| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
*
Gladstone Institute of Virology and Immunology, San Francisco, CA 94141, and Departments of Microbiology and Immunology and Medicine, University of California, San Francisco, CA 94110;
Chemocentryx, Mountain View, CA 94070; and
Department of Obstetrics, Gynecology, and Reproductive Sciences, University of California, San Francisco, CA 94110
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Given the difficulty of obtaining and studying bone marrow and thymic
tissue from humans, the heterochimeric SCID-hu mouse was developed (8, 9). This model is constructed upon implantation of either human fetal
bone marrow (10) or human fetal liver and fetal thymus (11) into the
immunodeficient C.B-17 scid/scid mouse. In the case of the
more extensively studied SCID-hu Thy/Liv mouse, multilineage human
hemopoiesis, including T lymphopoiesis, proceeds for a period of 
12
mo. Human thymopoiesis in the SCID-hu model appears indistinguishable
from normal human thymopoiesis, with normal distribution of thymocyte
subpopulations, appropriate induction of negative selection, and
development of a diverse TCR repertoire (11, 12, 13). Perhaps most
importantly, relatively large numbers (50) of SCID-hu Thy/Liv mice
can be prepared from the same donor of human tissue, facilitating the
design of controlled experiments that investigate the pathogenic
properties of HIV-1 in vivo.
Early studies evaluating the effects of HIV-1 on the SCID-hu Thy/Liv organ revealed qualitative interstrain variations in viral pathogenesis: tissue culture-adapted isolates (e.g., HXB2) were noninfectious while primary isolates or molecular clones that had not been extensively passaged in vitro (e.g., JR-CSF and NL4–3) were infectious (14, 15, 16, 17, 18, 19). Among the latter HIV-1 isolates, nonsyncytium-inducing (NSI)4 (generally, CCR5-using) viruses were observed to slowly replicate in the organ and to be minimally cytopathic. In contrast, syncytium-inducing (SI; generally, CXCR4-using) viruses were observed to rapidly induce thymocyte depletion (20), an effect due to preferential infection and destruction of intrathymic (CD3-CD4+CD8-) T progenitor cells (21). Notably, viruses such as NL4–3 have also been found to deplete mutilineage and lineage-restricted CD34+ hemopoietic progenitor cells in the Thy/Liv graft. Such depletion occurs even earlier than thymocyte depletion and appears to be effected by indirect means (22).
HIV-1 variants may have differential effects on the hemopoietic system
because they are tropic for different cells in the multilineage tree.
As has been demonstrated over the past several years, tropism is
determined in part by cell surface expression of CD4 and certain
chemokine receptors, such as CCR5 and CXCR4 (23). CXCR4 is the receptor
for SDF-1
, an -chemokine that attracts lymphocytes and monocytes
(24, 25, 26). CCR5 binds the ß-chemokines macrophage inflammatory
protein-1, macrophage inflammatory protein-1ß, and RANTES, which
attract monocytes, T cells, NK cells, basophils, eosinophils, and
dendritic cells (23). In the early stages of HIV-1 disease, most virus
particles can use only CCR5 as the coreceptor and thus infect only
those cells that express CCR5 (27, 28, 29). Later, in the symptomatic
stages of HIV-1 disease, virus particles often arise that can use both
CCR5 and CXCR4, expanding the target cell range to cells that express
CCR5 and/or CXCR4 (27, 28, 29, 30).
It is still not clear which cell subpopulations in the T cell lineage express CXCR4 or CCR5 and, hence, might be targets for infection with strains of HIV-1 recognizing one or both receptors. While inferences about receptor expression can be made based on transcript analyses (31) or responses to chemokines in vitro (23), cell surface CXCR4 and CCR5 proteins can be more directly measured using specific Abs in flow cytometry or immunohistochemistry. Using such Abs, T cells in the bloodstream exhibiting a naive phenotype have been shown to express CXCR4 but not CCR5, while a subset of T cells expressing a memory phenotype expresses CCR5 but not CXCR4 (32). In addition, variations in the levels of CCR5 on peripheral T cells correlate with variations in the infectability of those T cells by a CCR5-using strain of HIV-1 (33), suggesting that the density of surface CCR5 may be a rate-limiting factor for the infection of peripheral T cells. Visualization of CXCR4 or CCR5 on the surface of bone marrow hemopoietic cells has not been reported, but CXCR4 (34) has been found on CD3+CD4+CD8+ thymocytes.
