ABSTRACT
Pollution of aquatic environments is a concern and poses a health risk to aquatic and terrestrial life. Plastic pollutants act as surfaces to which microorganisms adhere, forming a community known as the plastisphere. Studies investigating the bacterial diversity of the plastisphere are plentiful, but less is known about the fungal communities, especially yeasts, within these biofilms. Given the increasingly recognised threat of yeast infections, it is important to investigate the environmental presence of potential pathogenic yeasts on possible vehicles that may contribute to their distribution. In this study, the fungal diversity of the plastisphere of various types of plastic polymers collected from an urban freshwater source was analysed, and possibly pathogenic yeast strains were identified. Isolates were also tested for susceptibility to the antifungal, fluconazole. Microscopy sheds light on the overall diversity found in the plastisphere, while metagenomic data revealed that fungal plastisphere communities are dominated by the phylum Ascomycota. Both metagenomic and culture-dependant analysis revealed the presence of possibly pathogenic yeast in the genera Candida (some with low fluconazole susceptibility) and Exophiala. This highlights the possibility that the plastisphere may harbour pathogenic yeasts and could contribute to their distribution in the environment and transmission between the environment and humans.
HIGHLIGHTS
Plastisphere harbours a diverse community of microorganisms.
The Ascomycota and Basidiomycota phyla dominate the fungal community of the plastisphere.
Pathogenic yeast from the Candida genus was frequently isolated from polymer samples.
Candida glabrata shows a varying susceptibility towards fluconazole.
Plastic pollutants act as a vehicle in the spread of various pathogens.
INTRODUCTION
Plastic is a cheap, lightweight, and durable (Laist 1987) polymer that is widely used for the manufacturing of a variety of products (Derraik 2002) from disposable packaging to medical equipment, such as catheters (Andrady & Neal 2009). Plastics Europe (2021) estimated that the global plastic product production in 2020 was 367 million tonnes in comparison to 1.5 million tonnes in the 1950s. Due to the dramatic increase in global plastic production, plastic pollutants that vary in size have become ubiquitous in freshwater and marine environments (Do Sul & Costa 2014). Polypropylene (PP), polyvinyl chloride (PVC), high-density polyethylene (PE), low-density polyethylene (LDPE), polystyrene (PS), and polyethylene terephthalate (PET) collectively make up 90% of overall plastic product production (Andrady & Neal 2009) and are thus common pollutants found in the environment. These compounds are treated with a variety of chemicals or additives to enhance durability and resistance to corrosion. This contributes to other characteristics of plastics, namely, the ability to be transported over great distances (Parthasarathy et al. 2019; Li et al. 2020; Metcalf et al. 2023).
The presence of plastic pollutants can pose a health risk to aquatic mammals due to ingestion, subsequent starvation, and strangulation, which may, in turn, reduce food security in various communities (De Tender et al. 2015; Guzzetti et al. 2018). In addition, plastic pollutants can act as a surface to which microorganisms can adhere. This microbial community or biofilm interacts with the plastic surface and is collectively referred to as the plastisphere (Zettler et al. 2013). It is suggested that the plastisphere provides protection to the plastisphere community as they move between different environmental matrices to areas that pose high risk to human health (Metcalf et al. 2023).
Studies that investigate the plastisphere tend to focus on bacterial diversity as well as the variations between the microbial diversity of the surrounding water versus the plastisphere or the differences in microbial diversity of the plastisphere in different geographical locations or seasonal times (Oberbeckmann et al. 2014; De Tender et al. 2015). Very little research is dedicated to the fungal diversity that forms part of the plastisphere (Douglas 2003; Du et al. 2022). However, in the few studies investigating the fungal diversity of the freshwater plastisphere (Zettler et al. 2013; Kettner et al. 2019; Lacerda et al. 2020), little is reported on the presence of pathogenic fungi or yeasts. Akinbobola et al. (2023) confirmed that Candida auris can proliferate on glass and plastic surfaces submerged in river and sea water sparking interest in the potential role that plastic vectors may play in increasing exposure to pathogenic yeast. However, the presence of yeast in freshwater sources has been investigated and common yeast genera found are Aureobasidium, Candida, Cryptococcus, Debaryomyces, Geotrichum, Hansenula, Kloekera, Kodamaea, Metschinikowia, Meyerozyma, Pichia, Rhodotorula, and Zygosaccharomyces (Hagler 2006; Ayanbimpe et al. 2012; Medeiros et al. 2012).
