Cowpox

* Objectives.-To determine the morphologic changes and disease progression of aerosolized cowpox virus infection in BALB/c mice and to ascertain the suitability of cowpox virus-infected BALB/c mice as a model of aerosol-transmitted, orthopoxviral respiratory disease.

Methods.-BALB/c mice were inoculated with cowpox virus, Brighton strain, by aerosol or intranasal route. Mice were killed at specified times after inoculation, necropsied, and tissues were collected for routine histology, immunohistochemistry, and electron microscopy.

Results.-Inoculation by both routes resulted in disease and death. Immunolabeled viral antigen and lesions predominated in the tissues associated with the inoculation route, that is, lungs, airways, trachea, and nasal passages and sinuses. Tracheitis was evident in the intranasally infected group only. Lesions were generally necrotizing and

hemorrhagic, neutrophilic, and increased in extent and severity in a time-dependent fashion. Viral intracytoplasmic inclusion bodies, immunolabeled viral antigen, or virions were readily seen in epithelial tissues, smooth muscle cells of airways and vessels, fibroblasts, periosteal cells, perineural cells, and macrophages. Although the extension of infection appeared to be primarily direct, lesions suggesting hematogenous dissemination were occasionally noted in bone marrow and skin. Transmission electron microscopy demonstrated features of cell injury or death, virion assembly and maturation, and both A-type and B-type inclusions.

Conclusions.-Aerosol inoculation of BALB/c mice with cowpox virus provides a reliable and facilitative model of aerosol-transmitted, orthopoxviral respiratory disease. (Arch Pathol Lab Med. 2000;124:362-377)

The family Poxviridae is composed of large, doublestranded DNA viruses comprising 2 subfamilies, Chordopoxvirinae and Entomopoxvirinae, which replicate in the cytoplasm of vertebrate or insect cells, respectively. 1-3 Within the subfamily Chordopoxvirinae, poxviruses in the genera Orthopoxvirus, Parapoxvirus, Yatapoxvirus, and Molluscipoxvirus can cause disease in humans.2 Orthopoxviruses that can cause disease in humans are variola, monkeypox, vaccinia, buffalopox, and cowpox (CPV) viruses.2,3 Cowpox in humans is a rare but potentially relatively severe zoonotic disease; the majority of recent cases have been traced to infected domestic cats .4,5 Variola (the etiologic agent of smallpox) and monkeypox viruses are responsible for disease of significant morbidity and mortality in humans.

Smallpox, a severe generalized disease in humans, was declared eradicated in 1980 by the World Health Organization (WHO) shortly after the last identified natural case in 1977, nearly 2 millennia since the earliest record of this disease.2,3,6 The WHO global Smallpox Eradication Program (vaccinations) ended shortly thereafter, resulting in an increasingly susceptible population.

Monkeypox, a re-emerging zoonotic Orthopoxvirus disease, maintains an enzootic cycle in the rain forests of central and western Africa, possibly involving squirrels as a natural reservoir.7 Monkeypox virus was first recognized in captive cynomolgus monkeys in 1958.8,9 The first case of monkeypox in humans was reported toward the end of the WHO global Smallpox Eradication Program in 1970 in the former Zaire.10 Monkeypox in humans is a smallpoxlike disease that is characterized by exanthema, fever, respiratory signs, and in 10% of cases, death." Vaccination for smallpox confers 85% protection against monkeypox. Person-to-person respiratory transmission is believed to be the primary means of transmission for Orthopoxvirus infections and has been substantiated epidemiologically for smallpox. A recent WHO-sponsored epidemiological investigation of an upsurge of monkeypox cases during 1996 and 1998 in the Democratic Republic of Congo indicated that secondary (person-to-person) rather than primary (animal-to-human) transmission accounted for the majority of cases (J.W.H., unpublished data, 1998).12-16 PUl_ monary disease and respiratory distress are sometimes seen in cases of human monkeypox and are associated with significant mortality.11 The scientific community has responded to the upsurge in monkeypox cases by actively supporting screening of drugs for antiviral activity against certain orthopoxviruses, including those responsible for smallpox and monkeypox. 17 Clearly, in vivo evaluation of antipoxviral drug efficacy depends on a facilitative, well-studied laboratory animal model in which an experimentally induced Orthopoxvirus infection approximates the disease expected in humans infected with aerosolized Orthopoxvirus. The model described herein may fulfill the criteria required by antiviral drug testing.

