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PHA synthesis by bacteria using low density polyethylene, starches and cellulosics§
Korean J. Microbiol. 2021;57(3):183-196
Published online September 30, 2021
© 2021 The Microbiological Society of Korea.

Rafeya Sohail, Rida Batool, and Nazia Jamil*

Institute of Microbiology and Molecular Genetics, University of the Punjab, Quaid-e-Azam Campus, Lahore 54590, Punjab, Pakistan
Correspondence to: E-mail:
§Supplemental material for this article may be found at
Received February 26, 2021; Revised August 27, 2021; Accepted August 28, 2021.
This study was conducted for optimization of polyhydroxyalkanoate (PHA) production using starches, cellulosics as carbon sources and mixed cultures. The capability of PHA producers to degrade and utilize LDPE (Low-density polyethylene) by carbon assimilation was also studied. Tannery effluents were used for isolation of PHA producers and screened using Nile blue and Nile red supplemented media. PHA production studies showed optimal PHA production occurred during 24 to 48 h time. Maximum PHA production was obtained at 24 h. After 48 h, gradual decline in PHA production was observed. Best candidate for PHA production was found to be strain PWF, based on number, size of granules inside cell, and %PHA production. PHA production was also optimized by use of mixed culture. Among starch-based sources, highest production rates were on pure starch. Among cellulose-based sources, maximum production rates were on dry wood powder instead of pure crystalline cellulose. In contrast, wood extract and wood shavings showed more pronounced PHA production rates, comparable to production on dry wood powder. LDPE utilization as sole carbon source in selective media showed that PHA producers were able to degrade synthetic plastic. Focus of future studies can be PHA production using these sources on industrial scale.
Keywords : Bacillus tequilensis, Pseudomonas aeruginosa, LDPE, polyhydroxyalkanoate, tannery

Bioplastics have gained an integral position in our daily lives since 1940s (Radecka et al., 2016). They have become popular due to their lightweight, strength, durability, elasticity, and easy utilization (Reddy et al., 2003). Biodegradable plastics such as polyhydroxyalkanoates made up of recyclable, easily available materials, are biocompatible and have less residence time in environment (Kubowicz and Booth, 2017). Polyhydroxyalkanoates (PHAs) optically active, piezoelectric, highly crystalline, and isotactic thermoplastics, showing similarity in properties to petrochemical based plastics can serve as their potential replacements (Radecka et al., 2016). PHA are accumulated intracellularly by bacteria cell as inclusion bodies and account for levels as high as 90% of cell dry weight (Reddy et al., 2003). Under extreme conditions, PHA serve not only as storage inside vegetative cells (Aljuraifani et al., 2019) but also as protection against osmotic shock, desiccation, thermal stress, UV irradiation and oxidative stress (Koller et al., 2015). Physical properties of PHA are dependent on their chemical composition. Short chain length PHA (scl-PHA) resemble conventional synthetic plastics more closely (Shah and Vasava, 2019). Medium chain length PHA (mcl-PHA) have higher elasticity being semi-crystalline and rubbery as compared to scl-PHA. Combination of different monomeric subunits also influences the thermal properties of PHA (Sharma et al., 2017; Aljuraifani et al., 2019). Due to the stereospecificity of PHA biosynthetic enzymes, all monomers are in ᴅ-configuration. Furthermore, depending on the microbial strain, carbon supply, culture conditions and nutrient regime provided during biosynthesis, PHA chains contain up to 102–105 3-hydroxyalkanoic acid monomers (Koller et al., 2015). Numerous pha genes encode different classes of PHA synthase enzymes and regulate PHA biosynthesis pathways. The diversity in pha loci shows the evolution of gene cluster organization in PHA operon structure over time. In various microbes, phaC gene is clustered with genes phaZ, phbA, or phaJ etc. that supply monomers (Solaiman et al., 2000; Numata et al., 2013). Both carbon source feed and diversity of bacterial strains affect the production and composition of PHA. Economically, the cost of feed and polymer recovery processes are the main issues slowing the pace of PHA research.

Tannery effluents containing high concentrations of biodegradable organic acids, antioxidants, treatment chemicals etc. are an excellent, cost-effective source for isolation of diverse bacterial strains capable of producing high value biopolymers (David et al., 2015). Whereas, utilization of substrates derived from natural, renewable, and economically cheap resources such as starches and cellulosics serves as a cost-effective alternative approach. Low value substrates including but not restricted to agro-forestry residues and industrial by byproducts such as cellulose and starch based carbon sources are also being repurposed for large scale PHA production (Queirós et al., 2015). Starch can be extracted from many crop plants such as rice, potato, corn etc (Nawrath et al., 1995). Agro-industrial bioprocesses yield high titers of starches (Shamala et al., 2012). While cellulosics such as wood shavings, sawdust, waste dried wood etc. are produced in tens of millions of tons per annuum by paper industries, furniture factories and other wood processing industrial sectors (Charis et al., 2019). Utilization of starches and cellulosics as carbon sources during production of PHA has high potential as an economic, environment friendly wood waste management and recycling innovative. Additionally, eco-biotechnology methodologies combining industrial biotechnology with environmental biotechnology i.e., employing the use of mixed microbial cultures. During the last decade, industrial production of PHA has been optimized by utilizing many different cost-saving strategies such as use of mixed microbial cultures (Queirós et al., 2015). However, so far, mixed cultures have been utilized only sparingly for biochemical production (Johnson et al., 2009).

