qPCR conditions.

Quantitative PCR assays for DNA were performed as previously described [11]. Quantitative PCR assays for DNA (qPCR) were performed using the Eppendorf realplex master gradient detection system. We used SYBR Green Quantitect (Qiagen) in a 20 µl volume reaction containing 10 µl of SYBR Green buffer, 150 nM of each primer and up to 500 ng DNA sample quantified by ND 8000 (Nanodrop Technologies).

The optimized qPCR conditions used were as follows: 95°C for 15min, 40 cycles of: 95°C for 15s, 60°C for 30s and 72°C for 30s. Samples were tested at least in duplicates in two independent assays. We checked in every individual assay the specificity of the primers by using a melting curve.

Normalization of cellular DNA.

The albumin DNA quantification method was the following: a 141bp-cellular albumin qPCR was done performed to normalize with cellular DNA content (albF-AAACTCATGGGAGCTGCTGGTT, albR- GCTGTCATCTCTTGTGGGCTGT) [4], using the same PCR conditions as for SFV quantification.

All assays included an independent albumin standard curve generated by adding 5-fold dilutions of human genomic DNA (fibroblastic MS5 cell-line) to the PCR reagents, ranging from 0.08ng DNA to 50ng DNA. All qPCR standard curves had correlation coefficients of >0.95.

The albumin DNA content of the samples was determined as follows: the amount of DNA was firstly quantified by Nanodrop technologies and samples of 500ng cellular DNA were used. To normalize samples input, a 250x dilution of the samples (2ng) was used. The amount of albumin DNA was determined using the albumin DNA standard curve, reported as cell equivalents (500ng DNA=75 000 cells), and multiplied by 250 to calculate the “effective” amount of cellular DNA in the samples. The minimum amount of “effective” DNA was 350ng DNA (5 x 10⁴ cells equivalent).

qPCR sensitivity.

FV sensitivity was thus given as 1 copy for 5 x 10⁴ cells equivalent. Of note, for some samples (as AKO394 PBMCs, BAD447 saliva, BAK74 saliva, LOBAK89 PBMCs), the input DNA, after normalization, was >350ng, giving a sensitivity >1 copy for 5 x 10⁴ cells equivalent.

RT-qPCR conditions.

Quantitative real-time reverse-transcription polymerase chain reaction (RT-qPCR) was performed using the same standard as qPCR. RNA samples quantified by ND 8000 (Nanodrop Technologies) were mixed with random hexamer primers (5µM final), dNTP (1mM final) and completed with DEPC-treated water until a final volume of 10 µl. The RNA mixture was subjected to denaturation: 75°C for 5 min, following by 4°C for 5 min (Mastercycler, epGradient ; Eppendorf). Ten µl of enzyme mix including the SuperScriptIII® Reverse Transciptase (Invitrogen) were added to the RNA mixture and subjected to 3 steps: 25°C for 10 min, 45°C for 90 min and 9°C for 5 min. cDNA obtained was added to 10 µl of SYBR Green buffer and 150 nM of each GF5qpcr and GR5qpcr primers in a final volume of 20 µl. The optimized RT-qPCR conditions used were as follows: 95°C for 15min, 40 cycles of: 95°C for 15s, 60.8°C for 30s and 72°C for 30s. Samples were tested at least in duplicates in two independent assays. We checked in every individual assay the specificity of the primers by using a melting curve. Our positive control was RNA extracted from the human microglial cell line CHME productively infected with SFV strain from BAK74.

Normalization of cellular RNA.

The GAPDH cDNA quantification method was the following: a 107bp-cellular GAPDH qRT-PCR was done to normalize with cellular RNA content (GAPDH-F 5′- ATGGGGAAGGTGAAGGTCG-3′ and GAPDH-R 5′- GGGTCATTGATGGCAACAATATC-3′) [15], using the same PCR conditions as for SFV quantification.

All assays included an independent GAPDH cDNA standard curve generated by adding 5-fold dilutions of human genomic cDNA (CHME microglial cell line) to the PCR reagents, ranging from 10pg to 100ng starting RNA. All RT-qPCR standard curves had correlation coefficients of >0.95.

To normalize samples input, a 4x dilution of the samples was used. The amount of GAPDH cDNA was calculated using the GAPDH cDNA standard curve, reported as cells equivalent (50ng RNA= 10⁴ cells), and multiplied by 4 to calculate the effective amount of cellular RNA in the samples. The minimal RNA content was 2.5ng RNA (500 cells equivalent), except for the lowest concentrated samples (100pg RNA, 20 cells equivalent).

RT-qPCR sensitivity.

The sensitivity of the assay was thus 1 copy/500 cells equivalent, except for the lowest concentrated samples (sensitivity of 1 copy/20 cells equivalent).

Quantification of SFV DNA load.

We studied 14 SFV-infected hunters, all of them infected by a gorilla SFV strain (Table 1). They have been determined SFV-positive in a previous epidemiological study by both serological and molecular findings [11].

