • 69.He C., Wang W., Hou J. Plant growth and soil microbial impacts of enhancing licorice with inoculating dark septate endophytes under drought stress. Front. Microbiol. 2019;10:2277. doi: 10.3389/fmicb.2019.02277. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 70.Liu Y., Wei X. Dark septate endophyte improves the drought-stress resistance of Ormosia hosiei seedlings by altering leaf morphology and photosynthetic characteristics. Plant Ecol. 2021;222:761–771. doi: 10.1007/s11258-021-01135-3. [DOI] [Google Scholar]
  • 71.Boukhris A., Laffont-Schwob I., Rabier J., Salducci M.D., El Kadri L., Tonetto A., Tatoni T., Chaieb M. Changes in mesophyll element distribution and phytometabolite contents involved in fluoride tolerance of the arid gypsum-tolerant plant species Atractylis serratuloides Sieber ex Cass. (Asteraceae) Environ. Sci. Pol. Res. 2015;22:7918–7929. doi: 10.1007/s11356-014-3957-6. [DOI] [PubMed] [Google Scholar]
  • 72.Chung Y.A., Sinsabaugh R.L., Kuske C.R., Reed S.C., Rudgers J.A. Spatial variation in edaphic characteristics is a stronger control than nitrogen inputs in regulating soil microbial effects on a desert grass. J. Arid Environ. 2017;142:59–65. doi: 10.1016/j.jaridenv.2017.03.005. [DOI] [Google Scholar]
  • 73.Li X., He X.L., Zhou Y., Hou Y.T., Zuo Y.L. Effects of dark septate endophytes on the performance of Hedysarum scoparium under water deficit stress. Front. Plant Sci. 2019;10:903. doi: 10.3389/fpls.2019.00903. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 74.He C., Zeng Q., Chen Y., Chen C., Wang W., Hou J., Li X. Colonization by dark septate endophytes improves the growth and rhizosphere soil microbiome of licorice plants under different water treatments. Appl. Soil Ecol. 2021;166:103993. doi: 10.1016/j.apsoil.2021.103993. [DOI] [Google Scholar]
  • 75.He C., Wang W., Hou J., Li X. Dark Septate Endophytes Isolated From Wild Licorice Roots Grown in the Desert Regions of Northwest China Enhance the Growth of Host Plants Under Water Deficit Stress. Front. Microbiol. 2021;12:522449. doi: 10.3389/fmicb.2021.522449. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 76.Mateu M.G., Baldwin A.H., Maul J.E., Yarwood S.A. Dark septate endophyte improves salt tolerance of native and invasive lineages of Phragmites australis. ISME J. 2020;14:1943–1954. doi: 10.1038/s41396-020-0654-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 77.Hou L., Li X., He X. Effect of dark septate endophytes on plant performance of Artemisia ordosica and associated soil microbial functional group abundance under salt stress. Appl. Soil Ecol. 2021;165:103998. doi: 10.1016/j.apsoil.2021.103998. [DOI] [Google Scholar]
  • 78.Tellenbach C., Sumarah M.W., Grunig C.R., Miller J.D. Inhibition of Phytophthora species by secondary metabolites produced by the dark septate endophyte Phialocephala europaea. Fungal Ecol. 2013;6:12–18. doi: 10.1016/j.funeco.2012.10.003. [DOI] [Google Scholar]