To help understand the basis of the differential infectivities of bone marrow cells and thymocytes, we used flow cytometry to determine the surface levels of CXCR4 and CCR5 on cells at different stages of T cell differentiation. This cell lineage included three discrete stages of CD34+ cells in the fetal bone marrow, five different subpopulations of fetal thymocytes, and naive and memory T cells in adult and cord blood. We found that the surface levels of CXCR4 and CCR5 vary widely among the cell types analyzed, indicating that the two coreceptors are cyclically up-regulated and down-regulated during the course of T cell differentiation. These findings may have important implications for the roles of chemokines in T cell production and function as well as for our understanding of HIV pathogenesis.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
A number of fluorescent mAbs were used in these studies. FITC-conjugated anti-human CD4, CD10, CD38, CD45, and CD45RA; phycoerythrin (PE)-conjugated anti-human CD2, CD4, CD8, CD16, CD19, CD20, and CD56; peridinin chlorophyll protein (PerCP)-conjugated anti-human CD3; allophycocyanin-conjugated anti-human CD34; and FITC-conjugated goat anti-mouse Ig were obtained from Becton Dickinson (San Jose, CA). Tri-color (TC)-conjugated anti-human CD8 and allophycocyanin-conjugated anti-human CD3 were obtained from Caltag (Burlingame, CA). PE-Texas Red (ECD)-conjugated anti-human CD8 was obtained from Coulter (Miami, FL). PE-conjugated anti-human glycophorin A was obtained from Immunotech (Westbrook, ME). PE-conjugated anti-human CXCR4 clone 47718 and unconjugated anti-human CXCR4 clones 47701 and 47712 were gifts from Monica Tsang (R&D Systems, Minneapolis, MN). These three mAbs recognize epitopes on the second extracellular loop of CXCR4, with contributions from the first extracellular loop (M. Tsang, unpublished observations), and do not stain cells expressing CXCR1, CXCR2, CXCR3, CCR2, or CCR5 (M. Tsang, unpublished observations, and data not shown). The anti-human CXCR4 clone 12G5 (35) was a gift from James Hoxie (University of Pennsylvania, Philadelphia, PA). The anti-human CCR5 clone 3A9 (33) was a gift from LeukoSite (Cambridge, MA); the mAb was conjugated with PE using a commercial kit (Prozyme, San Leandro, CA). Isotype control Abs for the CXCR4 and CCR5 mAbs consisted of unconjugated mouse IgG2a (Dako, Carpinteria, CA) and PE-conjugated mouse IgG2a and IgG2b (Caltag). Isotype control Abs for the anti-CD mAbs consisted of TC-conjugated mouse IgG2a (Caltag), FITC-conjugated mouse IgG1 and IgG2a, PE-conjugated mouse IgG1 and IgG2a, PerCP-conjugated mouse IgG1, and allophycocyanin-conjugated mouse IgG1 (Becton Dickinson).
Cell preparation
Human fetal bone marrow and thymus were obtained from fetuses at approximately 20 wk gestation at the time of elective termination of pregnancy. Thy/Liv implants were surgically removed from SCID-hu Thy/Liv mice, prepared as previously described (11). Thymi or Thy/Liv implants were ground between the membranes of a nylon mesh bag (Tetko, Kansas City, MO) submerged in PBS; the released cells were rinsed with PBS. Femurs were crushed, and the released cells were subjected to Ficoll-Hypaque density centrifugation. For depletion of cells expressing markers of mature lineage cells, mononuclear cells at the Ficoll interface were rinsed with PBS and incubated with a mixture of PE-conjugated Abs specific for human CD2, CD4, CD8, CD16, CD19, CD20, CD56, and glycophorin A for 30 min at 4°C. Cells coated with these Abs were removed with sheep anti-mouse Ig-conjugated magnetic beads (Dynal, Lake Success, NY). PE-negative cells were then purified from the remaining mixture of cells by FACS using a FACSVantage (Becton Dickinson).
SCID-hu Thy/Liv blood was obtained by cardiac puncture. Adult blood (age, 28–34 yr) was obtained by venipuncture. Cord blood was obtained from term deliveries without evidence of maternal or fetal infection. All procedures and practices were approved by the University of California-San Francisco Committee on Human Research or the University of California-San Francisco Committee on Animal Research.
FACS analysis
Staining of cells with the PE-conjugated CXCR4 or CCR5 mAbs was performed concurrently with mAbs against various CD markers using 106 to 107 purified cells or 50 µl of whole blood in a 100-µl reaction volume containing PBS and 2% FBS. Anti-CD markers included CD38-FITC and CD34-allophycocyanin mAbs for bone marrow mononuclear cells; CD10-FITC and CD34-allophycocyanin for lineage-depleted bone marrow cells; CD3-allophycocyanin, CD4-FITC, and CD8-TC or -ECD for thymocytes from fetal thymus or SCID-hu Thy/Liv implants; CD3-PerCP and CD45-FITC for whole blood from SCID-hu Thy/Liv mice; and CD3-PerCP and CD45RA-FITC for whole blood from adult humans or cord blood. The 47718-PE CXCR4 mAb and its IgG2b-PE isotype control mAb were used at 5 µg/ml, while the 3A9-PE CCR5 mAb and its IgG2a-PE isotype control mAb were used at 1 µg/ml. After 20 min, the cells were rinsed in PBS-2% FBS, fixed in 0.5% paraformaldehyde, and analyzed on a FACSVantage (bone marrow, thymus) or were exposed to FACS lysing solution (Becton Dickinson) for 5 min, fixed in 0.5% paraformaldehyde, and analyzed on a FACScan (blood samples).
