The Detection of Propionate Utilization by Bacteria Isolated from a Plastic Recycling Site
1
AgResearch Limited, Hopkirk Research Institute, Palmerston North 4442, New Zealand
2
New Zealand Food Safety Science and Research Centre, Tennent Drive, Massey University, Palmerston North 4474, New Zealand
*
Author to whom correspondence should be addressed.
Appl. Microbiol. 2024, 4(2), 856-874; https://doi.org/10.3390/applmicrobiol4020059
Submission received: 28 March 2024
/
Revised: 17 May 2024
/
Accepted: 20 May 2024
/
Published: 23 May 2024
Abstract
:(1) The study aims to utilize a reported approach for culturing mesophilic bacteria from a plastic waste environment; (2) The work revived mesophilic microbial population from an aged PET recycling site using a culture-based approach, and determined the purified isolates in genus level in 16S identification; (3) A total of 59 bacterial isolates were obtained, in which microbial species, including Pseudomonas spp, Rhodococcus spp, and Burkholderia spp were identified as abundance. It was observed that the surviving microbes favoured sodium propionate as a short-chain carbon source for growth, rather than the intended plastic substrate, PET. The preference of sodium propionate utilization by several bacterial isolates, including 5601W (detected as Rhodococcus spp.), 5601Y, 7801, and 7802 (detected as Burkholderia spp.), was confirmed through growth curve analysis and cell enumeration conducted in a medium where sodium propionate served as the sole carbon source.; (4) The microbial demonstration revealed the metabolic complex of microbial communities in the environment and indicated the challenges associated with bacterial isolation from environments with accumulated plastic waste.
1. Introduction
When plastic debris accumulates in various environments, it creates a unique ecological niche that becomes a thriving habitat for diverse microorganisms [1]. This plastic-induced habitat, referred to as the plastisphere, is characterized by the intricate association of various microbial communities, forming a dynamic ecosystem [2].
The durability of plastic substrates provides an enduring platform for microbial colonization and the development of complex microbial biofilms [2]. Numerous studies investigated the impact of plastic substrates on the microbial variance, like in seawater and sediment environments and [1,3]. Microbial nutrient uptake, particularly the ability of microbial consortia to utilize carbon sources ranging from long chain to short chain molecules, profoundly influences the microbial abundance and diversity [4,5]. Therefore, understanding these dynamics can offer valuable insights into the broader environmental consequences of plastic accumulation and can be essential for assessing the role of plastic as a vector for invasive or pathogenic species.
The study was to revive mesophilic bacteria from an aged PET recycling site using a culture-based approach. The in-vitro investigation didn’t retrieve any bacteria utilizing predominant plastic carbon source (PET), while the study isolated bacterial growth involving the addition of the antifungal agent sodium propionate [6]. Propionate as a short chain fatty acid can be directly used for chain elongation in fatty acid synthesis in the presence of other carbon source [7]. Propionate can be metabolised by specific bacteria through various pathways to generate the intermediate propionyl-CoA, which can be further metabolised to Propionyl-ACP (acyl carrier protein) that act as a primer for chain elongation [7]. The involvement of propionate in fatty acid synthesis resulted in a higher biomass production and fatty acid yield. Certain bacterial species, such as Cupriavidus necator [8] and Ralstonia eutropha [9], have been studied for their ability to produce PHA (polyhydroxyalkanoate) and PHB (Polyhydroxybutyrate) from propionate. When provided with propionate as a carbon source under suitable conditions, these bacteria accumulate PHA or PHB as intracellular granules.
The specific metabolic pathways for propionate metabolism can differ among various bacterial strains. However, it is known that intracellular Propionoyl-CoA likely increases resulting from elevated protein and amino acid degradation during nitrogen and glucose starvation as the physiology of these cells adapt to these nutrient limitations [10]. Propionate-utilizing bacteria may be present in plastic waste environments due to the limited availability of carbon source. The detection of propionate-utilizing bacteria in this study revealed the metabolic complex of microbial communities in such environments and indicated the challenges associated with bacterial isolation from environments with accumulated plastic waste.
2. Materials and Methods
2.1. Environmental Sampling
The environmental samples (92 in total) were collected from aged PET (polyethylene terephthalate) waste piles (2–3 years old) at a waste management site, Awapuni Resource Recovery Park in Palmerston North, New Zealand, 13 December 2022. The No. 1–46 samples mainly consisted of soil containing visibly fine plastic fragments or broken plastic pieces (2–3 cm) intermixed with soil, primarily located at the bottom of the piles. The No. 47–92 samples were particularly plastic piece samples, which were cut into smaller fragments to reach the expected weight for the following experiment. All samples were placed in 50 mL Falcon tubes and transported to the lab in a chilly bin with ice. They were promptly used for microbial enrichment on the same day.
