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Article

First Insights into the Biochemical and Metabolomic Characterization of Marine Fungi Isolated from Salt Marshes of an Argentine Estuary

by
Florencia Biancalana
1,2,
Marios Psarianos
3,
Paula C. P. Bueno
2,
Romina Sanchez
1,4,
Oliver K. Schlüter
3 and
Anna Fricke
2,*
1
Centro de Recursos Naturales Renovables de la Zona Semiárida (CERZOS), Universidad Nacional del Sur (UNS-CONICET), Bahía Blanca B8000, Argentina
2
Leibniz Institute of Vegetable and Ornamental Crops, Theodor-Echtermeyer-Weg 1, 14979 Großbeeren, Germany
3
Department Systems Process Engineering, Leibniz Institute for Agricultural Engineering and Bioeconomy (ATB), Max-Eyth-Allee 100, 14469 Potsdam, Germany
4
Departamento de Biología, Bioquímica y Farmacia, Universidad Nacional del Sur (UNS), Bahía Blanca B8000, Argentina
*
Author to whom correspondence should be addressed.
J. Mar. Sci. Eng. 2026, 14(12), 1106; https://doi.org/10.3390/jmse14121106
Submission received: 10 March 2026 / Revised: 5 June 2026 / Accepted: 8 June 2026 / Published: 16 June 2026
(This article belongs to the Section Marine Biology)
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Versions Notes

Abstract

The Bahía Blanca marshes are highly productive ecosystems that act as carbon sinks and support a diverse community of halophytic plants, including species of Distichlis, Spartina, Sarcocornia, among others. Marine fungi inhabit these ecosystems and play important roles in the decomposition of organic matter and the production of valuable biopolymers and metabolites. Despite their potential ecological and commercial importance, little is known about the identity and chemical profiles of these fungi associated with marine environments and halophytic plants in the Bahía Blanca estuary. To expand current knowledge of marine fungi in this system, three fungal species associated with marine waters and Sarcocornia perennis plants were isolated and identified: Aspergillus iizukae, Stemphylium sp. (Ascomycota), and Mucor sp. (Mucoromycota). Their biomass was analyzed to determine its biochemical and metabolic composition. Stemphylium sp. exhibited the highest protein (30.24 g/100 g dry mass) and lipid (2.65 g/100 g dry mass) contents, whereas Mucor sp. showed the highest levels of total sugars (26.13 g/100 g dry mass) and glucosamine (17.30 g/100 g dry mass). Aspergillus iizukae produced the greatest diversity of secondary metabolites. These findings provided a preliminary characterization of fungal species from this region and highlighted their potential for future biotechnological and industrial applications.

1. Introduction

Salt marshes are productive ecosystems that act as valuable blue carbon sinks, supporting a diverse halophyte plant community, including species of the genera Atriplex, Juncus, Sarcocornia, Spartina, among others, that serve as habitat and nursery grounds for a wide range of marine organisms [1,2]. Marine fungi represent one of the many groups associated with these ecosystems and constitute a diverse but still poorly studied group, with 2254 species documented to date [3], occurring in algae, halophytes, marine animals, driftwood, sediments and the water column [4,5,6]. They perform diverse and ecologically relevant roles, acting as saprotrophs that recycle organic matter (e.g., through the degradation of plankton-derived biopolymers); as symbionts that facilitate the life of other organisms (e.g., macroalgae); or as pathogens that may reduce host survival infecting phytoplankton, halophytes, and animals [7]. As primary decomposers, particularly the saprotrophic species, they secrete extracellular enzymes that degrade complex organic matter, releasing essential nutrients such as nitrogen and phosphorus and making them available to plants and plankton [8]. In salt marsh ecosystems, marine fungi have been reported to associate with various halophyte species [2,9] and to decompose lignocellulosic substrates more effectively than bacteria [10].
Beyond their ecological roles, marine fungi represent a promising source of natural bioactive compounds with potential applications in a broad range of industries, including food, agriculture, cosmetics, biotechnology, biochemistry, and pharmaceuticals [7,10]. Recent studies have focused on the chemical diversity of compounds produced by marine fungi, highlighting that fungal biomass is rich in proteins, fatty acids, and vitamins, thus making it a potentially valuable resource for human and animal nutrition [11]. Additionally, filamentous marine fungi produce polysaccharides, enzymes (e.g., cellulases, chitinases) and a wide array of secondary metabolites such as terpenes, steroids, polyketides, peptides, and alkaloids [10]. These compounds exhibit antimicrobial, anticancer, antiviral, antioxidant, and anti-inflammatory activities [6]. Consequently, marine fungi may complement conventional animal nutrition, such as fish feed [12], and contribute to human health-related applications [13,14]. In addition to their nutritional and biotechnological relevance, marine fungi have been widely studied for their ability to degrade synthetic polymers, such as polyethylene [15], and to produce natural polymers, including chitin, chitosan and glucans [16,17,18].
Composed of a diverse assemblage of marine macrophytes, the Bahía Blanca Estuary (BBE) salt marsh system generates large amounts of detritus. This material may decompose in situ or be transported by tides and wind to adjacent waters, where it plays a key role in estuarine food webs [19]. In addition, the BBE salt marsh supports a diverse fungal community (e.g., Alternaria, Cladosporium, Drechslera, Penicillium, and Trichoderma) [20] that contributes to organic matter degradation and biopolymer production that is dispersed throughout the estuarine system [20,21]. Despite the recognized ecological importance and commercial potential of these fungi, knowledge about the identity, function, and productivity of biopolymers, enzymes, and metabolites produced by marine fungi associated with salt marsh macrophytes remains scarce in the BBE.
This study forms part of an ongoing investigation into the fungal diversity of the BBE, in which fungal isolates were recovered and identified at the generic level. Based on these findings, selected taxa were chosen to explore the biotechnological potential of this natural resource, particularly for applications in food, health, and wastewater treatment. In this context, the main objective of the present study was to provide a preliminary biochemical and metabolomic characterization of the three fungal isolates obtained from the salt marshes of the BBE. Additionally, selected biological and ecological aspects of these fungi are discussed.

