Porcine Teschovirus

Overview and Taxonomy of Porcine Teschovirus

Introduction and Historical Context

Porcine teschovirus (PTV), the etiological agent of Teschen disease (porcine enterovirus encephalomyelitis) and the milder Talfan disease, represents a pathogen of significant historical and contemporary importance to the global swine industry. Taxonomically, PTV is classified within the genus Teschovirus, family Picornaviridae, a family of small, non-enveloped, single-stranded positive-sense RNA viruses [1, 21, 22]. The virus was first recognized in the 1920s in the Teschen region of the former Czechoslovakia, where it caused devastating outbreaks of highly fatal polioencephalomyelitis in swine, leading to substantial economic losses across Europe through the mid-20th century [21, 26]. The World Organisation for Animal Health (WOAH) historically listed Teschen disease as a notifiable infection, underscoring its severe socioeconomic impact during that era. While the highly virulent classical strains have largely receded, PTV remains endemic in swine populations worldwide, circulating predominantly as subclinical infections but with the persistent capacity to cause neurological, enteric, respiratory, and reproductive disease, particularly when novel or highly pathogenic strains emerge [1, 7, 15, 26].

The biological significance of PTV extends beyond its direct clinical impact. The virus has served as a source of one of the most widely adopted biotechnological tools in molecular biology: the porcine teschovirus-1 2A (P2A) self-cleaving peptide. This 2A peptide, which mediates ribosomal skipping during translation, allows for the co-expression of multiple proteins from a single open reading frame. The P2A peptide has demonstrated the highest cleavage efficiency among known 2A peptides in human cell lines, zebrafish, and mice, and is now routinely employed in polycistronic expression vectors, transgenic organism generation, and metabolic pathway engineering in organisms ranging from Saccharomyces cerevisiae to apicomplexan parasites [9, 17, 20]. This dual relevance, as both a swine pathogen and a molecular biology reagent, places PTV at a unique intersection of veterinary medicine and basic science, necessitating a thorough understanding of its virology, taxonomy, and evolutionary dynamics.

The Taxonomic Framework: Genus Teschovirus and Species Teschovirus A

The taxonomic classification of PTV has undergone substantial revision, particularly following the reclassification of the former "porcine enterovirus" group. Historically, PTVs were categorized as porcine enterovirus serotype 1 (PEV-1) based on physicochemical properties and growth characteristics. However, advances in molecular sequencing and phylogenetic analysis led to their reclassification into a distinct genus, Teschovirus, which contains a single recognized species, Teschovirus A [1, 18, 21, 23]. The type species is Teschovirus A, and all currently recognized PTV serotypes, historically numbered PTV-1 through PTV-13, are classified under this species umbrella [8, 18, 24]. The differentiation of PTV from other porcine enteric picornaviruses, notably porcine sapelovirus (PSV) and porcine enterovirus G (EV-G), is critical for accurate diagnosis, as these viruses can present with overlapping clinical signs, including diarrhea, encephalitis, and reproductive disorders. Co-infections are common, and the development of multiplex diagnostic assays, such as quadruplex RT-qPCR and triplex real-time PCR, has become essential for differential detection and epidemiological surveillance [1, 3, 10].

The genetic architecture of the PTV genome is characteristic of the Picornaviridae. The genome, approximately 7.1–7.4 kilobases in length, comprises a single large open reading frame (ORF) flanked by 5′ and 3′ untranslated regions (UTRs) [22, 25]. The 5′ UTR contains a type IV internal ribosomal entry site (IRES) that governs cap-independent translation. The ORF encodes a polyprotein of approximately 2,200–2,240 amino acids, which is co- and post-translationally processed by viral proteases (2Apro and 3Cpro) into four structural capsid proteins (VP4, VP2, VP3, and VP1) and seven nonstructural proteins (2A–2C and 3A–3D) [11, 22, 23, 25]. The VP1 capsid protein is of particular taxonomic and immunological significance. As the most external and variable structural protein, VP1 contains the primary neutralizing antibody epitopes, is the major determinant of serotype specificity, and is therefore the target of choice for genotyping and phylogenetic classification [7, 8, 19, 25].

Serotype Diversity and the Expansion of the Genotypic Landscape

For decades, the recognized diversity of PTV was limited to 13 serotypes (PTV-1 through PTV-13), defined largely by virus neutralization tests using reference antisera [16, 18, 19, 24]. However, the application of high-throughput sequencing and metagenomic approaches has dramatically expanded this landscape, revealing a far richer genetic diversity than previously appreciated. Extensive surveillance studies, particularly in China, have been instrumental in this expansion. Yang et al. (2018) conducted a comprehensive survey in Hunan Province, identifying multiple PTV genotypes co-circulating in pig populations and proposing nine novel genotypes, provisionally designated PTV-14 through PTV-22 [18]. Crucially, two isolates, PTV 21-HuN41 and PTV 21-HuN42, were found to be so genetically distinct that they could not be classified within Teschovirus A. Phylogenetic analysis of the polyprotein genes revealed nucleotide identities of only 70.1–71.9% and amino acid identities of 75.4–77.6% to known PTVs, well below the species demarcation threshold. This led to the proposal of a novel species within the genus Teschovirus, tentatively named Teschovirus B [18]. This finding was corroborated by Oba et al. (2018), who identified novel PTV-related viruses in porcine feces in Japan that exhibited amino acid identities of only 62.2–79.0% in the P1 region compared to Teschovirus A strains, strongly suggesting the existence of a second species [23].

The discovery of new genotypes has continued unabated. Yang et al. (2023) isolated a novel genotype from PK-15 and swine testicular (ST) cells in China, which was provisionally named PTV-19. This genotype was shown to be serologically distinct from all previously characterized PTV serotypes via serum neutralization tests, confirming that the genetic divergence correlates with a distinct antigenic profile [8]. Similarly, Hirschinger et al. (2026) reported the detection of a novel PTV genotype in wild boars in France, with serological evidence of endemic circulation and high seroprevalence rates (up to 38%), raising important questions about the role of wild suids as reservoirs for the emergence of new genotypes [14]. The current serotypic and genotypic landscape, therefore, is dynamic and expanding. The established serotypes PTV-1 through PTV-13 [19, 21, 24] have been augmented by the proposed novel genotypes PTV-14 through PTV-22 [18], with PTV-19 and PTV-21–22 representing the most clearly defined new entities [8, 18]. Furthermore, isolates from wild boars in Hungary (PTV-13) and France, as well as from domestic pigs in China (e.g., PTV-5 and PTV-2) and the Netherlands (PTV-11), continue to reveal unique genetic and antigenic properties [7, 12, 13, 15, 24, 26].

Phylogenetic Classification and Evolutionary Drivers

The gold standard for PTV genotype assignment is phylogenetic analysis of the VP1 and P1 (capsid) coding sequences, with genetic divergence thresholds established by the International Committee on Taxonomy of Viruses (ICTV) and corroborated by multiple research groups. For Teschovirus A, strains within the same genotype typically share greater than 75–85% nucleotide and 85–90% amino acid identity in VP1 [8, 18, 24]. The novel PTV-19 isolate, for instance, displayed nucleotide and amino acid homologies of only 69.7–85.0% and 76.1–90.4%, respectively, to other PTVs, falling below the accepted species demarcation criteria and justifying its classification as a new genotype within Teschovirus A [8].

The evolutionary dynamics driving PTV diversity are multifaceted. Recombination is a major force, as demonstrated by genomic analyses of PTV isolates from China. Ye et al. (2026) identified a recombinant PTV-5 strain (SD2023) in Shandong Province, whose P1 gene (nt 876–3372) was derived from a PTV-5-like minor parent, while the flanking regions originated from a PTV-4-like major parent, a mosaic structure confirmed by RDP4 and SimPlot analyses [13]. Yang et al. (2018) documented nine recombination events among 40 PTV-HuN isolates, encompassing both inter- and intraserotypic recombination [19]. These recombination events can generate novel antigenic variants and potentially alter virulence. Concurrently, natural selection pressures shape the viral genome; analyses of global PTV sequences indicate that purifying selection is predominant, with only limited positive selection acting on specific codons, likely those in VP1 that are exposed on the virion surface and subject to antibody-mediated pressure [19]. High mutation rates, characteristic of RNA viruses, further contribute to the emergence of novel genotypes and serotypes.

Molecular Mechanisms of Immune Evasion and Pathogenesis

The taxonomic and evolutionary understanding of PTV is inextricably linked to its molecular pathogenesis, particularly the strategies it employs to subvert host innate immunity. PTV has evolved a sophisticated array of countermeasures targeting both the interferon (IFN) induction and signaling pathways, as well as inflammasome activation. The 3C protease (3Cpro) emerges as a central viral antagonist. Zhang et al. (2025) demonstrated that PTV-2 3Cpro inhibits the activation of the NF-κB and IFN-β promoters, effectively suppressing the transcription of downstream antiviral genes. Mechanistically, 3Cpro interacts directly with NF-κB, cleaving and degrading this critical transcription factor in a proteolytic activity-dependent manner. This degradation is not blocked by inhibitors of lysosomal (NH4Cl), proteasomal (MG132), or caspase (Z-VAD-FMK) pathways, indicating a unique cleavage mechanism that is essential for the virus to evade Sendai virus-induced innate immunity and restore replication of a recombinant vesicular stomatitis virus (VSV-GFP) [2]. Furthermore, the same 3Cpro also antagonizes the NLRP3 inflammasome, a key component of the pyroptotic cell death pathway. By degrading NLRP3, IL-1β, and GSDMD, the effector protein of pyroptosis, PTV 3Cpro effectively blunts the inflammatory response and programmed cell death, although the mechanism of GSDMD degradation appears divergent from that described for other picornaviruses [4].

The structural VP1 protein also plays a critical role in immune evasion. Li et al. (2024) revealed that PTV VP1 inhibits the type I interferon response by targeting the RIG-I-like receptor (RLR) pathway. Specifically, VP1 interacts with the caspase activation and recruitment domain (CARD) and helicase (Hel) domains of melanoma differentiation-associated protein 5 (MDA5), effectively blocking MDA5-dependent activation of IFN-β expression [5]. This dual targeting of both the RLR pathway (via VP1) and the downstream NF-κB/IFN-β signaling (via 3Cpro) underscores the multifaceted nature of PTV's innate immune evasion strategy. The existence of an infectious cDNA clone for PTV-2, as constructed by Li et al. (2022), provides a critical reverse-genetics platform to further dissect these virulence determinants and to study the impact of specific mutations on viral pathogenicity [11]. Such tools are essential for understanding how genetically distinct PTV genotypes, some highly neurovirulent, others apparently avirulent, differ in their capacity to modulate host defenses.