Lately, the excessive use of non-degradable, synthetic plastics has made their eventual disposal quite difficult. In Europe, China, and USA, plastic wastes account for 7, 14, and 11.8% out of total generated municipal solid waste, respectively (Radecka et al., 2016). Recycling and disposal of plastics also raises many issues, such as need for landfills, incineration sites, dumping in oceans etc (Fossi et al., 2020). Around 60 to 80% of all debris in water bodies accounts for plastic waste. Microplastics; fragments of plastics less than 5 mm have infiltrated the food chain by entangled with sea life form by ingestion (Johnston et al., 2017). Even polyethylene (PE), most manufactured chemosynthetic plastic – about 29% of global production – has only 10% recycling, presently. The impediments in degradation of plastics wastes are a public concern eventually leading to environment pollution crisis (Verma et al., 2016). In view of all this, the demand for alternate recycling methodologies and production of eco-friendly plastics is increasing (Radecka et al., 2016; Muniyasamy et al., 2019). The hydrocarbon carbon rich backbone of many different kinds of plastics and plastic based waste products such as low-density polyethylene (LDPE) has potential for utilization in microbial metabolism (Radecka et al., 2016; Johnston et al., 2017). LDPE can, therefore, be bio-converted to value-added biopolymers such as PHA by utilization as a carbon source during growth and metabolism. This innovative way of synthetic polymer utilization for biopolymer production introduces a novel solution to plastic waste accumulation and degradation.

In this study, tannery effluent samples were collected for isolation of PHA producing bacteria. Bacterial isolates were screened for PHA production and best PHA producer was identified by 16S rRNA gene sequencing. The use of starch-based, cellulose-based carbon sources as well as known mixed culture was explored. Furthermore, low density polyethylene was used to understand the degradation and utilization of LDPE by PHA producers.

Materials and Methods

Isolation of strain DL3 and revival of produced water strains

Tannery samples were collected aseptically in sterile plastic containers from two different sites i.e., liming and deliming sites for tanning of buffalo hides. Samples were properly labelled with site, time, date of collection and collector’s name. During transport to laboratory, samples were held at 4°C. Both samples were analyzed appropriately and characterization parameters such as temperature, pH, color, texture, and odor of sample were noted precisely. Sample dilutions were prepared by serial dilution method described by da Silva et al. (2018) and 10 µl volume of each dilution was spread on LB agar plates, aseptically. Spread plates were incubated for 24 h at 37°C. Colony counter was used to estimate viable colony count using the methodology of Brown and Smith (2014), and Cappuccino and Sherman (2014). CFU/ml was calculated using formula:

CFU/ml=Number of coloniesDilution Factor×Volume plated

Discrete colonies with prominent distinguishing traits were selected from spread plates and sub cultured by quadrant streak method as described by Cappuccino and Sherman (2014) to obtain pure cultures. Streak plates were incubated for 24 h at 37°C. After incubation at 37°C for 24 h, distinguishing morphological characters and colonial parameters of pure colonies, for instance size, shape, texture, margins, appearance, elevation, pigmentation, and optical property etc. were observed and recorded in tabular form (Cappuccino and Sherman, 2014). Microscopic characterization of tannery isolates was done with the purpose of differentiation, identification, and visualization of bacterial specimen by Gram staining, spore staining, and capsule staining. Microscopic measurements of bacterial cells of tannery isolates were observed and recorded in tabular form (Brown and Smith, 2014; Cappuccino and Sherman, 2014). Biochemically diverse characteristics of tannery isolates were confirmed employing distinctive qualitative tests such as catalase activity test, citrate utilization test, DNase test, motility test, oxidase test, starch hydrolysis test, and urease activity test etc. All results were recorded in tabular form.

Stock cultures of bacterial strains PWA; Bacillus subtilis (MH142143), PWC; Pseudomonas aeruginosa (MH142144), PWF; Bacillus tequilensis (MH142145), and PWG; Bacillus safensis (MH142146) – isolated from produced water (hydrocarbon rich water, trapped in underground channels, brought up during drilling) in a previous study – were taken from Culture Collection of Research Lab II, Department of Microbiology and Molecular Genetics, University of the Punjab, Lahore, Pakistan (Sohail and Jamil, 2020; Sohail et al., 2020). Cultures were quadrant streaked on Nutrient Agar and incubated for 24 h at 37°C (Cappuccino and Sherman, 2014). Fresh cultures were maintained by streaking discrete colonies on Nutrient Agar and incubating for 24 h at 37°C.

