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5-2018
Investigations of Polyhydroxyalkanoate Secretion and Production Using Sustainable Carbon Sources
Chad L. Nielsen Utah State University
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This Thesis is brought to you for free and open access by the Graduate Studies at DigitalCommons@USU. It has been accepted for inclusion in All Graduate Theses and Dissertations by an authorized administrator of DigitalCommons@USU. For more information, please contact [email protected]. INVESTIGATIONS OF POLYHYDROXYALKANOATE SECRETION AND
PRODUCTION USING SUSTAINABLE CARBON SOURCES
by
Chad L. Nielsen
A thesis submitted in partial fulfillment of the requirements for the degree
of
MASTER OF SCIENCE
in
Biological Engineering
Approved:
______Charles D. Miller, Ph.D. Ronald C. Sims, Ph.D. Major Professor Committee Member
______Randolph V. Lewis, Ph. D Mark McLellan, Ph.D. Committee Member Vice President for Research and Dean of the School of Graduate Studies
UTAH STATE UNIVERSITY Logan, Utah
2018 ii
Copyright © Chad L. Nielsen 2018
All Rights Reserved iii
ABSTRACT
Investigations of Polyhydroxyalkanoate Secretion and Production Using Sustainable
Carbon Sources
by
Chad L. Nielsen, Master of Science
Utah State University, 2018
Major Professor: Dr. Charles D. Miller
Department: Biological Engineering
Polyhydroxyalkanoates (PHAs) are promising bioplastic polymers with the potential to replace petroleum-derived plastics in diverse applications. Production costs for PHAs are still very high, however, largely due to the cost of carbon feedstocks for the bacteria that produce the polymers and the cost of extracting the polymers after production. Waste streams that could be diverted to PHA production as inexpensive carbon sources include food wastes and methanol. Secretion-based recovery of PHA
granules was studied in the methylotrophic bacteria Methylobacterium to decrease the
cost of downstream processing of PHAs, while using methanol as the sole carbon source.
The hemolysin type I secretion pathway from Escherichia coli was transformed into a novel isolate of Methylobacterium. A synthetic biology approach was used to create a
phasin fusion protein that binds to the surface of PHA granules and the hemolysin iv
secretion signal peptide. This genetic construct was used in conjunction with the secretion system with the intent of inducing the cell to secrete PHA into the extracellular media. As currently constituted, however, the secretion system made no significant difference in the amount of PHA produced and secreted by the Methylobacterium isolate.
The information gathered in this work can be further optimized and applied to other methylotrophic bacteria to reduce costs in PHA manufacturing systems.
(198 pages)
v
PUBLIC ABSTRACT
Investigations of Polyhydroxyalkanoate Secretion and Production
Using Sustainable Carbon Sources
Chad L. Nielsen
Polyhydroxyalkanoates (PHAs) are a type of biologically-produced plastic known for their biocompatibility and biodegradability. They have the potential to replace petroleum-based plastics as an environmentally-friendly alternative. This is beneficial because the release of plastics into environments such as the ocean and the buildup of plastics in landfills are major concerns facing society today. Currently, however, PHAs are significantly more expensive than their petroleum-based counterparts. This is largely due to the cost of carbon sources and of extracting the bioplastics from bacteria. The goal of these studies was to examine replacing traditional carbon sources used in PHA production like sugar and oils with sustainable carbon sources and to improve extraction procedures by inducing secretion of PHAs in bacteria.
A few sustainable carbon sources were examined for use in PHA production.
First, studies focused on the conversion of food waste into PHAs were reviewed. It was shown that utilizing food wastes as carbon sources may be a viable approach to producing PHAs. A second carbon source examined was methanol. A novel isolate of
Methylobacterium that demonstrated the ability to produce PHAs from methanol was identified. A system of secreting PHAs that was constructed using synthetic biological engineering approach was introduced to this isolate. This secretion system was not shown to improve extraction of PHAs in Methylobacterium in its current form. vi
ACKNOWLEDGMENTS
This research would not have been possible without the help of many individuals.
First, I am grateful to my wife, Elise, for her continued support and encouragement. I am also grateful to my parents for their interest and support as I have pursued a graduate degree in biological engineering. My gratitude goes to Dr. Charles Miller for his guidance, mentoring, and patience throughout this process. I would also like to thank Dr.
Asif Rahman for his mentoring, support, advice, and contributions that helped make this research possible. My gratitude is also due to Dr. Ronald Sims for taking me under his wing as a research assistant many years ago, and continuing to provide support throughout my time at Utah State University. This research is also a result of contributions and input made by Dr. Asad-ur-Rehman, Dr. Marie K. Walsh, and Dr.
Randy Lewis. In addition, this research was made possible by the Research Catalyst grant program of the Utah State University Office of Research and Graduate Studies. Lastly, I would like to thank my fellow students who have worked with me and kept me company as I have worked and studied in the lab, including Alaric Siddoway, Anna Doloman,
Gabby Nielson, Alex Beeston, AJ Walters, James VanderMeyden, Emily Jesgarz, Sierra
Julander, and Timothy Kerns. Their friendship and support have made this process more enjoyable than it otherwise would have been.
Chad L. Nielsen
vii
CONTENTS
Contents
ABSTRACT ...... iii
PUBLIC ABSTRACT ...... v
ACKNOWLEDGMENTS ...... vi
CONTENTS ...... vii
LIST OF TABLES ...... x
LIST OF FIGURES ...... xi
LIST OF SYMBOLS, NOTATIONS, AND DEFINITIONS ...... xiii
INTRODUCTION ...... 1
1.1 Overview ...... 1 1.2 Hypotheses ...... 6 1.3 Objectives ...... 7 1.4 Thesis Outline ...... 7 References ...... 8
FOOD WASTE CONVERSION TO MICROBIAL POLYHYDROXYALKANOATES ...... 11
2.1 Summary ...... 11 2.2 Introduction ...... 12 2.3 PHAs from food waste using pure cultures ...... 15
2.3.1 PHAs from whey ...... 16 2.3.2 PHAs from starch ...... 18 viii
2.3.3 PHAs from waste oil...... 18 2.3.4 PHAs from spent coffee grounds...... 19 2.3.5 Lignocellulosic waste conversion to PHAs ...... 20 2.3.6 PHAs from sugar industry waste ...... 21 2.3.7 PHAs from legume waste ...... 23
2.4 Recombinant microbes for PHA production ...... 26 2.5 Production of PHAs from food waste using anaerobic digestion ...... 31 2.6 Indirect coupling of MMC to PHA production ...... 35 2.7 Conclusions and outlook ...... 37 References ...... 39
LITERATURE REVIEW ...... 56
3.1 Methanotrophic Bacteria ...... 56 3.2 PHA Biosynthesis in Methylotrophic Bacteria ...... 63 3.3 Current Methods for PHA Production in Methylotrophic Bacteria ...... 68 3.4 Current Methods for PHA Recovery ...... 75 3.5 Genetic Engineering in Methylobacterium species ...... 80 3.6 Recombinant Proteins ...... 82
3.6.1 HlyA Secretion Pathway ...... 82 3.6.2 Phasins ...... 86
References ...... 89
AN INVESTIGATION OF PHASIN TRANSLOCATION AND POLYHYDROXYALKANOATE PRODUCTION IN A NOVEL ISOALTE OF METHYLOBACTERIUM ...... 107
4.1 Abstract ...... 107
Background ...... 107 Results ...... 107
ix
4.2 Background ...... 108 4.3 Materials and Methods ...... 113
4.3.1 Strains and Plasmids ...... 113 4.3.2 BioBrick Design and Assembly ...... 114 4.3.3 Media Formulation and Growth Conditions ...... 116 4.3.4 PHA Production in Methylobacterium ...... 117 4.3.5 Recovery of Secreted PHA ...... 117 4.3.6 PHA Concentration Determination ...... 118 4.3.7 Scanning Electron Microscopy (SEM) ...... 118 4.3.6 Fluorescence Studies ...... 119
4.4 Results ...... 119
4.4.1 Isolation and Analysis of Methylobacterium sp...... 119 4.4.2 BioBrick Design and Construction ...... 122 4.4.3 Promoter strength analysis...... 125 4.4.4 PHB Production Analysis ...... 127
4.5 Conclusion ...... 131 References ...... 131
GENERAL CONCLUSIONS ...... 138
ENGINEERING SIGNIFICANCE AND FUTURE RESEARCH ...... 140
APPENDICES ...... 153
BIOBRICK PARTS AND SEQUENCES ...... 153 PROTOCOLS, REAGENTS, AND MISCELLANEOUS ...... 157 COPYRIGHT PERMISSION FORMS ...... 161 POLYHYDROXYBUTYRATE PRODUCTION USING A WASTEWATER MICROALGAE BASED MEDIA ...... 167
x
LIST OF TABLES
Table Page
2.1 Production of PHAs from food waste using pure microbial cultures……… 24
2.2 Production of PHAs from food waste using recombinant bacteria………… 31
2.3 Production of PHAs from anaerobically digested food waste……………... 35
2.4 Production of PHAs from organic acids derived from anaerobically 37
digested food waste…………………………………………………………
3.1 Summary of the methylotrophic and methanotrophic production of PHB… 67
4.1 Strains and plasmids used in this study…………………………………….. 114
4.2 Oligonucleotides…………………………………………………………… 116
4.3 Concentrations of PHB in cultures of Methylobacterium with nitrogen 122
starvation, nitrogen and magnesium starvation, and normal (unstressed)
conditions…………………………………………………………………...
4.4 Production of PHB in secreting (pCMHS3 + pLG575) and non-secreting 128
(pCMHS1) strains of Methylobacterium at 24 h……………………………
6.1 Advantages and disadvantages of different food wastes in PHA production 142
A.1 Nucleotide sequences of codon optimized protein coding regions for Ms. 153
trichosporium OB3b………………………………………………………..
A.2 Nucleotide sequences of Hemolysin Genes used in this study…………….. 155
D.1 Production and yields of PHB after 48 h from Escherichia coli…………… 178
xi
LIST OF FIGURES
Figures Page
1.1 Biorefinery proposed by the Sustainable Waste-to-Bioproducts 6
Engineering Center (SWBEC) at Utah State University……......
2.1 Conversion of food waste into PHAs using PHA-producing microbes…… 15
3.1 Pathways for the oxidation of methane and assimilation of formaldehyde.. 58
3.2 RuMP pathway for formaldehyde fixation……………………...... 59
3.3 Serine pathway for formaldehyde fixation…………………………...... 60
3.4 Chemical structure of PHAs……………………………………………...... 63
3.5 Dedicated pathway for P(3HB) monomer synthesis……………………..... 64
3.6 Purification strategy of PHAs……………………………………………… 75
3.7 Diagram of the type I secretion system (T1SS) and the closely-related 83
resistance-nodulated-division (RND) family of multidrux efflux pumps….
3.8 Scheme of the structure of PHA granules…………………………………. 87
4.1 Schematic for PHA secretion……………………………………………… 111
4.2 Diagram of the BioBrick standard assembly process……………………… 113
4.3 Schematic of PHA and sfGFP secretion genetic circuits………………….. 115
4.4 Cell Pellet of Methylobacterium strain……………………………………. 120
4.5 Phylogenetic tree showing the relationship of the Methylobacterium 121
isolate used in the current study to other methylotrophic bacteria…………
4.6 SEM images of Methylobacterium in various sizes of clusters……………. 121 xii
4.7 Image of Methylobacterium stained with Sudan Black to demonstrate the 122
presence of PHB……………………………………………………………
4.8 Schematic of the third contiguous region of the Ms. trichosporium OB3b 123
genome with locations of the four phasin proteins indicated………………
4.9 Gel electrophoresis depicting PhaP4MtO gene isolation……………………. 124
4.10 Gel electrophoresis depicting PhaP1Re genetic circuit…………………….. 125
4.11 Schematic of PHA and sfGFP secretion genetic circuits………………….. 125
4.12 Fluorescence comparison between Methylobacterium not transformed 126
with sfGFP (pCMHS3) and those with sfGFP (pCMHS 1 and pCMHS 2)..
4.13 Cell growth curve of Methylobacterium comparing cell densities of wild 127
type strains and transformed strains………………………………………..
6.1 Biorefinery proposed by the Sustainable Waste-to-Bioproducts 148
Engineering Center (SWBEC) at Utah State University. Section
highlighted in red is the potential area that production of PHAs using
methanotrophic bacteria would fit into the biorefinery…………………….
B.1 BioBrick assembly process………………………………………………... 160
D.1 Schematic for production of polyhydroxybutyrates in Escherichia coli 174
from wastewater microalgae……………………………………………….
4D.2 Growth of E. coli harboring the pBHR68 plasmid on 1%, 2%, 3%, and 175
10% microalgae media……………………………………………………..
D.3 Polyhydroxybutyrate production from different microalgae based cultures. 176
xiii
LIST OF SYMBOLS, NOTATIONS, AND DEFINITIONS
ABC transporter ATP-binding cassette transporter
AFEX Ammonia fiber expansion
ANOVA Analysis of variance
BTU British thermal units
cp8 Constitutive promoter 8
DCW Dry cell weight
EPA Environmental Protection Agency
FBP Fructose 1,6-bispohsphate
FF Feast-famine
FISH Fluorescence in situ hybridization
FMP Fructose-6-phosphate molecules
FP Flavoproteins
GAP Granule-associated protein
GC Gas chromatography
GFP Green fluorescence protein
Hly Hemolysin
HPH High pressure homogenization
HPR Hydroxypyruvate reducates iGEM International Genetically Engineered Machine
JGI Joint Genome Institute
KDPG 2-keto 3-deoxy 6-phosphogluconate mcl Medium chain length xiv
MCL Malyl coenzyme A lyase
MFP Membrane fusion protein
MMC Mixed microbial culture
MMO Methane monooxygenase
MTK Malate thiokinase
NCBI National Center for Biotechnology Information
PCR Polymerase chain reaction
PEP Phosphoenolpyruvate
PhaP Phasin proteins
PhaP4MtO Phasin protein from Methylosinus trichosporium OB3b
PhaP1Re Phasin protein from Cupriavidus necator H16
PHA Polyhydroxyalkanoates
PHB Polyhydroxybutyrate
PHBV Polyhydroxybutyrate-co-hydroxyvalerate
PHD Polyhydroxydecanoate
PHH Polyhydroxyhexanoate
PHO Polyhydroxyoctanoate
PHV Polyhydroxyvalerate pMMO Particulate methane monooxygenase
RABR Rotating algal biofilm reactor
REGWQ
RFU Relative fluorescent units
RND Resistance-nodulation-division xv
RuMP Ribulose monophosphate
SAS Statistical Analysis System
SBR Sequencing batch reactor
SC Supercritical
scCO2 Supercritical fluid extraction with CO2
SCG Spent coffee grounds
scl Short chain length
SDS Sodium dodecyl sulfate
Sec General secretory
SEM Scanning electron microscopy
sfGFP Strongly fluorescent protein
sMMO Soluble methane monooxygenase
STHM Serine hydroxymethyl transferase
SWBEC Sustainable Waste-to-Bioproducts Engineering Center
T1SS Type I Secretion System
T2SS Type II Secretion System
T3SS Type III Secretion System
T4SS Type IV Secretion System
T5SS Type V Secretion System
T6SS Type VI Secretion System
TAT Twin arginine translocation
USTAR Utah Science Technology and Research
VFA Volatile fatty acids
CHAPTER 1
INTRODUCTION
1.1 Overview
Petroleum-based synthetic plastics have a wide range of applications in
industrial and consumer products, and increasingly affect every sphere of human
life. As a result of this—according to an United States Environmental Protection
Agency study in 2011—there was an increase of non-biodegradable plastics
accumulation in municipal solid waste systems from 0.5% to 12.4% over the
period from 1960 to 2010 [1]. This increase in accumulation is problematic
because the amount of waste being generated has made acquiring sufficient area
for landfill sites difficult, especially in urban areas. Synthetic plastics are
recalcitrant to microbial degradation and remain in landfill sites semi-
permanently, shortening the life of landfill sites. In addition, once in the landfill,
harmful chemicals that have been added to the plastics leach into the soil and can
spread into groundwater [2].
Renewable, biodegradable plastics are of great interest as an
environmentally-friendly replacement for traditional fossil fuel-derived plastics
[3]. Polyhydroxyalkanoates (PHAs) are one type of bioplastics that have the
potential to replace traditional plastics in many applications. PHAs are
intracellular energy- and carbon-storage polyesters produced by a variety of
microoganisms. Once PHAs are extracted from bacterial cells, they show material
properties that are similar to synthetic polymers such as polypropylene [4]. They
are biodegradable [5,6] and biocompatible [7], breaking down into non-toxic 2
products—primarily carbon dioxide and water in aerobic conditions or methane in
anaerobic conditions [8]. PHAs are known to have melting temperatures between
50-180°C and crystallinity of 30-70% [9]. As such, some potential applications of
PHAs could include single-use commercial packaging (food packaging, hygiene
and cosmetic product containers, etc.) [10,11], agricultural purposes (single-use
plastic coverings and carriers for slow-release of herbicide and insecticide) [8], and medical uses (matrixes for tissue repair and artificial organ construction, implants, capsules or coatings for slow-release of drugs, and nutritional and therapeutic composites) [12–14]. Although it is hoped that PHAs will be able to replace petrochemical plastics in these areas as production processes improve, the cost of production is currently prohibitive to all applications except higher-value medical applications [3,15,16].
There are currently two significant bottlenecks that make PHA production expensive: the cost of downstream processing and the cost of carbon feedstocks for microorganisms that produce PHAs. The development of inexpensive and scalable processes that allow recovery of intracellular PHAs is necessary [17]. It is estimated that extraction and purification costs represent as much as 50% of the total process expense [18]. Current processes are invasive and involve lysing the cellular membrane prior to PHA isolation using mechanical or biological treatments and the use of caustic chemicals. In addition to being costly, these processes also account for >90% of the adverse environmental impact of PHA production [19]. Several alternative methods have been developed for PHA 3
recovery, but none of them have resulted in significant economic improvements
[20].
A promising alternative is using synthetic biology to induce the
microorganisms to secrete the bioplastic, which would eliminate the need for cell
disruption by mechanical or chemical means. One such system has previously
been successfully demonstrated in Escherichia coli using the type I secretion
system to translocate PHAs from the cytoplasm to the extracellular medium [21].
Phasin proteins that bind tightly to PHAs were tagged for translocation via the
Hemolysin (HlyA) pathway. The phasins bound tightly to and carried the PHA
granules with them as they were secreted from the cell. Using this system, 36% of
the total PHA produced in the secreting strain was collected in the secreted
fraction and 64% remained in the bacterial cell after 48 hours. In addition, total
productivity was increased relative to the non-secreting strain [21].
The secretion system was constructed using standardized BioBrick assembly. BioBricks are a synthetic biological engineering concept of standardized biological “parts.” Essentially, they are characterized DNA sequences that can be synthesized and pieced together using restriction enzyme sites found in the regions surrounding the gene of interest in standardized DNA plasmid vectors [22]. Using standardized BioBrick parts and assembly methods
provides a flexible and powerful platform for constructing and testing options for
type I secretion of PHAs in bacteria.
The cost of feedstock for the microorganisms producing PHAs is the other
primary factor in the cost of PHA production [17,19]. In fact, techno-economic 4
analysis estimates that carbon feedstock accounts for as much as 40% of the
product cost in pure culture production [23]. Many waste streams have been
studied for use as carbon sources with reduced costs, including wastewater
microalgae [24,25], agricultural waste [26], syngas [27], traditional plastic waste
[28], and others [29]. Currently, however, most of the feedstock sources used to produce PHAs are plant-derived sugars and oils. The disadvantage of these sources is that they tend to be expensive and they are tied to our food supply.
Food wastes can be used as one type of inexpensive carbon sources to produce PHAs. In Europe, it is estimated that approximately 88 million tons of food is wasted per year [30], while in the United States, the Environmental
Protection Agency (EPA) estimated that in 2013 approximately 37 million tons of food ended up in municipal solid waste system [31]. Several food wastes have been tested for PHA production, including dairy whey, starches, oils, lignocellulosic materials, legume wastes, and sugar wastes. Mixed microbial cultures have also been used to digest complex food wastes such as food scraps, spentwashes and wastewaters, and molasses into PHAs or PHA precursors. Food wastage provides another opportunity to valorize wastes by utilizing them as carbon sources for PHA production.
Another promising alternative carbon source is methanol. It is an inexpensive and widely-available carbon source that can be utilized as a carbon source by methylotrophic bacteria, many of which are known to produce PHAs
[23,32]. Species of Methylobacterium are among the best-characterized examples of methylotrophic bacteria that naturally produce PHAs. Its PHB production is 5
reported to be as high as 57% of cell dry weight in pure culture with a yield of up
to 0.59 g PHB/g Methanol [33–35]. Since sequenced genomes indicates that
Methylobacterium generally have at least one phasin protein gene and precursors for TolC (an outermembrane protein used in Type I secretion), it is an ideal organism for incorporation of the PHA secretion system.
Methane is another potential sustainable carbon source. The bacteria strain
Methylosinus trichosporium OB3b is an obligate methanotroph that produces
PHA and has many of the same advantages as Methylobacterium [23,36]. The ability of these bacteria to metabolize methane gas would allow the production of
PHAs to be connected to a proposed wastewater treatment biorefinery (see Figure
1.1). Initially, strain OB3b was the target organism of this study, however, difficulties in culturing the bacteria caused the focus to shift to Methylobacterium.
The two species of bacteria are close enough in relation, however, that is likely
that a secretion system that is functional in Methylobacterium would also be
functional in Methylosinus. Thus, research conducted in the Methylobacterium
provides foundational information that can be applied to Methylosinus
trichosporium and other related methanotrophs, linking PHB production to the
biological engineering concept of a biorefinery. 6
Figure 1.1. Biorefinery proposed by the Sustainable Waste-to-Bioproducts Engineering Center (SWBEC) at Utah State University.
1.2 Hypotheses
This research is being conducted to obtain foundational information for
secretion-based recovery of PHAs produced by methylotrophic bacteria using
synthetic biological methods. The hypothesis of the study is that by using a trial-
and-error synthetic biological engineering approach with phasin proteins fused to
a HlyA signaling peptide will lead to an optimized system for membrane
translocation of PHAs. It is believed that producing PHAs using a methylotroph
that secretes the bioplastics will provide a more stable and economically-viable
method for PHA production by reducing downstream processing complexity and
cost. 7
1.3 Objectives
The overall objective of this study is to investigate whether a method of
recovering PHA bioplastics via secretion through a phasin protein/PHA granule
interaction can be successfully applied to methylotrophic bacteria. More specific
project objectives include:
1. Analyze the potential of utilizing food waste as a carbon source for PHA
production
2. Characterize a novel isolate of Methylobacterium
3. Design and construct a BioBrick library of phasin proteins, signal peptides,
hemolysin transport proteins, and GFP parts, as well as composite devices for
investigating secretion in Methylobacterium
4. Monitor the translocation of PHAs in Methylobacterium using gas
chromatography (GC)
The completion of the outlined objectives would contribute novel information
to PHA research. It would provide a foundation for minimizing the obstacles to
more widespread use of biodegradable bioplastics, which are currently not an
economically viable alternative to synthetic plastics due to feedstock costs and
separation/purification costs
1.4 Thesis Outline Chapter II is a comprehensive overview of using food wastes as
inexpensive carbon sources for PHA production. This chapter was previously
published as a review entitled “Food Waste Conversion to Bacterial
Polyhydroxyalkanoates” in the journal Microbial Biotechnology [37]. Chapter III
provides a comprehensive review of PHA production in methylotrophic bacteria. 8
Chapter IV describes phasin and PHA production in Methylobacterium, as well as
the genetic systems constructed for bioplastic translocation. Chapter V
summarizes general conclusions for the entire study, while Chapter VI provides
insight into future directions in PHA secretion and production from inexpensive
carbon sources.
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Plastic waste as a novel substrate for industrial biotechnology. Microb. Biotechnol. 2015;8:900–3. 29. Gomez JGC, Méndez BS, Nikel PI, Pettinari MJ, Prieto M a, Silva LF. Making Green Polymers Even Greener : Towards Sustainable Production of Polyhydroxyalkanoates from Agroindustrial By-products. Adv. Appl. Biotechnol. [Internet]. 2012;41–62. Available from: http://www.intechopen.com/books/advances-in-applied- biotechnology/making-green-polymers-even-greener-towards-sustainable-production-of- polyhydroxyalkanoates-from-agro 30. Stenmarck Å, Jensen C, Quested T, Moates G. Estimates of European food waste levels. 2016. 31. US EPA. Advancing sustainable materials management: facts and figures 2013. United States Environ. Prot. Agency. 2015;1–16. 32. Schrader J, Schilling M, Holtmann D, Sell D, Filho MV, Marx A, et al. Methanol- based industrial biotechnology: current status and future perspectives of methylotrophic bacteria. Trends Biotechnol. 2009;27:107–15. 33. Zuniga C, Morales M, Le Borgne S, Revah S. Production of poly-beta- hydroxybutyrate (PHB) by Methylobacterium organophilum isolated from a methanotrophic consortium in a two-phase partition bioreactor. J. Hazard. Mater. 2011;190:876–82. 34. Kim SW, Kim P, Lee HS, Kim JH. High Production of Poly-P-hydroxybutyrate (PHB) from Methylobacterium organophilum under Potassium Limitation. Biotechnol. Lett. 1996;18:25–30. 35. Babel W. Pecularities of methylotrophs concerning overflow metabolism, especially the synthesis of polyhydroxyalkanoates. FEMS Microbiol. Rev. 1992;103:141–8. 36. Stein LY, Yoon S, Semrau JD, DiSpirito AA, Crombie A, Murrell JC, et al. Genome sequence of the obligate methanotroph Methylosinus trichosporium strain OB3b. J. Bacteriol. 2010;192:6497–8. 37. Nielsen C, Rahman A, Rehman AU, Walsh MK, Miller CD. Minireview: Food waste conversion to microbial polyhydroxyalkanoates. Microb. Biotechnol. 2017;
11
CHAPTER 2
MICROBIAL BIOTECHNOLOGY PAPER
REVIEW
FOOD WASTE CONVERSION TO MICROBIAL
POLYHYDROXYALKANOATES1
Chad Nielsen1, Asif Rahman2,3, Asad Ur Rehman1,4, Marie K. Walsh5 and Charles
D. Miller1
1 Department of Biological Engineering, Utah State University, 4105 Old Main Hill,
Logan, Utah, USA, 84322-4105
2 Bioengineering Branch, Space BioSciences Division, NASA Ames Research Center,
Moffett Field, CA, USA, 94035-1000
3 COSMIAC Research Center, University of New Mexico, Albuquerque, NM, 87106
4 Institute of Industrial Biotechnology, Government College University, Katchery Road,
Lahore, Pakistan
5 Department of Nutrition, Dietetics, and Food Sciences, Utah State University, 8700 Old
Main Hill, Logan, Utah, USA, 84322-8700
2.1 Summary
Polyhydroxyalkanoates (PHAs) are biopolymers with desirable material
properties similar to petrochemically-derived plastics. PHAs are naturally produced by a wide range of microorganisms as a carbon storage mechanism and can accumulate to significantly high levels. PHAs can potentially be an environmentally friendly alternative
1 Chad Nielsen, Asif Rahman, Asad Ur Rehman, Marie K. Walsh and Charles D. Miller 12
to their petroleum counterparts because they can be easily degraded, potentially reducing
the burden on municipal waste systems. Nevertheless, widespread use of PHAs is not
currently realistic due to a variety of factors. One of the major constraints of large scale
PHA production is the cost of carbon substrate for PHA-producing microbes. The cost of production could potentially be reduced with the use of waste carbon from food-related processes. Food wastage is a global issue and therefore harbors immense potential to create valuable bioproducts. This article’s main focus is to examine the state of the art of converting food-derived waste into carbon substrates for microbial metabolism and subsequent conversion into PHAs.
