Mutually bene cial FAB consortia fortify stress resistance of Euglena mutabilis: evidence from sequencing, antibiotics, and Cd challenges Emma Kaszecki Trent University Daniel Palberg Trent University Mikaela Grant Trent University Sarah Gri n Trent University Chetan Dhanjal Carnegie Mellon University Michael Capperauld Trent University R. J. Neil Emery Trent University Barry J. Saville (  barrysaville@trentu.ca ) Trent University Research Article Keywords: Bioremediation, biotechnology, heavy metals, algal symbiosis, co-culture, fungal-algal-bacterial (FAB) Posted Date: October 16th, 2023 DOI: https://doi.org/10.21203/rs.3.rs-3428948/v1 License:   This work is licensed under a Creative Commons Attribution 4.0 International License. Read Full License Additional Declarations: No competing interests reported. Page 1/30 Abstract Background Synthetic algal-fungal and algal-bacterial cultures have been investigated for technological applications because the microbe interactions enhance growth and improve stress tolerance of the co-cultures. Yet these studies often disregarded natural consortia due to the complexity of environmental samples. The protist Euglena mutabilis is found in association with other microbes in acidic environments with high heavy metal (HM) concentrations. This may suggest that microbial interactions are essential for the alga’s ability to tolerate these extreme environments. Our study assessed the Cd tolerance of a natural fungal-algal-bacterial (FAB) association where the algae is replaced by the photosynthetic protist E. mutabilis. Results This study provides the rst assessment of antimycotic and antibiotic agents on E. mutabilis. Our results indicate that suppression of associated fungal and bacterial partners signi cantly decreases the number of viable E. mutabilis cells upon Cd exposure. However, axenic Euglena gracilis recovered and grew well following antibiotic treatments. Interestingly, both Euglena species displayed increased chlorophyll production upon Cd exposure. Finally, the constituent organisms in the E. mutabilis FAB consortia were identi ed using PacBio sequencing to be a Talaromyces sp and Acidiphilium acidophilum. Conclusion This study uncovers a possible tripartite symbiotic relationship, a FAB consortia, that withstands exposure to high concentrations of HM. This unique fungus, bacterium, and E. mutabilis interaction strengthens the photobiont’s resistance to Cd and provides a model for the types of FAB interactions that could be used to create a self-sustaining bioremediation technology. Background The association of algae with fungi is perhaps best known from lichen symbioses, but these alliances occur in a wider range of systems in which bacteria and fungi exchange key metabolites with algae [1, 2, 3], or strengthen their stress responses [4, 5]. Algae have been co-cultured with bacteria or fungi to enhance levels of algal biomass [6, 7], the production of high-value products [8, 9, 10], or the tolerance to extreme environments [11]. The natural association between the euglenoid Euglena mutabilis and a fungus was proposed to occur in acid environments with high levels of heavy metals [11]. In the present investigation, potential associations between E. mutabilis and a fungus as well as bacteria are investigated to assess the contributions of the co-cultured species to cadmium tolerance of the Euglena. Page 2/30 Euglenoids are a group of unicellular agellates with a diverse ecological distribution [12, 13]. They ll similar ecological niches as algae and have been investigated for applications in pharmaceuticals, cosmetics, biofuels, bioremediation, and foodstuffs [10, 14–17]. Euglena gracilis is the model organism for this group as it is readily cultured and has been widely studied and characterized. However, its genome sequence remains unannotated [18, 19] and knowledge of gene content has relied heavily on transcriptome analyses [13, 20]. Insight regarding microbial interactions with Euglena in various environments has been gained by establishing synthetic associations in culture. Notably, E. gracilis is incapable of synthesizing vitamins B1 or B12 and must obtain these from exogenous sources [21]. When it is cultured with bacteria that can produce these vitamins, Lysinibacillus boronitolerans or Pseudobacillus badius, E. gracilis is able to grow for several generations without exogenous vitamin application [21]. Euglena biomass and compound production was also enhanced when grown with the growth promoting bacteria Emiticicia sp. EG3, [10] or with Vibrio natriegens, a bacterium capable of synthesizing the phytohormone indole-3-acetic acid [22]. While studying synthetic associations are informative and have led to many interesting ndings related to fungal-algal-bacterial (FAB) associations, the full potential of Euglena co-cultures and FABs would be better revealed by investigating naturally occurring interactions such as those between E. mutabilis and its inextricably associated partner organisms [23, 24, 25]. E. mutabilis is an extremophilic Euglenoid that is often found in toxic environments such as peat bogs, volcanic lakes and acid mine drainage (AMD) [25, 26, 27]. In fact, its ability to grow in environments where few other organisms can grow has led to it becoming a bioindicator for AMD [28, 29, 30, 31]. Acid drainage is naturally produced as water seeps through iron sul de-aggregated rocks; however, its production is dramatically enhanced by the disposal of industrial mining waste which creates acidic pools of toxic substances including high concentrations of heavy metals (HMs). AMD thus poses severe human and environmental health concerns [28]. When E. mutabilis grows in these environments it often associates with other microorganisms to form bio lms [11, 27, 32, 33, 34]. In an attempt to create an axenic culture of E. mutabilis from an acidic pond in the Northwest Territories (Canada) the resulting culture was found to contain a yeast, later identi ed as a Cryptococcus sp [11]. The fungus was originally considered a contaminate, but the culture could not be cured, and the authors concluded this was a probable E. mutabilis-fungal mutualism [11]. The di culty in obtaining axenic E. gracilis cultures was also noted during subsequent isolation attempts from toxic environments [11, 35]. This led us to hypothesize that one or more partner organisms augment the stress tolerance of E. mutabilis and aid its ability to survive in acidic metal-polluted environments. The objective of the presented research was to test this hypothesis by investigating the impact of antibiotic treatments on the growth and cadmium tolerance of a natural E. mutabilis, fungal, and bacterial co-culture. A culture of E. mutabilis originally from an AMD site in Timmins, Ontario, Canada was obtained. In this culture, extensive attempts to axenically separate fungi and bacterial from the E. mutabilis were not successful. Therefore, in the present study, an array of ve antibiotics and two antimycotics were used in systematic attempts to suppress the growth of the partner organisms in the Page 3/30 presence and absence of 100 µM CdCl2. Our ndings indicate that there is a signi cant decrease in numbers of viable E. mutabilis cells across all antibiotic and CdCl2 treatments compared to E. mutabilis co-cultures that are only exposed to CdCl2. This indicates the that associated microbes play a role in modulating the HM stress responses of Euglena. Suppressing bacterial and fungal growth also revealed the complexity of the interactions in this naturally occurring FAB association and supports the further investigation of natural interactions as model FABs for use in microbe technologies including bioremediation. Methods Culture Selection and Growth of E. mutabilis and E. gracilis Field samples of E. mutabilis were obtained from by the Canadian Phycological Culture Centre (CPCC, University of Waterloo, Canada). The strain of E. mutabilis (CPCC 657) contained unidenti ed bacteria and fungi. The CPCC did not have an axenic culture of E. mutabilis, therefore an axenic culture of E. gracilis (CPCC 95) was obtained to act as an experimental control. Both organisms were grown autotrophically in 250mL Pyrex Erlenmeyer asks capped with foam stoppers and aluminum foil under standard aeration (100RPM on a Thermo Fisher Scienti c MaxQ 3000), while cycling light and temperature (16:8 LD cycle at 260 µmol s− 1 m− 1; 24℃ ± 0.5℃ in light and 18℃ ± 0.5℃ in dark) in a Conviron PCG20 environmental chamber. Stock cultures were grown in Modi ed Acid Medium (MAM), a de ned inorganic medium [36], with modi cations recommended by the CPCC and adjusted to a pH between 4.3–4.5 [37]. Filter sterilized F/2 vitamin mix was added after the medium was autoclaved. Antibiotic Preparation The ve antibiotic and two antimycotic stock solutions were prepared with appropriate solvents. Kanamycin monosulfate (Bioshop Cat No. #KAN201) and tetracycline hydrochloride (Fisher Scienti c Cat No. #A39246) were reconstituted in sterile MilliQ (13.4MΩ·cm) water at a concentration of 1280 µg/mL. Rifampicin (Fisher Scienti c Cat No. #5573031GM), chloramphenicol (Fisher Scienti c Cat No. #AAB2084122), and cycloheximide (Fisher Scienti c Cat No. #AAJ6690103) were reconstituted in MeOH, 100% EtOH, and 95% EtOH, respectively, at a concentration of 1280 µg/mL. A penicillin-streptomycin blend was purchased in solution (Fisher Scienti c Cat No. #15140122) and diluted using sterile MilliQ (13.4MΩ·cm) water to a concentration of 1280 units/L of penicillin-streptomycin. Amphotericin B was also purchased as a solution (Fisher Scienti c Cat No. #15290026) and its concentration was not modi ed (250 mg/L). Antibiotics were stored at -20℃ in sterile polypropylene test tubes (VWR Cat No. #CA60819-761) with para lm around the lid. E. mutabilis and E. gracilis Antibiotic Treatments Cell counts of Euglena from stock cultures of E. mutabilis were performed using a hemocytometer (Hausser Scienti c). A 1mL volume of 500,000 cells were aliquoted into 1.5mL microfuge tubes which were centrifuged (6000RCF for 5 minutes), and the supernatant was removed using a micropipette. The Page 4/30 pelleted cells were inoculated into a well of a 12-well culture plate (VWR Cat No. #10861-556) containing 1mL of media per well. Separate 12-well culture plates (VWR Cat No. # 10861-556) were prepared for