Introduction
Simian immunodeficiency virus (SIV) infection in rhesus macaques mimics HIV infection in humans and is considered the best experimental system to study AIDS. Most SIV-infected monkeys develop a prolonged asymptomatic period followed by immunodeficiency, as in humans. Another subset of infected monkeys and humans control the virus for a protracted time. A relatively low percentage of HIV-infected humans develop rapid disease course, but the rate of rapid progression can be higher in some populations [1]. In SIV-infected monkeys, rapid progression (within 6 months of infection) to AIDS happens in approximately 25% of the animals. These animals notably have high viremia levels, poor antiviral antibody response and encephalitis [2-4].
In SIV-infected monkeys rapidly progressing to disease, the development of fatal immunodeficiency has been linked to insufficient production of tissue-homing memory CD4+ T cell [5,6]. Importantly, a subsequent study in chronically SIV-infected monkeys with normal progression revealed that such deficits in memory CD4+ T cells were a common pathway to AIDS [6].
The role of the CD8 T-cell response in CD4 deficits has not been examined. A crucial role for CD8 cells in controlling initial viremia and disease course has been well documented in macaques [7-9]. However, there are only a few reports on the role of CD8 cells, specifically cytotoxic T lymphocytes (CTLs), in the rapid progression of SIV to HIV. In monkeys, assessment of rapid progressors revealed poor CTL response to SIV [10-12]. Similarly, a study of HIV-infected individuals with rapid progression showed a failure to develop initial CTL response [13,14]. Two other studies identified initial development of CTLs in rapidly progressive HIV infection, which become functionally defective, independently of escape [15,16].
CTL reactivity to SIV has been best studied in monkeys bearing the MamuA*01 class I major histocompatibility complex (MHC) allele, which restricts immunodominant epitopes in the Gag (Gag181-189; CM9) [17] and Tat (Tat28-35; SL8) [18] proteins of SIV, allowing the study of epitope-specific CD8+ T-cell frequencies by staining with MHC-tetramer-peptide complexes [19]. In early infection, reactivity to these two epitopes comprises approximately 50% of the anti-SIV CTL response. Tat-specific cells are typically abundant at early phases but soon disappear, whereas Gag-specific cells are generally maintained for long periods [20].
In this study, we first analyze the peripheral kinetics and frequency of these immunodominant CTLs in rapid progressors and compare these with that we recently reported in regular progressors [21]. We observed that rapid progressors suffer a collapse of the anti-Gag and Tat CD8 response, accompanied by a decrease in both memory and activated CD8+ T cells. Loss of both memory and activated CD4 T cells was also found in these animals. In order to assess the relationship of the CD8 deficits with those in the CD4 compartment, we examined the experimental collapse of CD8 response in SIV-infected monkeys by using a CD8 cell-depleting antibody. This also led to losses of memory and activated CD4 cell. We propose an alternative model to explain rapid progression in which a collapse of CD8 responsiveness could lead to relatively uncontrolled virus replication, potentially driving the disease and eventual failure in maintaining the memory CD4 cell pool.
Material and methods
Animals and infection
Rhesus macaques, purchased from laboratories of Virginia and Charles River Laboratory, were free of SRV-type D and herpes B virus and infected with a SIVmac251-derived cell-free SIV stock [22]. Animal experiments were performed with Institutional Animal Care and Use Committee (IACUC) approval following National Institute of Health (NIH) guidelines. Blood was drawn from the femoral vein of ketamine-anesthetized animals into EDTA-treated vials. At necropsy, terminally anesthetized animals were intracardially perfused with sterile phosphate-buffered saline (PBS) containing 1 U/ml heparin.
Viral quantitation
Plasma and brain SIV RNA was calculated using quantitative branched DNA (bDNA) signal amplification performed by Siemens Clinical Laboratory (Emeryville, California, USA).
Cells
Cells separated by blood centrifugation, and 70 μm nylon mesh sieved cells from spleen, lymph nodes, and liver, were subjected to Histopaque (Sigma-Aldrich, St Louis, Missouri, USA) gradient for isolation of mononuclear fraction. The brain was removed and freed of meninges. Brain immune cells were isolated as described [23].
Major histocompatibility complex genotyping
Splenic cells were used for DNA extraction, using QIAamp DNA Mini Kit (Qiagen, Valencia, California, USA). Samples were assessed using a PCR-based class I MHC genotyping, at the University of Wisconsin AIDS Vaccine Research Laboratory. The class II genotyping was performed as described [24].
