production and characterization of polyhydroxyalkanoates (phas) from burkholderia cepacia atcc 17759...

8/2/2019 Production and Characterization of Polyhydroxyalkanoates (PHAs) From Burkholderia Cepacia ATCC 17759 Grown o

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PRODUCTION AND CHARACTERIZATION OF POLYHYDROXYALKANOATES (PHAS)

FROMBURKHOLDERIA CEPACIA ATCC 17759 GROWN ON RENEWABLE

FEEDSTOCKS

by

Chengjun Zhu

A dissertation

submitted in partial fulfillment

of the requirements for theDoctor of Philosophy Degree

State University of New YorkCollege of Environmental Science and Forestry

Syracuse, New York

August 2011

Approved: Department of Environmental and Forest Biology

James P. Nakas, Major Professor Emanuel J. Carter, Jr., Chair

Examining Committee

Donald J. Leopold, Department Chair S. Scott Shannon, Dean

The Graduate School


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2011

CopyrightC. J. Zhu

All rights reserved


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iii

Acknowledgements

I would like to thank my major professor, Dr. James P. Nakas, for his advice,

support and knowledge throughout my Ph.D study and with the manuscripts that we

have submitted and will submit for publication. I also would like to thank Dr. Nomura for

his guidance and discussion of our project and his kindness and great help for use and

maintenance of his equipment. Likewise, I am very grateful to Dr. Arthur Stipanovic for

his expert advice and assistance regarding physical-chemical property tests. I am also

grateful to Dr. Patrick Mather, Ms. Erika D. Rodriguez and Mrs. Xinzhu Gu for their

advice and support regarding mechanical property tests of PHAs. I wish to thank Mr.

David Kiemle for assistance with NMR analysis, Mr. Daniel Nicholson and Dr. Kun Cheng

for assistance with the physical characterization of PHAs. I want to thank Mr. David Sgroi,

Ms. Laura Mateya, Ms. Giselle Kathryn Schlegel, Mr Matthew Michael Cleere and Mr.

Sam Kogon for PHA isolation and purification from pilot-plant scale fermentations. I am

grateful to Mr. Joseph K. Gredder, Ms. Anna Elyse Karczewski, etc. for assistance with

the lab-scale research for PHA production. Lastly, I would like to thank my wife Qin for

her support of my research and my son Felix who has brought great joy into my life.

This research was supported by a grant from the New York State Energy Research

and Development Authority (NYSERDA), the Welch Allyn Corp. (Skaneateles, NY), the

Blue Highway LLC. (Syracuse, NY) and Tessy Plastics Corp. (Elbridge, NY).


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iv

Table of Contents

List of Tables..viii

List of Figures..ix

Thesis Abstract..xii

1. Introduction..1

1.1 Microbial polyhydroxyalkanoates..1

1.2 Physicochemical and mechanical properties of PHAs.3

1.3 History and development of PHAs.8

1.4 Metabolic pathway for biosynthesis of PHAs from various carbon sources.13

1.5 Biodegradation and thermal degradation of PHAs.17

1.5.1 Biodegradation of PHAs.18

1.5.2 Thermal degradation of PHAs21

1.6 Considerations and rationale of renewable feedstocks and downstream processing

1.6.1 Renewable feedstocks for PHA production.23

1.6.2 Production, economics and renewability of glycerol and levulinic acid.26

1.6.3 Downstream processing for PHA isolation29

2. Materials and Methods..33

2.1Microorganism and fermentation conditions.33

2.1.1 PHB production in shake flasks and fermentors.33

2.1.2 PHB-co-HV production in shake flasks and fermentors.342.1.3 P3HB -co-4HB production in shake flasks and fermentors..36

2.2Concentration determination of glycerol, xylose, lactose and levulinic acid..37

2.3Isolation and purification of PHAs from biomass.38

2.4Sample preparation with nucleating agents40


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v

2.5GC analysis for PHAs.41

2.6Physicochemical property and mechanical property tests of PHAs42

2.6.1 Molecular mass determination42

2.6.2 Thermal analysis42

2.6.3 Tensile test.43

2.6.4 Nuclear magnetic resonance46

3. Results47

3.1Renewable carbon sources for bacterial growth and PHA production by B. cepacia.47

3.2Biodiesel-derived glycerol as a carbon source for PHA production.49

3.3Production and characterization of PHB homopolymer using glycerol as a carbon

source by B. cepacia.52

3.3.1 Effects of glycerol content on bacterial growth52

3.3.2 Effects of glycerol content on molecular mass of PHB.53

3.3.3 Characterization of PHB end-capped with glycerol by1H NMR.55

3.3.4 Physical properties of PHB..56

3.3.5 Pilot scale (200-L) fermentation using biodiesel-glycerol..57

3.3.6 Injection-molding process for conversion of PHB to biodegradable eartips58

3.3.7 Thermal degradation of PHB..60

3.4 Production and characterization of PHB-co-HV copolymers using glycerol and

levulinic acid as substrates in shake flasks by B cepacia..61

3.4.1 Bacterial growth and production of PHB-co-HV copolymers when co-feeding

glycerol and levulinic acid in shake flasks.61

3.4.2 1H and 13C-NMR analysis for the structure and composition of PHB-co-HV

copolymers62

3.4.3 Physical property test of PHB-co-HV copolymers..65

3.4.4 Mechanical property test of PHB-co-HV copolymers..67


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3.5 Production and characterization of PHB-co-HV copolymers using glycerol and

levulinic acid as substrates in fermentors by B cepacia72

3.5.1 Bacterial growth and PHB-co-HV copolymers when co-feeding glycerol and

levulinic acid in fermentors by B. cepacia in fermentors..72

3.5.2 Crystallization temperatures of PHAs.74

3.5.3 Melting temperatures of PHAs75

3.5.4 Decomposition temperatures of PHAs..77

3.5.5 Decompositon temperature of PHAs with nucleating agents...78

3.5.6 Melting temperature and crystallization temperature of PHAs with

ULTRATALC60979

3.6 Effects of different aging periods on melting temperatures of PHAs.82

3.7 Production of P3HB-co-4HB by B. cepaciausing -butyrolactone/1,4-butanediol83

3.8 Confirmation of HV mol fraction in the PHB-co-HV copolymers by GC analysis and

1H-NMR.85

3.9 Solvent extraction of PHAs..86

3.9.1 Determination of the volume of chloroform for maximum extraction

efficiency 86

3.9.2 Determination of incubation temperature for maximum extractionefficiency.88

3.9.3 Determination of the incubation period for maximum extraction efficiency.89

4 Discussion.91

4.1 Renewable and inexpensive feedstocks for PHA production..91

4.2 Bacterial growth and Properties of PHB produced from glycerol99

4.2.1 Bacterial growth ofB. cepacia using glycerol as a carbon source..99

4.2.2 Properties of PHB produced from glycerol as a carbon source.100


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4.3 Production and characterization of the PHB-co-HV copolymers produced from

biodiesel-derived glycerol and levulinic acid103

4.3.1 Production of the PHB-co-HV copolymers using biodiesel-derived glycerol and

levulinic acid103

4.3.2 Physical properties of the PHB-co-HV copolymers produced from biodiesel-

derived glycerol and levulinic acid104

4.4 Mechanical properties of the PHB homopolymer and the PHB-co-HV copolymer107

4.5 Effects of nucleating agents on physical and mechanical properties of PHAs..111

4.6 Economic considerations of PHA production..112

4.6.1 Economic considerations of PHA production from renewable and inexpensive

feedstocks.113

4.6.2 Economic considerations of PHA production by selective downstream isolation

processes114

5 Conclusions118

6 References.123

7 Appendices134

8 Curriculum Vitae139


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List of Tables

Table 1. Comparison of physicochemical and mechanical properties of selected

polyhydroxyalkanoate (PHA) polymers and petroleum-derived plastics4

Table 2. Physical and mechanical properties of PHB-co-HV copolymers.6

Table 3. The effect of concentrations and exposure times of glycerol on the number-

average molecular weight (Mn) of PHB..54

Table 4. Physical-chemical properties of PHB produced from xylose and glycerol..56

Table 5. Comparison between the original PHB and the thermally degraded PHB..60

Table 6. Mechanical properties of PHB, PHB-co-HV copolymers and polypropylene..71

Table 7. Composition and molecular masses of PHB and copolymers of PHB-co-HV.73

Table 8. Comparison of melting temperatures (Tm) of PHAs detected at different aging

times.82

Table 9. Comparison of HV mol% in the PHB-co-HV copolymers detected by GC and1H

NMR.85

Table 10. Renewable and inexpensive feedstocks used for PHA production.98

Table 11. Comparison of dry cell mass and PHA content from different bacterial strains

grown on various carbon sources in shake flasks..100


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List of Figures

Figure 1. General structure of polyhydroxyalkanoates (a) and copolymers (b) poly-(3-

hydroxybutyrate-co-3-hydroxyvalerate) (abbr. PHB-co-HV) and poly-(3-hydroxybutyrate-

co-4-hydroxybutyrate) (abbr. P3HB-co-4HB)..1

Figure 2. Pathway of PHB biosynthesis.15

Figure 3. Proposed pathway for metabolism of various carbon sources and medium-

chain-length PHA production.16

Figure 4. Thermal degradation of PHB.. 22

Figure 5. Production of biodiesel and glycerol. Catalysts include alkali and acids.27