each antibiotic and antimycotic (Figure S1). Culture plates contained a sterile control (900µL of MAM and 100µL of antibiotic solvent), growth control (900µL of MAM, 100µL of sterile MilliQ (13.4MΩ·cm) water, and 500,000 cells), and growth “check” (900µL of MAM, 100µL of alcohol, and an aliquot of cells) if an antibiotic was reconstituted in an alcohol. Wells that contained antibiotics were prepared using serial dilutions beginning with 1800µL of MAM and 200µL of an antibiotic or antimycotic in well A4. Upon completion of serial dilutions each well contained a nal volume of 1000µL of media with antibiotic concentrations of 64, 32, 16, 8, 4, 2, 1, 0.5, and 0.25 µg/mL. Due to its stock concentration, modi cations were made to the volume of media and antimycotic in culture plates containing Amphotericin B, however nal antibiotic concentrations were the same (Figure S1). Cells were added to each well in the culture plate except for the sterile control. Culture plates were sealed with para lm and stored in the Conviron PGC20 environmental chamber under the same temperature and light conditions as the stock cultures with aeration (120RPM) for 72 hours. This time was previously determined to be the maximum length that amphotericin B and penicillin-streptomycin remained fully effective [38]. An identical method and analysis were used for E. gracilis. Each experiment was replicated 3 times. Euglena Cell Viability After Antibiotic and Antimycotic Exposure 72 hours post-inoculation 100µL aliquots were taken from each well containing cells. Then 400µL of a 0.4% solution of Trypan Blue (Bioshop Cat No. #TRY477.100) in PBS (Bioshop, Cat No. #PBS404.100) was added to each aliquot. Euglena cell viability was assessed by visualizing differential uptake of Trypan Blue stain, and cell counts were performed using a haemocytometer (Hausser Scienti c). This was carried out on both E. mutabilis and E. gracilis. Assessment of Chlorophyll Content A modi cation of the Methanol (MeOH) extraction procedure developed by Warren (2008) was used to extract chlorophyll [39]. In the modi ed procedure, 500µL aliquots from each well in the culture plates were added to 2mL Eppendorf tubes containing 200µL of glass beads (Sigma Aldrich Cat No. #G8772). Media was decanted after centrifugation at 4℃ (maximum speed for 2 minutes). 1mL of chilled MeOH was added to the Eppendorf tubes and samples were bead beat at 30Hz for 2 minutes (Retsch Mixer Mill MM 400). Tubes were then centrifuged at 4℃ (maximum speed for 2 minutes) and MeOH was aliquoted into a fresh 2mL Eppendorf tube. This method was repeated so the nal volume of extracted chlorophyll was 2mL in MeOH. 200µL of extract was added to each of three 96-well optic plates (ThermoFisher Scienti c Cat No. #165305) for triplicate readings on a BioTek Synergy HTX Multimode Reader at 652nm and 665nm wavelengths. Chlorophyll content was determine using the equations from Warren (2008) and chlorophyll content was normalized against the amount of chlorophyll in each Euglena cell. Page 5/30 Growth Check of Cultures After Antibiotic Treatment Reasoner’s 2A (R2A) and Potato Dextrose Agar (PDA) plates were prepared using a 1.4% (w/v) agar mixture. Solutions were autoclaved to sterilize, and then 15mL of media was added to each plate (Greiner Bio-One Cat No. #633181). 100µL aliquots were taken from wells in the culture plates containing cells and diluted with 900µL of sterile 0.9% NaCl in MilliQ (13.4MΩ·cm) water. 50µL of diluted sample was added to an agar plate, sealed with para lm, and incubated in darkness at 24℃ for 7 days. Cultures treated with antibiotics were plated on R2A while cultures treated with antimycotics were plated on PDA. The number of Euglena, fungal, and bacterial colony forming units (CFUs) were noted every 24 hours throughout the 7-day incubation period. Cadmium Tolerance of E. mutabilis and E. gracilis with Antibiotics Kanamycin, rifampicin, chloramphenicol, tetracycline hydrochloride, penicillin-streptomycin, and amphotericin B at concentrations of 64, 32, 16, 8, 4 and 2 µg/mL were selected as the antibiotics moving forward to assess CdCl2 tolerance of E. mutabilis and E. gracilis with the addition of antibiotics and antimycotics. The inoculation of 12-well plates was carried out in the same manner as indicated for the cell viability assessment with 500,000 cells being added to each well, with the antibiotics added through serial dilution as described (Figure S1). 10µL of CdCl2 with a stock concentration of 10,000 µM was added to each well in the plate containing antibiotics or antimycotics to yield a nal concentration of 100 µM CdCl2. One well in each plate was a CdCl2 control containing only CdCl2 at a concentration of 100 µM and cells. Culture plates were sealed with para lm and incubated in the environmental chamber under the same conditions (16:8 LD cycle at 260 µmol s− 1 m− 1; 24℃ ± 0.5℃ in light and 18℃ ± 0.5℃ in dark; shaking at 120RMP) as the antibiotic trials for 72 hours. After 72 hours assessments of cell count, cell viability, chlorophyll content, and agar plate growth checks were carried using the methods described. This method of assessing Cd tolerance, including subsequent analyses, were carried out on E. mutabilis and E. gracilis. All experimental were performed in triplicate. DNA Isolation Total genomic DNA (gDNA) was isolated from cultures of CPCC 657 grown in different medias to selectively enhance or limit growth of E. mutabilis and its unidenti ed bacterial and fungal partners. CPCC 657 was grown in MAM (pH 4.3), MAM with CdCl2 at a concentration of 100 µM (pH 4.3), MAM (pH 2.7), MAM with CdCl2 at a concentration of 100 µM (pH 2.7), Potato Dextrose Broth (PDB) grown in light, PDB grown in dark, TSB grown in light, and TSB grown in dark. Cultures were grown for 7 days in the Conviron PGC20 environmental growth chamber under standard aeration (100RPM on a Thermo Fisher Scienti c MaxQ 3000), while cycling light and temperature (16:8 LD cycle at 260 µmol s− 1 m− 1; 24℃ ± 0.5℃ in light and 18℃ ± 0.5℃ in dark). Page 6/30 After 7 days 1mL of culture was aliquoted into a microfuge tube, centrifuged (6000RCF for 5 minutes), and media decanted. Cells were resuspended in 300µL of Triton Solution and transferred to a 1.5mL screw cap tube containing 100µL of glass beads and 300µL 25phenol:24Chloroform:1 Isoamylalcohol. Samples were vortexed for 6 minutes, then 200µL of TE was added, and the tubes were centrifuged at 4℃ (maximum speed for 5 minutes). The aqueous layer was transferred to a fresh 1.5mL microfuge tube. Samples were precipitated in 1mL of ice-cold 100% EtOH and centrifuged at 4℃ (maximum speed for 2 minutes) before decanting the EtOH. 400µL of TE and 1.5µL of RNAs A solution (20 mg/mL) was added to each tube and samples were incubated at 37℃ for 30 minutes. After incubation, 10µL of 4M ammonium acetate and 1mL of ice-cold 100% EtOH was added to each tube, they were mixed, centrifuged at 4℃ (maximum speed for 2 minutes) and decanted. The pellet was washed with ice-cold 70% EtOH. Following centrifugation, the EtOH was decanted, and the pellets were air dried and dissolved in 50µL TE. Samples were visualized on a 0.8% EtBr agarose gel to assess the integrity of gDNA. 16S and ITS Full-Length Amplicon PacBio Sequencing High quality gDNA samples (at least 100 ng/µL and a 260/280 ratio of approximately 1.80 as determined using a Thermo Fisher Scienti c Nanodrop 8000 Spectrophotometer) were sent to the Integrated Microbiome Resource (IMR, Dalhousie University, Canada) for library preparation and sequencing on a PacBio Sequel II sequencer. Full 16S (forward primer: 27F(Paliy) = 5’-AGRGTTYGATYMTGGCTCAG-3’; reverse primer: 1492R = 5’-RGYTACCTTGTTACGACTT-3’) and full ITS (forward primer: 5’-ITS1FKYO2-3’ = 5’-TAGAGGAAGTAAAAGTCGTAA-3’; reverse primer: ITS4KYO1 = 5’-TCCTCCGCTTWTTGWTWTGC-3’) sequences were obtained and reported in the form of hi .fastq.gz le types. Bioinformatic Analysis 16S and ITS sequencing data were processed using the PacBio CCS pipeline [40] of the Quantitative Insights Into Microbial Ecology 2 (QIIME2) v 2022.11 software [41]. The dada2 algorithm was used to denoise raw sequences [42]. Taxonomic classi cation was performed against the SILVA 138.1 SSU Ref NR99 full-length database for 16S data [43] and the ITS custom classi er for all eukaryotes generated from the UNITE database [40], using the SK-Learn command from the q2-feature-classifer plugin [44]. Reads were then rari ed to lter out low depth samples and amplicon sequence variants (ASVs) with frequencies of less than 100, as well as mitochondrial, chloroplast, and unclassi ed sequences. Resultant tables from QIIME2 were imported into R-Studio for visualization and graphical analysis. The relative abundance of each ASV was plotted using the phyloseq package in R [45]. PCR Ampli cation PCR ampli cation was performed in a total reaction volume of 50µL containing 4µL of template DNA, 27.5µL of sterile deionized water, 4µL of dNTPs, 10µL of Phusion Buffer HF (Thermo Scienti c™), 0.5µL of Phusion DNA Polymerase (Thermo Scienti c™), and 2µL of each primer (5µM). The ITS region was ampli ed [46] (forward primer: ITSF = 5’-CTTGGTCATTTAGAGGAAGTAA-3’; reverse primer: ITS4 = 5’TCCTCCGCTTATTGATATGC-3’), and the 16S region was ampli ed [47] (forward primer: 68F = 5’TNANACATGCAAGTCGRRCG-3’; reverse primer: 518R = 5’-WITACCGCGGCTGCTGG-3’). Ampli cations Page 7/30 were performed with a Veriti™ Thermocycler (Applied Biosystem™), using the following program: denaturation for 30 seconds at 98°C; then 35 cycles consisting of 98°C for 10 seconds, 61°C for 30 seconds, and 72°C for 30 seconds; and a nal extension step at 72°C for 10 minutes. Each sample was ampli ed by 6 separate reactions with a volume of 50µL and subsequently pooled for a total sample volume of 300µL. Gel Extraction and DNA Puri cation 60µL of loading dye was added to each pooled sample and ampli cation products were analyzed by electrophoresis in a 0.8% (w/v) agarose (BioShop Canada Inc.) gel in 0.5 × TE with ethidium bromide in order to visualize DNA bands. ITS ampli cation resulted in a DNA fragment of approximately 460bp, while 16S ampli cation resulted in a DNA fragment of approximately 400bp. The DNA band visualized on the gel at the appropriate location was subsequently cut out and puri ed using the PureLink™ Quick Gel Extraction Kit (Invitrogen™). Sequencing Sanger Gel-puri ed DNA was prepared for Sanger Sequencing using the BigDye™ Terminator v3.1 Cycle Sequencing Kit with modi cations. Template DNA was normalized to approximately 70ng/µL and 3.00µL was added to a 96-well plate (Progene Cat. No. #87-C96-ABI-2) along with 3.20µL of primer (0.5µM), 1.33µL of Ready Reaction (Applied Biosystems™), 1.33µL of BigDye Buffer (Applied Biosystems™), and 1.14µL of deionized water for a total volume to 10.00µL in each well. 6 primers were selected for ITS identi cation [46] and 2 primers were used for 16S identi cation [47] (Table S2). Ampli cations were performed with a Veriti™ Thermocycler (Applied Biosystem™), using the following program: denaturation for 1 minute at 96°C; then 40 cycles consisting of 96°C for 10 seconds, 50°C for 5 seconds, and 60°C for 4 minutes. Following ampli cation, 2.5uL of 125mM EDTA and 30µL of 100% cold EtOH was added to each well. The plate was sealed, inverted 4 times to mix, covered with aluminum foil, incubated at room temperature for 15 minutes, and then centrifuged for 4℃ (2,500g for 30 minutes). The plate was then decanted by being placed upside down in a centrifuge and spun (190g for 60 seconds). 