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Immunogenetics of Small Ruminant Lentiviral Infections

Centre for the Genetic Improvement of Livestock, Department of Animal and Poultry Science, University of Guelph, Guelph, ON N1G 2W1, Canada
Department of Pathobiology, Ontario Veterinary College, University of Guelph, Guelph, ON N1G 2W1, Canada
Author to whom correspondence should be addressed.
Viruses 2014, 6(8), 3311-3333;
Original submission received: 23 May 2014 / Revised: 18 August 2014 / Accepted: 19 August 2014 / Published: 22 August 2014
(This article belongs to the Special Issue Small Ruminant Lentiviruses)


The small ruminant lentiviruses (SRLV) include the caprine arthritis encephalitis virus (CAEV) and the Maedi-Visna virus (MVV). Both of these viruses limit production and can be a major source of economic loss to producers. Little is known about how the immune system recognizes and responds to SRLVs, but due to similarities with the human immunodeficiency virus (HIV), HIV research can shed light on the possible immune mechanisms that control or lead to disease progression. This review will focus on the host immune response to HIV-1 and SRLV, and will discuss the possibility of breeding for enhanced SRLV disease resistance.

1. Introduction

The caprine arthritis encephalitis virus (CAEV) and the Maedi-Visna virus (MVV) are enveloped RNA viruses in the lentivirus genus of the Retroviridae family [1,2]. While small ruminant lentiviruses (SRLVs) were once considered to be species-specific, recent studies suggest that they can be transmitted between sheep and goats [3], and can recombine to form new CAEV-MVV strains [4]. These viruses primarily infect monocytes, macrophages, and dendritic cells [5], and like the human immunodeficiency virus (HIV), infection is lifelong and can persist for months or years in a latent or sub-clinical state [6]. When disease symptoms do emerge, goats predominantly show signs of arthritis or mastitis, while in sheep the disease tends to manifest as pneumonia or mastitis [7]. Encephalitis can also be a symptom in either lambs or kids but is less common [8].
SRLVs are predominantly vertically transmitted to offspring through the shedding of virus particles, and infected macrophages and epithelial cells in the colostrum and milk [9]. Horizontal transmission however, can occur through prolonged direct contact with bodily secretions, and sexual transmission of the virus may also be possible [10]. There is currently no effective treatment for SRLV infections and due to a high mutation rate, effective vaccine development has been and will continue to be challenging [11]. Therefore, the most effective means of controlling the virus is through herd management that prevents viral transmission.
The dynamics of the host immune response to SRLV infections remain unclear, but due to the similarities between SRLV and HIV, a great deal of our knowledge of the immune responses to HIV can be used to enhance our understanding of the host response to SRLV. This review will discuss the immune response to lentiviral infections, and the possibility of breeding for enhanced SRLV resistance will be addressed.

2. Lentiviral Characteristics

To fully understand the complex interaction between the host and virus, it is first necessary to understand the structural and genomic organization of the lentiviruses. Although an in-depth discussion of the structural and functional characteristics of lentiviruses is beyond the scope of this paper, a brief overview of SRLV and HIV-1 organization is given.

Genomic Organization

The SRLV genome is comprised of three structural genes, gag, pol and env, and three accessory genes, vif, tat, and rev (Figure 1) [12,13]. The gag gene encodes the capsid proteins, pol encodes the viral enzymes protease, reverse transcriptase, and integrase, and env encodes the envelope glycoproteins, gp135 (SU) and gp38 (TM) [14]. While the tat gene is dispensable for efficient viral replication [15], Vif is absolutely required for efficient in vivo virus replication and pathogenicity [16,17]. The flanking ends of the proviral DNA are regions of long terminal repeats (LTRs) that are divided into the U3, R, and U5 regions [18,19,20]. These regions provide the signals required for viral transcription and integration into the host genome [19]. Small ruminant lentiviruses differ from the primate lentiviruses in that their Tat proteins do not trans-activate the viral LTR promoters [21]. Rather, the SRLV Tat protein is functionally similar to the HIV type 1 (HIV-1) Vpr protein [22].
The HIV-1 genome is similar to the SRLV genome, but is more complex and contains additional genes [23]. In contrast to SRLV, HIV-1 contains nine genes encoding 15 proteins; these genes, include the structural gag, pol, and env genes that encode the capsid proteins, the viral enzymes, and the envelope glycoproteins, gp120 (SU) and gp41 (TM), respectively [24]. The HIV-1 non-structural genes consist of vif, tat, rev, nef, vpu, and vpr, and are associated with HIV-1 pathogenesis and immune evasion [25]. Like the SRLV genome, the flanking ends of the provirus contain LTRs consisting of U3, R, and U5 regions [26]. Additionally, a variety of promoter and enhancer elements have been identified in the LTR region of HIV-1. Some of these response elements include AP-1, NF-κB, and Sp1 [27], important host immune related transcriptional factors which all lead to transcriptional activation and virus replication.
Figure 1. Genomic organization of small ruminant lentiviruses (SRLV) and HIV-1 [23].
Figure 1. Genomic organization of small ruminant lentiviruses (SRLV) and HIV-1 [23].
Viruses 06 03311 g001

3. Host Immunity to Lentiviral Infections

There are substantial gaps in our knowledge of host innate and acquired immune responses to SRLV. Due to similarities between HIV-1 and SRLV, HIV-1 research can improve our understanding of SRLV immune responses. This section will provide an overview of the immune response to HIV-1, and discuss the current knowledge of the immune response to SRLV infection.

3.1. The Innate Immune Response to HIV-1

3.1.1. Toll-like Receptors and Antiviral Peptides

Toll like receptors (TLR) are host pattern recognition receptors (PRRs) that play a key role in innate recognition of a variety of conserved pathogen-associated molecular patterns (PAMPs). Ligation of PAMPS with PRRs induces intracellular signaling pathways that involve a series of phosphorylation events mediated through the common adaptor molecule, MyD88, reviewed by [28]. Downstream effects of these signaling pathways result in the activation and translocation of NF-κB or AP1 transcription factors to the nucleus, and subsequent pro-inflammatory cytokine production that facilitates the recruitment of innate effector cells such as neutrophils and macrophages to clear the pathogen [29].
During HIV-1 infection, the roles of TLR 7 and 8 have been investigated. Both TLR7 and 8 recognize single-stranded viral RNA (ssRNA), and this results in the induction of intracellular signaling cascades that involve NF-κB transcriptional activation, which ultimately results in the production of a variety of pro-inflammatory cytokines such as the type 1 interferons (IFN-α and -β), IL-6, and TNF-α [30,31]. Although this serves as an anti-viral defense mechanism through the induction of antiviral peptides such as APOBEC3G (A3G), tripartite motif 5-alpha (TRIM5α), and tetherin, persistent immune activation also induces viral replication through the activation of the NF-κB response element located within the HIV-1 LTR [27]. HIV-1 replication can also be induced through the activation of other TLR signaling pathways. For example, Ranjbar et al., [32], observed that HIV-1 replication can also be induced through activation of TLR2 during Mycobacterium tuberculosis infection. Since the recruitment of MyD88 and activation of either NF-κB or AP1 is conserved across TLRs 1, 2, 4, 5, 6, 7, 8, and 9 [28], recognition of a variety of bacterial and viral PAMPS may all contribute to HIV-1 transcriptional activation and disease progression.
Despite the induction of HIV-1 replication mediated through TLR signaling, there are host-adapted antiviral proteins that serve to control HIV-1 replication. Expression of these antiviral peptides is upregulated by type 1 interferons that are induced in response to TLR activation [33]. TRIM5α, for example, is a lentivirus restriction protein that mediates cellular restriction against retroviruses in a species-specific manner [34]. Although human TRIM5α poorly restricts HIV-1, polymorphisms within the human TRIM5α gene have allowed for the development of human TRIM5α with high HIV-1 restriction activity [34]. The exact mechanism by which TRIM5α works is not well understood, however, it does concentrate around the viral core and recognizes and binds to the viral capsid to facilitate rapid uncoating [35]. This disrupts the reverse transcription process since uncoating is a carefully regulated process, and early or delayed uncoating may negatively affect virus infectivity [36]. Once the capsid has been targeted by TRIM5α, the viral core is disassembled which is likely mediated by proteosomal association with TRIM5α-virus complexes [36].
A3G is another antiviral protein expressed by lymphocytes, macrophages and dendritic cells following cellular stimulation by IFN-α and IL-2 [37]. A3G restricts HIV-1 infection by two separate mechanisms. The first involves the packaging of A3G during viral assembly directly into viral capsids in the absence of vif, and this occurs through the interaction with the viral capsid proteins [38]. This provides A3G with direct access to the HIV-1 genome where it induces mutations during the reverse transcription process by editing cytosine residues to uracil residues in the proviral minus strand [37,38,39]. The mutated viral DNA is then degraded due to reduced stability or the inability to be incorporated into the host genome [40]. However, these mutations are not always sufficient to prevent proviral integration, if the viral DNA still integrates successfully, the mutations often alter viral open reading frames or introduce premature stop codons resulting in misfolded or truncated viral proteins that are unable to produce infectious particles [37,40]. These misfolded viral proteins provide host cells with a pool of viral peptides that can be presented onto major histocompatibility complex (MHC) I molecules for antigen presentation. Expression of A3G may also enhance MHC I presentation and promotes the activation of cytotoxic T lymphocytes (CTLs) [37]. The other mechanism by which A3G limits HIV-1 infectivity is by directly interfering with the reverse transcription process, which may occur through the direct interaction between A3G and the viral reverse transcriptase enzyme [41]. Although A3G is highly effective at controlling virus replication, HIV-1 has adapted mechanisms to counteract the effects of A3G [37,40]. The HIV-1 accessory gene, vif, is essential for virus replication, but also has a role in promoting A3G degradation [25,37,40]. Vif binds A3G and mediates its polyubiquitination, tagging it for proteosomal degradation [39], but may also inhibit A3G mRNA translation by directly binding A3G mRNA to alter its stability [39].
Tetherin is also induced by type 1 IFNs and restricts HIV-1 by preventing virion release from the cell surface [42]. Since tetherin is an integral membrane protein with cytoplasmic, transmembrane, and extracellular domains, it can be incorporated into the membrane of virion particles as they bud from the cell surface [43]. Consequently, this serves to anchor virion particles to both the cell surface and to other virion particles as they bud from the host cell [42]. Once anchored to the cell membrane, virion particles can be endocytosed and degraded in lysosomal compartments [44]. Interestingly, HIV-1 has adapted a mechanism to prevent viral tethering. The HIV-1 vpu gene for example, encodes an integral membrane protein that interacts with tetherin transmembrane domains [45]. The vpu protein prevents the incorporation of tetherin into the envelope of virion particles and down regulates tetherin expression at the cell surface by trafficking tetherin to the trans-golgi network and away from the sites of virion assembly prior to lysosomal degradation [45].
Other antiviral proteins include SAMHD1 and the zinc finger antiviral protein (ZAP). SAMHD1 is a host protein found in resting macrophages, dendritic cells, and CD4+ T cells that cleaves deoxynucleoside triphosphages (dNTP) into deoxynucleosides and inorganic triphosphates, which depletes the dNTP pool required for HIV-1 reverse transcription [46]. This prevents the synthesis of full-length double stranded viral DNA and therefore prevents integration of proviral DNA [46]. The vpx gene of HIV-2 and the simian immunodeficiency virus (SIV) has the ability to disrupt SAMHD1 by interacting with the C-terminus and promoting proteosomal degradation [46]. HIV-1, however, does not contain the vpx gene so HIV-1 replication is actively suppressed in resting CD4+ T cells [47]. ZAP has been identified for its role in restricting the murine leukemia virus (MLV) along with HIV-1, however, some viruses can replicate normally in ZAP-expressing cells [48]. ZAP restricts HIV-1 by depleting multiply spliced mRNA by recruiting poly(A)-specific ribonucleases that shorten the poly(A) tail and directs mRNA to exosomes for degradation [48].