From these studies, the genera Candida, Cryptococcus, and Rhodotorula are known to harbour major opportunistic pathogens implicated in invasive fungal infections. Since pathogenic yeast does occur in various water sources, and their presence in the plastisphere is expected. Candida spp. are known to form biofilms on medical implants and equipment like catheters and are causative agents for nosocomial infections (Douglas 2002, 2003; Kojic & Darouiche 2004). The materials used for medical equipment suspected to be involved in the origin of nosocomial infections are made up of the same polymers contained in plastic pollutants, including PET, PP, PVC (Hawser & Douglas 1994; Douglas 2003), and PS.
The increase in the occurrence of antifungal-resistant yeast species is of major concern considering that there are limited antifungals on the market, thus increasing the difficulty in effective therapeutic intervention in patients with invasive fungal infections. For instance, C. auris has already shown resistance to multiple antifungals including the three most commonly used antifungals (CDC 2019). Other yeasts, often displaying multidrug resistance, which are frequently isolated from water sources include Candida glabrata, Candida tropicalis, as well as the potential multidrug-resistant Candida krusei (Pfaller et al. 2012; Zuza-Alves et al. 2017; Jamiu et al. 2020). Antifungal resistance has also been attributed to the use of these agents in the environment/agriculture as fungicides (Snelders et al. 2008; Sanglard 2016). This has prompted the World Health Organisation to publish the first Fungal Pathogen Priority List in 2022 (WHO 2022).
The present study aims to investigate the fungal diversity of the plastisphere, with a focus on pathogenic yeasts, found on plastic pollutants isolated from urban freshwater sources, in Bloemfontein, South Africa, as a proof of concept. The diversity was investigated using microscopy, culture-dependent, and culture-independent techniques. Furthermore, the susceptibility of the isolated strains towards the antifungal, fluconazole, was also investigated. All of this was done with the aim of determining if plastic pollutants may act as reservoirs and vehicles of potentially pathogenic fungi.
METHODS
Sample collection and preparation
Five different types of macro plastics were collected from various locations along Bloemspruit, Bloemfontein, South Africa. The location and the type of polymer of each sample are listed in Table 1. From each sample, three random pieces (2 × 2 cm) of plastic were cut out: one piece was frozen at −80 °C before genomic DNA extraction, one piece was used for scanning electron microscopy (SEM), while the last piece was used for isolation of yeast strains.
Sampling locations and types of macro plastic collected
Sample number . | Location in Bloemfontein . | Type of plastic . |
---|---|---|
1 | Loch Logan 29°06′55.9″S 26°12'32.4″E | Transparent PE polymer bag |
2 | Linquinda 29°07'10.5″S 26°14'15.4″E | Black LDPE polymer bag |
3 | Linquinda 29°07'10.4″S 26°14'15.7″E | PS food container |
4 | Oos-Einde 29°07'36.4″S 26°15'07.3″E | Yellow PP polymer food packaging |
5 | Oos-Einde 29°07'36.3″S 26°15'07.1″E | Transparent PET polymer bottle |
Sample number . | Location in Bloemfontein . | Type of plastic . |
---|---|---|
1 | Loch Logan 29°06′55.9″S 26°12'32.4″E | Transparent PE polymer bag |
2 | Linquinda 29°07'10.5″S 26°14'15.4″E | Black LDPE polymer bag |
3 | Linquinda 29°07'10.4″S 26°14'15.7″E | PS food container |
4 | Oos-Einde 29°07'36.4″S 26°15'07.3″E | Yellow PP polymer food packaging |
5 | Oos-Einde 29°07'36.3″S 26°15'07.1″E | Transparent PET polymer bottle |
Scanning electron microscopy of biofilms
Plastic samples were fixed in 0.1 M (pH 7.0) sodium phosphate-buffered glutardialdehyde (3%) for 3 h followed by dehydration in a graded ethanol series (50, 70, and 95%) for 20 min in each phase. This was followed by two changes of 100% for 1 h in each phase change. Samples were then critical point dried (Tousimis Samdri-795 critical point dryer), whereafter they were mounted on aluminium pin stubs with double-sided carbon tape and coated with iridium (Ir, ±5 nm) (Leica EM ACE600 coater) for conductivity. Specimens were imaged using a JEOL JSM-7800F Extreme-Resolution Analytical Field Emission SEM (Zeiss, Germany).