30uL of ice cold 70% EtOH was added to each well, the plate was sealed, inverted 4 times to mix, and then centrifuged for 4℃ (1,650g for 15 minutes). The plate was then decanted by being placed upside down in a centrifuge and spun (190g for 60 seconds). 15µL of Hi-Di™ Formamide (Applied Biosystems™) was added to each well, the plate was sealed and quickly centrifuged, and covered in aluminum foil for a 15-minute incubation and room temperature. The plate was run on an ABI 3730 (Applied Biosystems™) and bases were called using SequencingAnalysis v5.4 to produce .phd1 les. The quality of the les was assessed using Seq Scanner 2 (v. 2.0; Applied Biosystems), and the data was assembled into contiguous (contig) sequences using SeqMan Pro (v. 11.2.1; DNASTAR). The average read length +/- the standard deviation was assessed for each contig assembly. Generated contigs were then analyzed using Nucleotide BLAST. Results Antibiotics Treatments and CdCl 2 Page 8/30 After 72 hours of growth, control E. mutabilis cells arrived at a concentration of approximately 9.4 x 105 ± 6.4 x 104 cells/mL, while cells exposed to 100 µM CdCl2 were approximately 5.5 x 105 ± 1.7x104 cells/mL (a 41% reduction in cells). Similarly, control E. gracilis cells grew to a concentration of approximately 8.0 x 105 ± 8.1 x 104 cells/mL, while cells exposed to 100 µM CdCl2 grew to approximately 5.4 x 105 ± 3.3 x 104 cells/mL (a 31% reduction in cells). A t-test revealed that the application of 100 µM CdCl2 statistically (p < 0.05) decreases the number of viable E. mutabilis and E. gracilis cells in culture, con rming that the application of CdCl2 inhibits the growth of Euglena. Cell counts of E. mutabilis and E. gracilis following antibiotic exposure revealed in uences of antibiotics on algal growth (Table S1). Applications of kanamycin, rifampicin, chloramphenicol, and amphotericin B produced similar growth responses in both Euglena species. Consistent with previous studies, neither species was affected by kanamycin [48, 49], as evidenced by the lack of statistical (t-test-; p < 0.05) difference in cell counts, while rifampicin, chloramphenicol, and amphotericin B resulted in signi cant decreases in cell counts when exposed to 8–64 µg/mL concentrations. Notably, the in uence of the penicillin-streptomycin blend and tetracycline differed between E. mutabilis and E. gracilis. Penicillin and streptomycin are antibiotics that have repeatedly been shown to bleach E. gracilis [48, 50, 51, 52] and treatment with the blend here showed a signi cant reduction in E. gracilis cell counts at every concentration used; however, these antibiotics did not have a signi cant impact E. mutabilis cell counts. This trend was also seen with the application of tetracycline, which despite previous studies stating that E. gracilis had tolerance at concentrations ranging from 100–300 µg/mL [49, 53], statistical decreases in cell counts were observed across all our concentrations of tetracycline while E. mutabilis was only signi cantly impacted at 8 µg/mL and 64 µg/mL concentrations. Although some antibiotics impacted growth of E. mutabilis and E. gracilis, the combination of antibiotics and CdCl2 reveals more substantial differences in response between the organisms (Fig. 1). The number of viable cells of E. mutabilis treated with both antibiotics and 100 µM CdCl2 were signi cantly less than all treatment combinations involving cells treated with only antibiotics (t-test, p < 0.05; denoted by * in Fig. 1). In contrast, statistical decreases in E. gracilis cells under the same treatments were only observed about 50% of the time. The addition of 100 µM CdCl2 resulted in decreases of viable E. mutabilis cells ranging from 31–92%, while decreases in E. gracilis cells ranged 1–44%. The biggest differences in cell counts were observed in samples treated with chloramphenicol and 100 µM CdCl2, which resulted in decreases in cell counts ranging from 64–92% in E. mutabilis and 9–44% in E. gracilis, compared to cells treated with only chloramphenicol. Additionally, E. mutabilis cells treated with all concentrations of antibiotics and 100 µM CdCl2 resulted in statistical decreases in cell counts compared to E. mutabilis cells treated with only 100 µM CdCl2 (t-test, p < 0.05; denoted by ฀ in Fig. 1). This differed from E. gracilis where approximately one-third of the antibiotic treatments revealed a decrease. A nal note of the differences between species is that, while there was no difference between the number of viable cells of E. mutabilis and E. gracilis when exposed to 100 µM CdCl2, the number of E. mutabilis cells was higher across every concentration of antibiotic in combination with 100 µM CdCl2. This suggested that the Page 9/30 addition of antibiotics had a greater impact on the CdCl2 tolerance of the E. mutabilis co-cultures compared to those of axenic E. gracilis. The antibiotics that had the greatest impact on Euglena cell viability were rifampicin and chloramphenicol for which 64 µg/mL treatments resulted in no viable E. mutabilis or E. gracilis cells (Fig. 1). We observed opposite growth patterns between E. mutabilis and E. gracilis following treatment with chloramphenicol. Treatment with 32 µg/mL of chloramphenicol revealed the greatest number of viable E. mutabilis cells, which remained as high as control conditions, while the least number of viable cells was seen at a concentration of 2 µg/mL. By contrast, the lowest number of viable E. gracilis cells was observed at 32 µg/mL while the greatest number appeared at 2 µg/mL. When E. mutabilis was treated with 100 µM CdCl2 it displayed the same trend as E. gracilis. Evaluation of Chlorophyll Content After Antibiotic Treatments and CdCl2 The total chlorophyll content (chl a + chl b) of E. mutabilis and E. gracilis cultures treated with CdCl2 was on average 1.95-fold and 1.82-fold higher, respectively, than cells only treated with antibiotic (Fig. 2). The concentrations of chlorophyll detected did not vary with antibiotic concentration, and the increase in the presence of CdCl2 was consistent across all antibiotic treatments except that of chloramphenicol. In the case of chloramphenicol, treatment with CdCl2 resulted different increases in chlorophyll content with antibiotic concentrations. Growth Recovery Following Antibiotic Treatments and CdCl 2 Exposure The ability of Euglena, and the co-cultured fungus in the case of E. mutabilis, to recover from treatment with antibiotic or antimycotic in the presence or absence of CdCl2 was assessed by plating on nonselective media. The plates were incubated in the dark for 7 days leading to heterotrophic growth only. In all instances, bacterial colonies too numerous to count were present on the plates and intimate growth was observed amongst the FAB consortia (Figure S2). The control for the experiment was phototrophic growth in MAM +/- 100 µM CdCl2, followed by heterotrophic growth. Growth in MAM followed by heterotrophic growth led to large numbers of E. mutabilis colonies, but no fungal CFUs. Phototrophic growth in MAM + CdCl2 followed by heterotrophic growth resulted in E. mutabilis and fungal colonies. Pregrowth in the presence of antibiotic, with and without CdCl2, led to reduced numbers of colonies for both Euglena species for most of the antibiotics. An exception for E. mutabilis occurred when pre-grown with tetracycline; whereby, it did not affect heterotrophic growth, but pre-growth with tetracycline and CdCl2 did. For E. gracilis pre-growth in kanamycin had no detectable difference in growth relative to the control, and pre-growth in amphotericin B led to less growth than the control; however, growth in amphotericin B with CdCl2 did not in uence growth. Page 10/30 While there was no growth of the fungus after phototrophic growth of the colony in the absence of antibiotics, fungal growth was noted following pre-growth in the presence of all antibiotics. However, growth was greatly reduced following pre-growth with kanamycin or amphotericin B. When CdCl2 was included in the pre-growth media, fungal growth was indistinguishable from the control. Table 1 Cell viability test (n = 3) comparing colony forming units of E. mutabilis (CPCC 657) and uncharacterized fungal growth, in addition to E. gracilis (CPCC 95) after 7 days incubation at 24oC in darkness. Bacterial CFUs are excluded as they were too numerous to count. E. mutabilis Fungi Media Only 100 µM CdCl2 Media Only 100 µM CdCl2 Media Only 100 µM CdCl2 +++ ++ - +++ ++ +++ 32 µg/mL 32 µg/mL + 100 µM CdCl2 32 µg/mL 32 µg/mL + 100 µM CdCl2 32 µg/mL 32 µg/mL + 100 µM CdCl2 Kanamycin ++ + + +++ ++ +++ Rifampicin + + +++ +++ + + Chloramphenicol + + +++ +++ + ++ Tetracycline +++ + +++ +++ + ++ PenicillinStreptomycin ++ + +++ +++ + ++ Amphotericin B ++ + + +++ + +++ Control E. gracilis (–) no CFU, (+) if < 50 CFU, (+ +) if 50 > < 150 CFU, and (+ + +) if > 150 or complete lawn present and CFU count impossible. PacBio Sequencing and Culture Identi cation A total of 372,830 reads were obtained from sequencing the full length 16S ribosomal RNA (rRNA) gene. Reads per sample ranged from 7,297 to 62,135. Sequencing the full length internal transcribed spacer (ITS) gene obtained a total of 154,027 reads, ranging from 165 to 29,136 per sample. Denoising left 229,321 reads (62%), with an average of 22,932 sequences per sample for 16S, and 141,901 reads (92%), with an average of 14,190 sequences per sample for ITS. Filtering of low-level amplicon sequence variants (ASVs), low depth samples, as well as unclassi ed, chloroplast, and mitochondrial sequences resulted in the exclusion of six samples and reserved a total of 34,857 reads (9%), with an average of 8,714 reads per sample for 16S. The ltering process resulted in the exclusion of three samples from ITS and reserved 140,242 sequences (91%), averaging 20,034 sequences per sample. The sequences that Page 11/30 passed the denoising and ltering processes were clustered into four ASVs for 16S and three ASVs for ITS. 91% of the 16S sequences were excluded due to low sample depth, low frequency, or were classi ed as mitochondrial, chloroplast or unclassi ed at the genus level. Of these, 187,716 sequences were classi ed as Euglena, alluding to the overrepresentation of this contributor. 