Staining of thymocytes for CXCR4 using the unconjugated mAbs 12G5, 44701, and 44712 was performed in a four-step procedure. First, the cells were incubated with the CXCR4 mAb or the IgG2a isotype control mAb at 2 µg/ml for 20 min and rinsed with PBS-2% FBS. The cells were then incubated with the FITC-conjugated goat anti-mouse Ig Ab for 20 min, rinsed with PBS-2% FBS, and blocked with normal mouse serum (Dako) for 20 min. After rinsing with PBS-2% FBS, the thymocytes were incubated with the CD4-PE, CD8-TC, and CD3-allophycocyanin mAbs for 20 min, rinsed with PBS-2% FBS, fixed in 0.5% paraformaldehyde, and analyzed on the FACSVantage.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
To determine the expression levels of CXCR4 during discrete stages
of human thymocyte differentiation (Fig. 1
), thymocytes from SCID-hu Thy/Liv
implants were analyzed by flow cytometry. The implants were derived
from different donors of fetal thymus tissue to assess the variability
in CXCR4 expression among individuals. Thymocytes were isolated from
the implants and incubated with Abs specific for human CD3, CD4, and
CD8 in combination with one of four different mAbs specific for CXCR4:
44701, 44712, 44718-PE (see Materials and Methods), and 12G5
(35). A CD4 vs CD8 plot discriminates four subpopulations of thymocytes
(Fig. 2A):
CD4-CD8-, CD4+CD8-,
CD4-CD8+ (single-positive CD8, or SP8), and
CD4+CD8+ (double-positive, or DP). Cells within
the CD4-CD8- and
CD4+CD8- subpopulations were further
subdivided based on their CD3 expression into
CD3-CD4-CD8- (triple-negative,
or TN), CD3+CD4-CD8-,
CD3-CD4+CD8- (intrathymic T cell
progenitor, or ITTP) (36), and
CD3+CD4+CD8- (single-positive CD4,
or SP4) cells.
|
|
A shows CXCR4 histograms for the TN, ITTP, DP, SP4,
and SP8 thymocyte subpopulations from a representative SCID-hu Thy/Liv
implant. CXCR4 was expressed at high levels, relative to an isotype
control mAb, on the ITTPs and on a subset of the TN cells. CXCR4 levels
on the DP cells varied widely, from high levels on those that are
CD3- to lower levels as CD3 is turned on (see contour
plot, top right of Fig. 2A). SP4s and SP8s
expressed relatively low levels of CXCR4, but SP8 cells expressed
higher levels than did SP4 cells in each implant.
Figure 2B shows the fold shifts in average mean fluorescence
intensity (MFI) for each of the Abs over the isotype mAb on each of the
Thy/Liv thymocyte subpopulations. The average percentages of TN cells
that were positive for CXCR4, as measured by the 12G5, 47701, and
47718-PE mAbs, were 24, 36, and 26% respectively. Each CXCR4 Ab
revealed that the expression of CXCR4 diminishes during thymocyte
maturation. In addition, the 44701 and 44718-PE Abs consistently bound
to the CXCR4+ cells to a greater extent than did the 44712
and 12G5 Abs, with the 44718-PE mAb consistently exhibiting the highest
MFIs.
To determine whether the pattern of expression of CXCR4 observed in the
SCID-hu Thy/Liv implants reflected the pattern of CXCR4 expression in
normal human fetal thymus, the 44701 or 44718-PE mAbs were used in
combination with the CD3, CD4, and CD8 mAbs to stain thymocytes from
three or four fetal thymus specimens, respectively. As seen in SCID-hu
Thy/Liv, CXCR4 levels in the fetal thymus were highest on a TN subset
(roughly 37%) and ITTPs, widely ranging but mostly intermediate on
DPs, lower in SP8s, and at background levels on SP4s (Fig. 2B). In the case of all the CXCR4+
subpopulations, MFI fold-shifts in fetal thymus were 1.5- to 3.6-fold
lower than those on their counterparts in the SCID-hu Thy/Liv implants
(Table I).
