Immunological alterations and associated diseases in mandrills (Mandrillus sphinx) naturally co-infected with SIV and STLV

Virology 454-455 (2014) 184–196

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Immunological alterations and associated diseases in mandrills (Mandrillus sphinx) naturally co-infected with SIV and STLV Sandrine Souquière a, Maria Makuwa a, Bettina Sallé b,1, Yves Lepelletier d, Franck Mortreux e, Olivier Hermine d, Mirdad Kazanji a,c,n a

Laboratoire de Rétrovirologie, Centre International de Recherches Médicales de Franceville, Franceville, Gabon Centre de Primatologie, Centre International de Recherches Médicales de Franceville, Franceville, Gabon c Réseau International des Instituts Pasteur, Institut Pasteur, Paris, France d Département d'Hématologie, Hôpital Necker, Paris, France e Université de Lyon 1, CNRS UMR5239, Oncovirologie et Biothérapies, Faculté de Médecine Lyon Sud, Pierre Bénite, France b

art ic l e i nf o

a b s t r a c t

Article history: Received 22 November 2013 Returned to author for revisions 16 January 2014 Accepted 18 February 2014 Available online 7 March 2014

Mandrills are naturally infected with simian T-cell leukaemia virus type 1 (STLV-1) and simian immunodeficiency virus (SIV)mnd. In humans, dual infection with human immunodeficiency virus (HIV) and human T-cell lymphotropic virus type 1 (HTLV-1) may worsen their clinical outcome. We evaluated the effect of co-infection in mandrills on viral burden, changes in T-cell subsets and clinical outcome. The SIV viral load was higher in SIV-infected mandrills than in co-infected animals, whereas the STLV-1 proviral load was higher in co-infected than in mono-infected groups. Dually infected mandrills had a statistically significantly lower CD4þ T-cell count, a lower proportion of naive CD8þ T cells and a higher proportion of central memory cells. CD4 þ and CD8 þ T cells from SIV-infected animals had a lower percentage of Ki67 than those from the other groups. Co-infected monkeys had higher percentages of activated CD4 þ and CD8 þ T cells. Two co-infected mandrills with high immune activation and clonal integration of STLV provirus showed pathological manifestations (infective dermatitis and generalised scabies) rarely encountered in nonhuman primates. & 2014 Elsevier Inc. All rights reserved.

Keywords: SIV STLV Co-infection Associated diseases Viral load Cell line Immune system Mandrillus sphinx

Introduction Human T-cell lymphotropic virus type 1 (HTLV-1), the causative agent of adult T-cell leukaemia/lymphoma (Yoshida, 1983) and of tropical spastic paraparesis/HTLV-1-associated myelopathy (Gessain et al., 1985), has also been implicated in other inflammatory diseases, such as paediatric infectious dermatitis (Lagrenade et al., 1990), uveitis (Mochizuki et al., 1992), arthropathy (Ijichi et al., 1990) and polymyositis (Morgan et al., 1989). It was reported previously that coinfection with other pathogens, such as parasites (e.g. Strongloides) or human immunodeficiency virus type 1 (HIV-1), affects the development of disease (Carvalho and Da Fonseca Porto, 2004; Casoli et al., 2007). Cohort studies of people co-infected with HIV-1 and HTLV-1 showed that they are at higher risk for more rapid development of more aggressive AIDS than people infected solely with HIV, with important implications for the progression and severity of HIVrelated disease (Beilke et al., 2005; Casoli et al., 2007). It has also been reported that co-infection with HIV-1 and HTLV-1 accelerates n

Correspondence to: Institut Pasteur de Bangui, Bangui, Central African Republic. E-mail address: [email protected] (M. Kazanji). 1 Deceased.

http://dx.doi.org/10.1016/j.virol.2014.02.019 0042-6822 & 2014 Elsevier Inc. All rights reserved.

diseases associated with HTLV-1 infection (Beilke et al., 2005; Casseb et al., 2008). Simian T cell leukaemia virus type 1 (STLV-1), the simian counterpart of HTLV-1, naturally infects Old World monkeys and shares virological, immunological, molecular and pathological features with HTLV-1 (Gessain et al., 1996; Hayami et al., 1985; Saksena et al., 1994). Natural STLV-1 infection has been associated with the development of diseases related to adult T-cell leukaemia/lymphoma, in some cases with co-infection with SIV. For example, neoplastic diseases in sooty mangabeys (Fultz et al., 1997) and lymphoproliferative disease in African green monkeys (Traina-Dorge et al., 1992) were found in animals naturally coinfected with STLV-1 and SIV. Furthermore, experimental coinfection of rhesus and pigtailed macaques with the two viruses (Fultz et al., 1999; McGinn et al., 2002; Traina-Dorge et al., 2007) led to a pre-leukaemic condition in a few animals. These findings provide support for clinical observations in humans that HIV– HTLV co-infection may up-regulate cell proliferation and HTLV-1 expression and increase disease potential. We showed recently that STLV-1 infection induces immune activation and increased proliferation of T cells in mandrills, suggesting that STLV-1 infection can induce T-cell activation and

S. Souquière et al. / Virology 454-455 (2014) 184–196

transformation and that this activation might be related to a high proviral load (Souquiere et al., 2009b). These observations are the first in monkeys naturally infected with STLV-1 in the wild. There was, however, no clinical manifestation or associated illness in these naturally infected mandrills. This species of monkeys has also been shown to be naturally infected with SIVmnd (Souquiere et al., 2001; Tsujimoto et al., 1989), but the infection does not appear to induce AIDS, despite a high plasma viral load (Onanga et al., 2002; Onanga et al., 2006). Recently, a functional phenotypic study of lymphocyte populations in SIV-infected mandrills showed limited activation and preserved immune regenerative capacity (Apetrei et al., 2011). Several studies have been conducted on SIV and STLV infections in nonhuman primates (Apetrei et al., 2011; Gabet et al., 2003; Souquiere et al., 2009a; Souquiere et al., 2009b; Sumpter et al., 2007), but none has explored SIV–STLV co-infection in mandrills. Natural co-infection of mandrills with both STLV-1 and SIV is a suitable model for studying the host–virus interaction and the role of co-infection in viral dynamics, immune system disruption and disease progression. We therefore followed up, characterised and compared infections and their effects on the immune system in four populations of mandrills: STLV-infected, SIV-infected, SIV– STLV co-infected and uninfected controls.