We first confirmed the SFV status of the 14 individuals. We performed a nested-PCR on the pol-in region of SFV using DNA isolated from PBMC samples (Table 2). As expected, all the individuals were found to be SFV-infected (Figure 1 “Nested-PCR” line under “PBMCs” bars). To explore the presence of SFV DNA in saliva samples, we performed the same nested-PCR on oral swabs. We detected SFV DNA in 8/14 saliva samples (Figure 1 “Nested-PCR” line under “Saliva” bars).

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Figure 1. Detection and quantification of Simian Foamy Virus (SFV) DNA in samples of PBMCs and saliva derived from 14 humans infected by a gorilla strain of SFV.

Detection of SFV DNA: Presence (+) or absence (-) of SFV DNA in PBMCs and saliva samples of each individual are indicated, based on nested-PCR results.

Quantification of SFV DNA: Blue and red bars represent mean SFV DNA levels from at least two independent assays run in duplicate in PBMCs and saliva samples, for each individual respectively. Viral loads were normalized to cell equivalents by q-PCR for albumin. Error bars represent the standard deviations from an individual participant. Mean of SFV DNA copies in PBMCs (blue) and saliva (red) of the 14 individuals are shown by the dotted lines.

https://doi.org/10.1371/journal.pone.0077072.g001

We then quantified SFV DNA load by performing q-PCR on the same samples. SFV DNA load could be quantified in 13/14 PBMC samples and in 6/14 saliva samples (Figure 1). Total SFV DNA load in PBMCs and in saliva were very low in most of the samples (7.1 ± 6.0 and 2.4 ± 4.3 SFV DNA copies/10⁵ cells respectively). No correlation was found between SFV DNA loads in the PBMCs and saliva samples Of note, only individuals with long time of infection (>16 years, which is the median) had more than 1 SFV copy/10⁵ cells in the saliva (data not shown).

The sensitivity of the technique used to detect SFV DNA is higher for our nested-PCR assay (1 copy/150 000 cells) [10] than for our q-PCR assay (1 copy/50 000 cells): 3/28 DNA samples (saliva from BAD456 and BAK177 and PBMCs from LOBAK2) were positive by nested-PCR but negative by q-PCR, probably due to a low viral load (Figure 1). Overall, in 8/14 cases, SFV nested-PCR was positive in DNA samples of both PBMCs and saliva derived from SFV infected-individuals (Figure 1 and Table 3).

Characterization of SFV clones.

We generated SFV clones of the pol-in fragment (mean number of 18 clones) when both saliva and PBMCs samples contained SFV DNA (8/14 individuals). At the individual level, the sequences of SFV clones derived from PBMCs versus saliva often differed by only 0.5% genetic variation (in 5/8 individuals, Figure 2A).

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Figure 2. Phylogenetic tree generated with sequences of 425 bp fragments of the SFV pol-in from blood and saliva of SFV-infected hunters BAK132, BAK177, BAD456 and LOBAK2 and different gorillas

Sequences from hunters BAK132 (A), BAK177 (B), BAD456 (C) and LOBAK2 (D) are indicated in red (saliva clones, "S") and blue (PBMC clones, "P"). Sequences are compared to 8 sequences available in Genbank from different central African NHP gorillas (green). Alignment was performed with the Dambe version 4.5.68 and Clustal W. Phylogenetic analysis was performed with PAUP, version 4.0b10 (Sinauer Associates, Sunderland, MA, USA) based on the Neighbor joining method applying the best model calculated by AIC: HKY (BAK132, BAK177), GTR (BAD456) or TrN (LOBAK2). Bootstrap analysis of 1000 replicates is shown as percentage at nodes. Only values greater than 50% are shown. The scale of the tree is 0.01 nucleotides replacement per site.

Of note, phylogenetic tree for BAK132 is representative of trees obtained for the other following hunters (BAK33, BAK74, BAK132, BAD447). Sequences accession numbers for BAK132 SFV clones are KF515432-KF515469. Phylogenetic tree for BAK177 indicates the presence of two distinct groups of SFV clones, suggesting a co-infection with two SFV strains. Sequences accession numbers for BAK177 SFV clones are KC602165 to KC602200. For BAD456 and LOBAK2, some clones displayed G to A mutations (G->A) present in a GG dinucleotide context and absent in the 8 NHP gorillas strains. Sequences accession numbers of SFV clones are KC602201 to KC602238 (BAD456) and KC602129 to KC602164 (LOBAK2).

Due to very low viral loads, a single cluster of SFV clones within one compartment (PBMCs or saliva) might derive from PCR amplification of a single SFV copy.

https://doi.org/10.1371/journal.pone.0077072.g002

In 1/8 case (BAK177), a higher genetic variation (3.3%) in the range of the inter-strain variability was found among SFV clones which segregated into two clusters (Figure 2B). Such data could indicate a co-infection by two SFV strains.