3.1.2. Natural Killer (NK) Cells

Further research investigating the innate control of HIV-1 has focused on identifying the roles of natural killer (NK) cells in controlling viral replication. The exact role of NK cells during HIV-1 infection is not well understood, however, it has been suggested that NK cells serve as a means of controlling viral replication prior to the induction of HIV-1-specific CD8+ T cell responses [49]. NK cells target HIV-1 infection by directly killing infected cells through the killer immunoglobulin-like receptor (KIR)-mediated recognition of target cells, degranulation resulting in granzyme and porforin release, the Fas-Fas ligand pathway, antibody-dependent cell-mediated cytotoxicity (ADCC), and modulating adaptive immune responses with IFN-γ production [50]. During acute HIV-1 infection, NK cells are known to rapidly proliferate [49], however, these cells may not be fully functional [48]. For example, Naranbhai et al., [51] observed reduced cytotoxic NK cell responses during acute HIV-1 infection. Similarly, antibody dependent cell mediated cytotoxicity (ADCC), the process by which antibodies bind to a specific pathogen and subsequently crosslink with Fc receptors on NK cells, macrophages, neutrophils, and mast cells, is also impaired during acute HIV-1 infection [50]. However, this may be attributed to the time required to generate HIV-1 specific antibody levels [50]. Once sufficient HIV-1 specific antibody levels were generated, strong NK cell responses were observed in chronic HIV-1 infected patients [52]. Although reduced cytotoxic responses and ADCC may be impaired during acute infection, the proportion of activating and inhibitory receptors on NK cells appears to be elevated [50]. As the disease progresses, a lower ratio of inhibitory to activating receptors has also been observed [50]. NK target cell lysis involves a balance between activating and inhibitory signals mediated through the ligation of MHC class I molecules. An increase in activating receptors during chronic HIV-1 infection implicates NK cells as important effector cells for controlling disease progression, however, further investigation into the diverse role of NK cells during HIV-1 infection is warranted.

3.1.3. γδ T Cells

γδT cells are a unique subset of innate immune effector cells that possess the γδT cell receptor. Unlike CD4+ and CD8+ T cell subsets, γδT cells do not require antigen presentation to become activated. The γδT cell subset can be further divided into Vδ1 and Vδ2 cells, which are localized to mucosal surfaces and peripheral blood, respectively [53]. Little is known about the role of γδT cells in HIV-1 infection; however, they do appear to play an important role in the control of HIV-1 viral replication. For example, Fenolilo et al., [53] observed an expansion of Vδ1 cells during HIV-1 infection, and these cells contained elevated gene expression levels for IFN-γ and IL-17. Similarly, in African patients, Vδ1 cells appeared to be expanded during HIV-1 and HIV-2 infection, and Vδ1 T cell counts appeared to be positively correlated with CD4+ T cell counts [54]. In addition to their cytotoxic activity and pro-inflammatory cytokine production, γδT cells have also been identified for their role in ADCC. In vitro studies have suggested that Vδ2 cells from HIV-1 patients are expanded and have potent ADCC activity [55]. Given that Vδ2 cells are present in the circulation, they have direct access to both circulating antibodies and circulating HIV-1 virion particles and may prove effective at controlling HIV-1 dissemination throughout the body.

3.2. Acquired Immunity to HIV-1

The acquired immune response involves cell-mediated (CMIR) and antibody-mediated immune responses (AbMIR). A balance between both responses is essential to maintaining overall health. The CMIR is primarily designed to combat intracellular pathogens such as viruses, and intracellular bacteria and parasites, whereas, the AbMIR has developed to combat extracellular pathogens such as extracellular bacteria and parasites. The CMIR involves antigen uptake and presentation by professional antigen presenting cells (APC) such as dendritic cells (DC) and macrophages that reside within epithelial cell layers. Following antigen uptake, the DCs migrate to a draining lymph node where they produce IL-12 to promote Th1 cell differentiation, and present antigens via MHC class II molecules to CD4+ T helper cells. This allows CD4+ T cells to become activated and produce the Th1 subset of cytokines, IFN-γ and IL-2. Production of IL-2 by CD4+ T cells is particularly important as it is necessary for the development and expansion of antigen-specific CD8+ T cells that recognize foreign antigen in the context of MHC class I molecules as a means of killing infected cells. The Th1 cytokine profiles also induce immunoglobulin (Ig) G2 production by B cells, which promotes complement activation and opsonization [56].
The AbMIR also involves antigen processing and presentation by DCs to CD4+ T cells, however, different cytokine profiles allow for the differentiation of Th2 cell subsets. These key cytokines include IL-3, 4, 5, 9, 10, and 13, which serve to suppress Th1 immune responses. In order to produce high levels of antigen specific antibodies, B cells must first be primed during initial antigen exposure. During this primary response, B cells differentiate and expand into memory or plasma cells. The plasma cells initially secrete IgM and small amounts of IgG. Upon subsequent antigen exposure, memory B cells become activated to rapidly produce high levels of IgG1 and other antibody isotypes.
Immune responses to HIV-1 infection tend to vary greatly from individual to individual; therefore, HIV-1 patients are generally classified based on their ability to control infection [57]. Although there is no standardized viral set point or CD4+T cell level to define a HIV-1 controller or progressor, in general, a controller is able to maintain low viral loads and CD4+ T cell levels in the absence of highly active antiretroviral therapy (HAART); whereas HIV-1 progressors have high viral loads and low CD4+ T cell counts [58]. However, many of the factors that differentiate a controller from a progressor are unclear, and despite over 30 years of HIV-1 research, there is still a great deal that is largely unknown.

3.2.1. Cell-Mediated Immune Response to HIV-1

The CMIR to HIV-1 is unique in that HIV-1 preferentially infects CD4+ CCR5+ T helper cells. During acute HIV-1 infection, the virus rapidly replicates and CD4+ T cell population numbers decline [59]. This decline in CD4+ T cells is attributed to both direct killing by the virus, and targeted CD8+ cytotoxic T lymphocyte (CTL) responses [59]. Maintenance of CD4+ T cells levels is essential to maintain HIV-1 specific CTL responses, which is associated with control of the infection [60]. During acute HIV-1 infection, HIV-1 specific CTLs emerge prior to neutralizing antibodies [61]. However, robust CTL responses to HIV-1 may not necessarily control the infection, as the viral nef protein is known to down-regulate MHC class I molecules to evade CTL responses [62]. It has also been suggested that CTL-mediated control of HIV-1 is specific to certain HIV-1 peptides. CTLs have been observed to mount the strongest cytotoxic responses to the gag and nef peptides [63,64]. However, this also induces selective pressure on HIV-1 to mutate these regions creating viral escape mutants [65]. It has been suggested that these HIV-1 escape mutants may be associated with the inability to control infection; however, this association is not always clear [66]. Some studies have suggested that these mutations come at a fitness cost to the virus [61,67,68], and escape tends to occur rapidly during acute infection and declines as the infection reaches the chronic state [61,69]. In some instances, the rate of mutation occurs so rapidly that after the initial infection, the transmitted virus or founder virus is completely lost [61]. Therefore, CTL responses that occur during acute infection must continuously adapt to changing HIV-1 peptides [61,70]. As the infection becomes chronic, immunological exhaustion becomes apparent due continuous immune activation in response to viral replication [71,72]. Consequently, CD8+ CTLs often exhibit an exhausted phenotype with increased CTLA-4 and PD-1 receptor expression and reduced cellular function [71,73]. HAART can help prevent this by limiting viral replication, however, if left untreated, the disease will continue to progress [73].

3.2.2. Antibody-Mediated Immune Response to HIV-1

Antibody responses to HIV-1 infection do not emerge until approximately 13 days post infection [74]. These antibodies include gp41-specific IgM and IgG, which are non-neutralizing [74]. As the infection progresses, IgG1 specific gp120 antibodies are produced [75]. These antibodies are specific to a variety of epitopes on gp120 including the CD4 binding site, glycan-containing regions, and the V3 loop [75]. The early production of non-neutralizing antibodies has little effect on viral infectivity and control of viral load; however, they do play a role in ADCC and viral opsonization [76]. During acute HIV-1 infection, IgG1-virus immune complexes are predominately comprised of gp41 antibodies and are present at relatively low levels compared to the viral load [76]. In contrast, during chronic HIV-1 infection, these virus-immune complexes were more abundant and are comprised of gp120-specific IgG1 [76,77]. This suggests that although HIV-1 specific antibodies emerge early after infection, they are not effective at controlling the viral load [76]. Neutralizing antibodies begin to emerge as early as 16 weeks post infection [75]. However, these early neutralizing antibodies may not be fully functional, as broad neutralizing ability does not emerge in HIV-1 patients until 2–3 years post infection [75]. Some HIV-1 infected patents, in contrast, never develop neutralizing antibodies, and it is unclear what factors contribute to these differences [78]. Additionally, the emergence of neutralizing antibodies may not necessarily control the infection. For example, Mikell et al. [78] observed a positive correlation between neutralizing ability and viral load, and Euler et al. [79] observed lower percentages of CD4+ T cells in patients with strong neutralizing activity; suggesting that HIV-1 may mutate env epitopes to escape virus neutralization.

