Elucidation of prognostic factors in the acute phase of feline severe fever with thrombocytopenia syndrome virus infection

Abstract

Severe fever with thrombocytopenia syndrome (SFTS) is a potentially fatal tick-borne zoonotic disease, endemic to Asian regions, including western Japan. Cats appear to suffer a particularly severe form of the disease; however, feline SFTS is not clinically well characterized. Accordingly, in this study, we investigated the associations of, demographic, hematological and biochemical, immunological, and virological parameters with clinical outcome (fatal cases vs. survivors) in SFTSV-positive cats. Viral genomic analysis was also performed. Viral load in blood, total bilirubin, creatine phosphokinase, serum amyloid A, interleukin-6, tumor necrotic factor-α, and virus-specific IgM and IgG differed significantly between survivors and fatal cases, and thus may have utility as prognosticators. Furthermore, survivor profiling revealed high-level of viremia with multiple parameters (white blood cells, platelet, total bilirubin, glucose, and serum amyloid A) beyond the reference range in the 7-day acute phase, and signs of clinical recovery in the post-acute phase (parameters returning to, or tending toward, the reference range). However, SFTSV was still detectable from some survived cats even 14 days after onset of disease, indicating the risk of infection posed by close-contact exposure may persist through the post-acute phase. This study provides useful information for prognostic assessments of acute feline SFTS, and may contribute to early treatment plans for cats with SFTS. Our findings also alert pet owners and animal health professionals to the need for prolonged vigilance against animal-to-human transmission when handling cats that have been diagnosed with SFTS.

Severe fever with thrombocytopenia syndrome (SFTS) is an emerging, and potentially fatal, zoonosis that has become a major challenge in One Health medicine. Its causative pathogen—the bunyavirus Dabie bandavirus [or STFS virus (SFTSV)]—was first identified in China in 2011 [35] and has since spread across East and South Asia [11, 16, 27, 31, 33, 36]. SFTSV reached Japan in 2013 [27] and is now endemic to the west of the country. The virus is known to circulate in and between ticks and humans and a range of domestic and wild animal species in endemic area [23]. Among these affected species, humans have inevitably received the most attention. Patients exhibit clinical signs including fever, thrombocytopenia, leukocytopenia, gastrointestinal dysfunction, muscular and neurological abnormalities, and coagulopathy [28], and the reported case fatality rate in Japan is 27–30% [12, 23].

In addition to human SFTS, SFTS with clinical signs has been reported in captive cheetah [18], domestic cats [19], and dogs [9] in Japan. The number of domestic cats with SFTS is higher than that of dogs [9], and cats appear to be highly susceptible to the virus, exhibiting severe clinical signs (including fever, leukocytopenia, thrombocytopenia, weight loss, inappetence, jaundice, and inactivity) following exposure to the virus [22]. The reported case fatality rate of feline SFTS is 62.5% [19]. Cats-to-human SFTS transmission has recently been formally confirmed in two reports, including one in which a veterinarian was infected after exposure to at least three SFTSV-infected cats within a 3-week period prior to disease onset [10, 32]. Considering the clinical severity and zoonotic nature of feline SFTS, cats could be a source of infection for humans and other animals. Cats may also be important sentinel animals for understanding the spread of the virus in endemic region.

SFTS has become widely recognized by clinical veterinarians in Japan, and the rate of diagnosis has increase. However, prognostic indicators have not been established and the disease has not been fully profiled through the acute and post-acute phases. Based on a 24-cat study in 2019 [19], we reported that disease severity appears to peak around day 7 after onset, and we defined the acute phase as within 7 days of onset, and the post-acute phase as the subsequent period. We suggested that acute-phase creatinine phosphokinase (CPK) level may predict outcomes and commented on the need for further investigation in high-powered studies. Since then, our laboratory has continued to perform feline SFTS diagnostic testing for affiliated animal hospitals across western Japan. Accordingly, in this retrospective study, we evaluated a range of demographic, clinical, immunological, and virological parameters to determine their ability to predict the outcomes in cats diagnosed with SFTS. Furthermore, we used the data obtained to clinically characterize feline SFTS as a risk factor through the acute and post-acute phases.