|
CCR5 expression on the thymocyte subsets of four SCID-hu Thy/Liv
implants and two fetal thymus specimens was analyzed using a
PE-conjugated form of the anti-human CCR5 mAb 3A9 (33). In contrast
to the pattern of expression of CXCR4, CCR5 was undetectable on TNs and
ITTPs, was expressed at low levels on DPs and SP8s, and was
undetectable on SP4s (Fig. 3A). On DPs and SP8s from
Thy/Liv implants and from fetal thymus, the MFI fold-shifts were
roughly equivalent (Fig. 3B and Table I).
|
To determine whether CXCR4 and/or CCR5 are expressed on
extrathymic subpopulations thought to include T cell progenitors, three
fetal bone marrow specimens (each about 20 wk gestation) were analyzed
by flow cytometry using 44718-PE or 3A9-PE mAb in combination with Abs
specific for human CD34 and CD38 (Fig. 4, A and C).
Bone marrow mononuclear cells were divided into four subpopulations
based on CD34 and CD38 staining, with cells in the
CD34+CD38- gate containing primitive
multilineage hemopoietic progenitors and cells in the
CD34+CD38+ gate containing more mature
lineage-restricted progenitors (37). The 44718-PE CXCR4 mAb stained
each subpopulation at levels higher than those with the isotype control
mAb but lower than those exhibited on fetal thymus TNs and ITTPs (Table I). In contrast, the 3A9-PE CCR5 mAb stained subsets of the
CD34+CD38- (10–13%) and
CD34-CD38- (6–14%) subpopulations in each
of the three marrow specimens but failed to stain the remaining bone
marrow mononuclear cells at levels above that of the isotype control
mAb.
|
, B and C). Expression of CXCR4 and CCR5 in the bloodstream
The 12G5 and 3A9 Abs have been used to show that naive
(CD45RA+) peripheral T cells, which are thought to be
thymic emigrants, express CXCR4 but not CCR5 (32, 33). To determine
whether CXCR4 is up-regulated and CCR5 is down-regulated upon transit
from the thymus to the bloodstream, we assayed the CXCR4 and CCR5
expression levels in peripheral T cells from three different sources:
SCID-hu Thy/Liv blood, in which many (about 74%) of the T cells are
naive (CD45RA+) in phenotype (46); cord blood isolated at
term, where peripheral T cells have had limited exposure to foreign Ags
and are mostly naive in phenotype; and blood from healthy adult humans,
where a mixture of naive and nonnaive (activated or memory) T cells
exists (Fig. 5 and Table I).
|
A, top
panel). The 44718-PE CXCR4 mAb stained
CD45+CD3+ cells in SCID-hu Thy/Liv blood at
relatively high levels. In contrast, the 3A9-PE CCR5 mAb did not stain
peripheral T cells over the isotype Ab.
Four cord blood specimens were stained with the 44718-PE or 3A9-PE mAb
in combination with Abs specific for human CD45RA and CD3 (Fig. 5A, middle panel). As in the case of SCID-hu
Thy/Liv blood, all CD45RA+ cells were CXCR4+.
Few if any CCR5+ T cells were detected, perhaps due to the
low numbers of truly CD45RA- cells.
Six age-matched human adult blood specimens were similarly stained with
44718-PE or 3A9-PE mAb in combination with CD45RA and CD3 mAbs (Fig. 5A, bottom panel). As in the case of cord blood
and SCID-hu Thy/Liv peripheral T cells, adult human peripheral
CD45RA+ T cells were found to bind the 44718-PE CXCR4 mAb
but not the 3A9-PE CCR5 mAb. In addition, the CD45RA- T
cell subset was CXCR4+, although the levels of CXCR4
expressed on these cells were approximately threefold lower than those
on CD45RA+ cells. The 3A9-PE CCR5 mAb stained only a subset
(3–23%) of CD45RA- adult human T cells.
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and Table I).
Qualitatively similar patterns of expression were observed both in
samples derived from human subjects and in samples analyzed in the
context of the SCID-hu Thy/Liv mouse. To the extent that HIV-1
pathogenesis can be studied in this animal model, these results serve
to illuminate the relationship between viral tropism and pathogenic
effects on hemopoietic, including T-lineage,
differentiation.
|
Detection of CXCR4 on CD34+ bone marrow cells is not
surprising, since these cells are attracted by SDF-1 in vitro (47)
and since CXCR4 mRNA was detected by RT-PCR in purified
CD34+ bone marrow cells from all nine persons analyzed
(31). We observed only low levels of CXCR4 on these cells despite the
fact that we used an Ab that exhibits higher MFIs than those of the
previously reported anti-CXCR4 mAb 12G5 (35). The low level of
CXCR4 in CD34+ bone marrow cells, which have been shown to
express low levels of CD4 (48, 49), may explain why they are poorly
infected by HIV-1 in vitro and in vivo (2, 50, 51). However, this level
of expression may be sufficient to trigger indirect effects of HIV-1,
e.g., those mediated by viral envelope proteins produced in adjacent
target cells.
