Journal of General Virology (2007), 88, 670–679
DOI 10.1099/vir.0.82434-0
Differential infection efficiencies of peripheral lung
and tracheal tissues in sheep infected with Visna/
maedi virus via the respiratory tract
Tom N. McNeilly,13 Peter Tennant,1 Lluı́s Luján,2 Marta Pérez2
and Gordon D. Harkiss1
Correspondence
Tom N. McNeilly
[email protected]
Received 31 July 2006
Accepted 30 October 2006
1
Royal (Dick) School of Veterinary Studies, University of Edinburgh, Easter Bush Veterinary
Centre, Easter Bush, Midlothian EH25 9RG, UK
2
Dipartamento de Patologı́a Animal, Universidad de Zaragoza, Facultad de Veterinaria, Miguel
Servet 177, 50013 Zaragoza, Spain
The main routes of transmission of Visna/maedi virus (VMV), an ovine lentivirus, are thought to be
through ingestion of infected colostrum and/or milk or through inhalation of respiratory secretions.
Whereas oral transmission appears to be mediated via epithelial cells within the small intestine,
the mechanism of virus uptake in the respiratory tract is unknown. In addition, it is not known whether
infection is mediated by cell-associated or cell-free VMV, previous studies having not addressed
this question. Intratracheal (i.t.) injection of VMV is known to be a highly efficient method of
experimental infection, requiring as little as 101 TCID50 VMV for successful infection. However,
using a tracheal organ culture system, we show here that ovine tracheal mucosa is relatively
resistant to VMV, with detectable infection only seen after incubation with high titres of virus (¢105
TCID50 ml”1). We also demonstrate that i.t. injection results in exposure of both trachea and the
lower lung and that the time taken for viraemia and seroconversion to occur after lower lung
instillation of VMV was significantly shorter than that observed for tracheal instillation of an identical
titre of virus (P=0.030). This indicates that lower lung and not the trachea is a highly efficient site for
VMV entry in vivo. Furthermore, cell-free virus was identified within the lung-lining fluid of naturally
infected sheep for the first time. Together, these results suggest that respiratory transmission of
VMV is mediated by inhalation of aerosols containing free VMV, with subsequent virus uptake in the
lower lung.
INTRODUCTION
Visna/maedi virus (VMV) is a member of the lentivirus
subgroup of retroviruses that causes interstitial pneumonia,
encephalitis, arthritis and mastitis in sheep (Sigurdsson et al.,
1952; Sigurdsson & Palsson, 1958). The main routes of
transmission of VMV are thought to be through ingestion of
infected colostrum and/or milk or through inhalation
of respiratory secretions (Pepin et al., 1998; Blacklaws et al.,
2004). Whereas oral transmission appears to be mediated
via uptake of virus by small-intestinal epithelial cells
(Preziuso et al., 2004), the mechanism of virus uptake in
the respiratory tract is as yet unknown.
Recent studies have identified intratracheal (i.t.) inoculation
as a highly efficient route of experimental infection. In one
study, the minimum infectious dose of VMV for the i.t.
3Present address: Moredun Research Institute, Pentlands Science
Park, Bush Loan, Penicuik EH26 0PZ, UK.
Primer details are available as supplementary material in JGV Online.
670
route was found to be 101 TCID50 delivered in a 1 ml
volume, whereas that for the intranasal route was 56106
TCID50 (Torsteinsdottir et al., 2003). This suggests that the
lower respiratory tract and not the upper respiratory tract
(nasal cavity and nasopharynx) is the major site of VMV
entry during respiratory transmission. It is unclear exactly
which parts of the respiratory tract are exposed by i.t.
instillates, although certainly trachea and possibly lower
lung (bronchioles and alveolar areas) may be involved.
Comparisons between trachea and lower lung for infection
efficiency have not been performed. Therefore, it is unclear
whether the high sensitivity of the tracheal route for
infection represents a unique feature of the tracheal mucosa
or is due to exposure of lower lung, or both.
In addition to the lack of knowledge regarding the site of
VMV uptake during respiratory transmission, the precise
nature of the infectious agent is unclear. It has long been
presumed that respiratory transmission of VMV is mediated
by inhalation of free virus or cell-associated virus, most
likely VMV-infected alveolar macrophages (AMs) (Pepin
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Visna/maedi virus infection of ovine respiratory tract
et al., 1998; Blacklaws et al., 2004; Peterhans et al., 2004).
