Establishment of a SFTSV Pathogenic Mouse Model.

In our initial study, to identify an infectious animal model that could mimic most clinical features during SFTSV infection, the susceptibilities of three commonly used laboratory rodent strains for phlebovirus (2⇓–4), C57/BL6 mice, BalB/C mice, and Syrian hamsters, were examined. The SFTSV strain HB29 was inoculated at 10⁵ TCID₅₀ (50% tissue culture infective dose) per mouse or 5 × 10⁵ TCID₅₀ per hamster through four different routes of infection, including intravenous, intramuscular, intraperitoneal, and intracerebral injections. The results showed that in C57/BL6 mice, inoculation through all four routes induced a reduction of white blood cells, and the inoculation intravenously and intramuscularly induced a reduction of platelets (Fig. S1), but abnormal medical signs and weight loss were not observed. Inoculation of SFTSV to BalB/C mice or hamsters through all of the above routes could not cause significant changes in weight, white blood cell counts, and platelet counts. However, pathology examination on day 14 showed that all these SFTSV-infected rodents developed similar pathological lesions in liver and kidney [Fig. 1, representative pathology at day 14 postintion (p.i.) from C57/BL6 mice]. Based on a comprehensive evaluation, we determined that SFTSV-infected adult C57/BL6 mice mimicked the major clinical features of leukocytopenia and thrombocytopenia in SFTS patients and, therefore, were selected for further investigations. Because SFTSV infection in the human is thought to be transmitted by tick bite (1), we chose intramuscular inoculation for subsequent animal studies.

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Fig. 1.

Dynamic profile of pathological changes in mice infected with SFTSV. Representative H&E-stained tissue sections from SFTSV-infected mice and mock mice are shown. Areas of interest (AOI) are enlarged, and quantitative graphs are presented at the right. (A) Decreased cellularity in the red pulp (RP) of the spleen at day 1 p.i.; no obvious changes are visualized in the white pulp (WP). (B) Increased megakaryocytes in the spleen at day 3 p.i.. Arrowheads indicate megakaryocytes. (C) Increased megakaryocytes in the bone marrow at day 3 p.i. Arrowheads indicate megakaryocytes. (D) In liver at day 14 p.i., arrowheads show hepatocyte necrosis with shrinking nucleic or hepatocyte degradation with balloon-like empty cytoplasm. (E) In the kidney at day 14 p.i., arrowheads show impaired renal capsules. In A and B, the images and AOI are 100× and 200×, respectively. In C–E, the images and AOI are 200× and 400×, respectively. In quantitative graphs, red dots and blue dots indicate SFTSV-infected mice and mock mice, respectively, and lines indicate the means. *P < 0.05.

Next, to thoroughly examine the complete disease progression of SFTSV infection in C57/BL6 mice, a group of 10 mice with intramuscular inoculation of 10⁵ TCID₅₀ SFTSV per mouse and a group of 5 mock-infected mice were killed at each time point of days 1, 3, 7, 14, 21, and 28 p.i. for analysis of body weight, temperature, various laboratory parameters, and pathology. Similarly, as in initial studies, SFTSV inoculation caused significantly reduced platelet count on day 3 p.i., as well as reduced white blood cell count on day 1 and day 3 p.i. (Table 1). Additionally, serum AST elevated on day 1 and lasted till day 7, and serum ALT and BUN elevated during 7–14 d (Table 1). Therefore, we showed that intramuscular inoculation of SFTSV into C57/BL6 mice could induce symptoms that mimic the thrombocytopenia, leukocytopenia, and dysfunction in liver and kidney that are key clinical presentations of SFTS patients. However, fever and weight loss were not induced in SFTSV-infected C57/BL6 mice (Table 1), and all tested mice survived. To evaluate if the immunocompromised animals could develop a more severe disease and even fatal outcome, we used immunosuppressive drug mitomycin C, which is known to inhibit the hematopoiesis in bone marrow (5), to treat SFTSV-infected mice. We found that with mitomycin C application, 50% of the SFTSV-infected mice died between day 9 and day 10, and the remaining surviving mice experienced a significant loss of body mass compared with uninfected mice administered mitomycin C in parallel (Fig. S2).

