https://orcid.org/0000-0002-4775-2280
Figures
Abstract
Marsupials of the genus Didelphis are highly adapted to urban environments and are widely distributed in the Americas. Streptococcus didelphis is a bacterium that has been isolated from, and associated with disease, in the Virginia opossum (Didelphis virginiana) and the white-eared opossum (Didelphis albiventris). This study describes pathological changes associated with S. didelphis infection in white-eared opossums (Didelphis albiventris) and Brazilian common opossums (Didelphis aurita) as well as microbiological and genomic characterization of isolates. Ten opossums underwent necropsy and had ulcerative dermatitis of probable traumatic origin had S. didelphis was isolated from cutaneous lesions (n = 16) or systemic sites (n = 4). In contrast, 34 free-ranging opossums that were captured had negative culture results for S. didelphis from nasal swabs. Other lesions in opossums naturally infected with S. didelphihs included splenitis (7/10), myocarditis (6/10), interstitial nephritis and pyelonephritis (7/10), and myositis (4/10). Most isolates were susceptible to the antimicrobial drugs tested and none of them were able to form biofilm in vitro. Whole genome analysis of six isolates revealed no resistance determinants, virulence factors or plasmids, and the isolates showed high genomic similarity.
Citation: Santos DOd, Castro YGd, Vieira AD, Campos BHd, Souza LdRd, Paula NFd, et al. (2026) Streptococcus didelphis infection in free-ranging white-eared opossum (Didelphis albiventris) and Brazilian common opossum (Didelphis aurita): pathology, microbiologic, and genomic characterization. PLoS One 21(4): e0348357. https://doi.org/10.1371/journal.pone.0348357
Editor: André Luiz Rodrigues Roque, Instituto Oswaldo Cruz, BRAZIL
Received: February 20, 2026; Accepted: April 15, 2026; Published: April 30, 2026
Copyright: © 2026 Santos et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the manuscript and its Supporting information files.
Funding: Work in RLS lab is supported by CNPq (Conselho Nacional de Desenvolvimento Científico e Tecnológico, Brazil; grant 310088/2023-2), FAPEMIG (Fundação de Amparo à Pesquisa do Estado de Minas Gerais, Brazil; grants APQ-01972-23 and APQ-01701-23), and CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior, Brazil; grant 88881.083133/2024-01 – PIPD). VA, ROSS and RLS have fellowships from CNPq (Brazil). There was no additional external funding received for this study. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Opossums are marsupials with generalist habits regarding diet and habitat, which favors their movement and adaptation to diverse environments. Thus, these animals are well adapted to urban areas, where they can interact with humans and domestic animals, as well as natural environments, where they interact with other wild animals [1–3]. Therefore, understanding the factors that lead to disease in these animals is important, as they can act as reservoirs of pathogens relevant to human and animal health [3–8]. Opossums are widely distributed across the American continent and the white-eared opossum (Didelphis albiventris) and the Brazilian common opossum (Didelphis aurita) are commonly found in Brazil.
The genus Streptococcus includes 212 species [9], characterized by gram-positive, spherical or ovoid cells, typically arranged in chains or pairs. They are found in the environment and can be commensal in both animals and humans [10]. S. didelphis was first described in the Virginia opossum (Didelphis virginiana) associated with dermatitis and septicemia [11]. More recently, S. didelphis infection was reported in a Virginia opossum coinfected with macropodid alphaherpesvirus 2 [12] and in a Virginia opossum with suppurative vaginitis and sepsis [13]. In the white-eared opossum, S. didelphis-associated infection was recently reported in only two animals with skin necrosis and pleuritis [14]. Given the scarcity of information about S. didelphis infection, this study aimed to describe the pathological aspects of naturally infected free-ranging white-eared and Brazilian common opossums, as well as the microbiological and genomic features of the isolates.
