Extracellular vesicles modulate growth and stress adaptation in Fusarium circinatum

doi:10.21203/rs.3.rs-5308708/v1

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Extracellular vesicles modulate growth and stress adaptation in Fusarium circinatum

https://doi.org/10.21203/rs.3.rs-5308708/v1

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The molecular mechanisms influencing Fusarium circinatum, an important pine tree pathogen, remain partially understood. We recently reported a biofilm-mediated response in this fungus, which supports its adaptation to harsh conditions including heat stress. Herein, we report that biofilm extracellular vesicles (EVs) play a key role in this adaptive response. The EVs were purified from planktonic and biofilm cells via differential ultracentrifugation and size exclusion chromatography. Their subsequent application to fungal cells revealed the capacity of biofilm-derived EVs (bEVs) to promote conidial viability and germination. When bEV-treated conidia were cultured in the presence of carbon sources (PM1), nitrogen sources (PM3B), and chemical sensitivity agents (PM21D), a delayed nutrient utilization and adaptation to antimicrobial agents such as nystatin, was observed. Furthermore, exogenous application of bEVs on mono- and polymicrobial biofilms significantly enhanced biomass and matrix production, with EVs derived from heat-stressed biofilm (45 ºC, 1 hour) showing more effectiveness at promoting biomass production and resistance to the antifungal agent, tebuconazole. This is consistent with the biofilm’s heat resistance previously reported for F. circinatumbiofilms. Taken together, our work provides novel insights into the EV-mediated molecular interactions that modulate environmental responses in F. circinatum.

Forestry

Pathology

Biofilm

extracellular vesicles (EVs)

microbial resilience

Pine Pitch Canker

Fusarium circinatum

polymicrobial biofilms

While extracellular vesicles (EVs) have been well-studied in bacterial and mammalian systems, their roles in fungal biofilms, especially in phytopathogens like Fusarium circinatum, remain largely unexplored. EVs include exosomes and microvesicles (MVs), both representing a group of heterogeneous spherical lipid-bilayer nanoparticles released by cells to the extracellular space (Yuana et al., 2013, Kwok et al., 2021). These EVs are secreted by all prokaryotes and eukaryotes so they can ferry biologically active compounds such as lipids, proteins, nucleic acids (i.e., DNA, RNA), and metabolites (Yáñez-Mó et al., 2015, Tkach and Théry, 2016, Van Niel et al., 2018). Consequently, the internalization of EVs by recipient cells can alter their pathological or physiological state (Kwok et al., 2021:), making the assessment of exogenously applied EVs possible.

Some of the biological functions previously reported for EVs analysed from mammalian studies include cell communication, cell homeostasis as well as pathogenesis (e.g., in cancer) (Bellingham et al., 2012, Yáñez-Mó et al., 2015, Maas et al., 2017, Sexton et al., 2019). In phytopathogenic filamentous fungi, EVs are not well studied, although a few previous studies have demonstrated their role in disease progression, transport of virulence factors, cell wall remodelling, antimicrobial resistance, and biofilm formation (Rodrigues et al., 2007, Bielska et al., 2018, Zarnowski et al., 2018, Kalra et al., 2019, Honorato et al., 2021). Similar studies are currently lacking in a wide range of economically important pathosystems involving filamentous pathogens of plants, and are absolutely non-existent in tree-infecting fungi such as Fusarium circinatum.

Fusarium circinatum is responsible for causing the Pine Pitch Canker (PPC) disease in Pinus species (pine) and Pseudotsuga menziesii (Mirb.) Franco (Wingfield et al., 2008, Martínez‐Álvarez et al., 2014). This pathogen is predominantly responsible for nursery seedling death and declining timber production, often associated with major economic losses. This makes PPC one of the most important limitations in commercial forestry (Wingfield et al., 2008, Mitchell et al., 2011, Bezos et al., 2018, Zamora-Ballesteros et al., 2019), and by default, circinatum an important pathogen to study. Recent studies have linked EVs with virulence of some of the most economically significant phytopathogenic fungi including Fusarium oxysporum f. sp. vasinfectum (Bleackley et al., 2020, Hill and Solomon, 2020, Garcia-Ceron et al., 2021), Botrytis cinerea (De Vallée et al., 2023), and Magnaporthe oryzae (He et al., 2023), and Colletotrichum higginsianum (Rutter et al., 2022). In addition, studies conducted in Candida albicans, an opportunistic human fungal pathogen, show that EVs are crucial in surface-associated microbial communities (biofilms) as they can promote matrix production and antifungal resistance (Zarnowski et al., 2018). Therefore, understanding the role of EVs in fungal biofilms could provide new insights into microbial resilience and open avenues for the development of novel biofilm management strategies. Given the agricultural and economic significance of F. circinatum (Wingfield et al., 2008, Mitchell et al., 2011, Nordström et al., 2022), studying biofilm-derived EVs may lead to the development of more effective control measures against this pathogen, and therefore, to improving forestry management.

In the present work, the aim was to study the role of exogenously applied EVs and their impact on F. circinatum growth using phenotypic analysis of spores that have internalized the EVs. The extent of these EV effects was further investigated via high throughput phenotypic microarray (PMs) (Boschner et al., 2001). The PM technique can assess the ability of microorganisms to metabolize hundreds of conditions simultaneously and, thus it is reliant on microtiter-plate-based substrate utilization (Bochner and Savageau, 1977, Mishra et al., 2022). Metabolic rate is measured using redox-sensitive dyes (e.g., tetrazolium-based dyes) to indicate metabolic activity by changing colour in response to growth or respiration on nutrient sources. As a result, PMs provide a fingerprint that is useful in determining the metabolism of a microbe in the presence of exogenous stimuli such as EVs (Bochner et al., 2001, Bochner, 2003).