Metagenomic characterisation of biofilm fungal diversity
To extract genomic DNA, the plastic was suspended in 10 ml sterile phosphate-buffered saline (PBS) overnight on a rocker. The samples were briefly vortexed, the plastic was removed, and the samples were centrifuged (twice at 5,000 rpm for 5 min). A volume of 9 ml of PBS was removed without disturbing the pellet. Sterile PBS (1 ml) was added to the pellet and then vortexed. A volume of 1 ml of the resuspended mixture was used for gDNA extraction using the Norgen Biotek Fungi/yeast genomic DNA isolation kit according to the manufacturer's protocol. A 0.8% agarose gel was run at 90 V for 25 min to confirm that DNA was extracted. Extracted DNA was stored at −20 °C.
For Internal Transcribed Spacer (ITS) Region amplification of each sample, two polymerase chain reaction (PCR) reactions containing 12.5 μl AccuStart II PCR ToughMix® [2X] (Quantabio), 1 μl ITS 4 primer (IDT 5′-TCCTCCGCTTATT3ATATGC-3′), 1 μl ITS 5 primer (IDT 5′-GGAAGTAAAAGTCGTAACAAGG-3′), as well as molecular grade water and either 2.5 μl gDNA template or 5 μl gDNA template were prepared.
The following PCR conditions were used: initial denaturation at 94 °C for 5 min, denaturation at 94 °C for 30 s, annealing at 50 °C for 30 s, extension at 72 °C for 30 s, and final extension at 72 °C for 5 min and hold at 4 °C, 25 cycles were completed. A 0.8% gel was run at 90 V for 30 min to confirm the amplification of the ITS sequence.
Amplicons were prepared for identification using the NEBNext® Ultra™ II End Repair/dA-Tailing Module (NEB, United States, E7546L). All ligation reactions were done using the Blunt/TA Ligase Master Mix from NEB (NEB, United States, M0367L). Amplicons were barcoded with native barcoding kits EXP-NBD104 and EXP-NBD114 (Oxford Nanopore Technologies, Oxford, UK), combined, and cleaned using AMPure XP Beads (Oxford Nanopore Technologies, Oxford, UK). The resulting eluate was collected, and AMII adapters (Oxford Nanopore Technologies, Oxford, UK) were ligated again and cleaned using AMPure XP beads. Library preparation was conducted according to the manufacturer's instructions and sequenced using R9.4.1 flow cells (Oxford Nanopore Technologies, Oxford, UK). Base calling and barcode demultiplexing were done using Guppy (version 6.1.2) adapters, and barcodes were also trimmed during this process. Read classification was done using megablast (Morgulis et al. 2008) against the non-redundant nucleotide database as previously reported (Langsiri et al. 2023). Reads were filtered based on the best bitscore, and only fungal sequences and sequence lengths longer than 400 base pairs were considered.
Yeast isolation and identification
The biofilms were scraped off the plastic samples using a sterile cell scraper and inoculated into 1 ml sterile PBS. This was vortexed, and 100 μl was spread over Yeast Malt Chloramphenicol plates (3 g/L yeast extract, 3 g/L malt extract, 10 g/L d-glucose, 5 g/L peptone powder, 16 g/L bacteriological agar, and 80 mg/L chloramphenicol powder) in duplicate. These plates were incubated at 30 °C for 2–3 days until possible yeast colonies were observed. The colonies were selected and streaked onto Yeast Malt Extract (YM) agar (3 g/L malt extract, 3 g/L yeast extract, 5 g/L peptone powder, 10 g/L d-glucose, 16 g/L bacteriological agar) plates and incubated at 30 °C until pure colonies were obtained. The cultures were confirmed to be yeast through microscopic analysis using a Zeiss Imager A2 Microscope.
Once pure cultures were obtained, yeast isolates were inoculated onto CHROMagar™ Candida plates (15 g/L of agar, 10.2 g/L peptone, 0.5 g/L chloramphenicol, and 22.0 g/L chromogenic mix), and incubated at 30 °C for 48 h.
In addition, for all pure cultures, an overnight grown colony on YM agar was suspended in molecular grade water and boiled at 94 °C for 10 min in a thermocycler. Colony PCR was performed using ITS 4 and ITS 5 primers and AccuStart II PCR ToughMix®. The PCR conditions were as follows: Initial denaturation at 94 °C for 5 min, denaturation at 94 °C for 30 s, annealing at 50 °C for 30 s, extension at 72 °C for 30 s, and final extension at 72 °C for 5 min and hold at 4 °C. An 0.8% gel was run at 90 V for 30 min to confirm the amplification of the ITS sequence.