To define the infectious status of the immunecompetent mouse model, the dissemination and replication of SFTSV in blood and tissues were examined by detection of both viral RNA copies and infectious viral titers (1, 6). The results showed that blood viral load peaked on day 1 p.i. and substantially decreased on day 3 p.i. (Table 1). Dynamic virus distribution in eight tissues, including the spleen, liver, kidney, lung, intestine, heart, muscle, and brain, were also analyzed. Viral RNA was detected in three organs, including the spleen, liver and kidney, and the viral RNA copies per milligram of tissue in the spleen were significantly higher than in the liver or the kidney. Infectious viral titers were only detected in the spleen and kidney. This difference between the viral copies and infectious titers might be a result of the small amount of virus in liver being below the limit of detection for assays to determine infectious titers. We also noticed that there was a marked increase of viral load in spleen on day 3 p.i., and the virus RNA copies could be detected at a low level till to day 21 p.i. (Table 1). This finding suggested active viral replication occurred and remained at certain level in spleen. In the liver and kidney, the viral RNA copies initially were elevated on day 1 p.i. and then cleared by day 7 p.i. On the whole, the dynamic analysis of SFTSV dissemination in the blood and tissues showed that intramuscularly inoculated SFTSV entered the blood on day 1 p.i. and then were enriched in spleen on day 3 p.i., where the virus underwent a short term of replication, followed by clearance to a low level, thereafter. The sustained detection of low viral RNA copies in the spleen suggests a possible long-term harboring of SFTSV in spleen, contrasting the fast kinetics of viral clearance in other organs.

To further define immune responses induced by SFTSV infection in C57/BL6 mice, SFTSV-specific IgM and IgG antibodies in sera were tested by Luminex assays (7). The data showed that SFTSV-specific IgM peaked on day 7 p.i., and the SFTSV-specific IgG started to increase on day 7 p.i. and peaked on day 28 p.i. (Table 1). The levels of neutralizing antibodies and splenic cellular responses in SFTSV-inoculated mice were significantly enhanced over mock-infected mice on day 28 p.i. (Table 1). This evidence of efficiently elevated humoral and cellular immune responses, combined with evidence of sustained viral replication in mice, support that C57/BL6 mice experienced a true infection with SFTSV and that they provide a useful parallel to SFTS patients for studying SFTSV pathogenesis.

The dose-dependent effect of SFTSV inoculation was also assessed by intramuscular inoculation of a low dose of 10³ TCID₅₀ SFTSV per C57/BL6 mouse. Low dose of SFTSV did not induce a significant decrease of circulating platelets on day 3, but caused a significantly decreased white cell count on day 1 (Table S1). Similarly, this low dose of SFTSV induced viremia on day 1, and the virus load peaked in spleen on day 3, but the levels of viral load were less than the levels observed in high dose of SFTSV infection (Table S1). Viral copies were hardly detected in liver or kidney tissue. Low-dose SFTSV induced similar levels of virus specific IgM and IgG as high dose of virus (Table 1). Because a low dose of SFTSV induced insignificant clinical signs, low viral load, and high humoral immune responses, this finding suggests that the viral load is associated with the severity of the disease and that a low amount of SFTSV could result in an asymptomatic infection.

Pathological Lesions in SFTSV-Infected Mice.