However, whereas productively infected AMs have been
identified in a number of studies of naturally infected sheep
(Lujan et al., 1994; Brodie et al., 1995; Gelmetti et al., 2000),
to date cell-free VMV has not been identified conclusively
within the lung-lining fluid of infected individuals. The
nature of the infectious agent may have important implications regarding the sites of initial virus uptake, as cellassociated VMV would be preferentially deposited within the
upper respiratory tract, whereas only smaller particles of
<5 mm diameter, such as cell-free VMV particles, which have
a diameter of approximately 100 nm (Lee et al., 1996), are
likely to reach the lower lung (Zhang et al., 2000; Gordon &
Read, 2002). Therefore, identification of free VMV would
appear to be crucial for lower lung involvement to occur
during natural respiratory transmission.
The aims of this study were twofold: firstly to identify the
sites of the respiratory tract that are responsible for virus
uptake during i.t. inoculation, and secondly to determine
whether cell-free VMV occurred within the lung-lining fluid
in naturally infected sheep. To assess the contribution of the
trachea to initial virus uptake, a tracheal organ culture
system was developed and subsequently used to determine
the sensitivity of the tracheal mucosa to VMV ex vivo. In
addition, patent blue, an inert protein-binding dye routinely
employed for in vivo tracking studies (Hirsch et al., 1982),
was used to identify areas of the respiratory tract exposed
during i.t. inoculation, and the relative sensitivities of areas
of the lung involved during i.t. inoculation were determined
using a novel in vivo infection model. Finally, lung-lining
fluid samples obtained from three naturally infected sheep
were analysed for the presence of cell-free VMV. The results
of this study will potentially identify sites of virus entry
during natural respiratory transmission of VMV and clarify
the nature of the infectious agent.
METHODS
Virus propagation and titration. Low-passage VMV strain EV1
(Sargan et al., 1991) was propagated routinely on ovine skin cells
(OSCs) in DMEM supplemented with 10 % FCS. Virus-containing
culture medium was clarified by centrifugation for 10 min at 400 g
and subsequently filtered through a 0.2 mm syringe-driven filter
(Nalgene).
Virus titres were determined as described previously using OSCs as
indicator cells (Ebrahimi et al., 2000). Titres were expressed as either
tissue culture infectious dose (TCID50) for samples containing ¢20
TCID50 ml21 or the percentage of wells with visible cytopathic
effect (CPE) after incubation with neat sample for samples <20
TCID50 ml21.
Ovine tracheal organ culture and infection with VMV. The
procedure was based on that of Campbell et al. (1979) and Lin et al.
(2001). Proximal trachea was removed aseptically from VMV-seronegative sheep immediately post-mortem and placed in cold PBS
containing (ml21) 100 U penicillin, 100 mg streptomycin, 4 mg
amphotericin B and 10 mg gentamicin (explant-PBS). After washing
three times in explant-PBS for 5 min each, tracheal tissue was cut
into 6 mm diameter discs using a sterile punch biopsy needle
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(Kruuse A/S). Tracheal discs were placed with epithelium surface
upward in 6-well plastic tissue culture dishes (Nunc) and allowed to
stick down at room temperature for 5 min prior to addition of 5 ml
DMEM supplemented with (ml21) 100 U penicillin, 100 mg streptomycin, 4 mg amphotericin B, 10 mg gentamicin, 100 U insulin and
2 % Ultroser-G (Ciphergen) (explant medium). Organ cultures were
subsequently incubated at 37 uC with 5 % CO2 in a humidified
atmosphere. After 4 h, the medium was removed and each organ
culture was incubated in 5 ml explant medium containing varying
concentrations of VMV (102 to 106 TCID50 ml21) for 2 h. Organ
cultures were then washed five times in PBS and subsequently incubated in fresh explant medium for 7 days. Negative controls
included harvesting organ cultures after the final PBS wash (day 0)
and incubation with no virus (mock infection). Each dilution of
virus was performed in duplicate for each source animal, and the
experiment was repeated using tracheas obtained from six animals.
After 7 days, organ cultures were halved. DNA was extracted from one
half using a DNeasy tissue kit (Qiagen) and samples were stored at
280 uC prior to PCR analysis for VMV provirus. The other half was
snap-frozen in isopentane/dry ice. Tissue sections were cut to 6 mm
thick, mounted on poly-L-lysine-coated slides (BDH) and stored at
280 uC prior to immunocytochemical (ICC) analysis for VMV capsid
protein. Virus titrations were performed on samples of wash fluid from
the last washing step after virus incubation and day 7 culture
supernatants.
To assess organ culture viability, the strength of epithelial cilial beating
in six preliminary organ cultures was assessed by direct visualization
using a Fluovert inverted light microscope (Leica) and scored
subjectively as either absent, weak, moderate or vigorous. In addition,
day 3 (n=6) and day 7 (n=6) organ cultures were fixed in 10 %
buffered formalin, paraffin embedded, sectioned and stained with
haematoxylin and eosin (H&E) to assess tissue morphology.
