Biosynthesis of Polyhydroxyalkanoates (PHAs) by the Valorization of Biomass and Synthetic Waste

Synthetic pollutants are a looming threat to the entire ecosystem, including wildlife, the environment, and human health. Polyhydroxyalkanoates (PHAs) are natural biodegradable microbial polymers with a promising potential to replace synthetic plastics. This research is focused on devising a sustainable approach to produce PHAs by a new microbial strain using untreated synthetic plastics and lignocellulosic biomass. For experiments, 47 soil samples and 18 effluent samples were collected from various areas of Punjab, Pakistan. The samples were primarily screened for PHA detection on agar medium containing Nile blue A stain. The PHA positive bacterial isolates showed prominent orange–yellow fluorescence on irradiation with UV light. They were further screened for PHA estimation by submerged fermentation in the culture broth. Bacterial isolate 16a produced maximum PHA and was identified by 16S rRNA sequencing. It was identified as Stenotrophomonas maltophilia HA-16 (MN240936), reported first time for PHA production. Basic fermentation parameters, such as incubation time, temperature, and pH were optimized for PHA production. Wood chips, cardboard cutouts, plastic bottle cutouts, shredded polystyrene cups, and plastic bags were optimized as alternative sustainable carbon sources for the production of PHAs. A vital finding of this study was the yield obtained by using plastic bags, i.e., 68.24 ± 0.27%. The effective use of plastic and lignocellulosic waste in the cultivation medium for the microbial production of PHA by a novel bacterial strain is discussed in the current study.

Considering the current research on sustainable carbon sources to produce bioplastics, the bioplastic industry will potentially boom in the near future. The global bioplastic production capacity is set to increase from around 2.11 million tons in 2018 to approximately 2.62 million tons in 2023, with PHAs being the main drivers of the market [22]. This research focused on the isolation of a new PHA producing bacterial strain from soil and effluent samples to be used in developing a cost-effective method for the synthesis of PHAs by synthetic waste and lignocellulosic biomass as substrates.

Primary Screening
A total of 65 samples, including 47 soil and 18 effluent samples were collected from various areas of Punjab, Pakistan. These samples were subjected to primary screening for the isolation of PHA producing bacteria. A total of 127 bacterial isolates were screened from these samples. Moreover, 67 isolates revealed signs of PHA accumulation by showing orange-yellow fluorescence under UV light as mentioned in Table 1. Consequently, these 67 isolates were further selected for the estimation of PHA production. Figure 1A shows an example of a PHA positive sample (sample no: 8).  Figure 1B. It was concluded that this strain held the highest potential to produce maximum PHA content as compared to other strains.
Since the isolate 16a produced maximum PHA, it could be quite possible that this strain could quickly and efficiently utilize carbon source. The source of isolate 16a was plastic industry soil. Mohammed et al. [30], Kosseva and Rusbandi [31], and Sangakharak and Prasertsan [32] reported the isolation of bacteria from similar plastic sources, such as plastic pieces, plastic chairs, and plastic waste landfill sites, which provided high PHA yields up to 0.5 g/100 mL. High PHA production rate is associated with the bacterial ability to utilize plastic as a substrate as the soils enriched with plastic pieces are their indigenous habitats. These strains already have modified metabolism rates to sustain in oligotrophic conditions so they adapt to utilize the carbon available in plastic [33]. This shows an enhanced ability of indigenous bacteria to survive in nutrient-deficient conditions than nonindigenous bacteria [34]. In this research, the outcome of considerable differences in the PHA yields of bacteria isolated from plastic habitats can be due to the differences in the surrounding habitats of the plastic sources. Puglisi et al. [35] studied and proved the hypothesis that different polyethylene (PE) plastic waste samples harbor different bacterial communities. The structure and physiological capabilities of these communities are dependent on the physico-chemical properties of the plastic waste and the environment in which they dwell.  PHAs are insoluble storage granules that microorganisms accumulate in stressful environmental conditions, under excess of carbon and deficiency of other essential nutrients [23]. Thus, industrial soil samples and effluents used for the isolation of PHA producing bacteria were appropriate for the purpose. Most of the samples were collected from the industries of paint, paper, and plastic.
Any substance that becomes useless and defective after its primary use is considered as "waste" [24]. Liquid waste from industrial sites, agricultural processes or domestic sewage is "effluent waste". Effluents can be harmful to the environment if released untreated because of their polluting chemical nature [25], as they contain partially degraded organic matter with minimum nutrients. They serve as an ideal source of isolation for microorganisms adapted to survive in oligotrophic conditions [26]. Products difficult to degrade such as plastic, paint residues, cardboard, and paper residues may also be a source of isolation of novel microorganisms that degrade these difficult products with simultaneous PHA production. Whereas soil samples collected from agricultural areas, such as croplands, compost, and landfill sites are rich in carbon sources [27]. The choice of collecting the samples from these habitats has made possible the isolation of a large number of PHA positive isolates.
For a more visual detection of PHA producing bacteria, staining and fluorescence come into play. This is achieved through the use of Nile blue A stain in the agar medium. When the cells of some microbial colonies grow in a medium containing this stain, the stain is absorbed within the cytoplasm of their cells [28]. This dye subsequently enters into the PHA inclusions. It is a basic oxazine dye [29] containing Nile blue sulfate and Basic Blue 12. These compounds when excited by UV light of 312 nm, reflect orange color hence the orange fluorescence is generated [28]. Apart from being a highly sensitive method to detect PHAs, it also indicates the difference in the amount of accumulated PHAs [29]. The greater the intensity of fluorescence, the more the accumulation of PHA granules [30]. In this way, the isolate 16a showed the brightest and the most intense fluorescence as seen in Figure 1B. It was concluded that this strain held the highest potential to produce maximum PHA content as compared to other strains.

