Reconstitution of Ion Channels in a Lipid Bilayer in aMicrofluidic Device using Cell-Free Protein Synthesis

Teresa Filipa Guerreiro Machado

Thesis to obtain the Master of Science Degree in

Biological Engineering

Supervisors: Prof. Gabriel Antonio Amaro Monteiro and Dr.ir Severine Le Gac

Examination CommitteeChairperson: Prof. Jorge Humberto Gomes LeitaoSupervisor: Prof. Gabriel Antonio Amaro MonteiroMember of the Committee: Prof. Maria Angela Cabral Garcia Taipa Meneses de Oliveira

July 2015

I have not failed. Ive just found 10,000 ways that wont work.Thomas A. Edison

Acknowledgments

I would like to express my gratitude to my supervisors Dr.ir Severine Le Gac and Dr. Alexander

Prokofyev for giving me the opportunity of developing my master thesis at the University of Twente

and for their excellent guidance and support, not only during my stay in The Netherlands but also

during the preparation of this manuscript.

My deepest acknowledgment is addressed to Dr. Alexander Prokofyev, for his constant guidance

and patience, for all the constructive advices and especially for his support and encouragement, which

were essential to keep me motivated throughout the development of this thesis.

To Yusuf Arik, Marleen Munsterman and everyone else who accompanied me at BIOS, my deepest

gratitude for being so friendly and for all the help provide during my stay. Furthermore, I would like to

thank Nathalie Schilderink for all the help and orientation during my work in the NBP facilities.

I also present my thankfulness to my supervisor Prof. Gabriel Monteiro for all the support provided.

To all my Friends, a huge thanks, for all the support and for always staying by my side, even when

I disappear for weeks and miss all our appointments. I could not have asked for better friends and I

know that I will count on you for the rest of my life.

To Pedro, a very special thanks, for all the love and unconditional support, for believing in me and

above all for always making me believe in myself.

Finally, my heartfelt gratitude to my family. To my parents, for their unconditional love and support

and for giving me the opportunity and encouragement for studying abroad. To my sister Catarina for

always, always put a smile on my face.

Muito obrigada!

Dankjewel!

iii

Abstract

Ion channels, which are transmembrane proteins, play a crucial role controlling a very wide spec-

trum of physiological processes. Mutations in genes encoding for ion channels results in the alteration

of their function often leading to diseases, making this type of proteins very attractive targets for the

development of new drugs.

Nonetheless, the expression and purification of ion channels has been a challenge. Cell-free

protein synthesis (CFPS) has emerged in the past years as a promising alternative to overcome the

limitations associated with the cellular systems. Additionally, an automable platform with sufficient

capacity for high throughput screening and high information content is not currently available. For this

reason, miniaturized bilayer lipid membranes (BLMs) platforms have gained more interest.

In this context, this work aimed to expressed ion channels, namely KcsA potassium channel and

bacteriorhodopsin (bR) channel, using a CFPS system and their subsequent electrophysiological

characterization in a microfluidic BLM device.

The proteins of interest were expressed in a CFPS systems based on an optimized E. coli ex-

tract, in the presence of nanolipoproteins (NLPs) and in a batch format. The expression of bR was

confirmed in a SDS-PAGE gel. However, for KcsA the results of the gel were not conclusive. The ineffi-

cient purification procedure not allowed to obtain suitable samples for electrophysiological recordings.

Hence, the results from this experimental work indicate that there is a need to revise the proto-

col used for the expression of the target proteins in the CFPS system and also to investigate other

methods of analysis and purification of the CFPS products.

Keywords

Ion channels; KcsA Potassium channel;, Bacteriorhodopsin channel; Cell-free protein synthesis;

Bilayer lipid membranes; Microfluidic

v

Resumo

A classe de protenas transmembranares, canais ionicos, desempenha um papel crucial no con-

trolo de um vasto espectro de processos fisiologicos. As mutacoes em genes que codificam estes

canais provocam a alteracao da sua funcao, resultando muitas vezes em doencas, o que os transfor-

mou em alvos muito atraentes para o desenvolvimento de novas drogas.

No entanto, a expressao e purificacao de canais ionicos tem sido um desafio. A cell-free protein

synthesis (CFPS) emergiu nos ultimos anos como uma alternativa promissora para ultrapassar as

limitacoes associadas aos sistemas celulares. Alem disso, nao existe actualmente disponvel, uma

plataforma autonoma, de alto rendimento e alto conteudo de informacao para testar novos farmacos.

Por esta razao, as plataformas miniaturizadas para membranas de bicamada lipdica (MBLs) tem

despertado muito interesse.

Neste contexto, este trabalho tem como objectivo expressar canais ionicos, nomeadamente o

canal de potassio KcsA e a bacteriorodopsina (bR), utilizando um sistema CFPS e sua posterior

caracterizacao eletrofisiologica num dispositivo microfludico MBL.

As protenas alvo foram expressas em sistemas de CFPS utilizando um extracto optimizado de E.

coli, na presenca de nanolipoproteinas (NLPs) e num formato batch. A expressao da bR foi obser-

vada num gel de SDS-PAGE. No entanto, para o KcsA os resultados do gel nao foram conclusivos.

O procedimento ineficiente de purificacao nao permitiu a obtencao de amostras adequadas para

registros eletrofisiologicos.

Assim, os resultados deste trabalho experimental indicam que existe a necessidade de rever o

protocolo utilizado para a expressao das protenas alvo no sistema CFPS e tambem investigar outros

metodos de analise e purificacao dos produtos de CFPS.

Palavras Chave

Canais ionicos; Canal de potassio KcsA; Bacteriorrodopsina; Cell-free protein synthesis; Mem-

branes de bicamada lipdica; Microfludico

vii

Contents

1 Introduction 1

1.1 Background and motivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1.2 Research goals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

1.3 Structure of thesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2 Literature Review 5

2.1 Ion channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

2.1.1 General definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

2.1.2 Voltage-gated Potassium channels . . . . . . . . . . . . . . . . . . . . . . . . . . 7

2.1.2.A KcsA Potassium channel . . . . . . . . . . . . . . . . . . . . . . . . . . 8

2.1.3 Bacteriorhosopsin channel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

2.2 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

2.2.1 CFPS Extract sources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

2.2.2 Configurations of CFPS reactions . . . . . . . . . . . . . . . . . . . . . . . . . . 14

2.2.3 Cell-free expression of membrane proteins . . . . . . . . . . . . . . . . . . . . . 15

2.2.3.A Cell-free synthesis of membrane proteins as precipitates . . . . . . . . 15

2.2.3.B Cell-free synthesis of membrane proteins in presence of detergents . . 15

2.2.3.C Cell-free synthesis of membrane proteins in the presence of vesicles

and liposomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

2.2.3.D Cell-free synthesis of membrane proteins in presence of nanodiscs . . 17

2.2.4 Applications and future perspectives of CFPS . . . . . . . . . . . . . . . . . . . . 19

2.3 Bilayer Lipid Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

2.3.1 Cell membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

2.3.2 Patch Clamp Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21

2.3.3 Bilayer Lipid Membrane Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

2.3.4 Miniaturized Bilayer Platforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

3 Materials and Methods 27

3.1 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

3.1.1 E. coli strains and vectors used . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

3.1.2 Transformation of competent cells . . . . . . . . . . . . . . . . . . . . . . . . . . 28

3.1.3 Purification of plasmid DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

ix

3.1.4 Primers design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.5 Polymerase chain reaction (PCR) . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.6 TOPO Cloning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.7 Concentration measurements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.8 Cell-Free Protein Synthesis expression . . . . . . . . . . . . . . . . . . . . . . . 30

3.1.9 Purification of CFPS products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32

3.1.9.A Acetone Precipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32

3.1.9.B Solubilization in buffer with detergent . . . . . . . . . . . . . . . . . . . 32

3.1.9.C Protein purification using Ni-NTA Spin Column . . . . . . . . . . . . . . 32

3.1.10 Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis (SDS-PAGE) . . . 33

3.2 Bilayer Lipid Membrane Experiments in a microfluidic device . . . . . . . . . . . . . . . 33

3.2.1 Fabrication of the Microfluidic Device for Bilayer Lipid Membrane Experimentation 33

3.2.2 Bilayer Lipid Membrane Experimental Set-up . . . . . . . . . . . . . . . . . . . . 35

3.2.3 Bilayer Lipid Membrane Formation and Characterization . . . . . . . . . . . . . . 35

3.2.4 Ion Channels Recording . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

3.2.4.A Gramicidin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

3.2.4.B KcsA Potassium Channel . . . . . . . . . . . . . . . . . . . . . . . . . . 36

3.2.4.C Bacteriorhodopsin Channel . . . . . . . . . . . . . . . . . . . . . . . . . 37

