1
An Analysis of Bovine immunodeficiency virus and Jembrana
disease virus Infections in Bos javanicus
Tegan Josephine McNab
BSc (Hons)
This thesis is presented for the degree of Doctor of Philosophy
of Murdoch University.
April 2010
i
Abstract
Two closely related bovine lentiviruses have been described, Jembrana disease virus
(JDV) and Bovine immunodeficiency virus (BIV), that produce very different clinical
manifestations in infected cattle. JDV causes an acute disease with a case fatality rate
of about 21% in Bos javanicus (Bali cattle) and is endemic in the cattle population of
parts of Indonesia. BIV produces a subclinical infection in Bos taurus and buffalo
and serological evidence has shown that this virus has a worldwide distribution,
possibly including Indonesia.
Attempts were made to confirm a previous report that BIV was present in the
B. javanicus population in Indonesia. BIV proviral DNA was not detected in any of
the animals although JDV proviral DNA was detected in 12 of 171 animals, only one
of which was seropositive.
To define the kinetics of BIV infection in B. javanicus and determine the optimal
time for sampling to detect BIV infection, 13 animals were experimentally infected
with the R29 strain of BIV. No clinical effects were detected but proviral DNA was
detected from 4-60 days post-infection (dpi) with peak titres 20 days dpi, and a
transient viraemia from 4 to 14 dpi. An antibody response to TM was detected 12 dpi
but an anti-capsid (CA) antibody response was detected in one animal only and not
until 34 dpi. The results indicated that detection of BIV in infected Bali cattle using
PCR would have a greater chance of success soon after infection and prior to the
onset of a CA antibody response.
To determine the effect of BIV infection on subsequent JDV infection in
B. javanicus, 15 cattle were infected with BIV-R29 and 9 of these were subsequently
infected 42 days later with JDV. The response to BIV was typical of that observed
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previously but BIV infection did not markedly modify the response to subsequent
infection with JDV. In response to JDV infection, all cattle previously infected with
BIV still developed an acute disease process typical of Jembrana disease. The results
suggested that despite the close genetic and antigenic relationship between BIV and
JDV, BIV infection does not confer protection against subsequent JDV infection.
The close antigenic relationship between BIV and JDV is a problem in the
development of specific serological tests and immunosurveillance of JDV infection.
To develop reagents capable of differentiating between antibody to BIV and JDV
infections, peptide mapping was used to define linear B cell epitopes on the matrix
(MA), CA and surface unit (SU) proteins of JDV. Short overlapping peptides that
spanned these regions were synthesised and used in an ELISA format to screen their
reactivity with a panel of bovine sera from animals experimentally infected with
JDVTab87, JDVPul01 or BIV-R29. Peptides representing potential immunoreactive
epitopes were identified that appeared to offer promise in the development of JDV-
specific serological tests and need to be tested further with a panel of sera taken from
naturally infected cattle.
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Declaration
I declare that this thesis is my own account of my research and contains as its main
content work which has not previously been submitted for a degree at any tertiary
education institution.
....................................
Tegan Josephine McNab.
iv
Contents
ABSTRACT........................................................................................................................................... I
DECLARATION................................................................................................................................III
ACKNOWLEDGEMENTS............................................................................................................... VI
ABBREVIATIONS ...........................................................................................................................VII
PUBLICATION AND INTERNATIONAL CONFERENCE PRESENTATIONS...................... IX
Publications arising from this thesis ............................................................................................ix Manuscripts submitted for publication .........................................................................................ix Oral presentations ........................................................................................................................ix Poster presentations .....................................................................................................................ix
CHAPTER 1: INTRODUCTION ....................................................................................................... 1
CHAPTER 2: LITERATURE REVIEW ........................................................................................... 4
Physiochemical characteristics of retroviruses ............................................................................ 4 Retrovirus taxonomy..................................................................................................................... 4 General structure of lentiviruses .................................................................................................. 5 The lentivirus genome................................................................................................................... 6 Lentivirus replication ................................................................................................................. 10 Common characteristics of lentiviruses...................................................................................... 14
PRIMATE LENTIVIRUSES................................................................................................................... 15 Human immunodeficiency virus.................................................................................................. 15 Simian immunodeficiency viruses............................................................................................... 17
NON-PRIMATE LENTIVIRUSES........................................................................................................... 19 Feline immunodeficiency virus ................................................................................................... 19 FIV in non-domestic feline species ............................................................................................. 21 Equine infectious anaemia virus................................................................................................. 22
SMALL RUMINANT LENTIVIRUSES .................................................................................................... 23 Visna maedi virus ....................................................................................................................... 24 Caprine arthritis encephalitis virus............................................................................................ 25
BOVINE LENTIVIRUSES..................................................................................................................... 25 Bovine immunodeficiency virus .................................................................................................. 27 Jembrana disease virus............................................................................................................... 29
IMMUNE RESPONSE TO INFECTION WITH A LENTIVIRUS.................................................................... 35 Humoral immune response to infection with bovine lentiviruses ............................................... 38
ASSAYS FOR THE DETECTION OF LENTIVIRUS INFECTIONS IN RUMINANTS ........................................ 40 Small ruminant lentiviruses ........................................................................................................ 40 Large ruminant lentiviruses........................................................................................................ 41 Difficulties with serological testing ............................................................................................ 42 Techniques to identify epitopes and distinguish between viral infections .................................. 43
LENTIVIRUS SUPERINFECTION.......................................................................................................... 45 Benefits of superinfection - superinfection resistance ................................................................ 45 Adverse consequences of superinfection..................................................................................... 47
CHAPTER 3: ATTEMPTS TO DETECT BOVINE IMMUNODEFICIENCY VIRUS INFECTION IN BALI CATTLE IN INDONESIA WITH A PCR-BASED ASSAY................... 49
SUMMARY ........................................................................................................................................ 49 INTRODUCTION................................................................................................................................. 50 MATERIALS AND METHODS.............................................................................................................. 51 RESULTS.......................................................................................................................................... 57 DISCUSSION..................................................................................................................................... 62
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CHAPTER 4: BOVINE IMMUNODEFICIENCY VIRUS PRODUCES A TRANSIENT VIRAEMIC PHASE SOON AFTER INFECTION IN BOS JAVANICUS.................................. 66
SUMMARY ........................................................................................................................................ 66 INTRODUCTION................................................................................................................................. 67 MATERIALS AND METHODS.............................................................................................................. 68 RESULTS.......................................................................................................................................... 73 DISCUSSION..................................................................................................................................... 80
CHAPTER 5: BOVINE IMMUNODEFICIENCY VIRUS INFECTION ALTERS THE DYNAMICS OF SUBSEQUENT JEMBRANA DISEASE VIRUS INFECTION......................... 83
SUMMARY ........................................................................................................................................ 83 INTRODUCTION................................................................................................................................. 84 MATERIALS AND METHODS.............................................................................................................. 85 RESULTS.......................................................................................................................................... 89 DISCUSSION................................................................................................................................... 100
CHAPTER 6: HUMORAL IMMUNE RESPONSES TO JEMBRANA DISEASE VIRUS DETECTED USING OVERLAPPING SYNTHETIC PEPTIDES SPANNING THE MA, CA AND SU REGIONS OF JDV .......................................................................................................... 105
SUMMARY ...................................................................................................................................... 105 INTRODUCTION............................................................................................................................... 106 MATERIALS AND METHODS............................................................................................................ 107 RESULTS........................................................................................................................................ 111 DISCUSSION................................................................................................................................... 119
CHAPTER 7: GENERAL DISCUSSION...................................................................................... 127
REFERENCES................................................................................................................................. 135
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Acknowledgements
First and foremost, I would like to thank my two supervisors, Dr. Moira Desport and
Professor Graham Wilcox, for their help throughout my PhD. I really appreciate all
the help they’ve given me. In particular, Moira for your input in designing my PhD,
for helping me in the lab and for all the wisdom you’ve passed on to me. To Graham,
for your help in editing my thesis and getting papers ready for submission. I hope
you catch many fish in your retirement!
Thankyou to Robert Dobson for your help with all things statistical, I am sure I still
owe you a few more cakes. Thanks must also be extended to other staff in the
Veterinary and Biomedical Sciences building for your help over the past few years.
Thankyou to the Australian Centre for International Agricultural Research for
providing the funding necessary to undertake this work and to the Australian
Government for providing my scholarship.
Thankyou to the members of the laboratory in Denpasar, particularly Nining and
Putu, for their help in running our animal trials and for the collection of samples.
Without this, it would not have been possible to complete my thesis.
A big thankyou to past and present members of our office: Will, Mark, Andrew,
Linda, Josh, Masa, Yudhi and honouree member Jill A. Thankyou so much for all
your help and good times shared around cake.
To all my friends, Laura G, Jacqui, Gael, Genevieve, Sarah, Nikki, Bry, Celia,
Laura H, Kendle, Michelle, and my entire family, thanks for all the good times. To
everyone at kickboxing, thankyou for helping to keep the stress levels low by letting
me punch and kick you on a regular basis.
Finally, a big thankyou to my parents, Marg and Ross, for being fantastic and so
supportive all the time.
To Bong for just being you.
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Abbreviations
AIDS Acquired immunodeficiency syndrome AGID Agar gel immunodiffusion APOBEC Apolipoprotein B mRNA-editing enzyme-catalytic polypeptide-like AUC Area under the curve BFL Bovine foetal lung BIV Bovine immunodeficiency virus BVDV Bovine viral diarrhoea virus CA Capsid CAEV Caprine arthritis encephalitis virus CE Cell equivalents CNS Central nervous system CCR5 C-C (beta) chemokine receptor 5 CD4 Cluster of differentiation 4 CD8 Cluster of differentiation 8 CTL Cytotoxic T-lymphocyte CXCR4 C-X-C (alpha) chemokine receptor 4 DNA Deoxyribonucleic acid dNTP Deoxynucleotide Triphosphate DMSO Dimethyl sulfoxide dpi Days post-infection ELISA Enzyme linked immunosorbent assay EIAV Equine infectious anaemia virus EDTA Ethylenediamine tetra-acetic acid FIV Feline immunodeficiency virus GAPDH Glyceraldehyde 3-phosphate dehydrogenase HRP Horse radish peroxidase HIV Human immunodeficiency virus HTLV-1 Human T-cell lymphotropic virus type 1 ID Immunodominant IgG Immunoglobulin G IR Immunoreactive IN Integrase JDV Jembrana disease virus LTR Long terminal repeat M-tropic Macrophage tropic VMV Visna maedi virus MHR Major homology region MA Matrix NC Nucleocapsid Nef Negative factor ORF Open reading frame PBMC Peripheral blood mononuclear cells PBS Phosphate-buffered saline PBS-T Phosphate-buffered saline-Tween 20 PCR Polymerase chain reaction pi Post-infection PR Protease
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qPCR Quantitative polymerase chain reaction qRT-PCR Quantitative reverse transcriptase polymerase chain reaction Rev Regulator of expression of virion proteins RT Reverse transcriptase RNA Ribonucleic acid RPMI Roswell Park Memorial Institute SIV Simian immunodeficiency virus SRLV Small ruminant lentivirus SU Surface unit T-tropic T-lymphocyte-tropic TCID50 Median tissue culture infective dose Tat Trans-activator of transcription protein TM Transmembrane glycoprotein U3 3’ Untranslated region U5 5’ Untranslated region Vif Viral infectivity protein VL Viral load Vpr Viral protein R Vpu Viral protein U Vpx Viral protein X WIB Western immunoblotting YT Yeast tryptone broth
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Publication and International Conference Presentations
Publications arising from this thesis
McNab, T., Desport, M., Tenaya, W. M., Hartaningsih, N., and Wilcox, G. E. (2010). Bovine immunodeficiency virus produces a transient viraemic phase soon after infection in Bos javanicus. Vet. Microbiol. 141, 216-223.
Desport, M., Ditcham, W. G., Lewis, J. R., McNab, T. J., Stewart, M. E.,
Hartaningsih, N., and Wilcox, G. E. (2009). Analysis of Jembrana disease virus replication dynamics in vivo reveals strain variation and atypical responses to infection. Virology. 386(2), 310-6.
Lewis, J., McNab, T., Tenaya, M., Hartaningsih, N., Wilcox, G., and Desport, M.
(2009). Comparison of immunoassay and real-time PCR methods for the detection of Jembrana disease virus infection in Bali cattle. J Virol Methods. 159(1), 81-6.
Manuscripts submitted for publication
McNab, T., Desprt, M., Dobson, R., Tenaya, I.W.M., Hartaningsih, N., Wilcox, G.E. (2010) Prior Bovine immunodeficiency virus infection does not inhibit subsequent superinfection by the acutely pathogenic Jembrana disease virus. Virology. Oral presentations
“Bovine immunodeficiency virus infection fails to provide protection against subsequent Jembrana disease virus infection” Presented at the European Society for Veterinary Virology Conference in Budapest, Hungary 2009. Poster presentations
“Towards a Jembrana disease virus specific diagnostic immunoassay- peptide mapping of gag and env proteins of bovine lentiviruses” Presented at the European Society for Veterinary Virology Conference in Budapest, Hungary 2009.
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Chapter 1: Introduction
Two bovine lentiviruses have been described, Jembrana disease virus (JDV) and
Bovine immunodeficiency virus (BIV). The 2 viruses, although genetically and
antigenically related, have been reported to have very different pathogenic effects in
cattle. JDV causes a severe, acute disease in Bali cattle (Bos javanicus) and a mild
disease or subclinical infection in other breeds of cattle, including B. taurus
(Soeharsono et al., 1990). The disease in Bali cattle is acute and associated with a
marked febrile response, very high titres of infectious virus in the blood and a case
fatality rate of about 21% (Soesanto et al., 1990). BIV infection is generally not
associated with significant clinical changes in B. taurus breeds of cattle, although
they have been reported: one study found an association between BIV and decreased
milk yield in dairy cattle (McNab et al., 1994) and another associated BIV infection
with marked weight loss and concurrent infections, suggesting immunosuppression
(Snider et al., 2003b). JDV appears to have a limited geographic distribution and is
restricted to Indonesia where Bali cattle are found (Hartaningsih et al., 1993), while
serological surveys have provided evidence for a worldwide distribution of BIV in
both cattle and buffalo (Bubalus bubalis) (Gonzalez et al., 2008; McNab et al., 1994;
Meas et al., 1998; Meas et al., 2000a; Meas et al., 2000b; Suarez et al., 1993). There
is also serological evidence of infection with a related non-pathogenic BIV-like virus
in Bali cattle in Indonesia on the island of Sulawesi and also in Bali (Barboni et al.,
2001; Desport et al., 2005). Although infection with a BIV-like virus has been
suspected in Bali cattle in Indonesia, this has not been confirmed. There is no
evidence of the nature of BIV infection in this cattle species and what effect BIV
infection might have on subsequent JDV infection.
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This thesis describes the development of a serological diagnostic assay to
differentiate JDV and BIV infection, the effects of BIV in Bali cattle and the
interaction between BIV and JDV infections in Bali cattle. As a background to these
investigations, a review of the literature relating to JDV and the other lentiviruses has
been undertaken and is reported in Chapter 2. The review includes the key features of
the various lentiviruses, comparing their genome arrangement, replication and host
immune responses to infection. It also includes the various techniques that have been
described for diagnosis of lentivirus infections, methods of differentiating between
closely-related lentivirus infections and the effects of infection with multiple strains
of closely related lentiviruses.
Chapter 3 describes an attempt to detect JDV and BIV in Bali cattle on the island of
Bali. Two quantitative PCR (qPCR) assays were developed to detect JDV and BIV
proviral DNA within peripheral blood mononuclear cells (PBMC) of naturally
infected animals.
Despite serological evidence for the presence of BIV in the Bali cattle population on
the island of Bali, the investigations reported in Chapter 3 failed to detect evidence of
BIV in the cattle that were sampled. Investigations were therefore undertaken to
determine the susceptibility of Bali cattle to BIV infection in an effort to better
understand the nature of the infection in this species, and these investigations are
reported in Chapter 4. Nineteen cattle were experimentally infected with the R29
strain of BIV and monitored for up to 65 days after infection. The presence of virus
in plasma and other tissues, the presence of proviral DNA in the PBMC and other
tissues, and the immune response to virus antigens was examined and is reported.
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The possibility that BIV infection of Bali cattle might modify the effect of
subsequent JDV infection was also investigated and these results are presented in
Chapter 5. These studies were undertaken to determine what might happen on the
island of Sulawesi, where BIV infection is suspected to occur in Bali cattle, if JDV
were to spread through the Bali cattle population of that island. If prior BIV infection
were to inhibit subsequent JDV infection, it was hypothesised that this might form
the basis of a method of vaccination for the control of Jembrana disease.
Due to the presence of cross-reactive epitopes between JDV and BIV proteins,
current serological assays are not capable of discriminating between antibody to the
2 bovine lentiviruses. During the studies undertaken and reported in Chapters 3, 4
and 5, the difficulty of distinguishing antibody to BIV and JDV made it difficult to
monitor the serological response to the individual virus infections. An attempt was
therefore made to develop a peptide antigen capable of differentiating between
antibody to the 2 viruses. Overlapping virus peptides were synthesized and used in
an enzyme linked immunosorbent assay (ELISA) format with serum samples taken
from JDV and BIV infected cattle to determine their reactivity to the peptides. The
results are reported in Chapter 6.
A general discussion of the results obtained and reported in the thesis is presented in
Chapter 7.
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Chapter 2: Literature Review
This review covers the general biological features of the retroviruses and
lentiviruses, in particular the lentiviruses that cause disease in animals. Also
reviewed are the various immune responses to animal lentivirus infection (including
cross-reactive epitopes) and the phenomenon of lentivirus superinfection resistance.
The nomenclature used in this thesis for viral genes and proteins is that suggested by
(Fauquet, 2005) where gene names are in lower case and italicised, eg. env, and
where the abbreviations for the encoded proteins have the initial letter in uppercase
and are not italicised, eg. Env and TM.
Retroviruses
Physiochemical characteristics of retroviruses
The key feature of the family Retroviridae is their mode of replication, involving
reverse transcription of the virion RNA into linear double-stranded DNA (Baltimore,
1970; Temin et al., 1970), and the integration of this double-stranded proviral DNA
into the genome of the cell. This reverses the normal flow of genetic information
from DNA to RNA, hence the name retrovirus (Coffin, 1997).
Retrovirus taxonomy
Many viruses have been classified within the family Retroviridae with a significant
proportion of them associated with disease (Coffin, 1997). On the basis of
evolutionary relatedness, retroviruses are separated into 7 genera: Lentivirus,
Spumavirus, Alpharetrovirus, Betaretrovirus, Gammaretrovirus, Deltaretrovirus and
Epsilonretrovirus (Buchschacher, 2001). The lentiviruses and spumaviruses are
distinct from the other 5 genera in that they do not have oncogenic potential. Some of
the well known oncogenic retroviruses include Rous sarcoma virus, Human T-
5
lymphotropic virus (HTLV-1) and Mouse mammary tumour virus. Less well
understood and researched are the spumaviruses which cause no known disease, of
which Human spumavirus is the type species (Coffin, 1997; Goff, 2001). The
lentiviruses have been studied extensively since the discovery of Human
immunodeficiency virus (HIV).
Lentiviruses
Most lentivirus infections are characterised by a long asymptomatic period before the
onset of chronic clinical disease with a slow but inevitable progression to death
(Campbell et al., 1998). Examples of lentiviruses inducing this type of chronic
infection include Visna maedi virus (VMV), HIV and Caprine arthritis encephalitis
virus (CAEV). Some, however, induce a rapidly progressive acute disease, including
JDV, Equine infectious anaemia virus (EIAV) and the Simian immunodeficiency
virus (SIV) SIVsmm-PBj. This section will review features of lentiviruses that are
common to the majority of members of the genus, including genome structure and
organisation, their mode of replication and common clinical features of infection.
General structure of lentiviruses
Lentiviruses are roughly spherical in shape with an average virion diameter of
approximately 100 nm including the surrounding bilayered lipoprotein envelope
(Fauquet, 2005). They are sensitive to heat, detergent and formaldehyde (Goff,
2001). The envelope contains 2 types of surface projections, the surface unit (SU)
and transmembrane (TM) glycoproteins (Fauquet, 2005; Wagner, 1999). The internal
virion is composed of the distinctively cone-shaped capsid (CA) which surrounds the
nucleocapsid (NC) and contains protease (PR), integrase (IN) and reverse
transcriptase (RT) enzymes (Figure 2.1). The RNA genome is located within the
nucleocapsid (Goff, 2001).
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The lentivirus genome
Lentiviruses have 2 identical linear, positive-sense, single-stranded RNA genomes
which range in size from 7-10 kb (Goff, 2001; Peterlin, 1995). Reverse transcription
takes place from one strand at a time. There are 3 major open reading frames (ORF)
in each strand that transcribe and translate 3 polyproteins which are then cleaved by
proteases into approximately 8 proteins. The 3 polyproteins are Gag, Pol and Env
(Figure 2.2). The gag ORF produces the structural proteins MA, CA and NC. The pol
ORF encodes the intravirion enzymes: RT responsible for copying the single-
stranded RNA genome into the double-stranded DNA, IN which is required to
incorporate the double stranded DNA into the host genome forming the provirus, and
PR, required to cleave the encoded polyproteins into smaller proteins (Miller et al.,
2000; Tobin et al., 1994). The env gene produces the 2 envelope proteins, SU and
TM, which play a vital role in receptor recognition and entry of the virus into the cell
(Fauquet, 2005; Wagner, 1999).
Figure 2.1. (A) Schematic representation of a mature HIV-1 virion, showing the location of the
major viral proteins, the lipoprotein envelope and genomic RNA. (B) Tomogram of a mature HIV-1
particle derived by electron cryotomography. Images from Ganser-Pornillos et al. (2008).
7
In addition to the obligatory gag, pol and env ORF common to every retrovirus,
lentiviruses also have a number of accessory genes (Table 2.1), including rev, tat, vif,
vpr, vpu and nef (Fauquet, 2005), which modulate the replication of the virus and
probably contribute to clinical latency and pathogenic mechanisms (Clements et al.,
1996). Rev plays an essential role in the replication cycle of all lentiviruses as it
facilitates the export from the nucleus of unspliced RNAs whose translated products
are later utilised in virus replication (Malim et al., 1989). Like Rev, Tat is produced
early in the replication cycle and plays a role in the expression of viral transcripts
from a promoter within the long terminal repeat (LTR) (Miller et al., 2000). HIV Tat
has also been shown to modulate the expression of cellular genes and has numerous
other roles in viral replication and pathogenesis (Chen et al., 2000). FIV is the only
lentivirus without the tat ORF but ORF2 encoded by orf2 is predicted to have Tat-
like activity and to act via cellular transcription factors during the expression of viral
transcripts from the promoter within the viral LTR (Chatterji et al., 2002; Miller et
al., 2000; Miyazawa et al., 1994) but requires additional elements within the LTR,
unlike other lentivirus Tat proteins (Chatterji et al., 2002). It has been proposed that
VMV and CAEV lack a tat ORF and that the ORF that is designated tat in these
viruses (named tat because of its similar position in the genome to the tat ORF in the
primate lentiviruses) instead encodes a Vpr-like accessory protein (Villet et al.,
2003).
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Figure 2.2. Genome organisation of lentiviruses. The location of the structural and accessory genes
are indicated, orientated 5’ to 3’. Each virus has 3 major open reading frames, gag, pol and env, which
transcribe and translate 3 polyproteins. Image from Craigo et al. (2010).
9
The vif ORF is transcribed and translated to Vif (viral infectivity factor), thought to
aid in the infectivity and spread of virus, although its mechanism of action is still
unclear (Clements et al., 1996; Miller et al., 2000). Vif has recently been implicated
in protecting virions against the actions of Apolipoprotein B mRNA-editing enzyme-
catalytic polypeptide-like (APOBEC) proteins. APOBEC3G and APOBEC3F are
cytidine deaminases which are packaged into HIV-1 virions and result in the
production of non-infectious virions due to the hypermutation of HIV proviral DNA.
Vif protects the virus against lethal incorporation of the APOBEC proteins by
marking them for ubiquitin-dependent degradation (Goila-Gaur et al., 2008, Romani
et al., 2009).
Vpr encodes Vpr (viral protein R) which, in HIV-1, mediates the nuclear import of
viral RT complexes in non-dividing cells and alters the cell cycle and proliferation
status of the infected host cell. In HIV-2 and SIVsm, Vpr inhibits cell cycle
progression while Vpx (encoded by vpx) is responsible for the nuclear import of the
viral RT complex (Fletcher et al., 1996; Stivahtis et al., 1997). The vpu ORF encodes
Vpu (viral protein U) that enhances the efficiency of virion production and induces
rapid degradation of CD4 (Maldarelli et al., 1993). The nef ORF encodes Nef
(negative factor) whose function is to decrease the expression of CD4 on T-cells; it is
thought to have this effect by increasing the rate of endocytosis of CD4 on the cell
surface which would ultimately prevent re-infection of cells that already harbour the
virus (Benson et al., 1993; Goff, 2001; Peterlin, 1995). The accessory ORF s2 is
unique to EIAV and its role in virus replication is unclear but it is thought to play a
part in replication and virulence of EIAV (Li et al., 1992; Nilsen et al., 1996).
10
Table 2.1. Accessory genes of each lentivirus, presence indicated by √ (Chakrabarti
et al., 1987; Chatterji et al., 2002; Clements et al., 1996; Dewhurst et al., 1990;
Freed, 2001; Li et al., 1992; Miller et al., 2000; Nilsen et al., 1996; Stivahtis et al.,
1997; Tobin et al., 1994). SIVsm has a gene encoding the Vpx protein, other SIVs
lack this gene (Stivahtis et al., 1997). The presence of tat within the VMV and
CAEV genomes has been debated (Villet et al., 2003).