3.2.3. Other T Cell Subsets

Recent studies have implicated roles of Th17 and Treg cells in the pathogenesis of lentiviral infections. Research investigating the role of Th17 cells in HIV-1, for example, found that long-term non-progressors had higher Th17 cell numbers compared to disease progressors, and higher Th17 cell numbers were associated with a lower viral load [80]. Additionally, it has been established that Th17 cells and Treg cells interact during HIV-1 infection to regulate immune responses [81]. Long-term non-progressors tended to have Th17/Treg ratios similar to uninfected controls, whereas, individuals that did not easily control the rate of viral replication had depleted Th17 numbers and increased Treg numbers suggesting a switch towards an anti-inflammatory immune response [81]. However, it is unclear if maintenance of Treg cell populations during HIV-1 infection is associated with rapid or delayed disease progression. It has been suggested that Treg-mediated IL-10 production is also associated with HIV-1 disease progression as it suppresses specific CD4+ T cell responses [82]. However, other studies have suggested that high levels of Treg cells can help prevent disease progression by limiting CD4+ T cell and CTL responses, thus limiting continuous immune activation and preventing immunological exhaustion [83,84]. Therefore, further research is required to understand the role of Tregs in HIV-1 disease progression.

3.3. Innate Immune Response to SRLV

3.3.1. Toll-Like Receptors and Antiviral Peptides

The role of viral-induced TLR signaling has not been widely studied in sheep and goats, however, during SRLV infection, TLR 7 and 8 become activated inducing IFN-α, IL-6, TNF-α production and subsequent antiviral protein expression [85]. It is unclear if TLR signaling pathways induce SRLV replication, or if the SRLV genome has a NF-κB transcriptional binding site in the promoter. However, given the importance of macrophages as innate immune effector cells, macrophage maturation and activation can induce SRLV replication.
There is considerably less research investigating the roles of the intrinsic restriction factors TRIM5α, A3G, and tetherin in SRLV infection, however, TRIM5α has recently been identified in sheep and goats, and has been found to be effective at restricting SRLV [86]. An A3G- like protein has also been identified in sheep, and has shown cytodine deaminase activity [87]. Like HIV-1, SRLV contains the accessory vif gene to combat the restrictive activity of A3G, and SRLV vif appears to restrict A3G across species [87]. Tetherin has been investigated in sheep due to its role in restricting endogenous retroviruses [88]. Since the SRLV genomes lack the accessory gene vpu, tetherin likely has high SRLV restriction activity. However, further investigation into the roles of TLR activation and intrinsic restriction factors in limiting SRLV infection is necessary.

3.3.2. NK Cells

The role of NK cells in SRLV infection has not been investigated; however, given the importance of NK cells for HIV-1 infection, it is likely that they play an important role in the control of SRLV. It is unclear how NK cells target SRLV-infected macrophages; however, we may speculate that they recognize and bind infected cells and virion particles through a number of mechanisms including KIR-mediated recognition, degranulation, complement activation, ADCC and the production of IFN-γ which serves to either kill infected cells, or modulate virus-specific immune responses. ADCC has been investigated as a possible control mechanism in MVV infected sheep [89]. Sheep vaccinated with a recombinant env protein had higher IgG2 antibody titers an IgG1 and polyclonal serum had ADCC activity compared to MVV-infected non-vaccinated animals suggesting that MVV specific IgG2 may have strong ADCC activity early in infection [89].

3.3.3. γδ T Cells

In ruminants, γδT cells comprise approximately 70% of all lymphocytes in young animals, and are an important part of the innate immune system [90]. CAEV-infected goats have a significantly higher proportion of γδT cells compared to healthy goats, which suggests that these cells may be important for controlling SRLV infection [90,91,92]. Given that γδT cells tend to localize to mucosal surfaces, it is possible that this cell type plays a crucial role in limiting SRLV entry and mediating early immune responses against these viruses.

3.4. Acquired Immunity to SRLV

During SRLV infection, both branches of the acquired immune system are activated though it remains unclear how each relates to either host protection or disease progression [93]. Like HIV-1, the degree of the immune response influences the viral load, which is correlated to the severity and presence of clinical disease symptoms [94,95]. Animals that respond with a CMIR are often referred to as long-term non-progressors because they exhibit a persistent viral infection but lack clinical symptoms and have a low viral load. These animals produce high levels of IgG2 antibodies specific to gp135, and a dominant subset of gp135 responsive Th cells displaying high levels of IFN-γ gene expression [94,95]. In contrast, arthritic animals tend to mount a type 2 or AbMIR, characterized by high polyclonal SRLV reactive IgG1 antibody titers and a dominant subset of Th2 cells with low proliferation and enhanced IL-4 gene expression [84,95].

3.4.1. Cell-Mediated Immune Response to SRLV

The CMIR is likely the most efficient response for controlling viral load. Since SRLV, unlike HIV-1, does not infect CD4+ T cells, the maintenance of CD4+ T cell populations during SRLV infection will allow for the development and maintenance of SRLV-specific CTLs. However, SRLV may interfere with CD4+ T cell proliferation as CAEV-infected arthritic goats had reduced CD4+ T cell proliferation compared to CAEV-infected asymptomatic goats [96]. Reduced lymphocyte proliferation was also observed in clinically affected sheep compared to MVV-infected asymptomatic animals [97]. Since the SRLV genome does not contain the viral nef gene, down-regulation of MHC I by SRLV likely does not occur. However, SRLV may down-regulate MHC class II molecule expression on SRLV-infected macrophages [91], and down-regulation of CD80 co-stimulatory molecules has been observed in sheep with clinical disease symptoms [97]. Overall, this suggests that SRLV infection appears to interfere with antigen processing and presentation, and thus limits the ability of antigen presenting cells to activate CD4+ T cells and induce CTL responses.
Although the CMIR is the preferential response for maintaining a low viral load, the presence of type 1 cytokines is not sufficient to control viral replication [94]. In fact, some of the Th1 cytokines, including IFN-γ, TNF-α and GM-CSF, activate the SRLV promoter and induce viral replication [98]. These cytokines activate the viral promoter in the U3 region 70 base pair repeat and this is mediated through the STAT1 pathway [99]. This also requires at least one gamma-activating site (GAS) within the viral promoter [100], indicating that monocyte differentiation and macrophage activation can induce viral transcription. Although not all SRLV strains contain a GAS, the presence of a GAS in the SRLV LTR is not necessary for viral replication [101]. However, high levels of viral replication in response to Th1 cytokines may lead to a cycle of continuous immune activation and to eventual immunological exhaustion and disease progression.

3.4.2. Antibody-Mediated Immune Response to SRLV

The antibody response to SRLV generally targets epitopes on the gp135, gp38, and capsid proteins [102]. Antibody responses can emerge as early as 2–4 weeks post infection, and tend to fluctuate during the first 6 months of infection [103]. Additionally, like HIV-1 these early antibodies are specific to linear epitopes and are thus non-neutralizing [94,96]. However, as discussed previously, these early antibodies may play an important role in ADCC [89]. Neutralizing antibodies can take as long as 2 years to emerge and can control virus infection [104]. However, like HIV-1, SRLV virus epitopes may mutate in response to selection pressure imposed by host immunoglobulins [104,105]. These mutations tend to occur in the fourth variable domain of gp135 and the mutation of a conserved cytosine was shown to change the neutralization epitope [106], these mutations likely contribute to disease progression. Antibody responses, in general, may contribute to SRLV disease progression, since asymptomatic animals also tend to mount a CMIR with a low titer of gp135-specific IgG2, while arthritic animals exhibit very high levels of IgG1 and a higher IgG1/IgG2 ratio than asymptomatic animals [107]. Although further research is necessary to better understand the roles of neutralizing antibodies in the control of SRLV infection, it is evident that an AbMIR is not sufficient to control infection [108], and readily contributes to disease progression.

3.4.3. Immune Dysregulation

The dynamics of immune evasion strategies of SRLV are unclear, however, SRLV infection can alter macrophage function, and immune dysregulation is apparent. This is particularly evident in animals that exhibit clinical signs of arthritis or mastitis, as these inflammatory conditions are characterized by dense mononuclear cell infiltration accompanied by necrosis and edema [108]. Arthritic animals also exhibit thickening and fibrosis of the articular capsule, and erosion and ulcer formation of the articular cartilage occurs in severe cases [109]. Consequently, lesions form in the joint synovial membranes as well as the mammary gland [5]. Histological analysis of these lesions revealed large numbers of macrophages, CD8+ T cells and B cells, and the proportion of B cells present in the lesions increases as the infection persists [110]. To investigate the immune dysregulation that occurs during SRLV infections, Lechner et al., [111] examined the cytokine profiles produced by CAEV-infected macrophages in culture. This study revealed that infected macrophages produced elevated IL-8 and monocyte chemotactic protein-1 (MCP1), but had reduced levels of TGF-β mRNA [101]. In addition to this, reduced levels of TNF-α, IL-1β, IL-6 and IL-12 mRNA, and increased levels of GM-CSF mRNA were observed in lipopolysaccharide (LPS)-stimulated CAEV-infected macrophages [111]. Higher levels of pro-inflammatory cytokine gene expression, such as IL-1β, IL-6, IL-12, and TNF-α likely contribute to increased immune activation and trafficking of effector cells to the infection site causing inflammation and lesion formation. Elevated levels of GM-CSF has also been observed in alveolar macrophages of MVV infected sheep [112], and SRLV infection can induce a phenotypic shift in macrophages from pro-inflammatory M1 to an anti-inflammatory M2 macrophages that favoured viral replication [113]. Interestingly, several reports have suggested that SRLV-infected goats are not immunocompromised [5,6,111,114], as is the case for HIV-1 patients. However, the altered cytokine profiles observed in CAEV- and MVV-infected macrophages, along with the reduced DTH response to mycobacterial antigens in MVV-infected sheep, suggest that altered cellular functions may directly affect immunity [115].