2. Materials and Methods

2.1. Research Area

The Bahía Blanca estuary (38°42′–39°25′ S, 61°50′–62°22′ W) (BBE) is located at the southwest of the Buenos Aires province in Argentina (Figure 1a–e). It is a mesotidal, temperate, and turbid estuary with a total area of 2300 km2 at high tide [22]. The salinity range is between 17.9 and 41.3, considering that it depends on temperature, as well as seasonal rainfall and winds [23]. The BBE is subject to high anthropogenic impact due to human settlements, commercial ports, petrochemical and chemical industries, and dredging projects that affect both the dynamics of the seston and its planktonic components [24]. The multi-use reserve known as Sustainable Water Resources Management (2023), where vulnerable species grow and are used as a nursery, occupies most of the BBE and overlaps with the activities [24]. The present study was carried out in the Rosales Port (RP) (38°55′ S, 62°04′ W), located on the northern bank of the BBE’s main channel (Figure 1a–e). The Rosales Port (RP) has moderate levels of nutrients and heavy metals, as well as high biological contamination due to the presence of Escherichia coli [25], indicating the impact of wastewater discharge from Punta Alta (population approximately 60,000). The BBE the intertidal zone is covered by saltmarshes, with an area of 836 km2 (Figure 1b–e). Spartina alterniflora mudflats (196 km2) consist of scattered plants up to 20 cm tall [26] to dense stands over 1 m tall in areas subject to high sedimentation rates [27] (Figure 1e). Intertidal mudflats, composed of Sarcocornia perennis, are less represented (72 km2) (Figure 1c,e), and the dominant species usually form circular mounds and may be associated with Heterostachys ritteriana or S. densiflora at higher altitudes [28]. The RP has an extensive intertidal zone (1000 m wide) with low slopes [29] consisting of S. perennis in the upper marsh and Spartina alterniflora in the lower marsh (Figure 1c–e).

2.2. Collection and Isolation of Fungal Species

The filamentous fungi were isolated from marine water and S. perennis from the RP in April 2024 [30,31]. The water sample was collected at the surface of the water coast using a 5 L sterilized glass bottle. The S. perennis specimens were extracted from the intertidal zone using sterile bags. Both types of samples were transported in ice-filled coolers and stored in a refrigerator until processing.
Seawater was collected from a pristine area (Monte Hermoso—38°59′24″ S and 61°17′28″ W—~100 km from the BBE); it was filtered using a GTTP polycarbonate filter of 0.2 µm pore size (GAMAFIL, Martinez, Buenos Aires, Argentina) and then used to prepare the culture media. The salinity of this water was ~30, the same as that of the water sample. All the strains were cultivated in Malt Extract Agar (MEA) and Potato dextrose broth (PDA) at 25 °C and in darkness for species identification [32].
To obtain filamentous fungi colonies, marine water samples were filtered through a 25 mm diameter type GTTP polycarbonate filter, with a 0.2 µm pore size (GAMAFIL). This filter was then placed on a Petri dish containing PDA with antibiotic (streptomycin 20 µg/mL) to observe the growth of fungal colonies at 25 °C. Between 1 and 3 days of incubation, each colony formed was individually transferred to a fresh PDA Petri dish to obtain pure fungal strains (replicates 2; Figure S1a,b). In the case of S. perennis samples, fungal isolates were obtained after examinations of the plants under a stereomicroscope by directly transferring spores with a sterile needle onto agar media in Petri dishes. When necessary, subcultures were transferred (between one and four times) to fresh PDA plates to maintain pure cultures (Figure S1c). These procedures were performed under sterile conditions and in a controlled environment.

2.3. Species Identification-Morphological and Phylogenic Analyses

After one and two weeks of incubation in darkness at 25 °C, the characteristics of the colonies were recorded and photographed. Microscopic observations were conducted on fungal structures mounted in tap water or 100% lactic acid and observations were performed using a LEICA DM2000 equipped with a LEICA ICC50 HD camera (Leica Microsystems, Wetzlar, Germany). Fungal structural measurements, including diameter and length, were obtained with the LEICA LAS EZ V3.4.0 software.
For DNA extraction, fungi were transferred into 100 mL of liquid medium (PDB Cultivation Medium-Potato dextrose broth with antibiotic (streptomycin 20 µg/mL) in 200 mL Erlenmeyer flacks and incubated in a shaker incubator (Eppendorf®Innova®S44i Shaker incubated, AC/DC input 120 V) at 25 °C only for 3–5 days. Then, pellets were obtained after transferring ≤100 mg wet weight of fungi to an Eppendorf of 1500 mL and grinding them with Retsch MM 400. The extraction of DNA (deoxyribonucleic acid) was done following the protocol of DNeasy® Plant Mini Kit (QIAGEN). Seven microliters of the PCR product were separated by electrophoresis in 1.5% agarose gel (25 min, 100 V, in 1.0 × TAE buffer). Gels were stained with SYBR Safe DNA Gel Stain (Invitrogen) and visualized under UV in a transilluminator. PCR products were sequenced at Eurofins Genomics Europe Shared Services GmbH.
ITS regions of the rDNA were amplified using the following primers: ITS1-ITS4 [33]. Other fungal gene regions were used for molecular identification was beta-tubulin 2 (TUB2) with primer set T1/Bt2b [34,35]. Sequences generated from the different primers of the different genes (ITS and TUB2) were analyzed along with sequences from previous studies that were retrieved from GenBank, visualized by the accession numbers shown in Figure 2. Data sets were initially aligned by using MEGA-X v. 10.2. [36] and manually corrected elsewhere using BioEdit v. 7.2.5 sequence alignment editor [37]. Gaps were treated as missing data. The species Aspergillus cervinus Massee (NRRL 5025) was selected as the outgroup species [38]. Nucleotide substitution models were determined with Partition Finder [39,40]. The concatenated ITS and TUB2 datasets were analyzed with K80+G and TRNEF+I model for each subset respectively. Phylogenetic trees were generated using maximum likelihood (ML) analyses. The ML trees were generated using IQ-TREE 2 [41].

