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Journal of Virology, October 1998, p. 7772-7784, Vol. 72, No. 10
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Effect of Immune Activation on the Dynamics of
Human Immunodeficiency Virus Replication and on the Distribution of
Viral Quasispecies
Mario A.
Ostrowski,1,*
David C.
Krakauer,2
Yuexia
Li,1
Shawn J.
Justement,1
Gerald
Learn,3
Linda A.
Ehler,1
Sharilyn K.
Stanley,1
Martin
Nowak,2 and
Anthony S.
Fauci1
Laboratory of Immunoregulation, National
Institute of Allergy and Infectious Diseases, National Institutes
of Health, Bethesda, Maryland1;
Department of Zoology, University of Oxford, United
Kingdom2; and
Department of
Microbiology, University of Washington, Seattle,
Washington3
Received 17 February 1998/Accepted 7 July 1998
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ABSTRACT |
Virus replication in a human immunodeficiency virus (HIV)-infected
individual, as determined by the steady-state level of plasma
viremia, reflects a complex balance of viral and host factors. We have
previously demonstrated that immunization of HIV-infected individuals
with the common recall antigen, tetanus toxoid, disrupts this steady
state, resulting in transient bursts of plasma viremia after
immunization. The present study defines the viral genetic basis for the
transient bursts in viremia after immune activation. Tetanus
immunization was associated with dramatic and generally reversible
shifts in the composition of plasma viral quasispecies. The viral
bursts in most cases reflected a nonspecific increase in viral
replication secondary to an expanded pool of susceptible CD4+ T cells. An exception to this was in a patient who
harbored viruses of differing tropisms (syncytium inducing and
non-syncytium inducing [NSI]). In this situation, immunization
appeared to select for the replication of NSI viruses. In one of three
patients, the data suggested that immune activation resulted in the
appearance in plasma of virus induced from latently infected cells.
These findings illustrate certain mechanisms whereby antigenic
stimulation may influence the dynamics of HIV replication, including
the relative expression of different viral variants.
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INTRODUCTION |
The pathogenic mechanisms
involved in the regulation of human immunodeficiency virus (HIV)
replication in vivo are multifactorial and complex (27).
Following primary HIV infection, most patients achieve a steady-state
level of plasma viremia within 4 months to 1 year, which is referred to
as the viral set point (39, 40, 46). However, HIV
replication is a dynamic process in which the half-life of virus in
plasma has been reported to be approximately 5.7 h
(44). It has been suggested that approximately 99% of virus
in the plasma is derived from recently infected CD4+ T
cells that have a life span of approximately 2.2 days (36, 44,
52). HIV type 1 (HIV-1) replication is influenced by a combination of viral and host factors (26). Viral
characteristics that sustain replication within a patient include
replicative fitness, predilection for mutation, and cell tropism
(26). Important host factors that can induce or
suppress viral replication include the HIV-specific immune
response, cellular activation, and the effects of endogenous cytokines
and chemokines (26). Thus, the net level of viral
replication, as reflected by plasma viremia, is the result of a complex
balance of multiple forces. However, the major influences that drive
HIV-1 replication in vivo within individual patients have not been
fully delineated.
HIV-1 replication is enhanced by cellular activation (3, 8,
54). We and others have reported that immune activation resulting
from exogenous stimuli such as vaccinations or coinfections induce
substantial increases in plasma viremia (7, 9, 32, 35,
42, 48, 49). In this regard, immunization of
HIV-infected individuals with the recall antigen tetanus
toxoid induced transient bursts in plasma viremia over a 6-week period
(48). The mechanism for this burst of virus in plasma is
uncertain but is presumably due to a transient increase in the
susceptible pool of activated CD4+ T cells and/or the
secretion of proinflammatory cytokines that are capable of inducing the
expression of HIV from already infected cells
(12).
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FIG. 1.
Kinetics of plasma viremia postimmunization in patients
2 through 4. Patient 1 was mock immunized. Viral RNA copies
(103/ml) of plasma are annotated on the ordinate, and days
postimmunization are annotated on the abscissa. Plasma samples were
obtained for sequencing of cDNA at baseline, during peak viremia, and
at last follow-up and are indicated ( ). PBMC samples obtained for
proviral sequencing at day 0 in patients 2 through 4 and at day 30 in
patient 2 are indicated (#). LNMC samples obtained for proviral
sequencing at days 0 and 30 postimmunization in patient 2 are indicated
(*). All patients were asymptomatic and were not receiving
antiretroviral agents.