2.2. Culture-Based Bacteria Isolation
Approximately 1 g of each sample (No. 1–92) was individually cultivated in a 20 mL glass vial containing 10 mL of a modified liquid medium carrying additional 0.4 g autoclaved virgin PET beads (Sigma, Tokyo, Japan). The medium used for the initial enrichment was modified basal medium (per 1 L of distilled water: 0.7 g of KH2PO4, 0.7 g of K2HPO4, 0.7 g of MgSO4·7H2O, 1.0 g of NH4NO3, 0.005 g of NaCl, 0.002 g of FeSO4·7H2O, 0.002 g of ZnSO4·7H2O, and 0.001 g of MnSO4·H2O, according to the ASTM standard (ASTM G22-76) [11]) supplemented with 0.04% sodium propionate (SP, antifungal agent) [6]. Samples were aerobically incubated in shaking (120 rpm) condition at 25 °C (Figure 1).
After an 80-day incubation period, 100 µL of a 100-fold dilution from the upper layer of each sample culture (above the sediments) was spread on the modified lignin-mineral salt agar [6]. This agar [12] is to promote the in vitro growth of polymer-remediating microbes, considering that natural heteropolymer (plant-derived lignin) and plastic polymers often coexisted in the waste environment.
The agar composition consisted of 1.0 g kraft lignin, 10 g D-glucose, 2.4 g Na2HPO4, 2 g KH2PO4, 5 g peptone, 0.10 g NH4NO3, 0.01 g MgSO4, 0.01 g CaCl2 and 15 g agar, supplemented with 0.04% SP and 0.5% PET powder (IML Plastic Ltd., Wellington, New Zealand). The inoculated agar plates were incubated at 25 °C, and incubated for up to 2 days.
Diverse microbial colonies emerged on each agar plate after 24–48 h of incubation. Distinct morphology, encompassing colonial shape, colour, gloss, form, and transparency, was observed for each sample. Colonies with unique characteristics were individually selected and streaked on new lignin-mineral salt agar plates to obtain pure cultures. Consequently, sample isolates were categorised into two groups: bacteria-like and fungi-like. A total of 101 pure cultures were established and individually incubated in a 20 mL glass vials under shaking conditions (130 rpm) at 25 °C.
These 101 environmental bacterial, capable of growing on the modified lignin agar, were purified by additional subculturing on the modified lignin agar at 25 °C (Figure 1). Fresh single colonies were retrieved at the 3rd or 4th subculture using sterile loops and individually inoculated into 10 mL of carbon-free basal medium [11] containing 0.04% SP and 10 PET beads (Sigma). The isolate ID, represented in four digits, indicates the sample number in the first two digits and the number of single colonies picked from each sample in the last two digits.
For further examination to determine the impact of SP, each single colony was individually inoculated into 10 mL of CFBM media supplemented with or without 0.04% SP/PET substrate, and the cultures were incubated under shaking conditions (130 rpm) at 25 °C. Changes in culture turbidity were used to estimate possible bacterial growth up to 10 days. A reference culture was conducted in medium without SP or PET substrates to serve as a control group.
2.3. Conserved Gene Amplification and Sequencing for Microbial Identification
Yeast colonies may pose challenges in differentiation when coexisting with bacterial cells in the samples [13]. The microscope proves inefficient and tedious in distinguishing yeasts from bacteria due to the significant size variation of yeast in various environments [13]. To mitigate the risk of inadvertently retrieving yeast cells, two different colony PCRs (polymerase chain reactions) were conducted on each isolate.
A pair of universal 16S rRNA primers, PA (5′-AGAGTTTGATCCTGGCTCAG-3′)/PH (5′-AAGGAGGTGATCCAGCCGCA-3′), was employed to detect the 16S rRNA gene (~1.5 kb) [14].
A pair of 18S rRNA gene primers, funSSUF (5′-TGGAGGGCAAGTCTGGTG-3′)/funSSUR (5′-TCGGCATAGTTTATGGTTAAG-3′) primers, were employed to target the 18S rRNA gene (varied sizes) [15].
Both bacterial and fungal identifications were conducted using the same preparation of the reaction mix and the PCR thermocycle program. Individual target PCRs comprised 12.5 μL of AmpliTaq Gold® 360 Master Mix (Applied Biosystems, Waltham, MA, USA), 0.1 μM of each primer, 2 μL of DNA template, and nuclease-free water, resulting in a final volume of 25 μL. Thermocycling conditions included an initial activation step of 95 °C for 5 min, followed by 35 cycles of 95 °C for 30 s, optimal annealing temperature (55–58 °C) for each primer for 45 s, 72 °C for 90 s, and a final extension of 72 °C for 5 min.
Amplicons were separated by agarose gel electrophoresis and purified using ROCHE high pure PCR product purification Kit for sequencing (Macrogen, Seoul, Korea). The base quality of sequence results was confirmed in Geneious Prime version 2021. The amplified sequences were compared with the GenBank (NCBI) database using the basic local alignment search tool (BLAST) algorithm. Using a calculated percent identity score, specimens were assigned to a genus and species [16]. A sequence was assigned to a species if the best matching reference sequence showed sequence similarity in percentage. Species identification was determined by the BLAST search with the highest bit score of the species.