4. Genetics of Lentiviral Resistance

There is little research investigating the genetic parameters that affect resistance or susceptibility to SRLV, however, research with HIV-1 patients has led to the discovery of a variety of genetic polymorphisms that appear to be associated with disease resistance, or slower disease progression. Most of the genes associated with HIV-1 resistance encode a variety of immune molecules such as MHC class I, chemokine receptor (CCR) 5, KIR, and TLRs. For example, the HLA-B*27, B*57, the Bw4 alleles as well as heterozygosity at these class I loci are associated with slower disease progression [116]. However, the effect of these alleles on HIV-1 disease progression is limited to Caucasian and African populations [117]. In the Japanese population, HLA-B*52, B*67, and C*12 have been associated with a lower viral load [117]. East Asian populations also have a high proportion of individuals carrying a deletion in the A3B coding region, which increases susceptibility to HIV-1 infection [118]. There are also significant population differences in the CCR5 Δ32 mutation that is associated with resistance to HIV-1 [119]. The Δ32 mutation, for example, is only found in European, West Asian, and North African populations and shows a north-to-south decline [120]. Individuals homozygous for this mutation have a non-functional CCR5 receptor, and protection has been observed in both Δ32 homozygous and heterozygous individuals [120].
Similarly, breed differences in resistance to SRLV, as well as polymorphisms in ovine TLR 7, 8, CCR5 and MHC genes, have been found to be associated with resistance to SRLV [114,121,122,123,124]. For example, a higher proportion of polymorphisms in the ovine TLR 7 and 8 leucine rich repeat have been identified in MVV-infected sheep [121], and a deletion in the ovine CCR5 gene was associated with a reduced proviral load in rambouillet, Polypay and Columbia breeds [124]. Given these genetic parameters, it may be possible to selectively breed sheep and goats to have enhanced SRLV resistance. Many of the genetic polymorphisms associated with HIV-1 resistance were identified using genome-wide association studies (GWAS). To date, few sheep GWASs, and no goat GWASs have been carried out to identify genes associated with SRLV resistance. One possible reason for this may be due to difficulty in reliably identifying the phenotype, using current diagnostic methods. For example, the agar gel immunodiffusion (AGID) test and the enzyme-linked immunosorbent assay (ELISA) are widely used to detect SRLV infection [125]. Both of these tests, however, depend on the presence of host antibodies that are specific to SRLV, which are influenced by variables such as age and health status of the animal [126]. It is also possible that the AGID and ELISA tests may be influenced by genetic differences among SRLV strains [7]. Polymerase chain reaction (PCR) is also being used to test for the presence of viral RNA and proviral DNA in different tissues. This test is more sensitive than serological testing, however, problems with primer binding heterogeneity can make detection difficult [127] and the increased cost of PCR makes this relatively inaccessible to most producers [11]. Despite these challenges, a recent ovine GWAS identified a transmembrane protein gene, TMEM154, as a candidate gene for SRLV resistance [128]. The role of this protein is currently unknown, but it is expressed at high levels in B cells and monocytes, which suggests that it may be of immunological importance [128]. Breed differences have also been identified for the TMEM154 gene, the Dalsebred, Herdwick, and Rough Fells breeds had a higher allele frequency for the TMEM154 mutation that is associated with SRLV resistance [123]. An additional ovine GWAS identified another transmembrane protein, TMEM38A, as a possible gene associated with SRLV resistance, whereas the DPPA2 gene was associated with susceptibility [129]. The DPPA2 gene is involved with embryonic lung development and suggests that altered lung development in sheep may increase susceptibility to SRLV infections [129].
Although breeding livestock for enhanced disease resistance is becoming an increasingly popular means of improving animal health, in the context of SRLV infections, using genomic selection to breed for SRLV resistance may not be practical. As observed in HIV-1 studies, resistance or slower disease progression is a polygenic trait, involving a complex interaction between a variety of different innate and acquired immune genes [130,131]. It may, however, be possible to breed for resistance using phenotypic rather than genotypic selection. In sheep, helminth resistance is also a polygenic trait and resistant sheep have been bred based on phenotypic parameters such as fecal egg count [132]. However, it is unclear if breeding for resistance to one disease will increase susceptibility to others. Also, if SRLV resistance was introduced based on phenotypic selection, it is unclear how the virus will adapt. Since SRLV mutation rates are high and the virus already mutates in response to immune-based selection pressure, it is possible that the virus will quickly adapt, rendering the breeding program redundant.
An additional approach to breeding for SRLV resistance is by breeding for overall enhanced immune responses (EIR). This approach involves measuring CMIR and AbMIR in response to various antigens. A recent GWAS in dairy cattle found several single-nucleotide polymorphisms (SNPs) in the MHC locus that were associated with both AbMIR and CMIR [133]. In goats, the MHC haplotype Be10-D2 was associated with rapid seroconversion and higher antibody titers compared to the Bel-D5 haplotype [114]. Given this association, it is possible that further investigation and SNP discovery will allow for the identification of EIR sheep and goats that readily control SRLV disease progression.

5. Conclusions

The immune response to lentiviral infections is a complex and dynamic response, and to date very little is known about how the immune system responds to these infections. Extensive research on HIV-1 has greatly contributed to our knowledge of the dynamics of the host–virus interaction; however, HIV and SRLV, despite their similarities are two distinct viruses and extrapolating knowledge from HIV research must be approached with caution. It is evident from the research presented here that gaps exist in our knowledge of SRLV, and before exploring the possibility of breeding for resistance, extensive research needs to be done to better understand SRLV immune responses. These knowledge gaps primarily lie in the areas that focus on how SRLV modulate the host immune response and the dynamics of early infection. As previously discussed, for example, the Th1/Th2 paradigm appears to apply to SRLV infection; however, if in fact SRLV infection steers macrophage polarization towards a M2 phenotype favoring the Th2 immune response, then one would expect that the disease would progress rapidly and clinical disease would be apparent in a large proportion of animals. However, it often takes years before clinical SRLV infection becomes apparent. Therefore, future studies should firstly investigate how SRLV infection alters macrophage function as a whole. To start, cytokine production and intracellular signaling should be investigated. It would also be beneficial to investigate how endogenous stress hormone levels affect disease resistance and progression, and how host macrophages respond to SRLV during co-infection with other pathogens. Since macrophages play an important role as both innate effector cells and antigen presenting cells, an improved understanding of SRLV-infected macrophage function will improve our understanding of how SRLV is controlled during early infection, how the acquired immune response is induced, and how SRLVs modulate the host immune response.

Conflicts of Interest

The authors declare no conflict of interest.

