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REVIEW ARTICLE
Laboratory diagnosis of CNS infections in children due to
emerging and re-emerging neurotropic viruses
Benjamin M. Liu
1,2,3,4,5,6
✉, Sarah B. Mulkey
2,5,7,8,9
, Joseph M. Campos
1,2,3,4
and Roberta L. DeBiasi
2,4,5,10
✉
© The Author(s), under exclusive licence to the International Pediatric Research Foundation, Inc 2023
Recent decades have witnessed the emergence and re-emergence of numerous medically important viruses that cause central
nervous system (CNS) infections in children, e.g., Zika, West Nile, and enterovirus/parechovirus. Children with immature immune
defenses and blood–brain barrier are more vulnerable to viral CNS infections and meningitis than adults. Viral invasion into the CNS
causes meningitis, encephalitis, brain imaging abnormalities, and long-term neurodevelopmental sequelae. Rapid and accurate
detection of neurotropic viral infections is essential for diagnosing CNS diseases and setting up an appropriate patient
management plan. The addition of new molecular assays and next-generation sequencing has broadened diagnostic capabilities
for identifying infectious meningitis/encephalitis. However, the expansion of test menu has led to new challenges in selecting
appropriate tests and making accurate interpretation of test results. There are unmet gaps in development of rapid, sensitive and
specific molecular assays for a growing list of emerging and re-emerging neurotropic viruses. Herein we will discuss the advances
and challenges in the laboratory diagnosis of viral CNS infections in children. This review not only sheds light on selection and
interpretation of a suitable diagnostic test for emerging/re-emerging neurotropic viruses, but also calls for more research on
development and clinical utility study of novel molecular assays.
Pediatric Research; https://doi.org/10.1038/s41390-023-02930-6
IMPACT:
●Children with immature immune defenses and blood–brain barrier, especially neonates and infants, are more vulnerable to viral
central nervous system infections and meningitis than adults.
●The addition of new molecular assays and next-generation sequencing has broadened diagnostic capabilities for identifying
infectious meningitis and encephalitis.
●There are unmet gaps in the development of rapid, sensitive and specific molecular assays for a growing list of emerging and
re-emerging neurotropic viruses.
INTRODUCTION
Viral infections are responsible for a large proportion of central
nervous system (CNS) infections in pediatric patients and can be
life-threatening.
1–3
Viral invasion into the CNS, especially in fetuses
and neonates, can cause meningitis, encephalitis, seizures, brain
imaging abnormalities, and long-term neurodevelopmental
sequelae, depending upon the timing of infection and other
factors.
1–6
Figure 1summarizes emerging/re-emerging, neurotro-
pic viruses that have caused outbreaks or epidemics since 1999
(www.who.int;www.cdc.gov;www.ecdc.europa.eu), including
Nipah virus, dengue virus (DENV), West Nile virus (WNV),
Chikungunya virus (CHIKV), enteroviruses (EV) D68, Hendra Virus,
Zika virus (ZIKV), yellow fever virus (YFV), tick-borne encephalitis
virus, human parechoviruses (HPeV), and Japanese encephalitis
virus (JEV).
1–9
Most of these viral infections are zoonotic diseases
transmitted via animals and vectors, except EV D68 and HPeV that
are transmitted through fecal-oral and respiratory routes. Besides,
congenital ZIKV infections are responsible for fetal brain
malformations following maternal infection during pregnancy,
especially in the first trimester.
5,6
The scope of the review will
focus on EVs, HPeV and arboviruses (e.g., ZIKV), which are global
public health concerns.
The incidence of emerging/re-emerging, neurotropic viruses is
on the rise,
4
which may be attributed to the following factors: (1)
novel molecular diagnostic tools have a higher sensitivity to
detect more positive cases; (2) geographic distribution of the viral
pathogens and their vectors has expanded. For example,
significant rise in international travel, shift in agriculture practices,
climate change, and growth of human population lead to
unprecedented spread of numerous arboviruses;
4
(3) increased
Received: 6 July 2023 Revised: 10 October 2023 Accepted: 5 November 2023
1
Division of Pathology and Laboratory Medicine, Children’s National Hospital, Washington, DC, USA.
2
Department of Pediatrics, The George Washington University School of
Medicine and Health Sciences, Washington, DC, USA.
3
Department of Pathology, The George Washington University School of Medicine and Health Sciences, Washington, DC,
USA.
4
Department of Microbiology, Immunology and Tropical Medicine, The George Washington University School of Medicine and Health Sciences, Washington, DC, USA.
5
Children’s National Research Institute, Washington, DC, USA.
6
The District of Columbia Center for AIDS Research, Washington, DC, USA.
7
Prenatal Pediatrics Institute, Children’s
National Hospital, Washington, DC, USA.
8
Division of Neurology, Children’s National Hospital, Washington, DC, USA.
9
Department of Neurology, The George Washington
University School of Medicine and Health Sciences, Washington, DC, USA.
10
Division of Pediatric Infectious Diseases, Children’s National Hospital, Washington, DC, USA.
✉email: bliu1@childrensnational.org; rdebiasi@childrensnational.org
www.nature.com/pr
1234567890();,:
immunocompromised and transplant population face risk of CNS
infection by established viruses that can cause neurological
syndromes; (4) decreased immunization rates, lack of surveillance,
and mass migration are responsible for outbreaks of vaccine-
preventable diseases, e.g., wild poliovirus type 1 outbreaks in
Pakistan and Mozambique since 2019 (https://www.who.int/
emergencies/disease-outbreak-news/item/2022-DON395).
10
Rapid and accurate detection of neurotropic viral infections is
important for setting up an appropriate patient management plan,
preventing inappropriate and costly treatments, counseling
patients and family members, and carrying out public health
interventions.