Furthermore, we explored the possibility that EVs released by fungi can be shared as a communal good, benefiting other members of a community from the biological activity of cargos contained on or inside these EVs. Our work also suggests that polymicrobial biofilms, which are likely an occurrence in compatible strains or species of a microbial community, may facilitate the exchange of EVs between community members. Taken together, our results suggests that polymicrobial biofilms, by facilitating the exchange of EVs, may promote communication, resistance and adaption in fungi. Therefore, our research on biofilm-derived EVs could lead to novel breakthroughs in controlling infections caused by F. circinatum.

2.1. Culture growth and maintenance

F. circinatum FSP34 (CMW350) and all of its nursery isolates (Table 1) was obtained from the Fusarium collection of the Tree Protection Co-operative Programme, Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, South Africa. The fungal isolates were routinely cultured on quarter-strength potato dextrose agar (¼ PDA) (Merck Group, Modderfontein GP, South Africa) for 7 days and kept in the dark at 25 ºC. For biofilm formation, a culture was flooded with 2 mL of phosphate-buffered saline (0.2 M PBS; 10 mM NaH₂PO₄, 10 mM Na₂HPO₄, 150 mM NaCl, pH 7.2) to obtain conidial cells. These cells were then counted in a Neubauer chamber and adjusted to the desired concentration before downstream analysis.

2.2. Extracellular vesicle isolation and purification

2.2.1 Isolation using ultracentrifugation

Isolation of EVs from planktonic and biofilm cells of F. circinatum FSP34 was performed using differential ultracentrifugation following a previous protocol with minor adjustments (Rodrigues et al., 2007). Briefly, conidia of F. circinatum were collected from 7-day-old cultures grown on ¼ PDA using 0.2 M PBS (pH 7.2). Conidia was inoculated into 200 mL of potato dextrose broth (PDB) to a final concentration of 1 X 10⁶ spores/mL. To maintain the planktonic state of the fungus, spores were allowed to grow under shaking conditions using the orbital shaker (100 rpm, Shake-O-Mat, Labotec, South Africa) while biofilm formation assay was conducted as previously described (Ratsoma et al., 2024), the spores were allowed to grow at static conditions for 72 h. To begin with EV isolation, cells, and debris were removed from the media of planktonic and biofilms by sequential centrifugation steps at 4,000× g for 15 min and 15,000× g for 30 min at 4 °C. The resulting supernatant was then filtered using a 0.45 μm membrane filter (Merck Millipore) to further remove any remaining cells or debris. The Amicon ultrafiltration system (100-kDa MWCO, Millipore) was then used to concentrate the supernatants to a final volume of approximately 20 mL. One part of the concentrated supernatants was subjected to ultracentrifugation (Beckman Coulter, Brea, CA, USA) at 100 000× g for 1 h at 4 °C to collect the EVs. The pellets enriched in EVs were washed twice with PBS at 100 000× g for 1 h at 4° C and stored in PBS as aliquots at -80° C until further use.

2.2.2. Purification of EVs using size exclusion chromatography

For size-exclusion chromatography (SEC), FSP34 EVs were concentrated using the Amicon® Ultra-4 Centrifugal system (100-kDa pore size) and purified as described previously by Kunene et al., (2023 ). This involved application of a mixture containing EVs with a cell membrane-specific fluorescent lipophilic dye (FM4-64; Thermo Fisher Scientific, South Africa) which was incubated for 15 min in the dark at room temperature. After incubation, the sample was added to a 10 mL plastic syringe stuffed with nylon stocking at the tip and stacked with 10 mL sepharose CL-2B (Sigma-Aldrich, South Africa), equilibrated with PBS. The sample was then eluted with PBS to collect 30 sequential fractions of 0.5 mL in black microtiter plates (Greiner, South Africa). Fraction fluorescence (excitation at 560 nm, and emission at 734 nm) was measured immediately using a Spectra Max M2 plate reader (SpectraMax paradigm, Multimode detection platform). Fractions with fluorescence levels above 3.0 relative fluorescence units (RFU) were pooled together, as "EV signal" after measurement and stored in PBS as aliquots at -80 ^oC until further use.

2.2.3. Physical characterization of EVs

Purified EVs of F. circinatum FSP34 were spotted on carbon-coated grids for adsorption for 5 min. The exposed vesicles were then negatively stained with 1% (w/v) uranyl acetate for 3 min. Finally, the EVs were visualized using transmission electron microscopy (TEM; JOEL JEM 2100F, JOEL Ltd., Tokyo, Japan).EVs were visually analysed using scanning electron microscopy. For this purpose, sterile 65 mm petri dishes containing glass coverslips and 200 mL PDB were inoculated with 20 µL conidial cells (to a final concentration of 2 x 10⁵ cells /mL) and statically incubated at RT for 7 days. Glass slides were then removed and flooded and rinsed with PBS prior to adding the pre-fixative solution containing 1 mL of 2.5% (v/v) glutaraldehyde (Merck, South Africa) / formaldehyde (Merck, South Africa). After another PBS rinse, biofilms were fixed with 1% osmium tetroxide for 1 h. Following a final PBS rinse, the fixed biofilms were dehydrated sequentially using a series of graded ethanol (i.e., 15 minutes rinses each in 1 mL of ethanol at concentrations of 30 %, 50%, 70%, 90%, and three rinses in absolute ethanol). The dehydrated samples were then treated with a 50:50 mixture of hexamethyldisilazane (HMDS) and absolute ethanol for 1 h, followed by a treatment with HMDS only, after which they were left to dry overnight. The glass slides were mounted on rectangle aluminium stubs and carbon coated for 15 min using Qourum Q150T ES sputter coater (Qourumtech, UK). The stubs were observed in a JEOL JSM 6490LV scanning electron microscope (GenTech Scientific Inc., Arcade, NY, USA).