References and Notes

  1. Fieni, F.; Rowe, J.; van Hoosear, K.; Burucoa, C.; Oppenheim, S.; Anderson, G.; Murray, J.; BonDurant, R. Presence of caprine arthritis encephalitis virus (CAEV) proviral DNA in genital tract tissues for superovulated dairy goat does. Theriogeneology 2003, 59, 1515–1523. [Google Scholar] [CrossRef]
  2. Konshi, M.; Tsuduku, S.; Haritani, M.; Murakami, K.; Tsuboi, T.; Kobayashi, C.; Yoshikawa, K.; Kimura, K.; Sentsui, H. An epidemic of caprine arthritis encephalitis in Japan: Isolation of the virus. Virology 2004, 66, 911–917. [Google Scholar]
  3. Pisoni, G.; Quasso, A.; Moroni, P. Phylogentic analysis of small ruminant lentivirus subtype B1 in mixed flocks: Evidence for natural transmission from goats to sheep. Virology 2005, 399, 147–152. [Google Scholar] [CrossRef]
  4. Pisoni, G.; Bertoni, G.; Puricelli, M.; Maccalli, M.; Moroni, P. Demonstration of coinfection with and recombination by caprine arthritis encephalitis virus and maedi visna virus in naturally infected goats. J. Virol. 2007, 81, 4948–4955. [Google Scholar] [CrossRef]
  5. Ravazzolo, A.; Nenci, C.; Vogt, H.; Waldvogel, A.; Obexer-Ruff, G.; Peterhans, E.; Bertoni, G. Viral load, organ distribution, histopathological lesions, and cytokine mRNA expression in goats infected with a molecular clone of the caprine arthritis encephalitis virus. Virology 2006, 350, 116–127. [Google Scholar]
  6. Dufour, S.; Zahno, M.; Vogt, H.; Peterhans, E.; Bertoni, G. Production of monospecific antibody to immunodominant epitopes of the caprine arthritis encephalitis virus transmembrane glycoprotein and analysis of their activity in vitro. Intervirology 2001, 45, 177–182. [Google Scholar]
  7. Olech, M.; Rachid, A.; Croise, B.; Kuzmak, J.; Valas, S. Genetic and antigenic characterization of small ruminant lentiviruses circulating in Poland. Virus Res. 2012, 163, 528–563. [Google Scholar]
  8. Polledo, L.; Gonzalez, J.; Benavides, J.; Morales, S.; Martinez-Fernandez, B.; Delago, L.; Reina, R.; Glaria, I.; Perez, V.; Ferreras, M.; et al. Patterns of lesion and local host cellular immune response in natural cases of ovine maedi visna. J. Comp. Path. 2011, 147, 1–10. [Google Scholar]
  9. Bolea, R.; Monleon, E.; Carrasco, L.; Vargas, A.; de Andres, D.; Amorena, B.; Badiola, J.; Jujan, L. Maedi-Visna virus infection of ovine mammary epithelial cells. Vet. Res. 2006, 37, 133–144. [Google Scholar]
  10. Ali Al Ahmad, M.; Chebloune, Y.; Chatagnon, G.; Pellerin, J.; Fieni, F. Is caprine arthritis encephalitis virus (CAEV) transmitted vertically to early embryo development stages (morulae or blastocyst) via in vitro infected frozen semen. Theriogenology 2012, 77, 1673–1678. [Google Scholar] [CrossRef]
  11. Modolo, J.; Stachissini, A.; Padovani, C.; Araujo Junior, J.; Castro, R.; Ravazzolo, A.; Leite, B. PCR associated with agar gel immunodifusion assay improve caprine arthritis encephalaitis (CAEV) control. Small Rumin. Res. 2009, 81, 18–20. [Google Scholar] [CrossRef]
  12. Reina, R.; Grego, E.; Bertolotti, L.; de Meneghi, D.; Rosati, S. Genome analysis of small ruminant lentivirus genotype E: A caprine lentivirus with natural deletions of the dUTPase subunit, vpr-like accessory gene, and 70-base-pair repeat of the U3 regions. J. Virol. 2009, 83, 1152–1155. [Google Scholar] [CrossRef]
  13. L’Homme, Y.; Ouardani, M.; Levesque, V.; Bertoni, G.; Simard, C.; Pisoni, G. Molecular characterization and phylogenetic analysis of small ruminant lentiviruses isolatd from Canadian sheep and goats. Virol. J. 2011, 8, 271–278. [Google Scholar] [CrossRef][Green Version]
  14. Abelson, M.; Schoborg, R. Characterization of the caprine arthritis encephalitis virus (CAEV) rev N-terminal elements required for efficient interaction with the RRE. Virus Res. 2003, 92, 23–35. [Google Scholar] [CrossRef]
  15. Harmache, A.; Bouyac, M.; Audoly, G.; Hieblot, C.; Peveri, P.; Vigne, R.; Suzan, M. The vif gene is essential for efficient replication of caprine arthritis encephalitis virus in goat synovial membrane cells and affects the late steps of the virus replication cycle. J. Virol. 1995, 69, 3247–3257. [Google Scholar]
  16. Harmache, A.; Russo, P.; Guiguen, F.; Vtu, C.; Vignoni, M.; Bouyac, M.; Heiblot, C.; Pepin, M.; Vigne, R.; Suzan, M. Requirement of caprine arthritis encephalitis virus vif gene for in vitro replication. Virology 1996, 224, 246–255. [Google Scholar] [CrossRef]
  17. Kristbjornsdottir, H.; Andresdottir, V.; Svansson, V.; Throsteinsdottir, S.; Matthiasdottir, S.; Andresson, O. The vif gene of madei-visna is essential for infectivity in vivo and vitro. Virology 2004, 318, 350–359. [Google Scholar] [CrossRef]
  18. Angelopoulou, K.; Brellou, G.; Greenland, T.; Vlemmas, I. A novel eletion in the LTR region of a Greek small ruminant lentivirus may be associated with low pathogenicity. Virus Res. 2006, 118, 174–184. [Google Scholar]
  19. Angelopoulou, K.; Poutahidis, T.; Brellou, G.; Greenland, T.; Vlemmas, I. 2008. A deletion in the R region of long terminal repeats in small ruminant lentiviruses is associated with decreased pathology in the lung. Vet. J. 2008, 175, 346–355. [Google Scholar] [CrossRef]
  20. Leroux, C.; Mornex, J. Retroviral infections in sheep and the associated diseases. Small Rumin. Res. 2008, 76, 68–76. [Google Scholar] [CrossRef]
  21. Villet, S.; Faure, C.; Bouzar, B.; Morin, T.; Verdier, G.; Chebloune, Y.; Legras, C. Lack of trans-activation function for madei visna virus and caprine arthritis encephalitis virus tat proteins. Virology 2003, 307, 317–327. [Google Scholar] [CrossRef]
  22. Villet, S.; Bouzar, B.; Morin, T.; Verdier, G.; Legras, C.; Chebloune, Y. Madei-Visna and caprine arthritis encelphalitis vurs genomes encode a vpr-like but no tat protein. J. Virol. 2003, 77, 9632–9638. [Google Scholar] [CrossRef]
  23. Valas, S.; Rolland, M.; Perrin, C.; Perrin, G.; Mamoun, R. Characterization of a new 5’ splice site within the caprine arthritis encephalitis virus genome: evidence for a novel auxiliary protein. Retrovirology 2008, 5, 1–17. [Google Scholar] [CrossRef]
  24. Watts, J.; Dang, K.; Gorelick, R.; Leonard, C.; Bess, J.; Swanstrom, R.; Burch, C.; Weeks, K. Architecture and secondary structure of an entire HIV-1 RNA genome. Nature 2009, 460, 711–719. [Google Scholar] [CrossRef]
  25. Malim, M.; Emerman, M. HIV-1 accessory proteins—Ensuring viral survival in a hostile environment. Cell Host Microbe 2008, 3, 388–398. [Google Scholar] [CrossRef]
  26. Arellano, E.; Alcami, J.; Lopez, M.; Soriano, V.; Holguin, A. Drastic decrease of transcription activity due to hypermutated long terminal repeat (LTR) region in different HIV-1 subtypes and recombinants. Antivir. Res. 2010, 88, 152–159. [Google Scholar] [CrossRef]
  27. Katagiri, D.; Hayashi, H.; Victoriano, A.; Okamoto, T.; Onozaki, K. Estrogen stimulates transcription of human immunodeficiency virus type 1 (HIV-1). Int. Immunopharmacol. 2006, 6, 170–181. [Google Scholar] [CrossRef]
  28. Kawai, T.; Akira, S. TLR signaling. Cell Death Differ. 2006, 13, 816–825. [Google Scholar] [CrossRef]
  29. Akira, S.; Takeda, K. Toll-Like receptor signaling. Nat. Rev. Immunol. 2004, 4, 499–511. [Google Scholar]
  30. Oh, D.; Baumann, K.; Hamouda, O.; Eckert, J.; Neumann, K.; Kucherer, C.; Bartmeyer, B.; Poggensee, G.; Oh, N.; Pruss, A.; et al. A frequent functional toll-like receptor 7 polymorphism is associated with accelerated HIV-1 disease progression. AIDS 2009, 23, 297–307. [Google Scholar] [CrossRef]
  31. Chang, J.; Lacas, A.; Lindsay, R.; Doyle, E.; Axten, K.; Pereyra, F.; Rosenberg, E.; Walker, B.; Allen, T.; Altfeld, M. Differential regulation of toll-like receptor pathways in acute and chronic HIV-1 infection. AIDS 2012, 26, 533–541. [Google Scholar] [CrossRef]
  32. Ranjbar, S.; Jasenosky, L.; Chow, N.; Goldfield, A. Regulation of mycobacterium tuberculosis-dependant HIV-1 transcription reveals a new role of NFAT5 in the toll-like receptor pathway. PLoS Pathog. 2012, 8, e1002620. [Google Scholar] [CrossRef]
  33. Goujon, C.; Malim, M. Characterization of the alpha interfearon-induced postentry block to HIV-1 infection in primary human macrophages and T cells. J. Virol. 2010, 84, 9254–9266. [Google Scholar] [CrossRef]
  34. Pham, Q.; Bouchard, A.; Grutter, M.; Berthoux, L. Generation of human TRIM5α mutants with high HIV-1 restriction activity. Gene Ther. 2010, 17, 859–871. [Google Scholar] [CrossRef]
  35. Campbell, E.; Perez, O.; Anderson, J.; Hope, T. Visualization of proteasome-independent intermediate during restriction of HIV-1 by rehesus TRIM5α. J. Cell Biol. 2008, 180, 549–561. [Google Scholar] [CrossRef]
  36. Lukic, Z.; Hausmann, S.; Sebastian, S.; Rucci, J.; Sastrji, J.; Robia, S.; Juban, J.; Campbell, E. TRIM5α associates with proteosomal subunits in cells while in complex with HIV-1 virons. Retrovirology 2011, 8, 1–15. [Google Scholar] [CrossRef]
  37. Casartelli, N.; Guivel-Benhassine, F.; Bouziat, R.; Brandler, S.; Schwartz, O.; Moris, A. The antiviral factor APOBEC3G improves CTL recognition of cultured HIV-infected T cells. J. Exp. Med. 2010, 207, 39–49. [Google Scholar] [CrossRef]