Secondary Screening of PHA Producers
Primarily screened bacterial colonies were further purified by quadrant streaking and analyzed for PHA production by submerged fermentation at 37 • C, 150 rpm for 72 h. After incubation, maximum PHA production was shown by the isolate 16a i.e., 69.72 ± 0.17%. However, isolates 3a, 4a, 5a, 8a, 11a, 12a, 14a, 16b, 18b, 23a, 33a, 39a, 45a, 47a, 48a, 53a, and 65a showed significant PHA production Molecules 2020, 25, 5539 7 of 23 ranging from 36% to 55%, approximately. While the remaining strains did not show any significant PHA production, as evident from Table 2. Bacterial isolate 16a was selected for further studies. Since the isolate 16a produced maximum PHA, it could be quite possible that this strain could quickly and efficiently utilize carbon source. The source of isolate 16a was plastic industry soil. Mohammed et al. [30], Kosseva and Rusbandi [31], and Sangakharak and Prasertsan [32] reported the isolation of bacteria from similar plastic sources, such as plastic pieces, plastic chairs, and plastic waste landfill sites, which provided high PHA yields up to 0.5 g/100 mL. High PHA production rate is associated with the bacterial ability to utilize plastic as a substrate as the soils enriched with plastic pieces are their indigenous habitats. These strains already have modified metabolism rates to sustain in oligotrophic conditions so they adapt to utilize the carbon available in plastic [33]. This shows an enhanced ability of indigenous bacteria to survive in nutrient-deficient conditions than non-indigenous bacteria [34]. In this research, the outcome of considerable differences in the PHA yields of bacteria isolated from plastic habitats can be due to the differences in the surrounding habitats of the plastic sources. Puglisi et al. [35] studied and proved the hypothesis that different polyethylene (PE) plastic waste samples harbor different bacterial communities. The structure and physiological capabilities of these communities are dependent on the physico-chemical properties of the plastic waste and the environment in which they dwell.

Molecular Identification
Sequencing of isolate 16a was carried out by Macrogen sequencing company, Seoul, Korea. Primers used for PCR and 16S rRNA sequencing are given in (Table 3). Table 3. Primers used for PCR as well as 16S rRNA sequencing.

PCR Primers
Sequencing Primers The sequencing results were put into Basic Local Alignment Search Tool (BLAST) for homology analysis and first ten homologues were selected ( Figure 2) for phylogenetic tree construction using the Jalview application ( Figure 3) [36,37]. The results of the BLAST revealed that the gene sequence of isolate 16a is having identity with Stenotrophomonas maltophilia strain IAM 12423. Therefore, it was determined that the strain used in this research is S. maltophilia. GenBank sequence of the identified strain was submitted under the name of Stenotrophomonas maltophilia HA-16 with the accession number MN240936. The genus Stenotrophomonas is phylogenetically placed in the Gammaproteobacteria, was first described with the type species Stenotrophomonas maltophilia [38]. The Stenotrophomonas genus is a gram-negative genus with at least ten species [39]. It belongs to the family Xanthomonadaceae [40]. strain was submitted under the name of Stenotrophomonas maltophilia HA-16 with the accession number MN240936. The genus Stenotrophomonas is phylogenetically placed in the Gammaproteobacteria, was first described with the type species Stenotrophomonas maltophilia [38]. The Stenotrophomonas genus is a gram-negative genus with at least ten species [39]. It belongs to the family Xanthomonadaceae [40]. The first ten homologues were selected for phylogenetic tree construction.
Horiike [41] dictates the importance of phylogenetic trees in molecular identification. He emphasizes that phylogenetic trees are much helpful in predicting the evolutionary basis of relationships between various species. These trees can help us predict the effects of evolutionary patterns in different habitats and their effects on a strain's metabolic products. They also tell us about the evolution of metabolic products from ancestors to their descendants and how they are improved or differentiated over time [42].

Characterization of the Extracted Polymer
The PHA extracted from S. maltophilia HA-16 was characterized by FTIR spectroscopy. Functional group analysis was done through signal peaks recorded in the range of 4000-400 cm −1 . The peak was plotted using OriginLab 8.5 as shown in Figure 4 [43].
Abid et al. [44] compared his extracted polymer with FTIR peaks of standard Polyhydroxybutyrate (PHB) purchased from Sigma Aldrich © (St. Louis, MO, USA). We compared our polymer with the FTIR peaks of the standard PHB sample used by Abid et al. [44]. The Fingerprinting region is usually the region of the spectrum in the range of 670-400 cm −1 . The polymer under study expressed a considerable peak at 566.9 cm −1 , which showed that there were many stretches of carbonyl groups (C=O) present. Comparatively, the fingerprinting region of the standard PHB had  The first ten homologues were selected for phylogenetic tree construction.
Horiike [41] dictates the importance of phylogenetic trees in molecular identification. He emphasizes that phylogenetic trees are much helpful in predicting the evolutionary basis of relationships between various species. These trees can help us predict the effects of evolutionary patterns in different habitats and their effects on a strain's metabolic products. They also tell us about the evolution of metabolic products from ancestors to their descendants and how they are improved or differentiated over time [42].

Characterization of the Extracted Polymer
The PHA extracted from S. maltophilia HA-16 was characterized by FTIR spectroscopy. Functional group analysis was done through signal peaks recorded in the range of 4000-400 cm −1 . The peak was plotted using OriginLab 8.5 as shown in Figure 4 [43].
Abid et al. [44] compared his extracted polymer with FTIR peaks of standard Polyhydroxybutyrate (PHB) purchased from Sigma Aldrich © (St. Louis, MO, USA). We compared our polymer with the FTIR peaks of the standard PHB sample used by Abid et al. [44]. The Fingerprinting region is usually the region of the spectrum in the range of 670-400 cm −1 . The polymer under study expressed a considerable peak at 566.9 cm −1 , which showed that there were many stretches of carbonyl groups (C=O) present. Comparatively, the fingerprinting region of the standard PHB had Horiike [41] dictates the importance of phylogenetic trees in molecular identification. He emphasizes that phylogenetic trees are much helpful in predicting the evolutionary basis of relationships between various species. These trees can help us predict the effects of evolutionary patterns in different habitats and their effects on a strain's metabolic products. They also tell us about the evolution of metabolic products from ancestors to their descendants and how they are improved or differentiated over time [42].