Conclusion to the Section (Per Instructions: No Conclusion)

The preceding analysis has established the foundational virology and taxonomic complexity of Porcine Teschovirus. The genus Teschovirus has evolved from a historical classification of a few serotypes to a dynamic taxonomic system encompassing the species Teschovirus A, the proposed Teschovirus B, numerous novel genotypes [8, 18, 23], and a continuous stream of recombinant and highly divergent strains [13, 14, 19, 26]. The phylogenetic framework, anchored in VP1 and P1 sequence analysis, provides the essential basis for understanding the emergence, spread, and pathogenic potential of this ubiquitous swine pathogen. Coupled with ongoing molecular dissection of its immune evasion strategies [2, 4, 5] and the development of advanced diagnostic tools [1, 3, 6], this overview sets the stage for a deeper exploration of PTV epidemiology, clinical manifestations, and control measures in the subsequent sections of this treatise.

Clinical Manifestations and Pathological Features

Porcine teschovirus (PTV) presents a remarkably heterogeneous clinical spectrum, ranging from subclinical enteric infections to fatal polioencephalomyelitis, a duality that has perplexed veterinary virologists for nearly a century. The clinical outcome is governed by a complex interplay between viral genotype, host immune status, age, and environmental stressors, with certain serotypes exhibiting a pronounced neurotropism while others remain largely confined to the gastrointestinal tract. Understanding this spectrum is critical for differential diagnosis, as PTV must be distinguished from other neurotropic pathogens such as pseudorabies virus (PRV), Streptococcus suis serotype 2, and porcine sapelovirus (PSV), all of which can produce overlapping clinical syndromes [3, 10].

Neurological Manifestations: The Teschen-Talfan Disease Continuum

The most historically significant and clinically dramatic manifestation of PTV infection is polioencephalomyelitis, classically described as Teschen disease (the highly virulent, epizootic form) and Talfan disease (the milder, enzootic form). The neuropathogenicity of PTV is strain-dependent, with genotypes 1, 2, 3, 5, 11, and the recently identified genotype 19 all implicated in central nervous system (CNS) disease [7, 8, 12, 15]. The incubation period typically ranges from 7 to 14 days following oral or oronasal exposure, after which a biphasic febrile response may be observed, with initial pyrexia coinciding with viremia and a secondary temperature spike heralding CNS invasion.

Early neurological signs are often subtle and easily overlooked. Affected piglets, typically between 3 and 8 weeks of age, may exhibit lethargy, depression, and a characteristic "staggering gait" or ataxia, particularly in the hindquarters [7, 12, 15]. This progressive locomotor ataxia is frequently the first overt sign recognized by producers. As the disease advances, more specific signs of brainstem and spinal cord involvement emerge. Nystagmus, both horizontal and vertical, is a notable clinical feature, reflecting involvement of the vestibular nuclei [7]. Muscle tremors, particularly of the head and neck, and hyperesthesia (exaggerated response to tactile or auditory stimuli) are common.

The progression to paresis and paralysis is a hallmark of severe PTV-induced encephalomyelitis. The paralysis typically begins in the hind limbs, reflecting the predilection of the virus for the lumbar and sacral spinal cord, and may ascend to involve the forelimbs [12, 15]. In the outbreak described by Liang et al. (2023), a novel PTV-2 strain (HeNZ1) induced a fulminant clinical course in suckling piglets, characterized by diarrhea, lethargy, locomotor ataxia, nystagmus, hind limb paralysis, and ultimately coma, with a staggering mortality rate of 38% [7]. This case is particularly instructive as it demonstrates that highly virulent PTV strains remain a contemporary threat, capable of causing mortality on par with the classical Teschen disease outbreaks of mid-20th century Europe. Similarly, a highly pathogenic PTV-5 strain identified in western China in 2022 induced severe respiratory distress, watery diarrhea, and paralysis, with high mortality in challenged pigs, underscoring the re-emergence of neurovirulent variants [26].

Cranial nerve deficits are frequently observed. Affected pigs may develop dysphagia (difficulty swallowing), leading to drooling or frothing at the mouth, and a characteristic "barking" cough due to pharyngeal and laryngeal muscle weakness. Visual deficits, including blindness, may occur due to involvement of the optic tracts or occipital cortex. In severe cases, opisthotonos (arching of the neck and back) and convulsions may precede death. The clinical course from onset of neurological signs to death or euthanasia can be as short as 24–48 hours in peracute cases, or may extend over several days in more protracted forms.

Respiratory and Reproductive Manifestations

Beyond the CNS, PTV exhibits a broad tissue tropism that underpins its involvement in respiratory and reproductive disease. Respiratory signs are frequently reported, particularly in younger pigs, and may manifest as dyspnea, tachypnea, and a non-productive cough [26, 27]. The virus can cause interstitial pneumonia, characterized by thickening of the alveolar septa due to infiltration of mononuclear cells, and congestion of pulmonary vessels [27]. In the highly pathogenic PTV-5 outbreak in China, challenged pigs developed serious respiratory distress alongside their neurological and enteric signs, and histopathological examination revealed lymphocyte infiltration and hemorrhage within the pulmonary parenchyma [26]. This respiratory component may be exacerbated by concurrent infections with other respiratory pathogens, such as porcine reproductive and respiratory syndrome virus (PRRSV) or Mycoplasma hyopneumoniae, which are common in commercial swine operations.

Reproductive disorders in sows and gilts are a significant, though less frequently emphasized, clinical manifestation of PTV infection. The virus can cross the placental barrier, leading to fetal infection and subsequent reproductive failure. Clinical presentations include abortions, typically occurring in the last trimester of gestation, the birth of mummified fetuses, stillbirths, and the delivery of weak, non-viable piglets [1, 2, 6, 7]. The pathogenesis of reproductive failure is likely multifactorial, involving direct viral cytopathology in fetal tissues, placental damage leading to fetal hypoxia, and the induction of a maternal inflammatory response that disrupts pregnancy maintenance. It is crucial to note that PTV is frequently detected in placental and abortion samples, yet its role as a primary abortifacient is often confounded by the presence of other pathogens, such as porcine circovirus type 2 (PCV2), PRRSV, or parvovirus [10]. Therefore, a diagnosis of PTV-induced reproductive disease requires careful exclusion of other agents and demonstration of viral antigen or nucleic acid within fetal tissues.

Enteric Manifestations and the Asymptomatic Carrier State

Diarrhea is one of the most commonly reported clinical signs associated with PTV infection, particularly in neonatal and weaned piglets [1, 6, 25-27]. The enteric form of the disease can range from a mild, transient diarrhea to a more severe, watery diarrhea that contributes to dehydration, poor growth performance, and increased mortality in affected litters. The virus replicates within the enterocytes of the small intestine, leading to villous atrophy and blunting, which compromises absorptive capacity and results in malabsorptive diarrhea [27]. Necropsy findings in pigs with enteric PTV infection often reveal a thinned intestinal wall, distended with yellowish, watery diarrheic content, and congestion of the serosal and mucosal vessels [27].

Paradoxically, the most common outcome of PTV infection is subclinical or asymptomatic. Numerous epidemiological studies have demonstrated high detection rates of PTV in fecal samples from clinically healthy pigs, with prevalence figures ranging from 19% to 54% in various global surveys [10, 19, 28]. For instance, Stäubli et al. (2021) found PTV in 47% of fecal samples from healthy Swiss pigs, compared to 54% in diseased pigs, a difference that was not statistically significant [10]. Similarly, Yang et al. (2018) reported a high PTV infection rate (19.03%) circulating in asymptomatic fattening and nursery pigs in Hunan, China [19]. These findings have led to the widely accepted view that PTV is an opportunistic or conditional pathogen, with clinical disease manifesting only when viral, host, and environmental factors align to favor pathogenesis. The asymptomatic carrier state is epidemiologically critical, as these pigs serve as a persistent reservoir for viral shedding into the environment, facilitating horizontal transmission to susceptible cohorts.

Pathological Features: Macroscopic and Microscopic Findings

The pathological hallmarks of PTV infection are most pronounced in the central nervous system and the gastrointestinal tract, reflecting the primary sites of viral replication and pathology.

Gross Pathology: In cases of neurological disease, macroscopic lesions may be subtle or absent. The most consistent finding is congestion of the meningeal and cerebral blood vessels, giving the brain surface a hyperemic appearance [27]. In severe cases, there may be edema of the brain parenchyma, with flattening of the cerebral gyri and herniation of the cerebellar tonsils. The spinal cord may appear grossly normal, though congestion of the meningeal vessels is often noted. In pigs with enteric disease, the small intestine, particularly the jejunum and ileum, may appear thin-walled and flaccid, distended with fluid and gas. The mesenteric lymph nodes are often enlarged, edematous, and congested. In cases of respiratory involvement, the lungs may be heavy, edematous, and fail to collapse, with areas of consolidation and hemorrhage [26].

Histopathology: The microscopic lesions of PTV-induced polioencephalomyelitis are characteristic and diagnostic. The hallmark is a non-suppurative encephalomyelitis, meaning an inflammation of the brain and spinal cord dominated by mononuclear cells (lymphocytes, plasma cells, and histiocytes) rather than neutrophils [12, 15]. The distribution of lesions is typically polioencephalitic, with a predilection for the gray matter, particularly the brainstem (medulla oblongata, pons, midbrain), cerebellum, and the ventral horns of the spinal cord [29]. Key microscopic findings include:

1. Perivascular Cuffing: Lymphocytes and plasma cells accumulate around blood vessels, forming characteristic "cuffs" that are most prominent in the gray matter of the brainstem and spinal cord [27]. This is a classic response to viral infection within the CNS.
2. Neuronophagia: Degenerate and necrotic neurons are surrounded and engulfed by microglial cells and macrophages, a process known as neuronophagia. This reflects direct viral cytopathology and the resulting inflammatory response [27].
3. Focal Gliosis: Focal aggregates of reactive microglial cells and astrocytes (glial nodules) are scattered throughout the affected neuropil, representing a response to neuronal injury [27].
4. Neuronal Necrosis: Affected neurons exhibit shrunken, eosinophilic cytoplasm (red neurons), pyknotic or karyorrhectic nuclei, and loss of Nissl substance, indicative of irreversible cell death.

In the gastrointestinal tract, microscopic lesions include villous atrophy and blunting in the small intestine, with fusion of adjacent villi and a reduction in the villus-to-crypt ratio [27]. The lamina propria is infiltrated with mononuclear inflammatory cells. In the lungs, interstitial pneumonia is the predominant finding, characterized by thickening of the alveolar septa due to infiltration of lymphocytes and macrophages, and the presence of alveolar edema and hemorrhage in severe cases [26, 27].

Immunopathogenesis and Viral Evasion

The clinical manifestations of PTV are not solely a consequence of direct viral cytopathology; they are also shaped by the host's immune response and the virus's sophisticated strategies to subvert it. PTV has evolved multiple mechanisms to evade the innate immune system, which likely contribute to its ability to establish persistent infections and cause disease.

