![[Feedback]](/icons/banner/feedbackACT.gif)
![[For Subscribers]](/icons/banner/subscriberACT.gif)
![[Archive]](/icons/banner/archiveACT.gif)
![[Search]](/icons/banner/searchACT.gif)
![[Contents]](/icons/banner/tocACT.gif)
Previous Article | Next Article 
Journal of Virology, November 1998, p. 8613-8619, Vol. 72, No. 11
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Dissemination of Lymphocytic Choriomeningitis Virus
from the Gastric Mucosa Requires G Protein-Coupled Signaling
Cheng
Yin,
Mahmoud
Djavani,
Alan R.
Schenkel,
Daniel
S.
Schmidt,
C. David
Pauza, and
Maria S.
Salvato*
Department of Pathology and Laboratory
Medicine, University of Wisconsin Medical School, Madison,
Wisconsin 53706
Received 23 April 1998/Accepted 28 July 1998
 |
ABSTRACT |
The gastric mucosa is an important portal of entry for lymphocytic
choriomeningitis virus (LCMV) infections. Within hours after
intragastric (i.g.) inoculation, virus appears in the gastric epithelia, then in the mesenteric lymph nodes and spleen, and then
in the liver and brain. By 72 h i.g.-inoculated virus is widely
disseminated and equivalent to intravenous (i.v.) infection (S. K. Rai, B. K. Micales, M. S. Wu, D. S. Cheung, T. D. Pugh, G. E. Lyons, and M. S. Salvato. Am. J. Pathol.
151:633-639, 1997). Pretreatment of mice with a G protein inhibitor,
pertussis toxin (PTx), delays LCMV dissemination after i.g., but not
after i.v., inoculation. Delayed infection was confirmed by plaque
assays, by reverse transcription-PCR, and by in situ hybridization. The differential PTx effect on i.v. and i.g. infections indicates that
dissemination from the gastric mucosa requires signals transduced through heterotrimeric G protein complexes. PTx has no direct effect on
LCMV replication, but it modulates integrin expression in part by
blocking chemokine signals. LCMV infection of macrophages up-regulates
CD11a, and PTx treatment counteracts this. PTx may prevent early LCMV
dissemination by inhibiting the G protein-coupled chemotactic response
of macrophages infected during the initial exposure, thus blocking
systemic virus spread.
| |
INTRODUCTION |
Lymphocytic choriomeningitis virus
(LCMV) is the prototype of the Arenaviridae, a family of
ambisense, bisegmented RNA viruses. Although laboratory studies of this
virus employ primarily parenteral routes of inoculation, virus
ingestion via the gastric route is the probable natural route of
infection (28). Our previous studies established that virus
delivered intravenously (i.v.) appears in several different organs
simultaneously, whereas virus delivered intragastrically (i.g.) appears
first in the stomach by 12 h, then in the spleen by 24 h,
then in the liver by 48 h, and finally, by 72 h, it is as
widely disseminated as i.v.-inoculated virus (29). Based on
these results we proposed that i.g. infection, unlike i.v. infection,
is likely to disseminate in a cell-associated manner through the
lymphatics. We used pertussis toxin (PTx), to disrupt mononuclear cell
(monocytes and lymphocytes) movement in order to test whether a block
to cell trafficking can delay LCMV dissemination. Our studies followed
the observation that PTx inhibits reovirus i.g. dissemination
(37) and extended this observation by linking the action of
PTx to virus-mediated changes in adhesion molecules and describing a
model for the events involved in virus dissemination.
PTx, a surface protein of Bordetella pertussis, catalyzes
ADP ribosylation of heterotrimeric GTP-binding proteins and disrupts signal transduction (24). Cytokines and chemokines, e.g.,
interleukin-8, monocyte chemotactic protein (MCP)-1, lymphotactin, and
fractalkine, are potent mediators of mononuclear cell migration
(14, 20, 33, 38), and they signal through G
protein-coupled seven-transmembrane receptors (9, 12).
PTx blocks G-coupled signaling and thereby blocks the rapid changes in
integrins posttranscriptionally activated by such signaling
(4). PTx-mediated leukocytosis can also be attributed to
inhibition of G protein-dependent cell migration (16, 22, 23,
36). We propose that LCMV is spread by migrating mononuclear
cells after mucosal inoculation and that PTx delays viral dissemination
by blocking cell migration. We show that LCMV infection up-regulates
cell surface adhesion molecule CD11a, but PTx down-regulates its
surface expression. We speculate that PTx is interfering with a
mechanism the virus has evolved to influence its tropism.
| |
MATERIALS AND METHODS |
Virus inoculation and PTx treatment of mice.
Six-week-old
male BALB/c mice were obtained from Harlan Sprague Dawley
(Indianapolis, Ind.). LCMV (Armstrong 53b strain) was plaque purified
and stored at 107 to 108 PFU/ml. Virus inocula
of 106 PFU in 0.1 ml of RPMI 1640 medium were administered
to mice either i.v. (tail vein) or by gastric intubation as described
previously (28).
Virus was titered by plaque assay on Vero cells (ATCC CCL-81) as
described previously (8). Briefly, dilutions of homogenized splenocytes were incubated at 37°C, 5% CO2 for 1 h
with Vero cell monolayers grown in 6-well plates (Costar, Cambridge,
Mass.). The plates were then overlaid with 1% agarose in minimal
essential medium 199 containing 10% fetal calf serum (FCS) and
incubated at 37°C, 5% CO2 for 5 days. The wells were
treated with 25% formaldehyde and stained with 0.1% crystal violet
for 30 min. The agarose overlay was removed, and plaques were counted.
