Severe Non–Donor-Derived Lymphocytic Choriomeningitis Virus Infection in 2 Solid Organ Transplant Recipients

Abstract

Lymphocytic choriomeningitis virus (LCMV) is a rodent-borne Old-World mammarenavirus with global distribution [1]. The main reservoir is the common house mouse, Mus musculus, which can carry and spread the virus throughout its life [2, 3]. An estimated 5% of house mice are infected throughout the United States; other rodents, including hamsters, guinea pigs, and gerbils, are also susceptible to infection [3–5]. Humans become symptomatic 8–13 days after exposure to secretions and/or excretions of infected rodents [6].

LCMV infection in humans may exhibit a range of symptoms from mild influenzalike illness to severe neurologic disease; death is uncommon [7]. Symptomatic patients typically present with a nonspecific febrile illness including headache, malaise, myalgia, nausea, and vomiting, which may last up to a week [6, 8]. After a brief recovery period, a small subset of patients may progress to aseptic meningitis, encephalitis, or meningoencephalitis, among other neurologic complications [6, 8]. Transplacental infection can also occur, causing spontaneous abortion or congenital abnormalities [6]. LCMV can be more severe in immunosuppressed individuals, such as solid organ transplant recipients [9, 10]. The immunocompromised host can present with fever, abdominal pain, hepatitis, altered mental status, or seizures and rapidly progress to multisystem organ failure [11]. The mortality rate for LCMV infection in immunosuppressed individuals can be >70% [9, 12–14]. Clusters of LCMV infection in transplant recipients have been linked to infected organ donors [9, 12].

In this report, we present findings of an investigation in 2 solid organ transplant recipients with diagnosed LCMV infection. LCMV was initially detected in cerebrospinal fluid (CSF) from the 2 transplant recipients with unique donors by clinical metagenomic next-generation sequencing (mNGS) testing performed at University of California, San Francisco (UCSF) between April and June 2023. This multistate investigation involved clinical, epidemiologic, and diagnostic evaluations of the transplant recipients and their donors to determine the likely source of infection.

CASE REPORTS

Clinical History of Confirmed LCMV Infection in Recipient 1A

Recipient 1A (R1A) was a man in his 60s, a resident of Arizona who underwent heart transplantation for ischemic cardiomyopathy in 2023, from organ donor 1 (OD1). Cytomegalovirus and Epstein-Barr virus testing were positive for both donor and recipient. His immune suppression included tacrolimus, mycophenolate, and steroid taper. Posttransplant antimicrobial prophylaxis included itraconazole, trimethoprim-sulfamethoxazole, and valganciclovir. He was readmitted 10 days later with watery diarrhea. Stool testing was positive for Cryptosporidium, and he received nitazoxanide.

R1A was hospitalized a month later for failure to thrive and Escherichia coli urinary tract infection. Mental status, fever and cytopenia developed altered. Initial lumbar puncture revealed 14 white blood cells (WBCs) (0 neutrophils, 88% lymphocytes, and 12% monocytes), a protein level of 366 g/L, and a glucose level of 55 mmol/L. Clinical diagnostic testing revealed negative results for Cryptococcus, Coccidioides, West Nile virus, Toxoplasma, Acanthamoeba, Balamuthia, Naegleria, bacteria, fungi, and mycobacteria by culture and Mycobacterium tuberculosis by polymerase chain reaction (PCR). A CSF BioFire FilmArray meningitis/encephalitis panel was also negative for 7 viruses, 6 bacteria, and cryptococcus. The patient’s mental status continued to worsen, and he experienced stool incontinence. Additional test results for Strongyloides, brucellosis, histoplasmosis, and CSF autoimmune encephalitis panel were negative. Magnetic resonance imaging of the brain with gadolinium revealed a T2 signal enhancement in the rostral midbrain, indicative of rhombencephalitis. Empiric therapy for listeriosis was initiated with ampicillin and gentamicin. Tacrolimus was discontinued given neurotoxicity concerns, and cyclosporin was initiated. Bronchoalveolar lavage was positive for Aspergillus antigen; itraconazole prophylaxis was switched to voriconazole.

