Bachelor of Applied Science (Honours)
Developing a Cell Culture System for Lagoviruses
Name: Roxanne Orford-Dunne
Student ID: 3043514
Supervisors: Dr Michael Frese, Dr Tanja Strive and Dr Markus Matthaei
Research Faculty: CSIRO Ecosystem Sciences
Clunies Ross Street, Black Mountain,
ACT 2601, Australia
Due Date: 21st of March 2012
Acknowledgements
I would like to thank Dr Michael Frese, Dr Tanja Strive and everyone from the CSIRO who
helped people during my studies. I would also like to thank Georg Koch for kindly suppling
me with the pGL3-Mx1P-Luc plasmid and Dr Michelle Gahan and Dr Peter Kerr for marking
my thesis. I would especially like to thank Dr Markus Matthaei for all the hours he put in to
teaching and helping me during my nine months at the CSIRO.
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Abstract
Rabbit haemorrhagic disease virus (RHDV) is a positive-stranded RNA virus belonging to the
Caliciviridae family and is used as a biological control agent to contain feral rabbit
populations in Australia. After RHDV’s initial introduction a significant decrease in rabbit
numbers (up to 90%) was observed, which allowed for the regeneration of many
endangered species detrimentally affected by the extreme rabbit infestation in Australia.
Unfortunately, rabbit numbers are steadily increasing, indicating a recently decreasing
effectiveness of the introduced RHDV strain (Czech strain V351). The underlying reasons for
the decreasing effectiveness of RHDV are still a matter of debate, partially due to a
significant lack of knowledge of RHDV biology as a consequence of the absence of a stable
cell culture system. Approaches to generate a stable cell culture system to study RHDV in
vitro have been unsuccessful so far, but recent advancements in calicivirus research indicate
that interferon (IFN)-induced anti-viral defence mechanisms may hinder effective calicivirus
growth.
Hence, we hypothesised that a type I IFN defective cell line may support RHDV propagation
in cell culture. To develop an IFN-defective cell line, we generated plasmids containing a
lethal gene under control of a type I IFN-inducible promoter to select for cells unable to
mount an IFN response from populations of otherwise IFN responsive cells.
To further study the interplay between RHDV and cellular innate immune responses, we
also established real time-PCR (rt-PCR) assays to measure the induction of several type I IFN-
inducible genes. We further developed an immunofluorescence assay to also examine the
expression of type I IFN-induced antiviral proteins. Both assays were shown to specifically
measure either IFN induced gene expression or IFN induced protein expression in RK13 cells.
The generated plasmids will be invaluable tools for future experiments to develop type I IFN
defective cell lines that may support RHDV growth in vitro. Furthermore, the assays
established are a necessity to study the interactions of RHDV and cellular innate immune
responses, which may critically determine virus growth and pathogenicity. While the assays
are rabbit specific, the plasmids can be used to generate IFN-defective cell lines from a wide
variety of mammalian species, potentially complementing attempts to establish cell culture
systems for other viruses as well.
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Table of contents
1 Introduction ........................................................................................................................ 1
1.1 The impact of feral rabbit populations in Australia .................................................... 1
1.2 Use of viral biological control agents in Australia ....................................................... 2
1.3 Decreasing effectiveness of RHDV on rabbits in Australia.......................................... 3
1.4 Caliciviridae biology ..................................................................................................... 4
1.4.1 Taxonomy ............................................................................................................. 5
1.5 Rabbit haemorrhagic disease virus ............................................................................. 7
1.5.1 Molecular characteristics ..................................................................................... 7
1.5.2 Pathology ............................................................................................................. 9
1.6 The role of the innate immune system in the prevention of viral replication ......... 10
1.6.1 Important innate immune system receptors .................................................... 11
1.6.2 Interferons ......................................................................................................... 19
1.7 Cell culture systems for caliciviruses ......................................................................... 25
1.7.1 Porcine enteric calicivirus (PECV) ...................................................................... 26
1.7.2 Murine norovirus (MNV) .................................................................................... 28
1.7.3 Feline calicivirus (FCV) ....................................................................................... 29
1.7.4 Tulane calicivirus (TCV) ...................................................................................... 29
1.7.5 Rabbit haemorrhagic disease virus (RHDV) ....................................................... 30
1.8 Development towards a new cell culture systems for RHDV ................................... 31
1.9 Hypothesis and Aims ................................................................................................. 33
1.9.1 Hypothesis.......................................................................................................... 33
2 Material and Methods ...................................................................................................... 35
2.1 Materials ................................................................................................................... 35
2.1.1 Plasmids ............................................................................................................. 35
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2.1.2 Oligonucleotides ................................................................................................ 37
2.1.3 Buffers, Solutions and Media ............................................................................. 38
2.1.4 Kits ...................................................................................................................... 39
2.1.5 Enzymes ............................................................................................................. 40
2.1.6 Cell Lines, Bacteria Strains and Viruses ............................................................. 40
2.1.7 Reagents ............................................................................................................. 41
2.1.8 Antibodies .......................................................................................................... 42
2.2 Methods .................................................................................................................... 43
2.2.1 Molecular Biology Methods ............................................................................... 43
2.2.2 Cell culture methods .......................................................................................... 53
3 Results ............................................................................................................................... 57
3.1 Generation of cells with a compromised type I IFN response .................................. 57
3.1.1 Construction of pcDNA3.1 Mx1 Promoter Plasmid ........................................... 57
3.1.2 Construction of TK1 Expression Plasmid ........................................................... 63
3.1.3 Construction of DTA Expression Plasmid ........................................................... 69
3.1.4 Generation of RK-13 clones ............................................................................... 73
3.2 Characterisation of Innate Immune Response ......................................................... 76
3.2.1 Real-Time Polymerase Chain Reaction Analysis ................................................ 76
4 Discussion ......................................................................................................................... 86
5 References ........................................................................................................................ 94
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1 Introduction
1.1 The impact of feral rabbit populations in Australia
European rabbits (Oryctolagus cuniculus) have caused major environmental and agricultural
damage in Australia since they were introduced with the arrival of the first fleet in the early
1800’s. Rabbits, intentionally released for hunting, spread rapidly throughout mainland
Australia from 1860 to 1930 were the rabbit population also increased in size dramatically
during this period, reaching approximately 3 billon (Fenner 2010). Today, rabbits occupy all
but the northern most regions (Figure 1-1) of the continent and are currently considered a
major national pest (Cooke 2002; Fenner 2010). Feral rabbits have a devastating effect on
Australia’s and New Zealand’s native fauna and flora, and cause major financial losses to the
agricultural industry. Rabbits damage crops, compete for pastures, feed on endangered
native plants, compete with native animals such as the Australian bilby for warrens and
cause soil erosion. Major efforts including trapping, shooting, poisoning, fencing and warren
ripping have been made to control and decrease feral rabbit numbers. While these methods
have the potential to lower local rabbit numbers, they will never solve the national rabbit
problem or be a permanent solution.
Figure 1-1 The distribution of feral European rabbits across the Australian mainland and Tasmania (http://invasive.boabdevelop.info/invasive-animals/rabbits/index.html).
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1.2 Use of viral biological control agents in Australia
The rabbit pathogen Myxoma virus, belonging to the family of Poxviridae was released in
response to the growing feral rabbit population in a Australia in the 1950s. The virus was
shown to be non-pathogenic in any host species tested excluding lagomorphs (Spiesschaert,
McFadden et al. 2011). Myxoma virus causes myxomatosis, a serious disease characterised
by the progressive development of conjunctivitis, mucopurulent secretion from the eyes
and mouth and a gradual breakdown of the host’s immune system, resulting in subsequent
supervening bacterial infections in the respiratory tract (Best and Kerr 2000; Jeklova, Leva et
al. 2008). Animals infected with Myxoma virus usually die within 10-14 days post infection
(Robinson, Muller et al. 1999). Myxoma virus has an estimated mortality rate of 99.5% in
rabbits (Spiesschaert, McFadden et al. 2011), which made it an appealing biological control
agent.
Myxoma virus was introduced into four local populations of wild rabbits on the southern
tablelands of New South Wales in the 1950s (Merchant, Kerr et al. 2003). Over the next two
years the virus spread throughout Australia and was identified in most areas occupied by
rabbits (Merchant, Kerr et al. 2003). Myxoma virus killed an estimated 400 million rabbits in
the first year after its introduction and within a decade the rabbit population in Australia
was reduced by 95% (Burnet 1952; Spiesschaert, McFadden et al. 2011). Annual outbreaks
of myxomatosis are observed throughout Australia, which appeared to be dependent on
rainfall and the availability of virus vectors such as mosquitoes and the European rabbit flea
(Fenner 1958; Merchant, Kerr et al. 2003). Unfortunately the high lethality of Myxoma virus
was not maintained, as co-evolutionary selection pressures resulted in a reduction of the
overall mortality caused by Myxoma virus to 30% seven years after release (Spiesschaert,
McFadden et al. 2011). The decrease in the effectiveness of Myxoma virus again brought
forth the need to find new strategies to control rabbit numbers.
Hence, RHDV was imported into Australia in 1995 for testing as a potential biological control
agent for the again growing European rabbit infestation (Mutze, Cooke et al. 1998). A series
of field trials were conducted on Wardang Island, 5 km off the coast of South Australia, to
assess the effectiveness of RHDV (Czech strain V351) (Cooke 2002). During these trials,
RHDV escaped from the island and spread through mainland Australia (Cooke 2002).
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Blowflies collected immediately outside the quarantine area tested positive for RHDV,
indicating that RHDV may have been transferred from the island via an insect vector (Mutze,
Cooke et al. 1998). The Czech RHDV strain V351 was introduced illegally from Australia into
New Zealand by farmers in 1997 and, similarly to Australia, RHDV rapidly spread through the
rabbit population causing high fatality rates (Forrester, Boag et al. 2003). It was estimated
that a month after the initial spread of RHDV, only 5% of pre-RHDV rabbit numbers
remained in South Australia (Mutze, Cooke et al. 1998). Although RHDV initially reduced the
number of feral rabbits in Australia and New Zealand significantly, rabbit population have
been steadily increasing, indicating the effectiveness of the present strain of RHDV has not
been maintained (Lugton 1999; Forrester, Boag et al. 2003). A consistent decrease in rabbit
numbers in Australia is desperately needed to allow for the regeneration of many
endangered trees, shrubs and animals and to decrease growing costs to the agricultural
industry (Lange and Graham 1983).
1.3 Decreasing effectiveness of RHDV on rabbits in Australia.
The initial impact of RHDV on the rabbit population numbers in Australia has not been
maintained. There has been a steady increase in rabbit numbers over the last 6-7 years to a
density where rabbits are again noticeably reducing Australia’s biodiversity (Saunders,
Cooke et al. 2010). Interestingly, pre-existing antibodies that cross-react to RHDV have been
found in rabbits from other parts of Australia and Europe (Collins, White et al. 1995;
Nagesha, Wang et al. 1995; Chasey, Trout et al. 1997; Moss, Turner et al. 2002; Robinson,
Kirkland et al. 2002). The effectiveness of RHDV was lower in cooler areas such as Victoria
were a higher percentage of rabbits survived infection (Ward, Cooke et al. 2010).
Furthermore antibodies cross reacting with RHDV were found in rabbit sera collected prior
to the release of RHDV (Cooke 2002).This lead to the hypothesis that the rabbits may have
acquired the antibodies after infection by non-lethal viruses, related to RHDV increasing
their survival rate (Cooke 2002; Ward, Cooke et al. 2010). A non-pathogenic virus belonging
to the Lagovirus genus termed ‘Rabbit Calicivirus-Australia 1’ (RCV-A1) was identified in
Australia, which may provide cross-protection against RHDV (Figure 1-2) (Strive, Wright et
al. 2009) and further RCV strains were discovered in Europe (Capucci, Frigoli et al. 1995).
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To overcome the deceasing effectiveness of originally released RHDV strain (Czech strain
V351), a screen for other strains of RHDV that are able come existing immunity to RHDV
and/or RCV-A1 and further lower rabbit numbers in Australia is currently underway (IA-CRC
2011).
Figure 1-2 Genbank sequences for the isolates were U54983, RHDV-V351 Czech; EU003579, RHDV Italy 90; M67473, RHDV FRG 91 Germany; Z29514, RHDV-SD 95 France; Z49271, RHDV-AST89 Spain; X87607, RHDV-BS89 Italy; EF363035, RHDV pJG Germany; DQ189077, RHDV 2006 Bahrain; EU003582, RHDV UT-01 USA; EU003581, RHDV NY-01 USA; DQ205345, RHDV JX/CHA/97 China; AF258618, RHDV Iowa USA; DQ280493, RHDV WHNRH China; EU003578, RHDV IN-05 USA; AY523410 RHDV CD/China 04; X96868, RCV Italy; AF454050, Ashington Isolate UK; EU871528, RCV-A1 Australia; NC_002615, EBHSV France; U09199, EBHSV pEB-2/4 Germany; U09199, EBHSV pEB-2/4 Germany; X9800, EBHSV BS89 Italy; AJ86699, rabbit Vesivirus. (Ward, Cooke et al. 2010)
The development of cell culture system for RHDV would help increase the knowledge of
RHDV biological aspects, such as molecular pathogencity determinants, which would be
useful information screening for a more favourable RHDV strain to be use in Australia’s
biological control efforts.
1.4 Caliciviridae biology
The name Caliciviridae is derived from the Latin word calix, meaning cup or chalice (Green,
Ando et al. 2000), as viruses belonging to this family have trademark cup-shaped
depressions on the virion surface (Figure 1-3) arranged in icosahedral symmetry (Green,
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Ando et al. 2000). The Caliciviridae family is comprised of small, non-enveloped positive
stranded RNA viruses, with a diameter of 27 to 35 nm (Clarke and Lambden 1997).
Figure 1-3 Negative-contrast electron micrograph of Norwalk-like virus in human stool specimen (Green, Ando et al.
2000).
1.4.1 Taxonomy
The ability to characterise and classify caliciviruses has been greatly limited by the absence
of effective cell culture systems, since many features commonly used to distinguish
between virus families such as proteins expressed in infected cells, antigenic relationships,
cell tropism and physicochemical properties cannot be easily analysed without it (Green,
Ando et al. 2000). Originally, caliciviruses were grouped into the Picornaviridae family, but
were listed as a distinct virus family in the Third Report of the International Committee on
Taxonomy of Viruses (ICTV) in 1978 (Green et al., 2000). The two virus families were
separated because of major differences in genome organisation. Calicivirus genomes are
organised into two to three major open reading frames, whereas picornaviruses only
contain one long open reading frame. Furthermore, Hepatitis E virus (HEV) was recently
removed from the Caliciviridae family because of the lack of phylogenetic relatedness
(Berke and Matson 2000) and differences in their replicative enzymes (Koonin and Dolja
1993). The decision to remove HEV from this family was not a unanimous by the calicivirus
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study group because of the apparent structural similarities between caliciviruses and HEV
(Green, Ando et al. 2000).
Figure 1-4 The phylogenetic relationship of among the Caliciviridae and Picornaviridae (Berke, Golding et al. 1997). Abbreviations: MX, Mexico Virus; RDHV, Rabbit hemorrhagic disease virus; EBHSV, European brown hare syndrome virus; FCV, Feline calicivirus; SMSV, San Miguel sea lion virus; Primate Pan-1, Primate calicivirus; VESV, Vesicular exanthema of swine virus; FMDV, foot-and-mouth disease virus; Polio 1, Poliovirus 1; HRV 14, Human rhinovirus 14; EMCV, Encephalomyocarditis virus; HAV, Hepatitis A virus (Picture taken from Cooke, 2002).
The Caliciviridae family is now comprised of four genera, Lagovirus, Norovirus, Sapovirus
and Vesivirus (Figure 1-4, Table 1-1). Caliciviruses affect a wide range of host species
including humans, primates, felines, swine and rabbits and cause a variety of different
diseases and symptoms. Viruses belonging to the Lagovirus genus such as RHDV and
European brown hare syndrome virus (EBHSV) cause serious diseases (rabbit haemorrhagic
disease and European brown hare syndrome respectively) with high mortality rates in
lagomorphs. The Lagovirus genus also includes Rabbit calicivirus (RCV), which infects rabbits
but is non-pathogenic (Capucci, Fusi et al. 1996; Strive, Wright et al. 2009). The Norovirus
and Sapovirus genuses include human pathogens, such Norwalk like viruses which are a
major cause of gastroenteritis in humans.
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Family Genus Species Strain
Caliciviradae Lagovirus European brown hare syndrome virus (EBHSV) EBHSV-BS89
EBHSV-FRG
EBHSV-GD
EBHSV-UK91
Rabbit hemorrhagic disease virus (RHDV) RHDV-AST89
RHDV-BS89
RHDV-FRG
RHDV-SD
RHDV-V351
Rabbit calicivirus (RCV)
Norovirus Norwalk virus (NV) Desert Shield
Lordsdale
Mexico
Norwalk
Hawaii
Snow Mountain
Southampton
Swine calicivirus
Sapovirus Sapporo virus (SV) Houston/86
Houston/90
London 29845
Manchester virus
Parkville virus
Sapporo virus
Vesivirus Feline calicivirus (FCV) FCV CFI/68
FCV F9
Vesicular exanthema of swine virus (VESV) Bovine
Cetacean s
Primate
Reptile
San Miguel sea lion virus (SMSV) serotype 1
serotype 4
serotype 17
Porcine enteric calicivirus (PEC)
Table 1-1 The Caliciviradae family contains four genera Lagovirus, Norovirus, Sapovirus, Vesivirus a. Also shown are the different virus species and strains.
1.5 Rabbit haemorrhagic disease virus
1.5.1 Molecular characteristics
RHDV is a single-stranded, positive-sense RNA virus, with a 7,437-nucleotide genome
(Wirblich, Thiel et al. 1996). In addition to the genomic RNA, a 2.2-kb subgenomic RNA
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covering the extreme 3’-third of the genomic RNA, is also produced during replication
(Wirblich, Thiel et al. 1996) and codes for the structural protein VP60 (Meyers, Wirblich et
al. 1991). Both RNA molecules are polyadenylated at the 3’ end, have a protein, named VPg,
covalently attached to the 5’end (Wirblich, Thiel et al. 1996; Goodfellow, Chaudhry et al.
2005) and are tightly packaged into non-enveloped icosahedral viral capsids that consist
largely of VP60 (Meyers, Wirblich et al. 1991).
The genomic RNA codes for two open reading frames, a larger 7 kb-open reading frame
(ORF1) and a 351-nucleotide open reading frame (ORF2) on its extreme 3’ end (Wirblich,
Thiel et al. 1996). The polypeptide transcribed from ORF1 contains seven autoproteolytic
cleavage sites and is cleaved to different degrees in vitro (Figure 1-5) (Wirblich, Thiel et al.
1996). Complete cleavage of the RHDV proprotein results in the production of 8 proteins
(Figure 1-5), including 3 helicases (P1, P2, P3), a cysteine protease (P5), VPg (P6), a
polymerase (P7) and the major capsid protein VP60 (Wirblich, Thiel et al. 1996).