  • 79.Deng X., Song X., Halifu S., Yu W., Song R. Effects of Dark Septate Endophytes Strain A024 on Damping-off Biocontrol, Plant Growth, and the Rhizosphere Soil Environment of Pinus sylvestris var. mongolica Annual Seedlings. Plants. 2020;9:913. doi: 10.3390/plants9070913. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 80.Narisawa K., Usuki F., Hashiba T. Control of Verticillium yellows in Chinese cabbage by the dark septate endophytic fungus LtVB3. Phytopathology. 2004;94:412–418. doi: 10.1094/PHYTO.2004.94.5.412. [DOI] [PubMed] [Google Scholar]
  • 81.Scervino J.M., Gottlieb A., Silvani V.A., Pergola M., Fernandez L., Godeas A.M. Exudates of dark septate endophyte (DSE) modulate the development of the arbuscular mycorrhizal fungus (AMF) Gigaspora rosea. Soil Biol. Biochem. 2009;41:1753–1756. doi: 10.1016/j.soilbio.2009.04.021. [DOI] [Google Scholar]
  • 82.Xiang L.B., Gong S.J., Yang L.J., Hao J.J., Xue M.F., Zeng F.S. Biocontrol potential of endophytic fungi in medicinal plants from Wuhan Botanical Garden in China. Biol. Control. 2016;94:47–55. doi: 10.1016/j.biocontrol.2015.12.002. [DOI] [Google Scholar]
  • 83.Jumpponen A. Dark septate endophytes—are they mycorrhizal? Mycorrhiza. 2001;11:207–211. doi: 10.1007/s005720100112. [DOI] [Google Scholar]
  • 84.Schadt C.W., Mullen R.B., Schmidt S.K. Isolation and phylogenetic identifi cation of a dark-septate fungus associated with the alpine plant Ranunculus adoneus. N. Phytol. 2001;150:747–755. doi: 10.1046/j.1469-8137.2001.00132.x. [DOI] [Google Scholar]
  • 85.Kernaghan G., Patriquin G. Host associations between fungal root endophytes and boreal trees. Microb. Ecol. 2011;62:460–473. doi: 10.1007/s00248-011-9851-6. [DOI] [PubMed] [Google Scholar]
  • 86.Mandyam K., Fox C., Jumpponen A. Septate endophyte colonization and host responses of grasses and forbs native to a tallgrass prairie. Mycorrhiza. 2012;22:109–119. doi: 10.1007/s00572-011-0386-y. [DOI] [PubMed] [Google Scholar]
  • 87.Mayerhofer M.S., Kernaghan G., Harper K.A. The effects of fungal root endophytes on plant growth: A meta-analysis. Mycorrhiza. 2013;23:119–128. doi: 10.1007/s00572-012-0456-9. [DOI] [PubMed] [Google Scholar]
  • 88.Heredia-Acuña C., Alarcón A., Hernández-Cuevas L.V., Ferrera-Cerrato R., Almaraz-Suarez J.J. Diversidad, Ecología e Importancia Potencial De Los Hongos Endófitos Septados Obscuros en México. Bot. Sci. 2014;92:321–333. doi: 10.17129/botsci.114. [DOI] [Google Scholar]
  • 89.Usuki F., Narisawa K. A mutualistic symbiosis between a dark septate endophytic fungus, Heteroconium chaetospira, and a nonmycorrhizal plant, Chinese cabbage. Mycologia. 2007;99:175–184. doi: 10.1080/15572536.2007.11832577. [DOI] [PubMed] [Google Scholar]
  • 90.Haselwandter K., Read D.J. The significance of a root-fungus association in two Carex species of high-alpine plant communities. Oecologia. 1982;53:352–354. doi: 10.1007/BF00389012. [DOI] [PubMed] [Google Scholar]
  • 91.Diene O., Wang W., Narisawa K. Pseudosigmoidea ibarakiensis sp. nov., a dark septate endophytic fungus from a cedar forest in Ibaraki, Japan. Microbes Environ. 2013;28:381–387. doi: 10.1264/jsme2.ME13002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 92.Mahmoud R.S., Narisawa K. A New Fungal Endophyte, Scolecobasidium humicola, promotes tomato growth under organic nitrogen conditions. PLoS ONE. 2013;8:e78746. doi: 10.1371/journal.pone.0078746. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 93.Richard C., Fortin J.A., Fortin A. Protective effect of an ec-tomycorrhizal fungus against the root pathogen Mycelium radicis atrovirens. Can. J. For. Res. 1971;1:246–251. doi: 10.1139/x71-034. [DOI] [Google Scholar]