Viral sequencing
Plasma was ultracentrifuged and viral RNA was obtained from the pellet, cDNA synthesized and sequences corresponding to the tat and gag genes determined as described [21].
Flow cytometry
Cells were labeled with antibodies in PBS containing 2% fetal calf serum (FCS) and 0.01% NaN3. The antibodies used were antimonkey CD3-biotin (clone FN-18; Invitrogen Biosource, Carlsbad, California, USA) followed by Streptavidin-PerCP or APC (BD Pharmingen, San Diego, California, USA); remaining antibodies were antihuman CD8-PE, fluorescein isothiocyanate (FITC) or PeCy5 (clone DK25; Dako, Carpinteria, California, USA), CD95-FITC, Ki67-FITC (Dako), CCR5-APC (BD Pharmingen), CD11a-FITC (clone 25.3.1; Immunotech), CCR5-APC, CD122-PE, CD25-PE, interferon γ phycoerythrin (IFNγ-PE), tumor necrosis factor α (TNFα), interleukin 2 phycoerythrin (IL2-PE) or isotype controls (BD Pharmingen). PE-labeled Tat and Gag tetramers (Beckman-Coulter, Fullerton, California, USA) were employed for specific CTL detection. Cells from SIV-infected non-MamuA*01, or uninfected MamuA*01 animals, were used as controls. For Ki67 and intracellular cytokine staining, cells were washed with fluorescence activated cell sorter (FACS) lysis buffer (BD Pharmingen), fixed with 3% paraformaldehyde and permeabilized with 0.3% Triton X-100. After incubation with antibody, cells were resuspended in 3% paraformaldehyde. Prior to the anticytokine antibody staining, cells were treated with 10 ng/ml paramethoxyamphetamine (PMA; Sigma, St Louis, Missouri, USA), 200 ng/ml ionomycin (Sigma) and 10 μg/ml brefeldin-A (BFA; Sigma), and incubated for 4 h. Stained cells were acquired in a FACSCalibur (BD Biosciences, San Jose, California, USA) and analyzed by FlowJo6.2.1 software (Tree Star Inc., San Carlos, California, USA).
CD8 depletion
SIV-inoculated, non-MamuA*01 monkeys were transiently depleted of CD8+ lymphocytes by administration of 10 mg/kg of body weight of antihuman CD8 monoclonal antibody (MAb), cM-T807, subcutaneously on day 6 postinoculation and 5 mg/kg intravenously on days 9 and 13, as reported earlier [7].
Results
Initial development and early loss of cytotoxic T lymphocytes in rapid progressors
We identified eight MamuA*01 animals in our studies: five had a regular disease course and three had rapidly progressive disease. In addition to MamuA*01, animals were typed for other disease progression-correlated MHC alleles [24]. However there was not a distinctive pattern of Mamu MHC class I or class II alleles associated with rapid progression (not shown). Clinically, the three MamuA*01-positive animals with rapid disease progression (RAP) showed wasting syndrome and symptoms of neurological disease. Terminal neuropathology revealed SIV encephalitis. We assessed Gag and Tat epitope reactivity using tetramers longitudinally in the blood and at necropsy in lymphoid and nonlymphoid organs in two of the animals (417 and 418) with rapid progression; the third animal (228) had been studied before these reagents were available. These data were compared with that found in four recently reported MamuA*01 regular progressors (REGs; 404, 406, 409 and 414; the fifth animal, 324, had been studied before these reagents were available). These were sacrificed following 6 months of infection in the absence of AIDS-like disease.
Following the acute stage, RAP animals 417 and 418 did not control viremia (Fig. 1a). At preinoculation and in the postacute phase, RAP animals tended to have higher number of blood lymphocytes (Fig. 1b). The percentage of CD4+ cells did not differ significantly over time between REG and RAP animals (not shown). However, CD4+ T-cell numbers were increased in RAP animals compared with that in the REG group (Fig. 1c). The percentage of CD8+ cells peaked acutely in REG animals but not in individual RAP monkeys (not shown). In absolute numbers, blood CD8+ T cells increased acutely (days 14 and 28 postinoculation, averaging three times more than their preinfection level) in the REG animals following infection, but not in the RAP animals, relative to their preinfection level (Fig. 1d). The absence of an acute increase in the proportion of CD8 cells in RAP animals suggests the importance of the CD8 cells in the control of the initial viremia.