Animals. For dye tracking studies, four adult greyface ewes were
used. For VMV infection studies, 10 adult Suffolk-cross ewes were
used. All sheep were commercially sourced. Prior to VMV infection
studies, sheep were determined to be free from VMV provirus and
seronegative for VMV-specific antibodies. All experimental procedures involving animals were approved by The University of
Edinburgh’s Biological Services Ethical Review Committee and were
performed under licence as required by the UK Animals (Scientific
procedures) Act 1986.
I.t. inoculation. Sheep were sedated by intramuscular injection of
xylazine hydrochloride (Rompun; Bayer) at a dose rate of approximately 0.5 mg kg21. One millilitre volumes of patent blue dye (0.1 %
w/v in PBS) or cell-free virus (106 TCID50) were injected into the
proximal third of the trachea via a 21 gauge needle, avoiding the tracheal cartilage. The neck was held in dorsiflexion for 1–2 min postinstillation. For patent blue dye tracking studies, sheep were allowed
to recover in a pen with free access to food and water for 1 h postinstillation prior to euthanasia by intravenous injection of pentobarbitone (Euthatal; Rhône Mérieux) at a dose rate of 100 mg kg21.
Lungs were removed and the location of dye was recorded.
Differential exposure of trachea and peripheral lung. Delivery
of inoculates to either trachea or lower lung was performed endoscopically under general anaesthesia. Induction of anaesthesia was
achieved by intravenous administration of thiopentone sodium
(Rhône Mérieux) at a dose rate of 20 mg (kg body weight)21. Sheep
were intubated and anaesthesia was maintained with gaseous
halothane (2–3 %) in oxygen and nitrous oxide under negative pressure ventilation.
For initial development of the differential exposure model, 1 ml
volumes of patent blue dye (0.1 % w/v in PBS) were instilled onto the
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T. N. McNeilly and others
lateral aspect of the distal one-third of the trachea of two adult greyface
ewes via a 21 gauge polyethylene catheter inserted into the lateral
channel of a flexible fibre-optic bronchoscope (5.3 mm o.d.) (model
FG-16X; Pentax). The position of the instillate was visualized at 15, 30,
45 and 60 min post-instillation. The sheep were then allowed to
recover in a pen with free access to food and water and were standing
within 15–20 min post-anaesthesia. After 1 h of recovery, sheep were
euthanized, the lungs were removed and the location of patent blue dye
was recorded.
For subsequent virus exposure experiments, 1 ml volumes containing
106 TCID50 cell-free VMV and 0.01 % patent blue dye (which had
previously been shown not to affect virus titre) were instilled into either
the distal trachea or the left cardiac lung lobe. Instillates were visualized
for 1 h post-instillation as described above and sheep were
subsequently allowed to recover. Sheep were bled weekly for the first
2 months and thereafter every 2 weeks until 6 months postinoculation. Blood samples were analysed for the presence of VMV
provirus and VMV-specific antibodies by PCR and ELISA, respectively,
and the times to PCR positivity and seroconversion were recorded.
Isolation of peripheral blood mononuclear cells (PBMCs).
Blood was collected into an EDTA-containing Vacutainer (Becton
Dickenson) and PBMCs were isolated using a Ficoll-paque density
gradient (GE Healthcare) according to the manufacturer’s instructions. Cells were pelleted by centrifugation at 100 g for 10 min and
the resultant cell pellet was resuspended in 200 ml PBS. DNA was
subsequently extracted using the DNeasy mini kit (Qiagen).
Detection of VMV provirus. VMV provirus was detected by semi-
nested PCR (snPCR) using the first-round primer pair gag 1 and gag
5 and second-round primer pair gag 1 and gag 2 (primer sequences
and positions within the VMV genome are shown in Supplementary
Table S1 available in JGV Online). Reactions of 50 ml final volume
contained 1.5 mM MgCl2, 0.2 mM dNTPs, 25 pmol each of sense
and antisense primer and 2.5 U HotTaq (Biogene). For organ culture experiments, 100 ng template DNA was added to first-round
reactions. For in vivo infection studies, 500 ng PBMC DNA was
added to first-round reactions. A 3 ml aliquot from the first round
of PCR was used in the second-round reaction.
For the first round of PCR, the thermal cycling profile involved a
10 min pre-incubation at 95 uC followed by 15 cycles each consisting of
denaturation at 94 uC for 1 min, primer annealing at 55 uC for 1 min
and extension at 72 uC for 2 min and a final extension at 72 uC for
5 min. For reamplification, the number of cycles was increased to 35
and denaturation, annealing and extension times were 30 s, 30 s and
1 min, respectively.
