Abstract
Introduction:
Marine macroalgae are increasingly recognized as promising sources of bioactive compounds for sustainable plant disease management. This study evaluated the anti-phytopathogenic potential of organic extracts from three intertidal macroalgal species, Ulva fasciata, Gracilaria khanjanapajiae, and Sargassum wightii, collected from Thalpe Reef, Sri Lanka.
Methods:
Chloroform, ethyl acetate, and methanol extracts were screened against phytopathogenic fungi Aspergillus niger, Colletotrichum lindemuthianum, Colletotrichum fructicola, Sclerotinia sclerotiorum, and Fusarium oxysporum using the poisoned plate method, and against phytopathogenic bacteria Xanthomonas cucurbitae, Pectobacterium carotovorum, Pseudomonas syringae, Dickeya chrysanthemi, and Ralstonia sp. using the well diffusion assay. Bioassay-guided fractionation was performed on selected active extracts, and the resulting fractions were analyzed by gas chromatography–mass spectrometry.
Results:
Antimicrobial activity varied significantly among macroalgal species, extraction solvents, concentrations, and target pathogens (ANOVA, p < 0.05). The ethyl acetate extract of G. khanjanapajiae exhibited the highest antifungal activity with 71.52 ± 1.60% inhibition against C. lindemuthianum. In contrast, the chloroform extract of U. fasciata showed the strongest antibacterial activity, producing a 24.2 ± 0.3 mm inhibition zone against D. chrysanthemi. Bioassay-guided fractionation further enhanced antimicrobial efficacy, with selected fractions demonstrating higher inhibition than crude extracts. GC–MS analysis of the most active fractions revealed diverse metabolites, including lipophilic phenolics, fatty acids, fatty amides, terpenes and terpenoid derivatives, long-chain alcohols, aldehydes, ketones, and long-chain hydrocarbons such as alkanes, alkenes, and α-olefins, many of which are known for antimicrobial properties.
Discussion:
The findings necessitate further investigation of Sri Lankan marine macroalgae species and solvent-dependent variation of their antimicrobial potential, as candidates for environmentally compatible plant disease control strategies.
1 Introduction
Plant diseases remain one of the major obstacles to global agricultural productivity, resulting in annual yield losses exceeding 30% worldwide (Savary et al., 2019). Beyond quantitative yield reductions, plant diseases also decline crop quality and market value, while increasing production costs through higher agrochemical inputs and labor requirements (Vurro et al., 2010). A wide range of biotic agents, including fungi, bacteria, nematodes, and viruses, are responsible for plant diseases. Especially, fungal and bacterial pathogens significantly impact a wide range of crops, particularly in the tropics, where warm and humid conditions favor pathogen proliferation.
A comprehensive checklist compiled by Adikaram and Yakandawala (2020) documented 404 species of plant pathogenic fungi and oomycetes, representing 110 genera, associated with horticultural, agricultural, plantation, forest plants, and harvested produce in Sri Lanka, signifying the threat posed by plant diseases to food security and agricultural sustainability. Notably, vegetable crops were reported to harbor the highest number of pathogenic fungal species. In addition to fungal pathogens, bacterial diseases also significantly contribute to crop losses in Sri Lanka. Therefore, in the present study, fungal pathogens (Fusarium oxysporum, Colletotrichum lindemuthianum, Aspergillus niger, Colletotrichum fructicola, and Sclerotinia sclerotiorum) and plant-pathogenic bacteria (Xanthomonas cucurbitae, Pectobacterium carotovorum, Pseudomonas syringae, Dickeya chrysanthemi, Ralstonia sp.) were selected for anti-phytopathogenic screening. These pathogens are frequently associated with major vegetable crops widely cultivated and consumed in Sri Lanka and cause a broad spectrum of plant diseases, including wilt, anthracnose, rots, mold, and bacterial blight. Collectively, these diseases cause substantial yield losses, reduce post-harvest quality, and have significant impacts on the national agricultural sector.
Many disease management strategies are utilized to mitigate the impact of such phytopathogens, including the use of resistant cultivars, improved agronomic practices such as certified seeds, balanced fertilizers, and biological and chemical control methods (Kalam et al., 2020). Among these, chemical control remains widely adopted due to its rapid and predictable outcomes. However, intensive and prolonged use of synthetic pesticides raises serious environmental and agronomic concerns, such as soil and water contamination, non-target organism toxicity, biodiversity loss, and the emergence of pesticide-resistant pathogens (Deising et al., 2008). These limitations, along with increasing consumer demand for organic produce, have shifted the focus to eco-friendly and biodegradable alternatives for crop disease management (Arora et al., 2021). Natural products are rich sources of diverse bioactive secondary metabolites with a wide range of biological activities. Compared to synthetic agrochemicals, they offer greater structural novelty and environmental compatibility (Gómez-Hernández et al., 2021). Among these, marine macroalgae have attracted growing interest as potential sources of bioactive compounds for plant disease control, with several studies reporting their antifungal and antibacterial activities against various phytopathogens (Esserti et al., 2017).
Marine macroalgae are multicellular, photosynthetic eukaryotes classified into three major groups: red algae (Rhodophyta), green algae (Chlorophyta), and brown algae (Phaeophyta) (Pereira, 2021). They produce a wide range of bioactive metabolites, including alkaloids, flavonoids, polysaccharides, terpenes, phlorotannins, sterols, quinones, polyphenols, tocopherols, carotenoids, and chlorophylls (Biris-Dorhoi et al., 2020). Such compounds have been reported to exhibit antimicrobial, antiviral, antioxidant, anticancer, and anti-inflammatory properties (Mickymaray and Alturaiki, 2018; Avila-Romero et al., 2023). Importantly, the chemical composition and bioactivity of macroalgal metabolites are strongly influenced by species, physiological status, environmental conditions (temperature, salinity, nutrient availability), seasonal variation, and epiphytic associations (Trigui et al., 2013).
Despite Sri Lanka’s rich marine biodiversity, systematic bioprospecting of marine macroalgae for anti-phytopathogenic applications remains limited. In particular, the macroalgae of the Thalpe reef, a biologically diverse coastal ecosystem along the southern coast of Sri Lanka, have received little scientific attention regarding their chemical composition and anti-phytopathogenic activity. Previous studies on macroalgae from the Thalpe reef are limited. Rodrigo et al. (2025) reported antifungal activity of Ulva, Sargassum, and Gracilaria extracts against phytopathogenic fungi (Lasiodiplodia theobromae, Pseudopestalotiopsis theae, and Diaporthe eugeniae) affecting Solanum melongena. However, their work was limited to pathogens associated with a single crop and lacked molecular identification of macroalgal samples. Similarly, Alahakoon et al. (2025) investigated antifungal compounds in methanolic extracts of U. fasciata and evaluated their inhibitory effects on the pectinase enzyme of P. theae using molecular docking approaches. Overall, existing studies primarily focus on antifungal screening and in silico analyses, and there are currently no studies that have addressed the antibacterial activity of molecularly identified macroalgae from the Thalpe reef.
The novelty of this study lies in its integrated evaluation of the antifungal and antibacterial activities of molecularly identified macroalgae, supported by chemical profiling and quantitative determination of minimum inhibitory concentrations (MICs). To the best of our knowledge, this study represents the first detailed investigation of the anti-phytopathogenic activity and chemical profiling of bioactive fractions derived from underexplored marine macroalgal species collected from the Thalpe reef, Sri Lanka. We hypothesize that marine macroalgae inhabiting the Thalpe reef produce chemically diverse secondary metabolites with significant antifungal and antibacterial activities against plant pathogenic fungi and bacteria, respectively, as a result of adaptation to a dynamic and competitive marine environment.
