ISSN 0355-1180

UNIVERSITY OF HELSINKI

Department of Food and Environmental Sciences

EKT Series 1714

Molecular cloning, expression and characterization of two potential antibacterial

fusion hydrophobins in Pichia pastoris

Xinyu Jiang

Helsinki 2015

HELSINGIN YLIOPISTO ––– HELSINGFORS UNIVERSITET ––– UNIVERSITY OF HELSINKI

Tiedekunta/Osasto – Fakultet/Sektion – Faculty

Faculty of Agriculture and Forestry

Laitos/Institution– Department

Department of Food and Environmental Sciences

Tekijä/Författare – Author

Xinyu Jiang

Työn nimi / Arbetets titel – Title

Molecular cloning, expression and characterization of two potential antibacterial fusion hydrophobins in Pichia

pastoris

Oppiaine /Läroämne – Subject

Food Safety (Food Chemistry)

Työn laji/Arbetets art – Level

M.Sc. Thesis

Aika/Datum – Month and year

September 2015

Sivumäärä/ Sidoantal – Number of pages

69

Tiivistelmä/Referat – Abstract

Biofilms constitute a successful protection mechanism for planktonic bacterial cells to survive in hostile

environments. To date, biofilm-associated infections in medical devices represent a major cause of morbidity and

mortality among patients. As a potential candidate for anti-biofilm therapy, fungal hydrophobins provide new

solutions to manipulate the physical and chemical properties of surfaces, which in turn may give protection against

bacterial colonization. However, in practice, native hydrophobin coatings generally have no impact on bacterial

surface colonization, because of the lack of being antibacterial by these fungal proteins themselves. The aim of this

study was to explore the feasibility of using recombinant fusion hydrophobins to control bacterial growth.

In this study, the class I hydrophobin hgfI gene isolated from the edible mushroom Grifola frondosa was in frame

fused with two antimicrobial peptide genes (bac8c and p11-5), respectively, and subsequently cloned into the

corresponding expression vectors with a view to obtain two recombinant fusion hydrophobins, Bac8c-HGFI and

P11-5-linker-HGFI. These two chimeric genes were separately expressed in Pichia pastoris under the regulation of

alcohol oxidase 1 promoter. SDS-PAGE and immunoblot analyses confirmed that these two fusion proteins were

successfully expressed and secreted into the culture medium. Minimum inhibitory concentration (MIC) test

demonstrated that the highly active antimicrobial peptide Bac8c became inactivated when it was fused with the

hydrophobin HGFI. Interestingly, the hydrophobin HGFI gained an acquired antibacterial nature when it was fused

with the antimicrobial peptide P11-5 through a 10-mer flexible polypeptide linker, with the MIC of 100 μg/ml

against Escherichia coli.

To the best of my knowledge, this study presents the first heterologous expression of an antibacterial fusion

hydrophobin in P. pastoris. This finding in combination with surface modification mediated by hydrophobin may

broaden the current approaches used for anti-biofilm therapies.

Avainsanat – Nyckelord – Keywords

Hydrophobin; antimicrobial peptide; fusion expression; polypeptide linker; self-assembly; Pichia pastoris

Säilytyspaikka – Förvaringställe – Where deposited

Viikki Science Library, University of Helsinki

Muita tietoja – Övriga uppgifter – Additional information

This study was supported by Magnus Ehrnooth Foundation; EKT Series 1714

PREFACE

This Master’s thesis project in molecular biology was carried out at the Department of

Food and Environmental Sciences at the University of Helsinki from May 2014 to

December 2014. This study was under the supervision of Prof. Per Saris in his laboratory.

Here I would like to express my sincere gratitude to my supervisor Prof. Per Saris for

providing instructions and comments, and offering strong support during the entire thesis

work. I also would like to thank Prof. Marina Heinonen and the program coordinator Tiina

Naskali for their support during my study.

My special thanks are given to my beloved parents and friends for their consistent

encouragement and support through all these years.

LIST OF ABBREVIATIONS

ABS

α-MF

AMP

Amp

AOX

AP

ATCC

ATP

ATPS

BCIP

BMG

BMM

bp

Da

ddH2O

DGGE

DNA

dNTP

dsDNA

EDTA

EG

Fc

FESEM

H+L

HAMBI

HBOEC

HPLC

IgG

kbp

LB

aqueous biphasic system

alpha-mating factor of Saccharomyces cerevisiae

antimicrobial peptide

ampicillin

alcohol oxidase

alkaline phosphatase

American Type Culture Collection

adenosine triphosphate

aqueous two-phase system

5-bromo-4-chloro-3-indolyl phosphate

buffered minimal glycerol

buffered minimal methanol

base pair

dalton

double-distilled water

denaturing gradient gel electrophoresis

deoxyribonucleic acid

deoxynucleotide

double-stranded deoxyribonucleic acid

ethylenediaminetetraacetic acid

endoglucanase

fragment crystallization region

field emission scanning electron microscopy

heavy + light chains (antibody)

culture collection center, University of Helsinki

human blood outgrowth endothelial cells

high performance liquid chromatography

immunoglobulin G

kilobase pair

Luria Bertani

LPS

MIC

NAD+/NADH

NBT

Ni-NTA

OD

PBS

PCR

PEG

RE

RNA

SDS

ssDNA

PAGE

SOC

T4

TAE

TEM

TFA

Tris

WCA

XPS

YPD

YPDS

lipopolysaccharide

minimum inhibitory concentration

nicotinamide adenine dinucleotide

nitro blue tetrazolium chloride

nickle-nitrilotriacetic acid

optical density

phosphate buffered saline

polymerase chain reaction

polyethylene glycol

restriction endonuclease

ribonucleic acid

sodium dodecyl sulfate

single-stranded deoxyribonucleic acid

poly acrylamide gel electrophoresis

super optimal broth with catabolite repression

T4 bacteriophage

Tris-acetic acid-EDTA

transmission electron microscopy

trifluoroacetic acid

tris-hydroxymethyl aminomethane

water contact angle

X-ray photoelectron spectroscopy

yeast extract-peptone-dextrose

yeast extract-peptone-dextrose-sorbitol

i

TABLE OF CONTENTS

1 INTRODUCTION .............................................................................................................. 1

2 LITERATURE REVIEW ................................................................................................... 3

2.1 Bacterial biofilms ..................................................................................................... 3

2.2 Hydrophobins ........................................................................................................... 6

2.3 Antimicrobial peptides ............................................................................................ 10

2.4 The Pichia pastoris protein expression system ...................................................... 16

EXPERIMENTAL RESEARCH ......................................................................................... 21

3 AIMS OF THE STUDY ................................................................................................... 21

4 MATERIALS AND METHODS ...................................................................................... 21

4.1 Materials ................................................................................................................. 21

4.1.1 Bacteria and yeast strains, plasmids, oligonucleotides and primers ............. 21

4.1.2 Antimicrobial peptides, chemical reagents and culture media components . 22

4.2 Methods .................................................................................................................. 22

4.2.1 Isolation and purification of plasmid DNA from Escherichia coli .............. 23

4.2.2 DNA agarose gel electrophoresis ................................................................. 23

4.2.3 Construction of pP-secSUMOpro3-bac8c-hgfI ............................................ 24

4.2.4 Construction of pPIC9-p11-5-linker-hgfI ..................................................... 25

4.2.5 Transformation of electrocompetent Escherichia coli DH5α cells .............. 26

4.2.6 Colony PCR .................................................................................................. 27

4.2.7 Transformation of electrocompetent Pichia pastoris cells ........................... 27

4.2.8 Inducible expression of recombinant fusion hydrophobins .......................... 29

4.2.9 Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE)

of proteins .............................................................................................................. 29

4.2.10 Western blot analysis of the recombinant fusion hydrophobin

SUMO3-Bac8c-HGFI ............................................................................................ 30

4.2.11 Purification of the recombinant fusion hydrophobin SUMO3-Bac8c-HGFI

............................................................................................................................... 31

4.2.12 Purification of the recombinant fusion hydrophobin P11-5-linker-HGFI .. 31

4.2.13 Dot blot analysis of the recombinant fusion hydrophobin

P11-5-linker-HGFI................................................................................................. 32

ii

4.2.14 Protein quantification ................................................................................. 32

4.2.15 Minimum inhibitory concentration test ...................................................... 32

4.2.16 Tools for data analyses................................................................................ 33

5 RESULTS ......................................................................................................................... 34

5.1 Construction of pP-secSUMOpro3-bac8c-hgfI ...................................................... 34

5.2 Expression of the recombinant fusion hydrophobin SUMO3-Bac8c-HGFI .......... 35

5.3 Minimum inhibitory concentrations of the recombinant fusion hydrophobin

Bac8c-HGFI ................................................................................................................. 36

5.4 Construction of pPIC9-p11-5-linker-hgfI ............................................................... 37

5.5 Expression of the recombinant fusion hydrophobin P11-5-linker-HGFI ............... 38

5.6 Minimum inhibitory concentration of the recombinant fusion hydrophobin

P11-5-linker-HGFI........................................................................................................ 39

6 DISCUSSION ................................................................................................................... 41

6.1 Molecular cloning, expression and characterization of the recombinant fusion

hydrophobin Bac8c-HGFI ............................................................................................ 41

6.2 Molecular cloning, expression and characterization of the recombinant fusion

hydrophobin P11-5-linker-HGFI .................................................................................. 44

6.3 Future work: specific-protease-enhanced antibacterial fusion hydrophobins ........ 46

7 CONCLUSIONS .............................................................................................................. 48

8 REFERENCES ................................................................................................................. 49

1

1 INTRODUCTION

Bacterial biofilms are well-organized bacterial cell aggregates on various surfaces, which

constitute a successful protection mechanism for planktonic bacterial cells to survive and

proliferate in hostile environments (Donlan 2002; Simões et al. 2010). To date,

biofilm-associated infections in medical devices represent a major cause of morbidity and

mortality among patients (Shirtliff and leid 2009; Simões et al. 2010; Chen et al. 2013a).

As a result, various biofilm control approaches, generally focus on changing the surface

characteristics, have been applied to practice (Champ et al. 1987), but often toxic

substances are used (Evans et al. 1995; Chen et al. 2013a). The use of biocompatible

bacteria-repellent agents such as hydrophobins, fluoropolymers and polydimethylsiloxanes

(Rieder et al. 2011; Brady 2000; Krishnan et al. 2008) to prevent bacterial colonization on

surfaces, thus negating the use of toxic substances is a new biofilm control strategy over

the last fifteen years.

Hydrophobins are a family of small adhesive proteins exclusively produced by filamentous

fungi and play a crucial role in fungal growth and development (Wösten and Scholtmeijer

2015; Wessels 1997). Apart from rich in hydrophobic amino acids, all of these fungal

proteins possess eight cysteine residues at conserved positions that form four

intramolecular disulfide bridges (Linder et al. 2005). Hydrophobins are extremely

surface-active proteins; they are able to self-assemble into stable monolayers at

hydrophilic-hydrophobic interfaces (Wösten 2001), and thereby altering the surface

wettability of solids, from hydrophilic to hydrophobic and vice versa (Wösten and de

Vocht 2000; Lugones et al. 1996). Due to these unique properties, many potential

applications have been proposed for hydrophobins, including uses as emulsifiers in food

products (Cox et al. 2008) and anti-biofilm agents (Rieder et al. 2011).

However, in practice, native hydrophobin coatings generally have no impact on bacterial

surface colonization, because of the lack of being antibacterial by these fungal proteins

themselves (Rieder et al. 2011). This finding has led to numerous proposals in developing

fusion hydrophobins that tethered with antibiotics, specific enzymes or antimicrobial

2

peptides to avoid or retard bacterial colonization on surfaces (Rieder et al. 2011). As a

potential candidate, antimicrobial peptides possess preferable characteristics in comparison

with antibiotics, including broad-spectrum antimicrobial activity, rapid onset of cell killing

(Hancock and Sahl 2006), and low propensity to cause drug resistance among pathogens

(Steckbeck et al. 2014).

The aim of this study was to explore the feasibility of using recombinant fusion

hydrophobins to control bacterial growth. In the first part of the thesis, bacterial biofilms

and the corresponding control strategies, hydrophobins and the potential applications,

antimicrobial peptides, especially antimicrobial peptides (Bac8c and P11-5) used in this

study, and the Pichia pastoris protein expression system are reviewed. In the second part of

the thesis, methods for molecular cloning, heterologous gene expression, protein

identification, purification and characterization are described. In the final sections, the

antibacterial potential of these recombinant fusion hydrophobins are evaluated and

discussed.

