in-situ observation and quantification of microalgae ......the direct observation of these complex...
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In-situ observation and quantification of microalgae
downstream processing on a microfluidic platform
by
Xiang Cheng
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Mechanical & Industrial Engineering
University of Toronto
© Copyright by Xiang Cheng 2018
ii
In-situ observation and quantification of microalgae downstream
processing on a microfluidic platform
Xiang Cheng
Doctor of Philosophy
Mechanical & Industrial Engineering
University of Toronto
2018
Abstract
Producing biofuels and bioproducts from microalgae is a promising path for low-carbon energy
and products. Microalgal biomass is an attractive feedstock for the generation of carbon neutral
biofuels and high-value bioproducts because of the high growth rate and lipid content of many
microalgae species. Understanding the downstream processing of converting microalgal biomass
to valuable products is a critical step in the biofuel industry. In this thesis, a novel microfluidic
platform capable of precise control of processing parameters and providing optical access to
reactions at high temperature and pressure was developed and applied to observe and quantify the
biomass-to-bioproducts conversions in three distinct studies.
First, for bioenergy application, hydrothermal liquefaction of microalgae was performed on this
microfluidic platform monitored using fluorescence microscopy. A strong shift in the fluorescence
signature from the algal slurry at 675 nm (chlorophyll peak) to a post-HTL stream at 510 nm is
observed for reaction temperatures at 260°C, 280°C, 300°C and 320°C (P = 12 MPa), and occurs
over a timescale on the order of 10 min. Biocrude formation and separation from the aqueous phase
into immiscible droplets is directly observed and occurs over the same timescale.
Second, many algal bioproduct efforts currently focus on high-value products such as astaxanthin
due to the much-improved economics over producing fuels. Hydrothermal disruption of the cell
wall for astaxanthin extraction from wet biomass using high temperature and pressure was
demonstrated and studied using this microfluidic platform. Hydrothermal disruption at a
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temperature of 200 °C was shown to be highly effective, resulting in near-complete astaxanthin
extraction from wet biomass - a significant improvement over traditional methods.
Third, supercritical CO2 has relatively low critical temperature and pressure (31.1 °C and 7.4 MPa)
is considered a greener solvent for bioactive compounds extraction. Supercritical CO2 extractions
of astaxanthin with and without co-solvents (ethanol and olive oil) were performed on the
microfluidic platform to study the extraction mechanism in each case. Astaxanthin extraction using
ScCO2 achieved 92% recovery at 55 °C and 8 MPa applied over 15 hours. With the addition of co-
solvents, ethanol and olive oil, the timescales of extraction process are reduced significantly from
15 hours to a few minutes, representing the fastest complete astaxanthin extraction at such low
pressures.
The direct observation of these complex reaction processes was made possible for the first time
here, allowing visual characterization, fluorescence spectroscopy, and quantitative imaging of the
conversion at the single-cell scale during all stages. This level of insight has simply not been
possible with previous conventional reactors. Although batch reactors have advantages in, for
instance, quantifying yields requiring large volumes of products, microfluidic reactors have
advantages with respect to process control and visualization at cellular level - providing high
resolution, real-time data on complex reactions. The innovative platform and results presented in
this thesis provide new insight in the challenging area of biomass-to-bioproduct conversion, and
provide insight that can inform larger scale operations.
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Acknowledgments
I would like to extend my gratitude and thanks to individuals who have supported me both
professionally and personally throughout my doctoral journey.
First, I would like to express my thanks to my thesis supervisor, Prof. David Sinton, for his
guidance, support and encouragement during the Ph.D. program. I am grateful for giving me the
opportunity to join his research group and encouraging me to strike out in a new direction of my
choice. I am also grateful for all I have learned from and inspired by him, not only the knowledge,
but also the professionalism, management and leadership that he shares on a daily basis.
I would also like to thank my examination committee: Prof. Grant Allen, Prof. Murray Thomson,
Prof. Shulin Chen and Prof. Axel Guenther for their time and effort in evaluating my work and
providing valuable and insightful comments. Also, thank you to those who wrote letters of
recommendation – Behraad Bahreyni, Farid Golnaraghi, Ahmad Rad, Scott Morgan and David
Sinton, your support and confidence have impacted me in so many ways. Moreover, I would like
to thank my lab mates that we shared amazing experience together over the last four years.
Although not exhaustive, I would like to thank Matthew Ooms, Bo Bao, Percival Graham, Brian
Nguyen, Jason Riordon, Seven Qi, Reza Nosrati, Pushan Lele and Tom Burdyny for your help,
encouragement, and inspiration.
Finally, I would like to thank my family for their love and support that was worth much more than
I can express on paper. A big thanks to my parents, Feng Cong and Qingzhong Cheng, for their
unconditional love and support along the way. Special thank you to my aunt Lam Cong who
encouraged me to study abroad and supported me all the way through. My deepest gratitude and
appreciation goes to my beautiful and lovely wife, Xia Zhang, for supporting me throughout my
career and my life in general.
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Table of Contents
Acknowledgments.......................................................................................................................... iv
Table of Contents .............................................................................................................................v
List of Figures .............................................................................................................................. viii
List of Tables ............................................................................................................................... xiv
List of Appendices .........................................................................................................................xv
Chapter 1. Forward ....................................................................................................................1
1.1 Motivation ............................................................................................................................1
1.2 Thesis Overview ..................................................................................................................4
Chapter 2. Introduction ..............................................................................................................7
2.1 Microalgae for biofuel production .......................................................................................7
2.1.1 Assessments for downstream conversions ...............................................................8
2.1.2 Water under subcritical conditions ........................................................................13
2.1.3 Characterization of microalgae and post-HTL products ........................................16
2.1.4 HTL of biomass and effects of processing conditions ...........................................19
2.2 Microalgae for high-value bioproducts ..............................................................................22
2.2.1 Cell structure of Haematococcus pluvialis ............................................................23
2.2.2 Cell wall disruption and extraction techniques ......................................................25
2.3 High temperature and pressure microfluidics ....................................................................26
Chapter 3. Current Downstream Processing and Emerging Technologies for Microalgae .....29
3.1 Optimizing downstream processing ...................................................................................29
3.2 Harvesting and dewatering processes ................................................................................30
3.2.1 Biomass thickening ................................................................................................30
3.2.2 Dehydration to dry biomass ...................................................................................31
3.3 Processing for dry microalgae ...........................................................................................32
3.3.1 Lipids extraction ....................................................................................................32
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3.3.2 Extracted lipids and other components ..................................................................33
3.3.3 Processing of whole dry microalgae ......................................................................34
3.4 Processing for wet microalgae biomass .............................................................................35
3.4.1 Hydrothermal liquefaction .....................................................................................36
3.4.2 Supercritical water gasification ..............................................................................39
3.5 Processing for microalgae in culture..................................................................................40
3.5.1 Fermentation ..........................................................................................................40
3.5.2 Anaerobic digestion ...............................................................................................41
3.5.3 Direct secretion ......................................................................................................42
3.5.4 Microalgae-microbial fuel cells .............................................................................42
Chapter 4. Hydrothermal Liquefaction of Microalgae on a Chip for Biocrude Production ....44
4.1 Introduction ........................................................................................................................45
4.2 Experimental ......................................................................................................................47
4.3 Results ................................................................................................................................50
4.4 Conclusion .........................................................................................................................54
4.5 Supplementary Information ...............................................................................................55
4.5.1 Fabrication of microfluidic chip ............................................................................55
4.5.2 Experimental apparatus ..........................................................................................56
4.5.3 High temperature and pressure packaging .............................................................57
Chapter 5. Hydrothermal Disruption of Algae Cells for Astaxanthin Extraction ...................59
5.1 Introduction ........................................................................................................................59
5.2 Experimental setup.............................................................................................................62
5.2.1 Device design and fabrication ................................................................................62
5.2.2 Cell culture and trapping ........................................................................................62
5.2.3 Cell wall disruption and extraction ........................................................................62
5.2.4 Quantifying astaxanthin content ............................................................................63
vii
5.2.5 HPLC analysis of extracted astaxanthin ................................................................64
5.3 Results and discussion .......................................................................................................64
5.4 Conclusion .........................................................................................................................71
5.5 Supplemental material .......................................................................................................71
Chapter 6. Astaxanthin Extraction from Algae using Supercritical CO2 with Co-solvent ......75
6.1 Introduction ........................................................................................................................75
6.2 Experimental Section .........................................................................................................78
6.3 Results and discussion .......................................................................................................79
6.4 Conclusions ........................................................................................................................86
6.5 Supplementary Information ...............................................................................................86
Chapter 7. Conclusions ............................................................................................................89
7.1 Summary ............................................................................................................................89
7.2 Future Outlook ...................................................................................................................90
References ......................................................................................................................................92
Appendices ...................................................................................................................................110
A1. The full thermodynamic phase envelope of a mixture in 1000 microfluidic chambers ..110
viii
List of Figures
Figure 1-1. Microalgae based biorefinery integrated with related industries to produce numerous
sustainable deliverables. Reproduced from ref9, © 2010, with permission from Elsevier. ............ 2
Figure 1-2. Sunlight-to-biomass conversion efficiency and losses along the path, as well as value
and market size of related microalgae products. Reproduced from ref15, © 2016, under CC-BY
license. ............................................................................................................................................ 4
Figure 2-1. A summary of life cycle assessments for biofuel production form microalgae. a) Net
energy ratio for microalgae biomass production and b) Illustrative estimates for carbon dioxide
emissions from algal biomass production. Reproduced from ref34, © 2013, with permission from
Elsevier. ........................................................................................................................................ 10
Figure 2-2. a) Energy balance and b) Global Warming Potential for biofuel production using wet
and dry lipid extraction. Reproduced from ref8, © 2013, with permission from American Chemical
Society........................................................................................................................................... 11
Figure 2-3. Techno-economic analysis of liquid fuel produced from hydrothermal liquefaction of
woody biomass a) Effect of improvement on MFSP from state-of-technology (SOT) case to goal
case. b) Sensitivity analysis of parameter variation on the MFSP of the goal case. a) and b) are
reproduced from ref37, © 2014, with permission from Elsevier. c) Sensitivity analysis of the MFSP
of biofuel produced from defatted microalgae via HTL. Reproduced from ref38, © 2014, with
permission from Elsevier. ............................................................................................................. 12
Figure 2-4. Density, static dielectric constant and ion dissociation constant (Kw) of water at 30
MPa as a function of temperature. Reproduced from ref46, © 2008, with permission from Royal
Society of Chemistry..................................................................................................................... 15
Figure 2-5. Characteristics for algae used for HTL. The green, black and red dashed arrows indicate
the mass fraction of lipids, proteins and carbohydrates, respectively. Reproduced from ref10, ©
2014, with permission from Elsevier. ........................................................................................... 17
Figure 2-6. HTL of microalgae procedure and product separation process. Reproduced from ref54,
© 2012, with permission from Elsevier. ....................................................................................... 18
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Figure 2-7. Yields of products from hydrothermal liquefaction of different types of feedstock in:
a) water, b) sodium carbonate, c) formic acid. Reproduced from ref12, © 2011, with permission
from Elsevier. ................................................................................................................................ 19
Figure 2-8. Biocrude yield of HTL as a function of heating rate. a) Temperature profiles with
corresponding biocrude yields from HTL of Nannochloropsis sp. for different set-point
temperatures of 300, 500, 400, and 600 °C. Reproduced from ref65, © 2013, with permission from
American Chemical Society. b) The effect of heating rate on the products yield from HTL of
macroalgae. (T = 350 °C; holding time = 15 min; and biomass/water ratio = 1/10, w/w).
Reproduced from ref66, © 2014, with permission from Elsevier. ................................................. 21
Figure 2-9. Market value of the microalgal components and total selling price of the biomass for
different market scenarios. Reproduced from ref69, © 2016, under CC-BY license. ..................... 23
Figure 2-10. a) Microscopic images of Haematococcus pluvialis in life cycle. (A) Green vegetative
motile cell; (B) Green vegetative palmella cell; (C) Astaxanthin accumulating palmella cell in
transition to aplanospore; (D) Astaxanthin accumulated aplanospore cell. Scale bar: 10 μm.
Reproduced from ref72, © 2016, under CC-BY license. b) Illustration of life cycle of H. pluvialis.
Reproduced from ref73, © 2013, under CC-BY license. ............................................................... 24
Figure 2-11. Summary of five developmental states of the cell wall during aplanospore morphogenesis
in Haematococcus pluvialis: I, 1-week-old flagellates ; II, flagellates at least 2 weeks old just rounding
off; III, 2- to 3-week-old aplanospores; IV, at least 3-week-old aplanospores; V, aplanospores in their
final state. CYP, cytoplasm; IS, interspace; PL, plasmalemma; PW, primary wall; SV, secretory
vesicles; SW, secondary wall; TCL, tripartite crystalline layer; TLS, trilaminar sheath; W1±W7, layers
of the extracellular matrix. Reproduced from ref74, © 2002, with permission from Taylor & Francis.
....................................................................................................................................................... 25
Figure 2-12. Examples of (a) metal, (b) glass and (c) silicon/glass microreactors. Reproduced from
ref20, © 2011, with permission from Elsevier. .............................................................................. 27
Figure 3-1. Summary of current downstream processing techniques for wet microalgae biomass (a-f)
and cells in culture (g-j). (a-b) Hydrothermal liquefaction156,157. Image (b) has been reproduced with
permission from Elsevier157, Copyright 2013. (c) Supercritical water gasification159. Reproduced with
permission from Elsevier, Copyright 2016. (d-e) Ionic liquid treatment for wet extraction194,195. Image
x
(d) has been reproduced with permission from Royal Society of Chemistry194, Copyright 2014. Image
(e) has been reproduced with permission from Royal Society of Chemistry196, Copyright 2015. (f)
Astaxanthin extraction for hydrothermal disrupted cells197. (g) Fermentation of pretreated wet
biomass161. Image reproduced under CC-BY license, Copyright 2014. (h-i) Anaerobic digestion of algal
biomass169,170. Image (h) has been reproduced with permission from Elsevier170, Copyright 2017. Image
(i) has been reproduced with permission from Elsevier169, Copyright 2016. (j) Microalgae-microbial
fuel cells198. Reproduced with permission from Elsevier, Copyright 2015. This figure is reproduced
from ref91 with permission from The Royal Society of Chemistry. ................................................... 38
Figure 4-1. a) Schematic of hydrothermal liquefaction of microalgae in the microfluidic chip with
in-situ observation of biocrude production using fluorescence microscopy. b) Distinct fluorescence
signatures of algae slurry at the inlet and biocrude at the outlet. Reproduced by permission of The
Royal Society of Chemistry. ......................................................................................................... 46
Figure 4-2. Schematic representation of the assembly of the water-cooled manifold, temperature
controlled heating chuck and the microfluidic chip using a separation glass and double seal O-ring
to prevent cracking from hard contact. Reproduced by permission of The Royal Society of
Chemistry. ..................................................................................................................................... 48
Figure 4-3. a) Normalized fluorescence intensity of algae slurry observed at viewing points along
the channel under 320°C indicating the formation of biocrude over time. b) The progression of
normalized fluorescence intensity of the 510nm peak at the reaction temperature of 260°C, 280°C,
300°C and 320°C indicating higher reaction temperature has higher reaction rate. Solid lines
included as a guide for the eye. Reproduced by permission of The Royal Society of Chemistry.51
Figure 4-4. a) Fluorescence images obtained at viewing points along the channel with increase in
reaction time indicating the progression of biocrude formation. b) Microscopic observation of
fluids at the inlet and outlet via both fluorescence and dark-field imaging. Scale bars: 50 μm.
Reproduced by permission of The Royal Society of Chemistry................................................... 53
Figure 4-5. a) Schematic illustration of achieving high temperature and pressure on a chip by
separating high pressure compression from high temperature area. b) Si/glass chip fabrication
process (from top to bottom, left to right)..................................................................................... 56
xi
Figure 4-6. Schematic diagram of experimental setup with flow direction indicated by arrows
along the processing path. The flow path of the switching valve at two positions is shown in green
lines. .............................................................................................................................................. 57
Figure 4-7. Detailed drawing of the compression sealing of the microfluidic chip (MF) with algae
slurry fluid indicated in green. O-rings and spacers are used to ensure a quality seal between the
manifold and the chip and to prevent overtightening and damage. .............................................. 58
Figure 5-1. Simplified schematic of on-chip astaxanthin extraction from H. pluvialis. Enlarged
schematics of the cell capture area show initial cell trapping, cell wall disruption and astaxanthin
extraction. Reproduced by permission of The Royal Society of Chemistry. ................................ 61
Figure 5-2. a) Dark field images of mature red cysts of H. pluvialis at both the initial stage and
after solvent-based extraction, for each of five tested cases. All images have the same scale bar,
and were obtained with identical settings using darkfield microscopy. b) Normalized extracted red
content for each case. Reproduced by permission of The Royal Society of Chemistry. .............. 66
Figure 5-3. Time-course images of red cysts treated with hydrothermal processes of 150 oC for 30
min, 200 oC for 30 min, 200 oC for 10 min and 200 oC for 5 min, respectively. The scale bar is
identical for all images. Reproduced by permission of The Royal Society of Chemistry. ........... 67
Figure 5-4. Normalized red content during acetone extraction for six cells treated by hydrothermal
processing at 200 °C for 10 min. Inset images shows corresponding images of one representative
cell at four times during the procedure. Reproduced by permission of The Royal Society of
Chemistry. ..................................................................................................................................... 68
Figure 5-5. HPLC analysis of cell extract products for (a) mechanical extraction using a mortar
and pestle and (b) hydrothermal extraction for 20 min at 200 °C treatment. Reproduced by
permission of The Royal Society of Chemistry. ........................................................................... 69
Figure 5-6. Schematic diagram of the experimental setup with flow direction indicated by arrows
along the processing path. The flow path of the switching valve at two positions is shown using
green lines. .................................................................................................................................... 71
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Figure 5-7. Irradiation spectrum of light used to induce astaxanthin accumulation in
Haematococcus pluvialis. ............................................................................................................. 72
Figure 5-8. Microscope images of red cysts, indicating coloration change during the heating up
phase in the hydrothermal disruption process. The scale bar applies to all images...................... 72
Figure 5-9. Dark field, bright field and fluorescence images of initial cells and after acetone
extraction. Fluorescence 1 and 2 images were taken using FITC (excitation filter: 475/50 nm;
emission filter: 540/50 nm) and TxRed (excitation filter: 559/34 nm; emission filter: 630/69 nm)
filter cubes respectively. The scale bar of 50 µm applies to all images. ...................................... 73
Figure 5-10. Cell wall deformation of red cysts under hydrothermal processes at 200 °C in 10 min
with and without flow. Green dashed lines indicate cell wall boundaries. The scale bar of 50 µm
applies to all images. ..................................................................................................................... 74
Figure 6-1. Single-cellular visualization and quantification of astaxanthin extraction in
Haematococcus pluvialis using supercritical CO2 and co-solvents. Post ScCO2 and ethanol
extraction using acetone provides an overall extraction efficiency metric. Images were taken from
pure ScCO2 extraction experiment at 70 °C. ................................................................................ 80
Figure 6-2. a) Darkfield images H.p cells before and after ScCO2 extraction at 40 °C and 70 °C. b)
The progression of normalized red content for ScCO2 extraction at 40 °C, 55 °C and 70 °C
indicating higher extraction temperatures resulted in higher extraction rates. Solid lines represent
1st order trendline fits to the experimental data with equation given by the side. c) The normalized
red content for ScCO2 extraction process at 55 °C for a complete extraction process over 900
minutes with dark-field snapshots along the process. ................................................................... 82
Figure 6-3. The progression of normalized red content for ScCO2 extraction with ethanol over
1000 seconds at 40 °C, 55 °C and 70 °C. Zoom-in plot for the first 30 s extraction time indicates
a rapid extraction of astaxanthin from ScCO2 with ethanol at 55 °C and 70 °C. ......................... 83
Figure 6-4. a) The comparison of normalized red content for ScCO2 extraction with ethanol and
olive oil at 55 °C over 200 seconds. Time-lapsed snapshots of the extraction process with olive oil
are provided. b) Snapshot of ScCO2 extraction with olive oil indicating three phases: ScCO2, olive
xiii
oil and water. The boundary of olive oil and CO2 phases are illustrated using yellow and green
dash lines. ...................................................................................................................................... 85
Figure 6-5. Schematic diagram of the experimental setup with the flow direction indicated by
arrows along the processing path. The flow of the switching valve at two positions is shown using
red lines. ........................................................................................................................................ 87
Figure 6-6. Illustration of typical colors of H. pluvialis cells with different concentration of
astaxanthin and the calculated RGB-based astaxanthin content. .................................................. 88
xiv
List of Tables
Table 3-1 Summary of extraction and conversion techniques for different concentrations of
microalgae biomass. ...................................................................................................................... 36
Table 4-1: Elemental composition and Higher Heating Value of dry algae and biocrude from 1
min, 5 min and 10 min reaction times........................................................................................... 52
Table 4-2: List of components in the apparatus and their purpose. .............................................. 57
Table 6-1: Summary of studies on ScCO2 extraction of astaxanthin from H.pluvialis ................ 77
Table 6-2: Summary of supercritical carbon dioxide extraction experiments and results. ........... 80
xv
List of Appendices
Appendix 1: The full thermodynamic phase envelope of a mixture in 1000 microfluidic
chambers
1
Chapter 1.