To the best of our knowledge, no well-characterized mouse model of aerosol infection with CPV currently exists. To determine the validity of the CPV-BALB / c mouse system as a model for aerosol-transmitted Orthopoxvirus disease, we infected mice with a predetermined dose of CPV, Brighton strain, via aerosol or intranasal inoculation, and sequentially examined the morphologic changes by histology, immunohistochemistry, and electron microscoPY.

MATERIALS AND METHODS

Animals

Female BALB / c mice obtained from the National Cancer Institute, Frederick, Md, were housed in filter-top microisolator cages and given commercial mouse feed and water ad libitum. Weanling mice were allowed to reach a weight of 13 g (approximately 4 weeks old) before commencement of the experiment and were transferred to a biosafety level 3 (BSL-3) containment area before infection with CPV. All work involving CPV was done under BSL-3 conditions at the US Army Medical Research Institute of Infectious Diseases (USAMRIID), Frederick, Md.

Virus

Cowpox virus, Brighton strain (ATCC VR302) was obtained from the Centers for Disease Control and Prevention reference collection and amplified twice in Vero cells. Vero C1008 (ATCC CRL 1586) and Vero clone E6 monkey kidney cells (VERO C1008, ATCC CRL 1586) were propagated in Eagle minimum essential medium (EMEM) with Earle salts, nonessential amino acids, 10% fetal bovine serum, glutamine, penicillin, and streptomycin at 37 deg C in a 5% carbon dioxide atmosphere. The same medium with 2% fetal bovine serum was used as replacement medium after cell infection.

Viral Dose and Inoculation Methods

Mice used for aerosol inoculation were exposed to virus suspended in EMEM, or to EMEM only, in a nose-only exposure system contained within a class III biological safety cabinet. Aerosol-infected mice were exposed to an estimated dose of 5 X 10 deg plaque-forming units (pfu). Aerosols were generated by a 3jet Collison nebulizer (BGL Inc, Waltham, Mass) at a flow rate of 7.5 L min-' and a predicted median particle diameter of 1.2 Rm. The aerosol was mixed with secondary air for a total system flow rate of 12 L min-' and then divided. Half flowed to the nose ports, where it was distributed evenly through metering orifices. The other half was sampled continuously by an all-glass impinger containing EMEM with 2% fetal bovine serum and an antifoaming agent. The amount of virus collected was determined by plaque assay, and the inhaled viral dose was calculated by Guyton's formula for minute volumes of rodents. Mice used for intranasal inoculation had 50 RL of virus delivered to each nostril with a micropipette, for a total of 100 (mu)L containing 10^sup 4^ to 10^sup 6^ pfu.

The median lethal dose (LD^sub 50^) for CPV by intranasal inoculation was determined by inoculating groups of 10 mice with 10^sup 4^ to 10^sub 6^ pfu of virus, observing them for 21 days for survival, and analyzing the results by using a linear regression program (SPSS, Inc, Chicago, III). Susceptibility to lethal aerosol infection was tested in 4-week-old mice, which were exposed to doses of approximately 5 X 10^sup 2^, 5 X 10^sup 4^, and 5 X 10^sup 6^ pfU.

Experimental Design

Aerosol-infected mice were killed (described below) on days 2, 4, 6, 8, 10, and 12 postinoculation (pi). Intranasally infected mice were killed on days 2, 3, 4, 5, 6, 7, and 8 pi. Groups of 5 mice from each inoculation group were killed on each day indicated. Tissues were collected for virology or pathology studies.

