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Bioplastics against Microplastics: Screening of Environmental Bacteria for Bioplastics Production

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Aisha S. Alwuhaib, Vitaly Zinkevich, Tamar Kartvelishvili, Nino Asatiani and Nelly Sapojnikova

Submitted: 09 November 2022 Reviewed: 03 January 2023 Published: 25 January 2023

DOI: 10.5772/intechopen.109756

Advances and Challenges in Microplastics IntechOpen
Advances and Challenges in Microplastics Edited by El-Sayed Salama

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Advances and Challenges in Microplastics

Edited by El-Sayed Salama

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Abstract

Polyhydroxyalkanoates (PHAs) are biopolymers produced by numerous bacteria and can be used in the production of bioplastics. PHAs are synthesized by microorganisms by fermentation of carbon sources. Due to the different monomer structures of PHAs, there are many kinds of PHAs, and their corresponding material properties are also very different. Thus, the search for bacteria producing the PHAs is of great interest. In this study, the bacteria isolated from the environment were analyzed for the presence of PHA. PHA production was tested with staining methods Sudan Black B, Nile Blue, and Nile Red. The presence of a PHA synthase gene (phaC) was confirmed by PCR amplification. PHAs were extracted from the strains and characterized by the FTIR spectroscopy method. A biochip for a fast screening of environmental samples for the presence of PHA-producing bacteria was designed. The biochip contained 11 probes for coding class 1, 2, and 3 PHA synthase genes.

Keywords

  • polyhydroxyalkanoates (PHAs)
  • PHA synthase gene
  • environmental bacteria
  • biochip
  • bioplastics

1. Introduction

Plastics (polymeric materials) are highly functional materials that have become an essential part of the products we use in our daily life due to their ease of production and robustness. Around 360 million tons of plastics have been produced in 2020 around the world [1]. As more plastics are used, especially with their short-use life span, more waste surfaces. Plastic trash photodegrades into smaller fragments (microplastics). As it was emphasized in Ref. [1], the worldwide use of disposable face masks during the pandemic time and still now is an additional source of microplastics in the environment. Plastics are majorly manufactured from petrochemical feedstock, accounting for 80% of the total produced plastics. The building blocks for the polymerization of bioplastics are biopolymers cultivated from renewable production pathways, such as polymers from microorganisms [2, 3]. Bio-based polymers have a lower carbon footprint than petrochemicals because they utilize biological materials and waste, which convert biocaptured CO2 into durable polymeric materials. It is a well-established fact that microorganisms are equipped with diverse metabolic activities that enable them to work as biorefineries for transforming a wide range of petrochemical and organic wastes into high-value specific products, such as polyhydroxyalkanoates biopolymers, while positively impacting the carbon cycle by consuming atmospheric CO2 [4]. The chemical structure of PHA is shown in Figure 1. PHA is a linear polyester that contains 3-hydroxy fatty acid monomers [5]. The most commonly produced PHA is poly 3-hydroxybutyrate (PHB), where the alkyl group is R = CH3. However, there are over 150 different monomers, such as polyhydroxyvalerate (PHV, R = C2H5), polyhydroxyhexanoate (PHH, R = C3H7), and polyhydroxyoctanote (PHO, R = C4H9). PHAs are high molecular weight linear polyesters, ranging in size from 50 to 10,000 kDa [2], characterized by a diversity of structures defined by the length of the carbon chain, referred to as short-chain length (SCL), medium-chain length (MCL) [6], and long-chain length (LCL) [7] PHAs.

Figure 1.

Chemical structure of polyhydroxyalkanoates (PHAs).

