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. 2026 May 4;49(10):e70193. doi: 10.1111/jfd.70193

Identification and Characterization of Flavobacterium psychrophilum as a Causative Pathogen in Farmed Grass Carp Ctenopharyngodon idella in China

Jiahao Li 1,2, Lingyun Lin 2, Jinyu Shen 2, Jiayun Yao 2, Xiao Liu 1,✉, Xiaoyi Pan 2,✉
PMCID: PMC13532608  PMID: 42083140

ABSTRACT

In March 2024, a severe mass mortality event occurred at a grass carp ( Ctenopharyngodon idella ) farm in Zhejiang, China, resulting in a cumulative mortality rate of 70%. To identify the causative agent, 12 bacterial isolates that exhibited the same phenotypic characteristics were isolated from diseased fish. A further phylogenetic (16S rDNA) characterization identified the bacterium as Flavobacterium psychrophilum . Genetic characterization revealed that all 12 bacterial isolates belonged to serotype Type‐1 and sequence type ST418, a newly documented ST closely associated with cyprinid fish hosts. In infection trials, the bacterium exhibited clear virulence in grass carp, producing clinical signs consistent with those observed during the outbreak. Furthermore, histopathology of diseased fish with F. psychrophilum revealed ulcerative dermatitis with associated myositis, proliferative branchitis, vascular congestion and focal necrosis in the liver, multifocal necrosis in the spleen, and tubular dilation with renal corpuscle necrosis in the kidney. Given the potential risk of cross‐species transmission among cyprinid fish, we also evaluated the pathogenicity of this bacterium in Prussian carp ( Carassius gibelio ), a widely cultured species in China. To our knowledge, this study represents the first report of F. psychrophilum causing mass mortality in farmed grass carp in China and provides essential information to support improved prevention and risk mitigation strategies for this emerging disease.

Keywords: antimicrobial susceptibility, Ctenopharyngodon idella , Flavobacterium psychrophilum , genotype, pathogen identification

1. Introduction

Flavobacterium psychrophilum is a Gram‐negative, rod‐shaped bacterium commonly found in freshwater environments and is well adapted to low temperatures (Madetoja et al. 2003; Duchaud et al. 2007; Barnes and Brown 2011; Knupp et al. 2024). It is best known as the causative agent of bacterial cold‐water disease and rainbow trout fry syndrome, which together result in substantial economic losses in salmonid aquaculture worldwide (Nematollahi et al. 2003; Nicolas et al. 2008; Rochat et al. 2017; Li et al. 2021; Saticioglu et al. 2025). The pathogen also significantly affects both farmed and wild populations of ayu Plecoglossus altivelis Temminck & Schlegel (Lee and Hee 1998; Imajoh 2022). Nearly all salmonid species are susceptible to infection, with rainbow trout Oncorhynchus mykiss (Walbaum), coho salmon Oncorhynchus kisutch (Walbaum) and Atlantic salmon Salmo salar Linnaeus being among the most vulnerable hosts (Nematollahi et al. 2003). Typical clinical signs include the development of characteristic external ulcerative lesions, particularly around the caudal peduncle and caudal fin (Starliper 2011). Additional symptoms may include exophthalmia, abdominal distension accompanied by increased ascitic fluid, and pale gills (Nematollahi et al. 2003; Barnes and Brown 2011; Starliper 2011; Jarau et al. 2018). The bacterium has also been reported to exhibit tropism for the lower jaw, fins and caudal peduncle (Kondo et al. 2002; Martínez et al. 2004; Papadopoulou et al. 2017). Outbreaks in salmonids generally occur at water temperatures between 3°C and 15°C (Nematollahi et al. 2003), whereas disease in ayu is more common at 14°C–21°C (Iida and Mizokami 1996; Imajoh 2022).

Flavobacterium psychrophilum has also been sporadically isolated from or detected in various non‐salmonid fish hosts. Reported cases include species from Cyprinidae, Anguillidae, Osmeridae, Gobiidae, Oxudercidae, Percidae, Centrarchidae, Petromyzontidae, Acipenseridae, Clariidae and Amblycipitidae (Lehmann et al. 1991; Amita et al. 2000; Madetoja et al. 2002; Izumi et al. 2003; Elsayed et al. 2006; Verma and Prasad 2014; Fujiwara‐Nagata et al. 2019; Soares et al. 2019; Chinchilla et al. 2023). Although F. psychrophilum lacks apparent host specificity, in non‐salmonid species (excluding ayu) the disease is significantly less severe, and to date there have been no documented cases of F. psychrophilum infection causing mass mortality in grass carp ( Ctenopharyngodon idella ) (Valenciennes).

Grass carp is the most widely cultured freshwater fish species in China. In 2023, national production reached 5.94 million tons, representing 21.30% of China's total freshwater fish yield (Bureau of Fisheries of the Ministry of Agriculture of the People's Republic of China 2024). Grass carp has also been introduced to multiple regions across Asia, Europe, the Americas and Africa, and is now a key aquaculture species in countries such as Vietnam and in regions including Western Siberia (Lin et al. 2022; Liu et al. 2023). However, the increasing scale and intensification of farming practices have led to more frequent and complex disease outbreaks. In recent decades, prevalent diseases in grass carp have included hemorrhagic disease caused by grass carp reovirus (Rao and Su 2015; Zhang et al. 2021), bacterial gill‐rot associated with F. columnare (Luo et al. 2016) and bacterial septicemia caused by Aeromonas hydrophila and A. sobria (Lv et al. 2025). However, since 2019, a newly emerging disease has been observed in farmed grass carp during late winter and early spring. Affected fish exhibit marked hemorrhage in the snout–jaw–cheek region, accompanied by cutaneous ulcerations at various body sites, with cumulative mortality of 50%–80%. Notably, outbreaks occur at water temperatures of 5°C–16°C, which differ substantially from the 20°C to 30°C temperatures typically associated with the aforementioned diseases. Moreover, the known pathogens responsible for those conditions have not been detected in these cases, indicating the involvement of an uncharacterized etiological agent. This underscores the need for further pathogen isolation and identification.

Therefore, this study aimed to identify the causative agent responsible for the emerging disease in grass carp and to investigate its phenotypic and genotypic characteristics. To this end, 12 bacterial isolates obtained from diseased grass carp were identified as F. psychrophilum based on phylogenetic analysis of the 16S rRNA gene and biochemical characterization. These isolates were further subjected to genotypic and serotypic analysis, and a representative isolate, CYL302‐1, was selected for subsequent investigations on physiological characteristics, pathogenicity and antimicrobial susceptibility. To the best of our knowledge, this is the first report identifying F. psychrophilum as a pathogen in grass carp, providing essential information for the prevention and control of this disease.

2. Materials and Methods

2.1. Sample Collection

In March 2024, 12 diseased grass carp (100–2000 g) and 10 healthy grass carp (100–2000 g) were collected from a farm in Zhejiang Province. The fish were transported to the laboratory within 1–2 h in oxygen‐supplemented bags at ambient temperature (12°C–15°C). Prior to dissection, the fish were anaesthetised with MS‐222 solution (100 mg/L; Solarbio, China) to assess pathological changes in internal organs. Six of the diseased fish were randomly selected and used for viral detection, bacterial isolation, parasitological examination and histopathological analysis.

2.2. Histopathology Examination

Ulcerated skin with underlying muscle, gill, liver, spleen, kidney and intestinal tissues were collected from the six selected diseased fish and immediately fixed in 4% paraformaldehyde. Following dehydration through an ethanol gradient and clarification with xylene, tissues were embedded in paraffin. Paraffin blocks were sectioned into 4 μm slices using a microtome, deparaffinized, rehydrated and stained with haematoxylin and eosin (H&E) or subjected to Gram staining. Histopathological lesions and the presence of bacterial cells in necrotic skin were examined under a light microscope and compared with tissues from healthy controls.

2.3. Pathogen Screening

2.3.1. Parasite Detection

Wet‐mount preparations were made from the gill filaments, caudal fin or pelvic fin and surface mucus of diseased fish. Parasitic infection was evaluated microscopically at 40–100× magnification.

2.3.2. Virus Detection

The spleen and kidney tissue (200 mg) from each fish were pooled, and total RNA was extracted using the RNAsimple Total RNA Kit (Tiangen, China). Grass carp reovirus nucleic acid was detected by RT‐PCR following the method of Zeng et al. (2013), targeting genotypes I, II and III. For viral infectivity assessment, spleen and kidney tissues from six diseased fish were homogenized, diluted 5–10‐fold in PBS, and sequentially filtered through 1.0, 0.45 and 0.22 μm filters (MF‐Millipore, Merck). Healthy grass carp (150 ± 25 g) were allocated into two groups (10 fish/group). The experimental group received 0.2 mL of filtered homogenate by intraperitoneal injection, and the control group received the same volume of PBS. Post‐injection mortality for 28 days was monitored to determine the presence of any additional viral pathogens.

