Next Article in Journal
Responses of Soil Enzyme Activities and Microbial Community Structure and Functions to Cyclobalanopsis gilva Afforestation in Infertile Mountainous Areas of Eastern Subtropical China
Previous Article in Journal
Ecosystem Services Evaluation of Mediterranean Woodlands: A Case Study of El Pardo, Spain
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Forests
Volume 17
Issue 2
10.3390/f17020150
forests-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Pathogenicity of Diplodia fraxini and Other Botryosphaeriaceae Identified on Fraxinus excelsior with Dieback Symptoms in Poland

by
Piotr Bilański
,
Bartłomiej Grad
and
Tadeusz Kowalski
*
Department of Forest Ecosystems Protection, University of Agriculture in Krakow, Al. 29 Listopada 46, 31-425 Krakow, Poland
*
Author to whom correspondence should be addressed.
Forests 2026, 17(2), 150; https://doi.org/10.3390/f17020150
Submission received: 6 December 2025 / Revised: 13 January 2026 / Accepted: 21 January 2026 / Published: 23 January 2026
(This article belongs to the Section Forest Health)
Browse Figures
Versions Notes

Abstract

In the current work, the analysis covered 70 isolates of fungi belonging to Botryosphaeriaceae obtained in the years 2007–2017 during research on the mycobiota of F. excelsior trees with dieback symptoms in various regions of Poland. Five botryosphaeriaceous species were identified: Diplodia fraxini, D. seriata, D. sapinea, Dothiorella omnivora, and Do. sarmentorum, supported by morphological characteristics and nucleotide sequence data from three genes. The effect of temperature on the in vitro growth of colonies of five identified botryosphaeriaceous species was assessed. Dothiorella omnivora achieved optimal growth at 19.0 °C, while the other four species have shown optimal growth between 22.8 °C (Do. sarmentorum) and 25.7 °C (D. seriata). The pathogenicity test was performed in field conditions on nine-year-old F. excelsior seedlings. In total, wound inoculation was performed on 176 shoots, using 22 isolates of five identified fungal species. Each isolate was inoculated onto eight F. excelsior shoots. The symptoms on shoots were examined at 12 weeks after the inoculation. Among the tested fungal species, necrotic lesion was caused by D. fraxini, D. seriata, and Do. sarmentorum. The extent of damage they caused showed statistically significant differences. The highest pathogenic properties were demonstrated by D. fraxini, which caused necrotic lesion with a length of 34.25–50.50 mm (mean 40.13 mm) on inoculated trees. D. seriata showed the lowest degree of virulence. Half of its strains caused necrotic lesions, which did not differ significantly from the control. Diplodia sapinea and Do. omnivora did not cause any visible lesions. None of the control shoots developed necrosis. The role of Botryosphaeriaceae species in intensifying disease symptoms in ash trees in the context of Hymenoscyphus fraxineus invasion and climate changes was discussed.

1. Introduction

European ash (Fraxinus excelsior L.), an ecologically and economically important tree species, has been suffering from serious ash dieback disease since the early 1990s [1,2,3,4,5,6,7,8]. In many European countries, it is caused by an invasive alien ascomycete, Hymenoscyphus fraxineus (T. Kowalski) Baral, Queloz & Hosoya. The disease spread from east to west and is today present in most parts of the ash distribution range across Europe [3,5,9,10,11,12]. F. angustifolia Vahl is also highly susceptible [13,14], and F. ornus L. is considered resistant [15]. In many regions of Europe, apart from H. fraxineus, many other species of fungi are also found on trees with ash dieback symptoms [8,11,16,17,18,19]. Among the fungal communities on European ash, species of the Botryosphaeriaceae have a significant share [8,11,18,19,20,21]. Members of this family appear to be an increasing economic problem on a large variety of other crops [22,23,24].
Within Botryosphaeriaceae, numerous changes have occurred in recent years in terms of species identification and their taxonomy [25,26,27]. The concept of some genus within this family has also changed over the years [28]. Some species are not easy to define because of the insufficiency of distinguishing morphological features [25,28,29]. Moreover, for a long time, species in Botryosphaeriaceae were defined on the basis of host association, which unnecessarily resulted in a proliferation of species names [28]. On the other hand, some species have been found to be composites of multiple cryptic species. Thus, some species are reduced to synonyms, while in other cases, new species are described [27,30]. It was suggested that the name D. mutila (Fr.) Fr. may have been applied to more than one species [31]. Recently, from this complex, the name D. fraxini (Fr.: Fr.) Fr. was re-instated and a neotype designated [30]. D. mutila s. lato has been identified in many European countries on Fraxinus spp. with dieback symptoms [3,9,16,17,32]. After making this taxonomic clarification, D. fraxini has already been identified on F. excelsior, F. ornus, and F. angustifolia with dieback symptoms [8,9,11,18,19,20,21,33,34,35]. On F. excelsior, in addition to Diplodia mutila s. str. and D. fraxini, the following Botryosphaeriaceae species were also found: Botryosphaeria dothidea (Moug.) Ces. & De Not, D. seriata De Not., Linald., Deidda & Scanu, D. subglobosa A.J.L. Phillips, Deidda & Linald., Diplodia sp., Dothiorella omnivora Linald., Deidda & Scanu, Do. parva Abdollahz., Zare & A.J.L. Phillips, Do. sarmentorum (Fr.) A.J.L. Phillips, A. Alves & J. Luque, Do. sempervirentis Abdollahz., Zare & A.J.L. Phillips, Do. symphoricarpicola W.J. Li, Jian K. Liu & K.D. Hyde, and Neofusicoccum parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips [9,17,18,19,21,23,30,34,36,37,38,39]. In addition to association with disease symptoms, some Botryosphaeriaceae species may also occur as endophytes in ash tissue [40]. Under stressful conditions, endophytes can switch to pathogenic activity [41,42,43].
To assess the role of Botryosphaeriaceae in ash dieback syndrome, it is necessary to know the pathogenicity of individual species. Most tests in this area have been carried out for D. mutila s. lato. Its pathogenicity has been confirmed, among others, for grapevines, apples, hazelnut trees, and oaks [22,24,44]. Pathogenicity tests of such isolates against F. excelsior yielded somewhat different results. In Sweden, D. mutila s. lato was unable to induce necrosis in young ash trees [16]. In Poland, the pathogenicity of D. mutila s. lato was confirmed on 2-year-old F. excelsior saplings [45]. On most 6-year-old F. excelsior saplings, the tested isolates caused extensive necrotic lesion. However, 26.7% of stems did not develop necrosis, and inoculation wounds healed [46]. Studies conducted in southern Europe indicate that D. fraxini can cause extensive necrotic lesions in F. excelsior and F. angustifolia [19,20,21,33,35]. The pathogenicity of other Diplodia species on Fraxinus has been evaluated only sporadically so far [19,20,21].
Therefore, the aims of this study were to (i) determine the incidence of Botryosphaeriaceae species on F. excelsior in Poland by means of morphological and molecular studies, (ii) evaluate the effect of temperature on their mycelial growth in vitro, and (iii) evaluate the pathogenicity of the defined fungal species to F. excelsior in field conditions.

2. Materials and Methods

2.1. Sampling of Cultures

In the current study, the analysis covered 70 isolates of fungi belonging to Botryosphaeriaceae (Table 1). They were obtained during research on the mycobiota of young F. excelsior trees with dieback symptoms in various regions of Poland in 2007–2017. These isolates were obtained as a result of three different activities. Some of the isolates were obtained randomly from studies whose results have already been published. These works determined the frequency of occurrence of individual species of fungi [17,32,39].
Some of the isolates were obtained as a result of additional studies, during which fragments were taken from dead or living stems and shoots with visible external necrotic lesion, as well as from ash leaf petioles. In the laboratory, disease symptoms were carefully described, and isolation was performed. For this purpose, the samples were surface-sterilized by soaking first for 1 min in 96% ethanol, then for 5 min in a solution of sodium hypochlorite (approx. 4% available chlorine), and finally for 30 s in 96% ethanol. After drying in layers of blotting paper and removal of the superficial tissue, small pieces consisting of inner bark and wood tissue were cut out and placed on the surface of malt extract agar (MEA; 20 g L−1 malt extract (Difco; Sparks, MD, USA), 15 g L−1 Difco agar supplemented with 100 mg L−1 streptomycin sulphate) in Petri dishes (diameter 9 cm). The incubation took place at 20 °C in darkness. The growing fungi were transferred to 2% MEA in tubes. Then, those isolates that were classified as potential members of Botryosphaeriaceae based on previous experience were separated.
The last group of isolates was obtained from samples in which species of Botryosphaeriaceae were identified based on fruitbodies produced in vivo (Table 1). Observations were made with a Zeiss Discovery stereomicroscope (Zeiss, Göttingen, Germany) and with a Zeiss Axiophot light microscope (Zeiss, Göttingen, Germany) using differential interference contrast (DIC) illumination. Fungal microstructures were observed in distilled water on glass slides. From measurements of at least 30 conidia, the mean and standard deviation were calculated. Dimensions of other structures are given as the range of at least 15 measurements. For the purpose of identifying fungal species, appropriate mycological monographs were used [28,30,44,45]. To obtain fungal cultures from such samples, isolation was made directly from conidia. A list of all analyzed cultures with the relevant data is provided in Table 1. The tested isolates are stored in the Department of Forest Ecosystem Protection, University of Agriculture, Kraków, Poland.
To obtain additional data on the production of conidiomata by Dothiorella omnivora in vivo, three isolates of this species (352K, 353K, 355K) were inoculated at the end of August 2023 onto excised shoots (diam. 0.8–1.2 cm) of Fraxinus excelsior and Juglans regia L., which belong to the known host plants of Dothiorella omnivora [46]. Two shoots were inoculated with each isolate. The same method was used as in the pathogenicity test (Section 2.4). The bottom and top ends of the shoots were sealed with parafilm to prevent drying and contamination. The inoculated shoots were placed in a shaded area in the garden, approximately 0.3 m above the ground. Microscopic analysis of the conidiomata was performed after 10 weeks.

2.2. DNA Extraction, PCR, Sequencing, and Phylogenetic Analyses

To verify the morphological identification, nucleotide sequences of three gene fragments from representative cultures were determined: the internal transcribed spacer regions ITS1 and ITS2 including the 5.8S gene (ITS), β-tubulin 2 (TUB2), and translation elongation factor 1-α (TEF1).
Genomic DNA extraction, PCR amplification, and sequencing reactions of the isolates were performed according to the procedure described by Bilański et al. [47]. The three loci (ITS, TUB2, and TEF1) were amplified using the primers [48,49,50,51] listed in the Supplementary Materials (Table S1). The nucleotide sequences determined in this study were deposited in GenBank, and the accession numbers are listed in Table 1.
The sequences obtained from representative cultures were used to query GenBank (http://www.ncbi.nlm.nih.gov, accessed on 1 May 2025) using the MegaBLAST algorithm [52,53]. This was performed to retrieve highly similar sequences of closely related taxa for subsequent phylogenetic analysis. Only ITS and TEF1 were used in phylogenetic analyses, which made it possible to compare the results with the work of other authors [22,30,46,54,55,56,57,58].
The ITS and concatenated ITS-TEF1 sequences of Diplodia spp. were used in phylogenetic analyses, with Lasiodiplodia theobromae (Pat.) Griffon & Maubl. included as an outgroup, while the ITS and concatenated ITS-TEF1 sequences of Dothiorella spp. were analyzed using Neofusicoccum luteum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips as an outgroup.
The ITS and TEF1 gene regions were combined and analyzed as a concatenated dataset, because they showed no conflicts in the grouping of isolates into terminal clades.
The datasets were compiled and edited in BioEdit v.2.7.5 [59] and aligned using the online version of MAFFT ver. 7 [60] with the default settings. The alignments were manually checked in BioEdit v.2.7.5 [59] and compared with the reference sequences to ensure proper alignment of introns and exons.
Phylogenetic analyses were performed using three different methods: maximum likelihood (ML), maximum parsimony (MP), and Bayesian inference (BI). For each dataset, the best-fitting substitution models were determined for ML and BI using the adjusted Akaike information criterion (AICc) in jModelTest 2.1.10 [61,62]. The best models for the ITS and ITS-TEF1 data of Diplodia spp. were HKY + I and GTR + G, respectively, and for the analogous datasets for Dothiorella spp., they were HKY + I + G and GTR + I + G.
ML analysis was conducted with PhyML 3.0 [63] using 1000 bootstrap pseudoreplicates to calculate the node support values. BI analysis based on a Markov chain Monte Carlo (MCMC) method was conducted with MrBayes v3.1.2 [64]. The chains were run for 10 million generations using the best-fitted model. Trees were sampled every 100 generations, resulting in 100,000 trees from both parallel runs. The default burn-in, first 25% of samples, was used. The remaining trees were used to generate a majority rule consensus tree and to determine the posterior probability node support values. MP analysis was conducted with PAUP* 4.0b10 [65]. Gaps were treated as fifth state characters. One thousand bootstrap pseudoreplicates were generated and analyzed to determine the levels of confidence for the nodes within the inferred tree topologies. The results of the phylogenetic analyses were combined and visualized using TreeGraph 2.10.1-641 beta [66] and FigTree v1.4.0 [67]. The alignments and resulting trees were placed in the TreeBASE database (http://purl.org/phylo/treebase/phylows/study/TB2:S32419, accessed on 6 December 2025).