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The present study investigates the viral genetic basis for the
transient bursts in viremia after immune activation. We asked whether perturbations of the steady-state plasma viremia
resulting from an activation stimulus such as immunization with a
common recall antigen are also reflected in changes in the relative
appearance of viral quasispecies. We also considered two
hypotheses: (i) the viral burst reflects an increase in a selectively
favored subpopulation of viruses, or (ii) the viral burst reflects a
nonspecific increase in viral replication due to an increase in the
pool of susceptible CD4+ T cells. In addition, we explored
the role of induction of latent viruses after immunization in the
perturbation of plasma viral quasispecies.
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MATERIALS AND METHODS |
Patients and samples.
The study was carried out according to
a clinical protocol approved by the institutional review board of
the National Institute of Allergy and Infectious Diseases. All patients
were asymptomatic HIV-1-infected individuals who were receiving no
antiretroviral medications. Briefly, participants were either given a
0.5-ml tetanus booster intramuscularly (patients 2 to 4) (Wyeth-Ayerst Laboratories, Marietta, Pa.) or were mock immunized (patient 1). Blood
was drawn on the day of the injection and sampled at least six times
thereafter over a 6-week period. Patient 2 underwent serial lymph node
biopsies before and after immunization. Of note, patients 2 and 3 in
this report correspond to patients 3 and 4, respectively, in a previous
report (48).
Virus isolation, quantitation of plasma viremia, and mononuclear
cell provirus.
Blood was collected in EDTA anticoagulant tubes
(Becton-Dickinson, Mountain View, Calif.) and was processed within
2 h of sampling. Peripheral blood mononuclear cells (PBMC) were
obtained by standard Ficoll centrifugation (LSM; Organon Teknika,
Durham, N.C.). The samples were CD8+ T-cell bead depleted
(Dynal, Lake Success, N.Y.) and cultured in the presence of 10 U of
interleukin-2 (Boehringer-Mannheim, Mannheim, Germany) per milliliter.
Cultures were monitored for 4 weeks for evidence of reverse
transcriptase (RT) activity, as previously described (48).
The ability of the isolated viruses to induce syncytia was assessed by
coculture of 100 50% tissue culture infective doses of the isolate
with 106 MT-2 cells as previously described (4).
Plasma HIV-1 RNA levels were measured by a quantitative competitive
reverse transcription-PCR assay as previously described
(48). Proviral burden was determined by a semiquantitative
PCR assay, using SK38 and SK39 primers to amplify the gag
region as previously described (48).
DNA preparation, cDNA synthesis, and PCR.
DNA was prepared
from PBMC by using the Puregene DNA isolation kit (Gentra Systems,
Minneapolis, Minn.). RNA was extracted from 500 µl of plasma with the
QIAamp HCV kit (Qiagen, Chatsworth, Calif.). RNA was incubated with
RNase-free DNAse (Boehringer-Mannheim, Mannheim, Germany) at 37°C for
20 min (1 U of RNA sample/µl) in DNase reaction buffer (50 mmol
Tris-Cl [pH 7.5]/liter, 10 mmol of MgCl2/liter, 40 U of
RNasin [Promega, Madison, Wis.]). The sample was then incubated at
80°C for 10 min to terminate the reaction. cDNA synthesis was carried
out in a total volume of 20 µl with 7 µl of the RNA sample, 40 U of
avian myeloblastosis virus (AMV) RT (Lifesciences, St. Petersburg,
Fla.), 40 U of RNasin, deoxynucleoside
triphosphate at a final concentration of 1 mM, and outer antisense
primer (ED12) at 1.5 µM final concentration in AMV RT buffer (at a
final concentration of 25 mM Tris-HCl, pH 8.3, 50 mM KCl, 2.0 mM
dithiothreitol, 5.0 mM MgCl2), with an incubation at 42°C
for 45 min, followed by 10 min at 70°C.
DNA and cDNA were amplified by using nested PCR by limiting dilution so
that single copies of DNA or cDNA could be amplified.
First-round
primers were ED5 (5'-ATGGGATCAAAGCCTAAAGCCATGTG-3',
nucleotides 6556 to 6581, HIV-1 HXB2) and ED12
(5'-AGTGCTTCCTGCTGCTCCCAAGAACCCAAG-3',
nucleotides
7822 to 7792, HIV-1 HXB2). Second-round reactions
used 1 µl of
first-round product as the template and primers DR7
(5'-TCAACTCAACTGCTGTTAAATGGCAGTCTAGC-3', nucleotides
6989 to 7020,
HIV-1 HXB2) and DR8
(5'-CACTTCTCCAATTGTCCCTCATATCTCCTCC-3', nucleotides
7637 to
7667, HIV-1 HXB2), as previously described (
18,
19).
The
final reaction amplifies ~650 bp of product spanning the C2
to V5
region of the HIV-1 envelope surface protein. PCRs and cycling
conditions were performed as previously described (
19).
Heteroduplex tracking assay (HTA).