Each dilution was done in duplicate, and the counts were averaged.
Infectious centers were similarly determined by plating murine
leukocytes at 105 to 106 cells per
50-mm-diameter plate containing Vero cells at 70% confluence.
Some of the mice received an i.v. injection of 25 ng of PTx (List
Biologicals, Campbell, Calif.) per g of body weight in 0.1 ml of
phosphate-buffered saline. Mock treatments utilized 0.1 ml of
phosphate-buffered saline. Cultured spleen mononuclear cells were
treated with 0.1 µg of PTx/ml, which is close to the estimated PTx
concentration in vivo and well below its mitogenic concentration (>1.0
µg/ml) (18).
Detection of viral nucleic acid.
Three, 7, or 10 days after
infection (or mock infection), the mice were sacrificed by cervical
dislocation. Blood was collected from the hearts and kept in
heparinized tubes on ice. Spleens, kidneys, and livers were removed for
use in virus titration, RNA extraction, flow cytometry, and/or cell
culture. The spleens were placed in RPMI 1640 containing 10% FCS and
kept on ice. The kidneys and livers were immediately processed for
extraction of viral RNA.
Kidneys and livers were homogenized and sonicated in guanidine
isothiocyanate solution and then layered on a CsCl cushion for density
gradient isolation of total RNA as described previously (5).
RNA was extracted from small amounts of tissue or cell cultures by
using TRIzol reagent (Life Technologies, Gaithersburg, Md.). Reverse
transcription followed by PCR (RT-PCR) was performed with 4 µg of
whole-cell RNA and 1 µg of random hexanucleotide primers (Promega,
Madison, Wis.) denatured at 65°C for 2 min, annealed on ice, and then
added to a solution containing 1 µl of 40-U/µl RNasin (Promega), 1 µl of deoxyribonucleotides at 1.25 mM each, 4 µl of 5× RT buffer
(50 mM Tris-HCl [pH 8.3], 50 mM KCl, 10 mM MgCl2, 10 mM
dithiothreitol, 0.5 mM spermidine), and 1 µl of 23-U/µl avian
myeloblastosis virus RT (Promega) in a total volume of 20 µl for
1 h at 42°C. RT-PCR template concentrations were used in the
range in which they are linearly related to the amount of product.
Amplification of the cDNA was performed as described previously
(32) with two oligonucleotide primers that anneal to the
LCMV glycoprotein (Gp) gene (5'-TCATCGATGAGGTGATCAAC-3' and
5'-CTTGGTGAACTCTCTAGACT-3'). Another set of oligonucleotide primers that anneal to the dihydrofolate reductase (DHFR) gene (5'-CTCAGGGCTGCGATTTCGCGCCAAACT-3' and
5'-TATCAGCCTCCGTCAAGACAAATGGTC-3') served as an internal
control (17). The oligonucleotide primers were made on an
automated synthesizer (Gene Assembler Plus; Pharmacia LKB, Piscataway,
N.J.). RT-PCR products were analyzed by 1% agarose gel
electrophoresis.
In situ hybridization to detect virus replication.
At 24 and
48 h after infection, the mice were sacrificed and the livers,
stomachs, spleens, kidneys, ilea, and mesenteric lymph nodes were
collected. The tissues were fixed in 10% neutral phosphate-buffered
formalin and embedded in paraffin, and 5-µm-thick sections were
prepared.
In situ hybridization was performed as described previously
(15) with digoxigenin-labeled RNA probes generated by SP6 or T7 polymerase transcription from the entire LCMV Gp gene. Briefly, tissue sections were hybridized with 1.5 ng of the riboprobes/ml at
52°C overnight, washed in 2× SSC-50% formamide solution (1× SSC
is 0.15 M NaCl plus 0.015 M sodium citrate) and then in 2× SSC, and
treated with RNase T1 and RNase A for 30 min at 37°C. The slides were
blocked for 1 h with a buffer containing 2% horse serum, 150 mM
NaCl, and 100 mM Tris, pH 7.4. After being blocked, the slides were
incubated for 1 h with sheep anti-digoxigenin-alkaline phosphatase
conjugate (Boehringer Mannheim, Indianapolis, Ind.) at a 1:500
dilution, rinsed in Tris, pH 7.4, and incubated overnight at room
temperature with nitroblue tetrazolium-BCIP (5-bromo-4-chloro-3 indolylphosphate) (Vector, Burlingame, Calif.) substrate in the dark.
The stained slides were rinsed in water, counterstained with nuclear
fast red, dehydrated, and sealed under a coverslip. LCMV Gp sense and
antisense probes were used, and uninfected tissues were included as
controls.
Flow cytometry to detect cell surface adhesion molecules and
lymphocyte subset distribution.
Peripheral blood mononuclear cells
(PBMC) were obtained from heparinized blood by Ficoll-Hypaque density
gradient centrifugation. Spleen mononuclear cells were obtained by
passage through an 80-mesh screen followed by Ficoll-Hypaque density
gradient centrifugation. The cells were fixed in 1% paraformaldehyde
and stained with fluorescein isothiocyanate-conjugated antibodies
against murine CD4, CD8, CD22, 
T-cell receptor, CD11a, CD18,
CD44, CD49d, or CD54 or double stained with phycoerythrin-conjugated
antibody against Mac-1 and fluorescein isothiocyanate-conjugated
antibodies against CD11a, CD44, or CD54 (Caltag, South San Francisco,
Calif.). Labeled isotype controls were included. Flow cytometry was
performed on a Becton-Dickinson FACScan, and data was processed with
Becton-Dickinson Lysis II and SAS JMP statistical software.
| |
RESULTS |
LCMV replication in cultured splenocytes is not inhibited by
PTx.
