ARTICLE   Open Access    

a novel coniothyrium-like genus in Coniothyriaceae ( Pleosporales) from salt marsh ecosystems in Thailand

More Information
  • In this study, a novel coniothyrium-like genus Coniothyrioides is introduced to Coniothyriaceae based on a fresh fungal collection from salt marsh habitats in Thailand. Coniothyrium-like taxa are taxonomically controversial and have been classified into different families in Pleosporales such as Didymosphaeriaceae ( Alloconiothyrium and Paraconiothyrium), Coniothyriaceae ( Coniothyrium) and Didymellaceae ( Microsphaeropsis). However, our novel genus shares similar morphology to some key characters in Coniothyriaceae in having dark, globose pycnidia, uni-locular conidiomata, a central ostiole, a peridium of textura angularis cells, and doliiform conidiogenous cells with a periclinal thickening at the apex, while conidial morphologies are diverse. The presence of setae arising from the outer peridial wall is the main difference between Coniothyrioides and other closely related Coniothyriaceae genera. Phylogenetically, LSU-SSU-ITS sequence analyses confirm the placement of this novel genus as a distinct lineage within Coniothyriaceae. Species boundaries were defined, based on morphology and multi-gene phylogenetic analyses using maximum likelihood and Bayesian inference analyses. The comprehensive descriptions and micrographs are provided. Our findings expand the taxonomic knowledge of Ascomycota in salt marsh ecosystems.
  • 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.
     | Show Table
    DownLoad: CSV

    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
    DownLoad: CSV

    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.

  • [1]

    Bertness MD. 2008. Atlantic Shorelines: Natural History and Ecology. Princeton University Press: Princeton & Oxford. 431 pp

    [2]

    Calabon MS, Jones EBG, Promputtha I, Hyde KD. 2021. Fungal biodiversity in salt marsh ecosystems. Journal of Fungi 7:648

    doi: 10.3390/jof7080648

    CrossRef   Google Scholar

    [3]

    da Luz Calado M, Carvalho L, Barata M, Pang K. 2019. Potential roles of marine fungi in the decomposition process of standing stems and leaves of Spartina maritima. Mycologia 111:371−83

    doi: 10.1080/00275514.2019.1571380

    CrossRef   Google Scholar

    [4]

    Jones EBG, Pang K, Abdel-Wahab MA, Scholz B, Hyde KD, et al. 2019. An online resource for marine fungi. Fungal Diversity 96:347−433

    doi: 10.1007/s13225-019-00426-5

    CrossRef   Google Scholar

    [5]

    Dayarathne MC, Wanasinghe DN, Devadatha B, Abeywickrama P, Gareth Jones EB, et al. 2020. Modern taxonomic approaches to identifying diatrypaceous fungi from marine habitats, with a novel genus Halocryptovalsa Dayarathne & K.D. Hyde, gen. nov. Cryptogamie, Mycologie 41:21−67

    doi: 10.5252/cryptogamie-mycologie2020v41a3

    CrossRef   Google Scholar

    [6]

    Devadatha B, Jones EBG, Pang KL, Abdel-Wahab MA, Hyde KD, et al. 2021. Occurrence and geographical distribution of mangrove fungi. Fungal Diversity 106:137−227

    doi: 10.1007/s13225-020-00468-0

    CrossRef   Google Scholar

    [7]

    Cooke WB. 1983. Coniothyriaceae. Revista de Biologia (Lisbon) 12:289

    [8]

    Corda AKJ. 1840. Icones Fungorum hucusque Cognitorum. 4:38

    [9]

    Cortinas MN, Burgess T, Dell B, Xu D, Crous PW, et al. 2006. First record of Colletogloeopsis zuluense comb. nov., causing a stem canker of Eucalyptus in China. Mycological Research 110:229−36

    doi: 10.1016/j.mycres.2005.08.012

    CrossRef   Google Scholar

    [10]

    Hongsanan S, Hyde KD, Phookamsak R, Wanasinghe DN, McKenzie EHC, et al. 2020. Refined families of Dothideomycetes: Dothideomycetidae and Pleosporomycetidae. Mycosphere 11:1553−2107

    doi: 10.5943/mycosphere/11/1/13

    CrossRef   Google Scholar

    [11]

    Kirk P, Cannon P, Minter D, Stalpers JA. 2008. Dictionary of the Fungi. 10th editionn. UK: CAB International

    [12]

    de Gruyter J, Aveskamp MM, Woudenberg JHC, Verkley GJM, Groenewald JZ, et al. 2009. Molecular phylogeny of Phoma and allied anamorph genera: towards a reclassification of the Phoma complex. Mycological Research 113:508−19

    doi: 10.1016/j.mycres.2009.01.002

    CrossRef   Google Scholar

    [13]

    Schoch CL, Crous PW, Groenewald JZ, Boehm EWA, Burgess TI, et al. 2009. A class-wide phylogenetic assessment of Dothideomycetes. Studies in Mycology 64:1−15

    doi: 10.3114/sim.2009.64.01

    CrossRef   Google Scholar

    [14]

    Schoch CL, Sung GH, López-Giráldez F, Townsend JP, Miadlikowska J, et al. 2009. The Ascomycota tree of life: a phylum–wide phylogeny clarifies the origin and evolution of fundamental reproductive and ecological traits. Systematic Biology 58:224−39

    doi: 10.1093/sysbio/syp020

    CrossRef   Google Scholar

    [15]

    Aveskamp MM, de Gruyter J, Woudenberg JHC, Verkley GJM, Crous PW. 2010. Highlights of the Didymellaceae: a polyphasic approach to characterise Phoma and related pleosporalean genera. Studies in Mycology 65:1−60

    doi: 10.3114/sim.2010.65.01

    CrossRef   Google Scholar

    [16]

    Hyde KD, Jones EBG, Liu JK, Ariyawansa H, Boehm E, et al. 2013. Families of Dothideomycetes. Fungal Diversity 63:1−313

    doi: 10.1007/s13225-013-0263-4

    CrossRef   Google Scholar

    [17]

    de Gruyter J, Woudenberg JHC, Aveskamp MM, Verkley GJM, Groenewald JZ, et al. 2013. Redisposition of phoma-like anamorphs in Pleosporales. Studies in Mycology 75:1−36

    doi: 10.3114/sim0004

    CrossRef   Google Scholar

    [18]

    Verkley GJM, da Silva M, Wicklow DT, Crous PW. 2004. Paraconiothyrium, a new genus to accommodate the mycoparasite Coniothyrium minitans, anamorphs of Paraphaeosphaeria, and four new species. Studies in Mycology 50:323−35

    Google Scholar

    [19]

    Verkley GJM, Dukik K, Renfurm R, Göker M, Stielow JB. 2014. Novel genera and species of coniothyrium-like fungi in Montagnulaceae ( Ascomycota). Persoonia: Molecular Phylogeny and Evolution of Fungi 32:25−51

    doi: 10.3767/003158514X679191

    CrossRef   Google Scholar

    [20]

    Sutton BC. 1980. The Coelomycetes. Fungi imperfecti with pycnidia, acervuli and stromata. Kew: Commonwealth Mycological Institute

    [21]

    Wijayawardene NN, Hyde KD, Wanasinghe DN, Papizadeh M, Goonasekara ID, et al. 2016. Taxonomy and phylogeny of dematiaceous coelomycetes. Fungal Diversity 77:1−316

    doi: 10.1007/s13225-016-0360-2

    CrossRef   Google Scholar

    [22]

