Leptospira and Leptospirosis: A Review of Species Classifications, Genomes, Morphological Structures, Antimicrobial Resistances, Transmissions, and Clinical Manifestations

Abstract

Leptospirosis, also known as “rat-urine disease”, is a neglected zoonotic and waterborne disease that is caused by Leptospira spp. This disease is transmitted by direct and indirect exposure to the urine and stool of infected animals. The current estimate has highlighted that leptospirosis has caused at least one million cases and 60,000 deaths, with high endemicity in tropical regions. With climate change, urbanisation, and increasing human-animal interaction, the threat of leptospirosis and other zoonotic diseases will continue to emerge. Investing in multidisciplinary research, technology, and global collaboration is critical to anticipate, detect, and respond effectively to these evolving threats.

Supplementary Information

The online version contains supplementary material available at 10.1007/s00284-026-04722-7.

Introduction

Leptospira, a genus of spirochete bacteria, causes zoonotic and waterborne disease called Leptospirosis, a neglected but emerging tropical disease that poses a significant public health risk worldwide. Leptospira spp. can be classified based on the pathogenicity into three groups, namely pathogenic, intermediate, and saprophytic, in which each pathogenicity group contains several species [1]. Known to adapt to a broad range of hosts, Leptospira spp. are acquired from direct or indirect contact with the urine of infected animals, or water and soil contaminated by these bacteria. Leptospira spp. thrive in tropical and temperate climates and have caused at least one million cases with 60,000 deaths annually [2]. Leptospira spp. were classically grouped into pathogenic, intermediate, and saprophytic clades [1]; however, advances in molecular taxonomy now recognise more complex and phylogenetic relationships and numerous species within these lineages. The global burden of leptospirosis is substantial, especially in low-resource settings where environmental conditions and limited healthcare infrastructure facilitates disease transmission [2]. This scenario causes delayed treatment when the MAT, which is the gold standard in diagnosing leptospirosis, is not practical in remote settings [3]. Given the significant global burden of leptospirosis, a thorough understanding of Leptospira spp. epidemiology is needed for developing more effective diagnostic strategies and making treatment development more accessible. Additionally, the seasonal epidemiological patterns of leptospirosis, characterised by increased incidence following heavy rainfall and flooding events, underscore the need for enhanced surveillance and timely intervention during these high-risk periods. Management of leptospirosis requires a multifaceted approach; therefore, understanding every aspect of the bacterial agent and disease is important before appropriate actions can be taken. This review consolidates contemporary insights into the species classifications, genomes, transmissions, and clinical manifestations of Leptospira spp., while evaluating current knowledge gaps and proposing new future research trajectories to more advanced evidence-based interventions for leptospirosis.

History of Leptospira

The history of Leptospira spp. dates back to Adolph Weil’s description of Weil’s disease in 1886 as an acute infectious disease with splenomegaly, jaundice, and inflammation of the kidneys [4]. Over the following decades, key discoveries included the first microscopic identification of the spirochete by Arthur Stimson in 1907, its isolation, and subsequent serovar classifications. He named the microorganism Spirocheta interrogans due to the hook at the end resembling a question mark. The microorganism was isolated by Inada and colleagues in Japan in 1915. They discovered this spirochete and found specific antibodies in the blood of miners who had contracted yellow fever [5, 6]. Almost simultaneously, two groups of doctors in Germany successfully transmitted the infection to guinea pigs and named as Spirochaeta nodosa and Spirochaeta icterogenes [7]. After detailed culture and microscopic observations, Hideyo Noguchi named it “Leptospira” (thin spirals) in 1918. From the 1920s to the 1950s, various serotypes of these spirochetes were discovered in Germany, Indonesia, and Japan. The detailed structure of Leptospira spp. was examined in the 1960s to 1970s with the aid of the electron microscope. Leptospirosis has been around for millennia, as some early disease outbreaks reported in ancient times were associated with leptospirosis. It was known before the typical aetiology was accepted. Names such as “rice field jaundice” in China, “seven-day fever” or “autumn fever” in Japan, and “schlammfieber (mud fever), “cane-cutter’s disease” or “swine fever” in Europe, Australia, and elsewhere were used to indicate leptospirosis [8, 9].

Taxonomy and Classification of Leptospira

Taxonomically, Leptospira belongs to the family Leptospiraceae, within the order Spirochaetales, class Spirochaetia, and phylum Spirochaetota [10]. The family Leptospiraceae was introduced in 1979 [11] and comprises three genera, namely Leptospira, Leptonema, and Turneriella, which were elucidated based on 16 S rRNA gene sequences, DNA-DNA relatedness, and differences in G + C content. Thus, the G + C contents of Leptospira, Leptonema, and Turneriella are 33 to 43, 54, and 53.6 mol%, respectively [12]. The taxonomy and classification of Leptospira spp. are complex and controversial [13] and are primarily classified through three main approaches: historical methods that rely on morphological characteristics and growth conditions (based on phenotypic and serological characteristics), genotypic classification based on DNA sequence data, and phylogenetic classification to elucidate evolutionary relationships [12, 14].

a. Historical (Phenotypic and Serological) Classifications

Traditionally, the genus Leptospira was divided into two species based on phenotypic classification: the pathogenic L. interrogans, comprising strains found in animals and humans, and the saprophytic L. biflexa, comprising environmental isolates Click or tap here to enter text. L. biflexa was distinguished from L. interrogans by its ability to grow at 13 °C in the presence of 8-azaguanine, and by its failure to form spherical cells in 1 M of sodium chloride [5]. However, the lack of distinguishing features often hinders phenotypic identification in routine clinical microbiological laboratories [14]. Considering the limitations, serological classification was then used to divide the two species into at least 24 serogroups and more than 300 serovars based on the expression of surface-exposed lipopolysaccharide (LPS) [15].

The structural differences in the carbohydrate component of LPS determine the antigenic diversity of the different serovars, which serve as a basic classification [13, 16]. The definition of serovar was then changed in 1986 by the Taxonomic Subcommittee on Leptospira. It is assumed that two strains belong to different serovars if, after cross-absorption with appropriate amounts of heterologous antigens in repeated tests, 10% or mre of the homologous titre routinely remains in at least one of the two antisera [12]. Antigenically related serovars are further categorised into serogroups. For example, some common serovars can be summarised under the serogroups of L. interrogans [17]. Although the term serogroup has no taxonomic meaning, it has been used to define antigenically related serovars due to the shared surface LPS structures. These serovars can be identified with the microscopic agglutination test (MAT) and are therefore widely accepted due to their epidemiological importance [18].