  38. Khan, M.; Golia-Gaur, R.; Kao, S.; Miyagi, E.; Walker, R.; Strebel, K. Encapsidation of APOBEC3G into HIV-1 virons involves lip raft assocaiation and does not correlate with APOBEC3G oligomerization. Retrovirology 2009, 6, 1–12. [Google Scholar] [CrossRef][Green Version]
  39. Feng, Y.; Love, R.; Chelico, L. HIV-1 viral infectivity factor (vif) alters possessive single stranded DNA scanning of the retroviral restriction factor, APOBEC3G. J. Biol. Chem. 2013, 288, 6083–6094. [Google Scholar] [CrossRef]
  40. Britan-Rosich, E.; Nowarski, R.; Kotler, M. Multifaceted counter-APOBEC3G mechanisms employed by HIV-1 vif. J. Mol. Biol. 2011, 410, 1065–1076. [Google Scholar] [CrossRef]
  41. Wang, X.; Ao, Z.; Chen, L.; Kobinger, G.; Peng, J.; You, X. The cellular antiviral protein APOBEC3G interacts with HIV-1 reverse transcriptase and inhibits its function during viral replication. J. Virol. 2012, 86, 3777–3786. [Google Scholar] [CrossRef]
  42. Perez-Caballero, D.; Zang, T.; Ebrahimi, A.; McNatt, M.; Gregory, D.; Johnson, M.; Bieniasz, P. Tetherin inhibits HIV-1 release by directly tethering virons to cells. Cell 2009, 139, 499–511. [Google Scholar] [CrossRef]
  43. Dube, M.; Roy, B.; Guiot-Guillain, P.; Binette, J.; Mercier, J.; Chiasson, A.; Cohen, E. Antagonism of thetherin restriction of HIV-1 release by Vpu involves binding and sequestration of the restriction factor in a perinuclear compartment. PLoS Pathog. 2010, 6, e1000856. [Google Scholar] [CrossRef][Green Version]
  44. Miyakawa, K.; Ryo, A.; Murakami, T.; Ohba, K.; Yamaoka, S.; Fukuda, M.; Guatelli, J.; Yammamoto, N. BCA2/Rabring7 promotes tetherin-dependant HIV-1 restriction. PLoS Pathog. 2009, 5, e1000700. [Google Scholar] [CrossRef]
  45. Kueck, T.; Neil, S. A cytoplasmic tail determinant in HIV-1 Vpu mediates targeting of tetherin for endosomal degradation and counteracts interfearon-induced restricton. PLoS Pathog. 2012, 8, e1002609. [Google Scholar] [CrossRef]
  46. Sze, A.; Olagnier, D.; Lin, R.; van Grevenyghe, J.; Hiscott, J. SAMHD1 host restriction factor: A link with innate immune sensing of retrovirus infection. J. Mol. Biol. 2013, 425, 4981–4494. [Google Scholar] [CrossRef]
  47. Baldauf, H.M.; Pan, X.; Erikson, E.; Schmidt, S.; Daddacha, W.; Burggraf, M.; Schenkova, K.; Abbiel, I.; Wabintz, G.; Gramberg, T.; et al. SAMHD1 restricts HIV-1 infection in resting CD4+ T cells. Nat. Med. 2012, 18, 1682–1689. [Google Scholar] [CrossRef]
  48. Zhu, Y.; Chen, G.; Lv, F.; Wang, X.; Ji, X.; Xu, Y.; Sun, J.; Wu, L.; Zheng, Y.; Gao, G. Zinc-finger antiviral protein inhibits HIV-1 infection by selectively targeting multiply spliced viral mRNAs for degradation. Proc. Natl. Acad. Sci. USA 2011, 108, 15834–15839. [Google Scholar]
  49. Alter, G.; Teigen, N.; Ahern, R.; Streeck, H.; Meier, A.; Rosenberg, E.; Atfeld, M. Evolution of innate and adaptive effector cell functions during acute HIV-1 infection. J. Infect. Dis. 2007, 195, 1452–1460. [Google Scholar] [CrossRef]
  50. Jia, M.; Li, D.; He, X.; Zhao, Y.; Peng, H.; Ma, P.; Hong, K.; Liang, H.; Shao, Y. Impaired natural killer cell induced antibody-dependant cell-mediated cytotoxicity is associated with human immunodeficiency virus-1 disease progression. Clin. Exp. Immunol. 2012, 171, 107–116. [Google Scholar]
  51. Naranbhai, V.; Altfeld, M.; Karim, S.; Ndung’u, T.; Karim, Q.; Carr, W. Changes in natural killer cell activation and function during primary HIV-1 infection. PLoS One 2013, 8, e53251. [Google Scholar]
  52. Thobakgale, C.; Fadda, L.; Lane, K.; Toth, I.; Pereyra, F.; Bazner, S.; Ndung’u, T.; Walker, B.; Rosenberg, E.; Alter, G.; et al. Frequent and strong antibody-mediated natural killer cell activation in response to HIV-1 env in individuals with chronic HIV-1 infection. J. Virol. 2012, 86, 6986–6993. [Google Scholar] [CrossRef]
  53. Fenogilo, D.; Poggi, A.; Catellani, S.; Battagila, F.; Ferrera, A.; Setti, M.; Murdaca, G.; Zocchi, M. Vδ1 T lymphocytes producing IFN-γ and IL-17 are expanded in HIV-1 infected patients and respond to candida albicans. Blood 2009, 113, 6611–6618. [Google Scholar] [CrossRef]
  54. Zheng, N.; McElrath, J.; Sow, P.; Mesher, A.; Hawes, S.; Stern, J.; Gottlieb, G.; De Rosa, S.; Kiviat, N. Association between peripheral γδ T-cell profile and disease progression in individuals infected with HIV-1 or HIV-2 in West Africa. J. Acquir. Immune Defic. Syndr. 2011, 57, 92–100. [Google Scholar] [CrossRef]
  55. Poonia, B.; Pauza, C. Gamma delta T cells form HIV+ donor can be extended in vitro by zoledronate/interlukin-2 to become cytotoxic effectors for antibody-dependant cellular cytotoxicity. Cytotherapy 2012, 14, 173–181. [Google Scholar] [CrossRef]
  56. Kindt, T.; Goldsby, R.; Osborne, B. Kuby Immunology, 6th ed.; W.H. Freeman and Company: New York, NY, USA, 2007; pp. 314–315. [Google Scholar]
  57. Munier, C.; Kelleher, A.; Kent, S.; De Rose, R. The role of T cell immunity in HIV-1 infection. Curr. Opin. Virol. 2013, 3, 1–9. [Google Scholar] [CrossRef]
  58. Buckheit, R.; Siliciano, R.; Lankson, J. Primary CD8+ T cells form elite suppressors effectively eliminate non-productively HIV-1 infected resting and activated CD4+ T cells. Retrovirology 2013, 10, 1–12. [Google Scholar] [CrossRef]
  59. Okoye, A.; Rohankhedkar, M.; Abana, C.; Pattenn, A.; Reyes, M.; Pexton, C.; Lum, R.; Sylwester, A.; Planer, S.; Legasse, A.; et al. Naive T cells are dispensable for memory CD4+ T cell homeostatis in progressive simian immunodeficiency virus infection. J. Exp. Med. 2012, 209, 641–651. [Google Scholar] [CrossRef]
  60. Ranasinghe, S.; Flanders, M.; Cutler, S.; Soghian, Z.; Ghebremichael, M.; Davis, I.; Lindquvist, M.; Pereyra, F.; Walker, B.; Heckerman, D.; et al. HIV-specific CD4 T cell responses to different viral proteins hve discordant assocaitions with viral load and clinical outcome. J. Virol. 2011, 86, 277–283. [Google Scholar]
  61. Goonetilleke, N.; Liu, M.; Salazar-Gonzalez, J.; Ferrari, G.; Giorgi, E.; Ganusov, V.; Keele, B.; Learn, G.; Turnbull, E.; Salazar, M.; et al. The first T cell response to transmitted/founder virus contributes to the control of acute virema in HIV-1 infection. J. Exp. Med. 2009, 206, 1253–1272. [Google Scholar] [CrossRef]
  62. Cohen, G.; Gandhi, R.; Davis, D.; Mandelboim, O.; Chen, B.; Strominger, J.; Baltimore, D. The selective downregulation of class 1 major histocompatibility complex proteins by HIV-1 protects HIV-infected cells from NK cells. Immunity 1999, 10, 661–671. [Google Scholar] [CrossRef]
  63. Frahm, N.; Korber, B.; Adams, C.; Szinger, J.; Draenert, R.; Addo, M.; Feeney, M.; Yusim, K.; Sango, K.; Brown, N.; et al. Constant cytotoxic T lymphocyte targeting of immunodominant regions in human immunodeficiency virus across multiple ethnicities. J. Virol. 2004, 78, 2178–2200. [Google Scholar]
  64. Chevalier, M.; Julg, B.; Pyo, A.; Flanders, M.; Ranasinghe, S.; Soghoian, D.; Kwon, D.; Rychert, J.; Lian, J.; Muller, M.; et al. HIV-1-specific interleukin-21+ CD4+ T cell responses contribute to durable viral control through the modulation of HIV-specific CD8+ T cell function. J. Virol. 2011, 85, 733–741. [Google Scholar] [CrossRef]
  65. Wang, Y.; Li, B.; Carlson, J.; Streeck, H.; Gladden, A.; Goodman, R.; Schneidewind, A.; Power, K.; Toth, I.; Frahm, N.; et al. Protective HLA class I alleles that restrict acute phase CD8+ T cell responses are associated with viral escape mutation located in highly conserved region of human immunodeficiency virus type 1. J. Virol. 2009, 83, 1845–1855. [Google Scholar] [CrossRef]
  66. Ferrari, G.; Korber, B.; Goonetilleke, N.; Liu, M.; Turnbull, E.; Salazar-Gonzalez, J.; Hawkins, N.; Self, S.; Watson, S.; Betts, M.; et al. Relationship between functional profile of HIV-1 specific CD8 T cells and epitope variability with the selection of escape mutants in acute HIV-1 infection. PLoS Pathog. 2011, 7, e1001273. [Google Scholar] [CrossRef]
  67. Sanchez-Merino, V.; Farrow, M.; Brewster, F.; Somasundaran, M.; Luzuriaga, K. Identification and characterization of HIV-1 CD8+ T cell escape variants with impaired fitness. JID 2008, 197, 300–308. [Google Scholar] [CrossRef]
  68. Troyer, R.; McNevin, J.; Liu, Y.; Zhang, S.; Krizan, R.; Abraha, A.; Tebit, D.; Zhao, H.; Avila, S.; Lobritz, M.; et al. Variable fitness impact of HIV-1 escape mutations to cytotoxic T lymphocyte (CTL) response. PLoS Pathog. 2013, 5, e1000365. [Google Scholar]
  69. Ganusov, V.; Goonetilleke, N.; Liu, M.; Ferrari, G.; Shaw, G.; McMicheal, A.; Borrow, P.; Korber, B.; Perelson, A. Fitness costs and diversity of the cytotoxic T lymphocyte response determine the rate of CTL escape during acute and chronic phases of HIV infection. J. Virol. 2011, 85, 10518–10528. [Google Scholar] [CrossRef]
  70. Akahoshi, T.; Chikatat, T.; Tamura, Y.; Gatanaga, H.; Oka, S.; Takiguchi, M. Selection and accumulation of an HIV-1 escape mutant by three types of HIV-1 specific cytotoxic T lymphocytes recognizing wild-type and/or escape mutant epitopes. J. Virol. 2012, 86, 1971–1981. [Google Scholar] [CrossRef]