The observation that CXCR4 expression was higher in ITTPs than in CD34+CD10+lin- bone marrow cells implies that CXCR4 expression is up-regulated during or after the T progenitor cell has moved to the thymus. The exact timing of CXCR4 up-regulation may be revealed by further analysis of the TN compartment, which we found to contain both CXCR4+ and CXCR4- cells. If the CXCR4- cells are the immediate predecessors of the CXCR4+ cells (as determined by lineage analysis), CXCR4 up-regulation must be occurring in TNs after, not during, thymus homing.
The data also suggest that CXCR4 is down-regulated during positive and negative selection in the thymus. CXCR4 down-regulation begins only after the thymocyte has become CD4+CD8+; from that point on, CXCR4 is down-regulated in a coordinated fashion with CD3 up-regulation. It is interesting that CXCR4 is down-regulated to a greater degree in SP4s than SP8s. The reason for this disparity is not obvious in light of the observation that both SP4s and SP8s up-regulate CXCR4 upon exit from the thymus.
Our data are in agreement with a previously published flow cytometric analysis of CXCR4 on fetal thymocytes (34). In that study, CXCR4 expression was found to be modulated during thymocyte maturation, with DP thymocytes showing higher levels than most SP4 and SP8 thymocytes. Of note, some SP4 thymocytes were observed to be CXCR4high. Based on our results, these are probably CD3-CD4+CD8- ITTPs, and not more mature SP4 thymocytes.
Our observation that CXCR4 is up-regulated and CCR5 is down-regulated
upon transit of mature thymocytes into the bloodstream is in agreement
with findings concerning CXCR4/CCR5 expression of human naive
peripheral T cells (32). However, our observation that
CD45RA- peripheral T cells express moderately high levels
of CXCR4 is not consistent with that report, in which the memory T cell
subset was found to be CXCR4- with the 12G5 mAb (32).
However, >80% of human T cells, including both CD45RA+
and CD45RO+ cells, undergo chemotaxis in response to
SDF-1 in vitro (24), implying that memory T cells do indeed express
CXCR4. The observed differences in CXCR4 expression levels between the
prior study and this one are probably due to differences in sensitivity
between the 12G5 and 44718-PE mAbs (see Fig. 2A).
The data also indicate that CCR5 is expressed minimally or not at all in the CD34+CD38+ bone marrow subpopulation, which typically accounts for >99% of all CD34+ bone marrow cells, and is expressed at low levels on only 10 to 13% of the cells in the CD34+CD38- subpopulation. These results are in line with those of previous related studies. First, >99% of purified CD34+ bone marrow cells were not significantly attracted to CCR5 ligands in in vitro chemotaxis assays (47). Second, CCR5 mRNA was detected by RT-PCR of purified CD34+ bone marrow cells from only a minor percentage of donors (31). Third, HIV-1 proviral DNA was detected only rarely in CD34+ bone marrow cells from subjects in the asymptomatic stage of HIV-1 disease (52, 53), when virus particles often use only CCR5 (27, 28, 29). The identity of the CD34+CD38-CCR5+ subset is not known but warrants further investigation.
Lastly, we observed that CCR5 is expressed at low but significant levels on SP8 and DP thymocytes, which comprise most of the T progenitor cells in the thymus. This observation implies that CCR5-using, NSI strains of HIV-1 might be able to infect the thymus during the asymptomatic stage of HIV-1 disease, when these strains of HIV-1 predominate (27, 28, 29). Studies are currently in progress to determine the pathogenic outcomes of infection of DP thymocytes with CXCR4- or CCR5-using viruses.
The relevance of these data to HIV-1 pathogenesis in humans must be considered with two caveats in mind. First, data obtained in SCID-hu Thy/Liv mice are correlative and subject to unknown perturbations specific to the animal model itself. Secondly, it is not known whether the levels of coreceptor expression observed in fetal organs of hemopoiesis (e.g., bone marrow and thymus) are similar or pertinent to those found in their adult counterparts. It is notable, however, that SI, CXCR4-using variants of HIV-1 often predominate in late stages of both pediatric and adult HIV-1 disease, whereas NSI, CCR5-using variants are encountered more frequently in early stages of disease (27, 28, 29, 30). In parallel, the former, but not the latter, types of viruses destroy the thymus in the SCID-hu Thy/Liv mouse (20). Furthermore, thymic function has been observed in a surprisingly large fraction of HIV-1-infected adults, but not in those in later stages of disease (CD4 counts of <200 cells/µl) (7). Together, these observations suggest that the switch to CXCR4-using variants in later stages of HIV-1 disease may presage and result in regenerative failure, precisely because such viruses abrogate the function of hemopoietic progenitor cells more efficiently than do CCR5-using viruses. Infection of these progenitor populations could thus be a pivotal event in the disruption of T lymphopoiesis during the course of HIV-1 disease in both children and adults.