Characterization of the Extracted Polymer
The PHA extracted from S. maltophilia HA-16 was characterized by FTIR spectroscopy. Functional group analysis was done through signal peaks recorded in the range of 4000-400 cm −1 . The peak was plotted using OriginLab 8.5 as shown in Figure 4 [43].
Abid et al. [44] compared his extracted polymer with FTIR peaks of standard Polyhydroxybutyrate (PHB) purchased from Sigma Aldrich© (St. Louis, MO, USA). We compared our polymer with the FTIR peaks of the standard PHB sample used by Abid et al. [44]. The Fingerprinting region is usually the region of the spectrum in the range of 670-400 cm −1 . The polymer under study expressed a considerable peak at 566.9 cm −1 , which showed that there were many stretches of carbonyl groups (C=O) present. Comparatively, the fingerprinting region of the standard PHB had multiple sharp peaks, the sharpest one being at 514.03 cm −1 . Absorption in the range of 1200-800 cm −1 indicated the presence of multiple C-C stretches with medium length bands which corresponded to an alkane group [45]. The extracted polymer expressed a sharp peak at 1017 cm −1 , whereas the standard showed a sharp peak at 1044.67 cm −1 . The sharpness of the peaks in both the polymers was almost the same which signified the same length of C-C stretches.
Pérez-Arauz et al. [46] mentioned that absorption peaks in the range of 3000-2850 cm −1 show a strong C-H bond stretch. The extracted polymer displayed a small peak at 2923 cm −1 in comparison to the standard which expressed a same sized peak at 2933.36 cm −1 . Since the C-O peaks are not very prominent in the main region, this difference could be due to the disturbances in the polymer structure occurred during extraction [47]. The comparison of the size and the location of peaks in between the standard PHB and our extracted polymer showed that our polymer might not be as pure. The presence of similar length patterns in the peaks at some places can indicate that the biosynthesized polymer is a medium chain length (mcl) PHA copolyester [48].

Optimization of Cultural Conditions
Incubation time, incubation temperature, pH of the fermentation medium, and carbon sources were optimized for maximum PHA production through duplicate fermentation experiments. The average percentage amount of PHA produced with intracellular cell dry weight (CDW g/100 mL), per experiment per parameter is expressed in Table 4.  Pérez-Arauz et al. [46] mentioned that absorption peaks in the range of 3000-2850 cm −1 show a strong C-H bond stretch. The extracted polymer displayed a small peak at 2923 cm −1 in comparison to the standard which expressed a same sized peak at 2933.36 cm −1 . Since the C-O peaks are not very prominent in the main region, this difference could be due to the disturbances in the polymer structure occurred during extraction [47]. The comparison of the size and the location of peaks in between the standard PHB and our extracted polymer showed that our polymer might not be as pure. The presence of similar length patterns in the peaks at some places can indicate that the biosynthesized polymer is a medium chain length (mcl) PHA copolyester [48].

Optimization of Cultural Conditions
Incubation time, incubation temperature, pH of the fermentation medium, and carbon sources were optimized for maximum PHA production through duplicate fermentation experiments. The average percentage amount of PHA produced with intracellular cell dry weight (CDW g/100 mL), per experiment per parameter is expressed in Table 5.   Figure 5). PHA percentage seems to be directly related to the amount of intracellular CDW (g/100 mL). Nonetheless, at 72 h incubation, 0.52 g/100 mL of intracellular CDW was extracted which was recorded the highest in this pool of experiments.  Figure 5). PHA percentage seems to be directly related to the amount of intracellular CDW (g/100 mL). Nonetheless, at 72 h incubation, 0.52 g/100 mL of intracellular CDW was extracted which was recorded the highest in this pool of experiments. Contrarily, Munir et al. [49] expressed a different trend for PHA production by Stenotrophomonas genus with highest PHA yield achieved after 48 h. They used 2% glucose as the sole carbon source compared to our study where only 0.45% of glucose was used. They recorded increasing growth until 72 h, but PHA production increased until 48 h, only and after that, it started declining. Additionally, this difference can be backed by the work of Alqahtani [50], where she describes 48 h as the optima for the highest metabolic activity for PHA production. Shaaban and Mowafy [51] described that the maximum PHA production by S. maltophilia occurred at 96 h and stayed stable until 144 h. They utilized 1% glucose in the medium. These differences in the optimum incubation time to produce maximum PHA might be attributed to differences in the nutrients and the carbon source (which in Contrarily, Munir et al. [49] expressed a different trend for PHA production by Stenotrophomonas genus with highest PHA yield achieved after 48 h. They used 2% glucose as the sole carbon source compared to our study where only 0.45% of glucose was used. They recorded increasing growth until 72 h, but PHA production increased until 48 h, only and after that, it started declining. Additionally, this difference can be backed by the work of Alqahtani [50], where she describes 48 h as the optima for the highest metabolic activity for PHA production. Shaaban and Mowafy [51] described that the maximum PHA production by S. maltophilia occurred at 96 h and stayed stable until 144 h. They utilized 1% glucose in the medium. These differences in the optimum incubation time to produce maximum PHA might be attributed to differences in the nutrients and the carbon source (which in our case was glucose).

