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Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China

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  • Received: 12 November 2024
    Revised: 05 December 2024
    Accepted: 05 December 2024
    Published online: 20 December 2024
    Studies in Fungi  9 Article number: e016 (2024)  |  Cite this article
  • During the investigation of ascomycetes on Poaceae in Yunnan Province, China, a fungus was collected from dead bamboo culms in a terrestrial habitat in the Ailaoshan subtropical evergreen broad-leaved forest. Based on morphological characterization and phylogenetic analyses, this fungus was introduced as Oxydothis ailaoshanensis sp. nov. Maximum likelihood and Bayesian inference analyses of a concatenated dataset of internal transcribed spacer (ITS), large subunits (LSU) of the nuclear ribosomal RNA gene, small subunit (SSU) of the nuclear ribosomal RNA gene, and translation elongation factor 1-alpha (tef1-α) sequences were performed to clarify the phylogenetic affinities of the new species. Phylogenetically, Oxydothis ailaoshanensis forms an independent lineage, basal to O. metroxylonis. Oxydothis ailaoshanensis differs from O. metroxylonis in having smaller, immersed ascomata that become raised and superficial with the long axis horizontal to the host surface, shorter asci that are mostly straight, and longer ascospores that are elongated and fusiform. The new species was justified based on morphological traits and multigene phylogenetic analyses in comparison with closely related species. A detailed description, micrograph, and a phylogenetic tree of the new species are provided.
  • Starting in the early 2000s, China has experienced rapid growth as an emerging wine market. It has now established itself as the world's second-largest grape-growing country in terms of vineyard surface area. Furthermore, China has also secured its position as the sixth-biggest wine producer globally and the fifth-most significant wine consumer in terms of volume[1]. The Ningxia Hui autonomous region, known for its reputation as the highest quality wine-producing area in China, is considered one of the country's most promising wine regions. The region's arid or semiarid climate, combined with ample sunlight and warmth, thanks to the Yellow River, provides ideal conditions for grape cultivation. Wineries in the Ningxia Hui autonomous region are renowned as the foremost representatives of elite Chinese wineries. All wines produced in this region originate from grapes grown in their vineyards, adhering to strict quality requirements, and have gained a well-deserved international reputation for excellence. Notably, in 2011, Helan Mountain's East Foothill in the Ningxia Hui Autonomous Region received protected geographic indication status in China. Subsequently, in 2012, it became the first provincial wine region in China to be accepted as an official observer by the International Organisation of Vine and Wine (OIV)[2]. The wine produced in the Helan Mountain East Region of Ningxia, China, is one of the first Agricultural and Food Geographical Indications. Starting in 2020, this wine will be protected in the European Union[3].

    Marselan, a hybrid variety of Cabernet Sauvignon and Grenache was introduced to China in 2001 by the French National Institute for Agricultural Research (INRA). Over the last 15 years, Marselan has spread widely across China, in contrast to its lesser cultivation in France. The wines produced from Marselan grapes possess a strong and elegant structure, making them highly suitable for the preferences of Chinese consumers. As a result, many wineries in the Ningxia Hui Autonomous Region have made Marselan wines their main product[4]. Wine is a complex beverage that is influenced by various natural and anthropogenic factors throughout the wine-making process. These factors include soil, climate, agrochemicals, and human intervention. While there is an abundance of research available on wine production, limited research has been conducted specifically on local wines in the Eastern Foot of Helan Mountain. This research gap is of significant importance for the management and quality improvement of Chinese local wines.

    Ion mobility spectrometry (IMS) is a rapid analytical technique used to detect trace gases and characterize chemical ionic substances. It achieves this through the gas-phase separation of ionized molecules under an electric field at ambient pressure. In recent years, IMS has gained increasing popularity in the field of food-omics due to its numerous advantages. These advantages include ultra-high analytical speed, simplicity, easy operation, time efficiency, relatively low cost, and the absence of sample preparation steps. As a result, IMS is now being applied more frequently in various areas of food analysis, such as food composition and nutrition, food authentication, detection of food adulteration, food process control, and chemical food safety[5,6]. The orthogonal hyphenation of gas chromatography (GC) and IMS has greatly improved the resolution of complex food matrices when using GC-IMS, particularly in the analysis of wines[7].

    The objective of this study was to investigate the changes in the physicochemical properties of Marselan wine during the winemaking process, with a focus on the total phenolic and flavonoids content, antioxidant activity, and volatile profile using the GC-IMS method. The findings of this research are anticipated to make a valuable contribution to the theoretical framework for evaluating the authenticity and characterizing Ningxia Marselan wine. Moreover, it is expected that these results will aid in the formulation of regulations and legislation pertaining to Ningxia Marselan wine in China.

    All the grapes used to produce Marselan wines, grow in the Xiban vineyard (106.31463° E and 38.509541° N) situated in Helan Mountain's East Foothill of Ningxia Hui Autonomous Region in China.

    Folin-Ciocalteau reagent, (±)-6-Hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox), 2,20-azino-bis-(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS), 2,4,6-tris (2-pyridyl)-s-triazine (TPTZ), anhydrous methanol, sodium nitrite, and sodium carbonate anhydrous were purchased from Shanghai Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). Reference standards of (+)-catechin, gallic acid, and the internal standard (IS) 4-methyl-2-pentanol were supplied by Shanghai Yuanye Bio-Technology Co., Ltd (Shanghai, China). The purity of the above references was higher than 98%. Ultrapure water (18.2 MΩ cm) was prepared by a Milli-Q system (Millipore, Bedford, MA, USA).

    Stage 1−Juice processing: Grapes at the fully mature stage are harvested and crushed, and potassium metabisulfite (5 mg/L of SO2) was evenly spread during the crushing process. The obtained must is transferred into stainless steel tanks. Stage 2−Alcoholic fermentation: Propagated Saccharomyces cerevisiae ES488 (Enartis, Italy) are added to the fresh must, and alcoholic fermentation takes place, after the process is finished, it is kept in the tanks for 7 d for traditional maceration to improve color properties and phenolics content. Stage 3−Malolactic fermentation: When the pomace is fully concentrated at the bottom of the tanks, the wine is transferred to another tank for separation from these residues. Oenococcus oeni VP41 (Lallemand Inc., France) is inoculated and malic acid begins to convert into lactic acid. Stage 4−Wine stabilization: After malolactic fermentation, potassium metabisulfite is re-added (35 mg/L of SO2), and then transferred to oak barrels for stabilization, this process usually takes 6-24 months. A total of four batches of samples during the production process of Marselan wine were collected in this study.

    Total polyphenols were determined on 0.5 mL diluted wine sample using the Folin-Ciocalteu method[8], using gallic acid as a reference compound, and expressed as milligrams of gallic acid equivalents per liter of wine. The total flavonoid content was measured on 0.05 mL of wine sample by a colorimetric method previously described[9]. Results are calculated from the calibration curve obtained with catechin, as milligrams of catechin equivalents per liter of wine.

    The antioxidative activity was determined using the ABTS·+ assay[10]. Briefly, the ABTS·+ radical was prepared from a mixture of 88 μL of potassium persulfate (140 mmol/L) with 5 mL of the ABTS·+ solution (7 mmol/L). The reaction was kept at room temperature under the absence of light for 16 h. Sixty μL samples were mixed with 3 mL of ABTS·+ solution with measured absorption of 0.700 ± 0.200 at 734 nm. After 6 min reaction, the absorbance of samples were measured with a spectrophotometer at 734 nm. Each sample was tested in triplicate. The data were expressed as mmol Trolox equivalent of antioxidative capacity per liter of the wine sample (mmol TE/L). Calibration curves, in the range 64.16−1,020.20 μmol TE/L, showed good linearity (R2 ≥ 0.99).

    The FRAP assay was conducted according to a previous study[11]. The FRAP reagent was freshly prepared and mixed with 10 mM/L TPTZ solution prepared in 20 mM/L FeCl3·6H2O solution, 40 mM/L HCl, and 300 mM/L acetate buffer (pH 3.6) (1:1:10; v:v:v). Ten ml of diluted sample was mixed with 1.8 ml of FRAP reagent and incubated at 37 °C for 30 min. The absorbance was determined at 593 nm and the results were reported as mM Fe (II) equivalent per liter of the wine sample. The samples were analyzed and calculated by a calibration curve of ferrous sulphate (0.15−2.00 mM/mL) for quantification.