3

2 LITERATURE REVIEW

2.1 Bacterial biofilms

Bacterial biofilms are well-organized bacterial cell aggregates on various surfaces, such as

metals, plastics, biological tissues, implant materials and dental surfaces (Donlan 2002),

which constitute a successful protection mechanism for planktonic bacterial cells to

survive and proliferate in hostile environments (Simões et al. 2010). There is no denying

that some bacterial biofilms (e.g. gut flora) are essential to human health, physiology and

development, like Eubacterium spp., Bifidobacterium spp. and Ruminococcus spp.

(Neufeld et al. 2009). However, under most circumstances, biofilms by human bacterial

pathogens on indwelling medical devices, artificial implants and daily care products that

are directly in contact with the human body, such as bile stents, urinary catheters, artificial

skeletons and joints, and contact lens pose a critical medical problem (Simões et al. 2010;

Costerton et al. 2005), which in turn may lead to acute infections and chronic illnesses

(Bryers 2008; Bjarnsholt 2013). Many pathogenic bacteria can form biofilms on surfaces,

including both gram-positive (e.g. Listeria monocytogenes, Staphylococcus aureus and

Enterococcus faecalis) and gram-negative (Escherichia coli, Pseudomonas aeruginosa and

Klebsiella pneumoniae) bacteria (Chen et al. 2013a).

One well-accepted process of bacterial biofilm development is characterized by 5 stages

(Hall-Stoodley et al. 2004) (Figure 1). (1) Reversible attachment of bacteria to surfaces. In

this stage, planktonic bacterial cells use a variety of extracellular organelles and sticky

membrane proteins for attaching to surfaces, including flagella (Daniels et al. 2004), pili

(Sauer and Camper 2001) and stalks (Renner and Weibel 2011). (2) Irreversible attachment

of bacteria to surfaces. Here bacteria are embedded within a ‘sticky scaffold’, a matrix of

extracellular polymeric substances (EPS) that mainly contain nucleic acids, peptidoglycan,

lipids/phospholipids and lipopolysaccharides (Sutherland 2001), and irreversibly cling to

surfaces (Flemming and Wingender 2010). (3) Early development of biofilm architectures.

Once bacterial cells are settled down on surfaces, they start to proliferate, accumulate and

recruit free-floating bacterial cells from the surrounding environment (Stoodley et al. 2002),

resulting in the formation of bacterial microcolonies with a diameter of tens or hundreds of

4

microns (Renner and Weibel 2011). Meanwhile, the EPS matrix acts as a ‘shield’ to protect

the encapsulated bacterial cells from adverse factors in the surrounding environment, such

as antibiotics, oxidizing agents, ultraviolet radiation (Flemming and Wingender 2010;

Simões et al. 2005). (4) Maturation of biofilm architectures. Bacterial cells are further

‘glued’ together by EPS, leading to the formation of complex three-dimensional biofilm

structures. For example, water channels that help to distribute nutrients and disposal of

waste products within or outside of the biofilm matrix (Stoodley et al. 1994). (5)

Detachment and dispersion of bacteria from mature biofilms. Bacterial cells detach from

mature biofilms and subsequently disperse into the surrounding environment, where they

may act as ‘germs’ for a new round of biofilm development (Costerton et al. 1987; Fey et

al. 2010).

Figure 1. Schematic diagram of bacterial biofilm development on a surface. (1) Reversible attachment of

bacteria to surfaces. (2) Irreversible attachment of bacteria to surfaces. (3) Early development of biofilm

architectures (microcolonies). (4) Maturation of biofilm architectures. (5) Detachment and dispersion of

bacteria from mature biofilms (Figure adapted from Stoodley et al. 2002).

Bacterial biofilm inhibitors as potential drugs

Bacterial cells growing in biofilms are highly resistant to antibacterial agents (Smith 2005;

Olson et al. 2002). This resistance is supposed to be due to a lower reproduction rate of

bacterial cells within biofilms and the barrier effect of EPS shell (Davies 2003; Lewis 2010;

Stewart and Costerton 2001). For example, Ito et al. (2009) found that bacteria Escherichia

coli showed an increased ampicillin resistance when it was grown as biofilms on an abiotic

surface. Usually these biofilm-associated bacterial cells can be between 100-1000 times

5

more resistant to antibiotics when compared with their planktonic counterparts (Gristina

1987), which makes them difficult to be eradicated with conventional antibiotic therapies

(Chen et al. 2013a). In addition, overuses of antibiotics in bacterial biofilm therapies may

give a rise to the antibiotic resistance among bacterial pathogens (Weinstein 2001).

Antibiotic resistances in Escherichia coli, Staphylococcus aureus and Klebsiella

pneumoniae such as multiple beta-lactam antibiotic resistance is one of the most urgent

public health problems nowadays (Muir and Weinbren 2010; Yong et al. 2009; Grundmann

et al. 2006).

The initial stage of bacterial biofilm development is a surface-dependent reversible step;

surface characteristics like wettability (Privett et al. 2011), roughness (Truong et al. 2010)

and protein adsorption capacity (Stallard et al. 2012) may play an important role in

facilitating bacterial colonization on surfaces. Based on these postulations, a research

group at Harvard University developed a novel bacteria-repellent surface coating agent

called SLIPS (Slippery-Liquid-Infused Porous Surfaces) (Wong et al. 2011). In the

practical study, significant inhibition of Pseudomonas aeruginosa, Escherichia coli and

Staphylococcus aureus biofilm development was observed on a SLIPS-coated surface,

under both static and realistic flow cultivation conditions (Wyss Institute, Harvard

University). Weickert et al. (2011) coated plastic bile stents with the recombinant fusion

hydrophobin H*Protein A (from BASF SE), decreased microbial adhesion and serum

protein adsorption were observed on these hydrophobin-treated bile stents.

Other attempts have been focused on giving surfaces with an acquired antibacterial activity.

This attempt is achieved primarily by incorporating antibacterial agents into/onto surface

materials, such as antibiotics (Hickok and Shapiro 2012; Raad et al. 1997; Liu et al. 2012),

chemicals (Jaramillo et al. 2012) and antimicrobial peptides (Costa et al. 2011; Tan et al.

2014).

6

2.2 Hydrophobins

Hydrophobins are a family of small (about 100 amino acids), amphipathic proteins, which

are exclusively produced by filamentous fungi (Bayry et al. 2012; Wösten and

Scholtmeijer 2015; Linder et al. 2005) and play a crucial role in fungal growth and

development (Wösten and Willey 2000; Wessels 1997). These fungal proteins have been

regarded as one of the most surface-active proteins known (Linder et al. 2005); they are

able to self-assemble into amphipathic monolayers at hydrophobic-hydrophilic interfaces,

for example, between air and water (Wösten et al. 1993), water and oil (Wang et al. 2004),

hydrophobic surfaces and water, and hydrophilic surfaces and air (Wösten et al. 1994), and

thereby altering the surface wettability of solids, from hydrophilic to hydrophobic

(Lugones et al. 1996) and vice versa (Wösten et al. 1993) (Figure 2).

Figure 2. Self-assembly of hydrophobins on surfaces. (A) Hydrophobin coating makes the hydrophobic

surface hydrophilic (water contact angle [WCA], ranging between 22 and 63°). (B) Hydrophobin coating

makes the hydrophilic surface hydrophobic (WCA, 110°). (1) If the WCA is smaller than 90°, the surface is

considered as hydrophilic. (2) If WCA is greater than 90°, the surface is considered as hydrophobic. (3) ○

hydrophilic region ● hydrophobic region (Figure adapted from Wösten and de Vocht 2000).

Hydrophobins show very low amino acid sequence homology in general, but all of them

contain eight cysteine residues at conserved positions that form four intramolecular

disulfide bridges with the conserved pattern (cys1-cys6, cys2-cys5, cys3-cys4, and

cys7-cys8) (Kwan et al. 2006; Linder et al. 2005) (Figure 3). There are two categories of

hydrophobins, class I and class II, which are firstly classified by Wessels (1994) based on

biophysical properties, hydropathy patterns (Kyte and Doolittle 1982), and film-forming

patterns of hydrophobins (Wessels 1997). Class I hydrophobins, such as SC3 of

7

Schizophyllum commune, EAS of Neurospora crassa and HGFI of Grifola frondosa can

form rigid and highly insoluble aggregates on hydrophilic surfaces (e.g. mica and

polystyrene plates) (Lugones et al. 1996), and can only be dissociated by strong acids (e.g.

100% formic acid or trifluoroacetic acid). In contrast, class II hydrophobins, including

HFBI and HFBII of Trichoderma reesei can form less stable monolayers on hydrophilic

surfaces, and can easily be dissociated by 60% ethanol or 2 % SDS (Linder et al. 2005;

Wösten et al. 1993).

Figure 3. Ribbon diagram of the class I hydrophobin EAS of Neurospora crassa (Figure adapted from Bayry

et al. 2012).

Application prospects of hydrophobins

Due to their amphipathic nature and self-assembly characteristics, many potential

applications of hydrophobins have been proposed (Linder et al. 2005; Linder 2009; Wösten

and Scholtmeijer 2015; Khalesi et al. 2012). Here applications in biotechnology and food

industry are briefly summarized, including uses in protein/peptide immobilization or

purification, food production and quality control, and anti-biofilm therapy.

Immobilization of proteins and peptides as hydrophobin fusions

The stable nanostructure of hydrophobin assembles on surfaces makes them suitable for

immobilization of proteins/peptides (Linder et al. 2005). Niu et al. (2012a) have studied the

surface modification characteristics of the class I hydrophobin HGFI of Grifola frondosa,

and its fusion with a short human blood outgrowth endothelial cells (HBOECs) binding

ligand, TPS (NH2-TPSLEQRTVYAK). The TPS-linker-HGFI fusion could be efficiently

and rigidly immobilized onto a hydrophobic surface, as analyzed by water contact angle

(WCA) and X-ray photoelectron spectroscopy (XPS) measurements. More importantly, the

HBOECs binding activity of TPS was highly retained upon the immobilization. Linder et

8

al. (2002) used two class II hydrophobins HFBI and HFBII of Trichoderma reesei to

immobilize the cellulose endoglucanase I catalytic core (EGI-core) of T. reesei, onto both

silanized glass and Teflon™ surfaces. The enzyme activity was remained unchanged after

the immobilization.

Purification of proteins as hydrophobin fusions in aqueous two-phase systems

Aqueous two-phase system (ATPS), also called aqueous biphasic system (ABS) is

frequently employed as a nondestructive method for purification of labile proteins (e.g.

membrane proteins and enzymes) at large scales. The amphipathic nature of hydrophobins

makes them suitable as a fusion partner for ATPS purification of proteins (Wösten et al.

1993).

Collén et al. (2002) used the class II hydrophobin HFBI of Trichoderma reesei for ATPS

purification of the cellulose endoglucanase I (EGI) of T. reesei, with both non-ionic

detergent and polymers. The ATPS purified EGI showed an unaffected catalytic activity

with a good recovery rate of 90% at pilot scale, which appeared to be more efficient than

EGI purified by other methods (Linder et al. 2005).

Food production and quality control

Gushing is an unpleasant quality index of carbonated beverages (e.g. beer, champagne and

sparkling wine), a phenomenon where a carbonated beverage, without agitation, vigorously

effervesces and jets out when the cap is removed (Linder et al. 2005; Sarlin et al. 2012).

Haikara et al. (2006) concluded that hydrophobins (class II hydrophobins) of barley/malt

microbiological contaminants, including pathogenic Fusarium poae and Nigrospora spp.,

and non-pathogenic Trichoderma reesei (HFBI and HFBII) are closely related with these

gushing problems (Figure 4). This finding has led to the development of an immunological

detection method for the evaluation of gushing potential of barley and malt materials

(Haikara et al. 2006).

9

Figure 4. Class II hydrophobin HFBII of Trichoderma reesei causes gushing of beer. In this laboratory

experiment, 50 μg purified hydrophobin HFBII was added to a 33 cl bottle of beer three days prior to opening

(Figure adapted from Linder et al. 2005).

On the other hand, due to their superior foaming abilities and foam stability. Hydrophobins

can thus be used in many aerated foods where foams are important, including chocolate,

ice cream and buttercream (Cox et al. 2009). Recently, a research group from Unilever UK

has successfully used hydrophobins from edible mushrooms to mimic the texture and

mouthfeel of fats. As a result, up to 50% of the fat could be removed from the food

products without affecting the mouthfeel (Green et al. 2013; Cox et al. 2008).

