Nipah Virus Infection in Swine

The etiological agent, Nipah virus (genus Henipavirus, family Paramyxoviridae), is an enveloped, negative-sense, single-stranded RNA virus. The virus is closely related toHenipavirus hendraense, the causative agent of Hendra virus infection.

In Malaysia, retrospective analysis of archival histological specimens has revealed that Nipah virus (NiV) began infecting pigs in 1996, causing low morbidity and mortality before being identified in 1999 as the agent responsible for a human encephalitis outbreak that had begun in 1998 (1). During the 1998–1999 NiV outbreak in Malaysia and Singapore, approximately 40% of occupationally exposed workers in the swine industry developed encephalitis. Genetic evidence indicates a single introduction of NiV from fruit bats into commercial pig farms, where the virus subsequently spread through densely housed animals. Although respiratory disease in infected pigs was often subclinical, the virus proved highly contagious under intensive farming conditions, amplifying transmission to the human workforce. Although secondary infections occurred in other domestic animals (dogs, cats, and horses), these infections did not contribute to the spread of the virus.

Annual outbreaks have occurred in Bangladesh and India (2003–present), with high mortality rates in humans. Direct transmission occurs from fruit bat reservoirs to humans, sometimes through consumption of contaminated date palm sap. In 2014 in Mindanao, Philippines, horses became infected after exposure to local fruit bats, and humans were subsequently infected through close contact (slaughtering sick horses) or consumption of infected horse meat. These outbreaks demonstrate variable transmission pathways, depending on the geographic region and available susceptible animal hosts. Pteropusfruit bats (also known as flying foxes) range from the western Pacific through Southeast Asia, South Asia, and African coastal islands, including Madagascar. Antibody detection by ELISA and positive PCR assay results in multiple Pteropus spp suggest the virus or related viruses are widespread across this range.

Nipah virus spillover from fruit bats to pigs occurs through contact with contaminated fruit and bat secretions (urine, saliva, uterine fluids). Within intensive swine facilities, nearly all pigs become infected through rapid respiratory and oral transmission, with spread between farms facilitated by poor biosecurity and animal movement. Infection in humans occurs via direct contact with infected pig secretions and excretions (occupational exposure in farm and abattoir workers), consumption of contaminated date palm sap or fruit in South Asia, and human-to-human transmission in Bangladesh and India. Horse-to-human transmission occurred in the 2014 Philippines outbreak through contact during slaughter and consumption of infected meat.

Nipah virus infects susceptible hosts through the naso-oral route, establishing initial replication in the nasal cavity and nasopharynx before systemic dissemination. The virus infects vascular endothelial cells, triggering generalized vasculitis that manifests as organ-specific disease (respiratory and neurological).

Clinical signs progress as predominantly respiratory and/or neurological. In pigs, incubation is 6–14 days with high morbidity rates but low mortality. Infected animals, including subclinically infected animals, shed virus in respiratory secretions and excretions (urine, blood). Pathological findings in swine include pulmonary edema, dilated lymphatic vessels, airway exudates, and pleural/pericardial fluid accumulation. In contrast to pigs, horses and cats develop high mortality rates.

During the Malaysian outbreak, Nipah virus infection primarily manifested as a febrile respiratory disease characterized by a severe, distinctive cough audible from up to 1 mile away—locally termed "barking pig syndrome" or "one-mile cough." Encephalitis also occurred, particularly in breeding animals (sows and boars); however, respiratory disease predominated. Overall mortality rates in affected facilities were poorly documented but likely did not exceed 5% across all age groups. A high incidence of subclinical infection was reported, with infected animals shedding virus without obvious clinical signs.

Clinical expression varies from subclinical infection to severe respiratory and neurological disease. Clinical signs in infected pigs include pyrexia, barking cough with labored or open-mouth breathing, pulmonary congestion and edema, terminal nasal discharge (white or blood-stained froth), neurological manifestations (behavioral changes, head pressing, nystagmus, champing, uncoordinated gait, spasms, myoclonus, hindlimb paresis), and early abortion. The combination of high morbidity rate, low mortality, prolonged clinical illness, and substantial subclinical shedding makes infected pig populations a sustained source of occupational exposure for farm and abattoir workers.

In humans, Nipah virus causes an influenza-like illness, encephalitis, and/or respiratory syndrome.

Differential diagnoses for Nipah virus infection include the following:

• Neurological signs:

  • Streptococcus suis infection

  • Glässer disease (Glaesserella parasuis)

  • edema disease

  • water intoxication

  • enteroviruses

  • West Nile virus infection

  • Japanese encephalitis

  • Aujeszky disease

  • rabies lyssavirus infection

• Respiratory and pyrexic signs:

  • influenza

  • classical swine fever (chronic form)

  • porcine reproductive and respiratory syndrome

  • postweaning multisystemic wasting syndrome (in weaned pigs)

Diagnosis is based on clinical suspicion in endemic settings and confirmed by molecular or serological testing performed under high-containment laboratory conditions. Samples of choice include EDTA blood, serum, and oral swabs. Primary diagnostic tests include real-time PCR assay for viral nucleic acid detection and ELISA for antibody detection. Virus isolation and serum neutralization testing require biosafety level 4 containment; pseudotype neutralization assays provide an alternative for antibody detection.