Materials and methods
Ethics statement
All procedures were previously approved by the Ethics Committee on the Use of Animals of the Universidade Federal de Minas Gerais (CEUA/UFMG) under protocol number 79/2022, by Instituto Chico Mendes de Conservação de Biodiversidade under protocol number 81680 and by Fundação de Parques Municipais e Zoobotânica de Belo Horizonte (FPMZ-BH) under protocol number FU003/2022. The study was also registered at Sistema Nacional de Gestão do Patrimônio Genético Tradicional Associado (SisGen) under protocol numbers ABB00DB and ACDCB02.
Necropsy and histopathology
Dead opossums were found at the Parque Américo Renné Gianetti (AR); Parque Aggeo Pio Sobrinho (AP); Universidade Federal de Minas Gerais campus (C); Parque Fazenda Lagoa do Nado (LN); Parque das Mangabeiras (PM); Parque Ursulina de Andrade Mello (U) and Belo Horizonte Zoo (Z), all located in Belo Horizonte, State of Minas Gerais, Brazil (coordinates 19° 55′ 00″ S, 43° 56′ 00″ W) (S1 Fig). Free-ranging opossums submitted to necropsy underwent gross examination, and all opossums from which S. didelphis was isolated were included in this study. Sex was recorded for each animal and age was estimated based on animal size and dentition development. Adults had bigger body sizes and fully developed dentitions and young had smaller body size and incomplete dentition. Samples from skin (ear and tail), superficial lymph nodes, salivary glands, brain, tongue, esophagus, stomach, small and large intestines, pancreas, trachea, lungs, heart, spleen, liver, gallbladder, kidneys, urinary bladder, testis, ovaries, uterus and bone marrow were systematically sampled and fixed in 10% buffers formalin solution for histopathology. Other samples such as bone, skeletal muscle and skin from other locations were also sampled when gross lesions were observed. Gross examination focused particularly on the presence of previous trauma, skin inflammatory infiltrate, extent of the lesions, systemic findings, and intravascular bacterial aggregates suggesting bacteremia. Formalin-fixed samples were routinely processed for paraffin embedding and 2–3 µm slides were then stained with hematoxylin and eosin or Gram stain. Based on gross examination, samples (swabs or tissue samples) from animals with suspected bacterial infection were collected for isolation.
Field procedures
In addition to pathological and microbiological analyses of opossums that were found dead, free-ranging opossum were captured in six urban parks in the city of Belo Horizonte (Parque das Mangabeiras – PM, Parque da Serra do Curral – SC, Parque Ursulina Andrade de Mello – U, Parque Aggeo Pio – AP, Parque Fazenda Lagoa do Nado – LN and Belo Horizonte Zoo – Z) with Tomahawk traps baited with bananas. Captured animals were physically and chemically restrained (Ketamine 10 mg/Kg and Midazolam 1 mg/Kg) prior to clinical evaluations and sample collection. Nasal swabs were sampled from all animals, and skin swabs were also sampled from individuals with skin lesions. All swabs were submitted to bacterial isolation. All captured animals were included in the study, identified with microchips, and released after sedation recovery.
Bacterial isolation and identification
Swabs were plated on brain heart infusion agar (BHI, Kasvi, Brazil) supplemented with 5% equine blood and MacConkey agar (BHI, Kasvi, Brazil) and incubated at 37°C for 48 h under aerobic conditions. Isolates were subjected to species identification by matrix-assisted laser desorption/ionization-time-of-flight mass spectrometry (MALDI-ToF MS; Bruker Daltonics, USA). Analyses were performed with pure colonies according to the manufacturer’s instructions, considering a confidence score ≥ 2.300 for species-level identification [15].
Antimicrobial susceptibility tests
Antimicrobial susceptibility of all S. didelphis isolates was evaluated by disk diffusion method [16]. Seven antimicrobials from five classes were tested: ampicillin (AM, 10 µg), cefotaxime (CTX, 30 µg), clindamycin (CM, 2 µg), chloramphenicol (CL, 30 µg), erythromycin (ERY, 15 µg), penicillin (PEN, 10 µg), and tetracycline (TET, 30 µg). The results were interpreted according to the criteria established in the CLSI [16] for Streptococcus spp. of the β-hemolytic group.