Figure 6. Route of levulinic acid from lignocellulosic biomass28

Figure 7. Bacterial growth and PHA production using renewable carbon sources (tall

oil fatty acids, biodiesel-derived glycerol and xylose) by B. cepacia in shake flask

experiments48

Figure 8. Bacterial growth and PHB production from various sources of biodiesel-

derived glycerol by B. cepacia in shake flask experiments50

Figure 9. Glycerol consumption during fermentation using pure glycerol, FutureFuel

glycerol, Twin River glycerol and ESF glycerol by B. cepaica in shake flask experiments.51

Figure 10. Dry biomass of B. cepacia grown in shake flasks on different

concentrations of pure glycerol...52

Figure 11. Number-average molecular weight (Mn) and weight-average molecular

weight (Mw) of PHB produced by B.cepacia grown with different concentrations of

glycerol..53

Figure 12.1H-NMR of PHB produced by B .cepacia grown on 7% (v/v) glycerol as a

carbon source. The expanded region indicates glycerol as the terminal end-group55

Figure 13. Changes in biomass, PHA% and glycerol concentration during a fed-batch

fermentation, in a 400-L fermentor, with periodic additions () of biodiesel-glycerol57


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x

List of Figures (Continued)

Figure 14. (a) Purification and vacuum drying process of PHB, (b) Schematic flowsheet

of injection-molding process, (c) Eartips made from polypropylene (black, left) and PHB(brown, right two). 59

Figure 15. Effects of levulinic acid concentration on bacterial growth and production of

PHB-co-HV copolymers61

Figure 16. (a) Chemical shift expansions of 300 MHz1H NMR spectra from PHB-co-HV

copolymers, illustrating a progressive increase in the mol% of HV and quantified by the

integrated areas of the HB doublet (methyl group of HB, 1.27 ppm) and the HV triplet

(methyl group of HV, 0.90 ppm). (b) Chemical shift assignments of 300 MHz13

C NMR

spectrum of PHB-co-35.8 mol% HV, which was produced by B. cepacia using 3% (v/v)

glycerol and 0.9% (w/v) levulinic acid..63

Figure 17a. DSC curves displaying melting temperatures of PHB-co-HV copolymers66

Figure 17b. Melting temperatures and glass transition temperatures of PHB-co-HV

copolymers with increasing HV mol%...............................................................................67

Figure 18a. Stress-strain curves of PHAs and polypropylene. Dot line: PHB (produced

by B. cepacia using xylose as a carbon source); Dash dot line: PHB-co-17.6 mol% HV;

Solid line: PHB-co-17.6 mol% HV; Dash line: polypropylene.69

Figure 18b. Real-time images of stretching for different types of PHAs in tensile testing.

i represents PHB homopolymer, of which the dogbone was pulled for less than 0.7 mmto break; ii displays PHB-co-29.5 mol% HV, of which the dogbone was stretched for

approximately 35 mm to break; iii represents for PHB-co-33 mol% HV, of which the

dogbone was pulled for around 66 mm to break.70

Figure 19. Growth (dry cell mass, DCM, ), PHA yield () and HV content in the

copolymer () produced by B. cepacia using glycerol and levulinic acid as carbon

sources in a 7 L fermentor72

Figure 20. Crystallization temperatures (Tc) of PHB () and copolymers of PHB-co-HV

() with increasing mol% HV..75

Figure 21. Melting temperatures (Tm) of PHB () and copolymers of PHB-co-HV ()

with increasing mol% HV76

Figure 22 Decomposition temperatures (Tdecomp,) and melting temperatures (Tm, )

of PHB and copolymers of PHB-co-HV with increasing mol% HV. Bars indicate

temperature differential (Tdecomp - Tm).77


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xi

List of Figures (Continued)

Figure 23. Effects of nucleating agents on decomposition temperatures of PHB (),

PHB with 5% ULTRATALC609 (

), and PHB with 1% HPN-68L (

).79

Figure 24. Effects of 5% ULTRATALC609 on melting temperature (Tm) of PHAs

containing increasing mol% HV (--PHB,--PHB-co-5.6 mol% HV,--PHB-co-11.4 mol%

HV,--PHB-co-14.7 mol% HV, --PHB-co-17.9 mol% HV,--PHB-co-30.5 mol% HV, --

PHB-co-32.6 mol% HV)...80

Figure 25. Crystallization temperatures of PHAs () and PHAs with 5% ULTRATALC609

(). Bars indicate temperature differentials.81

Figure 26.1H NMR spectrum of P3HB-co-3 mol% 4HB produced from 1,4-butanediol.

Peaks 2,3,4 are typical chemical shifts for P3HB and peaks 6,7,8 (highlighted by the

triangles) represent protons of P4HB84

Figure 27. Relationship between PHA extraction efficiency and dosage of chloroform.

Incubation of the mixture at 55 C overnight..87

Figure 28. Relationship between PHA extraction efficiency and incubation temperature.

Incubation of the mixture at room temp. when stirring or 55 C while standstill.88

Figure 29 Relationship between PHA extraction efficiency and incubation period.

Incubation of the mixture at room temperature when stirring.89


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Abstract

C. J. Zhu. Production and Characterization of Polyhydroxyalkanoates (PHAs) by Burkholderia

cepacia ATCC 17759 Grown on Renewable Feedstocks. 141 Pages, 11 tables, 29 figures, 2011

This thesis describes the microbial production of polyhydroxyalkanoates (PHAs)from bench- and pilot-plant scale fermentations by B. cepacia, followed by physical-

chemical and mechanical characterization of these polyesters. B. cepacia was evaluated

to utilize several renewable and inexpensive feedstocks, e.g. wood hydrolysate,

biodiesel-derived glycerol, cheese whey permeate and tall oil fatty acids from the paper

pulping process. Glycerol was found to be the best carbon source to support bacterial

growth and poly-3-hydroxybutyrate (PHB) production based on the highest dry cell mass

(DCM, 5.8 g/L) and the highest PHB yield (82% of DCM). Increasing the glycerol

concentration from 3% to 9% (v/v) resulted in a gradual reduction of biomass, PHB yield

and molecular mass (Mn and Mw) of PHB.1H-NMR revealed that molecular masses

decreased due to the esterification of PHB with glycerol resulting in chain termination

(end-capping). Supplementing levulinic acid, derived from lignocellulosic materials, with

glycerol led to production of the poly-3-hydroxybutyrate-co-3-hydroxyvalerate (PHB-co-

HV) copolymer. Based on the concentration and timing of levulinic acid added in the

medium, various mol fractions of HV were incorporated to form various PHB-co-HV

copolymers. The copolymers exhibited a typical isodimorphic behavior (V-typed shape),

where upon melting temperature decreased to a minimum point and then increased as

mol% HV increased. Increasing mol% HV in the copolymer resulted in enhanced

mechanical properties. Addition of heterologous nucleating agents improved industrial

processability of PHAs by increasing crystallization temperature. However, HPN-68L was

not used as a nucleating agent for the polyesters isolated in this study because it

decreased the decomposition temperature of PHAs. Production of the PHB homo-polymer and the PHB-co-HV copolymers were successfully scaled up for pilot-plant scale

fermentations. Large quantities of PHAs were isolated, purified and used in the

fabrication of eartips, which are biodegradable and environmentally friendly, through an

injection-molding process. These renewable and inexpensive carbon sources and

alternative downstream PHA isolation process may greatly reduce production cost of

PHAs, by which the market price of PHAs becomes more competitive with that of

conventional petroleum-derived plastics.

Key Words: renewable feedstocks, Burkholderia cepacia, biodiesel-derived glycerol, levulinic

acid, polyhydroxyalkanoates, physical properties, mechanical properties, pilot-plant scale

fermentation, downstream PHA recovery

Author: Chengjun Zhu

Candidate for the degree of Doctor of Philosophy, August 2011

Major professor: James P. Nakas, Ph.D.

Department of Environmental and Forest Biology

State University of New York College of Environmental Science and Forestry, Syracyse, New York

James P. Nakas, Ph.D.