4 Results 39

4.1 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

4.1.1 Construction of the pEXP5-CT/KcsA plasmid vector . . . . . . . . . . . . . . . . 40

4.1.2 Cell-free protein synthesis of bacteriorhodopsin . . . . . . . . . . . . . . . . . . . 43

4.1.3 Purification of bacteriorhodopsin . . . . . . . . . . . . . . . . . . . . . . . . . . . 44

4.1.4 Cell-free protein synthesis and purification of KcsA . . . . . . . . . . . . . . . . . 49

4.2 Bilayer Lipid Membrane Experiments in a microfluidic device . . . . . . . . . . . . . . . 54

4.2.1 Fabrication of the Microfluidic Device for Bilayer Lipid Membrane Experimentation 54

4.2.2 Bilayer Lipid Membrane Formation and Characterization . . . . . . . . . . . . . . 55

4.2.3 Ion Channels Recording . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

4.2.3.A Gramicidin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

4.2.3.B KcsA Potassium Channel . . . . . . . . . . . . . . . . . . . . . . . . . . 57

4.2.3.C Bacteriorhodopsin Channel . . . . . . . . . . . . . . . . . . . . . . . . . 60

5 Discussion 63

5.1 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

5.2 Bilayer Lipid Membrane Experiments in a microfluidic device . . . . . . . . . . . . . . . 67

6 Conclusion 69

6.1 General conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

6.2 Recommendations for future work . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

x

Bibliography 73

Appendix A KcsA Potassium channel I-V curve A-1

xi

List of Figures

2.1 Structure of Voltage-gated Potassium Channels for List of Figures. . . . . . . . . . . . . 7

2.2 Schematic view of KcsA Potassium Channel for List of Figures. . . . . . . . . . . . . . . 8

2.3 Schematic view of the Bacteriorhodopsin (bR) photo-cycle and the proton exchange

steps for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

2.4 Schematic comparison between the conventional cell-based expression systems and

the cell-free protein synthesis systems for List of Figures. . . . . . . . . . . . . . . . . . 12

2.5 Structure of a nanodisc for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . 18

2.6 Summary of the current strategies for the cell-free expression of membrane proteins

for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

2.7 Schematic representation of the patch clamp technique for the whole cell configuration

for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

2.8 Schematic representation of bilayer lipid membrane approaches for List of Figures. . . . 23

3.1 Schematic representation of the major steps of the standard protein synthesis reaction

for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

3.2 Diagram of the the fabrication of glass-Teflon-glass microfluidic devices for List of Figures. 34

3.3 Experimental set up for bilayer lipid membrane experimentation for List of Figures. . . . 35

4.1 Scheme of the process strategy followed to contruct pEXP5-CT/KcsA plasmid vector

for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

4.2 Results of sequencing for plasmid vector pQE60 /KcsA after amplification for List of

Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

4.3 PCR image for KcsA amplification for List of Figures. . . . . . . . . . . . . . . . . . . . . 41

4.4 Results of sequencing for plasmid vector pEXP5-CT with the KcsA gene inserted for

List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

4.5 Results of sequencing for plasmid vector pEXP5-CT with the KcsA gene inserted for

List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43

4.6 bR expression in the presence (Positive) or absence (Negative) of MembraneMax

Reagent and All-trans retinal for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . 44

4.7 SDS-PAGE of cell-free synthesized bacteriorhodopsin expressed in the presence (+)

or absence (-) of MembraneMax reagent for List of Figures. . . . . . . . . . . . . . . . . 45

xiii

4.8 SDS-PAGE of cell-free product reaction bacteriorhodopsin expressed in the presence

(+) of MembraneMax reagent and in optimized conditions for List of Figures. . . . . . . . 46

4.9 SDS-PAGE of bR expressed in standard conditions and in the presence (+) of Mem-

braneMax reagent and subsequently purified over a Ni-NTA spin column for List of

Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47

4.10 SDS-PAGE of FT fraction of the previous Ni-NTA purification purified again over a Ni-

NTA spin column for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

4.11 SDS-PAGE of bR expressed in the presence (+) of MembraneMax reagent and subse-

quently purified over a Ni-NTA spin column for List of Figures. . . . . . . . . . . . . . . . 49

4.12 SDS-PAGE for KcsA Potassium channel for List of Figures. . . . . . . . . . . . . . . . . 50

4.13 SDS-PAGE of KcsA expressed in the presence (+) of MembraneMax reagent at stan-

dard conditions and subsequently purified over a Ni-NTA spin column for List of Figures. 51

4.14 SDS-PAGE of cell-free synthesized KcsA expressed in the presence of MembraneMax

reagent, and purified with acetone (A), detergent DM and heated (DM 95) and unheated

(DM RT) for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

4.15 SDS-PAGE of KcsA expressed in the presence (+) of MembraneMax reagent at opti-

mized conditions and subsequently purified over a Ni-NTA spin column for List of Figures. 53

4.16 SDS-PAGE of cell-free synthesized bR expressed in the presence of MembraneMax

reagent (+) and cell-free synthesized KcsA expressed in optimized conditions for the

amount of DNA template used, 1 g and 2 g for List of Figures. . . . . . . . . . . . . . 54

4.17 Microfluidic device for bilayer lipid membrane experimentation for List of Figures. . . . . 55

4.18 Bilayer lipid membrane for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . 56

4.19 BLM recordings for pore forming gramicidin for List of Figures. . . . . . . . . . . . . . . 57

4.20 BLM recordings for KcsA Potassium channel for List of Figures. . . . . . . . . . . . . . . 58

4.21 BLM recordings for KcsA Potassium channel for CFPS products for List of Figures. . . . 60

4.22 BLM recordings for bacteriorhodospin for CFPS products for List of Figures. . . . . . . . 62

A.1 Kcsa I-V curve for List of Figures. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A-2

xiv

List of Tables

3.1 Detailed description of One Shot TOP10 Chemically Competent E. coli cells genome. . 28

3.2 Primers designed for PCR reaction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.3 Reagents for cell-free protein synthesis expression . . . . . . . . . . . . . . . . . . . . . 30

3.4 Cell-free protein synthesis Feed Buffer composition . . . . . . . . . . . . . . . . . . . . . 31

3.5 Description of samples used for bilayer lipid membrane experiments with cell-free pro-

tein synthesis products of KcsA expression. . . . . . . . . . . . . . . . . . . . . . . . . . 37

3.6 Description of samples used for bilayer lipid membrane experiments with cell-free pro-

tein synthesis products of negative (-) bacteriorhodopsin control expression. . . . . . . . 38

3.7 Description of samples used for bilayer lipid membrane experiments with cell-free pro-

tein synthesis products of negative (+) bacteriorhodopsin control expression. . . . . . . 38

4.1 Primers designed for PCR reaction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41

4.2 Membrane properties for control experiments. Bilayers are formed with DPhPC (25

mg/mL) in n-decane. The Buffer used was 150 nM KCl, 10 nM HEPES, at pH 7. The

measurements were recorded using a gain of 20 mV/pA. The BLMs are characterized

in terms of seal resistance (Rm), capacitance (Cm), surface area (ABLM ) and specific

capacitance (Cs). n - number of experiments performed; N - number of devices used. . 56

4.3 Membrane properties for gramicidin experiments. Bilayers are formed with DPhPC (25

mg/mL) in n-decane supplemented with 2 nM Gramicidin. The Buffer used was 1 M

KCl, 10 nM HEPES, at pH 7. The measurements were recorded using a gain of 20

mV/pA. The BLMs are characterized in terms of seal resistance (Rm), capacitance

(Cm), surface area (ABLM ) and specific capacitance (Cs). n - number of experiments

performed; N - number of devices used. . . . . . . . . . . . . . . . . . . . . . . . . . . . 57

4.4 Membrane properties for KcsA control experiments. Bilayers are formed with DOPC/CL

(25 mg/mL) in n-decane. The Buffer used for the bottom channel was 150 mM KCl,

10 nM HEPES, pH 4 and in the top channel was introduced the KcsA dissolved in

the Buffer 150 mM KCl, 10 nM HEPES, pH 7, in a ratio 1:300. The measurements

were recorded using a gain of 20 mV/pA. The BLMs are characterized in terms of

seal resistance (Rm), capacitance (Cm), surface area (ABLM ) and specific capacitance

(Cs). n - number of experiments performed; N - number of devices used. . . . . . . . . . 58

xv

4.5 Membrane properties for KcsA control experiments. Bilayers are formed with DOPC/CL

(25 mg/mL) in n-decane. The Buffer used for the bottom channel was 150 mM KCl,

10 nM HEPES, pH 4 and in the top channel was introduced the KcsA dissolved in

the Buffer 150 mM KCl, 10 nM HEPES, pH 7, in a ratio 1:300. The measurements

were recorded using a gain of 20 mV/pA. The BLMs are characterized in terms of

seal resistance (Rm), capacitance (Cm), surface area (ABLM ) and specific capacitance

(Cs). n - number of experiments performed; N - number of devices used. . . . . . . . . . 59

4.6 Membrane properties for CFPS bacteriorhodopsin experiments. Bilayers are formed

with DPhPC (25 mg/mL) in n-decane. The Buffer used for the bottom channel was 0.1

M MgCl2, 0.5 mM Tris, pH 7 and in the top channel was introduced the bR dissolved

in the same buffer in a ratio 1:10 - samples D.1, E.1 and F.3, 1:5 - sample F.2 and