Single-stranded probes
were generated by subjecting DR7-DR8-derived PCR products to an
additional five cycles of PCR in the presence of
[
-32P]dTTP, 33 µmol of each deoxynucleoside
triphosphate, primer DR8, and biotinylated primer DR7. After PCR, the
labeled DNA was bound to streptavidin-coated magnetic beads (Dynal,
Lake Success, N.Y.). Magnetic beads were washed, and the labeled strand
was disassociated with 8 µl of 0.1 N NaOH for 10 min. The probe was
neutralized by adding 40 µl of H2O, 4 µl of 0.2 N HCl,
and 1 µl of 1 M Tris-HCl (pH 7.4). Driver sequences were obtained by
nested PCR of cDNA or genomic DNA as described above. In PBMC samples
with low proviral load, multiple, independent nested PCR products were
pooled before analysis. Probes were mixed with the driver sequence at a
ratio of 1:100 in annealing buffer (100 mM NaCl, 10 mM Tris-HCl [pH 7.8], 2 mM EDTA), denatured at 94°C for 3 min, and then placed on ice for 5 min, followed by heating to 55°C for 5 min to form heteroduplexes. The resulting reaction mixtures were electrophoresed in
5% polyacrylamide gels (acrylamide/bisacrylamide ratio, 37.5:1) at 250 V for 3 h (19), stained with ethidium bromide to assure even migration of bands, dried, and scanned in a Molecular Dynamics PhosphorImager (Sunnyvale, Calif.).
Sequencing.
PCR products were directly sequenced by using
the ABI PRISM dye-terminator method and by using DR7 and DR8 primers
for forward and reverse reactions, respectively, according to the
manufacturer's specifications (Perkin-Elmer, Foster City, Calif.).
Automatic sequencing was performed on an Applied Biosystems Inc.
(Foster City, Calif.) 373 sequencer. Sequence editing and assembly were performed by using Sequencher, version 3.0 (Gene Codes Corporation, Inc., Ann Arbor, Mich.).
Phylogenetic analysis.
To root our sequences, envelope
(env) sequences from five patients from the United
States were used as outgroups in the analysis of the envelope genes
samples from each of the four immunized patients (three and one
control). Sequences were aligned, using Clustal W (50) with
default settings, and improved by hand. Phylogenetic reconstruction was
performed using MOLPHY (1), which first produces an
approximate phylogeny based on a neighbor-joining algorithm and then
improves the topology using local rearrangements under a maximum
likelihood model. The maximum likelihood algorithm uses the HKY85
matrix of nucleotide substitutions (34) and calculates bootstrap probabilities using the RELL procedure for local
rearrangements. Genetic distances between plasma-derived sequences for
each time point were calculated using DNADIST (28), based on
Kimura's two-parameter model. Mean diversity is calculated over all
pairwise distances for each time point. Genetic diversity along the C2 to V5 region was calculated for each patient's virus in plasma through
time. A standard entropy formula for diversity was employed in which
diversity = (ni/N) 
ln
(ni/N), where N is the total number
of nucleotides at a given site across sequences of an alignment, and
ni is the proportion of a nucleotide
i (i = A, C, T, and G).
Nucleotide sequence accession numbers.
Nucleotide sequences
obtained from patients 1 through 4 have been submitted to GenBank under
accession no. AF080698 through AF081019.
| |
RESULTS |
Immunization of subjects.
The clinical and laboratory profiles
of the four patients are summarized in Table 1. Figure 1 shows the
kinetics of plasma viremia in the mock-immunized (no. 1) and immunized
(no. 2 through 4) patients. Patients 2 through 4 demonstrated
significant increases in plasma viremia after immunization, ranging
from 7- to 14-fold above baseline. Expression of activation markers on
CD4+ T cells (i.e., HLA-DR and CD25) sampled from PBMC
of patients 2 through 4 increased after immunization (data not shown).
There were no significant changes in activation markers on
CD4+ T cells of the unimmunized, HIV-infected control
subject (data not shown).
In order to study HIV sequence populations in vivo we amplified single
molecules of virus (cDNA) or provirus after limiting
dilution of the
target sequence followed by a sensitive nested
PCR method (
18,
19,
47). The PCR product was then directly
sequenced. This
methodology has previously been validated by Simmonds
et al.
(
47) to study the relative proportions of virus variants
within patient samples. HIV cDNA sequences from plasma were obtained
in
all patients on the day of immunization, at the time of peak
viremia,
and at last follow-up when the level of plasma viremia
had returned to
baseline levels. As proviral DNA is largely composed
of latent or
nonreplicating viruses in a given patient (
47),
we were
interested in examining baseline proviral sequences to
determine if any
of these variants would be induced from the activation
of immunization.