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1. Introduction

1.1 Microbial polyhydroxyalkanoates

Polyhydroxyalkanoates (PHAs) are a class of polyesters which are accumulated as

microbial intracellular carbon and energy reserves. They exist as discrete inclusions in

the cell cytoplasm, typically 0.2-0.5 m in diameter1. A variety of PHAs is commonly

classified into two major groups which are referred to as short-chain-length (scl, 3-5

carbons) and medium-chain-length (mcl, 6-14 carbons)2. The scl-PHAs are semi-

crystalline thermoplastics, whereas mcl-PHAs are more elastomeric3. A large number of

microorganisms, encompassing Gram-positive 4, 5 and Gram-negative 2, 5-7 bacteria, have

been reported to produce various types of PHAs.

m=1, R=H poly-3-hydroxypropionate

R=CH3 poly-3-hydroxybutyrate

R=CH2CH3 poly-3-hydroxyvalerateR=(CH2)xCH3, x=2,3,,11 mcl-poly-3-hydroxyalkanoates

m=2, R=H poly-4-hydroxybutyrate

m=3, R=H poly-5-hydroxyvalerate

Figure 1. General structure of polyhydroxyalkanoates (a) and copolymers (b) poly-(3-

hydroxybutyrate-co-3-hydroxyvalerate) (abbr. PHB-co-HV) and poly-(3-hydroxybutyrate-

co-4-hydroxybutyrate) (abbr. P3HB-co-4HB)

O

On

O

O

m

5

6

7

8

4

3

21

O

O

n O

O

m

a b

O

CH

CH2

C

R O

nm


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Poly-3-hydroxyalkanoates are the most widely studied PHAs, which differ in the

length of side chain (R group depicted in Figure 1a) and can also differ in composition

(e.g. copolymers depicted in Figure 1b, terpolymers, etc.). However, poly-4-

hydroxyalkanoates8, 9

and poly-5-hydroxyalkanoates10

have also been observed in the

past several decades (Figure 1a). Over 150 different monomer units have been

identified as constituents of these storage molecules5, 11, 12

. The large variety of PHA

components results in an enormous diversity of material properties, which is beneficial

for various potential applications. PHAs have recently received increased attention due

to their physical and mechanical properties resembling those of petroleum-derived

plastics. These characteristics make PHAs ideal substitutes for petroleum-based plastics,

especially when global oil prices remain at relatively high levels. Also, PHAs are

completely degraded in the environment to CO2 and H2O13, 14

, compared to recalcitrant,

non-biodegradable conventional plastics, such as polypropylene, polyethylene and

polystyrene. Since PHAs are generally biosynthesized using photosynthetically-based

renewable carbon feedstocks and their end-use materials are biodegradable to carbon

dioxide and water by microbial extracellular PHA depolymerases15, 16

, production and

disposal of these PHA biopolymers constitute a sustainable, closed life cycle process

with much less energy consumption and greenhouse gas emission17-19

.

PHA polymers are synthesized as membrane-bound storage materials by a variety

of microorganisms when exogenous carbon sources are provided in excess and their

growth is impaired by the lack of at least one other nutrient, such as nitrogen, sulfate,

phosphate, magnesium and oxygen6, 20

. Therefore, microbial fermentation for PHA


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production is regulated largely by the concentrations and contents of these nutrients in

the medium. Optimal feeding strategies of these nutrients dramatically enhance

microbial growth and PHA production yields. As reported by Steinbuchel et al.21

, PHAs

are accumulated to levels as high as 90% (w/w) of the dry cell mass.

Furthermore, as a PHA-producing microbe enters the starvation stage due to a

carbon deficiency, PHA polymers will function as intracellular carbon and energy

reserves and initiate the degradation process of PHAs to provide carbon and energy for

microbial survival in a harsh environment22

. Therefore, PHAs, under natural rules of

survival, have been designed and developed by a large number of microorganisms as a

food and energy source during times of nutritional stress.

1.2 Physicochemical and mechanical properties of PHAs

PHAs are highly recommended as an ideal substitute for petroleum-derived

plastics in large part due to their similar material properties (Table 1), in terms of

processability, strength and industrial fabrication into commodity plastic products23, 24

.

Physicochemical and mechanical properties of PHA copolymers can be controlled and

regulated by variation of the mole fractions of the monomeric constituents (Figure 1b),

which could be achieved by careful selection and control of fermentation carbon

sources.


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Poly-3-hydroxybutyrate (PHB) is the most common type of PHA produced by

bacteria. The PHB homopolymer is a highly crystalline, stiff, yet relatively brittle,

material. Correspondingly, as shown in Table 1, PHB exhibits high tensile strength (43

MPa), but low elongation to break (5%)25

. The copolymer poly-3-hydroxybutyrate-co-20

mol%-3-hydroxyvalerate (PHB-co-20 mol% HV) exhibits lower crystallinity, less stiffness

(20 MPa), but higher elasticity and flexibility (50% elongation to break)25

compared to

the homopolymer PHB. Incorporation of different monomeric subunits, such as 4-

hydroxybutyrate (4HB)25

, 3-hydroxyhexanoate (HHx)25

and other mcl-

hydroxyalkanoates26

(e.g. 3-hydroxyoctanoate [3HO], 3-hydroxydecanoate [3HD], 3-

hydroxydodecanoate [3HDD]), with 3HB will result in copolymers with varying material

Table 1 Comparison of physicochemical and mechanical properties of selected

polyhydroxyalkanoate (PHA) polymers and petroleum-derived plastics25

PolymersCrystallinity

a

(%)

Tmb

(C)

Tgc

(C)

Tensile

strength

(MPa)

Elongation

to break

(%)

PHB 60 177 4 43 5

PHB-co-20%HV 56 145 -1 20 50

PHB-co-16%4HB 45 150 -7 26 444

PHB-co-10%HHx 34 127 -1 21 400

Polypropylene

50-70 176 -10 38 400

Polyethylene (LDPE)d

20-50 130 -36 10 620

Notes:a degree of crystallinity b melting temperature c glass transition temperatured LDPE:low density polyethylene


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properties to address numerous applications in medical and engineering fields.

Compared to commercially available polylactic acid (PLA) which is also renewable and

biodegradable27

, diverse combinations of PHA monomeric units are superior to PLA,

which only consists of a single monomer-lactic acid.

In addition to regulating material properties by controlling the composition of

PHAs, it is possible to change material properties of PHAs with the same composition by

incorporating various fractions of the co-monomer in the copolymers. This thesis mainly

describes the production and characterization of poly-3-hydroxybutyrate-co-3-

hydroxyvalerate (PHB-co-HV) copolymers, biosynthesized from glycerol and levulinic

acid as carbon sources. As the mol fraction of HV in the copolymer PHB-co-HV varies,

the physical and mechanical properties of PHB-co-HV will correspondingly change (Table

2). When HV ratios of PHB-co-HV increased from zero (the homopolymer of PHB) to 25%,

the melting temperatures (Tm) and glass transition temperatures (Tg) gradually

decreased from 179 C to 137 C and 10 C to -6 C, respectively. As shown in Table 2,

PHB-co-HV copolymers also became more flexible (as indicated by the decrease of

Youngs modulus) and tougher (as demonstrated by the increase in impact strength) as

the HV ratio increased.

Besides PHB-co-HV, similar patterns were observed with copolymers PHB-co-

HHx

28-30

, P3HB-co-4HB

20

, PHB-co-HO

31

, PHB-co-HD

31

, when the ratios of these co-

monomers, such as HHx, 4HB, HO or HD, in their copolymers increased.


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Table 2 Physical and mechanical properties of PHB-co-HV copolymers20

.

Mol fraction

(mol%) Tm (C) Tg (C)YoungsModulus

(GPa)

TensileStrength

(MPa)

Notched IzodImpact

Strength (J/m)3HB 3HV

100 0 179 10 3.5 40 50

97 3 170 8 2.9 38 60

91 9 162 6 1.9 37 95

86 14 150 4 1.5 35 120

80 20 145 -1 1.2 32 200

75 25 137 -6 0.7 30 400

PHB-co-HV copolymers with various HV mol fractions exhibit high degrees of

crystallinity (between 55% and 70%)32

. PHB-co-HV is unique among the PHA family of

copolymers in that the size and structure of HB and HV monomers are similar. Their

similarity allows HB and HV to participate in a co-crystallization process, in which HV can

be incorporated into the HB crystal lattice and vice versa. This phenomenon is termed

isodimorphism32, 33

. As a result, the melting temperatures of PHB-co-HV gradually

decrease to a minimum point, then increase as the HV mol fraction increases. Therefore,

isodimorphism and the transition between the HB crystal lattice and the HV crystal

lattice typically demonstrate a V-shaped pattern (see melting temperatures of PHB-co-

HV in results and references32, 33

). Incorporation of comonomers with PHB to form


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copolymers can lower melting temperatures, by which PHA copolymers demonstrate an

important advantage of industrial applications for melt processing at lower

temperatures.

However, the PHA family displays a rather slow crystallization process due to high

purity and limited heterogeneous nuclei34

. Basically, isolation of PHAs from cells using

solvents excludes most of the impurities from cell components. During the melt-

quenching process, PHAs at high purity crystallize at a relatively low rate, which causes a

longer manufacturing cycle and a less efficient industrial fabrication process for finished

products. PHB and PHB-co-HV have been extensively studied for their nucleation

behaviors34, 35

. Pure PHB exhibited a very slow nucleation, though self seeding to some

extent, when it was cooled from a melt34

. The nucleation density of pure PHB is too low

to initiate efficient crystallization. At the same time, limited nuclei formed limited

quantities of spherulites so that the size of each PHB spherulite is relatively large, which

makes PHB somewhat brittle and subject to cracking36, 37. PHB-co-HV also exhibited a

slow crystallization behavior, which resulted in the films made from the copolymers

with a higher HV ratio tacky and even sticky to themselves after cooling for an extended

period of time35, 38

.

Impurities usually behave as external nuclei for PHAs. They can increase not only

the rate of crystallization, but also the quantities of spherulites corresponding to the

numbers of nuclei. The large quantities of nuclei make the size of spherulites relatively

small, thus improving material mechanical properties36

. Accounting for the slow

crystallization process of PHB and PHB-co-HV, heterogeneous nucleating agents instead


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8

of impurities, which are difficult for the quantification and qualification, need to be

supplemented with these polymer melts to speed up the crystallization process by

increasing the crystallization temperature and forming small spherulites.