2,11,12
The addition of new molecular assays has
broadened diagnostic capabilities for identifying infectious menin-
gitis and encephalitis.
12
However, the expansion of test menu has
led to new challenges in selecting an appropriate test and making
accurate interpretation of test results. There are unmet gaps in
development of novel molecular assays for a growing list of
emerging and re-emerging neurotropic viruses. Herein we will
discuss the advances and challenges in laboratory diagnosis of viral
CNS infections. This review not only sheds light on selection and
interpretation of a suitable diagnostic test for emerging/re-
emerging neurotropic viruses, but also calls for more research on
development and clinical utility study of novel molecular assays.
SPECIAL CONSIDERATIONS FOR PEDIATRIC PATIENTS
Children, especially neonates and infants, are more vulnerable to
viral CNS infections and meningitis than adults. This is likely due to
their unique anatomical and physiological features (Fig. 2),
including, but not limited to, fast cell division, high basal
metabolism and respiratory rates, thin skin, large body surface
area, immature immune system and blood–brain barrier at an
early age (4 months) as well as distinct microbiota/virus
colonization in non-CNS body sites, e.g., respiratory and GI
systems.
13
Thin skin and large body surface area may enhance
the risk of infection, especially arboviral infection. The immature
blood-brain barrier and immune system may lead to vulnerability
to CNS infections (Fig. 2). There are some special considerations
one should include in the evaluation of pediatric patients,
especially sample collection for small children. For example, small
children have lower overall blood volume than adults, and can
develop anemia after collection of a high-volume blood for
diagnostic testing.
13
CONVENTIONAL DIAGNOSTIC MICROBIOLOGY/VIROLOGY
METHODS
Viral cultures
There are some conventional diagnostic microbiology/virology
methods that are useful in the diagnosis of CNS infections. One
example is traditional viral cultures (Fig. 3). To carry out this assay,
viruses-containing samples are used to inoculate cell lines, e.g.,
rhesus monkey kidney, African green monkey kidney, A549, and
MRC-5, which may yield cytopathic effects (CPE) after incubation
at 37 °C (up to 30 days).
14–16
An improvement of the CPE-based
viral culture is shell vial culture, a centrifuge enhanced tissue
culture assay followed by rapid detection of viral antigen using
monoclonal antibodies.
17
There are some advantages for viral cultures and shell vial
culture (Fig. 3). First, viral cultures are considered the gold
Dengue Chikungunya Zika Nipah virus
Nipah virus West Nile virus
Hendra virus
Enterovirus D68 Yellow fever Tick-borne
encephalitis
West Nile virus
Human Parechovirus
Japanese encephalitis
2000
There has been a dramatic
increase in the incidence
of dengue around the
world since 2000
In 2005–2006, a major
outbreak was reported
in the Indian Ocean
An epidemic of Zika
fever originated in Brazil
and spread to other
countries in 2015–2016
2018 Nipah virus
outbreak in Kerala
Clusters of parechovirus
CNS infections were reported
in young infants in the USA
Outbreaks of Japanese
encephalitis were reported
in Australia
Outbreaks of West
Nile were reported
in EU/EEA countries
and the USA
There were 25
EU/EEA countries
that reported 3411
cases of tick-
borne encephalitis
An outbreak of
hemorrhagic yellow
fever was reported
in Brazil
A total of fifty outbreaks of
Hendra virus were reported
in Australia
A nationwide outbreak of
respiratory illness due to
EV-D68 was reported in the
USA between August and
November in 2014
The first largest West
Nile neuroinvasive
disease in the USA
Was first discovered
during an outbreak
in the Sungai Nipah,
a village in Malaysia
1999 2002
2005
2014
2015
2016
2018
2019
2022
2023
Fig. 1 Timeline of emerging and re-emerging viral diseases that have caused central nervous system infections in the past two decades.
The year and geographic area of the indicated outbreaks or epidemics of viral diseases are shown (https://www.cdc.gov/westnile/statsmaps/
current-season-data.html;https://www.who.int/news-room/fact-sheets/detail/nipah-virus;https://www.who.int/news-room/fact-sheets/detail/
dengue-and-severe-dengue;https://www.cdc.gov/non-polio-enterovirus/about/ev-d68.html;https://www.who.int/health-topics/hendra-virus-
disease#tab=tab_1;https://www.ecdc.europa.eu/en/tick-borne-encephalitis/surveillance-and-disease-data/epidemiology;https://
www.who.int/emergencies/disease-outbreak-news/item/2022-DON365;https://www.cdc.gov/westnile/index.html;https://
www.ecdc.europa.eu/en/west-nile-fever/surveillance-and-disease-data/disease-data-ecdc).
7,8,14,54,55
EU European Union, EEA European
Economic Area.
B.M. Liu et al.
2
Pediatric Research
standard for the detection of viable viruses. Viral isolates from
clinical samples allows for further analysis of its virulence and
antiviral drug resistance. Second, this method can detect
cultivable neurotropic viruses from a broad range of sample
sources, including cerebrospinal fluid (CSF), tissue specimens and
acute-phase serum. For example, arboviruses such as Colorado
tick fever, DENV, YFV, and ZIKV are more likely to be detected
using culture in the early clinical course, probably due to delayed
immune response and decent duration of viremia.
18
Third, shell
vial culture has good specificity and showed relatively higher
sensitivity than indirect immunofluorescence techniques that
detect cell associated virus antigens.
17
Shell vial culture is
demonstrated as a rapid and efficient method aiding in the
detection of JEV, WNV and DENV-2 from CSF specimens.
17
However, viral cultures have been replaced by molecular tests in
most of the clinical virology labs in the U.S. due to the following
limitations (Fig. 3). First, viral cultures are limited to detecting
cultivable viruses. For example, some serotypes of EVs cannot
grow in viral culture and several coxsackievirus group A strains
require mouse inoculation for detection. Therefore, viral culture
for EVs has a sensitivity of approximately 65–75%.