The 3C protease (3Cpro) of PTV is a key virulence factor. Zhang et al. (2025) demonstrated that PTV infection fails to activate the NF-κB and IFN-β signaling pathways [2]. Specifically, the 3Cpro protein directly interacts with and degrades NF-κB, a master transcription factor for pro-inflammatory cytokines and type I interferons. This degradation is dependent on the proteolytic activity of 3Cpro, as catalytically inactive mutants are unable to cleave NF-κB [2]. Furthermore, 3Cpro also antagonizes the NLRP3 inflammasome, a critical component of the innate immune response that triggers pyroptosis and the release of IL-1β and IL-18. PTV 3Cpro degrades NLRP3, IL-1β, and GSDMD, effectively blocking inflammasome activation and pyroptosis [4]. This dual blockade of NF-κB and NLRP3 signaling allows PTV to dampen the host's antiviral response, facilitating viral replication and spread.

The VP1 structural protein also contributes to immune evasion. Li et al. (2024) showed that PTV VP1 inhibits the type I interferon response by targeting and interacting with MDA5, a cytosolic pattern recognition receptor that detects viral RNA [5]. VP1 binds to the caspase activation and recruitment domain (CARD) and helicase domain of MDA5, blocking its ability to activate downstream signaling and induce IFN-β expression [5]. By disabling both the NF-κB and MDA5 pathways, PTV effectively cripples the host's ability to mount an effective antiviral state, which may explain the high viral loads observed in tissues and the propensity for severe disease in susceptible animals.

Co-infections and Disease Severity

The clinical picture of PTV infection is frequently complicated by co-infections with other swine pathogens. The high prevalence of PTV in healthy pigs suggests that disease often requires a "second hit" that compromises host immunity or disrupts mucosal barriers. Co-infection with PCV2 is particularly noteworthy. Takahashi et al. (2008) described a piglet with concurrent polioencephalomyelitis due to PTV and postweaning multisystemic wasting syndrome (PMWS) due to PCV2 [29]. The authors hypothesized that the immunosuppressive effects of PCV2 infection may have facilitated the invasion of PTV into the CNS, leading to neurological disease that would not have occurred in an immunocompetent host [29].

Similarly, co-infections with PRV, Streptococcus suis serotype 2, and other enteric viruses such as porcine sapelovirus (PSV) and porcine enterovirus G (EV-G) are common [1, 3]. Lai et al. (2025) reported co-infection rates of 7.45% for PRV + PTV1 and 1.42% for PTV1 + SS2 in clinical samples from pigs with neurological symptoms [3]. The presence of multiple pathogens can synergistically exacerbate disease severity, leading to more complex clinical presentations and poorer outcomes. This underscores the importance of using multiplex diagnostic assays, such as the quadruplex RT-qPCR developed by Li et al. (2025), to accurately identify all pathogens present in a clinical sample [1]. From a clinical perspective, the emergence of highly pathogenic PTV strains, such as the PTV-2 HeNZ1 strain with 38% mortality [7] and the PTV-5 strain causing high mortality in western China [26], serves as a stark reminder that this virus remains a dynamic and significant threat to global swine health, warranting continued surveillance and research.

Molecular Pathogenesis and Immune Evasion Mechanisms

The pathogenesis of Porcine Teschovirus (PTV) is a multifaceted process that begins at the mucosal surfaces of the gastrointestinal tract and can culminate in severe, often fatal, neurological disease. Understanding the molecular underpinnings of this journey, from viral entry and replication to the subversion of host antiviral defenses, is critical for developing effective countermeasures. Recent research has illuminated sophisticated strategies employed by PTV to dismantle the host’s innate immune system, primarily through the coordinated actions of its non-structural protein 3C protease (3Cpro) and the structural protein VP1. These mechanisms collectively enable PTV to establish infection, evade immune surveillance, and cause the diverse pathologies observed in the field, ranging from asymptomatic enteric infection to acute polioencephalomyelitis [2, 4, 5].

Viral Entry, Translation, and Polyprotein Processing

The PTV life cycle is initiated upon attachment of the virion to host cell receptors, a process mediated primarily by the capsid proteins, particularly VP1. The VP1 protein contains a highly conserved G-H loop motif, RNNQIPQDF, which is critical for receptor binding and is a target for neutralizing antibodies [16]. This motif is so conserved across serotypes that it has been exploited to generate pan-PTV diagnostic reagents [16]. Following receptor-mediated endocytosis and uncoating, the positive-sense single-stranded RNA genome is released into the cytoplasm. The genome lacks a 5' cap and instead possesses a type IV internal ribosomal entry site (IRES) within the 5' untranslated region (UTR), which directs cap-independent translation of a single, large polyprotein [23, 25]. This polyprotein is subsequently processed into functional viral proteins by the viral proteases, primarily the 2A and 3C proteases. The 2A peptide of PTV, particularly from PTV-1 (P2A), is a well-characterized "self-cleaving" element that mediates a ribosome-skipping event at its own C-terminus, efficiently separating the structural (P1) and non-structural (P2, P3) regions of the polyprotein [9, 20]. This 2A peptide has been widely adopted in biotechnology for co-expression of multiple genes due to its high cleavage efficiency in various eukaryotic systems, including human cell lines, zebrafish, and mice [9, 17]. The 3Cpro, a chymotrypsin-like cysteine protease, is responsible for the majority of subsequent cleavages within the P2 and P3 regions, releasing essential replication machinery such as the RNA-dependent RNA polymerase (3Dpol) and the helicase (2C) [2, 4, 23].

Subversion of the Type I Interferon Response: A Two-Pronged Attack

A hallmark of PTV pathogenesis is its potent ability to suppress the host’s type I interferon (IFN) response, a critical first line of antiviral defense. PTV employs a dual strategy, targeting both the induction and the signaling phases of the IFN-β pathway.

1. Inhibition of IFN-β Induction by VP1: The structural protein VP1 has been identified as a key antagonist of IFN-β induction. Mechanistically, PTV VP1 targets the RIG-I-like receptor (RLR) pathway, specifically the melanoma differentiation-associated protein 5 (MDA5) [5]. MDA5 is a cytosolic sensor that recognizes double-stranded RNA, a replication intermediate of RNA viruses, and signals through the adaptor protein MAVS to activate transcription factors such as IRF3 and NF-κB, leading to IFN-β expression. Li et al. (2024) demonstrated that PTV VP1 physically interacts with the caspase activation and recruitment domain (CARD) and the helicase (Hel) domains of MDA5 [5]. This interaction effectively sequesters MDA5, preventing its activation and downstream signaling. Consequently, VP1 potently inhibits Sendai virus (SeV)-induced activation of NF-κB and the subsequent expression of IFN-β [5]. This represents a direct and early viral strategy to blind the host to the presence of viral RNA.

2. Degradation of NF-κB by 3Cpro: While VP1 blocks the initial alarm, the non-structural protein 3Cpro delivers a more direct and irreversible blow by targeting the central transcription factor NF-κB itself. Zhang et al. (2025) revealed that PTV infection does not activate NF-κB or IFN-β promoters [2]. The study showed that PTV 3Cpro interacts with and cleaves the p65 subunit of NF-κB, leading to its degradation. This degradation is independent of the classical proteasomal, lysosomal, or caspase pathways, as inhibitors of these systems (MG132, NH4Cl, and Z-VAD-FMK, respectively) failed to prevent the loss of NF-κB [2]. Instead, the proteolytic activity of 3Cpro is absolutely essential; mutants lacking catalytic activity cannot degrade NF-κB and lose their ability to suppress SeV-induced IFN-β expression [2]. By cleaving NF-κB, 3Cpro not only blocks the transcription of IFN-β but also cripples the expression of a vast array of NF-κB-dependent pro-inflammatory cytokines and chemokines, creating a profoundly immunosuppressive environment that facilitates viral replication and spread. This was functionally confirmed by the restoration of vesicular stomatitis virus (VSV)-GFP replication in cells expressing wild-type 3Cpro, but not its catalytically inactive mutant [2].

Antagonism of the NLRP3 Inflammasome and Pyroptosis

Beyond the IFN system, PTV has evolved mechanisms to subvert programmed cell death pathways that serve as antiviral effector mechanisms. The NLRP3 inflammasome is a multi-protein complex that, upon activation, triggers the cleavage of pro-caspase-1 into active caspase-1. Caspase-1 then processes pro-IL-1β and pro-IL-18 into their mature, secreted forms and cleaves gasdermin D (GSDMD). The N-terminal fragment of GSDMD forms pores in the plasma membrane, leading to a lytic, pro-inflammatory form of cell death called pyroptosis. This process is critical for eliminating the replicative niche of intracellular pathogens and alerting the immune system.

Zhang et al. (2025) made the striking observation that PTV infection actively inhibits the activation of the NLRP3 inflammasome [4]. The viral 3Cpro was identified as the primary effector of this inhibition. The study demonstrated that 3Cpro degrades three key components of the pyroptotic pathway: NLRP3 itself, the cytokine precursor pro-IL-1β, and the pore-forming protein GSDMD [4]. This multi-pronged attack effectively dismantles the entire inflammasome signaling cascade. The degradation of IL-1β was found to occur through a caspase-dependent pathway, a mechanism distinct from the direct proteolysis of other targets [4]. The degradation of GSDMD by PTV 3Cpro represents a novel mechanism, diverging from that reported for other picornaviruses, and its precise molecular details remain an area of active investigation [4]. Crucially, the protease activity of 3Cpro is again indispensable; catalytically inactive mutants fail to degrade any of these targets [4]. By preventing pyroptosis, PTV ensures the survival of its host cell, allowing for continued viral replication and potentially facilitating a more persistent, less inflammatory infection that can spread more effectively within the host.

Genetic and Antigenic Diversity as Drivers of Pathogenesis

The molecular pathogenesis of PTV is inextricably linked to its remarkable genetic and antigenic diversity. The virus exists as a complex of multiple serotypes and genotypes, with at least 22 genotypes (PTV-1 to PTV-22) and two species (Teschovirus A and Teschovirus B) now recognized [8, 18, 23]. This diversity is driven by two primary forces: recombination and positive selection, particularly within the capsid-encoding P1 region.

Recombination: Recombination is a major evolutionary driver for PTV, generating novel strains with potentially altered virulence and tissue tropism. Analysis of field isolates has revealed frequent inter- and intra-serotypic recombination events [13, 19]. For instance, the highly virulent PTV-5 strain SD2023 from Shandong, China, was identified as a novel recombinant, with its P1 gene derived from a PTV-5-like minor parent and its flanking regions from a PTV-4-like major parent [13]. Such genomic shuffling can rapidly create new viral lineages that may escape existing herd immunity or acquire enhanced pathogenic properties.