Thus proviral sequences from PBMC were obtained
in patients 2, 3, and 4 on the day of immunization. Lymph node
biopsies were performed before
and after immunization in patient
2, and proviral sequences were
determined on these samples on
the day of immunization and 30 days
later. At least 20 sequences
were obtained per tissue compartment
(e.g., plasma versus PBMC)
at each sampled time point in all
immunized patients. In some
patients, nested PCR of undiluted cDNA or
proviral DNA was performed
and the products were analyzed by a
heteroduplex tracking assay
(see below) in order to confirm the nature
of viral populations
obtained by sequencing.
Quasispecies analysis.
In the analysis of our phylogenetic
trees, we defined a phyletic group as a topological cluster of at least
three sequences that generally but not always had bootstrap values
exceeding 90%. In all instances, differing phyletic groups within a
patient corresponded to distinct differences in the amino acid length
and/or composition of V3, V4, and/or V5 regions, suggesting that
viruses belonging to a particular group have distinct phenotypes in
vivo.
The phylogenetic reconstruction of plasma cDNA sequences obtained from
patient 1, who was mock immunized, is shown in Fig.
2, left. The majority of
viruses in plasma samples at all time
points belonged to a large group
(A), and a minority of viruses
formed two smaller clusters (B and C). A
small number of sequences
did not clearly fit into any defined subgroup
(identified as ?).
There was overall stability in the distribution
of the predominating
variants in this unimmunized patient
over a 30-day period (Fig.
2, right), with group A viruses
making up 62, 72, and 77% of plasma
sequences sampled at 0, 14, and 30 days post-mock immunization,
respectively. The differences in
group A frequency were not statistically
significant between time
points as determined by Fisher's exact
test. Group B sequences
at the 14-day time point were not detected
by sequence analysis. This
was in all likelihood due to the fact
that fewer sequences (<15 per
time point) were sampled in this
patient. Of note, HTAs performed on
PCR products obtained from
plasma cDNA samples using probes derived
from variant B and C
sequences revealed the presence of these minor
groups at all time
points (data not shown).
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FIG. 2.
(Left) Phylogenetic analysis of the C2 to V5 region of
all sequences (cDNA) sampled from patient 1. Viral sequences are shown
by colored symbols according to their sampling time and tissue
compartment. Three groups are indicated as A, B, and C; unclassified
sequences are indicated by ?. Bootstrap probabilities are shown by
percentages. *, This sequence has an artifactually long branch length
due to a deletion of the V3 region. The following prototype HIV
env subtype B strains were used as outgroups: M79345
(derived from a primary isolate), M96155 (from isolate 89.6), M89973
(YU-2), M65024 (HIVSFAAA), and M60472 (ADA). (Right) Frequency of
detection of sequence variants in patient 1 (mock immunized) in
sequential plasma samples. All sequences were derived from RNA.
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A total of 130 sequences were studied in patient 2; these consisted of
cDNA from plasma (days 0 and 8 and 7 weeks postimmunization),
provirus from PBMC (days 0 and 30 postimmunization), and provirus
from lymph node mononuclear cells (LNMC) (days 0 and 30 postimmunization).
Alignment of all sequences revealed the presence of
two distinct
populations of variants (A and B) in each tissue
compartment at
all time points studied (Fig.
3, left). Both groups
could be easily
distinguished by comparison of amino acid translations
of the
V4 regions (data not shown). Prior to immunization, the relative
frequencies of groups A and B in plasma were significantly different
from those in proviral LNMC (see the legend to Fig.
3, right)
and PBMC
(
P < 0.01 for each, Fisher's exact test). Group A
made
up 75% of sequences in plasma but was the minor population in
sequences derived from proviral PBMC and LNMC (Fig.
3, right).
Conversely, group B comprised the major proviral population within
PBMC
and LNMC and thus was more likely a reflection of the archival
population of viruses (
47,
52). In this patient, group A
presumably
represented a more recently evolved variant circulating in
the
plasma, although viruses from group B were also actively
replicating
and circulating in the plasma at the time of immunization.
At
the time of peak viremia (day 8), there was a reversal in the
relative proportions of virus populations in the plasma, with
group
B predominating (
P < 0.025, Fisher's exact
test). In contrast
to group A which increased 4-fold in the
plasma following immunization,
group B had increased 18-fold. By 7 weeks after immunization,
at a time when viremia approached
baseline values, there was a
trend toward a return to the original
distribution of plasma viruses.
There was considerable similarity of
the V3 loop in all sequences,
which had a macrophage-tropic (M-tropic)
pattern consistent with
the isolation of non-syncytium-inducing (NSI)
viruses in culture
(data not shown).
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FIG. 3.