2.3. Effect of Temperature on Mycelial Growth

The temperature assay was performed with six isolates for each of the more abundant fungal species, D. fraxini, D. seriata, and Do. sarmentorum, three isolates of Do. omnivora, and one isolate of D. sapinea (Table 1). Mycelial plugs, 8 mm in diameter, from the edge of actively growing 7-day-old colonies were transferred onto 2% MEA in the center of a Petri dish and incubated at 5, 10, 15, 20, 25, 30, and 35 °C in the dark. Two replicated plates were prepared for each fungal isolate and temperature. Colony diameters were measured after 5 days with a millimeter ruler. The average diameter from two measurements in each replicate was calculated.

2.4. Pathogenicity Test

Pathogenicity tests were conducted on an experimental plot located in the Stary Sącz Forest District, southern Poland (49°33′49″ N, 20°39′20″ E), using Fraxinus excelsior seedlings that have reached 2.5–3.4 m in height and 1.8–3.1 cm in diameter at the stem basis. The trees grew from the seeds of ash trees of local provenance (Lipnica Forest Unit). During the first three years, they grew at a high density. After that time, they were planted in an open field at a spacing of approximately 1.0 m × 0.3 m. To determine the weather conditions prevailing during the pathogenicity test, meteorological data were obtained for the period August–October 2018 from the nearest meteorological station located in Nowy Sącz (49°37′38″ N, 20°41′19″ E, 292 m a.s.l.), approximately 7.5 km from the experimental plot. These data were used to construct the graph presented in Supplementary Figure S1. During the pathogenicity test, the minimum and maximum air temperatures were −1.0 °C and 32.2 °C, respectively. The average daily air temperature during this period was 15.4 °C, and the total precipitation amounted to 154.9 mm (Supplementary Figure S1).
Twenty-two isolates obtained from necrotic tissue on F. excelsior trees with ash dieback symptoms were used for the test: six isolates of D. fraxini, D. seriata, and Do. sarmentorum each, three isolates of Do. omnivora, and one isolate of D. sapinea (Table 1). Shoots (the previous year’s growth; diameter 5–8 mm at the inoculation site) were inoculated in the end of July 2018 on the internodes in wounds (ca. 7 mm long) made with a sterile scalpel. There were eight replicates (eight stems inoculated on eight plants) for each isolate. Inoculum production, inoculation, and re-isolation were carried out according to the methods described by Kowalski et al. [68]. In total, 176 shoots were inoculated. Control inoculations were carried out with sterile wood pieces applied to the wounds of eight shoots. The symptoms on the shoots were examined at 12 weeks after the inoculation. The lengths of superficially visible necrotic lesions were measured, and the occurrence of fungal fructification was recorded. Re-isolations were attempted from all inoculated and all control shoots within 24 h of harvesting. Three to six tissue pieces were taken from the inoculation wound, the advanced necrosis, and from the lesion edge. The isolation was considered positive if the tested fungus grew from at least one tissue sample.

2.5. Statistical Analyses

The effects of temperature on the in vitro growth of isolates from five Botryosphaeriaceae species were modeled via polynomial regression using PAST 4.17 [69], followed by the determination of local maxima. The Kruskal–Wallis test, followed by a non-parametric multiple comparison of mean ranks (Steel or Dunn’s post hoc tests; p-values were adjusted using the Benjamini–Hochberg method), was used to determine the significance of differences in lesion length and spore dimensions among the tested fungal species. The same procedure was used to determine the significance of differences in the dimensions of spores of the tested fungal species. The use of the Kruskal–Wallis and post hoc tests was necessitated by the lack of normal distribution and significant heterogeneity of variance in the data, which persisted despite transformation attempts.
The Kruskal–Wallis test and subsequent post hoc analyses were performed using R software (version 4.4.0). The ‘PMCMRplus’ package was used for Steel tests, while the ‘FSA’ package was employed for Dunn’s post hoc tests [70].
The Mann–Whitney U test was used only when comparing pairs of length or width of Diplodia fraxini conidia in relation to their origin. These statistical calculations were performed using Statistica 14 [71].
A classic cluster analysis was performed using the unweighted pair group method with arithmetic mean (UPGMA) in the PAST 4.17 program [69] to present the similarity of the colony growth patterns at different temperatures of five Botryosphaeriaceae species.

3. Results

3.1. Botryosphaeriaceae Species Identified on F. excelsior

The results of the phylogenetic analysis confirmed the species identification of the studied isolates, grouping them into separate well-defined clades corresponding to known species. The ITS sequence data were obtained for 70 isolates included in this study (Table 1). BLAST 2.17.0 and phylogenetic analyses of ITS sequences allowed 57 isolates to be placed within the genus Diplodia and 13 isolates within the genus Dothiorella (Supplementary Figures S2 and S3). The dataset containing aligned sequences of ITS and TEF1 for Diplodia spp. consisted of 551 and 308 characters, respectively. When ITS and TEF1 were combined, the resulting dataset contained 859 characters, with 278 of them being parsimony-informative (compared to 118 in the ITS dataset alone). For Dothiorella spp., the ITS and TEF1 gene regions contained 512 and 313 characters, respectively. The resulting ITS+TEF1 dataset, which gave a total of 825 characters, yielded 324 parsimony-informative sites, a significant increase compared to the 100 sites found in the ITS dataset alone.
Among the isolates obtained in the years 2007–2017 from F. excelsior in Poland, five distinct Botryosphaeriaceae species were identified based on the morphology and the concatenated sequence data for the ITS and TEF1 gene regions. These were: Diplodia fraxini, D. seriata, D. sapinea (Fr.) P. Karst., Dothiorella sarmentorum, and Do. omnivora (Figure 1 and Figure 2, Supplementary Figures S2 and S3 and Table 1). D. fraxini was the most represented, almost one third of which came from samples with developed conidiomata (Table 1). However, only three isolates of Do. omnivora and one D. sapinea isolate were found among the analyzed population (Table 1). Dothiorella sarmentorum isolates were mainly derived from necrotic shoot tissues without fruitbody. In the case of D. seriata, almost half of the isolates came from necrotic shoots with developed conidiomata (Table 1).

3.2. Morphological and Phylogenetical Aspects

The most characteristic morphological and phylogenetical features of Botryosphaeriaceae species identified on F. excelsior are given below.
Diplodia fraxini: Pycnidia were observed in Poland on ash shoots and stems and less frequently on ash petioles in the litter (Figure 3a,b and Table 1). Pycnidia were black, globose, 320 to 450 μm in diameter, solitary or in groups, immersed, and over time breaking through the bark (Figure 3a). Conidiogenous cells were hyaline, cylindrical, and 10–17 × 2.5–4.5 μm (Figure 3c). Conidia were 1-celled, thick-walled, hyaline with fine granular content, oblong to ovoid, rounded at both ends, brownish and one septate with age, and 20.0–30.0 × 8.7–14.0 μm [mean ± SD: 25.22 ± 2.02 × 12.01 ± 1.19 μm, n = 90] (Figure 3d). In humid conditions, the conidia collect as a white drop above the ostiole (Figure 3a). Colonies on MEA were woolly, initially hyaline, later dark olive–gray, and reverse olive–black. In cultures in which pycnidia were formed, white cirri and clusters with conidia were visible (Figure 3e). Conidia in vitro were hyaline, aseptate or with one septum, and turning brown with age, 20.0–30.0 × 10.0–14.0 μm [mean ± SD: 25.06 ± 2.08 × 12.27 ± 1.22 μm, n = 60] (Figure 3f). The analysis showed that D. fraxini produced conidia of similar length and width both in vivo and in vitro, and the differences found between them were not statistically significant (Supplementary Figure S4). Phylogenetic analysis (ITS and ITS+TEF1) resolved D. fraxini as a distinct highly supported monophyletic clade. This clade includes the ex-neotype sequence CBS 136010 (Figure 1 and Supplementary Figure S2).
Diplodia seriata: Pycnidia in vivo were solitary or in groups, black, globose, 220 to 400 μm in diameter, immersed, and, with time, breaking through the bark. Conidiogenous cells were hyaline, cylindrical, and 7–15 × 3–5 μm (Figure 3g). Conidia were one-celled, thick-walled, hyaline with fine granular content, oblong to ovoid, rounded at the apex, often truncate at the base (Figure 3h), becoming dark brown, and one septate with age (Figure 3h,i), 20.0–28.0 × 10.0–15.0 μm [mean ± SD: 23.12 ± 1.80 × 12.37 ± 1.16 μm, n = 60]. Conidia become brown when they are still attached to the conidiogenous cells (Figure 3g). Colonies were flocculate to woolly, initially hyaline, over time dark gray, and reverse olive–black. Pycnidia were not formed in vitro.
Isolates from this study clustered with D. seriata (including epitype CBS 112555), forming a coherent clade in all analyses. Intraspecific diversity within this clade was evident through several bootstrap-supported subgroups (Figure 1 and Supplementary Figure S2).
Diplodia sapinea: Colonies were fairly fluffy, even or with concentric zones of more abundant aerial mycelium, initially white-greyish, over time olive-blackish, and reverse hazy blackish. Hyphae were hyaline and olive-brown, smooth, 2–7.5 (10.0) μm thick with intercalary (single or in chains) chlamydospore-like cells, and 7.5–25.0 × 7.5–22.0 μm. Pycnidia were not detected on ash shoots or in cultures.
Combined ITS+TEF1 phylogeny confirmed that isolate 262K clustered with D. sapinea reference strains, including ex-epitype CBS 393.84. This clade was clearly distinct from D. scrobiculata and D. intermedia (Figure 1 and Supplementary Figure S2).
Dothiorella omnivora: Conidiomata of this species were not observed on the analyzed F. excelsior shoots (Table 1). However, after 10 weeks, on excised shoots inoculated with three isolates of Do. omnivora (Table 1), conidiomata were formed on four shoots of J. regia and two shoots of F. excelsior (out of six inoculated shoots of each tree species). On the shoots of J. regia, pycnidia were very numerous, both on the wounded part and above and below the wound, on a distance up to 3.2 cm (Figure 4a). On the shoots of F. excelsior, only a few pycnidia were produced on wounded parts (Figure 4b). Conidiomata in vivo were solitary, globose, 300–420 μm in diameter, immersed, and, with time, breaking through the bark (Figure 4a,b), with circular ostioles, and 30–50 μm in diameter. Conidiophores were absent. Conidiogenous cells were hyaline, subcylindrical, and 7–15 × 3–5 μm (Figure 4c). Conidia were initially one-celled and hyaline, starting to turn brown while still attached to conidiogenous cells (Figure 4c). Ripe conidia were dark brown, with one (Figure 4d) and occasionally two or three septa (Figure 4e), slightly constricted at the septum, oblong to ovoid, some truncate at the base, and 19.0–25.0 × 8.0–10.0 μm [mean ± SD: 21.83 ± 1.46 × 9.23 ± 0.63 μm, n = 30]. Colonies produced not high aerial mycelium, loosely woolly, initially whitish-gray, and later dark olive-gray. After 2 months, pycnidia began to form on the surface of the medium, with conidia collecting at the top in the form of brown–black drops. Conidia, as in vivo, were oblong to ovoid, brown, one septate, slightly constricted at the septum, some truncate at the base, and 19.0–22.0 × 7.5–10.0 μm [mean ± SD: 20.40 ± 0.93 × 8.50 ± 0.72 μm, n = 30]. In older cultures, after 4 months, most conidia became more swollen. They were oval, broadly elliptical with rounded ends, and quite clearly constricted at the septum (Figure 4f), with dimensions of 17.0–24.0 × 9.0–13.0 μm [mean ± SD: 20.53 ± 1.53 × 10.23 ± 0.90 μm, n = 30]. Occasionally, mainly the apical cell swelled, reaching even more than 20 μm in width (Figure 4g). The conidia of Do. omnivora reached a significantly longer length in vivo compared to in vitro (Supplementary Figure S5). The in vitro spore length was higher in older cultures (4 months) than in younger ones (2 months), but these differences were not significant (Supplementary Figure S5). Conidia of Do. omnivora became increasingly wider in vitro over time: spores in young cultures (2 months) were narrower than in old cultures (4 months). All differences in spore width found between the tested groups were statistically significant (Supplementary Figure S5).
While ITS phylogeny failed to distinguish Do. omnivora from Do. vidmadera, combined ITS+TEF1 analysis confirmed that isolates 352K, 353K, and 355K belong to Do. omnivora. These isolates formed a monophyletic clade with the ex-type CBS 140349, which was resolved as sister to Do. vidmadera, demonstrating clear taxonomic distinctness (Figure 2 and Supplementary Figure S3).
Dothiorella sarmentorum: Pycnidia were solitary, globose, 250–400 μm in diameter, immersed, and, with time, breaking through the bark (Figure 4h). Conidiogenous cells were hyaline, subcylindrical, and 6–15 × 2.5–5 μm. Conidia were elliptical to ovoid, initially hyaline, becoming brown and one-septate, slightly or not constricted at the septum, some with truncate base (Figure 4i), and 18.0–24.0 × 7.2–12.0 μm [mean ± SD: 20.77 ± 1.35 × 9.91 ± 1.13 μm, n = 60]. Occasionally, between pycnidia with macroconidia, small pycnidia filled with a mass of microconidia were developed. They formed on hyaline conidiophores with one or two short side branches in place of transverse septa, 8–18 × 2–3 μm. Microconidia were rod-shaped, hyaline, one-celled, with rounded tops, sometimes clearly truncate at the base, and 2.5–4.5 × 1.5–2.0 μm (Figure 4j). Colonies on MEA were initially hyaline, then olive-blackish from the center, fairly fluffy, with a prominent radial arrangement of hyphae, and the reverse was hazy blackish.
Neither ITS nor combined ITS+TEF1 phylogeny could distinguish Do. sarmentorum from Do. italica. Both species were interspersed within a single clade with short branch lengths, indicating high genetic similarity and minimal divergence (Figure 2 and Supplementary Figure S3).
Among the fungal species studied, D. fraxini and D. seriata produced in vivo conidia, whose length was significantly different from that of Do. sarmentorum spores. D. fraxini conidia were the longest, while Do. sarmentorum conidia were the shortest (Supplementary Figure S6). The studied species formed two groups in vivo that differed significantly in terms of the spore width. The conidia of Do. omnivora and Do. sarmentorum were significantly narrower than those of D. fraxini and D. seriata (Supplementary Figure S6).