Various nucleating agents, such as orotic acid39

, -cyclodextrin40

, boron nitride,

talc, terbium oxide, lanthanum oxide35

, saccharin and phthalimide41

have been tested

for enhancing the crystallization of PHAs. These nucleating agents will generally increase

crystallization temperatures, accelerate crystallization rates and enhance PHA stability

during heating. However, some nucleating agents have negative effects on PHAs by

decreasing their decomposition temperatures (Tdecomp.). Hydroxyapatite, which is a

naturally occurring mineral of calcium apatite, decreased the onset Tdecomp from 260oC

to 225oC when the hydroxyapatite content in the hybrid PHB-hydroxyapatite composite

increased from 0 to 10%42

. These side effects of nucleating agents have not been

described in many previous studies. Comprehensive physical property tests, including

crystallization temperature, melting temperature, glass transition temperature and

decomposition temperature, have been performed in this research (see results).

1.3 History and development of PHAs

Poly-3-hydroxybutyrate, as an intracellular storage material in Bacillus megaterium,

was first identified by Lemoigne in 192643

. At that time, PHB was discovered

unexpectedly when Lemoigne attempted to determine the cause of acidification in an

aqueous suspension of the bacterium Bacillus megaterium under oxygen-free conditions.


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9

Although methods for PHB detection and quantification were limited, Lemoigne and co-

workers published 27 papers from 1923 to 1951. Not until the late 1950s did microbial

physiologists finally recognize the importance of PHB in the overall metabolism of

bacterial cells44

.

The rediscovery of PHB occurred simultaneously and was reported independently

in 1958 and 1959. Williamson et al. 45 at the University of Edinburgh in Scotland treated

the cells of various Bacillus spp. with an alkaline solution of sodium hypochlorite, and

found large amounts of PHB (89%) with the ether-soluble lipid (11%) in some Bacillus

species, and elucidated the function of PHB in the cell. Doudoroff and Stanier46

at the

University of California at Berkeley discovered that PHB was the primary product of the

oxidative and photosynthetic assimilation of organic compounds by Pseudomonas

saccharophila and a phototropic bacterium, Rhodospirillum rubrum, respectively. When

external carbon sources were removed from the medium, there occurred a fairly rapid

intracellular breakdown of the storage PHB. Therefore, PHB was putatively thought to

play a role analogous to that of starch and glycogen in the metabolism of other

organisms. Doudoroff et al.46

attempted to clarify the biosynthesis and degradation

mechanism of PHB in microbial cells. In the 1960s, these aforementioned authors47, 48

isolated native PHB granules from a chemoheterotroph and phototroph, Bacillus

megaterium and Rhodospirillum rubrum. The native PHB granules from R. rubrum

retained the active PHB synthase and depolymerase that degrade PHB to the monomer,

and the isolated granules from B. megaterium only exhibited the synthase associated

with PHB. These authors also determined that PHB granules in bacteria actually serve as


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10

an intracellular energy and carbon reserve and that PHB is produced in response to a

nutrient limitation when at least one essential element in the environment is exhausted.

In 1965, Marchessault and his co-workers

49

at SUNY-ESF collected PHB samples

from various microorganisms and employed X-ray diffraction to explore the crystal

structure of PHB. All PHB samples precipitated from chloroform displayed an uniformity

of crystal structure and molecular structure due to identical X-ray diffractograms and

infrared spectra, respectively. The molecular mass of PHB isolated by alkaline

hypochlorite was also found to be uniformly low compared to that of PHB prepared by

direct solvent extraction.

In 1974, Wallen and Rohwedder50

reported that the polyesters, isolated from

activated sewage sludge, exhibited similar but not identical NMR and infrared spectra.

Gas chromatographic analysis indicated a mixture of C4, C5, C6 and C7 components in

these polyesters. The identification of a variety of PHAs in addition to PHB opened

avenues of research for PHAs regarding their material properties.

Since rising oil prices and an unstable oil supply became global issues and

environmental concerns over limited fossil reserves drew more and more attention, the

advantage of PHAs as biodegradable alternatives to petroleum-derived plastics was

gradually recognized by researchers and much effort has been devoted to investigation

of PHA properties. In addition, much time and effort has been dedicated to genetic

engineering of microorganisms or plants for biosynthesis of PHAs from renewable

feedstocks with improved material properties for different end-use applications.


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Prompted by the oil crisis in the 1970s, the first industrial production of PHB was

introduced in 1982 by Imperial Chemical Industries (ICI) in the UK, employing a two-

stage, fed-batch fermentation process and using a sugar-based carbon source for

growth of Ralstonia eutropha. However, PHB was produced at a relatively high

production cost due to expensive feedstocks and downstream processing for PHB

isolation. Compared to conventional petroleum-derived plastics, higher cost has placed

PHAs in a weaker economic position and mechanical properties, such as high

crystallinity and brittleness, have resulted in a rather limited range of applications20

.

Based on these limitations of the homopolymer PHB, PHB-co-HV copolymer, which

exhibits enhanced toughness and flexibility, attracted more attention and was first

produced on a commercial scale in the late 1980s by ICI, which marketed their PHB-co-

HV products under the trade name of Biopol6, 51

. Biopol (PHB-co-HV copolymer) was

produced by R. eutropha, using glucose and propionate as carbon sources, at a reported

production cost of US$ 16/kg, which was 18-fold higher than conventional

polypropylene52

. The prohibitively high price of PHAs and the relatively low prices of

commodity plastics made the commercial-scale production of PHAs unrealistic.

Eventually Biopol was acquired by Zeneca Ltd in 1990 until it was acquired in 1996 by

Monsanto (St. Louis, MO), which utilized their considerable expertise in plants and

initiated research to produce PHB and related copolymers photosynthetically in plants51.

Subsequently, Monsantos right to Biopol was sold to Metabolix (Cambridge, MA)

in 200153

. Recombinant Escherichia colistrains, which exhibit broad nutritional diversity


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12

and relatively fast growth, have been engineered to produce PHAs from inexpensive

carbon sources. Mirel (trademark of Telles, an affiliated company of Metabolix and

Archer Daniels Midland Company) bioplastics were announced to be produced using

corn starch sugars, starting in 2009 in Clinton, IA (www.mirelplastics.com). Several other

companies are also developing and commercializing various types of PHAs, including

Procter & Gamble (Cincinnati, USA) which introduced novel PHA copolymers with HB as

a monomeric unit and HHx (C6), HO (C8) or HD (C10) etc. as monomeric units, all of which

are under the trademark ofNodax54, 55

. The projections for Nodax marketing consist

of PHA production of 100-1000 metric ton in 2004 and the delivery plan of approximate

$ 1/lb ($ 2.2/kg) in 2005/200656

. Since the 1980s, BASF in Germany developed a pilot-

scale fermentation process for production of PHB and PHB-co-HV, which were

supplemented and blended with its biodegradable polymer Ecoflex, which are

petroleum-derived aliphatic-aromatic copolyesters57

. Tianan Biologic in China started to

produce PHB-co-HV in 2000 using glucose, fructose and organic acids as carbon sources

(www.tianan-enmat.com), and have currently scaled up to commercial production of

2000 metric tons per year57

.

Most of the above-mentioned commercial PHAs have been produced by various

microorganisms using pure carbon sources, such as glucose, sucrose, propionate and

lauric acid, which are always more expensive than renewable feedstocks, including

wood-based hydrolysate (xylose and levulinic acid)58

, cheese whey permeate (including

lactose)59

, sugarcane molasses (sucrose, fructose, glucose and trace amount of

maltose)60, 61

, corn steep liquor (nitrogen sources)60

, soybean oil62

, and crude glycerol


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13

from biodiesel-producing facilities63

. Regarding environmental concerns of

commercializing PHAs, the production of PHAs from renewable and biobased feedstocks,

e.g. wood hydrolysate and biodiesel-derived glycerol, need to be taken into account at a

higher priority.

1.4 Metabolic pathway leading to the biosynthesis of PHAs

from various carbon sources

Uptake of carbon sources by microorganisms supports their growth and other

basic vital functions. Carbon sources in the environment have to be translocated into

the cytosol inside the cell, the location for nutrient metabolism. In this translocation

process, transporters, which are transmembrane proteins, either actively transport the

molecules of carbon sources by utilizing ATP (adenosine triphosphate) energy (e.g ATP-

binding cassette transporters, ABC-transporters) or passively translocate these

molecules without energy expenditure (e.g glycerol diffusion facilitator protein, GlpF).

Therefore, whether a certain carbon source can be utilized for microbial growth is first

determined by the transporters which can recognize and transport this molecule.