HIV-1 HIV-2 SIV FIV VMV CAEV EIAV BIV JDV
vif √ √ √ √ √ √ √ √
vpx √ √
vpr √ √ √
tat √ √ √ √ √ √ √ √
rev √ √ √ √ √ √ √ √ √
vpu √ √
nef √ √ √
tmx √ √ √
Other
ORF 2/A S2
vpw,
vpy
Lentivirus replication
Infection of a cell by a lentivirus commences at the surface of the host cell (Figure
2.3). The SU interacts with specific receptors on the target cell. For example, HIV-1
SU interacts with the CD4 receptor located on the surface of T-lymphocytes and
monocytes/macrophages. The interaction brings about a conformational change in
the TM glycoprotein and ultimately leads to the fusion of the viral envelope with the
11
membrane of the target cell (Clapham et al., 2002; Clements et al., 1996). The virus
then moves into the cytoplasm where it is partially uncoated and virion enzymes
encoded by pol are released. The virion enzymes RT and RNase H copy the viral
RNA within the partially uncoated virion, generating a double-stranded DNA copy
of the viral RNA genome, referred to as the provirus. The provirus forms a complex
with a number of viral proteins including IN, MA, NC, RT, and possibly others, to
form the pre-integration complex, which translocates to the nucleus. Integrase then
catalyses the insertion of the linear, double-stranded viral DNA into the host cell
chromosome (Freed, 2001). At this point, the proviral DNA may remain integrated
and the cell can remain latently infected, or a productive infection may result. It
remains unclear as to what causes a cell to become productively infected, although it
is thought that the presence of specific transcription factors present in mature cells
may stimulate transcription of viral genes. For example, it has been proposed that the
binding of the transcription factor NF-κB, which is found in activated T-
lymphocytes, to the HIV-1 LTR, is important for transcriptional activation in vitro of
HIV-1 in T-cell lines (Clements et al., 1996; Wagner, 1999). Others have proposed
that defective HIV-1 particles preferentially activate CD4+ T-cells which render
them permissive for HIV replication and help to drive HIV pathogenesis (Finzi et al.,
2006).
Once transcription has been activated, expression of viral mRNA begins when RNA
polymerase II binds to the U3 region of the 5' LTR and transcription then proceeds
towards the 3' end of the provirus and into the host DNA. The RNA is cleaved and
polyadenylated at the R-U5 border of the 3' LTR (Figure 2.4), which yields a
complete unspliced viral genomic RNA suitable for incorporation into the virion
particle. A portion of the RNA produced at this level is then spliced by the cellular
12
splicing machinery to give rise to one or more subgenomic RNAs. Both the
unspliced and spliced RNAs are exported from the nucleus for translation (Goff,
2001).
Once in the cytoplasm, the RNAs transcribed from gag, pol and env are translated by
ribosomes into polyproteins (large precursor protein molecules). In HIV, the gag
ORF encodes a polyprotein precursor of 55 kDa, designated Pr55Gag, that is cleaved
by viral PR to produce MA (p17), CA (p24), NC (p7) and p6. The pol-encoded
enzymes are initially synthesised as part of a large polyprotein precursor, Pr160GagPol
that is cleaved by the virus encoded protease into PR, RT and IN. The Env precursor,
gp160, is cleaved by cellular protease into SU (gp120) and TM (gp41) (Egberink et
al., 1992; Freed, 2001).
Gag and Gag-Pol precursors assemble beneath the plasma membrane and incorporate
viral genomic RNA during the process of budding, while SU and TM glycoproteins
are also inserted into the viral envelope at this stage. After the virion has been
released, it matures when PR cleaves Gag precursors into their functional subunits.
The mature virion is then able to infect other cells (Tobin et al., 1994).
13
Figure 2.4. Schematic illustration of the changes to lentivirus long terminal repeats during the change
from viral RNA to proviral DNA to viral mRNA (transcript). Illustrations are orientated 5’ to 3’. The
positions of the R (repeat), U3 and U5 regions are shown. Viral mRNA is expressed when RNA
polymerase II binds to the U3 region of the 5’ LTR. Image from Coffin et al. (1997).
Figure 2.3. Schematic illustration of the lentivirus replication cycle depicting the major events in
the replication cycle. Image from Ganser-Pornillos et al. (2008).
14
Common characteristics of lentiviruses
Members of the genus Lentivirus share a number of features. They all replicate in
non-dividing terminally differentiated cells, have the ability to integrate their
genomes into the chromosomal DNA of non-dividing infected cells and they are
highly species-specific in terms of their ability to cause disease. Common clinical
features of infection shared by the majority of lentiviruses include long
asymptomatic incubation periods before the onset of a usually chronic disease,
persistence of virus infection in the face of vigorous immune responses including
neutralising antibody and cytotoxic T-lymphocytes (CTL), multi-organ disease and
replication in cells of the immune system and brain (Clements et al., 1996; Mealey et
al., 2004; Trautwein, 1992). It is often the immune response of the host against the
infected cells that results in the wide range of disease symptoms observed (Gonda,
1992).
The ability of the viruses to persist in the infected host is, at least in part, associated
with the capacity of lentiviruses to exhibit a wide array of genetic and antigenic
variations, particularly in env. Variations are produced in response to biological and
immunological selective pressures as the virus (usually) successfully avoids
clearance by defence mechanisms (Leroux et al., 2001; Mealey et al., 2004).
15
Primate lentiviruses
HIV and SIV cause disease in primates and it is the study of these lentiviruses that
has resulted in most of our knowledge of the molecular biology and mechanisms of
disease associated with lentivirus infections. This section will review HIV and SIV
with a particular focus on their genome organisation, cell tropism, clinical features of
diseases they induce, and species specificity.
Human immunodeficiency virus
Arguably the most studied virus in the world today, HIV was first identified in 1983
as the agent responsible for an acquired immunodeficiency syndrome (AIDS) which
led to opportunistic infections and eventually death (Barre-Sinoussi et al., 1983;
Gallo et al., 1984). It has assumed very significant pandemic proportions and it is
estimated that about 33 million people are infected with the virus (UNAIDS, 2008).
Based on epidemiological and genetic studies, HIV isolates form two clusters, HIV-1
and HIV-2, which are distinguished by variations primarily in env. They also show
differences in transmission rate and pathogenicity (Levy, 2009). HIV-1 is the type
species of the genus Lentivirus and although the published literature describing
aspects of the molecular biology of HIV-1 and the associated infection is very large,
a brief overview only will be given in this review.
Many lentiviruses evade the host immune system via their genetic variability (Lopez
et al., 2006). Phylogenetic analyses of HIV-1 isolates from around the world indicate
that more than 10 major groups of distinct genetic subtypes or clades of HIV-1 can
be distinguished. Clades are differentiated by genomic differences in env of 15% or
greater (Barker, 1995). The distribution of the clades tends to have a geographic
basis: Subtype C (clade C) infections are most commonly found in south Africa,
India, Ethiopia and east Africa. Subtype A clades are found in eastern Europe,
16
central Asia, west, east and central Africa while subtype B is present in the
Americas, western Europe and east Asia. Subtype D is present in north Africa, the
middle east, east and central Africa (Hemelaar et al., 2006).
In HIV the major determinants of cell tropism are the cell surface receptors used to
gain entry into the cell. HIV-1 uses the CD4 receptor and a co-receptor to enter the
host cell (Sattentau et al., 1988). The main cells that have the 2 receptors needed for
HIV-1 to gain entry are the CD4+ T-helper subset of lymphocytes, the CD4+ cells of
macrophage lineage and some dendritic cells (Clapham et al., 2002; Freed, 2001).
Major co-receptors for HIV-1 include the 2 chemokine receptors CCR5 (expressed
on macrophages) and CXCR4 (expressed on T-cells) but other co-receptors are used
(Deng et al., 1996; Doranz et al., 1996; Feng et al., 1996). CCR5-utilising HIV-1
variants dominate the early phases of HIV-1 infection while CXCR4-utilising HIV-1
variants dominate the latter phases of HIV-1 infection, and the switch from CCR5-
utilising to CXCR4-utilising is associated with accelerated disease progression (Ito et
al., 2003).
HIV-1 will establish infection in both humans and chimpanzees but will only cause
disease in humans, with a few rare exceptions (Freed, 2001, Keele et al., 2009).
There are 3 phases to the course of HIV infections. In the initial phase, there is a
period of rapid virus replication associated with influenza-like symptoms,
commencing about 2 weeks post infection (pi) and lasting for 2-3 weeks.
Subsequently, there is a variable asymptomatic period of weeks to years during
which lymphadenopathy can develop. In the third phase, the destruction of the T-cell
population causes the onset of AIDS, seen in about 30% of infected people within 5-
7 years and later in others (Campbell et al., 1998). The loss of T-cells results in the
body being unable to overcome opportunistic infections, and the immune system
17
finally fails, which without anti-viral therapy will ultimately lead to death (Wagner,
1999). The loss of CD4+ T-cells and the changes in HIV RNA levels are shown in
Figure 2.5. Key clinical manifestations of HIV-1 infection are immunodeficiency,
lymphadenopathy, opportunistic infections, encephalopathy, emaciation, Kaposi's
sarcoma and other cancers (Gonda, 1992).
Simian immunodeficiency viruses
Simian lentiviruses have been identified in several species of non-human primates by
epidemiological studies using serological assays and virus isolation. Each SIV is
named with a subscript that denotes the species from which the virus was first
isolated, for example those from sooty mangabeys as SIVsmm, from macaques as
SIVmac and from chimpanzees as SIVcpz (VandeWoude et al., 2006). In their natural
host to which they have adapted they do not normally cause disease but when they
infect species in which they do not normally occur, disease frequently results. These
Figure 2.5. The progression from infection with HIV to death in a victim. The patient was infected
at or near week 0. The diagram illustrates the levels of HIV RNA, the gradual decline in CD4+ T-
cells and the onset of clinical signs. Image from Pantaleo et al. (1995).
18
viruses share many features in common with HIV, particularly nucleotide homology,
genome organisation, size (approximately 9 kb in length), receptor usage as well as
the clinical manifestations of disease they induce in non-natural hosts (Clements et
al., 1996; Sattentau et al., 1988).
All SIV have a tropism for CD4+ macrophages or T-cells. Macrophage-tropic (M-
tropic) SIV can efficiently replicate in macrophages, primary CD4+ T-cells and a
variety of T-cell lines whereas T-cell-tropic (T-tropic) SIV replicate in primary
CD4+ T-cells and in T-cell lines but not macrophages (Edinger et al., 1999). Both M-
and T-tropic SIV strains use CCR5 to gain entry into CD4+ cells (Edinger et al.,
1997). The locations of viral replication are thought to be responsible for the varying
clinical manifestations of SIV infection: replication in cells of the
monocyte/macrophage lineage results in disease manifestations in the central nervous
system and lungs; replication in lymphocytes results in a loss of CD4+ lymphocytes
which in turn results in immunodeficiency and increased susceptibility to
opportunistic infections (Sharma et al., 1992).
While the majority of SIV cause disease after a relatively long period of infection,
there are strains which cause disease after only a short incubation period, such as
SIVsmmPBj14. This strain causes an cute disease and death within 6 to 10 days after
intravenous inoculation into pig-tailed macaques and rapidly replicates to high titres
(Fultz et al., 1994, Tao et al., 1995). The lethal nature of this phenotype is possibly
associated with insertions within the V1 region of env and within the NF-κB
enhancer element in the LTR, which enhances transcription and replication kinetics
(Tao et al., 1995).
19
SIV that originate in African primate hosts are thought to be relatively ancient
viruses that result in non-pathogenic infections in their native hosts. A similar
situation is seen with lentivirus infections of non-domestic cats (Hahn et al., 2000;
Terwee et al., 2005).
Non-primate lentiviruses
Feline immunodeficiency virus
FIV is associated with acquired immunodeficiency in cats and was first isolated in
California (Pedersen et al., 1987). Since its initial isolation, similarities in the disease
syndrome induced in cats to that of HIV in humans have created considerable interest
in the virus as a potential animal model for HIV infection (Dandekar et al., 1992;
Dua et al., 1994; Gardner et al., 1989; Olmsted et al., 1989). The virus infects several
species of Felidae, including the domestic cat Felis cattus, throughout the world but
particularly in Europe, East Asia, Australasia and North America (Brown et al.,
1994; Duarte et al., 2006; Little et al., 2009; Olmsted et al., 1992). Circumstantial
evidence suggests that the most efficient mode of virus transmission is horizontally
by biting (Yamamoto et al., 1989). Other studies have shown that it can be
transmitted through semen and vertically during pregnancy (Jordan et al., 1998;
Wasmoen et al., 1992).
FIV has a broader target cell range than HIV and replicates in both CD4+ and CD8+
lymphocytes, macrophages and immunoglobulin G-positive lymphocytes (Beebe et
al., 1994; Brown et al., 1991; English et al., 1993). The primary receptor of FIV is
CD134, a T-cell-activation antigen and co-stimulatory molecule (Shimojima et al.,
2004). Known co-receptors include CXCR4 (Willett et al., 1997). The progression
from infection to immunosuppression to death in domestic cats is well characterised
and mirrors HIV-1 infection in humans. There is a transient acute stage of the illness
20
a few weeks pi that is characterised by fever and lymphadenopathy. The next phase
is asymptomatic, lasting up to 5 years. The third and final phase of infection is
characterised by a gradual decline in CD4+ T-lymphocytes leading to
immunodeficiency. This stage, like AIDS in HIV-infected humans, is characterised
by the occurrence of opportunistic infections, neurological disorders and tumours of
various aetiologies, resulting in death usually within a few months (Dean et al., 1996;
Egberink et al., 1992; Ryan et al., 2003; Sauter et al., 2001).
Analogous to SIVsmmPBj14 inoculation into pig-tailed macaques, FIV-CPGammar also
causes an acute disease with a high case fatality rate. Mortalities range from 50 to
100% in kittens ≤12 weeks of age after a short incubation period following
intravenous inoculation However, in contrast to SIVsmmPBj14, acute phase virulence
was induced by acute-phase virus passage (Diehl et al., 1995) and the severe and
rapidly progressive disease could not be induced in young adult cats (Pedersen et al.,
2001).
21
FIV in non-domestic feline species
Serological surveys of a number of non-domestic feline species have revealed at least
17 species with FIV-like virus infection (Brown et al., 1994; Olmsted et al., 1989;
VandeWoude et al., 2006; VandeWoude et al., 2002). Isolates from pumas (also
referred to as cougar, mountain lion and panther; Puma concolor), lions (Panthera
leo), Pallas cat (Felis manul) and bobcats (Lynx rufus) have been genetically
characterised (Poss et al., 2006; Poss et al., 2008), and these isolates are distinct from
each other and are related to, but distinct from FIV of the domestic cat (Olmsted et
al., 1992). The standard nomenclature for designation of strains originating from
different species is to append genus and species identifiers for the feline species as a
subscript to FIV. For example FIV isolated from a lion is referred to as FIVple and
domestic cat as FIVfca (Brown et al., 1994; VandeWoude et al., 2006).
The available evidence indicates these viruses have been endemic within cat families
for a long period (Biek et al., 2006). Although the clinical effects of FIV from non-
domestic cat populations have not been well studied, it appears that infections due to
these viruses do not cause widespread disease (Biek, 2006; Brown et al., 1994;
VandeWoude et al., 2002; VandeWoude et al., 2003). The asymptomatic nature of
infection is most probably because of a long evolutionary association between virus
and host (Biek, 2006). Signs of neurologic disease have been reported in captive
lions (Brennan et al., 2006) but this may be attributed to the animals outliving their
“normal” life span. FIVpco from pumas is able to establish a productive infection in
domestic cats but does not cause T-cell dysregulation (unlike FIVfca) or clinical signs
and is able to be cleared from PBMC rapidly (Terwee et al., 2005; VandeWoude et
al., 2003). These cats also generate humoral and cell-mediated immune responses
22
reactive against both FIV and non-domestic cat isolates of FIV (VandeWoude et al.,
2003).
Equine infectious anaemia virus
EIAV, the cause of a chronic relapsing or intermittent anaemia in horses, was the
first lentivirus to be identified and the first non-plant virus to be discovered (Leroux
et al., 2004; Ligné, 1843; Vallée, 1904). The virus causes a persistent infection in
horses and closely related equids, such as donkeys and mules (O'Rourke et al., 1988;
Spyrou et al., 2003). It is transmitted mechanically mainly by blood-feeding biting
arthropods such as tabanids, or iatrogenically on contaminated needles, but contact
infection can also occur (Hawkins et al., 1976; Kemen et al., 1978; Li et al., 2003;
Williams et al., 1981). Because of its transmission by arthropods, the infection is
most common in geographic areas with long vector seasons (Issel et al., 1988).
EIAV is an exclusively tissue macrophage-tropic lentivirus which utilises the equine
lentivirus receptor-1 to gain entry into macrophages (Sellon et al., 1992; Zhang et al.,
2005). The virus causes a relapsing disease characterised by periods of depression,
fever, diarrhoea, anaemia, thrombocytopenia and haemorrhaging due to severe
depletion of platelets, which are associated with a high level viraemia (Hammond et
al., 1997; Harrold et al., 2000; O'Rourke et al., 1988; Oaks et al., 1998). The disease
cycles begin approximately 1-3 weeks pi, last 3-5 days and occur at irregular
intervals of weeks to months. The periodic disease cycles occur for 8-12 months pi,
after which infected horses normally progress to a subclinical infection lasting for the
life of the infected animal; these persistently infected animals serve as a source of
infection for other animals (Hammond et al., 1997; Issel et al., 1982; Montelaro et
al., 1984; Oaks et al., 1998; Payne et al., 1998; Salinovich et al., 1986). Between the
clinical episodes of disease, viral loads are greatly reduced (of the order of 4- to 733-
23
fold) and viral transcription within macrophages is restricted (Oaks et al., 1998). In
response to immune pressure, rapid genomic variations occur during replication
which results in altered glycoprotein structures and antigenic changes. Variants have
been detected within 28 days and this variation is responsible for the cyclical nature
of the disease, new antigenic variants having a replication advantage as they are not
susceptible to pre-existing immunity (Ball et al., 1992; Montelaro et al., 1984; Payne
et al., 1984; Salinovich et al., 1986).
Small ruminant lentiviruses
It was initially thought that VMV and CAEV were specific for sheep and goats,
respectively, but recent evidence has shown that VMV and CAEV are capable of
infecting both sheep and goats as well as some small ruminant species living in the
wild (Guiguen et al., 2000; Leroux et al., 1997; Narayan et al., 1980; Pisoni et al.,
2005; Ravazzolo et al., 2006; Shah et al., 2004). As a result, VMV and CAEV are
now collectively referred to as small ruminant lentiviruses (SRLV). The 2 viruses
share many similarities at both the genomic and antigenic level, particularly in gag
and pol (Brinkhof et al., 2007; Jolly et al., 1989; Pasick, 1998; Pyper et al., 1986;
Zanoni, 1998), and there is antigenic cross-reactivity between their CA proteins
(Clements et al., 1980). Some SRLV, however, show significant differences in their
LTR, env and regulatory genes (Jolly et al., 1989; Pyper et al., 1986; Zanoni, 1998).
They also differ phenotypically, with the prototypic Icelandic VMV strains inducing
syncytia and lysis of infected cell cultures whereas the prototypic CAEV-Cork strain
induces syncytia with a persistent but non-lytic infection of cell cultures (Narayan et
al., 1980; Pisoni et al., 2007; Querat et al., 1984).
24
Visna maedi virus
VMV is a natural pathogen of sheep (Zhang et al., 2000; Zink et al., 1987) and has
been isolated from the majority of sheep-rearing areas of the world, excluding
Australia and New Zealand (Dawson, 1988). VMV was initially reported as a cause
of progressive pneumonia of sheep in South Africa in 1915, then in Montana in the
1920's, then in sheep in Iceland in the 1950’s (Dawson, 1988; Jolly et al., 1989;
Sigurdsson et al., 1952; Sigurdsson et al., 1957). The virus name is derived from the
Icelandic language where “maedi” can be translated as “dyspnoea” (difficult
breathing due to pneumonia) and “visna” as “fading away” (due to a demyelinating
leukoencephalomyelitis), representing the 2 forms of the disease (Pepin et al., 1998).
The main route in which the virus is transmitted is thought to be via ingestion of
infected colostrum and/or milk or via the respiratory tract, involving direct inhalation
of infected respiratory secretions, including the inhalation of infected alveolar
macrophages (McNeilly et al., 2008; Pepin et al., 1998; Preziuso et al., 2004).
Monocytes, lung tissue macrophages and spleen tissue macrophages (but not T-
lymphocytes) are the major cell targets in vivo for VMV replication and viral
expression is greatly increased when the monocytes mature into macrophages
(Gendelman et al., 1986). In PBMC populations, dendritic cells are the most
permissive for viral replication (Gorrell et al., 1992; Zhang et al., 2000).
Clinical disease can take months to years to develop (Nilsen et al., 1996) and consists
of a multi-system disease characterised by a chronic infiltration and proliferation of
mononuclear cells in the lungs (chronic interstitial pneumonia, maedi) and the central
nervous system (affected by a chronic and progressive, paralytic disease
characterised by inflammatory and demyelinating lesions in the CNS leading to
wasting and paralysis, visna) (Narayan et al., 1980). Organs less commonly affected
25
include the joints (arthritis) and mammary glands (Bird et al., 1993; Deng et al.,
1986; Gorrell et al., 1992; Zink et al., 1987).
Caprine arthritis encephalitis virus
CAEV causes an economically significant disease in infected goats, particularly in
Europe, the Americas, Asia and Australia (Campbell et al., 1998; Herrmann et al.,
2003b; Mselli-Lakhal et al., 2007; Zanoni, 1998). The main route of virus
transmission is via colostrum but horizontal spread is also achieved via infected
secretions if there is direct contact between infected and susceptible animals
(Ravazzolo et al., 2006) or from doe to foetus either prior to or during the birth
process (East, 1993).
The virus, like VMV, infects cells of the monocyte/macrophage lineage and dendritic
cells and virus expression is activated during maturation of monocytes to
macrophages (Narayan et al., 1983). It does not infect lymphocytes and so does not
directly result in immunosuppression, unlike SIV and HIV. Infection results in a
chronic inflammatory disease affecting the joints, central nervous system, lungs and
mammary glands, and depending on the organs affected, can result in emaciation,
respiratory distress, mastitis, paralysis or arthritis (Dawson, 1988; Herrmann et al.,
2003b; Narayan et al., 1983) (Ravazzolo et al., 2006). Up to 40% of infected goats
develop chronic arthritis (Cheevers et al., 1997).
Bovine lentiviruses
Currently there are 2 lentiviruses known to infect cattle, BIV and JDV. The first
bovine lentivirus discovered was BIV (Tobin et al., 1994; Van der Maaten et al.,
1972) while JDV has more recently been identified as a lentivirus (Chadwick et al.,
1995b).
26
While the clinical and pathological syndromes associated with the 2 bovine
lentiviruses are markedy different, the viruses share a very close genetic and
antigenic relationship (Chadwick et al., 1995b). These are shown in Table 2.2.
Table 2.2. Comparison of putative protein products of JDVTab87 and BIV127.
JDV BIV
Protein Amino acids
Mol. Mass (kDa)
Amino acids
Mol. Mass (kDa)
Amino acid identity
gag precursor 436 48.8 476 53.4 62%
MA 125 14.3 126 14.6 60%
p2L - - 22 2.5
CA 226 25.3 219 24.6 75%
p3 - - 25 2.7
NC 85 9.2 66 7.3 63%
p2 - - 18 1.9
gag/pol precursor 1432 163 1475 168 66%
pol precursor 1027 118 1035 118 68%
env precursor 781 88.8 904 102 31%
SU 422 47.8 555 62.1 24%
TM 359 41.1 349 40.2 39%
vif 197 22.9 198 22.8 55%
tat 97 10.7 103 11.7 54%
(alt. tat)a (114) (12.5) - -
rev 213 23.8 186 20.7 35%
(alt. rev)a (201) (22.4) - -
tmx 164 18.5 159 18.0 29%
vpw - - 54 6.6
vpy - - 80 9.5 aAlternate forms of tat and rev generated by utilisation of alternative splice donor
site. Data sourced from Chadwick et al. (1995b).
27
Bovine immunodeficiency virus
BIV is a naturally-occurring lentivirus found in cattle and based on serological
surveys, infections occur worldwide (Campbell et al., 1998; McNab et al., 1994;
Meas et al., 2000a; Suarez et al., 1995). The virus may be transmitted horizontally by
iatrogenic routes, blood-sucking arthropods, via natural or artificial insemination or
via milk from infected cows (Egberink et al., 1992; Meas et al., 1998; Meas et al.,
2000a). BIV is best known as a virus infection of B. taurus but New Zealand white
rabbits (Oryctolagus cuniculus) and sheep have been infected experimentally,
resulting in a persistent infection (Gonda, 1992; Jacobs et al., 1994; Pifat et al.,
1992).
Genome structure
The first isolation of BIV was from a cow in Louisiana and this isolate, designated
R29 (Van der Maaten et al., 1972), has since been cloned (Braun et al., 1988) and
sequenced (Garvey et al., 1990). BIV-R29 is antigenically and genetically stable
during long-term, persistent infection (Carpenter et al., 2000).
The BIV proviral genome is 8.96 kb in size and consists of the 3 main ORF gag, pol
and env and 6 accessory genes rev, tat, vif, tmx, vpw and vpy (Figure 2.2). The
functions of vif, tmx, vpw and vpy in BIV are yet to be described (Egberink et al.,
1992; Gonda et al., 1987; Miller et al., 2000; Tobin et al., 1994). The BIV Gag
polyprotein that is translated is processed into a number of proteins, the 3 major
proteins found in all lentiviruses (MA, CA and NC) and 3 smaller proteins (p2L, p3
and p2) (Tobin et al., 1994).