Biofilm control

Adsorption of serum proteins plays an important role for bacterial colonization on surfaces

(Donlan 2002; Palmer et al. 2007), including dental plaque formation and the primary

stage of urinary tract infections (Bernsmann et al. 2008; Wösten and Scholtmeijer 2015).

Stallard et al. (2012) and Geoghegan et al. (2013) concluded that the fibrinogen loading of

a surface was closely related to its colonization potential of bacteria. Based on this fact,

von Vacano et al. (2011) successfully used the recombinant fusion hydrophobin protein

H*Protein B (from BASF SE), to prevent the non-specific absorption of serum proteins,

such as BSA, casein and collagen on a 1-octanethiol-coated gold surface. Sarparanta et al.

(2012) employed the class II hydrophobin HFBII of Trichoderma reesei to wrap thermally

hydro-carbonized porous silicon (THCPSi) nano-particles. Significant reduction in the

non-specific adsorption of plasma proteins, such as serum albumin and fibrinogen was

observed in these hydrophobin-treated nano-particles.

10

2.3 Antimicrobial peptides

Antimicrobial peptides (AMPs) are endogenous oligopeptide ‘antibiotics’, which are

widely distributed in all classes of life (Lehrer and Ganz 1999; Brodgen et al. 2003) and

generally execute a non-receptor mediated cell killing mechanism (Prenner et al. 1999)

against the competing microorganisms (Boman 1995). Usually these small molecules are

composed of less than 60 amino acid residues, with a net positive charge of +2 to +9

provided by lysine, arginine or, in acidic environments, histidine, and a dominant

proportion (>50%) of hydrophobic resides (Hancock and Sahl 2006; Brodgen 2005; Peters

et al. 2010).

Figure 5. Four major structural groups of antimicrobial peptides: (A) α-helical structure of magainin-2; (B)

β-sheeted structure of β-defensin 1; (C) extended structure of indolicidin; (D) looped structure of gramicidin

(Figure adapted from Peters et al. 2010).

The classification of AMP is difficult owing to the limited amino acid sequence homology

between these peptides, thus the further classification of AMPs is made on the basis of

their secondary structures (Epand and Vogel 1999; Hancock and Sahl 2006). Folded AMPs

are classified into four major structural groups (Figure 5): α-helical peptides (for example,

cecropins, magainins, pexiganans and the human cathelicidin LL-37); β-sheeted peptides

predominantly stabilized by more than one disulfide bridge (for example, human α- and

11

β-defensins and protegrins); extended peptides enriched with specific amino acids like

proline, tryptophan, arginine and histidine (for example, indolicidin, bactenecin-5 and

bactenecin-7); looped peptides coiled by one disulfide bridge (for example, gramicidin)

(Peters et al. 2010; Hancock and Sahl 2006).

Currently, more than 2,000 natural AMPs have been isolated from a wide range of

organisms (more information can be found at http://aps.unmc.edu/AP/main.php). Some of

them have been approved for clinical or food applications, including the cationic AMP

(cAMP) gramicidin S for treatment of genital ulcers and the lantibiotic nisin for

preservation of food products (Hancock and Sahl 2006). Of interest, AMPs exhibit a wide

range of activities against various organisms, including gram-negative and gram-positive

bacteria, fungi, enveloped viruses, parasites, and tumor cells with low propensity to cause

drug resistance among pathogens (Baker et al. 1993; Hancock and Scott 2000; Hancock

and Sahl 2006; Zasloff 2002; Chen et al. 2001; Hoskin and Ramamoorthy 2008). Due to

these unique properties, AMPs are considered as attractive substitutes for conventional

antibiotics to overcome the current drug resistance crisis (Hancock and Sahl. 2006; Lohner

2001).

Mechanisms of antimicrobial-peptide-mediated bacterial cell killing

Despite their vast structural varieties, most AMPs kill bacterial cells by disrupting the

cytoplasmic membrane integrity, resulting in the effusion of intracellular ions and organic

solutes (Prenner et al. 1999), which in turn may lead to cell death. Although the exact

mechanism by which AMPs disengage the bacterial cytoplasmic membrane is still not fully

understood. Indisputably, AMPs must be initially attracted to the bacterial surface, and this

step is thought to be facilitated by electrostatic interactions between AMPs and

lipopolysaccharides (LPS) on the surface of Gram-negative bacteria, and wall-associated

teichoic acids on the surface of Gram-positive bacteria (Figure 6) (Hancock 2001; Brogden

2005).

12

Figure 6. Cartoon of the electrostatic attraction of the positively charged AMPs to the negatively charged

bacterial envelopes (Figure adapted from Matsuzaki 2009).

Once non-covalently attached to the bacterial surface, AMPs must traverse the compact

bacterial outer wall before they can interact with the cytoplasmic membrane, by a process

described by Hancock and coworkers as ‘self-promoted uptake’ (Hancock and Sahl 2006;

Hancock 1997). This process is very important but rarely mentioned in many earlier

studies. Take gram-negative bacteria as an example, the bacterial outer wall stability is

maintained by native divalent cations (e.g. Mg2+

and Ca2+

) via salt bridges (Ledebo 1976).

AMPs show higher affinity to LPS than these native divalent cations and thus

competitively displacing them, resulting a disruption of the bacterial outer wall integrity.

This local disturbance in the bacterial out wall enables AMPs to pass through the bacterial

outer wall and reach the negatively charged cytoplasmic membrane.

Once reached the cytoplasmic membrane, AMPs start to interact with the anionic

phospholipid head groups of the cytoplasmic membrane (Figure 6). At low peptide/lipid

(P/L) molar ratios, AMPs are bound parallel onto the cytoplasmic membrane through

hydrophobic and electrostatic interactions, and slightly stretch the cytoplasmic membrane

(Yang et al. 2001; Brogden 2005). Once the P/L ratio surpasses a critical threshold, AMPs

start to aggregate into bundles and subsequently insert into the cytoplasmic membrane

interior, leading to the formation of transmembrane pores or detergent-like micelles

(Brogden 2005; Melo et al. 2009), which in turn may lead to cell death. Three modes of

action have been proposed based on the model membrane study with AMPs, they are: (1)

13

barrel-stave model (Ehrenstein and Lecar 1977); (2) carpet model (Pouny et al. 1992;

Jenssen et al. 2006) and (3) toroidal pore model (Yamaguchi et al. 2002) (Figure 7).

Figure 7. Modes of action of antimicrobial peptides: (A) barrel-stave model; (B) toroidal pore model; (C)

carpet model (Figure modified from Brogden 2005).

Alternative mechanisms of antimicrobial-peptide-mediated bacterial cell killing

Most of the existing AMPs (>90%) act directly on the bacterial cytoplasmic membrane, via

a non-receptor dependent membrane permeabilization process, as described in detail above.

However, some early studies revealed that there is a distinct group of AMPs, which can kill

bacteria in a non-lytic process (Bahar and Ren 2013). One example is AMP Buforin II,

which kills bacteria by inhibiting DNA and RNA synthesis (Park et al. 1998). Increasing

evidence indicates that there are several intracellular sites of AMP activity, including DNA

and RNA (Park et al. 1998), intracellular enzymes, and ribosomes (Boman et al. 1993)

(Figure 8).

Figure 8. Schematic diagram of intracellular sites of antimicrobial peptide activity in Escherichia coli

(Figure adapted from Brogden 2005).

14

Antimicrobial peptides P11-5 and Bac8c

AMP P11-5 (NH2-GKLFKKILKIL) is a novel 11-amino-acid peptide derived from AMP

BP76 (NH2-KKLFKKILKFL) with two amino acid substitutions (Qi et al. 2010). This

peptide showed a board-spectrum antimicrobial activity against a wide range of pathogenic

microorganisms, including gram-negative bacteria (E. coli and P. aeruginosa),

gram-positive bacteria (S. aureus) and fungi (C. albicans and F. solani) without significant

cytotoxicity to mammalian cells (Table 1).

AMP Bac8c (NH2-RRWIVWIR) is a truncated 8-amino-acid peptide derived from AMP

Bac2A (NH2-RAVRIVVIRALR), a C3A/C11A variant of bactenecin (also known as

bovine dodecapeptide), with four amino acid substitutions (Hilpert et al. 2005). This

peptide (Bac8c) is the smallest known antimicrobial peptide with broad-spectrum activity

against a range of microorganisms, such as gram-positive bacteria (S. aureus and

S. epidermidis), gram-negative bacteria (E. coli and P. aeruginosa) and a fungus

(C. albicans) (Table 1).

Table 1. Peptide properties: MICs (μg/ml) for five test pathogenic microorganisms

a values were obtained from Qi et al. 2010.

b values were obtained from Hilpert et al. 2005.

P11-5 is a cationic (+4) AMP with a substantial proportion (7/11) of hydrophobic amino

acid residues. The structural study of P11-5 showed that this peptide is mainly folded up

into an amphipathic α-helix structure, with the positively charged hydrophilic residues

localized on one side of the helix and the hydrophobic residues on the other side (Figure

9).

Peptide

Name

Peptide sequence

(NH2-)

Minimum inhibitory concentration (MIC, μg/ml)

E. coli P. aeruginosa S. aureus C. albicans F. solani

BP76a

P11-5a

Bac2Ab

Bac8cb

KKLFKKILKFL

GKLFKKILKIL

RAVRIVVIRAR

RRWIVWIR

8

3.1

17

2

16

12.5

50

8

62.3

12.5

17

2

25

3.1

9

8

50

12.5

-

-

15

Figure 9. Edmunson wheel projection of antimicrobial peptide P11-5. Black amino acids stand for

hydrophilic amino acid, and white amino acids stand for hydrophobic amino acids (Figure modified from Qi

et al. 2010).

Due to its amphipathic and cationic nature, P11-5 may spontaneously interact with the

negatively charged bacterial outer wall and rapidly disintegrate the cytoplasmic membrane

by the ‘carpet-like’ mode of action (Ferre et al. 2006; Melo et al. 2009). The field emission

scanning electron microscopy (FESEM) images of E. coli cells challenged by P11-5

showed conspicuous membrane fraying and surface blistering, together with the effusion of

intracellular materials (Figure 10).

Figure 10. Field emission scanning electron microscopy (FESEM) images of Escherichia coli ATCC 8739

cells untreated and treated with the antimicrobial peptide P11-5 for 15 min. (A) Untreated control; (B) 3.1

μg/ml (MIC) (Figure adapted from Qi et al. 2010). MIC: minimum inhibition concentration.

Bac8c is a cationic (+3) AMP with an abnormal low amount (< 50%) of hydrophobic

amino acid residues. The secondary structure of Bac8c is typically α-helix, with its specific

structure being anion-, pH- and detergent-dependent. For example, it adopts α-helical

16

structure in a zwitterionic solution and β-turn structure in anionic solutions (Wieczorek et

al. 2010).

Like the parent peptide Bac2A, Bac8c executes a two-stage mode of antimicrobial action,

in a way not solely dependent on membrane depolarization as the cause of cell death

(Spindler et al. 2011). In short, when Bac8c is present at concentrations that below the

minimum inhibitory concentration (MIC), the bacterial cytoplasmic membrane integrity is

well maintained (Figure 11B), while the cellular metabolism (e.g. ATP synthesis,

NAD+/NADH balance and protein synthesis) is severely impaired. Upon the accumulation

of a threshold concentration of Bac8c on microbial cytoplasmic membranes, the bacterial

cytoplasmic membrane is rapidly (less than 5 min) depolarized via an unknown mechanism,

resulting in the formation of transmembrane pores (between 2.2 and 3.3 nm) and effusion

of intracellular ions and organic solutes (Figure 11C & D) (Lee and Lee 2015).

Figure 11. Transmission electron microscopy (TEM) images of Escherichia coli ATCC 9637 cells untreated

and treated with the antimicrobial peptide Bac8c for 30 min. (A) Untreated control. (B) 3 μg/ml (IC50). (C) 6

μg/ml (MBC). (D) 30 μg/ml (5x the MBC) (Figure adapted from Spindler et al. 2011). MBC: minimum

bactericidal concentration. IC50: the half maximal inhibitory concentration.

