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Article

Diversity and Pathogenicity of Botryosphaeriaceae Species Isolated from Olives in Istria, Croatia, and Evaluation of Varietal Resistance

by
Elena Petrović
1,*,
Karolina Vrandečić
2,
Andreina Belušić Vozila
1,
Jasenka Ćosić
2 and
Sara Godena
1
1
Institute of Agriculture and Tourism, Karla Huguesa 8, 52440 Poreč, Croatia
2
Faculty of Agrobiotechnical Sciences Osijek, Josip Juraj Strossmayer University of Osijek, Vladimira Preloga 1, 31000 Osijek, Croatia
*
Author to whom correspondence should be addressed.
Plants 2024, 13(13), 1813; https://doi.org/10.3390/plants13131813
Submission received: 13 June 2024 / Revised: 27 June 2024 / Accepted: 28 June 2024 / Published: 1 July 2024
(This article belongs to the Special Issue Pathogenesis and Disease Control in Crops—2nd Edition)
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Versions Notes

Abstract

:
During 2021 and 2022, a field investigation was conducted in Istria, Croatia, searching for trees exhibiting signs of Botryosphaeria dieback. Samples of symptomatic trees were collected from 26 different locations and analysed. Isolates that morphologically corresponded to species from the Botryosphaeriaceae family were selected, and detailed morphological characterisation and molecular identification of the isolates were conducted. Based on morphological characteristics and phylogenetic analysis using the internal transcribed spacer (ITS), beta-tubulin (TUB2), and translation elongation factor 1-alpha (TEF1-α) regions, six species of fungi from the Botryosphaeriaceae family were identified: Botryosphaeria dothidea (Moug. ex Fr.) Ces. & De Not.; Diplodia mutila (Fr.) Fr.; Diplodia seriata De Not.; Dothiorella iberica A.J.L. Phillips, J. Luque & A. Alves; Dothiorella sarmentorum (Fr.) A.J.L. Phillips, Alves & Luque; and Neofusicoccum parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips. This is the first report of D. mutila, Do. sarmentorum, and Do. iberica causing Botryosphaeria dieback on olive trees in Croatia, and the first study investigating the resistance of Croatian olive varieties to species from the Botryosphaeriaceae family. Pathogenicity testing of selected isolates and assessment of variety resistance were conducted on four different olive varieties, namely Buža, Istarska bjelica, Leccino, and Rosinjola, using representative isolates of the mentioned species. The most aggressive species was found to be N. parvum. Olive varieties exhibited differences in susceptibility depending on the fungus they were infected with.

1. Introduction

The olive (Olea europea L.) is one of the oldest cultivated plants, spread across the Mediterranean from 45° north to 35° south [1]. Material evidence from ancient times indicates the significance of olive growing and olive oil, mostly used as food, fuel for oil lamps, cosmetic purposes, etc. [1]. Numerous tales suggest how olive production expanded. The prevalence of olives in Istria, as well as in Sicily and southern Spain, is attributed by historians to Aristeus, the ancient pastoral god of the Arcadians, Boeotians, and Thessalians, who was considered the inventor of olives and olive oil [2,3]. Pribetić [1] suggests the assumption that the olive’s homeland is Palestine or Asia Minor, from where it first spread to Egypt. Mythology tells of the goddess Minerva, challenged to a contest by Neptune, who plucked the first olive plant from the ground, already in bloom and bearing fruit, thus making it a symbol of peace [4]. It is interesting to note that a golden olive branch was left on the moon’s surface by Apollo 11 crew members as a symbol of peace. Olive cultivation in Istria dates back to the 1st century BC, mainly along the western and southern coastal areas [5]. During the period between 1500 and 1700, there was a significant decline in production, but in the 19th and 20th centuries, its production once again spread throughout the Istrian Peninsula [3]. As stated by Godena et al. [6], the Istrian Peninsula stands as the northernmost olive-growing region in Croatia and is also one of the northernmost olive-growing regions globally. Croatia’s olive strategy lies in producing extra-high-quality olive oil, precisely due to suitable climatic conditions [7]. According to the FAO [8], there are no data available for the annual olive production in the world for the year 2023, but in 2022, the production amounted to 21.4 million t. Olive production in Croatia amounted to approximately 40,100 t in 2022 [8], and 29,800 t in 2023 [9]. The production of a high-quality olive oil strongly depends on the quality of the fruit from which the oil is extracted. However, the quality of the fruit depends on numerous factors, such as agroecological conditions, disease and pest attacks, varieties, etc.
Olive varieties exhibit a vast range of diversity. A key question is whether this differentiation occurred post-domestication or whether olives have multiple origins [10]. Typically, olives are propagated through cuttings or grafts, resulting in varieties that are essentially clones [10]. It is assumed that over 1000 varieties and types of olives are cultivated in the Mediterranean region [1]. According to Rugini [11], the number of cultivated olive varieties is estimated to be around 2500. In studies investigating the pathogenicity of fungi and the susceptibility of olive varieties, two terms frequently appear: olive variety and olive cultivar. When discussing olives and olive trees, there is no difference between these terms, as the term “cultivar” is short for “cultivated variety” and is used frequently in olive growing to refer to the different varieties of olives produced. Therefore, the olive cultivar is a synonym for olive variety [12,13]. A significant advantage of Croatian olive cultivation is the indigenous assortment that distinguishes certain areas with the uniqueness of olive oil aroma and flavour, especially since nowadays, oil and olives with a geographical origin achieve twice the price of oil without an origin [7]. In Istrian olive groves, slightly more than a third of the trees belong to indigenous varieties: Buža (50.69%), Istarska bjelica (30.22%), Rosinjola (5.72%), Crnica (syn. Karbonera, 5.60%), and other less represented varieties (7.77%). New plantations primarily consist of foreign varieties (Leccino, Frantoio, Pendolino) [5].
The Buža variety (syn. Buga, Burgaca, Domaća, Gura, Morgaca) [14] is widespread in Istria. Pribetić [1] lists it as the most widespread variety in that area. It is highly valued inland for its excellent oil. It is sensitive to early autumn cold, which reduces oil yields [3]. It is known that in Istria entire plantations suffered from low temperatures in certain years [1]. The name “Buža” comes from the ancient term “bugio,” meaning pitted, hollow. Hugues [3] suggests that the name of this variety could originate from the frequent cavities or holes in its trunk near the stump. In the Vodnjan area (Istria), the Buža variety is often grown. This variety is susceptible to peacock spot disease caused by the species Venturia oleaginea (Castagne) Rossman & Crous and the appearance of sooty mould, and it is susceptible to olive fruit fly (Bactrocera oleae Rossi) and olive moth (Prays oleae Bernard) [1].
The Rosinjola variety (syn. Rošinjola, Rosulja, Rovinječka, Rušinjola) [14] is mostly grown in the southern part of Istria, around Vrsar, Rovinj, and Vodnjan [1]. It produces good-quality oil, with an intense aroma and a pronounced bitter taste [1]. Due to its dense canopy, it is often attacked by insects, which causes the appearance of sooty mould, but it shows good resistance to other pests and diseases [1].
The Istarska bjelica variety (syn. Bjelica, Bjankera, Bianchera) [14] is also widespread in Istria and on the Kvarner islands. Unlike Buža, it is somewhat more resistant to low temperatures and the wind [1]. It has a high oil yield, abundant productivity, and excellent oil quality, but it is susceptible to B. oleae [1].
The Leccino variety (syn. Leccio) [14] originates from Italy, and it has been cultivated in Istria since 1940 [1]. It is one of the most widespread varieties globally due to its significant adaptability to different agroecological conditions [1]. The Leccino variety is resistant to low temperatures, suitable for intensive plantations, shows good and constant productivity and, finally, produces outstanding-quality oil [1]. Moreover, it is noted to be more resistant to peacock spot disease [1,7], olive knot disease caused by the bacteria Pseudomonas savastanoi pv. savastanoi, and the pest P. oleae, but it is susceptible to B. oleae [1].
A large number of fungi and pests have been reported to cause damage to olive trees. Among the most significant pest of olive trees is B. oleae, which is the most economically significant pest in Croatian olive cultivation. On the other hand, in terms of diseases, there is peacock spot disease, as well as patula, caused by Botryosphaeria dothidea (Moug. ex Fr.) Ces. & De Not., and anthracnose of fruits caused by the species Colletotrichum gloeosporioides (Penz.) Penz. & Sacc. (syn. Gloeosporium olivarum) [7]. Fungi from the Botryosphaeriaceae family identified as pathogens of olive trees in Croatia include the species B. dothidea, Diplodia seriata De Not. and Neofusicoccum parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips [15,16,17,18]. In addition to the fungi from the Botryosphaeriaceae family, other pathogens of olive trees in Croatia include the species Armillaria mellea (Vahl) P. Kumm. [17]; Biscogniauxia mediterranea and Biscogniauxia nummularia (Bull.) Kuntze [19]; Colletotrichum spp. [15]; Comoclathris incompta (Sacc. & Martelli) Ariyaw. & K.D. Hyde (syn. Phoma incompta Sacc. & Mart.) [20]; Cytospora pruinosa Défago [21]; Diaporthe sp. [18]; Nigrospora gorlenkoana Novobr., Nigrospora osmanthi Mei Wang & L. Cai, and Nigrospora philosophiae-doctoris M. Raza, Qian Chen & L. Cai [22]; Phaeoacremonium iranianum L. Mostert, Gräfenhan, W. Gams & Crous [23]; Pleurostoma richardsiae (Nannfeldt) Réblová & Jaklitsch [24]; Pseudocercospora cladosporioides (Sacc.) U. Braun [15]; Sordaria fimicola (Roberge ex Desm.) Ces. & De Not [19]; Venturia oleaginea (Castagne) Rossman & Crous (syn. Spilocaea oleaginea) [25,26]; and Verticillium dahliae Klebahn [27].
Peacock spot disease and patula are among the first described fungal diseases of olives in Croatia. Peacock spot disease was first described in 1901 in Croatia [28]. Patula (syn. Dalmatian disease, Botryosphaeria dieback, escudete, maricume delle drupe, lepre des olives, etc.) was first described in 1883 in Dalmatia by the German–Australian botanist Thumen, and now it is present in almost all olive-growing countries in the Mediterranean region [7]. It compromises the quality of oil and diminishes the value of table olives. The vector of B. dothidea is the olive fruit fly and its predator Lasioptera berlesiana Paoli (syn. Prolasioptera berlesiana). L. berlesiana carries B. dothidea spores in a mycangium. While the mosquito deposits its egg adjacent to the fly egg, it also inoculates the puncture made by B. olea with the fungus [29]. The symptoms of the disease resemble an attack by C. gloeosporioides. Signs of the disease in commercial groves encompass necrotic, sunken, and distinctly demarcated lesions on fruits [30]. Besides B. dothidea, fungi from the Botryosphaeriaceae family are known as some of the most common pathogens of olive trees [31,32]. For instance, causal agents of Botryosphaeria dieback and fruit rot include species such as D. olivarum A.J.L. Phillips & Lazzizera; N. vitifusiforme (Van Niekerk & Crous) Crous, Slippers & A.J.L. Phillips; N. parvum; and N. mediterraneum Crous, M.J. Wingf. & A.J.L. Phillips [33,34]. Disease symptoms can be observed on olive fruits, leaves, branches, and trunks. They cause fruit rot, leaf wilting and defoliation, bark discolouration, branch dieback, canker formations, and the appearance of necrosis [35,36]. Fruit contamination with fungi is also a concern due to mycotoxins, which can develop through fruit decay and prolonged storage of damaged fruit [7].
The Botryosphaeriaceae family belongs to the class Dothideomycetes and the order Botryosphaeriales. It encompasses a range of morphologically diverse fungi that can be pathogens, endophytes, or saprobes, primarily on woody hosts [37]. This family was introduced by Theissen & Sydow in 1918 as a sub-family of Pseudosphaeriaceae. These species spread through conidia [38], with conidiomata (pycnidia), the fungi’s fruiting bodies, releasing conidia during rain and high humidity [39]. Fungi from the Botryosphaeriaceae family are also found in healthy tissue parts of the plant and usually cause diseases after the plant is exposed to stressful conditions (post-harvest and heavy rain). These species can remain in a latent stage until favourable conditions for development arise (biotic and/or abiotic stress) [40]. Species from this family are considered aggressive pathogens. According to Schoch et al. [41], 33 genera with over 1200 species of fungi in this family have been identified. The MycoBank database currently encompass 85 genera of fungi in this family [42].
The overall objectives of this study were to identify the causative agent responsible for symptoms observed in olive trees in Istria, Croatia; to conduct morphological characterisation and molecular identification of the fungal isolates through PCR and DNA sequencing of the ITS, TUB2, and TEF1-α gene regions; to evaluate the pathogenicity of fungal isolates via pathogenicity tests; and to investigate the resistance of various olive varieties to the identified fungal species.