  71. Rueda, C.; Veilla, P.; Chougnet, C.; Montoya, C.; Rugeles, M. HIV-induced T-cell activation/exhaustion in rectal mucosa is controlled only partially by antiretroviral treatment. PLoS One 2012, 7, e30307. [Google Scholar]
  72. Sauce, D.; Larsen, M.; Fastenackels, S.; Pauchard, M.; Ait-Mohand, H.; Schneider, L.; Guihot, A.; Boufassa, F.; Zaunders, J.; Iguertsira, M.; et al. HIV disease progression despite suppression of viral replication is associated with exhaustion of lymphopoesis. Blood 2011, 117, 5142–5151. [Google Scholar] [CrossRef]
  73. Yamamoto, T.; Price, D.; Casazza; Ferrari, G.; Nason, M.; Chattopadhay, P.; Roederer, M.; Gostik, E.; Katsikis, P.; Douek, D.; Jaubrich, R.; et al. Surface expression patterns of negative regulator molecules identify determinants of virus specific CD8+ T cell exhaustion in HIV infection. Blood 2011, 117, 4805–4815. [Google Scholar] [CrossRef]
  74. Tomaras, G.; Yates, N.; Liu, P.; Qin, L.; Fouda, G.; Chavez, L.; Decamp, A.; Parks, R.; Ashley, V.; Lucas, J.; et al. Initial B-cell responses to transmitted human immunodeficiency virus type 1, viron-binding immunoglobulin M (IgM) and IgG antibodies followed by plasma anti-gp41 antibodies with ineffective control of initial viremia. J. Virol. 2008, 82, 12449–12463. [Google Scholar] [CrossRef]
  75. Lynch, R.; Tran, L.; Louder, M.; Schmidt, S.; Cohen, M.; DerSimonian, R.; Euler, Z.; Gray, E.; Karim, S.; Kirchherr, J.; et al. The development of CD4 binding site antibodies during HIV-1 infection. J. Virol. 2012, 86, 7588–7595. [Google Scholar] [CrossRef]
  76. Liu, P.; Overman, R.; yates, N.; Alam, M.; Vandergrift, N.; Chen, Y.; Graw, F.; Freel, S.; Kappes, J.; Ochsenbauer, C.; et al. Dynamic antibody specificities and viron concentrations in circulating immune complexes in acute to chronic HIV-1 infection. J. Virol. 2011, 85, 11196–11207. [Google Scholar] [CrossRef]
  77. Smalls-Mantey, A.; Doria-Rose, N.; Klein, R.; Patamawenu, A.; Migueles, S.; Ko, S.; Hallahan, C.; Wong, H.; Liu, B.; You, L.; et al. Antibody-Dependent cellular cytotoxicity against primary HIV-infected CD4+ T cells is directly associated with the magnitude of surface IgG binding. J. Virol. 2012, 86, 8672–8680. [Google Scholar] [CrossRef]
  78. Mikell, I.; Sather, D.; Kalams, S.; Altfeld, M.; Alter, G.; Stamatatos, L. Characteristics of the earliest cross-neutralizing antibody response to HIV-1. PLoS Pathog. 2011, 7, e1001251. [Google Scholar] [CrossRef]
  79. Euler, Z.; Gils, M.; Bunnik, E.; Phung, P.; Schweighardt, B.; Wrin, T.; Schuitemaker, H. Cross-Reactive neutralizing humoral immunity does ot protect for HIV type 1 disease progression. J. Infect. Dis. 2010, 201, 1045–1053. [Google Scholar] [CrossRef]
  80. Salado, M.; Rallon, N.; Rodes, B.; Lopez, M.; Soriano, V.; Benito, J. Long-Term non-progressors display a greater number of Th17 cells than HIV-infected typical progressors. Clin. Immunol. 2011, 139, 110–114. [Google Scholar] [CrossRef]
  81. Li, D.; Chen, J.; Jia, M.; Hong, K.; Ruan, Y.; Liang, H.; Zhang, X.; Zhao, H.; Peng, H.; Ma, P.; et al. Loss of balance between T helper type 17 and regulatory T cells in chronic human immunodeficiency virus infection. Clin. Exp. Immunol. 2011, 165, 363–371. [Google Scholar] [CrossRef]
  82. Kwon, D.; Angin, M.; Hongo, T.; Law, K.; Johnson, J.; Porichis, F.; Hart, M.; Pavlik, D.; Tighe, D.; Kavanagh, D.; et al. CD4+ CD25+ regulatory T cell impair HIV-1 specific CD4 T cell responses by upregulating interlukin-10 production in monocytes. J. Virol. 2012, 86, 6586–6594. [Google Scholar] [CrossRef]
  83. Card, C.; McLaren, P.; Wachihi, C.; Kimani, J.; Plummer, F.; Fowke, K. Decreased immune activation in resistance to HIV-1 infection is associated with an elevated frequency of CD4+CD25+FOXP3+ regulatory T cells. J. Infect. Dis. 2009, 199, 1318–1322. [Google Scholar] [CrossRef]
  84. Prendergast, A.; Prado, J.; Kang, U.; Chen, F.; Riddell, L.; Luzzi, G.; Goulder, P.; Klenerman, P. HIV-1 infection is characterized by profound depletion of CD161+ Th17 cells and gradual decline in regulator T cells. AIDS 2010, 24, 491–502. [Google Scholar] [CrossRef]
  85. Blacklaws, B. Small ruminant lentiviruses: Immunopathogenesis of visna-maedi and caprine arthritis encephalitis virus. Comp. Immunol. Microbiol. Infect. Dis. 2012, 35, 259–269. [Google Scholar]
  86. Jauregui, P.; Crespo, H.; Glaria, I.; Lujan, L.; Conteras, A.; Rosati, S.; de Andres, D.; Amorena, B.; Towers, G.; Reina, R. Ovine TRIM5α can restrict visna/maedi virus. J. Virol. 2012, 86, 9504–9509. [Google Scholar] [CrossRef]
  87. LaRue, R.; Jónsson, S.; Silverstein, K.; Lajoie, M.; Bertrand, D.; El-Mabrouk, N.; Hotzel, I.; Andresdottir, V.; Smith, T.; Harris, R. The artiodactyl APOBEC3 innate immune repertoire shows evidence for a multi-functional domain organization that existed in the ancestor of placental mammals. BMC Mol. Biol. 2008, 9, 104. [Google Scholar] [CrossRef]
  88. Arnaud, F.; Black, S.; Murphy, L.; Griffiths, D.; Neil, S.; Spencer, T.; Palmarini, M. Interplay between ovine bone marrow stromal cell antigen 2/tetherin and endogenous retroviruses. J. Virol. 2010, 84, 4415–4425. [Google Scholar] [CrossRef]
  89. Singh, I.; McConnell, I.; Dalziel, R.; Blacklaws, B. Serum containing ovine IgG2 antibody specific for maedi visna virus envelope glycoprotein mediates antibody dependent cellular cytotoxicity. Vet. Immunol. Immunopathol. 2006, 113, 357–366. [Google Scholar] [CrossRef]
  90. Kaba, J.; Winnicka, A.; Zaleska, M.; Nowicki, M.; Bagnicka, E. Influence of chronic caprine arthritis encephalitis virus infection on the population of peripheral blood leukocytes. Pol. J. Vet. Sci. 2011, 14, 585–590. [Google Scholar]
  91. Jolly, P.; Gangpoadhyay, A.; Chen, S.; Gopal Reddy, P.; Weiss, H.; Sapp, W. Changes in the leukocyte phenotype profile of goats infected with the caprine arthritis encephalitis virus. Vet. Immunol. Immunopathol. 1997, 56, 97–106. [Google Scholar] [CrossRef]
  92. Ponti, W.; Paape, M.; Bronzo, V.; Pisoni, G.; Pollera, C.; Moroni, P. Phenotypic alteration of blood and milk leukocytes in goats naturally infected with caprine arthritis-encephalitis virus (CAEV). Small Rumin. Res. 2008, 78, 176–180. [Google Scholar] [CrossRef]
  93. Reina, R.; Barbezange, C.; Niesalla, H.; de Andres, X.; Arnarson, H.; Biescas, E.; Mazzei, M.; Fraisier, C.; McNeilly, T.; Liu, C.; et al. Mucosal immunization against ovine lentiviruses using PEI-DNA complexes and modified vaccine Ankara encoding the gag and/or env genes. Vaccine 2008, 26, 4494–4505. [Google Scholar]
  94. Cheevers, W.; Hotzel, I.; Beyer, J.; Kumpula-McWhirter, N. Immune response to caprine arthritis-encephalitis virus surface protein induced by coimmuization with recombinant vaccine viruses expressing the caprine arthritis-encephalitis virus envelope gene and caprine interlukin-12. Vaccine 2000, 18, 2494–2503. [Google Scholar]
  95. Cheevers, W.; Hotzel, I. Plasmid DNA encoding caprine interferon gamma inhibits antibody response to caprine arthritis encephalitis virus (CAEV) surface protein encoded by a co-administered plasmid expressing CAEV env and tat genes. Vaccine 2001, 19, 3209–3215. [Google Scholar] [CrossRef]
  96. Perry, L.; Wilkerson, M.; Hullinger, G.; Cheevers, W. Depressed CD4+ T lymphocyte proliferative response and enhanced antibody response to viral antigen in chronic lentivirus-induced arthritis. J. Infect. Dis. 1995, 171, 328–334. [Google Scholar] [CrossRef]
  97. Reina, R.; Glaria, I.; Benavides, J.; de Andrés, X.; Crespo, H.; Solano, C.; Perez, V.; lujan, L.; Perez, M.; Perez, J.; et al. Association of CD80 and CD86 expression levels with disease status of Visna/Maedi virus infected sheep. 2007. Viral Immunol. 2007, 20, 609–622. [Google Scholar]
  98. Murphy, B.; Hotzel, I.; Jasmer, D.; Davis, W.; Knowles, D. TNFα and GM-CSF-induced activation of the CAEV promoter is independent of AP-1. Virology 2006, 352, 188–199. [Google Scholar] [CrossRef]
  99. Murphy, B.; Jasmer, D.; Stephen, W.; Knowles, D. Localization of a TNF-activated transcpriton site and interactions with the gamma activated site within the CAEV U3 70 base pair repeat. Virology 2007, 634, 196–207. [Google Scholar]
  100. Murphy, B.; Hillman, C.; Castillo, D.; Vapniarsky, N.; Rowe, J. The presence or absence of the gamma-mediated transcriptional activation in CAEV prometers cloned from the mammary gland and joint synovium of a single CAEV-infected goat. Virus Res. 2012, 163, 537–545. [Google Scholar] [CrossRef]
  101. Juganaru, M.; Reina, R.; Grego, E.; Profiti, M.; Rosati, S. LTR promoter activity of SRLV genotype E, strain Roccaverano. Vet. Res. Commun. 2010, 34, 47–51. [Google Scholar] [CrossRef]
  102. Bertoni, G.; Hertig, C.; Zahno, M.; Vogt, H.; Dufour, S.; Cordano, P.; Peterhans, E.; Cheevers, W.; Sonigo, P.; Pancino, G. B-Cell epitopes of the envelope glycoprotein of caprine arthritis encephalitis virus and antibody response in infected goats. J. Gen. Virol. 2000, 81, 2929–2940. [Google Scholar]