2.4 The Pichia pastoris protein expression system

Over the past few decades, scientists have learned how to identify, amplify and place genes

into a variety of organisms that are different from the source organism (Macauley-Patrick

et al. 2005). Meanwhile, in this proteomics era, the demand of proteins for scientific and

17

pharmaceutical research is steadily on the increase. Therefore, a major application of these

transgenic organisms is to produce heterologous proteins (Macauley-Patrick et al. 2005;

Adrio and Demain 2010).

To date, bacterial protein expression systems are still predominately used in most

laboratories for heterologous protein production, due to their simplicity of genetic

manipulation, low cultivation costs and generally high yields, when compared with other

protein expression platforms (Chen 2012). However, insufficient and incorrect

post-translational modifications of complex proteins are two obvious drawbacks of these

bacterial systems. Take the class I hydrophobin HGFI of Grifola frondosa as an example,

this fungal protein has been successfully expressed in both Escherichia coli (Wang et al.

2010a) and Pichia pastoris (Wang et al. 2010b) protein expression systems. In E. coli,

HGFI is acquired as insoluble and miss-folded inclusion bodies, thus succeeding

solubilization and re-folding operations are needed (Wang et al. 2010a), which are either

technically difficult to achieve or time-consuming (Marston 1986; Daly and Hearn 2005;

Tsujikawa et al. 1996). In addition, E. coli produced heterologous proteins usually tend to

have an extra methionine residue (translation initiator) in their innate N-terminals, which

may pose a negative effect on protein activity and stability (Chaudhuri et al. 1999). In

contrast, in P. pastoris, HGFI is obtained in its biologically active form, and, more

importantly, its innate N-terminal sequence is well retained (Wang et al. 2010b). Other

fungal hydrophobins, initially produced as non-functional inclusion bodies through

production in E. coli, but obtained in their active forms when produced in P. pastoris,

including the class II hydrophobin HFBI of Trichoderma reesei (Nakari-Setälä et al. 1996;

Niu et al. 2012b) and the class I hydrophobin EAS of Neurospora crassa (Kwan et al. 2006;

Winefield 2004).

The P. pastoris protein expression system, except for allowing heterologous proteins to be

produced in their innate active forms, the increased popularity of this system can be

attributed to several additional factors: (1) the capability of performing eukaryotic

post-translational modifications, especially for target proteins with multiple disulfide

bridges or in need of glycosylation (Cereghino and Cregg 2000); (2) the ability of

18

producing heterologous proteins at high levels (Daly and Hearn 2005); (3) the availability

of systematic genetic manipulation techniques and kits, thus the expression system is easy

to set up and maintain in most laboratories.

Pichia pastoris host strains

P. pastoris is facultative methylotrophic yeast, which is capable of utilizing methanol as a

sole source of carbon and energy. The methanol utilization pathway involves several

enzymes, and the alcohol oxidase (AOX) is the most important one, catalyzing the

oxidation of methanol to formaldehyde (HCHO) and hydrogen peroxide (H2O2) in

peroxisomes.

The P. pastoris alcohol oxidase (AOX) is encoded by either the AOX1 or the AOX2 gene,

and the majority of AOX activity arises from the AOX1 gene (Tschopp et al. 1987; Cregg

et al. 1989). A wild type P. pastoris strain (e.g. P. pastoris X-33) generally contains both

AOX (AOX1 and AOX2) genes and primarily uses the AOX1 gene for its methanol

metabolism. This phenotype is designated as methanol utilization fast (Mut+). Once the

AOX1 gene is disrupted or deleted on purpose, as a result, P. pastoris cells (e.g. P. pastoris

KM71) are still capable of utilizing methanol via the transcriptionally weaker AOX2 gene

at a decreased rate, thus giving rise to the phenotype methanol utilization slow (Muts). In

addition, a rarely addressed phenotype exists for P. pastoris strains (e.g. P. pastoris MC

100-3) that have deletions at both AOX genes, which are unable to grow on methanol. This

phenotype is denominated as methanol utilization minus (Mut-) (Macauley-Patrick et al.

2005).

It is not entirely clear that which phenotype (Mut+, Mut

s and Mut

-) is the most suitable one

for expression of a given protein (Hellwig et al. 2001). However, for extracellular

production of heterologous proteins, Mut+ strains are more often used than Mut

s strains,

due to their higher growth rates and better environmental adaptabilities (Hohenblum et al.

2004; Slibinskas et al. 2004). This phenotype (Mut+) can easily be identified either by

growing P. pastoris cells on minimal dextrose (MD) and minimal methanol (MM) plates

19

sequentially (Vassileva et al. 2001), or by colony PCR with specific primers 5’ AOX1 and

3’ AOX1 (Linder et al. 1996).

Protein degradation has long been a problem associated with P. pastoris protein production

practices, due to the action of endogenous proteases (e.g. vacuole peptidase A). The use of

protease defect strains (e.g. SMD1168) has proven to reduce protein degradation in many

heterologous protein production studies with this system (Cereghino and Cregg 2000;

Macauley-Patrick et al. 2005).

Table 2. Pichia pastoris expression host strains

Strain Genotype Phenotype Reference

X-33

GS115

KM71

SMD1168

MC 100-3

wild-type

his4

△aox1:: SARG4 his4 arg4

△pep4::URA3 his4

△aox1: :SARG4△aox2: :Phis4

his4 arg4

Mut+, His

+

Mut+, His

-

Muts, His

-

Mut+, His

+

Mut-, His

-

Li et al. 2001

Cregg et al. 1985

Cregg and Madden 1987

White et al. 1995

Cregg et al. 1989

his4: Histidine dehydrogenase defective type. △aox1: Alcohol oxidase I defective type.

△aox2: Alcohol oxidase II defective type. △pep4: Vacuole peptidase A defective type.

△arg4: Argininosuccinate lyase defective type.

Pichia pastoris expression vectors

There is a wide range of commercially available vectors that can be used to express

heterologous proteins in P. pastoris (Table 3). All P. pastoris expression vectors have been

designed as an E. coli/P. pastoris shuttle vector. Therefore, an origin of replication for

vector maintenance and replication in E. coli, AOX1 promoter or alternative promoters,

one or more restriction sites for insertion of foreign genes, and selection markers functional

in one or both host organisms are needed (Cereghino and Cregg 2000).

pPIC9 (Figure 12) is a commercially available P. pastoris expression vector that has been

intensively used for the production of hydrophobins (Niu et al. 2012a; Wang et al. 2010b;

Niu et al. 2012b) and antimicrobial peptides (Zhang et al. 2005; Zhang et al. 2000). Except

for the basic features that cited above, this vector contains the functional histidine

dehydrogenase gene (HIS4), which can be used as a selection marker for the

transformation of histidine dehydrogenase defective (his4) strains (e.g. GS115, KM71,

20

SMD1168). In addition, to facilitate the extracellular production of heterologous proteins,

an extracellular secretion signal sequence (usually, alpha-mating factor of Saccharomyces

cerevisiae) is incorporated upstream of the multiple cloning site (MCS).

Figure 12. Vector diagram of pPIC9 (Figure adapted from Pichia expression kit user guide, MAN0000012,

Lifetechnologies)

Table 3. Commercially available P. pastoris expression vectors

Name Secretion signal Selection maker Promoter

pPIC9

pPIC9K

pPICZαA

pGAPZαA

pHIL-D2

pHIL-S1

pP-αSUMOstar

α-MF

α-MF

α-MF

α-MF

no

PHO1

α-MF

HIS4, AmpR

AmpR, KanR

ZeoR

ZeoR

HIS4, AmpR

HIS4, AmpR

ZeoR

AOX1

AOX1

AOX1

GAP

AOX1

AOX1

AOX1

Secretion signals: α-MF (S. cerevisiae α-mating factor); PHO1 (P. pastoris acid phosphatase).

Selection markers: AmpR (ampicillin resistance); Kan

R (kanamycin resistance); Zeo

R(zeocin resistance);

HIS4 (histidine dehydrogenase).

Promoters: GAP (glyceraldehyde-3-phosphate dehydrogenase); AOX1 (alcohol oxidase 1).

More information can be found at www.lifetechnologies.com.

To date, a wide range of fungal origin hydrophobins have been successfully expressed in

P. pastoris, including HFBI (Niu et al. 2012), HFBII (Kottmeier et al. 2011) and Hfb2

(Lutterschmid et al. 2011) of Trichoderma reesei, FcHyd5p and FcHyd3p of Fusarium

culmonrum (Lutterschmid et al. 2011; Stübner et al. 2010), HGFI of Grifola frondosa

(Wang et al. 2010b), RodA and RodB of Aspergillus fumigatusin (Pedersen et al. 2011) and

EAS of Neurospora crassa (Winefield 2004).

21

EXPERIMENTAL RESEARCH

3 AIMS OF THE STUDY

The aim of this study was to explore the feasibility of using recombinant fusion

hydrophobins to control bacterial growth. The specific aims were:

1. To construct expression vectors that can thus be used to transform P. pastoris with a

view to obtain two recombinant fusion hydrophobins, Bac8c-HGFI and

P11-5-linker-HGFI.

2. To express these two recombinant fusion hydrophobins in P. pastoris and purify them

from the culture medium.

3. To determine the minimum inhibition concentrations (MICs) of these two recombinant

fusion hydrophobins against bacteria.

4 MATERIALS AND METHODS

4.1 Materials

4.1.1 Bacteria and yeast strains, plasmids, oligonucleotides and primers

Pichia pastoris GS115 (Genotype: his4; Phenotype: His-, Mut

+) and X-33 (Wild type;

Phenotype: Mut+) strains were kindly provided by Dr. Kristiina Hildén (University of

Helsinki, Finland). Library Efficiency® DH5α™ Competent Cells (Genotype: F-

Φ80lacZΔM15 Δ (lacZYA-argF) U169 recA1 endA1 hsdR17 (rK-, mK

+) phoA supE44 λ-

thi-1 gyrA96 relA1) were purchased from Lifetechnologies (Amsterdam, Netherlands).

Reference strains Escherichia coli ATCC 8739, Staphylococcus aureus subsp. aureus

ATCC 12600, Pseudomonas aeruginosa ATCC 10145 and Micrococcus luteus ATCC 4698

were obtained from HAMBI culture collection center (University of Helsinki, Finland).

Reconstructed P. pastoris expression vectors pPIC9-linker-hgfI preserved in E. coli TG1

[Genotype: F' (traD36 proAB+ lacI

q lacZΔM15) supE thi-1 Δ(lac-proAB)

Δ(mcrB-hsdSM)5, (rK-, mK

-)] and pP-secSUMOpro3-hgfI preserved in E. coli DH5α

(Genotype: F- Φ80lacZΔM15 Δ (lacZYA-argF) U169 recA1 endA1 hsdR17 (rK-, mK

+)

22

phoA supE44 λ- thi-1 gyrA96 relA1) were kindly provided by Prof. Mingqiang Qiao

(Nankai University, China). All primers and oligonucleotides (Table 4) were custom

synthesized by Oligomer Oy (Helsinki, Finland).

Table 4. Overview of primers and oligonucleotides used in this study

Name Sequence 5’ – 3’

5’ AOX

3’AOX

α-factor

p11-5 forward

p11-5 reverse

AMP forward

hgfI reverse

hgfIa

bac8ca

GACTGGTTCCAATTGACAAGC

GCAAATGGCATTCTGACATCC

TACTATTGCCAGCATTGCTGC

TCGAGAAAAGAGGTAAGTTGTTCAAGAAGATCTTGAAGATCTTGG

AATTCCAAGATCTTCAAGATCTTCTTGAACAACTTACCTCTTTTC

CAATCAATGAAACTGACA

TGGTGGCCGGGTCGTTCG

CAACAGTGCACCACTGGCCA

TCTTCTCCAAATAACCCAAATTCTACCTCCCGTCTGCTGCTGGA

a 5’-phosphorylated

4.1.2 Antimicrobial peptides, chemical reagents and culture media components

Antimicrobial peptides Bac8c and P11-5 were custom synthesized by Thermoscientific

(Germany) using the solid phase method and standard 9-fluorenyl-methoxy-carbonyl

(FMOC) chemistry. These peptides were purified to > 95% purity using reversed-phase

high performance liquid chromatography (RP-HPLC). Mass spectrometry was used to

confirm the peptide identity.

All chemical regents and culture media components were purchased from Sigma-Aldrich

(Germany), Merck (Germany) or Lab M (Heywood, UK) unless otherwise stated.