2. Results

2.1. Field Symptoms

Symptoms observed during field research included branch and twig dieback, leaf wilting and defoliation, fruit rot, bark cracking, reddish-brown discolouration of bark, and the appearance of dark-brown necrotic lesions. Necrotic lesions were particularly evident in cross-sectional branch cuts (Figure 1).
Out of the 26 locations visited in total, species from the Botryosphaeriaceae family were identified in 10, representing 38.46% of the total. Among the 112 sampled trees, species from the Botryosphaeriaceae family were identified in 13, making up 11.6% of the total count. The species B. dothidea and N. parvum were found at four locations in Istria, while the species Do. sarmentorum was found at two locations. The species D. mutila, D. seriata, and Do. iberica were each found at one location. The highest number of isolates was collected in Vodnjan (5), followed by Rovinj (4), Poreč (3), and Novigrad (1).
None of the olive groves were equipped with irrigation systems. Pruning was carried out manually at every location, except for grove R18, where a combination of manual and mechanical methods was utilised. Moreover, pruning and burning of plant residues were standard practices across these olive groves. More information about the agricultural practices implemented in olive groves is presented in Table 1.

2.2. Morphological Characterisation

2.2.1. Botryosphaeria Dothidea

The colonies expanded to a diameter of nine cm within five days at 25 °C on potato dextrose agar (PDA) and after seven days on water agar (WA). On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour in the initial stages of growth was white-grey, gradually turning olive green to grey with white tips. As the colony aged, it developed a black-grey colour. The mycelium was dense and matte, with a woolly, cottony, and fluffy appearance (Figure 2). The reverse colonies were white to olivaceous green with dark grey spots. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy, olivaceous green with white tips, appearing individually or in clusters. Conidia were fusiform, thicker in the middle and thinner at the ends, aseptate, and hyaline to brownish. The dimensions of the conidia are shown in Table 2.

2.2.2. Diplodia Mutila

The colonies expanded to a diameter of nine cm within four days at 25 °C on PDA and after seven days on WA. On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour was olivaceous green. As the colony aged, it developed a black-grey colour. The mycelium was dense and matte, with a woolly, cottony, and fluffy appearance (Figure 3). The reverse colonies were white with black spots. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy and white, appearing individually or in clusters. Conidia were capsule shaped, hyaline, and aseptate, and over time changed from beige to dark brown, with one septum. The dimensions of the conidia are shown in Table 2.

2.2.3. Diplodia Seriata

The colonies expanded to a diameter of nine cm within four days at 25 °C on PDA and after seven days on WA. On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour was white-grey. As the colony aged, it developed a dark grey colour. The mycelium was dense and matte, with a woolly, cottony, and fluffy appearance (Figure 4). In certain areas, fluffy, whitish clumps of mycelium formed. The reverse colonies were dark grey. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy and brown with white tips, appearing in clusters. The conidia were ovoid, rounded or slightly pointed at the edges, aseptate, and hyaline, and over time turned dark brown. The dimensions of the conidia are shown in Table 2.

2.2.4. Dothiorella Iberica

The colonies expanded to a diameter of nine cm within four days at 25 °C on PDA and after eight days on WA. On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour was dark grey. The mycelium was dense and matte, with a cottony appearance (Figure 5). The aerial mycelium was more adherent to the substrate, forming rounded cushions, and was not as fluffy as in the previously mentioned species. The reverse colonies were dark grey. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy and grey, appearing in clusters. Conidia were capsule shaped, rounded at the edges, and hyaline, and over time turned light brown, with one septum. The dimensions of the conidia are shown in Table 2.

2.2.5. Dothiorella Sarmentorum

The colonies expanded to a diameter of nine cm within three days at 25 °C on PDA and after eight days on WA. On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour was dark grey with a white coating and brown edges. The mycelium was dense and matte, with a cottony appearance (Figure 6). The mycelium was adherent to the substrate with little cushion-like protrusions. The reverse colonies were black with brown edges. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy and brown with white tips, appearing in clusters. The conidia were capsule-shaped, rounded at the edges, hyaline, and aseptate, and over time turned light brown, with one septum. The dimensions of the conidia are shown in Table 2.

2.2.6. Neofusicoccum Parvum

The colonies expanded to a diameter of nine cm within four days at 25 °C on PDA and after six days on WA. On WA, the mycelium was poorly developed, ranging in colour from white to greyish. On PDA, the colony colour varied among isolates, ranging from light grey to brownish grey, and up to dark grey. The mycelium was dense and matte, with a cottony appearance (Figure 7). The aerial mycelium was adherent to the substrate. The reverse colonies were dark grey with beige-brown edges. The hyphae were septate, branched, and hyaline. Pycnidia formed on the nutrient medium WA + Pinus. The pycnidia were fluffy and grey with white tips, appearing in clusters. The conidia were ellipsoidal, with a round apex but sometimes slightly pointed at the ends, aseptate, and hyaline. The dimensions of the conidia are shown in Table 2.

2.2.7. Rate of Mycelial Growth

In tests of mycelial growth rate conducted at eight different temperatures, after 48 h, none of the isolates showed growth at 5 °C or 40 °C. At 10 °C, 15 °C, 20 °C, and 25 °C, the species Do. sarmentorum, Do. iberica, and D. seriata exhibited the fastest growth rates. At 25 °C, the highest growth was recorded for all isolates except for the species N. parvum, which had the highest growth at 30 °C (Table 3). According to empirically derived data, a temperature of 25 °C proved optimal for the growth of B. dothidea, D. seriata, D. mutila, Do. iberica, and Do. sarmentorum, while for N. parvum, the optimal temperature was 30 °C. At 30 °C, the growth rate of Do. iberica and Do. sarmentorum drastically decreased. At 35 °C, B. dothidea exhibited the fastest growth, followed by N. parvum, D. seriata, and D. mutila, whereas Do. iberica and Do. sarmentorum did not show any mycelial growth.
According to the empirical mathematical modelling (Table 4), optimal temperatures for species growth ranged between 21.3 °C and 28.1 °C, minimum temperatures between 5.1 °C and 5.4 °C, and maximum temperatures between 31.8 °C and 39.9 °C. As evident from the obtained data, the highest optimal growth temperature was recorded for the species N. parvum, while the lowest optimal growth temperature was recorded for the species Do. sarmentorum.

2.3. Molecular Phylogenetic Identification

Six fungal species were identified as belonging to the Botryosphaeriaceae family. All 39 nucleotide sequences obtained from 13 isolates, analysed via BLAST in this study, exhibited a 100% match for the ITS, TUB2, and TEF1-α gene regions, closely aligning with the species listed in the GenBank database. The sequences generated from this research were added to the GenBank database and are available under the accession numbers indicated in Table 5.
Phylogenetic analysis incorporated 100 nucleotide sequences per tree, with all ambiguous positions removed via the pairwise deletion option. The Biscogniauxia mediterranea isolates Bm04.001 and Bm10.019 were used as outgroups. The final dataset comprised 1189 positions for ITS sequence analysis, 1450 positions for TUB2 sequence analysis, 783 positions for TEF1-α sequence analysis, and 2838 positions for multilocus analysis. The optimal trees are shown in Figure 8, Figure 9, Figure 10 and Figure 11.
Based on the combination of morphological characteristics, sequence analysis using the BLAST system, and phylogenetic analysis, the isolates R8 NP, PL1 NP, N17 BJA3, and R19 F were identified as Botryosphaeria dothidea (Moug. ex Fr.) Ces. & De Not.; isolate IKB9 B2II as Diplodia mutila; isolate V16 K2II as Diplodia seriata De Notaris; isolate V16 BI as Dothiorella iberica A.J.L. Phillips, J. Luque & A. Alves; isolates V12 PEN and R18 PEN1 as Dothiorella sarmentorum (Fr.) A.J.L. Phillips, Alves & Luque; and isolates IMK9 IBVI, V16 K1, R18 B1, and V21 B5I as Neofusicoccum parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips.