Relevant to this research, xylose (a five carbon sugar) and/or glycerol were used as

carbon sources by Burkholderia cepacia. D-xylose is transported by XylFGH transporter

(possibly belonging to the ABC superfamily) into the cytosol and converted to D-xylulose

by xylose isomerase (encoded by xylA gene), subsequently phosphorylated to D-

xylulose-5-phosphate by xylulokinase (encode by xylB gene)64, 65

(Figure 3). Glycerol is


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14

translocated by GlpF into the cell and phosphorylated by glycerol kinase (encoded by

glpK), leading to glycerol-3-phosphate. Glycerol-3-phosphate dehydrogenase (encoded

by glpD) oxidizes glycerol-3-phosphate to dihydroxyacetone phosphate (DHAP)66

(Figure

3). Further metabolism of these intermediates (e.g. D-xylulose-5-phoaphate and DHAP)

derived from the carbon sources through either glycolysis (Embden-Meyerhofpathway)

or the pentose phosphate pathway or Entner-Doudoroff pathway results in the end

product pyruvate, which is converted through oxidative decarboxylation by pyruvate

dehydrogenase to acetyl-CoA67

.

In this research, levulinic acid (4-ketovaleric acid) was employed as the co-

substrate to provide the precursor propionyl-CoA for biosynthesis of poly-3-

hydroxyvalerate. Metabolism of levulinic acid is not clearly elucidated, but tentatively

considered to form one propionyl-CoA and one acetyl-CoA through -oxidation. The

enzymes involved are still ill-defined21

.

Three key enzymes -ketothiolase, acetoacetyl-CoA reductase and PHA synthase,

encoded by phaA, phaB and phaC, respectively, are involved in the last three steps of

PHA biosynthesis. Two acetyl-CoA moieties are condensed to acetoacetyl-CoA by -

ketothiolase, and acetoacetyl-CoA is reduced to (R)-3-hydroxybutyryl-CoA by

acetoacetyl-CoA reductase, followed by polymerizing the precursor 3-hydroxybutyryl-

CoA to the polymer poly-3-hydroxybutyrate68

. Similarly for poly-3-hydroxyvalerate

synthesis, one acetyl-CoA and one propionyl-CoA are condensed to 3-ketovaleryl-CoA,


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15

with subsequent reduction to 3-hydroxyvaleryl-CoA, followed by polymerization for PHV

synthesis (Figure 2).

acetyl-CoA

acetoacetyl-CoA

(R)-3-hydroxybutyryl-CoA

PHB

3-ketothiolase (PhaA)

acetoacetyl-CoA reductase (PhaB)

PHA synthase (PhaC)

CoASH

NADP+

NADPH

SCoA

O

SCoA

O O

SCoA

OH O

CoASH

O

O

n

2

Figure 2 Pathway of PHB biosynthesis

Biosynthesis of medium-chain-length PHAs, which are comprised of the

constituents C6-C14 chains, recruits PHA-specific enzymes such as (R)-specific enoyl-CoA

hydratase (PhaJ), putatively (R)-3-hydroxyacyl ACP thioesterase (PhaG) and acyl-CoA

ligase (AlkK) (Personal communication with Qin Wang and Christopher Nomura at SUNY-

ESF) to divert the intermediates (such as enoyl-CoA and (R)-3-hydroxyacyl acyl carrier

protein (ACP), respectively) of fatty acid -oxidation pathway and fatty acid biosynthesis

pathway for the formation of precursor (R)-3-hydroxyacyl-CoA (Figure 3), which leads to

biosynthesis of medium-chain-length PHAs by PHA synthase (PhaC)69

.


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Figure 3 Proposed pathway for metabolism of various carbon sources and medium-

chain-length PHA production69

. GAP: glyceraldehyde-3-phospahte. DHAP:

dihydroxyacetone phosphate. (1) or (2) represents one or more steps in glycolysis

pathway or pentose phosphate pathway, respectively. : cell membrane

While the pathway for PHA biosynthesis has been gradually elucidated in the last

several decades, construction of desired strains for PHA production seems to be a

promising path and has greatly stimulated and enhanced PHA research. These

engineered strains, such as E. coli70

, Pseudomonas putida71

and Cupriavidus necator62

(formerly known as Ralstonia eutropha or Alcaligenes eutrophus) demonstrate the

advantages of broad nutritional diversity (using various carbon sources, including

inexpensive feedstocks), relatively rapid growth to high cell density, accumulation of

high intracellular concentrations of PHAs and biosynthesis of novel PHAs which cannot

be accomplished in native strains and which exhibit diverse material properties for

various end-use applications.

1.5 Biodegradation and thermal degradation of PHAs

Although this thesis focuses on the production and characterization of PHAs using

inexpensive carbon sources, some consideration must be given to the biodegradability

of PHAs, which is a distinct advantage of PHAs over conventional plastics. Also, since

some applications of PHAs need to be conducted at high temperature, it is reasonable

to examine the thermal degradation of PHAs.


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1.5.1 Biodegradation of PHAs

Petroleum-derived plastics are xenobiotic compounds, which are recalcitrant to

degradation and take several decades, even over 100 years, to degrade in the natural

environment72

. One of the most important characteristics of PHAs as substitutes for

conventional plastics is that PHAs are biodegradable. In nature, a vast consortium of

microorganisms, by using intracellular or extracellular PHA depolymerases, will degrade

PHAs, which either are stored inside of the cells or exist in the natural environment. It is

noteworthy that intracellular PHA depolymerases cannot hydrolyze extracellular PHAs

and extracellular PHA depolymerases are also unable to degrade intracellular PHA

granules16

. Apparently, these differences result from the biophysical structures of

intracellular native PHA granules and extracellular denatured PHAs. The former are

completely amorphous elastomers73

, however, the latter are known for their high

crystallinities2. Native PHA granules with a particular surface layer containing protein

and phospholipids are denatured by losing this surface layer during the isolation

processes of PHAs, leading to semicrystalline polymers that exhibit an ordered helical

crystal structure, in which the remaining amorphous polymers are embedded13

.

In the remainder of this thesis, biodegradability of PHAs, if mentioned, will be

concerned with the extracellular denatured PHAs using extracellular depolymerases in

order to demonstrate the superiority of PHAs over petroleum-based plastics.

Volova et al74

studied the biodegradability patterns of PHB and PHB-co-11 mol%HV

in a tropical marine environment. After 160 days, the loss of mass of both PHA films,


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submerged to a depth of 120 cm, approached 50% of the original dry weight of the PHA

films. Due to a larger surface area, PHA films demonstrated a more rapid rate of

biodegradation than compacted PHA pellets. The polydispersity increasing in all PHA

samples suggested that the fragments of polymers with more diverse lengths were

growing due to random scission of the hydrolyzed polymer chains. Under their study

conditions, no significant differences were observed for the degradation rates between

PHB and PHB-co-11 mol%HV, and the degree of crystallinity of both PHAs remained

unchanged.

Mergaert et al.75

tested microbial degradation of PHB and PHB-co-10 mol% HV in

soils at 15 C, 28 C or 44 C for up to 200 days. These dog bone-shaped PHA samples

were degraded at an erosion rate of 0.03% to 0.64% weight loss per day, depending on

the polymer compositions, the types of soil and the incubation temperatures. In

summary, the degradation was enhanced by incubation at higher temperatures, and in

most cases the copolymer exhibited a higher erosion rate, based on weight loss, than

the homopolymer. The degradation also resulted in the loss of mechanical properties

(based on less elongation to break). In the studied soil (sandy soil, pH 6.5; clay soil, pH

7.1, loamy soil, pH 6.3; hardwood forest soil, pH 3.9; pinewood forest soil, pH 3.5), 295

dominant microbial strains capable of degrading PHBand PHB-co-HV in vitro were

isolated and identified. This same research team76 investigated PHB, PHB-co-10 mol%

HV and PHB-co-20 mol% HV in situ in natural waters. These polymers were degraded

rather slowly (less than 7% weight loss after 6 months) in two freshwater ponds.

However, after 358 days in a freshwater canal, 34% weight loss was recorded for PHB,


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77% for PHB-co-10 mol% HV, and 100% for PHB-co-20 mol% HV. In seawater, within 270

days, PHB lost 31% of the initial weight and the copolymers lost between 49-52%. The

degradation rate was observed to be more rapid during the summer due to higher

temperature. Mergaert et al.77

also studied biodegradation of PHB, PHB-co-10 mol% HV

and PHB-co-20 mol% HV in compost. PHB-co-20 mol% HV was degraded much faster (70%

weight loss) than PHB (6% weight loss) and PHB-co-10 mol% HV (4% weight loss) within

150 days. From this biodegradation study, as well as two other composts, 109 microbial

strains capable of degrading PHAs in vitro were isolated and identified.

Although hundreds of microorganisms are able to secrete extracellular PHA

depolymerases to degrade PHAs in the natural environment, the biodegradation rates of

PHAs varied dramatically from several months up to several years as a function of PHA

composition and shapes of the test samples, and environmental factors, such as

temperature, types of soil and water, sunlight (UV radiation)78

, pH16

, etc..

In a word, the biodegradability of PHAs in the natural environment makes PHA-

type thermoplastics ideal green substitutes for conventional plastics, which are resistant

to degradation in the environment and therefore result in severe environmental

pollution on a global scale.


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1.5.2 Thermal degradation of PHAs

Considering the injection molding process for industrial fabrication of finished

products (packaging, eartips, thermometer covers, medical implants, etc.), thermal

degradation of PHAs must be taken into account during the heating process due to their

potential thermal instability79

.