16S sequencing identi ed one bacterial genus, Acidiphilium (Fig. 3a). Of the sequences classi ed at the genus level as Acidiphilium, 46% were further classi ed at the species level as Acidiphilium acidophilum. The remaining sequences could not be classi ed to the species level. Thus, the major bacterial contributor is determined to be Acidiphilium acidophilum. ITS sequencing identi ed three fungal genera: Acidomyces, Exophiala, and Talaromyces (Fig. 3b); only one ASV could be classi ed to the species level (Exophiala oligosperma). 98% of the total ITS sequences that passed ltering were classi ed as Talaromyces, whereas Exophiala and Acidomyces contributed only 2.13% and 0.09% of sequences, respectively. Thus, the major fungal contributor is determined to be a species of the genus Talaromyces. Sanger Sequencing Results Sanger sequencing for ITS samples generate high-quality reads of 235 ± 119 bps with initially ambiguous bases being manually called. Talaromyces amestolkiae showed a 93.63% match with the raw data (562bp length), which was increased to 94.64% upon manually calling ambiguous bases (565bp length). 16S reads generated were of medium quality, with a mean ± standard deviation of 151 ± 27 base pairs. Contig assembly produced one single contig with a length of 281bp, approximately 62% of the target region. The top match for the sequence was a 94.58% match to an uncultured bacteria isolated from acidic soils, while the next result was a 94.22% match to Acidiphilium sp. Discussion The Cd tolerance of an E. mutabilis natural co-culture with unknown fungi and bacteria was investigated. The culture bank from which the E. mutabilis isolate was obtained had been unable to create an axenic E. mutabilis culture from any isolate. Therefore, we used an axenic E. gracilis culture as a control to determine the comparative impact of treatments on the E. mutabilis co-culture. E. gracilis is the model euglenoid and has been the focus of most the antibiotic and HM experiments using a euglenoid [48, 49, 54, 55]. Recent investigation revealed E. mutabilis had greater tolerance to Cd than E. gracilis, and that it responded differently to pre-growth under nutritional conditions [56]. The current study is the rst assessment of the impact of antibiotics on E. mutabilis. Rifampicin had a very toxic effect on both Euglena species in the presence and absence of CdCl2. In all other antibiotic or antimycotic treatments, the combined exposure to CdCl2 signi cantly decreased the number of viable E. mutabilis cells (Fig. 1). In contrast, the impact of the combined treatments on E. gracilis was either not signi cant or the reduction was substantially less than noted for E. mutabilis (Fig. 1). The assessment of antibiotic and CdCl2 by determining chlorophyll concentrations was consistent between Euglena species showing that cellular chlorophyll content generally increases after cells have been exposed to CdCl2 - with the exceptions of rifampicin and chloramphenicol where results varied (Fig. 2). The viability of E. mutabilis growing heterotrophically after antibiotic and CdCl2 treatments was substantially decreased, whereas the decrease Page 12/30 in E. gracilis was lower or non-existent. However, the fungus in E. mutabilis cultures remained viable and grew to cover the plates after these treatments (Table 1). Since fungal viability was signi cantly less following phototrophic growth of the culture, it is possible that suppression of bacterial growth enabled the fungus to survive and proliferate. Combined with the negative impact on CdCl2 tolerance in E. mutabilis following amphotericin B treatment, these results reveal several combinatorial interactions between the bacteria and E. mutabilis, the bacteria and the fungus, and the fungus and E. mutabilis. Consistent with synthetic co-culture work [57, 58, 59, 60] these results indicate that the fungus and bacteria assist in the resistance of E. mutabilis to Cd. Further, they reveal multiple organismal interactions in uence the response of the co-culture to HM challenge. This knowledge provides the necessary base for subsequent investigations into the mechanisms fungal-E. mutabilis-bacterial consortia resistance to HMs and provides a model for FAB consortia resistance to HMs. Current methods for treating waterbodies that have been polluted with HMs include neutralization processes, chemical precipitation, coagulation/ occulation, and adsorption; however, these processes suffer from high costs, ine cient HM removal, and may produce secondary pollution [28, 61, 63, 63]. As a result, biotechnological methods are being employed to enhance the bioremediation potential of microbes [59, 64, 65, 66, 67, 68]. Several algae have been investigated for bioremediation of textiles, organic pollutants, and HMs [69, 70, 71, 72], and recent studies have suggested that co-culturing algae with bacteria or fungi provides bene ts that include increased occulation e ciency, increased biomass, independent nutrient exchange, and enhanced tolerance to extreme environments [7, 67, 73, 74, 75, 76]. The model euglenoid E. gracilis has also been co-cultured with other microbes [21, 77], however the discoveries here show that E. mutabilis and its naturally associated microbial partners offer substantial insight into how organisms interact in a co-culture and provides a model to use for developing enhanced technological applications and potential use in bioremediation. Co-culturing algae with bacteria or fungi has several advantages, including enhanced nutrient exchange or protection factors for the alga [2, 4, 78]. It is widely known that many algae rely on exogenous vitamin B12 (cobalamin) and nitrogen, and that these can be supplied by bacteria [2, 79, 80] in exchange for the bacteria receiving a carbon source from the algae [81, 82]. As examples, the growth of marine diatom Ditylum brightwellii in nitrogen limiting media and a symbiotic bacterium in carbon limited media are both lower relative to co-cultures [83]. Similarly, an investigation of a symbiotic culture of Chlorella vulgaris and Bacillus subtilis revealed that co-culturing resulted in increased growth, photosynthetic activity, carbon xation, and vitamin B12 content of the alga [84]. Alternatively, fungi that are cultured with algae provide nutrients to the alga as well as mechanisms for protection [78, 85, 86]. When Chlamydomonas reindhardtii was cultured with Aspergillus nidulans and exposed to the algicide azalomycin F, algal cells grew within fungal hyphae avoiding contact with the algicide [78]. Additionally, the presence of azalomycin F prompted A. nidulans to produce polar lipids which attract the algicide and effectively neutralize it [78]. Fungi also provide protection from reactive oxygen species (ROS) generated during HM exposure and produce extracellular polymeric substances (EPS) and organic acids that can bind HMs reducing their toxicity when co-cultured with algae [87, 88, 89]. Our study found that suppression of Page 13/30 constituent bacterial and fungal organisms naturally associated in the E. mutabilis ‘FAB’ co-culture led to signi cantly lower numbers of viable Euglena cells after CdCl2 exposure. This is consistent with previous studies showing the importance of co-cultured organisms with algae; however, we did not have to screen for compatible interactions since we had a naturally compatible and e cient association that evolved in a toxic AMD environment. Further, the majority of research has used cultures of only two organisms: algae and bacteria or algae and fungi. Apart from lichen studies, FAB consortia have rarely been investigated for use in bioremediation despite reports of this tripartite interaction being considered a selfsustaining and cost-conscious system that exhibits superior HM removal e ciency [58]. Our results underscore the innate interaction of FAB co-cultures in extreme environments and establishes a foundation for investigating the association between natural FABs to inform future developments in biotechnology. Although we observed a decrease in the number of viable Euglena cells following treatment with antibiotics and CdCl2, there was an increase in chlorophyll production by Euglena in the presence of Cd (Fig. 2). This was unexpected because Cd is known to disrupt physiological and metabolic processes of phototrophic organisms including algae, cyanobacteria, and plants primarily by reducing photosynthetic rate and chlorophyll concentration [90, 91, 92, 93, 94]. Both E. mutabilis and E. gracilis display signi cantly greater chlorophyll per 100,000 cells under Cd exposure. Furthermore, the amount of chlorophyll being produced in the presence of Cd appears unaffected by the addition of most antibiotics suggesting that the impact does not involve the associated bacteria. This phenomenon has been reported in higher plants that have taken up Cd from their surroundings [95, 96]. The structural foundation of a chlorophyll molecule is Mg; however, Mg is also essential for several other metabolic processes including enzyme activation, sucrose transport, and energy metabolism [97]. It has been shown that Cd can replace Mg in the central position of the chlorophyll molecule [95]. Further, the Cd hyperaccumulator Sedum alfredii demonstrated no noticeable reduction in photosynthetic activity and simultaneous