    Crous PW, Summerell BA, Shivas RG, Romberg M, Mel'nik VA, et al. 2011. Fungal Planet Description Sheets 92–106. Persoonia: Molecular Phylogeny and Evolution of Fungi 27:130−62

    doi: 10.3767/003158511X617561

    CrossRef   Google Scholar

    [23]

    Crous PW, Wingfield MJ, Burgess TI, Hardy GS, Crane C, et al. 2016. Fungal Planet Description Sheets: 469–557. Persoonia: Molecular Phylogeny and Evolution of Fungi 37:218−403

    doi: 10.3767/003158516X694499

    CrossRef   Google Scholar

    [24]

    Crous PW, Wingfield MJ, Burgess TI, Carnegie AJ, Hardy GESJ, et al. 2017. Fungal Planet Description Sheets: 625–715. Persoonia: Molecular Phylogeny and Evolution of Fungi 39:270−467

    doi: 10.3767/persoonia.2017.39.11

    CrossRef   Google Scholar

    [25]

    Hyde KD, Dong Y, Phookamsak R, Jeewon R, Bhat DJ, et al. 2020. Fungal diversity notes 1151–1276: taxonomic and phylogenetic contributions on genera and species of fungal taxa. Fungal Diversity 100:5−277

    doi: 10.1007/s13225-020-00439-5

    CrossRef   Google Scholar

    [26]

    Quaedvlieg W, Verkley GJM, Shin HD, Barreto RW, Alfenas AC, et al. 2013. Sizing up Septoria. Studies in Mycology 75:307−90

    doi: 10.3114/sim0017

    CrossRef   Google Scholar

    [27]

    Crous PW, Groenewald JZ. 2017. The genera of fungi—G 4: Camarosporium and Dothiora. IMA Fungus 8:131−52

    doi: 10.5598/imafungus.2017.08.01.10

    CrossRef   Google Scholar

    [28]

    Thambugala KM, Wanasinghe DN, Phillips AJL, Camporesi E, Bulgakov TS, et al. 2017. Mycosphere notes 1–50: Grass ( Poaceae) inhabiting Dothideomycetes. Mycosphere 8:697−796

    doi: 10.5943/mycosphere/8/4/13

    CrossRef   Google Scholar

    [29]

    Wanasinghe DN, Hyde KD, Jeewon R, Crous PW, Wijayawardene NN, et al. 2017. Phylogenetic revision of Camarosporium ( Pleosporineae, Dothideomycetes) and allied genera. Studies in Mycology 87:207−56

    doi: 10.1016/j.simyco.2017.08.001

    CrossRef   Google Scholar

    [30]

    Valenzuela-Lopez N, Cano-Lira JF, Guarro J, Sutton DA, Wiederhold N, et al. 2018. Coelomycetous Dothideomycetes with emphasis on the families Cucurbitariaceae and Didymellaceae. Studies in Mycology 90:1−69

    doi: 10.1016/j.simyco.2017.11.003

    CrossRef   Google Scholar

    [31]

    Yuan HS, Lu X, Dai YC, Hyde KD, Kan YH, et al. 2020. Fungal diversity notes 1277–1386: taxonomic and phylogenetic contributions to fungal taxa. Fungal Diversity 104:1−266

    doi: 10.1007/s13225-020-00461-7

    CrossRef   Google Scholar

    [32]

    Li WJ, McKenzie EHC, Liu JK, Bhat DJ, Dai DQ, et al. 2020. Taxonomy and phylogeny of hyaline-spored coelomycetes. Fungal Diversity 100:279−801

    doi: 10.1007/s13225-020-00440-y

    CrossRef   Google Scholar

    [33]

    Wijayawardene NN, Hyde KD, Bhat DJ, Camporesi E, Schumacher RK, et al. 2014. Camarosporium-like species are polyphyletic in Pleosporales; introducing Paracamarosporium and Pseudocamarosporium gen. nov. in Montagnulaceae. Cryptogamie, Mycologie 35:177−98

    doi: 10.7872/crym.v35.iss2.2014.177

    CrossRef   Google Scholar

    [34]

    Wijayawardene NN, Hyde KD, Dai DQ, Sánchez-García M, Goto BT, et al. 2022. Outline of Fungi and fungus-like taxa - 2021. Mycosphere 13:53−453

    doi: 10.5943/mycosphere/13/1/2

    CrossRef   Google Scholar

    [35]

    Crous PW, Braun U, Schubert K, Groenewald JZ. 2007. Delimiting Cladosporium from morphologically similar genera. Studies in Mycology 58:33−36

    doi: 10.3114/sim.2007.58.02

    CrossRef   Google Scholar

    [36]

    Pem D, Hongsanan S, Doilom M, Tibpromma S, Wanasinghe DN, et al. 2019. https: //www. dothideomycetes. org: an online taxonomic resource for the classification, identification, and nomenclature of Dothideomycetes. Asian Journal of Mycology 2:287−97

    doi: 10.5943/ajom/2/1/19

    CrossRef   Google Scholar

    [37]

    Senanayake IC, Rathnayaka AR, Marasinghe DS, Calabon MS, Gentekaki E, et al. 2020. Morphological approaches in studying fungi: collection, examination, isolation, sporulation and preservation. Mycosphere 11:2678−754

    doi: 10.5943/mycosphere/11/1/20

    CrossRef   Google Scholar

    [38]

    Atlas RM. 2009. Experiencing displacement: Using art therapy to address xenophobia in South Africa. Development 52:531−36

    doi: 10.1057/dev.2009.74

    CrossRef   Google Scholar

    [39]

    Jayasiri SC, Hyde KD, Ariyawansa HA, Bhat DJ, Buyck B, et al. 2015. The Faces of Fungi database: fungal names linked with morphology, phylogeny and human impacts. Fungal Diversity 74:3−18

    doi: 10.1007/s13225-015-0351-8

    CrossRef   Google Scholar

    [40]

    Index Fungorum. 2022. www.indexfungorum.org/names/names.asp (Accessed on 28th September 2022)

    [41]

    Dissanayake AJ, Bhunjun CS, Maharachchikumbura SS, Liu JK. 2020. Applied aspects of methods to infer phylogenetic relationships amongst fungi. Mycosphere 11:2652−76

    doi: 10.5943/mycosphere/11/1/18

    CrossRef   Google Scholar

    [42]

    White TJ, Bruns T, Lee S, Taylor J. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenies. In PCR protocols: a guide to methods and applications, eds. Innis MA, Gelfand DH, Sninsky JJ, White TJ. San Diego: Academic Press. pp. 315–22. https://doi.org/10.1016/B978-0-12-372180-8.50042-1

    [43]

    Vilgalys R, Hester M. 1990. Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172:4238−46

    doi: 10.1128/jb.172.8.4238-4246.1990

    CrossRef   Google Scholar

    [44]

    Rehner SA, Samuels GJ. 1994. Taxonomy and phylogeny of Gliocladium analyzed from nuclear large subunit ribosomal DNA sequences. Mycological Research 98:625−34

    doi: 10.1016/s0953-7562(09)80409-7

    CrossRef   Google Scholar

    [45]

    Hall TA. 1999. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series 1:95−98

    Google Scholar

    [46]

    Katoh K, Rozewicki J, Yamada KD. 2019. MAFFT online service: multiple sequence alignment, interactive sequence choice and visualization. Briefings in Bioinformatics 20:1160−66

    doi: 10.1093/bib/bbx108

    CrossRef   Google Scholar

    [47]