Tetramer staining was used to evaluate the contribution of the SIV-specific CTL response. Early after inoculation, both Tat-reactive and Gag-reactive cells were found in the blood of both RAP and REG animals. As previously described, in both groups, Tat-specific cells disappeared from the blood soon after the acute phase (Fig. 1e). However, Gag-reactive cells, which are maintained in REG animals and dominant in chronically SIV-infected MamuA*01 animals [20], disappeared from the blood in RAP animals (Fig. 1f). Thus, there is a distinct extinction of virus-specific CD8 cells in animals with rapid progression.
Epitope escape has been suggested as a mechanism for virus-specific CTL decay, particularly within the Tat epitope in MamuA*01 animals with regular progression. Differences in sequence encoding Tat and Gag epitopes were assessed in viral genomic RNA isolated from plasma. All clones from animal 418 had escape mutations within the Tat epitope; the most frequent Tat SL8 epitope mutations were located at positions 26 (Ala to Asp) and 28 (Ser to Pro or Thr). However, in the plasma of animal 417, the virus did not exhibit escape mutations either in Tat or its flanking regions. In a similar manner, regardless of the degree of epitope escape, Tat-reactive cells disappear from the blood in REG animals [18]. In all the animals, no mutations were detected within the Gag epitope and flanking regions. Overall, although in animal 418 the CTLs were efficacious in their ability to select for escape mutants within Tat, the disappearance in CTLs in the RAP animals could not be explained by viral escape.
Global CD8 T-cell alterations in rapid progressors
The proliferative status of CD8 and CD4 T cells was assessed by the frequency of cells expressing Ki67, a nuclear marker of cell cycle, at acute phase and necropsy. Although the groups did not differ in the percentage of CD4 cells expressing Ki67 (not shown), the number of Ki67+ CD4 T cells decreased in both RAP animals and was relatively unchanged in the REG animals (Fig. 1g). In addition, following infection, REG animals had increased proportion (not shown) as well as numbers of Ki67+ CD8 T cells relative to RAP animals (Fig. 1h), suggesting that a failure to respond to SIV infection with T-cell proliferation is a hallmark of rapid progression.
In addition to the disappearance of CD8 cells reactive to the two major MamuA*01-restricted epitopes in RAP animals, a notable decrease in the surface expression of the chemokine receptor CCR5 and the activation marker/adhesion molecule CD11a/LFA-1 occurred in the total CD8 compartment (Fig. 2a and b). The lower expression of CCR5 that characterized RAP animals was observed also on the tetramer-positive fraction of CD8+ cells, especially early after infection (day 10) and as the cells have almost disappeared from the circulation (day 59) (Fig. 2c-f). Together, these results suggest that the loss of the immunodominant CTL in rapid progressors occurs in correlation with decreased expression of functionally relevant surface molecules on the totality of CD8 cells.
Rapid progressors lack cytotoxic T lymphocytes outside of the brain
The disappearance of virus-specific CTLs from the blood could be related to tissue demand and migration. This possibility was assessed by the quantification of T cells and tetramer-positive CTLs in various organs at necropsy. In the blood of RAP animals, both relative (Fig. 3a) and absolute (not shown) CD4 T-cell numbers were higher than those found in the REG group. In addition, an increased percentage of CD4 cells were observed in the liver of RAP animals in comparison with REG animals, but not in the spleen or the brain (the latter contained very few CD4+ cells in all animals) (Fig. 3a). Neither the percentage of CD8 cells in all sites (Fig. 3b) nor the absolute numbers of these cells in the blood differed between RAP and REG groups.
The CTL specificity within CD8+ T cells was then determined in the organs. Neither Tat nor Gag CTLs were found in the organs of RAP monkeys at necropsy, except for the brain (Fig. 3c and d). Such findings were qualitatively similar to our report of Tat-reactive cells in the central nervous system (CNS), but not elsewhere, in REG monkeys [21]. Here, RAP monkeys also did not have Gag-specific cells in peripheral sites but in the brain, similar to Tat-specific cells. Thus, virus-specific CD8 cells, which disappeared from the periphery of RAP animals, could be found in the CNS. However, although REG and RAP monkeys had similar levels of CD8+ T cells in the brain (Fig. 3b), in REG animals, Tat-specific and Gag-specific cells represented over one-third of the CD8 T cells in the brain [21], whereas in RAP animals, both specificities together were less than one-tenth of the CD8 T cells (Fig. 3c and d).