Particle size distribution and concentration of EV signals were measured using NanoSight NS500 (Malvern Panalytical, UK). The samples were vortexed for 1 minute, and a 5 mL sample was diluted with 5 mL of PBS (dilution factor of 2). Immediately after, a test volume of 1.5 mL of each sample was injected into the NTA and analysed in triplicate (each run = 30 s video). PBS was treated as a blank. The videos were captured and analysed using NTA 3.3 Dev Build 3.3.104 (Malvern Panalytical, UK). The camera sensitivity and detection threshold were optimized per video, and the temperature was set at 22 °C.

2.3. EV protein content quantification

The protein content of EVs was quantified to act as EV concentrations using the QuntiPro-BCA assay Kit (Walker, 2002). Therefore, following manufacturers' guidelines 20 μL of EV sample was added to a 96-well microplate followed by 200 μL of BCA reagent (Merck, South Africa). The plate was then incubated in the dark for 2 h at 37 °C. Post-incubation absorbance of the plate was measured at 560 nm and protein concentration was determined from a bovine serum albumin (BSA) standard curve.

2.4. Uptake analysis of biofilm derived-EVs by planktonic cells

2.4.1. EV membrane staining

The labelling of the biofilm derived-EVs (bEVs) was performed according to Regente et al., (2017). Briefly, purified vesicles were suspended in 40 μL of PBS and mixed gently with FM4–64 (Molecular Probes, Thermo Fisher Scientific, Argentina) to a final concentration 1 μg/mL and kept in the dark for 60 min on ice. After which, labelled samples were diluted with 3 mL PBS and ultracentrifuged at 100 000 x g to remove excessive dye, this was performed twice. The obtained pellet finally resuspended in 20 μL PBS.

2.4.2. Uptake of biofilm derived-EVs by FSP34 planktonic cells

A conidial suspension of F. circinatum adjusted to about 10 000 cells was incubated on a glass slide with 2 μL of bEVs labelled with 5 μg/mL of FM4–64. The cells were then visualized under confocal microscopy after 5 min of incubation at room temperature. Control treatments were performed by incubating conidia with PBS instead of FM4–64 labelled EVs. Following this, a microscopic examination of the cells was performed using a 63x oil immersion lens (ZEISS, CSLM), with FM4–64 excited at 488 nm and detected at 650-750 nm.

2.4.3. EV add-back assay of biofilm-derived EVs on FSP34 planktonic cells

Conidial suspension containing about 1500 cells was incubated with a mixture of 5 μL of purified bEVs (5 μg/mL) and 4% sucrose to a final volume of 20 μL. After 16 h of incubation at 25 °C, 5 μL of the mixture was evaluated for the presence and morphology of hyphae using a light microscope (LM, ZEISS, South Africa). Controls were performed by replacing EVs with the same volume of PBS. The experiment was performed in triplicate and repeated on different days. To perform the germ tube presence and cell viability analysis,the conidial suspension was co-incubatedwith bEVs as before, but this time around the cells were incubated for up to 24 h shaking at 25 °C in the dark. During the incubation period, cells were analysed every 24 h and at each time point, 20 μL of the cells were stained with tryphan blue (Merk, South Africa) and assessed for germ tube formation and cell viability. This assay was performed in triplicates using the countess 3FL cell counter (Invitrogen, ThermoFisher Scientific).

2.4.4. The effects of bEVs on pre-formed FSP34 biofilms

For this experiment, 72 h pre-formed biofilms of FSP34 were exposed to 5 μg/mL of either bEVs or heat-treated bEVs of FSP34 and incubated for 24 h. Post incubation, we quantified biomass, extracellular matrix (ECM), and metabolic activities by performing crystal violet, safranin assay, and XTT reduction assay, respectively with minor modifications, according to Ratsoma et al., (2024). Conidial suspension without bEVs was considered a positive control while the negative controls included conidial suspension with heat-treated (90 °C for 15 min) bEVs as well as ¼ PDB.

2.4.5. The effects of EVs purified from FSP34 heat-treated biofilm cultures on thebiofilm of nursery field isolates

*Fusarium circinatum isolates treated with vesicles released by heat treated biofilms (h-bEVs).

EVs were also purified from 72 h- old FSP34 biofilms that were subjected to heat at 45 ºC for an hour using SEC, as previously described. The FSP34 strain was then considered as an EV donor isolate. A total of 10 nursery field isolates of F. circinatum originating from diseased Pinus spp. trees found in South Africa were obtained from the Fusarium culture collection (CMWF) maintained at the Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, South Africa (Coutinho et al., 2007, Steenkamp et al., 2014) (Table 1). Among these, strains CMWF2597, CMWF2625, CMWF535, and CMWF568 were selected based on their antifungal response profiles to be recipients of EVs in order to understand the effects of EVs in promoting biofilm integrity and antifungal resistance. The source of EVs was prepared by forming FSP34 biofilms for 72 h under normal conditions (stationary at RT) and stressful conditions (72 h pre-formed biofilm) followed by 1 h exposure to heat at 45 ºC (Ratsoma et al., 2024). EVs from these sources were then purified using SEC as before and 5 µg/ml of these bEVs were applied to 72 h recipient biofilms formed by nursery isolates on microtiter plates to assess their impact on biofilm fortification.