Clean-up and sequencing preparations were done twice for all samples, once with the ITS 4 primer as the sequencing primer and again using the ITS 5 primer as the sequencing primer. A volume of 5 μl of the ITS amplicon for each isolate was mixed with 1 μl (1u) FastAP™ Thermosensitive alkaline phosphatase and 0.5 μl (10u) exonuclease I (Exo I) and incubated at 37 °C for 15 min and a further 15 min at 85 °C.
Clean-up products were prepared for sequencing using the Applied Biosystems BigDye™ terminator v. 3.1 Cycle Sequencing Kit in which the protocol for a 1/16 size reaction (total of 10 μl reaction volume) was followed: 0.5 μl premix, 1 μl sequencing primer (3,2 pmol.μl−1), 2 μl dilution buffer, 3 μl template, and 3.5 μl molecular grade water. The provided control reaction (1/16 reaction size) was also made. The PCR conditions were as follows: initial denaturation at 96 °C for 1 min, denaturation at 96 °C for 10 sec, annealing at 50 °C for 5 sec, extension at 60 °C for 4 min, and a hold phase at 4 °C, and the reaction was set for 25 cycles.
For post-reaction clean-up, 10 μl reaction volume was adjusted to 20 μl using molecular grade water. This was then transferred to 1.5 ml Eppendorf tubes containing 5 μl (125 mM) EDTA and 60 μl absolute ethanol. The tubes were vortexed and left to precipitate at room temperature for 15 min, and these tubes were then centrifuged at 20,000 g for 15 min at 4 °C. The supernatant was completely aspirated without disturbing the pellet, 200 μl of 70% ethanol was added, and the tubes were centrifuged at 20,000 g for 5 min at 4 °C. The supernatant was then completely aspirated, and the pellet was dried in a Speed-Vac for 5 min. Samples were stored at 4 °C to protect them from light.
Amplicons were sequenced using Applied biosystems™ 3,500 genetic analyser. Geneious Prime version 2022.2.2 was used to construct a consensus sequence between the ITS 4 and 5 sequences obtained. This consensus sequence was identified using NCBI's BLAST function.
Fluconazole susceptibility testing
Isolates were cultured overnight in YM broth at 37 °C in a shaker, streaked onto YM agar plates, and incubated for 24–48 h, until growth was observed. Cultures were then standardized in sterilised water up to a transmittance of 80–82% using a spectrophotometer. Standardised cultures were spread onto RPMI agar plates (10.4 g/L RPMI1640 powder without bicarbonate (Sigma Aldrich; Missouri USA), 34.5 g/L: 3-[N-morpholino] propanesulfonic acid buffer (MOPS) buffer (Sigma Aldrich), 20 g/L glucose, and 15 g/L agar) and left to dry in a laminar flow cabinet. A fluconazole strip (0.016–256 mg/L) (Liofilchem, Italy) was placed in the centre of each plate and incubated at 37 °C for 24–48 h, after which the minimum inhibitory concentration (MIC) values were read at the edge of the zone of inhibition (CDC 2020). C. krusei (ATCC 6258) was used as a quality control.
RESULTS AND DISCUSSION
Yeasts are visible in the plastisphere
SEM showed a mixed collection of microbial cells that were densely packed in each biofilm, with abundant microbial diversity present on all the polymer samples collected. Although most research on the plastisphere has focused on bacteria, it was evident that a rich eukaryotic diversity, including ciliates and diatoms (Figures S1 and S2 in the supplementary data), exists within these biofilms. The presence of diatoms is expected as many studies done on the plastisphere report their presence (Carpenter & Smith 1972; Zettler et al. 2013; Muthukrishnan et al. 2019; Amaral-Zettler et al. 2021). Diatoms act as phototrophs in these biofilms, and a previous study stated that they only make up a small portion of the eukaryotic diversity although they seem abundant in microscopy images (Bryant et al. 2016). Ciliates, including various freshwater ciliates of the genus Vorticella and the order Hypotrichida, were found on the PE polymer (Figure S1A and B). Other protists like dinoflagellates and amoeba were also seen and expected to form part of these biofilms (Zettler et al. 2013; Amaral-Zettler et al. 2021).
False colour micrograph of a possible yeast cell as evidenced by the size and shape of the cell. The arrow indicates a possible bud scar.
False colour micrograph of a possible yeast cell as evidenced by the size and shape of the cell. The arrow indicates a possible bud scar.
Pathogenic yeasts are part of the fungal metagenome of the plastisphere
Plot of fungal phyla detected on polymer samples. Circle size is indicative of abundance of each phylum.
Plot of fungal phyla detected on polymer samples. Circle size is indicative of abundance of each phylum.
