IMPROVED METHODS FOR PRODUCTION AND … · 2010. 12. 21. · Jembrana disease is an acute disease of Bali cattle (Bos javanicus) in Indonesia caused by Jembrana disease virus (JDV),

IMPROVED METHODS FOR PRODUCTION AND CHARACTERISATION OF

JEMBRANA DISEASE VIRUS PROTEINS

This thesis is presented for the degree of Doctor of Philosophy of Murdoch University

Judhi Rachmat

2010

i

DECLARATION

I declare that this is my own account of my research and contains as its main content work which has not previously been submitted for a degree at any tertiary

education institution

...................................... Judhi Rachmat

ii

Abstract

Jembrana disease is an acute disease of Bali cattle (Bos javanicus) in Indonesia

caused by Jembrana disease virus (JDV), a lentivirus most closely related to Bovine

immunodeficiency virus. Control of the disease in Bali cattle, which are important to

the economy of Indonesia, is dependent on the continued availability of protein

antigens for immunosurveillance procedures that have been developed. Further

investigation is also required to characterise of the proteins of JDV and to provide

methods of producing commercial quantities of recombinant proteins for vaccine

manufacture.

A problem with the large scale production of viral proteins using recombinant

technology was that the proteins have been mainly produced as insoluble products

within inclusion bodies in bacterial cells and in that insoluble format they were

unsuitable for use as antigens. A method for solubilisation of the insoluble proteins

was developed that involved solubilisation of the inclusion bodies with low

concentrations of urea in an alkaline solution and the method could be performed

easily and at low cost without any detectable loss of antigenicity. The solubilised

protein was successfully renatured without the formation of aggregates by dilution of

the urea in the presence of the reducing agent dithiothreitol. The method would be

suitable for use during the large scale production of recombinant viral proteins for

vaccine manufacture.

To provide an additional reagent for diagnosis of the disease, mice were immunised

with the recombinant capsid (CA) protein of JDV and a hybridoma was produced

that secreted monoclonal antibodies reactive with the CA protein. This monoclonal

antibody was effectively used in an immunoperoxidase assay to demonstrate virus

in tissues. As an alternative to this technology, recombinant antibody fragments,

scFv, reactive with the CA protein of JDV, were also produced by phage display

technology. These scFv were expressed as soluble products in the periplasmic

space of transfected host bacterial cells. The scFv reacted specifically with the CA

protein in western immunoblots and although further optimisation of the methods of

production of this scFv are required, the reagents developed can be for expression

of the antibody when required, without the need for maintaining liquid nitrogen

storage facilities that are necessary for storage of hybridomas.

The size and nature of the glycosylation of the envelope proteins SU and TM of JDV

harvested from infected cattle was determined. Two proteins of 75 and 60 kDa were

initially identified in SDS-PAGE as the SU and TM, respectively. They were initially

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identified using a specific glycoprotein stain and then matrix-assisted laser

desorption/ionisation-time of flight (MALDI-TOF) mass spectrometric (MS) analysis

of the protein sequence. Further investigation on the unmatched mass value from

MALDI-TOF/MS data suggested that post-translational glycosylation occurred on N-

linked glycosylation sites of the JDV-SU and on O-linked glycosylation sites of the

JDV-TM. This is the first report of the characteristics of the envelope proteins of JDV

and their identification will facilitate further studies of the nature of the immune

response to JDV infection using immunoblotting procedures.

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Acknowledgements

There are many people to thank. Without the help and understanding from the

people around me, the way to this thesis would have been impossible.

My PhD study at Murdoch University would not have been possible without help and

extraordinary support from my supervisor, Prof. Graham E. Wilcox who gave me the

opportunity to do my PhD in the School of Veterinary and Biomedical Sciences,

Murdoch University. I am especially grateful to him for supervising and giving me a

lot of valued advice and constant attention. I owe a great deal to his dedication and

thorough approach. I will never forget his kindness or his words of encouragement.

I would also like to take this opportunity to thank Moira Desport for stimulating

discussion and assistance in the theoretical aspects of the project in the virology

group. A special word of thanks to William Ditcham who has been of great

assistance in discussion of the necessary lab work. Linda Davies deserves a special

mention for her much appreciated assistance related to the production and

maintenance of the hybridoma cells, a crucial component of the study. Work would

have been far less enjoyable without the company of former and current group

members Meredith Stewart, Surachmi Setiyaningsih, Emilija Filipovska-Naumovska,

Mark O’Dea, Andrew Hughes, Joshua Lewis, Tegan McNab and I Wayan Masa

Tenaya. I have really enjoyed being around smart people in a stimulating

environment. I wish you all the best in your careers.

My Ph D work would not have been possible without the financial support from the

John Allwright, ACIAR (Australian Centre for International Agricultural Research)

Fellowships. I would like to acknowledge Dr. John W. Copland and Mrs. Sharon

Harvey from ACIAR, Ms. Anne Randell and Ms. Karen Olkowski from Murdoch

University for taking care of all the organization and administrative tasks needed for

my study at Murdoch University.

This page would not be complete without mentioning the great love and support I

have received from my parents, my brothers and my sisters. My debt to them is truly

without bound and is one that can never be repaid. I can only offer them all the

thanks and love that a son and a brother can give. Finally, this work is also

dedicated to my beautiful daughter, Anindya Aaqila Nurjannah Raisyaputri. May

God bless my daughter with wonderful life.

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Table of contents Page

Declaration i

Abstract ii

Acknowledgements iv

Table of contents v

List of abbreviations vi

Chapter 1. General introduction 1

Chapter 2. Review of the literature 3

Chapter 3. Solubilisation and purification of insoluble recombinant JDV-ΔSU present in inclusion bodies

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Chapter 4. Development of monoclonal antibody for the detection of recombinant JDV proteins

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Chapter 5. Production of single chain fragment antibody (scFv) against recombinant ΔSU and CA proteins of Jembrana disease virus

72

Chapter 6. Identification and preliminary analysis of JDV envelope glycoproteins by MALDI-TOF mass spectrometry

97

Chapter 7. General discussion 113

References 117

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List of abbreviations Viruses BIV : Bovine immunodeficiency virus BHV : Bovine herpesvirus BPV : Bovine papillomavirus CAEV : Caprine arthritis-encephalitis virus EIAV : Equine infectious anaemia virus FIV : Feline immunodeficiency virus HIV : Human immunodeficiency virus JDV : Jembrana disease virus MVV : Maedi-visna virus SIV : Simian immunodeficiency virus Reagents Amp : ampicillin DAB DMEM

: 3,3' diaminobenzidine tetrahydrochloride Dulbecco’s modified Eagle’s medium

DMSO : dimethyl sulfoxide DTT : dithiothreitol EDTA : ethylenediamine tetra-acetic acid FCS : foetal calf serum HAT : hypoxanthine-aminopterin-thymidine HT : hypoxanthine-thymidine IPTG : isopropylthiogalactoside LB PBS

: Luria-Bertani phosphate buffered saline

PEG : polyethylene glycol PMSF : phenylmethylsulfonyl fluoride HRP SDS

: horseradish peroxidase sodium dodecylsulfate

TBS : tris-buffered saline TEMED : N,N,N',N'-tetramethylene-ethylenediamine Tris : tris (hydroxymethyl) aminoethane Other aa : amino acid Ag : antigen bp : base pairs CA CDR

: capsid complementary determining region

cfu : colony forming units DNA : deoxyribonucleic acid dpi : day post inoculation ELISA : enzyme-linked immunosorbent assay Fab : fragment antibody GST : glutathione transferase

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HRP : horseradish peroxidase IN ISH LTR

: integrase in situ hybridisation long terminal repeat

MA MAb

: matrix monoclonal antibody

MALDI-TOF : matrix-assisted laser desorption/ionisation – time of flight MS : mass spectrometry NC PAGE PCR

: nucleocapsid polyacrylamide gel electrophoresis polymerase chain reactions

PR RNA

: protease ribonucleic acid

RT : reverse transcriptase RT-PCR : reverse transcription-PCR scFv : single chain variable fragment SDS SU TM VH

: sodium dodecyl sulfate surface unit transmembrane glycoprotein heavy chain variable domain

VL : light chain variable domain

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Chapter 1

General introduction

Cattle in Indonesia, and especially Bali cattle (Bos javanicus), contribute to poverty

reduction by improving efficacy in integrated farming systems. They are used as

draught animals for rice production, for meat production, for grazing on land otherwise

not used, and via manure contribute to soil fertility. The utilisation of cattle as draft

power enables smallholder farmers to double food production from their available

land. While Bali cattle have a number of advantages under Indonesian conditions,

and have been widely distributed throughout Indonesia, they have disadvantages.

A major disadvantage is their unique susceptibility to Jembrana disease and this

disease is therefore a major threat to the success of the various Bali cattle

distribution programs and consequently to the attempts to increase food production

in Indonesia.

The discovery that Jembrana disease is caused by Jembrana disease virus (JDV) a

lentivirus most closely related to Bovine immunodeficiency virus has led to the

development of a variety of diagnostic reagents and a tissue-derived vaccine that in

turn have led to better control of the disease. Molecular biological methods offer

extensive promise for improvement in the initial diagnostic reagents and vaccines

that were developed. The aims of the research reported in this thesis were to

improve the methods developed previously for the production of recombinant

proteins for antigens and for vaccine production, the development of monoclonal

antibody (MAb) reagents for diagnostic techniques, and further characterisation of

the envelope proteins of JDV that would enable enhanced diagnostic methods. As a

background to these investigations, a review of Jembrana disease and JDV, and a

review of literature related to the production of recombinant proteins, MAb and post-

translational glycosylation of viral proteins was undertaken and is incorporated in

Chapter 2.

Recombinant proteins of JDV have been produced for many years with a bacterial

expression system but a major problem during their production by this system has

been their production as insoluble proteins within inclusion bodies, and there has

been minimal production of soluble protein, resulting in poor yields of protein. To

increase yields, a method of solubilisation of the insoluble recombinant surface unit

(SU) and capsid (CA) proteins of JDV was investigated, which would potentially

increase the yield of these and other proteins produced by this expression system

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and enable their use as antigens in immunological assays and as experimental

vaccines. The results of this investigation are reported in Chapter 3.

A MAb against the CA protein of JDV has been produced previously and this has

provided extremely useful reagents for the development of diagnostic assays for the

detection of JDV. Unfortunately, maintenance of the hybridomas in Indonesia has

been difficult due to lapses in the supply of liquid nitrogen and the hybridoma was

lost. It was also hypothesised that additional MAbs against the envelope

glycoproteins (SU and TM) could provide a means of identifying the size of the

glycosylated SU and TM glycoproteins of JDV in SDS-PAGE gels. Attempts were

therefore made to produce additional MAb by conventional hybridoma technology

and these results are reported in Chapter 4. Recent developments in combinatorial

antibody libraries combined with the display of functional antibody fragments at the

tips of filamentous phage, known as phage display system, allows direct selection of

highly specific MAbs from naive combinatorial antibody libraries. It was

hypothesised that this technology would lead to the selection of stable clones that

could be used to express MAbs in a bacterial expression system and overcome

problems of long-term storage of hybridomas. Attempts were therefore made to

produce recombinant antibodies against the SU and CA protein of JDV using the

Tomlinson I and J libraries, and these investigations are described in Chapter 5.

Successful production of MAbs against the CA proteins was achieved but was not

successful against the envelope proteins of JDV. Therefore, to identify these

proteins in native JDV preparations, an attempt was made to characterise 2

glycosylated proteins present in JDV by mass spectrometric analysis and these

results are described in Chapter 6. Additional analysis of the mass spectrometric

data for 2 glycosylated proteins of 75 and 60 kDa, that were identified as SU and

TM, were undertaken to determine the nature of their polysaccharide moieties and

these results are also reported in Chapter 6.

A general discussion of the research results reported in the thesis and

recommendations for future research related to these results are presented in

Chapter 7.

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Chapter 2

Review of the literature

This Chapter contains a review of the literature relevant to the research undertaken

and reported in this thesis, which concerns the development of reagents for the

diagnosis and control of Jembrana disease and further characterisation of the

envelope glycoproteins of the bovine lentivirus, Jembrana disease virus (JDV). This

review is arranged into several sections: an initial section providing background

information on the family Retroviridae (retroviruses); a section on the bovine

lentiviruses including JDV and Bovine immunodeficiency virus (BIV); a section

describing technical aspects of the production of soluble recombinant viral proteins;

a section describing the technical aspects of the production of monoclonal antibody

using conventional and recombinant techniques.

2.1 Characteristics of the Retroviridae

The family Retroviridae comprises a large group of viruses that have been detected

in many vertebrate species. They have shared structural and genomic features and

similar modes of reproduction but vary considerably in the type of disease with

which they are associated.

All retroviruses have a genome consisting of 2 identical strands of single-stranded

RNA enclosed within a protein coat (capsid) which is again enclosed by a lipid

envelope. The viral envelope is formed from the plasma membrane of the cell as

the virus is released from the cell and it contains virus-encoded glycoproteins

important in attachment and penetration of the virus into the cell (Temin & Mizutani,

1970).

The retroviruses are unique in that their replication requires a reversal of the normal

flow of genetic information. In all living organisms and many viruses, genetic

information is stored as DNA and later transcribed into RNA, which serves as a

template for protein synthesis. In contrast, retroviruses store their genetic

information as RNA and they also contain the unique enzyme, reverse transcriptase

(RT) that catalyses the reverse transcription of the RNA genome into DNA. All

retroviruses contain 3 major open reading frames designated, from the 5’– to 3’-

ends of the genome, gag, pol and env genes that are translated as polyproteins:

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gag encoding the core proteins, which include the matrix (MA), the capsid (CA) and

the nucleocapsid (NC), pol encoding the enzymatic proteins RT, integrase (IN) and

protease (PR), and env encoding the envelope surface unit (SU) and

transmembrane (TM) glycoproteins. The SU contains the determinants that interact

with the host cell receptor and coreceptor, while TM not only anchors the SU/TM

complex in the membrane but also contains domains that are critical for catalysing

the membrane fusion reaction between viral and host lipid bilayers during virus

entry. Human immunodeficiency virus (HIV) is one of the members of this family and

a detailed understanding of the molecular biology of HIV is available in the review

by Wang et al. (2000).

The replication of retroviruses is divisible into early and late phases (Turner &

Summers, 1999) as depicted in Figure 2.1. Retroviruses recognise potential host

cells through specific interactions between a host cell receptor and the viral

envelope membrane glycoprotein SU. The main receptor for HIV-1 and HIV-2 is

CD4 on T lymphocytes whilst the main co-receptors are the α-chemokine CXCR4

and the β-chemokine CCR5 (Dragic et al., 1996). After binding, fusion of the viral

envelope with the cellular membrane occurs directly at the cell surface releasing the

virus core into the cytoplasm of the cell (early phase). In the late phase of

replication, the viral genome is reverse transcribed, via the action of RT, into

double-stranded DNA and integrated into the genome of the host cell where it

resides permanently as the “provirus”. Viral protein is expressed from spliced

messenger RNA transcripts of the proviral DNA and some such as SU and TM

require post-translational glycosylation; these glycoproteins are incorporated into

regions within the plasma membrane where the viral RNA genome is packed into

capsids that bud from the membrane along with the viral envelope proteins. The

cell-free particles undergo a further maturation process before they are capable of

productive infection in appropriate target cells.

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Figure 2.1. Schematic diagram illustrating the features of the retrovirus replication cycle. Following recognition of a specific cellular receptor by the viral envelope glycoprotein and adsorption of the virion to the cell surface, the viral core is released into the cell cytoplasm. The viral RNA is uncoated and reverse transcribed into a double-stranded DNA. The integrated provirus uses a combination of viral and cellular transcription factors to replicate new copies of its RNA genome. Some of these viral RNAs are exported to the cytoplasm to serve as new viral genomes, however a percentage of the viral RNA is spliced into smaller mRNA species that are translated by the host cell ribosomal machinery. The viral regulatory proteins Tat and Rev tightly control transcription and transport of viral mRNA to the cell cytoplasm and both proteins are essential for retrovirus replication. Figure from Turner & Summers (1999).

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Taxonomy

The retroviruses are subdivided into 7 genera: Alpharetrovirus, Betaretrovirus,

Gammaretrovirus, Deltaretrovirus, Epsilonretrovirus, Spumavirus and Lentivirus

(Table 2.1). Previously, these viruses were grouped according to the nature of the

diseases they produced and their electron microscopic appearance within infected

cells and the core in the mature virus: concentric nucleocapsids were attributed to

alpharetroviruses, gammaretroviruses, deltaretroviruses and spumaviruses; a rod or

truncated cone-shaped core was characteristic of lentiviruses and betaretroviruses.

Table 2.1. Genera in the family Retroviridae and their principal hosts and associated disease.

Genus Type species Hosts Genome of type species

Pathological features of disease

References

Alpharetrovirus Avian leukosis virus (ALV)

Avian 7200 bp

Oncogenic (Venugopal, 1999)

Betaretrovirus Mouse mammary tumor virus (MMTV)

Mammalian 8805 bp

Oncogenic (Cardiff & Wellings, 1999)

Gammaretrovirus Murine leukemia virus (MLV)

Mammalian 8256 bp Oncogenic (Chesterman et al., 1966)

Deltaretrovirus Bovine leukemia virus (BLV)

Mammalian 8000 bp Oncogenic (Van der Maaten et al., 1982, Willems et al., 2000)

Epsilonretrovirus Walleye dermal sarcoma virus (WDSV)

Fish 12700 bp

Oncogenic (Holzschu et al., 2003, Holzschu et al., 1995, Zhang et al., 1996)

Lentivirus Human immunodeficiency virus (HIV)

Humans 9869 bp

Lymphocyte-tropic associated with immunosuppression Macrophage-tropic strains associated with arthritis, pneumonia, encephalitis, anaemia

(Weber, 1989)

Spumavirus Human spumavirus

Simians 13246 bp

Subclinical infections

(Kupiec et al., 1991)

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Pathogenicity

The retroviruses may also be divided into 3 groups according to their pattern of

pathogenicity: the oncogenic retroviruses or oncoviruses, many of which are

associated with leukaemias and sarcomas; the spumaviruses that do not seem to

be associated with any disease; lentiviruses, many of which are associated with

immunodeficiencies, but some of which are associated with neurological lesions,

some with arthritis and some with pneumonia (Weiss, 1996).

Most retroviruses are transmitted laterally but the oncoviruses or cancer causing

viruses may also be transmitted vertically by integration of the provirus into the

genome of germ cells (Mims, 1981). The proviral form of these oncoviruses can be

associated with transformation of the host cells into cells that have a tumour

producing potential (Maeda et al., 2008).

Retroviruses characteristically persist in infected animals for the life of the animal

(Wells & Poiesz, 1990). Some produce clinical disease only after prolonged

incubation periods, some produce intermittent clinical disease associated with

reactivation of virus and expression of clinical disease after periods of latency

(Meiering & Linial, 2002). During the periods of latency the virus may be detectable

but it is often difficult to detect because it is present at only low levels, necessitating

indirect means such as the detection of antibody for the recognition of virus

infection.

In the period of clinical latency following HIV infection, there is still detectable virus

in the peripheral blood; it was suggested that this virus is derived from infected

CD4+ lymphoblasts that have reverted to a resting memory state (Marcello, 2006).

In FIV infection, it was suggested that CD4(+)CD25(-) cells provide latent viral

reservoirs for FIV infection and CD4(+)CD25(+) represent ideal candidates for a

productive FIV infection (Joshi et al., 2005a, Joshi et al., 2005b, Joshi et al., 2004).

In Equine infectious anaemia virus (EIAV) infections, the proviral DNA was present

in tissues regardless of disease status, particularly in macrophages that are the

primary cellular reservoir and site of viral replication of EIAV (Oaks et al., 1998).

Latency is not unique to retrovirus infections and is also seen in other virus types

such as herpesviruses. However, the mechanism of latency or persistence of the

virus in these groups differs from that in retroviruses. Herpesviruses persist

between periods of clinical disease in lymphoid cells or in ganglionic neurons of the

peripheral nervous system. For example, Bovine herpes virus type-1 (BHV-1)

localises and persists in ganglionic neurons of the peripheral nervous system

8

(Jones et al., 2006) and in germinal centres of pharyngeal tonsil (Winkler et al.,

2000). The virus within these sites exhibits limited replication: nested reverse

transcription-PCR (RT-PCR) on latently infected cattle showed a few cells contained

latency-related transcripts but not other immediate-early, and late transcripts

(Winkler et al., 2000). Latency has also been detected in the circulating lymphocyte

of Bovine papillomavirus-1 infected cattle (Campo et al., 1994) and in neoplastic

and non-neoplastic tissues of horses (Carr et al., 2001). Reactivation of viral

expression after periods of latency is often indicated by the appearance of infectious

virus at the site of the initial infection in an immune host.

Lentiviruses

The lentiviruses can be divided into 4 groups (Table 2.2) on the basis of the host

with which they have evolved: primate (including the 2 human virus types, HIV-1

and HIV-2, and multiple Simian immunodeficiency virus [SIV] types), equine (EIAV),

feline (Feline immunodeficiency [FIV]), ruminant lentiviruses including bovine

(including BIV and JDV), and small ruminant lentiviruses infecting sheep (Maedi-

visna virus [MVV]) and goats (Caprine arthritis encephalitis virus [CAEV] (Clements

& Zink, 1996).

Lentiviruses, like other retroviruses, contain the obligatory major open reading

frames gag, pol and env. The gag open reading frame encodes a precursor

polyprotein (Gag) that is cleaved to form the non-glycosylated structural proteins

MA, CA and NC. The NC is associated with the viral RNA genome and required for

packaging RNA into the virion (Coffin, 1979, Coffin, 1992). NC is also essential for

the 2 obligatory strand transfers during viral DNA synthesis via promotion of primer

binding site homo-dimer (Egele et al., 2004, Huthoff & Berkhout, 2001). The CA

forms the core of the virion, it is the most immunodominant viral protein (Coffin,

1979, Coffin, 1992) and the conserved C-terminus of the CA is important in virion

assembly and release (Melamed et al., 2004).

The pol open reading frame encodes 3 enzymatic proteins, RT, IN and PR that are

utilised during the replication process. RT is essential for the transcription of the

viral RNA genome to a dsDNA intermediate or provirus (Temin, 1993). IN is an

essential protein involved in incorporation of the provirus DNA into the host cell

genome, and PR is vital for the cleavage of the viral polyproteins into individual

subunit proteins during virus replication (Coffin, 1979, Coffin, 1992).

Table 2.2. Principle distinguishing characteristic of viruses in the genus Lentivirus

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Group Species Host Genome size/accessory genes

Tropism/disease Reference

Primate group

Human immunodeficiency virus (HIV)

Humans 9869 bp

Primarily lymphocyte-tropic/acquired immunodeficiency syndrome

(Weber, 1989)

Simian immunodeficiency virus (SIV)

Non-human primates

8816 bp Primarily lymphocyte-tropic/acquired immunodeficiency syndrome

(Ringler et al., 1988, Stephens et al., 1997)

Ruminant group

Caprine arthritis-encephalitis virus (CAEV)

Goat 9065 bp Macrophage-tropic/arthritis, encephalitis and pneumonia

(Olsen, 2001)

Maedi-visna virus

Sheep 9203 bp monocyte/macro-phage-tropic/ pneumonia, arthritis and mastitis

(Sargan et al., 1991)

Bovine immunodeficiency virus (BIV)

Cattle 8482 bp Lymphocyte and macrophage-tropic/possible immunosuppression

(Gonda et al., 1994)

Jembrana disease virus (JDV)

Cattle 7732 bp Lymphocyte-tropic/acute disease affecting

(Wilcox et al., 1995)

Feline group

Feline immunodeficiency virus (FIV)

Domestic and large cats

9891 bp Primarily lymphocyte-tropic/acquired immunodeficiency syndrome

(Parodi et al., 1994, Pecon-Slattery et al., 2008)

Equine group

Equine infectious anaemia virus (EIAV)

Horse 8249 bp

- lacks gene encoding Vif, but contains gene S2

Macrophage-tropic /intermittent anaemia

(Olsen, 2001, Yoon et al., 2000)

In lentiviruses, the polyprotein Env is translated and subsequently glycosylated in

the endoplasmic reticulum where it also oligomerises (Chan et al., 1997). In the

golgi, the attached oligosaccharides are processed and the glycoprotein is

proteolytically cleaved into 2 subunits, the SU and TM, by a cellular PC6 protease of

10

the subtilisin-like pro-protein convertase family. The glycoprotein complex is

subsequently transported to the plasma membrane via the secretory pathway

(Miranda et al., 2002). The TM subunit anchors the SU subunit to the viral surface

via a disulphide bond and together they form a homotrimer, which is situated on the

surface of viral particles. The external SU subunit determines receptor specificity

while the TM subunit is responsible for the fusogenic process during entry (Eckert &

Kim, 2001).

The lentiviruses possess unique properties that differentiate them from other

retroviruses, including their ability to infect both dividing and non-dividing cells

(Lewis et al., 1992, Weinberg et al., 1991) and by their possession of additional

accessory genes important in their replication. In addition to the major open reading

frames, lentiviruses have 2 regulatory genes and up to 4 accessory genes, the

number varying with the species of lentivirus, making them the most complex

members of the retroviruses. The regulatory and accessory genes are located

mainly between the pol open reading frame and the 3’ end of the genome, and may

include genes designated tat, rev, vif, nef, tmx, vpu, vpr, vpx, vpw, vpy, S2 and orfA.

They collectively regulate viral transcription, translation, and aspects of viral

pathogenicity (Cullen, 1991).

The primate lentiviruses are the most complex of all retroviruses, possessing at

least 6 accessory genes including tat, rev, vif, vpr, vpx or vpu and nef (Tristem et al.,

1992). All lentiviruses possess tat and rev, which are expressed from multiply

spliced transcripts, encode trans-acting regulatory proteins, and are essential for

viral replication (Dorn et al., 1990, Gonda, 1992, Narayan, 1990). Most lentiviruses

also possess vif, which is expressed from a singly spliced transcript, and encodes a

virion-associated protein also essential for viral replication (Schrofelbauer et al.,

2004, Volsky et al., 1995).

2.2 Bovine lentiviruses

2.2.1 Jembrana disease virus

History

Jembrana disease is an economically important infectious disease of Bali cattle that

emerged for the first time in the latter months of 1964 in the Jembrana district of

Bali, Indonesia (Pranoto & Pudjiastono, 1967). This outbreak spread throughout the

island over the ensuing 12 months and then the prevalence waned, perhaps a

11

consequence of herd immunity, and it was not reported again for several years.

Second and the third outbreaks of Jembrana disease were subsequently detected

that included one in the Tabanan district in 1972 and one in the Karangasem district

in 1981 (Hardjosworo & Budiarso, 1973, Putra et al., 1983). Despite attempts to

implement quarantine measures to prevent the movement of cattle from Bali, the

disease has subsequently spread to some other Indonesian islands: Lampung

province in South Sumatra (Soeharsono & Darmadi, 1976); the Banyuwangi district

of East Java (Tranggono, 1988); the Sawahlunto district of West Sumatra (Tembok,

1992); in the 1990s in South Kalimantan and then to West and East Kalimantan

(Hartaningsih, personal communication). The initial occurrence of the disease in

these areas was associated with high mortality rates and has since become

endemic with lower mortality rates (Soeharsono, 1997).