28
Cell tropism and clinical signs associated with disease
Two studies have indicated that BIV may be pantropic. An initial in vivo study with
BIV isolate FL112 found proviral DNA in CD3+, CD4+ and CD8+ cells, B cells,
monocytes and WC1 cells in both the acute and chronic stages of infection
(Whetstone et al., 1997). A subsequent study involving the R29 isolate detected
proviral DNA and infectious virus within CD2+ (located on CD4+ and CD8+ T-
cells), WC1+ (located on γδ T-cells), mature B cells and monocytes during the early
stages of infection (Heaton et al., 1998). However, it is not clear from either study
whether these cell types were productively infected, as only proviral DNA was
detected. BIV-R29 is able to productively infect primary cultures of embryonic
bovine spleen and lung, Madin-Darby bovine kidney cells and embryonic rabbit
epidermal cells producing syncytia, cell lysis and infectious virus (Ferens et al.,
2007; Gonda, 1992; Heaton et al., 1998; Pifat et al., 1992; Suarez et al., 1993;
Whetstone et al., 1990; Zhang et al., 1997a; Zhang et al., 1997b).
BIV is most efficiently transmitted via infected blood and cell-free and cell-
associated tissue culture-derived virus (Pifat et al., 1992). The re-use of contaminated
needles during vaccination and blood collection, communal sharing of colostrum fed
to calves and the failure to cleanse contaminated instruments, have also been
suggested to play a role in the spread of BIV (Gonda, 1992). Inoculation usually
results in a subclinical infection without significant clinical effects (Flaming et al.,
1993) but clinical signs have been reported and include a transient leucocytosis,
lymphoid hyperplasia, lymph node enlargement, wasting and in some cases immune
suppression (Carpenter et al., 1992; Egberink et al., 1992; Zhang et al., 1997b). The
impact of BIV infection to the overall health of herds has not been established (Tobin
et al., 1994), although there is one Canadian study which suggested BIV infection
29
had a significant effect on milk yield (McNab et al., 1994). Other studies have
suggested that persistent BIV infection plays a role in reducing functional immune
competence as shown by the frequent development of concurrent infections in BIV-
infected animals (Meas et al., 2000a; Snider et al., 2003a; Zhang et al., 1997b). It
would seem, however, that a direct role for BIV in chronic progressive disease or as
a cofactor in a specific disease is unlikely (Jacobs et al., 1994; Suarez et al., 1995).
Jembrana disease virus
In contrast to BIV, JDV causes a significant acute disease process in Bali cattle (Bos
javanicus) in Indonesia (Dharma, 1997; Kertayadnya, 1997). The first reported
outbreak of Jembrana disease occurred in Sangkargung, a village in the Jembrana
district of the island of Bali in Indonesia (Figure 2.6) in December 1964. The disease
only affected Bali cattle and while reference is sometimes made to an effect on
buffalo (Bubalus bubalis) this was anecdotal and has not been confirmed. The
disease spread through all districts of Bali by August 1965 with an estimated 20,000
to 70,000 deaths. The disease then disappeared but subsequent smaller outbreaks
were detected in 1972 in the Tabanan district and in 1981 in the Karangasem district
of Bali (Ramachandran, 1996; Soeharsono, 1997b).
30
Figure 2.6. (A) The location of Bali in relation to other Indonesian islands (D.F.A.T., 2008). Districts of
Bali province, adapted from Streetdirectory.com (2009). Jembrana disease was first reported in the district
of Jembrana. Antibodies to JDV have since been detected in Sumatra, Java and Kalimantan. Clinical
disease and virus have also been detected in Sumatra and Kalimantan.
A B
31
Early investigators proposed Rinderpest virus and then a rickettsia as probable causes
of Jembrana disease (Soeharsono, 1997b). A viral aetiology was subsequently
demonstrated, a conclusion based on filtration studies that suggested the infectious
agent in blood was too small for a rickettsia (Ramachandran, 1996). Based on its
estimated size of 50-100 nm, the probable presence of a lipid-containing envelope,
electron microscopic observations and reverse transcriptase activity, the agent was
then considered as a probable member of the family Retroviridae (Kertayadnya,
1997; Wilcox et al., 1992) and was subsequently identified, based on genome
structure and genomic nucleotide sequence analysis, as a lentivirus (Chadwick et al.,
1995b).
Significance of Jembrana disease
The case fatality rate resulting from experimentally induced Jembrana disease is
approximately 21% (Desport et al., 2009a). During the original outbreak
commencing in 1964, it was retrospectively estimated that 60% of the Bali cattle and
buffalo population on Bali island were affected in the following 12 months, of which
98.9% died (Ramachandran, 1996); this high prevalence and high case fatality rate is
unlikely and the evidence is obscure as at the time there were no veterinary services
on the island. The effect of JDV is widespread throughout Bali, as a consequence of
the important role that Bali cattle play in society. They are a source of employment
and help to sustain and generate profit in the agricultural system via the provision of
draught power and manure which in turn helps to improve soil fertility
(Wiryosuhanto, 1997).
Transmission and distribution of Jembrana disease
The probable modes of transmission of JDV include mechanical and contact
transmission. Virus can be mechanically transferred from infected cattle to
32
uninfected cattle by iatrogenic means or by arthropods. Examples of iatrogenic
routes include multi-use needles during vaccination programs, as has been shown to
occur in EIAV (Li et al., 2003). Arthropod transmission is thought to be limited to
the acute viraemic episode when there is a high titre of virus in blood of affected
cattle, but it requires reasonably close contact between animals and the restriction of
movement of infected animals by quarantine has helped to reduce the spread of the
virus. The likelihood of contact transmission is supported by the detection of the
virus in secretions such as saliva, urine and milk and the ability to reproduce the
disease by conjunctival and oral inoculation of the virus (Hartaningsih et al., 1993;
Soeharsono et al., 1995b).
The disease has now been detected in 4 Indonesian islands: Bali, Kalimantan
(Indonesian Borneo), West Sumatra and East Java (Figure 2.6). There are no reports
of JDV infection of Bali cattle in any other area of Indonesia and clinical Jembrana
disease has not been reported in other cattle types which has led to the belief that the
disease is specific to Bali cattle (Hartaningsih et al., 1993; Wilcox et al., 1995).
However, other cattle types and buffalo can be infected experimentally and become
infected under field conditions (Soeharsono et al., 1995a) although they do not seem
to develop clinical disease. Bos taurus, B. indicus, crossbred Bali cattle (B. javanicus
x B. indicus) and buffalo can be infected and develop viraemia (Soeharsono et al.,
1990; Soeharsono et al., 1995a). The clinical changes and lesions that occur in these
cattle types are consistent with those observed in Bali cattle, but they are much
milder and would be difficult to detect under field conditions.
33
The JDV genome
The genome of JDV is 7732 bp in length and is the smallest of all lentivirus
genomes. It contains the 3 major ORF typical of all retroviruses, gag, pol and env, as
well as accessory genes, vif, tat, rev and tmx (Figure 2.2) (Chadwick et al., 1995b;
Chen et al., 1999; Nilsen et al., 1996).
JDV is a unique lentivirus in that it is genetically stable, and there is a high level of
nucleotide conservation in gag, pol and env sequences taken from isolates throughout
Indonesia. Gag sequences in isolates from Bali and Sumatra had 97 to 100%,
nucleotide identity. Env sequences were also unexpectedly conserved with 96-99%
nucleotide identity, and 95 to 99% amino acid identity. The largest divergence was
seen in an isolate from South Kalimantan with only 88% identity to that of the
original JDVTab87 isolate (Desport et al., 2007).
This high level of nucleotide conservation is similar to the LTR regions of EIAV that
have been reported to be highly conserved during persistent infection of horses
(Maury et al., 2005; Reis et al., 2003). It is in contrast to other lentiviruses such as
HIV-1 (Balfe et al., 1990; Lamers et al., 1993; Wolfs et al., 1990), FIV (Brown et al.,
1994; Duarte et al., 2006) and CAEV (Valas et al., 2000) which exhibit greater levels
of genetic variation, particularly in envelope glycoprotein regions, in sequences
obtained from within one individual (Lamers et al., 1993; Simmonds et al., 1990),
between individuals and between different geographic locations (Balfe et al., 1990;
Maki et al., 1992).
34
Cell tropism and clinical signs associated with Jembrana disease
It has been suggested that during the early stages of JDV infection, initial rounds of
virus replication occur in lymphoid tissue before a rapid and widespread
dissemination of virus to other tissues during the second phase of replication
(Chadwick et al., 1995a). JDV has a tropism for mature IgG-containing cells
(Desport et al., 2009a) and JDV-infected cells are found within spleen, lymphoid
tissues, bone marrow, lung, liver and kidney, as shown by in situ hybridisation
techniques for the detection of JDV genomic RNA (Chadwick et al., 1998).
JDV is not a typical lentivirus in that it causes an acute disease syndrome after a
short incubation period (Chadwick et al., 1995a). The incubation period before the
onset of clinical signs is 4-12 days and the clinical signs continue for 5-12 days
(Soeharsono, 1997a). Major clinical signs include a febrile response, lethargy,
anorexia and enlargement of superficial lymph nodes. Less frequently observed
clinical signs include erosions of oral mucous membranes, hypersalivation, nasal
discharge, diarrhoea and blood in the faeces. JDV has also been implicated in the
suppression of the humoral immune response in infected cattle, perhaps not
surprising as the histopathological changes reflect a disease primarily affecting the
lymphoid system (Dharma et al., 1994; Wareing et al., 1999). The major
haematological changes associated with JDV infection include leucopenia,
lymphopenia, eosinopenia and a slight neutropenia. In addition to these changes, a
mild thrombocytopenia, a normocytic normochromic anaemia, elevated blood urea
concentrations and reduced total plasma protein are also observed during the acute
disease (Soesanto et al., 1990).
35
Immune response to infection with a lentivirus
The immune response to lentiviruses frequently takes longer to develop than in many
other virus types. For example, ponies infected with EIAV do not seroconvert to
envelope glycoproteins and CA until 3 weeks pi, with Env antibodies being
predominant. However, it requires 6-8 months for the EIAV humoral immune
response to fully mature into a high avidity, conformational epitope-specific response
(Hammond et al., 1997). The EIAV-specific CTL activity develops 3-4 weeks pi and
this correlates with the resolution of the primary viraemia (Hammond et al., 1997).
Studies involving the depletion of CD8+ lymphocytes in rhesus monkeys
subsequently infected with SIVmac support the role of these cells in controlling
viraemia (Schmitz et al., 1999) as do other studies of natural HIV-1 infections in
humans (Koup et al., 1994). Neutralising antibody is detected 2-3 months pi
(Hammond et al., 1997). In sheep infected with VMV, the seroconversion to CA also
occurs about 3 weeks pi (Singh et al., 2006) although serum neutralising antibodies
are not detectable until 1-3 months pi (Petursson et al., 1976), slightly earlier than in
EIAV. The time course of the immune response to HIV infection is summarised in
Figure 2.7 and its relationship to the development of other features of the virus
infection are shown in Figure 2.8.
36
Also of importance in limiting infection with lentiviruses are non-adaptive immune
responses. Studies of HIV and FIV have shown that natural killer cells have an
important role in controlling acute infection (Howard et al., 2010) while cytidine
deaminases such as the previously mentioned APOBEC family of proteins restrict
viruses shortly after they have entered the cell by interfering with viral DNA
formation (Goila-Gaur et al., 2008, Romani et al., 2009). Other important innate
immune responses include TRIM5α which is thought to inhibit HIV-1 transduction
in rhesus macaque cells, non-coding RNAs such as micro RNAs and silence inducing
RNAs (Strebel et al., 2009), anti-viral cytokines such as interferon and dendritic cells
(Williams et al., 2009).
37
Figure 2.8. Estimated time course for maturation of the host immune response in EIAV-infected
animals. Image from Leroux et al. (2004). Env-specific cytotoxic T-cells appear within the first month
after infection as do Env- and p26 (CA)-specific IgG. Neutralising antibodies appear after 2-3 months
after infection.
Figure 2.7. Estimated time course for host immune response to acute HIV infection. Image from
Levy (2007).
38
Humoral immune response to infection with bovine lentiviruses
A number of assays have been used to detect and quantify the immune response to
the bovine lentiviruses, most commonly Western immunoblotting (WIB) and ELISA.
Antigen can be prepared using cell culture systems (Burki et al., 1992; Hammond et
al., 1997; O'Rourke et al., 1988; Whetstone et al., 1991; Whetter et al., 1990),
sucrose-gradient purified plasma from infected animals (Hartaningsih et al., 1994) or
recombinant protein production (Bird et al., 1993; Burkala et al., 1999; Burkala et al.,
1998). Other techniques have been used to monitor the immune response to bovine
lentivirus infections, including immunofluorescent assays (O'Rourke et al., 1988),
agar gel immunodiffusion (AGID) (Burki et al., 1992; Coggins et al., 1972;
Hartaningsih et al., 1994; Whetter et al., 1990) and radioimmunoprecipitation (Beyer
et al., 2001; Morin et al., 2003; O'Rourke et al., 1988). Western immunoblot assays
were more sensitive than AGID tests for detection of antibody to BIV (Whetstone et
al., 1991) and for JDV an ELISA was also more sensitive than AGID (Hartaningsih
et al., 1994).
Experimental infection of Bali cattle with JDV results in the production of antibodies
detectable by ELISA and WIB to the CA protein 6-11 weeks pi, although
occasionally animals can seroconvert as early as 2 weeks pi (Desport et al., 2009a;
Hartaningsih, 1997). The antibody response to JDV peaks 23-33 weeks pi and
persists beyond 59 weeks (Hartaningsih et al., 1994). As JDV has not been cultured
in vitro, there have been limited investigations of the neutralising antibody response
to JDV. There is only a single report available of a neutralising antibody response to
JDV, which relied on the infection of cattle to determine virus neutralisation. The
evidence from this study suggested that neutralising antibody to JDV developed only
after a prolonged period following recovery, and therefore that neutralising antibody
39
did not seem essential in the recovery from acute Jembrana disease. Antibodies
capable of neutralising JDV were detected in cattle 4 and 28 months after cattle had
recovered from clinical disease and titres were low, in the range of 1:2 to 1:20
(Hartaningsih et al., 2001).
In experimental BIV infections, antibody develops earlier than in JDV infections.
Antibody to the CA protein can be detected as early as 14 days post infection (dpi)
with titres peaking 6-8 weeks pi. Multiple studies have confirmed the early CA
antibody response and determined that it persists for up to 2.5 years after initial
infection (Heaton et al., 1998; Suarez et al., 1995; Whetstone et al., 1990; Whetstone
et al., 1991) although there is a single report that found the CA response to BIV
infection decreased dramatically by 40 weeks pi (Isaacson et al., 1995). Antibodies
capable of neutralising BIV have been detected as early as 17 weeks pi and persist
for 44 months pi (Carpenter et al., 2000). Antibodies against BIV TM could be
detected 4 weeks pi, peaking 10-30 weeks pi and persisted for at least 50 weeks pi
(Scobie et al., 1999); in some cattle they were detected 4 years pi (Isaacson et al.,
1995). The antibody response to the SU protein of BIV was reported to take many
months to develop (Suarez et al., 1995).
The CA and TM proteins of BIV and JDV contain cross-reactive epitopes which has
made serological differentiation between infections with JDV and BIV impossible
(Desport et al., 2005; Kertayadnya et al., 1993; Lu et al., 2002). Attempts have been
made to develop reagents capable of differentiating between these infections but
these have been unsuccessful (Desport et al., 2005).
40
Assays for the detection of lentivirus infections in ruminants
Small ruminant lentiviruses
Serological methods such as ELISA (using whole virus, recombinant proteins or
synthetic peptides as antigens), competitive ELISA (using monoclonal antibodies),
AGID, radio-immunoprecipitation and WIB are most commonly performed to detect
SRLV infection and there are a number of kits available commercially (Brinkhof et
al., 2007; Brodie et al., 1993; de Andres et al., 2005; Herrmann et al., 2003a; Pasick,
1998). It is important to note, however, that no “gold standard” method of diagnosis
exists (Brinkhof et al., 2007; de Andres et al., 2005).
Of a number of assays recently evaluated by Brinkhof and van Maanen (2007) for
the serodiagnosis of SRLV infections in sheep and goats, the best performing assay
was an ELISA. This assay involved the sensitisation of the solid phase with a
combination of recombinant VMV CA protein produced in E. coli and a TM derived
peptide. The performance of this assay may be due to the simultaneous detection of
antibodies against CA and TM. Antibodies to CA are produced early after infection
but decline once clinical signs appear and TM antibodies are produced later but
persist into the clinical phase. Therefore any assay which combines the detection of
antibodies to these 2 proteins could be expected to cover a greater proportion of the
infection period (Boshoff et al., 1997; Brinkhof et al., 2007).
A number of supplementary tests are often performed to confirm or resolve
indeterminate ELISA results, including AGID, WIB and radioimmunoprecipitation,
and these can be supplemented by PCR assays for the detection of proviral DNA or
viral RNA in PBMC or plasma (Barlough et al., 1994; Brinkhof et al., 2007; Brodie
et al., 1993; de Andres et al., 2005; Wagter et al., 1998; Zanoni et al., 1992).
41
Large ruminant lentiviruses
Diagnosis of BIV infection plays a crucial role in controlling the spread of infection
and the pathogen-free preparation of vaccines prepared using cattle, for example tick
fever vaccines (Lew et al., 2004). Assays used for detecting BIV infection include
qPCR (Lew et al., 2004), nested PCR, cell culture syncytium assays, WIB assays
(Meas et al., 1998; Meas et al., 2000a; Snider et al., 2003b; Suarez et al., 1995;
Zhang et al., 1997a), and ELISA using recombinant CA and TM protein antigens
(Barboni et al., 2001; Burkala et al., 1999). Techniques to quantify proviral and viral
BIV loads in tissues have not been described.
All attempts to culture JDV in vitro have failed and it has not been possible to detect
JDV infections using virus isolation techniques (Kempster et al., 2002; Wilcox et al.,
1992). Two techniques have been described that enable the detection and
quantification of JDV virus load, the JDV gag quantitative Reverse Transcriptase-
PCR (qRT-PCR) assay and a JDV p26 antigen capture ELISA (Stewart et al., 2005).
The qRT-PCR technique was optimised with plasma samples taken from animals
experimentally infected with JDV, and it was found to be robust and sensitive, with
an apparent sensitivity 100-fold greater than that of a standard RT-PCR. The capture
ELISA was relatively insensitive when compared to the qRT-PCR, although it
provided an economical method for monitoring of virus in the absence of more
sensitive techniques (Stewart et al., 2005).
42
Difficulties with serological testing
A number of factors have been reported to be critical in the interpretation of
serological assays. In lentivirus infections, one problem encountered is the delayed
nature of the antibody response. For example, in donkeys experimentally infected
with EIAV, the animals respond weakly and produce antibodies 42 dpi while horses
produce antibodies which can be detected as early as 16 dpi (Spyrou et al., 2003).
Hence, testing donkeys less than 42 dpi will yield a false-negative result. Problems
with delayed antibody responses have also occurred in CAEV (Brinkhof et al., 2007;
Rimstad et al., 1993) and JDV infections (Desport et al., 2009a; Hartaningsih et al.,
1994). Another problem in lentivirus infections is that antibody titres to viral proteins
may fluctuate between positive and negative after infection, though reasons for this
are unclear (de Andres et al., 2005).
Problems can occur in the detection of neutralising antibody in non-primate
lentivirus infections (Pozzetto et al., 1986; Sahu et al., 1994). Some North American
strains of VMV do not appear to induce neutralising antibodies (Zink et al., 1987). In
other VMV infections, the time required to develop detectable levels of neutralising
antibody after infection may vary considerably, from 12-14 dpi (Bird et al., 1993)
through to 1-3 months (Petursson et al., 1976). The titre of neutralising antibody
reported in response to lentivirus infections has also varied considerably in different
systems: in response to CAEV in goats it was reported to be of low titre and low
affinity (Kennedy-Stoskopf et al., 1986; Pisoni et al., 2007) as was the response of
horses to EIAV infection (O'Rourke et al., 1988) but the titre of neutralising antibody
in sheep in response to VMV infection was reported to be as high as 1:640 (Narayan
et al., 1978).
43
Another important factor which should be considered in lentivirus serological tests is
cross-reactive epitopes on viral proteins, which can make serological differentiation
of antigenically related viruses difficult. Antigenic cross-reactivity between structural
proteins of lentiviruses, particularly the CA protein, is well known (Cheevers et al.,
1988; Gnann et al., 1987a). Cross-reactivity has been reported between the CA
protein of JDV and BIV (Burkala et al., 1998; Desport et al., 2005; Kertayadnya et
al., 1993), between the TM glycoprotein of BIV and JDV (Burkala et al., 1998) and
the SU glycoprotein of CAEV and VMV (Gogolewski et al., 1985; Valas et al.,
2000), among others.
Techniques to identify epitopes and distinguish between viral infections
Peptide mapping is a useful technique to identify specific epitopes that differ
between 2 viruses that cross-react antigenically (Valas et al., 2000; Van
Regenmortel, 1999b). The reactivity to peptides produced in this manner provides
information on the antigenic characteristics of similar viruses infecting an animal,
which permits differentiation between the virus infections (Gnann et al., 1987a;
Mordasini et al., 2006; Pisoni et al., 2007). Peptide mapping involves the generation
of a panel of overlapping peptides which cover the entire amino acid sequence of the
protein of interest. These peptides are chemically synthesised and used in immune
binding assays, usually in an ELISA format for high throughput, against a panel of
reference sera taken from natural and experimental infections (Ball et al., 1992;
Bertoni et al., 1994). Synthetic peptides have been used, for example, to identify
epitopes that differentiate HIV-1 and HIV-2 infections (Gnann et al., 1987a; Gnann
et al., 1987b).
Another technique to identify unique epitopes is phage display (Xiao et al., 2008).
This technique is made possible by the expression of peptide libraries on the surface
44
of filamentous phage particles. The first step in the process involves the insertion of
one random oligonucleotide into one phage. The peptide encoded by the
oligonucleotide is then expressed within the pIII or pVIII coat protein of the
filamentous phage fd or M13 (D'Mello et al., 1999). These random phage display
libraries are subsequently screened for reactive epitopes with an antibody of interest.
The peptide sequence can be determined after amplifying the DNA from the selected
phage by PCR and sequencing techniques (D'Mello et al., 1999; Van Regenmortel,
1999a; Westwood, 2000; Williams, 2000).
Another technique suitable for defining epitopes is the use of protein expression
libraries. To construct expression libraries, DNA from a coding sequence of interest
is digested with DNase I to generate random DNA fragments with an approximate
average of 200 bp in length. These fragments, which can encode peptides, are ligated
into vector DNA (for example λgt11) whilst preserving the reading frame of a fusion
protein such as β-galactosidase. Ligations are packaged into Escherichia coli and the
cloned fragments are expressed as fusion proteins from the recombinant phages. The
random libraries are subsequently screened for reactive epitopes with an antibody of
interest. The peptide sequence can be determined after amplifying the DNA from the
selected phage by PCR and then sequencing this product (Bertoni et al., 2000;
Bertoni et al., 1994; Pancino et al., 1993; Van Regenmortel, 1999a).
One traditional method of defining epitopes is the limited fragmentation of proteins
by chemical cleavage, using cyanogen bromide or enzymatic digestion.
Fragmentation is then followed by immunoblotting of protein fragments (Westwood,
2000). Other methods that have been used include crystallographic analysis of
antigen-antibody complexes and the binding of anti-peptide antibodies to either
natural or chemically modified proteins (Van Regenmortel, 1999a).
45
Lentivirus superinfection
Dual infection of an animal with 2 viruses can be the result of coinfection or
superinfection. Coinfection occurs when 2 viruses infect at or near the same time
prior to seroconversion. Superinfection is the infection of an animal (or a cell) by 2
genetically distinct viruses where the infection with one virus precedes infection with
the second virus, sometime after seroconversion (Blackard et al., 2002; Gottlieb et
al., 2004; Jurriaans et al., 2008).
A natural case of superinfection of goats with CAEV and VMV has recently been
documented (Pisoni et al., 2007). Several studies have reported superinfection of
HIV-1 infected individuals (Gottlieb et al., 2004; Jurriaans et al., 2008; Piantadosi et
al., 2007; Smith et al., 2006). Between 2002 and 2005, 16 cases of HIV-1
superinfection in humans were reported (Smith et al., 2005). Although this number
seems small, natural cases of superinfection have generated considerable interest as
they challenge the assumption that HIV-1 specific immune responses generated
against primary infection are protective against subsequent infection by different
strains of the same virus (Allen et al., 2003).
Benefits of superinfection - superinfection resistance
Experimental studies have shown that pre-infecting an animal with a relatively less
pathogenic virus protects against challenge with a second, more pathogenic
lentivirus, a situation referred to as superinfection resistance (Table 2.3). This has
been demonstrated in FIV (Terwee et al., 2008; VandeWoude et al., 2002), SIV
(Cranage et al., 1998; Nilsson et al., 1998; Stebbings et al., 2004) and heterologous
SHIV infections (Sealy et al., 2009).
Domestic species of cats that have been asymptomatically infected with non-
pathogenic lion lentivirus (FIVple) or puma lentivirus (FIVpco) have shown resistance
46
to subsequent challenge with pathogenic FIV (Brown et al., 1994). All cats became
infected with the pathogenic FIV but prior exposure to FIVple or FIVpco ameliorated
the normal clinical effects of FIV infection: CD4+ cell depletion was reduced and, in
some cases, plasma and PBMC FIV loads were reduced (Terwee et al., 2008;
VandeWoude et al., 2002).
Infection with live attenuated SIVmacC8 of cynomolgus macaques ameliorated the
effects of subsequent infection with pathogenic wild-type SIVmacJ5 (Stebbings et al.,
2004), its derivative SIVmac220 (Cranage et al., 1998) and SIVsm (Nilsson et al., 1998).
This was shown by a significant decrease of cell-associated virus and plasma viral
RNA loads (Cranage et al., 1998; Stebbings et al., 2004) and negative virus isolation
in a proportion of animals (Nilsson et al., 1998). Indian rhesus macaques infected
with SIVmacC8 also resisted superinfection with the virulence-reverted form of
SIVmacC8 (Sharpe et al., 1997). Likewise, Indian rhesus macaques infected with
SIVmacGX2 were completely protected against challenge with SIVmac220 (Sharpe et al.,
2004) and macaques infected with SIVmac239∆nef were protected against challenge
with SIVmac251 (Connor et al., 1998; Daniel et al., 1992).