Detection of VMV-specific serum antibodies. Serum samples
were tested for the presence of virus-specific antibodies to the major
core protein p25 of VMV and/or the viral transmembrane protein
gp46 using a commercial ELISA kit (ELITEST-MVV/CAEV; Hyphen
BioMed) (Saman et al., 1999). Absorbance values were measured
using a Bio-Tek Microplate Autoreader (Bio-Tek Instruments).
Bronchoalveolar lavage (BAL) collection and processing. The
lungs from three VMV-seropositive adult Rasa Aragonesa sheep
with diffuse, severe VMV lesions were subjected to BAL as follows: a
luer-tipped 50 ml syringe was wedged into selected segmental
bronchi exhibiting gross VMV lesions and a single 40 ml aliquot of
normal saline was used to lavage each lung segment. BAL samples
were passed through sterile gauze into sterile 50 ml Falcon tubes on
ice and subsequently centrifuged at 400 g for 7 min at 4 uC to separate out the cellular fraction. BAL supernatants were then passed
immediately through a 0.2 mm syringe-driven filter (Nalgene) and
672
stored at 280 uC prior to virus titration. Cytocentrifuge slides were
prepared from the resultant cell pellets using a Cytospin 3 cytocentrifuge (Thermo-Shandon).
ICC staining for VMV capsid protein p25. Slides were fixed in
100 % methanol for 10 min at 220 uC and then incubated in 0.3 %
hydrogen peroxide in PBS/0.5 % Tween 80 (PBS/T80) for 20 min at
room temperature. After washing in PBS, slides were incubated in
10 % normal goat serum (NGS) in PBS/T80 for 1 h at room temperature prior to incubation with the monoclonal antibody VPM70
(anti-VMV p25, mouse IgG1) (Reyburn et al., 1992) diluted 1 : 2 in
PBS/T80 containing 10 % NGS for 1 h at room temperature.
Negative controls were provided by replacing the primary antibody
with an identical dilution of isotype-matched control monoclonal
antibody VPM53 (McOrist et al., 1989). After washing, secondary
antibody (peroxidase-labelled polymer conjugated to goat antimouse immunoglobulins, EnVision Plus HRP system; Dako) was
applied to sections for 30 min at room temperature. Antibody binding was detected by incubation with 3,39-diaminobenzidine for
7.5 min at room temperature. Sections were counterstained with
haematoxylin, dehydrated in graded alcohols, cleared in xylene and
mounted in DPX mounting medium (Fischer Scientific). For cytocentrifuge preparations, 500 cells were examined and the number of
VMV-positive cells was recorded.
Statistical analysis. Non-parametric statistical analysis was carried
out using the Mann–Whitney procedure on Minitab version 14
for Microsoft Windows. Values of P<0.05 were determined to be
significant.
RESULTS
Assessment of tracheal organ culture viability
To evaluate the tracheal organ culture methodology used in
this study, assessment of epithelial ciliary activity and organ
culture morphology was performed. Cilial beating was
present in all organ cultures assessed throughout the 7 day
culture period and was either weak or moderate.
Histopathological evaluation of organ cultures revealed
preservation of a pseudostratified ciliated epithelium in
both day 3 and day 7 organ cultures (data not shown). Some
degenerative changes were observed in the submucosa at day
3 and day 7, including loss of structural integrity of the
connective tissue layers, sloughing and karryorrhexis of the
vascular endothelium and submucosal glandular epithelium
and degeneration of the submucosal cell nuclei.
Degenerative changes were more apparent in day 7 cultures.
Infection of ovine tracheal organ cultures with
VMV
To determine whether tracheal organ cultures were
infectable by VMV and the minimum infectious dose of
virus required, tracheal organ cultures were incubated with
varying titres of VMV strain EV1 and subjected to snPCR
analysis for VMV provirus. A summary of the snPCR results
is shown in Fig. 1(a). VMV gag proviral DNA was detected
in 100 % of explants incubated with 106 TCID50 VMV ml21
and in 66.6 % of explants incubated with 105 TCID50 VMV
ml21. No proviral DNA was detected in any other organ
culture incubated with virus. Day 0 and mock-infected
explants were negative. A representative image of an agarose
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Journal of General Virology 88
Visna/maedi virus infection of ovine respiratory tract
cultures but not in the 2 h cultures (data not shown),
indicating de novo provirus production and, subsequently,
that 7-day-old organ cultures were still metabolically active.