2.4. Pathogenicity Test and Evaluation of Variety Resistance

2.4.1. Pathogenicity Test

The first symptom observed on olive seedlings inoculated with species from the Botryosphaeriaceae family was leaf wilting, defoliation, and twig and branch dieback (Figure 12), which appeared after three months. This was most pronounced in seedlings inoculated with B. dothidea and D. mutila, and least pronounced in seedlings inoculated with D. seriata.
Other symptoms that appeared included a change in bark colour to reddish due to branch dieback, followed by the appearance of canker formations, bark cracking (Figure 13), and necrosis (Figure 14). The most aggressive species was N. parvum, with a total average necrotic lesion diameter of 93.45 mm. This was followed by D. mutila, with 33.2 mm; B. dothidea, with 17.8 mm; and Do. sarmentorum, with 11.3 mm. The least aggressive species were Do. iberica, with a total necrotic lesion diameter of 4.46 mm, and D. seriata, with 7.45 mm. For the olives inoculated with pure PDA (control group), there were no observed changes. The fungus that was re-isolated from the infected seedlings matched the originally inoculated species, thus validating Koch’s postulate.

2.4.2. Variety Resistance

The olive variety Buža exhibited the highest resistance to Do. sarmentorum, with a statistically significant difference compared to other species (Table 6). This was followed in descending order by resistance to Do. iberica, D. seriata, and B. dothidea. Conversely, Buža showed the greatest susceptibility to D. mutila and N. parvum. The Istarska bjelica variety demonstrated the greatest resistance to Do. iberica and Do. sarmentorum, followed by D. seriata and B. dothidea. Its highest susceptibility was observed for N. parvum, with a statistically significant difference compared to other species. Istarska bjelica also showed notable susceptibility to D. mutila. The Leccino variety had the highest resistance to Do. iberica and D. seriata. It was most susceptible to B. dothidea, followed by Do. sarmentorum and N. parvum, with significant susceptibility also noted for D. mutila. The Rosinjola variety exhibited the greatest resistance to Do. sarmentorum and Do. iberica, followed by D. seriata and B. dothidea. It displayed significant susceptibility to N. parvum and D. mutila.
When evaluating resistance by fungus, for B. dothidea, Buža demonstrated the highest resistance (9.30 ± 3.69), followed by Rosinjola (12.87 ± 6.26) and Istarska bjelica (31.75 ± 11.88), while Leccino was the most susceptible (65.41 ± 17.82). Similarly, for D. mutila, Buža demonstrated the highest resistance (19.16 ± 6.27), followed by Rosinjola (22.25 ± 6.36) and Leccino (44.95 ± 20.75), with Istarska bjelica being the most susceptible (84.33 ± 33.28). For D. seriata, Leccino demonstrated the highest resistance (5.10 ± 2.32), followed by Buža (5.50 ± 1.85) and Rosinjola (10.05 ± 5.41), with Istarska bjelica showing moderate susceptibility (11.15 ± 5.25). In the case of Do. iberica, Istarska bjelica was the most resistant (3.65 ± 1.93), followed by Leccino (3.75 ± 2.02) and Rosinjola (4.95 ± 2.85), while Buža was the most susceptible (5.50 ± 1.03). For Do. sarmentorum, Istarska bjelica also exhibited the highest resistance (4.55 ± 1.42), followed by Buža (4.70 ± 1.92) and Rosinjola (4.85 ± 0.75), with Leccino being significantly more susceptible (53.99 ± 20.04). For N. parvum, Buža again showed the greatest resistance (16.15 ± 5.32), followed by Rosinjola (24.11 ± 7.87) and Leccino (48.45 ± 18.69), whereas Istarska bjelica was highly susceptible (320.75 ± 87.39). This range of susceptibility highlights the varying resistance levels among different olive varieties to specific pathogenic fungi (Figure 15 and Figure 16).