MATERIALS AND METHODS

Patients and samples

SFTS diagnostic tests were performed on 796 cats at the Transboundary Animal Diseases Research Center, Joint Faculty of Veterinary Medicine, Kagoshima University between August 2017 and September 2021, using blood samples submitted from affiliated primary care veterinary hospitals across western Japan. The inclusion criteria were a confirmed diagnosis of SFTS and a known mortality status (survivor or fatal case) up to four weeks after initial presentation to the relevant animal hospital. SFTS was diagnosed based on the detection of viral RNA using reverse transcription polymerase chain reaction (RT-PCR) assay or anti-SFTSV IgM by enzyme-linked immunosorbent assay (ELISA), as described in our previous study [19]. We also included subsequent blood and swab samples from individuals in the study population through four weeks after the initial presentation. This study was approved by the Institutional Animal Care and Use Committee of Kagoshima University (approval number: JFVM20035) and was conducted in accordance with the guidelines of the committee.

Demographic characteristics and clinical pathology

Based on information from the affiliated veterinary hospital, age, sex, rearing environment, and clinical signs were recorded for each animal together with laboratory panel results at the acute phase, which was regarded as the period from onset of the disease (day 0) to day 7. The panel consisted of red blood cell (RBC), white blood cell (WBC), and platelet (PLT) counts, total bilirubin (T-Bil), CPK, alanine aminotransferase (ALT), aspartate aminotransferase (AST), glucose (GLU), blood urea nitrogen (BUN), creatinine (Cre), and total protein (TP).

We performed additional analyses for serum amyloid A (SAA), interleukin-6 (IL-6), interleukin-1β (IL-1β) and tumor necrosis factor-α (TNF-α). SAA was measured by a latex agglutination immuno-assay using Ramute Cat SAA Neo kits (SHIMA Laboratories Co., Ltd., Tokyo, Japan). The cytokine analyses were performed with commercial ELISA kits [DuoSet ELISA Feline IL-6, DuoSet ELISA TNF-α (R&D Systems, Minneapolis, MN, USA) or RayBio Feline IL-1 beta ELISA kit (RayBiotech, Peachtree Corners, GA, USA)]. Five plasma samples from healthy cats, which were seronegative for SFTSV, were examined for IL-1β, IL-6, and TNF-α as disease negative controls and determined to be below the detection limit for IL-1β (16 pg/mL), IL-6 (31.3 pg/mL) and TNF-α (50 ng/mL).

Antibody titers

Serum or plasma samples were analyzed for feline SFTSV-specific IgG and IgM using indirect ELISA, as described in our previous study, regarding values >0.5 as positive results.

Quantitative real-time PCR for SFTS viral RNA

The clinical samples used in this study comprised of sera, and oral and rectal swabs. Based on the nucleotide sequence of the S segment, we designed primers and probe for quantitative detection using real-time PCR. In this kit, the forward primer was 5′-ACCCTCACWGGRGTGATTGA-3′ (nt 1,383–1,402), the reverse primer was 5′-GTTGATGGCACTYCARGAGAAA-3′ (nt 1,457–1,436), and the probe was 5′-FAM-AGCCTGGTYTCWGCCCTCTC-BHQ-3′ (nt 1,404–1,423). Real-time PCR was performed using a Luna Universal probe one-step RT-qPCR kit (New England Biolabs, Ipswich, MA, USA), which contained 5 μL of 2× RT-PCR Buffer, 10 μM of primer mix, 10 μM of probe mix, 0.25 μL of 20× RT-PCR Enzyme Mix and 2 μL of RNA transcript or viral extracts as template at a final volume of 10 μL. Real-time PCR cycling was performed on a BioRad CFX connect system as follows: reverse transcription at 55°C for 10 min, Taq polymerase activation at 95°C for 1 min, amplification for 45 cycles consisting of a denature step at 95°C for 10 sec and annealing–extension step at 60°C for 30 sec. The emitted fluorogenic signals collected during annealing and extension. The cutoff cycle threshold (Ct) value for a positive sample was set at 1,000 RFU, based on the linear range of a typical standard curve from synthetic RNA of the target region. The assay was performed in triplicate and the values were evaluated within or between runs to assess reproducibility. The coefficient of variation was calculated for mean Ct values. This assay was used to determine the SFTSV copy number in each clinical sample. SFTS copy numbers were calculated per milliliter for each clinical sample, based on standard curves from synthetic RNA.