increase in chlorophyll content following Cd treatments, which negatively impacted leaf and root growth [96]. A similar result was observed in Chlamydomonas reinhardtii under excess Cu exposure where the correlation between increased chlorophyll and cell survivability suggested to be the result of chlorophyll accumulation in cells that do not divide [98]. Here we showed that Cd treatment of Euglena led to a reduction in cell viability and an increase in the amount of chlorophyll produced per cell. This could indicate that although exposure to divalent HMs debilitates overall culture health, Cd may be able to replace the Mg in chlorophyll leaving a less e cient but more plentiful photosynthetic apparatus, which, although less effective, still allows the Euglena to survive some HM toxicity. Despite the increasing chlorophyll content per cell, E. mutabilis is unable to recover from antibiotic and Cd exposure during subsequent heterotrophic growth (Table 1). In contrast the fungi and bacteria in the coculture adapt to the switch to heterotrophic growth and often grow to take over the entire plate. The effects of antibiotics on E. gracilis have been extensively studied [48, 50, 52, 53, 99, 100] and the results here are consitent with previous ndings; however, there has been no work on the impact of antibiotics on E. mutabilis. Based on CFU comparisons, E. mutabilis recovers after antibiotic exposure better than E. Page 14/30 gracilis, but E. gracilis shows better recovery following treatment with antibiotic and Cd (Table 1). This is notable because a synergistic effect can occur where toxicity is increased through the formation of antibiotic and di-valent HM complexes [101, 102]. The increased chlorophyll content in the presence of Cd suggests that both Euglena species are capable of generating energy and surviving the treatment. Then, upon transfer to nutrient rich media, they could both shift to heterotrophic growth and employ major facilitator superfamily (MFS) transporters, transmembrane (TrkA) transporters, HM pumps (P1 B ATPase) and other means to recover from the Cd stress [103, 104]. We postulate that while both Euglena species can recover, the recovery by the fungi and bacteria co-cultured with E. mutabilis occurs more quickly than that of E. mutabilis; consequently, substantial fungal and bacterial growth in heterotrophic conditions results in fewer E. mutabilis colonies being formed. Related to this are the observations that fungal growth is only visible following heterotrophic growth after antibiotic exposure; whereas, with antibiotic treatment under the phototrophic growth in MAM at pH 4.3, the fungus is not even microscopically observable. It is only upon introduction to nutrient rich media following antibiotic or Cd treatment that fungal colonies can be observed. This may indicate that bacteria interact with, or rely on, the fungus. Our results show that the number of fungal CFUs decreases with the concentration of antibiotics (Table S3) suggesting that the fungal-bacterial interaction is not simply a matter of presence or absence. In heterotrophic cultures containing fungi and microalgae, a similar over-growth by the fungus was observed [105, 106, 107, 108]. Together these results suggest that co-cultures, which lack a readily available carbon source, require a speci c light regimen and a speci c organism ratio to ensure the fungus does not overwhelm the photobiont. Sequence analysis of DNA extracted from the E. mutabilis FAB co-culture was used to identify the prominent constituent organisms in the co-culture. The fungus was determined to belong to the genus Talaromyces, and with a possible species identi cation being T. amestolkiae, while the bacteria were determined to belong to the genus Acidiphilium, and with a probable species identi cation being A. acidophilum (Fig. 3). Consistent with the fungus belonging to Talaromyces was the observation of yellow and red fungal colonies on PDA plates following amphotericin B and cycloheximide exposure (Figure S3) [109]. Sequence analysis also detected a low-level sequence with similarity to Exophiala oligosperma; however, this may have been an artifact since E. oligosperma colonies are black in colour and there is no evidence that E. oligosperma can tolerate HMs [110, 111]. Talaromyces sp., on the other hand, have been characterized by their ability to produce different coloured pigments based on its carbon source and in response to stress or predators [109, 112, 113]. Furthermore, they have been isolated from HM polluted areas, and can withstand high concentrations of Cr, As, Pb, Ni, Cu, and Cd [114, 115, 116]. Red pigment is produced by fungi that are susceptible to cycloheximide and carry the recessive ade2 gene which, when repressed, results in red pigmentation from the accumulation of phosphoribosylaminoimidazole, an intermediate in the biosynthesis of adenine [117, 118, 119, 120, 121]. The yellow pigmentation following amphotericin B and cycloheximide exposure is most likely comprised of products of the azaphilone family, namely mitorubrinol and mitorubrinic acid [109, 122, 123]. These compounds are found in numerous Talaromyces sp. and are proposed virulence factors regulated by polyketide synthesis genes pks11 and pks12 which become activated under stress [109, 122, 123, 124]. This cumulative information, Page 15/30 in combination with con rmatory Sanger sequencing results, indicates that the predominant fungus present in the E. mutabilis co-culture is a Talaromyces sp. Furthermore, the production of pigment by Talaromyces sp only when stressed or acting as a pathogen indicates that the lack of colour observed during control co-culturing conditions suggested the fungus is neither acting as a pathogen nor stressed. This would be consistent with it having a positive role in the FAB co-culture. Talaromyces sp. are known to produce a plant growth promoting hormone, indole-3-acetic acid (IAA), and they display tolerance to Cd when growing in soil [125]. As such, when this fungus was cultured with Arabidopsis thaliana it reduced the amount of Cd in the soil and underground plant tissues, and increased plant growth. It was postulated that the fungus promoted nutrient uptake and IAA production to promote plant development [126] and provided protection by increasing the essential nutrient bioavailability under low Cd concentrations, thereby effectively diluting the presence of Cd and enhancing plant HM tolerance [125]. These mechanisms are prevalent during other plant-fungal interactions [127, 128, 129] and may be fundamental to stress response in algal-fungal symbiosis [60, 87, 129, 130] as well as being present in the Euglena- Talaromyces interaction. Finding that the predominant bacterial species in the FAB co-culture is an Acidiphilium species is consistent with this species being found in acidic environments with high HM concentrations [131, 132, 133]. A. acidophilum could act as a nutrient source for other organisms in the FAB co-culture as it is a facultative, sulfur-reducing mixotroph [134, 135]; however, there is only evidence of heterotrophic growth in AMD [136, 137]. When growing heterotrophically the main energy source for A. acidophilum is ferrous iron, which it can reduce to generate a usable form [135, 137] and is abundant in AMD (28). The characteristics of Talaromyces sp. and A. acidophilum are consistent with their persistence in the FAB coculture and suggest they have a role in the stress protection of, and nutrient exchange with, E. mutabilis. Conclusions In this study, we investigated a natural FAB association between the extremophilic algae Euglena mutabilis, a bacterial acidophile Acidiphilium acidophium, and a HM tolerant, plant growth promoting, fungus Talaromyces sp. The FAB isolate originated from acid mining runoff in Timmins, Ontario, Canada. We found that the HM tolerance of E. mutabilis is dependent upon its interactions with the constituent organisms present, as suppression of the fungal and bacterial organisms with antibiotics and antimycotics decreased the viability and population growth of E. mutabilis when challenged with Cd. The characteristics of fungi and bacteria are consistent with them providing nutritional and stress protection bene ts to E. mutabilis which, in turn, could provide nutrients or a protective niche for the fungus and bacteria. This interaction combined with the inability to separate the organisms suggest an interdependence, and possibly a tripartite symbiotic relationship which we have coined as a FAB consortia, that can withstand exposure to high concentrations of HM. The unique interaction identi ed in this study between a fungus, a bacterium, and E. mutabilis shown to strengthen the alga’s resistance to Cd may offer insight into the types of FAB interactions that could be used to create a self-sustaining bioremediation technology. Page 16/30 Declarations Data Availability Statement Raw sequence data obtained from PacBio sequencing are available from the NCBI Sequence Read Archive (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1026301; accession number PRJNA1026301). Competing Interests The authors declare no competing interests. Funding This research was funded by Natural Sciences and Engineering Research Council of Canada (NSERC) grant number RGPIN/6029-2019 BJS. Authors’ Contributions BJS and RJNE contributed to the study conception, design, and funding. Material preparation, data collection, and analysis were performed by EK. DP assisted with experimental design and data collection. MG conducted bioinformatic analysis. SG, CD, and MC assisted with data collection. The rst draft of this manuscript was written by EK, DP, and MG, and edits were made by BJS and RJNE. All other authors commented on subsequent versions of the manuscript. All authors have read and approved the nal version. Acknowledgements The authors would like to extend sincere appreciation to Dr. Erin Morrison of Trent University for her support. References 1. Cantin NE, van Oppen MJH, Willis BL, Mieog JC, Negri AP. Juvenile corals can acquire more carbon from high performance algal symbionts. Coral Reefs. 2009;28(2):405–14. 10.1007/s00338-0090478-8. 2. Croft MT, Lawrence AD, Raux-Deery E, Warren MJ, Smith AG. Algae acquire vitamin B12 through a symbiotic relationship with bacteria. Nature. 2005;438(7064):90–3. 10.1038/nature04056. 