Keywords
Food waste, polyhydroxyalkanoates, microbes, bioplastics, polyhydroxybutyrate
2.2 Introduction
Microbially produced polyhydroxyalkanoates (PHAs) are among the most well- studied biologically derived plastics. This is due to their suitability as potential replacements for petrochemically-derived plastics because they are biodegradable and biocompatible [1,2]. PHAs are carbon-based polymers naturally created to store excess carbon sources and maintain energy balances [3]. Under certain conditions, such as nitrogen, phosphorus, or oxygen limitation in the presence of excess carbon sources, some microorganisms accumulate high concentrations of PHAs [4,5].
There are over 155 confirmed unique PHA monomer subunits, which demonstrates the diversity of potential PHA polymers that can be produced using microorganisms [6]. The diversity of available monomers could lead to many different applications, as each resulting polymer has different material properties. For example, the 13
melting temperatures of PHAs range from 50 °C to 180 °C and crystallinities of PHAs range from 30-70% [7]. Polyhydroxybutyrate (PHB), a short chain length (scl) PHA, is by far the most well-studied PHA polymer and is able to accumulate to high concentrations in cells growing on a variety of carbon substrates [8]. For example,
Cupriavidus necator has been recorded to have as high as 74% of its cell weight as PHB and recombinant Escherichia coli have been recorded to accumulate up to 85% of their dry cell weight as PHB [9,10]. Some potential applications of PHAs could include: commercial packaging [11,12], agricultural purposes [13], and medical uses [14–16].
Although it is hoped that PHAs will be able to replace petrochemical plastics in these areas as production processes improve, the cost of production is currently prohibitive to all applications except higher-value medical uses [17–19].
While the properties of PHAs seem suitable as potential petrochemical plastic replacements, there are still bottlenecks for scaling up microbial production systems. One of the major bottlenecks is the cost of carbon substrates, which have been estimated to be
28-50% of the total production process [20–22]. There are a number of complex waste streams that can potentially act as carbon substrates for microbial PHA manufacture, such as waste streams from biodiesel production [23,24], municipal wastewater [25,26], agricultural waste [27], syngas production [28], traditional plastic waste [29], and others
[30]. Food waste is a prime candidate for an inexpensive carbon source, due to its wide availability and the potential to solve a significant waste problem when used to produce
PHAs.
Food wastage is a global problem and occurs at different stages of food production systems, starting from the harvesting of food to storage, packaging, and end of 14
life [31]. In Europe, it is estimated that approximately 88 million tons of food is wasted and of this, 57 million tons is from households and food service [32]. In the United
States, the Environmental Protection Agency (EPA) estimated that in 2013 approximately
37 million tons of food ended up in municipal solid waste system, which was approximately 14% of all waste in the United States [33]. Another factor that is coupled to food waste is the energy that is lost in producing, processing, and transporting the waste. This energy has been reported to be upwards of 2000 trillion British thermal units
(BTU) in the United States, equivalent to 2.11x1012 megajoules (MJ) [34]. If an alternative means of transforming food waste into value-added products are developed, then energy is essentially being transformed into useful products. Food waste conversion to tangible bioproducts has gathered plenty of attention, with systems being developed to produce a wide range of value-added products such as biofuels, materials, and a variety of additional feedstock chemicals [35–37].
The main objective of this review is to summarize the current state of PHA production from food waste using microbes, as depicted in Figure 2.1. More specifically, the purpose of this investigation is to study systems that convert a variety of food wastes into microbe-derived PHAs. Some considerations taken into account were as follows: food waste pre-treatment steps, scalability, bioreactor design, microorganisms used, and final PHA polymer produced. 15
Figure 2.1. Conversion of food waste into PHAs using PHA-producing microbes. Conversion of food waste often requires a pre-treatment step where complex food waste is broken down into subcomponents. PHA-producing microbes can then metabolize the carbon substrate and accumulate the biopolymer PHA.
2.3 PHAs from food waste using pure cultures
There are many different systems that have been proposed to convert food waste into PHAs as there are many different waste streams generated by food production, processing, and use. Each food source has its own complexities and requires different pretreatments, bacterial strains, culturing conditions, and downstream processing. Often, the organics associated with food wastes are complex compounds that cannot be directly used by PHA-producing organisms [4]. In these cases, a pretreatment or processing method is necessary to convert the complex molecules found in food waste into PHA precursors. Precursors include simple sugars like glucose or lactose and fatty acids like 16
acetic or propionic acids. Many of the simpler food wastes are hydrolyzed to convert the food waste into suitable precursor molecules and then fed directly to a pure culture of an appropriate microorganism. Whey, starch, oils, lignocellulosic materials, legume, and sugar wastes each have methods proposed to create PHAs.
2.3.1 PHAs from whey
One of the food wastes of interest is dairy whey. Whey is a by-product of the cheese-making process that consists of lactose, proteins, fats, water-soluble vitamins, mineral salts, and other essential nutrients for microbial growth. Although whey may be used as a source for producing lactose, casein, and protein powder, it is estimated that approximately 50% of whey is still disposed of in wastewater treatment plants or used in animal feed [38]. One study indicated that 1.15x108 to 1.60x108 tons of whey were produced worldwide, surpassing the requirements for whey powder production [39].
When whey is used to produce proteins like lactoferrin or lactoferricin, lactose-rich whey retentate remains as a waste material that must be disposed. Furthermore, acid whey is a by-product of cottage cheese, cream cheese, and Greek yogurt manufacturing. While acid whey can be used as animal feed, it is difficult to process into traditional whey protein concentrates due to the high acid content [38,40–42]. Disposal of whey is currently a notable problem facing the dairy industry, making it a potentially inexpensive carbon source for PHA production [43].
In addition to being a low cost carbon source, whey has the advantage of not requiring extensive pretreatment for use in fermentation via hydrolysis using enzymes or acid methods [30]. A life cycle assessment demonstrated that optimized production of 17
PHAs from whey has a comparable ecological footprint to that of producing petroleum-
based plastics and is superior to producing whey powder [39]. This life cycle assessment
was based producing PHAs using the archaeal production strain Haloferax mediterranei
DSM 1411. Producing whey powder was inferior because it was a low market value
product that used high amounts of energy to concentrate whey by evaporation. The main
limitations on PHA production were found to be energy requirement for the fermentation
process and a low amount of PHA output per kg whey input.
Most studies that used whey to produce PHAs have involved recombinant
Escherichia coli, which will be described in a subsequent section. E. coli was selected
because many of the traditional PHA-producing microorganisms cannot directly
metabolize whey. Studies have also examined using organisms such as the halophile
archaeon H. mediterranei [44,45], an unidentified highly osmophilic organism [46];
Thermus thermophiles HB8 [47]; and Cupriavidus necator DSM 545 transformed to include the lacZ, lacI and lacO genes of E. coli (becoming C. necator mRePT) [48] (see
Table 2.1). Particularly notable is the fact that H. mediterranei produces the copolymer poly-(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV), while T. thermophiles HB8 produced a unique combination of 3-hydroxyvalerate (3HV) with medium-chain length
(mcl) PHAs. Both of these polymer blends have improved ductility compared to PHB, making them preferable for industrial use [7,8]. Apart from C. necator mRePT, these organisms are capable of utilizing whey to produce PHAs with more desirable properties than PHB.
18
2.3.2 PHAs from starch
Starch is another carbon source derived from food wastes that has been studied for production of PHAs. Starch is a glucose polymer produced by plants such as rice, wheat, potatoes, maize, and cassava. Although starch is readily consumed by humans, there are some significant starchy waste streams from food production that can be used by PHA-producing microorganisms. In the work of Poomipuk et al. [49], Cupriavidus sp.
KKU38 isolated from cassava starch wastewater was used to produce PHAs from cassava starch hydrolysate. This study demonstrated that under optimal conditions and nitrogen starvation, Cupriavidus sp. could produce a moderately high biomass concentration of
5.97 g/L, with a PHA content of 61.6% (Table 2.1). A study by Kim avoided using expensive enzymes to hydrolyze starch by using Azotobacter croococcum, a type of bacteria that can digest starch directly [50]. In this study, 54 g/L dry cell weight with 46%
PHB was obtained with oxygen limitation (Table 2.1). These studies indicate that high concentrations of cells containing PHAs are possible on starchy food wastes, even with minimal pretreatments in the case of the Kim study.
2.3.3 PHAs from waste oil
Waste oils from both household and industrial applications are potential carbon sources for producing PHAs. These oils generally require no pretreatment and may be added directly to media as a carbon substrate. A study by Taniguchi, et al. investigated the use of Cupriavidus necator H16 to convert waste oils and tallow to PHAs [51]. The highest amount of PHA produced in this study came from using palm oil and lard as carbon sources, achieving a dry cell weight of 6.8 g/L and a PHB accumulation of 83%. 19
Another notable finding is that when tallow was used as a carbon source, the copolymer
PHBV was produced instead of pure PHB (Table 2.1).
Obruca and colleagues also tested waste frying rapeseed oil, waste frying palm oil, and waste frying sunflower oil as carbon sources for PHA production via C. necator
H16. They demonstrated dry cell weights (g/L) and PHB contents (%) of 10.8 and 67.9%,
11.9 and 58.0%, and 10.8 and 52.4%, respectively for the different oil types [52].
Chaudhry et al. used corn oil and found that a Pseudomonas strain could achieve a dry cell weight of 12.53 g/L with a mcl-PHA content of 35.63% (Table 2.1) [53]. Although lower quantities of PHA were produced by the Pseudomonas species used by the
Chaudhry group when compared to the C. necator H16 used by the Obruca group, the mcl-PHA produced by the Pseudomonas strain when it was fed corn oil is more desirable than PHB. These studies indicate that using waste oils as a carbon source allow cells to produce high concentrations of PHAs relative to dry cell weight in low titers of cells.
2.3.4 PHAs from spent coffee grounds
An alternative oil waste from food is spent coffee grounds (SCG) oil. Spent coffee
grounds are produced during coffee processing and consumption. Approximately 9-15%
of the grounds is oil that can be extracted for use [54]. The remaining portion of the spent coffee grounds is primarily lignocellulosic materials that can be combusted for heat or hydrolyzed and converted into PHAs by Burkholderia cepacia [55]. Obruca, Petrik, et al. directly compared use of SCG oil to other waste oils in C. necator H16 and found that the
SCG oil was superior for PHB production [52]. In a shake flask experiment, the SCG oil produced a dry cell weight of 14.2 g/L with a PHB content of 70.3% compared to the 20
values for the waste oils discussed previously (Table 2.1). When scaled up, the SCG oil
achieved an impressive dry cell weight of 55 g/L with a PHB concentration of 89.1% in
fed-batch mode (Table 2.1). The main difficulty encountered is that the SCG oil is a
natural foaming agent, however, other plant oils, such as waste frying oils, can be added
to serve as both carbon sources and as antifoaming agents.
One study by Cruz et al. used supercritical fluid extraction with CO2 (scCO2) to extract the SCG oil, then fed it directly to C. necator DSM 428 in fed-batch mode [56].
The culture reached a dry cell weight of 10.7 g/L with a PHB content of 78.4% (Table
2.1). In the same study, batch mode operation was also used to produce PHB from spent
coffee ground oil, but this approach produced lower amounts of PHB in comparison with
the fed-batch mode. The maximum biomass accumulation observed in batch mode was
up to 55% (w/w) of PHB, which yielded a polymer concentration of 6 g/L, in comparison
with a polymer concentration of 13.1 g/L observed in the fed-batch mode. The main
difference between the study by the Cruz group and the study by the Obruca group was
that the Cruz group used an extraction method that avoided the use of hazardous organic
solvents like n-hexane. As such, the Cruz process may be superior for mass production processes despite achieving lower dry cell weights. Both studies indicate that spent coffee ground oils are a carbon source with great potential for PHA production.
2.3.5 Lignocellulosic waste conversion to PHAs
Lignocellulosic materials are tough plant-based materials that are made of cellulose, pectin, hemicellulose, and lignin. Examples of this type of waste from food industry include bagasse, rice straw, wheat straw, and bran. Waste streams of 21
lignocellulosic compounds generally require hydrolysis to convert them into fermentable
sugars, then detoxification to remove inhibitory compounds produced during hydrolysis,
as reviewed in Obruca et al., 2015 [21]. A variety of lignocellulosic materials have been investigated for PHA production, including oil palm empty fruit bunch [57], wheat and
rice straw [58,59], wheat bran [60], sugarcane bagasse [61], and tequila bagasse [62] (see
Table 2.1). Despite pretreatments, the lignocellulosic materials often resulted in low levels of cell growth. One of the more promising lignocellulosic processes, however, was investigated by Cesário et al. [58]. In this study, an Ammonia Fiber Expansion (AFEX) process was used as pretreatment followed by an enzymatic hydrolysis of the cellulose and hemicellulose fractions of ground wheat straw to produce glucose, xylose, and arabinose. The hydrolysate was fed to Burkholderia sacchari DSM 17165 in a fed-batch fermentation process. A biomass concentration of 146 g/L with a PHA concentration of
72% was achieved using this method (Table 2.1). While lignocellulosic materials generally require extensive pretreatment, they do offer some potential as carbon substrates.
2.3.6 PHAs from sugar industry waste
Several waste streams from the sugar industry have been investigated for their
PHA-producing potential. One example is low-grade molasses, which is a residual syrup generated in sugar-refining mills that is high in sucrose, but not suitable for food [30].
Most studies using molasses indicate that cell production and polymer content are not currently cost competitive. For example, Chaudhry, et al. used a Pseudomonas species to convert sugar industry wastes to PHAs, and found that the dry cell weight and PHA 22
contents were 7.02-12.53 g/L and 20.63-35.63%, respectively, with molasses functioning the best overall (Table 2.1) [53]. One promising study by Kulpreecha et al. [63],
however, used sugarcane molasses as the main carbon source for Bacillus megaterium
BA-019 to achieve a dry cell weight of 72.7 g/L in 24 h, with a PHB content of 42%.
This latter study does indicate that molasses may be used to produce a considerable
amount of PHB.
Sugar beet is another industrial waste with high sucrose content. Alcaligenes
latus (ATCC 29714) was demonstrated to grow in sugar beet juice with supplemental
nutrients to achieve optimal growth of 10.3 g/L and a PHB content 38.66% (Table 2.1)
[64]. The Italian company, Bio-on, has also developed a range of PHA polymers using
local sugar beet juice [65]. The company now uses sugar beet and sugar cane wastes from
around the world to produce PHAs for use in cosmetics and pharmaceuticals [66].
Bagasse, the lignocellulosic residue of crushed sugar beets or sugar cane stalks, has been examined as a source of xylose for PHA production [67,68]. As mentioned, bagasse needs pretreatment to convert it to digestible sugars and to remove inhibitory compounds like formic acid, acetic acid, and furfural. A study by Lopes et al., for example, acid treated sugarcane bagasse at 120°C and produced 3.264 g/L PHB in
Burkholderia sp. In addition, PHBV was produced when levulinic acid was added [69].
Yu and Stahl also used acid and moderate heat (100-130 °C) to pretreat sugarcane bagasse for use as a carbon source for C. necator. The inhibitory effects of the solution was overcome by using a large inoculum of a tolerant strain of C. necator and a diluted hydrolysate solution which yielded PHB to 57% dry cell weight [61]. This demonstrates 23
that bagasse can not only be used as a fuel for boilers and as a raw material for paper, but
also as a carbon source for PHA production.
Another significant sugar industry waste is vinasse, an acidic compost with a pH of 3.5-5.0, and high organic content. Recent research into using vinasse as a carbon source for PHA production has focused on using extremely halophilic archaea like
Haloarcula marismortui and H. mediterranei [70,71]. These organisms have the advantage of not requiring a sterile environment due to the high salinity of fermentation broth and are also notable for being able to produce PHBV without the addition of organic acids as precursors. One of the main drawbacks of using halophilic organisms is disposing the saline solution after fermentation, which Bhattacharyya and colleagues addressed by using a two-stage desalination of spent stillage medium to reuse medium salts, tested with rice-based ethanol stillage as carbon source [72]. In addition, vinasse
contains polyphenolic inhibitory compounds that make pretreatment such as adsorption
on activated carbon necessary to use vinasse as a carbon source in concentrations above
10% [70,71]. After pretreating vinasse, concentrations of up to 50% were used with H.
mediterranei to produce 17.4-19.7 g/L PHA [70] and 100% vinasse was used with H.
marismortui to produce 4.5±0.2 g/L PHA [71].
2.3.7 PHAs from legume waste
Legumes have also been demonstrated as suitable carbon sources for PHA production. In a study by Kumalaningsih et al. [73], liquid bean curd waste supplemented
with an initial sucrose concentration of 25 g/L was fed to A. latus. A dry cell weight of
3.73 g/L with a PHA content of 66.56% was observed after 60 h of culturing (Table 2.1).
Soy waste was used by Yu and colleagues to produce PHAs using Alcaligenus eutrophus 24
DSM 1124, but it was found that the organism was more successful at converting malt waste to PHAs than soy, with 32.57% PHA accumulated out of 18.42 g/L dcw for soy compared to 70% PHA out of 32.36 g/L dcw for malt [74,75] (Table 2.1).
Pure cultures grown on food wastes tend to promote high cell growth and accumulation of PHAs. Some substrates, such as starch or spent coffee grounds oil, are more promising than others, such as molasses or waste oils. While using waste food substrates to feed these cultures reduces costs due to carbon source, pretreatments are often necessary. Most of the traditional bacteria used for PHA production produce the
PHB polymer, and the cost of sterilization and oxygen supply for cultures are prohibitive.
Scale-up and optimization of processes may be able improve these difficulties. Using processes that require less energy expenditure may also reduce costs. For example, use the halophilic organism H. mediterranei has the dual benefits of making sterilization unnecessary and producing PHBV polymer instead of PHB alone. Even with these drawbacks, using pure cultures to produce PHAs from food wastes has promise.
Table 2.1. Production of PHAs from food waste using pure microbial cultures. Food Waste Microorganisms PHA Cultivation Dry cell Maximum Reference Source polymer weight PHA type (g/L) production reported (g PHA / g dcw) Whey Highly PHBV Fermenter, NA 8-10% [46] osmophilic batch organism Whey Haloferax PHBV Fermenter, 10.91 66% [44] mediterranei fed-batch Whey Haloferax PHBV Fermenter, 7.45 53% [45] mediterranei batch Whey Thermus PHV and Flask, batch 1.6 35.60% [47] thermophiles mcl- HB8 PHAs Whey Cupriavidus PHB Flask, batch 8 25% [48] permeate necator mRePT
25
Table 2.1. (Continued) Food Waste Microorganisms PHA Cultivation Dry cell Maximum Reference Source polymer weight PHA type (g/L) production reported (g PHA / g dcw) Cassava Cupriavidus sp. PHB Flask, batch 9.69 61.60% [49] starch KKU38 wastewater Starch Azotobacter PHB Fermenter, 54 46% [50] chroococcum batch Soy bean Cupriavidus PHB Fermenter, 6.1 57% [51] and rapeseed necator H16 two-stage oil batch Soy bean, Cupriavidus PHB Fermenter, 6.5 79% [51] rapeseed and necator H16 two-stage corn oil and batch lard Palm oil and Cupriavidus PHB Fermenter, 6.8 83% [51] lard necator H16 two-stage batch Tallow Cupriavidus PHBV Fermenter, 5.8 80% [51] necator H16 two-stage batch Waste frying Cupriavidus PHB Flask, batch 10.8 67.90% [52] rapeseed oil necator H16 Waste frying Cupriavidus PHB Flask, batch 11.9 58.00% [52] palm oil necator H16 Waste frying Cupriavidus PHB Flask, batch 10.8 52.40% [52] sunflower necator H16 oil Corn oil Psuedomonas mcl-PHA Flask, batch 12.53 35.63% [53] species Spent coffee Cupriavidus PHB Fermenter, 16.7 78.40% [56] grounds oil necator DSM fed-batch 428 Spent coffee Cupriavidus PHB Fermenter, 55.4 89.10% [52] grounds oil necator H16 fed-batch Spent coffee Burkholderia PHBV Flask, batch 4.91 54.79% [55] grounds cepacia hydrolysate Oil palm Bacillus PHB Flask, batch 24.29 51.60% [57] empty fruit megaterium R11 bunch Wheat straw Burkholderia PHB Fermenter, 146 72% [58] sacchari DSM fed-batch Rice straw Bacillus firmus PHB Fermenter, 1.9 89% [59] NII 0830 batch Wheat bran Halomonas PHB Fermenter, 1.08 34% [60] boliviensis LC1 batch Tequila Saccharophagus PHA NA NA >0% [62] bagasse Molasses Psuedomonas PHA Flask, batch 10.54 20.63% [53] species
26
Table 2.1. (Continued) Food Waste Microorganisms PHA Cultivation Dry cell Maximum Reference Source polymer weight PHA type (g/L) production reported (g PHA / g dcw) Fermented Psuedomonas PHA Flask, batch 7.02 23.56% [53] mash species Spent wash Psuedomonas PHA Flask, batch 8.56 25.46% [53] species Sugarcane Bacillus PHB Fermenter, 72.2 42% [63] molasses megaterium fed-batch BA019 Sugar beet Alcaligenus latus PHB Flask, two- 4.01 38.66% [64] juice stage batch Sugarcane Burkholderia sp. PHB Fermenter, 6.8 48% [69] bagasse fed-batch Sugarcane Cupriavidus PHB Flask, batch NA 57% [61] bagasse necator Sugarcane Haloarcula PHB Flask, batch 12 23% [71] vinasse marismortui Sugarcane Haloferax PHBV Flask, batch 28.1 70% [70] vinasse mediterrranei Rice-based Haloferax PHBV Flask, batch 23 71% [72] ethanol mediterrranei stillage Malt waste Alcaligenus PHB Fermenter, 32.36 70% [75] eutrophus batch DSM1124 Soy waste Alcaligenus PHB Fermenter, 18.42 32.57% [74] eutrophus batch DSM1124 Bean curd Alcaligenus latus PHB Flask, batch 3.73 66.56% [73] waste
2.4 Recombinant microbes for PHA production
Natively accumulating microbial strains are most commonly utilized for conversion of food based carbon substrates into PHAs. The use of recombinant organisms, however, could be advantageous as the microorganisms can be triggered to produce PHAs without inducing stressed conditions, such as nitrogen or phosphorus starvation, which could potentially lead to cost savings. In addition, recombinant microorganisms are well-defined and thus could be further engineered for optimization.
Bacteria known to be able to utilize certain substrates that the native PHA producers 27
cannot use may also be transformed with PHA-producing genes to produce PHA from food wastes like whey, starch, or oils. Furthermore, culturing recombinant microbes could allow faster growth and turnaround times for bioreactors.
E. coli is the standard organism used in genetic engineering and has been shown to be advantageous for producing PHAs. Some strains of E. coli are known to be able to utilize lactose, a substrate that many PHA-producing organisms like C. necator are not able to metabolize. As such, most studies using high lactose containing dairy whey as a carbon source have used E. coli with the PHA-producing genes (the pha operon) from C. necator. Traditional laboratory strains of E. coli like XL1-Blue, JM, or DH5α often lack the ability to utilize lactose as a nutrient source, which has made developing alternative strains from wild-type E. coli cells necessary. When nine different strains derived from wild-type cells were tested for their ability to produce PHA using lactose as a sole carbon source, it was documented that strains GCSC4401 and GCSC6576 transformed with a high-copy-number plasmid, pSYL107 containing the A. eutrophus PHA biosynthesis operon, were best able to produce PHAs. The maximum PHB concentration and PHB content obtained were 5.2 g/L and 81% of dry cell weight, respectively (Table 2.2) [76].