3. Discussion

In the past, species within the Botryosphaeriaceae family were identified mainly by their ascospores. However, relying solely on the sexual state for classification is inadequate, particularly because some species are known only in their asexual state, and in others, the sexual state is exceedingly rare. On the other hand, conidia of the Botryosphaeriaceae display great variation between genera and species [37]. Two types of conidia appear: thin-walled, narrow or spindle-shaped (fusicoccum-like), and thick-walled, broader (diplodia-like) conidia [37]. D. mutila, D. seriata, Do. iberica, and Do. sarmentorum belong to the first group, as they have diplodia-like conidia, while B. dothidea and N. parvum have fusicoccum-like conidia. In this study, isolates were morphologically characterised based on the appearance of mycelium and rate of mycelial growth, as well as the appearance and dimensions of spores, and the appearance of pycnidia. These morphological characteristics aligned with those reported for species in the relevant literature [37]. According to empirical mathematical modelling, the optimal temperatures for species growth varied depending on the species and ranged between 21.3 °C and 28.1 °C. The minimum temperatures ranged between 5.1 °C and 5.4 °C, and the maximum temperatures between 31.8 °C and 39.9 °C. N. parvum exhibited the highest values of optimal growth temperature, while Do. sarmentorum exhibited the lowest. Hernandez-Rodriguez et al. [32] examined the growth rates of various Botryosphaeriaceae species at six temperatures (10, 15, 20, 25, 30, and 35 °C). The optimal temperature was found to be 25 °C. All isolates grew at all temperatures evaluated between 10 and 30 °C. In this study, the isolates also grew at 35 °C, but none of the isolates grew at 5 °C or 40 °C. Kaliterna [43] tested the growth rates of B. dothidea, D. seriata, Do. sarmentorum, and N. parvum. Small deviations in temperature values estimated by the empirical mathematical modelling were recorded between the mentioned study and this study. Additionally, in the mentioned study, no mycelial growth was recorded at 5 °C or 40 °C, except for B. dothidea at 40 °C, which ranged between 1 and 4 mm. As the author notes, the exact reason for the difference in cardinal temperatures for mycelial growth, as well as mycelial growth rates, is not fully understood, likely due to intraspecific variability among isolates.
Phillips et al. [37] consider morphological characteristics alone to be inadequate for defining genera or identifying species, given the confusion it has caused in the past, their variation during development, and inevitable overlap as representation grows. The most accurate identification of fungi is achieved through a combination of morphological characteristics of fungi and molecular methods. As a standard for the molecular identification of fungi, the ITS region of the genome is commonly used [44]. However, in recent research, additional genes such as TUB2, TEF1-α, CALM, ACT, and COI are also utilised [44]. As highlighted by Kaliterna [43], a drawback of such research is its high cost. In studies of species from the Botryosphaeriaceae family, identification is most commonly performed based on the ITS, TUB2, and TEF1-α regions of the genome [32,36,37,43]. In this study, species were molecularly identified based on the ITS, TUB2, and TEF1-α regions of the genome, as well as utilising four phylogenetic trees derived from these gene regions. A total of six species were identified on olive trees in Croatia, namely, B. dothidea, D. mutila, D. seriata, Do. iberica, Do. sarmentorum, and N. parvum.
As pathogens on olives, the following species from the Botryosphaeriaceae family have been identified globally: B. dothidea; B. wangensis G.Q. Li & S.F. Chen; D. africana Damm & Crous; D. fraxini Fries; D. mutila; D. olivarum; D. seriata; D. subglobosa A.J.L. Phillips, Deidda & Linald; L. theobromae (Pat.) Griffon & Maubl.; Do. iberica; Do. omnivora Linaldeddu, Deidda & Scanu; Do. sarmentorum; Do. sempervirentis Abdollahz., Zare & A.J.L. Phillips; N. australe; N. cryptoaustrale Pavlic, Maleme, Slippers & M.J. Wingf.; N. luteum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips; N. mediterraneum; N. occulatum Sakalidis & T. Burgess; N. parvum; N. ribis; N. vitifusiforme; and Sardiniella urbana Linaldeddu, A. Alves & A.J.L. Phillips [16,32,35,36,45,46,47,48]. In Croatia, three species from the Botryosphaeriaceae family have been identified as pathogens on olive trees: B. dothidea, D. seriata, and N. parvum [15,16,17,18]. It is known that species from this family attack numerous plant species, predominantly woody ones, such as grapevine (V. vinifera) [49], European beech (Fagus sylvatica L.) [50], plum (Prunus salicina Lindl.) [51], etc. In Croatia, besides on olive trees, they have also been found on grapevine (V. vinifera) [43], walnut (Juglans sp.) [52], and giant sequoia (Sequoiadendron giganteum (Lindl.) J. Buchholz) [53]. On grapevines, the species B. dothidea, D. coryli, D. seriata, Do. sarmentorum, and N. parvum have been identified [43]. On walnut trees, B. dothidea and N. parvum have also been identified, while in giant sequoia, B. dothidea and N. yunnanense G.Q. Li & S.F. Chen have been recorded [52,53].
Various other fungi described in our previous studies were also identified at some locations. In grove R18 (Table 1), the fungal species B. mediterranea [19], N. philosophiae-doctoris [22], and P. iranianum [23] were identified, along with Do. sarmentorum and N. parvum detected in this study. In grove V16, fungi including B. nummularia (Bull.) Kuntze [19] and C. pruinosa [21] were identified, alongside D. seriata, Do. iberica, and N. parvum. In grove IMK9, the fungi B. mediterranea [19] and N. parvum were identified, while in grove N17 and R19, the species B. mediterranea [19] and B. dothidea were found. Mutual infection was not observed on all sampled trees. In this research, the species B. dothidea and N. parvum were the most frequently found Botryosphaeriaceae species on olive trees. In the study by Linaldeddu et al. [48], fungi from the Botryosphaeriaceae family were the main species isolated from olive trees with branch cankers and fruit rots. Similarly, Hernandez-Rodriguez et al. [32] predominantly isolated species from the genera Botryosphaeria (41%) and Neofusicoccum (51%) from olives, with Diplodia being less common (8%). In the study by Lazzizera et al. [33], Botryosphaeria and Neofusicoccum species were isolated from over 60% of the affected drupes, indicating they are the primary contributors to the disease. The most frequently isolated species was B. dothidea, found in 34% of the drupes. B. dothidea ranks among the most globally prevalent species [32]. The prevalence and distribution of these species may be influenced by climatic and geographical factors, which could be particularly pronounced in Croatia, as noted by Kaliterna [43]. When the climate conditions are optimal, phytopathogenic fungus can grow exponentially and devastate crops, which can cause high economic losses [54]. Humans and animals can also suffer the consequences of fungi attacks because of the mycotoxins produced by some fungi.
Most fungal species responsible for causing fruit rot in olives are typically common saprophytes or secondary invaders that usually enter through wounds inflicted by biotic or abiotic factors. Fungi of the Botryosphaeriaceae family have been recognised as some of the most aggressive and prevalent pathogens impacting olives in olive cultivation areas like California, Italy, South Africa, and others [16,32,35,36]. Different fungi can attack different plant organs, so fungal infections cause an enormous range of disease symptoms, such as colour and shape changes, rotting, wilting, and wounds. Cell death causes parts of the plant to decompose and turns plant tissues into a dark colour; this can appear as spots on leaves or rotten spots on fruits [54]. According to the literature, species from the Botryosphaeriaceae family cause symptoms that include fruit rot, leaf wilting and defoliation, branch and twig dieback, canker formation, and necrosis of internal tissue, as well as discolouration of the bark, among others [16,32,35,36]. These symptoms were also observed in this research, both in the field and on olive seedlings following pathogenicity tests.
In research conducted by Moral et al. [31], pathogenicity tests using unripe olive fruit and olive branches showed that D. seriata isolates were the least aggressive on both the fruit and branches, while N. mediterraneum isolates were the most aggressive in both tissues. Isolates of B. dothidea were not pathogenic on branches and only weakly aggressive on fruit. The most aggressive species in our research were N. parvum, D. mutila, and B. dothidea, whereas the least aggressive species was Do. iberica. As stated by Hernandez-Rodriguez et al. [32] in their studies, the Neofusicoccum isolates were also considerably more aggressive than the Botryosphaeria and Diplodia isolates. The same case was recorded in the pathogenicity tests in the study by Linaldeddu et al. [48], where N. parvum caused much larger lesions than species B. dothidea, D. fraxini, D. mutila, D. olivarum, and D. subglobulosa. Contrary to that, in the study by Godena et al. [17], D. seriata proved to be more aggressive than N. parvum.
The disease resistance of olive varieties offers an economically feasible alternative to chemical control, with minimal environmental impact, and can be integrated into pest management strategies [46]. Theophrastus [2] observed that the Greeks preferred to propagate olives through cuttings, knowing from experience that olives grown from seeds were inferior, thus requiring improvement through grafting and thereby creating more resistant trees. As a measure against B. dothidea, the use of varieties resistant to olive fruit fly attack may serve (since the parasite of the fly, L. berlesiana, cannot penetrate the fruit on its own), along with monitoring the fly occurrence using traps with attractants [46]. Varieties identified as more susceptible to olive fruit fly infestation include St. Catarina and Ascolara Tenera, while varieties such as Dužica [7], Nocellara etnea, Oliva di Cerignola, Orbetana, and Capolga have shown resistance [4]. As a preventive measure, varieties more resistant to disease-causing agents can be planted. Latinović et al. [30] tested the resistance of 17 olive varieties to B. dothidea. The most resistant were Crnjaka and Gloginja, along with Pendolino and Cassanesse. Moderately resistant varieties included Picholine, Grossa di Spagna, and Conserviola, while Rogganiella, Lumbardeška, Sant Agostino, Manzanilla, and Noccelara del Belice were rated as intermediate. Leccino, Coratina, and Žutica showed moderate susceptibility, while Giarraffa and Ascolana tenera were highly susceptible. Moral et al. [46] tested the resistance of the 11 most important table varieties to N. mediterraneum and B. dothidea. Testing results on branches showed that Gordal Sevillana, followed by Santa Caterina and San Agostino, were the most susceptible to N. parvum. Manzanilla Cacereña was the most resistant, followed by Verdial de Huévar and Morona. Concerning potted plant inoculation with N. mediterraneum, Manzanilla Cacereña and Gordal Sevillana were the most susceptible, while the most resistant were Verdial de Huévar, Hojiblanca, and Aloreña de Atarfe. The results of testing resistance on olive fruit showed that the most resistant varieties to B. dothidea were San Agostino and Hojiblanca, while Aloreña de Atarfe was the most susceptible. According to the authors, detached branches may experience significant stress and may not behave physiologically in the same manner as branches attached to a tree. Therefore, results obtained from detached branches may not be entirely indicative of varieties’ resistance. In this study, the susceptibility of varieties varied depending on the fungal species with which the olive seedlings were inoculated. When evaluating resistance by fungus, for B. dothidea, Buža demonstrated the highest resistance, while Leccino was the most susceptible. For D. mutila, Buža showed the highest resistance, whereas Istarska bjelica was the most susceptible. In the case of D. seriata, Leccino exhibited the highest resistance, with Istarska bjelica being most susceptible. For Do. iberica and Do. sarmentorum, Istarska bjelica showed the highest resistance, while Buža and Leccino were the most susceptible, respectively. For N. parvum, Buža had the highest resistance, while Istarska bjelica was highly susceptible. An important factor is not only the resistance of varieties but also the timing of fruit ripening, i.e., the occurrence of apparent resistance. The fruits of most varieties ripen between November and February, and the moment they must be harvested depends on the olive tree’s location, exposure, and meteorological conditions [4]. Early-ripening varieties harvested in late September and October avoid the most intense periods of olive fruit fly attacks [7]. Therefore, when establishing an olive grove, it is advisable to arrange varieties in separate rows to facilitate harvest segregation based on genotype and relative ripening period [4]. As noted by Latinović et al. [30], the identification of resistant varieties could represent the fundamental element of cost-effective disease management, especially for numerous small-scale growers unable to afford pesticide spraying for large olive trees.
Besides resistant varieties, disease control strategies include the removal of infected plant parts [55]. Pruning should be conducted during dry weather with clean and disinfected tools, and coating wounds after pruning is also crucial. Pitt et al. [56] and Díaz and Latorre [57] report that treating wounds with fungicides and horticultural wax helps reduce the incidence of Botryosphaeria dieback. However, it is important to carefully select the fungicide to be applied, as in vitro studies have shown that not all fungicides are equally effective against all species of this family [56,58]. Among the most effective fungicides are those containing the active ingredients benomyl, carbendazim, fluazinam, flusilazole, fludioxonil, iprodione, myclobutanil, penconazole, procymidone, pyraclostrobin, thiophanate–methyl, and tebuconazole [55,56]. When establishing an olive grove, it is also essential to consider the choice of location.

4. Materials and Methods

4.1. Fieldwork and Isolation of Fungi

As part of the olive disease research conducted in Istria County in 2021 and 2022, a total of 26 locations were surveyed, and samples were collected from a total of 112 olive trees. The Botryosphaeria dieback of olive was confirmed at 10 of these locations (Figure 17), i.e., the disease was observed on a total of 13 trees.
Information regarding the precise location and coordinates of the site where species from the family Botryosphaeriaceae were found, including the variety from which the sample was taken, collection date, olive grove area, and tree age, are presented in Table 7.
The trees exhibited symptoms such as leaf wilting and defoliation, twig and branch dieback, and the appearance of necrosis and cankers. A total of 10 branch samples per tree were collected. Samples were taken from the parts of the branches where the transition between healthy and infected parts were visibly apparent. The samples were placed in sterile black plastic bags, labelled, and stored in a portable refrigerator at a temperature of +4 °C. The collected samples were promptly transported to the Laboratory for Plant Protection at the Institute of Agriculture and Tourism in Poreč, Croatia, for analysis. Branch samples from affected trees were photographed and documented and then underwent a washing under tap water. With the use of a sterile surgical scalpel, the bark was removed from the branches, and subsequently, the samples were cut using fruit shears. The branch pieces (5 × 5 cm) were immersed in 70% ethanol for two minutes, followed by rinses in sterile distilled water for two minutes. After this process, they were carefully arranged on a sterile paper sheet within a laminar flow cabinet to facilitate surface drying. Once adequately dried, the pieces were placed on PDA supplemented with 35 mg/L of penicillin and incubated in a dark environment at 25 °C within an incubator. Upon the development of the culture on PDA, isolates were transferred to a medium containing WA and pine needles (Pinus L.) (WA + Pinus). The medium preparation involved cutting fresh green pine needles, washing them under tap water, and autoclaving them twice in a glass jar at 121 °C for 15 min. Two to three pieces of pine needles were placed on the surface of WA after the agar had solidified by 50%, submerging half of the needle in the medium while leaving the rest exposed. Fungal isolates were then inoculated into the nutrient medium WA + Pinus. After an incubation period of 20 days and the subsequent development of pycnidia, spores were extracted to create single-spore isolates. Pure cultures were preserved in 2 mL cryovial screw cap tubes containing a 50% glycerol solution at temperatures of −20 °C and −80 °C, as well as in sterilised water in plastic tubes at 4 °C. The preserved cultures are kept in the Laboratory for Plant Protection collection at the Institute of Agriculture and Tourism in Poreč.