Genotype analysis

The partial S segment of SFTSV was sequenced with Sanger sequencing, using samples in which viral RNA was detected. In the primer set, the forward primer was 5′-AGAAGACAGAGTTCACAGCA-3′ and the reverse primer was 5′-ACACAAAGAACCCCCAAAAAAG -3′ [34]. Sequence alignment was computed using the MUSCLE and built-in MEGAX software. Phylogenetic trees were constructed based on the partial S segment, using MEGAX software, and the association between the viral genotype and outcome was evaluated.

Statistical analysis

Data collected from SFTSV-positive cats between day 0 (onset of disease) and day 7 were analyzed to compare survivors and fatal cases. Continuous variables with abnormal distributions are shown as median, minimum, and maximum values. Comparisons were performed using the Wilcoxon signed-rank test for viral loads (blood and swab samples), laboratory parameters, and antibody titers, with P<0.05, considered statistically significant, and the χ² test for virus genotype (strain), age, sex, rearing environment, and clinical signs. When more than one sample was submitted for an individual cat, all data from days 0–7 were used for analysis. Acute-phase and non-acute-phase data from survivors were also compared against the relevant reference range [14, 29].

RESULTS

Study population

We included 85 cats that had been diagnosed with SFTS for which the outcome was known, in this study. Of these cats, 58.8% (50/85) were male and 35.3% (30/85) were female (the sex was indeterminate for five cats). Their ages ranged from 5 months to 18 years, with the majority (55.3%, 47/85) aged 1–5 years. In terms of outcomes, 32 cats (37.6%) were classified as survivors, and the remaining 53 cats (62.4%) were classified as fatal. Of the 85 cats included in this study, 68 had outdoor access or were free-roaming. All 85 (100%) cats showed acute onset of anorexia and lethargy, 28 (32.9%), 1 (1.2%), and 3 (3.5%) cats showed vomiting, diarrhea, and both of vomiting and diarrhea, respectively (Table 1). The infection route for each cat was not determined in this study.

Table 1. Demographic characteristics of cats with severe fever with thrombocytopenia syndrome (for total population and by clinical outcome).

Characteristic	Total n=85	Survivor n=32	Fatal case n=53	P-value
Sex-number (%)				0.521
Male	50 (58.8)	19 (59.4)	31 (58.5)
Female	30 (35.3)	10 (31.2)	20 (37.7)
Unknown	5 (5.9)	3 (9.4)	2 (3.8)
Age-number (%)				0.179
1 year	10 (11.8)	3 (9.4)	7 (13.2)
1–5 years	47 (55.3)	20 (62.5)	27 (50.9)
6–9 years	16 (18.8)	8 (25.0)	8 (15.1)
≥10 years	9 (10.6)	1 (31.2)	8 (15.1)
Unknown	3 (3.5)	0 (0)	3 (5.7)
Rearing environment-number (%)				0.108
Indoor	1 (1.2)	1 (3.1)	0 (0)
Free-roaming	68 (80.0)	28 (87.5)	40 (75.5)
Unknown	16 (18.8)	3 (9.4)	13 (24.5)
Clinical symptoms-number (%)				0.957
Anorecxia	85 (100)	32 (100)	53 (100)
Lethargy	85 (100)	32 (100)	53 (100)
Vomit	28 (32.9)	10 (31.3)	18 (34.0)
Diarrea	1 (1.2)	0 (0)	1 (1.9)
Vomit and diarrhea	3 (3.5)	1 (3.1)	2 (3.7)

*The χ² test was used to compare between survival and fatal group.