    The volatile compounds were analyzed on a GC-IMS instrument (FlavourSpec, GAS, Dortmund, Germany) equipped with an autosampler (Hanon Auto SPE 100, Shandong, China) for headspace analysis. One mL of each wine was sampled in 20 mL headspace vials (CNW Technologies, Germany) with 20 μL of 4-methyl-2-pentanol (20 mg/L) ppm as internal standard, incubated at 60 °C and continuously shaken at 500 rpm for 10 min. One hundred μL of headspace sample was automatically loaded into the injector in splitless mode through a syringe heated to 65 °C. The analytes were separated on a MxtWAX capillary column (30 m × 0.53 mm, 1.0 μm) from Restek (Bellefonte, Pennsylvania, USA) at a constant temperature of 60 °C and then ionized in the IMS instrument (FlavourSpec®, Gesellschaft für Analytische Sensorsysteme mbH, Dortmund, Germany) at 45 °C. High purity nitrogen gas (99.999%) was used as the carrier gas at 150 mL/min, and drift gas at 2 ml/min for 0−2.0 min, then increased to 100 mL/min from 2.0 to 20 min, and kept at 100 mL/min for 10 min. Ketones C4−C9 (Sigma Aldrich, St. Louis, MO, USA) were used as an external standard to determine the retention index (RI) of volatile compounds. Analyte identification was performed using a Laboratory Analytical Viewer (LAV) 2.2.1 (GAS, Dortmund, Germany) by comparing RI and the drift time of the standard in the GC-IMS Library.

    All samples were prepared in duplicate and tested at least six times, and the results were expressed as mean ± standard error (n = 4) and the level of statistical significance (p < 0.05) was analyzed by using Tukey's range test using SPSS 18.0 software (SPSS Inc., IL, USA). The principal component analysis (PCA) was performed using the LAV software in-built 'Dynamic PCA' plug-in to model patterns of aroma volatiles. Orthogonal partial least-square discriminant analysis (OPLS-DA) in SIMCA-P 14.1 software (Umetrics, Umeă, Sweden) was used to analyze the different volatile organic compounds in the different fermentation stages.

    The results of the changes in the antioxidant activity of Marselan wines during the entire brewing process are listed in Table 1. It can be seen that the contents of flavonoids and polyphenols showed an increasing trend during the brewing process of Marselan wine, which range from 315.71−1,498 mg CE/L and 1,083.93−3,370.92 mg GAE/L, respectively. It was observed that the content increased rapidly in the alcoholic fermentation stage, but slowly in the subsequent fermentation stage. This indicated that the formation of flavonoid and phenolic substances in wine mainly concentrated in the alcoholic fermentation stage, which is consistent with previous reports. This is mainly because during the alcoholic fermentation of grapes, impregnation occurred to extract these compounds[12]. The antioxidant activities of Marselan wine samples at different fermentation stages were detected by FRAP and ABTS methods[11]. The results showed that the ferric reduction capacity and ABST·+ free radical scavenging capacity of the fermented Marselan wines were 2.4 and 1.5 times higher than the sample from the juice processing stage, respectively, indicating that the fermented Marselan wine had higher antioxidant activity. A large number of previous studies have suggested that there is a close correlation between antioxidant activity and the content of polyphenols and flavonoids[1315]. Previous studies have reported that Marselan wine has the highest total phenol and anthocyanin content compared to the wine of Tannat, Cabernet Sauvignon, Merlot, Cabernet Franc, and Syrah[13]. Polyphenols and flavonoids play an important role in improving human immunity. Therefore, Marselan wines are popular because of their high phenolic and flavonoid content and high antioxidant capacity.

