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Arenavirus and West Nile Virus in Solid Organ Transplantation

2013; Elsevier BV; Volume: 13; Linguagem: Inglês

10.1111/ajt.12128

1600-6143

Neeraj Singh, Marilyn E. Levi,

Viral Infections and Vectors

Over the past several years, emerging pathogens such as arenaviruses and West Nile virus (WNV) have been identified as sources of both donor-derived and posttransplant infections. Thus far, data on these infections has been published primarily in case reports. Herein, we present discussions of WNV and arenavirus infections in solid organ transplant recipients. Arenaviruses are single-stranded enveloped RNA viruses associated with rodent-transmitted diseases of humans. Their family name is derived from the sandy appearance on electron microscopy (Latin arena, or “sand”). They are divided into two groups: The Old World complex (family Muridae, subfamily Murinae) which includes lymphocytic chorimeningitis virus (LCMV), Lassa virus and other closely related viruses and the New World complex (Family Muridae, subfamily Sigmodontinae) which includes Junin, Machupo, Guanarito, Sabia and other closely related New World viruses, commonly referred to as South American hemorrhagic fever viruses due to similar clinical presentation. Arenaviruses are maintained in nature through chronic asymptomatic infection in rodents. LCMV differs from other arenaviruses as common house mice (Mus domesticus and Mus musculus), a rodent with global distribution as opposed to geographically restricted field mice are its natural reservoir 1. Pet hamsters and guinea pigs are not the natural reservoirs for LCMV but pet and laboratory rodents can become infected if they come in contact with house mice (e.g. in a breeding facility, pet store or home). Among rodents, transmission may occur vertically or horizontally or both depending on the specific virus. The virus exhibits high species specificity with a given rodent species providing reservoir to a specific virus. The geographic distribution of the respective rodent species in turn determines the regional distribution of the disease. In United States, the seroprevalence of LCMV in rodents is quite variable with reported rates of 9% in Baltimore, Maryland 2, to up to 95% in a case study from Michigan 3. Humans are primarily infected by inhaling infectious aerosolized particles of rodent secretions (saliva, urine or droppings). In addition, contact with infectious rodent excreta, ingestion of contaminated food, rodent consumption and rodent bite have all been known to cause infection in humans. Lower socioeconomic status, substandard housing and agricultural activities have been associated with rodent infestation and a higher risk of infection 4. Transplant recipients may become infected with arenavirus if they are exposed to conditions conducive to contact with wild rodents or infected pet rodents. Isolated reported cases of LCMV infections have been reported in laboratory personnel after contact with infected hamsters or infected rodent cell lines 5, 6. Person-to-person transmission via aerosol spread and contact with infected fluid can occur in Lassa fever and some other South American viral hemorrhagic fevers. Transmission by sex and breast feeding can also occur in Lassa fever and potentially other viral hemorrhagic fevers even during recovery from acute illness. In case of LCMV, person-to-person transmission has occurred only through maternal–fetal transmission 7, 8 and organ transplantation 9. Anti-LCMV IgG antibody prevalence in healthy human populations ranging from 0.3% to 4.7% has been reported from different parts of the world 1, 10-13. The seroprevalence of latent Lassa virus infection in West African population is reported anywhere from 12% to 50% 14, 15. The seroprevalence of subclinical Junin virus infection in two rural populations in Argentina was found to be 1.9% and 4.4%, respectively 16. Table 1 describes the geographic distribution, incubation period, peak season and clinical features of arenavirus related diseases 8, 17, 18. LCMV infection in immunocompetent patients is asymptomatic or mild in most patients. When symptomatic, illness is often subtle with self-limiting symptoms of fever, malaise, headache, photophobia, listlessness, myalgia, confusion, memory deficits and abdominal pain. Some cases may progress to meningitis, encephalitis and other central nervous system manifestations 19 but the overall case fatality is 90%). To date, five clusters of transmission of LCMV and an LCMV-like arenavirus via organ transplantation 9, 20-22 have been described (Table 2). Fourteen of the 17 recipients died of multisystem organ failure, with LCMV-associated hepatitis as a prominent feature. A common donor was recognized in each cluster. In the 2005 cluster, the donor had contact in her home with a pet