Spleen mononuclear cells (splenocytes) are primarily
lymphocytes with approximately 4 to 5% macrophages (adherent
Mac-1+ cells). The lymphocytes are infectable by LCMV
(Armstrong 53b strain) at only 1/100 the efficiency of macrophages
(31a); hence, macrophages are the primary infected cell type
in this preparation. Splenocytes from four BALB/c mice were divided
into four groups: uninfected; infected; infected, with PTx
pretreatment; and infected, with PTx posttreatment. The cells were
cultured in RPMI 1640 containing 10% FCS, infected (or not) with LCMV
Armstrong at a multiplicity of 1 PFU (as determined on Vero
cells)/splenocyte, and treated (or not) with PTx (0.1 µg/ml) either 1 day before or 1 day after infection. The cells were cultured for 3 days, RNA was isolated, and RT-PCR was performed as described in
Materials and Methods to detect LCMV Gp mRNA. Neither pretreatment nor
posttreatment with PTx blocked the LCMV infection in cell culture (Fig.
1).
View larger version (34K):
[in this window]
[in a new window]
|
FIG. 1.
PTx does not affect LCMV mRNA production in splenocyte
cell culture. LCMV Gp mRNA was detected by RT-PCR, starting with total
RNA from splenocyte cultures. Agarose gel electrophoresis indicates
that the expected 1-kb LCMV Gp product is not detected in uninfected
splenocytes (lane 1) but is detected in samples from in vitro
LCMV-infected splenocytes (lane 2), despite pretreatment (lane 3) or
posttreatment (lane 4) with PTx (0.1 µg/ml). Lane N is a negative
control for PCR. The 447-bp DHFR gene product was used as an internal
control. Lane M is a 1-kb DNA ladder supplied by Gibco BRL (the upper
gel depicts only the 1-kb marker fragment, whereas the lower gel
depicts 0.5-kb and smaller marker fragments).
|
|
Portions of the splenocyte cultures were used for infectious-center
assays, with the result that both PTx-treated and untreated cultures
had 100 ± 10 infectious centers per 106 cells.
LCMV dissemination after i.g. inoculation but not after i.v.
inoculation can be delayed by PTx treatment.
Data on the tissue
levels of virus were collected by RT-PCR, by plaque assay, and by in
situ hybridization. For the RT-PCR experiments, there were four sets of
mice, three time points per set, and six mice per time point. The sets
were as follows: set 1, i.v.-infected mice; set 2, PTx-pretreated and
i.v.-infected mice; set 3, i.g.-infected mice; and set 4, PTx-pretreated and i.g.-infected mice. The PTx-pretreated mice received
PTx 3 days before LCMV inoculation and were sacrificed at 3, 7, or 10 days after infection. The mice inoculated without PTx treatment served as positive controls. Three days after inoculation, LCMV RNA was detected in the livers and the kidneys of all six mice from both the
i.v.- and i.g.-infected groups (Fig. 2,
lanes 1 and 2, and Table 1).
View larger version (29K):
[in this window]
[in a new window]
|
FIG. 2.
In vivo effects of PTx on virus. LCMV Gp mRNA was
detected by RT-PCR of total RNA isolated from kidneys and livers of
i.g.- or i.v.-infected mice. Gel electrophoresis of the RT-PCR product
of total RNA isolated from the kidneys of i.g.- and i.v.-infected mice
3 days after infection is shown. The expected 1-kb LCMV Gp product is
detected in samples from an i.g.-infected mouse (lane 1), an
i.v.-infected mouse (lane 2), and a PTx-pretreated, i.v.-infected mouse
(lane 4) but not in that from a PTx-pretreated, i.g.-infected mouse
(lane 3). Lane 5 is from the kidney of an uninfected mouse. Lane P is a
1-kb LCMV Gp fragment. Lane N is a negative control for PCR. The 447-bp
DHFR gene product was used as an internal control. Lane M is a 1-kb DNA
ladder supplied by Gibco BRL (the upper gel depicts only the 1-kb
marker fragment, whereas the lower gel depicts 0.5-kb and smaller
marker fragments).
|
|
PTx inhibited the dissemination of LCMV after i.g. inoculation in five
of six mice (Fig. 2, lane 3, and Table 1), but it did not inhibit
dissemination after i.v. infection (Fig. 2, lane 4, and Table 1). By
day 7, only three of six mice inoculated i.g. had detectable LCMV RNA
in the liver or kidneys, while all six of the mice inoculated i.v. had
LCMV RNA in those organs (Table 1).
We also determined the effect of PTx treatment on LCMV infection by
performing plaque assays in parallel with RT-PCR. Spleens were removed
3 days after infection and analyzed by plaque assay by using Vero cell
monolayers grown in 6-well plates. Plaques were counted after 5 days
and expressed as the number of plaques per gram of tissue (Table
2). Five of five i.g.-inoculated,
PTx-treated animals were RT-PCR negative for viral RNA and contained a
mean of 5.4 × 103 PFU/g of spleen, whereas untreated
animals were uniformly positive for viral RNA and had greater than
105 PFU/g (with a mean of 1.9 × 107
PFU/g). The results of the RT-PCR supported those of the large experiment reported above, as well as the results of the plaque assays
in the i.g.-infected, PTx-treated group (Table 2).
