Transmission of Lymphocytic Choriomeningitis Virus by Organ Transplantation
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《新英格兰医药杂志》
ABSTRACT
Background In December 2003 and April 2005, signs and symptoms suggestive of infection developed in two groups of recipients of solid-organ transplants. Each cluster was investigated because diagnostic evaluations were unrevealing, and in each a common donor was recognized.
Methods We examined clinical specimens from the two donors and eight recipients, using viral culture, electron microscopy, serologic testing, molecular analysis, and histopathological examination with immunohistochemical staining to identify a cause. Epidemiologic investigations, including interviews, environmental assessments, and medical-record reviews, were performed to characterize clinical courses and to determine the cause of the illnesses.
Results Laboratory testing revealed lymphocytic choriomeningitis virus (LCMV) in all the recipients, with a single, unique strain of LCMV identified in each cluster. In both investigations, LCMV could not be detected in the organ donor. In the 2005 cluster, the donor had had contact in her home with a pet hamster infected with an LCMV strain identical to that detected in the organ recipients; no source of LCMV infection was found in the 2003 cluster. The transplant recipients had abdominal pain, altered mental status, thrombocytopenia, elevated aminotransferase levels, coagulopathy, graft dysfunction, and either fever or leukocytosis within three weeks after transplantation. Diarrhea, peri-incisional rash, renal failure, and seizures were variably present. Seven of the eight recipients died, 9 to 76 days after transplantation. One recipient, who received ribavirin and reduced levels of immunosuppressive therapy, survived.
Conclusions We document two clusters of LCMV infection transmitted through organ transplantation.
Lymphocytic choriomeningitis virus (LCMV) is a rodent-borne, Old World arenavirus that has been reported to cause asymptomatic or mild, self-limited illness in otherwise healthy humans. It is a known cause of aseptic meningitis, but fatal infection is rare.1,2,3,4 Transmission of infection from a woman to a fetus can result in hydrocephalus, chorioretinitis, or microcephaly.5,6,7,8 Outside of vertical transmission during pregnancy, human-to-human transmission of LCMV has not been described.9 We describe two clusters of unexplained clinical syndromes in transplant recipients and the subsequent investigations to identify donor-transmitted infection as the cause of illness.
Methods
Case Reports
The 2003 Cluster
In December 2003, unexplained febrile illnesses developed in four recipients of solid organs from a common donor (Figure 1; additional information on clinical symptoms and laboratory findings for each recipient is listed in Table 1 of the Supplementary Appendix, available with the full text of this article at www.nejm.org).10 Kidney Recipient 1 was a 46-year-old man with diabetes. Diarrhea and mild, diffuse abdominal pain developed on post-transplantation day 5, but his condition was stable and he was discharged home the following day. He was readmitted on post-transplantation day 23 with fever, persistent watery diarrhea, and worsening abdominal pain. Laboratory studies revealed leukopenia with elevated aminotransferase and creatinine levels. Ganciclovir therapy was initiated because of concern about possible cytomegalovirus infection. Tacrolimus and mycophenolate mofetil were discontinued. Examination of kidney-, liver-, and bone marrow–biopsy specimens did not reveal an infectious cause. On day 40 after transplantation, seizures and polymyoclonus developed. The patient reported blurred vision, and chorioretinitis was noted on ophthalmologic examination. Cerebrospinal fluid studies revealed a markedly elevated level of protein (720 mg per deciliter), a normal glucose level (147 mg per deciliter ), and 4 white cells and 3 red cells per cubic centimeter. Polymerase-chain-reaction (PCR) tests for cytomegalovirus, herpes simplex virus, varicella–zoster virus, Epstein–Barr virus, human herpesvirus 6, enterovirus, adenovirus, Mycobacterium tuberculosis, and Borrelia burgdorferi were negative. Magnetic resonance imaging of the brain revealed bilateral, hemispheric, subdural fluid collections and diffuse dural thickening. Examination of a specimen obtained by dural biopsy on post-transplantation day 47 showed fibrosis; cidofovir and intravenous immune globulin were initiated for the suspected presence of an unknown viral pathogen. The patient's condition continued to deteriorate, and he died on post-transplantation day 53. An autopsy revealed bronchopneumonia and hepatic congestion without inflammation.
Figure 1. Clinical Course of Lymphocytic Choriomeningitis Virus (LCMV) Infection in the 2003 Cluster.
MMF denotes mycophenolate mofetil, TMP-SMX trimethoprim–sulfamethoxazole, IVIG intravenous immune globulin, and X death. Immunosuppressive agents are shown in red, and antimicrobial agents in blue.
Kidney Recipient 2 was a 56-year-old man with glomerulonephritis. He was discharged home on post-transplantation day 5 but was readmitted on post-transplantation day 22 with fever, leukopenia, and peri-incisional erythema and tenderness. A skin-biopsy specimen obtained at the wound edge revealed basal-cell vacuolation suggestive of viral infection; no viral inclusions were seen, and immunohistochemical stains were negative for cytomegalovirus and adenovirus. Tacrolimus and mycophenolate mofetil were discontinued. Altered mental status and seizures with myoclonus developed on post-transplantation day 31. Cerebrospinal fluid studies revealed a markedly elevated protein level (620 mg per deciliter), a low glucose level (57 mg per deciliter ), and 12 white cells (48 percent lymphocytes and 22 percent monocytes) and 6 red cells per cubic centimeter. PCR testing for the same infectious agents as in the case of Kidney Recipient 1 was unrevealing. Magnetic resonance imaging showed subdural fluid collections with diffuse dural enhancement. The patient's condition continued to deteriorate, with photophobia, nuchal rigidity, diarrhea, thrombocytopenia, diffuse erythroderma, respiratory failure, and atrial fibrillation, despite empirical administration of cidofovir and intravenous immune globulin. He died on post-transplantation day 76; an autopsy revealed meningoencephalitis and acute bronchopneumonia.
The liver recipient was a 40-year-old woman with alcoholic cirrhosis whose early postoperative course was marked by fever, lethargy, and hypotension. Markedly elevated aminotransferase levels, leukopenia, and bilateral pulmonary infiltrates with respiratory failure developed. Liver biopsy revealed focal centrilobular coagulative necrosis. On post-transplantation day 14, a peri-incisional petechial rash was noted; skin biopsy revealed chronic inflammation and hemorrhage. She died on day 17 after transplantation. An autopsy revealed extensive hepatic necrosis without evidence of infection.
A common source of infection in these recipients was suspected, all of whom underwent transplantation at the same facility. Therefore, the Department of Public Health of the Wisconsin Department of Health and Family Services and the organ-procurement organization (OPO) coordinating the transplantations were notified for assistance.
The lung recipient, who had undergone transplantation at a different facility, was a 46-year-old man with chronic obstructive pulmonary disease who had been receiving prednisolone daily for three months before transplantation. Within four days after transplantation, hypotension, bilateral pulmonary infiltrates, and leukocytosis developed; broad-spectrum antimicrobial agents were administered. Fever (temperature, 38.3°C), hypoxia, and refractory hypotension ensued. He died on day 9 after transplantation. An autopsy revealed diffuse alveolar damage without evidence of rejection.
The donor was a 51-year-old man who had been found unresponsive, with apparent head trauma. Computed tomography (CT) of the brain revealed a large, right-sided subdural hematoma with a midline shift. There was no improvement in his neurologic status, and he was declared brain-dead on hospital day 2. He received no blood products and was afebrile throughout his hospitalization. Donor-eligibility screening and testing detected no infection precluding organ or tissue donation. His liver, lungs, kidneys, and multiple musculoskeletal and vascular tissues were recovered for transplantation. Tissue specimens from the donor and recipients were submitted to the Centers for Disease Control and Prevention (CDC) for additional testing (Table 1).
Table 1. Summary of Laboratory Evaluations for Lymphocytic Choriomeningitis Virus Infection in the 2003 Cluster.
The 2005 Cluster
Kidney Recipient A was a 48-year-old man admitted to a hospital in Rhode Island in late April 2005, 17 days after undergoing cadaveric renal transplantation (Figure 2). He had been discharged home on post-transplantation day 7 with a creatinine level of 1.6 mg per deciliter (141 μmol per liter). At the time of readmission, he had had right-lower-quadrant pain in the area of the allograft for four or five days, as well as nausea, anorexia, diarrhea, fever, and chills. He was febrile (temperature, 38.7°C), and there was tender erythema over the area of the allograft, without incisional dehiscence or drainage. His creatinine level was 2.6 mg per deciliter (230 μmol per liter), and proteinuria, hematuria, a prolonged prothrombin time, and slightly elevated aminotransferase levels were present. Mycophenolate mofetil was discontinued, and administration of broad-spectrum antimicrobial agents was initiated. CT and ultrasonography of the abdomen and pelvis were unrevealing. Routine cultures of urine, blood, and stool were negative, as were studies of the stool for leukocytes, ova, parasites, Clostridium difficile, giardia, cryptosporidium, Yersinia enterocolitica, and rotavirus. Tacrolimus was discontinued because of concern about the worsening infection of uncertain cause. His temperature rose to 40.4°C, and he had copious diarrhea, dyspnea, and tender erythema extending from the area over the allograft to the right flank. Examination of biopsy specimens of the colon and kidney revealed no inflammation or viral inclusions.