hamster infected with an LCMV strain identical to that detected in the organ recipients. It is unclear if LCMV infection can happen in transplant recipients due to reactivation of latent virus posttransplant. LCMV is often underrecognized and underdiagnosed because the clinical characteristics of LCMV meningitis are similar to those of other viral meningitis. In addition, there is lack of awareness of the virus among physicians, and the diagnostic assays are not commercially available. Lassa fever is mild or asymptomatic in most of the infected individuals 15 but in a small proportion, it is severe and may progress to multisystem organ failure with shock, coma or death. The presentation of South American viral hemorrhagic fevers is similar to Lassa fever but with more frequent hemorrhagic and neurological complications. In contrast to South American hemorrhagic fevers, hepatitis is more frequent and severe in Lassa fever. The case fatality in Lassa fever and South American hemorrhagic fevers can be as high as 25–30% 23-25. There have no reported cases of Lassa fever and South American hemorrhagic fever in organ transplant recipients, but cases are likely missed and not reported. Diagnosis of LCMV should be strongly considered in organ transplant recipients presenting with aseptic meningitis and encephalitis, especially with unexplained fever, hepatitis or multisystem organ failure. Lassa fever and South American Hemorrhagic fever should be considered in travelers to the endemic areas with compatible clinical picture and potential exposure to rodents or a person with viral hemorrhagic fever. Possible alternate diagnosis including Yellow fever, dengue fever, malaria, Crimean-Congo hemorrhagic fever, Rift valley fever, Ebola and Marburg viral fevers, viral hepatitis and typhoid fever must also be considered. The laboratory diagnosis can be made by detection of anti-virus immunoglobulin M (IgM) and/or a fourfold rise in IgG in serum or CSF samples using enzyme-linked immunosorbent assay (ELISA) (preferred) and/or immunofluorescence assay (IFA; Refs. 26, 27). Reverse transcriptase polymerase chain reaction (PCR) can detect viral RNA rapidly and help identify strains but poses limitations due to natural genetic diversity of the virus and currently, remains as a research tool. Viral culture using cell lines can be confirmatory but is time consuming. Immunohistochemical staining of viral antigens in tissue specimens can be helpful in case of negative serological assays. Virus can be isolated from blood, CSF or throat swabs. “These tests are available at state and public health reference laboratories, such as the US Centers for Disease Control and Prevention (CDC), although they may also be available in few commercial laboratories.” There is currently no evidence of benefit of screening of potential organ donors for LCMV infection. Deceased donors may be asymptomatic at the time of death prior to donation. If the potential donor has died of aseptic meningitis or encephalitis of unknown cause, risks and benefits to potential transplant recipients in offering and accepting organs from such donors should be carefully considered. Supportive care with meticulous fluid balance and electrolyte management is the mainstay of therapy in arenavirus infection. One surviving transplant patient in the 2005 cluster of donor-transmitted cases was treated with ribavirin and reduction of immunosuppressive therapy 9. However, in 2011 cluster, 2 of the 4 infected recipients survived without receiving treatment with ribavirin 22. Although, ribavirin has in vitro activity against LCMV 7, it is not FDA approved for this indication. The surviving patient with LCMV infection in the 2005 cluster was treated with intravenous ribavirin (administered at a loading dose of 30-mg/kg, followed by 16-mg/kg every 6 h for 4 days; followed by 8-mg/kg every 8 h). After the patient's clinical condition stabilized, ribavirin was changed to oral route (400 mg every morning and 600 mg every evening). The drug was discontinued on day 37 when the virus became undetectable. Intravenous ribavirin is not available in United States for routine use, but it may be available through an Emergency Investigational New Drug (EIND) application as an investigational agent for patients with serious viral infections. IV ribavirin has also been used successfully to treat other arenavirus hemorrhagic fevers including Lassa fever 28, Argentine hemorrhagic fever 29, 30 and Bolivian hemorrhagic fever 31. It has been found to reduce the risk of developing oliguria in patients with confirmed hemorrhagic fever with renal syndrome 32. Convalescent plasma has been reported to reduce case fatality in Argentinine hemorrhagic fever when given within first 8 days of illness 33. Special care must be taken to prevent person-to-person spread of Lassa fever and potentially