View this table:
[in this window]
[in a new window]
|
TABLE 2.
Detection of LCMV infection by plaque assay and
RT-PCR in spleens from i.g.- or i.v.-infected mice with or without
PTx treatment
|
|
LCMV was detected in various organs at different time points by in situ
hybridization (Table 3). Three days after
intravenous injection of 25 ng of PTx/g, each mouse was inoculated i.v.
or i.g. with 106 PFU of LCMV. At 24 h after gastric
inoculation, viral RNA was detected in the stomach in all mice, whether
they were pretreated with PTx or not. Stomach infection appeared within
epithelial cells of the gastric mucosa (Fig.
3a). By 24 h virus was detectable in
the spleen and liver of one of the mice in the i.g.-infected, PTx-untreated group but not in the other three i.g.-infected mice. By
48 h after gastric infection, viral RNA was detected in the spleen
of one mouse and the livers of both mice from the PTx-untreated group,
but not in the PTx-treated mice (Fig. 3b and c). In contrast to the
i.g.-infected mice, virus appeared in i.v.-infected mice as early as
24 h in the stomach, liver, spleen, ileum, kidneys, and mesenteric
lymph nodes, and PTx pretreatment did not affect the infection (Fig. 3
and Table 3).
View this table:
[in this window]
[in a new window]
|
TABLE 3.
Summary of in situ hybridization for detection of viral
RNA with LCMV Gp cRNA probe in various mouse organs at different
time points after inoculationa
|
|
View larger version (90K):
[in this window]
[in a new window]
|
FIG. 3.
In situ hybridization of various mouse tissue
sections with an LCMV Gp probe. (a) Stomach sections at 24 h
showing infection of the gastric epithelium in the body of the
stomach. Magnification, ×100. (b and c) Spleen and liver sections,
respectively, at 48 h after infection. Magnification, ×100. IG,
i.g. inoculation; IV, i.v. inoculation.
|
|
PTx causes leukocytosis.
One week after PTx treatment, the
peripheral leukocyte counts of both i.v.- and i.g.-infected mice
increased significantly in comparison with those of their
PTx-untreated counterparts, with 7.65- and 10.18-fold
increases, respectively. However, 10 days after PTx treatment, the
peripheral leukocyte counts decreased, with only 1.99- and
2.46-times-higher concentrations of circulating leukocytes in the
infected, PTx-treated animals, respectively (Fig.
4).
View larger version (35K):
[in this window]
[in a new window]
|
FIG. 4.
Absolute leukocyte counts in peripheral blood from
LCMV-infected mice with or without PTx pretreatment. Leukocytes were
counted under a 20×-objective light microscope in the presence of
trypan blue to eliminate dead cell counts. Data are represented as
leukocytes/milliliter of whole blood at days 7 and 10 after toxin
administration. IV, i.v. inoculation; IG, i.g. inoculation.
|
|
LCMV infection affects adhesion molecule levels in vivo and in cell
culture, and PTx treatment also modulates adhesion molecule
expression.
Heparinized blood was obtained from 24 mice, half of
which were PTx treated and the other half untreated. By 3 to 7 days
after PTx treatment, lower levels of five adhesion molecules (CD11a, CD18, CD44, CD49d, and CD54) were detected on PBMC. However, the levels
of these adhesion molecules returned to near normal 10 days after PTx
treatment (Fig. 5).
View larger version (47K):
[in this window]
[in a new window]
|
FIG. 5.
Phenotypic analysis of mouse PBMC at different time
points (days 0, 3, 7, and 10) after PTx treatment in vivo. All five of
the adhesion molecules monitored show decreased expression on PBMC 3 to
7 days after PTx treatment and return to near normal by 10 days.
|
|
We also observed the effects of PTx on adhesion molecules in cell
culture with or without LCMV infection. Spleen mononuclear cells were
isolated from 24 BALB/c mice and divided into four groups, with
the cells of six mice in each group. Splenocytes were cultured
overnight, infected (or not) with LCMV at a multiplicity of 1 PFU/cell
overnight, and then treated (or not) with PTx (0.1 µg/ml) for 3 days. The amount of PTx used in vitro corresponded to the estimated
concentration of PTx in vivo and was below its mitogenic concentration
(>1 µg/ml) (18). Nonadherent cultured splenocytes were
collected, fixed, stained with fluor-conjugated antibodies, and
analyzed by flow cytometry. CD11a (LFA-1 
chain) was up-regulated on
murine splenocytes after LCMV infection (P < 0.05)
(Table 4). PTx treatment significantly
down-regulated the expression of CD11a, CD44, and CD54 (ICAM-1) on
mouse splenocytes (P < 0.001). Changes were not as
significant for CD18 (LFA-1 
chain), CD49d (VLA-4), and lymphocyte
subsets CD4 (helper T cells), CD8 (cytotoxic T cells), CD22 (B cells),
or T cells (P > 0.1) (Table 4).
View this table:
[in this window]
[in a new window]
|
TABLE 4.
Changes in adhesion molecule expression on
LCMV-infected and -uninfected murine splenocytes after
PTx treatmenta
|
|
Further study by two-color flow cytometry of splenocyte
cultures (which were a mixture of adherent and nonadherent cells) showed that LCMV infection up-regulated CD11a expression on
Mac-1+ monocyte/macrophages (P < 0.01)
(Table 5), and PTx treatment blocked this
up-regulation (P < 0.05) (Table 5). Two-color flow cytometry to detect expression of LCMV Gp and CD11a indicated that the
up-regulation of CD11a was occurring mostly in infected cells.
