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Blood, Vol. 91 No. 10 (May 15), 1998:
pp. 3841-3849
In Vivo Tropism of Hepatitis C Virus Genomic Sequences in
Hematopoietic Cells: Influence of Viral Load, Viral Genotype, and Cell
Phenotype
By
Hervé Lerat,
Sylvie Rumin,
François Habersetzer,
Françoise Berby,
Mary-Anne Trabaud,
Christian Trépo, and
Geneviève Inchauspé
From INSERM U271, Lyon, France.
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ABSTRACT |
Extrahepatic sites capable of supporting hepatitis C virus (HCV)
replication have been suggested. We analyzed the influence of
virological factors such as viral genotype and viral load, and cellular
factors such as cell phenotype, on the detection rate of HCV sequences
in hematopoietic cells of infected patients. Thirty-eight chronically
infected patients were included in the study: 19 infected by genotype 1 isolates (1a and 1b), 13 by nongenotype 1 isolates (including genotypes
2 a/c, 3a, and 4), and 6 coinfected by genotype 1 and 6 isolates.
Polymerase chain reaction (PCR) detection efficiency of viral genomic
sequences, both the positive and negative strand RNA, was evaluated
using RNA transcripts derived from genotype 1, 2, 3, and 4 cloned
sequences and found to be equivalent within one log unit. The serum
viral load, ranging from less than 2 × 105 Eq/mL to 161 × 105 Eq/mL, did not influence the detection rate of
either strand of RNA in patients' peripheral blood mononuclear cells
(PBMCs). Positive and negative strand RNA were found in PBMCs of all 3 cohorts of patients with a detection rate ranging from 15% to 100%
and from 8% to 83.3% for the positive and negative strand RNA,
respectively. Coinfected patients showed a detection rate in all cases
greater than 80%. Patients infected with genotype 1 isolates showed a
higher detection rate of either strands of RNA when compared with
patients infected with other genotypes (P < .001 and
P < .04). Both strands were found restricted to polymorphonuclear leukocytes, monocytes/macrophages, and B (but not T)
lymphocytes. These data show that HCV genomic sequences, possibly
reflecting viral replication, can be detected in PBMCs of chronically
infected patients independent of the viral load and that specific
associated cell subsets are implicated in the harboring of such
sequences.
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INTRODUCTION |
INFECTION CAUSED BY hepatitis C virus
(HCV), a single-stranded RNA virus belonging to the Flaviviridae
family,1 evolves in more than 70% of cases towards a
chronic carrier state that is responsible for liver cirrhosis in
approximately 20% of patients.2 HCV has also been found to
be closely associated with the development of hepatocellular
carcinomas.3
At least six different genotypes (1 to 6) and 52 subtypes of the virus
have been identified based on differences in the nucleotide sequences
(for review see Bukh et al4). Many studies suggest that the
course and severity of the disease may depend on the infecting
genotype, although it does remain a matter of
controversy.5,6 These studies point toward the development
of more aggressive liver disease when infection is due to subtype 1b.
Infection with this subtype has also repeatedly been found associated
with more severe graft injury in patients who have undergone
transplantation.7 Moreover, different HCV strains may vary
in their responsiveness to interferon therapy, subtypes 1b and possibly
1a being the poorest responders.8,9
HCV, like other hepatitis viruses, is predominantly a hepatotropic
virus. Nonetheless, previous work by several groups including ours
suggests that HCV genomic sequences, both the positive and negative
strand RNA (the presumed replicative intermediate),1 can be
detected in peripheral as well as medullar hematopoietic cells, mainly
peripheral blood mononuclear cells (PBMCs) from infected
patients.10-17 However, very few studies have attempted to
further discriminate the cell populations harboring these
sequences.18-20 Such studies implicate mainly B cells as a
potential reservoir for viral replication. A more recent study has, on
the contrary, failed to document any evidence of HCV replication in
hematopoietic cells or cells from any tissues other than the liver in
human and chimpanzee samples.21 All of these reports
illustrate the actual controversies and difficulties in providing clear
and specific evidence on the subject of the possible existence of
extrahepatic reservoirs capable of supporting HCV replication. A
disturbing observation is the wide variation in the detection rates of
HCV genomic sequences found in PBMCs of patients observed in the
different published studies. Such rates can range from 0% to
100%.10-21 Most of the conclusions reached to date have to
be interpreted with caution because they involved a very limited number
of patients or because detection techniques used may have been
subjected to artifacts resulting in erroneous
observations.10,21 In particular, most of these studies
used very poorly validated methods for detection of the negative strand
RNA. In addition, comparison of the different studies reported has been
hampered by the fact that the patient populations analyzed were not
well characterized, in particular with respect to the infecting viral
load and genotype. Analysis of such factors is important, not only to
better define a putative difference in the biology of HCV viruses
belonging to different genotypes, but also to allow for adequate
comparison of data provided by different studies.