A total of 154 samples from 85 cats were submitted to our laboratory for evaluation in this study. The samples comprised 111 blood samples, and 43 swab samples. A total of 33 samples from 32 survivors collected after day 7 (post-acute phase) were included in this study.

Demographic and clinical pathological data

Neither age nor sex significantly predicted outcomes (survivor vs. fatal case; Table 1). The clinical pathology data for the acute phase (days 0–7) are summarized in Table 2 for the basic panel. Data for individual fatalities and survivors through the acute and post-acute phases are shown as key parameters in Figs. 1

Fig. 2.

Transition in clinical parameters. Data on aspartate aminotransferase (AST), total bilirubin (T-Bil), glucose (GLU), creatine phosphokinase (CPK), serum amyloid A (SAA), and total protein (TP) during the acute and post-acute phases are presented chronologically.

–4. Leukopenia, thrombocytopenia, hyperbilirubinemia, hyperproteinemia, hyperglycemia, and elevated SAA and AST levels were observed in SFTSV-positive cats. All clinical pathology parameters were evaluated for their ability to predict outcome. Fatal cases showed significantly higher T-Bil (P=0.01), CPK (P=0.04), SAA (P=0.002), IL-6 (P=0.005), and TNF-α (P=0.028). No other clinical pathology parameters differed significantly between survivors and fatal cases (Table 2). Survivors showed values within or tended to return to the reference range for all acutely out-of-range parameters, except TP, in the post-acute phase (Figs. 1–3).

Table 2. Laboratory parameters in severe fever with thrombocytopenia virus-positive cats in the acute phase (day 0 to 7 after onset).

Laboratory tests	Reference range	Total		Survival		Fatal		P value††
		n	Median (Min, Max)	n	Median (Min, Max)	n	Median (Min, Max)
Viral load
In blood (copies/mL)	-	74	3.2E+07 (9,993.3, 3.4E+10)	29	1.4E+07 (1.0E+04, 8.1E+08)	45	2.3E+08 (1.1E+05, 3.4E+10)	<0.001
In swab§ (copies/mL)	-	28	1.2E+04 (383, 1.1E+08)	13	5.0E+03 (3.8E+02, 1.1E+08)	15	2.5E+04 (7.9E+02, 5.3E+05)	0.122
Blood test
RBC (×104/μL)	500⁓1,000	66	778 (121, 1,146)	26	782 (121, 1,146)	40	765 (127, 1,122)	0.135
WBC (×/μL)	5,500⁓19,500	77	0 (53, 2,000)	31	3,040 (53, 17,300)	46	1,999 (200, 13,340)	0.121
PLT (×104/μL)	30⁓80	74	2 (0, 15)	31	2 (0, 9)	43	1 (0, 15)	0.602
Biochemical test
T-Bil (mg/dL)	0.0⁓0.1	61	4.5 (0.1, 14.0)	25	3.0 (0.1, 11.3)	36	4.7 (1.1, 14.0)	0.011
CPK (U/L)	69⁓214	36	274.0 (0.6, 2,000**)	17	134.0 (0.6, 2,000**)	19	419.0 (1.7, 2,000**)	0.037
ALT (U/L)	25⁓97	56	56 (21, 2,000**)	24	56 (21, 2,000**)	32	59 (25, 1,000†)	0.247
AST (U/L)	7⁓38	35	78 (24, 1,000†)	15	68 (24, 150)	20	100 (36, 1,000)	0.072
BUN (mg/dL)	19⁓34	21	25.3 (1.2, 231.0)	9	25.3 (11.0, 231.0)	12	25.3 (1.2, 141.0)	0.374
Cre (mg/dL)	0.9⁓2.2	43	1.01 (0.32, 22.6)	20	1.05 (0.78, 22.6)	23	0.94 (0.32, 1.92)	0.156
TP (g/dL)	6.0⁓7.9	32	8.4 (5.3, 11.0)	10	9.4 (7.1, 10.4)	22	8.2 (5.3, 11.0)	0.207
Glu (mg/dL)	6.0⁓7.9	25	211 (18, 420)	6	205 (101, 345)	19	228 (18, 420)	0.356
SAA (μg/mL)	≤0.82	63	70.6 (10.4, 143.6)	23	61.4 (10.4, 112.5)	40	82.4 (37.8, 143.6)	0.002
IgM (OD)	-	82	0.38 (0, 2.38)	32	0.58 (0.02, 2.38)	50	0.34 (0.00, 1.67)	0.028
IgG (OD)	-	82	0.12 (0, 1.98)	32	0.23 (0.00, 1.98)	50	0.09 (0, 1.61	0.019
Cytokines
TNF-α (pg/mL)	-*	33	15.30 (1.03, 492)	13	14.45 (1.03, 27.7)	20	21.95 (2.31, 492)	0.034
IL-1β (pg/mL)	-*	57	7.39 (0, 15,568)	22	0.17 (0, 15,568)	35	16.65 (0, 3,682)	0.083
IL-6 (pg/mL)	-*	41	125.56 (0, 35,727)	15	0 (0, 35,727)	26	312.39 (0, 23,253)	0.005