A second lumbar puncture revealed 20 WBCs (88% lymphocytes and 12% monocytes), 227 red blood cells, a protein level of 1573 g/L and a glucose level of 101 mmol/L. Extensive repeated testing of this CSF sample returned negative results, including CSF studies for Lyme disease, broad-range bacterial PCR, Bartonella PCR, Brucella culture, Tropheryma whipplei, Histoplasma, and Blastomyces by PCR and Histoplasma CSF antigen. A CSF sample was sent to UCSF for mNGS, which detected LCMV. Cytopenia developed, along with significant liver enzyme elevation, serum ferritin levels up to 34 000 μg/dL and a soluble interleukin 2 receptor subunit α level of 6192 pg/mL (reference, 214–1910), confirming hemophagocytic lymphangiohistiocytosis. The patient received eculizumab and 3 doses of intravenous immunoglobin and methylprednisolone, followed by dexamethasone. High grade vancomycin-resistant Enterococcus faecium bacteremia was treated with intravenous daptomycin.

The condition of R1A continued to deteriorate, with progressive acidosis, increased ventilatory requirement, mental obtundation, and hypotension. Clinical status and prognosis were discussed with the family, who opted for comfort measures followed by withdrawal of care. The patient died.

Clinical History of Confirmed LCMV Infection in Recipient 2A

Recipient 2A (R2A) was a man in his 50s with end-stage renal disease due to diabetes mellitus and hypertension. He received a severe acute respiratory syndrome–positive deceased donor kidney transplant (donor's left kidney) in 2023 from organ donor 2 (OD2).

Induction therapy included alemtuzumab and rapid steroid withdrawal, followed by maintenance immune suppression with tacrolimus and mycophenolate. The patient was donor positive/recipient negative for cytomegalovirus and recipient positive for Epstein-Barr virus. Antimicrobial prophylaxis included valganciclovir, trimethoprim sulfamethoxazole, and fluconazole. The patient's sister reported that 1 week before transplantation, R2A experienced worse-than-usual flulike symptoms (compared with similar mild postdialysis symptoms).

R2A was hospitalized 2 weeks after receiving the transplant with nausea, vomiting, diarrhea, and acute kidney injury. Upper gastrointestinal endoscopy revealed grade A esophagitis. The stool BioFire FilmArray Gastrointestinal panel was negative. He was discharged within 48 hours but was readmitted 4 days later with ongoing nausea, vomiting, and leukopenia. Mycophenolate was switched to azathioprine. Computed tomography of the abdomen and pelvis demonstrated gastroenteritis, and flexible sigmoidoscopy was unrevealing. The patient’s diarrhea resolved within 4 days.

R2A was hospitalized again 2 days later with diarrhea, progressive confusion, agitation, and headache. He was unable to follow commands and had significant tremors in all extremities. Findings of brain magnetic resonance imaging with gadolinium were unremarkable. He subsequently became more somnolent with myoclonic jerks.

Initial lumbar puncture revealed 7 WBCs (35% lymphocytes and 61% monocytes), 0 red blood cells, a protein level of 563 g/L, and a glucose level 107 mmol/L. Subsequent lumbar punctures yielded 29 WBCs (80% lymphocytes and 20% monocytes) and protein and glucose levels of 667 g/L and 89 mmol/L, respectively, and 79 WBCs (87% lymphocytes and 8% monocytes) and protein and glucose levels of 442 g/L and glucose 75 mmol/g. Results of molecular and serologic CSF testing were negative; this testing included a BioFire FilmArray meningitis/encephalitis panel, West Nile virus reverse-transcription (RT) PCR and immunoglobulin (Ig) G/IgM enzyme-linked immunosorbent assay (ELISA), examination for protozoa, Toxoplasma RT-PCR, Eastern and Western equine encephalitis virus RT-PCR, St Louis encephalitis virus RT-PCR, JC virus RT-PCR, 2-tiered serology for Lyme disease, Coccidioides testing (antibodies and RT-PCR), RT-PCR for Histoplasma, Blastomyces, and M tuberculosis complex, and CSF autoimmune encephalitis panel. CSF bacterial, fungal, and mycobacterial smears and cultures were also negative. The patient’s empiric antimicrobial therapy was discontinued following negative test results.