With the development of modern molecular typing methods, relying solely on serovar designation may be insufficient for identifying epidemiologically important strains [19], especially when molecular classifications do not align with traditional serological classification, which has served clinical microbiologists for decades. Therefore, maintaining the serological classification remains relevant until a standardised molecular-based identification is developed [5].

b. Genotypic Classifications

Phenotypic classification was then superseded by genotypic classification, with the concept of “genomospecies” were introduced to encompass the diverse serovars of L. interrogans and L. biflexa. This genomospecies was first established using DNA-DNA hybridisation in the late 20th century, which showed that serologically similar isolates were often genetically different, leading to the description of several new species [20]. While DNA-DNA hybridisation was foundational, 16S rRNA gene sequencing and multilocus sequence typing (MLST) emerged as more practical and widely adopted molecular tools for Leptospira genotyping [21].

16S rRNA gene sequencing targets 16S ribosomal RNA, often a highly conserved gene present in all bacteria, supporting a three-group model in which Leptospira spp. can be categorised as pathogenic (Group I), intermediate or opportunistic (Group II), and saprophytic or non-infectious (Group III) (Fig. 1) [21]. Group I causes diseases of varying severity in humans, ranging from asymptomatic infections to severe complications and death. Group II pathogens, on the other hand, survive better in culture and lead to a mild, self-resolving disease without causing death. They are considered opportunistic because of insufficient information on the involvement of these species in human leptospirosis, unclear pathogenicity, and the phenotypic characteristics do not match those of pathogenic Leptospira spp [22]. The pathogenicity status of Group II Leptospira remains controversial. A review highlighted that animal models have primarily been developed for pathogenic Leptospira and that chronic or subclinical infections may occur without severe clinical disease in some hosts, indicating the complexity of disease manifestations across species [23]. Whilst early animal studies, such as those involving hamsters inoculated with intermediate L. inadai and L. licerasiae did not result in clinical manifestations [24, 25]. However, more recent phylogenomic and epidemiological work has emphasised the ambiguous nature of these species (i.e. genetically closer to pathogenic clades yet associated with milder human disease, thereby blurring the lines of their classification) [26]. Leptospira spp. of Group III are free-living environmental microorganisms and do not cause disease in humans and animals [25].

Fig. 1.

Previous classification of Leptospira species based on the 16S rRNA gene sequences [12, 25]

However, the conserved nature of the 16S rRNA gene lacks the resolution to differentiate between very closely related species within the pathogenicity group and is unsuitable for discriminating serovars or strains. MLST was developed to overcome this limitation by sequencing multiple “housekeeping’ genes (i.e. genes essential for basic cell function that are found in all strains). Due to a higher rate of evolution of these genes than the 16S rRNA gene, they can be used for better discrimination of the strains or serovars, allowing for intraspecies discrimination and clonal complex identification [20].

c. Phylogenetic Classifications

The phylogenomic classification of Leptospira spp. has evolved considerably with the advent of molecular and genome-wide analyses. The latest discovery through whole-genome sequencing (WGS) has significantly expanded the taxonomic landscape by identifying 74 validly described species (Table 1; Fig. 2) (instead of 21 species from genotypic classification) and enabling high-resolution comparisons of core and ancillary genomes [26]. Whole genome sequencing (WGS) has also revealed genomic signatures of virulence and ecological adaptation, highlighting the limitations of purely serological classification.

Table 1.

Updated list of 74 recognised Leptospira species classified into four established phylogenetic clades (P1, P2, S1, and S2) as described in [27]

Clade	Description	Leptospira species
P1 clade	Pathogenic, high-virulence, and responsible for the majority of severe human and animal leptospirosis cases	21 species: L. adleri, L. ainazelensis, L. ainlahdjerensis, L. alexanderi, L. alstonii, L. barantonii, L. borgpetersenii, L. ellisii, L. gomenensis, L. gorisiae, L. interrogans, L. kirschneri, L. kmetyi, L. mayottensis, L. noguchii, L. santarosai, L. sanjuanensis, L. stimsonii (synonym of L. putramalaysiae), L. tipperaryensis, L. weilii, and L. yasudae (synonym of L. dzianensis),
P2 clade	Intermediate pathogenic, capable of causing diseases but are generally associated with milder clinical outcomes.	22 species: L. andrefontaineae, L. broomii, L. cinconiae, L. dzoumogneensis, L. fainei, L. fletcheri, L. fluminis, L. haakeii, L. hartskeerlii, L. inadai, L. johnsonii, L. koniamboensis, L. langatensis, L. licerasiae, L. neocaledonica, L. perolatii, L. saintgironsiae, L. sarikeiensis, L. selangorensis, L. semungkisensis, L. venezuelensis, and L. wolffii
S1 clade	Non-pathogenic to humans and animals, free-living species that are common in water and soil. Widespread in soil and water. May have certain groups of genes that are not found in S2 clade.	26 species: L. abararensis, L. bandrabouensis, L. biflexa, L. bourretii, L. bouyouniensis, L. brenneri, L. chreensis, L. congkakensis, L. ellinghausenii, L. harrisiae, L. iowaensis, L. jelokensis, L. kanakyensis, L. kemamanensis, L. levettii, L. meyeri, L.mgodei, L. milleri, L. montravelensis, L. mtsangambouensis, L. noumeaensis, L. perdikensis, L. terpstrae, L. vanthielii, L. wolbachii, and L. yanagawae
S2 clade	Non-pathogenic to humans and animals. Smaller species group compared to S1 clade and may occupy more specialized niches.	5 species: L. idonii, L. ilyithenensis, L. kobayashii, L. ognonensis, and L. ryugenii.

Fig. 2.

The 74 currently recognised Leptospira species cluster into four subclades: P1 (pathogenic, high virulence), P2 (intermediate pathogenicity), S1 (non-pathogenic saprophytes), and S2 (non-pathogenic saprophytes). The phylogenetic tree was inferred from Mash distance matrices [28] using FastME [29]. Outgroup branch length (Leptonema illini) was shortened for visualisation purposes. The phylogenetic tree was constructed based on whole-genome assemblies retrieved from NCBI as provided in the Supplementary material

Subsequent application of WGS and phylogenomic analyses refined this taxonomy, dividing Leptospira spp. into four distinct clades: P1 (pathogenic, high virulence), P2 (intermediate pathogenicity) and the saprophytic clades S1 and S2 (Fig. 2) [26, 27]. S2 forms a distinct branch within the saprophytic lineage, separated from S1, and exhibits greater genetic divergence than S1 [26]. This four-class system is now widely accepted as the most robust representation of the evolutionary history of Leptospira spp., accounting for both genomic diversity and pathogenic potential, with approximately 74 validly described species distributed across the clades [27].

Genomic Features of Leptospira spp.