Pathological changes within organs were evaluated in the eight tissues indicated above, as well as in the bone marrow, by H&E staining of tissue samples collected at various time points postintramuscular inoculation. During SFTSV infection, pathological changes were mainly identified in the spleen and bone marrow within 3 d p.i. The lymphocyte cellularity of the red pulp was visually decreased in spleens of SFTSV infected mice during the first week after inoculation and gradually recovered 14 d later (Fig. 1A). This observation of lymphocyte depletion in the red pulp may coincide with the systemic decrease of white blood cells. In addition, at an early stage of SFTSV infection, a marked increase of megakaryocytes was observed in the spleen, a secondary hemapoietic organ in mice, based on their cellular morphology (Fig. 1B). This finding suggests extramedullary hematopoiesis, which has been reported to occur in conjunction with decreased lymphocyte cellularity of the red pulp (8). Similarly, megakaryocytes in the bone marrow, the principle organ for hematopoiesis, also significantly increased at the early stage of infection as they did in the spleen (Fig. 1C). Because megakaryocytes are progenitor cells for platelets, we presume that the rapid increase of megakaryocytes in the hemopoietic organs of the spleen and bone marrow functioned to compensate for the depletion of circulating platelets. Megakaryocytes have extended survival; therefore, once they have proliferated, they could persist in organs for a long period (9). During the late phase of SFTSV infection, pathological changes were noted in liver and kidney. The primary lesions in liver consisted of ballooning degeneration of hepatocytes and scattered necrosis, the latter indicated by appearing multifocal pyknosis, karyorrhexis, and karyolysis (Fig. 1D). TUNEL assays on liver tissues detected few TUNEL-positive nuclei, indicating that hepatocyte apoptosis was not induced. The kidney showed glomerular hypercellularity, mesangial thickening, and congestion in Bowman’s space, but infiltration of inflammatory cells was absent (Fig. 1E). Pathological changes within the liver and kidney tissues were maximal on day 14 p.i. but had nearly recovered on day 28 p.i. Therefore, the pathological features observed in the spleen and bone marrow during the early stage of infection were consistent with the hematological changes of thrombocytopenia and leukocytopenia. The transient pathologic changes observed in kidney and liver at the later stage of infection were indicative of acute glomerular nephritis and acute hepatitis with self-limiting outcomes.

Identification and Colocalization of SFTSV, Macrophages, and Platelets in the SFTSV-Infected Spleen.

The findings that viral RNA was enriched in the spleen and that remarkable pathological changes occurred in the spleen during the early stage of infection suggested that the spleen was the primary target organ of SFTSV. Therefore, we performed an immunohistochemistry assay using a polyclonal antibody against SFTSV virions to detect target cells for viral replication, using spleens collected on day 3 p.i. The results showed that, in the spleen of SFTSV-infected mice, virus-positive cells were large monocytes, suggested by their large cytoplasm, lightly stained nuclei, and their scattered location throughout the red pulp (Fig. 2A). Immunehistochemistry staining for SFTSV was negative in bone marrow, kidney, liver, lung, gastrointestinal tract, heart, muscle, and brain. Based on the morphology and location of virus-positive monocytes in the spleen, macrophages were hypothesized to be the target cell of SFTSV. Therefore, splenic macrophages were identified by staining for the macrophage-specific membrane marker F4/80 (10). These results showed a substantially increased number of macrophages in the red pulp of the spleen on day 3 p.i. It is known that splenic macrophages are responsible for clearing old or impaired platelets from blood (11, 12). Based on these observations, we hypothesized that the overwhelmingly increased number of macrophages appearing in the spleen function to clear platelets excessively from blood, resulting thrombocytopenia. We expected that, if this were the case, more platelets would be deposited in the spleens of SFTSV-infected mice. Therefore, splenic sections from tissues isolated day 3 p.i. were probed for platelets using an antibody against CD62P, an activation marker of platelets (11). The results showed an increased deposition of CD62P⁺ platelets in cytoplasm of the macrophage-like monocytes located in the red pulp of SFTSV-infected spleens (Fig. 2A), which coincided with increased macrophage density in the spleen.

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Fig. 2.

Identification and colocalization of SFTSV, macrophages, and platelets in SFTSV-infected spleen. (A) SFTSV, macrophages, and platelets were identified by immuno-histochemistry in the spleen of Mock mice or SFTSV-infected mice at day 3. Representative images and enlarged AOI are taken at an original magnification of 200× and 400×, respectively. (B) Confocal microscopy to examine colocalization of SFTSV (in green), macrophages (in red), and platelets (in blue) in the SFTSV-infected spleen collected on day 3 p.i. The representative fluorescent images and embedded AOI were originally taken at 200× and 400×, respectively.