A CSF sample sent to UCSF for mNGS testing returned positive for LCMV. Based on anecdotal data from the literature [9, 12–15], R2A received ribavirin, 600 mg orally 3 times a day for 10 days, and intravenous immunoglobulin, 500 mg/kg daily for 5 days. He had a prolonged hospital stay (10 weeks) and was then discharged to a skilled nursing facility. He continued to have intermittent confusion but was able to answer simple questions. He was discharged to home 6 months later with physical, occupational, and speech therapy. He had to relearn how to walk, talk, and eat, and cannot live independently due to ataxic gait and memory issues.

MATERIALS AND METHODS

Initial LCMV Laboratory Diagnosis by Clinical Metagenomics Sequencing

The UCSF CSF mNGS assay is a Clinical Laboratory Improvement Amendments–certified, clinically validated test conducted by the UCSF Clinical Microbiology Laboratory [16]. The mNGS approach leverages shotgun sequencing of DNA and RNA to enable agnostic detection of pathogens without the use of targeted primers or probes [17]. After enrichment using a kit that depletes human host methylated DNA, total nucleic acids were extracted, and both RNA/cDNA and DNA libraries were constructed using Nextera library construction (Illumina) with 2 rounds of PCR. Final libraries were sequenced on the Illumina NextSeq 550 Dx using 150–base pair single-end, dual-indexed sequencing, targeting 5–20 million sequences per library. The mNGS data were analyzed for pathogens using the SURPI+ bioinformatics pipeline (version 1.0.7-build.4). Near-complete LCMV genomes consisting of the L (large) and S (short) segments were assembled from the mNGS data by mapping trimmed LCMV reads to the reference sequences (NC_077807) and generating consensus segments using Geneious software (version 11.1.5) [18]. Pairwise nucleotide identities were 97.6% (S segment) and 97.7% (L segment) for R1A and 98.1% (S segment) and 98.2% (L segment) for R2A.

Epidemiologic Investigations

Transplant centers are required to report suspected donor derived infections to the Organ Procurement and Transplantation Network/Disease Transmission Advisory Committee (DTAC), which monitors the transmission of disease through organ transplantation. The DTAC examines potential transmission cases reported to the Organ Procurement and Transplantation Network to confirm transmissions. A subset of DTAC cases involving pathogens of special interest is referred to the Centers for Disease Control and Prevention (CDC) for additional investigation [19]. Since the present case included a pathogen of special interest, the CDC coordinated the investigation in collaboration with state and local health department(s), the organ procurement organizations, and transplant centers. All organ donor and recipient medical records were reviewed. Specimens from the organ donor and transplant recipients were provided for further testing and analysis. Transplant centers provided an up-to-date clinical history of the recipients.

Maricopa County Department of Public Health (MCDPH) interviewed the medical power of attorney for each patient regarding rodent exposure history. Interviews with R1A's wife and R2A's sister determined the timing and extent of exposure, documented in the Arizona state surveillance database.

LCMV Confirmatory Testing

Molecular Diagnostic Testing

Samples underwent inactivation using the MagMAX pathogen RNA/DNA kit (ThermoFisher) and were extracted on the KingFisher Duo Prime platform (ThermoFisher). The LCMV real-time RT-PCR (rRT-PCR) assay was developed using all available genomes in the National Center for Biotechnology Information (as of March 2021) to detect strains known to cause infections in humans [10]. Samples were tested using the Luna Probe One-Step rRT-PCR 4× Mix with UDG kit (New England Biolabs) following recommended cycling conditions on a Bio-Rad CFX system; results were analyzed using Bio-Rad CFX manager software (Bio-Rad).