3.3. Influence of Temperature on the Growth of Fungal Colonies In Vitro

Colonies of all the species tested showed growth on MEA in the temperature range of 5 to 30 °C, with some also at 35 °C (Figure 5). Do. omnivora, as the only species, achieved optimal growth at temperature below 20 °C (Figure 5). The remaining four species showed optimal growth between 22.8 °C (Do. sarmentorum) and 25.7 °C (D. seriata) (Figure 5). The result of the classic cluster analysis conducted on the basis of the growth rate of the tested colonies allows the distinction of two main groups of fungal species (Figure 6). The first group consisted of D. sapinea and D. seriata. Both of these species showed the highest similarity in colony growth pattern in the tested temperature range. A characteristic feature of these species was a gradual increase in the colony diameter at temperatures of 5 to 25 °C, and at higher temperatures (30 and 35 °C), there was a slow decrease in the colony growth rate (Figure 5). The second group of species consisted of Do. omnivora, D. fraxini, and Do. sarmentorum (Figure 6). These species were characterized by a sharp increase in colony diameter between temperatures of 15 and 20 °C and a sharp decrease in the colony growth rate at temperatures of 30 and 35 °C (Figure 5). The growth of colonies of species forming the second group almost completely stopped at a temperature of 35 °C (Figure 5).

3.4. Pathogenicity Assay

Evaluation of disease symptoms on F. excelsior shoots was performed 12 weeks after inoculation. Of the five fungal species tested, only three of them caused necrotic lesions: D. fraxini, D. seriata, and Do. sarmentorum. The extent of damage they caused showed statistically significant differences. The largest necroses were caused by D. fraxini and the smallest by D. seriata. Dothiorella sarmentorum caused necrotic lesions with intermediate values between D. fraxini and D. seriata (Table 2). All 48 F. excelsior shoots inoculated with D. fraxini developed necrotic lesions (Figure 7a–h, Table 2). The mean length of lesions reached from 34.25 mm (strain 986F) to 50.50 mm (987F) (Table 2). On most shoots, the edge of the necrosis was bluntly sharpened or evenly rounded (Figure 7a–c); less often, it showed an irregular course (Figure 7d,e). Necrotic areas were slightly depressed (Figure 7a–d), while for 16.7% of the shoots, the depression was distinct, accompanied by abundant callus formation from the wound edges and longitudinal cracking of the cortex (Figure 7e). All necroses on shoots resulted in discoloration of the tissue (Figure 7a–h). Necrotic tissue was uniformly dark brown (Figure 7a–c) or patchy in lighter areas (Figure 7d–f). On four shoots, light brown necrotic tissues were separated by dark gray transverse stripes, with pycnidia developing in both light and dark zones (Figure 7g). On six shoots, necrosis girdled the entire circumference of the shoot, and two of them developed a noticeable narrowing of the necrotic section (Figure 7h). On four shoots, dieback of the entire distal segments above the inoculation site was observed. Diplodia fraxini pycnidia developed on 81.3% of the necrotic lesion, while on 18.8% of the shoots, they were concentrated only on wounded tissue. Of the 48 shoots inoculated with D. seriata, 12 (25.0%) shoots did not develop necrotic lesions, and only the wounded tissue died (Figure 7i). Necrotic lesions were found in 36 (75.0%) shoots (Figure 7j–m, Table 2). Only on seven shoots was the necrosis more extensive and reached from 14 to 27 mm in length (Figure 7j). The size of necrosis on the remaining shoots was very limited and did not exceed 5 mm outside the inoculation wound (Figure 7k–m). The average length of a necrotic lesion caused by a given isolate ranged from 6.50 to 10.38 mm (Table 2). Some of them did not differ significantly from the control group (Table 2). On some shoots, although the tissues underwent longitudinal necrosis, intensive callus formation and scarring of wounds occurred. The necrotic lesions thus became poorly visible (Figure 7l,m). Despite this, the internal parts of the wood within the necrotic lesion became intensely gray in color. Diplodia seriata pycnidia developed on six shoots (Figure 7j), in four of them only on the wounded part (Figure 7k,m). Of the 48 shoots of F. excelsior inoculated with Do. sarmentorum, necrotic lesion occurred on 47 shoots (Table 2). However, in 10 (20.8%) shoots, necrosis was limited to an adjacent narrow zone (up to 5 mm) around the inoculation wound, which was partially or completely healed (Figure 7n). On the remaining shoots, the wound was unhealed, and the length of the necroses ranged from 13 to 42 mm (Figure 7o–q). Necrotic tissue was mostly characterized by a reddish-brown discoloration (Figure 7n–q). On most shoots, necrosis was accompanied by slight collapse of the tissues (Figure 7o–q). There was no dying of whole distal parts of shoots above the inoculation site. Do. sarmentorum pycnidia developed in the necrotic area on 11 (22.9%) shoots (Figure 7q). Do. omnivora did not cause necrotic lesion on any of the 24 inoculated F. excelsior shoots (Table 2). Only wounded tissue died. Inoculation wounds were partially (29.2%) or completely (70.8%) healed (Figure 7r). Pycnidia on inoculated shoots were not observed. Diplodia sapinea did not cause any disease symptoms (Table 2). The inoculation wounds were partially (18.7%) or completely (81.3%) healed (Figure 7s).
Based on re-isolation, the following species were found in inoculated shoots with or without necrotic lesion: D. fraxini in 91.7%, D. seriata in 65.1%, D. sapinea in 37.5%, Do. omnivora in 91.7%, and Do. sarmentorum in 82.3% of shoots. It should be emphasized that species such as D. seriata and Do. omnivora were numerously re-isolated from discolored wood located under completely healed inoculation wounds (Figure 7m,r). None of the control shoots developed necroses (Table 2). The control wounds on ash shoots healed completely at the time of evaluation (Figure 7t). None of the tested fungal cultures were isolated from the control shoots.

