Dynamics of Puumala hantavirus infection in naturally ... - Springer Link

Arch Virol (1999) 144: 2415–2428

Dynamics of Puumala hantavirus infection in naturally infected bank voles (Clethrinomys glareolus) A. D. Bernshtein,1 N. S. Apekina1 , T. V. Mikhailova1 , Yu. A. Myasnikov1 , L. A. Khlyap1 , Yu. S. Korotkov1 , and I. N. Gavrilovskaya2 1

Institute of Poliomyelitis and Viral Encephalitis, Russian Academy of Medical Sciences, Moscow, Russia 2 The Department of Medicine, Stony Brook University, Stony Brook, New York, U.S.A. Accepted May 17, 1999

Summary. Specific features of hantavirus infection in bank vole (Clethrionomys glareolus) were studied in the endemic area of hemorrhagic fever with renal syndrome (HFRS) in the foothills of the Ural mountains, using long-term observations on living animals by the capture-mark-recapture (CMR) method. The results demonstrated that the infection naturally circulating in the voles is chronic (lasting for up to 15 months) and asymptomatic, with a peak of Puumala virus accumulation and release from the organism during the first month after infection. It was shown that the bank vole population includes young animals with maternal immunity, which remain resistant to the Puumala virus infection for 3– 3.5 months. The infection rate in voles depended on the age and sexual maturity of animals. The greatest proportion of seropositive animals was observed among overwintered males. Seroconversion in voles was more frequent during the period of high reproductive activity. Introduction The genus Hantavirus, family Bunyaviridae, comprises at least 14 viruses, including those that cause hemorrhagic fever with renal syndrome (HFRS) and hantavirus pulmonary syndrome (HPS) [45]. Hantaviruses are primarily rodent-borne, although other animal species carrying hantaviruses have been reported [19, 27, 29, 39, 45, 47]. Unlike all other viruses in the family, hantaviruses are not transmitted by arthropod vectors but from inhalation of virus-contaminated aerosols of rodent excreta [29, 45]. Recently the possibility of human-to human transmission of hantavirus in Argentina is discussed [16].

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Hantaviruses fall into three distinct genetic groups that are parallel to the phylogens of rodent hosts in the subfamilies Murinae, Arvicolinae, and Sigmodontinae [24, 26, 42, 45]. Hantaan (HTN), Seoul (SEO), and Dobrava (DOB) viruses are carried murine-host, which cause severe forms of HFRS and are found primarily in East Asia and the Balkan region [29, 41, 45, 49]. The viruses hosted by arvicoline rodents group include the Puumala (PUU) virus which cause a milder form of HFRS in several European countries. The Sigmodontinae-associated hantaviruses were discovered in 1993 in the southwestern United States and has subsequently been identified in 28 states and Canada. Recently identified HPS cases in South America indicate that HPS-associated hantaviruses are widely distributed. At least seven serologically distinct hantaviruses caused MPS in both Americas [15, 16, 23, 36, 43, 44]. Today it is obvious that the bank vole (Clethrionomys glareolus) is the principal host of PUU hantavirus and the source of human infection of HFRS in Europe [6, 10, 11, 13, 19, 38, 47] and West Siberia [34]. After the isolation of PUU and PUU-like viruses from bank voles [3, 10, 20, 52], experimental studies on pathogenesis of infection in these animals were performed [9, 21, 53]. Moreover, certain ecological and epizootiological features of hantavirus infection in bank vole populations were analyzed [1, 2, 4, 6, 50, 51]. However, many aspects of PUU virus circulation in naturally infected hosts remain poorly studied. We performed long-term observations on bank voles in an endemic area of HFRS in foothills of the Ural Mountains. In such forest areas bank vole is the dominant species both among small mammals and particularly among infected animals [1, 4, 6, 19]. In this paper, we describe the results concerning the course of the infection process in the host organism. Dynamics of PUU virus at the host population level will be the subject of a separate paper. Materials and methods Field research Studies were performed from 1987 to 1992 in the HFRS endemic area in southern Udmurtia (56 ◦ 200 N and 52 ◦ 400 E), in the subzone of mixed conifer-broadleaf forests of the central foothills of the Ural Mountains. The study site (a plot 2.2 hectares in area in 1987–1989 and 3.2 hectares in 1990–1992) was located in an old tree stand with prevalence of linden (Tilia cordata) and spruce (Picea abies) and with well-developed undergrowth and herbaceous vegetation. During the study period, bank voles accounted for 70–80% of small mammals inhabiting the study area. Their density on the plot averaged at 29 animals per hectare in spring, increasing to 155 animals per hectare by the end of summer. The reproductive period continued from March-May to August- September in different years. Field collection Field research in 1987 was performed in September, and in the following years, from May to August. Animals were studied using the capture-mark-recapture (CMR) method. Live traps were situated at IO-m intervals. One trapping period lasted for 6–10 days with 2–3 trap checkings per 24 h. During the five years, trapping periods from May until August numbered as follows: two in 1988, seven in 1989, four in 1990, five in 1991 and four in 1992. Each newly