19
Second, for
some arboviruses (except those discussed above), viral culture was
found to have low yield even in the early phase of illness because
these viruses have a relatively short duration of viremia.
18
Third,
compared with molecular methods, the turnaround time and
sensitivity of CSF viral cultures are suboptimal, which does not
enable it to yield a rapid and reliable diagnosis required for
optimum clinical care of patients with CNS viral infections.
20
Last,
viral cultures require technical expertise of well-trained
technologists.
Detection of viral antigens
During active viral replication, viral antigens can be detected in
blood and fixed tissue by immunological methods, such as
immunohistochemical (IHC) staining and enzyme-linked immuno-
sorbent assays (ELISA) (Fig. 3).
14,18,21–25
Detection of viral antigen
from CSF is rarely used for arbovirus, EV or HPeV, though there are
some studies in which influenza virus antigen was detected in
CSF.
25
In contrast, blood samples have been used to detect DENV
nonstructural protein 1 (NS1) antigen by viral antigen ELISA.
14,18
For instance, Platelia DENV NS1 Ag (Bio-Rad) is a one-step
“sandwich”format microplate enzyme immunoassay for the
qualitative or semi-quantitative detection of DENV NS1 antigen
in human serum or plasma (Fig. 3). This assay has high sensitivity
(87.1–100% for different serotypes) and specificity (100%) per
package insert. With short turnaround time and relatively low cost,
this assay aids in early diagnosis within the first 1–7 days after
onset of illness. However, DENV NS1 antigen assays may have
some limitations (Fig. 3). First, positive results of some assays do
not provide serotypes while some may have the capacity to do
so.
14
Second, negative test results cannot rule out infection. Last,
some assays may be less sensitive in secondary DENV infections.
14
In addition, IHC enables visualization of infected cells with viral
antigen expression in situ (Fig. 3). Viral antigens can be detected
by performing IHC in brains of fatal cases infected with central
European tick-borne encephalitis virus.
21
Gelpi et al. found that
high levels of viral antigens can be found in the brain in varied
clinical durations ranging from 4 to 35 days from the onset of
infection.
21
Besides, viral antigens have been detected in the cell
bodies of neurons among patients infected with St. Louis
Encephalitis virus (SLEV).
22
Similarly, viral antigen of JEV can be
found in neurons of the cerebral cortex, thalamus, and brain
stem.
24
However, there are also some limitations for antigen
detection by IHC (Fig. 3). First, IHC may have relatively low
sensitivity compared to molecular methods. Second, some IHC
assays may suffer from poor specificity as antigen detection highly
depends upon the quality of antibodies. Third, the dependence
upon the temporal expression of viral antigen dictates that
antigen detection is limited to a specific phase of infections when
the antigens are expressed in tissue.
Serology
Besides identifying viruses by viral culture and antigen detection,
serology assays play a critical role in the diagnosis of emerging
and/or re-emerging neurotropic viral infections, which detect
human antibodies (e.g., IgM or IgG) elicited after humoral immune
response. Viral CNS infections can be confirmed by detection of
virus-specific IgM antibody during the acute phase in CSF or by
demonstration of at least fourfold or greater increase in virus-
specific neutralizing antibodies between acute- and convalescent-
phase CSF. For example, suspected cases of arboviral infection are
most rapidly diagnosed using serological methods.
18
Determina-
tion of CSF IgM or IgG for ZIKV, DENV, or WNV play an important
role in the definitive diagnosis of CNS infection of these
Fast cell division
High basal metabolism
High respiratory rates
Thin skin
Large
body surface area
Immature
immune system
Immature
blood-brain barrier
Vulnerability to
CNS infections
Enhanced risk
of infection
Distinct
microbiota/virus
colonization in
non-CNS body
sites
ATP
Fig. 2 Children, especially neonates and infants, are more vulnerable to viral CNS infections and meningitis than adults. This vulnerability
may be attributed to children’s unique anatomical and physiological features, including, but not limited to, fast cell division, high basal
metabolism, high respiratory rates, thin skin, large body surface area, immature blood-brain barrier and immature immune system. Among
them, fast cell division, high basal metabolism and respiratory rates as well as thin skin may be responsible for distinct microbiota/virus
colonization in non-CNS body sites.
13,56
Thin skin and large body surface area may enhance the risk of infection. The immature blood-brain
barrier and immune system may lead to vulnerability to CNS infections. ATP adenosine triphosphate.
B.M. Liu et al.
3
Pediatric Research
flaviviruses.
20
Although hemagglutination inhibition assay and
plaque reduction neutralization tests (PRNT) have been used,
ELISA has become the method of choice (Fig. 3). IgM antibody
capture ELISA is a sensitive method for the detection of IgM as it
can minimize the interference of high-avidity IgG and nonspecific
antibody binding.
14
For the detection of IgG, indirect IgG ELISA is
more sensitive than the direct method.
14
ELISA assays are cost-
effective and less expensive than molecular tests. In addition to
ELISA, indirect Fluorescent Ab (IFA), e.g., Arbovirus IFA IgM & IgG
(DiaSorin Molecular), is also widely used, which is cost-effective
and fast. IFA enables visualization and confirmation of the location
of fluorescence (Fig. 3).
However, serology assays have their own limitations (Fig. 3).
First, cross-reaction within the same viral family (e.g., flaviviruses)
may lead to false positive results for IgM testing and neutralizing
antibody assays for a specific virus. For example, a definitive
serological diagnosis for ZIKV infection may be affected by the
presence of a high-level cross-reactivity between ZIKV and other
flaviviruses such as DENV, and prior immunizations. This is
especially true to travelers to the endemic area of circulating
flaviviruses and populations with a high background of other
flavivirus infections, such as DENV, which may give rise to cross-
reaction in serologic assays for ZIKV.