Approximately 5% of the splenocytes were infected, and 90% of the
infected cells were doubly fluorescent for CD11a and Gp (data not
shown). PTx also down-regulated the expression of CD44 and CD54 on
Mac-1+ monocyte/macrophages (P < 0.001)
(Table 5).
View this table:
[in this window]
[in a new window]
|
TABLE 5.
Changes in adhesion molecule expression on cultured
Mac-1+ adherent splenocytes after LCMV infection and
PTx treatmenta
|
|
| |
DISCUSSION |
Previous studies have shown that the mucosa is the initial site of
LCMV infection after i.g. inoculation. Subsequently, the virus spreads
to the spleen and liver, then the ileum, and finally, the lungs,
kidneys, brain, and esophagus (29). Our results corroborate this order of events and show that treatment with the G protein inhibitor PTx blocks the initial step and delays dissemination from the
gastric mucosa to the spleen. Twenty-four hours after gastric
inoculation, viral RNA was first detected in the stomachs of all the
mice studied. By 48 h, viral RNA was detected in the spleen and
liver, as well as the stomach, in PTx-untreated mice but not in the
i.g.-inoculated, PTx-treated mice. In contrast, virus appeared at all
sites (stomach, liver, kidneys, and brain) as early as 24 h after
i.v. inoculation, irrespective of PTx treatment.
Chemokines, such as macrophage-derived chemokine (MDC) and
MCP-3, promote the chemotaxis of lymphocytes, monocytes/macrophages, polymorphonuclear leukocytes, NK cells, dendritic cells, and
endothelial cells both in vivo and in vitro (9, 13, 31)
through a PTx-sensitive G protein signaling pathway (7). In
murine cytomegalovirus infections, the loss of a virus-encoded G
protein-coupled chemokine receptor abrogates replication
in the salivary gland (6), suggesting that this receptor is
important for trafficking of infected leukocytes to the target organs.
In our studies, PTx was employed at a concentration sufficient to
inhibit G protein-coupled signaling but at 2 orders of magnitude below
the concentration needed to cause mitogenesis or toxicity in culture
(26). Arenaviruses primarily infect reticuloendothelial cells (monocyte/macrophages and dendritic cells) (41),
and whereas PTx does not affect their ability to replicate virus (Fig.
1), it directly affects the expression of integrins on their
surfaces (Table 5). We showed that PTx does not affect production of
interferon-, a major inhibitor of viral infection (data not shown),
and that it does not cause any significant changes in splenic
lymphocyte populations (Table 4). We have also shown that PTx acts
before LCMV immune responses come into play (28). Thus, the
most likely mechanism for the PTx inhibition of virus dissemination is
the inhibition of G protein-coupled chemotactic signals in infected reticuloendothelial cells.
To measure the effectiveness of PTx treatment, we monitored
leukocytosis. By 1 week after PTx injection, the peripheral leukocyte count increased significantly in PTx-treated mice and, at the same
time, LCMV RNA was detectable in the liver and kidneys in only one of
six i.g.-inoculated mice. However, 10 days after PTx injection, the
peripheral leukocyte count returned to near normal, and by this time,
LCMV RNA was detectable in three of six mice. Thus, we demonstrated a
correlation between PTx disruption of normal leukocyte trafficking and
PTx disruption of LCMV dissemination after mucosal inoculation.
Inhibition of G protein-coupled signaling blocks the ability of
chemokines to activate integrins (4) and leads to a decrease in integrin expression. Accordingly, PTx treatment of mice decreases the levels of five adhesion molecules within 3 to 7 days. The levels
return to normal by 10 days after treatment. The decrease in adhesion
molecules parallels the peak of leukocytosis, and circulating leukocyte
counts return to normal levels as the levels of adhesion molecules
return. Leukocytosis coincides with an observable depletion of cells in
the spleen and lymph nodes (as shown by histopathology done by C. Yin).
Adhesion molecules on leukocytes mediate early and reversible
interactions (rolling and margination) along the lumenal surfaces of
vascular endothelial cells. Certain members of the immunoglobulin
superfamily (VCAM-1 and ICAM-1) regulate later and irreversible steps
which lead to firm attachment and subsequent diapedesis
(10). Our current understanding of the role of adhesion
molecules in leukocyte trafficking allows us to surmise that the
down-regulation of adhesion molecules is connected to the depletion of
lymphoid tissues.
In order to investigate further the PTx effects on cell surface
adhesion molecules, we monitored the effects of PTx in cultured spleen
mononuclear cells with or without LCMV infection. Similar to what we
found in vivo, PTx significantly down-regulates the expression of
CD11a, CD44, and CD54 and has minor effects on CD18 and CD49d. We
discovered that CD11a is up-regulated on murine splenocytes after LCMV
infection, especially on murine splenic monocytes/macrophages, which
are the major cells infected by LCMV. Cultured splenocytes show a small
(4%) but significant increase in CD11a-positive cells. This can be
reconciled with the large effect on trafficking in vivo by the fact
that only 5% of the splenocytes are infected and that CD11a
up-regulation occurs preferentially in infected cells.