Necropsy and Sample Collection

Surgical anesthesia was induced with a combination of ketamine, xylazine, and acepromazine injected intraperitoneally; the mice were then killed by exsanguination. Tissue specimens for virus culture were harvested aseptically from 3 of the 5 mice. A postmortem examination was performed, and gross lesions were recorded for all mice. The lower respiratory tract was filled with neutral buffered formalin injected under low pressure through a blunt-ended needle (PrecisionGlide, Becton Dickinson & Co, Rutherford, NJ). A complete complement of tissues was collected from all mice and immersion-fixed in 10% neutral buffered formalin for routine histologic and immunohistochemical evaluation. After fixation, skulls and long bones were decalcified in a mixture of formic acid solution and Rexyn 101 resin (Fisher Scientific, Fair Lawn, NJ). Tissues that were examined histologically included cerebrum, cerebellum; brainstem; pituitary gland; nasal structures; eyes; the external, middle, and internal ear, parotid, submaxillary, and sublingual salivary glands; parotid/mandibular lymph node; thyroid gland; parathyroid gland; trachea; esophagus, lung; mediastinal lymph node; thymus; heart; spleen; liver; gallbladder, stomach; small and large intestine; mesenteric lymph node; kidney; adrenal gland; urinary bladder; ovary; uterus; long bones from a rear limb; skeletal muscle (skull and rear limb); bone marrow (skull and rear limb); haired skin; and tail. For recording purposes, histologic lesions were graded on a scale of minimal to severe, based on extent and severity.

For transmission electron microscopy, 1 mouse in each timepoint group of 5 was anesthetized and underwent whole-body perfusion. Perfusion via the left ventricle was achieved by using a fixative solution consisting of 4% paraformaldehyde and 1% glutaraldehyde in 0.1 mol/L Millonig buffer (pH 7.4).

Histochemistry and Immunohistochemistry

Fixed tissues were routinely processed in an automatic tissue processor, embedded in paraffin blocks, sectioned at 5 to 6 (mu)m on a standard rotary microtome, and mounted on glass microslides for automated staining with hematoxylin-eosin in a Sakura DRS 601 Slide Stainer (Sakura Finetek USA, Inc, Torrance, Calif).

Replicate tissue sections were assayed for viral antigen by using either a streptavidin alkaline phosphatase immunohistochemical technique or an immunoperoxidase method with a commercially available kit (Envision System, Dako Corporation, Carpinteria, Calif), according to the manufacturer's instructions. Replicate sections of formalin-fixed, paraffin-embedded tissue specimens were mounted on positively charged glass slides (Superfrost Plus, Fisher Scientific, Pa), deparaffinized, and rehydrated through graded alcohol. Rabbit polyclonal antiserum raised against whole vaccinia virus was used as the primary antibody preparation and was diluted 1:5000 for both procedures. Briefly, for streptavidin alkaline phosphatase immunohistochemistry, sections were blocked with normal goat serum (20 minutes at ambient temperature) and sequentially incubated at ambient temperature in a humidified chamber with the following reagents: rabbit polyclonal anti-vaccinia antibody (1 hour), biotinylated goat anti-rabbit immunoglobulin antibody (30 minutes) (Vector Laboratories, Burlingame, Calif), alkaline phosphatase-labeled streptavidin (30 minutes) (GIBCO BRL, Life Technologies, Gaithersburg, Md), and naphthol AS-BI phosphate/hexazotized new fuchsin (50 minutes in dark box) (HistoMark RED Phosphatase System, Kirkegaard and Perry Laboratories, Gaithersburg, Md). Sections were counterstained with Mayer hematoxylin.

Electron Microscopy

Perfusion-fixed tissue specimens were diced into 1- to 2-MM3 blocks, postfixed in osmium tetroxide, dehydrated in graded steps of ethanol, and embedded in Poly Bed 812 (Polysciences, Inc, Warrington, Pa). Ultrathin sections were stained with uranyl acetate and lead citrate and examined with a Philips CM100 (Philips, Mahwah, NJ) transmission electron microscope at 80 kV.

