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Start a Research on Biopolymer Polyhydroxyalkanoate (PHA): A Review

Residues and Resource Reclamation Centre, Nanyang Environment and Water Research Institute, Nanyang Technological University, 1 Cleantech Loop, 637141, Singapore
Division of Environmental and Water Resources, School of Civil and Environmental Engineering, Nanyang Technological University, 50 Nanyang Avenue, 639798, Singapore
Author to whom correspondence should be addressed.
Polymers 2014, 6(3), 706-754;
Submission received: 30 January 2014 / Revised: 21 February 2014 / Accepted: 27 February 2014 / Published: 12 March 2014
(This article belongs to the Special Issue Polymers from Biomass)


With the impending fossil fuel crisis, the search for and development of alternative chemical/material substitutes is pivotal in reducing mankind’s dependency on fossil resources. One of the potential substitute candidates is polyhydroxyalkanoate (PHA). PHA is a carbon-neutral and valuable polymer that could be produced from many renewable carbon sources by microorganisms, making it a sustainable and environmental-friendly material. At present, PHA is not cost competitive compared to fossil-derived products. Encouraging and intensifying research work on PHA is anticipated to enhance its economic viability in the future. The development of various biomolecular and chemical techniques for PHA analysis has led to the identification of many PHA-producing microbial strains, some of which are deposited in culture collections. Research work on PHA could be rapidly initiated with these ready-to-use techniques and microbial strains. This review aims to facilitate the start-up of PHA research by providing a summary of commercially available PHA-accumulating microbial cultures, PHA biosynthetic pathways, and methods for PHA detection, extraction and analysis.

1. Introduction

PHA is a family of naturally-occurring biopolyesters synthesized by various microorganisms. First discovered by Lemogine in 1926 [1], PHA has since attracted much commercial and research interests due to its biodegradability, biocompatibility, chemical-diversity, and its manufacture from renewable carbon resources [2]. A PHA molecule is typically made up of 600 to 35,000 (R)-hydroxy fatty acid monomer units [3]. Each monomer unit harbors a side chain R group which is usually a saturated alkyl group (Figure 1) but can also take the form of unsaturated alkyl groups, branched alkyl groups, and substituted alkyl groups although these forms are less common [4]. Depending on the total number of carbon atoms within a PHA monomer, PHA can be classified as either short-chain length PHA (scl-PHA; 3 to 5 carbon atoms), medium-chain length PHA (mcl-PHA; 6 to 14 carbon atoms), or long-chain length PHA (lcl-PHA; 15 or more carbon atoms) [3]. About 150 different PHA monomers have been identified and this number continues to increase with the introduction of new types of PHA through the chemical or physical modification of naturally-occurring PHA [5], or through the creation of genetically-modified organisms (GMOs) to produce PHA with specialized functional groups [6]. These features gave rise to diverse PHA properties which can be tailored for various applications ranging from biodegradable packaging materials to medical products. PHA is also considered as pharmaceutically-active compound and currently investigated as potential anti-HIV drugs, anti-cancer drugs, antibiotics, etc. [7,8]. The production of various types of PHA material, their properties and downstream applications was recently reviewed by Philip et al. [9], Olivera et al. [10], Chen [11], and Rai et al. [7].
The intense research and commercial interest in PHA is evident from the rapid increment in PHA-related publications. Web of Science citation report (Thomson Reuters, New York, NY, USA) revealed that in the last 20 years, PHA-related documents have increased by almost 10-fold while citations have increased by more than 500-fold with an average citation count of about 1100 citations per year. This has fuelled the growth of knowledge and development of techniques related to microbial PHA production. With this ready information, research work on PHA could be rapidly initiated either through using microbial strains previously deposited in culture collections or through isolating and characterizing novel PHA-producing microbes. Figure 2 is an illustration of the typical workflow processes for PHA research. This review aims to facilitate the start-up of PHA research by providing an overview of PHA-accumulating microbes currently available in culture collections, PHA biosynthetic pathways, techniques for microbial PHA detection and characterization, PHA polymer extraction, and polymer characterization.
Figure 1. Polyhydroxyalkanoate (PHA) chemical structure. The nonmenclature and carbon number for PHA compounds is determined by the functional alkyl R group. Asterisk denotes chiral center of PHA-building block.
Figure 1. Polyhydroxyalkanoate (PHA) chemical structure. The nonmenclature and carbon number for PHA compounds is determined by the functional alkyl R group. Asterisk denotes chiral center of PHA-building block.
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Figure 2. Schematic workflow processes for PHA research.
Figure 2. Schematic workflow processes for PHA research.
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2. PHA Biosynthetic Pathways

PHA plays a pivotal role in priming microorganisms for stress survival. PHA promotes the long-term survival of bacteria under nutrients-scarce conditions by acting as carbon and energy reserves for both non-sporulating and sporulating bacteria. Additionally, bacteria that harbor PHA showed enhanced stress tolerance against transient environmental assaults such as ultraviolet (UV) irradiation, heat and osmotic shock [12]. PHA biosynthetic pathways are intricately linked with the bacterium’s central metabolic pathways including glycolysis, Krebs Cycle, β-oxidation, de novo fatty acids synthesis, amino acid catabolism, Calvin Cycle, and serine pathway [4,13,14,15,16,17]. Many common intermediates are also shared between PHA and these metabolic pathways, most notably being acetyl-CoA. In some PHA-producing microbes such as Cupriavidus necator, Chromatium vinosum, and Pseudomonas aeruginosa, the metabolic flux from acetyl-CoA to PHA is greatly-dependent on nutrient conditions [18]. Under nutrient-rich conditions, the production of high amounts of coenzyme A from Krebs Cycle blocks PHA synthesis by inhibiting 3-ketothiolase (PhaA) such that acetyl-CoA is channeled into the Krebs Cycle for energy production and cell growth [19] (Figure 3). Conversely, under unbalanced nutrient conditions (i.e., when an essential nutrient such as nitrogen and phosphorus is limiting in the presence of excess carbon), coenzyme A levels are non-inhibitory allowing acetyl-CoA to be directed towards PHA synthetic pathways for PHA accumulation [19,20]. This metabolic regulation strategy in turn enables the PHA-accumulating microbes to maximize nutrient resources in their adaptation to environmental conditions.
To date, much insight has been gained on metabolic pathways for scl-PHA and mcl-PHA synthesis through studies using wild-type strains and heterologous expressions in recombinant strains [3]. In-depth reviews on these various PHA biosynthesis pathways and the enzymes involved have been provided by Chen [11], Lu et al. [4], Madison and Huisman [13], and Khosravi-Darani et al. [21]. Figure 3 shows the various routes of scl-PHA synthesis (pathways A to J) and mcl-PHA synthesis (pathways J to M) while Table 1 provides a summary of the enzymes involved. Although the biosynthesis of PHA from (R)-hydroxyalkyl-CoA ([R]-3-HA-CoA) precursors were most commonly reported, the diversity of PHA precursors is not restricted to (R)-3-HA-CoA alone [4]. Putative metabolic routes, such as pathways L and M (Figure 3), were recently proposed to expound for the metabolism of cyclohexanol to 6-hydroxyhexanoyl-CoA and 4,5-alkanolactone to 4,5-hydroxyacyl-CoA (4,5-HA-CoA) [11]. Nevertheless, the current knowledge on biosynthetic pathways is largely confined to (R)-3-HA-CoA precursors and falls short of accounting for the chemically-diverse PHA monomers and PHA monomers of lcl-PHA. There remains much about biosynthetic pathways waiting to be uncovered. Further studies to verify putative pathways as well as unraveling new biosynthetic pathways are anticipated to facilitate the creation of PHA materials that could be tailored for specific application needs.
Figure 3. PHA biosynthetic pathways. Dotted lines represent putative pathways. Numbers represent enzymes involved in the chemical reactions that are summarized in Table 1. (3-H2MB-CoA, 3-hydroxy-2-methylbutyryl-CoA; 3-H4MV-CoA, 3-hydroxy-4-methylvaleryl-CoA; 3-HV-CoA, 3-hydroxyvaleryl-CoA; 3-H2MV-CoA, 3-hydroxy-2-methylvaleryl-CoA; 4-HB-CoA, 4-hydroxybutyryl-CoA; 3-HB-CoA, 3-hydroxybutyryl-CoA; (R)-3-HA-CoA, (R)-3-hydroxyacyl-CoA; 4,5-HA-CoA, 4,5-hydroxyacyl-CoA).
Figure 3. PHA biosynthetic pathways. Dotted lines represent putative pathways. Numbers represent enzymes involved in the chemical reactions that are summarized in Table 1. (3-H2MB-CoA, 3-hydroxy-2-methylbutyryl-CoA; 3-H4MV-CoA, 3-hydroxy-4-methylvaleryl-CoA; 3-HV-CoA, 3-hydroxyvaleryl-CoA; 3-H2MV-CoA, 3-hydroxy-2-methylvaleryl-CoA; 4-HB-CoA, 4-hydroxybutyryl-CoA; 3-HB-CoA, 3-hydroxybutyryl-CoA; (R)-3-HA-CoA, (R)-3-hydroxyacyl-CoA; 4,5-HA-CoA, 4,5-hydroxyacyl-CoA).
Polymers 06 00706 g003
Table 1. Enzymes involved in PHA biosynthesis pathways.
Table 1. Enzymes involved in PHA biosynthesis pathways.
1Glyceraldehyde-3-phosphate dehydrogenase-Cupriavidus necator[22]
2Pyruvate dehydrogenase complex-Cupriavidus necator and Burkholderia cepacia[22]
33-KetothiolasePhaACupriavidus necator[23]
4NADPH-dependent acetoacetyl-CoA reductasePhaBCupriavidus necator[23]
5PHA synthasePhaCCupriavidus necator and various[12,23]
6Acetyl-CoA carboxylaseACCEscherichia coli K-12 MG1655[24]
7Malonyl-CoA:ACP transacylaseFabDEscherichia coli K-12 MG1655[24]
83-Ketoacyl carrier protein synthaseFabHEscherichia coli K-12 MG1655[24,25]
9NADPH-dependent 3-Ketoacyl reductaseFabGPseudomonas aeruginosa[26]
10Succinic semialdehyde dehydrogenaseSucDClostridium kluyveri[27]
114-Hydroxybutyrate dehydrogenase4HbDClostridium kluyveri[27]
124-Hydroxybutyrate-CoA:CoA transferaseOrfZClostridium kluyveri[27]
13Alcohol dehydrogenase, putative-Aeromonas hydrophila 4AK4[28]
14Hydroxyacyl-CoA synthase, putative-Mutants and recombinants of Cupriavidus necator[29]
15Methylmalonyl-CoA mutaseSbmEscherichia coli W3110[30]
16Methylmalonyl-CoA racemase-Nocardia corallina[31]
17Methylmalonyl-CoA decarboxylaseYgfGEscherichia coli W3110[30]
18Ketothiolase, putative--[32]
193-KetothiolaseBktBCupriavidus necator[33]
20Ketothiolase, putative--[32]
21NADPH-dependent acetoacetyl-CoA reductase-Rhizobium (Cicer) sp. CC 1192[34]
22Acyl-CoA synthetaseFadDPseudomonas putida CA-3 and Escherichia coli MG1655[35,36]
23Acyl-CoA oxidase, putative--[37]
24Enoyl-CoA hydratase I, putative--[37]
25(R)-Enoyl-CoA hydratasePhaJPseudomonas putida KT2440[38]
273-Ketoacyl-CoA thiolaseFadAPseudomonas putida KT2442[39]
283-Hydroxyacyl-ACP:CoA transacylasePhaGPseudomonas mendocina[40]
29Cyclohexanol dehydrogenaseChnAAcinetobacter sp. SE19 and Brevibacterium epidermidis HCU[41]
30Cyclohexanone monooxygenasesChnBAcinetobacter sp. SE19 and Brevibacterium epidermidis HCU[41]
31Caprolactone hydrolaseChnCAcinetobacter sp. SE19 and Brevibacterium epidermidis HCU[41]
326-Hydroxyhexanoate dehydrogenaseChnDAcinetobacter sp. SE19 and Brevibacterium epidermidis HCU[41]
336-Oxohexanoate dehydrogenaseChnEAcinetobacter sp. SE19 and Brevibacterium epidermidis HCU[41]
34Semialdehyde dehydrogenase, putative--[11]
356-hydroxyhexanoate dehydrogenase, putative--[11]
36Hydroxyacyl-CoA synthase, putative--[11]
37Lactonase, putative-Mutants and recombinants of Cupriavidus necator[29]