A very important property of biopolymers is hydrophobicity, which determines their solubility, biodegradability, and biocompatibility. As PHAs contain chains of hydrophobic groups with different lengths and structures, and at the same time PHAs are poorly hydrophilic due to the presence of carbonyl groups, the hydrophobicity-hydrophilicity balance is a very important point in the selection of appropriate bacteria that give the desired biomaterials [2]. PHAs can exist as homo or copolymers. PHB is a homopolymer, which has a linear isotactic structure, that is, highly crystalline making it brittle and unsuitable for many applications. This problem can be circumvented by forming a copolymer. The first commercially manufactured PHA, Biopol®, is copolymer produced from poly (3-hydroxybutyrate-co-3-hydroxyvalerate (PHB/PHV) that had an increased side chain length making it less crystalline and more ductile [8]. These biopolymers have been produced commercially since the 1980s, and are currently marketed as Mirel®. The combinations in different proportions of the available PHAs create copolymer plastics with various properties [9]. PHAs are synthesized by various types of bacteria in the form of water-insoluble granules, and are stored as carbonosomes within the cell cytoplasm [10]. Bacteria capable of producing PHAs include species of Alcaligenes, Bacillus, Burkholderia, Ralstonia, Pseudomonas, Aeromonas, and Thiococcus, among others [11, 12]. PHA synthases and depolymerases, which are the catalyst enzymes for PHA production and degradation, respectively, are located at the membrane of the storage organelles and control the amount of PHA stored by the cell. There are eight major pathways for PHA synthesis. The main metabolite, acetyl-CoA, provides the various lengths of 3-hydroxyalkanoyl-CoA, which act as substrates for PHA production [13]. These pathways are intricately linked to central metabolic pathways via glycolysis, Kreb’s cycle, β-oxidation, fatty acid synthesis, and others. Many enzymes and genes are involved in PHA synthesis. Bacterial cells normally grow when carbon (i.e., fructose and glucose) and nitrogen (ammonium) are present in the medium/environment. Polyhydroxybutyrate (PHB) is produced when the nitrogen source is depleted and carbon is present and will be utilized after refeeding cells with a nitrogen source [14].

The limiting step in the commercial development of non-petrochemical-based produced plastics is the biopolymer yield obtained after fermentation and the cost of its recovery. As the main drawback of bioplastics is the cost of the fermentation process, this has led to searches for cheaper carbon sources for fermentation, which has included activated sludge, paper and food wastes, wastewater, and various oil wastes [15]. Additionally, a lot of effort has been directed toward isolating PHA-producing bacteria from high carbon, low nitrogen, or phosphorus environments that might give greater yields in batch or continuous cultures [16]. The use of different carbon sources in fermentations and isolating producing strains from carbon-rich environments has led to the discovery of novel PHA polymers with different properties. In this study, the results of screening of environmental samples for PHA-producing organisms, isolation and characterization of the microorganisms, detection of genes, coding for enzymes involved in PHA synthesis, and physical characterization of the produced PHAs biopolymers are presented.

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2. Analysis of single bacterial isolates for PHA production using staining methods

Two types of samples were collected for this study. The sludge samples were collected from the Petersfield Southern Water Treatment plant (Hampshire, UK). The three compost samples were collected from (1) a graveyard, (2) food-based waste and (3) garden waste (Portsmouth, UK). A mass of 0.5 g of sludge and compost was added to the 50 ml of the mineral salt medium (MSM) and Lysogeny broth (LB) medium (Fisher BioReagents, UK); each sample was inoculated in duplicate for each media and was transferred to the Laboratory of Molecular Microbiology (University of Portsmouth, UK). These bottles were incubated overnight in Innova™ 4000 incubator shaker (New Brunswick Scientific, USA) at 150 rpm at 25°C. Overnight-grown cultures were used for the isolation of single bacterial colonies. Serial dilutions were made into six concentrations using 0.9% NaCl in dH2O for each culture after incubation. Six dilution tubes were generated for each media (two types of media) and sample (two types of samples), resulting in a total of 48 tubes. Each individual sample from each of the 48 tubes was used to inoculate one LB agar Petri dish. Some of the single colonies that grew from the two most diluted cultures were collected. The collected single colonies were regrown on separate LB agar to ensure that each Petri dish contains a pure colony. The colonies were picked up from Petri dishes and inoculated in individual universal tubes with 5 ml LB, rotating at 37°C overnight at 150 rpm. As a result, 73 bacterial isolates were recovered.

Different approaches have been used for the screening of PHA-producing microbes and/or the imaging of PHA granules [17]. The methods most widely used for detecting PHAs are staining techniques using Nile Red [18], Nile Blue A [19], and Sudan Black B [20]. Due to the lipophilicity of the dyes, the tests are very useful, but they have the ability to nonspecifically bind to other lipid droplets, membranes, and cell envelopes, leading to an incorrect answer [21, 22]. Sudan Black B (SBB) is a diazo fat-soluble dye. PHAs are observed as black granules with SBB by bright-field microscopy. Nile Blue A and Nile Red are highly fluorescent organic dyes belonging to the benzophenoxazine family. Nile Red is a neutral molecule that is poorly soluble in water, and its chromophore is highly susceptible to changes in solvent polarity, showing little or no fluorescence in most popular solvents. Nile Blue A is a cationic dye and is thus more soluble in water than Nile Red. Both fluorescent dyes are particularly useful for visualizing hydrophobic cell structures, such as membranes or lipid-like inclusions (PHA granules). PHA inclusions appear as brightly fluorescent red/orange granules with Nile Blue A and Nile Red [23].