4.2. Morphological Characterisation

Following incubation at 25 °C in the absence of light for 2, 15, and 30 days, pure fungal cultures were subjected to examination. The preliminary determination involved an inspection of colonies, considering their overall appearance and colour. Additionally, the observation of conidia included an assessment of colour, appearance, septation, and shape. Out of the 10 samples collected per tree (totalling 13 trees), the identical fungus was consistently isolated. For a detailed analysis of morphological characteristics, one representative isolate per tree was selected, and a total of 13 fungal isolates were analysed. This involved a detailed evaluation of colony traits, encompassing parameters such as colour, shape, elevation, margin, surface, and opacity. Macroscopic characteristics were observed using a BOECO zoom stereo microscope BSZ-405 and photographed with a fitted B-CAM16 industrial digital camera and B-View software (Boeckel, Hamburg, Germany) at the Institute of Agriculture and Tourism Poreč, as well as with an Olympus SZX10 microscope and Olympus N547 camera (Olympus, Tokyo, Japan) at the Faculty of Agrobiotechnical Sciences Osijek. Additionally, the determination of growth rate and cardinal temperatures for mycelial growth was performed. To determine the cardinal temperatures for growth, five-day-old isolates were inoculated in triplicate on PDA in Petri dishes with a diameter of 90 mm. A circular section of mycelium with a diameter of five mm was placed at the centre of the PDA. The isolates were incubated in darkness at eight different temperatures (5, 10, 15, 20, 25, 30, 35, and 40 °C), and measurements were taken after 48 h. Colony diameter was measured at two positions perpendicular to each other, and the obtained values were reduced by the diameter of the circular section of the mycelium. The average values were then calculated. Empirical mathematical modelling, employing the least squares method, as described by Sanchez et al. [59], was conducted within the suitable frameworks of third, fourth, fifth, and sixth-degree polynomial regressions using Microsoft Office Excel. This approach was used to generate graphs along with corresponding equations, serving to determine cardinal temperatures. Since the sixth-degree polynomial equation fits the examined data best and closely follows the points of determined growth rates, it was utilised for further determination of cardinal temperatures. In order to obtain the minimum, maximum, and optimal cardinal temperatures from the equation, the described empirical mathematical modelling using the least squares method was performed with the Wolfram Alpha (WolframAlpha LLC) mathematical program.
Furthermore, the features of spores, including characteristics such as colour, shape, the presence or absence of septum, and dimensional measurements, were examined. To determine the morphological characteristics of the conidia, they were extracted from pycnidia cut with a laboratory needle. Microscopic analysis was conducted using an LABOSGEN camera and LABOSGEN software at the Institute of Agriculture and Tourism Poreč, as well as with an Olympus BX41 microscope (Olympus, Tokyo, Japan) at the Faculty of Agrobiotechnical Sciences Osijek. Additionally, measurements of 30 conidia per isolate were performed. Subsequently, mean values, standard deviations of measured conidia and their minimum and maximum values, and the length-to-width ratio of conidia were calculated. Furthermore, 95% confidence intervals for the determined mean dimensions and length-to-width ratio were established using Microsoft Office Excel. The morphological profile derived from this analysis was systematically compared with relevant scientific literature sources [37].

4.3. DNA Extraction and Amplification

Molecular analysis was used to confirm the identification of all 13 isolates at the species level. Fungal isolates were cultured on PDA for seven days at 25 °C in the dark. Subsequently, a small portion of mycelium from the colony margins was aseptically sampled using a sterile laboratory needle for genomic DNA extraction. Maxwell® RSC Instrument (Promega, Madison, WI, USA) and Maxwell® RSC Plant DNA Kit (Promega, Madison, WI, USA) were used to extract total genomic DNA. The amount of genomic DNA in samples post-isolation was quantified using a Maxwell Promega Quantus fluorometer (Promega). The internal transcribed spacer (ITS) regions were subjected to amplification and subsequent sequencing using the primer pairs ITS1 (5′ TCCGTAGGTGAACCTGCGG 3′) and ITS4 (5′ TCCTCCGCTTATTGATATGC 3′) [60]. Amplification of a segment of the beta-tubulin (TUB2) gene was carried out utilising the oligonucleotide primers Bt2a (5′ GGTAACCAAATCGGTGCTGCTTTC 3′) and Bt2b (5′ ACCCTCAGTGTAGTGACCCTTGGC 3′) [61]. Furthermore, a part of the translation elongation factor 1-alpha gene (TEF1-α) was amplified and sequenced using the primer pairs EF-728F (5′ CATCGAGAAGTTCGAGAAGG 3′) and EF1-986R (5′ TACTTGAAGGAACCCTTACC 3′) [62]. Each PCR mixture, with a final volume of 25 µL, was composed of 12.5 µL of EmeraldAmp® GT PCR Master Mix, 0.5 µL of each primer (10 µM), 6.5 µL of nuclease-free water, and 5 µL of template DNA at a concentration of 5 ng/µL. PCR amplification was performed using a SureCycler 8800 Thermal Cycler (Agilent Technologies, Santa Clara, CA, USA). The amplification program comprised an initial denaturation step at 94 °C for two minutes, followed by 40 cycles of denaturation at 94 °C for 30 s, annealing at 55 °C for 45 s, elongation at 72 °C for one minute and 30 s, and a final extension step at 72 °C for five minutes [63]. Gel electrophoresis was performed utilising a 1% agarose gel at 110 V for 25 min in 1× TAE buffer, employing an omniPAC Midi CS-300V electropshoresis power supply (Cleaver Scientific, Rugby, Warwickshire, UK). Following electrophoresis, visualisation of PCR products was accomplished using an iBright CL1000 Imaging System (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA). After visualisation, purification of the PCR products was carried out using the GenElute™ PCR Clean-Up Kit (Sigma-Aldrich, Burlington, MA, USA).

4.4. DNA Sequence Assembly and Phylogenetic Analysis

Sequencing of the PCR products was carried out by the Macrogen Europe sequencing service (Amsterdam, the Netherlands). Sequencing was performed bidirectionally using the same primers that were used for amplification. Subsequently, nucleotide sequences were read and edited using Sequencher® software (Gene Codes Corporation, Ann Arbor, MI, USA). Comparative analysis was conducted against existing sequences from the Botryosphaeriaceae family available in the National Center for Biotechnology Information GenBank database. Consensus sequences resulting from this study were submitted to NCBI GenBank. Phylogenetic analyses were conducted for individual gene regions as well as for combined regions. Sequence data from isolates used in this study and relevant isolates from GenBank were utilised for phylogenetic analysis. The list of species/isolates included in the phylogenetic analysis is represented in Table 8. Sequence alignment was performed using ClustalX2 software (UCD, Dublin, Ireland) using the following multiple alignment parameters: gap opening = 15, gap extension = 6.66, delay divergent sequences = 30%, DNA transition weight = 0.5. The evolutionary history was deduced utilising the neighbour–joining method [64], with the optimal tree depicted. Bootstrap analysis (1000 replicates) indicated the percentage of replicate trees in which the associated taxa clustered together, shown next to the branches [65]. The evolutionary distances were calculated using the maximum composite likelihood method and are presented as the number of base substitutions per site [66]. MEGA11 (Pennsylvania State University, State College, PA, USA) was employed for the evolutionary analyses [67].

4.5. Pathogenicity Test and the Evaluation of Variety Resistance

For confirmation of species pathogenicity and evaluation of variety resistance to identified species, a greenhouse experiment was set up using olive seedlings. One representative isolate of each Botryosphaeriaceae species (a total of six isolates) identified in this study was selected for pathogenicity testing. Since multiple isolates were collected for certain species while only one for others, isolates with similar growth rates on PDA were chosen. The experiment utilised five-year-old seedlings of three Croatian indigenous olive varieties: Buža, Istarska bjelica, and Rosinjola, as well as one introduced variety: Leccino. The bark at the intended inoculation site was wiped with cotton soaked in 70% ethanol. Wounds measuring five mm in diameter were then created using a sterile cork borer. The outer bark was removed while preserving the inner bark. A 5 mm-diameter mycelium plug from a 10-day-old colony on PDA was inserted into the wound using a sterile cork borer. Inoculated wounds were coated with Vaseline and covered with Parafilm. Pure PDA plugs served as controls. Ten seedlings were inoculated per isolate. The inoculated plants were grown in the greenhouse for nine months, from December 2022 to October 2023. The seedlings were irrigated using a drip irrigation system. The average temperature during the experimental period in the greenhouse ranged between 24 and 25 °C, with a relative humidity of 85%. Changes were recorded over time. After the incubation period, samples were collected in black plastic bags, and the total length of surface necrotic changes above and below the inoculation site was measured. In accordance with Koch’s postulates, small necrotic tissue fragments from the periphery of lesions that had developed on each seedling were inoculated onto PDA medium to isolate the originally introduced fungus. The data obtained from the pathogenicity assay underwent analysis of variance (ANOVA), followed by Tukey’s test to identify significant differences between mean values at a significance level of 5% [43]. Statistical analysis was conducted using the SAS Enterprise Guide 8.4 statistical software.

5. Conclusions

In conclusion, six different species have been identified as causative agents of Botryosphaeria dieback of olive in Istria: Botryosphaeria dothidea, Diplodia mutila, D. seriata, Dothiorella iberica, Do. sarmentorum, and Neofusicoccum parvum. To our knowledge, D. mutila, Do. iberica, and Do. sarmentorum have not been previously identified in olive trees in Croatia, making this the first report of their presence. Species from the Botryosphaeriaceae family are economically significant pathogens due to their detrimental impact on olive trees, causing fruit rot, leaf wilting, necrosis, and other symptoms. They rank among the most aggressive pathogens attacking olive trees. Therefore, it is necessary to monitor olive groves and track the further movement of these pathogens to minimise the damage they cause. Preventive measures are of utmost importance in controlling the further spread of these pathogens. These measures include disinfection of tools, pruning of olive trees and burning of residues, selection of planting locations, selection of resistant varieties, etc. The varieties tested in this study showed differences in resistance depending on the fungus with which they were infected. In addition to preventive measures, it is important to protect olive groves by using preparations that have proven effectiveness against these pathogens.

Author Contributions

Conceptualisation, E.P. and S.G.; methodology, E.P., K.V. and A.B.V.; investigation, E.P., S.G., K.V. and J.Ć.; writing—original draft preparation, E.P.; writing—review and editing, S.G., K.V., A.B.V. and J.Ć. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Croatian Science Foundation Installation Research Project “Natural bioactive compounds as a source of potential antimicrobial agents in the control of bacterial and other fungal pathogens of olives”, Anti-Mikrobi-OL (AMO), UIP-2020-02-7413, and the “Young Researchers’ Career Development Project” DOK-2021-02-2882.