Clinical features of Jembrana disease

The disease can be readily transmitted to naïve cattle by the inoculation of tissues,

including blood and lymphoid tissues, from affected cattle; high titres of infectious

virus are present in the plasma and spleen of affected animals during the course of

the acute disease (Soeharsono et al., 1990). Experimental transmission studies

have confirmed field observations that the disease is an acute (transient) febrile

condition with characteristic clinical findings during the acute disease process

including anorexia, lethargy, fever, erosions of the oral mucous membranes and

enlargement of superficial lymph nodes. Other clinical signs less consistently

observed were hypersalivation, a nasal discharge, diarrhoea with blood in the

faeces and pallor of the mucous membranes (Soeharsono et al., 1990, Soesanto et

al., 1990). In experimentally infected cattle housed indoors the case fatality rate

associated with infection by the Tabanan/87 strain was 17% (Soesanto et al., 1990).

Recovered cattle are viraemic for at least 2 years after infection, possibly for the life

of the animal, and are therefore an important potential source of infection for other

cattle. However, in animals that recover there are no reports of the recurrence of

disease suggesting that these animals develop a solid immunity to infection.

Marked haematological changes are detected in affected Bali cattle: leukopenia as

a result of a lymphopenia, eosinopenia and neutropenia, thrombocytopenia,

anaemia, increased blood urea concentrations and diminished total plasma protein

levels; these changes occur principally during the febrile period (Soesanto et al.,

1990). Gross pathological changes include vascular damage such as mild exudates

12

and haemorrhages, but the most striking changes are lymphadenopathy and

splenomegaly. Lymphoid tissues of all organs, particularly in the enlarged lymph

nodes and spleen, feature proliferating lymphoblastoid cells predominantly

throughout parafollicular (T-cell) areas, and atrophy of follicles (B-cell areas). A

proliferative lymphoid infiltrate is also found in the parenchyma of most organs,

particularly the liver and kidneys and an infiltrate containing proliferative

macrophage-like cells is found in the lungs (Dharma, 1997).

Jembrana disease virus appears to have a particular affinity for Bali cattle and it is

only in this species that severe lesions and case fatalities appear to occur

(Soeharsono et al., 1995a). Other cattle types and buffalo, however, can be

infected experimentally and become infected under field conditions. Bos taurus,

Bos indicus and crossbred Bali (Bos javanicus x Bos indicus) cattle and buffalo

develop disease and a persistent viraemia (Soeharsono et al., 1995a). The clinical

changes and lesions that occur in these cattle types are consistent with those

observed in Bali cattle, but they are much milder and would be more difficult to

detect under field conditions (Wilcox et al., 1995).

Although only limited studies of the disease in Bos taurus have been conducted, the

studies have indicated that the effects of infection in Bos taurus are less severe than

in Bos javanicus. Infection of Friesian cattle induced an acute disease after a short

incubation period of about 4 days. Clinical signs were fever and concurrent

lymphadenopathy. Haematological changes included leukopenia as a result of

lymphopenia, neutropenia and thrombocytopenia (Soeharsono et al., 1995a).

Increased blood urea concentration and signs of anaemia consistently detected in

Bali cattle were not detected in Friesian cattle. Histological lesions consistent with a

mild form of Jembrana disease in Bali cattle were detected in some infected

animals. However, while there was follicular atrophy until 5 weeks after infection in

the spleen and lymph nodes of Bali cattle there was a marked follicular response in

the spleen and lymph nodes of one Friesian animal in the immediate post-febrile

period (Soeharsono et al., 1995a). Friesian cattle experimentally infected with JDV

had no parafollicular proliferation of mononuclear cells in intestinal lymphoid tissue

or haemorrhagic lesions (Soeharsono et al., 1995a).

Since the first outbreak of Jembrana disease in 1964, there have been several

hypotheses regarding the cause of the disease. The condition was initially

considered to be caused by rinderpest virus (Adiwinata, 1968, Pranoto &

Pudjiastono, 1967). Another theory was subsequently developed that the disease

was caused by a rickettsia-like agent (Hardjosworo & Budiarso, 1973) and this

13

persisted until the disease was demonstrated to be caused by a retrovirus (Wilcox

et al., 1992) and subsequent sequence analysis confirmed it was a lentivirus, most

closely related to BIV (Chadwick et al., 1995b, Lu et al., 2002a).

Genome of JDV

The genome of the Tabanan/87 strain of JDV, the only strain that has been

completely sequenced, is 7732 bp. A schematic representation of the genome of

JDV is shown in Figure 2.2 and the molecular characteristics of the JDV proteins

(Chadwick et al., 1995b) are shown in Table 2.3.

Table 2.3. Predicted characteristics of protein products encoded by the JDV genome. Data sourced from Chadwick at al. (1995b).

Coding area gene product

Protein Number of amino acid residues

Predicted Mr (kDa)

Gag Matrix (MA) 125 14.3

Capsid (CA) 226 25.3

Nucleocapsid (NC) 85 9.2

Gag/Pol Precursor

1432 163

Pol precursor 1027 118

Env Surface unit (SU) 422 47.8

Transmembrane (TM) 359 41.1

Vif 197 22.9

Tat 97 10.7

Rev 213 23.8

Tmx 164 18.5

The accessory genes of the bovine lentiviruses are less complex than that of the

primate lentiviruses. JDV contains 4 small putative accessory genes designated vif,

tat, rev and tmx that correspond to the accessory genes detected in BIV and are

assumed to have similar functions in both viruses (Chadwick et al., 1995a). While

the functions of the HIV-1 accessory genes have been extensively investigated and

reviewed (Cullen & Greene, 1990), in both bovine viruses their functions remain

largely uncharacterised. It is known that JDV Tat is very potent and can strongly

activate not only its own long terminal repeat (LTR) but also the HIV LTR. In

14

contrast, HIV Tat cannot reciprocally activate the JDV LTR (Chen et al., 1999) and

JDV Tat can functionally substitute for HIV Tat (Chen et al., 2000). Like HIV Tat,

JDV Tat transactivates the HIV LTR at least partially in a TAR-dependent manner.

However, the sequence in the loop region of TAR was not as critical for the function

of JDV Tat as it was for HIV Tat (Chen et al., 2000). The BIV vif (Oberste & Gonda,

1992) like the HIV-1 vif (Goncalves et al., 1996) facilitates the infectivity and spread

of virus.

Figure 2.2. The JDV genome resembles that of BIV and HIV type 1 with the typical 5’ to 3’ gag, pol and env gene organisation. It contains also the regulatory protein encoding genes between or overlapping the pol and env reading frames. From Burkala (2001).

Immunological reagents for the detection of bovine lentiviruses

There are no reports of the successful replication of JDV in cell culture and methods

of preparation of JDV proteins for antigens have been reliant on the extraction of

proteins from infected cattle tissues or the use of recombinant DNA techniques.

The JDV CA with 226 amino acid (aa) residues is a small and highly basic protein

with an estimated Mr of 25.3 kDa that is abundantly expressed, as detected by

SDS-PAGE and western immunoblotting techniques, in infected cells and can be

15

harvested from virus present in the plasma of infected animals during the acute

febrile phase of the disease (Kertayadnya et al., 1993).

The development of antigens able to differentiate BIV and JDV has been

problematic. The expression of antigenic and immunogenic JDV CA and TM

proteins as fusion proteins to the glutathione-s-transferase (GST) in Escherichia coli

has been reported previously (Burkala et al., 1998). Both recombinant proteins

reacted in western blots with JDV and BIV antisera. The Gag protein of JDV has

been cloned (Desport et al., 2005) as a series of overlapping fragments and when

analysed by western immunoblotting using JDV and BIV hyperimmune sera, the MA

and CA were recognised by both sera and the NC did not react with either of the

sera. This analysis suggested that the N-terminal domain of Gag might contain

more antigenic epitopes than the C-terminal domain.

Monoclonal antibodies (MAbs) against JDV have been produced by immunisation of

mice with whole virus (Kertayadnya et al., 1993) and with recombinant JDV CA

(Desport et al., 2005). Characterisation of these MAbs by determining their reactivity

with different truncated Gag proteins of JDV by western immunoblotting (Desport et

al., 2005) demonstrated that the antibodies produced from recombinant CA reacted

with different epitopes compared with those produced by immunisation of mice with

the whole virus. A BIV MAb that recognised epitopes specific to BIV was generated

(Zheng et al., 2001). This MAb, designated 10H1, was produced using a

recombinant fusion protein containing the CA of BIV. Based on the immunoreactivity

of the BIV CA, 3 domains of antigenic importance were identified on the N-terminus

of the protein and these domains were thought to be BIV-specific (Lu et al., 2002a,

Zheng et al., 2001). Unfortunately, this region was recognised by JDV-positive sera

(Desport et al., 2005).

Biological and physiochemical properties of JDV

The buoyant density of JDV purified from the plasma of infected animas was 1.15

g/ml in sucrose gradients (Kertayadnya et al., 1993); the diameter of particles at this

density in the gradients was determined by electron microscopy to be between 96

and 124 nm (Kertayadnya et al., 1993). The virus in plasma derived from infected

animals rapidly decline in infectivity at 4oC but was stable at –70oC (Kertayadnya et

al., 1993).

16

Geographic distribution of Jembrana disease

The population of Bali cattle represents 19% of the total cattle population of

Indonesia (Talib et al., 2002). The total population of Bali cattle in 2000 in the 5

major regions has been estimated as 718,000 in South Sulawesi, 443,000 in Nusa

Tenggara Timur, 377,000 in Nusa Tenggara Barat, 529,000 in Bali and 255,000 in

Lampung (Talib et al., 2002). There are other provinces with growing population

such as in Southeast Sulawesi and East Kalimantan (Talib et al., 2002). The unique

susceptibility of Bali cattle to both sheep-associated malignant catarrhal fever and

also to Jembrana disease is of concern (Talib et al., 2002).

Although precise reporting of cases of Jembrana disease is not undertaken in

Indonesia, it was estimated that there were about 2,000 cases of Jembrana disease

in Indonesia between 1989 and 1992 (Soeharsono, 1997). However, due to the

difficulty of recognising and diagnosing the disease even by trained staff, and as

milder non-fatal cases are unlikely to be reported, this is likely to be a gross

underestimate. It is only when there are high case fatality rates in association with

outbreaks that collection of data is likely. In 2000, 168 cases were reported, 331

cases were reported in 2002 (Peternakan, 2002) and 116 cases were reported in 3

provinces in 2003: 95 in Bengkulu, 2 in Lampung and 19 in South Sumatra. The

most recent cases have been identified in the Long Ikis district of East Kalimantan

(Hartaningsih et al., 2005) (Figure 2.3).

17

Figure 2.3. Distribution of Jembrana disease in Indonesia. The areas where Jembrana disease is endemic include the areas denoted by flags: 1, Bali; 2, Banyuwangi; 3, Lampung; 4, Bengkulu; 5, West Sumatra; 6, South Kalimantan; 7, East Kalimantan. The Data sourced from Soeharsono (1997) and Hartaningsih (2005).

Transmission of Jembrana disease

The mode of transmission of Jembrana disease under field conditions is not known

but certain assumptions have been made based on the level of virus in blood at

various stages of the disease process. During the acute disease the titre of

infectious virus in peripheral blood is about 108 per mL, and virus can also be

detected in secretions. During the acute disease, transmission probably occurs by

2 methods: through direct transmission of virus in secretions between cattle in close

contact, and through mechanical transmission of virus in the blood by

haematophagous insects (Soeharsono et al., 1995b). However, mechanical

transmission of JDV by arthropods, as seems likely, has not been responsible for

extensive spread of Jembrana disease from endemic to adjacent areas. For

example, the disease has not spread from Bali island to the adjacent islands of

Nusa Penida and Lombok since the disease initially occurred in Bali in 1964

(Wilcox, 1997). Recovered Bali cattle are persistently viraemic but the titre of virus

in blood by 60 days after recovery from the acute disease was only about 10

infectious doses per mL, virus could not be detected in secretions during this phase,

and mechanical transmission of these low levels of virus by haematophagous

7

6

5

4

3 2 1

18

insects was considered unlikely (Soeharsono et al., 1995b). Persistent infections in

recovered animals are a potential source of infection but how the virus is transmitted

from these animals is unknown.

It has been revealed through epidemiological studies that the risk of transfer was

intimately related to farming practices, and that an important risk factor was from

contact transmission associated with JDV present in secretions, including saliva and

possibly urine, and in lactating animals also in milk, during the acute febrile phase of

the disease (Soeharsono et al., 1995b).

Diagnosis of JDV infection

The diagnosis of Jembrana disease has traditionally been based on clinical signs

and the presence of characteristic pathological lesions in those animals that died,

although diagnosis was difficult and diagnosis of the disease by these methods was

often made only reluctantly. Hence the disease in East Java was initially referred to

as Banuwangi disease, and in Lampung province in Sumatra it was referred to as

Lampung disease (Soeharsono & Temadja, 1997). The development of an enzyme-

linked immunosorbent assay (ELISA) and western immunoblotting assays for the

detection of antibody to JDV in infected cattle (Hartaningsih et al., 1994) using a

whole viral antigen prepared from the plasma of infected cattle enabled the

distribution of the virus within Indonesia to be determined and consequent diagnosis

of the disease in infected areas made with greater confidence. The use of a

recombinant JDV CA antigen (Burkala et al., 1998) removed reliance on the use of

whole virus antigens prepared from infected cattle. ELISA and western

immunoblotting assays demonstrated that development of antibody against JDV in

recently infected cattle was delayed (Hartaningsih et al., 1994, Wareing et al.,

1999).

A major problem with the JDV serological assays that have been developed is that

they cannot distinguish between antibody to JDV and BIV (Desport et al., 2005).

There is a report that it is possible to differentiate antibody to BIV and JDV (Barboni

et al., 2001) but attempts to confirm this were unsuccessful (Desport et al., 2005). A

further problem, investigated in Chapter 6 (this thesis), is that the size and other

characteristics of the glycosylated envelope proteins have not been determined and

these glycoproteins have not been identifiable in western immunoblots

(Hartaningsih et al., 1994).

19

In situ hybridisation (ISH) is commonly used to detect the presence of virus in

tissues. Chadwick et al. (1998) used ISH with a digoxigenin (DIG)-labelled riboprobe

to detect viral RNA of JDV in formalin-fixed paraffin-embedded tissue sections,

concluding that JDV-infected cells were present in many tissues including spleen,

lymph nodes, lungs, bone marrow, liver and kidney.

The use of a quantitative PCR for JDV RNA detection in plasma samples has also

been reported (Stewart et al., 2005). The assay had a detection limit of 4.2 x 104

JDV genome copies per mL of plasma in experimentally infected cattle.

A monoclonal antibody developed against the CA of JDV (Kertayadnya et al., 1993)

has been used for the development of specific immunoperoxidase assays on frozen

and formaldehyde-fixed tissues.

2.2.2 Bovine immunodeficiency virus

BIV was first isolated in cell cultures from a dairy cow in Louisiana that had

lymphocytosis, lymphadenopathy, neuropathy, and progressive emaciation (Van der

Maaten et al., 1972). Based on serological evidence, the virus has since been

reported from cattle in several countries (Amborski et al., 1989, Forman et al., 1992,

McNab et al., 1994). Although the virus in these other countries appears to be non-

pathogenic, the genetic relationship between the original BIV isolate and the virus in

these other countries has not been determined.

BIV resembles HIV and other lentiviruses in its structural, genetic, antigenic and

biological properties (Gonda et al., 1987). The mature virions of BIV are bar-shaped

and 120-130 mm in diameter. The BIV genome contains the obligatory retrovirus

structural genes in the order gag, pol and env, flanked on the 5’ and 3’ ends by a

LTR. The core protein of the virus is encoded by gag, which produces a 53 kDa

precursor Gag protein that is further processed into MA (p17), CA (p26) and NC

(p15) (Battles et al., 1992, Rasmussen et al., 1990) and 3 small proteins, p2L, p3

and p2 (Tobin et al., 1994) in the mature virus (Gonda et al., 1994).

The BIV complete nucleotide sequence consists of 8,482 nucleotides (Garvey et al.,

1990) while JDV has been reported to contain 7,732 nucleotides (Chadwick et al.,

1995b). Apart from the nucleotide number, the difference between the 2 bovine

lentiviruses consisted of small deletions and insertions located throughout the

genomes (Chadwick et al., 1995a).

20

2.3 Potential Jembrana disease vaccines

Effective vaccines against a multitude of viral diseases have been produced by a

range of methods including whole, live attenuated or inactivated pathogens,

although the development of effective vaccines against lentiviruses has been

difficult. Control of Jembrana disease within Indonesia will most likely require the

development of not only efficacious vaccines but also vaccines of low cost.

Attenuated viral vaccines are effective in stimulating both humoral and cellular

immune responses (Young & Ross, 2003). They are usually attenuated in their

pathogenicity so that they still have ability to replicate but without causing overt

disease. Disadvantages of this type of vaccine are that they can potentially revert to

a virulent strain causing disease and they can be transmitted between individuals.

Modification of the antigenic properties of live attenuated FIV vaccines improved the

protection against FIV infection (Broche-Pierre et al., 2005). The close antigenic

relationship between the non-pathogenic BIV and the pathogenic JDV (Desport et

al., 2005) suggests that there may be some competitive interaction or cross-

protective immunity between these 2 viruses and prior infection of cattle with BIV

might protect against subsequent JDV infection.

Inactivated vaccines are safer than live attenuated vaccines because they cannot

replicate in the host, although they are frequently less effective in inducing

protective immunity (Chalmers, 2006, Ellis, 1999). Recently, a vaccine composed of

inactivated FIV was used and induced high titres of antibody to the Env proteins

(Hosie et al., 2005). A whole virus tissue-derived inactivated JDV vaccine has been

reported to induce a protective immunity against subsequent challenge with JDV but

this vaccine has several potential disadvantages including high cost that it was

inactivated with detergent and might therefore be contaminated with adventitious

infectious agents, and it was not amenable to commercial production methods

(Hartaningsih et al., 2001).

During the last decade, recombinant subunit vaccines have emerged as a promising

vaccine technology. A subunit vaccine can be produced in the form of synthetic

peptides (Audran et al., 2005, Lopez et al., 2001), recombinant proteins or gene

fragments (DNA or RNA) encoding the protein immunogens (Wang et al., 2006). By

using a small and defined part of a pathogen and producing that subunit in a non-

pathogenic host, the safety of vaccines will increase.

A potential target for a JDV vaccine is the envelope glycoproteins, and in a

subsequent section of this review the literature examining methods by which such

21

glycosylated proteins might be produced using recombinant DNA technology is

reviewed.

2.3.1 Recombinant protein vaccines

The first recombinant subunit vaccine to be produced was licensed in 1986 when

the Hepatitis B virus surface antigen (HbsAg) was successfully expressed in yeast

and this then replaced the plasma-derived hepatitis B vaccine used previously

(Valenzuela et al., 1982). This vaccine was initially tested and produced protective

antibodies in vaccinated chimpanzees (McAleer et al., 1984).

Recombinant DNA technology for the production of proteins potentially enables a

large amount of high value protein, which is often in limited supply due to its low

natural availability, to be produced. Several host systems are available for

production of recombinant proteins: eukaryotic cells such as yeasts, filamentous

fungi, insect cells, plants, mammalian cells and prokaryotes including bacteria such

as Escherichia coli, Bacillus and Staphylococcus sp. Each host system has its

advantages and disadvantages. The amount and quality of the produced

recombinant proteins are influenced by factors such as gene copy number,

transcription and translation efficiency, mRNA stability, stability and solubility of the

proteins as well as post-translational modification (Liljeqvist & Stahl, 1999).

The choice of expression system to be used will often depend on the characteristics

of the protein required. The heterologous host used may affect the immunogenicity

and protective efficacy of the protein. In practice, the choice is a combination of the

ease of growing the host, immunogenic and antigenic characteristics of the protein,

and yield and ease of purification of the protein. Several issues such as the gene

construct, solubility of the protein expressed and the nature of the fusion tag that is

often incorporated into the design of the plasmid, all have to be considered

(Hockney, 1994).

Escherichia coli has long been the primary prokaryotic host for heterologous protein

expression, and there is a lot of information available about the use of this

bacterium for the production of many different proteins. It has been successfully

utilised to produce many functional human proteins such as human growth hormone

(Singh & Panda, 2005), proinsulin (Winter et al., 2001), interferon-gamma

(Khalilzadeh et al., 2003, Khalilzadeh et al., 2004) and antibody fragments (Santala

& Lamminmaki, 2004). The advantages of E. coli include its relatively rapid growth,

its utilisation of inexpensive cultural techniques, its ease of transformation and its

22

ease of maintenance. Nevertheless, the use of E. coli also has major

disadvantages: it is unable to perform post-translational modifications such as

glycosylation, phosphorylation and disulfide bond formation, modifications which

occur in eukaryotic cells; the expression and accumulation of a recombinant protein

in E. coli frequently causes the formation of insoluble protein aggregates (Clark,

2001) and this raises issues concerning methods of subsequent solubilisation of the

expressed protein.

Expression in yeast such as Saccharomyces cerevisiae or Pichia pastoris offers

advantages but also disadvantages in comparison to bacterial systems. Many

recombinant proteins have been produced on a large scale using S. cerevisiae,

including human serum albumin (Kang et al., 2000, Okabayashi et al., 1991), the

HbcAg (Chen et al., 2004, Yoshida et al., 1991), insulin and hirudin (Mendoza-Vega

et al., 1994, Vai et al., 2000). Like E. coli, these unicellular eukaryotes grow rapidly

in relatively inexpensive and simple media and they are easy to transform and

maintain. Unlike E. coli, yeasts are eukaryotic and therefore express and process

proteins in a similar way to higher eukaryotes (Cereghino et al., 2002, Cereghino &

Cregg, 1999). The secretory pathway of yeasts closely resembles that of the

mammalian cells, thus they are capable of many posttranslational modifications,

although they are not capable of complex modifications such as prolylhydroxylation

and amidation, and the glycosylation of proteins in yeast can differ from that of

higher eukaryotes (Sudbery, 1996).

The baculovirus expression system is commonly used to express heterologous

proteins in insect cells (Kost & Condreay, 1999). Since insect cells are eukaryotic,

proteins expressed will be post-translationally modified in a manner similar to that of

mammalian cells (Kost & Condreay, 2002, Miller, 1993). Insect cells can be grown

as suspension cultures, which enable the use of large scale bioreactors for easier

production scale-up. Another significant advantage of insect cells is that unlike

mammalian cells, they can be grown in medium that does not need to be

maintained in CO2 incubators. The most commonly used baculovirus system utilised

Autographa californica multiple nuclear polyhedrosis virus (AcMNPV) (Jones &

Morikawa, 1996, Lu et al., 2002b) in a cell line derived from lepidopteran

Spodoptera frugiperda ovarian (Sf9) cells.

Mammalian cell systems may be the only way to produce appropriately processed

and active recombinant proteins. Gene transfer into mammalian cells may be

performed either by infection with a virus carrying the recombinant gene of interest

(Makrides, 1999) or by direct transfer of plasmid DNA (Geisse & Kocher, 1999).

23

Recombinant vaccinia virus vectors have been successfully used for the expression

of recombinant genes (Moss, 1996). Since vaccinia is infectious to humans, safety

aspects must be taken into considerations. Problems with vaccinia systems include

their cytopathic nature and dependence on efficient transfection rates (Moss, 1996).

Post-translational modification of viral glycoproteins

Post-translational modification is a common phenomenon in eukaryotic cells that is

associated with the chemical modification of one or more aa in a protein chain.

There are a number of modifications that may occur at the post-translational stage.

Glycosylation is the most necessary of these modifications as it is important in

secretion, antigenicity and clearance of glycoproteins (Jenkins et al., 1996).

Glycosylation during post-translational processing requires the addition of

carbohydrate structures, forming glycoproteins. The carbohydrate moieties of

glycoproteins, called oligosaccharides or glycans, are composed of individual sugar

residues or monosaccharides (Dell & Morris, 2001). Protein glycosylation is

important for many cellular processes including cell interactions, protein interactions

and protein folding. It is especially vital for the proper positioning and function of

surface receptors; glycosylation of the envelope proteins was shown to be a key

factor in the ability of HIV to evade recognition by the immune system (Rudd et al.,

2001). The presence of N-linked carbohydrates on the FIV Env is important for the

interaction of the virus and receptor (Willett et al., 2008). This is similar with both

HIV and SIV. Mutations in the variable V1/V2 region of HIV SU affect the interaction

between HIV SU and its receptor and co-receptor and alter the antigenicity of the

envelope glycoprotein (Kolchinsky et al., 2001, Srivastava et al., 2003).

There are 2 main classes of glycoproteins, N-linked and O-linked, based on the site

of attachment of the carbohydrate to the polypeptide chain (Medzihradszky, 2005,

Peter-Katalinic, 2005). N-linked glycans are attached to the protein only at the aa

sequences NXS or NXT and are covalently linked to the asparagine (N) residue. In

contrast, O-linked glycans are linked to any serine (S) or threonine (T) in the

polypeptide chain. The role of N-linked and O-linked carbohydrates in the function of

the viral envelope glycoproteins is different but poorly understood (Medzihradszky,

2005).

The glycans are monosaccharide residues and while there are approximately 20

monosaccharide residues (Kobata, 1992), 5 types are common (Table 2.4). Glycans

consist of one or more monosaccharide residues bonded by glycosidic bonds to

24

form polysaccharide chains. The glycan structure can be linear or branched

depending on the synthesis pathways (Marquardt & Denecke, 2003), making them

vary in terms of size, structure and composition. The glycans found on mammalian

N-linked glycoproteins have a common core composition of Glc3Man9GlcNAc2 (Dell

& Morris, 2001).

Table 2.4. The monosaccharides and additional residues found in glycoproteins. Data sourced from Kobata (1992).

Monosaccharide Monoisotopic mass (Da)

Hexose (Hex) Glucose (Glc), Galactose (Gal), Mannose (Man)

162.0528

N-acetylhexosamine (HexNAc)

N-acetylgalactosamine (GalNAc), N-acetylglucosamine (GlcNAc)

203.0794

Deoxyhexose (DeoxyHex)

Fucose (Fuc), Rhamnose (Rha)

146.0579

Pentose (Pent) Arabinose (Ara), Xylose (Xyl) 132.0423

Hexuronic acid (HexA) Glucuronic acid (GlcA) 176.0321

Acetyl (Ac) 42.0106

N-glycolylneuraminic acid (NeuGc)

307.0903

N-acetylneuraminic acid (NeuAc)

291.0954

Phosphate 79.9663

Sulfate 79.9568

2-Keto-3-deoxynonulosonic acid (KDN)

250.0689

Methyl (Me) 14.0157

Depending on the nature of the oligosaccharide chain, the glycan can be classified

as oligomannose, complex or hybrid. Oligomannose glycans contain only mannose

residues, whereas complex glycans have varied composition and a variable number

of antennae stemming from the core. Hybrid types have the characteristic of both

complex and oligomannose glycans (Dell & Morris, 2001). The structures of O-

linked glycans are less defined than that of the N-linked glycans. O-linked glycans

25

are composed of 7 different cores, 4 of them are found in mammalian glycoproteins,

with varied structures and compositions (Dell & Morris, 2001).