In this study, 73 bacterial isolates were tested for PHAs production using all above mentioned staining methods. When the ability of isolates to produce PHA was screened by Sudan Black B staining, cultures showed granules filled up with dark staining, as PHA granules can be observed as bluish dark spots under a light microscope (Figure 2). Out of the tested 73 isolates, 48 isolates tested positive for SBB staining and were considered to produce PHA granules.

Figure 2.

Screening of bacterial strains for PHA production using Sudan Black B dye. Panel A shows the positive result with different isolates from compost samples stained with SBB. Panel B shows the positive and negative results for different isolates from the sludge sample stained with SBB.

To test the possible presence of PHAs additional stains were used. The Sudan Black B-positive strains were stained by Nile Blue A and Nile Red. Both of these dyes are used to detect PHAs production in bacteria grown on solid media. Bacteria were grown on the LB agar plates in the presence of DMSO, incubated at 37°C overnight, and then exposed to UV light to check the fluorescence in the cells (Figure 3).

Figure 3.

Screening of different isolates for PHA production using Nile Blue A dye (Panel A) and Nile Red dye (Panel B).

Eighteen colonies out of 73 showed bright orange fluorescence by both fluorescent dye staining. Unfortunately, the staining methods cannot give an unambiguous answer. Therefore, other testing methods were essential to confirm the presence of PHAs in bacteria on the genetic and structural levels.

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3. PCR amplification of PHAs genes

The genomic DNA from the 48 strains that tested positive in the SBB staining test was analyzed for the presence of the PHA synthase gene (phaC) using the PCR amplification method. Genomic DNA was recovered using the DNeasy Blood and Tissue kit (Qiagen®) according to the manufacturer’s instructions. The phaC gene was amplified using F I-179 L forward primer and R I-179R reverse primer (Table 1) [24]. Final reactions contained: 2x GoTaq® Green Master Mix and 10 μM of each oligonucleotide primer. PCR was carried out as indicated: initially denatured at 94°C for 10 min; followed by 35 cycles of 94°C for 1 min, 53°C for 1 min, and 72°C for 1 min followed by the final extension step at 72°C for 10 min.

PrimerSequences of primers 5′ → 3′Gene targetSize of amplicon, bp
8FAGAGTTTGATCCTGGCTCAG16 S rRNA1500
1492RGGTTACCTTGTTACGACTT
F I-179LACAGATCAACAAGTTCTACATCTTCGACphaC & phbC540
R I-179RGGTGTTGTCGTTGTTCCAGTAGAGGATGTC
PhaCF1BOTCMYCTSKACTGCSCTGGYGphaC247
PhaCR2BOYWGCTRGACYAGACCTGGAT

Table 1.

Oligonucleotide primers are used in PCR.

The phaC gene was also amplified using the PhaCF1BO forward primer and PhaCR2BO reverse primer (Table 1) [25]. Final reactions contained: 2x GoTaq® Green Master Mix and 10 μM of each oligonucleotide primer. PCR was carried out as indicated: initially denatured at 95°C for 10 min; followed by 30 cycles of 95°C for 1 min, 57°C for 45 sec, and 72°C for 2 min followed by the final extension step at 72°C for 5 min.

The selective results of the amplification using the pair of primers F I-179 L/R I-179R are presented in Figure 4. The isolates tested positive for the presence of the phaC gene in case in their DNA correct amplicon size (540 bp) was revealed when run on the 1.2% agarose gel (Figure 4A and B).

Figure 4.

The DNA amplicons of phaC gene (540 bp) with primers: F I179L/ R I179R from the bacterial isolates. Panel A: Lane 1: 1 kb DNA Ladder; Lane 2: 3/1; Lane 3: 10/1; Lane 4: 12/1; Lane 5: 17/1; Lane 6: 19/1; Lane 7: 20/1; Lane 8: 22/1; Lane 9: 25/1; Lane 10: 100 bp DNA Ladder; Lane 11: 9/1; Lane 12: 6/2; Lane 13: 8/2; Lane 14:10/2; Panel B: Lane 1: 1 kb DNA Ladder. The isolates presented in the Lanes 2 to 13 are as follows: Lane 2: 13/2; Lane 3: 16/2; Lane 4: 17/2; Lane 5: 18/2; Lane 6 to 9 negative results for isolates 2/1, 4/1, 3/2, 14/2; Lane 10: 9/2; Lane 11: 19/2; Lane 12: 24-1/2; Lane 13: 24/2.