2.5. The effect of bEVs on mono-and polymicrobial biofilm of nursery field isolates

Monomicrobial biofilms were established by inoculating each conidial suspension from individual isolates, while for polymicrobial biofilms made up of two isolates were co-incubated to a final concentration of 2 × 10⁵ cells/mL in 96‐well flat‐bottom polystyrene plates containing 200 μL of PDB. At 72 h, pre-formed mono- and polymicrobial biofilms of nursery field isolates were exposed to 5 μg/mL of bEVs from FSP34 and incubated for 24 h. Post incubation, we quantified biomass, ECM, and metabolic activities by performing crystal violet, safranin assay, and XTT reduction assay, respectively with minor modifications, according to Ratsoma et al., (2024). Pre-formed mono- and polymicrobial biofilms without bEVs were considered a positive control while the negative control contained ¼ PDB only.

2.6. Global nutrient profiling of FSP34 spores treated with biofilm-derived EVs

F. circinatum utilization and assimilation profiles were generated for the FSP34 strain using the Filamentous fungi (FF) MicroPlates (Biolog®), namely, PM1, PM3B, and PM21D (Anatech, South Africa), respectively. For this analysis, we investigated the growth and metabolism of planktonic cells following co-incubation with bEVs. Each panel of the FF microplates contains 96 wells of which 95 wells represent specific carbon and nitrogen sources, while the one well contained water representing a negative control (Table S1). For chemical sensitivity analysis, each panel of the 96 well FF plate represented 24 different chemical agents (Table S1). For inoculum preparation, sterile swabs were soaked in PBS and then gently rolled over the mycelia of 7-day-old cultures. The spores on the swabs were then resuspended in 12 ml of FF inoculation fluid (FF-IF) (Anatech, South Africa) and mixed gently. The transmittance of this suspension was adjusted to 62% (0.1) using the spectrophotometer (SpectraMax paradigm, Multimode detection platform) at 590 nm. The FF inoculation fluid (IF) was prepared as shown in Table S1. A hundred microliters of FF-IF mixture were added to each well and incubated at 25 ^oC in the dark for 7 days. FF-IF mixture with bEV-treated spores was considered a positive treatment while the mixture with spores without bEVs was considered non-treatment (negative control), where PBS was substituted for bEVs. This experiment was performed in triplicate and readings were taken every 24 h for 7 days. Growth was measured using the spectrophotometer at an optical density (OD) of 750 nm to assess the cellular biomass (Tanzer et al., 2003).

2.7. Reproducibility and statistical analyses

Data was presented as mean ± standard error of the mean (SE). Statistical analyses were performed using GraphPad statistical software (GraphPad 8 Software, San Diego, CA, USA). The principal component analysis (PCA) plots were performed using R studios 4. 3 .2. For statistical analysis of viable counts, cultures containing 2 x 10⁵ and 1 X 10^6, etc. CFU/ mL, were respectively used for EV isolation, biofilm formation and uptake analysis. All experiments were performed in triplicates.

3.1. Physical properties of EVs

Using TEM, we observed spherical, rosette, and typical cup-shaped EV morphologies (Fig 1 A-C) secreted by F. circinatum. The size of the EVs ranged from 100-200 nm. We also observed the secretion of EV-like structures on the hyphae of the biofilm cells of F. circinatum (Fig 1 D). Size distribution results by NTA, revealed that both the planktonic-derived and biofilm-derived EVs (bEVs) had varying EV concentrations up to 1.8 x 10⁸ particles per mL, with a size range of up to 200 nm. We also observed that planktonic-derived EVs have smaller-sized EVs (50-90 nm), while larger EVs were observed from bEVs (90-120 nm).

3.2. Effects of exogenously applied bEVs during early planktonic cell growth

Florescence and LM uptake results of the co-incubation experiment between bEVs and planktonic cells were based on the FM4-64 fluorescence dye signal. These results revealed that the FSP34 planktonic cells were able to internalize its own bEVs after 5 min (Fig 2 A). This was illustrated by a red- fluorescence accumulation inside the cells and not on the surface of the spores, which indicated internalization of the EVs by the spores (Fig 2 B). Interestingly, LM analysis revealed that the fungal spores treated with bEVs germinated into hyphae whereas those not treated exhibited no signs of germination after 16 h of co-incubation with bEVs (Fig 2 D). When we compared the treatment with the control groups, the results of the germ tube assay, as depicted in Fig 3, revealed early germination of spores treated with bEVs at significant levels after 2 h (p < 0.0008) and 4 h (p < 0.007). Surprisingly, according to the cell viability assessment, planktonic cells treated with bEVs had a lower viability, especially after 2 (p < 0.001) and 4 h (p < 0.0001), compared to the negative control. However, viability increased after 24 h (p < 0.0001) (Fig 3 B).

3.3. Influence of FSP34 EVs on pre-formed biofilms

A significant increase in biomass and ECM production was observed after subjecting FSP34 pre-formed biofilms to its own bEVs (Fig 4 A-B; p < 0.005). Additionally, biofilms exposed to heat-treated bEVs showed a significant increase in both biomass and ECM production as compared to biofilms were no EVs were added (A-B). The pre-formed biofilms also exhibited reduced metabolic activities in the presence of heat-treated and non-heated bEVs as treatments (Fig 4 D). The treatment with heat-treated bEVs on pre-formed biofilms showed that they are heat resistant, as no significant differences were observed between the ECM produced by pre-formed biofilms treated with heat-treated bEVs compared to pre-formed biofilms treated with non-heat-treated bEVs. Combined, these data suggest that bEVs play an important role in influencing the biological response of F. circinatum to biofilm dynamics.