Figure 1-5 Schematic representation of the cleavage products of the ORF1 polyprotein and proposed genetic map of ORF1. (A) The genomic RNA of RHDV is represented below the scale bar. Open reading frames are shown as open or shaded bars. Cleavage sites in the ORF1 polyprotein are indicated by vertical lines and numbered 1 to 7. The nonstructural proteins are designated P1 to P7. The molecular masses of the proteins (in kilodaltons) are shown above the bars, and their established or putative functions are indicated below the bars. (Wirblich, Thiel et al. 1996).
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1.5.2 Pathology
RHDV is the cause of rabbit haemorrhagic disease (RHD), a highly infectious disease
characterised by high mortality and morbidity in adult rabbits (Xu and Chen 1989). The
typical mortality rate of RHDV is estimated to be between 60-90% (Mocsari, Meder et al.
1991; Ohlinger, Haas et al. 1993). After RHDV infection, viral RNA is first observed in the
liver and spleen and can then be detected in the lung, kidney, bile, thymus, lymph nodes
and white blood cells within 30 hrs post infection (Shien, Shieh et al. 2000). RHDV antigens
have also been detected in the rabbit’s liver, bile and spleen (Shien, Shieh et al. 2000). RHDV
is also found in macrophages and monocytes extracted from a variety of organs after
infection by RHDV (Ramiro-Ibanez, Martin-Alonso et al. 1999). Symptoms associated with
RHDV (Figure 1-6) include hepatic failure, an enlargement and discolouration of the spleen
and haemorrhages in multiple organs, including the liver (Marcato, Benazzi et al. 1991).
RHDV is usually fatal in adult rabbits within 48 to 72 c hrs post infection (Marcato, Benazzi et
al. 1991). Interestingly, rabbits younger than 10 weeks are more likely to survive an RHDV
infection then adult rabbits (Shien, Shieh et al. 2000).
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Figure 1-6 Pathology of RHD. (A) Typical posture of a rabbit that died from RHDV infection. (B) Bleeding from the nostrils is frequently observed in rabbits that die from acute RHD. (C) Internal organs of a rabbit that died from RHD. The haemorrhagic lungs and the discoloured liver are visible. (D) The internal organs of a healthy rabbit, with normal lungs and dark glossy liver. (E) Enlarged spleen of a rabbit that died from RHD and (F) a normal spleen for comparison (Ward, Cooke et al. 2010).
1.6 The role of the innate immune system in the prevention of viral
replication
The immune system is divided into two main branches, the adaptive immune system and
the innate immune system. The innate immune system is comprised of a variety of different
cells and defense mechanisms that protect the host from invading organisms in a non-
specific manner and is one of the bodies’ first lines of defense against the invasion of foreign
pathogens (Barral, Sarkar et al. 2009).
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1.6.1 Important innate immune system receptors
Viruses and other pathogens contain unique pathogen-associated molecular patterns
(PAMPs) which are not found naturally in the host (Kumar, Kawai et al.). The innate immune
system recognises these PAMPs via specific pattern-recognition receptors (PRRs), including
Toll-like receptors (TLRs), Nucleotide Oligomerization Domain (Nod)-like receptors (NLRs)
and retinoic acid-inducible gene I (RIG-I)-like receptors (RLRs) (Yoneyama and Fujita 2009)
(Table 2). The stimulation of these receptors results in the activation of different elements
of the innate immune system, through the production and release of pro-inflammatory
cytokines and interferons (IFN) (Yoneyama and Fujita 2009).
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PRRs (structure Adapters (structure) PAMPs/Activators Species
TLR TLR1 - TLR2 (LRR-TIR) MyD88 (TIR-DD), TIRAP (TIR)
Triacyl lipopeptides Bacteria
TLR2 - TLR6 (LRR-TIR) MyD88, TIRAP Diacyl lipopeptides Mycoplasma
LTA Bacteria
Zymosan Fungus
TLR2 (LRR-TIR) MyD88, TIRAP PGN Bacteria
Lipoarabinonmannan Mycobateria
Porins Bacteria (Neisseria)
tGPI-mucin Parasites (Trypanosoma)
HA proteins Virus (Measles virus)
TLR3 (LRR-TIR) TRIF (TIR) dsRNA Virus
TLR4 (LRR-TIR) MyD88, TIRAP, TRIF, TRAM (TIR)
LPS Bacteria
Envelope proteins Virus (RSV, MMTV)
TLR5 (LRR-TIR) MyD88 Flagellin Bacteria
TLR7 (LRR-TIR) MyD88 ssRNA RNA Virus
hTLR8 (LRR-TIR) MyD88 ssRNA RNA Virus
TLR9 MyD88 CpG DNA Bacteria
DNA DNA Virus
Malaria hemozoin Parasites
mTLR11 (LRR-TIR) MyD88 Not determined Bacteria (uropathogenic bacteria)
Profilin-like molecule Parasites (Toxoplasma gondii)
RLR RIG-I (CARDx2-helicase IPS-1 (CARD) RNA (5'-PPP ssRNA, short dsRNA)
Virus
MDA5 (CARDx2-helicase) IPS-1 RNA (poly IC, long dsRNA)
Virus
LGP2 (helicase) RNA Virus
NLR NOD1/NLRC1 (CARD-NBD-LLR) RICK (CARD), CARD9 (CARD)
iE-DAP Bacteria
NOD2/NLRC2 (CARDx2-NBD-LLR)
RICK, CARD9 MDP Bacteria
NALP3/NLRP3 (PYD-NBD-LRR) ASC (PYD-CARD) MDP Bacteria
CARDINAL (PYD-FIND) RNA Bacteria, Virus
ATP Bacteria? Host?
Toxin Bacteria
Uric acid, CPPD, amyloid-β
Host
NALP1/NLRP1 (CARD-FIND-NBD-LRR-PYD)
ASC Anthrax lethal toxin Bacteria
IPAF/NLRC4 (CARD-NBD-LRR) Flagellin Bacteria
NAIP5 (BIRx3-NBD-LRR) Flagellin Bacteria
β-Glucan Fungi
Table 1-2 PRRs are used by the innate immune system to detect PAMPs from a variety of different pathogens (Kawai and Akira 2009).
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1.6.1.1 Toll-like receptors (TLRs)
A wide range of different pathogens are detected by TLRs (Kawai and Akira 2009) and are
they best characterised group of receptors for the recognition of PAMPs (Kawai and Akira
2008). TLRs are expressed by a wide range of cells to detect invading pathogens including
viruses, bacteria, fungi and parasites (Kawai and Akira 2009). TLRs are comprised of three
major structural domains, a transmembrane domain, an intracellular domain that is
necessary for the activation of downstream signaling pathways (Kawai, 2008) and an
ectodomain, which is comprised of leucine-rich repeats (LRRs) that bind the respective
PAMP (Kawai and Akira 2009).
There are at least 12 known members belonging to the TLR family in mammals, and each
receptor detects a different PAMP (Akira 2009). The TLR family is commonly divided in two
subgroups, intracellular and extracellular TLRs, dependent on the localisation of the
receptor within the cell (Kawai and Akira 2009). TLR3, TLR7, TLR8 and TLR9 are located in
intracellular compartments within the cell, such as endosomes, lysosomes and endoplasmic
reticulum (ER) and detect microbial nucleic acids (e.g. viral RNA and DNA). TLR1, TLR2, TLR4,
TLR5, TLR6 and TLR11 are located on the surface of the cell and recognise pathogen
membrane components such as proteins, lipoproteins and lipids found in the bacterial cell
wall, viral particles and fungi (Kawai and Akira 2009). TLR9 and TLR11 are also found in
cellular compartments and a verity of detect microbes including bacteria and parasites. The
role of TLR10 (also expressed on the cell surface) is not currently known.
The activation of some TLRs triggers an anti-viral innate immune response resulting in the
production of type I IFNs, which play a major role in the cellular defense against viral
infection (Kawai and Akira 2009). The activation of TLRs results in the activation of myeloid
differentiation primary response gene (MyD88) dependent and/or TIR-domain-containing
adapter-inducing IFN-β (TRIF) dependent signaling pathways (Figure 1-7), (Akira 2009).
MyD88 dependent signaling has been shown to be activated by all TLRs except TLR3 (Honda,
Ouyang et al. 2007), whereas TRIF dependent signaling pathway is only activated by TLR3
and TLR4 (Honda, Ouyang et al. 2007). The initiation of the MyD88-dependent TLR signaling
pathway results in the recruitment of MyD88 to the receptor. MyD88 then forms a complex
with interleukin-1 receptor-associated kinases (IRAKs) 1, 2 and 4 (Akira 2009). During the
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formation of the complex, IRAK4 phosphorylates and activates IRAK1 and IRAK2 and recruits
tumor necrosis receptor-associated factor 6 (TRAF6) to the complex (Akira 2009). The
activation of downstream pathways by IRAK1 occurs within the first hour, after which IRAK1
is rapidly degraded, whereas IRAK2 activity is sustained for a longer period of time (Akira
2009). The IRAKs/TRAF6 complex then interacts with another complex consisting of TGF-β-
activated kinase 1 (TAK1) and TAK1-binding proteins (TAB) 1 and 2, resulting in the
phosphorylation and activation of TAK1. TAK1 subsequently phosphorylates and activates a
variety of different kinases, including IκB kinases (IKKs), Mitogen-activated protein kinases
(MAPK) and c-Jun N-terminal kinases (JNKs) (Akira 2009). The activation of IKK results in the
release of NF-κB (nuclear factor kappa-light-chain-enhancer of activated B cells) (Akira
2009), a major transcription factor that regulates genes involved in both the innate and
adaptive immune response (Livolsi, Busuttil et al. 2001).
The other signaling pathway initiated by TLR stimulation is the TRIF-dependent signaling
pathway that plays a crucial role in the innate immune response to viral infection through
the induction of type I IFN. TRIF has been shown to associate with TNF receptor-associated
factor 3 (TRAF3), TRAF6 and receptor-interacting protein 1 (RIP1) after stimulation of TLR3
and 4 (Meylan, Burns et al. 2004; Hacker, Redecke et al. 2006). TRAF6 and RIP1 activate NF-
KB, where as TRAF3 is responsible for the induction of type I IFN via IRF3 (Akira 2009). TRAF3
activates two related kinases, inducible IKB kinase (IKKi) and TANK-binding kinase 1 (TBK1),
which are both involved in the activation of IFN regulatory factor 3 (IRF3) and/or IRF7
(Fitzgerald, McWhirter et al. 2003; Hemmi, Takeuchi et al. 2004). IRF3, once phosphorylated
by IKKi or TBK1, translocates from the cytoplasm into the nucleus where it activates the
transcription of the type I IFN genes (Akira 2009).
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15
Figure 1-7 MyD88 (myeloid differentiation primary response gene)-dependent and TRIF (TIR-domain-containing adapter-inducing interferon-β)-dependent signalling pathways in TLR signaling. Abbreviations: TLR, Toll-Like
receptor; TRAM, TRIF-related adaptor molecule; IRAK, Interleukin-1 receptor-associated kinase; TRAF, TNF receptor associated factors; RIP1, receptor-interacting protein 1; TAK1, TGF-β-activated kinase 1; MAPK, mitogen-activated protein kinases (MAPK); IκB, kinases (IKK); NF-KB, nuclear factor kappa-light-chain-enhancer of activated B cells; TBK1, TANK-binding kinase 1; IRF, interferon regulatory factor (Akira 2009).
1.6.1.2 Nod-like receptors (NLRs)
The NLR (nucleotide-binding domain leucine-rich repeat containing) family of proteins
consists of cytosolic and membrane associated PRRs that function as regulators of the
innate immune system in response to microbial infection (Barnich, Aguirre et al. 2005;
Shaw, Reimer et al. 2008). The human NLR family currently consists of 23 known proteins
and at least 34 different NLRs have been identified in mice (Shaw, Reimer et al. 2008). NLR
proteins are mainly expressed in immune cells, although some NLRs are also present in non-
immune cells (Shaw, Reimer et al. 2008). Common structural features of NLRs include the
presence of a caspase recruitment domain (CARD), a baculovirus inhibitor domain (BIR) or a
pyrin domain (PYD), a C-terminal leucine-rich repeat (LRR) which is responsible for the
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16
detection/binding of PAMPs and a NOD domain that plays a role in self oligomerisation
during activation (Inohara, Koseki et al. 2000; Shaw, Reimer et al. 2008).
The two best characterised proteins belonging to the NLR family are NOD1 and NOD2
(Inohara, Ogura et al. 2001). NOD1 is expressed in most cell types, whereas NOD2 is only
expressed in some immune cells such as macrophages, dendritic cells, monocytes and
intestinal Paneth cells (Inohara, Chamaillard et al. 2005). It has been shown that both, NOD1
and NOD2, are capable of inducing NF-kB activation independent of TLR-signaling (Inohara,
Ogura et al. 2001). After ligand binding NOD1/2 undergoes self oligomerisation and
conformational change, after which a serine threonine kinase (RICK) is recruited and
activated, which is required for the activation of MAPKs and NF-kB (Shaw, Reimer et al.
2008). K63-linked regulatory ubiquitination of RICK then occurs (Shaw, Reimer et al. 2008),
which leads to the subsequent recruitment of TAK1 (Hasegawa, Fujimoto et al. 2008). TAK1
is also necessary for the activation of MAPKs, although the signaling intermediates required
for this process are not well characterised (Shaw, Reimer et al. 2008). NOD1 utilises TRAF6
appears, whereas TRAF2/5 seem to be used in the signaling pathway initiated be NOD2
(Shaw, Reimer et al. 2008). The stimulation of either NOD1 or NOD2 results in the
production of pro-inflammatory mediators via NF-κB (Shaw, Reimer et al. 2008).
1.6.1.3 RIG-I-like receptors (RLRs)
The RLR family consists of three cytosolic RNA helicase PRRs, which specifically detect single
stranded and double stranded viral RNA in immune and non-immune cells (Kawai and Akira
2009). The three receptors belonging to this family are RIG-I, melanoma differentiation
associated gene 5 (MDA5) and laboratory of genetics and physiology gene 2 (LPG2)
(Yoneyama and Fujita 2008). RIG-I contains two CARDs at its N-terminus, a central RNA
helicase domain and a repressor domain at its C-terminus (Johnson and Gale 2006;
Yoneyama and Fujita 2009). The C-terminal domain has been shown to bind to and
recognise non-self RNA (Saito, Owen et al. 2008) in virus infected cells and inhibit RIG-I
activity in the absence of viral RNA (Takahasi, Yoneyama et al. 2008). MDA5 also contains
tandem CARD-like regions and a RNA helicase domain, but unlike RIG-I the C-terminal
portion of MDA5 does not appear to play an inhibitory role (Yoneyama, Kikuchi et al. 2005).
In contrast to MDA5 and RIG-I, LPG2 does not contain any CARDs (Saito, Hirai et al. 2007)
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17
and in vitro experiments indicate LPG2 to play an inhibitory role in the MDA5/RIG-I
mediated signalling pathways (Rothenfusser, Goutagny et al. 2005; Komuro and Horvath
2006).
RLRs play a major role in the detection of viral infection and in the initiation of an antiviral
state in cells through the transcriptional activation of type I IFN-stimulated genes
(Yoneyama and Fujita 2007). Studies using RIG-I knockout mice have shown the importance
of RIG-I for the induction of type I IFN in response to viral infection in fibroblast and
dendritic cells (Kato, Takeuchi et al. 2008). RIG-I and MDA5 are capable of detecting foreign
dsRNA and initiating downstream (Figure 1-8) signaling pathways through the CARD-
containing mitochondrial adaptor molecule IFN-β promoter stimulator 1 (IPS-1) (Seth, Sun et
al. 2005). It appears that IPS-1 acts as a support molecule for the propagation of the
signaling cascade resulting in the activation of transcription factors including IRF-3 and NF-
κB (Seth, Sun et al. 2005; Saha, Pietras et al. 2006). Gene-silencing studies have shown that
IPS-1 is essential for the IFN induction after MDA5 and RIG-I stimulation (cite something).
IPS-1 acts upstream of TBK1 and IKKi, which are phosphorylated and activated after RLR
stimulation. The activation TBK1 and IKKi subsequently results in the activation of IRF-3
(Fitzgerald, McWhirter et al. 2003; Sharma, tenOever et al. 2003). NF-κB is also indirectly
activated by IRF-3. IRF-3 and NF-kB are responsible for the transcriptional activation of a
variety of antiviral effectors including type I IFN and inflammatory cytokines, leading to the
induction of an antiviral state in infected cells (Bamming and Horvath 2009).
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18
Figure 1-8 Signalling pathways initiated by MDA5 and RIG-I in infected cells. Abbrevations: IPS-1, interferon promoter
stimulator-1; RIP1, receptor-interacting protein 1; FADD, fas-associated protein with death domain; TRAF6, TNF receptor associated factor 6; NF-κB, nuclear factor κ-light-chain-enhancer of activated B cells; IKK, IκB kinases; TBK1, TANK-binding Kinase 1; IRF, Interferon regulatory factors (Johnson and Gale 2006).
Despite the domain structure and amino acid sequence similarities, RIG-I and MDA5 are not
redundant and induce IFN in response to different types of viruses (Bamming and Horvath
2009). RIG-I specifically binds short double stranded RNA molecules and 5’-
triphosphorylated single stranded RNA (Hornung, Ellegast et al. 2006; Kato, Takeuchi et al.
2006; Kato, Takeuchi et al. 2008; Saito, Owen et al. 2008). No specific natural ligand has
been confirmed for MDA5 but it shows a preference for larger double stranded RNA (>2kbp)
and can detect synthetic dsRNA (poly IC) (Gitlin, Barchet et al. 2006; Kato, Takeuchi et al.
2008). RIG-I has been shown to induce type I IFN expression in response to Newcastle
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19
disease virus, Sendai virus, Vesicular stomatitis virus and Japanese encephalitis virus,
whereas MDA5 induces type I IFN expression in response to Picornaviridae including Theilers
virus, Encephalomyocarditis and Mengo virus (Kato, Takeuchi et al. 2006). Detection of
Murine norovirus (MNV) and subsequent stimulation of IFN has also been shown to be
MDA5 dependent (McCartney, Thackray et al. 2008).
1.6.2 Interferons
The first line of defence against viral infection in higher eukaryotes is largely dependent on
the rapid activation of transcription factors that result in the production and secretion of a
family of cytokines referred to as IFNs (Johnson and Gale 2006). IFNs are commonly divided
into three subgroups, type I IFNs (including IFN-α, IFN-β and IFN-ω) which are induced by
viral infection and are produced by most cell types; type II IFN (IFN-γ) which is induced by
antigenic stimuli and is only produced by certain immune cells including natural killer cells,
cytotoxic T cells and helper T cells, and finally type III IFNs (IFN-λ1 to λ3) which are also
induced by viruses (Samuel 2001).