Red blood cell (RBC), white blood cell (WBC), platelet (PLT), total bilirubin (T-Bil), creatinine phosphokinase (CPK), alanine aminotransferase (ALT), aspartate aminotransferase (AST), blood urea nitrogen (BUN), creatinine (Cre), total protein (TP), glucose (Glu), serum amyloid A (SAA), tumor necrosis factor-α (TNF-α), interleukin-1β (IL-1β), and interleukin-6 (IL-6). *Cut off values were determined as <50 ng/mL for TNF-α, <16 pg/mL for IL-1β, and <31.3 pg/mL for IL-6, in this study. **Maximam value was determined as 2,000. †Maximam value was determined as 1,000. †† Wilcoxon signed-rank test was used to compare between survival and fatal group. §The swab samples comprised oral and rectal swabs.

Fig. 1.

Transition in complete blood counts. Data on red blood cell count (RBC), white blood cell count (WBC), and platelet count (PLT) during the acute and post-acute phases are presented chronologically. Where available, data from multiple blood tests performed during hospitalization are included. The first day on which a clinical abnormality occurred was defined as Day 0. Pink circles indicate fatal cases, and blue triangles indicate survivors.

Fig. 4.

Transition in specific antibodies and viral load in blood and swab samples. Optical density (OD) values are shown for IgM and IgG, and the viral load is shown for blood and swab samples (oral or rectal swab samples), chronologically (relative to the day of onset).

Fig. 3.

Transition in cytokines. Data on interleukin-1β (IL-1β), interleukin-6 (IL-6), and tumor necrosis factor-α (TNF-α) during the acute and post-acute phases are presented chronologically.

Antibody titers

Survivors showed significantly higher SFTS-specific IgM (P=0.03) and IgG (P=0.02) levels than the fatal cases (Table 2, Fig. 4).

SFTSV load (copy number)

We established a new quantification assay for the SFTSV copy number using real-time PCR. The reliability of the assay was confirmed by an intra-laboratory variability below 1.30% for all dilutions and an inter-laboratory variability between 0.30% and 2.41%. We used this assay to determine the SFTSV copy number in blood and swab samples. Fatal cases had significantly higher SFTSV copy numbers in blood samples than survivors (P<0.001), and the median of SFTSV copy number in the fatal cases in the blood samples was 16.8 times higher in basic arithmetic magnitude than that in survivors. Swab samples including both oral and rectal swabs, yielded no significant difference; the median copy numbers in the swab samples were 3.6 × 10⁻⁴ and 1.1 × 10⁻⁴ times lower than those in blood samples in fatal cases and survivors, respectively (Table2, Fig. 4).