The selective results of the amplification using the pair of primers PhaCF1BO/PhaCR2BO are presented in Figure 5. According to Figure 5, some of the isolates tested positive for the presence of the phaC gene in their DNA as shown in lanes (3, 4, 6, 7, and 9) (Figure 5A) and lanes (2, 7, 9, 11, and 12) (Figure 5B). These isolates produce the correct amplicon size (247 bp) when run on the 1.2% agarose gel.

Figure 5.

The DNA amplicons of phaC gene (247 bp) with primers: PhaCF1BO/PhaCR2BO from the bacterial isolates. Panel A: Lane 1: 1 kb DNA Ladder; Lane 2: 3/1; Lane 3: 10/1; Lane 4: 12/1; Lane 5: 17/1; Lane 6: 19/1; Lane 7: 20/1; Lane 8: 22/1; Lane 9: 25/1. Panel B: Lane 1: 1 kb DNA Ladder. The isolates presented in the Lanes 2 to 12 are as follows: Lane 2: 9/1; Lane 3: 6/2; Lane 4: 8/2; Lane 5: 10/2; Lane 6: 3/1; Lane 7: 10/1; Lane 8: 12/1; Lane 9: 17/1; Lane 10: 19/1; Lane 11: 20/1; Lane 12: 22/1.

Pseudomonas oleovorans NCTC 10692 (Bacteria Collection from Public Health England, ATCC 8062), which is the producer of PHA [26, 27] was used as a positive-control strain. Escherichia coli XL-1 Blue (Agilent Technologies), which is not able to produce PHA, was used as a negative control strain.

Out of the tested 48 strains, 18 strains tested positive for the presence of the phaC genes confirmed by one or both types of PCR and by the presence of PHA granules confirmed by the Nile Blue A and Nile Red staining methods. These 18 strains, in addition to two strains (9/1 and 9/2) that tested negative with PCR and positive with Sudan Black B were sent for sequencing after cloning their 16S rRNA PCR products into competent E. coli cells (Table 2). The 16S rRNA gene was amplified using the primer pair 8F and 1492R (Table 1) [28], the conditions were pointed in Ref. [29]. PCR products were ligated into a vector plasmid DNA and transformed into E. coli competent cells using the TOPO® TA Cloning® Kit (Invitrogen by Life Technologies, UK) following the manufacturer’s protocol. Sequencing was performed by GATC Biotech Ltd., London Bioscience Innovation Center, the identification was done through the BLAST search. The strains, which were isolated from sludge and compost samples, belonged to multiple bacterial families (Table 2), including the Enterobacteriaceae (7 strains, 35%), Micrococcaceae (2 strains, 10%), Alcaligenaceae (4 strains, 20%), Moraxellaceae (1 strain, 5%), Aeromonadaceae (1 strain, 5%), Bacillaceae (3 strains, 15%), and Pseudomonadaceae (2 strains, 10%). Most of the isolates from the environmental samples belong to Enterobacteriaceae (Citrobacter sp., Raoultella, Klebsiella, Serratia, and Acinetobacter), and they are commonly known as producers of polyhydroxyalkanoates [30]. The genomes of the identified isolates contain different classes of PHA synthase (I, II, and IV), producing different types of PHA based on the number of carbon atoms. Thus, Pseudomonas contains either class I or II of PHA synthase [12] and can produce medium-chain length PHA, but some strains are able to synthesize both SCL- and MCL-PHAs; Alcaligenes can produce short-chain length PHA.

Sample No.StrainsPresence of phaC gene, PCRSudan Black BNile Blue A and Nile RedPresence of plasmidNCBI BLAST matches (Strain identification name)Identity, %
13/1++++Citrobacter sp.97
29/1−+−+Arthrobacter sp.97
310/1++++Raoultella/Klebsiella99
412/1++++Raoultella/Klebsiella98
517/1++++Alcaligenaceae; Achromobacter97
619/1++++Citrobacter sp.99
720/1++++Raoultella/Klebsiella99
822/1++++Raoultella/Klebsiella99
925/1++++Serratia98
106/2++++Aeromonas99
118/2++++Alcaligenes98
129/2−+−+Arthrobacter99
1310/2++++Bacillus99
1413/2++++Bacillus99
1516/2++++Bacillus99
1617/2++++Alcaligenes99
1718/2++++Alcaligenes99
1819/2++++Acinetobacter99
1924-1/2++++Pseudomonas99
2024/2++++Pseudomonas99

Table 2.

