Biofilm characterisation of the maize rot-causing pathogen, Fusarium verticillioides

Biofilm characterisation of the maize rot-causing pathogen, Fusarium verticillioides | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL This is a preprint and has not been peer reviewed. Data may be preliminary. 27 January 2025 V1 Latest version Share on Biofilm characterisation of the maize rot-causing pathogen, Fusarium verticillioides Authors : Chizné Peremore , Cairin van ‘t Hof , Cebo-LeNkosi Nkosi , Kadima Tshiyoyo , Samkelo Malgas , Francinah Ratsoma , Wisely Kola , Quentin Santana , Brenda Wingfield , Emma Steenkamp T , and Thabiso Motaung 0000-0002-8813-7671 [email protected] Authors Info & Affiliations https://doi.org/10.22541/au.173798092.22701966/v1 Published Biofouling Version of record Peer review timeline 316 views 165 downloads Contents Abstract Introduction Fungal strains and maintenance Screening for biofilm formation Analyses of biofilm-like structures Analysis of biofilm-derived cells Quantification of biomass, EPS and metabolic activity The impact of abiotic conditions on biofilm growth Effect of DNase treatment on biofilm formation Statistical analysis Colony morphology of biofilm-like structures biofilm development biofilms and impact on cells therein 3.4. EPS/Biomass and metabolic activity as indicators of biofilm response 3.5. Biofilm formation in response to abiotic factors 3.6. The structural integrity role of extracellular DNA in biofilms Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Microorganisms often form biofilms—structured communities of microbial aggregates encased in self-produced extracellular polymeric substances (EPS). These biofilms enable adherence to surfaces and enhance microbial survival and interaction. Several plant-associated fungi, including Fusarium verticillioides, a mycotoxigenic fungus associated with maize, are known to create biofilms, necessitating research into their role in fungal persistence and disease development. This study aimed to investigate the biofilm formation capability of F. verticillioides under laboratory conditions. Our results indicated that stationary phase cultures developed a biofilm-like pellicle characterised by a cloudy, thin slime composed of hyphal aggregates. Microscopic analysis revealed a heterogeneous structure of dense, entangled hyphae alongside quantifiable EPS and extracellular DNA (eDNA) levels. The biofilms also exhibited responsiveness to factors such as pH and temperature, emphasizing their ecological relevance. Furthermore, we assessed the role of eDNA in maintaining biofilm structure through DNase treatment, which proved marginally effective in mature biofilms. This suggests complex interactions between eDNA and constituents in the EPS during maturation. The analysis of the Biofilm characterisation of the maize rot-causing pathogen, Fusarium verticillioides Chizné Peremore 1,2 , Cairin van ‘t Hof 1* , Cebo-LeNkosi Nkosi 1* , Kadima Tshiyoyo 1 , Samkelo Malgas 1 , Francinah Ratsoma 1,2 , Wisely Kola 1,2 , Quentin Santana 4 , Brenda Wingfield 1,2 , Emma T Steenkamp 1,2 , Thabiso E Motaung 1,2** 1 Division of Microbiology, Department of Biochemistry, Genetics and Microbiology, University of Pretoria, Private Bag X20, Hatfield Pretoria 0028, South Africa 2 Forestry and Agricultural Biotechnology Institute, University of Pretoria, Hatfield 0083, Pretoria, South Africa. 3 Biotechnology Platform, Agricultural Research Council, Onderstepoort, South Africa * Undergraduate students enrolled in the BSc Microbiology program at the University of Pretoria during the period of this study ** Correspondence: Email: [email protected] Thabiso E Motaung orcid.org/0000-0002-8813-7671 University of Pretoria Private Bag X20 Hatfield 0028 Pretoria South Africa Abstract Microorganisms, including fungal pathogens, often form biofilms—structured communities of microbial aggregates encased in self-produced extracellular polymeric substances (EPS). These biofilms enable adherence to surfaces and enhance microbial survival and interaction. Several plant-associated fungi, including Fusarium verticillioides , a mycotoxigenic fungus associated with maize, are known to create biofilms, necessitating research into their role in fungal persistence and disease development. This study aimed to investigate the biofilm formation capability of F. verticillioides under laboratory conditions. Our results indicated that stationary phase cultures developed a biofilm-like pellicle characterised by a cloudy, thin slime composed of hyphal aggregates. Microscopic analysis revealed a heterogeneous structure of dense, entangled hyphae alongside quantifiable levels of EPS and extracellular DNA (eDNA). The biofilms also exhibited responsiveness to factors such as pH and temperature, emphasizing their ecological relevance. Furthermore, we assessed the role of eDNA in maintaining biofilm structure through DNase treatment, which proved marginally effective in mature biofilms. This suggests complex interactions between eDNA and constituents in the EPS during maturation. The analysis of the EPS-extracted exopolysaccharide’s composition and structure was also analysed using nuclear magnetic resonance, ultraviolet spectrophotometry, viscometry, particle size analysis and Fourier transform infrared spectroscopy. The exopolysaccharide was found to have a hydrodynamic diameter of 4.19 nm and low viscosity (0.022 dl/g), and to be cationic (composed of amino sugars) and unordered, facilitating stability through complexation with the anionic eDNA. Based on these results, we propose that F. verticillioides forms a ‘true’ biofilm that may facilitate adaptation, survival, and persistence of this fungus in the field. Keywords: Biofilm, Fusarium verticillioides , extracellular polymeric substances, extracellular DNA, virulence factors Introduction After rice and wheat, maize ( Zea mays ) is the third most abundant grain crop, feeding millions of people, particularly in Sub-Saharan Africa. However, its growth yield is threatened by Fusarium verticillioides , which systemically colonises leaves, stems, roots, and kernels. The fungus can therefore induce serious damage that often manifests in Fusarium ear and stalk rot [1,2]. These diseases have food safety and security implications due to mycotoxin contamination that is associated with the fungus. Primarily, F. verticillioides secretes the mycotoxin, fumonisin B 1 , which contaminates symptomatic and asymptomatic maize kernels and stored grains [3]. The toxicity of this compound is due to the inhibition of ceramide synthase and subsequent toxic intracellular accumulation of sphingosine and other sphingoid bases [4], ultimately imposing detrimental health effects on consumer populations [5]. Indeed, studies from around the globe, including some from Africa, Asia, and Latin America, paint a disconcerting picture of how the prevalence of mycotoxins leads to a variety of human pathologies, including oesophageal and liver cancer in adults who consumed contaminated maize [5]. The mechanisms by which F. verticillioides invades maize have been outlined [1,2], providing important clues on the circumstances leading to infection symptoms and the precise anatomical locations of the maize plant that would probably harbour mycotoxins. The production of mycotoxins by Fusarium verticillioides has often been given far more attention than the production of virulence factors in many studies on mycotoxigenic fungi. The current study posits the development of biofilms is somehow closely related to the accumulation of mycotoxins in fungi. Tell-tale is the biofilm 3D structure and biomass, both of which are covered in extracellular polymeric substances (EPS). This structural design gives rise to emergent characteristics that are only seen in the biofilm mode of microbial life, such as surface adhesion, spatial organisation, physical and social interactions, chemical heterogeneity, and greater tolerance to antimicrobials [6]. In the case of mycotoxigenic fungi, the