4. Discussion

4.1. Diversity of Botryosphaeriaceae Species on F. excelsior

In recent years, the use of morphological characters and phylogenetic analyses has made it possible to determine the species diversity of fungi from the Botryosphaeriaceae family on various species of forest trees and crop plants [19,21,22,23,72,73,74,75]. During the current research in Poland, five Botryosphaeriaceae species were distinguished on F. excelsior with ash dieback symptoms: Diplodia fraxini, D. seriata, D. sapinea, Dothiorella omnivora, and Do. sarmentorum. Two of them, D. sapinea and Do. omnivora were detected in Poland on F. excelsior for the first time. The diversity of Botryosphaeriaceae on F. excelsior in Poland is therefore slightly lower than in the Mediterranean region, where ten Botryosphaeriaceae species have been found [19,34].
In general, the microscopic features of Botryosphaeriaceae currently identified on F. excelsior are consistent with those reported in the literature [26,28,30,37]. However, sometimes one can find different data on the dimensions of some fungal structures. For example, Rathnayaka et al. [76] report significantly smaller dimensions of conidiomata (70–105 × 16–30 µm) and significantly larger dimensions of conidia (25–31 × 10–15 µm) for Do. sarmentorum on Humulus lupulus L. It is worth noting in the current observations that, in Do. omnivora in vitro, the dimensions of spores change over time, becoming more swollen. In the analyzed population of D. fraxinea in Poland, no conidia of larger dimensions, characteristic of morphotype A, were observed [30].
Although fungal isolates from F. excelsior collected over a long period of time (2007–2017) were currently analyzed, they did not confirm the occurrence of D. mutila. This species was often found on F. excelsior in Poland in previous reports [2,17,36,39]. However, Kraj et al. [32] indicated that D. mutila is a heterogeneous species. All Polish isolates derived from F. excelsior formed a separate group compared to isolates from Malus, Vitis, and Quercus [32]. A few years ago, D. mutila as a fungal species was redefined, part of the population associated mainly with Fraxinus was excluded, and a separate species D. fraxini (Fr.: Fr.) Fr., previously known as Sphaeria fraxini Fr.: Fr. or Botryodiplodia fraxini (Fr.: Fr.) Sacc., was recreated [30]. The fact that D. mutila was not detected at all in the current study cannot be interpreted that F. excelsior does not belong to its range of host plants. The occurrence of D. mutila s. str. on F. excelsior has been confirmed in Italy [19]. D. subglobosa was also found there relatively often [19], which was not identified on F. excelsior in Poland.
Botryosphaeriaceae species found on F. excelsior differ in their host spectrum. D. fraxini occurs on F. excelsior and locally on F. angustifolia [8,9,11,14,18,19,20,33]. It has recently been shown to be an endophyte in European beech in central Germany [77]. The four remaining fungal species currently identified on F. excelsior are plurivorous. Diplodia seriata occurs on over 35 different woody hosts, fruit crops, and ornamental plants worldwide [22,28,30]. It has been found on F. excelsior and F. angustifolia [17,19,23,30]. Dothiorella sarmentorum is a cosmopolitan species, which has been identified on 18 host genera from 12 host families, in which identifications are supported by molecular data made post-2005 [78]. These authors do not list Fraxinus as a host plant for this fungus. However, in various European countries, Do. sarmentorum was confirmed on F. excelsior [16,17,18,37]. The host range of D. sapinea is wide, including Pinus species and other coniferous plants [28,42]. Recently, it has been shown that it can infect hosts other than conifer plants: Alnus incana (L.) Moench, Corylus avellana L., Fagus sylvatica L., Fraxinus excelsior, or Olea europaea L. [8,44,46,79]. European ash as a host tree was also confirmed in current research. The inoculum probably came from Pinus sylvestris L., which grows in the vicinity of the tested ash stands. Knowledge of the host range is important from an epidemiological point of view and for forest disease management [80]. This allows to determine where reservoirs of infectious material may be located in forest stands, which may enable appropriate actions to be taken [17]. Dothiorella omnivora has so far been found on Chamaecyparis lawsoniana (A. Murray) Parl., Corylus avellana, Fraxinus excelsior, Juglans regia, Malus domestica Borkh., Ostrya carpinifolia Scop., Thuja occidentalis L., and Vitis vinifera L. [46,58]). The currently performed inoculations on abscised shoots of F. excelsior and J. regia indicate that the latter plant is more suitable for the production of fruitbodies and conidia. In forest stands, ash trees are probably infected by inoculum created on other plants.
Sequence analysis of the ITS region alone proved insufficient to confirm the correctness of the identification based on morphological characteristics for three of the five studied species: Diplodia seriata, Dothiorella omnivora, and Dothiorella sarmentorum. This outcome indicates that, within the Botryosphaeriaceae family, the ITS region, although widely adopted as a primary barcode, is often too conservative for species-level differentiation. Consequently, additional genetic markers, such as TEF1 and TUB2, are required for reliable species identification in this group. Furthermore, molecular research on the genus Diplodia suggest that mating type (MAT) genes may offer even higher phylogenetic resolution than standard multilocus approaches, particularly when resolving closely related cryptic species [81].
Phylogenetic analysis of the genus Diplodia, based on combined ITS and TEF1 data, clearly distinguished the three studied species: Diplodia fraxini, Diplodia sapinea, and Diplodia seriata. Similar results were reported by Alves et al. [30]. The clades representing these species are characterized by the presence of several lineages, indicating significant intraspecific diversity. Furthermore, substructuring within the D. seriata clade is evident in studies by Olmo et al. [82], González-Domínguez et al. [56], and Lodolo et al. [58].
In this study, Dothiorella omnivora is clearly distinguished from other species based on phylogenetic analyses for the combined ITS+TEF1 dataset, including its sister species Do. vidmadera W.M. Pitt, Úrbez-Torr. & Trouillas. Similar results, confirming the distinct taxonomic status of Do. omnivora, were also obtained by Linaldeddu et al. [46] and Váczy et al. [57].
However, the same multigene analysis (ITS+TEF1) did not confirm taxonomic separation between Dothiorella sarmentorum and Dothiorella italica Dissan., Camporesi & K.D. Hyde. This result, where isolates of both taxa are interspersed within a common poorly resolved clade, suggests that Do. sarmentorum and Do. italica may be synonyms or that they belong to a species complex whose actual diversity is not fully captured by the genetic markers used (ITS and TEF1). Dissanayake et al. [55], when distinguishing Do. italica, similarly relied on phylogenetic analysis based on combined ITS and TEF sequence data. Unlike the present study, however, they used the maximum parsimony (MP) method instead of the maximum likelihood (ML) method to generate the phylogenetic tree. Consequently, they obtained a topology grouping isolates described as Do. italica with high statistical support.
As a result of multi-gene phylogenetic analysis, Zhang et al. [27] concluded that Do. italica should be considered a synonym of Do. sarmentorum. Our results appear to support this conclusion, as the ITS+TEF1 analysis failed to resolve these two taxa, showing the type strains of both species interspersed within the same clade. However, further studies are needed to definitively confirm their taxonomic status and species boundaries with a wider range of molecular markers.

4.2. Temperature Assay

Many Botryosphaeriaceae species can grow in a wide range of temperatures, from 5 to 30 or 35 °C, and the optimum growth of many of them occurs at 25 °C [21,28,30,74]. This scheme generally includes all Diplodia and Dothiorella species currently isolated from F. excelsior, except Do. omnivora, which showed optimal growth at 19 °C. It should be noted that some differences between isolates are observed. The currently tested D. seriata isolates showed quite good growth at 30–35 °C. Isolates of D. seriata from olive trees in Spain grew at 10 to 30 °C, but some of the isolates failed to grow at 5 or 35 °C. The optimal temperature for their mycelial growth was much higher than Polish isolates and ranged from 26.9 to 29.8 °C [83]. The optimal temperature for growth of D. seriata isolates collected from grapevines in New Zealand was 25.9 °C [72]. Isolates from Pistacia (USA) grew the fastest at 25 °C and showed some growth also at 40 °C [74]. In turn, isolates of Do. sarmentorum derived from Quercus spp. had an optimal temperature for growth of about 21 °C and did not grow at 35 °C [84]. This may be related to some intraspecific diversity reflected by phylogenetic analyses [22,23,72,74] or adaptation to the conditions of a given geographical zone, what has been found for other fungal species [85]. This was not observed in D. fraxini, isolates of which both from southern Europe [30] and those currently tested from various regions of Poland showed optimal growth at a temperature of about 25 °C. Good growth of many species at 25 °C and higher temperatures indicates that global warming may increase the occurrence of many botryosphaeriaceous species and the pathogenic role of some of them [3,41,86,87]. This fully applies to D. fraxini, which is gaining importance in relation to the health of ash stands. The opposite may be the reaction of the ash dieback causal agent H. fraxineus to climate warming [88].

4.3. Pathogenicity of Botryosphaeriaceae Species Towards Fraxinus excelsior

The pathogenicity test was carried out on 9-year-old F. excelsior seedlings growing in field conditions, similar to those found in young forest plantations. Other experiments indicate that the size of necrotic lesions caused by Botryosphaeriaceae species shows significant differences depending on whether the inoculated tree shoots were attached or detached [89]. The currently conducted experiments with five Botryosphaeriaceae species showed that D. fraxini should be considered the most pathogenic. It caused necrotic lesions on all inoculated ash shoots. It was also able to cause necrosis covering the entire circumference of the shoots in just 12 weeks, which resulted in the death of the top part of the shoot above the place of inoculation. In addition to uniformly brown necrotic tissue, there were discolorations with dark gray zones. This may be related to the intensive production of melanin, which is observed in some Diplodia species at high exposure to light [86]. In ash trees inoculated with H. fraxineus, similar grey–brown zones were formed along the border with living tissues and were associated with the defensive reactions of the host plant [90]. It was found that D. fraxini produced isochromanone, named fraxitoxin, which exhibits phytotoxic activity toward ash [91]. It is very likely that it is an important virulence factor involved in the pathogenesis leading to necrotic lesion and cankers observed in ash trees. The currently obtained results regarding the pathogenicity of D. fraxini are consistent with the results of studies in Italy, where D. fraxini caused larger lesions than D. subglobosa or D. mutila s. str. [19]. However, in general, in the ash dieback process in Poland, the role of D. fraxini or other Botryosphaeriaceae has so far been significantly smaller than Hymenoscyphus fraxineus. This is mainly indicated by the lower frequency of their occurrence on dying trees [17,39].
The current tests showed that the fungus causing the second most extensive necrotic lesion was Do. sarmentorum. It was distinctive that the necrotic tissue showed a reddish-brown discoloration. This may be the result of the interaction of specific metabolites. Studies on grapevines have shown that the infection mechanism of Do. sarmentorum may be implicated by the phytotoxic 6-methylpyridione [92]. Do. sarmentorum can cause disease symptoms in a wide host range, but its importance for plant health varies. Through pathogenicity tests, it was confirmed that Do. sarmentorum can cause canker and dieback in grapevines (Vitis vinifera), loguat (Eriobotrya japonica (Thunb.) Lindl.), and Quercus ilex L. [22,56,84]. However, this fungus turned out to be weakly virulent to pistachio (Pistacia vera L.) [74] and did not produce lesions on Forsythia europaea Degen & Bald. [75].
The tested D. seriata isolates caused only very limited necrotic lesions, and sometimes they did not cause any visible symptoms on F. excelsior shoots, which is similar to the tests performed on F. excelsior in southern Europe [20,21]. These observations correlate well with the relatively rare detection of this species on F. excelsior with dieback symptoms [17]. D. seriata occurs on many different host plants in association with various disease symptoms such as branch cankers, dieback twigs, leaf spots, and fruit rot [22,28,78]. Pathogenicity tests have shown, for example, that it is weakly virulent to almond, English walnut, olive, and pistachio [74,83,93]. However, it is an aggressive pathogen on vinegard and hazelnut [22,46]. In Ligustrum vulgare L. it caused vascular discoloration and did not produce lesions when inoculated into Quercus rubra L. or Prunus cerasus L. shoots [75]. Metabolites such as mellein, hydrophilic high-molecular-weight compounds, and exopolysaccharides with phytotoxic properties may play a significant role in the pathogenesis of D. seriata [94]. In the light of the results obtained, the remaining two tested species, D. sapinea and Do. omnivora, are currently of no importance in reducing the health status of F. excelsior. As stated above, D. sapinea is a known serious cause of the dieback of pine shoots and other Coniferae [42]. Observations on Do. omnivora indicate that it is not a dangerous pathogen. This species on inoculated twigs of Malus domestica produced mean necrotic lesions with an area of only 73 mm2 [58], and on inoculated excised hazelnut logs, necrotic lesions did not differ significantly from the control [46].
The re-isolation results obtained after 12 weeks indicate that the tested species, apart from D. sapinea, adapt well and survive in the bark and wood within the inoculation area, even in situations where there is no necrosis and when the wounds have been significantly healed. The high growth rate of the tested fungi may have contributed to the high percentage of successful re-isolations. In tissues densely colonized by them, there may not be much space left for secondary colonizers. The question arises as to whether they can move from such niches to pathogenic activity when environmental conditions change or the host plant weakens, as is known in endophytic Botryosphaeriaceae [41]. The currently conducted experiment does not allow for an assessment of whether the tested fungi are able to infect intact ash tissue [46]. These fungi in nature infect mainly with conidial spores and, to a lesser extent, with ascospores [28,30]. Given the pathogenicity of these canker-causing fungi, it is important to consider that wounding a tree for inoculation increases the ability of the pathogen to penetrate the host. This type of injury may often occur during various cultivation activities on crop plants [24,54]. In case of F. excelsior, frequent wounds may be the result of infection by primary pathogens causing necrotic lesion and canker on trunks and branches. Hymenoscyphus fraxineus has been such a pathogenic fungus in Europe for several decades [1,3,5]. The lifestyle of H. fraxineus shows that it does not require tissue damage to infect F. excelsior, which has not been studied in the case of D. fraxini [3,5,19,21]. In addition, plant pathogens from the Botryosphaeriaceae family may be favored by various abiotic factors that cause stress in plants [18,28,87,95,96]. They seem to become more aggressive during hot and dry periods. Some of them show high plasticity and adaptation to very different climatic conditions [34].