2.4. Cultivation for Biomass Production

To increase fungal biomass, cultivation was carried out on potato dextrose broth (PDB) 24 g/L, salt 20 g/L (NaCl as a principal macro-element—Tropic Marin Classic, Tropic Marin®, Wartenberg, Germany) and antibiotic (streptomycin 20 µg/mL). The normal pH for a PDB salt medium is typically between 6.0 and 7.0. For each fungus, three 1 L Erlenmeyer flacks for pouring with 400 mL of cultivation medium. The fungi were incubated in a shaker incubated (Eppendorf®Innova®S44i Shaker incubated, AC/DC input 120 V) at 25 °C, 120 rpm and in dark until the pellets were formed about 3–5 days for one month. All fungal strains were cultivated under identical conditions. This procedure was performed on the three filamentous fungi in duplicate.
At the end of the 3- or 5-day cultivation period, the fungal biomass was separated carefully by a sterile strainer (pore size 1 mm2) and washed several times with distilled water. After washing, fungal biomass was freeze dried in falcon tubes of 50 mL for three days to obtain stable dry weight. The dry fungi biomass was homogenized in the falcon tubes as obtained powder and placed in plastic bags that were vacuum sealed until samples were processed. This procedure was done between four and eight times in the month, depending on the growth of each fungus. Total dry biomass was used in different amounts to determine chemical composition.

2.5. Biochemical Characterization

The moisture content of the samples was quantified by drying the samples inside a drying oven at 105 °C for 3 h and measuring the weight difference due to the moisture loss (NFTA 2.1.4 official moisture method). The lipid content was measured gravimetrically after a Soxhlet extraction with petroleum ether for 6 h [42]. The protein content was estimated by quantifying nitrogen with the Kjeldahl method and calculating the crude protein content with the conversion factor 6.25 [43]. Total ash was measured gravimetrically after placing the samples inside a furnace operating at 550 °C for 4 h [44] (DIN 38414-EN 12879). Glucose, disaccharides, xylose, and arabinose were analyzed after acid hydrolysis of the samples using sulfuric acid. The analysis was performed using high-performance liquid chromatography (HPLC) (Knauer, Berlin, Germany) coupled with a refractive index detector (RI-71, SHODEX, Yokohama, Japan). The separation of sugars was performed with a Eurokat H column (300 mm × 8 mm × 10 μm; Knauer, Berlin, Germany), and samples were eluted with 5 mM H2SO4 at 0.8 mL/min [45]. The total content of non-chitin sugars was calculated as a sum of the number of individual sugars. Each chemical analysis was done for duplication (n = 2).
The total content of glucosamine and N-acetyl-glucosamine was measured using the method of [46]. Briefly, samples were ground to fine powder using a Retsch laboratory mill (Retsch GmbH, Haan, Germany) and 10 mg of each sample was hydrolyzed with 0.3 mL of sulfuric acid 72% at room temperature for 90 min. After the addition of 8.7 mL water, the samples were further hydrolyzed at 121 °C for 1 h. While the samples were still hot, an aliquot of 0.5 mL of hydrolysate was taken for the glucosamine estimation, while another aliquot of 0.5 mL from each sample was taken to be used as a blank. After the hydrolysates were cooled down, 0.5 mL of a NaNO2 solution 1 M was added to the aliquots for the glucosamine determination, while 0.5 mL of distilled water were added to the respective aliquot for each sample that would be used as a blank. After this step, both the mixture intended for glucosamine determination and the one intended as a blank were treated in the same way. The samples were capped and incubated at room temperature for 6 h. The caps were removed, and the samples were further incubated overnight. Afterwards, 0.5 mL of a 12% ammonium sulfamate solution was added to each sample and the samples were incubated for 4 min. Then an aliquot of 0.5 mL of a 0.5% MBTH solution was added to the samples that were then incubated for 1 h at room temperature. Finally, an aliquot of 0.5 mL of a 0.5% FeCl3 was added to the samples that were incubated for 1 h again at room temperature. The samples were diluted to the appropriate dilution, and the absorbance was measured at 650 nm using a UV/Vis spectrometer. A standard glucosamine solution was used for the calibration curve at a concentration range of 3.15–11.15 µg/mL. The concentration of glucosamine was done for duplication (n = 2).