Accordingly, the primary objective of this study was to evaluate the anti-phytopathogenic potential of selected macroalgal extracts through biological screening, bioassay-guided fractionation, and preliminary chemical characterization of active fractions. The conclusions of the study are based primarily on experimentally validated antimicrobial activity, while GC-MS analysis provides supporting evidence for the preliminary characterization of candidate bioactive metabolites in the active fractions. Definitive structural elucidation and confirmation of active compounds would require further targeted analyses using advanced techniques such as HRMS, LC-MS/MS, NMR spectroscopy, and purification of individual compounds, which are beyond the scope of the present investigation. Collectively, these findings contribute to the discovery of novel, environmentally sustainable agents for crop disease management.
2 Materials and methods
2.1 Sample collection, identification, and preparation
Three marine macroalgal species representing the three major algal phyla, U. fasciata (Chlorophyta), G. khanjapanajiae (Rhodophyta), and S. wightii (Phaeophyta), were collected from Thalpe reef, Galle, Sri Lanka (5° 59’ 44.54” N, 80° 17’ 10.34 “E) in July 2023 (Figure 1). Species identification was confirmed using standard references (Durairatnam, 1961; Coppejans et al., 2009) and molecular analysis targeting the tufA gene (plastid elongation factor Tu), cox1 gene (mitochondrial-encoded cytochrome oxidase subunit 1), and rbcL gene (ribulose-bisphosphate carboxylase). The generated sequences were deposited in GenBank under accession numbers PX512880 for U. fasciata and PX512889 for G. khanjanapajiae. Voucher specimens were deposited in the departmental herbarium of the University of Kelaniya for future reference. Immediately after collection, samples were rinsed thoroughly with seawater to remove sand and epibionts, followed by washing with distilled water to remove surface salts. The samples were shade dried at 28 ± 2 °C under ventilated conditions for 7–10 days until constant weight was achieved. Dried samples were ground into a fine powder using an electric grinder, transferred into sterilized, airtight glass containers, and stored at -4 °C in darkness until extraction.
Figure 1
2.2 Extraction of algal bioactive compounds
Finely powdered algal material (45 g) was extracted separately with 450 mL of chloroform, ethyl acetate, or methanol using a Soxhlet extractor (Electrothermal, Canada) for 6 h. Following the extraction, solvents were removed under reduced pressure using a rotary evaporator (BIOBASE, RE-201D, China). Evaporation was performed with the water-bath temperature maintained at 35–40 °C for chloroform and 40–45 °C for ethyl acetate and methanol, under reduced pressure (~100–250 mbar). Crude extracts were stored in sterilized glass vials at -20 °C until further analysis (Choudhury et al., 2005). The percentage yield (% yield) of crude extracts was calculated using the equation described by Agbaje-Daniels et al. (2020) and Zhang et al. (2024).
2.3 Microbial cultures
Antifungal activity was evaluated against five phytopathogenic fungi: F. oxysporum (PX517301) causing Solanum melongena leaf yellowing, C. lindemuthianum and C. fructicola (PX613666) causing anthracnose on Phaseolus vulgaris and Capsicum annum, respectively, A. niger (PX517287) causing black mold on Allium cepa, and S. sclerotiorum causing white mold on Brassica oleracea. Fungal cultures were maintained on potato dextrose agar (PDA). S. sclerotiorum was incubated at 23 °C, while others were incubated at 28 ± 2 °C.
Antibacterial assays were performed against five plant-pathogenic bacterial species: X. cucurbitae, causing black rot lesions on Brassica oleracea, P. carotovorum and D. chrysanthemi causing soft rot of Solanum tuberosum and B. oleracea L. var. gongylodes respectively, P. syringae causing bacterial speck symptoms of Solanum lycopersicum, and Ralstonia sp. causing S. lycopersicum bacterial wilt. Bacterial strains were cultured on nutrient agar (NA) at 28 ± 2 °C, and stock cultures were maintained at 4 °C with periodic sub-culturing.
2.4 Antifungal activity assay
Antifungal activity of macroalgal extracts was assessed using the poisoned food technique as described by Abhishek et al. (2021). Four concentrations, i.e., 0.25, 0.5, 1, and 2 mg mL-1, were prepared by dissolving the required amount of crude extracts in 500 µL of dimethyl sulfoxide (DMSO). Appropriate volumes of dissolved crude extracts (500 µL) were mixed with 15 mL of molten PDA (~ 40 °C), poured into petri plates, and allowed to solidify.
Mycelial disks of 6 mm diameter from the peripheral region of actively growing four days old cultures were placed at the center of each plate. Plates containing DMSO alone served as negative controls, while captan (0.5 mg mL-1) served as the positive control. Colony diameter (in mm) was measured after five days, and percentage inhibition was calculated using the following formula (Pourakbar et al., 2021).
Where I% is the percentage inhibition, dc is the mean diameter of the fungal colony in the control, and dt is the mean diameter of the fungal colony in treatment.
2.5 Antibacterial activity assay
Antibacterial activity was determined using the agar well diffusion method as described by Palaniyappan et al. (2023), with slight modifications. Twenty-four hour old bacterial broth cultures were adjusted to approximately 1 × 108 CFU mL-1 (OD600 ≈ 0.4) and uniformly spread on Muller-Hinton agar plates. Wells (~ 6 mm diameter) were cut using a sterilized cork borer and filled with 100 µL of algal extracts at concentrations of 0.25, 0.5, 1, and 2 mg mL-1. DMSO and streptomycin (0.5 mg mL-1) were used as the negative and positive controls, respectively. Zones of inhibition were measured after 24–48 h of incubation at 28 ± 2 0C.
2.6 Determination of minimum inhibitory concentration
MIC50 values were determined by broth microdilution method in 96-well plates. Nutrient broth (NB) and potato dextrose broth (PDB) were used for bacteria and fungi, respectively. Bacterial cultures were standardized to OD600 of approximately 0.4, and fungal conidial suspensions were adjusted to 1 × 105 conidia mL-¹. Extract concentrations ranged from 0.25 to 2 mg mL-1. Liquid culture medium containing the algal extract without microbial inoculation served as the negative control, while liquid culture medium inoculated with the microbial culture without algal extract served as the positive control. Absorbance was measured at 600 nm (bacteria) and 630 nm (fungi) using a UV-visible spectrophotometer (Thermo Scientific™ Multiskan™ GO Microplate Spectrophotometer). Percentage inhibition values at different extract concentrations were transformed into probit units using Finney’s table (Finney, 1952) and regressed against the logarithm of concentrations. The concentration corresponding to probit 5 (50% inhibition) was determined from the regression equation and reported as the MIC50, and all assays were conducted in triplicate (Čmiková et al., 2022).
2.7 Fractionation of algal extracts by column chromatography
Ethyl acetate extract of G. khanjapanajiae and the chloroform extract of U. fasciata, which showed the highest antifungal and antibacterial activity, respectively, were fractionated by column chromatography. The concentrated crude extracts were separately adsorbed onto silica gel (25 g, 100–200 mesh size) and subjected to elution using a gradient solvent system of increasing polarity: hexane< ethyl acetate< acetone< methanol. Elution was performed at a flow rate of 1 mL min-1,and a total of 61 fractions were collected separately for each extract into clean glass test tubes. All fractions were analyzed by Thin Layer Chromatography (TLC). TLC plates were visualized under UV light (254 nm and 356 nm). The retention factor (Rf) value was calculated using the equation Rf = d/D, where ‘d’ was the distance traveled by the spot, and ‘D’ was the distance traveled by the solvent front. Subsequent fractions exhibiting similar Rf values were pooled and stored at 4 °C in darkness until bioassay evaluation. Based on the TLC profiles, the 61 fractions obtained from the U. fasciata chloroform extract and G. khanjanapajiae ethyl acetate extract were combined into eight and five pooled fractions, respectively.