    Capella-Gutiérrez S, Silla-Martínez JM, Gabaldón T. 2009. trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25:1972−73

    doi: 10.1093/bioinformatics/btp348

    CrossRef   Google Scholar

    [48]

    Nguyen LT, Schmidt HA, von Haeseler A, Minh BQ. 2014. IQ-TREE: A Fast and Effective Stochastic Algorithm for Estimating Maximum-Likelihood Phylogenies. Molecular Biology and Evolution 32:268−74

    doi: 10.1093/molbev/msu300

    CrossRef   Google Scholar

    [49]

    Chernomor O, von Haeseler A, Minh BQ. 2016. Terrace aware data structure for phylogenomic inference from supermatrices. Systematic Biology 65:997−1008

    doi: 10.1093/sysbio/syw037

    CrossRef   Google Scholar

    [50]

    Miller MA, Pfeiffer W, Schwartz T. 2012. The CIPRES science gateway: enabling high-impact science for phylogenetics researchers with limited resources. Proceedings of the 1st Conference of the Extreme Science and Engineering Discovery Environment - Chicago, Illinois (2012.07.16–2012.07.20): Bridging from the extreme to the campus and beyond (XSEDE'12), Association for Computing Machinery, USA. pp. 1–8. https://doi.org/10.1145/2335755.2335836

    [51]

    Darriba D, Taboada GL, Doallo R, Posada D. 2012. jModelTest 2: more models, new heuristics and parallel computing. Nature Methods 9:772

    doi: 10.1038/nmeth.2109

    CrossRef   Google Scholar

    [52]

    Miller MA, Pfeiffer W, Schwartz T. 2010. Creating the CIPRES Science Gateway for inference of large phylogenetic trees. 2010 Gateway Computing Environments Workshop (GCE), New Orleans, LA, USA, 2010. USA: IEEE. pp. 1–8. https://doi.org/10.1109/GCE.2010.5676129

    [53]

    Rambaut A. 2012. FigTree. v. 1.4. 0. http://tree.bio.ed.ac.uk/software/figtree/

    [54]

    Grondona I, Monte E, Garcia-Acha I, Sutton B. 1997. Pyrenochaeta dolichi: an example of a confusing species. Mycological Research 101:1404−8

    doi: 10.1017/s0953756297004206

    CrossRef   Google Scholar

    [55]

    Stewart RB. 1957. An undescribed species of Pyrenochaeta on soybean. Mycologia 49:115−17

    doi: 10.1080/00275514.1957.12024619

    CrossRef   Google Scholar

    [56]

    Mohanty NN. 1958. An undescribed species of Pyrenochaeta on Dolichos biflorus Linn. Indian Phytopathology 8:85−87

    Google Scholar

    [57]

    Hartman GL, Sinclair JB. 1988. Dactuliochaeta, a new genus for the fungus causing red leaf blotch of soybeans. Mycologia 8:696−706

    doi: 10.2307/3807721

    CrossRef   Google Scholar

    [58]

    Hartman G, Murithi HM. 2022. Coniothyrium glycines (red leaf blotch). CABI Compendium

    doi: 10.1079/cabicompendium.17687

    CrossRef   Google Scholar

    [59]

    Chethana KW, Manawasinghe IS, Hurdeal VG, Bhunjun CS, Appadoo MA, et al. 2021. What are fungal species and how to delineate them? Fungal Diversity 109:1−25

    doi: 10.1007/s13225-021-00483-9

    CrossRef   Google Scholar

    [60]

    Pem D, Jeewon R, Chethana KWT, Hongsanan S, Doilom M, et al. 2021. Species concepts of Dothideomycetes: classification, phylogenetic inconsistencies and taxonomic standardization. Fungal Diversity 109:283−319

    doi: 10.1007/s13225-021-00485-7

    CrossRef   Google Scholar

    [61]

    Nag Raj TR. 1993. Coelomycetous anamorphs with appendage-bearing Conidia. Vancouver, Canada: Mycologue Publications

    [62]

    Crous PW, Wingfield MJ, Cheewangkoon R, Carnegie AJ, Burgess TI, et al. 2019. Foliar pathogens of eucalypts. Studies in Mycology 94:125−298

    doi: 10.1016/j.simyco.2019.08.001

    CrossRef   Google Scholar

    [63]

    Hawksworth DL, Lücking R. 2017. Fungal diversity revisited: 2.2 to 3.8 million species. Microbiology Spectrum 5:5.4.10

    doi: 10.1128/microbiolspec.FUNK-0052-2016

    CrossRef   Google Scholar

    [64]

    Hyde KD, Jeewon R, Chen YJ, Bhunjun CS, Calabon MS, et al. 2020. The numbers of fungi: is the descriptive curve flattening? Fungal Diversity 103:219−71

    doi: 10.1007/s13225-020-00458-2

    CrossRef   Google Scholar

    [65]

    Bhunjun CS, Niskanen T, Suwannarach N, Wannathes N, Chen YJ, et al. 2022. The numbers of fungi: are the most speciose genera truly diverse? Fungal Diversity 114:387−462

    doi: 10.1007/s13225-022-00501-4

    CrossRef   Google Scholar

    [66]

    Lücking R, Aime MC, Robbertse B, Miller AN, Aoki T, et al. 2021. Fungal taxonomy and sequence-based nomenclature. Nature Microbiology 6:540−48

    doi: 10.1038/s41564-021-00888-x

    CrossRef   Google Scholar

    [67]

    Species Fungorum. 2022. www.speciesfungorum.org/Names/Names.asp (Accessed on 14/12/2022)

    [68]

    Elsebai MF, Kehraus S, Lindequist U, Sasse F, Shaaban S, et al. 2011. Antimicrobial phenalenone derivatives from the marine-derived fungus Coniothyrium cereale. Organic & Biomolecular Chemistry 9:802−8

    doi: 10.1039/c0ob00625d

    CrossRef   Google Scholar

    [69]

    Elsebai MF, Nazir M, Kehraus S, Egereva E, Ioset KN, et al. 2012. Polyketide skeletons from the marine alga-derived fungus Coniothyrium cereale. European Journal of Organic Chemistry 2012:6197−203

    doi: 10.1002/ejoc.201200700

    CrossRef   Google Scholar

  • Cite this article

    Wijesinghe SN, Calabon MS, Xiao Y, Jones EBG, Hyde KD. 2023. A novel coniothyrium-like genus in Coniothyriaceae (Pleosporales) from salt marsh ecosystems in Thailand. Studies in Fungi 8:6 doi: 10.48130/SIF-2023-0006
    Wijesinghe SN, Calabon MS, Xiao Y, Jones EBG, Hyde KD. 2023. A novel coniothyrium-like genus in Coniothyriaceae ( Pleosporales) from salt marsh ecosystems in Thailand. Studies in Fungi 8:6 doi: 10.48130/SIF-2023-0006

Figures(2)  /  Tables(4)

Article Metrics

Article views(5957) PDF downloads(1335)

ARTICLE   Open Access    

a novel coniothyrium-like genus in Coniothyriaceae ( Pleosporales) from salt marsh ecosystems in Thailand

Studies in Fungi  8 Article number: 6  (2023)  |  Cite this article

Abstract: In this study, a novel coniothyrium-like genus Coniothyrioides is introduced to Coniothyriaceae based on a fresh fungal collection from salt marsh habitats in Thailand. Coniothyrium-like taxa are taxonomically controversial and have been classified into different families in Pleosporales such as Didymosphaeriaceae ( Alloconiothyrium and Paraconiothyrium), Coniothyriaceae ( Coniothyrium) and Didymellaceae ( Microsphaeropsis). However, our novel genus shares similar morphology to some key characters in Coniothyriaceae in having dark, globose pycnidia, uni-locular conidiomata, a central ostiole, a peridium of textura angularis cells, and doliiform conidiogenous cells with a periclinal thickening at the apex, while conidial morphologies are diverse. The presence of setae arising from the outer peridial wall is the main difference between Coniothyrioides and other closely related Coniothyriaceae genera. Phylogenetically, LSU-SSU-ITS sequence analyses confirm the placement of this novel genus as a distinct lineage within Coniothyriaceae. Species boundaries were defined, based on morphology and multi-gene phylogenetic analyses using maximum likelihood and Bayesian inference analyses. The comprehensive descriptions and micrographs are provided. Our findings expand the taxonomic knowledge of Ascomycota in salt marsh ecosystems.