Results

185

of STLV per 100 PBMC and mandrill A, 42.6. We have shown that STLV can infect CD4 þ and CD8 þ cells, and we therefore quantified the proviral load in these two compartments. Similar results were found in the two mandrills, which had the highest values ever recorded: mandrill T had 23.8% copies of STLV-1 in CD4 þ T cells and 35.3% in CD8 þ T cells, and mandrill A had a proviral load of 74.7% in CD4 þ T cells and 33.5% in CD8 þ cells. We next examined the effects of the dual viral burden on immunological parameters. The absolute numbers of CD4 þ T cells in blood were 224/mm3 in mandrill T and 184/mm3 in mandrill A; these values were in the normal range for the 58 uninfected mandrills. In contrast, remarkable differences were found in terms of immune activation. As shown in Table 2, activation of CD4 þ and CD8 þ T cells measured by expression of HLA-DR was 3–16 times higher than in the uninfected population. Confirmation of this result was found in the expression of CD25 in CD4 þ and CD8 þ T cells. Proliferation, evaluated from expression of Ki67, was increased only in mandrill A, but the proliferative capacity (13.5% in CD4 þ and 13.1% in CD8 þ T cells) was highly unusual. Il-2, IFN-γ, TNF-α, IL-4, IL-6 and IL-10 were quantified in plasma from these two mandrills. No significant differences from the values for 21 uninfected mandrills were observed (Table 2), except that the level of TNF-α in mandrill T, which had infective dermatitis, was three times higher than the average value in uninfected animals. It is of note that, in humans, infective dermatitis was recently associated with an increase in TNF-α (Nascimento et al., 2009).

Co-infection

Dynamics of viral loads

Four groups of mandrills were selected according to the serological results (Table 1): a control group of 58 mandrills uninfected with SIV or STLV; 38 mandrills infected with SIVmnd1 or SIVmnd-2 (SIVþ); 19 mandrills infected with STLV-1 subtype D (STLV þ); and 13 mandrills infected with both SIVmnd-1 and STLV-1 subtype D (SIVþ /STLV þ). The mean ages of each group were 10 76 years (range, 5–28) for controls, 12 75 years (3–23) for SIVþ, 12 75 years (4–19) for STLV þ and 13 75 years (6–20) for SIVþ /STLV þ.

To explore the influence of co-infection on the dynamics of the viral load, we measured the SIVmnd plasma viral load and the STLV-1 proviral load in the three infected groups, as described previously (Onanga et al., 2002; Onanga et al., 2006; Souquiere et al., 2009a; Souquiere et al., 2009c). As seen in Fig. 2A, the mean of viral load in SIVþ mandrills was somewhat higher (mean, 3.6  106 copies/ml; range, 3.7  104–2.5  107) than in SIVþ / STLV þ animals (mean, 8.7  105 copies/ml; range, 2.6  104– 2.5  106); however, this difference was not statistically significant (p ¼0.07). Quantification of the STLV-1 proviral load showed an average of 4.49 copies per 100 PBMC (range, 0.12–12.1) in the STLV þ group and 6.44 copies (range, 0.03–42.6) in SIVþ /STLV þ mandrills (Fig. 2B). The difference between the two groups was not statistically significant (p ¼0.23), but it is interesting that the highest proviral load value was found in the co-infected group. We showed previously that STLV-1 in mandrills is integrated into both CD4 þ and CD8 þ T cells (Souquiere et al., 2009a). We found no significant difference in the proviral load in CD4 þ and CD8 þ T cells in STLV þ and SIVþ /STLV þmandrills (Fig. S1).

Two co-infected mandrills showed severe immune perturbation and disease Two of the mandrills naturally co-infected with SIVmnd type 1 and STLV-1 had clinical signs that might have been related to the retroviral infections (Table 2). Mandrill T was a 12-year-old male, and, from data obtained during annual health examinations, we estimated that the duration of infection was 3 years for SIV and 5 years for STLV. Mandrill A was a 20-year-old male; the duration of infection could not be estimated, because it had been infected in the wild. Mandrill T had infective dermatitis, and mandrill A had generalised scabies (Fig. 1A and B), both conditions being resistant to treatment for several months. Blood smears showed the presence of flower cells in both animals and persistent hypereosinophilia in mandrill A (44800/mm3). In terms of virus replication, no difference from controls was found in SIV viral load, but the STLV proviral load was higher than any previously registered in mandrills: mandrill T had 22.5 copies Table 1 Distribution of mandrills in the four studied groups. Sex

Uninfected

STLV-1 infected

SIV infected

STLV/SIV co-infected

Total

Males Females Total

20 38 58

11 8 19

21 17 38

9 4 13

61 67 128

Dually infected mandrills have fewer CD4 þ T cells Although SIV and STLV infections do not appear to influence CD4 þ and CD8 þ T cells in mandrills independently, co-infection places greater pathogenic pressure on the immune system, which may modify the proportions of T cells. We evaluated the effect of retroviral infection by assessing the absolute numbers and percentages of CD4 þ and CD8 þ T cells in the four groups. As shown in Fig. 3A, the percentage of CD4 þ T cells in SIVþ /STLV þ animals (average, 11 74.7%) was significantly lower than that in the control group (21.77 6.2%; po 0.01), in the SIVþ group (17.5 79%; po 0.05) and in the STLV þ group (18.3 77.1%; p o0.01). The values for the SIVþ group were also significantly lower than in the control group. In absolute numbers, the values were significantly lower in the SIVþ /STLVþ group (mean, 2357207 cells/mm3) than in the control (4757315 cells/mm3) and the SIVþ group

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Table 2 Immunological and virological parameters of mandrill A and mandrill T compared to 58 uninfected mandrills. Mandrill T

Mandrill A

Uninfected mandrills (Mean of 58)

Age (years)/Sex Natural infections Age of infection SIV (years) Age of Infection STLV (years) SIVmnd-1 viral load (RNA copies/ml) STLV proviral load in total PBMC (%) STLV proviral load in CD4 þ T cells (%) STLV proviral load in CD8 þ T cells (%) Absolute number of CD4 þ T cells (/mm3) % HLADR in CD4 þ T cells % HLADR in CD8 þ T cells % Ki67 in CD4 þ T cells % Ki67 in CD8 þ T cells % CD25 in CD4 þ T cells % CD25 in CD8 þ T cells Cytokines Th1 (pg/ml) IL-2 IFN-γ TNF-α Cytokines Th2 (pg/ml) Il-4 Il-5 Il-6 Haematological anomaly