Dynamics of Puumala hantavirus infection

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trapped vole was marked with toe clipping, and its sex and age were determined. To determine the age, we weighed each marked animal to an accuracy of 1.0 g and described its external and reproductive status. According to these criteria, all the voles were divided into two groups: overwintered and yearlings. The first comprised of animals over nine months of age and weighing over 22 g, all of which were sexually mature. The second group included voles under five months of age. Some of them reached sexual maturity during summer, and their weight strongly varied depending on the time of birth and the reproductive condition. Within the latter group, we distinguished voles 20–40 days of age weighing 8–13 g, which usually had a juvenile pelage (sometimes moulting) and were sexually immature or just entered the reproductive cycle. If we had any doubt about the age of animals at first capture we did not include them in any categories. From 1987 till 1992, 133 overwintered, 1116 yearlings (older than 40 days), and 1105 voles 20–40 days old were collected using the CMR method. The age was not detected for 321 marked voles. In addition, 630 overwintered and 1083 yearling voles were collected using snap traps on the neighbouring territory. These animals were used for analysis of sex-related differences in the frequency of hantavirus infection in the different age groups. The age of bank voles was determined by the development of the molars [48]. Blood (approximately 0.1 ml) was drawn from the retroorbital sinus of each animal, diluted with four volumes of PBS (pH 7.4), and stored at 4 ◦ C until analysis for the presence of antibodies. Samples were taken from all the voles captured throughout the field season, from May to August, with intervals of 15 days in 1989, 30 days in 1991, and from 40 to 70 days in other years. In the case of overwintered voles, the maximal duration of sampling period was 15 months, and the interval between samples reached eight months (from September to April). During the study period, a total of 2675 bank voles was examined, 641 animals were bled twice, and 570 animals were bled 3–7 times. To analyse the presence of hantavirus antigen and virus infectivity, we used lungs dissected from some voles with known dates of acquiring infection, stored in liquid nitrogen. This material is described below (Results, Table 1). Lungs and hearts of the bank voles caught with snap traps were used for detection of PUU virus antigen and antibodies respectively. Laboratory studies 1. Antibodies to PUU virus were revealed by indirect immunofluorescent assay (IFA) as described previously [19]. We used acetone-fixed Vero E-6 cells infected by Puumala-like hantavirus, strain Cg-1820, and FITC-labeled rabbit gamma-globulin against mouse globulin as secondary antibodies. To obtain a contrasting background and reduce non-specific fluorescence, rhodamine-labeled albumin was added (reagents were manufactured by the Gamaleya Institute of Epidemiology and Microbiology, Moscow). Titers of positively reacting sera were determined by preparing their fourfold dilutions in PBS, pH 7.4 (1:8–1:1 024). A total of 4848 serum samples was tested for the presence of Puumala virus-specific antibodies. 2. The PUU antigen in organs of wild and laboratory raised voles was determined by a direct ELISA test described previously [18, 36]. Reactions were performed in polystyrene plates (Research Institute of Medical Equipment, Moscow). Wells were sensitized by adding IgG isolated from the immune human reference serum (titer of antibody to PUU virus > 1: 10,000 in IFA, 100 ␮l per well). The same serum was used to prepare IgG conjugated with horseradish peroxidase type VI, RZ = 3.1 (Sigma). Optical density at 492 nm was measured in a Titertek Multiscan photometer (Flow Laboratories). The reaction was regarded as positive if the ratio of OD492 in experimental and control samples (P/N) exceeded 2.1. [32]. Identified antigens were titrated by preparing twofold dilutions of the initial 10% tissue suspension (1: 2–1: 2 048). The highest dilution with P/N ≥ 2.1 was regarded as antigen titer. The controls of the assay were: 1. Positive reference antigen (suspension of the lungs