5. Conclusions

European ash (Fraxinus excelsior) has been suffering in Europe since the early 1990s from a serious ash dieback disease caused by an invasive alien ascomycete, Hymenoscyphus fraxineus. Among the fungal community on dying European ash, species of the Botryosphaeriaceae have a significant share. As a result of the analysis of 70 isolates collected for 11 years in Poland, five fungal species from this family were distinguished on F. excelsior with ash dieback symptoms: Diplodia fraxini, D. seriata, D. sapinea, Dothiorella omnivora and Do. sarmentorum. These studies did not confirm the presence of D. mutila sensu stricto on ash trees. Dothiorella omnivora has been detected in Poland for the first time, and D. sapinea, known in Poland on coniferous trees, has been identified for the first time on F. excelsior. Diplodia fraxini is only known to occur on Fraxinus spp. The remaining species are plurivorous, which means that ash trees can be infected from inoculum produced on other tree species in forest stands. Of the five fungal species tested, only three of them caused necrotic lesions: D. fraxini, D. seriata, and Do. sarmentorum. The extent of damage caused by them showed statistically significant differences. Among the fungal species currently studied, the highest pathogenic role should be assigned to D. fraxini. Colonies of the tested fungi showed optimal growth between 19.0 °C and 25.7 °C, but they can also grow quite well at higher temperatures (30.0 °C and some even up to 35.0 °C). Global climate change and associated stress conditions for trees may favor the development of many Botryosphaeriaceae species in infected ash tissues.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/f17020150/s1, Table S1: The primers used for DNA amplification; Figure S1. Daily air temperature (maximum, mean, minimum) and total precipitation at the Nowy Sącz meteorological station, August–October 2018. Data obtained from the Institute of Meteorology and Water Management (IMGW-PIB); Figure S2: Phylogram from maximum likelihood (ML) analyses of ITS data for Diplodia spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Lasiodiplodia theobromae represents the outgroup; Figure S3: Phylogram from maximum likelihood (ML) analyses of ITS data for Dothiorella spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Neofusicoccum luteum represents the outgroup; Figure S4. Variation in the length and width of Diplodia fraxini conidia in relation to their origin. Values marked with different letters within a single graph differ significantly at p < 0.05 according to the Mann–Whitney U test; Figure S5. Variation in conidial length and width of Dothiorella omnivora in relation to their origin and the age of the cultures. Values marked with different letters within a single graph differ significantly at p < 0.05 according to the Kruskal–Wallis test; Figure S6. Variation in the length and width of conidia of four Botryosphaeriaceae species in vivo. Values marked with different letters within a single graph differ significantly at p < 0.05 according to the Kruskal–Wallis test.

Author Contributions

Conceptualization, T.K. and P.B.; methodology, T.K. and B.G. (mycological and pathological aspects) and P.B. (molecular and statistical aspects); investigation, T.K., B.G., and P.B.; formal analysis, T.K. and P.B.; data curation, T.K. and P.B.; writing—original draft preparation, T.K. and P.B.; writing—review and editing, T.K. and P.B.; software, P.B.; supervision, T.K. and P.B.; visualization, P.B.; project administration and funding acquisition, T.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research was partly conducted in a project No. 2016/21/B/NZ9/01226, financed by the National Science Centre, Poland, and partly financed by the Ministry of Science and Higher Education of the Republic of Poland (SUB/040013-D019).

Data Availability Statement

The data presented in this study are available in the Supplementary Materials.