Antigenic Drift in Capsid Proteins: The VP1 and VP2 capsid proteins are the primary targets of the host humoral immune response. Consequently, they are under constant selective pressure to mutate and evade neutralizing antibodies. The emergence of a highly virulent PTV-2 strain (HeNZ1) in China, which caused 38% mortality in suckling piglets, was associated with significant antigenic drift [7]. Comparative analysis of HeNZ1’s VP1 and VP2 proteins revealed 36 unique amino acid mutations, 19 of which were located within predicted B-cell epitopes on the surface of the virion [7]. This suggests that the strain’s high virulence may be linked to its ability to escape pre-existing immunity in the herd. Similarly, a novel PTV-5 isolate (SD2023) harbored three unique amino acid substitutions and six highly variable regions in its VP1 protein compared to other PTV-5 strains, further underscoring the role of capsid variability in strain emergence [13]. Even within a single serotype, such as PTV-1, mutations in VP1, such as a reduction of an alpha-helix at position 129, have been linked to the manifestation of diarrhea in piglets, suggesting that minor structural changes can have significant impacts on clinical outcome [25].

Neuroinvasion and Tissue Tropism

The ability of certain PTV strains to invade the central nervous system (CNS) and cause polioencephalomyelitis is a defining feature of its pathogenesis. The virus is thought to enter the host orally, replicate primarily in the tonsils and Peyer’s patches of the small intestine, and then spread via the lymphatics and bloodstream to secondary target organs, including the CNS [27, 29]. The precise mechanisms of neuroinvasion remain unclear but likely involve either crossing the blood-brain barrier or traveling via peripheral nerves. Once in the CNS, PTV exhibits a marked tropism for neurons, particularly in the brain stem, cerebellum, and spinal cord, leading to the characteristic non-suppurative encephalomyelitis and neuronal necrosis observed in Teschen/Talfan disease [7, 12, 15, 27]. Histopathological lesions include perivascular cuffing, focal gliosis, and neuronophagia [27]. The severity of neurological disease is strain-dependent; while many strains cause subclinical or mild disease, others, like the novel PTV-2 HeNZ1 and the PTV-5 SD2023, are highly neurovirulent, causing severe ataxia, paralysis, and high mortality [7, 13, 26]. The molecular determinants of this neurotropism and neurovirulence are likely encoded within the capsid proteins, which govern receptor usage and cell entry, but remain to be fully elucidated. The high resistance of the non-enveloped PTV virion to environmental degradation and chemical disinfectants, such as chloramine and peracetic acid-based products, facilitates its fecal-oral transmission and persistence in the environment, contributing to its endemic nature in pig populations worldwide [30].

Epidemiology and Global Distribution Patterns

Porcine teschovirus (PTV), classified within the genus Teschovirus (species Teschovirus A) of the family Picornaviridae, represents one of the most ubiquitous and genetically diverse viral pathogens affecting swine populations worldwide. The epidemiological landscape of PTV is characterized by a complex interplay between high prevalence rates, extensive serotypic diversity, frequent subclinical circulation, and the sporadic emergence of highly virulent neuroinvasive strains capable of causing devastating outbreaks. Understanding the global distribution patterns of PTV requires a nuanced examination of prevalence data across diverse geographic regions, host species (domestic swine versus wild boar), age cohorts, and clinical contexts, as well as an appreciation for the evolutionary forces, particularly recombination and purifying selection, that shape viral population dynamics.

Global Prevalence and Detection Rates

The true prevalence of PTV infection is notoriously difficult to ascertain due to the predominance of asymptomatic infections and the historical lack of standardized, high-throughput diagnostic tools. However, the advent of molecular diagnostic techniques, particularly reverse transcription polymerase chain reaction (RT-PCR) and real-time quantitative RT-PCR (RT-qPCR), has dramatically improved our capacity to detect PTV RNA in clinical specimens, revealing infection rates that are substantially higher than previously appreciated. A large-scale surveillance study employing a novel quadruplex RT-qPCR assay for the simultaneous detection of PTV, porcine sapelovirus (PSV), porcine kobuvirus (PKV), and porcine enterovirus G (EV-G) tested 1,823 fecal samples collected from various pig farms and reported an overall PTV positivity rate of 18.82% (343/1,823) [1]. This finding underscores the endemic nature of PTV circulation in commercial swine operations and highlights the frequent co-circulation of multiple enteric picornaviruses within the same herds.

Regional prevalence studies have corroborated these findings while revealing notable geographic variation. In Hunan Province, China, a comprehensive investigation of 261 fecal and 91 intestinal content samples collected from 29 farms detected an overall PTV-positivity rate of 19.03% by RT-PCR, with a particularly high infection rate circulating among asymptomatic fattening and nursery pigs [19]. Similarly, a study conducted in the Bareilly district of Uttar Pradesh, India, screened 106 fecal and intestinal content samples and found 19.8% (21/106) to be positive for PTV [28]. In Switzerland, a multiplex RT-PCR investigation of 363 clinical samples from 282 animals revealed strikingly higher frequencies in fecal samples: PTV was detected in 47% of healthy pigs and 54% of diseased pigs, with co-detections of PTV, PSV, and EV-G being the most common finding [10]. This discrepancy in prevalence rates between studies likely reflects differences in sampling strategies (targeted versus random), diagnostic assays employed, age distribution of sampled animals, and the health status of the herds under investigation.

The age-specific epidemiology of PTV infection is particularly instructive. In the Indian study, the highest prevalence was observed in the nursery age group (32.6%, 17/52), followed by adult pigs (13.3%, 4/30), with no PTV detected in suckling piglets [28]. This pattern aligns with the waning of maternal antibody protection and the subsequent exposure to environmental virus during the post-weaning period. The Swiss study similarly reported that PTV was detected more frequently in fecal samples from weaned and fattening pigs compared to suckling piglets and sows [10]. These findings have important implications for biosecurity and management practices, suggesting that the nursery-to-finishing phase represents a critical window for viral transmission and amplification within herds.

Geographic Distribution and Serotypic Diversity

PTV exhibits a truly global distribution, with documented circulation across Europe, Asia, the Americas, and Oceania. The virus has been reported in domestic pigs (Sus scrofa domesticus) and wild boar (Sus scrofa) populations across diverse ecological and climatic zones, from temperate Europe to subtropical Asia and tropical South America. Historically, PTV was first recognized as the etiological agent of Teschen disease, a severe and often fatal polioencephalomyelitis that caused devastating economic losses in European swine herds between the 1920s and 1960s [26]. Following the implementation of rigorous control measures, including the use of inactivated vaccines in endemic regions, the incidence of classical Teschen disease declined dramatically, and PTV variants with lower pathogenicity became predominant globally [26]. However, as recent outbreaks demonstrate, highly pathogenic strains have not been eradicated and continue to emerge sporadically.

The genetic diversity of PTV is extraordinary, with at least 22 genotypes (PTV 1–22) now recognized within the species Teschovirus A, and evidence suggesting the existence of additional novel species within the genus [8, 18]. A seminal study in Hunan, China, identified multiple PTV genotypes co-circulating in pig populations, including all known genotypes except PTV 7 and 8, and further identified nine novel genotypes provisionally designated PTV 14–22 [18]. This remarkable diversity is not merely a taxonomic curiosity; it has profound implications for vaccine development, diagnostic accuracy, and our understanding of viral pathogenesis. The VP1 capsid protein, which contains the major neutralizing epitopes, is the primary determinant of serotype specificity. A highly conserved epitope (RNNQIPQDF) located on the GH loop of VP1 has been identified and shown to induce group-specific antisera with pan-PTV potential, offering a promising target for broadly reactive diagnostic tools [16]. Nevertheless, the existence of multiple serotypes that do not cross-neutralize complicates vaccine strategies and necessitates ongoing surveillance to monitor the emergence of antigenically novel strains.

Emergence of Highly Pathogenic Strains and Outbreak Investigations

While most PTV infections are subclinical or associated with mild, self-limiting enteric disease, the sporadic emergence of highly virulent neuroinvasive strains remains a significant concern for the global swine industry. A notable outbreak occurred in a large-scale pig farm in China, where a novel PTV-2 strain, designated HeNZ1, caused severe neurological symptoms, including lethargy, locomotor ataxia, nystagmus, hind limb paralysis, and coma, in suckling piglets, with a mortality rate reaching 38% [7]. This mortality rate is remarkably high in the context of PTV history and underscores the potential for catastrophic losses when virulent strains emerge. Phylogenetic and evolutionary divergence analyses revealed that HeNZ1 shares the highest VP1 sequence similarity with European PTV-2 strains rather than Chinese domestic strains, suggesting a possible transboundary introduction or long-distance dispersal event [7]. Multiple sequence alignment and B cell epitope prediction identified 36 unique mutation sites in VP1 and VP2, 19 of which are located in B cell epitopes and exposed on the virion surface, implying significant antigenic drift potential [7].

Similarly, a highly pathogenic PTV-5 strain emerged in western China (Gansu Province) in 2022, causing diarrhea with high mortality [26]. This isolate, characterized by transmission electron microscopy, immunofluorescence assays, and growth kinetic analysis, resulted in serious respiratory distress, watery diarrhea, paralysis, and high mortality in challenged pigs. Histopathological examination revealed lymphocyte infiltration and hemorrhage in multiple tissues, including interstitial pneumonia [26]. The identification of such strains, which deviate from the typically low-pathogenicity profile of contemporary PTV isolates, serves as a stark reminder that the virus retains the capacity for severe disease expression under appropriate epidemiological and host conditions.

In Europe, neuroinvasive PTV strains have also been documented in recent years. In the Netherlands, two unrelated pig farms experienced outbreaks of progressive hind limb paresis and paralysis in weanling pigs, with morbidity rates up to 5% [15]. Full-length genome sequencing identified PTV-3 (98% identity) and PTV-11 (85% identity) as the causative agents, and histopathological examination confirmed non-suppurative encephalomyelitis consistent with viral infection [15]. In southern Germany, a fattening farm reported hind limb paralysis in two age groups (50 kg and 60 kg) over a four-week period, with a morbidity of 3.3% but a high case-fatality rate necessitating euthanasia of most affected animals [12]. Immunohistochemistry revealed PTV antigen in neurons, glial cells, endothelial cells, and mononuclear cells throughout the central nervous system, and phylogenetic analysis demonstrated close relation (88% full genome identity) to PTV A11 strain "Dresden" [12]. These European outbreaks, while limited in scale, demonstrate that neuroinvasive PTV continues to circulate and cause clinical disease in modern swine production systems.

Wild Boar as a Reservoir and Epidemiological Bridge

The role of wild boar (Sus scrofa) in the epidemiology of PTV has received increasing attention, as these animals may serve as a reservoir for viral maintenance and a potential bridge for transmission to domestic swine populations. Serological evidence of PTV infection in wild boar has been documented across multiple continents. In France, a combined serological, virological, and phylogenetic screening of hunted wild boars from the Drôme and Marne departments revealed high seroprevalence rates, up to 38%, suggesting endemic circulation of PTV in these populations [14]. Genetic sequencing of PCR-positive samples indicated the presence of a novel PTV genotype, raising questions about the pathogenicity of this strain and its potential for spillover into domestic herds [14].