1:1 - sample G.1. The measurements were recorded using a gain of 20 mV/pA. The

BLMs are characterized in terms of seal resistance (Rm), capacitance (Cm), surface

area (ABLM ) and specific capacitance (Cs). n - number of experiments performed; N -

number of devices used. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

xvi

Abbreviations

ABLM membrane surface area

BLM bilayer lipid membrane

bR Bacteriorhodopsin

Cm membrane capacitance

Cs specific capacitance

CAT chloramphenicol acetyl transferase

CF Cell-free

CECF continuous-exchange cell-free

CFCF continuous-flow cell-free

CFPS Cell-Free Protein Synthesis

CMC critical micelle concentration

D-CF cell-free synthesis of membrane proteins in presence of detergents

DIBs Droplet interface bilayers

DM decyl maltoside

DOPC/CL 1,2-Dioleoyl-sn-glycero-3-phosphocholine / cardiolipin

DOPE/CL 1,2-Dioleoyl-sn-glycero-3-phosphoethanolamine / cardiolipin

DPhPC 1,2-Diphytanoyl-sn-glycero-3-phosphocholine

His-tag Histidine tag

MPs Membrane proteins

NLPs nanolipoprotein particles

NMR nuclear magnetic resonance

P-CF cell-free expression of membrane proteins as precipitates

xvii

PCR Polymerase chain reaction

PURE Protein synthesis Using Recombinant Element

SDS-PAGE Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis

S.O.C Medium Super Optimal Broth

TRP transient receptor potential

xviii

1Introduction

Contents1.1 Background and motivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2 Research goals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.3 Structure of thesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

1

1.1 Background and motivation

Ion channels, which are proteins located in cell membranes, play a crucial role controlling a very

wide spectrum of physiological processes such as nerve and muscle excitation, hormone secretion,

cell and lymphocyte proliferation, learning and memory, salt and water balance, regulation of blood

pressure and fertilization or cell death.[1] Mutations in genes encoding for ion channels results in the

alteration of the ion channel function often leading to diseases, called channelopathies, which include

cystic fibrosis, epilepsy, ataxia, myotonia and cardiac arrhythmia. [2][3][4] For these reasons, ion

channels have become very attractive targets for the development of new drugs, and approximately

13 % of known drugs have their primary therapeutic action on this class of proteins, representing a

market of more than 12 billion dollars in worldwide sales. [5]

Nonetheless, the natural abundance of ion channels, and membrane proteins in general, resulting

from conventional cellular expression systems is usually too low to purify enough material for high-

throughput therapeutic drug studies. Moreover, in vivo overexpression of this type of membrane

proteins is overtly problematic, resulting in low yield, cell toxicity, protein aggregation and misfolding.

Additionally, ion channels are embedded in a complex and dynamic lipid bilayer, which makes their

purification difficult, often leading to laborious and hard to handle processes. [6][7]

Cell-free protein synthesis has emerged in the past years as a promising alternatives to overcome

the previously mentioned obstacles, since target protein expression is independent of living cells

integrity in these systems. [8] In addition, the open nature of these systems allows high levels of

control, through the direct access and manipulation of the reaction conditions. [6]

However, even if the expression is successful, the high-throughput drug screening on ion channels

is still challenging. The currently employed pharmacological methods lack in precision. In contrast,

the standard techniques to study ion channels function, such as the patch clamp technique, provides

high content information but is far from being suitable to multiple compound screening, since it has a

very low throughput and is labour intensive. [5][9] Despite automation of the patch clamp technique,

the throughput is still not high enough and the associated costs are very high. [10] Furthermore, all

these methods use cell models, which is a limitation per se. [11] In this context, cell-free systems like

bilayer lipid membranes have gained more interest. The traditional bilayer set up is not suitable for

high throughput automated applications since they require skilled operators and the resulting bilay-

ers are not stable. Besides, to perform multiplex experiments large volumes will be needed, which

is not favourable. However the fields of microfluidics and microfabrication have contributed to the

creation of miniaturized platforms for bilayer lipid membrane experimentation that allow to overcome

the aforementioned limitations. [12][13] Decreasing the size of the structures used in these platforms

enables the controlled handling of sub-microliter volumes. Also the resulting smaller sized aperture

will increase the bilayer stability. Additionally these techniques have the potential for automation

and multiplexing. The horizontal aperture allow use of electrophysiological and optical methods as

well. Several microfluidic and miniaturized bilayer platforms have been reported in the past years.

[12][13][14] However there are still great challenges to overcome; new techniques should be devel-

2

oped and existing ones should be improved in order to develop reliable, automated, high-throughput

drug screening platform.

1.2 Research goals

The goal of this Master project is to optimize an in vitro Cell-Free Protein Synthesis (CFPS) for

expression of ion channels and their subsequent electrophysiological characterization in a microfluidic

bilayer lipid membrane (BLM) device. BIOS - Lab on a Chip group have recently developed a mul-

tiplex microfluidic platform that allows performing simultaneously electrophysiological and confocal

microscopy measurements on bilayer lipid membranes. [15][16] In this work, this microfluidic device

is used to test and characterize cell-free expressed ion channels, KcsA potassium channel and bacte-

riorhodopsin channel. This work has been performed in the BIOS - Lab on a Chip group, which is part

of MESA+ Institute of Nanotechnology and MIRA Institute for Biomedical Technology and Technical

Medicine, at University of Twente.

1.3 Structure of thesis

This master thesis is organized into six chapters. In the first chapter, the background and motiva-

tion, as well as the goals of this thesis are described.

The second chapter gives more detailed information on the topics presented in chapter 1. This

chapter is divided in three main sections. In the first section, a description about ion channels and their

general characteristics is given, followed by an introduction to the ion channels which are central in

this study, the KcsA potassium channel and bacteriorhodopsin channel. Secondly, a description about

cell-free protein synthesis, extract sources, reaction configurations and the application in membrane

proteins studies as well as the major applications of the technique is given. Finally, the techniques to

perform bilayer lipid membranes are reviewed.

Chapter 3 describes the experimental procedures and set-up as well as the analytical and char-

acterization techniques used in order to reach the proposed goals.

The fourth and fifth chapters unveil and discuss the obtain results.

Finally, in chapter 6, conclusions and future perspectives are presented.

3

4

2Literature Review

Contents2.1 Ion channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62.2 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102.3 Bilayer Lipid Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

5

2.1 Ion channels

2.1.1 General definitions

Ion channels are membrane proteins that produce and transduce electric signals in living cells,

regulating the flux of ions through biological membranes. They are found, not only in the external cell

membrane but also in the membranes of intracellular organelles. [17][5]

Ion channels are often highly specific for only one species, and they can be classified according

to the type of ions they allow to pass - sodium (Na+), potassium (K+), calcium (Ca2+), chloride (Cl)

channels. Although some ion channels are less selective - nonspecific cation or anion channels. [3]

The electrochemical gradient, established by combined action between ion channels, active pumps

and co-transporters, of the significant ionic species determines the direction of the net ion flow through

the membrane. The point at which the chemical and electrical driving forces are exactly balanced is

called the Nernst potential. Above or below this point of equilibrium, a particular species of ion flows in

the direction of the dominant force. Given the concentrations of ions and the number, conductances,

selectivities, and gating properties of the various ion channels the net flow of current/ions through the

cell membrane is foreseeable. [17][4] Most ion channels are regulated by external biological signals,

which when triggered causes conformational rearrangements and determines if the channel is open

and the ions can pass through, or if it is close and the flow of ions is prevent. This property is called

gating. [17][18][3] The signal can be the binding of a ligand to the channel like in the family of the

ligand gated ion channels [17][19], a difference of potential across the membrane (the voltage gated

ion channels [17][20], a change in temperature (members of the transient receptor potential (TRP)

channel family [21]), or pressure (the mechanosensitive channels [22]). Each time a channel opens,

up to 10 million ions per second flow into or out of the cell, a picoamper (10-12 pA) current is generate

by the flow of highly selected ions. [4]

Conductance indicates the facility with which ions move through a material and is measured in

Siemens (S). The ratio between the amplitude of current in a single channel (i) and the applied voltage

(V) defines the conductance of a single channel (). [17][4]

=i

V(2.1)

Ion channels are attractive therapeutic targets due to their high control in physiological systems.

Salt and water balance, nerve and muscle excitation, regulation of blood pressure, memory, hormone

secretion, cell proliferation and cell death are some example of the physiological processes where

they are involved. [1] Hence, mutations in genes that codified for ion channels may lead to modifi-

cations in their function and be the cause of several diseases, called channelopathies, which may

include diseases such as cystic fibrosis [23], epilepsy [24][25], heritable hypertension (Liddless syn-

drome) [26][27], the long-QT syndrome [28] familial hyperinsulinemic hypoglycemia of infancy [29][30]

and cardiac arrhythmia [31][32][33]. The number of known diseases caused by the malfunction of

ion channels has been increasing and have increased the interest in ion channels as drug targets.