Initial studies using recombinant E. coli showed potential, and subsequent studies
focused on improving production. The same group that developed E. coli strains
GCSC4401 and GCSC6576 published another study focusing on scale-up. E.coli
GCSC6576 (pSYL107) was grown on a high concentration of whey in a fed-batch
system, where a dry cell weight of 87 g/L and PHB content of 79% was achieved (Table
2.2) [77]. A follow-up study improved on these methods by controlling the timing of
PHB biosynthesis in recombinant E. coli using lactose concentrations. This allowed E. 28
coli strain GCSC6576 ( pSYL107) to accumulate PHA concentrations up to 80% of the
dry cell weight without removing culture broth (Table 2.2) [50]. As such, E. coli has been
successfully used to produce PHAs from whey using the pha operon from C. necator.
Other studies have also used E. coli to produce PHAs on whey using a PHB operon from A. latus rather than C. necator. Several strains of E. coli that were known to be able to utilize lactose were transformed with plasmid pJC4 with A. latus pha genes. It was found that strain CGSC 4401 was the ideal strain and dry cell weight and PHB content of 119.5 g/L and 80.5% were achieved (Table 2.2) [78]. This study and other early studies had problems with volumetric limitations of fermenter due to the low solubility of lactose in water and low PHB productivity. The former was partially addressed by using highly concentrated whey solution. A subsequent study by Ahn and colleagues used a cell recycle membrane fed-batch system to increase PHB productivity, achieving a final cell concentration and PHB content of 194 g/L and 87%, respectively
(Table 2.2) [79]. In a third study, the same recombinant strain grown in whey in a 30 L fermenter (26 h) and 300 L fermenter (20 h) produced 70% and 67% PHB respectively, demonstrating the process of using whey with this strain was scalable. [80].
Two other studies used recombinant E. coli to produce PHAs from whey. Pais et al. used proton suicide methodology to select for a recombinant strain of E. coli that synthesized a low amount of organic acids after it was transformed with the C. necator pha operon. The results indicated that the lower organic acid production resulted in slower growth, but a higher production of PHB (18.88 g PHB/L vs 7.8 g PHB/L in the original transformed strain) [81]. Another study used recombinant E. coli harboring the
PHB biosynthetic genes from Azotobacter sp. strain FA8 to produce PHB from whey and 29
corn steep liquor as the main carbon and nitrogen sources. The maximum cell density and
PHB concentrations attained were 70.1 g/L and 73% (Table 2.2) [82]. These studies demonstrate the use of metabolic engineering to improve production of PHA and another option for PHA producing genes that can be used in E. coli.
While whey is the primary food waste substrate investigated for recombinant E. coli, other nutrient sources have been pursued. A study by Hong and colleagues successfully cloned the pha operon from C. necator into E. coli XL1-Blue and demonstrated PHB production from soy waste. Soy waste was hydrolyzed in NaOH for 8 h then fed into a batch 3L fermenter and 27.83% PHB accumulation was observed after 9 h of culturing (Table 2.2) [83]. The same group later demonstrated recombinant production of PHBV using E. coli HMS174 with a plasmid containing the pha operon.
Malt and soy waste were obtained locally and recombinant E. coli accumulated up to
16% and 23% PHAs respectively (Table 2.2). As a comparison, when this strain was grown in glucose, it produced approximately 43% dry cell weight PHAs [84]. This demonstrates that soy and malt waste are potential carbon sources for PHA production.
In addition to malt and soy waste, another study used locally-procured restaurant waste that was anaerobically digested to produce lactic and acetic acids. Similar to processes mentioned previously, these precursors were fed to recombinant E. coli pnDTM2, which accumulated 44% PHB (Table 2.2) [85]. While using restaurant waste required an extra step to convert the complex mixture into organic acids, it provided an opportunity to use an inexpensive and widely-available carbon source.
Many studies that use recombinant bacteria to produce PHAs from food waste focus on using E. coli, which is the common workhorse of molecular biology. In addition 30
to E. coli, another Aeromonas sp. (strain KC007-1) has also been used. The strain was
chosen for its ability to directly use starch as a carbon source, and the pha operon from C.
necator H16 was added to increase production rates of PHAs. In this case, the bacteria
were able to accumulate 32.7% PHA [86], indicating that E. coli is not the only organism to be successfully modified to produce PHAs from food waste.
The examples mentioned here showed that non-native microorganisms were able
to successfully produce high amounts of PHAs. In all cases, pathways for PHA
production were transferred from a native host to the non-native microorganism. As
genetic tools increase, it will be possible to optimize pathways for non-native hosts to
consume substrates and produce PHAs. The idea to genetically engineer microbial strains
with a dual purpose of consuming inexpensive substrates and producing valuable
bioproducts has been mentioned previously [87], though relatively few studies in the past
ten years have used recombinant bacteria to produce PHAs from food waste. Future
research into scale-up and maintaining high productivity, cell concentrations, and PHA content are still necessary to make PHA production from food wastes using recombinant bacteria feasible on an industrial scale.
31
Table 2.2. Production of PHAs from food waste using recombinant bacteria.
Food Strain Plasmid PHA PHA Cultivation Dry Maximum Refer- waste operon polymer cell PHA ence source origin type weight production (g/L) reported (g PHA / g dcw) Whey E. coli pSYL107 C. PHB Shake flask 6.4 81% [76] GCSC necator 657 Whey E. coli pSYL107 C. PHB Fermenter, 87 80% [77] GCSC necator fed-batch 657 Whey E. coli pSYL107 C. PHB Fermenter, 31 80% [50] necator fed-batch Whey E. coli pMAB26 C. PHB Fermenter, 33.09 28.65% [81] CML3- necator fed-batch 1 Whey E. coli pJC4 Alcalige PHB Fermenter, 119.5 80.50% [78] CGSC nes fed-batch 4401 latus Whey E. coli pJC4 Alcalige PHB Fermenter, 194 87% [79] CGSC nes fed-batch 4401 latus Whey E. coli pJC4 Alcalige PHB Fermenter, 14.5 71% [80] nes fed-batch latus Whey E. coli pJP24K Azotoba PHB Fermenter, 70.1 72.90% [82] K24K cter sp. fed-batch FA8 Malt E. coli pUC19/P C. PHBV Fermenter, NA 16% [84] waste HA necator fed-batch Soy E. coli pUC19/P C. PHBV Fermenter, NA 23% [84] waste HA necator fed-batch Soy E. coli pKS C. PHB Fermenter, 3.025 27.83% [83] waste XL1- necator batch Blue Organic E. coli NA NA PHB Fermenter, 2.9 45% [85] acids pnDT batch M2 Starch Aerom pRK415 C. PHB Fermenter, 1.83 32.70% [86] onas H16 necator batch Sp. KC007 -R1
2.5 Production of PHAs from food waste using anaerobic digestion
One method that has been used to biologically convert complex food wastes to
PHA precursors is anaerobic digestion using open systems of mixed microbial cultures 32
(MMC). The general process that has been followed in most studies using food waste has
involved three steps: 1) acidogenic fermentation; 2) culture selection; and 3) PHA
accumulation. The primary advantage to this approach is that MMCs have lower
investment and operating costs, because substrate pretreatment processes are not
required, sterilization is typically not necessary and less expensive carbon sources can be
used [88,89]. This is important, since analyses have indicated that the high energy
expenditure used in sterilization, aeration, and agitation to produce PHAs in pure cultures
causes these bioplastics to have little advantage over traditional synthetic plastics in
environmental impact [30,39]. In addition, due to the greater variety of organisms
working together with complex substrates in MMCs, more diverse PHAs, such as
polyhydroxyhexanoate (PHH), polyhydroxyoctanoate (PHO), and polyhydroxydecanoate
(PHD) are produced [90]. This indicates that MMCs are a potentially viable option for
producing PHAs from food waste.
The first step of producing PHAs from mixed cultures is acidogenic fermentation.
During this step, complex wastes, such as food scraps, spentwash, and wastewater are
broken down into simpler and smaller fermentative acids, mainly C2-C4 acids such as
acetic, propionic, butyric, and lactic acids [91,92]. A variety of feedstocks have been used
for anaerobic production of volatile fatty acids (VFAs) from food waste, including food scraps from restaurants or kitchens [85,93–95], whey [96–98], jowar grain spentwash and rice spentwash [99], tomato cannery wastewater [100], olive oil mill pomace and wastewater [101,102], palm oil mill effluent [103,104], sugarcane molasses [91,105,106],
pea shell waste [107], condensate of food waste [90], and fermented brewery wastewater
[108,109] (see Table 2.3 and Table 2.4). Applying pretreatments such as filtering and 33
deproteinization to these waste streams has been examined, with mixed results [99,100].
Another pretreatment is buffering the waste solution to keep pH between 5.5-7.0, which has been shown to improve VFA production [85,101,110]. A variety of food waste sources have been used to produce VFAs for use by cultures of bacteria that accumulate
PHAs.
The second step of producing PHAs from mixed cultures is culture selection.
During this step, bacteria are subjected to alternating conditions to obtain a microbial community where almost all microorganisms have a high PHA storing capacity and production rate. This is often carried out in sequencing batch reactors (SBRs), compact systems where the full feast and famine cycle may be performed in one single reactor, and the length of each phase may be varied. The cycle may be either alternating conditions of external substrate excess (feast) and limitation (famine) in aerobic conditions or alternating anoxic and aerobic microenvironments. In both cases, limiting cell growth (through famine or anoxic conditions) increases PHA production and presents pressures that allow PHA-producing strains to become predominant [92]. By using microautoradiography and fluorescence in situ hybridization (FISH), Albuquerque et al.
(2012) found that in a culture grown on molasses, bacteria that their process selected for were primarily from the genera Azoarcus, Thauera, and Paracoccus. Each of these populations specialized in digesting specific products of the acidogenic fermentation.
Azoarcus and Thauera primarily consumed acetate and butyrate, respectively, while
Paracoccus consumed a broader range of substrates. Other studies on culture selection have indicated that PHA-storing bacteria in MMC are predominantly from the classes of 34
Alphaproteobacteria, Betaproteobacteria, and Gammaproteobacteria, as has been
reviewed elsewhere [92,111].
The third step of producing PHAs from mixed cultures is PHA accumulation.
This step is designed to maximize PHA production in cells harvested from the
enrichment bioreactor. The feast-famine cycle (FF) is known to increase synthesis and storage of PHA granules [112]. Alternating anoxic and aerobic conditions have also been used in SBRs to improve PHA production from food waste. It has been documented that anoxic microenvironments tend to promote higher PHA accumulation due to better access to VFAs and lack of an electron acceptor, while aerobic environments tend to promote PHA degradation, but better nutrient removal [113]. The alternating conditions pressure the cells to uptake nutrients and convert them to PHAs.
Several problems exist with MMCs. First, they generally have lower performance than pure cultures (when measured by volumetric productivity). This is due, in part, to lower cell concentrations that are usually found in MMC cultures. Often, concentrations are less than 10 g/L compared to values greater than 100 g/L that can be found in pure culture studies. Further, information on the quality of PHAs produced in MMCs is scarce
[105]. Another concern is that many of the organic acids produced during MMC fermentation inhibit bacterial growth, causing decreased productivity. These issues must be overcome to make MMCs a viable approach to producing PHAs from inexpensive carbon sources.
35
Table 2.3. Production of PHAs from anaerobically digested food waste.
Food Waste Microorganisms PHA Cultivation Maximum Reference Source polymer Production type Reported (g PHA / g dcw) Whey Wastewater PHBV Flask, batch [97] microbes N A Whey Pre-selected PHBV Flask, batch 81% [98] mixed microbial culture Whey Activated sludge PHBV Three-stage 65% [96] consortium reactors system Sugarcane Activated sludge PHBV Three-stage 56% [96] molasses consortium reactors system Brewery Activated sludge PHBV SBR 39% [109] wastewater consortium Food processing Activated sludge PHB Flask, batch 60.70% [99] wastewater consortium effluent Jowar grain-based Activated sludge PHB Flask, batch 42.30% [99] distillery consortium spentwash Rice grain-based Activated sludge PHB Flask, batch 40% [99] distillery consortium spentwash Condensate of Enriched PHBV VSMBR 1.80% [90] food waste activated sludge and mcl- consortium PHAs Olive oil mill Activated sludge PHBV SBR 39% [101] pomace consortia Olive oil mill Wastewater PHBV SBR 11.30% [102] wastewater microbes Tomato Activated sludge PHA Fermenter, 20% [100] wastewater consortium batch Fermented food Wastewater PHA Fermenter, 51% [114] waste microbes anaerobic/a erobic Fermented Mixed microbial PHBV Fermenter, 56% [105] molasses culture pulse feed Fermented Mixed microbial PHBV Fermenter, 60.50% [91] molasses culture batch
2.6 Indirect coupling of MMC to PHA production
One approach to overcoming some of the difficulties associated with MMCs is using an indirect coupling approach where complex food wastes are digested in an MMC 36
to produce VFAs, which are then harvested and fed to a pure-culture fermentation (see
Table 2.4). This increases the chance that PHA polymers will be consistent in their
quality and produced in a high concentration [115]. Like the MMCs, complex food
wastes and other inexpensive carbon sources like food scraps from restaurants or kitchens
[85,93,94,115,116] or pea shells [107] can be used as nutrient sources for PHA
production with minimal pretreatment steps.
One of the main difficulties of the indirect coupling process is efficiently
harvesting the VFAs and transferring them to a pure culture fermenter. This was initially
achieved using evaporation and ion exchange, but the processes were costly. In one of the
earliest alternative approaches, Du and colleagues compared using a silicone membrane
with a dialysis membrane to diffuse the acids into an air-bubbling reactor, while
preventing solids mixing. The dialysis membrane worked considerably better, allowing a
maximum dry cell weight and PHBV concentration of 22.7 g/L and 72.6%, respectively,
compared to a maximum of 11.3 g/L and 60.2% PHB (as opposed to the PHBV) when a
silicon membrane was used [115,116]. The primary problem was that the process still
consumed a high amount of operational energy due to small membrane pore sizes. A
subsequent study saw success with harvesting slurry once a week and filtering the VFAs
using a 0.45 μm filter [94]. Another difficulty with indirect coupling is that the VFAs are
inhibitory to growth in high concentrations, which means that it is often desirable to use
fed-batch approaches to keep concentrations low, while ensuring that VFAs are available for consumption [93]. Despite these problems, using MMC to produce VFAs that are
utilized by pure cultures is a way to combine many of the best aspects of pure culture and
MMC production of PHAs. 37
Table 2.4. Production of PHAs from organic acids derived from anaerobically digested food waste. Food Microorganism PHA Cultivation Dry Cell Maximum Reference waste polymer type Weight PHA source for (g/L) Production organic Reported (g acids PHA/g dcw) Restaurant Recombinant E. PHB Fermenter, 2.9 45% [85] waste coli pnDTM2 batch Restaurant C. necator H16 PHB Fermenter, 1.4 87% [94] waste continuous feeding Food C. necator PHBV Fermenter, 22.7 72.60% [115] scraps batch from cafeteria Kitchen C. necator PHB Fermenter, 4.6 52.79% [93] waste CCGUG 52238 batch Pea shells Bacillus cereus PHB Fermenter, 1.32 71% [107] Strain EGU3 batch
2.7 Conclusions and outlook
Utilizing food waste to create PHAs has potential for long term applications, but
is not without some hurdles. In most cases, many of the food waste sources were locally
procured and this is an important consideration since transportation of food waste to the
source of the microbial PHA production systems could be cost prohibitive. Furthermore,
the contributions of food waste pre-treatment to overall process costs of microbial PHA synthesis need to be examined in-depth. Ideally pre-treatment of food waste should be kept to a minimal to reduce time and cost. PHA extraction from microbial biomass should also be considered in a technoeconomic analysis. The most common PHA extraction methods are solvent based extraction, however there are a variety of different methods that can be used [117–119]. In addition, a biorefinery concept could be realized with food waste being the feedstock to producing PHAs and additional products, as has been proposed by others [35,36,65]. Biorefineries offer the advantages of not depending on a single product to be produced and thus are flexible and potentially sustainable. 38
The microbes used to convert food waste to PHAs are diverse, ranging from known, well-defined microorganisms to mixed microbial consortia. As mention, both natively accumulating PHA strains and genetically engineered strains could be used as platforms for PHA production. Strain selection is important aspect of PHA production and bioprospecting could lead to the discovery of additional microbes that can be used as
PHA production strains in the future [120]. In addition to engineering microbes to produce PHAs, microbes could also be optimized to use specific food substrates and generate defined chain-length PHAs.
Food wastage is a global issue, the ability to upgrade complex carbon substrates into a tangible product such as PHAs could help reduce the burden of waste processing by municipalities. As described here, many different routes on food waste conversion to
PHAs exist. There is not a one single solution to a specific type of food waste, rather there are multiple paths that could be taken, each with different pros and cons. New and innovative methods of food waste processing to PHAs will continue to grow in the future and these emerging technologies could make the economic production of microbial PHAs a reality in the not-to-distant future.
Conflict of interest
The authors declare no conflict of interests.
Acknowledgements
The authors would also like to acknowledge the Department of Biological
Engineering at Utah State University.
39
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56
CHAPTER 3
LITERATURE REVIEW
3.1 Methanotrophic Bacteria
Methanotrophic bacteria, or methanotrophs, are gram-negative bacteria known for their unique ability to utilize methane as a sole carbon and energy source. They are a subset of a physiological group of aerobic bacteria known as methylotrophs that utilize one-carbon compounds. The general category of methylotrophs utilize a wide variety of different one-carbon compounds including methane, methanol, methylated amines, halomethanes, and methylated compounds containing sulfur. Methanotrophs are unique in their use of the enzyme methane monoxygenase to catalyze the digestion of methane
[1].
The discovery and exploration of methanotrophic bacteria has been an ongoing process over the last century, as reviewed in Howard Dalton’s work [2]. Humans have known about the existence of methane since Alessandro Volta collected samples of it at
Lake Maggiore, Italy in 1778. It wasn’t until 1906, however, that the first report of a bacterium capable of growth on methane was published. Dutch microbiologist N. L.
Söhngen argued that since vast quantities of methane were produced on earth but only trace amounts of the gas could be found in the atmosphere that there must be organisms that consume methane. Through a series of experiments, he isolated Bacillus methanicus from aquatic plants and pond water. Despite this discovery, research on methanotrophic bacteria lagged for nearly six decades after Söhngen’s work. The next major study came in 1966, when Foster’s Texas laboratory published a report that detailed the isolation of a few methanotrophic bacteria [2]. 57
The major breakthrough in studies of methanotrophic bacteria came with the
Whittenbury group’s research, published in 1970. Using plate microscopes to detect tiny
colonies on plates, they isolated over 100 new species of methanotrophic bacteria.
Whittenbury and colleges devised a classification scheme for methanotrophs based on
morphological type, pathway of carbon assimilation, ability to fix dinitrogen, cyst or
spore formation and mol% G+C content [3,4]. Since that time, many more strains of
methanotrophic bacteria have been isolated from a variety of habitats, indicating that they
are likely ubiquitous in nature [2]. There is significant interest in using methanotrophs in biomanufacturing in order to avoid using a carbon source tied to the human food supply, resulting in an increasing number of studies into the physiology and capabilities of methanotrophs in recent decades.
As stated previously, the defining characteristic of methanotrophic bacteria is their use of methane monooxygenase enzymes (MMOs). The first step in assimilating methane as a carbon source is oxidation to methanol, which is catalyzed by MMOs.
There are two known types of MMOs—particulate methane monooxygenase (pMMO) and soluble methane monooxygenase (sMMO). The particulate variety is the more common type—all but one known species of methanotrophs produce pMMO. This enzyme is contained in the membrane of the cells and is thought to use copper active sites to catalyze methane oxidation. Under conditions of copper starvation, however, there are a few species of methanotrophs that express sMMO. This enzyme is contained in the cytosol of the cell and uses a di-iron active site instead of copper to catalyze methane oxidation.[5] The two types of MMOs (pMMO and sMMO) are what allow methanotrophic bacteria to metabolize methane. 58
Methane is assimilated as a carbon source over the course of several steps.
Methane is oxidized by methane monooxygenases, creating methanol. Methanol is then
converted into formaldehyde, which is the central intermediate in anabolism and
catabolism in methanotrophs (see Figure 3.1). Two main formaldehyde assimilation
pathways are possible for methanotrophic bacteria—the serine pathway and the ribulose
monophosphate (RuMP) pathway. Through these pathways, methane is assimilated by
methanotrophs.
Figure 3.1. Pathways for the oxidation of methane and assimilation of formaldehyde. Abbreviations: CytC, cytochrome c; FADH, formaldehyde dehydrogenase.
In the RuMP pathway, one molecule of C3 compound (generally pyruvate or
dihydroxyacetone phosphate) is synthesized from three molecules of formaldehyde. The
first part of the cyclic sequence (fixation) is the aldol condensation of formaldehyde with three molecules of ribulose 5-phosphate to give three hexulose-6-phosphate molecules.
These molecules then undergo isomerization to fructose-6-phosphate molecules (FMP). 59
The second part of the sequence (cleavage) begins with the conversion of one molecule of FMP into either fructose 1,6-bisphosphate (FBP) by phosphofructokinase, or to 2-keto
3-deoxy 6-phosphogluconate (KDPG) by the Entner/Doudoroff enzymes. These
molecules are then cleaved by aldolases to glyceraldehyde-3-phosphate plus the C3
molecule that is the “product” of the pathway. During the final part of the cycle
(rearrangement) the other two molecules of FMP and the remaining glyceraldehyde-3-
phosphate molecules undergo a series of reactions to regenerate three molecules of
ribose-5-phosphate (see Figure 3.2) [6]. The summary equation for the RuMP is as
follows:
2 + 3 + (1)
𝐻𝐻𝐻𝐻𝐻𝐻𝐻𝐻 𝐴𝐴𝐴𝐴𝐴𝐴 → 𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔𝑔ℎ𝑦𝑦𝑦𝑦𝑦𝑦 − − 𝑝𝑝ℎ𝑜𝑜𝑜𝑜𝑜𝑜ℎ𝑎𝑎𝑎𝑎𝑎𝑎 𝐴𝐴𝐴𝐴𝐴𝐴
Figure 3.2. RuMP pathway for formaldehyde fixation. The reactions catalyzed by unique enzymes of this pathway: hexulose-6-phosphate synthase and hexulose-phosphate isomerase, are indicated.
60
In the serine pathway of formaldehyde assimilation, two molecules of
formaldehyde plus two molecules of glyoxylate yield two molecules of 2- phosphoglycerate. One of the 2-phosphoglycerate molecules is incorporated into cell material by way of 3-phosphoglycerate, while the other is converted to phosphoenolpyruvate (PEP). The PEP molecule undergoes carboxylation, which yields oxaloacetate and subsequently malyl-CoA. The malyl-CoA is cleaved to glyoxylate plus acetyl-CoA, whose oxidation to glyoxylate completes the cycle (see Figure 3.3) [6]. The summary equation for the serine pathway is as follows:
2 + + 3 + 2 2 + 2 + + + 2 𝐻𝐻𝐻𝐻𝐻𝐻𝐻𝐻(2) 𝐶𝐶𝑂𝑂 𝐴𝐴𝐴𝐴𝐴𝐴 𝑁𝑁𝑁𝑁𝑁𝑁𝑁𝑁 → − 𝑝𝑝ℎ𝑜𝑜𝑜𝑜𝑜𝑜ℎ𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜𝑜 𝐴𝐴𝐴𝐴𝐴𝐴 𝑃𝑃𝑃𝑃 𝑁𝑁𝑁𝑁𝑁𝑁
Figure 3.3. Serine pathway for formaldehyde fixation. Unique reactions catalyzed by serine hydroxymethyl transferase (STHM), hydroxypyruvate reducates (HPR), malate thiokinase (MTK), and malyl coenzyme A lyase (MCL) are identified.
61
Methanotrophic eubacteria are sorted into three main groups, largely based on the
pathway used for formaldehyde fixation. Type I methanotrophs, or Methylococcaceae—
which include the genera Methylomona, Methylococcus, Methylomicrobium, and
Methylobacter—utilize the RuMP pathway. They are also distinguished by a membrane arrangement of bundles of vesicular disks and a G+C content of 49-60% (mol/mol). The cell morphology of type I methanotrophs is generally short rods with some cocci or ellipsoids. Group Ia methanotrophs are found in the gamma division of proteobacteria,
while group Ib methanotrophs are found in the beta division of proteobacteria.
Type X methanotrophs, such as Methylococcus capsulatus (Bath)—form an
intermediate group that shares characteristics with both type I and type II methanotrophs.
They are often classified as a subdivision of type I methanotrophs, due to use of the
RuMP pathway. What sets them apart from type I methanotrophs is their possession of low levels of serine-pathway enzymes, the presence of ribulose-1,5-biphosphate carboxylase, a G+C content of 59-65% (mol/mol), and a membrane structure of bundles of vesicular disks. Type X methanotrophs are found within the gamma division of proteobacteria.
Type II methanotrophs, or Methylocystaceae—which include the genera
Methylosinus, Methylocystis, and Methylopila—utilize the serine pathway for formaldehyde fixation. They are distinguished by a membrane arrangement of paired membranes aligned to the periphery of cells, a G+C content of 62-67% (mol/mol), and the ability to fix nitrogen. Cell morphology of type II cells is generally crescent-shaped rods, rods, and pear-shaped cells and all type II methanotrophs are found within the alpha subdivision of proteobacteria.[1] Types II methanotrophs have different survival 62
strategies than type I methanotrophs. Type I methanotrophs grow rapidly under favorable
conditions, but die off quickly under stress conditions. Type II methanotrophs grow slower, but are adapted to survive under stressed conditions. For example, type II methanotrophs outcompete type I methanotrophs under oxygen- and nitrogen-limiting conditions [7]. A few strains of type II methanotrophs are great interest for biological engineering applications.