South American viral hemorrhagic fever with both airborne and contact isolation of patients 34, 35. Household members should avoid close physical contact with infected person and their body fluids. Nursing mothers with viral hemorrhagic fever should avoid breast feeding even 2–3 months into recovery. Oral ribavirin may be considered for postexposure prophylaxis of Lassa fever in health care workers and close contacts that have been exposed to blood or body fluids of an infected person or animal. One of the recommended dose is: 35 mg/kg loading dose, maximum 2.5 g followed by 15 mg/kg, maximum 1 g three times a day for 10 days 36. If the exposed person develops manifestations of hemorrhagic fever, they should be immediately converted to intravenous ribavirin 36. The drug is however poorly tolerated 37, is teratogenic and cause hemolytic anemia. The vaccine against Lassa virus remains in the development phase 38, but is critically needed. A live attenuated vaccine against Junin virus (Argentine viral hemorrhagic fever) was found effective in a prospective, randomized, double-blind, placebo-controlled trial 39 and has drastically reduced the incidence of disease in Argentina 40. The vaccine has also been found effective against Machupo virus (Bolivian hemorrhagic fever)41. The vaccine is however not approved or available in the United States. Since live viral vaccines are not generally recommended posttransplant, solid organ transplant candidates in high risk areas may be vaccinated before transplant. However, its efficacy is unknown in transplant recipients. In experimental mice models, an immunosuppressive drug, sirolimus has been shown to enhance virus-specific CD8 T cells following acute LCMV infection as well as after immunization 42. The strategy can be potentially utilized to help in the future development of LCMV-specific vaccine. Rodent avoidance, control and elimination, safe disposal of rodent nests and droppings and rodent contaminated foods, proper hand washing and cleaning of rodent infested areas are important interventions for preventing spread of arenaviral infections. Future research should focus on making the molecular and serological assays for diagnosis of arenavirus infections commercially available, development of better and safer drugs and development of effective vaccines against LCMV and Lassa fever virus. Transplant recipients should avoid contact with house mice, and wild and pet rodents by taking adequate measures as outlined by CDC (http://www.cdc.gov/ncidod/dvrd/spb/mnpages/dispages/lcmv.htm). In general, patients should be advised to keep away from wild rodents, avoid close contact with pet rodents, observe proper hand hygiene after handling pet rodents, to ask another family member or friend to clean the cage and care for the pet, and maintaining adequate environmental cleaning. Although the risk of LCMV infection from pet rodents (i.e. mice, hamsters or guinea pigs) is generally low, proper precautions should be observed (Grade III). LCMV infection should be strongly considered in transplant recipients presenting with unexplained fever, hepatitis, encephalitis or multisystem organ failure with prompt reporting to CDC and initiation of testing (Grade III). Serum, CSF and tissue samples should be obtained for viral culture, serology and immunohistochemical staining. Universal organ donor screening for arenavirus is not recommended due to lack of evidence for its utility and lack of readily available diagnostic tests (Grade III). Donors with unexplained meningoencephalitis should be assessed for risk factors for arenaviruses, and organs from donors with suspected or proven arenavirus infection should not be used (Grade III). Intravenous ribavirin is the drug of choice for Lassa fever (Grade I), and should be considered for the treatment of Argentine and Bolivian hemorrhagic fever (Grade II-3). There is insufficient evidence for its efficacy and use in LCMV and other South American hemorrhagic fevers. WNV is a mosquito borne single-stranded RNA arbovirus that belongs to the Flaviviridae family, which also includes St. Louis Encephalitis (SLEV), Japanese B encephalitis virus, Dengue, Yellow Fever, Murray Valley encephalitis and Kunjin viruses. In 1937, the first human case of WNV was reported in Uganda 43. Since then, WNV outbreaks have occurred in Africa, Asia, Europe and the Middle East where the virus is endemic. In 1999, the first outbreak of WNV in the Western hemisphere occurred in New York City 43. Over the ensuing years, WNV has spread westward over the continental United States, northward to Canada and southward to the Caribbean islands and Latin America 44. WNV and SLEV are the only flaviviridae endemic in the United States 45 with WNV now the most common, and reported in 48 states. Infected mosquitoes, most commonly of the Culex genera, acquire WNV by feeding on infected birds who serve as the primary amplifying hosts of WNV 46. As the seasons progress from summer to fall, a bird-mosquito enzootic cycle develops with increasing viral amplification and infectivity of “bridge vector” mosquitoes 47, 48. The net result is the successful transmission of WNV to incidental hosts, including humans. The incidence and geographic location of WNV varies yearly depending on environmental conditions such as the presence of Culex spp. mosquitoes and their ability to grow in number and have access to bird vectors 80. The incubation period for WNV is between three and 14 days (average of 6 days). 