It is important to note that others have shown that LCMV infection
elicits CD11a expression on CD8+ lymphocytes (1, 2,
19). Since CD8+ lymphocytes are poorly infectable,
they account for most of the uninfected CD11a+ cells. The
marked and long-standing LFA-1 up-regulation on CD8+
lymphocytes may explain the rapid infiltration of lymphocytes into
infected sites and the accompanying immunopathogenesis that is a
hallmark of acute LCMV disease. Our new observation is that LFA-1 chain (CD11a) expression increases on the surface of
monocytes/macrophages, the primary cells infected with LCMV,
far more rapidly and extensively than on T cells. We surmise that this
increase has some effect on the trafficking of infected cells and
ultimately on the dissemination of infection within the host.
Virus-induced changes in adhesion molecule expression have
also been observed by many other laboratories. For example,
intercellular adhesion molecule-1 (ICAM-1) was induced on human
hepatocytes after transfection by hepatitis B virus DNA
(40). ICAM-1 expression was also up-regulated on BALB/c
mouse brain microvascular endothelial cells by measles virus and herpes
simplex type 1 virus and was responsible for increased lymphocyte
homing to the central nervous system (3). The expression of
LFA-1 and ICAM-1 is also stimulated by human immunodeficiency virus
type 1 (HIV-1) infection and suggests a mechanism for extravascular
dissemination of HIV-1-infected cells (30). Taken together,
these findings suggest that up-regulation of LFA-1 expression may be a
critical factor in virus tropism, directing infected cells towards
vascular adhesion, and that PTx counteracts this direction by
down-regulating LFA-1 and causing more leukocytes to remain in the
circulation.
In summary, we used PTx to interfere with the normal leukocyte
trafficking that is required for LCMV dissemination. These results
encourage further efforts to modify cell trafficking as an approach to
manipulating mucosal viral infections, such as LCMV infection. Other
viral infections that occur primarily via the mucosal route, e.g.,
simian immunodeficiency virus infection in rhesus macaques
(27), HIV infection in humans (11, 21, 34),
reovirus infection (39), and infections by picornaviruses (25), may all be affected by chemotaxis-inhibiting agents.
| |
ACKNOWLEDGMENTS |
This research was supported primarily by NIH grant AI 38491 (C.D.P.). Cheng Yin's stipend was from NIH grant RR00167 to the Wisconsin Regional Primate Research Center (for projects by C.D.P. and
M.S.).
We thank David A. Hildeman and Daniel Muller from the Department of
Microbiology, University of Wisconsin
Madison, for assistance in
performing plaque assays.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Pathology and Laboratory Medicine, University of Wisconsin Medical
School, 1300 University Ave., Madison, WI 53706. Phone: (608) 262-6058. Fax: (608) 262-9148. E-mail: msalvato{at}facstaff.wisc.edu.
| |
REFERENCES |
| 1.
|
Andersson, E. C.,
J. P. Christensen,
O. Marker, and A. R. Thomsen.
1994.
Changes in cell adhesion molecule expression on T cells associated with systemic virus infection.
J. Immunol.
152:1237-1245[Abstract].
|
| 2.
|
Andersson, E. C.,
J. P. Christensen,
A. Scheynius,
O. Marker, and A. R. Thomsen.
1995.
Lymphocytic choriomeningitis virus infection is associated with long-standing perturbation of LFA-1 expression on CD8+ T cells.
Scand. J. Immunol.
42:110-118[Medline].
|
| 3.
|
Brankin, B.,
M. N. Hart,
S. L. Cosby,
Z. Fabry, and I. V. Allen.
1995.
Adhesion molecule expression and lymphocyte adhesion to cerebral endothelium: effects of measles virus and herpes simplex 1 virus.
J. Neuroimmunol.
56:1-8[Medline].
|
| 4.
|
Campbell, J. J.,
J. Hedrick,
A. Zlotnik,
M. A. Siani,
D. A. Thompson, and E. C. Butcher.
1998.
Chemokines and the arrest of lymphocytes rolling under flow conditions.
Science
279:381-384[Abstract/Free Full Text].
|
| 5.
|
Chirgwin, J. M.,
A. E. Przybyla,
R. J. MacDonald, and W. J. Rutter.
1979.
Isolation of biologically active ribonucleic acid from sources enriched in ribonuclease.
Biochemistry
18:5294-5299[Medline].
|
| 6.
|
Davis-Poynter, N. J.,
D. M. Lynch,
H. Vally,
G. R. Shellam,
W. D. Rawlinson,
B. G. Barrell, and H. E. Farrell.
1997.
Identification and characterization of a G-protein-coupled receptor homolog encoded by murine cytomegalovirus.
J. Virol.
71:1521-1529[Abstract].
|
| 7.
|
del Pozo, M. A.,
P. Sanchez-Mateos,
M. Nieto, and F. Sanchez-Madrid.
1995.
Chemokines regulate cellular polarization and adhesion receptor redistribution during lymphocyte interaction with endothelium and extracellular matrix. Involvement of cAMP signaling pathway.
J. Cell Biol.
131:495-508[Abstract/Free Full Text].
|
| 8.
|
Doyle, M. V., and M. B. A. Oldstone.
1978.
Interactions between viruses and lymphocytes. I. In vivo replication of lymphocytic choriomeningitis virus in mononuclear cells during both chronic and acute viral infections.
J. Immunol.
121:1262-1269[Abstract/Free Full Text].
|
| 9.
|
Dunstan, C. A. N.,
M. N. Salafranca,
S. Adhikari,
Y. Xia,
L. Feng, and J. K. Harrison.
1996.
Identification of two rat genes orthologous to the human interleukin-8 receptors.