Species plot and number of reads of potentially pathogenic ascomycetous yeasts detected on the five polymer samples. The colour scale to the right visually indicates the increase in number of reads recorded, while the exact number of reads per sample is noted in the blocks.
Species plot and number of reads of potentially pathogenic ascomycetous yeasts detected on the five polymer samples. The colour scale to the right visually indicates the increase in number of reads recorded, while the exact number of reads per sample is noted in the blocks.
Species plot and number of reads of potentially pathogenic basidiomycetous yeasts detected on the five polymer samples. The colour scale to the right visually indicates the increase in number of reads recorded, while the exact number of reads per sample is noted in the blocks.
Species plot and number of reads of potentially pathogenic basidiomycetous yeasts detected on the five polymer samples. The colour scale to the right visually indicates the increase in number of reads recorded, while the exact number of reads per sample is noted in the blocks.
It should be noted that the number of reads per species was generally low. However, a notable exception is the relatively high numbers of C. auris and Rhodotorula mucilaginosa reads detected in sample 4. To the best of our knowledge, this is the first report of C. auris in biofilms on plastic pollutants from freshwater.
The relatively high numbers of C. auris is the cause for concern, considering that this pathogen is a global threat to human health (Chowdhary et al. 2023). This pathogen has an estimated mortality of 30–60% (Brown et al. 2012; CDC 2019) and is often multidrug resistant (CDC 2019), which hinders the effective treatment of infections. C. auris is halotolerant and thermotolerant with the ability to remain viable on plastic surfaces for 2 weeks (metabolically active for 4 weeks) (Welsh et al. 2017). The abundant presence of R. mucilaginosa was not surprising since it is well known that this yeast can degrade various hydrocarbons (Jarboui et al. 2012; Benmessaoud et al. 2022) including plastic (Vaksmaa et al. 2023), for which it has a strong affinity (Wirth & Goldani 2012). This yeast has also been isolated from various aquatic environments, including those with high temperatures and high salt concentrations. It is the cause of fungaemia, central nervous system infections associated with central venous catheters, as well as cutaneous and ocular and prosthetic joint infections, peritonitis, keratitis, and ventriculitis (Wirth & Goldani 2012; Jarros et al. 2020). Importantly, this emerging pathogen is intrinsically resistant to many azoles, including fluconazole, and echinocandins, with recent reports also identifying 5-flucytosine-resistant isolates (Huang et al. 2022). The presence of these yeasts on plastic pollutants and the dispersive nature of these pollutants further support the hypothesis that these pollutants are vehicles that can spread pathogens to different environments.
Due to the lack of complete fungal genome databases, the results obtained from metagenomic analysis should be carefully interpreted. Sequences obtained are compared to sequences deposited in a database, and thus, microorganisms can only be correctly identified if the database already contains a reference sequence. A study utilizing virtual microbiomes demonstrated how incomplete genomic databases can affect the reliability of microbiome analyses, by increasing the number of false positives and providing poor estimates of the abundance of genera and species (Serrano-Antón et al. 2023). Thus, the presence of potentially pathogenic yeast species in these biofilms was confirmed by culture-dependent techniques.
Pathogenic yeasts can be isolated from the plastisphere
Yeast isolates were obtained from samples 2 (six isolates), 3 (six isolates), 4 (seven isolates), and 5 (five isolates). These isolates were preliminarily identified as yeasts based on microscopic morphology as well as colony colour when grown CHROMagar™ Candida media (Perry 2017; Scharmann et al. 2020; de Jong et al. 2021).
Table 2 presents all the observed characteristics of the isolates, grouped based on the similarity between the isolates within each sample. Samples 2 and 3 are grouped together due to the samples being collected in the same general area, while samples 4 and 5 were also collected in the same general area.