12/M Yes 3 5 1.3  105 22.5 23.8 35.3 224 18.6 22.6 3.7 2.6 41.9 12.3

20/M Yes Unknown Unknown 1.2  106 42.6 74.7 33.5 184 67.2 62.5 13.5 13.1 87 35.5

6.1 375 39

1.3 154 17

10.6 7 6.5 – – – – – – – 476 7315 4.2 7 2.1 8.2 7 4.2 4.3 7 2 3.6 7 2.4 20.9 7 8.2 9.6 7 5.4 Mean of 21 5.8 7 6.4 2457 243 13.3 7 18.9

2.8 2.1 7.8 Presence of flower cells

Pathology

Infective dermatitis

0.9 0.6 3.7 Persistent hyper-eosinophilia: 4 4800/mm3 presence of flower cells Scabies resistant to therapy

1.9 7 2.2 2.6 7 2.5 4.3 7 3.6 – –

Fig. 1. Clinical manifestations observed in two SIVþ /STLV þ co-infected mandrills. (A) Dermatitis was observed in mandrill T and (B) scabies in mandrill A.

(5007372 cells/mm3) (po0.001) (Fig. 3B). In all groups, some mandrills had an absolute CD4 þ T cell numbero50/mm3. Significantly higher percentages of CD8 þ T cells (Fig. 3C) were found in SIVþ (average, 3379.4%) and STLVþ groups (32.776.3%) than in the control group (28.377.9%; po0.05). Evaluation of the absolute numbers of CD8 þ T cells confirmed that the highest number was in SIVþ mandrills (8987555 cells/mm3), which was significantly higher than in the control group (5897354 cells/mm3) and the STLVþ group (5617285 cells/mm3) (po0.05) (Fig. 3D). No significant difference in the number of CD8 þ T cells was found in the SIVþ/STLVþ group (Fig. 3C and D).

Dual infection and circulating naive, central memory and effector memory CD4 þ and CD8 þ T cells We next sought to determine whether retroviral infection is associated with specific changes in the proportions of circulating naive, central memory and effector memory CD4 þ and CD8 þ T cells. Characterisation of these three subsets in CD4 þ and CD8 þ T cells has been reported for macaques (Pitcher et al., 2002), African green monkeys (Pandrea et al., 2006), sooty mangabeys (Sumpter et al., 2007) and mandrills (Apetrei et al., 2011). These studies indicate that the subsets can be defined from the CD28/CD95

Median 25%-75% Min-Max Data

P= 0.07

108

P= 0.23

107

106

105

10

4

10

3

187

45 STLV proviral load (% in PBMC)

SIVmnd viral load (copies RNA /ml plasma

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40 35 30 25 20 15 10 5 0

SIV+

STLV+

SIV+/STLV+

SIV+/STLV+

Fig. 2. SIV and STLV-1 loads in three groups of mandrills. (A) Plasma SIVmnd viral load in mandrills infected with SIV (□) and co-infected with STLV and SIV (◊) (RNA copies/ml). (B) STLV proviral load in PBMCs of mandrills infected with STLV (Δ) and co-infected with STLV and SIV (◊) (per 100 PBMC). Mann-Witney test used for statistical analysis of each T-cell subset.

1800

** **

% CD4+T cells

35 30

20 15 10

Uninfected

*

SIV+

1000 800 600

0

STLV+ SIV+/STLV+

*

3500

Uninfected

SIV+

**

40 30 20 10

STLV+ SIV+/STLV+

**

3000

# CD8+T cells

% CD8+T cells

1200

200

50

0

Median 25%-75% Min-Max Data

400

5

60

**

1400

**

25

0

**

1600

* # CD4+T cells

40

2500 2000 1500 1000 500

Uninfected

SIV+

STLV+ SIV+/STLV+

0

Uninfected

SIV+

STLV+ SIV+/STLV+

Fig. 3. T-cell subsets in blood from four groups of mandrills. Mandrills infected with STLV (Δ), SIV (□), STLV and SIV (◊) and uninfected (○). (A) Percentages of CD3þ CD4þ cells. (B) Absolute numbers of CD4 þ T cells. (C) Percentages of CD3þ CD8 þ cells. (D) Absolute numbers of CD4 þ T cells. Mann-Witney test used for statistical analysis of each T-cell subset. Significance assumed at p o 0.05.

pattern of staining. We found no significant differences in the proportions of naive CD4 þ T cells (Fig. 4A) or of central or effector memory cells (data not shown). SIV infection decreased the proportion of naive CD8 þ T cells below that of the control group (po0.001) (Fig. 4B). Co-infection was associated with a decrease in the proportion of naive cells (po0.001) and an increase in that of central memory cells (po0.05) (Fig. 4B and C, respectively); no difference was observed in the proportion of effector memory CD8 þ T cells (Fig. 4D). Although SIV infection appears to affect naive T cells, SIV and STLV infections do not appear to cause major changes in the distribution of the T cell repertoire. Do retroviral infections perturb the activation and proliferation of CD4 þ and CD8 þ T cells? In a previous study, we demonstrated increased activation of CD4 þ T cells in mandrills naturally infected with STLV-1

(Souquiere et al., 2009b). Others have shown that T-cell activation and proliferation do not increase significantly during chronic SIV infection in mandrills (Apetrei et al., 2011). We therefore investigated whether mono and dual retroviral infections induce T cell activation and perturb their proliferative capacity. The fractions of proliferating CD4 þ and CD8 þ T cells clearly varied significantly with infection. As seen in Fig. 5A, a lower percentage of Ki67 was found in CD4 þ T cells from SIVþ animals and higher values in the STLVþ group. Proliferation was weaker in the SIVþ group than in the other three groups (po0.01), and that in the STLVþ group was higher than in the control group (po0.05) and the SIVþ group (po0.01). SIV infection appeared to have a negative effect on CD8 þ T cells (Fig. 5B), as the value in the SIVþ group was lower than in the controls, the STLVþ group (po0.01) and also the SIVþ /STLVþ group (po0.05). However, if we exclude the two co-infected monkeys that developed diseases, the difference with the SIVþ /STLVþ group was no longer statistically significant.