biofilm EPS may exert a substantial effect on mycotoxin production; it may permit mycotoxins to stably accumulate and persist for longer periods as it glues the cells together, in the process creating pockets and channels through which mycotoxins can be concentrated and distributed within a biofilm, respectively. Aspergillus fumigatus biofilms, for instance, augment the production of gliotoxin, a sulphur-containing mycotoxin with immunosuppressive properties [7]. An interesting area of research will be determining to what extent the components of a biofilm, including the EPS and its associated cell-free components such as the extracellular DNA (eDNA), influence mycotoxin production. Therefore, research on biofilms will provide a fresh perspective on mycotoxin synthesis in economically important fungi. As biofilms present a cross-sectoral challenge, affecting a wide range of sectors including healthcare, built environment, food and agriculture [8], our lack of understanding of how filamentous fungal biofilms originate, and how they adapt to their microenvironments once developed, will restrict our capacity to identify and counteract their detrimental impacts. This is apparent in the clinical setting, where most clinical infections associated with medical instruments, including indwelling devices (e.g., catheters, pacemakers, dentures, orthopedic prostheses, and heart valves) colonized by biofilms are difficult to treat due to antifungal resistance of these cell community structures [9–11] . Marine biofilms, on the other hand, which are created by both microorganisms and macroorganisms (such as algae), play a significant role in the environmental effects of biological fouling, also known as marine biofouling, which is the accumulation of undesirable biological matter on artificial submerged surfaces. Ships and underwater surfaces (such as undersea cables and acoustic instruments) are examples of colonized surfaces, and their treatment is difficult due to concurrent and intolerable environmental impacts on non-target species [12,13]. The same type of challenges might apply to agriculture, where some of the essential and most used tools and machinery are contaminated with harmful fungal biofilms that are challenging to remove. When employed across many fields, farming equipment colonized by biofilms has the potential to contaminate unaffected fields with biofilm-derived propagules that may have acquired novel traits within a biofilm, including resistance to fungicides. Somewhat formal descriptions of biofilms in plant fungal pathogens have started to emerge [14] , with the latest being provided for the Phaeomoniella chlamydospore [15], F. graminearum [16] and F. circinatum [17] . However, to date, the role of biofilm formation in many disease-causing plant fungi, including key biofilm components such as eDNA and EPS, has not been elucidated. The current study sought to elucidate how the morphology of surface-associated F. verticillioides might be included in current biofilm descriptions. Our research will contribute to a better understanding of how filamentous plant fungal pathogens coordinate survival by forming a community structure. Materials and Methods Fungal strains and maintenance Strains of F. verticillioides were obtained from a culture collection in the Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, following isolation from maize samples taken from fields in the Eastern Cape province of South Africa. Two strains (CMWF 1196, and CMWF 1197) were screened for their ability to form biofilms in this study. The strains were plated on ¼ strength PDA (Potato Dextrose Agar; 1% (w/v) PDA powder and 1.2% (w/v) Difco agar) and allowed to grow for two weeks at room temperature. The matured cultures were then used to develop biofilms in liquid media as described below. Screening for biofilm formation Rapid screening of biofilm formation was performed for all fungal isolates by cutting a block of agar (5 mm x 5 mm) from the sporulating culture and inoculating it into 15 ml of three different growth media in 50 ml falcon tubes; ¼ Potato Dextrose Broth (PDB), Roswell Park Memorial Institute-1640 broth (RPMI) and Sabouraud Dextrose Broth (SDB). After mixing, the inoculate solution was poured into petri dishes (100 mm 15 mm) and incubated at 25 and 30 °C for 24 – 72 hrs. The cultures were visually analysed every 24 hrs for biofilm formation which is a hyphal assemblage that appears in the form of a cloudy and thin slime material, and photographed with an Epson Perfection V700 Photo scanner. Biofilm formation for the remaining experiments was conducted using a cell counting method. To do this, the inoculum for counting cells was prepared by adding 2 ml of 1 phosphate-buffered saline (PBS, 137 mM NaCl, 2.7 mM KCl, 10 mM Na 2 HPO 4 , and 1.8 mM KH 2 PO 4 ) onto the sporulating culture of F. verticillioides that was grown on ¼ PDA for seven days. The plate was swirled to allow spores to be released into the PBS and then counted using a haemocytometer placed under a Zeiss Axioskop 2 plus Light Microscope. To distinguish between dead and living cells in the fungal culture, 40 μl of cell culture suspension was mixed with 40 μl of 0.4% (w/v) Trypan Blue solution (Sigma-Aldrich) into an Eppendorf tube. The concentration of the harvested conidia was adjusted to 1 x 10 5 conidia/ml in ¼ PDB. A volume of 200 μl of the adjusted harvested conidia culture was added to 48 wells plates (Corning® Costar® TC- Treated Multiple Well Plates from Sigma-Aldrich) or chamber slides (Lab- Tek® Chamber SlideTM System, 8 Well Permanox® Slide) and incubated for seven days at 25 °C . Once biofilms had matured, the culture was then analysed using different microscopic techniques as explained later. Analyses of biofilm-like structures To better describe the biofilms formed by F. verticillioides, colony and cell morphology were first analysed on plates containing liquid media (PDB) , where the former was examined visually every 24 hrs for 7 days. Following the visual screening, the spores were cultured in ¼ PDB with agitation (180 rpm) (Shake-O-Mat, LABOTEC) and stationary conditions for 72 hrs in chamber slides, harvested and then analysed under a light microscope (Zeiss Axioskop 2 plus Light Microscope (LM)). A scanning electron microscope (SEM) was used to study the ultrastructure of biofilms formed as previously described, in chamber slides or 48 well plates with glass coverslips (LASEC SA). Sample preparation for SEM was performed according to Harding et al. [18]. Samples were examined with Zeiss Ultra PLUS FEG SEM Confocal Laser Scanning Microscopy (CLSM). This was used to analyse of F. verticillioides biofilms by forming biofilms in chamber slides at different time points (24 and 7 days) and staining them for 30 mins in a dark room with 100 μl of FUN-1, which fluorescent viability probe for fungi. Biofilms were then visualized using Zeiss LM 880 CLSM with excitation wavelength at 488nm and emission at 650nm. Analysis of biofilm-derived cells This study considered whether biofilm-derived cells are different from planktonic cells in F. verticillioides. To do this, biofilms of F. verticillioides were formed as previously mentioned. After 7 days of incubation, cells were extracted from the biofilm by scraping and briefly agitated (30 s) to loosen the cells from the EPS matrix. This suspension was filtered through Mira cloth (Sigma-Aldrich) to separate the cells from the EPS material. In parallel, F. verticillioides planktonic cells were scraped from the sporulating cultures on ¼ PDA and added to PBS. The cells were then counted and adjusted to the desired concentration (1 x 10 5 spores/ml). A volume of 10 μl was aliquoted onto ¼ PDA and incubated at 25 °C for 7 days. The agar plates were then examined to study differences in growth and colony morphology, with three sections of each biofilm-derived and planktonic cells examined with Zeiss Ultra PLUS FEG SEM. Quantification of biomass, EPS and metabolic activity Biofilm biomass was determined by