2.8 Bioassay of fractions
Fractions from U. fasciata chloroform extract were tested for antibacterial activity using the agar well diffusion method as previously described, while fractions from the G. khanjanapajiae ethyl acetate extract were evaluated for antifungal activity using the broth microdilution method following the Clinical and Laboratory Standard Institute guidelines (CLSI, 2008). For the broth microdilution assay, C. lindemuthianum conidial suspensions were adjusted to 1 × 105 conidia mL-1. Each fraction was prepared as a 10 mg mL-1 stock solution in DMSO, and diluted to final concentrations of 0.25, 0.5, 1, and 2 mg mL-1 using PDB. DMSO and captan (0.5 mg mL-1) were employed as the negative and positive controls, respectively. In 96-well microtiter plates, 50 µL of diluted fraction, 50 µL of fungal inoculum, and 100 µL of PDB were added per well and incubated at 28 ± 2 °C for 72 h.
Fungal growth was quantified by measuring optical density at 630 nm using a UV-visible spectrophotometer, and percentage inhibition was calculated relative to the negative control using the following equation (Ing et al., 2012).
Where I% is the percentage inhibition, Ac is the optical density (OD) of the fungal mycelial growth in the control treatment, and At is the optical density (OD) of the fungal mycelial growth in the extract-containing treatment.
2.9 Gas chromatography-mass spectrometry analysis
The most active fractions were analyzed by GC-MS using an Agilent 7890B GC system coupled with a 5977A MSD mass spectrophotometer at the Sri Lanka Institute of Nanotechnology (SLINTEC). Separation was achieved on an HP-5MS capillary column (5%-phenyl)-methylpolysiloxane with a length of 30 m, an internal diameter of 0.25 mm, and a thickness of 0.25 µm. The injector and detector temperatures were set at 280 °C and 300 °C, respectively. The oven temperature was initially maintained at 80 °C for 1 min and then increased to 300 °C at a rate of 20 °C min-1. Helium was used as a carrier gas at a constant flow rate of 1.0 mL min-1. Samples (2 µL) were injected in splitless mode with a purge time of 1 min. The mass spectrometer operated in electron ionization mode at 70 eV, scanning a mass range of 50 to 650 m/z. Compound identification was carried out by comparing mass spectra with the NIST 14 and W9N11 libraries. Data acquisition and processing were performed using ChemStation software. The relative percentage of detected compounds was expressed as a percentage based on peak height (Shobier et al., 2016; Pourakbar et al., 2021).
2.10 In silico PASS prediction
Predicted antibacterial and antifungal activities of identified compounds were assessed using the PASS online server (Way2Drug), with probability of activity (Pa) values used to estimate biological relevance.
2.11 Statistical analysis
All experiments were conducted in triplicate. Data normality was tested using the Shapiro-Wilk test. Percentage inhibition data were log-transformed and analyzed using three-way ANOVA followed by Tukey’s post hoc test (p< 0.05) in R software (Version 4.4.2).
3 Results
3.1 Extraction yields of macroalgal crude extracts
Extraction yields differed significantly among macroalgal species and solvents (p< 0.05). Methanol extraction of U. fasciata produced the highest yield (17.8 ± 0.72%), followed by G. khanjanapajiae (12.4 ± 0.43%). In contrast, S. wightii yielded its highest extract yield with ethyl acetate (9.47 ± 0.55%), whereas chloroform consistently produced the lowest yields across species. These results indicate that extraction efficiency depends on both macroalgal species and solvent type (Figure 2).
Figure 2
3.2 Antifungal activity of crude macroalgal extracts
Crude extracts of the three macroalgal species exhibited statistically significant but variable antifungal activity against the tested phytopathogens, depending on algal species, extraction solvent, extract concentration, and target fungus (three-way ANOVA, p< 0.05). No measurable mycelial growth was observed in plates treated with the positive control, Captan (0.5 mg mL-¹), indicating 100% growth inhibition across all tested fungi. In contrast, the negative control (DMSO) exhibited no detectable antifungal activity. Significant main effects were observed for algal species (F(2,504) = 14.37, p< 0.001), solvent type (F(2,504) = 3.27, p = 0.0389), and extract concentration (F(3,504) = 21.70, p< 0.001), along with a significant species x solvent interaction (F(4,504) = 13.58, p< 0.001). Overall, extracts of G. khanjanapajiae and U. fasciata showed higher antifungal efficacy than those of S. wightii, and ethyl acetate extracts generally produced stronger inhibition than chloroform and methanol extracts. Furthermore, the efficacy of the extracts varied depending on the target pathogen, with a concentration of 2 mg mL-1 demonstrating the highest antifungal activity (Figure 3).
Figure 3
Methanol extracts of U. fasciata exhibited the strongest activity, showing 60.42 ± 3.33% inhibition against S. sclerotiorum, corresponding to approximately 60% of the inhibition produced by Captan. Conversely, the lowest efficacy was observed for methanol extracts of G. khanjanapajiae against A. niger (1.12 ± 0.37%). Ethyl acetate extracts exhibited varying levels of inhibition across the three algal species. The extract from G. khanjanapajiae showed the highest activity against S. sclerotiorum (71.92 ± 1.60%), whereas S. wightii showed negligible activity against F. oxysporum (0.24 ± 0.00%). Chloroform extracts further underscored these solvent-dependent differences. The extract of G. khanjanapajiae exhibited 75.00 ± 0.00% inhibition against S. sclerotiorum, representing the highest activity relative to the positive control, whereas S. wightii showed the lowest activity against F. oxysporum (0.95 ± 0.41%).
Across all 180 treatment combinations (3 algae × 3 solvents × 4 concentrations × 5 fungi), only 16 exhibited inhibition levels exceeding 50%, relative to the standard antifungal agent, Captan. Among these, the majority were effective against S. sclerotiorum, with inhibition values ranging from 55.83% to 75.00%. The strongest inhibition of S. sclerotiorum was observed with the chloroform extract of G. khanjanapajiae, reaching 75.00 ± 0.00% inhibition at 1 mg mL-¹ (Figure 3). Comparable inhibition was recorded for the chloroform extract of S. wightii (72.92 ± 2.08%) and the ethyl acetate extract of G. khanjanapajiae (70.00 ± 0.00%) at the same concentration. In contrast, the ethyl acetate extract of U. fasciata showed minimal activity against S. sclerotiorum (9.17 ± 2.08%).
Ethyl acetate extracts of G. khanjanapajiae at 2 mg mL-1 exhibited the highest antifungal activity against C. lindemuthianum and C. fructicola with inhibition values of 71.52 ± 1.60% and 56.61 ± 0.34%, respectively, corresponding to approximately 72% and 57% of the inhibition produced by the positive control. The respective MIC50 values were 0.78 mg mL-1 for C. lindemuthianum and 1.43 mg mL-1 for C. fructicola (Figure 4). Chloroform extracts of G. khanjanapajiae also showed strong antifungal activity against S. sclerotiorum, with an MIC50 value of 0.89 mg mL-¹. None of the tested algal extracts achieved ≥50% inhibition against A. niger or F. oxysporum. The highest inhibition for these fungi was observed at 2 mg mL-¹, with ethyl acetate and methanol extracts of U. fasciata inhibiting A. niger and F. oxysporum by 47.39 ± 1.71% and 49.40 ± 1.48%, respectively (Figure 4).