    Table 1.  GC-IMS integration parameters of volatile compounds in Marselan wine at different fermentation stages.
    No. Compounds Formula RI* Rt
    [sec]**
    Dt
    [RIPrel]***
    Identification
    approach
    Concentration (μg/mL) (n = 4)
    Stage 1 Stage 2 Stage 3 Stage 4
    Aldehydes
    5 Furfural C5H4O2 1513.1 941.943 1.08702 RI, DT, IS 89.10 ± 4.05c 69.98 ± 3.22c 352.16 ± 39.06b 706.30 ± 58.22a
    6 Furfural dimer C5H4O2 1516.6 948.77 1.33299 RI, DT, IS 22.08 ± 0.69b 18.68 ± 2.59c 23.73 ± 2.69b 53.39 ± 9.42a
    12 (E)-2-hexenal C6H10O 1223.1 426.758 1.18076 RI, DT, IS 158.17 ± 7.26a 47.57 ± 2.51b 39.00 ± 2.06c 43.52 ± 4.63bc
    17 (E)-2-pentenal C5H8O 1129.2 333.392 1.1074 RI, DT, IS 23.00 ± 4.56a 16.42 ± 1.69c 18.82 ± 0.27b 18.81 ± 0.55b
    19 Heptanal C7H14O 1194.2 390.299 1.33002 RI, DT, IS 17.28 ± 2.25a 10.22 ± 0.59c 14.50 ± 8.84b 9.11 ± 1.06c
    22 Hexanal C6H12O 1094.6 304.324 1.25538 RI, DT, IS 803.11 ± 7.47c 1631.34 ± 19.63a 1511.11 ± 26.91b 1526.53 ± 8.12b
    23 Hexanal dimer C6H12O 1093.9 303.915 1.56442 RI, DT, IS 588.85 ± 7.96a 93.75 ± 4.67b 92.93 ± 3.13b 95.49 ± 2.50b
    29 3-Methylbutanal C5H10O 914.1 226.776 1.40351 RI, DT, IS 227.86 ± 6.39a 33.32 ± 2.59b 22.36 ± 1.18c 21.94 ± 1.73c
    33 Dimethyl sulfide C2H6S 797.1 193.431 0.95905 RI, DT, IS 120.07 ± 4.40c 87.a02 ± 3.82d 246.81 ± 5.62b 257.18 ± 3.04a
    49 2-Methylpropanal C4H8O 828.3 202.324 1.28294 RI, DT, IS 150.49 ± 7.13a 27.08 ± 1.48b 19.36 ± 1.10c 19.69 ± 0.92c
    Ketones
    45 3-Hydroxy-2-butanone C4H8O2 1293.5 515.501 1.20934 RI, DT, IS 33.20 ± 3.83c 97.93 ± 8.72b 163.20 ± 21.62a 143.51 ± 21.48a
    46 Acetone C3H6O 836.4 204.638 1.11191 RI, DT, IS 185.75 ± 8.16c 320.43 ± 12.32b 430.74 ± 3.98a 446.58 ± 10.41a
    Organic acid
    3 Acetic acid C2H4O2 1527.2 969.252 1.05013 RI, DT, IS 674.66 ± 46.30d 3602.39 ± 30.87c 4536.02 ± 138.86a 4092.30 ± 40.33b
    4 Acetic acid dimer C2H4O2 1527.2 969.252 1.15554 RI, DT, IS 45.25 ± 3.89c 312.16 ± 19.39b 625.79 ± 78.12a 538.35 ± 56.38a
    Alcohols
    8 1-Hexanol C6H14O 1365.1 653.825 1.32772 RI, DT, IS 1647.65 ± 28.94a 886.33 ± 32.96b 740.73 ± 44.25c 730.80 ± 21.58c
    9 1-Hexanol dimer C6H14O 1365.8 655.191 1.64044 RI, DT, IS 378.42 ± 20.44a 332.65 ± 25.76a 215.78 ± 21.04b 200.14 ± 28.34b
    13 3-Methyl-1-butanol C5H12O 1213.3 414.364 1.24294 RI, DT, IS 691.86 ± 9.95c 870.41 ± 22.63b 912.80 ± 23.94a 939.49 ± 12.44a
    14 3-Methyl-1-butanol dimer C5H12O 1213.3 414.364 1.49166 RI, DT, IS 439.90 ± 29.40c 8572.27 ± 60.56b 9083.14 ± 193.19a 9152.25 ± 137.80a
    15 1-Butanol C4H10O 1147.2 348.949 1.18073 RI, DT, IS 157.33 ± 9.44b 198.92 ± 3.92a 152.78 ± 10.85b 156.02 ± 9.80b
    16 1-Butanol dimer C4H10O 1146.8 348.54 1.38109 RI, DT, IS 24.14 ± 2.15c 274.75 ± 12.60a 183.02 ± 17.72b 176.80 ± 19.80b
    24 1-Propanol C3H8O 1040.9 274.803 1.11042 RI, DT, IS 173.73 ± 4.75a 55.84 ± 2.16c 80.80 ± 4.99b 83.57 ± 2.34b
    25 1-Propanol dimer C3H8O 1040.4 274.554 1.24784 RI, DT, IS 58.20 ± 1.30b 541.37 ± 11.94a 541.33 ± 15.57a 538.84 ± 9.74a
    28 Ethanol C2H6O 930.6 231.504 1.11901 RI, DT, IS 5337.84 ± 84.16c 11324.05 ± 66.18a 9910.20 ± 100.76b 9936.10 ± 101.24b
    34 Methanol CH4O 903.6 223.79 0.98374 RI, DT, IS 662.08 ± 13.87a 76.94 ± 2.15b 61.92 ± 1.96c 62.89 ± 0.81c
    37 2-Methyl-1-propanol C4H10O 1098.5 306.889 1.35839 RI, DT, IS 306.91 ± 4.09c 3478.35 ± 25.95a 3308.79 ± 61.75b 3313.85 ± 60.88b
    48 1-Pentanol C5H12O 1257.6 470.317 1.25222 RI, DT, IS 26.13 ± 2.52c 116.50 ± 3.71ab 112.37 ± 6.26b 124.17 ± 7.04a
    Esters
    1 Methyl salicylate C8H8O3 1859.6 1616.201 1.20489 RI, DT, IS 615.00 ± 66.68a 485.08 ± 31.30b 470.14 ± 23.02b 429.12 ± 33.74b
    7 Butyl hexanoate C10H20O2 1403.0 727.561 1.47354 RI, DT, IS 95.83 ± 17.04a 62.87 ± 3.62a 92.59 ± 11.88b 82.13 ± 3.61c
    10 Hexyl acetate C8H16O2 1298.6 524.366 1.40405 RI, DT, IS 44.72 ± 8.21a 33.18 ± 2.17d 41.50 ± 4.38c 40.89 ± 4.33b
    11 Propyl hexanoate C9H18O2 1280.9 499.577 1.39274 RI, DT, IS 34.65 ± 3.90d 70.43 ± 5.95a 43.97 ± 4.39b 40.12 ± 4.05c
    18 Ethyl hexanoate C8H16O2 1237.4 444.749 1.80014 RI, DT, IS 55.55 ± 5.62c 1606.16 ± 25.63a 787.24 ± 16.95b 788.91 ± 28.50b
    20 Isoamyl acetate C7H14O2 1127.8 332.164 1.30514 RI, DT, IS 164.22 ± 1.00d 243.69 ± 8.37c 343.51 ± 13.98b 365.46 ± 1.60a
    21 Isoamyl acetate dimer C7H14O2 1126.8 331.345 1.75038 RI, DT, IS 53.61 ± 4.79d 4072.20 ± 11.94a 2416.70 ± 49.84b 2360.46 ± 43.29c
    26 Isobutyl acetate C6H12O2 1020.5 263.605 1.23281 RI, DT, IS 101.65 ± 1.81a 15.52 ± 0.67c 44.87 ± 3.21b 45.96 ± 1.41b
    27 Isobutyl acetate dimer C6H12O2 1019.6 263.107 1.61607 RI, DT, IS 34.60 ± 1.05d 540.84 ± 5.64a 265.54 ± 8.31c 287.06 ± 3.66b
    30 Ethyl acetate dimer C4H8O2 885.2 218.564 1.33587 RI, DT, IS 1020.75 ± 6.86d 5432.71 ± 6.55a 5052.99 ± 9.65b 5084.47 ± 7.30c
    31 Ethyl acetate C4H8O2 878.3 216.574 1.09754 RI, DT, IS 215.65 ± 3.58a 38.29 ± 2.37c 71.59 ± 2.99b 69.32 ± 2.85b
    32 Ethyl formate C3H6O2 838.1 205.127 1.19738 RI, DT, IS 175.48 ± 3.79d 1603.20 ± 13.72a 1472.10 ± 5.95c 1509.08 ± 13.26b
    35 Ethyl octanoate C10H20O2 1467.0 852.127 1.47312 RI, DT, IS 198.86 ± 36.71b 1853.06 ± 17.60a 1555.51 ± 24.21a 1478.05 ± 33.63a
    36 Ethyl octanoate dimer C10H20O2 1467.0 852.127 2.03169 RI, DT, IS 135.50 ± 13.02d 503.63 ± 15.86a 342.89 ± 11.62b 297.28 ± 14.40c
    38 Ethyl butanoate C6H12O2 1042.1 275.479 1.5664 RI, DT, IS 21.29 ± 2.68c 1384.67 ± 8.97a 1236.52 ± 20.21b 1228.09 ± 5.09b
    39 Ethyl 3-methylbutanoate C7H14O2 1066.3 288.754 1.26081 RI, DT, IS 9.70 ± 1.85d 200.29 ± 4.21a 146.87 ± 8.70b 127.13 ± 12.54c
    40 Propyl acetate C5H10O2 984.7 246.908 1.48651 RI, DT, IS 4.57 ± 1.07c 128.63 ± 4.28a 87.75 ± 3.26b 88.49 ± 1.99b
    41 Ethyl propanoate C5H10O2 962.1 240.47 1.46051 RI, DT, IS 10.11 ± 0.34d 107.08 ± 3.50a 149.60 ± 5.39c 167.15 ± 12.90b
    42 Ethyl isobutyrate C6H12O2 971.7 243.229 1.56687 RI, DT, IS 18.29 ± 2.61d 55.22 ± 1.07c 98.81 ± 4.67b 104.71 ± 4.73a
    43 Ethyl lactate C5H10O3 1352.2 628.782 1.14736 RI, DT, IS 31.81 ± 2.91c 158.03 ± 2.80b 548.14 ± 74.21a 527.01 ± 39.06a
    44 Ethyl lactate dimer C5H10O3 1351.9 628.056 1.53618 RI, DT, IS 44.55 ± 2.03c 47.56 ± 4.02c 412.23 ± 50.96a 185.87 ± 31.25b
    47 Ethyl heptanoate C9H18O2 1339.7 604.482 1.40822 RI, DT, IS 39.55 ± 6.37a 38.52 ± 2.47a 28.44 ± 1.52c 30.77 ± 2.79b
    Unknown
    1 RI, DT, IS 15.53 ± 0.18 35.69 ± 0.80 12.70 ± 0.80 10.57 ± 0.86
    2 RI, DT, IS 36.71 ± 1.51 120.41 ± 3.44 198.12 ± 6.01 201.19 ± 3.70
    3 RI, DT, IS 44.35 ± 0.88 514.12 ± 4.28 224.78 ± 6.56 228.32 ± 4.62
    4 RI, DT, IS 857.64 ± 8.63 33.22 ± 1.99 35.05 ± 5.99 35.17 ± 3.97
    * Represents the retention index calculated using n-ketones C4−C9 as external standard on MAX-WAX column. ** Represents the retention time in the capillary GC column. *** Represents the migration time in the drift tube.
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    This study adopted the GC-IMS method to test the volatile organic compounds (VOCs) in the samples from the different fermentation stages of Marselan wine. Figure 1 shows the gas phase ion migration spectrum obtained, in which the ordinate represents the retention time of the gas chromatographic peaks and the abscissa represents the ion migration time (normalized)[16]. The entire spectrum represents the aroma fingerprints of Marselan wine at different fermentation stages, with each signal point on the right of the relative reactant ion peak (RIP) representing a volatile organic compound detected from the sample[17]. Here, the sample in stage 1 (juice processing) was used as a reference and the characteristic peaks in the spectrum of samples in other fermentation stages were compared and analyzed after deducting the reference. The colors of the same component with the same concentration cancel each other to form a white background. In the topographic map of other fermentation stages, darker indicates higher concentration compared to the white background. In the 2D spectra of different fermentation stages, the position and number of peaks indicated that peak intensities are basically the same, and there is no obvious difference. However, it is known that fermentation is an extremely complex chemical process, and the content and types of volatile organic compounds change with the extension of fermentation time, so other detection and characterization methods are needed to make the distinction.

    Figure 1.  2D-topographic plots of volatile organic compounds in Marselan wine at different fermentation stages.

    To visually display the dynamic changes of various substances in the fermentation process of Marselan wine, peaks with obvious differences were extracted to form the characteristic fingerprints for comparison (Fig. 2). Each row represents all signal peaks selected from samples at the same stage, and each column means the signal peaks of the same volatile compound in samples from different fermentation stages. Figure 2 shows the volatile organic compounds (VOCs) information for each sample and the differences between samples, where the numbers represent the undetermined substances in the migration spectrum library. The changes of volatile substances in the process of Marselan winemaking is observed by the fingerprint. As shown in Fig. 2 and Table 2, a total of 40 volatile chemical components were detected by qualitative analysis according to their retention time and ion migration time in the HS-GC-IMS spectrum, including 17 esters, eight alcohols, eight aldehydes, two ketones, one organic acid, and four unanalyzed flavor substances. The 12 volatile organic compounds presented dimer due to ionization of the protonated neutral components before entering the drift tube[18]. As can be seen from Table 2, the VOCs in the winemaking process of Marselan wine are mainly composed of esters, alcohols, and aldehydes, which play an important role in the construction of aroma characteristics.