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100

60

*

50

80 % CM CD8+ T cells

% Naïve CD4+ T cells

90

70 60 50 40 30 20

40 30 20 10

10 0

80

Uninfected

SIV+

STLV+

0

SIV+/STLV+

**

STLV+

SIV+/STLV+

Uninfected

SIV+

STLV+

SIV+/STLV+

80 70

60

% EM CD8+ T cells

% Naïve CD8+ T cells

SIV+

90

**

70

50 40 30 20

60 50 40 30 20

10 0

Uninfected

10 Uninfected

SIV+

STLV+

SIV+/STLV+

0

Fig. 4. Repertoire of T cells in four groups of mandrills. Mandrills infected with STLV (Δ), SIV (□), STLV and SIV (◊) and uninfected (○). (A) Percentages of naive CD4 þ T cells. (B) Percentages of central memory (CM) CD8 þ T cells. (C) Percentages of naive CD8 þ T cells. (D) Percentages of effector memory (EM) CD8þ T cells. Mann-Witney test used for statistical analysis of each T-cell subset. Significance assumed at p o 0.05.

Interestingly, we found a direct correlation between the expression of Ki67 and STLV proviral load in CD4 þ T cells in the STLV þ and SIV þ/STLV þ groups (Fig. 5C and D). Taken together, these data suggest that STLV and SIV infections have different effects on the proliferation of CD4 þ T cells. We also evaluated the activation of CD4 þ and the CD8 þ T cells using expression of HLA-DR markers. In a previous study (Souquiere et al., 2009b), we showed that natural STLV infection in mandrills is associated with an increase in the activation of CD4 þ T cells. In the present study, we found no difference in HLADR expression in CD4 þ and CD8 þ T cells in the SIVþ group, whereas the SIV þ/STLV þ group had higher percentages of HLADR in CD4 þ and CD8 þ T cells than the other groups (p o0.01) (Fig. 6A and B). As the relevance of this marker has not been established in mandrills, we also evaluated the release of neopterin to confirm immune activation. Neopterin biosynthesis is closely associated with activation of the cellular immune system, and increased concentrations have been reported in patients with viral infections (Mildvan et al., 2005; Murr et al., 2002). We found a positive correlation between the expression of HLA-DR in CD8 þ T cells and secretion of neopterin in plasma only in the SIV þ/ STLV þ group (Fig. 6C). Thus, co-infection significantly increases cell activation and proliferation in mandrills. Do dual retroviral infections modify the immune regulation of T cells? One of the characteristics of adult T-cell leukaemia/lymphoma is the presence of abnormal lymphocytes that express CD25, and CD4 and CD8 T cells expressing the CD25 marker have been shown to be the reservoir of HTLV-1 (Nagai et al., 2001; Yamano et al., 2004). Furthermore, CD4 þ CD25 þ T cells, especially the Treg subpopulation with immunosuppressive functions, can be disrupted

by HTLV, resulting in immunodeficiency (Yamano et al., 2005; Yano et al., 2007). In cases of co-infection with HIV, the number of CD4 þ CD25 þ T cells decreased progressively, eliminating the Treg cell suppressor function and resulting in immune hyperactivation (Eggena et al., 2005). To assess the effect in mandrills, we first evaluated the percentages of CD25 þ in CD4 þ and CD8 þ T cells. In CD4 þ T cells, expression of CD25 þ was significantly higher in the SIVþ /STLV þ group than in the control (po 0.01) and the STLV þ groups (p o0.05); for the STLV þ group, the finding was no longer significant after removal of the extreme value for mandrill A. No significant difference was found from the SIVþ group (Fig. 7A). No difference among the groups was found in the percentage of CD25 in CD8 þ T cells (data not shown). The significance of CD25 þ expression in humans is uncertain, even though it is used in the phenotypic definition of Treg because it is strongly induced by the HTLV-1 Tax protein (Cross et al., 1987; Inoue et al., 1986). The marker used currently to define Treg cells is the forkhead transcription factor FoxP3 (Fontenot et al., 2003; Hori et al., 2003). Therefore, to determine the role of CD4 þ Treg cells precisely, we evaluated FoxP3 expression in CD25 þ CD4 þ T cells. As shown in Fig. 7B, no significant difference was found among the groups. Collectively, these results suggest that dual infection with SIV and STLV induces the activation mechanism in CD4 þ T cells rather than establishing a regulatory system. Characterisation of an STLV cell line derived from a co-infected mandrill Lymphocytes enriched with CD4 þ T cells from mandrill A were cultivated as described previously (Souquiere et al., 2009a), and HTLV-1 P19 antigen and SIV reverse transcriptase were evaluated

S. Souquière et al. / Virology 454-455 (2014) 184–196

**

16

** **

12

**

12 % Ki67 in CD8+T cells

% Ki67 in CD4+T cells

14

10 8 6 4

*

10 8 6 4 2

2 0

**

14

*

0 Uninfected

SIV+

STLV+ SIV+/STLV+

Uninfected

SIV+

STLV proviral load (% in CD4+ cells)

STLV proviral load (% in CD4+ cells)

Spearman r P=0.02

0

2

4

6

STLV+ SIV+/STLV+

SIV/STLV group

STLV Group 26 24 22 20 18 16 14 12 10 8 6 4 2 0 -2

189

8

10

12

% Ki67 in CD4+ T cells

80

Spearman r P=0.04

70 60 50 40 30 20 10 0 -10

0

2

4

6

8

10

12

14

% Ki67 in CD4+ T cells

Fig. 5. Proliferation of CD4þ and CD8þ T cells and correlation with STLV proviral load. Mandrills infected with STLV (Δ), SIV (□), STLV and SIV (◊) and uninfected (○). (A) Percentages of Ki67 in CD4þ T cells. (B) Percentages of Ki67 in CD8 þ T cells. (C) Correlation between STLV proviral load in CD4 þ T cells and percentage Ki67 in CD4þ T cells in STLV-infected mandrills. (D) Correlation between STLV proviral load in CD4 þ T cells and percentage Ki67 in CD4 þ T cells in mandrills co-infected with STLV and SIV. Mann–Witney test used for statistical analysis of each T-cell subset, and correlations sought with the Spearman rank test. Significance assumed at p o 0.05. Regression curves shown as red lines.