(Left) Phylogenetic analysis of sequences (cDNA and
proviral) from patient 2. Samples from plasma represent RNA sequences,
and those from PBMC and LNMC are proviral. Note two major groups, A and
B, which are found in all compartments and time points sampled. Variant
A was predominant in plasma, and variant B was predominant in PBMC at
baseline. (Right) Frequency distribution of RNA variants in plasma and
proviral variants in PBMC before and after immunization. For
comparative purposes the proviral distributions of LNMC and PBMC at day
30 are not illustrated but are as follows: day 30 PBMC, group A, 35%,
and group B, 65%; day 0 LNMC, group A, 35% and group B, 65%; day 30 LNMC group A, 45% and group B, 55%.
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Patient 3, who was a slow progressor, showed a much more complex viral
phylogenetic tree (Fig.
4,
left) that comprised six
distinguishable groups
(arbitrarily termed A through F). At least
five groups were identified
in the PBMC proviral compartment (A,
B, C, D, and F) as well as
sequences which did not clearly fit
into any group (Fig.
4, right).
Twenty-two percent of proviral
PBMC sequences were not found in plasma
and likely represented
archival virus within latently infected cells
(groups F and ?).
Since groups A, B, C, and D were also detected
in the plasma,
these proviral sequences were also likely contributing
to active
viral replication. At the time of immunization, five groups
could
be identified in plasma, with groups B and C predominating. The
composition of variants in plasma before immunization and at peak
viremia was somewhat similar, although group C variants decreased
and
group D variants expanded at peak viremia. By day 42, however,
a
dramatic change in quasispecies makeup was seen, in which 80%
of virus
variants that were circulating at day 21 (groups B, D,
and E) were
replaced by groups A and C. In order to confirm these
quasispecies
shifts in patient 3, we performed HTAs on PCR products
from undiluted
samples obtained from plasma and PBMC at multiple
time points from
immunization. We elected to perform HTAs by using
probes made from
group B and D viruses, since these variants comprised
70% of viruses
at day 21 and were not seen at day 42. Probes made
from representative
group B and D sequences could distinguish
sequences obtained from other
groups (Fig.
5a). These probes were
then
employed against PCR products from undiluted samples taken
from PBMC
(proviral DNA) and plasma (cDNA) at days 0, 21, and
42 (Fig.
5b). By
HTA, group B and D viruses were not identified
in plasma from day 42, as illustrated by slower migration of heteroduplexes
at these time
points (Fig.
5), thus confirming our sequence analysis.
Interestingly,
a reduced signal intensity for variant D was observed
in PBMC sampled
at day 42, suggesting early clearance of this
variant from
the PBMC compartment.
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FIG. 4.
(Left) Phylogenetic analysis of sequences (cDNA and
proviral) from patient 3. Major groups are indicated as A through F. (Right) Frequency distribution of viral variants in plasma and PBMC
preimmunization and in plasma postimmunization in patient 3.
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FIG. 5.
(a) HTA of viral sequences (C2 through V5) obtained from
patient 3. After referring to the phylogenetic tree, a representative
sequence from group B or D was selected as a probe against other
variants. Probes derived from group B or D sequences were able to
easily distinguish sequences of the same group (B or D) from a panel of
sequences obtained from different groups in the same patient or from an
unrelated sample (lane U). The experiment was repeated three times,
using different representative sequences for probe and panel, with
similar results. *, probe lane; arrow, position of homoduplex
migration. (b) The effect of tetanus immunization on viral quasispecies
turnover in patient 3, as displayed by HTA. In order to determine the
turnover of groups B and D, an HTA was performed with the probes
described above. For plasma samples, RT PCR products amplified from
approximately 100 copies of HIV-1 RNA templates were used as the
driver. For PBMC samples, PCR products amplified from about 40 copies
fo HIV-1 genomic DNA were used as the driver. Arrow, position of
homoduplex migration.
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The phylogenetic reconstruction of sequences obtained from patient 4 revealed the presence of two major variants, A and B
(Fig.
6, left). Groups A and B
were clearly distinguishable in
the V3 region: the group A consensus V3
loop was
CTRPNNNTRRGIHIGPGSAFYATGDIIGDIRQAHC;
the
group B consensus V3 loop was
CTRPSINKRRHIHIGPGRAFYATDITGDIRQAHC.
The V3 loop
of B variants differed from that of A variants (at
positions 5, 6, 8, 11, 18, 24, and 27 from the cysteine on the
5' side of the V3
region) by having a greater number of positively
charged amino acids
and the loss of a potential N-linked glycosylation
site (position 6) in
group B. These changes have previously been
shown to be consistent with
a switch from an envelope of an NSI
to that of a syncytium-inducing
(SI) virus (
11,
21,
23,
31). The existence of SI viruses in
patient 4 was confirmed
by PBMC culture (Table
1). One sequence
obtained from plasma
at day 0 was a recombinant of both variants,
comprising an A V3
region and a B V4/V5 region (data not shown and
Fig.