  • 94.Reininger V., Sieber T.N. Mycorrhiza reduces adverse effects of dark septate endophytes (DSE) on growth of conifers. PLoS ONE. 2012;7:e42865. doi: 10.1371/journal.pone.0042865. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 95.Li Y.C., Chen S.J., Li K.T. Symbiotic fungi in nature Finnish peat moss promote vegetative growth in rabbiteye blueberry cuttings. Hortic. Environ. Biotechnol. 2021;62:191–198. doi: 10.1007/s13580-020-00313-y. [DOI] [Google Scholar]
  • 96.Guo X., Wan Y., Shakeel M., Wang D., Xiao L. Effect of mycorrhizal fungi inoculation on bacterial diversity, community structure and fruit yield of blueberry. Rhizosphere. 2021;19:100360. doi: 10.1016/j.rhisph.2021.100360. [DOI] [Google Scholar]
  • 97.Jumpponen A., Mattson K.G., Trappe J.M. Mycorrhizal functioning of Phialocephala fortinii with Pinus contorta on glacier forefront soil: Interactions with soil nitrogen and organic matter. Mycorrhiza. 1998;7:261–265. doi: 10.1007/s005720050190. [DOI] [PubMed] [Google Scholar]
  • 98.Narisawa K., Kawamata H., Currah R.S., Hashiba T. Suppression of Verticillium wilt in eggplant by some fungal root endophytes. Eur. J. Plant Pathol. 2002;108:103–109. doi: 10.1023/A:1015080311041. [DOI] [Google Scholar]
  • 99.Vohník M., Albrechtova J., Vosatka M. The inoculation with Oidiodendron maius and Phialocephala fortinii alters phosphorus and nitrogen uptake, foliar C:N ratio and root biomass distribution in Rhododendron cv. Azurro. Symbiosis. 2005;40:87–96. [Google Scholar]
  • 100.Landolt M., Stroheker S., Queloz V., Gall A., Sieber T.N. Does water availability influence the abundance of species of the Phialocephala fortinii s.l.-Acephala applanata complex (PAC) in roots of pubescent oak (Quercus pubescens) and Scots pine (Pinus sylvestris)? Fungal Ecol. 2020;44:100904. doi: 10.1016/j.funeco.2019.100904. [DOI] [Google Scholar]
  • 101.Khan A.L., Hamayun M., Ahmad N., Waqas M., Kang S.M., Kim Y.H., Lee I.J. Exophiala sp. LHL08 reprograms Cucumis sativus to higher growth under abiotic stresses. Physiol. Plant. 2011;143:329–343. doi: 10.1111/j.1399-3054.2011.01508.x. [DOI] [PubMed] [Google Scholar]
  • 102.Berthelot C., Chalot M., Leyval C., Blaudez D. From darkness to light: Emergence of the mysterious dark septate endophytes in plant growth promotion and stress alleviation. In: Hodkinson T.R., Murphy B.R., Saunders M.J., Doohan F.M., editors. Endophytes for a Growing World. Cambridge University Press; Cambridge, UK: 2019. pp. 143–164. [Google Scholar]
  • 103.Farias G.C., Nunes K.G., Soares M.A., de Siquiera K.A., Lima W.C., Neves A.L.R., de Lacerda C.F., Filho E.G. Dark septate endophytic fungi mitigate the effects of salt stress on cowpea plants. Braz. J. Microbiol. 2020;51:243–253. doi: 10.1007/s42770-019-00173-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 104.Baltruschat H., Fodor J., Harrach B.D., Niemczyk E., Barna B., Gullner G., Janeczko A., Kogel K.H., Schafer P., Schwarczinger I., et al. Salt tolerance of barley induced by the root endophyte Piriformospora indica is associated with a strong increase in antioxidants. N. Phytol. 2008;180:501–510. doi: 10.1111/j.1469-8137.2008.02583.x. [DOI] [PubMed] [Google Scholar]