Phylogenetic analysis

Partial sequences of the SFTSV S segment were obtained from 63 cats, which allowed us to identify the viral strain. The phylogenetic tree is shown in Fig. 5. There were 55 and 6 viruses clustered within genotypes B2 and A according to A-F genotyping [6], which corresponded to the J1 and C4 genotypes respectively, by Chinese-Japanese genotyping [34]. One clustered within genotype B3 and the other clustered within getnotype E, corresponing to J3 and C1, respectively. There were no statistically significant associations between clinical outcomes and viral genotype.

Fig. 5.

Phylogenic analysis of Severe fever with thrombocytopenia syndrome (SFTS) S segment. Phylogenetic tree based on SFTS virus S segment partial sequences from SFTSV-positive cat specimens. The 63 sequences obtained in this study are shown in bold. The retrieved sequences from GenBank are indicated by the accession number, following isolate name, source, country (prefecture in Japan), and year. White circles indicate survivors, and red circles indicate fatal cases. The strains were classified according to the A-F [6] and C-J genotype [34]. *SFTS virus was isolated from an M segment, as described in our previous study [19].

DISCUSSION

To the authors’ knowledge, this is the first study to evaluate prognostic indicators in a sizable population (n=85) of SFTSV-positive cats from a region in, western Japan, where the virus is endemic. By following a subset of survivors in this population, we obtained evidence to characterize the clinical course of the infection during the acute and post-acute phases. All fatal cases in this study experienced acute death (between days 0–7), which is in line with the analysis period we set to investigating associations with outcomes, based on our previous observation of the acute phase in feline SFTS [19].

As key findings of this study, we identified several parameters that could predict the outcomes (significantly associated with survival or acute death). These parameters were antibody titers (IgM and IgG; lower in fatal cases), viral load in blood (higher in fatal cases), cytokine levels (IL-6 and TNF-α; higher in fatal cases), and some clinical pathology parameters (SAA, CPK, and T-Bil; higher in fatal cases).

SFTS-specific IgM and IgG titers and SFTSV loads are highly plausible prognostic indicators. Lower IgM and IgG titers and high viral loads in fatal cases could indicate animals whose immune response and ability to produce antibodies are overwhelmed by SFTSV. This is consistent with the reported disease mechanisms in fatal cases of human SFTS [13, 26] and other hemorrhagic viruses [5] and previous histological evidence in cats [22]. SFTSV is reportedly present in macrophages and B cell–lineage lymphocytes in fatal human infections, attacking and manipulating cells that initiate an antiviral response [26] in a way similar to the mechanisms reported for other hemorrhagic viruses, such as Ebola [5, 8]. Histologically, SFTSV has been shown to infect B cells in lymphoid organs, including the lymph nodes, spleen and Peyer’s patches [24]. However, it cannot be completely ruled out that the cats included in this study may have had underlying diseases that could have affected their immune response to SFTS.

Interestingly, two of the three cytokines that were investigated appeared to be associated to acute death (high IL-6 and TNF-α; IL-1β was not associated). These findings provide useful evidence for considering the role of cytokine storms,—in which a protective inflammatory response is converted into a detrimental response characterized by vascular permeability, tissue destruction, and organ dysfunction [20] — in fatal feline SFTS. Cytokine storms may occur in fatal human SFTS [26], where IL-6, IL-1β and TNF-α are three of the 17 elevated cytokines or chemokines that appear to predict a fatal outcome [3, 13, 25]. Our results suggest that cytokine elevation may be associated with disease severity in feline SFTS and is a fruitful area of research for improving prognostic predictions and possible therapeutic approaches. The measurement of cytokine panels may be useful in assessing the severity of SFTS in cats after a confirmed diagnosis and in predicting prognosis.