To assess productive virus replication in tracheal organ
cultures, virus titration of organ culture supernatants and
ICC for VMV capsid protein was also performed. ICC failed
to detect VMV capsid protein in all organ cultures. A
positive control sample consisting of in vitro-infected OSCs
was run in parallel with all samples and was consistently
positive. Positive virus titres were found in both last wash
(day 0) and day 7 supernatants of organ cultures incubated
with 106 TCID50 VMV ml21. However, no significant
difference in titre was seen between day 0 and day 7 time
points (Fig. 1c), suggesting that virus detected at day 7 was
most likely to be carry-over from input virus. All other
supernatants tested were negative.
Tracking of i.t. inoculates
To evaluate the lung distribution of fluid introduced via i.t.
inoculation, patent blue dye was instilled into the tracheas of
two adult greyface ewes by i.t. injection. Representative
images of the distribution of dye 1 h post-inoculation are
shown in Fig. 2. The majority of dye was located in the
trachea and left cardiac lung lobes of both sheep. In one
sheep, a small amount of dye was also present in the right
cardiac lung lobe. Dye was located throughout the whole
length of the trachea, including the region proximal to the
injection site. The dye in the trachea was primarily located
within the mucus layer overlying the tracheal mucosa. In
lung lobes, dye staining exhibited a patchy distribution and
was present in both smaller airways and lung parenchyma.
Fig. 1. Detection of VMV in ovine tracheal organ cultures after
in vitro infection. Organ cultures were incubated in duplicate
with 102 to 106 TCID50 VMV ml”1 for 2 h. After washing, cultures were incubated for 7 days before performing snPCR to
detect VMV proviral DNA. Virus titrations of last wash (day 0)
and day 7 supernatants (day 7) were performed to assess virus
release from organ cultures. (a) Summary of PCR results from
duplicate organ cultures from six source sheep. Day 0 represents incubation with 106 TCID50 VMV ml”1 and harvesting
after washing step. (b) Example of an agarose gel of PCR products generated from organ cultures derived from a single
sheep. (c) Summary of virus titrations. Titres are expressed as
the mean percentage of wells with visible CPE after incubation
with neat sample ±SEM (n=12).
gel of PCR products generated from organ cultures derived
from one source sheep is shown in Fig. 1(b).
To assess the metabolic activity of organ cultures at 7 days,
7-day-old organ cultures were infected with 106 TCID50
VMV ml21 as described above and provirus production was
assessed by snPCR at 2 h (n=4) or 24 h (n=4) postinfection. VMV provirus was detected in all 24 h organ
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Development of a differential lung exposure
model
To assess the relative sensitivities of trachea and lower lung
to VMV, a differential in vivo exposure model was created to
allow exclusive exposure of the trachea to 1 ml instillations
without involving the peripheral lung. One millilitre of
patent blue dye was introduced endoscopically onto the
lateral aspect of the distal one-third of the trachea of two
adult ewes and the location was recorded up to 2 h postinstillation. Representative images from one tracheal
exposure experiment are shown in Fig. 3. Dye pooled in
the ventral aspect of the distal one-third of the trachea just
proximal to the entrance to the right apical lung lobe
immediately post-instillation. During the 1 h of anaesthesia,
dye moved proximally up the trachea towards the larynx. By
45 min post-instillation, dye was present at the distal end of
the endotracheal tube. At post-mortem, dye was located at
the proximal end of the trachea and within the larynx. A
small amount of dye was present within the oral cavity. No
dye was located in lower trachea or within any lung lobe,
indicating good exposure of the trachea without any
detectable exposure of the peripheral lung.
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T. N. McNeilly and others
exposure group was traumatized with a 21 gauge needle to
account for tracheal trauma resulting from i.t. injection.
(a)
RC
Lower lung instillations remained within the left cardiac
lung lobe throughout anaesthesia. Tracheal instillations
pooled at the level of the distal trachea and moved
proximally up the trachea towards the larynx at 45 min
post-instillation. Two further sheep were inoculated with
106 TCID50 VMV strain EV1 to allow direct comparison
with tracheal and lower lung exposures. Weekly
blood samples were subsequently analysed for the presence
of viraemia and seroconversion by PCR and ELISA,
respectively.
LC
(b)
(c)
Weekly PCR results are summarized in Fig. 4(a). I.t.
injection resulted in viraemia at 3–4 weeks post-infection.
Lung instillation resulted in viraemia at 2–3 weeks postinfection. Tracheal instillation resulted in viraemia at
8–24 weeks post-instillation, significantly delayed compared with lung instillation (P=0.030). Results of ELISA
analysis are summarized in Fig. 4(b). Seroconversion
occurred at 4–6 weeks post-infection for i.t. injection,
2–8 weeks post-infection for lung instillation and
12–24 weeks post-infection for tracheal instillation. The
time to seroconversion was significantly greater for tracheal
instillation compared with lower lung instillation
(P=0.030). Traumatizing the tracheal epithelium prior to
tracheal instillation did not decrease the time taken for
viraemia or seroconversion to occur.