Data Availability Statement

All sequence data are available in NCBI GenBank in accordance with the accession numbers in the manuscript.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Symptoms of infection observed during field research caused by fungi from the Botryosphaeriaceae family: (a) branch and twig dieback and defoliation caused by the species Botriosphaeria dothidea; (b) reddish-yellow discolouration of bark caused by the species B. dothidea; (c) cracking of twig bark, branch dieback, and leaf wilting caused by the species Diplodia mutila; (d,e) necrotic changes in branch cross-sections: (d) Neofusicoccum parvum, (e) Diplodia seriata; (f,g) cracking of tree bark and the appearance of necrotic lesions: (f) Dothiorella sarmentorum, (g) N. parvum; (h) branch drying and bark discolouration caused by the species Dothiorella iberica; (i) bark cracking and the appearance of necrosis caused by the species B. dothidea.
Figure 1. Symptoms of infection observed during field research caused by fungi from the Botryosphaeriaceae family: (a) branch and twig dieback and defoliation caused by the species Botriosphaeria dothidea; (b) reddish-yellow discolouration of bark caused by the species B. dothidea; (c) cracking of twig bark, branch dieback, and leaf wilting caused by the species Diplodia mutila; (d,e) necrotic changes in branch cross-sections: (d) Neofusicoccum parvum, (e) Diplodia seriata; (f,g) cracking of tree bark and the appearance of necrotic lesions: (f) Dothiorella sarmentorum, (g) N. parvum; (h) branch drying and bark discolouration caused by the species Dothiorella iberica; (i) bark cracking and the appearance of necrosis caused by the species B. dothidea.
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Figure 2. (a,b) Upper and reverse view of Botryospheria dothidea isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) hyphae and conidia of B. dothidea isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 2. (a,b) Upper and reverse view of Botryospheria dothidea isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) hyphae and conidia of B. dothidea isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Figure 3. (a,b) Upper and reverse view of Diplodia mutila isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. mutila isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 3. (a,b) Upper and reverse view of Diplodia mutila isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. mutila isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Figure 4. (a,b) Upper and reverse view of Diplodia seriata isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) hyphae and conidia of D. seriata isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 4. (a,b) Upper and reverse view of Diplodia seriata isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) hyphae and conidia of D. seriata isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Figure 5. (a,b) Upper and reverse view of Dothiorella iberica isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. iberica isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 5. (a,b) Upper and reverse view of Dothiorella iberica isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. iberica isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Figure 6. (a,b) Upper and reverse view of Dothiorella sarmentorum isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. sarmentorum isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 6. (a,b) Upper and reverse view of Dothiorella sarmentorum isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of D. sarmentorum isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Figure 7. (a,b) Upper and reverse view of Neofusicoccum parvum isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of N. parvum isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
Figure 7. (a,b) Upper and reverse view of Neofusicoccum parvum isolate 15 days after incubation at 25 °C on potato dextrose agar (PDA) medium; (c) conidia of N. parvum isolate observed under the microscope, scale bar = 20 μm; (d) pycnidia developed on WA + Pinus.
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Plants 13 01813 g008
Figure 8. The phylogenetic tree, based on the alignment of internal transcribed spacer sequences, highlights the sequences identified in this research with red rectangles.
Figure 8. The phylogenetic tree, based on the alignment of internal transcribed spacer sequences, highlights the sequences identified in this research with red rectangles.
Plants 13 01813 g008
Plants 13 01813 g009
Figure 9. The phylogenetic tree, based on the alignment of beta-tubulin sequences, highlights the sequences identified in this research with red rectangles.
Figure 9. The phylogenetic tree, based on the alignment of beta-tubulin sequences, highlights the sequences identified in this research with red rectangles.
Plants 13 01813 g009
Plants 13 01813 g010
Figure 10. The phylogenetic tree, based on the alignment of translation elongation factor 1-alpha sequences, highlights the sequences identified in this research with red rectangles.
Figure 10. The phylogenetic tree, based on the alignment of translation elongation factor 1-alpha sequences, highlights the sequences identified in this research with red rectangles.
Plants 13 01813 g010
Plants 13 01813 g011
Figure 11. The multilocus phylogenetic tree, based on the alignment of the internal transcribed spacer, beta-tubulin, and translation elongation factor 1-alpha sequences, highlights the sequences identified in this research with red rectangles.
Figure 11. The multilocus phylogenetic tree, based on the alignment of the internal transcribed spacer, beta-tubulin, and translation elongation factor 1-alpha sequences, highlights the sequences identified in this research with red rectangles.
Plants 13 01813 g011
Plants 13 01813 g012
Figure 12. Defoliation and branch dieback symptoms on olive seedlings inoculated with (a) Diplodia mutila, (b) Neofusicoccum parvum, (c) Diplodia seriata, and (d) Botryosphaeria dothidea.
Figure 12. Defoliation and branch dieback symptoms on olive seedlings inoculated with (a) Diplodia mutila, (b) Neofusicoccum parvum, (c) Diplodia seriata, and (d) Botryosphaeria dothidea.
Plants 13 01813 g012
Plants 13 01813 g013
Figure 13. Canker formations, bark splitting and colour change on branches inoculated with the species (a) Botryosphaeria dothidea, (b) Diplodia mutila, (c,d) Neofusicoccum parvum, and (e) Diplodia mutila.
Figure 13. Canker formations, bark splitting and colour change on branches inoculated with the species (a) Botryosphaeria dothidea, (b) Diplodia mutila, (c,d) Neofusicoccum parvum, and (e) Diplodia mutila.
Plants 13 01813 g013
Plants 13 01813 g014
Figure 14. Necrosis observed in the cross-sections of branches caused by (a) Diplodia mutila, (b) Dothiorella iberica, (c) Neofusicoccum parvum, and (d) Botryosphaeria dothidea.
Figure 14. Necrosis observed in the cross-sections of branches caused by (a) Diplodia mutila, (b) Dothiorella iberica, (c) Neofusicoccum parvum, and (d) Botryosphaeria dothidea.
Plants 13 01813 g014
Plants 13 01813 g015
Figure 15. Results of pathogenicity test/variety resistance test. Average values of the length of necrotic changes (mm) per variety are shown in columns of different colours, ranging from light to dark blue. Each column represents the mean value of 10 measurements corresponding to Table 6. The vertical error bars marked in black indicate the standard deviation.
Figure 15. Results of pathogenicity test/variety resistance test. Average values of the length of necrotic changes (mm) per variety are shown in columns of different colours, ranging from light to dark blue. Each column represents the mean value of 10 measurements corresponding to Table 6. The vertical error bars marked in black indicate the standard deviation.
Plants 13 01813 g015
Plants 13 01813 g016
Figure 16. Results of pathogenicity test/variety resistance test: (a) samples collected from all inoculated olive seedlings; (b) differences between control plants and plants inoculated with species from the Botryosphaeriaceae family observed as follows: the white mark indicates control, the red mark indicates Botryosphaeria dothidea, the green mark indicates Diplodia mutila, the yellow mark indicates Diplodia seriata, the blue mark indicates Dothiorella iberica, the black mark indicates Dothiorella sarmentorum, and the yellow–green mark indicates Neofusicoccum parvum; (c–h) symptoms on branches per variety (from left to right: three branches from Buža, three from Istarska bjelica, three from Leccino, and three from Rosinjola): (c) B. dothidea, (d) D. mutila, (e) D. seriata, (f) Do. iberica, (g) Do. sarmentorum, (h) N. parvum.
Figure 16. Results of pathogenicity test/variety resistance test: (a) samples collected from all inoculated olive seedlings; (b) differences between control plants and plants inoculated with species from the Botryosphaeriaceae family observed as follows: the white mark indicates control, the red mark indicates Botryosphaeria dothidea, the green mark indicates Diplodia mutila, the yellow mark indicates Diplodia seriata, the blue mark indicates Dothiorella iberica, the black mark indicates Dothiorella sarmentorum, and the yellow–green mark indicates Neofusicoccum parvum; (c–h) symptoms on branches per variety (from left to right: three branches from Buža, three from Istarska bjelica, three from Leccino, and three from Rosinjola): (c) B. dothidea, (d) D. mutila, (e) D. seriata, (f) Do. iberica, (g) Do. sarmentorum, (h) N. parvum.
Plants 13 01813 g016
Plants 13 01813 g017
Figure 17. Locations of samples of species from the Botryosphaeriaceae family collected from olive trees in Istria, Croatia, are marked with green–white labels, with the isolate names next to the labels. The figure was made using Google Maps.
Figure 17. Locations of samples of species from the Botryosphaeriaceae family collected from olive trees in Istria, Croatia, are marked with green–white labels, with the isolate names next to the labels. The figure was made using Google Maps.
Plants 13 01813 g017
Table 1. Compilation of production methods, tree density, fertilisation practices, cultivation techniques, pre-culture, surrounding vegetation, and harvesting methods in olive groves.
Table 1. Compilation of production methods, tree density, fertilisation practices, cultivation techniques, pre-culture, surrounding vegetation, and harvesting methods in olive groves.
Location *Production Method
**		Tree
Density		Fertilisation ***Cultivation
****		Pre-CultureSurrounding VegetationHarvesting Method
R8O2 × 3NAUCLow shrubberyOlive groveManual
IKB9C6 × 6CANCGrains, vineyards, forage cropsOlive groveHandheld shakers
IMK9C6 × 6CANCGrains, vineyards, forage cropsVineyards, forestHandheld shakers
V12C5 × 5NAUCForest, meadowOlive groveTractor-mounted shakers
V16O6 × 6 and 8 × 8NACFruit treesOlive grove, forestHandheld shakers
N17 C6 × 7 and 5 × 6CANUCLow shrubbery, meadowVineyards, forestManual, handheld shakers
R18C6 × 6CANCVineyardsMeadow, field, forestHandheld shakers
R19C6 × 6StallaticoCVineyardsOlive grove, vineyards, meadow, forestManual, handheld shakers
PL1C10NAUCLow shrubbery, meadowOlive trees located by the sea, surrounded by a forestManual
V21C/I6 × 6, 5 × 7 and 7 × 7Sheep manure, compostC/UCOliveOlive groveHandheld shakers
* Location: The location tags are identical to the first half of the isolate names. ** C—conventional, I—integrated, O—organic. *** NA—not applicable, CAN—calcium ammonium nitrate. **** C—cultivated, UC—uncultivated.
Table 2. Conidial dimensions of fungal isolates.
Table 2. Conidial dimensions of fungal isolates.
SpeciesConidial Size (µm)
Length × Width *Average ± SD **95% Conf. ***
Botryosphaeria dothidea(20.0) 22.9–24.2 (27.3) ± (5.1) 6.0–6.6 (7.8)23.5 ± 1.9 × 6.3 ± 0.80.7–0.3
Diplodia mutila(20.4) 23.5–24.8 (27.4) ± (9.1) 10.8–11.6 (14.2)24.1 ± 1.8 ×11.2 ± 1.10.6–0.4
Diplodia seriata(16.2) 22.1–23.7 (27.6) ± (8.3) 10.7–11.7 (13.9)22.9 ± 2.1 × 11.2 ± 1.30.8–0.5
Dothiorella iberica(15.7) 20.4–21.7 (23.9) ± (7.1) 8.4–9.1 (10.7)21.0 ± 1.8 × 8.7 ± 0.90.7–0.4
Dothiorella
sarmentorum		(15.5) 18.9–20.1 (22.9) ± (7.3) 8.3–8.9 (10.9)19.5 ± 1.7 × 8.6 ± 0.90.6–0.3
Neofusicoccum parvum(9.2) 11.7–12.6 (15.5) ± (3.6) 5.4–6.0 (7.1)12.2 ± 1.3 × 5.7 ± 0.90.5–0.3
* Dimensions are expressed as a range from the lower to the upper limit of the 95% confidence interval, with the minimum and maximum values of the measured dimensions shown in parentheses. ** Mean dimension values (length × width) and standard deviation (SD) are presented. *** The length-to-width ratio of conidia is depicted as a range from the lower to the upper limit of the 95% confidence interval.
Table 3. The average mycelial growth rate (mm) after 48 h, incubated at eight different temperatures.