26–30
This issue made the
diagnosis of ZIKV infection very complicated during the Zika
epidemic due to short sensitivity windows of IgG and false
positives due to cross-reactivity with other viruses. PRNT that
detects virus-specific neutralizing antibodies can be used to
discriminate between cross-reacting antibodies in primary viral
infections, but it requires extra expertise and training and may not
be available at all laboratories. Second, though IgM is generally
considered as an acute-phase biomarker, a positive serum IgM test
may persist for long periods in some cases and reflect a prior
infection.
14
A compelling example is CSF IgM for WNV that can
persist for 30–90 days and complicate the diagnosis process. For
example, a patient presenting with neurological symptoms and
testing positive for WNV IgM could either be experiencing an
acute infection or have had a past infection and still have lingering
IgM antibodies. In addition, immunocompromised patients may
have a delayed or blunted serologic response, which can lead to
suboptimal sensitivity.
18
Third, it may take some time (window
period) to elicit antibody. Samples collected within 2 weeks after
onset may yield negative antibody IFA. Last, microscope optics
and light affect endpoint titers of IFA. Result interpretation of IFA
is subjective.
NUCLEIC ACID AMPLIFICATION TESTS (NAATS)
NAAT assays with ≤2 organism targets
While serology provides valuable information for clinical diagnosis,
NAATs are widely used in detection of CNS viral pathogens in CSF
since 1990s.
12
A NAAT assay consists of three separate steps:
nucleic acid extraction, target nucleic acid amplification, and
amplicon or nucleic acid detection. While nucleic acid detection is
an indispensable step, some assays do not have nucleic acid
extraction and amplification steps. Depending upon the numbers
of organism targets of interest, NAAT assays can be divided into
singleplex (one organism target) and multiplex (2 or more
organism targets) assays (Fig. 4). Due to their short turnaround
Targets
Antigen
(Ag)
Viral
nucleic
acids
Viruses
Antibody
(Ab)
(IgG/IgM
Detection methods Pros Cons
NAATs
e.g., DENV or ZIKV RT-PCR,
BioFire FilmArray ME Panel
e.g., EV typing
e.g., mNGS assays for CSF diagnostics
e.g., DENV NS1 Ag, serum or plasma
(Bio-Rad)
e.g., Dengue detect IgG and IgM
capture ELISA (InBios)
e.g., Arbovirus IFA IgM & IgG (DiaSorin)Enable visualization and confirmation of
the location of fluorescence
Cost-effective and short TAT
Indirect IgG ELISA is more sensitive than
direct method
IgM Ab capture (MAC) ELISA is sensitive
Shell vial culture has good specificity
and sensitivity
Accept a broad range of sample sources
Gold standard for virus detection
Short TAT and relatively low cost
Aid in early diagnosis within the first 1–7
days after onset of illness
High sensitivity and specificity
High sensitivity and specificity Prone to contamination
Cannot differentiate living vs dead organisms
Multiplexing tends to lower sensitivity
Can only sequence one fragment at a time
Fails to sequence samples with high CT value
Cannot detect minor (< 20%) variants
Relatively long TAT (1–2 days) and costly
May yield irrelevant organisms without
clinical significance
Lower sensitivity than NAATs
May have low sensitivity and specificity
Limited to clinical phases with Ag expression
Positive tests do not provide serotypes
Negative test results cannot rule out infection
May be less sensitive in secondary DENV
infections
Limited to detecting cultivable viruses
Viral culture may have low yield
Lower sensitivity & longer TAT than NAATs
May require technical expertise
Cross-reaction between viruses within family
Positive results need confirmation by PRNT
Positive IgM may persist for long periods
Negative Ab IFA within 2 weeks after onset
Microscope optics and light affect endpoint
titers. Result interpretation is subjective
Shorter turnaround time (TAT) than culture
+ive DENV or ZIKV PCR = Acute infection
Short TAT and relatively low cost
Enable to visualize infected cells in situ
Higher sequencing depth for minor
variants (down to 1–5%)
Pan-pathogen detection (agnostic)
Straightforward data analysis
Established workflow
Fast and cost-effective
Can detect unknown pathogens
Sanger sequencing
Metagenomic NGS (mNGS)
Immunohistochemistry
Ag ELISA
ELISA
Indirect Fluorescent Ab (IFA)
Viral culture
Shell vial culture
Fig. 3 Commonly used clinical diagnostic testing methods for emerging and re-emerging viral diseases. (Left panel) To diagnose
neurotropic viral infections, viral nucleic acids, antigens, viruses and antibodies (IgM and/or IgG) can be detected as targets in diagnostic
microbiology/virology assays. Pros (middle panel) and cons (right panel) of the indicated detection methods are shown. Viral nucleic acids
(red panel) can be detected by using nucleic acid amplification tests, Sanger sequencing and metagenomic next-generation sequencing. Viral
antigens (cyan panel) can be detected by using immunohistochemistry and antigen enzyme-linked immunosorbent assays (ELISA). Viruses
(purple panel) can be detected by viral culture and shell vial culture. Antibodies (IgM and/or IgG; orange panel) can be detected by ELISA and
indirect fluorescent antibody assay. Ab antibody, Ag antigen, CSF cerebrospinal fluid, C
T
cycle threshold, DENV dengue virus, EV enterovirus,
IFA indirect fluorescent antibody, MAC IgM Ab capture, ME meningitis/encephalitis, NAAT nucleic acid amplification test, NGS next-generation
sequencing, NS1 nonstructural protein 1, PRNT plaque reduction neutralization tests, RT-PCR reverse transcription-polymerase chain reaction,
TAT turnaround time, ZIKV Zika virus, +ive positive.