80% of immunocompetent individuals remain asymptomatic 46 while 20% commonly have mild symptoms such as fever, myalgias, malaise, nausea, vomiting, diarrhea and transient rash. Only one of 140 symptomatic patients develops neuroinvasive disease including meningitis, encephalitis or meningoencephalitis 46, a poliomyelitis-like flaccid paralysis, Parkinsonian cogwheel rigidity and profound cognitive impairment. Following acute infection, 50% of patients have residual difficulties with memory loss, fatigue, ambulation and depression 49. Groups at high risk for the development of WNV associated neuroinvasive disease are immunosuppressed individuals such as solid organ transplant recipients 51, 52 and recipients of chemotherapy 52 such as rituximab and B cell depleting agents 54-56, inferring the importance of humoral mediated immunity in controlling WNV infection. In 2002 and 2003, WNV epidemics in the United States and Canada identified nonmosquito borne transmission of WNV through solid organ donation, blood transfusions 46, breast milk and placental transmission during pregnancy 49. Between 2002 and 2009, a total of five cases of solid organ donor transmission of WNV have been identified (Table 3). In these cases, the mean duration of incubation period was 13.5 days (range 7–17 days; Ref. 50). While donor transmission of WNV has been of major concern, a majority of reports of WNV infection in transplant recipients are related to infected mosquito bites 51, 52. A seroprevalence study suggested that while less than 1% of immunocompetent individuals infected with WNV develop neuroinvasive disease, the incidence may be as high as 40% in solid organ recipients 57. However, this was not confirmed in another seroprevalence study 58. The diagnosis of WNV depends on a high index of suspicion and laboratory testing 46. The clinician should consider WNV in the differential diagnosis of a patient presenting with fevers, altered mental status, lower extremity paralysis, Parkinsonian cogwheel rigidity or other neurologic symptoms during the “typical WNV season”, defined as May 1 to November 30 59. To assist the clinician, local and state health departments and the CDC via Arbonet reporting (http://www.cdc.gov/ncidod/dvbid/westnile/index.htm) websites report cases of WNV infections in mosquito, bird and/or humans in specific locales. Laboratory studies for diagnosis include serum and CSF WNV IgM and IgG antibodies and viral PCR testing. Interpretation of the results is facilitated by review of the timeline of WNV infection (Figure 1). In most cases, WNV infected mosquito bites are followed by an average incubation period of 6 days. After the incubation period, asymptomatic viremia lasting for 5–14 days can be identified by serum and CSF WNV PCR testing. Longer periods of viremia may occur, especially in immunocompromised patients 59. Patients with defective humoral immunity, such as those treated with rituximab, may be unable to produce WNV IgM or IgG antibodies 54-56 and have a persistent WNV viremia. Therefore, serum and CSF PCR testing may be the primary means of diagnosing WNV infection in this population 54. Commonly, decline in WNV viremia is followed by the production of IgM antibodies. IgM is produced within 8 days after the initial WNV exposure 58 and an average of 3.9 days after the onset of viremia. Serum WNV IgG is then produced within the following 3.4 days in one study 61. Serum IgM may persist for over 500 days and therefore may not be indicative of acute infection 60. Serum IgG antibody confers lifelong protection against reinfection. Given the prolonged positive serologies, acute and convalescent serologies for IgM and IgG may be helpful in confirming the diagnosis of acute WNV infection. For the diagnosis of WNV neuroinvasive disease, CSF should be obtained for cell count with differential, protein, glucose, WNV IgM/IgG and PCR. Studies of solid organ transplant recipients with naturally occurring WNV disease showed CSF pleocytosis ranging from 5 to 540 cells with half of cases showing a lymphocytic pleiocytosis and the other half demonstrating a neutrophilic predominance. CSF protein levels have ranged between 41 and 142 with primarily normal glucose