at various times after culture. After 7 days of culture, p19 protein was detected in the supernatant, and evidence of SIV reverse transcriptase activity was found (Fig. 8A). After 6 months of culture, a continuous cell line was obtained, with extensive HTLV-1 P19 antigen production in the supernatant but with no evidence of SIV replication. After 6 months of culture, tax/rex mRNA was detected in each cell at a mean of two copies of provirus STLV (data not shown). A study of clonality in the cell line by linker-mediated PCR showed nine integration sites, three of which predominated (Fig. 8B). Our cell line, showed atypical morphology, with dendritic expansion (Fig. 8C). Electron microscopy confirmed the morphology and the production of STLV particles (Fig. 8D). To better characterise these cells, we performed flow cytometry with a panel of antibodies at various culture times. After 21 days, the majority of cells were CD3 þ and some expressed CD40, CD83 and CD86; they did not express CD11c or CD123 (Fig. 9A). After 6 months of culture, the phenotype indicated the absence of CD3 expression, and the majority of cells expressed CD40, CD83, CD86 and CD123 (Fig. 9B). A unique plasmacytoid dendritic cell (pDC) line was derived from a patient with pDC leukaemia (GEN cell line; patent 02–15927) (Chaperot et al., 2001). Our findings indicate that the phenotype of the cell line corresponds to a pDC lineage, which is immortalised and produces STLV particles.

Discussion We report here the results of a clinical, virological and immunological survey of SIV and STLV co-infection in 128 mandrills, representing the largest study of multiple lentiviral infections performed in mandrills.

Mandrills are known to have a high level of SIV replication, and they are natural hosts of SIV (for review, see Pandrea et al., 2008). We confirmed previously that STLV infection is characterised by a variable proviral load in infected animal (Souquiere et al., 2009b). We next investigated the effect of co-infection on CD4 þ and CD8 þ T cells. SIV and STLV mono-infected groups had elevated CD8 þ T cell levels, and animals with SIV infection showed moderate depletion of CD4 þ T cells. These results confirm those of previous studies in naturally SIV-infected sooty mangabeys and mandrills (Sumpter et al., 2007; Apetrei et al., 2011) and our results in mandrills naturally infected with STLV (Souquiere et al., 2009b). From the expression of CD28 and CD95 in T cells, we established that SIV and STLV had no effect on the distribution of CD4 þ TN, TM or TE cells; only SIV infection (either alone or with STLV co-infection) decreased the number of CD8 þ TN cells, as observed previously (Apetrei et al., 2011). Interestingly, coinfection tended to lower the number of CD4 þ T cells. This implies that co-infection involves two mechanisms, due to both SIV and STLV, targeting CD4 þ T cells and finally leading to depletion. Both SIV and STLV-1 integrated in CD4 þ and CD8 þ T cells preferentially target CD4 þ T cells. Perturbation of CD4 þ T cells in uninfected primate models is not clearly understood, but there is evidence that SIV is deleterious for these cells in vitro. The mechanism of action of STLV-1 has not yet been studied, but, as it is the simian counterpart of HTLV-1, some parallels could be drawn. Thus, in HTLV-1 infections, the viral genes Tax and HBZ are thought to play important roles in pathogenesis. Tax potently promotes the expression of its own viral genes and also stimulates the transcription of cellular genes, including cytokines (e.g. IL2), cytokine receptors (IL2Rα) and antiapoptotic genes (Yoshida, 2001). In addition, by binding to other protein complexes, Tax represses the transcription of genes involved in negative control of the cell cycle, activation of

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**

** **

60

** 50 40 30 20

60

*

50 40 30 20 10

10 0

Uninfected

SIV+

STLV+

0

SIV+/STLV+

Uninfected

SIV+

STLV+

SIV+/STLV+

SIV/STLV group

70 % HLADR in CD8+T cells

**

70

** % HLADR in CD8+T cells

% HLADR in CD4+T cells

70

Spearman r P<0.01

60 50 40 30 20 10 0

4

6

8

10

12

14

16

18

20

22

24

26

Neopterin

Fig. 6. Activation of CD4 þ and CD8 þ T cells. Mandrills infected with STLV (Δ), SIV (□), STLV and SIV (◊) and uninfected (○). (A) Percentages of HLADR in CD4þ T cells. (B) Percentages of HLADR in CD8 þ T cells. (C) Correlation between secretion of neopterin in plasma and percentage HLADR in CD8þ T cells in SIV- and STLV-infected animals. Mann–Witney test used for statistical analysis of each T-cell subset, and correlations sought with the Spearman rank test. Significance assumed at po 0.05. Regression curve shown as red lines.

**

100

7

% CD25 in CD4+T cells

% FoxP3 in CD25+ CD4+ T cells

*

90 80 70 60 50 40 30 20 10

6 5 4 3 2 1 0

0 Uninfected

SIV+

STLV+

SIV+/STLV+

Uninfected

SIV+

STLV+

SIV+/STLV+

Fig. 7. Expression of CD25 and FoxP3 in CD4þ T cells. Mandrills infected with STLV (Δ), SIV (□), STLV and SIV (◊) and uninfected (○). (A) Percentages of CD25 in CD4þ T cells. (B) Percentages of FoxP3 in CD4þ CD25 þ T cells. Mann-Witney test used for statistical analysis of each T-cell subset. Significance assumed at p o0.05.