Forward
Microalgae have great potential to help address critical global challenges on mitigating climate
change, producing sufficient fuels, food and chemicals in a sustainable manner. Understanding the
downstream conversions of microalgae biomass to biofuels and bioproducts is a crucial step –
particularly because these downstream conversion processes can be very energy intensive, eroding
both the environmental and economic motivations for microalgae. In this thesis, an innovative
microfluidic platform will be introduced with three industry-relevant applications demonstrated in
Chapter 4 to 6.
1.1 Motivation
In 2015, 195 nations have set a binding agreement, known as COP211 to keep global temperature
rise below 2 °C to prevent severe climate effects. In the same year, the planet temperature was
0.9 °C above the 20th-century average – nearly halfway to this globally agreed-on 2°C threshold.
Global temperature increase is believed to cause serious consequences, including sea level rise,
forced displacement, extreme weathers, reduced crop productivity and pandemics.2 In respect of
climate change mitigation, reducing greenhouse gas (GHG) emissions from fossil fuels which
contribute to 85.5% of global energy in 2016 (33.3% Oil, 24.1% Natural gas and 28.1 % Coal)
present the greatest potential. With the increasing energy demand from the expanding population,
along with the goal of mitigating GHG emissions, a significant amount of research in developing
renewable energy is required. Bioenergy presents the largest source of renewable energy today,
used for heat, electricity, and transport fuels.3 The used of biomass power play a key role in
decarbonizing electricity systems by replacing energy source from fossil fuels to carbon-neutral
fuels. More importantly, liquid biofuels can be used to replace petroleum-based transport fuels
without significant change of current infrastructure. It is projected that modern bioenergy could
contribute over 20 % of global final energy supply in 2030, doubling its share from 10% in 2010.4
Among all available sources for bioenergy, microalgae as the third generation of biomass have
several advantages including fast growing rate, rich lipid content, reduced land-use, synergy effect
in wastewater treatment and do not compete directly with food. The fast-growing microalgae is a
2
sustainable source to produce fuels, food, pharmaceuticals, chemicals and animal feed (Figure 1-
1). However, despite these advantages biofuel production from algae still faces serious challenges
especially the high energy and economic costs of current downstream processing techniques.
Unlike terrestrial plants, microalgae are harvested as wet biomass which contains a significant
amount of water than crop-based biomass. Conventional conversion process used for terrestrial
plants involving lipids extraction and transesterification to biodiesel requires an energy intensive
drying process which increases the carbon intensity of this approach. From a life cycle assessment
of biofuel production, drying algal biomass sufficiently for conventional lipids extraction
consumes more than 90% of the energy content in the algal oils.5–7 Therefore, the future algae-to-
biofuel conversion techniques must be adaptable to wet biomass to obtain a net positive energy
balance.8
Figure 1-1. Microalgae based biorefinery integrated with related industries to produce
numerous sustainable deliverables. Reproduced from ref9, © 2010, with permission from
Elsevier.
Similar to how nature generates fossil fuels from buried biomass under high temperature and high
pressure in a geological timescale, an emerging conversion approach adaptable to wet biomass
3
called Hydrothermal Liquefaction (HTL) has attracted great attention in recent years. HTL shares
the same mechanism to convert biomass into biocrude but with a duration as short as a few minutes.
At elevated temperature (200-380 °C) and pressure (5-28 MPa) with presence of water, HTL
chemically and physically cracks down large biomolecules into small fractions and simultaneously
transforms them into biocrude.10 HTL shows two prominent advantages over conventional
conversion technologies: drying treatment for feedstock is eliminated since a wet slurry is directly
input into the HTL process; and higher biocrude yields are usually found in HTL experiments
because HTL converts not only lipids but also other biomass such as carbohydrates and proteins
into biocrude.9–11 These advantages of HTL have made it a unique and promising path to produce
liquid biofuels from microalgae feedstock at a favorable energy return on investment. However,
HTL experiments have been conducted in laboratory scale batch reactors that (1) require
substantial heating times lead to ambiguity in the reported results and (2) lack of in-situ observation
of the reactions. Therefore, in order to understand the reaction mechanism of HTL and reach the
full potential of biocrude yield, precise control of processing conditions and in-situ visualization
of the reaction process are essential.
Producing high-value bioproducts from microalgae has attracted great attention in recent years due
to high processing costs and low market value for biofuels, with current low oil price further
driving this transition. Figure 1-2 indicates related microalgae products with their estimated market
value and size. Natural astaxanthin from microalgae is a highly valuable bioproduct with
tremendous health benefits and has become the primary source for the nutraceutical industry. The
extraction of astaxanthin from the microalgae is hindered by a thick cell wall that prevents solvent
extraction and digestion from direct consumption. This barrier is extremely robust to chemical and
physical disruption, making astaxanthin extraction difficult. Hydrothermal processes utilizing high
temperature and pressure have shown promise for extraction of bioactives, woody biomass
decomposition and biocrude formation which has strong potential for cell-wall disruption. In
addition, supercritical carbon dioxide (ScCO2) has attracted attention recently as a green solvent
for the extraction of bioactive compounds. The overall extraction efficiency of ScCO2 is highly
dependent on the characteristics of the feedstock, operating conditions (temperature, pressure,
duration) and addition of modifiers/co-solvents. Therefore, the development of effective cell wall
disruption techniques and understanding of the ScCO2 extraction mechanism are critical to the
design of commercial-scale astaxanthin extraction reactors and processes.
4
Figure 1-2. Sunlight-to-biomass conversion efficiency and losses along the path, as well as
value and market size of related microalgae products. Reproduced from ref15, © 2016, under
CC-BY license.
Microfluidic approaches with high degree of process control have been applied for bioenergy
applications.16–18 Recent applications of microfluidics in high temperature and pressure
processing, particularly in the context of chemical synthesis19,20 and phase analysis21,22 made it a
promising technique for microalgae downstream processing. In addition to precise control on
temperature and pressure, continuously flowing microfluidic reactors can achieve extremely high
heating rates avoiding ambiguity due to long heating times. Moreover, microfluidic devices with
in-situ observation using fluorescence enables direct, real-time monitoring of processes at the
cellular level. The ultimate goals of these works are to (1) develop a high temperature and pressure
microfluidic platform with in-situ observation capability and (2) perform microalgae to biofuels
and bioproducts conversions.
1.2 Thesis Overview
This thesis is focused on the development of high temperature and pressure microfluidic platform
and microalgae downstream processing techniques particularly biomass-to-biocrude conversion
via hydrothermal liquefaction, hydrothermal disruption of cell wall, and supercritical CO2
extraction of astaxanthin. The results and the microfluidic platform on which they were collected
5
represent the first of their kind in the field of microalgae downstream conversions. These
researches provide unprecedented insight into biomass to biofuels and bioproducts conversions
that will guide the design of large scale reactors and processes. Supplementary contributions
include projects which I contributed to as co-author and are included in appendices.
Chapter 1 briefly describes the research motivations of building a sustainable future by producing
energy and high-value bioproducts from microalgae biomass. This chapter discusses the key
aspects of these two downstream processes and briefly describes the demand for novel high
temperature and pressure microfluidic platform in that context.
Chapter 2 provides a background of related work in the field of hydrothermal processes for
biomass, supercritical CO2 extractions of bioactive compounds and microfluidics in general. These
concepts introduced in this chapter include processing evaluations using LCA and TEA, water
characteristics under subcritical conditions, biomass composition and conversion, supercritical
CO2 extraction and high temperature and pressure microfluidics.
Chapter 3 reviews the current microalgae biomass downstream processes and emerging techniques
mainly for bioenergy applications. This chapter is part of the review manuscript published in
Sustainable Energy and Fuels, and portions of this chapter have been reproduced with permission
from The Royal Society of Chemistry.
Chapter 4 demonstrates microalgae biomass-to-biocrude conversion on a chip via hydrothermal
liquefaction. A high temperature and pressure microfluidic platform with optical access and
precise control of processing parameters was first demonstrated in the research of HTL. This
chapter was published in Lab on a chip and featured as HOT article – reproduced by permission of
The Royal Society of Chemistry.
Chapter 5 presents the hydrothermal disruption of cell wall for astaxanthin extraction on a
microfluidic platform. Hydrothermal disruption at a temperature of 200 °C was shown to be highly
effective, resulting in near-complete astaxanthin extraction from wet biomass – a significant
improvement over traditional methods. This chapter was published in Green Chemistry –
reproduced by permission of The Royal Society of Chemistry.
6
Chapter 6 presents astaxanthin extraction from Haematococcus pluvialis using supercritical carbon
dioxide with and without co-solvents. The associated manuscript is in preparation. This chapter
was submitted for publication.
Chapter 7 summarizes the author’s contribution in the downstream conversion of biomass to
biofuels and bioproducts and provides potential directions for future studies.
7
Chapter 2.
Introduction
To reach a sustainable future, integrated biorefineries have been suggested as a means to produce
food, energy, chemicals and materials.23 Microalgae is particularly well positioned due to the many
benefits over other sources of biomass such as high growth rate, low land requirement, high oil
content and recycling of nutrients from wastewater. In this chapter, two major microalgae
applications are introduced, liquid biofuels and high-value bioproducts.
In context, current microfluidic devices used for high temperature and pressure reactions are
reviewed. There have been efforts to scale microfluidic methods for processing and scale
microfluidic reactors for emulsion production24–26 and the bulk production of chemicals under
challenging experimental conditions19,27–29. These reactors generally approach the challenge of
producing bulk product by 1) parallelizing (multiple parallel processes on a single chip) and 2)
numbering out (having many chips operating in parallel). There are several companies in this space
making commercial products such as Corning, Ehrfeld Mikrotechnik, Alfa Laval and Chemtrix,
who are developing modular microfluidic reactors. Other companies such as Uniqsis, Microinnova
and FutureChemistry are developing integrated continuous flow reactors for chemical synthesis
without using a chip. While some of the above-described methods may be applied to increase
output from methods developed in this thesis, production of bulk product is not the focus here.
Rather, this thesis focuses on developing and demonstrating microfluidic technologies to study
biomass conversion processes, with the goal to inform industry and the academic community by
resolving previously opaque processes at the single-cell scale. The insights herein can be applied
to improve large conversion systems already operating at scale.
2.1 Microalgae for biofuel production
With increasing energy demands resulting from accelerating population growth and global
development, the need to curb anthropogenic greenhouse gas emissions is increasingly important.
Biofuels from microalgae are both renewable and potentially carbon neutral and are therefore a
compelling alternative to fossil fuels for transportation, which accounts for 28% of global energy
8
consumption30. For biofuel production, microalgae have many benefits over other sources of
biomass. For instance, microalgae can be cultivated in brackish/wastewater environments which
do not interfere with food supplies or contribute to greenhouse gas emissions through land-use
change31. Perhaps the most significant advantage is the rapid growth rate of microalgae which can
double in as few as two hours.32 Moreover, microalgae cultivation can be integrated directly into
wastewater treatment plants and flue gas producing facilities to recycle wasted nitrogen,
phosphorus and CO2 as nutrients.7,33 Although these benefits make microalgae an outstanding
candidate for biofuel production, a major obstacle in producing carbon neutral and energy
favourable biofuel is the poor efficiency of converting the raw biomass into liquid biofuels.
2.1.1 Assessments for downstream conversions
Conventional methods to produce biofuel from biomass requires the extraction of lipids from cells
which can then separately be converted to biodiesel, also known as fatty acid methyl esters
(FAME). During this transesterification process, the major components in algal lipids,
triacylglycerides (TGA) are converted to FAME through a reaction involving alcohol catalyzed by
acids or alkalis at elevated temperature. This process is mature and commonly used in the
conversion of vegetable oils to biodiesel. In this process, lipids are the only intracellular
components contributing to the biofuel yield whereas carbohydrates and proteins are unused. As a
result, the overall biodiesel yield of this process is strongly correlated to the growth rate of
microalgae, lipid content of the cell, the efficiency of extraction method, and the lipids-to-biodiesel
conversion rate. Generally, for a specific algal strain, the growth rate and lipid content have an
inverse relationship, where the oil-rich microalgae tend to grow slower than most low-oil algal
strains. Even within the same species, the proportion of individual cell constituents largely depends
on environmental conditions. More importantly, the challenge for this method is that the biomass
must first be dried, which significantly increases the carbon intensity of the final fuel as well as its
cost. A more recent conversion approach, hydrothermal liquefaction (HTL), has attracted a great
deal of attention due to its ability to directly convert wet biomass to biofuels. Other downstream
conversion methods and emerging techniques are reviewed in Chapter 3. In this section, Life cycle
Assessment (LCA) and Techno-economic Analysis (TEA) are used to evaluate different
downstream processing routes.
9
Recently, a life cycle assessment (LCA) has been widely used to investigate energy balance and
greenhouse emissions on algae-derived fuels. Slade and Bauen34 reviewed several recent LCA
studies on algal biofuel production including cultivation, harvesting and oil extraction and found
that only open pond systems were able to achieve a positive net energy (Figure 2-1a). For raceway
pond systems, more than 80% of overall energy consumed was due to biomass drying and
dewatering with the exception of the LCA performed by “[11] Stephenson”. In this case the authors
assume the use of an effective oil extraction process adaptable to wet biomass which does not exist
yet. Analogous to these energy balance analyses, the carbon emission chart (Figure 2-1b) indicates
that only raceway pond systems can reduce emissions to less than 84g CO2e/MJ (similar to
petroleum-derived diesel) and the majority of those emissions are again associated with biomass
drying and dewatering. In agreement with other LCA studies, drying algal biomass sufficiently for
conventional lipids extraction consumes more than 90% of the energy content in the algal oils5–7.
10
Figure 2-1. A summary of life cycle assessments for biofuel production form microalgae. a)
Net energy ratio for microalgae biomass production and b) Illustrative estimates for carbon
dioxide emissions from algal biomass production. Reproduced from ref34, © 2013, with
permission from Elsevier.
11
A LCA from Sills et al.8 performed a direct comparison of the energy balance and GHG emissions
of biofuel production from microalgae using HTL and lipid extraction. The results indicated that
HTL approach generates a higher energy returns on investment (EROI) and lower global warming
potential than dry extraction methods (Figure 2-2). Although the EROI ratio of HTL is just slightly
higher than 1 in the base case, it is expected to increase over time with the development of
associated technologies. For petroleum-derived fuels the EROI used to be on the order of 100 but
has fallen to 4-5 nowadays35. It is then foreseeable that the EROI of algae-derived fuel could
surpass petroleum-derived fuel over time and the incentives to mitigate GHG emissions could
accelerate this transformation. Moreover, integrating HTL with wastewater treatments in algal
biofuel production brings additional environmental benefits into this assessment.36
Figure 2-2. a) Energy balance and b) Global Warming Potential for biofuel production using
wet and dry lipid extraction. Reproduced from ref8, © 2013, with permission from American
Chemical Society.
12
Figure 2-3. Techno-economic analysis of liquid fuel produced from hydrothermal
liquefaction of woody biomass a) Effect of improvement on MFSP from state-of-technology
(SOT) case to goal case. b) Sensitivity analysis of parameter variation on the MFSP of the
goal case. a) and b) are reproduced from ref37, © 2014, with permission from Elsevier. c)
Sensitivity analysis of the MFSP of biofuel produced from defatted microalgae via HTL.
Reproduced from ref38, © 2014, with permission from Elsevier.
A techno-economic analysis (TEA) is also commonly used to guide the decision making process
when commercializing new technologies. In terms of algal biofuel production, it is crucial to select
the pathways that show the most economic promise. The Pacific Northwest National Laboratory
13
(PNNL) has used a TEA to investigate the economic value of producing biofuels from woody
biomass via HTL37. Based on the laboratory scale testing results of current technologies, the
minimum fuel-selling price (MFSP) from HTL is $4.44/gallon of gasoline-equivalent value.
However, the technology improvement of HTL process alone is expected to reduce the MFSP by
$1.34/gallon or 31% of the MFSP (Figure 2-3a). A sensitivity analysis performed on an HTL
process indicates that the cost of feedstock and the efficiency of HTL process itself have the largest
effects on the MFSP (Figure 2-3b&c). With technology breakthroughs in strain selection, genetic
modification, cultivation and effective harvesting, however, it is possible that the price of
feedstocks could be dramatically reduced. Current price for dry ash free algae is estimated at
$430/ton including cultivation, harvesting and dewatering to 20 wt% dry solids39. This price is
much high than that of corn of $170/ton while the mass productivity of microalgae is about 10
times higher that of corn8. Since the cost reduction of microalgae feedstock is a foreseeable trend,
the performance of HTL as the largest influencing factor in the downstream processing is critical
to the commercialization of microalgae biofuel production.
Overall, results from TEA are generally in agreement with the LCA in suggesting that downstream
conversion processes using HTL is a promising approach due to its ability to directly convert wet
biomass. The development of this technology could then have a significant impact on the next
generation of biofuel production.
2.1.2 Water under subcritical conditions
Hydrothermal liquefaction, which directly converts biomass into biocrude at elevated temperatures
and pressure, has attracted much attention. This technique was originally called “pressurized hot
water extraction” and later known as “subcritical water extraction” because under subcritical
conditions, water is less polar, and more soluble to organic compounds. Later, water at subcritical
conditions was found to function as more than just a solvent and to classify the process better it
was divided into three categories: hydrothermal carbonization (< 200 °C), hydrothermal
liquefaction (200 - 380 °C) and hydrothermal gasification (> 380 °C). In these processes, water
plays an essential role and understanding the fundamentals of water under these subcritical
conditions (below 374 °C and 22MPa) is critical to the development of HTL technique for
producing biofuels from biomass. In this thesis, carbonization and gasification are not discussed
in detail as they are used mainly for solid and gaseous products respectively.
14
Under ambient conditions water is a polar solvent excellent at dissolving polar compounds and
salts. As temperature increases, the characteristics of water change substantially. In HTL, water is
not considered an inert medium but an active participant in the reaction, either as a reactant or
catalyst. The characteristics of water and the role of water in high-temperature reactions are well
reviewed by Akiya and Savage.40 At subcritical conditions, the dramatic changes in the properties
of water are mainly displayed in the changes to: 1) hydrogen bonding, 2) dielectric constant, 3)
ion product, and 4) density.
The strong hydrogen bonding of water is the source of its many unique properties. With increasing
temperature and decreasing density, the hydrogen bonding in water becomes weaker and less
persistent. For example, water at 300 °C and 0.5 g/cm3 retains only 30-45 % of the hydrogen
bonding that exists at ambient conditions.41 In addition, the hydrogen bond network in water at
high temperature exists in the form of small clusters, rather than the infinite percolating network
of hydrogen bonds found in ambient liquid water. In general, the average cluster size decreases
with increasing temperature and decreasing density which leads to a high “local concentration” of
H+ and OH- ions42 that enhance ionic reactions.
The dielectric constant (relative permittivity) is the ratio of the permittivity of a substance to the
permittivity of free space, which indicates a relative measure of its chemical polarity. The density,
static dielectric constant and ion dissociation constant (Kw) of water at 30 MPa as a function of
temperature is shown in Figure 2-4. In general, the static dielectric constant of water decreases
with increasing temperature and decreasing density. For example, the dielectric constant of water
decreases from 78.85 to 19.66 as temperature increases from 25 °C to 300 °C, resulting in water
molecules changing from highly polar to fairly nonpolar.43 As a result, small organic compounds
are highly soluble in subcritical water and completely miscible in supercritical water.44,45 In HTL,
water can dissolve organic compounds and enhance reaction with organic compounds.
15
Figure 2-4. Density, static dielectric constant and ion dissociation constant (Kw) of water at
30 MPa as a function of temperature. Reproduced from ref46, © 2008, with permission
from Royal Society of Chemistry.
The ion dissociation constant (Kw) is defined as the product of the concentration of H+ and OH- in
water and it also dramatically increases with the increasing temperature to a maximum of about
10-11 mol2/kg2 at around 300 °C (Figure 2-4). For the temperature in the HTL range (200-380 °C),
water is an effective medium for both acid- and base-catalyzed reactions due to the high
concentration of H+ and OH- ions. These effects are further enhanced due to smaller water cluster
sizes resulted from aforementioned weaker hydrogen bonding.42
Water density also varies greatly with temperature and pressure. The decrease in water density
contributes to chemical reactions mainly in two ways: 1) increase in diffusivity of water40 and 2)
increase in ion dissociation constant47. The increase in ion dissociation constant and diffusivity
enhance reactions by increasing the concentration of ions and mass transport, respectively. With
the change in density from 1 to 0.1 g/cm3, the diffusivity increases by an order of magnitude. These
changes in transport properties can affect reactions influenced by diffusion time scales and solvent
dynamics.