Superinfection resistance holds promise as a way to ameliorate the effects of
lentivirus infection and disease, in effect to act as a potential vaccine. It has been
proposed that if the mechanism of protection conferred could be better understood,
then a safe and effective HIV vaccine, for example, could be developed (Cranage et
al., 1998; Stebbings et al., 2004). The effect of challenge with a non-pathogenic virus
on the course of infection with a pathogenic virus also offers an opportunity to
examine host-virus and virus-virus interactions and their effect on pathogenicity and
resistance to virulent lentivirus infections (VandeWoude et al., 2003).
47
Mechanisms of superinfection resistance
The mechanisms responsible for protection against superinfection are not well
defined. Protection has not been directly correlated with humoral or cellular immune
responses including virus neutralising activity (Connor et al., 1998; Cranage et al.,
1998; Daniel et al., 1992; Nilsson et al., 1998; Sharpe et al., 1997; Stebbings et al.,
2004; Stebbings et al., 2002; VandeWoude et al., 2002) and in some cases protection
is not dependent on challenge-driven expansion of immunodominant epitope-specific
CD8+ T-cells (Sharpe et al., 2004). While CD8+ T-cells are important for control of
primary viraemia, they do not seem to play a central role in protection against
superinfection (Stebbings et al., 2005). Non-immunological phenomena such as virus
interference or antiviral factors such as CD8 suppression factors induced by defective
particles have been suggested as playing a role in superinfection resistance (Cranage
et al., 1998; Stebbings et al., 2004; Stebbings et al., 2002). Virus interference has
been detected in vitro when cell cultures were infected with a retrovirus and were
relatively resistant to infection by a related retrovirus; the phenomenon occurs only
when both viruses share the same receptor and results from a restricted penetration
into the cell (Corbin et al., 1993).
Adverse consequences of superinfection
While the positive effects of superinfection, namely resistance, have been well
described, superinfection or mixed infection of animals with 2 viruses has resulted in
potentially adverse effects. Goats infected naturally with both CAEV and VMV were
shown to contain chimeric viruses with CAEV-VMV envelope glycoproteins; this
was expected to have dramatic effects on the species specificity of the viruses and
their capacity to cross species barriers (Pisoni et al., 2007).
48
Table 2.3. Experimental superinfection studies conducted in a range of lentivirus animal model systems.
Model system Primary virus infection
Secondary pathogenic virus strain
Time between primary and secondary infections (weeks)
Effect on pathogenic virus Reference
FIVpco FIV-C 4 Reduced CD4+ T-cell depletion. (Terwee et al., 2008)
FIVpco or FIVple FIV-B 27 Reduced CD4+ T-cell depletion, reduced plasma and PBMC FIV load.
(VandeWoude et al., 2002) FIV in domestic cat
FIVPetaluma FIV-M2 28 Reduced total viral load, reduced CD4+ T-cell depletion.
(Pistello et al., 1999)
SIVmac251 or SIVsmE660
Reciprocal infection
Reduction in peak viraemia, amelioration of infection.
(Yeh et al., 2009)
SIV in rhesus macaques
SIVmacGX2 (nef-disrupted)
SIVmac220 89 or 122 Complete resistance (determined by negative virus isolation).
(Sharpe et al., 2004)
SIVmacC8 SIVmac32H/L28 3 or 20 Reduction of viral RNA and DNA load.
(Berry et al., 2008) SIV in cynomolgus macaques Attenuated
SIVmacC8 SIVmacJ5 3
Reduced cell-associated virus loads, reduced plasma virus load.
(Stebbings et al., 2004)
49
Chapter 3: Attempts to detect Bovine immunodeficiency virus infection in
Bali cattle in Indonesia with a PCR-based assay
Summary
Attempts were made to provide evidence for the occurrence of BIV in cattle in Indonesia.
One hundred and seventy one genomic DNA and serum samples were taken from Bali
cattle in the Bangli and Tabanan regions on the island of Bali. Genomic DNA samples
extracted from PBMC were screened for the presence of BIV or JDV proviral DNA using
both real time and conventional PCR methods and direct sequencing of any amplified
products to confirm their identity. Serum samples were screened for antibodies against
JDV using a range of antigens in a WIB or ELISA format and 21 of the 171 animals were
identified as being seropositive by a positive WIB reaction with the p26 CA protein of
JDV and at least one other positive serological test. BIV proviral DNA was not detected in
any of the cattle but JDV proviral DNA was detected in 12 animals, only one of which
was seropositive.
50
Introduction
Two bovine lentiviruses are suspected to circulate in the Bali cattle population of
Indonesia. The presence of Jembrana disease virus (JDV) is well documented and both
the disease and antibodies to the virus have been detected in cattle on the islands of Bali,
Sumatra and Java (Hartaningsih et al., 1993). The disease also now occurs in all
Kalimantan provinces in Indonesian Borneo (Hartaningsih, personal communication).
Despite the widespread distribution of Bali cattle in the eastern islands of Indonesia,
clinical Jembrana disease has not been reported in these areas. However, there are reports
of JDV antibody-positive cattle in some of the regions of Indonesia that are free of clinical
Jembrana disease, including on the island of Sulawesi (Desport et al., 2005), suggesting
the presence of a second non-pathogenic virus that is antigenically related to JDV,
possibly BIV. Serological evidence was also presented for the presence of a BIV-like
virus in Bali cattle in Bali where JDV is endemic (Barboni et al., 2001) although this has
not been confirmed by virus isolation.
The objectives of the investigations reported in this thesis were to attempt confirmation of
the presence of non-pathogenic BIV-like viruses in Bali cattle in Bali, to develop
methodology for the detection and differentiation of infection by these viruses and JDV,
perhaps enabling the determination of the distribution of each virus in Indonesia, and to
investigate the interaction between the 2 viruses in infected Bali cattle. Ideally, these
investigations required the isolation of the non-pathogenic BIV-like virus that is reputedly
present in Indonesia, that would then enable its experimental inoculation into animals not
only to determine the effects of these viruses in Bali cattle but also for the production of
reagents. Use of the local non-pathogenic virus would eliminate the need to import an
51
exotic strain of BIV for these investigations. Unsuccessful attempts were made previously
to isolate a non-pathogenic bovine lentivirus from cattle in Sulawesi: blood samples were
obtained from antibody-positive Bali cattle in Sulawesi, the samples were transported by
air to Bali and inoculated into Bali cattle on arrival. The inoculated cattle did not
seroconvert to BIV or JDV, suggesting the virus was not present in the inoculum. Further
attempts to detect the Sulawesi virus were abandoned due to political unrest in the area
from where cattle had been sampled and the expense involved (Hartaningsih, personal
communication).
This Chapter describes the results of attempts to confirm the report of Barboni et al.
(2001) that Bali cattle on the island of Bali were infected with BIV, and from which
attempts to isolate the virus could be made.
Materials and methods
Field samples
Peripheral blood samples were taken from 171 Bali cattle (Bos javanicus) in 2 districts
(Bangli and Tabanan) on the island of Bali, Indonesia. They were collected from the
jugular vein, into sterile 10 ml vacutainer tubes containing 15% EDTA (BD) for
extraction of genomic DNA, and into plain tubes for preparation of serum samples for
serological tests.
Preparation of PBMC genomic DNA
After collection into vacutainer tubes containing EDTA, the Bangli samples were
centrifuged (900 g for 10 min at 4°C) and the buffy coat was subsequently transferred into
10 ml tubes containing 6 ml distilled H2O, then mixed by inverting 3 times. Three ml of
2X PBS was then added and mixed by inverting 3 times after which tubes were
52
centrifuged again (250 g for 10 min at 4°C). The resulting supernatant was discarded and
the pellet washed twice with 10 ml PBS before the pellet was gently resuspended in 1 ml
Hank’s medium supplemented with 20% v/v heat inactivated foetal calf serum and 6% v/v
DMSO (Sigma) and stored in 200 µl aliquots at -20°C until required. Genomic DNA was
extracted from PBMC using the FlexiGene DNA Kit (Qiagen) according to the
manufacturer’s instructions and stored at -20°C until used. The concentration of PBMC
genomic DNA in each sample was determined using a spectrophotometer (Nanodrop ND-
1000).
After collection into vacutainer tubes, the Tabanan blood samples were centrifuged (900 g
for 10 min at 4°C) and the buffy coat was collected and mixed with 2 ml of PBS, then
overlayed onto 6 ml Ficoll (Amersham) in a sterile 10 ml tube and centrifuged at 400 g for
20 min at 4°C. The cells at the interphase were collected and washed 3 times in PBS by
mixing with PBS followed by centrifugation (400 g for 20 min at 4°C). The washed cells
were then resuspended with 1 ml PBS and stored at -20°C until required. Genomic DNA
was extracted from the purified PBMC using the QIAamp DNA Mini Kit (Qiagen)
according to the manufacturer’s instructions and the DNA was stored at -20°C until used.
The concentration of PBMC genomic DNA in the samples was determined using a
spectrophotometer.
Verification of DNA quality
Extracted DNA samples were initially tested by PCR using gene-specific glyceraldehyde
3-phosphate dehydrogenase (GAPDH) primers to test for integrity of the DNA. If no PCR
product was obtained with these primers then the sample was discarded. The GAPDH
primers used were as described previously (Mohan et al., 2001) and the reaction consisted
53
of 200 ng of PBMC genomic DNA, 1X PCR buffer, 1.25 mM MgCl2, 0.2 mM of each
dNTP, 0.8 mM of each primer (GAPDH F,
CCTTCATTGACCTTCACTACATGGTCTA, GAPDH R,
GCTGTAGCCAAATTCATTGTCGTTACCA; Invitrogen), 0.687 U Taq polymerase, and
ultrapure water to a final volume of 25 µl. All reagents were sourced from Fisher Biotec
unless otherwise stated. Thermal cycling conditions were 1 cycle of 94°C for 3 min, 35
cycles at 94 °C for 30 s, 60°C for 30 s and 72°C for 45 s, a final extension step of 72°C
for 7 min and they were then held at 14°C in a Bio-Rad thermal cycler.
qPCR for detection of JDV proviral DNA
The JDV proviral DNA genome copy number was quantified using a DNA plasmid-based
standard curve derived using JDV plasmid clone #139 as a template as previously
described (Stewart et al., 2005). The JDV sequence within the pT7T3 vector spanned JDV
nucleotides 19 to 2881 (U21603). The qPCR assay specifically amplified 118 bp in JDV
gag. All reactions consisted of 1X iQ Supermix (100 mM KCl, 40 mM Tris-HCl (pH 8.4),
1.6 mM dNTPs, 50 U/ml of iTaq DNA polymerase, 6 mM MgCl2, undefined stabilisers,
Bio-Rad), 0.6 mM of each primer (gag1f-GGGAGACCCGTCAGATGTGGA, gag1r-
TGGGAAGCATGGACAATCA; Invitrogen) prepared as described previously (Stewart et
al., 2005), 0.1 µM fluorogenic probe (Geneworks), 200 ng of extracted PBMC genomic
DNA and made up to a final volume of 10 µl using ultrapure water (Bio-Rad). Thermal
cycling conditions were the same as previously described (Stewart et al., 2005), except
that the reverse transcriptase step was omitted.
54
Conventional PCR for detection of JDV proviral DNA
A number of DNA samples which were tested by qPCR but provided results below the
cut-off for a positive result but where the results were clearly greater than that for the
JDV-negative control DNA, were re-analysed using conventional PCR. The primer pair
for detecting JDV proviral DNA by conventional PCR (JDV 1:
GCAGCGGAGGTGGCAATTTTGATAGGA, JDV 3:
CGGCGTGGTGGTCCACCCCATG) were located within the gag open reading frame
and specifically amplify a 360 bp fragment (Desport et al., 2007). Reactions conditions
consisted of 1X buffer, 1 mM MgCl2, 0.2 mM of each dNTP, 0.88 mM of each primer
(Invitrogen), 1.374 U Taq polymerase, 400 ng PBMC genomic DNA and made up to a
final volume of 50 µl with ultrapure water. Unless otherwise stated, all reagents were from
Fisher Biotec. Thermal cycling conditions were the same as those described (above) for
the GAPDH primer pair. Reaction conditions in the second round of amplification, where
necessary, were the same as the first except 1 µl of first round PCR product was added
into 25 µl reaction volumes.
Sequence analysis of PCR products
Direct DNA sequencing was performed to confirm and compare the proviral DNA
detected in the 2 cattle populations. If one band was produced by conventional PCR
amplification, the PCR product was purified using a PCR Purification Kit (Qiagen)
according to the manufacturer’s instructions. If multiple bands were produced, the PCR
product of the correct size was excised from the agarose gel and purified using a Gel
Extraction Kit (Qiagen) according to the manufacturer’s instructions. Ten ng of purified
product was sequenced using 1 µl of Big Dye Terminator (Applied Biosystems), 1.5 µl of
55
Big Dye Terminator sequencing buffer (Applied Biosystems), 3.2 pmoles of JDV 1 or
JDV 3 primer and made up to a volume of 10 µl using ultrapure water (Fisher Biotec).
The sequencing reaction consisted of a 2 min hold at 96°C and 25 cycles of 96°C for 10 s,
60°C for 30 s and 60°C for 4 mins. The sequencing reaction was purified by ethanol
precipitation according to the protocol supplied by Applied Biosystems. Samples were
then sequenced using an ABI 3730 48 capillary machine at the State Agricultural
Biotechnology Centre, Murdoch University.
Sequences were edited using Chromas Lite and aligned using the ClustalW program
(http://www.ebi.ac.uk/Tools/clustalw2/index.html). Phylogenetic analysis was performed
using Phylogeny.fr (http://www.phylogeny.fr/version2_cgi/index.cgi) and phylogenetic
trees were edited using MEGA (http://www.megasoftware.net/index.html).
qPCR for BIV proviral DNA
All animals were tested for the presence of BIV proviral DNA using a qPCR assay as
described previously (Lew et al., 2004), with the following modifications: 1X iQ
Supermix (Bio-Rad), 100 ng of each primer (BIVF1-
ACAAAAACTACGGGAATACCCTACA, BIVR1-
TCTTTTAGATCTCTGTGGGCTCTTTC; Invitrogen), 0.1 mM of probe (6FAM
CCACAATCCCAGGGAGT; Applied Biosystems) and 200 ng of PBMC genomic DNA.
The reaction volume was made up to 10 µl using ultrapure water (Fisher Biotec). The
qPCR assay specifically amplified 73 bp in BIV pol. The limit of quantification of the
assay was 20 copies per reaction (determined by amplification of specific known
quantities of cloned viral DNA).
56
Serological tests
Serum samples were tested for the presence of antibodies to JDV using ELISA or WIB
with a range of antigens. The assays were conducted by Dr J. Lewis (this laboratory)
using antigens that included a recombinant JDV p26-his, recombinant fused JDV
p26/TM-his, JDV TM peptide and plasma-derived JDV native antigen (kindly provided
by N. Hartaningsih). The WIB assay that utilised native JDV antigen derived from the
plasma of infected cattle was conducted as described previously by Hartaningsih et al.
(1994); results were recorded as positive if there was reactivity with the 26 kDa CA
protein. All other ELISA and WIB assays were conducted and interpreted as described
previously (Lewis, 2009).
The recombinant JDV p26-his construct expressed full-length JDVTab87 CA (Barboni et
al., 2001). The construct was kindly supplied by Dr. Margaret Collins, transformed into
BL21 (DE3) E. coli for protein expression and was purified using Ni-NTA agarose resin
in chromatography columns. The fused p26/TM construct was generated in a previously
described manner (de Andres et al., 2005; Rosati et al., 2004) whereby JDV capsid was
fused directly to the putative TM principle immunodominant domain (PID) epitope,
transformed into E. coli BL21 and purified as described for the recombinant JDV p26-his.
The JDV TM peptide ELISA was prepared as previously described (Ditcham et al., 2009)
and encompassed the PID of JDV TM (Barboni et al., 2001).
As there was no “gold standard” test for JDV antibody detection, samples were considered
“positive” when a positive result was obtained in WIB using the plasma-derived JDV
antigen and at least one other assay was also positive. Antibody to JDV and BIV cannot
be differentiated due to the presence of numerous cross-reactive epitopes on the CA
57
(Desport et al., 2005; Kertayadnya et al., 1993) and TM proteins (Burkala et al., 1998).
Therefore, the use ofthese antigens in WIB assays will equally detect antibody to both
JDV and BIV.
Results
Serology
Twenty one of the 171 cattle from which DNA was also analysed were seropositive for
antibody to the p26 CA of JDV by WIB utilising native JDV antigen and at least one other
assay (Table 3.1).
PCR
The GAPDH gene was detected by PCR in PBMC genomic DNA samples from 171 cattle
and these were deemed to have DNA of sufficient quality to test using qPCR or
conventional PCR. These 171 samples were then screened for the presence of JDV
proviral DNA by PCR and JDV-specific PCR products were detected in 12 of the 171
PBMC genomic DNA samples tested (Table 3.1). The sequences had between 99 and
100% homology with the reference JDVTab87 strain. The sequences generated are shown in
Figure 3.1.
With the exception of one animal (Tabanan Y26), all cattle in which JDV proviral DNA
was detected were seronegative (Table 3.1). In the 11 seronegative and PCR positive
cattle, 10 (83%) were negative in every serological assay and one (Bangli 35) was positive
in one of the 5 serological assays.
BIV proviral DNA was not detected in any of the 171 PBMC genomic DNA samples
tested.
58
Table 3.1. Results of PCR diagnostic assays and serological assays (ELISA and WIB) in
171 samples taken from Bali cattle (B. javanicus) in the Bangli and Tabanan region of
Bali, Indonesia. The results shown are those from 52 cattle where any assay provided a
positive result, and where samples provided negative results in all assays the results are
not shown. Italics indicate the sample was positive by both JDV PCR and serology.
Animal
identification
Age
(years)a
BIV
PCR
JDV
PCRb Antigen and serological assay
Final
serological
resultc
CA
ELISA
CA
WIB
Fused
CA/TM
ELISA
Fused
CA/TM
WIB
Native
antigen
WIB
Bangli 5 - - - - + - - -
Bangli 9 - + - - - - - -
Bangli 15 - - - - + - - -
Bangli 17 - - - - + - - -
Bangli 18 - - - - + - - -
Bangli 20 - - - - + - - -
Bangli 27 - + - - - - - -
Bangli 35 - + - - + - - -
Bangli 50 - + - - - - - -
Tabanan R3 5 - - + + - + + +
Tabanan R4 4 - - + + - + + +
Tabanan R9 6 - - + + - + + +
Tabanan R10 6 - - + + + + + +
Tabanan R12 6 - - - + - + - -
Tabanan R16 5 - - + + - + + +
59
Animal identification
Age (years)a
BIV PCR
JDV PCRb Antigen and serological assay
Final serological
resultc
CA ELISA
CA WIB
Fused CA/TM ELISA
Fused CA/TM
WIB
Native antigen WIB
Tabanan R17 4 - - - + - + - -
Tabanan R18 4 - - + + - + + +
Tabanan R21 4 - - + - - - - -
Tabanan R26 5 - - + - - + + +
Tabanan R28 8 - - + + - + + +
Tabanan R33 10 - - + - + + + +
Tabanan R39 7 - - - - - + - -
Tabanan R43 6 - - + + - + + +
Tabanan R46 3 - - - - - + - -
Tabanan R48 5 - - - - - + - -
Tabanan Y5 4 - - + - - - + +
Tabanan Y15 1.6 - - - - - + - -
Tabanan Y21 6 - - + + - - + +
Tabanan Y22 5 - - + + - + + +
Tabanan Y26 5 - + + + + - + +
Tabanan Y31 7 - - + + - - + +
Tabanan Y43 8 - - + + - - + +
Tabanan Y45 7 - - + + + - - -
Tabanan B5 5 - + - - - - - -
Tabanan B6 4 - - + - - + + +
Tabanan B11 8 - - - + - - - -
60
Animal identification
Age (years)a
BIV PCR
JDV PCRb Antigen and serological assay
Final serological
resultc
CA ELISA
CA WIB
Fused CA/TM ELISA
Fused CA/TM
WIB
Native antigen WIB
Tabanan B16 9 - - - - - + - -
Tabanan B26 8 - - + - + + + +
Tabanan B35 2 - - + - + - - -
Tabanan B37 7 - - + - + + + +
Tabanan G1 12 - - + - + - + +
Tabanan G4 7 - + - - - - - -
Tabanan G6 7 months - + - - - - - -
Tabanan G7 7 months - - - - + - - -
Tabanan G8 7 - + - - - - - -
Tabanan G11 6 months - + - - - - - -
Tabanan G15 10 - - + - - - - -
Tabanan G16 14 - - + - + - + +
Tabanan G26 4 - - - - + - - -
Tabanan G29 3 - + - - - - - -
Tabanan G31 5 - + - - - - - -
Tabanan G49 8 - - + - + - - +
Total 0 12 26 17 18 21 21 21
a Age data not available for Bangli cattle. b Positive status given where a positive result was obtained in the qPCR or conventional PCR assay and confirmed by direct sequencing. c Positive status given where a positive result was obtained with the JDV plasma-derived antigen WIB and at least one other serological assay.
61
Bangli 27 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 Bangli 35 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 Bangli 50 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 G31 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 Y26 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 59 G29 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 G11 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 G8 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 G6 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 B5 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 G4 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 Bangli 9 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 JDV/Tab 87 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCAGAAAAGTATTGGGAAGCATGG 60 JDV/Pul 01 CTGCTGGCGGGCCACCACCCAGAAAATTCAGATATGGCCGAAAAGTATTGGGAAGCATGG 60 BIV127 CTGCTGGCGGGGTACAAACCAGAGAGTACAGAAACGGCCCTAGGATATTGGGAGGCCTTT 60 ********** ** * ***** * * **** * *** * ******** ** * Bangli 27 ATAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 Bangli 35 ATAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 Bangli 50 ATAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 G31 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 Y26 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 119 G29 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 G11 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 G8 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 G6 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 B5 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 G4 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 Bangli 9 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 JDV/Tab 87 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 JDV/Pul 01 ACAATCAGAGAAAGGGAGTCCCAAAAGGAGGAGGAAGGAGAAATTACCAGCATCTACCCA 120 BIV127 ACATATAGAGAAAGGGAGGCCAGAGCTGATAAGGAAGGCGAAATTAAGAGTATTTACCCT 120 * * ************ ** * ** ******* ******* ** ** ***** Bangli 27 CAACTTAGAAAGAACT------------------------------------ 136 Bangli 35 CAACTTAGAAAGAACT------------------------------------ 136 Bangli 50 CAACTTAGAAAGAACT------------------------------------ 136 G31 CAACTTAGAAAGAACT------------------------------------ 136 Y26 CAACTTAGAAAGAACT------------------------------------ 135 G29 CAACTTAGAAAGAACT------------------------------------ 136 G11 CAACTTAGAAAGAACT------------------------------------ 136 G8 CAACTTAGAAAGAACT------------------------------------ 136 G6 CAACTTAGAAAGAACT------------------------------------ 136 B5 CAACTTAGAAAGAACT------------------------------------ 136 G4 CAACTTAGAAAGAACT------------------------------------ 136 Bangli 9 CAACTTAGAAAGAACT------------------------------------ 136 JDV/Tab 87 CAACTTAGAAAGAACT------------------------------------ 136 JDV/Pul 01 CAACTTAGAAAGAACT------------------------------------ 136 BIV127 TCCCTAACACAGAACACACAGAATAAGAAGCAGACATCGAATCAGACAAACA 172 ** * * *****
Figure 3.1. Sequence alignment of gag nucleotide sequences from 12 animals with JDV
taken from Bangli and Tabanan. Reference sequences JDVTab87 (accession number
U21603), JDVPul01 (accession number DQ229295) and BIV127 (accession number
NC_001413.1) are included for comparison. Sequences were aligned using ClustalW2.
“*” indicates all nucleotides are identical in that column.
62
Discussion
Although JDV proviral DNA was detected in the PBMC of 12 of 171 cattle examined,
which included cattle from 2 adjacent districts of Bali, BIV proviral DNA was not
detected in any of the animals tested. The result does not provide proof of the absence of
BIV in the Bali cattle population of Bali but it also does not provide any support for the
serological evidence reported by Barboni et al. (2001) that cattle within Bali are infected
not only by JDV but with a second antigenically related but presumably non-pathogenic
bovine lentivirus. The methodology used by Barboni et al. (2001) involved the production
of recombinant BIV CA and JDV CA and their use in a WIB format or a combination of
the recombinant CA with a BIV or JDV TM peptide in an ELISA format. Sera were
screened initially using the JDV antigens, then JDV antibody negative sera were screened
using the BIV antigens; if the serum reacted in the second set of assays the sample was
declared BIV seropositive and JDV seronegative. Given the high level of cross-reactivity
between JDV and BIV CA and TM antigens (Desport et al., 2005; Kertayadnya et al.,
1993), including the cross-reactivity of the antigens used in the study, these findings
should be interpreted with considerable caution. Results from this laboratory (Desport et
al., 2005) have shown that the reagents used by Barboni et al. (2001) would not
differentiate between JDV and BIV antibody.
The attempt to identify cattle infected with BIV by the detection of proviral DNA was
unsuccessful but not surprising. It is difficult to detect BIV in naturally infected cattle and
with the exception of the reports describing 3 successful attempts in the USA and Japan
(Suarez et al., 1993; Van der Maaten et al., 1972) and one from Japan (Meas et al., 1998),
this has not been reported, although there have been limited reports of the detection of
63
proviral BIV DNA in naturally infected cattle (Lew et al., 2004; Meas et al., 1998; Meas
et al., 2000b; Snider et al., 2003b). In the current study, both antibody-positive and
antibody-negative cattle were examined, and 2 specific PCR assays were used. Either both
assays lacked the sensitivity to detect the level of provirus present in cattle, or maybe
BIV-like viruses are present in Bali but were not present in the populations sampled, or
possibly BIV-like viruses do not occur in the cattle population of Bali. Modifications to
the PCR assays, such as degenerate primers and less stringent PCR reaction conditions,
may have assisted in the detection of proviral DNA. However, based on previous
experience in this laboratory, alterations such as these have a tendency to produce false
positive results. Further investigation of the pathogenesis of BIV infection in Bali cattle is
needed to examine the kinetics of virus replication and persistence after infection and the
optimal time for sampling.