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of experimentally infected bank voles with titer in ELISA I : 256). 2. Negative reference antigen (suspension of the lungs of noninfected laboratory raised bank voles). 3. Pretreatment of the plate with human IgG without antibody to PUU, HTN, SEO and PHV viruses (IFA). 4. Substrate solution. 3. Virus infectivity was estimated in biotests. Marked bank voles with known dates of infection, captured in the study area, served as donors of the virus. The group of recipients consisted of bank voles from the laboratory colony maintained in the Institute of Poliomyelitis and Viral Encephalitis since 1967 [14]. Before experiments, all the recipients were tested serologically. We used two variants of experimental procedure. The first consisted of placing one donor vole and two or three recipients in the same cage for one day. In the second variant, recipients were infected parenterally: 10% lung cell suspension from each donor was injected intramuscularly to five laboratory voles (0.3 ml per animal). In both cases, recipients were sacrificed after 20 days to analyze their lungs, spleens, and blood by means of IFA and ELISA. The presence of PUU antigen in organs and (or) antibodies in blood of recipients provided evidence that the donor was infectious. Parameters of infection in voles The presence of specific antibodies at titers of I: 8 and higher in blood sera of voles aged over 40 days was regarded as proof of their infection by PUU virus. Antibodies detected in younger voles were regarded as maternal if they subsequently disappeared [21, 54]. The time of acquiring the infection was determined by seroconversion occurring in twin serum samples and by IgG avidity assay [22]. Another criterion of infection was the presence of the antigen and (or) infective virus in organs. In bank voles captured in the field, only the lungs were analyzed, because previous studies showed that the PUU virus affects the lungs more severely than other organs [19, 21, 53]. The degree of virus release from animal organism was estimated only from horizontal virus transmission in experiments with animals placed in the same cage. Statistical analysis 2

0

Chi-square (χ ) test with Yates correction and Student t-test were used in the statistical comparisons [33].

Results The dynamics of infection process and horizontal transmission of PUU virus Bank voles retained PUU virus-specific antibodies, virus antigen, the infective virus, and the ability to transmit it for a maximum of 15 months after being infected (Table 1). Antibodies persisted in most of infected animals throughout this period, whereas the intensity of virus reproduction, the frequency of its horizontal transmission, and the accumulation of PUU antigen considerably decreased with time. Note that wild voles (donors) transmitted virus to their nonimmune cage mates more effectively during the early period after being infected: in the first month, horizontal transmission occurred in nine out of 14 cases (64.3%), whereas in the following months, in only four out of 32 cases i.e. 12.5% (χ 2 = 12.88, p < 0.001). Similar data given have been obtained for the virus-specific antigen (χ 2 = 6.31, p = 0.01). Prevalence of the virus in the lungs significantly decreased in the latest time post infection: 80.8% in the first four months and 30.7% in the following months (χ 2 = 9.42, p = 0.002) (Table 1).


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Table 1. Time course of PUU infection in wild bank voles Time p.i.

Antibody

Antigen

No.a

%

No.

%

Viral transmission 1b 2c No. % No.