    • In coastal ecosystems, salt marsh habitats are common and consist of diverse halophytes, grasses, herbs, and shrubs, as well as microorganisms [ 1, 2] . These species-rich ecosystems are highly productive, and investigating fungal diversity in these habitats is important while many areas are still being explored [ 2, 3] . Researchers are currently studying the taxonomy of fungi in marine and semi-marine environments and increasing the number of known taxa recorded [ 46] . Ascomycota was identified as the dominant group in world salt marsh ecosystems, including the highest diversity in Pleosporales, Dothideomycetes [ 2] . Coniothyriaceae is a pleosporalean family with a large number of terrestrial taxa, while taxa associated with salt marsh vegetation have rarely been reported [ 2] .

      Coniothyriaceae was established by Cooke [ 7] to accommodate Coniothyrium species. The type genus and species of Coniothyriaceae are Coniothyrium Corda and C. palmarum Corda, respectively [ 810] . This family was previously linked to Leptosphaeriaceae [ 11] and this was followed by several authors [ 1216] . Subsequently, molecular data analyses for phoma-like asexual morphs were performed by de Gruyter et al. [ 17] based on LSU and ITS sequence data and revealed that C. palmarum is phylogenetically distant from Leptosphaeriaceae and closely related to Coniothyriaceae. de Gruyter et al. [ 17] reinstated Coniothyriaceae as a distinct family in Pleosporales and transferred several Phoma and Pyrenochaeta species into this family. Thus, the morphological variations of Coniothyrium species were expanded by the addition of more characters, such as setose pycnidia and conidiogenesis with elongated conidiophores [ 17] . Several authors later updated the placements of many Coniothyrium species with generic level changes and novel genera placed in different families [ 18, 19] .

      Verkley et al. [ 19] studied morphology and phylogenetic relationships of coniothyrium-like and closely related taxa. These species are coelomycetous and characterized in having pycnidial or stromatic conidiomata and small, subhyaline to pigmented, 1- or 2-celled conidia [ 19] . The phenotypic plasticity of these coelomycetous species has made their taxonomic placements uncertain and, thus the majority of them have been placed in Coniothyrium [ 1921] . Both Coniothyrium and coniothyrium-like species were identified as polyphyletic within Pleosporales and recent taxonomic treatments were mainly treated with combined morphology and molecular data analyses [ 12, 16, 17, 19, 2132] . Coniothyrium sensu stricto is characterized by 1-septate conidia and grouping in Coniothyriaceae [ 17, 1921, 33] . Currently, Coniothyriaceae consists of five genera, such as Coniothyrium, Foliophoma Crous , Neoconiothyrium Crous, Ochrocladosporium Crous & U. Braun and Staurosphaeria Rabenh. (≡ Hazslinszkyomyces Crous & R.K. Schumach.) [ 10, 34] .

      Coniothyriaceae members have been identified as pathogens that cause necrotrophic and leaf spots on leaves, and saprobes on dead branches [ 10, 17] . The sexual morph is characterized in having cucurbitaria-like, black, globose ascomata, short central ostiole, textura angularis peridium cells, branched, septate, cellular pseudoparaphyses, 8-spored, cylindrical, bitunicate asci and muriform, ellipsoidal ascospores that are initially hyaline and brown at maturity [ 10] . Asexual morphs are coelomycetous and sometimes differentiated with phoma-like, camarosporium-like, coniothyrium-like, or cladosporium-like asexual characters. They are characterized in having dark, globose, pycnidial conidiomata, with central, sometimes papillate ostiole, cells of textura angularis or textura globulosa in the conidiomatal wall, hyaline macroconidiogenous and microconidial cells and conidia. Conidial morphology is varied as macroconidia and microconidia. Macroconidia are ellipsoid, red-brown, and septation is from the central transverse septum to muriformly septate, while microconidia are hyaline, globose to ellipsoid and aseptate [ 9, 10, 24, 27, 35, 36] .

    • In this study, we aim to expand the taxonomy of fungi associated with dead plant hosts in salt marsh ecosystems. We investigate salt marsh habitats in Thailand to collect fungal specimens and isolate them to find out the taxonomic novelties. Morphological illustrations, comprehensive descriptions, and multi-gene phylogenetic analyses are provided to confirm the placement of new findings.

    • Fungal specimens were collected from salt marsh habitats in Pranburi Province, Thailand, 2021. Samples were preserved in sterile Ziploc bags in the laboratory and incubated at room temperature 25 °C. Rehydrated specimens were observed to identify fungal fruiting bodies and macro-morphology was observed by using a Motic SMZ 168 compound stereomicroscope. Micro-morphologies (e.g., conidiomata, conidiogenous cells, conidia) were examined from hand-sectioned structures using a Nikon ECLIPSE 80i compound stereomicroscope, equipped with a Canon 600D digital camera. The measurements of photomicrographs were obtained using Tarosoft (R) Image Frame Work version 0.9.7. Images were edited with Adobe Photoshop CS6 Extended version 13.0.1 software (Adobe Systems, San Jose, California, USA).

      Single-spore isolation was carried out as described by Senanayake et al. [ 37] . Germinated spores were aseptically transferred into fresh malt extract agar medium (MEA) prepared in 50% or 100% concentrations of sterilized natural seawater [ 38] . Culture plates were incubated at 25 °C for six weeks and inspected every week. Herbarium specimens are preserved at Mae Fah Luang University Herbarium (MFLU) in Chiang Rai, Thailand. All living cultures are deposited at Mae Fah Luang Culture Collection (MFLUCC). Facesoffungi and Index Fungorum numbers for new taxa were obtained [ 39, 40] .

    • The methodologies for DNA extraction, PCR, gel electrophoresis, and sequencing were followed, as detailed in Dissanayake et al. [ 41] . The genomic DNA was extracted from fresh mycelium using the E.Z.N.A Fungal DNA Mini Kit- D3390-02 (Omega Bio-Tek, USA) following the guidelines of the manufacturer. DNA sequences were obtained for the internal transcribed spacer region (ITS1, 5.8S, ITS2), the small subunit (SSU), and the large subunit (LSU) of the nuclear ribosomal RNA gene. PCR thermal cycle programs for each locus region are presented in Table 1. Purification and sequencing were outsourced to the Bio Genomed Co. LTD laboratory (Biogenomed Co., Thailand). Newly generated sequences were submitted to NCBI GenBank ( www.ncbi.nlm.nih.gov/genbank).

      Table 1.  Gene regions, primers, and PCR thermal cycle programs used in this study, with corresponding reference(s).

