Previous Article | Next Article 
Journal of Virology, October 1999, p. 8587-8598, Vol. 73, No. 10
0022-538X/99/$04.00+0
African Swine Fever Virus Replication in the Midgut Epithelium
Is Required for Infection of Ornithodoros
Ticks
S. B.
Kleiboeker,1,*
G. A.
Scoles,1,2
T. G.
Burrage,1 and
J.-H.
Sur1
Plum Island Animal Disease Center,
Agricultural Research Service, U.S. Department of Agriculture,
Greenport, New York 11944,1 and
Department of Epidemiology and Public Health, Yale
University School of Medicine, New Haven, Connecticut
065202
Received 26 April 1999/Accepted 12 July 1999
 |
ABSTRACT |
Although the Malawi Lil20/1 (MAL) strain of African swine fever
virus (ASFV) was isolated from Ornithodoros sp. ticks, our attempts to experimentally infect ticks by feeding them this strain failed. Ten different collections of Ornithodorus porcinus
porcinus ticks and one collection of O. porcinus
domesticus ticks were orally exposed to a high titer of MAL. At 3 weeks postinoculation (p.i.), <25% of the ticks contained detectable
virus, with viral titers of <4 log10 50% hemadsorbing
doses/ml. Viral titers declined to undetectability in >90% of the
ticks by 5 weeks p.i. To further study the growth defect, O. porcinus porcinus ticks were orally exposed to MAL and assayed at
regular intervals p.i. Whole-tick viral titers dramatically declined
(>1,000-fold) between 2 and 6 days p.i., and by 18 days p.i., viral
titers were below the detection limit. In contrast, viral titers of
ticks orally exposed to a tick-competent ASFV isolate,
Pretoriuskop/96/4/1 (Pr4), increased 10-fold by 10 days p.i. and
50-fold by 14 days p.i. Early viral gene expression, but not extensive
late gene expression or viral DNA synthesis, was detected in the
midguts of ticks orally exposed to MAL. Ultrastructural analysis
demonstrated that progeny virus was rarely present in ticks orally
exposed to MAL and, when present, was associated with extensive
cytopathology of phagocytic midgut epithelial cells. To determine if
viral replication was restricted only in the midgut epithelium,
parenteral inoculations into the hemocoel were performed. With
inoculation by this route, a persistent infection was established
although a delay in generalization of MAL was detected and viral titers
in most tissues were typically 10- to 1,000-fold lower than those of
ticks injected with Pr4. MAL was detected in both the salivary
secretion and coxal fluid following feeding but less frequently and at
a lower titer compared to Pr4. Transovarial transmission of MAL was not
detected after two gonotrophic cycles. Ultrastructural analysis
demonstrated that, when injected, MAL replicated in a number of cell
types but failed to replicate in midgut epithelial cells. In contrast, ticks injected with Pr4 had replicating virus in midgut epithelial cells. Together, these results indicate that MAL replication is restricted in midgut epithelial cells. This finding demonstrates the
importance of viral replication in the midgut for successful ASFV
infection of the arthropod host.
| |
INTRODUCTION |
African swine fever (ASF) is a
lethal, hemorrhagic disease of domestic pigs for which animal slaughter
and area quarantine are the only methods of control. ASF virus (ASFV),
the causative agent of ASF, is a large, double-stranded DNA virus which
is the only member of the Asfarviridae family and the only
known DNA arbovirus (5, 6, 9). The genome of ASFV is
relatively large, consisting of approximately 180 kbp encoding at least
165 genes. Under a variety of experimental and natural conditions, ASFV
infectivity has been shown to be very resistant to inactivation (26). For example, ASFV remained viable for up to 140 days
in defibrinated blood held at room temperature (28). In
nature, ASFV infects both warthogs (Phacochoerus
aethiopicus) and bushpigs (Potamochoerus spp.), as well
as ticks of the genus Ornithodoros (34). The
natural arthropod host of ASFV is Ornithodoros porcinus porcinus (Walton) (31), a long-lived and nidicolous
(burrow-dwelling) argasid tick. Both the vertebrate and arthropod hosts
are likely to be required for maintenance of ASFV in the sylvatic
cycle, and persistently infected ticks serve as a natural reservoir of the virus. The mechanism of ASFV transmission from the sylvatic cycle
to domestic pigs is most likely through infected
Ornithodoros ticks feeding on pigs (32, 38),
since direct contact with infected warthogs rarely results in
transmission to pigs (7, 21, 28, 38). The virus is
transmitted between domestic pigs by either direct or indirect
contact (26).
Previous studies have described experimental infection of O. porcinus porcinus ticks with a number of different ASFV isolates (15, 20, 33). Although details of the pathogenesis vary in
these reports, ASFV infection of O. porcinus porcinus ticks is characterized by establishment of a long-term, persistent infection with relatively high levels of viral replication in a number of different tissues and organs. The initial site of viral replication is
the midgut, suggesting a critical role for this tissue in the establishment of infection. The infectious dose of ASFV has been reported to be less than 1 log10 50% hemadsorbing dose
(HAD50)/ml for larger ticks (33, 36). ASFV
infection of O. porcinus porcinus ticks has been associated
with very low mortality (15, 22, 31, 33), except during the
gonotrophic cycle (20, 36). These data suggest that ASFV
infection of the natural arthropod host represents a well-adapted and
possibly coevolved biological system. However, differences in infection
rate, infectious dose, or the proportion of ticks which became
persistently infected were observed when ticks from the same collection
were exposed to different ASFV isolates (15, 33), results
which suggest that virus-host adaptation plays a role in the infection
of a natural arthropod host with a given ASFV isolate.
Here we describe the results of oral exposure and intrahemocoelic
inoculation of O. porcinus porcinus ticks with Malawi Li 20/1 (MAL), an ASFV isolate made from Ornithodoros sp. ticks
(8, 18). MAL did not infect ticks exposed orally, although
virus entry into midgut cells, early gene expression, and limited late gene expression and viral DNA synthesis occurred. In contrast, when MAL
was inoculated intrahemocoelically, a persistent viral infection was
established although a slight generalized replication defect was
observed. These results indicate that midgut infection and escape
constitute important barriers to generalization of ASFV infection of ticks.
| |
MATERIALS AND METHODS |
Virus isolates.