161/163 72/74 47/48

98.8 97.3 97.9

20/31 13/37 12/30

64.5 35.1 40.0

9/14 3/22 1/10

%

100 76.2 30.7

(month) 0.05

0.08

> 0.05

0.13

> 0.05

Number of seropositive bank voles/number of tested bank voles in May–August

Table 3. Seroprevalence in different bank vole cohorts based on snap-trapped material Category

Group

Overwintered

Mature

Yearlings (over 1.5 months)

Mature Immature

a

females males females males females males

No.a

%

χ2

p

99/202 146/226 43/138 29/63 73/510 87/574

49.0 64.6 31.2 46.0 14.3 15.2

10.6

0.001

4.16

< 0.05

0.15

> 0.05

Number of seropositive bank vole/number on tested bank voles in April–September

snap traps (Table 3). The study of this larger material showed that the seroprevalence in mature males was significantly higher than in mature females. Sex differences were found both among overwintered and yearling mature bank voles. The seroprevalence was highest in the cohort of overwintered males. More than 80% of voles failing to reach sexual maturity in the year of birth remained seronegative until the end of summer. Any sex-related differences in the infection rate among immature voles were absent (Tables 2 and 3). Seroconversion in bank voles occurred both in the spring-summer season and during the period from autumn to spring. During the study period (1987– 1992), a total of 42 out 89 nonimmune voles in May (33 overwintered and 56 yearlings) became seropositive during the period from May to August (47.2%). Of 90 yearlings that were seronegative in late August and survived until the next spring, 53 animals (58.9%) were seropositive in the following May. Thus, the average monthly infection rate during the period from May to August was 11.8% and from September to April, 7.3% of all animals examined. In 1989 and 1991, we managed to follow the dynamics of vole infection during the spring-summer season (Table 4). Seroconversion occurred in any month, but its frequency was significantly higher in June, then in May, July and August (Total χ 2 = 6.80, 5.86, 6.87, respectively; p = 0.01 − 0.009). Thus, the peak of infection coincided with the time when young voles of the spring generation reached sexual maturity and entered in the reproductive cycle. In the two studied

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A. D. Bernshtein et al. Table 4. Infection of bank voles in spring and summer

Year

1989 1991 Total

May No.a

%

June No.

%

July No.

%

August No.

%

1/9 4/84 5/90

11.1 4.9 5.5

15/77 20/132 35/209

19.5 15.2 16.7

12/157 30/262 42/419

7.6 11.5 10.0

11/152 36/331 47/483

7.2 10.9 9.7

a

Number of new seropositive animals in the month/number of seronegative animals in the beginning of a month

Fig. 2. The dynamics of acquiring infection by bank voles inhabiting the study area from May to August in 1989 and 1991. Number of mature voles: 22 in May, 39 in June–August. Number of immature voles: 31 in May, 14 in June–August

years all voles that were born during the summer remained immature until the next reproductive season. It is such animals that were the majority of the tested voles in July and August [8]. Note that the data shown in Table 4 do not take into account the animals with congenital immunity. To study the effect reproductive status on the infection rate, we used a group of 53 animals (10 overwintered and 43 yearlings) regularly examined from May until late August of 1989 and 1991 (Fig. 2). All the animals were seronegative in early May. In this group, 22 voles had reached sexual maturity by May and 17 voles in June. From May to July seroconversion was registered at 21 out of 39 mature voles and 2 out of 14 immature ones (53.8% and 14.3% respectively, χ 2 = 6.56, p = 0.01). Two more voles became seropositive in August (Fig. 2). From these data it follows that mature animals can be infected more intensely than immature.