2.4. Evaluation of Bacterial Growth at Different Levels of Sodium Propionate
The tested isolates were initially streaked on Lignin plates to obtain individual colonies. A single colony was selected and inoculated into 10 mL of CFBM media supplemented with or without 0.04% SP to achieve a Log 7/mL cell suspension after 48 h incubation at 25 °C. Subsequently, this suspension was resuspended and diluted to attain a cell density of Log 5–6/mL in CFBM media, which contained varying levels (0%, 0.01%, 0.02%, 0.04%) of SP (specific substance). This targeted density ensures a sufficient initial cell population, crucial for maintaining cell viability under the low SP concentration of 0.01%. This approach ensures a reliable basis for comparing the effects of different SP levels in cell enumeration.
The cultivation was conducted in 96 well cell plate (200 µL in each well) for 72 h at 25 °C. Three replicates were tested for each isolate in each experiment, which was repeated twice. Cell growth was measured by optical density at 595 nm with a spectrophotometer (MultiScan Go, Thermo Scientific, Waltham, MA, USA), while cell viability was assessed by plate counting on Tryptic soy Agar (enumeration after 48 h culture at 25 °C).
2.5. Statistic Analysis
Grow testing and cell enumeration were conducted in triplicate. The average and standard deviation of growth OD values and viable cell numbers were calculated and analyzed using EXCEL 2026 (Microsoft, Washington, DC, USA). ANOVA One Way tests and post hoc tests (Tukey’s test) and 95% confidence were conducted using the Minitab 19 statistical software (Minitab Inc., Chicago, IL, USA). A “p” value of equal to or less than 0.05 was considered significant.
3. Results
3.1. Bacterial Phenotypic Observations during Culture-Based Isolation
Despite the addition of the antifungal agent 0.04% SP in the liquid culture to suppress fungal growth [6], fungi were still observed on the modified lignin-mineral salt agar plates in our investigation (Figure 2), suggesting that the SP level used was insufficient for effective fungal control. As depicted in Figure 2, plates dominated by mold present challenges in recovering bacterial colonies during subsequent bacterial isolation, with a high incidence of mold contamination. Consequently, these mold-dominant plates from the investigation were not employed for further isolation. Meanwhile, samples exhibiting bacterial dominance (Figure 2b) facilitated the retrieval of pure bacterial-like colonies during subsequent subculturing. After subculturing step, single isolates showing cell growth overnight in the modified CFBM media, as indicated by culture turbidity (OD 595 nm around 0.02–0.2) and viable cell enumeration (up to Log7–9 per 10 mL culture liquid), were considered positive. Among them, 59 isolates exhibited culture turbidity in SP-present groups, including both SP-only and SP plus PET groups. Negative isolates were defined by undetectable culture turbidity (OD 595 nm = 0) and no increase in viable cell numbers.
3.2. Molecular Identification Based on 16S Sequences
Genus or species-level identification of isolates was determined using BLAST analysis on 16S sequences. Out of the 59 isolates, 50 were identified as Pseudomonas spp., while three were identified as Achromobacter spp., 3 as Burkholderia spp., 1 as Rhodococcus spp., and 1 as Enterobacter spp. (refer to Figure 3 and Table 1). To minimise misidentification, 18S PCR was conducted for selected isolates, as yeast cells can sometimes exhibit similar colonisation on agar plates. However, the 18S examination did not detect any fungi among the isolates on amplicon and sequence levels.
The preliminary results from culture turbidity suggested that bacteria exhibited more significant growth on 0.04% SP compared to PET (Table A1). Specifically, three isolates (5601, 7801, and 7802) displayed substantial growth with a turbidity range of 0.15–0.2, contrasting with the rest of the isolates (0.02–0.08). The growth characteristics of these three isolates were further examined in subsequent analyses.
3.3. Bacterial Growth Dependency on Varied SP Concentrations
The results revealed the supportive role of low SP concentrations in promoting the cell growth of tested isolates, evident in sigmoidal growth curves based on OD values. Viable cell counts demonstrated a consistent 2-log/mL increase post-incubation, affirming the effectiveness of SP (as sole carbon source) at sustaining cellular proliferation (Figure 4d and Figure 5c). An additional observation was the isolate 5601, exhibiting distinct colonies with cream white and yellow pigmentation. Cell counts indicated a substantial >2-log increase in white colonies (5601W) compared to a 1-log increase in yellow colonies (5601Y) during mixed cultivation (Figure 4d). This could be seen through consecutive subcultures, leading to the identification of two subtypes, 5601W and 5601Y, subsequently streaked for individual examination.
Further examination revealed isolate 5601W displayed sigmoidal growth, while isolate 5601Y exhibited a non-sigmoidal growth pattern (Figure 5). Remarkably, the concentrations of SP had distinct impacts on cellular growth compared to the mixed culture. Initial cell proliferation occurred within the first 24 h at the three tested SP levels, with no significant differences (p > 0.05) (Figure 5c). In the subsequent 24-h incubation (from day 1 to day 2, as shown in Figure 5c), both isolates continued to grow at 0.04% SP, respectively (p > 0.05). However, at 0.01% and 0.02% SP, while the cell numbers of 5601W isolates decreased, 5601Y continued to grow, reaching significantly higher cell numbers at day 2 (p < 0.05). By the end of the incubation period (day 3), 5601Y isolates surpassed 5601W in cell numbers at all three SP levels (p < 0.05).