2.6. Secondary Metabolites Profiling

For the metabolite’s extraction, around 1 g of each fungus’s dried mass (two biological replicates of A. iizukae and Mucor sp. and one biological replicate of Stemphylium sp.), as well as the growth media (control media) were extracted with ethyl acetate. The samples were placed in 20 mL Falcon tubes, added alongside 15 mL of the solvent, vortexed in cycles of 1 min, and then centrifuged at 10,000 rpm for five minutes. After phase separation, the ethyl acetate phase was carefully collected and concentrated using a Speed Vac until dryness. The dried sample material was resuspended at the concentration of 10 mg/mL using MeOH/Water 1:1, with the aid of an ultrasonic bath in cold water for 30 s. Then samples were centrifuged at 4500 rpm for 5 min and filtered using 0.22 µm PTFE filters.
For metabolite analysis, samples were subjected to liquid chromatography coupled to mass spectrometry (UHPLC-DAD-ESI-MS/MS). The system consisted of a 1290 Infinity liquid chromatograph (Agilent Technologies, Santa Clara, CA, USA) coupled to a 6546 LC/Q-TOF mass spectrometer (Agilent Technologies). For each sample, a volume of 1 µL was injected into an InfinityLab Poroshell 120 C18 column (100 × 2.1 mm, 1.9 μm particle size, 120 Å pore size, Agilent Technologies) protected by a guard column of the same stationary phase. The column temperature was set to 40 °C and the flow rate at 0.4 mL·min−1. The mobile phase consisted of 0.1% formic acid in water (solvent A) and 0.1% formic acid in acetonirile (solvent B). Compounds were separated using a mobile-phase gradient starting at 15% of B from 0 to 3 min, then from 15 to 80% of B from 3 to 12 min, 80 to 98% of B from 12 to 15 min, keeping at 98% of B from 15 to 19 min, then from 98 to 2% of B from 19 to 19.1 min, and reconditioning at 2% of B from 19.1 to 22 min.
The mass spectrometric data was acquired in positive and negative ionization modes using an electrospray ionization source (Agilent Jet StreamESI, Santa Clara, CA, USA). The QTOF-MS operated in the data-dependent acquisition (DDA) mode. For both modes, the capillary voltage was set at 4 kV, the drying gas temperature at 270 °C, at a drying gas flow of 12 L·min−1. The nebulizer was set at 40 psi, sheath gas heater at 375 °C at a sheath gas flow of 12 L·min−1. The nozzle voltage (V) was set to 0, the fragmentor at 100, the skimmer at 65 and the Octopole RFPeak at 750. The full mass scan range was set from 60 to 1500 m/z for MS1 and MS2. The QTOF-MS operated in the data-dependent acquisition (DDA) mode. During DDA experiments, a maximum of 5 precursor ions were isolated and fragmented using collision-induced dissociation fragmentation energies of 10, 20 and 30 eV, at an acquisition rate/time of 5 spec/s. The intensity threshold was set at 200, number of precursors equal to 2, active exclusion enabled after 2 spectra and release after 0.5 min. The isolation and fragmentation settings were size and charge dependent, width 3–15 m/z and including charges state of 1z, 2z, and unknown. Accuracy of m/z values during data acquisition was guaranteed by continuous recalibration of the mass axis using reference ions including 121.0508, and 922.0098 Da for the positive acquisition mode and 112.9856 and 966.0007 for the negative acquisition mode.

2.7. Data Processing, Annotation and Visualization

Data for metabolic analyses were converted to. mzML using ProteoWizard msConvert (available at https://proteowizard.sourceforge.io/, installation 12 December 2024, version 3.0.24346-56bbd4d, installation 12 December 2024, version 3.0.24346-56bbd4d). Data extraction, processing, and matrix organization were made using Mzmine 4.3 (available at https://mzio.io/mzmine/, instalation 13 September 2024, version 4.3.0). For that, the positive and negative data acquisition modes were processed separately. Compound class predictions were performed using SIRIUS 6.0 (available at https://bio.informatik.uni-jena.de/software/sirius/, instalation 14 June 2023, version 5.7.3), uploading positive and negative mode data (.mgf) separatedly.
The metabolomic data were presented in the form of enriched classes to characterize the strains, as no biological activity studies were performed. The LC-MS (Liquid chromatography-mass spectrometry) data were acquired on MS1 and MS2, in both the positive and negative modes, for the chemical characterization described in the manuscript. Based on the MS2 spectral data, compound classes were predicted and processed using SIRIUS, aligning with the primary goal of obtaining a comprehensive characterization of the samples rather than a detailed metabolome annotation.
For data analysis, the features detected in the control media were subtracted from the samples. Overlapping and unique features detected in the samples were visualized using Venn diagrams (available at VIB/Ugent Bioinformatics & Evolutionary Genomics—https://bioinformatics.psb.ugent.be/webtools/Venn/, accessed at 29 April 2025). The chemical classification results from SIRIUS were visualized as sunburst plots using Excel (v. 2108, Microsoft® Excel® LTSC MSO 16.0.14332.21031, 64-bit). For that, the data matrices containing the positive and negative mode quantitative information (peak areas) obtained from Mzmine were combined with the output table from SIRIUS. The data points were sorted into one of the Natural Products pathways (inner ring), superclasses (middle ring) or compound classes (outer ring). Only probability scores greater than 0.7 were considered in the three classifications.

2.8. Statistical Analysis

Non-parametric statistical procedures were applied on biommass data (n ≥ 6), because the data did not meet the assumptions of normality and homoscedasticity. The Kruskal–Wallis H test was applied only to determine differences in fungal biomass. The significance level of the test was p = 0.05. Protein, total sugar, lipid, and ash content were measured in duplicate for each fungus (n = 2) to provide preliminary compositional data for fungi from an unexplored region of Argentina. For this reason, the concentrations of protein, total sugars, lipids, and ash for each fungus were presented as means and standard error (SE = Devest/ n 2 ) and plotted in Excel.

3. Results

3.1. Fungi Identity

Three filamentous fungi were selected, among others (Table S1), for identification (one of them at species level and two at the genus level) from the Bahía Blanca Estuary, two belonging to the Ascomycota phylum: Aspergillus iizukae (Eurotiales, Aspergillaceae) and Stemphylium sp. (Pleosporales, Pleosporaceae); and one to the Mucoromycota phylum: Mucor sp. (Mucorales, Mucoraceae). Two of them, A. iizukae and Mucor sp., were isolated from water samples along with Epicoccum sp. and Lasiodiplodia sp. (Table S1). On the other hand, Stemphylium sp. was found growing on S. perennis; the formation of fungal structures on the stems of this halophyte drew particular attention. The species found of the genus Aspergillus was selected for chemical composition analyses due to the abundance of previous studies reporting the isolation and characterization of compounds of biotechnological interest from this genus, thus enabling comparisons with other marine systems. In contrast, Mucor was selected based on the presence of chitosan as a major structural component of their cell walls (Table S1).
The determination to the species level of the Aspergillus strain, initially obtained from its morphology, was later supported by the phylogenetic analysis. The phylogenetic tree drawn from a combinate dataset of ITS and partial TUB2 (Figure 2) revealed a close relationship between the strain examined here and the ex-type of strain of A. iizukae (NRRL 3750) with good support. For Mucor sp. and Stempphylium sp., the use of ITS is limited to the genus level; in both cases, specific primers will be needed to confirm the species. The three descriptions, illustrations and comments are presented in the Supplementary Material (Figure S2).