Because total PBMCs represent a heterogeneous cellular population,
including lymphocytes and monocytes/macrophages (M/M) that show
different functions related to host defense against infection, it is
particularly important to gain further insights in the cellular as well
as the molecular events involved in the apparent infection of these
cells by HCV. In this study, we looked for evidence of HCV genomic
sequences (both the positive and the negative strand RNA) in PBMCs in
relation to the infecting viral load and genotype, as well as with the
phenotype of cell subsets harboring these sequences.
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MATERIALS AND METHODS |
Patients.
A total of 38 patients, 17 females and 21 males, ages 24 to 75 years
(mean 46.9 ± 2.1 years) were studied. All were positive for HCV
antibodies as detected by ELISA (third generation) and RIBA III assays
(both Ortho Diagnostic Systems Inc, Raritan, NJ). Viral infection was
confirmed in all patients by polymerase chain reaction (PCR) detection
of HCV RNA in patients' sera.10 Except for 1 case, all had
chronic hepatitis (n = 7, with cirrhosis). No evidence of other active
viral infection (HIV, HBV, HAV) was documented in these patients. The
infection mode was related to blood transfusion, parenteral drug abuse,
occupational exposure, or was unknown in 14, 9, 1, and 14 cases,
respectively. Three patients were under combination immunosuppressive
therapy (steroids and cyclosporines) because of a previous hepatic
transplantation (from the group of coinfected patients, see Results).
All patients underwent a complete clinical assessment including liver
biopsy.
Determination of genotypes.
The genotype of infecting strains was analyzed from sera, total PBMCs,
or hematopoietic cell subsets of infected patients by using three
different assays. Two assays are PCR based, the commercial HCV
INNO-LIPA assay (INNOGENETICS, Gent, Belgium) and the CAP-PCR assay.
This latter assay uses genotype-specific primers from the nucleocapsid
(CAP) region and was adapted from the original Okamoto's
technique.22 We designed a number of original primers, including 186 NTER, 132 N, 104 IIa, 134 N, 104 IIIa, 339N, and 104Va.
Complementary DNA (cDNA) synthesis was performed using the
186NTER primer (ATAGAGAAAGAGCAACCGGG) and the sense primer 256, as
described.22 The amplified product (1/50) was used for the
second (nested) PCR using a mix of 10 primers specific for the
amplification of 4 different HCV genotypes: 1 (a and b), 2, 3 (a), and
4. Detection of genotypes 5 and 6 is not achievable with this
technique. Four sense primers, 104 AGGAAGACTTCCGAGCGGTC, 104 IIa
AGGAAGACTTCGGAGCGGTC, 104 IIIa CGTAAAACTTCTGAACGGTC, and 104 IVa
CGAAAGACTTCGGAGCGGTC, and six antisense primers, 132 Nbis GCAGCCCTCATTGCCATA, 133 Nbis GCCATCCTGCCCACCCCATG, 134 Nbis1
ACTTGCCAGTGGAGCGCCG, 134 Nbis2 ATTTGCCAGTGGAGCGCCG, 339N
GCTGAGCCCAGGACCGGCCR, and 465 TCCCGTCCTCCACAGCCCRG were used. Primer
combinations, expected size of products, and corresponding detected
genotypes/subtypes were as follows: 104/132N, 125 bp, 1a; 104/133, 141 bp, 1b; 104 IIa/134N, 75 bp, 2; 104IIIa/339N, 87 bp, 3a; and 104 IVa/465, 336 bp, 4a. An illustration of the results
obtained with this technique is shown in
Fig 1.
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| Fig 1.
Analysis of the genotype distribution in the serum and
PBMC of a HCV patient harboring a dual infection. Nested RT-PCR was performed using Cap derived primers for the amplification of genotype specific products either from the positive or the negative strand viral
RNA. Control human sera included positive strand RNA amplified from
patients infected with genotypes 1a, 1b, 2(a/c), 3 and 4 (lanes 1, 2, 3, 4, and 5, respectively). Serum (S, lanes 6 and 8) and total PBMC
(PB, lanes 7 and 9) from a patient infected with a genotype 1a and a
genotype 2 were used for amplification of both the positive (+) and
the negative ( ) strand RNA. PCR products were fractionated on a 3%
agarose gel and stained by ethidium bromide. Markers are shown on the
left hand side (base pair, bp) while expected sizes of amplified
products specific for the different genotypes are shown on the right
hand side (bp).