    Figure 2.  Fingerprints of volatile organic compounds in Marselan wine at different fermentation stages.
    Table 2.  Antioxidant activity, total polyphenols, and flavonoids content of Marselan wine at different fermentation stages.
    Winemaking stage TFC (mg CE/L) TPC (mg GAE/L) FRAP (mM FeSO4/mL) ABTs (mM Trolox/L)
    Stage 1 315.71 ± 0.00d 1,083.93 ± 7.79d 34.82c 38.92 ± 2.12c
    Stage 2 1,490.00 ± 7.51c 3,225.51 ± 53.27c 77.32b 52.17 ± 0.95b
    Stage 3 1,510.00 ± 8.88a 3,307.143 ± 41.76b 77.56b 53.04 ± 0.76b
    Stage 4 1,498.57 ± 6.34b 3,370.92 ± 38.29a 85.07a 57.46 ± 2.55a
    Means in the same column with different letters are significantly different (p < 0.05).
     | Show Table
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    Esters are produced by the reaction of acids and alcohols in wine, mainly due to the activity of yeast during fermentation[19], and are the main components of fruit juices and wines that produce fruit flavors[20,21]. In this study, it was found that they were the largest detected volatile compound group in Marselan wine samples, which is consistent with previous reports[22]. It can be observed from Table 2 that the contents of most esters increased gradually with the extension of fermentation time, and they mainly began to accumulate in large quantities during the stage of alcohol fermentation. The contents of ethyl hexanoate (fruity), isoamyl acetate (banana, pear), ethyl octanoate (fruity, pineapple, apple, brandy), ethyl acetate (fruity), ethyl formate (spicy, pineapple), and ethyl butanoate (sweet, pineapple, banana, apple) significantly increased at the stage of alcoholic fermentation and maintained a high level in the subsequent fermentation stage (accounting for 86% of the total detected esters). These esters can endow a typical fruity aroma of Marselan wine, and played a positive role in the aroma profiles of Marselan wine. Among them, the content of ethyl acetate is the highest, which is 5,153.79 μg/mL in the final fermentation stage, accounting for 33.6% of the total ester. However, the content of ethyl acetate was relatively high before fermentation, which may be from the metabolic activity of autochthonous microorganisms present in the raw materials. Isobutyl acetate, ethyl 3-methyl butanoate, propyl acetate, ethyl propanoate, ethyl isobutyrate, and ethyl lactate were identified and quantified in all fermentation samples. The total contents of these esters in stage 1 and 4 were 255.28 and 1,533.38 μg/mL, respectively, indicating that they may also have a potential effect on the aroma quality of Marselan wine. The results indicate that esters are an important factor in the formation of flavor during the brewing process of Marselan wine.

    Alcohols were the second important aromatic compound in Marselan wine, which were mainly synthesized by glucose and amino acid decomposition during alcoholic fermentation[23,24]. According to Table 2, eight alcohols including methanol, ethanol, propanol, butanol, hexanol, amyl alcohol, 3-methyl-1-butanol, and 2-methyl-1-propanol were detected in the four brewing stages of Marselan wine. The contents of ethanol (slightly sweet), 3-methyl-1-butanol (apple, brandy, spicy), and 2-methyl-1-propanol (whiskey) increased gradually during the fermentation process. The sum of these alcohols account for 91%−92% of the total alcohol content, which is the highest content of three alcohols in Marselan wine, and may be contributing to the aromatic and clean-tasting wines. On the contrary, the contents of 1-hexanol and methanol decreased gradually in the process of fermentation. Notably, the content of these rapidly decreased at the stage of alcoholic fermentation, from 2,026.07 to 1,218.98 μg/mL and 662.08 to 76.94 μg/mL, respectively, which may be ascribed to volatiles changed from alcohols to esters throughout fermentation. The reduction of the concentration of some alcohols also alleviates the strong odor during wine fermentation, which plays an important role in the improvement of aroma characteristics.

    Acids are mainly produced by yeast and lactic acid bacteria metabolism at the fermentation stage and are considered to be an important part of the aroma of wine[22]. Only one type of acid (acetic acid) was detected in this experiment, which was less than previously reported, which may be related to different brewing processes. Acetic acid content is an important factor in the balance of aroma and taste of wine. Low contents of volatile acids can provide a mild acidic smell in wine, which is widely considered to be ideal for producing high-quality wines. However, levels above 700 μg/mL can produce a pungent odor and weaken the wine's distinctive flavor[25]. The content of acetic acid increased first and then decreased during the whole fermentation process. The content of acetic acid increased rapidly in the second stage, from 719.91 to 3,914.55 μg/mL reached a peak in the third stage (5,161.81 μg/mL), and decreased to 4,630.65 μg/mL in the last stage of fermentation. Excessive acetic acid in Marselan wine may have a negative impact on its aroma quality.

    It was also found that the composition and content of aldehydes produced mainly through the catabolism of amino acids or decarboxylation of ketoacid were constantly changing during the fermentation of Marselan wines. Eight aldehydes, including furfural, hexanal, heptanal, 2-methylpropanal, 3-methylbutanal, dimethyl sulfide, (E)-2-hexenal, and (E)-2-pentenal were identified in all stage samples. Among them, furfural (caramel bread flavor) and hexanal (grass flavor) are the main aldehydes in Marselan wine, and the content increases slightly with the winemaking process. While other aldehydes such as (E)-2-hexenal (green and fruity), 3-methylbutanol (fresh and malt), and 2-methylpropanal (fresh and malt) were decomposed during brewing, reducing the total content from 536.52 to 85.15 μg/mL, which might potently affect the final flavor of the wine. Only two ketones, acetone, and 3-hydroxy-2-butanone, were detected in the wine samples, and their contents had no significant difference in the fermentation process, which might not affect the flavor of the wine.

    To more intuitively analyze the differences of volatile organic compounds in different brewing stages of Marselan wine samples, principal component analysis was performed[2628]. As presented in Fig. 3, the points corresponding to one sample group were clustered closely on the score plot, while samples at different fermentation stages were well separated in the plot. PC1 (79%) and PC2 (18%) together explain 97% of the total variance between Marselan wine samples, indicating significant changes in volatile compounds during the brewing process. As can be seen from the results in Fig. 3, samples of stages 1, 2, and 3 can be distinguished directly by PCA, suggesting that there are significant differences in aroma components in these three fermentation stages. Nevertheless, the separation of stage 3 and stage 4 samples is not very obvious and both presented in the same quadrant, which means that their volatile characteristics were highly similar, indicating that the volatile components of Marselan wine are formed in stage 3 during fermentation (Fig. S1). The above results prove that the unique aroma fingerprints of the samples from the distinct brewing stages of Marselan wine were successfully constructed using the HS-GC-IMS method.

    Figure 3.  PCA based on the signal intensity obtained with different fermentation stages of Marselan wine.

    Based on the results of the PCA, OPLS-DA was used to eliminate the influence of uncontrollable variables on the data through permutation test, and to quantify the differences between samples caused by characteristic flavors[28]. Figure 4 revealed that the point of flavor substances were colored according to their density and the samples obtained at different fermentation stages of wine have obvious regional characteristics and good spatial distribution. In addition, the reliability of the OPLS-DA model was verified by the permutation method of 'Y-scrambling'' validation. In this method, the values of the Y variable were randomly arranged 200 times to re-establish and analyze the OPLS-DA model. In general, the values of R2 (y) and Q2 were analyzed to assess the predictability and applicability of the model. The results of the reconstructed model illustrate that the slopes of R2 and Q2 regression lines were both greater than 0, and the intercept of the Q2 regression line was −0.535 which is less than 0 (Fig. 5). These results indicate that the OPLS-DA model is reliable and there is no fitting phenomenon, and this model can be used to distinguish the four brewing stages of Marselan wine.

    Figure 4.  Scores plot of OPLS-DA model of volatile components in Marselan wine at different fermentation stages.
    Figure 5.  Permutation test of OPLS-DA model of volatile components in Marselan wine at different fermentation stages (n = 200).

    VIP is the weight value of OPLS-DA model variables, which was used to measure the influence intensity and explanatory ability of accumulation difference of each component on classification and discrimination of each group of samples. In previous studies, VIP > 1 is usually used as a screening criterion for differential volatile substances[2830]. In this study, a total of 22 volatile substances had VIP values above 1, indicating that these volatiles could function as indicators of Marselan wine maturity during fermentation (see Fig. 6). These volatile compounds included furfural, ethyl lactate, heptanal, dimethyl sulfide, 1-propanol, ethyl isobutyrate, propyl acetate, isobutyl acetate, ethanol, ethyl hexanoate, acetic acid, methanol, ethyl formate, ethyl 3-methylbutanoate, ethyl acetate, hexanal, isoamyl acetate, 2-methylpropanal, 2-methyl-1-propanol, and three unknown compounds.

    Figure 6.  VIP plot of OPLS-DA model of volatile components in Marselan wine at different fermentation stages.