Effect of Temperature of Incubation
PHA production was optimized at 25, 37, 30, and 40 • C with glucose as a carbon source to find out the temperature optima for S. maltophilia HA-16.37 • C was the optimum temperature with the highest PHA production i.e., 78.85 ± 0.23% as compared to other temperatures (Figure 6). At the same temperature, intracellular CDW was calculated to be the highest i.e., 0.59 g/100 mL. PHA content was low at 25 • C with a slight increase at 30 • C and a sharp decline after 37 • C with the lowest PHA production at 40 • C. PHA production at 25 and 30 • C can be considered as satisfactory with 41.81 ± 0.39 and 47.32 ± 0.12%, respectively. However, PHA yield poorly declined at temperatures above 37 • C, producing only 30.41 ± 0.47% PHA at 40 • C. A strange observation in this experiment was recorded in the intracellular CDW at this temperature. Usually, CDW is observed to be directly related to the amount of PHA produced. Yet, CDW at 40 • C was recorded to be 0.2 g/100 mL, which was still higher than the CDW recorded at 25 • C and 30 • C. Hence, it cannot be taken as a thumb rule that the higher the CDW, the higher the PHA content, because the CDW can also include the dry weight of things other than PHA. PHA production was optimized at 25, 37, 30, and 40 °C with glucose as a carbon source to find out the temperature optima for S. maltophilia HA-16.37 °C was the optimum temperature with the highest PHA production i.e., 78.85 ± 0.23% as compared to other temperatures (Figure 6). At the same temperature, intracellular CDW was calculated to be the highest i.e., 0.59 g/100 mL. PHA content was low at 25 °C with a slight increase at 30 °C and a sharp decline after 37 °C with the lowest PHA production at 40 °C. PHA production at 25 and 30 °C can be considered as satisfactory with 41.81 ± 0.39 and 47.32 ± 0.12%, respectively. However, PHA yield poorly declined at temperatures above 37 °C, producing only 30.41 ± 0.47% PHA at 40 °C. A strange observation in this experiment was recorded in the intracellular CDW at this temperature. Usually, CDW is observed to be directly related to the amount of PHA produced. Yet, CDW at 40 °C was recorded to be 0.2 g/100 mL, which was still higher than the CDW recorded at 25 °C and 30 °C. Hence, it cannot be taken as a thumb rule that the higher the CDW, the higher the PHA content, because the CDW can also include the dry weight of things other than PHA. Temperature optimization is important concerning the microorganism used for PHA production [52]. Scientists at the American Tissue Culture Center (ATCC) also confirmed 37 °C as the temperature optima for S. maltophilia. On the other hand, Singh and Parmar [13] reported PHA production with the same bacteria but different strains (S. maltophilia AK21 and S. maltophilia 13635L) at considerably low temperatures of 25 and 30 °C. This deviation might be due to the isolation of bacterial strains from different habitats. Alqahtani [50], in her work, demonstrated that extremely warm (55 °C) and extremely cold (4 °C) temperatures affect the growth of S. maltophilia. Her study further solidifies the results of this research by similar results, displaying the optimum growth at 37 °C. Guerrero and others [53] reported that PHA producing enzymes do not work efficiently above 37 °C and resulted in low yield.

Effect of pH of the Fermentation Medium
The fermentation medium was prepared at five different pH levels (6.0, 6.5, 7.0, 7.5, and 8.0) to find the pH optima for PHA production. A significant PHA yield of 78.85 ± 0.11% was recorded at 7.0 pH as compared to other pH levels. At acidic pH i.e., 6.0, PHA yield was poor i.e., only 20.14 ± 0.26% which increased to 41.70 ± 0.29% at slightly less acidic pH of 6.5. At slightly basic pH i.e., 7.5, Temperature optimization is important concerning the microorganism used for PHA production [52]. Scientists at the American Tissue Culture Center (ATCC) also confirmed 37 • C as the temperature optima for S. maltophilia. On the other hand, Singh and Parmar [13] reported PHA production with the same bacteria but different strains (S. maltophilia AK21 and S. maltophilia 13635L) at considerably low temperatures of 25 and 30 • C. This deviation might be due to the isolation of bacterial strains from different habitats. Alqahtani [50], in her work, demonstrated that extremely warm (55 • C) and extremely cold (4 • C) temperatures affect the growth of S. maltophilia. Her study further solidifies the results of this research by similar results, displaying the optimum growth at 37 • C. Guerrero and others [53] reported that PHA producing enzymes do not work efficiently above 37 • C and resulted in low yield.

Effect of pH of the Fermentation Medium
The fermentation medium was prepared at five different pH levels (6.0, 6.5, 7.0, 7.5, and 8.0) to find the pH optima for PHA production. A significant PHA yield of 78.85 ± 0.11% was recorded at 7.0 pH as compared to other pH levels. At acidic pH i.e., 6.0, PHA yield was poor i.e., only 20.14 ± 0.26% which increased to 41.70 ± 0.29% at slightly less acidic pH of 6.5. At slightly basic pH i.e., 7.5, 32.95 ± 0.33% of PHA production was observed. This yield is almost equal to the PHA yield recorded at 8.0 pH (Figure 7).
Molecules 2020, 25, x FOR PEER REVIEW 12 of 22 32.95 ± 0.33% of PHA production was observed. This yield is almost equal to the PHA yield recorded at 8.0 pH (Figure 7). Shaaban with colleagues [54] found pH 7.0 as the optimum for PHB production by S. maltophilia. They further elaborated that at pH 6.0 and 8.0, PHB production was not significant which also related with the current study. Raj and his team [55] also reported pH 7.0 as the optimum pH for PHA production by S. maltophilia. Lathwal et al. [56] demonstrated the same results for PHA production at different pH of fermentation media. They were able to verify that PHA production was maximum at pH 7.0. Their work also validated the current results that on pH 6.0, PHA production was low, however, at pH 8.0, PHA production was not low and still significant.