In contrast to the varied cell growth observed in the mixed culture, these findings suggest that the 5601W isolate may exert a suppressive effect on the growth of 5601Y, or potentially engage in a symbiotic relationship by utilizing converted nutrients and metabolites from 5601Y during coexistence.
Sigmoid growth patterns were consistently observed in the studied isolates (7801, 7802, and 5601W) at the 0.04% SP level (Figure 4a–c and Figure 5a). However, variations (difference on lag times) were noted across growth curves in triplicate experiment for each isolate (Appendix A Figure A1a,b), preventing the presentation of the growth features in average values. Utilizing growth curves from individual experiments was adequate to confirm the growth features for both sigmoid and non-sigmoid forms, which have been consistently seen in triplicate tests (Appendix A Figure A1).
Identification via 16S marked 5601W as Rhodococcus spp. and 5601Y as Burkholderia spp. In contrast to isolates 7801 and 7802, identified as Burkholderia spp. with sigmoidal growth, 5601Y displayed Burkholderia spp. identification without sigmoidal growth, adding a layer of complexity to the observed microbial responses.
4. Discussion
SP, the sodium salt of propionic acid, is a widely employed food preservative acknowledged for its efficacy in hindering mold and bacterial growth in food items. FDA classifies it as generally recognised as safe (GRAS). While its primary role in food preservation is preventing the growth of spoilage organisms and pathogens, propionate within the range of 0.1–5.0% delays the growth of various microorganisms [17]. Meanwhile, propionate can serve as an energy source for specific bacteria, such as Propionibacterium and Bacteroides, under strictly anaerobic conditions [18].
This study isolated several bacteria from plastic (PET predominant) waste environments with the capability to utilize SP as sole short-chain carbon source in aerobic condition. The discovery introduced a new layer of complexity to our understanding of microbial behaviour in plastic-rich settings, while also indicating the limitation of the adopted culture method involving the addition of the antifungal agent sodium propionate. The SP-utilizing bacteria were primarily retrieved from PET waste samples, specifically from plastic containing samples. However, the preference for SP utilization was not able to be detected in soil samples (No. 1–46), which may be attributed to potential missed results due to fungal dominance on plates unsuitable for investigation.
The initial objective of this study was to identify depolymerising bacteria within a plastic recycling environment using a culture-based approach. Depolymerizing bacteria with easy manipulation, cultivation ability, and model organisms with identified functional enzymes are preferred for research due to their suitability for genetic engineering, experimental studies, and biochemical characterization of depolymerization pathways. Lignin has also been used as potential inductors for biodegradation of synthetic polymers (Polyvinyl Alcohol/Polyethylene), in which the film degrading rate has been significantly improved [19].The potential routes of plastic polymer biodegradation are emerging due to the participation of microbial lignin degrading/modifying enzymes [20,21,22,23]. The waste environment at the sampling location contains natural heteropolymer (plant-derived lignin) and plastic wastes in the soil. Therefore, lignin in the mineral salt medium used in the study [12] is to promote the in vitro growth of polymer-remediating microbes from the soil microbial consortium, to sustain their polymer-degrading capabilities through subculturing under controlled laboratory conditions. Regrettably, depolymerising potential was not evident in this study. The presumed challenge lies in the high crystallinity of the tested plastics, posing difficulty for bacteria to degrade. Also, the investigation didn’t retrieve any bacteria with depolymerizing traits and uncovered a limitation in the adopted culture method, involving the addition of the antifungal agent sodium propionate [6].
However, microbial abundance in the sampling environment were detected, while the investigation didn’t retrieve any bacteria with depolymerizing traits in the adopted culture method [6]. Pseudomonas sp. was the dominant specie detected in this study, while Burkholderia were the second commonly isolated bacteria seen in the investigation. Pseudomonas and Burkholderia species areoxidative, chemoorganotrophic gram-negative bacilli that were previously classified in the same genus [24]. The genus Pseudomonas includes bacteria that are important pathogens causing diseases in humans, animals and plants, with example of Pseudomonas aeruginosa as a significant human pathogen, while Pseudomonas plecoglossicida is responsible for causing infection in fish. Many Burkholderia sp., known as endofungal bacteria, are known as pathogens causing onion soft rot, has evolved into a significant opportunistic pathogen in nosocomial infections [25]. Its prominence in individuals with cystic fibrosis or chronic granulomatous disease is notable [25]. The comprehension of microbial diversity within plastic waste environments could be key evidence for assessing the potential bio-risks posed by plastics serving as carriers for microorganisms in urban waste management zones. The microbial profiles observed within plastic contaminated niches showed discontinuous shifts unrelated to the sampling order, highlighting additional factors influencing microbial settlement or biofilm formation in other study [26]. Mechanical forces such as water-sand emulsion caused by rainfall or tide currents can impact biofilm development [26]. Understanding the long-term processes underlying biofilm formation, particularly, is therefore essential for a comprehensive assessment of the role of plastic as a vector for invasive or pathogenic species. Our preliminary study revealed relevance microbial diverse on the concern, however, further investigation is needed for understanding the dynamic changes, highlighting the importance of assessing the biorisks from this specific environment.