B.M. Liu et al.
4
Pediatric Research
time as well as high sensitivity and specificity, NAATs have
become the diagnostic standard for most viral CNS infections
(Fig. 3). For example, RT-PCR is the most sensitive and reliable
method for confirmation of ZIKV infections when the optimal
sample types are collected during the highest sensitivity window
of detection for ZIKV RNA.
30
Positive DENV or ZIKV NAATs confirm
acute infection, and no additional testing is indicated.
14,18
For ZIKV
infection, the duration of viremia is 3–14 days after the onset of
symptoms, but viremia can be prolonged up to 70 days in
pregnant women.
14,18,26–30
As shown in Table 1, there are four FDA-approved/cleared,
qualitative NAAT assays for the detection of microorganisms in
CSF specimens at the time of manuscript preparation. Three of
them are NAAT assays with ≤2 viral targets, including Xpert EV
Assay (Cepheid, Sunnyvale, CA) and NucliSens EasyQ Enterovirus
vl.1 Assay (bioMérieux, Durham, NC) for the detection of EVs in
CSF and Simplexa HSV 1&2 Direct (DiaSorin Molecular, Cypress,
CA) for detection of herpes simplex virus type 1 (HSV-1) and type 2
(HSV-2) in CSF, cutaneous and mucocutaneous swab samples.
20,31
In contrast, BioFire FilmArray Meningitis/Encephalitis (ME) Panel is
a NAAT assay with 14 organism targets (Table 1and Fig. 4), which
will be described in section 4.2. In March 2016, a triplex-PCR assay
was approved by the FDA under emergency use authorization for
the simultaneous detection of ZIKV, CHIKV and DENV in CSF, urine,
serum and amniotic fluid (Fig. 4).
20
Using dual labeled hydrolysis
probes, the RT-PCR assay has a LOD of 1.54 × 10
4
GCE/ml of ZIKV in
serum.
20
Though urine is not always routinely collected, this
specimen type may have longer windows of detection by
PCR.
26–29
Development of new NAAT assays for the detection of CNS viral
pathogens has evolved in recent years. The development of a pan-
HPeV RT-PCR test (covering HPeV 1–6) is particularly noteworthy
because HPeV, a common cause of sepsis-like illness and
meningitis in young infants, was previously difficult to diagnose
due to the lack of a sensitive and specific diagnostic test. In 2008,
Nix et al. reported a pan-HPeV RT-PCR assay for the detection of
HPeV1-6.
32
Subsequently, other laboratory-developed HPeV RT-
PCR tests, including a HPeV3-specific RT-PCR, and droplet digital
PCR assays were developed.
33–35
The addition of a sensitive and
specific method for the detection of HPeV to the current menu of
laboratory-developed RT-PCR viral pathogen detection tests on
CSF would be beneficial for appropriate patient care.
Given that various one-step and two-step RT-PCR kits can confer
different reverse transcription, amplification, and detection
efficiencies, Liu et al. compared the sensitivity of four commercial
one-step RT-PCR kits and found that SuperScript III One-Step RT-
PCR System was the best-performing one-step RT-PCR kit with
optimal amplification efficiency compared to GoTaq Probe 1-Step
RT-qPCR System, QuantiTect Probe RT-PCR Kit and PrimeScript
One Step RT-PCR Kit Ver. 2.
9,16,34,36–39
Subsequently, Liu et al.
developed and validated a novel pan-HPeV RT-PCR test (EliTech
HPeV RT-PCR Test) based on utilization of EliTech HPeV detection
reagent, optimization and standardization of RT-PCR master mix
and inclusion of MS2 internal control.
9
The new test was
demonstrated to have the limit of detection (LOD) of 570
copies/ml, broad coverage of HPeV 1–6, and excellent reprodu-
cibility and accuracy, with no cross-reactivity with other CNS
pathogens.
9
Multiplex CNS panels with >3 organism targets
With the development of NAAT assays, multiplexed syndromic
NAAT panels emerge, which employ multiple primer/probe sets to
simultaneously detect multiple organisms associated with a
similar and overlapping clinical symptomatology. For example, a
Xpert EV assay
Flexibility
Simplexa
HSV-1&2 direct
ZIKV, CHIKV & DENV
BioFire FilmArray
Meningitis/Encephalitis Panel
Metagenomic
NGS assay
Extraction
RNA/DNA
Library
Prep NGS
Cell
lysis PCR1PCR2
Sequence-
based ID
Virus B
Virus C
Virus A
Report
Target numbers
significantly increase and
can report > 1000 targets
14 targets
3 targets
2 targets
1
target
Throughput
Fig. 4 Classification of molecular methods for infectious diseases diagnostics based on number of targets. Nucleic acid amplification tests
(NAATs) can be divided into singleplex (one organism target) and multiplex (>1 organism targets) tests.
14,20,57
(Top panel) Xpert EV Assay
(Cepheid) is an example of singleplex NAAT. (Middle panels) Simplexa HSV-1&2 Direct (DiaSorin Molecular) is an example of duplex (two
organism targets) PCR panel. In addition, a molecular panel for the detection of Zika, Chikungunya, and dengue virus in CSF was approved by
the FDA under emergency use authorization, which is a triplex (three organism targets) PCR assay.
20
BioFire FilmArray Meningitis/Encephalitis
Panel detecting 14 CNS pathogens is the only FDA-approved syndromic panel for CNS infection, which uses a closed pouch to perform cell
lysis and nested PCR (PCR 1 and 2). (Bottom panel) Metagenomic next-generation sequencing (NGS) assays were developed to aid in pan-
pathogen detection from CSF, which consists of total nucleic acid (RNA/DNA) extraction, library prep, NGS and sequence-based identification.
Target numbers of metagenomic NGS assay significantly increase and can report >1000 organism targets in some instances.