3.2. Fungi Biomass Production and Biochemical Characterization

After one month of measuring the fungal biomass every 3 or 5 days, total dry mass was estimated for each fungus (Figure 3). After the growth period, Mucor sp. presented the highest amount of harvested dry biomass (2.75 ± 0.37 g d.m.) followed by A. iizukae (2.51 ± 0.38 g d.m.) and Stemphylium sp. (1.65 ± 0.06 g d.m.). Non-significant differences were detected in dry mass between fungi (Kruskal–Wallis H test, p ≤ 0.05).
Figure 4a shows the biochemical characterization of the three different fungi. Overall, Stemphylium sp. showed in sum the highest amount of the measured nutritional macromolecules (64.72 g/100 g d.m.). This was mainly due to the highest proteins (30.24 ± 0.32 g/100 g d.m.) and contents of lipids (2.65 ± 0.49 g/100 g d.m.) being measured in Stemphylium sp. Whereas total sugars contents were similar in Stemphylium sp. (15.17 ± 1.04 g/100 g d.m.) and A. iizukae (15.28 ± 7.52 g/100 g d.m.), the highest content was observed in Mucor sp. (26.13 ± 2.09 g/100 g d.m.) (Figure 4a).
Figure 4b shows the contents of sugars of three fungal species. All strains showed a very low amount of arabinose (<1%), while exhibiting great amounts of glucosamine, saccharose, glucose and fructose. Mucor sp. showed the overall highest concentrations of total sugar monomers, mainly due to high amounts of glucosamine (equal to 17.30 ± 1.56 g/100 g d.m.) and saccharose (6.73 ± 1.13 g/100 g d.m.) (Figure 4b). In contrast, almost no fructose was detected in Mucor sp., which peaked in A. iizukae (5.18 ± 4.62 g/100 g d.m.), whereas glucose was highest (5.65 ± 1.06 g/100 g d.m.) in Stemphylium sp. (Figure 4b).

3.3. Metabolomic Characterization

The highest metabolite diversity was observed in A. iizukae in both the positive and negative ionization modes, as visualized in the Venn diagrams (total number of features) and sunburst plots (Figure 5a,b and Figure 6a–c). It is important to note that the metabolite diversity shown in the sunburst plots was assessed based on predicted compound classes; therefore, no compound-level validation was performed in the present study.
In A. iizukae, the main putatively annotated chemical classes, based on class prediction, included alkaloids, polyketides, amino acids, and fatty acids (Figure 6a), whereas Stemphylium sp. and Mucor sp. were mainly characterized by fatty acids and carbohydrates (Figure 6b,c). Alkaloids, including pseudoalkaloids and tyrosine-derived alkaloids, were detected in all samples and were more readily detected in the positive ionization mode (Figure 6a–c). In contrast, polyketides, including aromatic, cyclic, and linear polyketides, macrolides, oligopeptides, polycyclic aromatic polyketides and polyethers, were more efficiently detected in negative ionization mode and were particularly abundant in A. iizukae (Figure 6a). Interestingly, amino acids and small peptides, which were also better detected in negative ionization mode, were mainly observed in Stemphylium sp. and Mucor sp. (Figure 6b,c).

4. Discussion

4.1. Initial Information on A. iizukae, Stemphylium sp. and Mucor sp. in the Bahía Blanca Estuary

The fungal strains studied in this research included A. iizukae and Stemphylium sp., both belonging to the phylum Ascomycota, and Mucor sp., which belong to the phylum Mucoromycota. Recently, filamentous fungal species from Alternaria, Aspergillus, Cladosporium, Drechslera, Penicillium, Ramichloridium, Rhacodiella, and Trichoderma genera were isolated from sediments of the BBE [20]. Mucor sp. was exclusively recovered from water samples, whereas Stemphylium sp. was only found associated with S. perennis from the RP. The three genera investigated in this study are commonly reported as inhabitants of salt marsh ecosystems [2].
Aspergillus is a very diverse genus, with more than 440 species classified into sections and series ranks, categories formally established by [47]. Its species are distributed world-wide, growing on plants, animals and fungal tissues, different types of indoor environments, soils and water samples, and in the air [48]. Marine environments have also been colonized by Aspergillus species found living in sediments, seawater, seaweed and marine animals. It has been associated with extreme marine environments, including those with high salinity (30%) [49,50]. Aspergillus iizukae is a ubiquitous species that has been reported on a wide variety of substrates, including soil, indoor air, cave sediments, herbivore excrement, food, peat soil, and salt marshes [38,50,51]. Thermal tolerance up to 37 °C (this study, [50]) and growth at 40 °C has also been reported by [38]. Particularly, A. iizukae was isolated from coastal saline soil in Kenli, Shandong Province, China, in August 2008 using an OSMAC (one strain, many compounds) approach [52].
Despite the cosmopolitan distribution of the genus Aspergillus, A. iizukae was isolated exclusively from seawater in the present study, despite temperature and salinity conditions in RP being suitable for fungal growth in both seawater and sediments. First, it is important to note that sampling was conducted at a single time point (April 2024), which may not fully reflect the temporal dynamics of the fungal community. Additionally, environmental factors other than temperature and salinity, such as dissolved oxygen, organic matter availability, heavy metal concentrations, and other pollutants, may influence fungal community structure and distribution [53]. In this context, the anoxic and highly polluted sediment conditions in the BBE may affect different ecological and biological aspects of A. iizukae, potentially limiting its occurrence in this substrate. Indeed, RP is a polluted area receiving both autochthonous and allochthonous organic matter, which contributes to oxygen depletion and anoxic conditions in the sediments [25]. Further studies addressing the temporal and spatial dynamics of fungal communities in the BBE would help to better understand the environmental factors shaping their distribution.
The genus Stemphylium was revised by Woudenberg and coll. who distinguished 28 species based on morphological characteristics and multigene molecular data combining partial ITS, gapdh, and cmdA gene regions [54]. Subsequent descriptions of new species have followed these taxonomic criteria. Currently, Stemphylium is recognized as a well-established monophyletic genus comprising more than one hundred saprophytic and plant pathogenic species distributed worldwide [55,56,57]. Until gapdh and cmdA sequences are obtained for the specimen isolated from S. perennis in the BBE, its specific identity will remain unresolved. Stemphylium is a fungal genus commonly found in salt marsh environments [2], and some species have demonstrated an ability to adapt to high salinity conditions [9]. Although generally regarded as parasitic or saprophytic, certain Stemphylium species may also establish mutualistic interactions with salt marsh vegetation, particularly under specific environmental conditions [9]. For instance, [58] demonstrated that inoculation of Salicornia (=Sarcocornia) with a Stemphylium isolate increased total biomass production and root nitrogen concentration under optimal salinity conditions (150 mM NaCl), suggesting that some species may behave as beneficial endophytes that promote host plant performance. Based on these observations, we propose that the Stemphylium species found in the present study may also behave as an endophyte of S. perennis, remaining latent in healthy tissues and expressing sporulation only when the host entered senescence. Notably, no fungal expression or sporulation attributable to Stemphylium was detected in healthy plant specimens examined during this study.
Mucor species are fungi commonly found in soils, freshwater environments, and decaying organic matter. Although they are less frequently reported in marine ecosystems [2], they have also been reported in salt marshes, particularly in areas with lower salinity or high organic matter accumulation [59]. In these environments, members of the genus have been recovered mainly from soils across different tidal levels, including the rhizosphere of salt marsh vegetation [2,9]. The sampling site examined in the present study receives substantial inputs of allochthonous organic matter, largely associated with wastewater discharges from the nearby city [19,60], which may partially explain the occurrence of Mucor at this location.
As mentioned previously, species belonging to the genera Aspergillus, Stemphylium, and Mucor have been reported in association with Spartina spp. and other halophytes worldwide, mainly exhibiting saprophytic, but occasionally endophytic, interactions [2,59]. These observations, together with the findings of the present study, suggest that the environmental conditions of temperature, salinity, and organic matter availability in the water, sediment, and halophyte-associated substrates of the BBE may provide suitable conditions for the establishment, development, and persistence of these fungi.