3.4. Cross-isolate effects of FSP34 EVs on biofilm fortification

Previously we observed that the addition of bEVs from FSP234 strain on its own pre-formed biofilm re-establishes ECM accumulation and increases biomass (Fig 5). As EVs are released into the environment, we suspected that these EVs could be taken up by other members of a community, who can in turn benefit from this interaction. To test this hypothesis, we cultured FSP34 biofilms for 72 h at RT and derived two groups of preformed biofilm cultures, one where there was no application of heat and one exposed to heat (45 ºC) for one hour (Ratsoma et al., 2024). From the supernatant of these cultures, we purified bEVs via SEC. The EVs (5 µg/ml) were then exogenously applied as before to a 72-h biofilm formed by F. circinatum nursery isolates (Table 1). The data suggests that bEVs purified from heat-exposed biofilms are more effective at enhancing biofilm traits, including biomass and metabolic activity (e.g., in strain CMWF2597 and CMWF2625) (Fig 5 A-B). In strain CMWF2597, tebuconazole induced significant inhibition of ECM and, interestingly, the addition of bEVs restored ECM production. In several strains, bEVs imposed an antagonistic effect when combined with tebuconazole, which was evident during biomass production (CMWF259 and CMWF568) and ECM production (CMWF2597 and CMWF535).

3.4. The impact of bEVs on polymicrobial biofilms

We further investigated whether FSP34 EVs can impact multi-strain biofilms, since in a biofilm context where cells are close to one another, the EVs can easily be exchanged. To test this hypothesis, FSP34 bEVs from a nonheated biofilm were introduced to mono- and polymicrobial biofilms formed by the nursery isolates to examine their effects on the biomass, metabolic activity and ECM production. The results showed that, compared to the untreated biofilm, the presence of FSP34 bEVs significantly increased the biomass and ECM of the monomicrobial biofilms in 30% and 50% and of polymicrobial biofilms in 50% and 0% of nursery field isolates (Fig 6 A and B), respectively. This suggests that in polymicrobial settings, the role of bEVs might be more directed to promoting cell growth and biomass accumulation rather than enhancing the biofilm's ECM. Except in 30% of isolates that formed a monomicrobial biofilms, bEVs inhibited metabolic activity in all scenarios (Fig 6 C). It is possible that different strains within the biofilm compete for resources, or that the vesicles influence biomass differently in mixed-species communities, where ECM production might not be the primary focus for survival.

3.5. Global nutrient profiling of bEVs treated Fusarium circinatum

3.5.1. Effects of bEVs on carbon source assimilation

Since we observed the morphological effects of bEVs following exposure to fungal spores including the ability of spores to germinate, cell viability, and pre-formed biofilms (i.e., biomass and ECM production) (Motaung et al., 2023), we were interested to determine from what metabolic pathways could these effects originate. For this, we used the high-throughput PM analysis using PM1, PM3B, and PM21D biolog 96-well microtiter plates. Based on growth (biomass) analysis the substrates were divided into three categories by optical density (OD_750nm) readings as follows; low-poor (0-0.4), moderate (0.4-0.7), and robust growth (> 0.7).

The analysis of growth in carbon sources present in the PM1 plate showed that fungal cells treated with bEVs exhibited positive biomass production indicated by high optical density (0.8-1) values on eight out of the 95 substrates tested, while the remainder of the substrates had moderate to low growth (0.7- 0.4) (Fig 7 A). These include significant growth enhancement observed at 168 hours (7 days) (Fig 6 B) across a variety of simple sugars and derivatives such as D-trehalose (p = 0.0010), D-glucuronic acid (p = 0.0320), and D-galacturonic acid (p = 0.0171). Similarly, polyols, including D-mannitol (p = 0.0001) and adonitol (p < 0.00001), also resulted in significant growth enhancement. The same trend was observed with complex carbohydrates such as maltotriose (p < 0.00001) and D-cellobiose (p < 0.00001). Furthermore, treatment by the non-sugar surfactant, Tween 20 (p = 0.0208), also resulted in increased growth. All the substrates tested led to increased growth with bEV treatment when compared to non-treated controls.

The negative control, which is F. circinatum spores without bEV treatment, showed a significant increase in biomass production (p < 0.001) in the presence of carbon sources (OD_750nm 0.4 - 0.7) L-arabinose, N-acetyl-D-glucosamine, L-aspartic acid, D-trehalose, D-mannose, dulcitol, D-ribose, L-rhamnose, α-D-glucose, D-melibiose, α-D-lactose, α-methyl-D-galactoside, sucrose, β-methyl-D-glucoside, L-alanyl-glycine, and glycyl-L-proline. It is noteworthy that spores without bEV treatment were unable to completely utilize some of the carbon sources including D-galactonic acid- ϒ-lactone (p < 0.001) and glyoxylic acid (p < 0.005) as compared to the positive control (bEVs) treatment (Fig 6 A). Compared to the negative control, bEV treatment led to a general delay of in the utilization of carbon sources (Fig 7 A, Fig S1). Interestingly, of the sugars producing significant biomass production in the presence of bEVs, D-mannitol sugar alcohol was highly assimilated (Fig S2).

3.5.2. Effects of bEVs on carbon nitrogen source assimilation

Looking at 11 nitrogen sources present in PM3B, as depicted in Fig 8 A-B, treatment with bEVs appeared to enhance biomass production. More specifically, ammonium formate, ethylamine, adenosine, and uridine resulted in increased biomass production due to bEVs, all with p-values below 0.05 as compared to the negative control devoid of EVs. bEV treatment also led to in significant biomass increases (p < 0.05) compared to the negative control in the presence of amino acids including L-cysteine, L-glutamic acid, L-threonine, L-citrulline, and amino sugars including D-glucosamine, and D-mannosamine. Conversely, significant biomass increase in the non-treatment group was observed in the presence of nitrogenous compounds (e.g., nitrite, nitrate, urea, bluret, L-glutamine, and guanine), peptides (e.g., Ala-Glh, Ala-Glu, and Ala-Leu), proteinogenic amino acids (e.g., L-histidine, L-glycine, L-arginine, and L-alanine) - all of which demonstrating p-values less than 0.05.