Screening of polyhydroxyalkanoate (PHA) producers

Nile Blue (Spiekermann et al., 1999) or Nile Red (Oshiki et al., 2011; Phanse et al., 2011) supplemented selective media; PHA detection media was used for screening of PHA producers. Polyhydroxyalkanoate detection media containing MgSO4•7H2O, 1.2 g/L; K2HPO4, 13.3 g/L; (NH4)2SO4, 2.0 g/L; citric acid, 1.7 g/L; agar, 15.0 g/L; and trace elements solution; 10.0 ml was prepared and pH was adjusted to 6.84. Trace elements solution was prepared by dissolving (NH4)6Mo7O24, 0.1 g/L; CaCl2•2H2O, 2.0 g/L; CuSO4•5H2O, 1.0 g/L; MnSO4•5H2O, 0.5 g/L; FeSO4•7H2O, 10.0 g/L; Na2B4O7•10H2O, 0.23 g/L; ZnSO4•7H2O, 2.25 g/L; and 35% HCl, 10 ml/L in DW and adjusting pH to 7.0 (Ostle and Holt, 1982; Kitamura and Doi, 1994). PHA binds with Nile Blue or Nile Red dye and bacterial culture with PHA inside cells fluoresces under UV. PHA detection media supplemented with Nile Blue or Nile red dye was prepared and dispensed in petri plates. Plates were streaked with bacterial culture, and incubated for 24 h at 37°C. Culture plates were visualized under UV illuminator, after incubation, for absence or presence of fluorescence (Greenspan et al., 1985). For further confirmation of PHA production, Sudan Black staining was done for visualization of PHA granules inside bacterial cells (Rehman et al., 2007; Chaudhry et al., 2011). Bacterial smears of all tannery isolates were prepared, slides were labelled accordingly, smears were stained for 15 min with Sudan Black dye. Stained slides were washed using xylene for decolorization and washed again using distilled water to remove xylene. Slides were counterstained for 1 min with safranin, washed using distilled water and air dried. Stained smears were observed, using 4×, 10×, 40×, and 100× magnification, under light microscope (Serafim et al., 2002).

16S rRNA sequencing of tannery bacteria

Bacterial strain was streaked on Nutrient agar plates and incubated for 14 h at 37°C. 16S rRNA gene sequencing was done commercially by Macrogen Inc ( Complementary sequence was obtained by converting reverse sequence using Chromas 2.6.6. software (McCarthy, 1996). Forward and reverse sequence alignment and assembly was done using Cap3 software to obtain consensus sequence (Huang and Madan, 1999). BlastN was used to inspect maximum homology of sequence against GenBank and sequence was submitted to GenBank (Dumontier and Hogue, 2002). Evolutionary relationship for sequence similarity to similar records was mapped out, constructing dendrograms by neighbor joining method using MEGA 10.1.7 (Kumar et al., 2008, 2018).

Polyhydroxyalkanoate production

Polyhydroxyalkanoate (PHA) detection media supplemented with 2% glucose as carbon source was prepared and 200 ml media was dispensed in 500 ml Erlenmeyer flasks. Preliminary 10% culture of PHA producers was added to PHA detection media aseptically. Culture media was incubated for 96 h at 37°C in a rotary incubator. Each analysis was conducted in triplicate. Growth kinetics of PHA producers were evaluated by measuring optical densities of bacterial cultures at regular intervals using IRMECO U2020 UV-Vis spectrophotometer at 600 nm (Teeka et al., 2010). Growth kinetics of PHA producers were analyzed by plotting optical densities against y-axis and time against x-axis. Optical density values were mean of data collected during triplicate experiments. Standard error and standard deviation were calculated. After regular intervals i.e., 24, 48, 72, and 96 h, 40 ml volume of culture media was taken and centrifuged at 6,000 rpm for 15 min to obtain pellet containing cell biomass. PHA production statistics were analyzed by plotting time against x-axis, biomass against primary y-axis and percentage PHA against secondary y-axis. All values were mean of experiments conducted in triplicate. Standard error and standard deviation (SD) were calculated.

Extraction of polyhydroxyalkanoates

Culture broth was aseptically collected in 50.0 ml falcons and falcons were centrifuged at 6,000 rpm for 15 min, to harvest bacterial biomass pellet. Pellet was air-dried. Dry weight of biomass was recorded. Dried pellet was resuspended in 5.0 ml DW to rehydrate bacterial cells. Rehydrated pellet was then treated with 0.25% SDS (at 25°C and pH 10.0) for 15 min and 5.25% sodium hypochlorite (at room temperature and pH 10.0) for 5 min, to achieve complete cell lysis. Cell lysate – containing cell debris and intracellular PHA – was centrifuged at 6,000 rpm for 15 min and supernatant was discarded. Pellet was washed using ice cold acetone and centrifuged again at 6,000 rpm for 15 min, for thorough removal of cell debris from intracellular PHA. Supernatant was discarded. Pellet containing crude PHA and cell debris was transferred to 100 ml glass Erlenmeyer flasks. Chloroform (10 volumes the volume of pellet) was dispensed in the flasks to dissolve pellet. The chloroform/biopolymer mixture was incubated on shaker, at room temperature for 48 h, to dissolve PHA. After 48 h, biomass layer was separated from chloroform by filtration (Whatman Grade 1 filter paper). Chloroform/PHA mixture was collected in glass vials. Solvent-cast dry films of PHA were obtained after evaporation of chloroform. Dry weight PHA films was recorded (Van Doan et al., 2015). Percentage of PHA was calculated using formula:

%PHA=Weight of PHAWeight of biomass×100

Optimization of polyhydroxyalkanoate production using different carbon sources

Polyhydroxyalkanoate (PHA) production was optimized using different structurally diverse, non-fossil fuel based, eco-friendly, renewable carbon sources such as starch based polysaccharides e.g., pure starch, rice starch, potato starch, corn starch, cellulose based polysaccharides e.g., dry wood powder, wood extract, wood shaving, crystalline cellulose, and polyolefin resin based carbon sources LDPE. Carbon sources were used as 2% supplementation in PHA detection media. Media was incubated for 96 h at 37°C. Each study was conducted in triplicate. Growth kinetics of PHA producers were evaluated (Teeka et al., 2010) and PHA production statistics for each carbon source by PHA producers were measured (Van Doan et al., 2015).