RESULTS

Gross Findings

Neither aerosol- nor intranasally infected mice (hereinafter referred to as "aerosol" and "intranasal" mice, respectively) had pulmonary lesions by day 8 pi. By day 10 pi, however, the aerosol mice developed pulmonary consolidation, which eventually involved up to 85% of the visceral pleural surface by day 12 pi (Figure 1). The mediastinum was often hemorrhagic and the thymus indiscernible by day 12 pi. Cutaneous tenting and enophthalmos indicating dehydration became evident by day 8 pi in the aerosol group. Tail ulcers were noticed in some aerosol mice by day 12 pi, and 1 mouse had erythematous and swollen digits with black, desiccated nailbeds by day 12 pi.

Histologic Findings

The histologic morphology of the virus-induced lesions in the lower and upper respiratory tracts was similar between the aerosol and intranasal mice. Progressive bronchiolitis/bronchopneumonia and rhinitis/sinusitis were seen in both groups. Tracheitis / tracheobronchitis was evident only in the intranasal group. Lesions in both groups were characteristically degenerative, necrotizing, and hemorrhagic, and predominately contained neutrophilic and histiocytic infiltrates. Typically, the extent and severity of respiratory lesions increased in a time-dependent fashion. Mucosal epithelial necrosis was usually preceded by marked intracellular and intercellular edema, which contributed to a mucosa that often was 1.5 to 3 times normal thickness. Mitotic figures were seen occasionally in the epithelium of early lesions, but were noted more frequently in longer-standing lesions. Viral eosinophilic intracytoplasmic inclusion bodies (hereinafter referred to as "inclusions") were evident in epithelial cells, fibroblasts, smooth muscle cells of airways and vessels, and macrophages. Extension of infection appeared to be direct, and resulting distribution was regional; lesions suggesting dissemination were uncommon. Routine light microscopic and immunohistochemical findings in the aerosol and intranasal groups are summarized in the Table.

Bronchiolitis and bronchopneumonia were seen in both the aerosol and intranasal groups (Figure 2, A, B, and C). Viral antigen was readily demonstrated in bronchiolar epithelial and smooth muscle cells, macrophages, and alveolar septal lining cells (most likely epithelial) by day 2 pi in both aerosol and intranasal mice. By day 4 pi, viral antigen was also evident in fibroblasts, vascular smooth muscle cells, and alveolar macrophages (Figure 2, D). Viral antigen persisted in both groups examined at all later time points. Intracellular viral antigen was exclusively cytoplasmic. Immunohistochemistry did not significantly label the eosinophilic viral inclusions. Pulmonary lesions included occasional segmental vascular degeneration and necrosis with inclusions (Figure 2, E). Bronchiolar hyaline membranes were rarely noted. Although both groups had similar histopathology, the lesions differed in the apparent time of onset, distribution, progression, and incidence. Pulmonary lesions in aerosol mice were seen much more frequently in the peripheral regions of the lung lobes (Figure 2, F), whereas in the intranasal mice, lesions were central (hilar region). In the intranasal group, lesion extensiveness was greater at any one time point than in the aerosol group, when comparing similar anatomic sites. Pulmonary lesions in the intranasal mice were moderate to marked by day 4 pi, whereas lesions of similar severity in the aerosol mice were not evident until days 10 through 12 pi. The incidence of lung lesions also appeared to vary between the 2 groups; beyond day 2 pi, lesions were seen in all of the aerosol mice, and although all intranasal mice had also developed nasal lesions, only about half of the intranasal mice had pulmonary lesions. Viral tracheitis or bronchitis was evident in the intranasal group as early as day 3 pi, but similar lesions were never seen in the aerosol group.

Rhinitis was also seen in both groups (Figure 3, A, B, and C). Nasal epithelium was affected before structures in the lamina propria, and by days 3 and 4 pi in intranasal and aerosol mice, respectively, viral antigen had already extended into the lamina propria and involved cells and structures interpreted as macrophages, fibroblasts, vascular smooth muscle cells, Bowman glands, and periosteum. Viral antigen was subsequently seen in olfactory perineural cells, olfactory bulb meninges, and periodontium in both groups. Viral antigen and inflammation, coursing along olfactory nerves and periosteum, extended through the cribriform plate and along the meninges investing the olfactory bulbs (Figure 3, D, E, and F). Viral meningitis was characterized by a neutrophilic infiltrate, hemorrhage, fibrin and edema, and inclusions (Figure 4, A and B). Viral antigen was not seen in neurons.