2.3.3. Bacterial Isolation

The external surface of each diseased fish was disinfected with alcohol. Under sterile conditions, tissues from the liver, spleen, kidney and external lesions were streaked onto trypticase soy agar (TSA), brain heart infusion agar (BHI) and blood agar plates (TSA supplemented with 5% defibrinated sheep blood; Hopebio, China) to isolate non‐fastidious bacteria. Additionally, 200 mg of each tissue was placed into a 1.5 mL Eppendorf tube containing 200 μL of PBS. The tissue was homogenized on ice using an electric grinder for 30–60 s, followed by the addition of 600 μL of PBS and thorough mixing. An aliquot of 100–200 μL of the homogenate was then spread onto Shieh agar (Shanghai Ruichu Biotech Co. Ltd., China). Plates were incubated at 15°C for 5–7 days. The predominant colonies were sub‐cultured for purification and subsequently stored in the same liquid medium used for isolation and purification, supplemented with 20% (v/v) glycerol, at −80°C until further use.

2.4. Pathogen Identification

2.4.1. 16S rDNA Molecular Identification

The molecular identification of one representative isolate (CYL302‐1) was performed using standard 16S rRNA gene sequencing (Weisburg et al. 1991). A 2 mL aliquot of fresh CYL302‐1 culture was collected, and genomic DNA was extracted using a bacterial genomic DNA extraction kit (Tiangen, China) according to the manufacturer's instructions. The almost complete 16S rRNA gene was amplified by PCR using universal primers 27F (AGAGTTTGATCCTGGCTCAG) and 1492R (GGTTACCTTGTTACGACTT) (Weisburg et al. 1991). The 50 μL PCR reaction mixture contained 25 μL 2× Premix Taq (TaKaRa, Dalian, China), 2 μL primer pair (10 μM), 1 μL DNA template (20–100 ng), and nuclease‐free water. The amplification conditions were: initial denaturation at 94°C for 5 min; followed by 35 cycles of denaturation at 94°C for 30 s, annealing at 54°C for 30 s, and extension at 72°C for 100 s; with a final extension at 72°C for 10 min. PCR products were verified on a 1.0% agarose gel stained with Ultra GelRed (Vazyme, China) and subsequently sequenced by ShangYa Biotech Co. Ltd. The obtained 16S rRNA gene sequence was compared with reference sequences in the NCBI database using the Basic Local Alignment Search Tool (BLAST). Multiple sequence alignment of highly similar strains retrieved from GenBank was conducted using the ClustalW algorithm in MEGA 12 (Kumar et al. 2024). A phylogenetic tree was constructed using the Maximum Likelihood method with the Tamura–Nei model and evaluated with 1000 bootstrap replicates in MEGA 12. Chryseobacterium piscium served as the outgroup.

2.4.2. Morphological Observation and Biochemical Tests

Bacterial morphology and biochemical properties of the 12 purified bacterial isolates (CYL302‐1 to CYL302‐12) were performed using conventional methods as described by Bernardet (1989). For morphological studies, Gram staining, colony morphology, growth on several agar media, and gliding motility were examined. Furthermore, one representative isolate (CYL302‐1) was selected for scanning and transmission electron microscopy. For biochemical tests, the following characteristics of the 12 isolates were analysed including production of flexirubin‐type pigments, Congo red absorption, respiratory metabolism (cytochrome oxidase and catalase activities, nitrate reduction, hydrogen sulphide production), carbohydrate metabolism (β‐galactosidase activity, degradation of carboxymethyl cellulose, chitin, starch and esculin; acid production from glucose, mannose, fructose, galactose, etc.), lipid metabolism (degradation of lecithin, tween 20 and tween 80), protein metabolism (degradation of gelatin, casein and tyrosine; activities of ornithine decarboxylase, lysine decarboxylase and arginine dihydrolase), DNase production and susceptibility to the vibriostatic compound O/129 (Bernardet 1989; Lehmann et al. 1991). All assays were performed on enriched Anacker–Ordal (EAO) agar or EAO broth (Oplinger and Wagner 2012) and were incubated at 15°C ± 1°C. Additionally, F. psychrophilum type strain DSM 3660 was used as a control strain for this phenotypic study.

2.4.3. Physiological Characteristics

The growth tests of the representative isolate CYL302‐1 to temperature, NaCl concentration and pH was studied as followed. For solid cultures, 5 μL of bacterial culture in logarithmic phase (OD525 ≈ 0.5) was inoculated onto EAO agar plates. Colony diameters were measured after 5 days of incubation at different temperatures (0°C–25°C). For liquid cultures, 0.5 mL of bacterial culture in logarithmic phase was inoculated into flasks containing 50 mL EAO broth and incubated at different temperatures (2°C–25°C) in a shaking incubator (200 rpm) for 6 days. OD525 was recorded every 12 h to generate growth curves.

To assess the effects of salinity and pH, NaCl was added to EAO broth to obtain final concentrations of 0%–2.0% (w/v), and the pH was adjusted to 4.0–9.0 using HCl or NaOH. Cultures were incubated at 15°C in a shaking incubator (200 rpm) for 5 days, and growth was monitored by OD525. All assays were performed in triplicate. Statistical differences among groups were analysed by one‐way ANOVA using GraphPad Prism version 9.5.1, followed by Tukey's post hoc test for multiple comparisons.

2.5. Experimental Infection

Healthy grass carp (150 ± 25 g) were obtained from a fish farm in Huzhou, Zhejiang Province, and maintained in a recirculating system at 15°C ± 2°C for at least 10 days for acclimation. Prior to challenge, three fish were randomly selected and tested using standard microbiological procedures to confirm the absence of bacterial infection. The remaining grass carp were assigned to three groups (two treatment groups and one control group, n = 10 per group). The bacterial concentration of the logarithmic‐phase culture of isolate CYL302‐1 was estimated based on a pre‐established standard curve correlating OD525 with colony‐forming units (CFU). The suspension was then diluted with sterile pond water to final concentrations of 6.0 × 108 and 6.0 × 107 CFU/mL, respectively. The two treatment groups were intraperitoneally injected with 0.2 mL of the bacterial suspension per fish, respectively, whereas the control group received the same volume of sterile pond water.

To further evaluate pathogenicity toward Prussian carp Carassius gibelio (Bloch) (a widely cultured cyprinid species in China), a second infection trial was conducted. The healthy Prussian carp (125 ± 25 g) were maintained in a recirculating system at 15°C ± 2°C for at least 10 days for acclimation. Prior to the challenge, three fish were randomly selected for pathogen detection, including bacterial isolation and qPCR detection of CyHV‐2 (Goodwin et al. 2006), to confirm their health status. Twenty healthy Prussian carp were randomly selected and evenly divided into two groups: one group was intraperitoneally injected with 0.2 mL of 1.2 × 107 CFU/mL bacterial suspension per fish, whereas the control group received the same volume of sterile pond water.

Prior to pathogen detection and challenge experiments, fish were anaesthetised with MS‐222 (50–100 mg/L, 1–2 min). Fish in each group were maintained in a 200 L aquarium with continuous aeration at 15°C ± 2°C, and one‐third of the water volume was replaced with fresh, oxygenated water every other day. No feeding was conducted during the challenge period. Disease signs and mortality were recorded daily for 25 days post‐challenge. Dead fish were dissected, and kidney tissues were streaked on EAO agar for bacterial isolation and cultured to confirm the cause of death. After incubation at 15°C for 5–7 days, yellow colonies were identified by F. psychrophilum‐specific real‐time PCR assay (Strepparava et al. 2014) and partial isolates were further confirmed by 16S rRNA gene sequencing. Survival curves were generated using GraphPad Prism version 9.5.1.

2.6. Multiplex PCR Serotyping

The molecular serotype of all 12 isolates (CYL302‐1 to CYL302‐12) was determined following the serotyping scheme for F. psychrophilum Types 0–3 described by Rochat et al. (2017), with additional identification of serotype 4 based on Avendaño‐Herrera et al. (2020). The multiplex PCR was performed using the Multiplex PCR Assay Kit (TaKaRa, Dalian, China) with the following reaction mixture: 25 μL of 2× Multiplex PCR Buffer, 0.25 μL of Multiplex PCR Enzyme Mix, 2 μL of each of the five primer sets (10 μmol/L), 5 μL of DNA template (20–100 ng), and nuclease‐free water to a final volume of 50 μL. PCR cycling conditions were: initial denaturation at 95°C for 5 min; followed by 30 cycles of 95°C for 30 s, 52°C for 30 s, and 72°C for 60 s; and a final extension at 72°C for 10 min. PCR products were separated on a 1.2% agarose gel stained with Ultra GelRed (Vazyme, China) to determine fragment sizes. The 188 bp amplicon served as an internal positive control. Strains displaying only the 188 bp band were classified as serotype 0, whereas serotypes 1–4 were identified based on characteristic band patterns: serotype 1 (188 and 549 bp), serotype 2 (188 and 841 bp), serotype 3 (188 and 361 bp) and serotype 4 (188 and 992 bp).