The genome of Leptospira is relatively large, as it normally ranges in size from approximately 4.6 to 5.1 megabases (Mb) (Table 2). One of the unique genomic structures of Leptospira is the presence of two circular chromosomes, which are rather unusual among spirochetes and most prokaryotes, reflecting the ecological and evolutionary complexity of Leptospira [16]. The two-chromosomal structure is most likely the result of genome plasticity and the retention of niche-associated characteristics, which may impact host interactions and virulence. Chromosome I is ~ 4.3 Mb in size and contains most housekeeping genes and many virulence-associated genes, whilst chromosome II is ~ 350–400 kilobases (kb) and behaves as a secondary chromosome (chromid) that carries genes related to accessory functions, environmental sensing, and host adaptation [29].

Table 2.

Comparative genomic features of P1, P2, and S1/S2 Leptospira spp.

Genomic feature	Strains of P1 clade*	Strains of P2 clade*	Strains of S1 or S2 clades*	Reference
Genome size	~4.6–5.1 Mb	~4.0–4.3 Mb	~3.9–4.1 Mb	[5, 30]
Chromosome I (carries housekeeping and many virulence genes)	Present	Present	Present	[29]
Chromosome II (carries genes of accessory and essential functions)	Present	Present	Present	[30]
Pan-genome feature	Most open pan-genome	Open	Least open	[16, 26]
	Frequent gene acquisition via horizontal gene transfer.	Slower rate of gene acquisition via horizontal gene transfer than P1 clade.	Limited gene acquisition via horizontal gene transfer.
Virulence factors	LipL32, LigA/B, hemolysins, adhesins, CRISPR-Cas	Partial presence	Absent	[16, 25, 31]
Genomic islands/pathogenicity islands	Abundant, encode virulence and immune evasion genes	Occasional	Absent	[16]
Mobile elements (insertion sequence and transposons)	The most abundance of mobile elements.	Fewer abundance of mobile elements than P1 clade.	Least abundance of mobile elements.	[32]
Antimicrobial resistance genes	At least 32 genes	Rare	Absent	[16]
Pseudogenes	Numerous, due to genome decay linked to host adaptation	Moderate, reflecting transitional ecology	Very few, stable genome needed for diverse environmental survival	[32]
Environmental adaptation genes	Osmoregulation, oxidative stress, nutrient uptake	Some	Basic core only	[33]
Host adaptation mechanisms	Plasminogen binding (e.g., LIC11711), sialic acid, B12 biosynthesis	Limited	Absent	[32]
Genomic plasticity	High; associated with ecological versatility	Moderate	Low; high stability	[34]
Genetic tools	Scarce; improved via transposon mutagenesis	Minimal, genetic manipulation remains poorly developed	Moderate, more tractable for genetic manipulation	[35, 36]
Unique genes	The most abundance of strain-specific genes. 627 genes not found in L. biflexa, 500 + uncharacterised, many linked to virulence and host adaptation	Fewer abundance of strain-specific genes than P1 clade. Partial overlap pathogenic strains; reduced virulence repertoire; transitional adaptation	The least abundance of strain-specific genes. Lacks pathogen-specific genes, enriched in metabolic and environmental survival functions	[37, 38]

^*as representated by the Leptospira strains reported by Dellagostin et al. [39]

The role of some genes responsible for the pathogenesis of pathogenic Leptospira spp. is still not fully understood. For example, L. interrogans comprises 627 genes that are not found in the genome of L. biflexa. As the functions of more than 500 of these genes are unknown, it is hypothesised that additional genetic traits are required for survival in mammalian hosts and in the environment [37]. Although research on Leptospira spp. has entered the post-genomic era, genetic studies still lag behind other pathogenic bacteria [35]. Nevertheless, access to genome sequences and the development of mutagenesis systems, especially transposon mutagenesis, have enabled the elucidation of the phases of pathogenesis [36]. Comparative genomic analyses demonstrated that pathogenic strains have large repertoires of virulence determinants, including adhesins, hemolysins, CRISPR-Cas loci, and complex LPS biosynthesis clusters [16].

These virulence determinants are normally harboured in genomic islands and associated with insertion sequence (IS) mobilisation capability. Saprophytic L. biflexa, on the other hand, did not possess any of the pathogen-specific genes and had a lower number of pseudogenes and mobile elements [32]. Pathogenic Leptospira strains (P1 and P2 clades) also show a more open pan-genome structure indicative of frequent horizontal gene transfer, with elements such as vitamin B12 biosynthesis and sialic acid biosynthesis increasing host adaptation potential. In contrast, non-pathogenic strains (S1 and S2 clades) have the least open genome that corresponds to their stable environmental niches [16]. These genetic patterns reflect their evolutionary trajectory in which pathogenic strains evolve genome plasticity and specialisation through the acquisition of virulence encoding modules while saprophytic species maintain genome integrity to support their free-living lifestyle [34]. Recognising these distinctions is important for identifying potential diagnostic targets, vaccine antigens (i.e. Lig and LipL proteins), and further describing environmental versus host-adapted in Leptospira spp.

Comparative genomics is important in understanding the pathogenicity, resistance, and environmental adaptation in Leptospira spp [40]. By comparing the genomes of pathogenic versus saprophytic strains, several genomic regions have been identified as putative pathogenicity islands (PAls), revealing virulence-associated features that are predominantly found in highly pathogenic species such as L. interrogans and L. kirschneri [25]. These PAls have genes encoding surface-exposed adhesins (e.g., LipL32, LigA/B), as well as hemolysins, metalloproteases, and other proteins that interfere with the host’s bactericidal pathways and facilitate immune invasion. These proteins are important for successful host-pathogen interactions [31]. One notable example is LIC11711, a gene characteristic of pathogenic L. interrogans. Heterologous expression of this gene in the saprophyte L. biflexa markedly increased bacterial adhesion to host laminin and plasminogen, suggesting a role of this gene in pathogenic strains in tissue colonisation and evasion of the immune responses [32].

Antimicrobial Resistance

Studies over several decades have investigated the antimicrobial susceptibility of Leptospira species using both culture-based and minimum-inhibitory concentration methods (Table 3), as well as in vitro and in silico methods (using CARD, RAST, PATRIC/BV-BRC for AMR detections) to understand the resistance mechanisms (Table 4). Early studies reported different resistance patterns: for example, L. biflexa showed resistance to streptomycin despite sensitivity to oxytetracycline, chloramphenicol, kanamycin, dihydrostreptomycin, and ampicillin [41], while L. borgpetersenii isolated from cattle was resistant to sulfamethazine but sensitive to penicillin, tetracycline, ampicillin, erythromycin, and streptomycin [42]. More comprehensive assessments later confirmed that pathogenic species such as L. interrogans, L. borgpetersenii, L. kirschneri, L. noguchii, L. santarosai, and L. weilii generally remain sensitive to β-lactams, macrolides, tetracyclines, chloramphenicol, and fluoroquinolones, but show reduced sensitivity to aztreonam [38]. Recent surveys of clinical and environmental isolates show consistent susceptibility to β-lactams, doxycycline, and fluoroquinolones, although resistance to neomycin, fosfomycin, vancomycin, and especially sulfamethoxazole-trimethoprim has been increasingly reported in L. interrogans and L. borgpetersenii [43–46]. Regional studies have also shown differences in resistance, with some isolates showing lower susceptibility to polymyxin B, gentamicin, chloramphenicol, and rifampicin [52, 53]. A large-scale review of isolates collected between 1948 and 2016 confirmed that while penicillins, cephalosporins, tetracyclines, and fluoroquinolones remain largely effective, resistance to polymyxins is widespread in various Leptospira species and hosts [47]. Taken together, these results show that while traditional first-line antibiotics remain largely effective against Leptospira, emerging resistance, particularly to aminoglycosides, folate pathway inhibitors, and polymyxins, warrants continued surveillance. However, there is no standardised clinical breakpoint for Leptospira. Resistance interpretations vary across the studies.