On the other side, the coexistence of Pseudomonas, Rhodococcus, and Burkholderia has been previously reported by other studies, highlighting its significant bioremediation and biodegradation capabilities as consortium in a particular niche [27]. Vaidya et al. [28] identified a bacterial consortium, comprised three bacterial species, Pseudomonas sp. ASDP1, Burkholderia sp. ASDP2, and Rhodococcus sp. ASDP3, was developed from a long-term polluted site, and effectively metabolised high-molecular weight polycyclic aromatic hydrocarbons (pyrene) through phthalic acid pathway. Jiang et al. investigation [29] unveiled BHET-degrading strains, including Rhodococcus biphenylivorans GA1 and Burkholderia sp. EG1, which exhibit the capability to utilize Bis (2-hydroxyethyl) terephthalate (BHET) as their exclusive carbon source. A novel esterase gene, betH, was successfully cloned from strain GA1, encoding an esterase that demonstrates maximal activity at 30 °C and pH 7.0, specifically targeting BHET hydrolysis [29]. The co-cultivation of strain GA1 and strain EG1 resulted in the complete degradation of high concentrations of BHET, mitigating the inhibitory effects of intermediate metabolite ethylene glycol (EG) accumulation on strain GA1 [29]. This innovative approach not only identifies potential microbial strains but also presents a viable strategy for PET bio-upcycling, as BHET is a prominent compound derived from the enzymatic or chemical depolymerization of polyethylene terephthalate (PET). The current evidence underscores the significance of comprehending the catabolism process from long-chain to short-chain carbon sources at the microbial consortium level.
Importantly, the surviving microbes in this study exhibited an unexpected preference for utilising sodium propionate as a short-chain carbon source for growth, possibly diverting the consortia metabolism away from the intended plastic substrate, poly(ethylene terephthalate) or PET. Rhodococcus is known for its potential to utilize various carbon sources and accumulate oils. Rhodococcus grown on propionate generates odd carbon number fatty acids, like C15:0, C17:0, and C17:1 fatty acids [30,31]. Biochemical characterization of the propionate (prp) locus [32] revealed that the 2-methylcitric acid pathway could be an alternative pathway for catabolism of propionic acid, which was detected in B. sacchari [33,34]. Burkholderia sacchari IPT101T has been observed to induce the formation of 2-methylcitrate synthase and 2-methylisocitrate lyase when cultivated in the presence of propionic acid [33]. The prp locus of B. sacchari IPT101T is crucial for propionic acid utilization as a sole carbon source and plays a role in the incorporation of 3-hydroxyvalerate (3HV) into copolyesters [33]. Through cloning and sequencing, five genes (prpR, prpB, prpC, acnM, and ORF5) were identified, showing homology to genes in the prp loci of other gram-negative bacteria [33]. Subsequent analysis revealed lower activity levels of PrpC and PrpB in the propionate-negative mutant IPT189 obtained from IPT101T [33]. This study sheds light on the genetic basis underlying copolyesters accumulated by a B. sacchari mutant, revealing a defect in prpC. Pereira et al. [34] reported the involvement of the 2-methylcitric acid cycle (2MCC) in the catabolism of propionate by Burkholderia sacchari. They generated two B. sacchari mutants, one with a disruption in acnM and the other with deletions in both acnM and prpC, which were unable to grow on propionate [34]. Their findings revealed that an intact 2MCC significantly reduces the bacterial capacity to incorporate 3-hydroxyvalerate (3HV) into a biodegradable copolyester synthesized from carbohydrates and propionate [34]. The efficiency of the mutants in converting propionate to 3HV units (Y3HV/prp) increased substantially from 0.09 g·g–1 to 0.81–0.96 g·g–1, highlighting the essential roles of acnM and prpC in propionate utilization. However, none of the mutations led to the attainment of the maximum theoretical Y3HV/prp (1.35 g·g–1) [34]. Furthermore, increasing propionate concentrations resulted in decreased Y3HV/prp values [34]. These findings support the hypothesis of the existence of alternative propionate catabolic pathways in B. sacchari. While the 2MCC appears to be the predominant pathway, a secondary pathway, yet to be elucidated, becomes more significant at propionate concentrations of 1 g·L–1 or higher. Notably, reducing propionate concentrations can enhance the efficiency of converting propionate to 3HV units, underscoring the role of the 2MCC in this process.
Understanding the metabolic pathways involved in propionate utilization provides insights into the microbial ecology of different environments, such as the soil ecosystems, and industrial processes. The knowledge contributes to our understanding of nutrient cycling and energy metabolism in complex microbial communities. Moreover, propionate is a common intermediate in various metabolic pathways, including those involved in the degradation of organic matter and the synthesis of biodegradable polymers. By elucidating the mechanisms by which bacteria metabolize propionate, researchers can develop strategies to harness these pathways for biotechnological applications, such as bioremediation, biofuel production, and the synthesis of value-added chemicals. Additionally, studying propionate utilization can also shed light on microbial interactions and competition in diverse ecological niches, further expanding our understanding of microbial ecology and evolution. Overall, investigating propionate-utilizing bacteria has broad implications for fields ranging from environmental microbiology to biotechnology and can lead to the development of novel solutions for various environmental and industrial challenges. Therefore, our work will continue to elucidate the possible role of propionate utilizing bacteria in the plastic waste environment.