      Genes/loci PCR primers (forward/reverse) PCR conditions Reference (s)
      ITS and LSU ITS5/ITS4 and LR0R/LR5 94 °C; 2 min (95 °C; 30 s, 55 °C; 50 s, 72 °C; 90 s) × 35 thermal cycles, 72 °C; 10 min [ 4244]
      SSU NS1/NS4 95 °C; 3 min (95 °C; 30 s, 55 °C; 50 s, 72 °C; 30 s) × 35 thermal cycles, 72 °C; 10 min [ 42]

      BioEdit v 7.0.9.0 program [ 45] was used to check the quality of the newly generated sequence chromatograms. For primary identification, contig sequences were checked with BLAST searches in NCBI. Sequences for phylogenetic analyses were downloaded from GenBank ( Table 2) following Hyde et al. [ 25] . Each gene matrix was aligned with MAFFT version 7 [ 46] with default parameters and manually adjusted for improvement where necessary using BioEdit v. 7.2 [ 45] . The trimAl v1.4 software was used for the automated removal of spurious sequences or poorly aligned regions in each single gene alignment, and gappyout was selected as the automated trimming method [ 47] . Two separate phylogenetic analyses were conducted: Maximum Likelihood (ML) and Bayesian Inference (BI). LSU, SSU, and ITS concatenated dataset was analyzed for Coniothyriaceae and selected families in Pleosporales.

      Table 2.  Taxa used in the phylogenetic analyses and their GenBank accession numbers. Sequences of new taxon generated in this study are in blue-bold and type strains are in black-bold.

      Species Strain/voucher number GenBank accession numbers
      ITS LSU SSU
      Amarenographium ammophilae MFLUCC 16–0296 KU848196 KU848197 KU848198
      Ascochyta pisi CBS 126.54 GU237772 EU754137 EU754038
      Bipolaris microstegii CBS 132550 NR_120160 NG_042690 NA
      Bipolaris victoriae CBS 327.64 NR_147489 NG_069233 NA
      Comoclathris arrhenatheri MFLUCC 15–0465 NR_165855 NG_068240 NG_068374
      Coniothyrioides thailandica MFLUCC 22-0193 OQ023276 OQ023277 OQ025050
      Coniothyrium carteri LG1401 MS6E KX359604 KX359604 NA
      Coniothyrium cereale CBS 157.78 MH861123 JX681080 NA
      Coniothyrium chiangmaiense MFLUCC 16–0891 KY568987 KY550384 KY550385
      Coniothyrium dolichi CBS 124140 JF740183 GQ387611 GQ387550
      Coniothyrium glycines CBS 124455 JF740184 GQ387597 GQ387536
      Coniothyrium palmarum CBS 400.71 AY720708 EU754153 AY720712
      Coniothyrium palmarum CBS 758.73 NA JX681085 EU754055
      Coniothyrium sp. B9-10-9 MW764153 NA NA
      Coniothyrium sp. P16-10-4 MW764259 NA NA
      Coniothyrium telephii CBS 188.71 JF740188 GQ387599 GQ387538
      Coniothyrium telephii CBS 856.97 JF740189 GQ387600 GQ387539
      Coniothyrium telephii UTHSC:DI16–189 LT796830 LN907332 NA
      Coniothyrium triseptatum MFLU 19–0758 NR_171948 NG_073674 NA
      Curvularia heteropogonis CBS 284.91 JN192379 JN600990 NA
      Didymella azollae A1 MT514913 MT514910 NA
      Foliophoma camporesii MFLUCC 18–1129 KY929151 KY929181 NA
      Foliophoma fallens CBS 161.78 KY929147 GU238074 GU238215
      Foliophoma fallens CBS 284.70 KY929148 GU238078 GU238218
      Libertasomyces myopori CPC 27354 NR_145200 NG_058241 NA
      Libertasomyces platani CPC 29609 NR_155336 NG_059744 NA
      Libertasomyces quercus CBS 134.97 NR_155337 DQ377883 NA
      Melnikia anthoxanthii MFLUCC 14–1010 NA KU848204 KU848205
      Neoconiothyrium hakeae CPC 27616 KY173397 KY173490 NA
      Neoconiothyrium hakeae CPC 27620 KY173398 KY173491 NA
      Neoconiothyrium multiporum CBS 353.65 NR_111617 JF740268 NA
      Neoconiothyrium multiporum CBS 501.91 JF740186 GU238109 GU238225
      Neoconiothyrium persooniae CBS 143175 NR_156386 NG_058509 NA
      Neoconiothyrium viticola CPC 36397 NR_165929 NG_068326 NA
      Neoplatysporoides aloeicola CPC 24435 NR_154230 NG_058160 NA
      Ochrocladosporium elatum CBS 146.33 EU040233 EU040233 NA
      Ochrocladosporium frigidarii CBS 103.81 NR_156512 NG_064123 NA
      Phaeosphaeria chiangraina MFLUCC 13–0231 KM434270 KM434280 KM434289
      Phaeosphaeria musae MFLUCC 11–0133 KM434267 KM434277 KM434287
      Phaeosphaeria thysanolaenicola MFLUCC 10–0563 NR_155642 NG_069236 NG_063559
      Phaeosphaeria oryzae CBS 110110 NR_156557 NG_069025 NG_061080
      Phaeosphaeriopsis dracaenicola MFLUCC 11–0157 NR_155644 NG_059532 KM434292
      Pleospora herbarum MFLUCC 14-0920 KY659560 KY659563 KY659567
      Querciphoma carteri CBS 101633 JF740180 GQ387593 GQ387532
      Querciphoma carteri CBS 105.91 JF740181 GQ387594 GQ387533
      Querciphoma carteri Gv5 MT819903 NA NA
      Querciphoma carteri UASWS2031 MN833930 NA NA
      Shiraia bambusicola NBRC 30753 AB354987 AB354968 NA
      Shiraia bambusicola NRBC 30771 AB354990 AB354971 NA
      Shiraia bambusicola NRBC 30772 AB354991 AB354972 NA
      Staurosphaeria aloes CBS 136437 KF777142 KF777198 NA
      Staurosphaeria aloes CPC 21572 NR_137821 NG_067283 NA
      Staurosphaeria aptrootii CBS 483.95 NR_155186 NA NA
      Staurosphaeria lycii CPC 30998 KY929150 KY929180 NA
      Staurosphaeria lycii CPC 31014 KY929151 KY929181 NA
      Staurosphaeria rhamnicola MFLUCC 17–0813 MF434200 MF434288 MF434376
      Staurosphaeria rhamnicola MFLUCC 17–0814 NR_154461 MF434289 NG_063659
      Stemphylium vesicarium CBS 191.86 MH861935 MH873624 GU238232
      NA: Sequences not available in GenBank.
    • In the phylogenetic analyses, maximum likelihood (ML) was executed using IQ-Tree web server ( http://iqtree.cibiv.univie.ac.at/) with bootstrap support obtained from 1,000 pseudoreplicates [ 48, 49] . Bayesian Inference (BI) analysis was performed on the CIPRES Science Gateway portal under MrBayes on XSEDE (3.2.7a) [ 50] . Six simultaneous Markov chains were run for 1,000,000 generations, and trees were sampled every 1,000 th generation, ending the run automatically when the standard deviation of split frequencies dropped below 0.01. The best nucleotide substitution models for each genetic marker were evaluated using jModelTest2 on XSEDE in the online CIPRES Portal ( www.phylo.org/portal2) [ 51, 52] . The best-fit models under the AIC criterion were revealed to be GTR+I+G for ITS and LSU regions while GTR+I for SSU region. Phylogenetic trees were visualized with FigTree version 1.4.0 [ 53] and edited in Microsoft PowerPoint (2019).