Serology

LCMV IgM and IgG were determined using 2 laboratory-developed ELISA, as described elsewhere [7]. The anti-LCMV IgM ELISA plates were coated with goat anti-human IgM antibodies, and then patient samples were added and incubated. LCMV-Armstrong and negative antigens then followed. Mouse anti-LCMV hyperimmune ascetic fluid and conjugated goat anti-mouse IgG were subsequently added as separate steps and incubated. For the IgG ELISA, plates were initially coated with LCMV antigen, with the subsequent addition of samples. Mouse anti-human IgG was followed with another incubation period. Positive samples were identified if the titer exceeded 1:400 and the sumOD (sum of adjusted optical density) of the entire sample was greater than 0.45 (IgM) or 0.95 (IgG).

Next-Generation Sequencing, Bioinformatics, and Phylogenetic Analysis

RNA was treated and libraries were prepared using the NEBNext Ultra II Directional RNA Library prep kit (New England Biolabs). Libraries were sequenced on an Illumina MiSeq instrument with a 300-cycle paired-end sequencing (PE) cartridge. Reads were trimmed using prinseq-lite (-min_qual_mean 25 -trim_qual_right 20 -min_len 50 -out_good; version 0.20.3), and contigs were built by mapping to the human genome with bwa (version 0.7.17), taking unmapped reads and building contigs with SPAdes (-k auto; version 3.15.3). The L-segment genomes were built by mapping reads to scaffold genomes originally produced by UCSF and replacing regions with missing coverage using NC_077806. The S-segment genome was built by mapping trimmed reads and contigs to the LCMV reference sequence (NC_077807). Consensus L and S segments were called using Geneious Prime software (0% threshold—majority alleles). Phylogenetic trees were constructed using available full-length LCMV genomes from GenBank (obtained December 2023) and in-house scripts (https://github.com/evk3/hantavirus_US_distribution) and aligned using MAFFT software (version 1.5.0), and trees were built using Raxml software (version 8.2.12). Phylogenetic trees were constructed using R software (version 4.1.3) and the ggtree library (version 3.2.0).

RESULTS

Epidemiologic Investigation Findings

Findings in R1A

The MCDPH performed a telephone interview with R1A's wife (who had medical power of attorney). She reported a 3-year history of rodent exposure within the home following fumigation of a neighboring home. She described small house mice and reported frequently finding rodent droppings in the home. Most recently, she reported seeing 2 mice and feces the month before R1A's transplant. She also revealed that R1A was exposed to rodents at her sister's home in Mexico a month before the transplant, where they noticed droppings and assisted with cleaning. She denied any contact with hamsters, guinea pigs, or other rodents in the 6 months before the interview (Figure 1).

Figure 1.

Timeline of symptom onset and laboratory testing in 2 unrelated lymphocytic choriomeningitis virus infections in solid organ transplant recipients. Plus and minus signs represent positive and negative results, respectively. Abbreviations: CSF, cerebrospinal fluid; Ig, immunoglobulin; mNGS, metagenomic next-generation sequencing; OD1, organ donor 1; OD2, organ donor 2; PCR, polymerase chain reaction; R1A, recipient 1A; R2A, recipient 2A.

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Findings in R2A

The MCDPH performed a telephone interview with R2A's sister (who had medical power of attorney). She confirmed that the patient had exposure to house mice and droppings within his New Mexico residence 3 months before his transplant, including rodent droppings in his bedroom. These were sprayed with bleach and removed by his mother, and he reportedly returned to the room the following day. The sister was unaware of any other rodent or rodent dropping exposures and denied any outdoor recreational activities or exposure to pet or wild rodents.

Initial LCMV Laboratory Diagnosis

mNGS was performed on CSF samples from patients R1A and R2A, collected during hospitalization. A high number of LCMV reads were identified in CSF specimens from R1A and R2A and, near-complete LCMV consensus genomes were assembled. Other viruses, bacteria, fungi, and parasites were not detected with mNGS.