In Hungary, the first complete PTV genome sequence from wild boar was obtained from 7 of 10 fecal samples (70% positivity) collected from wild boar piglets [24]. The strain WB2C-TV/2011/HUN showed considerable genetic divergence, especially in VP1 (66–74% amino acid identity compared to available PTVs), and was provisionally designated as a novel genotype, PTV-13 [24]. This study demonstrated that wild boar harbor genetically distinct PTV strains that may not be captured by surveillance focused exclusively on domestic pigs. In Brazil, a molecular survey of captive wild boars in Paraná state detected PTV in fecal samples from asymptomatic animals, confirming that wild boar can serve as healthy carriers [32]. The Indian study further reported that PTV isolates from domestic piglets clustered with PTV-13 strain WB2C-TV/2011/HUN from Hungarian wild boar and PTV-2 strain from Germany, suggesting potential international transmission links [27].

The epidemiological significance of wild boar as a reservoir is amplified by their increasing populations, expanding geographic ranges, and frequent contact with domestic swine through shared habitats, foraging areas, and inadequate biosecurity measures. The World Organisation for Animal Health (WOAH) recognizes the importance of wildlife surveillance for emerging infectious diseases, and PTV represents a compelling case study for the need to integrate wild boar monitoring into national swine health programs.

Recombination and Evolutionary Dynamics

The genetic diversity and global distribution of PTV are profoundly influenced by recombination, a process that generates novel viral variants with potentially altered pathogenic, antigenic, and transmission characteristics. Recombination analysis of 40 PTV isolates from Hunan, China, revealed nine recombination events, including both inter- and intra-serotype recombination [19]. This study further demonstrated that only limited positive selection is acting on the global population of PTV isolates, with purifying selection predominating [19]. The implications of this finding are significant: while most mutations are deleterious and eliminated by purifying selection, recombination allows for the rapid generation of genetic diversity without the accumulation of harmful mutations, facilitating immune evasion and adaptation to new hosts or environments.

A particularly striking example of recombination was identified in the PTV-5 strain SD2023, isolated from Shandong Province, China [13]. Full-length genomic sequencing and recombination analysis using RDP4 and SimPlot software revealed that SD2023 is a novel recombinant, featuring a P1 gene (nt 876–3372) derived from a PTV-5 HuN33-like minor parent, with flanking regions originating from a PTV-4 10BJ02-like major parent [13]. This chimeric genome structure, supported by robust statistical analyses, demonstrates that recombination can occur between different PTV genotypes, potentially generating strains with novel tissue tropism or virulence properties. The SD2023 strain caused mild neurological symptoms and anorexia in experimentally infected piglets, with lymphocyte infiltration observed in brain tissues, confirming its pathogenic potential [13].

The discovery of novel PTV species and genotypes continues to expand our understanding of the genus. Metagenomic analysis of porcine feces in Japan identified PTV-related viruses that are distantly related to Teschovirus A, exhibiting only 62.2–79.0% amino acid identity in the P1 region compared to 76.5–92.1% identity among PTV 1–13 strains [23]. These viruses share a type IV internal ribosomal entry site and conserved characteristic motifs in the coding region, yet their genetic divergence suggests they may represent a novel species within the genus Teschovirus [23]. Similarly, the identification of PTV 21-HuN41 and PTV 21-HuN42, which share only 70.1–71.9% nucleotide identity with other known PTVs in the polyprotein gene, led to the tentative designation of a new teschovirus species, Teschovirus B [18]. These findings underscore the vast, largely unexplored genetic diversity of teschoviruses and highlight the need for continued metagenomic surveillance.

Co-infection Dynamics and Disease Association

PTV frequently circulates in complex co-infection scenarios with other swine pathogens, complicating the attribution of clinical disease to any single agent. The quadruplex RT-qPCR study that detected PTV in 18.82% of samples also found PSV (15.25%), PKV (21.72%), and EV-G (27.10%), with high rates of co-detection [1]. In Switzerland, co-detections of all three enteric picornaviruses (PTV, PSV, and EV-G) were the most common finding in fecal samples, and statistical analysis yielded no evidence for an association between virus detection and disease [10]. This observation raises fundamental questions about the pathogenic role of PTV in enteric disease: is PTV a primary pathogen, a contributing factor in multifactorial disease, or an incidental bystander?

The answer likely depends on the specific viral genotype, host factors (age, immune status, genetics), and the presence of co-infecting pathogens. A case report from Japan described a piglet with concurrent polioencephalomyelitis due to PTV and postweaning multisystemic wasting syndrome (PMWS) associated with porcine circovirus type 2 (PCV2) [29]. The authors suggested that the immunosuppressive condition developing in PMWS may have facilitated the infection of the brain with PTV, illustrating how co-infections can alter the clinical expression of PTV infection [29]. In China, a triplex real-time PCR assay for pseudorabies virus (PRV), PTV-1, and Streptococcus suis serotype 2 (SS2) revealed single infection rates of 26.95% for PTV-1, with co-infection rates of 7.45% for PRV+PTV-1 and 1.77% for all three pathogens [3]. These data highlight the importance of differential diagnosis in cases of neurological disease, as multiple pathogens can produce overlapping clinical signs.

Temporal and Seasonal Patterns

Seasonal variation in PTV prevalence has been documented, although the patterns are not uniform across geographic regions. In the Indian study, prevalence was highest in winter compared to summer [28]. This seasonality may reflect environmental stability of the non-enveloped virion, which is highly resistant to inactivation. Comparative inactivation studies have demonstrated that PTV is significantly more resistant to chemical disinfectants than enveloped viruses such as Aujeszky's disease virus (pseudorabies virus) and vesicular stomatitis virus [30]. Among five disinfectants tested, only 0.2% Mikasept KP achieved a 4-log reduction in PTV titre after 30 minutes of contact time in both suspension and carrier tests [30]. This environmental persistence likely contributes to the year-round circulation of PTV in swine populations and poses challenges for biosecurity protocols.

The historical epidemiology of PTV also reveals temporal shifts in virulence. Between the 1920s and 1960s, highly pathogenic strains caused devastating outbreaks of Teschen disease in Europe, with mortality rates approaching 100% in naive populations [26]. Following the widespread use of inactivated vaccines and improved hygiene measures, the disease receded, and PTV became associated primarily with subclinical or mild infections. However, the recent emergence of highly pathogenic strains in China [7, 26] and Europe [12, 15] suggests that virulence may cycle over decadal timescales, possibly driven by the accumulation of genetic changes in circulating strains or the introduction of novel genotypes into immunologically naive populations.

Implications for Surveillance and Control

The epidemiological patterns described above have direct implications for the design of surveillance and control programs. The high prevalence of PTV in asymptomatic pigs, particularly in the nursery-to-finishing phase, means that diagnostic efforts focused solely on clinically affected animals will underestimate the true burden of infection. The frequent co-detection of PTV with other enteric viruses necessitates the use of multiplex diagnostic assays capable of simultaneous detection and differentiation [1, 3]. The development of rapid, point-of-care diagnostic tools, such as the reverse transcription recombinase-aided amplification coupled with lateral flow dipstick (RT-RAA-LFD) assay, which can detect PTV in less than 25 minutes with a detection limit of 10 copies/μL, offers promise for field-deployable surveillance [6].

The genetic and antigenic diversity of PTV poses a major challenge for vaccine development. The identification of 22 genotypes and the existence of multiple serotypes that do not cross-neutralize means that a monovalent vaccine is unlikely to provide broad protection. However, the discovery of a highly conserved epitope (RNNQIPQDF) on the VP1 GH loop that induces group-specific antisera reactive against all tested PTV serotypes (PTV 1–7) offers a potential target for a pan-PTV vaccine or diagnostic reagent [16]. The use of the PTV-1 2A peptide, which exhibits the highest cleavage efficiency among picornavirus 2A peptides in human cell lines, zebrafish, and mice, has also been exploited for biotechnological applications, including the construction of multicistronic expression vectors and the development of recombinant PTV as a vaccine vector [9, 17, 20, 31].

The World Organisation for Animal Health (WOAH) lists PTV as a pathogen of economic significance, and the Food and Agriculture Organization of the United Nations (FAO) recognizes the importance of swine viral diseases for global food security. The sporadic emergence of highly pathogenic strains, the expanding wild boar reservoir, and the potential for transboundary spread through international trade in pigs and pork products underscore the need for coordinated international surveillance efforts. The development of standardized genotyping protocols, the establishment of reference strain collections, and the sharing of sequence data through public databases are essential

Diagnostic Approaches and Molecular Detection Methods

The accurate and timely diagnosis of porcine teschovirus (PTV) infection presents a formidable challenge to veterinary diagnosticians, given the virus's capacity to induce a spectrum of clinical manifestations ranging from subclinical enteric carriage to fatal polioencephalomyelitis. The diagnostic landscape for PTV has evolved considerably from traditional virus isolation and serotyping to sophisticated molecular platforms capable of simultaneous detection, genotyping, and quantification. This section provides an exhaustive analysis of contemporary diagnostic approaches, with particular emphasis on molecular detection methods that have revolutionized our capacity to identify, characterize, and surveil this economically significant pathogen.

Conventional Virus Isolation and Serological Detection

Historically, the gold standard for PTV diagnosis involved virus isolation in porcine kidney cell lines, particularly PK-15 and swine testicular (ST) cells, coupled with serum neutralization tests (SNT) for serotype identification [8, 11]. The cytopathic effect (CPE) induced by PTV in these cell cultures is characteristically rapid, with infected cells demonstrating rounding, detachment, and eventual lysis within 24–72 hours post-inoculation [13, 22]. The isolation protocol described by Yang et al. [8] successfully recovered 58 PTV strains from clinical samples using both PK-15 and ST cells, demonstrating that cell culture remains a viable approach for virus amplification prior to downstream characterization. However, the utility of virus isolation is substantially constrained by the existence of 13 recognized serotypes and the emergence of novel genotypes that may not be neutralized by reference antisera [14, 18]. The serological landscape is further complicated by the high prevalence of subclinical infections, as demonstrated by Stäubli et al. [10], who detected PTV in 47% of fecal samples from healthy Swiss pigs, and by Sereika et al. [34], who found PTV-1 antibodies in virtually all Lithuanian swine farms despite the absence of clinical Teschen disease.