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The third assay, the MUREX 1-6 test (MUREX Inc, Dartford, UK) is based
on the detection of genotype-specific antibodies. All genotypes
presented in the study were deduced from concordant results obtained
with at least two of the three assays run. Distribution of the
different genotypes/subtypes was: 1a, n=5; 1b, n=14; 2a/c, n=4; 3a,
n=7; 4, n=2; and 1a + 2 a/c, n=5; 1b + 2 a/c, n=1.
Assessment of viral loads.
Positive strand HCV RNA was quantified by the branched
(bDNA) assay (Chiron HCV RNA 2.0 Assay; Chiron Corp,
Emeryville, CA) using 50 µL of serum. This second-generation assay
has been shown to correct for genotype differences in amplification
efficiency. Patients' viral loads varied from less than 2 × 105 to greater than 107 genomic equivalents
(Eq)/mL of serum.
Extraction of nucleic acids and cDNA synthesis.
They were essentially performed as described.10 RNA was
extracted from 250 µL of serum and different amounts of
total PBMCs (see Table 1) or hematopoietic
cell subsets using two phenol/acid guanidium thiocyanate extraction
steps, followed by a chloroform extraction step and precipitation with
ethanol. cDNAs were synthesized with specific primers as described
below and as reported elsewhere. Distilled water, normal sera, and
normal total PBMCs or cell subsets were used as negative controls.
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Table 1.
Detection of HCV Positive and Negative Strand RNA in
Different Subsets of Peripheral Hematopoietic Cells
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Cloning, sequencing, and in vitro transcription.
Briefly, total RNA was extracted from 4 patients' sera (250 µL23) harboring subtypes 1b, 2a/c, 3a, and 4 strains as
determined by the INNO-LIPA assay. cDNA was synthesized with
genotype-specific primers located in the C-terminal end of CAP and
designed according to Bukh et al.24 RNA samples were
preheated first for 10 minutes at 70°C, with 0.1 µm of primer and
10 U RNasin (Promega, Madison, WA), and cDNA synthesis was
then performed as described.10 Samples were then heated to
95°C for 30 minutes. A unique-sense primer (nt 37-56), from the
highly conserved 5 UTR was used for PCR amplification of the cDNA.
Final products encompassed nt 37-901, 37-900, 37-904 for genotype 1b-,
3a-, and 4-derived sequences, respectively. For the genotype 2a/c
sequence, a nested PCR was performed with 1/25 of the first
amplification product using internal primers. The final product
encompassed nt 45-884. PCR products were cloned in the pGEM-T vector
(TA Cloning Kit; Promega) according to the manufacturer's
instructions. Plasmid DNA was purified (QIAgen plasmid preparation
system, Qiagen) and sequenced, using a DNA Sequencer
Stretch (Applied Biosystems, Foster City, CA). Sequences were compared with published databases to confirm the genotype/subtype of infecting isolates (courtesy of L. Jarvis and P. Simmonds). The
sequence from the isolate 2a/c was confirmed to be 2c and for isolate 4 to be 4c. A clone encompassing the entire structural region of a genotype 1a strain, strain H, was also used.25
GenBank accession numbers for the sequences derived in this study are U94722-U94724.
Three µgs of plasmid DNA were linearized with SphI or NotI and in
vitro transcription was performed with SP6 or T7 RNA polymerases, respectively, using a T7-SP6 Riboprobe System (Promega). A 15-minute digestion with Klenow DNA polymerase (GIBCO, Grand Island,
NY) of the 3 protruding ends generated by SphI was
performed at 22°C before in vitro transcription, in order to avoid
synthesis of RNA molecules that were initiated at the terminus of the
template.26 Digestion of template DNA was performed using
RQ1 DNase (Promega) at the concentration of 1 U/µg DNA twice at
37°C for 15 minutes. The obtained synthetic RNAs were quantitated
using a UV spectrophotometer at 260 and 280 nm. Integrity of the
products as well as predicted concentrations were controlled on
ethidium bromide-stained agarose gel. These analyses were
systematically performed in duplicate. RNAs were serially diluted in
DEPC-treated water before reverse transcription-polymerase chain
reaction (RT-PCR) amplification. Absence of residual plasmid DNA was
controlled by performing PCR amplification without prior cDNA synthesis
using primers located in the 5 NCR. In all cases, residual DNA could
be detected at a concentration of 109 to 106
RNA molecules per assay and thus did not interfere with the
quantitative analysis described thereafter. All analyses were performed
in duplicate, independent experiments.
PCR amplification of cDNA.
One-eighth of the generated cDNA was amplified as previously
described10 (Inchauspé et al,
submitted) using primer pairs and conditions specific for
the amplification of positive (using primers from the 5 noncoding
region, 5 NCR primers) and negative (using primers from the
nucleocapsid region, CAP primers) strand HCV RNA.