4.2 Methods

All techniques used for the molecular biology experimental work were based upon these

standard protocols found in Molecular Cloning: A Laboratory Manual (Green and

Sambrook, 2012) and Cold Spring Harbor Online Protocols (http://cshprotocols.cshlp.org/)

unless otherwise stated.

23

4.2.1 Isolation and purification of plasmid DNA from Escherichia coli

Glycerol stock of E. coli TG1 cells containing the reconstructed P. pastoris expression

vector pPIC9-linker-hgfI was streaked onto a freshly prepared LB agar plate (1% [w/v]

trypton, 0.5% [w/v] yeast extract, 1% [w/v] sodium chloride, 1.5% [w/v] agar) containing

100 μg/ml ampicillin. In the same way, glycerol stock of E. coli DH5α cells containing the

reconstructed P. pastoris expression vector pP-secSUMOpro3-hgfI was streaked onto a

freshly prepared low salt LB agar plate (1% [w/v] trypton, 0.5% [w/v] yeast extract, 0.5%

[w/v] sodium chloride, 1.5% [w/v] agar) containing 50 μg/ml zeocin (Invitrogen, Carlsbad,

CA, USA). These plates were incubated overnight at 37℃ until single colonies appeared.

A single colony from the LB agar plate was inoculated into 5 ml LB broth (1% [w/v]

trypton, 0.5% [w/v] yeast extract, 1% [w/v] sodium chloride) containing 100 μg/ml

ampicillin (for E. coli DH5α, 5 ml low salt LB broth [1% [w/v] trypton, 0.5% [w/v] yeast

extract, 0.5% [w/v] sodium chloride] containing 50 μg/ml zeocin was used), and incubated

at 37℃ with shaking at 200 rpm until OD600 reached 0.4 (log-phase growth, approximately

8 hours). Bacterial cells were collected by centrifugation (Sigma 1-14 Microfuge) for 5

min at 12000g. Plasmid DNA was extracted from bacterial pellets and purified with

GeneJET Plasmid Miniprep Kit (Thermoscientific) according to the manufacturer’s

instructions, and stored in a sterile nuclease-free Eppendorf tube at -20℃ . The

concentrations of DNA samples were measured by NanoDrop-1000 Spectrophotometer

(Thermoscientific).

4.2.2 DNA agarose gel electrophoresis

Separation and visualization of DNA fragments was performed by agarose gel

electrophoresis. The percentage (w/v) of agarose (Lonza, Basel, Switzerland) used was

varied between 1% and 3% depending on the size of the DNA fragments. Gel solutions

were prepared with 1x TAE buffer (40 mM tris-HCl, 40 mM glacial acetic acid, 1 mM

EDTA, pH 8.0) and ethidium bromide was added to a final concentration of 0.5μg/ml.

DNA samples were pre-mixed with 6x DNA Loading Dye (Thermoscientific) prior to gel

loading. 1 kbp GeneRuler™ DNA Ladder (Thermoscientific) or 100 bp plus GeneRuler™

DNA ladder (Thermoscientific) was used as a marker of molecular mass and run parallel to

24

the DNA samples. Electrophoresis was carried out in 1x TAE buffer at 80 V. Complete gels

were observed and image captured with Gel Doc XR+ imaging system (Bio-Rad

Laboratories, Hercules, CA, USA).

4.2.3 Construction of pP-secSUMOpro3-bac8c-hgfI

The inverse fusion PCR (IF-PCR) (Figure 13) was conducted with Phusion® Hot Start II

High-Fidelity DNA polymerase (Thermoscientific) (Table 5). 5’ phosphorylated primers

(bac8c and hgfI) were designed using a protocol described by Spiliotis (2012). Yield

optimization was performed with respect to the annealing temperature via gradient PCR in

Bio-Rad C1000 Thermal Cycler under the following conditions: 98℃ for 1 min, 30 cycles

of 98℃ for 10 s, (63℃-75℃) for 30 sec, 72℃ for 2 min, and a final extension step at 72℃

for 10 min. The PCR products were analyzed on 1% agarose gel.

Table 5. Pipetting instructions for the inverse fusion PCR reaction

Component Volume (μl) Final concentration

5X Phusion HF Buffer

Phusion Hot Start II DNA polymerase (2 U/μl)

hgfI (10 μM)

bac8c (10 μM)

pP-secSUMOpro3-hgfI (template, 1 ng/μl)

dNTPs (10 mM)

ddH2O

10

0.5

2.5

2.5

1

1

32.5

1x

0.02 U/μl

0.5 μM

0.5 μM

0.02 ng/μl

200 μM each

The required DNA band was excised from the agarose gel with x-tracta™ Gel Extractor

(Promega, Madison, WI, USA) on an UV transilluminator (Bio-Rad Laboratories) and

subsequently purified with GeneJET Gel Extraction Kit (Thermoscientific) following the

manufacturer’s instructions.

For self-circularization of blunt-ended PCR products, the reaction mixture was prepared as

follows: 50 ng blunt-ended linear DNA, 2 μl 10x T4 DNA ligase buffer, 1 μl T4 DNA

ligase (5 Weiss/μl, Thermoscientific), 2 μl PEG 4000 (50% [w/v]) and ddH2O to a total

volume of 50 μl. The reaction mixture was incubated at 22℃ for 2 h, and the ligase was

heat inactivated at 65℃ for 10 min.

25

Figure 13. Outline of inverse fusion PCR (IF-PCR). For IF-PCR, a PCR master mix containing 5’

phosphorylated primer bac8c with the insert, circular plasmid template and 5’ phosphorylated primer hgfI

was prepared. The bac8c primer was annealed with the complementary sequence (black) within the plasmid

forward strand (2.1 & 2.2), the insert (white) was elongated by overlap extension (2.3), thus generating the

single-stranded DNA (ssDNA) template with the insert (2.4). The complementary strand of the ssDNA

template was generated by primer extension of primer hgfI, finalizing the double-stranded DNA (dsDNA)

template (2.5), which was now exponentially amplified via primers bac8c and hgfI. The linear insert-plasmid

fusions were self-circularized by T4-ligation (2.6) (Figure adapted from Spiliotis 2012).

4.2.4 Construction of pPIC9-p11-5-linker-hgfI

Restriction endonuclease (RE) digestion of pPIC9-linker-hgfI

The freshly prepared vector pPIC9-linker-hgfI from the plasmid isolation step (section

4.2.1) was double-restricted with FastDigest® EcoRI and XhoI restriction endonucleases

(Thermoscientific). The double REs digestion was performed under the optimal reaction

condition described by the manufacturer.

One step phosphorylation and annealing of oligonucleotides

As oligonucleotides (p11-5 forward and p11-5 reverse) used in this study are supplied 5’

non-phosphorylated. Therefore, to improve the ligation efficiency, T4 polynucleotide

kinase (T4 PNK, Thermoscientific) was used to phosphorylate oligonucleotides. The

reaction mix was prepared and the one-step phosphorylation and annealing reaction was

carried out in Bio-Rad C1000 Thermal Cycler (Table 6).

26

Table 6. Pipetting instructions and thermocycler program for the one-step phosphorylation and annealing

reaction of oligonucleotides

Reaction mix composition Thermocycler program

Component

10X T4 PNK buffer A

T4 PNK (10 U/μl)

p11-5 forward (10 μM)

p11-5 reverse (10 μM)

ATP (10 mM)

ddH2O

Volume (μl)

2

2

1

1

4

10

Step

1

2

3

4

5

6

Temperature (℃)

37

95

95 (-1/cycle)

74

74(-1/cycle)

4

Time

30 min

5 min

1 min/cycle

30 min

1 min/cycle

∞

Ligation of DNA fragments

The 5’ phosphorylated double-stranded oligo (ds oligo) fragment synthesized from the

previous step was ligated into the prepared pPIC9-linker-hgfI vector. For DNA insert

ligation into vector DNA, the reaction mixture was prepared as follows: 100 ng linear

vector DNA, 10: 1 molar ratio of insert DNA over vector, 2 μl 10x T4 DNA ligase buffer,

0.2 μl T4 DNA ligase (5 Weiss/μl, Thermoscientific), and ddH2O to a total volume of 20 μl.

The reaction mixture was incubated overnight at room temperature (20℃).

Figure 14. Outline of annealed oligo cloning. A short double-stranded oligo (ds oligo) fragment was

synthesized by annealing oligonucleotides p11-5 forward and p11-5 reverse, with sticky ends specific to XhoI

and EcoRI restriction endonucleases in 5’- and 3’-terminus, respectively. The double-stranded oligo fragment

was ligated into the prepared pPIC9-linker-hgfI vector by T4-ligation.

4.2.5 Transformation of electrocompetent Escherichia coli DH5α cells

Electrocompetent E. coli DH5α cells from -80℃ storage were thawed on ice. 1 μl ligation

mix (from section 4.2.3 or section 4.2.4) was added to 50μl cells, gently mixed and

transferred to a pre-chilled 2 cm disposable electroporation cuvette (VWR International,

Germany). Electroporation was conducted with a Gene Pulser™ apparatus (Bio-Rad

Laboratories) set at 2.5 kV, 200 Ω and 25 μF. Immediately after the electroporation, 950 μl

SOC broth (Lifetechnologies) was added to the cuvette and a total volume of 1 ml bacterial

27

suspension was transferred to a sterile Eppendorf tube and placed on a shaking incubator at

37℃ for 1 hour. The bacterial cells were then plated out on selection agar plates with the

corresponding antibiotics and incubated overnight at 37℃ until single colonies appeared.

4.2.6 Colony PCR

Colony PCR was used as a rapid method to screen ligation reactions for positive clones.

Visible bacterial colonies were picked from selection agar plates with pipette tips,

suspended in 10 μl nuclease-free ddH2O and thoroughly mixed by vortex. 1 μl bacterial

liquid was directly added to the PCR master mix as the DNA template (Table 7).

The colony PCR was conducted with vector specific primers in Bio-Rad C1000 Thermal

Cycler under the following conditions: 94℃ for 5 min, 30 cycles of 94℃ for 1 min, 53℃

for 1 min, 72℃ for 8 sec, and a final extension step at 72℃ for 10 min. The PCR

products were analyzed on 3% agarose gel. Two colony PCR verified positive

transformants were sent for sequencing (Institute of Biotechnology, University of Helsinki,

Finland).

Table 7. Pipetting instructions for E. coli colony PCR

Component Volume (μl) Final concentration

10X optimized DyNAzyme buffer

10 mM dNTPs

α-factor/AMP forward (10 μM)a

hgfI reverse (10 μM)

Bacterial liquid (DNA template)

DyNAzyme II polymerase (2 U/μl)

H2O

2

0.4

1

1

1

0.2

14.4

1x

200 μM each

0.5 μM

0.5 μM

0.02 U/μl

a primers AMP forward and hgfI reverse were used for E. coli transformants that transformed with the

pP-secSUMOpro3-bac8c-hgfI ligation mix, and primers α-factor and hgfI were used for E. coli transformants

that transformed with the pPIC9-p11-5-linker-hgfI ligation mix.

4.2.7 Transformation of electrocompetent Pichia pastoris cells

The sequencing-confirmed P. pastoris expression vector pPIC9-p11-5-linker-hgfI was

linearized with StuI (Thermoscientific) (for pP-secSUMOpro3-bac8c-hgfI, PmeI was used).

The linearized vector DNA was concentrated by isopropanol precipitation, suspended in a

small amount of nuclease-free Milli-Q water and subsequently used to transform

electrocompetent P. pastoris GS115 cells (for pP-secSUMOpro3-bac8c-hgfI,

28

electrocompetent P. pastoris X-33 cells were used). Electroporation was conducted with a

Gene Pulser™ apparatus set at 2.0 kV, 200 Ω and 25 μF. Immediately after the

electroporation, 1 ml 1M ice cold sterile sorbitol was added to the cuvette and the cell

suspension was transferred to a 15 ml Falcon™ tube and left to incubate for 2 hours at 28℃

without shaking. Electrocompetent P. pastoris GS115 and X-33 cells were prepared using a

protocol described by Cereghino et al. (2005).