The analysis of protein glycosylation remains difficult due to the complexity of the

process and the heterogeneity of the structures that are formed. This heterogeneity

arises from the variation in the use of the glycosylation sites on the protein and the

variations in the monosaccharides used. The complexity is increased further as

different cells, tissues, organs and organisms exhibit different glycosylation patterns

(Rudd et al., 2001). Therefore, analysis of glycoproteins needs multiple analytical

methods and requires several analytical steps and a large amount of sample that

can make it a very expensive process.

There are a few publicly available glycoprotein analysis tools. GlycoSuiteDB is one

of the few glycoprotein databases available on the internet and it has only been

available since 2001 (Cooper et al., 2001b). It is a database that collates information

on glycoproteins from the scientific literature only, and consequently there are only a

small number of entries (Cooper et al., 2003).

Another publicly available database is O-Glycbase maintained by the Centre for

Biological Sequencing from the Technical University of Denmark. It is a database of

O-linked glycoproteins only (Gupta et al., 1999). O-Glycbase is coordinated with

NetOGlyc, a tool for predicting O-linked glycosylation sites (Julenius et al., 2005).

Both GlycoSuiteDB and O-Glycbase are extensively cross-linked to various

nucleotide and protein databases.

GlycoMod is another tool that predicts possible oligosaccharide structures from their

experimentally determined masses (Cooper et al., 2001a).

The identification and characterisation of post-translational modifications has

become achievable with improvements in mass spectrometry (MS). MS was

formerly useful in organic chemistry for the analysis of low molecular weight

molecules but with new methods of ionization of proteins and peptides, MS now

permits analysis of proteins (Domon & Aebersold, 2006), high molecular weight

carbohydrates (Mutenda & Matthiesen, 2006), and nucleotides (Banoub et al., 2005,

Miketova & Schram, 1997) on a routine basis. MS has become the most powerful

technique to determine the mass of a biomolecules and has several advantages

over techniques like gel filtration or SDS-PAGE, principallly the high accuracy, high

sensitivity, and speed (Domon & Aebersold, 2006). The major uses of MS for

protein analysis were reviewed by (Mann & Pandey, 2001).

26

Two desorption techniques, fast atom bombardment (FAB) (Dell et al., 1981) and

plasma desorption (PD) (Maugh, 1985) initiated the development of new strategies

for protein characterization by mass spectrometry. Although these two methods

were successful, the breakthrough came with the development of matrix-assisted

laser desorption/ionization (MALDI) (Hillenkamp et al., 1991) and electrospray

ionisation (ESI) (Andersen et al., 1996, Gaskell, 1997, Ho et al., 2003). The

techniques are fast, simple, and accurate, and have made mass spectometric

analysis of biomolecules a routine analytical activity. MALDI has been coupled to

many different mass analysers but is mostly coupled to a time-of-flight (TOF)

analyser.

For the localization of modifications and mutations or for identification by peptide

mass fingerprinting, the protein needs to be fragmented into smaller peptides. The

use of in-gel digestion (Huynh et al., 2009, Jimenez et al., 2001) is a good

alternative for the generation of peptide fragments of a membrane protein. An in-gel

approach has several advantages. First, only few materials are needed. Second,

the purity of sample is not critically important because SDS-PAGE separates the

protein of interest from contaminants. Third, the membrane protein is partially

unfolded in SDS, providing better accessibility for the protease, whereas the

denaturant is removed by washing the excised gel. Several examples have been

published in which the identification of a modification was successful using a

specific isolation procedure without having a complete peptide map (Lennon &

Walsh, 1999, Qin & Chait, 1997, Tsur et al., 2005).

Peptide mass fingerprinting is an identification method first described by James and

Yates (James et al., 1993, Yates et al., 1993). The peptide mass fingerprinting

method is based on the idea to divide a whole protein into smaller fragments to

measure the mass of each fragment and to then use the resulting list of masses, ie.

a fingerprint, to identify the protein. The analysed protein is then compared with a

database of proteins, and for characterisation, ie. to detect which variant of an

identified protein was analysed or if the protein was modified by post-translational

modification (Cottrell, 1994, Wise et al., 1997). The peptide mass fingerprinting

method is also widely used to identify and characterise unknown samples (Cottrell,

1994, Wilkins et al., 1997).

27

Selection of appropriate gene constructs for expression of proteins

Selecting a minimal region of the protein required to elicit a strong immune

response could reduce the length of the gene to be inserted into the expression

vector and in some cases the gene needs to be truncated for the purpose of

facilitating expression (Johne et al., 2004). Promoter sequences may also be altered

to make production more efficient, especially in the case of toxic proteins (Chevalet

et al., 2000). The secretion of expressed protein can be intracellular or directed into

the medium or periplasm; secretion of the protein into the growth medium may

protect the protein from cytoplasmic proteases and it can simplify the purification

process. Extracellular production is often preferred to express proteins containing

transmembrane regions (Sorensen & Mortensen, 2005) or that have a tendency to

aggregate (Sorensen et al., 2004) as intracellular expression may lead to the

formation of inclusion bodies, which requires solubilisation and refolding to obtain a

soluble and active form.

Formation of recombinant proteins within inclusion bodies

Production of recombinant proteins by bacterial cells is usually the most convenient

and cost-effective method of recombinant protein production and E. coli has

become the most extensively used bacterial host. It is a genetically and

physiologically well-characterised organism that grows rapidly and requires simple

medium but its use is also affected by 2 problems: overproduction of recombinant

proteins leading to the formation of inclusion bodies, and protein degradation

(Baneyx & Mujacic, 2004, Cabrita & Bottomley, 2004, Markossian & Kurganov,

2004). Foreign proteins expressed in E. coli can contain regions that are recognised

by specific host proteases, leading to cleavage and sometimes to subsequent

degradation (Baneyx & Georgiou, 1990, Rozkov & Enfors, 2004). Inclusion bodies

are very dense particulate amorphous structures (Bowden et al., 1991) that almost

exclusively contain over-expressed proteins (Cabanne et al., 2005, Carrio et al.,

1998, Razeghifard, 2004, Rinas & Bailey, 1992, Sakono et al., 2004, Valax &

Georgiou, 1993). Formation of inclusion bodies can be beneficial if the protein

product is susceptible to host proteases or is toxic to the cells in its active form; in

this case, inclusion bodies are advantageous and strategies for increasing inclusion

body formation have been developed. Although recombinant proteins with

secondary structure and native-like conformations have been found in inclusion

28

bodies (Khan et al., 1998, Oberg et al., 1994) the proteins aggregated in inclusion

bodies are normally inactive, unfolded or folded incorrectly and need refolding in the

functional active format (Buchner & Rudolph, 1991, Misawa & Kumagai, 1999). The

presence of high concentrations of the protein in inclusion bodies, however, up to

95% of the mass of the inclusion body, can simplify downstream processing (Han et

al., 2004, Lilie et al., 1998). Inclusion bodies can be easily removed from host cell

proteins and debris by low speed or gradient centrifugation (Haelewyn & De Ley,

1995, Taylor et al., 1986) or an expanded bed adsorption (Cabanne et al., 2005).

Inclusion bodies form not only with expression of heterologous recombinant proteins

but also from homologous proteins (Rudolph & Lilie, 1996), indicating that the

formation is not specific for “foreign proteins“. The formation of Inclusion bodies is

also not specific for E. coli since they have been found also in Bacillus subtilis

(Wang et al., 1989), in yeast such as Saccharomyces cerevisiae (Binder et al.,

1991) and in mammalian cells (Broido et al., 1991). They can also form from

proteins like ß-lactamase secreted into the periplasm (Chalmers et al., 1990).

Reasons given for the formation of inclusion bodies include the heterologous nature

of the protein (especially for proteins that require post-translational modifications),

high protein synthesis rates, proteins with high hydrophobicity that aggregate

intermolecularly as a result of non-covalent association, and a lack of available

chaperones (Mukhopadhyay, 1997). Point mutations which change the

hydrophobicity of a protein can alter the stability and solubility of the protein (Luck et

al., 1992) and change considerably the amount of protein deposited in inclusion

bodies (Wetzel et al., 1991).

To produce a product from inclusion bodies that is similar to the native form, a

renaturation strategy is required (Clark, 2001, Rudolph & Lilie, 1996). General

strategies for recovering native proteins from inclusion bodies normally involve 3

steps: (1) isolation of Inclusion bodies including washing several times to remove

undesired co-precipitation proteins, (2) solubilisation of inclusion bodies in a strong

denaturant such as urea or guanidinium chloride to break intermolecular

interactions, and (3) renaturation of solubilised proteins by dialysis or dilution to

remove the denaturant. For renaturation of proteins containing disulfide bonds, the

supplementation of redox systems to the renaturation buffer is needed (Creighton,

1986). These redox systems provide the appropriate redox potential to allow

formation and reshuffling of disulfide bonds.

This strategy to prepare soluble proteins from inclusion bodies is possible only for

small proteins. Large multi-domain proteins normally fail to fold properly, often

29

leading to kinetically trapped intermediates that aggregate, even in diluted solutions

and at low temperature (Jaenicke & Bohm, 1998). Moreover, the folding process in

vitro can take a long time, up to several days. Due to competition between

aggregation and folding in vitro (Kiefhaber et al., 1991), the protein concentrations

must be kept low. Thus, to get a given amount of soluble protein, large volumes of

buffers are needed. The yield of protein refolding can be improved by coupling

denatured proteins to a matrix (Stempfer et al., 1996), adding low molecular weight

additives (Rudolph & Lilie, 1996), genetically engineering hydrophobic patches

(Wetzel, 1994), shifting the temperature rapidly (Betts & King, 1998), or adding

molecular chaperones (reviewed by (Thomas & Baneyx, 1997)). However, the

yields are still likely to be modest, especially for large multi-domain proteins.

Moreover, refolding in vitro is an expensive and time consuming process that

sometimes is not easy to scale up. To simplify the downstream processing,

maximisation of the yield of soluble active product during production in vivo is an

attractive alternative.

Tag protein fusions

The tags on most fusion proteins are commonly used to optimise the production and

purification procedures of the target protein but they also can be used for detection

and immobilisation purposes (Nilsson et al., 1997). The fusion tag can improve the

solubility (Nallamsetty & Waugh, 2006) or proteolytic stability (Murby et al., 1991,

Murby et al., 1996) of the overall protein. There are many well-defined affinity tags

that have been described such as glutathione-S-transferase (GST), polyhistidine

(His), maltose-binding protein (MBP) and these have been reviewed by (Terpe,

2003). No single affinity tag is ideal for all expression and purification systems.

2.4 Production of monoclonal and recombinant antibody

The use of virus-specific antibodies, and in particular the use of MAb, has been of

considerable value in the development of various assay systems for the detection of

microbial antigens and diagnosis of disease.

Monoclonal antibodies

Antibodies are composed of immunoglobulins that are produced by plasma cells.

There are 5 isotypes of immunoglobulins, namely IgG, IgM, IgA, IgD and IgE. Once

30

a B-lymphocyte has been stimulated by foreign antigen to differentiate into antibody-

producing plasma cells, they may produce only one single type of antibody

molecule, binding to a particular site (epitope) on the antigen. The clonal selection

of individual antibody producing B cells and continued propagation of the cloned

cells is the basis of producing MAb.

Attempts to culture single antibody-producing cells in vitro were met with limited

success until Kohler and Milstein (1975) developed a technique for the reliable long-

term production of cloned antibody producing lymphocytes. Their methods enabled

the production of individual antibodies of invariant specificity and selectivity, and the

immortalisation of the antibody-producing cells, ensuring a virtually infinite supply

(Kohler & Milstein, 1975). The procedure for the production of MAb involves

immunisation of a mouse or a rat, fusion of the plasma cells from the immunised

mouse or rat with myeloma cells to create hybridomas, then cloning the fused cells

and harvesting the antibody product.

Structure and function of antibody molecules

A schema of a complete antibody molecule is illustrated in Figure 2.4. The basic

dimeric antibody structure comprises 2 heavy chains (VH) of about 440 aa and 2

light chains (VL) of about 220 aa residues. These chains fold into 3 domains. Two of

the domains (Fab domains) are identical and form the arms of the Y-shaped

molecule. One light chain associates with the amino-terminal region of one heavy

chain to form an antibody-binding site. The third domain (Fc) forming the base of the

Y shape is folded together by the carboxy-terminal regions of the 2 heavy chains

and is responsible for aspects of the immune response. The 4 polypeptide chains

are held together by disulfide (S-S) bridges and non covalent bonds. The light chain

can be divided into 2 regions, variable (V) and constant (C) region. The heavy chain

contains one variable and 3 constant regions. Within the variable regions, 4

framework regions (F) and 3 hypervariable regions (CDR) can be discriminated.

These CDRs form the majority of contact residues for the binding of the antibody to

the antigen (Padlan, 1994).

31

Figure 2.4. Schematic drawings of the organisation of a natural antibody and derived recombinant fragment scFv and Fab. The structure of antibody is composed of the 2 variable regions of heavy and light chain (VH and VL), which include CDR. In recombinant antibodies; a peptide linker connects the 2 antigen binding regions of the light and the heavy chain.

The repertoire of antibodies is created during B-cell development by DNA

rearrangement and combinatorial assembly of different genetic segments. The V-,

D- and J-segments form the heavy chain gene and V- and J-segments form the light

chain gene (Winter, 1998, Winter et al., 1994).

Recombinant antibodies Current molecular techniques enable the construction of recombinant antibody

fragments that have antigen-binding properties. The general strategy taken is to

isolate the mRNA from mouse splenocytes or MAb-producing hybridoma cells and

to reverse transcribe the immunoglobulin encoding mRNA with a single primer,

which binds either the heavy or light chain antibody gene near the beginning of the

constant domain.

Reverse transcription of the mRNA results in the production of immunoglobulin

variable region cDNA. The cDNA is then amplified by PCR with the original 3'

constant region primers and the 5' leader signal primers, binding upstream from the

variable region. The immunoglobulin heavy or light chain cDNA amplified by PCR

can be cloned into a plasmid or phagemid vector. Once the immunoglobulin genes

Fab

COOH COOH

NH2 NH2

V L

V H

CH1

CH3

CH2

CDR3 CDR2

CDR1

Fab

CH1

Fc

S-S

ScFv

NH2 NH2

COOH COOH

V H V L

32

are cloned, they can be sequenced and produced in large scale by simple E. coli

fermentation methods (Buchner & Rudolph, 1991, Su et al., 2003).

Single chain variable fragment (scFv) antibody ScFv is a product resulting from the development of the current biotechnology and

antibody engineering. The 2 antigen binding variable regions of the heavy (VH) and

light chain (VL) are artificially connected by a linker peptide, designated as single-

chain variable fragment or single chain antibody (Bird et al., 1988, Orlandi et al.,

1989). This resulting scFv facilitates the equal expression of both variable fragments

in heterologous micro-organisms, mammalian cells and plants (Schirrmann et al.,

2008).

The antigen-binding site is made up of the variable domains of light and heavy

chains of a MAb. The smallest portion containing an antigen binding site is the

variable fragment (Fv) of an antibody. The Fv fragment has the full intrinsic antigen

binding affinity of one binding site similar to the binding pattern of the whole parental

antibody (Takemura et al., 2000). The relative affinity of VH and VL fragments for

each other depends on the particular sequence of the antibody. Low affinity may

result in dissociation of the Fv fragment into its components (Horne et al., 1982).

To stabilise the association of the recombinant Fv fragments, they are joined with a

short peptide linker and expressed as a single polypeptide chain; linker peptides, of

12-25 aa do not normally disturb the proper folding of the VH and VL domains (Bird

et al., 1988, Huston et al., 1988). The most frequently used linker for scFv

antibodies is (Gly4Ser)3 (a 15 aa peptide of 12 glycines and 3 serines) that bridges

the ~4.5 nm gap between the C-terminus of one domain and the N-terminus of the

other and has a flexible structure with enhanced mobility (Freund et al., 1993,

Huston et al., 1988). This construction facilitates chain pairing and minimises

refolding and aggregation encountered when the 2 chains are expressed

individually. Comparison of the unlinked Fv fragment of the antibody McPC603 with

the corresponding scFv containing a linker (Gly3Ser)4 (VH-linker-VL constructs) has

shown no perturbation of the folding of the variable domains by the linker (Freund et

al., 1993).

There are different forms of scFv; a soluble scFv antibody is the most popular form

as it shares all the advantages of the MAb (Blazek et al., 2004). The scFv genes

can also be fused to gene3 protein (g3p) of the phage to form recombinant phage

display scFv antibody and leads to the expression of scFv on the surface of the

33

phage. An advantage of this form of scFv is that the supernatant of the phage-

infected bacterial culture can be directly used in ELISA (Xu et al., 2004). This is

useful for screening specific scFv to target antigens from a scFv library through

multiple rounds of selection. However, this kind of scFv may exhibit some cross-

reactivity (McElhiney et al., 2002).

Dimeric or mini-antibody is another form of scFv. Such molecules preserve the

bivalency of native antibody molecules, which is a very effective means of

increasing the functional affinity (avidity) to the surface of polymeric antigens (Kortt

et al., 2001, Muller et al., 1998). ScFv fragments can be linked by a small modular

dimerisation domain in the form of one or 2 amphipathic helices. These mini-

antibodies assemble in dimeric form in E. coli and the avidity of the best of them is

indistinguishable from a native antibody (Kortt et al., 2001).

Biotinylated antibodies are commonly used reagents in research and molecular

diagnostics. The traditional approach to biotinylate antibodies is to conjugate a

chemically active biotin to scFv antibody. This genetically biotinylated scFv enables

the production of the antibody conjugate in bacteria, which could decrease costs of

ELISA reagents (Grimm et al., 2004, Santala & Lamminmaki, 2004).

The small scFv fragments are considered promising for medical and biological

applications because of superior tissue penetration, absence of side reactions

involving the constant domains, as well as easy engineering of fusion proteins, such

as scFv-coupled toxins, the creation of multivalent or bi-specific proteins (Hudson &

Kortt, 1999, Leath et al., 2004).

Fab antibody fragments The construction of Fab fragments (50 kDa) is similar to that of scFv (30 kDa). The

difference is that both variable and constant regions of light chain and heavy chain

still remain in Fab (Figure 2.4). Therefore, a molecule of recombinant Fab fragments

has the complete antigen binding site and structure of a natural antibody, which may

result in a relatively higher avidity than that of the same scFv. The thermodynamic

stability of the corresponding Fab fragments is also higher than that of scFv

(Rothlisberger et al., 2005). This is due to the 2 interchain disulphide bonds typically

present in the variable domains (VH and VL) and additional disulphide bonds in the

constant domains (CH1 and CL). Although Fab fragments have these advantages,

they are usually produced in the periplasmic space of E. coli at lower functional

yields and do not have the same advantages of small size, especially in multivalent

34

formats (Skerra & Pluckthun, 1988). Production of functional antibody Fab

fragments in an oxidising bacterial cytoplasm has overcome those limitations in

terms of yield, folding and functionality and is economic, rapid and efficient (Venturi

et al., 2002). A number of Fab fragments have been developed and used in

immunotherapy and diagnostics (Zhang et al., 2001) and have found some

applications in clinical toxicology (Flanagan & Jones, 2004).

Generation of scFv antibodies

Established hybridoma clones are a rich source for generating scFvs. The variable

domains of the antibodies expressed by the hybridoma cells can be derived

relatively easily, and cloned into the scFv format. However, a major disadvantage is

that not all scFv derived from existing antibodies are active. Another disadvantage is

the non-human origin of most hybridoma clones, which may render the derived

scFvs immunogenic (Blazek & Celer, 2003, Blazek et al., 2004).

ScFvs can also be selected from large naïve scFv repertoires or smaller

“immunised” repertoires. The techniques and methods for such selections include

phage display that is widely used due to its simplicity and ability to be adapted to

many specific conditions, including selection on whole cells and tissues (Hust &

Dubel, 2005) and the system has been extensively reviewed (Kotz et al., 2004, Pini

& Bracci, 2000), ribosome display that has the capacity to screen libraries of greater

size as well as facilitating diversity and efficient antibody maturation (He et al., 2004,

Lee et al., 2004b), bacterial display (Bessette et al., 2004, Su et al., 2003) and yeast

display (Boder & Wittrup, 2000, Feldhaus & Siegel, 2004, Powers et al., 2001,

Siegel et al., 2004) that have several advantages over the phage system including

use of flow-cytometry and sorting techniques to enable finer affinity discrimination of

selected antibodies. Other techniques are periplasmic expression cytometric sorting

(PECS) (Chen et al., 2001) and potentially some 2-hybrid systems to screen for

intracellular antibodies.

Display on phage particles

The first paper describing phage display (Smith, 1985) described display of a

fragment of the EcoRI enzyme in the central region of pIII, and showed that the

recombinant phage could be enriched in a phage pool by panning the pool on

antibodies recognising EcoRI. The display of an anti-lysozyme scFv on the surface

35

of filamentous phage particles by fusion of the antibody variable genes to one of the

phage coat proteins paved the way for the generation of antibody repertoires on

phage (McCafferty et al., 1990).

Over the years, most of the phage coat proteins have been tested for display of

antibody fragments, with pIII as the most preferred choice (Ding et al., 2005, Gupta

et al., 2001, Han et al., 2003, Yang & Shiuan, 2003). The scFv genes are inserted

directly upstream of the gene for pIII, which results in fusion proteins upon

expression. The phage particles produced from a phage vector are homogeneous in

their content of pIII, whereas the phage particles produced from a phagemid vector

in combination with a helper phage have pIII from both the phagemid (fusion

proteins) and from the helper phage (non-fusion). The phage vectors offer the

advantage that all phage particles display scFvs and therefore enable a more

efficient selection. In contrast, the phage particles produced from phagemid systems

are monovalent. On average the phage particles displaying a fusion protein display

only one, whereas the vast majority do not display any (O'Connell et al., 2002); this

is a clear advantage when high affinity is desired. The other advantage of phagemid

vectors is their relative high transformation efficiency allowing the generation of

libraries with higher diversity and the accurate measurement of the antibody library

size (Hong et al., 2004).

Antibody repertories can be divided into categories based on the source used for

their generation (Hoogenboom & Chames, 2000) (Figure 2.5). Natural antibody

repertoires can be created from B-lymphocyte mRNA obtained from either

immunised or non-immunised donors. The “immune” repertoires (Duggan et al.,

2001, Okamoto et al., 2004) will be biased by the immune response of the donor

and will therefore be enriched for antibodies directed against the antigen used for

immunisation. In addition, several of the antibody genes may contain somatic

hypermutations, and therefore encode high affinity antibodies (Hoet et al., 2005).

However, there are several drawbacks: immunisation may not result in an immune

response, toxic antigens may kill the donor, tolerance may limit or prevent an

immune response toward self-antigens, ethical concerns may hinder active

immunisation of humans, and the applicability range of a repertoire is limited to

selections against the immunising antigen. In contrast, naïve repertoires (reviewed

by (Winter et al., 1994)) can be applied in the selections against a wide range of

targets, including those that cannot be used in immunisations. The caveats are the

diversity requirement and the potential bias by the donors’ immunoglobulin

repertoire.

36

Figure 2.5. The source of antibody genes. The model sourced from Hoogenboom & Chames (2000).

To extend the repertoire diversity, “synthetic” repertoires have been generated in

which parts of the natural antibodies are randomised (Sidhu et al., 2004). The

classic approach is to exchange the CDR3 regions, which are normally created by

the recombination of the V(D)J gene segments, with synthetic oligonucleotides. This

strategy can be further extended to cover the other CDR regions (Fellouse et al.,

2004). The synthetic repertoires are not biased, and can generally be applied in

selections against any antigen, including self-antigens and toxic antigens. The

synthetic repertoires may contain high fractions of non-active or unfolded antibodies

(Rubinstein et al., 2003).

Morphology and life cycle of the filamentous bacteriophage Ff

The filamentous bacteriophage Ff is a rod-shaped particle (Figure 2.6) about 900

nm in length and 6.5 nm in diameter. It can be propagated by infection of E. coli that

contain the F conjugative plasmid. The genome of the Ff bacteriophage is a

covalently closed single-stranded DNA molecule (ssDNA) of about 6,400

nucleotides, encoding 11 proteins (Table 2.5) that can be grouped according to their

37

functions in the phage morphology or to their roles in the phage lifecycle (Russel,

1991, Russel et al., 1997).

Table 2.5. F-specific filamentous phage genes/proteins and properties. Data sourced from Russel et al. (1991).

Gene Protein Size (aa)

Function Location Used for display?

I I 348 Assembly Inner membrane

XI 108 Assembly Inner membrane

II II 409 Replication (nickase) Cytoplasm X 111 Replication Cytoplasm III III 406a Virion component Virion tip

(end) Yes (N-terminus)

IV IV 405a Assembly (exit channel) Outer membrane

V V 87 Replication (ssDNA bp) Cytoplasm VI VI 112 Virion component Virion tip

(end) Yes (C-terminus)

VII VII 33 Virion component Virion tip (start)

Yes (N-terminus)

VIII VIII 50 Virion component Virion filament

Yes (N- and C- termini)

IX IX 32 Virion component Virion tip (start)

Yes (N-terminus)

a: Mature protein without signal sequence.

The phage capsid consists of 5 different proteins of which pVIII (50 aa) is the most

abundant. pIII, pVI, pVII and pIX (406, 112, 33 and 32 aa, respectively) are all

present in a few copies at the ends. The actual wall of the rod-like phage particle is

composed of pVIII (2,700-2,800 copies). All the capsid proteins have hydrophobic

membrane spanning domains and reside in the inner membrane until they are

assembled into the mature phage particle (Willats, 2002) (Figure 2.6).

The infection process is initiated when the phage particle binds to the first of the 2

bacterial receptors needed for infection, the F-pilus, via an interaction with domain

N2 of the pIII (Karlsson et al., 2003) as shown in Figure 2.7. Subsequently, domain

N1 of pIII can interact with the second bacterial receptor, TolA, and complete the

infection (Click & Webster, 1997, Deng & Perham, 2002, Deprez et al., 2002,

Lubkowski et al., 1999, Nilsson et al., 2000).

38

Figure 2.6. Schematic illustration of M13 bacteriophage particle showing the organisation of the 5 coat proteins. The approximate number of copies of each M13 coat protein is indicated. The model was sourced from Willats (2002).

Three proteins encoded by the phage genome are involved in the replication of

phage DNA. pII (410 aa) binds to the double-stranded replicative-form of the phage

genome and introduces a specific cleavage on the (+) strand at the origin of

replication. Upon completion of the rolling circle replication, the ssDNA is

circularised by pII, and a stabilising complex is formed with pV (87 aa). pX (111 aa)

is the product of an internal initiation of gene II at methionine 300, and probably

controls DNA replication (Horiuchi, 1997).

The last 3 proteins encoded by the genome are required for the assembly of the

phage coat. The pI (348 aa), pXI (108 aa) (Haigh & Webster, 1999) and pIV

(405[+21] aa) (Marciano et al., 2001) proteins form a channel in the bacterial

membrane and are required for the phage assembly and release. It has been

speculated that this channel may set the restrictions on maximum size of proteins

that can be displayed on phage (Marvin, 1998, Russel, 1991, Russel et al., 1997,

Stassen et al., 1994).