    • The combined LSU, SSU, and ITS alignment was used to construct the final phylogenetic analysis ( Fig. 1) of maximum likelihood (ML) and Bayesian inference (BI).

      Figure 1. 

      Phylogram generated from maximum likelihood analysis based on combined LSU, SSU, and ITS sequenced data. Fifty-eight strains were included in the combined sequence analyses, which comprised 2251 characters with gaps (LSU = 800, SSU = 948, ITS = 503). Single gene analyses were also performed, and topology and clade stability were compared from the combined gene analyses. Ascochyta pisi Lib. (CBS 126.54) and Didymella azollae E. Shams, F. Dehghanizadeh, A. Pordel & M. Javan-Nikkhah (A1) were used as the outgroup taxa. The final ML optimization likelihood is -10163.644. The matrix included 494 distinct alignment patterns including undetermined characters. Estimated base frequencies were obtained as follows: A = 0.245, C = 0.219, G = 0.274, T= 0.262; substitution rates AC = 2.73290, AG = 3.93954, AT = 2.73290, CG = 1.0, CT = 7.93321, GT = 1.0 and the gamma distribution shape parameter α = 0.439534. Bootstrap support values for ML (first set) equal to or greater than 75% and BYPP equal to or greater than 0.95 are given above or below the nodes. The strains from the current study are in red bold and the type strains are in black bold. The scale bar represents the expected number of nucleotide substitutions per site.

    • Coniothyrioides Wijes., M.S. Calabon, E.B.G. Jones & K.D. Hyde, gen. nov.

      Index Fungorum number: 555045; Facesoffungi number: 13901 Fig. 2

      Etymology – Resembling Coniothyrium taxa

      Saprobic on a submerged decaying wood in salt marsh ecosystems. Sexual morph: Undermined. Asexual morph: Coelomycetous. Forming conspicuous, round to irregular, black pycnidia. Conidiomata semi-immersed, erumpent through the host substrate, globose to subglobose, solitary, scattered to aggregated, uni-loculate, ostiolate, covered in setae, rigid when dehydrated, black. Setae originated from the outermost layers of conidiomatal wall, divergent, brown, with hyaline apex, septate, smooth-walled, uniformly wide from base to apex. Conidiomatal wall composed of several layers, from outer to inner layers black, dark brown, pale brown to hyaline cells of textura angularis. Conidiophores reduced to conidiogenous cells. Conidiogenous cells lining the inner cavity, doliiform to subcylindrical, smooth-walled, hyaline, enteroblastic, phialidic conidiogenesis with periclinal thickening at the apex. Conidia solitary, ellipsoidal to obovoid, rounded at the apex, aseptate, initially hyaline, becoming pale to dark brown at maturity, smooth-walled, sometimes finely verruculose, with smaller guttules at young and indistinct at maturity.

      Figure 2. 

      Coniothyrioides thailandica sp. nov. (MFLU 22-0276, holotype). (a) & (b) Appearance of conidiomata on a submerged decaying woody substrate. (c) Longitudinal section of conidioma. (d) Conidiomatal wall. (e) The appearance of setae. (f) & (g) Conidiogenous cells with developing conidia. (h) Conidia. Scale bars: a = 200 μm, b = 100 μm, c = 50 μm, d = 20 μm, e, h = 10 μm, f, g = 5 μm.

      Type species – Coniothyrioides thailandica

      Note – Coniothyrioides gen. nov. is a monotypic genus associated with decaying woody substrates in salt marsh habitats in central Thailand. This genus is characterized in having pycnidial conidiomata with the cells of textura angularis wall surrounded by distinct setae, doliiform to subcylindrical, hyaline conidiogenous cells, and ellipsoidal to obovoid, aseptate and hyaline to brown conidia. Based on some conidial characteristics such as aseptate, hyaline to brown conidia the genus shares similar morphologies to coniothyrium-like taxa [ 19] , by ellipsoidal to subcylindrical conidia sharing similar characters to Coniothyrium and Neoconiothyrium [ 9, 16, 20, 24] . However, other accepted genera in Coniothyriaceae differ from this genus in conidial morphologies: Foliophoma has only hyaline conidia except for F. camporesii D. Pem & K.D. Hyde; Hazslinszkyomyces has muriformly septate conidia [ 27] ; Ochrocladosporium has cladosporium-like conidia [ 35] . Moreover, phylogenetically Coniothyrioides forms a distinct lineage within Coniothyriaceae ( Fig. 1). Coniothyrium carteri (Gruyter & Boerema) Verkley & Gruyter (LG1401_MS6E) was the closest species based on BLAST result of ITS (94.33% similarity) and C. telephii (Allesch.) Verkley & Gruyter (UTHSC:DI16-189) was the closest species LSU sequence data (99.31% similarity) and sequences are lacking for SSU in the GenBank. The genus is known from its asexual morph and the sexual morphology was not observed.

      In our phylogenetic analyses, Foliophoma species were grouped outside of Coniothyriaceae with closer to Libertasomycetaceae and Pleosporaceae species. Foliophoma was introduced by Crous & Groenewald [ 27] to accommodate F. fallens (Sacc.) Crous, in Coniothyriaceae based on the parsimony analyses of single LSU and ITS sequence data. Foliophoma camporesii was later introduced based on morphology and maximum likelihood analyses of LSU-SSU- ITS sequence data by Hyde et al. [ 25] . Based on morphology, Foliophoma species share similar characteristics to the species of Coniothyriaceae in having dark brown conidiomata, conidial wall with textura angularis cells, phialidic conidiogenesis sometimes with periclinal thickening or percurrent proliferation, and mainly ellipsoidal shaped conidia. However, the type species of the genus, F. fallens differs other Coniothyriaceae taxa in having eustromatic conidiomata. Based on this taxonomic uncertainty, more fresh collections with additional coding genes are required to clarify the accurate placement of Foliophoma.

      Coniothyrioides thailandica Wijes., M.S. Calabon, E.B.G Jones & K.D. Hyde, sp. nov.

      Index Fungorum number: 555050; Facesoffungi number: 13902

      Etymology – The name reflects the county Thailand, from where the species was isolated.

      Saprobic on a submerged and decaying woody substrate. Sexual morph: Undermined. Asexual morph: Coelomycetous. Conidiomata 150–200 μm high, 100–150 μm diam. (x̄ = 160 × 130 µm), pycnidial, semi-immersed, erumpent through the host substrate, globose to subglobose, solitary, scattered to aggregated, uni-loculate, ostiolate, covered by setae, rigid when dehydrated, black. Setae 3–5 µm wide, originating from the outermost layers of conidiomatal wall, divergent, brown, with hyaline apex, septate, smooth-walled, uniformly wide from base to apex. Conidiomatal wall 15–20 µm wide, equally thickened, composed of several layers, outermost layers dark brown to black, towards inside pale brown to hyaline cells of textura angularis, surrounded by setae. Conidiophores reduced to conidiogenous cells. Conidiogenous cells 4–5 μm long × 2.5–3.5 μm wide, lining the inner cavity, doliiform to subcylindrical, smooth-walled, hyaline, enteroblastic, phialidic conidiogenesis with periclinal thickening at the apex. Conidia 3–5 × 2.5–3 μm ( ¯x = 4.5 × 2.7 µm, n = 20), solitary, ellipsoidal to obovoid, rounded at the apex, aseptate, initially hyaline, becoming pale to dark brown at maturity, smooth-walled, sometimes finely verruculose, with smaller guttules at young and indistinct at maturity.