For P. pastoris GS115 transformants, cells were initially plated on MD (minimal dextrose)

agar plates (1.34% [w/v] YNB, 4 × 10-5

% [w/v] biotin, 2% [w/v] dextrose, 1.5% [w/v]

agar) and incubated at 30℃ for 3-4 days until single colonies appeared. Subsequently,

thirty colonies (His+) were picked from MD agar plates and pasted on MM (minimal

methanol) agar plates (1.34% [w/v] YNB, 4 × 10-5

% [w/v] biotin, 0.5% [v/v] methanol,

1.5% [w/v] agar) to identify the methanol utilization type. For P. pastoris X-33

transformants, cells were initially plated on YPDS agar plates (1% [w/v] yeast extract, 2%

[w/v] peptone, 2% [w/v] dextrose, 1 M sorbitol, 2% [w/v] agar) containing 100 μg/ml

zeocin and incubated at 30℃ for 3-4 days until single colonies appeared. Subsequently,

twenty zeocin-resistant colonies were picked from YPDS agar plates and further polished

on YPD agar plates (1% [w/v] yeast extract, 2% [w/v] peptone, 2% [w/v] dextrose, 2%

[w/v] agar) containing 100 μg/ml zeocin.

Table 8. Pipetting instructions and thermocycler program for P. pastoris colony PCR

PCR master mix composition Thermocycler program

Component

10X standard Taq buffer

10 mM dNTPs

5’ AOX1 (10 μM)

3’ AOX1 (10 μM)

Hot start Taq DNA polymerasea

Template DNA

ddH2O

Volume (μl)

2

0.4

1

1

0.2

1

to 20

Step

1

2

3

4

5

6

7

Temperature (℃)

95

95

54

68

go back to 2

68

4

Time

3 min

30 s

30 s

2.2 min

35 cycles

5 min

∞

a New England Biolabs(NEB)

The fast-growing/zeocin-resistant transformants were further analyzed by colony PCR with

primers 5’ AOX and 3’ AOX (Table 8). To obtain the genomic DNA template for PCR,

29

single colonies were picked and lysed with 0.2% hot SDS treatment (microwave oven set

at full power) for 2 min. Cell debris was then pelleted by a pulse spin in a bench top

centrifuge and the supernatant was directly used for PCR analysis.

4.2.8 Inducible expression of recombinant fusion hydrophobins

The colony PCR-validated P. pastoris transformants together with the corresponding

control strains (P. pastoris GS115 transformed with pPIC9, P. pastoris X-33 transformed

with pP-secSUMOpro3) were used for a small-scale trial expression of recombinant fusion

proteins.

For optimal expression, P. pastoris transformants and control strains were initially

inoculated into 25 mL buffered minimal glycerol (BMG) broth (100 mM potassium

phosphate, 1.34% [w/v] yeast nitrogen base, 4 x 10-5

% [w/v] biotin, 1% [v/v] glycerol, pH

6.0) and grown at 30℃ with shaking at 280 rpm until OD600 reached 2-6 (log-phase

growth, approximately 16-18 hours). These cultures were centrifuged for 5 min at 3000g

and pelleted cells were suspended in 50 mL buffered minimal methanol (BMM) broth (100

mM potassium phosphate, 1.34% [w/v] yeast nitrogen base, 4 x 10-5

% [w/v] biotin, 0.5%

[v/v] methanol, pH 6.0). Cultures were grown for 96/120 h at 30℃ with shaking at 280

rpm, filter-sterilized methanol was added to a final concentration of 0.5% (v/v) every 24 h.

Meanwhile, 1 ml culture samples were taken at 24 h intervals for protein expression

analysis. After 96 h/120 h of cultivation, culture supernatants were collected by

centrifugation (8000g, 10 min, JA-14, Beckman Avanti™ J-251) and stored in sterile

SCHOTT® glass bottles (Duran, Germany) at -80℃.

4.2.9 Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) of

proteins

Tricine-SDS-PAGE with the discontinuous pH system described by Schägger (2006) was

used to separate and visualize small proteins/peptides. Culture supernatant samples from

the previous step were concentrated by trichloroacetic acid (TCA) precipitation, mixed

with Tricine Sample Buffer (Bio-Rad) containing 125 mM dithiothreitol (DTT) and heated

immediately for 10 minutes at 95℃. Spectra Multicolor Low Range Protein Ladder

30

(Thermoscientific) was loaded as a marker of size. Gels were run at 30 V through the

stacking gel and 200 V through the separating gel. When the electrophoresis had finished,

gels were stained with Silver Stain Plus Kit (Bio-Rad).

Glycine-SDS-PAGE with the discontinuous pH system originally described by Laemmli

(1970) was used to separate and visualize proteins. Culture supernatant samples were

concentrated by acetone precipitation, mixed with 2x Laemmli Sample Buffer (Bio-Rad)

containing 355 mM 2-mercaptoethanol and heated immediately for 10 minutes at 95℃.

Spectra Multicolor Low Range Protein Ladder (Thermoscientifc) was loaded as a marker

of size (for Western blot analysis, 7-175 kDa prestained protein marker [New England

Biolabs] was used). Gels were run at 100 V through the stacking gel and 200 V through the

separating gel. When the electrophoresis had finished, complete gels were stained for three

hours and de-stained, using laboratory stocks of filtered Coomassie staining solution (0.04%

[w/v] Coomassie Brilliant Blue R-250, 40% [v/v] methanol, 10% [v/v] glacial acetic acid)

and de-staining solution (10% [v/v] glacial acetic acid), respectively.

4.2.10 Western blot analysis of the recombinant fusion hydrophobin

SUMO3-Bac8c-HGFI

Western blot originally reported by Towbin et al. (1979) was used as a more sensitive and

specific approach for visualizing proteins. Culture supernatant samples were initially

separated with Glycine-SDS-PAGE as described in section 4.2.9. Instead of staining,

proteins on SDS-PAGE gel were electro-blotted to a polyvinylidene fluoride (PVDF)

membrane (Pall, Port Washington, NY, USA) with a wet-transfer apparatus (Bio-Rad) set

at 30 V for 2 h. After transfer, the PVDF membrane was blocked in the blocking solution

(WesternBreeze®, Invitrogen) at room temperature for 30 minutes.

Transferred proteins were probed with the primary antibody anti-Penta·His IgG1 (Mouse,

1:1000 dilution) (Qiagen, Netherlands) for 1 h at room temperature. After rinsed three

times with the antibody wash solution (WesternBreeze®, Invitrogen), alkaline phosphatase

(AP) conjugated Goat anti-Mouse IgG (H+L) secondary antibody (1:5000 dilution) (Pierce,

31

Rockford, IL, USA) was added and left to bind for 1 h and visualized with BCIP/NBT

Color Development Substrate (Promega).

4.2.11 Purification of the recombinant fusion hydrophobin SUMO3-Bac8c-HGFI

The recombinant fusion hydrophobin SUMO3 [6x His-tag]-Bac8c-HGFI was purified with

Ni-NTA slurry (Invitrogen) following the manufacturer’s instructions. The purified protein

solution was transferred into an Amicon Ultra-15 ultrafiltration unit (Millipore) and buffer

exchanged against phosphate buffered saline (PBS) solution (137 mM NaCl, 2.7 mM KCl,

10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4) to remove imidazole and chemical impurities.

SUMO protease 2 (Lifesensors, Malvern, PA, USA) was added to the substrate (1 unit

enzyme to 10-100 μg target fusion protein) to remove the SUMO3 fusion tag. The reaction

was performed overnight at 4℃. In the same way, the SUMO3 fusion tag and SUMO

protease 2 in the digestion mixture was removed with Ni-NTA slurry and buffer exchanged

against ultrapure Milli-Q water. The purified protein solution was then lyophilized and

stored at -80℃.

4.2.12 Purification of the recombinant fusion hydrophobin P11-5-linker-HGFI

Culture supernatant samples were separated with 16% Tricine-SDS-PAGE as described in

section 4.2.9, instead, gel was stained with Coomassie staining solution (10% acetic acid,

40% methanol and 0.1% Coomassie Brilliant Blue-R250). The required protein band was

excised from the gel and eluted with Model 422 Electro-Eluter apparatus (Bio-Rad)

following the manufacturer’s instructions. In short, the excised protein gel slice was placed

into an Electro-Eluter capsule and the electroelution was performed at 50 V for 12 h at 4℃

in Tris-glycine buffer (25 mM Tris-HCl, 250 mM glycine, 0.1% [v/w] SDS). Eluent in the

membrane cap was collected and transferred into an Amicon Ultra-4 ultrafiltration unit

(Millipore) and buffer exchanged against ultrapure Milli-Q water to remove chemical

impurities. The protein solution was then lyophilized and stored at -80℃.

32

4.2.13 Dot blot analysis of the recombinant fusion hydrophobin P11-5-linker-HGFI

The purified recombinant fusion hydrophobin P11-5-linker-HGFI was analyzed by dot blot,

2 μl protein solution was directly applied to a PVDF membrane (Pall) via a micropipette

and air-dried at room temperature for at least 30 min. The PVDF membrane was then

blocked in the blocking solution (WesternBreeze®, Invitrogen) at room temperature for 30

minutes.

Proteins on the PVDF membrane were probed with the primary antibody anti-rHGFI

(1:1000 dilution) (from a male New Zealand white rabbit, Nankai University, China) for 1

h. After rinsed three times with the antibody wash solution (WesternBreeze®, Invitrogen),

alkaline phosphatase (AP) conjugated secondary antibody anti-Rabbit IgG (Fc) (1:3000

dilution) (Promega) was left to bind for 1 h and visualized with BCIP/NBT Color

Development Substrate (Promega).

4.2.14 Protein quantification

Protein quantification was performed with Modified Lowry Protein Assay Kit (Pierce,

Waltham, MA, USA) following the manufacturer’s instructions.

4.2.15 Minimum inhibitory concentration test

Minimum inhibitory concentrations (MICs) were determined using a modified microplate

dilution method described by Wiegand et al. (2008). Reference bacteria were grown to a

mid-log phase (OD600 ≈ 0.4) in Mueller-Hinton broth (BD Difco™) and diluted to 2-7 x

105

CFU/ml inoculum sizes.

Antimicrobial peptides and purified fusion proteins were serial diluted to obtain final

concentrations of 200, 100, 50, 25, 12.5, 6.25 and 3.1 μg/ml, respectively (Qi et al. 2010;

Hilpert et al. 2005). Meanwhile, bacterial growth control with no test peptides/proteins and

environmental sterility control with no bacterial cells were prepared. Plates were incubated

at 37℃ for 24 h and bacterial growth was determined with Infinite 200 microplate

reader (Tecan, Switzerland) set at 600 nm. The experiment was independently repeated two

33

times. MIC was defined as the lowest concentration that needed to reduce bacterial growth

by more than 50% after 24 hours (Wiegand et al. 2008).

4.2.16 Tools for data analyses

1. Thermoscientific Tm calculator was used to predict the annealing temperature for PCR

or sequencing.

2. NEBcutter V2.0 was used to determine the potential endonuclease cleavage sites of

vectors.

3. NCBI Nucleotide BLAST was used to generate sequence alignments for DNA

sequence comparison.

34

5 RESULTS

5.1 Construction of pP-secSUMOpro3-bac8c-hgfI

Yield optimization of inverse fusion PCR (IF-PCR) was performed with respect to the

annealing temperature via gradient PCR. In lanes 1-5, a band was apparent above 4 kbp.

The no-template control in lane C did not display any bands (Figure 15).

Figure 15. 1% agarose gel displaying inverse fusion PCR products. Lane L: Generuler™ 1 kbp DNA ladder.

Lanes 1-5: gradient PCR products (from left to right, 65℃, 67℃, 69℃, 71℃, 73℃). Lane C: No-template

control.

The >4 kbp DNA fragment was purified from the agarose gel, self-circularized and

transformed into E. coli DH5α cells. Eight zeocin-resistant transformants growing on

selection plates were picked and analyzed by colony PCR with vector specific primers. In

lanes 5, 6 and 9, a band was apparent below 200 bp. In lanes 2-4, 7 and 8, three bands were

apparent below 1000 bp, between 1000 bp and 1200 bp, and above 1200 bp, respectively.

The positive control (E. coli DH5α transformed with pP-secSUMOpro3-hgfI) in lane 1

displayed a band around 150 bp. The negative control (E. coli DH5α cells) in lane C did

not show any bands (Figure 16C).

35

Figure 16. (A) Schematic representation of the reconstructed P. pastoris expression vector

pP-secSUMOpro3-bac8c-hgfI. (B) The amino sequence of the recombinant fusion hydrophobin Bac8c-HGFI.