Poor to no utilization (0-0.4 OD_750nm) of the proteinogenic amino acids L-phenylalanine (p > 0.5), L-proline (p = 0.4111), was observed in both the negative control (without bEVs) and bEV-treatment groups, while L-methionine (p < 0.1) was fairly utilized by the negative control. Assessment of the nitrogen sources also revealed that the bEVs treatment delayed biomass production compared to the negative control, of which the bEVs treatment only increased biomass production after 24 h (Fig 8 A, Fig S3). Although ammonium formate is second best to L-arginine with regards to biomass production (Fig 8 B), it results in the best assimilation over time (Fig S4).

3.5.3. Effects of bEVs on chemical sensitivity

We also examined whether bEV treatment can protect fungal cells against harsh chemical agents, including antifungal agents (Fig 9, Fig S5-7). For this purpose, we generated a chemical sensitivity fingerprint based on the PM21D plate, which contains agents such as antibiotics, osmolytes, and potentially toxic compounds. Growth of cells treated with bEVs on this plate reveals that bEVs may result in F. circinatum adapting to a wide range of harmful chemicals (Fig 9 A-B). The analysis showed that in comparison to the non-treatment groups (negative control), treating spores with bEVs results in significant amounts of biomass production in the presence of eight out of 24 chemicals in at least two or more concentrations of the active substrates (Fig 9 A-B). Specifically, this was the case in the presence of guanidine hydrochloride (0.1 < p > 0.001), 2,2-dipyridyl (p < 0.001), promethazine (p < 0.0001), EDTA (0.1 < p > 0.01), sodium dichromate, and magnesium chloride (0.01 < p > 0.0001). We also observed significant variations in growth patterns in the presence of antimicrobial agents including neomycin, nystatin, and protamine sulphate. In comparison to chemical compounds showing increased fungal growth from the negative control, the assimilation of the E05 manganese (II) chloride was slightly higher over time when compared to the other substrates such as D12 sodium dichromate which shows relatively low assimilation (Fig S7).

3.5.4 Impact of bEVs on metabolic diversity in Fusarium circinatum

At this stage, it was apparent given the above results that bEVs lead to more metabolic capacity in F. circinatum. To look further into this, and to obtain a visual overview of the effects of treatment conditions, we generated Principal Component Analysis (PCA) plots for nutrient and chemical compound assimilation in the PM plates with spores exposed to bEVs and nonexposed groups (Fig 10). The nontreated groups, shown as (-)bEVs (Fig 10 A, Left plot) exhibited an obvious dense clustering for PM1 (Red), dispersed clustering for PM21D (Blue), and a more tight clustering for PM3B (Green). The introduction of EVs, as demonstrated in (+)bEV (Fig 10 B, Right plot), had a great influence, resulting in the cluster spreading in PM1 (Red), suggesting that EVs may results in diverse metabolic response. In the case of PM21D (Blue), the grouping of the samples was maintained, however, there was a noticeable spread along the PC2, which was almost similar to PM3B along PC1. In short, the introduction of EVs resulted in the spreading of data points across the principal component axes (PC1 and PC2), especially when it comes to PM1 and PM21D, suggesting the molecules inside these vesicles have the potential to increase metabolic diversity in F. circinatum.

Studying EVs from plant fungal pathogens is important given they have been reported to affect a broad range of cellular functions (Albuquerque et al., 2008, Lin et al., 2017, Bielska et al., 2018, Bitencourt et al., 2022). In this study, EVs isolated from planktonic and biofilm cells displayed typical spherical and cup-shaped morphological characteristics as well as the size distribution (50-200 nm), consistent with previous studies (Garcia‐Ceron et al., 2021, Zarnowski et al., 2021). Similar observations were reported in other studies investigating EVs from plant pathogenic filamentous fungi (e.g., Zymoseptoria tritici, F. oxysporum f. sp. vasinfectum, and F. graminearum) (Bleackley et al., 2020, Garcia‐Ceron et al., 2021, Hill & Solomon, 2020), suggesting similar mechanisms of EV generation in diverse fungal species. Furthermore, the correct purification and physical characterization of EVs is crucial for downstream analyses, which paves the way for better understanding their role in various functions. As previously reported in our recent study (Ratsoma et al., 2024), F. circinatum displays a biofilm-like lifestyle that is responsive to a wide variety of abiotic conditions including azole antifungals. Additionally, bEVs have previously been reported in a limited number of fungal pathogens to influence biofilm responses (Zarnowski et al., 2018, 2021, 2022a), including competition in mixed-Candida biofilm interactions (Zarnowski et al., 2022b). As a result, we wanted to better understand the metabolic response of F. circinatum to self-produced EVs, which may reflect the complex regulatory processes induced by EVs. In addition, we were keen to know whether EVs can be shared among members of a community and influence mixed-strain/species biofilm interactions.

Our results show that EVs associated with both planktonic cells and biofilms of the FSP34 strain influence fungal growth. However, bEVs were found to be more efficient in influencing planktonic cell biology by inducing hyphal growth and cellular viability. The latter was affected in different phases of exposure, with initial decrease occurring within 2-4 hours and recovery and increased viability within 16-24 hours. This suggests that bEVs carry signals or components that initially induce short-term metabolic suppression or cellular reprogramming and later promote cell survival or even activating protective pathways in response to stress. Similar responses were reported in C. albicans, where EVs supported fungal growth in specific growth mediums and further altered the pathogen’s metabolic profile by influencing cellular metabolism and environmental stress responses (Trentin et al., 2023, Wei et a., 2023).