Table 3.

Antimicrobial susceptibility and resistance profiles of different Leptospira spp. from various sources

Leptospira species	Year	Main source	Method	Susceptible antibiotics	Resistant antibiotics	Reference
L. biflexia	1988	Culture strain	MIC	Oxytetracycline, chloramphenicol, kanamycin, dihydrostreptomycin, ampicillin	Streptomycin	[41]
L. borgpetersenii	1988	Cattle	MIC	Penicillin, tetracycline, ampicillin, erythromycin, streptomycin	Sulfamethazine	[42]
L. interrogans, L. borgpetersenii, L. kirschneri, L. noguchii, L. santarosai, L. weilii	2004	Culture strain	MIC	Penicillin and other β-lactams, macrolides, chloramphenicol, tetracycline, doxycycline, fluoroquinolones, telithromycin	Aztreonam	[38]
L. interrogans, L. borgpetersenii	2010	Rodent	MIC	Ampicillin, cefotaxime, fluoroquinolones, doxycycline, erythromycin, streptomycin	Neomycin, fosfomycin, sulfamethoxazole, trimethoprim, vancomycin	[43]
L. interrogans	2013	Human, dog, rodent, cattle	MIC	Penicillin and other β-lactams, fluoroquinolones, tetracyclines, gentamicin	Sulfamethoxazole-trimethoprim, neomycin	[44]
L. interrogans	2015	Rodent, dog	MIC	Amoxicillin, ceftriaxone, ciprofloxacin, clindamycin, doxycycline, erythromycin, imipenem, penicillin, polymyxin B	Polymyxin B, gentamicin, sulfamethoxazole-trimethoprim	[35]
L. interrogans, L. borgpetersenii, L. kirschneri, L. weilii	2015	Not available	Disc susceptibility test	Amoxicillin, azithromycin, cefoxitin, ceftazidime, ceftriaxone, chloramphenicol, ciprofloxacin, clindamycin, doripenem, doxycycline, gentamicin, linezolid, nitrofurantoin, penicillin, piperacillin/tazobactam, tetracycline	Sulfamethoxazole-trimethoprim, fosfomycin, nalidixic acid, rifampicin	[35]
L. interrogans, L. borgpetersenii, L. meyeri, L. santarosai	2016	Human, dog, rodent, cattle, swine	MIC	Penicillin, ampicillin	Sulfamethoxazole-trimethoprim	[45]
L. interrogans, L. borgpetersenii	2017	Dog, rodent, human, swine, water	MIC	Penicillin, ampicillin, doxycycline	Sulfamethoxazole-trimethoprim, chloramphenicol	[46]
L. borgpetersenii, L. broomi, L. interrogans, L. kirschneri, L. noguchii, L. santarosai	1948–2016	Various (swine, cattle, human, deer, and donkey)	MIC	Penicillin, amoxicillin, clavulanate, cephalexin, ceftriaxone, doxycycline, tetracycline, streptomycin, enrofloxacin, spectinomycin	Polymyxin	[47]

Table 4.

Antibiotic resistance mechanisms found across Leptospira spp. based on in silico and in vitro studies

Type of antimicrobial resistance	Resistance mechanism	Antibiotic class affected	Study type	Leptospira group/clade	Reference
Aminoglycoside-modifying enzyme	Resistant Leptospira produces aminoglycoside acetyltransferase (N6’ac gene) that inactivates aminoglycoside antibiotics (e.g. gentamycin, tobramycin) by acetylation, therefore preventing them from binding ribosomes that can interfere protein synthesis	Aminoglycosides	In silico	P1, S1	[48]
Folate-pathway bypass	Mutations of dihydropteroate synthase (DHPS) or dihydrofolate reductase (DHFR) or acquisition of alternative enzymes to bypass inhibition of folate synthesis induced by trimethoprim and sulfonamides. The reduced binding of trimethoprim and sulfonamides causes continued folate synthesis.	Trimethoprim and Sulfonamides	In silico	P1	[49]
Multidrug efflux pump system	Efflux pumps are natural transport proteins in bacterial membranes, responsible for exporting antibiotics out of bacterial cell. This efflux pump system consists of: - Resistance-Nodulation-Division (RND) pump - Multidrug And Toxic compound Extrusion (MATE) pump - Pump components: TolC, AcrB, MacB, NodT - Regulators (Reg)	Multiple classes	In silico	P2, S1, S2	[48]
Ribosomal target mutation	Mutations in the gene (encoding ribosomal protein S12) or the rpsJ gene (encoding ribosomal protein S10) reduce aminoglycoside binding, thereby preventing disruption of protein synthesis.	Aminoglycosides	In vitro	Saprophytic	[41]
			In silico	Pathogenic	[49]
Vancomycin resistance genes	vanT, vanW, vanY genes alter peptidoglycan precursors (D-Ala-DlLac instead of D-Ala-DlAla), reducing vancomycin binding	Glycopeptides (Vancomycin)	In silico	P1, P2, S1, S2	[48]
β‑lactamase enzyme production	β-lactamase is an enzyme that hydrolyses the β-lactam ring of penicillins, cephalosphorins, and carbapenems, that can render them inactive.	β‑Lactams	In silico	P1, P2, S1, S2	[48]
			In silico	Pathogenic	[50]
Cell-wall target alteration	Mutations in alr (alanine racemase) and ddl (D-Ala–D-Ala ligase) reduce binding of D-cycloserine, impairing inhibition of peptidoglycan biosynthesis	Cycloserine (cell-wall synthesis inhibitors)	In silico	P1	[51]
Cell-wall target alteration	Alteration in murA (UDP-N-acetylglucosamine enolpyruvyl transferase) reduces binding of fosfomycin, preventing inhibition of cell-wall precursor synthesis	Fosfomycin	In silico	P1	[51]
DNA gyrase/topoisomerase target mutation	Mutations in gyrA and gyrB reduce fluoroquinolone binding to DNA gyrase, impairing inhibition of DNA replication	Fluoroquinolones	In silico	P1	[51]
RNA polymerase target mutation	Mutations in rpoB and rpoC alter rifampicin binding to RNA polymerase β-subunits, preventing inhibition of transcription	Rifamycins	In silico	P1	[51]
Translation factor alteration	Alterations in ef-G (elongation factor G) and ef-Tu (elongation factor Tu) impair interaction with protein-synthesis inhibitors	Protein synthesis inhibitors (various)	In silico	P1	[51]
Loss of rRNA methyltransferase	Absence or inactivation of gidB (16 S rRNA methyltransferase) causes low-level streptomycin resistance by reducing ribosomal binding	Aminoglycosides (streptomycin)	In silico	P1	[51]
Cell-wall charge modification	Genes gdpD and pgsA modify outer-membrane lipopolysaccharides (LPS), altering surface charge and reducing uptake of cationic antibiotics	Polymyxins, Aminoglycosides	In silico	P1	[51]