Carbon sources based on starch

Pure starch, rice starch, potato starch and corn starch were used as carbon source for optimization of PHA production. Pure starch was taken as control against extracted starches. Rice starch, potato starch, and corn starch were extracted using the methodology described by with some modifications (Schoch and Maywald, 1968). Rice, potato, and corn were weighed at 100.0 g, washed thoroughly, and added to 100.0 ml of autoclaved distilled water in separate flasks. Flasks were sealed and kept at room temperature for a week on shaker. Mixture was ground in a mortar and filtered. DW (100.0 ml) was added to mixture, stirred thoroughly, ground in mortar, and filtered. Above step was repeated three times more and 500.0 ml filtrate was obtained. Filtrate was allowed to settle down and water was decanted. DW (100.0 ml) was added to remove impurities. Mixture was agitated softly, and water containing impurities was removed. Above step was repeated three to four times to achieve a white suspension (Marichelvam et al., 2019). Suspension was centrifuged and pellet was dried in hot air oven at 45°C to obtain starch extract as white powder (Santana et al., 2018). Starch extracts were not subjected to any further chemical or physical treatments for separation of other crude chemical complexes.

Carbon sources based on cellulose

Utilization of unprocessed cellulosics such as dry wood powder, wood extract and wood shavings for PHA production was explored as a low-cost innovative (Koller et al., 2015). PHA production on pure cellulose, in form of crystalline cellulose, was taken as archetype against unprocessed cellulosics. Waste dry wood and wood shavings were obtained from wood industries, and separated from other contaminants. Dry wood was shredded and crushed to obtain powder. To prepare wood extract, 100 g of wood was obtained by cutting a section of Syzygium cumini (Jamun) tree and submerged it in 1,000 ml of distilled water for a week. After a week, mixture was heated at 65°C for 3 h, and decanted. Wood extract was further purified by filtration (Whatman grade 1 filter paper). Wood extract was not subjected to any further treatments for separation of hemicellulose, lignin, and other crude chemical complexes. Crystalline cellulose was dissolved in 8% NaOH/water at -4°C to obtain a homogenous solution (Egal et al., 2008). In optimization experiments, all cellulose based sources were used as a 2% carbon source supplement in PHA production media.

Preparation of low density polyethylene (LDPE)

Recyclable waste LDPE was obtained and scanned for separation from other contaminants. LDPE sheets were shredded into small pieces of 1 cm × 1 cm size. Pieces were sealed in a glass petri plate and autoclaved for 15 min at 121°C. LDPE pieces were stored at room temperature for use without any further processing. In optimization experiments, LDPE was used as a 2% carbon source supplement in PHA production media.

Optimization of polyhydroxyalkanoate production using mixed culture

Polyhydroxyalkanoate (PHA) production was optimized using mixed bacterial culture of strains PWA, PWC, PWF, PWG and DL3. PHA detection media containing each carbon source were prepared. Mixed culture was added to each media and was incubated for 96 h at 37°C. Each study was conducted in triplicate. Growth kinetics (Teeka et al., 2010) and PHA production statistics by mixed culture for each carbon source were observed and measured, respectively (Van Doan et al., 2015).

PCR amplification and sequencing of gene phaC1 and phaC2

Genes phaC and phaC1 were amplified and sequenced to corroborate the biological PHA production capability bacterial strains with highest PHA production. Gene phaC1 of Pseudomonas was amplified using 179-L (ACAGATCAACAAGTTCTACA TCTTCGAC) and 179-R (GGTGTTGTCGTTCCAGTAGAG GATGTC) primers to corroborate PHA production. PCR program was run for 30 cycles consisting of denaturation at 95°C for 1 min, annealing at 56°C for 1 min, and extension at 72°C for 2 min. Initial denaturation was done at 95°C for 10 min, and final extension was done at 72°C for 5 min. PhaC gene operon of Pseudomonas comprising of phaC1 and phaC2 genes was amplified using ORF1 (CCAYGACAGCGGCCTGTTCACCTG) and 179-R primers to obtain amplicons starting at orf. Initial denaturation was at 95°C for 10 min while final extension was at 72°C for 10 min. PCR program was run for 33 cycles; denaturation at 95°C for 30 sec, annealing at 60°C for 45 sec, and extension at 72°C for 1 min. Primers phal-1-Forward (CARACNTA[Y][Y] TNGCNTGGMGNAARGA) and phal-2-Reverse (TARTTRTTNACCCARTARTTCCADAT) were used for gene phaC amplification of Bacillus and Pseudomonas, to obtain high molecular weight PCR product. Initial denaturation was at 95°C for 5 min, and final extension was at 72°C for 5 min. For Bacillus, PCR program was run for 35 cycles; denaturation at 95°C for 30 sec, annealing at 45.8°C for 45 sec, and extension at 72°C for 1 min. For Pseudomonas, PCR program was run for 30 cycles; denaturation at 95°C for 30 sec, annealing at 48°C for 1 min, and extension at 72°C for 30 sec.

PHA synthase gene phaC of Pseudomonas aeruginosa amplified using phal-1-Forward and phal-2-Reverse primers was purified using Vivantis GF-1 Nucleic Acid Extraction kit and sequenced by Sanger dideoxy sequencing. BlastN was used to inspect similarity index of sequence against GenBank. Nucleotide sequence was aligned with PHA synthase gene sequences of closely related taxa by ClustalW (Kumar et al., 2018). Sequence was submitted to GenBank (Dumontier and Hogue, 2002). BioEdit 7.2.6 software was used to align the sequence with related taxa and ExPASy was used to translate sequence (Artimo et al., 2012). Sequence was analyzed against NCBI conserved domain database for determination of conserved domains (Marchler-Bauer et al., 2015).