3. PHA-Producing Microbial Strains from Culture Collections

The PHA bioaccumulation trait is widespread among the bacterial and archaeal domains with PHA-producing microbes occurring in more than 70 bacterial and archaeal genera [4,42]. Bioaccumulated PHA is stored in the form of intracellular lipid granules in these microbes [43]. Acting as biocatalysts, these PHA-producing microorganisms enable the coupling of a myriad of carbon catabolic pathways together with PHA anabolic pathways, thereby playing a key role in the diversification of PHA production from various carbon sources. These carbon sources include saccharides (e.g., fructose, maltose, lactose, xylose, arabinose, etc.), n-alkanes (e.g., hexane, octane, dodecane, etc.), n-alkanoic acids (e.g., acetic acid, propionoic acid, butyric acids, valeric acid, lauric acid, oleic acid, etc.), n-alcohols (e.g., methanol, ethanol, octanol, glycerol, etc.), and gases (e.g., methane and carbon dioxide) [1,44]. Wastestreams, which provide a free source of carbons, have also been identified for PHA production [45]. These include waste frying oil, vinegar waste, waste fats, food waste, agricultural waste, domestic wastewater, plant oil mill effluents, crude glycerol from biodiesel production, plastic waste, landfill gas, etc. The deposition of some PHA-producing microbial strains in culture collections has made these strains commercially available. Microbial strains from culture collections are generally well-documented in terms of their genetics and biochemistry underlying carbon assimilation and PHA accumulation. With this knowledge, it enables the appropriate microbes to be selected according to the targeted carbon source, facilitating the rapid start-up of PHA-related research and/or industrial production. Table 2 provides a summary of carbon substrate utilization and PHA production by deposited bacterial and archaeal strains.
Table 2. PHA-producing microbial strains available in culture collections.
Table 2. PHA-producing microbial strains available in culture collections.
MicroorganismCulture collection number bCarbon sourcePHA monomer or polymer cPHA content
Average PHA productivity
(g L−1 h−1)
Gram-negative bacteria
Azohydromonas australica
(formerly Alcaligenes latus)
ATCC 29713,
DSM 1124, IAM 12664,
LMG 3324
Malt wasteP3HB70.10.445[46]
Azohydromonas lata
(formerly Alcaligenes latus)
ATCC 29714,
DSM 1123,
IAM 12665,
LMG 3325
Fructose, glucoseP3HB76.5–79.40.121–0.128[50]
Azotobacter beijerinckiiDSM 1041,
NCIB 11292
Burkholderia cepacia
(formerly Pseudomonas multivorans and Pseudomonas cepacia)
ATCC 17759,
DSM 50181,
NCIB 9085
Fructose, glucose, sucroseP3HB50.4–59.0NG[50]
Burkholderia sp. USMJCM 15050Lauric acid, myristic acid, oleic acid, palmitic acid, stearic acidP3HB1.0–69.0NG[54]
Caulobacter vibrioides
(formerly Caulobacter crescentus)
DSM 4727GlucoseP3HB18.30.008[55]
Cupriavidus necator H16
(formerly Hydrogenomonas eutropha H16, Alcaligenes eutrophus H16, Ralstonia eutropha H16 and Wautersia eutropha H16)
ATCC 17699,
DSM 428,
KCTC 22496,
NCIB 10442
Fructose, glucoseP3HB67.0–70.50.052–0.067[50]
4-Hydroxyhexanoic acidP3HB76.3–78.5NG[56]
Corn oil, oleic acid, olive oil, palm oilP3HB79.0–82.00.041–0.047[57]
Acetate, butyrate, lactic acid,
propionic acid
3HB, 3HV3.9–40.70.001–0.037[58]
Cupriavidus necator
(formerly Hydrogenomonas eutropha, Alcaligenes eutrophus N9A, Ralstonia eutropha N9A and Wautersia eutropha)
DSM 5184-Hydroxyhexanoic acidP3HB65.8–66.2NG[56]
Cupriavidus necator
(formerly Hydrogenomonas eutropha, Alcaligenes eutrophus TF93, Ralstonia eutropha TF93 and Wautersia eutropha)
ATCC 17697, DSM 5314-Hydroxyhexanoic acidP3HB67.2NG[56]
Cupriavidus necator a
(formerly Hydrogenomonas eutropha, Alcaligenes eutrophus, Ralstonia eutropha and Wautersia eutropha)
CECT 4623, KCTC 2649, NCIMB 11599GlucoseP3HB76.02.420[61]
Potato starch, saccharified wasteP3HB46.01.470[62]
Cupriavidus necator
(formerly Hydrogenomonas eutropha, Alcaligenes eutrophus, Ralstonia eutropha and Wautersia eutropha)
DSM 545MolassesP3HB31.0–44.00.080–0.120[63]
Glucose, propionic acidP3HB3HV80.00.820[64]
Waste glycerolP3HB14.8–36.10.330–4.200[65]
Halomonas boliviensis LC1ATCC BAA-759,
DSM 15516
Hydrolyzed starchP3HB56.0NG[66]
Hydrogenophaga pseudoflavaATCC 33668, DSM 1034Lactose, sucroseP3HB3HV20.2–62.50.018–0.117[67]
Hydrolyzed whey and valerateP3HB3HV40.00.050[68]
Methylobacterium extorquensATCC 55366MethanolP3HB40.0–46.00.250–0.600[69]
Methylobacterium extorquensATCC 8457, DSM 1340, NCIB 2879, NCTC 2879MethanolP3HB35.0–62.30.183–0.980[70,71]
Methylocystis sp. GB25 aDSM 7674MethaneP3HB51.0NG[72]
Novosphingobium nitrogenifigens Y88DSM 19370, ICMP 16470GlucoseP3HB81.00.014–0.021[73]
Paracoccus denitrificansATCC 17741, DSM 413n-PentanolP3HV22.0–24.0NG[74]
Pseudomonas aeruginosaNCIM 2948Cane molasses, fructose, glucose, glycerol, sucroseP3HB12.4–62.00.012–0.110[75]
Pseudomonas aeruginosa PAO1ATCC 47085Oil and wax products from polyethylene (PE) pyrolysismcl-PHA25.0NG[76]
Pseudomonas frederiksbergensis GO23 aNCIMB 41539Terephthalic acid from polyethylene
terephthalate (PET) pyrolysis
Pseudomonas marginalisDSM 502761,3-butanediol, octanoatescl-mcl-PHA,
Pseudomonas mendocinaATCC 25411,
DSM 50017
1,3-butanediol, octanoatescl-mcl-PHA13.5–19.3NG[78]
Pseudomonas oleovoransATCC 8062, DSM 10454-Hydroxyhexanoic acidscl-mcl-PHA18.6NG[56]
Pseudomonas putida CA-3 aNCIMB 41162Styrenemcl-PHA31.80.063[79]
Styrene from polystyrene (PS) pyrolysismcl-PHA36.40.033[80]
Pseudomonas putida GO16 aNCIMB 41538Terephthalic acid from polyethylene
terephthalate (PET) pyrolysis
mcl-PHA27.0~0.005, 0.008 d[77]
Pseudomonas putida GO19 aNCIMB 41537Terephthalic acid from polyethylene
terephthalate (PET) pyrolysis
mcl-PHA23.0~0.005, 0.008 d[77]
Pseudomonas putida GPo1
(formerly Pseudomonas oleovorans)
ATCC 29347Alkenes, n-alkanesmcl-PHA2.0–28.0NG[81]
Pseudomonas putida KT2440ATCC 47054Nonanoic acidmcl-PHA26.8–75.40.250–1.110[84]
4-Hydroxyhexanoic acidmcl-PHA25.3–29.8NG[56]
Pseudomonas putida F1ATCC 700007, DSM 6899Benzene, ethylbenzene, toluenemcl-PHA1.0–22.0NG[86]
Pseudomonas putida mt-2NCIMB 10432Toluene, p-xylenemcl-PHA22.0–26.0NG[86]
Acetic acid, citric acid, glucose, glycerol, octanoic acid, pentanoic acid, succinic acidmcl-PHA4.0–77.0NG[87]
Thermus thermophilus HB8ATCC 27634, DSM 579Wheyscl-mcl-PHA35.60.024[88]
Gram-Positive bacteria
Bacillus megateriumDSM 90Citric acid, glucose, glycerol, succinic acidP3HB9.0–50.0NG[87]
Bacillus megateriumCCM 1464,
DSM 509,
IFO 12109, NBRC 12109
Citric acid, glucose, glycerol, succinic acid, octanoic acidP3HB, scl-mcl-PHA, mcl-PHA3.0–48.0NG[87]
Various Bacillus spp. type strainsRefer to [89]Acetate, n-alkanoate,
3-Hydroxybutyrate, propionate, sucrose, valerate
3HB, 3HV, 3HHx2.2–47.6NG[89]
Corynebacterium glutamicumATCC 15990,
DSM 20137, NCIB 10337
Acetic acid, citric acid, glucose, glycerol, succinic acidP3HB, mcl-PHA4.0–32.0NG[87]
Corynebacterium hydrocarboxydansATCC 21767Acetate, glucose3HB, 3HV8.0–21.0NG[90]
Microlunatus phosphovorusDSM 10555,
JCM 9379
Glucose3HB, 3HV20.0–30.0NG[91]
Nocardia lucidaNCIMB 10980Acetate, succinate3HB, 3HV7.0–20.0NG[90]
Rhodococcus sp. aNCIMB 40126Acetate, 2-alkenoate,
5-chlorovalerate, fructose, glucose, hexanoate,
4-Hydroxybuytrate, lactate, molasses, succinate, valerate
Various Streptomyces spp. type cultureRefer to [89]GlucoseP3HB1.2–82.0NG[89]
Haloferax mediterraneiATCC 33500,
CCM 3361,
DSM 1411
Hydrolyzed wheyP3HB3HV72.80.090[93]
Glycerol and crude glycerol from biodiesel productionP3HB3HV75.0–76.00.120[94]
Various archaeal strainsRefer to [95]Fructose, glucose, glycerolP3HB, P3HB3HV0.8–22.9<0.001–0.021[95]
a Refers to patent strain; b ATCC, American Type Culture Collection (Manassas, VA, USA); CCM, Czech Collection of Microorganisms (Masaryk University, Brno, Czech Republic); CECT, Colección Española de Cultivos Tipo (Universidad de València, Edificio de Investigación, Burjassot, Spain); DSM, German Collection of Microorganisms and Cell Cultures GmbH (Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, Braunschweig, Germany); IAM, Institute of Molecular and Cellular Biosciences (The University of Tokyo, Japan [collection transferred to JCM]); ICMP, International Collection of Microorganisms from Plants (Plant Diseases Division, DSIR, Auckland, New Zealand [also known as PDDCC]); IFO, Institute for Fermentation, Osaka (Yodogawa-ku, Osaka, Japan [collection transferred to NBRC]); JCM, Japan Collection of Microorganisms (RIKEN BioResource Center, Tsukuba, Ibaraki, Japan); KCTC, Korean Culture Center of Microorganisms (Department of Food Engineering, Yonsei University, Seoul, Republic of Korea); LMG, Laboratorium voor Microbiologie, Universiteit Gent (Gent, Belgium); NBRC, NITE Biological Resource Center (Department of Biotechnology, National Institute of Technology and Evaluation, Kisarazu, Chiba, Japan); NCIB, National Collection of Industrial Bacteria (Torry Research Station, Aberdeen, Scotland, UK [incorporated with NCIMB]); NCIM, National Collection of Industrial Microorganisms (National Chemical Laboratory, India); NCIMB, National Collections of Industrial Food and Marine Bacteria (Aberdeen, Scotland, UK); NCTC, National Collection of Type Cultures (Public Health England, UK); c PHA is indicated as monomers unless nuclear magnetic resonance (NMR) verification was performed; d Refers to the maximal PHA productivity rate reported; CDM: cell dry mass; NG: not given.

3.1. Gram-Negative Bacteria

To date, most PHA-producing bacteria were found to be Gram-negative bacteria [4]. Scl-PHA is usually synthesized in high amounts by bacterial species from Azohydromonas, Burkholderia, and Cupriavidus. A. lata (ATCC 29714) is reportedly capable of producing between 50% and 88% cell dry mass (%CDM) of poly(3-hydroxybutyrate) (P3HB) from various sugars including glucose, fructose and sucrose [47,48,49,50] while Burkholderia sp. USM (JCM 15050) could synthesize up to 69 %CDM of P3HB from fatty acids [54]. C. necator is a hydrogen-oxidizing (Knallgas) bacterium which could produce PHA by fixing CO2 through Calvin Cycle [21]. The well-characterized C. necator H16 (ATCC 17699) could shift between heterotrophic and autotrophic mode for growth and PHA production. Emerging evidence also suggested that both heterotrophic and autotrophic PHA biosynthesis can occur concurrently in this bacterium [16]. The unique physiology of C. necator H16 (ATCC 17699) meant that it was able to utilize chemically-diverse carbon substrates such as CO2, sugars (i.e., glucose and fructose), n-alkanoic acids (i.e., 4-hydroxyhexanoic acid), vegetable oils (i.e., olive oil, corn oil, and palm oil) for P3HB accumulation in the range of 67 to 88.9 %CDM [50,56,57,59] (Table 2). Owing to the high scl-PHA production capacity, wild-type and mutant bacterial species from the aforementioned genera are widely employed in industrial PHA production. Recent updates on the application of Gram-negative bacteria in industrial-scale PHA production could be found in review articles by Chen [96] and Chanprateep [97].
Scl-PHA was also produced by Gram-negative methylotrophs. Methylobacterium extorquens (ATCC 55366) and Paracoccus denitrificans (ATCC 17741) produced up to 46 %CDM P3HB from methanol and up to 24 %CDM poly(3-hydroxyvalerate) (P3HV) from n-pentanol, respectively [69,74]. Due to the cheaper cost of methanol compared to pure sugar substrates, the use of methylotrophs for industrial scl-PHA production could reduce PHA cost. However, further studies would be required to enhance the PHA content and PHA productivity of methylotrophs before they can be considered as attractive inoculum alternatives for industrial scl-PHA production [3].
The production of mcl-PHA is often reported in Pseudomonas sp., and usually occurs at PHA contents between about 1 and 30 %CDM (Table 2). Higher mcl-PHA contents have also been observed in P. putida mt-2 (NCIMB 10432), which could produce up to 77 %CDM mcl-PHA from octanoic acid [87]; and P. putida KT2440 (ATCC 47054), a mutant of P. putida mt-2 lacking the TOL plasmid, which could produce up to 75.4 %CDM using nonanoic acid [84]. Aside from mcl-PHA, some Pseudomonad species have also been reported to synthesize scl-mcl-PHA copolyesters. These species include P. marginalis (DSM 50276), P. mendocina (ATCC 25411), P. putida GPo1 (ATCC 29347), and P. oleovorans (ATCC 8062) when n-alkanoates and 1,3-butanediol were provided as carbon sources [56,78,82]. For most of these Pseudomonad species, 3-hydroxybutyrate (3HB) is typically incorporated as a minor constituent in scl-mcl-PHA (between less than 1 mol% and 7.8 mol%) [78,82]. An exception is P. oleovorans (ATCC 8062) where its cultivation on 4-hydroxyhexanoic acid resulted in a copolymer predominated by 3HB (92.4 mol%) [56]. Pseudomonads are also well-known for their bioremediation properties including the biodegradation recalcitrant and/or toxic aromatic carbon substrates [98], and have been successfully applied in the treatment of contaminated effluents, exhaust gas and soils [99,100,101]. Recent studies demonstrated that aromatic-degraders P. putida F1 (DSM 6899), P. putida mt-2 (NCIMB 10432), and P. putida CA-3 (NCIMB 41162) could bioconvert toxic pollutants benzene, toluene, ethylbenzene, xylene (BTEX) and styrene to mcl-PHA [86]; P. putida CA-3 (NCIMB 41162) and other Pseudomonads including P. aeruginosa PAO1 (ATCC 47085), P. frederiksbergensis GO23 (NCIMB 41539), P. putida GO16 (NCIMB 41538) and P. putida GO19 (NCIMB 41537) could utilize crude pyrolysis products from various plastics (i.e., polystyrene [PS], polyethylene [PE] and polyethylene terephthalate [PET]) for mcl-PHA production [76,77,80], which offers the potential benefit to off-set waste treatment cost through PHA recovery.
PHA accumulation has been observed in Gram-negative extremophilic bacteria as well. These bacteria accumulate PHA under unique cultivation conditions with either high salinity or elevated temperatures. The halophilic Halomonas boliviensis LC1 (DSM 15516) could grow and produce 56 %CDM of scl-PHA P3HB from starch hydrolysate under moderately saline conditions (0.77 M NaCl) [66] while the thermophilic Thermus thermophilus HB8 (ATCC 27634) synthesized up to 35.6 %CDM of scl-mcl-PHA copolymer from whey at a high cultivation temperature of 70 °C [88]. Compared to other Gram-negative bacteria, extremophiles are advantageous in terms of their lower sterility demand as well as their potential for direct application with waste effluents originally high in salt concentrations or temperatures, eliminating the need and cost involved for pre-treatment of waste effluents.
The main concern with Gram-negative bacteria however, is the presence of lipopolysaccharide (LPS) endotoxins in the bacteria’s outer cell membrane, which may co-purify with crude PHA polymer during the extraction process [7]. LPS endotoxin is a pyrogen which can elicit a strong inflammatory response [102], rendering the PHA polymer unsuitable for biomedical applications. Removal of LPS endotoxin can be achieved through the treatment of PHA polymer with oxidizing agents (i.e., sodium hypochlorite and NaOH, ozone, hydrogen peroxide, and benzoyl peroxide), with repeated solvent extractions, or with solvent extraction followed by purification with activated charcoal [7,103,104]. These methods however, increase the overall cost of PHA production and lead to changes in PHA polymer properties (i.e., reduction in molecular mass and polydispersity).

3.2. Gram-Positive Bacteria

PHA production in Gram-positive bacteria has been reported in genera Bacillus, Caryophanon, Clostridium, Corynebacterium, Micrococcus, Microlunatus, Microcystis, Nocardia, Rhodococcus, Staphylococcus, Streptomyces [4]. Compared to Gram-negative bacteria, Gram-positive bacteria were mostly found to produce scl-PHA [89] and at lower PHA contents between about 2 and 50 %CDM (Table 2), which is why Gram-positive bacteria have yet to be adopted for commercial PHA production. A high scl-PHA content of 82 %CDM has been previously reported for Streptomyces sp. (ATCC 1238) growing on glucose, but the value may be an overestimation by crotonic acid assay [89,105]. Some microbial strains are able to synthesize mcl-PHA or scl-mcl-PHA copolymers if suitable carbon substrates and conditions are provided. A study by Shahid et al. [87] demonstrated that B. megaterium (DSM 509) formed exclusively P3HB from glycerol and succinic acid in a mineral medium supplemented with nitrogen, but started to synthesize scl-mcl-PHA upon sub-culturing to the same medium and in the absence of nitrogen. In the same bacterium, the formation of exclusively mcl-PHA (48 %CDM) was observed when it was cultured on octanoic acid in the absence of nitrogen [87].
Despite generally accumulating lower amounts of PHA, Gram-positive bacteria are advantageous over Gram-negative bacteria owing to their lack of LPS which may make them a better source of PHA raw material for biomedical applications [89]. However, some Gram-positive bacteria are known to produce lipidated macroamphiphiles including lipoglycans and lipoteichoic acids (LTA), which have immunogenic properties similar to LPS [102]. The bacterial genera Corynebacterium, Nocardia, Rhodococcus reportedly produce lipoglycans [106,107,108]; and while LTA production occurs in the genera Bacillus, Clostridium, and Staphylococcus [109,110], some PHA-producing strains from these genera lack LTA [111,112]. Further investigation will be required to verify if there are alternative lipidated macroamphiphiles in Gram-positive and PHA-producing bacteria. At present, the immunogenic effects of lipidated macroamphiphiles in PHA remain unknown. Future in vitro or in vivo evaluation studies would be imperative to evaluate the suitability of PHA, derived from Gram-positive bacteria, for biomedical applications [89].

3.3. Archaea

PHA is also found in archaea but to date however, its discovery has been limited to haloarchaeal species, specifically the genera Haloferax, Halalkalicoccus, Haloarcula, Halobacterium, Halobiforma, Halococcus, Halopiger, Haloquadratum, Halorhabdus, Halorubrum, Halostagnicola, Haloterrigena, Natrialba, Natrinema, Natronobacterium, Natronococcus, Natronomonas, and Natronorubrum [95]. Haloarchaea are the extremely halophilic members of the archaea domain, which require high salt concentrations for normal enzyme activity, growing at saturation conditions of up to 6 M NaCl [113]. Haloarchaea have been reported to synthesize PHA from glucose, volatile fatty acids and more complex carbon sources such as starch, whey hydrolysate, vinasse and crude glycerol from biodiesel production [92,93,94,95]. The type of PHA synthesized appeared to be exclusively scl-PHA homopolymer containing either 3HB or 3HV monomers, and/or scl-PHA heteropolymer containing both 3HB and 3HV monomers [42,95]. Many PHA-producing haloarchaeal cultures are currently available from culture collections but most of them produce PHA at low cellular contents between 0.8 to 22.9 %CDM (Table 2) [95].
At present, the best PHA producer is Haloferax mediterranei (DSM 1411), which requires 2 to 5 M NaCl for growth and can accumulate high PHA levels between 50 and 76 %CDM [92,93,94]. H. mediterranei (DSM 1411) could be an attractive candidate for PHA production as the hypersaline conditions, required for its growth and PHA cultivation, meant that that very few contaminating organisms can survive thereby reducing the sterility requirements and its associated cost (i.e., process piping, instrumentation and insulation, electricity for steam generation, etc.) [92]. However, when compared to moderately halophilic bacteria such as H. boliviensis LC1 (DSM 15516), the extreme salinity required by haloarchaea can be a bane to PHA production as the high salt concentration incur higher chemical cost and accelerates the corrosion of stainless steel fermentors [66]. Nevertheless, haloarchaea are advantageous over halophilic bacteria in the ease of PHA recovery. PHA recovery from halophilic bacteria typically requires the use of chemical, enzymatic or mechanical method for cell wall disruption to release intracellular PHA granules, and these methods could account for up to 50% or more of the overall PHA production cost [114]. Extraction solvents such as chloroform and acetone also posed potential environmental hazards if their utilization and disposal are mismanaged. Conversely, haloarchaea undergo cell lysis in distilled water and release PHA granules that can be recovered by low speed centrifugation [42]. This makes PHA recovery from haloarchaea a relatively easy, less chemical- and energy-intensive process, which translates into lower extraction cost, and has lower ecological footprint.