Postmortem examination—requiring full personal protective equipment (PPE)—allows immunohistochemical (IHC) staining of formalin-fixed tissues with specific antibodies to detect viral antigen. Useful tissues include urine, brain, lung, kidney, and spleen, collected with appropriate biosafety precautions. In pregnant animals or cases of abortion, uterus, placenta, and fetal tissues should be included.

• No treatment available

Supportive care is the only option for Nipah virus infection in swine, because no effective treatment exists. Treatment of swine with Nipah virus infection was not attempted during the 1998–1999 Malaysian emergency.

In the Malaysian outbreak, human patients required intensive care with ventilation support to manage the encephalitis. Ribavirin was administered to some patients; however, subsequent studies in laboratory animals suggest that it is ineffective (2). A monoclonal antibody has been used therapeutically in subsequent outbreaks.

Control of the Malaysian Nipah virus outbreak required strict quarantine procedures and culling of all swine from affected facilities. An active surveillance and slaughter program successfully eliminated Nipah virus from the national commercial swine population, which has remained free of infection since 1999. The persistence of NiV in bat reservoirs across a wide geographic range underscores the critical importance of robust disease surveillance and biosecurity practices to enable early detection and containment should reintroduction occur.

Nipah virus has a lipid envelope, making it susceptible to desiccation and temperature fluctuations. Survival in the environment ranges from several hours to several days, depending on conditions; for disease control purposes, survival of > 4 days represents the presumed maximum under optimal conditions (3). Effective disinfectants include soap and detergents; broad-spectrum, oxidizing disinfectants; hypochlorites; iodophors/iodine; biguanides (eg, chlorhexidine); and quaternary ammonium compounds.

A subunit vaccine for horses is based on Hendra virus soluble G protein and has proven effective in preventing Hendra virus infection in horses (4). The vaccine has demonstrated efficacy against NiV in experimental challenge studies in nonhuman primates and swine (5, 6). Several vaccines for swine are in development.

In humans, early preclinical and Phase 1 clinical data for an mRNA vaccine suggest cross-protection against both NiV and Hendra virus infections, consistent with the cross-protective responses observed with other vaccine platforms (7).

During the Malaysian outbreak, transmission of NiV from infected pigs to humans occurred almost exclusively in occupational settings, with close contact with infected swine being the primary route of infection. Similar patterns have emerged with Hendra virus in Australia, where sporadic clusters in horses led to subsequent human cases with serious disease outcomes. These experiences underscore the critical importance of appropriate PPE during veterinary clinical examinations and postmortem procedures when Hendra or Nipah virus infection is suspected.

When NiV infection is suspected, minimize contact with suspect animals, isolate cases, and implement biocontainment protocols (including farm movement controls) until advised by government veterinary authorities. Use PPE, including gloves, disposable coveralls, rubber boots, goggles or safety glasses, and P3 or N95 respirators for any close contact with infected pigs. Consult relevant state or territory health authorities immediately, noting that Nipah virus is a WOAH-listed disease with international reporting obligations. NiV is also a WHO priority pathogen of concern.

• WHO Health Emergency Dashboard.

• McLean RK, Pedrera M, Thakur N, et al. Nipah virus vaccines evaluated in pigs as a "One Health" approach to protect public health. NPJ Vaccines. 2025;10(1):163.

1. Nipah virus. Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Terrestrial Manual) of the World Organisation for Animal Health. 2022. https://www.woah.org/en/disease/nipah-virus/

2. Goh KJ, Tan CT, Chew NK, et al. Clinical features of Nipah virus encephalitis among pig farmers in Malaysia. N Engl J Med. 2000;342(17):1229-1235. doi:10.1056/NEJM200004273421701

3. Fogarty R, Halpin K, Hyatt AD, Daszak P, Mungall BA. Henipavirus susceptibility to environmental variables. Virus Res. 2008;132(1-2):140-144. doi:10.1016/j.virusres.2007.11.010

4. Middleton D, Pallister J, Klein R, et al. Hendra virus vaccine, a one health approach to protecting horse, human, and environmental health. Emerg Infect Dis. 2014;20(3):372. doi:10.3201/eid2003.131159

5. Pickering BS, Hardham JM, Smith G, et al. Protection against henipaviruses in swine requires both, cell-mediated and humoral immune response. Vaccine. 2016;34(40):4777-4786. doi:10.1016/j.vaccine.2016.08.028

6. Satterfield BA, Mire CE, Geisbert TW. Overview of experimental vaccines and antiviral therapeutics for Henipavirus infection. In: Freiberg AN, Rockx B, eds. Nipah Virus: Methods and Protocols. Humana Press, Inc; 2023:1-22. Methods in Molecular Biology; vol 2682. doi:10.1007/978-1-0716-3283-3_1

7. Ploquin A, Mason RD, Holman LA, et al. A structure-based mRNA vaccine for Nipah virus in healthy adults: a phase 1 trial. Nat Med. 2026. doi:10.1038/s41591-026-04265-1