Optimization of Carbon Source for PHA Production
Some unconventional carbon sources such as undegraded wood chips, cardboard cutouts, shredded plastic bottles, wasted polystyrene cups, and waste plastic bags were used for increased PHA production. These carbon sources were used without any pre-treatments. Surprisingly, plastic bags proved the most optimum among these carbon sources and produced PHA content of 68.24 ± 0.27%. Whereas the other two plastic carbon sources (shredded polystyrene cups and plastic bottle cutouts) did not prove to be much efficient in PHA production. They produced the lowest PHA content of 43.75 ± 0.30% and 38.19 ± 0.22%, respectively. PHA content extracted from the wood chips and cardboard was almost equal i.e., 53.15 ± 0.17% and 51.76 ± 0.48%, respectively (Figure 8).
The cost of carbon sources is one of the prime difficulties in PHA commercialization as these sources contribute to more than 50% of the total industrial production costs. Glucose, among the optimized carbon sources, proved to be the best carbon source for PHA production, as also confirmed by Singh and Parmar [13]. However, the use of industrially produced glucose adds to the costs considerably. Replacing glucose in the fermentation medium with waste sources in the current research was an attempt to address this concern. When the same approach will be industrially adapted, the costs can be further cut down by integrating the production lines with waste streams of paper industries, plastic industries and packaging industries. However, the collection of plastic at the end of PHA production cycle does not ensure its complete breakdown. There are still concerns that need more attention. Since the use of plastics to produce PHAs is a new approach, extensive research to study all related industrial parameters are needed to make it a reality. Jimenez's team [57] found S. maltophilia associated with the gut of Bark Beetle Dendroctonus rhizophagus (Curculionidae: Scolytinae). In its gut, it plays a role in the degradation, hydrolysis, fermentation, and oxidation of lignin and cellulose derived aromatic products. Their findings can elaborate the current results of Shaaban with colleagues [54] found pH 7.0 as the optimum for PHB production by S. maltophilia. They further elaborated that at pH 6.0 and 8.0, PHB production was not significant which also related with the current study. Raj and his team [55] also reported pH 7.0 as the optimum pH for PHA production by S. maltophilia. Lathwal et al. [56] demonstrated the same results for PHA production at different pH of fermentation media. They were able to verify that PHA production was maximum at pH 7.0. Their work also validated the current results that on pH 6.0, PHA production was low, however, at pH 8.0, PHA production was not low and still significant.