Comparing these phenotypic data with genomic features provides a basis for identifying mechanisms of AMR (Table 4). Several in silico and in vitro studies demonstrated that AMR in Leptospira is multifactorial and includes enzymatic inactivation, target site modification, efflux-mediated resistance, cell envelope changes, and metabolic evasion. Aminoglycoside-modifying enzymes such as acetyltransferases encoded by N6’ac prevent ribosomal binding of gentamicin and tobramycin, while β-lactamases, which can hydrolyse penicillins, cephalosporins, and carbapenems, render β-lactams ineffective [48, 50]. Resistance is also mediated by mutations in ribosomal proteins (rpsL, rpsJ), deletion of gidB, and alterations in alr, ddl, murA, gyrA/gyrB, and rpoB/rpoC, which impair the activity of aminoglycosides, streptomycin, cycloserine, fosfomycin, fluoroquinolones, and rifamycins, respectively. Rifamycins; modifications of the elongation factors EF-G and EF-Tu additionally impair the inhibition of protein synthesis [41, 49]. Efflux-mediated resistance is supported by transporters of the RND and MATE family as well as accessory proteins (TolC, AcrB, MacB, NodT), which excrete various antimicrobial substances and reduce intracellular drug levels [48]. Structural defence mechanisms include LPS modifications that limit polymyxin uptake, phospholipid alterations by gdpD and pgsA mutations that alter surface charge and confer cross-resistance to polymyxins and aminoglycosides, and glycopeptide resistance mediated by vanT, vanW, and vanY that remodel peptidoglycan termini [51]. Finally, intrinsic resistance to inhibitors of the folate pathway arises from changes in DHPS and DHFR or from the acquisition of alternative enzymes that bypass the inhibition induced by trimethoprim and sulfonamides and maintain folate biosynthesis despite drug exposure [49]. Plasmids and prophages are also responsible in the horizontal gene transfer in bacteria, enhancing the spread of such resistance determinants [35].

Morphological Structures of Leptospira spp.

Leptospira spp. (from the Greek leptos, meaning “fine”, and “spira”, meaning “a spiral”) are finely coiled, filamentous spirochetes. They are extremely thin, helically coiled spirochetes typically measuring 6–20 μm in length and approximately 0.1 μm (100 nm) in diameter [49, 54]. Certain species have been reported to reach lengths of about 15.1 μm and diameters of about 0.12 μm, with a wavelength of ~ 0.6 μm [55]. Leptospira spp. stain poorly with conventional aniline dyes, including Gram stain, and are too thin to be visible under a normal light microscope [4]. They are typically visualised using dark-field or phase-contrast microscopy. Leptospira spp. are corkscrew-shaped bacteria with “question mark”-hook-shaped ends that distinguish them from other spirochetes [22].

Leptospira spp. have a double-membrane envelope, similar to other Gram-negative bacteria but with distinct spirochetal features (Fig. 3A). The outer membrane contains numerous lipopolysaccharides (LPS) and a variety of outer-membrane proteins: LPS of many Gram-negative bacteria exhibits endotoxin activity that can activate TLR4; leptospiral LPS, however, exhibits low endotoxin activity and structurally unusual lipid A that signals through TLR2 instead of TLR4. This alters the innate immune recognition (whilst LPS of most Gram-negative bacteria usually triggers TLR4) and potentially enables the pathogen to partially evade typical endotoxin-driven responses while still inducing inflammation [55, 56]. The outer-membrane proteins include classical barrel porins (e.g., OmpL1), which contribute to nutrient exchange and host interaction [55], as well as many surface-exposed proteins involved in adhesion, immune evasion, and virulence. These include LipL41, LigA-LigC, LenA-LenF, and Lsa lipoproteins that bind a range of extracellular matrix (ECM) components such as fibronectin, laminin, collagen, and fibrinogen [57–60].

Fig. 3.

A Schematic representation of the Leptospira cell envelope. The outer membrane contains surface-exposed and internal lipopolysaccharides (LPS), and the transmembrane porin OmpL1. Beneath it, the periplasm contains the endoflagellum, peptidoglycan layer. B Cross-sectional view showing the outer membrane, peptidoglycan layer, cytoplasm, and the distinctive periplasmic flagellum characteristic of Leptospira. C Schematic representation of the spiral Leptospira cell with the periplasmic endoflagella (green, purple, yellow) anchored to the cell pole and extending along the protoplasmic cylinder under the outer membrane. D Organisation of periplasmic endoflagella originating from flagellar motors located at both ends of the cell and spreading towards the centre of the cell, giving rise to the characteristic helix and hook morphology. E Molecular architecture of the endoflagellum, consisting of a core (FlaB1–FlaB4) surrounded by a sheath composed of the proteins FlaA1, FlaA2, FcpA, and FcpB, which together regulate the stability and function of the filament. F Model of the asymmetric organisation of the envelope, with convex and concave envelope proteins promoting curvature and helical cell shape (Fig. 3C−F were adapted from Gibson et al. [56] and used under the terms of the Creative Commons Attribution License (CC BY) license)

Within the periplasm, several internal lipoproteins of the outer membrane, such as LipL32 and LipL36, are located in the periplasmic face of the outer membrane; these proteins trigger inflammatory responses and are hypothesised to contribute to envelope stability [61]. Another periplasmic lipoprotein, LipL21, binds tightly to the peptidoglycan layer and protects it from enzymatic fragmentation, thereby preventing the release of muropeptides that would otherwise activate innate immune receptors [62]. The peptidoglycan layer itself lies unusually close to the inner membrane, is more reminiscent of a Gram-positive structural envelope, that plays an essential structural role in stabilising the elongated helical cell body, resisting mechanical deformation during motility, and providing an anchoring scaffold for the periplasmic endoflagella (Fig. 3A&B) [59].