PHB was extensively studied during melt processing and found to be rather

unstable at temperatures above or even close to its melting temperature (177 C). Doi

and his co-workers80

confirmed that PHB polyester suffered increasing reduction of

molecular mass within 20 min at 175 C, even 2 C below its melting temperature. In

addition, all copolyester samples (PHB-co-HV, HV=0-71 mol%; P3HB-co-4HB, 4HB=0-82

mol%) tested were thermally unstable at temperatures above 170 C, based on rapid

loss of molecular masses. Below or equal to 160 C, all copolymer samples tested

demonstrated thermal stability and their molecular masses decreased at a very slow

rate within 20 min. Especially for the first 2 min of heating at 180 C , most of PHA

polymers did not show significant decrease in molecular masses. Therefore, PHAs are

suggested to be heated to melt and kept at that temperature for a short time period (2

min suggested in the study by Kunioka et al.80

).

It was reported that thermal degradation of PHB occurred by random chain

scission, a widely accepted six-membered ring ester decomposition process, yielding

crotonic acid (one product from PHB pyrolysis, Figure 4), 2-pentenoic acid (one product

from PHV pyrolysis) and other related oligomers81-83

.


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O CH

R1

H2C

CH3

C

O

H

CH

CH

OCH3

C

O C

O

R2

O CH

R1

H2C

CH3

C

OH

O

+

HC

HC

CH3

C HC

CH

C R2

O

OH

+

H

CH

CH

CH3

CH3

O

Crotonic acid

pyrolysis

Figure 4 Thermal degradation of PHB

A major concern of PHB and other related PHAs for use as a thermoplastic is

thermal sensitivity during melt processing82. A potential solution is grafting of certain

compounds with PHAs. It has been reported that none of the conventional polyolefin

stabilizers enhanced PHB stability84

. However, radiation grafting of methyl methacrylate

(MMA)85

, 2-hydroxyethyl methacrylate (HEMA)85

, acrylic acid (AAc)86

and styrene87, 88

onto PHB and its copolymers was found to enhance the thermal stability of PHAs. Also,

grafting maleic anhydride onto PHB remarkably enhanced its thermal decomposition

temperature79

. In most cases, a low degree of grafting not only enhanced the thermal

stability of PHAs, but also promoted the biodegradability of PHAs due to the wettability


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23

between the polymer and the enzyme solutions. It was also noted that a high degree of

grafting steeply decreased the biodegradability of PHAs.

Therefore, it is necessary to strictly control the temperature to melt PHAs during

heating. Because of the loss of molecular mass of PHAs, mechanical properties of PHAs

definitely change after heating. In addition, a low degree of grafting helps stabilize PHAs

when the processing temperature reaches the melting temperature.

1.6 Considerations and rationale of renewable feedstocks

and downstream processing

Compared to the price of conventional petrochemical plastics (less than US $ 1/kg),

the price of PHAs was prohibitively high (ca. US $ 16/kg Biopol PHAs)89

, which places

PHAs at a distinct disadvantage in the market place. By examining the entire flowsheet

of PHA production, the high price of PHAs is driven by high production costs of PHAs in

two major areas. One is the cost of feedstocks, and secondly, the downstream recovery

process of PHAs.

1.6.1 Renewable feedstocks for PHA production

A large number of microorganisms have been shown to produce various types of

PHAs. Although it is feasible to use pure carbon sources, such as glucose, galactose,

xylose, fatty acids, etc., for PHA production in the laboratory, high production costs for

carbon feedstocks hinder the scale-up of PHA production towards the commercial level.


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Consequently, the cost of carbon sources accounts for as much as 50% of the overall

production costs89, 90

. In order to reduce the production costs of PHAs, renewable and

inexpensive feedstocks have to be taken into consideration to substitute for pure and

expensive carbon sources at the commercial level.

As reviewed in 2009 by Chen57

, the microbial production of PHAs at the industrial

scale still uses some high purity sugars and fatty acids, such as glucose, sucrose, lauric

acid, and propionate, during the fermentation process. Although the price of PHAs is

expected to be approximately US $3-4/kg in the near future, it is obvious that PHAs are

still more expensive than conventional plastics, especially when the price of oil is

depressed.

In order to further reduce the production costs of PHAs, some progress has been

achieved in the last several years by using inexpensive agricultural or industrial wastes

for PHA production.

Khardenavis et al.91

evaluated waste activated sludge generated from a combined

dairy and food processing industry wastewater treatment plant for PHB production.

Jowar or rice grain-based distillery spent wash was used as a carbon source for a PHB

yield of 42.3% (w/w) and 40% (w/w), respectively. Addition of di-ammonium hydrogen

phosphate further increased PHB production to 67% (w/w). Meanwhile, mixed culture

production of PHAs from waste water was evaluated to be financially attractive in

comparison to pure culture production of PHAs92

.


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Gouda et al.60

used sugarcane molasses and corn steep liquor, two of the most

inexpensive substrates available in Egypt, as sole carbon and nitrogen sources for PHB

production. The best growth of Bacillus megaterium was obtained with 3% molasses,

while the maximum yield of PHB at 46.2% (w/w) occurred with 2% molasses. Corn steep

liquor with a concentration equivalent to 0.05% NH4Cl was the best nitrogen source for

PHB synthesis (32.7%, w/w).

Yellore et al.59

isolated a strain ofMethylobacterium sp. ZP24 from a local pond,

and tested this strain by using lactose from cheese whey, a byproduct of the dairy

industry, for PHB production. Pure lactose at a concentration of 12 g/L resulted in

biomass and PHB yield of 5.25 g/L and 59% (w/w) in 40 h, respectively. Cheese whey,

used as a carbon source, led to 1.1 g/L PHB and further addition of ammonium sulphate

increased PHB production from whey 2.5-fold. Nath et al.93

employed a fed batch

process to attain a PHB yield of 4.58-fold using limiting dissolved oxygen in the

fermentor with processed cheese whey supplemented with ammonium sulfate.

Keenan et al.58

used aspen and maple wood hydrolysate, which contains xylose

and glucose as major sugar components and low amounts of galactose, arabinose and

mannose, supplemented with levulinic acid for PHB-co-HV production. These detoxified

hydrolysates amended with 0.25%-0.5% levulinic acid were used as feedstocks, and

resulted in PHB-co-HV yields, PHA% of dry cell mass and mol fraction of HV at 2.0 g/L, 40%

(w/w), and 16 mol% - 52 mol%, respectively. From an economic standpoint, the

substrate cost of hemicellulosic hydrolysate was reduced to US $ 0.34/kg PHB,


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compared to US $ 0.58/kg PHB from hydrolyzed corn starch and US $ 1.30/kg PHB from

pure glucose.

1.6.2 Production, economics and renewability of glycerol and

levulinic acid

In this research, waste glycerol, a byproduct from the biodiesel-producing process

(Figure 5), was tested as a carbon source for PHA production. Biodiesel-derived glycerol

has been produced in huge quantity as the production of biodiesel has significantly

increased. Biodiesel production increased dramatically from 500,000 gallons in 1999 to

450 million gallons in 2007 (National Biodiesel Board, 2008). The major byproduct of the

biodiesel industry is glycerol, which is product of approximately 10% of the final weight

of biodiesel94

. Consequently, in 2007, glycerol was produced in a quantity of 45 million

gallons in the United States, and this crude glycerol is not suitable for use in the food,

pharmaceutical, cosmetics and other industries due to low purity. It is expensive to

refine crude glycerol to the purity needed for these applications95

. Biodiesel production,

as well as glycerol, are at an all-time high. Since the market is glutted, the price of

glycerol has decreased and raw glycerol from biodiesel-producing companies is now at

US $ 2.5 cents/lb96

. Therefore, conversion of crude glycerol into higher-value products,

such as PHAs, improves the economic viability of the biofuel industry by producing a

value-added product as well as eliminating the cost of treatment for glycerol disposal.


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O

O

O

R1

R2

R3

O

O

O

Alkali,Acids

OH

OH

O

R1

O

R2

O

R3

OH +

O

O

O

Triacylglycerol GlycerolBiodiesel

(Fatty acid methyl ester)

Methanol

Figure 5 Production of biodiesel and glycerol. Catalysts include alkali and acids.

Besides glycerol, levulinic acid was employed as a cosubstrate for PHB-co-HV

copolymer production by Burkholderia cepacia in this research. Levulinic acid (4-

ketovaleric acid, 4-oxopentanoic acid), an important biomass-derived feedstock, could

be produced cost-effectively from a wide-array of renewable, hexose-containing

materials, including forest-based lignocellulosic biomass97, 98

. Hexoses, such as glucose,

fructose and mannose, from forest residues are converted under acid dehydration to 5-

hydroxymethylfurfural (HMF) and subsequently hydrated resulting in the final products

of levulinic acid and formic acid (Figure 6).


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O

HO

HOOH

OH

OH

OHO

O

OH

O

O

H2O H2O

+H OH

O

Cellulose,Hemicellulose

Glucose 5-hydroxymethylfurfural,HMF

levulinic acid formic acid

Figure 6 Route of levulinic acid from lignocellulosic biomass

Levulinic acid, in addition to glycerol, is also among the top 12 value-added

chemicals from biomass described by the US DOE99

. It is especially attractive because a

variety of lignocellulosic biomass, such as rice straw, wood, pulp slurry, corn starch,

switch grass, sugarcane, etc., can be used for the direct production of this

thermodynamically stable molecule100. The dehydration of C6 sugars by acids was

reported to generate levulinic acid with a theoretical maximum yield of 64.5% (w/w)

because of the concurrent production of equimolar amounts of formic acid101

.