3.4. Formulation of Defined Co-Cultures Using Deposited Microbial Strains

It has been estimated that at least 30% or more of PHA cost is attributed to carbon, nutrients and aeration cost [115]. This has prompted intensive research to diversify PHA production from cheaper carbon sources and waste carbon as a means to lower PHA cost [45]. Using waste resources for PHA production is particularly challenging as wastestream is often a complicated mixture of carbon substrates, some of which cannot be assimilated for PHA production or are inhibitory to a single microbial culture. Defined co-culture on the other hand, is a culture involving two or more microbes, and has been successfully applied to bioconvert more complex carbon feed into PHA [86,116]. The commercial availability of culture collection microbes enables the rapid formulation of defined co-culture using deposited microbial strains in accordance to the characteristics of the carbon feedstock for PHA production.
Co-culturing of microbial strains, with different carbon utilization, enables the synergistic bioconversion of carbon substrate mixture into PHA. This was exemplified in the bioconversion of synthetic plastic pyrolysis oil, containing toxic compounds BTEX and styrene, into PHA by a co-culture system consisting of three P. putida strains (i.e., strains F1, mt-2 and CA-3) [86]. On their own, P. putida F1 (DSM 6899) could metabolize BTE to form mcl-PHA while P. putida mt-2 (NCIMB 10432) and P. putida CA-3 (NCIMB 41162) could do so with TX and styrene, respectively. Monoculture cultivations using single assimilable monoaromatic substrate led to bacterial growth of between 0.3 and 0.8 g L−1 and cellular PHA yields of between 0.048 and 0.26 g L−1 while no bacterial growth was observed for non-assimilable monoaromatic substrate. As a co-culture growing on BTEX and styrene mixture however, higher biomass yield (1 g L−1) was attained and overall PHA yield (0.25 g L−1) was equivalent to that observed in the best-performing monoculture. This suggests that under co-culture conditions, the cooperative metabolism amongst bacterial members not only maximizes the utilization of various carbon substrates for PHA production but may also enhance the robustness of individual strains through the removal of inhibitory compounds, providing co-culture an advantage over monoculture systems.
Co-culturing of microbial strains can be applied to expand the repertoire of carbon substrates for PHA formation by using one microorganism to convert carbon substrate into a metabolite which can be efficiently consumed by a second microorganism for PHA production. This was demonstrated for co-cultures of C. necator H16 (ATCC 17699) with lactate-producing bacteria. While C. necator H16 (ATCC 17699) is capable of high PHA accumulation, it can only readily metabolize and accumulate PHA from organic acids such as acetate, butyrate and lactate, and was unable to do so for common sugars such as glucose and xylose [117]. To overcome this problem, Lactobacillus delbrueckii (IAM 1928) and Lactococcus lactis IO-1 (JCM 7638) were used to convert glucose and xylose, respectively, into lactate which can be easily converted to P3HB by C. necator H16 (ATCC 17699) [116,118]. While a two-stage fermentation system was used for PHA production from xylose-derived lactate [118], the feasibility of using single-stage fermentation system has also been demonstrated by Ganduri et al. [116] for PHA production from glucose-derived lactate, achieving up to 36.6 g L−1 of P3HB and displaying notably higher PHA productivity over monoculture system [50].
Co-culture system may also have the potential to mitigate biogas, produced from anaerobic digesters or landfills, while achieving aerobic PHA production without aeration supply. In a proof-of-concept study by van der Ha et al. [119], a gas mixture of 60% CH4 and 40% CO2 was photosynthetically fixed by an algal Scenedesmus sp. monoculture to produce 60% CH4 and close to 40% O2. The resultant gas components O2 and CH4 provided the aerobic condition and carbon substrate, respectively, required for P3HB accumulation by a second monoculture of methane-oxidizing bacteria Methylocystis parvus (NCIMB 11129). Under three cycles of feast-and-famine regime, cellular PHA content in M. parvus (NCIMB 11129) reached a maximum of 29.5 %CDM with 243 mg P3HB produced for every 1 g of CH4–C consumed. Co-culturing of Scenedesmus sp. and M. parvus (NCIMB 11129) led to a conversion of 98% of CH4–C and CO2–C as algal and bacterial biomass but the PHA yield was not reported. Hence, the efficacy of the co-culture system for simultaneous biogas treatment and PHA production remains to be verified.
At present, the application of defined co-cultures for PHA production is still in its infancy. While the existing knowledge and commercial availability of deposited microbial strains facilitates the rapid formulation of defined co-cultures, there are still technical challenges that need to be circumvented. For single-stage co-culture fermentation systems, one of the main challenges is providing cultivation parameters for efficient and effective bioconversion of carbon substrates into PHA. Parameters such as inoculum concentration, dissolved oxygen, pH, temperature, cultivation time, carbon and nutrients feed rate, and secondary metabolites production rate would need to be fine-tuned in order to maximize the bioconversion process. For an example, L. delbrueckii (IAM 1928) requires anaerobic conditions for conversion of glucose into lactate while C. necator H16 (ATCC 17699) requires aerobic conditions for conversion of lactate into PHA. To overcome this problem, Ganduri et al. [116] employed an imperfectly mixed bioreactor to create non-uniform spatial distribution of dissolved oxygen, achieving 91.5% of P3HB theoretical yield (i.e., 36.6 g L−1 of P3HB) within 30 h.
Another challenge is the harvesting and separation of PHA-containing biomass from non-PHA-containing biomass, particularly for co-cultures comprising of PHA-accumulating and non-PHA-forming microorganisms as the presence of non-PHA-containing biomass would increase the extraction cost of PHA. Compared to single-stage fermentation approach, two-stage fermentation approach may be more advantageous as it enables finer control over cultivation parameters and harvesting of PHA-accumulating biomass. However, higher capital and operation cost are associated with two-stage fermentation system. Ultimately, the type of systems chosen would greatly depend on the microbial characteristics of the co-culture as well as the economic viability of the bioprocess.

4. Techniques for Detecting PHA and PHA Production Potential in Microbes

Various methods are available for the detection and analysis of intracellular microbial PHA. These methods are useful in identifying novel PHA-producing microbes or for routine monitoring of PHA production bioprocesses. Table 3 provides a summary of these methods, sample characteristics and preparation, method execution, as well as their strengths and limitations.
Table 3. Methods for detection of PHA in biomass and PHA production capacity.
Table 3. Methods for detection of PHA in biomass and PHA production capacity.
MethodCharacteristicSampleSample preparationTypical conditionsAdvantageLimitationReference
Polymerase chain reaction (PCR) gene detectionphaC gene encoding enzyme PHA synthase50–500 ng of DNA material or a single bacterial colonyDNA extraction or freeze/thaw cells to release DNA materialPCR thermal cycler temperature program for specific primer setsRequires small sample size, high sensitivity and specificity, high throughputPrimers are inadequate for detection of all phaC genes, and prone to detection errors[120,121]
Nile red and Nile blue A stainingIntracellular PHA granule structuresBacterial colonies on agar mediumAdd 0.5 µg mL−1 of Nile red or Nile blue
A to sterilized agar growth medium
Expose the agar plates to ultraviolet light (312 nm) after appropriate cultivation periodsEnables direct observation of live and actively-growing cells, requires small sample size, rapid analysis, allows differentiation between scl- and mcl-PHA under flow cytometry analysis, high throughputMethod cannot discriminate between lipids and PHAs, and is also less effective at distinguishing between PHA-negative and PHA-positive strains of Gram-positive bacteria[122,123,124,125]
Microscope slide containing heat-fixed bacterial cells smearStain slide with 1% Nile blue A at 55 °C for 10 min. Remove excess stain with tap water before staining with 8% acetic acid for 1 min. Rinse slide with tap water and blot dry with bibulous paperExamine slide with an epifluorescence microscope with an excitation wavelength of 460 nm
1 mL of cell culture with optical density at 600 nm (OD600) of 1.0 or lessAdd 2.0–10 µg mL−L of Nile red to 1 mL cell culture and incubate in the dark for 15 minEpifluorescence microscopy imaging with FITC filter with an excitation wavelength of 470–490 nm and an emission wavelength of 505 nm or fluorescence spectroscopy analysis at excitation wavelength of 488 nm and an emission wavelength of 590 nm and 575 nm for scl-PHA and mcl-PHA, respectively
Transmission electron microscopy (TEM)Intracellular PHA granule structures1–3 mL of exponential or stationary phase cell cultureCell fixation with glutaraldehyde in phosphate buffer, followed by post-fixation with osmium tetroxide. Dehydrate fixated cells through a graded acetone series before acetone-resin infiltration and resin polymerization. Cut resins into ultrathin sections (70–100 nm thickness) with an ultramicrotomeView with an accelerating voltage of 200 kV and perform imaging at magnifications of 25,000–40,000×High magnification enables direct visualization and size measurements of PHA granulesTedious sample preparation involving radioactive and hazardous chemicals, cells are killed during sample preparation[43]
Crotonic acid assayQuantitative determination of P3HB5–50 µg P3HBAdd 10 mL concentrated H2SO4, and heat at 100 °C for 10 min to form crotonic acidMeasure UV absorbance at 235 nmEasy operation, inexpensive per analysis, specific to P3HB determinationResult can be interfered by other endogenous components and matrix interferences can result in overestimation of P3HB content. Method is limited to P3HB determination[89,105,126]
Fourier transform infrared spectroscopy (FTIR)Cellular PHA content0.4-10 mg biomassSpread cells on thallium bromoiodide (KRS-5) window and air-dryFTIR was used to record the PHA spectrum at ambient temperature (25 °C), at a spectra range of 400–4000 cm−1, for 10–64 scans and a resolution of 4 cm−1Requires small sample size, short analysis time, solvent usage is optional, can provide quantitative information, enables online and real-time PHA analysis, high throughputMethod cannot discriminate between different PHA monomeric units, unable to distinguish between homogenous PHA and PHA copolymer, low sensitivity, quantification limited to scl-PHA[127,128]
Liquid chromatography (LC)PHA monomeric units0.01–500 mg biomass or 0.01–14 µg P3HBHydrolytic digestion with concentrated sulfuric acid 90 °C for 30 min, cool on ice before adding 0.014 N of sulfuric acid with rapid mixing to yield crotonic acidsHigh performance liquid chromatography (HPLC) analysis with an ion-exclusion organic acid analysis column and a UV detector at 210 nmDoes not require cell lyophilization, requires small sample size, short sample preparation time, provides both quantitative and qualitative information. Coupling with mass spectrometer (MS) detector enables tentative identification of novel PHA monomers, applicable for quantitative and qualitative analysis of mcl-PHA monomersLow separation power that is currently limited to analysis of scl-PHA monomers unless coupled to MS detector, unable to distinguish between homogenous PHA and PHA copolymer[129,130]
10–25 mg biomass or 2 mg PHAPropanolic digestion with propanol and concentrated sulfuric acid at 90 °C for
1 h to yield a mixture of
monomeric acids and propionyl esters
Ion chromatography (IC) analysis with an anion trap column and a conductivity detector
2 mg PHAReductive depolymerization by dissolution of PHA in toluene, followed by addition of lithium aluminum hydride in tetrahydrofuran (THF) with 15 min of gentle agitation at room temperature to yield 1,3-diolsHPLC-MS analysis with a C18 column
Gas chromatography (GC)PHA monomeric units5–15 mg biomass or 0.15–15 mg PHAMethanolysis with either sulfuric acid/methanol or boron trifluoride/methanol at 100 °C for 2 h–4 h to yield methyl esters or propanolysis with hydrochloric acid/propanol at 80 °C for 20 h to yield propyl estersAnalysis with a Supelco SPB-35 or DB-5 column using a flamed ionization detector (FID), or with a HP-5MS column using a MS detectorHigh separation power, high sensitivity, provides both quantitative and qualitative information, and can be applied for tentative identification of novel PHA monomers when coupled to MS detectorRequires cell lyophilization, long sample preparation time requiring the use of hazardous and volatile solvents, unable to distinguish between homogenous PHA and PHA copolymer[43,131,132,133,134]

4.1. Detection of PHA in Biomass and PHA Production Capacity

Methods which detects for PHA or PHA-producing capacity in microbes include colony/cell staining, polymerase chain reaction (PCR) gene detection, and transmission electron microscopy (TEM). Colony/cell staining and gene detection are often used as front-line methods for high throughput screening and identification of novel microbes with PHA production potential owing to the relative ease of sample preparation and short analysis time. In colony/cell staining method, Nile red or Nile blue A dye is directly added into the solid growth medium [122], liquid cell cultures [123,124] or onto heat-fixed smeared cells [125]. Under UV illumination, Nile red and Nile blue A dye stains PHA to give a pink/red/yellow/orange appearance enabling the PHA-producing microbes to be identified and isolated. In PCR gene detection, primer pairs are designed to specifically amplify phaC gene, which encodes for PHA synthase, an enzyme responsible for PHA synthesis. The gene is typically present in PHA-accumulating microbes and absent in non-PHA-accumulating microbes, serving as the basis for identifying potential PHA-producers [120]. A review on the current research status of phaC gene detection and the available PCR primers is provided by Solaiman and Ashby [121].
Although staining and genetic detection methods provide a simple way to screen for PHA-producing microbes efficiently, these methods are also prone to detection errors. False positives may arise from the staining of other non-PHA lipid storage compounds and non-specific PCR amplification while false negatives could result from unfavorable conditions for biomass PHA accumulation and unsuitable detection primers or PCR conditions used in PCR amplification [120,122]. Hence, these two methods could only be employed as a presumptive test of PHA production potential. TEM on the other hand, enables direct visualization of PHA that appears as intracellular granules, providing affirmation of PHA bioacummulation [43]. However, TEM sample preparation is time-consuming and involves the use of radioactive chemicals [43], making it unsuitable for screening purposes. While Nile red/Nile blue staining, phaC gene detection and TEM are effective at providing evidence for PHA production or PHA-producing capacity, the downside of these methods is that they could neither quantify PHA nor provide qualitative information about PHA monomeric composition.

4.2. Quantification and Characterization of PHA in Biomass

Traditionally, crotonic acid assay was commonly used as a viable method for quantitative determination of P3HB [126,135,136]. In crotonic acid assay, P3HB is dissolved in concentrated sulphuric acid and converted into crotonic acid. As crotonic acid has a strong UV absorption at 235 nm in concentrated sulfuric acid, it can be measured by UV spectrophotometer. While this method is an easy and fast way to quantify P3HB, this method tends to overestimate P3HB content and is limited to P3HB determination [89,105]. Currently, quantification and characterization of various microbial intracellular PHAs can be achieved using modern analytical techniques, including fourier transform infrared spectroscopy (FTIR), liquid chromatography (LC) and gas chromatography (GC).
FTIR has been applied to detect and distinguish between the different types of PHA (i.e., scl-PHA, mcl-PHA, and scl-mcl-PHA), present within intact cells or as purified polymers. Characteristic ester carbonyl band for intracellular scl-PHA, mcl-PHA, and scl-mcl-PHA were observed at 1732 cm−1, 1744 cm−1 and 1739 cm−1, respectively whereas the same band for purified polymer scl-PHA, mcl-PHA, and scl-mcl-PHA were observed at 1728 cm−1, 1740 cm−1 and 1732 cm−1, respectively [127]. FTIR has also been proposed as a quantification tool for scl-PHA. Arcos-Hernandez et al. [137] showed that biomass scl-PHA content, between 0.03 and 0.58 weight/weight (w/w), could be coupled to FTIR spectra using a partial least squares model, allowing scl-PHA content determination within a standard error of prediction value of 0.023 w/w. The solvent-less nature of the FTIR technique and short analysis time eliminates risk exposure to hazardous chemicals while providing fast data output. However, FTIR-based methods have lower detection sensitivities, are inapt at describing or detecting changes in PHA monomeric composition, and cannot discriminate between PHA blends and copolymers [127]. Hence, FTIR-based methods tend to be more suitable for routine monitoring of PHA production for standard bioprocesses with well-characterized PHA products.
LC- and GC-based methods are the most frequently used analytical techniques due to automated sample analysis, and ability to provide accurate PHA quantification and qualitative information about PHA monomeric composition. Compared to FTIR-based methods, chromatography-based methods have higher detection sensitivities ranging from 0.014 to 14 μg for HPLC and 0.05 pg to 15 mg for GC depending on the type of detectors and chemical derivatization methods used [138]. The advent of automated LC platforms has enhanced scl-PHA analysis. Improvement in the measurement accuracy of P3HB-derived crotonic acid was made possible with ion-exchange HPLC coupled with UV detection [105,139,140]. Simultaneous analysis and quantification of 3HB and 3HV monomers could also be achieved through ion chromatography (IC) equipped with an anion trap column and a conductivity detector [129]. The advanced liquid chromatography-mass spectrometry (LC-MS) technique, using a combination of higher pressure and small diameter particles as column packing for separation with specific MS detection, could also be a sufficiently rapid and robust approach for the routine analysis of PHA monomers. However, to date, LC-MS has been applied only to a limited extent as a complementary technique to analyze PHA monomers [65]. There remains much potential to expand the capabilities of LC platforms beyond quantitative analysis of scl-PHA through utilizing LC-MS for quantification of other PHA monomers.
At present, GC remains the preferred method for qualitative and quantitative analysis of PHA monomers owing to its high separation power and detection sensitivity [138]. One of the earliest works on GC determination of PHA was reported by Braunegg et al. [134], who developed a method for accurate and reproducible determination of P3HB content in bacterial biomass using GC with flamed ionization detector (GC-FID). They showed that after subjecting P3HB-containing bacterial biomass to methanolysis, P3HB could be completely recovered in the form of its methyl esters derivatives and quantified to levels as low as 10−5 g L−1. GC-FID analysis was subsequently expanded to P3HV and mcl-PHA [81,141]. The robustness of GC-FID determination however, is dependent on the inclusion of appropriate PHA analytical standards. Conversely, coupling GC to mass spectrometry detector (GC-MS) ensures more reliable detection, identity confirmation and quantification of PHA monomers, as well as enables tentative identification of novel PHA monomers in the absence of analytical standards [133]. Figure 4 shows representative GC-MS chromatograms of PHA monomers [37]. A recent study by Tan et al. [132] showed that PHA monomers with carbon number between 4 and 16 have strong linear correlations with their respective retention times and response factors (R2 ≥ 0.987) under GC-MS, which enabled the retention time and response factor of other PHA monomers to be predicted. This method allowed a wide series of PHA, ranging from scl-PHA to lcl-PHA, to be reliably detected and quantified in the absence of reference standards. It is also expected that the coupling of GC to tandem mass spectrometry (GC-MS/MS) could offer improvements in sensitivity and specificity for determination of PHA monomers [142].
Despite their advantages, chromatography-based methods require PHA to be depolymerized and chemically converted into acids, diols or methyl ester derivatives prior to analysis [105,130,132]. This meant that chromatography-based methods are unable to distinguish if different PHA monomers are part of several homogenous PHA polymers or if they are part of a single PHA copolymer.
Figure 4. (A) Gas chromatography-mass spectrometry (GC-MS) total ion chromatogram of PHA after acidic methanolysis; (B) GC-MS extracted ion chromatogram of m/z 103. (Adapted from [37], with permission; Copyright 1998, National Academy of Sciences, USA). (C6, 3-hydroxyhexanoate; C8, 3-hydroxyoctanoate; C8:1, 3-hydroxyoctenoate; C9, 3-hydroxynonanoate; C10, 3-hydroxydecanoate; C11, 3-hydroxyundecanoate; C12, 3-hydroxydodecanoate; C12:1, 3-hydroxydodecenoate; C12:2, 3-hydroxydodecadienoate; C13, 3-hydroxytridecanoate; C14, 3-hydroxytetradecanoate; C14:1, 3-hydroxytetradecenoate; C14:2, 3-hydroxytetradecadienoate; C14:3, 3-hydroxytetradecatrienoate; C16, 3-hydroxyhexadecanoate; C16:1, 3-hydroxyhexadecenoate; C16:2, 3-hydroxyhexadecadienoate; C16:3, 3-hydroxyhexadecatrienoate)
Figure 4. (A) Gas chromatography-mass spectrometry (GC-MS) total ion chromatogram of PHA after acidic methanolysis; (B) GC-MS extracted ion chromatogram of m/z 103. (Adapted from [37], with permission; Copyright 1998, National Academy of Sciences, USA). (C6, 3-hydroxyhexanoate; C8, 3-hydroxyoctanoate; C8:1, 3-hydroxyoctenoate; C9, 3-hydroxynonanoate; C10, 3-hydroxydecanoate; C11, 3-hydroxyundecanoate; C12, 3-hydroxydodecanoate; C12:1, 3-hydroxydodecenoate; C12:2, 3-hydroxydodecadienoate; C13, 3-hydroxytridecanoate; C14, 3-hydroxytetradecanoate; C14:1, 3-hydroxytetradecenoate; C14:2, 3-hydroxytetradecadienoate; C14:3, 3-hydroxytetradecatrienoate; C16, 3-hydroxyhexadecanoate; C16:1, 3-hydroxyhexadecenoate; C16:2, 3-hydroxyhexadecadienoate; C16:3, 3-hydroxyhexadecatrienoate)
Polymers 06 00706 g004