Figure 4
3.3 Antifungal activity of isolated ethyl acetate fractions of G. khanjanapajiae
The ethyl acetate extract of G. khanjanapajiae at 2 mg mL-1 showed the highest antifungal activity in preliminary screening and was therefore selected for bioassay-guided fractionation. Five fractions obtained through column chromatography were subsequently evaluated against C. lindemuthianum, which showed greater susceptibility compared to the other tested fungal species. Overall, antifungal activity varied significantly among fractions and concentrations (two-way ANOVA; fraction: F(4,40) = 24.25, p<0.05; concentration F(3,40) = 88.11, p<0.05), with a significant fraction x concentration interaction (F(12,40) = 11.85, p<0.05). Among the fractions, fraction 3 exhibited the strongest inhibition of C. lindemuthianum, achieving 76.06 ± 2.85% inhibition at 2 mg mL-1, which was significantly higher than that of the other fractions (p< 0.001; Figure 5). In contrast, fraction 1 showed the lowest antifungal activity, with 43.05 ± 2.39% inhibition at 0.5 mg mL-1. The antifungal activity of the crude ethyl acetate extract of G. khanjanapajiae (71.52 ± 1.60% inhibition at 2 mg mL-¹) was lower than that of fraction 3 at the same concentration (76.06 ± 2.85%) (Figure 5), indicating enhanced antifungal efficacy following fractionation.
Figure 5
3.4 Antibacterial activity of crude macroalgal extracts
The antibacterial activity of chloroform, ethyl acetate, and methanol extracts of U. fasciata, G. khanjanapajiae, and S. wightii was tested at concentrations of 0.25, 0.5, 1, and 2 mg mL-1 against D. chrysanthemi, X. cucurbitae, P. syringae, P. carotovorum, and Ralstonia sp. Distinct solvent and species-dependent activity patterns were observed (Figure 6). The antibacterial activity of the extracts was quantitatively compared with the standard antibiotic streptomycin (0.5 mg mL-¹), which exhibited inhibition zones ranging from 19.2 ± 0.2 to 28.1 ± 0.3 mm across the tested bacterial strains. Statistical analysis revealed significant effects of algal species (F(2, 504) = 6.88, p = 0.0011), solvent type (F(2, 504) = 5.29, p = 0.0053), and extract concentration (F(3, 504) = 7.77, p< 0.001), as well as a significant interaction between algal species and solvent (F(4, 504) = 3.73, p = 0.0053), indicating that antibacterial activity varied depending on both species and extraction solvent.
Figure 6
Overall, extracts from U. fasciata and G. khanjanapajiae exhibited significantly higher antibacterial activity than those of S. wightii. Ethyl acetate extracts demonstrated a clear concentration-dependent response, with the highest inhibition observed for G. khanjanapajiae at 2 mg mL-1 against D. chrysanthemi (25.5 ± 1.0 mm; MIC50 = 0.52 mg mL-1), followed by U. fasciata (21.7 ± 0.8 mm) and S. wightii (20.8 ± 0.7 mm) at the same concentration. In contrast, the positive control, streptomycin, produced a significantly larger inhibition zone (28.1 ± 0.3 mm) against D. chrysanthemi, while no inhibition was detected for the negative control (DMSO). Ethyl acetate extracts of all three species showed no detectable inhibition against P. syringae at any tested concentrations (Figure 6). Methanol extracts exhibited antibacterial activity against most tested pathogens. The methanol extract of U. fasciata at 2 mg mL-1 showed the highest inhibition against D. chrysanthemi (22.0 ± 0.5 mm) (Figure 6). However, methanol extracts of U. fasciata and G. khanjanapajiae did not inhibit P. syringae growth. Chloroform extracts exhibited variable antibacterial activity. The chloroform extract of U. fasciata at 2 mg mL-1 produced a high inhibition zone against D. chrysanthemi (24.2 ± 0.7 mm) and was the only extract exhibiting activity against P. syringae, with an inhibition zone of 21.5 ± 0.3 mm and an MIC50 value of 0.63 mg mL-1 (Figure 7). In contrast, chloroform extracts of S. wightii and G. khanjanapajiae showed no detectable inhibition against P. syringae at any concentration. Similarly, the chloroform extract of U. fasciata at 2 mg mL-1 produced the highest inhibition against P. carotovorum (12.33 ± 0.4 mm), whereas ethyl acetate extracts of S. wightii showed no activity. The remaining extracts exhibited comparable inhibition zones ranging from 10.0 to 11.5 mm, with MIC50 values exceeding 2 mg mL-1 (Figure 7).
Figure 7
All extracts inhibited the growth of Ralstonia sp., with inhibition zones ranging from 11.8 ± 0.4 to 16.5 ± 0.8 mm, except for the ethyl acetate extract of U. fasciata at 0.25 mg mL-1. The highest inhibition was observed for the ethyl acetate extract of U. fasciata at 2 mg mL-1 (16.5 ± 0.8 mm; MIC50 = 1.6 mg mL-1), compared with 19.2 ± 0.2 mm for streptomycin (0.5 mg mL-¹). Although the extracts exhibited lower activity than the standard antibiotic, they demonstrated moderate antibacterial efficacy, corresponding to approximately 85% of the activity of streptomycin against Ralstonia sp. (Figure 7). Growth of X. cucurbitae was inhibited by all extracts at all tested concentrations. The chloroform extract of G. khanjanapajiae at 2 mg mL-1 showed the highest activity, with an inhibition zone of 21.8 ± 1.2 mm and an MIC50 value of 0.64 mg mL-1 (Figure 7).
3.5 Antibacterial activity of isolated chloroform fractions of U. fasciata
Based on Tukey’s post-hoc analysis, the chloroform extract of U. fasciata at 2 mg mL-1 exhibited the highest antibacterial activity across the tested bacterial strains and was therefore selected for fractionation by column chromatography. Eight fractions were obtained and subsequently evaluated for antibacterial activity against D. chrysanthemi, which showed the highest susceptibility to the crude extracts.
Among the eight fractions, fraction 5 at 2 mg mL-1 exhibited the highest antibacterial activity, producing an inhibition zone of 25.2 ± 0.2 mm against D. chrysanthemi (Figure 8). In contrast, fraction 7 at 0.25 mg mL-1 showed the lowest activity, with an inhibition zone of 12.0 ± 0.0 mm. The chloroform crude extract of U. fasciata at 2 mg mL-¹ produced an inhibition zone of 24.1 ± 1.6 mm, which was lower than that observed for the most active isolated fraction, indicating enhanced antibacterial activity following fractionation.
Figure 8
3.6 Characterization of compounds in the most active antimicrobial fractions
The most active antifungal fraction derived from the ethyl acetate extract of G. khanjanapajiae (Fraction 3), and the most active antibacterial fraction derived from the chloroform extract of U. fasciata (Fraction 5) were subjected to GC–MS analysis to obtain a preliminary chemical profile. Compound annotation was performed by comparing the obtained mass spectra with reference spectra in the NIST 14 and Wiley (W9N11) libraries. Only compounds with library match values ≥ 80% were considered for tentative identification and included in the analysis. As the GC–MS instrument used in this study operated at unit mass resolution, accurate mass measurements and ppm-level mass error calculations were not available. Therefore, all detected compound were assigned as tentative annotations based on spectral library matching.
3.6.1 GC-MS analysis of the third fraction of the ethyl acetate extract of G. khanjanapajiae
The chromatogram of the third fraction of the ethyl acetate extract of G. khanjapajiae revealed multiple peaks corresponding to putatively annotated metabolites belonging to diverse chemical classes, including phenolics, terpenes, fatty acids, hydrocarbons, ketones, esters, alcohols, aldehydes, fatty amides, aromatic amines, and monocarboxylic acids. Among the major peaks, compounds tentatively identified as 2,4-di-tert-butylphenol and n-hexadecanoic acid were detected, with predicted antifungal activity probabilities (Pa) of 0.356 and 0.407, respectively. Several minor peaks corresponded to metabolites previously reported in marine macroalgae and were detected at lower relative abundances (0.25–0.66%), with predicted antifungal probabilities greater than 0.500 (Supplementary Table 1).