39

Figure 2.7. Schematic illustration of the bacteriophage infection process via the 2 E. coli receptors needed: Pilus and TolA. Pilus is usually involved in the conjugal transfer of DNA between E. coli. It consists of a protein tube assembled by polymerisation of pilin subunits from the bacterial inner membrane. The pilin subunits are encoded by the traA operon on the F conjugate plasmid (Manchak et al., 2002). Upon interaction with N2 of pIII, pilus retracts and N1 of pIII can interact with the TolA leading to completion of infection. The model was sourced from Karlsson et al. (2003).

40

Chapter 3 Solubilisation and purification of insoluble recombinant JDV-ΔSU present in inclusion bodies

Summary

Methods were developed to express, solubilise and refold a truncated recombinant

surface unit (ΔSU) protein of Jembrana disease virus that was expressed as an

insoluble product in Escherichia coli. The protein was engineered with a glutathione-

s-transferase (GST) tag to facilitate purification and was truncated by deletion of the

N-terminal region of the protein to eliminate the hydrophobic transmembrane

domain and cytoplasmic tail of the full-length protein but this truncation of the

protein failed to increase the solubility of the protein expressed in E. coli and it was

produced as an insoluble product within inclusion bodies. To purify the insoluble

protein present within inclusion bodies, the inclusion bodies were partially purified

from lysed bacterial cells by an initial washing step to separate the inclusions from

cellular lipids and membranes. The effect of different buffers on the solubilisation of

ΔSU inclusion bodies was examined and a denaturing buffer of 2 M Tris-HCl, pH 12,

containing 2 M urea and 10 mM DTT was selected as an optimal solubilising buffer.

The solubilised denatured proteins were then refolded by slowly diluting the protein

solution with 10 volumes of refolding buffer (20 mM Tris pH 8.0 containing 0.1%

CHAPS, 10 mM DTT and 1 mM PMSF). Final purification of the ΔSU was achieved

by affinity column purification of the tagged protein with glutathione-Sepharose 4B

resin (BioRad) and cleavage of the GST tag with PreScission™ protease (GE

Healthcare). The purified solubilised ΔSU protein could be considered for use as a

vaccine protein or antigen for serological tests.

41

Introduction

Recombinant proteins are widely used in medicine for production of vaccines and

therapeutic proteins for human (Green & Gaston, 2006) and animal diseases

(Weerasinghe et al., 2006). They are also used as antigens for immunological tests

(Zhang et al., 2006).

Production of any recombinant protein in a heterologous host cell or organism is a

complex multi-step process and is often problematic. Protein expression levels can

be low or the purified protein may be insoluble or unstable. To generate large

amounts of recombinant proteins, optimal conditions must be determined which

require selection of an appropriate expression vector and expression host, and the

ability of the fusion partner to facilitate purification of the expressed protein.

Glutathione S-transferase (GST) fusion vectors with expression in an Escherichia

coli vector provide high transformation efficiency, simplicity and low cost of

production, and are frequently chosen and used for recombinant protein production

and purification (Terpe, 2006). GST fusion vectors provide GST fusion proteins that

can be purified by binding to immobilised glutathione followed by competitive elution

of the immobilised protein with reduced glutathione (Smith & Johnson, 1988).

However, a significant problem with GST fusion proteins is that they are often

expressed as insoluble proteins that are difficult to purify by standard procedures

(Hollenbach et al., 1999, Iwata et al., 2000, Mercado-Pimentel et al., 2002, Terpe,

2003, Wang et al., 1997).

Recombinant JDV capsid (CA) and transmembrane (TM ) proteins have been

expressed in Escherichia coli as a fusion protein with GST (Burkala et al., 1998).

The soluble fraction of the expressed proteins was purified using affinity

chromatography with reduced glutathione and then used as an antigen to develop

serological tests for JDV (Burkala et al., 1998). The majority of the expressed

protein, however, was present as an insoluble product and yields would have been

markedly enhanced if the insoluble product had been solubilised. An effort to

increase the solubility of the expressed JDV fusion proteins by truncation of the

protein still resulted in a considerable proportion of the expressed protein being

present as an insoluble product within inclusion bodies (Burkala et al., 1998). This

Chapter reports the development of a simple process for solubilisation and

purification of the insoluble JDV recombinant ΔSU protein present in inclusion

bodies that considerably enhanced the yield of soluble protein.

42

Material and methods

Protein expression

The map of the expression cassette for a fusion of the surface unit protein is shown

in Figure 3.1. E. coli DH1 (F-, endA1, hsdR17 (rK-, mK+), supE44, thi-1, λ-, recA1,

gyrA96, relA1) transformed with the pGEX-6P-1 plasmid containing JDV nucleotides

5556 – 6463 (numbering as described by (Chadwick et al., 1995b)) was grown

overnight with shaking 225 rpm in 5 ml of Luria-Bertani (LB) medium supplemented

with 100 µg of ampicillin per ml as a selection agent at 37oC. The overnight culture

was diluted 1:40 in 200 ml of fresh LB-ampicillin medium, grown again at 37oC to an

OD600 of 0.6 – 0.8, when pre-induction samples were taken as controls and protein

production was induced with 0.5 mM isopropyl β-D-thiogalactopyranoside (IPTG) for

4 h at 37oC. During incubation, post-induction samples were taken every 1 h.

Samples were then centrifuged (4,000 g for 10 min at 4oC) and cell pellets were

resuspended in reducing SDS-PAGE loading buffer prior to electrophoresis.

Figure 3.1. Map of vector pGEX-6P-1 used to produce recombinant ΔSU in E coli (from Amersham website http://www4.amershambiosciences.com/pdfs/970004M2-01.pdf).

PreScission Protease SU protein RE GST Ptac Amp

43

After 4 h of induction, the bacteria were harvested by centrifugation (4000 g for 10

min at 4oC) and the bacterial pellets were suspended in 10 ml of ice cold

phosphate-buffered saline (PBS; 2 mM KCl, 140 mM NaCl, 10 mM K2HPO4, 2 mM

KH2PO4, pH 7.4). Lysis of the bacterial cells was achieved by the addition of

lysozyme (1 mg/ml final concentration) and incubation on ice for 15 min with

occasional mixing, followed by freeze-thawing using liquid nitrogen, which resulted

in a dramatic increase in viscosity. The viscosity was then reduced by treatment

with benzonase (Novogen) (10 µg/ml final concentration) in the presence of 10 mM

MgCl2. The soluble and insoluble fractions of the E. coli lysate were separated by

centrifugation (10,000 g for 20 min at 4oC). The supernatant fractions were collected

and stored at –20oC for further analysis and the residual pellets were used for

solubilisation.

Solubilisation of insoluble proteins in inclusion bodies

The pelleted inclusion bodies were washed 3 times with washing buffer (50 mM

Tris-HCl buffer pH 8.0, containing 5 mM EDTA and 1% Triton X-100) by

centrifugation (10,000 g for 15 min) after each wash step. The pelleted material was

then washed once with distilled water to remove salt and detergent and centrifuged

again (10,000 g for 30 min). The pelleted material was then resuspended in 50 mM

Tris-HCl buffer at pH 8.0.

To determine the effect of different solubilisation buffers, 100 µl volumes of

resuspended washed inclusion bodies were placed in microcentrifuge tubes and

centrifuged (4,000 g for 10 min at 4oC). The supernatant was discarded and 1 ml of

the various solubilisation buffers was added to each of the pellets. The suspensions

were vortexed and left for 1 h at room temperature. The solubilisation buffers tested

included 100 mM Tris-HCl buffer with or without additional 2% urea at pH 7 to pH 13

and other solubilisation buffers listed in Table 3.1. The relative soluble protein

concentration of the suspensions was determined by centrifugation (10,000 g for 10

min) of the suspension, followed by filtration of the supernatant through a 0.45 µm

filter and then determination of the relative protein concentrations by measuring the

absorbance of the suspensions at 280 nm (A280), where A280 = 1.00 is assumed to

correspond to 1 mg of protein per ml (Lowry et al., 1951).

44

Table 3.1. Buffers tested for solubilisation of insoluble JDV ΔSU within inclusion bodies.

Buffer

A. 2 M Tris-HCl pH 12

B. 2 M Tris-HCl pH 12, plus 2 M urea and 10 mM DTT

C. 2 M Tris-HCl pH 12, plus 10 mM DTT

D. 50 mM Tris-HCl pH 8.5, plus1% Sarkosyl

E. 50 mM Tris-HCl pH 8.5 plus 8 M urea

F. 50 mM Tris-HCl pH 8.5 plus 6 M guanidine-SCN

G. 50 mM Tris-HCl pH 8.5 plus 1% SDS

H. 100 mM Tris-HCl buffer pH 12.5, plus 2 M urea and 10 mM DTT

Refolding of solubilised protein

The solubilised ΔSU-GST was refolded by diluting the protein solution slowly with

10 volumes of refolding buffer (20 mM Tris pH 8.0 containing 0.1% CHAPS, 10 mM

DTT and 1 mM phenyl methyl sulfonyl fluoride, a serine protease inhibitor [PMSF])

followed by storage overnight at 4oC.

Cleavage of GST-JDV ΔSU

After refolding, the fusion protein was applied to a disposable gravity column (Bio-

Rad) containing 2 ml of glutathione-Sepharose 4B resin (Amersham Biosciences)

equilibrated previously with binding buffer (PBS, pH 7.4) at 4oC. The column matrix

was washed twice with the binding buffer and then that buffer was substituted with

cleavage buffer (50 mM Tris HCl, 100 mM NaCl, 1 mM EDTA, 1 mM DTT, pH 8.0)

to pre-equilibrate the column, altering the buffer composition required for efficient

cleavage by PreScission™ protease (GE Healthcare). PreScission protease™

(2 U/100 μg of ΔSU-GST protein) was diluted in cleavage buffer and injected onto

the column. Following the injection, the column was placed in a closed flow status at

4oC overnight. An elution buffer containing reduced glutathione (50 mM Tris-HCl

and 10 mM reduced glutathione, pH 8.0) was then applied in a one-step gradient

(100%) to elute the GST and PreScission™ protease. A series of 1 ml eluates were

collected and stored on ice, then the proteins were precipitated with acetone and

45

ethanol, and stored at -20oC until subjected to sodium dodecyl sulphate-

polyacrylamide gel electrophoresis (SDS-PAGE).

SDS-polyacrylamide gel electrophoresis

Fractions collected during the expression and purification procedures were

subjected to SDS-PAGE in a Mini Protean III cell (Bio-Rad) for 1 h at 200 V by using

a 4% stacking gel and a 12.5% polyacrylamide separation gel as described by

Laemmli (1970). The samples were heated at 95oC for 10 min before

electrophoresis. After electrophoresis, gels were stained overnight on a rocking

platform at room temperature in a solution containing 0.1% (w/v) Coomassie brilliant

blue R-250 (Bio-Rad) in 40% methanol, 50% water and 10% acetic acid. Excess

dye was removed by multiple washing in 40% methanol, 50% water and 10% acetic

acid.

Results

Expression and solubilisation of inclusion body protein

The recombinant fusion protein ΔSU-GST of ~65 kDa (ΔSU at ~40 kDa plus GST at

~26 kDa) was detected after induction of the cultures with IPTG and was present in

the insoluble fraction of the cell lysate, presumably within inclusion bodies. The

proteins present within the centrifuged and washed pelleted material are shown in

Figure 3.2 (lane 2). This material contained not only ΔSU-GST but also other

cellular proteins.

46

Figure 3.2. Coomassie brilliant blue stained SDS-PAGE of fractions collected during solubilisation, refolding, and cleavage of GST from to purify recombinant ΔSU-GST fusion protein. A: lane 1, MW markers; lane 2, pellet after cell lysis and washing; lane 3 and 4, supernatant and pellet after solubilisation in buffer B (2 M Tris pH 12 supplemented with 2 M urea and 10 mM DTT); lane 5, supernatant after refolding in refolding buffer (20 mM Tris pH 8.0 containing 0.1% CHAPS, 10 mM DTT and 1 mM PMSF); lane 6, 7 and 8, serial eluates from a glutathione Sepharose 4B column loaded with refolded protein (as illustrated in lane 5) and eluted with binding buffer. B: lane 1, MW markers in kDa; lanes 2-9, serial eluates from 10 mM reduced glutathione column containing bound ΔSU-GST fusion protein after treatment with PreScission™ protease.

The solubilisation of the insoluble ΔSU-GST product within the inclusion bodies was

pH dependent in 100 mM Tris-HCl buffer. The solubilisation increased as the pH

increased, increasing particularly from pH 10 to a maximum at pH 13. The addition

of 2% urea to the 100 mM Tris-HCl buffer did not significantly affect the

solubilisation (Figure 3.3).

1 2 3 4 5 6 7 8

1 2 3 4 5 6 7 8 9

25

37 50

75

25

37 50

75 ΔSU-GST

GST

ΔSU

GST

A

B

47

00.20.40.60.8

11.21.41.61.8

7 8 9 10 11 12 13pH

Abso

rban

ce 2

80nm

00.20.40.60.8

11.21.41.61.8

7 8 9 10 11 12 13

pH

Abs

orba

nce

280n

m

Figure 3.3. Effect of pH (top) and the effect of pH combined with 2% urea (bottom) on the solubility of recombinant ΔSU-GST fusion protein in 100 mM Tris-HCl buffer. The relative soluble protein concentration was determined by A280 ().

Comparison of the solubilisation of the washed inclusion bodies and other insoluble

cell products with the buffers shown in Table 3.1 is shown in Figure 3.4. The

greatest solubilisation was achieved with 50 mM Tris-HCl buffer at pH 8.5

containing 6 M guanidine (Buffer F), followed by 2 M Tris-HCl at pH 12 containing

2 M urea and 10 mM DTT (Buffer B). An example of the product solubilised in 2 M

Tris-HCl at pH 12 containing 2 M urea and 10 mM DTT is shown in Figure 3.2A,

lane 2.

Without addition of 2% urea

Without addition of 2% urea

With addition of 2% urea

48

0

0.2

0.4

0.6

0.8

1

1.2

1.4

A B C D E F G H

Buffers

Abs

orba

nce

units

Figure 3.4. Relative yield of soluble recombinant ΔSU-GST protein consequent of treatment of washed inclusion bodies with various solubilising buffers, determined by measurement of the A280. A, 2 M Tris-HCl pH 12; B, 2 M Tris-HCl pH 12 plus 2 M urea and 10mM DTT; C, 2 M Tris-HCl pH12 plus 10 mM DTT; D, 50 mM Tris-HCl pH 8.5 plus1% sarkosyl; E, 50 mM Tris-HCl pH 8.5 plus 8 M urea; F, 50 mM Tris-HCl pH 8.5 plus 6 M guanidine; G, 50 mM Tris-HCl pH 8.5 plus 1% SDS; H, 100 mM Tris-HCl pH 12.5 plus 2 M urea and 10 mM DTT.

As it was deemed that 6M guanidine could not be used for preparation of vaccine

material (see Discussion), further solubilisation was conducted with Buffer B (2M

Tris pH 12, 2 M urea and 10 mM DTT). SDS-PAGE of the supernatant and the non-

solubilised pellet resulted in weak bands due to a diluted protein samples used.

Some impurities from the host cell proteins were detected (Figure 3.2A, lane 3 and

4). The supernatant was then refolded overnight (Figure 3.2A lane 5) and applied

directly to a glutathione Sepharose 4B column. Unattached material was eluted with

binding buffer (PBS, pH 7.4) as shown in Figure 3.2A, lane 6). The buffer applied to

the column was then changed to the cleavage buffer which provided more stringent

wash conditions to release the ΔSU-GST fusion proteins, and PreScission™

protease to cleaves the GST-tag. After incubation period in a closed flow situation,

the bound ΔSU-GST was eluted as 2 separate proteins of 40 kDa (ΔSU) and 26

kDa (GST) as shown in Figure 3.2B.

Discussion

The expression of recombinant JDV ΔSU-GST in E. coli resulted in mostly insoluble

protein. Protein insolubility may be caused by a variety of factors including the need

49

for general molecular chaperones, cofactors, or protein partners for proper folding

(Luo & Hua, 1998, Sun et al., 2005). The properties of proteins, such as the

hydrophobic and hydrophilic regions of the aa in the protein sequences govern the

protein structure and folding. In Figure 3.5 the distribution pattern along the aa

sequences of full length JDV SU shows a highly hydrophobic region in the N-

terminal region. This hydrophobic N-terminal end may be the decisive factor that

contributes to the formation of full length-SU inclusion bodies. The proteins are

more predisposed to aggregation due to increased intermolecular interactions of

regions of the folding polypeptide chain. The truncation of the SU (ΔSU ) protein in

the current study was designed to eliminate the hydrophobic region near the N-

terminus of the full-length protein but this still did not prevent expression in an

insoluble form.

Figure 3.5. Hydrophobicity analysis of the N-terminus of the 422 aa SU protein of JDV. The profile was determined using the Kyte and Doolittle scale; hydrophilic residues have a negative score (Kyte & Doolittle, 1982).

The formation of protein aggregates within cells is a general cellular response to the

presence of mis-folded proteins when the proteins are expressed in a concentration

exceeding the degrading capacity of the cells (Markossian & Kurganov, 2004). The

expression of recombinant proteins within inclusion bodies in E. coli is affected by

the specific growth conditions including growth temperature and time of induction

with inducers, such as IPTG (Kang et al., 2007, Vera et al., 2007, Yeo et al., 2009)

50

but it is difficult to prevent their formation (Peternel et al., 2008). When present, it is

often necessary to convert the insoluble recombinant proteins and possibly

improperly folded protein into a soluble and correctly folded product to obtain a

product that is biologically active (Singh & Panda, 2005). Purification of the inclusion

bodies containing the protein of interest can assist in the removal of contaminating

cell debris and host cell proteins from the final recombinant protein; impure inclusion

body preparations result in a less effective refolding and necessitate further

purification steps (Clark, 2001).

In this current study, advantage was taken of the presence of the expressed protein

in an insoluble form in inclusion bodies to separate the inclusion bodies from

bacterial cell proteins, resulting in partial purification of the expressed recombinant

protein. This was easily achieved following disruption of the host cells by washing

the pelleted inclusion bodies with 50 mM Tris buffer (pH 8) containing 1% Triton X-

100 to remove lipid and E. coli membrane-associated proteins. The success of this

initial step was illustrated by the presence of a high proportion of ΔSU protein,

relative to other proteins, in the solubilized inclusion body preparations (Figure 3.2A,

lane 2).

A number of methods and strategies have been used previously for converting

inclusion body proteins into their soluble form (reviewed by Fischer, 1994, and

Misawa & Kumagai, 1999). The suitability and use of each method has varied.

Some have resulted in very low yields and some are incompatible with protein

purification procedures that require the protein to remain in solution as they may

result in re-aggregation and precipitation of the protein after removal of the

solubilising agent (Goldberg et al., 1991, Sunitha et al., 2000). The various

methods used have included high concentration of chaotropic agents such as urea

(Cabanne et al., 2005) and guanidine hydrochloride (Tsumoto et al., 2003b), the use

of detergents such as sodium dodecyl sulphate (SDS) (Lechtzier et al., 2002) and

sarkosyl (Mercado-Pimentel et al., 2002, Zhuo et al., 2005) with reducing agents like

dithiothreitol (DTT).

The need to consider the end use of the protein during the selection of the

solubilising agents is illustrated by 6M guanidine-hydrochloride. This is used

extensively to regenerate active proteins from inclusion bodies, but is not

acceptable for proteins to be employed as vaccines since guanidine may be

deleterious to the immunogenicity of the protein, and during attempts to remove the

guanidine from the protein, either by dialysis (Tsumoto et al., 2003a) or ion-

exchange chromatography (Ejima et al., 1999), significant re-aggregation of the

51

protein may occur. Meanwhile, urea has been generally favoured over the use of

guanidine, providing for better refolding and improved yields (Wang et al., 2005a)

and an improved immune response (Rothel et al., 1997). Detergents such as SDS

are highly effective denaturing agents but bind tenaciously to the denatured

proteins, making its complete removal from the protein problematic (Arakawa et al.,

1994).

The method chosen for solubilisation of the insoluble SU protein involved the use of

2 M urea in 2 M Tris buffer pH 12, with the addition of 10 mM DTT. This provided

considerable greater solubilisation of the insoluble proteins in inclusion bodies than

other buffers investigated, with the exception of 6 M guanidine (see Figure 3.4,

buffer B). Urea at a concentration of 2 M was less than the 8 M concentration that

has been used in many previous studies (Tsumoto et al., 2003a, Villaverde &

Carrio, 2003). The solubilisation of inclusion bodies expressed in E coli by alkaline

solutions has been previously reported (Jin et al., 1994, Lavallee et al., 1993,

Suttnar et al., 1994). In the current study a high pH appeared to have a crucial role

in destabilizing the aggregated proteins in the inclusion bodies. The addition of

10 mM DTT, a thiol containing reducing agent, allowed reduction of inter-chain

disulfide bonds by thiol-sulfide exchange and increased solubilisation of the

aggregated protein. The use of DTT has the advantage that it does not have to be

removed prior to initiation of the refolding.

The molecular basis for the denaturation of proteins by urea remains unknown,

despite its widespread use. Urea may act directly, by binding to the protein, or

indirectly, by altering the solvent environment. Most versions of the direct interaction

model (Auton et al., 2007, Dotsch et al., 1995, Zou et al., 1998) postulate that urea

binds to and then stabilizes the denatured protein, thereby favouring unfolding (Liu

et al., 1997). Alternatively, urea acts indirectly by altering the solvent environment,

thereby mitigating the hydrophobic effect and facilitating the exposure of residues in

the hydrophobic core (Courtenay et al., 2001, Tobi et al., 2003). The mechanism of

urea-promoted unfolding may depend on the urea concentration (Timasheff & Xie,

2003).

A decrease in the concentration of Tris-HCl buffer from 2 M in the solubilisation

buffer to 20 mM in the refolding buffer, in the presence of 0.1% CHAPS and 10 mM

DTT, maintained the protein in a soluble format without aggregation. This was

fortunate as aggregation is a phenomenon common to many refolded proteins

(Bondos & Bicknell, 2003). The addition of the zwitterionic detergent CHAPS to the

refolding buffer may have contributed to the continued solubility of the ΔSU protein;

52

this is one of a series of useful kosmotropes (salts that stabilize protein structures),

chaotropes (salts that destabilize protein structures), aa, sugars, and detergents

that can contribute to maintaining proteins in a soluble state (Bondos & Bicknell,

2003).

An attempt was made to cleave the GST tag from the ΔSU protein in the current

study. Sometimes, it is necessary to remove the tag since it can cause unwanted

immunological responses, inactivate the fused target protein or influence the quality

of binding properties (Buning et al., 1996, Nilsson et al., 1997). Since GST is a

relatively large tag (26 kDa), it is preferable to remove it to determine whether the

truncated SU protein is soluble and properly folded. While several methods have

been described for site-specific cleavage treatment of the fusion partner, enzymatic

methods are preferred due to their specificity and milder reaction conditions

required. The GST vector system (pGEX-6P-1) used in this study allows for

purification of the GST fusion protein and subsequent cleavage of GST from the

protein using the PreScission protease, which specifically recognizes the aa

sequence Leu-Glu-Val-Leu-Phe-Gln-Gly-Pro, cleaving between the Gln and Gly

residues (Cordingley et al., 1990). Upon removal of the GST-tag with protease,

SDS-PAGE identified 2 bands (~40 kDa and ~26 kDa), correlated with the ΔSU and

GST proteins. Further testing of the eluted ΔSU is required as a possible

complication associated with the use of reduced glutathione for elution is that it can

affect target proteins containing disulfides (Sassenfeld, 1990) and the proteases

that cleave after a basic residue such as thrombin and factor Xa, sometimes

produce non-specific and unwanted products (Jenny et al., 2003).

53

Chapter 4

Development of monoclonal antibody for the detection of recombinant JDV proteins

Summary

This chapter reports the attempted generation of hybridomas producing monoclonal

antibodies (MAbs) against the recombinant capsid (CA), truncated SU (ΔSU) and

Tat proteins of JDV using hybridoma technology. BALB/c mice were immunised 4

times subcutaneously at 2-week intervals with the proteins excised from SDS-PAGE

gels. Hybridomas secreting monoclonal antibodies against the proteins were

developed by fusion of the spleen cells of immunised mouse with SN0 myeloma

cells and detection of antibody secretion by western immunoblotting. Antibody

secreting hybridomas were cloned by limiting dilution and the isotypes of the ΔSU

and Tat MAb were determined as IgM and the CA MAb as IgG. A CA MAb

designated LD1 was shown to be an effective reagent for use in a capture ELISA for

quantitation of viral antigen and for an immunoperoxidase assay for the detection of

viral antigen in tissue sections.

54

Introduction

Following the discovery of hybridoma technology (Kohler & Milstein, 1975) and the

consequent development of monoclonal antibody (MAb) production technology, the

use of MAbs has been extremely useful in medical and biological research, enabling

a multitude of tasks such as the detection of viral antigens (Keuser et al., 2004,

Oldoni et al., 2004), tissue and blood typing (Lee et al., 2004a, Noda et al., 2002),

the identification of hormones (Fortunati & Frairia, 2003) and the identification of

tumour antigens (Nustad et al., 2004).

A range of MAbs against JDV proteins would be extremely valuable for diagnostic

purposes and particularly to aid in the identification of individual viral proteins. One

technique that has been successfully used for the detection of JDV in tissues of

cattle is immunoperoxidase staining, valuable as it can be conducted using fixed

tissue sections. For immunoperoxidase staining, monoclonal antibodies produced

previously against the 26 kDa CA protein (Kertayadnya et al., 1993) from mice

immunised with whole viral preparations, have been used but unfortunately the

hybridomas producing these MAbs were lost due to the difficulty of maintaining

liquid nitrogen supplies in Indonesia. Monoclonal antibodies would also be

extremely useful reagents for the characterisation of JDV proteins. Although the

size of the native non-glycosylated proteins can be predicted and therefore

identified with confidence in western immunoblotted viral preparations, the

glycosylated native SU and TM proteins have not been characterised with regard to

size. Monoclonal antibodies produced against recombinant SU and TM proteins

would be extremely useful for characterisation of the native viral envelope

glycoproteins and would then have application for improved diagnostic

methodology. The MAbs would also have application during the development of

Jembrana disease vaccines incorporating individual recombinant viral proteins.

This Chapter reports the production of MAb against the JDV-CA protein that was

used for the development of an antigen capture ELISA and for immunoperoxidase

staining for the detection of JDV antigen in tissues. It also describes the production

of MAb against recombinant ΔSU and Tat fusion proteins to assist in the further

characterisation of these proteins.

55


Preparation of antigens for immunisation of mice and serological tests

GST-fusion proteins

Recombinant glutathione-S-transferase (GST) JDV CA, ΔSU and Tat proteins were

expressed from JDVTab/87 genes ligated into the pGEX-6-P system (Pharmacia).

The JDV CA constructs were prepared as previously described (Burkala et al.,

1998) and contained the full-length JDV capsid gene. JDV Tat was prepared as

previously described from exon 1 of the tat gene (Setiyaningsih, 2006). JDV ΔSU

was prepared as previously described in Chapter 3. All proteins were purified using

the methods described in Chapter 3 except that GST was not cleaved from the final

purified fusion protein (as these studies were undertaken prior to the studies

reported in Chapter 3). A summary of the size of the proteins is shown in Table 4.1.

Table 4.1. Recombinant fusion proteins produced and their determined molecular weight.