(C) 3% high-resolution agarose gel (Sigma-Aldrich, Germany) displaying E. coli colony PCR products. Lane

L: Generuler™ 100 bp Plus DNA ladder. Lane 1: positive control (E. coli DH5α transformed with

pP-secSUMOpro3-hgfI). Lane C: negative control (E. coli DH5α cells). Lanes 2-9: eight zeocin-resistant E.

coli transformants.

5.2 Expression of the recombinant fusion hydrophobin SUMO3-Bac8c-HGFI

The sequencing-confirmed expression vector pP-secSUMOpro3-bac8c-hgfI was linearized

with PmeI and subsequently transformed into P. pastoris X-33 cells. Six zeocin-resistant

P. pastoris transformants growing on selection plates were picked and analyzed by colony

PCR with primers 5’AOX1 and 3’AOX1. In lanes 1-6, two clear bands were apparent

above 2 kbp and above 1 kbp, respectively (Figure 17).

Figure 17. 1% agarose gel displaying P. pastoris colony PCR products. Lane L: Generuler™ 1 kb DNA

ladder. Lanes 1-6: six zeocin-resistant P. pastoris X-33 transformants.

36

Six P. pastoris X-33 clones identified as positive transformants by colony PCR together

with the control strain (P. pastoris X-33 transformed with pP-secSUMOpro3) were

subjected to a small-scale trial expression. After the expression was induced with 0.5%

methanol for 96 h, supernatant samples from each induced culture were collected,

concentrated and analyzed by SDS-PAGE. Two protein bands with apparent molecular

weights of 40 kDa and > 40 kDa were observed in all six P. pastoris X-33 transformants

(lanes 1-6, Figure 18A), but not in the control strain at the corresponding positions (lane C,

Figure 18A). Western blot showed a protein with apparent molecular weight of 80 kDa

could react positively (purple band) with the protein specific primary antibody, which was

observed in all six P. pastoris X-33 transformants (lanes 1-6, Figure 18B), but not in the

control strain at the corresponding position (lane C, Figure 18B).

Figure 18. (A) Coomassie stained 12% SDS-PAGE analysis of the recombinant fusion hydrophobin

SUMO3-Bac8c-HGFI expressed in P. pastoris X-33. Lane L: Spectra™ multicolor low range protein ladder.

Lanes 1-6: six P. pastoris X-33 transformants. Line C: control strain (P. pastoris X-33 transformed with

pP-secSUMOpro3). (B) Western blot analysis of the recombinant fusion hydrophobin SUMO3-Bac8c-HGFI

expressed in P. pastoris. Lane L: Prestained protein marker (7-175 kDa, NEB). Lane C: control strain (P.

pastoris X-33 transformed with pP-secSUMOpro3). Lanes 1-6: six P. pastoris X-33 transformants. Primary

antibody: 1:1000 anti-Penta·His IgG1 (Mouse). Secondary antibody: 1:3000 Goat anti-Mouse IgG (H+L), AP

conjugate. Color development substrate: NBT/BCIP.

5.3 Minimum inhibitory concentrations of the recombinant fusion hydrophobin

Bac8c-HGFI

The SUMO3 fusion tag was removed from the fusion protein SUMO3-Bac8c-HGFI by

protease digestion followed by an affinity chromatography cleaning step. Minimum

inhibitory concentrations (MICs) of the antimicrobial peptide Bac8c and the recombinant

37

fusion hydrophobin Bac8c-HGFI against E. coli, S. aureus, P. aeruginosa and M. luteus

were measured. A summary of results can be found in Table 9. The MIC of Bac8c was 3.1

μg/ml against E. coli, S. aureus and M. luteus, respectively, and 12.5 μg/ml against P.

aeruginosa. There was no detectable growth inhibition effect of the recombinant fusion

hydrophobin Bac8c-HGFI in all tested bacteria strains.

Table 9. Minimum inhibitory concentrations (μg/ml) of the recombinant fusion hydrophobin Bac8c-HGFI

and the antimicrobial peptide Bac8c

Minimum inhibitory concentration (MIC), μg/ml

Name E. coli P. aeruginosa S. aureus M. luteus

Bac8c

Bac8ca

Bac8c-HGFI

3.1

2

N.D

12.5

8

N.D.

3.1

2

N.D.

3.1.

N.A.

N.D.

a Values were obtained from Hilpert et al. 2005.

N.A.: not available. N.D.: not detectable.

5.4 Construction of pPIC9-p11-5-linker-hgfI

The double-stranded oligo (ds oligo) fragment containing the encoding sequence of the

antimicrobial peptide P11-5 was ligated into the expression vector pPIC9-linker-hgfI and

transformed into E. coli DH5α cells. Seven ampicillin-resistant colonies growing on

selection plates were picked and analyzed by colony PCR with vector specific primers. In

lanes 2-8, a band was apparent below 200 bp. The positive control (E. coli DH5α

transformed with pPIC9-linker-hgfI) in lane 1 displayed a band around 150 bp. The

negative control (E. coli DH5α cells) in lane C did not show any bands (Figure 19C).

Figure 19. (A) Schematic diagram of the reconstructed P. pastoris expression vector pPIC9-p11-5-linker-hgfI.

(B) The amino acid sequence of the recombinant fusion hydrophobin P11-5-linker-HGFI. (C) 3%

high-resolution agarose gel displaying the colony PCR products. Lane L: Generuler™ 100 bp Plus DNA

38

ladder. Lane 1: positive control (E. coli DH5α transformed with pPIC9-linker-hgfI). Lanes 2-8: seven

ampicillin-resistant E. coli transformants. Lane C: Negative control (E. coli DH5α cells).

5.5 Expression of the recombinant fusion hydrophobin P11-5-linker-HGFI

The sequencing-confirmed expression vector pPIC-p11-5-linker-hgfI was linearized with

StuI and transformed into P. pastoris GS115 cells. Six fast-growing P. pastoris GS115

transformants on MM agar plates were picked and analyzed by colony PCR with primers

5’AOX1 and 3’AOX1. In lanes 1-6, two clear bands were apparent above 2 kbp and above

750 bp, respectively (Figure 20).

Figure 20. 1% agarose gel displaying the colony PCR amplification products. Lane L: Generuler™ 1kb DNA

ladder. Lanes 1-6: Six P. pastoris GS115 transformants.

Six P. pastoris GS115 clones identified as positive transformants by colony PCR together

with the control strain (P. pastoris GS115 transformed with pPIC9) were subjected to a

small-scale trial expression. After the expression was induced with 0.5% methanol for 96 h,

supernatant samples from each induced culture were collected, concentrated and analyzed

by Tricine-SDS-PAGE. A band with apparent molecular weight of <15 kDa was observed

in all six P. pastoris GS115 transformants (lanes 1-6, Figure 21A), but not in the control

strain at the corresponding position (lane C, Figure 21A). Dot blot analysis showed that the

eluent from the <15 kDa protein band could react positively (purple) with the protein

specific antibody (Figure 22).

39

Figure 21. (A) Silver-stained 16% Tricine-SDS-PAGE analysis of the recombinant fusion hydrophobin

P11-5-linker-HGFI expressed in P. pastoris GS115. Lane L: Spectra™ multicolor low range protein ladder.

Lanes 1-6: six P. pastoris GS115 transformants. Line C: control strain (P. pastoris GS115 transformed with

pPIC9). (B) Silver-stained 4-15% SDS-PAGE (Mini-PROTEAN® TGX™ Precast Protein Gel, Bio-Rad)

analysis of the recombinant fusion hydrophobin P11-5-linker-HGFI expressed in P. pastoris GS115

transformant one. Lane L: Spectra™ multicolor low range protein ladder. Lanes 1-5: culture supernatant

samples collected after 24, 48, 72, 96, 120 hours of induction, respectively. Lane C: control strain (P. pastoris

GS115 transformed with pPIC9).

Figure 22. Dot blot analysis of the eluent from the <15 kDa band. 1: P. pastoris GS115 transformant one.

Dot 2: P. pastoris transformant two. Dot C: control strain (P. pastoris GS115 transformed with pPIC9).

Primary antibody: anti-rHGFI antibody (rabbit source). Secondary antibody: Anti-Rabbit IgG (Fc), AP

conjugate. Color development substrate: NBT/BCIP.

Meanwhile, time course analysis to determine the optimal induction time for the

production of P11-5-linker-HGFI was performed, daily supernatant samples of

transformant one were collected and analyzed by SDS-PAGE. A protein band with

apparent molecular weight of <15 kDa was observable after 24 h, increasing in intensity up

to 96 h or 120h (Figure 21B).

5.6 Minimum inhibitory concentration of the recombinant fusion hydrophobin

P11-5-linker-HGFI

Minimum inhibitory concentration (MICs) of the antimicrobial peptide P11-5 and the

recombinant fusion hydrophobin P11-5-linker-HGFI against E. coli were measured. A

40

summary of results can be found in Table 10. The MIC of P11-5 was 3.1 μg/ml against

E. coli. The MIC of the fusion protein P11-5-linker-HGFI was 100 μg/ml against E. coli.

Table 10. Minimum inhibitory concentrations (μg/ml) of the recombinant fusion hydrophobin

P11-5-linker-HGFI and the antimicrobial peptide P11-5

Minimum inhibitory concentration (MIC, μg/ml)

Name E. coli

P11-5

P11-5a

P11-5-linker-HGFI

3.1 (3.1μM)

3.1

100 (9.1μM)

a Value was obtained from Qi et al., 2010. N.D.: no detectable.

As shown in Figure 23, abnormal cell clumps were observed, when E. coli cells were

treated with the fusion protein P11-5-linker-HGFI at concentrations (25 and 50 μg/ml)

below the MIC (100 μg/ml). The treated E. coli culture had a cloudy and yellowish

appearance.

Figure 23. Light microscopic images of E. coli ATCC 9027 untreated, and treated with 50 μg/ml and 25

μg/ml purified recombinant fusion hydrophobin P11-5-linker-HGFI for 24 h. Macroscopic appearances of

E. coli culture untreated and treated with 50 μg/ml purified recombinant fusion hydrophobin

P11-5-linker-HGFI for 24 h.

41

6 DISCUSSION

6.1 Molecular cloning, expression and characterization of the recombinant fusion

hydrophobin Bac8c-HGFI

In this study, Pichia pastoris X-33 strain was used to express the recombinant fusion

hydrophobin Bac8c-HGFI, as it secretes extremely low levels of endogenous proteins,

which in turn could facilitate purification of the expressed fusion protein from the culture

medium (Cereghino and Cregg 2000). More importantly, being a fungus, P. pastoris shares

the similar post-translation modification (PTM) strategies and extended glycosylation

modes of hydrophobins as Grifola frondosa (Wang et al. 2010b), and thus may ensure that

the hydrophobin HGFI domain of the fusion protein Bac8c-HGFI could be processed to its

optimally active form (Niu et al. 2012a; Huang et al. 2013).

Heterologous expression of antimicrobial peptides is facing two challenges. Firstly, the

peptide’s antimicrobial nature makes it potentially lethal to the expression host (Parachin et

al. 2012; Li 2009). Secondly, the peptide’s linear structure and high base amino acid

content makes it highly vulnerable to proteolytic degradation in all steps of the expression

(Raimondo et al. 2005). Thus, in order to achieve successful heterologous expression, the

antimicrobial peptide is usually fused to a chaperone protein to mask its cytotoxicity

against the host cells (Lee et al. 2000; Parachin et al. 2012). To date, many fusion

chaperone proteins have been used successfully to express a wide range of

host-unfavorable antimicrobial peptides, including thioredoxin (Xie et al. 2013), small

ubiquitin-like modifiers (SUMO) (Li et al. 2013; Wang et al. 2014) and

phosphoribosyltransferase (PurF) (Lee et al. 2000). Following this approach, the

reconstructed P. pastoris expression vector pP-secSUMOpro3-hgfI that carries the human

SUMO3 gene and the hgfI gene isolated from Grifola frondosa (GenBank Accession

NO.EF486307) was used to achieve secretory expression of the recombinant fusion

hydrophobin Bac8c-HGFI in P. pastoris.