47
(Right) With the
increase of target numbers from singleplex tests to multiplex tests to metagenomic NGS assays, test throughput increases but flexibility
decreases. The bigger a panel/assay is, the more organisms they can detect. However, some organisms on the panel but not on the differential
are still tested if BioFire FilmArray Meningitis/Encephalitis Panel or a metagenomic NGS assay is performed. Unexpected organisms detected
from the BioFire panel or the metagenomic NGS assay may lead to some confusion and challenges in interpretation of test results. CHIKV
Chikungunya virus, DENV dengue virus, EV enteroviruses, HSV herpes simplex virus, ID identification, ZIKV Zika virus.
B.M. Liu et al.
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Pediatric Research
multiplex CNS panel is a NAAT assay that uses different primer/
probe sets to simultaneously detect multiple organisms causing
CNS infection. Though there have been numerous FDA-approved/
cleared automated multiplexed syndromic NAAT panels for
simultaneous detection of organisms implicated in blood stream,
respiratory, gastrointestinal, joint infections, the BioFire FilmArray
ME Panel is the only FDA-approved multiplexed syndromic testing
for CNS infections.
31,40,41
The BioFire FilmArray ME Panel can
simultaneously detect 14 pathogens directly from CSF specimens
collected by lumbar puncture, with rapid turnaround time
(∼1 h).
40,41
Among the target organisms, there are 7 viruses,
including cytomegalovirus, EV, HSV-1, HSV-2, human herpesvirus 6
(HHV-6), HPeV, and varicella zoster virus. This panel has an overall
94.2% sensitivity and 99.8% specificity (https://www.biofiredx.com/
products/the-filmarray-panels/filmarrayme/). The BioFire FilmArray
ME Panel demonstrated positive percentage of agreement of 100%
for 9 of 14 analytes; for example, EV and HHV-6 demonstrated
agreements of 95.7% and 85.7%, respectively (https://
www.biofiredx.com/products/the-filmarray-panels/filmarrayme/).
40,41
The use of this panel likely led to more infants being diagnosed
with HPeV meningitis during outbreaks in the U.S. in 2022.
7,8
Between January and December of 2022, we identified 20 infants
with HPeV meningitis by testing CSF specimens using BioFire
FilmArray ME Panel at Children’s National Hospital in Washington,
DC (unpublished data). Four of the positive patients were
eventually admitted to our neonatal intensive care unit.
However, the BioFire FilmArray ME Panel has several limitations
(Fig. 3). First, as a molecular test, the BioFire FilmArray ME Panel
has common limitations as other molecular assays, e.g., high cost
and contamination, and failure to differentiate living vs dead
organisms. Second, it has varied sensitivity for detecting different
target organisms. A high proportion of false-negative results were
observed for both HSV-1 (7/26) and HSV-2 (7/55) by Leber et al.,
which suggests that negative panel results do not rule out CNS
HSV infections.
41,42
Of note, BioFire FilmArray ME Panel has a high
specificity (99.9%) for HSV-1 (1556/1558) and HSV-2 (1548/1550).
43
Third, multiplexing tends to lower sensitivity. When comparing
with NAAT assays with ≤2 organism targets, BioFire FilmArray ME
Panel has a lower sensitivity for HSV, EV and HPeV. For example,
Simplexa HSV 1&2 Direct has ~10-fold lower LOD (more sensitive)
than that of BioFire FilmArray ME panel.
44
This is also true for EV
and HPeV that are more easily detected by some singleplex
molecular assays due to a higher sensitivity than BioFire FilmArray
ME panel.
9,44
Fourth, the BioFire FilmArray ME Panel has some
targets which may yield hard-to-interpret results, e.g., HHV-6, the
only human herpesvirus known to be integrated into germline
chromosomal telomeres.
43
Given the existence of chromosomally
integrated HHV-6, positive HHV-6 results may not indicate active
viral replication or infection, and therefore, should be carefully
interpreted with correlation with clinical symptoms and other
laboratory testing results.
43
This highlights the importance of
correlating of molecular tests with other tests. Last, the BioFire
FilmArray ME Panel is a “one for all”test detecting 14 pathogens at
the same time. With the number of targets increases, the test
throughput increases but flexibility decreases (Fig. 4). Some
organisms on the BioFire FilmArray ME Panel but not on the
differential are still tested if the panel is used. Unexpected
organisms detected from the BioFire FilmArray ME Panel may lead
to some confusion and challenges in interpretation of test results.
SEQUENCING-BASED MOLECULAR ASSAYS
Previously Sanger sequencing has shown to have useful applica-
tion for EV genotyping and detection of antiviral-resistant CSF
CMV isolate.
12,45
Sanger-based EV typing relies on the proper
design of universal primers for multiple EV genotypes, which has
turned out to be a critical step for direct sequencing-based EV
genotyping.
45
Sanger sequencing has its pros and cons (Fig. 3). It
Table 1. Commercially available, FDA-cleared/approved nucleic acid amplification tests for the detection of microorganisms associated with central nervous system infections.