Fusion protein Molecular weight (kDa)

CA-GST ~ 50

CA-biotin ~ 37

ΔSU-GST ~ 65

ΔSU-biotin ~ 53

Tat-GST ~ 38

Tat-biotin ~ 27

GST ~ 26

GST protein

Preparation of GST for use as a control reagent was as previously described by

Ditcham (2007). Escherichia coli transformed with the pGEX plasmid with no insert

were grown in 2YT supplemented with ampicillin (100 µg/ml) and induced with IPTG

(final concentration 0.1 mM) at a culture OD600 of 0.6. After 4 h of further growth, the

bacteria were harvested by centrifugation (18,000 g, 3 min), washed in PBS by

centrifugation and resuspended in 50 µl of lysis buffer/ml of original culture, and

lysed by freezing and thawing. After clarification of the lysate by centrifugation, the

56

supernatant was passed 3 times through a column of Glutathione-Sepharose resin

(0.2 ml volume, equilibrated with PBS). Bound and soluble GST was displaced with

10 column volumes of 30 mM glutathione in PBS.

Biotinylated fusion proteins

Biotinylated fusion proteins for immunisation of mice that would be recognised only

by antibody to the JDV protein and not the GST fusion tag, were produced as

previously described by Ditcham (2007). The same genes cloned into a pGEX-6P

expression vector for the production of GST fusion proteins were ligated into the

PinPoint Xa-1 T-tailed vector (Promega) for the expression of biotinylated proteins

in E. coli. Briefly, the recombinant plasmids were transformed by heat-shock into

JM109 competent E. coli. The transformed cells were then plated onto LB agar

plates supplemented with 100 µg/ml ampicillin and incubated (37ºC, overnight). The

colonies were screened for transformants with inserts in the correct orientation by

directional PCR with standard thermocycling conditions and a Tm of 50ºC. Positive

clones were verified by sequencing. Protein expression was induced by the addition

of IPTG to a final concentration of 0.1 mM in LB broth, supplemented with 100 µg/ml

ampicillin and biotin at a concentration of 2 µM. The expression of the proteins was

monitored by SDS-PAGE and western immunoblotting using an alkaline

phosphatase labelled streptavidin probe and western Blue chromogenic substrate

(Promega).

For immunisation of mice, fusion proteins were first separated by SDS-PAGE (30%

acrylamide, 0.3% bisacrylamide) and extracted from gels as described by Prussack

(1989), with slight modification. Briefly, after SDS-PAGE, gels were placed in ice-

cold 0.25 M KCl for 20 min to identify the protein bands. The bands of the

appropriate size were then excised and eluted from the gel slices in 700 µl of elution

buffer (0.01% SDS, 20 mM Tris-HCl pH 7.9, 1 mM CaCl2) overnight at 37oC. The

eluate was then centrifuged and unbound SDS was precipitated by the addition of

KCl to a final concentration of 50 mM on ice for 10 min. The mixtures were

centrifuged (10,000 g for 5 min) and the supernatant containing recombinant

proteins were stored at -20oC until required.

57

Production of fragments of Gag protein

Biotin tagged truncated gag fusion proteins as shown schematically in Figure 4.1

were produced in E. coli JM109 following the procedure described by (Desport et

al., 2005). Briefly, early log phase cultures were induced by the addition of IPTG

and recombinant fusion proteins were harvested by pelleting the cultures by

centrifugation. Pelleted cells were resuspended in cold lysis buffer, sonicated and

centrifuged again. The biotinylated proteins were purified in batch by mixing the cell

lysates with equilibrated Steptavidin Softlink resin (Promega). The bound proteins

were washed with cold lysis buffer and then eluted from the resin by the addition of

5mM biotin in cold lysis buffer. Total and purified protein concentrations were

determined using an assay based on the Bradford method (BioRad) with bovine

serum albumin standards.

Figure 4.1. Schematic representation of primer combinations used to amplify JDV gag sequences for cloning in-frame into Pinpoint Xa-1 for protein expression. The picture sourced from Desport et al. (2005).

Generation of MAb-producing hybridomas

Immunisation of mice

A 500 µl volume of each biotinylated CA, ΔSU, and Tat recombinant protein

extracted from SDS-PAGE gels was homogenised with an equal volume of

incomplete Freund’s adjuvant (Sigma) and 200 µl of the suspension was injected

subcutaneously into 8-week old female BALB/c mice. Further injections with freshly

prepared emulsions were given at 2-week intervals. Four days after the fourth

injection the mice were bled, the blood was allowed to clot at 4oC, the serum was

removed from the clot following centrifugation (10,000 g for 10 min) and specific

58

antibody was detected by ELISA. If antibody was detected, the mice required for

fusions were re-injected with a further 200 µl of freshly prepared protein emulsion

and used for fusions 4-7 days later.

Preparation of spleen cells

When required for fusions, immunised mice were euthanised and the heart was

punctured to obtain about 300 µl of blood, which was subsequently examined for

antibody. In a laminar flow hood, the skin of the mouse was swabbed with 70%

alcohol, the abdominal skin cut and resected, and the peritoneal cavity was opened

and the spleen was removed and placed in a plastic 50 mm cell culture dish (Nunc)

containing 10 ml Dulbecco’s modified Eagle’s medium (DMEM). The spleen was

cut into several pieces and disrupted using small gauge needles on sterile 10 ml

disposable syringes. After removal of the connective tissue and other large pieces

of cellular debris, the cell suspension containing an estimated 108 splenocytes was

placed in a centrifuge tube. An aliquot of the suspension was stained with 0.4%

trypan blue and counted in a haemocytometer to ensure at least 90% viability. The

spleen suspension was gently centrifuged (500 g for 5 min) to pellet the cells and

the cells were then resuspended in 10 ml of DMEM and incubated at 37oC for 15

min. For the preparation of feeder cells (splenocytes), spleens from normal BALB/c

mice were processed as described above.

Preparation of myeloma cells

Two weeks prior to fusion, a vial of NS0 myeloma cells was recovered from liquid

nitrogen and thawed in a 37°C water bath. The thawed cells were immediately

transferred into 10 ml of DMEM (serum free) before centrifugation (500 g for 5 min)

to gently pellet the cells. The cells were resuspended in growth medium (DMEM

supplemented with 10% FCS, 0.58 mg/ml L-glutamine, 0.22 mg/ml sodium

pyruvate, 1 ml/L fungizone, 200 IU/ml penicillin and 200 μg/ml streptomycin). The

cells were then distributed into culture flasks and incubated in a 5% CO2-in-air

atmosphere at 37°C. To ensure the cells were in active growth phase for cell

fusion; the cells were subcultured daily for 3 days prior to fusion. The concentration

of viable cells at the time of fusion was determined by counting the cells with a

haemocytometer following staining with 0.4% trypan blue.

59

Fusions

Cell fusions were performed by techniques similar to those described by Chan &

Mitchison (1982). The mouse spleen cell suspension (above) containing an

estimated 108 cells was mixed with the NS0 cell suspension (above) containing an

estimated 2 x 107 cells in a 50 ml round bottom sterile centrifuge tube and gently

centrifuged (100 g for 5 min). The supernatant was discarded and the cell pellets

were disrupted by gently tapping the side of the tube. One ml of 43% PEG (MW.

1300-1600; Sigma) solution was added drop-wise over a 1 min period, at the same

time shaking the tube in water bath at 37oC. The cell suspension was then

incubated at 37°C for 1 min, then 1 ml of serum-free DMEM was added over a 1 min

period, and the suspension was shaken gently for a further 1 min. Subsequently, 10

ml of serum-free DMEM was added over 1 min. The fused cells were gently

centrifuged (100 g for 5 min) and the cell pellet gently resuspended in 50 ml of

DMEM-HAT medium (DMEM supplemented with 20% FCS, 100 µM hypozanthine,

0.4 µM aminopterin, 16 µM thymidine, 0.58 mg/ml glutamine, 0.22 mg/ml sodium

pyruvate, 200 IU/ml penicillin and 200 µg/ml streptomycin). An estimated 106

splenocytes from another freshly killed normal mouse were added to the cell

suspension as feeder cells, mixed gently, and distributed into 96-well flat bottomed

cell culture microplates (Nunc) in 100 μl/well volumes (resulting in about 5 plates per

fusion). Trays were placed in a 5% CO2 incubator at 37°C.

Four days after fusion, an additional 2 drops of HAT medium containing 106 feeder

cells (spleen cells from a normal mouse) was added to each well with a sterile

Pasteur pipette, followed 2 days later by a further 2 drops of feeder cells in HT

medium (HAT medium without aminopterin). After 7 days, plates were examined

daily for evidence of hybridoma formation. Ten to 15 days after fusion, the medium

from wells containing hybridomas was screened by western immunoblotting for

antigen-specific antibodies.

Cloning hybridoma cells by limiting dilution

To assure monoclonality, antibody-producing hybridoma clones were subcloned 3

times by a limiting dilution method. The cells in antibody-positive wells were diluted

in 50 µl of cell suspension and mixed with 50 µl of DMEM-HT. The cells were

serially diluted (2-fold) in DMEM-HT in wells of a 96-well plate. Wells were

supplemented with an equal volume of feeder spleen cells and any positive wells

containing single hybridomas were checked for antibody production by western

60

immunoblotting. Any positive wells containing single hybridomas were re-cloned by

the same method. Cloned hybridomas were expanded in number by first cultivating

them in 24-well and then in 6-well plates (Nunc) and finally in 25 cm2 culture flasks

(Nunc).

Storage and recovery of hybridoma cells

Hybridomas were stored in liquid nitrogen when not required. Approximately 106

cells from a 25 cm2 flask were centrifuged (500 g for 5 min) and then resuspended

in a mixture of 90% (v/v) FCS and 10% dimethylsulfoxide (DMSO). The cells were

pipetted into 2 ml cryogenic vials and frozen slowly overnight by placing them in an

insulated container at -80oC, and they were then transferred into liquid nitrogen.

When required, the frozen cells were removed from the liquid nitrogen and thawed

rapidly in a 37°C water bath, and then immediately transferred aseptically into 10 ml

of DMEM before centrifugation (500 g for 5 min) to gently pellet the cells. The

pelleted cells were resuspended in growth medium (DMEM-HAT) supplemented

with feeder spleen cells. The cells were then distributed into wells and cultured in a

5% CO2 incubator at 37°C.

Western immunoblot analysis for antibodies

Fifteen days after fusion, the supernatants of the wells that contained large

hybridomas (that covered minimal 50% of the bottom of the wells) were screened

for protein-specific antibody by western immunoblotting using GST-ΔSU, CA and

Tat proteins produced using the pGEX expression system. The proteins were

fractionated by SDS-PAGE and then transferred to nitrocellulose membranes

overnight at 4ºC. Non-specific binding of antibodies to the membrane was blocked

by incubating the membranes in TBS (200 mM NaCl, 50 mM Tris-HCl, pH 8.0)

containing 0.05% (v/v) Tween-20 (TBST) and 5% (w/v) commercial skim milk

powder for 1 h at room temperature. After washing 3 times with PBST (PBS

containing 0.05% Tween-20), membranes were incubated with 100 µl of the

medium (diluted 1:10) from wells containing hybridomas for 3 h at room

temperature. The membranes were washed again 3 times with TBST and then

incubated with 1:2,000 dilution of anti-mouse IgG horseradish peroxidise (HRP)

conjugate for 1 h at room temperature. After 3 additional washings, HRP colour

development reagent (Bio-Rad) was added until the colour developed. The reaction

was stopped by washing the membranes in deionised water.

61

Determination of Ig isotype

The Ig subclass of MAb was determined with a Mouse Typer Sub-Isotyping Kit

(BioRad) according to the manufacturer’s instructions. One hundred µl of antigen

(GST-ΔSU, GST-Tat and GST–CA) diluted in a carbonate buffer (pH 9.6) was

added to each well of the 96-well cell culture microplates. After incubation overnight

at 4oC, the solution was discarded and the plate was washed 4 times with PBST

(PBS containing 0.05% Tween-20). The wells were dried by tapping them hard onto

tissue paper. Two hundred µl of blocking solution (5% skim milk powder in PBS)

was then added to all the wells and the plates incubated at room temperature for 1

h. The blocking solution was then discarded, the wells were washed and dried as

before, and 100 µl of hybridoma supernatant was added (ΔSU and Tat supernatants

were diluted 1:10 and CA supernatants were diluted 1:250 in PBST) and incubated

for 1 h at room temperature. The wells were washed and dried again and 100 µl of

the appropriate rabbit anti-mouse panel reagent was added for 1 h at room

temperature. The wells were then washed and dried and 100 µl of goat anti-rabbit

horseradish peroxidase conjugate was added and incubated for 1 h. After washing

the plate, 100 µl of ABTS substrate solution was added and incubated for 15 min at

room temperature when the colour development reaction was stopped by the

addition of 100 µl of 2% oxalic acid.

ELISA for detection of antibody

To obtain an optimum combination of CA antigen and CA MAb for ELISA, a

checkerboard titration of antigen and antibody was performed by indirect ELISA

using a Jgag6 antigen (a full length capsid protein; Figure 4.1) at concentrations

ranging from 8.8 to 87.5 ng/well and a MAb dilution ranging from 1:100 and 1:3,000.

The optimum combination was selected on the basis of the antigen: MAb

combination producing the highest absorbance above 0.5.

The 96-well cell culture microplates were coated with 100 µl/well of 2-fold dilutions

of Jgag6 protein in a carbonate buffer (pH 9.6) and allowed to stand at 4oC

overnight. The plates were then washed 4 times with PBST to remove the non-

adsorbed antigen. Sites not coated with the proteins were blocked with 200 µl/well

of 5% (w/v) skim milk in PBST. After incubation at room temperature for 1 h, the

plates were washed and dried as described previously. Then 100 µl/well of 2-fold

dilutions of MAb (diluted with PBST) was added to the plate, which was then

reacted at room temperature for 1 h and washed and dried. One hundred µl/well of

62

goat anti-mouse IgG conjugated to horseradish peroxidase diluted 1:4,000 with

PBST was added to each well and the plates were incubated at room temperature

for 1 h. The plates were then washed and 100 µl/well of a substrate solution (TMB;

tetramethylbenzidine, Bio-Rad) was added to each well. After colour development at

room temperature, the reaction was stopped by the addition of 100 µl/well of 2%

oxalic acid and the absorbance read at a wavelength of 405 nm.

Antigen capture ELISA for quantitation of viral antigen

An antigen capture ELISA was performed as previously described (Stewart et al.,

2005). Briefly, 96-well flat bottom cell culture microplates were coated with either a

1:200 dilution of the produced MAb or with a 1:1,000 dilution of the previously

produced BC10 CA MAb (Kertayadnya et al., 1993) in 100 µl carbonate buffer pH

9.6 per well, then incubated at room temperature for 1 h. The microplates were

washed 3 times with PBST and blocked with 200 µl of blocking solution (2% casein

and 1-5% skimmed milk powder in PBST) and incubated at room temperature for

1h. After the plates were washed 3 times with PBST, 100 µl of Jgag6 protein (100

ng/ml) in PBST containing 1% skim milk powder was added. The plates were again

washed 3 times with PBST and 100 µl of a 1:4,000 dilution of polyclonal rabbit anti-

JDV CA (from J. Brownlie) was added. After another 3 washes with PBST, 100 µl of

goat anti-rabbit HRP conjugate (ICN, Australia) (1:4,000) was added to each well

and incubated at room temperature for another 1 h. After another wash with PBST,

100 µl of TMB colour development reagent was added to each well, the microplates

were incubated at room temperature until colour development and the absorbance

of the colour reaction was determined at 450 nm.

Immunoperoxidase staining of virus in fixed tissue sections

The CA MAb was analysed by immunoperoxidase staining of formaldehyde-fixed

paraffin-embedded issue sections from JDV-infected and control animals. The

antigen retrieval was performed by microwave treatment of 6 µM tissue sections for

2 x 4 min treatments at high wattage (805 watt), followed by 2 x 4 min treatments at

low wattage (230 watt). The sections were then left to cool in distilled water. The

tissue sections were rinsed in PBS, blocked in 3% H2O2 for 5 min to remove

endogenous peroxidases and then rinsed again before incubation for 1.5 h at room

temperature with anti-CA (MAb LD1 or MAb BC10) diluted 1:50, 1:100 and 1:200 in

PBS. The sections were then washed in PBS before the addition of secondary

63

antibody (HRP-conjugated goat anti-mouse IgG) (EnVision) for 30 min at room

temperature. After washing, the bound antibodies were visualised as a brown

insoluble precipitate by the addition of 3,3’-diaminobenzidine (DAB; DAKO) solution

for 5-10 min at room temperature. The slides were counterstained with Harris

haematoxylin and dehydrated before they were mounted with DePex (BDH

Chemicals).

Results

Production of MAb-secreting hybridomas

BALB/c mice immunised 4 times with biotinylated ∆SU, CA and Tat extracted from

acrylamide gels developed serum antibody that was detectable by western blot

utilising the homologous GST-fused protein as an antigen.

One week after fusion, hybridoma cell growth was visible microscopically in the

96-well microplates and at about day 14, the number of hybridoma cells increased

and grew large enough to where they occupied one half or more of the surface of

each well (Figure 4.2).

Figure 4.2. Microscopic image of hybridoma cells 14 days after fusion. Magnification X400.

From the mice immunised with the biotin-tagged ΔSU, western immunoblotting

detected antibody production in 68 wells containing hybridomas from 5 plates. Many

64

of the these supernatants reacted with more than one band, and to achieve mono-

specificity, the hybridoma cells from all the positive wells were then cloned by

limiting dilution and after 3 cloning procedures, 7 of the 68 hybridomas produced

MAb reactive with a single band of about 65 kDa in western immunoblots (Figure

4.3). The MAbs were all of IgM subclass and had kappa light chains.

Fusion of spleen cells obtained from BALB/c mice immunised with biotinylated-CA

recombinant proteins resulted in 74 wells containing hybridomas, which detected a

mixture of antibodies by western immunoblotting. After cloning by a series of 3

limiting dilutions, the supernatant from one hybridoma was selected that reacted

with one protein only of ~ 50 kDa (Figure 4.4). Cells from one well were expanded in

25 cm2 culture flasks. This cloned hybridoma was selected for further analysis and

designated as LD1.

Figure 4.3. The supernatant from hybridoma-containing wells which reacted with recombinant ΔSU-GST proteins in western immunoblots. After cloning, 7 of the hybridomas produced MAb that reacted to a ~65 kDa protein only (positive lanes indicated by arrows). The control molecular weight markers and their size are shown on the left.

MW

75

50

37

25

65

Figure 4.4. The supernatant from hybridoma-containing wells was screened against recombinant CA-GST fusion proteins by western immunoblotting. The positive hybridomas were cloned 3 times by a limiting dilution method and the supernatant from several clones that reacted with one protein only of ~ 50 kDa. The control molecular weight markers and their size are shown on the left.

Following fusion of spleen cells obtained from BALB/c mice immunised with

biotinylated-Tat recombinant proteins, 34 wells containing hybridomas were

produced but only one was detected that contained antibody against Tat. This

positive hybridoma was re-cloned 3 times (Figure 4.5). The MAb against

recombinant Tat protein was of IgM isotype and possessed kappa light chains.

Figure 4.5. The supernatant from hybridoma containing wells was screened against recombinant Tat-GST fusion protein by western immunoblotting. The positive wells were cloned by a limiting dilution method and one of the cloned hybridomas produced antibody reactive with the recombinant protein. The gel depicts the reactivity of multiple wells containing the same cloned hybridoma. The control molecular weight markers and their size are shown on the left.

75

50

37

25

MW

75 50 37 25

MW

66

Determination of optimal antibody and antigen dilution

A checkerboard titration was conducted to determine the optimal working dilution of

the LD1 CA MAb and the optimal concentration of Jgag6 protein for maximum

sensitivity. The optimum conditions were selected on the basis of the highest

absorbance above 0.5. The optimal antigen concentration was 87.5 ng/well of

Jgag6 and the optimal LD1 CA MAb dilution was 1:500 (Figure 4.6).

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

87.5 ng 58.3 ng 35 ng 23.3 ng 17.5 ng 14 ng 11.7 ng 8.8 ng

Concentration of Jgag6

Abs

orba

nce

(405

nm)

1:100 1:200 1:250 1:500 1:7001:1000 1:1500 1:2000 1:2500 1:3000

Figure 4.6. A checkerboard titration to determine the reactivity of various dilutions of the LD1 CA MAb with varying dilutions of the Jgag6 protein by ELISA.

Analysis of reactivity of LD1 with different proteins

The MAb LD1 CA was reacted in western immunoblots against ΔSU, CA and Tat

fusion protein products that were produced (Figure 4.7). The MAb LD1 reacted with

CA-GST and CA-His (a gift from William Ditcham) but not with the ΔSU and Tat

proteins.

67

Figure 4.7. Reactivity by western blotting of the anti-CA MAb LD1 with various recombinant proteins. Lane 1, GST; lane 2, Tat-GST; lane 3, biotinylated-ΔSU; lane 4, CA-GST; lane 5, biotinylated Tat; lane 6, Tat-His; lane 7, CA-His; lane 8, ΔSU-GST. The control molecular weight markers and their sizes are shown on the left.

The CA MAb LD1 was reacted in western immunoblots against the 9 overlapping

fragments of Gag proteins (Figure 4.1) and the results are shown in Figures 4.8 and

4.9. The MAb LD1 recognised Jgag2, Jgag3, Jgag4, Jgag5, Jgag6, and Jgag11 but

not Jgag1 (MA protein) and Jgag 7 confirming its specific reactivity to the CA

protein.

This reactive epitope for this MAb was a region of CA that was encoded by a region

of the JDV genome between nucleotides 604 and 810. The reactivity of LD1

monoclonal antibody was different to the hyperimmune sera when tested against

the same truncated Gag proteins (Figure 4.9); the hyperimmune serum did not

recognise the Jgag11 product whereas it was recognised by the MAb LD1,

indicating a difference in reactivity between native and recombinant CA protein

epitopes.

MW 1 2 3 4 5 6 7 8

75 50

37

25

68

Figure 4.8. Schematic diagram of recombinant Gag fragments expressed in E. coli and reacted in western immunoblots with truncated proteins encompassing regions of the MA, CA and NC. The top (multi-coloured) bar represents the full length of the region covered by the individual peptides that were produced. Numbers below the bar show nucleotide numbers of the regions of gag encoding the truncated fragments Jgag1 to Jgag13. Those fragments shown in red were reactive to the LD1 anti-CA MAb, those in black were non-reactive. The schematic was adapted from Desport et al. (2005).

Figure 4.9. Reactivity of different truncated Gag proteins detected by hyperimmune sera (A) and anti-CA LD1 (B). Lane 1, Jgag1; lane 2, Jgag2; lane 3, Jgag4; lane 4, Jgag4; lane 5, Jgag5; lane 6, Jgag6; lane 7, Jgag7; lane 8, Jgag11; lane 9, Jgag13.

Comparison LD1 and BC10

To determine if the MAb LD1 was suitable for the antigen capture ELISA to

quantitate virus, the reactivity of MAb LD1 was compared with the MAb BC10; as

whole virus was unavailable for this, the Jgag6 peptide was used instead. The MAb

Matrix Capsid MHR NC

Jgag1

Jgag2

Jgag3

Jgag4

Jgag5

Jgag6

Jgag7 Jgag11

Jgag13

229 604 1030 810 1123 1282 1540

75 50 35

25

MW

1 2 3 4 5 6 7 8 9 1 2 3 4 5 6 7 8 9

75

35

25

50

(A) (B)

69

LD1 (at 1:200 dilution) was able to detect 1,750 ng/ml Jgag6 whereas the MAb

BC10 (at 1:1,000 dilution) detected 875 ng/ml Jgag6 (Figure 4.10). Evaluation with

different concentration of blocking solutions showed that the results were not

significantly different, and 5% skim milk powder in PBS was arbitrarily selected for

use in all serological assays.

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

3500 1750 875 437.5 218.75 109.375 54.688

Jgag6 (ng/ml)

Abs

orba

nce

at 4

50 n

m

Casein 1%SM 2%SM 3%SM 4%SM 5%SM

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

3500 1750 875 437.5 218.75 109.375 54.688

Jgag6 (ng/ml)

Abs

orba

nce

450

nm

Casein 1%SM 2%SM 3%SM 4%SM 5%SM

Figure 4.10. Comparison of antigen capture ELISA with MAb BC10 at 1:1,000 dilution (top) and MAb LD1 at a 1:200 dilution using Jgag6 protein as antigen and with various blocking agents including casein and skim milk (SM) powder (bottom).

70

Immunoperoxidase staining of tissue sections

The IHC procedure was conducted with spleen tissue from uninfected control cattle

and from cattle on the 2nd day of fever after experimental infection with JDVTab/87.

The staining pattern was observed with 1:50, 1:100, and 1:200 dilutions of the CA

MAb LD1. Granular brown staining in the cytoplasm was detected with all dilutions;

however, the 1:200 dilution resulted in less background staining than the other

dilutions. Comparison of the CA MAb LD1 with the MAb CA BC10 for

immunoperoxidase staining of virus in formaldehyde-fixed paraffin-embedded

sections showed that while both were effective reagents for the detection of virus

antigen, the MAb BC10 provided more intense brown staining of tissues than did the

MAb LD1 (Figure 4.11).

Figure 4.11. Immunoperoxidase staining using MAb CA LD1 (B) and MAb CA BC10 (C) of formaldehyde-fixed paraffin-embedded spleen tissue section from uninfected cattle (A) and JDV-infected cattle (B and C). Magnification X20.

Discussion

Only one MAb was produced against each of the 3 proteins that were selected.

While further investigation may have resulted in a greater range of MAbs covering

multiple epitopes, the aim of this project was to quickly produce a single MAb

(A) (B)

(C)

71

against each of the 3 proteins, that could then be used to identify each of the

proteins and expand the range of diagnostic reagents available for Jembrana

disease. As the immunising protein was in a denatured state it was not expected

that MAb would be produced against all epitopes on native proteins.

As the proteins used for this study were fusion proteins, to facilitate the process of

selection of specific antibodies the fusion tag on the immunising protein was

different to the fusion protein used as antigen in the serological test used for

screening for MAbs. western immunoblotting was used as a screening assay and

while it was possibly less sensitive than ELISA, it had the advantage of identifying

the reactivity of the antibodies with proteins of the appropriate size. Additionally,

certain antigens that bind to ELISA plates may also not be accessible for the

antibodies due to steaic hindrance (Mohammad & Esen, 1989). It would have been

appropriate to test the MAbs against native protein antigens but these were not

available.

The anti-CA MAb LD1 was shown to react specifically with the CA as it reacted to

the CA-specific construct Jgag11. Desport et al. (2005) suggested that the Jgag11

region must contain a reactive epitope since it was recognised by the hyperimmune

JDV sera and to a lesser extent hyperimmune BIV sera. In the previous

investigation reported by Desport et al. (2005) the CA-specific MAb BC10 was

reactive with the Jgag13 construct as was the MAb LD1. However, some CA-

specific MAbs investigated by Desport et al. (2005), which included BC1 and BC10

that were produced by Kertayadnya et al. (1993), did not react with Jgag11 whereas

the MAb produced in this study (LD1) reacted with Jgag11 indicating it was different

to the other MAbs previously available for JDV research (Kertayadnya et al., 1993).