Sample collection and characterization of isolated strains

Samples collected from liming and deliming sites of tannery were labelled as B 1 and B 2, respectively. Sample B 1 was grayish brown. Sample temperature and pH were noted as 28°C and 12.5, respectively. Sample B 2 had 8.4 pH and 35°C temperature with brown coloration. Burnt leather odor was recorded for both samples. For sample B 1, highest CFU/ml (1.59 × 101) was in dilution 10-1 with 159 colonies, lowest CFU/ml (1.21 × 10-3) was in dilution 10-5 with 121 colonies while dilution 10-7 had 0 colonies. For sample B 2, highest CFU/ml (1.23 × 101) was in dilution 10-1 with 123 colonies while lowest CFU/ml (9.0 × 10-7) was in dilution 10-7 with 9 colonies. Fifteen discrete bacterial colonies were selected from sample B 1 and labelled as Liming strains L1–L15. Ten discrete bacterial colonies were selected from sample B 2 and labelled as Deliming strains DL1–DL10. Isolated bacterial colonies were characterized morphologically (Supplementary data Table S1). Out of 25 bacterial strains, one colony was gram negative cocci (pink cell walls), five colonies were gram positive cocci and the others were gram positive bacilli (purple cell walls). 15 bacterial strains were spore formers (green spores visualized against pink background) while 10 were non spore former. Fourteen bacterial strains were capsular (clear capsules against dark back ground) while other 11 were non capsular (Supplementary data Table S2). Biochemical characterization results of bacteria isolated from tannery were recorded in tabular form. Out of 25 bacteria, 21 were catalase positive (bubble formation) while four were catalase negative. All 25 strains were oxidase positive (change in color; filter paper turned blue). Fourteen strains were DNase positive (presence of yellowish halo around colonies) while 11 strains were DNase negative. 4 strains were urease positive (color change) while 21 strains were urease negative. Nineteen strains were citrate positive (color change) while 6 strains were negative. Twenty strains were positive for starch hydrolysis (presence of colorless halo around colonies) while 5 strains were negative. 16 strains were motile (deviation of growth from streak line) while 9 were non motile (Supplementary data Table S3).

Screening for polyhydroxyalkanoate (PHA) producers

Among all tannery isolated bacterial strains, only strain DL3 (isolated from deliming sample) was able to grow on PHA detection media supplemented with Nile blue A or Nile red dye and gave fluorescence under UV. Other tannery isolated strains showed minimal or no growth on PHA detection media and did not give fluorescence under UV. On Sudan Black B staining, black PHA granules were observed against pink background in strain DL3, in which almost 40–50% of cellular volume was found to be occupied by PHA granules. Strain DL3 was identified as Bacillus subtilis by 16S rRNA gene sequencing and sequence was submitted to GenBank under accession number MT043898 and evolutionary history of strain DL3 with closely related taxa was inferred using MEGA 10.1.7 by neighbor joining method. Optimal tree is shown with branch length sum = 0.00623495. The tree was drawn to scale using Maximum Composite Likelihood method to compute evolutionary distances (Fig. 1).

Fig. 1. Evolutionary relationship of closely related taxa with strain DL3. The evolutionary distances between strain DL3 and closely related taxa are shown by drawing a dendrogram to scale using Maximum Composite Likelihood method. The sum of branch length is 0.00623495.

Polyhydroxyalkanoate production

Overall highest PHA production was shown by mixed culture, followed by bacterial strains PWF (Bacillus tequilensis), PWC (Pseudomonas aeruginosa), PWA (Bacillus subtilis), and PWG (Bacillus safensis). While lowest PHA production was by strain DL3 (Bacillus subtilis). In terms of carbon source, highest growth rate, biomass and PHA production was on glucose, followed by that on pure starch, rice starch, potato starch, dry wood powder, corn starch, wood extract, wood shaving, crystalline cellulose, and LDPE. Optimum PHA production was during 24 to 48 h while maximum production was obtained at 24 h. Overall, PHA production increased from 24-h mark to 48-h mark. On all carbon sources, after 24 h, highest biomass was by PWC while highest PHA production was by strain PWF. The highest percentage of PHA production was by mixed culture. On the other hand, the lowest production of biomass and PHA were by strain PWG and DL3, respectively. Biomass and PHA production increased exponentially till 96 h, however percentage of PHA (%PHA) production decreased after 48 h.

Among all carbon sources based on starch, highest production of biomass and PHA followed the trend; pure starch > rice starch > potato starch > corn starch (Table 1). On pure starch, highest production of PHA was by mixed culture, followed closely by single cell cultures of strain PWF and strain PWC, PWG, and DL3 (Fig. 2). Among carbon sources based on cellulose, biomass and PHA production followed the trend; dry wood powder > wood extract > wood shavings > crystalline cellulose (Table 2). Highest PHA production among all cellulosic carbon sources was on dry wood powder (Fig. 3). After 24 h, production trend of PHA among single cell cultures was as follows: PWF > PWC > PWA > PWG > DL3. High production of PHA was obtained using mixed culture. Successful LDPE utilization by PHA producers was corroborated using LDPE as sole carbon in nutrient-limited, selective media (Supplementary data Fig. S1). Using LDPE as sole carbon source, highest PHA production after 24 h was by strain PWF (3.27 g/L; 51%). These production rates were followed by bacterial strains PWC (0.74 g/L; 26%), PWA (0.64 g/L; 18%), and DL3 (0.32 g/L; 18%) (Table 3). Lowest PHA production (0.28 g/L, 18%) was by strain PWG. Mixed culture gave 1.95 g/L i.e., 59% PHA production (Fig. 4).