2.7. Multilocus Sequence Typing (MLST)

For MLST of all 12 isolates (CYL302‐1 to CYL302‐12), the typing scheme of Nicolas et al. (2008) was applied, which targets partial sequences of seven housekeeping genes (atpA, dnaK, fumC, gyrB, murG, trpB and tuf). Each PCR reaction contained 25 μL of EmeraldAmp MAX PCR Master Mix (2× Premix), 2 μL of each primer (10 μmol/L), 5 μL of DNA template (20–100 ng) and nuclease‐free water to a final volume of 50 μL. Cycling conditions were as follows: initial denaturation at 94°C for 3 min; followed by 35 cycles of 94°C for 30 s, 52°C for 30 s and 72°C for 60 s; with a final extension at 72°C for 8 min. PCR products were verified by electrophoresis, purified, and submitted to ShangYa Biotech Co. Ltd. for bidirectional sequencing. Chromatograms were manually inspected, and allele types (ATs) were assigned by comparison with entries in the PubMLST database (https://pubmlst.org/fpsychrophilum/; accessed April 2025). Finally, the sequence type (ST) was determined based on the combination of the seven ATs.

2.8. Antimicrobial Susceptibility Testing

Antimicrobial susceptibility testing of the representative isolate CYL302‐1 was conducted according to the broth microdilution method recommended in the Clinical and Laboratory Standards Institute (CLSI) guidelines VET03 (CLSI 2020a), using diluted cation‐adjusted Mueller‐Hinton broth (CAMHB) (Hopebio, China) at 18°C ± 2°C and a bacterial suspension equivalent to 0.5 McFarland. Minimum inhibitory concentrations (MICs) were determined for enrofloxacin, erythromycin, florfenicol, flumequine, oxytetracycline and oxolinic acid, all purchased from Aladdin (Shanghai, China). Antibiotic preparation, dilution procedures, and quality control ranges followed VET03 guidelines. Escherichia coli ATCC 25922 and Aeromonas salmonicida subsp. salmonicida ATCC 33658 served as quality control strains. Susceptibility interpretations for CYL302‐1 were based on epidemiological cutoff values (ECVs) provided in CLSI VET04 (CLSI 2020b).

2.9. Identification of Mutations in the Quinolone Resistance Determining Regions (QRDRs)

To determine whether quinolone resistance in isolate CYL302‐1 involved amino acid substitutions in QRDRs of DNA gyrase (GyrA, GyrB) and topoisomerase IV (ParC, ParE), PCR amplification was performed using primers specific for F. psychrophilum QRDRs (Izumi and Aranishi 2004; Shah et al. 2012). Amplicons were sequenced and compared with the reference strain NCIMB 1947 (GenBank ID: CP007207).

3. Results

3.1. Clinical Symptoms of Diseased Fish

The most common clinical presentation in diseased grass carp was severe haemorrhaging around the snout and cheek (Figure 1A,B). In addition, skin lesions characterized by raised, pus‐filled vesicles (containing light red blood or serous fluid resulting from muscle necrosis) and cutaneous ulcerations at various body sites—particularly in the caudal region—were frequently observed (Figure 1C–E). Some affected fish also exhibited pronounced ascites, pale liver, enlarged spleen, and numerous red petechiae across the mesentery (Figure 1F). The disease typically occurred when water temperatures ranged from 9°C to 15°C, affecting grass carp weighing between 100 and 2000 g. Approximately 80% of fish developed visible clinical signs, and the cumulative mortality reached up to 70% within 15 days of onset.

FIGURE 1.

FIGURE 1

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The clinical signs of diseased grass carp. (A) Snout hyperemia and redness. (B) Sunken orbit, exophthalmos, and periorbital depigmentation. (C) Ulceration of the caudal fin. (D) Scale loss with epidermal congestion and mild ulceration. (E) Cutaneous pustules and extensive superficial skin ulceration along the trunk (from the anal fin to the caudal fin), with increased mucus on the caudal fin. (F) Petechiation of the mesentery, and mild splenomegaly.

3.2. Histopathology Examination

Histopathological examination revealed marked lesions in multiple organs. The skin in the ulcerated regions exhibited partial to complete loss of the epidermal layer (Figure 2A), with numerous rod‐shaped bacteria occupying the exposed dermal layer (Figure 2B). Adjacent skeletal muscle showed myofiber degeneration and lytic necrosis, with an inflammatory infiltrate composed of lymphocytes and occasional granulocytes (Figure 2C). Myofiber degeneration was observed in five of the six examined fish; the remaining fish had no external ulcerative lesions and was not sampled for skin or muscle tissue. In the gills, diffuse epithelial hyperplasia resulted in fusion of the secondary lamellae, accompanied by prominent lamellar mucous cells (Figure 2D). The hyperplastic areas exhibited a mild inflammatory infiltration of granulocytes characterized by eosinophilic cytoplasm and irregular nuclei (Figure 2E). These gill lesions were observed in three of the six examined fish. The liver presented vascular congestion and focal necrotic areas, with hepatocytes displaying karyopyknosis and karyolysis (Figure 2F). Renal changes included scattered necrosis of renal corpuscles, the lumina of which contained cellular debris and brownish granular material (Figure 2G), as well as dilation of renal tubules (Figure 2H). The spleen contained multifocal necrosis within the parenchyma (Figure 2I).

FIGURE 2.

FIGURE 2

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Histological analysis of diseased grass carp. Histological sections stained with H&E (A, C–I) or the Gram stain (B). (A) Loss of the epidermal layer (black arrow). (B) Numerous rod‐shaped bacteria in the dermis (black arrow). (C) Myofiber fragmentation and lytic necrosis (blue circle) in skeletal muscle adjacent to ulcer, with an inflammatory infiltrate composed of lymphocytes (green arrow) and occasional granulocytes (black arrow). (D) Diffuse hyperplasia of gill filament epithelium (green arrow) and increased mucous cells (blue arrow); blood vessels entering the gill (black arrows). (E) Infiltration of eosinophilic inflammatory cells (red arrow) in the fused secondary lamellae. (F) Vascular congestion (green circle) and necrotic foci (red circle) in liver; hepatocytes showing karyopyknosis (blue arrow) and karyolysis (red arrow). (G) Abnormal renal corpuscle with cellular necrosis and dissolution; cellular debris and brownish granular material within the corpuscular lumen (red arrow). (H) Dilated and abnormal renal tubules (blue arrow) compared to normal tubules (red arrow). (I) Necrotic foci in spleen (green circle); red blood cells (red arrow); lymphocytes and immature blood cells (green arrow).

3.3. Pathogen Screening

No parasites were observed, or only a few were present, in the gills, caudal fin and skin mucus of the diseased grass carp as examined under an optical microscope. Additionally, RT‐PCR testing did not detect grass carp reovirus. When sterile homogenates prepared from tissues of the diseased fish were injected into healthy grass carp, no clinical signs or mortality occurred.

No dominant bacterial growth was observed on TSA, BHI or blood agar plates. In contrast, a dominant colony type with consistent morphology was isolated on 12 of 20 Shieh agar plates (Figure 3A). A total of 12 isolates were obtained from the tissues of all six diseased fish (five isolates from kidney, three from spleen, two from liver and two from external lesions) (Table S1). These were designated as isolates CYL302‐1 through CYL302‐12. The purified predominant colonies were yellow, smooth, moist and circular with regular margins (Figure 3B). Gram staining showed that the bacteria were Gram‐negative. Cells measured 1–5 μm in length and 0.3–0.5 μm in width, and neither flagella nor fimbriae were observed (Figure 3C,D).

FIGURE 3.

FIGURE 3

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Bacterial isolation using Shieh agar and observation of microbial morphology. (A, B) Colony morphology on Shieh agar plates. (C, D) Electron microscopy of cell morphology (C: Scanning electron microscopy, SU8010, ×20,000; D: Transmission electron microscopy, 80 kV, ×8000).

3.4. Pathogen Identification

3.4.1. Phylogenetic Analysis Based on 16S rDNA

The 16S rRNA gene sequence of the representative isolate CYL302‐1 has been deposited in GenBank under accession number PX419286 (www.ncbi.nlm.nih.gov/genbank). BLAST analysis revealed 100% identity with F. psychrophilum reference strain 160401‐1/5N (GenBank ID: CP039120.2; 957536–959008). A phylogenetic tree comparing CYL302‐1 with closely related taxa confirmed that the isolate clusters within the F. psychrophilum clade (Figure 4).

FIGURE 4.

FIGURE 4

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Phylogenetic tree of the representative isolate CYL302‐1 based on 16S rDNA sequences using the Maximum Likelihood method, with Chryseobacterium piscium as the outgroup. Numbers next to branches indicate bootstrap values based on 1000 replicates. Sequence accession numbers are shown in parentheses next to strain names, and scale bars represent genetic distance.

3.4.2. Biochemical Characteristics

Physiological and biochemical characteristics of 12 isolates from diseased grass carp are summarized in Table 1. All 12 isolates exhibited identical phenotypic profiles and were consistent with those described for F. psychrophilum (Bernardet et al. 1996). Results from the F. psychrophilum type strain DSM 3660 were also included.

TABLE 1.

Phenotypic characteristics of grass carp's isolates and control strain.