Pretoriuskop/96/4/1 (Pr4) was isolated from
O. porcinus porcinus ticks collected as previously described
(22). MAL was isolated in 1983 from Ornithodoros
sp. ticks collected from domestic pig structures during an ASFV
epizootic (8, 18). Chiredzi/83/1 (Ch1) was isolated from
Ornithodoros sp. ticks collected from warthog burrows near
Chiredzi, Zimbabwe, in 1983 (20).
Ticks.
Eight collections of ticks were made from warthog
burrows at various locations in Kruger National Park and the Northern
Transvaal region of the Republic of South Africa in 1996. A single
collection of ticks was made from warthog burrows in the Masai Mara
Reserve in Kenya in 1996. All of these ticks were classified as
O. porcinus porcinus in accordance with the criteria of
Walton (39). A single collection of ticks was made from
domestic pig structures in Chalaswa (Mchinji district), Malawi, in
1997. This collection has been identified as O. porcinus
domesticus in accordance with the criteria of Walton
(39). Ticks used for intrahemocoelic inoculations were from
an O. porcinus porcinus colony established from uninfected ticks in the original collection which yielded the Pr4 isolate. Ticks
used for reverse transcription (RT)-PCR experiments were from an
O. porcinus porcinus colony maintained for an indeterminate period of time at Plum Island Animal Disease Center. All ticks were
held at 26°C and a relative humidity of 76% with 12 h of light
per 24-h cycle.
RT-PCR analysis.
Ticks were exposed by feeding on an
artificial membrane feeder placed over a pool of heparinized pig blood
containing either Ch1 or MAL at 7.0 log10
HAD50/ml. Dissected tick midguts or whole ticks were frozen
in liquid nitrogen and then ground to a powder before thawing. Total
cellular RNA was prepared by using Tri Reagent (Sigma Chemical
Company). Samples were harvested at 2, 4, 10, and 24 days postfeeding.
Total cellular RNA (~5 µg) was treated with 100 U of RNase-free
DNase I (Boehringer Mannheim) for 4 h at 37°C. Samples were then
extracted with the RNeasy Mini Kit (Qiagen) and then denatured in the
presence of 2 µg of random hexamers (Gibco BRL Life Technologies) at
94°C for 5 min. RT was performed with 200 U of SuperScript RNase
H
reverse transcriptase (Gibco BRL Life Technologies) for
1 h at 45°C. The negative control for each time point was an
aliquot of each sample without addition of reverse transcriptase.
Resulting cDNAs were amplified by PCR for 30 cycles (94°C, 10 s;
58°C, 30 s; 72°C, 30 s) with a final 10-min incubation at
72°C. A second PCR amplification was performed by using the same
protocol with nested primer sets. The primers used for first-round
amplification were p72f1, (5'-GCGTTGTGACATCCGAACTA-3'),
p72r1 (5'-CAAGATTATATTGGCCCAAG-3'), p30f1
(5'-CCATGAGTCTTACCACCTCT-3'), and p30r1
(5'-GGAGGTCATCTTCAAAACGG-3'). The primers used for
second-round amplification were p72f2
(5'-CTCTAAAGGTGTTTGGTTGTC-3'), p72r2
(5'-ATTTTAAGCCTTATGTTCCAG-3'), p30f2
(5'-GAGGGGTTCCATGAATGGTT-3'), and p30r2
(5'-GTAGAATTGTTACGACCGCT-3').
Tick inoculations.
Ticks were exposed by feeding on an
artificial membrane feeder placed in heparinized pig blood. For the
18-day time course (Fig. 1), immunohistochemistry (IHC) and in situ
hybridization (ISH) experiments (see Fig. 2), and ultrastructural
experiments (see Fig. 3), the titer of the bloodmeal for both MAL- and
Pr4-exposed ticks was 7.3 log10 HAD50/ml. For
the experiment which compared orally exposed Pr4 to injected Pr4 (see
Fig. 4), orally exposed ticks were fed on a viremic pig. The viremic
titer of Pr4 on the day of tick feeding was 8.3 log10
HAD50/ml.
Intrahemocoelic inoculations were performed by injecting 3.9 log10 HAD50 of either MAL or Pr4 in a volume of
2 µl with a finely pulled, calibrated glass capillary pipet.
Injections were made in the membranous region located between the coxa
and trochanter of the second leg of either N4, N5, or adult ticks
weighing 10 to 25 mg. Viral stocks were prepared from dissected midguts
of 10 MAL-injected ticks 42 to 98 days postinoculation (p.i.) by inoculation of primary porcine peripheral blood mononuclear cell cultures (14, 30). Naive ticks were then fed on artificial membrane feeders placed in heparinized pig blood which contained the
resulting viral stocks at approximately 6.75 log10
HAD50/ml.
Virus titrations.
Individual whole ticks were ground in 0.5 ml of cell culture medium in sterile tubes with plastic pestles (Pellet
pestle; Kontes). The samples were stored at 
70°C. Immediately prior
to titration, samples were thawed at 37°C and sonicated for 1 min. Samples were serially diluted and then added to porcine peripheral blood mononuclear cells as previously described (22) and
endpoint titers were calculated (35).
To determine the virus titers in isolated organs, ticks were dissected
by using a binocular microscope and tissues were titrated
as previously
described (
22). The titer of hemolymph was obtained
by
clipping the distal tarsus of the second leg and collecting
1 to 2 µl
of hemolymph in a sterile tube. All samples were diluted
to 0.5 ml and
titrated as described
above.
For data presented in Table
3, individual ticks were fed on a membrane
feeding apparatus as previously described (
22,
25).
Following feeding, coxal fluid (if any was produced during a 2-h
postfeeding observation period) and blood (containing salivary
secretions) from beneath the membrane were collected, diluted
to 1 ml
with cell culture medium containing 20% fetal bovine serum,
and then
held at
70°C until assayed by virus titration. Values
were
normalized to the volume of the diluted sample. The undiluted
sample
and the first serial dilution were blind passaged for any
samples that
were negative for ASFV on the initial
titration.
Ultrastructural procedures.