measuring the production of EPS and metabolic activity. To measure the biofilm biomass, crystal violet was used as it is known to be a good indicator of the amount of cellular biomass [19]. Biomass was then quantified according to a previously described method [20]. Biofilms in 96 well plates (Corning® Costar® TC- Treated Multiple Well Plates (Sigma-Aldrich) were fixed for 15 min in 200 μl of 99% methanol. The supernatant was then removed, and the biofilm that was attached in microtiter plates was air-dried for 5 min before adding 200 μl of 0.3% crystal violet solution (stock solution diluted in PBS; Sigma-Aldrich) to each well. The stained biofilms in microtiter plates were then incubated for 20 min at room temperature before being rinsed twice with PBS to remove excess dye. The biomass in each well was then decolourised with 200 μl of 99% ethanol for 5 min. A volume of 100 μl of this solution was then transferred to a new 96-well plate and the absorbance was measured at 590 nm using a microplate reader (SpectraMax ® Paradigm® Multi-Mode Detection Platform). The extracellular matrix was quantified according to the method described by Mello et al. [20]. Biofilms in 96 well plates (Corning® Costar® TC- Treated Multiple Well Plates (Sigma-Aldrich) were stained for 5 min at room temperature with 20 μl of 0.1% safranin (stock solution diluted in PBS: Sigma-Aldrich). After that, the stained biofilms were rinsed with PBS till the supernatants became transparent. With 200 μl of 99% ethanol, the extracellular matrix was decolorized. A volume of 10 μl of the supernatant was transferred to a fresh 96-well plate, and the absorbance at 530 nm was measured using a microplate reader as previously reported. To measure the metabolic activity of the biofilm, biofilms were formed in 96 well plates as before and incubated at 25 °C. Once biofilms had been formed, their metabolic activity was then quantified using a colorimetric assay, XTT (sodium 3 ̵́- [1- (phenylaminocarbonyl)- 3,4- tetrazolium]-bis (4- methoxy6-nitro) benzene sulfonic acid hydrate) (Sigma-Aldrich), according to the manufacturer’s recommended specifications. The activity of fungal mitochondrial dehydrogenase converts XTT tetrazolium salt to XTT formazan resulting in a colour change that can be measured using the microplate reader as previously described, with the absorbance from each well measured at 492 nm. The impact of abiotic conditions on biofilm growth The impact of abiotic conditions on biofilm production by F. verticillioides was assessed at different pH values (2, 3, 4, 5, 6, 7, 8) and temperatures (10, 15, 20, 25 and 35 °C) in ¼ PDB. The pH was adjusted using HCl and NaOH. To conduct these experiments, standardized spore suspensions were inoculated as previously described and biofilms were allowed to develop under the aforementioned abiotic conditions for 7 days. Quantification of biomass, EPS, and metabolic activity was then performed using the microplate reader as previously described. Following this, EPS and biomass were determined, with EPS expressed per biomass (i.e., EPS/Biomass) and metabolic activity percentages calculated to compare the response of biofilms under the different physical factors. Effect of DNase treatment on biofilm formation The eDNA release was measured using a microplate-based fluorescence assay using a DNA binding dye (SYBR® Green I), as previously described by Rajendran et al. (2014). For this study, biofilms were cultivated in ¼ PDB for 7 days at 25 °C before being scraped from the plates using sterile cell scrapers and rinsed with PBS. The EPS was extracted from the disaggregated biofilm using 0.2 M EDTA. Following this, the samples were centrifuged at 10,000 x g for 30 min, the EDTA supernatant was collected and filtered through a 0.45 m syringe filter (Millipore). SYBR® Green I (Invitrogen), whose binding results in fluorescence that is directly proportional to DNA content, was applied at a 1:4 ratio to biofilm supernatants in a black well microtiter plate (Costar3603; Corning). The levels of eDNA were then quantified using a fluorescence plate reader (SpectraMax ® Paradigm ® Multi-Mode Detection Platform) at 485 and 525 nm, respectively. The concentration of eDNA in the sample was quantified using the DNA standard curve as previously described by Leggate et al. [21]. The role of eDNA in F. verticillioides biofilm formation was investigated by depletion of eDNA within the biofilm using the hydrolytic enzyme [22], and DNase I from bovine pancreas (Sigma-Aldrich). The DNase I was prepared in 0.15 M NaCl supplemented with 5 mM of MgCl 2 . To assess the effect of DNase I on biofilm formation, biofilms were formed in ¼ PDB as described above and were incubated with DNase I at the concentration of 0.25, 1, and 2 mg/ml, and incubated at 25°C for 72 hrs and 7 days. Untreated controls were included for direct comparison. After each treatment, the biofilms were washed in PBS and their metabolic activity, biomass, and EPS were quantified as mentioned previously. Exopolysaccharide composition analysis Protein content determination The Bradford assay was employed to detect protein in the exopolysaccharide, as outlined previously, with bovine serum albumin (BSA) serving as a standard [23] Spectrophotometric measurements were conducted at room temperature and protein concentration was estimated in terms of BSA equivalents. Phenolic content determination The Folin-Coicalteu assay was employed to detect phenolics in the exopolysaccharide, as outlined previously, with gallic acid serving as a standard [24]. Spectrophotometric measurements were conducted immediately after the heating step at 40 °C and phenolic concentration was expressed gallic acid equivalents per dry mass of the exopolysaccharide. Total sugar content determination The monosaccharide composition of the exopolysaccharide was determined after hydrolysis of 10 mg dry powder with 1 ml of 2 M trifluoroacetic acid (TFA) in a dry digital bath (Eins Sci, Johannesburg, South Africa) for 2 h at 120 °C. The residual TFA was evaporated from the sample using nitrogen gas. The sample was resuspended in 1 ml of deionised water for sugar analysis. The sulphuric acid-UV assay was used to estimate the total sugar content of the exopolysaccharide, using N-acetyl-glucosamine as a suitable standard. The method was developed and optimised using a previously described method [25]. Spectrophotometric measurements at 290 nm were made at room temperature and the total sugar was estimated and expressed as total sugar per dry mass of the exopolysaccharide. UV spectroscopic determination of the degree of acetylation (DA) Ultraviolet (UV) spectroscopy was employed according to a previously reported method whereby 25 mg of the exopolysaccharide was dissolved in 5 ml of 85% phosphoric acid at 60 °C for 40 min. One milliliter of the resulting solution was diluted to 100 ml with distilled water and incubated at 60 °C for 2 h before measurements. The determination of the degree of N-acetylation was carried out by using the UV value at 203 nm after calibration with monomer mixtures of glucose and N-acetyl glucosamine prepared in 0.85% phosphoric acid at concentrations of 0, 10, 20, 30, 40, 50 and 100 µg/ml [26,27]. Sulphate content determination The sulphate content of the F. verticillioides exopolysaccharide was determined using the gelatine‑barium method developed as described previously [28]. Spectrophotometric measurements were conducted at room temperature and sulphate ion concentration was estimated in terms of Na 2 SO 4 equivalents. Conducting the barium-chloride assay on both non- and hydrolysed exopolysaccharides allowed for the determination and discrimination between organic and inorganic sulphates contained in the sample. Exopolysaccharide physical and structural analysis Functional group analysis by FTIR Exopolysaccharide functional group composition was investigated by FTIR using a Perkin Elmer Frontier spectrophotometer, equipped with a universal ATR diamond crystal sampling accessory (Waltham, United