Figure 2. Clinical Course of Lymphocytic Choriomeningitis Virus (LCMV) Infection in the 2005 Cluster.
MMF denotes mycophenolate mofetil, TMP-SMX trimethoprim–sulfamethoxazole, CVVH continuous venous–venous hyperfiltration, and X death. Immunosuppressive agents are shown in red, and antimicrobial agents in blue.
Kidney Recipient B was a 54-year-old man who was admitted to the same hospital with fever. He had undergone cadaveric renal transplantation 17 days previously and had been discharged home on post-transplantation day 8 with a creatinine level of 3.3 mg per deciliter (292 μmol per liter). On post-transplantation day 14, mycophenolate mofetil was discontinued because of diarrhea. Fever and pain developed in the right lower quadrant over the area of the allograft, and he was readmitted for evaluation. He was lethargic and febrile (temperature, 38.4°C) and had tender erythema overlying the allograft, without incisional drainage. His creatinine level was 4.0 mg per deciliter (354 μmol per liter), with a platelet count of 113,000 per cubic millimeter and an alanine aminotransferase level of 298 IU per liter. Routine cultures of urine, blood, and stool were negative, and ultrasonography and CT of the abdomen and pelvis were unrevealing. Fever, diarrhea, and pain persisted, despite empirical use of broad-spectrum antimicrobial agents. A percutaneous liver biopsy was performed on post-transplantation day 23 because of worsening hepatitis and leukocytosis. Multiple foci of hepatocellular necrosis without inflammation or viral inclusions were noted. Later that day, he had a cardiac arrest and died. An autopsy revealed coronary artery disease and diffuse cerebral edema without meningitis or encephalitis.
A review of the records of the hospital's transplantation center revealed that the kidney recipients shared a common donor who had also been hospitalized there. The donor was a 45-year-old woman with hypertension who had presented to the emergency department with a five-day history of right-sided headache and acute left-sided weakness. She was alert and afebrile and had left-sided hemiparesis. CT of the brain revealed an infarct in the distribution of the right middle cerebral artery, and tissue plasminogen activator was administered. Throughout her hospitalization, she was afebrile and received no blood products or antimicrobial agents. She had a normal white-cell count on admission, with normal hepatic enzyme levels and platelet counts throughout her course. Intracerebral and subarachnoid hemorrhages with uncal herniation subsequently developed, and she was declared brain-dead. The donor met the screening criteria for organ and tissue donation, and surgical teams procured the lungs, kidneys, liver and associated blood vessels, skin, and corneas. Cultures of urine and blood performed at the time of organ procurement were negative. Examination of preimplantation biopsy specimens of the liver and kidney revealed no inflammation or granulomas. An autopsy revealed infarction in the distribution of the right middle cerebral artery, subarachnoid and left frontal intracerebral hemorrhages, and a patent foramen ovale. There was no evidence of infection.
The OPO coordinating the transplantations was contacted to obtain additional information about the donor. Physicians caring for the liver and lung recipients were contacted for information on their clinical status.
The liver recipient was a 54-year-old man with cirrhosis and chronic hepatitis B and C. In the initial days after the transplantation, he had headache, fever (temperature, 39.1°C), and abdominal and right-shoulder pain. Leukopenia, thrombocytopenia, rising aminotransferase levels, and prolongation of the prothrombin time were noted. Administration of broad-spectrum antimicrobial agents was initiated; multiple cultures were negative. Exploratory laparotomy revealed intraabdominal hematoma without evidence of infection. The patient had a single, generalized seizure with hypotension and subsequent worsening of renal, hepatic, and respiratory function. No seizure focus was identified on radiologic imaging or electroencephalography. Examination of a liver-biopsy specimen revealed mild portal inflammation, liver-cell regeneration, cholestasis, and mild steatosis; these findings were interpreted as transplant-associated ischemia. Left bundle-branch block and atrial fibrillation developed. The patient became obtunded, with worsening coagulopathy and multiorgan failure. On post-transplantation day 21, high-dose methylprednisolone and antithymocyte globulin were administered for suspected acute graft rejection. Fever and hypotension persisted, with increases in aminotransferase and lactate dehydrogenase levels. The cause of his multiorgan failure was unclear. He had a cardiac arrest and died on post-transplantation day 26. An autopsy revealed extensive hepatic necrosis, bronchopneumonia, pulmonary edema, and subarachnoid hemorrhage.
The lung recipient was a 41-year-old man with cystic fibrosis. He was extubated on post-transplantation day 2 but became delirious the following day and had leukocytosis and thrombocytopenia. Chest radiographs revealed right-lower-lobe infiltrates. His temperature rose to 37.9°C, and diffuse abdominal pain and respiratory distress developed. CT of the chest on post-transplantation day 16 showed bilateral air-space disease, a finding interpreted as evidence of acute rejection, and high-dose methylprednisolone was administered. His creatinine level, prothrombin time, and aminotransferase levels steadily increased, and intermittent atrial fibrillation developed. On post-transplantation day 19, a pustular rash was noted on the face and trunk; examination of a skin-biopsy specimen revealed folliculitis, and aerobic, fungal, and viral cultures were negative. Symmetric effusions of the knees, elbows, and ankles developed. The hypoxemia and acidemia progressed, and the patient died on post-transplantation day 23. An autopsy revealed organizing diffuse alveolar damage and extensive geographic hepatic necrosis without inflammation or viral cytopathic changes.
Because of concern about transplant-transmitted infection in the organ recipients, the Rhode Island Department of Health, the Massachusetts Department of Public Health, and the CDC were contacted for assistance in investigating the causes of the recipients' illnesses. Causes considered included acute hepatitis A, leptospirosis, toxoplasmosis, enterovirus infection, and flavivirus infection. Tissue specimens from the donor and recipients were submitted to the CDC for additional testing.
Epidemiologic Investigations
For both clusters, epidemiologic investigations were conducted at the transplantation centers and coordinating OPOs. For the 2005 cluster, epidemiologic investigations were also performed at the donor's home and workplace by state public health authorities and the CDC.
Virus Isolation and Indirect Fluorescence Microscopy
Virus isolation was attempted by inoculation of Vero E6 cells with cerebrospinal fluid, serum, blood, and 10 percent fresh-tissue suspensions. Cultures were examined by thin-section electron microscopy. "Spot" slides of culture cells were also evaluated by indirect fluorescent antibody testing with the use of specific mouse hyperimmune ascitic fluids prepared against the Armstrong strain of LCMV.11
In the 2003 investigation, virus isolation was also attempted by inoculating suckling mice (Mus musculus) with fluid specimens (0.03 ml intracranially and 0.1 ml intraperitoneally) or with 10 percent homogenates of frozen tissue. Mice were killed by exposure to isoflurane, and tissues were fixed in 10 percent neutral buffered formalin and evaluated with the use of immunohistochemical stains for LCMV, as described below.
Studies in animals were performed at the CDC and were approved by the CDC Laboratory Animal Care and Use Committee. Animal research was conducted in compliance with the Animal Welfare Act and other federal statutes and regulations relating to animals and experiments involving animals, and it adhered to the principles stated in the Guide for the Care and Use of Laboratory Animals, by the U.S. National Research Council.
Histopathological and Immunohistochemical Analyses
Multiple formalin-fixed, paraffin-embedded tissue specimens, including biopsy and autopsy tissues from the donors and recipients, were stained with hematoxylin and eosin and various immunohistochemical stains by means of an immunoalkaline phosphatase technique.12 For both clusters, initial tissue specimens were evaluated with immunohistochemical stains for a number of viral agents, including flaviviruses, adenoviruses, and herpesviruses 1, 2, 3, and 5. In the investigation of the 2003 cluster, immunohistochemical testing for LCMV was performed after identification of this virus by culture, indirect fluorescent antibody testing, and electron microscopy. In the investigation of the 2005 cluster, immunohistochemical testing for LCMV was included in the initial evaluation of tissue specimens. The primary antibodies used for LCMV immunohistochemical detection included hyperimmune rabbit and mouse anti-LCMV antibodies specific for LCMV and an anti–Lassa virus monoclonal antibody reactive with both Lassa virus and LCMV.13
Genetic Detection and Characterization of Virus
RNA was extracted from clinical specimens or viral isolates, and specific molecular targets were amplified by reverse-transcriptase–PCR (RT-PCR) assays with the use of broadly reactive polymerase gene–specific primers for the detection of arenavirus RNA. The resulting complementary DNA products were purified, and nucleotide sequences were determined and analyzed. In the investigation of the 2005 cluster, after LCMV-specific sequences had been obtained from PCR products amplified from clinical specimens, a more sensitive, LCMV-specific, quantitative real-time RT-PCR (TaqMan) technology was developed and used for more extensive analysis of specimens.