  • 105.Abdelaziz M.E., Kim D., Ali S., Fedoroff N.V., Al-Babili S. The endophytic fungus Piriformospora indica enhances Arabidopsis thaliana growth and modulates Na+/K+ homeostasis under salt stress conditions. Plant Sci. 2017;263:107–115. doi: 10.1016/j.plantsci.2017.07.006. [DOI] [PubMed] [Google Scholar]
  • 106.Gonçalves D.R., Pena R., Zotz G.M., Albach D.C. Effects of fungal inoculation on the growth of Salicornia (Amaranthaceae) under different salinity conditions. Symbiosis. 2021;84:195–208. doi: 10.1007/s13199-021-00783-3. [DOI] [Google Scholar]
  • 107.Tanha S.R., Ghasemnezhad A., Babaeizad V.A. Study on the effect of endophyte fungus, Piriformospora indica, on the yield and phytochemical changes of globe artichoke (Cynara scolymus L.) leaves under water stress. Int. J. Adv. Biol. Biom. Res. 2014;2:1907–1921. [Google Scholar]
  • 108.Zhang Q., Gong M., Yuan J., Hou Y., Zhang H., Wang Y., Hou X. Dark Septate Endophyte Improves Drought Tolerance in Sorghum. Int. J. Agric. Biol. 2017;19:53–60. doi: 10.17957/IJAB/15.0241. [DOI] [Google Scholar]
  • 109.Panke-Buisse K., Cheng L., Gan H., Wickings K., Petrovic M., Kao-Kniffin J. Root fungal endophytes and microbial extracellular enzyme activities show patterned responses in tall fescues under drought conditions. Agronomy. 2020;10:1076. doi: 10.3390/agronomy10081076. [DOI] [Google Scholar]
  • 110.Valli P.P.S., Muthukumar T. Dark Septate Root Endophytic Fungus Nectria haematococca Improves Tomato Growth Under Water Limiting Conditions. Indian J. Microbiol. 2018;58:489–495. doi: 10.1007/s12088-018-0749-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 111.Potisek M., Likar M., Vogel-Mikuš K., Arčon I., Grdadolnik J., Regvar M. 1,8-dihydroxy naphthalene (DHN)-melanin confers tolerance to cadmium in isolates of melanised dark septate endophytes. Ecotoxicol. Environ. Saf. 2021;222:112493. doi: 10.1016/j.ecoenv.2021.112493. [DOI] [PubMed] [Google Scholar]
  • 112.Ban Y., Xiao Z., Wu C., Lv Y., Meng F., Wang J., Xu Z. The positive effects of inoculation using arbuscular mycorrhizal fungi and/or dark septate endophytes on the purification efficiency of CuO-nanoparticles-polluted wastewater in constructed wetland. J. Hazard. Mater. 2021;416:126095. doi: 10.1016/j.jhazmat.2021.126095. [DOI] [PubMed] [Google Scholar]
  • 113.Shadmani L., Jamali S., Fatemi A. Effects of root endophytic fungus, Microdochium bolleyi on cadmium uptake, translocation and tolerance by Hordeum vulgare L. Biologia. 2021;76:711–719. doi: 10.2478/s11756-020-00598-5. [DOI] [Google Scholar]
  • 114.Su Z.Z., Dai M.D., Zhu J.N., Liu X.H., Li L., Zhu X.M., Wang J.Y., Yuan Z.L., Lin F.C. Dark septate endophyte Falciphora oryzae-assisted alleviation of cadmium in rice. J. Hazard. Mater. 2021;419:126435. doi: 10.1016/j.jhazmat.2021.126435. [DOI] [PubMed] [Google Scholar]