Aerosol infection resulted in less severe nasal lesions than intranasal inoculation. By day 6 pi, lesions in the aerosol group were mild, while those in the intranasal group were moderate to severe. The earliest time point at which upper respiratory lesions were observed in the intranasal mice was day 3 pi, and these lesions rapidly progressed in severity by day 6 pi before appearing to stabilize. Early nasal lesions developed in the aerosol group on day 4, and these lesions developed slower than those of the intranasal mice, not matching those of the day 6 intranasal group until days 10 or 12 pi.

Other lesions resulting from extension of infection from the mucosal lining of the nasal passages or sinuses included dacryosolenitis, myositis involving cranial muscle, otitis media, and eustachitis. These lesions, when evident, were typically noted in the later time-point groups, that is, days 8 through 12 pi in the aerosol group and days 6 through 8 pi for the intranasal group.

Uncommon lesions suggesting hematogenous dissemination of virus were noted in both inoculation groups. By days 10 through 12 pi, tail dermatitis was seen in the aerosol mice. These lesions were characterized by epidermal or dermal necrosis, rare epidermal inclusions, and occasional dermal vascular thrombus formation. Viral antigen was not demonstrable in replicate sections. In the intranasal group, similar tail lesions with viral antigen present extensively in the epidermis, superficial dermis, and deep dermis were present in 1 mouse by day 8 pi. This same mouse also exhibited viral antigen in its femoral bone marrow.

Transmission Electron Microscopic Findings

Examination of lung (Figures 5 and 6), respiratory (Figure 7) and olfactory (Figure 8) nasal mucosae, and olfactory nerves (Figure 9) confirmed that both epithelial and mesenchymal cells supported productive viral infection, which resulted in cell injury. Replicating virus was evident in several cell types, including ciliated and nonciliated bronchiolar epithelium, fibroblasts, perineural fibroblasts, periosteal fibroblasts, bronchiolar and vascular smooth muscle cells, macrophages, nasal respiratory epithelium, and the sustentacular and basal cells of the nasal olfactory epithelium. Neither bipolar olfactory neurons nor vascular endothelial cells contained virions.

Bronchiolar epithelial cell injury was typical of that seen in other epithelial cells. Intracellular swelling was characterized by cytoplasmic rarefaction; dilatation of membranous structures, such as endoplasmic reticulum, nuclear membrane, and mitochondria; myelin figures; and distortion and loss of microvilli. Nuclei were often pleomorphic in infected cells. Progressive swelling resulted in cytolysis and release of virions. Intercellular edema varied from slightly expanded intercellular spaces to loss of cellto-cell adhesion and separation; edema distorted and thickened the normal architecture of the lamina epithelialis mucosae.

Transmission electron microscopy demonstrated developing and mature stages of CPV in infected cells (Figure 10). Viral factories, consistent with type-B inclusions, were characterized by granular viroplasm and crescents of viral bilayer membranes, many of which contained viroplasm. The morphology of the membranous crescents varied from shallow cuplike structures to nearly completely circular structures containing viroplasm. Intermediate stages of maturing virions were characterized by complete closure of the incipient membrane, condensation of nucleoprotein, formation of a dense core, a progression from round to a brick shape, and a slight reduction in size. Typically, further development resulted in intracellular mature virus averaging 300 nm in length, which featured the characteristic "dumbbell-shaped" core, lateral bodies, and external membrane. A-type inclusions (ATIs) consisted of uniformly granular, moderately electron-dense material, which was often surrounded by virions in varying stages of maturation. The ATIs ranged from less than 1 @Lrn to several micrometers in diameter and were consistent with the eosinophilic inclusions seen histologically. Mature virus was occasionally occluded within ATIs and typically consisted of many mature viral particles compactly embedded in the granular ATIs. When the polyclonal antibody preparation previously used for immunohistochemistry was used for immunoelectron microscopy, we found that viral membrane was consistently and specifically labeled, whereas the granular material of B- or A-type inclusions was not (M.J.M. and J.W.H., unpublished data, 1998).