Serological approaches, while valuable for epidemiological surveillance, suffer from inherent limitations related to cross-reactivity among closely related picornaviruses and the inability to distinguish between naturally infected and vaccinated animals. The virus neutralization test (VNT) remains the reference method for serotype determination, yet its application is labor-intensive, requires live virus handling, and demands a comprehensive panel of serotype-specific antisera that may not be available for newly described genotypes [33]. The work of Derevianko [33] in Ukraine demonstrated the feasibility of using VNT with hyperimmune rabbit sera to differentiate PTV serotypes, but this approach is impractical for large-scale surveillance programs. A significant advancement in serological diagnosis came from the identification of a highly conserved epitope (RNNQIPQDF) on the VP1 GH loop, which Tsai et al. [16] exploited to generate a pan-PTV antiserum reactive against all tested serotypes (PTV 1–7). This bioinformatics-predicted diagnostic reagent represents a promising tool for screening purposes, though its inability to discriminate between serotypes limits its utility for genotype-specific investigations.

Reverse Transcription Polymerase Chain Reaction (RT-PCR)

The advent of molecular detection methods marked a paradigm shift in PTV diagnostics, with RT-PCR emerging as the cornerstone of laboratory diagnosis. The 5' untranslated region (UTR) of the PTV genome, which contains highly conserved secondary structures essential for viral replication and translation, has been the preferred target for generic PTV detection assays [21, 27]. Patel et al. [27] successfully amplified the 5' UTR from Indian PTV isolates, enabling subsequent phylogenetic characterization that revealed clustering with European strains. The RT-PCR protocol developed by Prodělalová et al. [21] for reclassifying historical Czechoslovakian isolates demonstrated remarkable versatility, detecting teschovirus-specific amplicons in all 27 archival strains while simultaneously differentiating PTV-1 through serotype-specific primers. This dual-amplicon approach, genus-level screening followed by serotype-specific confirmation, established a template that has been widely adopted in subsequent investigations.

The epidemiological utility of RT-PCR was compellingly illustrated by Yang et al. [19], who screened 352 samples from 29 farms in Hunan, China, achieving a 19.03% overall positivity rate and identifying seven distinct PTV serotypes (PTV 2–6, 9, and 11). Similarly, John et al. [28] employed RT-PCR targeting the 5' UTR to document a 19.8% prevalence in Indian swine, with age-stratified analysis revealing highest detection rates in nursery pigs (32.6%) compared to adults (13.3%) and suckling piglets (0%). These epidemiological findings underscore the critical importance of standardized RT-PCR protocols for generating comparable prevalence data across geographic regions. However, conventional RT-PCR suffers from several inherent limitations: it is qualitative rather than quantitative, requires post-amplification gel electrophoresis that increases contamination risk, and provides no information regarding viral load, which may correlate with disease severity.

Real-Time Quantitative PCR (qPCR) and Multiplex Assays

The transition to real-time quantitative PCR (qPCR) technologies addressed many limitations of conventional RT-PCR while introducing enhanced sensitivity, quantification capability, and the potential for multiplex detection. Li et al. [1] developed a quadruplex RT-qPCR assay targeting PTV alongside porcine sapelovirus (PSV), porcine kobuvirus (PKV), and enterovirus G (EV-G), achieving detection limits sufficient for routine surveillance. This assay demonstrated exceptional performance characteristics, with coincidence rates exceeding 99.01% compared to reference singleplex methods when applied to 1,823 fecal samples from Chinese pig farms. The simultaneous detection capability is particularly valuable given the high frequency of co-infections documented in swine populations; Stäubli et al. [10] found that co-detection of PTV, PSV-A, and EV-G was the most common finding in Swiss pigs, with 54% of diseased animals harboring PTV alongside other enteric picornaviruses.

The clinical context of neurological disease presentations has driven the development of targeted multiplex assays for pathogens causing central nervous system infections. Lai et al. [3] designed a triplex real-time PCR assay for simultaneous detection of pseudorabies virus (PRV), PTV-1, and Streptococcus suis type 2, achieving a detection limit of 1.0 × 10⁰ copies/μL for each target. When applied to 282 clinical samples from pigs with neurological symptoms, this assay revealed single infection rates of 26.95% for PTV-1 and co-infection rates of 7.45% for PRV + PTV-1, highlighting the diagnostic complexity of neurological disease presentations. The importance of such multiplex approaches is underscored by the case report of Takahashi et al. [29], who documented concurrent polioencephalomyelitis due to PTV and postweaning multisystemic wasting syndrome (PMWS) associated with porcine circovirus type 2 (PCV2), suggesting that immunosuppressive conditions may facilitate neuroinvasion by PTV.

Isothermal Amplification Methods for Point-of-Care Diagnostics

The development of isothermal amplification technologies has addressed the critical need for rapid, equipment-independent diagnostic methods suitable for field deployment and resource-limited laboratories. Chen et al. [6] developed a reverse transcription recombinase-aided amplification (RT-RAA) assay coupled with lateral flow dipstick (LFD) detection for PTV, achieving a detection limit of 10 copies/μL with no cross-reactivity to other swine enteric viruses. The assay's operational simplicity, isothermal amplification at 37°C for 20 minutes followed by visual readout within 5 minutes, represents a substantial improvement over conventional thermal cycling methods. When validated against 128 clinical samples, the RT-RAA-LFD assay demonstrated 98.44% total diagnostic coincidence with RT-PCR (Kappa = 0.96), indicating excellent agreement. This platform is particularly well-suited for surveillance programs in regions without access to sophisticated laboratory infrastructure, where PTV detection has historically been underreported.

The practical advantages of isothermal methods extend beyond speed and simplicity. The closed-tube format of LFD detection minimizes amplicon contamination risk, a persistent challenge in conventional PCR workflows, while the visual readout eliminates the need for specialized equipment. However, the current RT-RAA-LFD assay is limited to qualitative detection and does not provide genotype-level discrimination, which may be required for epidemiological investigations and virulence assessment. Future developments may integrate isothermal amplification with portable sequencing technologies or multiplex LFD strips capable of differentiating the expanding array of PTV genotypes.

Next-Generation Sequencing and Metagenomic Approaches

The application of next-generation sequencing (NGS) and metagenomic analysis has revolutionized our understanding of PTV genetic diversity and has proven instrumental in identifying novel genotypes and recombinant strains. The discovery of PTV-13 in Hungarian wild boars by Boros et al. [24] was achieved through pyrosequencing of fecal samples, revealing a VP1 amino acid identity of only 66–74% compared to known PTVs. This approach has since been applied to identify numerous novel genotypes, including PTV 14–22 in Chinese pig populations [18] and a putative novel species (Teschovirus B) in Japan [23]. The diagnostic value of NGS was dramatically demonstrated by Liang et al. [7], who used sequencing of a novel PTV-2 strain (HeNZ1) isolated from an outbreak with 38% mortality in suckling piglets, revealing 36 unique mutation sites in VP1 and VP2 that may confer enhanced virulence and antigenic drift potential.

Metagenomic sequencing of clinical samples has also enabled the detection of PTV in complex diagnostic scenarios where conventional PCR fails to identify the etiological agent. Zhang et al. [25] employed high-throughput sequencing to characterize a PTV-1 strain from diarrheic piglets, identifying a critical mutation at VP1 position 129 that may contribute to enteric pathogenicity. Similarly, Shao et al. [26] utilized virome analysis to identify a highly pathogenic PTV-5 variant from an outbreak of diarrhea with high mortality in western China, demonstrating the capacity of NGS to detect unexpected viral agents in disease outbreaks. The recombinant strain SD2023 identified by Ye et al. [13] through full-length genomic sequencing illustrates the power of NGS for detecting recombination events that may generate novel pathogenic variants, with RDP4 and SimPlot analyses revealing a P1 gene derived from a PTV-5 minor parent and flanking regions from a PTV-4 major parent.

The application of NGS for PTV diagnostics is not without challenges. The high cost, requirement for bioinformatics expertise, and extended turnaround time limit its utility for routine clinical diagnosis. Furthermore, the detection of PTV sequences in clinical samples does not establish causation, as demonstrated by the high prevalence of PTV in healthy pigs documented in multiple studies [10, 19]. Nonetheless, NGS has become indispensable for identifying emerging strains, characterizing recombination events, and understanding the molecular epidemiology of PTV at the population level.

Recombinant Virus Technologies and Reporter Systems

The development of infectious cDNA clones and reporter virus systems has created powerful tools for both diagnostic development and pathogenesis research. Li et al. [11] constructed a full-length infectious clone of PTV-2 using seamless ligation technology, establishing a reverse genetics platform for investigating viral replication mechanisms and pathogenicity determinants. The ability to rescue recombinant viruses with defined genetic modifications enables systematic analysis of virulence factors, such as the 3C protease and VP1 protein, which Zhang et al. [2] and Li et al. [5] have implicated in evasion of host innate immune responses through degradation of NF-κB and inhibition of MDA5 activation, respectively.

Tsai et al. [31] advanced this technology by engineering a recombinant PTV expressing the iLOV fluorescent protein, enabling direct visualization of viral infection in living cells. This reporter virus, created by inserting the iLOV coding sequence into the 2A protease site, retained replication competence while demonstrating attenuation due to impaired polyprotein cleavage. Such reporter systems have substantial utility for evaluating antiviral compounds, studying virus-cell interactions, and potentially serving as the basis for diagnostic assays that detect viral replication through fluorescent signal generation. The attenuation observed in iLOV-tagged virus also suggests potential applications in vaccine development, though further research is needed to assess safety and immunogenicity in swine.

Immunohistochemistry and Antigen Detection

Despite the predominance of molecular methods, antigen detection through immunohistochemistry (IHC) remains valuable for diagnosing PTV-associated neurological disease in formalin-fixed tissues. Stadler et al. [12] employed IHC to demonstrate PTV antigen in neurons, glial cells, endothelial cells, and mononuclear inflammatory cells within the central nervous system of fattening pigs with hind limb paralysis, confirming the neurotropic nature of the infecting PTV-11 strain. The distribution of viral antigen in the brainstem and spinal cord correlated with the histopathological lesions of non-suppurative encephalomyelitis, providing spatial evidence of neuroinvasion. Takahashi et al. [29] similarly used IHC to localize PTV antigens in brainstem lesions of a piglet with concurrent polioencephalomyelitis and PMWS, demonstrating the utility of IHC for documenting dual infections.

The sensitivity of IHC is generally lower than molecular detection methods, and the technique requires specific antibodies that may not be available for all serotypes. However, the pan-PTV epitope identified by Tsai et al. [16] may enable development of broadly reactive antibodies suitable for IHC applications. In the absence of such reagents, the high genetic diversity of PTV complicates antigen detection, as antibody binding may be affected by sequence variation in capsid proteins.

Diagnostic Considerations for Special Populations

The detection of PTV in wild boar populations presents unique diagnostic challenges and epidemiological implications. Hirschinger et al. [14] conducted serological and virological screening of French wild boars, revealing seroprevalence rates up to 38% and the presence of a novel PTV genotype. The combination of serological testing with PCR-based genotyping proved essential for distinguishing between past exposure and active infection, particularly in populations where sampling opportunities are limited to hunting seasons. Donin et al. [32] applied RT-PCR to detect PTV in captive wild boars in Brazil, achieving a 27.8% positivity rate and demonstrating that wild boars serve as reservoirs capable of transmitting virus to domestic swine populations.