Acknowledgments

The authors would like to thank Paweł Szczygieł, the Head of the Stary Sącz Forest District, for his help during the field research.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Kowalski, T. Ozamieraniu jesionów [About the dying of ash trees]. Tryb. Leśnika 2001, 4, 6–7. [Google Scholar]
  2. Przybył, K. Fungi associated with necrotic apical parts of Fraxinus excelsior shoots. For. Pathol. 2002, 32, 387–394. [Google Scholar] [CrossRef]
  3. Gross, A.; Holdenrieder, O.; Pautasso, M.; Queloz, V.; Sieber, T.N. Hymenoscyphus pseudoalbidus, the causal agent of European ash dieback. Mol. Plant Pathol. 2014, 15, 5–21. [Google Scholar] [CrossRef]
  4. Coker, T.L.R.; Rozsypálek, J.; Edwards, A.; Harwood, T.P.; Butfoy, L.; Buggs, R.J.A. Estimating mortality rates of European ash (Fraxinus excelsior) under the ash dieback (Hymenoscyphus fraxineus) epidemic. Plants People Planet 2019, 1, 48–58. [Google Scholar] [CrossRef]
  5. Enderle, R.; Stenlid, J.; Vasaitis, R. An overview of ash (Fraxinus spp.) and the ash dieback disease in Europe. CABI Rev. 2019, 2019, 1–12. [Google Scholar] [CrossRef]
  6. Pastircáková, K.; Adamcíková, K.; Barta, M.; Pažitnỳ, J.; Hot’ka, P.; Sarvašová, I.; Horáková, M.K. Host range of Hymenoscyphus fraxineus in Slovak Arboreta. Forests 2020, 11, 596. [Google Scholar] [CrossRef]
  7. Agan, A.; Tedersoo, L.; Hanso, M.; Drenkhan, R. Traces of Hymenoscyphus fraxineus in Northeastern Europe extend further back in history than expected. Plant Dis. 2023, 107, 344–349. [Google Scholar] [CrossRef]
  8. Peters, S.; Fuchs, S.; Bien, S.; Bußkamp, J.; Langer, G.J.; Langer, E.J. Fungi associated with stem collar necroses of Fraxinus excelsior affected by ash dieback. Mycol. Prog. 2023, 22, 52. [Google Scholar] [CrossRef]
  9. Bakys, R.; Pliūra, A.; Bajerkevičienė, G.; Marčiulynas, A.; Marčiulynienė, D.; Lynikienė, J.; Menkis, A. Mycobiota associated with symptomatic and asymptomatic Fraxinus excelsior in post-dieback forest stands. Forests 2022, 13, 1609. [Google Scholar] [CrossRef]
  10. Migliorini, D.; Luchi, N.; Nigrone, E.; Pecori, F.; Pepori, A.L.; Santini, A. Expansion of Ash Dieback towards the scattered Fraxinus excelsior range of the Italian peninsula. Biol. Invasions 2022, 24, 1359–1373. [Google Scholar] [CrossRef]
  11. Soldi, E.; Tiley, A.; O’hanlon, R.; Murphy, B.R.; Hodkinson, T.R. Ash dieback and other pests and pathogens of Fraxinus on the island of Ireland. Biol. Environ. 2022, 122, 85–122. [Google Scholar] [CrossRef]
  12. Marçais, B.; Giraudel, A.; Husson, C. Ability of the ash dieback pathogen to reproduce and to induce damage on its host are controlled by different environmental parameters. PLoS Pathog. 2023, 19, e1010558. [Google Scholar] [CrossRef] [PubMed]
  13. Kirisits, T.; Matlakova, M.; Mottinger-Kroupa, S.; Halmschlager, E.; Lakatos, F. Chalara fraxinea associated with dieback of narrow-leafed ash (Fraxinus angustifolia). Plant Pathol. 2010, 59, 411. [Google Scholar] [CrossRef]
  14. Kranjec Orlović, J.; Moro, M.; Diminić, D. Role of root and stem base fungi in Fraxinus angustifolia (Vahl) dieback in Croatian floodplain forests. Forests 2020, 11, 607. [Google Scholar] [CrossRef]
  15. Kirisits, T. Further observations on the association of Hymenoscyphus fraxineus with Fraxinus ornus. Balt. For. 2017, 23, 60–67. [Google Scholar]
  16. Bakys, R.; Vasaitis, R.; Barklund, P.; Thomsen, I.M.; Stenlid, J. Occurrence and pathogenicity of fungi in necrotic and non-symptomatic shoots of declining common ash (Fraxinus excelsior) in Sweden. Eur. J. For. Res. 2009, 128, 51–60. [Google Scholar] [CrossRef]
  17. Kowalski, T.; Kraj, W.; Bednarz, B. Fungi on stems and twigs in initial and advanced stages of dieback of European ash (Fraxinus excelsior) in Poland. Eur. J. For. Res. 2016, 135, 565–579. [Google Scholar] [CrossRef]
  18. Pastirčáková, K.; Ivanová, H.; Pastirčák, M. Species diversity of fungi on damaged branches and leaves of ashes (Fraxinus spp.) in different types of stands in Slovakia. Cent. Eur. For. J. 2018, 64, 133–139. [Google Scholar] [CrossRef]
  19. Linaldeddu, B.T.; Bottecchia, F.; Bregant, C.; Maddau, L.; Montecchio, L. Diplodia fraxini and Diplodia subglobosa: The Main Species Associated with Cankers and Dieback of Fraxinus excelsior in North-Eastern Italy. Forests 2020, 11, 883. [Google Scholar] [CrossRef]
  20. Linaldeddu, B.T.; Bregant, C.; Montecchio, L.; Brglez, A.; Piškur, B.; Ogris, N. First report of Diplodia fraxini and Diplodia subglobosa causing canker and dieback of Fraxinus excelsior in Slovenia. Plant Dis. 2022, 106, 26–29. [Google Scholar] [CrossRef]
  21. Lyubenova, A.; Dimitrova-Mateva, P. Severe Dieback of European Ash Shelterbelts in Northeastern Bulgaria Associated with Diplodia fraxini. Forests 2025, 16, 1701. [Google Scholar] [CrossRef]
  22. Carlucci, A.; Cibelli, F.; Lops, F.; Raimondo, M.L. Characterization of Botryosphaeriaceae species as causal agents of trunk diseases on grapevines. Plant Dis. 2015, 99, 1678–1688. [Google Scholar] [CrossRef] [PubMed]
  23. Kazemzadeh Chakusary, M.; Mohammadi, H.; Khodaparast, S.A. Diversity and pathogenicity of Botryosphaeriaceae species on forest trees in the north of Iran. Eur. J. For. Res. 2019, 138, 685–704. [Google Scholar] [CrossRef]
  24. Wiman, N.G.; Webber, J.B.; Wiseman, M.; Merlet, L. Identity and pathogenicity of some fungi associated with hazelnut (Corylus avellana L.) trunk cankers in Oregon. PLoS ONE 2019, 14, e0223500. [Google Scholar] [CrossRef] [PubMed]
  25. Phillips, A.J.L.; Lopes, J.; Abdollahzadeh, J.; Bobev, S.; Alves, A. Resolving the Diplodia complex on apple and other Rosaceae hosts. Persoonia Mol. Phylogeny Evol. Fungi 2012, 29, 29–38. [Google Scholar] [CrossRef]
  26. Dissanayake, A.J.; Phillips, A.J.L.; Li, X.H.; Hyde, K.D. Botryosphaeriaceae: Current status of genera and species. Mycosphere 2016, 7, 1001–1073. [Google Scholar] [CrossRef]
  27. Zhang, W.; Groenewald, J.Z.; Lombard, L.; Schumacher, R.K.; Phillips, A.J.L.; Crous, P.W. Evaluating species in Botryosphaeriales. Persoonia Mol. Phylogeny Evol. Fungi 2021, 46, 63–115. [Google Scholar] [CrossRef]
  28. Phillips, A.J.L.; Alves, A.; Abdollahzadeh, J.; Slippers, B.; Wingfield, M.J.; Groenewald, J.Z.; Crous, P.W. The Botryosphaeriaceae: Genera and species known from culture. Stud. Mycol. 2013, 76, 51–167. [Google Scholar] [CrossRef]
  29. Nagel, J.H.; Wingfield, M.J.; Slippers, B. Next-generation sequencing provides important insights into the biology and evolution of the Botryosphaeriaceae. Fungal Biol. Rev. 2021, 38, 25–43. [Google Scholar] [CrossRef]
  30. Alves, A.; Linaldeddu, B.T.; Deidda, A.; Scanu, B.; Phillips, A.J.L. The complex of Diplodia species associated with Fraxinus and some other woody hosts in Italy and Portugal. Fungal Divers. 2014, 67, 143–156. [Google Scholar] [CrossRef]
  31. Zhou, S.; Stanosz, G.R. Relationships among Botryosphaeria species and associated anamorphic fungi inferred from the analyses of ITS and 5.8S rDNA sequences. Mycologia 2001, 93, 516–527. [Google Scholar] [CrossRef]
  32. Kraj, W.; Kowalski, T.; Zarek, M. Structure and genetic variation of Diplodia mutila on declining ashes (Fraxinus excelsior) in Poland. J. Plant Pathol. 2013, 95, 499–507. [Google Scholar] [CrossRef]
  33. Elena, G.; León, M.; Abad-Campos, P.; Armengol, J.; Mateu-Andrés, I.; Güemes-Heras, J. First Report of Diplodia fraxini Causing Dieback of Fraxinus angustifolia in Spain. Plant Dis. 2018, 102, 2645. [Google Scholar] [CrossRef]
  34. Benigno, A.; Bregant, C.; Aglietti, C.; Rossetto, G.; Tolio, B.; Moricca, S.; Linaldeddu, B.T. Pathogenic fungi and oomycetes causing dieback on Fraxinus species in the Mediterranean climate change hotspot region. Front. For. Glob. Change 2023, 6, 1253022. [Google Scholar] [CrossRef]
  35. Orlović, J.K.; Bregant, C.; Linaldeddu, B.T.; Montecchio, L.; Volenec, I.; Uidl, K.; Diminić, D. Diplodia fraxini: The Main Pathogen Involved in the Ash Dieback of Fraxinus angustifolia in Croatia. Microorganisms 2025, 13, 1238. [Google Scholar] [CrossRef]
  36. Kowalski, T.; Kraj, W.; Szeszycki, T. Badania nad zamieraniem jesionu w drzewostanach nadleśnictwa Rokita. The studies on ash decline in Rokita Forest District stands. Acta Agrar. Silvestria Ser. Silvestris 2012, 50, 3–22. [Google Scholar]
  37. Ivanová, H. Identification and characterization of the fungus Dothiorella sarmentorum on necrotic shoots of declining ash in Slovakia. Folia Oecologica 2018, 45, 53–57. [Google Scholar] [CrossRef]
  38. Kowalski, T.; Łukomska, A. Badania nad zamieraniem jesionu (Fraxinus excelsior L.) w drzewostanach Nadleśnictwa Włoszczowa. Studies on Fraxinus excelsior L. dieback in Włoszczowa Forest Unit stands. Acta Agrobot. 2005, 58, 429–440. [Google Scholar] [CrossRef]
  39. Kowalski, T.; Czekaj, A. Symptomy chorobowe i grzyby na zamierających jesionach (Fraxinus excelsior L.) w drzewostanach Nadleśnictwa Staszów. Disease symptoms and fungi on dying ash trees (Fraxinus excelsior L.) in Staszów Forest District stands. Leśne Pr. Badaw. 2010, 71, 357–368. [Google Scholar]
  40. Schlegel, M.; Queloz, V.; Sieber, T.N. The endophytic mycobiome of European ash and sycamore maple leaves—Geographic patterns, host specificity and influence of ash dieback. Front. Microbiol. 2018, 9, 2345. [Google Scholar] [CrossRef]
  41. Slippers, B.; Wingfield, M.J. Botryosphaeriaceae as endophytes and latent pathogens of woody plants: Diversity, ecology and impact. Fungal Biol. Rev. 2007, 21, 90–106. [Google Scholar] [CrossRef]
  42. Blumenstein, K.; Bußkamp, J.; Langer, G.J.; Terhonen, E. Diplodia tip blight pathogen’s virulence empowered through host switch. Front. Fungal Biol. 2022, 3, 939007. [Google Scholar] [CrossRef]
  43. Rafiqi, M.; Kosawang, C.; Peers, J.A.; Jelonek, L.; Yvanne, H.; McMullan, M.; Nielsen, L.R. Endophytic fungi related to the ash dieback causal agent encode signatures of pathogenicity on European ash. IMA Fungus 2023, 14, 10. [Google Scholar] [CrossRef]
  44. Lazzizera, C.; Frisullo, S.; Alves, A.; Lopes, J.; Phillips, A.J.L. Phylogeny and morphology of Diplodia species on olives in Southern Italy and description of Diplodia olivarum sp. nov. Fungal Divers. 2008, 31, 63–71. [Google Scholar]
  45. Phillips, A.J.L.; Alves, A.; Pennycook, S.R.; Johnston, P.R.; Ramaley, A.; Akulov, A.; Crous, P.W. Resolving the phylogenetic and taxonomic status of dark-spored teleomorph genera in the Botryosphaeriaceae. Persoonia Mol. Phylogeny Evol. Fungi 2008, 21, 29–55. [Google Scholar] [CrossRef]
  46. Linaldeddu, B.T.; Deidda, A.; Scanu, B.; Franceschini, A.; Alves, A.; Abdollahzadeh, J.; Phillips, A.J.L. Phylogeny, morphology and pathogenicity of Botryosphaeriaceae, Diatrypaceae and Gnomoniaceae associated with branch diseases of hazelnut in Sardinia (Italy). Eur. J. Plant Pathol. 2016, 146, 259–279. [Google Scholar] [CrossRef]
  47. Bilański, P.; Grad, B.; Kowalski, T. Pyrenochaeta fraxinina as colonizer of ash and sycamore petioles, its morphology, ecology, and phylogenetic connections. Mycol. Prog. 2022, 21, 74. [Google Scholar] [CrossRef]
  48. White, T.J.; Bruns, T.; Lee, S.; Taylor, J.W. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In PCR Protocols: A Guide to Methods and Applications; Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J., Eds.; Academic Press Inc.: New York, NY, USA, 1990; pp. 315–322. [Google Scholar]
  49. Gardes, M.; Bruns, T.D. ITS primers with enhanced specificity for Basidiomycetes—Application to the identification of mycorrhizae and rusts. Mol. Ecol. 1993, 2, 113–118. [Google Scholar] [CrossRef]
  50. Glass, N.L.; Donaldson, G.C. Development of primer sets designed for use with the PCR to amplify conserved genes from filamentous ascomycetes. Appl. Environ. Microbiol. 1995, 61, 1323–1330. [Google Scholar] [CrossRef] [PubMed]
  51. Alves, A.; Crous, P.W.; Correia, A.; Phillips, A.J.L. Morphological and molecular data reveal cryptic speciation in Lasiodiplodia theobromae. Fungal Divers. 2008, 28, 1–13. [Google Scholar]