The characteristics and sequence identification of the selected bacterial isolates from the environmental samples.

Note: (−) indicates a negative test result; and (+) indicates a positive test result.

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4. Confirmation of the presence of a PHA pathway

The isolates were screened for the presence of plasmids, as they often carry the PHA biosynthetic pathway. Plasmids were detected in all 20 selected strains for PHA production (Table 2). To show that the plasmid and PHA production were co-inherited, a non-PHA-producing strain of E. coli XL-1 Blue (Agilent Technologies) was transformed with a plasmid. This will allow the identification of plasmids that can express the PHA operon in E. coli. As a control, a known PHA-producing plasmid from Pseudomonas oleovorans NCTC 10692 was transformed to show that the insertion was successful. The plasmid DNA was purified from the bacterial isolates grown in culture in the LB media using the Zyppy miniprep kit (ZYMO Research, Irvine, CA) according to the manufacturer’s protocol. Purified plasmid DNA was used for the transformation of E. coli XL-1 Blue by electroporation (MicroPulser Electroporator, BioRad). Before performing the transformation experiments, the PHA-producing strains were characterized for their antibiotic resistance profiles. This was done to identify possible determinants that could be used as counter-selective agents, assuming that it was plasmid bourn. The 20 strains were tested for their susceptibility to 16 different antibiotics, delivered in discs to check the resistance profiles of these strains grown on solid media. Four of the PHA-producing strains (6/2, 8/2, 10/2, 13/2) were sensitive to all the tested antibiotics. As all the PHA-producing strains were resistant to ampicillin and/or nalidixic acid, but the E. coli XL-1 Blue host strain was not, these two antibiotics were then used as a selection for the plasmid when it was transformed into the PHA-negative strain (E. coli XL-1 Blue). After the electroporation, transformed cells were inoculated on LB agar containing ampicillin or nalidixic acid. After overnight incubation, the colonies obtained were picked up and grown in LB medium with the appropriate antibiotics chosen (ampicillin and nalidixic acid). These cultures were then screened for PHA-producing genes by using the staining methods, PCR amplification, and sequencing. Four strains (3/1, 10/1, 17/1, and 19/1) in addition to the positive-control strain were successfully transferred into E. coli XL-1 Blue, and were tested positive for PHA production and the presence of the phaC gene. These four strains were isolated from the sludge samples collected from the Petersfield Southern Water Treatment plant. The pointed bacterial strains provide plasmids, which have the potential to synthesize PHA for industrial/commercial purposes when co-introduced into the same genetic background.

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5. Physical characteristics of PHAs

The PHA extracted from the 20 environmental samples that tested positive for phaC genes and/or the staining methods were analyzed using the FTIR spectroscopy method.

Pure bacterial overnight culture grown in LB media (10 ml) at 37°C was used for the PHA extraction and purification. Bacterial cells were pelleted at 4000 rpm for 25 min at RT using glass conical centrifuge tubes (Sigma, UK). The pellet was suspended in 10 ml of 0.1% sodium hypochlorite and centrifuged for 30 min at 4000 rpm. The supernatant was removed, and the pellet was washed with 5 ml of sterile water, 5 ml of acetone, and 5 ml of methanol consequently. The pellet was dissolved in 5 ml of chloroform and left overnight at RT to evaporate on a glass Petri dish (Figure 6) [31]. The PHA biopolymer was collected, weighed, and analyzed using a physical method.

Figure 6.

PHA was obtained as a white powdery substance on a glass Petri dish.

The PHAs extracted from the isolates were analyzed using PerkinElmer®'s Spectrum 1™ FT-IR Spectrometer (USA); spectral range 4000–400 cm−1 with a spectral resolution of 4 cm−1 [32]. The characteristic absorption peaks were used to interpret the presence of specific functional groups in the extracted polymers. The yield of PHA in mg/10 ml extracted from the environmental pure bacterial isolates varied from 0.1 mg (24/2) to 8.8 mg (20/1). The main FTIR spectral peaks for the presumptive PHAs extracted from 20 studied bacterial isolates; from the PHA-producing bacterial strain (+ve), used as a positive control, along with peaks for the commercial PHB (Sigma, USA) are presented in Table 3.