The potential application of the MAbs produced, particularly the anti-CA MAb LD1,

was illustrated by its ability to be used as an alternative to MAb BC10 for the

quantitation of virus (Stewart et al., 2005) and the staining of JDV in paraffin-

embedded tissue sections. This was facilitated as this MAb was of IgG type. The

MAbs against the SU and Tat proteins also have the potential to provide diagnostic

and research reagents for the detection and characterisation of these 2 proteins but

unfortunately, these MAbs were of IgM type and this will limit their usefulness; most

commercially available labelled secondary immunoglobulins are directed against

IgG.

72

Chapter 5

Production of single chain fragment antibody (scFv) against recombinant ΔSU and CA proteins of Jembrana disease virus

Summary

Single chain fragment antibody (scFv) that specifically bound to recombinant

truncated SU (∆SU)-GST fusion protein and capsid (CA) proteins from Tomlinson I

and J phage display antibody libraries were produced. Two different methods of

panning, a colony lift assay (CLA) and microtiter plate technique, were used to

screen and identify the scFv: the CLA was used to detect scFv against the less

soluble ΔSU protein and the microplate technique was used for the more soluble CA

proteins. The phage-positive clones from both methods were transformed into

E. coli HB2151 and the expression of soluble scFv was induced by the addition of

IPTG. The scFv were then identified by ELISA. Comparison of scFv binding

affinities assessed by indirect and capture ELISA tests indicated that the scFv

produced against the SU was reactive not with the ∆SU component but the GST

component of the ∆SU-GST fusion protein. The microplate panning generated

colonies reactive with a truncated CA protein (Jgag6) and recombinant CA-Tat

polyproteins. Four Jgag6 clones with the highest affinity were expressed as soluble

scFv in the cell periplasmic area.

73

Introduction

The technology for the large scale production of highly specific antibody reagents

has developed significantly with recent advances in recombinant antibody

technology that may overcome many of the problems associated with the production

of monoclonal antibodies (MAb) by conventional hybridoma technology (see

Chapter 4). Phage display antibody technology provides an in vitro selection

technique in which antibody fragments are genetically fused to the coat protein of a

filamentous bacteriophage, resulting in display of antibody fragments on the exterior

surface of the phage virion. The DNA encoding the fusion product resides within the

virion. Once cloned, it is possible to improve the affinity and specificity of antigen

binding by mimicking somatic hypermutation during an immune response (Gram

et al., 1992). The method was introduced for the first time by Smith (1985) who

fused a portion of the gene encoding the EcoRI endonuclease to the minor CA

protein pIII present on the surface of bacteriophage M13. For reviews of the

technique see Hoogenboom et al. (1998) and Winter et al. (1994).

The use of phage display technology has many applications, including the isolation

of antigen-specific antibody fragments either as fragment-antigen-binding (Fab) or

single-chain antibody fragments (scFv). Recent reports of these applications include

the production of scFv against the SARS-associated coronavirus (Liu et al., 2005b),

human fibrin clots (Yan et al., 2004) and the capsid (CA) protein and

transmembrane (TM) glycoprotein (gp46) of Maedi-visna virus (Blazek et al., 2004,

Celer et al., 2003). ScFvs have also been expressed intracellularly to inhibit the

function of intracellular proteins (Lobato & Rabbitts, 2004). These intracellular

antibodies or intrabodies have been applied to engineer a human MAb F240, which

recognised the disulfide loop-bonded immunodominant epitope of HIV-1 gp41 (Liu

et al., 2005a) and human MAb F102, which recognised the conserved CD4-binding

region of HIV-1 gp120 (Wang et al., 2005b). These results suggested that the

constructed scFv F240 and scFv F102 may inhibit viral replication and may be

useful for gene therapy. Other research has demonstrated that these intrabodies

can be directed to multiple HIV-1 target proteins that are present in cells, including

the major CA p24 protein (de Haard et al., 1998), Gag p17 protein (Tewari et al.,

1998), Vif (Goncalves et al., 2002) and Vpr (Krichevsky et al., 2003).

There are different methods of panning to isolate high affinity scFv against a wide

variety of target antigens. Immunotube-based selection has resulted in production of

scFV with high affinity for the target antigen but it requires a high volume of purified

74

antigen (Mi et al., 2005). Similar techniques have been described that use

microplates or tagged antigens that can be isolated using magnetic beads (Siegel et

al., 1997). However, those techniques may not be suitable for antigens with poor

solubility or that tend to aggregate. An improved method for phage display panning

was developed employing whole cells that produced specific phage after several

rounds of panning (Radosevic et al., 2003, Siva et al., 2008, Sui et al., 2004).

Storage of hybridomas for conventional MAb production has always been a problem

within Indonesia because of difficulties in maintaining liquid nitrogen storage

facilities due to supply problems. To overcome the limitation represented by the

storage of hybridomas producing MAb against JDV proteins prepared by

conventional methods as described in Chapter 4, antibody phage display was

attempted to produce specific scFv directed towards the recombinant truncated SU

(ΔSU) and CA proteins of JDV. Due to the nature of the recombinant proteins used

in this study, 2 methods of panning were employed for each of the 2 proteins. For

the insoluble ΔSU protein, a colony lift assay (CLA) was used. This method is

based on the production of scFv by phage-infected bacteria plated on IPTG-

containing agar plates (Siegel et al., 1997). The scFv diffuse through the bacterial

membrane and bind to the antigen-coated membrane that is placed under the

bacterial membrane. The scFv that bind to the antigen-coated membrane are

detected using antibodies specific for the tag fused to the scFv. An alternative

microplate technique was also used, where phages are absorbed onto ELISA plates

coated with CA that are subsequently eluted with trypsin. The eluted phages are

used to infect E. coli of the TG1 genotype to produce scFv-phages, followed by

soluble scFv expression in E. coli of the HB2151 genotype. The synthetic phagemid

library, the human single-fold scFv libraries I+J (Tomlinson I+J), used for the

generation of scFv antibodies directed against recombinant JDV SU and CA is

described.


Tomlinson I and J phage antibody libraries

Two phage-display antibody libraries generated at the MRC Centre for protein

engineering (Cambridge, UK) were used. Both libraries were based on a single

human framework for VH(V3-23/DP-47 and JH4b) and VK(012/02/DPK9 and JK1),

with side chain diversity (either NKK or DVT encoded library J and I, respectively)

incorporated at positions in the antigen binding site that make contact with antigen

75

and are highly diverse in the mature repertoire (Rubinstein et al., 2003). The vector

used for the expression of scFv antibody fragments is depicted in Figure 5.1.

Phagemid vector pIT2 contains a lac promoter and a pelB leader sequence up-

stream of the scFv insert, which is then followed by hexa-histidine and myc tags, an

amber stop codon and the gene encoding the pIII phage coat protein. After helper

phage KM13 infection, scFv up-stream of the amber stop codon is displayed on the

phage particle with the use of a suppressor strain such as TG1. However, scFv with

a His6-myc tag is secreted into the periplasmic space with the use of a non-

suppressor strain such as HB2151.

colE1 ori M13 oriamp

pIT2

lac promoter

RBS pelBleader

linker myctag

gIII

NotISfiI/Ncol

XhoI

SaII

amber

colE1 ori M13 oriamp

pIT2

lac promoter

RBS pelBleader

linker myctag

gIII

NotISfiI/Ncol

XhoI

SaII

amber

Figure 5.1. Schematic representation of the scFv expression vectors: amp, ampicillin resistance encoding gene; colE1 ori, origin of DNA replication; pelB, signal peptide sequence of bacterial pectate lyase; myc tag, sequence encoding an epitope recognised by the MAb 9E10. Adapted from the Tomlinson I+J protocol, http://www.geneservice.co.uk/products/proteomic/ scFv_tomlinsonIJ.jsp.

In conjunction with the phage display antibody library, the E. coli strain TG1 (K12,

∆(lac-pro), supE, thi, hsd∆5/F’traD36, proA+B+, lac Iq, lacZ∆M15) used for the phage

rescue and the non-suppressor E. coli strain HB2151 (K12, ara, ∆(lac-pro),

thi/F’proA+B+, lacIqZ∆M15) used for the preparation of single-chain Fv fragments

were supplied by I. M. Tomlinson, Laboratory of Molecular Biology (MRC,

Cambridge).

76

Preparation of phage library stocks

Tomlinson I and J libraries were delivered as glycerol stocks. One hundred µl of the

E. coli (TG1 genotype) glycerol stock was inoculated into 200 ml of pre-warmed 2YT

broth containing 1% glucose and 100 µg/ml of ampicillin and incubated with shaking

at 37oC to an optical density at 600 nm (OD600) of 0.4. KM13 helper phage (2 x 1011

pfu) was added to 50 ml of each culture and incubated without shaking in a 37oC

water bath for 30 min. The remaining 150 ml of each culture was incubated for a

further 2 h with shaking at 37oC and used to make secondary bacterial stocks. After

incubation, the phage-infected culture was centrifuged (3,000 g for 10 min) and the

resulting pellet was resuspended in 100 ml of 2YT broth containing 100 µg/ml of

ampicillin, 50 µg/ml of kanamycin and 0.1% glucose and re-incubated overnight at

30oC. The resulting culture was clarified by centrifugation (3,000 g for 30 min) and

phage particles were concentrated from the supernatant by precipitation with 20 ml

of PEG/NaCl (20% [w/v] polyethylene glycol 6,000 in 2.5 M NaCl) for 1 h on ice.

The suspension was then centrifuged (3,000 g for 30 min), the supernatant was

discarded and the pellet resuspended in 4 ml PBS and re-centrifuged (3,000 g for

30 min) and the supernatant containing phage was kept at 4oC for short term

storage or at –70oC in PBS with 15% glycerol for longer storage.

To titrate the phage, 10-fold serial dilutions of the phage suspension were prepared

in PBS. To 100 µl of each dilution, 900 µl of TG1 cells at an OD600 of 0.4 was added

and the tubes then incubated at 37oC in a water bath for 30 min. Ten µl of each

dilution was then spotted on a 2YT agar plate containing 1% glucose and 100 µg/ml

ampicillin.

Protein preparations

A ΔSU-GST protein was prepared as previously described (Chapter 3). A

biotinylated CA fragment (Jgag6) was expressed and solubilised as described

previously (Chapter 4). A recombinant CA-Tat fused polyprotein (Lewis, 2008) was

kindly provided by Dr Joshua Lewis (this laboratory). Ubiquitin (Sigma) was used as

a control antigen for assessment of the quality of the panning process.

77

Preparation of scFV by panning using CLA

The detection of ΔSU scFv was attempted using CLA; the principle of the assay

utilised is demonstrated in Figure 5.2 (Radosevic et al., 2003).

Figure 5.2. Schematic representation of the colony lift assay. YTAG denotes 2YT agar containing 100 µg/ml ampicillin and 1% glucose. YTAI denotes 2YT agar containing 100 µg/ml ampicillin and 1 mM IPTG. Diagram sourced from Radosevic et al. (2003).

Selection of ΔSU phage

Solubilised ΔSU was separated by SDS-PAGE and electro-transferred to a

nitrocellulose membrane. The membrane was stained with Ponceau red and after

washing, non-reactive sites on the membrane were blocked by immersing it in 5%

skimmed milk powder in TBST for 1 h at 4oC. The membrane was washed with

TBST and the band that corresponded to ΔSU was excised. The excised ΔSU

protein was then incubated with ~1012 phage in 1 ml of 5% skimmed milk powder in

78

TBST at 4oC for overnight. After 3 washes with TBST the bound phages were eluted

by adding 500 µl of trypsin-PBS (50 µl of 10 mg/ml trypsin stock in 450 µl PBS) for

10 min and propagated in fresh TG1 cells by adding 250 µl of the eluted phage

particles to 1.75 ml of exponentially growing TG1 cells at an OD600 of 0.4 and

incubating them for 30 min at 37oC in a water bath without shaking. A sample was

removed and the phage titrated. The remaining TG1 culture was centrifuged

(microcentrifuge for 5 min) to pellet the cells and the pellet resuspended in 50 µl of

2YT and plated on a 100 mm 2YT agar plate containing 100 µg/ml ampicillin and

1% glucose. The plates were then incubated overnight at 37oC. After overnight

growth, 2 ml of 2YT medium was added to the plate and the bacteria were collected

by scraping with a glass spreader. Fifty µl of the scraped bacteria suspension was

added to 50 ml of 2YT containing 100 µg/ml ampicillin and 1% glucose and grown at

37oC while shaking until the OD600 was 0.4. Then 5 x 1010 helper phage was added

to 10 ml of this culture and incubated at 37oC for 30 min without shaking. The

bacteria were centrifuged (3,000 g for 10 min) and the pellet resuspended in 50 ml

of 2YT medium containing 100 µg/ml ampicillin, 50 µg/ml kanamycin and 0.1%

glucose and incubated with shaking at 30oC overnight. The overnight culture was

centrifuged (15 min at 3,300 g at 4oC) and 40 ml of the phage-containing

supernatant was transferred to a new tube and precipitated with 10 ml of ice-cold

PEG/NaCl (20% polyethylene glycol 6,000, 2.5 M NaCl) for 1 h. The phage

suspension was centrifuged (3,300 g for 30 min) and the pellet resuspended in 2 ml

PBS and centrifuged (~11,000 g for 10 min in a micro centrifuge) to remove

bacterial debris.

The phage selection protocol was repeated a further 3 times (a total of 4 times) and

the enrichment of specific phages in each round of the selection was assessed by

polyclonal phage ELISA.

Coating ΔSU proteins onto nitrocellulose membranes

Two ml of ΔSU-GST protein (100 µg/ml) was further diluted in 4 ml of PBS and

poured onto the surface of a nitrocellulose membrane (pore size 0.45 µm; Bio-Rad)

in a culture dish and left overnight at 37oC. The excess suspension was then

removed and the ΔSU-coated membrane was rinsed with PBS. Non-reactive sites

on the membrane were then blocked by immersing it in 10 ml of PBS containing 3%

(w/v) skimmed milk powder for 1 h at room temperature. The blocked membrane

was rinsed twice with PBS and then air-dried.

79

Panning for ΔSU scFV

The phage rescued from each round of selection (as described above) were used to

infect E. coli HB2151 cells (non-suppressor strain) for the attempted expression of a

soluble scFv antibody. The bacteria were grown in 2YT culture medium at 37oC until

OD600 of 0.4 was achieved. Two hundred ml of the culture was infected with 10 µl of

phage from each round of panning (above) and then incubated at 37oC for 30 min

without shaking. Serial 10-fold dilutions of the phage-infected HB2151 bacteria were

plated on YTAG plates (2YT agar containing 100 µg/ml ampicillin and 1% glucose)

and grown overnight at 37oC. The dilution which resulted in isolated bacterial

colonies was used for the CLA (further referred to the original bacterial plate).

The nitrocellulose ΔSU-coated membrane (prepared as described above) was

soaked in 2YT medium containing 100 µg/ml ampicillin and 1 mM IPTG and then

placed, coated side upwards, onto the surface of an YTAI agar plate (2YT agar

containing 100 µg/ml ampicillin and 1 mM IPTG). Another non-coated membrane

was placed on top of the original bacterial plate (referred to as the master

membrane). The master membrane was removed from the original bacterial plate

and placed, colony side up, on the antigen-coated membrane on the YTAI agar

plate. The YTAI agar plate was then incubated overnight at 30oC. Both membranes

were marked simultaneously by stabbing them with a needle for orientation

purposes.

The master membrane was removed and placed, colony side up, onto a fresh

YTAG agar plate. The plate together with the master membrane was stored at 4oC

and later used for picking selected clones. The ΔSU-coated membrane was

removed from the YTAI plate, washed 3 times with 10 ml PBST (PBS containing

0.05% Tween-20) for 10 min with gentle shaking, blocked with PBS/3% dried

skimmed milk powder for 1 h at room temperature and rinsed twice with PBS/0.05%

Tween-20. The membrane was incubated with 10 ml mouse anti-His antibody

(Sigma) (recognising the His-tag fused to scFv) diluted 1:2,000 in PBS/1% dried

skimmed milk powder, for 1 h at room temperature. After incubation, the filter was

washed 3 times with 10 ml PBST for 10 min each time, with shaking, followed by

incubation with 10 ml of 1:3,000 dilution of goat anti-mouse Ig-HRP (ICN) in

PBS/1% dried skimmed milk powder for 1 h at room temperature. The membrane

was washed 3 times with 10 ml PBST for 10 min each time with gentle shaking and

then rinsed twice with PBS. The membrane was then treated with 10 ml of HRP

colour development solution (Bio-Rad) until spots were visible when the membrane

was rinsed with water to stop the reaction. The spots from the ΔSU-coated

80

membrane were used to identify the corresponding colonies responsible for the

production of specific scFv on the master membrane and the original plate.

Preparation of soluble ΔSU scFv

Individual colonies were picked from each positive agar plates and resuspended in

100 µl 2YT medium containing 100 µg/ml ampicillin and 1% glucose in 96-well

microplates and incubated overnight at 37oC with shaking. The overnight cultures

were then diluted 1:20 and added to wells of another 96-well microplate containing

200 µl fresh 2YT medium supplemented with 100 µg/ml ampicillin and 0.1% glucose

and grown at 37oC with shaking until the OD600 was about 0.9. IPTG was added to a

final concentration of 1 mM and the cultures were incubated with shaking overnight

at 28oC. Bacteria were pelleted by centrifugation (1,800 g at 4oC for 15 min) and 50

µl of the supernatant was used for ELISA.

ELISA

To evaluate the specificity of the scFv against the ΔSU protein, 2 kinds of ELISA

were used: an indirect phage ELISA and a capture phage ELISA.

Indirect ELISA. ELISA microplates (Maxisorp, Nunc) were coated overnight at 4oC

with 100µl/well of 10 µg/ml ΔSU-GST fusion protein and GST alone in 100 mM

carbonate buffer (pH 9.6). After 3 washes with PBST, the plates were subsequently

blocked with 200 µl/well of 5% skimmed milk powder in PBS for 2 h at room

temperature. The plates were washed 3 times with PBST and then 75 µl/well of the

scFv supernatants were added to the plates and they were incubated at room

temperature for 2 h. PBS was used as negative control. After 3 washes with PBST,

the plates were incubated for 1 h with 100 µl/well of a 1:5,000 dilution of HRP-

conjugated protein L (Sigma) in PBS. The plates were washed a further 3 times with

PBST and once with PBS before they were reacted with 50 µl/well of TMB (3,3’,5,5’-

tetramethylbenzidine) colour development reagent for 5-15 min at room

temperature. The colour development was stopped with 50 µl/well of 1 M H2SO4 and

the absorbance was determined at 450 nm and 650 nm.

Capture ELISA. ELISA microplates (Maxisorp, Nunc) were coated overnight at 4oC

with 100 µl/well of goat anti-GST antibody (diluted 1:5,000 in 100 mM carbonate

buffer pH 9.6). After 3 washes with PBST the plates were subsequently blocked with

100 µl/well of goat anti-GST antibody (diluted 1:5,000 in 100 mM carbonate buffer

81

pH 9.6). After 3 washes with PBST the plates were subsequently blocked with 200

µl/well of 5% skimmed milk powder in PBS for 1 h at room temperature. The plates

were then washed 3 times with PBST and 100 µl/well of 10 µg/ml ΔSU-GST or GST

only were added to the wells and incubated for 1 h at room temperature, then PBS

was added to blank wells as a negative control. After 3 washes with PBST, 75

µl/well of the scFv-containing supernatant was added to the plates for 2 h at room

temperature. After washing as described above, the plates were further incubated

for 1 h with 100 µl/well of a 1:5,000 dilution in PBS of HRP-conjugated protein L

(Sigma). The plates were washed a further 3 times with PBST and once with PBS

before the addition of 50 µl/well of substrate solution containing TMB for 5-15

minute at room temperature. The colour development was stopped with 50 µl/well of

1 M H2SO4 and the absorbance determined at 450 nm and 650 nm.

Preparation of CA scFV by panning with microplate method

Selection of CA scFv-phage

The panning procedure was conducted using 96-well microplates. One hundred µl

of Jgag6 (10 µg/ml), CA-Tat polyprotein (10 µg/ml) or ubiquitin (100 µg/ml) was

added to wells of a 96-well microplate (Nunc). The plates were incubated for 3 h at

room temperature and then the non-reactive sites were blocked by the addition of

with 2% skimmed milk powder in PBS. The plates were then washed 3 times with

PBS and 100 µl of the supernatant containing the phage library stocks was added to

each well. Plates were incubated at room temperature for 2 h and washed 10 times

with PBST. To the wells containing bound phage, 100 µl of 10 mg/ml trypsin in PBS

was added and incubated at room temperature for 10 min. The eluted phage were

used to infect 1 ml of log-phase TG1 cells at an OD600 of 0.4 in 2YT medium and

grown at 37°C for 1 h. Ampicillin was added to a final concentration of 100 µg/ml

and glucose to a final concentration of 1% (w/v). Helper phage (5x1010 cfu) was

then added to the TG1 cell suspension and the culture was incubated at 37°C for 30

min then centrifuged (3,000 g for 10 min at room temperature) to pellet the bacteria.

The bacterial cells were then resuspended in 1 ml 2YT containing 100 µg/ml

ampicillin, 50 µg/ml kanamycin and 0.1% glucose and cultured overnight at 28°C.

The phage panning protocol was repeated a further 3 times.

82

Preparation of scFv-phage for monoclonal ELISA

Serial dilutions (10-1, 10-2, 10-4 and 10-6) of E. coli TG1 infected with eluted scFv-

phages from each round of selection were plated on 2YT agar with 1% glucose and

100 µg/ml ampicillin to enable the selection of individual separated colonies. These

individual colonies were picked and inoculated into individual wells of a microplate

with 100 µl of 2YT, 1% glucose and 100 µg/ml ampicillin and cultured at 37oC

overnight with shaking (250 rpm). Then, 2 µl of the overnight cultures from each well

was transferred into a new plate with 200 µl of 2YT, 1% glucose and 100 µg/ml

ampicillin per well and incubated for 2 h at 37oC with shaking. The remainder of the

bacterial culture from the overnight plate was stored frozen after the addition of 15%

glycerol. Next, 109 pfu of the helper phage KM13 was added per well and shaken

for 60 min. This was centrifuged (1,800 g for10 min) and samples of the supernatant

and the pellet were cultured in 200 µl of 2YT, 50 µg/ml kanamycin and 100 µg/ml

ampicillin and cultured at 28oC overnight with shaking. The overnight culture was

centrifuged (1,800 g for 10 min) and 75 µl of the phage containing supernatant was

used in monoclonal phage ELISA.

Production of soluble scFv

The selected scFv-phage from each round of selection was inoculated into the

E. coli HB2151 non-suppressor strain to produce soluble scFv antibody. HB2151

cells were grown in 2YT media at 37oC until the OD600 was 0.4. Two hundred µl of

this culture was infected with 10 µl of scFv-phage, incubated at 37oC for 30 min

without shaking and plated on 2YT agar with 100 μg/ml ampicillin. From this plate, a

single colony was picked and inoculated into 5 ml 2YT, 1% glucose and 100 μg/ml

ampicillin, and cultured at 37oC overnight with shaking (250 rpm). The overnight

cultures were then diluted 1:20 in fresh 2YT medium containing 100 µg/ml ampicillin

and 0.1% glucose and grown at 37oC with shaking until the OD600 was 0.9. The

culture was then induced by the addition of isopropyl β-D-thiogalactopyranoside

(IPTG), to a final concentration of 1 mM. The culture was incubated at 28oC

overnight with shaking (250 rpm). The cells were centrifuged (3,300 g for 30 min)

and the soluble scFv were recovered from the pellet by osmotic shock. The pellet

was resuspended in ice-cold osmotic shock buffer [20% sucrose, 200 mM Tris-HCl

pH 7.6 and 1 mM EDTA] and incubated on ice for 5 min. The suspension was

pelleted by centrifugation in microcentrifuge (14,000 g for 5 min): the supernatant

was collected and stored (sucrose fraction), the pellet was resuspended in ice-cold

83

water, incubated on ice for 5 min, centrifuged in a microcentrifuge (14,000 g, 5 min)

and the resulting supernatant containing the periplasmic fraction was collected. All

osmotic shock fractions were either stored at 4oC and analysed immediately by

ELISA or frozen at -20oC and analysed later.

Polyclonal and monoclonal scFv-phage ELISA

Wells of a 96-well microplate were coated with 50 µg/ml of ubiquitin, 10 µg/ml of

Jgag6 or 10 µg/ml of CA-Tat polyprotein diluted in carbonate buffer (pH 9.6), 100 µl

per well, and left at 4oC overnight. The plate was washed with PBS and blocked with

200 µl of PBS/2% skimmed milk powder for 60 min at room temperature, washed

with PBS, then 100 µl of scFv-phage (about 2x1012 pfu) of either amplified

polyclonal (polyclonal ELISA) collected after each round of selection or MAb

(monoclonal ELISA) in PBS/2% skimmed milk was added and incubated at room

temperature for 60 min. PBS was used as negative control. Bound scFv-phage was

detected with the monoclonal anti-M13gp8 conjugated to HRP (Pharmacia) diluted

1:5,000 in PBS, 100 µl per well, and incubated for 60 min at room temperature. After

each incubation step, the microplates were washed 3 times with PBST and finally

with PBS. The ELISA was developed with 100 µl ABTS (BioRad) per well, and the

absorbance was read after 5 min incubation at 405 nm with a microplate reader

(BioRad).

SDS-PAGE and western blotting of soluble scFv

SDS polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting were

performed as described previously (Chapter 3). Briefly, 20 µl of all fractions of scFv

were separated by 12.5 % SDS-PAGE, and then transferred electrophoretically to

nitrocellulose membrane. The membrane was blocked with PBST containing 5%

skimmed milk powder for 1 h before washing with PBST. The membrane was then

incubated with 1:5,000 diluted HRP-conjugated anti-His antibody in 10 ml of PBST

containing 2% skimmed milk powder for 1 h at room temperature. Bound anti-His

was detected by the addition of HRP colour development reagent (Bio-Rad).

84

Results

Screening for ΔSU scFv by CLA

The panning procedure with ΔSU-GST proteins bound to a nitrocellulose membrane

was used to selectively capture ΔSU-positive recombinant phage from 2 phage

antibody display libraries (I and J). During 4 panning cycles, the number of ΔSU-

GST specific phage increased significantly from the third round (Figure 5.3).

1.00E+00

1.00E+01

1.00E+02

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1.00E+08

1 2 3 4

Panning round

Elut

ed p

hage

(cfu

/ml)

Figure 5.3. ELISA results demonstrating enrichment of ΔSU-GST scFv during panning. Four rounds of panning were conducted against ΔSU-GST proteins immobilised on nitrocellulose membranes. The bound phage from Library I (blue bars) and Library J (violet bars) after each round of selections were eluted with trypsin and were titrated. An increase in number of eluted phages could be seen as the panning progressed.

HB2151 bacteria that were infected with phage recovered after the third and fourth

panning rounds were plated on individual YTAG agar plates in different dilutions and

screened by CLA using nitrocellulose membranes coated with ΔSU-GST fusion

protein (Figure 5.2). This screening resulted in isolated colonies at a 10-4 dilution.

These single bacterial colonies were tested for their ability to bind to ΔSU-GST

immobilised on nitrocellulose membranes and a number of scFv-positive-colonies

from both phage libraries were identified (Figure 5.4).