3.6.2 GC-MS analysis of the fifth fraction of the chloroform extract of U. fasciata
The chromatogram of the fifth fraction of the chloroform extract of U. fasciata revealed a complex mixture of putatively annotated metabolites, including fatty acids, aldehydes, hydrocarbons, phenolic compounds, terpenoids, esters, ketones, and alcohols. Major peaks were tentatively identified as n-hexadecanoic acid, phytol, and long-chain alkenes, each exhibiting a predicted antifungal activity probabilities (Pa) greater than 0.300 (Supplementary Table 2).
Information on the molecular formulae, molecular weights, theoretically calculated exact masses, chemical classifications, and previously reported biological activities of the compounds tentatively identified in the third fraction of the ethyl acetate extract of G. khanjapajiae and the fifth fraction of the chloroform extract of U. fasciata was compiled and is presented in Supplementary Tables 1 and 2 to support and facilitate the interpretation of the tentative compound annotations.
4 Discussion
Marine macroalgae are rich sources of structurally diverse and biologically active metabolites, shaped by their adaptation to complex and often extreme habitats (O’Keeffe et al., 2019a). Species inhabiting the intertidal zone experience daily fluctuations in temperature, salinity, light, and desiccation, which likely drive the biosynthesis of defence metabolites with antimicrobial properties (Freitas et al., 2020). In this study, species collected from the upper intertidal region, U. fasciata, G. khanjanapajiae, and S. wightii, encounter particularly harsh conditions during low tide, including intense solar radiation and osmotic stress, potentially enhancing their bioactive profiles.
Consistent with previous reports, Chlorophyta and Rhodophyta showed the highest antimicrobial activity, followed by Phaeophyta (Osman et al., 2013; Sheikh et al., 2018). Notably, higher activity of U. fasciata (Chlorophyta) and Gracilaria corticata (Rhodophyta) extracts aligns with earlier studies demonstrating the strong antimicrobial potential of these algae (Padmakumar and Ayyakkannu, 1997; Fareed and Khairy, 2008). Nevertheless, deviations exist for certain brown algae, which often exhibit strong antifungal activity in other studies (Ambika and Sujatha, 2014; Vicente et al., 2021), reflecting species-specific metabolite profiles and methodological variability. These discrepancies underscore the influence of environmental factors, extraction methods, and pathogen selection on observed bioactivity (Radhika et al., 2012; Mickymaray and Alturaiki, 2018).
Our data reveals pronounced solvent- and species-dependent activity. The ethyl acetate extract of G. khanjanapajiae displayed the strongest antifungal activity, whereas the chloroform extract of U. fasciata exhibited potent antibacterial effects. In contrast, S. wightii extracts exhibited lower efficacy. This aligns with the findings of Rodrigo et al. (2025), who found that the Ulva-ethyl acetate extract had the highest inhibition against D. eugeniae and P. theae, while Gracilaria-ethyl acetate extracts showed over 50% inhibition at the lowest concentrations (250 ppm), and Sargassum-methanol extracts had minimal activity (0.81%). These findings support prior observations that non-polar to moderately polar compounds are often responsible for antimicrobial effects (Toledo et al., 2023; Wasihun et al., 2023). Dose-dependent responses were evident in this study, aligning with previous studies on Gracilaria edulis and other macroalgae, where higher extract concentrations produced larger inhibition zones (Kolanjinathan et al., 2009; O’Keeffe et al., 2019b).
Pathogen specific susceptibility patterns were also notable in our study. S. sclerotiorum and C. lindemuthianum were highly sensitive to selected extracts, whereas F. oxysporum and A. niger showed lower inhibition, reflecting trends observed in earlier work (Sheikh et al., 2018). These resistance patterns highlight intrinsic microbial defenses that limit algal metabolite efficacy and underscore the need for targeted screening in antimicrobial discovery. The pronounced antibacterial activity of the chloroform extract of U. fasciata against P. syringae, in contrast to the lack of activity observed in S. wightii, likely reflects differences in their metabolite profiles, particularly the presence of compounds such as palmitic acid and sulfated polysaccharides, which are known for their antibacterial properties and plant defense inducing properties (Arunkumar et al., 2010; Vera et al., 2011).
Conversely, Padmakumar and Ayyakkannu (1997) and Mashjoor et al. (2016) reported that Pseudomonas aeruginosa was resistant to all tested extracts. This suggests that there are specific differences in susceptibility among species due to their robust outer membrane, which may limit penetration of algal metabolites (Michalak and Chojnacka, 2015). Notably, methanol extracts consistently inhibited X. cucurbitae, likely due to the solubility of polar compounds, such as phenolic acids or carotenoids, which disrupt bacterial membranes through oxidative stress (Christaki et al., 2013). The current findings showed that the most susceptible strain was X. cucurbitae, since the majority of macroalgal extracts reduced the growth of this pathogen to some extent. These results were in accordance with those of O’Keeffe et al. (2019b).
Although the antimicrobial activity observed in this study was detected at milligram-level concentrations, which is characteristic of crude natural product extracts. According to established criteria for marine natural products, MIC values for purified compounds are categorized into four groups as inactive (>32 µg mL-¹), weak (8–32 µg mL-¹), moderate (1–8 µg mL-¹), and potent (<1 µg mL-¹) (Holland and Carroll, 2023). However, these thresholds are not directly applicable to crude extracts. Furthermore, studies investigating crude seaweed extracts frequently report MIC values in the mg mL-¹ range. For instance, McGurrin et al. (2025) reported antimicrobial activity of Alaria esculenta extracts with MIC values of 6.25–12.5 mg mL-¹. The present study quantitatively evaluated the antimicrobial activity of crude macroalgal extracts using MIC50 values, and the observed activity levels are consistent with previous reports. Fractionation of the most potent crude extracts further enhanced antimicrobial efficacy. The third fraction of G. khanjanapajiae and the fifth fraction of U. fasciata showed increased inhibition compared to their corresponding crude extracts, supporting the evidence that fractionation concentrates bioactive compounds while removing inhibitory components (Plouguerné et al., 2006; Kumar et al., 2008). This approach addresses the limitation of crude extracts, which may mask potent metabolites through antagonistic interactions (Padmakumar and Ayyakkannu, 1997). Thus, the enhanced activity observed in these fractions suggests the presence of bioactive compounds that could be further isolated through bioassay-guided purification for improved efficacy.
GC-MS analysis revealed that the third fraction of G. khanjanapajiae and the fifth fraction of U. fasciata contain diverse bioactive compounds, including fatty acids, aldehydes, hydrocarbons, phenolics, terpenoids, esters, ketones, amines, amides, carboxylic acids, terpenes, and alcohols, contributing to a broad spectrum of antimicrobial activity. 2,4-di-tert-butylphenol, n-hexadecanoic acid, tetradecanoic acid, palmitoleic acid, neophytadiene, and tetradec-2-ene bioactive compounds exhibit overlapping presence in both fractions, highlighting their potential as broad-spectrum antimicrobial agents, likely acting synergistically to enhance efficacy. n-Hexadecanoic acid (palmitic acid) and 2,4-di-tert-butylphenol emerged as major constituents, supporting reports of their antibacterial and antifungal efficacy across multiple macroalgal species (Selim et al., 2015; Barot et al., 2016; Pourakbar et al., 2021; Alahakoon et al., 2025). Other compounds, such as phytol, 1-docosene, and E-15-heptadecenal, were consistent with prior studies demonstrating membrane-disruptive and growth-inhibitory effects against fungi and bacteria (Sivakumar et al., 2014; El-Din and El-Ahwany, 2016; Hassan and Shobier, 2018).