Strain IDBond name and functional group
carbonyl group C=OEster group stretching C–ON–H amide protein in the polymerC–H stretching methyl and methylene–CH group–OH group
Wavelength cm−1
PHB1720.862——2976.1331278.590—
+ve—1650.0541536.8892961.9771275.3343282.092
3/1—1650.2121532.6062961.5071285.7253283.034
9/1——1531.962———
10/1—1649.8841537.6002163.5951234.149—
12/1—1632.6071531.0582922.2351258.489—
17/1—1633.1901533.5942961.5551258.5943282.994
19/1—1645.3821537.1592962.1021285.469—
20/1—1634.3941530.9412962.5791285.4543284.661
22/1—1643.3541532.3212961.3081259.4013280.077
25/1—1650.2011533.1002962.1691285.9533282.528
6/2—1650.2681537.788—1233.206—
8/2—1653.2751532.0552357.763——
9/2—1637.7331532.606———
10/2—1646.2771530.9202922.6921259.232—
13/2—1650.3931538.0622357.8901215.4173272.019
16/2—1650.6501536.9012159.8991236.901—
17/2—1638.0421526.1252965.5481229.0053297.117
18/2—1643.0421537.9812161.1821234.6793282.213
19/2—1635.1901530.7882361.2361248.5393277.935
24-1/2—1632.858—2933.5811226.658—
24/2—1635.579—2359.4621249.1153347.794

Table 3.

Functional groups identified by the FTIR method for PHA analysis.

According to Table 3, the spectra of the standard PHB show the peak at 1720.88 cm−1, which corresponds to the C=O stretch of the ester group, and the peak at 1278.59 cm−1, which corresponds to –CH group [33]. These peaks are similar to the published spectra peaks of PHB [34]. The positive PHA production bacterial strain (+ve) showed the peak at 1650.05 cm−1, which corresponds to ester carbonyl group (C–O), and accompanying peak at 1275.33 cm−1, which corresponds to –CH group. The results of the FTIR analysis of the PHA extracted from the 20 environmental samples showed a resemblance to the positive strain’s characteristic absorption peaks, which indicates the presence of the PHA functional groups in the samples. As it was mentioned above, the positive-control strain was confirmed as a PHA producer by PCR and the staining methods. The strain 9/1 was tested positive by Sudan Black B, negative for the phaC gene by the PCR method, and it did not show any characteristic absorption peaks corresponding to PHA production. As a result, it was concluded that this strain was not a PHA producer.

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6. Biochip analysis of environmental samples for the presence of PHA-producing bacteria

A low-density biochip as a collection of DNA probes arranged on a solid matrix was the first time applied for rapid screening of PHA-producing organisms in the environment. This technique performs detection using a single undivided environmental sample and allows the identification of different genes in one reaction. The development of a biochip includes the following steps: design of the appropriate probes; equalization of the hybridization capacity of the probes for their inclusion in the biochip prototype; environmental DNA preparation for hybridization; and the fine-tuning of a biochip for the optimization conditions of bacteria detection.

Eleven oligonucleotide probes for the phaC genes responsible for the metabolic pathways involved in PHA production were designed (Table 4). The phaC gene codes a subunit common to the four known PHA synthase classes (class I, class II, class III, and class IV) [35], which are found in different species of bacteria; the PHA C polypeptide subunit varies in molecular weights. The BLAST search in the NCBI database (http://www.ncbi.nlm.nih.gov) was used to derive the sequences used for the probe design. The complete nucleotide sequence for the genomic DNA of PHA-producing microorganisms was downloaded and saved in FASTA format. The sequence was loaded into the SeaView software, which is a sequence alignment editor allowing manual or automatic alignment through an interface with the CLUSTALW program. Region variations in the sequences were identified from multiple alignments generated by the Clustalo option in SeaView [36] or creating multiple alignments of protein sequences by Muscle [37]. Based on final alignments, conserved parts of the sequences of each gene were chosen and used for probe designing using the OLIGO 7 program [38]. The phylogenetic tree generated after a distance-based analysis using the Kimura 2 model and neighbor-joining algorithm showed a clear distinction between the classes of phaC genes. After the phylogenetic analyses, the probes were designed to cover different classes of PHA synthase.