4.2. Biomass, Biochemical and Metabolites Characterization, and Potential Applications

Obtaining filamentous fungal biomass is essential for subsequent biochemical analyses. In the present study, the selected strains were cultured under the same temperature, salinity, and pH conditions using PDA as a carbon source and progressively increasing culture volumes. However, despite being exposed to identical culture conditions, the fungi did not produce the same biomass yields or concentrations of biochemical compounds, as has also been reported in previous studies [12,61,62]. In our study, Mucor sp. produced the greatest amount of biomass, followed by A. Iizukae and Stemphylium sp. Although no statistically significant differences were detected among strains, the total dry mass of each fungus (see Figure 3) suggests differences in biomass accumulation patterns, potentially reflecting species-specific growth responses that warrant further investigation.
Marine fungi are recognized as a valuable source of proteins, with several species exhibiting protein contents ranging from 28% to 63% (w/w) on a dry weight basis [11,12,61,62]. In the present study, protein contents were comparatively low for all strains analyzed, although proteins still represented the dominant macromolecular component of fungal biomass. Among the studied strains, Stemphylium sp. appeared to produce the highest protein content. Previous studies on Stemphylium species associated with halophytes have reported the production of exopolysaccharides (EPSs) under specific cultivation conditions, including pH 6–7 and salinity levels around 150 mM NaCl [10,63]. These EPSs may contain proteins associated with sugar polymers in their structure. Furthermore, all three strains evaluated in the present study have been reported to produce them [63]. Therefore, the culture conditions applied here may have favored EPS production by Stemphylium sp., which could partially explain its relatively higher protein content.
Beyond proteins, fungal biomass also contains a wide variety of sugars involved in metabolism and cell wall structure [64,65,66]. These sugars are often organized into polysaccharide structures, although some may also occur as monomeric compounds depending on the fungal species and physiological conditions. In the present study, glucosamine was detected in all strains, with the highest concentration observed in Mucor sp. Among fungal polysaccharides, chitin and glucans are major structural components of the cell wall. Typically, fungal cell walls are composed of a chitin–glucan complex adjacent to the plasma membrane and an outer layer predominantly consisting of glycoproteins [63]. The methodology used to quantify glucosamine involved biomass hydrolysis, during which chitin is deacetylated to chitosan and subsequently hydrolyzed to glucosamine [44]. Therefore, glucosamine concentration cannot be used to directly quantify chitin or chitosan content but may serve as an indirect indicator of the presence of these biopolymers. In this context, the higher glucosamine concentration observed in Mucor sp. may be associated with the predominance of chitin and chitosan as major structural components of the cell wall in Mucoromycota, in contrast to Ascomycota species, whose cell walls are typically dominated by chitin and β-glucans. [64,65]. Therefore, the differences observed among strains suggest distinct sugar composition profiles, particularly regarding glucosamine-related structural polymers. Several species of the Mucoromycota, including M. rouxii, M. circinelloides, and Rhizopus oryzae, have been widely recognized as fungal sources of chitin and chitosan [64,65]. Moreover, fungal-derived chitosan can be produced under controlled cultivation conditions, resulting in more homogeneous physicochemical properties than chitosan obtained through conventional chemical extraction methods [67]. In contrast, Stemphylium species have been reported to produce extracellular polysaccharides (EPSs), complex sugar-rich compounds involved in several biological functions [65]. For example, an endophytic Stemphylium species isolated in India was reported to produce mannoglucan-type polysaccharides [63], suggesting that a comparable glucan composition could also occur in the strain examined in the present study. All strains contained the sugars analyzed, and arabinose was also consistently detected at low concentrations.
Regarding fructose detection in A. iizukae, species of the genus Aspergillus are known to produce fructose through the degradation of plant inulin mediated by inulinase activity [68]. However, because all strains in the present study were cultured under identical conditions and in the absence of an inulin source, active fructose production through this pathway is unlikely to fully explain the observed concentrations. Moreover, the acid hydrolysis procedure used for sugar extraction (sulfuric acid) may have contributed to the breakdown of larger carbohydrate structures or glycoconjugates into smaller sugars, including fructose. Therefore, the fructose detected in A. iizukae may reflect hydrolysis-derived products and/or intracellular sugar pools retained within fungal hyphae rather than direct metabolic production under the culture conditions tested. The relatively high variability among replicates (n = 2; Figure 4) further suggests that these results should be interpreted cautiously.
With respect to lipid content, all strains exhibited very low levels (<3% on a dry basis). Lipid accumulation in fungi varies considerably among different taxa, typically ranging from 1% to 50% of dry biomass [11], and can also be strongly influenced by the developmental stage and culture conditions, including temperature, pH, carbon source, and inorganic salt availability [69]. Although fungal lipids have attracted considerable interest in various biotechnological applications, particularly biofuel production [70], the low levels detected in the present study suggest a limited potential for lipid-based applications under the culture conditions applied. However, given the recognized ability of Mucor species to produce high-value polyunsaturated fatty acids (PUFAs), including γ-linolenic acid (GLA) [71], further analyses of lipid composition could provide additional insight into the biotechnological potential of this strain.
The fungal strains investigated in the present study exhibited a biochemical composition consistent with that previously reported for fungal biomass, suggesting potential applicability in selected biotechnological contexts. Such applications may include the recovery of specific macromolecules, such as polysaccharides (e.g., chitin or chitin–glucan complexes), proteins, and enzymes, as well as their use as substrates in fermentation or anaerobic digestion processes to produce biochemicals, organic acids and biofuels.
Although biological activity assays were not performed, the secondary metabolite composition was assessed to provide an initial overview of the chemical diversity of the studied strains and to compare their metabolomic profiles. To this end, metabolomic data were presented as enriched predicted chemical classes, in accordance with the primary objective of achieving a broad characterization of the samples rather than a detailed annotation of individual metabolites.
The metabolomic analysis revealed the presence of distinct chemical classes among the analyzed strains, with A. iizukae showing the greatest chemical diversity. In particular, this species was characterized by a higher abundance of alkaloids and the presence of classes commonly associated with bioactive compounds, including polyketides, shikimate-derived metabolites, and phenylpropanoids. These findings are consistent with previous reports describing the production of biotechnologically relevant metabolites by A. iizukae, such as diphenyl derivatives (iizukines), flavonolignans (silybines and isosilybines), and oxidative enzymes with industrial applications, including laccase, manganese peroxidase, and lignin peroxidase [72,73]. In contrast, neither Stemphylium sp. nor Mucor sp. exhibited a broad diversity of the predicted secondary metabolites. Instead, these strains were characterized by a relatively higher abundance of primary metabolism-related compounds, including fatty acid derivatives (fatty acyls, fatty amides, glycerolipids, octadecanoids, and sphingolipids) and carbohydrate-associated metabolites such as amino sugars, aminoglycosides, and nucleosides.
Overall, these results highlight species-specific differences in both biomass composition and metabolomic profiles, providing a foundation for future studies aimed at evaluating the biotechnological potential of these marine-derived fungi and identifying specific compounds of industrial or pharmaceutical interest.