41,44,51–53
Organisms Assay name Manufacturer Technology Specimen type Target Comments TAT (h)
Enterovirus Xpert EV Assay Cepheid,
Sunnyvale, CA
Real-time PCR CSF 5’UTR Fully integrated and
random access
2.5
Enterovirus NucliSens EasyQ
Enterovirus vl.1
Assay
bioMérieux,
Durham, NC
Nucleic acid
sequence-based
amplification
CSF 5’UTR Nucleic acid
separation and
amplification/
detection
5
HSV 1 & 2 Simplexa HSV-1&2
Direct
DiaSorin Molecular,
Cypress, CA
Real-time PCR CSF, cutaneous and
mucocutaneous swab
samples
DNA
Polymerase
Semi-automated; no
extraction
1
Escherichia coli K1, Haemophilus
influenzae,Listeria monocytogenes,
Neisseria meningitidis,Streptococcus
agalactiae,Streptococcus pneumoniae,
Cryptococcus neoformans/gattii,
Cytomegalovirus, Enterovirus, Herpes
simplex virus type 1, Herpes simplex
virus type 2, Human herpes virus 6,
Human parechovirus, Varicella zoster
virus
FilmArray
Meningitis/
Encephalitis Panel
BioFire Diagnostics,
Salt Lake City, UT
Multiplex PCR
followed by solid
array
CSF Not available Fully integrated and
random access
1
CSF cerebrospinal fluid, EV Enterovirus, HSV herpes simplex virus, TAT turnaround time, UTR untranslated region.
B.M. Liu et al.
6
Pediatric Research
is cost-effective, and has established workflow and straightforward
data analysis. However, Sanger sequencing can only sequence one
fragment at a time, and fails to sequence samples with high C
T
value or detect minor (<20%) variants.
The advent of next-generation sequencing (NGS) has resulted in
elimination of the requirement for the “primer walking”steps, as
used in Sanger sequencing. Rather, NGS has higher throughput
(i.e., sequencing multiple fragments at a time) and higher
sequencing depth for minor variants (down to 1–5%) (Fig. 3).
The past decade has witnessed agnostic, metagenomic NGS (i.e., a
NGS assay allowing for comprehensive detection of all genes in all
organisms in a given sample) emerging as a promising pan-
pathogen detection method for clinical specimens (Fig. 4).
46,47
The
hypothesis-free features of metagenomic NGS have made it a
valuable addition to a suite of molecular assays for detection of
CNS infections (Fig. 3). Simner et al. developed and optimized a
metagenomic NGS test for pan-pathogen detection in CSF, which
correctly detected pathogens compared to stand-of-care assays in
a proof-of-concept study.
47
In addition, Wilson et al. conducted a
1-year, multicenter, prospective study to investigate the utility of
metagenomic NGS of CSF for the diagnosis of infectious
meningitis and encephalitis in hospitalized patients.
46
They found
that metagenomic NGS of CSF obtained from patients with
meningitis or encephalitis improved diagnosis of neurologic
infections and provided actionable information in some cases,
e.g., those with SLEV.
46
Metagenomic NGS also has the promise
and capacity to detect unknown pathogens (Fig. 3).
However, it is challenging to implement and interpret the
metagenomic NGS assay for pan-pathogen detection in CSF
(Fig. 3). First, current NGS workflow may take 1–2 days and would
delay the turnaround time of the diagnosis of CNS infections
compared with targeted NAATs or the BioFire FilmArray ME Panel
whose turnaround time is one to several hours.
40
Second, due to
the metagenomic nature of the existing metagenomic NGS assays,
they may yield irrelevant organisms without clinical significance,
which may lead to unnecessary treatment. Therefore, clinical
microbiology consultation may be necessary to provide a better
interpretation on metagenomic NGS results for CNS infections.
Third, metagenomic NGS assays may lead to false negative results
given that their sensitivity for some targets may be lower than
targeted NAATs. Therefore, negative metagenomic NGS results
cannot completely rule out the presence of CNS infection.
PERSPECTIVES AND CONCLUSIONS
The emergence and re-emergence of neurotropic CNS viruses, e.g.,
ZIKV, WNV, EV and HPeV, have constituted a major threat to the
health of young children. Infection due to these viruses can lead to
meningitis, encephalitis, seizures, brain imaging abnormalities, and
long-term neurodevelopmental sequelae, especially in the most
vulnerable populations, e.g., neonates and immunocompromised
hosts. Rapid and accurate detection of these viruses from CSF and
other specimens is critical for diagnosis of CNS diseases and for
preventing inappropriate and costly treatments. The advent of
multiplex meningitis/encephalitis panels and metagenomic next-
generation sequencing assays for pan-pathogen detection have
been demonstrated as useful additions to a suite of molecular
assays for detection of viral pathogens in CSF. However, the
expansion of test menu has led to new challenges in selecting an
appropriate test and making accurate interpretation of test results.
The new assays have their own advantages and limitations due to
their varied sensitivities, specificities and the best detection
window. Clinicians should consider these important features of
different assays to choose the right tests at the right time for the
right patients and make accurate interpretation of the test results.
Further, more research is needed to best determine the clinical
utility of the BioFire FilmArray ME Panel and metagenomic NGS CSF
assays and optimize their implementation into clinical practice.
There are still unmet gaps in the development of rapid, sensitive
and specific molecular assays for a growing list of emerging and re-
emerging neurotropic viruses, such as EV-D68. This review calls for
more research on development and clinical utility study of novel
molecular assays. With the growing usage of host immune
biomarkers (e.g., TRAIL, IP-10, CRP) and other cytokines/chemokines
in infectious diseases,
48–50
their clinical applications in emerging
and re-emerging viral diseases are worth further investigations.
DATA AVAILABILITY
Not applicable.
REFERENCES
1. Autore, G., Bernardi, L., Perrone, S. & Esposito, S. Update on viral
infections involving the central nervous system in pediatric patients. Children 8,
782 (2021).
2. Tunkel, A. R. et al. The management of encephalitis: clinical practice guidelines by
the Infectious Diseases Society of America. Clin. Infect. Dis.47, 303–327 (2008).
3. Abdullahi, A. M., Sarmast, S. T. & Jahan, N. Viral infections of the central nervous
system in children: a systematic review. Cureus 12, e11174 (2020).
4. Chow, F. C. & Glaser, C. A. Emerging and reemerging neurologic infections.
Neurohospitalist 4, 173–184 (2014).