Biofilm formation test
Biofilm production capacity of the isolates was performed in microtiter plates according to Stepanović et al. [17]. Samples of Staphylococcus epidermidis ATCC 35984 (SE35984) and Staphylococcus aureus ATCC 6538 (SA6538) were used as positive control, while wells containing Tryptic Soy Broth (TSB, Oxoid, United Kingdom) were used as negative control. The reading was performed in a spectrophotometer (492 nm) (Thoth, Brazil).
DNA extraction and genomic sequencing
Six isolates from different cases underwent whole-genome sequencing. Strains were incubated on brain-heart infusion agar (BHI, Kasvi, Brazil) supplemented with 5% equine blood at 37°C for 48 h. Genomic DNA was extracted using the Wizard Genomic DNA Purification Kit (Promega, USA). Genome sequencing was performed using the Illumina MiSeq platform with the MiSeq Reagent Kit v2, generating paired-end reads of 2 x 150 pb. The generated raw data was analyzed with FastQC (Babraham Bioinformatics, Cambridge, England), retaining only paired-end reads with a Phred score ≥ 30. Assembly was performed using the de novo method with SPAdes v3.13.0 in careful mode [18,19]. Gaps were filled with Pilon [20]. Genomic sequences were submitted to the GenBank (https://www.ncbi.nlm.nih.gov/genbank/) under BioProject ID: PRJNA1265247. Individual sample identifiers are detailed in Table 1.
Genomic analysis
Bacterial species prediction from genomes was performed using Kmerfinder 3.2 [21]. ResFinder 4.1 [22] was used to identify the determinants of acquired antimicrobial resistance, adopting minimum criteria of 90% identity and 60% coverage. The search for virulence factors was performed using ABRicate, using the VFDB (Virulence Factors of Pathogenic Bacteria) database [23]. Single nucleotide polymorphism (SNP) analysis was performed using CSIPhylogeny 1.4 [24] with a minimum Z-score of 1.96 and a minimum depth at the SNP position of 10x. For this analysis, the genome of S. didelphis DSM 15616 (Accession number: SAMN02256413) was used as a reference, and the genomes of LBVP100_21 (Accession number: SAMN30684696) and LBVP101_21 (Accession number: SAMN30684697) were included for genetic comparison purposes. The phylogenetic tree was visualized using the online software iTOL applying automatic midpoint rooting [25].
Results
Animals
From July 2021 to April 2025, ten free-ranging white eared opossums were submitted to necropsy and diagnosed with S. didelphis infection based on bacterial isolation and pathologic findings. All animals were adults, three females and seven males (Table 2).
Additionally, free-ranging live opossums were captured in six urban parks to investigate S. didelphis as a part of nasal microbiota and possible association with skin lesions. Nasal swabs from 34 opossum (32 white-eared opossums and two Brazilian common opossums) were submitted to bacterial isolation and no samples were positive for S. didelphis. However, ten of the 34 captured free-ranging opossums presented skin lesions, consisting of varying degrees of ulceration and purulent exudate (Fig 1), from which S. didelphis was isolated. Eight of the S. didelphis positive opossum were adults and the other two were young. Seven of them were females and three were males. Only one of the ten positive animals was a Brazilian common opossum, the nine remaining positive animals were white-eared opossum. Table 2 describes general data (species, sex, estimated age, origin of the animal and year of sampling) from all S. didelphis positive opossum, both necropsied and captured.
Locally extensive area of ulceration with elevated borders and central purulent exudate.