  52. Zhang, Z.; Schwartz, S.; Wagner, L.; Miller, W. A Greedy Algorithm for Aligning DNA Sequences. J. Comput. Biol. 2000, 7, 203–214. [Google Scholar] [CrossRef]
  53. Morgulis, A.; Coulouris, G.; Raytselis, Y.; Madden, T.L.; Agarwala, R.; Schäffer, A.A. Database indexing for production MegaBLAST searches. Bioinformatics 2008, 24, 1757–1764. [Google Scholar] [CrossRef] [PubMed]
  54. Olmo, D.; Armengol, J.; León, M.; Gramaje, D. Characterization and Pathogenicity of Botryosphaeriaceae Species Isolated from Almond Trees on the Island of Mallorca (Spain). Plant Dis. 2016, 100, 2483–2491. [Google Scholar] [CrossRef]
  55. Dissanayake, A. Saprobic Botryosphaeriaceae, including Dothiorella italica sp. nov., associated with urban and forest trees in Italy. Mycosphere 2017, 8, 1157–1176. [Google Scholar] [CrossRef]
  56. González-Domínguez, E.; Alves, A.; León, M.; Armengol, J. Characterization of Botryosphaeriaceae species associated with diseased loquat (Eriobotrya japonica) in Spain. Plant Pathol. 2017, 66, 77–89. [Google Scholar] [CrossRef]
  57. Váczy, K.Z.; Németh, M.Z.; Csikós, A.; Kovács, G.M.; Kiss, L. Dothiorella omnivora isolated from grapevine with trunk disease symptoms in Hungary. Eur. J. Plant Pathol. 2018, 150, 817–824. [Google Scholar] [CrossRef]
  58. Lódolo, X.V.; Lutz, M.C.; Mondino, P.; Ousset, J.; Sosa, M.C. First report of Diplodia seriata, Diplodia mutila, and Dothiorella omnivora associated with apple cankers and dieback in Rio Negro, Argentina. Plant Dis. 2022, 106, 325–327. [Google Scholar] [CrossRef]
  59. Hall, T.A. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp. Ser. 1999, 41, 95–98. [Google Scholar]
  60. Katoh, K.; Standley, D.M. MAFFT Multiple Sequence Alignment Software Version 7: Improvements in Performance and Usability. Mol. Biol. Evol. 2013, 30, 772–780. [Google Scholar] [CrossRef]
  61. Guindon, S.; Gascuel, O. A simple, fast and accurate method to estimate large phylogenies by maximum-likelihood. Syst. Biol. 2003, 52, 696–704. [Google Scholar] [CrossRef]
  62. Darriba, D.; Taboada, G.L.; Doallo, R.; Posada, D. jModelTest 2: More models, new heuristics and parallel computing. Nat. Methods 2012, 9, 772. [Google Scholar] [CrossRef] [PubMed]
  63. Guindon, S.; Dufayard, J.-F.; Lefort, V.; Anisimova, M.; Hordijk, W.; Gascuel, O. New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Syst. Biol. 2010, 59, 307–321. [Google Scholar] [CrossRef] [PubMed]
  64. Ronquist, F.; Huelsenbeck, J.P. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 2003, 19, 1572–1574. [Google Scholar] [CrossRef]
  65. Swofford, D.L. PAUP* 4.0. In Phylogenetic Analysis Using Parsimony (*and Other Methods); Sinauer Associates: Sunderland, MA, USA, 2003. [Google Scholar]
  66. Stöver, B.C.; Müller, K.F. TreeGraph 2: Combining and visualizing evidence from different phylogenetic analyses. BMC Bioinform. 2010, 11, 7. [Google Scholar] [CrossRef] [PubMed]
  67. Rambaut, A. FigTree, Tree Figure Drawing Tool, Version 1.4.0; University of Edinburgh: Edinburgh, UK, 2006.
  68. Kowalski, T.; Bilański, P.; Kraj, W. Pathogenicity of fungi associated with ash dieback towards Fraxinus excelsior. Plant Pathol. 2017, 66, 1228–1238. [Google Scholar] [CrossRef]
  69. Hammer, Ø.; Harper, D.A.T.; Ryan, P.D. Past: Paleontological statistics software package for education and data analysis. Palaeontol. Electron. 2001, 4, 9. [Google Scholar]
  70. R Core Team. R: A Language and Environment for Statistical Computing; R Foundation: Vienna, Austria, 2024. [Google Scholar]
  71. TIBCO Software Inc. Data Science Workbench, Statistica Version 14.0; TIBCO Software Inc.: Palo Alto, CA, USA, 2020.
  72. Baskarathevan, J.; Jaspers, M.V.; Jones, E.E.; Ridgway, H.J. Incidence and distribution of botryosphaeriaceous species in New Zealand vineyards. Eur. J. Plant Pathol. 2012, 132, 549–560. [Google Scholar] [CrossRef]
  73. Pitt, W.M.; Úrbez-Torres, J.R.; Trouillas, F.P. Dothiorella vidmadera, a novel species from grapevines in Australia and notes on Spencermartinsia. Fungal Divers. 2013, 61, 209–219. [Google Scholar] [CrossRef]
  74. Chen, S.F.; Morgan, D.P.; Michailides, T.J. Botryosphaeriaceae and Diaporthaceae associated with panicle and shoot blight of pistachio in California, USA. Fungal Divers. 2014, 67, 157–179. [Google Scholar] [CrossRef]
  75. Zlatković, M.; Wingfield, M.J.; Jami, F.; Slippers, B. Host specificity of co-infecting Botryosphaeriaceae on ornamental and forest trees in the Western Balkans. For. Pathol. 2018, 48, e12410. [Google Scholar] [CrossRef]
  76. Rathnayaka, A.R.; Thilini Chethana, K.W.; Phillips, A.J.L.; Jones, E.B.G. Two new species of Botryosphaeriaceae (Botryosphaeriales) and new host/geographical records. Phytotaxa 2022, 564, 8–38. [Google Scholar] [CrossRef]
  77. Tropf, J.; Bien, S.; Bußkamp, J.; Langer, G.J.; Langer, E.J. Fungi associated with Vitality loss of European beech in central Germany. Mycol. Prog. 2025, 24, 53. [Google Scholar] [CrossRef]
  78. Dissanayake, A.J.; Camporesi, E.; Hyde, K.D.; Phillips, A.J.L.; Fu, C.Y.; Yan, J.Y.; Li, X. Dothiorella species associated with woody hosts in Italy. Mycosphere 2016, 7, 51–63. [Google Scholar] [CrossRef]
  79. Bußkamp, J.; Blumenstein, K.; Terhonen, E.; Langer, G.J. Differences in the virulence of Sphaeropsis sapinea strains originating from Scots pine and non-pine hosts. For. Pathol. 2021, 51, e12712. [Google Scholar] [CrossRef]
  80. Jactel, H.; Nicoll, B.C.; Branco, M.; Gonzalez-Olabarria, J.R.; Grodzki, W.; Långström, B.; Moreira, F.; Netherer, S.; Orazio, C.; Piou, D.; et al. The influences of forest stand management on biotic and abiotic risks of damage. Ann. For. Sci. 2009, 66, 701. [Google Scholar] [CrossRef]
  81. Lopes, A.; Linaldeddu, B.T.; Phillips, A.J.L.; Alves, A. Mating type gene analyses in the genus Diplodia: From cryptic sex to cryptic species. Fungal Biol. 2018, 122, 629–638. [Google Scholar] [CrossRef]
  82. Olmo, D.; Gramaje, D.; Armengol, J. Evaluation of fungicides to protect pruning wounds from Botryosphaeriaceae species infections on almond trees. Phytopathol. Mediterr. 2017, 56, 77–86. [Google Scholar] [CrossRef]
  83. Moral, J.; Muñoz-Díez, C.; González, N.; Trapero, A.; Michailides, T.J. Characterization and pathogenicity of Botryosphaeriaceae species collected from olive and other hosts in Spain and California. Phytopathology 2010, 100, 1340–1351. [Google Scholar] [CrossRef]
  84. Sánchez, M.E.; Venegas, J.; Romero, M.A.; Phillips, A.J.L.; Trapero, A. Botryosphaeria and Related Taxa Causing Oak Canker in Southwestern Spain. Plant Dis. 2003, 87, 1515–1521. [Google Scholar] [CrossRef] [PubMed]
  85. Kraj, W.; Kowalski, T. Genetic variation in Polish strains of Gremmeniella abietina. For. Pathol. 2008, 38, 203–217. [Google Scholar] [CrossRef]
  86. Thompson, S.; Alvarez-Loayza, P.; Terborgh, J.; Katul, G. The effects of plant pathogens on tree recruitment in the Western Amazon under a projected future climate: A dynamical systems analysis. J. Ecol. 2010, 98, 1434–1446. [Google Scholar] [CrossRef]
  87. Langer, G.J.; Bußkamp, J. Vitality loss of beech: A serious threat to Fagus sylvatica in Germany in the context of global warming. J. Plant Dis. Prot. 2023, 130, 1101–1115. [Google Scholar] [CrossRef]
  88. Grosdidier, M.; Ioos, R.; Marçais, B. Do higher summer temperatures restrict the dissemination of Hymenoscyphus fraxineus in France? For. Pathol. 2018, 48, e12426. [Google Scholar] [CrossRef]
  89. López-Moral, A.; Lovera, M.; Antón-Domínguez, B.I.; Gámiz, A.M.; Michailides, T.J.; Arquero, O.; Trapero, A.; Agustí-Brisach, C. Effects of Cultivar Susceptibility, Branch Age, and Temperature on Infection by Botryosphaeriaceae and Diaporthe Fungi on English Walnut (Juglans regia). Plant Dis. 2022, 106, 2920–2926. [Google Scholar] [CrossRef]
  90. Kowalski, T.; Holdenrieder, O. Pathogenicity of Chalara fraxinea. For. Pathol. 2009, 39, 1–7. [Google Scholar] [CrossRef]
  91. Cimmino, A.; Maddau, L.; Masi, M.; Linaldeddu, B.T.; Pescitelli, G.; Evidente, A. Fraxitoxin, a new isochromanone isolated from Diplodia fraxini. Chem. Biodivers. 2017, 14, e1700325. [Google Scholar] [CrossRef]
  92. Salvatore, M.M.; Nicoletti, R.; Russo, M.T.; Mahamedi, A.E.; Berraf-Tebbal, A.; DellaGreca, M.; Anna, A. First report of 6-methylpyridione analogues from Dothiorella sarmentorum, a botryosphaeriaceous fungus associated with grapevine trunk diseases. Nat. Prod. Res. 2023, 12, 2748–2755. [Google Scholar] [CrossRef]
  93. Inderbitzin, P.; Bostock, R.M.; Trouillas, F.P.; Michailides, T.J. A six locus phylogeny reveals high species diversity in Botryosphaeriaceae from California almond. Mycologia 2010, 102, 1350–1368. [Google Scholar] [CrossRef]
  94. Reveglia, P.; Billones-Baaijens, R.; Savocchia, S. Phytotoxic Metabolites Produced by Fungi Involved in Grapevine Trunk Diseases: Progress, Challenges, and Opportunities. Plants 2022, 11, 3382. [Google Scholar] [CrossRef] [PubMed]
  95. Fabre, B.; Piou, D.; Desprez-Loustau, M.; Marçais, B. Can the emergence of pine Diplodia shoot blight in France be explained by changes in pathogen pressure linked to climate change? Glob. Change Biol. 2011, 17, 3218–3227. [Google Scholar] [CrossRef]
  96. Piškur, B.; Pavlic, D.; Slippers, B.; Ogris, N.; Maresi, G.; Wingfield, M.J.; Jurc, D. Diversity and pathogenicity of Botryosphaeriaceae on declining Ostrya carpinifolia in Slovenia and Italy following extreme weather conditions. Eur. J. For. Res. 2011, 130, 235–249. [Google Scholar] [CrossRef]
Forests 17 00150 g001aForests 17 00150 g001b
Figure 1. Phylogram from maximum likelihood (ML) analyses of the combined datasets of ITS+TEF1 for Diplodia spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Lasiodiplodia theobromae represents the outgroup.
Figure 1. Phylogram from maximum likelihood (ML) analyses of the combined datasets of ITS+TEF1 for Diplodia spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Lasiodiplodia theobromae represents the outgroup.
Forests 17 00150 g001aForests 17 00150 g001b
Forests 17 00150 g002aForests 17 00150 g002b
Figure 2. Phylogram from maximum likelihood (ML) analyses of the combined datasets of ITS+TEF1 for Dothiorella spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Neofusicoccum luteum represents the outgroup.
Figure 2. Phylogram from maximum likelihood (ML) analyses of the combined datasets of ITS+TEF1 for Dothiorella spp. Sequences obtained in this study are in bold. Bootstrap values ≥ 75% for ML and maximum parsimony (MP) analyses are presented at nodes as follows: ML/MP. Bold branches indicate posterior probabilities values ≥ 0.95 obtained from Bayesian Inference (BI) analyses. * Bootstrap values < 75%. The tree is drawn to scale (see bar) with branch lengths measured in the number of substitutions per site. Neofusicoccum luteum represents the outgroup.
Forests 17 00150 g002aForests 17 00150 g002b
Forests 17 00150 g003
Figure 3. Diplodia fraxini (a–f): (a) conidia oozing from pycnidia in vivo, (b) pycnidia on dead ash petiole, (c) conidiogenous cells and conidia, (d) hyaline conidia developed in vivo, (e) conidia oozing from pycnidia in vitro, (f) conidia developed in vitro, hyaline and pale brown, with one septum; Diplodia seriata (g–i): (g) conidiogenous cells and conidia, (h,i) ripe conidia developed in vivo, without septa (h) and with septa (i). Scale bars: (a) = 5 mm, (b) = 1 mm, (c,d) = 10 µm, (e) = 10 mm, (f) = 25 µm, (g–i) = 10 µm.
Figure 3. Diplodia fraxini (a–f): (a) conidia oozing from pycnidia in vivo, (b) pycnidia on dead ash petiole, (c) conidiogenous cells and conidia, (d) hyaline conidia developed in vivo, (e) conidia oozing from pycnidia in vitro, (f) conidia developed in vitro, hyaline and pale brown, with one septum; Diplodia seriata (g–i): (g) conidiogenous cells and conidia, (h,i) ripe conidia developed in vivo, without septa (h) and with septa (i). Scale bars: (a) = 5 mm, (b) = 1 mm, (c,d) = 10 µm, (e) = 10 mm, (f) = 25 µm, (g–i) = 10 µm.
Forests 17 00150 g003
Forests 17 00150 g004