Assessment of brain viral load revealed a likely consequence of this decrease in brain CTLs. Each RAP animal had over 10 000-fold higher levels of virus in brain (frontal lobe) than did REG animals (not shown). Therefore, although the antiviral-specific response is still present in the brain, it is insufficient to control the virus.
The deficiencies in the CD8 compartment in RAP animals were further evidenced by a lower percentage of cells expressing CD25 (IL2 receptor α chain) and CD122 (IL2 receptor β chain) in terminal blood (Fig. 3e), suggesting a poor responsiveness to stimulation through IL2 and IL15. In addition, the ability of blood CD8 T cells to produce IFNγ, TNFα and IL2 was determined both at the peak of CD8 cell increase in REG monkeys (28 days; Fig. 1d) and at termination. A lower percentage of cells from RAP animals produced the proinflammatory cytokines IFNγ and TNFα (Fig. 3f). In both cases, the difference was greater at the terminal point than at 28 days postinoculation. Together, these data indicate that the CD8 T-cell population in RAP animals is deficient in several functional aspects following SIV infection.
CD8 T-cell deficits parallel those found in CD4 T cells
Despite the relative abundance of the total CD4 cell compartment, RAP animals had a lower percentage of CD4 cells expressing CD95, a correlate of memory phenotype in macaques, than did REG animals in the blood, spleen (Fig. 4a) and other sites such as lymph nodes (not shown), as demonstrated on FACS histograms (Fig. 4b). Moreover, a decrease in CD4 cells expressing high CD11a was observed in the organs (Fig. 4c and d). Interestingly, longitudinal analysis revealed that the decrease in CD11a on CD4 cell surface occurred early in both RAP animals (Fig. 4e and f). This was also detectable in lymph nodes, in both MamuA*01 and other non-MamuA*01 animals (Fig. 4g).
CD8 collapse can drive the characteristic CD4 alterations
We hypothesized that the decrease in memory CD4 cells in SIV-infected monkeys may be secondary to intrinsic defects in the CD8 compartment. To assess this hypothesis, we examined the phenotype of CD4 cells in monkeys (not carrying the MamuA*01 allele), which were depleted of CD8 cells (by anti-CD8 MAb) to mimic the physiological collapse in the CD8 population. Indeed, these animals developed a drastic fall in CD11a-high CD4+ cells. Upon incomplete recovery of CD8 levels, the CD11a expression on CD4+ cells exhibited a partial regain before diminishing again (Fig. 5a). In contrast, mock-depleted animals did not manifest this compromise of CD4 cells (Fig. 5b). Similar to the RAP group, as reported earlier, CD8-depleted animals were unable to control viral load (Fig. 5c), quickly developed disease and exhibited encephalitis at sacrifice (93-111 days postinoculation). At necropsy, CD8-depleted animals had decreased CD8 proportions in all sites compared with REG non-MamuA*01 animals (Fig. 5d). Furthermore, CD4 cells from different sites of these animals had lower CD95 levels (Fig. 5e and f) compared with CD4 cells from non-CD8-depleted non-MamuA*01 regular progressors, indicating memory CD4 cell deficiency. Similarly, in the different sites, lower levels of CD11a were found in CD8-depleted animals (Fig. 5g). Therefore, mimicking early CD8 deficits by CD8 depletion leads to CD4 changes and AIDS, suggesting that CD8 collapse can be instructive of CD4 dysfunction.
Discussion
Here we find that in rapid progressors, SIV-specific CTLs develop initially, but quickly disappear from the body, except from the CNS. In the general CD8 T-cell compartment, striking changes develop - a decrease in CCR5+ and CD11a high cells, in temporal association with a loss of virus-specific CTL, and a failure to proliferate. In addition to the characteristic decrease in memory CD4 cells, we also find a loss of CD11a+ in the CD4 population. These changes in CD4 cells are closely correlated to changes in CD8 cells in rapid progressors and inability to control viral load. We asked whether these CD4 changes could be driven by the CTL loss and the resulting high viral load. In infected animals in which CD8 cells were experimentally depleted at infection, rapid progression was accompanied by similar CD4 changes, leading us to conclude that changes in CD4 cells can result from a failure of CD8 responsiveness.
The alterations in the CD8 T-cell compartment were beyond the inability to recognize immunodominant epitopes Gag and Tat, reaching the totality of the CD8 population regarding the expression of surface molecules (CCR5 and CD11a). In addition, CD8 proliferation was lower in RAP animals than in REG animals. In the CD4 compartment, we observe a loss of memory CD4 cells, confirming previous reports [5]. In addition, there is a loss of CD4 cells expressing high CD11a levels.