For detection of the positive strand RNA, one-fourth of the PCR product
was analyzed by agarose gel and Southern hybridization with a 32 P-labeled probe internal to the PCR primers.10 The probe
was derived from a genotype 1a sequence. All samples were analyzed at
least in duplicate experiments. Hybridizations were done as previously
described.10
For the detection of the negative strand RNA, typically a RT-PCR nested
amplification of CAP viral sequences was performed. Primers for cDNA
synthesis and first PCR are those described for the CAP-assay (used in
a reverse order). These primers show a high degree of conservation
ranging from 85% to 100% between genotypes. For the second PCR, the
cocktail of 10 different primers (5 pairs) described for the CAP-assay
were used. Primers displayed 100% homology with the corresponding
prototype sequences.24 PCR conditions consisted of an
initial cycle at 95°C for 5 minutes, 30 cycles at 95°C for 1 minute, 63°C for 1 minute (negative strand); or 55°C for 1 minute (positive strand), 72°C for 1 minute 30 seconds, and a final
extension cycle of 72°C for 10 minutes. One-fourth of the PCR
product was analyzed by agarose gel. In some cases (dual infection), a
RT-PCR using the reverse set of primers was performed for amplification
and direct genotyping from the positive strand RNA.
Purification of PBMCs and peripheral hematopoietic cell subsets.
Peripheral venous blood cells were collected in EDTA-treated tubes.
Mononuclear cells were obtained as described,10 ie, after
Ficoll separation at 300g for 15 minutes (the collected fraction still contained granulocytes and erythrocytes, up to 20%).
When specified, peripheral hematopoietic cell subsets were separated
and purified using immunomagnetic positive selection with antibodies
directed against specific surface molecules. Mononuclear cells were
washed and the cell pellet was resuspended in 80 µL phosphate-buffered saline (PBS) buffer containing 0.5% bovine serum
albumin (BSA) (wt/vol) and 5 mmol/L EDTA (PBBE) per 107
cells. Twenty µL of anti-CD3 (Leu4 clone, Becton Dickinson, San Jose,
CA) coupled with magnetic microbeads (Miltenyi Biotech
Inc, Sunnyvale, CA) per 107 cells was added and the mixture
was gently mixed by rotation for 15 minutes at 4°C. Cells were then
washed and resuspended in 100 µL of PBBE per 107 cells.
Depletion of T lymphocytes (CD3+ cells) was performed using
a VarioMacs column-type BS (Miltenyi Biotech Inc)
according to the manufacturer's instructions.
CD3 cells were further separated using MiniMacs
columns (Miltenyi Biotech Inc). CD3 cells were first
incubated 20 minutes at 4°C with an anti-CD15 antibody (Leu-M1;
Becton Dickinson), washed, and incubated 15 minutes at 4°C with
goat antimouse Ig-conjugated microbeads (Miltenyi Biotech Inc).
CD15+ cells were purified and collected.
CD15 cells were then incubated with an anti-CD14
antibody (LeuM3, Becton Dickinson) and further separated as mentioned
above. CD14 cells were finally incubated with an
anti-CD19 antibody (LeuM12; Becton Dickinson). Those sequential
separations allowed us to purify four different cell subsets: CD3,
CD15, CD14, and CD19+ cells. The last negative fraction
consisted mainly of platelets and dead cells. The viability of cells
was tested after each purification by the mean of trypan blue
exclusion. Purity of each cell subset was assessed by morphological
(May Grunwald Giemsa staining) and phenotypical analysis by
immunofluorescence study of membrane antigens with a FACscan flow
cytometer (Becton Dickinson, Baltimore, MD). Results of a typical
purification are summarized in Table 2 and
show that all fractions contained more than 95% of the expected cells.
Dry pellets of those cell subsets were stored at -80°C before RNA
extraction.
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Table 2.
FACscan Analysis of Immunofluorescence Assays Performed
on PBMC and Cell Subsets Purified From HCV-Infected Patients
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Statistical analysis.
Genotype distribution was analyzed with the  2
or test.
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RESULTS |
Efficiency of PCR-based detection assays for the positive and negative
strand RNA.