At various times p.i., ticks
were fixed and embedded as previously described (22). For
analysis, 70- to 90-nm sections were collected on single-slot grids
coated with Formvar and stabilized with carbon (Electron Microscopy
Sciences) and then observed and photographed with a Philips 410 electron microscope operated at 80 kV.
IHC and ISH.
At the indicated times p.i., ticks were cut
along the sagittal plane to allow infiltration of the fixative
solutions (10% neutral buffered formalin), embedded in paraffin, and
sectioned. Sections were allowed to adhere to Superfrost/plus slides
(Fisher Scientific, Pittsburgh, Pa.), heated for 20 min at 65°C, and
then deparaffinized by using xylene. Sections were rehydrated through graded alcohol and washed with phosphate-buffered saline (PBS) (pH 7.4)
for IHC and ISH. Four-micrometer-thick sections were used for IHC and ISH.
IHC was performed essentially as described previously (
37).
Briefly, sections were first treated with 3% hydrogen peroxide
in PBS
for 20 min, followed by washes in PBS and digestion with
0.05%
Protease XIV (Sigma Chemical Co., St. Louis, Mo.) for 2
min at 37°C.
After several washes in PBS, sections were incubated
in a blocking
solution (5% normal goat serum in PBS) for 30 min
at room temperature
and then incubated for 2 h at 4°C with specific
polyclonal
antiserum (diluted 1:200 in PBS) directed against ASFV
structural
protein p30 or p72. Following washes with PBS, slides
were incubated
with alkaline phosphatase-conjugated, goat anti-rabbit
antibody for 20 min at room
temperature.
ISH was performed as described previously (
37), with minor
modifications. Probes containing both the p30 and p72 genes were
labeled by a random priming reaction with digoxigenin-dUTP (DIG;
Boehringer Mannheim Corp., Indianapolis, Ind.). Labeled DNA probes
were
diluted in the prehybridization mixture and then heated for
10 min at
95°C and placed on ice before being applied to the sections.
Coverslips were applied, and the target DNA was denatured by placing
the slides for 5 min on a hot plate preheated to 96°C. The slides
were then placed on ice for 3 min. Hybridization was performed
overnight at 37°C. An anti-DIG-alkaline phosphatase conjugate
(diluted to 1:500) was then added to the tissue sections and incubated
for 2 h at room temperature. Sections were then incubated with
color substrate solution for 2 to 3 h in the dark, and then the
reaction was stopped with distilled water. Sections were counterstained
with 0.5% methyl
green.
| |
RESULTS |
MAL does not infect ticks following oral inoculation.
Ten
different collections of O. porcinus porcinus ticks and one
collection of O. porcinus domesticus ticks were orally
exposed to high titers of MAL (Table 1).
Based on the size of the bloodmeal, a minimum of 6 to 7 log10 HAD50 was inoculated into each tick. By 3 weeks postfeeding, less than 25% of the ticks contained detectable virus and in the ticks in which virus could be detected, titers had
fallen by at least 100-fold. By five weeks p.i., virus could be
detected in less than 10% of the ticks and the titers had fallen by
10,000-fold.
In a second experiment, ticks from the Pr4 colony were orally exposed
to either MAL or Pr4 and sampled regularly between 0
and 18 days p.i.
(Fig.
1). One group was fed an inoculum
containing
swine blood, and the second group was fed with fetal bovine
serum
in place of the blood. In both groups, MAL titers declined
1,000-fold
between 2 and 6 days p.i. By 14 to 18 days p.i., no virus
was
detected in MAL-exposed ticks. In contrast, titers of Pr4 increased
by 10-fold at 10 days p.i. and 50-fold by 14 days p.i. In an
independent
replicate, MAL titers were not detected at 14, 21, 28, or
60 days
following oral exposure (data not shown). As a control for the
stability of the virus remaining in the midgut lumen, a portion
of each
group was killed by momentary freezing in liquid nitrogen
and then
returned to the same holding conditions as the live ticks
(Fig.
1). For
ticks exposed to virus in blood and then killed
immediately p.i., viral
titers did not decline appreciably over
the sampling period. For ticks
exposed to virus in serum and then
killed, viral titers were stable
until 14 days p.i., after which
they declined 100- to 1,000-fold.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 1.
Viral replication in O. porcinus porcinus
ticks following oral exposure. Ticks were exposed by membrane feeding.
The inoculum contained MAL or Pr4 diluted in either heparinized pig
blood (A) or fetal bovine serum in place of pig blood (B). Immediately
postfeeding, a portion of each group was killed by momentary freezing
in liquid nitrogen. At the indicated times p.i., individual ticks
(n = 4) from each group were ground in cell culture
medium and titrations were performed. The values are mean titers ± the standard errors of the means.
|
|
Limited MAL gene expression and viral DNA synthesis occur following
oral inoculation.
ASFV gene expression in ticks orally exposed to
MAL was assessed by RT-PCR at various times p.i. (Table
2). Assays were performed for expression
of genes encoding structural proteins p30 and p72, which are encoded by
an early and a late gene in ASFV infection of swine macrophages,
respectively (1, 4). Following oral inoculation with MAL,
p30 expression was detected in the midguts of 90% of the ticks at 2 days p.i., 60% of the ticks at 4 days p.i., and none of the ticks at
10 days p.i. In contrast, p30 expression in ticks orally exposed to
Ch1, an ASFV isolate which has been previously shown to replicate in
ticks (22), was detected in a majority of the ticks sampled
at each time point. Expression of the late gene encoding p72 was
detected at low frequency (22% of dissected midguts) only at 10 days
p.i. following oral exposure to MAL but was detected at a considerably
higher frequency in Ch1-exposed ticks.
IHC performed at 3, 6, and 11 days p.i. using polyclonal antibodies
against p30 and p72 corroborated the RT-PCR results. At
6 days p.i.,
p30, but not p72, could be readily detected in midgut
epithelial cells
following oral inoculation of MAL (Fig.
2A).
At 11 days p.i., p72 expression
could be detected, but only in
an occasional section (Fig.
2C). In
contrast, both p30 and p72
were expressed in a majority of midgut cells
at both 6 and 11
days p.i. following oral inoculation with Pr4
(Fig.
2B and D).