States). About 16 scans were collected over the 4000 – 650 cm -1 spectral range. Automatic baseline correction and normalisation of the spectra were conducted with Spectrum One software (Perkin Elmer). The degree of acetylation (DA) is the share of nitrogen sites occupied by acetyl groups (each nitrogen atom can react with one acetyl group). The DA of the exopolysaccharide was determined using the absorbance ratios of the amide II-band to the CH- stretching band as described previously [29]. Glycosidic linkage analysis by NMR The sugar sequence and types of glycosidic linkages constituting the exopolysaccharide were analysed by 1 H and 13 C nuclear magnetic resonance (NMR) spectroscopy using a Bruker Avance III 400 MHz spectrometer with a BBI probe (Bruker, Karlsruhe, Germany). For NMR analysis, the exopolysaccharide was suspended in D 2 O (99.96%) (Merck, Darmstadt, Germany) and the spectra were recorded at ambient temperature. The spectra were processed and analysed using TopSpin NMR software (Bruker, Karlsruhe, Germany). For structural elucidation of the exopolysaccharide, the NMR chemical shifts obtained from 1 H and 13 C spectra were used for input to the CASPER webserver [30,31]. Rheological and viscosity-average molecular mass analysis To determine the intrinsic viscosity and viscosity-average molecular weight (Mv) of the F. verticillioides exopolysaccharide, solutions were prepared by dissolving the polysaccharide at 0.1 – 20 mg/ml in a 50 mM NaCl solution. The intrinsic viscosity ([η]) of the exopolysaccharide was determined by the average efflux time of each sample measured at 25 °C using a semi-micro viscometer (size: 50) (CANNON Instrument Company, State College, United States). First, the specific viscosity (ηₛₚ) of the exopolysaccharide, which is calculated from the increment in viscosity due to the polymer, was calculated using Equation (1): \(\eta ₛₚ=\ \frac{tₐ-t₀}{t₀}\) (1) where tₐ is the efflux time of the solution and t₀ is the efflux time of the solvent. The intrinsic viscosity ([η]) is defined as specific viscosity extrapolated to an exopolysaccharide concentration (C) of zero as follows (Equation 2): \(\left[\eta\right]=(\frac{\eta ₛₚ}{C})_{C\rightarrow 0}\)(2) where C is in g/ml. The viscosity-average molecular weight (Mv) of the exopolysaccharide was calculated using the Mark-Houwink equation (Equation 3). The K (0.00016) and α (0.79) are constants for a given solute-solvent system and temperature (Chandrasekharan et al., 2019; Rinaudo, 2006). \(\left[\eta\right]=K\text{Mv}^{\alpha}\) (3) Particle size analysis by DLS The particle size of the exopolysaccharide was determined by dynamic light scattering (DLS) using a Genizer Dual-Light Nano Particle Sizer (Irvine, United States), with a standard green laser (30 mW and 570 nm). Scattering was analysed at 25°C using a 4 ml quartz cuvette, with samples diluted to 0.1 – 10 mg/ml concentrations in 50 mM NaCl. For all the samples, the mean value of three measurements was taken at a photon counting rate of around 40 for a green laser with a delay time of 5 µsec. The LPSA software was used to obtain the hydrodynamic diameter using the cumulant analysis with a repeatability of 5% and size distribution (polydispersity index, PdI) of the exopolysaccharide. Structural conformation determination by Congo red assay The conformational structure of the exopolysaccharide in an aqueous solution was determined by characterising the Congo red-polysaccharide complex as described previously [32]. Solutions of the exopolysaccharide (1 mg/ml), 80 μM Congo red and NaOH with different concentrations (0, 0.1, 0.2, 0.3, 0.4 and 0.5 M) were prepared. Meanwhile, water instead of the polysaccharide solution was used as the control. After being kept for 10 min at room temperature, λmax was measured at a wavelength range of 400 to 600 nm using a plate reader. Water solubility and conductivity determination The solubility of the exopolysaccharide was determined by dissolving 1 g of the polysaccharide in 100 ml of dH 2 O with constant stirring for 5 minutes at 25°C. Prepared suspensions were then centrifuged for 5 minutes at 12 000 g. The supernatant was removed, while the non-dispersible pellet of the exopolysaccharide was dried at 40°C overnight. The tubes were weighed, and the mass of the dispersible fraction was determined. To determine the conductivity, a 1% (w/v) exopolysaccharide solution was prepared in dH 2 O. The conductivity of the exopolysaccharide solution was determined using an XS Tester PC5 Tester (Carpi MO, Italy). The reference temperature for the experiment was kept at 25°C, and the results were expressed in µS/cm. Statistical analysis All experiments were performed in duplicate, in two independent experimental sets. The data were expressed as mean ± standard deviation (SD). The results were evaluated using the GraphPad Prism 9 computer program. A p -value of 0.05 or below was deemed statistically significant in all analyses (ns= p > 0.05; *= p ≤ 0.05; **= p ≤ 0.01; *** = p ≤ 0.001; ****= p ≤ 0.0001). In addition, a one-way ANOVA was used to determine significant differences between samples. Results Colony morphology of biofilm-like structures The ability of F. verticillioides to form biofilm-like structures was observed in both strains examined in this study (Figure 1). Then, using visual inspection of liquid cultures, colonies emerging from a biofilm culture were observed in PDB, SDB and RMPI, incubated at 25 °C from 24 hrs to 7 days (data not shown). The biofilm is normally distinguished from planktonic cells by its dense, highly hydrated clusters of cells enmeshed in a gelatinous matrix [33–35]. Indeed, F. verticillioides biofilm-like colonies displayed a dense, thin, and cloudy material (Figure 1). Based on these morphological traits, the biofilm-like formations will be referred to as simply biofilms from this point on. biofilm development Since, morphologically, both the strains appeared to be forming similar biofilms, only CMW 1196 was then selected and utilized for subsequent studies. It was anticipated that a biofilm would form most effectively under stationary conditions and assumed that the shear stress from shaking would prevent the formation of the EPS matrix, a distinguishing feature of microbial biofilms. Therefore, the cells from F. verticillioides CMW 1196 were cultured under both shaking and stationary conditions. Following this, it was found that planktonic cells incubated under shaking conditions did not typically clump together when observed under a light microscope (Figure 2A), but those incubated under stationary conditions developed a community of cells resembling a biofilm (Figure 2B). When these cells were analysed under SEM, little to no EPS formation around cells incubated under shaking conditions was observed (Figure 2C). However, cells that were incubated without shaking formed a visible EPS (Figure 2D), which was not surprising given that EPS has been observed often after growth under non-shaking conditions in biofilm investigation studies [36]. biofilms and impact on cells therein The development of a biofilm in F. verticillioides may influence the metabolic status of cells and by extension, their phenotype [37]. Our observations were in agreement with this, as the spores at the dispersion stage (Figure 3D) appeared to be morphologically distinct from the normal microconidia spores initially used as inoculum to initiate a biofilm (indicated in Figure 3A). Usually, microconidia of F. verticillioides are club- or elliptical-shaped or pointed at both ends (Figure 3A). However, the biofilm-derived spores were more globose/lemon-shaped and slightly larger than typical conidial cells (Figure 3D) . Therefore, biofilm-derived cells, as indicated in Figure 3D, may influence phenotypic diversity in F. verticillioides . For instance, when these cells were harvested from a biofilm and plated on ¼ strength PDA , they displayed a colony morphology that is different from cells not derived from a biofilm i.e., they formed a colony smaller than that of their planktonic counterpart (Figure 3). The physiological responses of cells within a biofilm are likely shaped by the dynamics of the biofilm ecosystem. However, the morphology of cells derived from a biofilm had no apparent differences when compared to the morphology of normal cells (Supplementary Figure 2), suggesting that the differences between these cells might largely be in their response to environmental signals, as in the case with observations in Figure 3, as opposed to their morphology. 