6

Presently, 13-15% of the drugs on the market target this class of proteins. [5][18][3] However, con-

sidering their crucial role in the cell life, ion channels are still under-targeted, mainly due to difficulties

associated to expression, purification protocols and procedures of the class of proteins as well as for

the huge functional and structural diversity. [7][34]

2.1.2 Voltage-gated Potassium channels

Voltage-gated potassium channels (Kv) belong to a diverse family of cation channels that includes

for example voltage-gated sodium (Nav) and voltage-gated calcium (Cav) channels [35][36][37], Ca2+-

activated K+ channels [38], inward rectifier K+ channels [39][40] and two-pore K+ channels [41][42].

At their simplest, they are composed of four identical (or similar) subunits [43], each containing a

single six-transmembrane region (S1 to S6), with both amino and carboxy termini on the intracellular

side of the membrane. [44] The subunits are assembled to form the central pore in a process that

also determines the basic properties of gating and permeation characteristic of the channel type. The

topology and structure of the S5-S6 region is the best defined because it is well conserved across

K+ channels [45] and the peptide chain (H5 or P loop) between them projects into and lines the

water-filled channel pore. [46][47] Mutations in this region affect the permeation properties of the

channel. S4 region is the major voltage sensor in the ion channel and contains a cluster of positively

charged amino acids (lysines and arginines). [48] The gate region in K+ channels controls the flux of

ions through the ion conduction pore. [45] The permeation pathway is blocked by a tethered amino-

terminal-blocking particle (the ball and chain) leading to a voltage-dependent fast inactivation of

the channel. [48]

Figure 2.1: Structure of Voltage-gated Potassium Channels. A - Subunit containing the six transmembran-spanning motifs, (S1 to S6), which forms the core structure of potassium channels. S4 is the voltage sensor andthe circles containing plus signs are lysine and arginine residues. The ball and chain structure at the N-terminalof the protein is the region that participates in N-type fast inactivation, obstructing the permeation pathway.The channel pore (H5) are found between S5 and S6. The genes for voltage activated potassium channels (Kv)encode a protein with only a single subunit. B - Four subunits assembled to form a potassium channel. [4]

7

2.1.2.A KcsA Potassium channel

The determination of the structure of the KcsA potassium channel from Streptomyces lividan [49] is

of particular interest due to its structural proximity to eukaryotic potassium channels [50] and is widely

investigated to understand the mechanism of transport of potassium across the membrane. KcsA

is a tetrameric protein, each subunit contains two transmembrane domains holding a pore region

comprising the selectivity filter, responsible for the permeation and selectivity of the ion channel.

[51][52][53] KcsA channel shares most of the mechanistic properties of C-type inactivation in voltage-

dependent K+ channels [54][55][56][57]. The crystal structures of open/inactivated KcsA reveals a

remarkable correlation between the ion occupancy of the selectivity filter and the degree of opening at

the activation gate. [54] KcsA selectivity filter is stabilized by a hydrogen bound [58] and the disruption

of this ligation favours the conductive conformation of the filter [55][57]. Besides, key interactions were

observed between residues Glu71, Asp80, and Trp67 and a bound water molecule. Glu71 and Asp80

residues stabilize the water molecule [58] and the strength of the hydrogen bound between these

two residues is directly responsible for the initiation of the C-type inactivated state.[57] Additionally

the hydrogen bonding between Trp67 and Asp80 residues determines the rate and the extent of the

C-type inactivation. [57] Also, the Asp80 side chain is accessible to the external solution and its

neutralization severely compromises the KcsA oligomeric stability.[59]

This protein is activated by changes in the intracellular pH controlled by the cytoplasmic domain

of the channel. KcsA is highly selective for potassium ions. Differences between potassium ions and

other cations free energy combined with structural arrangements favour the transport of potassium

ions over other cations.[60][61]

Figure 2.2: Schematic view of KcsA Potassium Channel. a - Top view of the KcsA channel showing four subunits,each in a different colour. A K+ ion is shown in the center. b - Side view of two diagonally opposed subunits.Inner helices (yellow) delimit the permeation pathway. Three K+ ions in the permeation pathway are shown. Inthe selectivity filter are shown the oxygens involved in the coordination of K+ ions [62]

Electrophysiological measurements are the most common approach to study the single-channel

8

behaviour of KcsA through the recording of the current caused by the ion flow across the pore. The

majority of the studies performed resorts in the fusion of liposomes, containing the purified protein

from the cell, with a planar bilayer, [61][63][64][65][66] or the direct transfer of the target protein from

the cell to the planar bilayer. [67][68]

KcsA has been expressed in mammalian cell-lines leading to the understanding of its activation-

inactivation mechanism. [56] More recently, cell-free approaches have been successfully applied for

the expression of KcsA. [69][70][71][72] This advance, has brought improvements for working with

this ion channel, allowing the design of genes with several mutations for structural variants in order to

get a better understanding of its mechanism.

2.1.3 Bacteriorhosopsin channel

Bacteriorhodopsin (bR) is a membrane protein that functions as a light-driven proton pump in

Halobacterium halobium,[73] that converts light energy into an electrochemical proton gradient. As all

retinal proteins, bR fold into a seven-transmembrane helix (named A to G) with short interconnecting

loops, and additionally forms patches of two-dimensional crystals yielding a channel-like structure in

the cell membrane, called purple membrane. [74] The channel is formed by helices B, D, F and G

and is divided into a cytoplasmic and an extracellular half channel by a retinal molecule that forms

a Schiff base with a conserved lysine on helix G.[74][75][76][77] The absorption of a photon, makes

the bR undergo into a catalytic photo cycle, which is intrinsically linked to transport cycle, leading to

the movement of a proton to out of the cell. The cycle is composed of six steps of isomerization, ion

transport, and accessibility change. Thermal intermediates of the cycle are termed J, K, L, M1, M2,

N, O. First, the retinal isomerizes from all-trans to 13-cis configuration, followed by the transfer of the

proton of the Schiff base to Asp-85 and the releasing of a proton to the extracellular side.[74][78][79]

As a result, the blue-shifted M state is observed due to its absorbance at 410 nm. [80][8] Next, the

Schiff base needs to change its accessibility from extracellular to intracellular and the reprotonation

occurs via Asp96 from the cyytoplasmatic side. After the reprotonation, the Schiff base switches back

its accessibility to the extracellular side and the red-shifted N state is formed, absorbing at 570 nm.

Aftewards, the retinal reisomerizes leading to a return to the bR state. [74][77][80]

9

Figure 2.3: Schematic view of the bR photo-cycle and the proton exchange steps. A - Photocycle of bR with thespectral intermediates (K to O) and their approximate time-scales. B - Seven transmembrane helical structureof bR, the location of key residues along the proton-translocation channel, and the accepted sequence of protonexchange events[81]

Bamberg et al. demonstrated the ability of incorporating bR in planar lipid bilayers and successfully

recorded the photocurrent of the channel. [82] Nagel et al. also expressed bR heterologously in

Xenopus laevis oocytes plasma membrane, allowing the study of the electrical properties of the pump

using both voltage clamp and patch clamp methods. [83]

Bacteriorhodopsin is a well-studied integral membrane protein [84] and had been widely expressed

in cell-free systems. Originally, it was reconstituted from precipitated material into membranes [85][86]

but more recent studies demonstrated that bR can be expressed by cell-free translation using E. Coli

extracts in a folded state in the presence of liposomes [87], detergents [88] or nanodiscs [89].

2.2 Cell-Free Protein Synthesis

Membrane proteins (MPs) are currently one of the most important drug targets.[5][8] However the

knowledge about MPs structure and function is limited, and overexpression of this class of proteins in

vivo has been a challenge, often resulting in low yield, cell toxicity, protein aggregation, misfolding and

even cell lysis. [6] Another difficulty is that MPs are embedded in the lipid bilayer, a very complex and

dynamic structure. This limits the use of many standard biophysical techniques to determine structure

and function of MPs, and further complicates their purification, often leading to laborious and hard to

handle processes. [90][91][92] Finally, MPs are generally not soluble in aqueous solutions and need

to surrounded by a medium that satisfy their high hydrophobicity and special synthetic systems, such

as mixed lipid-detergent systems or detergent-lipid micelles and bicelles, are required for in vitro work.

Reconstitution of purified MPs in this type of systems has proven to be challenging.[92] So far, the

10

functional and structural analyses of this extremely difficult class of proteins have been hampered due

to the previously described reasons.

Furthermore, the activity and function of a given MP is not only related to its high-yield produc-

tion, but is strongly dependent on a proper membrane environment. Certain parameters which are

involved in regulation of MPs function must be considered, such as the lipid composition (as well as

hydrophobic chain length, lipid charge, shape, and hydrogen bonding potential of the head group),

and the associated tension, fluidity and curvature of the membrane. Altogether, the membrane will

be the matrix for MPs and provide the framework for the correct adjustment of protein structure and

function.

CFPS offers potential to overcome these limitations. The ability to produce a functional protein in

a test-tube instead of in cells, is the basic principle behind this methodology. [93] CFPS is a simple

method that provides high levels of control, direct access and manipulation of the reaction conditions,

allowing, for example, the addition of accessory reagents such as lipids, folding catalysts, unnatural

animo acids and a variety of amphiphilic molecules due to the completely open nature of the system.