Serologic Testing
An enzyme-linked immunosorbent assay was used to detect LCMV-specific IgM and IgG. The assay was performed as previously described,11 with some modifications in commercially available components, such as microtiter plates and conjugates.
Results
Investigation of the 2003 Cluster
Results of testing in the 2003 cluster are summarized in Figure 3 and Table 1. Electron microscopy of Vero E6 cell cultures inoculated with cerebrospinal fluid from Kidney Recipient 1 revealed viral particles compatible with an Old World arenavirus. Indirect fluorescent antibody testing of these cultures confirmed the identity of the arenavirus as LCMV. Subsequently, LCMV was identified by immunohistochemical analysis of the brain tissue of mice inoculated with cerebrospinal fluid from the same patient. LCMV was identified in multiple tissues from each of the four recipients. Nucleotide analysis of PCR products revealed that the viral isolates obtained from the two kidney recipients were identical and were distinct from previously described LCMV strains (Figure 1 of the Supplementary Appendix). Extensive testing of multiple donor tissues by means of immunohistochemical analysis, cell culture, and RT-PCR revealed no evidence of LCMV. Neither IgM nor IgG antibodies against LCMV were detected in the donor's serum. Interviews with the donor's family revealed no known rodent exposures. Investigation of procedures, materials, and personnel in hospitals at which the donor and recipients had received care revealed no likely route of LCMV transmission.
Figure 3. Pathological Studies Revealing Lymphocytic Choriomeningitis Virus (LCMV) in the 2003 Cluster.
Panel A, an electron micrograph of LCMV isolated in Vero E6 cells from the cerebrospinal fluid of Kidney Recipient 1, shows highly pleomorphic, 50-to-300-nm virions containing electron-dense particles. Panel B shows immunohistochemical staining of LCMV (red) in neurons and choroid-plexus ependymal cells of a mouse inoculated with the virus (immunoalkaline phosphatase with naphthol–fast red and hematoxylin counterstain; monoclonal anti-LCMV antibody). Panel C shows immunohistochemical staining of LCMV antigens in lung tissue from the lung recipient (monoclonal anti-LCMV antibody). The image in Panel D reveals extensive hepatocellular necrosis with minimal inflammatory-cell infiltrates in the liver recipient (hematoxylin and eosin). Panel E shows immunohistochemical staining of viral antigens in the transplanted liver (monoclonal anti–Lassa virus antibody). Panel F shows immunohistochemical staining of LCMV antigens in the donor kidney of Kidney Recipient 1 (monoclonal anti-LCMV antibody). Panel G shows immunohistochemical staining of viral antigens in the skin of Kidney Recipient 2 (monoclonal anti–Lassa virus antibody). All micrographs are shown at low magnification.
Investigation of the 2005 Cluster
Results of testing in the 2005 cluster are summarized in Figure 4 and Table 2 (and Figure 1 of the Supplementary Appendix). LCMV was detected by immunohistochemical staining, cell culture, and quantitative real-time RT-PCR in multiple tissues from all four recipients. Both kidney recipients had IgM antibodies reactive to LCMV. Extensive testing of multiple donor tissues revealed no evidence of LCMV. No IgM or IgG antibodies against LCMV were detected in the serum of the donor.
Figure 4. Immunohistochemical Staining for Lymphocytic Choriomeningitis Virus (LCMV) in Tissue Samples from the Donor, the Donor's Household Hamster, and Organ Recipients in the 2005 Cluster.
Red staining indicates the presence of LCMV antigens. The image in Panel A contains no immunohistochemical evidence of LCMV in choroid plexus from the donor. Panel B shows antigens in the kidney tubules of the donor's household hamster. Panel C shows LCMV antigens in lung tissue obtained at autopsy from the lung recipient; there are extensive hyaline-membrane formation and viral antigens in the interstitium. Panel D shows LCMV antigens in liver tissue obtained at autopsy from the liver recipient; viral antigens delineate the hepatocyte cytoplasmic membrane. Panel E shows LCMV antigens in a kidney specimen obtained at autopsy from Kidney Recipient B; viral antigens in endothelial cells are entering and exiting the glomerulus. Panel F shows LCMV antigens in a colon sample obtained at autopsy from Kidney Recipient B, with viral antigens in the muscularis mucosae and mucous cells of colonic glands. Panel G shows LCMV antigens in a kidney-biopsy specimen from Kidney Recipient A, who survived; viral antigens are in endothelial cells of the renal interstitium. (The studies shown in Panels A, B, E, and G used a rabbit anti-LCMV antibody, those in Panels C and D a mouse anti–Lassa virus antibody, and that in Panel F a mouse ascitic-fluid anti-LCMV antibody in an immunohistochemical assay with naphthol–fast red substrate and hematoxylin counterstain.) All micrographs are shown at low magnification.
Table 2. Summary of Laboratory Evaluations for Lymphocytic Choriomeningitis Virus Infection in the 2005 Cluster.
The epidemiologic investigation revealed that a member of the donor's household had brought home a pet hamster three weeks before the donor died. Although the donor had not been the primary caretaker of the hamster, she had had contact with the rodent's environment on multiple occasions. Her home and work environment contained no evidence of active rodent infestation. Testing of multiple hamster tissues by immunohistochemical analysis, quantitative real-time RT-PCR, and viral culture detected evidence of LCMV infection (Figure 4 and Table 2). The primary caretaker of the hamster was asymptomatic but had LCMV-reactive IgG and IgM antibodies present in the serum. Nucleotide analysis of PCR products identified that the viral isolates from all four organ recipients and the hamster were identical, but that they differed from the strain identified in the 2003 cluster and previously described LCMV strains (Figure 1 of the Supplementary Appendix).
After identification of LCMV as the etiologic agent, intravenous ribavirin (a loading dose of 30 mg per kilogram of body weight, followed by 16 mg per kilogram every six hours for four days and then 8 mg per kilogram every eight hours) was initiated in Kidney Recipient A beginning on post-transplantation day 26 (Figure 2 of the Supplementary Appendix).14 The fever and diarrhea decreased; the pain, tenderness, and erythema in the area of the allograft diminished; and the hypoxemia, elevation of aminotransferase levels, thrombocytopenia, and coagulopathy decreased. After the patient's clinical condition had stabilized, ribavirin administration was changed to the oral route (400 mg every morning and 600 mg every evening), and it was discontinued after a renal-biopsy specimen was found to be LCMV-negative by RT-PCR and immunohistochemical staining and after serum IgM became detectable, 63 days after transplantation. During 37 days of ribavirin treatment, clinically significant hemolytic anemia developed and required the transfusion of 19 units of packed red cells. Three hundred eleven days after transplantation, the patient had stable graft function without evidence of infection, after he had restarted immunosuppressive therapy with tacrolimus, mycophenolate mofetil, and prednisolone.
Tissue Disposition
In the 2003 cluster, no additional tissues from the donor were transplanted. In the 2005 cluster, the corneas were transplanted into a 4-year-old girl and a 29-year-old woman in Algeria, neither of whom required systemic immunosuppression. Two hundred thirteen days after transplantation, neither patient had reported symptoms of infection or graft loss. The skin and liver-associated blood vessels were not transplanted.
Discussion
We describe the transmission of LCMV by solid-organ transplantation. In both clusters, disseminated infection developed in the recipients of organs from a common donor who had no clinical or laboratory evidence of infection. In the 2003 cluster, no rodent exposures could be identified, whereas the 2005 cluster was associated with recent donor exposure to an LCMV-infected pet hamster. The isolation of identical virus strains from the two kidney recipients in the 2003 cluster and from all the organ recipients and the donor household's pet hamster in the 2005 cluster indicates that in both clusters, LCMV was transmitted through organ transplantation.15
LCMV infection in humans is sporadic, is generally benign, and can be asymptomatic.4,16,17 Serologic surveys suggest that up to 5 percent of adults in the United States have been infected with LCMV.18,19 Humans become infected with LCMV by direct contact with rodents or through aerosolized droplets from rodent secretions and excretions, including urine or feces.20,21 Reports of sporadic infection have most frequently implicated the common house mouse, although outbreaks of infection have been reported after exposure to infected pet hamsters and laboratory rodents.17,20,21,22,23,24,25,26,27,28
In clinically apparent infection, the incubation period is 5 to 13 days, with subsequent fever, headache, and myalgias. Abdominal pain, diarrhea, and rash have been described.16,20,22,26,29 A second phase of illness may be seen five to nine days after convalescence, with meningitis or, rarely, encephalomyelitis, orchitis, parotitis, pneumonitis, arthritis, myocarditis, or alopecia.16,22,30 Recognized LCMV infection carries a mortality rate of less than 1 percent.3
The marked severity of LCMV-related illness in transplant recipients is probably the result of intensive immunosuppression, including T-cell depletion, coincident with direct viral inoculation by way of the transplanted organs. There are few data on the clinical behavior and outcomes of LCMV infection in immunocompromised patients. However, three patients with advanced lymphoma experimentally inoculated with the virus in a trial investigating its antitumor effect died with disseminated infection within 14 to 45 days after inoculation31 — a pattern similar to that seen in the current clusters after solid-organ transplantation.