JDV proviral DNA was detected in 12 animals sampled, suggesting recent infection, even
though there was no evidence of clinical Jembrana disease immediately preceding and
following the sampling. This is the first reported detection of JDV in clinically normal
cattle. The lack of variation between the PCR amplicons suggests a common virus strain
was circulating in the cattle, although the lack of variation is also consistent with previous
studies reporting a high level of nucleotide conservation in gag sequences of JDV detected
in Bali (Desport et al., 2007) and they may not have arisen from a single source. All the 21
JDV-seropositive cattle were 4 years or older indicating minimal transmission of JDV
between the cattle in the preceding 4 years, and this is consistent with the lack of reports
of Jembrana disease outbreaks in Bali during this period (Hartaningsih, personal
communication).
64
Proviral DNA positive-seronegative animals are rare but not uncommon after natural
lentivirus infections, and have been reported in cattle infected with BIV (Meas et al.,
2000a), cats infected with FIV (Dandekar et al., 1992) and sheep infected with VMV
(Leginagoikoa et al., 2009). These observations, and the detection of JDV provirus in 12
cattle, only one of which was seropositive, highlights the difficulty associated with
conclusively detecting natural lentivirus infections. There are a number of possible
reasons for the lack of concordance between serological and PCR assays. Firstly, the
antibody response to JDV is delayed and has been reported to be detectable only from 11
weeks post infection (Hartaningsih et al., 1994), hence serological assays would only
detect antibodies in the period after this time and this is a period of active virus replication
(Stewart et al., 2005). The detection of JDV proviral DNA in seronegative animals
suggests recent infection with JDV, prior to seroconversion. Secondly, there are a
proportion of animals infected with JDV that do not mount an antibody response to the
CA antigen (Desport et al., 2009a; Ditcham et al., 2009). These animals are referred to as
atypical responders and account for 15% of all animals experimentally infected but it
seems unlikely that this would account for the lack of antibody in 11 of the 12
PCR-positive animals detected. Thirdly, JDV proviral DNA levels in the PBMC are
hypothesised to persist at very low levels in naturally infected cattle, similar to SRLV
infections (de Andres et al., 2005). JDV proviral DNA has been readily detectable in
experimentally infected cattle, even many months after infection but is normally detected
only with difficulty in naturally infected cattle (Desport, personal communication).
Further studies are required to characterise the response of Bali cattle to inoculation with
BIV, including whether the animals can become actively infected and whether the virus
65
will persist in the animals over time. It will also be of interest to determine if and at what
level BIV proviral DNA and viral RNA loads occur. The development of reagents that are
able to distinguish between BIV and JDV infections in an ELISA will help in the
identification of BIV infections and in Indonesia may clarify discrepancies between
genomic and antibody based assay results.
66
Chapter 4: Bovine immunodeficiency virus produces a transient
viraemic phase soon after infection in Bos javanicus
Summary
Infection of Bali cattle in Indonesia with a non-pathogenic bovine lentivirus similar
to BIV is suspected but efforts to detect the virus have been unsuccessful. To define
the kinetics of BIV infection and seroconversion in Bali cattle and determine the
optimal time for sampling for detection of virus in infected cattle, 13 cattle were
infected with the R29 strain of BIV and monitored for up to 65 days. No clinical
signs were observed in the infected cattle following infection. Proviral DNA was
detected in PBMC from 4-60 days with peak titres 20 dpi. There was a transient
viraemia from 4 to 14 dpi with a maximum titre of 1 x 104 genome copies/ml plasma.
An antibody response to the TM glycoprotein commenced 12 dpi but an antibody
response to the CA protein was detected in one animal only and not until 34 dpi. The
results indicated that detection of BIV in infected Bali cattle is similar to B. taurus
with levels of proviral DNA detectable during the early stage of infection. Based on
these results, a CA based serological assay would not identify the majority of
infected cattle.
67
Introduction
Bali cattle are particularly susceptible to JDV and develop an acute disease process
soon after infection. The acute disease is characterised by a transient febrile
response, enlargement of superficial lymph nodes, high virus titres in the plasma and
a number of haematological changes including leucopenia and thrombocytopenia.
The case fatality rate is about 21% and recovered animals are resistant to re-
challenge with the virus (Desport et al., 2009a; Soeharsono et al., 1990; Soesanto et
al., 1990). In contrast, breeds such as B. taurus (Fresian cattle) and B. indicus
(Ongole cattle) develop a mild febrile response but no other clinical signs of disease
(Soeharsono et al., 1990).
The effects of BIV in Bali cattle are unknown but the pathogenesis of BIV in
B. taurus has been investigated by several groups. Many of the studies that were
undertaken assumed that the virus would produce effects long after infection, akin to
many other lentiviruses, and they therefore followed the infections for long time
intervals. In experimentally infected B. taurus, the original R29 isolate caused no
major clinical signs in the period up to 27 months after infection (Flaming et al.,
1993; Zhang et al., 1997b) although subclinical changes in experimentally infected
cattle were reported by others. Subclinical changes reported have included
lymphocytosis and follicular hyperplasia (Carpenter et al., 1992) and immune
suppression at 3-7 weeks post-infection (Zhang et al., 1997b). Other isolates of BIV
(FL491 and FL112) were reported to cause a transient increase in PBMC (Suarez et
al., 1993). The FL112 isolate caused a transient B-cell lymphocytosis that peaked 14
dpi (Whetstone et al., 1997) and is also reported to cause lymphadenopathy and non-
suppurative meningoencephalitis 12 months post-infection (Munro et al., 1998).
Serological studies have also reported no major associations with significant clinical
68
changes but one study based in North America found associations between BIV and
decreased milk yield in dairy cattle (McNab et al., 1994) while another reported
marked weight loss with frequent and severe concurrent infections (Snider et al.,
2003b).
This Chapter reports the experimental infection of Bali cattle with BIV that was
conducted with 2 objectives. First, to determine if Bali cattle were more susceptible
to infection with BIV than B. taurus, similar to their greater susceptibility to JDV
than other cattle species (Soeharsono et al., 1995a). Second, to determine the kinetics
of virus replication and persistence of BIV and the development of the antibody
response after infection, so as to provide insights into the optimal periods in which to
sample naturally infected cattle to detect the virus. The experiments were conducted
with the R29 strain of BIV as attempts to detect BIV in Indonesian cattle were not
successful (Chapter 3).
Materials and methods
Animals
Nineteen cattle were obtained from Nusa Penida, an island adjacent to Bali where
Jembrana disease has never been detected and the cattle have been consistently
negative for antibody to JDV and BIV. The cattle were housed indoors as previously
described (Soeharsono et al., 1990). Six weeks prior to infection with BIV the cattle
were vaccinated against Bovine viral diarrhoea virus using Pestigard (Pfizer) twice
at 0 and 4 weeks apart as the FBL cell culture was contaminated with BVDV. BVDV
is a common contaminate of BIV cell cultures and foetal bovine serum (Makoschey
et al., 2003).
69
Virus
BIV-R29 was obtained from J Brownlie and M Collins, Royal Veterinary College,
UK, passaged in primary bovine foetal lung (BFL) cell cultures grown in RPMI
medium (Invitrogen) supplemented with 10% foetal bovine serum (Thermo
Scientific) and antibiotics in 75 cm2 flasks (Nunc). For inoculation into cattle,
infected cells exhibiting marked syncytium formation were scraped from the surface
of flasks 24 h after infection and the cells suspended in RPMI medium. The titre of
the virus was retrospectively estimated by titration in BFL cell cultures and the total
inoculum administered to each animal determined to be 1.38 x 106 50% tissue culture
infectious doses (TCID50).
Experimental infection and sampling of cattle
The infectious inoculum was divided into 2 equal amounts, one administered
intravenously and the other subcutaneously. Seven virus infected cattle (CB169-
CB175) were monitored for 42 days after infection and 6 (CB177-CB182) for 65
days after infection. An additional 6 cattle (CB183-CB189) were inoculated with an
equivalent volume of uninfected BFL cells in RPMI medium and monitored for 65
days as controls.
Animals were observed daily for the development of clinical signs of disease. Rectal
temperatures were measured daily for the duration of the study. Blood samples were
obtained as required by venipuncture of the jugular vein and used for determination
of total leukocyte counts, extraction of DNA from PBMC, extraction of RNA from
plasma and serum for serological tests.
The 6 cattle monitored for 65 days were killed and a complete post-mortem
examination conducted. Tissue samples (retropharyngeal and prescapular lymph
70
nodes, spleen, bone marrow, kidney, lung and thymus) were collected into RNAlater
(Ambion) for DNA and RNA extraction.
Extraction of DNA and RNA from tissues
Total DNA was extracted and purified from tissue samples using a DNeasy Tissue
Kit (Qiagen) according to the manufacturer’s instructions.
Total RNA was extracted and purified from tissues using the RNeasy Mini Kit
(Qiagen) according to the manufacturer’s instructions. Disruption and
homogenisation was performed using the TissueLyser (Qiagen) and an optional on-
column DNase digestion step using an RNase-Free DNase Set (Qiagen) was included
to remove any contaminating genomic DNA.
A Ficoll-Paque Plus (GE Healthcare) gradient was used to purify PBMC from
heparinised blood according to the manufacturer’s directions. Genomic DNA was
subsequently extracted from the cells using a QIAamp DNA Mini Kit (Qiagen).
Viral RNA was extracted from plasma using the QIAamp Viral RNA Extraction Kit
(Qiagen) as recommended by the manufacturer.
Quantitation of BIV proviral DNA
Extracted DNA samples were initially tested by PCR using gene-specific (GAPDH)
primers to test for sample integrity. If no product was obtained then the sample was
not used in the analysis. The GAPDH primers used were as described in Chapter 3.
Proviral DNA loads were determined with GAPDH-positive samples by a qPCR
assay as described in Chapter 3.
BIV Proviral DNA loads were normalised to GAPDH copy number according to
previously published methods (Terwee et al., 2008). Briefly, the number of cell
equivalents (CE) in 200 ng of PBMC genomic DNA or 200 ng of tissue DNA was
71
determined by qPCR using GAPDH-specific primers. All reactions consisted of 1X
iQ SYBR® Green Supermix (Bio-Rad), 400 µM of each primer (GAPDH156F:
GGTGATGCTGGTGCTGAGTA, GAPDH342R: GGCACTGCTGACAATCTTGA;
Invitrogen), 200 ng of extracted DNA and were made up to a final volume of 10 µl
using ultrapure water (Fisher Biotec). Thermal cycling conditions were 1 cycle of
95°C for 3 min, 40 cycles of 95°C for 15 s, 60°C for 30 s, 72°C for 30 s and a final
extension step of 72°C for 10 min. The DNA-based plasmid standard was generated
using the Mohan et al. (2001) GAPDH primers. The BIV proviral DNA copy number
generated using the protocol from Lew et al. (2004) was then expressed as the
number of BIV proviral DNA copies/100 000 cell equivalents (CE). The number of
CE was determined using the GAPDH qPCR protocol.
When DNA yields from PBMC were less than 50 ng/ul and insufficient for qPCR,
samples were tested by conventional PCR using primers (Heaton BIV F, 5’
CCCCAGGTCCCATCAACATTCATC and Heaton BIV R, 5’
GTCTTCCCACATCCGTAACATCTCC) as previously described (Heaton et al.,
1998).
Quantification of BIV RNA
Viral RNA loads were determined using qRT-PCR with primers and probes as
described for the qPCR for provirus except that 2 µl of viral RNA, 0.2 µl iScript
Reverse Transcriptase for One-Step RT-PCR (Bio-Rad) and 1X iScript RT-PCR
reaction mix for probes was included per reaction and a 10 min reverse transcription
incubation step at 50°C was included at the start of the reaction. To determine viral
loads within the tissues, the above protocol was followed except 200 ng of tissue
RNA was included in each reaction instead of 2 µl of viral RNA. Results were
expressed in BIV viral RNA genome copies/µg of total RNA.
72
Enzyme linked immunosorbent assays
The BIV TM antibody response was determined by ELISA with 1:25 dilutions of
sera and a peptide antigen as previously described (Barboni et al., 2001; Ditcham et
al., 2009), except that the 36 amino acid TM peptide
RVSYLEYVEEIRQKQVFFGCKPHGRYCHFDFGPEEV (Proteomics
International) of BIV-R29 was converted from linear to cyclic form (TMc) as
described previously (Scobie et al., 1999). BIV hyperimmune serum was included as
a positive control and serum from uninfected cattle was used as a negative control in
every plate.
The CA antibody response was determined by ELISA using a recombinant protein
antigen. To produce the CA protein, full length BIV CA protein was cloned into the
pTrcHisB vector and transformed into JM109 chemically competent E. coli. Bacteria
were grown overnight at 37°C at 225 rpm in a 50 ml culture of standard 2X YT
medium with ampicillin (50 µg/ml, Sigma). Forty ml of overnight culture was added
to 1 L of 2X YT medium plus ampicillin and grown at 37°C for 4 h at 225 rpm.
Protein production was induced by the addition of 1 mM isopropyl-beta-D-
thiogalactopyranoside (Sigma) for 4 h. The protein was purified as recommended in
the QIAExpressionist Handbook (Qiagen) and an ELISA protocol was optimised.
One hundred ng of protein was coated onto each well of a Maxisorp 96-well plate
(Nunc) and incubated overnight at 4°C. Plates were then washed 3 times with PBS-T
(137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, 0.05% Tween 20).
One hundred µl of serum diluted 1:100 in PBS-T with 5% w/v skim milk (Fonterra)
was added to each well for 1 h at 37°C. Plates were then washed 3 times with PBS-T
and 100 µl of a 1:2 000 dilution of anti-bovine IgG-HRP (ICN) in PBS-T-skim milk
was added to each well for 1 h at 37°C after which they were washed 3 times with
73
PBS-T and once with PBS. Colour development was induced by the addition of
peroxidase substrate (Bio-Rad) and the reaction stopped by the addition of 2% w/v
oxalic acid and absorbance read at 405 nm. BIV hyperimmune serum was included
in every plate as a positive control and serum from uninfected cattle was used as a
negative control.
Results
Clinical observations
The 13 Bali cattle infected with BIV-R29 and the 6 BFL-only controls did not
develop any clinical signs of disease during the observation period. Rectal
temperatures and total leukocyte counts remained normal throughout the experiment
and no gross lesions were observed during post-mortem examination of the 7 cattle
killed 65 days after infection.
Quantification of BIV proviral DNA load
Proviral DNA was detected in PBMC of all cattle inoculated with virus during the 65
days after infection. Proviral DNA was first detected 8 dpi in 4 of the 13 animals
(Table 1), it was detected in all 13 cattle 20 dpi but in only 2/13 at 40 dpi and 3/6 at
60 dpi. Maximum proviral DNA titres were 6.2 x102 proviral genome
copies/100 000 CE.
Quantification of BIV RNA load
Plasma viral RNA was detected in 8 of the 13 cattle from 4 to 14 dpi (Table 4.2).
Viral titres ranged from 4.2 x 101 to 1.0 x 104 genome copies/ml plasma.
74
Table 4.1. BIV proviral DNA detection and quantification by conventional and
qPCR in cattle inoculated with BIV-R29.
Days after infection
Animal
Identification 0 2 4 6 8 12 14 20 40 56 60
CB169 - - - NTa +b + 1.6x101 4.5 x101 - NAd NA
CB170 - - - - - - + 5.1 x101 - NA NA
CB171 - - - NT - + + + - NA NA
CB172 - - NT NT + - + + - NA NA
CB173 - - - - - 4.9x101 c - 4.0 x102 + NA NA
CB174 - - - NT - - 6.6x101 1.6 x102 - NA NA
CB175 - - NT - - - + 1.9 x102 + NA NA
CB177 - - NT NT + + + 3.3 x101 - NT -
CB178 - - - - - - 1.2x102 3.2 x101 - + -
CB179 - - - - NT - + + - + -
CB180 - - - - - - - + - - +
CB181 - - - - - + 7.7x101 6.2 x102 - - +
CB182 - - NT NT + + 6.8x101 2.2 x101 - + +
Percent
positive 0 0 0 0 33.3 46.1 84.6 100 15.3 60.0 50.0
aNT, not tested.
b+, positive result by conventional PCR.
cNumber of BIV proviral genome copies/100 000 peripheral blood mononuclear cell-
equivalents.
dNA, not available
75
Table 4.2. BIV viral RNA genome copies/ml of plasma determined by qPCR in cattle inoculated with BIV-R29.
Days after infection
Cattle 0-2a 4 6 7 8 9 10 11 12 14 15-34b 15-62c
CB169 - - - - - - - - - - -
CB170 - - - - - - - - - - -
CB171 - - - - - - - - - - -
CB172 - - - - - - - - - - -
CB173 - - - - 4.7 x103 - 5.9 x103 - - -
CB174 - - - - - 5.7 x103 - 5.4 x103 - - -
CB175 - - - - 4.2 x101 2.1 x101 - 3.2 x101 - - -
CB177 - - - - - - - - - - -
CB178 - - - - 2.0 x102 - - 3.2 x102 - - -
CB179 - - - 6.1 x101 - - 2.6 x102 6.3 x102 1.3 x102 - -
CB180 - - - - - 4.8 x102 - - - - -
CB181 - 1.0 x104 - 4.4 x101 1.0 x103 3.1 x103 6.5 x102 8 x101 - - -
CB182 - - - - - - - 3.4 x103 5.6 x103 6.0 x102 -
arepresents results from days 0,1 and 2 p.i. brepresents results from days 15, 16, 18, 20, 27 and 34 p.i. crepresents results from days 15, 16, 18, 20, 27, 34, 40, 41, 43, 45, 47 to 53, 55 to 58, 60 and 62 p.i.
76
Virus RNA and proviral DNA in tissues 65 days after infection
Proviral DNA and/or viral RNA were detected in all 6 cattle killed 65 dpi
(Table 4.3). Proviral DNA was detected in at least 1 tissue type of each animal and
most commonly in lymphoid tissues. Viral RNA was detected less frequently than
proviral DNA and only in the spleen and prescapular lymph nodes. The levels of
viral RNA detected by qRT-PCR were very low, bordering on undetectable.
Serological response to BIV-R29 infection
Antibody to the BIV TMc and CA antigens were detected by ELISA (Figures 4.1 and
4.2). Increased ELISA absorbance readings with the BIV TM c peptide were detected
12 dpi and were detected in most cattle by 20 dpi. The antibody response to the CA
antigen was markedly less than to the TMc peptide and a strong antibody response to
this protein was detected in one animal only between 34 and 56 dpi (Figure. 4.2).
None of the control group developed detectable TM or CA antibody.
77
Table 4.3. Viral RNA and proviral DNA quantification in tissues after necropsy of
experimentally infected Bali cattle.
Cattle Tissue
CB177 CB178 CB179 CB180 CB181 CB182
Provirusa + (<20c) + (<20) + (<20) - 21.8 21.1 Spleen
Viral RNAb + (<20) + (<20) 204.0e - - -
Provirus - + (<20) + (<20) + (<20) - + (<20) Lung
Viral RNA - - - - - -
Provirus - - - 36.6 - - Thymus
Viral RNA - - - - - -
Provirus - - - 3849.4 - - Bone marrow
Viral RNA - - - - - -
Provirus - + (<20) 3086.4 - + (<20) + (<20) Retropharyngeal
lymph node Viral RNA - - - - - -
Provirus - 6800.6d + (<20) + (<20) + (<20) - Prescapular lymph
node Viral RNA + (<20) 63.5 - - - -
Provirus - 29.28 - - - - Kidney
Viral RNA - - - - - -
aProviral genome in tissues quantified using qPCR. bViral RNA genome copies in tissues quantified using qRT-PCR. cSamples with values less than 20 but consistently above the negative control. dNumbers refer to BIV proviral genome copies/100 000 cell equivalents. eNumbers refer to BIV viral RNA genome copies/µg total RNA.
78
0 8 12 14 16 18 20 27 34 56 60-1
0
1
2
3
4
Days post infection
Abs
orba
nce
(405
nm
)
10 20 30 40 50 60-1
0
1
2
3
4
CB181
CB169CB170CB171CB172CB173CB174CB175CB177CB178
CB180
CB182
CB179
CB183
CB186CB187
CB184
Days post infection
Abs
orba
nce
(405
nm
)
Figure. 4.1. TM IgG in serum of cattle after infection with BIV-R29. Antibody was
detected by ELISA with a BIV TMc peptide antigen. Absorbances were normalised
to day 0 readings. ELISA absorbances from mock infected cattle (CB183 – CB187)
are also shown. Top: individual results in 17 cattle. Bottom: box and whisker plot of
the values in the 13 cattle infected with BIV.
79
10 20 30 40 50 60-1
0
1
2
3
4 CB169CB170CB171CB172CB173CB174CB175
CB177CB178
CB179
CB181CB182CB183
CB186CB187
CB180
CB184
Days post infection
Abs
orba
nce
(405
nm
)
0 8 12 14 16 18 20 27 34 56 60
-1
0
1
2
3
4
Days post infection
Abs
orba
nce
(405
nm
)
Figure. 4.2. CA IgG in serum of cattle after infection with BIV-R29. Antibody was
detected by ELISA with a BIV CA antigen. Absorbances were normalised to day 0
readings. ELISA absorbances from mock infected cattle (CB183 – CB187) are also
shown. Top: individual results in 17 cattle. Bottom: box and whisker plot of the
values in the 13 cattle infected with BIV.
80
Discussion
The detection of BIV provirus in PBMC in all 13 cattle over the course of the
experiment and within tissues at the end of the experiment 65 dpi, the transient
detection of viral RNA in plasma from 8 of 13 infected cattle and an antibody
response to BIV in inoculated cattle, confirm that the Bali cattle were productively
infected with BIV.
No clinical signs of infection were observed in any of the infected Bali cattle. The
absence of significant clinical effects is similar to other experiments where BIV had
been inoculated into B. taurus (Carpenter et al., 1992; Heaton et al., 1998; Isaacson
et al., 1995; Zhang et al., 1997a). The greater susceptibility to JDV of Bali cattle than
B. taurus is not reflected in their susceptibility to BIV. This absence of clinical signs
in Bali cattle in response to BIV would be consistent with the presence of a BIV-like
virus in Bali cattle on the island of Sulawesi in Indonesia where antibody to JDV has
been detected but there is no evidence of Jembrana disease (Hartaningsih, personal
communication).
BIV proviral DNA was detected in PBMC from 8 dpi until the conclusion of the
experiment at 65 dpi but was not detected in all animals at all sampling occasions.
The highest proportion of BIV proviral DNA positive animals was at 14 and 20 dpi
when 84.4 and 100% of animals were positive, suggesting this was a peak period of
virus replication and indicating an acute phase for BIV infection. After this period
the virus appeared to persist in PBMC, at a level that was often undetectable.
Modifications to the PCR assays may have assisted in the detection of very low
proviral DNA copy numbers.
Provirus was detected in a wide variety of tissues at 65 dpi when the experiment was
concluded, similar to a previous report of infection in B. taurus (Zhang et al., 1997a).
81
The levels of proviral DNA were low but highest in the lymphoid tissues examined,
spleen and lymph nodes. The low titres were similar to those reported in other
lentivirus infections, including African green monkeys persistently infected with
SIVagm (Gueye et al., 2004) and in horses asymptomatically infected with a cell-
adapted pathogenic EIAV (Harrold et al., 2000).
Although previous studies involving experimental infection of B. taurus with BIV
have detected viral RNA within PBMC subpopulations, either by RT-PCR (Baron et
al., 1995; Wu et al., 2003) or by in situ hybridisation (Carpenter et al., 1992), this
study appears to be the first report documenting the transient nature of the plasma
viraemia in the period soon after infection, similar to that reported in many other
lentivirus infections (Langemeier et al., 1996; Miyake et al., 2006; Ryan et al., 2003;
Stewart et al., 2005). While provirus was detected in all 13 infected cattle, viral RNA
was detected in 8 of 13 cattle and only during the period from 4 to 14 dpi. The levels
of virus RNA detected never exceeded 1 x 104 genome copies/ml plasma, much less
than those detected during infection of Bali cattle with the genetically related JDV
where titres of up to 1.6 x 1012 genome copies/ml plasma have been detected during
the acute disease (Stewart et al., 2005). The transient nature of the plasma viraemia
could be viewed as escape of the virus from host control. While provirus was
widespread in the tissues that were tested, viral RNA was detected in the spleen and
prescapular lymph node only of some animals and was at low levels. Perhaps the
level of replication of BIV relative to JDV may be associated with the relative lack of
pathogenicity of BIV compared to JDV.
There was a rapid and strong response against TM but a poor antibody response to
the CA protein in most infected cattle. The TM response to BIV infection was
similar to that reported by Scobie et al. (1999) in cattle infected with the BIV FL112
82
isolate and an antibody response was detected as early as 2 weeks pi. The weak CA
response detected in all but one of the 13 infected Bali cattle was surprising as in
B. taurus there is normally a strong response to CA between 2 and 4 weeks pi
(Isaacson et al., 1995; Whetstone et al., 1991). Seroconversion to a gag precursor
also occurred between 2 and 4 weeks pi in rabbits infected with BIV (Pifat et al.,
1992). The reason for the poor antibody response to CA in Bali cattle is unknown. In
response to infection with the genetically and antigenically related JDV, Bali cattle
normally mount a strong albeit delayed immune response against CA (Hartaningsih
et al., 1994) but as in the Bali cattle infected with BIV, a subset of cattle infected
with JDV mount a poor antibody response to the JDV CA and a strong antibody
response to JDV TM (Ditcham et al., 2009). A 10 to 100 times greater antibody
response to envelope proteins compared to CA proteins was also observed in horses
in response to EIAV (O'Rourke et al., 1988).