PHA production on carbon sources based on starch

Strains Time (h) Starch (pure) Rice water Potato starch Corn starch
PWA 24 10.29 3.58 35 9.34 3.18 34 6.21 2.14 34 4.77 1.36 29
48 16.96 5.24 31 13.64 4.14 30 9.67 2.90 30 8.12 2.14 26
72 19.81 5.08 26 23.40 5.15 22 18.74 4.16 22 17.34 3.59 21
96 30.31 7.77 26 35.18 7.28 21 35.01 7.45 21 31.84 6.62 21
PWC 24 8.55 4.70 55 8.98 4.70 52 6.07 3.15 52 4.69 2.35 50
48 21.10 10.90 52 20.51 10.00 49 15.92 7.67 48 14.46 6.27 43
72 33.12 9.40 28 33.83 8.40 25 28.03 7.00 25 27.38 6.49 24
96 47.15 8.90 19 46.78 7.90 17 39.80 6.88 17 37.14 5.82 16
PWF 24 11.90 8.90 75 9.40 6.85 73 8.31 5.70 69 7.09 4.64 65
48 18.70 13.62 73 13.20 8.96 68 13.20 8.21 62 11.27 6.72 60
72 23.13 7.84 34 23.45 7.21 31 25.45 7.80 31 24.89 6.40 26
96 40.04 14.90 37 40.04 14.84 37 40.31 13.61 34 39.47 12.22 31
PWG 24 3.98 1.25 31 2.89 0.90 31 2.20 0.69 31 2.28 0.64 28
48 9.17 1.97 22 7.30 1.39 19 6.62 1.27 19 6.05 1.14 19
72 25.75 4.83 19 23.09 3.94 17 22.83 3.36 15 17.65 2.28 13
96 37.48 6.88 18 33.79 5.64 17 32.14 5.56 17 26.45 4.34 16
DL3 24 6.01 1.53 25 4.93 1.23 25 4.84 1.20 25 3.82 0.97 25
48 11.87 2.41 20 9.88 1.90 19 9.00 1.42 16 8.24 1.11 14
72 25.71 4.25 17 23.06 3.46 15 19.27 2.90 15 17.58 2.50 14
96 34.17 6.88 20 30.64 5.84 19 27.18 5.00 18 27.42 4.57 17
Mixed culture 24 9.53 7.86 82 8.01 6.56 82 6.33 5.04 80 4.85 3.74 77
48 17.59 9.91 56 17.20 9.25 54 17.04 9.10 53 15.63 8.01 51
72 29.39 9.91 34 29.79 9.48 32 27.55 8.52 31 23.53 6.82 29
96 43.47 13.44 31 46.54 12.13 26 45.12 11.71 26 43.81 10.44 24

A, Biomass (g/L); B, PHA (g/L); C, %PHA (%).

PHA production on carbon sources based on cellulose

Strains Time (h) Crystalline cellulose Dry wood powder Wood extract Wood shavings
PWA 24 3.97 0.96 24 5.43 1.85 34 4.44 1.19 27 4.20 1.06 25
48 6.24 1.32 21 9.12 2.39 26 7.96 2.10 26 7.21 1.90 26
72 14.91 2.65 18 18.52 3.99 22 17.31 3.48 20 15.08 3.04 20
96 26.76 5.77 22 32.10 6.74 21 30.66 6.52 21 27.26 6.05 22
PWC 24 2.92 1.31 45 5.76 2.94 51 4.68 2.24 48 3.60 1.70 47
48 8.33 3.32 40 14.52 6.67 46 13.82 5.88 43 9.11 3.72 41
72 17.71 3.65 21 27.43 6.94 25 25.27 5.77 23 18.11 4.00 22
96 26.12 3.49 13 38.04 5.94 16 34.45 5.31 15 26.86 4.10 15
PWF 24 6.80 3.70 54 7.75 5.05 65 7.09 4.49 63 6.83 4.00 59
48 10.20 5.00 49 12.20 7.30 60 10.45 6.03 58 10.27 5.89 57
72 21.60 5.49 25 25.04 7.36 29 23.45 6.21 26 22.59 5.90 26
96 36.40 10.60 29 40.17 12.73 32 37.82 11.84 31 37.69 11.59 31
PWG 24 2.10 0.46 22 2.37 0.69 29 2.14 0.60 28 2.11 0.52 25
48 5.71 0.90 16 6.48 1.18 18 5.85 1.12 19 5.10 0.99 19
72 14.44 1.85 13 18.48 2.74 15 17.39 2.22 13 16.42 2.20 13
96 19.82 2.83 14 28.77 4.93 17 26.30 4.03 15 25.56 3.93 15
DL3 24 2.15 0.44 21 4.03 1.00 25 3.56 0.86 24 2.88 0.68 24
48 6.11 0.69 11 9.07 1.24 14 7.59 0.97 13 6.87 0.84 12
72 14.39 1.74 12 18.63 2.70 14 17.36 2.33 13 15.27 1.86 12
96 20.10 2.82 14 28.17 4.74 17 26.73 4.01 15 23.90 3.45 14
Mixed culture 24 3.57 2.18 61 5.53 4.30 78 4.20 3.05 73 3.91 2.72 70
48 13.54 6.11 45 16.60 8.52 51 14.60 7.05 48 14.17 6.73 48
72 22.42 5.57 25 26.59 7.83 29 23.01 6.24 27 22.67 5.93 26
96 38.23 8.09 21 44.64 11.00 25 42.92 9.96 23 41.93 9.76 23

A, Biomass (g/L); B, PHA (g/L); C, %PHA (%).