85

Figure 5.4. Screening of diffused scFv antibodies secreted by the bacterial colonies grown on master filter to the membrane coated with recombinant protein ΔSU-GST. The scFv-antibodies were detected by anti-His antibody. (A) Antibodies identified after third round of selection and (B) antibodies identified after the fourth round of selection.

Forty single colonies obtained using CLA from the third and the fourth panning

round of libraries I and J were selected and subcultured on YTAG plates and then

tested for the production of antibody by CLA. Multiple scFv-binding spots,

corresponding to colonies, were visualised on the membrane and an example of the

results obtained during the third selection round of Library J is shown in Figure 5.5.

A few positive single colonies were also obtained using Library I.

Figure 5.5. Bacteria colonies expressing potential antibody fragments (scFv). Forty selected colonies were tested for antibody production by CLA. Antibody fragments that diffused from master filter (A) and bound to the ΔSU-GST coated filter (B) could be detected by an enzyme colorimetric reaction. The corresponding positive colonies could be identified from the master filter (A).

A B

A B

86

Four positive colonies derived from the third round of panning using the J-library

and 8 positive colonies from the fourth round of panning (4 using the J-library and 4

using the I-library) that showed a strong band on western immunoblotting, were

further subcultured and re-screened by CLA for positive scFv antibody reactions

and the results are shown in Table 5.1. Numerous positive colonies were identified

although 2 of the 12 parental clones produced a failed to react when retested after

subculture.

Table 5.1. Results of screening of parental clones with colony lift assay (CLA).

Plate Number of colonies Number of positive colonies determined by CLA

Percentage positive

3J-1 200 98 49 3J-1A 196 49 25 3J-1B 192 57 30 3J-9 140 43 31 4J-1 158 17 11 4J-2 154 Negative N/A 4J-6 260 10 4 4J-7 132 Negative N/A 4I-1 180 75 42 4I-2 284 43 15 4I-3 115 24 21 4I-4 240 24 10

Five isolated colonies from each positive plate were subcultured and 10 colonies

from each subculture (total of 50) were used to express soluble scFv antibody,

determined by indirect and capture ELISA against ΔSU-GST and GST proteins.

Most of the 50 colonies produced scFv that reacted with both ΔSU-GST and GST

by both ELISA methods (Figure 5.6) but scFv produced by 2 colonies (referred to as

clones 32 and 34) showed a relatively stronger affinity to antigen. However, there

was a strong reactivity to both the ΔSU-GST and GST indicating that the selected

scFv had affinity to the GST and not ΔSU protein.

87

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

1

1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49

clones

Abso

rban

ce

SU-GST

GST

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

1

1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49

clones

Abso

rban

ce

SU-GST

GST

Figure. 5.6. Results of indirect (A) and capture (B) ELISA of scFv from individual colonies against ΔSU-GST and GST proteins. The scFv were produced from individual colonies after detection by CLA. Fifty colonies from the 10 positive plates were analysed by indirect and capture ELISA and the bound scFv was detected with protein L-HRP. The absorbance readings shown were the average of 3 replicate reactions.

Screening for CA scFv by microplate method

Four rounds of panning were undertaken on ubiquitin, Jgag6 and CA-Tat proteins

bound to microplates. After each round of panning, the titre of the eluted phage was

measured to monitor the efficiency of the selection process (Figure 5.7). After the

third round, phage recovery increased almost 100-fold compared to the second

round of panning, indicating the library was already enriched in each protein binder.

However, during the fourth round the recovery decreased slightly in comparison to

the third round.

A

B

88

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1 2 3 4

Panning round

Phag

e (c

fu/m

l)

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1 2 3 4

Panning round

Phag

e (c

fu/m

l)

1.00E+03

1.00E+04

1.00E+05

1.00E+06

1.00E+07

1 2 3 4

Panning round

Phag

e (c

fu/m

l)

Figure 5.7. The numbers of eluted scFv-phage after each of 4 rounds of selection that bound to ubiquitin (A), Jgag6 (B) and CA-Tat polyprotein (C). The titre was determined as cfu/ml with library I (brown bars) and the library J (green bars).

The increase in the number of protein-specific binders in the phage population was

confirmed with polyclonal scFv-phage ELISA after each panning round. Higher

absorbance values were obtained with each round of panning than in the previous

round, indicating a progressive increase of the number of specific scFv-phage

A

B

C

89

binders in each panning round (Figure 5.8). The enrichment with Library J was

greater than with Library I for all 3 proteins, including ubiquitin.

To identify individual specific scFv-phage binders, the eluted scFv-phage after the

third round of selection were used for infection of TG1, which were then cultured to

obtain single colonies. Twenty colonies from each library were selected and tested

for their ability to bind to ubiquitin, Jgag6 and CA-Tat proteins with the monoclonal

scFv-phage ELISA. When the value of absorbance of all colony samples for each

binding protein was averaged, the average absorbance of scFv-phage clones for

ubiquitin, Jgag6 and the CA-Tat polyprotein were 2.6049, 2.1529 and 1.4442 for

library I and 2.4628, 2.3820 and 1.7622 for library J (Figure 5.9). Based on these

average values, colonies producing absorbance values that exceeded the average

absorbance were selected as potentially protein-specific scFv-phages (Table 5.2).

90

0

0.5

1

1.5

2

2.5

3

1 2 3 4

Panning round

Abs

orba

nce

405n

m

0

0.5

1

1.5

2

2.5

1 2 3 4

Panning round

Abs

orba

nce

405n

m

0

0.5

1

1.5

2

2.5

3

3.5

1 2 3 4

Panning round

Abs

orba

nce

405n

m

Figure 5.8. Results of polyclonal phage ELISA after each round of panning. The precipitated phages were reacted with ubiquitin (A), Jgag6 (B) or CA-Tat polyprotein (C) and detected with an anti-M13/HRP conjugated antibody at an absorbance of 405 nm. Library I (green bar) and library J (yellow bar).

Table 5.2. The number of clones (colonies) specific for each protein binder used.

Ubiquitin Jgag6 CA-Tat

Library I 15 9 7

Library J 14 15 12

A

B

C

91

Twenty selected clones reacted better to the truncated CA protein Jgag6 than the

CA-Tat polyprotein (Figure 5.9) even though the CA-Tat polyprotein contained the

full-length JDV CA, plus Tat encoded by exon 1 of the tat1 gene (Lewis, 2008).

0

0.5

1

1.5

2

2.5

3

3.5

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

Clones

Abs

orba

nce

405n

m

Ubiquitine Jgag6 Cap-Tat

0

0.5

1

1.5

2

2.5

3

3.5

4

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

Clones

Abs

orba

nce

405n

m

Ubiquitine Jgag6 Cap-Tat

Figure 5.9. The binding of 20 clones detected by a monoclonal phage ELISA to ubiquitin, Jgag6 and CA-Tat proteins. The 20 colonies (clones) from library I (top) and library J (bottom) were selected from the third round of panning.

92

In order to obtain specific scFv against the CA protein, 4 colonies of Jgag6 scFv-

phage, 2 each from each library, designated as IF3, JG8, JB1 and ID6 were

selected and used to infect non-suppressor strain E. coli HB2151 for expression of

soluble scFv. The culture was grown overnight with shaking at 28oC and expression

was induced with IPTG when the absorbance was 0.9. The distribution of the

soluble scFv in the bacterial periplasm and in the culture supernatant was evaluated

in 12.5% SDS-PAGE gels stained with Coomassie Blue and in western

immunoblots reacted with anti-His antibody (Figure 5.10). western immunoblotting

indicated that in the supernatant, the sucrose fraction, the periplasmic fraction, a

distinct band of 30 kDa within the expected molecular weight range of soluble scFv

was present and was of stronger reactivity in the sucrose and periplasmic fractions.

The results indicated that most of soluble scFv was secreted into the periplasm but

that a substantial amount of soluble scFv was also released into the medium (Figure

5.10). A major band of about 30 kDa corresponding to the monomeric form of the

scFv was also detected in the pellet fraction of JB1 scFv by Coomassie Blue

staining and western immunoblotting with anti-His antibody.

93

Figure 5.10. 12.5% SDS-PAGE (A and B) and western blot analysis (C and D) of scFv in the supernatant, sucrose fraction, periplasmic fraction and pellet of E. coli HB2151 after induction of expression with IPTG. The gels were stained with Coomassie Brilliant Blue and scFv were detected using His antibody. Lane 1, IF3 supernatant; lane 2, JG8 supernatant; lane 3, JB1 supernatant; lane 4, ID6 supernatant; lane 5, IF3 sucrose fraction; lane 6, JG8 sucrose fraction; lane 7, JB1 sucrose fraction; lane 8, ID6 sucrose fraction; lane 9, IF3 periplasmic fraction; lane 10, JG8 periplasmic fraction; lane 11, JB1 periplasmic fraction; lane 12, ID6 periplasmic fraction; lane 13, IF3 pellet; lane 14, JG8 pellet; lane 15, JB1 pellet; lane 16, ID6 pellet.

Discussion

Recombinant antibody can be produced in E. coli or other expression systems from

cloned cDNA and due to the stability of the DNA constructs and ease of production

compared with conventional monoclonal antibody technique, unlimited supplies of

specific antibodies can be produced at a reasonable cost (Foord et al., 2007)

A B

C D

75

50

37

25

75

50

37

25

75

50

37

25

75

50

37

25

Supernatant Sucrose fraction Periplasmic fraction

Pellet

1 2 3 4 5 6 7 8

1 2 3 4 5 6 7 8

9 10 11 12 13 14 15 16

9 10 11 12 13 14 15 16

94

without the need for long term storage of hybridomas. The technique does have the

disadvantage of producing only a monovalent product that may have a fast off-rate

and poor retention time on the target (Cheng et al., 2005). In addition, there are

challenges to overcome when isolating antibodies from the phage display library.

Isolation of phage displayed antibodies is dependent on a number of parameters

including the size of the library, phage preparation, conditions and time of

incubation, blocking and washing during selection, and the method of elution of

bound scFv (Chen et al., 2009).

In this current study, phage display technology was used to facilitate the selection of

the scFv against recombinant ΔSU and CA proteins of JDV using the synthetic

phagemid Tomlinson I and J libraries. Protein-specific scFv would have application

in methods for the detection of protein either by ELISA or western immunoblotting

and could have many applications in the development of diagnostic methods. This

phage display technology has not been used previously to produce antibody against

JDV proteins. The selection of the 2 JDV proteins ΔSU-GST and CA for which it

was attempted was based on the premise that they would be useful proteins to

explore as vaccines against Jembrana disease. They also have a role during the

production and purification of these proteins and could be targets of immunological

tests for the detection of virus during routine diagnosis.

Technical problems associated with the technology included problems associated

with the selection of methods used for screening. Proteins have different tendencies

of adsorption to nitrocellulose membrane and polystyrene. According to the

manufacturer’s specification for the polystyrene microplate used in this work (Nunc

MaxiSorp) the microplates have hydrophilic binding groups that attract hydrophilic

regions on molecules. Since the recombinant CA proteins used were hydrophilic

(soluble in water) they were likely to be well adsorbed onto the surface of the plate.

In contrast, the solubilised ΔSU protein was more insoluble and probably has few

hydrophilic aa residues exposed as a consequence it was likely to attach poorly to

the surface of microplates whereas the presence of hydrophobic interactions and

weak hydrogen bonds would allow its adsorption to the surface of nitrocellulose

membranes (Ahmad et al., 2009). For that reason, in this study the microtitre plate

method was used for panning and screening the phage antibody library using CA

proteins and nitrocellulose membrane was used with the ΔSU protein. The nature

of the solid binding surface, however, is not the only factor in selection of an

appropriate matrix for protein binding, and the protein structure, size of a molecule,

95

charge, density and distribution in the protein molecule, and conformational

rearrangements all must be considered (Kamyshny et al., 1999).

The production of scFv that reacted specifically with CA proteins appeared to be

successful although additional studies will be required to optimise their expression

and demonstrate their value for diagnostic purposes and in protein identification and

purification. At least 4 clones were produced that expressed soluble scFv in E. coli

and reacted with the Jgag6 ΔCA protein. western immunoblotting results indicated

the soluble scFv were expressed into the bacterial periplasm, under the culture

conditions used, although some leakage of scFv fragments to the extracellular

medium did occur. Antibody fragments expressed under the control of the lac

promoter are known to be released to the extracellular medium after several hours

of induction, even though signal sequences have directed them to the periplasm

(Shibui & Nagahari, 1992, Takkinen et al., 1991). Control of leakage would be

required to produce a single location for scFv fragments, which will improve

downstream processing for recovery. One of the Jgag6-scFv clones, JB1, did not

remain exclusively in the periplasmic space and was not released into the culture

medium: a small proportion of the JB1-scFv was also present in the pellet as

insoluble scFv. Possibly this may have been the result of a high concentration of

scFv produced in the bacterial periplasmic space where the accumulation of scFv

may then form insoluble aggregates. If this were the case, this insoluble scFv could

be solubilised with urea and refolded (as described in Chapter 3) using a buffer

containing aa, which destabilizes wrongly folded structures (Huston et al., 1994), or

its solubility could be influenced by the mutation within the framework regions of the

scFv (Duenas et al., 1995).

The expression of the scFv reactive with the JDV CA was limited and further

optimisation of expression is required. It was, however, expressed into the

periplasm which is preferred for large-scale preparation because this is of smaller

volume than the extracellular fraction. Specific release of scFv using a technique

such as osmotic shock, which would not rupture the inner cell membranes, could

offer an attractive option for purification. If, however, it was instead chosen to

promote extracellular scFv production, this might avoid the need for cell disruption.

Attempts to produce clones that would express scFv against the ΔSU protein with

the phage display library were unsuccessful. The CLA method used in this

experiment did result in the production of clones that expressed scFv that reacted

with the ΔSU-GST but analysis of the results suggested they reacted against the

GST component of the ΔSU-GST and not the ΔSU. The 26 kDa GST tag is 220 aa,

96

larger than other tags such as myc- or FLAG. This large size of GST may potentially

influence the panning or may render it more immunogenic than the target protein

ΔSU. Zhang et al. (2001) also described selection of false positives when GST

fusion proteins were used for selection of antibodies specific to viral movement

protein. A similar phenomenon was also reported by Murthy et al. (1999) during

attempts to identify a consensus motif specific for the PDZ2 domain of a cytosolic

protein tyrosine phosphatase by peptide phage display.

In conclusion, the results presented demonstrate successful production of a scFv

with the capacity to bind to CA antigens of JDV. Further development of this scFv

antibody could enable it to be used for immunodiagnosis of JDV infections and for

identification of CA proteins during future attempts to produce Jembrana disease

vaccines.

97

Chapter 6

Identification and preliminary analysis of JDV envelope glycoproteins by MALDI-TOF mass spectrometry

Summary

Matrix-assisted laser desorption/ionisation–time of flight (MALDI-TOF) mass

spectrometry of the proteins of JDV derived from infected cattle was conducted to

determine the size and potential glycosylation sites in the SU and TM glycoproteins.

Virus extracted from the plasma of JDV-infected cattle was purified and subjected to

SDS-PAGE and 2 proteins of 60 and 75 kDa were identified as potential

glycosylated proteins by staining with a Gelcode® Glycoprotein Staining Kit (Pierce

Biotechnology). MALDI-TOF/MS examination of 77 tryptic peptide fragments of the

75 kDa protein and 80 peptide fragments of the 60 kDa protein revealed significant

identity of the aa sequence of these proteins to the predicted aa sequence of JDV

SU and TM. Further analysis identified glycosylation sites in the fragments of the 2

proteins: 2 potential sites of N-linked glycosylation in the 75 kDa band and 3

fragments with 6 potential O-linked glycosylation sites in the 60 kDa protein. This

study has confirmed the identity of the 75kDa and 60kDa as SU and TM

glycoproteins, respectively.

98

Introduction

Although the predicted sizes of the non-glycosylated forms of the Env polyprotein

and the SU and TM have been determined as 88.8, 47.8 and 41.1 kDa,

respectively, and a number of potential glycosylation sites have been predicted from

analysis of the sequence (Chadwick et al., 1995b), information about the size or the

degree of glycosylation of the native forms of these proteins has not been reported.

Membrane glycoproteins are responsible for many important functional properties of

not only eukaryotic cell surfaces but also virus-cell interactions (Parekh, 1991) and

their post-translational modification by glycosylation is critical to their function. This

glycosylation is a complex process varying in different cell types and cannot be

easily predicted by simple rules. Two types of carbohydrate-protein linkages are

known: O-linked and N-linked (Medzihradszky, 2005, Peter-Katalinic, 2005).

Although some characteristics of glycosylation can be obtained by metabolic

labeling with radioactive saccharides or affinity chromatography with lectins such as

concanavalin A (Bundy & Fenselau, 1999, Bundy & Fenselau, 2001), mass

spectrometry (MS) offers several advantages and it has been used widely to

characterise biomolecules such as carbohydrates (Siemiiatkoski & Lyubarskaya,

2005), peptides and proteins (Stutz, 2005, Wysocki et al., 2005), DNA (Mauger et

al., 2006), metabolites (Xu et al., 2006), phospholipids (Pulfer & Murphy, 2003) and

also whole cells (Easterling et al., 1998). It provides a rapid method for identification

of biomolecules based upon molecular weight and offers better resolution and

accuracy than other techniques such as SDS-PAGE. In recent years, matrix-

assisted layer desorption/ionisation (MALDI) MS has been used to characterise

glycoproteins (Ayers et al., 2002, Barrientos et al., 2004, Kim et al., 2001,

Serebryakova et al., 2006). MS procedures offer the opportunity for peptide mass

fingerprinting and de novo sequencing. Peptide mass fingerprinting is a useful

technique used to identify proteins whose aa sequences are known or have been

predicted from DNA analysis of the encoding gene (Thiede et al., 2005). MALDI MS

in combination with proteolytic assays has been used to identify the functional

epitope on envelope glycoprotein gp41 of HIV-1 (Parker et al., 2001).

To provide information about the size of the native glycosylated proteins of JDV an

attempt was made to identify the proteins present in native viral preparations

obtained from the plasma of JDV-infected cattle by utilising matrix-assisted laser

desorption/ionisation–time of flight (MALDI-TOF) MS. The data generated was also

99

used to identify glycosylation sites (both N- and O-linked) as well as the types of

glycans attached to the glycosylated proteins.


Identification of potential glycoproteins in SDS-PAGE preparations of JDV

A plasma-derived preparation of JDV (30 µl) from JDV infected cattle was a kind gift

from Dr. Nining Hartaningsih (Indonesia). The plasma was prepared as described

by (Hartaningsih et al., 1994). Briefly, plasma collected from JDV-infected cattle and

containing an estimated 108 infectious virions per mL of plasma was clarified by low

speed centrifugation. The virus was then pelleted by centrifugation and the

resuspended pellet subjected to sucrose gradient centrifugation. Visible bands in

the sucrose gradient were harvested and resuspended in Tris-HCl buffer to 1% of

the original plasma volume. The partially purified virus sample obtained was

subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-

PAGE) as described previously (Hartaningsih et al., 1994) but using a 4% stacking

gel and 14% resolving gel. The protein bands were detected by staining with

Coomassie Blue R20 in 40% methanol and 10% acetic acid or with a GelCode®

Glycoprotein Staining Kit (Pierce Biotechnology) according to the manufacturer’s

instructions. This glycoprotein detection system selectively stains glycoproteins in

polyacrylamide gels using a modified Periodic Acid-Schiff (PAS) staining method

(Zacharius et al., 1969). Staining of sugar moieties of the glycoproteins yields

magenta bands with a light pink background. The Periodic Acid-Schiff reagent

stains vicinal diol groups found mainly on peripheral sugars and sialic acids and is

used as a general glycoprotein stain (Thornton et al., 1994).

Preparation of potential glycoproteins for MALDI-TOF MS analysis

Two protein bands of 75 kDa and 60 kDa visualised with both Coomassie Blue and

the Glycoprotein Staining Kit were excised using a sterile scalpel blade and placed

in separate Eppendorf tubes. Each gel segment was washed by covering it with 500

µl of PBS at 4oC for 30 min, then discarding the PBS and repeating the washing

step. The gel pieces were then washed a further 2 times with 500 µl of 100 mM

NH4HCO3 / acetonitrile (1:1, v/v) for 15 minute and dehydrated the gel in 200 µl of

100% acetonitrile. Digestion of the proteins was conducted by the addition of 200 µl

of 50 mM NH4HCO3 containing 4 ng/µl of trypsin to fully immerse the gel pieces and

100

incubated for 30 min at room temperature before the tubes were incubated

overnight at 37oC. The tubes were spun down for 10 s. The gels were washed with

200 µl of 25 mM NH4HCO3/acetonitrile and the peptides were extracted with 50 µl

of 5% formic acid (HCOOH)/acetonitrile by sonicating the gel in a ultrasonic bath for

2 min and then incubated for 30 min at room temperature. The supernatant was

transferred to a clean Eppendorf tube and dried in a speedvac. The dried peptides

were reconstituted in 20 µl of 0.5% HCOOH in 65% methanol.

MALDI-TOF MS analysis of tryptic peptides

One µl of each peptide solution was mixed with an equal volume of a saturated

solution of α-cyano-4-hydroxycinnamic acid in 0.5% HCOOH/65% methanol, and

dried at room temperature. The molecular masses of the tryptic peptides were then

determined by MALDI-TOF MS (Voyager, Applied Biosystems) with the assistance

of Proteomics International Inc. at Murdoch University. MALDI spectra were

calibrated using a peptide mixture provided by the manufacturer. Monoisotopic

peptide masses obtained from mass spectra were searched against the NCBI non-

redundant protein database (NCBInr) and MSDB databases using the MASCOT

protein identification system (Matrix Science Ltd. London, UK,

http://www.matrixscience.com). The same monoisotopic peptide masses were also

subjected to FindPept (http://au.expasy.org/tools/findpept.html) and GlycoMod

(http://au.expasy.org/tools/glycomod/) tools to predict the theoretical peptides and

the glycosylation sites, respectively. The data was also compared to the predicted

aa sequence of the JDV genomic sequence (GenBank accession no. U21603.1 gi:

733067) and the JDV Env precursor protein sequence (accession number gi:

733070). This Env precursor is assumed to be post-translationally processed to

produce the SU and TM proteins (Chadwick et al., 1995b).

Results

Identification of glycoproteins in SDS-PAGE gels

When the native JDV proteins were separated by SDS-PAGE and stained with

Coomassie Blue, 4 relatively abundant proteins with a MW of >41.1 kDa, the

predicted size of the smallest non-glycosylated envelope protein (Chadwick et al.,

1995), were identified with a MW of approximately ~250, ~75, ~60 and ~50 kDa

(Figure 6.1). When stained with the GelCode® Glycoprotein Staining Kit (Pierce

101

Biotechnology), magenta bands with a light pink background indicated 3 of these

proteins were glycosylated. The protein with a predicted mass of ~75 kDa was the

most abundant while the 2 bands at ~250 kDa and ~60 kDa stained weakly.

Figure 6.1. Coomassie Blue (RHS) and GelCode® Glycoprotein Staining (LHS) of JDV proteins in 14% SDS-PAGE gels. The protein markers and their size are shown in the extreme LHS of the gel.

MALDI-TOF/MS analysis

Two potentially glycosylated protein bands of MW 75 and 60 kDa (Figure 6.1) were

excised from the SDS-PAGE gel and subjected to in-gel digestion with trypsin,

which cleaves after lysine and arginine residues, and the peptide digests were then

analysed by MALDI-TOF MS. The m/z spectra of the mixture of peptides resulting

from tryptic digestion of the 75 kDa proteins and the 60 kDa protein are shown in

Figures 6.2 and 6.3.

35

MW

75

50

100 150 250

~75 kDa

~60 kDa

~250 kDa

~50 kDa

102

Figure 6.2. Single MALDI-TOF/MS spectrum of the peptide mixture obtained from a tryptic digests of the 75 kDa protein. The labels associated with the peaks indicate the measured mass for the tryptic peptides. The ordinate axis represents the intensity of the detection signal of peptide detected by the MS at a specific mass to charge (m/z).

Figure 6.3. MALDI-TOF/MS spectra of a tryptic digests of 60 kDa protein excised from a preparative PAGE gel. The peak labels indicate the measured mass (m/z) for all the tryptic peptides that contribute to the whole protein.

103

The sequence of the monoisotopic peptide masses were examined for sequence

similarity to proteins in the MSDB and NCBInr databases using the MASCOT

search program. With this program, the protein hits were scored using a probability

based Mowse (molecular weight search) score. The ion score is –10*Log (P), where

P is the probability that the observed match is a random event. Individual ions that

scored more than 51 indicated identity or extensive homology (p<0.05). The scores

obtained ranged from 24 to 40, so that neither protein was identified as having a

significant match to any protein in the MSDB database. However, the proteins with

greatest identity to the 75 kDa peptides in the NCBInr database were envelope

glycoproteins of HIV-1, Saimiriine herpesvirus-2 and the envelope glycoprotein of

Caprine arthritis-encephalitis virus (Table 6.1). Proteins of greatest identity with the

60 kDa protein in the NCBInr database were of Porcine reproductive and respiratory

syndrome virus, the envelope glycoprotein of HIV-1 and Hepatitis C virus (Table

6.2). The databases did not contain data for any bovine lentivirus protein.

Table 6.1. Comparison of peptides found in tryptic digests of the 75 kDa protein with proteins in the NCBInr database using the MASCOT search program. Only Mowse scores of more than 51 indicate identity or extensive homology. Mowse score

Protein Virus Molecular weight

Peptide sequence

30 Envelope glycoprotein gp120

HIV-1 826.42 QPVSKLR


HIV-1 1035.53 KLSDQFGNK


HIV-1 939.41 KSINIGPGR

25 Glycoprotein Saimiriine herpesvirus-2

1414.78 VITSSLQTTSSYK

24 Glycoprotein 120

HIV-1 516.51 VGLTK

24 Surface glycoprotein

Caprine arthritis-encephalitis virus

516.51 VIGTK

104

Table 6.2. Comparison of peptides found in tryptic digests of the 60 kDa protein with proteins in the NCBInr database using the MASCOT search program. Only Mowse scores of more than 51 indicate identity or extensive homology. Mowse score

Protein name Species Molecular Weight

Peptide Sequence

40 Polypeptide precursor NSP1 – NSP8

Porcine reproductive and respiratory syndrome virus

821.50 VAAEIYR

34 Envelope glycoprotein, V3-V5 region

HIV-1 913.50 DPITLPCR

29 Reverse transcriptase

HIV-1 1069.53 WGFYTPDGK

29 Polyprotein Hepatitis C virus

989.54 TRVTGGSSAR

The peptide mass fingerprinting data obtained by MALDI-TOF MS were compared

with the theoretical peptides calculated by Findpept software program. The

FindPept program deduced the theoretical peptides that would be obtained by

trypsin digestion of the aa sequence of the envelope proteins of JDV derived from

the DNA sequence and then calculated the theoretical masses of the fragments.

The 75 kDa peptide mass fingerprinting data generated by MALDI-TOF/MS resulted

in masses for 77 peptides, 66 of which matched the theoretical peptide masses of

JDV SU (aa 1-422) calculated by the FindPept program and 11 were unmatched

and assumed to be glycosylated (Figure 6.4 and Table 6.3). The 60 kDa peptide

mass fingerprinting data generated by MALDI-TOF/MS resulted in a mass for 94

peptides, 80 of which had high sequence identity to the deduced masses of the

peptides of JDV TM (Env aa 423-781) calculated by the FindPept program and 14

were unmatched (Figure 6.5 and Table 6.4) and were assumed to be glycosylated.