16
The dramatic change in the physiochemical properties of water at high temperature suggests the
promising path of converting biomass to biofuels via hydrothermal liquefaction. In addition,
subcritical water can serve as a hydrogen source in reactions which can greatly affect the product
distribution.40 In summary, water at high temperature has the ability to carry out a unique set of
chemical reactions and it may help to explain why HTL is more likely to convert biomass into oil
than pyrolysis. Since water is the cheapest and most environmentally benign solvent and is present
with the microalgae feedstock, HTL of biomass with other additives such as alcohols and catalysts
are avoided and discussed elsewhere.48
2.1.3 Characterization of microalgae and post-HTL products
Characterizing the physical and chemical properties of microalgae is critical to the design of
appropriate downstream processes as well as an understanding of reaction mechanism in HTL.
Also, classifying and quantifying the post-HTL products is essential for the understanding of HTL
processes. In this section, major algal components and analytical methods to quantify them will be
introduced.
The major organic components of algae including microalgae and macroalgae are lipids, proteins
and carbohydrates. Unlike woody biomass, most species of algae do not contain lignin. This is
considered a benefit for algae because lignin is relatively resistant to chemical degradation and
produces a significant amount of solid residue in HTL.49 Lipids are non-polar aliphatic compounds
namely triacylglycerides which is the main screening criterion in biodiesel production.32 However,
algae with high lipid contents usually have slow growth rates and low biomass productivity.50
Since HTL converts all organic components of biomass into biocrude, rather than being purely
limited to lipids, the algae used for HTL can have low lipids content as indicated in Figure 2-5.
Nevertheless, high-lipid algae tend to have higher biocrude yields because the conversion rate
follows the trend of lipids > proteins > carbohydrates.12,51 Determination and quantification of
lipids are commonly performed through solvent extraction followed by Gas Chromatography –
Mass Spectroscopy (GC-MS). For lipids extraction, a modified Bligh-Dyer extraction method
utilizing chloroform and methanol is preferred due to the higher extraction efficiency as oppose to
Soxhlet extraction, which utilizes only non-polar organic solvent such as hexane. Acid hydrolysis
of lipids to produce isolated fatty acids is usually used prior to the GC-MS.
17
Figure 2-5. Characteristics for algae used for HTL. The green, black and red dashed arrows
indicate the mass fraction of lipids, proteins and carbohydrates, respectively. Reproduced
from ref10, © 2014, with permission from Elsevier.
Proteins are major components of algae and are the primary component containing sulfur and
nitrogen. The proteins content in the dry biomass is therefore estimated based on the nitrogen
composition. A thermogravimetric analysis (TGA) based on continuous measurement of biomass
can be used to determine the moisture and ash content before HTL. The first-stage mass loss (up
to 110 °C) generally refers to the moisture content of the mix while the mass leftover at
temperatures greater than 900 °C is considered as ash. Dry biomass generated after moisture
removal is subjected to an elemental analysis to determine the amounts of constituent elements:
carbon (C), hydrogen (H), nitrogen (N) and sulfur (S). In the elemental analysis, a complete
combustion process with sufficient oxygen is used to convert biomass into gases such as CO2, SO2,
N2 and NOx for quantification. Oxygen (O) content is usually calculated by difference (%O =
100% − %C − %H − %S − %N − %𝐴𝑠ℎ ). The protein contents are estimated from the
nitrogen content with a conversion factor of 6.25.52
18
The carbohydrates in biomass are mainly polysaccharides, starch, cellulose and hemicellulose. The
carbohydrate contents in biomass are determined by subtracting the lipid, protein and ash content
from 100%.53 Trace amounts of minerals are also contained in biomass and can be determined by
inductively coupled plasma atomic emission spectrometer (ICP-AES).
Figure 2-6. HTL of microalgae procedure and product separation process. Reproduced from
ref54, © 2012, with permission from Elsevier.
The previously described analysis techniques describe the contents of wet biomass. It is similarly
important to be able to quantify the outputted products after biomass has undergone HTL. These
post-HTL products consist of liquid products, gaseous products and solid residue. The portion of
liquid products that is dissolvable in dichloromethane is defined as biocrude and the rest is
considered as aqueous products. Within the biocrude, the portion that is dissolvable in hexane is
considered as light biocrude while the remaining is heavy biocrude. The separation of each
component is shown in Figure 2-6. Biocrude is a complex mixture. The easiest and most interesting
characteristic of biocrude is the higher heating value (HHV) which is estimated from the elemental
composition using a modified Dulong’s formula.10,54 A more detailed analysis of biocrude can be
performed using GC-MS and a large number of compounds are found mainly consist of cyclic
nitrogenates, cyclic oxygenates and cyclic nitrogen and oxygen compounds.55 Products in the
aqueous phase have been quantified using elemental analysis on water-evaporated residue.
Gaseous products are also quantifiable using gas chromatography with a thermal conductivity
detector (TCD).56 The solid residue is separable by centrifuge and mainly contains ash and
inorganics.
19
2.1.4 HTL of biomass and effects of processing conditions
Hydrothermal liquefaction has been studied on multiple species of algae and woody biomass as
well as individual biomass components such as lipids, proteins and carbohydrates. In general, the
biocrude yield varies significantly (20% to 87%) with different types of feedstock and processing
conditions10, but follows the trend according to the composition in biomass: lipids > proteins >
carbohydrates (Figure 2-7).12,51,57
Figure 2-7. Yields of products from hydrothermal liquefaction of different types of feedstock
in: a) water, b) sodium carbonate, c) formic acid. Reproduced from ref12, © 2011, with
permission from Elsevier.
20
Lipids can be readily hydrolyzed by HTL at greater than 90% conversion rates to produce fatty
acids that are relatively stable in subcritical water58. King et al. obtained free fatty acid yields of
90-100 % from hydrolysis of soybean oil in 10-15 min with subcritical water at 330-340 °C and
13.1 MPa.59 Proteins are polymers of amino acids that are linked through peptide bonds connecting
carboxyl and amine groups. In HTL, proteins are hydrolyzed slowly to produce amino acids which
will further be degraded in subcritical water via decarboxylation and deamination reactions.60 The
hydrolysis rate of proteins and the decomposition rate of amino acids highly depends on the
reaction temperature. Rogalinski et al. performed hydrolysis of bovine serum albumin and found
the amino acid decomposition rate is higher than the hydrolysis rate at temperatures above 250 °C
with the highest amino acid yield obtained at 290 °C in 65 s.61 HTL of several carbohydrates have
also been well studied in the past including monosaccharides, such as glucose and fructose, and
polysaccharides, such as starch and lignocellulose.60 The destruction of monosaccharides in
particular is drastic under hydrothermal conditions.62 For instance, Kabyemela et al.63 observed a
55% conversion of glucose after 2 s at 300 °C and 90% conversion after 1 s at 350 °C.
Polysaccharides including starch, cellulose and hemicellulose are fundamentally polymers of
monosaccharides suggesting that the HTL process consists of two stages: polysaccharide
depolymerization and monosaccharide degradation. Starch and cellulose are polymers of glucose
that are linked by different bonds. Rogalinski et al.64 found that 100 % conversion of cellulose was
achieved within 2 min at 280 °C while starch hydrolysis was faster than cellulose hydrolysis in
hydrothermal conditions. Hemicellulose is a heteropolymer composed of monosaccharides
including xylose, mannose, glucose, galactose and others. It was found that nearly 100% of
hemicellulose could be hydrolyzed at 230 °C and 34.5 MPa in 2 min.13 The timescales of the above
HTL of carbohydrates were mainly over a span of a few minutes, suggesting that rapid heating is
crucial to study the specific effects of hydrolysis and degradation under hydrothermal conditions.
21
Figure 2-8. Biocrude yield of HTL as a function of heating rate. a) Temperature profiles with
corresponding biocrude yields from HTL of Nannochloropsis sp. for different set-point
temperatures of 300, 500, 400, and 600 °C. Reproduced from ref65, © 2013, with permission
from American Chemical Society. b) The effect of heating rate on the products yield from
HTL of macroalgae. (T = 350 °C; holding time = 15 min; and biomass/water ratio = 1/10,
w/w). Reproduced from ref66, © 2014, with permission from Elsevier.
22
Several research sources indicate that biocrude yield is tightly related to the processing
conditions67,68 such as temperature, pressure, residence time and especially heating rate65,66 (Figure
2-8). Under current system optimizing the many parameters involved in HTL remains a challenge.
For instance, Faeth et al.65 reported that their highest biocrude yield (66 wt%) from HTL was
produced when operating at their maximum heating rate of 230 °C/min for a duration of 1 min,
meaning the temperature over the course of the experiment was constantly changing. Higher
heating rates were not investigated due to the limited heating capability of their experimental setup.
To date, HTL experiments have been conducted in laboratory scale batch reactors using sandbaths,
ovens or heating coils for temperature control. These approaches have two critical issues. First,
due to the large fluid volumes and large physical size of the apparatus, substantial heating times
are required, which are often ignored in the subsequent analyses and reported reaction times. These
long heating times lead to ambiguity in the reported optimal values for temperature, heating rate,
and reaction time since the heating and pressurization delays blur the results. Second, in situ
monitoring of the reaction process is not possible in current reactors, precluding real-time
quantification of the reaction process. In order to understand the reaction mechanism of HTL and
reach the full potential of biocrude yield, precise control of processing conditions and in-situ
visualization of the reaction process are essential.
2.2 Microalgae for high-value bioproducts
The potential commercialization of microalgae mainly falls into two categories: 1) large-volume,
low-value bulk commodities such as biofuels and food/feed and 2) low-volume, high-value special
products such as pigments, health care and additives. Although HTL shows promise in biofuel
production, further development of this technology is required to lower the production cost to
compete with fossil fuels. On the other hand, the production of high-value products from
microalgae could be currently profitable69 and the maturation of associated technology will also
benefit the development of biofuel production. The market value of microalgae products is
indicated in Figure 2-9.
23
Figure 2-9. Market value of the microalgal components and total selling price of the
biomass for different market scenarios. Reproduced from ref69, © 2016, under CC-BY
license.
Among all high-value products, natural astaxanthin from microalgae – Haematococcus pluvialis
is one of the most promising candidates showing high market value and large market volume.
Astaxanthin is a strong antioxidant, a pigment and has important applications in the nutraceuticals,
cosmetics, food and aquaculture industries.70,71 However, critical bottlenecks and major challenges
also exist in the commercial scale production of astaxanthin especially in astaxanthin recovery
from biomass. The accumulation of astaxanthin is strongly associated with the formation of a rigid
cell wall that prevents solvent extraction. Also, for astaxanthin extraction, a chemically inert and
greener solvent is required in terms of product quality and environment concerns. Therefore,
development of effective cell-wall disruption and ScCO2 extraction techniques are crucial to the
commercial implementation of astaxanthin production at large scale.
2.2.1 Cell structure of Haematococcus pluvialis
H. pluvialis is a freshwater unicellular green microalgae with four distinguishable stages in the life
cycle: macrozooid (during cell division), microzooid (after germination), palmella, and cyst
(aplanospore) (Figure 2-10b). The first three stages are generally referred as green vegetative phase
((A) and (B) in Figure 2-10a) and the cyst (aplanospore) stage is called red phase ((C) and (D) in
Figure 2-10a) suggesting an accumulation of astaxanthin in the cell.
24
Figure 2-10. a) Microscopic images of Haematococcus pluvialis in life cycle. (A) Green
vegetative motile cell; (B) Green vegetative palmella cell; (C) Astaxanthin accumulating
palmella cell in transition to aplanospore; (D) Astaxanthin accumulated aplanospore cell.
Scale bar: 10 μm. Reproduced from ref72, © 2016, under CC-BY license. b) Illustration of
life cycle of H. pluvialis. Reproduced from ref73, © 2013, under CC-BY license.
Harvesting of astaxanthin is performed on mature aplanospores resulting in the maximum amount
of astaxanthin during the life cycle (2-5 % of dry weight). At this stage, cells become resistant to
extreme environmental conditions and show two distinct structures: 1) a thick and right trilaminar
sheath and 2) a secondary cell wall. The development of the rigid cell wall and its structure during
the life cycle is shown in Figure 2-11. Hagen et al.74 reported the aplanospore cell wall contained
70 % carbohydrates (66% hexoses), 3% cellulose, 6% proteins, and 3% acetolysis-resistant
material. Due to the fast reaction rates of carbohydrates under hydrothermal conditions, subcritical
water could be an effective solution to disrupt cell walls.
25
Figure 2-11. Summary of five developmental states of the cell wall during aplanospore
morphogenesis in Haematococcus pluvialis: I, 1-week-old flagellates ; II, flagellates at least
2 weeks old just rounding off; III, 2- to 3-week-old aplanospores; IV, at least 3-week-old
aplanospores; V, aplanospores in their final state. CYP, cytoplasm; IS, interspace; PL,
plasmalemma; PW, primary wall; SV, secretory vesicles; SW, secondary wall; TCL,
tripartite crystalline layer; TLS, trilaminar sheath; W1±W7, layers of the extracellular
matrix. Reproduced from ref74, © 2002, with permission from Taylor & Francis.
2.2.2 Cell wall disruption and extraction techniques
Astaxanthin accumulates inside a thick, rigid cell wall of H.p cysts and the performance of
recovery is highly dependent on the cell-wall disruption and subsequent solvent extraction
processes. Intact cysts of H.p without any cell-wall disruption treatment only releases ∼20% of
the internal astaxanthin using an acetone extraction solvent over a 16 hour period.75 Both chemical
disruptions using acid or base and biological disruption using enzyme followed by acetone
extraction recovered 25-40 % of astaxanthin, a slight improvement over the control. Therefore,
having an effective cell-wall disruption treatment is critical to the subsequent solvent extraction
process. Kim et al.76 recently reviewed different astaxanthin recovery processes and separated
them into four groups based on cell-wall disruption methods: chemical, physical, physico-
chemical, and biological methods. Physical disruptions use grinding, bead beating and French-
26
pressure-cell while physico-chemical disruptions are a combination of physical (bead beating,
grinding, milling or ultrasound) and chemical (organic solvent, acid or breaking buffer) processes.
They are reported with higher yields than chemical disruptions but there is ambiguity in the
comparison due to three issues: 1) most physical disruptions use dry biomass which might be
processed differently; 2) different solvents are used in the subsequent extraction processes such as
methanol/dichloromethane mixture, acetone and ethyl acetate; 3) recovery rates were reported as
mg/g cell which is strongly influenced by the feedstock as opposed to the percentage of total
astaxanthin extracted. Therefore, a platform that can quickly and visually assess the effectiveness
of different cell-wall disruption methods and extraction conditions is desired to provide a fair
comparison and insights into this important process.
Among all extraction solvents, supercritical carbon dioxide (ScCO2) has attracted much attention
as a green solvent for the extraction of bioactive compounds.77,78 Supercritical CO2 features three
major advantages over organic solvents: 1) it is abundant and benign to human health and the
environment, 2) a solvent-free extract can easily be produced and extraction by CO2 evaporation
at room temperature and pressure, 3) bioactive compounds are well-preserved due to the inert
chemical property of CO2 and relatively low critical temperature (31.1 °C). However, the overall
extraction efficiency of ScCO2 is highly dependent on the characteristics of the feedstock75,76,79,80,
operating conditions (temperature, pressure, duration)81,82 and addition of modifiers/co-solvents83–
85. Understanding the extraction mechanism of ScCO2 with and without co-solvents is critical to
the development of astaxanthin production from microalgae. In addition, to avoid extremely high-
pressure operation while remaining the high extraction rate and efficiency, a platform with optical
access is desired to bring insights into the extraction mechanism and the relationship of co-solvents
to ScCO2.
In summary, the extraction efficiency, extraction rate, scalability along with the energy
consumption should be considered in the development of cell-wall disruption and solvent
extraction techniques for industrial-scale astaxanthin production form H.pluvialis biomass.
2.3 High temperature and pressure microfluidics
Microfluidic technologies have been used widely in chemical synthesis, health care, fundamental
physics and bioenergy production mainly due to the following advantages: 1) they require less
reagent as compared to bulk systems; 2) they provide a high degree of control over processing
27
parameters; and 3) they provide safer working environments when performing dangerous
chemistry.19,86 In addition, microfluidic reactors have often claimed to produce higher yields than
bulk reactor due to the scale-dependent processes of heat and mass transfer and large surface-area-
to-volume ratio. As a result, there have been some examples of people using parallel microfluidic
reactors (numbering up) to increase productivity rather than increasing the characteristic
dimension of the channel (scaling up). Given the low volume size of microfluidic channels but
high throughput on information from a rapid screening of reaction conditions, the actual product
of most microfluidic reactors is the information.
Figure 2-12. Examples of (a) metal, (b) glass and (c) silicon/glass microreactors. Reproduced
from ref20, © 2011, with permission from Elsevier.
The advantages of using microfluidic reactors are even more pronounced in chemical synthesis
involving harsh process conditions such as high temperature and pressure.87,88 Commonly used
microfluidic reactor made of polymers such as polydimethylsiloxane (PDMS) and
polymethyacrylate (PMMA) cannot withstand high temperature and pressure. Metal-based
microreactors fabricated using conventional machining are good for harsh condition processing
but are limited by the lack of optical access. Glass microfluidic reactors have the advantage of
utilizing in-situ optical characterization techniques but are limited by the low thermal conductivity.
Microfluidic reactors fabricated out of silicon and glass have advantages in both high temperature
and pressure reactions by allowing: 1) optical access through the glass, 2) high thermal
28
conductivity through the silicon, and 3) both high pressure and temperature. Examples of different
microfluidic reactors are shown in Figure 2-12.
Silicon/glass microfluidic reactors have been used for materials synthesis88, fuel conversions21,
phase behavior studies89 and supercritical fluid extractions90. The silicon/glass chip is fabricated
by silicon etching using solutions or deep reactive ion etching (DRIE) followed by anodic bonding.
By matching the thermal expansion coefficient of glass to silicon, the chip can withstand high
pressure at various temperatures. The high-pressure sealing is achieved using modular packaging
at room temperature by cutting a silicon window to isolate the high-temperature region. Achieving
high temperature and pressure conditions in microfluidic reactors suggests promising applications
of water under hydrothermal conditions and supercritical fluid extractions.
29
Chapter 3.
Current Downstream Processing and
Emerging Technologies for Microalgae
This chapter is a portion of a review published in Sustainable Energy and Fuels and reproduced
from ref91 – reproduced by permission of The Royal Society of Chemistry. The candidate was one of
the equally contributing authors in this work and played the primary role in reviewing the emerging
techniques in downstream processing. Additional authors for the work include Dr. Scott C.
Pierobon, Dr. Percival Graham, Mr. Brian Nguyen, Mr. Evan G. Karakolis, and Prof. David
Sinton.
3.1 Optimizing downstream processing
Unlike terrestrial plants, microalgae are harvested as wet biomass which requires different
downstream processing from crop-based biomass. To complete the microalgae-to-biofuel
conversion route, pretreatment of biomass such as harvesting, thickening and dehydration are
typically applied to simplify conversion processes92–96. However, given the high latent heat
vaporization for water (2,265kJ/kg) the energy intensity and associated cost of drying is a
fundamental challenge for microalgal biofuels. Given the fact that most microalgae have a total
energy content on the order of 18,000 kJ/kg97,98, drying a microalgae slurry at a concentration
below ~20 wt% (which contains ~3600 kJ/kg of energy) is of limited practicality for biofuel
application. As such, the strategy to operate a microalgae operation is to either avoid massive
drying or produce high-value products to offset the high processing costs. In this section, we will
first discuss the dewatering and drying processes, then describe how dry biomass, wet biomass,
microalgae in culture and direct secretion can be implemented. Microalgae in culture involve
directly using cells, wet biomass is typically concentrated from harvested cells and dry biomass is
30
obtained by drying wet biomass. Each area will be briefly reviewed while highlighting emerging
technologies.
3.2 Harvesting and dewatering processes
Initial harvesting is typically accomplished by either screening or sedimentation, where the
concentration is increased from 0.1% of total suspended solids (TSS) to a slurry of about 2-7%
TSS95,96,99,100. In terms of screening, microstraineres and vibrating screens are the most commonly
used. Standalone sedimentation by gravity is highly energy efficient for large cells99,101 but is
unsuitable for many types of microalgae. Sedimentation can be enhanced by forming larger
aggregates of cells, typically by chemical flocculation and coagulation. Ultrasound-assisted
harvesting forces cell aggregation at acoustic nodes.102 This technique is free of chemical additives,
operated continuously and has the flexibility of adopting different types and sizes of cells by
adjusting the operating parameters such as acoustic energy density, contrast factor, and ultrasound
frequency. Lab-scale experiments have demonstrated a concentration factor of 11.6 can be
achieved at a flow rate of 25 mL/min.102 Other initial harvesting techniques includes air
flotation103,104, and electric field assisted harvesting95,96. As discussed, biofilm cultivation105,106 has
unique harvesting advantages but further development is required to make them cost and energy
effective for biofuel production.