5. PHA Polymer Extraction Methods

Microbial PHA is stored as insoluble intracellular granules. Methods to recover PHA would typically involve cell wall/cell membrane lysis, solubilization and purification of PHA component, and precipitation of PHA polymer. Common methods for PHA polymer recovery from microbial biomass are solvent extraction methods, chemical- and enzyme-based digestions methods, which will be briefly reviewed herein. A comprehensive review on this topic is provided by Kunasundari and Sudesh [143].
For scl-PHA, various PHA isolation methods have been developed using P3HB as a model polymer. A summary of the common extraction methods for scl-PHA is provided in Table 4. Among all recovery methods, solvent extraction is the most well-established and commonly used to obtain PHA polymer from biomass due to its high purity. In a study by Ramsay et al. [144], the solvents chloroform, methylene chloride or 1,2-dichloroethane were evaluated for P3HB recovery under various conditions (chloroform: 61 °C, 1 h; methylene chloride: 40 °C, 24 h; and 1,2-dichloroethane: 83 °C, 0.5 h). After solvent extraction, cellular debris was removed via filtration and the solution was concentrated by rotary evaporation before the P3HB polymer was precipitated by dropwise addition of ice-cold ethanol. Chloroform and 1,2-dichloroethane were observed to achieve high P3HB recovery (68%) with high purity (96% to 98%) compared to methylene chloride (recovery: 25%, purity: 98%). A novel method was developed for extraction of scl-PHA by “scl-PHA anti-solvent” acetone at elevated temperature and pressure in a closed system combining components for extraction, filtration, and product work-up. The quality of acetone-extracted polyesters showed no significant difference from chloroform-extracted ones, providing a promising substitute in terms of higher recyclability of the solvent without negatively impacting the structural features of the biopolyester [145]. The preponderance of solvent extraction lies on its high P3HB recovery purity, however there are much concerns about the high cost of operation as well as negative environmental impact caused by the generation of hazardous waste. One of the ways to minimize this problem is by using waste solvents for P3HB extraction. This has been demonstrated in Brazil where a pilot-scale P3HB production plant was integrated with a sugarcane mill and the solvent by-products of ethanolic fermentation from the mill was subsequently used as extraction solvents for P3HB recovery [146], thus presenting a feasible solution for achieving high P3HB purity at lower environmental cost.
Digestion method is a well-established alternative to solvent extraction for PHA recovery. In chemical-digestion, sodium hypochlorite is used to solubilize non-P3HB biomass thus achieving separation of P3HB content which can be recovered by centrifugation [147]. While the method is simple and effective, P3HB polymers obtained through hypochlorite digestion are generally of lower molecular masses due to the severe polymer degradation [147]. To resolve this problem, a combination approach was developed using dispersions of sodium hypochlorite solution as cell solubilizer, and chloroform to protect P3HB from further degradation after its release from cells, thus taking advantage of both hypochlorite digestion and solvent extraction [148].
Compared to solvent extraction and chemical digestion, enzymatic digestion requires milder operating conditions while achieving negligible product degradation [149]. Biomass was suspended in a specialized buffer and incubated at a specific temperature, which were optimized for enzymatic activity. After enzymatic hydrolysis, P3HB polymer was recovered by centrifugation. A polymer purity of up to 90% could be attained with this method [149]. Enzyme-based PHA recovery methods are safer in terms of operation, posed lower health risks, and has lower environmental footprint. Nevertheless, the high cost of enzymes may drive up the overall extraction cost.
For mcl-PHA, the recovery methods are based on similar principles and procedures as those for scl-PHA. Table 5 provides a summary of the common extraction methods for mcl-PHA. Like scl-PHA, mcl-PHA could also be recovered through solvent extraction and enzyme digestion, albeit with modifications to optimize mcl-PHA recovery. For solvent extraction of mcl-PHA, chloroform is commonly used according to procedures similar to those for scl-PHA. The main difference is in the final polymer precipitation step where ethanol is replaced with ice-cold methanol [150]. Apart from chloroform, dichloromethane is a less hazardous choice as an extraction solvent and together with ice-cold methanol as precipitant, up to 98% purity of mcl-PHA could be attained [151]. Solvent extraction however, tends to result in significant polymer degradation [144]. Jiang et al. [152] showed that this issue could be circumvented. Using P. putida biomass sample, the combination of biomass pretreatment with methanol and using acetone as the extraction solvent resulted in no detectable molecular mass loss of mcl-PHA with an overall purity of 94% [152].
Table 4. Poly(3-hydroxybutyrate) (P3HB) recovery methods (modified from [143]).
Table 4. Poly(3-hydroxybutyrate) (P3HB) recovery methods (modified from [143]).
MethodChemicalSpeciesConditionsPurity and recoveryReference
Solvent extractionChloroformCupriavidus necator
(DSM 545)
Mixing continuously at 25 °C for 12 hPurity: 94.0%-96.0%
Recovery: 65.0%-70.0%
Methylene chlorideCupriavidus necator
(DSM 545)
Mixing continuously at 25 °C for 12 hPurity: 95.0%-98.0%
Recovery: 24.0%-25.0%
1,2-DichloroethaneCupriavidus necator
(DSM 545)
Mixing continuously at 25 °C for 12 hPurity: 93.0%-98.0%
Recovery: 66.0%-70.0%
AcetoneHaloferax mediterranei
(DSM 1411)
Mixing continuously at 120 °C, 7 bar for 20 min under anaerobic conditions followed by filtering hot solution and cooling it down at 4 °C to precipitate polymerPurity: 98.4%
Recovery: 96.8%
Medium-chain-length alcoholsCupriavidus necator and Burkholderia sp.Multi-stage extraction process in continuous-stirred tank reactors. Remove cell debris from the extract and cool extract to recover polymerPurity: > 98.0%
Recovery: 95.0%
Hypochlorite digestionSodium hypochloriteCupriavidus necator
(DSM 545)
Biomass concentration: 10-40 g/L;
pH: 8-13.6; Temperature: 0-25 °C;
Digestion time: 10 min-6 h; Hypochlorite concentration: 1%-10.5% weight/volume (w/v)
Purity: 90.0%-98.0%
Recovery: 90.0%-95.0%
Sodium hypochlorite and chloroformCupriavidus necator
(NCIMB 11599) and
recombinant Escherichia coli
Biomass concentration: 1% (w/v);
Temperature: 30 °C; Digestion time: 1 h;
Hypochlorite concentration: 3%-20% volume/volume (v/v)
Purity: 86.0%
Recovery: NG
Purity: 93.0%
Recovery: NG
Enzyme digestionTrysin, bromelain, pancreatinCupriavidus necator (DSM 545)Digestion with 2% trypsin (50 °C, pH 9.0, 1 h) or 2% bromelain (50 °C, pH 4.75, 10 h) or 2% pancreatin (50 °C, pH 8.0, 8 h), followed by centrifugation and washing with 0.85% saline solutionPurity: 87.7%-90.3%
Recovery: NG
NG, not given.
Table 5. Medium-chain length PHA (mcl-PHA) recovery methods.
Table 5. Medium-chain length PHA (mcl-PHA) recovery methods.
MethodChemicalSpeciesConditionsPurity and recoveryReference
SolventChloroformPseudomonas oleovorans (strains NRRL B-14682, NRRL B-14683, and NRRL B-778)30 °C overnight at 250 rpmNG[150]
ChloroformPseudomonas oleovorans (NRRL B-14683), Pseudomonas resinovorans (NRRL B-2649), Pseudomonas citronellolis (NRRL B-2504), and Pseudomonas putida KT2442Soxhlet extraction for 24 hNG[154,155]
ChloroformPseudomonas putida IPT 046Soxhlet extraction for 6 hNG[156]
ChloroformPseudomonas aeruginosa 42A2 (NCIMB 40045)100 °C for 3 h in screw cap tubes for small quantities or in a soxhlet apparatus for large amounts of cell materialNG[157]
Dichlorome-thanePseudomonas oleovorans (ATCC 29347)Soxhlet extraction at 60 °C for 5 hPurity: > 98.0%
Recovery: NG
AcetonePseudomonas putida KT2440 (ATCC 47054)22 °C for 24 h at 170 rpmPurity: 80.0%–90.0%
Recovery: 60.0%–80.0%
Enzyme digestionAlcalase, SDS, EDTA, lysozymePseudomonas putidaDigestion with alcalase and SDS at pH 8.5 and 55 °C followed by further treatments with EDTA and lysozyme at pH 7 and 30 °CPurity: 92.6%
Recovery: nearly 90.0%
Pseudomonas putida KT2442Digestion with excess alcalase, EDTA and SDS at pH 8.5 and 55 °C followed by diafiltrationPurity: > 95.0%
Recovery: NG
NG, not given.
Table 6. Techniques for PHA polymer characterization.
Table 6. Techniques for PHA polymer characterization.
CharacteristicIndexMethodSampleSample preparationTypical conditionsReference
PHA monomeric compositionChemical derivative of PHA monomersLCRefer to Table 3
GCRefer to Table 3
PHA polymeric compositionTopology and functional groups of PHA molecule1D-Nuclear magnetic resonance (NMR)5–10 mg PHA for 1H-NMR and 20– 30 mg PHA for 13C-NMRDissolution of PHA polymer in 0.7 mL deuterated chloroform (CDCl3) containing 0.03% (v/v) tetramethylsilane (TMS)1H-NMR at 200 or 300 MHz and 13C-NMR measurements at 75.4 MHz at 20 °C with a sampling pulse of 3 s. Chemical shifts were referenced to the residual proton peak of CDCl3 at 7.26 ppm and to the carbon peak of CDCl3 at 77 ppm[82]
2D-NMR10 mg PHA for homonuclear 2D-NMR and 40–50 mg PHA for heteronuclear 2D-NMRRefer to above “1D-NMR”For homonuclear COSY and TOCSY, 16 scans were accumulated per increment over a spectral width of 7.8 ppm. For heteronuclear HSQC, 48 scans were accumulated per increment over a spectral width of 7.8 ppm for 1H and 75 ppm for 13C. For heteronuclear HMBC spectrum, 64 scans were acquired with the long-range coupling delay set for 8 Hz[161]
PHA polymeric compositionTopology and functional groups of PHA moleculeMatrix assisted laser desorption ionization-time of flight-mass spectrometry (MALDI-TOF-MS)1 µg–1 mg PHAThe matrix used was either dithranol or dihydroxybenzoic acid (DHB) at a concentration of 10 mg mL−1 in THF. 1 mg mL−1 PHA solution (in chloroform) was mixed with equal volume THF. The matrix solution and the PHA solution were subsequently mixed in a 5:2 ratio (matrix/sample). 1 µL mixture was deposited onto the stainless steel sample holder. The solvent was allowed to air-dry before loading the sample plate into the MALDI ion sourceMALDI-TOF-MS with 25 kV acceleration and detection in the positive-ion high-resolution reflection mode[162]
Molecular distributionPolydispersity, molecular mass and molecular mass distributionGel permeation chromatography (GPC)0.1–1 mg PHADissolution of PHA polymer in 1 mL of THFAnalysis conducted with a refractive index detector (47 °C, 2.0 bar) and a solvent-compatible GPC column. THF, containing 250 ppm of 2,6-di-tert-butyl-4-methylphenol (BHT) as inhibitor, was used as an eluent at a flow rate of 0.5 mL min−1 and 40 °C[43]
Dissolution of PHA polymer in chloroformAnalysis conducted with a differential refractive index detector (30 °C), a UV dual wavelength absorbance detector, and a combination of four GPC columns series. Chloroform was used as an eluent with a flow rate of 1.0 mL min−1[163]
MALDI-TOF-MSRefer to above “PHA polymeric composition”
Thermal propertiesGlass transition temperature and melting temperatureDifferential scanning calorimetry (DSC)10 mg PHA-Heat sample from −100 °C– 400 °C at a heating rate of 10 °C min−1 under purified air or nitrogen gas with a flow rate of 80 mL min−1[43]
Differential thermal analysis (DTA)5 mg PHA-Crystallization was carried out isothermally by abruptly quenching the samples from melt to the crystallization temperature, at which the samples were annealed for 10 min. Melting of semicrystalline samples was performed by heating at a rate of 5 °C min−1[164]
Thermodegradation temperatureThermogravimetric analysis (TGA)10 mg PHA-Heat sample from room temperature to 700 °C at a heating rate of 10 °C min−1 under purified air or nitrogen gas with a flow rate of 50 mL min−1[43]
CrystallinityMelting enthalpyDSCRefer to above “Thermal properties”
Infrared absorption bands correlated to crystallinityFTIR5–10 mg PHADissolve PHA in chloroform, apply onto KRS-5 window and blow dry to evaporate solvent. Alternatively, mix PHA with potassium bromide (KBr) powder and pelletizeRefer to Table 3[127,165]
Place PHA sample between two pieces of barium fluoride slidesMelt sample at 100 °C for 2 min in FTIR hot stage under the protection of dry nitrogen gas. Quench the amorphous sample to 58 and 28 °C by a flow of liquid nitrogen and maintain at these temperatures for 30 min for isothermal melt-crystallization before re-heating at 1 °C min−1[166]
CrystallinityDiffraction intensity correlated to crystallinityX-ray diffractionDry polymer powder-Diffractogram of the sample powder were measured at room temperature by an imaging plate diffractometer with Cu-Kα radiation (wavelength = 0.1542 nm) as an incident X-ray source emitted by a X-ray generator with a Ni filter. The scattering angle range of 2θ = 10°–40° at a scan speed of 3° min−1[156]
Mechanical propertiesTensile strength, tensile stress, percent elongation, modulus of elasticityMechanical testing machine of the constant-rate-of-crosshead-movement type with extensometer and micrometersPolymer thickness 1–14 mm, width 19–29 mm,
length 165–246 mm
Test samples were prepared using a hydraulic press at 150 °C and conditioned at a relative humidity of 50% ± 5% for 24 h prior to measurementsPerform stress-strain test at room temperature with a strain rate of 20 mm min−1[167]
For mcl-PHA recovery through enzyme digestion, enzymes alcalase and lysozyme, together with sodium dodecyl sulfate (SDS) and ethylenediaminetetraacetic acid (EDTA) were used. This combination of enzymes and chemicals were successfully applied to mcl-PHA isolation for fed-batch fermentations of up to 200 L, where the cells were first ruptured by thermal treatment and the resultant debris was exposed to excess protease (alcalase), EDTA and SDS for solubilization. After cross-flow microfiltration, the final mcl-PHA latex had a purity exceeding 95%, demonstrating potential commercial applicability [159,160]. In another study [158], the PHA granules, present in water suspension after enzymatic treatment, were recovered by removing the solubilized non-PHA cell material through ultrafiltration system and purified through continuous diafiltration process. The final purity of PHA was 92.6% and recovery was nearly 90% [158]. While enzyme digestion is a more environmental-friendly approach than solvent extraction, the purity of polymer attained is lower. Biomedical application requires a final purity of 99% or more which is currently not achievable through enzymatic mcl-PHA recovery and a second purification using solvent extraction is necessary [160].

6. Techniques for PHA Polymer Characterization

Purified PHA polymers are diverse in their chemical composition and material properties due to the myriad of PHA monomeric units available as well as the incorporation of these monomers at varying amounts. To identify suitable downstream applications for PHA, characterization of the biomaterial is imperative. A summary of these techniques and their execution is provided in Table 6.