Three methanotrophic bacteria have been the primary focus of efforts to characterize methanotrophs. Methylosinus trichosporium OB3b is a type II methanotroph that has been called the “work horse” organism for research on the physiology, biochemistry, and molecular biology/genetics of methanotrophy.[8] A draft genome has been published for OB3b [8], and it has been shown that it is capable of producing fairly high amounts of polyhydroxybutyrate (PHB) [9–12]. Methylocystis are also type II methanotrophs. They tend to dominate mixed microbial cultures, and the genus is most often studied for non-sterile production of the bioplastic polyhydroxybutyrate (PHB) using methanotrophic bacteria [12–14]. Optimized protocols exist for Methylocstis parvus OBBP [15]. Methylococcus capsulatus Bath is a type X methanotroph that has also been studied. In the European Union, a mixed culture dominated by M. capsulatus
Bath has been approved for production of proteins for nutritional feeds for salmon and livestock [16]. Together, these three bacteria are among the most notable methanotrophs.
A significant genus of methylotroph that is related to the type II methanotrophs is
Methylobacterium. Although most strains lack the ability to metabolize methane due to the absence of MMO genes, Methylobacterium species are considered facultative methylotrophs that can metabolize a variety of C1 feedstocks, including methanol. 63
Methylobacterium are ubiquitous, simple and inexpensive to culture, and have been
studied extensively. The many studies that have been performed have provided optimized
fermentation protocols [17–19], and genomic information and genetic tools are available
[20–26]. They have been shown to produce valuable products like bioplastics, vitamin
B12, pyrroloquinoline, and carotenoids [22]. Thus, Methylobacterium are considered an
ideal organism for biomanufacturing and are one of the best-characterized types of methylotrophs.
3.2 PHA Biosynthesis in Methylotrophic Bacteria
Polyhydroxyalkanoates (PHAs) are a class of intercellular energy- and carbon- storage molecules produced by a variety of microorganisms. In general, PHAs are linear, head-to-tail polyesters composed of 3-hydroxy fatty acid monomers (see Figure 3.4).
Poly(3-hydroxybutyrate) [P(3HB)] is by far the best-characterized and the most common type of PHA [27]. PHA accumulation is a response to an essential nutrient limitation such as nitrogen, phosphorus, or oxygen limitation in the presence of excess carbon sources
[28,29]. PHAs are a way for the cell to store carbon and reducing potential in a way that doesn’t affect osmotic pressure when growth is limited [30,31].
Figure 3.4. Chemical structure of PHAs. PHAs are generally composed of (R)-β-hydroxy fatty acids, where the pendant group (R) varies from methyl to tridecyl. In P(3HB), the R group is methyl. 64
PHB serves as a model compound of how PHAs are assembled. The basic pathway for PHB monomer construction starts with the condensation of two acetyl-CoA by β-ketothiolase followed by the formation of a hydroxyl group by acetoacetyl-CoA reductase, producing the monomer unit of PHB. These monomer units are then linked together by PHB polymerase into a polymer [32]. There are three genes that work to code the enzymes that convert acetyl-CoA into PHB: phbC (polymerase), phbA (β- ketothiolase) and phbB (acetoacetyl-CoA reductase), which form the phbCAB operon
(see Figure 3.5). The carbon source has an influence on which type of PHA is produced by the cells, since different S-CoA-containing molecules may be condensed with acetyl-
CoA by β-ketothiolase, becoming incorporated into the PHA molecules [32].
Figure 3.5. Dedicated pathway for P(3HB) monomer synthesis. This pathway involves the condensation of two acetyl-CoA by β-ketothiolase (PhaA) and formation of a hydroxyl group by acetoacetyl-CoA reductase (PhaB).
There are several methanotrophs that are known to produce PHB. The production of PHB in type I methanotrophs has been reported in literature [33,34], but these reports are controversial. Usually, only qualitative analysis of PHB production was reported. 65
Moreover, the genes that are generally required to create PHB are not present in type I or
type X methanotrophs [35]. Type II methanotrophs of the genera Methylocystis and
Methylosinus, however, have consistently tested positively for both the presence of genes
associated with PHB production and quantitative analysis of PHB [7,13,35–40]. Thus,
most studies involving PHA production from methanotrophs utilize Type II
methanotrophic bacteria.
It is thought that PHB production in methanotrophs may be linked to the serine
cycle. Babel hypothesized that serine-pathway methylotrophs produce PHB as a carbon storage polymer, while RuMP-pathway methylotrophs instead produce exopolysaccharides under unbalanced growth conditions [41]. Korotkova and Lidstrom found that when PHB producing genes were mutated in Methylobacterium extorquens
AM1 that growth on C1 and C2 compounds was defective. Further research indicated that
β-hydroxybutyryl-CoA (an intermediate in PHB production) may also be an intermediate
in the unknown pathway that converts acetyl-CoA to glyoxylate in methylotrophs [42].
These results point towards the theory that PHB is essential to assimilation of C1 carbon
sources in methylotrophs.
Further research has shown that PHB is utilized primarily as a sink for reducing
power in type II methanotrophs instead of energy. Pieja and colleagues observed PHB
consumption in the type II methanotroph Methylocystis parvus OBBP in a variety of
conditions. They found that when Methylocystis was starved of methane, the bacteria did
not consume significant amounts of PHB. Instead, the bacteria only consumed PHB when
methane and nitrogen were also present. When formate (a source of reducing power) was
added, PHB consumption was delayed. They theorized that PHB was used as a source of 66
reducing power to assimilate nitrogen in type II methanotrophs [14]. Likewise, Xin and
colleagues found that when Methylosinus trichosporium IMV 3011 accumulated higher
amounts of PHB that cells had increased capacity to produce methanol from CO2 because
PHB provided intracellular reducing equivalents for the cell [43]. In these cases, PHB primarily served as a sink for reducing potential rather than as a source of C2 units.
The overall equation for PHB accumulation for methanotrophs that use the serine cycle was calculated by Asenjo and Suk as follows:
8 + 12 + ( ) + 4 + 12 + (3)
4 2 4 6 2 2 2 where𝐶𝐶𝐻𝐻 FPH2 is𝑂𝑂 reduced𝐹𝐹𝐹𝐹 →form𝐶𝐶 𝐻𝐻of 𝑂𝑂flavoproteins𝑃𝑃𝑃𝑃𝑃𝑃 𝑚𝑚𝑚𝑚𝑚𝑚 (FP𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚) succinate𝐶𝐶𝑂𝑂 dehydrogenase.𝐴𝐴𝐴𝐴𝐴𝐴 𝐹𝐹𝐹𝐹 This𝐻𝐻 equation has a theoretical yield for bioconversion of methane to PHB that can be estimated to be 67% (86g/128g) [40]. Yamane suggested that if the regeneration of NADP+ of acetoacetyl-CoA reductase in the PHB biosynthetic pathway was taken into consideration the equation for PHB accumulation would become:
+ 4 + (5 3) + (5 3) + + 1 + 𝐶𝐶𝐻𝐻4 𝑂𝑂2 ( 𝑞𝑞 − 𝐴𝐴𝐴𝐴𝐴𝐴 ) +𝑞𝑞 − +𝑃𝑃𝑃𝑃4 𝑁𝑁𝑁𝑁𝑁𝑁𝑁𝑁+ (5 𝐻𝐻 3)→ + (4) + 4 6 2 2 2 Where� �𝑛𝑛�𝐶𝐶 q 𝐻𝐻is 𝑂𝑂the number𝑃𝑃𝑃𝑃𝑃𝑃 𝑚𝑚 𝑚𝑚of𝑚𝑚 𝑚𝑚moles𝑚𝑚𝑚𝑚𝑚𝑚 of ATP𝐶𝐶𝑂𝑂 assumed𝐻𝐻 𝑂𝑂 to be𝑞𝑞 produced− 𝐴𝐴𝐴𝐴𝐴𝐴 through𝑁𝑁𝑁𝑁𝑁𝑁 the conversion of CH3OH to HCHO. The theoretical yield for this equation is 54% (kg/kg).[44]
Studies have been performed to determine the amount of PHB accumulated in methanotrophs and the yield of PHB from methane or methanol. On average, those studies that used methanol tended to display higher levels of PHB Content (% of total biomass). This is likely due to methanol dissolving into water more easily, making it more accessible to bacteria. These findings are summarized in Table 3.1.
67
Table 3.1. Summary of the methylotrophic and methanotrophic production of PHB. Adapted from Strong et al.[39] Microorganisms (% in Mixed Carbon PHB Content Yield (g References Culture) Source (% of Total PHB/g Biomass) Carbon Source) Methylocystis sp. GB25 DSMZ Methane 28.3--51.3 0.54 [13,37,45] 7674 (>90%) Methylocystis sp. GB25 DSMZ Methane 10.4--33.6 0.45 [36] 7674 (>86%) Methylosinus/Methylocystis Methane 7—46 n/a [35] (percent unknown) + Type I genera Mixed consortium including Methane 34 0.8 [46] Methylobacterium organophilum (percent unknown) Methylocystis (~77%) Methane 39 0.64 [47] Methylocystis (>76%) Methane 40 0.49 [48] Methylomicrobium (96%) Methane 23 n/a [49] Methylocystis sp. (71.6%) Methane 2.5--8.5 n/a [50] Microorganisms (Pure Carbon PHB Content Yield (g References Culture) Source (% of Total PHB/g Biomass) Carbon Source) Methylocystis parvus OBBP Methane 68 n/a [40] Methylosinus trichosporium Methane 30—50 n/a [11] OB3b Type II methanotrophic strain Methane 3 n/a [7] MTS Methylosinus trichosporium Methane 30 n/a [9] OB3b Methylosinus trichosporium Methane 25 n/a [46] OB3b Methylobacterium Methane 38—57 0.48-0.59 [46] organophilum strains Methylocystis spp. Methane 7—36 n/a [35] Methylosinus spp. Methane 9—38 n/a [35] Methylocystis hirsuta Methane 51.6 n/a [52] Methylocystis parvus OBBP Methane 60 0.88 [12] Methylosinus trichosporium Methane 29 1.13 [12] OB3b Methylocystis parvus OBBP Methane 49.4 n/a [15] 68
Table 3.1. (Continued)
Microorganisms (Pure Carbon PHB Content Yield (g References Culture) Source (% of Total PHB/g Biomass) Carbon Source) Pseudomonas sp. K Methanol 66 0.18 [54] Methylobacterium rhodesianum Methanol 45-55 n/a [41] Methylobacterium extorquens K Methanol 44 0.11 [55] + n-amyl alcohol Paracoccus denitrifans Methanol 57 0.97 [55] + n-amyl alcohol Pseudomonas 135 Methanol 55 n/a [56] Methylobacterium extorquens Methanol 7—21 [57] NCIMB Methylobacterium extorquens Methanol 40—46 0.09--0.12 [19] ATCC 55366 Methylobacterium Methanol 52—56 0.19 [58] organophilum Methylobacterium sp V49 Methanol 11 n/a [59] Methylobacterium extorquens Methanol 34—42 n/a [42] AM1 Methylobacterium extorquens Methanol 22—25 n/a [60] AM1 Methylobacterium sp. GW2 Methanol 40 n/a [61] Methylobacterium extorquens Methanol 27 n/a [62] AM1 Methylobacterium extorquens Methanol 40 n/a [63] G10 Methyloligella halotolerans C2 Methanol 17 n/a [63]
3.3 Current Methods for PHA Production in Methylotrophic Bacteria
Bacteria that naturally produce PHAs generally do so under stress conditions such as nitrogen or phosphorous limitation in the presence of an excess carbon source. Due to this, most approaches to increase PHA production in microorganisms generally follow a pattern of growing a high concentration of cells and then creating a stressed environment that favors PHA production. This is largely true of methylotrophic bacteria. Much 69
research has also gone into optimizing nutrient conditions to produce high-quality and high-quantities of PHB in methylotrophs.
Economic analyses of PHA production using methane show that it is a more promising carbon source than sugars or oils. Raw material costs are reduced from 30-
50% of production costs to 22%, though temperature removal, air compression, and biomass drying remain significant factors in production costs [64]. A cradle-to-gate life cycle assessment by Rostkowski’s group indicated that using biogas methane is a preferable source of methane and that PHB recovery and demands for energy are the biggest challenges to production [65]. Utilization of biogas for PHB production also has the advantage of increasing revenue from anaerobic digesters used in wastewater remediation [16]. For these reasons, methanotrophs are of great interest in PHB production, though some hurdles remain.
Methanol is another compound that shows great promise for use as a carbon source in producing PHAs. Like methane, methanol is widely available and independent from seasonal variations and weather conditions. It can also be produced using biological organisms or manufactured from waste products like glycerol. While methane has economic advantages as a carbon source for PHA production, there are practical limitations that make methanol a superior carbon source for actual production. In one of the earliest studies on utilizing methanotrophs for PHA production, Asenjo and Suk observed that methane has a low solubility in water and that conversion of methane leads to high oxygen demands. Their suggestion was to use methanol, which has a higher solubility in the liquid phase, or to operate the system under pressure [40]. Methane is also expensive to store and transport, while methanol is easier to work with in those 70
regards [66]. Both the amount and the molecular weight of PHB produced by methanotrophs fed with methanol or a combination of methane and methanol have been shown to be superior to those fed by methane alone [11,67,68]. In addition,
Methylobacterium species have been intensely studied, but are not able to metabolize
methane directly. They, like most methylotrophs, are capable of metabolizing methanol,
adding incentive to using methanol instead of methane.
One method that has been used to reduce energy-associated costs for
methanotroph production is using non-sterile mixed microbial cultures (MMC).
Wentlandt et al., found that using Methylocystis sp. GB 25 DSM 7674 in a short non-
sterile process allowed accumulation of high-quality PHB that neared theoretical yields at
51% [37]. Another study found that in an open system of MMC, that Methylocystis sp.
GB 25 was dominant, with 7 other species present (2 methylotrophs and 5 other species).
This MMC was stable for long periods of time and produced high-quality PHB. It was
theorized that mixed cultures were beneficial because they allowed some bacteria to
remove toxic byproducts of methylotrophic metabolism, while also providing essential
vitamins or growth supplements to the PHB-producing methylotrophs [13]. It has been
suggested that investigation of a mixed culture of PHB-producing methanotrophs and
PHB-producing cyanobacteria may result in a particularly viable co-culture for PHB
production [10]. MMCs of methanotrophic bacteria are likely to be viable for industrial
production of PHB.
A large amount of research has gone into optimizing media for PHB production
using a variety of methanotrophic bacteria. Much of this has been reviewed elsewhere
[10,39,66], but it has been discovered that several environmental factors affect 71
accumulation and molecular weight of PHAs. Among these factors are pH, temperature,
CH4 concentration, and trace elements. Variations in optimizing media exist across
specific strains, and attempts at optimizing media must focus on individual species. For
example, a few major attempts have been made to optimize cell growth and PHA
production in Methylobacterium extorquens over the years, and have found that EDTA is
inhibitory to growth [18], and that starvation of both magnesium and nitrogen have been
shown to increase PHA production [17]. Optimization of media and protocols for cell growth and production in methylotrophic bacteria is an important step to industrial use of these species.
Most studies that have tried to optimize the parameters of these nutrients have done so in small-scale batch reactors, though a few have examined the use of larger bioreactors. At a larger scale, Stottmeister’s group used pressurized bioreactors at 7 L and
70 L volumes to produce PHAs in a mixed consortia of bacteria [13,36,37,45] and
Pfluger et al. used a fluidized bed reactor with a total volume of 15.2 L under non-sterile
conditions to produce PHB in a mixed culture of methanotrophs [69]. Both of these
mixed-culture conditions were shown to be viable. Rahnama et al. examined using a pure
culture of Methylocystis hirsuta in a 1 L bubble column reactor and a forced liquid loop
reactor to produce PHB [52], Daniel et al. produced PHB with Pseudomonas 135 in a 15
L bottle with a 10 L initial working volume in fed-batch mode [56], and Bourque et al. utilized a 14 L bioreactor [19]. Finally, three studies looked at using pure cultures of methylotrophic bacteria in 5 L stirred tank reactors, including Song et al. [53], Ghatnekar et al., [59], and Ueda et al. [55]. Of the larger-scale pure cultures, PHB concentrations (% of total biomass) of greater than 40% were observed in all the studies listed except 72
Ghatnekar et al. (see Table 3.1). Thus, some work has been done with PHB production
using methanotrophic bacteria in larger bioreactors.
One problem with PHA production in methylotrophic bacteria is that they often
produce low-quality PHAs. It is reported that PHA granule size varies from 50 to 1,000
kDa based on growth parameters and host strain [27]. In general, longer polymer chains
of PHAs yield higher molecular weights and better properties. Early research into
methylotrophic production of PHAs using Methylobacterium extorquens and
Pseudomonas 135 indicated that methylotrophic bacteria were unable to produce PHAs
of a sufficient molecular weight to be usable in commercial applications [56,57]. More
recently, however, it has been reported that in certain growing conditions,
methanotrophic bacteria Methylocystis sp. GB 25 DSM 7674, Ms. trichosporium OB3b,
and Ms. trichosporium IMV 3011 can produce high molecular weight PHBs (above
1x106 Da) [36–38,67]. Studies in Mtb. extorquens indicated that molecular weights of
PHB produced were lower when methanol was used as a carbon source, compared to
succinate (at best 6 x 105 Da compared to 1.7 x 106 Da). Anderson and colleagues
discovered, however, that when lower concentrations of methanol were available that the
molecular weights of PHB produced were higher and had a lower polydispersity
compared to those produced when methanol was abundantly available (6 x 105 Da
compared to 2 x 105 Da) [70]. Thus, methylotrophic bacteria are capable of producing
high-quality PHB under certain conditions.
It is known that many types of PHAs can be created when different carbon
sources are metabolized. When propionate is introduced as a feedstock, it is possible to
produce 3-hydroxyvalerate (3HV) using the PHB pathway. This occurs due to one of the 73
two acetyl-CoA units that are condensed into PHB being replaced with propionyl-CoA, producing polyhydroxybutyrate-co-valerate (PHBV) [32]. Until recently, only PHB production was reported in obligate methanotrophic bacteria, but ongoing research by
Myung and colleagues has shown that a greater variety of PHAs may be synthesized in methanotrophs. In a mixed microbial culture that was dominated by Methylocystis, they found that when valerate was added, PHBV was produced [48]. It was later confirmed that both M. parvus OBBP and Ms. trichosporium OB3b were capable of producing
PHBV when valerate and propionate were added to pure cultures with methane [71].
When other ω-hydroxyalkanoate monomers were added to pure cultures of M. parvus
OBBP, PHAs containing repeating units beyond 3-hydroxybutyrate (3HB) and 3HV were found [72]. These studies represent the first times that PHAs other than PHB have been reported in obligate methanotrophic bacteria.
Multiple types of PHAs have also been detected in Methylobacterium strains.
PHV has been detected in Mtb. extorquens and Methylobacterium sp. GW2 when propionate or valerate are added as co-substrates [24,73]. Zuñiga et al. discovered that the
Mtb. organophilum CZ-2 strain produced six different monomers when methane was used along with either citrate or propionate as co-substrates. These monomers included
3HB, 3HV, 4HV, 4-hydroxyheptanoate (4HH), 3-hydroxyoctanoate (3HO), and 4HO
[74]. Höfer and colleagues have investigated using genetic engineering to transform the phaC1 or phaC2 gene from Pseudomonas fluorescens GK13 into Mtb. extorquens ATCC
55366 to introduce double bonds into PHAs produced. They found that they were able to do so, which allowed functional groups to be added to the PHAs using chemical steps 74
after production, creating more desirable properties [24,75]. Thus, Methylobacterium can
produce multiple types of PHAs when specific substrates are made available.
These findings are significant because variations of PHAs may have greater value
and use. PHAs vary in chemical structure and in number of monomer subunits. There are
155 different confirmed types of PHA monomer subunits, each with varying monomer
repeat number and side groups [32]. The variations among these polymers impact their
crystallinity, mechanical strength, and biodegradability, making some types of PHAs
more suitable for commercial use than others [67,76]. For example, on its own, PHB is
brittle (glass transition temperature (Tg)=4°C, melting temperature (Tm)=175°C), but
when synthesized into a copolymer 3HV, a polymer with better properties for consumer
products is created [32]. Each type of PHA has its own advantages and disadvantages, and the variety that exists allows final bioplastic products that can be tailored for many different applications.
Very little literature has been devoted to economic analyses of PHA production
using methane and methanol as carbon sources. As previously mentioned, Rostkowski’s
group did some analyses on types of methane and indicated that using biogas methane is
a preferable. They also indicated that PHB recovery and demands for energy are the
biggest challenges to production [65]. Further cost analyses on the influence of methane
gas, methanol, and sugar as carbon sources on the final cost of production would be
important to demonstrate the viability of these feedstocks for PHA production. 75
3.4 Current Methods for PHA Recovery
Recovery of PHAs from bacteria is a costly process. PHAs are intracellular molecules that accumulate inside of the cell and are usually not ejected into the surrounding environment. Thus, the cells must be disrupted to recover PHA. Three general steps are involved in the process of recovering and purifying PHA (see Figure
3.6). First, a pretreatment step is applied to help lyse the cell. Second, the PHA is extracted from the cells. Third, further purification or polishing is often necessary after extraction to obtain high purity. The goal of optimizing these processes is to achieve a high purity and low cost.
Figure 3.6. Purification strategy of PHAs. The recovery of PHAs could be composed of three steps: pretreatment, extraction, and purification.
76
The first step is pretreatment, which is applied after centrifugation to make cell
disruption easier. This step is not always used; however, it can lead to a better recovery of
PHA granules. One pretreatment option is heat treatment, which destabilizes the outer
cell membrane while denaturing genetic material and proteins. [77,78]. A second option
is alkaline treatment, often using a solution of sodium hydroxide. For example, one study
using 0.12 kg NaOH/kg biomass of Alcaligenes latus found that most of the PHB could
be released within three passes through a bead mill, compared to at least ten passes with
untreated cells [79]. Salt pretreatment is another option that has been investigated. For this treatment, cells are placed in saline solution, putting osmotic pressure on the cells and causing them to shrivel and dehydrate [79,80] Finally, a freezing pretreatment can be
used to rupture cells. This is often performed by performing a deionizied water washing
and then freezing. The important disadvantage to this method is that it has high energy
demands [81–83]. Each of these pretreatment methods is intended to loosen up cell
membranes to make extraction more successful.
The second, and most important, step in PHA recovery is extraction. One classic
approach to extraction is to use solvents to disrupt the cell membrane and solve PHAs.
Chlorinated hydrocarbons solvents are one category that has been used, including
chloroform, 1,2-dichloroethane, and methylene chloride [84], sometimes in combination
with methanol or ethanol. Liquid halogenated solvents, such as chloroethanes and
chloropropanes, have also been used [85], as have diols, acetalized triols, di- or
tricarboxylic acid esters, butyrolactone [86], and many other compounds [87,88]. The
PHAs are generally separated from the solvent by evaporation of the solvent or
precipitation in a non-solvent, such as cool water [84,89,90]. Benefits of using solvents 77
include removal of endotoxins [91] and minimal degradation of the polymer. Problems
facing solvent extraction include cost [92,93], viscosity of the extracted polymer solution
and a resulting difficulty of separation from cell debris [94], destruction of the natural
morphology of PHA granules [95], and hazards to operators and the environment [96].
Thus, solvent extraction is often used in lab studies, but not in industrial extraction of
PHAs.
Another extraction method is digestion. Chemical digestion using surfactants like sodium dodecyl sulfate (SDS) or synthetic palmitoyl carnitine breaks up the cell membrane and solubilizes proteins and other cell material [97–100]. While this allows recovery of PHAs from high cell densities, surfactant digestion does not lead to high purities on its own and can cause problems in wastewater treatment and reuse [101].
Alternatively, digestion using sodium hypochlorite breaks down non-PHA cellular material, allowing for high-purity levels of PHAs [81,82]. Unfortunately, sodium hypochlorite is known to degrade PHB, resulting in low molecular weights [81,102,103].
Combining chloroform with sodium hypochlorite overcomes some of this problem, since
the chloroform solves and sequesters the PHAs (reducing degradation) while the sodium
hypochlorite breaks down cell materials [82]. Lytic enzymes like bromelain or pancreatin
have also been used for digestion. Methods of enzyme digestion generally lead to good recovery levels, but high costs [77,104]. Digestion techniques are often used to extract
PHAs from bacteria.
Mechanical disruption techniques like bead mills and high pressure homogenization are also widely used in extraction. Bead mills involve grinding cell slurries in a rotating chamber filled with beads. The performance of the chamber is 78
consistent and predictable, making the process easy to scale-up [101]. A second approach to mechanical disruption is high pressure homogenization (HPH). An air-driven positive displacement pump forces cell slurry through two parallel slots under high pressure. The two streams impinge on a vertical plate, recombine into one stream and then flow out.
The shear forces on the liquid during this process disrupt cells. This approach works particularly well with higher concentrations of biomass, but often has issues with blockages [59]. Ultrasonication has also been used on a lab scale, often in combination with centrifugation or chemical treatments [105]. Bead mills and HPH, however, remain the most common mechanical disruption techniques used of PHA extraction.
Biological extraction methods for PHAs have been investigated in recent years.
One group investigated using the predatory bacterium Bdellovibrio bacteriovorus HD100 to lyse the cells of three PHA-producing species of bacteria (P. putida, C. necator, and E. coli). A strain of B. bacteriovorus was mutated to knock out a PHA depolymerase gene in order to prevent unwanted breakdown of PHA granules, and up to 80% of the accumulated PHA was recovered from the prey bacteria [106]. In another novel biological approach, Murugan and colleagues fed PHA-containing C. necator to mealworm beetle (Tenebrio molitor) larvae and purified the bioplastic granules from the insect’s feces using water, detergent and heat. They observed almost 100% pure PHA with minimal loss of molecular weight as a result of their process [107]. These cases demonstrate that biological methods can be used to release PHAs from bacterial cells.