SpeciesGeneProbeSequence 5′ → 3′Probe length, bp
Burkholderia spp.PHA synthase (phaC) gene
poly-beta-hydroxybutyrate polymerase gene		Probe 1TCA ACA AGT TCT ACA TCC TCG21
Burkholderia spp.poly-beta-hydroxybutyrate polymerase geneProbe 2CGT GCA TCA ACA AGT TCT ACA21
Burkholderia pseudomalleipoly-beta-hydroxybutyrate polymerase geneProbe 3TCT GCG GAA TAC CTA TCT CGA21
Synechococcus sp. MA19PhaC (phaC) geneProbe 4TTT AGG TAA CAT TCG CAT GCC21
Synechococcus sp. MA19PhaC (phaC) geneProbe 5TTT AAT GCT CAA ACC CCG ACA21
Pseudomonas sp.Poly(R)-hydroxyalkanoic acid synthaseProbe 6GCC ACA GAT CAA CAA GTT CTA21
Pseudomonas sp.Poly(R)-hydroxyalkanoic acid synthaseProbe 7TGA TCT GGA ACT ACT GGG TCA21
Pseudomonas chlororaphisPoly(R)-hydroxyalkanoic acid synthaseProbe 8CCG GGT ACT TAT GTC CAT GAA21
Aquabacterium sp. A7-YPHA synthase (phaC) geneProbe 9TAT CTC GAA AAC AAG CTC AGC21
Cupriavidus sp.polyhydroxyalkanoate synthase (phaC) geneProbe 10ATG GCG ACC GGC AAG GGT GCG21
Uncultured bacterium clone, Cupriavidus sp.PHA synthase (phaC) gene
poly-beta-hydroxybuterate polymerase		Probe 11CGT GCA TCA ACA AGT ACT ACA21

Table 4.

Probes used in the biochip.

Probes (4–10) target PHA synthase (phaC) gene, probes (2,3) target poly-beta-hydroxybutyrate polymerase gene, and probes 1 and 11 target both genes.

The 3D dendrimeric matrixes for biochip preparation were manufactured at Tbilisi State University, Andronikashvili Institute of Physics, Georgia. All procedures concerning dendrimeric matrix activation for probes immobilization, and dendrimeric matrix deactivation for environmental DNA hybridization are presented in our previous publication [39].

In order to check and evaluate that the chosen probes have similar hybridization capacities, the cassette approach developed and published in Ref. [40] has been used. If the probes exhibit the same hybridization characteristics, they should give equal fluorescent signals in hybridization reactions. The result of the estimation of the hybridization capacity of the proposed probes showed that the probes exhibited comparable hybridization signals and were characterized by the mean signal-to-noise ratio S/N = 12 (data not shown). These probes were tested against DNA obtained from environmental samples (sludge and compost) samples. Total DNA was purified from the environmental samples (sludge or compost) using MO BIO PowerSoil® DNA Isolation Kit (MO BIO technologist, Inc.). The procedures for DNA preparation for hybridization that include DNA amplification, fragmentation, and fluorescent labeling are described in Ref. [40]. Two sludge and three compost environmental samples were analyzed for the bacterial assemblage, revealing PHAs producing capacity using a biochip method. Figure 7 represents the proportion/interrelation of the issued by the probes´ bacterial species in the functional assemblage of sludge T2, as the equalized biochip is the basis for the estimation of the microbial ratio. The hybridization signal S/N above 1.5 was counted as a reliable result.

Figure 7.

The results of DNA from T2 sludge sample hybridization with the biochip. Panels A and B represent the arrangement of the probes on the biochips. The red circles and PCM are the position control markers. The white circles mark the position of the reliably visible signals detected after hybridization with the probes for phaC gene. Panel C shows the hybridization signal, presenting a signal-to-noise ratio, for the probes on the biochip. The data presented are mean values ± SD.

According to Figure 7 and Table 4,

  • Probes 1, 2, and 3 are probes for the identification of Burkholderia sp. For all of them, the identification on the biochip is under the threshold of detection (S/N < 1.5).

  • Probes 4 and 5 are different probes for the identification of Synechococcus sp. MA19. The probes are characterized by identical hybridization capacity (cassette assay); however, probe 5 is more efficient in the case of real ecological samples.

  • Probes 6, 7, and 8 are probes for the identification of Pseudomonas sp. All of them are characterized by S/N > 1.5.

  • Probe 9 is a probe for the identification of Aquabacterium sp. A7-Y. The hybridization signal is reliable (S/N > 1.5).

  • Probes 10 and 11 are probes for the identification of Cupriavidus sp. The identification of this bacteria on the biochip by probe 10 is reliable, but by probe 11 under the threshold of detection (S/N < 1.5).

Therefore, the bacteria species present in sample T2 are Synechococcus; Pseudomonas; Cupravidus, and Aquabacterium. The first three species are very well-known PHA producers.