To further clarify whether the SFTSV colocalized monocytes were macrophages and whether the increased splenic macrophages were engaged in platelet clearance, confocal microscopy was used to examine the colocalization of SFTSV, macrophages, and platelets. The results showed that almost all increased CD62P⁺ platelets in spleen colocalized with F4/80⁺ macrophages and, moreover, SFTSV was found to colocalize with CD62P⁺ platelets and F4/80⁺ macrophages in spleen (Fig. 2B). Therefore, splenic macrophages were identified as target cells for SFTSV replication, and the increased numbers of splenic macrophages during the early stage of SFTSV infection appear to promote thrompocytopenia as a result of their responsibility of phagocytosing platelets.

SFTSV Adherence on Platelets Enhances Phagocytosis of Platelets by Macrophages.

To investigate the mechanistic interactions between SFTSV, platelets, and macrophages, mouse primary macrophages were used to test virus infection in vitro. The results showed that SFTSV could efficiently infect and replicate in mouse primary macrophages (Fig. 3A). Because some Bunyaviruses have been reported to bind to platelets (13), we hypothesized that platelet-binding by SFTSV could promote their activation and phagocytosis by macrophages. To clarify this hypothesis, serially diluted SFTSV virions were cocultured with a fixed amount of mouse platelets, after which unbound virions were washed away. By using quantitative real-time PCR to determine viral copies in platelet pellets, a gradient of viruses were found to adhere on platelets corresponding to the serial dilutions (Fig. 3B, Left). The virions that had adhered to platelets were further cultured in medium for studies to address their infectivity after platelet binding. Our results showed that the platelet-adherent virus dissociated into the supernatants after culturing for 1 d (Fig. 3B, Right). Additionally, the total amount of virus detected in the platelets and supernatants did not increase over time, suggesting that no virus replication occurred in platelets.

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Fig. 3.

Phagocytosis of SFTSV-bound platelets by macrophages. (A) The confocal image shows viral N protein (in green) in mouse primary macrophages (in blue). The graph shows dynamic virus replication in mouse primary macrophages and the virus released into culturing supernatants. (B) Real-time PCR quantification detected a gradient of adherence of SFTSV on mouse platelets corresponding to the amount of virus added into platelets. The graph on the right shows the virus amount detected in cultured virus-adhered platelets as well as in culturing supernatants at the indicated time points. (C) Confocal images show mouse primary macrophages cocultured with normal platelets (Top), mouse primary macrophages cocultured with platelets with adherent SFTSV (Middle), mouse macrophages previously infected by SFTSV for 3 d before coculturing with normal platelets (Bottom). All images were taken at an original magnification of 400×. Quantification of macrophage phagocytosis of fluorescent platelets is shown in Fig. S2D.

Moreover, mouse primary macrophages were mixed with mouse platelets that were previously cultured with or without SFTSV, respectively. After 30 min of culturing, macrophages were washed to remove unbound platelets. The results showed that, although normal platelets could adhere on the surface of uninfected macrophages, they were rarely internalized (Fig. 3C, Top). In contrast, platelets with adherent SFTSV were rapidly phagocytosed by uninfected macrophages (Fig. 3C, Middle). The SFTSV-infected mouse macrophages did not show a significant difference in their phagocytosis of platelets (Fig. 3C, Bottom). We also tested the macrophage phagocytosis of virus-adhered platelets using human macrophage cell line THP-1, differentiated by phorbol-12-myristate-13-acetate (11, 12, 14). Similarly, we found that SFTSV could infect human THP-1 cells and adhered on human platelets. Furthermore, only SFTSV-adhered platelets but not normal platelets could be phagocytosed by THP-1 cells, and SFTSV-infected THP-1 cells were incapable of phagocytosing normal platelets (Fig. S3). Therefore, the adherence of SFTSV on platelets is necessary for platelets to be phagocytosed by macrophages. Although we observed that macrophages could be directly infected by SFTSV, we expect that the phagocytosis of virus-decorated circulating platelets would deliver larger amounts of virus into macrophages than direct infection, while concurrently promoting the internalization of platelets.