For individuals BAD456 and LOBAK2, consensus sequences of PBMCs and saliva SFV clones were not identical and displayed a G/A polymorphism in a GG dinucleotide-context (representing 5/5 and 4/8 nucleotide differences respectively), with the A polymorphism being absent in the 8 SFV strains found in gorillas and available in GenBank (Figures S1 and S2 provide alignments of SFV sequences, and Figure 2C and 2D provide corresponding phylogenetic trees).

SFV DNA load in PBMCs of SFV-infected individuals.

In NHP, few studies are available on viral load in blood samples. In 11 healthy, non-immunosuppressed macaques (Macaca mulatta), SFV DNA is present at a low copy number in PBMCs with however a relatively wide range (mean 98, range 19-223 copies/10⁵ cells) [5]. In another series of 18 macaques (Macaca tonkeana), we previously found a very low viral load, ranging from 1 to 100 SFV DNA copies in 500 ng of peripheral blood buffy-coat DNA (equivalent to 0.7-70 SFV DNA copies /10⁵ cells) [18]. In chimpanzees, the situation in the buffy-coat was found to be quite similar [4].

In humans, Switzer et al. reported SFV infection (by an Angolan colobus or red-tailed monkeys strains) in 3 women from Democratic Republic of Congo, with viral loads ranging from 14-1755 copies/10⁵ blood cells [12]. In natural settings, our laboratory reported that viral loads range from 1 to 100 copies/10⁵ blood cells in 40 humans infected with a gorilla SFV strain, and 100-1000 copies/10⁵ blood cells in 2 individuals infected with a chimpanzee strain [10,11].

In this report, we extended these findings demonstrating low SFV DNA loads (less than 20 copies/10⁵ cells) in the same individuals, with variation possibly linked to sampling date and the use of PBMCs instead of buffy-coat.

Of note, putative fluctuation of SFV load should be considered with the potential for onward transmission if exposure occurs at that moment in time, even though epidemiological data, with no cases of human to human transmission reported so far, suggests this is not a major route of infection. We previously reported the case of BAD447, whose wife BAD460 was found to be positive for SFV by serology, but not by nested-PCR [11]. We note that SFV DNA loads are not especially high for this individual, either in PBMCs or in saliva. Further studies looking at SFV RNA and DNA loads in sexual compartments could be interesting.

Molecular characterization of SFV in the saliva of SFV-infected individuals.

It is interesting that, if the blood is presumably the first site of SFV infection after a bite, 8/14 saliva samples also contained SFV DNA as shown by nested-PCR positivity. Detection and/or isolation of SFV in the buccal cavity have been shown in NHP and in 3 humans [6,14]. This could be due to the trafficking of PBMCs to the oral mucosa, contamination when sampling or infection of epithelial cells in the oral mucosa [19]. However, the genetic differences between SFV clones derived from samples of saliva and PBMCs as shown for LOBAK2 and BAD456 individuals suggest that SFV DNA found in the saliva does not simply mirror SFV DNA derived from PBMCs. Additional SFV cloning and sequencing experiments based on available material (BAD456 PBMCs and saliva samples) confirmed that the pools of clones of these two compartments were distinct, with a G/A polymorphism.

SFV intra-individual variability.

The low intra-individual SFV variation is in agreement with the well-known genetic stability of SFV [20]. This is also in agreement with the low intra-individual SFV genetic diversity found in NHP [21]. Of note, inferred from intrinsic SFV RT fidelity [22], G/A polymorphism (accounting for 35% of in vitro RT substitution) in the G-G dinucleotide context (accounting for 27% of the A/T/G/C-G dinucleotide context) should represent 9% of the total viral genetic substitutions. The bias towards G/A polymorphism in a GG dinucleotide-context in SFV clones derived from BAD456 and LOBAK2 samples, which are absent in the available SFV gorilla strains from the same regions, could be indicative of APOBEC3 activity, as previously reported in few other individuals from another series [23]. It would have been of course interesting to have the sequence of the infecting gorilla SFV strain, which is obviously not possible to obtain, and to look at a more variable gene. This higher genetic variability allows to appreciate the compartmentalization of SFV in one individual. Indeed, SFV compartmentalization could also occur in other individuals but might not be visible, because the genetic variability of SFV clones is too low.

Detection of SFV DNA does not predict SFV replicative status.

SFV DNA is not a dead-end product reminiscent of SFV primary infection and is still functional even decades after primary infection, as SFV has already been isolated from blood cells and saliva of SFV-infected humans [9], including from PBMCs of 2 hunters involved in this work [24]. This raises the question of the replication status of SFV within the human host. SFV DNA detected can come from proviral DNA (present in latent and productive infections) and/or viral DNA (present in productive infections). Indeed, SFV DNA can be integrated into the host genome (proviral DNA) and can also be produced upon productive infection (viral DNA), in a later reverse-transcription step. Production of viral DNA is preceded by a SFV RNA step during the life cycle of SFV [25]. In order to search for productive infection in humans, we investigated SFV RNA levels, in humans.