      Culture characteristics – On MEA, colony circular with a filamentous margin, reaching 40–45 mm diam. in 25 d at 25 °C, light gray from above, brown from center becoming light gray in the margin below, surface rough, dry, flat, with dense mycelia, edge filiform.

      Material examined – Thailand, Pranburi Province, on a submerged decaying wood, 23 March 2021, Mark S. Calabon, SPAR26 (MFLU 22-0276, holotype), ex-type living cultures, MFLUCC 22-0193.

      GenBank numbers – ITS = OQ023276, LSU = OQ023277, SSU = OQ025050.

      Notes – Coniothyrioides thailandica sp. nov. shares morphological characters with other representatives in Coniothyriaceae in having pycnidial, globose, uni-locular conidioma with a central ostiole, peridial wall with the cells of textura angularis, and doliiform to subcylindrical conidiogenous cells, phialidic conidiogenesis with a periclinal thickening at the apex. The synopsis of asexual morphological characters for the generic types of the family including their hosts and localities is presented in Table 3. Based on the presence of conidiomatal setae, our species (MFLU 22-0276) resembles Neoconio thyrium [ 24] . In addition, our species resembles Foliophoma camporesii (MFLU 17-1006) in having hyaline to brown and aseptate conidia but differs in having larger conidiomata (150–200 × 100–150 vs 40–47 × 40–69 μm) and the presence of setae on the wall ( Table 3). Phylogenetically, our strain (MFLUCC 22-0193) formed an independent lineage within Coniothyriaceae with 94% ML and 0.99 BI statistical support ( Fig. 1). The base pair differences between our stain and the strains represent type species of other genera in Coniothyriaceae are listed ( Table 4). Thus, the evidence based on both morphology and phylogeny, we establish Coniothyrioides as a new genus in Coniothyriaceae with C. thailandica as the type species.

      Table 3.  Synopsis of asexual morphological characters of related genera of Coniothyriaceae.

      Species Conidiomata (µm) Conidiomata wall (µm) Conidiogenous cells (µm) Conidia (µm) Habitat(s) and host(s) Locality Reference
      Coniothyrioides thailandica (holotype: MFLU 22-0276) 150–200 high, × 100–150 diam., pycnidial, semi-immersed, erumpent, dark brown to black, globose to subglobose,
      uni-locular, ostiolate
      15–20 wide, black, dark brown to hyaline cells of textura angularis,
      Brown, septate setae (3–5 µm wide,) with hyaline apex
      4–5 long × 2.5–3.5 wide, hyaline, doliiform to subcylindrical, enteroblastic, phialidic conidiogenesis with periclinal thickening 3–5 × 2.5–3, ellipsoidal to obovoid, aseptate, rounded at apex, initially hyaline, becoming pale to dark brown at maturity On decaying wood in salt marsh habitat Thailand This study
      Coniothyrium
      palmarum (CBS 400-71)
      Immersed, dark brown, globose, pale to uni-locular brown, thick-walled cells of textura angularis hyaline, phialidic conidiogenesis, doliiform to cylindrical Subcylindrical, spherical, ellipsoid or broadly clavate, 0(–1)-septate, apex obtuse, brown, base truncate, sometimes minute marginal frill On Chamaerops humilis ( Arecaceae) Italy [ 16]PP,
      [ 20]GN
      Foliophoma fallens (holotype: CBS 284.70) 120–250 wide, eustromatic, globose, uni-multi locular,
      1–3 ostiolate
      3–6 layers,
      brown textura angularis
      5–7 × 4–5, hyaline, phialidic conidiogenesis with thickening or proliferation at apex, dolliform to subcylindrical,
      periclinal
      (5–)5.5–6(–7) × (3–)4(–5),

      broadly ellipsoidal, aseptate, hyaline, guttulate or granular, apex obtuse, base truncate to bluntly rounded
      Leaf spot on Nerium oleander ( Apocynaceae) Italy [ 27]
      *Foliophoma camporesii (holotype: MFLU 17-1006) 40–47 × 40–69, pycnidial, immersed to semi-immersed, globose to subglobose, ellipsoidal or irregular, carbonaceous 15–40, 1–2-layered of cells of textura angularis 2–4 × 2–3, hyaline, globose to short cylindrical, phialidic conidiogenesis with periclinal thickening or percurrent proliferation at apex 2–6 × 3–5, ovoid to ellipsoidal, aseptate, hyaline when immature, brown at maturity On dead stems of Maclura pomifera ( Moraceae) Italy [ 25]
      Hazslinszkyomyces aloes (≡ Camarosporium aloes: ex-type - CPS 21572) 250 diam, pycnidial, erumpent, brown, globose, central ostiolate 3–6 layers of brown textura angularis
      5–10 × 4–5, hyaline,
      ampulliform to doliiform, apex with several inconspicuous percurrent proliferation,
      (9–)11–13(–14) × (4–)6–7(–8), ellipsoid, initially hyaline, aseptate, becoming pale brown, subcylindrical
      to clavate or obovoid with 3 transverse eusepta, constricted at median septum or not, apex obtuse, base bluntly rounded to truncate
      Dead bark of Aloe dichotoma ( Xanthorrhoeaceae) South Africa [ 27]
      Neoconiothyrium persooniae (ex-type CPC 32021 = CBS 143175) 100–200 diam, superficial, ellipsoid to obpyriform, 1–2 papillate ostioles, 10–15 diam, with or without setae 3–6 layers, hyaline textura angularis 5–8 × 4–5, hyaline, doliiform to ampulliform, phialidic, with periclinal thickening or percurrent proliferation (5–)6–7(–8) × 3(–4), ellipsoid to subclavate, aseptate, initially hyaline medium brown, becoming cylindrical and at times 1-septate, apex subobtuse, base bluntly rounded On leaves of Persoonia laurina subsp. laurina ( Proteaceae) Australia [ 24]
      Ochrocladosporium elatum (CBS 146.33) Integrated as lateral peg-like loci on hyphal
      cells, or erect, subcylindrical, up to 25 µm long, 2.5–4 µm wide,
      with 1–3 terminal loci, occasionally lateral, 1–1.5 µm wide
      Ramoconidia, 10–40 × 3–5, subcylindrical to ellipsoid, hyaline to pale brown, 0(–1)-septate, giving rise to branched
      chains of conidia that are subcylindrical to ellipsoid, aseptate, (7–)8–10(–14) × (3–)4(–4.5), olivaceous brown
      Wood pulp Sweden [ 35]
      '–' observed morphologies on cultures, therefore conidiomata and wall characters are not recorded. '*' species which is not represent a generic type. GN- based on the generic description. PP- based on the photographic plate provided.

      Table 4.  The base pair comparisons of our strain (MFLUCC 22-0193) with the strains representing type species of other genera in Coniothyriaceae.