apoptosis and DNA repair. The effect of these properties is that T cells accumulate DNA mutations, resulting in transformation and monoclonal expansion (Jeang et al., 2004). In the present study, expression of CD25 in CD4 þ T cells was enhanced in co-infected animals. We demonstrated previously that CD25 in CD4 þ T cells was correlated with STLV proviral load (Souquiere et al., 2009b), which was confirmed in this study in the STLV þ and SIVþ /STLV þ groups. This finding is in accordance with the fact that Tax protein activates the transcription of gene coding for CD25 (Yoshida, 2001) in HTLV-1 infection, which could explain the increase in this subset of cells. As Treg have been identified among CD4 þ T cells expressing CD25 in humans (Fehervari and Sakaguchi, 2004), it could be hypothesised that the increased activation involves generation of more Treg cells. We did not, however, confirm this with FOxP3

expression in CD4 þ CD25 þ T cells. There is no direct evidence that all Treg cells express FoxP3 in mandrills, however, as in humans (Mori et al., 1996; Thompson and Powrie, 2004). In particular, Yamano et al. (2005) demonstrated the role of Tax in decreased expression of FoxP3 mRNA in CD4 þ CD25 þ T cells. Further investigations are needed to determine the mechanism of overexpression of CD25 in mandrills. We found that proliferation was increased and positively correlated with STLV proviral load in both the group infected with STLV and in the co-infected group. This implies that STLV integration also affects proliferation of T cells, as established in humans (Nicot, 2005). We further showed a direct correlation between the proviral load in CD4 þ T cells and their proliferative capacity in vivo. In humans infected with HTLV, Asquith et al. (2007) demonstrated a correlation between Tax expression and proliferation

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P19 kinetic RT kinetic

4000 2000 3000 2000 1000 1000

Uninfected DNA

Mandrill A

5000 RT in supernatant (pg/ml)

P19 in supernatant (pg/ml)

3000

191

MW

517

396 344 298

0

0 7

14 21 28 35 42 49 56 70 84 98 112 126 140 154 168 182

Days of culture

220 201

154

134

74

FSC

CD3

CD11c

CD86 CD3

CD3

CD123

CD3

CD11c

CD83

SSC

CD40

CD3

CD86

FSC

CD83

SSC

CD40

Fig. 8. Morphological and virological characteristics of cell line from mandrill with scabies. (A) Time course of detection of p19 viral antigen and SIV reverse transcriptase activity in the supernatants of culture-enriched CD4þ T cells from mandrill A. (B) Clonality performed by linker-mediated PCR of mandrill A cell line at 6 months of cultivation. (C) Atypical clumps of transformed cells after Giemsa staining. (D) Visualisation of one STLV particle in the cell line by electron microscopy (100  ).

CD3

CD123

Fig. 9. Phenotypic characterisation of cell line from mandrill A with scabies by flow cytometry. Cells were gated on the live lymphocyte population and analysed for the presence of CD3, CD40, CD83, CD86, Cd11c and CD123 antibodies. (A) Cells sampled at 21 days of culture. (B) Cells sampled at 6 months of culture.

of CD4 þ CD45RO þ (the CD4 þ T cell memory subset in humans) in vivo. STLV infection, but not SIV infection, in mandrills clearly involves increased activation and proliferation of T cells. Activation and proliferation phenomena are particularly important in understanding the pathogenesis of SIV. It was reported recently that limited activation and proliferation protect the natural SIV host (Silvestri et al., 2003; Paiardini et al., 2006; Sumpter et al., 2007). In our study, co-infected animals showed significantly increased activation of T cells, as confirmed by the presence of neopterin. Thus, STLV appears

to perturb the immune system, while SIV plays the role of a deleterious cofactor. Two co-infected mandrills in this study had pathological manifestations, infective dermatitis and scabies, that are unusual in our colony. Both conditions have been associated with HTLV infection in endemic zones (Bittencourt et al., 2006; Blas et al., 2005; Brites et al., 2002; LaGrenade et al., 1995; Mahé et al., 2004). As we saw no clinical signs in mandrills naturally infected with STLV in a previous study (Souquiere et al., 2009b), we hypothesise that the development of these conditions was due to the

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STLV proviral load (% of PBMC)

45

STLV proviral

40 35 30 25 20 15 10 5 0 2004

STLV infection

2005

2006

SIV infection

2007

Pathology

2009

Years

Remission

Fig. 10. Evolution of STLV proviral load in mandrill with infective dermatitis over 6 years.

co-infection. The two monkeys had high STLV proviral loads, but their SIV load was no different from that of controls. An elevated proviral load of HTLV is known to enhance the risk for disease in humans (Casseb et al., 2008; Gabet et al., 2003). We demonstrated previously that the duration of infection of mandrills is correlated with an increase in integrated virus (Souquiere et al., 2009b). The dates of infection were unknown for the mandrill with scabies, which died 3 months after its arrival; necropsy showed chronic granulomatousm eosinophilic serositis and generalised amyloidosis. Mandrill T, which had infective dermatitis, had been infected with STLV 5 years previously, which can be considered recent. Three years previously, however, this monkey was also infected with SIV, which may have perturbed immune control. Consistent with this observation, we examined the evolution of its proviral load during the past 5 years (Fig. 10) and found that the increase corresponded to the date of co-infection with SIV. This implies that in the primo-infection state moderate activation and proliferation due to SIV could promote clonal expansion of STLVinfected T cells. Most importantly, disease occurred concomitantly with co-infection by SIV. CD4 þ T cells were not depleted in these animals, with numbers in the normal range. During a survey of sooty mangabeys, two animals were found to have fewer than 50 cells/mm3 CD4 þ T cells (Sumpter et al., 2007); however, these two animals did not develop disease, and no activation or proliferation was evident. In our study, several mandrills in all groups had low CD4 þ T cell numbers; all were characterised only by old age and were healthy. This implies that the number of CD4 þ T cells is not a criterion of immunodeficiency in nonhuman primates, as it is for HIV and SIV infection in human and macaques (Gordon et al., 2008; Klatt et al., 2008; Milush et al., 2007). The level of immune activation in our study was the highest ever reported in this species, indicating that high immune activation exists in African nonhuman primates. Nevertheless, none of these factors, which are unusual in mandrills, affected the overall number of CD4 þ T cells or the level of SIV replication. Examination of the IFNγ response to the entire overlapping Tax gene in the two mandrills that developed disease showed results comparable to those in mandrills infected only with STLV (Souquiere et al., 2009b). No particular peptide in the Tax region appeared to be the target for the cellular response in these mandrills (Fig. S2), and no correlation was found between the STLV proviral load and the level of cell response to the Tax protein. We also assessed immortalised cell lines from mandrills coinfected with SIV and STLV. Those from STLV-infected animals have a CD4 þ T cell phenotype that produces STLV virions and expresses Tax mRNA (Souquiere et al., 2009a). After 6 months,