3.2.1 Biomass thickening
After initial harvesting, thickening techniques can be applied to concentrate the algae suspension
to above 15% of TSS95,96,99. Two major thickening techniques are centrifugation and filtration.
Thickening saves a significant amount of energy compared to directly drying the harvested
biomass6,107,108 but requires extra capital investment. The energy cost for this process depends on
the characteristics of the cells, system design, and desired output concentration. Moreover, as the
31
desired output concentration increases, the energy cost associated with incremental percentage of
dry biomass climb steeply97,108. Centrifugation is commonly used in the lab and provides rapid
water removal but it is very energy intensive, thus not practical at industrial scale for bioenergy
applications, but is widely used to produce high-value products such as pigments, polyunsaturated
fatty acids (PUFAs), phycobiliproteins, enzymes and toxins96,109,110. Filtration is conceptually
simple but potentially difficult to operate due to the two major issues: 1) the pore size needs to be
small to increase efficiency but not too small to cause clogging issues and reduce flow rates; 2)
easy recovery of algal biomass from the filter is desired without using the backwash which will
lead to re-dilution of the product95,97,111. Advantages of alternative filtration techniques such as
dead-end filtration, tangential flow filtration, cross-flow membrane filtration, axial vibration
membrane filtration and ultrafiltration have been demonstrated for specific applications95,99,112–114.
3.2.2 Dehydration to dry biomass
Dehydration is used to achieve the final high level of biomass concentration so that conventional
extraction methods and infrastructure used for terrestrial plants can be used effectively. Typical
dehydration techniques include solar drying, spray drying, freeze drying and belt drying. Solar
drying either directly uses sunlight, or converts sunlight into heat for drying. This approach is
relatively easy to implement but it is hindered by the lack of control (overheating, weather
dependency), long drying time and possible biomass degradation100,115–118. Spray drying is
commonly used to dry microalgae for food production but it still requires a significant amount of
energy which make it impractical for biofuel production99,118. Freeze drying is used to produce
pharmaceuticals from microalgae to maintain the chemical. physical, and biological characteristics
of intracellular components119–121. However, this process is far too costly and energy intensive to
be applicable to less valuable chemical outputs. Belt drying transports biomass on a conveyor belt
and dehydrate the biomass either by passing through a heating chamber or by the heat conducted
32
from the belt. Other drying techniques such as rotary drying, cross-flow drying, vacuum shelf
drying and flash drying also been used in the industry but they haven’t reached commercial scale
for biofuel production due to high energy consumption118,122.
3.3 Processing for dry microalgae
Once microalgae biomass is completely dried through dehydration techniques, conventional
physical disruption followed by chemical extraction and conversion techniques can be directly
applied. Effective physical disruption can offset the need for harsh processing conditions required
in the process of chemical extraction93,110,118. Cell homogenization and bead milling are commonly
used in the industry, and emerging disruption techniques such as microwave, pulsed electric field
and ultrasonic are still under development101,118,123. Among these techniques, microwave-assisted
extraction (MAE) has attracted lots of attention due to its advantage of easy operation, high energy
transfer efficiency, rapid heating and relatively low cost. MAE has been investigated to be highly
effective for pigments124 and lipids125 extraction from microalgae. MAE of fucoxanthin from
Cylindrotheca closterium for 5 min enabled maximal extraction equivalent to 60 min of
conventional solvent extraction method.124 With the addition of ionic liquids, MAE has been
applied to extract lipids from wet microalgal biomass where extraction rates were increased by an
order of magnitude in most cases.126 However, the cost associated with these disruption techniques
is still too high for biofuel production. The requirements for physical disruption depend on the
characteristics of the microalgae and specific extraction techniques could be simplified, or even
eliminated, by genetic engineering or extraction process enhancement.
3.3.1 Lipids extraction
The most commonly used lipids extraction technique for dry microalgal biomass is organic solvent
extraction which is found to be highly effective and cost efficient101,123. Supercritical CO2
33
extraction is not suitable for biofuel production due to its high initial and maintenance cost, but its
application has increased in the nutraceutical and biochemical industry in recent years78,85,127,128.
The major advantage of using supercritical CO2 as solvent is to produce contamination-free
products and the associated cost can be offset by the high marginal value bioproducts. Other
extraction techniques such as accelerated solvent, two-phase solvents and switchable-solvent
extractions have been proposed and demonstrated to improve the extraction efficiency of
intracellular components from microalgae129–131.
3.3.2 Extracted lipids and other components
Lipids extracted from microalgae can be further converted to biodiesel through several pathways:
chemical transesterification, enzymatic conversion and catalytic upgrading. Among these,
chemical transesterification is relatively mature and has been used widely for biodiesel production
with conversion efficiencies above 90%132–134. However, algal oils are typically highly complex
in contrast with vegetable oils, containing fatty acids, phospholipids, carotenoids, chlorophyll and
other components in various composition97,135. Therefore, in order to accelerate this reaction,
obtain higher conversion rate and reach the full potential of commercialization, a full
understanding of strain specific oil composition is required. In contrast, enzymatic conversion uses
biological catalyst (lipases) instead of acids or bases to convert lipids into biodiesel with less
processing energy, easier removal of glycerol and catalysts and less alkaline wastewater pollution.
Although, enzymatic approaches have advantages over conventional methods, other challenges
associated with the cost of lipase, operational life, tolerance of the environment for enzymes and
harvesting strategies for products need to be addressed for this path to run at a commercial scale136–
138. Catalytic upgrading of algal lipids into renewable gasoline, jet fuel and diesel can also be
achieved by hydrotreating processes which react oils with hydrogen at high temperature and
pressure in the presence of catalysts139–141. This conversion process is commonly used in the
34
petroleum industry to upgrade crude oil to produce a wide multitude of performance specified
fuels. Ideally, this process leverages existing techniques and infrastructure albeit with catalysts
and process parameters tuned for algal feedstocks.
Recent research in algal downstream processing, has shifted from a lipid-centric approach to a
more holistic approach. If one can obtain value from all feedstock components the
commercialization of microalgae technology will be more economic viable. For example, residual
carbohydrates and proteins from microalgae can be used to enrich the nutrition in animal feed or
further processed to produce biofuels through other conversions such as fermentation, anaerobic
digestion and hydrothermal liquefaction. Another approach to achieve commercially feasibility in
microalgae industry is to produce high-value bioproducts.69,142–145. These bioactive components
present a great interest in pharmaceutical, cosmetic and nutraceutical industries and can easily
offset the high costs associated with biomass processing. The understanding of microalgae and
development of technology gained in this approach will accelerate the accomplishment of
producing sustainable energy from microalgae.
3.3.3 Processing of whole dry microalgae
Without extracting lipids, direct combustion and pyrolysis have been investigated to directly
produce energy from whole algae. Direct combustion of dry microalgae biomass releases the
largest amount of energy. However, multiple life cycle assessments5,6,8 indicate drying the
microalgae into powders costs more energy than the biomass contained, plus the challenges
associated with ash content and emission control make direct combustion of microalgae is
impractical on a commercial scale146–148. Moreover, the energy released from direct combustion is
in the form of heat which is less desirable than liquid biofuels used for transportation. Pyrolysis is
a thermal decomposition process in absence of oxygen at high temperature (300 °C – 1000 °C),
35
converting organic components into a wide range of products with hydrocarbon rich liquid (bio-
oil) as the desired product.149–151 Recent research152 indicated that fast or flash pyrolysis of
microalgae with reaction time of 2-3 s, reaction temperature around 500 °C with heating rate of
600 °C/s is capable of achieving 18 – 24 % liquid yields with higher heating value (HHV) of 29
MJ/kg. Pyrolysis bio-oil from microalgae has lower oxygen content and viscosity and higher
heating value (HHV) than that from woody biomass but still requires extensive refining for use in
conventional fuel engines. More importantly, similar to direct combustion, the major roadblock in
pyrolysis is removing the moisture content in the biomass which cause the overall process to be
energy negative.
3.4 Processing for wet microalgae biomass
Avoiding the energy and cost intensive thickening and drying processes dramatically alleviates the
energy burden for microalgae biofuel applications. Two promising conversion processes
applicable to wet biomass at a concentration above 15% TSS are hydrothermal liquefaction (HTL)
and supercritical water gasification (SCWG). A summry of extraction and conversion techniques
for different concentration of microalgal biomass is shown in Table 2-1.
36
Table 3-1 Summary of extraction and conversion techniques for different concentrations of
microalgae biomass.
Feedstock Type Conversion/Extraction
Techniques
Main Products Reference
Dry powder
Mechanical disruption and
chemical extraction
Antioxidants, pigments,
PUFA, additives 153,154
Direct Combustion Heat 155
Pyrolysis Bio-oil 152
Microalgae slurry
(> 15% TSS)
Hydrothermal Liquefaction Biocrude 7,65,156,157
Supercritical water
gasification Syngas 158–160
Microalgae in culture
(< 10% TSS)
Fermentation Bioethanol 161–167
Anaerobic digestion Methane 168–172
Direct secretion Hydrogen, alcohols or
alkanes 173–175
Microbial fuel cells Electricity 176–184
3.4.1 Hydrothermal liquefaction
HTL is one of the most promising conversion pathway for biofuel production due to the ability to
employ raw wet feedstock, a fast reaction rate, a high energy return on investment, good
characteristics of biocrude, a low production of char, and the potential for recycling of nutrients
(Fig 3-1a&b). Similar to natural formation of petroleum-based fossil fuels, HTL converts biomass
into biocrude at high temperature (200-380 °C) and pressure (5-28 MPa) in the presence of water,
albeit on a timescale of minutes by chemically and physically cracking down large biomolecules
into small fractions7,10,65,156,185. Compared to other thermochemical conversion techniques such as
pyrolysis, the higher heating value of HTL biocrude is about 35 MJ/kg, much higher than typical
pyrolysis bio-oil with a value of 20-25 MJ/kg186.
A fundamental challenge in HTL of biomass is optimizing the processing conditions (temperature,
pressure, residence time and heating rate) to obtain optimal yield and efficiency10,12,187. To this
37
end, HTL65,66,188 with a fast heating rate was investigated and resulted in higher biocrude
productivity. However, the heating rate in this experiment was limited at 230 °C/min which was
not able to explore the peak productivity with respect to heating rate. By eliminating the limits
from conventional batch reactors, microdevices156 that allow direct observation of HTL (Figure 3-
1a) have been developed to precisely control processing parameters and perform real-time
monitoring. The results indicate the higher heating values of biocrude approached saturation within
1 min due to early mechanical disruption of cells that enabled solvent extraction. Therefore, the
HHV alone does not present the quality of biocrude whereas chemical composition and physical
characteristics should also be considered.
A sequential HTL process189 was also performed to maximize the biocrude productivity and
minimize the bio-char formation. A low temperature (140 – 200 °C) HTL was used as the first step
to disrupt the cell and release intracellular products, polysaccharides in this case. The second step
utilize a higher temperature (220 – 300 °C) HTL that converts the remaining biomass to biocrude
which resulted in overall higher biocrude productivity and less bio-char. Lower polysaccharide
content resulting in decreased bio-char production also agrees with HTL of individual categories
of biomass feedstocks which indicates polysaccharides generates more solid products than protein
and lipids.12 Similar to HTL, Hydrothermal carbonization (HTC) uses reaction temperatures in the
lower range (180 – 250 °C) with slightly elevated pressure (2 – 10 MPa) which tends to produce
more solid products instead of liquid products190–192. A recent comparison of torrefaction and HTC
of lignocellulosic biomass indicates that HTC as the wet torrefaction method is the more favorable
process in terms of energy content, hydrophobicity, and inorganic components of the solid
products193.
38
Figure 3-1. Summary of current downstream processing techniques for wet microalgae biomass
(a-f) and cells in culture (g-j). (a-b) Hydrothermal liquefaction156,157. Image (b) has been
reproduced with permission from Elsevier157, Copyright 2013. (c) Supercritical water
gasification159. Reproduced with permission from Elsevier, Copyright 2016. (d-e) Ionic liquid
treatment for wet extraction194,195. Image (d) has been reproduced with permission from Royal
Society of Chemistry194, Copyright 2014. Image (e) has been reproduced with permission from
Royal Society of Chemistry196, Copyright 2015. (f) Astaxanthin extraction for hydrothermal
disrupted cells197. (g) Fermentation of pretreated wet biomass161. Image reproduced under CC-
BY license, Copyright 2014. (h-i) Anaerobic digestion of algal biomass169,170. Image (h) has been
reproduced with permission from Elsevier170, Copyright 2017. Image (i) has been reproduced
with permission from Elsevier169, Copyright 2016. (j) Microalgae-microbial fuel cells198.
Reproduced with permission from Elsevier, Copyright 2015. This figure is reproduced from ref91
with permission from The Royal Society of Chemistry.
39
3.4.2 Supercritical water gasification
Analogous to HTL, SCWG reacts at higher temperature (400 - 700 °C) and pressure (> 22 MPa)
to decompose biomolecules to produce syngas containing hydrogen, carbon monoxide and
methane with a small quantity of solid and liquid products158–160,199(Figure 3-1c). Gasification is
commonly combined with Fischer-Tropsch Synthesis (FTS) to convert syngas into liquid fuels200–
202. The major advantage of this pathway is the flexibility to produce a wide variety of fuels and
products with known properties. Recent research indicates the issues for developing this process
are: precipitation of inorganic salts, an unclear reaction mechanism associated with reaction
temperature, pressure, heating rate and wall effects, the requirement of effective catalyst and high-
temperature-resistant materials, and high energy costs150,192,199. Conventional gasification (in an
environment of insufficient oxidizer) of microalgae in a temperature range of 800 – 1000 °C was
also studied and due to the requirement of dry biomass, this pathway for biofuel production usually
resulted in negative net energy100,148,203.
Another interesting application of subcritical water is to lyse cells for extraction of high-value
intracellular products197,204,205. A particular challenge is breaking robust cell walls to allow
extraction, particularity with wet biomass197. Lab scale tests have shown that hydrothermal (Figure
3-1f) and ionic liquids (ILs) (Figure 3-1d&e) can achieve comparable efficiencies as mechanical
processing to dry biomass. Hydrothermal disruption of the cell wall at 200 °C effectively enabled
solvent extraction of astaxanthin to achieve more than 95% efficiency as opposed to control with
a 7.5% in extraction efficiency.197 ILs have been used as extracting agents to replace organic
solvents or pre-treatment material to disrupt the cell wall to enhance extraction. An extraction
efficiency of 82% was achieved using 1-Ethyl-3-methylimidazolium ethylsulfate, but a
germination process of H. pluvialis cysts is required to weaken the cell wall.194 ILs are also applied
40
in a pre-treatment process to achieve > 70% extraction efficiency, but the experiment utilized dry
cell powders as feedstock which normally has weaker cell walls and the reusability of ILs needs
improvement for large scale production.196
There are also other approaches to directly extract intracellular lipids from wet biomass94,206–208,
but due to either high cost or low extraction efficiency none of them have been commercialized
yet. For biofuel productions, since the current wholesale price for diesel is down to about 70 cents
per liter, bringing additional steps or expensive additives into the process is most likely a non-
started.
3.5 Processing for microalgae in culture
To minimize the energy used in removing water from biomass, biologic conversion processes
involving fermentation, anaerobic digestion, and direct secretion have been studied. In these
pathways, biofuels can be either produced directly from microalgae culture or after initial
harvesting with minimal energy cost. Lastly this section includes microalgae-microbial fuel cell
approaches whereby microalgae are employed for direct electricity production.
3.5.1 Fermentation
Fermentation is a well-established method used in alcohol production to convert carbohydrates
into ethanol by yeast. There are two major approaches to produce ethanol from microalgae: 1) like
yeast, some microalgae such as Chlorella and Chlamydomonas can produce alcohols through
heterotrophic fermentation162,163,209,210; 2) microalgae containing a significant amount of
carbohydrates can be used as a sugar source for yeast fermentation164,165,167. In the first approach,
sugars can either be generated internally from the synthesis of microalgae or fed to algae
externally, however the process lacks the productivity of yeast fermentation. The second approach
usually requires pretreatment to promote hydrolysis of the cell wall to: 1) access to intracellular
41
components; and 2) release fermentable sugars. Although research203 indicated the sugar released
from wet microalgal biomass was lower than that from dried biomass, drying the biomass prior to
conversion does not provide an energy return on investment5,8,148. Recent research161 (Figure 3-
1g) demonstrated that acid-catalyzed pretreatment prior to lipids extraction released more than
90% of fermentable sugars for wet microalgae biomass. Although noticeable achievements have
been made in this pathway, significant breakthroughs are required to produce biofuels
economically at industry scale with this approach.
3.5.2 Anaerobic digestion
Anaerobic digestion used in wastewater treatment to produce methane has the potential to exploit
the entire organic carbon content of microalgae without the requirement for drying (Figure 3-1
h&i). It can also significantly benefit from the direct use of existing infrastructure and experience
in the field of wastewater treatment168–170,211,212. Methane as the main product in the process not
only can be used as fuel but can also be converted to bioplastics to increase the value171,172,213.
However, this pathway is hindered by the low practical methane yields mainly due to the resistance
of microalgal cell walls. Rigid cell walls of microalgae not only reduce the amount of digestible
substrate but also limit the access of microorganisms to intracellular components resulting in low
reaction rate211,212,214,215. Thermal, chemical and mechanical pretreatments of microalgae were
used to improve methane yields but the energy cost involved in the extra steps is higher than the
energy gain from increased methane production168,212,216. By tuning the pH and retention time,
providing heat treatment and addition of methanogen inhibitors, this process can be altered to
produce hydrogen210,217 instead of methane but it requires significant development to be
commercially available. The more likely role for anaerobic digestion is in combination with other
methods to fully harvest the remaining value of biomass and therefore maximize the biofuel
production.
42
3.5.3 Direct secretion
Direct secretion of products such as hydrogen218,219, alcohols174,220,221 and alkanes173,175,222 from
microalgae culture can be achieved through genetic engineering. Hydrogen production from
Scenedesmus obliquus was first observed by Gaffron and Rubin223 and having an anaerobic
environment was later found to be critical for this process.224 Sulfur deprivation was used to inhibit
the activity of PSII and enhance the activity of hydrogenase enzymes for hydrogen production.
Breakthroughs in genetic engineering processing parameter control on sulfur quantity and
immobilization of cells are expected to increase hydrogen productivity by further inhibiting PSII
activity and increasing the activity of hydrogenase enzymes. Secreted hydrogen can be easily
collected, however the low conversion efficiency remains a major roadblock for this pathway.
Direct secretion of alcohols and alkanes appears to be a promising alternative but most information
in this area is proprietary. Secretion of triterpene from Botryococcus braunii was reported to reach
a volumetric productivity of 22.5 mg/L/photo-h at a high cell concentration of 20 gDW/L.102 These
pathways usually require cheap source of CO2 or sugars as feedstock and development to boost
the feedstock-to-product conversion rates.
3.5.4 Microalgae-microbial fuel cells
Microalgae-microbial fuel cells biologically convert solar energy to electrical energy and while
the technology is maturing, the development of this conversion strategy is early
stage177,178,180,181,198. Microbes at the anode generate electrons, protons and CO2 by digestion of
organics and the microalgae at the cathode takes CO2, light, proton and electrons to grow through
photosynthesis (Figure 3-1j). Also known as biological photovoltaics, or BPVs, this approach has
seen many recent advances. Microfluidic approaches were applied to perform a qualitative
investigation of several key factors including cell density, electron mediator concentration and
light intensity and indicated the major obstacle for power production from BPVs is the transport
43
of reducing equivalents across the cytoplasmic membrane.176,182 Understanding of electron
transport mechanism is also believed to be essential for selection of photosynthetic microbes to
enhance electrical output.179,184 Furthermore, the overall performance of BPVs is strongly
associated with surface morphology and corresponding material characteristics225, manipulation
of thylakoid terminal oxidases226,227 and types of feedstocks228. One of the challenges with this
approach is the need for electrodes which tend to block light paths in a way similar to the CO2
delivery mechanisms discussed earlier. A proof-of-concept cell was developed whereby both light
and electrons were delivered via a metallic/plasmonic surface183 , however, that approach is not
well suited to large scale production for a number of reasons. In general, there are many challenges
with microalgal-microbial fuel cell approaches including technical obstacles, high operation costs
and low power output that need to be solved for this process to be economically feasible. Similar
to some other approaches above, this strategy can be more readily adopted in a wastewater
treatment context, where the primary objective is remediation and power production is a side
benefit. We note however, that this research area is adapting quickly and may well produce
surprises 229.
44
Chapter 4.
Hydrothermal Liquefaction of Microalgae on
a Chip for Biocrude Production
Hydrothermal liquefaction uses high temperatures and pressures to break organic compounds into
smaller fractions, and is considered the most promising method to convert wet microalgae
feedstock to biofuel. Although, hydrothermal liquefaction of microalgae has received much
attention, the specific roles of temperature, pressure, heating rate and reaction time remain unclear.