These experiments reveal a correlation between rapid progression in SIV-infected animals and a failure to enrich and maintain major antiviral CTL. Others have reported dysfunctions in the CD4 cell population in rapid progressors as related to defects in memory replenishment [5] as well CD8 T cells [11]. Although these facts are interconnected, the instructive events were never well clarified. Our findings reveal that the wheel of rapid progression may reside primarily in the CD8 compartment.
The expression of CD95 and CD11a on CD4 cells was lower in the RAP group than in the REG group, confirming the correlation of a restricted availability of memory cells with the development of disease [5,25]. We determined that such findings could develop following CD8 failure, experimentally induced by CD8 cell depletion. Overall, our data indicate the connection between collapse of the epitope recognition and immunodominance patterns in CD8 cells and CD4 memory availability. The nature of the CD8 requirements during the early acute phase is not clear and may not be restricted to priming, recognition and antiviral effector activity. The outcome, in terms of CD4 changes and rapid disease, was identical in natural and experimentally induced rapid progression. Thus, the dysfunction identified by the decline in memory and activated CD8+ T cells is as profound as if such cells were eliminated.
A similar outcome was previously obtained by co-stimulatory signal blockage, which attenuates SIV-specific CTL response and decreases CD8 proliferation, leading to increased viremia and rapid progression [26]. In fact, rapid progressors failed to maintain proliferation in the whole CD8 population following acute infection. This could be due to functional exhaustion or impaired induction of CD8 cells, classically attributed to defective CD4 help [27].
The use of CD8 depletion to accelerate disease strongly suggests CD8 cells are pivotal in early viral load control. Moreover, CD8-depleted animals developed CD4 cells that phenotypically resemble those found in animals with spontaneous rapid progression. However, CD8 depletion potentially disturbs homeostatic and antigen-driven proliferation of CD4+ cells and eliminates natural killer cells (CD8+ in rhesus monkeys). Nevertheless, mathematical models that take into account the balance between virus and target cells, examining both nonmanipulated infection and animals in which co-stimulatory blockage was performed to modulate CD8 response, indicate that at early periods of infection (first month after inoculation) viral control is due to CTL, in a positive correlation either with CD8+ T cells as a whole or the Tat and Gag CTLs individually in MamuA*01 animals [28]. From our study, the CD8 functional aspects that define progression at initial time points result from the viral-host interactions, but are in fact not restricted to the virus-specific population.
As rapid progression is correlated with encephalitis, the brain findings are particularly interesting. In REG animals, the MamuA*01-restricted immunodominant anti-Tat and anti-Gag CTLs represent close to 50% of the brain accumulating CD8 cells [21]. Interestingly, in RAP animals, the brain environment still supports CD8 accumulation; however, the proportion of SIV Tat-specific and Gag-specific CD8 cells among the brain-infiltrating CD8 cells is strikingly lower. Thus, possibly, the proportion of immunodominant CTLs rather than the brain CD8 accumulation per se affects the viral control and protection from encephalitis.
The development of CD4 deficits and rapid progression to AIDS as a result of loss of control over viral load due to CD8 defects represent an alternative perspective to other concepts in which loss of CD4 memory and helper activity leads to AIDS. Our results indicate that in SIV-infected macaques, a collapse of CD8 cells can be a potential sign for the damage to the CD4 memory pool related to rapid progression. The basis of the global defect in CD8 cells should be investigated in depth and may give important clues for the development of therapeutic strategies aimed at maintaining functional CTLs. The importance of these findings resides in the out-of-the-ordinary perspective and understanding of the necessity of therapeutic approaches that focus on better virus-specific CD8 expansion, as well as on the search for mechanisms of global CD8 response regulation and biomarkers that identify the CD8 collapse related to progression.
Acknowledgements
This work was supported by NIH grants MH062261 and MH073490. We thank Drs Bianca Mothe' (California State University San Marcos) and Daniel Mucida (La Jolla Institute of Allergy and Immunology) for interesting and helpful discussions, and Nancy Delaney for administrative assistance. M.C.G.M. made the initial observation of the phenomenon described, performed experiments and wrote the manuscript; S.S. and U.S. performed class II MHC phenotyping of the monkeys and participated in the discussions, T.H.B. sequenced the viral epitopes, D.D.W. and M.Z. performed technical assistance, and H.S.F. participated in experiment planning, interpretation, discussions and writing, and obtained funding. This is manuscript 19185 of The Scripps Research Institute.
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