however, we identified a new population of cells, which produced only STLV, with no trace of SIV. Moreover, the morphology and immunophenotyping strongly suggested that this cell line consists of pDCs. Our pDC line was grown in the presence of IL-2, which appears to be necessary to modulate its activation and survival (Naranjo-Gomez et al., 2007). It is difficult to explain this phenomenon. Only a few pDCs are present in the bloodstream; for example, only one pDC line was obtained from tumour cells from a patient with an unclassified leukaemia (Chaperot et al., 2001). Therefore, complex events must have to occur in order to produce such cells. The mandrill with generalised scabies presented some biological anomalies. First, co-infection induced replication of SIV in vitro, which involved the destruction of most CD4 þ T cells and made expansion of these cells at term impossible: 74.7% of the CD4 þ enriched cells used for cell culture were infected with STLV. This implies that CD4 þ T cells can produce a large amount of STLV-free virions in the first phase of culture. In the HTLV model, primary dendritic cells are very permissive to HTLV virions (Jones et al., 2008). We therefore assume that pDCs from mandrills have similar properties with regard to STLV and can also produce free virions. This raises the possibility that STLV also uses pDCs to infect target cells in vivo efficiently. This mandrill also had a high level of eosinophils in its blood, and eosinophils have been reported to induce maturation of dendritic cells (Lotfi and Lotze, 2008). These observations are consistent with a disorder in the performance of dendritic cells, in addition to the effects of HTLV itself (Hishizawa et al., 2004; Makino et al., 2000). Further investigations will reveal the exact mechanisms of pDC infection in mandrills. In conclusion, our study shows that retroviral infection can induce immune disturbances in mandrills, especially after coinfection with SIV, which increases activation of the immune system, leading to disease. Dual infections are not rare. Data obtained by our laboratory show that 85% of wild mandrills living in the Lopé reserve in the centre of Gabon are carriers of both infections (Makuwa et al., 2004; Souquiere et al., 2001). In this study, however, STLV infection was the initial infection in almost all cases of co-infection. As we have shown previously, STLV infection can increase immune activation under certain conditions and thus lead to disease conditions that are worsened by coinfection with SIV. SIV can therefore be considered a cofactor in the development of disease related to STLV in mandrills. Moreover, as we have clearly demonstrated, activation and proliferation determine the development of disease, independently of the number of CD4 þ T cells. These findings lead to reconsideration of the effect of multiple retroviral infections in non-human primates. Many questions arise about the interactions in the same host between an oncovirus (STLV) with proliferative properties and a lentivirus (SIV) that is intrinsically deleterious for each CD4 cell. Further studies are needed to explore the mechanisms by which STLV increases proliferation and activation of T cells in mandrills and what occurs in the presence of SIV.

Materials and methods Ethics statement The animals were handled in accordance with standard national operating procedures in the Centre International de Recherches Médicales de Franceville (CIRMF), in accordance with the United States National Institutes of Health guidelines for the Care and Use of Laboratory Animals and in accordance with the recommendations of a working group chaired by Sir David Weatherall in December 2006. The animal protocols and procedures were approved by the

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Gabonese ethics committee for animal experimentation at the CIRMF, and registered under No. CE0 08-006 CIRMF. The housing conditions are in strict accordance with European Union guidelines for animal care (European Union Directive 86/609/EEC). The primate centre has three veterinarians specialized in primates – an ethologist, a primatologist and an ecologist, and all experiments were conducted under their supervision. Each primate housed in the CIRMF Primate Centre has an annual health check under anaesthesia (ketamine at 10 mg/kg body weight). The blood samples taken for this study were collected under strict health control by Primate Centre veterinarians. Specimen collection Blood samples from SIVmnd- and STLV-1-positive mandrills were collected in EDTA K2 tubes under ketamine-HCl and used for flow cytometry. Peripheral blood mononuclear cells (PBMCs) were isolated by Ficoll-Hypaque gradient centrifugation (SigmaAldrich), and plasma was centrifuged at 3000  g for 10 min, dispensed into 1-ml aliquots and frozen at –80 1C. Serological screening Plasma from mandrills was screened for the presence of HTLV1/2 antibodies by two enzyme-linked immunosorbent assays (ELISAs), HTLV-1 Platelia New (Biorad, Marnes-la-Coquette, France) and Vironostica (Biomerieux, Marcy l'Etoile, France). HTLV-1 infection was confirmed by western blot (HTLV blot 2.4, Genelabs Diagnostic). A peptide-based ELISA was used to detect antibodies against SIVmnd-specific peptides mapping the V3 region of the env glycoproteins (Simon et al., 2001). SIV infection was confirmed by western blot HIV-2 (New LAV-Blot II, Biorad, Marnes-la-Coquette, France). Flow cytometry for surface and intracellular markers Four-colour flow cytometry was performed on whole blood according to the standard procedure. The monoclonal antibodies used were CD4-fluorescein isothiocyanate (FITC) (clone MT4-77), HLA DR-phycoerythrin (PE) (clone G46-6), CD25-PE (clone 2A3), CD3-allophycocyanin (APC) (clone SP34-2), CD8-peridine chlorophyll protein (PerCP) (clone SK1), CD28-PE (clone L293) and CD95FITC (clone DX2) (all from BD Bioscience, Le Pont de Claix, France). For intracellular staining, CD4-PE (clone L200), CD3-APC (clone SP34-2), CD8-PerCP (clone SK1) and Ki67-FITC (clone B56) were used. Flow cytometric acquisition was performed on at least 10,000 events in lymphocyte squares on a FACScalibur flow cytometer driven by the CellQuest software package (Becton Dickinson, Heidelberg, Germany). The acquired data were analysed with FlowJo software v7.2 (Tree Star, Inc., Ashland, Oregon, USA). Immunophenotyping of the STLV-1 cell line Cells were sampled on day 21 and month 6 of culture. After washing in phosphate-buffered saline 0.1% bovine serum albumin buffer, cells were stained with CD3-APC (clone SP34-2), CD25-PE (clone 2A3), CD83-PE (clone HB15e), CD86-FITC (clone 2331), CD11c-APC (clone S-HCL-3) and CD123-PE (clone 7G3) (all from BD Bioscience) and CD40-PE (clone MAB89) (Immunotech, Marseille, France). Clonality of cell lines The clonality of STLV-1-infected cell lines (sampled at 6 months) was assessed by the sensitive quadruplicate elongation-mediated