Table 3. The average mycelial growth rate (mm) after 48 h, incubated at eight different temperatures.
Temperature (°C)Botryosphaeria dothideaDiplodia mutilaDiplodia seriataDothiorella ibericaDothiorella sarmentorumNeofusicoccum parvum
Growth of Mycelium (mm)
5000000
102.585.168.511.018.337.03
158.8712.019.1623.538.010.21
2024.2933.345.3345.6657.028.16
2539.0448.3355.050.3357.4140.95
3030.1731.1644.667.1616.8351.95
3516.471.834.16005.04
40000000
Table 4. The cardinal temperature for mycelium isolate growth derived from empirically determined growth rate values and mathematical modelling outcomes.
Table 4. The cardinal temperature for mycelium isolate growth derived from empirically determined growth rate values and mathematical modelling outcomes.
SpeciesCardinal Temperatures (°C) for Mycelial Growth Based on Empirically Determined Growth Rate Values Cardinal Temperatures (°C) for Mycelial Growth Estimated through Mathematical Modelling
MinimumOptimalMaximumMinimumOptimalMaximum
Botryosphaeria dothidea5–102535–405.125.739.9
Diplodia mutila5–102535–405.124.935.3
Diplodia seriata5–102535–405.125.435.6
Dothiorella iberica5–102530–355.126.931.8
Dothiorella sarmentorum5–102530–355.221.334.3
Neofusicoccum parvum5–103035–405.428.135.5
Table 5. List of accession numbers of isolates deposited in GenBank.
Table 5. List of accession numbers of isolates deposited in GenBank.
IsolateSpeciesGenBank Accession Number
ITSTUB2TEF1-α
R8 NPBotryosphaeria dothideaOQ338370OQ348378OQ348385
PL1 NPBotryosphaeria dothideaOQ352832OQ361692OQ553927
N17 BJA3Botryosphaeria dothideaOQ353073OQ361697OQ553926
R19 FBotryosphaeria dothideaOQ354201OQ361700OQ361701
IKB9 B2IIDiplodia mutilaOQ338569OQ348379OQ348386
V16 K2IIDiplodia seriataOQ352870OQ361695OQ361696
V16 BIDothiorella ibericaOQ339205OQ348381OQ348388
V12 PENDothiorella sarmentorumOQ339150OQ348380OQ348387
R18 PEN1Dothiorella sarmentorumOQ341230OQ348383OQ348390
IMK9 IBVINeofusicoccum parvumOQ352837OQ361693OQ361694
V16 K1Neofusicoccum parvumOQ341191OQ348382OQ348389
R18 B1Neofusicoccum parvumOQ353087OQ361698OQ361699
V21 B5INeofusicoccum parvumOQ341428OQ348384OQ553928
Table 6. The results of the pathogenicity test/variety resistance test with average values of the length of necrotic changes (mean ± standard deviation, in mm) of six isolates of fungi from the Botryosphaeriaceae family on olive seedlings, with sterile PDA as a negative control.
Table 6. The results of the pathogenicity test/variety resistance test with average values of the length of necrotic changes (mean ± standard deviation, in mm) of six isolates of fungi from the Botryosphaeriaceae family on olive seedlings, with sterile PDA as a negative control.
SpeciesVariety *
BužaIstarska BjelicaLeccinoRosinjola
Botryosphaeria
dothidea		9.30 ± 3.69 b31.75 ± 11.88 c65.41 ± 17.82 a12.87 ± 6.26 b
Diplodia
mutila		19.16 ± 6.27 a84.33 ± 33.28 b44.95 ± 20.75 b22.25 ± 6.36 a
Diplodia
seriata		5.50 ± 1.85 b11.15 ± 5.25 c5.10 ± 2.32 c10.05 ± 5.41 bc
Dothiorella
iberica		5.50 ± 1.03 b3.65 ± 1.93 c3.75 ± 2.02 c4.95 ± 2.85 cd
Dothiorella
sarmentorum		4.70 ± 1.92 bc4.55 ± 1.42 c53.99 ± 20.04 ab4.85 ± 0.75 cd
Neofusicoccum
parvum		16.15 ± 5.32 a320.75 ± 87.39 a48.45 ± 18.69 ab24.11 ± 7.87 a
Control0.0 ± 0.0 c0.0 ± 0.0 c0.0 ± 0.0 c0.0 ± 0.0 d
Minimum
significant
difference		4.8748.6219.996.90
* Mean values with the same letters in a row are not significantly different, according to Tukey’s honestly significant difference test (p < 0.05).
Table 7. Isolate label, location, and coordinates of Botryosphaeriaceae species site, olive variety from which the sample was taken, collection date, olive grove area, and tree age.
Table 7. Isolate label, location, and coordinates of Botryosphaeriaceae species site, olive variety from which the sample was taken, collection date, olive grove area, and tree age.
IsolateLocationCoordinatesVariety from Which Sample Was TakenCollection DateOlive Grove Area (Ha)Tree Age (Years)
R8 NPRovinj45°05′20″ N, 13°38′51″ EUnknown6 September 20210.0115
IKB9 B2IIPoreč45°13′19.9″ N, 13°36′07.7″ EBuža13 September 20211.49>30
IMK9 IBVIPoreč45°13′13″ N 13°36′09.8″ EIstarska bjelica13 September 20211.49>30
V12 PENVodnjan44°57′65″ N, 13°50′19″ EPendolino24 September 20210.120–100
V16 BIFažana near
Vodnjan		44°56′21″ N; 13°50′18″ EBuža14 October 2021113
V16 K1Karbonaca
V16 K2IIKarbonaca
N17 BJA3Novigrad45°20′08.8″ N, 13°33′33.6″ EIstarska bjelica14 October 2021320–25
R18 B1Rovinj45°03′02.2″ N 13°42′43.9″ EBuža14 October 20210.4339
R18 PEN1Pendolino
R19 FRovinj45°03′46″ N, 13°42′71″ EFrantoio14 October 20210.3825–30
PL1 NPPoreč45°12′26″ N 13°35′29″ EUnknown23 March 20220.0210
V21 B5IVodnjan44°57′34″ N, 13°50′37″ EBuža24 March 20228.212–300
Table 8. List of species/isolates included in the phylogenetic analysis, detailing their host, country of origin, GenBank accession numbers, and references.
Table 8. List of species/isolates included in the phylogenetic analysis, detailing their host, country of origin, GenBank accession numbers, and references.
SpeciesIsolateHostCountryGenBank Accession NumberReferences
ITSTUB2TEF1-α
Biscogniauxia mediterraneaBm04.001Quercus suber L.PortugalKM216752KM267202KM216788[68]
B. mediterraneaBm10.019Q. suberPortugalKM216761KM267210KM216797[68]
Botryosphaeria corticis (Demaree & Wilcox) Arx & E. Mull.ATCC 22927Vaccinium corymbosum L.USADQ299247 EU673108 EU673291 [33,69]
Bo. corticisCBS119047V. corymbosumUSADQ299245 EU673107 EU017539 [33,69]
Bo. corticisCBS119048V. corymbosumUSADQ299246 MT592464 EU017540 [33,69]
Bo. dothidea (Moug. ex Fr.) Ces. & De Not.CBS110302Vitis vinifera L.PortugalAY259092 EU673106AY573218[70,71,72]
Bo. dothideaCMW8000Prunus sp.SwitzerlandAY236949 AY236927 AY236898 [63]
Bo. dothidea BJS-01sV. viniferaChinaJX275778 JX462259JX462285 [49]
Bo. pseudoramosa G.Q. Li & S.F. ChenCERC 2001Eucalyptus urophylla × Eucalyptus grandisChinaKX277989 KX278198KX278094[73]
Bo. pseudoramosaCERC 3455E. urophylla × E. grandisChinaKX277997 KX278206 KX278102 [73]
Bo. pseudoramosaCERC 3472E. urophylla × E. grandisChinaKX277999 KX278208 KX278104 [73]
Bo. qingyuanensis G.Q. Li & S.F. ChenCERC 2947E. urophylla × E. grandisChinaKX278001 KX278210 KX278106 [73]
Bo. qingyuanensisCERC 2946E. urophylla × E. grandisChinaKX278000 KX278209 KX278105 [73]
Bo. qingyuanensisTKBQ16-30Carya cathayensis Sarg.ChinaMW561677 MW561777 MW561687[74]
Diplodia bulgarica A.J.L. Phillips, J. Lopes & S.G. BobevUCD11351Malus domestica (Suckow) Borkh.USAOR631210 OR637362 OR637364 [75]
D. bulgaricaUCD11350 M. domesticaUSAOR631209 OR637361 OR637363 [75]
D. bulgaricaCBS124136M. sylvestris (L.) Mill.BulgariaMH863355MT592475GQ923822 [76,77,78]
D. corticola A.J.L. Phillips, A. Alves & J. LuqueCBS 112546Q. ilex L.SpainAY259090 EU673117 EU673310[70,72]
D. corticolaCBS 112549Q. suberPortugalAY259100 DQ458853 AY573227[70,71]
D. corticolaUCD1260SoV. viniferaCaliforniaGU799470GU799464 GU799467[79]
D. cupressi A.J.L. Phillips & A. AlvesCBS168.87 Cupressus sempervirens L.IsraelDQ458893 DQ458861 DQ458878 [80]
D. cupressiCBS261.85 C. sempervirensIsraelDQ458894 DQ458862 DQ458879 [80]
D. cupressiCBS121027C. sempervirensCyprusMT587340 MT592487 MT592045 [78]
D. mutilaCBS112553V. viniferaPortugalAY259093MZ073931 AY573219[70,71,81]
D. mutilaCBS230.30 Phoenix dactylifera L.USADQ458886DQ458849 DQ458869[80]
D. mutilaPD75HollyUSAGU251119 GU251779GU251251 [82]
D. seriata De NotarisUCD244MaV. viniferaCaliforniaDQ008314 DQ008337 EU012406[79]
D. seriataCBS 112555V. viniferaPortugalAY259094 DQ458856AY573220[70]
D. seriataSDZ-01 V. viniferaChinaHQ629954 HQ629956 HQ629958 [49]
Dothiorella brevicollis Jami, Gryzenh., Slippers & M.J. Wingf.CMW36463Vachellia karroo (Hayne) Banfi & GalassoSouth AfricaJQ239403 JQ239371 JQ239390 [83]
Do. brevicollisCMW36464V. karrooSouth AfricaJQ239404 JQ239372JQ239391[83]
Do. brevicollisCBS130412V. karrooSouth AfricaMT587395 MT592578 MT592107 [78]
Do. dulcispinae Jami, Gryzenh., Slippers & M.J. Wingf.CMW36460V. karrooSouth AfricaJQ239400 JQ239373 JQ239387 [83]
Do. dulcispinaeCMW36461V. karrooSouth AfricaJQ239401JQ239374 JQ239388 [83]
Do. dulcispinaeCMW36462V. karrooSouth Africa JQ239402JQ239375 JQ239389 [83]
Do. iberica A.J.L. Phillips, J. Luque & A. AlvesCBS115041Q. ilexSpainNR111165 EU673096 AY573222 [72,84]
Do. ibericaUCRDI3Prunus dulcis (Mill.) D.A.WebbCaliforniaKP012591 KP067201 KP828802 [85]
Do. ibericaUCRDI1 P. dulcisCaliforniaKP012590 KP067200 KP828801 [85]
Do. oblonga F.J.J. van der Walt, Slippers & G.J. MaraisCBS130414V. karrooSouth AfricaMT587408 MT592598 MT592120 [78]
Do. oblongaCBS121766Senegalia mellifera (Vahl) L.A. Silva & J. FreitasSouth AfricaEU101301 KX464863 EU101346 [86,87]
Do. oblongaCBS121765S. melliferaSouth AfricaKF766163 KX464862 EU101345 [86,87,88]
Do. sarmentorum (Fr.) A.J.L. Phillips, Alves & LuqueCMW39366Aesculus hippocastanum L.SerbiaKF575009 KF575105 KF575047 [89]
Do. sarmentorumPD78P. dulcisCaliforniaGU251169 GU251829 GU251301 [82]
Do. sarmentorumCBS115038Malus pumila Mill.NetherlandsAY573206 EU673101 AY573223 [71]
Lasiodiplodia crassispora T.I. Burgess & P.A. BarberUCD27CoV. viniferaCaliforniaGU799457 GU799480 GU799488 [90]
L. crassispora CMW13488Eucalyptus urophylla S.T. BlakeVenezuelaDQ103552 KU887507 DQ103559 [91,92]
L. crassisporaUCD24CoV. viniferaCaliforniaGU799456 GU799479 GU799487 [90]
L. gonubiensis Pavlic, Slippers & M.J. Wingf CBS115812Syzygium cordatum Hochst.South AfricaDQ458892 DQ458860DQ458877[80]
L. gonubiensis CMW36240 Adansonia sp.AfricaKU887124 KU887502KU887001 [92]
L. gonubiensisCMW43763Bruguiera gymnorhiza (L.) SavignySouth AfricaKU587955 KU587865KU587944[93]
L. iranensis Abdollahz., Zare & A.J.L. PhillipsPCoCo10Theobroma cacao L.TaiwanOR534188 OR551954 OR552312 [94]
L. iranensisPCoCo11T. cacaoTaiwanOR534189 OR551955OR552313 [94]
L. iranensisPCoCo12 T. cacaoTaiwanOR534190 OR551956 OR552314 [94]
L. pseudotheobromae A.J.L. Phillips, A. Alves & CrousCBS116459Gmelina arborea Roxb. ex Sm.Costa RicaEF622077 EU673111 EF622057 [72,95]
L. pseudotheobromaeMFLUCC 18-1120 Magnolia liliifera (L.) L.ChinaMK496933MK524719MK521585 [96]
L. pseudotheobromaeMFLUCC 18-0950M. liliiferaChinaMK501818 MK550605MK521586 [96]
L. theobromae (Pat.) Griffon & Maubl.CAA006 V. viniferaUSADQ458891 DQ458859 DQ458876[80]
L. theobromaeGX-5-5AV. viniferaChinaJX275780 JX462262 JX462288 [49]
L. theobromaeTJXHS1S1V. viniferaChinaJX275790 JX462278JX462304[49]
N. arbuti (D.F. Farr & M. Elliott) Crous, Slippers & A.J.L. PhillipsCBS116576Arbutus menziesii PurshUSAKX464156KX464928 KX464651[87]
N. arbuti CBS116574A. menziesiiUSAKX464154 KX464926KX464649 [87]
N. arbutiCBS116573A. menziesiiUSAKX464153 KX464925 KX464648[87]
N. australeCMW6837Acacia sp.AustraliaAY339262 AY339254 AY339270 [97]
N. australeCMW9073Acacia sp.AustraliaAY339261 AY339253AY339269 [97]
N. australeCMW6853Sequoiadendron giganteum (Lindl.) J. BuchholzAustraliaAY339263 AY339255 AY339271[97]
N. brasiliense M.W. Marques, A.J.L. Phillips & M.P.S. CamaraCMM1285Mangifera indica L.BrazilJX513628 KC794030 JX513608[98]
N. brasilienseCMM1338M. indicaBrazilJX513630 KC794031 JX513610 [98]
N. brasilienseCMM1269 M. indicaBrazilJX513629 KC794032 JX513609 [98]
N. hongkongense G.Q. Li & S.F. ChenCERC 2967Araucaria cunninghamii MudieChinaKX278050KX278259 KX278155 [73]
N. hongkongenseCERC 2968A. cunninghamiiChinaKX278051 KX278260 KX278156 [73]
N. hongkongenseCERC 2973A. cunninghamiiChinaKX278052 KX278261 KX278157 [73]
N. mangroviorum J.A. Osorio, Jol. Roux & Z.W. de Beer CPC32482Diospyros dichrophylla (Gand.) De WinterSouth AfricaMT587494MT592701 MT592209[78]
N. mangroviorumCMW41365Avicennia marina (Forssk.) Vierh.South AfricaKP860859KP860779KP860702 [93]
N. mangroviorumCMW42481Bruguiera gymnorhiza (L.) Savigny,South AfricaKP860848 KP860770 KP860692 [93]
N. mangiferae (Syd. & P.Syd.) Crous et al.CMW7024 Magnifera indica L.AustraliaAY615185 AY615172DQ093221 [99]
N. mangiferaeCMW7797M. indicaAustraliaAY615186 AY615173 DQ093220[99]
N. mangiferaeCMW7081M. indicaAustraliaAY615187 AY615174 KF766425[99]
N. microconidiumCERC 3497E. urophylla × E. grandisChinaKX278053 KX278262 KX278158 [73]
N. microconidiumCERC 3498E. urophylla × E. grandisChinaKX278054 KX278263 KX278159[73]
N. microconidiumCBS118821Syzygium cordatum Hochst.South AfricaMT587497 MT592704 MT592212 [78]
N. luteum (Pennycook & Samuels) Crous, Slippers & A.J.L. PhillipsCBS110299V. viniferaPortugalAY259091 DQ458848 AY573217 [70,71]
N. luteumCBS110497V. viniferaPortugalEU673311 EU673092 EU673277 [72]
N. luteumCBS133502Persea americana Mill.USAMT587483 MT592689 MT592197 [78]
N. parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. PhillipsCBS110301V. viniferaPortugalAY259098 EU673095 AY573221 [70,71,72]
N. parvumCMW9081Populus nigra L.New Zealand AY236943 AY236917AY236888[97]
N. parvumCBS123652S. cordatumSouth AfricaKX464184 KX464996 KX464710 [87]
N. parvumZai-5Syzygium samarangense Merr. & L.M. PerryTaiwanOR534045 OR551811OR552360 [94]
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MDPI and ACS Style