J. Biol. Chem.
271:32770-32776[Abstract/Free Full Text].
|
| 10.
|
Foster, C. A.,
M. Dreyfuss,
B. Mandak,
J. G. Meingassner,
H. U. Naegeli,
A. Nussbaumer,
L. Oberer,
G. Scheel, and E. M. Swoboda.
1994.
Pharmacological modulation of endothelial cell-associated adhesion molecule expression: implications for future treatment of dermatological diseases.
J. Dermatol.
21:847-854[Medline].
|
| 11.
|
Frankel, S. S.,
B. M. Wenig,
A. P. Burke,
P. Mannan,
L. D. Thompson,
S. L. Abbondanzo,
A. M. Nelson,
M. Pope, and R. M. Steinman.
1996.
Replication of HIV-1 in dendritic cell-derived syncytia at the mucosal surface of the adenoid.
Science
272:115-117[Abstract].
|
| 12.
|
Godiska, R.,
D. Chantry,
C. J. Raport,
V. L. Schweickart,
H. L. Trong, and P. W. Gray.
1997.
Monocyte chemotactic protein-4: tissue-specific expression and signaling through CC chemokine receptor-2.
J. Leuko. Biol.
61:353-360[Abstract].
|
| 13.
|
Godiska, R.,
D. Chantry,
C. J. Raport,
S. Sozzani,
P. Allavena,
P. D. Leviten,
A. Mantovani, and P. W. Gray.
1997.
Human macrophage-derived chemokine (MDC), a novel chemoattractant for monocytes, monocyte-derived dendritic cells, and natural killer cells.
J. Exp. Med.
185:1595-1604[Abstract/Free Full Text].
|
| 14.
|
Hedrick, J. A.,
V. Saylor,
D. Figueroa,
L. Mizoue,
Y. Xu,
S. Menon,
J. Abrams,
T. Handel, and A. Zlotnik.
1997.
Lymphotactin is produced by NK cells and attracts both NK cells and T cells in vivo.
J. Immunol.
158:1533-1540[Abstract].
|
| 15.
|
Hirsch, V. M.,
G. Dapolito,
P. R. Johnson,
W. R. Elkins,
W. T. London,
R. J. Montali,
S. Goldstein, and C. Brown.
1995.
Induction of AIDS by simian immunodeficiency virus from an African green monkey: species-specific variation in pathogenicity correlates with the extent of in vivo replication.
J. Virol.
69:955-967[Abstract].
|
| 16.
|
Huang, K.,
S. Y. Im,
W. E. Samlowski, and R. A. Daynes.
1989.
Molecular mechanisms of lymphocyte extravasation. III. The loss of lymphocyte extravasation potential induced by pertussis toxin is not mediated via the activation of protein kinase C.
J. Immunol.
143:229-238[Abstract].
|
| 17.
|
Kennedy, M. K.,
D. S. Torrance,
K. S. Picha, and K. M. Mohler.
1992.
Analysis of cytokine mRNA expression in the central nervous system of mice with experimental autoimmune encephalomyelitis reveals that IL-10 mRNA expression correlates with recovery.
J. Immunol.
149:2496-2505[Abstract].
|
| 18.
|
Kong, A. S., and S. I. Morse.
1977.
The in vitro effects of Bordetella pertussis lymphocytosis-promoting factor on murine lymphocytes. I. Proliferative response.
J. Exp. Med.
145:151-162[Abstract/Free Full Text].
|
| 19.
|
Marker, O.,
A. Scheynius,
J. P. Christensen, and A. R. Thomsen.
1995.
Virus-activated T cells regulate expression of adhesion molecules on endothelial cells in sites of infection.
J. Neuroimmunol.
62:35-42[Medline].
|
| 20.
|
Martin, T.,
P. M. Cardarelli,
G. C. Parry,
K. A. Felts, and R. R. Cobb.
1997.
Cytokine induction of monocyte chemoattractant protein-1 gene expression in human endothelial cells depends on the cooperative action of NF-kappa B and AP-1.
Eur. J. Immunol.
27:1091-1097[Medline].
|
| 21.
|
Milman, G., and O. Sharma.
1994.
Mechanisms of HIV/SIV mucosal transmission.
AIDS Res. Hum. Retroviruses
10:1305-1312[Medline].
|
| 22.
|
Morse, S., and B. Barron.
1970.
Studies on the leukocytosis and lymphocytosis induced by Bordetella pertussis. III. The distribution of transfused lymphocytes in pertussis treated and normal mice.
J. Exp. Med.
132:663-672[Abstract].
|
| 23.
|
Morse, S., and S. Riester.
1966.
Studies on the leukocytosis and lymphocytosis induced by Bordetella pertussis.
J. Exp. Med.
125:401-408[Abstract].
|
| 24.
|
Munoz, J. J.
1988.
Action of pertussigen (pertussis toxin) on the host immune system, p. 173-192.
In
A. C. Wardlaw, and R. Parton (ed.), Pathogenesis and immunity in pertussis. John Wiley & Sons, New York, N.Y.
|
| 25.
|
Norkin, L. C.
1995.
Virus receptors: implications for pathogenesis and the design of antiviral agents.
Clin. Microbiol. Rev.
8:293-315[Abstract].
|
| 26.
|
Pauza, C. D.,
P. W. Hinds,
C. Yin,
T. S. McKechnie,
S. B. Hinds, and M. S. Salvato.
1997.