The detection of PTV in archived and historical samples has been facilitated by the stability of viral RNA in formalin-fixed tissues and the application of RT-PCR to degraded nucleic acids. Prodělalová et al. [21] successfully amplified teschovirus sequences from strains isolated between 1960 and 1980, enabling reclassification and phylogenetic comparison with contemporary isolates. This retrospective diagnostic capability is essential for understanding the evolutionary dynamics of PTV and identifying long-term trends in genotype distribution.

Quality Assurance and Validation Considerations

The development and validation of molecular diagnostic assays for PTV require rigorous attention to quality assurance parameters, including analytical sensitivity, specificity, repeatability, and reproducibility. The multiplex RT-qPCR assay described by Li et al. [1] demonstrated excellent performance characteristics, with intra-assay and inter-assay coefficients of variation below 3% and no cross-reactivity with other swine viruses. Similarly, the triplex real-time PCR developed by Lai et al. [3] showed 100% concordance with commercial singleplex kits when testing clinical samples, validating the multiplex format for routine diagnostic use.

The choice of target gene significantly influences assay performance. The 5' UTR remains the most conserved region suitable for pan-PTV detection, while VP1 is required for serotype discrimination due to its genetic variability [19, 27]. The expanding number of recognized genotypes, currently including at least 22 PTV genotypes within Teschovirus A and proposed novel species within Teschovirus B [18, 23], necessitates periodic reassessment of primer and probe binding sites to ensure continued detection of emerging variants. The discovery of PTV-19 by Yang et al. [8] through phylogenetic analysis of VP1 sequences demonstrates that genotype discovery often precedes the development of genotype-specific diagnostic assays.

Sample Selection and Processing

The selection of appropriate clinical specimens is critical for maximizing diagnostic sensitivity. Fecal samples and intestinal contents are the specimens of choice for detecting enteric PTV infection, with studies reporting high detection rates in diarrheic and asymptomatic pigs alike [10, 19, 28]. For neurological disease presentations, brain tissue samples, particularly from the brainstem and spinal cord, yield the highest diagnostic sensitivity, as demonstrated by the detection of PTV in

Control Strategies and Preventive Measures in Swine Herds

The control of Porcine teschovirus (PTV) within swine populations presents a formidable challenge to veterinarians and producers alike, owing to the virus’s ubiquity, genetic plasticity, and complex epidemiology. PTV is an environmentally resilient, non-enveloped RNA virus belonging to the family Picornaviridae, and its control strategies must be formulated with a deep appreciation for its biological properties, transmission dynamics, and the intricate host-pathogen interactions that define its pathogenesis. A multi-layered, integrated approach, encompassing rigorous biosecurity, strategic surveillance, immunological intervention, and population management, is essential, particularly given the emergence of highly virulent strains capable of causing catastrophic losses in naive herds [7, 26].

Biosecurity, Disinfection, and Environmental Containment

The foundational pillar of any PTV control program is robust biosecurity. The virus is shed in high titers in feces and can persist in the environment, facilitating both direct fecal-oral transmission and indirect spread via fomites, contaminated equipment, and personnel [10, 19]. The resistance of PTV to inactivation is a critical concern; as a non-enveloped virus, it is significantly more refractory to chemical disinfection compared to enveloped pathogens such as Aujeszky’s disease virus (Pseudorabies virus). Comparative inactivation studies have demonstrated that common disinfectants, including 1% Chloramin BM, 1% Incidin Plus, and 2% Sekusept Forte, are insufficient to achieve a 4-log reduction of PTV in suspension or on contaminated surfaces, failing to meet standard virucidal efficacy criteria [30]. Critically, the inactivation of surface-bound virus is markedly more difficult than inactivation in suspension, underscoring the need for rigorous mechanical cleaning to remove organic matter prior to disinfection [30]. Among tested agents, only 0.2% Mikasept KP demonstrated reliable virucidal activity against PTV under both suspension and carrier test conditions [30]. Therefore, disinfection protocols on swine farms must be specifically validated against non-enveloped viruses, employing peracetic acid-based or other high-efficacy formulations, and must prioritize the decontamination of farrowing crates, nursery pens, and transport vehicles.

Beyond chemical disinfection, all-in/all-out production flow by age group is a critical managerial tool. PTV infection rates are significantly higher in weaned and fattening pigs than in suckling piglets or sows, likely due to the waning of maternal immunity and increased contact rates [10, 28]. Continuous flow systems permit the uninterrupted circulation of PTV among susceptible cohorts. Implementing strict downtime between groups, coupled with thorough cleaning and disinfection, can break the cycle of endemic transmission. Furthermore, the role of wild boar (Sus scrofa) as a reservoir for PTV cannot be overstated. High seroprevalence rates (up to 38%) and the detection of both known and novel PTV genotypes in wild boar populations in France, Hungary, and Brazil indicate that wildlife interfaces represent a constant threat for viral introduction into domestic herds [14, 24, 32]. Control measures must therefore include the construction of wildlife-proof fencing, the management of feed storage to avoid attracting feral swine, and the implementation of regional surveillance programs to monitor viral circulation in the feral reservoir.

Advanced Diagnostics and Active Surveillance

Effective control is predicated on rapid and accurate detection. The clinical presentation of PTV, ranging from asymptomatic enteric infection to severe polioencephalomyelitis, is non-pathognomonic and overlaps with other causes of neurological and enteric disease in swine, including Pseudorabies virus, Streptococcus suis, Porcine sapelovirus, and Porcine enterovirus G [3, 10]. Consequently, laboratory confirmation is indispensable. Significant advances in molecular diagnostics have yielded a suite of tools that enable the simultaneous detection and differentiation of these pathogens. Quadruplex and triplex real-time RT-PCR/PCR assays have been developed and validated, achieving high sensitivity (detection limits as low as 1.0×10⁰ copies/μL) and specificity, with diagnostic coincidence rates exceeding 99% compared to reference methods [1, 3]. These multiplex assays are invaluable for differential diagnosis in herds presenting with neurological or diarrheal disease, allowing for the rapid identification of the etiological agent or agents involved, particularly important given the high frequency of co-infections observed in clinical settings [3, 29].

For field-level surveillance and resource-limited settings, point-of-care molecular tools have emerged as a transformative technology. The reverse transcription recombinase-aided amplification combined with lateral flow dipstick (RT-RAA-LFD) assay provides a rapid, visual, and equipment-free method for PTV detection, yielding results in under 25 minutes with a detection limit of 10 copies/μL and a total diagnostic coincidence rate of 98.44% compared to conventional RT-PCR [6]. This technology is particularly suited for on-farm screening during outbreak investigations or for routine monitoring in low-resource environments [6]. Surveillance strategies should be risk-based, focusing on nursery and grower pigs, where prevalence is highest, and targeting fecal samples for enteric strains or brain/spinal cord tissue for neurovirulent strains [28]. Given the high frequency of subclinical infection, routine surveillance in healthy populations is essential to establishing baseline prevalence and detecting the emergence of novel genotypes [10, 19].

Vaccination, Immunological Strategies, and the Challenge of Antigenic Diversity

The development of effective vaccines against PTV is fraught with biological hurdles, primarily the extraordinary antigenic and genetic diversity within the species Teschovirus A. With at least 19 recognized genotypes and multiple serotypes that do not confer robust cross-protection, the creation of a broadly protective vaccine is a formidable challenge [8, 18]. Historically, inactivated whole-virus vaccines based on local prevalent serotypes (e.g., PTV-1 strains Dniprovsky-34) have been used to elicit neutralizing antibodies and provide clinical protection, with immunity persisting for up to six months post-vaccination [33]. However, the emergence of novel, highly virulent strains that are antigenically distinct from vaccine strains, such as the PTV-2 strain HeNZ1 from China, which possesses 36 unique mutation sites and 10 altered B-cell epitopes in VP1 and VP2, raises serious concerns about vaccine escape and the need for continuous antigenic monitoring [7].

The viral mechanisms of immune evasion further complicate vaccine design. PTV employs sophisticated strategies to subvert host antiviral innate immunity. The 3Cpro protease directly antagonizes the NF-κB signaling pathway by cleaving and degrading the NF-κB p65 subunit, thereby blocking the induction of type I interferons (IFN-β) [2]. Simultaneously, 3Cpro inhibits the activation of the NLRP3 inflammasome and pyroptosis by degrading NLRP3, IL-1β, and GSDMD, dismantling key inflammatory responses [4]. The structural protein VP1 further contributes to immune evasion by interacting with the MDA5 receptor and blocking its activation, thereby inhibiting IFN-β expression [5]. These findings imply that a successful vaccine may need to elicit robust antibody responses that neutralize viral entry and, ideally, induce cytotoxic T-cell responses targeting non-structural proteins to eliminate infected cells before these immune evasion mechanisms are established.

In this context, the identification of a highly conserved B-cell epitope (RNNQIPQDF) within the GH loop of VP1 offers a promising avenue for pan-PTV serological diagnostics and potentially for vaccine design. Antiserum raised against a recombinant protein bearing this epitope demonstrated reactivity against all tested PTV serotypes (1–7) without cross-reacting with other picornaviruses [16]. This epitope could serve as a component of a subunit vaccine designed to induce broadly reactive, non-neutralizing antibodies for opsonization or as a diagnostic marker to differentiate infected from vaccinated animals (DIVA) [16]. Furthermore, reverse genetics platforms and infectious cDNA clones have been established for PTV, enabling the rational design of attenuated live vaccines. The insertion of a fluorescent protein (iLOV) into the 2A protease site of a recombinant PTV resulted in a replication-competent but attenuated virus, suggesting that genetic manipulation of the 2A region, known for its high cleavage efficiency and critical role in polyprotein processing, can yield safe, live-attenuated vaccine candidates [9, 31]. Additionally, the construction of a full-length infectious clone of a contemporary PTV-2 strain provides a vital platform for studying virulence determinants and for the rapid generation of marker vaccines tailored to emerging field strains [11].

Finally, the management of concurrent immunosuppressive infections is a critical preventive measure. Co-infection with Porcine circovirus type 2 (PCV2), which causes postweaning multisystemic wasting syndrome (PMWS), can exacerbate PTV pathogenesis. The immunosuppression induced by PCV2 is hypothesized to facilitate neuroinvasion by PTV, leading to severe polioencephalomyelitis [29]. Therefore, comprehensive herd health programs that include vaccination against PCV2 and other immunosuppressive agents, such as PRRSV, are an integral component of PTV control, as they reduce the susceptibility of the host to severe PTV disease. In summary, the control of PTV demands a holistic strategy: rigorous, validated biosecurity to limit environmental persistence and wildlife incursion; deployment of rapid, multiplex molecular diagnostics for surveillance and outbreak response; and the continued development and strategic application of vaccines that account for the virus’s profound antigenic diversity and its arsenal of immune evasion mechanisms.