Viral infection has been shown to induce the transcriptional activation of a large number of
cellular genes (Zhu, Cong et al. 1998; Chang and Laimins 2000). Type I IFNs signal cells in an
autocrine and paracrine manner (Heim 1999) via binding to a common cell surface receptor
comprised of two subunits, IFN-α/β receptor (IFNAR)-1 and IFNAR-2 (Samuel 2001). The
type II IFN receptor complex also consists of two subunits, the IFN-γ ligand-binding IFN-γ
receptor-1 (IFNGR) subunit and the accessory subunit IFNGR-2 (Heim 1999). The initiation of
IFN signaling involves the IFN-mediated heterodimerisation of the respective cell surface
receptor subunits (Bach, Aguet et al. 1997). The activation of the type I IFN receptor (Figure
1-9) results in the intracellular activation/phosphorylation of the receptor associated Janus
kinase 1 (Jak1) and tyrosine kinase 2 (Tyk2) and the subsequent phosphorylation of the
latent transcription factors signal transducer and activator of transcription 1 (STAT1) and
STAT2 in the cytoplasm (Samuel 2001). After phosphorylation, STAT1 and STAT2 proteins
form a heterodimer and translocate into the nucleus. Once in the nucleus, the STAT
heterodimer forms a complex with IRF9 and binds to IFN-stimulated response elements
(ISREs), located in the promoter regions of IFN-stimulated genes (ISG) (Samuel 2001). In this
way, more than 100 known ISGs are transcriptionally activated and expressed, resulting in
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20
the establishment of an antiviral state in the type I IFN-stimulated cell. Since ISGs include a
whole set of directly anti-viral acting proteins, the ability of viruses to propagate in type I/III
interferon stimulated cells is severely reduced. The importance of type I IFNs in the cellular
defense to virus infection has been demonstrated in type I IFN receptor-knockout mice,
which are unable to establish an antiviral state and hence are much more susceptible to a
range of viruses including Poxviridae, Arenaviridae, Rhabdoviridae and Togaviridae (Samuel
1985).
Figure 1-9 Type I IFN induced Jak/STAT signaling pathway. Abbreviations: IFNAR, interferon-α/β receptor-1; IFNGR, interferon-γ receptor-1; Jak1, Janus kinase 1; Tyk2, tyrosine kinase 2; P, phosphate group; STAT, signal transducer and activator of transcription; ISRE, IFN-stimulated response elements (adapted from Windisch et al, 2005).
IFN-α/β also play an important role in the mediation of apoptosis in virus infected cells
(Samuel 2001). Early destruction of virus infected cells can greatly reduce the
production/yield of progeny viruses and slow virus spread. Primary mouse embryonic
fibroblasts (MEF) undergo apoptosis after infection by Encephalomyocarditis virus (EMCV),
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21
Vesicular stomatitis virus (VSV) and Herpes simplex virus (HSV) in cell culture, although
infection by these viruses did not induce apoptosis in MEFs treated with anti-IFN-α/β
antibodies or in type I IFN receptor/STAT-1 defective cells (Tanaka, Sato et al. 1998).
Type II IFN plays an important role in the cellular protection against microbial pathogens,
although a decreased resistance to some viruses such as Herpes simplex virus and Vaccinia
virus has been observed in IFN-γ receptor-knockout mice (Huang, Hendriks et al. 1993; Bach,
Aguet et al. 1997; Cantin, Tanamachi et al. 1999; Samuel 2001).
1.6.2.1 IFN-induced antiviral proteins
The production of IFN in response to viral infection inhibits the replication of viruses due to
the expression of IFN-induced proteins, some of which elicit a variety of different antiviral
effects (Samuel 1991; Stark, Kerr et al. 1998). Genes up regulated in response to IFN
stimulation with a well studied antiviral function include protein kinase R (PKR), 2’,5’-
oligoadenylate synthetase (OAS), RNA-specific adenosine deaminase (ADAR1) and the Mx
protein family (Mx GTPases) (Figure 1-10).
Figure 1-10 Antiviral proteins contributing to the establishment of the antiviral state in IFN-stimulated human cells. From left to right: The MxA GTPase inhibits viral replication by missorting and trapping of viral components into large membrane-associated complexes (the role of GTP hydrolysis in this process is not
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22
fully understood). The different IFN-induced oglioadenylate synthetases include OAS1, OAS2 and OAS3. Binding to double-stranded RNA (dsRNA) leads to hetero- and/or homo-oligomerisation and subsequently to the production of oligoadenylates with a 2’-5’phosphodiester bond linkage. These 2-5A oligonucleotides activate the latent endo-ribo-nuclease RNase L, which leads to the degradation of viral and cellular RNAs (in some cell types, the expression of RNase L is also regulated by IFNs). ADAR1 binds to double stranded RNA and catalyses the conversion of adenosine to inosine (A to I). Such editing may occur selectively at one or few positions, or more frequently at a large number of sites. Editing of viral RNA may change the coding sequence, activate an inosine-specific RNase and/or destroy RNA secondary structures by distrupting adenosine/uracil base pairing. The double-stranded RNA-activated protein kinase PKR blocks viral protein translation via phosphorylation and thereby inactivation of eukaryotic initiation factor (eIF-)2α. Furthermore, PKR activates intracellular signaling pathways that contribute to the establishment of a robust antiviral response (adapted from Frese and Dazert, 2008).
1.6.2.1.1 Protein kinase R (PKR)
PKR is an IFN inducible, RNA-dependent kinase found predominantly in the cytoplasm of
type I IFN stimulated cells (Thomis, Floyd-Smith et al. 1992; Samuel 2001). It is expressed in
an inactive form and undergoes autophosporylation, dimerisation and activation in
response to dsRNA produced during viral replication (Clemens and Elia 1997; Randall and
Goodbourn 2008). It is also activated in response to a cellular stress-activated protein, the
PKR-activating protein (Ito, Yang et al. 1999). Post activation, PKR phosphorylates six or
more known substrates including inactive PKR (Thomis and Samuel 1993), the α subunit of
the eukaryotic initiation factor 2 (eIF-2α) and transcription inhibitor IKB (Kumar, Haque et al.
1994). The best characterised substrate for PKR is eIF-2α. The phosphorylation of eIF-2α
results in a decrease in mRNA translation in the host cell blocking mRNA synthesis and viral
protein expression (Samuel 1993).
1.6.2.1.2 2’,5’-Oligoadenylate synthetase (OAS) and RNase L
OAS and RNase L are both IFN-inducible antiviral enzymes that aid in the protection of cells
from viral infection by degrading RNA. Similar to PKR, OAS is synthesised in its inactive form
and requires dsRNA as a cofactor for activation (Randall and Goodbourn 2008). After
binding to dsRNA, OAS undergoes oligomerisation and becomes active. Active OAS catalyses
the conversion of ATP into 2’,5’-linked oligomers of adenosine (2-5A oligonucleotides) (Kerr
and Brown 1978). RNase L, which is constitutively present in most cell types is activated
after binding to the 2-5A oligonucleotides (Samuel 2001). The binding of 2-5A
oligonucleotides to RNase L triggers the dimerisation of RNase L monomers, allowing RNase
L to cleave viral RNA and decrease viral protein expression (Samuel 2001). RNase L deficient
cells are more susceptible to viral infection by EMCV, Picornaviruses, West Nile virus and
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23
Paramyxoviruses (Dong, Xu et al. 1994). Active RNase L also degrades some self-mRNA,
which results in the production of small RNA molecules that may act as ligands for
intercellular receptors such as RIG-I and MDA-5 thereby amplifying the antiviral innate
immune response (Malathi, Dong et al. 2007).
1.6.2.1.3 RNA-specific adenosine deaminase (ADAR1)
ADAR1 is an IFN inducible RNA-specific adenosine deaminase. The transcription of ADAR1 is
induced by both IFN-α and IFN-γ and up to a fivefold increase in transcripts can be observed
in cells treated with type I IFN (Patterson and Samuel 1995; Patterson, Thomis et al. 1995).
Structural and functional studies of ADAR1 have shown that the nucleic acid binding
domains are located in the N-terminal region and the C-terminal region constitutes the
catalytic domain of the deaminase (Lai, Drakas et al. 1995; Samuel 2001). The central region
of the ADAR1 open reading frame contains three copies of the dsRNA binding motif
(dsRBMI-III) which are highly similar to each other and to the dsRBM identified in PKR
(Green and Mathews 1992; Kim, Wang et al. 1994; Liu, George et al. 1997).
The modification of mRNA by ADAR1 occurs via the site-specific deamination of adenosine
to inosine. The conversion of adenosine to inosine destabilises double stranded RNA by
disrupting base paring. Adenine-uracil base pairs are exchanged with inosine-uracil pairs,
which are considerably less stable and as a result the RNA becomes more single stranded in
character (Bass and Weintraub 1988). Further, such RNA modification has potential to
change the protein-coding capacity of the edited transcript, as polymerases and ribosomes
recognise inosine as guanine, not adenine (Samuel 2001). This can result in a change in the
amino acids that are coded for in the RNA and potentially prevent the synthesis of
functional viral proteins (Samuel 2001).
1.6.2.1.4 Mx protein family
Mx proteins were originally identified in an inbred mouse strain, that showed exceptionally
high levels of resistance to influenza A virus (Horisberger, Staeheli et al. 1983). The protein
responsible for the increased viral resistance seen in mice was later identified as Mx1
(orthomyxovirus resistance gene 1) (Arnheiter, Skuntz et al. 1990). High levels of Mx1
expression at the initial sites of viral replication were shown to significantly decrease virus
spread and greatly increase survival (Arnheiter, Skuntz et al. 1990). Mx proteins now have
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24
been identified in most vertebrates including humans (MxA and MxB). Interestingly, Mx
genes are polymorphic in most species (Haller, Staeheli et al. 2007), which may account for
the different anti-viral mechanisms used by different Mx proteins (Samuel 2001).
The Mx proteins comprise a small family of high molecular weight guanosine
triphosphatases (GTPases) that belong to the super-family of dynamin-like GTPases (Haller,
Staeheli et al. 2007). Structural characteristics of Mx proteins include the presence of a
GTPase domain in the N-terminal region, which is highly conserved and an effector domain
in the C-terminal region containing a leucine zipper (LZ) motif (Haller, Staeheli et al. 2007). A
common trait of large dynamin-like GTPases is their ability to self-assemble into highly
ordered oligomers and to cooperatively hydrolyse GTP (Haller, Staeheli et al. 2007). Mx
proteins have been shown to form homo-oligomers and assemble into ring-like structures in
vitro (Figure 1-11), which appears crucial for GTPase activity and viral recognition
(Nakayama, Yazaki et al. 1993). A mutant MxA with an amino acid exchange in the proximal
part of the LZ region failed to self-assemble and prevented the MxA protein from
hydrolysing GTP, indicating self-oligomerisation is vital for the regulation of MxA GTPase
activity (Janzen, Kochs et al. 2000; Haller, Staeheli et al. 2007). It has also been hypothesised
that the self-oligomerisation maybe prevent protein degradation, as the LZ region mutant
MxA was degraded very rapidly compared to wild-type MxA, which has a half life of over 24
hrs (Haller, Staeheli et al. 2007).
Figure 1-11 MxA GTPase self-assembles into highly ordered oligomers. GTP binding causes a change in conformation, leading to self-assembly of MxA in to ring-like structures (Kochs, Haener et al. 2002).
Orford-Dunne, Roxy (CES, Black Mountain)
25
Unlike other IFN induced proteins such as OAS and PKR, Mx proteins are not constitutively
expressed in normal cells. Basal levels of Mx are up regulated in response to IFN-α and IFN-β
stimulation, but not to IFN-γ (Arnheiter, Frese et al. 1996). The antiviral activity of Mx
proteins appears to be species and cell-specific and dependent on virus type (Haller,
Staeheli et al. 2007). MxA has been shown to have an antiviral effect on members of several
different virus families including Bunyaviruses, Orthomyxoviruses, Paramyxoviruses,
Rhabdoviruses, Togaviruses and Picornaviruses (Haller, Frese et al. 1998; Janzen, Kochs et al.
2000; Haller, Staeheli et al. 2007). The exact mechanism through which the antiviral action
of Mx proteins is executed is not entirely understood, but it has been shown that Mx
proteins are able to bind important viral components and thereby inhibit their cellular
movements and function (Kochs and Haller 1999).
1.7 Cell culture systems for caliciviruses
Most caliciviruses cannot be propagated in standard cell culture systems, which has
hampered many areas of calicivirus research and there is currently no cell culture system
available that supports RDHV replication. The same is true or the replication of human
Noroviruses, which are the major cause of non-bacterial gastroenteritis worldwide,
accounting for 50,000 hospitalisations and 23 million cases per year (Mead, Slutsker et al.
1999). Most efforts to generate calicivirus cell culture models presumably have been made
to establish a cell culture system to propagate human noroviruses and a wide variety of
different cell lines have been tested in this regard, but without any success (Duizer, Schwab
et al. 2004).
Currently one can only speculate why most Caliciviruses fail to replicate in cell cultures.
Caliciviruses appear to be very species specific, indicating the need to grow RHDV in Rabbit
cell lines. Furthermore, some animal caliciviruses can be propagated in cell culture and their
respective requirements to replicate in vitro may indicate the best way to start the
establishment of a cell culture system for RHDV. For example, Feline calicivirus has been
successfully propagated in feline kidney cortex cells (Slomka and Appleton 1998; Thumfart
and Meyers 2002), Porcine enteric virus can be grown in kidney epithelial cells (Flynn and
Saif 1988; Parwani, Flynn et al. 1991), the newly discovered primate calicivirus Tulane virus
has also been shown to effectively replicate in monkey kidney cells (Farkas, Sestak et al.
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26
2008). Murine Norovirus is currently used as a virus models for human norovirus studies, as
MNV can be propagated to some extent in primary macrophages and dendritic cells of mice
(Wobus, Karst et al. 2004). Investigating effective cell culture systems currently used to
propagate caliciviruses may give insight into the requirements needed to propagate and
study RHDV in cell culture (as outlined in the following paragraphs).
1.7.1 Porcine enteric calicivirus (PECV)
Porcine enteric calicivirus (PECV) successfully grows in cell culture and has therefore been
used as a model to study the replication of enteric caliciviruses. It is currently the only
calicivirus that causes a gastrointestinal disease for which a cell culture system is available
(Chang, Sosnovtsev et al. 2004). The Cowden strain of PECV successfully replicates in
primary and continuous porcine kidney epithelial cells (LLC-PK) (Flynn and Saif 1988;
Parwani, Flynn et al. 1991). Interestingly, PEC growth is dependent on the addition of
intestinal content fluid filtrate from uninfected gnotobiotic pigs (pigs in which only certain
known strains of bacteria and other microorganisms are present) (Miniats and Jol 1978;
Chang, Sosnovtsev et al. 2005). In the absence of intestinal content, no viral protein or RNA
synthesis has been observed in cell culture, which suggests that some substance(s) in the
intestinal content may play an essential role in PEC replication (Chang, Sosnovtsev et al.
2004). Bile acids may be the active factors in the IC that allow PECV growth in vitro, as PECV
growth is inhibited by the addition of cholestyramine resin (bile acid-binding resin) to
intestinal content-treated growth medium (Chang, Sosnovtsev et al. 2004).
A G-protein-linked receptor for bile acids was recently identified and it was shown that the
interaction between this receptor and bile acids results in an increase in the concentration
of intracellular cyclic AMP (cAMP) (Kawamata, Fujii et al. 2003). Bile acids have also been
shown to activate protein kinase A (PKA) pathways in cells (Chang, Sosnovtsev et al. 2004). It
has been suggested that the activation of the PKA and the up regulation of cAMP may play a
role in the immunosuppressive actions associated with bile acids (David, Petricoin et al.
1996; Sengupta, Schmitt et al. 1996; Lee and Rikihisa 1998). The addition of bile acids to the
growth medium of LLC-PK cells causes a significant decrease in IFN-α and IFN-γ mediated
STAT1 phosphorylation (by 30-50%) and the addition of intestinal content (1%) inhibits
STAT1 phosphorylation up to 75% (Figure 1-12) (Chang, Sosnovtsev et al. 2004). The
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27
inhibitory role of bile acids on the innate immune system might be essential to allow for the
replication of PECV in cell culture and in vivo.
Figure 1-12 The relative values (as represented by the value of IFN treatment as 100%) of IFN-γ induced luciferase activity after cells (LLC-PK) were transfected with pGAS-TA-luc plasmid and treated with medium only, IFN-γ, IFN-γ + individual bile acids (GCDCA, TCDCA, TCA, TDCA, TUDCA) and IFN-γ + IC (Chang, Sosnovtsev et al. 2004).
An effective reverse genetics system for PECV has also been developed (Chang, Sosnovtsev
et al. 2005). Infectious PECV RNA was produced in LLC-PK cells after being transfected with a
plasmid containing a full-length PECV genome. Viral capsid PECV proteins were detectable
48 hrs post transfection and infective progeny virus was recovered and passaged (Chang,
Sosnovtsev et al. 2005). The production of viral RNA after transfection with PECV genome
encoding plasmids, was also dependent on the addition of bile acids (such as GCDCA) or IC
(Chang, Sosnovtsev et al. 2005). Recovered recombinant PECV from transfected cells was
shown to be infectious for gnotobiotic pigs via oral inoculation, although reduced virulence,
less severe clinical symptoms and delayed virus shedding was observed in comparison to
infections with wild-type PECV (Chang, Sosnovtsev et al. 2005). These results warrant
further investigation if the use of bile acids could help to establish stable cell culture
systems for other caliciviruses, especially those that grow in tissues were bile fluids are
present.
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28
1.7.2 Murine norovirus (MNV)
The strongest argument that calicivirus replication is severely inhibited by type I IFNs comes
from experiments using MNV. MNV was originally identified in immune-compromised mice
(Karst, Wobus et al. 2003). MNV is highly virulent and lethal in mice with defective IFN type I
or IFN type II receptors and STAT1 or recombination-activating gene 2 deficient mice (Karst,
Wobus et al. 2003). STAT1 knockout mice suffer a fatal disease after infection with MNV,
which is not observed in wild type mice (Wobus, Karst et al. 2004; Changotra, Jia et al.
2009). Higher levels of MNV RNA were also observed in the knockout mice (Figure 1-13)
indicating suppression of elements of the innate immune system might be a prerequisite for
efficient MNV replication (Karst, Wobus et al. 2003).
Figure 1-13 The level of viral RNA in the tissues of wild-type and STAT1-/- mice (Karst, Wobus et al. 2003).
A cell culture system for MNV was developed shortly after its discovery, making MNV the
only norovirus with a robust cell culture system and a popular model for studying human
noroviruses. MNV propagates successfully in primary bone marrow-derived macrophages
and dendritic cells from wild-type mice (Wobus, Karst et al. 2004). An increase in viral
replication is seen in MDA5-defective dendritic cells when compared to wild-type dendritic
cells (McCartney, Thackray et al. 2008). Interestingly, type I IFN signalling has been shown to
prevent the accumulation of MNV non-structural proteins and a late step in the replication
cycle of MNV in macrophages in cell culture (Changotra, Jia et al. 2009). These studies again
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29
highlight that disabling part(s) of the innate immune system such as the type I IFN signalling
may significantly enhance calicivirus replication, as is true for many other viruses.