Two RT-PCR assays were typically used in this study; for the detection
of the positive strand RNA, primers were located in the 5 NCR, whereas
for the negative strand RNA, they were located in the CAP
region. Although the 5 NCR is the most conserved domain in the HCV genome (>97%),25 its use as a target for the
amplification of HCV negative strand RNA has been limited because it
has been shown that artifactual priming of the template could result in erroneous interpretation of the results.10,21,27 Most of
the described artifacts can nonetheless be eliminated when primers for
the PCR-amplification are derived from the less structured CAP
region.10 This region, although less conserved than the 5
NCR, is the most conserved region in the HCV genome open-reading frame
(90% to 95%).25
Primers used for cDNA synthesis and PCR amplification of the positive
strand RNA displayed 100% homology with genotype 1 to 4 sequences
found in published data bases. The primer used as probe in the Southern
blotting analysis displayed 91% to 100% homology with genotype 1 to 4 described sequences from the same databases (Gene
Bank24,28). Primers used for the detection of the negative
strand RNA had been derived from conserved regions deduced from
published consensus sequences (see Materials and Methods). CAP primers
used for cDNA synthesis and first PCR showed 85% to 100% homology
when compared with genotype 1a, 1b, 2a/c, 3a, and 4 templates,
respectively, for the antisense primer, and 0 nt, 1 nt, 1 nt,
3 nt, and 2 nt mismatch, respectively, for genotypes 1a, 1b, 2a/c, 3a,
and 4 templates for the sense primer. Primers used for the nested
CAP-PCR showed 100% homology when compared with genotype consensus
sequences described by Bukh et al.25
It was particularly important in the frame of our study and with
respect to the above remarks to assess the detection efficiency of both
strands of RNA specific for each genotype included in our study. Such
evaluation was performed using synthetic RNAs. Figure 2 illustrates the sensitivity of
detection of the positive strand (Fig 2A) or the negative strand (Fig
2B) RNA using transcripts derived from genotype 1 (a or b), 2c, 3a, and
4c cloned sequences. The following observations could be made:
sensitivity of detection of the positive strand RNA observed after
single PCR amplification ranged from 102 to 103
template copies per reaction. It was the lowest (103) for
genotypes 1a- and 3a-derived RNA. The level of detection was unchanged
after Southern blot hybridization (data not shown). The sensitivity of
detection of the negative strand RNA also ranged from 102
to 103 template copies per reaction after nested
amplification, the lowest sensitivity (103) being detected
for the genotype 4c-derived RNA. For genotype 4c-derived sequences,
duplicate bands could be observed, after nested PCR, possibly
reflecting annealing of primers at two different positions on the
synthetic template yielding lower size products than the expected ones.
These bands were nonetheless never detected using biological samples
(see, for example, Fig 1, lane genotype 4 below) and did not affect
sensitivity of the assays. In presence of biological contexts (such as
serum or liver extracts), an additional loss of half a log to one log
in sensitivity was noticed independent of the genotype tested and as
previously reported.10
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| Fig 2.
Efficiency of detection of positive (A) and negative (B)
strand RNA from genotype 1 to 4 derived templates. Synthetic RNA, encompassing the near full length 5' NCR and CAP sequences from genotypes 1 to 4 HCV sequences, were derived as described in Materials and Methods and used in serial dilution assays. Assays included amplification of 0 to 10 6 RNA copies per reaction. PCR products obtained after single rounds of amplification (positive strand RNA, 1A)
or nested rounds of amplification (negative strand RNA, 1B) were
fractionated on 2.5% agarose gel and stained with ethidium bromide.
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Thus, under experimental conditions used in our study, a maximal
difference of 1 log unit in sensitivity of RNA template detection could
be observed for either the positive or the negative strand RNA and for
all the genotypes tested.
Detection of HCV genomic RNA in total PBMCs of chronic carriers:
Association with serum viral load and genotype.
The presence of positive and negative strand HCV RNA in total PBMCs was
analyzed from 38 chronically infected patients and results interpreted
with respect to the viral load and viral genotype of the patients. All
genotypes indicated were confirmed by at least 2:3 independently run
genotyping assays (see Materials and Methods). Overall results are
shown in Fig 3A and 3B.
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| Fig 3.
Percent of patients harboring HCV RNA sequences in total
PBMC: correlation with viral load and viral genotype. Serum (250 µL)
and PBMC (2 × 106 cells) from 38 patients were processed
as described in Materials and Methods. Titration of HCV RNA in serum
was performed by means of the bDNA assay (HCV RNA 2.0 assay). All
indicated genotypes were deduced fom concordant results obtained from
three genotyping assays. Titers < 2 × 105 genome
equivalents/ml (Eq/mL) were considered equal to 2 × 105 Eq/ml for representation in the figure. (2A): detection
of the positive strand RNA; (2B): detection of the negative strand RNA. Subtype distribution in co-infected patients was: subtype 1a, n = 5, subtype 1b, n = 1. Open circles indicate individual viral titers for
each patient while black symbols represent the mean titers for each
group of patients represented.
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Detection of the positive strand RNA (2A).