When midgut sections were analyzed by ISH for ASFV DNA
at 3, 6,
and 11 days p.i., a small percentage of midgut cells were
found
to be weakly positive in MAL-exposed ticks (Fig.
2E) at 11 days
p.i. However, for Pr4-exposed ticks, many more midgut cells were
positive for ASFV DNA by this technique and the intensity of staining
was greater in Pr4-exposed ticks than in MAL-exposed ticks (Fig.
2F).
View larger version (114K):
[in this window]
[in a new window]
|
FIG. 2.
IHC and ISH analysis of midgut epithelial cells. Ticks
were exposed by membrane feeding on an inoculum containing either MAL
(A, C, and E) or Pr4 (B, D, and E). Analysis for ASFV protein p30 (A
and B) or p72 (C and D) or ASFV DNA (E and F) was performed at 6 (A and
B) or 11 (C, D, E, and F) days p.i. H, hemocoel; L, midgut lumen.
|
|
Ultrastructural analysis of orally exposed ticks was performed at 4, 6, and 10 days p.i. In MAL-exposed ticks, viral replication
factories were
rarely observed in midgut epithelial cells. When
present, MAL factories
were comparatively smaller and contained
few fully formed
viral particles (Fig.
3B). These
particles often
had an atypical morphology characterized by either an
acentric
or absent condensed viral nucleoid structure (Fig.
3D). When
MAL
factories were observed, the cells which contained them had a
fragmented cellular membrane, mitochondrial condensation, pulling
away
of the nuclear membrane, and extensive vacuolization (Fig.
3B).
Sloughed digestive midgut epithelial cells and free nuclei
(Fig.
3B)
were commonly observed in the midgut lumen. These observations
indicated considerable cytopathology in phagocytic midgut epithelial
cells following oral inoculation of MAL. In Pr4-exposed ticks,
virus
factories were commonly observed at 6 days p.i. Numerous
large virus
factories with a large number of fully formed particles
were present in
the midgut epithelial cells of these ticks (Fig.
3A and C). In
Pr4-exposed ticks, viral factories were similar,
if not identical, to
those observed following oral exposure to
the Ch1 isolate
(
22). The phagocytic midgut epithelial cells
which contained
Pr4 viral factories showed minimal evidence of
cellular pathology, and
sloughed midgut epithelial cells were
rarely observed in the lumen of
Pr4-infected ticks.
View larger version (326K):
[in this window]
[in a new window]
|
FIG. 3.
ASFV in an O. porcinus porcinus midgut. Ticks
were exposed by membrane feeding. The inoculum contained Pr4 (A and C)
or MAL (B and D). Analysis was performed 6 days p.i. (A) Virus factory
(VF) in a midgut epithelial cell from a Pr4-exposed tick. (B) Virus
factory in a midgut epithelial cell from an MAL-exposed tick. (C)
Higher magnification of a virus factory from a Pr4-exposed tick. (D)
Higher magnification of a virus factory from an MAL-exposed tick. N,
nucleus; L, midgut lumen; NF, nucleus free in the midgut lumen.
|
|
A persistent infection is established by intrahemocoelic
inoculation of MAL.
To determine if MAL replication was restricted
in tissues other than the midgut, parenteral inoculations into the
hemocoel were performed. In contrast with oral exposure to MAL,
intrahemocoelic inoculation resulted in a generalized, persistent
infection. Although total tick MAL titers did not increase through 27 days p.i., by 50 days p.i., the total-tick viral titer and most tissue
viral titers had increased 10- to 100-fold (Fig.
4). Maximum MAL tissue titers, which
occurred at 98 days p.i., were at least 100-fold higher than titers
measured immediately p.i. However, even at their maximum, most MAL
tissue titers were 10- to 100-fold lower than Pr4 titers. The only
exception was the hemolymph titer, which was 50-fold higher in
MAL-injected ticks than in Pr4-injected ticks at 98 days p.i. After 98 days p.i., MAL tissue including hemolymph, titers declined
approximately 10- to 100-fold but did not decrease further during the
period of study (364 days). The infection rate for MAL-injected ticks
was 100% (40 of 40 ticks sampled). Intrahemocoelic injection of Pr4
resulted in rapid dissemination of infection to all of the tissues
assayed. By the first sampling at 12 days p.i., total tick Pr4 titers
had increased 1,000-fold to 6.1 log10 HAD50/mg
and did not decrease appreciably for the duration of the study. Most
tissue viral titers peaked by 12 days p.i. In contrast to all of the
other tissues assayed, viral titers of the hemolymph declined
dramatically in Pr4-injected ticks after the initial sampling at 12 days p.i. The infection rate of Pr4-injected ticks was 100% (40 of 40 ticks sampled). As a control for the intrahemocoelic route of
infection, tissue viral titers from ticks exposed to Pr4 by this route
were compared to the titers of ticks exposed orally to Pr4 (Fig. 4).
Although intrahemocoelic inoculation is an unnatural route of
infection, once generalization of infection occurred, viral titers in
injected ticks were not different than those in orally exposed ticks.
View larger version (23K):
[in this window]
[in a new window]
|
FIG. 4.
Viral replication in O. porcinus porcinus
ticks following intrahemocoelic (intrahemo.) or oral inoculation
(inoc.). Ticks were infected by intrahemocoelic injection of either Pr4
or MAL or by feeding on a pig infected with Pr4. Individual ticks
(n = 4) from each group were dissected at the indicated
times p.i., and tissue viral titers were determined. Total-tick
viral titers represent the sum of all dissected and undissected
tissues. The values are mean titers ± the standard errors of the
means.
|
|
At 98 days p.i., MAL midgut titers were approximately 3.2 log
10 HAD
50/mg and midgut viral titers in
Pr4-injected ticks were
nearly equivalent to those in ticks orally
exposed to Pr4 (Fig.
4). These data suggested that following injection,
virus may have
crossed the basal lamina of the midgut and was
replicating in
midgut epithelial cells. To determine if this had
occurred, ultrastructural
analysis of injected ticks was performed.
Analysis of 1,195 midgut
epithelial cells (in 24 cecae from three
ticks) of MAL-injected
ticks failed to identify a single infected
midgut epithelial cell.