Characteristics Results Characteristics Results
1 2 3 a 1 2 3 a
Morphology of colonies on AOA Low convex, round, with entire margins Low convex, round, with entire margins Low convex, round, with entire or uneven margins Acid produced from
Congo red absorption − − − Glucose − − −
Growth on seawater media − − − Lactose − − −
Growth on nutrient agar − + V Maltose − − −
Growth on trypticase soy agar − − − Mannitol − − −
Gliding motility (+) + (+) Mannose − − −
Flexirubin type of pigments + + + Sucrose − − −
Glucose used as a sole carbon and energy source − − − Xylose − − −
Degradation of Fructose − − −
Gelatin + + + Rhamnose − − −
Casein + + + Galactose − − −
Starch − − − Trehalose − − −
Carboxymethyl cellulose − − − Production of
Chitin − − − Arginine dihydrolase − − −
Esculin − − − Lysine decarboxylase − − −
DNA + + (+) Ornithine decarboxylase − − −
Lecithin + + + β‐Galactosidase − − −
Tween 20 + + + Cytochrome oxidase (+) + V
Tween 80 + + + Catalase (+) + (+)
Tyrosine + − v H2S − − −
Brown diffusible pigment produced on tyrosine agar − − − Susceptibility to vibriostatic compound O/129 + + +
Nitrate reduction − − −
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Note: 1, DSM 3660 strain; 2, twelve grass carp isolates; 3, Flavobacterium psychrophilum .

Abbreviations: (+), weakly positive; −, negative; +, positive; V, variable among references; v, variable among strains.

a

Data from Bernardet et al. (1996).

3.4.3. Physiological Characteristics

The growth curve of the representative isolate CYL302‐1 in EAO broth is shown in Figure 5A. The isolate exhibited significant temperature‐dependent growth. Optimal growth occurred at 15°C, both in terms of growth rate and final biomass. Growth at 10°C and 18°C was slightly reduced, but the growth curves remained similar to that observed at 15°C, with a lag phase of 0–36 h followed by a logarithmic phase from 36 to 72 h. At 10°C, the stationary phase was extended by 24 h relative to 15°C and 18°C. At 5°C, the lag phase extended to 48 h and the growth rate was slower, reaching stationary phase at approximately 108 h; however, the maximum cell density was comparable to that at 15°C. No growth occurred at 2°C, 20°C or higher temperatures in liquid culture. Interestingly, the isolate demonstrated broader temperature tolerance on EAO agar plates, with growth observed up to 20°C–23°C (Figure 5B).

FIGURE 5.

FIGURE 5

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Growth test of Flavobacterium psychrophilum CYL302‐1. (A) Growth curves of CYL302‐1 at different temperatures. (B) Effect of temperature on colony diameter expansion. Statistical significance was determined versus the 18°C group. (C) Growth of CYL302‐1 at different NaCl concentrations. Statistical significance was determined versus the 0 concentration group. (D) Growth of CYL302‐1 at different pH values. Statistical significance was determined versus the pH 7.0 group. Data are presented as mean ± SD (n = 3). ****p < 0.0001; ***p < 0.001; **p < 0.01; *p < 0.05; ns, not significant.

The salinity test results (Figure 5C) indicated that the isolate is highly sensitive to salinity. Growth was significantly inhibited at NaCl concentrations of 0.1%–0.3%, with a marked reduction in bacterial proliferation. At salinities of 0.5%–1.0% NaCl, growth was very weak, and no growth occurred when the NaCl concentration exceeded 1.0%.

The pH test results (Figure 5D) showed that CYL302‐1 can grow within a pH range of 6.0–8.0, with cell density slightly decreased at pH 6.0. No growth was observed at pH values < 6.0 or > 8.0.

3.5. Experimental Infection

In the 1.2 × 108 CFU dose group, the grass carp developed darkened body coloration, slight congestion around the snout, cheeks and eyes, and a red, swollen, and protruding anus by Day 3 after infection. By Day 5, mortality began to occur. In the later stages (14–21 dpi), the internal organs, particularly the kidneys, showed distinct histolysis, with progressive loss of structural integrity and indistinct anatomical boundaries. In the 1.2 × 107 CFU dose group, the grass carp exhibited symptoms similar to natural infections, including snout congestion, and some fish displayed spiral swimming within 5–10 days after infection. However, the symptoms of most diseased fish (4/6) gradually subsided over time. As shown in Figure 6, the final mortality rates for the two dose groups were 80.0% and 20.0%, respectively. No disease or mortality occurred in the control group. Koch's postulates were fulfilled, as pure bacterial cultures were re‐isolated from the tissues of the infected grass carp and identified as F. psychrophilum through 16S rDNA sequencing.

FIGURE 6.

FIGURE 6

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Survival curves of grass carp and Prussian carp challenged with CYL302‐1 via intraperitoneal injection. For grass carp, two infection doses were tested: 1.2 × 108 CFU/fish and 1.2 × 107 CFU/fish. For Prussian carp, a single dose of 1.2 × 107 CFU/fish was used. Both control groups were injected with sterile pond water.

For Prussian carp, at the dose of 1.2 × 107 CFU/fish, severe disease symptoms developed, including extensive congestion from the snout to the lower jaw, around the cheek, accompanied by scale loss and slight body ulceration, leading to a 70% mortality rate. No disease symptoms or mortality were observed in the control group (Figure 6).

3.6. Genetic Characterization

The mPCR results showed that all 12 isolates produced two target bands (188 bp and 549 bp), indicating they all belong to serotype Type‐1. MLST analysis confirmed that all isolates possessed identical allele types at the seven loci (atpA, dnaK, fumC, gyrB, murG, trpB and tuf), which were assigned as 8, 8, 6, 108 (newly assigned), 8, 5 and 89 (newly assigned), respectively. This unique and consistent allelic profile defines a novel sequence type, ST418, for the outbreak strain. All sequence data and background information for the representative isolate (CYL302‐1) have been submitted to the PubMLST database (https://pubmlst.org/organisms/flavobacterium‐psychrophilum).

3.7. Drug Sensitivity and Mutations in the QRDRs

Based on the epidemiological cutoff values for F. psychrophilum in CLSI VET04 (2020b), the susceptibility of CYL302‐1 to six antimicrobial agents was determined (Table 2). The isolate was classified as wild‐type (WT) for erythromycin, tetracycline, florfenicol, non‐wild‐type (NWT) for enrofloxacin, flumequine and norfloxacin. Using the nucleotide and corresponding amino acid sequences of the quinolone‐sensitive reference strain NCIMB1947, mutations in the QRDRs of CYL302‐1 were analysed. The results showed a T82A substitution in GyrA, whereas no mutations were detected in the QRDRs of GyrB, ParC or ParE.

TABLE 2.

Antimicrobial susceptibility pattern of isolate CYL302‐1.

MIC (WT/NWT, μg/mL) of drug
Erythromycin Florfenicol Oxytetracycline Enrofloxacin Flumequine Oxolinic acid
Interpretive categories and MIC ECVs
WT ≤ 8 ≤ 2 ≤ 0.12 ≤ 0.03 ≤ 0.12 ≤ 0.25
NWT ≥ 16 ≥ 4 ≥ 0.25 ≥ 0.06 ≥ 0.25 ≥ 0.5
CYL302‐1 2 (WT) 1 (WT) 0.12 (WT) 0.25 (NWT) 2 (NWT) 8 (NWT)
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Abbreviations: ECV, epidemiological cutoff value; MIC, minimal inhibitory concentration; NWT, non‐wild‐type, manifest a reduced susceptibility compared to those of WT isolates; WT, wild‐type, fully susceptible members of their species.

4. Discussion

Flavobacterium psychrophilum was first isolated in the United States in 1948 (Borg 1960). By the late 1980s and early 1990s, this bacterium had been widely reported in salmonid species and ayu in several European countries and in Japan (Nematollahi et al. 2003). It is now considered one of the most important bacterial pathogens affecting freshwater‐farmed salmonids worldwide. In northern China, several provinces (Qinghai, Gansu, Liaoning and Heilongjiang) have reported diseases in salmonid species caused by F. psychrophilum (Li et al. 2021), whereas such reports remain rare in warmer southern regions or in other fish species. In recent years, however, mass mortality events have frequently occurred in cultured grass carp during late winter and early spring in China, although the causative agent has been unclear. Jiang et al. (2024) reported F. psychrophilum ‐associated disease in Prussian carp, with disease onset and water temperature conditions similar to those observed in the current grass carp outbreak. Therefore, during the spring of 2024, when large‐scale mortality occurred in grass carp in Zhejiang Province, we added Shieh medium to our routine pathogen detection procedures to selectively culture F. psychrophilum . We subsequently obtained 12 yellow‐pigmented bacterial isolates. Physiological and biochemical characteristics, together with 16S rDNA phylogenetic analysis, confirmed these isolates as F. psychrophilum . Artificial infection experiments using a representative isolate reproduced the symptoms observed in naturally infected grass carp, and the isolate also exhibited pathogenicity toward Prussian carp. To our knowledge, this represents the first identification of F. psychrophilum as a bacterial pathogen of grass carp.