COMMENT

The results of this study show that inoculating BALB/c mice with aerosolized CPV leads to lethal disease and, in our opinion, reflect the predicted effects of aerosol delivery or transmission of Orthopoxvirus. The morphologic features seen in the aerosol group were similar in many respects to those seen in the intranasal group; however, we observed interesting differences between the 2 groups in the frequency, distribution, and severity of lesions. We characterized the morphologic changes in both inoculation groups by histologic, immunohistochemical, and electron microscopic methods.

Aerosolized CPV consistently infected the distal respiratory tract, that is, the bronchiolo-alveolar junctions; this pattern of distribution is expected from an aerosol exposure to an infectious agent. Aerosolized virus, not already exhaled, that fell short of the distal respiratory tract was most likely cleared by mucociliary action, and that reaching beyond the bronchiolo-alveolar junctions was rapidly cleared by alveolar macrophages.

Although all intranasal mice developed nasal lesions, only half developed pulmonary lesions, which tended to develop in the central hilar region. The intranasal inoculum may have fallen short of the distal respiratory tract, and the concentrated virus overwhelmed the protective mucus blanket normally present in larger airways, resulting in a quickly developing, rapidly progressive lesion that extended from central (hilar) foci of initial infection. Additionally, the intranasal mice, but not the aerosol mice, developed tracheal and bronchial lesions. Similar interactions between host and virus may have contributed to the quicker onset and progression of nasal lesions as well.

The disparities in the location and frequency of lesions may be explained by multiple factors, including the interaction between the inoculum and the protective mucus blanket, alveolar clearance, and the differential deposition, retention, or clearance of particles of various sizes within the respiratory tract.

Both aerosol and intranasal groups developed nasal lesions. Viral antigen extended, in a time-dependent fashion, from nasal epithelium into the subjacent lamina propria and along olfactory nerves in cells interpreted to be perineural fibroblasts. Infection appeared to be supported also in cells interpreted as meningothelial cells of the dura mater, which blends imperceptibly with the periosteum investing the cribriform plate and the calvaria. Viral antigen eventually extended to the leptomeninges that invest the olfactory bulb. Turner18 worked with neurovirulent strains of vaccinia virus and has already suggested that infection of the central nervous system could occur directly via the nasal mucosa and cribriform plate. The apparent cell-to-cell extension of virus along olfactory nerves, periosteum, and meninges in our study most likely involved cell-associated enveloped virus.' In some tissue sections, we observed viral antigen coursing along olfactory nerve bundles after having entered the olfactory bulb. Whether this represented infected perineural fibroblasts or glial cells was not investigated. Viral antigen was not observed in neurons in this study, and this apparent resistance of neurons to infection was previously suggested by Mims,19 who infected mice intracerebrally with mousepox virus. The findings herein regarding meningitis and the apparent ability of meningothelial cells to support infection agree with those of others working with mousepox or vaccinia viruses in BALB/c mice.20-21 The possible relation between the development of meningitis in this study and the cases of encephalitis reported in some cases of smallpox in humans is uncertain. Although uncommon, smallpox-associated encephalitis, described as a nonsuppurative demyelinating process, was seen more frequently than postvaccinial encephalitis.3,23 Ultrastructurally, our findings fundamentally agreed with earlier descriptions of orthopoxviruses.24-29 The demonstration of increasingly large ATIs unassociated with virions, a feature also referred to as the V character, is characteristic of certain strains of cowpox, ectromelia, raccoonpox, and fowlpox viruses. Many ATIs had virions around their periphery, a feature referred to as the Vi character, whereas only occasional ATIs contained embedded (occluded) virions within the ATI (V+ character).1,30

In nasal olfactory epithelium, our electron microscopic findings are somewhat similar to those reported by Kraft et al3l in a study of an unidentified poxvirus in laboratory rats. The fact that the rats in that study originated from the former Soviet Union begs the question of whether these rats were infected with the same Orthopoxvirus that Marennikova et al described.32 Whereas the report by Kraft et al was inconclusive regarding the presence of virus in olfactory neurons, in our study we failed to see viral replication in olfactory neurons. Our electron microscopic finding of viral replication in perineural fibroblasts confirms the demonstration of viral antigen around and within olfactory nerve bundles and supports the concept of cell-to-cell extension of virus from olfactory epithelium to meninges.