(d)
Identification of cell-free VMV within lung-lining
fluid of naturally infected sheep
Fig. 2. Distribution of patent blue dye after i.t. inoculation. (a)
Large amounts of dye were located in the left cardiac lung
lobe (LC). A small amount of dye was also present in the right
cardiac lung lobe (RC) of one sheep (arrowhead). Arrow indicates the injection site. (b) Dye was present throughout the
whole length of the trachea within the overlying mucus layer.
(c) Dye was present throughout the airways and parenchyma of
the left cardiac lung lobe. (d) Tissue section of the left cardiac
lung lobe demonstrating dye staining in alveolar areas. Bar,
50 mm.
Comparison between tracheal and peripheral
lung exposure to VMV
To compare the relative sensitivities of trachea and lower
lung to VMV, seronegative adult sheep were inoculated
under general anaesthesia with 1 ml VMV strain EV1
containing 106 TCID50 either directly into the left cardiac
lung lobe (n=4) or into the trachea using the tracheal
exposure model developed previously (n=4). Virus
inoculates were spiked with 0.01 % patent blue dye to
allow direct visualization of virus instillations by endoscopy.
In addition, the trachea from one sheep from the tracheal
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Cellular and cell-free fractions of post-mortem BAL samples
obtained from three naturally infected sheep were analysed
for the presence of VMV using a combination of virus
titration and ICC. Within bronchoalveolar cell (BAC)
populations, VMV-positive cells were identified with typical
AM morphology (Fig. 5a). Virus was detected in all cell-free
BAL supernatant samples, with co-cultivated OSCs exhibiting syncytial formation typical of VMV CPE within 7 days
(Fig. 5b). The presence of VMV within these multinucleate
cells was confirmed by ICC staining for VMV capsid protein
(Fig. 5c, d), which revealed widespread staining throughout
the syncytia.
The results of BAL supernatant virus titrations and VMVpositive cell counts are shown in Table 1. Virus titres within
BAL supernatants ranged from 1.16102 to 1.46103 TCID50
[ml BAL fluid (BALF)]21. Assuming the BAL procedure
resulted in a dilution factor of approximately 1 : 100
(McGorum et al., 1993), these titres equate to 1.16104 to
1.46105 TCID50 (ml lung-lining fluid)21. VMV capsid
protein was detected in 1.4–4.4 % of BACs. There was
no direct correlation between the percentage of positive
BACs and virus titre in the BALF, although the lowest virus
titre was associated with the lowest percentage of positive
BACs.
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Visna/maedi virus infection of ovine respiratory tract
(a)
(b)
(c)
C
RA
(d)
ET
(e)
(f)
(g)
RMB
LMB
RA
DISCUSSION
Tracheal organ cultures have been used in many ex vivo
experimental models to examine the initial interactions
between the host and various viruses and bacteria (Campbell
et al., 1979; Dhinakar Raj & Jones, 1996; Rutman et al., 1998;
Lund et al., 2001; Lin et al., 2001; Anderton et al., 2004).
Organ cultures offer the advantage that both epithelial and
interstitial cells are present in their normal anatomical
arrangement, allowing realistic interactions between the
cells in each compartment. In the case of in vitro infection
with VMV, most cell types isolated and grown in primary
cell culture have been shown to support productive VMV
replication, unlike the situation in vivo, in which virus
replication is highly restricted (Haase et al., 1977;
Gendelman et al., 1985; Brodie et al., 1995). This loss of
the normal restricted replication pattern as a result of in vitro
culturing is a significant problem when trying to study
normal in vivo VMV behaviour and, to date, in vitro studies
of VMV infection have concentrated on primary cell
cultures (Leroux et al., 1995; Craig et al., 1997; Lerondelle
et al., 1999). Using an organ culture system, it was hoped
that the observed VMV replication pattern more truly
represented the situation in vivo.
In this study, it was shown that ovine tracheal organ cultures
were capable of supporting VMV replication up to the level
of proviral synthesis. No evidence of productive virus
infection was found, with no release of virions into the
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Fig. 3. Representative images from one tracheal exposure experiment. One millilitre
patent blue dye was deposited into the
distal trachea under general anaesthesia via
an endoscope. After 1 h of anaesthesia and
1 h of recovery, sheep were euthanized. (a)
Deposition of dye onto the lateral aspect of
the distal tracheal wall via a catheter (C)
inserted into the lateral channel of an endoscope. (b) Dye pooled immediately in the
ventral aspect of the distal trachea, just
proximal to the entrance to the right apical
lung lobe (RA). (c) At 45 min post-instillation, dye was present at the distal end of
the endotracheal tube (ET). (d–g) At postmortem, no dye was visible in any lung lobe
(d) and was located at the proximal end of
the trachea (arrow) and within the larynx (e).