[8][6]

11

Figure 2.4: Schematic comparison between the conventional cell-based expression systems and the cell-freeprotein synthesis systems. Typical steps, from the DNA template until the expressed target protein, are illustrated.The bottlenecks of cell-based systems are given in red. In green are shown examples of additives that can beprovided to CF reactions. [94]

12

2.2.1 CFPS Extract sources

The preparation of a CFPS system requires a cell lysate, an environment full of active biomolecules

capable of supporting many cellular functions, like metabolic pathways and transcription and transla-

tion. [93] Lysates for CFPS systems have been prepared from different sources including mammalian

cells (e.g., rabbit reticulocytes [95][96], HeLa [97], CHO [98], mouse embryonic fibroblasts [99]), wheat

germ [100][101][102][103], insect cells [104][105] and bacteria cells (E. coli [106][107][108][109][110]

[111], Pseudomonas fluorescens [112], Thermococcus kodakaraensis [113]), Drosophila [114], Xeno-

pus oocytes [115] or yeast [116]. However, most of these systems have showed low productivity which

has harmed their applications.

The high productivity and flexibility of wheat germ and E. coli extracts have made them the two

main sources of cell-free extracts. [117] The eukaryotic wheat germ extracts demonstrate higher

stability allowing extended reaction times up to several days. Also, they have very low endogenous

RNase activity, which makes them suitable for translation systems with purified mRNA as a template

and the synthesis of eukaryotic proteins. However, the genetic constructs required for effective ex-

pression are often understudied and complex [7][118]. On the other side, prokaryotic E. coli extracts

are most commonly used in coupled transcription-translation systems with double-stranded DNA as

template. They also shown higher translation rates. Simple and universal translation vectors are

available for this type of extracts, as well as variety of mutants with reduced degradative (nucleases,

proteases, or phosphatases) activities. Yet, E. coli extracts exhibit high rates of degradation of ge-

netic messages (plasmids, PCR products or mRNA), proteins and energy, and they are also more

likely to form protein products aggregates. [7][118] Both systems have similar productivities. [118]

Commercial suppliers of wheat germ extracts or expression systems, such as Cell-Free Sciences or

Roche Applied Science, are available, as well as for E. coli systems, where Roche Applied Science,

Invitrogen, or Qiagen have CF expression systems.

Over the past years, the CFPS extracts have been tremendous improved in terms of buffer compo-

sition [119], energy recycling [120][121][122], the utilization of various mutated cell strains [123][124],

the supplementation of small molecules [125][126], the addition of proteins such as T7 RNA poly-

merase (RNAP) to generate a transcription/translation coupled system [119][127][128] and chaper-

ones or combined chaperones with disulphide isomerase [118] to improve the proper folding of target

proteins.

It is also important to mention that, besides these conventional prokaryotic and eukaryotic CFPS

systems, Ueda and coworkers had developed a promising and interesting approach for producing

recombinant proteins: Protein synthesis Using Recombinant Element (PURE). [129][130] This sys-

tem is based on the use of all recombinant His-tagged translation factors, energy enzymes, energy

sources and aminoacyl-tRNA synthetases, T7 RNA polymerase and ribosomes. These His-tagged

proteins are removed at the end of the synthesis reaction through an affinity chromatography, leav-

ing the expressed recombinant protein as the major synthesized product. [129] This technology is

already commercially available as the PURExpress kit by New England Biolabs [131][132] and has

several advantages over the lysate-based systems, one of which is the capability to easily control the

13

synthesis reaction and to produce highly pure recombinant proteins in one step. Besides, it is possible

to think that a whole artificial protein produced from a synthetic gene could be achieved resorting this

technique. [117] However, due to the use of E. coli extracts, these kits are not well suitable for the

expression of mature mammalian proteins. Glycosylation and other post-translational modifications

of synthesized proteins typical of mammalian cells do not take place in this type of bacterial systems.

2.2.2 Configurations of CFPS reactions

The first generation of cell-free translation and transcription-translation systems was projected in

the batch format. Due to the rapid depletion of the high-energy phosphate pool, that happens even

in the absence of protein synthesis, and the accumulation of free phosphates that inhibit the protein

synthesis, this systems performed in a test-tube are characterized by short lifetimes (approximately

1 hour) and consequently by low yields of production. [118][121] This problem was first overcome

by Spirin and coworkers by the development of the continuous-flow cell-free (CFCF) system, which

relies on the principle of the living cell: continuous supply of the consumable substrates and energy

and continuous removal of the reaction by-products. [133] To realize this strategy reactors, where

the high molecular weight components of the reaction mixture (ribosomes, mRNA, tRNA, translation

factors, aminoacyl-tRNA synthetases and other enzymes) are retained by a permeable membrane

through which the reaction by-products are removed, were developed. Amino acids and nucleotides

were supplied to the reaction chamber by a direct flow. [133][134][135][136][137] The reaction time

was extended to 20 hours and the product yield was increased by two orders of magnitude. [133]

Additionally, the CFCF system offers a high purity of the target protein as well as a high control of the

reaction mixture during the run of the process. [118] However the operational complexities and the

high costs of the reactor make this system extremely impractical.

To address these limitations, a semi-continuous or continuous-exchange cell-free (CECF) system

was developed, in which the exchange between the substrates and the products occurs passively

though a dialysis bag or a flat dialysis membrane. [138][139][140][102] Recently membrane-free

reactors were proposed. For example, the reaction mixture for the translation was encapsulated

into small vesicles. [141] Another example of a membrane-free CECF was described by Endo and

coworkers, consisting in a bilayer system in which the exchange process occurs across the boundary

between two liquid layers, the top one containing the feed solution and the bottom one consisting in

the reaction mixture. [142] This methodology is compatible with a high-throughput format.

Although the CECF system maintains the initial rate of proteins synthesis constant over long pe-

riods, the traditional batch format still offers some advantages, for example, excellent reproducibil-

ity, easy scale-up, convenience of operation and ability to be operated in a multiplex format. [121]

For these reasons, this traditional system have been improved over the past years. [136][139][143]

Swartzs group has continuously been looking for more efficient and economical energy alternatives

to the conventional ATP and/or GTP regeneration systems with some successful results. They de-

veloped the Cytomim system, a prokaryotic in vitro translation method that mimics the cytoplasmic

composition of the host organism, E. coli, and uses pyruvate to regenerate ATP instead of the expen-

14

sive high-energy phosphate compound (PEP), avoiding thereby the formation of inhibitory byprod-

ucts. Besides, the Cytomim systems shows improved economics, greater yields (700 g/mL of

chloramphenicol acetyl transferase (CAT), when only 120 g/mL of the same protein was previously

achieved in a batch reaction system, which also utilizes pyruvate to regenerate ATP [144]), phos-

phate homeostasis, and pH stability. [109] The same group, has also shown that by mutating genes

involved in the catabolism of certain amino acids the efficiency of the cell-free system can also be

increased.[145]

2.2.3 Cell-free expression of membrane proteins

2.2.3.A Cell-free synthesis of membrane proteins as precipitates

A standard CF expression lacks substantial amount of membranes and lipids and as a result no

sufficient amounts of hydrophobic environments are present to keep the MPs soluble after translation

and MPs precipitates are formed. This precipitate reminds the inclusion bodies formed during the

MP synthesis in E. coli cells. [7][146] However, there are some significant differences. For instance,

the precipitates formed via cell-free expression of membrane proteins as precipitates (P-CF) can ef-

ficiently be solubilized with detergents by smooth shaking for few hours [147] and can already result

in functionally folded MPs (multidrug transporter EmrE [147][148] and human histamine-1 receptor

[149]). In contrast, inclusion bodies require strong denaturants for solubilisation, like high concentra-

tions of urea or guanidinium hydrochloride, followed by laborious refolding procedures with excessive

dialysis steps and buffer exchanges in high dilutions. The successful refolding of MPs from inclusion

bodies is so far confined to outer MPs. [146] Precipitates formed through P-CF reactions seem to

present less denatured and aggregated MPs when compared with inclusion bodies. In fact, folded

proteorhodopsin was already observed in a P-CF generated precipitate. [150]

Various mild detergents have been tested for their suitability to solubilize P-CF generated precip-

itates, and some, like dodecyl phosphocholine (DPC) [151], n-Dodecyl--D-maltopyranoside (DDM)

[148][151][152], 1-myristoyl-2-hydroxy-sn-glycero-3-[phospho-rac-(1-glycerol)] (LMPG) [151] and sodium

dodecyl sulfate (SDS) [151], have shown high efficiency in the solubilisation as well as a functional

folding. Additionally, these detergents are appropriate for structural analysis of MPs by nuclear mag-

netic resonance (NMR) spectroscopy. [153] However, the functional solubilization of MPs from pre-

cipitates might not always occur.