5. Conclusions

This study provides the first characterization of the biochemical composition and metabolomic profile of Aspergillus iizukae, Stemphylium sp., and Mucor sp. isolated from the Bahía Blanca Estuary. The occurrence of these fungi in environments with high-salinity, elevated temperatures, and high organic matter content highlights their ability to thrive under stressful and anthropogenically influenced conditions. The results also emphasize the need for further taxonomic, ecological, and physiological studies, particularly for Stemphylium sp. and Mucor sp., whose ecology in marine environments remains poorly understood. Since all strains were cultivated under the same synthetic media, which likely influenced biomass production and biochemical composition in different ways, future research should evaluate the effects of culture parameters on metabolite production and biochemical biomass composition for each strain. Metabolomic analyses revealed marked differences among the studied fungi. Aspergillus iizukae exhibited the greatest chemical diversity, highlighting the need for targeted studies focused on the identification and characterization of its individual metabolites. In contrast, Mucor sp. stood out for its high glucosamine content, suggesting potential applicability in processes involving chitin- and chitosan-rich biomass. Further characterization of the polysaccharide fractions of Stemphylium sp. and A. iizukae may also help clarify their potential value in biotechnological applications. Overall, the studied marine fungi represent underexplored biological resources with both ecological significance and biotechnological potential. The present work provides a baseline for future investigations aimed at understanding their ecological roles, optimizing their cultivation and exploring their potential as sources of valuable biomolecules and metabolites.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/jmse14121106/s1. Table S1. Taxonomic table of fungi found in seawater from Rosales Port in the Bahía Blanca Estuary. Figure S1. (a–c). Procedure for isolating and culturing fungi from seawater and Sarcocornia perennis. (a) Filtration of seawater, observation of S. perennis and culture colonies under stereomicroscopy. (b) Colonies of three isolates from point (a): Aspergillus iizukae (I), Stemplylium sp. (II) and Mucor sp. (III). (c) Two replicates for each fungus, transferred between one to four times to obtain pure colonies. Figure S2. (a–d)—Aspergillus iizukae BBB:BF5. (a) Conidial heads in MEA culture, view from above. (b) Conidial head, view of metulae (black arrow) and conidiogenous cells (white arrow). (c) Conidia. (d) Accessory conidia, sessile (white arrows) and short conidiophores (black arrows). (e,f)—Mucor sp. BBB:BF7. (e) Young sporangia. (f) Mature sporangia. (g) Columellae. (h) Sporangia borne on circinate branches. (i)—Sporangiospores. (j–n)—Stemphylium sp. BBB:BF3. (j) Conidiogenous cells and hyaline conidium. (k) Ornamented hypha. (l) Mature dictyoseptate conidium. (m) Intercalary conidium. (n) Chlamydospores chains. Scales bars: b, c, d, e, f, i, n = 10 µm; g, h: 15 µm; j, k, l, m: 5 µm [74,75,76,77,78].