Several technologies have been developed for the industrial-scale continuous

production of levulinic acid, with yields reported between 20% and 48% (w/w)100

. The

most promising technology proposed for the large-scale production of levulinic acid was

patented by the Biofine Corporation (South Glens Falls, NY)102, 103

. This approach used a

double-reactor system and minimized the formation of byproducts. The carbohydrate

feedstocks and sulfuric acid catalyst (1-5 wt% of the feedstocks) were mixed at 210-

230 C for a short time of 13 to 25 s in the first reactor, in which C 5 and C6 sugars are


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produced. Meanwhile, dehydration of C6 sugars in the first reactor produces HMF.

Subsequently, HMF is continuously removed from the first reactor and fed into the

second reactor at 195-215 C for 15 to 30 min for a final conversion to levulinic acid at a

yield of 50%-70% (w/w). As a result, levulinic acid can be produced at a low cost (US

$ 0.04-0.1/kg)104

, which depends on the scale of production and the economic climate of

the cellulosic feedstocks.

Besides levulinic aicd which can be obtained from lignocellulosic biomass, tall oil

fatty acids, mainly consisting of C18 (i.e. 52% oleic acid and 45% linoleic acid58

), are

byproducts of the paper/Kraft pulping process, which is a technology for conversion of

wood into almost pure cellulose fibers. These free fatty acids are isolated at a

percentage of 30-40% from crude tall oil by distillation105, 106

and could be potential

inexpensive precursors for forest-based PHA production107, 108

.

1.6.3 Downstream processing for PHA isolation

In addition to the production costs contributed by feedstocks, another major cost

contributor is the recovery process of PHAs. PHAs are known to be stored inside cells

and the native granules are surrounded by phospholipid membranes. However, only

purified PHA polymers exhibit the desired physical and mechanical properties of

thermoplastics. Therefore, removal of other cell components from PHAs presents a

technical challenge and this isolation process serves to increase the production cost of

PHAs.


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30

Solvent extraction is the most common PHA recovery technique. Usually,

halogenated solvents, such as chloroform and dichloromethane, display good solubility

of PHAs and are widely used in the laboratory. In terms of high efficiency (around 90%)

and high purity (over 99%), solvent extraction exhibits undoubted advantages over

other recovery methods109

. Meanwhile, this method can also remove bacterial

endotoxins and causes negligible degradation of PHAs. Unfortunately, solvent extraction

is not viewed as an environmentally benign method, and high costs of these solvents

also hamper large-scale application in the industry. In addition to the solvents for

dissolution of PHAs, anti-solvents need to be used for precipitation of PHAs from the

solvents. Methanol, ethanol and heptane are usually employed to precipitate and purify

PHAs in this extraction process, in which these solvents might contribute up to 50% of

the entire production costs of PHAs110

and are not commercially economical if used at

an industrial scale for PHA production. However, some non-chlorinated solvents could

be acceptable alternatives in terms of environmental and economic concerns. Non-

chlorinated solvents, including ethyl acetate, acetone (hot), cyclic carbonate, methyl

tertiary butyl ether (MTBE), etc., can be used to partly reduce costs and water could also

be used as an inexpensive anti-solvent to precipitate PHAs110, 111

.

Besides solvent extraction, another strategy for PHA isolation is to remove non-

PHA cell mass (NPCM) from PHAs and keep PHAs in the solid state during the isolation

process. In this regard, cell disruptions are necessary and can be performed in three

ways: chemical digestion, enzyme disruption and mechanical disruption.


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Chemical agents, such as NaOH, hypochlorite and surfactants, can destroy the cells

and dissolve cell components into the supernatant, followed by centrifugation or

filtration to collect PHA pellets112

. However, this method is a non-selective process and

polymer degradation occurs simultaneously. Compared to solvent extraction, the purity

of PHAs isolated by chemical digestion is lower and varies from 68% to 98% based on

the digestion conditions and the types of microorganisms109, 111

. A combination of

heating, H2SO4, NaOH and sodium hypochlorite at proper concentrations and duration

could improve the quality of PHAs with purity higher than 97% and a recovery yield

higher than 95%113

.

Enzymatic disruption, employing proteolytic enzymes, will break cells by hydrolysis

of proteins through cleavage of peptide bonds. The proteases help to lyse the cells

efficiently at 50 C and pH 9.0 and release PHAs from the cell at a purity of 88.8%.

Supplemented with chloroform or sodium dodecyl sulphate-ethylenediaminetetraacetic

acid (SDS-EDTA), higher purity was achieved with a recovery yield of 90%109, 111

.

Nevertheless, the high cost of enzymes and complexity of the recovery process

outweigh advantages of this method.

Mechanical disruption, using a bead mill, high-pressure homogenization (including

microfluidization) or supercritical fluid (SCF), is favored mainly due to the elimination of

harsh chemicals, which minimizes environmental effects. The recovery yield of this

process can reach 89%111

. Pretreatments using SDS or NaOH help to further purify PHAs.


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The major drawbacks of this method are high capital investment cost and long

processing time.

Currently, a green, low-cost, highly efficient, and environmentally friendly PHA

recovery process has not generally been accepted or implemented. Therefore, based on

the characteristics and requirements for the end use of PHAs, a combination of several

aforementioned recovery methods might be beneficial to reduce the production costs

of PHAs.


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2. Materials and Methods

2.1 Microorganism and fermentation conditions

2.1.1 PHB production in shake flasks and fermentors

Burkholderia cepacia ATCC 17759 was used in both shake flask as well as pilot

scale (200 L) fermentations. The nitrogen-limited mineral salts medium used in the

fermentations was initially described by Bertrand et al.114

. The recipe for this medium

was slightly changed in our experiments as follows: 1.5 g of (NH4)2SO4, 3.6 g of Na2HPO4 ,

1.5 g of KH2PO4, 75.5 mg of CaCl2, 60 mg of NH4-Fe(III) citrate, 200 mg of MgSO4 7H2O,

and 1 ml of trace elements solution per liter. The trace elements solution contained 100

mg of ZnSO4 7H20, 30 mg of MnCl2 4H20, 300 mg of H3BO3, 200 mg of CoCl2 6H20, 20

mg of CuSO4 5H20, 20 mg of NiCl2 6H20, 30 mg of NaMoO4 2H20 per liter.

For shake-flask experiments, all cultures were shaken at 30C and 150 rpm.

Glycerol (99.5%, EMD) and xylose (99% purity, Acros) were used to produce PHB for

physical- chemical characterizations and were autoclaved separately in a solution of 50%

(v/v) or 50% (w/v), respectively. All shake flask experiments were conducted in 500 mL

baffled flasks containing 100 mL of medium with metal enclosures

In experiments utilizing both xylose and glycerol as carbon sources, xylose (2.2%)

was initially added into the medium. At 24 h or 48 h, 2% or 5% glycerol was added to the

medium, and cultures were harvested at 72 h. Only 2.2% xylose was used as a carbon

source for a control. When using the 400 L fermentor (Model No: IF 400, New Brunswick

Scientific Co. Inc., NewBrunswick, NJ), a fed-batch method was used and glycerol (85%


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purity) from a biodiesel-producing facility (Twin Rivers Technologies, Quincy, Mass.) was

added as the primary carbon source for production of the homopolymer PHB.

Tall oil fatty acids (MWV L-1A, MeadWestvaco Corporation, Richmond, VA) were

autoclaved as received. Two milliliters of tall oil fatty acids were added as a carbon

source into the 100 ml medium for bacterial growth and PHA production.

Lactose (>99%, Sigma-Aldrich) was prepared as a stock solution at 20% (w/v) and

sterilized separately. Cheese whey permeate (15%-20%, w/v) from Crowley Foods, a

local subsidiary of HP Hood LLC in Arkport, NY, was adjusted to pH 7.0 and filtered by

Whatman paper to remove the precipitate, followed by filter sterilization using Nalgene

disposable filter units with 0.22 m PES membrane (ThermoFisher Scientific Inc.).

The purities of other biodiesel-glycerol sources (from FutureFuel Corporation,

Clayton, MO and SUNY-ESF biodiesel-producing facility, Syracuse, NY) were 93% and

40%, respectively. The ESF crude glycerol was adjusted to neutral pH before sterilization.

All sources of biodiesel-derived glycerol were autoclaved at 121 C for 20 min before use

in fermentation experiments.