    This study focuses on the change of volatile flavor compounds and antioxidant activity in Marselan wine during different brewing stages. A total of 40 volatile aroma compounds were identified and collected at different stages of Marselan winemaking. The contents of volatile aroma substances varied greatly at different stages, among which alcohols and esters were the main odors in the fermentation stage. The proportion of furfural was small, but it has a big influence on the wine flavor, which can be used as one of the standards to measure wine flavor. Flavonoids and phenols were not only factors of flavor formation, but also important factors to improve the antioxidant capacity of Marselan wine. In this study, the aroma of Marselan wines in different fermentation stages was analyzed, and its unique aroma fingerprint was established, which can provide accurate and scientific judgment for the control of the fermentation process endpoint, and has certain guiding significance for improving the quality of Marselan wines (Table S1). In addition, this work will provide a new approach for the production management of Ningxia's special wine as well as the development of the native Chinese wine industry.

  • The authors confirm contribution to the paper as follows: study conception and design: Gong X, Fang L; data collection: Fang L, Li Y; analysis and interpretation of results: Qi N, Chen T; draft manuscript preparation: Fang L. All authors reviewed the results and approved the final version of the manuscript.

  • The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.

  • This work were supported by the project of Hainan Province Science and Technology Special Fund (ZDYF2023XDNY031) and the Central Public-interest Scientific Institution Basal Research Fund for Chinese Academy of Tropical Agricultural Sciences in China (Grant No. 1630122022003).

  • The authors declare that they have no conflict of interest.

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    Dissanayake LS, Phookamsak R, Xu J, Wanasinghe DN. 2024. Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China. Studies in Fungi 9: e016 doi: 10.48130/sif-0024-0016
    Dissanayake LS, Phookamsak R, Xu J, Wanasinghe DN. 2024. Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China. Studies in Fungi 9: e016 doi: 10.48130/sif-0024-0016

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Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China

Studies in Fungi  9 Article number: e016  (2024)  |  Cite this article

Abstract: During the investigation of ascomycetes on Poaceae in Yunnan Province, China, a fungus was collected from dead bamboo culms in a terrestrial habitat in the Ailaoshan subtropical evergreen broad-leaved forest. Based on morphological characterization and phylogenetic analyses, this fungus was introduced as Oxydothis ailaoshanensis sp. nov. Maximum likelihood and Bayesian inference analyses of a concatenated dataset of internal transcribed spacer (ITS), large subunits (LSU) of the nuclear ribosomal RNA gene, small subunit (SSU) of the nuclear ribosomal RNA gene, and translation elongation factor 1-alpha (tef1-α) sequences were performed to clarify the phylogenetic affinities of the new species. Phylogenetically, Oxydothis ailaoshanensis forms an independent lineage, basal to O. metroxylonis. Oxydothis ailaoshanensis differs from O. metroxylonis in having smaller, immersed ascomata that become raised and superficial with the long axis horizontal to the host surface, shorter asci that are mostly straight, and longer ascospores that are elongated and fusiform. The new species was justified based on morphological traits and multigene phylogenetic analyses in comparison with closely related species. A detailed description, micrograph, and a phylogenetic tree of the new species are provided.

    • China is recognized as the most diverse country for bamboo, with 43 genera and 728 species[1]. Southwest China, particularly Yunnan Province is the richest area of bamboo diversity in the country, accounting for 50% of all bamboo species diversity[2]. Yunnan has three types of bamboo forests viz. cold-temperate, temperate, and tropical bamboo forests[3]. Since 2017, many studies of bambusicolous fungi have been conducted in Yunnan[2,410]. However, studies on bambusicolous fungi in this region remain limited compared to other regions such as Hong Kong and Taiwan[2,11]. Among these studies, Xylariales has been relatively under-researched in Yunnan (comprising 14% of known species), compared to Pleosporales (39% of known species). Nonetheless, Sordariomycetes has emerged as the largest group of Ascomycota found on bamboo[2].

      The genus Oxydothis was introduced by Penzig & Saccardo[12] to initially accommodate three species (viz. O. grisea, O. maculosa, and O. nigricans) within the Amphisphaeriaceae, with O. grisea designated as the type species. The taxonomic placement of Oxydothis has been subject to extensive historical discussion[1321]. A comprehensive study of Oxydothis was carried out by Konta et al.[20] who introduced Oxydothidaceae to accommodate the genus Oxydothis within Xylariales. Species of Oxydothis are characterized by solitary or aggregated ascomata that form in large groups, appearing as darkened, raised regions or dots on the host surface, and cylindrical asci with a J+ (rarely J-) subapical apparatus. The ascospores are filiform to fusiform, hyaline, 1-septate, and have pointed or blunt ends. In some species, ascospores produce appressoria upon germination[20,22]. The asexual morph of Oxydothis has been linked to Selenosporella species by Samuels & Rossman[23], although the sexual-asexual connection between Oxydothis and Selenosporella remains unproven. Furthermore, Samuels & Rossman[23] reported that the asexual morph of O. selenosporellae sporulated in vitro, producing black stromatic masses with perithecia developing throughout the colony. This morph displays macronematous, mononematous conidiophores that are (1–)2–3 septate, unbranched or branched, brown to olivaceous, with finely denticulate conidiogenous cells. A minute refractive scar remains after the conical dehiscence, and the conidia are arcuate, hyaline, and unicellular.

      Oxydothis is a common genus mainly found on monocotyledons, such as bamboo (Poaceae), palms (Arecaceae), and Pandanus (Pandanaceae) where it exists mainly as a saprobe[1820,22,2429]. A few species have been reported as phytopathogens[30] or as endophytes[31]. Hyde et al.[22] suggested that the Oxydothis species may serve as biological control agents against plant pathogens. Species in this genus with appressorium formation are involved in protein and amino acid degradation, as well as secondary metabolite synthesis i.e. melanin biosynthesis[32]. Neoxylaria, Oxydothis, and Xylaria species collected from palms produce appressoria indicating they may have the ability to produce secondary metabolites especially when they live as endophytes[24,32]. Ninety-one epithets are currently listed under the Oxydothis in Index Fungorum (https://indexfungorum.org/Names/Names.asp; accessed on 11 November 2024), though more than half of the described species lack molecular data to clarify their phylogenetic relationships. Among those that do have sequence data, most species are represented only by ITS, LSU, and SSU sequences. Recently, Zhang et al.[29] introduced five new Oxydothis species (O. caryotae, O. foliata, O. palmae, O. pyriforme, and O. sinensis) and improved the phylogenetic resolution of the genus using multigene phylogeny based on a concatenated dataset of ITS, LSU, SSU, tef1-α, and rpb2 sequences. Unfortunately, only a few protein coding genes (i.e. rpb2 and tef1-α) are currently available for Oxydothis species in GenBank, resulting in incomplete classification of the phylogenetic relationships within the genus.

      During ongoing studies of Xylariomycetidae in Southwest China[3337], numerous new species have been reported. As part of this continuing effort, the new species Oxydothis ailaoshanensis was discovered in the Ailaoshan rain forest of Yunnan Province, China on dead bamboo culms. This is the second report of an Oxydothis species associated with a bamboo host in China. The new species is described based on its morphological characteristics and supported by multi-gene phylogenetic analyses.

    • Dead bamboo culms were collected from the Ailaoshan subtropical evergreen broad-leaved forest (24.536944° N, 101.019444° E, 2,500 m), in Yunnan Province, China during the rainy season (April 2023). Specimens were processed according to the methods outlined by Senanayake et al.[38]. Observation and photography of the morphological features followed the procedure described in Dissanayake et al.[37]. Melzer's reagent was used to examine the apical apparatus of the asci. Measurements were taken using the Tarosoft (R) Image Framework software and photo plates representing the morphology were edited with Adobe Photoshop CS6 software (Adobe Systems, USA). Type specimens (holotype and isotype) were deposited in the Herbarium of Cryptogams, Kunming Institute of Botany, Chinese Academy of Sciences (KUN-HKAS), Kunming, China. The new taxon was registered in MycoBank (www.mycobank.org) and Facesoffungi[39].

    • Fungal DNA was extracted from 15–20 fresh fruiting bodies of the fungus as described in Wanasinghe et al.[40] using Forensic DNA Kit (Omega®, Norcross, GA, USA). LSU, SSU, ITS, and tef1 gene were amplified with primer pairs LR0R/LR5[41], NS1/NS4[42], ITS5/ITS4[42], and EF1-983F/EF1-2218R[43] respectively. The thermal cycling program was followed by Konta et al.[20] and Hu et al.[26]. The amplified PCR fragments were sent to a commercial sequencing provider (BGI, Ltd Shenzhen, PR China). All the sequences generated in this study were deposited in GenBank (Table 1).

      Table 1.  Names, strain numbers, and GenBank accession numbers of the taxa used in the phylogenetic analyses.