193

polymerase chain reaction (linker-mediated PCR) method described elsewhere (Cavrois et al., 1996; Leclercq et al., 1999). Isolation of CD4 þ and CD8 þ T cells Peripheral blood lymphocytes were separated by Ficoll gradient density centrifugation. Negative depletion was performed on 10 million lymphocytes with magnetic beads coupled to CD4 or CD8 antibody, as recommended by the manufacturer (Dynabeads, Invitrogen Dynal, Oslo, Norway). The phenotype of depleted cells was confirmed by three-colour flow cytometry with monoclonal antibodies CD4-FITC (clone MT4-77), CD3-APC (clone SP34-2) and CD8-PerCP (clone SK1). Cell culture After washing, the remaining CD4 þ - and CD8 þ -enriched cells were suspended in RPMI 1640 growth medium (Cambrex Bioscience, Walkersville, Maryland, USA) supplemented with 20% heat-inactivated foetal bovine serum (Gibco BRL, Eragny, France), 1% penicillin–streptomycin mixture (Gibco BRL, Eragny, France), 1% L-glutamine 200 mmol/l (Gibco BRL, Eragny, France) and 20 U/ml human recombinant IL-2 (Roche Diagnostics, Mannheim, Germany). The lymphocytes were stimulated with 3 mg/ml of the mitogen concanavalin-A (Sigma-Aldrich, Saint Quentin Fallavier, France) and then incubated at 37 1C in 5% CO2. To maintain the cells, 90% of the medium was changed twice a week. SIV reverse transcriptase determination and HTLV-1 P19 antigen detection Reverse transcriptase (RT) activity produced by SIV was quantified in supernatant with the Lenti-RT kit (Cavidi Tech AB, Uppsala, Sweden), as recommended by the manufacturer. The kinetics of STLV production was determined by detection of P19 antigen in supernatant with the Retrotek HTLVI/II p19 Antigen ELISA kit (ZeptoMetrix Corp.) according to the manufacturer's instruction. The concentrations of P19 in specimens were determined by interpolation from a standard curve. Electron microscopy STLV cells line were fixed in glutaraldehyde and then in osmium tetroxide and embedded in epoxy resin. Thin sections were stained with uracyl acetate and lead citrate and examined under a JEOL 100 S electron microscope. SIV viral load RNA was extracted from 150 ml of plasma with the QiaAmp viral RNA mini kit (Qiagen, Courtaboeuf, France) and eluted in 50 ml of Tris EDTA buffer, as recommended by the manufacturer. Quantification was performed by real-time SYBR Greens RT-PCR as described previously (Souquiere et al., 2009c). STLV proviral load determined with real-time PCR DNA was extracted from PBMC with phenol/chloroform, and STLV proviral loads were quantified by real-time PCR, as described previously (Souquiere et al., 2009b). A standard curve was generated with the MT4 cell line containing seven copies of provirus HTLV per cell by 10-fold serial dilutions. To standardise the cell number, the albumin gene was quantified by the TaqMan PCR method with iTaq Supermix and ROX (Biorad, Marnes-la-Coquette,

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France). The TaqMan Probe 5'FAM-CCTGTCATGCCCACACAAATCTCTCC-TAMRA3' (ALBT) and the primers GCTGTCATCTCTTGTGGGCTGT (ALBF) and ACTCATGGGAGCTGCTGGTTC (ALBR) were used as described previously (Souquiere et al., 2009b). The PCR protocol consisted of 3 min at 95 1C, 50 cycles for 10 s at 95 1C and 45 s at 60 1C. The results for the STLV proviral load are presented as copy number per 100 cells.

for technical help. We also thank Professor C. De Giuli Morghen, University of Milan, Italy, for electron microscopy observation. This manuscript is especially dedicated to the memory of our colleague and friend Dr Bettina Sallé.

Detection of tax/rex mRNA by RT-PCR

Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.virol.2014.02.019.

Total cellular RNA was extracted from the cultured T cells with the RNeasy Protect Mini Kit (Qiagen, Courtaboeuf, France). RT and cDNA synthesis were determined as described previously (Kazanji, 2000). Immediately after cDNA synthesis, semi-nested PCR was performed on the pX region, with RPX3 and RPX5 as the outer primers and RPX3 and RPX4 as the inner primers (Kazanji, 2000; Kinoshita et al., 1989). To confirm the presence of amplifiable cDNA in the cultured cells, a portion of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA was amplified with primers 18SL (sense) and 20SL (antisense), as described previously (Kazanji, 2000). Neopterin assay Neopterin in plasma was quantified with a commercial ELISA kit (Neopterin ELISA, IBL-Hamburg GmbH, Hamburg, Germany) according to the manufacturer's instructions. The concentrations of neopterin in specimens were determined by interpolation from a standard curve. Detection of cytokine in plasma by the Luminex method The Bio-Plex human cytokine 27-Plex assay was carried out according to the manufacturer's instructions (Bio-Rad Laboratories, Marnes-la-Coquette, France). A 50-ml volume of each plasma sample (diluted four times in serum) and of the standard was added to a 96-well plate containing 50 ml of antibody-coated fluorescent beads. Biotinylated secondary and streptavidin-PE antibodies were added to the plate with alternate incubation and washing. After the last washing, 125 ml of assay buffer were added to the wells, and the plate was incubated and read on the Bio-plex array reader, with a 5 PL regression curve to plot the standard curve. Samples and controls were read at a low RP1 target setting (to maximise assay sensitivity when the expected concentrations are o3200 pg/ml). Data were analysed with Bioplex manager software, version 3. The results are expressed in pg/ml. Statistical analysis For comparison of groups, Mann–Whitney U tests were performed. Correlations between different sets of data for the same group were analysed by either the standard Pearson correlation coefficient or the Spearman rank correlation test. Significance was assumed at p o0.05. All analyses were performed with Statistica software v7.1.

Acknowledgments The CIRMF is funded by the Gabonese Government, TotalGabon and the French Foreign Ministry. The funders had no role in study design, data collection or analysis, decision to publish or preparation of the manuscript. The authors declare no competing interests. We thank all the staff of the Primatology Centre at the CIRMF for animal care and providing blood samples. We thank Paul Ngari

Appendix A. Supporting information

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