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Survival of infected bank voles in the study area In the spring-summer season, the life expectancy in the study area was virtually the same for infected and noninfected bank voles: 55.4 days (S.E. = 5.4, n = 43) and 56.0 days (S.E. = 2.3, n = 114), respectively, (t = 0.1) according to the data obtained in 1989. We took into account voles of all age and sex groups trapped in the study area beginning from May or June of the current year. Animals of different immune status similarly survived the period from autumn to spring. In the years of our observation, the proportion of seronegative and seropositive voles surviving from August until next May averaged at 8.1% (n = 1233) and 8.3% (n = 265), respectively. Discussion Our long-term studies on bank voles in an endemic area confirmed the existing experimental data on the chronic progression of hantavirus infection in natural hosts [9, 17, 21, 30, 31, 46, 53]. In certain cases, bank voles remain infected for 15 months (time of the observation), which is close to their maximal lifespan in nature [40]. This is evidence that the natural circulation of PUU virus in bank voles may be lifelong. The virus in the lungs of voles was revealed more frequently than the virus-specific antigen. As demonstrated previously [9], the PUU antigen is not necessary detected in all virus carriers: in some animals the concentration in lungs may be below the limits of ELISA sensitivity. In such cases, however, the antigen is sometimes possible to reveal in other organs [1]. Virus reproduction and release from naturally infected voles reach a peak during the first month after the onset of infection, as under experimental conditions [21, 53]. Our experiments [21] showed that not only infected animals but also materials from their nests serve as a source of infection during this period, so that virus is transmitted both during the contact and with aerosols, to a distance of up to 1.5 m. In the following months, virus concentration in most voles apparently decreases, because they transmit infection to healthy animals more rarely even at direct contact. The probability of airborne infection in “excretory points” and nests at this stage is even lower. It is obvious that to provide for infection of voles under such conditions, virus concentration in the environment should exceed that achieved during behavioral contacts. We cannot exclude that long-term circulation of hantavirus in the host organism may lead to changes in its biological properties, but this issue requires special investigation. The data discussed above show that one of the conditions necessary for the development of intense epizootic process in HFRS endemic areas is that the principal host population, besides high total density, should include the sufficient number of freshly infected individuals. Today, we definitely conclude that the natural hantavirus infection in bank voles goes on without any apparent symptoms. In previous works, we have already noted that HFRS epizootics affect neither the physiological state of these animals nor their reproduction and population dynamics [5, 6]. The data given in the article indicate that survival of voles has no relation to the infection by

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PUU virus. Verhagen et al. [50] also noted that the survival of bank voles in an endemic area does not depend on the PUU infection. Moreover, the mobility of seropositive and seronegative bank voles and sizes of their home ranges proved to be similar as well [8]. All these facts, along with the absence of any serious pathomorphological changes in organs of infected individuals [9], suggest that PUU virus has no adverse influence on vital activities of bank voles. Hosts of other hantaviruses also proved to be free from any clinical symptoms of disease [25, 30, 31, 47]. We found that the bank vole population of HFRS endemic areas includes young animals with congenital immunity. As in previous studies [21, 50, 54], the progeny of infected females usually retained the inherited (passively acquired) antibodies until the age of no more than 1.5 months. Nevertheless, these animals remained resistant to infection for the additional 45 to 60 days. In certain cases, the accumulation of young immune animals in the bank vole population may apparently impede the development of epizootic process. The results of this work and the data published previously [6, 50, 51] provide evidence for the existence of a positive correlation between the age of bank voles and the seroprevalence. Nonimmune voles of any age are apparently equally susceptible to PUU virus infection. Hence, the probability of acquiring infection increases with age. Another interesting aspect is the relationship between the seroprevalence and the reproductive status of bank voles. Seroconversion occurs more frequently during the reproductive season and in sexually mature voles, and the “group at risk” largely consists of recently mature nonimmune individuals. It is possible that among overwintered voles in which antibodies were first detected in May, many animals became seropositive soon after the beginning of reproductive season, in March or April, rather than in autumn or winter. This assumption is partly confirmed by the findings based on the snap-trapped material of a similar HFRS endemic area. For three years seroprevalence among bank voles increased two or three times from February or March to May (our unpubl. data). A special role in the maintenance and spread of infection may belong to mature males. As their mobility in reproductive season markedly increases, they travel over considerable area and can widely disseminate the pathogen. In addition, they frequently visit “excretory points” and shelters of other animals [28] and, hence, are at greater risk of acquiring infection. Reproducing females are less mobile, remain within their relatively defined home ranges, and may play a less important role in spatial dissemination of the virus. During the reproductive period, the frequency of direct contacts between voles of different sexes drastically increases. Females in estrus can mate with several males in a row, and males, in turn, usually mate with several females within a short period of time (12). The possibility of sexual transmission of hantavirus has not yet been proved, but, nevertheless, behavioral contacts during mating provide one of the most feasible natural pathways of PUU virus transmission between animals of different sex. The probability of horizontal transmission between mature females during the reproductive period is low, because their home ranges are largely