No dye was present at the entrances to the
right main bronchus (RMB), left main
bronchus (LMB) or right apical lung lobe
(RA) (f). Sectioning of the left cardiac lung
lobe revealed the lobe to be free of dye (g).
culture medium and no viral protein detected using ICC.
The permissive life cycle of the virus in vitro is usually
complete within 3 days (Brahic et al., 1981; Haase et al.,
1982; Vigne et al., 1987), and maintenance of organ cultures
for 7 days post-infection prior to analysis was thought to
allow sufficient time for productive VMV infection to take
place. This suggests that VMV replication in tracheal organ
cultures is restricted at some level between proviral synthesis
and translation of structural viral proteins, thus mimicking a
typical in vivo VMV replication pattern (Gendelman et al.,
1985; Brodie et al., 1995). This is of interest, since organ
cultures are without an adaptive immune system, implying
that the restricted replication observed is not due to specific
anti-VMV immune responses. It is possible that infected
cells are not in the correct state of maturation or activation,
as productive VMV replication is known to be closely linked
to the maturation/activation state of the host cell
(Gendelman et al., 1986). The restricted replication
observed could result from a suboptimal environment for
virus replication due to reduced cell viability. However, 7day-old cultures were still capable of de novo provirus
production and therefore were still metabolically active.
Detectable infection was seen only in organ cultures
incubated with high titres of virus (¢105 TCID50 ml21).
The 2 h contact time between virus and trachea in these
studies was long relative to the rate of binding of VMV to its
cellular receptor(s), which has been estimated to occur
within 15 min in vitro (Kennedy-Stoskopf & Narayan,
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T. N. McNeilly and others
lower lung exposure, and not tracheal exposure, is the
reason for the high efficiency of infection observed for i.t.
inoculation with VMV.
The relative inefficiency of VMV uptake by the trachea in
vivo may be explained firstly by an inherent insensitivity of
the tracheal mucosal cells to VMV, as suggested by the
results of tracheal organ culture experiments performed in
this study, and secondly by the action of innate mucosal
defence mechanisms. For example, tracheal virus inoculations were transported efficiently from the distal trachea to
the larynx by the mucociliary escalator within 45 min,
resulting in only limited contact time between VMV and the
tracheal mucosa. Given that infection of tracheal organ
cultures was inefficient despite 2 h continuous exposure to
VMV of identical titre and strain, it is likely that mucociliary
escalator function plays an additional role in limiting VMV
uptake in vivo.
Fig. 4. Comparison of the efficiencies of i.t., tracheal and lower
lung inoculation routes for VMV. Sheep were inoculated with
106 TCID50 VMV strain EV1 via i.t. injection (i.t.), endoscopic
instillation into trachea (Trachea) or endoscopic instillation into
the left cardiac lung lobe (Lung). The trachea of one sheep in
the tracheal instillation group (denoted by a large circle) was
traumatized using a 21 gauge needle prior to virus instillation.
The time taken for viraemia (detected by snPCR) (a) and seroconversion (detected by ELISA) (b) to occur was recorded for
each route. Horizontal lines indicate mean values. Asterisks
denote significance in the Mann–Whitney test.
1986). Taken together, these results indicate that uptake of
VMV by ovine tracheal organ cultures is inefficient.
Furthermore, as processing of tracheal organ cultures
prior to virus incubation is likely to compromise a
number of innate immune defence mechanisms severely,
including loss of mucus and antimicrobial peptides from the
epithelial surface and disruption of the mucociliary escalator
(Zhang et al., 2000; Welsh & Mason, 2001), these results
suggest an inherent insensitivity of tracheal cell populations
to VMV.
It has been shown that i.t. inoculation in sheep results in
exposure of both trachea and all parts of the lower lung to
the instillation. Using a differential exposure model for
lower lung and trachea, it was demonstrated that lower lung
instillation is a highly efficient route of infection with VMV
compared with tracheal exposure in vivo. The time taken for
viraemia and seroconversion to occur after lower lung
instillation was similar to that observed for i.t. inoculation.
In addition, trauma to the tracheal epithelium prior to
tracheal exposure to virus did not appear to increase the
efficiency of virus infection. Therefore, it is concluded that
676
The reasons for the high efficiency of lower lung instillation
of cell-free VMV may be twofold. Firstly, there are abundant
populations of a number of potential target cell types for
VMV in the lower lung. These include respiratory tract
dendritic cells, AMs, interstitial macrophages and bronchial
epithelial cells, all of which have been implicated in VMV
infection in vivo (Gendelman et al., 1985; Staskus et al., 1991;
Brodie et al., 1995; Ryan et al., 2000; Carrozza et al., 2003).