  • 115.De Beeck M.O., Lievens B., Busschaert P., Rineau F., Smits M., Vangronsveld J., Colpaert J.V. Impact of metal pollution on fungal diversity and community structures. Environ. Microbiol. 2014;17:2035–2047. doi: 10.1111/1462-2920.12547. [DOI] [PubMed] [Google Scholar]
  • 116.Utmazian M., Schweiger P., Sommer P., Gorfer M., Strauss J., Wenzel W.W. Influence of Cadophora finlandica and other microbial treatments on cadmium and zinc uptake in willows grown on polluted soil. Plant Soil Environ. 2007;53:158–166. doi: 10.17221/2310-PSE. [DOI] [Google Scholar]
  • 117.Berthelot C., Zegeye A., Chalot M., Kovács G.M., Franken P., Leyval C., Blaudez C. Unravelling the Role of Melanin in Cd and Zn Tolerance and Accumulation of Three Dark Septate Endophytic Species. Microorganisms. 2020;8:537. doi: 10.3390/microorganisms8040537. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 118.Zhu L., Li T., Wang C., Zhang X., Xu L., Xu R., Zhao Z. The effects of dark septate endophyte (DSE) inoculation on tomato seedlings under Zn and Cd stress. Environ. Sci. Pollut. Res. 2018;25:35232–35241. doi: 10.1007/s11356-018-3456-2. [DOI] [PubMed] [Google Scholar]
  • 119.Haruma T., Yamaji K., Masuya H. Phialocephala fortinii increases aluminum tolerance in Miscanthus sinensis growing in acidic mine soil. Lett. Appl. Microbiol. 2021;73:300–307. doi: 10.1111/lam.13514. [DOI] [PubMed] [Google Scholar]
  • 120.Shen M., Schneiderb H., Xua R., Caoc G., Zhanga H., Lia T., Zhaoa Z. Dark septate endophyte enhances maize cadmium (Cd) tolerance by the remodeled host cell walls and the altered Cd subcellular distribution. Environ. Exp. Bot. 2020;172:104000. doi: 10.1016/j.envexpbot.2020.104000. [DOI] [Google Scholar]
  • 121.Yung L., Blaudez D., Maurice N., Azou-Barré A., Sirguey C. Dark septate endophytes isolated from non-hyperaccumulator plants can increase phytoextraction of Cd and Zn by the hyperaccumulator Noccaea caerulescens. Environ. Sci. Pollut. Res. 2021;28:16544–16557. doi: 10.1007/s11356-020-11793-x. [DOI] [PubMed] [Google Scholar]
  • 122.Ban Y., Xu Z., Yang Y., Zhang H., Chen H., Tang M. Effect of Dark Septate Endophytic Fungus Gaeumannomyces cylindrosporus on Plant Growth, Photosynthesis and Pb Tolerance of Maize (Zea mays L.) Pedosphere. 2017;27:283–292. doi: 10.1016/S1002-0160(17)60316-3. [DOI] [Google Scholar]
  • 123.Berthelot C., Blaudez D., Beguiristain T., Chalot M., Leyval C. Co-inoculation of Lolium perenne with Funneliformis mosseae and the dark septate endophyte Cadophora sp. in a trace element-polluted soil. Mycorrhiza. 2018;28:301–314. doi: 10.1007/s00572-018-0826-z. [DOI] [PubMed] [Google Scholar]
  • 124.Berthelot C., Leyval C., Foulon J., Chalot M., Blaudez D. Plant growth promotion, metabolite production and metal tolerance of dark septate endophytes isolated from metal-polluted poplar phytomanagement sites. FEMS Microbiol. Ecol. 2016;92:fiw144. doi: 10.1093/femsec/fiw144. [DOI] [PubMed] [Google Scholar]
  • 125.Suelgaray F.J.U., Beltramo D.M.A., Lavado R.S., Chiocchio V.M. Dark septate endophytes (DSE): Potential bioremedial promoters of oil derivatives. Int. J. Phytoremediation. 2021;23:1–8. doi: 10.1080/15226514.2021.1932733. [DOI] [PubMed] [Google Scholar]