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isolated from one another. In mature males, such transmission is only possible during aggressive encounters. Compared to adults, sexually immature voles are less aggressive and usually remain more aggregate throughout the year. However, their individual ranges are small, and contacts are limited to close neighbours. Beyond the reproductive season, a similar spatial structure is also characteristic for the entire bank vole population [40]. As a result of this an early onset of the bank vole reproduction season could intensify the epizootic process. Such a phenomenon was observed in endemic areas with the optimum of bank vole distribution where breeding under snow occurred regularly [2, 6, 7, 35]. References 1. Apekina NS, Myasnikov YuA, Gavrilovskaya IN, Ryltseva EV, Bernshtein AD, Ozhegova ZE, Kopylova LF, Gorbachkova EA (1982) Detection of HFRS-antigen in autonomous natural foci of infection in Udmurtskaya ASSR. Ecology of viruses, Moscow, pp 99–103 (in Russian) 2. Apekina NS, Bernshtein AD, Myasnikov YuA, Kopylova LF, Gavrilovskaya IN (1991) The nature of functioning of the hemorrhagic fever with renal sindrome (HFRS) foci with the principal host of virus – Clethrionomys glareolus. International Symposium on HFRS, Leningrad, p 8 3. Bashkirtsev VN, Tkachenko EA, Dzagurova TK, Ryltseva EV (1984) Isolation of strains of haemorrhagic fever with renal syndrome virus of cell culture. Vopr Virusol 4: 497–502 (in Russian) 4. Bashkirtsev VN, Ryltseva EV, Tkachenko EA, Stepanenko AG (1985) On some features of epizootic process in haemorrhagic fever with renal syndrome. Vopr Virusol 4: 463–467 (in Russian) 5. Bernshtein AD, Ryltseva EV, Zubri GL, Sonkin VD, Myasnikov YuA (1971) Morphophysiological characteristics of two populations of bank vole in natural foci of haemorrhagic fever with renal syndrome. Viral hemorrhagic fevers, Moscow, pp 301–313 (in Russian) 6. Bernshtein AD, Apekina NS, Kopylova LF, Myasnikov YuA. (1987) Comparative ecological and epizootological characteristics of Clethrionomys from the Middle Ural region. Zool Zh 66: 1 397–1 407 (in Russian) 7. Bernshtein AD, Kopilova LF, Apekina NS, Mikhailova TV (1997) Prognosis of hemorrhagic fever with renal syndrome outbreak. RAT-INFO 2: 10 (in Russian) 8. Bernshtein AD, Mikhailova TV (1999) Social organization of bank vole (Clethrionomys glareolus) in natural foci of hemorrhagic fever with renal syndrome. Zool Zh (in Russian) 9. Bogdanova SB, Gavrilovskaya IN, Boiko YA, Prokhorova IA, Linev MB, Apekina NS, Gorbachkova EA, Rymalov IV. Bernshtein AD, Chumakov MP (1987) Persisting infection induced by hemorrhagic fever with renal syndrome in bank voles (Clethrionomys glareolus), natural host of the virus. Microbiol Zh 49: 99–106 (in Russian) 10. Brummer-Korvenkontio M, Vaheri A, Huvi T, von Bonsdortf CH, Vuorimies J, Manni T, Penttinen K, Oker-Blom N, Lahdevirta J (1980) Nephropathia epidemica: detection of antigen in bank voles and serologic diagnosis of human infection. J Infect Dis 141: 131–134 11. Brummer-Korvenkontio M, Henttonen H, Vaheri A (1982) Hemorrhagic fever with renal syndrome in Finland: ecology and virology of nephropathia epidemica. Scand J Infect Dis [Suppl] 36: 88–91

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Authors’ address: Dr. A. Bernshtein, Institute of Poliomyelitis, Moscow Region, 142782 Russia. Received December 12, 1997