LCMV Diagnostic Testing in Solid Organ Donors and Recipients

Confirmatory Testing of Samples From R1A

After the positive LCMV result with mNGS, the CDC received specimens, including the 2 CSF samples forwarded by USCF (collected on day 58 and day 65 after transplantation), a CSF sample (collected on day 61), a pretransplant serum sample (collected the day before transplant [day −1]), and a whole-blood (ethylenediaminetetraacetic acid [EDTA]) sample (collected on day 87). The CSF samples were positive, with titers of 1:400 and 1:1600 for anti-LCMV IgM antibodies. The pretransplant serum sample was negative for LCMV. A blood sample collected on day 87 was positive at the CDC for LCMV by rRT-PCR and mNGS and negative by IgM and IgG testing (Table 1).

Testing of Sample From Donor OD1

The Viral Special Pathogens Branch (VSPB) laboratory at the CDC received a serum sample from OD1 from the day of organ harvest (day 0). This sample was negative by rRT-PCR and IgM and IgG testing (Table 1 and Figure 1).

Testing of Samples From Other Recipients of OD1 (R1B and R1C)

Due to concerns about the potential transmission of LCMV from OD1 to R1A, samples were requested from the other 2 recipients associated with this donor, who included R1B (left kidney and liver recipient) and R1C (right kidney recipient). LCMV testing was not possible for R1B as samples were not collected before the recipient's non–LCMV-related death within 3 weeks after transplantation. A whole-blood (EDTA) sample from recipient R1C (collected on day 163) revealed a negative rRT-PCR result for LCMV. Six months after transplantation, the IgM result was positive, with a low titer of 1:400; there was no evidence of IgG seroconversion. Serology testing (performed twice) consistently revealed isolated IgM positivity without IgG seroconversion. R1C is alive without evidence of LCMV infection.

Confirmatory Testing of Samples From R2A

After the positive LCMV result reported by USCF for R2A, the VSPB laboratory received specimens from this patient, including the CSF sample tested by USCF (collected on day 45), a pretransplant serum (collected on day −3), and a follow-up serum sample (collected on day 57). The CSF sample and the pretransplant serum sample were negative for LCMV by rRT-PCR, IgM, and IgG ELISA testing. The serum sample (collected on day 57) had a 1:1600 LCMV IgM titer but was negative by IgG and rRT-PCR (Figure 1).

Testing of Samples From Donor OD2

A pretransplant serum sample collected from OD2 on the date of death was negative by rRT-PCR, IgM and IgG testing (Table 1).

Testing of Samples From Other Recipients of OD2 (R2B and R2C)

Other recipients from OD2 included R2B (right kidney recipient) and R2C (liver recipient). Blood samples collected from recipient R2B on day 67 and from recipient R2C on day 63 were negative for LCMV by all assays (rRT-PCR and IgM and IgG ELISA) (Table 1). Both R2B and R2C are alive without evidence of LCMV infection.

Genomic Analysis

Genetic relationships of LCMV sequences from R1A and R2A were determined by whole-genome sequencing of the L and S segments (Figure 2 and Supplementary Figure 1). L-segment sequences from R1A and R2B were not closely related and were located within distinct clades, demonstrating that viral infections in these individuals likely originated from 2 different and unrelated sources. LCMV sequences from R1A whole-blood (PP465627) and CSF (PP445004) specimens were identical and were most closely related to an LCMV sequence from a mouse collected in France in 2009 (blue highlight in Figure 2). Sequences derived from patient R2A were most closely related to a sequence from an individual from Massachusetts in the United States in 2008 (yellow highlight in Figure 2). The S segments for R1A and R2A exhibited genetic relationships similar to those for the L segments (Supplementary Figure 1). Overall, LCMV L- and S-segment sequences derived from historic and current US patients and rodents exhibited a wide range of viral diversity (Figure 2 and Supplementary Figure 1; orange vs green sequence names).

Figure 2.