Although aerosol infection of BALB/c mice with CPV has not been characterized morphologically until now, the findings here are compatible with previous reports of natural or experimental Orthopoxvirus infections in other species. Thompson et a133 showed that intranasal inoculation of wild-type CPV leads to both upper and lower respiratory disease in BALB / c mice.

The prominent hemorrhagic nature of the lesions in our study agrees with the observations of others.33-35 Although the routes of CPV inoculation differed, our findings of viral replication in vessel walls and hemorrhage confirm those of Wallnerova and Mims; however, the formation of hemal lymph nodes, as observed by Wallnerova and Mims, was not evident in our mice.35 Instead, regional hemorrhage sometimes resulted in sinus erythrocytosis in draining lymph nodes. The hemorrhage often associated with CPV-induced lesions may be associated with impaired coagulation due to a CPV-elaborated protein with homology to plasma protein inhibitors of serine proteases (serpins), as indicated by Pickup et al.34 Outbreaks of CPV infection characterized by dermal and pulmonary disease in white rats were described by Marennikova et al.32 Subsequently, Maiboroda, 36 experimentally infected (by intranasal, intradermal, or environmental exposure) wildcaught Norwegian rats with CPV, and observed nasal, pulmonary, intestinal, and cutaneous lesions. Kraft et al31 described the inflammatory and desquamating nasal lesions in clinically normal laboratory rats naturally infected with an unidentified poxvirus.

Interestingly, respiratory disease has not been associated with CPV that has been injected intracerebrally, intravenously, intradermally, cutaneously (scarification), subcutaneously, intrafollicularly (feather), or intraperitoneally in mice (J.WH. and M.J.M., unpublished data, 1997), guinea pigs, chicks, or rabbits.20,35,37-40 However, the most significant feature of CPV in domestic cats, whether induced naturally or experimentally, is respiratory disease.41,42 other than CPV, orthopoxviruses have been aerosolized and used by several investigators to infect a variety of laboratory animals, and in these cases, a broad spectrum of pulmonary lesions resulted. For example, the pathogenesis of ectromelia acquired by aerosolized mousepox virus was studied; rabbits were infected with aerosolized rabbitpox virus; cynomolgus monkeys were infected with aerosolized strains of vaccinia, variola, rabbitpox, cowpox, monkeypox, and alastrim viruses; and rhesus monkeys were infected with aerosolized variola virus. 43-411 Vaccinial pneumonia was characterized in mice and rabbits infected intranasally and intratracheally, respectively.21,49

Although the cutaneous tail lesions occasionally seen in either group of mice suggested hematogenous dissemination of virus, other more direct modes of infection, such as wound contamination with virus-laden secretions or excretions, must be considered. Our findings are largely consistent with those Of MiMS,311 who observed that CPV and certain strains of vaccinia virus cause a localized or regional infection at the site of experimental inoculation, rather than a systemic infection in immunocompetent animals. Although mediastinal lymph nodes were inconsistently present in the tissue sections in our study, the absence of virus-induced lesions or viral antigen, when nodes were present for examination, supports the localization of infection observed by Mims. Regardless, the reason the infection remained localized in our mice is uncertain, considering the morphology. The infection of cells interpreted as macrophages and their proximity to lymphatics and vessels would lead one to expect dissemination of virus and disease.44 It could be argued that the rapidly fatal outcome in both groups of mice, despite localized infection and disease, may have precluded the eventual development of systemic lesions. Rapidly lethal diseases caused by other orthopoxviruses similarly demonstrated that widely disseminated lesions may not have the opportunity to develop (eg, mousepox virus in mice, rabbitpox virus in rabbits, monkeypox in orangutans, and perhaps certain toxic forms of classical smallpox in humans). However, our observations from other experiments indicate that those mice that survived beyond expectation and that did not develop systemic lesions did so as a result of an effective immune response. Consideration should be given to other possible explanations for the apparent absence of widely disseminated lesions in our mice, such as the influence of major histocompatibility complex, strain or gender of mice used, strain of virus used, and the role of cell-associated or extracellular enveloped CPV in cellto-cell spread or systemic dissemination. 1,20,50-53 Further studies, perhaps involving scid mice, are needed to clarify the role of these factors in the pathogenesis of CPV.