Another interesting finding was the acute phase protein SAA, which was higher in fatal cases. SAA is widely used as a prognostic marker for neoplastic or inflammatory diseases in feline medicine (and more widely in multiple species including humans) [30]. In SFTS, increased cytokine levels are thought to cause high levels of SAA. The elevated levels in this study clearly suggest its utility for the prognostic monitoring of feline SFTS and are consistent with a recent report suggesting an association with fatal outcomes in human SFTS [7, 15].

The other predictive clinical parameters in this study (CPK and T-Bil) illustrated both the similarities and contrasts between feline and human SFTS. CPK levels are significantly elevated soon after the onset of SFTS in human patients [2]. The CPK is primarily used in veterinary medicine to assess skeletal muscle damage. However, an increase in CPK activity can be observed as a non-specific finding in unwell and anorexic cats. High serum CPK activity in anorexic cats is postulated to be result from muscle catabolism rather than from muscle necrosis or inflammation [1, 4]. Although the presence of acute viral myositis associated with SFTS viral infection in cats is unknown, it was significantly elevated in more severe (fatal) cases in our study. Jaundice or hyperbilirubinemia is frequently observed in cats with SFTS [19, 20, 22], however, the specific mechanism of hyperbilirubinemia in feline SFTS has not been clarified in previous pathological reports [19, 21, 23]. T-Bil was the only liver-related parameter associated with a fatal outcome in cats in our study; however, fatal human SFTS is characterized by liver damage secondary to shock, hypercytokinemia, or hemophagocytosis, and progressively elevated ALT and AST level until death [7]. SFTS progresses faster in cats than in humans; thus, hepatic abnormalities may not manifest before death from feline disease. Jaundice is often prominent in feline SFTS, suggesting a role of adverse effects on the liver in determining the outcome; a suggestion supported by the elevated T-Bil levels in fatal cases in this study. The mechanism for such hepatic effects is unclear; however, T-Bil values may better reflect the severity of feline SFTS than other liver-related parameters.

In this study, a subset of the sample of survivors was monitored for up to four weeks after the initial onset of the disease. These results produced important observations regarding the clinical and virological profile of SFTS in feline cases. The overall population of SFTSV-positive cats was highly viremic, with several parameters departing from the reference ranges (WBC, PLT, T-Bil, Glu, and SAA). The target cells infected with SFTSV were reported to be B cells, which impair the host immune response to invading pathogen, and may lead to a decrease in white blood cells [17, 19]. Thrombocytopenia is a hallmark of SFTSV infection in human and cats with SFTSV infection, which is thought to be involved in several mechanisms, such as increased consumption of peripheral platelets from virus-induced activation of the coagulation pathway, or by clearance of SFTSV-bound platelet by splenic macrophages. Acute inflammation due to viral infection is thought to be associated with hyperglycemia and high SAA. When subsequent samples were submitted from survivors, we were able to perform further analyses, and found that the relevant values returned to the reference range or were moving towards the reference range in the post-acute phase, demonstrating signs of clinical recovery.

No one in contact with SFTSV-infected cats, including animal owners and veterinarians showed any SFTS like symptoms. We monitored 20 survived cats and 5 cats showed positive SFTSV RNA, even day 14 after onset. These findings are important from a One Health perspective, as the risk of transmission of this zoonosis is not eliminated when the cat enters the recovery phase. Despite the lack of a definitive correlation between viral genotypes and difference in disease outcomes following natural SFTSV infection, caution is warrantied for any SFTSV infection.

In conclusion, this study demonstrates that SFTS-specific IgM and IgG, IL-6, TNF-α, T-Bil, and CPK are potential prognostic indicators for feline SFTS at the acute stage. Our findings highlight the importance of anti-infection measures beyond the acute phase in SFTSV-positive cats.

CONFLICT OF INTEREST

The authors declare no conflicts of interest associated with this manuscript.