However, mature MAL particles and small viral
factories were observed
in connective tissue cells and hemocytes which
adhered to (or
were closely associated with) the hemocoelic side of the
midgut
(Fig.
5A) and thus likely
contributed to the titers observed in
dissected midguts (Fig.
4). In
contrast, ultrastructural analysis
of Pr4-injected ticks demonstrated
that Pr4 had crossed the basal
lamina from the hemocoel and was
replicating in midgut epithelial
cells (Fig.
5B). Approximately 7% (93 of 1,223 cells in 24 cecae
from three ticks) of midgut epithelial cells
from Pr4-injected
ticks were observed to have virus factories.
View larger version (157K):
[in this window]
[in a new window]
|
FIG. 5.
ASFV in an O. porcinus porcinus midgut.
Analysis was performed 14 weeks after intrahemocoelic inoculation. (A)
Mature virions (arrow) budding from a connective tissue cell adjacent
to the basal lamina (BL) of the midgut in an MAL-injected tick. (B)
Virus factory (VF) in a midgut epithelial cell from a Pr4-injected
tick. H, hemocoel; L, lumen; M, muscle; N, nucleus.
|
|
Virus isolated from midguts of MAL-injected ticks was refed to naive
ticks, but this failed to result in an infection. Viral
stocks were
prepared from the midguts of 10 ticks (dissected from
42 to 98 p.i.) and then refed to ticks from the same colony. At
29 days p.i., 12 ticks per midgut sample were assayed individually
for virus. All of the
ticks from 8 of the 10 groups contained
no detectable virus while two
groups had a single positive tick
with a titer of
2.5
log
10 HAD
50. A subsequent passage of virus
isolated from these two ticks failed to infect any naive ticks.
Thus,
even after a period which would allow adaptation to midgut
epithelial
cells, MAL could not establish an infection when administered
by the
oral
route.
To assess the ability of MAL-injected ticks to transmit the virus
during feeding, individual ticks were fed on artificial
membrane
feeders (Table
3). Prior to 50 days p.i.,
only a single
sample from an MAL-injected tick contained the virus and
the titer
of this coxal fluid sample was low. Sixty-six percent of the
salivary
secretion samples and all of the coxal fluid samples from
Pr4-injected
ticks contained considerable levels of the virus. Between
65 and
101 days p.i., 60% of MAL-injected ticks secreted the virus in
the coxal fluid; however, the presence of the virus in the salivary
secretion was still rare (1 of 11 samples). In contrast, all of
the
salivary secretion and coxal fluid samples from Pr4-injected
ticks
contained the virus after 65 days p.i. After 117 days p.i.,
38% of the
salivary secretion samples and 91% of the coxal fluid
samples from
MAL-injected ticks were positive. In addition to
a delay before
transmission and a lower frequency of positive
samples from
MAL-injected ticks, positive samples had a mean titer
that was
typically 50- to 100-fold lower than that of Pr4-injected
controls.
Assessment of transovarial transmission demonstrated that MAL-injected
female ticks were not capable of transmitting the virus
to their
offspring. Approximately 120 days p.i., MAL- and Pr4-injected
females
were mated with uninfected males from the same colony
and then fed
individually on uninfected blood. All of the resulting
first-stage
nymphs (
n = 24 per egg mass) from MAL-injected females
were negative when assayed for the virus after both the first
(
n = 8 egg masses) and second (
n = 6
egg masses) gonotrophic cycles.
In contrast, all (
n = 5) egg masses from the first gonotrophic
cycle of Pr4-injected
females contained first-stage nymphs that
were positive for the virus
and the mean infection rate was 49%
of the nymphs (
n = 24 per egg
mass).
| |
DISCUSSION |
MAL is an ASFV isolate that is highly pathogenic for domestic
pigs. Although the MAL we used had been isolated from a pool of four
adult male Ornithodoros sp. ticks collected in a domestic pig structure during an ASF epizootic (8, 18), our attempts to infect ticks by oral exposure to MAL failed. In other studies, oral
exposure of O. porcinus porcinus ticks to ASFV resulted in an infection characterized by primary replication in the midgut, followed by dissemination, with relatively high tissue viral titers and
persistence of infection for at least several months or, in some
studies, for several years (15, 22, 33). While the data
presented in Table 1 suggest that a portion of the ticks may have been
infected, this infection is likely to have been abortive
(nonproductive) since both the virus titers and the number of ticks
containing virus (for all collections) declined between 21 and 35 days
postfeeding. Based on data collected in all other experiments, the
titers detected in this initial experiment were most likely due to the
high titer of the inoculum coupled with the stability of the virus in
the midgut lumen. This may also be the reason why MAL was originally
isolated from ticks, since it is likely that ticks had ample
opportunity to feed on pigs with viremic titers in excess of 8 log10 HAD50/ml. Alternatively, we have observed
that gut contents will occasionally (<2% of the ticks fed) leak into
the hemocoel during or shortly after feeding without tick mortality
(36). This occurrence would have the same effect as
intrahemocoelic inoculation of MAL and could have resulted in
persistent infection of the field-collected ticks from which MAL was
originally isolated. Leakage of midgut contents into the hemocoel
following blood feeding is known to occur in mosquitos (17).
However, the incidence documented in one study (16% of the mosquitos
fed) (41) was much higher than what we have observed in
Ornithodoros ticks.
Following oral exposure to MAL, there is a rapid decline in viral
infectivity and after 18 days p.i., no virus was detected in these
ticks. The results of RT-PCR, IHC, and ISH experiments all suggest that
MAL enters midgut epithelial cells following oral exposure. The decline
in MAL titers is not due to degradation of the virus in the midgut
lumen (following failure or delay of virus entry into midgut epithelial
cells), since titers in ticks which were killed immediately following
feeding declined at a much slower rate than in live ticks.