Characteristics of yeast isolates grouped by similarity
Isolate number . | Microscopic morphology . | CHROMagar ™ Candida colour after 48 h . |
---|---|---|
2A1 2A2 2A3 2B2 2B3 2A2 3B1 3 B2 | Ovoid | Pink-purple with white margin![]() |
2B1 3A3 | Ellipsoidal | Dark green-blue![]() |
3B3 | Ovoid single cells in chains | Bright green![]() |
3A1 | Ovoid | Light green![]() |
4B1 | Elongated cells, hyphal formation | Bright pink with white margin![]() |
4A1 4B2 5A1 5B1 5B2 | Ovoid | Pale purple/pink![]() |
4A2 4A3 | Round to ovoid | Pink-purple with white margin![]() |
4B3 5A2 5B3 | Ovoid to elongated cells | Purple with white margin![]() |
4A4 | Elongated cells | Dark olive green![]() |
Isolate number . | Microscopic morphology . | CHROMagar ™ Candida colour after 48 h . |
---|---|---|
2A1 2A2 2A3 2B2 2B3 2A2 3B1 3 B2 | Ovoid | Pink-purple with white margin![]() |
2B1 3A3 | Ellipsoidal | Dark green-blue![]() |
3B3 | Ovoid single cells in chains | Bright green![]() |
3A1 | Ovoid | Light green![]() |
4B1 | Elongated cells, hyphal formation | Bright pink with white margin![]() |
4A1 4B2 5A1 5B1 5B2 | Ovoid | Pale purple/pink![]() |
4A2 4A3 | Round to ovoid | Pink-purple with white margin![]() |
4B3 5A2 5B3 | Ovoid to elongated cells | Purple with white margin![]() |
4A4 | Elongated cells | Dark olive green![]() |
By using these data, some of the yeast isolates could be presumptively identified as C. albicans or C. dubliniensis (green), C. glabrata (pink-purple), or C. tropicalis (blue to blue-green) (Agrawal et al. 2014; de Jong et al. 2020; Lu et al. 2021). Interestingly, isolate 4A4 had an olive-green colony colour, which could be possibly the thermophilic black yeast, Exophiala dermatitidis (Lu et al. 2021).
The yeast isolates were further identified based on their ITS sequences, and it was found that the molecular identification matched the grouping done in Table 2, with several pathogenic yeast species identified (Table 3). Similar to our results, using in situ assays with virgin plastic material, Forero-López et al.(2022) were able to isolate C. tropicalis and R. mucilaginosa from a polluted estuary.
Yeast isolates identification after DNA sequencing as well as the expect (E) value and percentage identity according to NCBI BLAST
Yeast isolate . | Yeast species . | E value . | % ID . |
---|---|---|---|
2A1 | Candida glabrata | 0.0 | 99.40 |
2A2 | Candida glabrata | 0.0 | 100 |
2A3 | Candida glabrata | 0.0 | 100 |
2B1 | Candida tropicalis | 0.0 | 95.81 |
2B2 | Candida glabrata | 0.0 | 100 |
2B3 | Candida glabrata | 0.0 | 100 |
3A1 | Candida albicans | 0.0 | 100 |
3A2 | Candida glabrata | 0.0 | 99.74 |
3A3 | Candida troplicalis | 0.0 | 100 |
3B1 | Candida glabrata | 0.0 | 100 |
3B2 | Candida glabrata | 0.0 | 100 |
3B3 | Candida albicans | 0.0 | 100 |
4A1 | Saccharomyces cerevisiae | 0.0 | 100 |
4A2 | Candida glabrata | 0.0 | 99.87 |
4A3 | Candida glabrata | 0.0 | 100 |
4A4 | Exophiala dermatitidis | 0.0 | 100 |
4B1 | Candida krusei | 0.0 | 99.78 |
4B2 | Saccharomyces cerevisiae | 0.0 | 100 |
4B3 | Candida pseudoglaebosa | 0.0 | 100 |
5A1 | Saccharomyces cerevisiae | 0.0 | 100 |
5A2 | Candida pseudoglaebosa | 0.0 | 97.31 |
5B1 | Saccharomyces cerevisiae | 0.0 | 100 |
5B2 | Saccharomyces cerevisiae | 0.0 | 100 |
5B3 | Candida pseudoglaebosa | 0.0 | 100 |
Yeast isolate . | Yeast species . | E value . | % ID . |
---|---|---|---|
2A1 | Candida glabrata | 0.0 | 99.40 |
2A2 | Candida glabrata | 0.0 | 100 |
2A3 | Candida glabrata | 0.0 | 100 |
2B1 | Candida tropicalis | 0.0 | 95.81 |
2B2 | Candida glabrata | 0.0 | 100 |
2B3 | Candida glabrata | 0.0 | 100 |
3A1 | Candida albicans | 0.0 | 100 |
3A2 | Candida glabrata | 0.0 | 99.74 |
3A3 | Candida troplicalis | 0.0 | 100 |
3B1 | Candida glabrata | 0.0 | 100 |
3B2 | Candida glabrata | 0.0 | 100 |
3B3 | Candida albicans | 0.0 | 100 |
4A1 | Saccharomyces cerevisiae | 0.0 | 100 |
4A2 | Candida glabrata | 0.0 | 99.87 |
4A3 | Candida glabrata | 0.0 | 100 |
4A4 | Exophiala dermatitidis | 0.0 | 100 |
4B1 | Candida krusei | 0.0 | 99.78 |
4B2 | Saccharomyces cerevisiae | 0.0 | 100 |