Relying solely on culturing methods and PCR primer sets introduces potential biases, as not all bacteria or yeasts may yield amplicons. For future research, the inclusion of metagenomic shotgun sequencing is crucial, as it could have provided a comprehensive understanding of the microbial community engaged in carbon source utilization and their associated metabolic pathways. Moreover, the phenomenon of “cross-feeding” within complex ecosystems underscores the need for a broader investigation into microbial interactions. While this study focuses on SP-metabolizing bacteria, future research should consider incorporating metagenomic sequencing to explore a wider spectrum of microbial interactions and metabolic dynamics.
5. Conclusions
Generally, diverse microbial isolates demonstrated a preference for metabolizing sodium propionate, an antifungal agent used in the screening culture method. Preliminary results showed enhanced bacterial growth with 0.04% SP compared to PET, particularly in isolates 5601, 7801, and 7802. The results underscore the supportive role of short chain carbon source (SP) at different low concentrations in promoting cell growth, evident in sigmoidal growth curves. Further examination revealed distinct growth patterns in 5601W and 5601Y, with varying impacts of SP concentrations on cellular growth compared to mixed culture. The detection of propionate-utilizing bacteria in this study revealed the metabolic complex of microbial communities and indicated the challenges associated with bacterial isolation from such environments. The microbial demonstration in the plastic waste environment suggested a possible influence of microbial variance on broader aspects of ecosystem health and functioning with the accumulation of plastic waste. Accordingly, the study uncovered a limitation of the adopted culture method involving the addition of the antifungal agent sodium propionate, as surviving microbes utilized sodium propionate as a short-chain carbon source for growth instead of the intended plastic substrate (polyethylene terephthalate, PET), diverging from the initial objective of screening out depolymerizing bacteria from a plastic recycling environment using a culture-based approach.
Supplementary Materials
The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/applmicrobiol4020059/s1, Table S1: Tukey Pairwise Comparisons for group data of Figure 4d. Grouping Information Using the Tukey Method and 95% Confidence; Table S2: Tukey Pairwise Comparisons for group data of Figure 5c. Grouping Information Using the Tukey Method and 95% Confidence.
Author Contributions
Investigation, S.W.; Resources, F.P.; Data curation, P.S.; Writing—original draft, S.W.; Writing—review & editing, S.W., J.M. and G.B.; Funding acquisition, J.M. and G.B. All authors have read and agreed to the published version of the manuscript.
Funding
This work is funded by AgResearch Ltd., Strategic Science Investment Fund (PRJ0126336, 2022-2023); AgResearch Ltd., FY21 Curiosity Plus Fund (PRJ0331443, 2021-2022).
Data Availability Statement
Data are contained within the article and Supplementary Materials. The data supporting this study are available upon request to the corresponding author.
Acknowledgments
We would like to express our gratitude to Chris Evans, for providing access to and facilitating our sampling activities at the Awapuni Resource Recovery Park in Palmerston North, New Zealand.
Conflicts of Interest
The authors declare no conflicts of interest.
Appendix A
Table A1.
Growth of isolates on 0.04% sodium propionate or PET determined by culture turbidity. Growth is indicated by variant OD values: 0.02–0.1 (+, growth); 0.1–0.2 (++, significant growth); 0 (-, no growth).
Table A1.
Growth of isolates on 0.04% sodium propionate or PET determined by culture turbidity. Growth is indicated by variant OD values: 0.02–0.1 (+, growth); 0.1–0.2 (++, significant growth); 0 (-, no growth).
| Isolates ID | Growth on 0.04% SP | Growth on PET |
|---|---|---|
| 0111 | + | - |
| 0211 | + | - |
| 0304 | + | - |
| 0311 | + | - |
| 0312 | + | - |
| 0321 | + | - |
| 0601 | + | - |
| 1302 | + | - |
| 1304 | + | - |
| 1903 | + | - |
| 0811 | + | - |
| 0821 | + | - |
| 0831 | + | - |
| 0832 | + | - |
| 1311 | + | - |
| 1421 | + | - |
| 1911 | + | - |
| 2303 | + | - |
| 2303 | + | - |
| 2511 | + | - |
| 2521 | + | - |
| 2801 | + | - |
| 2901 | + | - |
| 2902 | + | - |
| 3301 | + | - |
| 3303 | + | - |
| 3321 | + | - |
| 3521 | + | - |
| 3701 | + | - |
| 3703 | + | - |
| 4112 | + | - |
| 4121 | + | - |
| 4221 | + | - |
| 4401 | + | - |
| 4402 | + | - |
| 5001 | + | - |
| 5101 | + | - |
| 5501 | + | - |
| 5502 | + | - |
| 5601W | ++ | - |
| 5601Y | ++ | - |
| 6001 | + | - |
| 6002 | + | - |
| 6302 | + | - |
| 6303 | + | - |
| 6402 | + | - |
| 6412 | + | - |
| 6502 | + | - |
| 6712 | + | - |
| 6911 | + | - |
| 6931 | + | - |
| 7401 | + | - |
| 7402 | + | - |
| 7501 | + | - |
| 7502 | + | - |
| 7801 | ++ | - |
| 7802 | ++ | - |
| 8321 | + | - |
| 8322 | + | - |
| 8803 | + | - |
| 8821 | + | - |
Figure A1.