Author Contributions

Conceptualization; methodology; software, validation; formal analysis; investigation; data curation; writing—original draft preparation; writing—review and editing; visualization; supervision; resources: F.B. Methodology; formal analysis; investigation; writing—original draft preparation; writing—review and editing, supervision: M.P. Methodology; formal analysis; investigation; writing—original draft preparation; writing—review and editing: P.C.P.B. and R.S. Validation; project administration: O.K.S. Conceptualization; validation; investigation; resources; writing—review and editing; visualization; supervision; project administration; funding: A.F. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported by DAAD, Deutscher Akademischer Austausch Dienst (German Academic Exchange Service), grant number: 91695828. This study was supported by PICT-2021-I-A-00779 I-A (FONCYT, project lead: Florencia Biancalana), PIP 11220210100830 (CONICET, project lead: Florencia Biancalana) and the project “Sustainable functional integration in composite materials” (NaFuVer) (project lead: Dreyer, TH Wildau; project coordination at the IGZ: Anna Fricke; project number: 86000999), funded by the European Regional Development Fund and the state of Brandenburg (EFRE).

Data Availability Statement

The original contributions presented in this study are included in the article/Supplementary Material. Further inquiries can be directed to the corresponding author.

Acknowledgments

The authors would like to thank the assistant Andrea Maikath for her work and support in laboratory work. I would particularly like to thank the DAAD, Deutscher Akademischer Austausch Dienst (German Academic Exchange Service) for trusting in my professional abilities and once again granting me the opportunity to work in the project titled Marine Biopolymers: Chitin and chitosan, a potential source from the associated fungal microbiome of wetland macrophytes, in the Leibniz Institute for Vegetable and Ornamental Crops (IGZ)-Germany (2024). Furthermore, the authors would like to thank the organizing staff of the INSECTS Plus international congress for the invitation to present this work.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Map of the Bahía Blanca Estuary (BBE) (a), showing Rosales Port (RP, upper right side) (b); salt marshes from Rosales Port (b–e); Sarcocornia perennis and Spartina alterniflora patch in the Rosales Port (c–e).
Figure 1. Map of the Bahía Blanca Estuary (BBE) (a), showing Rosales Port (RP, upper right side) (b); salt marshes from Rosales Port (b–e); Sarcocornia perennis and Spartina alterniflora patch in the Rosales Port (c–e).
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Figure 2. Maximum likelihood phylogenetic tree of Aspergillus species section Flavipedes, based on the combined data set of ITS and BenA. Bootstrap values >70 are presented at the nodes. The scale bar represents the number of nucleotide substitutions per site.
Figure 2. Maximum likelihood phylogenetic tree of Aspergillus species section Flavipedes, based on the combined data set of ITS and BenA. Bootstrap values >70 are presented at the nodes. The scale bar represents the number of nucleotide substitutions per site.
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Figure 3. Total dry mass (g) corresponded to Aspergillus iizukae, Stemphylium sp. and Mucor sp.
Figure 3. Total dry mass (g) corresponded to Aspergillus iizukae, Stemphylium sp. and Mucor sp.
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Figure 4. The biochemical characterization: (a) Composition of macromolecules and (b) sugar monomers of fungal strains. Results are expressed as g/100 g in dry mass (d.m.). The error bars correspond to the SE (standard Error) of means.
Figure 4. The biochemical characterization: (a) Composition of macromolecules and (b) sugar monomers of fungal strains. Results are expressed as g/100 g in dry mass (d.m.). The error bars correspond to the SE (standard Error) of means.
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Figure 5. Venn diagrams showing differences in metabolic composition detected in ethyl acetate extract from five isolated fungi (with replication of A. iizukae and Mucor sp.). (a) Refers to the data acquired in the positive mode; (b) refers to the data acquired in the negative mode.
Figure 5. Venn diagrams showing differences in metabolic composition detected in ethyl acetate extract from five isolated fungi (with replication of A. iizukae and Mucor sp.). (a) Refers to the data acquired in the positive mode; (b) refers to the data acquired in the negative mode.
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Figure 6. Sunburst plots of compound classes predicted by SIRIUS based on MS2 spectral data. On the left, data acquired in positive ionization mode (ESI+). On the right, data acquired in negative ioniation mode (ESI−). (a) A. iizukae; (b) Stemphylium sp.; (c) Mucor sp.
Figure 6. Sunburst plots of compound classes predicted by SIRIUS based on MS2 spectral data. On the left, data acquired in positive ionization mode (ESI+). On the right, data acquired in negative ioniation mode (ESI−). (a) A. iizukae; (b) Stemphylium sp.; (c) Mucor sp.
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MDPI and ACS Style

Biancalana, F.; Psarianos, M.; Bueno, P.C.P.; Sanchez, R.; Schlüter, O.K.; Fricke, A. First Insights into the Biochemical and Metabolomic Characterization of Marine Fungi Isolated from Salt Marshes of an Argentine Estuary. J. Mar. Sci. Eng. 2026, 14, 1106. https://doi.org/10.3390/jmse14121106

AMA Style

Biancalana F, Psarianos M, Bueno PCP, Sanchez R, Schlüter OK, Fricke A. First Insights into the Biochemical and Metabolomic Characterization of Marine Fungi Isolated from Salt Marshes of an Argentine Estuary. Journal of Marine Science and Engineering. 2026; 14(12):1106. https://doi.org/10.3390/jmse14121106

Chicago/Turabian Style

Biancalana, Florencia, Marios Psarianos, Paula C. P. Bueno, Romina Sanchez, Oliver K. Schlüter, and Anna Fricke. 2026. "First Insights into the Biochemical and Metabolomic Characterization of Marine Fungi Isolated from Salt Marshes of an Argentine Estuary" Journal of Marine Science and Engineering 14, no. 12: 1106. https://doi.org/10.3390/jmse14121106

APA Style

Biancalana, F., Psarianos, M., Bueno, P. C. P., Sanchez, R., Schlüter, O. K., & Fricke, A. (2026). First Insights into the Biochemical and Metabolomic Characterization of Marine Fungi Isolated from Salt Marshes of an Argentine Estuary. Journal of Marine Science and Engineering, 14(12), 1106. https://doi.org/10.3390/jmse14121106

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