1.7.3 Feline calicivirus (FCV)
Feline calicivirus (FCV) is one of the best studied members of the Vesivirus genus because it
replicates effectively in Crandel Reese Feline Kidney (CRFK) cells (Slomka and Appleton
1998; Thumfart and Meyers 2002). No specific innate immune defects or no special
additives to the cell growth medium are necessary to enable FCV replication in CRFK cells.
FCV grows in CRFK cells in medium containing non-essential amino-acids and 10% fetal calf
serum (Slomka and Appleton 1998; Thumfart and Meyers 2002). This cell culture system has
already been used in FCV reverse genetic studies, were infectious FCV particles were
successfully recovered after transfection of CRFK cells with cDNA constructs (Thumfart and
Meyers 2002).
1.7.4 Tulane calicivirus (TCV)
Three monkey kidney cell lines (Vero, MA104 and LLC-MK2) and one human colon
carcinoma cell line (Caco-2) were used in an attempt to cultivate the newly discovered
primate Tulane calicivirus (TCV). The cells were inoculated with sterile filtrate collected from
rhesus macaques stool samples that tested positive, by a TCV-specific RT-PCR (Farkas,
Sestak et al. 2008). The cells were grown in Dulbecco’s modified Eagles medium (MA104,
Vero and Caco-2) or in M199 medium (LLC-MK2) supplemented with 10% fetal bovine
serum, penicillin, streptomycin and amphotericin B (Farkas, Sestak et al. 2008). Cells and
medium were collected 5 days post inoculation and LLC-MK2 cells were the only cells in
which TCV was detectable by RT-PCR (Figure 1-14) (Farkas, Sestak et al. 2008).
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Figure 1-14 Detection of TCV-specific RNA after the sixth passage of CaCo-2, MA-104, Vero and LLC-MK2 cells inoculated with a TCV-positive stool sample. Mock-infected cells were passaged in parallel with stool-inoculated cells for each cell line. RNA was extracted from 100μl of cell-free tissue culture medium from mock/infected cell culture. RNA extracted from 100μl of TCV-positive stool sample that was used to inoculate the cells originally was used as a positive control (+contr) (Farkas, Sestak et al. 2008).
The replication of TCV was then studied by infecting LLC-MK2 cells with plaque-purified TCV.
TCV caused a visible cytopathic effect on the cells within 24 hrs post inoculation and by 36
to 48 hrs, all cells were rounded and starting to detach (Farkas, Sestak et al. 2008). The viral
titers peaked between 36 to 48 hrs (Farkas, Sestak et al. 2008). The results from this study
indicate the replication cycle of TCV is quite rapid and that TCV can be grown in cell culture.
1.7.5 Rabbit haemorrhagic disease virus (RHDV)
1.7.5.1 Current cell culture systems
Various methods have been attempted to create cell culture systems for RHDV, but so far
efforts have been unsuccessful. An effective cell culture system needs to support several
processes including the entry of the virus into the cell, the production of viral proteins,
replication of the viral genome and the assembly and release of infectious progeny virus.
None of the cell cultures used to propagate RHDV so far have resulted in systems that
support all of these processes and lead to the production of infectious progeny virus (Liu,
Zhang et al. 2006; Liu, Ni et al. 2008).
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Primary hepatocytes have been used to examine RHDV replication and particle structure,
but this method is not ideal as it is very expensive, laborious and has not been shown to
result in the generation of infectious progeny virus (Konig, Thiel et al. 1998). The infection of
primary hepatocytes with RHDV resulted in the production of VP60 in the cytoplasm of
some cells, indicating at least some RHDV transcription may occur (Konig, Thiel et al. 1998).
A considerable amount of cells do not become infected with RHDV using this method and
infected cells show signs of apoptosis including a decrease in size and condensation of the
nuclei after 48 to 72 hrs post infection (Konig, Thiel et al. 1998).
DNA-based reverse genetic systems have also been used in an attempt to characterise RHDV
protein function and structure. Rabbit kidney (RK-13) cells have been transfected with RNA
transcripts generated in vitro from full-length RHDV genome cDNA clones (Liu, Zhang et al.
2006) and with the cDNA clones (Liu, Ni et al. 2008), which resulted in the production of
viral antigens and had a cytopathic effect on the cells. However, consecutive infection and
re-infection of RK-13 cells was not shown.
1.8 Development towards a new cell culture systems for RHDV
Choosing an appropriate cell line for the propagation of RHDV is difficult as very little is
known about many of the biological aspects of RHDV replication, including potential
receptor specificity or cell tropism. Many different factors need to be considered when
developing a new cell culture system for viruses, for example whether virus replication is
species specific or whether the virus shows tissue tropism.
So far there is no evidence to suggest that RHDV replicates in any species other then
Oryctolagus cuniculus (European Rabbit), proven by extensive trials using a wide range of
animals prior to the planned release of RHDV in Australia (Lenghaus, Westbury et al. 1994).
Also most, if not all of the cell culture models that have been developed for caliciviruses use
cell lines derived from the virus’ natural host and propagation of caliciviruses in non-species
specific cells has so far been unsuccessful (Farkas, Sestak et al. 2008). Therefore, using a
rabbit-derived cell line for the development of RHDV may have a greater potential for
success, than the use of cell lines derived from other species.
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Many caliciviruses that are currently propagated in cell culture replicate in kidney cell lines,
even though they do not appear to necessarily replicate in these tissues in vivo, and animal
studies have detected higher titers of FCV in alveolar cells (Langloss, Hoover et al. 1978) and
PECV in the proximal small intestine (Guo, Hayes et al. 2001). Using a rabbit kidney cell line
may be a good starting point for the development of a successful cell culture system, as a
rabbit kidney continuous cell line (i.e. RK-13) is readily available, whereas rabbit hepatocytes
are not. Immune cells, such as macrophages and dendritic cells may also enable RHDV
propagation. Large quantities of RHDV can been found in macrophages and monocytes in a
variety of organs after intravascular infection by RHDV (Ramiro-Ibanez, Martin-Alonso et al.
1999), indicating the virus may show a tropism for these immune cells.
The innate immune system has been shown to be a strong cellular defense against viral
infections. Thus, Vero cells which do not produce type I IFN, are commonly used to grow a
variety of viruses, such as Vaccinia virus (Benhnia, McCausland et al. 2009). Cells that do not
produce an effective IFN-induced antiviral response may allow for RHDV growth, which so
far has not been observed in vitro. This hypothesis is supported by the fact that MNV growth
in cell culture is dependent on disabling elements of the IFN signalling pathway and PECV
replication may also rely on a down regulation of the type I IFN response by bile acids.
Interestingly, RHDV has been shown to replicate very rapidly and effectively in vivo, in the
face of a functional innate immune system. It is possible that RHDV grows rapidly enough to
outrun the innate immune response. Other viruses such as highly pathogenic influenza
viruses have been shown to be capable of replicating to very high levels before the antiviral
defense of the innate immune system is fully activated (Haller, Staeheli et al. 2007). RHDV
may not be able to replicate as rapidly in cell culture because available continuous cells do
not support such a rapid rate of replication. RHDV does not replicate successfully in primary
cells in vivo either, which could be the result of sub-optimal growth conditions. Hence, IFN
incompetent cells may provide an advantage that counteracts the potential disadvantage of
suboptimal growth conditions/cells.
It is difficult to determine whether continuous or primary cells are more likely to support
RHDV replication. The use of a continuous cell as a potential cell culture system could be
problematic as continuous cells are derived from tumor cells and may lack specific elements
Orford-Dunne, Roxy (CES, Black Mountain)
33
that allow for the growth of RHDV in vivo. Primary cells may more accurately represent the
in vivo replication environment and therefore enhance the chance of replication, although
continuous cell culture systems have many advantages over primary systems as they tend to
be more economical, less laborious to use and more readily available. Also using primary
cells will make the selection of immune incompetent cells difficult, if not impossible, as
primary cells are hard to transfect and to manipulate and show a limited number of
population doubling, only. Therefore it is unlikely that primary cells will ever become the
favourite model. Studies using liver tissue cultures to propagate RHDV have also been
unsuccessful, indicating that RHDV growth in vitro is not only limited in continuous cell lines
(Konig, Thiel et al. 1998).
Taking into account the current literature on caliciviruses and the role of the innate immune
system in the prevention of viral replication, the use of a continuous cell line derived from
rabbit kidney cells (RK-13 cells) was used as a starting point for the development of a new
cell culture system for RHDV in this study.
1.9 Hypothesis and Aims
1.9.1 Hypothesis
It has been shown that the type I IFN response effectively inhibits calicivirus growth (e.g.
that of MNV) (Karst, Wobus et al. 2003; Changotra, Jia et al. 2009). Earlier infection studies
in primary hepatocytes and RK-13 cells showed that these can be infected with RHDV in
vitro, but no viral replication was observed (M. Matthaei, personal communication; (Konig,
Thiel et al. 1998). Hence, the hypothesise that RHDV may replicate more effectively in rabbit
cells with a compromised type I IFN response will be examined in this study.
To address this hypothesis, expression plasmids for DTA and TK1 under the control of the
murine Mx1 promoter were constructed. The Mx1 promoter was chosen as it is specifically
and strongly induced by type I IFNs and has a very low base line activity level (Samuel 2001).
The Mx1 promoter-DTA/TK1 plasmid was then transfected into RK-13 cells. Successfully
transfected cells could be selected for using the antibiotic G418, as both plasmids contain a
neomycin resistance gene, allowing cells containing the plasmid to survive G418 treatment.
The surviving cells will then be treated with a hybrid human type I IFN to stimulate a type I
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34
IFN response. Any cell that in response to type I IFN stimulation initiates the type I IFN
Jak/STAT signaling pathway is expected to activate the Mx1 promoter, and express DTA or
TK1. Cells that express DTA should die without further treatment, whereas cells that express
TK1 require a substrate (Ganciclovir) for cell death to occur. Cells surviving this selection
process should have a defect in the type I IFN signaling cascade and can then be tested if
they support RHDV replication
Figure 1-15 Flow chart illustrating the process that will be used in this study to select for RK13 cells with a defect in the type I IFN signaling cascade
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2 Material and Methods
2.1 Materials
2.1.1 Plasmids
Construct Description
pGL3-Mx1P-Luc Expression vector for firefly luc (luciferase) reporter gene, under
the control of the murine Mx1 promoter. Vector also contains
ampicillin resistance gene.
pKO-Scrambler-DTA Expression vector for DTA gene, under the control of the
phosphoglycerate kinase (PGK) promoter. Vector also contains
ampicillin resistance gene.
pcDNA3.1/V5-His Mammalian expression vector, containing a cytomegalovirus
(CMV) promoter, an ampicillin resistance gene and a neomycin
resistance gene (Invitrogen).
pBluescript-TK1 Expression vector for TK1 gene, under the control of the Lac
promoter. Vector also contains ampicillin resistance gene.
peGFP-N1 Encodes a red-shifted variant of wild-type GFP, which has been
optimized for brighter fluorescence and higher expression in
mammalian cells. Also contains a Neomycin resistance gene.
Table 2-1 Plasmids used in this study.
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Construct Description
pcDNA3.1-Mx1p-9 Vector plasmid containing the Murine Mx1 promoter and an
ampicillin and neomycin resistance gene.
pcDNA3.1-Mx1p-13 Vector plasmid containing the Murine Mx1 promoter and an
ampicillin and neomycin resistance gene.
pcDNA3.1-Mx1p-DTA-8 Expression vector for DTA ‘death’ gene, under the control of the
Murine Mx1 promoter. Vector also contains ampicillin and
neomycin resistance gene.
pcDNA3.1-Mx1p-DTA-24 Expression vector for DTA ‘death’ gene, under the control of the
Murine Mx1 promoter. Vector also contains ampicillin and
neomycin resistance gene.
pcDNA3.1-Mx1p-TK1-1 Expression vector for TK1 ‘death’ gene, under the control of the
Murine Mx1 promoter. Vector also contains ampicillin and
neomycin resistance gene.
pcDNA3.1-Mx1p-TK1-2 Expression vector for TK1 ‘death’ gene, under the control of the
Murine Mx1 promoter. Vector also contains ampicillin and
neomycin resistance gene.
Table 2-2 Plasmids generated in this study.
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2.1.2 Oligonucleotides
Name Sequence (5’-3’)
rGAPDH1_fw_278 TCGAAGACGATCAGATACCG
rGAPDH1_rev_425 CCCTTCCGTCAATTCCTTTA
rGAPDH2_fw_470 CCACCTAGAGGAGCCTGTTC
rGAPDH2_rev_633 AATGTAGCCCATTGCTCTCC
rIP10_299fw GCCATCAAGAAGTTGCTGAA
rIP10_446rev CTGCAAACTGAGGCCAATTA
rIP10_55fw AACCATGAACCAAAGTGCAA
rIP10_237rev CATTGTGGCAATGATCTCAA
rMx1_522fw TGACTAAAGCCCAGAACGTG
rMx1_720rev GACCAGGTTGATGGTCTCCT
rMx1_51fw TGGTGGAGAAGAGCACACA
rMx1_203rev GCGTACAGGTTGTTCTCAGG
pGL3_4967fw GCAGGTGCCAGAACATTTCTCTATCG
pGL3_123rev GCGGAACTGGGCGGAGTTAGG
pcDNA3.1_98fw GTTGGAGGTCGCTGAGTAGCG
pcDNA3.1_1039rev ACTCAATGGTGATGGTGATGATGACC
DTA_cl_revNot1 AGGTCCTCGCGCGGCCGCCTGATGAGTTGTTGATTCTTCTAAATC
DTA_cl_fwXho1 TTTGCCGAGCTCCGAGGTCGAGCCCCAGCTGGTTC
DTA_cl_fwNot1 CCTTTTGGCGGCCGCCGAGGTCGAGCCCCAGCTGGTTC
DTA_seqfw1 GCCGCCTGATGATGTTGTTGATTC
DTA_seqfw518 CCATATACTCATACATCGCATCTTGG
DTA_seqrev658 TCAGCCTTCCCTTCGCTGAGG
DTA_417rev AGGCTGAGCACTACACGC
DTA_cl_rev2Not CTGACCTGCGGCCGCTCGCCATGGATCCTGATGATG
Mx1P-int_rev CTCTTCTTACCCTGTCATGCGG
Mx1P-int_fw TGTCCTTCCACCATGTGGCC
pBlueSc_seq_fw GTAAAACGACGGCCAGTG
pBlueSc_seq_rev CAGGAACAGCTATGACC
TK1-int194-fw CCACGCAACTGCTGGTGGC
TK1pA-int188-rev GCCTTCACCCGAACTTGGG
DTA_cl_fwNot1-2 CCTTTTGGCGGCCGCCGAGGTCGAGCCCCAGCTGGTTC
DTA_cl_rev2Not-2 CTGACCTGCGGCCGCTCGCCATGGATCCTGATGATG
Table 2-3 Primers used for analytical and cloning procedures during this study.
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2.1.3 Buffers, Solutions and Media
Buffer/Solution Composition
TAE (50x) 2 M Tris, 5.7% glacial acetic acid, 50 mM EDTA
DNA loading dye (6x) 2.5 mg/ml bromophenol blue, 400 mg/ml
sucrose
PBS 50 mM potassium phosphate, 150 mM NaCl, pH
7.2
Table 2-4 Buffers and solutions used in this study.
Media Composition
Minimum Essential Medium Eagle (MEM) MEM; 10% fetal calf serum (FCS), 7.5% w/v
sodium bicarbonate, 200 mM L-glutamine, 2 mM
glutamax, penicillin 100 units/l, streptomycin
100 µg/l.
Minimum Essential Medium Eagle (MEM)
/minus FCS
as above, without FCS
FCS Invitrogen
Normal Goat Serum (NGS) NGS diluted to 5% in PBS
Luria-Bertani (LB) 5 g/l yeast extract, 10 g/l tryptone, 5 g/l NaCl.
LBA LB; 100 µg/l ampicillin.
LB Agar LB; 15 g/l agar, 100 µg/l ampicillin (added after
autoclaving).
Table 2-5 Culture mediums used for bacterial and cell culture.
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2.1.4 Kits
Kit Name Company
RNeasy RNA extraction Kit QIAGEN
QIAprep Spin Miniprep Kit QIAGEN
QIAquick Gel Extraction Kit QIAGEN
QIAquick PCR purification Kit QIAGEN
Nucleobond® Plasmid Purification (mini-
midi-maxi) kit
Nucleobond
Table 2-6 Commercial kits used in this study.
Enzyme kits Company
Tetro cDNA synthesis Kit Bioline
iScript One-Step RT-PCR Kit with SYBR
Green
BIO-RAD
Platinum® Taq DNA polymerase Invitrogen
Platinum® Pfx DNA polymerase Invitrogen
Superscript®III reverse transcriptase Invitrogen
RNase-Free DNase Promega
T4 DNA Ligase Promega
LigaFast™ Rapid DNA Ligation System Promega
T4 DNA polymerase Promega
Calf intestinal alkaline phosphatase (CIAP) Promega
QuantiFast SYBR® Green PCR Kit QIAGEN
QuantiTect® Reverse transcription Kit QIAGEN
Table 2-7 Commercial enzymes kit used in this study
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2.1.5 Enzymes
Restriction endonucleases Use Company
ApaII Analytical Promega
BamHI Analytical Promega
BglII Cloning Promega
BstzI Cloning Promega
DraI Analytical/cloning Neb
EarI Cloning Neb
EcoRI Analytical/cloning Promega
EcoRV Cloning Promega
HindII Analytical/cloning Promega
HindIII Analytical/cloning Promega
NotI Cloning NEB/Promega
MfeI Analytical/cloning NEB
SacI Analytical Promega
XbaI Analytical Promega
XhoI Cloning Promega
Table 2-8 Restriction endonucleases used for analytic restriction and cloning approaches in this study.
2.1.6 Cell Lines, Bacteria Strains and Viruses
Bacteria Genotype Company
One Shot® TOP10
Chemically Competent E. coli
F- mcrA Δ(mrr-hsdRMS-mcrBC),
φ80lacZΔM15 ΔlacX74 recA1
araD139 Δ(araleu) 7697 galU
galK rpsL (StrR) endA1 nupG
Invitrogen
E.coli XL1blue supE44 hsdR17 recA! endA1
gyrA46 thi relA1 lac- F[proAB+ lacI
lacZΔM15 Tn10(tet)]
Stratagene
Table 2-9 Bacterial strains used in this study
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Cell line Characteristics
RK-13 A continuous rabbit kidney cell line
SIRC A continuous rabbit cornea epithelial cell line
Table 2-10 Cell lines used in this study.