Petrović, E.; Vrandečić, K.; Belušić Vozila, A.; Ćosić, J.; Godena, S. Diversity and Pathogenicity of Botryosphaeriaceae Species Isolated from Olives in Istria, Croatia, and Evaluation of Varietal Resistance. Plants 2024, 13, 1813. https://doi.org/10.3390/plants13131813

AMA Style

Petrović E, Vrandečić K, Belušić Vozila A, Ćosić J, Godena S. Diversity and Pathogenicity of Botryosphaeriaceae Species Isolated from Olives in Istria, Croatia, and Evaluation of Varietal Resistance. Plants. 2024; 13(13):1813. https://doi.org/10.3390/plants13131813

Chicago/Turabian Style

Petrović, Elena, Karolina Vrandečić, Andreina Belušić Vozila, Jasenka Ćosić, and Sara Godena. 2024. "Diversity and Pathogenicity of Botryosphaeriaceae Species Isolated from Olives in Istria, Croatia, and Evaluation of Varietal Resistance" Plants 13, no. 13: 1813. https://doi.org/10.3390/plants13131813

APA Style

Petrović, E., Vrandečić, K., Belušić Vozila, A., Ćosić, J., & Godena, S. (2024). Diversity and Pathogenicity of Botryosphaeriaceae Species Isolated from Olives in Istria, Croatia, and Evaluation of Varietal Resistance. Plants, 13(13), 1813. https://doi.org/10.3390/plants13131813

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