Phylogenetic tree for lymphocytic choriomeningitis virus (LCMV) long segment of samples from the 2 independent US transplant recipients who contracted LCMV. The phylogenetic history of the LCMV short segment was inferred using maximum-likelihood estimation (GTR + I). Samples from R1A are highlighted in blue, and the sample from R2A is highlighted in yellow. Bootstrap support (n = 1000) is indicated at internal nodes in red, and scale bars are in units of substitutions per site. Phylogenetic trees were generated using a nucleotide alignment of LCMV long segments. GenBank accession numbers are provided. An additional phylogenetic tree constructed from the LCMV short segment is included in the Supplementary materials.

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DISCUSSION

The majority of published cases of adult human LCMV infection in immunocompromised hosts pertain to donor-derived transmission. An initial report detailed 2 donor-derived clusters from 2003 and 2005 in which LCMV was transmitted to all 8 organ recipients; 7 of 8 died [9]. A member of the 2005 donor household had brought home a pet hamster 3 weeks before the donor died; the hamster tested positive for LCMV. This hamster was traced to a pet store and subsequently to a parent facility in another state. All infected rodents and the transplant recipients had identical LCMV sequences by phylogenetic analysis [4]. The next report from 2008 reported severe multisystem illness, leading to death in 2 kidney recipients from a common donor who was hospitalized with fever, seizures, and confusion and had CSF pleocytosis before death [13]. In 2011, a common donor transmitted LCMV to all 4 organ recipients; the liver and right kidney recipients survived [12].

The most recent publication details LCMV donor transmission to 3 recipients in 2013 [14]. Both kidney recipients survived, while the liver recipient died. Overall, of these 5 donor-derived LCMV clusters in the United States, 5 donors transmitted LCMV to 17 recipients, with 12 deaths, a 71% mortality rate. Patients presented within the first few weeks after transplantation with rapidly progressive multisystem illness and prominent neurologic manifestations. The only published case of non–donor-derived LCMV infection was in a kidney transplant recipient who apparently acquired LCMV meningoencephalitis after cleaning his basement with mouse excreta 5 years after transplantation [20]. He was alive at the time of discharge and was lost to follow up.

Similar to the experience in transplant recipients, human LCMV infection in the general population remains uncommon. LCMV has been associated with severe congenital infection causing microcephaly, hydrocephalus, periventricular calcifications, cerebellar hypoplasia, visual impairment, and other abnormalities [6]. Spontaneous abortion and fetal death can ensue. Several human outbreaks related to infected pet hamsters and laboratory rodents have been reported [9]. In a recent study from Hungary, investigators evaluated CSF samples for LCMV from patients with central nervous system infections of unknown etiology over a 12-year period [21]. Of samples from 74 patients, only 2 samples were positive for LCMV by RT-PCR and sequencing. One was from a 5-year-old who had been bitten by a hamster, and the second was from an elderly man with known dementia. Investigators from Iraq screened cases of febrile neurologic infections for LCMV [22]. Two serum and 2 CSF samples tested positive for LCMV with phylogenetically distinct strains, for a seroprevalence of 8.8%. A man with well-controlled human immunodeficiency virus infection made full recovery from LCMV meningoencephalitis acquired after cleaning mouse feces and urine 2 weeks before symptom onset [10].

In this report, presumptive LCMV infection was identified by mNGS in 2 solid organ transplant recipients from separate donors. LCMV-specific serologic and molecular testing at the CDC confirmed LCMV infection in both cases. Collaborative laboratory and epidemiologic investigations involving multiple states and investigating partners failed to confirm donor-derived transmission. On the contrary, both recipients with LCMV infection were noted to have significant rodent exposure leading up to transplantation.

Samples were obtained from the 2 donors and pretransplant and/or posttransplant specimens for 2 of the 3 organ transplant recipients associated with OD1 and all 3 recipients associated with OD2. CSF samples from R1A and R2A were initially sent to UCSF for mNGS and subsequently forwarded to the CDC for rRT-PCR and anti-LCMV IgM and IgG antibody testing. Limited CSF samples received by the CDC were aliquots that had undergone 2 freeze-thaw cycles at UCSF, which likely contributed to negative rRT-PCR results due to viral RNA degradation. It is worth noting that mNGS was not repeated on CSF samples at the CDC. However, results of whole-blood mNGS and rRT-PCR testing for R1A on day 87 at the CDC were positive. Both CSF and blood mNGS testing for R1A at different laboratories yielded similar LCMV sequences.