Optimization of Carbon Source for PHA Production
Some unconventional carbon sources such as undegraded wood chips, cardboard cutouts, shredded plastic bottles, wasted polystyrene cups, and waste plastic bags were used for increased PHA production. These carbon sources were used without any pre-treatments. Surprisingly, plastic bags proved the most optimum among these carbon sources and produced PHA content of 68.24 ± 0.27%. Whereas the other two plastic carbon sources (shredded polystyrene cups and plastic bottle cutouts) did not prove to be much efficient in PHA production. They produced the lowest PHA content of 43.75 ± 0.30% and 38.19 ± 0.22%, respectively. PHA content extracted from the wood chips and cardboard was almost equal i.e., 53.15 ± 0.17% and 51.76 ± 0.48%, respectively (Figure 8).
The cost of carbon sources is one of the prime difficulties in PHA commercialization as these sources contribute to more than 50% of the total industrial production costs. Glucose, among the optimized carbon sources, proved to be the best carbon source for PHA production, as also confirmed by Singh and Parmar [13]. However, the use of industrially produced glucose adds to the costs considerably. Replacing glucose in the fermentation medium with waste sources in the current research was an attempt to address this concern. When the same approach will be industrially adapted, the costs can be further cut down by integrating the production lines with waste streams of paper industries, plastic industries and packaging industries. However, the collection of plastic at the end of PHA production cycle does not ensure its complete breakdown. There are still concerns that need more attention. Since the use of plastics to produce PHAs is a new approach, extensive research to study all related industrial parameters are needed to make it a reality. Jimenez's team [57] found S. maltophilia associated with the gut of Bark Beetle Dendroctonus rhizophagus (Curculionidae: Scolytinae). In its gut, it plays a role in the degradation, hydrolysis, fermentation, and oxidation of lignin and cellulose derived aromatic products. Their findings can elaborate the current results of PHA production by S. maltophilia HA-16 (MN240936) through wood chips and cardboard cutouts. In another study by Kirtania et al. [58], it was found that S. maltophilia has a significant ability to naturally degrade cellulose and hemicellulosic materials. Furthermore, Ali Wala'a et al. [59] reported numerous pretreated cellulosic and lignocellulosic sources giving maximum PHA production yield up to 90% which does not align with this research. In the current study, cellulosic and lignocellulosic materials gave lower yields as compared to one synthetic source i.e., plastic bag. PHA production by S. maltophilia HA-16 (MN240936) through wood chips and cardboard cutouts. In another study by Kirtania et al. [58], it was found that S. maltophilia has a significant ability to naturally degrade cellulose and hemicellulosic materials. Furthermore, Ali Wala'a et al. [59] reported numerous pretreated cellulosic and lignocellulosic sources giving maximum PHA production yield up to 90% which does not align with this research. In the current study, cellulosic and lignocellulosic materials gave lower yields as compared to one synthetic source i.e., plastic bag. Plastics are of various types. One of the most abundant types of plastics is polyethylene terephthalate (PET). PET plastics are made of repeating units of polymer ethylene terephthalate [60]. Plastic bags are another very common plastic products which are made up of low density poly ethylene (LDPE) and/or high density poly ethylene (HDPE) [61]. The reason why PET and most plastics do not easily biodegrade is because the entire plastic structure has very strong C-C bonds that require too much energy to breakdown and plastics do not dissolve in water [62]. However, with the increased plastic accumulation in our environment, microbial life forms have evolved to degrade plastic products to some extent. There are many bacteria that have the ability to degrade PET, Polyethylene (PE), and Polystyrene (PS) but their enzymes only have been able to give moderate turnouts. The enzymes involved in PET degradation are known as PET hydrolases [20].
While working with synthetic plastics, Kenny and his research team [63] used pre-degraded PET as a carbon source and received a yield of only up to 21% with Pseudomonas frederiksbergensis GO23. Whereas, S. maltophilia HA-16 (MN240936) produced up to 38% of PHA with undegraded PET bottles. In another study, Dai and Reusch [64] utilized synthetic plastics by using the pyrolysis oil of polystyrene and reported PHA production up to 48% in tryptic soy broth (TSB) medium with Cupriavidus necator H16. Contrarily, S. maltophilia HA-16 gave a yield of 43% with undegraded or pretreated polystyrene fragments, which is still impressive. These yields indicate an evident ability of S. maltophilia HA-16 to degrade PET bottles and polystyrene. Current research is the first case of PHA production reported from the strain S. maltophilia HA-16 (MN240936).
A surprising finding of this study is the high yield of PHA, i.e., 68.24% in plastic bags. S. maltophilia is a gram-negative, non-fermentative bacterium that is present ubiquitously in various anthropogenic and natural environmental habitats [36]. It is a frequent colonizer of the rhizosphere Plastics are of various types. One of the most abundant types of plastics is polyethylene terephthalate (PET). PET plastics are made of repeating units of polymer ethylene terephthalate [60]. Plastic bags are another very common plastic products which are made up of low density poly ethylene (LDPE) and/or high density poly ethylene (HDPE) [61]. The reason why PET and most plastics do not easily biodegrade is because the entire plastic structure has very strong C-C bonds that require too much energy to breakdown and plastics do not dissolve in water [62]. However, with the increased plastic accumulation in our environment, microbial life forms have evolved to degrade plastic products to some extent. There are many bacteria that have the ability to degrade PET, Polyethylene (PE), and Polystyrene (PS) but their enzymes only have been able to give moderate turnouts. The enzymes involved in PET degradation are known as PET hydrolases [20].
While working with synthetic plastics, Kenny and his research team [63] used pre-degraded PET as a carbon source and received a yield of only up to 21% with Pseudomonas frederiksbergensis GO23. Whereas, S. maltophilia HA-16 (MN240936) produced up to 38% of PHA with undegraded PET bottles. In another study, Dai and Reusch [64] utilized synthetic plastics by using the pyrolysis oil of polystyrene and reported PHA production up to 48% in tryptic soy broth (TSB) medium with Cupriavidus necator H16. Contrarily, S. maltophilia HA-16 gave a yield of 43% with undegraded or pretreated polystyrene fragments, which is still impressive. These yields indicate an evident ability of S. maltophilia HA-16 to degrade PET bottles and polystyrene. Current research is the first case of PHA production reported from the strain S. maltophilia HA-16 (MN240936).
A surprising finding of this study is the high yield of PHA, i.e., 68.24% in plastic bags. S. maltophilia is a gram-negative, non-fermentative bacterium that is present ubiquitously in various anthropogenic and natural environmental habitats [36]. It is a frequent colonizer of the rhizosphere and, hence, this species is present in various types of soils worldwide [65]. S. maltophilia has also been found as a part of the natural microbiome of various amoebal genera that are free-living [66]. S. maltophilia holds bioremediation capability of sites polluted with hydrocarbons and various xenobiotics [67].
What is more surprising about S. maltophilia is that, not only it is a potential PHA producer, but it is also a potential PHB degrader, as demonstrated by Wani et al. [68]. This bacterium is kind of like hitting two targets with one bullet, where it degrades its creation as well. It can immensely increase recycling in PHA production processes, hence, promoting a circular economy. The use of plastic bags as a carbon source for PHA production by any bacteria is yet unreported. This study could be the first of its kind to report such impressive PHA yields by the undegraded plastic bag as a carbon source, opening up new horizons in the field of plastic bag biodegradation and bioconversion by S. maltophilia HA-16 (MN240936). However, their effectiveness still needs to be improved with the aid of genetic engineering [69].

PHA Film Preparation
PHA polymer film was prepared by adding chloroform into the extracted polymer and evaporating it at 60 • C. The film obtained after preparation with chloroform was somewhat in the shape of fragments as compared to the quality of standard PHB film. Its durability still needs further improvement.
The film obtained after drying with chloroform was very brittle and fragile ( Figure 9). Mohammed et al. [30] reported a similar kind of brittle film made entirely of PHB polymers. He further added that these films are delicate unless made in combination with copolymers. Our lab-scale experiment was carried out with minimum resources so the copolymerization to achieve a proper film out of the extracted polymer was not possible.
Molecules 2020, 25, x FOR PEER REVIEW 14 of 22 producer, but it is also a potential PHB degrader, as demonstrated by Wani et al. [68]. This bacterium is kind of like hitting two targets with one bullet, where it degrades its creation as well. It can immensely increase recycling in PHA production processes, hence, promoting a circular economy.
The use of plastic bags as a carbon source for PHA production by any bacteria is yet unreported. This study could be the first of its kind to report such impressive PHA yields by the undegraded plastic bag as a carbon source, opening up new horizons in the field of plastic bag biodegradation and bioconversion by S. maltophilia HA-16 (MN240936). However, their effectiveness still needs to be improved with the aid of genetic engineering [69].