  • 126.Gehring C., Sevanto S., Patterson A., Ulrich D.E.M., Kuske C.R. Ectomycorrhizal and Dark Septate Fungal Associations of Pinyon Pine Are Differentially Affected by Experimental Drought and Warming. Front. Plant Sci. 2020;11:582574. doi: 10.3389/fpls.2020.582574. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 127.Berthelot C., Leyva C., Chalot M., Blaudez D. Interactions between dark septate endophytes, ectomycorrhizal fungi, and root pathogens in vitro. FEMS Microbiol. Lett. 2019;366:fnz158. doi: 10.1093/femsle/fnz158. [DOI] [PubMed] [Google Scholar]
  • 128.Morita S., Azuma M., Aoba T., Satou H., Narisawa K., Hashiba T. Induced systemic resistance of Chinese cabbage to bacterial leaf spot and Alternaria leaf spot by the root endophytic fungus, Heteroconium chaetospira. J. Gen. Plant Pathol. 2003;69:71–75. doi: 10.1007/s10327-002-0005-z. [DOI] [Google Scholar]
  • 129.Bartholdy B.A., Berreck M., Haselwandter K. Hydroxamate siderophore synthesis by Phialocephala fortinii, a typical dark septate fungal root endophyte. BioMetals. 2001;14:33–42. doi: 10.1023/A:1016687021803. [DOI] [PubMed] [Google Scholar]
  • 130.Su Z., Mao L.J., Li N., Feng X.X., Yuan Z.L., Wang L.W., Lin F.C., Zhang C.L. Evidence for Biotrophic Lifestyle and Biocontrol Potential of Dark Septate Endophyte Harpophora oryzae to Rice Blast Disease. PLoS ONE. 2013;8:e61332. doi: 10.1371/journal.pone.0061332. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 131.Khastini R.O., Ogawara T., Sato Y., Narisawa K. Control of Fusarium wilt in melon by the fungal endophyte, Cadophora sp. Eur. J. Plant Pathol. 2014;139:339–348. doi: 10.1007/s10658-014-0389-6. [DOI] [Google Scholar]
  • 132.Yakti W., Kovacs G.M., Franken P. Differential interaction of the dark septate endophyte Cadophora sp. and fungal pathogens in vitro and in planta. FEMS Microbiol. Ecol. 2019;95:164. doi: 10.1093/femsec/fiz164. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 133.Harsonowati W., Marian M., Surono , Narisawa K. The Effectiveness of a Dark Septate Endophytic Fungus, Cladophialophora chaetospira SK51, to Mitigate Strawberry Fusarium Wilt Disease and With Growth Promotion Activities. Front. Microbiol. 2020;11:585. doi: 10.3389/fmicb.2020.00585. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 134.Chu H., Wang C., Li Z., Wang H., Xiao Y., Chen J., Tang M. The dark septate endophytes and ectomycorrhizal fungi effect on Pinus tabulaeformis Carr. Seedling growth and their potential effects to pine wilt disease resistance. Forests. 2019;10:140. doi: 10.3390/f10020140. [DOI] [Google Scholar]
  • 135.Chu H., Wang H., Zhang Y., Li Z., Wang C., Dai D., Tang M. Inoculation With Ectomycorrhizal Fungi and Dark Septate Endophytes Contributes to the Resistance of Pinus spp. to Pine Wilt Disease. Front. Microbiol. 2021;12:687304. doi: 10.3389/fmicb.2021.687304. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 136.Terhonen E., Kerio S., Sun H., Asiegbu F.O. Endophytic fungi of Norway spruce roots in boreal pristine mire, drained peatland and mineral soil and their inhibitory effect on Heterobasidion parviporum in vitro. Fungal Ecol. 2014;9:17–26. doi: 10.1016/j.funeco.2014.01.003. [DOI] [Google Scholar]