2.2.3.B Cell-free synthesis of membrane proteins in presence of detergents

The addition of detergents directly to the reaction vessel could be performed in order to avoid the

precipitation of the synthesized MPs. Supplementation of detergents will result in the formation of pro-

teomicelles, mainly due to nonspecific hydrophobic interactions. In the reaction mixture, detergents

are available at the level of ribosomes making the transport of the synthesized MPs to membranes,

in order to get the appropriate hydrophobic environment, unnecessary. This allows to overcome the

problems originating from the transport and translocations processes that are common in the conven-

tional living cells systems. [154][155]

15

Although the final outcome of cell-free synthesis of membrane proteins in presence of deter-

gents (D-CF) systems strongly depends on the protein being synthesized, some common charac-

teristics to several detergents seems to result in a successful expression. [6][7][117][146][151][156]

The use of mild detergents with relatively low critical micellar concentrations, such as polyoxyethylene-

alkyl-ethers (Brij-derivatives), digitonin, 1,2-diheptanoyl-sn-glycero-3-phosphocholine (DHPC) and n-

dodecyl--D-maltoside (DDM), appears to result in triumphant expression in terms of yield and sol-

ubilisation. [6][146][151] As a general rule, in order to prevent aggregation and non-uniform pro-

teomicelles, the molar ratio between the synthesized proteins and the micelles should not exceed

1.0. [6] However, the type and the final concentration of the supplied detergents will have an im-

portant influence on the quality and the yield of solubilized MPs and each D-CF system should

be optimized for these parameters. Detergents with high critical micelle concentration (CMC), de-

fined as the concentration of detergents above which micelles are spontaneously formed, are usually

not recommended because they must be present in rather high concentrations, which may inhibit

the expression machinery of cell extracts. [6][146][151] However, some exceptions have been re-

ported, and PomA and PomB, which together comprise the stator complex of the motor and func-

tion as a Na+ channel to generate torque, were expressed in the presence of the detergent 3-[(3-

cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS), a zwitterionic detergent with high

CMC. [157]

The major disadvantage of using detergents to solubilize MPs is the low knowledge of how well

proteomicelles mimic their natural environment. For example, the lack of a bilayer environment may

have strong consequences on the structure and the function/functionality of MPs. [92] Additionally,

the denaturating function of detergents makes it difficult to study protein macromolecular structure

and function. Despite these limitations, using detergents to solubilize MPs in CFPS is a convenient

methodology. However, a strategy that allows the synthesis in vitro of MPs in a native-like environment

will be a better solution. [6]

2.2.3.C Cell-free synthesis of membrane proteins in the presence of vesicles and liposomes

The expression of MPs in an environment similar to the natural lipid bilayer has been one of the

major goals of CFPS and a few strategies have been developed. Incorporation of the MPs into lipo-

somes is one of them. This approach requires that the MP has been previously purified and solubilized

in detergent micelles and then reconstituted into liposomes. However the reconstitution must be re-

alized in non-equilibrium conditions which are very difficult to stage. [6] MPs had been reconstituted

into liposomes using the freeze-thaw technique, based on the demonstration that freeze-thawing of

liposomes mixed with proteins enhances the formation of proteoliposomes. [147][158][159] Another

successful methodology is the use of detergent adsorbent particles in the presence of liposomes.

[160][149] Liposomes can be made of a variety of lipids, such as phosphatidylserine, phosphatidyl-

choline, cholesterol, E. Coli lipids or azolectin. [6] The major shortcoming of this approach is the lack

of a single strategy for all MPs, meaning that the protocol must be adjusted for every protein studied.

Another strategy is the addition of unilamellar liposomes to the CFPS reaction vessel. The in-

16

sertion mechanism of proteins into naked membranes is not clear but it is evident that its efficiency

is strongly dependent on the phase transition temperature, the length of the lipid and the hydropho-

bic thickness of the bilayer. [87] Several MPs have been synthesized using this technique such as

stearoyl-CoA desaturase, bacteriorhodopsin, apoCytochrome b5 and the catalytic core of the NADPH

oxidase complex [87][161][162][163][164] and some of them have also been considered good candi-

dates for the delivery of therapeutic proteins. [163][164] The major drawback of unilamellar liposomes

is the inhomogeneity of the samples and the presence of residual multilamellar vesicles. [165] Pro-

cedures to prepare homogeneous liposome populations exist but are not commonly used due to the

need of unusual equipment.[166]

Based on the rationale that most of MPs from Gram-negative bacteria have their final location

defined by processes that depend on the signal recognition particle (SRP) receptor, the SecYEG

translocon and YidC, which are components embedded in the natural target membranes, Kuruma

and coworkers proposed a system where these components were added to the cell-free reaction as

part of E. coli inverted vesicle. [167]

Wuu and Swartz reported a high-yield cell-free methodology for expression of functional inte-

gral membrane proteins using an energy-efficient system supplemented with E. coli inner membrane

vesicles. [168] However, the method relies in high-quality inverted E. coli vesicle samples, which are

difficult to prepare and not commercially available. On the other side, crude vesicles derived from dog

pancreas have been widely commercialized but they have an intense negative effect on the trans-

lational rate of the resulting lysate.[169] Again, the major disadvantage of the use of vesicles is the

heterogeneity of the samples.

2.2.3.D Cell-free synthesis of membrane proteins in presence of nanodiscs

To overcome the limitations from the use of liposomes and vesicles, new strategies based on the

use of nanodiscs or nanolipoprotein particles (NLPs) have been recently developed. NLPs are dis-

coidal nanoparticles consisting on a planar phospholipid membrane bilayer comprised between two

copies of a membrane scaffold protein, which itself constitutes a modified apolipoprotein. [165][170]

The diameters of nanodiscs can vary from 9 to 20 nm, depending on the length of the membrane

scaffold protein, and when compared with liposomes and vesicles, samples are remarkably monodis-

perse and homogeneous (molecular mass deviation less than 5%). [170][171][172][173] A variety of

scaffold proteins have been used to prepare NLPs, such as apolipoprotoeins A1, E and C and insect

lipophorins.[170][172]

17

Figure 2.5: Structure of a nanodisc. Nanodiscs consist in a lipid bilayer (grey) wrapped between two copiesof a scaffold protein (blue). The CFPS synthesized membrane proteins can be incorporated in the lipid bilayer(green). [174]

NLPs present several advantages over the conventional solubilization approaches, previously de-

scribed. [170][175] The membrane scaffold protein provides stability to the MP. NLPs are a better

representation of the phase transition behaviour of biological membranes. Proteins can be extracted

from the nanodiscs in a native functional form. They are accessible from both sides of the lipid bilayer

which could be helpful for ligand binding studies. [8][6] The membrane composition, a factor that can

influence the functionality of the MP, can be defined by a wide range of lipids.[176][177] Additionally,

the preparation of NLPs is a simple procedure and no unconventional instrumentation is required.[6]

An important advantage of NLPs is the fact that after their formation they sustain the soluble tar-

get MP in a detergent-free environment, which makes the purification and functional analysis free of

detergent-based systems limitations. Besides, the protein purification in this system is simplified by

the introduction of a Histidine tag (His-tag) in the membrane scaffold protein.[8]

In one of the cell-free NLP approaches the preformed (empty) NLPs are added to the CFPS

reaction mixture and serve as a membrane support for the MPs. [178][179] Many MPs with relevant

biological importance and various topologies, origins, sizes and proposed roles, have been expressed

in the presence of NLPs and the results showed that all of them exhibit enhanced solubility and

proper folding was observed. [178][179] This methodology has been applied using cell-free extracts

from wheat germ, rabbit reticulocytes and E. Coli cells[178], but only the last ones are available as

commercial kits [179] (MembraneMax Protein Expression Kit, Life Technologies). The major limitation

of this system is the absence of a translocon and a transmembrane potential that are required for the

18

correct folding and insertion of some integral MPs. [180]

In a second approach for CFPS with NLPs, the membrane scaffold protein is co-expressed with

the target MP in the presence of liposomes, resulting in the in situ assembly of the NLPs. [181] The

advantage of this technique is its simplicity and the fact that the NLPs do not need to be manufactured

previously. However, the lipids need to undergo a pre-treatment step to be able to form unilamellar

liposomes. The disadvantage of this method is the complicate purification procedure of the NLP-

protein complexes due to a large set of reaction products with different sizes. Also, the ratio between

the DNA template for the scaffold protein and the DNA template for the target MP needs to be adjusted

for each protein. This system has been implemented in batch and continuous exchange formats.[181]

Figure 2.6: Summary of the current strategies for the cell-free expression of membrane proteins. The Keyhave represented the reagents and products involved in each of the approaches. Not in scale. (a) Expressionin the presence of detergents. If no additional reagents are added, the protein will be integrated in detergentmicelles after expression. (b) Expression of membrane proteins as a precipitate. The solubilization of the proteinprecipitate with detergent can lead to the formation of proteoliposomes. The protein can also be solubilizedin detergent micelles upon the addition of a mild detergent. (c) Expression in the presence of liposomes orvesicles. Liposomes are added directly to the reaction. (d) Expression in the presence of nanodisks or NLPs.(e) Co-expression with scaffold proteins. The target and scaffold proteins are expressed in vitro and protein-NLPcomplexes are then formed in situ. [6]