Additionally, since nearly identical declines in MAL titers were
obtained for ticks exposed either with or without erythrocytes, the
loss of infectivity is not due to virus being internalized via
hemadsorption to erythrocytes, which are subsequently phagocytosed and
digested within midgut epithelial cells. The results of RT-PCR and IHC
experiments also demonstrate that early gene expression occurred at
levels similar to that detected after inoculation with a virus that can
infect ticks by the oral route. However, both late gene expression and viral DNA synthesis occurred at comparatively lower levels in MAL-exposed ticks. Thus, the restriction of MAL replication in midgut
epithelial cells is likely to occur late in the viral replication cycle. As would be predicted from these results, the appearance of
progeny virus was quite rare in midgut epithelial cells of MAL-exposed ticks.
When MAL viral factories were observed in midgut epithelial cells, the
mature particles often had atypical morphology, suggesting that they
are noninfectious. Interestingly, the phagocytic midgut epithelial cells which contained MAL particles exhibited cellular pathology which included condensation of chromatin, pulling away of the
nuclear membrane, roughening of the cell border, and sloughing of cells
into the midgut lumen. Cytopathology has been rarely observed following
arboviral infection of the natural host. In previous studies,
pathologic effects were not detected following infection of ixodid
ticks with Thogoto virus (2) or Dugbe virus (3)
but have been reported following infection of mosquitos with eastern
equine encephalomyelitis virus (40) and western equine
encephalomyelitis virus (42). Additional work is
required to establish a correlation between nonproductive
infection with MAL and the death of midgut cells. Early death of midgut
cells may be a mechanism which prevents productive infection of MAL in
this tissue and could possibly be due to failure of the virus to
express specific genes which allow the production of large quantities
of progeny virus without damaging the host cell. Alternatively, early cell death may be an active mechanism of host defense and MAL may
lack the viral gene(s) required to counter this response. It is
interesting that, despite obvious cytopathology, we have observed no
increase in the mortality of MAL-exposed ticks over that of Pr4-exposed
controls (24).
There are several possible explanations for the failure of MAL to
infect ticks following oral exposure. First, the natural arthropod host
for MAL may be a species or subspecies of Ornithodoros ticks
which was not tested in this study and/or MAL may have a very narrow
arthropod host range. However, the 11 tick collections in which MAL
infectivity was tested represent at least four geographic regions of
sub-Saharan Africa. Most significantly, the O. porcinus domesticus ticks tested were collected in the same village from which MAL was isolated 13 years earlier. Based on the work of others
and our unpublished results, it appears that all of the other ASFV
isolates studied to date will infect most, but not all, collections
of O. porcinus porcinus ticks (24, 33), as well
as a large number of distantly related congeners, such as O. savignyi (27), O. coriaceus (16,
19), O. turicata (19), O. puertoricensis (10, 11, 19), and O. marocanus (12, 13). Second, although the MAL virus
stock was subjected to a limited number of subpassages, these passages
were in either domestic pigs or porcine peripheral blood mononuclear
cells. Thus, rearrangements or deletions of genes important for
infection of midgut epithelial cells may have occurred since MAL was
isolated from ticks. However, it is significant that Pr4 has been
subjected to numerous successive rounds of plaque purification on
porcine peripheral blood mononuclear cells without any detectable loss
of the ability to infect ticks (24), a result which suggests
that this is not a likely explanation for the MAL defect. Finally, as
suggested above, MAL may be a virus that is not capable of
infecting ticks following oral inoculation but was isolated from
ticks following feeding on a pig with a high viremic titer.
To determine if the inability to replicate was specific to midgut
epithelial cells, MAL was injected intrahemocoelically into O. porcinus porcinus ticks. When MAL was inoculated by this route, a
generalized and persistent viral infection occurred in 100% of the
ticks. MAL-injected ticks secreted the virus during feeding but did not
transmit the virus transovarially. While the tissue distribution of the
virus was similar to that in Pr4-injected ticks, differences in viral
tissue titers and the time course of infection were observed. On
average, MAL tissue titers were 10- to 100-fold lower than Pr4 titers
and there was a delay of approximately 40 days before MAL titers began
to increase. As expected, transmission of MAL during feeding was also
delayed and both the frequency and the quantity of the virus excreted were reduced compared to those of Pr4. Therefore, MAL has a slight generalized replication defect in O. porcinus porcinus
ticks. However, the midgut is the only tissue in which MAL has an
absolute replication defect, suggesting a narrow genetic basis for this phenotype. The midgut replication defect appears to be stable, since
even after a period which would allow adaptation (up to 98 days
following intrahemocoelic injection), MAL was not detected in midgut
epithelial cells and reinoculation of MAL isolated from injected ticks
failed to infect ticks by the oral route. These results demonstrate
conclusively that midgut infection and escape constitute barriers to
dissemination of MAL and, presumably, other ASFV isolates following
oral exposure of O. porcinus porcinus ticks. An earlier ASFV
pathogenesis study suggested that a midgut barrier did not exist in
O. porcinus porcinus ticks, but the supporting data were
indirect (15).
In summary, we have characterized the phenotype of an ASFV isolate that
does not replicate in the midgut epithelium of the arthropod host,
O. porcinus porcinus, and thus cannot establish an infection
when these ticks feed on viremic blood. MAL differs from other ASFV
isolates previously studied in its inability to infect O. porcinus porcinus ticks following oral exposure. The MAL midgut
defect is likely to be due to missing or defective viral genes, and
future experiments can now be designed to elucidate the gene(s)
required for infection of the midgut epithelium, the first and most
important tissue encountered by a blood-borne pathogen infecting a
hematophagous arthropod. Identification of the gene(s) important for
ASFV infection of the midgut is feasible since the ASFV genome can be
manipulated through deletion, addition, or substitution of specific
genes and/or regions of the viral genome by either reverse genetics or
marker rescue techniques (23, 29, 30, 43, 44). Since
Ornithodoros ticks are an important route by which ASFV
moves from the sylvatic cycle to domestic pigs in sub-Saharan Africa,
knowledge of the viral genes required to infect ticks may suggest novel
control methods.
| |
ACKNOWLEDGMENTS |
We thank Daniel L. Rock and Douglas M. Moore for valuable
discussions and critical reviews of the manuscript. We also thank Gray
Matita (Central Veterinary Laboratory, Lilongwe, Malawi) for
collecting ticks from the Chalaswa village in Malawi, Stefan Swanepoel (Onderstepoort Institute for Exotic Animal Diseases) for assistance in collecting ticks from South Africa, and Cherise Rohr
(Yale University School of Medicine) for collecting ticks from the
Masai Mara Reserve, Kenya.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Plum Island
Animal Disease Center, P.O. Box 848, Greenport, NY 11944-0848. Phone:
(516) 323-3337. Fax: (516) 323-2507. E-mail:
skleiboe{at}asrr.arsusda.gov.
| |
REFERENCES |
| 1.
|
Afonso, C.,
C. Alcaraz,
A. Brun,
M. D. Sussman, and D. L. Rock.
1992.