Positive strand RNA was detected in all three groups of patients: the
genotype 1 infected group, the nongenotype 1 infected group (others),
and the groups of coinfected patients. Detection rates were 79%
(15:19) for genotype 1 infected patients, 15% (2:13) for nongenotype 1 infected patients, and 100% (6:6) for patients from the coinfected
group. These latter patients harbored a dual infection due to genotype
1a/b and 2a/c viruses. More precisely, positive signals were detected
in 2:5, 13:14, 1:4, 1:7 genotype 1a, 1b, 2a/c, and 3a infected
patients. The difference in detection rates between genotype 1 and
other genotype infected patients was statistically significant
(P < .001). As shown in the figure, viral loads
between the groups of patients harboring HCV sequences in their PBMCs
and the ones who did not were comparable. They ranged from less than
2.105 Eq/mL to 161 × 105 Eq/mL, whereas
mean titers varied from 11 ± 15 to 26 ± 7 105 Eq/mL
for the groups displaying positive strand RNA in their PBMCs and from
25 ± 10 to 30 ± 15 105 Eq/mL for the groups who did
not display positive strand RNA in these cells. Patients with titers
below the level of detection (ie, <2.105 Eq/mL)
represented a very limited number (n=6) and were mostly found in the
group 2a/c infected patients (n=3). The mean titers of coinfected
patients were comparable to those from other groups of infected
patients (42 ± 17 105 Eq/mL).
Detection of negative strand RNA (2B).
Negative strand RNA could also be detected in all three groups of
patients although to a lower frequency than observed for detection of
the positive strand RNA. Detection rates were 42%, 8%, and 83.3% in
the genotype 1, the nongenotype 1, and the coinfected group of
patients, respectively. More precisely, positive signals were found in
8:14 and 1:7 genotype 1b- and 3a-infected patients. The difference in
detection rates between the two first groups of patients was
statistically significant (P < .04). Viral loads for all
groups of patients were comparable. Excluding patients with titers
below the level of detection, mean titers varied from 33 105 Eq/mL for patients harboring an HCV negative strand in
their PBMCs and from 21 ± 7 to 29 ± 14 105 Eq/mL
for patients who did not harbor the HCV negative RNA.
Overall results indicate that, independent of the viral load and viral
genotype, HCV genomic sequences can be found in PBMCs of chronically
infected patients. Under the experimental conditions used in our study,
genotype 1-derived sequences were predominantly detected.
Analysis of the genotype distribution in PBMCs of coinfected
patients.
There were 6 patients in our study who were infected by two genotypes,
genotype 1a (n = 5) or 1b (n = 1) and 2a/c viruses, as confirmed with
two of the three genotyping assays used. To identify whether sequences
from one genotype or both were present in the PBMCs, we determined
genotypes of the PBMC-associated RNA using two genotyping assays, the
INNO-LIPA and CAP-PCR assays. Both assays were performed using both the
positive as well as the negative strand viral RNA as templates.
Genotypes were also similarly and concomitantly analyzed from
patients' sera. Results obtained with all samples were concordant
between the two assays used.
RT-PCR results obtained with the CAP-derived assay for one
representative patient (infected with a genotype 1a and 2a/c) are shown
in Fig 1. Lanes 1 through 4 illustrate the different size products
obtained when HCV positive strand RNA is amplified using this assay
from genotype 1a, 1b, 2a/c, 3a, and 4, respectively, containing sera
(chosen randomly). The expected size product for genotype 1a is 125 bp;
for 1b, 141 bp; for 2, 75 bp; for 3a, 87 bp; and for 4, 336 bp (see
Materials and Methods). In the serum as well as in the PBMCs of this
patient, products specific for genotype 1a and 2 were amplified from
the positive strand RNA (lanes 6 and 7, respectively). No product was
obtained after amplification of the negative strand RNA from the serum
of the patient (lane 8). On the other hand, a product specific for
genotype 1a (125 bp) could be amplified using the negative strand RNA
from PBMCs of the patient (lane 9) but no bands corresponding to
genotype 2 amplified products could be detected when amplification was performed from the negative strand RNA. Overall, identical results were
obtained for all 6 coinfected patients; the negative strand RNA for
genotype 2 viruses was never documented in PBMCs, whereas the one for
genotype 1 was systematically found. In contrast, products specific for
both genotypes were amplified from PBMCs of the patients when PCR was
performed from the positive strand RNA.
In conclusion, the data indicate that different genotypes/subtypes can
be identified in a biological sample depending on the (genomic) RNA
template used in the assay.
Detection of HCV RNA sequences in different subsets of hematopoietic
cells.
In hematopoietic cells, three cellular subsets
theoretically have the capacity to
phagocytose viral particles: monocytes, macrophages, and
granulocytes. Because the presence of HCV genomic RNA in PBMCs was
clearly documented, we investigated which hematopoietic cell type may
actually harbor HCV sequences and whether viral sequences could be
documented in cells without phagocytic skills. We purified three
different peripheral hematopoietic cell subsets from total PBMCs of 11 different patients: monocytes (and macrophages), B lymphocytes, and T
lymphocytes. The purification protocol that we followed resulted in the
purification of a granulocyte-rich fraction in addition to the CD15
fraction (a fraction that could still contain some activated B cells
and monocytes). In our study, natural killer cell
populations (CD2+, CD3) were not
selected and would likely be found in the CD3 population as well as the
negative fraction. Results of the detection of HCV sequences are shown
in Table 1.