The clinical presentation of LCMV infection in the recipients was variable and included fever, diarrhea, peri-incisional erythema and tenderness, altered mental status, and respiratory insufficiency (as noted in the table of the Supplementary Appendix). Leukopenia or leukocytosis, thrombocytopenia, coagulopathy, renal insufficiency, and progressive liver dysfunction dominated the laboratory findings. Histopathological findings in all the recipients were characterized by necrotic and occasionally hemorrhagic foci in multiple tissues, with a notable absence of inflammatory infiltrates and viral inclusions. LCMV antigens present in some tissues (e.g., the gastrointestinal tract and skin) correlated with clinical symptoms (e.g., diarrhea and erythema or pustular rash, respectively). Antigens were identified in the leptomeninges of some patients in both clusters. However, signs of meningeal inflammation, though prominent in the 2003 cluster, were generally absent in the 2005 cluster. Clinical manifestations and pathological findings in all the cases were probably altered by immunosuppression.
To our knowledge, there have been no trials of antiviral agents in human LCMV disease. Ribavirin has demonstrated efficacy in the treatment of Lassa fever and possesses in vitro activity against LCMV infection, which prompted its use in the renal-transplant recipient who survived.14,32 The role of ribavirin in this patient's improvement is unclear, since the level of immunosuppression was also considerably reduced, as it was for all four kidney recipients in the two clusters.
Rapid evaluation of organ-donor suitability is essential in transplantation to minimize the duration of ischemia and to preserve allograft function. Therefore, assays used to screen potential donors for transmissible infections must be rapid, sensitive, reproducible, and readily available to OPOs. The Food and Drug Administration has not approved any diagnostic tests for LCMV infection. Furthermore, the sensitivity of currently available assays is not adequate for routine donor screening, as demonstrated by the negative results of tests on a wide array of clinical specimens from the donors in both clusters. The use of information pertaining to recent rodent exposure for donor-suitability screening may exclude healthy donors from an already limited organ-donor pool. However, the collection of additional epidemiologic information on donors' exposures may be useful, notably for the investigation of unusual outcomes after transplantation. Zoonotic-disease transmission after transplantation is also a concern; immunosuppressed persons should take special care and limit exposure to some animals, including certain pets (additional information is available from the CDC at www.cdc.gov/healthypets).
Transplant recipients are susceptible to infection with a variety of donor-derived pathogens, including West Nile virus, Trypanosoma cruzi, rabies virus, and now LCMV.33,34,35,36 Although such infections are probably uncommon, outcomes can be fatal, and diagnosis is feasible with specialized laboratory testing. Diagnosis of LCMV infection is usually made by serologic testing, isolation of the virus from the blood or cerebrospinal fluid, or PCR testing.37,38,39 Because immunohistochemical staining revealed LCMV in multiple biopsy specimens obtained to evaluate unexplained post-transplantation symptoms in the clusters described in the current report, such testing might be beneficial in the early diagnosis of LCMV infection.
Each year, approximately 25,000 organ transplantations are performed at more than 250 transplantation centers throughout the United States.40 Allocation policies commonly result in the distribution of organs from a single donor to multiple transplantation centers. It is unlikely that either LCMV-illness cluster would have been identified without the allocation of kidneys to two recipients in whom similar symptoms simultaneously developed after undergoing transplantation at the same hospital. Similar chance clinical observations have been critical in the recognition of recent transplant-associated outbreaks of rabies and West Nile virus infection.33,35
The Organ Procurement and Transplantation Network (OPTN), which is operated by the United Network for Organ Sharing, requires transplantation centers to report certain outcomes, including allograft failure and the death of transplant recipients, in a timely manner. In April 2005, the OPTN revised its policies to require the reporting of suspected donor-transmitted medical conditions (including cancers and infections) to the procuring OPO, which is then responsible for investigating and communicating with the transplantation centers caring for the recipients of other transplants from the donor and the involved tissue and eye banks.41
Investigation of potential donor-transmitted infection requires rapid communication among physicians in multiple transplantation centers, OPOs, and public health authorities. An immediate system for tracking and disseminating pertinent patient data is needed. Until such a system can be established, clinicians must recognize that the presence of an unusual constellation of symptoms, particularly during the first few weeks after transplantation, should raise the possibility of donor-transmitted infection. Prompt notification of the OPO and public health authorities can help facilitate rapid investigation and discovery of these events.
Dr. Kotton reports having received grant support from Wyeth and the Massachusetts General Hospital Clafin Award. Dr. Fishman reports having received consulting fees or lecture fees from Astellas, Enzon, Novartis, Pfizer, and Roche. No other potential conflict of interest relevant to this article was reported.
The views expressed herein are those of the authors and not necessarily those of the Department of Health and Human Services.
We are indebted to Jennifer Betts, Kimberly Slaughter, Deborah Cannon, and Thomas Stevens for specimen processing, serologic testing, and virus isolation and characterization; to Nicole Lundstrom and Sherry Hayes for manuscript preparation; and to Donita Croft, James Kazmierczak, Mark Sotir, and Mark Wegner for investigation assistance.
* The members of the Lymphocytic Choriomeningitis Virus (LCMV) in Transplant Recipients Investigation Team are listed in the Appendix.
Source Information
From Rhode Island Hospital and Brown Medical School, Providence (S.A.F.); the Medical College of Wisconsin, Milwaukee (M.B.G.); the Divisions of Viral and Rickettsial Diseases (M.J.K., J.A.C., J.G., C.D.P., W.-J.S., B.R.E., B.P., A.L., M.J.V., T.K.S., C.S.G., P.E.R., M.M.P., M.P., C.R., R.F.H., S.T.N., T.K., S.R.Z.) and Healthcare Quality Promotion (A.S., D.B.J.), National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta; Massachusetts General Hospital and Harvard Medical School (C.N.K., F.L.D., J.A.F.) and Brigham and Women's Hospital and Harvard Medical School (F.M.M., D.L.D.) — all in Boston; the Rhode Island Department of Health, Providence (U.B.); the Massachusetts Department of Public Health, Boston (A.D.); the Wisconsin Department of Health and Family Services, Madison (J.P.D.); and the New England Organ Bank, Newton, Mass. (F.L.D.).
Drs. Ksiazek and Zaki contributed equally to this article.
Address reprint requests to Dr. Kuehnert at the Division of Viral and Rickettsial Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, 1600 Clifton Rd., Mailstop A-30, Atlanta, GA 30333, or at mkuehnert@cdc.gov.
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Enders G, Varho-G?bel M, L?hler J, Terletskaia-Ladwig E, Eggers M. Congenital lymphocytic choriomeningitis virus infection: an underdiagnosed disease. Pediatr Infect Dis J 1999;18:652-655.
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Paddock C, Ksiazek T, Comer JA, et al. Pathology of fatal lymphocytic choriomeningitis virus infection in multiple organ transplant recipients from a common donor. Mod Pathol 2005;18:Suppl 1:263A-263A.
Ksiazek TG, Erdman D, Goldsmith CS, et al. A novel coronavirus associated with severe acute respiratory syndrome. N Engl J Med 2003;348:1953-1966.
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Puglielli MT, Browning JL, Brewer AW, et al. Reversal of viral-induced systemic shock and respiratory failure by blockade of the lymphotoxin pathway. Nat Med 1999;5:1370-1374.
McCormick JB, King IJ, Webb PA, et al. Lassa fever: effective therapy with ribavirin. N Engl J Med 1986;314:20-26.
Lymphocytic choriomeningitis virus infection in organ transplant recipients -- Massachusetts, Rhode Island, 2005. MMWR Morb Mortal Wkly Rep 2005;54:537-539.
Rousseau MC, Saron MF, Brouqui P, Bourgeade A. Lymphocytic choriomeningitis virus in southern France: four case reports and a review of the literature. Eur J Epidemiol 1997;13:817-823.
Baum SG, Lewis AM, Rowe WP, Huebner RJ. Epidemic nonmeningitic lymphocytic-choriomeningitis-virus infection: an outbreak in a population of laboratory personnel. N Engl J Med 1966;274:934-936.
Stephensen CB, Blount SR, Lanford RE, et al. Prevalence of serum antibodies against lymphocytic choriomeningitis virus in selected populations from two U.S. cities. J Med Virol 1992;38:27-31.
Childs JE, Glass GE, Ksiazek TG, Rossi CA, Oro JG, Leduc JW. Human-rodent contact and infection with lymphocytic choriomeningitis and Seoul viruses in an inner-city population. Am J Trop Med Hyg 1991;44:117-121.