Results from this study indicate not only that BIV is non-pathogenic in Bali cattle but
that maximum virus replication occurred soon after infection and prior to the onset of
a significant antibody response, certainly prior to the onset of a significant antibody
response to the CA protein. It is likely that future attempts to detect BIV infection in
Bali cattle and possibly other cattle species using PCR based assays, would have
greatest chance of success soon after infection and before the onset of a significant
antibody response.
83
Chapter 5: Bovine immunodeficiency virus infection alters the
dynamics of subsequent Jembrana disease virus infection
Summary
To determine whether BIV infection is capable of protecting against superinfection
with JDV, 15 animals were infected with BIV-R29 and 42 days after BIV infection,
9 of the BIV infected and 4 mock BIV infected animals were superinfected with
JDVTab87. All cattle were successfully infected with BIV, shown by the presence of
proviral DNA and, in a subset of cattle, a transient viraemia. Strong antibody
responses against the TM glycoprotein and poor antibody responses against the CA
protein were also detected. Despite the development of immune responses against
TM, a region known to contain cross-reactive epitopes, all cattle became infected
with JDV, as indicated by the development of typical clinical signs of Jembrana
disease and an acute phase viraemia.
84
Introduction
The 2 bovine lentiviruses, BIV (Gonda et al., 1987) and JDV (Chadwick et al.,
1995a; Kertayadnya et al., 1993), are genetically and antigenically related but differ
markedly in pathogenicity. The incidence of clinical Jembrana disease and
serological surveys indicate JDV is common in the Bali cattle population in parts of
Indonesia (Hartaningsih et al., 1993). A BIV-like non-pathogenic bovine lentivirus is
also suspected to occur in the cattle population on Bali island (Barboni et al., 2001)
and antibody to JDV has been detected in Bali cattle on the island of Sulawesi where
there is no clinical evidence of Jembrana disease in the Bali cattle population
(Desport et al., 2005). Antibody to JDV and BIV cannot be differentiated due to the
presence of numerous cross-reactive epitopes on the CA (Desport et al., 2005;
Kertayadnya et al., 1993) and TM proteins (Burkala et al., 1998).
A protective immunity against JDV infection has been induced by vaccination with
inactivated whole virus antigens (Ditcham et al., 2009). It was considered possible
that infection of Bali cattle with a non-pathogenic BIV-like lentivirus might also
induce a protective immune response against Jembrana disease. Non-pathogenic
strains of other lentiviruses have been shown to induce protective immunity against
pathogenic strains of the same virus. Infection of domestic cats with non-pathogenic
lion lentivirus or puma lentivirus ameliorated the effects of subsequent wild-type FIV
infection (Terwee et al., 2008; VandeWoude et al., 2002). Similar results were
obtained in domestic cats pre-infected with chimeric FIV (generated by substituting
part of env of clade A FIVPET with a corresponding region of clade B FIVM2, or vice-
versa) and subsequently inoculated with FIVPET or FIVM2 (Giannecchini et al., 2007).
Infection of macaques infected with attenuated SIVmac also ameliorated the effects of
challenge with pathogenic SIVmac (Cranage et al., 1998; Sharpe et al., 2004) and
85
SIVsm (Nilsson et al., 1998) although attenuated SIVmac did not provide protection
against the more divergent HIV-2 (Nilsson et al., 1998).
This Chapter describes an experiment to determine whether prior infection of Bali
cattle with BIV would provide protection against superinfection with pathogenic
JDV infection 42 days after the initial BIV infection. This experiment was expected
to provide information that would increase our understanding of the effect JDV
would have on the Bali cattle population if it were introduced onto the island of
Sulawesi where BIV is suspected to occur in the cattle population of that island.
Materials and methods
Animals
Nineteen cattle approximately 18 months of age were obtained from Nusa Penida, an
island adjacent to Bali where Jembrana disease has never been detected and the cattle
have been consistently negative to antibody to JDV and BIV (Hartaningsih et al.,
1993). The cattle were housed indoors as previously described (Soeharsono et al.,
1990). Cattle were screened by PCR and ELISA to ensure they were not infected
with bovine lentiviruses prior to challenge (Chapter 3). Six weeks prior to infection
with BIV the cattle were vaccinated against BVDV with Pestigard® (Pfizer), as
previously described (Chapter 4).
Viruses
The BIV-R29 for infection of cattle was obtained from J Brownlie and M Collins,
Royal Veterinary College, UK. The virus was cultured in primary BFL cells and
titrated in BFL cells as previously described (Chapter 4). An estimated 1.82 x 104
TCID50 was inoculated into each animal, half subcutaneously and half intravenously.
86
The JDVTab87 (Chadwick et al., 1995b) used for infection of cattle was prepared by
infection of antibody-negative cattle with a suspension of frozen spleen harvested
from infected cattle as described previously (Soeharsono et al., 1990). Plasma from
the infected cattle was obtained 2 days after the development of a febrile response
typical of Jembrana disease, and the approximate ID50 in the plasma determined
using an antigen-capture ELISA as described previously (Stewart et al., 2005). An
estimated 1 000 ID50 of the virus was inoculated intravenously into each animal.
Experimental infection and sampling of cattle
Nine cattle were inoculated with BIV at day 0 and with JDV 42 days later (CB190–
CB197 and CB205). Six cattle were inoculated with BIV at day 0 and were not
subsequently inoculated with JDV (CB198–CB202 and CB204). Four cattle were
inoculated with uninfected BFL cells in RPMI medium at day 0 and subsequently
with JDV 42 days later (CB203, CB206, CB208 and CB210).
Animals were observed daily for clinical signs of disease. Rectal temperatures were
measured daily for the duration of the study. Heparinised blood samples were
obtained as required by venipuncture of the jugular vein and used for extraction of
DNA from PBMC, extraction of RNA from plasma and serum for serological tests.
Extraction of DNA and RNA from peripheral blood
PBMC were purified from 10 ml of heparinised blood using a Ficoll-Paque PlusTM
(GE Healthcare) gradient as recommended by the manufacturer. DNA was extracted
from the PBMC using a QIAamp DNA Mini Kit (Qiagen) as recommended by the
manufacturer. Viral RNA was extracted from plasma using the QIAamp Viral RNA
Extraction Kit (Qiagen) as recommended by the manufacturer.
87
Detection of BIV proviral DNA
The integrity of genomic DNA was confirmed using gene-specific GAPDH primers
as previously described (Chapter 4). BIV proviral DNA loads were determined using
a conventional PCR assay as described previously (Chapter 4).
Quantification of BIV and JDV RNA in plasma
BIV RNA was quantified as described previously (Chapter 4) and JDV RNA as
described by Stewart et al. (2005), with the following exceptions: 1X RT-PCR
Reaction Mix for Probes (Bio-Rad), 0.2 µl iScript Reverse Transcriptase for One-
Step RT-PCR (Bio-Rad) and 2 µl RNA extracted from plasma was added to each
reaction. The reaction was made up to 10 µl using nuclease free water (Bio-Rad).
ELISA
The BIV TM antibody response was determined by ELISA using a cyclic BIV TM
(BIV TM c) peptide as described previously (Chapter 4). The JDV TM antibody
response was determined by ELISA using a cyclic JDV TM (JDV TMc) peptide as
previously described (Ditcham et al., 2009) as while there was extensive cross-
reactivity between JDV and BIV TM antigens, our experience was that the JDV TMc
peptide provided greater sensitivity in the detection of JDV TM antibody than did the
BIV TM c antigen. The BIV CA antibody response was determined using an ELISA
with a recombinant BIV CA antigen as previously described (Chapter 4) except that
100 ng of protein was coated onto each well and serum was tested at a dilution of
1:100.
88
Analysis of data
To determine whether prior BIV infection would induce protection against
subsequent JDV infection, differences between the viral load (VL) on the first and
second day of the febrile period when the rectal temperature exceeded 39.3°C, the
peak VL, the duration and magnitude of the VL where the area under the curve
(AUC) was >106 genomes/ml, and the duration of the febrile period, were compared
in JDV-infected cattle that had been previously infected with BIV and in cattle not
previously infected with BIV, as described previously (Desport et al., 2009a;
Ditcham et al., 2009). The AUC where the VL was > 106 for each animal was
estimated and linear interpolation between consecutive observations, in combination
with the previously described piecewise-linear model was used to supply missing
data caused by variations in sampling intervals (Ditcham et al., 2009). The duration
of the infectious period was defined as the period where VL was > 106 genomes/ml.
A baseline of 106 was chosen (Ditcham et al., 2009) as bloodmeal residues of
between 4–10 nl have been reported on the mouthparts of tabanid flies and EIAV at
106 ID/ml in blood can be transmitted by a single fly (Foil et al., 1987). For analysis
of the febrile response, rectal temperatures were divided into ranges adapted from a
previously described method (Muraguri et al., 1999) as low fever (>39.3 °C – 40.2
°C), moderate fever (40.3 °C – 41.2 °C) and high fever (> 41.2 °C). Student’s t-test
was used for statistical comparison of the magnitude and duration of plasma VL and
febrile responses and P-values < 0.05 were considered statistically significant.
89
Results
Clinical and virological observations in cattle after BIV inoculation
Fifteen BIV inoculated Bali cattle and 4 mock-infected control cattle were monitored
for 42 days and then superinfected with JDV. During the initial 42 day period after
BIV infection, the BIV-infected cattle did not exhibit any change in rectal
temperature or other clinical signs of disease.
BIV proviral DNA was detected in PBMC initially 7 dpi in one animal and
subsequently in PBMC of all 15 cattle inoculated with BIV, confirming that all
animals were infected (Table 5.1). BIV proviral DNA was detected in most cattle
after 17 dpi but at no single time point was BIV proviral DNA detected concurrently
in all 15 cattle. BIV proviral DNA was not detected in the mock-infected cattle.
Plasma viral RNA was detected in 5 of the 15 cattle in the period from 8 to 13 dpi
(Table 5.2) and plasma viral titres ranged from 1.04 to 4.25 log10 genome copies/ml
plasma.
An antibody response to the BIV TMc and CA antigens was detected by ELISA
(Figure 5.1). Antibody to the BIV TMc peptide was detected from 10 dpi and in the
majority of cattle by 41 dpi (Figure 5.1A). Antibody to the BIV CA antigen was
detected from 19 dpi (Figure 5.1C) but the CA response was markedly less than the
response to the BIV TMc peptide (Figure 5.1A). Only some of the BIV-infected
cattle developed CA antibody and only one seroconverted strongly in the 42 day
period after infection. None of the mock-infected cattle developed BIV TMc or CA
antibody (Figure 5.1B and 5.1D).
90
0 7 10 13 17 21 28 35 41
0.0
0.5
1.0
1.5
2.0
2.5
Days after BIV inoculation
Abs
orba
nce
(405
nm
)
0 7 10 13 17 21 28 35 41
0.0
0.5
1.0
1.5
2.0
2.5
Days after mock BIV inoculation
Abs
orba
nce
(405
nm
)0 1 5 11 19 41
0.0
0.5
1.0
1.5
Days after BIV inoculation
Abs
orba
nce
(405
nm
)
0 1 5 11 19 41
0.0
0.5
1.0
1.5
Days after mock BIV inoculation
Abs
orba
nce
(405
nm
)
A B
C D
Figure 5.1. TMc and CA IgG response in serum of cattle after inoculation with BIV,
shown as box and whisker plots of the ELISA absorbance values. (A) Response
detected by BIV TMc peptide in all 15 cattle inoculated with BIV at day 0.
(B) Response detected by His-BIV CA antigen in all 15 cattle inoculated with BIV at
day 0. (C) Response detected by BIV TMc peptide antigen in all 8 cattle mock
inoculated at day 0. (D) Response detected by His-BIV CA antigen in all 8 cattle
mock inoculated at day 0.
91
Table 5.1. BIV proviral DNA detected by PCR in cattle inoculated with BIV and
prior to infection with JDV.
Days after BIV infection Animal
0 7 10 17 19 21 42
Cattle inoculated with BIV at day 0 and then JDV at day 42
CB190 -a - - +a - + - CB191 - - + + - + - CB192 - - NT + - + - CB193 - - NT + + + + CB194 - - - + + - + CB195 - - - + + + + CB196 - - - + + - - CB197 - - - + - + - CB205 - - - + + + + Cattle inoculated with BIV only at day 0 CB198 - - - - - - + CB199 - + + - + - + CB200 - - - NTb + - + CB201 - - + + + - - CB202 - - - + - - + CB204 - - + + - - + Cattle inoculated with JDV only at day 42 CB203 - - NT - - - - CB206 - - NT - NT - - CB208 - - - - - - - CB210 - - - - - - - Percent BIV-inoculated cattle positive 0 7 15 85 53 47 60 Cumulative % positive 0 7 15 93 93 93 100 Percent mock-inoculated cattle positive 0 0 0 0 0 0 0
a - and + denote negative and positive PCR results, respectively, for BIV proviral
DNA.
b not tested.
92
Table 5.2. BIV viral RNA genome copies/ml (log10) of plasma determined by qPCR in cattle inoculated with BIV at day 0 (all cattle)
and JDV at day 42 (CB190 – CB205 only).
Days after BIV inoculation Days after JDV inoculation
Animal 0 - 1a 7 8 9 10 11 13
14-
42c
44-
50d 51 52 53 54 55
56-
63e
Cattle inoculated with BIV at day 0 and JDV at day 42 CB190 -b - - - 1.20 - 4.25 - - - - - - - - CB191 - - - - - - 1.89 - - 1.40 - - - - - CB192 - - - - - - - - - - - - - - - CB193 - - - - 1.79 - - - - - - - - - - CB194 - - - - - - - - - - - 1.86 - - - CB195 - - - - - - - - - 2.18 - 2.57 2.51 1.48 - CB196 - - - - - - - - - - - - - - - CB197 - - - - - - - - - 1.79 - - - - - CB205 - - 1.04 2.45 - - - - - - - - - - - Cattle inoculated with BIV at day 0 only CB198 - - - - - - - - - - - - - - - CB199 - - - - - - - - - - - - - - - CB200 - - - - - - - - - - - - - - - CB201 - - - - - - - - - - - - - - - CB202 - - - - - - - - - - - - - - - CB204 - - - 1.04 - - - - - - - - - - - a represent results from 0 and 1 dpi. b negative. c represents results from 14 - 17, 19, 21, 28, 35 and 41 - 42 dpi. d represents results from 44, 46 and 48 - 50 dpi. e represents results from 56 - 59, 61 and 63 dpi
93
Clinical and virological observations after JDV inoculation
Forty two days after BIV infection, 9 of the 15 BIV-infected cattle and 4 of the
mock-infected cattle were inoculated with JDV and monitored for a further 15 to 19
days. All the JDV-inoculated cattle developed a febrile response but there were
differences in the responses in the previously BIV-infected and non-BIV-infected
groups (Figure 5.2). Excluding 2 animals CB190 and CB205 that responded
uncharacteristically to JDV infection and were considered atypical responders (see
below), the febrile response in the BIV-infected group started 2 days earlier (a mean
of 7 dpi compared to 9 dpi, P = 0.058), the peak of the febrile response occurred
earlier (a mean of 10 dpi versus 12 dpi, P = 0.067), the peak VL also occurred
significantly sooner (a mean of 10 dpi versus 12 dpi, P = 0.008) and there was a
significantly earlier conclusion of the febrile response (a mean of 13 dpi compared to
15 dpi, P = 0.033, Table 5.3). There were no significant differences between groups
in regards to the duration and severity of the febrile response (Table 5.3). The 6 BIV-
only controls did not develop a febrile response or other clinical signs during the
observation period (Figure 5.2).
JDV plasma RNA was detected in all cattle inoculated with JDV from 3 to 19 dpi
(Figure 5.3). The maximum VL was 2.18 x 1011 genome copies/ml plasma.
Excluding the 2 atypical responders, the cattle infected with JDV, regardless of
whether they had been challenged previously with BIV, developed a viraemia typical
of that reported previously (Desport et al., 2009a; Stewart et al., 2005). As shown in
Figure 5.3, the pattern and magnitude of the viraemia was also similar between
groups, but the viraemia started and finished earlier in the cattle infected previously
with BIV, although these differences were not statistically significant; accordingly,
in cattle previously infected with BIV there was a reduction in the mean duration of
94
the JDV viraemia that was >106 genomes/ml plasma (P = 0.068). There were no
significant differences between previously BIV-infected and non-BIV-infected
groups in the VL on the first or second day of the febrile period, the peak VL or the
total AUC when the viraemia was ≥106 genomes/ml plasma (Table 5.4).
Two animals, CB190 and CB205, responded atypically to JDV infection. The cattle
had a late onset fever (Figures 5.2 and Table 5.3) and the dynamics of their viraemia
were different to those in the other cattle (Figure 5.3), similar to atypical responders
previously reported (Desport et al., 2009a). These animals had an erratic viraemia
increasing to a maximum titre of approximately 1010 genome copies/ml which
continued through to the conclusion of the experiment. As the viraemia in these
animals did not decrease before the experiment concluded, the data provided for the
duration of the viraemia in Table 5.4 is an underestimation of the actual values.
After inoculation with JDV, plasma BIV RNA was again detected in 4 of the 9
superinfected cattle (Table 5.2). The titre of BIV RNA during this period ranged
from 1.40 to 2.57 log10 genome copies/ml plasma.
Antibody to the JDV TMc antigen was detected (Figure 5.4) in the 15 day period
after JDV infection in a majority of cattle previously infected with BIV (Figure
5.4A), but was not detected in any of the JDV-infected cattle that had not been
infected previously with BIV (Figure 5.4B). ELISA absorbances to the CA protein
increased at 4 days after JDV infection in the cattle that had been previously infected
with BIV (Figure. 5.4D). BIV CA ELISA absorbances remained low in the cattle
infected with JDV only (Figure 5.4F).
95
0 2 4 6 8 10 12 14 16 18 20 22
37.5
38.0
38.5
39.0
39.5
40.0
40.5
41.0
41.5
42.0
Days after JDV inoculation
Rec
tal t
empe
ratu
re (
°° °°C)
Figure 5.2. Mean rectal temperatures in cattle inoculated with JDV. Y-error bars
represent the standard deviations of rectal temperatures. Shown are the rectal
temperatures in cattle which had been previously inoculated with BIV and responded
to JDV in a typical fashion (○; CB191 – CB197), in cattle which had been
previously infected with BIV and responded to JDV in an atypical fashion (▼;
CB190 and CB205), JDV-only control cattle (●; CB203, CB206, CB208 and CB210)
and BIV-only control cattle JDV (■).
96
Table 5.3. Effect of previous BIV infection of cattle on the febrile response
following JDV infection.
Days after infection Duration of febrile response
(days)
Animal
Peak
VL
Onset of
febrile
response
Peak
febrile
response
End of
febrile
response
Low Moderate High Total
Cattle infected with BIV 42 days previously
CB190 18 17 18 21 1 3 0 4
CB191 8 5 8 13 4 2 2 8
CB192 9 8 10 12 3 1 0 4
CB193 12 11 13 16 4 1 0 5
CB194 9 4 10 12 8 0 0 8
CB195 11 8 10 15 3 3 1 7
CB196 9 7 10 12 1 3 1 5
CB197 9 7 9 13 3 3 0 6
CB205 18 17 20 21 0 3 1 4
Mean 11
(10)a
9
(7)a
12
(10)a
15
(13)a
3 2 1 6
Variance 15.28 22.75 17.75 13.50 5.50 1.36 0.53 2.75
Cattle not previously infected with BIV
CB203 12 8 10 15 2 5 0 7
CB206 12 10 13 15 4 1 0 5
CB208 11 10 11 14 1 2 1 4
CB210 12 9 12 17 4 4 0 8
Mean 12 9 12 15 3 3 0 6
Variance 0.25 0.92 1.67 1.58 2.25 3.33 0.25 3.33
P-valueb 0.441
(0.008)c
0.487
(0.058)c
0.412
(0.067)c
0.449
(0.033)c
0.425 0.153 0.233 0.376
a mean calculated excluding atypical responders CB190 and CB205. b P-values represent statistical differences between animals infected with BIV and then JDV and those infected with JDV only. c P-value calculated excluding atypical responders CB190 and CB205.
97
0 2 4 6 8 10 12 14 16 18 20
0
2
4
6
8
10
12
Days after JDV inoculation
log1
0 JD
V v
iral
RN
Age
nom
e co
pies
/ml p
lasm
a
Figure 5.3. Mean plasma viral loads (JDV RNA genome copies/ml plasma) in
animals inoculated with JDV at day 0. Y-error bars represent the standard deviations
of viral loads. Shown are viral loads in cattle which had been previously infected
with BIV and responded to JDV in a typical fashion (o; CB191 – CB197), JDV-only
control cattle (●; CB203, CB206, CB208 and CB210), and cattle which had been
previously infected with BIV and responded to JDV in an atypical fashion
(▼; CB190 and CB205).
98
Table 5.4. Effect of previous BIV infection on the dynamics of the JDV viral load
(VL) in plasma after JDV inoculation.
Animal VL 1st day of
febrile period
(log10)
VL 2nd day of
febrile period
(log10)
Peak VL
(log10)
AUC ≥106
genome
copies/ml (log10)
Total
days
VL≥106
Cattle infected with BIV 42 days previously
CB190 6.61 9.77 11.34 10.15 5.60
CB191 7.35 10.63 10.12 11.76 11.20
CB192 8.49 9.59 9.59 9.71 7.40
CB193 8.89 9.26 9.26 9.68 8.60
CB194 1.91 4.64 9.98 10.25 7.60
CB195 9.23 10.12 10.57 10.98 10.10
CB196 9.22 9.25 10.31 10.44 9.60
CB197 9.36 9.93 10.72 11.00 8.50
CB205 11.25 10.68 10.68 10.70 9.20
Mean 8.04 9.06 10.08 10.55 9.00
Variance 7.00 4.04 0.27 0.57 1.91
Cattle not previously infected with BIV
CB203 9.98 10.28 10.73 11.18 16.40
CB206 9.40 10.01 10.18 10.55 10.30
CB208 7.54 8.95 10.02 10.22 10.70
CB210 7.46 9.16 10.13 10.60 13.40
Mean 8.59 9.60 10.26 10.64 12.53
Variance 1.65 0.41 0.10 0.15 7.77
P-valuea 0.624 0.692 0.931 0.404 0.068
a P-values represent statistical differences between groups of animals infected with
BIV 42 days prior to infection with JDV and those infected with JDV only.
99
A D
B E
C F
-1 2 6 10 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after JDV inoculation
Abs
orba
nce
(405
nm
)
-1 4 6 10 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after JDV inoculation
Abs
orba
nce
(405
nm
)
-1 2 6 10 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after mock JDV inoculation
Abs
orba
nce
(405
nm
)
-1 6 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after mock JDV inoculation
Abs
orba
nce
(405
nm
)-1 2 6 10 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after JDV inoculation
Abs
orba
nce
(405
nm
)
-1 6 15
0.0
0.5
1.0
1.5
2.0
2.5
Days after mock JDV inoculation
Abs
orba
nce
(405
nm
)
Figure 5.4. IgG response detected by JDV TMc peptide and His-BIV CA after
inoculation with JDV, shown as box and whisker plots of the ELISA absorbance
values. (A) Response detected by JDV TMc peptide in cattle infected with JDV at
day 0 and infected with BIV 42 days earlier. (B) Response detected by JDV TMc
peptide in cattle infected with JDV at day 0 but not infected previously with BIV.
(C) Response detected by JDV TMc peptide in cattle not infected with JDV at day 0
but infected with BIV 42 days earlier. (D) Response detected by His-BIV CA in
cattle infected with JDV at day 0 and infected with BIV 42 days earlier.
(E) Response detected by His-BIV CA in cattle infected with JDV at day 0 only.
(F) Response detected by His-BIV CA in cattle not infected with JDV at day 0 but
infected with BIV 42 days earlier.
100
Discussion
Under the conditions with which the experiment was conducted, prior BIV infection
did not prevent subsequent JDV infection or result in significant amelioration of the
normal response of Bali cattle to JDV infection. Although it did not alter the normal
clinical response to JDV infection, prior infection of cattle with BIV altered the
dynamics of their response to JDV and resulted in an earlier onset of Jembrana
disease. Although many of the effects were not significantly different, cattle
previously infected with BIV developed an earlier onset of fever, an earlier peak
febrile response, a significantly earlier peak VL and a significantly earlier resolution
of fever following superinfection with JDV. Although prior BIV infection did not
cause a difference in the total AUC, there was (in cattle previously infected with
BIV) a reduction in the duration of the viraemia that exceeded 106 genome copies/ml
of plasma. The differences were only evident when 2 atypical responders, identified
in a small percentage of all cattle infected with the JDVTab87 strain of JDV (Desport et
al., 2009a), were removed from the analysis. The lack of statistical significance in
those animals that responded typically was probably at least partly due to the wide
variance in the response to JDV of the cattle previously infected with BIV. A case
fatality rate of about 21% is normally evident in experimentally JDV-infected Bali
cattle (Desport et al., 2009a) but no fatalities occurred in any of the JDV-infected
cattle during the 14 day observation period after infection so no effect on the JDV-
associated case fatality rate of prior BIV infection was determined.
The enhanced early replication of a superinfecting virus seen here has not been
reported previously in superinfections with other related lentiviruses. An earlier
onset of disease has, however, been reported with CAEV, JDV, EIAV and FIV
following vaccination. Goats vaccinated with inactivated CAEV developed arthritis
101
more rapidly after CAEV infection than control animals (McGuire et al., 1986).