3. Matthews JL, Crowder CM, Oakley CA, Lutz A, Roessner U, Meyer E, et al. Optimal nutrient exchange and immune responses operate in partner speci city in the cnidarian-dino agellate symbiosis. Proc Natl Acad Sci. 2017;114(50):13194–9. 10.1073/pnas.1710733114. 4. Ramanan R, Kim BH, Cho DH, Oh HM, Kim HS. Algae-bacteria interactions: Evolution, ecology and emerging applications. Biotechnol Adv. 2016;34(1):14–29. 10.1016/j.biotechadv.2015.12.003. Page 17/30 5. Zan J, Li Z, Tianero MD, Davis J, Hill RT, Donia MS. A microbial factory for defensive kahalalides in a tripartite marine symbiosis. Science. 2019;364(6445):eaaw6732. 10.1126/science.aaw6732. . Berthold DE, Shetty KG, Jayachandran K, Laughinghouse HD, Gantar M. Enhancing algal biomass and lipid production through bacterial co-culture. Biomass Bioenergy. 2019;122:280–9. 10.1016/j.biombioe.2019.01.033. 7. Fuentes JL, Garbayo I, Cuaresma M, Montero Z, González-Del-Valle M, Vílchez C. Impact of microalgae-bacteria interactions on the production of algal biomass and associated compounds. Mar Drugs. 2016;14(5):100. 10.3390/md14050100. . Lutzu GA, Dunford NT. Interactions of microalgae and other microorganisms for enhanced production of high-value compounds. Front Biosci (Landmark Ed). 2018;23(8):1487–504. 10.2741/4656. 9. Magdouli S, Brar SK, Blais JF. Co-culture for lipid production: advances and challenges. Biomass Bioenergy. 2016;92:20–30. 10.1016/j.biombioe.2016.06.003. 10. Toyama T, Hanaoka T, Yamada K, Suzuki K, Tanaka Y, Morikawa M, et al. Enhanced production of biomass and lipids by Euglena gracilis via co-culturing with a microalga growth-promoting bacterium, Emticicia sp. EG3. Biotechnol Biofuels. 2019;12:205. 10.1186/s13068-019-1544-2. 11. Nakatsu C, Hutchinson TC. Extreme metal and acid tolerance of Euglena mutabilis and an associated yeast from Smoking Hills, Northwest Territories, and their apparent mutualism. Microb Ecol. 1988;16(2):213–31. 10.1007/BF02018915. 12. Kostygov AY, Karnkowska A, Votýpka J, Tashyreva D, Maciszewski K, Yurchenko V, et al. Euglenozoa: taxonomy, diversity and ecology, symbioses and viruses. Open Biol. 2021;11(3):200407. 10.1098/rsob.200407. 13. Ebenezer TE, Low RS, O’Neill EC, Huang I, DeSimone A, Farrow SC, et al. Euglena International Network (EIN): driving euglenoid biotechnology for the bene t of a challenged world. Biol Open. 2022;11(11):bio059561. 10.1242/bio.059561. 14. Khatiwada B, Hasan MT, Sun A, Kamath KS, Mirzaei M, Sunna A, et al. Probing the role of the chloroplasts in heavy metal tolerance and accumulation in Euglena gracilis. Microorganisms. 2020;8(1):115. 10.3390/microorganisms8010115. 15. Li J, Zheng Z, Du M, Chen J, Zhu H, Hu Z, et al. Euglena gracilis and its aqueous extract constructed with chitosan-hyaluronic acid hydrogel facilitate cutaneous wound healing in mice without inducing excessive in ammatory response. Front Bioeng Biotechnol. 2021;9:713840. 10.3389/fbioe.2021.713840. 1 . Nakashima A, Yamada K, Iwata O, Sugimoto R, Atsuji K, Ogawa T, et al. β-Glucan in foods and its physiological functions. J Nutr Sci Vitaminol (Tokyo). 2018;64(1):8–17. 10.3177/jnsv.64.8. 17. Yamamoto FY, Sutili FJ, Hume M, Gatlin DM III. The effect of β-1,3-glucan derived from Euglena gracilis (Algamune™ ) on the innate immunological responses of Nile tilapia (Oreochromis niloticus L). J Fish Dis. 2018;41(10):1579–88. 10.1111/jfd.12871. Page 18/30 1 . Borst P, Sabatini R, Base J. Discovery, Biosynthesis, and Possible Functions. Annu Rev Microbiol. 2008;62:235–51. 10.1146/annurev.micro.62.081307.162750. 19. O’Neill EC, Trick M, Henrissat B, Field RA. Euglena in time: evolution, control of central metabolic processes and multi-domain proteins in carbohydrate and natural product biochemistry. Perspect Sci. 2015;6:84–93. 10.1016/j.pisc.2015.07.002. 20. Ebenezer TE, Zoltner M, Burrell A, Nenarokova A, Novák Vanclová AMG, Prasad B, et al. Transcriptome, proteome, and draft genome of Euglena gracilis. BMC Biol. 2019;17:11. 10.1186%2Fs12915-019-0626-8. 21. Lukáčová A, Beck T, Koptašiková L, Benda A, Tomečková L, Trniková M, et al. Euglena gracilis can grow in the mixed culture containing Cladosporium westerdijkiae, Lysinibacillus boronitolerans and Pseudobacillus badius without the addition of vitamins B1 and B12. J Biotechnol. 2022;351:50–9. 10.1016/j.jbiotec.2022.04.013. 22. Kim JY, Oh J-J, Jeon MS, Kim G-H, Choi Y-E. Improvement of Euglena gracilis Paramylon Production through a Cocultivation Strategy with the Indole-3-Acetic Acid-Producing Bacterium Vibrio natriegens. Appl Environ Microbiol. 2019;85(19):e01548–19. 10.1128/AEM.01548-19. 23. Sabater S, Buchaca T, Cambra J, Catalan J, Guasch H, Ivorra N, et al. Structure and function of benthic algal communities in an extremely acid river. J Phycol. 2003;39(3):481–9. 10.1046/j.15298817.2003.02104.x. 24. Searles PS, Kropp BR, Flint SD, Caldwell MM. In uence of solar UV-B radiation on peatland microbial communities of southern Argentinia. New Phytol. 2001;152(2):213–21. 10.1046/j.0028646X.2001.00254.x. 25. Casiot C, Bruneel O, Personné JC, Leblanc M, Elbaz-Poulichet F. Arsenic oxidation and bioaccumulation by the acidophilic protozoan, Euglena mutabilis, in acid mine drainage (Carnoulès, France). Sci Total Environ. 2004;320(2–3):259–67. 10.1016/j.scitotenv.2003.08.004. 2 . Gross W, Oesterhelt C, Tischendorf G, Lederer F. Characterization of a non-thermophilic strain of the red algal genus Galdieria isolated from Soos (Czech Republic). Eur J Phycol. 2002;37(3):477–82. 10.1017/S0967026202003773. 27. Sittenfeld A, Mora M, Ortega JM, Albertazzi F, Cordero A, Roncel M, et al. Characterization of a photosynthetic Euglena strain isolated from an acidic hot mud pool of a volcanic area of Costa Rica1. FEMS Microbiol Ecol. 2002;42(1):151–61. 10.1111/j.1574-6941.2002.tb01004.x. 2 . Akcil A, Koldas S. Acid mine drainage (AMD): causes, treatment and case studies. J Clean Prod. 2006;14(12–13):1139–45. 10.1016/j.jclepro.2004.09.006. 29. Halter D, Casiot C, Heipieper HJ, Plewniak F, Marchal M, Simon S, et al. Surface properties and intracellular speciation revealed an original adaptive mechanism to arsenic in the acid mine drainage bio-indicator Euglena mutabilis. Appl Microbiol Biotechnol. 2012;93(4):1735–44. 10.1007/s00253011-3493-y. 30. Halter D, Goulhen-Chollet F, Gallien S, Casiot C, Hamelin J, Gilard F, et al. In situ proteo-metabolomics reveals metabolite secretion by the acid mine drainage bio-indicator, Euglena mutabilis. ISME J. Page 19/30 2012;6(7):1391–402. 10.1038/ismej.2011.198. 31. Valente T, Gomes C. The role of two acidophilic algae as ecological indicators of acid mine drainage sites. J Iber Geol. 2007;33:283–94. 10.5209/JIGE.33916. 32. Brake SS, Hasiotis ST, Dannelly HK. Diatoms in acid mine drainage and their role in the formation of iron-rich stromatolites. Geomicrobiol J. 2004;21(5):331–40. 10.1080/01490450490454074. 33. Ňancucheo I, Barrie Johnson D. Acidophilic algae isolated from mine-impacted environments and their roles in sustaining heterotrophic acidophiles. Front Microbiol. 2012;3:325. 10.3389/fmicb.2012.00325. 34. Yanagawa K, Haraguchi A, Yoshitake K, Asamatsu K, Harano M, Yamashita K, et al. Ubiquity of Euglena mutabilis population in three ecologically distinct acidic habitats in southwestern Japan. Water. 2021;13(11):1570. 10.3390/w13111570. 35. Prigent S, Frioux C, Dittami SM, Thiele S, Larhlimi A, Collet G, et al. Meneco, a topology-based gaplling tool applicable to degraded genome-wide metabolic networks. PLOS Comput Biol. 2017;13(1):e1005276. 10.1371/journal.pcbi.1005276. 3 . Olaveson MM, Stokes PM. Reponses of the acidiphilic alga Euglena mutablilis (Euglenophyceae) to carbon Eerichent at pH 3. J Phycol. 1989;25(3):529–39. 10.1111/j.1529-8817.1989.tb00259.x. 37. Canadian Phycological Culture Centre. : Modi ed acid medium (MAM). https://uwaterloo.ca/canadian-phycological-culture-centre/cultures/culture-media/modi ed-acidmedium (2017). Accessed 14 Sep 2020. 3 . Warren CR. Rapid Measurement of Chlorophylls with a Microplate Reader. J Plant Nut. 2008;31(7):1321–32. 10.1080/01904160802135092. 39. Comeau AM, Douglas GM, Langille MGI. Microbiome helper: a custom and streamlined work ow for microbiome research. mSystems. 2017;2(1):e00127–16. 10.1128/mSystems.00127-16. 40. Bolyen E, Rideout JR, Dillon MR, Bokulich NA, Abnet CC, Al-Ghalith GA, et al. Reproducible, interactive, scalable and extensible microbiome data science using QIIME 2. Nat Biotechnol. 2019;37(8):852–7. 10.1038/s41587-019-0209-9. 41. Callahan BJ, McMurdie PJ, Rosen MJ, Han AW, Johnson AJA, Holmes SP. DADA2: high-resolution sample inference from Illumina amplicon data. Nat Methods. 2016;13(7):581–3. 10.1038/nmeth.3869. 42. Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2012;41(D1):D590–6. 10.1093/nar/gks1219. 43. Bokulich NA, Kaehler BD, Rideout JR, Dillon M, Bolyen E, Knight R, et al. Optimizing taxonomic classi cation of marker-gene amplicon sequences with QIIME 2’s q2-feature-classi er plugin. Microbiome. 2018;6(1):90. 10.1186/s40168-018-0470-z. 44. McMurdie PJ, Holmes S. phyloseq: an R package for reproducible interactive analysis and graphics of microbiome census data. PLoS ONE. 2013;8(4):e61217. 10.1371/journal.pone.0061217. Page 20/30 45. Raja HA, Miller AN, Pearce CJ, Oberlies NH. Fungal identi cation using molecular tools: a primer for the natural products research community. J Nat Prod. 2017;80(3):756–70. 10.1021/acs.jnatprod.6b01085. 4 . Thijs S, De Beeck MO, Beckers B, Truyens S, Stevens V, Van Hamme JD, et al. Comparative evaluation of four bacteria-speci c primer pairs for 16S rRNA gene surveys. Front Microbiol. 2017;8:494. 10.3389/fmicb.2017.00494. 47. He J, Liu C, Du M, Zhou X, Hu Z, Lei A, et al. Metabolic responses of a model green microalga Euglena gracilis to different environmental stresses. Front Bioeng Biotechnol. 2021;9:662655. 10.3389/fbioe.2021.662655. 4 . Shao Q, Jiang Y, Lei A, Hu Z, Wang J. Research progress on genetic transformation technology of Euglena. Acta Hydrobiol Sin. 2018;42(4):655–62. 10.7541/2018.081. 49. Ebringer L, Mego JL, Jurášek A. Mitomycins and the bleaching of Euglena gracilis. Arch Mikrobiol. 1969;64(3):229–34. 10.1007/BF00425628. 50. Ebringer L, Nemec P, Santová H, Foltínová P. Changes of the plastid system of Euglena gracilis induced with streptomycin and dihydrostreptomycin. Arch Mikrobiol. 1970;73(3):268–80. 10.1007/BF00410628. 