Assessment of a Pulsed Electric Field based Treatment for ... · Assessment of a Pulsed Electric...

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

i

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.

iii

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

v

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.

vii

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.

ix

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

x

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

xi

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

xiii

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

xiv

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

xv

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

xvi

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



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



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


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



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

xvii

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

xix

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

xx

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

1

Chapter 1

Introduction

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.

7

Chapter 2

Theoretical Background

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).


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)


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

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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

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