PASS analysis of GC–MS–identified metabolites in the active ethyl acetate fraction of G. khanjanapajiae revealed several compounds with predicted antifungal probabilities (Pa) exceeding their probabilities of inactivity (Pi), including minor constituents with Pa values greater than 0.50. Similarly, metabolites detected in the chloroform fraction of U. fasciata exhibited predicted antibacterial potential. Previous studies on freshwater macrophytes and marine macroalgae have shown that metabolites such as fatty acids and volatile compounds with high predicted antifungal probabilities often correspond to experimentally validated antimicrobial activities (Kurashov et al., 2016; Pourakbar et al., 2021; El-Sheekh et al., 2022).
In the present study, the detection of n-hexadecanoic acid, along with other fatty acids and phenolic derivatives in the most active fraction, is consistent with these reports. Such compounds have been frequently associated with inhibitory effects against plant pathogenic fungi. Similar findings have been reported in marine macroalgae, including Ulva, Gracilaria, and Sargassum, where bioactive extracts analyzed by GC–MS and evaluated using the PASS identified compounds with Pa > Pi, such as dihydroactinidiolide, 4-hydroxy-2-butanone, and 6,10,14-trimethylpentadecan-2-one, as potential contributors to antifungal activity (Rodrigo et al., 2025).
Although PASS predictions do not confirm biological activity, the observed agreement between predicted activities and experimental inhibition of phytopathogens in this study provides supportive evidence for the potential role of these metabolites. The antimicrobial activity is likely attributed to the combined or synergistic action of multiple compounds, particularly fatty acids and phenolic derivatives. Overall, integrating GC–MS profiling with PASS-based bioactivity prediction provides supportive evidence for identifying candidate bioactive compounds in macroalgal extracts (Amiranashvili et al., 2020; Kawsar et al., 2022).
Given the speculative nature of GC–MS-based annotation, the observed antimicrobial activity likely results from the combined or synergistic effects of multiple metabolites rather than a single dominant compound. Therefore, further purification and structural elucidation are required to determine the specific bioactive constituents responsible for the observed effects. This can be achieved using advanced analytical techniques such as Liquid Chromatography–High-Resolution Electrospray Ionization Tandem Mass Spectrometry (LC-HR-ESI-MS/MS). The latter provides high mass accuracy and enables confident compound identification through exact mass measurements with low mass error (ppm-level), in combination with Nuclear Magnetic Resonance (NMR) spectroscopy, including one- and two-dimensional approaches, for definitive structural characterization. Collectively, these findings emphasize the importance of ecological context, species selection, and extraction methodology in evaluating marine macroalgal antimicrobial potential, while highlighting fractionation and chemical profiling as key strategies for identifying potent bioactive compounds with potential applications in plant protection.
5 Conclusions
Marine macroalgae serve as a valuable natural source of structurally diverse secondary metabolites with demonstrated antimicrobial potential relevant to sustainable crop protection. This study shows that G. khanjanapajiae and U. fasciata demonstrate significantly higher antimicrobial activities than S. wightii, emphasizing clear species-specific differences between green, red, and brown macroalgae. Antimicrobial activity varied markedly with extraction solvent, concentration, and target pathogen, reflecting the specificity and complexity of macroalgal chemical defenses. Chloroform and ethyl acetate extracts consistently showed greater bioactivity than methanolic extracts, indicating that non-polar to moderately polar metabolites play a dominant antimicrobial role. Bioassay-guided fractionation further enhanced antimicrobial efficacy compared to crude extracts, supporting its utility for isolating active constituents.
GC-MS analysis of the most potent fractions revealed a large number of active metabolites, providing a chemical basis for the observed antimicrobial effects. However, the detection of a compound in an extract does not confirm its direct role in antimicrobial activity. Therefore, further studies involving bioactivity-guided fractionation, isolation, and testing of pure compounds are necessary to validate their bioactive potential.
Overall, these findings highlighted marine macroalgae as promising candidates for the development of environmentally compatible antimicrobial agents and support further investigation into their application in crop protection. However, it is important to note that these results are based on controlled in vitro experiments. Therefore, these findings need to be extended through in vivo evaluations under greenhouse and subsequently field conditions to confirm efficacy, stability, and safety in ecosystems. Such stepwise validation is essential to account for complex biological interactions and to ensure that laboratory-scale observations reliably translate into practical agricultural or environmental applications.
Statements
Data availability statement
DNA sequences of the macroalgal species (accession numbers: PX512889, PX512880) and fungal pathogens (accession numbers: PX517301, PX517287, PX613666) used in this study are available in GenBank. Data will be made available on request.
Author contributions
AA: Data curation, Investigation, Methodology, Formal analysis, Software, Visualization, Writing – original draft, Writing – review & editing. BR: Data curation, Methodology, Writing – review & editing. PE: Conceptualization, Formal analysis, Investigation, Methodology, Resources, Software, Supervision, Writing – review & editing. HH: Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Writing – review & editing. RW: Conceptualization, Methodology, Investigation, Formal analysis, Validation, Resources, Software, Visualization, Funding acquisition, Project administration, Supervision, Writing – review & editing.
Funding
The author(s) declared that financial support was received for this work and/or its publication. This research was supported by the National Research Council of Sri Lanka (NRC IDG 22-062) and the Research Council of University of Kelaniya (RC/2025/PPRP02).
Acknowledgments
We would like to express our gratitude to W.A.V.S. Jayathilaka, L.M.T.N. Kabral, C.J.M.S.T. Chandrasekara, and M.P.C.N. Pathirana for their invaluable support during the field collections.
Conflict of interest
The author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declared that generative AI was not used in the creation of this manuscript.
Any alternative text (alt text) provided alongside figures in this article has been generated by Frontiers with the support of artificial intelligence and reasonable efforts have been made to ensure accuracy, including review by the authors wherever possible. If you identify any issues, please contact us.
Publisher’s note
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.
Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmars.2026.1816866/full#supplementary-material
References
1
AbhishekD.SanjayS.JadejaB. A. (2021). Cytotoxicity, antioxidant and antimicrobial activity of marine microalgae (Iyengaria stellata and Padina boryana) from the Gujarat coast. J. Maharaja Sayajirao Univ. Baroda55, 130–140.
2
AdikaramN. K. B.YakandawalaD. M. D. (2020). A checklist of plant pathogenic fungi and Oomycota in Sri Lanka. Ceylon J. Sci.49, 93–123. doi: 10.4038/cjs.v49i1.7709
3
Agbaje-DanielsF.AdeleyeA.NwankwoD.AdeniyiB.SekuF.BeukesD. (2020). Antibacterial activities of selected green seaweeds from West African coast. EC Pharmacol. Toxicol.8, 84–92.