Pathological findings
The S1 Table describes pathological findings associated with S. didelphis infection in the 10 necropsied opossum. Most animals (9/10) had gross lesions attributed to trauma, all of them compromised the skin integrity (lacerations or perforations). Bacteremia in these cases was characterized by the presence of bacteria intravascularly in multiple organs (Fig 2) and was observed in nine out of the 10 necropsied opossum. Most lesions were neutrophilic and necrotizing. Necrotizing and/or neutrophilic splenitis were observed in seven opossums, often associated with intralesional Gram-positive cocci (5/7) (Fig 3). Myocarditis was found in six opossums, mainly neutrophilic in most cases (5/6) it is also associated with lymphocytes and histiocytes, and in three cases with intralesional Gram-positive cocci (Fig 4). Interstitial nephritis and pyelonephritis were observed in seven animals, five of them with intralesional Gram-positive cocci (Fig 5). Cutaneous lesions were ulcerative and neutrophilic (4/10) and extended deeply to the subcutaneous tissues (Fig 6). Skeletal muscles were affected in four cases (Fig 7). In three of them large amounts of suppurative exudate were observed in the subcutaneous tissues adjacent to skin perforations (most likely due to predation). In one case (animal 2), myositis was associated with multiple rib fractures and pyothorax. All morphologic diagnosis of the necropsied opossum including lesions not associated with S. didelphis infection are described in the S1 Table.
(A) Mild multifocal gram-positive cocci intravascular in sinusoids and intracytoplasmic in Kupffer cells, liver, Didelphis albiventris, Gram stain; bar = 50 μm. (B) Moderate multifocal gram-positive cocci intravascular in glomeruli capillaries, kidney, Didelphis albiventris, Gram stain; bar = 50 μm.
(A) Severe diffuse splenomegaly with multifocal to coalescent white areas, bar = 1 cm. (B) Locally extensive necrosis delimited by fibrosis, HE; bar = 500 μm. (C) Necrosis with neutrophilic infiltrate, HE; bar = 50 μm. (D) Multifocal gram-positive cocci in an area of necrosis, Gram stain; bar = 50 μm.
(A) Moderate focal loss of cardiomyocytes and neutrophilic infiltrate, heart, HE; bar = 100 μm. (B) Gram-positive cocci associated with neutrophilic myocarditis, Gram stain; bar = 50 μm.
(A) Multiple tubules with intraluminal neutrophilic infiltrate (pyelonephritis), HE; bar = 100 μm. (B) Gram-positive cocci associated with pyelonephritis, Gram stain; bar = 50 μm.
(A) Subcutaneous tissue of the nasal planum with purulent exudate on the cut surface. (B) Focal ulcer with purulent exudate in the skin of the distal thoracic limb. (C) Dermis and subcutaneous with diffuse neutrophilic infiltrate, HE; bar = 100 μm. (d) Gram-positive cocci associated with cutaneous neutrophilic infiltrate, Gram stain; bar = 50 μm.
(A) Locally extensive purulent exudate between external and internal layers of skeletal muscle in the thoracic wall. (B) Multiple complete rib fractures with purulent exudate and hemorrhage. (C) Severe diffuse neutrophilic myositis, HE; bar = 200 μm. (d) Higher magnification of neutrophilic infiltrate in the skeletal muscle with intralesional aggregates of coccoid bacteria, HE; bar = 50 μm.
Isolation, identification, and antimicrobial susceptibility of Streptococcus didelphis
Streptococcus didelphis was isolated from many organs. Table 3 describes the isolation source and isolated bacteria from tested samples. From necropsied animals S. didelphis was mostly isolated from skin samples (6/10) as was for all captured animals (10/10).
Most of the S. didelphis isolates were sensitive to most of the antimicrobial drugs tested (Fig 8). None of the isolates were able to form biofilm in vitro (Fig 9).
AM, ampicillin; CTX, cefotaxime; CM, clindamycin; CL, chloramphenicol; ERY, erythromycin; PEN, penicillin; TET, tetracycline.
ODc – optical density cut-off value for biofilm production. Staphylococcus aureus ATCC 6538 (SA6538) and Staphylococcus epidermidis ATCC 35984 (SE35984) were used as positive controls. NC: negative control.
Genomic comparative analysis
No genetic resistance determinants, virulence factors, or plasmids were identified in the isolates from this study. SNP analysis revealed high genomic similarity among some isolates from the same location (Fig 10): Isolates from cases 1 and 3 differed by only five SNPs, while isolates 5 and 16 differed by two SNPs.