A microfluidic screening platform to precisely control and observe reaction conditions at high
temperature and pressure. In-situ observation using fluorescence enables direct, real-time
monitoring of this process. A strong shift in the fluorescence signature from the algal slurry at 675
nm (chlorophyll peak) to a post-HTL stream at 510 nm is observed for reaction temperatures at
260°C, 280°C, 300°C and 320°C (P = 12 MPa), and occurs over a timescale on the order of 10
min. Biocrude formation and separation from the aqueous phase into immiscible droplets is
directly observed and occurs over the same timescale. The higher heating values for the sample
are observed to increase over shorter timescales on the order of minutes. After only 1 minute at
300°C, the higher heating value increases from an initial value of 21.97 MJ/kg to 33.63 MJ/kg.
The microfluidic platform provides unprecedented control and insight into this otherwise opaque
process, with resolution that will guide the design of large scale reactors and processes.
This chapter was published as technical innovation in Lab on a Chip and reproduced from156.
Copyright © 2016, Rights Managed by Royal Society of Chemistry. This work was featured in
Lab on a Chip as HOT article. The candidate was the first author in this work and played the
primary role in designing the research, performing the experiments, analyzing the data, and writing
the paper. Additional authors for the work include Dr. Matthew D. Ooms, and Prof. David Sinton.
Their contributions were central to the publication of this work and are gratefully acknowledged
and appreciated.
45
4.1 Introduction
Microalgal biomass is an attractive feedstock for the generation of carbon neutral biofuels
because of the high growth rate and lipid content of many microalgae species.32 Conversion
of the raw algal biomass into biocrude however, remains an energy intensive6 and costly
process230 which is in part why algal biofuels have not yet achieved commercial success.
To produce biodiesel from microalgae using conventional methods, the fatty components
of the cells (lipids), must first be extracted and then converted to biodiesel via
transesterification. Traditionally this is accomplished through mechanical pressing or
solvent extraction. A challenge for both these approaches is that the biomass must first be
dried, which significantly increases the carbon intensity of the final fuel as well as its cost.8
From a life-cycle assessment perspective, drying of the biomass can consume more than
90% of the energy content of the final algal oil.7 Consequently, the potential benefit of
microalgal biofuels is severely undermined by the high energy and financial cost associated
with biomass-to-biofuel conversion.
An emerging conversion approach that does not require pre-drying is hydrothermal
liquefaction (HTL).11,46,48 HTL uses high temperatures and pressures to break organic
compounds into smaller fractions to produce biocrude which can be further upgraded into
a variety of fuels. Optimizing the many parameters involved in HTL processing, however,
remains a challenge. To date, HTL experiments have been conducted in laboratory scale
batch reactors using sand-baths, ovens or heating coils for temperature control. These
approaches have two critical issues. First, due to the large fluid volumes and large physical
size of the apparatus, substantial heating times are required, which are often ignored in the
subsequent analyses. These long heating times lead to ambiguity in the reported optimal
values for temperature, heating rate, and reaction time since the heating and pressurization
delays blur the results. Second, in-situ monitoring of the reaction process is not possible in
current reactors, precluding real-time quantification of the reaction process.
Microfluidic and lab-on-a-chip methods have recently been applied to bioenergy
generation16–18, particularly with respect to microalgae. The cellular scale of microalgae
makes them well suited to manipulation and analysis using microfluidic platforms.183,231
Ensuring cells receive both light and fluids is an optofluidic challenge that has been
addressed, for instance, with integrated waveguides to deliver light to cultures232–234 and
micro-reactor arrays integrated onto individual LCD displays for parallelized illumination
46
and growth studies.235 These efforts have focused primarily on cultivation of microalgae,
but have not addressed the downstream challenge of converting biomass into useful
products.
In this work, we demonstrate a high temperature and high pressure, continuous flow,
microfluidic reactor to perform controlled HTL on a glass and silicon chip (Fig. 1a). The
small length-scales of the microfluidic chip enable effectively immediate heating of the
algal slurry eliminating the ambiguity associated with conventional reactors. This work
leverages established advantages of using high-pressure high-temperature silicon-glass
microfluidic reactors10,21,86,88,236,237, for bioenergy applications. Our microfluidic chip
makes possible real-time in-situ observation including fluorescence imaging and analysis
as shown in Fig. 1. The fluorescence signatures from both chlorophyll in algae238 and
aromatics in the produced oil239 provide direct indicators of chemical composition during
the reaction.
Figure 4-1. a) Schematic of hydrothermal liquefaction of microalgae in the microfluidic chip
with in-situ observation of biocrude production using fluorescence microscopy. b) Distinct
fluorescence signatures of algae slurry at the inlet and biocrude at the outlet. Reproduced by
permission of The Royal Society of Chemistry.
47
4.2 Experimental
The chip was fabricated out of glass and silicon because common chip materials such as
polydimethylsiloxane (PDMS) and polymethyacrylate (PMMA) cannot support the high
temperatures and pressures used in HTL and metal-based chips do not permit in-situ observation.
The combination of silicon and glass allowed for (i) optical access through the glass, (ii) high
thermal conductivity through the silicon, and (iii) both high pressure and temperature (over 10
MPa and 300°C respectively).
The square cross-section channel dimensions were 200 μm x 200 μm with a total length of 1320
mm, 1250-mm of which were located in a serpentine heating region. Raw algae slurry was
continuously pumped into the chip at a flow rate of 5 μL/min at 12 MPa. At this rate, the slurry
required 10 min to flow through the length of the heated region. Under steady state flow, the
reaction time (time spent in the heated region) at a given location along the channel could be
calculated. The heating rate of 15 °C/s, the fastest reported heating rate for HTL experiments in
literature, was calculated based on the 35 mm length of channel between the inlet at 50 °C and the
beginning of the heating region at 300 °C. The other pump was running at constant pressure to
provide a reference pressure for the back pressure regulator (Equilibar Inc.). The ultra-low flow,
back pressure regulator was used at the outlet to maintain steady flow at a constant pressure. It was
essential to minimize dead volume downstream of the chip in order to collect representative
samples for off-chip product analysis. Here, a separate fixed-volume sample collector loop
(300μL) was incorporated into the outlet stream, immediately downstream of the chip and
upstream of the backpressure regulator. The outlet line was switched using a 6-port valve with
minimal dead volume (Rheodyne®7030). When isolated, the contents of the collector loop could
be dispensed into a collection vial without depressurizing the entire chip. During the experiment,
the switching valve was in collection position to ensure fluids exiting the microfluidic reactor
quickly entered the fixed-volume sample isolator. Once the sample isolator is filled with
representative samples collected at steady state, the switching valve is quickly turned to the eluting
position. Then a large amount of DI water is pumped in from the syringe to eject the sample into
the sample vial. This enabled inline sampling directly from the output stream. This procedure
ensures sample quality while maintaining high pressure in the chip during collection.
48
The heated region of the chip was mounted in a temperature controlled stainless steel heating
chuck, as shown in Figure 4-2. Precise temperature control was accomplished through a
proportional-integral-derivative temperature controller (Omega CNI3222) with three cartridge
heaters (Omega CSH-102135/120) maintaining a constant temperature over the entire heating
region of the chip. A thermocouple inserted in the heating chuck provided closed-loop feedback
and kept the temperature variation to less than +/- 1°C during steady state operation at a reaction
temperature of 300°C. The heating chuck was also equipped with a borosilicate glass viewing
window which enabled in-situ observation of the HTL reactions that occurred in the channel. The
chip manifold was maintained at a lower temperature (~ 50°C) to (i) prevent O-ring material failure
and leakage at the ports21 and (ii) to allow rapid and controlled on-chip heating as the fluid
transitioned from the cold to the hot zone (Figure 4-4).
Figure 4-2. Schematic representation of the assembly of the water-cooled manifold,
temperature controlled heating chuck and the microfluidic chip using a separation glass and
double seal O-ring to prevent cracking from hard contact. Reproduced by permission of The
Royal Society of Chemistry.
The biomass injected into the chip consisted of Nannochloropsis oculata with an ash content of
5.9% (Reed Mariculture Inc.). The as-received biomass was cleaned through centrifugation and
suspended in DI water prior to use. Algal slurry at a concentration of 2 wt% was then injected into
the chip using a high pressure pump with constant flow rate. This concentration was chosen in
49
order to minimize clogging and fouling issues while still providing enough material to allow real-
time observation of the chemical reactions on-chip and for subsequent off-chip analysis.
During operation, the chip was monitored with a fluorescence microscope for changes in the
fluorescence signature of the algal slurry. The sample was excited with ultraviolet light (λ = 375-
400 nm) and fluorescence detected through a long-pass filter (λ > 405 nm) using a
spectrofluorometer (ocean optics USB2000) attached to the microscope.
Produced samples were collected off-chip for analysis of their higher heating values (HHV) by
isolating the reaction products in the sample collector and eluting them to small glass vials. The
product at the outlet had a variety of components including water-soluble compounds and
biocrude, with other smaller amounts of solid particulates and gas. To isolate the biocrude, 2 mL
of dichloromethane was added to the recovery vial followed by vigorous shaking for several
minutes, completely dissolving the biocrude into the dichloromethane. The vials were then set
aside to allow complete phase separation. Once separated the dichloromethane layer was
withdrawn using a glass syringe and stored in a separate vial. This extraction process was
performed multiple times to ensure more than 95% of biocrude was recovered from the sample.
The extracted biocrude was then heated in an oven at 40°C for 8 hours to remove the solvent. The
higher heating values of each sample were calculated using the modified Dulong’s formula as
follow:
𝐻𝐻𝑉 (𝑀𝐽/𝑘𝑔) = 0.335𝐶 + 1.423𝐻 − 0.154𝑂 − 0.145𝑁 (4 − 1)
A carbon-hydrogen-nitrogen elemental analyser was used to measure the carbon (C), hydrogen
(H), nitrogen (N) composition of the initial dry algae sample and the produced biocrude. The
oxygen (O) content was estimated according to:
%O = 100% − %C − %H − %N − %𝐴𝑠ℎ (4 − 2)
where the sulphur content is assumed to be negligible, as is typical for microalgae.57,240 Prior to
elemental analysis, the raw algae was dried in an oven at 105 °C for an hour.
50
4.3 Results
Algae slurry was continuously pumped through the chip for 15 min to achieve steady state. By
observing the fluorescence signature at different points along the channel (Figure 4-3a), the
progression of the HTL process as a function of time spent in the reactions chamber was quantified.
Here, a series of viewing points along the channel were chosen to correspond to reaction times at
0.4-min intervals, ranging from 0 to 10 min. Near the inlet, raw algal slurry mainly showed
chlorophyll fluorescence with a peak at 675 nm (Figure 4-1b). By 1.2 min, the chlorophyll peak
had significantly dropped and an emerging peak at 510 nm began to rise. Over the next 2 min, the
510 nm peak became dominant and continued to grow. After 10 min, the original chlorophyll peak
was no longer visible and the normalized peak intensity at 510 nm approached a saturation point
as shown in Fig. 3b. The evolution of the peak at 510 nm indicated the formation of aromatic
compounds which are a characteristic component of crude oils as well as other processed plant
based oils. The progression of the fluorescence signature from one dominated by chlorophyll
fluorescence to one resembling conventional crude oils tracked the progression of the HTL
conversion process.
51
Figure 4-3. a) Normalized fluorescence intensity of algae slurry observed at viewing points
along the channel under 320°C indicating the formation of biocrude over time. b) The
progression of normalized fluorescence intensity of the 510nm peak at the reaction
temperature of 260°C, 280°C, 300°C and 320°C indicating higher reaction temperature has
higher reaction rate. Solid lines included as a guide for the eye. Reproduced by permission
of The Royal Society of Chemistry.
The effect of reaction temperature was investigated by performing identical experiments at 260°C,
280°C, 300°C and 320°C. For each temperature, the normalized peak intensity at 510 nm over the
course of the reaction is shown in Figure 4-3b. The fluorescence signals were normalized to the
52
projected saturation intensity based on a 1st order exponential curve fit to the experimental data.
The results in Figure 4-3b clearly show that higher reaction temperatures resulted in higher
reaction rates. Specifically, the characteristic times (time required for the fluorescence intensity at
510 nm to reach 63% of its maximum value) were: 6.0 min, 4.6 min, 3.4 min, and 1.9 min for the
reactions run at 260 °C, 280 °C, 300 °C, and 320 °C respectively.
The measured HHVs of dry algae and biocrude are shown in Table 4-1 and correspond well with
previously reported values12,65,241 using similar microalgae species and reaction conditions. As
shown, the most significant increase in HHV occurred within the first few minutes of the reactions.
Beyond reaction times of 1 minute, the variation of HHV was less than 5%.
Table 4-1: Elemental composition and Higher Heating Value of dry algae and biocrude
from 1 min, 5 min and 10 min reaction times.
Element Dry Algae Biocrude (1 min RT)
Biocrude (5 min RT)
Biocrude (10 min RT)
C [%] 51.08 70.11 70.35 70.63
H [%] 7.23 8.75 9.23 9.22
N [%] 8.65 4.46 6.00 6.56
O [%] 27.14 10.78 8.52 7.69
HHV [MJ/kg] 21.97 33.63 34.52 34.65
Combined with the fluorescence data collected, this analysis of HHV suggests that while the
energy content of the biocrude approached saturation at very early times (~ 1 min), other reactions
continued to occur that were not accounted for by the HHV alone. The sharp increase in HHV
during the first minute of the reaction was likely due to rupturing of the cells which made lipid
extraction by dichloromethane more efficient.56 As such, the increase in HHV should be used as
an indicator of mechanical disruption rather than chemical conversion which can be observed more
directly with the fluorescence signal.
Direct observation of the channel during operation provided additional insight into this process.
Most notably, non-fluorescent droplets were observed forming and adhering along the length of
the channel and increased in size towards the outlet (Figure 4-4a). These droplets are expected to
53
be comprised of aliphatic compounds which are immiscible with water and do not fluoresce. The
formation of these droplets progressed along the length of the channel which suggests a correlation
between the formation of immiscible oil droplets and the parallel change in fluorescence signature
resulting from the formation of aromatic compounds in the aqueous phase. Additionally, the
degradation of the microalgae could be observed between the inlet where whole cells were clearly
visible and the outlet where individual cells were no longer discernable (Figure 4-4b).
Figure 4-4. a) Fluorescence images obtained at viewing points along the channel with
increase in reaction time indicating the progression of biocrude formation. b) Microscopic
observation of fluids at the inlet and outlet via both fluorescence and dark-field imaging.
Scale bars: 50 μm. Reproduced by permission of The Royal Society of Chemistry.
54
It was also observed that a significant amount of cell debris and solid particulates (and in some
cases, clogging) resulted from operating with short reaction times. Specifically, we observed that
at 320oC when the flow rate was increased such that the maximum reaction time was only two
minutes, a significant amount of solid debris remained in the effluent. In contrast, less solid debris
in the output was observed for longer reaction times achieved with lower flow rates and otherwise
similar conditions. This finding is likely a result of more complete disruption of the biomass during
the first few minutes of the reaction (also indicated by the increase in HHV described earlier).
Furthermore, clogging at the outlet was exacerbated by the rapid cooling of the effluent which
promoted the separation of the oil phase from the aqueous phase, which was directly observed as
an increase in the number and size of oil droplets in the cooled outlet line (Figure 4-4a). Lastly,
the multiphase nature of the generated products, visible as channel-adhered droplets in Fig. 4, will
influence to some extent the residence time, accumulation and ultimate production of different
components. The HHV analysis here, however, is largely unaffected by this issue as the oil is
separated from the produced fluid and based on the relative elemental composition of the biocrude.
These observations, which were made possible only through the direct visualization afforded by
our chip design, have implications for the optimum processing parameters of continuous flow HTL
reactors. Specifically, these findings indicate optimal reaction times between 2 and 10 minutes to
both maximize the conversion of biomass to biocrude, and minimize the amount of debris in the
effluent to prevent fouling. Gradual cooling of outlet stream is also recommended to avoid
clogging.
4.4 Conclusion
In summary, our microfluidic reactor provides unprecedented insight and control over the high
temperature and high pressure cracking of biomass via hydrothermal liquefaction. It allows for in-
situ observation of hydrothermal liquefaction reactions using fluorescence microscopy and
convenient and precise control of reaction temperature, pressure and reaction time in a continuous
flow reactor. These advantages enable the study of high temperature and pressure cracking of
biomass on a platform with a high degree of control which will allow improved understanding of
the reactions taking place during hydrothermal liquefaction. The significant change of
fluorescence signature between the algal slurry (peak at 675 nm) and converted biomass (peak at
510 nm) was observed as an indicator of the progression of hydrothermal liquefaction. Biocrude
formation and separation from the aqueous phase into immiscible droplets was directly observed
55
and occurred over timescales of ~10 min. The rapid increase of higher heating values was observed
over the timescales of ~1 min and was correlated to observations of particulate matter in the
effluent which manifested as partially clogged channels. These results and the microfluidic
platform on which they were collected represent the first of their kind in the field of hydrothermal
liquefaction research. Lab-on-a-chip methods offer a unique toolset to probe high temperature and
high pressure reaction dynamics and inform large scale reactor design.
4.5 Supplementary Information
4.5.1 Fabrication of microfluidic chip
To achieve both high temperature and pressure on a chip, the high pressure connection between
the chip and the tubing needed to remain at low temperature (Figure 4-5a). A window cut in silicon
was used to reduce heat transfer from high temperature area to high pressure compression ports
and therefore a large temperature gradient was generated along the arms. The chip fabrication
followed the process indicated in Figure 4-5 b. The features on the chip was generated on the
silicon and bonded to glass at the end. The whole fabrication process included two sets of
spincoating, patterning, and Deep reactive-ion etching (DRIE) and therefore two types of
photoresists (S1818 and AZ4620) and two photomasks were used. The silicon wafer was 4” in
diameter,1mm thick, and double side polished. The channel was patterned by photomask #1 and
generated by DRIE for a depth of 200um. The holes and the window cut in silicon were generated
by etching through using DRIE. The pattern on the other side of the channel (called back side in
the figure) was aligned using Back Side Alignment technique with designed markers on both
photomasks. Once the features were completed on the silicon wafer, both silicon and glass were
thoroughly cleaned using Piranha solution before anodic bonding.
56
Figure 4-5. a) Schematic illustration of achieving high temperature and pressure on a chip
by separating high pressure compression from high temperature area. b) Si/glass chip
fabrication process (from top to bottom, left to right).
4.5.2 Experimental apparatus
The experimental apparatus contains a number of components and the flow path among them is
indicated in the Figure 4-6. All critical components in the apparatus have pressure rating higher
than 20 MPa and are mainly connected using Yor-Lok fittings. Two high pressure pumps (ISCO
260D) were used in this experiment. One was running at constant flow rate to continuously pump
algae slurry in the piston cylinder to the chip for a desired reaction time. The manufacturing
numbers and the application purposes of major components are provided in Table 4-2.
57
Figure 4-6. Schematic diagram of experimental setup with flow direction indicated by arrows
along the processing path. The flow path of the switching valve at two positions is shown in
green lines.
Table 4-2: List of components in the apparatus and their purpose.
Number Part Name Mfg Number Purpose
1 High Pressure
Pump
ISCO 260D 1a. Pumping algae slurry at constant flow rate
1b. Maintaining constant pressure for BPR
2 Piston cylinder HIP TOC3-10 Algae slurry container
3 Switching valve Rheodyne®7030 Switching flow paths without depressurizing the
whole system
4 Fixed-volume
sample isolator
Radel® R Tubing
1220
Temporarily storing fixed-volume sample in the
loop ensures sample quality
5 Back pressure
regulator
Equilibar
EB1ULF1 - SS316
Maintaining constant pressure for a continuous
flow at ultra-low flow rate
4.5.3 High temperature and pressure packaging
The manifold module is used to provide high pressure sealing between the microfluidic chip and
the rest of the apparatus. This manifold uses a modular design (Figure 4-7) and can interface with
any chip sharing the same port pattern and thickness. Compression between the O-ring (Double-
Seal Viton® 004) and the chip is achieved through screw fasteners and care must be taken when
58
tightening to avoid fracturing the chip. To assist with tightening, a spacer is placed between the
clamp and the manifold to prevent fracture by overstressing. Also, a layer of polished glass is
placed between the chip and the clamp to provide even clamping pressure. The manifold was
fabricated out of stainless steel (SS316).
Figure 4-7. Detailed drawing of the compression sealing of the microfluidic chip (MF) with
algae slurry fluid indicated in green. O-rings and spacers are used to ensure a quality seal
between the manifold and the chip and to prevent overtightening and damage.
59
Chapter 5.
Hydrothermal Disruption of Algae Cells for
Astaxanthin Extraction
We demonstrate a hydrothermal method of astaxanthin extraction from wet biomass using a high
temperature and high pressure microfluidic platform. Haematococcus pluvialis cysts are trapped
within the device and visualized in-situ during the cell wall disruption and astaxanthin extraction
processes. The device provides a highly controlled environment and enables direct comparison of
chemical vs. hydrothermal processes at the cellular level. Hydrothermal disruption at a temperature
of 200 °C was shown to be highly effective, resulting in near-complete astaxanthin extraction from
wet biomass - a significant improvement over traditional methods.