Biggar RJ, Woodall JP, Walter PD, Haughie GE. Lymphocytic choriomeningitis outbreak associated with pet hamsters: fifty-seven cases from New York State. JAMA 1975;232:494-500.
Dykewicz CA, Dato VM, Fisher-Hoch SP, et al. Lymphocytic choriomeningitis outbreak associated with nude mice in a research institute. JAMA 1992;267:1349-1353.
Vanzee BE, Douglas RG, Betts RF, Bauman AW, Fraser DW, Hinman AR. Lymphocytic choriomeningitis in university hospital personnel: clinical features. Am J Med 1975;58:803-809.
Biggar RJ, Schmidt TJ, Woodall JP. Lymphocytic choriomeningitis in laboratory personnel exposed to hamsters inadvertently infected with LCM virus. J Am Vet Med Assoc 1977;171:829-832.
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Hirsch MS, Moellering RC Jr, Pope HG, Poskanzer DC. Lymphocytic-choriomeningitis-virus infection traced to a pet hamster. N Engl J Med 1974;291:610-612.
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Iwamoto M, Jernigan DB, Guasch A, et al. Transmission of West Nile virus from an organ donor to four transplant recipients. N Engl J Med 2003;348:2196-2203.
West Nile infections in organ transplant recipients -- New York and Pennsylvania, August-September 2005. MMWR Morb Mortal Wkly Rep 2005;54:1021-1023.
Srinivasan A, Burton EC, Kuehnert MJ, et al. Transmission of rabies virus from an organ donor to four transplant recipients. N Engl J Med 2005;352:1103-1111.
Chagas disease after organ transplantation -- United States, 2001. MMWR Morb Mortal Wkly Rep 2002;51:210-212.
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OPTN/SRTR 2004 annual report. Richmond, Va.: Organ Procurement and Transplantation Network, United Network for Organ Sharing, 2004. (Accessed May 1, 2006, at http://www.optn.org/data/annualReport.asp.)
Acquired immune deficiency syndrome (AIDS), human pituitary derived growth hormone (HPDGH), and reporting of potential recipient diseases or medical conditions, including malignancies, of donor origin. Richmond, Va.: Organ Procurement and Transplantation Network, United Network for Organ Sharing, 2004. (Accessed May 1, 2006, at http://www.optn.org/PoliciesandBylaws/policies/docs/policy_16.doc.)(Staci A. Fischer, M.D., M)
Background In December 2003 and April 2005, signs and symptoms suggestive of infection developed in two groups of recipients of solid-organ transplants. Each cluster was investigated because diagnostic evaluations were unrevealing, and in each a common donor was recognized.
Methods We examined clinical specimens from the two donors and eight recipients, using viral culture, electron microscopy, serologic testing, molecular analysis, and histopathological examination with immunohistochemical staining to identify a cause. Epidemiologic investigations, including interviews, environmental assessments, and medical-record reviews, were performed to characterize clinical courses and to determine the cause of the illnesses.
Results Laboratory testing revealed lymphocytic choriomeningitis virus (LCMV) in all the recipients, with a single, unique strain of LCMV identified in each cluster. In both investigations, LCMV could not be detected in the organ donor. In the 2005 cluster, the donor had had contact in her home with a pet hamster infected with an LCMV strain identical to that detected in the organ recipients; no source of LCMV infection was found in the 2003 cluster. The transplant recipients had abdominal pain, altered mental status, thrombocytopenia, elevated aminotransferase levels, coagulopathy, graft dysfunction, and either fever or leukocytosis within three weeks after transplantation. Diarrhea, peri-incisional rash, renal failure, and seizures were variably present. Seven of the eight recipients died, 9 to 76 days after transplantation. One recipient, who received ribavirin and reduced levels of immunosuppressive therapy, survived.
Conclusions We document two clusters of LCMV infection transmitted through organ transplantation.
Lymphocytic choriomeningitis virus (LCMV) is a rodent-borne, Old World arenavirus that has been reported to cause asymptomatic or mild, self-limited illness in otherwise healthy humans. It is a known cause of aseptic meningitis, but fatal infection is rare.1,2,3,4 Transmission of infection from a woman to a fetus can result in hydrocephalus, chorioretinitis, or microcephaly.5,6,7,8 Outside of vertical transmission during pregnancy, human-to-human transmission of LCMV has not been described.9 We describe two clusters of unexplained clinical syndromes in transplant recipients and the subsequent investigations to identify donor-transmitted infection as the cause of illness.
Methods
Case Reports
The 2003 Cluster
In December 2003, unexplained febrile illnesses developed in four recipients of solid organs from a common donor (Figure 1; additional information on clinical symptoms and laboratory findings for each recipient is listed in Table 1 of the Supplementary Appendix, available with the full text of this article at www.nejm.org).10 Kidney Recipient 1 was a 46-year-old man with diabetes. Diarrhea and mild, diffuse abdominal pain developed on post-transplantation day 5, but his condition was stable and he was discharged home the following day. He was readmitted on post-transplantation day 23 with fever, persistent watery diarrhea, and worsening abdominal pain. Laboratory studies revealed leukopenia with elevated aminotransferase and creatinine levels. Ganciclovir therapy was initiated because of concern about possible cytomegalovirus infection. Tacrolimus and mycophenolate mofetil were discontinued. Examination of kidney-, liver-, and bone marrow–biopsy specimens did not reveal an infectious cause. On day 40 after transplantation, seizures and polymyoclonus developed. The patient reported blurred vision, and chorioretinitis was noted on ophthalmologic examination. Cerebrospinal fluid studies revealed a markedly elevated level of protein (720 mg per deciliter), a normal glucose level (147 mg per deciliter ), and 4 white cells and 3 red cells per cubic centimeter. Polymerase-chain-reaction (PCR) tests for cytomegalovirus, herpes simplex virus, varicella–zoster virus, Epstein–Barr virus, human herpesvirus 6, enterovirus, adenovirus, Mycobacterium tuberculosis, and Borrelia burgdorferi were negative. Magnetic resonance imaging of the brain revealed bilateral, hemispheric, subdural fluid collections and diffuse dural thickening. Examination of a specimen obtained by dural biopsy on post-transplantation day 47 showed fibrosis; cidofovir and intravenous immune globulin were initiated for the suspected presence of an unknown viral pathogen. The patient's condition continued to deteriorate, and he died on post-transplantation day 53. An autopsy revealed bronchopneumonia and hepatic congestion without inflammation.
Figure 1. Clinical Course of Lymphocytic Choriomeningitis Virus (LCMV) Infection in the 2003 Cluster.
MMF denotes mycophenolate mofetil, TMP-SMX trimethoprim–sulfamethoxazole, IVIG intravenous immune globulin, and X death. Immunosuppressive agents are shown in red, and antimicrobial agents in blue.
Kidney Recipient 2 was a 56-year-old man with glomerulonephritis. He was discharged home on post-transplantation day 5 but was readmitted on post-transplantation day 22 with fever, leukopenia, and peri-incisional erythema and tenderness. A skin-biopsy specimen obtained at the wound edge revealed basal-cell vacuolation suggestive of viral infection; no viral inclusions were seen, and immunohistochemical stains were negative for cytomegalovirus and adenovirus. Tacrolimus and mycophenolate mofetil were discontinued. Altered mental status and seizures with myoclonus developed on post-transplantation day 31. Cerebrospinal fluid studies revealed a markedly elevated protein level (620 mg per deciliter), a low glucose level (57 mg per deciliter ), and 12 white cells (48 percent lymphocytes and 22 percent monocytes) and 6 red cells per cubic centimeter. PCR testing for the same infectious agents as in the case of Kidney Recipient 1 was unrevealing. Magnetic resonance imaging showed subdural fluid collections with diffuse dural enhancement. The patient's condition continued to deteriorate, with photophobia, nuchal rigidity, diarrhea, thrombocytopenia, diffuse erythroderma, respiratory failure, and atrial fibrillation, despite empirical administration of cidofovir and intravenous immune globulin. He died on post-transplantation day 76; an autopsy revealed meningoencephalitis and acute bronchopneumonia.
The liver recipient was a 40-year-old woman with alcoholic cirrhosis whose early postoperative course was marked by fever, lethargy, and hypotension. Markedly elevated aminotransferase levels, leukopenia, and bilateral pulmonary infiltrates with respiratory failure developed. Liver biopsy revealed focal centrilobular coagulative necrosis. On post-transplantation day 14, a peri-incisional petechial rash was noted; skin biopsy revealed chronic inflammation and hemorrhage. She died on day 17 after transplantation. An autopsy revealed extensive hepatic necrosis without evidence of infection.
A common source of infection in these recipients was suspected, all of whom underwent transplantation at the same facility. Therefore, the Department of Public Health of the Wisconsin Department of Health and Family Services and the organ-procurement organization (OPO) coordinating the transplantations were notified for assistance.