2.2.4 Applications and future perspectives of CFPS

CFPS in a larger scale can provide unique possibilities for structural analysis of proteins. NMR

spectroscopy and X-ray crystallography can benefit from the advantages of cell-free expression. The

effective incorporation of stable-isotope (13C,15N)-labeled amino acids into proteins synthesized in

CFPS [140][147] allows structure determination of the target proteins in solution by high-resolution

NMR. In a similar way, the cytotoxic unnatural amino acid, selenomethionine, incorporated in a pro-

19

tein via a cell-free system [182] provides an important advantage for the phasing of crystal protein

structure and the atomic structure determination of that protein using X-ray crystallography. In fact,

labelling protocols for NMR-based structural proteomics resorting on wheat germ extracts have al-

ready been established.[183][184]

The incorporation of unnatural and chemically modified amino acids into target proteins through

CFPS, brings new opportunities of development in functional studies, protein engineering and phar-

maceutical studies.[118] A variety of labels, such as fluorescent dyes for functional studies, biotiny-

lated moieties to simplify purification processes, and the ones for structural studies and for posttrans-

lational modifications, can be sequence-specifically incorporated into target proteins.[185] Protein

interaction validation studies could also benefit from the incorporation of labels, for instance through

the combination of a dual labelling with fluorescence cross-correlation spectroscopy. [186]

CFPS could offer an alternative approach for oligomerization studies of MPs by providing two

different DNA templates, with one encoding for the wild-type MP and the other for a derivative that

has been modified by a translation tag. Plasmids of identical incompatibility groups can be used as

DNA template as no replication or selection is needed and even linear DNA fragments generated by

standard PCR would be applicable. CFPS allows controlling the ratios of both protein species just by

titration of the correspondent DNA templates. It would also be possible to add already purified proteins

isolated from other sources in order to analyse their interaction with the cell-free synthesized target

proteins. This enables accurate quantitative experiments that are independent from the expression

rates and proper folding conditions of the assumed interaction partners.[7]

Since the first reports on the direct use of DNA PCR products for programming protein synthesis

in cell-free transcription-translation systems, functional mapping of genomes through expression of

genomic libraries and construction and screening of protein libraries is one of the most important

application of the CFPS.[187][188][189][190] A 96 well format for CFPS with robotic automation was

proposed for this approach, [191] as well as a smaller nano-well chip format. [192] MPs, which

usually have been excluded from common high-throughput proteome expression due to their complex

production, can also benefit from the CFPS systems and develop membrane proteomics.[7] The high-

throughput screening of cloning-independent and engineered proteins can also be improved through

the use of cell-free systems.[193][194]

2.3 Bilayer Lipid Membranes

2.3.1 Cell membrane

The cell membrane is a highly complex and dynamic structure. Biological membranes form an

impermeable barrier between the intracellular cytoplasm and the exterior environment. Biological

membranes consist of a self-assembled lipid bilayer, with sorbed contains proteins, carbohydrates

and their complexes.[195] Phospholipids consist of a hydrophilic polar head and two hydrophobic

hydrocarbon chains. The tails of the phospholipids orient towards each other creating a hydrophobic

environment within the lipid bilayer. This leaves the phosphate groups facing out into the aqueous

20

environment. In the interior of the bilayer, hydrocarbon chains are held together by van der Waals

forces and are in a liquid-crystalline state, in physiological conditions.[196]

The phospholipids show different characteristics regarding the tail length, number of unsaturations

and spatial arrangement, and the head group in terms of charge, shape and size. All these character-

istics will determine the phospholipid shape. Membranes have several types of phospholipids in their

composion such as phosphatidylcholine, phosphatidylserine, phosphatidylethanolamine, and sphin-

gomyelin. The membrane composition is heterogeneous and is responsible for its properties, also

depending on the temperature. Besides phospholipids, there are two other common types of lipids

present in the membrane, cholesterol and glycolipids. Structural functions and permeability, fluidity,

charge and deformability are related to the phospholipids and cholesterol content, and protection from

harsh conditions is a functional role played by glycolipids and sphingolipids.[12][10]

The main components of cell membranes are proteins and, on average, 50% of the membrane

mass is due to them. [197] Membrane proteins regulate the cell communication with the environment

and the transport across the membrane. Ion channels are one type of integral membrane proteins that

are permanently bound to the lipid bilayer via their hydrophobic domains. Besides integral proteins,

biological membranes contain peripheral proteins, temporarily associated to parts of the lipid bilayer

by non-covalent interactions, and lipid-anchored proteins, bounded to fatty acid. [12][13] MPs are the

main target of the majority of the current pharmaceutical drugs, ion channels being the major class of

interest.

2.3.2 Patch Clamp Technique

Currently the patch clamp technique, introduced by Neher and Sakmann in 1976, [198] is still

considered the Gold standard for ion channel measurements.[199] In this method, a glass pipette

is pressed against the cell membrane, while a suction is applied to establish a tight seal (so-called

gigaohm seal) between them. High resolution electrophysiological recordings are possible by placing

one electrode in the pipette buffer solution and the other one in the bath medium around the cell. The

electric current and voltage through the membrane can be accurately controlled and recorded using

a patch-clamp amplifier, which combined with low noise measurements, achieved by the cleanliness

of the pipette tip and the precise shape of the apertures, allows the study of single ion channel

function (figure 2.6). [198] After a mechanically stable seal is formed, several geometric configurations

can be used. The current can be measured when passing through an attached patch (cell-attached

configuration) giving information about ion channels within the environment of the cell, a detach patch

(inside-out or outside-out configurations) where information about ion channels in isolation from the

rest of the cell is obtained, or the whole cell can be measured, providing information about the entire

cell.[200][201] However, patch clamp is a laborious and time-consuming technique that demands a

high skilled operator leading to high costs. Also the technique suffers from a very low throughput,

allowing only the analysis of 10 compounds per week.[202]

21

Figure 2.7: Schematic representation of the patch clamp technique for the whole cell configuration. [5]

To overcome these limitations, in the past years, automated patch-clamp systems have been

developed. [202] Here, individual cells are sucked into a micrometer-size aperture etched in a planar

substrate, instead of into the glass pipette, to form a cell patch. Although this technique preserves

the high information content of electrophysiological recordings, while increasing the throughput of the

analyses and decreasing the level of technicality, it still utilizes cell models.[10][202]

2.3.3 Bilayer Lipid Membrane Models

A bilayer lipid membrane is a simplified and controlled model of the cell membrane and can be arti-

ficially created due to the property of lipids to spontaneous self-assemble and remain in a bilayer form

through hydrophobic interactions, in an aqueous environment. The use of bilayer lipid membranes

models offers several advantages over cell based models. They allow full control and manipulation of

the experimental conditions, such as lipid composition, solutions on both sides of the membrane and

the ion channel introduced. All these parameters could affect the membrane properties and conse-

quently the ion channel function and the use of a bilayer lipid membrane model allows the study of the

interactions between ion channels and the membrane environment by systematically varying these

parameters. Artificial models do not require cellular cultures, decreasing the cost and the technicality

of the assays.[10][13]

Several approaches have been developed using bilayer lipid membranes, such as suspended and,

supported membranes, vesicles, and droplet bilayer.

22

Figure 2.8: Schematic representation of bilayer lipid membrane approaches. a) Vesicles, b) supported bilayersseparated from the substrate by a hydrophilic polymer layer, c) droplet interface bilayers (DIBs), and suspendedbilayer achived by d) painting technique that forms solvent-containing bilayers, or e) Montal-MA 1

4ller technique,

which bilayers are solvent-less. [10]

Suspended

Suspended bilayers can be formed by the painting or Muller-Rudin technique. [203] Lipids are

dissolved in an organic solvent, like n-decane, and manually deposited across a microaperture with a

brush, in a hydrophobic substrate, such as Teflon, separating two reservoirs. Painting the lipid-solvent

solution in the aperture leads to the formation of a thick film, that spontaneously thins down to form

a lipid bilayer in the center of the aperture with a annular region at the edge of the aperture,, that

stabilizes the membrane. The resulting bilayers can incorporate solvent molecules since the lipids

are dissolved in a non-volatile solution. This can change the membrane thickness, fluidity and be

prejudicial in the reconstitution of ion channels. Additionally, this method is often irreproducible.

Solvent-free lipid membranes can be formed using the Montal-Mueller technique. [204] In the

method, lipids, in a solution with a volatile solvent, self-assemble into a monolayer at the liquid-air

interface on top of the buffer solution in the two reservoirs, by evaporation of the solvent. Then, the

level of buffer in the reservoirs is slowly raised above the aperture, where the two lipid monolayers

contact with each other and assemble into a bilayer membrane. The formed bilayers have no solvent

between the two leaflets, and with this technique is possible to form asymmetric bilayer. However, the

method is still irreproducible and the absence of annulus affects the stability of the membrane.[12]

Finally, lipid vesicles can be used to create suspended bilayer, by flushing a solution with this vesi-

cles through a planar hydrophobic substrate containing apertures. After the rupture of the vesicles,

a bilayer is formed across the aperture. This result in solvent-free membranes. Also, the great ad-

23

vantage of this technique is that proteins can be incorporated in the vesicles so that they are directly

integrated in the final bilayer.[10][12]

Suspended lipid bilayers allow the access to both sides of the membrane, making electrophys-

iological measurements possible with the insertion of an electrode in each of the reservoirs. The

accessibility of the two reservoirs also allows the control of both buffer solutions and consequently the

lipid composition of both sides of the membrane. The major drawback of suspended membranes is

their low stability.