Discussion
Here we describe several cases of S. didelphis infection in free-ranging opossums with pathological, microbiological and genomic characterization. Species of the genus Streptococcus are responsible for a wide range of diseases affecting various species, including strangles, mastitis, streptococcal meningitis, pneumonia, endocarditis, sepsis, and skin infections [26–30]. Suppurative lesions and abscess formation have frequently been observed in previous reports [31–33]. S. didelphis has been isolated from the Virginia opossum (D. virginiana) [11–13] and the white-eared opossum (D. allbiventris) [14]. Here, S. didelphis was also isolated from the Brazilian common opossum.
In this study, S. didelphis infection in necropsied animals affected several organs and frequently presented with neutrophilic and necrotizing lesions accompanied by bacteremia. Most necropsied opossums had traumatic lesions compromising skin integrity, which most likely represented an entry point for S. didelphis infection. Bacteremia was a significant feature of S. didelphis infection in our cases, as also described in previous studies in Virginia opossums and white-eared opossums [11,13,14]. We also observed lesions in other organs, including myocarditis and pyelonephritis, in most animals, which were associated with intralesional Gram-positive bacteria. In live-captured opossums, S. didelphis was isolated from 10 animals with cutaneous ulcerative and purulent lesions, supporting the hypothesis that skin lesions can serve as an entry point for infection that may progress to systemic disease.
As expected, most isolates in this study were sensitive the majority of antimicrobials tested and no resistance genes were identified in the sequenced isolates, corroborating the phenotypic results obtained in this study, and the first report of the species [11]. These findings contrast with those of a recent study, in which resistance genes to β-lactams, fluoroquinolones, lincosamides, and macrolides were detected in S. didelphis isolates [14]. The absence of resistance genes and the high sensitivity observed may reflect a low level of exposure of opossums to antimicrobials, especially in less urbanized areas, such as the ecological parks from which most of the samples in this study were obtained.
Although biofilm formation capability is an important virulence factor in several Streptococcus spp. [34], such as S. mutans [35], S. pneumoniae [36], S. agalactiae [37], S. suis [38], and S. pyogenes [39], its absence in S. didelphis does not exclude its pathogenic potential, as other mechanisms may be involved in the pathogenesis of the observed infections. The presence of the bacterium in different sites in this study reinforces this possibility and indicates the need for further research on its virulence factors. Furthermore, the absence of virulence genes in the sequenced isolates may be related to the scarcity of genomic data for the species and limited annotation in databases, which hinders the identification of genetic determinants associated with its pathogenicity.
A limitation of this study is that we were unable to define the source of infection. Streptococcus spp. are well-recognized commensals of mucosal surfaces, particularly the oral cavity and upper respiratory tract, where they commonly establish stable colonization [40]. Thus, we hypothesized that the likelihood of detecting S. didelphis in swabs from these sites would be higher. In contrast, the presence of Streptococcus spp. on the skin is generally considered transient and often reflects contamination from mucosal sources rather than true colonization. Moreover, the cutaneous microbiota is predominantly composed of other genera, including Staphylococcus and Corynebacterium [41], which, in the absence of selective media for Streptococcus spp., may further reduce the likelihood of isolating Streptococcus colonies, thereby decreasing detection sensitivity.
Genomic analysis of the isolates from this study has advanced our knowledge of S. didelphis, as available data on this species remain scarce. The low variation observed in the SNP analyses of two pairs of samples from Parque Américo Renné Giannetti (1 and 3) and the Parque Fazenda Lagoa do Nado (5 and 16) suggests the possible circulation of the same clone in these locations. In a study also conducted in Brazil, high phylogenetic similarity was observed between clinical and reference strains of S. didelphis, but without evidence of clonality [14], in contrast to the results of our study. These findings reinforce the importance of molecular surveillance, even in the absence of resistance or virulence genes.