This chapter was published as communication in Green Chemistry and reproduced from [ref].
Copyright © 2017, Rights Managed by Royal Society of Chemistry. The candidate was the first
author in this work and played the primary role in designing the research, performing the
experiments, analyzing the data, and writing the paper. Additional authors for the work include
Dr. Jason Riordon, Mr. Brian Nguyen, Dr. Matthew D. Ooms and Prof. David Sinton. Their
contributions were central to the publication of this work and are gratefully acknowledged and
appreciated.
5.1 Introduction
Astaxanthin is a highly valuable microalgal bioproduct with uses ranging from human health to
aquaculture. While synthetic production is currently favoured in industry due to lower production
costs, natural astaxanthin is considered more beneficial than synthetic astaxanthin due to its
superior antioxidant activity.69,70,242 The microalgae Haematococcus pluvialis is by far the richest
natural source of astaxanthin243 and has become the primary source for the nutraceutical industry.72
However, extraction of astaxanthin remains a challenge; the stress conditions that induce
astaxanthin accumulation in H. pluvialis cells also induce cell wall thickening. This barrier is
extremely robust to chemical and physical disruption,76 making astaxanthin extraction difficult,
particularly for mature cysts which are larger and have thicker cell walls compared to cells at other
60
life-cycle stages. Effective cell wall disruption increases the effectiveness of post-disruption
recovery approaches, which include organic solvents75, ionic liquids,194,196 and supercritical
CO2153,244. Chemical disruption using acids or bases typically results in low (<40 %) extraction
efficiencies.75 Mechanical disruption is therefore the primary method used in industry, but requires
dry biomass.72 Biomass drying in combination with mechanical disruption enhances the extraction
efficiency (>80 %) but also significantly increases the energy and financial costs of
processing245,246.
Hydrothermal processes – where wet biomass is subject to high pressures (5 – 20 MPa) and
temperatures (150 – 350 ºC) – have shown promise for several applications, including pressurized
hot water extraction of bioactives247,248, woody biomass decomposition249 and biocrude
formation57,187,250,251. Hydrothermal processes leverage the physiochemical characteristics of water
at elevated temperature and pressure, including: (i) a significantly higher fraction of water ions
which aids acid or base-catalysed reactions to break cell walls; (ii) a lower dielectric constant,
which enhances the hydrothermal conversion of carbohydrate biomass252; and (iii) energy savings
by avoiding the need for water evaporation.10 Importantly, hydrothermal disruption represents an
environmentally friendly, chemical-free approach, in sharp contrast to traditional chemical
methods.253 However, despite the strong potential of hydrothermal processes for cell wall-
disruption, such an approach has not been applied to astaxanthin extraction.
Current approaches to studying cell disruption and extraction processes of this nature generally
involve relatively large opaque batch reactors.75,254 Significant chemical and thermal gradients are
inherent in such reactors given their size, and pressure vessel requirements generally preclude
direct observation of the process86,88,255. Operating at smaller length scales, as with silicon-glass
microfluidic reactors, can minimize gradients, improve control, and provide access to high
temperatures and pressures, all while allowing direct optical access to the process.21,156 While such
microfluidic systems are themselves impractical for the production of bulk product, they are
ideally suited to screening conditions and quantifying unit process efficiencies – valuable
information for commercial-scale processing.
61
Figure 5-1. Simplified schematic of on-chip astaxanthin extraction from H. pluvialis.
Enlarged schematics of the cell capture area show initial cell trapping, cell wall disruption
and astaxanthin extraction. Reproduced by permission of The Royal Society of Chemistry.
In this work, we demonstrate a hydrothermal approach to cell wall disruption for astaxanthin
extraction from wet biomass, where extraction is visualized on a microfluidic screening platform.
Extraction was performed using a microfluidic device to provide real-time visualization of the
disruption and extraction processes. The glass/silicon microfluidic chip was designed to allow
optical access, while providing uniform heating and both chemical and thermal resiliency. The
device features parallel reaction channels with trapping posts, where H. pluvialis cysts are trapped
and monitored during wall disruption and astaxanthin extraction (Figure 5-1). Direct visualization
of the process enables quantitative comparison of chemical and hydrothermal methods at the cell
level. It was found that biomass treated hydrothermally, at a temperature of 200 ºC and a pressure
of 6 MPa, demonstrated near-complete astaxanthin extraction efficiency – a significant
improvement over traditional approaches.
62
5.2 Experimental setup
5.2.1 Device design and fabrication
The silicon-glass microfluidic device featured 40 parallel reaction channels, each 200 µm wide
and 100 µm tall. Each of these channels was lined with cylindrical posts of 20 µm diameter
separated by a distance of 40 µm. These posts served to trap cells while allowing fluid flow. The
device was fabricated by deep reactive ion etching (DRIE) of the silicon, with subsequent bonding
to borosilicate glass of 1.75 mm thickness. The full fabrication procedure for devices of this type
is reported elsewhere.156
5.2.2 Cell culture and trapping
Haematococcus pluvialis cultures were obtained from Algae Analytics (Las Cruces, New Mexico)
and cultured at 25 ºC under continuous white light illumination at ~30 µmol m−2s−1 in media
according to the recipe of Fábregas et al.256. Encystment was induced by exposure to red and blue
light at ~60 µmol m−2s−1 until the cultures developed red coloration indicative of astaxanthin
accumulation (irradiation spectrum shown in Figure 5-7). For each experiment, a suspension of H.
pluvialis cysts in media was injected into the chip using a pump and trapped in the post arrays (a
video of initial cell trapping is presented in ESI† - Video S1). Cysts with thickened cell walls were
selected by using a 70 µm cell strainer. After the cells were loaded, deionised water was flowed
into the chip at a rate of 1 mL min-1 for 5 min to flush the media prior to cell disruption. During
cell loading, and at all stages of astaxanthin recovery, the cells were observed using darkfield
microscopy (Olympus BXFM microscope, 10x objective).
5.2.3 Cell wall disruption and extraction
The cell wall disruption method was varied to test both chemical treatments (HCl 0.1M, NaOH
0.1M) and hydrothermal treatments (30 min @ 150 ºC, 30 min @ 200 ºC, 10 min @ 200 ºC and 5
min @ 200 ºC), and compared to a control case (no treatment). For chemical disruption, solutions
were pumped into the device at a flow rate of 1 mL min-1 for 1 min to rapidly fill the volume of
the chip before stopping the flow for 30 mins. The hydrothermal processes were performed at a
pressure of 6 MPa, chosen to be sufficiently high to maintain water at liquid phase at the selected
temperatures. The device was then heated to the desired temperature at a heating rate of ~60 ºC
min-1 using a custom heating chuck and maintained at the set temperature for 5-30 min by a PID
63
temperature controller. Once the hydrothermal treatment was completed, the heating chuck was
removed to allow the device to cool to room temperature.
The temperature of 200 °C was chosen to correspond to the approximate temperature where
structural cell wall polysaccharides rapidly depolymerize without degradation of other biomass as
a result of secondary reactions.13,257 After cell wall disruption, acetone was flowed at a rate of 1
mL min-1 for 1 min, before being reduced to a constant rate of 100 µL min-1 for another 20 min to
fully purge the microchannels. The solvent extraction process was identical in all cases.
5.2.4 Quantifying astaxanthin content
To compare the efficiency of astaxanthin extraction methods, the change in coloration of
individual H. pluvialis cells was quantified by image processing for all cases. An equation was
devised which converts the 8-bit RGB values measured for each pixel within the area of a cell,
into a coloration-based global “extracted red content” metric, indicative of the relative astaxanthin
concentration:
𝐴 = (100 −0.59𝐵 + 0.41𝐺
2.55) ∗ (1.36
255−𝑅45 ) (5 − 1)
where R, G and B represents the red, blue and green value at that pixel, respectively. The first
factor was determined using the 475 nm absorbance peak of pure astaxanthin,[258] which
corresponds to R, G and B values of 0, 178, and 255 respectively. Blue and green values are
weighed based on their relative importance to astaxanthin absorption. The second factor was
determined based on the observed drop of the R value in control experiments, without cell-wall
disruption, due to cell shrinkage and associated increase in astaxanthin pigmentation within that
smaller area. Cell shrinkage under exposure to hypertonic media during preliminary control
experiments was used to establish a relationship between R value and astaxanthin concentration –
R value within a cell with a constant quantity of astaxanthin, albeit at different times with different
volumes. Such shrinkage was not induced during hydrothermal disruption experiments presented
herein. It was determined that in this case, a drop of 45 in R value after solvent extraction
corresponded to a 36% decrease in area and corresponding increase in local astaxanthin
concentration. Individual cells in images were selected and tracked to calculate the extraction
efficiency according to (Ai-AF)/Ai where Ai and AF represent the initial and final red content.
64
Images taken during hydrothermal disruption processes were slightly affected by the heat but the
effect on RGB values was negligible. Notably, our evaluation based on relative red content
provides a rapid, in-situ measure of cell disruption and extraction efficiency. The observed red
pigment is expected to be 80-99% astaxanthin (free form, 4-5%; monoesters, ~70%; and diesters
15-20%) out of overall carotenoids based on previous studies72,259,260. The dark red astaxanthin
pigment within otherwise largely transparent cells provides a high contrast handle for rapid, real-
time screening of conditions at the individual cell level, not possible with off-chip methods. The
method is thus well suited to rapidly screen a large number of conditions for disruption and
extraction, with the best performing conditions candidates for more detailed analysis.
5.2.5 HPLC analysis of extracted astaxanthin
The acetone extracts from the hydrothermally treated (200 ºC for 10 min) cells and mechanically
disrupted cells (by mortar and pestle) were analysed using a high-performance liquid
chromatograph (HPLC) (Shimadzu SPD-10A) equipped with a reversed phase C18 (Supelco, 25
cm × 4.6 mm) column. The detection wavelength was set to 475 nm and the analysis protocol was
identical to previous research.258
5.3 Results and discussion
Initial and post-extraction cell images are presented for each wall disruption method in Figure 5-
2a, enabling direct comparison of different wall disruption methods. Each post-extraction image
was taken immediately after acetone extraction. Figure 5-2b shows normalized extracted red
content, as obtained with equation (1). Notably, hydrothermal disruption methods demonstrated
the largest change in coloration, with cells transitioning from a dark red hue to bright red (at 150
ºC) or white (200 ºC), indicative of near-complete astaxanthin extraction (>95% red content
reduction for the 200 ºC cases). Shorter duration tests at 200 °C indicate that the disruption process
is rapid at this temperature and insensitive to duration. A hydrothermal disruption temperature of
200 °C was sufficient to quickly decompose the cell wall in a span of a few minutes, which is in
agreement with previous research.13 Cells that were not subjected to any wall disruption treatment
(control case) show very little coloration change (7.5% red content reduction). The exchange of
media from deionized water to acetone causes water to exit via the water-permeable cell, resulting
in a 34% reduction in cell area. That cells retain their circular shape is indicative that the cell wall
and plasmalemma were not significantly compromised during acetone exposure. As the cell wall
65
contracts, however, light scattering at the cell wall - responsible for the white halos seen in the
initial images – is greatly reduced, with only light blue outlines remaining. This change is
attributed to water loss within the cell, and the related contraction of the plasmalemma and outer
cell wall (disappearance of inner bright ring, corresponding to the plasmalemma, confirmed by
brightfield imaging, ESI). The extraction efficiencies for acid and base treatments were higher than
the control, achieving 19.3% and 13.2% recovery, respectively. Cell size reduction was similar to
the control case (36%, 34%). During the HCl disruption process, astaxanthin bleeding was
observed in one of the cells, indicting minor cell wall disruption in isolated cases. For the NaOH
disruption process, no discernible structural changes other than size reduction occurred. The
relative extraction efficiencies achieved are slightly improved and shows similar improvement
(control, 20%; acid treatment, 35%; and base treatment 40%) to those published previously in bulk
reactors with longer extraction time.75 Collectively, these result demonstrate that short duration
treatment at 200 ºC is very effective for subsequent astaxanthin extraction, in sharp contrast to
traditional acid and base disruption methods.
66
Figure 5-2. a) Dark field images of mature red cysts of H. pluvialis at both the initial stage
and after solvent-based extraction, for each of five tested cases. All images have the same
scale bar, and were obtained with identical settings using darkfield microscopy. b)
Normalized extracted red content for each case. Reproduced by permission of The Royal
Society of Chemistry.
67
Figure 5-3 shows detailed time-course images of cells during the full extraction protocol, for each
of the four hydrothermal cell wall disruption recipes (detailed intermediate steps in the processes
of Figure 5-2). As shown, a treatment temperature of 150 ºC was sufficient to disrupt the
plasmalemma, but it was insufficient to disrupt the cell wall. In the 150 ºC case run for 30 min, the
cells darkened during heating followed by continuous fading of the outer regions (between the
“initial” and “beginning of the disruption process” images). After a return to room temperature, a
5 µm layer of interspace between the plasmalemma and cell wall was observed. A video of acetone
extraction is presented in ESI† - Video S2.
Figure 5-3. Time-course images of red cysts treated with hydrothermal processes of 150 oC
for 30 min, 200 oC for 30 min, 200 oC for 10 min and 200 oC for 5 min, respectively. The
scale bar is identical for all images. Reproduced by permission of The Royal Society of
Chemistry.
Figure 5-3 also shows the high efficiency of hydrothermal astaxanthin recovery at 200 ºC, for all
tested disruption times – with slight improvements at longer times. For the 30 min at 200 ºC case,
initial fading around the cell periphery was observed during heating (as observed in the 30 min at
150 °C case). However, rapid growth of the interspace and significant astaxanthin bleeding
occurred 1 min into the disruption process (Figure 5-3), indicating disruption of both the
68
plasmalemma and the cell wall. Rapid cell wall disruption within 1 min of treatment is also
supported by our study of the role of flow rate, presented in Figure 5-9. During the 200 °C
disruption, the polysaccharide wall is expected to slowly depolymerise into furan compounds49,
however, no discernible color change was observed on the cell wall, and furan-based RGB value
modifications are considered negligible. Astaxanthin flow from the cell to the channel was
observed 20 min into the disruption process. The majority of the pigmentation change occurred 1
min into acetone extraction, indicating high cell permeability, in contrast to that observed for other
cell disruption conditions. After 20 min of solvent extraction the cells showed virtually no
pigmentation indicating near complete astaxanthin recovery. The other tested times for the 200 ºC
case (10 min, 5 min) produced similar results, with the 5 min case showing a pigmentation change
during extraction that was less complete. These observations, which were made possible through
the direct visualization enabled by our microfluidic platform, have implications for the optimum
processing parameters of cell wall disruption techniques in larger commercial systems. These
findings indicate an optimal disruption duration between 5 and 10 min to both maximize the
extraction efficiency, and minimize the amount of time and energy required for processing.
Figure 5-4. Normalized red content during acetone extraction for six cells treated by
hydrothermal processing at 200 °C for 10 min. Inset images shows corresponding images of
one representative cell at four times during the procedure. Reproduced by permission of The
Royal Society of Chemistry.
69
The first 25s of the extraction profile for six individual cells disrupted for 10 min at 200 °C is
shown in Figure 5-4. The plot demonstrates the speed at which cell respond to acetone extraction
after hydrothermal treatment. Six cells were tracked to monitor the astaxanthin content which was
normalized to the post-treatment conditions of each cell. The first 25 seconds of solvent extraction
show a rapid decrease in astaxanthin (> 90%) in each cell (a video showing rapid extraction is
presented in ESI† - Video S3).
Figure 5-5. HPLC analysis of cell extract products for (a) mechanical extraction using a
mortar and pestle and (b) hydrothermal extraction for 20 min at 200 °C treatment.
Reproduced by permission of The Royal Society of Chemistry.
The HPLC analysis in Figure 5-5 demonstrates that the extracted red content for the case of 200 °C
for 10 min is similar in profile to that of cells mechanically disrupted using a mortar and pestle –
and correspond to expected astaxanthin HPLC peaks. Such a similarity demonstrates the
effectiveness of rapid hydrothermal treatment as an attractive alternative to mechanical disruption
methods. There are, however, two notable differences between the spectra. First, diester peaks are
clearly identifiable in the mechanical extraction case, but not in the hydrothermal case. Such a
70
difference could be due to a portion of diesters being hydrolyzed into monoesters and further
converted to free form astaxanthin. Second, the magnitude of peaks overall is much lower in the
hydrothermal on-chip case due to a level of dilution (a few nanograms of astaxanthin dissolved in
1mL of acetone), and is only an artefact of the visualization method here – trapping a small number
of cells in a flow – and is not a practical limitation of high temperature hydrothermal disruption at
larger scales. Notably, the presence of monoester and carotenoid HPLC peaks suggests the high
temperature, high pressure hydrothermal process did not have a significant adverse effect on
astaxanthin in general, perhaps due to the presence of largely intact rigid cell walls, which have
been shown to protect astaxanthin from thermal degradation at higher temperatures (eg. spray
drying at 220 ºC).261 The absence of oxidising agents within the cells, and inability for external
oxygenating species to enter the reactor, led to effective astaxanthin extraction.
Whereas the presence of strong astaxanthin HPLC peaks in Figure 5-5b demonstrates the
effectiveness of our hydrothermal disruption method, there remains uncertainty as to any
temperature-induced conformational changes, such as trans-cis isomerization of extracted
carotenoids. Kaczor and Baranska performed in-situ Raman spectroscopy to monitor astaxanthin
structural change in a single cell with thermal stress up to 150 °C.262 It was found that astaxanthin
experienced only minor conformational change. While more study is required to elucidate
astaxanthin structural changes under high pressure and high temperature conditions, strong HPLC
peaks here demonstrate the potential of the hydrothermal extraction method.
There is potential to adapt the hydrothermal astaxanthin extraction method presented herein for
commercial-scale reactors. To achieve industrial-scale hydrothermal astaxanthin recovery, the
results of our chip-scale experiments suggest three criteria that must be met: (i) an inert
environment is required to prevent astaxanthin oxidation, (ii) temperature, pressure and residence
time must be precisely controlled, and (iii) low shear conditions must be satisfied to prevent cell
wall rupture and subsequent astaxanthin degradation (as observed in Figure 5-10 in flow
experiments). Such a combination of criteria could be met by with a continuous flow-through
reactor approach, similar in principle to the experiments performed herein, albeit with macroscale
reactor tubes in parallel. In short, while the chip allows unprecedented cell-scale resolution of the
process, the reaction conditions achieved here are in no way unique to the chip-based reactor. The
observations here can be readily applied to engineer scaled processes, with careful engineering to
ensure similar conditions and residence times.
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5.4 Conclusion
In this chapter, we have demonstrated a hydrothermal method of cell wall disruption for
astaxanthin extraction uniquely enabled by a high-pressure, high-temperature microfluidic device.
Individual H. pluvialis cells were trapped and visualized throughout one of several procedures,
both chemical and hydrothermal, and extracted red content was quantified optically. Hydrothermal
disruption at 200 ºC was the most effective wall disruption technique, enabling near complete
astaxanthin extraction from wet biomass, a significant improvement over traditional methods.
5.5 Supplemental material
The experimental apparatus is adapted from our previous research with slight modification
upstream of the microfluidic reactor (fabrication details are shown in Chapter 4). The algae
solution, chemicals and solvent stored within piston cylinders [2a & 2b] were separately injected
into the microfluidic reactor by controlling the switching valve [3a]. Switching valve [3b] was
fixed at position 1 until the solvent extraction stage. After 1 min of solvent extraction, the switching
valve [3b] was quickly switched to position 2 to unload the extracted astaxanthin to the fixed-
volume sample isolator.
Figure 5-6. Schematic diagram of the experimental setup with flow direction indicated by
arrows along the processing path. The flow path of the switching valve at two positions is
shown using green lines.
72
Figure 5-7. Irradiation spectrum of light used to induce astaxanthin accumulation in
Haematococcus pluvialis.
Figure 5-8. Microscope images of red cysts, indicating coloration change during the heating
up phase in the hydrothermal disruption process. The scale bar applies to all images.
73
Figure 5-9. Dark field, bright field and fluorescence images of initial cells and after acetone
extraction. Fluorescence 1 and 2 images were taken using FITC (excitation filter: 475/50 nm;
emission filter: 540/50 nm) and TxRed (excitation filter: 559/34 nm; emission filter: 630/69
nm) filter cubes respectively. The scale bar of 50 µm applies to all images.
74
Figure 5-10. Cell wall deformation of red cysts under hydrothermal processes at 200 °C in
10 min with and without flow. Green dashed lines indicate cell wall boundaries. The scale
bar of 50 µm applies to all images.
Video S1: Real-time video showing H. pluvialis cells being trapped by posts in a microfluidic
channel. Cells in this video were not fully transformed into mature red cysts and are here used as
a demonstration.
Video S2: Real-time video showing the in-situ acetone extraction of red cysts of H. pluvialis
treated by hydrothermal disruption at a temperature of 150 °C and pressure of 6 MPa for 30 min.