Figure 4. Dothiorella omnivora (a–g): (a) pycnidia on inoculated abscised shoot of Juglans regia, (b) pycnidia on inoculated abscised shoot of Fraxinus excelsior, only on wounded tissue, (c) conidiogenous cells and young conidia developed in vivo (on J. regia), (d,e) conidia developed in vivo with one or three septa, (f) swollen conidia in 4-month-old culture, (g) conidium in 4 months old culture with a strongly swollen upper cell; Dothiorella sarmentorum (h–j): (h) pycnidia on dead shoot of Fraxinus excelsior, (i) conidia developed in vivo, (j) microconidia developed in vivo. Scale bars: (a) = 2.5 mm, (b) = 1 mm, (c,d) = 20 µm, (e–g) = 10 mm, (h) = 5 mm, (i) = 20 µm, (j) = 10 µm.
Figure 4. Dothiorella omnivora (a–g): (a) pycnidia on inoculated abscised shoot of Juglans regia, (b) pycnidia on inoculated abscised shoot of Fraxinus excelsior, only on wounded tissue, (c) conidiogenous cells and young conidia developed in vivo (on J. regia), (d,e) conidia developed in vivo with one or three septa, (f) swollen conidia in 4-month-old culture, (g) conidium in 4 months old culture with a strongly swollen upper cell; Dothiorella sarmentorum (h–j): (h) pycnidia on dead shoot of Fraxinus excelsior, (i) conidia developed in vivo, (j) microconidia developed in vivo. Scale bars: (a) = 2.5 mm, (b) = 1 mm, (c,d) = 20 µm, (e–g) = 10 mm, (h) = 5 mm, (i) = 20 µm, (j) = 10 µm.
Forests 17 00150 g004
Forests 17 00150 g005
Figure 5. Mean colony diameters of five Botryosphaeriaceae species after five days of growth on MEA at different temperatures. Growth data for each species were fitted using polynomial regression to identify the local maxima of the resulting functions.
Figure 5. Mean colony diameters of five Botryosphaeriaceae species after five days of growth on MEA at different temperatures. Growth data for each species were fitted using polynomial regression to identify the local maxima of the resulting functions.
Forests 17 00150 g005
Forests 17 00150 g006
Figure 6. Result of classical cluster analysis of five Botryosphaeriaceae species by unweighted pair group method with arithmetic mean (UPGMA). The analysis was performed using the Euclidean distances between the average values of colonies diameters after 5 days of growth at various temperatures from 5 to 35 °C, in steps of 5 °C, as an index similarity. The percentage number of bootstrap replicates (n = 1000) is presented in each node.
Figure 6. Result of classical cluster analysis of five Botryosphaeriaceae species by unweighted pair group method with arithmetic mean (UPGMA). The analysis was performed using the Euclidean distances between the average values of colonies diameters after 5 days of growth at various temperatures from 5 to 35 °C, in steps of 5 °C, as an index similarity. The percentage number of bootstrap replicates (n = 1000) is presented in each node.
Forests 17 00150 g006
Forests 17 00150 g007
Figure 7. Appearance of the necrotic lesions on shoots of Fraxinus excelsior twelve weeks after wound inoculation with five botryosphaeriaceous fungi; Diplodia fraxini: (a–c) uniformly brown discolored extensive necrotic lesions with a regular edge, (d,e) necrotic lesions irregularly incised, (f,g) gray brown necrotic lesion with light patches or stripes, (h) necrosis covering the entire circumference of the shoot with constriction, Diplodia seriata: (i) wound closed by callus formation and no necrosis of the surrounding tissue, (j) necrotic lesion without callus formation, (k) small necrosis mostly closed by callus formation with numerous pycnidia on wounded tissue, (l) small necrosis mostly closed by callus formation, (m) small necrosis fully closed by callus formation, Dothiorella sarmentorum: (n) small necrosis partly closed by callus formation, (o–q) reddish brown necrotic lesion without callus formation, Dothiorella omnivora: (r) fully closed inoculation wound, Diplodia sapinea: (s) fully closed inoculation wound, (t) control shoot with completely closed wound. Arrows indicate the presence of ripe pycnidia. Scale bars denote length of 0.3 cm.
Figure 7. Appearance of the necrotic lesions on shoots of Fraxinus excelsior twelve weeks after wound inoculation with five botryosphaeriaceous fungi; Diplodia fraxini: (a–c) uniformly brown discolored extensive necrotic lesions with a regular edge, (d,e) necrotic lesions irregularly incised, (f,g) gray brown necrotic lesion with light patches or stripes, (h) necrosis covering the entire circumference of the shoot with constriction, Diplodia seriata: (i) wound closed by callus formation and no necrosis of the surrounding tissue, (j) necrotic lesion without callus formation, (k) small necrosis mostly closed by callus formation with numerous pycnidia on wounded tissue, (l) small necrosis mostly closed by callus formation, (m) small necrosis fully closed by callus formation, Dothiorella sarmentorum: (n) small necrosis partly closed by callus formation, (o–q) reddish brown necrotic lesion without callus formation, Dothiorella omnivora: (r) fully closed inoculation wound, Diplodia sapinea: (s) fully closed inoculation wound, (t) control shoot with completely closed wound. Arrows indicate the presence of ripe pycnidia. Scale bars denote length of 0.3 cm.
Forests 17 00150 g007
Table 1. Botryosphaeriaceae species isolated in 2007–2017 from Fraxinus excelsior with ash dieback symptoms in Poland used in present study. Sequences were deposited in GenBank.
Table 1. Botryosphaeriaceae species isolated in 2007–2017 from Fraxinus excelsior with ash dieback symptoms in Poland used in present study. Sequences were deposited in GenBank.
Fungal SpeciesIsolate 1Location (Forest District) 2Sampling DateS. T. 3ITSTEF1BT
Diplodia fraxini48EMiechów08.10.2016fPX658893
51ERokita10.10.2012cPX658894
74EOjców23.11.2013cPX658895
247KPińczów24.08.2007cPX658896PX663858
249KGryfice18.08.2009bPX658897PX663859
250KStaszów20.10.2009aPX658898PX663860
251KWłoszczowa02.06.2010cPX658899PX663861
252KWłoszczowa02.06.2010cPX658900PX663862
253KRokita16.08.2010cPX658901PX663863
254KRokita16.08.2010cPX658902PX663864
255KResko12.08.2010cPX658903PX663865
256KResko12.08.2010dPX658904PX663866
257KResko12.08.2010bPX658905PX663867
258KGryfice21.08.2009aPX658906PX663868
259KStaszów05.08.2009aPX658907PX663869
260KMiechów23.05.2010aPX658908PX663870
266KOjców29.05.2010aPX658909PX663871
267KStary Sącz12.12.2013cPX658910PX663872
268KStary Sącz12.12.2013dPX658911PX663873
269KStary Sącz12.12.2013cPX658912PX663874
270KStary Sącz12.12.2013cPX658913PX663875
307EPuławy18.08.2015ePX658914
347KGryfice08.08.2007cPX658915PX663876
348KOjców22.09.2009aPX658916PX663877
349KOjców22.09.2009aPX658917PX663878
350KMiechów25.05.2010aPX658918PX663879
351KMiechów25.05.2010aPX658919PX663880
354KPrzedbórz12.06.2010bPX658920PX663881
357KOjców27.03.2017aPX658921PX663882
358KOjców27.03.2017aPX658922PX663883
974F 1Przedbórz12.06.2010bPX658923PX663884
975FWłoszczowa02.06.2010cPX658924PX663885
976FRokita16.08.2010cPX658925
977FResko12.08.2010bPX658926
978F 1Gryfice18.08.2010bPX658927PX663886
979F 1Staszów23.09.2009aPX658928PX663887PX663920
980FMiechów29.10.2010aPX658929
981FMiechów25.05.2010aPX658930PX663888
983F 1Limanowa12.06.2010bPX658931PX663889
985FOjców23.11.2013dPX658932PX663890
986F 1Stary Sącz12.12.2013cPX658933PX663891PX663921
987F 1Rokita30.08.2009bPX658934PX663892
994FOjców22.09.2009aPX658935PX663893
998FMiechów01.06.2010bPX658936
Diplodia seriata59KLimanowa12.06.2010aPX658937PX663894
71KRokita18.06.2008cPX658938PX663895
248KPińczów24.08.2007cPX658939PX663896
265KMiechów01.06.2016aPX658940PX663897
982F1Stary Sącz10.08.2009cPX658941PX663898
996F1Staszów30.06.2009cPX658942PX663899PX663922
997FRokita16.11.2009cPX658943PX663900
1000F 1Stary Sącz26.08.2010bPX658944PX663901PX663923
1001F 1Miechów02.12.2009aPX658945PX663902
1002F 1Miechów29.05.2010aPX658946PX663903
1003F 1Miechów29.05.2010aPX658947PX663904PX663924
1004FResko12.08.2010aPX658948PX663905PX663925
Diplodia sapinea262K 1Miechów01.06.2010bPX658949PX663906PX663926
Dothiorella omnivora352K 1Stary Sącz10.08.2009cPX658950PX663907PX663927
353K 1Stary Sącz10.08.2009cPX658951PX663908PX663928
355K 1Przedbórz12.06.2010bPX658952PX663909PX663929
Dothiorella sarmentorum261K 1Przedbórz12.06.2010bPX658953PX663910
263KResko12.08.2010aPX658954PX663911
264K 1Miechów25.05.2010cPX658955PX663912PX663930
984FOjców23.08.2015cPX658956PX663913
988F1Przedbórz12.06.2010cPX658957PX663914PX663931
989FGryfice06.08.2010cPX658958PX663915
990F 1Przedbórz12.06.2010bPX658959PX663916
991F 1Włoszczowa02.06.2010bPX658960PX663917
992FWłoszczowa02.06.2010bPX658961PX663918
993F 1Resko12.08.2010cPX658962PX663919
1 Isolates used in temperature assay and in pathogenicity test. 2 The coordinates are given together with the accession number in Genbank, 3 Substrate type: a—dead shoot, conidiomata present, b—dead shoot, conidiomata absent, c—necrotic lesion on living shoot, conidiomata absent, d—necrotic lesion on living shoot, conidiomata present, e—dead petiole in the litter, conidiomata present, f—necrotic lesion on living petiole, conidiomata absent.
Table 2. Number of shoots of Fraxinus excelsior with necrotic lesion 12 weeks post inoculation with five Botryosphaeriaceae species.
Table 2. Number of shoots of Fraxinus excelsior with necrotic lesion 12 weeks post inoculation with five Botryosphaeriaceae species.
Strains 1Inoculated Shoots [n]Occurrence of Longitudinal Necrosis [n]Mean Length of Necrosis [mm] 2,3Standard DeviationStandard Error of Mean
Diplodia fraxini
974F a8838.38 a7.392.61
978F a8839.00 a7.732.73
979F a8842.00 a22.978.12
983F a8836.63 a3.501.24
986F a8834.25 ab14.195.02
987F a8850.50 a29.1910.32
Total484840.13 A16.792.42
Diplodia seriata
982F a867.00 ab4.411.56
996F a867.00 ab4.341.54
1000F a8710.00 a5.131.81
1001F a868.88 a6.292.22
1002F b856.50 ab5.662.00
1003F a8610.38 a8.913.15
Total48368.29 C5.870.85
Dothiorella sarmentorum
261K a8823.13 a12.044.26
264K a8822.00 a5.451.93
988F a8815.63 a4.631.64
990F a8816.75 a6.582.33
991F a8712.25 ab5.341.89
993F a8818.88 a6.622.34
Total484718.10 B7.781.12
Dothiorella omnivora
352K b800.00 b0.000.00
353K b800.00 b0.000.00
355K b800.00 b0.000.00
Total2400.00 D0.000.00
Diplodia sapinea
262K b800.00 b0.000.00
Total800.00 CD0.000.00
Control
Control b800.00 b0.000.00
Total800.00 CD0.000.00
1 Isolates marked with lowercase letters different from the control group caused lesion lengths significantly higher than the control (p < 0.05), according to the Kruskal–Wallis test followed by the Steel post hoc test. 2 Values for species and controls marked with different capital letters are significantly different (p < 0.05) according to the Kruskal–Wallis test followed by Dunn’s post hoc test; p-values were adjusted using the Benjamini–Hochberg method. 3 Values within species and controls marked with different lowercase letters are significantly different at p < 0.05 according to the Kruskal–Wallis test followed by Dunn’s post hoc test; p-values were adjusted using the Benjamini–Hochberg method.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Bilański, P.; Grad, B.; Kowalski, T. Pathogenicity of Diplodia fraxini and Other Botryosphaeriaceae Identified on Fraxinus excelsior with Dieback Symptoms in Poland. Forests 2026, 17, 150. https://doi.org/10.3390/f17020150

AMA Style

Bilański P, Grad B, Kowalski T. Pathogenicity of Diplodia fraxini and Other Botryosphaeriaceae Identified on Fraxinus excelsior with Dieback Symptoms in Poland. Forests. 2026; 17(2):150. https://doi.org/10.3390/f17020150

Chicago/Turabian Style

Bilański, Piotr, Bartłomiej Grad, and Tadeusz Kowalski. 2026. "Pathogenicity of Diplodia fraxini and Other Botryosphaeriaceae Identified on Fraxinus excelsior with Dieback Symptoms in Poland" Forests 17, no. 2: 150. https://doi.org/10.3390/f17020150

APA Style

Bilański, P., Grad, B., & Kowalski, T. (2026). Pathogenicity of Diplodia fraxini and Other Botryosphaeriaceae Identified on Fraxinus excelsior with Dieback Symptoms in Poland. Forests, 17(2), 150. https://doi.org/10.3390/f17020150

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop