-
Assessment of a Pulsed Electric Field based Treatment for Marine
Microalgae Cell Disruption
Diana Alexandra Santana da Silva Fernandes
Thesis to obtain the Master of Science Degree in
Biological Engineering
Supervisors: Prof. Drª Maria Manuela Regalo da Fonseca MSc,
Gerard ‘t Lam
Examination Committee
Chairperson: Prof. Dra Helena Maria Rodrigues Vasconcelos
Pinheiro Supervisors: Prof. Drª Maria Manuela Regalo da Fonseca
Members of the Committee: Dr. Alberto José Delgado dos Reis
June 2015
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In loving memory of my Grandfather who passed away the day I was
accepted in the internship and
was not able to see me graduate.
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Acknowledgements
Closing my 7-month internship at Bioprocess Engineering Research
Group (BPE, Wageningen
University, Netherlands), I would like to express my never
ending gratitude to all who made it possible.
First and foremost, I would like to thank René Wijffels for
granting me with this beyond valuable
opportunity to develop my thesis at BPE.
I would like to thank my supervisors Gerard ‘t Lam, Giuseppe
Olivieiri (BPE, Wageningen University)
and Professor Manuela Fonseca (Institute for Bioengineering and
Biosciences - IBB, IST). Gerard,
thank you for all the continuous motivation, guidance and time
spent explaining things clearly and
simply. Giuseppe, thank you for all the learning experiences
that challenged me to think outside the
box when approaching and trying to understand different
problems. Professor Manuela, thank you for
all the good advice that helped improve my thesis, for all the
patience and engagement, despite the
physical distance.
I also would like to thank my fellow students at BPE for
providing me with such a friendly environment
where we all could learn something with each other, from cooking
recipes to lab tricks and tips.
Herein, a special thank to Lucia not only for the great
companionship while we were both PEFing
around but for all the help and support given whenever needed
and regardless the circumstances.
Above all, I would like to thank my family for always
encouraging me in every single decision, this one
included. Dad, I cannot thank you enough for always doing more
than you should. I look up to you in
all your different roles, first as a Father, then as a Mentor
and finally as a Professor. I hope you are
able to trace that in this work.
Last but not least (by all means), I would like to thank (more
congratulate) Filipe for doing a
remarkable job putting up with me on a daily basis, even when I
could not stand myself. I am forever
grateful for having shared this experience with you, and I know
many more like this will follow.
Thank you all! Dank an alle! Grazzie a tutti! Obrigada a
todos!
Diana, June 2015
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Resumo
A exploração acelerada de recursos fósseis e o seu impacto
lesivo no ambiente tem estimulado a
investigação de fontes alternativas de matéria-prima renovável
tais como as microalgas. Contudo, o
seu cultivo apenas para a obtenção de percursores de (bio)
combustível não compensa o elevado
investimento inicial nem os custos operacionais associados a
este processo. Uma forma de contornar
este problema prende-se com a valorização dos restantes produtos
intracelulares que as microalgas
podem proporcionar. Para que estes possam ser recuperados sem
que as suas propriedades
funcionais sejam afetadas é necessário um método de rutura
celular energeticamente eficiente e
suave que permita em simultâneo elevados rendimentos de
extração.
Na presente tese, Pulsed Electric Field (PEF) é avaliado como
potencial solução, com o objetivo de
recuperar proteínas solúveis e pigmentos (carotenoides e
clorofila) de duas espécies marinhas nunca
antes testadas com PEF, A e B. Efetuou-se um estudo de prova de
conceito assim como uma análise
detalhada sobre o efeito de parâmetros elétricos e de processo
de modo a otimizar a eficiência de
PEF. Além disso, um novo protocolo de Pré-tratamento foi
desenvolvido de forma a permitir a
aplicação de PEF para ambas as estirpes marinhas. Finalmente,
realizou-se pela primeira vez um
estudo comparativo entre PEF e Bead milling com base em
rendimentos de proteína solúvel e
respetivo consumo de energia.
Ao contrário de A, B não resistiu ao Pré-tratamento, e como tal
não se qualificou para PEF. O
rendimento máximo de proteína solúvel obtido com PEF para A foi
de 13%, não se tendo observado
libertação de pigmentos, o que sugere que PEF não foi capaz de
provocar a rutura desta espécie. No
entanto, foi possível melhorar a eficiência de PEF através da
redução da energia específica por
quarenta vezes mantendo o mesmo rendimento máximo de proteína
solúvel. Em comparação com
Bead milling, PEF revelou não ser ainda competitivo, exigindo um
valor de energia específica vinte
vezes superior para um rendimento de proteína solúvel quatro
vezes inferior. Algumas
recomendações são também propostas de modo a otimizar PEF como
método de rutura celular de
microalgas marinhas.
Keywords: Pulsed Electric Field, Bead milling, Rutura celular de
microalgas, Microalgas marinhas, A,
B.
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Abstract
The rampant exploitation of fossil based resources and its
negative environmental impact is triggering
the demand for renewable feedstock alternatives, such as
microalgae. However, cultivating
microalgae only to obtain (bio)fuel precursors does not offset
the demanded investment and
operational costs. One way of circumventing this problem is by
extending the extraction process to
other microalgae added-value intracellular compounds. To recover
them, a mild and energy-efficient
cell disruption method is required.
In this thesis, Pulsed Electric Field (PEF) is assessed as such
method, aiming at intracellular water
soluble proteins and pigments from two never tested marine
strains, A and B. A Proof-of-Concept is
performed along with an in-depth study on the effect of process
and electrical parameters towards
PEF efficiency improvement. Furthermore, a novel Pre-treatment
protocol is developed to enable the
application of PEF to both marine strains. Finally, a
comparative analysis between PEF and Bead
milling is performed for the first time based on protein yields
and respective energy consumption.
Unlike A, B could not withstand the Pre-treatment, so it did not
qualify for PEF. The maximum protein
yield achieved with PEF for A was 13% but no pigment release was
observed, providing evidence that
PEF was not able to disrupt this species. Nonetheless, it was
possible to improve PEF efficiency by
reducing the theoretical energy input by fourty-fold while
maintaining the same maximum protein yield.
Compared to Bead milling, PEF revealed not to be yet
competitive, requiring a twenty-fold higher
energy input range for a four-fold lower protein yield.
Follow-up studies and recommendations are
presented to enhance PEF process for marine microalgae
intracellular content recovery.
Keywords: Pulsed Electric Field, Bead milling, Microalgae cell
disruption, Marine microalgae, A, B.
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Contents
ACKNOWLEDGEMENTS III
RESUMO V
ABSTRACT VII
CONTENTS IX
LIST OF FIGURES XIII
LIST OF TABLES XVII
LIST OF ABBREVIATIONS XIX
LIST OF NOTATIONS XX
Chapter 1 1
INTRODUCTION 1
1.1 Introduction 3
1.2 Objectives of this thesis 4
1.3 Thesis Outline 5
Chapter 2 7
THEORETICAL BACKGROUND 7
2.1 Microalgae 9
2.1.1 Morphology and Biodiversity 9
2.1.2 Potential of Microalgae… 10
2.1.2.1 …as whole cells 11
2.1.2.2 …extracellular compounds 11
2.1.2.3 …intracellular compounds 12
2.2 Microalgae Cell Disruption Methods 14
2.2.1 Non-Mechanical Cell Disruption Methods 15
2.2.1.1 Chemical Cell Disruption 15
2.2.1.2 Enzymatic Cell Disruption 16
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2.2.1.3 Physical Cell Disruption 17
2.2.2 Mechanical Cell Disruption Methods 17
2.2.2.1 Solid Shear Cell Disruption 18
2.2.2.2 Liquid Shear Cell Disruption 19
2.2.2.3 Other 20
2.3 Pulsed Electric Field 21
Chapter 3 29
MATERIALS AND METHODS 29
3.1 Study Design 31
3.2 Strains and Culture conditions 32
3.3 Pre-treatment step 33
3.3.1 Washing Treatment 33
3.3.2 Concentration Treatment 34
3.4 PEF Treatment 34
3.4.1 PEF Experimental Set-up 34
3.4.2 Experimental Conditions 38
3.4.3 “Before PEF” Sample Preparation and Sample Handling after
PEF 41
3.4.4 Temperature Control 41
3.5 PEF versus Bead milling experiment 41
3.6 Analytical Methods 42
3.6.1 Conductivity 42
3.6.2 Protein analysis 42
3.6.2.1 Water soluble Protein Content 42
3.6.2.2 Total Protein Content 43
3.6.3 Pigment analysis 43
3.7 Calculations 44
3.7.1 Pre-treatment Step 44
3.7.2 PEF treatment 44
Chapter 4 45
RESULTS AND DISCUSSION 45
4.1 Pre-treatment Step 47
4.1.1 Washing treatment 47
4.1.2 Concentration treatment 50
4.2 PEF Treatment 51
4.2.1 Effect of Biomass Concentration 51
4.2.2 Effect of the Electrical Parameters 56
4.2.3 Effect of the Temperature 61
4.3 PEF versus Bead milling 63
Chapter 5 65
CONCLUSIONS AND RECOMMENDATIONS 65
5.1 Conclusions 67
5.2 Recommendations 69
5.2.1 In a short run… 69
5.2.2 In a long run… 69
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REFERENCES 71
APPENDIX 77
A1. Pre-culture and Reactor Medium Conditions 79
A2. A Reactor Set-Up 79
A2.1 Light Calibration Curve 79
A2.2 Pump Calibration Curve 81
A2.3 Daily Analysis 81
A2.3.1 Optical Density 81
A2.3.2 Quantum Yield 82
A2.3.3 pH 83
A2.3.4 Dry weight 83
A3. B Reactor Set-Up 84
A3.1 Light Calibration Curve 84
A3.2 Pump Calibration Curve 85
A3.3 Daily Analysis 85
A3.3.1 Optical Density 85
A3.3.2 Quantum Yield 86
A4. Modified Lowry Protein Assay Calibration Curve 87
A5. PEF using Freeze Dried Biomass 87
A6. Pigment Analysis for fresh Biomass 90
A6.1 …Varying Biomass Concentration 90
A6.1 …Varying Electrical Parameters 91
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List of Figures
Figure 2.1 – Schematic structure of a prokaryotic [39]
and an eukaryotic microalgal cell [40]
. .................. 9
Figure 2.2 – Protein content of different feed and food sources
(in black) against microalgae (in
green). Reproduced based on [44]
..........................................................................................................
12
Figure 2.3 – Carotenoid contents of raw carotenoid-rich fruits
and vegetables, petals of marigold
flowers and some microalgae (blue box). Adapted from [52]
.
.................................................................
14
Figure 2.7 – Classification of cell disruption methods.
Reproduced based on [2, 3]
. .............................. 15
Figure 2.8 – Extraction of total carotenoids into acetone from
processed Haematococcus biomass. (1)
Control; (2) Autoclave (physical, Non-Mechanical); (3) HCl 15
min (chemical, Non-Mechanical); (4)
HCl 30 min (chemical, Non-Mechnical); (5) NaOH 15 min (chemical,
Non-Mechanical); (6) NaOH 30
min (chemical, Non-Mechanical); (7) Enzyme (enzymatic,
Non-Mechanical); (8) High Speed
Homogenization (solid-shear, Mechanical) [36]
;
.....................................................................................
17
Figure 2.9 – Bead milling typical experimental set-up [3]
.
......................................................................
18
Figure 2.10 – High speed homogenizer typical experimental set-up
[67]
. .............................................. 19
Figure 2.11 – High speed homogenizer typical experimental
set-up. ................................................... 20
Figure 2.12 – Lipid bilayer rearrangement in a cell membrane
subjected to an electric field (E) of
increasing strength. ER corresponds to the minimum electric
field required for pores formation. A. The
cell membrane has a potential TMPm and it is being charged to a
potential TMP > > TMPm causing its
thinning; B. The membrane has reached breakdown potential TMPR
with resulting reversible pore
formation; C: Pore enlargement resulting in irreversible
membrane breakdown. [24, 34]
........................ 22
Figure 2.13 – Examples of batch and continuous electroporation
chambers. A. Lab-scale batch
electroporation chamber (Micro and Milli-electrodes);
Flow-through chamber with a B. coaxial
geometry, C. a collinear geometry and D. a planar geometry. Red
and blue regions match opposite
electrodes. [35]
........................................................................................................................................
24
Figure 2.14 – A. Exponential, B. square and C. bell wave pulses.
[35]
.................................................. 24
Figure 3.1 – Study design schematic representation.
...........................................................................
31
Figure 3.2 – Washing treatment schematic representation.
..................................................................
33
Figure 3.3 - Concentration treatment schematic representation.
.......................................................... 34
Figure 3.4 – A. Batch PEF system used and its components. B. PEF
cuvettes used (Gene Pulser
MicroPulser Cuvettes 165-2082, BIO-RAD, USA). The PEF chamber
presents two parallel-plate
aluminum electrodes with two different gap widths, 2 mm (400 μL
volume capacity) – green cap – and
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4 mm (800 μL volume capacity) – blue cap – housed in a
sterilized polycarbonate container. C. Layout
on how the PEF chamber fits the SchokPodTM
chamber.
.....................................................................
35
Figure 3.5 – Equivalent electric circuit for PEF. A) Equivalent
circuit of the pulse forming circuit; B)
When the sample is conductive an equivalent resistance can be
defined in parallel with . .......... 36
Figure 3.6 - Time response of the RC parallel circuit. A)
General example for two different values of
time constant ; B) Square wave produced as a chopped exponential
with long time constant. ....... 37
Figure 3.7 – Transmembrane potential TMP induced by the electric
field, adds on to the initial
transmembrane potential TMPm. Breakdown potential TMPR is first
reached at the pole, where
potential vectors, TMP and TMPm point to the same direction.
.............................................................
39
Figure 3.8 – Time dependence of the applied voltage V(t) and
corresponding cell response TMP(t), for
a situation where the cell rupture potential TMP is exceeded.
Typical values are TMPmax = 4.5 V for V0
= 3000 V.
...............................................................................................................................................
39
Figure 3.9 – Expected temperature variation along the PEF
treatment. ............................................... 41
Figure 4.1 – Conductivity measured in the supernatant initially,
after one and two washing cycles with
Milli-Q water in A. linear and B. logarithmic scale, for A
(black bars) and B (brown bars) cells
suspensions. All measurements were performed in duplicate
(technical) and error bars represent
standard deviations. The concentration in the reactor at
steady-state was 1.15 cX for A and 1.73 cX for
B.
...........................................................................................................................................................
47
Figure 4.2 – A. Total protein content %DW, of A (black solid
bars) and B (brown solid bars)
determined in replete conditions. Measurement performed in
triplicate (technical). Protein content
%(DW), released in the supernatant, after one and two washing
cycles for both species. All
measurements were performed in duplicate (technical) and error
bars represent standard deviations.
B. Percentage of total protein loss in the supernatant after one
and two washing cycles, for both
species...................................................................................................................................................
48
Figure 4.3 - A. Conductivity, B. Pigments concentration and C.
Protein content in %(DW) measured in
the supernatant, after the concentration step. All measurements
were performed in duplicate
(technical) and the error bars represent standard deviations. D.
Percentage of total protein loss in the
supernatant after one washing cycles and the concentration step
for both species. ............................ 51
Figure 4.4 – Conductivity measured in the supernatant before
(dotted bars) and after applying PEF
(solid black bars) for different concentrations. All experiments
were performed in duplicates (technical)
and the error bars represent standard deviations.
................................................................................
53
Figure 4.5 – A. Total protein content in %DW (striped bars), of
A determined in replete conditions.
Measurement performed in triplicate (technical). Protein content
in %DW released in the supernatant,
before (dotted bars) and after PEF treatment (black solid bars).
All experiments were performed in
technical duplicates and the error bars represent standard
deviations. B. Percentage of total protein
recovered in the supernatant before and after PEF treatment.
.............................................................
54
Figure 4.6 – Voltage (v) forms when the sample resistance is
much smaller than the electroporator
resistance . A. The applied voltage at the electrodes terminals
decreases significantly during the
duration of the pulse length (10-3
t) (vertical red dashed line). B The response of the
transmembrane
potential TMP(t) consequently descreases during the pulse
length, possibly under the TMPR value
thus reducing the probability of membrane rupture.
..............................................................................
54
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Figure 4.7 – Specific energy input computed according to
Equation (4.1) as a function of biomass
concentration.
........................................................................................................................................
55
Figure 4.8 – Conductivity measured in the supernatant before
(dotted bars) and after applying PEF for
0.005 t and 0.0005 t (solid brown and black bars). All
measurements were performed in duplicate
(technical) and the error bars represent standard
deviations................................................................
57
Figure 4.9 – Total protein content in %DW of A determined in
replete conditions (striped bars).
Measurement performed in triplicate (technical). Protein content
in %DW released in the supernatant,
before (dotte bars) and after PEF treatment (solid brown and
black bars). All measurements were
performed in duplicate (technical) and the error bars represent
standard deviations. .......................... 58
Figure 4.10 – A. Conductivity and B. Water soluble protein
yields after applying PEF, as a function of
the number of pulses for A, after applying 7.5 e and 0.005 t
(black solid line), 7.5 e and 0.0005 (grey
solid line), 15 e and 0.005 t (brown solid line), 15 e and
0.0005 t (green solid line). ............................ 58
Figure 4.11 – Water soluble protein yields as a function of
specific energy input computed according to
Equation (4.1).
.......................................................................................................................................
60
Figure 4.12 –Temperature variation, ΔT, after PEF treatment for
A. different concentrations at fixed
electrical parameters (15 e, 10 pulses, 0.005 t). and for B.
different electrical parameters at fixed
biomass concentration (14 cX), after applying 7.5 e and 0.005 t
(black solid line), 7.5 e and 0.0005
(grey solid line), 15 e and 0.005 t (brown solid line), 1 e and
. t green solid line)., C. ........... 62
Figure A1 – A. Top and B. side view of the reactor positions for
light intensity measurement. C. Light
calibration curve obtained for A reactor.
................................................................................................
80
Figure A2 – Volumes measured for a fixed time period of 10
minutes and respective pump calibration
curve obtained.
......................................................................................................................................
81
Figure A3 – OD daily analysis for A reactor run.
...................................................................................
82
Figure A4 - QY daily analysis for A reactor run.
....................................................................................
82
Figure A5 – DW (cx) - OD curve obtained for A reactor.
.......................................................................
83
Figure A6 – Light calibration curve obtained for B reactor.
...................................................................
84
Figure A7 – Volumes measured for a fixed time period of 10
minutes and respective pump calibration
curve obtained.
......................................................................................................................................
85
Figure A8 – OD daily analysis for B reactor run.
...................................................................................
86
Figure A9 - QY daily analysis for B reactor run.
....................................................................................
86
Figure A10 – Modifield Lowry Protein Assay Calibration curve
obtained using BSA as a standard
representing OD750 as a function of cprot.
...............................................................................................
87
Figure A11 – A. Conductivity, B. Total protein content %DW of A
and B sp determined in replete
conditions (striped bars). Measurement performed in triplicate
(technical). Water soluble protein
content %DW released in the supernatant, C. Carotenoids and D.
Chlorophyll before PEF treatment
for 7 biological multicates. It is possible to observe that only
the freeze-drying process already partially
damages microalgal cells. The water soluble protein detected in
the supernatant before applying PEF
for P. sp is more than 50% of the total protein content of this
species. This also provides evidence that
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A is more resistant than P. sp due to the lower loss of proteins
(less than 50%) in the supernatant
before PEF is applied.
...........................................................................................................................
87
Figure A12 – A. Conductivity, B. Total protein content %DW of A
and B sp determined in replete
conditions (striped bars). Measurement performed in triplicate
(technical). Water soluble protein
content %DW, C. Carotenoids and D. Chlorophyll released in the
supernatant before and after PEF for
different post-pulse incubation times (0-3h) ( = 7.5 e ; = 2; =
0.005 t) The post-pulse
incubation time seems not to affect the release of ions, water
soluble proteins and pigments in the
supernatant for the tested range.
..........................................................................................................
88
Figure A13 – A. Conductivity, B. Total protein content %DW of A
and B sp determined in replete
conditions (striped bars). Measurement performed in triplicate
(technical). Water soluble protein
content %DW, C. Carotenoids and D. Chlorophyll released in the
supernatant before and after PEF for
different electric fields (7.5 e, 15e) ( = 2; = 0.005 t). There
was no effect of the electric field in
the release of ions, water soluble proteins and pigments in the
supernatant for the tested range ....... 89
Figure A14 – A. Conductivity, B. Total protein content cprotDW
of A and B sp determined in replete
conditions (striped bars). Measurement performed in triplicate
(technical). Water soluble protein
content cprotDW, C. Carotenoids and D. Chlorophyll released in
the supernatant before and after PEF
for different number of pulses (2, 10) ( = 7.5 e ; = 2; = 0.005
t). There was no effect of the
number of pulses in the release of ions, water soluble proteins
and pigments in the supernatant for the
tested range.
..........................................................................................................................................
89
Figure A15 – A. Conductivity, B. Total protein content %DW of A
and B sp determined in replete
conditions (striped bars). Measurement performed in triplicate
(technical). Water soluble protein
content %DW, C. Carotenoids and D. Chlorophyll released in the
supernatant before and after PEF by
pre-cooling the sample for 5 minutes before applying PEF
treatment (2, 10) ( = 7.5 e ; = 2; =
0.005 t). It seems that there is no negative or positive effect
of this pre-cooling in the recovery of the
intracellular compounds.
........................................................................................................................
90
FigureA16 – Release of pigments in the supernatant before and
after applying PEF at = 15 e ;
= 10; = 0.005 t)
.................................................................................................................................
91
Figure A17 – Release of pigments in the supernatant before and
after applying PEF at = 14 cX for
A. 2, B. 10 and C. 18 pulses. The release of pigments increases
with the number of pulses, as
expected, since the conditions become harsher but the amount
released is negligible when
considering the units.
.............................................................................................................................
91
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List of Tables
Table 2.1 - Summary of the main microalgae group features.
..............................................................
10
Table 2.4 – PEF advantages over Non-Mechanical and other
Mechanical microalgae Cell Disruption
Methods based on literature [2-5, 63, 64]
. BM – Bead milling, HSP – High Speed Homogenization, HPH –
High Pressure Homogenization, US – Ultrasounds, MW – Microwave.
............................................... 25
Table 2.5 – Literature review on the use of PEF for microalgae
cell disruption. (T– Temperature; DCW
– Dry Cell Weigth; EF– Electric Field; PL– Pulse Length; f –
frequency; ET – Energy Treatment). ..... 25
Table 3.1 – Lower and upper limits of the adjustable parameters
of the used electroporator. ............. 35
Table 4.1 - values for the initial, after washing 1x and after
washing 2x samples computed
according to Equation (3.3).
..................................................................................................................
49
Table 4.2 – Experimental conditions used to study the effect of
different biomass concentrations on
PEF performance.
.................................................................................................................................
52
Table 4.3 – Specific energy input of PEF treatment computed
according to Equation ((4.1) for both
biomass concentrations with = 2.78 10-7
, = 15 e, = 0.005 t, = 10,
= 0.04198 j, = 0.06675 j................................ 56
Table 4.4 – Experimental conditions used to study the effect of
different electrical parameters on PEF
performance.
.........................................................................................................................................
56
Table 4.5 – Difference in the initial energy input by reducing
the value of the electric field, , and
pulse length, , for a fixed biomass concentration, . was chosen
based on the lowest value
required to assure the higher water soluble protein yield.
.....................................................................
59
Table 4.6 – Comparision of PEF water soluble protein yields
between the only two studies available in
open literature featuring another marine microalga,
Nannochloropsis salina, and this work. ............... 61
Table A1 - Pre-cultutre and Reactor Medium composition for both
species used in this thesis. .......... 79
Table A2 – Light intensity values measured in μmolm2s
-1for all the different positions at different light
intensity percentages.
............................................................................................................................
80
Table A3 – Light intensity values measured in μmolm2s
-1for all the different positions at different light
intensity percentages.
............................................................................................................................
84
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List of Abbreviations
PEF Pulsed Electric Field
PUFA Polyunsaturated Fatty Acid
EPA Eicosapentaeonic
DHA Docosahexaenoic
TAG Triacylglycerides
BM Bead Milling
HSP High Speed Homogenization
HPH High Pressure Homogenization
US Ultrasonication
MW Microwaves
TMP Transmembrane potential
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List of Notations
T Temperature
DCW Dry Cell Weigth
DW Dry Weight
Electric Field
Pulse Length
f Pulse frequency
Number of pulses
σ Conductivity
Biomass Concentration
Energy Treatment
ODXXX Absorbance at a wavelength XXX
Protein concentration
Dilution factor
TMP Transmembrane potential
TMPm Intrinsic transmembrane potential
TMPR Critical transmembrane potential
TMPmax Settling transmembrane potential
PEF Treatment time
Theoretical PEF treatment time
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Chapter 1
Introduction
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3
1.1 Introduction
The European Environment Agency states on their State and
Outlook 2015 report that “Oil still
accounts for the largest share in total primary energy supply
(31%) followed by coal (29%) and natural
gas 21%)” [1-5]
. The continued exploitation of fossil based resources and its
negative environmental
impact is triggering the demand for renewable feedstock
alternatives for sustainable fuel production [6,
7]. Terrestrial crops such as sugarcane, maize, palm oil, rape
seed, beat corn and soy have already
been pointed out as potential candidates to meet this challenge,
due to their ability to produce (bio)fuel
precursors. However, the energy input required for this process
is even more demanding as the one
required for fossil fuels production [8, 9]
. In addition, the competition for arable land and raw
materials
also suitable for the food industry raises an ethical dilemma
between using it as a food or a different
energy supply (food vs. fuel) [6, 7, 10-12]
.
To overcome these limitations a great deal of research is being
channelled to microalgae as a
possible alternative for (bio)fuel production. Their (i) high
triacylglycerides (TAG) content, (ii) their
ability to recycle nutrients from waste water (N, P), (iii) to
uptake carbon dioxide from air emissions
and (iv) to thrive in non-arable environments, sometimes within
extreme pH and salt conditions, render
them attractive to fulfil this purpose [9, 13]
. Despite these encouraging features, feasibility studies
reveal
that cultivating microalgae only to produce biofuel does not
offset both investment and operational
costs [14, 15]
. One possible way of addressing this problem is by extending
the extraction process not
just to lipids but to other intracellular compounds that
microalgae are able to provide [10, 11, 15]
.
Besides lipids, microalgae also host proteins, pigments,
antioxidants, vitamins and sugars that are
valuable for food, feed and pharmaceutical industries. Wijffels
et al.[15]
claim that after biorefining,
these compounds are able to retrieve up to 1496 € ton-1
Algal Biomass, while biofuels are just able to
contribute with 150 € ton-1
Algal Biomass. By replacing the “fuel-only” approach with a
microalgae
biorefinery concept, the value of the same biomass increases by
approximately ten times.
This microalgae biorefinery concept generally comprises four
steps: harvest, cell disruption, product
extraction and product purification [6]
.
To accomplish a high product recovery while preserving its
quality after biorefining, the second stage -
cell disruption - plays a substantial role. Conventional cell
disruption methods based on non-
mechanical and mechanical treatment usually require harsh
conditions such as high temperatures and
towering pressures that only allow the selective extraction of a
specific component while damaging the
remaining ones [3, 6, 16]
. The use of such severe conditions also impacts the energy
input resulting in
higher operating costs [3, 6]
. Thus, novel cell disruption technologies that are milder and
less energy
intensive need to be studied.
One potential candidate that might feature the above
requirements is Pulsed Electric Field (PEF) [17-22]
.
Nowadays, PEF is a widely acknowledged alternative to the
traditional thermal pasteurization
processing of liquid foods. When using PEF, microbial
inactivation is achieved by applying high
intensity, periodic and short duration electric pulses (micro to
milliseconds) to the processed stream,
instead of applying heat [23]
. These pulses cause the formation of irreversible pores in the
cell
membrane of contaminant microorganisms (electroporation). This
results in the deactivation of the
microorganisms while the organoleptic properties of food are
preserved [24-26]
.
The same principle could be applied for microalgae cell
disruption. The underlying mechanism of PEF
over microalgae cell membranes has not been fully elucidated up
until to now. However, a general
-
4
consensus supports the theory that in the presence of an
external electric field, the cell membrane
potential shifts, resulting in the rearrangement of the
phospholipid bilayer and consequent increase in
membrane permeability. This increase in permeability not only is
strain-specific but it also depends on
the fine tuning of particular process parameters like biomass
concentration, sample conductivity, field
strength, number of pulses, pulse shape and duration [24, 25,
27]
.
According to the available literature, PEF has been regarded as
a promising microalgae cell disruption
technique for the recovery of intracellular valuables, such as
proteins and lipids [19-22, 28-32]
. However,
issues concerning the application of the method like species
variety and extraction yields, need to be
tackled before placing PEF among other well-known cell
disruption methods.
The majority of the studies published so far on the use of PEF
for microalgae cell disruption, only
feature freshwater species [19-22, 28-32]
.
The use of freshwater to grow microalgae obviously increases the
water and nutrient footprint [8, 9]
.
Thus, from a process point of view for large scale
implementation, microalgae grown on a marine
environment are preferred. Although marine microalgae seem to be
a good alternative, they entail an
additional hurdle when applying PEF. One of the parameters that
can influence PEF performance is
the sample conductivity. Microalgae grown on a marine
environment present a considerably higher
conductivity (5 Sm-1) than freshwater’s 5-50 mSm
-1)
[33]. The higher the conductivity, the greater is the
energy consumption rendering the process less efficient [24, 25,
34, 35]
.
Besides the marine aspect, the lack of data in the literature
regarding yields of extraction and
respective energy consumption makes it hard to place PEF in
perspective against benchmark
technologies. Before a qualitative conclusion on the feasibility
of PEF can be drawn, these gaps in the
existing literature should be cleared up.
1.2 Objectives of this thesis
This thesis seeks to fill the previously described gaps in the
literature. The aims of this thesis are therefore, to carry out: 1.
A Proof-of-Concept study to aasssseessss tthhee aabbiilliittyy ooff
PPEEFF ttoo ssuucccceessssffuullllyy ddiissrruupptt mmaarriinnee
mmiiccrrooaallggaaee
ssttrraaiinnss..
- Is it possible to use PEF to disrupt marine microalgae? - Do
different marine microalgae strains respond differently to PEF?
2. A pprroocceessss ooppttiimmiizzaattiioonn towards PEF
efficiency improvement.
- Is it possible to maintain the same optimal yields with a
lower energy input?
3. A ccoommppaarraattiivvee aannaallyyssiiss bbeettwweeeenn
PPEEFF aanndd ootthheerr bbeenncchhmmaarrkk
tteecchhnnoollooggiieess, based on protein
extraction yields and respective energy consumption ffoorr
ooppttiimmiizzeedd pprroocceessss ppaarraammeetteerrss..
To attain the first two objectives,, pprroocceessss (sample
conductivity, species cell wall, biomass
concentration) and eelleeccttrriicc (electric field, number of
pulses, pulse length) ppaarraammeetteerrss are explored to
understand their role;
-
5
1.3 Thesis Outline
This thesis is mainly structured in 5 chapters.
Chapter 1 provides an introduction to the research focus and the
objectives of this thesis.
Chapter 2 presents an overview on microalgae, showcasing their
main features, current opportunities
and constraints. The main characteristics and commercial
interest of the marine microalgae species
used in this thesis are also evidenced. In addition, different
microalgae cell disruption methods are
introduced giving special focus to PEF. Since this technology is
the core of the thesis, its
fundamentals, advantages and drawbacks are illustrated as well.
A literature survey on different
studies performed with PEF hitherto, is also presented.
Chapter 3 delineates the study design featuring the approach to
meet the objectives of this thesis. The
strains and culture conditions are characterized and a novel
Pre-treatment step for marine microalgae
before applying PEF is proposed. PEF experimental Set-Up is
presented along with the set of
conditions used varying process (biomass concentration) and
electrical parameters (electric field,
number of pulses, pulse length). The analytical methods
performed and the main calculations used to
process the data acquired are also explained within this
Chapter.
Chapter 4 features the results obtained regarding the impact of
the Pre-treatment and PEF set of
experiments. The effect of process and electrical parameters is
discussed and a process optimization
based on this study is reported. Also, a comparative analysis
between PEF and Bead milling set as a
benchmark technology is established not only to place PEF among
other cell disruption methods but
also as a preliminary assessment of PEF feasibility in a larger
scale.
Chapter 5 presents the main conclusions of this work and
recommendations to improve its outcomes
in a short and long run. Also, suggestions for future research
featuring PEF are also provided.
The Annex contains all supplementary information regarding a
preliminary study using the freeze-dried
marine microalgae species studied in this work, to help
establishing the PEF set-up and protocol. In
addition it is also provided complementary information
concerning microalgae cultivation, PEF related
electrical circuit concepts, analytical methods and results.
Author’s note: The name of the used species and the units of the
experimental conditions ofthis work
have been replaced by variables which are specified in the
Confidential Appendix available upon
request.
-
6
-
7
Chapter 2
Theoretical Background
-
8
-
9
2.1 Microalgae
Microalgae are one of the most ancient and resilient living
structures on Earth. Eons ago, with brazing
temperatures and harsh atmosphere on Earth, these single-cell
microorganisms managed to grow,
absorbing atmospheric gases and reducing CO2 levels to a
fraction of a percent. By doing so,
microalgae pioneered an environment in which higher life forms
could develop [12, 36, 37]
.
Nowadays, microalgae are still at the basis of freshwater and
marine aquatic system food chains,
fuelling a wide variety of organisms including humans. This is
possible due to their ability to
incorporate inorganic carbon and to convert light energy into
chemical building blocks while releasing
oxygen (photosynthesis). In addition, they are also able to
reduce nitrogen enriched substrates (NO3-;
urea) [38]
.
2.1.1 Morphology and Biodiversity
Microalgae do not exhibit roots, stems or leaves. Two typical
prokaryotic (cyanobacteria) and
eukaryotic microalgal cells are depicted in Figure 2.1.
The cell wall composition and rigidity are species-dependent in
both types of cells. Like plant cells,
eucaryotic microalgae also present one or multiple chloroplasts
that host the photosynthetic machinery [36]
.
Figure 2.1 – Schematic structure of a prokaryotic [39]
and an eukaryotic microalgal cell [40]
.
The morphology and the internal structure of a microalgal cell
vary according to the belonging strain
and life stage. Their size, on the other hand, can also depend
on the light and nutrient growing
conditions [36, 37, 41]
.
Taxonomically, microalgae are diverse. Estimates suggest that
over 800,000 species exist, yet only
tens of thousands of these have been described in the literature
[38, 42]
. Their systemic classification is
-
10
primarily based on the pigment composition [37]
. Currently, the following groups are considered: Green
Algae (Chlorophyceae), Red Algae (Rhodophyceae), Diatoms
(Bacillariophyceae), Brown Algae
(Phaeophyceae), Gold Algae (Chrysophyceae), Yellow-green algae
(Xanthopyceae) and Blue algae
(Cyanobacteria) [36-38, 43]
. Table 2.1 showcases their occurrence and key features.
Table 2.1 - Summary of the main microalgae group features.
Main Microalgae Groups Occurence (sp) Environment Special
Features and Applications Ref
Green Algae
≈7
Fresh/Marine
Eucaryote; unicellular or multicellular; contain high
chlorophyll levels and a large amount of proteins; produce starch
and oil under light and nitrogen limited conditions.
[38, 44]
Red Algae
≈
Marine
Eucaryote; multicellular; contain high levels of
phycoerythrin.
[37, 38, 43,
44]
Diatoms
≥ 1 ,000
Fresh/Marine
Eucaryote; unicellular; display a skeleton of silica that fits
together like two halves of a sphere; produces most of earth´s
biomass; regulate their buoyancy by producing oil; indispensable
food source for zooplankton;
[36-38]
Brown Algae
≈1 -2000
Marine
Eucaryote; multicellular; contain high levels of
fucoxanthin.
[36-38]
Gold Algae
≈1
Fresh/Marine
Eucaryote; unicellar; possess flagella used for
displacement.
[36-38, 43]
Yellow-green Algae
≈6
Fresh
Eucaryote; unicellular; produce large amounts of oil.
[36-38, 43]
Blue Algae/Cyanobacteria
2000-8000
Fresh/Marine
Procaryote; unicellular or multicellular; may produce toxins
that can compromise the water quality when in high concentrations;
store food reserves in form of starch.
[36-38, 43]
2.1.2 Potential of Microalgae…
The heterogeneity and versatility of microalgae is also
reflected in their many possible applications as
whole cells or their extracellular or intracellular
metabolites.
Regardless the final end-product, using microalgae over fossil
and terrestrial crops based resources,
consists in a more sustainable option due to their (i) fast
growing nature that is not seasonally limited,
(ii) lower land footprint (no necessity for arable land), (iii)
ability to grow within extreme pH and salt
conditions (seawater) and (iv) from renewable light energy,
carbon and nitrogen enriched substrates [6,
8, 9, 11, 13, 45].
-
11
This section presents an overview on the potential of microalgae
as whole cells and of microalgae
extracellular and intracellular compounds. Despite the many
different applications addressed herein,
not all of them are being commercially exploited at the moment.
To render microalgae competitive,
large scale cultivation systems, genetic engineering tools, and
downstream processing are still being
studied in order to optimize growth conditions, metabolite
productivity and product recovery yields with
the lowest possible energy consumption and thus with the lowest
operating costs [8-10, 45, 46]
.
2.1.2.1 …as whole cells
Microalgae as whole cells are being used as feed in aquaculture,
in human nutrition and in wastewater
treatment [45-47]
.
To counteract the decrease of natural fish catches, to face the
need for more fish as a food source for
the increasing population, and to reduce the pressure on energy
demand and carbon dioxide
emissions in the fishery sector, sustainable aquaculture has to
be developed [46]
. For this to be
possible, a solid food chain has to be established and
maintained until fish is matured to enter the
consumer market. Polyunsaturated fatty acids (PUFA) based feed
can be a solution since it
constitutes a key growth factor [45, 46]
. Eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA)
are two different PUFA currently produced as oil fish from
bycatch (fish caught unintentionally when
going for another fish) [46]
but marine microalgae are regarded as the primary producer of
these two
PUFA in marine food webs [45]
. Replacing oil fish by microalgae in aquaculture feed, would
not only
minimize the sustainability problem but also render superior the
quality/stability of the provided PUFA,
since in this case, these are encapsulated by the algal cell
wall. Furthermore, it is noteworthy that
microalgae are naturally rich in antioxidants that help
preventing lipids oxidation [45-47]
.
These antioxidants, such as carotenoids and phenolic compounds,
together with EPA and DHA are
also claimed to have a positive effect on the reduction of
cancer and heart coronary diseases
incidence. DHA is also important for brain and retinal tissues
development of human embryos and
newborns. It is absent in cow milk so it is considered a
valuable supplement to baby food [46, 47]
.
Microalgae can also be used for waste water and flue gas
treatment due to their ability to uptake
carbon dioxide, sulphur, nitrogen and phosphorus oxides and thus
purify these waste streams [9, 10, 13,
46].
2.1.2.2 …extracellular compounds
Some of the extracellular products derived from microalgae are
also commercially interesting [46]
.
Because these metabolites are excreted, no energy has to be
invested to disrupt the rigid cell walls
and so the process becomes easier to translate into a larger
scale [46]
. In addition, the purification
process is simpler since there are no cell debris hampering the
extraction of these products [9, 15, 46]
.
Microalgae mainly excrete their metabolic end-products such as
toxins, antibacterial substances,
polysaccharides and hydrocarbons. These toxins can have
anti-cancer properties whereas
antibacterial substances can be used as antibiotics. Chlorella
pyrenoidosa and Phaeocystis pouchetii
-
12
are two microalgae that are able to provide such compounds, but
both offer low metabolite productivity [44]
.
Polysaccharides may act as precursors of gelling agents,
thickeners, stabilizers and emulsifiers while
hydrocarbons can be used as a feedstock for fuels and plastics
[44, 46]
. Botryococcus braunii is an
example of a microalgae species that is able to produce
hydrocarbons up to 80% of its biomass dry
weight. However, low growth rates and hydrocarbon biosynthesis
are slowing the implementation of
this microalga as an alternative source for fuels and plastics
[46]
.
2.1.2.3 …intracellular compounds
Microalgae usually accumulate intracellular compounds such as
proteins, carbohydrates and lipids
(membrane lipids and triacylglycerides also known as TAG).
Although the biochemical composition of
microalgae varies according to the species and growth conditions
(nutrient, light, temperature and
growth stage) these fractions together make up to 70-90% of the
biomass dry weight [46]
. Under
optimal growth conditions, the protein fraction can go up to 50%
of the total biomass dry weight but
under nitrogen starvation this portion can be reduced, while
increasing carbohydrates and lipids
fractions substantially up to 60-80% [8, 13, 15, 46]
. The remaining percentage matches other constituents
such as pigments, antioxidants, sterols, glycerol, toxins and
minerals (ash).
The following subsections focus more specifically on the
potential of microalgae proteins,
carbohydrates and lipids in the present framework.
PPrrootteeiinnss
Microalgae could be a possible source of protein owing to its
high content under optimal growth
conditions. Also when compared to other feed and food sources
their protein content is comparable or
two times higher as depicted in Figure 2.2 [44, 46]
.
Figure 2.2 – Protein content of different feed and food sources
(in black) against microalgae (in green).
Reproduced based on [44][44][44][44]
.
-
13
Additionally, an improved and valuable feed can be formulated
from microalgae due to their larger
diversity in essential amino acids composition [48, 49]
. Soybean contains all essential amino acids for
animals, but their ratios are not optimal for every type of
feed. A balanced soybean feed thus requires
an extra addition of expensive refined essential amino acids,
which could be provided by algae [13, 46].
CCaarrbboohhyyddrraatteess
Like proteins, carbohydrates also represent a large percentage
of dry weight biomass.
Glucans such as glycogen, starch, cellulose, laminarin and
chrysolaminarin are the most abundant.
These are responsible for the storage of energy when cells are
exposed to suboptimal growth
conditions [50]
. Next to these carbohydrates, also cellulose, which mainly has
a structural function, is
found in larger amounts [46]
.
Due to the elevated carbohydrate content of some of the strains,
these show potential in many
applications, namely for bioethanol production in animal feed
ingredients, gelling agents in foods,
biofertilizers, cation chelators in both food and waste water
treatment and as building blocks for
bioplastics [38]
.
LLiippiiddss
Microalgae contain a broad range of lipids such as sterols, some
vitamins, free fatty acids, TAG,
phosphodiacylglycerides, glycodiacylglycerides and pigments [36,
37, 43, 46]
.
However the commercial relevance of microalgae lipids lies
mainly in TAG for biofuels and pigments [50, 51]
.
Fresh microalgae TAG display a fatty acid composition similar to
vegetable oils which makes the
conversion into an effective and clean fuel possible. This fatty
acid composition also enables the
replacement of edible oils such as palm oil [46]
.
Pigments are molecules that absorb a significant part of the
visual light spectrum. They can be
obtained from petroleum, organic acids and inorganic chemicals
(so called synthetical route) or from
insects, vegetables, fruits, petals of flowers and from
microalgae (so called natural route) [51]
. At
present, producing pigments synthetically is less expensive than
producing them naturally, but the
consumer demand for natural based products is rising [52]
.
Of all possible natural pigment sources, microalgae provide a
large variety of these molecules that can
be grouped in chlorophylls (green), carotenoids (yellow to red)
and phycobiliproteins (red, blue, purple
and yellow) [52]
.
Carotenoids are the most naturally occurring type of pigment
being produced by all photosynthetic and
several non-photosynthetic organisms. Their color ranges from
yellow to pink and it is mainly due to it
that this type of pigments are considered interesting for
industrial applications such as dyes in food,
feed and cosmetics fields [9, 51]
. They are also claimed to have antioxidant properties which
makes
them suitable for nutraceuticals. These kind of pigments can be
produced in concentrations exceeding
those found in higher plants by one or more orders of magnitude
as one can observe in Figure 2.3 [52]
.
-
14
Figure 2.3 – Carotenoid contents of raw carotenoid-rich fruits
and vegetables, petals of marigold flowers and
some microalgae (blue box). Adapted from [52]
.
Currently, microalgae are being commercially exploited for only
three pigments, namely β-carotene,
astaxanthin and phycocyanin [51]
.
2.2 Microalgae Cell Disruption Methods
As described in the previous sections microalgae are able to
provide a large range of products useful
as alternative feedstock for many different industries. Despite
their great potential there are still some
bottlenecks that need to be overcome in order to render
microalgae competitive [8-11, 38, 45]
.
As highlighted before, one of those is the low revenue after
high initial investment costs, only to get a
specific product out of microalgae [6, 11, 15]
. A possible approach to address this problem is by adopting
a biorefinery concept. It consists in fractioning the biomass
(microalgae) not into one, but several
added-value co-products using downstream processes that help
making the whole process more
feasible [6, 7, 11]
. These include harvesting, cell disruption, product isolation
and product purification [3, 5,
6].
To successfully implement this strategy, the downstream
processes used to fractionate microalgae
must be such so that a balance between product recovery yields
and energy input is reached to obtain
a maximum financial profit.
A major challenge in this biorefinery approach is the cell wall
disruption. Microalgae cell walls are
relatively thick and rigid. In addition, most of the valuable
products are located within globules or
bound to cell membranes [3, 5]
. For this reason, a considerable amount of energy has to be
invested
but at the same time the energy applied should be low enough not
to compromise the quality of the
recovered intracellular content. An appropriate cell disruption
method should thus be energy-efficient
and simultaneously mild to ensure a low operating cost, high
product recovery and high quality of the
extracted products.
In general, the available microalgae cell disruption methods can
be categorized in two main groups,
according to their working mechanism of microalgal cell
disintegration (Figure 2.4) – Non-Mechanical
and Mechanical methods [2-5, 62-64]
.
-
15
Figure 2.4 – Classification of cell disruption methods.
Reproduced based on [2, 3]
.
This section presents an overview on the current
state-of-the-art of microalgae Non-Mechanical and
Mechanical cell disruption methods focusing on their
fundamentals, advantages and drawbacks.
Unlike the other cell disruption methods depicted in Figure 2.4,
Pulsed Electric Field will not be
discussed within this section. Since PEF disruption method is
the core of this thesis, an independent
section is dedicated to discuss it thoroughly, including the
underlying mechanism, features,
applications, advantages, drawbacks/challenges, and microalgae
PEF case-studies can be found after
the present one.
2.2.1 Non-Mechanical Cell Disruption Methods
Non-Mechanical methods usually involve cell lysis with chemical
agents, enzymes or physical
phenomena such as osmotic shock [2, 3, 5]
. These methods are not as harsh as mechanical processes
since cells are only perforated or permeabilized rather than
being completely destroyed. Additionally,
the energy required for these non-mechanical methods to act is
also lower, resulting in a lower energy
input [2]
. Further, these methods are recognized to be more selective
than mechanical methods owing
to their specific interaction either with the cell wall,
weakening it, or with membrane components,
affecting their function as barriers, causing products to leak
[62-64]
.
2.2.1.1 Chemical Cell Disruption
Cell disruption can be induced by a great variety of chemical
compounds that usually act specifically
on the cell wall. However, the selectivity, suitability,
efficiency and cell disintegration mechanism of
these compounds are dependent on the cell wall composition of
the microorganism [2, 3, 5]
.
-
16
Hitherto, the chemical agents tested for microalgae cell
disruption are mainly surfactants, oxidizing
agents (ozone, chlorine), organic solvents (toluene, alkanes, or
alcohols), acids (hydrochloric and
sulfuric acid) and alkali (sodium hydroxide) [2-5, 62-64]
.
Surfactants form micelles together with membrane molecules,
while oxidizing agents affect the
metabolic and physiological processes, damaging the cell
membrane [62, 64]
.
Organic solvents hydrolyze or are absorbed by the lipids in the
cell-wall causing them to swell and
ultimately to break [65]. Acids cause the formation of pores
within the cell membrane/wall and bases
saponify the membrane lipids [2]
.
Despite their selectivity and low energy requirement, the use of
chemical agents entails some
limitations. Although chemicals selectively attack the cell
wall, the same chemicals can also attack
accessible intracellular compounds after disrupting the biomass.
From this perspective, chemical
treatment is not selective and the value of intracellular
products may decrease due to loss of
functionality. In addition, the product quality can also be
highly affected due to oxidation or high
temperatures that are usually combined with these chemicals to
enhance cell disruption. Finally, after
introducing those chemicals, they will remain present in the
subsequent downstream process acting
as a contaminant [2-5, 64]
. These set of drawbacks requires the use of low concentrations
of chemical
agents, which impacts the yields of product recovery and so
economical feasibility.
2.2.1.2 Enzymatic Cell Disruption
Enzymatic cell disruption features enzymes that, in a general
way, specifically bind to cell
membrane/wall constituents hydrolyzing their intermolecular
bonds [62, 64]
.
Lysozyme, protease K, driselase, α-amilase and cellulase are
some of the enzymes that were already
tested for microalgae cell disruption. The disruption yields in
enzymatic processes are mainly
dependent on the enzyme selected, its mechanism of action and
configuration (free or immobilized),
reagents concentrations (enzyme and microalgae), and so reactive
bonds [3, 62, 64]
.
To render this process efficient a balance between the above
factors has to be established together
with other enzyme intrinsic parameters such as optimal
temperature, pH and salt concentration [13, 59]
.
Additionally, microalgae biochemical composition (mainly lipid
content) should also be considered
since it can influence the morphology of the microalgal cell,
thus affecting the interaction with the
enzyme and ultimately the disruption efficiency [2-5]
.
This Non-Mechanical method has been regarded as an interesting
microalgae cell disruption
technique due to its biological specificity, mild operating
conditions (absence of shear forces unlike
Mechanical methods) and low energy requirements [2, 3, 5]
.
Despite their potential, their application costs are too high
and their reproducibility quite low since the
same enzyme might not be able to disrupt microalgae with
different cell wall composition [2-5]
.
-
17
2.2.1.3 Physical Cell Disruption
Osmotic shock has already been reported for microalgae lipids
recovery. It consists in a sudden
change in the solute concentration around the cells, resulting
in the abrupt change of water movement
across its cell membrane. Under high concentrations of salts,
substrates or any other solute (neutral
polymers such as polyethylene glycol, dextran) in the
supernatant, water is drawn out of the cells
through osmosis, preventing the transport of substrates and
cofactors into the cell, ultimately causing
its breakage [2, 5]
.
Like enzymatic extraction, the osmotic shock method of microbial
cell disruption has found no large-
scale applications owing to high cost, water foot print and long
treatment times. In addition its
efficiency depends on the microalgae strain which also affects
its robustness [2, 4, 5]
.
2.2.2 Mechanical Cell Disruption Methods
Mechanical cell disruption methods can act by solid-shear (e.g.,
bead mill, high speed
homogenization), liquid-shear forces (e.g., high pressure
homogenization, microfluidization) and
energy transfer through waves (e.g., ultrasonication, microwave)
or currents (e.g., pulsed electric field) [3, 5]
. They usually result in the complete destruction of the cell
wall in a non-specific manner due to
their harshest nature, when compared to non-mechanical methods
[36]
. For instance, Figure 2.5
compares different non-mechanical and mechanical methods for
total carotenoids extraction from the
microalga Haematococcus. It can be observed that mechanical
methods yield a higher release for this
pigment [36]
.
Besides the higher product recovery, mechanical disruption of
cells is generally preferred also
because no additional impurities are introduced into the system,
avoiding further contamination of the
algal preparation while preserving the functionality of the
material within the cell [3, 5, 36]
.
Figure 2.5 – Extraction of total carotenoids into acetone from
processed Haematococcus biomass. (1) Control;
(2) Autoclave (physical, Non-Mechanical); (3) HCl 15 min
(chemical, Non-Mechanical); (4) HCl 30 min (chemical,
Non-Mechnical); (5) NaOH 15 min (chemical, Non-Mechanical); (6)
NaOH 30 min (chemical, Non-Mechanical); (7)
Enzyme (enzymatic, Non-Mechanical); (8) High Speed
Homogenization (solid-shear, Mechanical) [36]
;
-
18
2.2.2.1 Solid Shear Cell Disruption
BBeeaadd MMiilllliinngg ((BBMM))
Bead milling or bead beating is a simple and effective method to
disrupt microalgae [2, 3, 5, 66]
. Its set-up
can display various configurations. The most basic consists in a
grinding chamber with a rotating shaft
through its centre. This shaft is fitted with agitators of
different designs (concentric or eccentric discs,
rings) that impart kinetic energy to small beads in the chamber,
forcing them to collide with each other [2]
. The energy input is dissipated as heat and so the units are
jacketed with high-capacity cooling
systems [2, 3]
. The beads, typically < 1.5 mm steel or glass, are retained
in the grinding chamber by a
sieve or an axial slot smaller than the bead size [2, 3, 5]
.
The mechanism of disruption is not yet completely understood but
the general consensus is that cells
break in the contact zones of the beads by compaction or
shearing action and by energy transfer from
beads to cells [2, 5, 66]
. The degree of disruption relies on many different factors:
contact area between
biomass and beads; bead size, shape and composition; agitator
peripheral velocity; flow rate;
temperature; and rigidity of the microalgal cell walls [3]
.
Algal cell disruption in bead mills is regarded as one of the
most efficient methods for mechanical cell
disruption with potential for industrial implementation [2-5,
66]
. It shows high disruption efficiency in
single-pass operations, high throughput, high biomass loading,
good temperature control,
commercially available equipment, easy scale up procedures and
low labor intensity [3, 5]
.
Figure 2.6 – Bead milling typical experimental set-up [3]
.
However, this technique requires a large amount of energy. The
energy transfer from the rotating shaft
to individual cells is claimed to be inefficient and the
increase in temperature requires intensive,
energy demanding cooling to allow the recovery of functional
fragile ingredients [65]. In addition, the
formation of cell debris and non-selective distribution of
biochemicals over the soluble and solid phase
hamper downstream processing, contributing to the increase of
further operating costs. [2, 3, 5, 66]
HHiigghh SSppeeeedd HHoommooggeenniizzaattiioonn ((HHSSHH))
A high-speed homogenizer is a stirring device, consisting of a
stator-rotor assembly with many
different possible configurations, usually made of stainless
steel [2]
. At a specific rpm value (speed
-
19
critical value), hydrodynamic cavitation is generated due to a
local pressure drop to a value near the
vapor pressure of the liquid and shear forces at the
solid-liquid interphase are formed, resulting in cell
disruption. [3, 5]
Figure 2.7 – High speed homogenizer typical experimental set-up
[67]
.
The degree of cell disruption depends on the blade design,
speed, microalgae species, biomass
concentration and treatment time [3]
.
High speed homogenization is considered a very interesting cell
disruption method due to short
contact times and the potential to process suspensions with
relatively high biomass concentration,
thus reducing the water footprint and downstream process costs
[2, 3, 5]
.
The shear stress induced and the consequent temperature
increase, though, lead to protein
denaturation. In addition, it is still a high energy demanding
technique [2, 3, 5]
.
2.2.2.2 Liquid Shear Cell Disruption
HHiigghh PPrreessssuurree HHoommooggeenniizzaattiioonn
((HHPPHH))
A high pressure homogenizer consists of a positive-displacement
pump that forces a cell suspension
through the centre of a valve seat and radially across the seat
face. Fluid pressure is controlled by a
spring-loaded or hydraulically controlled valve. The fluid flows
radially across the valve, striking an
impact ring, and the suspension then exits the valve assembly to
either a second valve or to discharge [2]
. The shear forces that result from high-pressure impact and
hydrodynamic cavitation causes the cell
disruption [2, 16]
.
In this case, cell disruption is mainly dependent on the
homogenizer design (configuration of the valve
seat), pressure, flow rate, biomass concentration and microalgal
species [3, 5]
.
High pressure homogenization, like bead milling is considered
one possible candidate for the industrial
scale cell disruption of microalgae [16, 36]
. However it has its shortcomings such as the use of low
-
20
biomass concentrations resulting in large treatment volumes,
difficulties to break microalgae cell walls,
generation of very fine cell debris and product (protein) damage
[3]
. This entails the increase of water
footprint and downstream costs [3, 5]
.
Figure 2.8 – High speed homogenizer typical experimental
set-up.
UUllttrraassoonniiccaattiioonn ((UUSS))
Ultrasonication or ultrasounds generate intensive microbubbles
in liquid medium. These microbubbles
grow and collapse violently (cavitation) giving rise to a shock
wave with enough energy not only to
disrupt cell membranes but also to break covalent bonds. These
harsh conditions result in vigorous
and effective cell disruption leading to substantial release of
intracellular materials into the bulk liquid [2, 3]
.
The degree of disruption is mainly influenced by the equipment
power and the microalgal species [3, 5]
.
This mechanical method is claimed not be applicable on a large
scale. Product recovery is poor and
during the process, considerable amount of heat is produced
during the process, destroying
intracellular metabolites such as proteins. In addition, very
reactive hydroxyl radicals can be formed
during sonication, ultimately interacting with most
biomolecules. Finally, a high power input is required
and the method is not very robust since it is not equally
successful in disrupting various microalgal
species. [3, 5, 16]
2.2.2.3 Other
MMiiccrroowwaavvee ((MMWW))
Microwaves interact selectively with the dielectric or polar
molecules (e.g., water) resulting in local
heating due to frictional forces between inter- and
intramolecular movements. The free water
concentration in cells contributes to the microwave efficiency
for cell disruption [16]
. Water exposed to
microwaves reaches the boiling point fast, expanding within the
cell thus increasing the internal
pressure [3, 5]
. The local heat and pressure, together with the microwave
induced damage to the cell
membrane/wall, allow the recovery of intracellular metabolites
[3]
.
-
21
Cell disruption is mostly dependent on the equipment power,
treatment time and microalgal species [3,
5].
Microwave is considered an effective, robust, and of easy
scale-up cell disruption method. However, it
is limited to polar solvents and not suitable for volatile
target compounds. The formation of free
radicals, the temperature increase and the triggered chemical
reactions can interfere with the recovery
of functional compounds. [3, 5, 16]
2.3 Pulsed Electric Field
Like Microwave, Pulsed Electric Field can be placed within the
“Other” category. This method
underlying mechanism of disruption does not rely neither in
solid nor liquid shear. Instead, it consists
in a cell membrane phenomenon usually defined as electroporation
or electropermeabilization. It
consists in the application of periodic, high intensity and
brief electric pulses to cell suspensions,
resulting in the increase of cells membrane permeability and so
in a greater access into or out of the
cell cytoplasm. [24, 25, 27]
To date, it is known that the cell membrane of most biological
cells is made up of phospholipids
assembled by non-covalent interactions in a non-planar bilayer.
Proteins acting as channels or pumps
for specific molecules transportation can also be found along
with it [34]
. Electrically, both cell
membrane surfaces (internal and external) display free charges
of opposite signs, but not inside the
phospholipid bilayer [24, 25, 34]
. Thus, an intrinsic transmembrane potential (TMPm) across the
cell
membrane already exists.
Spontaneous aqueous pore formation takes place whenever the cell
membrane is exposed to
sufficiently high temperature, surface tension or any other
scenario that might affect the conformation
of the lipids within it. However, these pores are too small and
unstable with radii below a nanometer
and lifetimes below a nanosecond. [34]
On the other hand, if the same cell is exposed to an external
electric field, the energy required for
spontaneous formation of aqueous pores is reduced, the number of
pores increases, as they are
larger and more stable, with radii above a nanometer and
lifetimes ranging from milliseconds up to
minutes. [34]
Although the belonging mechanism has not been fully elucidated
yet, a general consensus supports
the idea that by applying an external electric field,
accumulation of surface charges is induced and so
an increase of the local TMP is observed [23-26, 34]
. The simultaneous attraction between opposite
charges on both sides of the membrane causes its thinning until
electrocompression exceeds its
elastic resistance causing it to break. This breakdown results
in the rearrangement of membrane lipids
hydrophilic regions, ultimately giving rise to pores. [24,
34]
The number/distribution, length and thus the stability of the
pores created rely on the intensity of the
electric field applied and its time of exposure to cells. The
combination of these two factors can entail
three different possible outcomes [24-26, 34]
:
-
22
A B C
Figure 2.9 – Lipid bilayer rearrangement in a cell membrane
subjected to an electric field (E) of increasing
strength. ER corresponds to the minimum electric field required
for pores formation. A. The cell membrane has a potential TMPm and
it is being charged to a potential TMP > > TMPm causing its
thinning; B. The membrane has reached breakdown potential TMPR with
resulting reversible pore formation; C: Pore enlargement resulting
in irreversible membrane breakdown.
[24, 34]
1. If the electric field applied and its time of exposure are
such that the resulting potential goes
below the TMPR, then, there is no formation of pores (Figure
2.9A).
2. If the electric field applied and its time of exposure are
such that the resulting potential goes
above the TMPR, reaching the critical value (breakdown
potential), then formation of reversible
pores takes place (reversible electroporation). In this
scenario, the size and number of pores
is not sufficient to maintain membrane breakdown. Thus, as soon
as the electric field is
removed, the pores cease to exist and the original cell membrane
conformation is restored to
its viable state. (Figure 2.9B)
3. If the electric field applied and its time of exposure are
such that the resulting potential goes
above the TMPR and beyond the critical value, then formation of
irreversible pores takes place
(irreversible electroporation). As soon as the application of
the electric field ceases, the pores
still exist due to their high number and larger size when
compared to reversible electroporation
and so, cell membrane original conformation can be no longer
restored to its viable state.
(Figure 2.9C).
Besides the intensity of the electric field and time of
exposure, the degree of disruption also depends
on biological and other physical factors [24, 25]
. Biological factors include sample conductivity, type and
physiological state of biological cell and load concentration,
while physical factors consist on electric
field, number of pulses, pulse shape and length and
electroporation chamber.
-
23
Sample conductivity is an important and critical parameter in
the electroporation efficiency. The higher
the conductivity of the cell suspension, the lower is its
electrical resistance to current flow [24-26, 34]
. This
current leak through the electrolyte corresponds to a waste of
energy in the form of heating and so
process efficiency becomes lower. In the limit, high sample
conductivity may favor arcing (electric
current surge through the sample) between the electrodes which
results in its degradation, thus
limiting the maximum applicable electric field.
The biological cell type and physiological state are also
important [24]
. Animal, bacterial, fungal, plant
and microalgal cells have different features. Unlike animal
cells, bacterial, fungal, plant and some
microalgal cells exhibit a cell wall, the biochemical
composition and thickness of which changes
according to the species [36]
. The presence of a cell wall influences electroporation
performance since
it is an additional and thicker barrier to overcome.
Gram-negative bacteria or yeasts are more
sensitive to pulsed electric fields than gram-positive bacteria,
which have a thicker cell wall [23]
.
In general, the physiological state affects cells morphology and
size. Size has been reported in
literature as a key-factor in electric fields efficiency [19,
21, 28]
. For smaller cells and their organelles
permeabilization, higher intensity electric fields have to be
applied than for larger cells. Coustets et al. [28]
studied the effect of the electric field applied in the
permeabilization of three microalgae species
with different sizes – Nannochloropsis salina (diameter=2,5 μm),
Chlorella vulgaris (diameter=3-6 μm)
and Haematococcus pluvialis (diameter= 5-25 μm). They were able
to observe that the electric field
required to attain successful electroporation would increase
with microalgae size reduction.
The cell load, i.e, the concentration of biological cells in the
suspension under treatment, can also
impact electroporation. The amount of released intracellular
content after electroporation is higher for
higher concentrations thus resulting in the increase of sample
conductivity and consequent reduction
of sample electrical resistance and, once again, in energy
dissipation as heat. [3, 28]
Electroporation is also markedly affected by the electric
process itself [24, 25]
. The importance of the
electric field, number of pulses and pulse length has been
showcased already. Depending on these,
electroporation can be reversible or irreversible. Another
relevant aspect is the electric chamber used,
namely its electrodes configuration and pulse generator.
Electrodes configuration determine the
distribution of the electric field in the test chamber. Pulse
generators are responsible for reproducibility
and electroporation efficacy [25, 35]
.
There are four main configurations for the chamber and
electrodes [35]
: single-cell chambers, micro-
electrodes, mili-electrodes and flow-through chambers. Micro and
milli-electrodes are frequently used
in laboratory scale for batch mode operations, while
flow-through chambers are designed to process
larger volumes of cells for continuous mode operations (Figure
2.10).
A B C D
-
24
Figure 2.10 – Examples of batch and continuous electroporation
chambers. A. Lab-scale batch electroporation
chamber (Micro and Milli-electrodes); Flow-through chamber with
a B. coaxial geometry, C. a collinear geometry
and D. a planar geometry. Red and blue regions match opposite
electrodes. [35]
Pulse generators are responsible for the shape, amplitude and
polarity of the pulses [25, 35]
. In Figure
2.11 some typical pulse shapes such as exponential decay, square
and bell wave are depicted.
Square pulse generators are the most used for (reversible and
irreversible) electroporation
applications because they are claimed to have a greater lethal
effect when compared to the other two
mentioned [24, 25]
. Such can be explained by the fact that square wave pulse
maintains a maximum
voltage and field for the entire pulse duration, whereas an
exponential decay pulse has a major portion
of the pulse as a tail with low voltage and field [25]
. In addition, square wave pulses also enable a good
control and reproducibility of the electrical features of the
pulses being applied which enhances
electroporation [35]
.
Polarity can also influence electroporation efficacy. Polarity
inversion (bipolar pulses) is believed to
change the direction of moving charges within the cell membrane
thus facilitating the electroporation
process [23, 25, 35]
.
Both reversible and irreversible electroporation have steadily
gained ground in many different fields,
since its discovery [24, 26, 34]
.
Reversible electroporation finds wide applications in medicine,
from introducing chemotherapeutic
drugs into tumor cells to gene therapy dropping the risks caused
by viral vectors. Moreover, it is also a
useful tool in biotechnology for genetic information transfer
into cells to artificially over express an
intracellular or extracellular valuable product. [26, 34]
Irreversible electroporation can also play a role in medicine as
a method for tissue ablation
[34]. In
biotechnology it has been widely used as an alternative to the
traditional thermal pasteurization
processing of liquid foods [26]
. Instead of applying heat, contaminant microorganisms
deactivation is
achieved by the formation of irreversible pores within their
cell membrane, while conserving the
organoleptic properties of food [23, 26]
.
A B C
Figure 2.11 – A. Exponential, B. square and C. bell wave pulses.
[35]
The same principle is now being envisioned to extract valuable
intracellular components from
microalgae, by creating irreversible pores within their cell
wall and membrane. Table 2.2 points
out the advantages of using PEF over the methods described in
the previous section in order to attain
this goal.
-
25
Table 2.2 – PEF advantages over Non-Mechanical and other
Mechanical microalgae Cell Disruption Methods
based on literature [2-5, 63, 64]
. BM – Bead milling, HSP – High Speed Homogenization, HPH – High
Pressure
Homogenization, US – Ultrasounds, MW – Microwave.
Non-Mechanical Methods Mechanical Methods
vs Chemical Enzymatic Physical Solid-Shear Liquid Shear
Other
BM HSH HPH US MW
PEF
No additional impurities to hamper product recovery Shorter
treatment times
Less energy intensive Less cell debris to hamper product
recovery
Less energy intensive Less shear stress and so higher product
quality
Less cell debris to hamper product recovery Lower temperature
increase and so high product quality
Less energy intensive Lower temperature increase and so high
product quality No radicals formation
Not-limited by polar solvents or volatile target compounds Lower
temperature increase and so high product quality No radicals
formation
Despite its promising features over the other cell disruption
methods, PEF also faces some challenges
as a microalgae cell disruption method. Its efficiency
dependence on sample conductivity might
represent an additional hurdle when dealing with marine
microalgae. This is a real drawback since the
use of the latter is ideal to reduce freshwater and nutrient
footprint. A marine microalgae suspension
has a conductivity value higher than a freshwater microalgae
suspension so in order to apply PEF, this
conductivity has to be reduced as will be discussed ahead, in
section 3.4.1. A washing pre-treatment
step is thus required to allow the use of PEF with marine
microalgae.
Cell load can also stand a problem. The higher it is, the more
products are released within the cell
suspension, sample conductivity increases and more energy gets
dissipated through heat rendering
the process less energy efficient.
Although a few studies on the use of PEF for microalgae cell
disruption, are already available in the
literature (Table 2.3), these did not assess the impact of these
limitations before placing PEF among
other cell disruption methods.
Table 2.3 – Literature review on the use of PEF for microalgae
cell disruption. (T– Temperature; DCW – Dry Cell
Weigth; EF– Electric Field; PL– Pulse Length; f – frequency; ET
– Energy Treatment).
Microalgae species
Environment Product Conditions PEF treatment
Main outcomes Ref
Synechocystis PCC 6803
Fresh
Lipids
36-54°C outflow T; 0.03%DCW
Continuous mode 59.67-239 kWh/kgDW. EF, PL and f not
specified
Lipid recovered intact after PEF; Less organic solvent required
for lipid extraction.
Sheng et al. 2011 [31]
Auxenochlorella
Fresh
Protein
T outflow not
Continuous
Biomass concentration does not
Goettel et al. 2013
-
26
protothecoides Carbohydrates Lipids
specified 3.6-16.7% DCW
mode 0.014-0.059 kWh/kgDW. EF=23-43 kV/cm PL=1 μs f=not
specified
affect PEF efficiency; No evident effect of EF in cell
disruption; Only soluble substances leaked spontaneously; Less
organic solvent required for lipid e