Relative Humidity Influences Aureobasidium pullulans Degradation of Polyester Polyurethane Foam
Amanda Stickney, Nicole Renninger, Dominique Wagner, John Van Dusen, Victor A. Roman, Vanessa Varaljay, Blake W. Stamps, Nancy Kelley‐Loughnane, Karen C. Dannemiller
TL;DR
This study shows that higher humidity increases the ability of a fungus to break down plastic foam, which could help in managing plastic waste.
Contribution
The study identifies how relative humidity affects fungal degradation of polyurethane foam and links it to cutinase gene expression.
Findings
Higher ERH leads to greater foam weight loss, with up to 5.1% loss at 100% ERH.
Cutinase genes are upregulated at higher ERH, with one showing significant Impranil clearing.
Fungal growth and degradation are visible at high ERH via SEM imaging.
Abstract
Microbial‐induced corrosion costs billions of dollars, including replacing plastics degraded by fungi. Fungal growth is moisture dependent, but we need to better understand how equilibrium relative humidity (ERH) affects plastic degradation. The goal of this project was to measure how ERH impacts degradation of polyester polyurethane foam by Aureobasidium pullulans and identify potential genetic pathways. We incubated three environmental strains of A. pullulans on foam at 50%, 85%, and 100% ERH and evaluated degradation through foam weight loss, Scanning Electron Microscopy (SEM), external nutrient availability, RNA sequencing, and proxy Impranil clearing. Higher ERH after 1 week of incubation corresponded to greater weight loss in foam (p = 0.002), with percent weight loss ranging from 0.11% to 5.1%. SEM foam imaging shows signs of fungal growth and degradation at high ERH while…
Genes, proteins, chemicals, diseases, species, mutations and cell lines named across the full text — each resolved to its canonical identifier and authoritative record.
Click any figure to enlarge with its caption.
FIGURE 1
FIGURE 2
FIGURE 3
FIGURE 4
FIGURE 5
FIGURE 6
FIGURE 7
FIGURE 8
FIGURE 9| Strain | Genome size (Mb) | BUSCO duplicate score |
|---|---|---|
| AFRL64 | 27 | 0.3% |
| AFRL76 | 77 | 93.1% |
| OSU3 | 49 | 0.4% |
| Most similar cluster representative sequence (percent identity %) (Accession) | Most similar | |
|---|---|---|
| Cutinase 1 | Hypothetical Protein in | Hypothetical Protein (92.82%) ( |
| Cutinase 2 | Cutinase Protein in | Cutinase Protein (90.22%) ( |
| Cutinase 3 | Cutinase Protein in | Cutinase Protein (87.56%) ( |
| Cutinase 4 | Cutinase Protein in | Cutinase Protein (87.64%) ( |
| Cutinase 5 | Cutinase Protein in | Cutinase Protein (90.22%) ( |
| Cutinase 6 | Cutinase Protein in | Cutinase Protein (87.50%) ( |
| Cutinase 7 | Hypothetical Protein in | Hypothetical Protein (92.82%) ( |
| Cutinase 8 | Cutinase Protein in | Cutinase Protein (92.27%) ( |
| Cutinase 9 | Cutinase Protein in | Cutinase Protein (100.00%) ( |
| Cutinase 10 | Cutinase Protein in | Cutinase Protein (99.11%) ( |
- —Alfred P. Sloan Foundation10.13039/100000879
- —Ohio Department of Higher Education (ODHE) Strategic Ohio Council for Higher Education Defense Associated Graduate Student Innovators/Air Force Research Labs10.13039/100016942
Peer Reviews
No public reviews on file for this paper yet. If you reviewed it on a platform where reviews are public (OpenReview, ICLR, NeurIPS, ICML), you can paste yours below so the community can read it here.
Videos
No videos yet. Explain this paper in a talk, walkthrough, or lecture? Add one.
Taxonomy
TopicsMicroplastics and Plastic Pollution · biodegradable polymer synthesis and properties · Polymer Foaming and Composites
Introduction
1
Polyurethane is used in homes, cars, boats, and aircraft as insulation, furniture, coatings, adhesives, apparel, gaskets, life jackets, and other products (Di Bisceglie et al. 2022). Polyurethanes have a wide range of uses because the polymers can be aligned to create ridged or flexible structures (Eceiza et al. 2005). Polyurethanes are largely resistant to biodegradation due to their crystalline structure, but amorphous regions in the plastic are vulnerable to degradation by microorganisms (Howard 2002; Huang and Roby 1986). For over 30 years, the US Department of Defence has used polyurethanes in aircraft topcoats (Iezzi 2022), which, if compromised, can lead to corrosion of the underlying metallic substrate that requires maintenance or corrosion management. Corrosion management in 2018 cost the US Air Force $5.67 billion, some of which could be attributed to degradation or damage of protective topcoats (Li et al. 2021). One potential method for lowering the cost of corrosion and biodegradation is to create environments unsuitable for growth of microbes such as bacteria and fungi that induce biodegradation.
Plastics have high versatility and durability and are now common in everyday life. Plastics are everywhere in the built environment and plastic pollution has become a growing issue in the outdoor environment as well. Daily plastic consumption ranges from 20 to 100 kg per capita (Kibria et al. 2023). This plastic is disposed of in landfills or directly into the environment. This can cause a wide range of environmental and human health issues. For instance, exposure of zebrafish to nanoplastics caused impairments impacting survival and reproductive behaviours (Sarasamma et al. 2020). In humans, nanoplastics can negatively impact gut microbiota (Fackelmann and Sommer 2019) and cause damage to cells (Hoffman and Lumpkin 2018). Biodegradation may be one solution to the challenges with plastic waste because it uses organisms such as fungi and bacteria to break down plastics.
Aureobasidium pullulans is one example of a common fungus capable of biodegradation. A. pullulans is a plant pathogen and polymorphic fungus that lives in a wide range of natural and built environments (Gostinčar et al. 2014; Prasongsuk et al. 2018). A. pullulans is a saprophytic fungus (Prasongsuk et al. 2018), a type of fungus that acquires nutrients as monomers dissolved in water that result from enzymatic reactions (Puiggené et al. 2022; Shah et al. 2008). Saprophytic fungi can excrete different enzymes to cleave molecules in the surrounding environment. One enzyme produced by Aureobasidium spp. is cutinase, which can break ester bonds found in cutin (i.e., the waxy protective layer of plants). Breaking down the protective layer of cutin can allow for A. pullulans and other microbes to infect the plant (Schäfer,1993). This can result in root or stem rot as well as leaf spots (Cooke 1959). The ester bonds in cutin can also be found in the carbon backbones of plastic polymers (Di Bisceglie et al. 2022; Kaushal et al. 2021; Schäfer 1993; Shah et al. 2008). This means that cutinase is naturally able to degrade both cutin and certain plastics. With A. pullulan's ability to survive in many environments, the natural opportunistic use of cutinase to break down ester bonds found in plastics is of interest. Cutinases, EN 3.1.1.74, break the ester bonds in plastics leaving monomers that are dissolved into water and taken up into the fungal cell (Puiggené et al. 2022; Shah et al. 2008). A. pullulans potentially uses cutinases to degrade materials to a water‐soluble metabolizable form that the fungus uses as a nutrient source (Webb et al. 1999; Wright et al. 2020).
Moisture, including from elevated relative humidity in the air, is critical to support degradation driven by microbes. High relative humidity increases the microbial growth of fungi such as Aspergillus fumigatus , Penicillium sp., and Eurotium (Nielsen et al. 2004; Pasanen et al. 1991). While the impact of humidity on microbial growth may be well documented, less is known about the interactions with specific fungi and their enzymes. Relative humidity impacts the fungal production of enzymes, by‐products, and physical masses that impact degradation. More potential metabolic pathways are expressed in indoor fungal communities at elevated relative humidity compared to lower levels, and there is an up‐regulation of fungal genes that corresponds to the production of secondary metabolites and allergens (Hegarty et al. 2018). Biosynthesis pathways in Aspergillus flavus also become upregulated at elevated water activity (Zhang et al. 2014). In addition to upregulating synthesis pathways, water is used in the chemical processes of degradation. Biotic and abiotic hydrolysis uses water to break longer polymers into shorter monomers (Parekh et al. 2011; Singh et al. 2021). In sum, relative humidity impacts growth, gene expression, and chemical processes that play an important role in understanding biodegradation.
What is not known is how ERH, the relative humidity when both a material and the surrounding air are at equilibrium, impacts A. pullulans degradation of polyurethane or how ERH impacts A. pullulans genetic degradation pathways. These pathways include the cutinase pathway. Understanding the impact of environmental conditions on fungal processes is critical to controlling material degradation. This information can be used to encourage degradation in landfills or prevent degradation in environments in which plastics are used, such as aircraft.
The goal of this work was to measure how relative humidity impacts the degradation of polyester polyurethane foam by A. pullulans and identify the associated biochemical pathways. Strains were isolated from environmental sampling from aircraft and house dust. We incubated inoculated commercial polyurethane foam with A. pullulans under different ERH levels and then measured the degradation of polyurethane foam in weight loss trials and with scanning electron microscopy (SEM) imaging. These experiments were also supported by nutrient trials to determine if carbon sources outside of the foam were needed for fungal growth. Identification of cutinases and their gene expression across different ERHs was evaluated through whole genome RNA sequencing and Impranil plate clearing.
Materials and Methods
2
Strain Selection
2.1
Isolates of A. pullulans were isolated from aircraft and house dust from a home in Ohio, USA. Swabs from sites in the aircraft visibly contaminated with fungi were streaked onto a plate as described in Hung et al. (Hung et al. 2019). Dust from the home was added directly to the initial plate made of potato dextrose agar (PDA) (Sigma Aldrich, St. Louis, MO, USA) with 0.025 g of chloramphenicol (Sigma Aldrich, St. Louis, MO, USA). Chlorampenicol was used to prevent unwanted bacterial growth allowing for fungal isolation and focus on the strains of fungi from environmental swabs. Colonies were isolated from mixed culture plates using a sterile technique. A metal inoculation loop was sterilised by submersion in 100% ethanol followed by 5 s in the flame of a Touch‐O‐Matic. The loop was allowed to cool before scraping a single colony off the mixed plate and streaking onto a fresh PDA plate with 0.025 g of chloramphenicol. Colonies for each strain were streaked onto their own plate. The plates were then incubated at 25°C for 5 days. After 5 days, the sterile loop technique was repeated until each strain was isolated on its own plate without visual contaminants.
Strains were tested for hydrolytic potential based on ImpranilDLN (Material Number 57822612, Covestro, Leverkusen, Germany) clearing. Plates were created with 12.5 g ThermoFisher Scientific LB Agar (ThermoFisher, Waltham, MA, USA), 7.5 g Agar (Durable Industrial Finishing Co., Tucker, GA, USA), and 0.1 g of CaCl_2_ dihydrate (Sigma Aldrich, St. Louis, MO, USA) in 500 mL of distilled water. The mixture was then autoclaved for 30 min at 121°C. To prevent bacterial growth, 0.025 g of chloramphenicol was added into the cooled mixture. Impranil was then added as 3.75 mL of Impranil DLN‐SD. Once the mixture cooled, 28 mL was added into each petridish. The petridishes were allowed to cool before covering and storing at 4°C. Isolated A. pullulans colonies were transferred from its PDA plate onto an Impranil plate using a sterile loop technique. Each plate was then incubated for 2 weeks at 25°C with clearing documented every 1 week. Based on the initial weight loss, Impranil results, and isolate morphological characteristics, three strains were selected for inclusion in this study: OSU3, AFRL76, and AFRL64. The strain OSU3 was isolated from house dust. The AFRL76 and AFRL64 strains were isolated from aircraft.
Cell Collection
2.2
We created solutions of spores from each strain to inoculate onto foam pieces for testing. Each of the strains was grown on plates of PDA and 10^7^ spore/mL concentrated spore solutions were collected using the modified plate flooding method (ASTM G26) described previously (Nastasi et al. 2020). A solution of PowerBead Solution (PBS) (Ricca Chemical Company, Batesville, IN, USA) and 0.1% Tween‐20 (Sigma Aldrich, St. Louis, MO, USA) was added to the plates of each strain. Each plate was then scraped with a sterilised inoculating loop to loosen spores off the plate and into the PBS solution. This resulting solution was collected, placed into a flask of 2 mm garnet beads, and vigorously shaken by hand for 10 min. Next, the solution was filtered through sterile glass wool. The concentration of spores in the filtered solution was determined by the method described previously (Nastasi et al. 2020). Briefly, the spore solution was diluted and stained with crystal violet. The stained spores were then counted under a Labomed microscope using an InCyto DHC‐N01‐5 Neubauer Improved C‐Chip. Cell counts were completed in triplicate and then the mean was used as the concentration value for each separate solution. The spore solutions were stored at −20°C until inoculation of foam.
Cleaning Foam
2.3
Before use in experiments, commercial blue polyester polyurethane foam without antistatic additives or flame retardant was cut and cleaned. Polyester polyurethane foam pieces were cut into 1.27 × 1.27 × 2.54 cm blocks. These blocks were then submerged in 70% ethanol for 30 s followed by another 30 s in deionised water. To improve permeation, each foam block was squeezed as it was placed into and out of the ethanol and the deionised water. The foam was then placed into a furnace set to 50°C for 8 h or until dry. After drying, the foam was covered in baked aluminium foil and stored at room temperature.
Weight Loss Incubations
2.4
Cleaned foam was weighed, embedded with 2 mg of Difco Potato Dextrose Broth powder (BD Diagnostics, Franklin Lakes, NJ, USA), then inoculated using nebulized A. pullulans strain spores. Potato Dextrose Broth powder was used to mimic the nutrients that might be found in dust or other environments. Nutrient sources were added using a modified ASTM method F608‐13 (Dannemiller et al. 2017). A steel pipe sterilised with 70% ethanol and wrapped in baked aluminium foil was rolled over the nutrients and foam samples to spread nutrients into the foam. The foam was then inoculated with spores through nebulization using the Medline Aeromist Compact Nebuliser compression kit and a modified method (Nastasi et al. 2020). Nebulization is the process of turning liquid into a mist. During the spore nebulization process, air bubbles flow through a spore solution to create a mist of spores. The mist of spores flows through tubing into a sealed container of samples. In the sealed container, the mist of spores settles, and spores are deposited onto the samples. This process simulates the natural settling of spores onto materials such as foam or indoor dust. The nebulization setup consists of two containers, an inoculation chamber and a nebulization cup, which is used to hold the spore solution. These containers are connected by a large tube. A smaller tube also connects the nebulization cup to the nebuliser compressor. For the weight loss trials, 2 mL of 10^7^ cells per mL spore solution were placed into the nebulization cup and foam samples were placed into a 13.97 × 13.97 × 17.15 cm glass nebulization jar. All samples were prepared in triplicate for each nutrient and ERH variation. The compressor was turned on for 12 min then turned off for another 12 min, allowing the aerosolized spores to settle. Then, the samples were removed from the nebulization jar and placed into 16.51 × 16.51 × 20.32 cm glass incubation jars.
ERH within the incubation jars was controlled at 50%, 85%, and 100% using salt solutions or deionised water. MgCl_2_ was used to make the 50% ERH solution, NaCl for the 85% ERH solution, and pure deionised water was used to make the 100% ERH solution (Dannemiller et al. 2017). Open beakers containing 100 mL of the appropriate salt solution were placed into the incubation jar along with an Onset HOBO logger (Bourne, MA USA) which was used to measure ERH conditions in the incubation jar. All incubation jars were then sealed with a single layer of parafilm. Parafilm was used as a cover to allow for the exchange of gases while maintaining ERH. Samples were incubated in triplicate for 1 and 2 weeks at 25°C in an incubator at each ERH level (VWR, Radnor, PA, USA).
Weight Loss Evaluation
2.5
Each foam piece was cleaned after incubation to remove fungal debris. To clean, each foam piece was cut into four pieces to improve the cleaning penetration into the foam. The pieces were placed into a tube containing 5 mL of 70% ethanol and then the tube was vortexed for 30 s before the foam was removed. The foam pieces were placed into a second tube containing 5 mL of 70% ethanol and vortexed for another 30 s. The foam was then removed from the second tube and all of the wash liquid was vacuum filtered using a 55 mm cellulose filter with 11 μm particle retention to collect any small foam pieces that might have broken off during the cleaning process. Filtrate was discarded and both the foam and the filters were dried in a 50°C furnace for 8 h or until dry before a final weighing of the foam sample. Starting weights and post‐incubation weights were compared. Foam that was incubated without inoculation was used as a control. All weights were recorded on a Mettler Toledo XSE105 Balance. Cleaning method efficacy was confirmed through SEM imaging. Weights were also recorded on all foam pieces prior to inoculation and incubation.
Evaluation of the Impact of Nutrient Availability on Degradation
2.6
For the nutrient evaluation, polyurethane foam was cut into 1.27 × 2.54 × 2.54 cm blocks and then cleaned using the pre‐incubation ethanol bath process described for the weight loss evaluation. Larger foam blocks were used to increase the number of fungi present to levels more easily detected by qPCR. Either 2 mg Potato Dextrose Broth powder or an equivalent amount of a minimal media mixture was added to the cleaned foam. The minimal media contained ammonium chloride, potassium phosphate, and magnesium sulfate in amounts corresponding to the amount of N, C, P, and S found in 2 mg of dust based on the findings in Dannemiller et al. Table S2 (Dannemiller et al. 2017). Minimal media was used as a nutrient source without carbon, and Potato Dextrose Broth powder was used as the carbon‐containing nutrient source.
Samples comparing nutrient availability were inoculated with AFRL64 and used a modified nebulization method. To increase spore concentration on the foam, 3 mL of spore solution was used rather than the 2 mL used for the weight loss samples. More spore solution was used to increase the present fungal concentrations. All incubation jars for this trial were incubated for 5 weeks at 25°C in an incubator.
Fungal Quantification
2.7
Quantification of the 18S rRNA gene via quantitative polymerase chain reaction (qPCR) was used as a proxy for A. pullulans concentration in the foam. DNA was extracted from foam cut into quarters and placed into tubes with 3 mL of PBS. The tubes were vortexed, and then the foam was removed from the PBS and squeezed to expel as much solution as possible. Next, the solution underwent extraction using the Maxwell RSC PureFood GMO and Authentication Kit and Promega instrument (Madison, WI, USA). The kit instructions were modified to begin with 5 min of bead beating before using the Maxwell RSC PureFood GMO and Authentication Kit. All samples then underwent qPCR on a 96‐well plate using a QuantStudio 6 Flex System (Applied Biosystems, Carlsbad, CA, USA) with SYBR Green qPCR master mix (Applied Biosystems, Carlsbad, CA, USA) and a reaction volume of 25 μL, 23 μL of master mix and 2 μL of extracted DNA. Both forward and reverse primers were diluted to 0.3 μM in the final master mix solution. The fungal forward‐primer FF2 (5′‐GTTAAAAAGCTCGTAGTTGAAC‐3′) and reverse primer FR1 (5′‐CTCTCAATCTGCAATCCTTATT‐3′) were used to determine fungal growth (Zhou et al. 2000). The qPCR run method consisted of two sets of cycles. First, a single cycle for 2 min at 50°C followed by 10 min at 95°C. Second, 40 cycles with 15 s at 95°C then 1 min at 60°C. Each qPCR run contained a standard curve, an extracted negative sample, an extracted positive sample, and a no template control. The standard curve was extracted from AFRL64 of a known concentration serial diluted 1:10 from 3.1 × 10^5^ to 3.1 spores per mL.
Scanning Electron Microscopy
2.8
SEM was performed to check for visual changes in the polyurethane foam before, during, and after A. pullulans growth. Foam samples were inoculated and incubated in duplicate for each strain at 50, 85, and 100% ERH. All samples were incubated in an incubator set to 25°C for 2 weeks before imaging. One of each duplicate was imaged with the fungal growth while the other was cleaned before imaging to observe the foam surface with no microbial obstructions. Samples imaged without A. pullulans growth were cleaned using 70% ethanol using the post‐incubation protocol described in the section “Weight Loss Evaluation” above. To prepare for imaging, foam pieces were attached to autoclaved SEM pin stubs and gold sputtered. The samples were then imaged using a Thermo Scientific Apreo l LoVac Scanning Electron Microscope (Thermo Fisher Scientific Inc., Waltham, MA) at the Center for Electron Microscopy and Analysis.
Nucleic Acid Sequencing
2.9
All 3 A. pullulans strains were grown on the foam at the 3 ERH levels and RNA was sequenced to identify gene expression potentially associated with degradation. Foam samples were cut, cleaned, embedded with nutrients, and inoculated as described for the weight loss trials. The inoculated samples were incubated for 2 weeks at 50%, 85%, and 100% ERH. Strain AFRL64 was incubated in triplicate to test for variance. Strains OSU3 and AFRL76 only had one sample for each ERH. After incubation, the foam samples were cut into four pieces each and washed in 3 mL of Buffer RLT from the Zymo Quick DNA Fungal Kit (Irvine, CA, USA). This was done to remove fungal growth from the foam into a solution. The wash liquid was placed into 2 mL tubes of 0.1 mm glass beads. The tubes with samples for RNA extraction were bead beaten for 5 min in a cold block and centrifuged. The supernatant in the tubes was then extracted using the Qiagen RNeasy Mini Kit (Qiagen, Hilden, Germany) procedure. The resulting RNA extracts each totaled 2–5 μg. The tubes with samples for DNA extraction were bead beaten for 3 min and centrifuged and then extracted following the Zymo Quick DNA Fungal Kit procedures. DNA isolates were each 1 μg or more. Both the RNA and DNA extracts were quality tested using a bioanalyzer before being sequenced. Extracted samples were sequenced using an Illumina NovaSeq (Illumina, CA, USA) at the University of Kansas Medical Center. DIN ranged from 6.6 to 6.8 and RIN ranged from 5.3 to 8.9 (Table S2). The Illumina DNA library was prepared using Unique Dual Index adapters (UDI) (Illumina 20018704) and the stranded mRNA library was prepared using the Universal Plus mRNA‐seq with NuQuant + UID library prep (Tecan Genomics 0520‐A01). The gDNA and mRNA libraries were pooled for sequencing using the NovaSeq 6000 S1 Reagent Kit v1.5 (200 cycles) (Illumina 20028318). The read depth was 100X and targeted the gDNA library assuming an A. pullulans genome size of 30 Mbp. Stranded mRNA libraries targeted 10M paired end reads.
Genomic DNA Sequencing Data
2.10
Genomic DNA data from sequencing were used to identify genes that may be associated with degradation. The reads from sequencing were first checked for quality using FastQC (Andrews 2010) before trimming with Trimmomatic (Bolger et al. 2014). A secondary quality check was conducted on the trimmed reads using FastQC. The trimmed reads were then de novo assembled into a complete genome with SPAdes (Bankevich et al. 2012). A “careful” flag was used in SPAdes with all other parameters set to the default settings. De novo assembly created scaffolds. Any repeats in the scaffolds were masked using RepeatMasker (Smit et al. 2015). The assembled scaffolds were structurally annotated by BREAKER (Hoff et al. 2019). The structural annotations then underwent functional gene annotation to predict cutinase genes. Cutinase genes were predicted using InterProScan (Jones et al. 2014; Zdobnov and Apweiler 2001).
Genomic RNA Sequencing Data
2.11
RNA sequencing data were processed and used to determine differential gene expression. First, the RNA data was checked for quality using FastQC before trimming with Trimmomatic and another quality check with FastQC. Trimmed reads were then aligned to the DNA assembly made previously using SPAdes (Langmead and Salzberg 2012). From the RNA assembly, a transcript count table was generated using featureCounts in subread (Liao et al. 2013). In this process, an annotation table was also generated. Differential expression analysis was performed in R (RStudio Team 2020). In R, the DESeq2 package (Love et al. 2014) was used for strain AFRL64. Default settings were used alongside a significance threshold with a multiple test correction of FDR < 0.1. DESeq2 normalised using the median of ratios method accounting for RNA composition and sequencing depth (Mistry et al. 2017). AFRL76 and OSU3 did not have biological replicates so edgeR (Robinson et al. 2010) was used for differential expression analysis. edgeR default settings for an exact test between groups was used with an assumed dispersion value of 0.2^2^. The assembled genomes were run in BUSCO (Benchmarking Universal Single‐Copy Orthologs) to measure genome quality (Manni et al. 2021). BUSCO was run with default settings with Dothideomycetes reference orthologs. Genomic data can be found in the NCBI BioProject database. OSU3 data can be found in PRJNA1186706: Built Environment Isolates SAMN44769747: Aureo_osu3 while AFRL64 and AFRL76 data can be found in PRJNA811524: Aircraft Sampling as SAMN44749444: Aureo_64 and SAMN44752121: Aureo_76, respectively.
Cutinase Expression in Bacteria
2.12
Strain AFRL64 cutinases were cloned and their expression was examined as Impranil clearing by Escherichia coli , BL21 (DE3) (Studier and Moffatt 1986), a standard host for studying fungal cutinase expression (Chen et al. 2008). The corresponding open reading frame (ORF) Cutinase 1 of AFRL64 was cloned from amino acids 431–847 which includes the full predicted cutinase domain. The complete cutinase was not sequenced due to the complexity and ability to make a gBlock. For cutinase 2, the complete ORF was cloned. Gene fragments were obtained from Integrated DNA Technologies (IDT) (Newark, New Jersey, USA). Both cutinases were cloned using restriction enzyme digestion into the bacterial expression vector pET28a using BamHI and SalI. The gene fragments were amplified using PCR to verify the constructs. PCR was conducted following the manufacturer's instructions. The PCR products were gel purified, digested using BamHI and SalI and ligated using the T4 DNA Ligase (New England Biolabs, Ipswich, MA, USA) into pET28a. The genetically modified E. coli were then plated on 50 mg/mL LB Kanamycin for construct selection. DNA constructs were verified by both PCR and restriction digestion. After verification of the construct, correct clones were transformed into BL21 (DE3) cells. Expression was induced by adding isopropyl‐β‐D‐thiogalactopyranoside (IPTG) 0.1 mM to cultures during the exponential growth phase. Expression of the cutinase genes was verified using Coomassie blue staining or Western Blot using an anti‐his antibody. All predicted cutinases were run in NCBI's Blastx algorithm using the ClusteredNR (nr_cluster_seq) database to check homology and identity (Altschul et al. 1997). Their completeness was also analysed using SignalP 6.0 (Neilsen et al. 2024) to check for signal peptides.
Impranil Hydrolysis
2.13
E. coli cultures containing AFRL64 cutinase 1 gene, AFRL64 cutinase 2 gene, and an empty control vector were grown overnight at 37°C. Then 10 μL of the culture were spotted onto LB Kan plates supplemented with 1% Impranil. The Impranil was supplemented with and without 0.1 mM IPTG. The plates were incubated at 27°C for 2 days. Plates were inspected for halos around biofilm spots to define the preliminary indicator of hydrolysis of the Impranil polymer.
Confocal Microscopy
2.14
Confocal microscopy was used to determine if any of the three strains of A. pullulans are heterokaryons (contain multiple nuclei), which has been documented previously (Durrell 1968; Zabel and Morrell 2020). Slides of the strains were fixed, then stained with DAPI and TryPan Blue. DAPI stained the nuclei and TryPan Blue stained the cell wall. Spores were grown on PDA plates at 25°C for 2 weeks. Spores were then placed onto sterilised glass slides and rinsed with PBS. After the initial rinse, the slides were fixed using 4% paraformaldehyde. The paraformaldehyde was applied to the slides and incubated at room temperature for 15 min. The slides were then rinsed with PBS before adding 1% tween‐20 to the slide to increase cell permeability (Amidzadeh et al. 2014). After 5 min at room temperature, the slides were rinsed again with PBS before TryPan Blue was added and incubated at room temperature for 15 min. The slides were once again rinsed before staining with DAPI for 15 min. The slides were rinsed a final time and mounted using a solution of 50% glycerol and 50% PBS. All slides were then covered with a number 1 glass coverslip secured by clear nail polish. The slides were then imaged using an Olympus FV3000 multi‐confocal with a 60× oil objective at the OSU Center for Microscopy Imaging Facility by one of their staff members.
Statistics and Software Utilised
2.15
Bioinformatics were run as described above while all other statistics were done in Stata/BE17.0 (StataCorp 2021). ANOVA tests were used to check for significant associations. If significant associations were present, Tukey–Kramer pairwise comparisons or t‐tests were calculated. Statistical significance was defined as p‐values less than the alpha significance level of 0.05. This corresponds to 95% confidence. All weight loss and nutrient samples were prepared and assessed in triplicate. The foam weight preincubation and post‐incubation were compared using a two‐sample paired t‐test to determine the significance of the difference in the foam weight. Then the weight loss ANOVA compared the foam weight loss across strain, incubation time, and ERH. Then Tukey–Kramer pairwise comparisons were used to compare weight loss across the incubation times and ERHs. The nutrient trial ANOVA compared the relative quantity of fungus based on qPCR across the nutrients added to the foam and the ERH. AFRL64 RNA and DNA sequencing samples were prepared in triplicate but the AFRL76 and OSU3 sequencing samples were not prepared with replicates. A one‐tailed two‐sample t‐test assuming equal variance was conducted to determine if the number of counts present in the AFRL64 samples incubated at 85% ERH and 100% ERH was statistically greater than the number of counts present in the AFRL64 samples incubated at 50% ERH. Due to the limited replicates, AFRL76 and OSU3 did not undergo this comparison.
Results
3
Weight Loss and Nutrient Evaluation Reveal That RH Impacts Fungal Degradation and Growth
3.1
Indoor environmental A. pullulans strains AFRL64, AFRL76, and OSU3 were incubated on foam at three ERH levels to measure fungal growth and foam weight loss. After both one and two week incubations, there was a significant decrease in foam mass relative to pre‐inoculation weights (two sample paired t‐test p < 0.0001) (Figure 1). In the weight loss trials, higher ERH was significantly associated with increased percent weight loss in polyurethane foam (ANOVA p = 0.0002). The greatest amount of weight loss across all strains was observed at 100% ERH. There were no significant differences in weight loss between strains at 100% ERH (ANOVA p = 0.30). The 100% and 85% ERH samples had significantly higher weight loss compared to the 50% ERH samples (Tukey–Kramer pairwise comparison to 50% p < 0.001 and p = 0.007 respectively). The 100% and 85% ERH samples were not significantly different from each other (Tukey–Kramer pairwise comparison p = 0.19). A higher percent of weight loss was also observed at 2 weeks compared to 1 week in the 100% ERH samples (Tukey–Kramer pairwise comparison p = 0.002). Tukey–Kramer pairwise comparison p‐values can be found in Table S1. The significance of time's impact on foam degradation requires further exploration.
Percent weight loss in (A) one‐week and (B) two‐week incubations of AFRL64, AFRL76, and OSU3 at 50%, 85%, and 100% ERH significant differences are noted with bars and ** for (p < 0.05) and *** for (p < 0.005). The difference between one and two week incubations was significant with a p‐value < 0.005.
While ERH significantly impacted weight loss, across the three ERH levels there was no significant difference in A. pullulans growth, as determined by qPCR genome equivalents, in foam samples grown with minimal media compared to Potato Dextrose Broth powder (ANOVA p‐value with nutrient type and ERH variables = 0.11) (Figure 2).
Carbon influence on A. pullulans growth. Nutrient trials where samples were embedded with Potato Dextrose Broth, added carbon, or Minimal Media, no carbon, and incubated at 50%, 85%, and 100% ERH for 5 weeks at 25°C using strain AFRL64.
Scanning Electron Microscopy Reveals Signs of Foam Degradation
3.2
After a 2 weeks incubation, SEM images of all A. pullulans strains showed visual signs of presumed degradation at elevated ERH levels and fungal presence at all ERH levels. Cleaned foam that never contained A. pullulans was used as a control that, in SEM images, appeared smooth with no pitting or tears (Figure 3A). On top of the smooth foam, nutrients added to the foam appeared as crystalline structures in these images and A. pullulans spores appeared rounded (Figures 3B, 4A,C,E, 5A,C, and 6A,C,E). Images from strain AFRL64 showed increased biomass on the foam as the ERH increases (Figure 4A,C,E). At the 100% ERH foam cleaned of AFRL64 had visual signs of degradation with cracking and holes visible in the cleaned foam imaged after growth at 100% ERH (Figure 4F). The foam cleaned of this AFRL64 strain imaged after growth at 50% and 85% ERH still appeared smooth at the scale imaged (Figure 4B,D). AFRL76 showed a similar increase in growth across ERH and visible cracking at 100% ERH (Figure 5A,C,E,F) with no visible cracking or pitting by AFRL76 at 85% ERH (Figure 5D). The AFRL strains showed no obvious signs of cracking or pitting at 50% ERH (Figures 4B and 5B). However, OSU3 showed pitting at both 50% and 100% ERH (Figure 6B,F), not visible at 85% ERH (Figure 6D). OSU3 also appeared to have similar increases in growth related to increases in ERH (Figure 6A,C,E). All three strains had visible round fungal cells at 50% ERH that were visibly denser at 85% ERH (Figures 4A,C, 5A,E, and 6A,C). At 100% ERH the fungal cells of all three strains appeared to coat the foam as a biofilm (Figures 4E, 5E, and 6E). Cracking and pitting in the foam are visible signs of degradation similar to those found in polyurethane foam after 3 weeks in a degradation chamber set for 50% RH and 90°C (Pellizzi et al. 2014). These images suggest that A. pullulans degrades the foam with the greatest foam degradation occurring at 100% ERH.
SEM of (A) clean foam with no growth or nutrients and (B) clean foam with nutrients and no growth. Arrow points to nutrients that can be seen on the foam surface.
SEM of (A) AFRL64 fungal growth at 50% ERH, (B) cleaned foam after AFRL64 fungal growth at 50% ERH, (C) AFRL64 fungal growth at 85% REH, (D) cleaned foam after AFRL64 fungal growth at 85% ERH, (E) AFRL64 fungal growth at 100% ERH, and (F) cleaned foam after AFRL64 fungal growth at 100% ERH. Arrows in A, C, and E point to cells growing on the foam surface. The arrow in F points to cracks in the foam's surface. Control foam with no nutrients or fungi and control foam with nutrients and no fungi can be seen in Figure 3.
SEM of (A) AFRL76 fungal growth at 50% ERH, (B) cleaned foam after AFRL76 fungal growth at 50% ERH, (C) AFRL76 fungal growth at 85% ERH, (D) cleaned foam after AFRL76 fungal growth at 85% ERH, (E) AFRL76 fungal growth at 100% ERH, and (F) cleaned foam after AFRL76 fungal growth at 100% ERH. Arrows in A, C, and E point to cells growing on the foam surface. The arrow in F points to pitting in the foam's surface. Control foam with no nutrients or fungi and control foam with nutrients and no fungi can be seen in Figure 3.
(A) OSU3 fungal growth at 50% ERH, (B) cleaned foam after OSU3 fungal growth at 50% ERH, (C) OSU3 fungal growth at 85% ERH, (D) cleaned foam after OSU3 fungal growth at 85% ERH, (E) OSU3 fungal growth at 100% ERH, and (F) cleaned foam after OSU3 fungal growth at 100% ERH. The arrows in B and F point to pitting in the foam's surface. Control foam with no nutrients or fungi and control foam with nutrients and no fungi can be seen in Figure 3.
A. pullulans Genome Size
3.3
In order to determine if degradation was due to enzymatic processes, we sequenced the DNA and RNA of our A. pullulan strains. Strains of A. pullulans have a wide range of genome sizes (Table 1). AFRL64 had the smallest genome with only 27 Mb. AFRL76 genome was about 77 Mb in size, more than twice the size of AFRL64 and other previously reported A. pullulans (Wei et al. 2023). OSU3 was between the two AFRL strains containing 49 Mb. The large differences in genome size may suggest that AFRL76 and OSU3 have more than one nucleus or possible contamination. AFRL76 has a BUSCO duplicate score of 93.1%. This indicates a high amount of replicated orthologs within the genome, further suggesting the presence of an additional heterogeneous nucleus. To determine the presence of multiple nuclei, confocal microscopy was conducted. The two‐week incubation of the samples for confocal microscopy mirrored the 2‐week incubations for the DNA and RNA sequencing. A full summary of BUSCO genome data can be seen in Table S3.
Another possible source for these results could be contamination. To test for contamination, each strain was streaked on 5 PDA plates and allowed to incubate at 25°C for 2 weeks. As plates aged, the colonies turned darker, which corresponds with the chlamydospores morph of A. pullulans which produces melanin and is more likely to occur in older colonies (Gadd and Mowll 1985; Prasongsuk et al. 2018). No obvious contamination was observed.
Cutinase Gene Identity and Signal Cleavage Sites
3.4
Through Blastx analysis the predicted cutinase genes have strong ties with the Aureobasidium genus and cutinases. Table 2 shows the results indicating that all 10 cutinases had percent identities of 87% or more with an A. pullulans protein or cutinase. Only cutinase 1 and 7 were most similar to a hypothetical protein, with all other predicted cutinases having the greatest percent identity with cutinases. Cutinase 9 and 10 both had the highest percent identity with A. pullulans, while the other eight predicted cutinases shared the greatest percent identity with other species in the Aureobasidium genus. These include Aureobasidium melanogenum and Aureobasidium namibiae. A. melanogenum, once considered a type of A. pullulans, is currently classified as its own species (Zalar et al. 2008). This aligns with our prediction that these sequences are cutinase genes from Aureobasidium fungus. A total of 4 of the 10 cutinases, cutinases 1, 7, 9, and 10, have a high probability of having a signal cleavage site (> 0.97). All other cutinases had a lower probability of having a signal, with probabilities ranging from < 0.01 to 0.59. This could potentially mean that some cutinases are being held within the cells rather than being transported to outside the cell, which could be studied in future work. This may cause discrepancies between sequencing results and weight loss results.
Confocal Microscopy Nucleus Evaluation
3.5
Confocal microscopy of AFRL64, AFRL76, and OSU3 imaged layers of A. pullulans cells. Nuclei are stained blue by PureBlu DAPI, and cell walls are stained red by TryPan Blue (Figure 7). DAPI has previously been used for nuclei staining including finding multiple nuclei in the stages of fungal cell division (Ko et al. 2016). All images suggested that each of the three strains contains only one nucleus. Large blotches of blue in the OSU3 images may be residual DAPI stain that was not removed with rinsing. While there were no observed heterokaryons, another possible reason for the large genome size could be that AFRL76 was composed of more than one organism and not pure.
Confocal microscopy images using an Olympus FV3000 multi‐confocal with a 60× oil objective for strains (A) AFRL64, (B) AFRL76, and (C) OSU3 after 2 weeks of incubating at 25°C on PDA plates. Nuclei are stained blue with DAPI and cell walls are stained red with TryPan Blue.
Predicted Cutinases
3.6
Genome sequencing data was used to find genes predicted to be associated with cutinase production, one of many enzymes that could be the cause of plastic degradation. From the three A. pullulans strains, 10 cutinase genes were predicted by InterProScan (v5.56‐89.0); (Jones et al. 2014; Zdobnov and Apweiler 2001) based on the assembled genomes. AFRL64 and OSU3 each have 2 unique putative cutinase genes. AFRL76 contains 6 putative cutinase genes. These genes can be classified as long, ranging from 1896 to 2574 base pairs, or short, ranging from 666 to 678 base pairs. Cutinase 1 in AFRL64 and Cutinase 7 in AFRL76 contain identical nucleotide sequences. Cutinase 2 in AFRL64 and Cutinase 5 in AFRL76 are also identical. This shows some shared cutinases between the strains isolated from aircraft. Cutinase 9 and 10 from OSU3 do not appear to be present in either of the AFRL strains. While not identical cutinase 10 from OSU3 is similar to the M438DRAFT_264999 cutinase previously reported from A. pullulans (Watanabe and Hashimoto 2021). Using NCBI BLAST the percent identity between the two cutinase genes is 98.92% (using OSU3 as a reference has 100% coverage while using M438DRAFT_264999 as a reference has 86% coverage) (Sayers et al. 2022; Watanabe and Hashimoto 2021).
A majority of the cutinases have the most counts at 85% ERH or 100% ERH. Cutinases of different lengths demonstrated different patterns of expression at the different ERH conditions (ANOVA p = 0.0195) (cutinase lengths can be found in Table S4). ERH significantly impacted the cutinase expression (ANOVA p = 0.0196) with the difference between 85% ERH and 50% ERH being statistically different (pairwise tukey p = 0.015). The longer cutinases (Figure 8A) such as cutinase 1 and cutinase 7 showed increases in counts at elevated ERH. At 85% ERH there were 610 counts of cutinase 1 and 800 counts of cutinase 7. At 50% ERH cutinase 1 and 7 had 120 counts and 6 counts, respectively. These data show increased gene expression of cutinases at higher ERH (Figure 8A,B and S1).
Heat map of cutinase expression (shown in red). Expression is determined by counts, the number of overlapping sequence fragments. (A) Cutinases 1, 4, 7, 8, and 9 are long ranging from 1896 to 2574 base pairs. (B) Cutinases 2, 3, 5, 6, and 10 are short ranging from 666 to 678 base pairs. The AFRL64 cutinase counts were measured in triplicate while the AFRL76 and OSU3 cutinase counts had no replicates. Identical cutinase pairs are noted using * and +. See Figure S1 for a bar chart of this data.