Positive strand RNA was detected in three different cell subsets:
granulocytes, monocytes, and B lymphocytes from respectively 89%,
56%, and 63% of patients. Even when a low number of cells were used
in the PCR (3 × 104 to 6 × 104)
positive signals could be detected. Positive strand RNA was not
detected in T lymphocytes (0/7), even when more than 4 × 106 cells were tested, or in thrombocytes (0/6). Similarly,
the negative strand RNA could be amplified from granulocytes,
monocytes, and B lymphocytes from respectively 3:10, 1:10, and 2:9
patients. The two patients harboring negative strand RNA in B
lymphocytes displayed well-characterized severe cryoglobulinemia. Thus,
different PBMC-associated cell subsets are capable of harboring HCV
genomic sequences, either the positive or the negative strand RNA.
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DISCUSSION |
Two main conclusions can be drawn from our study: first, that PBMCs
from HCV chronic carriers can harbor HCV genomic sequences whatever the
infecting viral genotype and viral load; second, that such sequences
can be found in different hematopoietic cell subsets. Because of the
typically low viral loads associated with HCV infections, PCR remains
to date the most reproducible and sensitive technique to track the
putative presence of virions, passively absorbed or replicating, in
cells. We provide evidence that both genomic strands, detected using
highly specific assays, are present in specific hematopoietic cell
subsets of patients infected with at least three genotypes (1, 2, and
3) and four specific subtypes (1a, 1b, 2a/c, and 3a). This constitutes
an original observation compared with previous studies reported in the
literature as none of these studies included the systematic characterization of patients' viral genotypes. HCV's well-described genomic diversity (for review see Bukh et al4) makes it
particularly difficult to develop experimental conditions equally
sensitive and specific for all the existing genotypes. We took
particular care in combining conditions providing the most specific
amplification of sequences (in particular for the negative strand RNA)
together with keeping comparable sensitivity. This could be achieved
within a difference of one log factor when templates from the different genotypes (1, 2, 3, and 4) were tested. Both assays used for
amplification of the positive and negative strand RNA had an identical
sensitivity (102 template copies) for all genotypes tested,
except in only three instances for which a sensitivity of
103 template copies was reached: for genotypes 1a and 3a
positive strand RNA and for genotype 4c negative strand RNA as
evaluated using synthetic transcripts (Fig 2). In our study, there was
a trend toward a higher detection rate for both the positive and the
negative strand RNA in patients from the genotype 1 (a or b) infected
group (ranging from 42% to 78.9%). This difference was statistically
significant when compared with the detection rate found in the
nongenotype 1 infected group (P < .001). Although a larger
number of patients from the nongenotype 1 infected type should be
studied, it remains nonetheless that this observation is intriguing.
Genotype 1b isolates have been reported to be associated with a more
common and active disease on the graft after liver transplant7 and a poorer response to interferon
treatment29 (although they are characterized
by viral loads equivalent to those found for other
genotypes).30 It is tempting to speculate that extrahepatic
reservoirs could be favored by viruses from this genotype. Such
replication advantage appears independent of the serum viral load
because we could not show any influence of such viral load on detection
of HCV sequences in PBMCs of the patients (see Fig 3). Our observation
further confirms, in that respect, that there is no association between
viral load and viral genotype as was previously
suggested.30
Although our data are only representative of the conditions and
patients used in our study, we observed that in cases of multiple infections, at least with genotypes 1a or b and 2a/c, the genotypes identified in the serum of the patients may not reflect the genotype(s) of viral sequences found in the PBMCs of the same patient. Because the
sensitivity of detection of genotype 2 negative strand RNA is as good
as the one observed for genotype 1a and 1b sequences (102
template copies, see Fig 2), it is difficult to attribute the lack of
detection of genotype 2 sequences to a poorer sensitivity of the
detection assay. Another explanation could be due to a difference in
the viral loads of PBMC-associated HCV sequences depending on the
genotype involved. Alternatively, if detection of HCV negative strand
RNA is indeed a reflection of active replication, the data would
reflect a preferential tropism of genotype 1 isolates for PBMCs
compared with genotype 2 isolates. Shimizu et al31 have
recently described the existence of a preferential tropism of specific
HCV viral quasispecies for PBMCs, an observation that reinforces the
concept of PBMCs versus hepatocyte-adapted isolates. Cases of dual
infections are not commonly found even in populations with increased
risks of infection.32,33 Although additional such cases
should be studied, extension of our analysis remains a difficult task
to perform.