      Species Strain LSU SSU ITS
      Coniothyrium palmarum CBS 400-71 14/800 (1.75%) 3/948 (0.3%) 69/487 (14.10%)
      Foliophoma fallens CBS 284.70 8/800 (1%) 2/948 (0.2%) 66/497 (13.27%)
      Hazslinszkyomyces aloes CPC:21572 6/800 (0.75%) 52/497 (10.46%)
      Neoconiothyrium persooniae CBS:143175 20/800 (2.5%) 49/497 (9.85%)
      Ochrocladosporium elatum CBS 146.33 13/800 (1.62%) 53/497 (10.66%)

      Members of Coniothyriaceae have high morphological plasticity and it is not adequate to use only morphology for identification at the genus level. Coniothyrium dolichi (Mohanty) Verkley & Gruyter (≡ Pyrenochaeta dolichi Mohanty, CBS 124140) and C. glycines (R.B. Stewart) Verkley & Gruyter (≡ P. glycines R.B. Stewart, CBS 124455) form a separate clade within Coniothyriaceae ( Fig. 1). Based on the morphology observed from corn meal agar medium (CMA) by Grondona et al. [ 54] , C. dolichi differs to our species by having two types of conidiogenous cells including discrete, ampulliform conidiogenous cells and integrated, cylindrical conidiogenous cells on filiform, septate conidiophores in the same conidioma, while our species has doliiform to subcylindrical conidiogenous cells and conidiophores are reduced to conidiogenous cells [ 54] . The pycnidial conidiomata and the ostiole of C. dolichi covered by dark brown, septate setae resembles our species and conidia are aseptate, ellipsoid, and hyaline with more or fewer guttules while our species has brown conidia at maturity [ 54] . The original description of C. dolichi, mentioned that conidia were greenish-yellow in mass similar to coniothyrium-like conidia, as well as to our species [ 55]. Also, a monodictys-like synanamorph was reported in C. dolichi based on its dark brown to black, dictyosporous conidia by differs from our species [ 17, 54, 56] . Coniothyrium glycines produces monophialidic, ampulliform, conidiogenous cells and aseptate, ellipsoidal conidia (4–8 × 1–3 µm), while our species has doliiform to subcylindrical conidiogenous cells [ 57, 58] . The unique character of C. glycines is well-defined, dark brown to black, melanized sclerotia covered with setae which differs from our species and other Coniothyrium taxa. Based on the multi-gene phylogeny provided by de Gruyter et al. [ 17] and the results of our study, the placements of these two species were confirmed in Coniothyriaceae. Also, Coniothyrium triseptatum Dayar., Thyagaraja & K.D. Hyde (MFLU 19-0758) creates a separate lineage in Coniothyriaceae ( Fig. 1) and only sexual morph was reported for this fungus. Therefore, we could not compare the morphology of C. triseptatum with our species [ 31] .

    • In this study, we introduced the novel genus Coniothyrioides in Coniothyriaceae, with C. thailandica as the type species, following the guidelines and major criteria for defining generic and species boundaries in Dothideomycetes by Chethana et al. [ 59] and Pem et al. [ 60] . The coelomycetous asexual morph of Coniothyrioides was associated with the decaying and submerged wood in the salt marsh habitats. Traditionally, morphology is used to delimit coelomycetes by considering the characteristics of conidiomata, conidiophores, conidiogenesis, and conidia including host associations [ 20, 32, 61] . However, accurate taxonomy of most coniothyrium-like species is challenging because of their simplicity, plasticity, and morphological variations [ 19] . In our study, genera in Coniothyriaceae differ in some conidial morphologies. For instance, Coniothyrium is characterized by aseptate to 1-septate, ellipsoidal to clavate or cylindrical, brown conidia [ 9, 17, 62] , Foliophoma with aseptate, ovoid ellipsoidal, only hyaline or hyaline-brown conidia, and Hazslinszkyomyces ellipsoidal to obovoid, transversely and muriformly septate, uniformly brown conidia [ 27] . Neoconiothyrium species have aseptate or 1-septate, ellipsoid to subclavate or subcylindrical, hyaline to medium brown conidia [ 24] while Ochrocladosporium species have cladosporium-like pale brown, aseptate or 1-septate conidia that occurring in branched chains [ 35] . Coniothyrioides thailandica is characterized by aseptate, ellipsoidal to obovoid, hyaline to pale or dark brown conidia. However, the phylogenetic analyses in this study reveal these morphological differences are not strong enough for generic delimitation of the family. Some characteristics of Coniothyrioides overlap with those of other accepted genera in the family, such as the conidiogenous cell morphology of Coniothyrium and Foliophoma which have doliiform to cylindrical or subcylindrical, hyaline, phialidic conidiogenesis with periclinal thickening and conidia show aseptate, ellipsoid-associated shapes, and hyaline to brown pigmentation ( Table 3).

      The number of fungi was estimated at between 2.2 to 3.8 million [ 63] , with about 100,000-150,000 known species and fungus-like taxa [ 64 66] . There are 151,834 species listed in Species Fungorum [ 67] . An up-to-date online database ( https://coelomycetes.org/) for coelomycetes is being implemented [ 21, 32] . As coniothyrium-like taxa are frequently collected and morphologically similar, it is likely that they will remain unidentified. Therefore, it is to be expected that if molecular data are incorporated in morpho-taxonomic studies of these groups, will help identify many more novel taxa. This has occurred in other genera, which are plant pathogens and ecologically more important. According to Bhunjun et al. [ 65] Coniothyrium is one of the most speciose genera listed in Species Fungorum in 2021 and studies of coniothyrium-like taxa may yield more novel species. A few records of Coniothyriaceae taxa have been identified in salt marsh ecosystems, such as Coniothyrium obiones Jaap (India and Portugal) and as unidentified Coniothyrium species (USA) [ 2] . However, Wanasinghe et al. [ 29] referred the placement of C. obiones in Neocamarosporiaceae based on multi-gene phylogeny. In marine habitats, C. cerealis E. Müll. was isolated as an alga-derived fungus in the Baltic Sea by Elsebai et al. [ 68, 69] .

      In this study, we discussed the morphology and multi-gene phylogenetic analyses results of our new collection to verify its identity and phylogenetic placement in Coniothyriaceae. Based on ecological and geographical data on salt marsh fungi, we noted lack records of Coniothyriaceae worldwide (see Calabon et al. [ 2] ). Thus, we propose that additional collections be conducted in order to identify other Coniothyriaceae taxa and improve our understanding of fungal diversity in salt marsh ecosystems, a topic that is currently understudied.

      • S.N. Wijesinghe would like to thank Mae Fah Luang University on behalf of the Graduate Studies Support Grant (Grant No. Oh 7702(6)/125) for financial support and Mushroom Research Foundation (MRF), Thailand. Further, S.N. Wijesinghe would like to thank Dr. Udeni Jayalal, Dr. Samantha C. Karunarathna, and Dr. Gothamie Weerakoon for their precious advice during this study. M.S. Calabon is grateful to the Department of Science and Technology – Science Education Institute (DOST-SEI). K.D. Hyde would like to thank the National Research Council of Thailand (NRCT) grant 'Total fungal diversity in a given forest area with implications towards species numbers, chemical diversity and biotechnology' (Grant No. N42A650547).

      • Kevin David Hyde is the Editorial Board member of Journal Studies in Fungi. He is blinded from reviewing or making decisions on the manuscript. The article was subject to the journal's standard procedures, with peer-review handled independently of Kevin David Hyde and his research groups.

      • Copyright: © 2023 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 (4) References (69)
  • About this article
    Cite this article
    Wijesinghe SN, Calabon MS, Xiao Y, Jones EBG, Hyde KD. 2023. A novel coniothyrium-like genus in Coniothyriaceae (Pleosporales) from salt marsh ecosystems in Thailand. Studies in Fungi 8:6 doi: 10.48130/SIF-2023-0006
    Wijesinghe SN, Calabon MS, Xiao Y, Jones EBG, Hyde KD. 2023. A novel coniothyrium-like genus in Coniothyriaceae ( Pleosporales) from salt marsh ecosystems in Thailand. Studies in Fungi 8:6 doi: 10.48130/SIF-2023-0006

Catalog

  • About this article

/

DownLoad:  Full-Size Img  PowerPoint
Return
Return