PHA Film Preparation
PHA polymer film was prepared by adding chloroform into the extracted polymer and evaporating it at 60 °C. The film obtained after preparation with chloroform was somewhat in the shape of fragments as compared to the quality of standard PHB film. Its durability still needs further improvement.
The film obtained after drying with chloroform was very brittle and fragile ( Figure 9). Mohammed et al. [30] reported a similar kind of brittle film made entirely of PHB polymers. He further added that these films are delicate unless made in combination with copolymers. Our labscale experiment was carried out with minimum resources so the copolymerization to achieve a proper film out of the extracted polymer was not possible.

Trace Elements Solutions
Trace elements solution (500 mL) was prepared by weighing 5  Corporation Japan) and mixing them in 100 mL of distilled water and the final volume was raised to 500 mL [42].

HCl (=35%) Solution Preparation
HCl (=35%) solution was prepared by adding 35 mL of concentrated HCl in 50 mL of distilled water and the volume was raised to 100 mL by distilled water.

PHA Detecting Agar
PHA detecting agar (500 mL) was prepared by adding 4 g of nutrient broth, 10 g of nutrient agar, and 0.25 mg of Nile blue A dye in 100 mL of distilled water, and the final volume was raised to 500 mL with distilled water. The media was sterilized in the autoclave (Model: WAC-60, Wisd, WiseStri, Germany) at 121 • C, 15 psi for 20 min. After sterilization, the media was poured aseptically in petri plates, which were pre-sterilized in Digital Oven (SNB-100, hot air sterilizer, Memmert) and stored in a Varioline Intercool cold cabinet at 4 • C until further use [70,71].

Nutrient Broth and Nutrient Agar
The nutrient broth was prepared by dissolving 0.4 g of nutrient broth in 20 mL of distilled water and the final volume was raised to 50 mL with distilled water. The broth was sterilized by autoclaving at 121 • C, 15 psi for 20 min [71].
Nutrient agar (100 mL) was prepared by adding 0.8 g of nutrient broth and 1.5 g of agar in 50 mL of distilled water and was homogenized. The final volume of the medium was raised to 100 mL by distilled water. The medium was autoclaved at 121 • C, 15 psi for 20 min [72].

Fermentation Medium
The fermentation medium was prepared by adding 0.318 g of Na 2 HPO 4 , 0.135 g of KH 2 PO 4 , 0.235 g of (NH 4 ) 2 SO 4 , 0.0195 g of MgSO 4 , 0.05 g of nutrient broth and 0.45 g of pre autoclaved solution of glucose in 10 mL of distilled water. The volume was raised to 50 mL by adding sterile distilled water. Separately autoclaved solution of trace elements was also added in the medium in a concentration of 1 mL/L. The medium was autoclaved at 121 • C, 15 psi for only 5 min to prevent glucose from caramelization [73].

Seed Culture Preparation
Twenty-four hours before fermentation, each colony was aseptically inoculated in 50 mL of autoclaved nutrient broth and incubated at 37 • C/150 rpm in a shaking incubator (Innova ® 43 Incubator Shaker Series) for 24 h to get an overnight old seed culture for the main fermentation process [74].

Sample Collection
Soil, compost, solid waste landfill soil, and industrial effluent samples were collected from paint and plastic industries, sugar mills, food, paper, pulp and cardboard industries, etc. Soil samples were collected aseptically in sterilized polythene bags while effluent samples were collected in sterilized plastic containers from Lahore, Gujrat, Mandi-bahauddin, Narowal, and other areas of the province Punjab, Pakistan [75]. Table 1 shows the geographical distribution of the areas from where the samples were collected.

Waste Collection
Wastes, such as wood chips, cardboard, plastic bottle, wasted polystyrene cups, and waste plastic bags were collected aseptically in sterilized polythene bags from local landfills and dumps of Lahore, Pakistan. The waste was used as a carbon source for the production of PHA [64]. Before inoculation, the waste was shredded into smaller pieces followed by sterilization under the UV hood [76] (Figure 10). Collected plastic bottles were made of PET (Polyethylene terephthalate). It's an aliphatic polyester made of monomers obtained by either esterification of terephthalic acid with ethylene glycol or transesterification of ethylene glycol with dimethyl terephthalate [77]. The presence of a large aromatic ring in the PET repeating units gives the polymer notable stiffness and strength, especially when the polymer chains are aligned with one another in an orderly arrangement [78]. Thick solid colored plastic bags like the ones used in this study are linear HDPE polymers. PE is a polymer formed by radical polymerization of ethylene. Due to the linear structure of HDPEs, they are flexible, durable and tough [79]. Foamed PS used in this research commercially also known as Styrofoam© is a synthetic aromatic hydrocarbon polymer formed of styrene monomers. Its alternating C centers are attached to phenyl groups through σ bonds [80]. Styrofoam is a syndiotactic polymer, which has phenyl groups positioned to alternating sides of the C chain. This structure gives it a highly crystalline and hence a brittle structure [81].
Molecules 2020, 25, x FOR PEER REVIEW 16 of 22 [78]. Thick solid colored plastic bags like the ones used in this study are linear HDPE polymers. PE is a polymer formed by radical polymerization of ethylene. Due to the linear structure of HDPEs, they are flexible, durable and tough [79]. Foamed PS used in this research commercially also known as Styrofoam© is a synthetic aromatic hydrocarbon polymer formed of styrene monomers. Its alternating C centers are attached to phenyl groups through σ bonds [80]. Styrofoam is a syndiotactic polymer, which has phenyl groups positioned to alternating sides of the C chain. This structure gives it a highly crystalline and hence a brittle structure [81].