For the first time in a plant fungal pathogen, we observed that EVs can be shared as a public good among field isolates forming a biofilm either in a monomicrobial or polymicrobial context. This share resource (bEVs) also appears to come with several benefits including enhanced biomass and reduced metabolic activity. High biomass will likely contribute to the longevity and tenacity of biofilms by ensuring a larger population of cells remains in a protected state. Furthermore, high biomass can shield the inner cells from plant immune system attacks and protect the biofilm being detected or targeted by the host's immune system (Mannan et al., 2024). Beyond this, reduced metabolic activity allows the biofilm to remain viable for long periods without needing a constant supply of nutrients (Flemming et al., 2016). This persistence is particularly beneficial for pathogens, as it allows them to survive in hosts or environmental niches until favourable conditions return or immune defences weaken.

In clinical and environmental settings, microbes rarely co-exist as monomicrobial communities. Instead, they predominantly co-exist as polymicrobial communities, with a range of synergistic interactions between constituent populations (Ramírez Granillo et al., 2015, Bowen et al., 2018). For instance, during infection, the effects of synergistic interactions within polymicrobial communities can result in a more severe infection than monomicrobial communities, thus resulting to increased antimicrobial resistance (Murray et al., 2014). EVs are not only well suited to coordinate cell growth and metabolic responses as observed in this study. However, EVs can mediate cell-cell communications within microbial communities and play role in the pathogenesis of infection (Bitencourt et al., 2022). Hendricks and co-workers (2021), using cystic fibrosis model, showed that EVs are involved in trans-kingdom communication in a polymicrobial infection setting by acting as a nutrient source for bacterial co-infections during viral infection. Although EVs seem to play role in virus-associated polymicrobial infections, their role in filamentous phytopathogenic fungal polymicrobial biofilms have never been analysed. In this study, we therefore investigated the possibility that the different nursery isolates of F. circinatum form a polymicrobial biofilm. Furthermore, we also investigated the role of bEVs in this polymicrobial context. Our results revealed three interesting situations: firstly, in comparison to a monomicrobial biofilm, there seemed to be generally more biomass and ECM produced in a polymicrobial biofilm, with the exclusion of a polymicrobial biofilm formed between CMWF535 and CMWF568, suggesting more biofilm formation in this context. Secondly, bEVs selectively enhance biomass more significantly in polymicrobial biofilms and ECM production more in monomicrobial ones. This could reflect differences in how the vesicles influence biofilm dynamics depending on the microbial community composition. Thirdly, different strains or species might have different mechanisms for receiving or responding to bEV signals, leading to diverse outcomes in terms of biomass and ECM production. Overall, bEVs seem to contribute to biofilm stability, either by increasing biomass (ensuring the biofilm can grow and expand) or by enhancing ECM production (strengthening the biofilm structure). In polymicrobial biofilms, where interactions between species are likely more complex, bEVs may focus more on promoting growth rather than matrix production. It would be interesting to study these responses in strains from other fungal species to better understand the effects of these EVs.

As demonstrated by PMs, bEVs enhanced several key phenotypes even when cells were growing outside a biofilm environment. For instance, these EVs enhanced the utilization of various carbon sources, increasing metabolic diversity. The utilization of nutrients is an essential basic requirement for the reproduction, development, and growth of organisms (Dickie, 2007, Brock, 2009, Ene et al., 2014). The fact that bEVs could enhance phenotypes in PMs, even when the cells were not in a biofilm state, indicates that bEVs are not exclusively biofilm-specific in their function. bEVs may also precondition planktonic cells to better handle stress or nutrient challenges. Even though we are unsure of the occurrence of biofilm formation in the microarray plates, the bEVs might still prepare cells for such conditions by activating protective mechanisms or optimizing metabolic pathways. Overall, our data suggests that bEVs benefit both biofilm and planktonic cells by enhancing their ability to adapt to external stimuli, such as diverse nutrient sources or chemical agents.

Further supporting the aforementioned hypothesis, the presence of bEVs exhibited protective effects against toxic compounds. Various toxic chemicals interfere with numerous cellular pathways such as protein synthesis, cell wall synthesis, DNA replication, and cell membrane synthesis, and are used on a broad range of microorganisms (Bochner 2008, Chojniak et al., 2015). Our findings indicate that bEVs from F. circinatum may serve as a protective mechanism against antimicrobial agents, potentially aiding in fungal survival against these chemicals. This is in accordance, with other research that suggests EVs can transport virulence factors, helping pathogens evade antimicrobial effects and promote resistance (Rizzo et al., 2020, Lai et al., 2023, Jiang et al., 2024). We also noted that bEVs can enhance biomass production in the presence of antibiotics, suggesting a complex interaction between fungal growth and antibiotic exposure. Specific compounds such as nystatin and promethazine, exhibit varying effects on fungal growth, indicating the adaptive nature of F. circinatum and the specific cargo enriched in its bEVs, therefore further investigations in the identification of these cargos should be explored.

We also noted that bEVs are resistant to increased temperatures (i.e., 90 ^oC), exhibiting a similar contribution of increased ECM production as non-heated bEVs. This suggests that bEVs are highly thermostable and functionally resilient. This thermostability may allow bEVs to play a crucial role in biofilm formation and maintenance, even in harsh environments where heat is present. It also highlights the potential challenges in controlling biofilms formed by organisms that produce heat-resistant vesicles, emphasizing the need for multi-faceted approaches to biofilm management and eradication. In the context of pine trees where F. circinatum is a major fungal threat, under harsh environmental conditions, bEVs can effectively progress the infection rate of F. circinatum through the metabolic and exopolysaccharide biosynthesis pathways (Vila et al., 2017).