Other methods of extraction exist, such as supercritical fluid or utilization of cell fragility. Supercritical (SC) fluids are substances held at a temperature and pressure above their critical point, where distinct liquid and gas phases do not exist. The most 79
widely-used SC fluid is CO2, since it is widely available and inexpensive, has low toxicity, is nonflammable, and has moderate critical temperature and pressure (31 °C and
73 atm) [80,108]. One study used SC fluids to recover PHB and found that the amount recovered was similar to other studies, at 89% [108]. Cell fragility is another approach to extraction of PHAs. Some bacteria become fragile after accumulating large amounts of
PHA, and addition of compounds like NaOH or NH3 at raised temperatures released PHA into the medium [109,110]. These approaches represent a few other techniques that have been used for extraction.
One other significant method of extraction is spontaneous liberation or secretion of PHAs. One study found that when E. coli cells harboring Alcaligenes phbCAB genes were grown in a 2 x LB medium containing 21% glucose, that up to 80% of the cells spontaneously secreted PHB granules [111]. Resch’s group used a combination of the phaCAB genes and the cloned lysis gene E of bacteriophage PhiX174 from plasmid pSH2 in another study. They showed that small PHB granules exited the cells through the E- lysis tunnel structure [112]. Sabitova et al. discovered a mutant of the marine oil- degrading bacterium Alcanivorax borkumensis SK2 that hyperproduced PHAs. The result of the hyperproduction was that the bacteria pushed PHA into the extracellular fluid when grown on alkanes [113]. Finally, Rahman and colleagues created fusion proteins of a haemolysin secretion tag with phasin proteins (PhaP) that bind to PHAs. When transformed E. coli that contained the fusion proteins, C. necator phaCAB genes, and a plasmid with the hemolysin BD pore proteins were cultivated, it was found that up to
36% of PHB was secreted after 48 hours [114]. Thus, secretion is a possible method for liberation of PHAs. 80
The third step of that may be applied to PHA recovery is a final purification or
polishing step. Most often, methods combine a hydrogen peroxide treatment combined
with the action of enzymes of chelating agents. These methods have drawbacks,
including high operation temperature (80-100 °C), instability of peroxides in the presence
of high levels of cellular biomass and a decrease of polymer molecular weight [101,115].
One process proposed by Horowitz and Brennan overcame some of these problems by
using an ozone treatment to increase the purity of the polymer [115]. When applied,
polishing results in bioplastics suitable for applications that require high purities, such as
coating.
Many different approaches to purifying PHAs from the cells that produce them
have been developed. Unfortunately, they are generally still too expensive and inefficient
to produce sufficient quantities of PHA to be widely used. Although many alternative
methods have been developed for PHA recovery, none of them have resulted in
significant economic improvements in downstream processing [116]. It is estimated that
extraction and purification costs represent as much as 50% of the total process expense
[111]. In addition to being costly, these processes also account for >90% of the adverse
environmental impact of PHA production [65]. The development of inexpensive and
scalable processes that allow recovery of intracellular PHAs is still deemed necessary
[117,118].
3.5 Genetic Engineering in Methylobacterium species
Many of the genetic tools used with methylotrophic bacteria were developed for use in Methylobacterium. In the early 2000s, Figueira and colleagues developed a 81
protocol for electroporation in Methylobacterium using wide-host expression vectors pJB3KmD, pRK310, and pVK101 and were able to express GFP proteins [23]. Soon afterwards, Marx and Lidstrom developed a set of improved plasmids for use in
Methylobacterium. Expression vectors for use in Methylobacterium extorquens AM1 used the strong methanol dehydrogenase (PmaxF) promoter isolated from
Methylobacterium, since lac promoters from E. coli proved ineffective [26]. Using one of
these plasmids, Choi and colleagues were able to express an esterase gene from
Lactobacillus casei CL96 in Mtb. They did so at levels of enzyme activity 1.2- to 1.5-fold
higher than those found in E. coli [21]. The same group were also able to use the mini-
Tn7 transposon system to integrate three different genes into a specific locus on the
bacterial chromosome of Methylobacterium. These tools have formed a basis for genetic
engineering in Methylobacterium.
Further research has continued to provide tools for metabolic engineering of
Methylobacterium. The PmaxF promoter that is commonly used in Mtb. is regulated by the
presence of methanol, causing it to effectively be a constitutive promoter when used in
Methylobacterium. To increase flexibility of use, Choi’s group developed an inducible
regulatory system to the PmaxF promoter using a regulatory element from Pseudomonas
putida F1. The cym element was added to the PmaxF promoter, creating a cumate-
inducible promoter for Methylobacterium [22]. Research by Höfer and group used this
system to clone phaC1 and phaC2 genes from Pseudomonas fluorescens GK13 to
produce PHAs with C-C double bonds [24,75]. Another recent study worked to create a
library of standardized plasmid vectors and promoters for use in Mtb. extorquens AM1 82
specifically, and alphaproteobacteria generally [119]. These studies represent ongoing
developments in the genetic manipulation of Methylobacterium.
3.6 Recombinant Proteins
3.6.1 HlyA Secretion Pathway
Secreted proteins are the primary means by which bacteria interact with their
external environments. Functions of secreted molecules are vital to cells, and include
adhesion, pathogenicity, adaptation, and enzymes related degradation in some cases. Due
to this, bacteria have developed a variety of methods used to secrete proteins into the
extracellular matrix. For example, there are six secretory pathways by which Gram-
negative bacteria are known to translocate proteins to the extracellular matrix [120,121].
Proteins secreted via these pathways tend to have three fates: to remain associated with the bacterial outer membrane, to be released into the extracellular space, or to be injected into a target cell. The difficulty associated with secretion in Gram-negative bacteria compared to Gram-positive bacteria is that Gram-negative strains have two lipid bilayers
(making them didermic bacteria) while Gram-positive only have one lipid bilayer
(making them monodermic bacteria). Each of the six known secretion systems in didermic bacteria handle this issue in different ways.
Type I Secretion Systems (T1SSs) are composed of three segments that interact together to transport proteins into the extracellular matrix. In this system, proteins with a glycine-rich C-terminus signal peptide are transported across the inner membrane via an
ATP-binding cassette transporter (ABC transporter), then fed through a membrane fusion protein (MFP) to an outer membrane polypeptide known as the TolC protein, which 83
secretes the proteins into extracellular space (see Figure 3.7) [122]. These systems are
closely related to the resistance-nodulation-division (RND) family of multidrug efflux
pumps, and are often used to secrete toxins and virulence factors [120]. An example of a
T1SS found in E. coli is the hemolysin system, in which the toxin protein HlyA is secreted through the HlyB (ABC transporter), the HlyD (MFP), and the TolC into the extracellular environment, where the HlyA can lyse red blood cells [123–126]. Several
other double-membrane-spanning systems are also classified as T1SSs.
Figure 3.7. Diagram of the type I secretion system (T1SS). OM is outer membrane, IM is inner membrane, MFP is membrane fusion protein, and IMC is inner membrane channel.
Type II secretion systems (T2SSs) are one of the most ubiquitous secretions
systems known in Gram-negative bacteria. They are two-step systems involving four
parts wherein a cytoplasmic ATPase, such as the SecYEG complex, exports proteins with 84
the proper N-terminus recognition sequence into the periplasmic space [127,128].
Following this, a periplasmic pseudopilus mounted on an inner membrane platform
transports the protein to an outer membrane complex that secretes the protein into
extracellular space [120,129]. The two most well-known T2SSs are the Sec (general secretory) pathway and the twin arginine translocation (TAT) protein export pathway
[127,130,131]. Of the two, the Sec pathway is more common, but limited to transporting unfolded peptides into the periplasm [131]. The TAT export pathway is capable of transporting folded proteins across the inner membrane, sparking a greater amount of interest for use in biological engineering than the Sec pathway [130,132,133]. The T2SSs are well-characterized and have potential for use in secreting proteins.
Type III, IV, and VI secretion systems allow direct transport of cell materials into a host cell. Type III secretion systems (T3SSs) are double-membrane embedded nanomachines that allow transfer of bacterial effector proteins to the cytoplasm or plasma membrane of a target cell. They are formed primarily as a needle-like filament on a base that forms a secretion conduit from the cell membrane of the bacterial cell to the interior of the host cell.
Type IV secretion systems (T4SSs) mediate the conjugation of plasmid DNA and some proteins between bacterial cells via pili. These systems are found across the spectrum in bacteria, and are famous as a means of conferring antibiotic resistance in bacteria. Type VI secretion systems (T6SSs) are cell envelope-spanning machines that translocate toxic effector proteins into target cells, and are important in pathogenesis.
They are built of two main complexes: a membrane complex made of inner membrane proteins and a tall, bacteriophage tail-like structure that can contract when necessary 85
[120,134]. These three types of secretion systems (T3SSs, T4SSs, and T6SSs) are known for forming a conduit to transport proteins and DNA from the bacterial cell of origin to a host cell.
Type V secretion systems (T5SSs) are different from other secretion systems in that they are single-membrane-spanning secretion systems, while the other five system types are double-membrane-spanning secretion systems. Most famously, the T5SSs include autotransporter systems in which the substrate and the outer membrane pore are a single peptide, though there are other types of T5SSs. These systems are often involved in bacterial virulence [120,135]. T5SSs are often simpler in composition than the other secretion systems, due to the fewer individual proteins involved in secretion.
The T1SSs and T2SSs are the most commonly-used secretion systems in biological engineering research. The fact that each system recognizes a known sequence on one terminus of the target peptide means that the signal sequence may be fused to other proteins, causing the cell to target the new fusion protein for secretion. This is advantageous for downstream processing of proteins because they are subjected to less degradation via protease activity outside of the cytoplasm, the oxidative environment of the periplasm allows the correct formation of disulfide bonds, and when secreted into the extracellular environment, disruption of the cell membrane is no longer necessary to purify the protein [133]. The use of secretion tags in recombinant protein production has been reviewed extensively elsewhere [133,136]. In short, the T1SSs and T2SSs are most commonly used with recombinant proteins in E. coli due to their ability to export proteins to the periplasm or extracellular environment. 86
Fusion proteins involving secretion peptides have been used to indirectly secrete
PHB molecules. A foundational study was performed by Linton and colleagues that
tested the efficacy of four of the most common secretion tags in E. coli. The signal
peptides utilized were the HlyA (T1SS), TorA (T2SS, TAT), GeneIII (T2SS, Sec), and
PelB (T2SS, Sec) peptides. The PelB secretion system was ineffective at translocating
GFP, while the other two T2SSs successfully exported GFP to the periplasm. They HlyA
system was able to successfully secrete GFP into the extracellular media [137]. Building
on this research, the Miller group used a fusion protein of the HlyA signal peptide and a
phasin protein that associates with PHB granules to link the signal sequence to PHB. This
resulted in the secretion of PHB granules into extracellular media via a T1SS in E. coli
[114].
3.6.2 Phasins PHA granules are not purely composed of the bioplastic polymer, but are covered
with a layer of granule-associated proteins (GAPs). This proteinaceous surface layer of
GAPs makes up approximately 2% (w/w) of the granules and is a network-like surface of structural, metabolic and regulatory polypeptides (see Figure 3.8) [138]. The GAPs that are involved in PHA granules have been divided into four categories: PHA synthases, which catalyze the formation of ester linkages between the monomeric units of PHAs;
PHA depolymerases, which degrade PHAs; phasin proteins (which will be discussed in greater detail); and other proteins (transcriptional regulators, hydrolases, reductases, etc.)
[139]. Phospholipids are also potentially included in the external layer, though it is possible that their involvement is an experimental artefact of PHA extraction and 87
preparation rather than a part of PHA granules in vivo [138]. The most abundant polypeptides in PHA granules are the phasin proteins.
Figure 3.8. Schematic of the structure of PHA granules.
Phasin proteins are known to play an important role in the regulation of PHA synthesis, morphology, and degradation. Phasin proteins are amphiphilic molecules that bind to the surface of PHA granules. The seven phasin proteins in the Cupriavidus necator H16 (formerly Ralstonia eutropha) have been studied most intensely, particularly
PhaP1Reu. Mutant C. necator cells that lack PhaP1Reu have been observed to produce larger granules and less PHB compared to the wild type strains [140–142], while overexpression of PhaP1Reu results in a high number of smaller granules [143]. The PhaM has specifically been identified as an activator of PHB synthase activity in C. necator 88
[144]. It has also been noted that phasin protein deletion mutants of C. necator exhibit
some PHB autodegradation in vivo, indicating that phasin proteins are important to granule stability [142]. Ironically, PhaP1Reu is also known to be essential to PHB
degradation [145,146]. Thus, many functions have been observed for phasin proteins
relative to PHA granules.
Phasin proteins are also known to regulate PHA granule distribution during cell
division. In Pseudomonas species, the PhaF phasin protein is composed of two domains,
one like PhaP1Reu in structure and function on the N-terminal, and a highly-charged, histone-like domain on the C-terminal [147,148]. Working together, these two domains link PHA molecules to nucleoid DNA during cell division, ensuring an equal distribution between daughter cells [149,150]. Similar behavior has been observed with the phasin- like polypeptide PhaM and the Pha5 protein in C. necator [151,152]. These studies demonstrate the capability of phasin proteins and phasin-like proteins linking PHA granules to other molecules in the cytosol.
Most PHA-producing organisms are known to produce some form of phasin proteins. In addition to C. necator and Pseudomonas species, Paracoccus denitrificans
[153,154], Rhodococcus ruber NCIMB 40126 [155,156], Azotobacter species [157,158],
Aeromonas species [159,160], Rhodospirillum rubrum [161,162], Magnetospirillum
[161,163], Bradyrhizobium [164,165], Sinorhizobium meliloti [166], Haloferax
mediterranii [167], and Herbaspirillum serepedicae [168,169] have all been noted for
phasin protein or phasin-like protein production related to PHA synthesis. In most cases,
phasin protein production has a positive correlation to PHA production and have been 89
noted for similar functions to those discussed previously relative to C. necator and
Pseudomonas.
Phasin proteins have been used for a few different functions in biological
engineering. In addition to their importance in producing PHAs, the amphiphilic nature of phasins allows them to function as natural surfactants. The PhaPAh from A. hydrophila
4AK4 has been shown to form emulsions with lubricating oil, diesel, and soybean oil that
are relatively stable when compared to those formed by bovine serum albumin, sodium
dodecyl sulfate (SDS), Tween 20, and sodium oleate [170]. Taking advantage of the
native abilities of PhaF from Pseudomonas to link PHA molecules to DNA, one study
used the phasin to anchor proteins to PHA granules, immobilizing the enzymes [148]. As
has been discussed previously, phasin proteins have also been used to anchor signaling
peptides to PHB granules in recombinant E. coli, inducing secretion of PHB into the
extracellular milieu [114]. The amphiphilic nature of phasin proteins has been utilized in
several biological engineering applications.
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sequence-like elements associated with putative polyhydroxybutyrate regulatory genes in Azotobacter sp . FA8. Plasmid. 2003;50:36–44. 158. Mezzina MP, Wetzler DE, Almeida A de, Dinjaski N, Prieto MA, Pettinari MJ. A phasin with extra talents: a polyhydroxyalkanoate granule-associated protein has chaperone activity. Environ. Microbiol. 2015;17:1765–76. 159. Fukui T, Kichise T, Iwata T, Doi Y. Characterization of 13 kDa Granule- Associated Protein in Aeromonas caviae and Biosynthesis of Polyhydroxyalkanoates with Altered Molar Composition by Recombinant Bacteria. Biomacromolecules. 2001;2:148–53. 160. Qiu Y-Z, Han J, Chen G. Metabolic engineering of Aeromonas hydrophila for the enhanced production of poly (3-hydroxybutyrate-co-3-hydroxyhexanoate). Appl. Microbiol. Biotechnol. 2006;69:537–42. 161. Handrick R, Reinhardt S, Schultheiss D, Reichart T, Jendrossek V, Jendrossek D. Unraveling the Function of the Rhodospirillum rubrum Activator of Polyhydroxybutyrate (PHB) Degradation: the Activator Is a PHB-Granule-Bound Protein (Phasin). J. Bacteriol. 2004;186:2466–75. 162. Handrick R, Technow U, Reichart T, Reinhardt S, Sander T, Jendrossek D. The activator of the Rhodospirillum rubrum PHB depolymerase is a polypeptide that is extremely resistant to high temperature (121 °C) and other physical or chemical stresses. FEMS Microbiol. Lett. 2004;230:265–74. 163. Schultheiss D, Handrick R, Jendrossek D, Hanzlik M, Schüler D. The Presumptive Magnetosome Protein Mms16 Is a Poly (3-Hydroxybutyrate) Granule- Bound Protein (Phasin) in Magnetospirillum gryphiswaldense. J. Bacteriol. 2005;187:2416–25. 164. Yoshida K, Takemoto Y, Sotsuka T, Tanaka K, Takenaka S. PhaP phasins play a principal role in poly- -hydroxybutyrate accumulation in free-living
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CHAPTER 4
AN INVESTIGATION OF PHASIN TRANSLOCATION AND
POLYHYDROXYALKANOATE PRODUCTION IN A NOVEL ISOALTE OF
METHYLOBACTERIUM
4.1 Abstract
Background
Polyhydroxyalkanoates (PHAs) are a type of biodegradable plastic that are produced by several microorganisms as a carbon and energy storage molecule when the cells are under stress. They are a promising alternative to synthetic plastics but are currently too expensive for widespread use. The two largest hurdles to economic production are the cost of carbon feedstock and the cost of isolating and purifying PHAs.
Using methylotrophic bacteria that can utilize methanol as their sole carbon sources may reduce the cost of carbon feedstock. Indirectly secreting PHAs using phasin proteins that bind to PHAs and type I secretion system signal peptides to translocate PHAs across the cell membrane may reduce the cost of downstream processing.
Results
A novel strain of Methylobacterium was isolated and tested to show its ability to produce PHAs using methanol as a sole carbon source. A constitutive promoter known as cp8 was tested for use with a synthetic biology system in the Methylobacterium. The promoter demonstrated expression of superfolding green fluorescent protein (sfGFP) and was used in phasin protein genetic constructs to create a secreting strain of
Methylobacterium. When tested, there was no significant difference in the amount of
PHB secreted by the secreting and non-secreting strains. This may be because the 108
promoter was not strong enough to produce a noticeable amount of the secretion-inducing
fusion proteins, issues with the protocol used to prepare samples for analysis, or due to an
unidentified secretion mechanism in the bacteria.
4.2 Background
Polyhydroxyalkanoates (PHAs) are a class of biodegradable, biologically- produced plastics that accumulate in many microorganisms [1]. In native organisms, they act as carbon reserves that accumulate in response to nutrient deficiency or environmental stress [2]. When extracted, PHAs display properties similar to synthetic plastics, such as polypropylene [3]. Unlike synthetic plastics, however, PHAs are non-toxic, renewable and biodegradable [4,5]. In addition, the chemical structures of PHAs are variable, allowing them to be tailored for a range of applications in packaging, the medical field, agriculture, and elsewhere [6]. Consequently, there is great interest in using PHAs as an environmentally-friendly alternative to synthetic plastics [7].
Currently, the most significant hurdles to industrial-scale production of PHAs are
the costs associated with carbon feedstock for the microorganisms producing the PHAs
and the cost of extracting and purifying the PHAs from the microorganisms once they
have accumulated in the cell cytoplasm [8,9]. One possible solution to the cost of carbon
feedstock is to use methanol or methane rather than the traditional approach of using
plant-derived oils and sugars. Both methane and methanol are inexpensive, stable and
readily available feedstocks that are independent from seasonal variations and weather
conditions. They can be utilized by methylotrophic bacteria, many of which are known to 109
produce PHAs [10,11]. Thus, methane and methanol have been suggested as an
alternative, sustainable carbon sources for PHA production.
One well-known type of methylotrophic bacteria is Methylobacterium.
Methylobacterium are serine-pathway utilizing facultative methylotrophs that are simple
and inexpensive to culture. There are optimized fermentation protocols [12–14], genomic
information, and genetic tools [15–21] available for use with these bacteria. They have
also been shown to produce valuable products like PHAs, vitamin B12, pyrroloquinoline,
and carotenoids [17]. Strains of the most well-characterized species, Methylobacterium
extorquens, have been known to achieve PHA concentrations as high as 46% dry cell
weight (DCW) [14]. Methylobacterium strains normally produce polyhydroxybutyrate
(PHB), but can produce polyhydroxyvalerate (PHV) when propionate or valerate are
added as co-substrates [19,22]. These characteristics make Methylobacterium strains
organisms of interest for methanol-based biomanufacturing [23].
Current approaches to extracting and purifying PHAs are invasive, involving the disruption of cell membranes by mechanical, biological, or chemical means [24,25]. It is estimated that extraction and purification costs represent as much as 50% of the total process expense [26] and that these processes account for >90% of the adverse environmental impact of PHA production [8]. Although many alternative methods have been developed for PHA recovery, none of them have resulted in significant economic improvements in downstream processing [27]. Thus, there is a need for discovering a system that can improve isolation of PHAs.
One process for extraction of PHA granules that has been investigated in recent years is spontaneous liberation or secretion of PHAs. One study found that when E. coli 110
cells harboring Alcaligenes phbCAB genes were grown in a 2 x LB medium containing
21% glucose, that up to 80% of the cells spontaneously secreted PHB granules [26].
Another study used a combination of the phaCAB genes and the cloned lysis gene E of
bacteriophage PhiX174 from plasmid pSH2. It was shown that small PHB granules
exited the cells through the E-lysis tunnel structure [28]. Sabitova and colleagues
discovered a mutant of the marine oil-degrading bacterium Alcanivorax borkumensis SK2 that hyperproduced PHAs. The hyperproduction caused the bacteria to push PHA into the extracellular fluid when grown on alkanes [29]. Secretion of PHA granules, such as was
achieved in these studies, would negate the need to use harsh chemicals or energy-
intensive processes to lyse the cells open during an extraction process.
Another secretion system was developed by our group for E. coli by taking
advantage of a type I secretion system to translocate PHAs from the cytoplasm to the
extracellular medium [30]. Type I secretion systems, such as the hemolysin system in E. coli, are composed of ATP-binding cassette transporter (ABC transporter) and a
membrane fusion protein (MFP) that interact with an outer membrane polypeptide known
as the TolC protein [31]. Proteins, often toxins and virulence factors, that have a glycine-
rich C-terminus signal peptides are transported from the cytosol of the bacteria into
extracellular milieu [32]. Type I secretion systems are of interest as a method to secrete molecules produced within the cell. 111
The hemolysin system has been used to
secrete proteins and PHA granules. Previously, our
group demonstrated that when fusion proteins of
green fluorescent proteins (GFP) and the C-terminus
signal peptide of the hemolysin A (HlyA) protein
were expressed in a strain of E. coli that also
expressed the appropriate ABC transporter (HlyB)
and MFP (HlyD), GFP was secreted [33] . In a Figure 4.1. Schematic for PHA secretion. Phasin (blue) with attached HylA signal peptide (green) binds to PHA granule surface and the PHA-phasin-signal peptide subsequent study, a fusion protein of phasin proteins complex is targeted for type I secretion.
that bind to the surface of PHA granules and the HlyA signal was created. The phasins
bound tightly to and carried the PHA granules with them as they were secreted from the
cell (see Figure 4.1). Using this system, 36% of the total PHA produced in the secreting
strain was collected in the secreted fraction and 64% remained in the bacterial cell after
48 hours. In addition, total productivity was increased relative to the non-secreting strain
[30].
The secretion system was assembled using a synthetic biological engineering
method. Synthetic biological engineering is an approach to genetic engineering that seeks
to improve our ability to engineer life. A scientific understanding of the nature of DNA,
restriction enzymes, ligation, and polymerase chain reactions (PCR) combined in the late
20th century to make genetic engineering of organisms possible. Generally, however, the
process of combining genes of multiple organisms has involved expensive, unreliable,
and ad hoc approaches. A push to integrate engineering principles more fully into genetic
engineering since the early 21st century has led to ideas for simpler, more accessible 112
approaches [34]. One of the most significant ideas put forward is that of creating standardized, biological “parts” that can be assembled into genetic circuits [35].
Standardization allows streamlined research, as information related to DNA sequences can be shared rapidly and DNA can be synthesized to order. This allows for testing our understanding of biological systems; the creation of novel molecules and molecular systems for diagnostic assays and drugs; and development of new technologies that make it easier and more efficient to satisfy human needs [34,36]. One example of this type of standardization in biotechnology is the concept of BioBricks.
BioBricks are defined sequences of DNA used as biological parts. The standardized BioBrick assembly uses a DNA sequence flanked by the restriction sites
EcoRI and XbaI upstream and BcuI (formerly SpeI) and PstI downstream with no repetition of the restrictions sites in the DNA sequence. Since BcuI and XbaI are compatible, BioBrick segments may be pieced together sequentially due to the regeneration of the prefix and suffix restriction sites during each step of the process (see
Figure 4.2). The BioBrick system enables systematic assembly of DNA sequences, making it a convenient approach for the construction of the DNA for the PHA secretion systems used in this study [37]. To facilitate sharing of information, sequences of
BioBrick parts are available in an online registry maintained by the International
Genetically Engineered Machine (iGEM) Foundation at http://parts.igem.org/Main_Page.
The current study investigates the possibility of using a secretion system constructed from BioBricks in Methylobacterium.
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Figure 4.2. Diagram of the BioBrick standard assembly process. Two BioBrick parts, shown in the blue and green, are assembled by cutting the appropriate plasmid using restriction endonucleases that cut in the prefix and suffix surrounding the gene of interest. The appropriate linearized DNA fragments can be isolated, mixed, and ligated together to form a composite part of the individual DNA fragments. The restriction endonuclease sites in the prefix are EcoRI (E) and XbaI (X) and the restriction sites in the suffix are BcuI (B) and PstI (P).