The growth curves in triplicate tests. (a) growth curve of isolate 5601W; (b) growth curve of isolate 5601Y.
Figure A1.
The growth curves in triplicate tests. (a) growth curve of isolate 5601W; (b) growth curve of isolate 5601Y.
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Figure 1.
Flow chart of microbial cultivation conducted in the study.
Figure 2.
Microbial enrichment examples from 1st generation of sample isolation. (a) Sample No. 63; (b) Sample No. 83; (c) Sample No. 68; (d) Sample No. 56.
Figure 2.
Microbial enrichment examples from 1st generation of sample isolation. (a) Sample No. 63; (b) Sample No. 83; (c) Sample No. 68; (d) Sample No. 56.
Figure 3.
Percentage of culturable bacteria identified using 16S PCR amplicons from the sampling survey.
Figure 3.
Percentage of culturable bacteria identified using 16S PCR amplicons from the sampling survey.
Figure 4.
Bacterial growth over 72 h incubation in the presence of different levels of SP. (a) Growth curve of 5601 mixed isolates (5601W and 5601Y); (b) Growth curve of isolate 7801; (c) Growth curve of isolate 7802; (d) Density of viable cells in cultivation with 0.04% SP at the endpoint of the growth curve. Control, initial viable cell counts at the start of the 72-h incubation test. Sample groups that did not share a letter were significantly different (p < 0.05). Grouping Information Using the Tukey Method and 95% Confidence has been provided in Table S1.
Figure 4.
Bacterial growth over 72 h incubation in the presence of different levels of SP. (a) Growth curve of 5601 mixed isolates (5601W and 5601Y); (b) Growth curve of isolate 7801; (c) Growth curve of isolate 7802; (d) Density of viable cells in cultivation with 0.04% SP at the endpoint of the growth curve. Control, initial viable cell counts at the start of the 72-h incubation test. Sample groups that did not share a letter were significantly different (p < 0.05). Grouping Information Using the Tukey Method and 95% Confidence has been provided in Table S1.
Figure 5.
Growth characteristics of 5601W and 5601Y isolates. (a) Growth curve of 5601W. (b) Growth curve of 5601Y. (c) Cell enumeration at incubation times: D0 (0 h), D1 (24 h), D2 (48 h), and D3 (72 h). Significant differences were detected between no-SP group and SP group at different levels (p < 0.05). Grouping Information Using the Tukey Method and 95% Confidence has been provided in Table S2.
Figure 5.
Growth characteristics of 5601W and 5601Y isolates. (a) Growth curve of 5601W. (b) Growth curve of 5601Y. (c) Cell enumeration at incubation times: D0 (0 h), D1 (24 h), D2 (48 h), and D3 (72 h). Significant differences were detected between no-SP group and SP group at different levels (p < 0.05). Grouping Information Using the Tukey Method and 95% Confidence has been provided in Table S2.
Table 1.
Identification of Isolates Based on 16S PCR Amplicons.
| Isolates ID | Sequencing Result: BLAST 0.7–1.0 kb E = 0 | Colony Feature | |
|---|---|---|---|
| Sequenced from the PA End (Forward Sequencing) | Sequenced from the PH End (Reverse Sequencing) | ||
| 0111 | Pseudomonas nitritireducens | Pseudomonas knackmussi | Cream coloured colonies |
| 0211 | Pseudomonas monteilii | Pseudomonas monteilii | Cream coloured colonies |
| 0304 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Mucoid colonies |
| 0311 | Pseudomonas aestus | Pseudomonas aestus | Cream coloured colonies |
| 0312 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 0321 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Cream coloured colonies |
| 0601 | Pseudomonas multiresinivorans | Pseudomonas knackmussii | Mucoid colonies |
| 1302 | Pseudomonas monteilii | Pseudomonas putida | Brown colonies |
| 1304 | Pseudomonas helmanticensis | Pseudomonas helmanticensis | Cream coloured colonies |
| 1903 | Achromobacter kerstersii | Achromobacter deleyi | Cream coloured colonies |
| 0811 | Pseudomonas aeruginosa | Pseudomonas aeruginosa | Mucoid colonies |
| 0821 | Pseudomonas aeruginosa | Pseudomonas aeruginosa | Cream coloured colonies |
| 0831 | Pseudomonas aeruginosa | Pseudomonas aeruginosa | Mucoid colonies |
| 0832 | Pseudomonas plecoglossicida | Pseudomonas plecoglossicida | Mucoid colonies |
| 1311 | Pseudomonas koreensis | Pseudomonas koreensis | Mucoid colonies |
| 1421 | Rhodococcus sp. | Rhodococcus sp. | Cream coloured colonies |
| 1911 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Cream coloured colonies |
| 2303 | Pseudomonas putida | Pseudomonas putida | Cream coloured colonies |