In short, our study supports limited research already conducted in membrane vesicles produced by fungal pathogens, especially their crucial role in promoting biofilm resilience and survival under harsh conditions, including high temperatures. To a preformed biofilm, these EVs can be beneficial for pathogen survival as they can enhance biomass and ECM production in both monomicrobial and polymicrobial biofilms, despite reducing metabolic activity. But they can exert detrimental effects when it comes to plant health as they can be shared between biofilm and non-biofilm (planktonic) cells, thereby enhance their phenotypes and nutrient utilization. Therefore, bEVs have a broader role in microbial adaptation. The thermostability of these EVs and their dynamic impact on cellular viability present significant challenges for biofilm control, emphasizing the need for more comprehensive biofilm management strategies for forestry pathogens.

Credit authorship contribution statement

Francinah Ratsoma: Conceptualization, methodology, investigation, analysis, writing-original draft, writing manuscript and editing. Nthabiseng Mokoena: Methodology, investigation, analysis, writing manuscript and editing. Sokunene Mpupa: Methodology, investigation, analysis, writing manuscript and editing. Quentin Santana: Methodology, investigation, analysis, writing manuscript, and editing. Brenda Wingfield: Conceptualization, methodology, investigation, analysis, writing manuscript and editing. Emma Steenkamp: Conceptualization, methodology, investigation, analysis, writing manuscript and editing Thabiso Motaung: Conceptualization, Methodology, writing manuscript and editing, resources, supervision, and funding acquisition.

Acknowledgments

We thank the Council for Scientific and Industrial Research (CSIR) water research lab which assisted with the Nano tracking particle analysis (NTA). We thank Thuthuka funding instrument (Grant no. 129580) of the South African National Research Foundation (NRF), and the South African National Department of Science and Innovation-NRF Centres of Excellence programme and South African Research Chairs Initiative (Grant No. 98353). We are thankful to Silindile Maphosa who helped generate the PCA plots.

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Table 1. Fusarium circinatum nursery field isolates from different parts of South Africa.

CMWF number	Province/Origin	Region	Host	Reference
CMWF2650	Eastern Cape	Maclear	P. greggii	Santana et al., 2016
CMWF2651	Eastern Cape	Maclear	P. greggii	Santana et al., 2016
CMWF2652	Eastern Cape	Maclear	P. greggii	Santana et al., 2016
CMWF2654	Eastern Cape	Maclear	P. greggii	Santana et al., 2016
*CMWF2597	KwaZulu Natal	Tweefontein	P. greggii	Fru et al. 2017
CMWF2601	KwaZulu Natal	Tweefontein	P. greggii	Fru et al., 2017
*CMWF2625	Mpumalanga	Sabie	P. patula	Fru et al., 2017
CMWF2626	Mpumalanga	Sabie	P. patula	Fru et al., 2017
*CMWF535	Western Cape	George	P. radiata	Santana et al., 2016
*CMWF568	Western Cape	George	P. radiata	Santana et al., 2016
*CMWF350 (FSP34)	USA	California	Pinus spp.	Gordon et al., 1996:

*Fusarium circinatum isolates treated with vesicles released by heat treated biofilms (h-bEVs).

The authors declare no competing interests.

FigureS1nl.jpg
Figure S1: Time-course analysis of biomass production with different carbon sources in the presence and absence of biofilm-derived extracellular vesicles (bEVs). The graphs show the growth kinetics of F. circinatum in response to various carbon sources over a period of 168 hours, comparing treatments with bEVs (black lines) to those without bEVs(green lines).
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Figure S2: Time-dependent carbon assimilation by Fusarium circinatum in response to biofilm-derived extracellular vesicles (bEVs) treatment. In each bar chart, the growth or biomass production was measured at specific time points (0, 72, 168 hours) (OD490 nm), reflecting the assimilation rate of different carbon sources by F. circinatumplanktonic cells over a period of 168 hours.
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Figure S3: Enhanced biomass production in Fusarium circinatumspores with biofilm-derived extracellular vesicle (bEV) treatment across various nitrogen sources. Presented is a series of growth curves showing OD490 nm for F. circinatumspores cultured with different nitrogen sources over a period of 168 hours. Each plot compares the growth kinetics between conditions with bEVs (red lines) and without bEVs (black lines).
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Figure S4: Temporal nitrogen assimilation dynamics in Fusarium circinatumwith biofilm-derived extracellular vesicles (bEVs) treatment. The chart illustrates the assimilation rates of various nitrogen sources by F. circinatum over 168 hours, measured by OD490 nm.
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Figure S5: Effect of biofilm-derived extracellular vesicles on growth under chemical stress in Fusarium circinatum. The graphs show the growth kinetics of F. circinatum in response to various chemical agents, measured over a period of 168 hours. Each graph represents a different agent and compares growth rates between samples treated with biofilm-derived extracellular vesicles (bEVs, red lines) and those without bEVs (black lines).
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Figure S6: comparative analysis of Fusarium circinatum growth response to biofilm-derived extracellular vesicles (bEVs) under various chemical stress conditions. The graphs depict the response of F. circinatum to different chemical agents, monitored over a period of 168 hours. The graphs compare the OD490 nm for cultures treated with bEVs (red lines) versus those without bEV treatment (black lines).
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Figure S7: Assessment of chemical sensitivity in Fusarium circinatum treated with biofilm-derived extracellular vesicles (bEVs). The bar chart showcases the assimilation rates of F. circinatum under exposure to various chemical agents, measured at OD490 nm over a period of 168 hours.
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Extracellular vesicles modulate growth and stress adaptation in Fusarium circinatum

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