6, left).
Prior to immunization, patient 4 had an approximately
equal distribution
of variants A and B in the plasma (Fig.
6, right).
At the time
of peak viremia (day 21), variant A (NSI) made up 95% of
viruses
(
P < 0.005, Fisher's exact test). By day 41, as plasma viremia
returned to baseline levels, the original
distribution of viruses
was observed. By extrapolating from absolute
plasma viral levels
(Fig.
1), the NSI variants had increased
13-fold following immunization,
whereas the SI variants
had essentially remained unchanged. Of
note, viruses of variant B (SI)
made up the greatest proportion
in the proviral PBMC compartment.
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FIG. 6.
(Left) Phylogenetic analysis of sequences from patient
4. Two major groups, A and B, are identified. *, a recombinant of
groups A and B. (Right) Frequency distribution of groups A and B in
plasma and PBMC preimmunization and in plasma postimmunization.
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We did not observe the emergence of new groups of variants during the
course of immunization in any of the patients studied.
We also did not
observe the emergence of SI-type envelopes in
those patients who had
NSI viruses in culture at baseline (patients
1, 2, and 3).
Diversity analysis.
No consistent trends in the mean virus
population diversity within plasma samples was observed with time (Fig.
7); however, reversible changes were
noted in two of three immunized patients. For example, in patient 2, virus diversity had significantly increased at peak viremia (3.1% at
day 0 to 3.5% at day 8), whereas in patient 4, virus diversity had
decreased at the time of peak viremia (3.3% at day 0 to 2.1% at day
21). These patterns of diversity were also reflected in the topology of
the phylogenetic trees. In patient 2, viruses from group B, which was
predominant at peak viremia, had longer branch lengths than group A
viruses, thus reflecting the mean diversity of the plasma virus
populations at this time point. In patient 4, a single group made up
the majority of the plasma viral population at peak viremia, which
accounted for the decrease in diversity at day 21.
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FIG. 7.
Mean diversity of plasma viral sequences over time in
patients 1 through 4. Error bars represent standard error of the
mean.
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In order to identify regions of the envelope that were responsible for
overall changes in mean diversity within samples, entropy
was plotted
against nucleotide position for all within sample
sequences. No
specific region along the envelope in patients 1
and 3 clearly
accounted for the majority of changes in mean diversity.
In patient 2, the increased diversity at peak viremia was due
to changes in the V4
region (data not shown). Interestingly, in
patient 4, the decrease in
diversity seen at peak viremia was
localized to the V3 region (Fig.
8) and reflected the decrease
of SI
variants.
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FIG. 8.
Diversity along env over time in patient 4. Entropy (ordinate) is plotted against nucleotide position.
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Accumulation rates of synonymous and nonsynonymous
substitutions.
Selection pressures on HIV-1 quasispecies were
determined during the course of immunization by analyzing the
synonymous (amino acid preserving) and nonsynonymous (amino acid
changing) nucleotide substitution patterns (41). The mean
numbers of nucleotide substitutions per synonymous site,
ds, and per nonsynonymous site,
dn, for all pairwise comparisons within
sequences sampled at each time point were determined. Thus, all
comparisons were of intrasample sequences against a consensus sequence
from the same sample. Generally, a
dn/ds ratio of <1 implies that
purifying selection for replication fitness predominates, whereas a
dn/ds ratio of >1 implies strong positive selection for amino acid diversification, as might be imposed
by the immune system or by the availability of specific target cells
(37).
The unimmunized patient, no. 1, revealed
dn/ds ratios that indicated no
selection (neutrality) at variable sites (Fig.
9).
In patient 2,
dn/ds ratios at all time points in
plasma and lymph
nodes were consistent with a pattern for purifying
selection throughout
the course of immunization. Patient 3 revealed a
pattern similar
to that of mock-immunized patient 1. The most
interesting findings
were in patient 4, who had both NSI and SI
viruses. Prior to immunization
and at last follow-up, when the viremia
was at steady state,
dn/ds ratios
were >1, indicating strong positive selection for amino
acid change.
During peak viremia, the
dn/ds ratio
was close to
unity, implying neutral drift at variable sites and
suggesting
a loss of the selective pressure that was driving the
fixation
of nonsilent mutations prior to immunization. We considered
the
possibility that the high
dn/ds
ratios observed at days 0 and
42 were a consequence of the presence of
T-tropic (SI) viruses
within the sample. It has been previously
postulated that SI viruses
may be under greater immunologic pressure
than NSI viruses within
a given patient, thus driving a higher
nonsynonymous substitution
rate in SI viruses (
6). However,
upon comparison of
dn and
ds values between all M-tropic (group A) and
T-tropic variants
(group B) obtained from all time points in this
patient, both
groups showed
dn/ds
ratios consistent with neutral evolution (Fig.