51. Zahalsky AC, Hutner SH, Keane M, Burger RM. Bleaching Euglena gracilis with antihistamines and streptomycin-type antibiotics. Arch Mikrobiol. 1962;42:46–55. 10.1007/BF00425189. 52. Ebringer L, Krajčovič J, Polónyi J, Lahitová N, Doupovcová M, Dobias J. Tetracycline reduces uoroquinolones-induced bleaching of Euglena gracilis. Mutat Res. 1996;340(2–3):141–9. 10.1016/S0165-1110(96)90045-7. 53. Gumińska N, Płecha M, Walkiewicz H, Hałakuc P, Zakryś B, Milanowski R. Culture puri cation and DNA extraction procedures suitable for next-generation sequencing of euglenids. J Appl Phycol. 2018;30:3541–9. 10.1007/s10811-018-1496-0. 54. Bishop DG, Smillie RM. The effect of chloramphenicol and cycloheximide on lipid synthesis during chloroplast development in Euglena gracilis. Arch Biochem Biophys. 1970;139(1):179–89. 10.1016/0003-9861(70)90059-7. 55. Kennedy V, Kaszecki E, Donaldson ME, Saville BJ. The impact of elevated sulfur and nitrogen levels on cadmium tolerance in Euglena species. 2023. [Manuscript submitted for publication]. 5 . Boivin MEY, Greve GD, García-Meza JV, Massieux B, Sprenger W, Kraak MHS, et al. Algal–bacterial interactions in metal contaminated oodplain sediments. Environ Pollut. 2007;145(3):884–94. 10.1016/j.envpol.2006.05.003. 57. dos Santos VZ, Vieira KR, Nass PP, Zepka LQ, Jacob-Lopes E. Application of microalgae consortia/cocultures in wastewater treatment. In: Ahamed I, Prasad MI R, editors. Recent advances in microbial degradation. Environmental and Microbial Biotechnology. Singapore: Springer; 2021. pp. 131–54. 10.1007/978-981-16-0518-5_5. 5 . Wang J, Chen R, Fan L, Cui L, Zhang Y, Cheng J, et al. Construction of fungi-microalgae symbiotic system and adsorption study of heavy metal ions. Sep Purif Technol. 2021;268(1):118689. Page 21/30 10.1016/j.seppur.2021.118689. 59. Li B, Zhang T, Yang Z. Immobilizing unicellular microalga on pellet-forming lamentous fungus: can this provide new insights into the remediation of arsenic from contaminated water? Bioresour Technol. 2019;284:231–9. 10.1016/j.biortech.2019.03.128. 0. Hlabela P, Maree J, Bruinsma D. Barium carbonate process for sulphate and metal removal from mine water. Mine Water Environ. 2007;26:14–22. 10.1007/s10230-007-0145-7. 1. Skousen JG, Sexstone A, Ziemkiewicz PF. Acid mine drainage control and treatment. In: Barnhisel RI, Darmody RG, Daniels WL American Soceity of Agronomy, Crop Science Society of America, Soil Science Society of America, editors. Reclamation of Drastically Disturbed Lands. Wisconsin: American Society of Agronomy; 2000. p. 131–68. 10.2134/agronmonogr41.c6. 2. Yang W, Tang Q, Wei J, Ran Y, Chai L, Wang H. Enhanced removal of Cd(II) and Pb(II) by composites of mesoporous carbon stabilized alumina. Appl Surf Sci. 2016;369:215–23. 10.1016/j.apsusc.2016.01.151. 3. Das PK, Rani J, Rawat S, Kumar S. Microalgal co-cultivation for biofuel production and bioremediation: current status and bene ts. BioEnergy Res. 2022;15(1):1–26. 10.1007/s12155-02110254-8/Published. 4. Hassan Zul, Ali S, Rizwan M, Ibrahim M, Nafees M, Waseem M. Role of bioremediation agents (bacteria, fungi, and algae) in alleviating heavy metal toxicity. In: Kumar V, Kumar M, Sharma S, Prasad R, editors. Probiotics in Agroecosystem. Singapore: Springer; 2017. pp. 517–37. 5. Higgins BT, Gennity I, Fitzgerald PS, Ceballos SJ, Fiehn O, VanderGheynst JS. Algal–bacterial synergy in treatment of winery wastewater. Npj Clean Water. 2018;1:6. 10.1038/s41545-018-0005-y. . Nazari MT, Freitag JF, Cavanhi VAF, Colla LM. Microalgae harvesting by fungal-assisted bio occulation. Rev Environ Sci Biotechnol. 2020;19:369–88. 10.1007/s11157-020-09528-y. 7. Pei XY, Ren HY, Liu BF. Flocculation performance and mechanism of fungal pellets on harvesting of microalgal biomass. Bioresour Technol. 2021;321:124463. 10.1016/J.BIORTECH.2020.124463. . Rehman A. Heavy metals uptake by Euglena proxima isolated from tannery e uents and its potential use in wastewater treatment. Russ J Ecol. 2011;42:44–9. 10.1134/S1067413611010085. 9. Salgueiro JL, Pérez L, Maceiras R, Sánchez A, Cancela A. Bioremediation of wastewater using Chlorella Vulgaris microalgae: phosphorus and organic matter. Int J Environ Res. 2016;10(3):465–70. 70. Waluyo L, Prihanta W, Bachtiar Z, Permana TI. Potential bioremediation of lead (Pb) using marine microalgae Nannochloropsis oculata. AIP Conf Proc. 2020;2231:040088. 10.1063/5.0002441. 71. Znad H, Al Ketife AMD, Judd S, AlMomani F, Vuthaluru HB. Bioremediation and nutrient removal from wastewater by Chlorella vulgaris. Ecol Eng. 2018;110:1–7. 10.1016/j.ecoleng.2017.10.008. 72. Ayangbenro AS, Babalola OO, Aremu OS. Bio occulant production and heavy metal sorption by metal resistant bacterial isolates from gold mining soil. Chemosphere. 2019;231:113–20. 10.1016/j.chemosphere.2019.05.092. Page 22/30 73. Lal A, Banerjee S, Das D. Aspergillus sp. assisted bio occulation of Chlorella MJ 11/11 for the production of biofuel from the algal-fungal co-pellet. Sep Purif Technol. 2021;272:118320. 10.1016/J.SEPPUR.2021.118320. 74. Mujtaba G, Rizwan M, Lee K. Simultaneous removal of inorganic nutrients and organic carbon by symbiotic co-culture of Chlorella vulgaris and Pseudomonas putida. Biotechnol Bioprocess Eng. 2015;20(6):1114–22. 10.1007/s12257-015-0421-5. 75. Zhang Z, Nair S, Tang L, Zhao H, Hu Z, Chen M, et al. Long-term survival of Synechococcus and heterotrophic bacteria without external nutrient supply after changes in their relationship from antagonism to mutualism. mBio. 2021;12(4):e0161421. 10.1128/mBio.01614-21. 7 . Rubiyatno MK, Inoue D, Kim S, Yu J, Lee T, et al. Isolation and characterization of Euglena gracilisassociated bacteria, Enterobacter sp. CA3 and Emticicia sp. CN5, capable of promoting the growth and paramylon production of E. gracilis under mixotrophic cultivation. Microorganisms. 2021;9(7):1496. 10.3390/microorganisms9071496. 77. Krespach MKC, García-Altares M, Flak M, Hanno Schoeler, Scherlach K, Netzker T, et al. Lichen-like association of Chlamydomonas reinhardtii and Aspergillus nidulans protects algal cells from bacteria. ISME J. 2020;14(11):2794–805. 10.1038/s41396-020-0731-2. 7 . Kazamia E, Czesnick H, Nguyen TTV, Croft MT, Sherwood E, Sasso S, et al. Mutualistic interactions between vitamin B12-dependent algae and heterotrophic bacteria exhibit regulation. Environ Microbiol. 2012;14(6):1466–76. 10.1111/j.1462-2920.2012.02733.x. 79. Liba CM, Ferrara FIS, Man o GP, Fantinatti-Garboggini F, Albuquerque RC, Pavan C, et al. Nitrogenxing chemo-organotrophic bacteria isolated from cyanobacteria-deprived lichens and their ability to solubilize phosphate and to release amino acids and phytohormones. J Appl Microbiol. 2006;101(5):1076–86. 10.1111/j.1365-2672.2006.03010.x. 0. Amin SA, Hmelo LR, Van Tol HM, Durham BP, Carlson LT, Heal KR, et al. Interaction and signalling between a cosmopolitan phytoplankton and associated bacteria. Nature. 2015;522(7554):98–101. 10.1038/nature14488. 1. Seymour JR, Amin SA, Raina JB, Stocker R. Zooming in on the phycosphere: The ecological interface for phytoplankton-bacteria relationships. Nat Microbiol. 2017;2:17065. 10.1038/nmicrobiol.2017.65. 2. Brussaard C, Riegman R. In uence of bacteria on phytoplankton cell mortality with phosphorus or nitrogen as the algal-growth-limiting nutrient. Aquat Microb Ecol. 1998;14:271–80. 3. Zhou Y, Li X, Chen J, Li W, Wang F. Carbon sequestration performance, enzyme and photosynthetic activity, and transcriptome analysis of algae-bacteria symbiotic system after antibiotic exposure. Sci Total Environ. 2023;902:166486. 10.1016/j.scitotenv.2023.166486. 4. Kranner I, Cram WJ, Zorn M, Wornik S, Yoshimura I, Stabentheiner E, et al. Antioxidants and photoprotection in a lichen as compared with its isolated symbiotic partners. Proc Natl Acad Sci. 2005;102(8):3141–6. 10.1073/pnas.0407716102. 5. Grube M, Wedin M. Lichenized fungi and the evolution of symbiotic organization. Microbiol Spectr. 2016;4(6). 10.1128/microbiolspec.FUNK-0011-2016. Page 23/30 . Wang J, Tian Q, Cui L, Cheng J, Zhou H, Peng A, et al. Effect of extracellular proteins on Cd(II) adsorption in fungus and algae symbiotic system. J Environ Manage. 2022;323:116173. 10.1016/j.jenvman.2022.116173. 7. Shen L, Li Z, Wang J, Liu A, Li Z, Yu R, et al. Characterization of extracellular polysaccharide/protein contents during the adsorption of Cd(II) by Synechocystis sp. PCC6803. Environ Sci Pollut Res. 2018;25(21):20713–22. 10.1007/s11356-018-2163-3. . Park D, Yun Y-S, Hye Jo J, Park JM. Mechanism of hexavalent chromium removal by dead fungal biomass of Aspergillus niger. Water Res. 2005;39(4):533–40. 10.1016/j.watres.2004.11.002. 9. Trevors JT, Stratton GW, Gadd GM. Cadmium transport, resistance, and toxicity in bacteria, algae, and fungi. Can J Microbiol. 1986;32(6):447–64. 10.1139/m86-085. 90. Jamers A, Blust R, De Coen W, Gri n JL, Jones OAH. An omics based assessment of cadmium toxicity in the green alga Chlamydomonas reinhardtii. Aquat Toxicol. 2013;126:355–64. 10.1016/j.aquatox.2012.09.007. 91. Choppala G, Saifullah, Bolan N, Bibi S, Iqbal M, Rengel Z, et al. Cellular mechanisms in higher plants governing tolerance to cadmium toxicity. Crit Rev Plant Sci. 2014;33(5):374–91. 10.1080/07352689.2014.903747. 92. Du J, Qiu B, Pedrosa Gomes M, Juneau P, Dai G. In uence of light intensity on cadmium uptake and toxicity in the cyanobacteria Synechocystis sp. PCC6803. Aquat Toxicol. 2019;211:163–72. 10.1016/j.aquatox.2019.03.016. 93. Andresen E, Küpper H. Cadmium toxicity in plants. Met Ions Life Sci. 2013;11:395–413. 10.1007/978-94-007-5179-8_13. 94. Grajek H, Rydzyński D, Piotrowicz-Cieślak A, Herman A, Maciejczyk M, Wieczorek Z. Cadmium ionchlorophyll interaction – examination of spectral properties and structure of the cadmiumchlorophyll complex and their relevance to photosynthesis inhibition. Chemosphere. 2020;261:127434. 10.1016/j.chemosphere.2020.127434. 95. Zhou W, Qiu B. Effects of cadmium hyperaccumulation on physiological characteristics of Sedum alfredii Hance (Crassulaceae). Plant Sci. 2005;169(4):737–45. 10.1016/j.plantsci.2005.05.030. 9 . Ishfaq M, Wang Y, Yan M, Wang Z, Wu L, Li C, et al. Physiological essence of magnesium in plants and its widespread de ciency in the farming system of China. Front Plant Sci. 2022;13:802274. 10.3389/fpls.2022.802274. 97. Jiang Y, Zhu Y, Hu Z, Lei A, Wang J. Towards elucidation of the toxic mechanism of copper on the model green alga Chlamydomonas reinhardtii. Ecotoxicology. 