Characterization of p30, a highly antigenic membrane and secreted protein of African swine fever virus.
Virology
189:368-373[Medline].
|
| 2.
|
Booth, T. F.,
C. R. Davies,
L. D. Jones,
D. Staunton, and P. A. Nuttall.
1989.
Anatomical basis of Thogoto virus infection in BHK cell culture and in the Ixodid tick vector, Rhipicephalus appendiculatus.
J. Gen. Virol.
70:1093-1104[Abstract/Free Full Text].
|
| 3.
|
Booth, T. F.,
G. M. Steele,
A. C. Marriott, and P. A. Nuttall.
1991.
Dissemination, replication, and trans-stadial persistence of Dugbe virus (Nairovirus, Bunyaviridae) in the tick vector Amblyomma variegatum.
Am. J. Trop. Med. Hyg.
45:146-157.
|
| 4.
|
Borca, M. V.,
P. Irusta,
C. Carrillo,
C. L. Afonso,
T. Burrage, and D. L. Rock.
1994.
African swine fever virus structural protein p72 contains a conformational neutralizing epitope.
Virology
201:413-418[Medline].
|
| 5.
|
Brown, F.
1986.
The classification and nomenclature of viruses: summary of results of meetings of the International Committee on Taxonomy of Viruses in Sendai, September 1984.
Intervirology
25:141-143.
|
| 6.
|
Costa, J. V.
1990.
African swine fever virus, p. 247-270.
In
G. Darai (ed.), Molecular biology of iridoviruses. Kluwer Academic Publishers, Norwell, Mass.
|
| 7.
|
DeTray, D. E.
1963.
African swine fever.
Adv. Vet. Sci. Comp. Med.
8:299-333.
|
| 8.
|
Dixon, L. K.
1988.
Molecular cloning and restriction enzyme mapping of an African swine fever virus isolate from Malawi.
J. Gen. Virol.
69:1683-1694[Abstract/Free Full Text].
|
| 9.
|
Dixon, L. K.,
D. L. Rock, and E. Vinuela.
1995.
African swine fever-like viruses.
Arch. Virol.
10:92-94.
|
| 10.
|
Endris, R. G.,
T. M. Haslett, and W. R. Hess.
1991.
Experimental transmission of African swine fever virus by the tick Ornithodoros (Alectorobius) puertoricensis (Acari: Argasidae).
J. Med. Entomol.
28:854-858[Medline].
|
| 11.
|
Endris, R. G.,
T. M. Haslett, and W. R. Hess.
1992.
African swine fever virus infection in the soft tick, Ornithodoros (Alectorobius) puertoricensis (Acari: Argasidae).
J. Med. Entomol.
29:990-994[Medline].
|
| 12.
|
Endris, R. G.,
W. R. Hess, and J. M. Caiado.
1992.
African swine fever virus infection in the Iberian soft tick, Ornithodoros (Pavlovskyella) marocanus (Acari: Argasidae).
J. Med. Entomol.
29:874-878[Medline].
|
| 13.
|
Endris, R. G., and W. R. Hess.
1992.
Experimental transmission of African swine fever virus by the soft tick, Ornithodoros (Pavlovskyella) marocanus (Acari: Ixodoidea: Argasidae).
J. Med. Entomol.
29:652-656[Medline].
|
| 14.
|
Genovesi, E. V.,
F. Villinger,
D. J. Gerstner,
T. C. Whyard, and R. C. Knudson.
1990.
Effect of macrophage-specific colony-stimulating factor (CSF-1) on swine monocyte/macrophage susceptibility to in vitro infection by African swine fever virus.
Vet. Microbiol.
25:153-176[Medline].
|
| 15.
|
Greig, A.
1972.
The localization of African swine fever virus in the tick Ornithodoros moubata porcinus.
Archiv. Gesamte Virusforsch.
39:240-247.
|
| 16.
|
Groocock, C. M.,
W. R. Hess, and W. J. Gladney.
1980.
Experimental transmission of African swine fever virus by Ornithodoros coriaceus, an argasid tick indigenous to the United States.
Am. J. Vet. Res.
41:591-594[Medline].
|
| 17.
|
Hardy, J. L.,
E. J. Houk,
L. D. Kramer, and W. C. Reeves.
1983.
Intrinsic factors affecting vector competence of mosquitos for arboviruses.
Annu. Rev. Entomol.
28:229-262[Medline].
|
| 18.
|
Haresnape, J. M.,
P. J. Wilkinson, and P. S. Mellor.
1988.
Isolation of African swine fever virus from ticks of the Ornithodoros moubata complex (Ixodoidea: Argasidae) collected within the African swine fever enzootic area of Malawi.
Epidemiol. Infect.
101:173-185[Medline].
|
| 19.
|
Hess, W. R.,
R. G. Endris,
T. M. Haslett,
M. J. Monahan, and J. P. McCoy.
1987.
Potential arthropod vectors of African swine fever virus in North America and the Caribbean basin.
Vet. Parasitol.
26:145-155[Medline].
|
| 20.
|
Hess, W. R.,
R. G. Endris,
A. Lousa, and J. M. Caiado.
1989.
Clearance of African swine fever virus from infected tick (Acari) colonies.
J. Med. Entomol.
26:314-317[Medline].
|
| 21.
|
Heuschele, W. P., and L. Coggins.
1969.
Epizootiology of African swine fever in warthogs.
Bull. Epizoot. Dis. Afr.
17:179-183[Medline].
|
| 22.
|
Kleiboeker, S. B.,
T. G. Burrage,
G. A. Scoles,
D. Fish, and D. L. Rock.
1997.
African swine fever virus infection in the argasid host, Ornithodoros porcinus porcinus.