      Species name Strain no. GenBank accession no.
      ITS LSU SSU tef1-α
      Oxydothis ailaoshanensis HKAS 130464T PQ635200 PQ587528 PQ587530 PQ584438
      O. ailaoshanensis HKAS 130465IST PQ635201 PQ587529 PQ587531 PQ584439
      O. calamicola MFLUCC 14-1165T NA KY206761 KY206767 NA
      O. caryotae HKAS 115712T PP592449 PP621075 PP639207 PP761002
      O. chinensis ZHKUCC 22-0134T OR164912 OR164957 NA NA
      O. coperniciae CMUB 40043T PP278359 PP278849 PP278850 NA
      O. cyrtostachicola MRC0007T DQ660334 DQ660337 NA NA
      O. daemonoropsicola MRC 0005 DQ660335 DQ660338 NA NA
      O. dehongensis ZHKU 23-0986T PP580831 PP002130 PP002127 PP001172
      O. dehongensis ZHKU 23-0987 PP580832 PP002131 PP002128 PP001173
      O. dehongensis ZHKU 23-0988 NA PP002132 PP002129 PP001174
      O. foliata MFLU 24-0165T PP592450 PP621076 PP639208 PP761003
      O. fortunei GMBC0315T NR_187011 NG_228961 NA NA
      O. fortunei GMBC0389 ON510944 ON510945 NA NA
      O. frondicola HKUCC 3173/Mt14 AF009803 AY083835 AY083818 NA
      O. garethjonesii MFLUCC 15-0287T KY206773 KY206762 KY206768 KY206777
      O. hohnelli HKUCC 3854 NA DQ810227 DQ810259 NA
      O. inaequalis MRC0004T DQ660336 DQ660339 NA NA
      O. metroxylonicola MFLUCC 15-0281T KY206774 KY206763 KY206769 NA
      O. metroxylonis MFLUCC 15-0283T KY206775 KY206764 KY206770 KY206779
      O. narathiwatensis MFLUCC 24-0085T PP824654 PP824658 PP824659 NA
      O. palmae HKAS 115711T PP592451 PP621077 PP639209 PP761004
      O. palmicola MFLUCC 15-0806T KY206776 KY206765 KY206771 NA
      O. phoenicis MFLUCC 18-0269T MK088065 MK088061 MK088063 MK087667
      O. phoenicis MFLUCC 18-0270IS MK088066 MK088062 MK088064 MK087668
      O. pyriforme HKAS 115710T PP592452 PP621078 PP639210 PP761005
      O. rhapidicola MFLUCC 14-0616T NA KY206766 KY206772 NA
      O. sinensis GZCC21-0240T PP592453 PP621079 PP639211 PP761006
      Oxydothis sp. JHGB17 3A MH268015 NA NA NA
      Oxydothis sp. IFO 32218 NA DQ810225 DQ810261 NA
      Oxydothis sp. E04B-2 PP592454 PP621080 PP639212 NA
      O. yunnanensis GZUCC 0127T ON176681 ON176684 NA NA
      Vialaea mangiferae MFLUCC 12-0808T KF724974 KF724975 NA NA
      V. minutella BRIP 56959 KC181926 KC181924 NA NA
      Superscripts 'T' and 'IST' represent the type and isotype strains. Newly generated sequences are indicated in bold. 'NA' sequences are unavailable.
    • Newly generated sequences were subjected to BLAST search in the NCBI GenBank database and sequences of closely related taxa were downloaded. Phylogenetic analysis was performed using ITS, LSU, SSU, and tef1-α sequences (Table 1). Multiple alignments, including both consensus sequences and reference sequences, were generated using MAFFT v. 7[44] and manually refined using BioEdit v. 7.0.5.2[45]. The individual datasets were combined into a concatenated dataset and further refined with BioEdit. Combined and individual datasets were subjected to maximum likelihood (ML) and Bayesian inference (BI) analyses. The best-fit substitution models were evaluated using MrModeltest v. 2.3[46] with the Akaike Information Criterion (AIC) as the selection criteria executed in PAUP v. 4.0b10[47]. ML and BI analyses were performed on the CIPRES Science Gateway platform[48]. For ML analyses, RAxML-HPC2 on XSEDE v. 8.2.10[49,50] was used, applying the GTR + I + G model with 1000 bootstrap repetitions. The BI analysis was executed with MrBayes on XSEDE v.3.2.7a[5153] under the GTR + I + G, with one million generations and sampling every 100 generations. The analysis stopped automatically once the standard deviation of split frequencies fell below 0.01, with a burn-in fraction of 0.25. Phylogenetic results were considered significant if ML bootstrap values (MLB) ≥ 60% and Bayesian posterior (BYPP) ≥ 0.95, which were displayed above each node in the resulting tree. The phylogram was visualized using the FigTree v1.4.0 program[54], and final reorganization was done in Microsoft PowerPoint (2019).

    • The combined ITS, LSU, SSU, and tef1 matrix comprised 34 strains, including Vialaea mangiferae (MFLUCC 12-0808) and V. minutella (BRIP 56959) as outgroup taxa. The concatenated alignment consisted of 3,421 characters (ITS: 1–680 bp, LSU: 681–1,510 bp, SSU: 1,511–2,512 bp, tef1-α: 2,513–3,421 bp), including gaps. The species-level relationships within Oxydothis in both ML and BI trees were similar in topology. The best-scoring RAxML tree was selected to represent the taxa relationship with a final likelihood value of –15,291.651757 (Fig. 1). The phylogenetic tree obtained from ML analysis in Fig. 1 showed a topology consistent with previous work[28,29]. The matrix had 990 distinct alignment patterns with 33.64% of characters being undetermined or gaps. The proportion of invariable sites I = 0.453933, the gamma distribution shape parameter alpha = 0.576006 and the Tree-Length = 1.338729. The Bayesian analysis ran for one million generations, with the average standard deviation of split frequencies reaching below 0.01 (0.009728). This analysis generated 1,922 trees, from which 721 were sampled after discarding the 25% as burn-in. The alignment contained a total of 995 unique site patterns.

      Figure 1. 

      RAxML tree based on a combined dataset of analyzed ITS, LSU, SSU, and tef1-α sequences. Bootstrap support values for ML equal to or greater than 60%, Bayesian posterior probabilities (BYPP) equal to or greater than 0.95 are shown as MLB/BYPP above the nodes. The new isolate is in red; ex-type strains are in bold. The scale bar represents the expected number of nucleotide substitutions per site.

      Based on the results of multigene phylogeny, two strains of the new collection (Oxydothis ailaoshanensis sp. nov.) formed a robust subclade within the monophyletic clade of Oxydothis (60% MLB, 0.99 BYPP, Fig. 1). Oxydothis ailaoshanensis (HKAS 130464 and HKAS 130465) formed an independent lineage that is basal to O. calamicola (MFLUCC 14-1165), O. coperniciae (CMUB 40043), O. caryota (HKAS 115712), O. cyrtostachicola (MRC-007), O. metroxylonicola (MFLUCC 15-0281), O. metroxylonis (MFLUCC 15-0283), O. palmicola (MFLUCC 15-0806), O. palmae (HKAS 115711), O. phoenicis (MFLUCC 18-0269, MFLUCC 18-0270), and O. rhapidicola (MFLUCC 14-0616).

    • Oxydothis ailaoshanensis L.S. Dissan., Phookamsak & Wanas. sp. nov. (Fig. 2)

      Figure 2. 

      Oxydothis ailaoshanensis (HKAS 130464, holotype). (a) Substrate. (b), (c) Ascomata on the host surface. (d) Section of an ascoma. (e) Close up of ostiole. (f) Peridium. (g) Paraphyses. (h)–(j) Asci (j = Asci in Melzer's reagent showing J+, apical apparatus, arrow showing short pedicel). (k)–(o) Ascospores. (o = Ascospore in Congo Red). Scale bars: (c) 200 μm, (e) 50 μm, (f, g) 10 μm, (h–j) 50 μm, (k–o) 20 μm.

      MycoBank: MB856009; Facesoffungi number: FoF 14917

      Etymology − The specific epithet is derived from the locality, Ailaoshan, where the holotype was collected

      Holotype − HKAS 130464

      Saprobic on dead bamboo culms (Poaceae). Sexual morph: Ascomata 450–550 μm high × 130–150 μm diameter (x¯ = 508 × 140 μm, n = 5), solitary or aggregated in groups, immersed in host's exodermis, becoming raised, superficial, visible as black, shiny knobbed, long axis horizontal to flat on the host, hemispherical (dome-shaped) to subconical, with flattened, wedge-shaped base, uniloculate, somewhat clustered, forming pseudostromatic, with 3–6 in groups of ascomata, glabrous, ostiolate, papillate. Ostiole 50–100 μm long × 40–50 μm diameter (x¯ = 75 × 140 μm, n = 5), central, broad neck, open-ended. Peridium 15–20 μm wide, composed of several layers of flattened, inside thin cell layers, composed, hyaline, textura prismatica, outside, thin, light brown to dark brown, textura prismatica, merged with host tissues. Paraphyses are cylindrical, fragmented, hyaline, branched, or non-branched. Asci 100–120 × 10–15 μm (x¯ = 110 × 13 μm, n = 15), 8-spored, unitunicate, cylindrical to elongated fusiform, with blunt apex, slightly tapering towards both ends, mostly straight, short pedicillate, with a J+, apical apparatus. Ascospores 55–75 × 3–5 μm (x¯ = 65 × 4 μm, n = 15), overlapping 1–3-seriate, elongated fusiform, with acute ends, hyaline, obliquely 1-septate, tapering gradually from the center to the ends, with multi-guttules in each cell, pointed processes. Asexual morph: Undetermined.