6.1. Monomeric Composition and Distribution

The monomeric composition and distribution of PHA polymer could be determined from GC, LC and nuclear magnetic resonance spectroscopy (NMR). For chromatography-based methods, the analysis of PHA polymer is similar to that for intracellular PHA, requiring depolymerization of the polymer, usually combined with chemical derivatization before it can be analyzed [43,130,132], which meant that chromatography methods cannot analyze PHA as an intact polymer. NMR on the other hand, could study the chemical makeup of an intact PHA polymer and differentiate between PHA blends and PHA copolymer though providing details on the topology and functional group in molecules [150,153,154,168]. Typically, two types of NMR techniques are available and they are 1H-NMR and 13C-NMR. The high proton abundance in nature meant that 1H-NMR is more sensitive and requires shorter analytical time (within one hour). In contrast, owing to low sensitivity and natural abundance of 13C, it may require longer analysis (up to 24 h) to accumulate enough signal intensity for 13C-NMR spectrum. Despite its shortcoming, 13C-NMR performs better than 1H-NMR at the analysis of macromolecule as well as long carbon chain of monomers. As PHA polymers contain hydrogen and carbon, 1H-NMR and 13C-NMR are usually applied in combination to provide a more comprehensive analysis of the polymer. NMR is broadly used for saturated and unsaturated PHA analysis. Functional groups such as methane protons, methylene protons, –CH=CH– can be identified from both NMR spectra while microstructures like 3-hydroxypropionate (3HP) and 4-hydroxybutyrate (4HB) can be obtained by analyzing both 1H-NMR and 13C-NMR spectra [156,169]. Quantitative estimation of PHA monomers can also be performed with NMR using the intensity ratio of the signals [156]. NMR is also a powerful non-destructive tool that could be applied to the analysis of novel functionalized PHAs for which analytical standards are currently unavailable [170,171]. Typical 1H-NMR and 13C-NMR spectra of mcl-PHA are shown in Figure 5 [172].
Figure 5. (A) Typical 1H-NMR spectrum of PHA. Protons in the polymers are numbered and assigned to the peaks in the spectrum; (B) Typical 13C-NMR spectrum of PHA. Carbon atoms in polymers are numbered and assigned to peaks in the spectrum. (Adapted from [172], with permission). (3HHx, 3-hydroxyhexanoate; 3HO, 3-hydroxyoctanoate; 3HD, 3-hydroxydecanoate; 3HDD, 3-hydroxydodecanoate).
Figure 5. (A) Typical 1H-NMR spectrum of PHA. Protons in the polymers are numbered and assigned to the peaks in the spectrum; (B) Typical 13C-NMR spectrum of PHA. Carbon atoms in polymers are numbered and assigned to peaks in the spectrum. (Adapted from [172], with permission). (3HHx, 3-hydroxyhexanoate; 3HO, 3-hydroxyoctanoate; 3HD, 3-hydroxydecanoate; 3HDD, 3-hydroxydodecanoate).
Polymers 06 00706 g005
In addition, two-dimensional (2D) NMR methodologies (such as correlation spectroscopy [COSY], nuclear Overhauser effect spectroscopy [NOESY], heteronuclear multiple-quantum correlation [HMQC] and heteronuclear multiple bond coherence [HMBC]) are very useful in the characterization of all kinds of specialized PHA, such as unsaturated, branched, halogenated or acetylated PHA [43,56,161,173,174]. 2D-NMR provides information about the environment where each carbon/hydrogen is positioned. With the aid of 2D homonuclear or heteronuclear NMR techniques, the exact position of double bonds and the cis/trans configuration of monomers can be determined [43,173]. 1H-NMR and 2D homonuclear NMR were used to unambiguously identify the position of the hydroxyl group and the positions of the double bonds in mcl-PHA [174]; while 2D heteronuclear COSY NMR was successfully applied to show that 4-hydroxyvaleric acid was a constituent of the PHA polymer [56]. Moreover, 2D heteronuclear HMBC NMR could also clearly reveal the presence of the block microstructure in PHA [175].
Besides, it is notable that MS techniques, such as fast-atom bombardment (FAB)-MS [176], pyrolysis/MS [177] and matrix assisted laser desorption ionization-time of flight-mass spectrometry (MALDI-TOF-MS) [162,178,179] have been applied for the characterization of PHA composition. Among them, MALDI-TOF-MS offers a cost-effective, straightforward and high-throughput alternative to the well-established GC-MS methods [162,179]. Compared with GC-MS, MALDI-TOF-MS is a more straightforward method, and no chemical derivatization is required during sample preparation. As the differences in monomer composition and detailed PHA structural information can easily be identified by MALDI-TOF-MS, it can be used as complementary technique to NMR [178]. The total amount of sample deposited onto the target can be in the pico- to femtomole range, which makes MALDI-TOF-MS extremely sensitive with minimal sample requirement. The significant advantages provided by MALDI-TOF-MS technology therefore could facilitate routine analysis of PHA with high accuracy and precision.

6.2. Molecular Mass (Mw), Molecular Mass Distribution (Mn), and Polydispersity Index (PDI)

A PHA polymer’s average molecular mass (Mw), molecular mass distribution (Mn), and polydispersity index (PDI; Mw/Mn) could be determined through a gel permeation chromatography (GPC) system, calibrated with polystyrene standards [150]. The Mw of PHA spans over a wide range from 50 kDa to 10,000 kDa and depending on Mn value, PDI could be between 1.1 and 6.0 [13,97,180]. GPC columns such as Styragel HMW 6E (5 to 10,000 kDa) [150] and PLgel MIXED-C (0.2 to 2,000 kDa) [181] have been applied for PHA analysis. Often, the broad Mw range of PHA makes the analysis of an unknown polymer more challenging particularly when the biomaterial is a mixture of several PHA molecules with vastly different Mw and Mn values. Two or more GPC columns, connected in series, are therefore necessary to fully reveal the polymer’s Mw and Mn [153].
MALDI-TOF-MS, on the other hand, is the new and promising method for PHA characterization. It could potentially be used for evaluating the Mw and Mn of PHAs and their oligomers [162,178,179]. Unlike GPC that can only be used in determining apparent Mn values, MALDI-TOF-MS can offer accurate mass measurement of PHA [182]. Because of its high molecular resolution, excellent accuracy, reproducibility, and automation properties, MALDI-TOF-MS can make a significant contribution to the study of PHA.

6.3. Thermal Properties

Thermal properties such as glass transition temperature (Tg), melting temperature (Tm), and thermodegradation temperature (Td) are commonly examined for PHA material to determine the temperature conditions at which the polymer can be processed and utilized. The Tg, Tm, and Td values for PHA are usually in the range of −52 to 4 °C, non-observable to 177 °C, and 227 to 256 °C, respectively [7,180]. Information on Tg and Tm could be obtained using differential scanning calorimetry (DSC) and differential thermal analysis (DTA). The difference which sets them apart is that DTA is able to measure mass loss and qualitatively provide thermal information [183] while direct heat flow measurement enables DSC to provide not only qualitative results, but also quantitative thermal information, making DSC the preferred method in PHA studies [153,156]. The Td value of PHA is obtained using thermogravimetric analysis (TGA), a technique where a sample is heated in a controlled atmosphere at a defined rate while sample mass loss is measured [153,156]. The development of the simultaneous thermal analysis (STA) combines TGA and DSC/DTA measurement techniques, enabling Tg, Tm, and Td values determination on a single instrument, which provides a more productive and simpler means to analyze PHA [156].

6.4. Crystallinity

PHA polymers can range from non-crystalline to highly crystalline with crystallinity values between 0% and 70% [7,97]. Crystallinity could be measured by structural analysis instruments including FTIR, DSC and X-ray diffraction. In FTIR analysis, PHA displays characteristic infrared absorption bands at certain wavenumbers which can be correlated to crystallinity. The exact band locations vary according to the chemical composition of the polymer. For scl-PHAs such as P3HB and P3HB3HV, bands around 1279, 1228, and 1185 cm−1 are sensitive to the degree of crystallinity [184,185]. Band 1725 cm−1 and bands in the range of 1500 to 1300 cm−1, 1300 to 1000 cm−1 and 1000 to 800 cm−1 are revealing of the conformational changes of mcl-PHA and scl-mcl PHA in both the crystalline phase and amorphous phase [166]. In DSC analysis, melting enthalpy (∆Hm) provides an estimated value for heat of fusion (∆Hf) under the analysis conditions, which could be related to PHA’s crystallinity. PHA polymers with very low crystallinity typically have low to non-observable ∆Hf while highly-crystalline polymers such as P3HB can have ∆Hf values up to 146 J g−1 [7,186]. On their own, both FTIR and DSC are only adequate at measuring relative crystallinities within a given material. Measurement of the absolute crystallinity using FTIR and DSC can only be performed for PHA polymer with known crystallinity [187,188]. Absolute crystallinity could be obtained using methods based on X-ray diffraction. X-ray diffraction analysis is able to shed light on the polymer’s rate of crystallinity, as well as atomic structures such as chemical bonds, their disorder [189]. Crystallinity percentage can be calculated according to semi-crystalline and amorphous polymer areas in the diffractogram using Lorentzian and Gauss functions [156].

6.5. Mechanical Properties

Young’s modulus, elongation at break and tensile strength are mechanical properties commonly evaluated for PHA polymers. The Young’s modulus provides a measure of PHA’s stiffness and ranges from the very ductile mcl-PHA (0.008 MPa) to the stiffer scl-PHA (3.5 × 103 MPa) [7]. Elongation at break measures the extent that a material will stretch before it breaks and is expressed as a percentage of the material’s original length. PHA polymers can take the form of a hard rigid material or a soft elastomeric material, displaying a wide elongation at break values of between 2% and 1000% [180]. Tensile strength measures the amount of force required to pull a material until it breaks, and is typically in the range of 8.8 to 104 MPa for PHA polymers [7]. Measurement of the aforementioned mechanical properties can be performed with tensile tester instrument by standardized test methods such as the ASTM standards [167].

7. Conclusions

This review paper provided a summary of PHA biosynthetic pathways, PHA-producing microbial strains commercially available from culture collections and their application, as well as techniques for PHA analysis and polymer extraction. It is evident that there are many avenues through which PHA could be produced depending on the type of microorganisms employed, choice of carbon source, and cultivation conditions. These aforementioned factors also influenced the type of PHA produced, which in turn determines their downstream applications. Using this wealth of knowledge, future development in commercial PHA production could adopt a more “top-down” approach where the targeted carbon source and desired PHA product are decided a priori together with economic considerations before the appropriate microorganism or group of microoganisms is selected for the purpose as a means to achieve economic viability for the bioprocess. The formulation of microbial co-cultures for PHA production is largely considered as unexplored territories but may have the potential to produce PHA cheaply from organic waste streams. A fast-growing area in PHA research is the biosynthesis of tailored PHA for specific application needs. Existing microbial strains from culture collections serve as an excellent platform for genetic modification to produce specialized PHAs and enhancing PHA yield. The elucidation of PHA biosynthetic pathways is also likely to complement such research efforts. Many well-established methods are currently available for PHA analysis but each of them come with their own strengths and limitations. On the basis of the reports that have been gathered to date, GC-MS in conjunction with NMR remains a pre-eminent analytical tool in PHA investigations. Well-established analytical methods such as FTIR, GPC and X-ray diffraction can provide general information on the overall structures, molecular mass distribution and rate of crystallinity, respectively. However, it is important to further develop efficient technologies (e.g., LC-MS and MALDI-TOF-MS) for characterization of PHA. It is expected that the advanced analytical approaches will provide us with further insights about the physical properties and degradation mechanisms of PHA.