2.1.2 PHB-co-HV production in shake flasks and fermentors

PHB-co-HV copolymers were produced by B. cepacia ATCC 17759 in shake flasks, a

7 L fermentor or a 400 L fermentor. The mineral salts-trace elements medium described

previously was employed in these experiments, except for further addition of


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ammonium sulfate to 10 g/L in the fermentor experiments. Reagent-grade glycerol

(99.5%, EMD, Gibbstown, NJ) and levulinic acid (98%, Alfa Aesar, Ward Hill, MA) were

used as carbon sources for bacterial growth and polymer production. Glycerol and

levulinic acid were dissolved into ddH2O for a stock solution of 50% (v/v) and 50% (w/v),

respectively. The stock solution of levulinic acid was adjusted to pH 7.0 before

autoclaving. In shake flask experiments, Fernbach flasks (2800 ml in volume) were

employed for copolymer production. Each Fernbach flask was prepared with 500 ml

mineral salts medium and 3% (v/v) glycerol. Levulinic acid at a concentration of 0.1%

(w/v) was added, except for the control group (only containing glycerol as a carbon

source), when inoculated with a 5% (v/v) seed culture. At 24 h, the remaining levulinic

acid at concentrations of 0.2%, 0.4%, 0.6% or 0.8% were added into the medium, to

achieve final concentrations of levulinic acid in the medium of 0.3%, 0.5%, 0.7% and

0.9%, respectively, along with the control group without levulinic acid and the group

with only 0.1% levulinic aicd. All Fernbach experiments were incubated at 30 C and

shaken at 150 rpm for 72 h.

Copolymers of PHB-co-HV, with a ratio of HV between 5% and 32.6%, were

produced by B. cepacia in a 7-L fermentor (BIOFLO410, New Brunswick Scientific Co.,

Edison, NJ) with a working volume of 5 L. The concentration of glycerol was kept

between 1% and 3% (v/v) during the fermentation and pure levulinic acid was

continuously pumped into the fermentor at the rate of 0.5 g/Lh. Dissolved oxygen (DO)

and pH were maintained at 40% and 7, respectively, by adjusting agitation/aeration and

pumping 10 M sodium hydroxide. For this study, two stages of the fermentation were


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employed: 1) growth ofB. cepacia to as high an OD540 (optical density) as possible with

sufficient glycerol; 2) initiating levulinic acid addition at the point of highest OD540 for HV

production and providing additional glycerol for bacterial growth and PHB production.

Scaling up to pilot-plant level, copolymers were produced in a 400 L fermentor

(Model No: IF 400, New Brunswick Scientific Co.,Inc. NewBrunswick, NJ), using a fed-

batch method and glycerol (85% purity), from a biodiesel-producing facility (Twin Rivers

Technologies, Quincy, MA), was added as the primary carbon source and levulinic aicd

( 97% purity, Sigma-Aldrich) was added periodically at concentrations of approximately

1% (v/v) whenever the concentration of levulinic acisd was determined by HPLC to be

less than 0.2%.

2.1.3 P3HB-co-4HB production in shake flasks and

fermentors

P3HB-co-4HB copolymers were produced by B. cepacia ATCC 17759 in either shake

flasks or a 7-L fermentor. In shake flask experiments, 0.1%, 0.3%, 0.5%, 0.7% and 0.9%

(v/v) of 1,4-butanediol (99% ReagentPlus, Sigma-Aldrich) or -butyrolactone ( 99%

ReagentPlus, Sigma-Aldrich) was co-fed with 3% (w/v) xylose or 3% (v/v) glycerol as

major carbon sources in mineral-salts medium, described previously in 2.1.1. After

inoculation with a 5% (v/v) seed culture shaken for 48 h, all flasks were shaken at 30 C

and 150 rpm for 72 h. In a 7 L fermentor, Luria-Broth (LB) medium, containing 10 g/L

BactoTM

tryptone (BD, Sparks, MD), 5 g/L yeast extract (Sensient Technologies


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Corporation, Junean, WI) and 5 g/L sodium chloride (EM Science, Gibbstown, NJ), was

used for bacterial growth, supplemented with 1,4-butanediol or -butyrolactone at a

total concentration of 2% (v/v) for 72 h at 30 C with 30% dissolved oxygen.

Cell harvest and polymer isolation followed the procedure described in section 2.3.

2.2 Concentration determination of glycerol, xylose, levulinic

acid and lactose

Glycerol concentration was measured using the Free Glycerol Reagent kit (Catalog

No. F6428, Sigma, St,Louis, MO), basically containing glycerol kinase, glycerol phosphate

oxidase, peroxidase, ATP, 4-aminoantipyrine (4-AAP) and sodium N-ethyl-N-(3-

sulfopropyl) m-anisidine (ESPA). The reactions were incubated for 5 min at 37C and the

absorbances were recorded spectrophotometrically at 540 nm (CARY 300

Spectrophotometer, Varian Inc. USA) as described in details per the instructions of the

manufacturer (http://www.sigmaaldrich.com/etc/medialib/docs/Sigma/Bulletin/f6428

bul.Par.0001.File.tmp/f6428bul.pdf). Briefly, the Free Glycerol Reagent was pipetted at a

volume of 0.8 ml into each cuvet. Water, Glycerol Standard (Catalog No. G7793, Sigma),

and sample were added at a volume of 10 l to each cuvet labeled as blank, standard,

and sample, respectively. The solution in each cuvet was mixed by gentle shake and all

samples tested were incubated at 37 C for 5 minutes, and the absorbances at 540 nm

were recorded of Blank, Standard, and Sample versus water as reference.


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Note: the glycerol standard solution concentration is 0.26 mg glycerol/ml, with an

equivalent triolein concentration of 2.5 mg/ml.

Sugars, including xylose and lactose, and organic acids, including levulinic acid,

were analyzed by HPLC (Waters Corporation, Milford, MA) equipped with a Waters 717

plus autosampler, a 2996 photodiode array detector, a 2414 refractive index detector

and a 1525 binary HPLC pump. For sugar analysis, a carbohydrate analysis column

(WAT084038, 3.9 x 300 mm, Waters, Milford, MA) was employed with a mobile phase

containing 20% Milli-Q water and 80% acetonitrile which had been filtered through a

0.22 m membrane before use. A Jordi Gel DVB organic acid column (Cat. 17001, 250

mm x 10 mm, The Nest Group, Inc., Southborough, MA) was used to determine the

concentration of levulinic acid. The mobile phase (containing 0.02 M phosphoric

acid/acetonitrile/methanol at a ratio of 90/5/5) was run at 1 ml/min and the spectrum

was recorded at 214 nm. Empower Pro software (Waters, Milford, MA) was used to

collect and analyze data.

2.3 Isolation and purification of PHAs from biomass

Cultures were harvested from shake flasks by centrifugation at 7000 x g for 10 min

(Sorvall model SS-3). The biomass was subsequently washed with distilled H2O, if


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necessary, using additional methanol (10 ml CH3OH per 100 ml culture) for removal of

residual long chain fatty acids, and centrifuged again to remove the supernatant and

stored at -20C until lyophilization. The biomass could be washed up to three times for

further removal of residual glycerol, which could interfere with biomass drying after

lyophilization. For cultures grown in the 400 L fermentor, the broth was centrifuged at

16,000 rpm using a continuous flow centrifuge (CEPA High Speed Centrifuge Z81 G, New

Brunswick Scientific, USA).

The harvested biomass was then frozen and lyophilized at -80C and 200 millitorr

for 24 h. The dry biomass was subsequently ground to a powder with a mortar and

pestle, mixed with chloroform (10 ml chloroform / 1 g dry biomass), and either stirred at

room temperature for 24 h for the biomass from the 400 L fermentor or placed in an

incubator at 55-60 C overnight for the biomass from shake flask experiments and 7 L

fermentor runs.

For the purification of bench-scale quantities of PHAs, the general procedure of

Keenan et al.115

was used for all biomass samples. In general, after PHA extraction, the

mixture of biomass and chloroform was filtered through Whatman #1 filter paper

seated in a porcelain Buchner funnel apparatus to remove cell debris. Subsequently, the

PHA-containing filtrate solution was decanted into cold methanol or ethanol (at 4 C) by

a ratio of 1: 10 (v/v) during stirring to precipitate PHAs. The powder-like PHAs were

collected by filtration through a Pyrex glass Buchner funnel with a fritted disc (fine-

grade porosity with a maximum pore size of 4-5.5 m).


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For the extraction and purification of polymers from cells produced at the pilot-

plant scale, lyophilized cell mass (1 kg) was stirred with 9 L of chloroform in a 10-L Pyrex

glass container at room temperature for 24 h. This mixture was filtered as described

previously for bench-scale PHA extraction. The polymer solution was concentrated by

rotary evaporation using a Flash Evaporator (Buchler Instruments, Fort Lee, NJ) at 70C

to a highly viscous solution and a distillation system (Buchler Instruments, Fort Lee, NJ)

was used to recover chloroform. The polymer was precipitated from this solution via

slow addition to cold methanol (1 volume of PHA solution / 10 volumes of methanol).

Precipitated polymer samples exhibited a fibrous, noodle-shaped morphology and, if

necessary, were washed with additional methanol for further purification. After the

large noodle-shaped PHA polymers were screened and removed directly by a wire sieve,

the remaining floc of PHAs was filtered through a Pyrex glass Buchner funnel with a

fritted disc (coarse-grade porosity with a pore size of 40-60 m).

All purified PHA samples were air dried in glass petri dishes in a hood overnight,

followed by vacuum drying in a desiccator until use.

2.4 Sample preparation with nucleating agents

The purified homopolymer PHB and various copolymers of PHB-co-HV (between

0.5 g and 1 g) were di

production and characterization of polyhydroxyalkanoates (phas) from burkholderia cepacia atcc 17759...

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