Although ulcerative lesions on the caudal peduncle and caudal fin are a classic characteristic of bacterial cold‐water disease in salmonids (Starliper 2011), grass carp in this study also developed ulcers on the body trunk (including the caudal peduncle); however, the most prominent clinical sign was hemorrhage in the snout‐jaw‐cheek region. Similar variations in tissue tropism have been documented in other non‐salmonid hosts. For example, in cultured ayu with cold‐water disease, affected fish frequently develop hemorrhagic lesions or ulcers on the lower jaw, whereas ulcerative lesions on the body trunk are typically absent (Miwa and Nakayasu 2005). These observations suggest that F. psychrophilum may exhibit varying tissue tropism and clinical manifestation profiles across different host species.

The biochemical characteristics of F. psychrophilum indicate its capacity to degrade various substrates including proteins, lipids and nucleic acids. Genomic studies further reveal that this bacterium encodes at least 14 putative extracellular proteases (Duchaud et al. 2007). Following colonization of the host epidermis, these enzymes likely contribute to progressive damage to connective and muscle tissues, providing a plausible explanation for the skin ulcers and muscle necrosis observed in the diseased grass carp in this study. In the skeletal muscle adjacent to ulcerative lesions, a mixed inflammatory infiltrate consisting predominantly of lymphocytes with occasional granulocytes was observed. This cellular composition suggests a host immune response transitioning from an acute to a subacute/chronic phase. In the low‐temperature aquaculture environment, F. psychrophilum infections typically follow a protracted course lasting several weeks. The inflammatory cell profile observed in this case is consistent with this clinical feature, reflecting the sustained interaction between the pathogen and the host under cold‐water conditions.

After infection with isolate CYL302‐1 at concentrations of 1.2 × 108 and 1.2 × 107 CFU/fish, 80% and 20% of the healthy grass carp died, respectively. It is noteworthy that some fish in the lower‐concentration group displayed typical symptoms, including snout congestion, swelling and spiral swimming, but eventually recovered. Therefore, it could be concluded that the isolated bacterium is indeed pathogenic, although the mortality rate was lower than that during field outbreaks. Similar patterns of disease caused by low‐pathogenicity strains have been reported in bacterial cold‐water disease of salmonids (Jarau et al. 2018; Sundell et al. 2019). As Sundell et al. (2019) emphasized, disease severity depends not only on pathogen virulence but also on host condition and environmental factors.

Regarding the frequent outbreaks of low‐temperature diseases in grass carp farming in recent years, we propose that they result from an interaction between the pathogen's psychrotolerance and host immunosuppression under cold conditions. F. psychrophilum has evolved multiple low‐temperature adaptation mechanisms, and its secreted extracellular proteases maintain high activity at low temperatures (Duchaud et al. 2007; Secades et al. 2001, 2003). Concurrently, low temperatures can cause pathological damage to the spleen and liver in grass carp, and suppress the expression of immune‐related genes (Chen et al. 2026). The spleen, as a key target organ for F. psychrophilum (Rangdale et al. 1999), plays a critical role in the host's anti‐infective immunity (Hadidi et al. 2008). Under cold conditions, the pathogen's pathogenic potential and the host's immunosuppressive state interact, collectively increasing grass carp's susceptibility to this disease. Furthermore, the intensive high‐density farming models commonly used in current grass carp aquaculture can significantly elevate physiological stress levels and reduce immunity, thereby contributing to increased pathogen susceptibility (Li et al. 2023). Similarly, moderate stocking density has been shown to effectively reduce mortality from bacterial cold‐water disease in ayu (Iguchi et al. 2003).

Additionally, stress or handling is known to trigger outbreaks of bacterial cold‐water disease (Miwa and Nakayasu 2005; Barnes and Brown 2011). Consistent with our observations, when water temperatures drop below 20°C, especially below 15°C, even minor handling of grass carp can lead to disease outbreaks within 1–2 weeks, with mortality continuing for up to a month. Currently, this recurrent disease poses a serious challenge to the cyprinid aquaculture industry in China. For example, farmers increasingly avoid netting, pond transfer and pond combining operations during winter, and spring stocking—previously carried out from February to March—has been delayed until mid‐to‐late April, when water temperatures rise above 20°C.

Although the intraperitoneal injection model in this study confirmed the pathogenicity of the F. psychrophilum isolate in grass carp, this route bypasses natural barriers such as skin, mucus and the intestinal tract, differing from the natural infection process. To simulate natural infection, we also attempted immersion challenges. The results were consistent with previous reports: only grass carp with artificially abraded skin developed ulcerative lesions and exhibited approximately 20% mortality (immersion concentration 107 CFU/mL, 30 min, fish size 150 ± 25 g), whereas fish with normal skin showed no obvious signs of infection.

Similarly, several studies have reported that establishing a reliable immersion infection model for F. psychrophilum is challenging and that minor skin damage significantly promotes disease development and increases mortality (Decostere et al. 2000; Garcia et al. 2000; Madetoja et al. 2000), suggesting that skin injuries may be the critical portals of entry for this pathogen (Miwa and Nakayasu 2005). Therefore, future systematic immersion challenge studies are necessary to further elucidate the natural entry portals and early colonization mechanisms of F. psychrophilum in grass carp.

Several studies have examined the growth temperature of F. psychrophilum under laboratory conditions. These studies reported an optimal growth range of 10°C–20°C, with growth slowing at 5°C–8°C and becoming sparse above 20°C; most strains show no growth at 25°C (Bernardet and Kerouault 1989; Lehmann et al. 1991; Iida and Mizokami 1996; Lee and Hee 1998). Similarly, in this study, the growth temperature range of isolate CYL302‐1 in EAO broth was 5°C–18°C, with optimal growth observed between 10°C and 18°C. Notably, in EAO agar, no visible colonies formed at 0°C and 2°C after 5 days of incubation; however, when incubation was extended to 7–10 days, clear colonies developed. Therefore, the actual growth temperature range on solid medium should be considered 0°C–23°C. The broader temperature adaptation range observed on solid medium compared with liquid medium may be attributed to the marked osmotic sensitivity of F. psychrophilum (Michel et al. 1999). The physical structure and microenvironmental buffering of solid substrates can shield cells from temperature‐related stress, enabling survival and slow growth under conditions that would inhibit or kill planktonic cells in liquid culture (Panikov and Sizova 2007; Jeanson et al. 2015). Additionally, the optimal growth temperature of this is corresponds to the water temperature at which disease outbreaks occur, further supporting its role as the causative agent of grass carp disease. Previous studies have shown that F. psychrophilum grows well at NaCl concentrations of 0%–0.5% (Bernardet and Kerouault 1989; Lee and Hee 1998), with some strains tolerating up to 1% (Iida and Mizokami 1996) and a few up to 2% (Nematollahi et al. 2003). In contrast, the salt tolerance of CYL302‐1 was narrower, showing marked growth inhibition at 0.1% NaCl and only sparse growth at 0.5% NaCl. These results suggest that modulating salinity may represent a viable strategy for controlling bacterial proliferation and preventing disease outbreaks.

Previous research has demonstrated an association between specific ST types of F. psychrophilum and host fish species. For example, strains responsible for bacterial cold‐water disease in rainbow trout are predominantly classified as CC‐ST10 (Nilsen et al. 2014; Knupp et al. 2019), whereas strains isolated from ayu are more frequently distributed within CC‐ST52 and CC‐ST48/CC‐ST56 (Fujiwara‐Nagata et al. 2013). The isolate CYL302‐1 represents a newly identified ST (ST418) and belongs to CC‐ST56, a clonal complex associated with ayu and cyprinids. To further explore the phylogenetic relationships, we performed a comparative phylogenetic analysis using strains deposited in the PubMLST database (https://pubmlst.org/fpsychrophilum/), positioning ST418 alongside the major STs from salmonids, as well as all known STs from ayu and cyprinid fish. As shown in Figure S1, the strains could be divided into four major clusters (Clusters 1–4). ST418, the sequence type identified in this study, falls within Cluster 1, which contains strains isolated from salmonids, ayu and cyprinids. Notably, ST418 and CC‐ST10, the predominant clonal complex in salmonids (located in Cluster 4), belong to distinct major branches. Although the subclade containing ST418 includes both ayu‐ and cyprinid‐derived strains, ST418 is more closely related to the cyprinid strains, suggesting that it may represent a lineage associated with cyprinid host adaptation.

Previous studies have also indicated a strong correlation between serological typing and host species in this bacterium. Rochat et al. (2017) reported a significant association between mPCR serotypes and host fish species. Isolates from coho salmon are mainly Type‐0, those from ayu are predominantly Type‐3 and isolates from rainbow trout are mostly Type‐1 and Type‐2. To date, a limited number of F. psychrophilum isolates from cyprinid fish have been characterized for their molecular serotypes and sequence types (STs). Most of these isolates, including 10 recently deposited F. psychrophilum isolates from cyprinid fish in China available in the PubMLST database (https://pubmlst.org/fpsychrophilum), belong to serotype Type‐1 or Type‐4, followed by Type‐0, with fewer assigned to Type‐2 and there are no reports of Type‐3 among cyprinid hosts (Fujiwara‐Nagata et al. 2024). In the present study, all 12 isolates obtained from the diseased grass carp were determined as ST418 and serotype Type‐1, suggesting that this type may represent a prevalent genotype in this particular disease outbreak, though further surveillance is needed to determine the dominant genotypes in grass carp aquaculture in China.