Figure 8 represents the proportion of the bacterial species in the functional assemblage of compost T5.

Figure 8.

The results of DNA from T5 compost sample hybridization with the biochip. Panels A and B represent the arrangement of the probes on the biochips. The red circles and PCM are the position control markers. The white circles mark the position of the reliably visible signals detected after hybridization with the probes for phaC gene. Panel C shows the hybridization signal, presenting as a signal-to-noise ratio, for the probes on the biochip. The data presented are mean values ± SD.

According to Figure 8 and Table 4,

  • Probes 1, 2, and 3 are probes for the identification of Burkholderia sp. For all of them, the identification on the biochip is above the threshold of detection (S/N > 1.5).

  • Probes 4 and 5 are different probes for the identification of Synechococcus sp. MA19. The probes are characterized by identical hybridization capacity (cassette assay); however, probe 5 is again more efficient in the case of real ecological samples.

  • Probes 6, 7, and 8 are probes for the identification of Pseudomonas sp. All of them are characterized by S/N > 1.5; however, the signals from probes 6 and 7 are much stronger than for probe 8.

  • Probe 9 is a probe for the identification of Aquabacterium sp. A7-Y. The hybridization signal is reliable (S/N > 1.5).

  • Probes 10 and 11 are probes for the identification of Cupriavidus sp. The signal from both probes is reliable; however, the signal of probe 10 on the biochip is again much stronger than of probe 11.

Therefore, all studied types of bacteria are present in sample T5. Dominant bacteria species in sample T5 are Synechococcus, Pseudomonas, and Cupravidus, well-known PHA producers.

Table 5 summarizes the result obtained with biochips for the studied environmental samples.

T2 sludgeT4 sludgeT5 compost (food-based waste)T6 compost (warm garden waste)T7 compost (graveyard)
Probe 1−+++−
Probe 2−−++−
Probe 3−−+−−
Probe 4−−+++
Probe 5+++++
Probe 6++++−
Probe 7+−+++
Probe 8−−++−
Probe 9+−++−
Probe 10+++++
Probe 11−−++−

Table 5.

Summary of the phaC gene detected in the biochip from environmental samples.

The compost samples T5 and T6 contain all studied bacteria species with phaC gene. The bacteria spp. Synechococcus and Cupravidus (probes number 5 and 10) were detected in all environmental samples (sludge and composts). The absence of hybridization signals on the biochip for some bacteria might be explained by the absence of these bacteria in the samples.

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7. Conclusions

Bioplastics are dramatically favored over oil-based plastics due to their ability to degrade. The use of such natural polymers would help counteract the current accumulation of standard nonbiodegradable polymers in the global environment. Bioplastics have many applications, such as packaging materials, biomedical implants, drug delivery systems, and biofuels. The screening of the indigenous bacteria in the environmental samples is the first key step on the way of the selection of bioplastic-producing organisms. In this study, the conventional chemical, molecular biological, and physical methods were successfully applied for the screening of the environmental bacteria-producing PHAs, biopolymers that can be used to produce bioplastics. Twenty bacterial strains have tested positive for phaC genes using the PCR method and/or the Sudan Black B, Nile Blue, and Nile Red staining methods. The PHAs were extracted from these bacterial strains and characterized using the FTIR method. The main FTIR spectral peaks for the PHAs extracted from 19 studied bacterial isolates resemble the peaks for the PHAs isolated from PHA-producing bacterial strain (+ve) that was used as a positive-control strain, indicating the presence of the PHA functional groups in the samples. Diagnostic biochip was first time explored as a fast method for primary screening of PHA-producing microorganisms in environmental samples. The hybridization results showed that the designed probes successfully detected the phaC genes, which covered three classes of PHA synthase in the environmental samples. Class I resides in Cupriavidus sp.; classes I and II are detected in Pseudomonas sp., and class III are found in the Synechococcus and Burkholderia spp. The bacterial composition in the environmental samples revealed the frequently encountered Synechococcus, Pseudomonas, and Cupravidus species, well-known PHA producers. Moreover, the use of biochips and staining procedures for the detection of biopolymers-producing bacteria will allow the screening of microorganisms for novel PHA production pathways.

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Written By

Aisha S. Alwuhaib, Vitaly Zinkevich, Tamar Kartvelishvili, Nino Asatiani and Nelly Sapojnikova

Submitted: 09 November 2022 Reviewed: 03 January 2023 Published: 25 January 2023

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution 3.0 License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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