Cattle vaccinated with a tissue-derived JDV vaccine had earlier peak VL than control
animals (Ditcham et al., 2009). Horses vaccinated with a recombinant EIAV subunit
vaccine and then challenged with EIAV displayed severe enhancement of viral
infection and exacerbation of disease (Montelaro et al., 1996). Goats vaccinated with
a T-cell priming Gag peptide from CAEV had transiently enhanced virus replication
after CAEV infection compared with control animals, potentially via T-cell
enhancement of virus replication (Nenci et al., 2007) and similar findings have been
reported in FIV vaccine studies (Richardson et al., 1997). The presence of an active,
cross-reactive T-helper cell immune response may explain the earlier start of
viraemia and accelerated febrile response. JDV has recently been identified as
replicating in IgG-containing cells (Desport et al., 2009b) and alternatively, the
earlier replication of JDV in BIV-infected cattle may be related to the B-cell
stimulatory activity of BIV (Whetstone et al., 1997).
The lack of amelioration of the febrile response and the replication of JDV in the
cattle previously infected with BIV occurred despite a strong antibody response to
the BIV TM and, in a proportion of the cattle, an antibody response to the CA at the
time of JDV inoculation. Due to the close antigenic relationship between JDV and
BIV (Desport et al., 2005; Kertayadnya et al., 1993) the result was not expected as
other lentiviruses have been shown to offer protection against infection with closely
related heterologous viruses. Domestic cats infected with non-pathogenic puma or
lion lentiviruses developed humoral and cell-mediated immune responses against
both homologous and heterologous FIV isolated from domestic cats (VandeWoude et
al., 2003), suppressed FIV-induced CD4+ T-cell depletion (Terwee et al., 2008;
VandeWoude et al., 2002) and FIV-induced plasma and PBMC viral loads
102
(VandeWoude et al., 2002). Domestic cats pre-infected with a chimeric FIV and
subsequently infected with fully virulent FIV ameliorated the clinical effects of the
virulent challenge virus in some challenged cats, and reduced viral RNA and proviral
DNA loads in others (Giannecchini et al., 2007). A lack of neutralising antibody
response against the TM glycoprotein could explain why JDV infection occurred
despite a strong anti-TM antibody response.
It is possible that the R29 strain of BIV may have become attenuated since its
isolation in 1969 (Whetstone et al., 1997) and that this may have affected the result
obtained. It is also possible that the 42 day period between BIV infection and
subsequent JDV infection was too short for an effective protective immunity to
develop. Although there was a strong antibody response to the TM protein of BIV at
the time of JDV infection, and a rapid antibody response to TM and CA after JDV
infection, which is normally delayed until at least 6 weeks after infection
(Hartaningsih et al., 1994), this response may be unrelated to the development of a
protective immune response which may require a longer period to develop with the
bovine lentiviruses. However, in a similar experiment, protection against SIV in
macaques could be achieved within 21 days of infection with a live attenuated SIV,
with partial protection against wild-type SIV provided within 10 days of inoculation
(Stebbings et al., 2004). In contrast, after wild-type FIV infection there was no
protection against the effects of heterologous virus challenge until 2 to 3 years after
infection when animals exhibited a reduced virus load and sometimes a reduced
decline of CD4+ T-cells (Pistello et al., 1999). The possibility that longer term
infections with BIV may induce a protective response to JDV infection needs further
investigation as it would perhaps offer a low cost means of immunizing cattle against
Jembrana disease.
103
The dose of BIV used for infection and the challenge dose of JDV may also be
factors responsible for lack of resistance to JDV infection following BIV infection.
Studies of SIVmac in macaques have shown that the dose of primary virus affects the
level of resistance to infection with a pathogenic virus (Cranage et al., 1998).
Macaques given higher doses, 2 000 - 20 000 TCID50, of SIVmacC8 were shown to
completely resist infection with SIVmac, shown by a lack of virus detection by PCR
and a lack of virus isolation, whereas animals given lower doses of primary virus, 2 –
200 TCID50, were protected against a loss of CD4 cells only.
While there was no evidence that prior BIV infection provided protection against the
pathological effects of subsequent JDV infection, there was also no evidence that the
prior BIV infection exacerbated subsequent JDV infection, as reported in cases of
HIV superinfection. Epidemiological observations suggest that infection with a
second heterologous strain of HIV-1 has in the majority of cases, accelerated disease
progression after infection, reviewed previously (Smith et al., 2005). It was observed
in the current experiments that JDV infection in cattle previously infected with BIV
reactivated replication of the BIV, and this appears to be an observation seen during
heterologous lentivirus infections. Reactivation of primary infection virus has also
been seen in HIV-2 infected macaques superinfected with SIVmac (Petry et al., 1995)
and in HIV-2 infected baboons superinfected with heterologous HIV-2 (Locher et al.,
1997). Reactivation might be associated with the potent transactivation function of
the JDV Tat protein, which was shown to be a potent transactivator not only of its
own LTR but also the BIV and HIV LTR in vitro (Chen et al., 2000; Chen et al.,
1999). JDV Tat may function similarly in vivo in Bali cattle.
There is no evidence of the outcome of mixed infections with JDV and non-
pathogenic BIV-like viruses under field situations in the Bali cattle population of
104
Indonesia. Although mixed infection with JDV and a second BIV-like virus has been
reported to occur in Bali cattle on Bali island (Barboni et al., 2001), the difficulty of
detecting BIV-like virus in cattle and the close antigenic relationship of the 2 viruses
has so far precluded the epidemiological study of mixed infection by JDV and
possible non-pathogenic BIV-like viruses. BIV proviral DNA is only transiently
present within PBMC of experimentally infected Bali cattle (Chapter 4), making the
virus difficult to detect even when the animals are known to be infected.
105
Chapter 6: Humoral immune responses to Jembrana disease virus
detected using overlapping synthetic peptides spanning the MA, CA
and SU regions of JDV
Summary
The mapping of linear B-cell epitopes on the MA, CA and SU regions of Jembrana
disease virus is described. One hundred and fifty five overlapping peptides that
spanned these regions were synthesised and used in an ELISA format to screen a
panel of bovine sera from animals experimentally infected with JDVTab87, JDVPul01 or
BIV-R29. Six immunoreactive (IR) peptides, representing 6 potential epitopes, were
identified when the set of peptides was screened with sera taken following JDVTab87
infections; 1 in MA, 1 in CA and 5 in SU. Numerous IR peptides were identified
when the set of peptides was screened with JDVPul01 sera. BIV-R29 sera also reacted
with many peptides, including the IR peptides identified with the JDVTab87 sera.
However, BIV-R29 sera did not react with some peptides and a combination of these
peptides would enable detection of JDV-only seropositive cattle. These peptides
include MA18, MA19, SU93, SU95, SU103, SU119 and SU135.
106
Introduction
The occurrence of clinical Jembrana disease and serological surveys for antibody
reactive to JDV indicate that JDV infection is widespread in some islands of
Indonesia including Bali, Java, Sumatra and Kalimantan (Indonesian Borneo) but the
absence of clinical Jembrana disease and the occurrence of antibody to JDV suggests
the occurrence of a non-pathogenic bovine lentivirus, possibly related to BIV, on the
island of Sulawesi (Hartaningsih, personal communication). A BIV-like, non-
pathogenic bovine lentivirus was also reported to occur in the cattle population on
Bali island (Barboni et al., 2001) but this has not been confirmed and efforts to detect
BIV proviral DNA in PBMC of cattle on Bali island were unsuccessful (Chapter 3).
While JDV and BIV are sufficiently different genetically that they can be
differentiated utilising a number of PCR assays (Lew et al., 2004; Lewis et al., 2009)
low proviral DNA loads in PBMC after infection precludes their use as a screening
tool. They are very similar antigenically and attempts to differentiate antibody to the
2 viruses using ELISA and WIB procedures have been unsuccessful due to the
presence of numerous cross-reactive epitopes on the CA, MA and TM proteins
(Desport et al., 2005). Using recombinant, overlapping JDV Gag proteins, previous
attempts have been made to define specific epitopes that differentiate between the 2
bovine lentiviruses antigenically, however these were unsuccessful (Desport et al.,
2005). A number of recombinant proteins were produced and reacted with JDV and
BIV hyperimmune sera as well as monoclonal antibodies in a WIB. These attempts
identified at least three epitopic domains within MA and CA, including the Major
Homology Region (MHR) but could not identify an epitope(s) that differentiated
between the two viruses (Desport et al., 2005). A monoclonal antibody directed
against an epitope which spans the cleavage site between BIV MA and the p2L
107
protein was reported to differentiate between BIV and JDV antibody using a WIB
assay (Lu et al., 2002; Zheng et al., 2001) but the epitope involved does not appear to
be immunogenic in cattle and linear forms of this epitope can not be used to
differentiate between bovine lentivirus infections (Desport et al., 2005). Reliable
serological surveillance for JDV infection in cattle in Indonesia requires the use of
serological techniques that can differentiate the 2 infections.
Various epitope mapping studies have employed the use of expression libraries and
recombinant proteins to map epitopes and then fine mapping using synthetic peptide
strategies (Bertoni et al., 1994; Chong et al., 1991a; Chong et al., 1991b; Rosati et
al., 1999). These approaches were used to map the antigenicity of the SU
glycoproteins of EIAV (Ball et al., 1992) and CAEV (Valas et al., 2000) and the
antigenicity of CAEV TM (Bertoni et al., 1994). In this Chapter, synthetic peptides
were used to map the epitopes of JDV MA, CA and SU proteins to find epitopes
which can be used for differentiating the 2 viruses in serological assays.
Materials and methods
Source of animal sera
A panel of bovine sera from experimentally infected Bali cattle was used in this
study. The cattle were infected with either JDVTab87, JDVPul01 or BIV-R29 during
various studies of the response of cattle to these viruses (Desport et al., 2009;
Ditcham et al., 2009; Chapters 4 and 5) and serum samples were acquired at various
dpi. Pre-infection serum samples from the same cattle were used to test the
background level of reactivity to each peptide.
Hyperimmune JDV and BIV sera, a gift from N. Hartaningsih (Indonesia) were also
tested. JDV hyperimmune serum was created by inoculating a bovine lentivirus-free
108
animal with plasma taken from an animal during the febrile phase after infection with
JDVTab87. The animal then received 7 booster inoculations intramuscularly at 2 week
internals with tissue-derived JDV vaccine. Serum was taken 2 weeks after the final
inoculation. BIV hyperimmune serum was produced in a bovine lentivirus-free
animal with cell-culture preparations of BIV-R29 prepared as described in Chapter 4.
This cell culture material was emulsified in Freund’s incomplete adjuvant and
inoculated 3 times at 2-week intervals and serum was collected 2 weeks after the
final inoculation. The first inoculation was from freshly harvested cell culture
material while the final 2 inoculations were from frozen cell culture material. The
reactivity of the JDV hyperimmune serum has previously been reported (Desport et
al., 2005). The BIV hyperimmune serum reacted with JDV CA and BIV CA and SU
proteins on a WIB.
109
Overlapping synthetic peptides
One hundred and fifty five overlapping synthetic peptides were constructed to cover
the entire amino acid sequence of MA, CA and SU of JDVTab87 (accession number
U21603). The peptides were designed to be 16 amino acids long and to overlap each
other by 11 amino acids. This length allows for coverage of the three regions while
the minimal overlap enables mapping to a fine specificity and takes into account the
significant expense of peptide production. The peptides were synthesised by
automated 9-fluorenylmethyloxycarbonyl chemistry (Mimotopes, Australia) as
previously described (Valas et al., 2000). An amino-terminal biotinylated
tetrapeptide (Ser-Gly-Ser-Gly) was added to all peptides to facilitate epitope
accessibility and absorption to streptavidin-coated wells. All peptides were dissolved
in 200 µl of cell culture grade, 100% DMSO (Sigma-Aldrich). For use in the ELISA,
peptides were further diluted 1:200 in PBS-T (PBS containing 0.1% Tween-20
[Sigma] and 0.1% sodium azide [Sigma]).
ELISA
Serum samples were tested at a dilution of 1:50 against each peptide in a standard
ELISA according to the peptide manufacturer’s instructions. Plates were coated with
100 µl of 5 µg/ml stock streptavidin (Sigma-Aldrich) and left to evaporate overnight
at 37°C. Plates were blocked for 2 h with 200 µl of blocking solution (0.01 M PBS
containing 2.5% gelatin [Bio-Rad], 5% rabbit serum [Invitrogen], 1% sodium
caseinate [MP Biomedicals], 0.1% Tween 20 [Sigma]). All incubation steps were
conducted at room temperature (~25°C) on a rocker platform. Plates were then
washed 4 times with PBS-T. Peptides were further diluted 1:4 in PBS-T and 100 µl
of peptide solution were added to each ELISA well, in duplicate, and incubated for
1 h. Plates were again washed 4 times. Serum was diluted 1:50 using PBS-T and
110
100 µl added to each well. Plates were incubated for 1 h then washed a further 4
times. Goat anti-bovine IgG HRP-conjugated (MP Biomedicals) was diluted 1:2 000
in 0.01 M PBS with additional 1% sheep serum (Invitrogen), 0.1% Tween 20 and
0.1% sodium caseinate, 100 µl was added to each well. Plates were incubated for 1 h
then washed 4 times with PBS-T and 2 times with PBS only. Substrate (SigmaFast
OPD; Sigma) was prepared as per the manufacturer’s instructions and 100 µl was
added to each well. After colour development, the reactions were stopped with 3 M
H2SO4 and read at 450 nm.
Statistical analysis
A model previously developed and described to map linear epitopes in HIV-1 was
used to analyse the data produced in the ELISA (Loomis-Price et al., 1997). The cut-
off for reactivity of individual peptides was determined as follows: the median and
first quartile values (of the optical densities) were determined separately for each
serum sample and for each block of peptides. The data were normalised by
subtracting the median reactivity of the set from each value in the set and the
standard deviation was calculated:
Standard deviation (σ) = (median – first quartile)/0.675
The data were then divided by the calculated standard deviation and expressed as
normalised reactivity (σ) compared to the median. Measurements above a cut-off of
5 σ were considered positive.
The overlapping peptide set was screened with pre-infection sera to determine the
level of background binding present for each serum sample. Unless the level of
reactivity increased over time, the peptide was scored as non-reactive.
Immunoreactive (IR) peptides were defined as those which reacted with 75% or
111
greater of the serum samples tested and immunodominant (ID) peptides defined as
those which reacted with 100% of the serum samples tested, as previously reported
(Ball et al., 1992).
Results
When the overlapping peptides were tested against sera from cattle after infection
with JDVTab87, a number of peptides were not reactive against any of the sera tested,
including peptide MA18 in MA (Figure 6.1A), peptides CA34, CA35, CA42 and
CA62 in CA (Figure 6.2A) and peptides SU78, SU85, SU92, SU121, SU138,
SU145, SU148 and SU149 in SU (Figure 6.3A). One peptide (MA24) was identified
as IR in MA (Figure 6.1A), one peptide (CA70) was identified as IR in CA
(Figure 6.2A) and 4 peptides (SU112, SU152, SU154 and SU155) were as IR in SU
(Figure 6.3A). The locations of these peptides on the linear amino acid sequence of
JDVTab87 are shown in Figures 6.5 and 6.6. No peptides were identified as ID in MA,
CA or SU using the Ball et al. (1992) definitions. Using the less stringent definitions
reported in a separate study (Valas et al., 2000), whereby IR peptides were defined
by greater than 20% reactivity with bovine sera and ID peptides defined by greater
than 58% reactivity, 22 MA peptides were IR and 1 was ID, 27 CA peptides were IR
and 4 were ID, and 44 SU peptides were IR and 8 were ID.
When the overlapping peptides were screened with serum taken from cattle
experimentally infected with JDVPul01, a number of peptides were not reactive
against the JDVPul01 sera, including peptides MA6, MA10, MA13, MA14 and MA22
in MA (Figure 6.1B), peptides CA26, CA27, CA38, CA39, CA43, CA44, CA55,
CA62, CA63, CA67, CA68 and CA70 in CA (Figure 6.2B) and peptides SU78-80,
SU83, SU84, SU86- 88, SU90, SU94, SU95, SU111, SU112, SU114, SU116,
SU131, SU135, SU136, SU147 and SU154 in SU (Figure 6.3B). Numerous peptides
112
were IR against the sera from JDVPul01 infected animals including peptides MA4,
MA15, MA21 and MA25 in MA (Figure 6.1B), peptides CA41, CA49, CA52 and
CA69 in CA (Figure 6.2B) and peptides SU100, SU106, SU108, SU110, SU115,
SU118, SU120, SU121, SU126, SU145 and SU150 in SU (Figure 6.3B). Peptides
CA53, SU96 and SU102 were ID.
When the overlapping peptides were screened with serum taken from cattle
experimentally infected with BIV, peptides not reactive with the BIV sera tested
included peptide MA10, MA12 and MA18 in MA (Figure 6.1C), CA33, CA42 and
CA63 in CA (Figure 6.2C) and peptides SU74, SU83, SU85, SU86, SU88, SU92,
SU93, SU95, SU103, SU105, SU111, SU114, SU119, SU135 and SU137 in SU
(Figure 6.3C). Peptide 25 in MA (Figure 6.1C), peptides CA29, CA53 and CA55 in
CA (Figure 6.2C) and peptides SU126, SU129, SU100, SU118 and SU123 in SU
(Figure 6.3C) were all IR and no peptides were ID against these sera.
The 2 hyperimmune serum samples tested reacted against different peptides.
JDVTab87 hyperimmune serum reacted particularly strongly against peptide SU83 as
well as peptide MA5 (Figure 6.4). In contrast, BIV hyperimmune serum reacted
strongly to peptides in 3 regions of CA: CA57 and 58, peptides CA44 and CA45, and
CA65 (Figure 6.4).
113
1 5 9 13 17 21 25
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
1 5 9 13 17 21 25
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
1 5 9 13 17 21 25
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
A
B
C
Figure 6.1. Reactivity to peptides derived from the MA sequence of JDVTab87 to
serum samples taken (A) 56-159 days after infection of cattle with JDVTab87, (B)
71-102 days after infection of cattle with JDVPul01 and (C) 35-101 days after infection
of cattle with BIV.
114
26 31 36 41 46 51 56 61 66 71
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
25 30 35 40 45 50 55 60 65 70
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
25 30 35 40 45 50 55 60 65 70
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
A
B
C
Figure 6.2. Reactivity to peptides derived from the CA sequence of JDVTab87 to
serum samples taken (A) 56-159 days after infection of cattle with JDVTab87, (B) 56-
159 days after infection of cattle with JDVTab87 and (C) 35-101 days after infection of
cattle with BIV.
115
72 82 92 102 112 122 132 142
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
72 82 92 102 112 122 132 142
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
72 82 92 102 112 122 132 142
0
25
50
75
100
Peptide number
Rea
ctiv
ity
( σσ σσ)
A
B
C
Figure 6.3. Reactivity to peptides derived from the SU sequence of JDVTab87 to
serum samples taken (A) 56-159 days after infection of cattle with JDVTab87, (B) 71-
102 days after infection of cattle with JDVPul01 and (C) 35-101 days after infection of
cattle with BIV.
116
10 20 30 40 50 60 70 80 90 100 110 120 130 140 150
-5
0
5
10
15
20
25
Peptide number
Rea
ctiv
ity
( σσ σσ)
10 20 30 40 50 60 70 80 90 100 110 120 130 140 150
-5
0
5
10
15
Peptide number
Rea
ctiv
ity
( σσ σσ)
A
B
Figure 6.4. Reactivity of hyperimmune serum to overlapping peptides spanning the
MA (peptides MA1-25), CA (peptides CA26-71) and SU (peptides SU72-155) of
JDV. A: reactivity with JDVTab87 hyperimmune sera. B: reactivity with BIV-R29
hyperimmune sera. Connecting lines are shown for clarity and are not meant to imply
continuous data. Horizontal line (-----) indicates the cut-off of 5 σ.
117
JDV MKLSKLEKALKKVRVTPQRDDTYTIGNVLWAIRMCRLMGLDCCIDEAT-AAEVAILIGRF 59
Start gag ►
BIV MKRRELEKKLRKVRVTPQQDKYYTIGNLQWAIRMINLMGIKCVCDEECSAAEVALIITQF 60
** :*** *:*******:*. *****: ***** .***:.* ** *****::* :*
JDV QSLDLQDSPLKGKDEKAILTTLKVLWSLLAGHHPENSDMAEKYWEAWTIRERESQKEEEG 119
Peptide 24
BIV SALDLENSPIRGKEEVAIKNTLKVFWSLLAGYKPESTETALGYWEAFTYREREARADKEG 120
.:***::**::**:* ** .****:******::**.:: * ****:* ****:: ::**
JDV EITSIYPQLRKN---------------FPAVSTSDGSPRYDPDLTKQLKIWADATEKHGV 164
MA♦♦♦♦CA
Peptide 24
BIV EIKSIYPSLTQNTQNKKQTSNQTNTQSLPAITTQDGTPRFDPDLMKQLKIWSDATERNGV 180
**.****.* :* :**::*.**:**:**** ******:****::**
JDV DHHAVNILGVITANLTQSEIRLLLQSTPQWRLDIQLIESKLNAREHAHRVWKESHPEAPK 224
BIV DLHAVNILGVITANLVQEEIKLLLNSTPKWRLDVQLIESKVREKENAHRTWKQHHPEAPK 240
* *************.*.**:***:***:****:******:. :*:***.**: ******
JDV TDEIIGKGLTAAEQATLTTQECRDTYRQWVLEAALEVAQGKHDRPGPINIHQGPKEPYPE 284
BIV TDEIIGKGLSSAEQATLISVECRETFRQWVLQAAMEVAQAKHATPGPINIHQGPKEPYTD 300
*********::****** : ***:*:*****:**:****.** **************.:
JDV FVNKLVTALEGMAAPETTKQYLLDHLSVDHANEDCRAVLLPLGPSAPMERKLEACRAVGS 344
BIV FINRLVAALEGMAAPETTKEYLLQHLSIDHANEDCQSILRPLGPNTPMEKKLEACRVVGS 360
*:*:**:************:***:***:*******:::* ****.:***:******.***
JDV SKQKMQFLAEAFAAINVK-------------------GDGEVQRCYGCGKPGHIRRDCKN 385
CA♦♦♦♦NC
Peptide 70
BIV QKSKMQFLVAAMKEMGIQSPIPAVLPHTPEAYASQTSGPEDGRRCYGCGKTGHLKRNCKQ 420
.*.*****. *: :.:: * : :*******.**::*:**:
JDV QKCFKCGKPGHLQRNCKSKNGRRSSAPSGQRS----GYHQEKTS-VTPSAPPLVLD 436
◄ End gag
BIV QKCYHCGKPGHQARNCRSKNGKCSSAPYGQRSQPQNNFHQSNMSSVTPSAPPLILD 476
***::****** ***:****: **** **** .:**.: * ********:**
Figure 6.5. Linear representation of JDV MA and CA epitopes. A panel of sera taken from animals experimentally infected with JDVTab87 were reacted in ELISA against the panel of JDV MA and CA synthetic peptides. Bovine humoral epitopes are portrayed along the linear amino acid sequence of the JDVTab87 gag precursor protein. Grey highlighted sequences delineate immunoreactive peptides which are recognized by 75% or more of the bovine sera. The BIV gag precursor protein sequence is also shown below the JDV sequence for comparison. The sequences were aligned using ClustalW2 (Larkin, 2007). “*” indicates identical residues, “:” indicates conserved substitutions and “.” indicates semi-conserved substitutions.
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JDV MMEEGRKEEPEERGEKSTMRDLLQRAVDKGHLTAREALDRWTLEDHGEIHPWIILFCFAG 60
Start env ►
BIV MDQDLDGAERGERGGGS--EELLQEEINEGRLTAREALQTWINN--GEIHPWVLAGMLSM 56
* :: * *** * .:***. :::*:*******: * : ******:: ::
JDV AIGVIGGWGLRGELNVCMLIVLVVLVPIYWGIGEAARNIDSLDWKWIRKVFIVIIFVLVG 120
BIV GVGML--LGVYCQLPDTLIWILMFQLCLYWGLGETSRELDKDSWQWVRSVFIIAILGTLT 114
.:*:: *: :* :: :*:. : :***:**::*::*. .*:*:*.***: *: :
JDV LLG--------------------------------------------------------- 123
BIV MAGTALADDDQSTLIPNITKIPTKDTEPGCTYPWILILLILAFILGILGIILVLRRSNSE 174
: *
JDV ------------------------------------------------------------
BIV DILAARDTIDWWLSANQEIPPKFAFPIILISSPLAGIIGYYVMERHLEIFKKGCQICGSL 234
JDV ----------------------------------------GCSAQRQHVAMLLSPPGIRL 143
BIV SSMWGMLLEEIGRWLARREWNVSRVMVILLISFSWGMYVNRVNASGSHVAMVTSPPGYRI 294
.*. .****: **** *:
JDV P-STVDIPWFCISNAPIPDCVHWTVQK---PDQKHQQIENVMELQEVLDNATFFEVPDLF 199
BIV VNDTSQAPWYCFSSAPIPTCSSSQWGDKYFEEKINETLVKQVYEQAAKHSRATWIEPDLL 354
.* : **:*:*.**** * . :: :: : : : * . .. : : ***:
JDV DRVYLELARLDANSTGVPVNIPPTGISQVKGDCSTGDIQGMNETLSTRGTLGERTFLSIR 259
Peptide 112
BIV EEAVYELALLSANDS-----------RQVVVENGTDVCSSQNSSTNKGHPMTLLKLRGQV 403
:.. *** *.**.: ** : .*. .. *.: .. .: .: .
JDV PGGWFTNTTVWFCVHWPFGFIQRKEN-----LSEGSAQVRNCLDPINVTEPRVANYSYCP 314
Peptide 134
BIV SETWIGNSSLQFCVQWPYVLVGLNNSDSNISFNSGDWIATNCMHPITLNKS--------- 454
. *: *::: ***:**: :: ::. :..*. . **:.**.:.:.