A defining feature of Leptospira spp. is the presence of two periplasmic endoflagella, one located at each pole (Fig. 3C and F) [39]. The molecular architecture of these endoflagella is well characterised, consisting of a core (composed of FlaB1–FlaB4 subunits) surrounded by a sheath (composed of FlaA1, FlaA2, FcpA, and FcpB), which together regulate the stability, coiling, and mechanical properties of the filament [4, 5]. These periplasmic endoflagella are responsible for the motility of Leptospira spp. and are associated with pathogenicity, through coordinated rotation of both the endoflagella and the helical cell body, enabling the bacteria to switch between swimming in the liquid (undulatory or wave-like movement) and crawling on surfaces [58, 60, 61].

Isolation, Growth, and Cultivation of Leptospira spp.

The primary isolation of pathogenic Leptospira spp. from clinical samples (e.g. mammalian tissues, blood, or urine) is notoriously difficult. Cultures may require prolonged incubation of 6 to 13 weeks due to slow growth rate [5], low bacterial loads in clinical samples, and heightened risks of contamination [62, 63]. Environmental samples (e.g. water and soil) can be more challenging because saprophytic Leptospira naturally grow easily and rapidly compared to pathogenic Leptospira, affecting the downstream data analysis. For example, a study using Oxford Nanopore Technology (ONT) metagenomic sequencing on enriched cultures from soil and water found most sequencing reads (~ 98–99% reads) comprised saprophytic Leptospira compared to pathogenic species (~ 1–2% reads) [64, 76]. Once isolated, Leptospira typically grow more rapidly in subsequent in-vitro cultures, often reaching maximal cell density within 3 to 4 weeks under optimal conditions. In a semi-solid medium, growth concentrates in a distinct subsurface zone that progressively becomes more turbid with increasing incubation time. This growth is known as Dinger’s ring or disc, an opaque region associated with optimal oxygen tension [5].

Outside the host, the most optimal conditions for the survival of Leptospira spp. are a humid environment with neutral pH and a temperature range of 20 to 32 °C [65]. In the environment, they can survive from a few weeks to almost a year in moist soil during the dry season or in surface water during the rainy season [17]. Leptospira spp. are obligate aerobes that require aeration for maximum growth [4]. When grown in suitably aerated culture media at pH 7.2–7.6 and temperatures of 28–30 °C, the generation time of Leptospira spp. varies considerably among species. Pathogenic L. interrogans typically exhibits a generation time of approximately 8–12 h [4], whereas many other species replicate more slowly under comparable conditions.

Growth and cultivation characteristics of Leptospira spp. also differ considerably based on their pathogenicity. Pathogenic Leptospira are more fastidious, with slower growth rates, and demonstrate limited ranges of temperature and pH tolerance [5], while saprophytic Leptospira grow more steadily and proliferate at lower temperatures [66]. Pathogenic Leptospira are susceptible to 8-azaguanine, whilst saprophytic species are resistant [67]. Taken together, these explain why screening and isolation of pathogenic species are more challenging.

Leptospira spp. can be cultivated using a range of liquid (broth), semi-solid (0.1–0.2% agar) [68], or fully solid media (1–1.2.2% agar) [69], which these physical forms are based on the type of media used, since some media are limited to certain forms only (e.g. only fully solid or only liquid and semi-solid). Liquid media are used for routine selective enrichment and maintenance, biomass production, metabolic studies, and preparation of antigens or nucleic acids. Semi-solid media [typically EMJH (Ellinghausen-McCullough-Johnson-Harris)] support primary isolation since motile leptospires form a characteristic subsurface Dinger’s zone, enabling early detection from low inocula and separation from contaminants. Fully solid media [e.g. EMJH, Leptospira Vanaporn Wuthiekanun (LVW), and Hornsby-Alt-Nally (HAN) agars [47, 70, 71] enable visible colony formation that is essential for the selection of clonal isolates, genetic mutants, and antimicrobial susceptibility testing, as well as allow more rapid growth and long-term preservation.

Among these, the most widely used formulation is EMJH medium, supplied with bovine serum albumin or prepared in a protein-free form. EMJH medium is routinely used as a broth or as a semi-solid, for primary isolation and maintenance [72]. EMJH medium supplemented with selective antimicrobial agents has been reported to perform as well as or better than other commonly used media. Several comparative studies have justified that EMJH medium outperforms other media, including Gorman et al. [64] who found EMJH medium supplemented with antimicrobial agents outperforms other media in isolating and detecting Leptospira spp. from water and soil samples. Zarantonelli et al. [73] found that EMJH medium outperforms Fletcher’s medium for isolating Leptospira in rodents. Steinparzer et al. mentioned that EMJH medium supplemented with STAFF antibiotics (sulfamethoxazole, trimethoprim, amphotericin, fosfomycin, and 5-fluorouracil) effectively enhances leptospiral isolation from swine urine. Since Leptospira spp. are intrinsically tolerant to 5-fluorouracil, this antibiotic is usually added to the culture media to prevent contamination and enhance the isolation of Leptospira spp [70]. Other than EMJH, there are alternative culture media: i) classical serum-rich formulations (e.g. Stuart, Korthof, Fletcher, and Noguchi media); and modern EMJH-derived formulations (e.g. EMJH-supplemented with pyruvate/serum, LVW, and HAN media) [70, 71, 74]. These media with improved modern EMJH-derived formulations have been developed to improve the isolation of fastidious or clinical strains. Therefore, the choice of media (considering the form and formulations of media) depends on the objectives of the laboratory procedure, the Leptospira species/serovars, and feasibility.

The nutritional requirements of Leptospira spp. are minimal, with long-chain fatty acids serving as the principal carbon and energy sources via β-oxidation, supplemented by vitamins B₁, B₁₂ as growth factors, and ammonium salts as cellular nitrogen [75]. Leptospira spp. consistently exhibit catalase and oxidase activity, whereas urease production is restricted to only some species, and lipase activity is widely observed across the genus [71]. The cultures are usually preserved by repeated subculturing or storage in semi-solid agar with haemoglobin. Other than fully solid medium (LVW agar), liquid nitrogen is widely used for the long-term storage of Leptospira spp. because it preserves viability, motility, antigenicity, and virulence over extended periods [47].

Transmission Pathways of Leptospira Involving Animals and the Environment to Humans

Rodents play a central role as an important maintenance host of Leptospira spp. that can contaminate soil and surface water (Fig. 4) [1, 76, 77]. A wide range of domestic animals, such as dogs, cattle, pigs, goats, and horses, and diverse wild animals, such as bats, small mammals, primates, and ungulates, also serve as reservoirs or incidental hosts, contributing further to environmental contamination through urinary and faecal shedding [78]. Pathogenic Leptospira survive for a prolonged duration of weeks to months in warm, humid environments with neutral pH, and their persistence is enhanced by biofilm formation in water and soil [79, 80]. Human infection occurs predominantly through indirect exposure to contaminated water, mud, or soil during flooding, agricultural activities, or recreational freshwater contact, whilst direct exposure to infected animals remains important among high-risk occupational groups such as farmers, veterinarians, and sewage workers [77]. Humans are incidental hosts and rarely contribute to onward transmission, as person-to-person spread is extremely uncommon [81].