3.4. EPS/Biomass and metabolic activity as indicators of biofilm response The complexity of the biofilm is for the most part brought about by the release of EPS [38]. This means that the biofilm may possess the ability to affect the physiology of the cells within it by virtue of holding them in place, thus maintaining the biofilm’s 3D structure while also optimising the exchange of nutrients and genetic material. As previously indicated, the results showed that unlike cells cultured under shaking conditions (Figure 2A, C), the generation of a visible EPS occurs concurrently with the establishment of a mature biofilm (Figure 2B, D). Since establishing that F. verticillioides biofilms produce EPS, in this study, we were also interested in how much of this material was being produced during biofilm formation and to what extent metabolically active cells contributed to the total biofilm biomass. For this reason, colourimetric assays were used, namely crystal violet and XTT, to analyse the biomass and EPS by the absorption of safranin and to evaluate the metabolic activity (cell viability) of the biofilm, respectively. In the case of the XTT reduction assay, the production of soluble coloured formazan salts by sessile cells is a direct reflection of cellular metabolic activity. According to the results, an increase in cell mass and EPS production (Figure 4A) was accompanied by an increase in metabolic activity of the cells inside the biofilm (Figure 4B); EPS/biomass increased significantly from 13% (3 days) to 45% (7 days) ( p = 0,002), suggesting the more the biofilm matures the more EPS is produced. 3.5. Biofilm formation in response to abiotic factors Having shown that F. verticillioides asexual cells could develop into a biofilm, this study then investigated how the biofilm reacts to different environmental conditions, namely different temperature and pH conditions. As shown in Figure 5A, EPS/biomass was highest at pH 5, suggesting that F. verticillioides may prefer pH 5 for biofilm formation. The media that was used in the initial experiments (¼ PDB) has a pH of around 5, and it was in this medium that all the stages of a biofilm were observed (Figure 2). The EPS and metabolic activity were essentially the same at acidic pH levels (i.e., 2, 3, and 4, below the optimal pH of 5), whereas at pH levels higher than the optimal (i.e., pH 6, 7, and 8), the EPS was produced at lower levels and plateaued at these levels. From the optimal pH (pH 5) point of view, F. verticillioides biofilms seem to produce significantly more EPS/Biomass than at pH 6 and pH 8 ( p ≤ 0.05). However, this biofilm had a significantly lower metabolic activity at a range of pHs from 2-8 (2, 3, 4, 6, 7, and 8) ( p ≤ 0.001). This suggests that the biofilm response to pH is versatile, which could influence the adaptability of this pathogen to a range of field conditions. Figure 5C and D depict the influence of temperature on the production of F. verticillioides biofilms. The best temperature tested for metabolic activity is 25 °C, and the fungus displayed similar metabolic activities and formed robust biofilms at 20 and 25 °C with no significant differences . The fungus also formed biofilms at 10, 15 and 35 °C, but these were not as robust as 20 and 25 °C and their cells had lower metabolic activity. Different temperatures do not seem to significantly affect the EPS/biomass yield except for 10 °C where the biomass was undetectable. 3.6. The structural integrity role of extracellular DNA in biofilms The biofilm was treated with DNase I to elucidate the role of eDNA in maintaining the structure of the biofilm. The DNAse abolished biofilm formation during the early stages of development, i.e., at 72 hrs (Figure 6A). Also, the structural integrity of the biofilm was revealed to be significantly impacted by the addition of DNase I in a concentration-dependent manner. Unfortunately, the EPS/biomass yield could not be determined during the early biofilm maturation phase (72 hrs) as the biofilm was had no substantial formation integrity to perform the relevant assays. In comparison to the biofilm growth control at 7 days, the application of 0.25 and 1 mg/ml DNase I slightly reduced EPS/biomass formation by 17% ( p > 0.05) while 2 mg/ml of DNase I significantly reduced it by 40% ( p = 0.0095) (Figure 6B). Interestingly, while DNase I influenced the formation of EPS and biomass, it did not significantly affect the metabolic activity of the biofilm (data not shown). This suggests that eDNA plays a more crucial role in the structure of EPS in the biofilm of F. verticillioides rather than in its cellular activity, a finding that has also been demonstrated in other fungal species. Exopolysaccharide composition analysis The general compositional analysis of the F. verticillioides exopolysaccharide was determined after freeze-drying and the constituents of dry matter weight are reported (Table 8). Overall, the composition analysis showed that the exopolysaccharide was exclusively composed of carbohydrates, with minor quantities of phenolic content (5.33% on a dry mass basis). Exopolysaccharide functional group and structural monomeric units’ analysis The FTIR spectrum of the exopolysaccharide displayed the characteristic absorption peaks of functional peaks found in polysaccharides (Figure 7A). The exopolysaccharide displayed a large and strong absorption peak at 3380 cm -1 , attributed to the vibration of -OH stretching [39]. Its appearance at such a high frequency reflects increased hydrogen bonding interactions. The pronounced absorption peak at 3012 cm -1 may be due to the stretching vibrations of C–H bonds in CH 3 , CH 2 and CH [40]. The peak displayed around 1615 cm -1 , with a shoulder at 1590 cm -1 , indicates the N-H bending vibration (amide II) [41] and the amide III band at 1327 cm -1 was also detected [29]. The splitting of the amide I band is due to the influence of hydrogen bonding or the presence of an enol form of the amide moiety usually observed in α-chitinous polysaccharides. Finally, additional peaks were also observed at 1395 cm -1 (asymmetrical deformation of CH 2 ) [41], 1213 cm -1 (symmetric stretching vibration due to S=O group, indicating the presence of sulfate group) [42] and 1096 cm -1 (pyranose residue) [39], and 921 cm -1 (C-H bending or C-O stretching vibrations) [40]. The anomeric configuration of the monosaccharides in the F. verticillioides exopolysaccharide was determined using 1 H (Figure 7B) and 13 C NMR (Figure 7C) spectroscopy. The faint doublet signal at δ1.34 and δ1.36 was attributed to H6 of the methyl group in 2-acetamido-2-deoxy-L-quinovose (Qui3N) [43,44]. The signals at δ3.14 and δ3.34 in 1 H NMR were assigned to H2 of a hexosamine [45] Similarly, glucosamine (GlcN) residues in chitosan exhibit a signal at δ3.15 [46]. The signal at δ3.82 was attributed to H6 of the hexosamine Qui3N residue in the exopolysaccharide [43]. Since there was no signal at δ2.0 – 2.1, it was concluded that most of the hexosamine residues in the exopolysaccharide did not contain an acetyl-H [31,46]. Still, it is noteworthy that the exopolysaccharide exhibited a signal at δ1.79 that could be attributed to an acetyl-H. In the F. verticillioides exopolysaccharide spectra, a signal at δ5.3 could match with an α-anomer present at the reducing end of the polysaccharide and no β-anomer-associated signal (δ4.7) was present [47]. A faint amide proton signal at δ8.33 was observed in the 1 H NMR spectra of the exopolysaccharide. The signals