The lung recipient, who had undergone transplantation at a different facility, was a 46-year-old man with chronic obstructive pulmonary disease who had been receiving prednisolone daily for three months before transplantation. Within four days after transplantation, hypotension, bilateral pulmonary infiltrates, and leukocytosis developed; broad-spectrum antimicrobial agents were administered. Fever (temperature, 38.3°C), hypoxia, and refractory hypotension ensued. He died on day 9 after transplantation. An autopsy revealed diffuse alveolar damage without evidence of rejection.
The donor was a 51-year-old man who had been found unresponsive, with apparent head trauma. Computed tomography (CT) of the brain revealed a large, right-sided subdural hematoma with a midline shift. There was no improvement in his neurologic status, and he was declared brain-dead on hospital day 2. He received no blood products and was afebrile throughout his hospitalization. Donor-eligibility screening and testing detected no infection precluding organ or tissue donation. His liver, lungs, kidneys, and multiple musculoskeletal and vascular tissues were recovered for transplantation. Tissue specimens from the donor and recipients were submitted to the Centers for Disease Control and Prevention (CDC) for additional testing (Table 1).
Table 1. Summary of Laboratory Evaluations for Lymphocytic Choriomeningitis Virus Infection in the 2003 Cluster.
The 2005 Cluster
Kidney Recipient A was a 48-year-old man admitted to a hospital in Rhode Island in late April 2005, 17 days after undergoing cadaveric renal transplantation (Figure 2). He had been discharged home on post-transplantation day 7 with a creatinine level of 1.6 mg per deciliter (141 μmol per liter). At the time of readmission, he had had right-lower-quadrant pain in the area of the allograft for four or five days, as well as nausea, anorexia, diarrhea, fever, and chills. He was febrile (temperature, 38.7°C), and there was tender erythema over the area of the allograft, without incisional dehiscence or drainage. His creatinine level was 2.6 mg per deciliter (230 μmol per liter), and proteinuria, hematuria, a prolonged prothrombin time, and slightly elevated aminotransferase levels were present. Mycophenolate mofetil was discontinued, and administration of broad-spectrum antimicrobial agents was initiated. CT and ultrasonography of the abdomen and pelvis were unrevealing. Routine cultures of urine, blood, and stool were negative, as were studies of the stool for leukocytes, ova, parasites, Clostridium difficile, giardia, cryptosporidium, Yersinia enterocolitica, and rotavirus. Tacrolimus was discontinued because of concern about the worsening infection of uncertain cause. His temperature rose to 40.4°C, and he had copious diarrhea, dyspnea, and tender erythema extending from the area over the allograft to the right flank. Examination of biopsy specimens of the colon and kidney revealed no inflammation or viral inclusions.
Figure 2. Clinical Course of Lymphocytic Choriomeningitis Virus (LCMV) Infection in the 2005 Cluster.
MMF denotes mycophenolate mofetil, TMP-SMX trimethoprim–sulfamethoxazole, CVVH continuous venous–venous hyperfiltration, and X death. Immunosuppressive agents are shown in red, and antimicrobial agents in blue.
Kidney Recipient B was a 54-year-old man who was admitted to the same hospital with fever. He had undergone cadaveric renal transplantation 17 days previously and had been discharged home on post-transplantation day 8 with a creatinine level of 3.3 mg per deciliter (292 μmol per liter). On post-transplantation day 14, mycophenolate mofetil was discontinued because of diarrhea. Fever and pain developed in the right lower quadrant over the area of the allograft, and he was readmitted for evaluation. He was lethargic and febrile (temperature, 38.4°C) and had tender erythema overlying the allograft, without incisional drainage. His creatinine level was 4.0 mg per deciliter (354 μmol per liter), with a platelet count of 113,000 per cubic millimeter and an alanine aminotransferase level of 298 IU per liter. Routine cultures of urine, blood, and stool were negative, and ultrasonography and CT of the abdomen and pelvis were unrevealing. Fever, diarrhea, and pain persisted, despite empirical use of broad-spectrum antimicrobial agents. A percutaneous liver biopsy was performed on post-transplantation day 23 because of worsening hepatitis and leukocytosis. Multiple foci of hepatocellular necrosis without inflammation or viral inclusions were noted. Later that day, he had a cardiac arrest and died. An autopsy revealed coronary artery disease and diffuse cerebral edema without meningitis or encephalitis.
A review of the records of the hospital's transplantation center revealed that the kidney recipients shared a common donor who had also been hospitalized there. The donor was a 45-year-old woman with hypertension who had presented to the emergency department with a five-day history of right-sided headache and acute left-sided weakness. She was alert and afebrile and had left-sided hemiparesis. CT of the brain revealed an infarct in the distribution of the right middle cerebral artery, and tissue plasminogen activator was administered. Throughout her hospitalization, she was afebrile and received no blood products or antimicrobial agents. She had a normal white-cell count on admission, with normal hepatic enzyme levels and platelet counts throughout her course. Intracerebral and subarachnoid hemorrhages with uncal herniation subsequently developed, and she was declared brain-dead. The donor met the screening criteria for organ and tissue donation, and surgical teams procured the lungs, kidneys, liver and associated blood vessels, skin, and corneas. Cultures of urine and blood performed at the time of organ procurement were negative. Examination of preimplantation biopsy specimens of the liver and kidney revealed no inflammation or granulomas. An autopsy revealed infarction in the distribution of the right middle cerebral artery, subarachnoid and left frontal intracerebral hemorrhages, and a patent foramen ovale. There was no evidence of infection.
The OPO coordinating the transplantations was contacted to obtain additional information about the donor. Physicians caring for the liver and lung recipients were contacted for information on their clinical status.
The liver recipient was a 54-year-old man with cirrhosis and chronic hepatitis B and C. In the initial days after the transplantation, he had headache, fever (temperature, 39.1°C), and abdominal and right-shoulder pain. Leukopenia, thrombocytopenia, rising aminotransferase levels, and prolongation of the prothrombin time were noted. Administration of broad-spectrum antimicrobial agents was initiated; multiple cultures were negative. Exploratory laparotomy revealed intraabdominal hematoma without evidence of infection. The patient had a single, generalized seizure with hypotension and subsequent worsening of renal, hepatic, and respiratory function. No seizure focus was identified on radiologic imaging or electroencephalography. Examination of a liver-biopsy specimen revealed mild portal inflammation, liver-cell regeneration, cholestasis, and mild steatosis; these findings were interpreted as transplant-associated ischemia. Left bundle-branch block and atrial fibrillation developed. The patient became obtunded, with worsening coagulopathy and multiorgan failure. On post-transplantation day 21, high-dose methylprednisolone and antithymocyte globulin were administered for suspected acute graft rejection. Fever and hypotension persisted, with increases in aminotransferase and lactate dehydrogenase levels. The cause of his multiorgan failure was unclear. He had a cardiac arrest and died on post-transplantation day 26. An autopsy revealed extensive hepatic necrosis, bronchopneumonia, pulmonary edema, and subarachnoid hemorrhage.
The lung recipient was a 41-year-old man with cystic fibrosis. He was extubated on post-transplantation day 2 but became delirious the following day and had leukocytosis and thrombocytopenia. Chest radiographs revealed right-lower-lobe infiltrates. His temperature rose to 37.9°C, and diffuse abdominal pain and respiratory distress developed. CT of the chest on post-transplantation day 16 showed bilateral air-space disease, a finding interpreted as evidence of acute rejection, and high-dose methylprednisolone was administered. His creatinine level, prothrombin time, and aminotransferase levels steadily increased, and intermittent atrial fibrillation developed. On post-transplantation day 19, a pustular rash was noted on the face and trunk; examination of a skin-biopsy specimen revealed folliculitis, and aerobic, fungal, and viral cultures were negative. Symmetric effusions of the knees, elbows, and ankles developed. The hypoxemia and acidemia progressed, and the patient died on post-transplantation day 23. An autopsy revealed organizing diffuse alveolar damage and extensive geographic hepatic necrosis without inflammation or viral cytopathic changes.
Because of concern about transplant-transmitted infection in the organ recipients, the Rhode Island Department of Health, the Massachusetts Department of Public Health, and the CDC were contacted for assistance in investigating the causes of the recipients' illnesses. Causes considered included acute hepatitis A, leptospirosis, toxoplasmosis, enterovirus infection, and flavivirus infection. Tissue specimens from the donor and recipients were submitted to the CDC for additional testing.
Epidemiologic Investigations
For both clusters, epidemiologic investigations were conducted at the transplantation centers and coordinating OPOs. For the 2005 cluster, epidemiologic investigations were also performed at the donor's home and workplace by state public health authorities and the CDC.
Virus Isolation and Indirect Fluorescence Microscopy
Virus isolation was attempted by inoculation of Vero E6 cells with cerebrospinal fluid, serum, blood, and 10 percent fresh-tissue suspensions. Cultures were examined by thin-section electron microscopy. "Spot" slides of culture cells were also evaluated by indirect fluorescent antibody testing with the use of specific mouse hyperimmune ascitic fluids prepared against the Armstrong strain of LCMV.11
In the 2003 investigation, virus isolation was also attempted by inoculating suckling mice (Mus musculus) with fluid specimens (0.03 ml intracranially and 0.1 ml intraperitoneally) or with 10 percent homogenates of frozen tissue. Mice were killed by exposure to isoflurane, and tissues were fixed in 10 percent neutral buffered formalin and evaluated with the use of immunohistochemical stains for LCMV, as described below.