Impranil Clearing Is Observed in Bacteria With Cloned AFRL 64 Cutinase 2
3.7
Impranil is a polyester‐polyurethane that can be used as a proxy for studying preliminary indicators of polyurethane degradation. The long cutinase (cutinase 1) and the short cutinase (cutinase 2) from AFRL64 were cloned in bacteria to determine their polyester degradation potential via Impranil clearing on plates after 2 days of incubation (Figure 9). Impranil clearing tests showed a visible halo or clearing of Impranil surrounding E. coli colonies containing AFRL64 cutinase 2 only on all 4 plates, including ones containing the inducer isopropyl‐β‐D‐thiogalactopyranoside (IPTG) or not. The relatively small degree of Impranil degradation seen in Figure 3B may be due to leakiness, when genes are uninduced but gene expression is observed. The colonies containing the AFRL64 cutinase 1 gene and the empty vector exhibit no clearing in any of the 4 plates. The lack of visible clearing expressed by cutinase 1 may be due to complications in gene expression or enzyme secretion. The influence of pH on Impranil sensitivity makes these results a preliminary indicator of degradation requiring quantitative molecular analysis for confirmation (Biffinger et al. 2015). These clearing patterns are preliminary indicators that the AFRL64 short cutinase 2 gene has the ability to hydrolyze polyester polyurethane‐based compounds such as Impranil.
Impranil clearing of Escherichia coli strains expressing AFRL cutinase 1 or cutinase 2. Colony 1 is E. coli containing the AFRL cutinase 1 gene, colony 2 is E. coli with an empty vector, and colony 3 is E. coli containing the AFRL cutinase 2 gene. (A) LB control (IPTG‐) plates after 2 days incubating at 27°C. (B) Shows the same plate as in A, but following incubation of 1 week at 4°C. (C) LB (IPTG+) plates after 2 days incubating at 27°C. (D) Shows the same plate as in C, but following incubation of 1 week at 4°C.
Discussion
4
The goal of this study was to measure the impact of relative humidity on polyester polyurethane foam degradation by A. pullulans and identify the associated biochemical pathways. We found that higher ERH was associated with more foam degradation by A. pullulans as demonstrated by weight loss, SEM imaging, and analysis of enzymes. At 100% ERH, images show visual signs of degradation for all three strains. Cutinase expression in strain AFRL64 was also impacted by ERH levels, which is in agreement with the results observed in the SEM and weight loss trials where there is increased degradation at higher ERH if this enzyme is involved in degradation. Cutinase 2, isolated from AFRL64, showed Impranil hydrolysis as seen by environmental isolates of A. pullulans previously (Crabbe et al. 1994). Cutinase P1 cut1 isolated from the yeast Papiliotrema laurentii caused Impranil clearing (Roman et al. 2024). NMR (Nuclear Magnetic Resonance) confirmed Impranil hydrolysis at rates above that of some commercial lipases (Biffinger et al. 2015; Roman et al. 2024). This makes cutinases like those found in A. pullulans especially important to understand as they may have a large impact on polyurethane degradation.
Apparent degradation was also observed through weight loss and SEM results. Other studies found microbial degradation to result in 10% weight loss in hard polyurethane foam after foam pieces were incubated with 300 mL of landfill leachate water for 3 months at 50°C (Filip 1978). All strains in our experiments showed signs of degradation with the greatest amount of observed degradation occurring at 100% ERH. This degradation was visible in SEM images of foam before and after A. pullulans growth. Fungal polyester polyurethane esterases have been shown to cause visible degradation in polyester polyurethane (Zhang et al. 2022).
All three A. pullulans genomes are different sizes with 2 shared and 6 unique cutinases. The cutinases are shared between AFRL64 and AFRL76, while OSU3 was found to have a cutinase similar to one previously recorded (Watanabe and Hashimoto 2021). These cutinases could be sorted into short (about 670 bp) and long (over 1800 bp). The length of these genes is significant due to the influence of gene length on factors such as gene duplication and alternative splicing (Grishkevich and Yanai 2014). The results presented here are suggestive, and future research would be necessary to definitively link cutinase expression to polyurethane degradation in the environment.
Two of the strains, AFRL76 and OSU3, showed signs of impurities or possibly being another type of Aureobasidium. AFRL76's large 77 Mb genome size with a BUSCO score of 93.1% might imply that it is not a pure strain and may be composed of more than one organism while OSU3 might not be A. pullulans, but another Aureobasidium spp. The genome size of the A. pullulans var. aubasidani DH177 was 31.34 Mbp (Wei et al. 2023), compared to AFRL64 at 27 Mbp, AFRL76 at 77 Mbp, and OSU3 at 49 Mbp. Aureobasidium can be difficult to define because of their versatility both under stress and with different nutrient availability (Gostinčar et al. 2014). Genome size alone is not enough of an indicator to identify between Aureobasidium. The need to redefine Aureobasidium can occur even with strain genomes ranging from 25.43 to 29.62 Mbp (Gostinčar et al. 2014). Another possible explanation is that AFRL76 is a heterokaryon, but confocal microscopy suggests that the strains all only contain a single nucleus. Future work might explain the variation in genome size of these strains, the presence of similar cutinases among A. pullulans strains, and the differences in ERH influence on cutinase expression.
Moisture is generally the limiting factor for fungal growth in the indoor environment (Nastasi et al. 2020). We demonstrated that ERH (moisture availability) also influenced material degradation and expression of associated genes. A. pullulans is a saprophytic fungus that receives its nutrient from absorbing water containing dissolved monomers (Prasongsuk et al. 2018). An increase in humidity would result in a greater likelihood of interactions with water that can act as a transport and reservoir for enzymes and monomers. This knowledge can help better manage plastics. Plastics are persistent and can cause harm to human and environmental systems. If plastic production and management continue at current levels, by 2050 there will be 12,000 Mt. of plastics in landfills or the natural environment (Geyer et al. 2017). There are few ways to sustainably recycle and degrade plastics, and plastic recycling and incineration can cause the release of pollution to the environment and requires energy. Moist environments may lead to better promotion of polyurethane degradation, which may have important implications in landfills to promote plastic degradation. Alternatively, maintaining sufficiently low ERH in built environments may be critical to limit cutinase production and undesirable fungal degradation.
Beyond plastics, cutinases can also cause agricultural issues by increasing the virulence of fungal plant pathogens (Lee et al. 2010). Other enzymes are also effective at both plant tissue degradation and plastic degradation. These enzymes include chitinases, lipases, and laccases (Chua et al. 2013; Safdar et al. 2024; Santo et al. 2013). To control the presence and production of these enzymes, it is important to control fungal growth. Water is often the limiting factor for fungal growth (Nastasi et al. 2020), and therefore we need to understand the impact of relative humidity on enzymatic activity related to plastic degradation. Other papers have considered the impact of relative humidity on fungal growth and genetic pathways, but this paper focuses specifically on the impact of cutinase isolated from A. pullulans on polyurethane foam and its relationship with ERH. This focus can help relate broader findings to specific applications that aim to control degradation in order to save time and financial resources maintaining environments. Those responsible for maintaining buildings and aircraft, as well as those who maintain landfills, should consider the environmental conditions to control the polymer's longevity and state.