Note added in proof.
We have found in subsequent experiments that, unlike the CCR5 mAb 3A9 used in this study, the CCR5-specific mAb 2D7 (Pharmingen, San Diego, CA) does not stain DP or SP8 thymocytes by flow cytometry. However, in a recently published study (54), a novel CCR5-specific mAb (clone 45502.111; R&D Systems) stained DP and SP8, but not DN or SP4, thymocytes by flow cytometry, confirming our results with the CCR5 mAb 3A9.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
2 Address correspondence and reprint requests to Dr. Joseph M. McCune, Gladstone Institute of Virology and Immunology, P.O. Box 419100, San Francisco, CA 94141-9100.
3 M. Hellerstein, M. B. Hanley, D. Cesar, C. Papageorgopoulos, E. Wieder, D. Schmidt, S. Siler, R. Hoh, R. Neese, D. Macallan, S. Deeks, and J. M. McCune. Direct measurement of T lymphocyte kinetics in humans using a stable isotope-mass spectrometric technique: effects of HIV-1 infection and anti-retroviral therapy. Submitted for publication.
4 Abbreviations used in this paper: NSI, nonsyncytium inducing; SI, syncytium inducing; SDF-1, stromal cell-derived factor-1; PE, phycoerythrin; PerCP, peridinin chlorophyll protein; TC, Tri-color; ECD, phycoerythrin-Texas Red; TN, triple negative; SP8, single-positive CD8; DP, double positive; ITTP, intrathymic T progenitor; SP4, single-positive CD4; MFI, mean fluorescence intensity.
Received for publication February 6, 1998. Accepted for publication May 28, 1998.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
This article has been cited by other articles:
|
|
 |
|
  E. Coakley, J. D. Reeves, W. Huang, M. Mangas-Ruiz, I. Maurer, A. M. Harskamp, S. Gupta, Y. Lie, C. J. Petropoulos, H. Schuitemaker, et al. Comparison of Human Immunodeficiency Virus Type 1 Tropism Profiles in Clinical Samples by the Trofile and MT-2 Assays Antimicrob. Agents Chemother., November 1, 2009; 53(11): 4686 - 4693. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Melar, D. E. Ott, and T. J. Hope Physiological Levels of Virion-Associated Human Immunodeficiency Virus Type 1 Envelope Induce Coreceptor-Dependent Calcium Flux J. Virol., February 15, 2007; 81(4): 1773 - 1785. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. B. Gurney and C. H. Uittenbogaart Human Immunodeficiency Virus Persistence and Production in T-Cell Development Clin. Vaccine Immunol., November 1, 2006; 13(11): 1237 - 1245. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Swainson, S. Kinet, N. Manel, J.-L. Battini, M. Sitbon, and N. Taylor Glucose transporter 1 expression identifies a population of cycling CD4+CD8+ human thymocytes with high CXCR4-induced chemotaxis PNAS, September 6, 2005; 102(36): 12867 - 12872. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. K. Choudhary, N. R. Choudhary, K. C. Kimbrell, J. Colasanti, A. Ziogas, D. Kwa, H. Schuitemaker, and D. Camerini R5 Human Immunodeficiency Virus Type 1 Infection of Fetal Thymic Organ Culture Induces Cytokine and CCR5 Expression J. Virol., January 1, 2005; 79(1): 458 - 471. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. C. Yopp, S. Fu, S. M. Honig, G. J. Randolph, Y. Ding, N. R. Krieger, and J. S. Bromberg FTY720-Enhanced T Cell Homing Is Dependent on CCR2, CCR5, CCR7, and CXCR4: Evidence for Distinct Chemokine Compartments J. Immunol., July 15, 2004; 173(2): 855 - 865. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. A. Reyes, D. R. Canfield, U. Esser, L. A. Adamson, C. R. Brown, C. Cheng-Mayer, M. B. Gardner, J. M. Harouse, and P. A. Luciw Induction of Simian AIDS in Infant Rhesus Macaques Infected with CCR5- or CXCR4-Utilizing Simian-Human Immunodeficiency Viruses Is Associated with Distinct Lesions of the Thymus J. Virol., February 15, 2004; 78(4): 2121 - 2130. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. M. Brenchley, B. J. Hill, D. R. Ambrozak, D. A. Price, F. J. Guenaga, J. P. Casazza, J. Kuruppu, J. Yazdani, S. A. Migueles, M. Connors, et al. T-Cell Subsets That Harbor Human Immunodeficiency Virus (HIV) In Vivo: Implications for HIV Pathogenesis J. Virol., February 1, 2004; 78(3): 1160 - 1168. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