105

Table 6.3. Comparison between the masses of experimentally observed with the theoretical masses and assignment of the tryptic peptide mass fingerprint MALDI-TOF/MS of the 75 kDa protein.

Peptide m/z (detected)

m/z (calculated)

Residue position Peptide sequence

P1 1649.77 1648.72 1-13 MMEEGRKEEPEER P2 1192.58 1191.60 13-22 RGEKSTMRDL P3 1589.73 1588.84 17-30 STMRDLLQRAVDKG P4 2095.10 2094.11 25-42 RAVDKGHLTAREALDRWT P5 2338.37 2337.16 43-63 LEDHGEIHPWIILFCFAGAIG P6 1739.02 1737.95 59-76 AGAIGVIGGWGLRGELNV P7 2556.33 2555.44 71-93 RGELNVCMLIVLVVLVPIYWGIG P8 2247.07 2246.20 94-111 EAARNIDSLDWKWIRKVF P9 1445.65 1444.75 98-108 NIDSLDWKWIR P10 2440.29 2439.38 111-133 FIVIIFVLVGLLGGCSAQRQHVA P11 1595.89 1594.88 134-148 MLLSPPGIRLPSTVD P12 1633.82 1632.83 142-155 RLPSTVDIPWFCIS P13 2408.21 2397.12 152-172 FCISNAPIPDCVHWTVQKPDQ P14 2094.22 2093.02 169-185 KPDQKHQQIENVMELQE P15 1036.55 1035.49 184-192 QEVLDNATF P16 1038.55 1037.51 190-198 ATFFEVPDL P17 1266.59 1265.63 196-205 PDLFDRVYLE P18 2220.03 2219.19 206-227 LARLDANSTGVPVNIPPTGISQ P19 2435.29 2434.28 208-231 RLDANSTGVPVNIPPTGISQVKGD P20 2398.20 2397.09 227-249 QVKGDCSTGDIQGMNETLSTRGT P21 1034.54 1033.54 243-252 TLSTRGTLGE P22 1475.85 1474.77 251-263 GERTFLSIRPGGW P23 1783.92 1782.80 262-275 GWFTNTTVWFCVHW P24 1665.77 1664.80 269-281 VWFCVHWPFGFIQ P25 1019.53 1018.50 282-290 RKENLSEGS P26 2511.26 2510.25 285-307 NLSEGSAQVRNCLDPINVTEPRV P27 1412.69 1411.67 301-312 NVTEPRVANYSY P28 1634.81 1633.78 307-320 VANYSYCPLEYKGK P29 1635.81 1634.86 321-335 NYINKGLKCVGGRVD P30 1226.58 1225.56 334-344 VDLSSNPEQHT P31 2408.03 2407.02 345-365 DLLACGTFCQNFRNCDMVSRD P32 1635.76 1634.75 358-371 NCDMVSRDILIGYH P33 1485.80 1484.72 369-380 GYHPSQQKQHIY P34 1052.54 1051.56 376-383 KQHIYINH P35 1635.82 1634.81 384-396 TFWEQANTQWILV P36 1855.92 1855.00 392-407 QWILVQVPNYGFVPVP P37 2527.38 2526.36 401-422 YGFVPVPDTERPWKGGKPRGKR

106

10 20 30 40 50 MMEEG RKEEP EERGE KSTMR DLLQR AVDKG HLTAR EALDR WTLED HGEIH 60 70 80 90 100 PWIIL FCFAG AIGVI GGWGL RGELN VCMLI VLVVL VPIYW GIGEA ARNID 110 120 130 140 150 SLDWK WIRKV FIVII FVLVG LLGGC SAQRQ HVAML LSPPG IRLPS TVDIP 160 170 180 190 200 WFCIS NAPIP DCVHW TVQKP DQKHQ QIENV MELQE VLDNA TFFEV PDLFD 210 220 230 240 250 RVYLE LARLD ANSTG VPVNI PPTGI SQVKG DCSTG DIQGM NETLS TRGTL 260 270 280 290 300 GERTF LSIRP GGWFT NTTVW FCVHW PFGFI QRKEN LSEGS AQVRN CLDPI 310 320 330 340 350 NVTEP RVANY SYCPL EYKGK NYINK GLKCV GGRVD LSSNP EQHTD LLACG 360 370 380 390 400 TFCQN FRNCD MVSRD ILIGY HPSQQ KQHIY INHTF WEQAN TQWIL VQVPN 410 420 YGFVP VPDTE RPWKG GKPRG KR Figure 6.4Interpretation of the MALDI peptide mass fingerprint results of the 75 kDa protein. The arrows indicate the tryptic peptide fragment (P1 – P37) produced by MALDI-TOF/MS.

P2

P

P4 P5

P6 P7

P9

P8 P10 P11

P12

P13

P14 P15

P16

P17

P19

P18 P20

P21

P22 P23 P24

P27

P25

P26

P28 P29 P30 P31

P32 P33

P34 P35

P36

P37

107

Table 6.4. Comparison of the masses of the experimentally produced peptides of the 60 kDa protein generated by MALDI-TOF/MS with the theoretical masses of the tryptic peptide mass fingerprint generated by FindPept software.

Peptide m/z (detected)

m/z (calculated)

Residue position Peptide sequence

P1 1358.77 1357.84 423-435 AVGMVIFLLVLAI P2 1479.85 1478.82 429-442 FLLVLAIMAMTASV P3 1036.60 1035.53 438-448 MTASVTAAATL P4 1209.69 1208.69 442-453 VTAAATLVKQHA P5 1567.81 1566.85 452-466 HATAQVVGRLSTNLT P6 1194.67 1193.67 463-472 TNLTYITKIQ P7 1283.75 1282.69 467-476 YITKIQNQYL P8 1355.83 1354.71 477-487 HLFQNLNTRVN P9 1266.66 1265.66 487-496 NNLHHRVTYL P10 927.53 926.49 492-498 RVTYLEF P11 1855.11 1853.95 498-514 FLAEVHEVQTGLGCVPR P12 1249.70 1248.58 512-521 VPRGRYCHFD P13 2435.29 2434.08 518-537 CHFDWRPEEVGLNMTLWNST P14 1567.70 1566.69 533-544 LWNSTTWQQWMS P15 1490.77 1489.62 540-550 QQWMSYYDQIE P16 804.31 803.34 550-555 EENIWN P17 1423.60 1422.69 554-564 WNLKYNWSEAL P18 1789.96 1788.84 561-577 SEALEKGKSNTDGLEPD P19 1311.71 1310.61 571-581 TDGLEPDVFRY P20 1469.68 1468.72 576-588 PDVFRYLADLSSS P21 2045.11 2044.01 587-603 SSFTWGSWVDKLVWLAY P22 1866.15 1865.05 594-608 WVDKLVWLAYILLAY P23 2393.46 2392.27 600-619 WLAYILLAYFAFKVLQCIMS P24 1597.90 1596.78 616-629 CIMSNLGAQTRYQL P25 1633.89 1632.77 626-639 RYQLLNAQEDTDPA P26 832.34 831.32 633-640 QEDTDPAG P27 1471.58 1470.56 640-653 GDGDQPDDHRSGDT P28 1398.73 1397.63 646-658 DDHRSGDTPRSGV P29 833.34 832.37 656-664 SGVPSGGWS P30 1247.73 1246.67 662-672 GWSQKLSEGKK P31 1732.98 1731.93 668-682 SEGKKIGCLILRTEW P32 1019.56 1018.46 680-686 TEWQNWR P33 1800.10 1799.00 685-698 WRNDLRTLRWLTLG P34 1640.01 1639.03 689-711 LRTLRWLTLGGKIL P35 1293.79 1292.84 699-710 GKILQLPLSLLV P36 1485.88 1484.98 710-722 VLLVRILLHILSP P37 1220.66 1219.57 721-730 SPTFQNQRGW P38 1644.83 1643.82 731-746 TVGRKGTGGDDRELSP P39 1097.52 1096.51 747-755 ELEYLSWTG P40 1926.03 1924.87 753-768 WTGSSQEMVEMRDLKE P41 993.61 987.44 766-774 KEEDIPEE P42 1059.55 1058.51 773-781 EEGIRPVEM

108

430 440 450 ***** ***** ***** ***** **AVG MVIFL LVLAI MAMTA SVTAA ATLVK 460 470 480 490 500 QHATA QVVGR LSTNL TYITK IQNQY LHLFQ NLNTR VNNLH HRVTY LEFLA 510 520 530 540 550 EVHEV QTGLG CVPRG RYCHF DWRPE EVGLN MTLWN STTWQ QWMSY YDQIE 560 570 580 590 600 ENIWN LKYNW SEALE KGKSN TDGLE PDVFR YLADL SSSFT WGSWV DKLVW 610 620 630 640 650 LAYIL LAYFA FKVLQ CIMSN LGAQT RYQLL NAQED TDPAG DGDQP DDHRS 660 670 680 690 700 GDTPR SGVPS GGWSQ KLSEG KKIGC LILRT EWQNW RNDLR TLRWL TLGGK 710 720 730 740 750 ILQLP LSLLV LLVRI LLHIL SPTFQ NQRGW TVGRK GTGGD DRELS PELEY 760 770 780 LSWTG SSQEM VEMRD LKEED IPEEG IRPVE M Figure 6.5. Interpretation of the MALDI peptide mass fingerprint results of the 60 kDa protein. The arrows indicate the tryptic peptide fragment (P1 – P42) produced by MALDI-TOF/MS.

Glycoprotein analysis

The GlycoMod program was used to predict the mass of the glycopeptides and the

numbers of glycosylation sites obtaining from the tryptic digest of the 75 and 60 kDa

proteins. Analysis of the 11 unmatched masses of the 75 kDa protein predicted

there was one glycosylated peptide of 2013.98 Da with 2 potential sites of N-linked

P33 P32

P20

P16

P6

P5

P3

P4

P11

P12

P13

P7 P8 P9

P10

P14

P15

P19

P26

P25

P2 P1

P18 P17

P21 P22

P23

P24 P27

P28

P31 P30

P29

P37 P39 P38

P40 P41

P42

P35 P36

P34

109

glycosylation at asparagine (N) residues at aa 285 and 301 (Table 6.5). Analysis of

the 14 unmatched masses of the 60 kDa protein predicted there were 3

glycosylated peptides (1596.01 Da, 1802.11 Da and 1894.13 Da) and 6 potential

sites of O-linked glycosylation at serine residues at aa positions 650, 668 and 707,

and at threonine (T) residues at aa positions 653, 737 and 691 (Table 6.6).

Table 6.5. Prediction of unmatched masses of 75 kDa protein containing the motif 'N-X-S/T/C (X not P)'. Coloured (red) letters indicates the predicted position of glycosylated aa.

Peptide mass (exp.observed)

Peptide sequence Peptide mass (calc.)

Δm or predicted residual carbohydrate mass

Predicted carbohydrate side chain structure

2013.98 284-294 ENLSEGSAQVR

1188.57 825.41 1. (Hex)1 (NeuAc)2 (Sulph)1

2. (Hex)1 (NeuAc)2 (Phos)1

295-306 NCLDPINVTEPR

1369.67 644.31 (Hex)1 (HexNAc)1 (Deoxyhexose)1 (Pent)1

110

Table 6.6. Predictions of unmatched masses of 60kDa protein containing S or T. Coloured (red) letters indicate the predicted position of glycosylated amino acids. Peptide mass (exp.observed)

Peptide sequence Peptide mass (calc.)

Δm or predicted residual carbohydrate mass

Predicted carbohydrate side chain structure

1596.01 650-655 SGDTPR

631.29 964.72 (Pent)1 (Sulph)6 (HexA)2

736-742 GTGGDDR

676.28 919.73 (NeuGc)1 (Pent)1 (Sulph)6

1802.11 650-655 SGDTPR

631.29 1170.82 1. (Pent)2 (Sulph)6 (KDN)1 (HexA)1

2. (Hex)1 (Deoxyhexose)1 (Pent)1 (Sulph)6 (KDN)1

3. (Pent)2 (Phos)6 (KDN)1 (HexA)1

667-671 LSEGK

532.29 1269.82 1. (Deoxyhexose)1 (NeuAc)1 (Sulph)6 (HexA)2

2. (HexNAc)1 (Deoxyhexose)1 (Pent)2 (Sulph)6 (HexA)1

701-714 ILQLPLSLLVLLVR

1589.06 213.05 1. (Pent)1 (Sulph)1

2. (Pent)1 (Phos)1 736-742

GTGGDDR 676.28 1125.83 1. (Hex)1 (NeuGc)1 (Sulph)6

(HexA)1

2. (Hex)1 (NeuGc)1 (Phos)6 (HexA)1

1894.13 650-655 SGDTPR

631.29 1262.84 (HexNAc)1 (NeuGc)1 (Sulph)5 (HexA)2

667-671 LSEGK

532.29 1361.84 (Hex)2 (NeuGc)1 (Sulph)6 (KDN)1

691-693 TLR

388.24 1505.89 1. (NeuAc)1 (Pent)1 (Sulph)6 (KDN)1 (HexA)2

2. (Deoxyhexose)2 (NeuGc)1 (Sulph)6 (KDN)1 (HexA)1

3. (Hex)1 (Deoxyhexose)1 (NeuAc)1 (Sulph)6 (KDN)1 (HexA)1

4. (Hex)2 (HexNAc)1 (Deoxyhexose)1 (Sulph)6 (HexA)2

5. (Deoxyhexose)1 (NeuGc)1 (Pent)3 (Sulph)6 (HexA)1

6. (HexNAc)1 (Deoxyhexose)1 (Sulph)6 (KDN)2 (HexA)1

7. (Hex)1 (NeuAc)1 (Pent)3 (Sulph)6 (HexA)1

8. (HexNAc)1 (Pent)3 (Sulph)6 (KDN)1 (HexA)1

9. (NeuAc)1 (Pent)1 (Phos)6 (KDN)1 (HexA)2

Carbohydrate symbols used are: Hex, hexose; Pent, Pentose; Sulph, sulphate; HexA, β-hexosaminidase A; NeuAc, N-acetylneuraminic acid; NAc, N-acetyl; NeuGc, N-glycolylneuraminic acid; KDN, 2-keto-3-deoxy-nonulosonic acid.

111

Discussion

The HIV envelope glycoproteins have a central role in the induction of the host

immune response and are targets for protective immunity (Bukrinskaya, 2004, Chan

& Kim, 1998, Gummuluru & Emerman, 2002) and it is assumed that this would also

be true for JDV. Some preliminary study of the role of the JDV envelope proteins as

potential vaccine immunogens has been undertaken utilizing recombinant SU and

TM expressed in a bacterial expression system, and hence with non-glycosylated

forms of these proteins (Ditcham, 2007), with minimal effect. Further work with

these proteins as potential vaccines requires an improved understanding of the

comparative structure of the non-glycosylated recombinant JDV proteins and the

structure and glycosylation characteristics of the native proteins. While the size of

the JDV Env polyprotein has been predicted (781 aa), this is considerably smaller

than that of the other lentivirus envelope glycoproteins including the other bovine

lentivirus BIV (904 aa) (Garvey et al., 1990), HIV (856 aa) (Martoglio et al., 1997),

SIV (854 aa) (Fomsgaard et al., 1991), FIV (856 aa) (Olmsted et al., 1989) and

EIAV (859 aa) (Rushlow et al., 1986) and the predicted structure of those envelope

proteins is varied and it is possible that these differences could be even greater for

JDV.

The use of peptide mass fingerprinting with the MALDI-TOF system, whereby the

proteins were digested into smaller peptide fragments and then the resulting

peptides were compared to those in protein databases, enabled identification of 2

native JDV proteins, one of 75 kDa and the other of 60 kDa, as the SU and TM

proteins, respectively. Analysis of peptide mass fingerprinting data enabled 66 of 77

peptides of the 75 kDa protein to be matched to the JDV Env aa positions 1-422

(SU region) and 80 of 94 peptides of the 60 kDa protein matched to the JDV Env aa

positions 423-781 (TM region). A search of the NCBInr database using Mascot

software also revealed similarities of both proteins to other viral envelope

glycoproteins, particularly lentiviral proteins, but there were no definitive matches as

the scores obtained were less than 51. However, none of the other databases

searched contained data for JDV proteins.

Further analyses of the 2 proteins using GelCode® Glycoprotein Staining Kit

suggested that the 75 and the 60 kDa proteins were glycosylated and the

glycosylation of these 2 proteins was confirmed by analysis of the MALDI-TOF data.

One peptide fragment of the JDV 75 kDa SU was identified that had 2 potential sites

of N-linked glycosylation. Three peptide fragments the 60 kDa TM protein were

112

identified with a total of 6 potential O-linked glycosylation sites. A lower number of

potential glycosylation sites in the JDV Env than in other lentiviruses was predicted

by examination of the sequence data (Chadwick et al., 1995b) and the low number

of glycosylation sites identified by MALDI-TOF analysis was consistent with this

analysis. The position and number of potential N- and O-glycosylation sites in the

JDV 75kDa and 60 kDa proteins seems quite different to that in the envelope

glycoprotein of the other lentiviruses. The envelope proteins of HIV-1 contain

approximately 30-38 potential asparagine-linked (Asn-X-Ser/Thr) glycosylation sites

(Ratner, 1992). HIV-1 Env gp120 contains 18 to 33 potential N-glycosylation sites

(Korber et al., 2001) and is among the most heavily glycosylated envelope proteins

of any retrovirus. HIV-1 has also 3-6 potential N-glycosylation sites in the TM

protein, the number varying in different strains of the virus (Dedera et al., 1992).

Potential O-glycosylation sites in different strains of HIV-1, HIV-2, and SIV were

examined by Hansen et al. (1998) and demonstrated that SIV Env (gp110)

contained more O-glycosylated (4–18 sites) than HIV-1 or HIV-2 (less than 5 sites).

FIV has 20 potential N-linked glycosylation sites present in the deduced Env aa

sequence (Olmsted et al., 1989).

The probable structures of the carbohydrate side chain at the glycosylation site

predicted with the GlycoMod program used for the analysis were heterogeneous

and possibly complex. It might be possible to use the molecular mass of

carbohydrate side chain to search for their probable glycan structures, thus

providing useful information towards the structural characterization of JDV

glycoproteins. Other analytical methods such as complete cleavage of the

oligosaccharide chains from the glycoproteins and mass spectrometry analysis of

each purified oligosaccharide after separation of the carbohydrate mixture by high

performance liquid chromatography would be required for the complete

characterisation of the glycan structure.

113

Chapter 7

General discussion

Jembrana disease is a disease specific to the Indonesian archipelago and there are

2 major reasons for this: the geographic isolation of Indonesia and limited trade in

live cattle from Indonesia to other countries, and the specificity of the disease for

Bali cattle. One consequence of this is that research on the disease, especially

studies involving the use of live animals, has been restricted to Indonesia and a

small research group only. While our understanding of Jembrana disease and has

improved markedly since the recognition that a lentivirus was the aetiological agent

(Chadwick et al., 1995a), progress has been restricted by infrastructure and funding

restraints.

Attempts to control the disease by limiting the movement of Bali cattle from Bali

have slowed the spread of the disease from Bali to other islands but despite these

restrictions it has also occurred on the islands of Java, Sumatra and Indonesian

Borneo (Kalimantan). How this occurred is not fully understood and documented but

it may well have, at least in the case of Java and Sumatra, involved the illegal

movement of virus-infected Bali cattle from Bali. Future control of Jembrana disease

will likely require more than quarantine, and will need the development of improved

diagnostic and immunosurveillance procedures to monitor the distribution of the

disease and provide rapid diagnosis, and the development of an effective vaccine.

To support the development of improved diagnostics and vaccines, an improved

understanding of native JDV proteins and their production using recombinant

techniques is required. The studies reported in this thesis have helped to improve

our knowledge in this area. In this current study, only selected proteins were

examined: the CA, the major and immunodominant protein of JDV that is likely to be

a key protein in the development of diagnostics, and the 2 envelope proteins SU

and TM involved in attachment and entry of virus into cells. There is remarkably little

information available about the envelope glycoproteins of JDV, not even

fundamental characteristics of the size of the native proteins and the degree and

type of glycosylation. The studies reported in this thesis have added to our

understanding of these glycoproteins and will facilitate their future use as antigens

and vaccines.

114

For many years in this laboratory, recombinant JDV proteins and including CA and

the envelope proteins SU and TM have been produced as fusions to GST in E. coli

under the control of the tac promoter and used as antigens for ELISA and western

blotting procedure for detection of antibody (Burkala et al., 1999, Burkala et al.,

1998, Desport et al., 2005). Cloning and expression of recombinant protein in E. coli

has been favoured because this bacterium has relatively simple genetics, a rapid

growth rate and it is well characterised. The methodology needed for cloning and

expression of proteins is relatively simple compared to many other expression

systems, requiring only basic equipment, enabling it to be used in Indonesia. Most

JDV recombinant proteins were expressed, however, in the form of insoluble

proteins within inclusion bodies which has limited their use in serological assays and

in experimental vaccines.

One of the aims of this thesis was to establish a simple method to solubilise the

inclusion body proteins while retaining their antigenicity. The studies reported in

Chapter 3 showed that SU-GST could be reliably and efficiently expressed in E. coli

as inclusion bodies with good yields. The solubilisation of the SU-GST inclusion

bodies was achieved with a combination of a low concentration of urea in an

alkaline solution, with the addition of the reducing agent dithiothreitol (DTT) and the

serine protease inhibitor phenylmethanesulfonyl fluoride, providing an acceptable

yield of denatured protein for purification. The solubilised SU-GST protein was

refolded by slow 10-fold dilution in 20 mM Tris pH 8.0 containing CHAPS and DTT

and the refolded protein reacted in western immunoblots with bovine antisera. The

technique developed for solubilisation of the proteins present in inclusion bodies is

simple and the technology has been successfully used in Indonesia for the

solubilisation not only of recombinant SU but also for the solubilisation of other JDV

proteins present in inclusion bodies, including CA, Tat and TM. These solubilised

proteins have been used successfully for the development of serological antigens

and experimental vaccines (Ditcham, 2007, Lewis, 2008). The technique is a

significant advance in the preparation of antigens and vaccines for JDV and

removes the dependency on the need for expression of soluble proteins, which is

always difficult using such bacterial expression systems.

The addition of the reducing agent DTT seemed to be important in the solubilisation

process, probably because of the presence of disulfide bonds in the proteins. The

SU protein of JDV contains 13 cysteine residues predicted to form 5 disulfide bonds

(Chapter 6) and supposing that the aggregation of the protein is mainly caused by

the creation of the intra- and interchain disulfide bridges, resulting in formation of

115

disulfide bonded aggregates. The addition of DTT probably prevented aggregation

by inhibiting the formation of non-native disulfide bonds and dissolve disulfide

bonded aggregates (Rothel et al., 1997).

During western immunoblotting procedures for the detection of antibody against

JDV, reliance has been placed on the detection of the immunodominant 26 kDa CA

protein as many of the other proteins that are observed with whole JDV

preparations have not been identified (Hartaningsih et al., 1994, Kertayadnya et al.,

1993). In Chapter 6, a study utilising a specific glycoprotein staining method and

MALDI-TOF mass spectrometric analysis identified the 2 JDV envelope

glycoproteins SU and TM in SDS-PAGE gels as being of 75 and 60 kDa,

respectively. Identification of the size of these glycosylated proteins in SDS-PAGE

gels will permit their use for better interpretation of western immunoblotting results

with sera from JDV-infected cattle.

The MALDI-TOF mass spectrometric analysis of the glycosylated Env proteins SU

and TM (Chapter 6) also provided information on the secondary glycan structure of

these glycoproteins. MALDI-TOF mass spectrometry has proven to be a method of

choice for glycoprotein analysis because it is one of the most sensitive methods for

the analysis of glycoproteins (de Laurentiis et al., 2006). The GlycoMod method of

determining the mass value from MALDI-TOF/MS data provided initial

characterisation of the sugar moiety decorating at least 2 putative N-glycosylation

site of the JDV-SU and 6 putative O-glycosylation sites of the JDV-TM.

To assist in the characterisation of JDV proteins and to provide methods for the long

term production of MAbs in Indonesia, their production by conventional and

recombinant techniques was investigated in Chapters 4 and 5 of this thesis.

Monoclonal antibodies have been used widely for diagnostic purposes for many

conditions and infections and for many years a JDV CA MAb BC10 (Kertayadnya et

al., 1993) produced in Indonesia provided an invaluable reagent in

immunoperoxidase assays for the detection of JDV in tissues and as a control

reagent in western immunoblotting. Unfortunately, the hybridoma secreting this MAb

was lost when liquid nitrogen supplies in which it was maintained were lost. During

the current study, an alternative hybridoma was produced (Chapter 4) against the

JDV CA which was of IgG2 isotype and this was identified as binding to the aa

encoded by nucleotides 604-810 of the JDV capsid gene.This MAb was

successfully used to identify JDV in formalin-fixed paraffin embedded tissue

sections. Attempts were made to also produce a monoclonal against a recombinant

SU but these attempts were unsuccessful. The attempts to produce a SU MAb

116

were undertaken prior to the identification of the whole virus SU and additional

attempts using the glycosylated whole virus SU may have been worthwhile.

To overcome the problem of storage of hybridomas in liquid nitrogen in Indonesia,

the use of recombinant antibody fragments was investigated as a means of

providing a solution to the problem of long-term storage. These attempts to produce

recombinant MAb (Chapter 5) utilised the phage display antibody method because

the technique has been previously used for generating recombinant antibodies

against the TM glycoprotein gp46 of Maedi visna virus (Blazek et al., 2004) and CA

(Celer et al., 2003). Recombinant antibody fragments, scFv, with specificity for the

CA protein of JDV were generated using the Tomlinson I and J phage display

antibody libraries. Attempts were also made to produce scFv against a ΔSU with a

GST fusion tag but the only scFv detected were against the GST component. Using

an E. coli expression system, soluble scFv against the CA was expressed in the

bacterial periplasm but it was produced in small quantities only. While clones are

now available as a consequence of these studies that will allow the expression of

CA scFv, further work is now required to optimise this system and obtain adequate

stocks for further investigation, perhaps by modifying the bacterial growth conditions

or modifying the method of expression. The success with the method shows that it

does offer the potential for the production not only of CA scFv but also those against

other viral proteins. The main difficulty and most time consuming part of attempts to

produce scFv with the Tomlinson I and J libraries was the panning process. It is

likely that with experience, greater efficiency of the phage technique could be

obtained. Another problem was an apparent low affinity of the scFv to the JDV ΔSU

and CA proteins and this is a noticeable characteristic of scFv isolated from this

library (Kjaer et al., 2001, Wang et al., 2004) that can be improved by introducing

diversity to the V genes of the original antibody. Diversity can be introduced by

random mutations either by error-prone PCR (Meyer et al., 2002) or by applying

bacterial mutator strains (Coia et al., 2001).

117

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IMPROVED METHODS FOR PRODUCTION AND … · 2010. 12. 21. · Jembrana disease is an acute disease of Bali cattle (Bos javanicus) in Indonesia caused by Jembrana disease virus (JDV),