This study expands knowledge of S. didelphis infection in free-ranging opossums, focusing on its pathological, microbiological and genomic aspects. A wide range of lesions was described in association with S. didelphis infection in free-ranging opossums; however, cutaneous lesions appear to be an important early feature of the disease. The high antimicrobial susceptibility, together with the absence of resistance genes, suggests a low impact from selective pressure in the environments studied. In addition, the absence of virulence factors highlights the need for studies addressing the pathogenicity mechanisms of this bacterium. Genomic analysis suggests the circulation of clones and reinforces the importance of molecular surveillance in monitoring pathogens relevant to wild animals.
Supporting information
S1 Fig. Geographic location of areas of capture or dead opossum in Belo Horizonte (Minas Gerais, Brazil).
Map of the city of Belo Horizonte indicating: Belo Horizonte Zoo (Z); Parque Fazenda Lagoa do Nado (LN); Parque Ursulina de Andrade Mello (U); Universidade Federal de Minas Gerais campus (C); Parque Américo Renné Gianetti (AR); Parque das Mangabeiras (PM); and Parque Aggeo Pio Sobrinho (AP). Insets: bottom, Brazilian map indicating the State of Minas Gerais; middle, State of Minas Gerais indicating the area of Belo Horizonte; top, map of Belo Horizonte.
https://doi.org/10.1371/journal.pone.0348357.s001
(TIF)
S1 Table. Morphologic diagnosis of Streptococcus didelphis-associated lesions in naturally infected white-eared opossum (Didelphis albiventris).
https://doi.org/10.1371/journal.pone.0348357.s002
(DOCX)
Acknowledgments
We thank Dr. Elaine Maria Seles Dorneles from the Universidade Federal de Lavras for technical support.
References
- 1. Cáceres NC. Food habits and seed dispersal by the white-eared opossum, Didelphis albiventris, in southern Brazil. Stud Neotrop Fauna Environ. 2002;37:97–104.
- 2.
Faria MB. Guia dos marsupiais no Brasil: guia de identificação com base em caracteres morfológicos externos e cranianos. Amélie Editorial; 2019.
- 3. Braga M do SCO, Costa FB, Calchi AC, de Mello VVC, Mongruel ACB, Dias CM, et al. Molecular detection and characterization of vector-borne agents in common opossums (Didelphis marsupialis) from northeastern Brazil. Acta Trop. 2023;244:106955. pmid:37236334
- 4. Silva EM, Alves LC, Guerra NR, Farias MPO, Oliveira ELR, Souza RC. Leishmania spp. in Didelphis spp. from northeastern Brazil. J Zoo Wildl Med. 2016;47:942–4.
- 5. Bezerra-Santos MA, Nogueira BCF, Yamatogi RS, Ramos RAN, Galhardo JA, Campos AK. Ticks, fleas and endosymbionts in the ectoparasite fauna of the black-eared opossum Dipelphis aurita in Brazil. Exp Appl Acarol. 2020;80(3):329–38. pmid:31927646
- 6. Bezerra-Santos MA, Ramos RAN, Campos AK, Dantas-Torres F, Otranto D. Didelphis spp. opossums and their parasites in the Americas: a One Health perspective. Parasitol Res. 2021;120(12):4091–111. pmid:33788021
- 7. Muller G, Brum JGW, Langone PQ, Michels GH, Sinkoc AL, Ruas JL, et al. Didelphis albiventris Lund, 1841, parasitado por Ixodes loricatus Neumann, 1899, e Amblyomma aureolatum (Pallas, 1772) (Acari: Ixodidae) no Rio Grande do Sul. Arq Inst Biol. 2005;72:319–24.
- 8. Ferreira-Machado E, Conselheiro JA, Bernardes da Silva BE, Matsumoto PSS, Castagna CL, Nitsche A, et al. Naturally acquired rabies in white-eared opossum, Brazil. Emerg Infect Dis. 2023;29(12):2541–5. pmid:37987590
- 9.
Toit MD, Huch M, Cho GS, et al. The genus Streptococcus. In: Lactic acid bacteria: biodiversity and taxonomy; 2014. p. 457–505.