Serum RHDV +/-
Serum 453 RHDV positive
Serum K31 RHDV positive
Serum 305 RHDV negative
Table 2-11 Serum samples collected from rabbits (either positive or negative for RHDV) which were used in this study
2.1.7 Reagents
Reagent Company
Antibiotic G418 Invitrogen
SYBR® Safe DNA Gel Stain Invitrogen
Lipofectamine® 2000 Invitrogen
Poly(I:C)-Low Molecular Weight (Synthetic
analog of dsRNA – TLR3 ligand)
InvivoGen
Ganciclovir InvivoGen
Universal Type I Interferon [Human
Interferon Alpha A/D (BglII)]
PBL InterferonSource
DAPI (kind gift of Michael Frese)
TritonX (0.25%) Sigma
Formaldehyde Polysciences
Table 2-12 Reagents and treatments used in cell culture experiments in this study.
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2.1.8 Antibodies
Antibody Dilution Source
Mouse monoclonal antibody to Mx1 protein 1:250 (kind gift of Michael Frese)
Table 2-13 Primary antibodies used in this study for immunofluorescence experiments.
Antibody Dilution Source
Goat polyclonal anti-Mouse IgG (Dylight 488) 1:2000 GeneTec
Table 2-14 Secondary antibodies used for immunofluorescence experiments.
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2.2 Methods
2.2.1 Molecular Biology Methods
2.2.1.1 Restriction Digestions
Restriction digestions were used to verify the identity of original and constructed plasmids
and during cloning produces. Between 200-300ng of DNA was used to determine during
analysis experiments and up to 5 µg of DNA were used during cloning procedures
(preparative restriction digestions). The units of restriction endonuclease required for a
particular reaction was determined using the following equation (Equation 2-1).
Equation 2-1 Equation used to determine the amount of enzyme (with 100% enzyme activity) required to completely digest template DNA within one hour. *Substrate DNA is the DNA used by supplier to define one unit of the enzyme.
DNA was digested using a suitable restriction endonuclease in the presence of BSA (1
mg/ml) and the buffer recommend by the manufacturer following the manufacturer’s
protocol. All digestions were incubated for 1 -16 hrs and enzymes known to posses star
activity were not used in excess or over extended periods.
2.2.1.2 Polymerase Chain Reaction
DNA amplification for analytical and cloning purposes was performed via polymerase chain
reaction (PCR). For analytical purposes Platinum Taq DNA polymerase was used and for
preparative scale PCRs performed during the construction of plasmids Platinum DNA Pfx
polymerase and platinum DNA Taq polymerase was used. The reaction components and PCR
cycles for both polymerases are described below (Table 2-15, Table 2-16).
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Compents Amt (µl)
10x PCR buffer 1
5 mM dNTP mixture 0.4
50 mM MgCl2 0.3
Platinum® Taq polymerase
(Invitrogen)
0.1
Primer 1 (10 pmol/µl) 0.5
Primer 2 (10 pmol/µl) 0.5
Template DNA (approx 10 ng) 1
RNase free H2O 6.2
Total 10
Table 2-15 PCR reaction components for DNA taq polymerase used for analytical purposes.
DNA Taq polymerase Amt (µl) DNA Pfx polymerase Amt(µl)
10x PCR buffer 5 10x Pfx amplification buffer 5
5 mM dNTP mixture 2 5 mM dNTP mixture 3
50 mM MgCl2 1.5 50 mM MgCl2 1
Platinum® Taq polymerase 0.2 Platinum® Pfx polymerase 0.4
Primer 1 (10 pmol/µl) 1 Primer 1 (10 pmol/µl) 1.5
Primer 2 (10 pmol/µl) 1 Primer 2 (10 pmol/µl) 1.5
Template DNA 1 Template 1
RNase free H2O 38.3 RNase free H2O 39.6
Total 50 Total 53
Table 2-16 PCR reaction components for DNA Taq/Pfx polymerase used for preparative scale PCR.
The above reactions were prepared on ice and PCR reaction was carried out using a PCR
cycler (eppendorf).
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Taq DNA polymerase PCR cycle:
1. Initial denaturation: 94°C for 2 min
2. Denature: 94°C for 15secs
3. Anneal*: 55-60°C for 30 secs
4. Extend: 72°C for 1 min per 1kb of PCR product
5. Repeat steps 2 – 4 for 35 cycles
6. Final extension: 72°C for 5 min
*Annealing temperature was optimised for each individual primer set.
Pfx DNA polymerase PCR cycle:
1. Initial denaturation: 94°C for 2min
2. Denature: 94°C for 15secs
3. Anneal: 58°C for 30secs
4. Extend: 68°C for 1min per 1kb of PCR product
5. Repeat steps 2 – 4 for 35 cycles
6. Final extension: 68°C for 5min
All primers were tested before used for any analysis on known samples to ensure their
applicability and occurrence of only neglectable non-specific amplification. Each primer set
was also shown to produce fragments of expected sizes after PCR amplification. The
separation and analysis of PCR products were performed using gel electrophoresis. Agarose
gels between 0.5 – 2% where prepared using TAE buffer and SYBR safe DNA stain (0.01%).
The DNA samples were separated at approx. 80V for 45 min – 2 hrs (depending on the size
of the expected fragments) using a horizontal gel apparatus. DNA fragments were visualised
under UV illumination using a transilluminator (Alpha Innotech, FluorChem).
2.2.1.3 Gel Extraction and DNA Clean up
DNA digest fragments were separated by electrophoresis on agarose gel. The bands of DNA
were visualised using UV light and fragments of interest were excised using a sterile scalpel.
The isolation of DNA from the agarose was then performed using the QIAquick Gel
Extraction Kit (all buffers and columns used were provided with the kit). The manufactures
protocol was followed; firstly the agarose was solubilised at 50°C in solubilisation binding
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buffer. Isopropanol was added and the solution was transferred to QIAquick column. The
column was then centrifuged at 12,000rpm for 1 min and washed twice using two separate
wash buffers. The column was then placed in a clean 1.5ml microcentrifuge tube and DNA
was eluted using 30-50 µl of RNase free H2O. DNA samples were stored at -20°C until
required.
2.2.1.4 Vector Dephosphorylation
Linear vector plasmids were dephosphorylated before undergoing ligation to minimise the
occurrence of self ligation. The components used for the desphosphorylation reactions are
listed below.
Component Amount
DNA 30-40 µl
CIAP 10X reaction buffer 5 µl
Calf intestinal alkaline phosphatase (CIAP)
(0.01u/µl)
5 µl
RNase free H2O Up to 50 µl
Table 2-17 Components used in dephosphorylation reactions in this study.
The above components were prepared on ice and incubated at 37°C for 30min. After 30min
another 5µl of CIAP was added and the mixture was incubated at 37°C for an additional
30min. The reaction was terminated by performing a gel extraction (2.2.1.3).
2.2.1.5 Converting 5’ and 3’ overhangs to blunt ends
T4 DNA polymerase was used to fill 5’ over hangs and remove 3’ over hangs generated
during restriction digests, which were then used in blunt end ligation reactions. The
reaction components are described below (Table 2-18).
Component Amount
Plasmid DNA 1 µg
T4 DNA polymerase 5 U
10x T4 DNA polymerase buffer 5 µl
5 mM dNTP 5 µl
H2O Up to 45 µl
Table 2-18 Components used into fill 5’ and 3’ over hangs in this study.
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The reactions components were prepared on ice and incubated at 37°C for 5 min. The
reaction was terminated via incubation at 75°C for 10 min. The product was cleaned up via
agarose gel extraction using the QIAGEN gel extraction kit (2.2.1.3).
2.2.1.6 Plasmid Purification
Plasmid purification was either performed by running DNA on agarose gel using
electrophoresis and extracting the DNA (see 2.2.1.3). The manufactures protocol was
followed (all buffers and columns used were provided in the kit); firstly a membrane binding
buffer was added to the sample, which was then transferred to a QIAquick spin column. The
column was centrifuged at 13,000xg for 1 min to bind DNA and then washed twice using the
wash buffer supplied. The filter column was placed into a clean 1.5ml microcentrifuge tube
and DNA was eluted using 30-50 µl of RNase free H2O. DNA samples were stored at -20°C
until further analysed.
2.2.1.7 Ligation
DNA ligations were performed using T4 DNA ligase (standard ligation) or T4 DNA ligase –
LigaFast Rapid DNA Ligation System (rapid ligation). The amount of vector and insert DNA
differed depending on the size of vector to insert (Equation 2-2).
Equation 2-2 Used to determine the ng of vector required for the ligation reaction. A 3:1 molar ratio of vector:insert was used for ligations reactions using T4 DNA ligase and 1:2 molar ratio was used for rapid ligation.
The ligation reaction components for each system are listed below.
T4 DNA ligase (standard ligation)
Component Amount
Vector DNA ng*
Insert DNA ng*
T4 DNA ligase (3 u/µl) 0.5 µl
Ligase 10x buffer 1 µl
RNase free H2O Up to 10 µl
Table 2-19 Reaction components used for T4 DNA ligation (standard ligation) procedures in this study.
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T4 DNA ligase (LigaFast Rapid DNA Ligation System)
Component Amount
Vector DNA ng*
Insert DNA ng*
T4 DNA ligase (3 u/µl) 1.5
Ligase 2x buffer 5 µl
RNase free H2O Up to 10 µl
Table 2-20 Reaction components used for T4 DNA ligation using the LigaFast Rapid DNA Ligation System.
The above components were prepared on ice. The standard ligation components were
incubated at room temperature for 3 hours and the rapid ligation were incubated at room
temperature for 10 – 15 min. The ligations were directly used in the transformation
procedures (2.2.1.8).
2.2.1.8 Transformation of E.coli
The chemical transformation of E.coli was performed in accordance to the manufactures
protocol. 0.2 – 1 µl of plasmid maxi prep or 1 – 5 µl of ligation reaction was combined with
50 µl of bacteria (OD600=0.6). The mixture was incubated on ice for 30 min, after which it
was heated at 42°C for exactly 75 sec (‘heat shock treatment’). 250 µl of pre-warmed SOC
medium (Invitrogen) was then added to the mixture and it was incubated at 37°C for 1 hr.
Vials were centrifuged at 6000rpm for 2 min, after which 100 µl of supernatant was
removed. The pellet was then resuspended in the remaining 200 µl and 50-200 µl were
plated on LB-agar plates containing ampicillin (100µg/L). Plates were incubated over night at
37°C. Obtained colonies were resuspended in 5 ml of LBA and incubated over night at 37°C
for subsequent DNA extraction (mini prep).
2.2.1.9 Sequencing
Sequencing was used to confirm the identity of some of the original plasmids and generated
plasmids. All sequencing was performed by the Australian Genome Research Facility (AGRF)
Brisbane. The sequencing components sent to AGRF are listed below (Table 2-21).
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Component Amount
Double stranded plasmid 1000-1500 ng
Primer (10 pmol/µl) 1 µl
RNase free H2O up to 12 µl
Table 2-21 Reaction components sent to AGRF for sequencing.
2.2.1.10 Reverse transcription
The reverse transcription of RNA was performed using several different commercially
available reverse transcription kits, Superscript®III reverse transcriptase kit, Tetro cDNA
synthesis Kit and QuantiTect® Reverse transcription Kit. The reaction components and
procedures followed for each kit are listed below (Table 2-22, Table 2-23, Table 2-24).
Superscript®III reverse transcriptase kit
Step 1.
Components Amt (µl)
Oligo(dT) (50 μM) 1
5 mM dNTP mix 2
Template RNA 2
RNase free H2O 8
Step 2.
Components Amt (µl)
5X First-Strand Buffer 4
0.1 M DTT 1
RNaseOUT (40 U/μl) 1
SuperscriptIII RT (200 U/µl) 1
Table 2-22 Composition of reverse transcription reactions using the Superscript®III reverse transcriptase kit. All components were provided by the manufacturer.
The components for step 1 were prepared on ice and incubated at 65°C for 5 min before
being chilled on ice for 2 min. The components from step 2 were prepared on ice and added
to step 1 and mixed gently by pipetting. The reaction mixture was then incubated at 50°C
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for 45 min, after which the reaction was terminated by heating mixture at 70°C for 15 min.
Samples were stored at -20°C for further analysis.
Tetro cDNA synthesis Kit
Step 1.
Components Amt (µl)
Random Hexamer primer mix 1
5 mM dNTP 2
Template RNA 2
RNase free H2O 6
Step 2.
Components Amt (µl)
5x RT buffer 4
RNase Inhibitor (10 U/μl) 1
Reverse Transcriptase (200 U/µl) 1
RNase free H2O 4
Table 2-23 Components required for reverse transcription reactions using the Tetro cDNA synthesis transcription Kit. All components were provided by the manufacture.
The components from step 1 were prepared on ice, and then incubated at 65°C for 10 min
before being placed on ice for 2 min. The components from step 2 were then prepared on
ice, after which the component from step 1 and step 2 were combined and incubated for a
further 45 min at 4°C. The reaction was then terminated by heating the reaction mixture to
70°C for 15 min. Samples were then stored at -20°C until required.
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QuantiTect® Reverse transcription Kit components
Step 1.
Components Amt (µl)
7x gDNA Wipeout buffer 2
Template RNA 2
RNase free H2O 10
Step 2.
Components Amt (µl)
Quantiscript Reverse transcriptase 1
5x Quantiscript RT buffer 4
RT buffer mix 1
Table 2-24 Components required for reverse transcription reactions using the QuantiTect Reverse transcription Kit. All components were provided by the manufacture.
Components from step 1 were prepared on ice and then incubated at 42°C for 2 min. During
this time components from step 2 were prepared on ice. After the 2 min incubation period
components from step 1 and step 2 were combined and incubated for a further 15 min at
42°C. The reverse transcriptase was then inactivated by heating the reaction mixture to 95°C
for 3 min. The samples were diluted using 82µl of RNase free H2O and kept at -20°C until
required.
2.2.1.11 Real-Time Polymerase Chain Reaction
Several commercially available real-time PCR kits were tested during this study. QuantiFast
SYBR® Green PCR Kit and iScript One-Step RT-PCR Kit with SYBR® Green were both trialled
during experiments performed to optimise the real-time PCR reactions and the QuantiFast
SYBR® Green PCR Kit was used in all following experiments (because it was more cost
effective and showed comparable performance). The components required for each kit are
listed below (Table 2-25).
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QuantiFast SYBR® Green PCR Kit iScript One-Step RT-PCR Kit with SYBR®
Green
Component Amt (µl) Component Amt (µl)
2x QuantiFast SYBR Green PCR
Master Mix
5 2x SYBR Green RT-PCR reaction
mix
5
Primer 1 (10 pmol/µl) 0.5 Primer 1 (10 pmol/µl) 0.5
Primer 2 (10 pmol/µl) 0.5 Primer 2 (10 pmol/µl) 0.5
H2O 3 H2O 3
Table 2-25 RT-PCR reaction components required for RT-PCR reactions using QuantiFast SYBR® Green PCR Kit and iScript One-Step RT-PCR Kit with SYBR® Green.
96 well plates were used for all RT-PCR reactions. 96 well plates were kept on ice while RT-
PCR reactions were being prepared. Master mixes of above reagents (Table 2-25) were
prepared on ice and 9 µl of master mix was used per well. 1 µl of template (c)DNA was
added to each well after the addition of the master mix. The plates were then sealed and
briefly centrifuged, before being analysed using an rt-PCR thermal cycler (model: CFX96
Real–Time System, C1000 Thermal Cycler, Biorad). The absorbance of ROX and SYBR dyes
were measured.
Several experiments were performed to optimise the RT-PCR reaction conditions and to
make sure the primer pairs chosen worked effectively and no non-specific amplification
occurred. A range of different annealing temperatures (55-65°C) were tested for each
primer set to determine at which temperature optimal amplification occurred. The finally
used rt-PCR conditions are given below.
Rt-PCR cycle (for both QuantiFast SYBR® Green PCR Kit and iScript One-Step RT -PCR Kit with
SYBR® Green):
1. Initial denaturation: 95°C for 5 min
2. Denature: 15°C for 15 secs
3. Anneal: 60°C for 30 secs
4. Extend: 76°C for 10 secs
5. Repeat steps 2 – 4 for 39 cycles
6. Melt curve analysis (65-95°C ) of reaction products
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2.2.2 Cell culture methods
2.2.2.1 Culturing cell lines
The continuous rabbit kidney cell line (RK-13) and the continuous rabbit cornea epithelial
cell line (SIRC) used were cultured in MEM (Sigma) supplemented with 10% heat-inactivated
foetal calf serum (FCS, Gibco), sodium bicarbonate 7.5% w/v, 200 mM L-glutamine, 2 mM L-
glutamax, penicillin 100 U/ml and streptomycin 100 µg/ml. The cells were cultured in T75
cell culture flasks with 10 ml of MEM, in a humidified incubator (Sanyo CO2 incubator, model
MCO-17AI) at 37°C with 5% CO2 atmosphere. The cell cultures were passaged every 3-4
days. Cells were passaged after being washed once with 10 ml of sterile PBS, after which 1.5
ml of trypsin (0.5%) was applied to cells. The cells were then placed back into the incubator
for 2 min (or until cells detached completely from the bottom of the flask) and were
resuspended and diluted in fresh MEM + FCS. Cells were also passaged and grown in 6 and
12 well plates and 100mm dishes during different experiments
2.2.2.2 Generation of type I IFN defective RK-13 cell line
In an attempt to generate a type I IFN defective RK-13 cell line, RK-13 cells were passaged
and grown in 100 mm cell culture dishes and transfected with the final Mx1 promoter
DTA/TK1 plasmids generated (Table 2-2). G418 was then applied to select for successfully
transfected cells, after which cells were treated with either recombinant IFN-α or
recombinant IFN-α 500 U/ml and 40 mM ganciclovir (1:10,000) in order to trigger the
transcription of DTA or TK1, respectively.
2.2.2.2.1 Transfection of eukaryotic cells
RK-13 cell transfection was performed using lipofectamine 2000 (Table 2-12) following the
manufactures protocol (Invitrogen). RK-13 cells were seeded into 100 mm cell culture dishes
and incubated over night at 37°C in 5% CO2 atmosphere to obtain 50-60% confluent cells for
transfection. 24 µg of either DTA or TK1 expression plasmids were diluted in 1.5 ml of FCS-
free MEM and subsequently combined with 1.5 ml of lipofectamine (diluted in FCS-free
MEM and incubated at room temperature for 5 min). The mixture of lipofectamine and DNA
was incubated at room temperature for 20 min before being applied drop-wise to the cells.
Transfected cells were then incubated for 5–6 hrs, after which the medium was removed,
and fresh MEM was applied. A plasmid from which eGFP is constitutively expressed in
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transfected cells was used as to approximate transfection experiments to measure the
transfection efficiencies.