  • 137.Terhonen E., Sipari N., Asiegbu F.O. Inhibition of phytopathogens by fungal root endophytes of Norway spruce. Biol. Cont. 2016;99:53–63. doi: 10.1016/j.biocontrol.2016.04.006. [DOI] [Google Scholar]
  • 138.Martínez-Arias C., Plata J.S., Moncalvillo S.O., Gil L., Calcerrada J.R., Martín J.A. Endophyte inoculation enhances Ulmus minor resistance to Dutch elm disease. Fungal Ecol. 2021;50:101024. doi: 10.1016/j.funeco.2020.101024. [DOI] [Google Scholar]
  • 139.Narisawa K., Tokumasu S., Hashiba T. Suppres-sion of clubroot formation in Chinese cabbage by the rootendophytic fungus, Heteroconium chaetospira. Plant Pathol. 1998;47:210–216. doi: 10.1046/j.1365-3059.1998.00225.x. [DOI] [Google Scholar]
  • 140.Ohtaka N., Narisawa K. Molecular characterization and endophytic nature of the root-associated fungus Meliniomyces variabilis (LtVB3) J. Gen. Plant Pathol. 2008;74:24–31. doi: 10.1007/s10327-007-0046-4. [DOI] [Google Scholar]
  • 141.Surono , Narisawa K. The inhibitory role of dark septate endophytic fungus Phialocephala fortinii against Fusarium disease on the Asparagus officinalis growth in organic source conditions. Biol. Cont. 2018;120:159–167. doi: 10.1016/j.biocontrol.2018.02.017. [DOI] [Google Scholar]
  • 142.Khastini R.O., Ohta H., Narisawa K. The Role of a Dark Septate Endophytic Fungus, Veronaeopsis simplex Y34, in Fusarium Disease Suppression in Chinese Cabbage. J. Microbiol. 2012;50:618–624. doi: 10.1007/s12275-012-2105-6. [DOI] [PubMed] [Google Scholar]
  • 143.Tellenbach C., Sieber T.N. Do colonization by dark septate endophytes and elevated temperature affect pathogenicity of oomycetes? FEMS Microbiol. Ecol. 2012;82:157–168. doi: 10.1111/j.1574-6941.2012.01415.x. [DOI] [PubMed] [Google Scholar]
  • 144.Reininger V., Grünig C.R., Sieber T.N. Host species and strain combination determine growth reduction of spruce and birch seedlings colonized by root-associated dark septate endophytes. Environ. Microbiol. 2012;14:1064–1076. doi: 10.1111/j.1462-2920.2011.02686.x. [DOI] [PubMed] [Google Scholar]
  • 145.Hidayat I. Dark Septate Endophytes and Their Role in Enhancing Plant Resistance to Abiotic and Biotic Stresses. In: Sayyed R., Arora N., Reddy M., editors. Plant Growth Promoting Rhizobacteria for Sustainable Stress Management. Volume 12. Springer; Singapore: 2019. pp. 35–63. [Google Scholar]
  • 146.Berek-Nagy P.J., Tóth G., Bősze S., Horváth L.B., Darcsi A., Csikos S., Knapp G., Kovács G.M., Boldizsár I. The grass root endophytic fungus Flavomyces fulophazii: An abundant source of tetramic acid and chlorinated azaphilone derivative. Phytochemistry. 2021;190:112851. doi: 10.1016/j.phytochem.2021.112851. [DOI] [PubMed] [Google Scholar]
  • 147.Lin L.C., Tan Y.L., Lin W.R., Ku K.L., Ho S.T. The effect of dark septate endophytic fungi on Mahonia oiwakensis. Plants. 2021;10:1723. doi: 10.3390/plants10081723. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 148.Spagnoletti F.N., Chiocchio V.M. Tolerance of dark septate endophytes (DSE) to agrochemicals in vitro. Rev. Argent. Microbiol. 2020;52:43–49. doi: 10.1016/j.ram.2019.02.003. [DOI] [PubMed] [Google Scholar]

Articles from Journal of Fungi are provided here courtesy of Multidisciplinary Digital Publishing Institute (MDPI)

RESOURCES