The T82A substitution in the quinolone resistance‐determining region (QRDR) of GyrA detected in isolate CYL302‐1 is a well‐validated quinolone resistance mechanism in F. psychrophilum (Izumi and Aranishi 2004; Shah et al. 2012; Saticioglu et al. 2019). The non‐wild‐type (NWT) phenotypes observed for enrofloxacin, flumequine and oxolinic acid in this isolate are highly consistent with the presence of this QRDR mutation. Furthermore, no mutations were detected in the QRDRs of GyrB, ParC or ParE, further supporting GyrA as the primary determinant of quinolone resistance in this isolate.

For warm‐water fish, feeding behaviour declines markedly below 10°C and remains very low between 10°C and 15°C. This anorexia becomes more severe following F. psychrophilum infection. Therefore, isolating the pathogen, conducting drug sensitivity testing, identifying effective antibiotics, and administering medicated feed are generally difficult to implement in practice. This is one of the key factors contributing to the increasing incidence of disease outbreaks in grass carp in recent years. Currently, reducing stress and handling, and appropriately increasing water temperature and salinity during transport, remain the most effective prevention and control.

5. Conclusion

In this study, a novel disease emerging in grass carp, characterized by outbreaks under low‐temperature conditions, was associated with F. psychrophilum infection. To the best of our knowledge, this is the first report of severe disease caused by F. psychrophilum in grass carp. The pathogen induces systemic infection in grass carp and also shows pathogenicity to Prussian carp. All isolates obtained in this study belonged to serotype Type‐1 and were assigned to a novel sequence type, ST418. Further characterization of the representative isolate, CYL302‐1, revealed that it grows at 0°C–23°C, is highly sensitive to salinity, and shows reduced sensitivity to quinolones associated with a mutation in GyrA. These findings provide useful reference data for aquaculture practitioners and veterinarians in disease prevention and management.

Author Contributions

Jiahao Li: investigation, validation, data curation, visualization, writing – review and editing. Xiao Liu: writing – review and editing, formal analysis. Lingyun Lin: conceptualization, methodology, visualization, writing – original draft, writing – review and editing, funding acquisition. Jinyu Shen: conceptualization, investigation, resources, project administration, funding acquisition. Xiaoyi Pan: investigation, formal analysis, project administration, funding acquisition, writing – review and editing. Jiayun Yao: resources, formal analysis.

Funding

This study was supported by the Science and Technology Cooperation Project of Zhejiang Province (2025SNJF093), Zhejiang Provincial Natural Science Foundation of China (ZCLTGN24C1901) and National Key Research and Development Program of China (2024YFD2401102).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Phylogenetic tree based on the allelic sequences of seven housekeeping genes used in Flavobacterium psychrophilum MLST.

JFD-49-e70193-s002.zip (118.1KB, zip)

Table S1: Bacterial isolates from different tissues of the diseased grass carp.

JFD-49-e70193-s001.docx (16.4KB, docx)

Acknowledgements

The authors are grateful to Professor Zhenrong Song of Jimei University for providing valuable advice on histopathological sectioning and staining techniques, and particularly for guidance in the interpretation of histopathology.

Contributor Information

Xiao Liu, Email: liuxiao@zjou.edu.cn.

Xiaoyi Pan, Email: panxiaoyi@163.com.

Data Availability Statement

The 16S rDNA sequence of strain CYL302‐1 is available in GenBank (accession no. PX419286), and the associated AT and ST are available in PubMLST.