In R1C, a serum sample collected 6 months after transplantation resulted in a weak IgM response of 1:400 and negative IgG result. A clinical illness compatible with LCMV infection did not develop in this individual. Isolated anti-LCMV IgM positivity without anti-LCMV IgG seroconversion 6 months after transplantation suggests probable cross-reactivity of the IgM ELISA. This weak cross-reactivity is unreported with the CDC's anti-LCMV IgM assay, warranting further investigation.

Laboratory findings in the remaining individuals, including findings of mNGS, rRT-PCR, and anti-LCMV IgM and IgG antibody testing, did not reveal evidence of LCMV infection in the donor or the other tested recipients of the donor’s solid organs. These results, along with epidemiologic investigations, suggest that LCMV infections in R1A and R2A were most likely due to pretransplant rodent exposure, rather than donor-derived infections. However, the possibility of an unknown rodent exposure after transplantation cannot be entirely ruled out.

Surveillance and diagnostic detection of LCMV is challenging for multiple reasons. Symptoms of LCMV infection are nonspecific and resemble those of common viral infections; most cases are asymptomatic. Laboratory diagnosis of LCMB can be challenging due to the lack of commercial assays for serology and molecular detection of LCMV. An approach combining serologic and molecular testing is one strategy for enhancing detection of LCMV infection. Pathogen-agnostic testing with mNGS can be helpful in the diagnosis of rare infections such as LCMV for which targeted assays are lacking [17] but is generally confined to specialized reference laboratories. LCMV-associated disease is not a nationally notifiable illness, and, as a result, CDC receives only sporadic domestic case reports.

The findings from these investigations underscore the importance of raising awareness among transplant recipients and healthcare professionals about LCMV and the potential risks associated with rodent and pet hamster exposure before or after solid organ transplantation. Severe and progressive systemic illness with features of meningoencephalitis in the early posttransplant setting should include consideration of LCMV infection. It is of utmost importance to enhance clinician and transplant recipient awareness about LCMV and promote public health case reporting. No-cost LCMV testing is available through the CDC's VSPB, and reporting of LCMV cases to the CDC is encouraged to enhance surveillance and to elucidate the true burden and distribution of this important but underappreciated pathogen in the United States.

Supplementary Data

Supplementary materials are available at Open Forum Infectious Diseases online. Consisting of data provided by the authors to benefit the reader, the posted materials are not copyedited and are the sole responsibility of the authors, so questions or comments should be addressed to the corresponding author.

Acknowledgments

The authors thank Brian W. Hardaway in the Division of Heart Failure and Transplant, Mayo Clinic, Phoenix, Arizona, Curtis Fritz in the California Department of Public Health, Sacramento, California, and Monica Brady in the VSPB laboratory, Atlanta, Georgia.

Author contributions. H. R. V., L. E. S., K. L. S., K. S. S., and C. M. C. conceived the study and drafted the manuscript. L. E. S., S. W., D. C., I. K., M. M. B., and C. Y. C. conducted the virologic assays and sample processing. M. J. C., P. A., I. R., M. K., N. G., K. Z., C. A., E. S., V. S., A. F., S. V. B., N. S., B. A. A., H. A. K., C. C. J., T. E. G., A. J., M. F. G., C. Y. C., and H. R. V. provided expert epidemiologic and clinical consultation. J. M. M., T. S., and J. D. K. provided guidance on all aspects of the study. All authors assisted with data analysis, critically revised the manuscript for important intellectual content, and approved the final version of the manuscript.

Disclaimer. The findings and conclusions in this report are those of the authors and do not necessarily represent the official position of the Centers for Disease Control and Prevention.

Financial support. This work was supported by CDC Emerging Infectious Disease funds.

References

Author notes