| 2303 | Pseudomonas putida | Pseudomonas putida | Cream coloured colonies |
| 2511 | Pseudomonas monteilii | Pseudomonas monteilii | Cream coloured colonies |
| 2521 | Pseudomonas monteilii | Pseudomonas putida | Mucoid colonies |
| 2801 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Cream coloured colonies |
| 2901 | Pseudomonas plecoglossicida | Pseudomonas cremoricolorata | Cream coloured colonies |
| 2902 | Pseudomonas cremoricolorata | Pseudomonas knackmussii | Mucoid colonies |
| 3301 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 3303 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Mucoid colonies |
| 3321 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Cream coloured colonies |
| 3521 | Pseudomonas monteilii | Pseudomonas plecoglossicida | Cream coloured colonies |
| 3701 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Mucoid colonies |
| 3703 | Pseudomonas nitroreducens | Pseudomonas knackmussii | Cream coloured colonies |
| 4112 | Cupriavidus metallidurans | Cupriavidus metallidurans | Cream coloured colonies |
| 4121 | Pseudomonas monteilii | Pseudomonas monteilii | Cream coloured colonies |
| 4221 | Achromobacter insuavis | Achromobacter deleyi | Cream coloured colonies |
| 4401 | Pseudomonas nitroreducens | Pseudomonas citronellolis | Mucoid colonies |
| 4402 | Pseudomonas plecoglossicida | Pseudomonas cremoricolorata | Cream coloured colonies |
| 5001 | Pseudomonas protegens | Pseudomonas protegens | Cream coloured colonies |
| 5101 | Pseudomonas putida | Pseudomonas cremoricolorata | Cream coloured colonies |
| 5501 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Cream coloured colonies |
| 5502 | Enterobacter ludwigii | Enterobacter ludwigii | Mucoid colonies |
| 5601W | Rhodococcus qingshengii | Rhodococcus qingshengii | Creamy white colonies |
| 5601Y | Burkholderia cenocepacia | Burkholderia diffusa | Yellow colonies |
| 6001 | Pseudomonas nitroreducens | Pseudomonas knackmussii | Mucoid colonies |
| 6002 | Pseudomonas nitroreducens | Pseudomonas knackmussii | Mucoid colonies |
| 6302 | Pseudomonas mediterranea | Pseudomonas corrugata | Cream coloured colonies |
| 6303 | Achromobacter kerstersii | Achromobacter deleyi | Cream coloured colonies |
| 6402 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Mucoid colonies |
| 6412 | Pseudomonas nitroreducens | Pseudomonas nitroreducens | Cream coloured colonies |
| 6502 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 6712 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Cream coloured colonies |
| 6911 | Achromobacter deleyi | Achromobacter deleyi | Mucoid colonies |
| 6931 | Pseudomonas nitritireducens | Pseudomonas knackmussii | Cream coloured colonies |
| 7401 | Pseudomonas monteilii | Pseudomonas monteilii | Cream coloured colonies |
| 7402 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 7501 | Pseudomonas nitroreducens | Pseudomonas knackmussii | Mucoid colonies |
| 7502 | Pseudomonas monteilii | Pseudomonas putida | Mucoid colonies |
| 7801 | Burkholderia ambifaria | Burkholderia ambifaria | Mucoid colonies |
| 7802 | Burkholderia pyrrocinia | Burkholderia metallica | Light yellow colonies |
| 8321 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 8322 | Pseudomonas monteilii | Pseudomonas putida | Cream coloured colonies |
| 8803 | Pseudomonas nitroreducens | Pseudomonas knackmussii | Mucoid colonies |
| 8821 | Pseudomonas knackmussi | Pseudomonas knackmussi | Cream coloured colonies |
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Wu, S.; Subharat, P.; Palevich, F.; Mills, J.; Brightwell, G. The Detection of Propionate Utilization by Bacteria Isolated from a Plastic Recycling Site. Appl. Microbiol. 2024, 4, 856-874. https://doi.org/10.3390/applmicrobiol4020059
AMA Style
Wu S, Subharat P, Palevich F, Mills J, Brightwell G. The Detection of Propionate Utilization by Bacteria Isolated from a Plastic Recycling Site. Applied Microbiology. 2024; 4(2):856-874. https://doi.org/10.3390/applmicrobiol4020059
Chicago/Turabian StyleWu, Shuyan, Pornchanok Subharat, Faith Palevich, John Mills, and Gale Brightwell. 2024. "The Detection of Propionate Utilization by Bacteria Isolated from a Plastic Recycling Site" Applied Microbiology 4, no. 2: 856-874. https://doi.org/10.3390/applmicrobiol4020059
APA StyleWu, S., Subharat, P., Palevich, F., Mills, J., & Brightwell, G. (2024). The Detection of Propionate Utilization by Bacteria Isolated from a Plastic Recycling Site. Applied Microbiology, 4(2), 856-874. https://doi.org/10.3390/applmicrobiol4020059