The authors gratefully acknowledge the financial support (ETRP 0901 161) from the National Environment Agency, Singapore.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Anderson, A.J.; Dawes, E.A. Occurrence, metabolism, metabolic role, and industrial uses of bacterial polyhydroxyalkanoates. Microbiol. Rev. 1990, 54, 450–472. [Google Scholar]
  2. Shah, A.A.; Hasan, F.; Hameed, A.; Ahmed, S. Biological degradation of plastics: A comprehensive review. Biotechnol. Adv. 2008, 26, 246–265. [Google Scholar] [CrossRef]
  3. Khanna, S.; Srivastava, A.K. Recent advances in microbial polyhydroxyalkanoates. Process Biochem. 2005, 40, 607–619. [Google Scholar] [CrossRef]
  4. Lu, J.; Tappel, R.C.; Nomura, C.T. Mini-review: Biosynthesis of poly(hydroxyalkanoates). Polym. Rev. 2009, 49, 226–248. [Google Scholar] [CrossRef]
  5. Zinn, M.; Hany, R. Tailored material properties of polyhydroxyalkanoates through biosynthesis and chemical modification. Adv. Eng. Mater. 2005, 7, 408–411. [Google Scholar] [CrossRef]
  6. Escapa, I.F.; Morales, V.; Martino, V.P.; Pollet, E.; Avérous, L.; García, J.L.; Prieto, M.A. Disruption of β-oxidation pathway in Pseudomonas putida KT2442 to produce new functionalized PHAs with thioester groups. Appl. Microbiol. Biotechnol. 2011, 89, 1583–1598. [Google Scholar] [CrossRef]
  7. Rai, R.; Keshavarz, T.; Roether, J.A.; Boccaccini, A.R.; Roy, I. Medium chain length polyhydroxyalkanoates, promising new biomedical materials for the future. Mater. Sci. Eng. R. Rep. 2011, 72, 29–47. [Google Scholar] [CrossRef]
  8. De Roo, G.; Kellerhals, M.B.; Ren, Q.; Witholt, B.; Kessler, B. Production of chiral R-3-hydroxyalkanoic acids and R-3-hydroxyalkanoic acid methylesters via hydrolytic degradation of polyhydroxyalkanoate synthesized by Pseudomonads. Biotechnol. Bioeng. 2002, 77, 717–722. [Google Scholar] [CrossRef]
  9. Philip, S.; Keshavarz, T.; Roy, I. Polyhydroxyalkanoates: Biodegradable polymers with a range of applications. J. Chem. Technol. Biotechnol. 2007, 82, 233–247. [Google Scholar] [CrossRef]
  10. Olivera, E.R.; Arcos, M.; Naharro, G.; Luengo, J.M. Unusual PHA biosynthesis. In Plastics from Bacteria: Natural Functions and Applications; Chen, G.-Q., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 14, pp. 133–186. [Google Scholar]
  11. Chen, G.-Q. Plastics completely synthesized by bacteria: Polyhydroxyalkanoates. In Plastics from Bacteria: Natural Functions and Applications; Chen, G.-Q., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 14, pp. 17–37. [Google Scholar]
  12. Kadouri, D.; Jurkevitch, E.; Okon, Y.; Castro-Sowinski, S. Ecological and agricultural significance of bacterial polyhydroxyalkanoates. Crit. Rev. Microbiol. 2005, 31, 55–67. [Google Scholar] [CrossRef]
  13. Madison, L.L.; Huisman, G.W. Metabolic engineering of poly(3-hydroxyalkanoates): From DNA to plastic. Microbiol. Mol. Biol. Rev. 1999, 63, 21–53. [Google Scholar]
  14. Rothermich, M.M.; Guerrero, R.; Lenz, R.W.; Goodwin, S. Characterization, seasonal occurrence, and diel fluctuation of poly(hydroxyalkanoate) in photosynthetic microbial mats. Appl. Environ. Microbiol. 2000, 66, 4279–4291. [Google Scholar] [CrossRef]
  15. Peplinski, K.; Ehrenreich, A.; Döring, C.; Bömeke, M.; Reinecke, F.; Hutmacher, C.; Steinbüchel, A. Genome-wide transcriptome analyses of the “Knallgas” bacterium Ralstonia eutropha H16 with regard to polyhydroxyalkanoate metabolism. Microbiology 2010, 156, 2136–2152. [Google Scholar] [CrossRef]
  16. Shimizu, R.; Chou, K.; Orita, I.; Suzuki, Y.; Nakamura, S.; Fukui, T. Detection of phase-dependent transcriptomic changes and Rubisco-mediated CO2 fixation into poly (3-hydroxybutyrate) under heterotrophic condition in Ralstonia eutropha H16 based on RNA-seq and gene deletion analyses. BMC Microbiol. 2013, 13, 169. [Google Scholar] [CrossRef]
  17. Yamane, T. Yield of poly-D(-)-3-hydroxybutyrate from various carbon sources: A theoretical study. Biotechnol. Bioeng. 1993, 41, 165–170. [Google Scholar] [CrossRef]
  18. Steinbüchel, A.; Hein, S. Biochemical and molecular basis of microbial synthesis of polyhydroxyalkanoates in microorganisms. In Biopolyesters; Babel, W., Steinbüchel, A., Eds.; Springer: Berlin/Heidelberg, Germany, 2001; Volume 71, pp. 81–123. [Google Scholar]
  19. Ratledge, C.; Kristiansen, B. Basic Biotechnology, 2nd ed.; Cambridge University Press: Cambridge, UK, 2001. [Google Scholar]
  20. Jung, Y.M.; Lee, Y.H. Utilization of oxidative pressure for enhanced production of poly-β-hydroxybutyrate and poly(3-hydroxybutyrate-3-hydroxyvalerate) in Ralstonia eutropha. J. Biosci. Bioeng. 2000, 90, 266–270. [Google Scholar]
  21. Khosravi-Darani, K.; Mokhtari, Z.-B.; Amai, T.; Tanaka, K. Microbial production of poly(hydroxybutyrate) from C1 carbon sources. Appl. Microbiol. Biotechnol. 2013, 97, 1407–1424. [Google Scholar] [CrossRef]
  22. Raberg, M.; Bechmann, J.; Brandt, U.; Schlüter, J.; Uischner, B.; Voigt, B.; Hecker, M.; Steinbüchel, A. Versatile metabolic adaptations of Ralstonia eutropha H16 to a loss of PdhL, the E3 component of the pyruvate dehydrogenase complex. Appl. Environ. Microbiol. 2011, 77, 2254–2263. [Google Scholar] [CrossRef]
  23. Peoples, O.P.; Sinskey, A.J. Poly-β-hydroxybutyrate (PHB) biosynthesis in Alcaligenes eutrophus H16. Identification and characterization of the PHB polymerase gene (phbC). J. Biol. Chem. 1989, 264, 15298–15303. [Google Scholar]
  24. Lee, S.; Jeon, E.; Yun, H.S.; Lee, J. Improvement of fatty acid biosynthesis by engineered recombinant Escherichia coli. Biotechnol. Bioprocess Eng. 2011, 16, 706–713. [Google Scholar] [CrossRef]
  25. Nomura, C.T.; Taguchi, K.; Taguchi, S.; Doi, Y. Coexpression of genetically engineered 3-ketoacyl-ACP synthase III (fabH) and polyhydroxyalkanoate synthase (phaC) genes leads to short-chain-length-medium-chain-length polyhydroxyalkanoate copolymer production from glucose in Escherichia coli JM109. Appl. Environ. Microbiol. 2004, 70, 999–1007. [Google Scholar] [CrossRef]
  26. Ren, Q.; Sierro, N.; Witholt, B.; Kessler, B. FabG, an NADPH-dependent 3-ketoacyl reductase of Pseudomonas aeruginosa, provides precursors for medium-chain-length poly-3-hydroxyalkanoate biosynthesis in Escherichia coli. J. Bacteriol. 2000, 182, 2978–2981. [Google Scholar] [CrossRef]
  27. Valentin, H.E.; Dennis, D. Production of poly(3-hydroxybutyrate-co-4-hydroxybutyrate) in recombinant Escherichia coli grown on glucose. J. Biotechnol. 1997, 58, 33–38. [Google Scholar] [CrossRef]
  28. Xie, W.P.; Chen, G.-Q. Production and characterization of terpolyester poly(3-hydroxybutyrate-co-4-hydroxybutyrate-co-3-hydroxyhexanoate) by recombinant Aeromonas hydrophila 4AK4 harboring genes phaPCJ. Biochem. Eng. J. 2008, 38, 384–389. [Google Scholar] [CrossRef]
  29. Valentin, H.E.; Steinbüchel, A. Accumulation of poly(3-hydroxybutyric acid-co-3-hydroxyvaleric acid-co-4-hydroxyvaleric acid) by mutants and recombinant strains of Alcaligenes eutrophus. J. Polym. Environ. 1995, 3, 169–175. [Google Scholar] [CrossRef]
  30. Aldor, I.S.; Kim, S.W.; Jones Prather, K.L.; Keasling, J.D. Metabolic engineering of a novel propionate-independent pathway for the production of poly(3-hydroxybutyrate-co-3-hydroxyvalerate) in recombinant Salmonella enterica serovar Typhimurium. Appl. Environ. Microbiol. 2002, 68, 3848–3854. [Google Scholar] [CrossRef]
  31. Valentin, H.E.; Dennis, D. Metabolic pathway for poly(3-hydroxybutyrate-co-3-hydroxyvalerate) formation in Nocardia corallina: Inactivation of mutB by chromosomal integration of a kanamycin resistance gene. Appl. Environ. Microbiol. 1996, 62, 372–379. [Google Scholar]
  32. Satoh, H.; Mino, T.; Matsuo, T. PHA production by activated sludge. Int. J. Biol. Macromol. 1999, 25, 105–109. [Google Scholar] [CrossRef]
  33. Slater, S.; Houmiel, K.L.; Tran, M.; Mitsky, T.A.; Taylor, N.B.; Padgette, S.R.; Gruys, K.J. Multiple β-ketothiolases mediate poly(β-hydroxyalkanoate) copolymer synthesis in Ralstonia eutropha. J. Bacteriol. 1998, 180, 1979–1987. [Google Scholar]
  34. Chohan, S.N.; Copeland, L. Acetoacetyl coenzyme a reductase and polyhydroxybutyrate synthesis in Rhizobium (Cicer) sp. strain CC 1192. Appl. Environ. Microbiol. 1998, 64, 2859–2863. [Google Scholar]
  35. Hume, A.R.; Nikodinovic-Runic, J.; O’Connor, K.E. FadD from Pseudomonas putida CA-3 is a true long-chain fatty acyl coenzyme A synthetase that activates phenylalkanoic and alkanoic acids. J. Bacteriol. 2009, 191, 7554–7565. [Google Scholar] [CrossRef]
  36. Yuan, M.-Q.; Shi, Z.-Y.; Wei, X.-X.; Wu, Q.; Chen, S.-F.; Chen, G.-Q. Microbial production of medium-chain-length 3-hydroxyalkanoic acids by recombinant Pseudomonas putida KT2442 harboring genes fadL, fadD and phaZ. FEMS Microbiol. Lett. 2008, 283, 167–175. [Google Scholar] [CrossRef]
  37. Mittendorf, V.; Robertson, E.J.; Leech, R.M.; Krüger, N.; Steinbüchel, A.; Poirier, Y. Synthesis of medium-chain-length polyhydroxyalkanoates in Arabidopsis thaliana using intermediates of peroxisomal fatty acid β-oxidation. Proc. Natl. Acad. Sci. USA 1998, 95, 13397–13402. [Google Scholar] [CrossRef]
  38. Sato, S.; Kanazawa, H.; Tsuge, T. Expression and characterization of (R)-specific enoyl coenzyme A hydratases making a channeling route to polyhydroxyalkanoate biosynthesis in Pseudomonas putida. Appl. Microbiol. Biotechnol. 2011, 90, 951–959. [Google Scholar] [CrossRef]
  39. Ouyang, S.-P.; Luo, R.C.; Chen, S.-S.; Liu, Q.; Chung, A.; Wu, Q.; Chen, G.-Q. Production of polyhydroxyalkanoates with high 3-hydroxydodecanoate monomer content by fadB and fadA knockout mutant of Pseudomonas putida KT2442. Biomacromolecules 2007, 8, 2504–2511. [Google Scholar] [CrossRef]
  40. Zheng, L.Z.; Li, Z.; Tian, H.-L.; Li, M.; Chen, G.-Q. Molecular cloning and functional analysis of (R)-3-hydroxyacyl-acyl carrier protein:coenzyme A transacylase from Pseudomonas mendocina LZ. FEMS Microbiol. Lett. 2005, 252, 299–307. [Google Scholar] [CrossRef]
  41. Brzostowicz, P.B.; Blasko, M.B.; Rouvière, P.R. Identification of two gene clusters involved in cyclohexanone oxidation in Brevibacterium epidermidis strain HCU. Appl. Microbiol. Biotechnol. 2002, 58, 781–789. [Google Scholar] [CrossRef]
  42. Poli, A.; Di Donato, P.; Abbamondi, G.R.; Nicolaus, B. Synthesis, production, and biotechnological applications of exopolysaccharides and polyhydroxyalkanoates by archaea. Archaea 2011, 2011, 1–13. [Google Scholar]
  43. Galia, M.B. Isolation and analysis of storage compounds. In Handbook of Hydrocarbon and Lipid Microbiology; Timmis, K.N., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; pp. 3725–3741. [Google Scholar]
  44. Verlinden, R.A.J.; Hill, D.J.; Kenward, M.A.; Williams, C.D.; Radecka, I. Bacterial synthesis of biodegradable polyhydroxyalkanoates. J. Appl. Microbiol. 2007, 102, 1437–1449. [Google Scholar] [CrossRef]
  45. Koller, M.; Atlić, A.; Dias, M.; Reiterer, A.; Braunegg, G. Microbial PHA production from waste raw materials. In Plastics from Bacteria: Natural Functions and Applications; Chen, G.-Q., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 14, pp. 85–119. [Google Scholar]
  46. Yu, P.H.; Chua, H.; Huang, A.L.; Lo, W.; Chen, G.-Q. Conversion of food industrial wastes into bioplastics. Appl. Biochem. Biotechnol. 1998, 70, 603–614. [Google Scholar]
  47. Wang, F.; Lee, S.Y. Poly(3-hydroxybutyrate) production with high productivity and high polymer content by a fed-batch culture of Alcaligenes latus under nitrogen limitation. Appl. Environ. Microbiol. 1997, 63, 3703–3706. [Google Scholar]
  48. Grothe, E.; Moo-Young, M.; Chisti, Y. Fermentation optimization for the production of poly(beta-hydroxybutyric acid) microbial thermoplastic. Enzyme Microb. Technol. 1999, 25, 132–141. [Google Scholar] [CrossRef]
  49. Yamane, T.; Fukunaga, M.; Lee, Y.W. Increased PHB productivity by high-cell-density fed-batch culture of Alcaligenes latus, a growth-associated PHB producer. Biotechnol. Bioeng. 1996, 50, 197–202. [Google Scholar] [CrossRef]
  50. Gomez, J.; Rodrigues, M.; Alli, R.; Torres, B.; Netto, C.B.; Oliveira, M.; Da Silva, L. Evaluation of soil Gram-negative bacteria yielding polyhydroxyalkanoic acids from carbohydrates and propionic acid. Appl. Microbiol. Biotechnol. 1996, 45, 785–791. [Google Scholar] [CrossRef]
  51. Lasemi, Z.; Darzi, G.N.; Baei, M.S. Media optimization for poly(β-hydroxybutyrate) production using Azotobacter Beijerinckii. Int. J. Polym. Mater. 2012, 62, 265–269. [Google Scholar] [CrossRef]
  52. Pan, W.; Perrotta, J.A.; Stipanovic, A.J.; Nomura, C.T.; Nakas, J.P. Production of polyhydroxyalkanoates by Burkholderia cepacia ATCC 17759 using a detoxified sugar maple hemicellulosic hydrolysate. J. Ind. Microbiol. Biotechnol. 2012, 39, 459–469. [Google Scholar] [CrossRef]
  53. Zhu, C.; Nomura, C.T.; Perrotta, J.A.; Stipanovic, A.J.; Nakas, J.P. Production and characterization of poly-3-hydroxybutyrate from biodiesel-glycerol by Burkholderia cepacia ATCC 17759. Biotechnol. Prog. 2010, 26, 424–430. [Google Scholar]
  54. Chee, J.-Y.; Tan, Y.; Samian, M.-R.; Sudesh, K. Isolation and characterization of a Burkholderia sp. USM (JCM15050) capable of producing polyhydroxyalkanoate (PHA) from triglycerides, fatty acids and glycerols. J. Polym. Environ. 2010, 18, 584–592. [Google Scholar] [CrossRef]
  55. Qi, Q.S.; Rehm, B.H.A. Polyhydroxybutyrate biosynthesis in Caulobacter crescentus: Molecular characterization of the polyhydroxybutyrate synthase. Microbiology 2001, 147, 3353–3358. [Google Scholar]
  56. Valentin, H.E.; Lee, E.Y.; Choi, C.Y.; Steinbüchel, A. Identification of 4-hydroxyhexanoic acid as a new constituent of biosynthetic polyhydroxyalkanoic acids from bacteria. Appl. Microbiol. Biotechnol. 1994, 40, 710–716. [Google Scholar] [CrossRef]
  57. Fukui, T.; Doi, Y. Efficient production of polyhydroxyalkanoates from plant oils by Alcaligenes eutrophus and its recombinant strain. Appl. Microbiol. Biotechnol. 1998, 49, 333–336. [Google Scholar] [CrossRef]
  58. Chakraborty, P.; Gibbons, W.; Muthukumarappan, K. Conversion of volatile fatty acids into polyhydroxyalkanoate by Ralstonia eutropha. J. Appl. Microbiol. 2009, 106, 1996–2005. [Google Scholar]
  59. Sonnleitner, B.; Heinzle, E.; Braunegg, G.; Lafferty, R.M. Formal kinetics of poly-β-hydroxybutyric acid (PHB) production in Alcaligenes eutrophus H 16 and Mycoplana rubra R 14 with respect to the dissolved oxygen tension in ammonium-limited batch cultures. Eur. J. Appl. Microbiol. 1979, 7, 1–10. [Google Scholar] [CrossRef]
  60. Ishizaki, A.; Tanaka, K. Production of poly-β-hydroxybutyric acid from carbon dioxide by Alcaligenes eutrophus ATCC 17697T. J. Ferment. Bioeng. 1991, 71, 254–257. [Google Scholar] [CrossRef]
  61. Kim, B.S.; Lee, S.C.; Lee, S.Y.; Chang, H.N.; Chang, Y.K.; Woo, S.I. Production of poly(3-hydroxybutyric acid) by fed-batch culture of Alcaligenes eutrophus with glucose concentration control. Biotechnol. Bioeng. 1994, 43, 892–898. [Google Scholar] [CrossRef]
  62. Haas, R.; Jin, B.; Zepf, F.T. Production of poly(3-hydroxybutyrate) from waste potato starch. Biosci. Biotechnol. Biochem. 2008, 72, 253–256. [Google Scholar]
  63. Beaulieu, M.; Beaulieu, Y.; Melinard, J.; Pandian, S.; Goulet, J. Influence of ammonium salts and cane molasses on growth of Alcaligenes eutrophus and production of polyhydroxybutyrate. Appl. Environ. Microbiol. 1995, 61, 165–169. [Google Scholar]
  64. Du, G.C.C.; Chen, J.; Yu, J.; Lun, S.Y. Feeding strategy of propionic acid for production of poly(3-hydroxybutyrate-co-3-hydroxyvalerate) with Ralstonia eutropha. Biochem. Eng. J. 2001, 8, 103–110. [Google Scholar] [CrossRef]
  65. Cavalheiro, J.M.B.T.; de Almeida, M.C.M.D.; Grandfils, C.; da Fonseca, M.M.R. Poly(3-hydroxybutyrate) production by Cupriavidus necator using waste glycerol. Process Biochem. 2009, 44, 509–515. [Google Scholar] [CrossRef]