PHA production on LDPE

Strains Time (h) A B C
PWA 24 3.60 0.64 18
48 5.35 1.13 21
72 9.25 1.68 18
96 21.60 4.37 20
PWC 24 2.88 0.74 26
48 7.70 2.92 38
72 15.58 2.41 15
96 25.59 2.20 9
PWF 24 6.45 3.27 51
48 9.36 3.94 42
72 18.09 4.55 25
96 30.09 8.64 29
PWG 24 1.54 0.28 18
48 3.90 0.58 15
72 13.33 1.73 13
96 15.04 2.18 15
DL3 24 1.72 0.32 18
48 5.57 0.55 10
72 14.13 1.37 10
96 18.73 2.30 12
Mixedculture 24 3.33 1.95 59
48 11.49 5.17 45
72 16.93 4.13 24
96 31.27 5.98 19

A, Biomass (g/L); B, PHA (g/L); C, %PHA (%).

Fig. 2. Graphical representation for percentage PHA production on pure starch.
Fig. 3. Graphical representation for percentage PHA production on dry wood powder.
Fig. 4. Graphical representation for percentage PHA production on LDPE.

PCR amplification of phaC and phaC1 gene

PCR amplification for phaC1 gene of strain PWC (Pseudomonas aeruginosa) using 179-L and 179-R primers was carried out and PCR amplicon of 540 bp length was obtained. PCR was done to amplify phaC gene of strain PWC (Pseudomonas aeruginosa) using ORF1 and 179-R primers. PCR amplicon of 1,700 bp length was obtained. However, when using ORF1 and 179-R primers, aside from specific band at 1,700 bp, some non-specific bands were also observed. Using phal-1-Forward and phal-2-Reverse primers to amplify gene phaC, PCR amplicons of 800 bp and 420 bp were obtained for Pseudomonas and Bacillus, respectively. Gene sequence obtained for phaC gene of strain PWC amplified using phal-2-Reverse primer was submitted to GenBank under accession number MT340498. Blast results showed similarity to gene phaC. Analyzing sequence against NCBI conserved domain database for determination of conserved domains showed one hit for PHA_synth_II super family with accession number CL31144 within sequence interval 1 to 453 with 4.85e-86 error value. Evolutionary history of gene phaC of strain PWC with closely related taxa was inferred using MEGA 10.1.7 by neighbor joining method (Fig. 5). Optimal tree is shown with sum of branch length = 1.16144686. The tree was drawn to scale using Maximum Composite Likelihood method to compute evolutionary distances.

Fig. 5. Dendrogram of closely related taxa with gene phaC of strain PWC. The evolutionary distances between strain DL3 and closely related taxa are shown by drawing a dendrogram to scale using Maximum Composite Likelihood method. The sum of branch length is 1.16144686.

Current study was aimed at two objectives. First, the isolation, identification and revival of bacterial strains – from tannery and the revival of produced water – capable of PHA production and plastic degradation (Sohail and Jamil, 2020; Sohail et al., 2020). Secondly, the use of low cost, easily replenishable, non-fossil fuel-based carbon source and mixed cultures for optimization of PHA production over a period of 96 hours. PHA production capability of bacterial strains was corroborated by amplification of PHA synthase gene phaC.

Bacterial strain DL3 demonstrating PHA production ability was isolated from tannery. Tannery effluent samples collected from leather tanning industries were found to be dark in color with obnoxious odor. In a similar study, David et al. (2015) reported high concentrations of biodegradable organic acids such as hemicellulose, carbohydrate, dextrin, lignin, etc. with an obnoxious odor. This composition makes tannery samples a suitable environment for PHA producers. Strain DL3 gave florescence on Nile blue and/or Nile red dyes supplemented media, due to presence of PHA granules and was able to produce PHA successfully, indicating that the recalcitrant and antioxidant polymers present in tannery effluents were utilized by bacterial strain DL3 for PHA production at its source of isolation. Studies conducted by Merugu et al. (2012) and David et al. (2015) – reporting 26 mg/L PHB production by Rhodobacter capsulatus and isolation of Pseudomonas aeruginosa from tannery effluents, respectively – corroborate the hypothesis that strain DL3 could have produced PHA as a carbon assimilation product during nutrient limited stress provided by tannery samples.

Results for optimization of PHA production indicated that maximum production of PHA occurred at 24 h (during exponential growth phase). During this phase, high productions of PHA were recorded using bacterial strain PWF (single cell culture) and mixed cell culture.

Using carbon sources based on starch, high productions of PHA were given by strain PWF (Bacillus tequilensis) and strain PWC (Pseudomonas aeruginosa) respectively (Fig. 2). Halami also reported high PHA production by cell dry weight i.e. 48% using starch by Bacillus cereus (Halami, 2008). Whereas, Aljuraifani et al. reported 15 g/L (w/v) or 90.9% PHA production by Pseudomonas sp. Strain P(16) on rice bran (Aljuraifani et al., 2019). Using mixed culture also optimized PHA production and high yields can, therefore, be obtained industrially through fed-batch fermentation using these bacterial strains on starches (Koller and Braunegg, 2015). Huang et al. (2006) reported 140.0 g/L cell biomass and 77.8 g/L (55.6%) PHA production by Haloferax mediterranei using extruded rice starch as main carbon source in fed-batch fermentation.