at δ51.1 and δ56.3 in the 13C NMR spectrum were assigned to a carbon (C2) substituted by the amino group in monosaccharide residues. These signals were similar to the two nitrogen-bearing carbons at δ49.15 and δ55.09 (GalN C-2 and Qui3N C-3, respectively) reported for the lipopolysaccharide from Aeromonas veronii Strain Bs19, Serotype O16 [44,45]. The 13 C NMR spectrum of the exopolysaccharide lacked a signal at δ21 – 25, where the -CH 3 of acetamide-containing hexosamine residues is displayed [31], concurring with the 1 H NMR spectrum regarding a majority of the hexosamines in the exopolysaccharide structure containing an amine (-NH 2 ) at this position. Finally, the signal at δ171 was attributed to carbonyl carbon atoms, indicating some degree of acetylation in the exopolysaccharide [48]. Based on all the data obtained from FTIR and NMR, it was concluded that the biological repeating unit of the exopolysaccharide from F. verticillioides had the following structure: α-d-QuiN (1→4) α-d-GalNAc (1→4) α-d-GalN Physical properties determination of the F. verticillioides exopolysaccharide The physico-chemical properties of the F. verticillioides exopolysaccharide, such as its intrinsic viscosity, viscosity-average molecular weight, hydrodynamic diameter, solubility and conductivity, were determined (Table 3). The exopolysaccharide had a low molecular mass of 0.505 kDa. The size distribution analysis suggested that the average particle diameter of the exopolysaccharide was 4.19 nm with a d90 (the particle size with 90% of particles ≤ d90) of 5.26 nm (Figure 8). The polydispersity or distribution (PdI) of the exopolysaccharide was 0.18, showing relative homogeneity of its size. Conformation of the F. verticillioides exopolysaccharide structure The conformation of polysaccharides in solution can be analysed by observing the shift in the maximum absorption wavelength (λmax) of Congo red dye in the presence of varying concentrations of NaOH. Triple-helix polysaccharides show a bathochromic shift in their λmax when they form a complex with Congo red [32] . Therefore, the Congo red assay was used to determine the structural conformation of the F. verticillioides exopolysaccharide (Figure 9A). At 0.0 – 0.20 M NaOH, λmax for Congo red alone and the exopolysaccharide-Congo red complex exhibited a sharp hypsochromic shift and tended to be stable at higher NaOH concentrations. These results indicated that the exopolysaccharide does not have a triple-helix conformation, but rather has a random coil conformation since triple-helix transitions (>15 nm) were not observed for the exopolysaccharide-Congo red complex compared to Congo red alone [32,49]. The UV-Vis absorption spectrum of the exopolysaccharide was also evaluated and displayed maximum absorption in the 205 – 215 nm wavelength range, which often results from n−σ* and π−π* transitions, which are found in many functional groups, such as amine, carboxyl, carbonyl and ester [50], in polysaccharides (Figure 9B). Similarly, the amino sugar, acetyl-glucosamine, constituting chitinous polysaccharides has been reported to absorb in the range of 190 – 220 nm [27]. No absorption peak was observed in 260 – 280 nm in the spectrum of the exopolysaccharide (Figure 11). This indicated the absence of nucleic acids or protein content in the extracted polysaccharide, consistent with the results in Table 1 and Table 2. Discussion Members of the genus Fusarium cause economically burdensome and hard-to-control diseases, including cankers, crown rot, head blight, scabs, and wilts. Many of these diseases are strongly linked to biofilm formation [14,18,51], as microbes primarily exist in a biofilm state in their natural environments. However, biofilm formation has been formally described in a few Fusarium species including F. oxysporum f. sp. cucumerinum and F. graminearum [16,52]. Therefore, for many fungal pathogens of plants, including those belonging to Fusarium , it is unclear how biofilms form, let along how they impact infections and disease outcomes. Ten years ago, Miguel and his colleagues observed what looked like F. verticillioides biofilms, in which the mycelium was structured in an extracellular material around the hyphae [53]. These researchers also discovered a flocculating substance over the cells or small fibrils of hyphae connecting to one another, like a biofilm. To the best of our knowledge, this is the first-time evidence of a biofilm-like structure for F. verticillioides has been reported in vitro . In the current study, we addressed this knowledge deficit by describing how F. verticillioides forms biofilms under in vitro conditions . Information that describes biofilm colony morphologies in stationary liquid cultures is generally lacking in fungal plant pathogen biofilm studies. Filamentous fungi including F. graminearum and F. circinatum have recently been described as being able to form biofilm colonies at the air/liquid interface that grow as pellicle–floating masses of cells that cling to each other and move as a unit [16,17]. In this study, we observed floating masses for F. verticillioides that are distinguishable from free-living (planktonic) cells by forming colonies displaying a dense, thin, and cloudy material. These results are consistent with several other studies also reporting similar features of fungal and bacterial biofilms [33,34,54,55]. It was also observed that biofilms in F. verticillioides also developed most efficiently under stationary conditions, while shear stress from shaking conditions prevented proper biofilm formation. Cells incubated in the stationary conditions without shaking had hyphal cells tangled with EPS that appeared to behave like a matrix binding the hyphae together, and cells cultured under agitated conditions seldom clumped together or not at all. The findings are analogous to those reported by Hawser et al. [56], who demonstrated that, under shaking conditions, only a limited number of cells are observable on the surface of cultures. In contrast, cultures maintained under stationary conditions are characterized by the presence of dense networks of hyphae. This might be due to the severe shaking influencing cell architecture, matrix deposition, and biofilm formation [57]. In C. albicans , shaking at a speed of 60 rpm prevents biofilm growth, with biofilms exposed to shear stress being thinner than those exposed to non-shaking conditions[36]. Hawker et al. [56] also found that lower agitation speeds result in the production of biofilms with no extracellular matrix but only hyphae, while shaking at higher-speed results in a biofilm which consists of a few cells on the surface. In this study, it was observed that some EPS material is present in planktonic cultures; however, this was not as abundant as in biofilm cells, suggesting that under conditions causing agitation of the fungal cells, the cells struggle to produce the EPS matrix. In light of the above, our findings align with the biofilm concept previously proposed [14,58,59]. The biofilms of F. verticillioides seem to develop through spore adhesion, microcolony formation, maturation, and dispersion. Although many fungal and bacterial species have recorded comparable developmental stages for biofilm formation, filamentous fungal biofilm formation seems to differ from strain to strain [37,38,52,60]. In contrast to unicellular life forms such as yeast and bacteria, most fungi contain many planktonic forms that can disperse and continue the cycle (e.g., sporangia asexual spores, sexual spores, and hyphal fragments), and these dispersive forms most usually float in the air rather than water [59]. The dispersal phase of biofilms leads to a substantial number of free-living cells in the form of conidia, but in F. verticillioides biofilms these cells appeared to be morphologically distinct from normal microconidial spores initially used as inoculum to initiate biofilm formation (Supplementary Figure 1). This suggests that the dispersed biofilm cells differ from normal microconidia. Similar