  66. Quillaguamán, J.; Hashim, S.; Bento, F.; Mattiasson, B.; Hatti-Kaul, R. Poly(β-hydroxybutyrate) production by a moderate halophile, Halomonas boliviensis LC1 using starch hydrolysate as substrate. J. Appl. Microbiol. 2005, 99, 151–157. [Google Scholar] [CrossRef]
  67. Povolo, S.; Romanelli, M.G.; Basaglia, M.; Ilieva, V.I.; Corti, A.; Morelli, A.; Chiellini, E.; Casella, S. Polyhydroxyalkanoate biosynthesis by Hydrogenophaga pseudoflava DSM1034 from structurally unrelated carbon sources. New Biotechnol. 2013, 30, 629–634. [Google Scholar] [CrossRef]
  68. Koller, M.; Hesse, P.; Bona, R.; Kutschera, C.; Atlic, A.; Braunegg, G. Potential of various archae- and eubacterial strains as industrial polyhydroxyalkanoate producers from whey. Macromol. Biosci. 2007, 7, 218–226. [Google Scholar] [CrossRef]
  69. Bourque, D.; Pomerleau, Y.; Groleau, D. High-cell-density production of poly-β-hydroxybutyrate (PHB) from methanol by Methylobacterium extorquens: Production of high-molecular-mass PHB. Appl. Microbiol. Biotechnol. 1995, 44, 367–376. [Google Scholar] [CrossRef]
  70. Mokhtari-Hosseini, Z.B.; Vasheghani-Farahani, E.; Heidarzadeh-Vazifekhoran, A.; Shojaosadati, S.A.; Karimzadeh, R.; Darani, K.K. Statistical media optimization for growth and PHB production from methanol by a methylotrophic bacterium. Bioresour. Technol. 2009, 100, 2436–2443. [Google Scholar] [CrossRef]
  71. Mokhtari-Hosseini, Z.B.; Vasheghani-Farahani, E.; Shojaosadati, S.A.; Karimzadeh, R.; Heidarzadeh-Vazifekhoran, A. Effect of feed composition on PHB production from methanol by HCDC of Methylobacterium extorquens (DSMZ 1340). J. Chem. Technol. Biotechnol. 2009, 84, 1136–1139. [Google Scholar] [CrossRef]
  72. Wendlandt, K.D.; Jechorek, M.; Helm, J.; Stottmeister, U. Production of PHB with a high molecular mass from methane. Polym. Degrad. Stabil. 1998, 59, 191–194. [Google Scholar] [CrossRef]
  73. Smit, A.M.; Strabala, T.J.; Peng, L.; Rawson, P.; Lloyd-Jones, G.; Jordan, T.W. Proteomic phenotyping of Novosphingobium nitrogenifigens reveals a robust capacity for simultaneous nitrogen fixation, polyhydroxyalkanoate production, and resistance to reactive oxygen species. Appl. Environ. Microbiol. 2012, 78, 4802–4815. [Google Scholar] [CrossRef]
  74. Yamane, T.; Chen, X.; Ueda, S. Growth-associated production of poly(3-hydroxyvalerate) from n-pentanol by a methylotrophic bacterium, Paracoccus denitrificans. Appl. Environ. Microbiol. 1996, 62, 380–384. [Google Scholar]
  75. Tripathi, A.D.; Yadav, A.; Jha, A.; Srivastava, S.K. Utilizing of sugar refinery waste (cane molasses) for production of bio-plastic under submerged fermentation process. J. Polym. Environ. 2012, 20, 446–453. [Google Scholar] [CrossRef]
  76. Guzik, M.W.; Kenny, S.T.; Duane, G.F.; Casey, E.; Woods, T.; Babu, R.P.; Nikodinovic-Runic, J.; Murray, M.; O’Connor, K.E. Conversion of post consumer polyethylene to the biodegradable polymer polyhydroxyalkanoate. Appl. Microbiol. Biotechnol. 2014. In Press. [Google Scholar]
  77. Kenny, S.T.; Nikodinovic-Runic, J.; Kaminsky, W.; Woods, T.; Babu, R.P.; Keely, C.M.; Blau, W.; O’Connor, K.E. Up-cycling of PET (polyethylene terephthalate) to the biodegradable plastic PHA (polyhydroxyalkanoate). Environ. Sci. Technol. 2008, 42, 7696–7701. [Google Scholar] [CrossRef]
  78. Lee, E.; Jendrossek, D.; Schirmer, A.; Choi, C.; Steinbüchel, A. Biosynthesis of copolyesters consisting of 3-hydroxybutyric acid and medium-chain-length 3-hydroxyalkanoic acids from 1,3-butanediol or from 3-hydroxybutyrate by Pseudomonas sp. A33. Appl. Microbiol. Biotechnol. 1995, 42, 901–909. [Google Scholar] [CrossRef]
  79. Nikodinovic-Runic, J.; Casey, E.; Duane, G.F.; Mitic, D.; Hume, A.R.; Kenny, S.T.; O’Connor, K.E. Process analysis of the conversion of styrene to biomass and medium chain length polyhydroxyalkanoate in a two-phase bioreactor. Biotechnol. Bioeng. 2011, 108, 2447–2455. [Google Scholar] [CrossRef]
  80. Ward, P.G.; Goff, M.; Donner, M.; Kaminsky, W.; O'Connor, K.E. A two step chemo-biotechnological conversion of polystyrene to a biodegradable thermoplastic. Environ. Sci. Technol. 2006, 40, 2433–2437. [Google Scholar] [CrossRef]
  81. Lageveen, R.G.; Huisman, G.W.; Preusting, H.; Ketelaar, P.; Eggink, G.; Witholt, B. Formation of polyesters by Pseudomonas oleovorans: Effect of substrates on formation and composition of poly-(R)-3-hydroxyalkanoates and poly-(R)-3-hydroxyalkenoates. Appl. Environ. Microbiol. 1988, 54, 2924–2932. [Google Scholar]
  82. Gross, R.A.; DeMello, C.; Lenz, R.W.; Brandl, H.; Fuller, R.C. The biosynthesis and characterization of poly(β-hydroxyalkanoates) produced by Pseudomonas oleovorans. Macromolecules 1989, 22, 1106–1115. [Google Scholar] [CrossRef]
  83. Elbahloul, Y.; Steinbuchel, A. Large-scale production of poly(3-hydroxyoctanoic acid) by Pseudomonas putida GPo1 and a simplified downstream process. Appl. Environ. Microbiol. 2009, 75, 643–651. [Google Scholar] [CrossRef]
  84. Sun, Z.; Ramsay, J.A.; Guay, M.; Ramsay, B.A. Carbon-limited fed-batch production of medium-chain-length polyhydroxyalkanoates from nonanoic acid by Pseudomonas putida KT2440. Appl. Microbiol. Biotechnol. 2007, 74, 69–77. [Google Scholar] [CrossRef]
  85. Davis, R.; Kataria, R.; Cerrone, F.; Woods, T.; Kenny, S.; O’Donovan, A.; Guzik, M.; Shaikh, H.; Duane, G.; Gupta, V.K.; et al. Conversion of grass biomass into fermentable sugars and its utilization for medium chain length polyhydroxyalkanoate (mcl-PHA) production by Pseudomonas strains. Bioresour. Technol. 2013, 150, 202–209. [Google Scholar] [CrossRef]
  86. Nikodinovic, J.; Kenny, S.T.; Babu, R.P.; Woods, T.; Blau, W.; O’Connor, K.E. The conversion of BTEX compounds by single and defined mixed cultures to medium-chain-length polyhydroxyalkanoate. Appl. Microbiol. Biotechnol. 2008, 80, 665–673. [Google Scholar] [CrossRef]
  87. Shahid, S.; Mosrati, R.; Ledauphin, J.; Amiel, C.; Fontaine, P.; Gaillard, J.-L.; Corroler, D. Impact of carbon source and variable nitrogen conditions on bacterial biosynthesis of polyhydroxyalkanoates: Evidence of an atypical metabolism in Bacillus megaterium DSM 509. J. Biosci. Bioeng. 2013, 116, 302–308. [Google Scholar] [CrossRef]
  88. Pantazaki, A.A.; Papaneophytou, C.P.; Pritsa, A.G.; Liakopoulou-Kyriakides, M.; Kyriakidis, D.A. Production of polyhydroxyalkanoates from whey by Thermus thermophilus HB8. Process Biochem. 2009, 44, 847–853. [Google Scholar] [CrossRef]
  89. Valappil, S.P.; Boccaccini, A.R.; Bucke, C.; Roy, I. Polyhydroxyalkanoates in Gram-positive bacteria: Insights from the genera Bacillus and Streptomyces. Antonie Van Leeuwenhoek 2007, 91, 1–17. [Google Scholar]
  90. Haywood, G.W.; Anderson, A.J.; Roger Williams, D.; Dawes, E.A.; Ewing, D.F. Accumulation of a poly(hydroxyalkanoate) copolymer containing primarily 3-hydroxyvalerate from simple carbohydrate substrates by Rhodococcus sp. NCIMB 40126. Int. J. Biol. Macromol. 1991, 13, 83–88. [Google Scholar] [CrossRef]
  91. Akar, A.; Akkaya, E.U.; Yesiladali, S.K.; Celikyilmaz, G.; Cokgor, E.U.; Tamerler, C.; Orhon, D.; Cakar, Z.P. Accumulation of polyhydroxyalkanoates by Microlunatus phosphovorus under various growth conditions. J. Ind. Microbiol. Biotechnol. 2006, 33, 215–220. [Google Scholar] [CrossRef]
  92. Bhattacharyya, A.; Pramanik, A.; Maji, S.K.; Haldar, S.; Mukhopadhyay, U.K.; Mukherjee, J. Utilization of vinasse for production of poly-3-(hydroxybutyrate-co-hydroxyvalerate) by Haloferax mediterranei. AMB Express 2012, 2, 1–10. [Google Scholar] [CrossRef]
  93. Koller, M.; Atlić, A.; Gonzalez‐Garcia, Y.; Kutschera, C.; Braunegg, G. Polyhydroxyalkanoate (PHA) biosynthesis from whey lactose. Macromol. Symp. 2008, 272, 87–92. [Google Scholar] [CrossRef]
  94. Hermann-Krauss, C.; Koller, M.; Muhr, A.; Fasl, H.; Stelzer, F.; Braunegg, G. Archaeal production of polyhydroxyalkanoate (PHA) co-and terpolyesters from biodiesel industry-derived by-products. Archaea 2013, 2013. [Google Scholar]
  95. Han, J.; Hou, J.; Liu, H.; Cai, S.; Feng, B.; Zhou, J.; Xiang, H. Wide distribution among halophilic archaea of a novel polyhydroxyalkanoate synthase subtype with homology to bacterial type III synthases. Appl. Environ. Microbiol. 2010, 76, 7811–7819. [Google Scholar] [CrossRef]
  96. Chen, G.-Q. Industrial production of PHA. In Plastics from Bacteria: Natural Functions and Applications; Chen, G.-Q., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 14, pp. 121–132. [Google Scholar]
  97. Chanprateep, S. Current trends in biodegradable polyhydroxyalkanoates. J. Biosci. Bioeng. 2010, 110, 621–632. [Google Scholar] [CrossRef]
  98. Poblete-Castro, I.; Becker, J.; Dohnt, K.; dos Santos, V.M.; Wittmann, C. Industrial biotechnology of Pseudomonas putida and related species. Appl. Microbiol. Biotechnol. 2012, 93, 2279–2290. [Google Scholar] [CrossRef]
  99. Greene, E.A.; Voordouw, G. Biodegradation of C5+ hydrocarbons by a mixed bacterial consortium from a C5+-contaminated site. Environ. Technol. 2004, 25, 355–363. [Google Scholar] [CrossRef]
  100. Jung, I.-G.; Park, C.-H. Characteristics of styrene degradation by Rhodococcus pyridinovorans isolated from a biofilter. Chemosphere 2005, 61, 451–456. [Google Scholar] [CrossRef]
  101. Gąszczak, A.; Bartelmus, G.; Greń, I. Kinetics of styrene biodegradation by Pseudomonas sp. E-93486. Appl. Microbiol. Biotechnol. 2012, 93, 565–573. [Google Scholar] [CrossRef]
  102. Ray, A.; Cot, M.; Puzo, G.; Gilleron, M.; Nigou, J. Bacterial cell wall macroamphiphiles: Pathogen-/microbe-associated molecular patterns detected by mammalian innate immune system. Biochimie 2013, 95, 33–42. [Google Scholar] [CrossRef]
  103. Chen, G.-Q.; Wu, Q. The application of polyhydroxyalkanoates as tissue engineering materials. Biomaterials 2005, 26, 6565–6578. [Google Scholar] [CrossRef]
  104. Wampfler, B.; Ramsauer, T.; Rezzonico, S.; Hischier, R.; Kohling, R.; Thony-Meyer, L.; Zinn, M. Isolation and purification of medium chain length poly(3-hydroxyalkanoates) (mcl-PHA) for medical applications using nonchlorinated solvents. Biomacromolecules 2010, 11, 2716–2723. [Google Scholar] [CrossRef]
  105. Karr, D.B.; Waters, J.K.; Emerich, D.W. Analysis of poly-β-hydroxybutyrate in Rhizobium japonicum bacteroids by ion-exclusion high-pressure liquid chromatography and UV detection. Appl. Environ. Microbiol. 1983, 46, 1339–1344. [Google Scholar]
  106. Misaki, A.; Azuma, I.; Yamamura, Y. Structural and immunochemical studies on D-arabino-D-mannans and D-mannans of Mycobacterium tuberculosis and other Mycobacterium species. J. Biochem. 1977, 82, 1759–1770. [Google Scholar]
  107. Nigou, J.; Gilleron, M.; Puzo, G. Lipoarabinomannans: From structure to biosynthesis. Biochimie 2003, 85, 153–166. [Google Scholar] [CrossRef]
  108. Sutcliffe, I.; Brown, A.; Dover, L. The Rhodococcal cell envelope: Composition, organisation and biosynthesis. In Biology of Rhodococcus; Alvarez, H.M., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 16, pp. 29–71. [Google Scholar]
  109. Ruhland, G.J.; Fiedler, F. Occurrence and structure of lipoteichoic acids in the genus Staphylococcus. Arch. Microbiol. 1990, 154, 375–379. [Google Scholar]
  110. Sutcliffe, I.C. The lipoteichoic acids and lipoglycans of Gram-positive bacteria: A chemotaxonomic perspective. Syst. Appl. Microbiol. 1995, 17, 467–480. [Google Scholar] [CrossRef]
  111. Iwasaki, H.; Shimada, A.; Yokoyama, K.; Ito, E. Structure and glycosylation of lipoteichoic acids in Bacillus strains. J. Bacteriol. 1989, 171, 424–429. [Google Scholar]
  112. Sutcliffe, I.C.; Shaw, N. Atypical lipoteichoic acids of Gram-positive bacteria. J. Bacteriol. 1991, 173, 7065–7069. [Google Scholar]
  113. Danson, M.J.; Hough, D.W. The structural basis of protein halophilicity. Comp. Biochem. Physiol. Part. A Physiol. 1997, 117, 307–312. [Google Scholar] [CrossRef]
  114. Chen, G.-Q.; Zhang, G.; Park, S.; Lee, S. Industrial scale production of poly(3-hydroxybutyrate-co-3-hydroxyhexanoate). Appl. Microbiol. Biotechnol. 2001, 57, 50–55. [Google Scholar] [CrossRef]
  115. Choi, J.; Lee, S.Y. Factors affecting the economics of polyhydroxyalkanoate production by bacterial fermentation. Appl. Microbiol. Biotechnol. 1999, 51, 13–21. [Google Scholar] [CrossRef]
  116. Ganduri, V.; Ghosh, S.; Patnaik, P. Mixing control as a device to increase PHB production in batch fermentations with co-cultures of Lactobacillus delbrueckii and Ralstonia eutropha. Process Biochem. 2005, 40, 257–264. [Google Scholar] [CrossRef]
  117. Shi, H.; Shiraishi, M.; Shimizu, K. Metabolic flux analysis for biosynthesis of poly(β-hydroxybutyric acid) in Alcaligenes eutrophus from various carbon sources. J. Ferment. Bioeng. 1997, 84, 579–587. [Google Scholar] [CrossRef]
  118. Tanaka, K.; Katamune, K.; Ishizaki, A. Fermentative production of poly(β-hydroxybutyric acid) from xylose via L-lactate by a two-stage culture method employing Lactococcus lactis IO-1 and Alcaligenes eutrophus. Can. J. Microbiol. 1995, 41, 257–261. [Google Scholar] [CrossRef]
  119. van der Ha, D.; Nachtergaele, L.; Kerckhof, F.-M.; Rameiyanti, D.; Bossier, P.; Verstraete, W.; Boon, N. Conversion of biogas to bioproducts by algae and methane oxidizing bacteria. Environ. Sci. Technol. 2012, 46, 13425–13431. [Google Scholar] [CrossRef]
  120. Romo, D.M.R.; Grosso, M.V.; Solano, N.C.M.; Castaño, D.M. A most effective method for selecting a broad range of short and medium-chain-length polyhidroxyalcanoate producing microorganisms. Electron. J. Biotechnol. 2007, 10, 348–357. [Google Scholar]
  121. Solaiman, D.K.; Ashby, R.D. Rapid genetic characterization of poly(hydroxyalkanoate) synthase and its applications. Biomacromolecules 2005, 6, 532–537. [Google Scholar] [CrossRef]
  122. Spiekermann, P.; Rehm, B.H.; Kalscheuer, R.; Baumeister, D.; Steinbüchel, A. A sensitive, viable-colony staining method using Nile red for direct screening of bacteria that accumulate polyhydroxyalkanoic acids and other lipid storage compounds. Arch. Microbiol. 1999, 171, 73–80. [Google Scholar] [CrossRef]
  123. Melnicki, M.R.; Eroglu, E.; Melis, A. Changes in hydrogen production and polymer accumulation upon sulfur-deprivation in purple photosynthetic bacteria. Int. J. Hydrog. Energy 2009, 34, 6157–6170. [Google Scholar] [CrossRef]
  124. Wu, H.A.; Sheu, D.S.; Lee, C.Y. Rapid differentiation between short-chain-length and medium-chain-length polyhydroxyalkanoate-accumulating bacteria with spectrofluorometry. J. Microbiol. Meth. 2003, 53, 131–135. [Google Scholar] [CrossRef]
  125. Ostle, A.G.; Holt, J. Nile blue A as a fluorescent stain for poly-beta-hydroxybutyrate. Appl. Environ. Microbiol. 1982, 44, 238–241. [Google Scholar]
  126. Law, J.H.; Slepecky, R.A. Assay of poly-β-hydroxybutyric acid. J. Bacteriol. 1961, 82, 33–36. [Google Scholar]
  127. Hong, K.; Sun, S.; Tian, W.; Chen, G.-Q.; Huang, W. A rapid method for detecting bacterial polyhydroxyalkanoates in intact cells by fourier transform infrared spectroscopy. Appl. Microbiol. Biotechnol. 1999, 51, 523–526. [Google Scholar] [CrossRef]
  128. Gumel, A.; Annuar, M.; Heidelberg, T. Effects of carbon substrates on biodegradable polymer composition and stability produced by Delftia tsuruhatensis Bet002 isolated from palm oil mill effluent. Polym. Degrad. Stabil. 2012, 97, 1224–1231. [Google Scholar] [CrossRef]
  129. Hesselmann, R.P.; Fleischmann, T.; Hany, R.; Zehnder, A.J. Determination of polyhydroxyalkanoates in activated sludge by ion chromatographic and enzymatic methods. J. Microbiol. Meth. 1999, 35, 111–119. [Google Scholar] [CrossRef]