9). Thus, the increased
dn/ds ratios observed in plasma
sequences
obtained from baseline and at last follow-up are not the sole
consequence of the presence of SI viruses in the samples but reflect
selection in some subset of the sequences analyzed.
View larger version (25K):
[in this window]
[in a new window]
|
FIG. 9.
Effect of tetanus immunization on synonymous
(ds) and nonsynonymous
(dn) substitution rates. Intrasample
ds and dn values for
patients 1 through 4. Significant
dn/ds ratios away from neutrality,
i.e., dn/ds either <1 or >1, are
defined as those plots with error bars which have significantly
deviated either below or above the x = y line,
respectively.
|
|
| |
DISCUSSION |
The present study has characterized viral sequence variation and
evolutionary relationships in three HIV-infected patients after
immunization with a common recall antigen. Antigen-specific immune
activation induces changes in the host characterized by proliferation
of antigen-responsive cells, elaboration of cytokines, and subsequent
proliferation of bystander cells (10, 33, 51). Such a state
of immune activation results in an increase in the number of available
activated CD4+ T cells that serve as susceptible targets
for HIV. We tested this hypothesis in some detail by searching for any
systematic trends in the growing virus population. We asked whether
immunization had temporally influenced the source of plasma virus
(phylogenetic analysis), virus diversity (diversity plots), selection
pressures (ds/dn), and precise
positions of new substitutions along the env sequence
(entropy).
A common feature was that the perturbation in plasma viremia induced by
the activating stimulus of the immunization was consistently associated
with significant perturbations in the relative predominance of viral
quasispecies detectable in the plasma of immunized subjects. Although
the unimmunized patient (no. 1) demonstrated a shift in a minor plasma
virus variant over time, this change was not as dramatic as that seen
in the immunized patients. The combined observations of the relative
stability of the major quasispecies variant in the mock-immunized
subject (patient 1) and the transient and apparently reversible nature
of the plasma quasispecies shifts in patients 2 and 4, strongly suggest
that the immunization had actually influenced the quasispecies
distributions in our patients. The quasispecies changes observed in
patient 3 are noteworthy. We were able to confirm, using the HTA, that
80% of virus variants that were present during the time of peak
viremia (day 21) were cleared from the plasma by day 42. This rate of
clearance of variants over a 3-week period is not altogether surprising
given our current understanding of viral turnover (36, 44,
52).
We asked whether immune activation had provided a replicative
advantage to certain virus variants. In most cases we were unable to demonstrate a consistent pattern of selection. We could not demonstrate consistent changes in diversity, nor were we able to
demonstrate an increase in the dn/ds
ratio as might be expected with an increase following the emergence of
an escape mutant. No common set of amino acid substitutions along
env could explain the increase in viremia. A possible
explanation for this is that during immunization a subset of
CD4+ cells becomes activated and that virus release from
these cells is amplified, irrespective of the virus genotypes they
contain. This leads to a change in the distribution of variants but not to any evidence of selection and raises the notion that the shifting waves of diversity observed during infection are attributable to
various selection pressures on the CD4+ T cells themselves
and not on the specific virus that they harbor. For example,
immunization will activate tetanus-specific CD4+ T cells
and resident virions will incidentally become amplified. This idea has
recently been supported by Cheynier et. al. (13) who
demonstrated in the macaque model that the dynamics of simian immunodeficiency virus quasispecies after BCG immunization was a
reflection of those viruses found in circulating BCG-specific CD4+ T cells.
Specific selection of viruses, may, however, be operating in those
patients who harbor variants of differing tropisms, as seen in patient
4. Prior to immunization, equal proportions of NSI and SI viruses were
circulating in the plasma, whereas after immunization the induced burst
in viremia was due to an expansion of NSI viruses. This was an
unexpected finding since SI viruses generally replicate to a greater
degree than NSI viruses in tissue culture (4). In this
patient, diversity of the V3 region transiently decreased after
immunization. The higher rates of nonsynonymous substitutions observed
(dn/ds of >1) in plasma prior to
immunization and at last follow-up (day 42) suggest that the selective
pressures that might have been driving the fixation of new mutants at
baseline were relaxed during the burst of viremia. These findings
suggest that prior to immunization, strong selective pressures were
acting on both the NSI and SI viruses and that the sudden availability of a new population of activated CD4+ T cells following
immunization allowed the NSI population to expand without constraint.