J. Virol.
72:1711-1724[Abstract/Free Full Text].
|
| 23.
|
Kleiboeker, S. B.,
G. F. Kutish,
J. G. Neilan,
Z. Lu,
L. Zsak, and D. L. Rock.
1998.
A conserved African swine fever virus right variable region gene, l11l, is nonessential for growth in vitro and virulence in domestic swine.
J. Gen. Virol.
79:1189-1195[Abstract].
|
| 24.
| Kleiboeker, S. B., G. A. Scoles, and D. L. Rock. Unpublished data.
|
| 25.
|
Mango, C. K. A., and R. Galun.
1977.
Ornithodoros moubata: breeding in vitro.
Exp. Parasitol.
42:282-288[Medline].
|
| 26.
|
Mebus, C. A.
1988.
African swine fever.
Adv. Virus Res.
35:251-269[Medline].
|
| 27.
|
Mellor, P. S., and P. J. Wilkinson.
1985.
Experimental transmission of African swine fever virus by Ornithodoros savignyi (Audouin).
Res. Vet. Sci.
39:353-356[Medline].
|
| 28.
|
Montgomery, R. E.
1921.
On a form of swine fever occurring in British East Africa (Kenya Colony).
J. Comp. Pathol.
34:159-191.
|
| 29.
|
Moore, D. M.,
L. Zsak,
J. G. Neilan,
Z. Lu, and D. L. Rock.
1998.
The African swine fever virus tymidine kinase gene is required for efficient replication in swine macrophages and for virulence in swine.
J. Virol.
72:10310-10315[Abstract/Free Full Text].
|
| 30.
|
Neilan, J. G.,
Z. Lu,
G. F. Kutish,
L. Zsak,
T. G. Burrage,
M. V. Borca,
C. Carrillo, and D. L. Rock.
1997.
A BIR motif containing gene of African swine fever virus, 4CL, is nonessential for growth in vitro and viral virulence.
Virology
230:252-264[Medline].
|
| 31.
|
Plowright, W.,
J. Parker, and M. A. Pierce.
1969.
African swine fever virus in ticks (Ornithodoros moubata, Murray) collected from animal burrows in Tanzania.
Nature
221:1071-1073[Medline].
|
| 32.
|
Plowright, W.,
J. Parker, and M. A. Pierce.
1969.
The epizootiology of African swine fever in Africa.
Vet. Rec.
85:668-674[Medline].
|
| 33.
|
Plowright, W.,
C. T. Perry,
M. A. Pierce, and J. Parker.
1970.
Experimental infection of the argasid tick, Ornithodoros moubata porcinus, with African swine fever virus.
Archiv. Gesamte Virusforsch.
31:33-50.
|
| 34.
|
Plowright, W.,
G. R. Thomson, and J. A. Neser.
1994.
African swine fever, p. 568-599.
In
J. A. W. Coetzer, G. R. Thomson, and R. C. Tustin (ed.), Infectious diseases of livestock with special reference to southern Africa. Oxford University Press, Cape Town, South Africa.
|
| 35.
|
Reed, L. J., and H. Muench.
1938.
A simple method of estimating fifty per cent endpoints.
Am. J. Hyg.
27:493-497.
|
| 36.
| Scoles, G. A., S. B. Kleiboeker, and D. L. Rock. Unpublished data.
|
| 37.
|
Sur, J.-H.,
V. L. Cooper,
J. A. Galeota,
R. A. Hesse,
A. R. Doster, and F. A. Osorio.
1996.
In vivo detection of porcine reproductive and respiratory syndrome virus RNA by in situ hybridization at different times postinfection.
J. Clin. Microbiol.
34:2280-2286[Abstract].
|
| 38.
|
Thomson, G. R.
1985.
The epidemiology of African swine fever: the role of free-living hosts in Africa.
Onderstepoort J. Vet. Res.
52:201-209[Medline].
|
| 39.
|
Walton, G. A.
1979.
A taxonomic review of the Ornithodoros moubata (Murray) 1877 (sensu Walton, 1962) species group in Africa.
Rec. Adv. Acarol.
11:491-500.
|
| 40.
|
Weaver, S. C.,
T. W. Scott,
L. H. Lorenz,
K. Lerdthusnee, and W. S. Romoser.
1988.
Togavirus-associated pathologic changes in the midgut of a natural mosquito vector.
J. Virol.
62:2083-2090[Abstract/Free Full Text].
|
| 41.
|
Weaver, S. C.,
T. W. Scott,
L. H. Lorenz, and P. M. Repik.
1991.
Detection of eastern equine enchephalomyelitis virus deposition in Culiseta melanura following ingestion of radiolabeled virus in blood meals.
Am. J. Trop. Med. Hyg.
44:250-259.
|
| 42.
|
Weaver, S. C.,
L. H. Lorenz, and T. W. Scott.
1992.
Pathologic changes in the midgut of Culex tarsalis following infection with western equine encephalomyelitis virus.
Am. J. Trop. Med. Hyg.
47:691-701.
|
| 43.
|
Zsak, L.,
Z. Lu,
G. F. Kutish,
J. G. Neilan, and D. L. Rock.
1996.
An African swine fever virus virulence-associated gene NL-S with similarity to the herpes simplex virus ICP34.5 gene.
J. Virol.
70:8865-8871[Abstract].
|
| 44.
|
Zsak, L.,
E. Caler,
Z. Lu,
G. F. Kutish,
J. G. Neilan, and D. L. Rock.
1997.
A nonessential African swine fever virus gene UK is a significant viral virulence determinant in domestic swine.
J. Virol.
72:1028-1035[Abstract/Free Full Text].
|
Journal of Virology, October 1999, p. 8587-8598, Vol. 73, No. 10
0022-538X/99/$04.00+0
This article has been cited by other articles:
-
Basto, A. P., Nix, R. J., Boinas, F., Mendes, S., Silva, M. J., Cartaxeiro, C., Portugal, R. S., Leitao, A., Dixon, L. K., Martins, C.
(2006). Kinetics of African swine fever virus infection in Ornithodoros erraticus ticks. J. Gen. Virol.
87: 1863-1871
[Abstract]
[Full Text]
Top
Abstract
Introduction