N. Schmitt, L. Chene, D. Boutolleau, M.-T. Nugeyre, E. Guillemard, P. Versmisse, C. Jacquemot, F. Barre-Sinoussi, and N. Israel Positive Regulation of CXCR4 Expression and Signaling by Interleukin-7 in CD4+ Mature Thymocytes Correlates with Their Capacity To Favor Human Immunodeficiency X4 Virus Replication J. Virol., May 15, 2003; 77(10): 5784 - 5793. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. B. Gurney, O. O. Yang, S. B. Wilson, and C. H. Uittenbogaart TCR{gamma}{delta}+ and CD161+ Thymocytes Express HIV-1 in the SCID-hu Mouse, Potentially Contributing to Immune Dysfunction in HIV Infection J. Immunol., November 1, 2002; 169(9): 5338 - 5346. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. Pedroza-Martins, W. J. Boscardin, D. J. Anisman-Posner, D. Schols, Y. J. Bryson, and C. H. Uittenbogaart Impact of Cytokines on Replication in the Thymus of Primary Human Immunodeficiency Virus Type 1 Isolates from Infants J. Virol., June 14, 2002; 76(14): 6929 - 6943. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Fukada, Y. Sobao, H. Tomiyama, S. Oka, and M. Takiguchi Functional Expression of the Chemokine Receptor CCR5 on Virus Epitope-Specific Memory and Effector CD8+ T Cells J. Immunol., March 1, 2002; 168(5): 2225 - 2232. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Igarashi, C. R. Brown, R. A. Byrum, Y. Nishimura, Y. Endo, R. J. Plishka, C. Buckler, A. Buckler-White, G. Miller, V. M. Hirsch, et al. Rapid and Irreversible CD4+ T-Cell Depletion Induced by the Highly Pathogenic Simian/Human Immunodeficiency Virus SHIVDH12R Is Systemic and Synchronous J. Virol., January 1, 2002; 76(1): 379 - 391. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. Guillemard, M.-T. Nugeyre, L. Chene, N. Schmitt, C. Jacquemot, F. Barre-Sinoussi, and N. Israel Interleukin-7 and infection itself by human immunodeficiency virus 1 favor virus persistence in mature CD4+CD8{-}CD3+ thymocytes through sustained induction of Bcl-2 Blood, October 1, 2001; 98(7): 2166 - 2174. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. R. Taylor Jr., K. C. Kimbrell, R. Scoggins, M. Delaney, L. Wu, and D. Camerini Expression and Function of Chemokine Receptors on Human Thymocytes: Implications for Infection by Human Immunodeficiency Virus Type 1 J. Virol., September 15, 2001; 75(18): 8752 - 8760. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
H. Xiao, C. Neuveut, H. L. Tiffany, M. Benkirane, E. A. Rich, P. M. Murphy, and K.-T. Jeang Selective CXCR4 antagonism by Tat: Implications for in vivo expansion of coreceptor use by HIV-1 PNAS, October 10, 2000; 97(21): 11466 - 11471. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Gobbi, C. A. Stoddart, G. Locatelli, F. Santoro, C. Bare, V. Linquist-Stepps, M. E. Moreno, N. W. Abbey, B. G. Herndier, M. S. Malnati, et al. Coinfection of SCID-hu Thy/Liv Mice with Human Herpesvirus 6 and Human Immunodeficiency Virus Type 1 J. Virol., September 15, 2000; 74(18): 8726 - 8731. [Abstract] [Full Text]
|
|
|
|
|
|
S. Lee, H. L. Tiffany, L. King, P. M. Murphy, H. Golding, and M. B. Zaitseva CCR8 on Human Thymocytes Functions as a Human Immunodeficiency Virus Type 1 Coreceptor J. Virol., August 1, 2000; 74(15): 6946 - 6952. [Abstract] [Full Text]
|
|
|
|
|
|
S. Lee, C. K. Lapham, H. Chen, L. King, J. Manischewitz, T. Romantseva, H. Mostowski, T. S. Stantchev, C. C. Broder, and H. Golding Coreceptor Competition for Association with CD4 May Change the Susceptibility of Human Cells to Infection with T-Tropic and Macrophagetropic Isolates of Human Immunodeficiency Virus Type 1 J. Virol., June 1, 2000; 74(11): 5016 - 5023. [Abstract] [Full Text]