The detection of HCV sequences, either the positive or the negative
strand RNA, does not constitute a sufficient argument to prove
replication. The detection of positive strand RNA could simply reflect
adsorption of viral particles on the cellular surface membrane. Such
adsorption could be due to Fc-receptor-mediated binding of
antibody-coated viral particles. This is consistent with our
observation that monocytes, granulocytes, and B cells of the patients
were indeed found to harbor HCV positive strand RNA. It is also
consistent with the observation that immune complexes are typically
found in sera of HCV-infected patients.34,35 Detection of
HCV negative strand RNA being the theoretical intermediate of
replication could be a stronger indication for putative replication in
a cell reservoir. However, one should keep in mind that phagocytic cells could take up cellular debris or even entire cells with actively
replicating HCV. Nonetheless, because the negative strand RNA was
absent from the sera of all patients, trapping of this putative
replication intermediate from serum would be unlikely. We observed that
clinical features showing liver damage (enzyme levels, histological
score) were similar in the group of patients harboring negative strand
RNA in their PBMCs compared with the one without evidence of negative
strand RNA in PBMCs (Lerat, personal observation). This
could suggest that the release of infectious virus particles
and/or negative strand RNA from damaged hepatocytes may not be
the source of the negative strand RNA detected in PBMCs and that
detection of such an intermediate may indeed reflect active
replication.
We were able to detect HCV genomic sequences in phagocytic cells (PMNL
and M/M) as well as in B lymphocytes. Previous reports had described
evidence of HCV RNA in B lymphocytes18-20,33 and monocytes20 of chronic carriers. Examples of viruses
replicating in phagocytic cells have been reported, eg,
HIV36; Dengue virus37,38; HSV-139;
and CMV.40,41 A favored tropism of HCV for B cells may
clearly be a factor participating in the unusual high-association rate that has been reported between HCV infection and mixed
cryoglobulinemia.42 In our study, both patients harboring
HCV negative strand sequences in B cells suffered from severe
cryoglobulinemia. Such tropism may also have implications in the
development of lymphomas recently described in cases of HCV-infected
carriers.43 We could not document HCV RNA in T lymphocytes
and platelets. Although T lymphocytes constitute the most abundant
population in PBMCs, they thus appear to play no role in the tracking
of extrahepatic viral sequences.
Replication of viruses in PBMCs of patients has been proposed
and/or shown for members of other hepatitis viruses such as hepatitis A and B viruses (HBV).44-47 A recent study, in
particular, clearly describes the presence of all encoded HBV
transcripts in PBMCs of chronically infected
patients.48 The capacity of HCV to enter and
possibly replicate in cells from the immune system and the possible
dysregulation of normal cell functions associated with such an invasion
could be one factor responsible for the establishment of chronicity
characterizing infection by this virus.
| |
FOOTNOTES |
Submitted July 21, 1997;
accepted December 31, 1997.
H.L. and S.R. have equally contributed to the study.
Supported in part by l'Association pour la Recherche sur le Cancer;
H.L. is supported by a grant from INSERM.
Address correspondence to Geneviève Inchauspé, INSERM U271,
151 Cours Albert Thomas, 69424 LYON Cedex 03, FRANCE; e-mail: inchauspe{at}lyon151.inserm.fr.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
| |
ACKNOWLEDGMENT |
We thank F. Zoulim (INSERM U271, Lyon) for assisting in the recruitment
of patients and JC. Tremisi (Blood Center, Lyon) and C. Biron (Bone
Marrow Transplant Unit, Lyon) for providing some patient samples and A. Fatmi for performing the sequencing of samples. We are grateful to C. Bain, S. Lemon, A.M. Prince, and M. Beard for critical review of the
manuscript and P. Simmonds and L. Jarvis for help with the genotyping
of cloned sequences. The authors are thankful to the Chiron Corporation
and J.P. Bonn for providing us with the bDNA Chiron HCV RNA 2.0 assay.
| |
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M. Hsu, J. Zhang, M. Flint, C. Logvinoff, C. Cheng-Mayer, C. M. Rice, and J. A. McKeating
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A. Dolganiuc, K. Kodys, A. Kopasz, C. Marshall, T. Do, L. Romics Jr., P. Mandrekar, M. Zapp, and G. Szabo
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M. S. Sulkowski and D. L. Thomas
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T. Laskus, M. Radkowski, A. Bednarska, J. Wilkinson, D. Adair, M. Nowicki, G. B. Nikolopoulou, H. Vargas, and J. Rakela
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M. Radkowski, J. Wilkinson, M. Nowicki, D. Adair, H. Vargas, C. Ingui, J. Rakela, and T. Laskus
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F. Penin, C. Combet, G. Germanidis, P.-O. Frainais, G. Deléage, and J.-M. Pawlotsky
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Abstract
Introduction
Abstract