4
AlahakoonA. H. D.RodrigoB. D.BalasooriyaB. C. M.HerathH. M.WanigatungeR. (2025). Molecular docking of potential antifungal compounds from Ulva fasciata methanolic extract against Pseudopestalotiopsis theae. J. Sri Lanka Assoc. For. Advancement Sci.7, 3–15. doi: 10.5281/zenodo.17488742
5
AmbikaS.SujathaK. (2014). Comparative studies on brown, red and green alga seaweed extracts for their antifungal activity against Fusarium oxysporum f. sp. udum in Pigeon pea var. CO (Rg) 7 (Cajanus cajan (L.) Mills.). J. Biopesticides7, 167–176. doi: 10.57182/jbiopestic.7.2.167-176
6
AmiranashviliL.NadaraiaN.MerlaniM.KamoutsisC.PetrouA.GeronikakiA.et al. (2020). Antimicrobial activity of nitrogen-containing 5-α-androstane derivatives: in silico and experimental studies. Antibiotics9, 224. doi: 10.3390/antibiotics9050224
7
AroraH.SharmaA.SharmaS.HaronF. F.GafurA.SayyedR. Z.et al. (2021). Pythium damping-off and root rot of Capsicum annuum L.: impacts, diagnosis, and management. Microorganisms9, 823. doi: 10.3390/microorganisms9040823
8
ArunkumarK.SivakumarS. R.RengasamyR. (2010). Review on bioactive potential in seaweeds (marine macroalgae): a special emphasis on bioactivity of seaweeds against plant pathogens. Asian J. Plant Sci.9, 227–240. doi: 10.3923/ajps.2010.227.240
9
Avila-RomeroM.García-BoresA. M.Garduño-SolorzanoG.Avila-AcevedoJ. G.Serrano-ParralesR.Orozco-MartínezJ.et al. (2023). Antimicrobial activity of some macroalgae of the Veracruzano Reef System (SAV), Mexico. Saudi J. Biol. Sci.30, 103496. doi: 10.1016/j.sjbs.2022.103496
10
BarotM.KumarN. J. I.KumarR. N. (2016). Bioactive compounds and antifungal activity of three different seaweed species Ulva lactuca, Sargassum tenerrimum and Laurencia obtusa collected from Okha coast, Western India. J. Coast. Life. Med.4, 284–289. doi: 10.12980/jclm.4.2016J5-185
11
Biris-DorhoiE. S.MichiuD.PopC. R.RotarA. M.TofanaM.PopO. L.et al. (2020). Macroalgae—A sustainable source of chemical compounds with biological activities. Nutrients12, 3085. doi: 10.3390/nu12103085
12
ChoudhuryS.SreeA.MukherjeeS. C.PattnaikP.BapujiM. (2005). In vitro antibacterial activity of extracts of selected marine algae and mangroves against fish pathogens. Asian fisheries Sci.18, 285–294.
13
ChristakiE.BonosE.GiannenasI.Florou‐PaneriP. (2013). Functional properties of carotenoids originating from algae. J. Sci. Food Agric.93, 5–11. doi: 10.1002/jsfa.5902
14
CLSI (2008). Performance standards for antimicrobial susceptibility testing. 15th informational supplement, M100- S15, clinical and laboratory standards institute, wayne.
15
ČmikováN.GalovičováL.MiškejeM.BorotováP.KluzM.KačániováM. (2022). Determination of antioxidant, antimicrobial activity, heavy metals and elements content of seaweed extracts. Plants11, 1493. doi: 10.3390/plants11111493
16
CoppejansE.LeliaertF.DargentO.GunasekaraR.De ClerckO. (2009). Sri Lankan seaweeds: Methodologies and field guide to the dominant species Vol. 6 (Brussels, Belgium: Belgian Development Cooperation Brussels).
17
DeisingH. B.ReimannS.PascholatiS. F. (2008). Mechanisms and significance of fungicide resistance. Braz. J. Microbiol.39, 286–295. doi: 10.1590/S1517-83822008000200017
18
DurairatnamM. (1961). Contribution to the study of the marine algae of Ceylon. Fisheries Res. Station Ceylon Bull.10, 181.
19
El-DinS. M. M.El-AhwanyA. M. (2016). Bioactivity and phytochemical constituents of marine red seaweeds (Jania rubens, Corallina mediterranea and Pterocladia capillacea). J. Taibah Univ. Sci.10, 471–484. doi: 10.1016/j.jtusci.2015.06.004
20
El-SheekhM. M.GalalH. R.MousaA. S.FarghlA. (2022). Screening of antifungal activity of bioactive chemical constituents in some brown marine macroalgae from the Red Sea, Egypt. Egypt. J. Bot.62, 865–878. doi: 10.21608/ejbo.2022.141379.2008
21
EssertiS.SmailiA.RifaiL. A.KoussaT.MakroumK.BelfaizaM.et al. (2017). Protective effect of three brown seaweed extracts against fungal and bacterial diseases of tomato. J. Appl. Phycol.29, 1081–1093. doi: 10.1007/s10811-016-0996-z
22
FareedM. F.KhairyH. M. (2008). In vitro antimicrobial activities of seaweeds collected from abu qir bay alexandria, Egypt. World Appl. Sci. J.5, 389–396.
23
FinneyD. J. (1952). Probit analysis: a statistical treatment of the sigmoid response curve. (Cambridge: Cambridge University Press).
24
FreitasR.MartinsA.SilvaJ.AlvesC.PinteusS.AlvesJ.et al. (2020). Highlighting the biological potential of the brown seaweed Fucus spiralis for skin applications. Antioxidants9, 611. doi: 10.3390/antiox9070611
25
Gómez-HernándezM.Rodríguez-GarcíaC. M.Peraza-EcheverríaL.Peraza-SánchezS. R.Torres-TapiaL. W.Pérez-BritoD.et al. (2021). In vitro antifungal activity screening of beach-cast seaweeds collected in Yucatan, Mexico. J. Appl. Phycol.33, 1229–1237. doi: 10.1007/s10811-021-02384-5
26
Hassan SaharW. M.Shobier AidaH. (2018). GC/MS identification and applications of bioactive seaweed extracts from Mediterranean coast of Egypt. Egypt. J. Aquat. Biol. Fish.22, 1–21. doi: 10.21608/ejabf.2018.17952
27
HollandD. C.CarrollA. R. (2023). Marine indole alkaloid diversity and bioactivity. What do we know and what are we missing? Nat. Prod. Rep.40, 1595–1607. doi: 10.1039/d2np00085g
28
IngL. Y.ZinN. M.SarwarA.KatasH. (2012). Antifungal activity of chitosan nanoparticles and correlation with their physical properties. Int. J. Biomater.2012, 632698. doi: 10.1155/2012/632698
29
KalamS.BasuA.AhmadI.SayyedR. Z.El-EnshasyH. A.DailinD. J.et al. (2020). Recent understanding of soil acidobacteria and their ecological significance: a critical review. Front. Microbiol.11, 580024. doi: 10.3389/fmicb.2020.580024
30
KawsarS. M.KumerA.MuniaN. S.HosenM. A.ChakmaU.AkashS. (2022). Chemical descriptors, PASS, molecular docking, molecular dynamics and ADMET predictions of glucopyranoside derivatives as inhibitors to bacteria and fungi growth. Organic Commun.15, 203. doi: 10.25135/acg.oc.122.2203.2397
31
KolanjinathanK.GaneshP.GovindarajanM. (2009). Antibacterial activity of ethanol extracts of seaweeds against fish bacterial pathogens. Eur. Rev. Med. Pharmacol. Sci.13, 173–177.
32
KumarC. S.SaradaD. V.RengasamyR. (2008). Seaweed extracts control the leaf spot disease of the medicinal plant Gymnema sylvestre. Indian J. Sci. Technol.1, 93–94.
33
KurashovE. A.FedorovaE. V.KrylovaJ. V.MitrukovaG. G. (2016). Assessment of the potential biological activity of low molecular weight metabolites of freshwater macrophytes with QSAR. Scientifica2016, 1205680. doi: 10.1155/2016/1205680
34
MashjoorS.YousefzadiM.EsmaeiliM. A.RafieeR. (2016). Cytotoxicity and antimicrobial activity of marine macro algae (Dictyotaceae and Ulvaceae) from the Persian Gulf. Cytotechnology68, 1717–1726. doi: 10.1007/s10616-015-9921-6
35
McGurrinA.Suchintita DasR.SoroA. B.MaguireJ.Flórez FernándezN.DominguezH.et al. (2025). Antimicrobial activities of polysaccharide-rich extracts from the Irish seaweed Alaria esculenta, generated using green and conventional extraction technologies, against foodborne pathogens. Mar. Drugs23, 46. doi: 10.3390/md23010046
36
MichalakI.ChojnackaK. (2015). “ Production of seaweed extracts by biological and chemical methods,” in Marine Algae Extracts: Processes, Products, Applications. Eds. KimS. K.ChojnackaK. ( WILEY–VCH Verlag GmbH & Co. KGaA, Weinheim), 121–144. doi: 10.1002/9783527679577.ch7
37
MickymarayS.AlturaikiW. (2018). Antifungal efficacy of marine macroalgae against fungal isolates from bronchial asthmatic cases. Molecules23, 3032. doi: 10.3390/molecules23113032
38
O’KeeffeE.HughesH.McLoughlinP.TanS. P.McCarthyN. (2019a). Methods of analysis for the in vitro and in vivo determination of the fungicidal activity of seaweeds: A mini review. J. Appl. Phycol.31, 3759–3776. doi: 10.1007/s10811-019-01832-7
39
O’KeeffeE.HughesH.McLoughlinP.TanS. P.McCarthyN. (2019b). Antibacterial activity of seaweed extracts against plant pathogenic bacteria. J. Bacteriol. Mycol.6, 1105.