2.2.2.2.2 Antibiotic treatment
An antibiotic titration was performed using G418 to determine the amount of antibiotic
required to kill all RK-13/SIRC cells within 1-2 weeks. RK-13 cells were seeded into a 24 well
plate and different concentrations of G418 were applied to the wells in duplicate. The
concentrations of G418 trialled were 0.25 mg/ml, 0.375 mg/ml, 0.5 mg/ml, 0.75 mg/ml, 1
mg/ml, 1.5 mg/ml and 2 mg/ml. The cells were washed every 3 days, after which fresh MEM
and G418 was applied. The cells were monitored for two weeks to determine the G418
concentration that killed all cells between 10-12 days.
2.2.2.3 Infections
RK-13 cells were infected with different rabbit sera which were either positive or negative
for RHDV (Table 2-11). A variety of different dilutions of the K31 serum (1:3, 1:5 and 1:10)
was used during this study as the original concentration of K31 caused cell death in >90% of
cells after transfection.
RK-13 cells were seeded into 6 or 12 well plates one day prior to infection so that they were
approximately 60% confluent (for RNA extraction) or 80% confluent (for
immunofluorescence staining) the following day. Before infection the cell culture medium
was removed and cells were washed once wit PBS (1 ml of PBS for 12 well plates and 2 ml of
PBS for 6 well plates). After cells were washed the sera were applied (RHDV positive and
RHDV negative sera were applied to separate sides of the plates to minimise the chance of
contamination). 200 µl of serum was applied to wells on the 12 well plates and 400 µl of
serum was applied to wells on the 6 well plates. 1:3, 1:5 and 1:10 dilutions of serum K31
were used. After the sera were applied plates were incubated at 37°C in 5% CO2
atmosphere for 45 min – 1 hrs, after which cells were washed once with PBS and new MEM
was applied (FCS 2%). Cells were incubated for additional 16 hrs before being analysed
further (2.2.2.5, 2.2.2.6).
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2.2.2.4 Cell treatments
A variety of different treatments were applied to RK-13 during this study in an attempt to
generate an immune response. The different cell treatments are listed below.
2.2.2.4.1 dsRNA transfection
During this study 6 and 12 well plates of RK-13 cells were transfected with dsRNA as
described for plasmid transfections.using 2.5 – 5 µl of dsRNA and Lipofectamine2000. The
amount of lipofectamine2000 used is indicated below.
6 well plate 12 well plate
dsRNA (2.5/5µl) + FCS-free MEM (250µl) dsRNA (2.5µl) + FSC free MEM (100µl)
Lipofectamine (10µl) + FSC free MEM (250µl) Lipofectamine (4µl) + FSC free MEM (100µl)
Table 2-26
2.2.2.4.2 Treatment of cells with dsRNA and IFNα
RK-13 cells were treated with dsRNA and IFN in 6 and 12 well plates. 20 µl of dsRNA was
applied in a drop wise fashion directly to the cell medium or IFNα (500 U/ml) of was used
(also applied directly to the cell culture medium). Treated cells were incubated at 37°C in 5%
CO2 atmosphere over night (approx 16 – 20 hrs) before being used in further experiments
(2.2.2.5, 2.2.2.6)
2.2.2.5 Cellular RNA extraction
Cellular RNA was extracted from RK-13 cells that had undergone a variety of different
treatments (2.2.2.3, 2.2.2.4) in 6 well plates. RNeasy RNA extraction Kit (QIAGEN) was used
for all RNA extractions and the manufacturers’ procedure was followed. The cell culture
medium was removed and all cells were washed twice with 2 ml of PBS, prior to application
of 600 µl cell lysis buffer (RLT with β-Mercaptoethanol, 10 μl/ml). Cells were scrapped from
the wells and collected in 1.5ml centrifuge tubes. Cell lysates were homogenised using
QIAshredder spin columns before undergoing RNA extraction. RNA was extracted using the
buffers and columns provided by the manufacturer and following the manufacturer’s
protocol. RNA samples were kept at -20°C until required (2.2.1.10).
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2.2.2.6 Immunofluorescence staining
Prior to immunofluorescence staining, RK-13 cells were seeded onto cover slips in 12 well
plates and treated with dsRNA or IFNα as described above (2.2.2.3, 2.2.2.4). Cells were
washed with PBS, fixed via incubation using 2,5% formaldehyde in PBS for 10 min at room
temperature and subsequently permeabilised by applying 1 ml of 0.25% Triton-X100 in PBS
for 10 min at room temperature and washed twice with PBS. 0.5 ml FCS (cells being stained
for Vp60) or NGS (cells being stained for Mx1) was then applied to and left on cells for 30
min before being removed. Cells were washed with PBS and glass cover slides were carefully
removed from wells and place face (cell side) down onto 40 µl of primary antibody (Table
2-13) on parafilm. Cells on the cover slides were kept in the dark for 1 hr before being
placed back in the wells and washed 3 times with PBS. 200 µl of secondary antibody (Table
2-14) was then applied directly to wells and plates were wrapped in aluminium foil (to
prevent light from reaching the cells) and placed on a plate shaker (Ratek Instruments,
model no.MPS1) for 45 min. The secondary antibody was then removed and cells were
washed once before 200 µl of DAPI (1:40,000 (w/vol)) was applied to the wells. Cells were
incubated for exactly 1.5 min before the DAPI was removed and cells were washed 5 times
with PBS, leaving PBS on cells for 1 min each wash. H2O was used to wash cells and excess
H2O was blotted using a clean tissue, before the cover slides were mounted cell bearing side
down to glass slides using Fluoromount (Invitrogen). Slides were stored over night at 4°C
over night to allow Fluoromount to solidify.
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3 Results
3.1 Generation of cells with a compromised type I IFN response
3.1.1 Construction of pcDNA3.1 Mx1 Promoter Plasmid
We chose to generate the pcDNA3.1 Mx1 promoter plasmid (pcDNA3.1-Mx1P) by replacing
the CMV (Cytomegalovirus) promoter in the pcDNA3.1/V5-His-Mx1P plasmid with the
Murine Mx1 promoter from the pGL3-Mx1P-Luc plasmid. To do that, the CMV and Mx1
promoter had to be excised from the plasmids using MfeI/HindIII and EcoRV/HindIII,
respectively. MfeI and EcoRV are producing complementary overhangs allowing site specific
insertion of the Mx1 promoter into the pcDNA3.1/V5-His plasmid. The Mx1 promoter could
then be ligated into the pcDNA3.1/V5-His plasmid as described in section 2.2.1.7and
transformed into E.coli (2.2.1.8).
Figure 3-1 Vector map of pcDNA3.1/V5-His-Mx1P plasmid, an mammalian expression vector, containing a cytomegalovirus (CMV) promoter, an ampicillin resistance gene and a neomycin resistance gene (Invitrogen).
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3.1.1.1 Verification of plasmids
The pcDNA3.1/V5-His (kind gift from June Liu, Ecosystems Sciences, CSIRO) and the pGL3-
Mx1P-Luc (kind gift from Georg Koch, Freiburg/Germany) were retransformed in E.coli and
appropriate clones were selected after DNA preparation and restriction digestion analysis
(Table 3-1, Figure 3-2).
Plasmid RE Cutting sites (bp) Expected Fragments
(bp)
pcDNA3.1/V5-His SnabI
HindIII
590
902
312, 5188
pGL3-Mx1P plasmid EcoRI
HindIII
60 (approx)
2328
2268, 5000
Table 3-1 Restriction enzymes (RE) used to digest pcDNA3.1/V5-His and pGL3-Mx1P. Cutting sites in the plasmids and expected fragment sizes are also shown.
Figure 3-2 Restriction digest of pcDNA3.1/V5-His and pGL3-Mx1P as described in section 2.2.1.1 with indicated restriction enzymes (Table 3-1). Fragments were separated using a 1% TBE agarose gel (0.01% SYBR safe DNA stain), visualised under UV illumination. 1a – restriction digest of maxi prep of pcDNA3.1/V5-His using SnabI and HindIII, S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the right), 1b – 5b restriction digest of E.coli colonies tested for to contain pGL3_Mx1P_Luc.
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Restriction digest of the prepared pcDNA3.1/V5-His plasmid (QIAGEN Maxi Prep kit) showed
one band at the expect size (300bp) and one band slightly higher than expected (5,500-
8,000), which, presumably, is a result of the high amount of plasmid loaded onto the
agarose gel. Restriction digest of DNA preparations from E.coli transformed with pGL3-
Mx1P-Luc showed one positive clone (4b), with fragments of the expected sizes (Figure 3-2).
These positive clones were further propagated to amplify sufficient amounts of plasmid for
subsequent experiments (Qiagen maxi prep kit).
3.1.1.2 Preparation Mx1 Promoter Fragment/Vector Backbone
The prepared and verified preparations of pGL3-Mx1P plasmid and pcDNA3.1/V5-His
plasmid were digested in a preparative scale to generate the Mx1 promoter insert and to
linearise pcDNA3.1/V5-His vector plasmid and to remove the CMV promoter upstream of
the multiple cloning site. The restriction endonucleases used are listed below.
Plasmid RE Cutting sites (bp) Expected Fragments
(bp)
pcDNA3.1/V5-His MfeI
HindII
161 (downstream of CMV
promoter)
902 (upstream of CMV promoter)
741 CMV promoter
4759 plasmid backbone
pGL3-Mx1P plasmid EcoRI
HindIII
60 (approx) (downstream of Mx1
promoter)
2328 (upstream of Mx1 promoter)
2268 Mx1 promoter
5000 plasmid backbone
Table 3-2 Restriction enzymes (RE) used to digest pcDNA3.1/V5-His and pGL3-Mx1P. Cutting sites in the plasmids and expected fragment sizes are also shown.
The digestion products were run on an agarose gel and the Mx1 promoter and the linearised
pcDNA3.1/V5-His (minus CMV promoter) were excised (Figure 3-3). The DNA fragments
were purified using the Qiagen Gel extraction kit, as described (2.2.1.3).
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Figure 3-3 Restriction digest of pcDNA3.1/V5-His and pGL3-Mx1P as described in section 2.2.1.1 with the indicated restriction enzymes (Table 3-2). 1a – Restriction digest of pcDNA3.1/V5-His to linearise plasmid and remove pCMV, 1b – Restriction digest of pGL3-Mx1P performed to obtain Mx1 promoter from plasmid, S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the left). The linearised pcDNA3.1/V5-His plasmid and the Mx1 promoter were extracted from the 1% TBE gel using a sterile scalpel before photograph was taken.
3.1.1.3 Preparation of Mx1 promoter plasmid
After being excised from the agarose gel, the vector plasmid was dephosphorylated (2.2.1.4)
to minimise self ligation. To create the pcDNA3.1-Mx1P plasmid, the Mx1 promoter (150ng)
was inserted into the linear pcDNA3.1/V5-His plasmid (104ng) using the T4 DNA ligase
(2.2.1.7). Chemically competent E.coli were transformed with 5 µl of the ligation reaction
and plated on LB agar containing ampicillin (2.2.1.8). The bacteria were grown up over night
at 37°C and DNA was prepared from 16 of the obtained clones (Qiagen mini prep kit).
Identification of positive pcDNA3.1-Mx1P clones was done via digestion as described
(2.2.1.1) using the restriction endonuclease given in Table 3-3.
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Plasmid RE Cutting sites (bp) Expected Fragments
(correct orientation)
(bp)
pcDNA3.1-Mx1P XbaI Approx 600bp after start of Mx1
promoter
Approx 50bp after end of Mx1
promoter
Approx 1750bp
Approx 5400bp
Table 3-3 Restriction sites and expected fragment sizes of a restriction digest of the pcDNA3.1-Mx1P plasmid with XbaI.
Figure 3-4 Analytical restriction digest (2.2.1.1) of pcDNA3.1-Mx1P clones using the restriction endonuclease XbaI. Fragments were separated using a 1% TAE agarose gel (0.01% SYBR safe DNA stain) and visualised under UV illumination. 1-16 – Analytical Restriction digest of pcDNA3.1-Mx1P clones, all clones showed fragments close to the approximated sizes, S – GeneRuler 1kb DNA ladder (relative fragment sizes are indicated on the left).
All restriction digests of DNA preparations from E.coli transformed with pcDNA3.1-Mx1P
plasmid resulted in fragments of the expected sizes, indicative of pcDNA3.1-Mx1P plasmids
containing the Mx1 promoter in the correct orientation. Plasmid pcDNA3.1-Mx1P was
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prepared from clones 9 (pcDNA3.1-Mx1P-9) and 13 (pcDNA3.1-Mx1P-13) (chosen randomly)
using the Qiagen maxi prep kit.
PcDNA3.1-Mx1P-9 and pcDNA3.1-Mx1P-13 were then sequenced (AGRF) using the primers
given in Table 3-4 to confirm that the plasmids contained the Mx1 promoter in the correct
orientation. The pGL3-Mx1P was also sequenced to obtain the Mx1 promoter sequence to
verify the absence of point mutations in the pcDNA3.1-Mx1P constructs.
DNA Primer Area sequenced
pcDNA3.1-Mx1P-13 pcDNA3.1_98fw Segment of vector plasmid
before Mx1 promoter and
beginning of Mx1 promoter.
pcDNA3.1-Mx1P-13 pcDNA3.1_1039rev Segment of vector plasmid
after Mx1 promoter and end of
Mx1 promoter.
pcDNA3.1-Mx1P-9 pcDNA3.1_98fw Segment of vector plasmid
before Mx1 promoter and
beginning of Mx1 promoter.
pcDNA3.1-Mx1P-9 pcDNA3.1_1039rev Segment of vector plasmid
after Mx1 promoter and end of
Mx1 promoter.
pGL3-Mx1P pGL3_4967fw Segment of vector plasmid
before Mx1 promoter and
beginning of Mx1 promoter.
pGL3-Mx1P pGL3_123rev Segment of vector plasmid
after Mx1 promoter and end of
Mx1 promoter.
Table 3-4 Plasmids and primers sent to agrf for sequencing. Also shows the regions of the DNA plasmids that should be sequenced using the chosen primers.
Analysis of the obtained sequences showed that both pcDNA3.1-Mx1P-9 and pcDNA3.1-
Mx1P-13 contained the Mx1 promoter in the correct ordination.
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3.1.2 Construction of TK1 Expression Plasmid
The pcDNA3.1-Mx1P-TK1 construct (pcDNA3.1-Mx1P-TK1) (Figure 3-5) was generated by
inserting the TK1 gene from the pBluescript plasmid (kind gift from Michael Frese, University
of Canberra) into the pcDNA3.1-Mx1P-9/13 plasmids downstream from the Mx1 promoter.
The TK1 gene was excised from the pBlusecript plasmid using BglII and DraI/EarI and over
hangs were removed/filled in using T4 DNA polymerase (2.2.1.5). The vector plasmids
pcDNA3.1-Mx1P-9/13 was linearised using EcoRV, which creates blunt ends. The TK1 could
then be ligated into the pcDNA3.1/V5-His plasmid as described section 2.2.1.7 and
transformed into E.coli (2.2.1.8).
Figure 3-5 Vector map of pcDNA3.1/V5-His-Mx1P-TK1 plasmid, an expression vector for TK1 ‘death’ gene, under the control of the Murine Mx1 promoter. Vector also contains ampicillin and neomycin resistance gene
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3.1.2.1 Preparation TK1 Gene Fragment/Mx1 Promoter Plasmid
The prepared and verified preparations of pcDNA3.1-Mx1P-9, pcDNA3.1-Mx1P-13 and pKO-
Scrambler-TK1 plasmids were digested in a preparative scale to generate the TK1 gene
insert and to linearise pcDNA3.1-Mx1P vector plasmids as described (2.2.1.1) using the
restriction endonucleases given in Table 3-5.
Plasmid RE Cutting sites (bp) Expected size of fragment
containing TK1 gene (bp)
pKO-Scrambler-TK1 BglII 328 before start of TK1
gene
517 after TK1 polyA tail
Approx 2500
pKO-Scrambler-TK1 DraI
EarI
225 before start of TK1
gene
458 after TK1 polyA tail
Approx 2300
pcDNA3.1-Mx1P EcoRV 2753 7150
Table 3-5 Restriction enzymes used to digest pcDNA3.1/V5-His and pGL3-Mx1P. Respective restriction sites in the plasmids and expected fragment sizes are also shown.
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Figure 3-6 Preparative restriction digest of pBluescript-TK1 as described in section 2.2.1.1 with indicated restriction enzymes (Table 3-5). Fragments were separated using a 1% TBE agarose gel (0.01% SYBR safe DNA stain) and visualised under UV illumination. 1 – Restriction digest of pBluescript_TK1 using BglII, performed to obtain to obtain TK1 gene 2– Restriction digest of pBluescript_TK1 using DraI and EarI, performed to obtain TK1 gene, 4 – Undigested pBluescript_TK1, S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the left).
Separation of the pBluescript-TK1 restriction digestion products (TBE agarose gel) indicated
fragments of the expected sizes for both digestion reactions. The TK1 gene fragments were
excised from the gel using a sterile scalpel and DNA was purified using a Qiagen Gel
extraction kit, as described (2.2.1.3).
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Figure 3-7 Preparative Restriction digest of pcDNA3.1_Mx1_(3/9) as described in section 2.2.1.1 with indicated restriction enzymes (Table 3-5). Fragments were separated using a 1%TBE agarose gel (0.01% SYBR safe DNA stain) and visualised under a UV illumination. 1 – Undigested pcDNA3.1-Mx1P-13, 2– Undigested pcDNA3.1-Mx1P-9, 3 – Restriction digest of pcDNA3.1-Mx1P-13 using EcoRV, to obtain linear vector plasmid, 4 – Restriction digest of pcDNA3.1-Mx1P-9 using EcoRV, to obtain linear vector plasmid, S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the left).
The pcDNA3.1-Mx1P clones 3 and 9 were digested using restriction endonuclease EcoRV
(produces blunt ends). The linearised plasmids were separated on a gel and excised using a
sterile scalpel. The DNA was then extracted from the gel and purified using a Qiagen Gel
extraction kit, as described (2.2.1.3).
3.1.2.2 Preparation of TK1 expression plasmid
The overhangs on the TK1 gene fragments were filled prior to ligation using T4 DNA
polymerase as described section 2.2.1.5., to create blunt ends. The vector plasmids
(pcDNA3.1-Mx1P-(3/9)) were also dephosphorylated (2.2.1.4) to minimise the occurrence of
self ligation during ligation. The TK1 gene was then ligated into the vector plasmid using T4
DNA ligase (rapid ligation protocol) as described in section 2.2.1.7. Chemically competent
E.coli were transformed with 5 µl of the ligation reaction and plated on LB agar containing
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ampicillin (2.2.1.8). The bacteria were grown up over night at 37°C and the following day
clones were selected (Table 3-6) and DNA was prepared using Qiagen mini prep kit.
Vector backbone TK1 fragment
(digest)
Number of clones
obtained
Number of clones
selected
pcDNA3.1-Mx1P-13 BglII 3 1
pcDNA3.1-Mx1P-13 EarI/DarI 6 4
pcDNA3.1-Mx1P-9 BglII Approx 20 - 50 10
pcDNA3.1-Mx1P-9 EarI/DarI Approx 20 - 50 9
Table 3-6 Number of E.coli obtain and selected after transfection with pcDNA3.1-Mx1P constructs.