JDV LEYKGKNYINKGLKCVGGRVDLSSNPEQHTDLLACGTFCQNFRNCDMVSRDILIG-YHPS 373
Peptide 134
BIV AQDLGKNFP--RLTFLDGQLSQLKN-----TLCGHNTNCLKFGNKSFSTNSLILCQDNPI 507
: ***: *. :.*::. .* * . .* * :* * .: :..::: :*
JDV QQKQHIYINHTFWEQANTQWILVQVPNYGFVPVPDTERPWKGGKPRGKRAVGMVIFLLVL 433
SU♦TM
Peptide 152 Peptide 154 Peptide 155
BIV GNDTFYSLSHSFSKQASARWILVKVPSYGFVVVNDTDTPP-SLRIRKPRAVGLAIFLLVL 566
:. . :.*:* :**.::****:**.**** * **: * . : * ****:.******
Figure 6.6. Linear representation of JDV SU epitopes. A panel of sera taken from animals experimentally infected with JDVTab87 were reacted in ELISA against the panel of JDV SU synthetic peptides. Bovine humoral epitopes are portrayed along the linear amino acid sequence of the JDV Env precursor protein. Grey highlighted sequences delineate immunoreactive peptides which are recognized by 75% or more of the bovine sera. Three immunoreactive peptides are shown at the carboxyl end of the protein; these are distinguished by the grey highlighted text (1), the underlined text (2) and the bolded text (3). The BIV Env precursor protein sequence is also shown below the JDV sequence for comparison. The sequences were aligned using
119
ClustalW2 (Larkin, 2007). “*” indicates identical residues, “:” indicates conserved substitutions and “.” indicates semi-conserved substitutions.
Discussion
The investigations conducted were designed to use synthetic peptides to identify
antigenic sites on the MA, CA and SU that would react differentially to antibody to
JDV and BIV. Synthetic peptides have been used previously to map antigenic sites in
a number of lentiviruses, including EIAV (Ball et al., 1992; Grund et al., 1996;
Soutullo et al., 2007), the SRLV CAEV and VMV (Mordasini et al., 2006; Rosati et
al., 1999; Valas et al., 2000) and HIV-1 (Loomis-Price et al., 1997; Neurath et al.,
1990). They have also been used with other virus systems including human
cytomegalovirus (Greijer et al., 1999), Foot and mouth disease virus (Geysen et al.,
1987) and Epstein-Barr virus (Middeldorp et al., 1988). The peptides were able to
identify specific linear sites which were immunogenic in the native protein antigens.
It is recognised, however, that a limitation of these studies as well as the current
study is that the use of synthetic peptides cannot identify discontinuous
conformation-dependent epitopes which may represent important antigenic
determinants of viral proteins.
Another limitation of this current study was that it was necessary to use serum from
BIV-R29 infected animals to screen the peptides and examine for differences in the
reactions comparative to those in JDV-infected cattle. This BIV strain may not
necessarily have a close antigenic relationship to the putative non-pathogenic bovine
lentivirus in Indonesia. However, this was the only BIV serum available and sera
from antibody-positive cattle in Sulawesi, which would have been a suitable
alternative, were unavailable for this study.
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Sera from cattle experimentally infected with JDVTab87 or JDVPul01 were used to
screen the peptides for JDV-reactive epitopes as these are 2 reasonably well defined
strains that have been detected in Indonesia and the samples had a well documented
history. Of a number of JDV strains sequenced, JDVPul01 is the most divergent from
JDVTab87 in env and gag regions (Desport et al., 2007) and we sought to determine
whether the differences extended to the humoral response. The amino acid sequence
homology between the two strains in the entire env region and part of the gag regions
is 97% (Desport et al., 2007). Differences also existed between the reaction of cattle
infected with these 2 JDV strains in regards to peak viral loads and duration of
viraemia (Desport et al., 2009a). Differences were detected in the reactivity of the
peptides between the JDVTab87 and JDVPul01 sera. The JDVPul01 sera had a larger
number of IR peptides, with 3 ID peptides CA53, SU96 and SU102. These
differences could be attributed to the fewer number of cattle analysed and it would be
of interest to screen the peptides with more JDVPul01 sera to confirm these results.
The synthetic peptides used in this study encompassed the complete JDVTab87 MA,
CA and SU regions. The MA, CA and SU proteins were chosen for investigation for
a number of reasons. Firstly, the strongest and earliest immune responses against
JDV are directed at CA (Hartaningsih et al., 1994; Kertayadnya et al., 1993) although
a subset of cattle do not mount an immune response against CA (Desport et al.,
2009a; Ditcham et al., 2009). Matrix was chosen as BIV was reported to contain at
least one unique epitope in the CA that is absent in JDV, at the 6.4-kDa N terminus
of the 29-kDa CA adjacent to MA (Zheng et al., 2001) and strong antibody responses
against MA have been previously identified in experimentally infected animals
(Desport et al., 2005; Kertayadnya et al., 1993). SU was included as anti-Env
responses have been reported to persist beyond 190 weeks after BIV infection whilst
121
the Gag response wanes 40 weeks after infection (Isaacson et al., 1995), env
sequences between JDV strains are reasonable well conserved (Desport et al., 2007)
and there are a significant number of differences between the sequence of the SU
regions of JDV and BIV (Chadwick et al., 1995b, Figure 6.6, Table 2.2). Little
information is currently available detailing the antigenicity of JDV SU due to
problems expressing recombinant SU.
In the ELISA used to examine differential reactivity of the peptides, a cut-off of 75%
and greater positive reactivity to represent a significant B-cell epitope was used, as
reported by others (Ball et al., 1992). Other studies have utilised lower cut-offs of
33% (Kusk et al., 1992) and 50% (Valas et al., 2000) but a higher cut-off was used in
the current study to identify highly reactive B-cell epitopes in the JDV proteins.
There is potential for the use of IR peptides, those reacting with a high percentage of
serum samples tested, and ID peptides reacting with all serum samples, to facilitate
the development of a peptide-based ELISA able to identify most animals infected
with JDV or both JDV and BIV depending on the peptides used. Several potentially
useful IR peptides were detected.
One IR MA peptide was identified which spanned amino acids 116 to 131 of the CA
and MA proteins. This peptide spanned the MA-CA border and indicates that the
antibodies which recognise this peptide are produced in response to the uncleaved
Gag precursor protein.
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Sera from long-term JDV infections (>12 months after infection) were reported to
recognise a protein of the same size as MA (Kertayadnya et al., 1993) and these
findings identify an epitope within MA. Some JDV-infected cattle (CB83-86) have
been reported not to respond to MA within 175 dpi when tested in an ELISA using
recombinant MA (Ditcham et al., 2009) but MA24 reacted with sera from all of these
cattle (Table 6.1). The difference between the responses reported by Ditcham et al.
(2009) and those in the current study may be due to differences associated with the
sequences covered by the antigens. The responses reported by Ditcham et al. (2009)
were those generated when using a full length recombinant MA while MA24 spans
the MA-CA border, encompassing additional sequence. Peptide ELISA has been
shown to be more sensitive than an ELISA using recombinant proteins (Rosati et al.,
1999). Coincidentally, most variation in JDV Gag occurs just before the predicted
cleavage point between the MA and CA proteins (Desport et al., 2007) and this,
combined with the identification of an IR peptide in this region, suggests this may be
a region under pressure from the immune system.
The IR CA peptide reported here is within amino acids 346 – 360 of the Gag
precursor protein and spans the CA-NC border. Like the MA response, it also
suggests that the antibodies are produced in response to the uncleaved Gag precursor
polyprotein. Hyperimmune sera against whole JDV and BIV were reported to not
recognise recombinant proteins encompassing this region, presumable because of the
differences in the way hyperimmune sera is raised compared to sera from natural or
experimental infections (Desport et al., 2005). As shown in Figure 6.1, JDV
hyperimmune serum did react to this peptide while BIV hyperimmune sera did not.
A highly immunogenic domain has been reported in a similar location for EIAV
(Chong et al., 1991b).
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There were no IR peptides found within the MHR in CA when the peptides were
screened with JDVTab87 sera. The lack of an JDVTab87 IR peptide in this region is
surprising given that previous studies have shown that it is likely to be an epitopic
domain within JDV (Desport et al., 2005). The MHR is conserved among
retroviruses and is essential for virus assembly, maturation and infectivity
(Mammano et al., 1994) and is assumed to account for some of the cross-reactivity
observed between the Gag proteins of HIV, EIAV and BIV. Previous studies have
shown cross-reactivity between BIV hyperimmune sera and the JDV MHR (Desport
et al., 2005), although confirmation is required. Peptides CA52-59 spanned the MHR
and although they were not identified as IR in the current study, a number of peptides
had high reactivities against JDVTab87 sera, including CA53–55, 57 and CA58.
Peptide CA57 had the second highest reactivity of all the CA peptides with a
reactivity of 62.5%. The CA53 peptide was a JDVPul01 ID peptide and when screened
with BIV sera, peptides CA53 and CA55 were found to be IR, confirming that this is
one region of cross-reactivity. The BIV hyperimmune serum used in this study also
strongly reacted to this region (Figure 6.4). JDV hyperimmune serum also reacted to
the MHR, albeit more weakly than the BIV hyperimmune serum.
Surprisingly, the major homology region identified as an immunodominant domain
in the CA proteins of many of the lentiviruses was not identified as IR or ID using
this method of analysis.This may indicate that the level of stringency was too high or
may be due to the characteristics of the sera chosen or the conformation of the
peptides. Previous studies have shown that conformational changes to peptides, such
as converting the peptides to a cyclical form, can help to increase reactivity (Scobie
et al., 1999). Further studies are required to clarify this discrepancy.
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Five IR peptides were found within SU. Peptide SU112 spanned amino acids 201–
216 in the central portion of the protein. The most IR peptide, reactive with 87.5% of
the sera tested, was SU134 and this spanned amino acids 311–326. The remaining 3
peptides, SU152, SU154 and SU155, clustered around the carboxyl end of SU and
spanned amino acids 401–430. The reduction in reactivity for peptide SU153
compared with the reactivity for peptides SU152 and SU154 indicated that there may
be 2 different epitopes at this location, as reported in CAEV SU (Valas et al., 2000).
The clustering of IR peptides around the carboxyl end of SU is similar to CAEV
(Bertoni et al., 2000; Valas et al., 2000), HIV-1 (Palker et al., 1987) and EIAV (Ball
et al., 1992; Grund et al., 1996), although these viruses also have IR regions at the
amino end of SU which were not identified in this current study of JDV. FIV also has
an epitope at the carboxyl end of SU, progressing into the start of TM (Pancino et al.,
1993), similar to the region encompassed by peptides SU154 and SU155 in the
current study. It was suggested that the antigenicity of terminal segments of proteins,
including lentiviral SU glycoproteins, is because these regions are frequently surface
orientated and thus exposed and immunogenic in their native state (Ball et al., 1992;
Valas et al., 2000; Van Regenmortel, 1999b).
No ID peptides were identified with the JDVTab87 serum. This might be associated
with the outbred nature of the Bali cattle population, as has been suggested in the T-
cell epitope mapping in FIV (Dean et al., 2004). A number of ID peptides were
identified with the JDVPul01 serum including 1 in CA and 2 in SU (Figures 6.2 and
6.3). Reactivities with the JDVTab°/87 sera to these peptides ranged from 25 to 50%
(Figures 6.2 and 6.3). Further investigations are required to determine whether the
differences are attributed to strain variation or to the number of samples analysed.
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The differential reaction of some peptides to JDV sera compared to BIV sera
suggests some of these peptides could be used to form antigens for potential JDV-
specific and broadly reactive bovine lentivirus serological assays. Peptides which
were reactive against 50% of sera taken from JDVTab87, JDVPul01 and BIV-R29 would
be useful to include in a bovine lentivirus serological assay as these should detect a
majority of cattle infected with these bovine lentiviruses. The areas encompassed by
these peptides are significant regions of cross-reactivity since a majority of both JDV
and BIV sera reacted to these peptides.
Peptides that were not recognised by BIV sera are potential candidates for inclusion
in a JDV-specific ELISA antigen, an approach previously suggested in CAEV (Valas
et al., 2000). Unfortunately, none of these peptides were IR or ID against the JDV
sera only. However, a combination of 5 of these peptides, SU93, SU95, SU103,
SU119 and SU135 from SU, would have reacted with 87.5% of the JDV-only sera,
although this specific combination would not have detected animals infected with
JDVPul01. The addition of 2 extra peptides, MA18 and MA19 would provide
reactivity with the JDVPul01 sera. It is important to note, however, that this approach
would potentially provide a JDV-specific serological assay and not a BIV-specific
assay. Previous diagnostic assays have combined multiple antigens in an ELISA
format (Khan et al., 2006) and it would be practical to combine these peptides in this
format.
This study needs to be extended by testing sera from naturally infected cattle as well
as by testing the longitudinal responses to the IR peptides in experimentally infected
cattle, and applying the tests in a clinical setting in Indonesia. Consideration could
also be given to using selected peptides in the formulation of vaccines capable of
protecting against JDV, as has been reported in a number of lentivirus systems
126
including SIV (Belyakov et al., 2001; Nehete et al., 2008) and HIV-1 (Hovanessian
et al., 2004). Due to the persistent nature of the anti-TM antibody response in BIV
infections (Isaacson et al., 1995, Scobie et al., 1999), the TM glycoprotein may also
be a promising linear antigenic target and therefore extending this study to the TM
glycoprotein may yield a potential antigen for inclusion in a differential serological
assay. It would also be of interest to test the response of cattle superinfected with
both BIV and JDV and their pattern of reactivity over time.
127
Chapter 7: General discussion
The close genetic and antigenic relationship between BIV and JDV raised 2 issues
that were investigated and are reported in this thesis. First, in animals that are
infected with both viruses, it was hypothesised that there might be cross-protective
immunity or other interaction between the 2 viruses that could modify their
pathogenesis. Second, there is a need for serological tests that will differentiate
antibody to the pathogenic JDV and other non-pathogenic bovine lentiviruses. The
presence of a BIV-like virus in cattle on the island of Sulawesi where Jembrana
disease does not occur could have a marked effect on the events that might occur if
JDV spreads to that island. In an endemic area, previous infection with a non-
pathogenic bovine lentivirus like BIV might ameliorate the effect of subsequent
infection with JDV, resulting in subclinical JDV infections. Immunosurveillance for
JDV infection would be affected by the presence of a second antigenically cross-
reactive but non-pathogenic bovine lentivirus in the cattle population of Indonesia.
The investigations reported in this thesis have provided information that clarifies
these issues.
Evidence was provided previously for the occurrence of a non-pathogenic bovine
lentivirus in Bali cattle in Indonesia, although this was based on serological evidence
only and the virus has not been detected. This serological evidence, particularly that
presented by Barboni et al. (2001), is difficult to evaluate because other
investigations have demonstrated a very close antigenic relationship between JDV
and BIV and there are no reagents or tests available that allow differentiation of
antibody to the 2 types of virus in cattle sera (Desport et al, 2005). Antibody to JDV
was detected in B. javanicus in Sulawesi where there is no evidence of Jembrana
disease (Hartaningsih, personal communication) suggesting the presence of a non-
128
pathogenic bovine lentivirus antigenically related to JDV. Blood from a seropositive
animal in Sulawesi was inoculated into Bali cattle in an attempt to transmit the non-
pathogenic bovine lentivirus but these attempts were unsuccessful (Hartaningsih,
personal communication). For further investigation of the effects of the Indonesian
strain of this non-pathogenic bovine lentivirus in Bali cattle, an attempt was made to
detect the virus in cattle on the island of Bali (Chapter 3). A large number of cattle
were screened using PCR and serological assays which detect both BIV and JDV.
While a number of cattle were identified that contained proviral JDV DNA, BIV
proviral DNA was not detected and the investigations reported in Chapter 3 therefore
provide no evidence for the occurrence of a second non-pathogenic bovine lentivirus
in these cattle. Isolation of virus in cell culture was not attempted in this study due to
a lack of suitable facilities.
The lack of evidence of a BIV-like virus reported in Chapter 3, however, was
insufficient to eliminate the possibility that there is a second non-pathogenic bovine
lentivirus in the cattle population of Bali as only a limited number of samples from
one area were screened and the seroprevalence was lower than expected. Previous
investigators have concluded that BIV is difficult to detect in cattle. While numerous
studies have reported serological evidence for BIV infection (Barboni et al., 2001;
Bhatia et al., 2008; Horzinek et al., 1991; Meas et al., 1998; Meas et al., 2000a;
Whetstone et al., 1990) there have been only 3 reports of isolation of the virus from
cattle (Meas et al., 1998; Suarez et al., 1993; Van der Maaten et al., 1972). Suarez et
al. (1993) were only successful in the isolation of virus on 2 occasions from many
samples and only after 4 blind passages of samples in cell culture. A very exceptional
result was reported from Japan, where an unusually high isolation rate of BIV in cell
culture was reported from PBMC and milk-derived leukocytes from BIV-antibody-
129
positive dairy cattle. These isolates had 99.0 to 99.7% nucleotide sequence identity
with the R29 strain of BIV within a 258 nucleotide amplicon from the pol region.
These authors were also able to identify BIV proviral DNA by nested PCR and
Southern blot in cattle and buffalo samples from Japan, Cambodia and Pakistan
(Meas et al., 1998; Meas et al., 2000a; Meas et al., 2000b). The reports by Meas and
colleagues are interesting as they suggest that infection with BIV is widespread and
reasonably easy to detect in buffalo and cattle in Asian countries, and that virus
isolation was possible from antibody-positive cattle.
The study of BIV infection in Bali cattle reported in Chapter 4 is the first report of
the pathogenesis of BIV infection in this species. Because of the unusual
susceptibility of Bali cattle to JDV and Malignant catarrhal fever virus (Soesanto et
al., 1990) it was hypothesised that these cattle might also show a greater
susceptibility to BIV than do B. taurus but their susceptibility appeared generally
similar to the effects reported in B. taurus (Scobie et al., 1999; Suarez et al., 1993;
Whetstone et al., 1990). We also considered that BIV in Bali cattle might have very
different effects to those observed in B. taurus, as some lentiviruses do not cause
disease in their natural host (VandeWoude et al., 2006) but do so in heterologous
hosts, a phenomenon best described with SIV strains (Apetrei et al., 2004) but also
recognised with other lentiviruses such as puma and lion lentivirus infection in
domestic cats (VandeWoude et al., 1997). It has been reported that infection of B.
taurus with JDV resulted in a mild disease only, of much less severity that observed
in B. javanicus (Soeharsono et al., 1995a), and the reciprocal event, greater
susceptibility of Bali cattle to BIV was a potential outcome.
Bali cattle inoculated with BIV did not develop clinical signs within 62 days of
infection and this suggested that if the putative bovine lentivirus present in Sulawesi
130
is indeed related to BIV-R29, then it should have no significant effect on animal
health in the Bali cattle population. A result that is significant in terms of the optimal
method for detection of natural BIV infection in Bali cattle, and possibly B. taurus,
was that there was a transient period of viraemia detected after infection and it is
during this period that the virus would be most easily detected. However, in natural
infections, the timing of infection is unknown and therefore these results show that
the choice of assay is an important one. If a recent infection is suspected, then a
genome based assay would be most likely to detect infection but if a longer term
infection is suspected then a ELISA using a TM peptide would be more appropriate.
This transient period of viraemia was analogous to that which occurs in other animal
lentivirus infections such as JDV infection of Bali cattle (Stewart et al., 2005), EIAV
infection of horses (Harrold et al., 2000) and SIVsmm-PBj14 infection of pig-tailed
macaques (Dewhurst et al., 1990; Fultz et al., 1989), although BIV was found to have
significantly lower titres than these viruses. It is suggested that BIV infection of B.
taurus should be re-examined using sensitive techniques such as qRT-PCR to see
whether a similar acute phase viraemia is produced. Sensitive techniques need to be
used in such an investigation as the level of viraemia in Bali cattle infected with BIV
was low and virus was not detected consistently in any animal, reflecting the
difficulty of detecting BIV infection even in experimentally infected cattle. The
majority of infected cattle had a significant TM antibody response but a poor CA
response but this occurred after the transient viraemic period, suggesting that
detection of BIV proviral DNA would be most successful in antibody-negative cattle,
similar to the result obtained for JDV in field cattle (Chapter 3). The ability to detect
BIV prior to the onset of an antibody response but not afterwards is in contrast to the
131
report of the high frequency of isolation of BIV from antibody-positive cattle in Asia
(Meas et al., 1998).
A limitation of this current study of BIV-R29 in Bali cattle is that an Indonesian BIV
isolate was not discovered and could not be used for experimental studies, and the
BIV-R29 isolate used might not reflect what would occur with the putative BIV-like
virus present in Indonesia. However, other Asian isolates have been shown to be
closely related to BIV-R29 (Meas et al., 1998). The R29 strain of BIV may have a
different pathogenesis to the BIV-like virus present in Indonesian cattle. The results
obtained may also not be typical of BIV infection under field conditions as it is
considered by some that the BIV-R29 strain may have, since its isolation in 1969
(Van der Maaten et al., 1972), become attenuated during its prolonged storage and
passage in cell culture (Whetstone et al., 1997). However, an unsuccessful attempt
was made to increase the virulence of BIV-R29 by its serial passage through cattle
(Whetstone et al., 1990), a technique known to increase the virulence of EIAV
(Orrego et al., 1982), and perhaps this inability to increase its virulence reflects a
natural low pathogenicity. Other BIV strains isolated from Florida were thought to be
more pathogenic than BIV-R29, causing a B-cell lymphocytosis in experimentally
infected cattle, but the effects of the Florida isolates in cattle were still very mild
(Suarez et al., 1993; Whetstone et al., 1997). It is possible, however, that the
inoculation of the Florida isolates into Bali cattle may produce results different to
that observed with the R29 strain.
The lower antibody response to the CA versus the TM antigen was an unexpected
finding in BIV-infected Bali cattle. Animals infected with lentiviruses normally
develop a strong antibody response to CA and this response is the first to be detected
(Rosati et al., 2004; Soutullo et al., 2007). A strong response to CA has also been
132
documented in Bali cattle infected with JDV (Hartaningsih et al., 1994) although
there is a subset of JDV infected Bali cattle that do not develop a detectable CA
antibody response (Desport et al., 2009a; Ditcham et al., 2009). A loss of Gag-
reactivity in the presence of a sustained Env response has previously been reported in
B. taurus infected with BIV (Isaacson et al., 1995).
In several lentivirus systems, infection with a non-pathogenic lentivirus is able to
protect against subsequent superinfection with a closely related pathogenic lentivirus
(Cranage et al., 1998; VandeWoude et al., 2002). An investigation of whether the
same was true for BIV and JDV in Bali cattle showed that BIV-R29 infection did not
induce a protective immune response against subsequent infection 42 days later with
JDV. All BIV-infected animals inoculated with JDV became infected and had a
viraemic phase and a febrile response typical of JDV infection. However, while
previous BIV infection did not protect against subsequent JDV infection, there was
no enhancement of JDV infection even though it did cause an earlier onset and
resolution of fever that was associated with JDV infection. The result suggests that if
BIV-R29 is related to the BIV-like lentivirus circulating within the cattle population
of Sulawesi, if JDV were introduced into the cattle population of Sulawesi it would
not lead to disease enhancement.
The lack of any evidence of a protective immunity against JDV as a consequence of
previous BIV infection was unexpected, considering the extensive antigenic cross-
reactivity of JDV and BIV proteins and evidence of an antibody response to BIV at
the time of superinfection with JDV. It will be necessary to determine if protection
against JDV can be achieved by altering the parameters of the superinfection.
Perhaps if the dose of BIV used to infect the cattle was increased this would affect
the result: higher doses of SIVmacC8 provided greater protection against challenge
133
with SIVmac (Cranage et al., 1998). Alternatively, the time interval between BIV and
JDV infection could be important: greater protection has been provided by a
decreased time of 21 days between superinfection with closely related simian
lentiviruses (Stebbings et al., 2004) but it is also possible that a longer interval may
be required for the development of a cell-mediated immune response. Although
desirable, analysis of cell-mediated immune responses was not possible and so the
type of immune response required to control infection could not be adequately
assessed.
An important technical requirement for immunosurveillance of JDV infection in
Indonesia is the development of reliable and specific serological tests. JDV and BIV
contain an extensive array of cross-reactive epitopes on several proteins (Desport et
al., 2005; Kertayadnya et al., 1993) and using serological assays, differentiation of
infection by the 2 viruses using serological assays is currently not possible. The
investigation reported in Chapter 6 where a series of overlapping peptides spanning
the MA, CA and SU regions of JDV was used to identify epitopes that would react
specifically to JDV and not BIV has identified potential peptides that could be used
for this purpose. Seven peptides were identified within the MA-CA and the CA-NC
junctions, while 5 were identified in SU, that reacted with >75% of sera from JDV-
infected cattle. Unfortunately, these peptides also reacted with some sera from BIV-
infected cattle and could not therefore be used for the development of a JDV-specific
serological assay. A combination of these peptides could, however, be used as an
alternative antigen for the development of a broadly-reactive bovine lentivirus
serological test. A number of peptides were identified which reacted with sera from
JDV-infected cattle only and hence have potential for development of a JDV-specific
serological assay. These peptide combinations must now be tested using an extensive
134
array of sera from naturally infected cattle from areas of Indonesia where infection
with both BIV and JDV have been suspected, such as Bali, and in regions where
infection with a non-pathogenic BIV is suspected and where Jembrana disease has
not been detected, such as Sulawesi. A longitudinal series of serum samples from
individual cattle also needs to be tested to determine how the antibody response to
the peptides changes over time.
In conclusion, the results reported in this thesis have made a significant original
contribution to our understanding of BIV infection in Bali cattle that indicate BIV is
unlikely to be a significant pathogen in these cattle. The results obtained have also
provided insights into the interaction of BIV and JDV in Bali cattle which suggest
that previous BIV infection will not provide significant cross-protective immunity
against subsequent JDV infection, and that BIV infection is unlikely to provide a
simple method of vaccinating cattle against JDV infection. The results have also
provided information about epitopes of the Gag and SU proteins of JDV that indicate
there are potential epitopes on these proteins that could be used for the development
of JDV-specific serological tests needed in Indonesia.
135
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