References

  1. Amita, K. , Hoshino M., Honma T., and Wakabayashi H.. 2000. “An Investigation on the Distribution of Flavobacterium psychrophilum in the Umikawa River.” Fish Pathology 35: 193–197. 10.3147/jsfp.35.193. [DOI] [Google Scholar]
  2. Avendaño‐Herrera, R. , Tapia‐Cammas D., Duchaud E., and Irgang R.. 2020. “Serological Diversity in Flavobacterium psychrophilum : A Critical Update Using Isolates Retrieved From Chilean Salmon Farms.” Journal of Fish Diseases 43: 877–888. 10.1111/jfd.13199. [DOI] [PubMed] [Google Scholar]
  3. Barnes, M. E. , and Brown M. L.. 2011. “A Review of Flavobacterium psychrophilum Biology, Clinical Signs, and Bacterial Cold Water Disease Prevention and Treat.” Open Fish Science Journal 4: 40–48. 10.2174/1874401x01104010040. [DOI] [Google Scholar]
  4. Bernardet, J. F. 1989. “‘ Flexibacter columnaris ’: First Description in France and Comparison With Bacterial Strains From Other Origins.” Diseases of Aquatic Organisms 6: 37–44. 10.3354/dao006037. [DOI] [Google Scholar]
  5. Bernardet, J. F. , and Kerouault B.. 1989. “Phenotypic and Genomic Studies of ‘ Cytophaga psychrophila ’ Isolated From Diseased Rainbow Trout ( Oncorhynchus mykiss ) in France.” Applied and Environmental Microbiology 55: 1796–1800. 10.1128/aem.55.7.1796-1800.1989. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bernardet, J. F. , Segers P., Vancanneyt M., Berthe F., Kersters K., and Vandamme P.. 1996. “Cutting a Gordian Knot: Emended Classification and Description of the Genus Flavobacterium, Emended Description of the Family Flavobacteriaceae, and Proposal of Flavobacterium hydatis nom. nov. (Basonym, Cytophaga aquatilis Strohl and Tait 1978).” International Journal of Systematic Bacteriology 46: 128–148. 10.1099/00207713-46-1-128. [DOI] [Google Scholar]
  7. Borg, A. F. 1960. “Studies on Myxobacteria Associated With Diseases in Salmonid Fishes.” Wildlife Disease 8: 85. [Google Scholar]
  8. Bureau of Fisheries of the Ministry of Agriculture of the People's Republic of China . 2024. The China Fisheries Statistical Yearbook. Chinese Agricultural Press. [Google Scholar]
  9. Chen, G. , Xu Y., Lu Y., Pan J., Feng H., and Su J.. 2026. “High Temperatures Deteriorate Grass Carp ( Ctenopharyngodon idella ) Response to GCRV‐II Infection: Survival Rates, Viral Loads, Histopathology, and Gene Expressions.” Aquaculture 613: 743409. 10.1016/j.aquaculture. [DOI] [Google Scholar]
  10. Chinchilla, B. , Vázquez‐Fernández E., Rebollada‐Merino A., Pérez‐Sancho M., Domínguez L., and Rodríguez‐Bertos A.. 2023. “First Detection of Flavobacterium psychrophilum in Juvenile Siberian Sturgeons ( Acipenser baerii ) and Description of the Pathological Findings.” Journal of Fish Diseases 46: 887–894. 10.1111/jfd.13801. [DOI] [PubMed] [Google Scholar]
  11. CLSI , ed. 2020a. Methods for Antimicrobial Broth Dilution and Disk Diffusion Susceptibility Testing of Bacteria Isolated From Aquatic Animals. 2nd ed. CLSI Guideline VET03. Clinical and Laboratory Standards Institute. [Google Scholar]
  12. CLSI , ed. 2020b. Performance Standards for Antimicrobial Susceptibility Testing of Bacteria Isolated From Aquatic Animals. 3rd ed. CLSI Supplement VET04. Clinical and Laboratory Standards Institute. [Google Scholar]
  13. Decostere, A. , Lammens M., and Haesebrouck F.. 2000. “Difficulties in Experimental Infection Studies With Flavobacterium psychrophilum in Rainbow Trout ( Oncorhynchus mykiss ) Using Immersion, Oral and Anal Challenges.” Research in Veterinary Science 69, no. 2: 165–169. 10.1053/rvsc.2000.0408. [DOI] [PubMed] [Google Scholar]
  14. Duchaud, E. , Boussaha M., Loux V., et al. 2007. “Complete Genome Sequence of the Fish Pathogen Flavobacterium psychrophilum .” Nature Biotechnology 25: 763–769. 10.1038/nbt1313. [DOI] [PubMed] [Google Scholar]
  15. Elsayed, E. E. , Eissa A. E., and Faisal M.. 2006. “Isolation of Flavobacterium psychrophilum From Sea Lamprey, Petromyzon marinus L., With Skin Lesions in Lake Ontario.” Journal of Fish Diseases 29: 629–632. 10.1111/j.1365-2761.2006.00756.x. [DOI] [PubMed] [Google Scholar]
  16. Fujiwara‐Nagata, E. , Chantry‐Darmon C., Bernardet J.‐F., Eguchi M., Duchaud E., and Nicolas P.. 2013. “Population Structure of the Fish Pathogen Flavobacterium psychrophilum at Whole‐Country and Model River Levels in Japan.” Veterinary Research 44: 34. 10.1186/1297-9716-44-34. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Fujiwara‐Nagata, E. , Rochat T., Lee B.‐H., Lallias D., Rigaudeau D., and Duchaud E.. 2024. “Host Specificity and Virulence of Flavobacterium psychrophilum : A Comparative Study in Ayu ( Plecoglossus altivelis ) and Rainbow Trout ( Oncorhynchus mykiss ) Hosts.” Veterinary Research 55: 75. 10.1186/s13567-024-01326-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fujiwara‐Nagata, E. , Shindoh Y., Yamamoto M., Okamura T., and Eguchi M.. 2019. “Distribution of Flavobacterium psychrophilum and Its gyrA Genotypes in a River.” Fisheries Science 85: 913–923. 10.1007/s12562-019-01355-7. [DOI] [Google Scholar]
  19. Garcia, C. , Pozet F., and Michel C.. 2000. “Standardization of Experimental Infection With Flavobacterium psychrophilum , the Agent of Rainbow Trout Oncorhynchus mykiss Fry Syndrome.” Diseases of Aquatic Organisms 42, no. 3: 191–197. 10.3354/dao042191. [DOI] [PubMed] [Google Scholar]
  20. Goodwin, A. E. , Merry G. E., and Sadler J.. 2006. “Detection of the Herpesviral Hematopoietic Necrosis Disease Agent (Cyprinid herpesvirus 2) in Moribund and Healthy Goldfish: Validation of a Quantitative PCR Diagnostic Method.” Diseases of Aquatic Organisms 69: 137–143. 10.3354/dao069137. [DOI] [PubMed] [Google Scholar]
  21. Hadidi, S. , Glenney G. W., Welch T. J., Silverstein J. T., and Wiens G. D.. 2008. “Spleen Size Predicts Resistance of Rainbow Trout to Flavobacterium psychrophilum Challenge.” Journal of Immunology 180: 4156–4165. 10.4049/jimmunol.180.6.4156. [DOI] [PubMed] [Google Scholar]
  22. Iguchi, K. I. , Ogawa K., Nagae M., and Ito F.. 2003. “The Influence of Rearing Density on Stress Response and Disease Susceptibility of Ayu ( Plecoglossus altivelis ).” Aquaculture 220: 515–523. 10.1016/S0044-8486(02)00626-9. [DOI] [Google Scholar]
  23. Iida, Y. , and Mizokami A.. 1996. “Outbreaks of Coldwater Disease in Wild Ayu and Pale Chub.” Fish Pathology 31: 157–164. 10.3147/jsfp.31.157. [DOI] [Google Scholar]
  24. Imajoh, M. 2022. “Bacterial Cold‐Water Disease in Ayu ( Plecoglossus altivelis altivelis ) Inhabiting Rivers in Japan.” Frontiers in Cellular and Infection Microbiology 12: 1073966. 10.3389/fcimb.2022.1073966. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Izumi, S. , and Aranishi F.. 2004. “Relationship Between gyrA Mutations and Quinolone Resistance in Flavobacterium psychrophilum Isolates.” Applied and Environmental Microbiology 70: 3968–3972. 10.1128/AEM.70.7.3968-3972.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Izumi, S. , Liu H., Aranishi F., and Wakabayashi H.. 2003. “A Novel Serotype of Flavobacterium psychrophilum Detected Using Antiserum Against an Isolate From Amago, Oncorhynchus masou rhodurus Jordan & Gilbert, in Japan.” Journal of Fish Diseases 26: 677–680. 10.1046/j.1365-2761.2003.00502.x. [DOI] [PubMed] [Google Scholar]
  27. Jarau, M. , Di Natale A., Huber P. E., Macinnes J. I., and Lumsden J. S.. 2018. “Virulence of Flavobacterium psychrophilum Isolates in Rainbow Trout Oncorhynchus mykiss (Walbaum).” Journal of Fish Diseases 41: 1505–1514. 10.1111/jfd.12861. [DOI] [PubMed] [Google Scholar]
  28. Jeanson, S. , Floury J., Gagnaire V., Lortal S., and Thierry A.. 2015. “Bacterial Colonies in Solid Media and Foods: A Review on Their Growth and Interactions With the Micro‐Environment.” Frontiers in Microbiology 6: 1284. 10.3389/fmicb.2015.01284. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Jiang, H. , Li R., Zhao Y., et al. 2024. “Isolation, Identification, and Histopathological Observation of Flavobacterium psychrophilum From Diseased Carassius auratus gibelio .” Journal of Fisheries of China 48: 94–106. 10.11964/jfc.20230914162. [DOI] [Google Scholar]
  30. Knupp, C. , Soto E., Call D., and Loch T.. 2024. “Persistence of Heterologous Flavobacterium psychrophilum Genetic Variants in Microcosms Simulating Fish Farm and Hatchery Environments.” Environmental Microbiology 26: e16581. 10.1111/1462-2920.16581. [DOI] [PubMed] [Google Scholar]
  31. Knupp, C. , Wiens Gregory D., Faisal M., et al. 2019. “Large‐Scale Analysis of Flavobacterium psychrophilum Multilocus Sequence Typing Genotypes Recovered From North American Salmonids Indicates That Both Newly Identified and Recurrent Clonal Complexes Are Associated With Disease.” Applied and Environmental Microbiology 85: e02305‐18. 10.1128/AEM.02305-18. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Kondo, M. , Kawai K., Kurohara K., and Oshima S.‐I.. 2002. “Adherence of Flavobacterium psychrophilum on the Body Surface of the Ayu Plecoglossus altivelis .” Microbes and Infection 4: 279–283. 10.1016/S1286-4579(02)01539-3. [DOI] [PubMed] [Google Scholar]
  33. Kumar, S. , Stecher G., Suleski M., Sanderford M., Sharma S., and Tamura K.. 2024. “MEGA12: Molecular Evolutionary Genetic Analysis Version 12 for Adaptive and Green Computing.” Molecular Biology and Evolution 41: msae263. 10.1093/molbev/msae263. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Lee, K. B. , and Hee G. J.. 1998. “First Isolation and Identification of Cytophaga psychrophila From Cultured Ayu in Korea.” Fish Pathology 33: 37–38. 10.3147/jsfp.33.37. [DOI] [Google Scholar]
  35. Lehmann, J. , Mock D., Sturenberg L.‐J., and Bernardet J.‐F.. 1991. “First Isolation of Cytophaga psychrophila From a Systemic Disease in Eel and Cyprinids.” Diseases of Aquatic Organisms 10: 217–220. 10.3354/dao010217. [DOI] [Google Scholar]