The lymphocytosis-promoting agent pertussis toxin affects virus burden and lymphocyte distribution in the SIV-infected rhesus macaque.
AIDS Res. Hum. Retroviruses
13:87-95[Medline].
|
| 27.
|
Pauza, C. D.,
M. Malkovsky, and M. S. Salvato.
1994.
SIV transmission across the mucosal barrier.
AIDS Res. Hum. Retroviruses
10:S7-S10.
|
| 28.
|
Rai, S. K.,
D. S. Cheung,
M. S. Wu,
T. F. Warner, and M. S. Salvato.
1996.
Murine infection with lymphocytic choriomeningitis virus following gastric inoculation.
J. Virol.
70:7213-7218[Abstract/Free Full Text].
|
| 29.
|
Rai, S. K.,
B. K. Micales,
M. S. Wu,
D. S. Cheung,
T. D. Pugh,
G. E. Lyons, and M. S. Salvato.
1997.
Timed appearance of lymphocytic choriomeningitis virus after gastric inoculation of mice.
Am. J. Pathol.
151:633-639[Abstract].
|
| 30.
|
Rossen, R. D.,
C. W. Smith,
A. H. Laughter,
C. A. Noonan,
D. C. Anderson,
W. M. McShan,
M. Y. Hurvitz, and F. M. Orson.
1989.
HIV-1 stimulated expression of CD11/CD18 integrins and ICAM-1: a possible mechanism for extravascular dissemination of HIV-1 infected cells.
Trans. Am. Assoc. Physicians
102:117-130[Medline].
|
| 31.
|
Rubbert, A.,
D. Weissman,
C. Combadiere,
K. A. Pettrone,
J. A. Daucher,
P. M. Murphy, and A. S. Fauci.
1996.
Human monocyte chemotactic proteins-2 and -3: structural and functional comparison with MCP-1.
J. Leuko. Biol.
59:67-74[Abstract].
|
| 31a.
| Salvato, M. Unpublished data.
|
| 32.
|
Salvato, M. S., and E. M. Shimomaye.
1989.
The completed sequence of lymphocytic choriomeningitis virus reveals a unique RNA structure and a gene for a zinc finger protein.
Virology
173:1-10[Medline].
|
| 33.
|
Schall, T.
1997.
Fractalkinea strange attractor in the chemokine landscape.
Immunol. Today
18:147[Medline].
|
| 34.
|
Schmitt, D., and C. Dezutter-Dambuyant.
1994.
Epidermal and mucosal dendritic cells and HIV1 infection.
Pathol. Res. Pract.
190:955-959[Medline].
|
| 35.
|
Shimomaye, E., and M. Salvato.
1989.
Use of avian myeloblastosis virus reverse transcriptase at high temperature for sequence analysis of highly structured RNA.
Gene Anal. Tech.
6:25-28[Medline].
|
| 36.
|
Spangrude, G. J.,
B. A. Araneo, and R. A. Daynes.
1985.
Site-selective homing of antigen-primed lymphocyte populations can play a crucial role in the efferent limb of cell-mediated immune responses in vivo.
J. Immunol.
134:2900-2907[Abstract].
|
| 37.
|
Sugimoto, M.,
A. H. Sharpe,
Y. Sato,
M. I. Greene, and B. N. Fields.
1990.
Reovirus transportstudies using lymphocytosis promoting factor.
Pathobiology
58:185-192[Medline].
|
| 38.
|
Takahashi, M.,
U. Ikeda,
T. Kasahara,
S. Kitagawa,
Y. Takahashi,
K. Shimada,
S. Kano,
C. Morimoto, and J. Masuyama.
1997.
Activation of human monocytes for enhanced production of interleukin-8 during transendothelial migration in vitro.
J. Clin. Immunol.
17:53-62[Medline].
|
| 39.
|
Tyler, K. L.,
M. A. Mann,
B. N. Fields, and H. W. Virgin, IV.
1993.
Protective anti-reovirus monoclonal antibodies and their effects on viral pathogenesis.
J. Virol.
67:3446-3453[Abstract/Free Full Text].
|
| 40.
|
Volpes, R.,
J. J. van den Oord,
V. J. Desmet, and S. H. Yap.
1992.
Induction of intercellular adhesion molecule-1 (CD54) on human hepatoma cell line HepG2: influence of cytokines and hepatitis B virus-DNA transfection.
Clin. Exp. Immunol.
87:71-75[Medline].
|
| 41.
|
Walker, D. H., and F. A. Murphy.
1987.
Pathology and pathogenesis of arenavirus infections.
Curr. Top. Microbiol. Immunol.
133:89-113[Medline].
|
Journal of Virology, November 1998, p. 8613-8619, Vol. 72, No. 11
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Yang, Y., Tikhonov, I., Ruckwardt, T. J., Djavani, M., Zapata, J. C., Pauza, C. D., Salvato, M. S.
(2003). Monocytes Treated with Human Immunodeficiency Virus Tat Kill Uninfected CD4+ Cells by a Tumor Necrosis Factor-Related Apoptosis-Induced Ligand-Mediated Mechanism. J. Virol.
77: 6700-6708
[Abstract]
[Full Text]
-
Yin, C., Wu, M. S., Pauza, C. D., Salvato, M. S.
(1999). High Major Histocompatibility Complex-Unrestricted Lysis of Simian Immunodeficiency Virus Envelope-Expressing Cells Predisposes Macaques to Rapid AIDS Progression. J. Virol.
73: 3692-3701
[Abstract]
[Full Text]
Top
Abstract
Introduction