Inverse fusion PCR cloning (IFPC) is a seamless in-frame cloning strategy, requires simple

starting materials, one step routine PCR, and generally results in the insertion of the

42

foreign gene into a wide range of plasmids with high success rates (Spiliotis 2012). To

perform a successful IFPC, high-fidelity DNA polymerases (with proofreading activity)

that generate blunt-ended PCR products should be used, since blunt ends are mandatory for

the final self-circularization step (Garrity and Wold 1992; Spiliotis 2012). In this study, the

encoding sequence of the antimicrobial peptide Bac8c (bac8c) was successfully cloned

into the vector (pP-secSUMOpro3-hgfI) backbone by IFPC, which was confirmed by

colony PCR as the corresponding band (174 bp fragment) was observed by gel

electrophoresis (lanes 5, 6 and 9, Figure 16C) and sequencing. Although the primer pair

(hgfI and bac8c) used in this study was calculated (Tm Calculator, Thermoscientific) to

have an average annealing temperature of 72℃, PCR with either lower or higher annealing

temperatures did also provide very good amplification performances (Figure 15),

demonstrating that IFPC could be successfully carried out even under some non-ideal

conditions.

On the other hand, two drawbacks of IFPC emerged in this study: (1) non-specific PCR

amplification products formed (data not shown) probably due to primer mismatching

(Arnheim and Erlich 2000; Spiliotis 2012). This problem was successfully solved with a

hot start PCR protocol to allow for more specific primer binding during the annealing step

(Zhang and Gurr 2000); (2) high background of false positive clones (5/8, lanes 2-4 and

7-8, Figure 16C), which was likely due to poly-circularization of the blunt-ended PCR

products.

To perform a reliable colony PCR screening of P. pastoris positive transformants, the

proposed protocols usually involve a lyticase pretreatment followed by extraction and

purification of genomic DNA template for PCR analysis (Linder et al. 1996; Lin et al.

2012). However, in practice, preparation of genomic DNA template is either tedious or

time-consuming (Haaning et al. 1997). Instead of this, I developed a hot SDS method for

rapid preparation of crude genomic DNA template for PCR analysis (section 4.2.7). This

method was reliably used in the whole study (Figure 17 and 20). The colony PCR results

confirmed the integration of pP-secSUMO3pro-bac8c-hgfI into all six P. pastoris X-33

43

transformants tested (1093 bp fragment, Figure 17). Meanwhile, all six transformants were

also confirmed with the phenotype Mut+ (2.2 kbp fragment, Figure 17).

Dimerization and tetramerization of hydrophobins in an SDS-PAGE gel due to the

self-assembly of hydrophobins into stable polymers in aqueous solutions (de Vries et al.

1993) was reported in many studies with hydrophobins (Hakanpää et al. 2006; Kisko et al.

2008; Wang et al. 2004; Szilvay et al. 2007). In the expression study, two unique proteins

with apparent molecular masses of 40 kDa and >40 kDa were identified in all P. pastoris

X-33 transformants (Figure 18A). These two values were inconsistent with the theoretical

value (21 kDa) of the fusion protein SUMO3-Bac8c-HGFI, which would probably be a

dimer (40 kDa ≈ 2 x 21 kDa) and a tetramer of the fusion protein SUMO3-Bac8c-HGFI.

The >40 kDa protein could react positively with the fusion protein specific primary

antibody (Figure 18B). These results indicate that the fusion protein SUMO3-Bac8c-HGFI

was expressed and secreted into the culture medium, but subsequently self-assembled into

polymers.

Protease cleavage of the SUMO3 fusion tag followed by an affinity chromatography

cleaning step was not successful as expected, only 1.04 mg fusion protein Bac8c-HGFI

was recovered from 100 mL culture broth (data not shown), probably because the

purification protocol (more details can be seen in the Ni-NTA purification system user

manual [25-0496], Invitrogen) that I used has not been optimized for hydrophobins yet. In

the following minimum inhibitory concentration test, the results showed that the highly

active antimicrobial peptide Bac8c became inactivated when it was fused with the

hydrophobin HGFI (Table 9). Similarly, in an earlier study, Rieder et al. (2014) found that

a highly active antimicrobial peptide became inactivated when it was fused with the class I

hydrophobin DewA of Aspergillus nidulans. To the best of my knowledge, steric hindrance

between the adjacent domains (Bac8c and HGFI) might have been responsible for this

inactivation (Linder et al. 2002; Iwanaga et al. 2011).

44

6.2 Molecular cloning, expression and characterization of the recombinant fusion

hydrophobin P11-5-linker-HGFI

Flexible polypeptide linkers, generally composed of small, non-polar amino acids (e.g.

glycine, serine and threonine) are often used in fusion protein studies when the adjacent

domains require a certain degree of spacing to reduce unfavorable inter-domain

interferences (Chen et al. 2013b). Following this approach, a 10-mer flexible polypeptide

linker (NH2-GGGGSGGGGS), which has been used successfully in an earlier fusion

hydrophobin study (Niu et al. 2012a) was employed to construct the recombinant fusion

hydrophobin P11-5-linker-HGFI.

The double-stranded oligo (ds oligo) fragment (p11-5) encoding the antimicrobial peptide

P11-5 was successfully subcloned into the expression vector pPIC9-linker-hgfI, which was

confirmed by colony PCR as the corresponding band (188 bp fragment) was observed by

gel electrophoresis (lanes 2-9, Figure 19C) and sequencing. When the p11-5 fragment and

the hgfI gene were fused, the linker sequence was laid between them, in order to reduce the

steric hindrance between the adjacent peptide (P11-5) and protein (HGFI), when the fused

gene (p11-5-linker-hgfI) was translated into protein (Figure 19B). In addition, the innate

nucleotide sequence encoding the STE13 protease cleavage site (Glu-Ala-Glu-Ala) was

deleted when the p11-5 fragment was inserted into the XhoI site of pPIC9-linker-hgfI.

Although Glu-Ala repeats has been recommended in many studies, in order to ensure that

the protein secretory signal (in this study, α-MF of saccharomyces cerevisiae) could be

processed correctly in the endoplasmic reticulum of P. pastoris (Sreekrishna and Kropp

1996). However, there are numerous cases where STE13 cleavage of Glu-Ala repeats is not

sufficient enough as expected, and thus Glu-Ala repeats are left on the N-terminus of the

expressed protein (Kim et al. 1997; Emberson et al. 2005), which in turn may pose a

negative effect on the activity and stability of the expressed protein (Wu and Hancock

1999). For example, Cabral et al. (2013) found that the P. pastoris produced Pisum sativum

defensin 1 (Psd1), which contained Glu-Ala repeats on its innate N-terminal region

presented at least 10-fold decrease in antifungal activity, as compared with the native

counterpart. In this study, the KEX2 protease cleavage site (Lys-Arg) was designed to be

45

located after the XhoI site, in order to ensure that the fusion protein P11-5-linker-HGFI

could be secreted into the culture medium with an intact N-terminus.

The colony PCR results confirmed the integration of pPIC9-p11-5-linker-hgfI into all six

P. pastoris GS115 transformants tested (784 bp fragment, Figure 20). All tested

transformants were also confirmed with the phenotype Mut+ (2.2 kbp fragment, Figure 20).

In the expression study, all six P. pastoris GS115 transformants could produce a unique

protein with apparent molecular mass of <15 kDa (Figure 21A). This value was

inconsistent with the theoretical value (10.3 kD) of the fusion protein P11-5-linker-HGFI.

This phenomenon was similar with those found in earlier studies (Tagu et al. 2001; Wang

et al. 2010a). For example, Wang et al. (2010a) found that the apparent molecular weight

(14 kDa) of the recombinant hydrophobin HGFI was slightly higher than its theoretical

size (10.9 kDa). This change in gel mobility is proposed to be due to an insufficient

reduction of the intramolecular disulfide bonds within hydrophobins during the sample

pretreatment step, thus resulting in a retarded gel mobility (Cumming et al. 2004; Tagu et

al. 2001). The eluent from the <15 kDa protein band could react positively with the

primary antibody raised against the hydrophobin HGFI (Figure 22), which indicates that

the fusion protein P11-5-linker-HGFI was successfully expressed and secreted into the

culture medium.

The antibacterial activity of the antimicrobial peptide P11-5 against E. coli as determined

in this study was well conformed to the literature value (Table 10). In contrast, the peptide

represented 3-time reduced antibacterial activity when it was fused with the hydrophobin

HGFI, in terms of the molecular concentration (Table 10). It is well acknowledged that the

length of polypeptide linker may affect the cytoplasmic membrane penetration efficiency

of tethered antimicrobial peptides (Qi et al. 2011). Many of the existing studies have

demonstrated that a tethered antimicrobial peptide would exhibit better activities, if a long

and flexible polypeptide linker is used (Bagheri et al. 2009). More specifically, Robinson et

al. (1998) concluded that polypeptide linkers of more than 10-mer in length are excellent

candidates for fusion proteins. In this fusion protein construct, a relatively short

46

polypeptide linker (10-mer) was used, which may still not sufficient long enough to

prevent the reciprocal interference between the adjacent domains.

There is currently no clear explanation for this abnormal aggregation behavior of E. coli

cells, when they were treated with the fusion protein P11-5-linker-HGFI at concentrations

that below the MIC (Figure 23). It is well known that the cell-wall-associated

hydrophobins play an important role in facilitating the aggregation of filamentous hyphal

cells into large clumps (Talbot 1997). Based on this fact, Nakari-Setälä et al. (2002)

engineered the cell wall of Saccharomyces cerevisiae with the class II hydrophobin HFBI

of Trichoderma reesei, increased cell-cell adhesion (clumping) ability was observed among

these hydrophobin-modified yeast cells, as compared with the unmodified counterpart. The

proposed cell-killing mechanism of the antimicrobial peptide P11-5 is the ‘carpet-like’

mode of action. Therefore, if the concentration of peptides is not sufficient enough to

disintegrate the bacterial cytoplasmic membrane, they are bound parallel onto the bacterial

cytoplasmic membrane (Brogden 2005), thus leading to the immobilization of the attached

hydrophobins on the bacterial surface. In the aqueous environment, these bacterial surface

immobilized hydrophobins would self-assembly into polymers, and thereby resulting in the

aggregation of bacterial cells.

6.3 Future work: specific-protease-enhanced antibacterial fusion hydrophobins

Many pathogenic microorganisms possess unique proteases to invade host organisms

(Aoki and Ueda 2013). For example, Listeria monocytogenes, the bacterium that causes the

invasive infection listeriosis, secretes a stress-induced serine protease that plays a crucial

role in intracellular survival of this pathogen (Gaillot et al. 2000). On the basis of this,

designing fusion antimicrobial peptides that can be activated by species-specific proteases

may provide high selectivity toward each target pathogenic microorganism. Following this

postulation, Aoki et al. (2012) designed a fungicidal fusion protein (Figure 24) that could

be exclusively activated by a unique aspartic protease of Candida albicans. This peptide

showed a highly selective in vitro activity against C. albicans, but not against other tested

microorganisms, which do not possess the corresponding aspartic protease.

47

Figure 24. Schematic diagram of the aspartic protease-activated fungicidal fusion protein. This peptide is

composed of three functional domains: an antimicrobial peptide, a protective peptide and a

protease-cleavable linker. Generally, the fungicidal activity of the tethered antimicrobial peptide is inhibited

by the protective peptide. Species-specific protease cleaves the linker and subsequently releases the

antimicrobial peptide, causing its activation (Figure adapted from Aoki and Ueda, 2013).

The potential cleavage site analysis of the fusion protein P11-5-linker-HGFI demonstrated

that this protein contains an endoprotease AspN cleavage site, which is located in the

C-terminal region of the antimicrobial peptide. Therefore, upon the enzymatic digestion

with AspN, the antimicrobial peptide P11-5 could be neatly released from the fusion

protein. In the future work, once we confirmed that the enzymatic digestion could improve

the antibacterial activity of the fusion protein. By using the design approach described by

Aoki et al. (2012), a series of specific-protease-enhanced antimicrobial hydrophobins could

thus be designed.

48

7 CONCLUSIONS

In summary, two recombinant fusion hydrophobins, Bac8c-HGFI and P11-5-linker-HGFI

were successfully expressed and secreted into the culture medium. Similar to hydrophobins,

the fusion protein Bac8c-HGFI could form stable polymers in the aqueous environment.

The highly active antimicrobial peptide Bac8c became inactivated when it was fused with

the hydrophobin HGFI. Interestingly, the hydrophobin HGFI gained an acquired

antibacterial nature when it was fused with the antimicrobial peptide P11-5 through a

10-mer flexible polypeptide linker, with the minimum inhibitory concentration of 100

μg/ml against Escherichia coli. However, there is currently no clear explanation for the

abnormal aggregation behavior of E. coli cells in this study. Further experiments are

needed to clarify this issue.

49

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