Fig. 4.

Leptospira transmission occurs through a complex interplay of animal reservoirs, environmental persistence, and human exposure, with rodents serving as the primary maintenance hosts that chronically shed pathogenic Leptospira in urine and faeces, contaminating soil and surface water. Blue arrows indicate transmissions from rodents (the maintenance hosts) to other animals and the environment. Grey arrows indicate transmissions from animals to the environment, or bidirectional transmission between wild animals to domestic animals. Green arrows indicate environmental transmission to the animals. Dashed green arrows indicate indirect transmission from a contaminated environment to humans. Red arrows indicate direct transmissions from infected animals to humans

The most common entry points are cuts and abrasions on the skin, intact mucous membranes of the eyes, nose, or throat, and waterlogged skin. Leptospira spp. can also enter the body by inhaling aerosols containing Leptospira or through drinking water, especially in high-risk environments (e.g. contaminated water and animal facilities) [25, 82]. Once established in the renal tubules, particularly the proximal convoluted tubules, different animal hosts shed leptospires for markedly different durations. Rodents, especially rats, can shed large numbers of leptospires throughout their lifespan [83], whereas livestock such as cattle can shed organisms for months to years [84], and domestic pets, such as dogs, can shed intermittently even when clinically asymptomatic [85–87]. In the pathogenesis of leptospirosis, the first step is the penetration of tissue barriers to enter the human body [88, 89]. Environmental factors strongly modulate transmission dynamics, such as heavy rainfall, flooding, and warm humid conditions, which enhance leptospiral survival in soil and water, facilitate their spread across wider areas, and amplify human exposure during outbreak periods [90]. Collectively, these ecological, host-related, and environmental factors interact to sustain the transmission cycle and drive leptospirosis emergence in endemic regions. The multi-host, multi-reservoir transmission system highlights the importance of a One Health approach integrating environmental management, rodent control, livestock vaccination, and improved sanitation to disrupt transmission pathways linking wildlife, domestic animals, contaminated environments, and human infection.

Clinical Manifestations of Leptospirosis

Leptospirosis is characterised by a broad spectrum of clinical manifestations that often mimic other infectious diseases (Table 5) [79]. In humans, it is usually misdiagnosed as influenza, aseptic meningitis, encephalitis, and dengue fever [14]. Jaundice (icterus) is a well-known symptom of leptospirosis that can also be observed in other liver-related diseases such as hepatitis [80]. The severity of leptospirosis is influenced by the size of the inoculum, strains of Leptospira spp., and the patient’s health status and age [81]. One of the difficulties in eliminating the consequences of a Leptospira infection is the condition of the kidney carrier, which can last from months to years. Although uncommon, leptospirosis may progress to a fulminant and severe disease state, in which case mortality can range between 5 and 40% [91]. More than 90% of leptospirosis cases are mild and self-limiting, whilst multi-organ failure occurs in less than 10% of individuals [23].

Table 5.

Clinical phases, symptoms, and key features of leptospirosis

Phase/form		Symptoms and clinical features	Notes
General/overall		- Broad spectrum, mimics influenza, dengue, aseptic meningitis, encephalitis - Jaundice (can mimic hepatitis) - Multi-organ failure (< 10% cases) - Mortality: 5–40% in severe cases		> 90% mild & self-limiting Mortality higher in fulminant cases
Acute phase (Septicaemic phase) (2–9 days)		- Sudden high fever - Headache - Chills - Myalgia (muscle pain) - Rash - Nausea, vomiting - Fatigue - Conjunctival suffusion (reddening without pus or discharged) - Leptospira in blood, cerebrospinal fluid, urine		Biphasic course (two distinct phase): fever → short afebrile phase → fever returns
Immune phase (Leptospiruric phase)				- Divided into anicteric and icteric forms - Leptospira appears in urine
a) Anicteric form (mild)		- Absence of jaundice - Severe headache - Neck stiffness (meningitis) - Uveitis (may occur months later)		- ~90% of cases - Usually self-limiting and resolve with proper treatment.
b) Icteric form (severe)		- Jaundice - Liver dysfunction - Renal insufficiency - Haemorrhagic episodes - Multi-organ failure (kidney, liver, lung, brain) - Severe pulmonary haemorrhage syndrome (> 50% mortality)		− 5–10% of cases - Weil’s Disease - High risk of death (virulence linked to lipopolysaccharide hemolysins, outer membrane proteins of Leptospira)
Kidney involvement		- Non-oliguric nephropathy with potassium loss - Impaired sodium reabsorption - Oliguria if dehydrated → renal failure	Kidney is main target organ. Dialysis may be required
Liver involvement		- Jaundice (bilirubin increase) - Hepatocellular damage from intercellular junction disruption - Direct bilirubin ↑ (bile leakage) - Indirect bilirubin ↑ (haemolysis/breakdown of red blood cells)	Major organ target along with kidneys
Pregnancy-related		-Vertical transmission (breastfeeding, placenta invasion) - Miscarriage, fetal disorders	Human-to-human transmission is rare

As humans are a dead-end host for the spread of Leptospira spp., human-to-human transmission of leptospirosis is rare [84]. However, Leptospira spp. can be transmitted vertically from infected mothers to susceptible infants through breastfeeding. Infection of pregnant women can cause various foetal disorders and even miscarriage through invasion of the placenta from the mother to the foetus [4]. On the other hand, infected livestock such as cattle, dogs and pigs are prone to stillbirths, miscarriages, mastitis and a reduced milk yield [85, 86]. Reproductive disorders in livestock represent a serious economic concern, with abortion-associated losses valued at USD 97 − 2,611 per case, and herd outbreaks capable of generating financial risks that may reach USD 150,000 per year [87].