findings were reported in a study on Bacillus cereus where the cells in the biofilm have different cell-surface characteristics than their planktonic counterparts. For example, the structure of a polysaccharide linked to peptidoglycan in B. cereus could change during biofilm development (Majed et al., 2016). Additionally, Boles et al. [61] showed that the short-term development of P. aeruginosa in biofilms causes considerable genetic diversity in the resident bacteria. The researchers discovered that genetic diversity creates specialized bacterial subpopulations in biofilms, enhancing their ability to withstand physiological stress. However, similar diversity in fungal biofilms has not been observed and needs further study. The production of the matrix has a high energetic cost, which may be evolutionary justified given the matrix’s structural and physicochemical significance in the growth and operation of the biofilm, without which the beneficial emergent properties of biofilms would not be possible [62]. This goes to show that the formation of biofilms is closely linked to the formation of the matrix, the bulk of which is extracellular material [63,64]. As biofilms grow, metabolic activity and EPS production also increase, suggesting that they may become more resistant to abiotic stress [14] . In the current study, it was found that F. verticillioides developed biofilms under a range of pH and temperature conditions (optimum pH and temperature of 5 and 25 °C, respectively), and a similar trend was previously reported [17]. Under field conditions, it has been reported that the optimum temperature for spore development is 27 °C, a temperature at which biofilm development could occur. Evaluation of biofilm formation under different pH conditions showed that biofilms were much more robust in somewhat acidic (pH 4 – 5). These findings are consistent with other reports [65] since the pH range for fungal development is fairly broad, ranging from pH 3 to more than pH 8, with the optimum at pH 5.0 assuming nutritional needs are met. The capacity to develop a biofilm under varying physical conditions may provide the fungus with survival benefits in inhospitable environments. Unlike other components of the biofilm EPS matrix, the eDNA has attracted the most attention and is considered an interesting component in the study of biofilms. Many studies have uncovered that eDNA plays many important roles in bacteria such as in biofilm structural maintenance, assisted by the action of DNA binding proteins [66–68], antimicrobial resistance [22,69], and acting as a reservoir for the interexchange of genes through natural transformation [70]. Furthermore, eDNA assumes more unusual roles, including acting as a source of energy and nutrients (e.g., carbon, nitrogen and phosphorus) [71–73], and forming higher-order conformations (e.g., G-quadruplex DNA) that further strengthen the biofilm through extracellular EPS-eDNA networks [74]. Given these roles, eDNA is an attractive target for antimicrobial drugs to manage biofilm-related infections. However, only a few studies have explored the existence and function of eDNA in filamentous fungi, with no studies conducted in plant pathogenic fungi. In the current study, it was demonstrated that DNase I has an impact on biofilm stability. In a similar study using A. fumigatus, DNase I was effective at all concentrations (0.25, 1, and 4 mg/ml) [22,75], but the maximal effect was observed with 4 mg/ml of DNase I ( p value < 0.001) which is similar to the observations in this study. The discovery that eDNA contributes significantly to the biofilm EPS in both bacteria and fungi suggests that this may be a conserved and possibly active microbial biofilm process. Nucleic acids are incorporated early in the development of the EPS matrix in F. graminearum , and they similarly appear to function as a scaffold, likely regulating the entire matrix structure of the biofilm [16]. The findings presented in the current study imply that eDNA plays a significant structural maintenance role during biofilm development in a filamentous plant fungal pathogen and may contribute to the severity of the disease. The F. verticillioides exopolysaccharide had a low molecular mass (0.505 kDa) compared to exopolysaccharides from other fungal species, such as that from F. solani SD5 (187 kDa) [76] and Cryptococcus laurentii AL 100 (4.2 kDa) [77]. It is worth noting that a majority of exopolysaccharide molecular masses reported in the literature were reported as weight average molecular mass, as reported by size exclusion chromatography (SEC) [78] . It is worth noting that SEC excels in providing detailed size distributions and absolute molecular weights but requires careful calibration, while viscometry offers a quicker and simpler approach but relies on empirical relationships that may introduce uncertainty. Lastly, weight-average molecular weight (Mw), as determined by SEC, emphasizes larger molecules and their contributions to physical properties, while Mv focuses on how polymers behave in solution regarding viscosity. Therefore, Mv may not emphasize larger molecules as strongly, leading to molecular weight values lower than those reported by Mw. The presence of –NH 2 and OH groups in the amino sugars composing the exopolysaccharide appear to be important factors that influence the solubility and conductivity of the exopolysaccharide. It is known that the presence of acetyl groups in amino sugars constituting polysaccharides such as chitin increases intermolecular hydrogen bonding, which stabilizes the structure and prevents water from penetrating and dissolving it, therefore, the protonation of these amino groups enhances the F. verticillioides exopolysaccharide’s solubility in aqueous solutions, particularly in acidic environments. The particle diameter of the exopolysaccharide isolated from F. verticillioides was significantly larger than that isolated from F. solani SD5, which was reported to exhibit a diameter of approximately 1 nm using transmission electron microscopy (TEM) [78]. Considering that F.verticillioides exhibited a significantly lower molecular mass than that from F. solani SD5 (0.505 versus 187 kDa), we suppose that the F. verticillioides exopolysaccharide may exhibit increased entanglement among polysaccharide chains in solution. This entanglement may have affected the viscosity and flow properties of the solution, further influencing how the exopolysaccharides arrange themselves, leading to an underestimation of their size. The cationic nature of the F. verticillioides exopolysaccharide may offer a survival advantage to the fungus as it can likely bind to eDNA and this interaction has the potential to impact the virulence of the fungus and protect eDNA within the extracellular matrix from digestion. To support this, the cationic Pseudomonas aeruginosa Pel exopolysaccharide was shown to bind to eDNA and it was postulated that this interaction likely impacts current therapies by increasing antimicrobial tolerance and protecting eDNA from digestion [79]. The production of exopolysaccharides containing α-1,4–linked acetyl-galactosamine (GalNAc) and GalN have also been confirmed in both Aspergillus and non- Aspergillus spp., including Neurospora crassa , Penicillium frequentans , Paecilomyces sp., and Trichosporon asahi , wherein they have been linked to adherence to surfaces or flocculation [80]. The F. verticillioides exopolysaccharide characterized in this study was highly de-N-acetylated. De-N-acetylation of biofilm exopolysaccharide has been linked to cell aggregation, surface attachment, exopolysaccharide secretion, and biofilm maturation depending on the organism [80]. Conclusions In the natural environment, plant stem and leaf surfaces can be sparsely or densely colonized by diverse fungal biofilms and are likely more complex than conditions used in this study. Laboratory-grown biofilms are a simple surface-covering, frequently exhibiting confluent and compact uniformity that is consistent