|
|
|
|
|
|
H. Nagase, M. Miyamasu, M. Yamaguchi, T. Fujisawa, K. Ohta, K. Yamamoto, Y. Morita, and K. Hirai Expression of CXCR4 in Eosinophils: Functional Analyses and Cytokine-Mediated Regulation J. Immunol., June 1, 2000; 164(11): 5935 - 5943. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. Camerini, H.-P. Su, G. Gamez-Torre, M. L. Johnson, J. A. Zack, and I. S. Y. Chen Human Immunodeficiency Virus Type 1 Pathogenesis in SCID-hu Mice Correlates with Syncytium-Inducing Phenotype and Viral Replication J. Virol., April 1, 2000; 74(7): 3196 - 3204. [Abstract] [Full Text]
|
|
|
|
|
|
R. M. Scoggins, J. R. Taylor Jr., J. Patrie, A. B. van't Wout, H. Schuitemaker, and D. Camerini Pathogenesis of Primary R5 Human Immunodeficiency Virus Type 1 Clones in SCID-hu Mice J. Virol., April 1, 2000; 74(7): 3205 - 3216. [Abstract] [Full Text]
|
|
|
|
|
|
S. G. Kitchen, S. Killian, J. V. Giorgi, and J. A. Zack Functional Reconstitution of Thymopoiesis after Human Immunodeficiency Virus Infection J. Virol., March 15, 2000; 74(6): 2943 - 2948. [Abstract] [Full Text]
|
|
|
|
|
|
H. Blaak, A. B. van't Wout, M. Brouwer, B. Hooibrink, E. Hovenkamp, and H. Schuitemaker In vivo HIV-1 infection of CD45RA+CD4+ T cells is established primarily by syncytium-inducing variants and correlates with the rate of CD4+ T cell decline PNAS, February 1, 2000; 97(3): 1269 - 1274. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Shankarappa, J. B. Margolick, S. J. Gange, A. G. Rodrigo, D. Upchurch, H. Farzadegan, P. Gupta, C. R. Rinaldo, G. H. Learn, X. He, et al. Consistent Viral Evolutionary Changes Associated with the Progression of Human Immunodeficiency Virus Type 1 Infection J. Virol., December 1, 1999; 73(12): 10489 - 10502. [Abstract] [Full Text]
|
|
|
|
|
|
L. Chêne, M.-T. Nugeyre, E. Guillemard, N. Moulian, F. Barré-Sinoussi, and N. Israël Thymocyte-Thymic Epithelial Cell Interaction Leads to High-Level Replication of Human Immunodeficiency Virus Exclusively in Mature CD4+ CD8- CD3+ Thymocytes: a Critical Role for Tumor Necrosis Factor and Interleukin-7 J. Virol., September 1, 1999; 73(9): 7533 - 7542. [Abstract] [Full Text]
|
|
|
|
|
|
R. D. Berkowitz, A. B. van 't Wout, N. A. Kootstra, M. E. Moreno, V. D. Linquist-Stepps, C. Bare, C. A. Stoddart, H. Schuitemaker, and J. M. McCune R5 Strains of Human Immunodeficiency Virus Type 1 from Rapid Progressors Lacking X4 Strains Do Not Possess X4-Type Pathogenicity in Human Thymus J. Virol., September 1, 1999; 73(9): 7817 - 7822. [Abstract] [Full Text]
|
|
|
|
|
|
C. Chelucci, I. Casella, M. Federico, U. Testa, G. Macioce, E. Pelosi, R. Guerriero, G. Mariani, A. Giampaolo, H.J. Hassan, et al. Lineage-Specific Expression of Human Immunodeficiency Virus (HIV) Receptor/Coreceptors in Differentiating Hematopoietic Precursors: Correlation With Susceptibility to T- and M-Tropic HIV and Chemokine-Mediated HIV Resistance Blood, September 1, 1999; 94(5): 1590 - 1600. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. M. Harouse, A. Gettie, R. C. H. Tan, J. Blanchard, and C. Cheng-Mayer Distinct Pathogenic Sequela in Rhesus Macaques Infected with CCR5 or CXCR4 Utilizing SHIVs Science, April 30, 1999; 284(5415): 816 - 819. [Abstract] [Full Text]
|
|
|
|
|
|
R. D. Berkowitz, S. Alexander, C. Bare, V. Linquist-Stepps, M. Bogan, M. E. Moreno, L. Gibson, E. D. Wieder, J. Kosek, C. A. Stoddart, et al. CCR5- and CXCR4-Utilizing Strains of Human Immunodeficiency Virus Type 1 Exhibit Differential Tropism and Pathogenesis In Vivo J. Virol., December 1, 1998; 72(12): 10108 - 10117. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