Pregnant or lactating cows show a decrease in milk production as Leptospira spp. require a pregnant uterus and a lactating mammary gland to reproduce [71]. Leptospirosis typically has a biphasic course in humans, in which the first phase corresponds to the multiplication and spread of Leptospira spp. in the body, while the second phase is characterised by the development of circulating antibodies and the shedding of Leptospira spp. in the urine [92]. The incubation period from exposure to the onset of symptoms is typically 7 to 12 days but can vary from 2 to 20 days [22].

a. Acute Phase (Septicaemic Phase)

The first phase of leptospirosis, known as the acute phase (septicaemic phase), typically lasts for about 3–7 days (Table 5). This phase is characterised by the appearance of Leptospira spp. in the bloodstream, in some cases, the cerebrospinal fluid (CSF), and is therefore also referred to as the leptospiraemic phase. Clinically, patients present with non-specific, flu-like symptoms, including sudden onset of high fever, headache, chills, myalgia, rash, nausea, vomiting, fatigue, and in many cases, conjunctival suffusion (reddening of the conjunctival vessels without purulent exudate) [6]. During this phase, Leptospira can be isolated from blood and CSF. Conjunctival suffusion is considered a particularly useful clinical clue because it is common in leptospirosis but uncommon in many other acute febrile illnesses [22]. The septicaemic phase is followed by a brief afebrile period (asymptomatic), after which fever returns, marking the onset of the second or “immune” phase of the disease. This classic biphasic pattern consists of the septicaemic phase of 3–7 days, followed by an immune phase characterised by the development of complications and recurrent fever [51].

b. Immune Phase (Leptospiruric Phase)

The second phase of leptospirosis is the immune phase (also known as the leptospiruric phase) in which Leptospira spp. can be isolated from the urine (Table 5). This phase can be divided into an anicteric and an icteric form. Clinically, most cases of leptospirosis (~ 90%) are anicteric and mild in nature, while the more severe icteric presentation, known as Weil’s disease, is seen in approximately 5–10% of patients [93]. Accounting for approximately ~ 90% of cases, the anicteric form of leptospirosis is the predominant presentation, distinguished by the absence of jaundice and the possible occurrence of leptospiral meningitis [94]. It is also characterised by severe headaches and neck stiffness. Uveitis (inflammation inside the eyes) may develop at this time or two weeks to one year after the onset of the disease, with an average of six months [95].

Icteric leptospirosis, also known as Weil’s syndrome, is the most severe manifestation of the Leptospira spp. infection. The symptoms can be triggered by the release of some virulent factors such as lipopolysaccharides, haemolysins, and outer membrane proteins [80]. It is usually characterised by jaundice, liver dysfunction, renal insufficiency, haemorrhagic episodes and multi-organ failure, including kidney, liver, lung, and brain, with a high risk of fatal outcome [6, 96]. Severe pulmonary haemorrhage syndrome (SPHS) leads to a mortality rate of over 50% and is th main cause of death [39, 97]. Leptospirosis leads to non-oliguric nephropathy with potassium loss, often characterised by impaired sodium reabsorption [98].

When inadequate oral intake leads to dehydration, patients are exposed to oliguria and renal failure, leading to death in areas where haemodialysis is not available. Pathogenic Leptospira primarily target the kidneys, especially the proximal convoluted tubules, where they can form biofilm-like aggregates that enable persistent colonisation and, in animals, lead to a chronic carrier state. Besides the kidney, another important target organ of leptospirosis is the liver. Leptospira spp. is known to preferentially attach in the perijunctional region between the hepatocytes. The disruption of the intercellular junctions between the hepatocytes and the damage to the liver cells lead to the leakage of bile. This leads to the higher levels of direct bilirubin seen in the icteric form. Sometimes the increase in indirect bilirubin levels can also occur in haemolysis caused by leptospirosis [22].

Research Gaps and Forward Directions

Despite substantial advances in our understanding of Leptospira spp., critical gaps remain in the effective management, diagnosis, and prevention of leptospirosis, particularly within veterinary and zoonotic contexts. Addressing these challenges requires a multidisciplinary, One Health approach. Key priorities include: (i) the expansion of Leptospira spp. diversity through discovery of new species and serovars highlights the need for more comprehensive taxonomic studies, particularly using state-of-the-art genomic approaches; (ii) In many countries, leptospirosis often goes unreported because the clinical symptoms are very similar to other communicable diseases [3]. Developing tools with sufficient sensitivity and specificity to detect and identify pathogenic Leptospira spp. strains in environmental samples are critical for risk assessment and targeting public health interventions. This includes multiplex PCR assays targeting clinically relevant species; (iii) Further studies are needed to fully discover the virulence mechanisms of Leptospira spp. and the specific role of different virulence factors, aligned with the evolving Leptospira genome, including genetic manipulation techniques such as CRISPRi which is powerful to study these mechanisms [99] (iv) Since leptospirosis is a neglected zoonotic disease, there is a need to improve diagnostic capabilities with consideration of rapid, sensitive and specific tests that can be readily deployed in marginal and resource-limited settings [3]. The use of machine learning algorithms, alongside traditional diagnostic methods, is promising to improve diagnostic accuracy and identify high-risk populations; (v) Developing effective, broadly protective vaccines against leptospirosis remains a major challenge. Continued research on the immunology of Leptospira spp. and the development of new vaccine candidates are important. Research into the use of recombinant proteins and the development of DIVA strategies is essential to improve vaccine efficacy that does not cross-react with many diagnostic tests in vaccinated individuals; (vi) The combination of the lack of standardised diagnostic and harmonised reporting procedures further contributes to the difficulty in obtaining a reliable global estimate of leptospirosis mortality. The disparity in reported mortality rates underscores the urgent need for strengthened surveillance systems, including systematic monitoring of animal reservoirs and serogroup distribution at the regional level, alongside standardised case definitions, to generate more accurate and reliable estimates of the true global mortality burden of leptospirosis.

Conclusions

Leptospirosis is an example of the complexity and urgency of controlling emerging zoonotic diseases in a rapidly changing global health landscape. As this review highlights, advances in our understanding of Leptospira spp., from epidemiology and pathogenesis to diagnostics and genomics, have significantly deepened scientific insight, yet major challenges remain. Gaps in surveillance, diagnostic sensitivity, and vaccine development continue to hamper effective control, particularly in resource-poor settings where the disease burden is highest. A coordinated One Health approach is essential to reduce these gaps. Priorities include expanding genomic surveillance to capture the full diversity of Leptospira spp., improving the resolution and accessibility of diagnostic tools, deciphering virulence mechanisms with modern molecular tools, and developing next-generation vaccines that are both effective and compatible with surveillance systems. Equally important is the standardisation of reporting and case definitions to enable accurate estimates of the global burden. With climate change, urbanisation, and increasing human-animal interaction, the threat of leptospirosis and other zoonotic diseases will continue to emerge. Investing in multidisciplinary research, technology, and global collaboration is critical to anticipate, detect, and respond effectively to these evolving threats [100–117].

Supplementary Information

Below is the link to the electronic supplementary material.

Authors’ Contributions

Lesley Maurice Bilung and Kasing Apun conceptualised the study. Ahmad Syatir Tahar, Chai Fung Pui, Muhammad Khairil Syamri Bakeri, and Rosdi Kira performed the literature search and interpreted relevant articles. Ahmad Syatir Tahar, Chai Fung Pui, and Muhammad Khairil Syamri Bakeri prepared the first manuscript draft. Lesley Maurice Bilung, Lela Su’ut, Romano Ngui, and Kasing Apun provided critical feedback. All authors read and approved the final manuscript.

Funding

Open access funding provided by The Ministry of Higher Education Malaysia and Universiti Malaysia Sarawak. This work did not receive any funding.

Declarations

Conflict of interest

All authors report no conflict of interest in this work.

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