Limitations
5
This study was limited by the controlled setting and number of strains, replicates, temperatures, light exposures, nutrients, and ERHs tested. In the environment, fungi commonly experience variations in temperature and ERH, which our samples were not exposed to. Foam in the built environment also withstands long‐term use and stress, which were not evaluated in this study. Our 1 versus 2 week foam weight loss results indicate time plays an important role in degradation. Future work could consider more extended periods of time. Other parameters may also influence degradation and add to the variability of results. Future research can consider the contribution of degradation differences due to both environmental and phenotypic effects. Our strains might not be representative of all A. pullulans, all phases of A. pullulans growth, or the microbiome interactions. Organisms exist in complex communities, including not only other fungi but also bacteria. These community dynamics were not considered here. Growth phase and nutrient availability are both factors that have been shown to impact A. pullulans colonies. In addition to microbial degradation, foams in built environments would likely experience other types of degradation, such as mechanical degradation and photodegradation. In this study, we focused on cutinase genes that may be associated with degradation. However, we are unable to prove causality that these genes are associated with degradation. Other enzymes, such as chitinases, lipases, and laccases, could also potentially play a role in foam degradation (Chua et al. 2013; Safdar et al. 2024; Santo et al. 2013). A. pullulan, cutinase, and environmental degradation remain areas of interest as we move forward.
Limitations were also present in imaging, sequencing, and qPCR. Imaging and many laboratory processes only capture one small part of the sample and might not be representative of a whole. SEM imaging also provides qualitative evidence of degradation. Taxa resolution and reference database completeness limit standard whole genome sequencing (Somervuo et al. 2016). Due to these limitations, sequencing is suggestive without the use of radio labelling to determine protein production. Though sequencing has improved over recent years, it is suggestive, and future research might require RNA confirmation of the interactions between cutinase production and relative humidity.
Conclusions
6
This study demonstrated that ERH influences A. pullulans degradation of polyurethane foam. High ERH resulted in higher rates of A. pullulans growth and increases in degradation observed through weight loss and SEM. The three strains of A. pullulans in this study had a variable number of predicted cutinase genes. Of the two short cutinases observed to be impacted by ERH in AFRL64 and OSU3, one cutinase exhibited upregulation at both 85% and 100% ERH; however, the other was up‐regulated only at 85% ERH. Similar patterns were also seen in two of the short cutinases predicted in AFRL76. Future research should investigate the impact of longer incubation periods as well as environmental factors such as light, radiation, and oxygen availability on cutinase expression and the promotion or prevention of foam degradation. This will aid in engineering better environments and buildings for more sustainable plastic use as people look to create long‐standing structures and communities that require minimal maintenance.
Author Contributions
Amanda Stickney: investigation, writing the original draft, validation, visualisation, reviewing and editing, software, formal analysis, and data curation. Nicole Renninger: conceptualization, investigation, funding acquisition, writing the original draft, reviewing and editing, and data curation. Dominique Wagner: investigation, methodology, visualisation, reviewing and editing, software, and formal analysis. John Van Dusen: investigation, funding acquisition, reviewing, and editing. Victor A. Roman: investigation, methodology, reviewing and editing, formal analysis, and resources. Vanessa Varaljay: conceptualization, investigation, funding acquisition, methodology, reviewing and editing, project administration, and supervision. Blake W. Stamps: investigation, methodology, reviewing and editing, formal analysis, project administration, supervision, and resources. Nancy Kelley‐Loughnane: investigation, funding acquisition, reviewing and editing, project administration, and supervision. Karen C. Dannemiller: conceptualization, investigation, funding acquisition, writing the original draft, methodology, validation, visualisation, reviewing and editing, formal analysis, project administration, data curation, supervision, and resources.
Funding
This work was supported by the Alfred P. Sloan Foundation, G‐2018‐11240; Ohio Department of Higher Education (ODHE) Strategic Ohio Council for Higher Education Defense Associated Graduate Student Innovators/Air Force Research Labs, RX29‐OSU‐21‐1, RX36‐OSU‐20‐3.
Conflicts of Interest
The authors declare no conflicts of interest.
Supporting information
Table S1: Tukey–Kramer pairwise comparison p‐values. Table S2: RNA and DNA quality scores. Table S3: Summary of BUSCO genome data. Table S4: Summary of predicted cutinase lengths. Raw sequences for Cutinases 1–10. Figure S1: Bar chart of cutinase expression determined by counts, the number of overlapping sequence fragments. (A) Cutinases 1, 4, 7, 8, and 9 are long ranging from 1896 to 2574 base pairs. (B) Cutinases 2, 3, 5, 6, and 10 are short ranging from 666 to 678 base pairs. The AFRL64 cutinase counts were measured in triplicate while the AFRL76 and OSU3 cutinase counts had no replicates. Identical cutinase pairs are noted using * and ^+^. Codon Optimised Sequences for Cut1 and Cut2.
The reference list from the paper itself. Each links out to its DOI / PubMed record.
- 1Altschul, S. F. , T. L. Madden , A. A. Schäffer , et al. 1997. “Gapped BLAST and PSI‐BLAST: A New Generation of Protein Database Search Programs.” Nucleic Acids Research 25: 3389–3402.9254694 10.1093/nar/25.17.3389 PMC 146917 · doi ↗ · pubmed ↗
- 2Amidzadeh, Z. , A. B. Behbahani , N. Erfani , et al. 2014. “Assessment of Different Permeabilization Methods of Minimizing Damage to the Adherent Cells for Detection of Intracellular RNA by Flow Cytometry.” Avicenna Journal of Medical Biotechnology 6, no. 1: 38–46.24523954 PMC 3895578 · pubmed ↗
- 3Andrews, S. 2010. “Fast QC: A Quality Control Tool for High Throughput Sequence Data (0.11.9).” [Computer software]. Babraham Bioinformatics. http://www.bioinformatics.babraham.ac.uk/projects/fastqc/.
- 4Bankevich, A. , S. Nurk , D. Antipov , et al. 2012. “SP Ades: A New Genome Assembly Algorithm and Its Applications to Single‐Cell Sequencing.” Journal of Computational Biology: A Journal of Computational Molecular Cell Biology 19, no. 5: 455–477.22506599 10.1089/cmb.2012.0021 PMC 3342519 · doi ↗ · pubmed ↗
- 5Biffinger, J. C. , D. E. Barlow , A. L. Cockrell , et al. 2015. “The Applicability of Impranil DLN for Gauging the Biodegradation of Polyurethanes.” Polymer Degradation and Stability 120: 178–185.
- 6Bolger, A. M. , M. Lohse , and B. Usadel . 2014. “Trimmomatic: A Flexible Trimmer for Illumina Sequence Data.” Bioinformatics (Oxford, England) 30, no. 15: 2114–2120.24695404 10.1093/bioinformatics/btu 170PMC 4103590 · doi ↗ · pubmed ↗
- 7Chen, S. , X. Tong , R. W. Woodard , G. Du , J. Wu , and J. Chen . 2008. “Identification and Characterization of Bacterial Cutinase.” Journal of Biological Chemistry 283, no. 38: 25854–25862.18658138 10.1074/jbc.M 800848200 PMC 3258855 · doi ↗ · pubmed ↗
- 8Chua, T.‐K. , M. Tseng , and M.‐K. Yang . 2013. “Degradation of Poly(ε‐Caprolactone) by Thermophilic Streptomyces thermoviolaceus Subsp. Thermoviolaceus 76T‐2.” AMB Express 3, no. 1: 8.23360778 10.1186/2191-0855-3-8PMC 3844369 · doi ↗ · pubmed ↗