4.3 Materials and Methods
4.3.1 Strains and Plasmids
Descriptions of strains and plasmids that were used to study PHA and phasin production and translocation are provided in Table 4.1. Electrocompetent XL 1-Blue
Escherichia coli (Stratagene, La Jolla, CA) were used as the host for assembly of
BioBrick parts and devices. Plasmids pSB1C3, pSB1A3 and pSB3K3 are BioBrick standard vectors for assembly and expression of BioBrick genetic devices [38]. The 114
broad-range vector plasmid pCM66 was used to transfer the competed BioBrick devices
into Methylobacterium via electroporation. Cultures of the Methylobacterium were
isolated from samples discovered in our laboratory.
Table 4.1. Strains and plasmids used in this study Strain/Plasmid Relevant Characteristics Reference E. coli Strains XL1-Blue endA1 gyrA96(nalR ) thi-1 recA1 relA1 lac glnV44 Stratagene F'[ ::Tn10 proAB+ lacIq Δ(lacZ)M15] hsdR17(rK- mK+ ) Methylobacterium Plasmids R pSB1C3 High copy BioBrick vector, pMB1 origin, Cm Registry pSB3K3 Medium copy BioBrick standard vector, p15A Registry origin, KanR pSB1A3 High copy BioBrick vector, pMB1 origin, AmpR Registry pLG575 pACYC184 derivative, HlyBD, p15A origin, CmR [39] pBHR68 pBluescript SK-, phbCAB genes from C. necator, [40] AmpR pRK2073 Helper plasmid, SpecR [] pCM66 High copy broad host range cloning vector in Addgene bacteria, ColE1 origin, KanR #45827 pCMHS1 pCM66 with constitutive promoter and superfolder This study GFP (sfGFP) pCMHS2 pCM66 with constitutive promoter, sfGFP, and This study HlyA pCMHS3 pCM66 with constitutive promoter, PhaP1Re, and This study HlyA pCMHS4 pCM66 with constitutive promoter, PhaP4MtO, and This study HlyA
4.3.2 BioBrick Design and Assembly
BioBrick parts utilized in this study included two phasin proteins (PhaP4MtO from
the methanotrophic bacteria Methylosinus trichosporium OB3b and PhaP1Re from
Cupriavidus necator H16 (formerly Ralstonia eutropha)), one signal peptide (HlyA); two
secretion system proteins (HlyB and HlyD), one reporter protein (sfGFP), a constitutive promoter, a ribosome binding site, and a terminator (see Figure 4.3). The PhaP4MtO 115
phasin protein gene was obtained using PCR by annealing forward and reverse
oligonucleotides to isolate DNA from Ms. trichosporium OB3b. PCR was also used to
add the BioFusion prefix and suffix to the phasin gene as overhanging ends on a second set of primers (see Table 4.2 for primers used). The PCR product was isolated by gel
electrophoresis and extracted using a Thermo Scientific GeneJET Gel Extraction Kit. The
DNA product was digested with EcoRI and BcuI, and ligated into pSB1C3. The ribosome
binding site and terminator (BBa B0030 and BBa B0015, respectively) were obtained
from the 2015 iGEM distribution kit. The promoter (cp8) was a constitutive promoter
created for use in Lactococcus lactis, but has been shown to be active in multiple gram-
negative bacteria [41]. Step-wise assembly of composite BioBrick devices was primarily
carried out in pSB1C3.
Figure 4.3 Schematic of PHA and sfGFP secretion genetic circuits. Promoter is a constitutive promoter; RBS is ribosome binding site; PhaP4MtO and PhaP1Re are phasin protein coding sequences from Ms. trichosporium OB3b and Cupriavidus necator, respectively; HlyA is the hemolysin A secretion. tag; B0015 is a double-stop terminator.
116
Table 4.2. Oligonucleotides (prefix/suffix overhangs are bolded; restriction enzyme sites are underlined) Oligo/Primer Description 5'->3' Source PhaP4MtOFOR Atggcaacccatttcaaagg This study PhaP4MtOREV Tcacgcgacgaaagtcgaga This study PhaP4MtOFOR_P Tctgaattcgcggccgcgttctagaatggcaacccatttcaaagg This study PhaP4MtOREV_S Tctctgcagcggccgctactagttcacgcgacgaaagtcgaga This study
4.3.3 Media Formulation and Growth Conditions
Overnight cultures of E. coli for genetic transformation were grown in a standard
LB Media (containing, per liter: 10 g Bacto-Tryptone, 5 g Bacto-yeast extract, 10 g
NaCl). Cultures of Methylobacterium were grown in 150-ml serum bottles with 50 mL of
NMS media solution (containing, per liter: 1.0 g KNO3, 1.0 g MgSO4 x 7H2O, 0.23 g
CaCl2 x 2H2O, 0.0005 g Na2MoO4 x 2 H2O, 0.002 g Ferric III ammonium citrate, 0.004
g EDTA, 0.006 ml HCl, 0.0014 g CuCl2 x 2 H2O, and 1 mL trace solution); trace solution
(containing, per liter: 0.5 g FeSO4 x 7 H2O, 0.4 g ZnSO4 x 7 H2O, 0.02 MnCl2 x 7 H2O,
0.05 g CoCl2 x 6 H2O, 0.01 g NiCl2 x 6 H2O, 0.250 g EDTA, and 0.015 g H3BO3); and
0.5 mL of phosphate solution (containing, per liter: 26 g KH2PO4 and 62 g Na2HPO4 x 7
H2O). To sterilize, the base NMS solution, trace solution, and phosphate solution were all
autoclaved at 121°C for 30 minutes. For solid media plates, 15 g of agar per 1 L of media
solution was added to trace solution prior to autoclaving. The media solution and
phosphate solution were mixed in a laminar flow hood after cooling. As a carbon source,
250 µL of methanol were added to the Methylobacterium cultures. Cultures were
incubated at 30°C, shaking at 200 rpm.
117
4.3.4 PHA Production in Methylobacterium
To asses PHB concentrations, cell growth was measured via optical density at 600
nm using a spectrophotometer. Cultures that enter the exponential growth phase and achieve an optical density of greater than 0.30 OD600 were assayed for PHB production.
PHB production was induced by incubation with 0.6% methanol in the absence of
nitrogen and magnesium. Cultures were centrifuged at 3452 rcf for 10 min, washed once
with nitrogen- and magnesium-free medium, re-centrifuged, and re-suspended in 50 mL
of nitrogen- and magnesium-free NMS medium. After 24 h of incubation, cultures were
harvested, immediately frozen at -80°C, and then lyophilized for PHA analysis.
4.3.5 Recovery of Secreted PHA
Techniques for secreted PHA recovery followed as described in Rahman et al.
2013 [30]. At 24 h after starvation was initiated, 0.01 M CaCl2 (final concentration,
Avantor Performance Materials, Inc. Center Valley, PA) was added to the bacterial
culture and mixed by inverting the tube several times. The tubes were then allowed to sit
for 10 mins at room temperature and then centrifuged at 54 rcf for 5 min. The supernatant
was removed and transferred to a fresh tube and the pellet was freeze dried. The supernatant was centrifuged at 3452 rcf for 10 mins and the pellet from the second centrifugation contained bacterial mass and non-secreted PHA. Secretion studies and
PHA analysis were conducted in triplicate samples. Statistical tests were performed using the Statistical Analysis System (SAS) software.
118
4.3.6 PHA Concentration Determination
PHA concentrations were determined using gas chromatography (GC). Sample
preparation was carried out in accordance with reported acid methanolysis procedures, as
described in Linton, et al. 2012 [42]. Dried cell pellets of approximately 15 mg
constituted a sample. Equal volumes (0.7 mL) of acidified methanol (0.03% H2SO4) and chloroform were added to each sample. These mixtures were vortexed and incubated at
100 °C for 3 h. After cooling to room temperature, 0.4 mL of distilled water was added, followed by vortexing for 10 min. Following a phase separation time of 20 min, the organic phase was transferred to a new vial for GC analysis.
GC instrument parameters were as follows. An HP 6890 Series II gas chromatography system was used with an HP-INNOWax cross-linked polyethylene
glycol capillary column with dimensions of 30 m x 0.25 mm ID x 0.25 µm film
thickness. The injection ratio was split to 1:20, with carrier gas argon flow rate of 3.8
mL/min. The initial temperature of 60 °C was held constant for 4 min, followed by a 15
°C/min increase to a final temperature of 250 °C, which was held constant for 5 minutes.
The flame ionization detector and injection port temperatures were maintained at 250 °C.
4.3.7 Scanning Electron Microscopy (SEM)
SEM protocols were followed as mentioned in Mortensen 2012 [43]. Briefly,
bacterial cultures were grown until they achieved exponential phase and fixed onto glass
cover slips. Samples were mounted on aluminum stubs and sputter coated with 10 nm
gold. SEM was carried out using a Hitachi S4000 SEM. 119
4.3.6 Fluorescence Studies
Cultures of bacteria were grown to an OD600 of approximately 0.3 A. 1 mL
samples were harvested from each culture. As a control, three 300µL samples of the
uninoculated media were placed into wells of a 96-well plate. Three 300µL samples each
of the bacterial cultures were also placed in wells of the same 96-well plate. Fluorescence
was read using a BioTek Synergy 2 well plate reader instrument, with an excitation
wavelength of 485/20 and an emission wavelength of 528/20. To adjust for bacteria
levels and auto fluorescence in the media, the following equation was used to produce a
corrected value:
= 𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑒𝑒 𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚 𝐹𝐹𝐹𝐹𝐹𝐹𝐹𝐹𝐹𝐹 𝐹𝐹 − 𝐹𝐹 𝐹𝐹 𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠𝑠 𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚 Where F is the level of fluorescence in 𝑂𝑂𝑂𝑂relative fluorescence− 𝑂𝑂𝑂𝑂 units (RFU) and OD is the
optical density of the sample at 600 nm.
4.4 Results
4.4.1 Isolation and Analysis of Methylobacterium sp.
During our research, we isolated a novel strain of Methylobacterium. The isolate
was initially found as a contaminant in cultures of another type of methylotrophic
bacteria, Methylosinus trichosporium OB3b. The OB3b strain was initially used due to its
ability to metabolize methane gas, but difficulties in culturing led to a greater focus on
Methylobacterium instead. Cell cultures that were contaminated with Methylobacterium
were identified by a red color in the cell pellet (see Figure 4.4) and a lack of the
particulate methane monoxogenase (pMMO) gene when tested using PCR. It is likely
that the Methylobacterium survived by metabolizing methanol produced by Ms. 120
trichosporium OB3b. The Methylobacterium was isolated from the initial culture using streaking on agar plates. Analysis of the 16s-region narrowed the isolate down to a few different species, including Methylobacterium extorquens (see Figure 4.5). Several SEM images of the new isolate were generated to demonstrate the morphology of the bacteria.
The images indicated that the isolate often exists in clusters of the bacteria, though it also grows as individual cells (see Figure 4.6). It is believed that this naturally-occurring strain of Methylobacterium can serve as a platform for production of PHAs from methanol.
Figure 4.4. Cell pellet of Methylobacterium isolate.
121
Figure 4.5. Phylogenetic tree showing the relationship of the Methylobacterium isolate used in the current study to other methylotrophic bacteria based on 16s sequencing. All species displayed except E. coli are from the rhizobiales order.
Figure 4.6. SEM images of Methylobacterium in various sizes of clusters.
The Methylobacterium isolate was analyzed for PHB production. The bacteria’s ability to synthesize PHB was initially confirmed using Sudan Black staining (see Figure
4.7). It is known that some strains of Methylobacterium have been reported to accumulate
PHAs in association with cell growth [22], while most require induction through nitrogen and magnesium limitation [14,44–49]. Initial analysis of PHB production indicated that higher quantities of PHB were accumulated with 24 hours of nitrogen and magnesium 122
limitation compared to nitrogen limitation and unstressed conditions (see Table 4.3). This indicates that the isolate is capable of producing PHB when placed in stressed conditions, most successfully when starved of both nitrogen and magnesium.
Figure 4.7. Image of Methylobacterium stained with Sudan Black to demonstrate the presence of PHB.
Table 4.3. Concentrations of PHB in cultures of Methylobacterium with nitrogen starvation, nitrogen and magnesium starvation, and normal (unstressed) conditions. Differences between the amount of PHB produced between all conditions were statistically significant (p < 0.05). Production mg/L Condition % mass of PHB in dry mass PHB Unstressed 0.30 ± 0.55 0.03 ± 0.05 Nitrogen Starved 4.54 ± 4.20 9.02 ± 13.097 Nitrogen and Magnesium 13.46 ± 5.14 100.95 ± 38.72 Starved
4.4.2 BioBrick Design and Construction
Initially, the methanotrophic bacteria Ms. trichosporium OB3b was identified as a
candidate for use in this study. Four phasin or phasin-like proteins were identified in the
sequences of the OB3b strain available on the Joint Genome Institute (JGI) and the
National Center for Biotechnology Information (NCBI) websites (see Figure 4.8), and
PCR primers were designed to isolate each of the four phasin proteins. One of four phasin
protein candidates from the type II methanotroph Ms. trichosporium OB3b was 123
successfully isolated from the genomic DNA and tagged with the BioBrick prefix and suffix using PCR. This phasin protein gene (PhaP4MtO) was then cloned into the vector plasmid pSB1C3 and transformed into E. coli using electroporation (see Figure 4.9).
Figure 4.8. Schematic of the third contiguous region of the Ms. trichosporium OB3b genome with locations of the four phasin proteins indicated.
124
Figure 4.9. (A) Gel electrophoresis depicting PhaP4MtO gene isolation (607 bp) from Ms. trichosporium by PCR; MW – Molecular Weight Marker (GeneRuler 1 k DNA Ladder, Thermo Fisher Scientific, Waltham, MA). (B) Gel electrophoresis depicting successful ligation of PhaP4MtO into pSB1C3. Extracted from cell culture and cut using EcoRI and BcuI.
The genetic circuits that include a phasin protein from Ms. trichosporium OB3b
(PhaP4MtO) and a phasin protein from C. necator (PhaP1Re) were constructed using step- wise assembly in E. coli plasmid pSB1C3 and then were inserted into pCM66 (see Figure
4.10). Two other genetic circuits were constructed with the same constitutive promoter and ribosome binding site as the phasin constructs, one with a sfGFP gene to test the strength of the promoter in Methylobacterium and one with the sfGFP gene fused to the
HlyA secretion tag. These four plasmids constitute the pCMHS series utilized in this study (see Figure 4.11). 125
Figure 4.10. A) Gel electrophoresis depicting PhaP1Re genetic circuit (987 bp) digested from pCM66 using NotI and EcoRI (digested fragment size is 3068 bp). B) PhaP4MtO genetic circuit (1015 bp) digested from pCM66 using NotI and EcoRI (digested fragment size is 3096 bp). MW – Molecular Weight Marker (GeneRuler 1 k DNA Ladder, Thermo Fisher Scientific, Waltham, MA).
Figure 4.11. Schematic of PHA and sfGFP secretion genetic circuits. The promoter is a constitutive promoter (cp8); RBS is ribosome binding site; PhaP4MtO and PhaP1Re are phasin protein coding sequences from Ms. trichosporium OB3b and Cupriavidus necator, respectively; HlyA is the hemolysin A secretion tag; B0015 is a double-stop terminator.
4.4.3 Promoter strength analysis
The pCMHS1 and pCMHS2 plasmids were used to investigate whether the
constitutive promoter cp8 was functional in methylotrophic bacteria. The cp8 promoter is 126
part of a series of synthetic promoters constructed for use in Lactococcus lactis, but which were shown to function in a variety of bacteria [41]. Fluorescence levels were measured in wild-type strains and in transformed strains of Methylobacterium. Statistical analysis of three separate cultures of each type indicated that there was a significant difference between the wild-type bacteria and the transformed strain (p < 0.05). This indicates that the constitutive promoter is functional in Methylobacterium (see Figure
4.12). Levels of sfGFP were significantly lower in Methylobacterium when compared with E. coli transformed with the same plasmid, however, with fluorescence registering at an average of 1,378,000 relative fluorescence units (RFU) in E. coli compared to an average of 4512 RFU in Methylobacterium. Thus, the cp8 promoter is active in
Methylobacterium, but not as effective at generating protein products as it is in E. coli.
Fluorescence Comparison
5000 B B 4500 4000 3500
3000 PhaP1_hlyA 2500 sfGFP 2000 sfGFP_hlyA 1500 1000
Relative Relative Fluorescence Units (RFU) 500 A 0 Samples
Figure 4.12. Fluorescence comparison between Methylobacterium not transformed with sfGFP (PhaP1_hlyA [pCMHS3]) and those with sfGFP (sfGFP [pCMHS 1] and sfGFP_hlyA [pCMHS 2]). Different letters indicate groups of data that were statistically significant in differences (p < 0.05) 127
In addition, growth rates of transformed and wild-type strains of
Methylobacterium were investigated using optical density. This was performed to
determine if the plasmids would drain a significant amount of cell resources, affecting
growth. The results indicated that there was no significant difference between strains of
cells that were transformed and the wild-type strain of Methylobacterium (see Figure
4.13).
Growth Curve Comparison 0.3 0.25 0.2 Wild Type 0.15 sfGFP
OD600 (A) OD600 0.1 PhaP1_hlyA 0.05 PhaOB4_hlyA 0 0 50 100 150 200 Time (h)
Figure 4.13. Cell growth curve of Methylobacterium comparing cell densities of wild type strains and transformed strains.
4.4.4 PHB Production Analysis
To investigate secretion of bioplastics from the methylotrophic bacteria, the pCMHS3 plasmid was co-transformed with the pLG575 plasmid into the new isolate and
PHB production was measured. This was compared with the strain containing an alternative plasmid (pCMHS1) that did not include the secretion machinery. PHB 128
measured inside the cell is defined as the internal fraction and PHB collected by the
CaCl2 precipitation of the media is defined as the secreted fraction. The non-secreting strain and secreting strain produced similar amounts of PHB in the internal fraction at
16.20 ± 1.78% of PHB in the dry cell weight (DCW) for the non-secreting strain compared to 12.09 ± 6.06% DCW PHB for the secreting strain (see Table 4.4). This was not a statistically significant difference (p > 0.05) in the amount of PHB present. The amount of PHB present in the internal fraction of both strains was comparable to previous studies with Methylobacterium strains [44,46,50].
Table 4.4. Production of PHB in secreting (pCMHS3 + pLG575) and non-secreting (pCMHS1) strains of Methylobacterium at 24 h % mass of PHB in dry mass Production mg/L PHB Strain Secreted Non- secreted Secreted Non-secreted Total fraction fraction fraction fraction production Non- 12.36 ± 2.56 16.20 ± 1.78 82.10 ± 4.22 152.06 ± 13.78 234.16 ± secreting 16.57 Secreting 10.68 ± 2.36 12.09 ± 6.06 31.26 ± 9.28 75.40 ± 21.11 106.66 ± 19.01
The PHB secreted fractions were analyzed for the secreting and non-secreting strains. PHB harvested in the secreted fractions of the non-secreting strains was 12.36 ±
2.56% DCW PHB, while the secreting strain produced 10.68 ± 2.36% DCW PHB in the secreted fraction. The amount of PHB present in the secreted fraction of the secreting strain was not statistically significant compared to the non-secreting strain (p > 0.05).
Relative to the total amount of PHB produced (mg/L PHB), 30% of the PHB in the secreting strain was collected in the secreted fraction while the remaining 70% was collected in the internal fraction. This was similar to the percentage of PHB collected in the secreted fraction in E. coli during our previous study, at 36% [30]. However, 35% of 129
the PHB in the non-secreting strain of Methylobacterium was collected in the secreted
fraction. This also indicates that the secreting and non-secreting strains had similar
amounts of PHB in the secreted fraction.
There are at least two viable explanations for why the secreting and non-secreting strains of Methylobacterium were similar. First, although it has not been reported in literature thus far, it is possible that the cells have a native system for secretion of PHAs.
Second, it could be that all PHB detected in the secreted faction was contained inside of cells that were lysed during preparation for GC analysis. A previous study using the same secretion system also observed PHB released by lysed E. coli cells in the non-secreting strain’s secreted fraction [30]. During this study, pellets of samples collected using CaCl2
precipitation and low speed centrifugation were a vivid red, the color of the cell pellets in
the non-secreted fraction. This likely means that the majority of matter collected was cell
biomass. In addition, it is possible that the Methylobacterium cell walls may be more
fragile than the cell walls of the E. coli strains used in previous studies, rendering the
harvesting procedure ineffective at discerning differences between secreted and non-
secreted fractions due to cell lysis. Thus, PHB in the secreted faction of non-secreting
strains were mostly likely either present due to lysed cells or because of native secretion
systems.
If it is the case that all PHB was present due to lysed cells, then it would indicate
that the secretion system is not effective in Methylobacterium in its present form. Future
studies would need to evaluate if using promoters known to have a higher efficiency in
Methylobacterium, like the PmxaF promoter, would prove more effective for expressing
both the HlyBD secretion system and the fusion proteins of phasins and HlyA signaling 130
peptides. Since the phasin gene used in this study was originally obtained from E. coli,
codon optimization for optimal protein expression in the new isolate could also lead to
improved production of phasin proteins. This is achieved by altering the DNA sequence
so that the mRNA produces the same amino acid sequence, but uses the tRNAs that the
species of bacteria favors. Thus, the resources for producing the protein are more likely to
be available, allowing higher quantities of protein to be assembled. Improved expression
could lead to a more visible demonstration of secretion.
Using native phasin proteins could also improve secretion in methylotrophic
bacteria. Since these proteins are naturally produced by the bacteria, they may be
produced more effectively than non-native proteins, or may interact more successfully
with PHA granules in their native environment. The Joint Genome Institute (JGI)
database indicates that there is at least one native phasin protein present in
Methylobacterium extorquens AM1, which is closely related to the isolate characterized
in this study. A phasin protein (PhaP4MtO) from the related type II methanotroph
Methylosinus trichosporium OB3b was isolated using PCR and converted into a
BioBrick. Testing of the hlyA- PhaP4MtO fusion protein in a well-characterized system
like E. coli will give greater insight into the protein’s ability to induce secretion of PHAs.
Isolation or synthesis of other potential phasin proteins from Mtb. extorquens AM1 or the
Methylobacterium isolate used in this study would also allow characterization of their
interactions with PHA granules and analysis of their effectiveness in inducing secretion.
It is possible that the native phasin protein would prove more effective than phasin
proteins from C. necator at linking the hlyA secretion tag to PHA granules. 131
4.5 Conclusion
A novel strain of Methylobacterium was isolated and tested for PHB producing
capabilities. A constitutive promoter (cp8) was tested in the isolate using the pCM66
plasmid and a sfGFP protein. Gene expression was measured by fluorescence levels,
which indicated that the promoter worked in Methylobacterium, though not as well as in
E. coli. Once expression of recombinant proteins was confirmed, a recombinant synthetic
biology system was assembled and tested with the intent of causing secretion of PHB
granules. There was, however, no significant differences between PHB produced and
secreted by the secreting and non-secreting strains, necessitating future work to improve
the secretion system in Methylobacterium. If achieved, secretion of PHB should help in
downstream processing, negating the need to use expensive methods to lyse cells,
reducing costs in downstream processing.
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CHAPTER 5
GENERAL CONCLUSIONS
The overall objective of this research was to investigate the viability of alternative carbon sources, like methanol and food waste, and a novel secretion system for use in production of polyhydroxyalkanoate (PHA) bioplastics.
Specifically, we aimed to analyze the potential of utilizing food waste as a carbon source for PHA production; characterize a novel strain of Methylobacterium; design and construct a BioBrick library of phasin proteins, signal peptides, hemolysin transport proteins, green fluorescent protein (GFP) parts, and composite devices for investigating secretion in Methylobacterium; and monitor the translocation of PHAs in
Methylobacterium using gas chromatography (GC). These aims were all addressed.
A novel strain of Methylobacterium was isolated and characterized for PHA production. The isolate was identified as a Methylobacterium using 16s region sequencing and comparison. SEM images were generated to demonstrate cell morphology. Assays were performed to analyze the bacteria for PHA production and conditions necessary to produce noticeable amounts. It was discovered that the bacteria produce polyhydroxybutyrate (PHB) under stressed conditions of nitrogen and magnesium starvation.
A series of BioBricks and composite parts were constructed to test secretion in
Methylobacterium. These were constructed in E. coli, then transformed by electroporation into Methylobacterium in the wide-host vector plasmid pCM66. First, the sfGFP constructs indicated that the cp8 promoter that was used in these constructs could 139 drive expression of proteins in Methylobacterium, though not as well as in E. coli. Then, the PhaP1Re and HlyA signal peptide fusion protein construct was used in conjunction with the secretion system plasmid pLG575 to test for secretion in Methylobacterium.
There was no significant difference between the amount of PHB produced and secreted by the secreting and non-secreting strains. These results indicate that the phasin protein- based secretion system made no noticeable difference in PHA secretion in
Methylobacterium as it is currently constituted. Future work is necessary to evaluate if improvements in the secretion system will render it viable in Methylobacterium. 140
CHAPTER 6
ENGINEERING SIGNIFICANCE AND FUTURE RESEARCH
This section outlines specific ideas for working with food wastes as carbon sources for polyhydroxyalkanoate (PHA) production and improving our understanding of the new isolate of Methylobacterium and secretion of PHAs with this isolate. Some of the most promising food wastes and processes for producing PHAs from them are discussed and some future directions are pointed out for studies using these carbon sources. As for the methylotrophic bacteria, characterization of the novel isolate of Methylobacterium used in this study will allow us to evaluate its potential for bioplastic production.
Improving the efficiency of the phasin protein secretion system is important in engineering applications because it could lead to improved economic viability of an important class of bioplastic. There are several ways that cellular product recovery by secretion can be improved and that the isolate of Methylobacterium can be characterized.