levels 51, 52. The presence of WNV IgM in the CSF is pathognomonic for central nervous system disease, as the IgM antibody does not cross the blood–brain barrier. A complicating factor in the interpretation of WNV serologies is the cross reactivity with other flaviviridae, such as St. Louis and Japanese Encephalitis and Dengue viruses. Furthermore, the Yellow Fever vaccine may also result in false positive serologies for WNV. To assist in differentiation, the CDC utilizes IgM-ELISA microsphere assays that are specific to the different flaviviridae. For specific confirmation, plaque reduction neutralization testing (PRNT) may be obtained through the CDC 76, although results are not available prior to organ harvesting. The primary treatment of WNV is supportive care such as hydration, hospitalization and use of ventilatory support, if needed. Temporary reduction in immunosuppression should be considered in order to allow for restoration of natural immunity to WNV. There are no solid data to support use of specific antivirals for treatment, but several agents have shown encouraging results 70. Intravenous immuno-globulin (IVIG) containing WNV specific antibodies has shown promise in the treatment of acute infection 61-64. WNV appears to have greater susceptibility to humoral mediated, as opposed to cell mediated immunity 55. Passive transfer of monoclonal or polyclonal virus-specific antibodies has been shown to play a key role in both prophylaxis and treatment 65, 66. This was shown in a mouse model where WNV infection was lessened or completely aborted in a dose dependent manner with transfer of passive antibodies 67. The presence of adequate WNV antibodies in the IVIG product initially required use of high titer WNV-specific immunoglobulin (Omr-IgG-am®, Omrix Biopharmaceutical Ltd, Kiryat-Ono, Israel) from the Middle East, where there are areas of high endemicity for WNV 62, 67 and was granted orphan drug status by the FDA in 2007 68. However, the seroprevalence of WNV in the United States has increased, resulting in the presence of high titer WNV antibodies in US plasma derived products 69 although the concentrations may vary from region to region depending on WNV endemicity. Successful use of U.S. derived IVIG for the treatment of acute WNV infection has been reported, with two doses of 0.4 g/kg administered four days apart in one report 62 and 1000 mg/kg followed by 500 mg/kg in a second report 63. Early administration of IVIG at the time of viremia may improve the outcome of WNV infection 62, 65, 66. A delay in dosing has been shown to decrease survival benefit 70, so that empiric early administration may be prudent if there is a high level of suspicion for the presence of WNV infection, prior to obtaining results of studies. A small randomized controlled trial of Omr-IgG-am® versus standard IVIG failed to show a clinical benefit in adults with symptomatic disease 76. Further studies with higher doses or prophylaxis of donor infections will need to be performed utilizing either the United States derived IVIG or the Omr-IgG-am product. Interferons restrict viral replication by activation of cytotoxic T cell responses 71 with animal studies suggesting improved survival with WNV infection 72. Reports of successful treatments with interferon-α 2b 3 million units daily × 14 days have been published 70, 72, 73. However, due to concern that interferon may be associated with organ rejection, its use in transplant recipients has not been studied. Due to the significant false positive rate and potential organ loss, WNV NAT screening for organ donor screening is not routinely performed unless specifically requested. In comparison, WNV NAT screening is performed routinely for blood donation with serologic confirmation of positive results. Patients with positive results are asked to hold on further donations for 120 days. In the posttransplant population, prevention of WNV infection focuses on avoidance of mosquito bites, specifically with the use of long sleeves and long pants, and application of topical insecticides on exposed skin, such as DEET, picardin, oil of lemon eucalyptus or IR3535 in concentrations between 10% and 50%. As mosquitoes are most active in the evenings, they should be advised to avoid outdoor activities from dusk to dawn whenever possible. A brochure specifically designed for transplant patients can be downloaded through the CDC website 82. Human to human transmission of WNV does not occur through contact, respiratory or droplet exposure. Patients with WNV who are hospitalized require standard universal infection control precautions. The authors thank Drs. Steven Geier and Michael Sauer for their valuable input with regard to WNV NAT testing. The authors of this manuscript have no conflicts of interest to disclose as described by the American Journal of Transplantation.