4B3 | Candida pseudoglaebosa | 0.0 | 100 |
5A1 | Saccharomyces cerevisiae | 0.0 | 100 |
5A2 | Candida pseudoglaebosa | 0.0 | 97.31 |
5B1 | Saccharomyces cerevisiae | 0.0 | 100 |
5B2 | Saccharomyces cerevisiae | 0.0 | 100 |
5B3 | Candida pseudoglaebosa | 0.0 | 100 |
It is well known that certain Candida species are important opportunistic pathogens, where C. albicans is the most frequently isolated and best-studied Candida species causing infectious candidiasis (Rodrigues et al. 2014). C. glabrata is a non-albicans Candida species (NAC) that can cause serious fungal infections (including pneumonia, septicaemia, pyelonephritis, or endocarditis) in especially immunocompromised humans (Corrin & Nicholson 2011; Rodrigues et al. 2014). C. tropicalis and C. krusei are other NAC species that cause nosocomial infections and are currently increasing in prevalence as causative agents of candidaemia. C. tropicalis can rapidly disseminate in the immunocompromised host and can lead to high mortality rates (Chai et al. 2010), while C. krusei is recognised as potentially multidrug resistant and can also cause endophthalmitis, onycholysis, endocarditis, and osteomyelitis in infected patients (Pfaller et al. 2008; Jamiu et al. 2020).
According to the updated WHO fungal priority pathogens list to guide research, development, and public health action (2022), C. albicans is classified in the critical group, C. tropicalis in the high group, and C. krusei in the medium group. This further highlights the importance of studies investigating the presence of these pathogens on environmental plastic pollutants.
E. dermatitidis is a melanised ascomycete with a significant opportunistic potential. It is adapted to life at higher temperatures and is commonly found in household dishwashers, kitchen sinks, and steam baths and occurs at low levels in biofilms (Zalar et al. 2011; Heinrichs et al. 2013). E. dermatitidis can cause a spectrum of infections, subclinical pulmonary colonisation of cystic fibrosis patients, and in rare cases, disseminating infections in patients who are not immunocompromised (Machouart et al. 2011).
Saccharomyces cerevisiae is best known for its use in baking and brewing and is rarely thought of as pathogenic. However, there have been reports of S. cerevisiae causing fungaemia in individuals who are healthy, immunocompromised, or critically ill (Piarroux et al. 1999; Lherm et al. 2002). These infections were typically detected in individuals who received probiotic therapy with an S. cerevisiae strain invalidly referred to as Saccharomyces boulardii (Muñoz et al. 2005).
Candida pseudoglaebosa is a non-pathogenic yeast, frequently found in raw milk, including the milk of goats and cows (Delavenne et al. 2011; Neubeck et al. 2015), and has also been implicated in the spoilage of white brine cheeses like feta and halloumi (Geronikou et al. 2022).
Fluconazole susceptibility of isolated yeasts
Antifungal susceptibility tests for various Candida species as well as the comparison of E-test strip results to the reference method results are well documented (Sewell et al. 1994; Maxwell et al. 2003; Metin et al. 2011), with a high level of agreement (>90%) between the results obtained from commercial E-test strips and reference broth microdilution results (Mallié et al. 2005). Thus, E-test strips were used for rapid antifungal susceptibility testing of yeast isolates.
The results presented in Table 4 are generally within the ranges previously reported (Sewell et al. 1994; Chryssanthou 2001; Metin et al. 2011; Song et al. 2015). However, it is evident that C. glabrata isolates exhibited the lowest susceptibility towards fluconazole, ranging from an MIC of 16 μg/mL to complete resistance. It is known that C. glabrata has intrinsically low susceptibility to fluconazole and can quickly become resistant upon exposure (Arendrup 2010; Arendrup & Patterson 2017; Won et al. 2021). The incidence of fluconazole-resistant blood stream infections is also increasing (Won et al. 2021), indicating the potential risks of these yeasts present in the plastisphere.