As fluid introduced into lung lobes has been shown in this
study to come into contact with all levels of the respiratory
tract, including the alveolar air space, all of these cell types
may theoretically be exposed to virus during lung lobe
instillations. However, whether these cell types are involved
in initial virus uptake is unclear, as studies to date have
concentrated on analysis of lungs with pre-existing VMV
lesions rather than tracking initial virus uptake.
Secondly, pulmonary clearance of inhaled particles and
pathogens in the lower lung is primarily mediated via
phagocytic uptake by AMs (Lehnert, 1992; Gordon & Read,
2002). It can therefore be seen that innate defence
mechanisms of the lower lung may actually result in
enhanced uptake of VMV, as interaction between AMs, a
natural target cell, and free virus within the lower lung is
likely to result in virus infection of AMs rather than virus
inactivation. Thus, pulmonary clearance mechanisms in the
lower lung would increase rather than reduce virus uptake.
Indeed, the intracellular bacterium Legionella pneumophila
has been shown to utilize AM uptake to facilitate
colonization of the lung (Yamamoto et al., 1994).
Instillation of VMV into a lung lobe and subsequent
analysis of AM populations for evidence of infection at an
early (pre-lesional) time point would be required to confirm
initial uptake of VMV by AMs.
The high sensitivity of the lower lung to cell-free VMV
suggests that uptake of virus at this site may play a role in
natural respiratory transmission. As cell-free virus was
identified in the lung-lining fluid of naturally infected sheep
in this study, it is possible that coughing by VMV-infected
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Journal of General Virology 88
Visna/maedi virus infection of ovine respiratory tract
(a)
(b)
(c)
Fig. 5. Representative images from a naturally VMV-infected sheep demonstrating the
presence of VMV within BAL cells and cellfree BAL supernatant. (a) ICC staining of
BAL cells for VMV capsid protein. A VMVpositive cell with AM morphology is indicated by the arrow (bar, 20 mm). (b) Giemsa
stain of OSCs co-cultivated with cell-free
BAL supernatant at 7 days. Many multinucleate cells are present (arrowheads) (bar,
50 mm). (c–d) Cytocentrifuged multinucleate
OSCs co-cultivated with cell-free BAL
supernatant. (c) ICC staining for VMV
capsid protein demonstrating positive staining throughout two multinucleate cells
(arrows) (bar, 20 mm). (d) Isotype-matched
negative control. The arrow indicates a multinucleate cell free of staining (bar, 20 mm).
(d)
animals may generate free-virus-containing aerosols that are
small enough to reach the lower lung. Indeed, in a study of
cough-generated aerosols in humans, the majority of
aerosolized particles were found to be between 0.65 and
3.3 mm in diameter (Fennelly et al., 2004), which would be
small enough in theory to reach the lower regions of the
lung. Therefore, inhalation of aerosols containing cell-free
VMV into the lower lung may be an important mechanism
of respiratory transmission in the natural situation. In
addition, as aerosol transmission would require close
contact between infected and non-infected animals, this
may partly explain the recent observation that VMV
transmission appears to be minimal in extensively reared
flocks (Leginagoikoa et al., 2006).
In conclusion, this study has demonstrated that the tracheal
mucosa is relatively resistant to infection and that the high
Table 1. BAC immunohistochemistry for VMV and freevirus titrations of corresponding BALF from three naturally
infected sheep
BACs were stained for the presence of VMV capsid protein. Five
hundred cells were examined per sheep and the percentage of cells
positive for VMV was recorded. Virus titrations were performed
on cell free-BALF.
Sheep
efficiency of infection via the i.t. route results from lower
lung exposure. The identification of cell-free VMV within
the lung-lining fluid of naturally infected sheep, together
with the high sensitivity of the lower lung to VMV, suggests
that inhalation of aerosols containing free virus particles and
subsequent exposure of the lower lung may play a significant
role in natural respiratory transmission of VMV.
Identification of cell types within the lower lung that are
involved in initial virus uptake is required to elucidate the
exact mechanism of virus entry in the lower lung.
ACKNOWLEDGEMENTS
T. N. McN. was supported by a PhD studentship from the Royal (Dick)
College of Veterinary Studies, University of Edinburgh, and funding
from the European Union (contract no. QLK2-CT-2002-00167). We
would like to thank Alison Baker for technical assistance during the in
vivo infection studies, Joan Docherty for provision of ovine tracheal
tissue and Paul Wright for his excellent care of the experimental
animals during this study.
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