Video S3: Real-time video showing the in-situ acetone extraction of red cysts of H. pluvialis
treated by hydrothermal disruption at a temperature of 200 °C and pressure of 6 Mpa for 10 min.
75
Chapter 6.
Astaxanthin Extraction from Algae using
Supercritical CO2 with Co-solvent
Supercritical CO2 is an attractive green-solvent for the extraction of astaxanthin from microalgae,
but current processes require prohibitively high pressures and long extraction times. This study
demonstrates the efficacy of low pressure supercritical CO2 extraction of astaxanthin from
disrupted Haematococcus pluvialis. We employ a microfluidic reactor that enables excellent
control and allows direct monitoring of the whole process at the single cell level, in real time.
Astaxanthin extraction using ScCO2 achieved 92% recovery at 55 °C and 8 MPa applied over 15
hours. With the addition of co-solvents, ethanol and olive oil, the extraction rates in both
experiments were significantly improved reaching full recovery within a few minutes. Notably,
for the ethanol case, the timescales of extraction process are reduced 1800-fold from 15 hours to
30 seconds at 55 °C and 8 MPa, representing the fastest complete astaxanthin extraction at such
low pressures.
This chapter was submitted as original paper to Bioresource Technology. The candidate was the
first author in this work and played the primary role in designing the research, performing the
experiments, analyzing the data, and writing the paper. Additional authors for the work include
Mr. Zhenbang Qi, Dr. Thomas Burdyny, Ms. Tian Kong and Prof. David Sinton. Their
contributions were central to the publication of this work and are gratefully acknowledged and
appreciated.
6.1 Introduction
The production of high-value products from microalgae is currently the most economically viable
option for the industry to offset the high overall costs of growth and extraction.15,69,242,263
Astaxanthin naturally occurs in a number of aquatic species including the microalgae
Haematococcus pluvialis which accumulates the highest levels per cell. Due to its unique structure
and strong antioxidant properties in humans70,242, natural astaxanthin from microalgae has
76
cultivated a large nutraceutical industry264–266. Unfortunately, the accumulation of astaxanthin
within H. pluvialis is strongly linked to the cell’s rigid wall that is extremely resistant to both
chemical and physical disruptions.73,74,118,267 Also, once the cell wall is breached, organic solvents
are generally needed to extract astaxanthin from the interior. The range of available disruption and
extraction methods and solvents is limited by the requirements for human product consumption.268
A detailed understanding of factors affecting astaxanthin extraction using green solvents at the
cellular level are needed to help inform future large-scale astaxanthin production processes.
Recently, supercritical carbon dioxide (ScCO2) has attracted attention as a green solvent for the
extraction of a variety of bioactive compounds.78,269 Supercritical CO2 features three major
advantages over organic solvents: 1) it is abundant and benign to human health and the
environment; 2) CO2 solvent can be removed easily by evaporation at room temperature and
pressure; and 3) bioactive compounds are well-preserved due to the inertness of CO2 and relatively
low critical temperature (31.1 °C).77,128 However, previous studies using ScCO2 to extract
astaxanthin show large variations in the overall recovery rates, even with similar processing
conditions (Table 6-1). The overall extraction efficiency with ScCO2 depends greatly on the
preparation of the feedstock and the addition of any modifiers/co-solvents, while operating
conditions (temperature, pressure, duration) and extraction cycles are parameters which could also
be adjusted. With regard to commercial feasibility, temperatures for astaxanthin extraction are
limited to a relatively low 70 °C81,83, while the minimum pressure needed for ScCO2 is 7.39 MPa
(although the majority of work to date employs much higher pressures).
Due to the rigid cell wall, direct extraction of wet microalgae biomass using ScCO2 is generally
not feasible. A common strategy is then to dehydrate and mechanically disrupt the feedstock prior
to CO2 extraction, easing the downstream extraction process. However, reported astaxanthin
recovery rates vary widely due to the range of disruption tools used and the ambiguity in
quantifying the degree of cell wall disruption. For example some researchers found the overall
recovery rates were below 50% after cell wall disruption, even when the operating pressure was
as high as 30 MPa.79,80 Furthermore, dehydration and mechanical cell disruption add considerable
time, energy and cost to the process. Intracellular water and disrupted cell debris can also present
challenges such as caking, further reducing extraction rates and/or yield over time.270
77
Another strategy to aid the extraction process is to employ co-solvents81,84 such as ethanol and
vegetable oils in ScCO2. Although this strategy allowed for a higher recovery rate of 71 %, high
extraction pressures (31 MPa) and multiple cycles of batch processing (8 cycles) were required
(Table 6-1 - #5). The combination of mechanical cell disruption and co-solvent extraction has also
been tested79,84, indicating the potential to greatly speed up extraction and enhance recovery using
ethanol co-solvent. However, the high associated capital and operating costs present barriers to
large-scale production271,272. Large discrepancies between reports in literature present additional
uncertainty. Process control and cell-scale resolution are essential to quantify the effectiveness of
extraction processes, and to inform ScCO2-based extraction of astaxanthin from microalgae at
industrial scales.
Table 6-1: Summary of studies on ScCO2 extraction of astaxanthin from H.pluvialis
# Feedstock Solvent Extraction
conditions Duration
Recovery
rate (%) Reference
1 Wet
Biomass
Acetone STP 16 h 20 Mendes-Pinto et
al., 2001
2 CO2 27.6 Mpa, 60 °C 30 min 1 Pan et al., 2012
3
Dry powder
CO2 40 Mpa, 70 °C 5 h 25 Krichnavaruk et
al., 2008
4 CO2 50 Mpa, 70 °C 4 h 19 Machmudah et
al., 2006
5 Ethanol in CO2
= 9.23 mL/g 31 Mpa, 50 °C
160 min (20 min x 8)
71 Pan et al., 2012
6 Ethanol in CO2
= 2.3 mL/g 43.5 Mpa, 65 °C 3.5 h 87 Wang et al., 2012
7 Ethanol in CO2
= 10 % (v/v) 40 Mpa, 70 °C 5 h 51
Krichnavaruk et
al., 2008
8 Ethanol in CO2
= 5% (v/v) 40 Mpa, 70 °C 4 h 80
Machmudah et
al., 2006
9
Disrupted dry powder
CO2 35 Mpa, 55 °C 2 h 5
Reyes et al., 2014
10 Ethanol in CO2 = 13 % (w/w)
35 Mpa, 55 °C 2 h 56
11 Ethanol in CO2 = 50 % (w/w)
7 Mpa, 45 °C 2 h 124
12 Ethanol in CO2 = 70 % (w/w)
7 Mpa, 45 °C 2 h 65
78
13
Disrupted dry powder – degree 1
CO2 30 Mpa, 60 °C NA 47
Nobre et al., 2006 14
Disrupted dry powder – degree 1
Ethanol in CO2 = 10 % (v/v)
30 Mpa, 60 °C NA 59
15
Disrupted dry powder – degree 2
Ethanol in CO2 = 10 % (v/v)
30 Mpa, 60 °C NA 92
*NA: not reported
In this paper, we screened the extraction conditions of astaxanthin from H. pluvialis cells using
ScCO2 with in-situ observation on a pressure- and temperature-controlled microfluidic platform.
For the first time, direct monitoring in real time and quantification of the astaxanthin extraction
process on a single cell is realized. Applying this method we demonstrate potential for rapid, low-
pressure astaxanthin extraction with ScCO2 and co-solvent.
6.2 Experimental Section
Haematococcus pluvialis (H. pluvialis) cells were obtained from Iconthin Biotech Corp. and
maintained in aplanospore form at 25 ºC under continuous white light illumination at ~60 µmol
m−2s−1 in media according to the recipe described elsewhere.256 The mature H. pluvialis cysts with
cell diameters from 40 to 70 µm were selected by using two cell strainers. CO2 (99.9%) was
purchased from Praxair. Ethanol (96%) was purchased from Sigma-Aldrich, and the olive oil was
commercially available extra virgin, food grade.
The microfluidic chip was fabricated out of glass and silicon to provide optical access to the
reaction channel and high thermal conductivity for precise temperature control. This device
featured a long single reaction channel, 300 μm wide, 100 μm deep and 680 mm long, with
multiple C-shape traps located in the temperature-controlled region. Each trap has an inner
diameter of 100 μm and outer diameter of 160 μm with a front opening of 80 μm and back opening
of 20 μm. These traps served to immobilize cells while allowing fluid flow. The device was
fabricated by deep reactive ion etching (DRIE) of the silicon, with subsequent anodic bonding to
borosilicate glass of 1.75 mm thickness. The temperature-controlled region of the chip was inserted
into a stainless steel heating chuck with temperature controlled by a PID controller and three
79
cartridge heaters. The full fabrication procedure for devices of this type was reported in a previous
study.156
For each experiment, a suspension of selected cysts in media was injected into the chip using a
syringe and trapped within C-shape walls. After the cells were loaded, deionized water was flowed
into the chip at a rate of 1 mL min-1 for 5 min to flush the media prior to cell disruption.
Hydrothermal disruption of cell wall at 200 ºC and 8 MPa for 10 mins was used as a standard
disruption process that provided uniform and thorough disruption to all cells. The full
hydrothermal disruption procedure for H. pluvialis cells was reported elsewhere.197
All supercritical CO2 extractions were carried out in the microfluidic chip on disrupted cells. For
all experiments, the heating chuck was adjusted to steady experimental temperatures before
pumping CO2 into the chip. The schematic diagram of the experimental setup and procedure are
illustrated in the supplementary material. The flow path of the system was controlled by a 6-port
valve with minimal dead volume (Rheodyne®7030). When switched, high pressure CO2 could be
slowly pumped into the channel without depressurizing the entire chip and avoid back flow. The
flow rate of CO2 was controlled by a back-pressure regulator (Swagelok KCP1GRA2C1P10000).
During extraction, and at all stages of astaxanthin recovery, the cells were observed using darkfield
microscopy (Olympus BXFM microscope, 10× objective).
To compare the extraction rate and efficiency of different processing conditions, the change in
coloration of individual H. pluvialis cells was quantified by image processing for all cases. The
relative astaxanthin content was determined directly from the colour images through analysis of
RGB pixel values as reported elsewhere197 (additional details in supplementary material).
6.3 Results and discussion
To allow astaxanthin to be extracted from the cell into the extraction medium, the cell wall was
first disrupted. Hydrothermal disruption was chosen here instead of chemical (acid and base),
biologic (enzyme), or mechanical (eg. grinding, milling and bead beating) approaches due to the
uniformness and completeness of the disruption process achieved.75,76,197 As outlined in Fig. 1, all
trapped cells undergo hydrothermal disruption prior to the extraction processes. The disrupted cells
were then extracted with ScCO2 at various temperatures and solvent conditions. At each stage in
the process, the astaxanthin content was approximated from the colour image data.197 To provide
80
a baseline reference point for 100% extraction, all test processes were followed by acetone
extraction which has been proven as a highly effective approach to remove residual
astaxanthin79,82,84. A summary of experimental conditions and results of this study are shown in
Table 2.
Figure 6-1. Single-cellular visualization and quantification of astaxanthin extraction in
Haematococcus pluvialis using supercritical CO2 and co-solvents. Post ScCO2 and ethanol
extraction using acetone provides an overall extraction efficiency metric. Images were taken
from pure ScCO2 extraction experiment at 70 °C.
Table 6-2: Summary of supercritical carbon dioxide extraction experiments and results.
Experiment Solvents Extraction
conditions Duration
Reduction in
normalized red
content (%)
Red
content
drop rate
(%/min)
1 CO2 8 MPa 40 °C
no flow 120 min 11.9 0.07
2 CO2 8 MPa 55 °C
no flow 120 min 22.4 0.23
3 CO2 8 MPa 70 °C
no flow 120 min 63.1 0.57
4 Ethanol in CO2
= 20 % (v/v)
8 Mpa 40 °C
120 µL min-1 15 min 84.4 31.4*
5 Ethanol in CO2
= 20 % (v/v)
8 MPa 55 °C
120 µL min-1 15 min 98.3 168.5*
81
6 Ethanol in CO2
= 20 % (v/v)
8 MPa 70 °C
120 µL min-1 15 min 97.8 167.6*
7 Olive oil in CO2
= 20 % (v/v)
8 MPa 55 °C
120 µL min-1 15 min 98.6 168.9*
*This rate is estimated based on the change of normalized red content in 35 seconds.
We first investigated the effects of temperature on pure ScCO2 extraction by setting the
temperature of the reaction chamber to 40 °C, 55 °C and 70 °C. For all pure CO2 extractions it was
necessary to flow CO2 (at atmospheric pressure) to remove the water surrounding the cells.
Without this drying step, water surrounding the cells effectively shielded them from ScCO2,
resulting in no noticeable extraction over long periods. Once the water was removed, we applied
an operating pressure of 8 MPa, just above the critical pressure (7.39 MPa), and a relatively low
pressure as compared to previous work in this area.79,80 No significant change in colour was
observed in the 40 °C case but a noticeable change in the amount and colour of the remaining
astaxanthin in cell was observed in the 70 °C case (the real time images are presented in Figure 6-
1). By calculating the change in normalized red content, the extraction in terms of overall
percentage is plotted over time in Figure 6-2a. In this two-hour interval, the 40 °C case resulted in
very little astaxanthin recovery (12 %) with a slow extraction rate; the 55 °C case showed improved
extraction rates with 22% of the red content being extracted; the 70 °C case showed a significant
improvement by achieving 63% drop over the same time period. The trend observed here for higher
extraction rates at higher temperatures is well supported in the literature.81,83–85
To further characterize the removal over time, the full extraction process was visualized over a 15-
hour period at 55 °C as shown in Figure 6-2b. The near-complete extraction of astaxanthin using
pure ScCO2 was achieved over this period indicating that the reported extraction efficiency is also
tightly related to extraction time. These results show that relatively low-pressure supercritical CO2
is capable of effectively complete astaxanthin extraction, and the rates are higher than those
achieved previously. However, even at the maximum temperature suitable for astaxanthin (70 °C),
the process requires hours to complete.
82
Figure 6-2. a) Darkfield images H.p cells before and after ScCO2 extraction at 40 °C and 70
°C. b) The progression of normalized red content for ScCO2 extraction at 40 °C, 55 °C and
70 °C indicating higher extraction temperatures resulted in higher extraction rates. Solid
lines represent 1st order trendline fits to the experimental data with equation given by the
side. c) The normalized red content for ScCO2 extraction process at 55 °C for a complete
extraction process over 900 minutes with dark-field snapshots along the process.
Co-solvents present an opportunity to increase extraction rates beyond that of pure ScCO2. A
popular organic co-solvent used is ethanol, which is inexpensive and commonly used in food
processing. Adopting a similar experimental approach as pure ScCO2, we observed the cellular
extraction over time of disrupted H. pluvialis cells using a 4:1 CO2:ethanol flow rate ratio (100 µL
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min-1 and 25 µL min-1) with temperatures again varying between 40 °C and 70 °C. Following
disruption of the cell wall (hydrothermally in pure CO2 as before), Figure 6-3 shows the relatively
rapid removal of astaxanthin at the three test process temperatures. With addition of ethanol, the
timescales of extraction process are reduced 1800-fold from 15 hours to 30 seconds for a
ScCO2/ethanol mixture at 55 °C (video 1). Even at 40 °C, the final extraction is over 84% of the
measured red content after 1000 seconds.
Figure 6-3. The progression of normalized red content for ScCO2 extraction with ethanol
over 1000 seconds at 40 °C, 55 °C and 70 °C. Zoom-in plot for the first 30 s extraction time
indicates a rapid extraction of astaxanthin from ScCO2 with ethanol at 55 °C and 70 °C.
The rapid extraction of astaxanthin was observed around 4 minutes after we started pumping
ethanol which is approximately the amount of time required to replace the dead volume (~500 µL)
at the given flow rate (125 µL/min). The enhanced extraction with ethanol is attributed to two
factors. First, water is miscible with the ethanol phase, removing the two-phase interfacial barrier
between the solvent and the water-wet surface of the cell. Secondly, the solubility of astaxanthin
in ethanol is 5 orders of magnitude higher than that in ScCO2 .273
Olive oil is another attractive ScCO2 co-solvent due its potential to be directly combined with
astaxanthin for use in food products, avoiding the need for the solvent separation step required
84
with ethanol. Olive oil has been shown to achieve extraction rates comparable to ethanol.81 As
shown in Figure 6-4a the addition of olive oil to the extraction fluid instead of ethanol also
performed significantly better than pure ScCO2 and reached complete extraction (~98%) in just
180 seconds. Images at various stages of the extraction process are shown in Figure 6-4a for an
olive oil co-solvent at 55 °C.
In contrast to the ethanol case, the removal of the water barrier by olive oil is observed as oil
immediately displacing the surrounding water phase. In this process, three phases can be observed
within the microchannel: ScCO2, oil and a small amount of water left from hydrothermal process.
Upon interaction with the disrupted cell-wall, the introduction of olive oil results in a number of
emulsions spontaneously appearing immediately around the cell. Rather than the formation of an
oil-water-astaxanthin mixture, the remaining water in the cell instead emulsifies into smaller
droplets and can be seen during the initial extraction phase (Figure 6-4b). Similar to the ethanol
case the increased solubility of astaxanthin versus ScCO2 is readily apparent by the rapid increase
in red content in the co-solvent phase envelope around the C-trap (video 2). While the colour
increases within this region, there is no evidence of colour loss to the surrounding CO2 fluid until
the fluid flow forces the oil droplet forward along the channel, leaving a depleted H. Pluvialis cell.
85
Figure 6-4. a) The comparison of normalized red content for ScCO2 extraction with ethanol
and olive oil at 55 °C over 200 seconds. Time-lapsed snapshots of the extraction process with
olive oil are provided. b) Snapshot of ScCO2 extraction with olive oil indicating three phases:
ScCO2, olive oil and water. The boundary of olive oil and CO2 phases are illustrated using
yellow and green dash lines.
86
Some differences between the results of the experiments here and those of previous studies are
noteworthy. First, with respect to the pure ScCO2 test cases, the results here show 92% extraction
in 15 hours at 55 ºC and 8 MPa, with extraction rates increasing with temperature. Other
studies81,84,85 have shown the highest total extraction level plateaued at 25% over 5 hours, at 70 ºC
and 40 MPa. We attribute our improved performance in these pure CO2 cases to the high degree
of disruption achieved prior to extraction and the precise control of conditions. Second, with
respect to co-solvent tests, we achieved full extraction within minutes which is in marked contrast
to other results showing comparable results only after several hours of extraction81,84,85,255. Here we
attribute the improved performance to within the reactor – namely that all cells had access to co-
solvents (in addition to the high degree of prior cell disruption noted earlier). These results point
to the possibility of extraction with significantly lower pressures, provided the reactor achieves a
high degree of cell-to-co-solvent contact. In the context of larger scale reactors, these results point
the potential to reduce pressures and processing time - key drivers of capital and operational costs.
6.4 Conclusions
In this work we demonstrate low pressure supercritical CO2 extraction of disrupted
Haematococcus pluvialis in a microfluidic reactor with optical access to monitor the extraction
process in real time. A near-complete (~98%) rapid extraction of astaxanthin using ScCO2 and an
ethanol co-solvent was achieved in 30 seconds at 8 MPa, representing the fastest complete
astaxanthin extraction at such low pressures. Our results using a single cell reactor provide time
resolved and direct evidence of practical astaxanthin extraction using supercritical CO2 at low
pressures in a matter of minutes, highlighting the potential for fast and full recovery of this valuable
bio-product.
6.5 Supplementary Information
The experimental apparatus was adapted from our previous research with different arrangements
specifically designed for supercritical CO2 extraction. The fluid flow path used is shown in Figure
6-5. At the beginning of the experiment, Haematococcus pluvialis (H.p) cells were loaded into the
microfluidic chip before connecting the chip to the flow system. Prior to the hydrothermal
disruption process, the channels downstream to the microfluidic reactor were pressurized to 8 MPa
with CO2 to avoid abrupt fluid flow due to depressurization when turning the switching valve [3].
Three high pressure pumps were used in the experiment: two water pumps [1a and 1c] and a CO2
87
pump [1b]. Deionised water in pump [1a] was used to flush the media and pressurize the system
to 8 MPa for the hydrothermal disruption process After hydrothermal disruption, the switching
valve [3] was switched to position 2 to supply supercritical CO2 for the extraction stage. Co-solvent
(ethanol or olive oil) was stored in the piston cylinder [2] and added to the system using a
controlling valve [5a]. Both pumps [1b & 1c] were running at constant flow rates during the
extraction process.
Figure 6-5. Schematic diagram of the experimental setup with the flow direction indicated
by arrows along the processing path. The flow of the switching valve at two positions is shown
using red lines.