- 10. Haenni M, Lupo A, Madec J-Y. Antimicrobial resistance in Streptococcus spp. Microbiol Spectr. 2018;6(2). pmid:29600772
- 11. Rurangirwa FR, Teitzel CA, Cui J, French DM, McDonough PL, Besser T. Streptococcus didelphis sp. nov., a streptococcus with marked catalase activity isolated from opossums (Didelphis virginiana) with suppurative dermatitis and liver fibrosis. Int J Syst Evol Microbiol. 2000;50 Pt 2:759–65. pmid:10758886
- 12. Debrincat S, Rejmanek D, Wünschmann A, Crossley BM, Jelinski J, Armién AG. Detection of macropodid alphaherpesvirus 2 infection and lesions in sudden death of a captive Virginia opossum and a water opossum. J Vet Diagn Invest. 2024;36(4):515–21. pmid:38721879
- 13. Rebollada-Merino A, Chan TC, Guarino C, Taylor RP, Hitchener GR. Pyovagina and sepsis due to Streptococcus didelphis in a Virginia opossum. J Vet Diagn Invest. 2025;37(3):479–81. pmid:39939837
- 14. Breyer GM, Rocha Jacques da Silva ME, Slaviero M, Albuquerque de Almeida B, Machado Sousa da Silva E, de Queiroz Schmidt VR, et al. Genotypic characterization of Streptococcus didelphis causative of fatal infection in white-eared opossums. Lett Appl Microbiol. 2023;76(12):ovad131. pmid:37968138
- 15. Assis GBN, Pereira FL, Zegarra AU, Tavares GC, Leal CA, Figueiredo HCP. Use of MALDI-TOF mass spectrometry for the fast identification of Gram-positive fish pathogens. Front Microbiol. 2017;8:1492. pmid:28848512
- 16.
Clinical and Laboratory Standards Institute. Performance standards for antimicrobial susceptibility testing. 35th ed. CLSI supplement M100. Wayne (PA): Clinical and Laboratory Standards Institute; 2025.
- 17. Stepanović S, Vuković D, Hola V, Di Bonaventura G, Djukić S, Cirković I, et al. Quantification of biofilm in microtiter plates: overview of testing conditions and practical recommendations for assessment of biofilm production by staphylococci. APMIS. 2007;115(8):891–9. pmid:17696944
- 18. Bankevich A, Nurk S, Antipov D, Gurevich AA, Dvorkin M, Kulikov AS, et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. J Comput Biol. 2012;19(5):455–77. pmid:22506599
- 19. Prjibelski A, Antipov D, Meleshko D, Lapidus A, Korobeynikov A. Using SPAdes de novo assembler. Curr Protoc Bioinformatics. 2020;70(1):e102. pmid:32559359
- 20. Walker BJ, Abeel T, Shea T, Priest M, Abouelliel A, Sakthikumar S, et al. Pilon: an integrated tool for comprehensive microbial variant detection and genome assembly improvement. PLoS One. 2014;9(11):e112963. pmid:25409509
- 21. Larsen MV, Cosentino S, Lukjancenko O, Saputra D, Rasmussen S, Hasman H, et al. Benchmarking of methods for genomic taxonomy. J Clin Microbiol. 2014;52(5):1529–39. pmid:24574292
- 22. Bortolaia V, Kaas RS, Ruppe E, Roberts MC, Schwarz S, Cattoir V, et al. ResFinder 4.0 for predictions of phenotypes from genotypes. J Antimicrob Chemother. 2020;75(12):3491–500. pmid:32780112
- 23. Chen L, Yang J, Yu J, Yao Z, Sun L, Shen Y, et al. VFDB: a reference database for bacterial virulence factors. Nucleic Acids Res. 2005;33(Database issue):D325-8. pmid:15608208
- 24. Saltykova A, Wuyts V, Mattheus W, Bertrand S, Roosens NHC, Marchal K, et al. Comparison of SNP-based subtyping workflows for bacterial isolates using WGS data, applied to Salmonella enterica serotype Typhimurium and serotype 1,4,[5],12:i:. PLoS One. 2018;13(2):e0192504. pmid:29408896