The explanation for why NSI viruses would have an apparent replicative
advantage over SI viruses in the setting of immune activation is not
clear. A similar situation has been described in patients who, during
acute HIV infection, were initially infected with mixtures of SI and
NSI viruses and then were subsequently shown to harbor only NSI strains
(17). The NSI virus phenotype has been correlated with
macrophage tropism and usage of the CCR5 HIV coreceptor, whereas the SI
phenotype has been correlated with T-cell-line tropism and usage of the CXCR4 HIV coreceptor (2, 14, 16, 22, 24, 25, 29, 45). It has
been demonstrated that CCR5 expression is markedly upregulated and
CXCR4 expression is downregulated on activated CD4+ T cells
(5, 43). This observation lends credence to the possibility
that tetanus immunization transiently increased the pool of activated
CD4+ T cells that express CCR5 and little or no CXCR4, thus
favoring the replication of viruses that solely use CCR5 as a
coreceptor, i.e., M-tropic (NSI) viruses. Although material was not
available to study coreceptor expression in patient 4, we have
subsequently observed modest transient increases in CCR5 expression and
decreases in CXCR4 expression in both HIV-infected individuals
and healthy volunteers after tetanus immunization (data not
shown). In addition, we are currently determining the exact coreceptor
usage of the envelope variants obtained in patient 4 in order to
address this issue further. It is also unclear what role the recent
discovery of other coreceptors that appear to be used by NSI and SI
strains play in the setting of immune activation (2, 14, 20, 24, 29, 38).
We asked what contribution latent viruses play in the induced viremia
postimmunization. Studies by Simmonds et al. (47) and Wei et
al. (52) have demonstrated that turnover of proviral DNA in
PBMC lags behind turnover of plasma cDNA and that the majority of
proviral DNA in PBMC reflects archival or latent viruses that had
previously appeared in the plasma. It is also important to recognize
that sources of provirus within lymph nodes include latently infected
CD4+ T cells, long-lived macrophages, and dendritic cells,
which may all potentially be induced to produce virus (15,
36). Of the three immunized patients described in this report,
patients 2 and 4 manifested a quasispecies distribution in PBMC (and
LNMC in patient 2) proviral samples that could be easily distinguished from that in plasma, thus confirming previous findings (47). In patient 3, a small group of proviral variants (~20%) could be
distinguishable from those in plasma. If immune activation did indeed
reactivate latently infected cells to a significant extent, one might
then have expected to detect the predominating proviral PBMC variants
in the plasma after immunization. This was in fact the case with
patient 2, in whom the predominant variant (B) within the proviral PBMC
(and LNMC) compartment emerged as the predominant variant in the plasma
during peak viremia. The higher baseline proviral load seen in patient
2 (Table 1) might have allowed a greater opportunity for proviruses to
become reactivated and to significantly contribute to the actively
replicating pool of viruses in the plasma. This pattern of quasispecies
shift was not observed in either patient 3 or patient 4; thus, in these two patients, the preexisting major variants in the plasma were expanded after immunization, suggesting that the transient increase in plasma viremia that resulted from immune activation was
merely acceleration of a process that was already accounting for
the bulk of plasma virus as opposed to a shift from one source of virus
replication to another.
These observations may have considerable relevance to the pathogenesis
of HIV disease. Our observations confirm the dynamic nature of HIV
replication in vivo (36, 44, 52). The fact that immune
activation appears to, in most cases, nonspecifically amplify viruses
of differing genotypes likely contributes to the great diversity
observed in vivo. The exception to this may be in patients who harbor
viruses of differing tropisms; the observation that immune
activation favors viruses of an NSI phenotype over SI strains may
explain in part the paucity of SI strains observed during the early
stages of HIV. Finally, the fact that latently infected cells may
contribute to plasma viremia under certain circumstances such as immune
activation is of particular interest in light of the recent
observations that a stable reservoir of resting, latently infected
CD4+ T cells that can be induced to express
replication-competent virus persists over extended periods of time in
patients whose plasma viremia has been driven to below detectable
levels by highly active antiretroviral therapy (15, 30, 53).
| |
ACKNOWLEDGMENTS |
We thank Tae-Wook Chun, Susan Moir, Lin Qi Zhang, Raj
Shankarappa, and Jim Arthos for helpful discussions. We thank MaryBeth Daucher and Colombe Chappey for help with DNA sequencing. We are also
grateful to the four patients who committed their time and effort to
this study.
This work was supported by NIH grants to G.L. (A132885 and
A127757).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Laboratory of
Immunoregulation, National Institute of Allergy and Infectious
Diseases, NIH, Bldg. 10, Rm. 6A11, Bethesda, MD 20892. Phone: (301)
402-2618. Fax: (301) 402-4122. E-mail: mostrowski{at}nih.gov.
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Journal of Virology, October 1998, p. 7772-7784, Vol. 72, No. 10
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