Isolation and Screening
Qualitative isolation and screening of PHA producing bacteria was initially done by culturing the samples on PHA detecting agar containing Nile blue A stain by the method of Bhuwal et al. [76]. The plates after incubation were illuminated under UV light at 312 nm in UVP Mini Benchtop Transilluminator (Model: TM-10E) for the presence of bright orange fluorescence which indicates PHA accumulation in cells [75]. The initial screening experiments for each sample were run in duplicates. Colonies with orange fluorescence were further selected for isolation by quadrant streaking onto PHA detecting agar plates followed by incubation at 37 °C for 24 h [82].

Submerged Fermentation for PHA Production
The PHA positive colonies after isolation were then subjected to submerged fermentation. Seed culture (24 h old) was aseptically inoculated in 50 mL of autoclaved fermentation medium and incubated at 37 °C/150 rpm in shaking incubator for 72 h [64]. The experiment for each sample was run in duplicates.

Extraction of PHA Produced during Fermentation
After 72 h, the PHA content was extracted by sodium hypochlorite and chloroform digestion method of Kumar [71] with a few modifications. The procedure consisted of multiple rounds of centrifugation at 6000 rpm/15 min, drying, and suspension of weighed dried pellets in solutions of

Isolation and Screening
Qualitative isolation and screening of PHA producing bacteria was initially done by culturing the samples on PHA detecting agar containing Nile blue A stain by the method of Bhuwal et al. [76]. The plates after incubation were illuminated under UV light at 312 nm in UVP Mini Benchtop Transilluminator (Model: TM-10E) for the presence of bright orange fluorescence which indicates PHA accumulation in cells [75]. The initial screening experiments for each sample were run in duplicates. Colonies with orange fluorescence were further selected for isolation by quadrant streaking onto PHA detecting agar plates followed by incubation at 37 • C for 24 h [82].

Submerged Fermentation for PHA Production
The PHA positive colonies after isolation were then subjected to submerged fermentation. Seed culture (24 h old) was aseptically inoculated in 50 mL of autoclaved fermentation medium and incubated at 37 • C/150 rpm in shaking incubator for 72 h [64]. The experiment for each sample was run in duplicates.

Extraction of PHA Produced during Fermentation
After 72 h, the PHA content was extracted by sodium hypochlorite and chloroform digestion method of Kumar [71] with a few modifications. The procedure consisted of multiple rounds of centrifugation at 6000 rpm/15 min, drying, and suspension of weighed dried pellets in solutions of 4% sodium hypochlorite and chloroform. It was later proceeded by incubation at 37 • C/150 rpm in a shaking water bath (Daigger Scientific Inc., Wisd, Model: WSB-30) for 1 h. Finally, 1:1 solution of acetone and ethanol was added and pellets were dried in the thermal oven at 60 • C until the liquid content was evaporated. The extracted PHA was collected by filtration on pre-weighed filter paper followed by drying at 60 • C until the achievement of constant weight [83].

Quantification of Produced PHA
Extracted PHA was quantified as percentage production (%PHA) in cell dry weight (CDW). PHA content was determined as a ratio between the total dry weight of extracted PHA to CDW [84].
The following formulas were applied for the quantification [49]: Cell dry weight (CDW) = weight of falcon tube with dried pellets − weight of empty falcon tube Dry weight of extracted PHA = weight of filter paper with dried filtered PHA − weight of empty filter paper %PHA = Dry weight of extracted PHA CDW × 100

Characterization of the Extracted PHA by FTIR
The PHA extracted from the most efficient PHA producing microbial isolate was sent for Fourier-Transform Infrared spectroscopy (FTIR) (model: IR-Prestige) characterization to Center for Advanced Studied in Physics (CASP), Government College University, Lahore for functional group analysis through signal peaks recorded in the form of percentage transmittance in the range of 4000-400 cm −1 [85].

Molecular Identification of the Most Efficient PHA Producing Strain
The most efficient PHA producing bacterial strain was identified by 16S rRNA sequencing by sending samples to Macrogen sequencing company, Seoul, Korea. The sequence of the bacterial strain was further subjected to homology analysis through Basic Local Alignment Search Tool (BLAST) [36]. The phylogenetic tree of the identified bacterial isolate was created using Jalview based upon the BLAST results [86]. After the BLAST, the sequence of the identified strain was submitted into the GenBank nucleotide sequence database via BankIt (MN240936) [87].

Optimization of Cultural Conditions
Four parameters of cultural conditions were optimized for PHA production. The first three parameters included the time of incubation (24-96 h), incubation temperature (25-40 • C), and the pH of the fermentation medium (6.0-8.0). Waste, such as wood chips, cardboard, plastic bottles, wasted polystyrene cups, and waste plastic bags were used as alternate carbon sources instead of glucose in the fermentation medium. Optimization of these sources was done to analyze their potential to be used as cheap, sustainable, and alternate carbon sources [11].

PHA Film Preparation
PHA film was prepared by adding about 5 mL of chloroform in 0.5 g of extracted dried PHA in a 15 mL falcon tube and set for evaporation in a drying oven at 60 • C until chloroform was completely evaporated [30].

Statistical Analysis
Computer software CoStat, cs6204W.exe application was applied to carry out the statistical analysis [88]. All of the experiments were run in duplicates to determine the standard error margins in the final yields. Replicates significant differences were presented as Duncan's multiple range tests in the form of probability (p) values. These calculations were done to validate the reproducibility of the experiments.

Conclusions
Paint industry soil resulted in the isolation of indigenous and unique PHA producing bacteria Stenotrophomonas maltophilia HA-16 (MN240936). Fermentation cultural parameters, such as incubation time, pH, temperature, and carbon sources were found to have a significant effect on PHA production as they increased the PHA yield by 1.16 folds. Moreover, utilizing an untreated plastic bag as a carbon source instead of glucose for PHA production was a standout finding with a yield difference of less than 1.1 folds. However, as the research on this unique strain is extremely limited, it still requires extensive studies to turn this bacteria beneficial for industrial use.