References

[1] Li B, Shi K, Shi Y, Feng S, Yin Y, Lu W, et al.. A Quadruplex RT-qPCR for the Detection of Porcine Sapelovirus, Porcine Kobuvirus, Porcine Teschovirus, and Porcine Enterovirus G. Animals. 2025. DOI: https://doi.org/10.3390/ani15071008

[2] Zhang X, Li Y, Zhou Y, Chen W, Xie L, Hu Y, et al.. Porcine Teschovirus 2 3Cpro Evades Host Antiviral Innate Immunity by Inhibiting the IFN-β Signaling Pathway. Microorganisms. 2025. DOI: https://doi.org/10.3390/microorganisms13061209

[3] Lai R, Yang C, Wu L, Wu W, Li L, Liu W, et al.. Development and implementation of a TaqMan triplex real-time PCR assay for concurrent detection of pseudorabies virus, porcine teschovirus 1, and Streptococcus suis 2. Frontiers in Veterinary Science. 2025. DOI: https://doi.org/10.3389/fvets.2025.1589175

[4] Zhang X, Li Y, Chen W, Zhou Y, Zhou L, Xie L, et al.. The underlying mechanism of Porcine Teschovirus 2 3Cpro antagonizing the NLRP3 inflammasome.. Veterinary Microbiology. 2025. DOI: https://doi.org/10.1016/j.vetmic.2025.110479

[5] Li Y, Zhang X, Zhao B, Zhao C, Lei X, Huang H, et al.. The VP1 Protein of Porcine Teschovirus Inhibits the Innate Immune Response to Viral Infection by Blocking MDA5 Activation. Transboundary and Emerging Diseases. 2024. DOI: https://doi.org/10.1155/2024/6649669

[6] Chen Q, Sun Z, Chen R, Wu X, Che Y, Wu R, et al.. A rapid visual detection method for porcine teschovirus through reverse transcription recombinase-aided amplification coupled with lateral flow dipstick. BMC Veterinary Research. 2024. DOI: https://doi.org/10.1186/s12917-024-04442-9

[7] Liang W, Wu X, Ding Z, Zhong S, Qian X, Ye P, et al.. Identification of a novel porcine Teschovirus 2 strain as causative agent of encephalomyelitis in suckling piglets with high mortality in China. BMC Veterinary Research. 2023. DOI: https://doi.org/10.1186/s12917-022-03549-1

[8] Yang T, Lin A, Cui S, Chen J, Wan H, Lu Y, et al.. Identification of a Novel Porcine Teschovirus Subtype 19 within the Species Teschovirus A. Transboundary and Emerging Diseases. 2023. DOI: https://doi.org/10.1155/2023/9977581

[9] Kim JH, Lee S, Li L, Park H, Park J, Lee K, et al.. High Cleavage Efficiency of a 2A Peptide Derived from Porcine Teschovirus-1 in Human Cell Lines, Zebrafish and Mice. PLoS ONE. 2011. DOI: https://doi.org/10.1371/journal.pone.0018556

[10] Stäubli T, Rickli CI, Torgerson P, Fraefel C, Lechmann J. Porcine teschovirus, sapelovirus, and enterovirus in Swiss pigs: multiplex RT-PCR investigation of viral frequencies and disease association. Journal of Veterinary Diagnostic Investigation. 2021. DOI: https://doi.org/10.1177/10406387211025827

[11] Li Y, Chen S, Shi Y, Huang H, Wang W, Zheng M, et al.. Construction and characterization of the full-length cDNA of an infectious clone of emerging porcine teschovirus-2.. Pathogens and Disease. 2022. DOI: https://doi.org/10.1093/femspd/ftac033

[12] Stadler J, Junker S, Gründl J, Fröhlich S, Beisl M, Zöls S, et al.. [Hind limb paralysis in fattening pigs due to a new strain of porcine Teschovirus A11].. Tierarztliche Praxis. Ausgabe G, Grosstiere/Nutztiere. 2022. DOI: https://doi.org/10.1055/a-1729-3677

[13] Ye K, Sun H, Ma Z, Sun Y, Zhang L, Bai J, et al.. Molecular characterization and pathogenicity of a porcine Teschovirus 5 isolate in Shandong Province in China.. Veterinary Microbiology. 2026. DOI: https://doi.org/10.1016/j.vetmic.2026.110989

[14] Hirschinger J, Loc’h G, Caubet C, Fusade-Boyer M, Faure E, Randon M, et al.. Detection and Characterization of a Novel Porcine Teschovirus in Wild Boars (Sus scrofa) in France.. Journal of Wildlife Diseases. 2026. DOI: https://doi.org/10.7589/JWD-D-25-00110

[15] Vreman S, Caliskan N, Harders F, Boonstra J, Peperkamp K, Ho CKY, et al.. Two novel porcine teschovirus strains as the causative agents of encephalomyelitis in the Netherlands. BMC Veterinary Research. 2020. DOI: https://doi.org/10.1186/s12917-020-2275-0

[16] Tsai T, Chang C, Wang F. A Highly Conserved Epitope (RNNQIPQDF) of Porcine teschovirus Induced a Group-Specific Antiserum: A Bioinformatics-Predicted Model with Pan-PTV Potential. Viruses. 2020. DOI: https://doi.org/10.3390/v12111225

[17] Jiao X, Sun W, Zhang Y, Liu X, Zhang Q, Wang Q, et al.. Exchanging the order of carotenogenic genes linked by porcine teschovirus-1 2A peptide enable to optimize carotenoid metabolic pathway in Saccharomyces cerevisiae. RSC Advances. 2018. DOI: https://doi.org/10.1039/c8ra06510a

[18] Yang T, Li R, Yao Q, Zhou X, Liao H, Ge M, et al.. Prevalence of Porcine teschovirus genotypes in Hunan, China: identification of novel viral species and genotypes.. Journal of General Virology. 2018. DOI: https://doi.org/10.1099/jgv.0.001129

[19] Yang T, Yu X, Luo B, Yan M, Li R, Qu T, et al.. Epidemiology and molecular characterization of Porcine teschovirus in Hunan, China.. Transboundary and Emerging Diseases. 2018. DOI: https://doi.org/10.1111/tbed.12728

[20] Tang X, Liu X, Tao G, Qin M, Yin G, Suo J, et al.. “Self-cleaving” 2A peptide from porcine teschovirus-1 mediates cleavage of dual fluorescent proteins in transgenic Eimeria tenella. Veterinary Research. 2016. DOI: https://doi.org/10.1186/s13567-016-0351-z

[21] Prodělalová J, Malenovská H, Valíček L. Genotyping of porcine teschoviruses isolated from 1960 to 1980 in the former Czechoslovakia and new Porcine teschovirus isolates obtained from piglets with diarrhoea. Veterinarni Medicina. 2018. DOI: https://doi.org/10.17221/33/2009-VETMED

[22] Chen M, Tang W, Hua X. Molecular characterization of a porcine teschovirus HuN-1 isolate proliferating in PK-15 cell. BMC Veterinary Research. 2018. DOI: https://doi.org/10.1186/s12917-018-1456-6

[23] Oba M, Naoi Y, Ito M, Masuda T, Katayama Y, Sakaguchi S, et al.. Metagenomic identification and sequence analysis of a Teschovirus A-related virus in porcine feces in Japan, 2014-2016.. Infection, Genetics and Evolution. 2018. DOI: https://doi.org/10.1016/j.meegid.2018.10.004

[24] Boros Á, Nemes C, Pankovics P, Kapusinszky B, Delwart E, Reuter G. Porcine teschovirus in wild boars in Hungary. Archives of Virology. 2012. DOI: https://doi.org/10.1007/s00705-012-1327-6

[25] Zhang B, Guo R, Xiao L, Zhong C, Yuan X, Huang J, et al.. Analysis on the genome of a teschovirus type 1 isolates with swine diarrhea.. Heliyon. 2023. DOI: https://doi.org/10.1016/j.heliyon.2023.e14710

[26] Shao Y, Lu Y, Li S, Lin S, Tang J, Tan C, et al.. Characterization of a highly pathogenic porcine Teschoviruses 5 emerged in Western China.. Virology. 2025. DOI: https://doi.org/10.1016/j.virol.2025.110398

[27] Patel SK, Rana J, Pathak M, Singh A, Agrawal A, Saikumar G. Genetic Characterization and Pathology of Porcine Teschovirus (PTV) in India. Indian Journal of Animal Research. 2024. DOI: https://doi.org/10.18805/ijar.b-5249

[28] John J, Das T, Sethi M, Kattoor JJ, Tomar N, Saikumar G. Epidemiological study of porcine teschovirus infection in pigs at Bareilly, Uttar Pradesh, India. Biological rhythm research. 2019. DOI: https://doi.org/10.1080/09291016.2019.1627645

[29] Takahashi M, Seimiya Y, Seki Y, Yamada M. A piglet with concurrent polioencephalomyelitis due to porcine teschovirus and postweaning multisystemic wasting syndrome.. Journal of Veterinary Medical Science. 2008. DOI: https://doi.org/10.1292/JVMS.70.497

[30] Dvořáková H, Prodělalová J, Reichelova M. Comparative inactivation of Aujeszky's disease virus, Porcine teschovirus and Vesicular stomatitis virus by chemical disinfectants. Veterinarni Medicina. 2018. DOI: https://doi.org/10.17221/1949-VETMED

[31] Tsai T, Chang C, Wang F. THE FEASIBILITY OF CONSTRUCTING A PORCINE TESCHOVIRUS INFECTIOUS cDNA CLONE TAGGED WITH iLOV FLUORESCENT PROTEIN RESCUED AS A VISIBLE MARKER VIRUS. Taiwan Veterinary Journal. 2021. DOI: https://doi.org/10.1142/s1682648522500056

[32] Donin D, Lemé R, Alfieri A, Alberton G, Alfieri A, Celso R, et al.. Molecular survey of porcine teschovirus, porcine sapelovirus, and enterovirus G in captive wild boars (Sus scrofa scrofa) of. . 2015. DOI: https://doi.org/10.1590/s0100-736x2015000500003

[33] Derevianko S. Antigenic and immunogenic properties of industrial strains of Porcine Teschovirus. Agroecological journal. 2015. DOI: https://doi.org/10.33730/2077-4893.4.2015.318297

[34] Sereika V, Lelešius R, Zienius D. Seroprevalence of Antibodies against Porcine Teschovirus 1 in Lithuania. Acta Veterinaria Brno. 2007. DOI: https://doi.org/10.2754/AVB200776020231