      Material examined – CHINA, Yunnan Province, Ailaoshan Forest Mountain (24.536944° N, 101.019444° E, 2,500 m), on dead culms of bamboo, 7 April 2023, L.S. Dissanayake, ALF23-10 (HKAS 130464, holotype), ibid., ALF23-10A (HKAS 130465, isotype).

      Note: The multigene phylogenetic analyses indicate that Oxydothis ailaoshanensis is closely related to O. metroxylonis (MFLUCC 15-0283), with 60% MLB, 0.99 BYPP statistical support (Fig. 1). The nucleotide difference between O. ailaoshanensis and O. metroxylonis in ITS, LSU, SSU, and tef1-α are 124/555 bp (22.3%), 37/826 bp (4.5%), 13/981 bp (1.3%), and 95/906 bp (10.5%) respectively. Oxydothis ailaoshanensis differs from O. metroxylonis in having smaller, immersed ascomata (450–550 × 130–150 μm) that become raised and superficial with the long axis horizontal to the host surface, shorter asci (100–120 × 10–15 μm) that are mostly straight, and longer ascospores (55–75 × 3–5 μm) that are elongated and fusiform. In contrast, O. metroxylonis has larger ascomata (716–1,580 μm diam), with an axis oblique or perpendicular to the host surface, longer asci (165–181 × 9–15μm) with a cylindrical-clavate shape, and shorter ascospores (47–57 × 4–6 μm) with central curve[20]. Morphologically, O. ailaoshanensis is similar to O. bambusicola sharing characteristics such as solitary or aggregated ascomata with a central papilla, 8-spored, unitunicate, cylindrical asci with a J+, apical apparatus and elongated fusiform, 1-septate, hyaline ascospores[18]. Both species were isolated from bamboo hosts[18]. However, O. ailaoshanensis can be distinguished from O. bambusicola by its larger ascomata (450–550 × 130–150 μm vs 130–375 × 90–160 μm), shorter asci (100–120 × 10–15 μm vs 240 × 23 μm), and shorter ascospores (55–75 × 3–5 μm vs 93 × 7 μm) which taper gradually from the center to the pointed ends. In contrast, O. bambusicola ascospores are gradually tapering to the rounded apices and are covered with small amounts of mucilage. Based on the phylogenetic evidence and morphological differences, we describe our new collection as a distinct species, O. ailaoshanensis.

    • This study listed 89 accepted Oxydothis species in Table 2 with their hosts. Among them nearly all host species for Oxydothis belong to Arecaceae, except Oxydothis aequalis, O. bambusicola, and O. miscanthicola, recorded from Poaceae[18,31,55]. Notably, O. aequalis and O. bambusicola were recorded on bamboo hosts in the Philippines and the Hong Kong region in China respectively. In the current study, O. ailaoshanensis is introduced also from a bamboo host in China marking a new record for Poaceae hosts in Yunnan Province. Other Oxydothis species, i.e. Oxydothis caryotae, O. chinensis, O. fortunei, O. palmae, O. pyriforme, and O. sinensis were introduced from an Arecaceae host in China, Guangdong Province and Guizhou Province[26,27,29]. Based on the findings in Table 2, Oxydothis appears to exhibit host specificity primarily within Arecaceae and Poaceae with a distribution across both temperate and tropical regions.

      Table 2.  Host occurrences and distribution of all known Oxydothis species.

      Species name Host Family Country Ref.
      Oxydothis acutata Orania spp. Arecaceae Philippines [31]
      O. aequalis Bamboo, Calamus sp. Arecaceae, Poaceae Australia, Brunei, Malaysia, Philippines [31,57]
      O. alexandrarum Archontophoenix alexandrae Arecaceae Australia [31,58]
      O. angustispora Licuala ramsayi Arecaceae Australia, Brunei, Thailand [5759]
      O. asiatica Calamus flabellatus, Daemonorops sparsiflorus, Licuala sp. Arecaceae Australia, Brunei, China (Hong Kong) [57,60,61]
      O. asymmetrica Calamus conirostris Arecaceae Brunei [57]
      O. atypica Licuala longicalycata Arecaceae Thailand [62]
      O. atypica Licuala longicalycata Arecaceae Thailand [59]
      O. australiensis Archontophoenix sp. Arecaceae Australia [31]
      O. bambusicola Indocalamus sp. Poaceae China (Hong Kong) [18]
      O. batuapoiensis Daemonorops oxycarpa Arecaceae Brunei [57]
      O. belalongensis Licuala sp. Arecaceae Brunei [57]
      O. bruneiensis Calamus sp., Licuala sp. Arecaceae Brunei [57]
      O. calami Calamus sp., Salacca wallichiana Arecaceae Australia, Burma, China (Hong Kong), Indonesia, Myanmar, Philippines [31,57,60,61,63]
      O. calamicola Calamus sp. Arecaceae Thailand [20]
      O. caryotae Caryota sp. Arecaceae China (Guangdong Province) [29]
      O. chinensis Pandanus sp. Arecaceae China (Guangdong Province) [27]
      O. circularis Myrsine sp. Primulaceae Brazil [31]
      O. coperniciae Copernicia alba Arecaceae Thailand [28]
      O. cyrtospora Licuala ramsayi Arecaceae Australia [57]
      O. cyrtostachicola Cyrtostachys renda Arecaceae Thailand [19]
      O. daemonoropis Daemonorops sp. Arecaceae Philippines [31]
      O. daemonoropsicola Daemonorops margaritae Arecaceae Australia, China (Hong Kong), Malaysia, Thailand [57,58,60,61]
      O. dispariapicis Daemonorops oxycarpa Arecaceae Brunei [57]
      O. elaeicola Calamus sp., Elaeis sp., Livistona sp., Pandanus sp. Arecaceae Brazil, China (Hong Kong, Taiwan), Democratic Republic of the Congo, Honduras, Nigeria, Sierra Leone, Tanzania [31,60,6467]
      O. elaeidis Elaeis sp. Arecaceae China (Taiwan), Democratic Republic of the Congo, Tanzania, Zaire [31,60,65,66]
      O. extensa Licuala ramsayi Arecaceae Australia [57]
      O. foliata Licuala sp. Arecaceae Thailand [29]
      O. fortunei Trachycarpus fortunei Arecaceae China (Guizhou Province) [26]
      O. froehlichii Calamus radicalis Arecaceae Australia [31]
      O. froehlichiae Calamus sp., Linospadix sp. Arecaceae Australia [31]
      O. frondicola Licuala sp., Archontophoenix sp. Arecaceae Australia, Malaysia, Thailand [31,58,59]
      O. garethjonesii Eleais sp. Arecaceae Thailand [20]
      O. gigantea Palm Arecaceae Australia, Indonesia [31,58]
      O. grisea Arenga sp., Calamus sp., Heliconia sp., Licuala sp., Ptychosperma sp. Arecaceae China (Taiwan), Malaysia, Indonesia, Venezuela [31,59,66,68]
      O. hoehnelii Arenga sp., Calamus sp., Licuala sp. Arecaceae Philippines [31,5961]
      O. hongkongensis Daemonorops sp., Calamus sp. Arecaceae Australia, China (Hong Kong) [57,60,61]
      O. ianei Trachycarpus sp. Arecaceae China (Hubei Province), UK [58]
      O. inaequalis Wallichia siamensis Arecaceae Thailand [19]
      O. insignis Eugenia sp. Myrtaceae Brazil [31]
      O. licualae Archontophoenix sp., Calamus sp., Jessenia sp., Licuala sp. Arecaceae Australia, China (Hong Kong), Ecuador, Malaysia, Philippines, Thailand [31,5760]
      O. licualicola Licuala sp. Arecaceae Brunei, Myanmar [57,63]
      O. linospadicis Linospadix microcarya Arecaceae Australia [30,31]
      O. livistonae Licuala sp., Livistona sp. Arecaceae Brunei, Philippines, Thailand [31,57,59,69]
      O. livistonica Calamus sp., Licuala sp., Livistona sp. Arecaceae China (Hong Kong), Japan, Thailand [31,57,5961]
      O. livistonicola Licuala sp. Arecaceae Australia [57]
      O. luteaspora Calamus sp. Arecaceae Australia [31]
      O. maculosa Palm Arecaceae Indonesia [31,70]
      O. magnicolla Calamus sp., Licuala sp. Arecaceae Brunei [56]
      O. manokwariensis Calamus sp., Daemonorops sp. Arecaceae China (Hong Kong), Indonesia [31,57,60]
      O. maquilingiana Daemonorops sp. Arecaceae Philippines [31]
      O. mauritiae Mauritia flexuosa Arecaceae Ecuador [57]
      O. megalospora Calamus sp. Arecaceae Brunei [57]
      O. metroxylonis Metroxylon sagu Arecaceae Thailand [20]
      O. miscanthicola Miscanthus floridulus Poaceae China (Hong Kong) [54]
      O. narathiwatensis Eleiodoxa conferta Arecaceae Thailand [56]
      O. nigra Archontophoenix sp., Licuala sp., Livistona sp. Arecaceae Australia, China (Hong Kong), Malaysia [31,58,60,61]
      O. nigricans Ptychosperma sp. Arecaceae Indonesia [12]
      O. nonamyloidea Livistona sp. Arecaceae Indonesia [31]
      O. nonspecifica Calamus sp., Licuala sp. Arecaceae Australia, Brunei [57]
      O. nontincta Licuala sp. Arecaceae Brunei [57]
      O. nypae Nypa fruticans Arecaceae Brunei [31]
      O. nypicola Nypa fruticans Arecaceae Brunei [31]
      O. obducens Calamus sp., Linospadix microcarya Arecaceae Australia, China (Hong Kong) [31,57,60,61]
      O. oedema Mauritia flexuosa Arecaceae Brunei, China (Hong Kong), Guiana, Malaysia, Papua New Guinea, Seychelles [31,57,58,60,61]
      O. opaca Rhopalostylis sp., Ripogonum sp. Arecaceae New Zealand [31,73]
      O. oraniopsidis Calamus sp., Laccospadix sp., Licuala sp., Oraniopsis sp. Arecaceae Australia, India, Thailand [31,57,59]
      O. palmae Licuala sp. Arecaceae China (Guangdong Province) [29]
      O. palmicola Eleais guineensis Arecaceae Thailand [20]
      O. pandani Pandanus sp. Arecaceae China (Hong Kong), French Polynesia, Tubuai, United States [31,60,61]
      O. pandanicola Livistona chinensis, Pandanus sp., Pritchardia sp. Arecaceae Indonesia, Philippines, United States [31,57,60,61]
      O. parasitica Licuala ramsayi Arecaceae Australia [30,31,57]
      O. parvula Calamus sp., Orania sp., Phoenix sp. Arecaceae China (Hong Kong), Philippines [31,57,60,61]
      O. perangusta Licuala sp. Arecaceae Brunei [57]
      O. phoenicis Phoenix paludosa Arecaceae Thailand [22]
      O. poliothea Palm Arecaceae Venezuela [31]
      O. pusillispora Licuala sp. Arecaceae Brunei [57]
      O. pyriforme Licuala sp. Arecaceae China (Guangdong Province) [29]
      O. ragae Palm Arecaceae Indonesia [31]
      O. ragae Arenga sp. Arecaceae China (Hong Kong), Irian Jaya [31,60,61]
      O. rattanica Calamus sp., Daemonorops sp., Eleiodoxa sp. Arecaceae Brunei, China (Hong Kong), Thailand [57,60]
      O. rattanicola Calamus sp., Daemonorops sp. Arecaceae Australia, China (Hong Kong) [57]
      O. rhapidicola Rhapis excelsa Arecaceae Thailand [20]
      O. rhopalostylidis Rhopalostylis sapida Arecaceae New Zealand [23]
      O. rimicolla Calamus pogonacanthus Arecaceae Brunei [57]
      O. rubella Calamus sp. Arecaceae Australia [31,57,70]
      O. sabalensis Sabal sp., Serenoa sp. Arecaceae USA [71,31]
      O. saltuensis Archontophoenix sp., Cocos nucifera, Licuala sp., Livistona sp. Arecaceae Australia, Brunei, Indonesia, Papua New Guinea, Seychelles, Sri Lanka [31,57,58]
      O. selenosporellae Rhopalostylis sapida Arecaceae New Zealand [23,31,72,73]
      O. sinensis Livistona chinensis Arecaceae China (Guangdong Province) [29]
      O. tayabensis Calamus sp. Arecaceae Philippines [31]
      O. uniseriata Calamus radicalis Arecaceae Australia [57]
      O. wallichianensis Wallichia siamensis Arecaceae Thailand [19]