- 25. Letunic I, Bork P. Interactive Tree Of Life (iTOL) v5: an online tool for phylogenetic tree display and annotation. Nucleic Acids Res. 2021;49(W1):W293–6. pmid:33885785
- 26. Mallicote M. Update on Streptococcus equi subsp equi infections. Vet Clin North Am Equine Pract. 2015;31(1):27–41. pmid:25600455
- 27. Haas B, Grenier D. Understanding the virulence of Streptococcus suis: a veterinary, medical, and economic challenge. Med Mal Infect. 2018;48(3):159–66. pmid:29122409
- 28. Liu X, Kimmey JM, Matarazzo L, de Bakker V, Van Maele L, Sirard J-C, et al. Exploration of bacterial bottlenecks and Streptococcus pneumoniae pathogenesis by CRISPRi-seq. Cell Host Microbe. 2021;29(1):107-120.e6. pmid:33120116
- 29. Lannes-Costa PS, de Oliveira JSS, da Silva Santos G, Nagao PE. A current review of pathogenicity determinants of Streptococcus sp. J Appl Microbiol. 2021;131(4):1600–20. pmid:33772968
- 30. Alexander NG, Cutts WD, Hooven TA, Kim BJ. Mechanisms and manifestations of Group B Streptococcus meningitis in newborns. J Pediatric Infect Dis Soc. 2025;14(2):piae103. pmid:39927629
- 31. Tran MP, Caldwell-McMillan M, Khalife W, Young VB. Streptococcus intermedius causing infective endocarditis and abscesses: a report of three cases and review of the literature. BMC Infect Dis. 2008;8:154. pmid:18992173
- 32. Nakashima R, Kotoku M, Gamachi A, Inagaki N, Kasaoka S. An autopsy case of fulminant, suppurative bacterial myocarditis caused by Group B Streptococcus. Intern Med. 2022;61(6):907–12. pmid:34544955
- 33. Hua C-Z, Shen Z-P, Zhou M-M, Lai C, Bai G-N, Gu W-Z, et al. Brain abscess caused by Streptococcus pyogenes with atypical symptoms: a case report and literature review. BMC Pediatr. 2024;24(1):722. pmid:39528984
- 34. Speziale P, Geoghegan JA. Biofilm formation by staphylococci and streptococci: structural, functional, and regulatory aspects and implications for pathogenesis. Front Cell Infect Microbiol. 2015;5:31. pmid:25905046
- 35. Krzyściak W, Jurczak A, Kościelniak D, Bystrowska B, Skalniak A. The virulence of Streptococcus mutans and the ability to form biofilms. Eur J Clin Microbiol Infect Dis. 2014;33(4):499–515. pmid:24154653
- 36. Domenech M, García E, Moscoso M. Biofilm formation in Streptococcus pneumoniae. Microb Biotechnol. 2012;5(4):455–65. pmid:21906265
- 37. Rosini R, Margarit I. Biofilm formation by Streptococcus agalactiae: influence of environmental conditions and implicated virulence factors. Front Cell Infect Microbiol. 2015;5:6. pmid:25699242
- 38. Wang H, Fan Q, Wang Y, Yi L, Wang Y. Rethinking the control of Streptococcus suis infection: biofilm formation. Vet Microbiol. 2024;290:110005. pmid:38280304
- 39. Brouwer S, Barnett TC, Rivera-Hernandez T, Rohde M, Walker MJ. Streptococcus pyogenes adhesion and colonization. FEBS Lett. 2016;590(21):3739–57. pmid:27312939
- 40. Nobbs AH, Lamont RJ, Jenkinson HF. Streptococcus adherence and colonization. Microbiol Mol Biol Rev. 2009;73(3):407–50, Table of Contents. pmid:19721085
- 41. Seo JY, You SW, Gu K-N, Kim H, Shin J-G, Leem S, et al. Longitudinal study of the interplay between the skin barrier and facial microbiome over 1 year. Front Microbiol. 2023;14:1298632. pmid:38033568