Studies in animals were performed at the CDC and were approved by the CDC Laboratory Animal Care and Use Committee. Animal research was conducted in compliance with the Animal Welfare Act and other federal statutes and regulations relating to animals and experiments involving animals, and it adhered to the principles stated in the Guide for the Care and Use of Laboratory Animals, by the U.S. National Research Council.
Histopathological and Immunohistochemical Analyses
Multiple formalin-fixed, paraffin-embedded tissue specimens, including biopsy and autopsy tissues from the donors and recipients, were stained with hematoxylin and eosin and various immunohistochemical stains by means of an immunoalkaline phosphatase technique.12 For both clusters, initial tissue specimens were evaluated with immunohistochemical stains for a number of viral agents, including flaviviruses, adenoviruses, and herpesviruses 1, 2, 3, and 5. In the investigation of the 2003 cluster, immunohistochemical testing for LCMV was performed after identification of this virus by culture, indirect fluorescent antibody testing, and electron microscopy. In the investigation of the 2005 cluster, immunohistochemical testing for LCMV was included in the initial evaluation of tissue specimens. The primary antibodies used for LCMV immunohistochemical detection included hyperimmune rabbit and mouse anti-LCMV antibodies specific for LCMV and an anti–Lassa virus monoclonal antibody reactive with both Lassa virus and LCMV.13
Genetic Detection and Characterization of Virus
RNA was extracted from clinical specimens or viral isolates, and specific molecular targets were amplified by reverse-transcriptase–PCR (RT-PCR) assays with the use of broadly reactive polymerase gene–specific primers for the detection of arenavirus RNA. The resulting complementary DNA products were purified, and nucleotide sequences were determined and analyzed. In the investigation of the 2005 cluster, after LCMV-specific sequences had been obtained from PCR products amplified from clinical specimens, a more sensitive, LCMV-specific, quantitative real-time RT-PCR (TaqMan) technology was developed and used for more extensive analysis of specimens.
Serologic Testing
An enzyme-linked immunosorbent assay was used to detect LCMV-specific IgM and IgG. The assay was performed as previously described,11 with some modifications in commercially available components, such as microtiter plates and conjugates.
Results
Investigation of the 2003 Cluster
Results of testing in the 2003 cluster are summarized in Figure 3 and Table 1. Electron microscopy of Vero E6 cell cultures inoculated with cerebrospinal fluid from Kidney Recipient 1 revealed viral particles compatible with an Old World arenavirus. Indirect fluorescent antibody testing of these cultures confirmed the identity of the arenavirus as LCMV. Subsequently, LCMV was identified by immunohistochemical analysis of the brain tissue of mice inoculated with cerebrospinal fluid from the same patient. LCMV was identified in multiple tissues from each of the four recipients. Nucleotide analysis of PCR products revealed that the viral isolates obtained from the two kidney recipients were identical and were distinct from previously described LCMV strains (Figure 1 of the Supplementary Appendix). Extensive testing of multiple donor tissues by means of immunohistochemical analysis, cell culture, and RT-PCR revealed no evidence of LCMV. Neither IgM nor IgG antibodies against LCMV were detected in the donor's serum. Interviews with the donor's family revealed no known rodent exposures. Investigation of procedures, materials, and personnel in hospitals at which the donor and recipients had received care revealed no likely route of LCMV transmission.
Figure 3. Pathological Studies Revealing Lymphocytic Choriomeningitis Virus (LCMV) in the 2003 Cluster.
Panel A, an electron micrograph of LCMV isolated in Vero E6 cells from the cerebrospinal fluid of Kidney Recipient 1, shows highly pleomorphic, 50-to-300-nm virions containing electron-dense particles. Panel B shows immunohistochemical staining of LCMV (red) in neurons and choroid-plexus ependymal cells of a mouse inoculated with the virus (immunoalkaline phosphatase with naphthol–fast red and hematoxylin counterstain; monoclonal anti-LCMV antibody). Panel C shows immunohistochemical staining of LCMV antigens in lung tissue from the lung recipient (monoclonal anti-LCMV antibody). The image in Panel D reveals extensive hepatocellular necrosis with minimal inflammatory-cell infiltrates in the liver recipient (hematoxylin and eosin). Panel E shows immunohistochemical staining of viral antigens in the transplanted liver (monoclonal anti–Lassa virus antibody). Panel F shows immunohistochemical staining of LCMV antigens in the donor kidney of Kidney Recipient 1 (monoclonal anti-LCMV antibody). Panel G shows immunohistochemical staining of viral antigens in the skin of Kidney Recipient 2 (monoclonal anti–Lassa virus antibody). All micrographs are shown at low magnification.
Investigation of the 2005 Cluster
Results of testing in the 2005 cluster are summarized in Figure 4 and Table 2 (and Figure 1 of the Supplementary Appendix). LCMV was detected by immunohistochemical staining, cell culture, and quantitative real-time RT-PCR in multiple tissues from all four recipients. Both kidney recipients had IgM antibodies reactive to LCMV. Extensive testing of multiple donor tissues revealed no evidence of LCMV. No IgM or IgG antibodies against LCMV were detected in the serum of the donor.
Figure 4. Immunohistochemical Staining for Lymphocytic Choriomeningitis Virus (LCMV) in Tissue Samples from the Donor, the Donor's Household Hamster, and Organ Recipients in the 2005 Cluster.
Red staining indicates the presence of LCMV antigens. The image in Panel A contains no immunohistochemical evidence of LCMV in choroid plexus from the donor. Panel B shows antigens in the kidney tubules of the donor's household hamster. Panel C shows LCMV antigens in lung tissue obtained at autopsy from the lung recipient; there are extensive hyaline-membrane formation and viral antigens in the interstitium. Panel D shows LCMV antigens in liver tissue obtained at autopsy from the liver recipient; viral antigens delineate the hepatocyte cytoplasmic membrane. Panel E shows LCMV antigens in a kidney specimen obtained at autopsy from Kidney Recipient B; viral antigens in endothelial cells are entering and exiting the glomerulus. Panel F shows LCMV antigens in a colon sample obtained at autopsy from Kidney Recipient B, with viral antigens in the muscularis mucosae and mucous cells of colonic glands. Panel G shows LCMV antigens in a kidney-biopsy specimen from Kidney Recipient A, who survived; viral antigens are in endothelial cells of the renal interstitium. (The studies shown in Panels A, B, E, and G used a rabbit anti-LCMV antibody, those in Panels C and D a mouse anti–Lassa virus antibody, and that in Panel F a mouse ascitic-fluid anti-LCMV antibody in an immunohistochemical assay with naphthol–fast red substrate and hematoxylin counterstain.) All micrographs are shown at low magnification.
Table 2. Summary of Laboratory Evaluations for Lymphocytic Choriomeningitis Virus Infection in the 2005 Cluster.
The epidemiologic investigation revealed that a member of the donor's household had brought home a pet hamster three weeks before the donor died. Although the donor had not been the primary caretaker of the hamster, she had had contact with the rodent's environment on multiple occasions. Her home and work environment contained no evidence of active rodent infestation. Testing of multiple hamster tissues by immunohistochemical analysis, quantitative real-time RT-PCR, and viral culture detected evidence of LCMV infection (Figure 4 and Table 2). The primary caretaker of the hamster was asymptomatic but had LCMV-reactive IgG and IgM antibodies present in the serum. Nucleotide analysis of PCR products identified that the viral isolates from all four organ recipients and the hamster were identical, but that they differed from the strain identified in the 2003 cluster and previously described LCMV strains (Figure 1 of the Supplementary Appendix).
After identification of LCMV as the etiologic agent, intravenous ribavirin (a loading dose of 30 mg per kilogram of body weight, followed by 16 mg per kilogram every six hours for four days and then 8 mg per kilogram every eight hours) was initiated in Kidney Recipient A beginning on post-transplantation day 26 (Figure 2 of the Supplementary Appendix).14 The fever and diarrhea decreased; the pain, tenderness, and erythema in the area of the allograft diminished; and the hypoxemia, elevation of aminotransferase levels, thrombocytopenia, and coagulopathy decreased. After the patient's clinical condition had stabilized, ribavirin administration was changed to the oral route (400 mg every morning and 600 mg every evening), and it was discontinued after a renal-biopsy specimen was found to be LCMV-negative by RT-PCR and immunohistochemical staining and after serum IgM became detectable, 63 days after transplantation. During 37 days of ribavirin treatment, clinically significant hemolytic anemia developed and required the transfusion of 19 units of packed red cells. Three hundred eleven days after transplantation, the patient had stable graft function without evidence of infection, after he had restarted immunosuppressive therapy with tacrolimus, mycophenolate mofetil, and prednisolone.