      Despite the substantial diversity within this genus, only 23 species currently have sequence data available in GenBank. Some species lack informative genetic markers such as ITS for Oxydothis calamicola, O. hohnelli, and O. rhapidicola, or LSU and SSU sequences data for O. chinensis, O. cyrtostachicola, O. daemonoropsicola, O. fortunei, O. inaequalis, and O. yunnanensis. Previous studies on Oxydothis taxonomy relied on ITS, LSU, and SSU sequences[20,22,26,28,56]. However, recent advancements in phylogenetic studies such as that by Zhang et al.[29], introduced five new Oxydothis species using a multigene dataset including ITS, LSU, SSU, tef1-α, and rpb2. Only nine species incorporate tef1 in their datasets, further supporting the phylogenetic placement of Oxydothis species in Oxydothidaceae. In the present study, we contribute to this approach by providing a combined ITS, LSU, SSU, and tef1-α phylogeny for introducing O. ailaoshanensis. The use of multiple gene markers in phylogenetic analyses has enhanced our ability to resolve species-level relationships within Oxydothis.

      While ITS, LSU, and SSU have previously been used as the primary markers for differentiating species, the addition of protein-coding genes such as tef1-α and rpb2 has improved resolution by increasing phylogenetic signal and reducing ambiguities in species placement[29]. For Oxydothis ailaoshanensis, the multigene analysis including tef1-α, coupled with morphological data such as the unique ascomata and spore characteristics (Fig. 2), has successfully differentiated it from close relatives such as O. metroxylonis. Nevertheless, some Oxydothis species still lack sequences for key informative gene regions, limiting comprehensive phylogenetic analyses. Future studies should focus on obtaining missing sequence data, especially for protein-coding genes, across more species in this genus. This could further clarify phylogenetic relationships and species boundaries within Oxydothis.

      An ITS BLAST search of the sequences linked Oxydothis to some leaf-litter-based ascomycetes (i.e., AF502894, AF502896, AF502740) and various uncultured fungal strains (i.e., KT328718, GU174316, AM999626, KC222801). However, these strains lack associated morphological data, which limits deeper insights into their morpho-phylogenetic relationships. This absence of morphological connections emphasizes the need for comprehensive morphological and molecular characterization of these strains. The present findings suggest that Oxydothis diversity remains underexplored in this region, with potentially numerous species still awaiting discovery.

      • Rungtiwa Phookamsak sincerely acknowledges the Introducing Talents Start-up Fund of Kunming Institute of Botany, Chinese Academy of Sciences, Yunnan Revitalization Talent Support Program 'Young Talent' Project (Grant No. YNWR-QNBJ-2020-120), Yunnan Revitalization Talent Support Program "High-end Foreign Expert" Project and the Independent Research Department of Economic Plants and Biotechnology, Yunnan Key Laboratory for Wild Plant Resources, Kunming Institute of Botany, Chinese Academy of Sciences (Grant No. Y537731261). Jianchu Xu thanks the Yunnan Department of Sciences and Technology of China (Grant No. 202302AE090023, 202303AP140001). Dhanushka N. Wanasinghe is funded by the Distinguished Scientist Fellowship Program (DSFP), King Saud University, Kingdom of Saudi Arabia.

      • The authors confirm contribution to the paper as follows: conceptualization, formal analysis, writing – original draft: Dissanayake LS; data curation: Dissanayake LS, Wanasinghe DN; methodology: Dissanayake LS, Phookamsak R; resources, project administration: Wanasinghe DN, Xu J; supervision: Xu J; writing – review and editing: Wanasinghe DN, Phookamsak R. All authors reviewed the results and approved the final version of the manuscript.

      • The datasets generated for this study can be found in the NCBI, GenBank and MycoBank.

      • The authors declare that they have no conflict of interest.

      • Copyright: © 2024 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (2)  Table (2) References (73)
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    Dissanayake LS, Phookamsak R, Xu J, Wanasinghe DN. 2024. Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China. Studies in Fungi 9: e016 doi: 10.48130/sif-0024-0016
    Dissanayake LS, Phookamsak R, Xu J, Wanasinghe DN. 2024. Oxydothis ailaoshanensis sp. nov. (Oxydothidaceae, Xylariales) from dead bamboo culms in Yunnan Province, China. Studies in Fungi 9: e016 doi: 10.48130/sif-0024-0016

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