On cellulosic carbon sources, highest PHA yields obtained were on dry wood powder (Fig. 3) after 24 h. It was hypothesized that decline in growth and PHA production after 48 h was due to utilization of PHA inclusions present inside vegetative cells for growth rather than the use of extracellular cellulosic carbon source. It is also possible that organic inhibitors such as furfural, acetic acid, acid soluble lignin, and formic acid, etc. in wood extract had an inhibitory effect on growth which was overcome by tolerant strains – as is the case with bacterial strains PWC (Pseudomonas aeruginosa) and PWF (Bacillus tequilensis). Yu and Stahl (2008) reported similar findings while studying utilization of bagasse hydrolysates – rich in cellulosic biomass – by Ralstonia eutropha for PHA production. They observed an inhibitory effect of hydrolysates on microbial activity and PHA production which could be overcome by use of large inoculum, tolerant strain, or diluted hydrolysate. They reported simultaneous organic inhibitors utilization and PHA production (57%) by Ralstonia eutropha, during potential recycling of process water (Yu and Stahl, 2008).

Bacterial growth and PHA production occurred comparatively slower on crystalline cellulose than on unprocessed cellulosics. It is hypothesized that in contrast to unprocessed cellulosics – containing many simple monomeric constituents like glucose – degradation of crystalline cellulose to levulinic acid occurred during bacterial metabolism. Unprocessed cellulosics, in contrast, had much lower cellulose content and were utilized more effectively. However, levulinic acid is produced in low levels during growth on simpler carbon sources i.e., glucose and is available as carbon source. Keenan et al. (2004) reported 9.5 g/L biomass and 4.2 g/L P (3HB-co-3HV) yield by Burkholderia cepacia using 0.52% levulinic acid. Jang and Rogers (1996) also detected low levels of levulinic acid when using glucose as sole carbon source for PHA production by Alcaligenes sp. and during experimentation to study effects of levulinic acid reported 38.3% PHA production with 23.5% polyhydroxyvalerate (PHV) content on 0.5 g/L/h addition of levulinic acid.

Low density polyethylene (LDPE) was used as sole carbon source in nutrient-limited, selective media to produce high value biopolymer PHA from low value recyclable waste materials, as polyethylene (PE) is the most used plastic. Since, PE is composed of highly stable covalent hydrocarbon bonding, it is likely that LDPE was utilized in the fatty acid PHA biosynthesis pathway. Thus, bioconversion of LDPE to PHA happened via β-oxidation. In a related study by Johnston et al. (2017) 32% PHA production and 1.42 g/L cell biomass using N-PEW (non-oxidized polyethylene wax) by Cupriavidus necator was reported. While Guzik et al. (2014) in another study, 25% production of lcl PHA by Pseudomonas aeruginosa using pyrolyzed PE wax was reported. Overall LDPE bioconversion to PHA and growth rate on LDPE increased up to 48 h (Fig. 4). After 48 h, a decline in growth and PHA production was observed indicating that the presence of chemically stable C-C and C-H bonding in high molecular weight, hydrophobic LDPE renders its complete bacterial degradation difficult. Similar findings were reported by Leja and Lewandowicz (2010) and Shah et al. (2008). The overall increase at 48 h and the subsequent gradual decrease after 48 h could also be due to substrate limitation i.e., digestion of shorter alkane chains in LDPE occurred up to 48 h while after 48 h, the concentration of longer alkane chains in media becomes high. These longer alkane chains, however available, are inaccessible to bacteria for metabolism resulting in gradual decline in growth and PHA production. Radecka et al. (2016) reported similar finding while studying use of oxidized polyethylene wax (O-PEW) as carbon source for PHA production by Ralstonia eutropha. It is also possible that after 48 h, degradation products of LDPE showed low antimicrobial activity against microbial growth resulting in its decline. In a study by Zhang et al. (2008) low antimicrobial activity of PE was reported. Seyfriedsberger et al. (2006) also reported low antimicrobial activity of linear low-density polyethylene (LLDPE) against Staphylococcus aureus. However, they did not report antimicrobial activity against Escherichia coli (Seyfriedsberger et al., 2006). Gregorova et al. (2011) on the other hand, reported no antimicrobial activity against either E. coli or S. aureus. Altogether, utilization of LDPE and PHA production by bacteria using said plastic waste was observed. Over 29% of plastics manufactured worldwide are made up of PE (Johnston et al., 2017). This study – focusing on utilizing the waste of non-degradable polymer; plastic for production of biodegradable biopolymer; PHA, offers a green alternative to plastic waste accumulation. PHA production through use of LDPE, therefore, also presents a possible solution to waste treatment costs (Radecka et al., 2016).


Current study demonstrated the successful utilization of easily replenishable, low cost, non-fossil fuel-based carbon sources by produced water and tannery bacteria – especially Bacillus tequilensis (PWF) and Pseudomonas aeruginosa (PWC). These bacterial strains can produce PHA using such carbon sources as an alternative to non-fossil fuel-based, non-replenishable carbon sources. This study also signified the ability of PHA producers to degrade plastics mainly polyolefin resins like LDPE and utilize their degradation products to produce biopolymers of high value i.e., PHA.

Supplementary Materials



§Supplemental material for this article may be found at

Conflict of Interest

The authors have no conflict of interest to report.

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