MIC values of isolated yeasts as determined using fluconazole E-test strips
Yeast isolates . | Yeast species . | Fluconazole MIC (μg/ml) . |
---|---|---|
2A1 | Candida glabrata | 16 |
2A2 | Candida glabrata | 16 |
2A3 | Candida glabrata | 24 |
2B1 | Candida tropicalis | 0.5 |
2B2 | Candida glabrata | >256 |
2B3 | Candida glabrata | 32 |
3A1 | Candida albicans | 0.19 |
3A2 | Candida glabrata | 16 |
3A3 | Candida tropicalis | 1.5 |
3B1 | Candida glabrata | 64 |
3B2 | Candida glabrata | >256 |
3B3 | Candida albicans | 0.19 |
4A1 | Saccharomyces cerevisiae | 48 |
4A2 | Candida glabrata | 24 |
4A3 | Candida glabrata | 24 |
4A4 | Exophiala dermatitidis | Not determined |
4B1 | Candida krusei | 32 |
4B2 | Saccharomyces cerevisiae | 48 |
4B3 | Candida pseudoglaebosa | 24 |
5A1 | Saccharomyces cerevisiae | 16 |
5A2 | Candida pseudoglaebosa | 6 |
5B1 | Saccharomyces cerevisiae | 32 |
5B2 | Saccharomyces cerevisiae | 48 |
5B3 | Candida pseudoglaebosa | 12 |
ATCC 6258 | Candida krusei | 48 |
Yeast isolates . | Yeast species . | Fluconazole MIC (μg/ml) . |
---|---|---|
2A1 | Candida glabrata | 16 |
2A2 | Candida glabrata | 16 |
2A3 | Candida glabrata | 24 |
2B1 | Candida tropicalis | 0.5 |
2B2 | Candida glabrata | >256 |
2B3 | Candida glabrata | 32 |
3A1 | Candida albicans | 0.19 |
3A2 | Candida glabrata | 16 |
3A3 | Candida tropicalis | 1.5 |
3B1 | Candida glabrata | 64 |
3B2 | Candida glabrata | >256 |
3B3 | Candida albicans | 0.19 |
4A1 | Saccharomyces cerevisiae | 48 |
4A2 | Candida glabrata | 24 |
4A3 | Candida glabrata | 24 |
4A4 | Exophiala dermatitidis | Not determined |
4B1 | Candida krusei | 32 |
4B2 | Saccharomyces cerevisiae | 48 |
4B3 | Candida pseudoglaebosa | 24 |
5A1 | Saccharomyces cerevisiae | 16 |
5A2 | Candida pseudoglaebosa | 6 |
5B1 | Saccharomyces cerevisiae | 32 |
5B2 | Saccharomyces cerevisiae | 48 |
5B3 | Candida pseudoglaebosa | 12 |
ATCC 6258 | Candida krusei | 48 |
The presence of antifungals in water is expected due to their frequent use in personal care products, and oral and topical medications as well as from runoff from farms (Hof 2001; Yao et al. 2016; Dalhoff 2017; Li et al. 2019; Toda et al. 2019), with azoles as the most used (Vanreppelen et al. 2023). Antifungals from all these point sources frequently spill into various bodies of water, establishing an environment in which yeasts and antifungals co-exist in close association with one another, potentially leading to the development of antifungal resistance. This occurrence is often seen in bacteria found in water containing antibiotics (Osińska et al. 2020; Kotlarska et al. 2015). Although some increased Minimum Inhibitory Concentrations (MICs) were seen, the current study is not able to elucidate if the specific aquatic environment may be aiding in antifungal resistance development, and more studies are required to clarify this.
CONCLUSIONS
The fungal diversity found in the plastisphere is abundant and dominated by the phyla Ascomycota and Basidiomycota. Potentially pathogenic yeasts belonging to the genera Candida (including strains with elevated MICs for fluconazole) and Exophiala were identified using culture-dependent and culture-independent methods. This is not surprising as many pathogenic yeasts are able to form biofilms on commonly used plastic medical equipment (Douglas 2002, 2003; Azevedo et al. 2014). This increases the need to investigate the presence of pathogenic yeasts on environmental plastic pollution and the possible health impact that their presence may have on the health of the surrounding community who come into frequent contact with these pollutants. From this study, we can conclude that plastic pollutants may act as both reservoirs and vectors that aid in the spread of potentially pathogenic yeast species to different environments, with these yeasts showing similar susceptibility to fluconazole as isolates from clinical samples.
ACKNOWLEDGEMENTS
The authors wish to thank Jade Hastings for assistance with adding false colour to Figure 1 and Linda Basson for assistance with identification of the ciliates. This work was funded by the National Research Foundation of South Africa (grant 115566 to CHP). The funders had no input into the study design, generation or interpretation of the data or the writing of the manuscript.
DATA AVAILABILITY STATEMENT
All relevant data are included in the paper or its Supplementary Information.
CONFLICT OF INTEREST
The authors declare there is no conflict.