with the original definition of biofilms [14,59]. Documenting biofilm formation in the natural environment by analysing heavily infected plant tissues and a population of field strains, as opposed to a few ones, is needed to better understand what is happening under field conditions. This would enable the development of novel antibiofilm drugs and treatment alternatives to decrease the prevalence of fungal infections. This study thus establishes a baseline with regards to F. verticillioides biofilms, showing its intricate structure and response to the environment. The anionic exopolysaccharide of this fungus may aid survival by binding to eDNA. This polymer seems to be essential for biofilm integrity, consistent with analyses conducted in human fungal pathogens. Taken together, F. verticillioides ability to form biofilms may give it an ecological edge in its battle to keep its place as a commensal and pathogen of maize. The biofilm might enable this fungus to evade host immunity, withstand antifungal treatment and competition from other microbes. The current study, together with earlier studies, therefore, will deepen the understanding of the relationship between disease outcomes and biotic interactions in F. verticillioides . Declaration of competing interest The authors affirm that they have no known financial or personal interests. Acknowledgements The work was funded by the The South African National Department of Science and Innovation-NRF under the Thuthuka funding instrument (Grant no. 129580) and the Centres of Excellence programme and South African Research Chairs Initiative (Grant No. 98353). 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Nature Communications, 2020, 11 , 2450. Figures Figure 1: Fusarium verticillioides strains that formed biofilm-like cultures in Petri-dishes containing ¼ strength Potato Dextrose Broth. The strains were left to grow, without shaking for 7 days at 25 °C. Figure 2 : Fusarium verticillioides cultured for 7 days under shaking conditions ( A, C ), remained in the planktonic (free-living) state, and without shaking (B, D), formed biofilms with observable extracellular polymeric substances (EPS, indicated by red arrows in D ). Figure 3: Biofilm-derived and planktonic cultured Fusarium verticillioides in chamber slides for (A) 24 hrs and (C) 7 days . The cells were stained with FUN-1 to determine biofilm developments. F. verticillioides cells from shaking (B) and non-shaking (D) cultures established colonies on ¼PDA. Colonies derived from biofilm cells of asexual cells that were formed were significantly smaller (**= p ≤ 0.01) than those derived from planktonic cells (E). This is despite the fact that the respective cells were plated at the same concentrations and incubated under the same conditions . Each dot on the bar graphs represents an independent biological replicate. ns= p > 0.05; *= p ≤ 0.05; **= p ≤ 0.01; *** = p ≤ 0.001; ****= p ≤ 0.0001 Figure 4 : Fusarium verticillioides biofilm formation was assessed based on (A) metabolic activity, evaluated by XTT reduction assay and (B) biomass and extracellular polymeric substances (expressed as EPS/Biomass) evaluated using crystal violet (OD 590nm ) and safranin (OD 530nm ), respectively ( p -value=0.0013). An increase in cell mass and EPS production is observed. Each dot on the bar graphs represents an independent biological replicate. ns= p > 0.05; *= p ≤ 0.05; **= p ≤ 0.01; *** = p ≤ 0.001; ****= p ≤ 0.0001 Figure 5 : Fusarium verticillioides biofilm formation assessed at various pH and temperature conditions by measuring (A, C) metabolic activity (XTT reduction assay (OD 475nm; p value<0.05) (expressed as metabolic activity percentage)), and (B, D) biomass and extracellular polymeric substances (expressed as EPS/Biomass percentage), evaluated using crystal violet (OD 590nm ) and safranin (OD 530nm ), respectively ( p value<0.05). This data shows that pH 5 permits better biofilm formation, and the biofilm response to pH is versatile, spanning a range of pH conditions. The optimum temperatures evaluated for metabolic activity were 20 and 25°C. It also generated biofilms at 10, 15, and 35°C, but these were less robust and had lower metabolic activity. Different temperatures do not appear to have a significant impact on the EPS/Biomass %, except for 10°C, when the biomass could not be quantified. Each dot on the bar graphs represents an independent biological replicate. ns= p > 0.05; *= p ≤ 0.05; **= p ≤ 0.01; *** = p ≤ 0.001; ****= p ≤ 0.0001 Figure 6 : Fusarium verticillioides biofilm response to DNase treatment. (A) The response of a 72hr-old biofilm to DNase at different concentrations (0.25, 1 and 2 mg/ml). (B) The response of biofilms measured in biomass and extracellular polymeric substances (expressed as EPS/Biomass percentage). This data shows that DNase I caused the collapse of biofilm formation at the early stages of growth, i.e., at 72 hrs, and the structural integrity of the biofilm was found to be strongly influenced by DNase I in a concentration-dependent manner. Each dot on the bar graphs represents an independent biological replicate. ns= p > 0.05; *= p ≤ 0.05; **= p ≤ 0.01; *** = p ≤ 0.001; ****= p ≤ 0.0001 Figure 7: Determination of functional groups constituting the F. verticillioides exopolysaccharide by FTIR. Figure 8: Determination of sugar sequence and types of glycosidic linkages constituting the F. verticillioides exopolysaccharide by (A) 1 H NMR and (B) 13 C NMR. Figure 9: Particle size distribution of the F. verticillioides exopolysaccharide in 50 mM saline at 25 °C. Figure 10: Maximum absorption wavelengths (λmax) of Congo red solution and Congo red‑exopolysaccharide complex at different NaOH concentrations. Figure 11: UV spectrum of the F. verticillioides exopolysaccharide. Table 1 . Composition analysis of the F. verticillioides exopolysaccharide, where the constituents are represented as a percentage on a dry mass basis. 93.8±0.6 Nd 5.33±0.01 Nd 3.4±0.0 Where Nd = not detected. Table 2. 1 H and 13 C NMR chemical shifts (ppm) of the F. verticillioides exopolysaccharide. →4) α-d-GalN 5.29 - 4.17 51.1 4.01 - 4.17 - 4.15 - 3.82 - - - - - →4) α-d-GalNAc 5.11 - 4.15 51.1 4.01 - 4.51 – 4.53 - - - - - 1.79 - - 171 α-d-QuiN - - 3.82 56.3 3.69 - 3.34 - 4.15 - 1.34 – 1.36 - - - - - Where - = corresponding signal not detected. Table 3. Physico-chemical characteristics of the F. verticillioides exopolysaccharide. 0.022 0.505 4.19 100% 3.58±0.40 Information & Authors Information Version history V1 Version 1 27 January 2025 Peer review timeline Published Biofouling Version of Record 9 Jun 2025 Published Copyright This work is licensed under a Non Exclusive No Reuse License. Keywords fusarium verticillioides biofilm extracellular dna extracellular polymeric substances virulence factors Authors Affiliations Chizné Peremore University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Cairin van ‘t Hof University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Cebo-LeNkosi Nkosi University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Kadima Tshiyoyo University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Samkelo Malgas University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Francinah Ratsoma University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Wisely Kola University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Quentin Santana Agricultural Research Council Onderstepoort Veterinary Research View all articles by this author Brenda Wingfield University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Emma Steenkamp T University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Thabiso Motaung 0000-0002-8813-7671 [email protected] University of Pretoria Department of Biochemistry Genetics and Microbiology View all articles by this author Metrics & Citations Metrics Article Usage 316 views 165 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Chizné Peremore, Cairin van ‘t Hof, Cebo-LeNkosi Nkosi, et al. 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