The pulmonary disease in the BALB / c mice of this study is similar to that seen in experimentally transmitted aerosolized monkeypox virus in nonhuman primates (G. M. Zaucha, unpublished data, 1995). Additionally, in cynomolgus monkeys, respiratory disease has been shown to result from infection with aerosolized variola virus.46,47 In cases of smallpox in humans, Councilman et al reported bronchopneumonia, and Jezek et al as well as others reported clinically diagnosed respiratory disease in cases of fatal monkeypox in humans. 11,54-57

Given the localized nature of infection and disease in the mice used in this study, we could not always determine the mechanism of death. Although compromised pulmonary function due to pneumonia was the mechanism of death in the majority of aerosol-infected cases, examination of lung sections from half of the lethally infected intranasal mice revealed an absence of significant pulmonary disease. The reason for this disparity is unclear. However, both groups of mice did develop rhinitis and meningitis, and these lesions may have contributed to death. Further studies elucidating the role(s) of virokines, host cytokines, or meningitis are needed.

The CPV-BALB/c mouse system characterized in this study makes unnecessary the extraordinary animal husbandry or biocontainment facilities and practices that are indicated for infection of nonhuman primates with variola or monkeypox virus. BALB/c mice are a convenient, facilitative, and well-characterized strain that has already been used to study orthopoxviruses, and a large amount of related data has been amassed. Moreover, CPV shares with other orthopoxviruses many of the viral immunomodulatory proteins that are believed to weaken host resistance against infection, and this knowledge further contributes to the understanding of orthopoxviral pathogeneSiS.21,58-74

We showed that infection of BALB/c mice with aerosolized CPV supports the CPV-BALB / c mouse system as a reliable and suitable model of aerosol-transmitted, orthopoxviral respiratory disease and complements existing data from experimental Orthopoxvirus infections in other species.

We thank Bernard Moss, PhD, National Institutes of Health, Bethesda, Md, for kindly providing the antivaccinia virus antibody, and Dave Fritz, DVM, Keith Steele, DVM, and Gary Zaucha, DVM, US Army Medical Research Institute of Infectious Diseases (USAMRIID), Fort Detrick, Md, for their valuable contributions. We also thank (alphabetically) D'Angelo Austin, Jim Barth, Jeff Brubaker, Lorraine Farinick, Steve Ferendo, Tom Geisbert, Debbie Kefauver, Kathy Kuehl, Lynda Miller, Rosie Moxley, and Ralph Tammariello, USAMRIID, for their excellent technical assistance.

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Accepted for publication July 30, 1999.

From the Pathology Division (Dr Martinez) and Virology Division Qrs Bray and Huggins), US Army Medical Research Institute of infectious Diseases, Ft Detrick, Md.

In conducting research using animals, the investigators adhered to the Guide for the Care and Use of Laboratory Animals, prepared by the Committee on Care and Use of Laboratory Animals of the Institute of Laboratory Animal Resources, National Research Council (National Institutes of Health publication 86-23, revised 1996). The facilities are fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care, International.

The views, opinions, and/or findings contained herein are those of the authors and should not be construed as an official Department of the Army position, policy, or decision unless so designated by other documentation.

Reprints: LTC Mark J. Martinez, US Army Medical Research Institute of Infectious Diseases, Attn: MCMR-UIP, 1425 Porter St, Fort Detrick, MD 21702-5011.

Copyright College of American Pathologists Mar 2000
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