Identification of pcDNA3.1-Mx1P-TK1 plasmids which contained the TK1 gene in correct
orientation was done via digestion (2.2.1.1) using the restriction endonuclease given in
Table 3-3. A restriction endonuclease that cuts in the vector backbone and in the TK1 gene
was chosen so that the orientation of the inserted TK1 gene could be determined.
Plasmid RE Expected
Fragments (correct
orientation)
Expected Fragments
(incorrect correct
orientation)
Expected Fragments
(empty vector)
pcDNA3.1-Mx1P-
TK1
XbaI 146, 590, 600, 1400,
6630
146, 590, 1400, 2120,
5050
146, 590, 1400, 5140
Table 3-7 Restriction enzyme (RE) used for the analytical digest of pcDNA3.1-Mx1P-TK1. Restriction sites in the plasmid and expected fragment sizes are shown.
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Figure 3-8 Analytical restriction digest of a selected pcDNA3.1-Mx1P-TK1 clones (2.2.1.1) using restriction endonuclease SacI. Fragments were separated using a 1% TBE agarose gel (0.01% SYBR safe DNA stain), visualised under a UV illumination. 1 – undigested pBluescript_TK1 plasmid, 2 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-13, TK1 digested with BglII), 3 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-13, TK1 digested with Eari and DraI), 4 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-13, TK1 digested with BglII), 5 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-9, TK1 digested with BglII), 6 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-9, TK1 digested with BglII), 7 - pcDNA3.1-Mx1P-TK1 clone (vector backbone pcDNA3.1-Mx1P-9, TK1 digested with BglII), S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the right).
The restriction digests of pcDNA3.1-Mx1P-TK1 rows 4 and 7 (Figure 3-8) showed fragments
of the expected size for positive clones that contained the TK1 in the correct orientation.
One of the positive clones contained pcDNA3.1-Mx1P-13 as the vector backbone and the
TK1 gene was digested using EarI and DraI (pcDNA3.1-Mx1P-TK1-1) and the other contain
pcDNA3.1-Mx1P-9 as the vector backbone and the TK1 gene was digested using BglII
(pcDNA3.1-Mx1P-TK1-2). DNA preparations of these clones were performed using a Qiagen
maxi prep kit.
PcDNA3.1-Mx1P-TK1-1 and pcDNA3.1-Mx1P-TK1-2 were sent to agrf for sequencing using
primers given in Table 3-8, to confirm that the plasmids contained the TK1 gene in the
correct orientation and with the correct sequence.
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DNA Primer Area sequenced
pcDNA3.1-Mx1P-TK1-1 TK1-int194-fw Internal section of TK1 gene
pcDNA3.1-Mx1P-TK1-1 pcDNA3.1_1039rev Segment of vector plasmid
after the Tk1 gene and end of
the TK1 gene.
pcDNA3.1-Mx1P-TK1-2 TK1-int194-fw Internal section of TK1 gene
pcDNA3.1-Mx1P-TK1-2 pcDNA3.1_1039rev Segment of vector plasmid
after the Tk1 gene and end of
the TK1 gene.
Table 3-8 Plasmids and primers sent to agrf for sequencing. Also shows the regions of the DNA plasmids that should be sequenced using the chosen primers.
The sequencing results obtained from agrf showed that both plasmids contained the TK1
gene and that the gene had been inserted in the correct orientation.
3.1.3 Construction of DTA Expression Plasmid
The construction of the DTA expression plasmid (pcDNA3.1-Mx1P-DTA) took longer than
originally planned and was successfully generated after the TK1 plasmid. The generation of
the Mx1 promoter plasmid used as the backbone plasmid was the same for the DTA and the
TK1 expression plasmids (3.1.1.3).
The pcDNA3.1-Mx1P-DTA construct (pcDNA3.1-Mx1P-DTA) (Figure 3-9) was generated by
inserting the DTA gene from the pKO-scrambler-DTA plasmid (kind gift from Michael Frese,
University of Canberra) into the pcDNA3.1-Mx1P-9/13 plasmid downstream from the
murine Mx1 promoter. Large amounts of the DTA gene were generated from the pKO-
scrambler-DTA plasmids via preparative scale PCR, using the DTA_cl_fwNot1 and
DTA_cl_revNot1 primers. The DTA PCR products were then digested using restriction
endonuclease NotI and over hangs were removed/filled in using T4 DNA polymerase
(2.2.1.5). The vector plasmids pcDNA3.1-Mx1P-9/13 was linearised using EcoRV (same as
outlined in TK1 expression generation approach 3.1.2.1), which creates blunt ends. The DTA
could then be ligated into the pcDNA3.1/V5_His plasmid as described section 2.2.1.7 and
transformed into E.coli (2.2.1.8).
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Figure 3-9 Vector map of pcDNA3.1-Mx1 promoter-DTA plasmid, expression vector for DTA ‘death’ gene, under the control of the Murine Mx1 promoter. Vector also contains ampicillin and neomycin resistance gene.
3.1.3.1 Preparation DTA Gene Fragment/Mx1 Promoter Plasmid
The Mx1 promoter plasmids (pcDNA3.1-Mx1P-9/13) were prepared in the same manner as
describe in section 3.1.2.1 and will not be described again in this section.
The DTA gene was amplified via preparative PCR, using using the DTA_cl_fwNot1 and
DTA_cl_revNot1 primers. The PCR products were then purified using QIAquick PCR
purification kit (2.2.1.6). The DTA PCR products were the digested to generate the DTA gene
insert as described (2.2.1.1) using the restriction endonuclease given in Table 3-5Table 3-9.
Template DNA RE Expected fragment size (bp)
DTA PCR product NotI Approximately 1000
Table 3-9 Restriction enzymes used to DTA PCR products, generated using DTA_cl_fwNot1 and DTA_cl_revNot1 primers. The expected fragment size is also shown.
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Figure 3-10 Preparative Restriction digest of DTA PCR products as described in section 2.2.1.1 with indicated restriction enzymes (Table 3-9). Fragments were separated using a 1%TBE agarose gel (0.01% SYBR safe DNA stain) and visualised under a UV illumination. 1-2 – Digested DTA PCR product, to obtain DTA gene, S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the left).
The digested DTA products were run on a agarose gel and excised using a sterile scalpel. The
DNA was then extracted from the gel and purified using a Qiagen Gel extraction kit, as
described (2.2.1.3).
3.1.3.2 Preparation of DTA expression plasmid
The overhangs on the DTA gene fragments were filled prior to ligation using T4 DNA
polymerase as described section 2.2.1.5 to create blunt ends. The vector plasmids
(pcDNA3.1-Mx1-(3/9)) were also dephosphorylated (2.2.1.4) to minimise the occurrence of
self ligation during ligation. The DTA gene was then ligated into the vector plasmid using T4
DNA ligase (rapid ligation protocol) as described in section 2.2.1.7. E. coli XL1blue were
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transformed with 5 µl of the ligation reaction and plated on LB agar containing ampicillin
(2.2.1.8). The bacteria were grown up over night at 37°C and the following day clones were
selected (Table 3-6) and DNA was prepared using Qiagen mini prep kit.
Identification of pcDNA3.1-Mx1P-DTA plasmids which contained the DTA gene in correct
orientation was performed via digestion (2.2.1.1) using the restriction endonuclease given in
Table 3-3. A restriction endonuclease that cuts in the vector backbone and in the TK1 gene
was chosen so that the orientation of the inserted TK1 gene could be determined.
Plasmid RE Expected Fragments (correct orientation)
pcDNA3.1-Mx1P-DTA NocI 735, 2123, 2475, 3035
Table 3-10 Restriction enzyme (RE) used for the analytical digest of pcDNA3.1-Mx1P-DTA. Restriction sites in the plasmid and expected fragment sizes are shown.
Figure 3-11 Analytical restriction digest of a selected pcDNA3.1-Mx1P_DTA clones (2.2.1.1) using restriction endonuclease NocI. Fragments were separated using a 1% TBE agarose gel (0.01% SYBR safe DNA stain), visualised under a UV illumination. 1 – digested pcDNA3.1-Mx1P_DTA (clones 8/24) plasmid (expected fragment sizes on the right), S – GeneRuler 1kb DNA ladder (relative sizes in basepairs (bp) indicated on the left).
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The restriction digests of pcDNA3.1-Mx1P-DTA clones 8 and 24 (Figure 3-12Figure 3-8)
showed fragments of the expected size for positive clones that contained the DTA in the
correct orientation. DNA preparations of these clones were performed using a Qiagen maxi
prep kit.
3.1.4 Generation of RK-13 clones
The antibiotic G418 was used in this study to select for cells that were successfully
transfected with DTA/TK1 expression plasmids generated during this study (pcDNA3.1-
Mx1P-TK1 and pcDNA3.1-Mx1P-DTA). An antibiotic titration was performed to determine
the concentration of G418 at which all untransfected RK-13 and SIRC cells died within a 7 –
14 day period. To do that, RK-13 and SIRC cells were seeded in 24 well plates (approx 100%
confluent) and treated with different concentrations of G418 (0.25, 0.37, 0.5, 0.75, 1, 1.5
and 2 mg/ml cell culture supernatant). At a concentration of 1 mg/ml, G418 killed both RK-
13 and SIRC after approximately nine days and was used at this concentration in future
experiments.
100mm cell culture dishes of RK-13 cells (approx 60% confluent) were transfected in
duplicate with the final plasmid constructs pcDNA3.1-Mx1P-TK1-1, pcDNA3.1-Mx1P-TK1-2,
pcDNA3.1-Mx1P-DTA-8 and pcDNA3.1-Mx1P-DTA-24. Approximately 24 µg of DNA was used
for the transfection and a non-transfected plate was kept as a control for the antibiotic
selection process. One plate was transfected with peGFP-N1 was used to measure the
transfection efficiency. Fluorescence was noticeable in 20-30% of cells transfected with
peGFP-N1.
The transfected RK-13 cells and the control RK-13 cells were treated with G418 one day post
transfection to select for successfully transfected cells.
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Days post
G418
treatment
TK1_1 TK1_2 DTA_8 DTA_24 Control
Starting
Confluency
(pre G418
treatment)
90% 100% 100% 100%
2 days 100% confluent 100% confluent 100% confluent 100% confluent 100% confluent
3 days 40% confluent 50% confluent 70% confluent 60% confluent 30% confluent
4 days 40% confluent 40% confluent 70% confluent 70% confluent 20% confluent
6 days 5-10% confluent 10% confluent 90% confluent
(most cells rounded)
80% confluent
(most cells rounded)
<10% confluent
(all cells rounded)
10 days 10-20% confluent
(growing in
colonies)
10-20% confluent
(growing in
colonies)
70% confluent
(most cells rounded)
60% confluent
(most cells rounded)
All cells dead
11 days 40% growing in
colonies
30% growing in
Colonies
40% confluent
(Over 80% of cells
rounded)
50%
(Over 80% of cells
rounded)
13 days 80% confluent 70-80% confluent 70% confluent 80% confluent
15 days 100% confluent 100% confluent 100% confluent 100% confluent
Table 3-11 Shows confluency of RK13 cells transfected with pcDNA3.1-Mx1P-TK1-1, pcDNA3.1-Mx1P-TK1-2, pcDNA3.1-Mx1P-DTA-8 and pcDNA3.1-Mx1P-DTA-24 plasmids and treated with 1 mg/ml G418 over a 15 day period.
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Days post G418
treatment
TK1_1 TK1_2 DTA_24
Starting confluency
(pre G418 treatment)
50% 40% 50%
2 days 20-30% confluent 10% confluent 60% confluent
(some rounded cells)
4 days 10-20% confluent <5% confluent 60% confluent
(Most cells rounded)
7 days 10% confluent
(growing in colonies)
<5% confluent
(growing in colonies)
30% confluent
(Most cells rounded)
9 days 20% confluent
(growing in colonies)
10-20% confluent
(growing in colonies)
30% confluent
(Most cells rounded)
10 days 30% confluent
(growing in colonies)
20% confluent
(growing in colonies)
20% confluent
(Most cells rounded)
13 days 60-70% confluent 70% confluent 60% confluent
15 days 100% confluent 100% confluent 90% confluent
Table 3-12 Shows confluency of RK13 cells transfected with pcDNA3.1-Mx1P-TK1-1, pcDNA3.1-Mx1P-TK1-2, and pcDNA3.1-Mx1P-DTA-8 plasmids and treated with 1 mg/ml G418 over a 15 day period.
All of the RK13 cells in the control plate were killed in approximately 10 days post G418
treatment. After approximately 6-7 days RK-13 cells transformed with TK1 expression
plasmids reached their lowest confluency (<10%) and formed colonies. RK13 transformed
with DTA expression plasmids did not reach confluency lower then approximately 30%.
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3.2 Characterisation of Innate Immune Response
3.2.1 Real-Time Polymerase Chain Reaction Analysis
3.2.1.1 Optimisation of Real-Time Polymerase Chain Reaction
3.2.1.1.1 Optimisation of Mx1 primer sets
Figure 3-12 a) amplification of dsRNA transfected RK13 cells cDNA using Mx1 primer set rMx1_522fw/rMx1_720rev (0.3µl) over a temperature gradient of 55-65°C b) melt curves for data shown in a), c) amplification of dsRNA transfected RK13 cDNA using Mx1 primer set rMx1_522fw/rMx1_720rev (0.5µl) over a temperature gradient of 55-65°C and d) melt curves for data shown in c).
The temperature that produced optimal amplification for Mx1 primer set 1
(rMx1_522fw/rMx1_720rev) was 61°C and temperatures below 57°C resulted in slightly less
specific amplification (difference in melting peak seen Figure 3-12 b)). Using a primer
concentration of 500 nM resulted in products with a more consistent melt temperature
than reactions using a primer concentration of 300 nM.
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Figure 3-13 a) amplification of dsRNA transfected RK13 cells cDNA using primer set Mx1 primer set rMx1_51fw / rMx1_203rev (0.3µl) over a temperature gradient of 55-65°C b) melt curves for data shown in a), c) amplification of dsRNA transfected RK13 cDNA using Mx1 primer set rMx1_51fw / rMx1_203rev (0.5µl) over a temperature gradient of 55-65°C and d) melt curves for data shown in c).
No large difference in amplification was observed over the annealing temperature gradient
trialled, for Mx1 primer set 2 (rMx1_51fw / rMx1_203rev) using 300 nM of primer. A slightly
larger difference was observed when using 500 nM of the primers, although this also in the
production of a second product (two melting temperatures were observed Figure 3-13 d)).
Amplification appeared to be slightly better between 59°C and 61.4°C for Mx1 primer set 2.
Mx1 primer set 1 (using 500 nM of primer) was chosen to measure Mx1 expression in all
subsequent RT-PCR experiments.
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3.2.1.1.1 Optimisation of GAPDH primer sets
Figure 3-14 a) amplification of dsRNA transfected RK13 cells cDNA using GAPDH primer set rGAPDH1_fw_278/rGAPDH1_rev_425 (0.3µl) over a temperature gradient of 55-65°C b) melt curves for data shown in a), c) amplification of dsRNA transfected RK13 cDNA using GAPDH primer set rGAPDH1_fw_278/rGAPDH1_rev_425 (0.5µl) over a temperature gradient of 55-6°C and d) melt curves for data shown in c).
The optimal amplification temperature observed for GAPDH primer set 1
(rGAPDH1_fw_278/rGAPDH1_rev_425) was 61°C for both concentrations of primers trailed
(300 nM and 500 nM). 500 nM of primer produced a more consistent amplification over the
trialled temperature range.
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Figure 3-15 a) amplification of dsRNA transfected RK13 cells cDNA using GAPDH primer set rGAPDH2_fw_470/ rGAPDH2_rev_633 (0.3µl) over a temperature gradient of 55-65°C b) melt curves for data shown in a), c) amplification of dsRNA transfected RK13 cDNA using GAPDH primer set rGAPDH2_fw_470/ rGAPDH2_rev_633 (0.5µl) over a temperature gradient of 55-65°C and d) melt curves for data shown in c).
The optimal amplification temperature for GAPDH set 2
(rGAPDH2_fw_470/rGAPDH2_rev_633) was also 61°C and more consistent amplification
across the temperature range was obtained by using a higher concentration of the primers
(500 nM).
GAPDH primer set 1 was slightly more sensitive then GAPDH primer set 2 (had lower Cq
values) and was therefore used in all subsequent RT-PCR reactions.
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3.2.1.1.1 Optimisation of Ip10 primer sets
Figure 3-16 a) amplification of serum 453 transfected RK13 cells cDNA using Ip10 primer set rIP10_299fw / rIP10_446rev (0.3µl) over a temperature gradient of 55-65°C b) melt curves for data shown in a), c) amplification of dsRNA transfected RK13 cDNA using primer set rIP10_299fw/rIP10_446rev (0.5µl) over a temperature gradient of 55-65°C and d) melt curves for data shown in c).
The temperature range for Ip10 primer set 1 (rIP10_299fw / rIP10_446rev) that produced
optimal amplification as 55 - 61°C. Using a lower concentration of the primers (300 nM)
produced a more consistent of amplification across the annealing temperatures trialled.
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Figure 3-17 a) amplification of serum 453 transfected RK13 cells cDNA using Ip10 primer set rIP10_55fw/rIP10_237rev (0.3µl) over a temperature gradient of 55-65°C and b) melt curves for data shown in a).
Optimal amplification for Ip10 occurred between 59 - 61°C for Ip10 primer set 2
(rIP10_55fw/rIP10_237rev). Annealing temperatures above and below 59 - 61°C resulted in
noticeable decrease amplification.
Ip10 primer set 1 was used for all subsequent RT-PCR experiments because the product
produced by this primer set had a similar melting temperature to the products produced by
the other primer sets.
3.2.1.2 Type-I IFN response in RK-13 cells
To characterise the type I IFN response in RK13 cells, the rt-PCR assays (3.2.1.1) were used
to measure Ip10, Mx1 and GAPDH expression after treatment with different stimuli. RK13
cells were treated with pIC, IFN or with rabbit serum (2.2.2.4) and RNA was extracted
approximately 16 – 24 hours later (2.2.2.5). The RNA was then reverse transcribed using the
QuantiTect® Reverse transcription Kit (2.2.1.10) to obtained cDNA that could be used in the
rt-PCR assays. This process was performed three times.
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Figure 3-18 Relative increase in Mx1 mRNA observed in RK13 cells treated with dsRNA, IFN and RHDV compared to control samples. The data obtained was analysed using the delta-delta-Cq quantification model.
Figure 3-19 Relative increase in Ip10 mRNA observed in RK13 cells treated with dsRNA, IFN and RHDV compared to control samples. The data obtained was analysed using the delta-delta-Cq quantification model.