Tissue Disposition
In the 2003 cluster, no additional tissues from the donor were transplanted. In the 2005 cluster, the corneas were transplanted into a 4-year-old girl and a 29-year-old woman in Algeria, neither of whom required systemic immunosuppression. Two hundred thirteen days after transplantation, neither patient had reported symptoms of infection or graft loss. The skin and liver-associated blood vessels were not transplanted.
Discussion
We describe the transmission of LCMV by solid-organ transplantation. In both clusters, disseminated infection developed in the recipients of organs from a common donor who had no clinical or laboratory evidence of infection. In the 2003 cluster, no rodent exposures could be identified, whereas the 2005 cluster was associated with recent donor exposure to an LCMV-infected pet hamster. The isolation of identical virus strains from the two kidney recipients in the 2003 cluster and from all the organ recipients and the donor household's pet hamster in the 2005 cluster indicates that in both clusters, LCMV was transmitted through organ transplantation.15
LCMV infection in humans is sporadic, is generally benign, and can be asymptomatic.4,16,17 Serologic surveys suggest that up to 5 percent of adults in the United States have been infected with LCMV.18,19 Humans become infected with LCMV by direct contact with rodents or through aerosolized droplets from rodent secretions and excretions, including urine or feces.20,21 Reports of sporadic infection have most frequently implicated the common house mouse, although outbreaks of infection have been reported after exposure to infected pet hamsters and laboratory rodents.17,20,21,22,23,24,25,26,27,28
In clinically apparent infection, the incubation period is 5 to 13 days, with subsequent fever, headache, and myalgias. Abdominal pain, diarrhea, and rash have been described.16,20,22,26,29 A second phase of illness may be seen five to nine days after convalescence, with meningitis or, rarely, encephalomyelitis, orchitis, parotitis, pneumonitis, arthritis, myocarditis, or alopecia.16,22,30 Recognized LCMV infection carries a mortality rate of less than 1 percent.3
The marked severity of LCMV-related illness in transplant recipients is probably the result of intensive immunosuppression, including T-cell depletion, coincident with direct viral inoculation by way of the transplanted organs. There are few data on the clinical behavior and outcomes of LCMV infection in immunocompromised patients. However, three patients with advanced lymphoma experimentally inoculated with the virus in a trial investigating its antitumor effect died with disseminated infection within 14 to 45 days after inoculation31 — a pattern similar to that seen in the current clusters after solid-organ transplantation.
The clinical presentation of LCMV infection in the recipients was variable and included fever, diarrhea, peri-incisional erythema and tenderness, altered mental status, and respiratory insufficiency (as noted in the table of the Supplementary Appendix). Leukopenia or leukocytosis, thrombocytopenia, coagulopathy, renal insufficiency, and progressive liver dysfunction dominated the laboratory findings. Histopathological findings in all the recipients were characterized by necrotic and occasionally hemorrhagic foci in multiple tissues, with a notable absence of inflammatory infiltrates and viral inclusions. LCMV antigens present in some tissues (e.g., the gastrointestinal tract and skin) correlated with clinical symptoms (e.g., diarrhea and erythema or pustular rash, respectively). Antigens were identified in the leptomeninges of some patients in both clusters. However, signs of meningeal inflammation, though prominent in the 2003 cluster, were generally absent in the 2005 cluster. Clinical manifestations and pathological findings in all the cases were probably altered by immunosuppression.
To our knowledge, there have been no trials of antiviral agents in human LCMV disease. Ribavirin has demonstrated efficacy in the treatment of Lassa fever and possesses in vitro activity against LCMV infection, which prompted its use in the renal-transplant recipient who survived.14,32 The role of ribavirin in this patient's improvement is unclear, since the level of immunosuppression was also considerably reduced, as it was for all four kidney recipients in the two clusters.
Rapid evaluation of organ-donor suitability is essential in transplantation to minimize the duration of ischemia and to preserve allograft function. Therefore, assays used to screen potential donors for transmissible infections must be rapid, sensitive, reproducible, and readily available to OPOs. The Food and Drug Administration has not approved any diagnostic tests for LCMV infection. Furthermore, the sensitivity of currently available assays is not adequate for routine donor screening, as demonstrated by the negative results of tests on a wide array of clinical specimens from the donors in both clusters. The use of information pertaining to recent rodent exposure for donor-suitability screening may exclude healthy donors from an already limited organ-donor pool. However, the collection of additional epidemiologic information on donors' exposures may be useful, notably for the investigation of unusual outcomes after transplantation. Zoonotic-disease transmission after transplantation is also a concern; immunosuppressed persons should take special care and limit exposure to some animals, including certain pets (additional information is available from the CDC at www.cdc.gov/healthypets).
Transplant recipients are susceptible to infection with a variety of donor-derived pathogens, including West Nile virus, Trypanosoma cruzi, rabies virus, and now LCMV.33,34,35,36 Although such infections are probably uncommon, outcomes can be fatal, and diagnosis is feasible with specialized laboratory testing. Diagnosis of LCMV infection is usually made by serologic testing, isolation of the virus from the blood or cerebrospinal fluid, or PCR testing.37,38,39 Because immunohistochemical staining revealed LCMV in multiple biopsy specimens obtained to evaluate unexplained post-transplantation symptoms in the clusters described in the current report, such testing might be beneficial in the early diagnosis of LCMV infection.
Each year, approximately 25,000 organ transplantations are performed at more than 250 transplantation centers throughout the United States.40 Allocation policies commonly result in the distribution of organs from a single donor to multiple transplantation centers. It is unlikely that either LCMV-illness cluster would have been identified without the allocation of kidneys to two recipients in whom similar symptoms simultaneously developed after undergoing transplantation at the same hospital. Similar chance clinical observations have been critical in the recognition of recent transplant-associated outbreaks of rabies and West Nile virus infection.33,35
The Organ Procurement and Transplantation Network (OPTN), which is operated by the United Network for Organ Sharing, requires transplantation centers to report certain outcomes, including allograft failure and the death of transplant recipients, in a timely manner. In April 2005, the OPTN revised its policies to require the reporting of suspected donor-transmitted medical conditions (including cancers and infections) to the procuring OPO, which is then responsible for investigating and communicating with the transplantation centers caring for the recipients of other transplants from the donor and the involved tissue and eye banks.41
Investigation of potential donor-transmitted infection requires rapid communication among physicians in multiple transplantation centers, OPOs, and public health authorities. An immediate system for tracking and disseminating pertinent patient data is needed. Until such a system can be established, clinicians must recognize that the presence of an unusual constellation of symptoms, particularly during the first few weeks after transplantation, should raise the possibility of donor-transmitted infection. Prompt notification of the OPO and public health authorities can help facilitate rapid investigation and discovery of these events.
Dr. Kotton reports having received grant support from Wyeth and the Massachusetts General Hospital Clafin Award. Dr. Fishman reports having received consulting fees or lecture fees from Astellas, Enzon, Novartis, Pfizer, and Roche. No other potential conflict of interest relevant to this article was reported.
The views expressed herein are those of the authors and not necessarily those of the Department of Health and Human Services.
We are indebted to Jennifer Betts, Kimberly Slaughter, Deborah Cannon, and Thomas Stevens for specimen processing, serologic testing, and virus isolation and characterization; to Nicole Lundstrom and Sherry Hayes for manuscript preparation; and to Donita Croft, James Kazmierczak, Mark Sotir, and Mark Wegner for investigation assistance.
* The members of the Lymphocytic Choriomeningitis Virus (LCMV) in Transplant Recipients Investigation Team are listed in the Appendix.
Source Information
From Rhode Island Hospital and Brown Medical School, Providence (S.A.F.); the Medical College of Wisconsin, Milwaukee (M.B.G.); the Divisions of Viral and Rickettsial Diseases (M.J.K., J.A.C., J.G., C.D.P., W.-J.S., B.R.E., B.P., A.L., M.J.V., T.K.S., C.S.G., P.E.R., M.M.P., M.P., C.R., R.F.H., S.T.N., T.K., S.R.Z.) and Healthcare Quality Promotion (A.S., D.B.J.), National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta; Massachusetts General Hospital and Harvard Medical School (C.N.K., F.L.D., J.A.F.) and Brigham and Women's Hospital and Harvard Medical School (F.M.M., D.L.D.) — all in Boston; the Rhode Island Department of Health, Providence (U.B.); the Massachusetts Department of Public Health, Boston (A.D.); the Wisconsin Department of Health and Family Services, Madison (J.P.D.); and the New England Organ Bank, Newton, Mass. (F.L.D.).
Drs. Ksiazek and Zaki contributed equally to this article.
Address reprint requests to Dr. Kuehnert at the Division of Viral and Rickettsial Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, 1600 Clifton Rd., Mailstop A-30, Atlanta, GA 30333, or at mkuehnert@cdc.gov.
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