Production of PHAs using food waste is likely to prove both beneficial and challenging. Associated costs of production using the food wastes discussed are difficult to determine, since most studies did not perform life cycle assessments or other economic analyses. The best-characterized food waste carbon source for PHA production was dairy whey, which was the subject of one life cycle assessment [1]. Some of the most promising studies using pure cultures involved digesting whey with the halophilic archaeal species Haloferax mediterranei. This species was shown to thrive in pure cultures without sterilization due to a high saline content in the media used, producing moderate to high amounts of polyhydroxybutyrate (PHB) and polyhydroxybutyrate-co- 141
valerate (PHBV) [2,3]. When whey was used to feed strains of recombinant E. coli, very high levels of PHB production were observed, including 96.2 g PHB / L in a 6.6 L bioreactor during one study [4]. The primary disadvantages of using whey were generally the high energy requirement for fermentation and a relatively low yield of PHB compared to the amount of whey available (see Table 6.1). Use of Haloferax mediterranei and recombinant E. coli, however, did seem to be capable of producing high yields and may warrant further consideration for PHA production from dairy whey.
Table 6.1. Advantages and disadvantages of different food wastes in PHA production. Food waste Advantages Disadvantages Dairy whey Extensive pretreatment not required Energy requirement for fermentation Comparable ecological imprint to Relatively low amounts of PHA petroleum plastics output per kg whey input Some studies showed high volumetric productivities of PHAs Some studies resulted in production of PHAs other than PHB Tested at larger scales Starch Minimal pretreatments are necessary Moderate levels of biomass Waste oil No pretreatment necessary Moderate to low levels of biomass Tallow resulted in PHBV production Natural antifoaming agents Spent coffee High volumetric productivity Natural foaming agent grounds oil Use of n-hexane in most extraction procedures Lignocellulosic One study showed high Extensive pretreatment necessary waste productivities Generally experienced low levels of cell growth Sugar industry waste Moderate levels of biomass Moderate levels of PHA production Legume waste Moderate levels of biomass Food scraps Can produce PHAs and their Composition varies precursors in anaerobic cultures Complex waste--cannot be used directly by pure cultures Food processing Can produce PHAs and their Complex waste--cannot be used wastewater precursors in anaerobic cultures directly by pure cultures
142
There were a few other carbon sources and processes that showed good potential.
One promising food waste was spent coffee grounds oil, which was used to feed strains
of Cupriavidus necator. Production as high as 49.3 g PHB / L was observed in 2 L
bioreactors during one study [5], among the highest amount observed with native-PHA producing organisms using food waste. The main challenges with spent coffee grounds oil was that the oil is a natural foaming agent and that hazardous materials, like n-hexane, were generally used during the oil extraction process (see Table 6.1). Antifoaming agents can be used to deal with the foaming tendencies of the oil, and one study demonstrated that supercritical fluid extraction could be used to extract the oil using CO2 instead of n-
hexane, which reduces both concerns. Producing PHAs using Azotobacter chroococcum,
a bacteria that could digest starch directly, negated the need for pretreatments on starch,
and displayed relatively high volumetric productivities, thus showing promise for future
investigation [6]. Lignocellulosic wastes proved to generally be the least promising food waste carbon sources for PHA production, due to the need for intense pretreatments to reduce toxicity and low levels of cell growth (see Table 6.1). One study, however, used sugars extracted from ground wheat straw and saw production levels as high as 105.1 g
PHB / L. Though promising, extensive pretreatment was still required to render the straw digestible to the Burkholderia sacchari strain used in the study [7]. While spent coffee
grounds, starch, and ground wheat straw show promise for producing PHAs and deserve
further examination, most other studies using pure cultures demonstrated that a variety of
food sources could be used to produce PHAs, but that relatively low total amounts of
PHAs were produced. 143
Studies in anaerobic mixed microbial cultures (MMCs) were valuable because
they did not consume large amounts of energy for sterilization and because they could
digest complex wastes without extensive pretreatment. Lower cell concentrations
(resulting in lower volumetric productivity) and concerns over the quality of PHAs
produced in anaerobic digestion are the major concerns with MMC production of PHAs.
An approach to blending the benefits of both pure cultures and MMCs involved using
anaerobic digestion in MMCs as a pretreatment for food wastes. During the initial stages
of anaerobic digestion, food wastes were converted into volatile fatty acids (VFAs).
Those VFAs were then harvested and fed to pure cultures, generally resulting in higher
volumetric productivities compared to those seen in MMCs. This indirect coupling
approach faces challenges in harvesting VFAs and transferring them to the pure culture
fermenters without incurring prohibitive expenses in energy and time. It is also
challenged by the fact that VFAs in high concentrations are toxic to most bacteria. Fed
batch fermentation helped alleviate this latter concern by making only small amounts of
VFA available at first, then occasionally adding more VFAs via the feed media as the
previous VFAs were consumed by the bacteria. By so doing, concentrations of VFAs
were kept low, but consistently present throughout the fermentation process. Using
indirect coupling approaches to digest food wastes and producing PHAs shows promise
and deserves further attention, particularly fed-batch approaches and attempts to implement this approach at larger scales.
An important aspect of creating PHAs using food wastes that needs further investigation is scale-up of production. Most pure-culture studies examined used either shaking flasks or bioreactors that contained less than 10 L of media. Among the native- 144
PHA-producing pure culture studies, the largest volume used was with the halotolerant bacteria Haloferax mediterranei, in a 330 L bioreactor with a working volume of 220 L
[2]. The only other study using native PHA producers to use a larger volume than 10 L involved an unidentified highly osmophilic organism in a 42 L fermenter [8]. Among studies using recombinant bacteria, most were performed in bioreactors, but at sizes less than 10 L. One study, however, did scale up production using recombinant Escherichia coli with dairy whey, operating at volumes of 30 L and 300 L. Volumetric production as high as 10.3 g PHB / L was observed, demonstrating that the process could successfully be scaled up [9]. Another study used recombinant E. coli with malt waste and soy waste as the carbon sources in a 15 L fermenter, though relatively low concentrations of PHAs were produced [10]. In general, more studies need to be performed at larger scales using food wastes as carbon sources for PHA production with pure cultures.
Mixed microbial cultures (MMCs) faced similar issues with scale. Sequencing batch reactors (SBRs) generally were operated at a 1 L working volume [11,12], as were at least two other reactor types [13,14]. One SBR used to digest olive oil mill pomace was 12 L, however, which saw moderate success in producing PHAs [15]. The MMC
reactor with the largest volume was a vertical submerged membrane bioreactor operating
at a working volume of 1,333 L [16]. Otherwise, all MMCs were operated in bioreactors
with a capacity less than 5 L or in shaking flasks, as were all studies using volatile fatty
acids produced by MMCs to feed pure cultures. More studies with larger bioreactors
should be performed with MMCs.
Shifting discussion to the production of PHAs from methane and methanol,
studies to further characterize the isolate of Methylobacterium and its capabilities need to 145
be performed to assess its usefulness in bioplastic production. In-depth DNA fingerprinting could clarify the isolate’s relationship to other strains of
Methylobacterium. In addition, it would be advisable to test the bacteria’s ability to metabolize alternative carbon sources and examine their impact on polyhydroxyalkanoates (PHA) production. Many strains of Methylobacterium are capable of synthesizing hydroxyvalerate when propionate or valerate are added as co-substrates
[17,18]. One strain has even been shown to also produce hydroxyheptanoate and hydroxyoctanoate when citrate or propionate were added as co-substrates [19]. Similar studies could be conducted with our isolate to test what types of PHAs are produced, particularly once its relationship to well-characterized strains of Methylobacterium is established.
Continuing the development of a synthetic biology toolbox for alphaproteobacteria based on Methylobacterium species will provide an improved platform for a methanol-based biomanufacturing. Promoters are a sequence that regulates transcription of DNA into RNA. In this study, we utilized the cp8 constitutive promoter, which had not previously been used in methylotrophic bacteria. Characterizing promoters for use in Methylobacterium species, such as the constitutive promoters from the same set as cp8 or native promoters from Methylobacterium and other methylotrophic bacteria would increase our ability to control levels of production of desired compounds.
Inducible promoters have been developed using the PmxaF promoter from
Methylobacterium extorquens AM1, however, more could be developed. Having an array of inducible promoters would be useful for fine tuning control over production in methylotrophs. Promoters that proved highly effective in Methylobacterium could then be 146
tested with our secretion study to see if improved production results in a noticeable
amount of PHA secretion.
Another useful component for use in a synthetic biology toolkit would be the
discovery and characterization of more origins of replication beyond the phage-based
options currently in use. Origins of replication are a part of plasmid vectors that allow
plasmid DNA to replicate in bacteria. Most often, origins of replication that are based on
viral DNA sequences have been used in methylotrophic bacteria, since they work in
many different types of bacteria. The pLG575 plasmid used in this study and at least one
previous study used the E. coli p15A and pBR322 origins, respectively, indicating that E.
coli origins of replication may be utilized in methylotrophic bacteria [20]. Since E. coli is
a model Gram-negative bacterium for synthetic biology, plasmid vectors that can work in both E. coli and methylotrophic bacteria would allow construction of genetic constructs in the former and an easy transition to use in the latter.
Improvement of secretion could be achieved through codon optimization. Codon optimization is a method of DNA design where sequences for the same amino acids are changed to match the codon option that the specific organism in question is most likely to use. This has been shown to improve protein expression and functionality, and could increase the production of foreign phasin proteins like the Cupriavidus necator phasin protein PhaP1Re in Methylobacterium. Type I secretion systems have not been investigated in methylotrophic bacteria before, and so the hemolysin secretion system components are also a target for codon optimization that could improve the bacteria’s ability to secrete PHA granules. Both the phasin protein and the type I secretion system 147
components have an impact on the functionality of the secretion system, and codon optimization could improve expression of these important proteins.
In addition to codon optimization, characterization of native phasin proteins could lead to optimized secretion in methylotrophic bacteria. In Methylobacterium extorquens
AM1, which is closely related to the isolate characterized in this study, the JGI database indicates that there is at least one native phasin protein present. The type II methanotroph
Methylosinus trichosporium OB3b has four coding sequences that may potentially be for phasin proteins. Strain OB3b was originally the target organism of this study, and one phasin protein (PhaP4MtO) has already been successfully isolated from this organism and converted into a BioBrick. Further testing of the hlyA- PhaP4MtO fusion protein in a well- characterized system like E. coli would give greater insight into its ability to induce secretion of PHAs. Isolation or synthesis of other potential phasin proteins would also allow characterization of their interactions with PHA granules and analysis of their effectiveness in inducing secretion. It is possible that the native phasin protein would prove more effective than phasin proteins from C. necator at linking the hlyA secretion tag to PHA granules.
If achieved, mastery of the type I secretion system in Methylobacterium will enable application in other methylotrophic bacteria. Methylobacterium cannot digest methane gas, however, limiting their flexibility. Methanotrophic bacteria that produce
PHAs can utilize methane gas, including methane from waste sources like biogas, that
Methylobacterium cannot use to produce bioplastics. Using their methanotrophic capabilities in combination with the secretion system would create another sustainable platform for PHA production that would be connected to the concept of a biorefinery (see 148
Figure 6.1). The type II methanotrophs Methylosinus trichosporium OB3b and
Methylocystis species are potential targets that utilize methane gas and produce PHAs. If the secretion system works in both Methylobacterium and either Methylosinus or
Methylocystis, further studies could be carried out with co-cultures of multiple secreting methylotrophic bacteria, which could be a possible approach for improved production of
PHAs from methane and methanol.
Figure 6.1. Biorefinery proposed by the Sustainable Waste-to-Bioproducts Engineering Center (SWBEC) at Utah State University. Section highlighted in red is the potential area that production of PHAs using methanotrophic bacteria would fit into the biorefinery.
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APPENDICES 153
APPENDIX A
BIOBRICK PARTS AND SEQUENCES
Table A.1. Nucleotide sequences of codon optimized protein coding regions for Ms. trichosporium OB3b Gene Nucleotide Sequence Origin Pha1 ATGGTTGCCACTTTCGAGACCGTTCAGCAGCTGGGCAA Ms. GGAGCAGTTCGAGGCGGTGTCCGCGGCTACCGCCGCCG trichosporium TCACCAAGGGCTGGCAGACGATCGCCGCCGAGGCCACC OB3b GATTATTCGAAAAAGTCGTTCGAGAAGAGCCGCCTGCT CGCGGAAAAGCTTATCGGCGTCAAGAAGATCGACGAAG CTCTTCAGCTTCAGTCGGACTTCGCCAAGGGCGCCTATG AGGATTTCATCGCCGAGGCGACCAAGATCGGCGAGCTC TATTCCAGCCTCGCCAAGGAAGCCTTCAAGCCCGTCGA GACGGCAGCGTCCAAGGTCTACACCTCGGCCGAGTGA Pha2 ATGGCCAATCTGAACTTCGATGATTTTCGCGAGCTCGGA Ms. AAATTTCAGCTCGAAGCCATGAACTCGGCAGCGAGCAC trichosporium GACGACGAGGGGCCTCAAGGCGATCGCCGCCGAAGCG OB3b ACCGATTATTCCAAGCAGTCGCTCGACAACAGCCGCGC CTATTTCGAGAAGCTGCTGCGCGTGCAGAAGATCGACG ACGTGGTCGAGCTGCAATCCGAGTTCTGCCGCGCCGCC TATGGCGACTTCTTCGCGCGCGCCTCGCGCGTCGGCGA GCTGTGCTCCAATCTCGCCAAGGAAGCTTTCGTCAATGC GCAGACGACGGGCGAGAGCGCGACCAAGGCCGCCACG GAGGCGACGACCAAGGCGCTCTCGGGGCTCGGCGAGCA GACCGAGCAATTCGTCTCGAAGGCGCAGCAGGCCGCCG GCGGTCAGCGCTGA Pha3 GTGAGCGAACCGATCTTCCAGATTCCGAACGAAGTGCG Ms. CGATTTCGCCGAAAAGAGCATGGAGCAGGCTCGCAAGG trichosporium CGTTCGAAGGCTATGCCGGCGCCGCGCAGAAGGCGATC OB3b AGCTCCCTCGAGACGAGCACCTCGACCTTTCAGACCGG CGCGACCGATGTGAGCGCCAAGGCGCTCGGCTACGCCG AGGCCAATGTCAACGCCGCCTTCGAGCTCGCGCGCAAG CTGCTGCATGCGAAAGACCCGCAGGAAGTGCTCACCCT GCAAACGGAATATGTGAAGACCCAGGTCGAATCGATTC AGTCGCAGGCCAAGGATCTCGGCGCAGCGCTGCAAAAG GCCGCGACCGGTCGGTGA 154
Table A.1 continued. Gene Nucleotide Sequence Origin Pha4 ATGGCAACCCATTTCAAAGGTCCCGCGAAACCGCGCGCGCCCA Ms. AGGTCGCGCCGATCTCCCTGAAAGACGAGCTCGACGCCGACGA trichosporium GCCGGTCGCCCCGCCCGCCGACACGCCCTTCGCCGCTCCGTCG OB3b GACACGACCTTCGCCGACACGATCGTGAGCGCGCTCGAGGCCG AGCCGGTCGCGAGCGAAGCTGTCGCCCCGGCCGAGATCCTCGA GCCCGCGACCGCGGCGAATATCATCATCGAGCCGCTCGCCGCG GTCGAGCCCGAGCCGAGTCCCGCCGCGGCCGAGCCGCTCGTCA TCGTCGCCGCCGTGACGAGCGCCGCGCCGGCGACGCTCGACGC CAGCGCGCTGCCGCTGAAGACGCTCGATCTCTTCAACGAGAAC GCCGCCGCCATGATGGACTTCGCGCTGGCTCTGGGCGCCGCGA AAACGGTCGGCGACGCGCTCGAGCTGCAGTCGCGCTTCGCCAG CGAGCGCTATTCGAGCCTCGTTCGCCAGGCCGGCGAGGTCGCC GAGCTGACGCGGCGGCTGGCGTTCCAGCACGCGCCGTTCAAGC TGCGGGTCTCGACTTTCGTCGCGTGAACTAGTAGCGGCCGCTG CAGTCT PhaP1 ATGATCCTCACCCCGGAGCAGGTCGCCGCGGCGCAGAAGGCC Ralstonia AATCTGGAGACCCTCTTCGGCCTGACCACGAAGGCGTTCGAAG eutropha GCGTCGAGAAACTGGTCGAGCTGAATCTCCAGGTCGTCAAAAC CTCGTTCGCGGAGGGCGTGGACAATGCCAAGAAGGCCCTGTCG GCCAAGGATGCGCAGGAGCTGCTGGCCATCCAGGCCGCGGCC GTGCAGCCCGTCGCGGAAAAGACCCTGGCCTATACCCGTCACC TGTACGAGATCGCGTCGGAGACGCAGTCGGAGTTCACCAAGGT TGCGGAGGCCCAGTTGGCCGAGGGCAGCAAGAACGTCCAGGC CCTGGTCGAGAACCTCGCCAAGAATGCGCCGGCGGGCTCCGAG TCCACGGTCGCTATCGTGAAAAGCGCGATCAGCGCCGCCAACA ACGCCTATGAGTCGGTCCAGAAGGCCACCAAGCAGGCGGTCG AAATCGCGGAGACAAACTTCCAGGCCGCCGCGACCGCGGCGA CCAAAGCTGCGCAGCAGGCCTCGGCGACGGCGCGGACCGCGA CGGCGAAGAAGACCACCGCGGCC GFPuv ATGGCCAGCAAGGGCGAGGAGCTCTTCACAGGCGTGGTGCCG ATCCTGGTCGAACTCGATGGCGACGTCAATGGCCATAAGTTCA GCGTCAGCGGAGAGGGTGAGGGCGACGCGACCTACGGCAAGC TCACGCTGAAATTCATCTGCACCACGGGCAAGCTGCCGGTGCC GTGGCCAACACTCGTCACCACCTTCTCGTACGGCGTCCAGTGC TTCTCGCGCTACCCGGATCACATGAAACGTCACGATTTCTTCAA GTCGGCCATGCCGGAAGGCTATGTGCAGGAGAGAACGATCTC GTTCAAAGACGACGGCAACTATAAGACTCGCGCCGAGGTCAA GTTCGAGGGCGACACGCTCGTCAATCGCATCGAGTTGAAGGGG ATCGACTTCAAAGAGGACGGCAACATCCTCGGCCATAAGCTGG AGTATAATTATAACTCGCACAACGTGTATATCACCGCCGACAA GCAAAAGAATGGCATCAAGGCGAACTTCAAAATCCGCCACAA CATCGAGGACGGCTCCGTCCAGCTGGCGGATCACTACCAGCAG AACACCCCGATTGGCGACGGCCCGGTCCTCCTCCCCGACAATC ATTATCTCAGCACTCAGTCGGCGCTCTCGAAGGACCCCAACGA GAAGCGCGACCACATGGTCCTGCTGGAGTTCGTCACCGCGGCG GGCATCACCCATGGCATGGATGAGCTGTATAAGTAG
155
Table A.2. Nucleotide sequences of Hemolysin Genes used in this study
Gene Nucleotide Sequence References HlyA CTCGCCTATGGCTCGCAGGGTGACCTCAATCCGCTCATCAACGAGATCTCT AAGATCATCTCGGCGGCGGGCTCGTTCGACGTCAAAGAGGAGCGTACGGC GGCGTCGCTGCTCCAGCTGTCGGGAAACGCGTCGGACTTCTCGTATGGCCG CAACTCGATCACCCTCACCACCTCGGCC HlyB ATGATGTCGAAGTGCTCGTCGCATAACTCCCTGTACGCTCTGATCCTCCTC GCCCAGTATCATAACATCACGGTGAACGCTGAGACGATCCGTCATCAGTAT AACACCCACACCCAGGACTTCGGCGTCACGGAATGGCTGCTGGCGGCGAA GAGCATCGGACTGAAGGCCAAGTATGTCGAAAAGCATTTCTCGCGCCTCTC GATCATCTCCCTGCCGGCTCTGATCTGGCGCGATGATGGCAAGCACTACAT CCTCAGCAGGATCACCAAAGACTCATCGCGCTATCTGGTCTATGATCCCGA GCAGCACCAGTCCCTCACGTTCTCGCGCGACGAGTTCGAGAAGCTGTACCA AGGTAAGGTGATCCTGGTCACGTCGCGCGCCACCGTCGTCGGAGAGCTGG CCAAGTTCGATTTTTCCTGGTTCATCCCCTCGGTCGTCAAGTACCGCCGCAT CCTGCTCGAGGTGCTCACCGTGTCCGCCTTCATCCAGTTCCTCGCGCTCATC ACGCCCCTCTTCTTCCAGGTGGTCATGGACAAGGTGCTGGTCCATCGTGGC TTCTCCACCCTCAACATCATTACCATCGCCTTTATCATCGTCATCCTGTTCG AGGTCATCCTGACGGGCGCGCGCACCTATATCTTCTCTCACACCACCTCGC GCATCGACGTTGAGCTGGGCGCGAAATTGTTCCGGCACCTCCTCGCGCTCC CAGTCAGCTATTTCGAGAACCGCCGCGTGGGCGAGACCGTCGCGCGGGTG CGCGAACTGGAGCAGATCCGCAACTTCCTGACTGGTCAGGCTCTCACCAGC GTCCTGGACCTCTTCTTCAGTGTGATCTTCTTCTGCGTGATGTGGTACTATT CCCCCCAGCTGACGCTGGTCATCCTGCTCTCCCTCCCCTGCTACGTTATCTG GAGCCTGTTCATCAGCCCCCTCCTGCGCCGCCGCCTGGACGATAAGTTCCT GCGCAATGCCGAGAATCAGGCTTTCCTCGTGGAGACGGTGACCGCGATCA ACACGATTAAGAGCATGGCCGTGTCGCCCCAGATGATTGCCACGTGGGAC AAGCAGCTGGCCGGCTATGTGGCCAGCAGCTTCCGCGTGAATCTCGTCGCG ATGACCGGCCAGCAGGGCATCCAGCTCATTCAGAAGAGCGTGATGGTGAT CTCGCTCTGGATGGGCGCGCATCTGGTGATCTCGGGTGAAATCTCGATCGG CCAGCTGATTGCCTTCAACATGCTCGCCGGCCAGGTGATTGCCCCGGTGAT CCGGCTCGCGCACCTCTGGCAGGACTTCCAGCAGGTCGGGATCTCGGTCGA GCGCCTGGGCGACGTGCTCAACACCCCGGTCGAGAAGAAGTCGGGCCGCA ACATCCTGCCGGAGATCCAGGGCGATATCGAGTTTAAGAACGTGCGCTTCC GCTACTCCTCGGACGGCAACGTCATCCTGAATAACATCAACCTCTATATTT CGAAGGGCGACGTGATCGGTATCGTCGGCCGTTCGGGCTCGGGCAAGTCG ACCCTCACGAAGCTGCTCCAGCGCTTCTATATCCCGGAGACGGGCCAGATC CTCATTGATGGGCATGACTTGTCGCTGGCGGACCCGGAGTGGCTCCGCCGC CAAATTGGCGTCGTCCTCCAGGAGAACATTCTGCTCAATCGCTCGATCATT GATAATATCACGCTGGCCTCGCCCGCCGTCAGCATGGAGCAGGCCATCGA AGCGGCGCGGCTCGCCGGCGCCCATGACTTCATCCGGGAACTCAAGGAGG GCTACAACACCATCGTCGGCGAGCAGGGCGTGGGCCTCAGCGGCGGCCAG CGGCAGCGCATCGCGATCGCCCGCGCTCTCGTGACGAATCCCCGCATCCTC ATCTTCGACGAGGCGACCTCCGCCCTGGATTACGAATCGGAGAACATCATC ATGAAAAATATGTCGCGCATCTGCAAGAATCGCACCGTGATCATCATCGCT CATCGCCTGAGCACGGTCAAGAACGCGAACCGCATCATCGTGATGGACAA CGGCTTCATCTCGGAGGATGGCACGCATAAAGAGCTCATCTCGAAGAAGG ATTCGCTGTACGCCTACCTGTATCAGTTGCAGGCCTGA 156
Table A.2 continue. Gene Nucleotide Sequence References HlyD ATGCGTTTCTATATGAAGGGCCTGTGGGATCTGGTCTGCCGCTACAAGACG GTCTTCTCGGACGTCTGGAAGATCCGCCACACCCTCGACGCGCCGGTCCGT GAAAAAGACGAGTACGCTTTCCTCCCGGCCCACCTGGAGCTGATCGAAAC GCCGGTGTCGCGTCGCTCCCACTTCGTCGTCTGGTCGATCCTGCTTTTCGTC ATCATTTCGCTCCTGCTCTCGGTGCTCGGCAAGGTCGAGGTGGTGAGCGTC GCGAATGGCAAGTTCACGCACTCCGGTCGCTCCAAGGAGATCAAGCCGAT CGAGAACGCCATCGTCGAGAAGATCATGGTGAAGGACGGCTCGTTCGTGA AGAAGAACGACCCGCTGGTCGAACTCACGGTGCCGGGCGTGGAGTCGGAC ATCCTGAAGTCGGAGGCCTCGCTGCTGTATGAGAAAACCGAGCAGTACCG CTATGCGATTCTCTCGGAGTCGATCCAGCGCAACGAGCTCCCGGAGATCCG CATCACCGATTTCCCCGGTGGGGAGGACAATGCCGGCGGCGAGCATTTCC AGCGCGTGAGCTCGCTCATCAAGGAGCAGTTCATGACTTGGCAGAATCGC AAGAACCAGAAGCAGCTGACACTGAATAAGAAGATCGTGGAGCGCGACG CGGCCCTGGCCCGCGTCAGCCTCTACGAGCATCAGGTCAGTCAGGAGGGC CGCAAGCTGAATGATTTCAAGTACCTCCTCAACAAGAAGGCCGTGTCGCA GCATTCGGTGATGGAGCAGGAGAATAGCTATATCCAGGCGAAGAATGAGC ATGCGGTCTGGCTCGCGCAGGTCAGCCAGCTCGAGAAGGAGATCGAACTG GTGCGCGAGGAGCTGGCCCTCGAGACCAATATCTTCCGCTCGGAAATCATC GAGAAGCATCGCAAGAGCACGGACAACATCGTGCTGCTGGAGCATGAGTT GGAAAAGAACCGCCAGCGCAAAGCGTCGTCGTTCATCAAGGCGCCGGTCT CCGGCACCGTGCAGGAGCTGAACATCCATACAGAAGGGGGCGTCGTGACG ACGGCCGAGACGCTGATGATCATCGTCCCGGATAATGACATCCTCGAGGTC ACCGCCTCGGTGCTGAATAAGGACATCGGCTTCATCCAGCCGGGCCAGGA GGTGGTCATCAAGGTGGATGCGTACCCCTACACCCGCCATGGCTATCTGAC GGGCAAGGTCAAGAATATTACGGCCGACTCAGTGAGCGTGCCGGATACGG GCCTGGTCTTCAATGTCATCATCTCGGTCGACCGTAACGACATCCAGGGCG AGCGCAAGAAGATCCCCGTCACCGCCGGCATGACGGTCATGGCCGAGATC AAGACCGGCGTGAGGAGCGTTATCTCCTATCTGCTCAGCCCACTCAAGGAG ACGATCAACGAGTCGCTCCGCGAGCGCTAG
157
APPENDIX B
PROTOCOLS, REAGENTS, AND MISCELLANEOUS PCR
Add the following reagents to a tube (50 µl reaction) in the following volumes and order: • 29µl sterile H2O • 5 µl 10X buffer • 2µl dNTP Mix • 6µl MgCl2 • 6µl cells/DNA • 0.25µl Taq Polymerase • 1µl Primer 1 • 1µl Primer 2
These can be adjusted as necessary. Higher MgCl2 concentrations mean less specific binding, while lower concentrations mean more specific binding. DMSO may be added to lower the annealing temperatures of the primers (usually 0.5-2 µL). DNA is based on concentration of the sample being used. Adjust the amount of water added to bring the total to 50µL.