  103. Rachid, A.; Croise, B.; Russo, P.; Vignoni, M.; Lacerenza, D.; Rosati, S.; Kuzmak, J.; Valas, S. Diverse host-pathogen interactions following caprine arthritis encephalitis virus infection in sheep and goats. J. Gen. Virol. 2013, 94, 634–642. [Google Scholar] [CrossRef]
  104. Narayan, O.; Clements, J.; Griffin, D.; Wolinsky, J. Neutralizing antibody spectrum determines the antigenic profiles of emerging mutants of visna virus. Infect. Immun. 1891, 32, 1045–1050. [Google Scholar]
  105. Narayan, O.; Griffin, D.; Clements, J. Virus mutation during slow infection: Temporal development and characterization of mutants of visna virus recovered form sheep. J. Gen. Virol. 1978, 41, 343–352. [Google Scholar] [CrossRef]
  106. Haflidadóttir, B.; Matthíasdóttir, S.; Agnarsdóttir, G.; Torsteinsdóttir, S.; Pétursson, G.; Andrésson, Ó.; Andrésdóttir, V. Mutational analysis of a principal neutralization domain of visna/maedi virus envelope glycoprotein. J. Gen. Virol. 2008, 89, 716–721. [Google Scholar] [CrossRef]
  107. Trujillo, J.; Hotzel, K.; Snekvik, K.; Cheevers, W. Antibody response to the surface glycoprotein of caprine arthritis-encephalitis lentivirus: Disease status is predicted by SU antibody isotype. Virology 2004, 325, 129–136. [Google Scholar] [CrossRef]
  108. González, B.; Reina, R.; García, I.; Andrés, S.; Glaria, I.; Alzueta, M.; Mora, M.; Jugo, B.; Arrieta-Aguirre, A.; Perez, J.; et al. Mucosal immunization of sheep with a Maedi-Visna virus (MVV) env DNA vaccine protects against early MVV productive infection. Vaccine 2005, 23, 4342–4352. [Google Scholar]
  109. Pérez, M.; Biescas, E.; Reina, R.; Glaria, I.; Marín, B.; Marquina, A.; Salazar, E.; Alvarez, N.; de Andres, D.; Fantova, E.; et al. Small ruminant lentivirus–induced arthritis clinicopathologic findings in sheep infected by a highly replicative SRLV B2 genotype. Vet. Pathol. Online 2014. [Google Scholar] [CrossRef]
  110. Bodungen, U.; Lechner, F.; Pifster, H.; Vogt, H.; Cheevers, W.; Bertoni, T.; Jungi, W.; Peterhans, E. Immunohistology of the early course of lentivirus-induced arthritis. Clin. Exp. Immunol. 1998, 111, 384–390. [Google Scholar] [CrossRef]
  111. Lechner, F.; Machado, J.; Bertoni, G.; Seow, H.; Dobbelaere, D.; Peterhans, E. Caprine arthritis encephalitis virus disregulates the expression of cytokines in macrophages. J. Virol. 1997, 71, 7488–7497. [Google Scholar]
  112. Zhang, Z.; Harkiss, G.; Hopkins, J.; Woodall, C. Granulocyte macrophage colony stimulating factor is elevated in alveolar macrophages from sheep naturally infected with maedi-visna virus and stimulates maedi-visna virus replication in macrophages in vitro. Clin. Exp. Immunol. 2002, 129, 240–246. [Google Scholar] [CrossRef]
  113. Crespo, H.; Bertolotti, L.; Juganaru, M.; Glaria, I.; de Andrés, D.; Amorena, B.; Rosati, S.; Reina, R. Small ruminant macrophage polarization may play a pivotal role on lentiviral infection. Vet. Res. 2013, 44, 83. [Google Scholar] [CrossRef]
  114. Fluri, A.; Nenci, C.; Zahno, M.; Vogt, H.; Charan, S.; Busato, A.; Pancino, G.; Peterhans, E.; Obexer-Ruff, G.; Bertoni, G. The MHC-haplotype influence primary, but not memory, immune responses to an immunodominant peptide containing T- and B-cell epitopes of the caprine arthritis encephalitis virus gag protein. Vaccine 2006, 24, 597–606. [Google Scholar]
  115. Pyrah, I.; Watt, N. Immunohistological study of the depressed cutaneous DTH response in sheep naturally infected with an ovine lentivirus (Maedi-Visna virus). Clin. Exp. Immunol. 1996, 104, 32–36. [Google Scholar]
  116. Kuniholm, M.; Gao, X.; Xue, X.; Kovacs, A.; Anastos, K.; Marti, D.; Greenblatt, R.; Cohen, M.; Minkoff, H.; Gange, S.; et al. Human leukocyte antigen genotype and risk of HIV disease progression before and after initiation of antiretroviral therapy. J. Virol. 2011, 85, 10826–10833. [Google Scholar] [CrossRef]
  117. Naruto, T.; Gatanaga, H.; Nelson, G.; Sakai, K.; Carrington, M.; Oka, S.; Takiguchi, M. HLA class I-mediated control of HIV-1 in the Japanese population, in which the protective HLA-B*57 and HLA-B*27 alleles are absent. J. Virol. 2012, 86, 10870–10872. [Google Scholar] [CrossRef]
  118. An, P.; Johnson, R.; Phair, J.; Kirk, G.D.; Yu, X.F.; Donfield, S.; Buchbinder, S.; Goedert, J.; Winkler, C.A. APOBEC3B deletion and risk of HIV-1 acquisition. J. Infect. Dis. 2009, 200, 1054–1058. [Google Scholar] [CrossRef]
  119. Marmor, M.; Sheppard, H.; Donell, D.; Bozeman, S.; Celum, C.; Buchbinder, S.; Koblin, B.; Seage, G. Homozygous and heterozygous CCR5-Δ32 genotypes are associated with resistance to HIV infection. J. Acquir. Immunodefic. Syndr. 2001, 27, 472–481. [Google Scholar] [CrossRef]
  120. Novembre, J.; Galvani, A.; Slatkin, M. The geographic spread of CCR5 Δ32 HIV-resistance allele. PLoS Biol. 2005, 3, e339. [Google Scholar] [CrossRef]
  121. Mikula, I.; Bhide, M.; Pastorekova, S.; Mikula, I. Characterization of ovine TLR7 and TLR8 protein coding regions, detection of mutations and Maedi Visna virus infection. Vet. Immunol. Immunopathol. 2010, 138, 51–59. [Google Scholar] [CrossRef]
  122. Larruskain, A.; Bernales, I.; Luján, L.; de Andrés, D.; Amorena, B.; Jugo, B.M. Expression analysis of 13 ovine immune response candidate genes in Visna/Maedi disease progression. Comp. Immunol. Microbiol. Infect. Dis. 2013, 36, 405–413. [Google Scholar]
  123. Bowles, D.; Carson, A.; Isaac, P. Genetic distinctiveness of the herdwick sheep breed and two other locally adapted hill breeds of the UK. PLoS One 2014, 9, e87823. [Google Scholar] [CrossRef]
  124. White, S.; Mousel, M.; Reynolds, J.; Lewis, G.; Herrmann-Hoesing, L. Common promoter deletion is associated with 3.9-fold differential transcription of ovine CCR5 and reduced proviral level of ovine progressive pneumonia virus. Anim. Gen. 2009, 40, 583–589. [Google Scholar] [CrossRef]
  125. Brajon, G.; Mandas, D.; Liciardi, M.; Taccori, F.; Meloni, M.; Corrias, F.; Montaldo, C.; Coghe, F.; Casciari, C.; Giammarioli, M.; et al. Development and field testing of a real-time PCR assayfor caprine arthritis encephalitis virus (CAEV). Open Virol. J. 2012, 6, 82–90. [Google Scholar]
  126. Herrmann-Hoesing, L.; Broughton-Neiswanger, L.; Gouine, K.; White, S.; Mousel, M.; Lewis, G.; Marshall, K.; Knowles, D. Evaluation of caprine arthritis encephalitis virus/maedi-visna virus indirect enzyme-linked immunosorbent assay in the serological diagnosis of ovine progressive pneumonia virus in U.S. sheep. Clin. Vacc. Immunol. 2010, 17, 307–310. [Google Scholar]
  127. Santry, L.A.; de Jong, J.; Gold, A.C.; Walsh, S.R.; Menzies, P.I.; Wootton, S.K. Genetic characterization of small ruminant lentiviruses circulating in naturally infected sheep and goats in Ontario, Canada. Virus Res. 2013, 175, 30–44. [Google Scholar] [CrossRef]
  128. Heaton, M.; Clawson, M.; Chitko-McKown, C.; Leymaster, K.; Smith, T.; Harhay, G.; White, S.; Herrmann-Hoesing, L.; Mousel, M.; Lewis, G.; et al. Reduced lentivirus susceptibility in sheep with TEMEM154 mutations. PLoS Gen. 2012, 8, 1–12. [Google Scholar]
  129. White, S.; Mousel, M.; Herrmann-Hoesing, L.; Reynolds, J.; Leymaster, K.; Neibergs, H.; Lewis, G.; Knowles, D. Genome-Wide association identifies multiple genomic regions associated with susceptibility to and the control of ovine lentivirus. PLoS One 2012, 7, e47829. [Google Scholar] [CrossRef]
  130. Petrovski, S.; Fellay, J.; Shianna, K.; Carpenetti, N.; Kumwenda, J.; Kamanga, G.; Kamwendo, D.; Letvin, N.; McMicheal, A.; Haynes, B.; et al. Common human genetic variants and HIV-1 susceptibility: A genome-wide survey in a homogenous African population. AIDS 2011, 25, 513–518. [Google Scholar] [CrossRef]
  131. Luo, M.; Sainsbury, J.; Tuff, J.; Lacap, P.; Yuan, X.; Hirbod, T.; Kimani, J.; Wachihi, C.; Ramdahin, S.; Bielawny, T.; et al. A genetic polymorphism of FREM1 is associated with resistance against HIV infection in the Pumwani sex worker cohort. J. Virol. 2012, 86, 11899–11905. [Google Scholar] [CrossRef]
  132. Bishop, S. Possibilities to breed for resistance to nematode parasite infections in small ruminants in tropical production systems. Animal 2012, 6, 741–747. [Google Scholar] [CrossRef]
  133. Thompson-Crispi, K.; University of Guelph, Department of Pathobiology, Guelph, Ontario, Canada. Personal Communication 2013.

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Stonos, N.; Wootton, S.K.; Karrow, N. Immunogenetics of Small Ruminant Lentiviral Infections. Viruses 2014, 6, 3311-3333.

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Stonos N, Wootton SK, Karrow N. Immunogenetics of Small Ruminant Lentiviral Infections. Viruses. 2014; 6(8):3311-3333.

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Stonos, Nancy, Sarah K. Wootton, and Niel Karrow. 2014. "Immunogenetics of Small Ruminant Lentiviral Infections" Viruses 6, no. 8: 3311-3333.

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