  130. Grubelnik, A.; Wiesli, L.; Furrer, P.; Rentsch, D.; Hany, R.; Meyer, V.R. A simple HPLC-MS method for the quantitative determination of the composition of bacterial medium chain-length polyhydroxyalkanoates. J. Sep. Sci. 2008, 31, 1739–1744. [Google Scholar] [CrossRef]
  131. Furrer, P.; Hany, R.; Rentsch, D.; Grubelnik, A.; Ruth, K.; Panke, S.; Zinn, M. Quantitative analysis of bacterial medium-chain-length poly([R]-3-hydroxyalkanoates) by gas chromatography. J. Chromatogr. A 2007, 1143, 199–206. [Google Scholar] [CrossRef]
  132. Tan, G.-Y.A.; Chen, C.-L.; Ge, L.; Li, L.; Wang, L.; Zhao, L.; Mo, Y.; Tan, S.N.; Wang, J.-Y. Enhanced gas chromatography-mass spectrometry method for bacterial polyhydroxyalkanoates (PHAs) analysis. J. Biosci. Bioeng. 2014, 117, 379–382. [Google Scholar] [CrossRef]
  133. Lee, E.Y.; Choi, C.Y. Gas chromatography-mass spectrometric analysis and its application to a screening procedure for novel bacterial polyhydroxyalkanoic acids containing long chain saturated and unsaturated monomers. J. Ferment. Bioeng. 1995, 80, 408–414. [Google Scholar] [CrossRef]
  134. Braunegg, G.; Sonnleitner, B.; Lafferty, R. A rapid gas chromatographic method for the determination of poly-β-hydroxybutyric acid in microbial biomass. Eur. J. Appl. Microbiol. 1978, 6, 29–37. [Google Scholar] [CrossRef]
  135. Ward, A.C.; Dawes, E.A. A disk assay for poly-β-hydroxybutyrate. Anal. Biochem. 1973, 52, 607–613. [Google Scholar] [CrossRef]
  136. Slepecky, R.A.; Law, J.H. A rapid spectrophotometric assay of alpha, beta-unsaturated acids and beta-hydroxy acids. Anal. Chem. 1960, 32, 1697–1699. [Google Scholar] [CrossRef]
  137. Arcos-Hernandez, M.V.; Gurieff, N.; Pratt, S.; Magnusson, P.; Werker, A.; Vargas, A.; Lant, P. Rapid quantification of intracellular PHA using infrared spectroscopy: An application in mixed cultures. J. Biotechnol. 2010, 150, 372–379. [Google Scholar]
  138. de Rijk, T.C.; van de Meer, P.; Eggink, G.; Weusthuis, R.A. Methods for analysis of poly(3-hydroxyalkanoate) (PHA) composition. In Biopolymers Online; Doi, Y., Steinbüchel, A., Eds.; Wiley-VCH: Weinheim, Germany, 2005; Volume 3b, pp. 1–12. [Google Scholar]
  139. Korotkova, N.A.; Ashin, V.; Doronina, N.V.; Trotsenko, Y.A. A new method for quantitative determination of poly-3-hydroxybutyrate and 3-hydroxybutyrate-3-hydroxyvalerate copolymer in microbial biomass by reversed-phase high-performance liquid chromatography. Appl. Biochem. Microbiol. 1997, 33, 302–305. [Google Scholar]
  140. Zhang, S.; Norrlöw, O.; Wawrzynczyk, J.; Dey, E.S. Poly(3-hydroxybutyrate) biosynthesis in the biofilm of Alcaligenes eutrophus, using glucose enzymatically released from pulp fiber sludge. Appl. Environ. Microbiol. 2004, 70, 6776–6782. [Google Scholar] [CrossRef]
  141. Comeau, Y.; Hall, K.J.; Oldham, W.K. Determination of poly-β-hydroxybutyrate and poly-β-hydroxyvalerate in activated sludge by gas-liquid chromatography. Appl. Environ. Microbiol. 1988, 54, 2325–2327. [Google Scholar]
  142. Gumel, A.M.; Annuar, M.S.M.; Heidelberg, T. Biosynthesis and characterization of polyhydroxyalkanoates copolymers produced by Pseudomonas putida Bet001 isolated from palm oil mill effluent. PLoS One 2012, 7, e45214. [Google Scholar]
  143. Kunasundari, B.; Sudesh, K. Isolation and recovery of microbial polyhydroxyalkanoates. Express Polym. Lett. 2011, 5, 620–634. [Google Scholar] [CrossRef]
  144. Ramsay, J.A.; Berger, E.; Voyer, R.; Chavarie, C.; Ramsay, B.A. Extraction of poly-3-hydroxybutyrate using chlorinated solvents. Biotechnol. Tech. 1994, 8, 589–594. [Google Scholar] [CrossRef]
  145. Koller, M.; Bona, R.; Chiellini, E.; Braunegg, G. Extraction of short-chain-length poly-[(R)-hydroxyalkanoates] (scl-PHA) by the “anti-solvent” acetone under elevated temperature and pressure. Biotechnol. Lett. 2013, 35, 1023–1028. [Google Scholar] [CrossRef]
  146. Nonato, R.; Mantelatto, P.; Rossell, C. Integrated production of biodegradable plastic, sugar and ethanol. Appl. Microbiol. Biotechnol. 2001, 57, 1–5. [Google Scholar] [CrossRef]
  147. Berger, E.; Ramsay, B.A.; Ramsay, J.A.; Chavarie, C.; Braunegg, G. PHB recovery by hypochlorite digestion of non-PHB biomass. Biotechnol. Tech. 1989, 3, 227–232. [Google Scholar] [CrossRef]
  148. Hahn, S.K.; Chang, Y.K.; Kim, B.S.; Chang, H.N. Optimization of microbial poly(3-hydroxybutyrate) recover using dispersions of sodium hypochlorite solution and chloroform. Biotechnol. Bioeng. 1994, 44, 256–261. [Google Scholar] [CrossRef]
  149. Kapritchkoff, F.M.; Viotti, A.P.; Alli, R.C.P.; Zuccolo, M.; Pradella, J.G.C.; Maiorano, A.E.; Miranda, E.A.; Bonomi, A. Enzymatic recovery and purification of polyhydroxybutyrate produced by Ralstonia eutropha. J. Biotechnol. 2006, 122, 453–462. [Google Scholar] [CrossRef]
  150. Ashby, R.; Solaiman, D.; Foglia, T. Poly(ethylene glycol)-mediated molar mass control of short-chain- and medium-chain-length poly(hydroxyalkanoates) from Pseudomonas oleovorans. Appl. Microbiol. Biotechnol. 2002, 60, 154–159. [Google Scholar] [CrossRef]
  151. Durner, R.; Zinn, M.; Witholt, B.; Egli, T. Accumulation of poly[(R)-3-hydroxyalkanoates] in Pseudomonas oleovorans during growth in batch and chemostat culture with different carbon sources. Biotechnol. Bioeng. 2000, 72, 278–288. [Google Scholar]
  152. Jiang, X.; Ramsay, J.A.; Ramsay, B.A. Acetone extraction of mcl-PHA from Pseudomonas putida KT2440. J. Microbiol. Meth. 2006, 67, 212–219. [Google Scholar] [CrossRef]
  153. Hahn, S.K.; Chang, Y.K.; Lee, S.Y. Recovery and characterization of poly(3-hydroxybutyric acid) synthesized in Alcaligenes eutrophus and recombinant Escherichia coli. Appl. Environ. Microbiol. 1995, 61, 34–39. [Google Scholar]
  154. Cromwick, A.M.; Foglia, T.; Lenz, R.W. The microbial production of poly(hydroxyalkanoates) from tallow. Appl. Microbiol. Biotechnol. 1996, 46, 464–469. [Google Scholar] [CrossRef]
  155. Ashby, R.D.; Foglia, T.A. Poly(hydroxyalkanoate) biosynthesis from triglyceride substrates. Appl. Microbiol. Biotechnol. 1998, 49, 431–437. [Google Scholar] [CrossRef]
  156. Sánchez, R.J.; Schripsema, J.; da Silva, L.F.; Taciro, M.K.; Pradella, J.G.C.; Gomez, J.G.C. Medium-chain-length polyhydroxyalkanoic acids (PHAmcl) produced by Pseudomonas putida IPT 046 from renewable sources. European Polymer Journal 2003, 39, 1385–1394. [Google Scholar] [CrossRef]
  157. Fernández, D.; Rodríguez, E.; Bassas, M.; Viñas, M.; Solanas, A.M.; Llorens, J.; Marqués, A.M.; Manresa, A. Agro-industrial oily wastes as substrates for PHA production by the new strain Pseudomonas aeruginosa NCIB 40045: Effect of culture conditions. Biochem. Eng. J. 2005, 26, 159–167. [Google Scholar] [CrossRef]
  158. Yasotha, K.; Aroua, M.K.; Ramachandran, K.B.; Tan, I.K.P. Recovery of medium-chain-length polyhydroxyalkanoates (PHAs) through enzymatic digestion treatments and ultrafiltration. Biochem. Eng. J. 2006, 30, 260–268. [Google Scholar] [CrossRef]
  159. De Koning, G.J.M.; Kellerhals, M.; van Meurs, C.; Witholt, B. A process for the recovery of poly(hydroxyalkanoates) from Pseudomonads-Part 2. Bioprocess Eng. 1997, 17, 15–21. [Google Scholar] [CrossRef]
  160. De Koning, G.J.M.; Witholt, B. A process for the recovery of poly(hydroxyalkanoates) from Pseudomonads-Part 1. Bioprocess Eng. 1997, 17, 7–13. [Google Scholar] [CrossRef]
  161. Dai, Y.; Lambert, L.; Yuan, Z.; Keller, J. Characterisation of polyhydroxyalkanoate copolymers with controllable four-monomer composition. J. Biotechnol. 2008, 134, 137–145. [Google Scholar] [CrossRef]
  162. Saeed, K.A.; Ayorinde, F.O.; Eribo, B.E.; Gordon, M.; Collier, L. Characterization of partially transesterified poly(β-hydroxyalkanoate)s using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 1999, 13, 1951–1957. [Google Scholar]
  163. Li, Z.; Yang, X.; Wu, L.; Chen, Z.; Lin, Y.; Xu, K.; Chen, G.-Q. Synthesis, characterization and biocompatibility of biodegradable elastomeric poly(ether-ester urethane)s based on poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) and poly(ethylene glycol) via melting polymerization. J. Biomater. Sci., Polym. Ed. 2009, 20, 1179–1202. [Google Scholar]
  164. Katime, I.; Cadenato, A. Compatibility of peo/poly(iso-butyl methacrylate) and peo/poly(tert-butyl methacrylate) blends by DTA. Mater. Lett. 1995, 22, 303–308. [Google Scholar] [CrossRef]
  165. Shamala, T.; Divyashree, M.; Davis, R.; Kumari, K.L.; Vijayendra, S.; Raj, B. Production and characterization of bacterial polyhydroxyalkanoate copolymers and evaluation of their blends by fourier transform infrared spectroscopy and scanning electron microscopy. Indian J. Microbiol. 2009, 49, 251–258. [Google Scholar] [CrossRef]
  166. Chen, S.; Liu, Q.; Wang, H.; Zhu, B.; Yu, F.; Chen, G.-Q.; Inoue, Y. Polymorphic crystallization of fractionated microbial medium-chain-length polyhydroxyalkanoates. Polymer 2009, 50, 4378–4388. [Google Scholar] [CrossRef]
  167. Wu, C.-S.; Liao, H.-T. The mechanical properties, biocompatibility and biodegradability of chestnut shell fibre and polyhydroxyalkanoate composites. Polym. Degrad. Stabil. 2014, 99, 274–282. [Google Scholar] [CrossRef]
  168. Teeka, J.; Imai, T.; Reungsang, A.; Cheng, X.; Yuliani, E.; Thiantanankul, J.; Poomipuk, N.; Yamaguchi, J.; Jeenanong, A.; Higuchi, T.; et al. Characterization of polyhydroxyalkanoates (PHAs) biosynthesis by isolated Novosphingobium sp. THA_AIK7 using crude glycerol. J. Ind. Microbiol. Biotechnol. 2012, 39, 749–758. [Google Scholar] [CrossRef]
  169. Meng, D.-C.; Shi, Z.-Y.; Wu, L.-P.; Zhou, Q.; Wu, Q.; Chen, J.-C.; Chen, G.-Q. Production and characterization of poly(3-hydroxypropionate-co-4-hydroxybutyrate) with fully controllable structures by recombinant Escherichia coli containing an engineered pathway. Metab. Eng. 2012, 14, 317–324. [Google Scholar] [CrossRef]
  170. Hartmann, R.; Hany, R.; Geiger, T.; Egli, T.; Witholt, B.; Zinn, M. Tailored biosynthesis of olefinic medium-chain-length poly[(R)-3-hydroxyalkanoates] in Pseudomonas putida GPo1 with improved thermal properties. Macromolecules 2004, 37, 6780–6785. [Google Scholar] [CrossRef]
  171. Hany, R.; Hartmann, R.; Böhlen, C.; Brandenberger, S.; Kawada, J.; Löwe, C.; Zinn, M.; Witholt, B.; Marchessault, R.H. Chemical synthesis and characterization of POSS-functionalized poly[3-hydroxyalkanoates]. Polymer 2005, 46, 5025–5031. [Google Scholar] [CrossRef]
  172. Wang, Q.; Tappel, R.C.; Zhu, C.; Nomura, C.T. Development of a new strategy for production of medium-chain-length polyhydroxyalkanoates by recombinant Escherichia coli via inexpensive non-fatty acid feedstocks. Appl. Environ. Microbiol. 2012, 78, 519–527. [Google Scholar] [CrossRef]
  173. De Waard, P.; van der Wal, H.; Huijberts, G.; Eggink, G. Heteronuclear NMR analysis of unsaturated fatty acids in poly(3-hydroxyalkanoates). Study of beta-oxidation in Pseudomonas putida. J. Biol. Chem. 1993, 268, 315–319. [Google Scholar]
  174. Eggink, G.; de Waard, P.; Huijberts, G.N. Formation of novel poly(hydroxyalkanoates) from long-chain fatty acids. Can. J. Microbiol. 1995, 41, 14–21. [Google Scholar] [CrossRef]
  175. Tripathi, L.; Wu, L.-P.; Dechuan, M.; Chen, J.; Wu, Q.; Chen, G.-Q. Pseudomonas putida KT2442 as a platform for the biosynthesis of polyhydroxyalkanoates with adjustable monomer contents and compositions. Bioresour. Technol. 2013, 142, 225–231. [Google Scholar] [CrossRef]
  176. Ballistreri, A.; Garozzo, D.; Giuffrida, M.; Impallomeni, G.; Montaudo, G. Sequencing bacterial poly(β-hydroxybutyrate-co-β-hydroxyvalerate) by partial methanolysis, HPLC fractionation, and fast-atom-bombardment mass spectrometry analysis. Macromolecules 1989, 22, 2107–2111. [Google Scholar] [CrossRef]
  177. Ballistreri, A.; Montaudo, G.; Garozzo, D.; Giuffrida, M.; Montaudo, M.S. Microstructure of bacterial poly(β-hydroxybutyrate-co-β-hydroxyvalerate) by fast atom bombardment mass spectrometry analysis of the partial pyrolysis products. Macromolecules 1991, 24, 1231–1236. [Google Scholar]
  178. Abate, R.; Ballistreri, A.; Montaudo, G.; Garozzo, D.; Impallomeni, G.; Critchley, G.; Tanaka, K. Quantitative applications of matrix-assisted laser desorption/ionization with time-of-flight mass spectrometry: Determination of copolymer composition in bacterial copolyesters. Rapid Commun. Mass Spectrom. 1993, 7, 1033–1036. [Google Scholar] [CrossRef]
  179. Saeed, K.A.; Ayorinde, F.O.; Eribo, B.E.; Gordon, M.; Collier, L. Characterization of partially transesterified poly(beta-hydroxyalkanoate)s by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. J. AOAC Int. 2001, 84, 1109–11015. [Google Scholar]
  180. Chen, G.-Q. Introduction of bacterial plastics PHA, PLA, PBS, PE, PTT, and PPP. In Plastics from Bacteria: Natural Functions and Applications; Chen, G.-Q., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; Volume 14, pp. 1–16. [Google Scholar]
  181. Koller, M.; Bona, R.; Braunegg, G.; Hermann, C.; Horvat, P.; Kroutil, M.; Martinz, J.; Neto, J.; Pereira, L.; Varila, P. Production of polyhydroxyalkanoates from agricultural waste and surplus materials. Biomacromolecules 2005, 6, 561–565. [Google Scholar] [CrossRef]
  182. Li, L. MALDI-MS for polymer characterization. In MALDI MS: A Practical Guide to Instrumentation, Methods, and Applications, 2nd ed.; Hillenkamp, F., Peter-Katalinic, J., Eds.; Wiley-VCH Verlag GmbH & Co. KGaA: Weinheim, Germany, 2013; pp. 313–365. [Google Scholar]
  183. Aoyagi, Y.; Yamashita, K.; Doi, Y. Thermal degradation of poly[(R)-3-hydroxybutyrate], poly[ε-caprolactone], and poly[(S)-lactide]. Polym. Degrad. Stabil. 2001, 76, 53–59. [Google Scholar] [CrossRef]
  184. Bloembergen, S.; Holden, D.A.; Hamer, G.K.; Bluhm, T.L.; Marchessault, R.H. Studies of composition and crystallinity of bacterial poly(β-hydroxybutyrate-co-β-hydroxyvalerate). Macromolecules 1986, 19, 2865–2871. [Google Scholar] [CrossRef]
  185. Porter, M.; Yu, J. Monitoring the in situ crystallization of native biopolyester granules in Ralstonia eutropha via infrared spectroscopy. J. Microbiol. Meth. 2011, 87, 49–55. [Google Scholar] [CrossRef]
  186. Barham, P.J.; Keller, A.; Otun, E.L.; Holmes, P.A. Crystallization and morphology of a bacterial thermoplastic: Poly-3-hydroxybutyrate. J. Mater. Sci. 1984, 19, 2781–2794. [Google Scholar] [CrossRef]
  187. Simon-Colin, C.; Raguénès, G.; Crassous, P.; Moppert, X.; Guezennec, J. A novel mcl-PHA produced on coprah oil by Pseudomonas guezennei biovar. tikehau, isolated from a “kopara” mat of French Polynesia. Int. J. Biol. Macromol. 2008, 43, 176–181. [Google Scholar] [CrossRef]
  188. Cheng, S.-T.; Chen, Z.-F.; Chen, G.-Q. The expression of cross-linked elastin by rabbit blood vessel smooth muscle cells cultured in polyhydroxyalkanoate scaffolds. Biomaterials 2008, 29, 4187–4194. [Google Scholar] [CrossRef]
  189. Dufresne, A.; Kellerhas, M.B.; Witholt, B. Transcrystallization in mcl-PHAs: Cellulose whiskers composites. Macromolecules 1999, 32, 7396–7401. [Google Scholar] [CrossRef]

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MDPI and ACS Style

Tan, G.-Y.A.; Chen, C.-L.; Li, L.; Ge, L.; Wang, L.; Razaad, I.M.N.; Li, Y.; Zhao, L.; Mo, Y.; Wang, J.-Y. Start a Research on Biopolymer Polyhydroxyalkanoate (PHA): A Review. Polymers 2014, 6, 706-754.

AMA Style

Tan G-YA, Chen C-L, Li L, Ge L, Wang L, Razaad IMN, Li Y, Zhao L, Mo Y, Wang J-Y. Start a Research on Biopolymer Polyhydroxyalkanoate (PHA): A Review. Polymers. 2014; 6(3):706-754.

Chicago/Turabian Style

Tan, Giin-Yu Amy, Chia-Lung Chen, Ling Li, Liya Ge, Lin Wang, Indah Mutiara Ningtyas Razaad, Yanhong Li, Lei Zhao, Yu Mo, and Jing-Yuan Wang. 2014. "Start a Research on Biopolymer Polyhydroxyalkanoate (PHA): A Review" Polymers 6, no. 3: 706-754.

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