Supported

Supported lipid bilayers are formed on the surface of a solid substrate. These bilayers can be

created by the Langmuir-Blodgett [205] and Langmuir-Schaefer techniques. Another process that

leads to the formation of these membranes is the rupture of a vesicle over a smooth and hydrophilic

substrate.[206] Usually the bilayer is separated from the substrate by a thin water layer (1-2 nm thick),

and the transmembrane proteins can interact with the surface, which can alter their properties and

functions. Supported lipid bilayers are very stables; however, only one side of the bilayer is accessible,

not allowing control of on both sides of the buffer solutions and making electrophysiological analysis

difficult. Besides, as already mentioned, proteins can see their function and properties altered which

disrupts their study in this models. [10][12]

Vesicles

Lipid vesicles or liposomes are spherical lipid membranes that encloses an aqueous solution.

They can be used as model cell membrane system and as cell-size container. For example, in image

phase separation phenomena in the bilayer with respect to lipid composition. [207]

Liposomes are formed by self-assembly of lipids making it complicate to control their proper-

ties, such as size, and their round shape makes impossible the direct access to the inner compart-

ment, either for changes in the solution either insertion of electrodes for electrophysiological measure-

ments. However their round shape mimics the shape of a cell, which does not happen with planar

membranes.[12][13]

Droplet interface bilayer

Droplet interface bilayers (DIBs) are created by bringing two aqueous droplets in contact in an

oil-lipid mixture. A lipid monolayer spontaneously forms at the water-oil interface of each droplet. An-

other method, consists in filling the aqueous droplets with liposomes, which fuse with the water-oil

interface to form a monolayer and finally symmetric or asymmetric bilayer, without external interven-

tion. Alternatively, droplet-on-hydrogel bilayers can be formed by replacing one droplet for a planar

hydrophilic substrate covered by a lipid monolayer. [208]

Although the solution inside the droplet is not directly accessible, these systems have very high

stabilities and the formation of the bilayer is automatable.

24

2.3.4 Miniaturized Bilayer Platforms

Miniaturized and microfluidic devices are a particularly interesting platform for bilayer lipid mem-

branes experiments. Due to their high level of integration, easy parallelization of the assays, com-

patibility with automated interfaces, the reduced sample volumes leading to the decrease of assays

costs, miniaturized devices exhibit several advantages for drug screening in general and for bilayer

experimentation and membrane proteins study in particular. [209] Smaller apertures, fabricate using

micro and nanofabrication techniques, leads to increased stability of suspended bilayers and reduced

noise level. Furthermore, microfluidic platforms allow the preparation of suspended bilayers in a hori-

zontal configuration, which makes combined electrophysiological measurements and high-resolution

imaging possible. In the past decade, various miniaturized devices for BLMs have been developed.

Multiplexed, automatable, stable and reproducible devices are the major goal for this approach. Be-

sides, analysis of important ion channels in the microfluidic bilayer platform must be validate.

25

26

3Materials and Methods

Contents3.1 Cell-Free Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283.2 Bilayer Lipid Membrane Experiments in a microfluidic device . . . . . . . . . . . 33

27

3.1 Cell-Free Protein Synthesis

3.1.1 E. coli strains and vectors used

One Shot TOP10 Chemically Competent E. coli cells (Invitrogen) (table 3.1) were used for DNA

amplification and cloning reaction of KcsA into a new plasmid.

Table 3.1: Detailed description of One Shot TOP10 Chemically Competent E. coli cells genome.

E. coli Genome Description

One Shot TOP10 ChemicallyCompetent E. coli

F- mcrA (mrr-hsdRMS-mcrBC)80lacZM15 lacI74 recA1araD139 (ara-leu) 7697 galU galKrpsL (StrR) endA1 nupG -

KcsA was expressed in pQE60 with C-terminal His-tag. In order to perform the cell-free protein

synthesis (CFPS) experiments, the pEXP5 expression vector with C-terminal His-tag (pEXP5-CT)

was used. This type of expression vector was used due to the necessity of a T7 promoter for the

CFPS reaction.

3.1.2 Transformation of competent cells

A vial of One Shot TOP10 chemically competent E. coli cells was thawed on ice. Cells were gently

re-suspended and DNA solution (1 g) was added to the cells. The tube with cells was incubated

on ice for 20 min and then heat-shocked for 30 s in a 42oC water bath. Afterwards, the cells were

incubated another 2 min on ice. Then, 250 L of Super Optimal Broth (S.O.C Medium) was added and

the vial was placed in a shaking incubator (37oC, 225 rpm) for 1 h. At the end of the incubation period,

aliquots of 25-50 L of cells were plated out on a pre-warmed LB-agar selective plate containing 100

g/mL ampicillin. The LB-agar plates were incubated overnight at 37oC.

3.1.3 Purification of plasmid DNA

A single colony, from one LB-agar plate, was isolated and inoculated with 10 mL of LB medium

containing 100 g/mL ampicillin. Then the tube was incubated in a shaking incubator (37oC, 225 rpm)

overnight.

The plasmid DNA was purified resorting to QIAprep Spin Miniprep Kit (Qiagen). Bacterial culture

was pelleted by centrifugation (8000 rpm) for 5 min at room temperature and resuspended in 250 L

of Buffer P1, previously supplemented with LyseBlue reagent (ratio 1 to 1000) and RNase A solution,

then transferred to a microcentrifuge tube. Successively, 250 L of Buffer P2 and 350 L of Buffer

N3 were added, and gently mixed. The solution was centrifuged for 10 min at 13000 rpm and the

supernatant was applied to the QIAprep spin column and centrifuged for 45 s at 13000 rpm, the flow-

through was discarded. Afterwards, 500 L of Buffer PB were added, followed by 750 L of Buffer

PE. Each addition step was followed by a centrifugation (13000 rpm) for 45 s. To elute the DNA, 50

28

L of Buffer EB (10 mM Tris.Cl, pH 8.5) were added to the center of the QIAprep spin column and

centrifuged for 1 min.

3.1.4 Primers design

Taking into account the results of the sequencing of the vector pQE60/KcsA two pairs of primers

(A and B; C and D) were designed.

Table 3.2: Primers designed for PCR reaction.

Identification of primer Primer Description

A - Forward primer 1 5 ACCATGGCACCCATGCTG 3

B - Reverse primer 1 5 CCAAGCTCAGCTAATTAAGC 3

C - Forward primer 2 5 GTTTCTTACCATGGCACCCATGCTG 3

D - Reverse primer 2 5 GTTTCTTGCTCAGCTAATTAAGC 3

3.1.5 Polymerase chain reaction (PCR)

A PCR reaction was realized to obtain an appropriate product with the KcsA gene from the plasmid

vector, pQE60/KcsA, and posterior cloning into the suitable vector for CFPS reaction, pEXP5-CT. The

pQE60/KcsA plasmid (10 ng) were mixed with 10X PCR Buffer (100 mM Tris-HCl, pH 8.3 (at 42oC),

500 mM KCl, 25 mM MgCl2, 0.01% gelatin) (5 L), dNTP Mix (12.5 mM dATP, 12.5 mM dCTP, 12.5

mM dGTP, 12.5 mM dTTP, neutralized at pH 8.0 in water) (0.5 L), PCR primers (1 M each), Taq

Polymerase (1 U/L) and water to a final volume of 50 L. The mixture was pre-heated at 95oC for

5 min and then subjected to a PCR cycle, denaturation - 30 s at 95oC, annealing - 30 s at 53oC,

and extension - 1 min/kb at 72oC, repeated 30 times. After the last cycle a final extension step was

performed at 72oC for 7 min. The resulting PCR product was then placed at 4oC for short-term

storage.

3.1.6 TOPO Cloning

After production of the desired PCR product, this product was cloned into the pEXP5-CT/TOPO

vector. Fresh PCR product (2 L), salt solution (1.2 M NaCl, 0.06 M MgCl2) (1 L), water to a final

volume of 5 L and TOPO vector (1 L) were added in this order to the reaction vessel and gently

mixed and incubated for 5 minutes at room temperature (22-23oC). The reaction was then placed on

ice and was followed by the transformation in competent cells as described in section 3.1.2.

3.1.7 Concentration measurements

The concentration of DNA template after the purification of the plasmid vector was determined

using a NanoDrop 1000 Spectrophotometer (Thermo Scientific). With the sampling arm open, 1.5 L

of the sample was pipetted onto the lower measurement pedestal. The sampling arm was then closed

and the spectral measurement was initiated using the operating software on the computer

29