The data obtained from the rt-PCR analysis of the RK13 cell line indicated that Mx1 and Ip10
gene expression is up-regulated in response to transfection with dsRNA and that Mx1 gene
-2
0
2
4
6
8
10
12
14
dsRNA(transfected)
IFNa (500IU/ml)
dsRNA applied RHDVcontaining
rabbit serum(K31)
RHDVcontaining
rabbit serum(453)
rela
tive
incr
eas
e in
Mx1
-mR
NA
Treatment
-1
-0.5
0
0.5
1
1.5
2
2.5
3
3.5
4
dsRNA(transfected)
IFNa (500IU/ml)
dsRNA applied RHDVcontaining
rabbit serum(K31)
RHDVcontaining
rabbit serum(453)
rela
tive
incr
eas
e in
Ip1
0-m
RN
A
Treatment
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83
expression is also up-regulated in response to IFN stimulation. There were large differences
in the results obtained for the other treatments (shown by the large error bars) making it
difficult to determine whether the treatment actually causes an up regulation in Ip10 and
Mx1 gene expression.
3.2.1.3 Immunofluorescence Analysis
To determine whether Mx1 protein expression was up-regulated in response to a variety of
different stimuli, an immunofluorescence assay was developed using an Mx1 specific
antibody.
Figure 3-20 Light microscope images of untreated RK13 cells stained with Mx1 specific antibody and DAPI. 1 – Image taken using DIC filter, showing DAPI stain, 2 – Image taken using a brightfield filter set, 3 – Image taken using a GFP filter, Mx1 antibody staining shown in green.
Figure 3-21 Light microscope images of pIC treated RK13 cells stained with Mx1 specific antibody and DAPI. 1 – Image taken using DIC filter, showing DAPI stain, 2 – Image taken using a brightfield filter set, 3 – Image taken using a GFP filter, Mx1 antibody staining shown in green.
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Figure 3-22 Light microscope images of IFN treated RK13 cells stained with Mx1 specific antibody and DAPI. 1 – Image taken using DIC filter, showing DAPI stain, 2 – Image taken using a brightfield filter set, 3 – Image taken using a GFP filter, Mx1 antibody staining shown in green.
Figure 3-23 Light microscope images of untreated (transfected) RK13 cells stained with Mx1 specific antibody and DAPI. 1 – Image taken using DIC filter, showing DAPI stain, 2 – Image taken using a brightfield filter set, 3 – Image taken using a GFP filter, Mx1 antibody staining shown in green.
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Figure 3-24 Light microscope images of pIC transfected RK13 cells stained with Mx1 specific antibody and DAPI. 1 – Image taken using DIC filter, showing DAPI stain, 2 – Image taken using a brightfield filter set, 3 – Image taken using a GFP filter, Mx1 antibody staining shown in green.
No significant difference in fluoresence was observed between the controls and the
treatments, as all samples stained positive for Mx1 protein expression
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4 Discussion
Currently, there is no robust cell culture system available for RHDV, which greatly impedes
studies of this virus. As a result the majority of studies involving RHDV have be done using
an in vivo system (rabbits), which is more expensive and laborious than using an in vitro
alternative. Using animals for studies also involves ethical and regulatory problems, which
can be easily avoided by using a cell culture system. Studies in vitro are also easier to
standardise than studies using animals, as differences between animals, including genetic
differences and differences in infection history and immune status can greatly impact
results. Cell culture systems are commonly used for the study of many aspects of virus
biology including replication cycle, gene function and pathogenicity. Gaining a better
understanding of the biological aspects of RHDV, such as genetic markers associated with
high virulence could also help to improve RHDV mediated biological control strategies
currently used in Australia to contain the impact of rabbits on Australias Flora and Fauna.
RHDV is only one of many viruses belonging to the Calicivirdae family for which a working
cell culture system has not yet been established (Radford, Gaskell et al. 2004). There are
currently no cell culture systems available for any of the caliciviruses affecting humans, even
though the calicivirus family contains several important human pathogens, including human
Noro- and Sapporo viruses (Li, Predmore et al. 2012). Human Norovirus is one of the leading
causative agents of food-borne disease worldwide and currently there is no in vitro system
or small animal model available in which the virus can be studied (Li, Predmore et al. 2012).
As a result most of the insights into human calicivirus pathophysiology have been obtained
through extensive studies using healthy human volunteers (Dolin, Blacklow et al. 1971;
Dolin, Blacklow et al. 1972; Atmar, Opekun et al. 2008; Vashist, Bailey et al. 2009). A cell
culture system for human Norovirus would aid in the development of antiviral drugs,
potential vaccines production and would be a useful diagnostic tool. The development of a
cell culture system for RHDV may provide insights into what is required for the propagation
of other caliciviruses in vitro, such as caliciviruses affecting humans.
A few caliciviruses have been successfully cultured in vitro, including PECV, MNV, FCV and
TCV. An interesting recurring feature in the development of cell culture systems for some of
these virus is need to suppress the type I IFN response (Flynn and Saif 1988; Parwani, Flynn
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87
et al. 1991; Slomka and Appleton 1998; Farkas, Sestak et al. 2008). Suppression of
components of innate immune responses, namely the type I IFN system has been shown to
result in increased viral propagation for many viruses, including Polio Virus, Herpes Simplex
Virus, Hepatitis B Virus, Human immunodeficiency virus and Murine Norovirus (Pierce,
DeSalvo et al. 2005; Barnes, Kunitomi et al. 2008; McCartney, Thackray et al. 2008; Akhtar,
Qin et al. 2010; Li, Hu et al. 2011; Belloni, Allweiss et al. 2012). The stimulation of type I IFN
is the first line of defence cells have against viral infection and results in expression of many
IFN induced proteins known to inhibit viral replication (Samuel 2001). Type I IFNs were
shown to effectively inhibit the propagation of MNV in vitro and in vivo in macrophages and
dendritic cells (Karst, Wobus et al. 2003; Chang, Sosnovtsev et al. 2004; McCartney,
Thackray et al. 2008). It is also believed that type I IFN induced antiviral proteins prevents
PECV propagation in vitro. PECV replication in cell culture depends on the addition of
intestinal filtrate, which subsequently has been shown to suppress the type I and type II
response and enable efficient viral replication (Chang, Sosnovtsev et al. 2004). Bile acids, are
the active compounds found in intestinal filtrate which suppress the type I IFN response
through the up-regulation of cAMP and the activation PKA, but the downstream effectors
have not been indentified (David, Petricoin et al. 1996; Sengupta, Schmitt et al. 1996; Lee
and Rikihisa 1998). This lead to the hypothesis that RHDV growth, that hasn’t been observed
in vitro so far, may be enhanced significantly in cells with a compermised type I IFN
response.
To test the hypothesis the main aim of this study was to develop a method for selecting type
I IFN defective cells and whether this cell line supported the growth of RHDV. The chosen
approach to generate a cell line with a defect in the type I IFN response was based on the
assumption that due to mutations which occur consistently, even though rarely, such cells
will be present in of cultured cells. To select these few cells we generated expression
plasmid containing a death gene under the control of an IFN inducible promoter. The two
plasmids used carried either DTA or TK1 under the control of the murine Mx1 promoter. A
vector plasmid was chosen for the construction of these plasmids that contained a
neomycin resistance gene so to allow for selection of transfected cells using the antibiotic
G418. The mouse Mx1 promoter from the pGL3-Mx1P reporter plasmid (kindly provided by
Georg Koch, Freiburg/Germany) was used as it has been shown to be strongly activated in
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IFN stimulated cells (Jorns, Holzinger et al. 2006). Further. the murine Mx1 promoter shows
extremely low basal activity in unstimulated cells, unlike other Mx promoters, such as the
human MxA promoter (Holzinger, Jorns et al. 2007). DTA and TK1 have been extensively
studied in regard to their ability to induce cell death in tumour or Cytomegalovirus infected
cells. (Yamaizumi, Mekada et al. 1978; Fillat, Carrio et al. 2003).
This approach has the advantage of resulting in the generation of cells that are unable to
mount a type I IFN induced antiviral state, due to inability to stimulate type I IFN inducible
promoters (Samuel 2001). Other methods considered to obtain IFN defective cells, including
RNA interference or suppression of the IFN system via chemical inhibitors only disable the
IFN immune response temporally. On the other hand, the more direct approach of targeted
gene knock-out as routinely done in mice, was considered to be too laborious, expensive
and neglected due to the lack of established procedures for rabbit cells.
After transfection of RK13 cells with one of the two constructs and subsequent selection
with G418, IFN or IFN and ganciclovir will be used to treat cells transfected with the DTA and
TK1 expression plasmids, respectively. The DTA gene was originally chosen as the selector
gene for IFN defective cells, as DTA has been shown to have a very efficient cytopathic on
cells, even at concentrations as low as one DTA molecule per cell (Yamaizumi, Mekada et al.
1978; Bell and Eisenberg 1996). A potential problem of using DTA in this approach is that the
bystander effect of DTA may result in cell death of all cells surrounding cells with intact IFN
signalling pathway. If this turned out to be correct the DTA plasmid would be of no use as a
selector for type I IFN defective cells. To address this problem a TK1 (from herpes simplex
virus) expression plasmid was also constructed in addition to the DTA expression plasmid.
TK1 has also been shown to be capable of causing cell death but unlike DTA, TK1 requires
the addition of a specific substrate ganciclovir to initiate a cytopathic effect (Solaroli,
Johansson et al. 2008). The combination of TK1 and ganciclovir has been used to target
cancer cells in several human trials and in a wide variety of animal tumour models (Fillat,
Carrio et al. 2003; Solaroli, Johansson et al. 2008). Unlike DTA the bystander effects of TK1
can be controled to a certain degree by using different concentrations of IFN/ganciclovir,
which is why it was chosen in case the DTA expression plasmid does not produce any clones.
Orford-Dunne, Roxy (CES, Black Mountain)
89
TK1 requires the addition of a substrate (ganciclovir) in order to cause cell death, whereas
DTA does not (Fillat, Carrio et al. 2003).
As a result of the work performed during this study an expression plasmids for DTA and TK1
were successfully constructed that can be used for the selection of type I IFN defective cell
lines. RK13 cells were effectively transfected with both constructs and the best conditions to
select successfully transfected cells via G418 were determined during this study. This
approach has the advantage of producing large pools of cells with selectable markers
(DTA/TK1). The next step would be to treat the transfected RK13 cells with IFN and
IFN/Ganciclovir and see if cellular death occurred. Cells surviving the selection could then be
infected with RHDV to examine if viral propagation is improved in the absence of a
functional type I IFN response.
The second aim of our study was to verify our hypothesis that the type I IFN system plays a
role in the suppression of RHDV growth in vitro, by characterising the type I IFN-mediated
innate immune response in RK13 cells. We also wanted to generate data describing the
interplay between RHDV infection in RK13 cells and the innate immune response. One way
of gauging the IFN response in RK13 cells was to develop rt-PCR assays that could be used to
measure the level of mRNA expression of different IFN inducible genes, such as Ip10 and
Mx1 (Samuel 2001) in RK13 cells. Primers were developed for Ip10, Mx1 and GAPDH (Table
2-3) and used in preliminary PCR reactions to determine whether the DNA products
produced were of the expected size (data not shown). GAPDH mRNA levels were used as an
endogenous control during this study, as GAPDH expression is constant and should be not
affected by the experimental manipulation performed during this study (Bustin 2000). The
primers were then used in preliminary rt-PCR experiments to optimise conditions including
annealing temperatures and primer concentrations and to choose the primer sets that
produced specific and sensitive amplification. We were able to successfully detect Mx1, Ip10
and GAPDH mRNA in the cDNA samples and optimise the rt-PCR conditions to increase
sensitivity and decrease non-specific product amplification (see section 3.2.1.1).
The rt-PCR assays were then used to characterise the IFN response in RK13 cells. To
measure the IFN response we treated RK13 cells with two separate stimuli known to induce
type I IFN induction, pIC and hybrid human IFN-α (Baum and Garcia-Sastre 2010). RK13 cells
Orford-Dunne, Roxy (CES, Black Mountain)
90
were also infected with two different rabbit sera known to contain RHDV virus and a mock
serum (Table 2-11). RNA was then extracted from the cells. Remarkably, even though a
different stimuli known to induced an IFN response (pIC and IFN) were applied to the cells
(Liu, Sanchez et al. 2012), little induction of IFN-stimulated genes transcription (Mx1, IP10)
was observed, indicating either an intrinsically low ability of RK13 cells to respond to IFN or
a possible contamination of the used cells (viruses, mycoplasma, bacterial).
The rt-PCR assays allowed for the expression of mRNA of type I IFN inducible proteins to be
measured, but do not indicate whether the mRNA is transcribed into proteins. Many viruses
have developed mechanisms that inhibit or suppress various IFN induced antiviral
mechanisms, to allow for increased viral replication. For example Influenza A Virus encodes
a non structural protein NS1, which is capable of blocking posttranscriptional processing of
cellular mRNAs (Kochs, Garcia-Sastre et al. 2007). This decreases the up regulation of IFN
induced antiviral proteins, counteracting the host cells antiviral response and increases viral
replication (Kochs, Garcia-Sastre et al. 2007). The rt-PCR assays do not allow us to determine
whether RHDV was capable of decreasing the antiviral effect of IFN by suppressing antiviral
protein mRNA transcription. To counter act this problem we developed a second assays to
measure the expression of Mx1 proteins in RK13 cells.
An Mx1 specific antibody shown to work previously was used in this study to gauge the level
of Mx1 expression in stimulated RK13 cells. Different concentrations of the Mx1 antibody
and secondary antibodies were trialled to optimise staining conditions before Mx1
characterisation experiments were conducted. An immunofluorescence staining procedure
for Mx1 was successfully established. The Mx1 antibody was shown to specifically stain
Mx1, producing very little background staining. RK13 cells were treated with pIC, IFN and
infected with RHDV containing rabbit serum and mock serum, to determine whether these
treatments caused an up regulation of Mx1.
The results obtained from the Mx1 immunofluorescence were similar to the results
obtained from the rt-PCR experiments. An up regulation of Mx1 protein expression was
observed in the positive controls (pIC and IFN treated cells), the negative controls
(untreated and mock serum infected cells) and the in RHDV infected cells. This again
indicated that either RK13 cells have a low intrinsically response to IFN or that the cell
Orford-Dunne, Roxy (CES, Black Mountain)
91
culture was contaminated and that Mx1 expression was being up regulated in response to
the contamination.
Indicative of a cellular contamination was the fact that unstimulated had a comparable
abundance of Mx1 and IP10 mRNA to the treated cells. Also similar expression levels of Mx1
were observed in the control and treatment groups after staining with the Mx1 antibody.
Previous studies using the same Mx1 specific antibody did not show Mx1 staining in
untreated cells, so it was very unusual that the untreated RK13 were positive for Mx1
expression.
It was speculated that the contrasting findings were mostly likely the result of cell culture
contamination, so Bovine Viral Diarrhea Virus was tested for. BVDV was the first possible
contaminant tested for as is it commonly found in a variety of different cell culture lines
from species including, rabbits, deer, sheep, goats and bovine (Bolin, Ridpath et al. 1994).
Non-cytopathic strains of BVDV virus are capable of persistently infecting cells in culture
without inducing apoptosis or showing signs of viral infection (Schweizer and Peterhans
2001). The non-cytopathic strains of BVDV are frequently found in fetal bovine serum (FBS)
(Molander, Boone et al. 1971), which is commonly used in mammalian cell cultures. A
survey was conducted in the early 90s for the presence of BVDV in a variety of cell lines and
showed positive results of BVDV infection in the RK13 cell line tested, indicating RK13 cells
are susceptible BVDV infection (Bolin, Ridpath et al. 1994). BVDV studies have also shown
that BVDV infection decreases the expression of IFN and suppresses apoptosis in response
to pIC in bovine macrophages (Schweizer and Peterhans 2001).
The RK13 cell line used during this study was tested for BVDV by Andrew Read, Elizabeth
MacArthur Agricultural Institute, NSW and results showed the RK13 cells were positive for
BVDV. This could explain why no signification up regulation of the IFN-stimulated genes
(Mx1 and Ip10) for was observed in the stimulated RK13 cells. To accurately characterise the
type I IFN response in RK13 cells and determine whether they are a good cell choice for the
develop of a RHDV cell culture system a non contaminated batch of RK13 cells would need
to be obtained and tested.
Unfortunately because of time constraints the designed approach to develop a cell culture
system for RHDV could not be completed during this study, we are therefore unable to say
Orford-Dunne, Roxy (CES, Black Mountain)
92
whether or not it would have been successful. If the approached out lined in this study
proved to be unsuccessful, several other approaches could be used as alternatives or in
addition our approach to develop a cell culture system for RHDV.
Several other rabbit cell lines are readily available (such as SIRC cell line) and could be
trialled as an alternative if the RK13 cell line proved to be a poor choice of cell lines. Rabbit
hepatocytes or macrophages may support RHDV growth, as large quantities of the virus are
found in the liver tissue after in vivo infection and macrophages were used to successfully
propagate related MNV in cell culture. Potential cell lines could be screened for factors
other than defectives in the type I IFN response that may enhance the chance of viral
replication occurring. For example Sialic acids (SA) have recently have been identified as
attachment factors for RHDV and MNV (Nystrom et al, 2011, Taube et al., 2009). In the case
of RHDV these SAs are histo blood group antigen associated. Potential cell lines could be
screened in advance to determine whether or not they express the respective Glykosides, in
addition to characterising there type I IFN response. This could increase the likelyhood of
cell types supporting RHDV growth.
Different cell treatments that suppress the IFN mediated antiviral response could also be
trialled if our approach proved to be unsuccessful. Bile acids were shown to be curial for the
growth of PECV in cell culture and are believed to play a role in the down regulation of the
IFN response (ref). In this regard, pre-treatment of virus inoculum with bile extracts should
be tried as well. Cells could be treated with bile acids and then infected with RHDV to see if
this increases virus propagation. Another alternative is the use of siRNAs, which are double
stranded, sequence-specific inhibitors of gene expression, which can be used to down
regulate the type I IFN response (Rossi 2009).
Although the original aims of this study were not met due to time constraints, the
constructs and methods developed will be very helpfully in future attempts to develop a cell
culture system for RHDV and other viruses which do not have a working cell culture system.
As a result of the BVDV contamination the information on the RK13 cell IFN response could
not be obtained, but the methods developed to measure IFN response in cells will help
future efforts to characterise the innate immune response in RK13. The assays developed
can be used to screen for other cell lines if RK13 cells are shown to be a poor choice. Time
Orford-Dunne, Roxy (CES, Black Mountain)
93
constraints prevented us from using the DTA and TK1 expression plasmids to select for IFN
defective cells in this study, but they will be a useful tool to screen for IFN defective cells in a
range of different cell types in future studies. As a result of this study, there is a good
foundation to try to generate the type I IFN defective cells and study the involvement of
innate immune responses in rabbit cells in regard to RHDV/RCV infection, in any cell line
deemed promising
Orford-Dunne, Roxy (CES, Black Mountain)
94
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