40
OsmanM. E. H.AbushadyA. M.ElshobaryM. E. (2013). Production and characterization of antimicrobial active substance from some macroalgae collected from abu-qir bay (Alexandria) Egypt. Afr. J. Biotechnol.12, 6847–6858. doi: 10.5897/AJB10.2150
41
PadmakumarK.AyyakkannuK. (1997). Seasonal variation of antibacterial and antifungal activities of the extracts of marine algae from southern coasts of India. Bot. Mar.40, 507–515. doi: 10.1515/botm.1997.40.1-6.507
42
PalaniyappanS.SridharA.KariZ. A.Téllez-IsaíasG.RamasamyT. (2023). Evaluation of phytochemical screening, pigment content, in vitro antioxidant, antibacterial potential and GC-MS metabolite profiling of green seaweed Caulerpa racemosa. Mar. Drugs21, 278. doi: 10.3390/md21050278
43
PereiraL. (2021). Macroalgae. Encyclopedia1, 177–188. doi: 10.3390/encyclopedia1010017
44
PlouguernéE.Le LannK.ConnanS.JechouxG.DeslandesE.Stiger-PouvreauV. (2006). Spatial and seasonal variation in density, reproductive status, length and phenolic content of the invasive brown macroalga Sargassum muticum (Yendo) Fensholt along the coast of Western Brittany (France). Aquat. Bot.85, 337–344. doi: 10.1016/j.aquabot.2006.06.011
45
PourakbarL.MoghaddamS. S.EnshasyH. A. E.SayyedR. Z. (2021). Antifungal activity of the extract of a macroalgae, Gracilariopsis persica, against four plant pathogenic fungi. Plants10, 1781. doi: 10.3390/plants10091781
46
RadhikaD.VeerabahuC.PriyaR. (2012). Antibacterial activity of some selected seaweeds from the Gulf of Mannar Coast, South India. Asian J. Pharm. Clin. Res.5, 9–90.
47
RodrigoB. K. D. M.AlahakoonA. H. D.BalasooriyaB. C. M.EdirisingheP.HerathH. M.WanigatungeR. P. (2025). Antifungal activity of extracts from Ulva, Sargassum, and Gracilaria against three fungal pathogens and GC-MS analysis of the most effective extracts. Int. J. Secondary Metabolite12, 331–342. doi: 10.21448/ijsm.1506431
48
SavaryS.WillocquetL.PethybridgeS. J.EskerP.McRobertsN.NelsonA. (2019). The global burden of pathogens and pests on major food crops. Nat. Ecol. Evol.3, 430–439. doi: 10.1038/s41559-018-0793-y
49
SelimR. E.AhmedS. M.El-ZemityS. R.RamsesS. S.MoustafaY. T. (2015). Antifungal activity and seasonal variation of green alga (Ulva lactuca) extracts. Asian J. Agric. Food. Sci.3, 419–427.
50
SheikhH.El-NaggarA.Al-SobahiD. (2018). Evaluation of antimycotic activity of extracts of marine algae collected from Red Sea Coast, Jeddah, Saudi Arabia. J. Biosci. Medicines6, 51–68. doi: 10.4236/jbm.2018.64004
51
ShobierA. H.GhaniS. A. A.BarakatK. M. (2016). GC/MS spectroscopic approach and antifungal potential of bioactive extracts produced by marine macroalgae. Egypt. J. Aquat. Res.42, 289–299. doi: 10.1016/j.ejar.2016.07.003
52
SivakumarK.KannappanS.DineshkumarM.PatilP. K. (2014). Evaluation of marine macro alga, Ulva fasciata against bio-luminescent causing Vibrio harveyi during Penaeus monodon larviculture. Afr. J. Microbiol. Res.8, 803–813. doi: 10.5897/AJMR2013.6233
53
ToledoE.FélixC.VicenteT. F.AugustoA.FélixR.ToledoB.et al. (2023). Seaweed extracts to control postharvest phytopathogenic fungi in Rocha pear. J. Fungi9, 269. doi: 10.3390/jof9020269
54
TriguiM.GasmiL.ZouariI.TounsiS. (2013). Seasonal variation in phenolic composition, antibacterial and antioxidant activities of Ulva rigida (Chlorophyta) and assessment of antiacetylcholinesterase potential. J. Appl. Phycol.25, 319–328. doi: 10.1007/s10811-012-9866-5
55
VeraJ.CastroJ.GonzalezA.MoenneA. (2011). Seaweed polysaccharides and derived oligosaccharides stimulate defense responses and protection against pathogens in plants. Mar. Drugs9, 2514–2525. doi: 10.3390/md9122514
56
VicenteT. F.LemosM. F.FelixR.ValentaoP.FélixC. (2021). Marine macroalgae, a source of natural inhibitors of fungal phytopathogens. J. Fungi7, 6. doi: 10.3390/jof7121006
57
VurroM.BoncianiB.VannacciG. (2010). Emerging infectious diseases of crop plants in developing countries: impact on agriculture and socio-economic consequences. Food Secur.2, 113–132. doi: 10.1007/s12571-010-0062-7
58
WasihunY.Alekaw HabteweldH.Dires AyenewK. (2023). Antibacterial activity and phytochemical components of leaf extract of Calpurnia aurea. Sci. Rep.13, 9767. doi: 10.1038/s41598-023-36837-3
59
ZhangY.HawboldtK.MacQuarrieS.ThomasR.GebregiworgisT. (2024). Alkaline subcritical water extraction of bioactive compounds and antioxidants from beach-cast brown algae (Ascophyllum Nodosum). Chem. Eng. J.494, 153109. doi: 10.1016/j.cej.2024.153109
Summary
Keywords
antimicrobial potential, bioactive compounds, plant disease management, poisoned plate method, well diffusion assay
Citation
Alahakoon AHD, Rodrigo BKDM, Edirisinghe P, Herath HM and Wanigatunge RP (2026) Anti-phytopathogenic activity and GC–MS profiling of bioactive fractions derived from three marine macroalgae of Sri Lanka. Front. Mar. Sci. 13:1816866. doi: 10.3389/fmars.2026.1816866
Received
24 February 2026
Revised
10 June 2026
Accepted
10 June 2026
Published
01 July 2026
Volume
13 - 2026
Edited by
Prof. Vaibhav A. Mantri, Central Salt & Marine Chemicals Research Institute (CSIR), India
Reviewed by
Lijian Ding, Ningbo University, China
Çağla Yarkent, Ege University, Türkiye
Badr-ddine El Mouns, Chouaïb Doukkali University, Morocco
Updates
Copyright
© 2026 Alahakoon, Rodrigo, Edirisinghe, Herath and Wanigatunge.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: R. P. Wanigatunge, rasikaw@kln.ac.lk
Disclaimer
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.