The thermocyler is setup beforehand with the desired protocol. Typically: 94°C for denaturing, 50-60°C for primer annealing, and 72°C for polymerase extending. Example setup: 1. 94°C 2 min 1x 2. 94°C 45 sec 3. 55°C 45 sec 4. 72°C 1min 15 sec 5. Repeat Step 2. thru Step 4. 35x 6. 72°C 5min 1x 7. 4°C indefinitely
Restriction Enzyme Digestion
1. Prepare and label 1.5 µl tubes for each reaction to be carried out 2. Add the following to each tube (keeping all materials on ice): a. 3 µl 10x FD Green buffer b. 1 µl FD restriction enzyme 1 c. 1 µl FD restriction enzyme 2 d. 4 µg DNA i. Calculate the volumes of DNA to add based on the stock concentration using the digest calculation spreadsheet on the lab computer 158
e. Bring the solution to 30 µl using sterile ddH2O (the volume of DNA and H2O will be 25 µl when added together) 3. Incubate the restriction digests at 37°C for 1 hour 4. Run the digest on a gel to isolate the desired pieces or to check fragment lengths. Electrophoresis
1. Gel Preparation: a. Weight out agarose i. For large gel, add 200 ml 1X TAE; ii. For small gel, add 50 ml 1X TAE . iii. For 1% gel, add 2 g agarose to large and 0.5 g agarose to small gel. iv. For 2% gel, add double the agarose. b. Heat in 20 second increments in microwave until boiling, stir carefully c. Let cool until comfortable to touch and add 1 μl of EtBR for every 10 μl of soln. d. After cooling period, pour into electrophoresis chamber and let solidify 2. Gel Loading: a. Add 10 μl of DNA ladder (GeneRuler™ DNA Ladder Mix, ready-to-use) b. Add 6X loading dye (6X = sample volume + X; solve for X) 3. Use Thermo Scientific GeneJET gel extraction kit to isolate DNA bands
TAE (50X) Buffer: 242 g Tris base to around 600 ml 57.1 ml glacial acetic acid (1 mole) 100 ml 0.5 M EDTA (pH 8.0) ddH2O to 1000 ml
Ligation Reaction
1. Prepare and label 1.5 ml tubes for each reaction to be carried out 2. Calculate the amount of insert and vector DNA to add using the ligation spreadsheet on the lab computer. 3. Add the following to each tube: 1. Insert DNA 2. Vector DNA 3. 5µl 10X ligation buffer 4. 1µl T4 DNA ligase 5. H2O to bring the solution to 50 µl 4. Allow reaction to occur at room temperature for 30 min to 1 hour. Alternatively, reactions may be placed on ice and allowed to carry out over a longer period of time (an afternoon, overnight) before use in transformation 5. Ligation reactions not used in transformations may be stored in the fridge for a day or two.
Transformation using XL1 Blue E. coli cells 159
1. Take competent cells out of the -80 °C freezer and thaw on ice. 2. Add 25 µL chilled 10% glycerol to each competent cell tube. 3. Add 5 µL of DNA to the cell solution. 4. Incubate on ice for 5-8 minutes. 5. Turn on the electroporation machine. It should be set to 2500 V, 200 O, and 25 µF for E. coli. 6. Carefully, add one of the cell/DNA/glycerol solutions into a cuvette. 7. Before electroporation, lightly tap the cuvette down on the counter to ensure that the solution is filling the bottom portion of the cuvette. 8. Dry the cuvettes off with a KimWipe. 9. Place the cuvette so that the metal sides are touching the metal sides of the electroporator. 10. Pulse the cells, and then quickly add 1 ml pre-warmed LB media with no antibiotic. 11. Remove the solution and transfer it to the original competent cell tube 12. Incubate the cell solutions at 37°C for 1-2 hours. 13. Plate the cells on plates containing the correct antibiotic. Add 500 µL-1000 µL of solution to one plate, spread with the spreading stick. 14. Let the plates dry in the bio-safety hood for no longer than 30 minutes with the cap off. 15. Invert the plates and place them in the 37°C incubator overnight
CTAB Plasmid DNA Extraction
1. Inoculate 5 ml of LB media with antibiotics contained in a 12 ml tube with one colony or with a flake from a freezer stock. Grow overnight. 2. Centrifuge the 12 ml tubes containing the 5 ml cultures in the large centrifuge at 3500 RPM for 10 min. Discard supernatant liquid. 3. Re-suspend cells in 200 µl of “STET for CTAB” buffer. Transfer to 1.5 ml tubes. 4. Add 10 µl Lysozyme (50 mg/ml) and incubate at room temperature for 5 min. 5. Boil for 45 seconds in blue plastic holder and then centrifuge for 20 min at 13,000 RPM in the bench top centrifuge (or until pellet gets tight). 6. Use a sterile toothpick or pipette tip to remove the pellet by dragging it (it should be somewhat slimy, but if pelleted well enough, it will hold together), if it doesn’t hold together, re-centrifuge and retry. 7. Add 5 µl RNase A (10 mg/ml) and incubate at 68°C for 10 minutes. 8. Add 10 µl of 5% CTAB and incubate at room temperature for 3 min. 9. Bench top centrifuge for 5 min at 13,000 RPM, discard supernatant, and re- suspend in 300 µl of 1.2 M NaCl by vortexing. 10. Add 750 µl of 95% ethanol to precipitate DNA. 11. Incubate for 30 mins in -20°C freezer (or 15 min in -70°C freezer) to help DNA precipitate. 12. Bench top centrifuge for 5 min at 13,000 RPM to compact DNA pellet. Make sure the hinge is away from the center of the rotor, this will make the pellet form on the bottom of the tube on the side of the hinge (it might be hard to see or invisible, so this way you know where it should be). 160
13. Discard supernatant, rinse pellet in 750 µL 80% ethanol, and let tubes dry upside down with caps open. 14. Re-suspend DNA in ddH2O (50 µl). Vortex or pipette up and down to ensure re- suspension of DNA. 15. Measure DNA concentrations using the NanoDrop 1000 instrument and record them on the tube holding the DNA and in the lab notebook.
BioBrick Assembly
Figure B.1. BioBrick assembly process. Samples are cut with restriction enzymes. Fragments are purified using gel electrophoresis and gel extraction. Isolated DNA is mixed and ligated together. This process is repeated for each added part.
References 1. http://parts.igem.org/Assembly:Standard_assembly 161
APPENDIX C
COPYRIGHT PERMISSION FORMS
The manuscript entitled “Food waste conversion to microbial polyhydroxyalkanoates” was published in the Journal of Microbial Biotechnology, with John Wiley and Sons as the publisher. According to the Copyright Clearance Center, “This article is available under the terms of the Creative Commons Attribution License (CC BY) (which may be updated from time to time) and permits use, distribution and reproduction in any medium, provided that the Contribution is properly cited.” 162 163 164 165
The article entitled “Polyhydroxybutyrate production using a wastewater microalgae based media,” presented in Appendix D, was published in the journal Algal Research, with Elsevier as the publisher. According to the Copyright Clearance Center, “as the author of this Elsevier article, you retain the right to include it in a thesis or dissertation, provided it is not published commercially. Permission is not required, but please ensure that you reference the journal as the original source.” 166 167
APPENDIX D
POLYHYDROXYBUTYRATE PRODUCTION USING A WASTEWATER
MICROALGAE BASED MEDIA
Asif Rahman, Ryan J. Putman, Kadriye Inan, Fulya Ay Sal, Ashik Sathish, Terence
Smith, Chad Nielsen, Ronald C. Sims, Charles D. Miller
Algal Research (2015), 8: 95-98
Abstract
Bioproduct production from wastewater microalgae has the potential to contribute to societal needs with value added chemicals. Microalgae can remediate wastewater to remove nitrogen, phosphorus, and heavy metals and can be processed to produce biofuels and bioproducts. It was previously demonstrated that recombinant Escherichia coli could produce polyhydroxybutyrates (PHBs) when cultured on a wastewater microalgae wet lipid extracted media. In this present study, microalgae were harvested from the effluent of a wastewater treatment facility via centrifugation and hydrolyzed to create a liquid medium for recombinant E. coli growth and PHB production. Standard E. coli growth media was supplemented with various concentrations of hydrolyzed algal extract to produce a maximum of 31% PHB of the E. coli dry cell weight.
Introduction
Microalgae have been well studied for production of biodiesel [1] and recently microalgae have been proposed to be the basis for a biorefinery model where multiple 168
chemicals can be produced simultaneously [2]. Producing several chemicals from the
same microalgae feedstock could potentially make the production of multiple commodity
chemicals from a biological resource economically viable. The limitations to microalgae
culturing are well-documented, including but not limited to: nutrient supply, water
scarcity, harvesting, and dewatering [3].
The City of Logan, UT has a 460 acre seven pond facultative lagoon system to treat weak domestic wastewater. Weak domestic wastewater contains approximately 20 mg/L nitrogen and 4 mg/L phosphorus, and is ideal for microalgae growth [4].
Facultative lagoon systems can be used to culture mixed consortia of microalgae to remediate the wastewater by removal of phosphorus and nitrogen. There are a wide range of methods previously employed to harvest microalgae from an open pond system that include: rotating algal biofilm reactor (RABR) [5,6], biological and chemical flocculants
[2,7,8], and centrifugation [9]. Harvested microalgae can then be processed and used as a feedstock for production of bioproducts [10,11]. It has been demonstrated that
Escherichia coli can be cultured on microalgae based substrates for production of biofuels and bioplastics [12,13].
E. coli can be easily cultured and has a fast doubling time making it an ideal candidate for production of recombinant bioproducts. Polyhydroxybutyrates (PHBs) are bioplastics that can be recombinantly produced in E. coli [14] and cyanobacteria [15].
PHB is a potentially useful polymer, in addition to being completely biodegradable, it has similar properties to traditional petrochemically derived plastics such as polypropylene and polystyrene [16]. Three genes are needed for the conversion of acetyl-CoA to PHB in 169
E. coli. The pBHR68 plasmid contains the lac promoter and three genes (phaA, phaB,
and phaC) needed for production of the short chain length (scl) polymer PHB [17].
Bacterial PHB production is not widespread in part due to the cost of the carbon
substrate. It has been estimated that the carbon substrate in a large scale manufacturing
context would constitute approximately 37% of the total production cost [18]. Due to the
high cost of carbon, an alternative low cost substitute is needed to culture E. coli in order
to make PHB production economically viable. In a previous study, it was demonstrated
that E. coli harboring the pBHR68 plasmid was able to successfully grow on a
Scenedesmus obliquus microalgae based media [2]. In a different study, various harvesting methods were used to collect microalgae grown in photobioreactors [19] and
then the harvested microalgae was processed via the wet lipid extraction procedure
(WLEP) to generate a variety of side streams and bioproducts [2,20]. One of the side
streams, termed ‘aqueous phase’ was used to culture E. coli and it was established that
the upstream harvesting method of S. obliquus affected the growth of the E. coli in the
aqueous phase media. The most successful microalgae harvesting method for high levels
of E. coli growth after 48 h (1012–1013 CFU/mL) was observed when the S. obliquus was centrifuged [2]. The same experiment was extended to harvesting wastewater mixed culture microalgae from the City of Logan, UT treatment plant pond E. Different harvested wastewater microalgae samples (approximately 11% lipids as fatty acid methyl esters, FAME [20]) were subjected to the WLEP to generate the aqueous phase. Results showed that harvesting via centrifugation gave the best E. coli growth (~1013 CFU/mL) and PHB production (7.8% PHB dry cell weight) [13]. 170
Centrifugation was selected as the preferred harvesting method because centrifuged microalgae processed via the WLEP demonstrated the highest levels of E. coli growth and PHB production. Additionally, the previous studies [2,13] used an unmodified aqueous phase media to culture E. coli, and a subsequent study used a fraction of the aqueous phase with standard E. coli media and obtained promising PHB yields [21]. The main objectives of the study reported here were to demonstrate E. coli growth and PHB production using hydrolyzed microalgae from wastewater effluent of the City of Logan, UT facultative pond treatment facility.
Materials and Methods
All chemicals and reagents were purchased from Thermo Fisher Scientific
(Pittsburgh, PA) unless stated otherwise.
Microalgae harvesting and processing
Wastewater microalgae were harvested from the City of Logan, UT wastewater treatment facility from the effluent stream leaving the facultative lagoon system.
Microalgae were centrifuged using a continuous centrifuge (Alfa Laval Clara 80, Lund,
Sweden) and dried in a temperature controlled oven. After drying, microalgae were stored at −20 °C.
A modified microalgae hydrolysis method was used similar to Ellis et al., where hydrolyzed microalgae was used to culture Clostridia to produce Acetone, Butanol, and
Ethanol [22]. Briefly, 10 g of dry microalgae was dissolved into 0.5 M (final concentration) sulfuric acid (H2SO4) with a total volume of 100 mL. The solution was 171 placed on a stir plate and heated to 90 °C for 30 min with constant stirring. After cooling to room temperature, the solution was neutralized to pH 7 with sodium hydroxide
(NaOH). This neutralized solution was then centrifuged at 3500 rpm for 30 min to clarify the solution. The supernatant was then used for culturing E. coli.
Bacterial Growth
Supernatant from hydrolyzed microalgae was used as the sole carbon source and was added to standard E. coli M9 growth media [23] in 1%, 2%, and 3% ratios (weight dry hydrolyzed microalgae to culture volume ratio, where 1% is 0.5 g dry algae in
50mL). An additional study was conducted that consisted of culturing E. coli in a 10% hydrolyzed microalgae solution (w/v). The hydrolyzed microalgae supernatant was not autoclaved in order to demonstrate that E. coli growth and PHB production could occur from a non-traditional carbon source. In addition to the liquid algal extract, growth media also contained M9 salts (Becton, Dickinson and Co, Sparks, MD), 0.002 M MgSO4, and
50 μg/mL ampicillin (IBI Scientific, Peosta, IA) [24,25].
The E. coli strain, XL1 Blue (Agilent Technologies, Santa Clara, CA) harboring the pBHR68 plasmid [17] was grown in LB media [23] overnight (~15 h). Cultures were then used to start larger 50 mL cultures with an initial optical density (OD600) of 0.05.
Isopropyl β-D-1thiogalactopyranoside (0.1 mM) (Gold Biotechnology, Inc. St. Louis,
MO) was added at 0 h to induce expression of the phaCAB genes. Bacterial growth was measured using optical density (OD600 nm) at 0, 4, 8, 12, 24, and 48 h. 172
Sugar Analysis
Total sugar was determined using a modified phenol–sulfuric acid method [22].
Briefly, 3 μL of 85% (w/v) phenol solution and 150 μL of 12 M sulfuric acid were added to the samples and the mixture was heated for 5 min at 90 °C. After cooling to room temperature for 5 min in an ice bath, absorbance (A490 nm) was measured using a
Synergy 2 microtiter plate reader (BioTek, Winooski, VT). Sugar concentrations were calculated based on a glucose standard.
Polyhydroxybutyrate analysis
PHB analysis was conducted on samples after 48 h of bacterial culturing.
PHB concentration was determined from a 1H NMR/GC correlation as described previously [26]. Briefly, approximately 15mg of lyophilized sample were dissolved in 1 mL deuterated chloroform (CDCl3 with 0.03% TMS (v/v), Cambridge Isotope
Laboratories, Inc. Andover, MA) and 5% sodium hypochlorite solution. Samples were vortexed, incubated, and centrifuged to promote phase separation. PHB phase was run on a Jeol ECX-300 NMR (Jeol USA, Inc. Peabody, MA) and a standard NMR/GC correlation was used for PHB quantification.
Statistical analysis
Data was processed with Statistical Analysis Software (SAS 9.4, SAS Institute
Inc., Cary, NC). One-way analysis of variance (ANOVA) was conducted on the data collected at 48 h for % PHB for the three different media types. REGWQ post hoc 173
comparison was performed on significant results with a confidence level of 95%. All %
PHB experiments were conducted in triplicate.
Results and Discussion
The steps conducted in this study to remove microalgae from wastewater effluent
of the City of Logan, UT facultative pond treatment facility, to hydrolyze the dried
microalgae, and to grow E. coli to produce the PHB polymer are depicted in Fig. D.1.
E. coli harboring the pBHR68 plasmid was grown on 1%, 2%, 3%, and 10% microalgae-
M9 media (w/v). The maximum optical density (OD600) for E. coli grown in 1% media was approximately 1.3, where stationary phase was reached at approximately 12 h post- induction (Fig. 2). The 2%, 3%, and 10% samples reached stationary phase at 24 h and achieved a maximum OD600 of 2.5, 3.61, and 7.6 respectively. E. coli growth in microalgae-M9 media was typical of that observed in traditional glucose-M9 media in other studies [25]. Cultures were allowed to continue growing until 48 h to allow time for
PHB accumulation.
174
Figure D.1. Schematic for the production of polyhydroxybutyrates in Escherichia coli from wastewater microalgae.
Results of the total sugar analysis indicated that simple sugars were present in the microalgae extract and were consumed during the course of bacterial growth. The 1% sample had 152 mg/L total sugar at 0 h and after 48 h of bacterial growth had 132 mg/L.
The 2% sample had 320 mg/L sugar at 0 h and after 48 h had 267 mg/L. The 3% sample had 445 mg/L sugar at time 0 and 287mg/mL after 48 h. The 10% sample had 1890 mg/L total sugar and finished with 1344 mg/L sugar. The amount of sugar consumed by the bacteria in each experiment (1%, 2%, 3%, and 10%) was comparable to the growth observed in Fig. D.2. 175
Figure D.2. Growth of E. coli harboring the pBHR68 plasmid on 1%, 2%, 3%, and 10% microalgae media.
Comparing PHB yields 48 h post-induction demonstrated that the M9 media containing 1% and 2% microalgae extract had the most PHB (as a percentage of E. coli dry cell weight). The 1% and 2% samples produced 31± 8.9% and 28.2 ± 2.1% PHB, respectively (Fig. D.3). In comparison, E. coli cultured in 3% and 10% microalgae extract media that had an average PHB accumulation of 11.2 ± 2.6% and 4.6 ± 0.7%, respectively. The statistical analysis yielded a p-value of 0.0026 indicating significant results (confidence of 95%, p < 0.05). Post hoc comparison using REGWQ showed that the PHB production levels at 1% and 2% were not significantly different from each other
(indicated with ‘a’ in Fig. D.3). 176
Figure D.3. Polyhydroxybutyrate production from different microalgae based cultures (where % PHB is a proportion of E. coli dry cell weight). Data sets with same letter (a or b) demonstrated no statistically significant difference (p N 0.05), error bars represent standard deviation (n= 3).
Additionally, the statistical analysis also demonstrated that PHB accumulation in
3% and 10% samples were not statistically different from each other (indicated with ‘b’ in Fig. D.3). The PHB production from 1% and 2% media samples was both significantly different from the PHB yields observed from E. coli grown in 3% and 10% algal extract.
These results demonstrate that there is a drop in PHB production from the 2% to the 3% microalgae media. The drop in PHB production at the higher percent microalgae media could be attributed to the increase in salts present in the media. In this study microalgae media was neutralized from a low pH with sodium hydroxide, thereby generating sodium sulfate. In a previous study, wastewater microalgae harvested with Aluminum Sulfate
(Alum) and used to culture E. coli yielded no PHB [13].
As described, E. coli growth in the 3% and 10% samples reached stationary phase later than then 1% and 2% samples, suggesting that E. coli in the 3% and 10% samples 177
could also have had less time to accumulate PHB. It has been suggested previously that acetyl-CoA is required for cell synthesis in log phase but is diverted to PHB production in stationary phase [27]. Future work could be conducted to best determine if extending the culturing time of the 3% and 10% experiments improves yields of PHB and whether or not this is economically viable.
The percentage of PHB accumulated in bacterial cells cultured in the 1% and 2% microalgae-M9 media was slightly lower than that seen in E. coli harboring pBHR68 grown inM9-glucosemedia (Table 1). In a previous study, it was found that E. coli harboring the pBHR68 plasmid grown in M9 media supplemented with 1.5% glucose could accumulate up to 47.24 ± 6.0%, 48 h post-induction [24]. Achieving approximately
31 ± 8.9% demonstrates the potential of using microalgae as the sole carbon source in media for E. coli culturing and bioproduct production.
Addition of an external carbon substrate such as glucose to the microalgae based media could potentially increase the growth and yields of PHB in E. coli. It was demonstrated in a previous study that 1% glucose addition to a microalgae based media to culture Clostridia tripled the yield of solvent production [22].
Comparing the 10% microalgae extract media (with noM9 addition) to that of a similar study [13], it was found that this 10% media did not perform as well as the media used in the previous study. In the previous study, approximately 9.6 g of microalgae (dry weight equivalent) was extracted (via centrifugation) from the City of Logan, UT wastewater treatment facility pond E and subjected to the wet lipid extraction procedure
(WLEP) to produce approximately 7.8% PHB [13]. In the present study, microalgae extracted from the effluent of the wastewater treatment plant generated 4.6% PHB. The 178
lower PHB yield could be attributed to the fact that in this study the microalgae harvested could have already been lysed, resulting in a lower yield of sugars extracted. There could have also been some inhibitory effect from the media that was more concentrated, thus reducing the production capacity of the E. coli.
Table D.1 shows the yields of PHB obtained from the different experiments carried out in this study compared to that of another study with the same strain of E. coli
[24]. In addition to PHB% as a fraction of dry cell weight, the total carbon substrate needed to produce 1 kg PHB was also estimated. It was determined that using the 1% microalgae inM9 media would need approximately 4.3 kg of dry microalgae to produce 1 kg of PHB. This is comparable to a standard 1.5% glucose M9 media in the previous study that was predicted to need 2.5 kg of glucose to make 1 kg of PHB. This estimate assumed a linear scaling from shaker flask volume of 50 mL to a large scale bioreactor, however, in order to get a more accurate measurement additional parameters would need to be considered.
Table D.1. Production and yields of PHB after 48 h from Escherichia coli XL1 Blue harboring the pBHR68 plasmid grown in M9 media supplemented with wastewater microalgae hydrolyzed fraction. Standard deviations are based on triplicate (n=3). Carbon source PHB % g PHB / L g PHB / Carbon needed Reference in M9 media g carbon (kg) to produce substrate 1 kg PHB 1% microalgae 30.97 ± 8.9 2.30 ± 1 0.232 4.3 This study media 2% microalgae 28.19 ± 5.1 2.09 ± 0.5 0.104 9.5 This study media 3% microalgae 11.24 ± 2.6 0.77 ± 0.5 0.026 38.7 This study media 10% 4.60 ± 0.7 0.32 ± 0.1 0.003 305.2 This study microalgae media 1.5% glucose 47.24 ± 6.0 5.43 ± 1.7 0.4 2.5 [24]
179
Data sets with same letter (a or b) demonstrated no statistical significant difference (p > 0.05).
Conclusions
This study built upon work previous work that demonstrated the production of
PHB from a microalgae feedstock. From this study, growth of recombinant E. coli
harboring the pBHR68 plasmid (containing the phaCAB operon) on wastewater
microalgae based media was observed for all samples. It was found that the maximum
PHB accumulation in E. coli was approximately 31 ± 8.9% seen on the 1% microalgae-
M9 media 48 h post-induction. Future work could include determining the effects of production of additional bioproducts using recombinant E. coli grown on wastewater microalgae media and the addition of a traditional carbon source such as glucose or a carbon-rich waste to the microalgae based media. Additionally, a technoeconomic analysis could be conducted to determine the most cost effect means of production of various recombinant products in E. coli from a wastewater microalgae feedstock.
Acknowledgments
The authors would like to acknowledge the following groups: Environmental
Department of the City of Logan, UT, Utah Science Technology and Research (USTAR),
Sustainable Waste to Bioproducts Engineering Center (SWBEC), Synthetic
Biomanufacturing Institute (SBI), the Huntsman Environmental Research Center
(HERC), the Scientific and Technological Research Council of Turkey (TÜBITAK), and
the Turkey Council of Higher Education (YÖK). The authors would also like to thank
Reese Thompson, Neal Hengge, and Brian Smith for their contributions.
180
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