To compare the extraction efficiency of astaxanthin for different processes, the change in
coloration of individual H. pluvialis cells was quantified by image processing for all cases. The
area of interest (AOI) was manually selected based on the size of observed cells and used
throughout the entire extraction process. The equation used to convert the 8-bit RGB values of
each pixel in the AOI into a coloration-based global “extracted red content” was adopted from
previous study and shown below:
𝐴 = (100 −0.59𝐵 + 0.41𝐺
2.55) ∗ (1.36
255−𝑅45 )
88
where R, G and B represents the red, blue and green value at that pixel, respectively. The first and
second factors were determined by the maximum astaxanthin absorbance peak at 475 nm and
empirical correlation between R-value and astaxanthin concentration (a drop of 45 in the R-value
corresponds to a 36% increase in astaxanthin concentration), respectively. The AOI was tracked
to calculate the extraction efficiency with respect to time according to (Ai-AF)/Ai where Ai and
AF represent the initial and final red content. Typical colors of a H. pluvialis cell observed during
extraction were selected and are illustrated in Figure 6-6 with RGB values, assigned astaxanthin
content and calculated astaxanthin content based on the given equation showing on the same row.
The small difference between the assigned astaxanthin content and calculated astaxanthin content
indicates a fair approximation using the given equation.
Figure 6-6. Illustration of typical colors of H. pluvialis cells with different concentration of
astaxanthin and the calculated RGB-based astaxanthin content.
89
Chapter 7.
Conclusions
7.1 Summary
Maximizing productivity from microalgal biomass requires innovative approaches to address the
central challenges in downstream processing. I have developed and applied a microfluidic platform
capable of performing reactions at high temperature (350 °C) and high pressure (20 MPa), precise
control on processing parameters, and providing optical access for in-situ observation and
quantification. The direct observation of microalgae downstream processes was made possible for
the first time and allowed visual characterization, fluorescence spectroscopy, and quantitative
imaging of the conversion at the single-cell scale during all stages of the reaction.
Besides the engineering achievement of successfully developed this microfluidic platform, a
significant contribution in science was also made to improve microalgae downstream processes.
In the hydrothermal liquefaction of microalgae project, biomass-to-biocrude conversion was
directly observed in a microfluidic channel and the reaction mechanism was indicated by a high
time-resolution change of fluorescence signature in 10 mins. Formation of oil droplets comprised
of aliphatic components was directly observed in a short reaction time suggesting that cell wall
disruption takes place within the early stages of these reactions. Inspired by the findings from HTL
processes, a hydrothermal method was introduced to disrupt rigid cell walls in Haematococcus
pluvialis for astaxanthin recovery. Hydrothermal disruption at 200 °C was the most effective wall
disruption technique, enabling near complete astaxanthin extraction from wet biomass, a
significant improvement over traditional methods. For ScCO2 extraction of astaxanthin, the degree
of disruption, water film surrounding the cells and mass transport showed dominating effects on
the performance of extraction process. Astaxanthin extraction using ScCO2 achieved 92%
recovery at 55 °C and 8 MPa applied over 15 hours. With the addition of co-solvents, ethanol and
olive oil, the timescales of extraction process are reduced dramatically from 15 hours to a few
minutes, representing the fastest complete astaxanthin extraction at such low pressures.
90
These results and the microfluidic platform on which they were collected represent the first of their
kind in the field of microalgae downstream processing. Although, conventional batch reactors have
competitive advantages on quantifying product yields which requires relatively large volumes of
products, microfluidic methods presented here offer a unique toolset to better understand the
processes with high-resolution information at cellular level, at specific operating conditions such
temperature and pressure. This level of insight has simply not been possible with previous
conventional reactors. The results presented in this thesis provide new insight into important
biomass-to-bioproducts conversion processes – insight that can be applied to improve large scale
operations. Perhaps most notably, the fast reaction rates achieved with this reactor (in some cases
at lower pressures than previously employed) highlight what is possible in larger reactors –
provided that excellent transport is achieved.
7.2 Future Outlook
As discussed earlier, converting biomass into valuable products is a critical step for producing
bioproducts from microalgae. The results presented in this thesis provided unprecedented insights
into these downstream processes but also proposed future challenges. Effectively use this
information to achieve the same mass and thermal transport shown in the microfluidic reactors at
large scale is the next challenge. Also, going down the path of bioproducts production, there are
many other challenges remained. Successfully making biocrude at low cost is just halfway through
and upgrading the biocrude to drop-in fuels at industrial scale still require major breakthroughs.
Perhaps most notably, the biocrude produced in this thesis work is highly oxygenated and thus
requires more downstream processing than, say, conventional crude oil. The purity of high value
products from microalgae is also a rising concern due to different downstream pathways were
used. Standardized testing protocols are expected for future microalgae products.
For both bioenergy and high-value bioproduct production, a more holistic approach over a lipid-
centric approach is needed to maximize all the potential value contained in the biomass. Looking
forward I see two major opportunities for improving the performance of biomass-to-biofuel and
biomass-to-bioproducts conversions respectively.
First, a clearer understanding of the reaction mechanism of HTL will become available by
investigating the effects of processing parameters on individual biomass components such as
carbohydrates and proteins. In particular, the reaction time associated with reaction temperature
91
and pressure is critical to the design of a sequential process such that each step is finely tuned to
optimize the yield of a targeted product from a specific feedstock. For example, fast hydrothermal
processes could be used as a cell wall disruption technique that enables access to high energy and
high value components – lipids. Once the lipids are recovered, the remaining biomass could then
be used as animal feed or further converted to bioenergy using a more target-specific conversion
method for proteins and carbohydrates. Work presented in Chapter 4 will help provide vital
insights towards optimizing the output of hydrothermal processes, particularly in cases where high
time-resolution information is required to carefully tune the processing parameters.
Second, technology developed during the commercialization of high-value bioproducts from
microalgae will help drive the development of microalgal biofuel. The downstream processes of
both applications share similar challenges – high water content in the feedstock. The breakthroughs
in cell-wall disruption and product extraction made from commercial cases will help to reduce the
costs of biofuel production helping make it an economically viable alternative to fossil fuels.
Further integration and development of utilizing waste streams, such as nutrients from wastewater
and energy from waste heat or geothermal, can further provide value to the competition with fossil
fuels.
Finally, challenges and exciting opportunities both exist for improving the performance of
downstream processing. The markets open to microalgal products are vast and diverse. Low-
volume high-value bioproduct markets are the best initial target with biofuels being the ultimate
long-term high-volume market. Collectively these opportunities motivate near-term advances to
overcome challenges in the downstream processing, and to maximize the overall recoverable value
from microalgae biomass.
92
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Appendices
The following appendix is a manuscript for which the applicant was co-author and has been
published in Angewandte Chemie, reprinted274 with permission from John Wiley and Sons. While
it is not to be considered part of the core contributions of this thesis, they are nevertheless
worthwhile considering since it provides additional breadth to the application of high temperature
and pressure microfluidics.
A1. The full thermodynamic phase envelope of a mixture in 1000
microfluidic chambers
Authors: Yi Xu, Jason Riordon, Xiang Cheng, Bo Bao and David Sinton*
Abstract: Knowing the thermodynamic state of complex mixtures – liquid, gas,
supercritical or two-phase – is essential to industrial chemical processes. Traditionally,
phase diagrams are compiled piecemeal from individual measurements in a pressure-
volume-temperature cell performed in series, where each point is subject to a long fluid
equilibrium time. Herein, 1,000 microfluidic chambers, each isolated by a liquid piston
and set to a different pressure and temperature combination, provide the complete
pressure-temperature phase diagram of a hydrocarbon mixture at once, including the
thermodynamic phase envelope. Measurements closely match modelled values, with a
standard deviation of 0.13 MPa between measurement and model for the dew and bubble
point lines, and a difference of 0.04 MPa and 0.25 ºC between measurement and model
for the critical point.
Long a fixture within chemistry textbooks, phase diagrams are crucial to understanding chemical
processes – phase diagrams identify how complex fluid mixtures behave under a wide range of
pressures and temperatures.275 Detailed knowledge of a multi-component fluid’s two-phase
envelope and associated key parameters – dew and bubble point lines, liquid volume lines and
critical point – is critical to understanding a variety of chemical processes,276 and is often difficult
to obtain using equation-of-state models.277 For many fluid mixtures, even small changes in
composition can have a profound impact on thermodynamic properties.278 Measuring fluid phase
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properties is routine in the oil and gas industry, where complex hydrocarbon mixtures279,280 – raw
natural gas, gas condensates and oil – are analyzed during recovery, treatment, transportation and
consumption processes. Measurement of the phase diagram is typically performed using traditional
pressure-volume-temperature (PVT) cells.281 Compiling even a sparse phase diagram in this
manner is a costly and time-consuming process, often requiring hours with costly large-scale
experimentation to determine the phase state at a single point on the pressure-temperature plane.
Recently, specialized microfluidic approaches have been applied to measure isolated phase
properties, including the dew point,22,282 bubble point,22,283,284 and liquid-to-vapor ratios.284 These
methods each use direct visual observation to determine the formation of droplets or bubbles
within a confined planar surface or microfluidic channel and benefit from short equilibrium times,
reduced sample volume requirements and precise control over pressure and temperature offered
by microfluidic systems.16,285 Approaches to date, however, involve measuring single pressure-
temperature conditions in series, and fail to exploit parallelization opportunities as recently
demonstrated in other phase-mapping microfluidics applications, for example in concentration-
concentration or temperature-concentration phase mapping of salts,286 polymers287 or protein
crystallization.287–289 Bao et al. recently demonstrated phase diagram mapping of pure CO2 by
using an array of microwells subject to different pressure-temperature conditions.290 Visualizing
the two-phase envelope of mixtures, however, was not possible due to the interconnected nature
of the device. Specifically, fluid communication between microwells precluded the resolution of
the mixture phase envelope, limiting application to pure substances. Mixtures, however, are the
norm in chemical applications and pure substances are the exception.
Herein, we report full mapping of the phase diagram of a fluid mixture with an array of 1,000
microfluidic chambers, each isolated and pressurized by liquid pistons. The liquid piston fluid
compartmentalizes the test fluid, fully isolating and pressurizing each microfluidic chamber and
enabling accurate measurement of vapor-liquid ratios therein. A temperature gradient is applied
perpendicular to the pressure gradient by using external temperature controls. Our method enables
rapid one-step measurement of a fluid mixture’s phase diagram including dew and bubble point
lines, liquid volume lines and critical point.
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Figure 1. a) Schematic of the rapid pressure-temperature phase mapping device for mixtures. Insets show enlarged
regions of the device, Liquid piston operation is demonstrated in (b) and (c), respectively. d) Corresponding phase
diagram.
Our approach is shown schematically in Figure 1. Trapped pockets of test fluid are isolated by the
piston fluid within long vertical interdigitated microfluidic chambers (2 mm long × 100 μm wide
× 15 μm tall) in a silicon-glass chip by sequentially loading first the mixture and then the piston
fluid. These microfluidic chambers are each connected at their base to dead-end horizontal
channels which are positioned parallel to each other across the device, and connect to the flow
channel at their leftmost point (see Figure S1 in the Supporting Information for full details). During
regular operation, the piston fluid is continually flowing through the device – entering at the inlet,
flowing vertically along the flow channel, and exiting at the outlet; fluid throughout the remainder
of the channel network is stagnant (Figure 1a). The pressure in each horizontal channel is fixed
based on the location of the connection point to the flow channel, and pressure measurements at
both ends. At low pressure, the liquid pistons are near the entrance, whereas at high pressure, the
liquid pistons extend deep into the chamber (Figure 1b and c). In addition, an orthogonal linear
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temperature gradient is achieved in the horizontal direction via temperature controlled copper
blocks in direct contact with the silicon. The high thermal conductivity of silicon ensures a linear
temperature gradient and precise control over local temperature conditions within the gradient-
orthogonal microfluidic chambers (see Figure S2 in the Supporting Information for temperature
calibration). The result is a pair of orthogonal, linear pressure and temperature gradients across the
device, which enable mapping of the pressure-temperature phase diagram of a fluid mixture (see
Table S1 in the Supporting Information). The temperature variation across a given chamber is
negligible, less than 1% of the full temperature range and the temperature range is tunable to the
region of interest.
The ideal piston fluid should (i) be insoluble with the test mixture (i.e. use a polar piston fluid for
a non-polar test mixture), (ii) have strong wall-wetting properties, (iii) remain liquid at all test
pressures and temperatures, (iv) have a low vapor pressure, (v) have a low viscosity and (vi)
provide sharp visual contrast with the test fluid. Here, to contain our test fluid – an 80.0% propane
+ 20.0 % methane mixture (mole fraction, Praxair Canada, Inc.) representative of thermogenic
natural gas.291–293 – we used ethylene glycol (99.99% purity, Shell Chemical, LP.), which readily
satisfies these criteria and has been deployed successfully in related applications in industry.294,295
Notably, the solubility of natural gas within ethylene glycol is low, and has been well characterized
over a wide range of temperature and pressures.294–297 . For other applications involving, for
instance, aqueous test fluids, mineral oil could be a suitable piston fluid.
The structure of our device ensures a linear gradient between inlet and outlet pressures and
temperature. The user controls the range, effectively setting the scale of both axes on the phase
diagram of interest, as well as the corresponding resolution. We first demonstrate application over
a wide range of pressures and temperatures, P = 0.89 ± 0.013 MPa – 6.75 ± 0.013 MPa and T =
20.9 ± 0.23 ºC – 89.8 ± 0.23 ºC, respectively (error bars are representative of instrument
uncertainty and fluctuations observed over time). As shown in Figure 2a, this broad, industrially
relevant pressure-temperature range was chosen to encompass a broad region of the two-phase
envelope of the mixture. A complete set of microscope images of each microfluidic chamber were
collected in sequence immediately after sample loading over the course of 3 min. These images
were processed similarly (contrast enhanced, cropped, and realigned) and for cases where two
phases were present (two-phase envelope), images were overlaid with a red or blue filter based on
phase state as determined through semi-automated analysis (see Figure S3 in Supporting
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Information image processing example and protocol). All images were positioned within a grid at
coordinates corresponding to their pressure-temperature location to form the mosaic in Figure 2a.
Enlarged regions of interest (i-vii) are shown to highlight typical test fluid within key areas of the
phase diagram. For all pressure and temperature cases, the test mixture was successfully isolated
by ethylene glycol within individual chambers. The piston extension length, and the volume of the
test fluid, is dependent on applied pressure as well as the state of the test fluid. The two-phase
envelope, highlighted here with a red/blue filter overlay, is rich in thermodynamic information.
Within the two-phase envelope, vapor bubbles (red) increase in size from left to right (low
temperature to high temperature). At low pressures, a dark hue can often be seen within the vapor
regions, indicative of the presence of a liquid film along the chamber wall. Figure 2b shows a color
map of the vapor volume percentage within the two-phase region, as obtained through image
analysis. Such analysis was performed by measuring the liquid region area, and dividing by the
total confined area. Given that the microfluidic chambers are shallow, the area ratio is taken as a
measure of the volume ratio – the role of edge “rounding” and film formation is minimal. Whereas
identification of the presence of both liquid and vapor phases is straightforward at the center of the
envelope, measurements are more challenging near the bubble point line (where bubbles are small)
and dew point line (where the liquid region is confined to a thin film). Whereas quantifying the
precise liquid content (vol. %) at these locations was not feasible, optical microscopy was fully
capable of discerning the location of the dew/bubble point boundary, which is of interest here.
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Figure 2. Wide-range pressure and temperature mapping of the phase diagram of an 80.0% propane + 20.0% methane
mixture. a) Image mosaic of an array of microfluidic chambers, each light-corrected, contrast-enhanced, cropped,
realigned and overlaid with a red or blue filter based on phase. Enlarged areas of interest are also shown. The test fluid
nearest the critical point is overlaid with a yellow filter. b) Liquid content within the two-phase envelope. c) Measured
phase envelope boundary positions, and points intersecting 25, 50 and 75 % liquid volume lines, as well as curve fits
and SRK model.
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Figure 2c shows how the liquid fill percentage data can be further analyzed to yield the
thermodynamic two-phase envelope with liquid volume lines. The red points here are plotted along
the boundary of the envelope in Figure 2b, at a temperature point half-way to the next microfluidic
chamber along the same pressure row. The error bars represent thus not only the experimental
uncertainty based on pressure and temperature measurements, but also the temperature difference
between two adjacent microfluidic chambers – our resolution limit. Determination of liquid
volume lines, however, is not bound by this resolution limit, as all liquid content data points along
a single pressure are fit to a curve, and the precise location of 25 %, 50 % and 75 % liquid volume
line intersects determined. The error bars on these liquid volume line points (green, magenta, cyan)
are based on the standard deviation of these fits, as wells as the experimental uncertainty based on
pressure and temperature measurements, as above. These positions were then fit to low-degree
polynomials to obtain dew point, bubble point and liquid volume lines. To determine the critical
point, the 50 % liquid-volume points nearing critical were fit to a first order polynomial function.
The intersection point between this line (magenta) and the phase envelope boundary (red)
corresponds to the measured critical point. The 25 % and 75 % liquid volume lines were fitted to
a low degree polynomial with a forced intersect to the critical point (further details in Table S2 in
Supporting Information).
To validate the measured phase diagram, we used a Soave-Redlich-Kwong equation of state (SRK)
model, widely used in industry to analyze the phase state of hydrocarbon mixtures,298,299 which
yielded the dashed blue curve on Figure 2c. Overall, a standard deviation of 0.13 MPa was obtained
between measurement and model for the dew and bubble point lines. The critical point was 6.07
MPa and 82.1 ºC, which is near the modeled critical point of 5.86 MPa and 82.7 ºC, respectively.
As is typical for instrumentation, our critical point resolution depends on the full scale. In the
Figure 2 example the full scale is very large (7 MPa and 90 ºC).
To demonstrate the device’s ability to zoom-in to region of interest within the phase diagram,
improve resolution, and demonstrate reproducibility, a series three consecutive mapping
experiments were performed around the critical point. Similar pressure and temperature ranges
were applied in each run, and collectively span the narrowed ranges of 5.63 ± 0.013 MPa – 6.52 ±
0.013 MPa and 68.9 ± 0.23 ºC - 88.2 ± 0.23 ºC. Figure 3a shows a mosaic of microfluidic chambers
for one of the three runs, illustrating fluid behavior near the critical point. In contrast to the Figure
2a mosaic, the zoom-in better resolves the fluid behavior near the critical point. The pressure
117
difference between rows in Figure 2 is an order of magnitude smaller than in Figure 3. The phase
trends are immediately visible, such as the bubble size increasing as temperature is increased left
to right.
Figure 3. a) Narrow-range pressure and temperature mapping of the phase diagram of an 80.0 % methane + 20.0 %
propane mixture at the critical point. a) Image mosaic of an array of microfluidic chambers each light-corrected,
contrast-enhanced, cropped, realigned and overlaid with a red or blue filter based on phase state. Yellow highlights
the supercritical chamber nearest the critical point. b) Liquid content of microfluidic chambers within the two-phase
envelope. c) Measured phase envelope boundary positions, and points intersecting 25 %, 50 % and 75 % liquid volume
lines, as well as corresponding curve fits for a series of three identical experiments, each performed with the same
device. SRK model is also shown.
Figure 3b shows the measured phase envelope boundary locations and the fit envelope, 25 %, 50 %
and 75 % liquid volume line points and liquid volume lines, as well as curve fits for all three
consecutive mapping experiments combined. The experimental envelope and critical point are
compared to the SRK model. With the heightened resolution provided by the zoom-in, the
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measured critical point of 5.90 MPa and 81.8 ºC is now offset by 0.04 MPa and 0.25 ºC from the
model prediction. All three runs led to liquid volume and phase envelope boundary points that
were closely matched, which further indicates the reproducibility of the device.
Overall, the phase diagram mapping device provides flexibility, with the option to map the mixture
phase over a wide range of temperatures and pressures at once, or to zoom-in to a narrower region
of interest with high resolution. It is important to note that while the thermodynamic phase data
measured are time-independent, the data should be collected quickly to limit any potential for
dissolution of a mixture component within the piston fluid (see Figure S4 in Supporting
Information). Potential for solubility is greatest at high pressures. For the highest pressures, the
bubble point line can deviate from the initial measurement if left over time. In practice, we first
established a stable temperature gradient and uniform filling of the mixture in gas phase throughout.
Then we pressurized with the piston fluid, and immediately scanned the chip. The pressure and
phase data stabilized rapidly, within a few seconds, and the resulting bubble point line agreed well
with theory (Figure 2), and showed no more deviation than that the dew point line (unaffected by
solubility).
In conclusion, full thermodynamic phase envelope mapping of a fluid mixture has been
demonstrated by precisely varying temperature and pressure over 1,000 liquid piston-isolated
microfluidic chambers simultaneously. The approach is tunable, offering both wide and narrow
condition ranges while resolving the full suite of phase envelope data: dew and bubble point lines,
liquid volume lines and the critical point. Measurements closely matched expected results based
on an established equation of state model, with a standard deviation of 0.13 MPa between
measurement and model for the dew and bubble point lines, and a difference of 0.04 MPa and 0.25
ºC between measurement and model for the critical point. In stark contrast to the established state
of the art (serial point-by-point measurement) this method provides full characterization of the
phase envelope of a fluid mixture over the pressure-temperature plane of interest, informing a
broad range of chemical applications where fluid mixtures are the norm.