2016;25(7):1417–25. 10.1007/s10646-016-1692-0. 9 . Scott NS. Precursors of chloroplast ribosomal RNA in Euglena gracilis. Phytochemistry. 1976;15(8):1207–13. 10.1016/0031-9422(76)85079-0. 99. Ebringer L. Are plastids derived from prokaryotic micro-organisms? action of antibiotics on chloroplasts of Euglena gracilis. J Gen Microbiol. 1972;71(1):35–52. 10.1099/00221287-71-1-35. Page 24/30 100. Fu Y, Zhu Y, Dong H, Li J, Zhang W, Shao Y, et al. Effects of heavy metals and antibiotics on antibiotic resistance genes and microbial communities in soil. Process Saf Environ Prot. 2023;169:418–27. 10.1016/j.psep.2022.11.020. 101. Khurana P, Pulicharla R, Kaur Brar S. Antibiotic-metal complexes in wastewaters: fate and treatment trajectory. Environ Int. 2021;157:106863. 10.1016/j.envint.2021.106863. 102. Grenson M. Physiology and cytology of chloroplast formation and loss in Euglena. Int Rev Cytol. 1964;37–59. 10.1016/s0074-7696(08)60293-4. 103. Khatiwada B, Hasan MT, Sun A, Kamath KS, Mirzaei M, Sunna A, et al. Proteomic response of Euglena gracilis to heavy metal exposure – identi cation of key proteins involved in heavy metal tolerance and accumulation. Algal Res. 2020;45:101764. 10.1016/j.algal.2019.101764. 104. Gultom S, Hu B. Review of microalgae harvesting via co-pelletization with lamentous fungus. Energies. 2013;6(11):5921–39. 10.3390/en6115921. 105. Wyatt KH, Seballos RC, Shoemaker MN, Brown SP, Chandra S, Kuehn KA, et al. Resource constraints highlight complex microbial interactions during lake bio lm development. J Ecol. 2019;107(6):2737– 46. 10.1111/1365-2745.13223. 10 . Bhattacharya A, Malik A, Malik HK. A mathematical model to describe the fungal assisted algal occulation process. Bioresour Technol. 2017;244(Pt 1):975–81. 10.1016/J.BIORTECH.2017.08.062. 107. Bans eld D, Spilling K, Mikola A, Piiparinen J. Bio occulation of Euglena gracilis via direct application of fungal laments: a rapid harvesting method. J Appl Phycol. 2022;34:321–34. 10.1007/s10811-021-02651-5. 10 . Morales-Oyervides L, Ruiz-Sánchez JP, Oliveira JC, Sousa-Gallagher MJ, Méndez-Zavala A, Giuffrida D, et al. Biotechnological approaches for the production of natural colorants by Talaromyces/Penicillium: Aareview. Biotechnol Adv. 2020;43:107601. 10.1016/j.biotechadv.2020.107601. 109. Kennes C, Veiga MC. Fungal biocatalysts in the bio ltration of VOC-polluted air. J Biotechnol. 2004;113(1–3):305–19. 10.1016/j.jbiotec.2004.04.037. 110. Tokuhisa Y, Hagiya Y, Hiruma M, Nishimura K. Phaeohyphomycosis of the face caused by Exophiala oligosperma: facial phaeohyphomycosis caused by E. oligosperma. Mycoses. 2011;54(4):e240–3. 10.1111/j.1439-0507.2009.01845.x. 111. Yilmaz N, Houbraken J, Hoekstra ES, Frisvad JC, Visagie CM, Samson RA. Delimitation and characterisation of Talaromyces purpurogenus and related species. Persoonia. 2012;29:39–54. 10.3767/003158512X659500. 112. Koolen HHF, Menezes LS, Souza MP, Silva FMA, Almeida FGO, Souza AQLD, et al. Talaroxanthone, a novel xanthone dimer from the endophytic fungus Talaromyces sp. associated with Duguetia stelechantha (Diels) R. E. Fries. J Braz Chem Soc. 2013;24(5):880–3. 10.5935/0103-5053.20130104. 113. Priyanka DSK. Hexavalent chromium (Cr + 6) detoxi cation potential and stress response of fungus Talaromyces pinophillus isolated from tannery wastewater. Environ Adv. 2023;13:100417. 10.1016/j.envadv.2023.100417. Page 25/30 114. Guerra Sierra BE, Arteaga-Figueroa LA, Sierra-Pelaéz S, Alvarez JC. Talaromyces santanderensis: a new cadmium-tolerant fungus from cacao soils in Colombia. J Fungi. 2022;8(10):1042. 10.3390/jof8101042. 115. Nam I-H, Murugesan K, Ryu J, Kim JH. Arsenic (As) removal using Talaromyces sp. KM-31 isolated from As-contaminated mine soil. Minerals. 2019;9(10):568. 10.3390/min9100568. 11 . Ajimura M, Leem SH, Ogawa H. Identi cation of new genes required for meiotic recombination in Saccharomyces cerevisiae. Genetics. 1993;133(1):51–66. 10.1093/genetics/133.1.51. 117. Vyas VK, Barrasa MI, Fink GR. A Candida albicans CRISPR system permits genetic engineering of essential genes and gene families. Sci Adv. 2015;1(3):e1500248. 10.1126/sciadv.1500248. 11 . Parry JM, Zimmerman FK. The detection of monosomic colonies produced by mitotic chromosome non-disjunction in the yeast Saccharomyces cerevisiae. Mutat Res. 1976;36(1):49–66. 10.1016/0027-5107(76)90020-8. 119. Granot D, Snyder M. Carbon source induces growth of stationary phase yeast cells, independent of carbon source metabolism. Yeast. 1993;9(5):465–79. 10.1002/yea.320090503. 120. Mano Y, Kobayashi TJ, Nakayama J, Uchida H, Oki M. Single cell visualization of yeast gene expression shows correlation of epigenetic switching between multiple heterochromatic regions through multiple generations. PLoS Biol. 2013;11(7):e1001601. 10.1371/journal.pbio.1001601. 121. Tam E, Tsang C-C, Lau S, Woo P. Polyketides, toxins and pigments in Penicillium marneffei. Toxins (Basel). 2015;7(11):4421–36. 10.3390/toxins7114421. 122. Woo PCY, Lam C-W, Tam EWT, Leung CKF, Wong SSY, Lau SKP, et al. First discovery of two polyketide synthase genes for mitorubrinic acid and mitorubrinol yellow pigment biosynthesis and implications in virulence of Penicillium marneffei. PLoS Negl Trop Dis. 2012;6(10):e1871. 10.1371/journal.pntd.0001871. 123. Wang F, Han R, Chen S. An overlooked and underrated endemic mycosis—Talaromycosis and the pathogenic fungus Talaromyces marneffei. Clin Microbiol Rev. 2023;36(1):e00051–22. 10.1128/cmr.00051-22. 124. Xiao Y, Chen R, Chen L, Yang B, Jiang L, Fang J. Endophytic fungus Talaromyces sp. MR1 promotes the growth and cadmium uptake of Arabidopsis thaliana L. under cadmium stress. Curr Microbiol. 2023;80(11):346. 10.1007/s00284-023-03453-3s. 125. Emery RJN, Longnecker NE, Atkins CA. Branch development in Lupinus angustifolius L. II. relationship with endogenous ABA, IAA and cytokinins in axillary and main stem buds. J Exp Bot. 1998;49(320):555–62. 10.1093/jxb/49.320.555. 12 . Yang G-L, Zheng M-M, Tan A-J, Liu Y-T, Feng D, Lv S-M. Research on the mechanisms of plant enrichment and detoxi cation of cadmium. Biology (Basel). 2021;10(6):544. 10.3390/biology10060544. 127. Velivelli SLS, Sessitsch A, Prestwich BD. The role of microbial inoculants in integrated crop management systems. Potato Res. 2014;57(3–4):291–309. 10.1007/s11540-014-9278-9. Page 26/30 12 . Sudová R, Vosátka M. Differences in the effects of three arbuscular mycorrhizal fungal strains on P and Pb accumulation by maize plants. Plant Soil. 2007;296(1–2):77–83. 10.1007/s11104-007-92918. 129. Du Z-Y, Zienkiewicz K, Pol NV, Ostrom NE, Benning C, Bonito GM. Algal-fungal symbiosis leads to photosynthetic mycelium. eLife. 2019;8:e47815. 10.7554/eLife.47815.001. 130. Xu L, Gu W, Bai J. Thiobacillus acidophilus removes lead and copper from wastewater containing low concentration of heavy metals. IOP Conf Ser Earth Environ Sci. 2021;769:022039. 10.1088/17551315/769/2/022039. 131. Yang W, Song W, Li J, Zhang X. Bioleaching of heavy metals from wastewater sludge with the aim of land application. Chemosphere. 2020;249:126134. 10.1016/j.chemosphere.2020.126134. 132. Li L, Liu Z, Zhang M, Meng D, Liu X, Wang P, et al. Insights into the metabolism and evolution of the genus Acidiphilium, a typical acidophile in acid mine drainage. mSystems. 2020;5(6):e00867–20. 10.1128/mSystems.00867-20. 133. Rawlings DE. Heavy metal mining using microbes. Annu Rev Microbiol. 2002;56:65–91. 10.1146/annurev.micro.56.012302.161052. 134. Priya A, Hait S. Biometallurgical recovery of metals from waste printed circuit boards using pure and mixed strains of Acidithiobacillus ferrooxidans and Acidiphilium acidophilum. Process Saf Environ Prot. 2020;143:262–72. 10.1016/j.psep.2020.06.042. 135. Johnson DB, Bridge TAM. Reduction of ferric iron by acidophilic heterotrophic bacteria: evidence for constitutive and inducible enzyme systems in Acidiphilium spp. J Appl Microbiol. 2002;92(2):315– 21. 10.1046/j.1365-2672.2002.01535.x. 13 . Liu H, Yin H, Dai Y, Dai Z, Liu Y, Li Q, et al. The co-culture of Acidithiobacillus ferrooxidans and Acidiphilium acidophilum enhances the growth, iron oxidation, and CO2 xation. Arch Microbiol. 2011;193(12):857–66. 10.1007/s00203-011-0723-8. Figures Page 27/30 Figure 1 Antibiotic and CdCl2 treatment reveal differences in cell viability between E. mutabilis and E. gracilis. Cultures were grown in MAM for 72 hours. Light coloured bars indicate the inclusion of a dilution series of antibiotic alone while darker colour bars indicate exposure to antibiotic and a xed concentration,100 μM, of CdCl2. Colours indicating different antibiotic concentrations are indicated in the legend at the bottom of the gure. Error bars represent standard deviation (n = 3). A star (*) indicates statistical difference (tPage 28/30 test) between cells exposed to antibiotics and cells exposed to the same concentration of antibiotics with the addition of 100 μM CdCl2 (* = p < 0.05, ** = p < 0.01, *** = p < 0.001). A dagger (฀) indicates statistical difference (t-test) between control cells exposed to 100 μM CdCl2 and those exposed to antibiotics and CdCl2 (฀ = p < 0.05, ฀฀ = p < 0.01, ฀฀฀ = p < 0.001). Figure 2 Page 29/30 Cadmium exposure increases chlorophyll content of Euglena cultures. Total chlorophyll content (chl a + chl b) per 100,000 Euglena cells was determined after 72-hours of exposure to varying concentrations of antibiotics (light coloured bars), and varying concentrations of antibiotics plus 100 μM CdCl2 (dark coloured bars). Error bars represent standard deviation (n = 3). Statistical difference between total chlorophyll content in control and treatments conditions was assessed using a t-test, and signi cant differences are denoted by a star (* = p < 0.05, ** = p < 0.01, *** = p < 0.001). Figure 3 PacBio sequencing identi es a major bacterial and a major fungal partner for E. mutabilis. Relative abundance taxa bar plot of E. mutabilis co-culture (CPCC 657) grown in MAM at pH 4.3 (A samples), MAM at pH 2.7 (B samples), TSB grown in light (C samples), and TSB grown in dark (D samples). Taxa (targeting the full 16S region of bacterial rDNA and full ITS region of fungal rDNA) are represented at the species level where possible. Supplementary Files This is a list of supplementary les associated with this preprint. Click to download. EKaszeckiAntibitoticsSupplementaryv1.1.docx Page 30/30