  36. Li, S. , Chai J., Knupp C., et al. 2021. “Phenotypic and Genetic Characterization of Flavobacterium psychrophilum Recovered From Diseased Salmonids in China.” Microbiology Spectrum 9: e0033021. 10.1128/Spectrum.00330-21. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Li, W. , Li D., Yang Q., et al. 2023. “Long‐Term Crowding Stress Induces Chronic Inflammatory Response and Declines the Immunity of Grass Carp ( Ctenopharyngodon idella ).” Aquaculture 577: 739976. 10.1016/j.aquaculture.2023.739976. [DOI] [Google Scholar]
  38. Lin, S. , Milardi M., Gao Y., and Wong M. H.. 2022. “Sustainable Management of Non‐Native Grass Carp as a Protein Source, Weed‐Control Agent and Sport Fish.” Aquaculture Research 53: 5809–5824. 10.1111/are.16080. [DOI] [Google Scholar]
  39. Liu, X.‐G. , Shen H.‐Y., Gu Z.‐J., Cheng G., Wang J., and Zhu H.. 2023. “The Environmental Impact and Development Direction of Grass Carp, Ctenopharyngodon idella , Aquaculture.” Journal of the World Aquaculture Society 54: 1354–1366. 10.1111/jwas.12990. [DOI] [Google Scholar]
  40. Luo, Z. , Liu Z., Fu J., Zhang Q., Huang B., and Nie P.. 2016. “Immunogenicity and Protective Role of Antigenic Regions From Five Outer Membrane Proteins of Flavobacterium columnare in Grass Carp Ctenopharyngodon idella .” Chinese Journal of Oceanology and Limnology 34: 1247–1257. 10.1007/s00343-016-5096-z. [DOI] [Google Scholar]
  41. Lv, W. , Zhou Z., Xie L., et al. 2025. “Pathological and Molecular Characterization of Grass Carp Co‐Infected With Two Aeromonas Species.” Animals 15: 263. 10.3390/ani15020263. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Madetoja, J. , Dalsgaard I., and Wiklund T.. 2002. “Occurrence of Flavobacterium psychrophilum in Fish‐Farming Environments.” Diseases of Aquatic Organisms 52: 109–118. 10.3354/dao052109. [DOI] [PubMed] [Google Scholar]
  43. Madetoja, J. , Nyman P., and Wiklund T.. 2000. “ Flavobacterium psychrophilum , Invasion Into and Shedding by Rainbow Trout Oncorhynchus mykiss .” Diseases of Aquatic Organisms 43: 27–38. 10.3354/dao043027. [DOI] [PubMed] [Google Scholar]
  44. Madetoja, J. , Nystedt S., and Wiklund T.. 2003. “Survival and Virulence of Flavobacterium psychrophilum in Water Microcosms.” FEMS Microbiology Ecology 43: 217–223. 10.1111/j.1574-6941.2003.tb01061.x. [DOI] [PubMed] [Google Scholar]
  45. Martínez, J. L. , Casado A., and Enríquez R.. 2004. “Experimental Infection of Flavobacterium psychrophilum in Fins of Atlantic Salmon Salmo salar Revealed by Scanning Electron Microscopy.” Diseases of Aquatic Organisms 59: 79–84. 10.3354/dao059079. [DOI] [PubMed] [Google Scholar]
  46. Michel, C. , Antonio D., and Hedrick R. P.. 1999. “Production of Viable Cultures of Flavobacterium psychrophilum : Approach and Control.” Research in Microbiology 150: 351–358. 10.1016/S0923-2508(99)80061-8. [DOI] [PubMed] [Google Scholar]
  47. Miwa, S. , and Nakayasu C.. 2005. “Pathogenesis of Experimentally Induced Bacterial Cold Water Disease in Ayu Plecoglossus altivelis .” Diseases of Aquatic Organisms 67: 93–104. 10.3354/dao067093. [DOI] [PubMed] [Google Scholar]
  48. Nematollahi, A. , Decostere A., Pasmans F., and Haesebrouck F.. 2003. “ Flavobacterium psychrophilum Infections in Salmonid Fish.” Journal of Fish Diseases 26: 563–574. 10.1046/j.1365-2761.2003.00488.x. [DOI] [PubMed] [Google Scholar]
  49. Nicolas, P. , Mondot S., Achaz G., Bouchenot C., Bernardet J.‐F., and Duchaud E.. 2008. “Population Structure of the Fish‐Pathogenic Bacterium Flavobacterium psychrophilum .” Applied and Environmental Microbiology 74: 3702–3709. 10.1128/AEM.00244-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Nilsen, H. , Sundell K., Duchaud E., et al. 2014. “Multilocus Sequence Typing Identifies Epidemic Clones of Flavobacterium psychrophilum in Nordic Countries.” Applied and Environmental Microbiology 80: 2728–2736. 10.1128/AEM.04233-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Oplinger, R. W. , and Wagner E. J.. 2012. “Effects of Media Ingredient Substitution and Comparison of Growth of Flavobacterium psychrophilum Among Four Media.” Journal of Aquatic Animal Health 24, no. 1: 49–57. 10.1080/08997659.2012.668510. [DOI] [PubMed] [Google Scholar]
  52. Panikov, N. , and Sizova M.. 2007. “Growth Kinetics of Microorganisms Isolated From Alaskan Soil and Permafrost in Solid Media Frozen Down to −35°C.” FEMS Microbiology Ecology 59: 500–512. 10.1111/j.1574-6941.2006.00210.x. [DOI] [PubMed] [Google Scholar]
  53. Papadopoulou, A. , Dalsgaard I., Lindén A., and Wiklund T.. 2017. “In Vivo Adherence of Flavobacterium psychrophilum to Mucosal External Surfaces of Rainbow Trout ( Oncorhynchus mykiss ) Fry.” Journal of Fish Diseases 40: 1309–1320. 10.1111/jfd.12603. [DOI] [PubMed] [Google Scholar]
  54. Rangdale, R. E. , Richards R. H., and Alderman D. J.. 1999. “Histopathological and Electron Microscopical Observations on Rainbow Trout Fry Syndrome.” Veterinary Record 144: 251–254. 10.1136/vr.144.10.251. [DOI] [PubMed] [Google Scholar]
  55. Rao, Y. , and Su J.. 2015. “Insights Into the Antiviral Immunity Against Grass Carp ( Ctenopharyngodon idella ) Reovirus (GCRV) in Grass Carp.” Journal of Immunology Research 2015, no. 1: 670437. 10.1155/2015/670437. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Rochat, T. , Fujiwara‐Nagata E., Calvez S., et al. 2017. “Genomic Characterization of Flavobacterium psychrophilum Serotypes and Development of a Multiplex PCR‐Based Serotyping Scheme.” Frontiers in Microbiology 8: 1752. 10.3389/fmicb.2017.01752. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Saticioglu, I. B. , Duman M., Ajmi N., Altun S., Rochat T., and Duchaud E.. 2025. “Phylogenomic Characterization of Flavobacterium psychrophilum Isolates Retrieved From Turkish Rainbow Trout Farms.” Journal of Fish Diseases 48: e13961. 10.1111/jfd.13961. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Saticioglu, I. B. , Duman M., Smith P., Wiklund T., and Altun S.. 2019. “Antimicrobial Resistance and Resistance Genes in Flavobacterium psychrophilum Isolates From Turkey.” Aquaculture 512: 734293. 10.1016/j.aquaculture.2019.734293. [DOI] [Google Scholar]
  59. Secades, P. , Ãlvarez B., and Guijarro J. A.. 2001. “Purification and Characterization of a Psychrophilic, Calcium‐Induced, Growth‐Phase‐Dependent Metalloprotease From the Fish Pathogen Flavobacterium psychrophilum .” Applied and Environmental Microbiology 67, no. 6: 2436–2444. 10.1128/AEM.67.6.2436-2444.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Secades, P. , Ãlvarez B., and Guijarro J. A.. 2003. “Purification and Properties of a New Psychrophilic Metalloprotease (Fpp2) in the Fish Pathogen Flavobacterium psychrophilum .” FEMS Microbiology Letters 226, no. 2: 273–279. 10.1016/S0378-1097(03)00599-8. [DOI] [PubMed] [Google Scholar]
  61. Shah, S. Q. A. , Nilsen H., Bottolfsen K., Colquhoun D. J., and Sørum H.. 2012. “DNA Gyrase and Topoisomerase IV Mutations in Quinolone‐Resistant Flavobacterium psychrophilum Isolated From Diseased Salmonids in Norway.” Microbial Drug Resistance 18: 207–214. 10.1089/mdr.2011.0142. [DOI] [PubMed] [Google Scholar]
  62. Soares, S. M. C. , Walker A., Elwenn S. A., et al. 2019. “First Isolation of Flavobacterium psychrophilum Associated With Reports of Moribund Wild European Eel ( Anguilla anguilla ) in Scotland.” Journal of Fish Diseases 42: 1509–1521. 10.1111/jfd.13069. [DOI] [PubMed] [Google Scholar]
  63. Starliper, C. E. 2011. “Bacterial Coldwater Disease of Fishes Caused by Flavobacterium psychrophilum .” Journal of Advanced Research 2: 97–108. 10.1016/j.jare.2010.04.001. [DOI] [Google Scholar]
  64. Strepparava, N. , Wahli T., Segner H., and Petrini O.. 2014. “Detection and Quantification of Flavobacterium psychrophilum in Water and Fish Tissue Samples by Quantitative Real Time PCR.” BMC Microbiology 14: 105. 10.1186/1471-2180-14-105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Sundell, K. , Landor L., Nicolas P., et al. 2019. “Phenotypic and Genetic Predictors of Pathogenicity and Virulence in Flavobacterium psychrophilum .” Frontiers in Microbiology 10: 1711. 10.3389/fmicb.2019.01711. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Verma, V. , and Prasad Y.. 2014. “Isolation and Immunohistochemical Identification of Flavobacterium psychrophilum From the Tissue of Catfish, Clarias batrachus .” Journal of Environmental Biology 35: 389–393. [PubMed] [Google Scholar]
  67. Weisburg, W. G. , Barns S. M., Pelletier D. A., and Lane D. J.. 1991. “16S Ribosomal DNA Amplification for Phylogenetic Study.” Journal of Bacteriology 173: 697–703. 10.1128/jb.173.2.697-703.1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Zeng, W. , Wang Q., Wang Y., et al. 2013. “Establishment of Multiplex PCR for Detection of Grass Carp Reovirus and Its Application.” Journal of Fishery Sciences of China 20: 419–426. 10.3724/SP.J.1118.2013.00419. [DOI] [Google Scholar]
  69. Zhang, K. , Ma J., and Fan Y.. 2021. “Epidemiology of the Grass Carp Reovirus.” In Aquareovirus, edited by Fang Q., 133–148. Singapore. [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1: Phylogenetic tree based on the allelic sequences of seven housekeeping genes used in Flavobacterium psychrophilum MLST.

JFD-49-e70193-s002.zip (118.1KB, zip)

Table S1: Bacterial isolates from different tissues of the diseased grass carp.

JFD-49-e70193-s001.docx (16.4KB, docx)

Data Availability Statement

The 16S rDNA sequence of strain CYL302‐1 is available in GenBank (accession no. PX419286), and the associated AT and ST are available in PubMLST.

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