5. Mulkey, S. B. & DeBiasi, R. L. New insights into Zika in infants and children. Trop.
Med. Infect. Dis. 7, 158 (2022).
6. Pierson, T. C. & Diamond, M. S. The emergence of Zika virus and its new clinical
syndromes. Nature 560, 573–581 (2018).
7. Tao, L., Humphries, R. M., Banerjee, R. & Gaston, D. C. Re-emergence of par-
echovirus: 2017-2022 national trends of detection in cerebrospinal fluid. Open
Forum Infect. Dis. 10, ofad112 (2023).
8. Tao, L., Fill, M. A., Banerjee, R. & Humphries, R. M. Notes from the field: cluster of
parechovirus central nervous system infections in young infants - Tennessee,
2022. MMWR Morb. Mortal. Wkly. Rep. 71, 977–978 (2022).
9. Liu, B., Forman, M. & Valsamakis, A. Optimization and evaluation of a novel real-
time RT-PCR test for detection of parechovirus in cerebrospinal fluid. J. Virol.
Methods 272, 113690 (2019).
10. Al Awaidy, S. T. & Khamis, F. Wild poliovirus type 1 in Oman: a re-emerging threat
that requires urgent, targeted and strategic preparedness. Sultan Qaboos Univ.
Med. J. 20,e1–e4 (2020).
11. Tang, Y. W. Laborat ory diagnosis of CNS infections by molecular amplification
techniques. Expert Opin. Med. Diagn. 1, 489–509 (2007).
12. Debiasi, R. L. & Tyler, K. L. Molecular methods for diagnosis of viral encephalitis.
Clin. Microbiol. Rev. 17, 903–925 (2004).
13. Doern, C. D., Dunn, J. J. & McAdam, A. J. Pediatric clinical microbiology: it’s the
little things. J. Clin. Microbiol. 54, 1412–1413 (2016).
14. Jorgensen, J. H. et al. Manual of Clinical Microbiology 11th edn, Vol. 2 (ASM Press,
2015).
15. Yang, D. et al. The E3 ligase TRIM56 is a host restriction factor of Zika virus and
depends on its RNA-binding activity but not miRNA regulation, for antiviral
function. PLoS Negl. Trop. Dis. 13, e0007537 (2019).
16. Liu, B. et al. Overlapping and distinct molecular determinants dictating the
antiviral activities of TRIM56 against flaviviruses and coronavirus. J. Virol. 88,
13821–13835 (2014).
17. Jayakeerthi, R. S., Potula, R. V., Srinivasan, S. & Badrinath, S. Shell vial culture assay
for the rapid diagnosis of Japanese encephalitis, West Nile and Dengue-2 viral
encephalitis. Virol. J. 3, 2 (2006).
18. Kimberlin, D. W., Barnett, E. D., Lynfield, R. & Sawyer M. H. Red Book: 2021-2024
Report of the Committee on Infectious Diseases/Committee on Infectious Diseases,
American Academy of Pediatrics (American Academy of Pediatrics, 2021).
19. Ginocchio, C. C. et al. Development, technical performance, and clinical evalua-
tion of a NucliSens basic kit application for detection of enterovirus RNA in
cerebrospinal fluid. J. Clin. Microbiol. 43, 2616–2623 (2005).
20. He, T., Kaplan, S., Kamboj, M. & Tang, Y. W. Laboratory diagnosis of central
nervous system infection. Curr. Infect. Dis. Rep. 18, 35 (2016).
21. Gelpi, E. et al. Visualization of Central European tick-borne encephalitis infection
in fatal human cases. J. Neuropathol. Exp. Neurol. 64, 506–512 (2005).
22. Reyes, M. G., Gardner, J. J., Poland, J. D. & Monath, T. P. St Louis encephalitis.
Quantitative histologic and immunofluorescent studies. Arch. Neurol. 38, 329–334
(1981).
23. Nagel, M. A. & Gilden, D. Neurological complications of varicella zoster virus
reactivation. Curr. Opin. Neurol. 27, 356–360 (2014).
24. Ludlow, M. et al. Neurotropic virus infections as the cause of immediate and
delayed neuropathology. Acta Neuropathol. 131, 159–184 (2016).
B.M. Liu et al.
7
Pediatric Research
25. Forest, F. et al. Lethal human herpesvirus-6 encephalitis after cord blood trans-
plant. Transpl. Infect. Dis. 13, 646–649 (2011).
26. Rossini, G., Gaibani, P., Vocale, C., Cagarelli, R. & Landini, M. P. Comparison of Zika
virus (ZIKV) RNA detection in plasma, whole blood and urine—case series of
travel-associated ZIKV infection imported to Italy, 2016. J. Infect. 75, 242–245
(2017).
27. Joguet, G. et al. Effect of acute Zika virus infection on sperm and virus clearance
in body fluids: a prospective observational study. Lancet Infect. Dis. 17,
1200–1208 (2017).
28. Suy, A. et al. Prolonged Zika virus viremia during pregnancy. N. Engl. J. Med. 375,
2611–2613 (2016).
29. Lozier, M. J. et al. Prolonged detection of zika virus nucleic acid among symp-
tomatic pregnant women: a cohort study. Clin. Infect. Dis. 67, 624–627 (2018).
30. Voermans, J. J. C. et al. Whole-blood testing for diagnosis of acute zika virus
infections in routine diagnostic setting. Emerg. Infect. Dis. 25, 1394–1396 (2019).
31. Hanson, K. E. The first fully automated molecular diagnostic panel for meningitis
and encephalitis: how well does it perform, and when should it be used? J. Clin.
Microbiol. 54, 2222–2224 (2016).
32. Nix, W. A. et al. Detection of all known parechoviruses by real-time PCR. J. Clin.
Microbiol. 46, 2519–2524 (2008).
33. Esposito, S. et al. Pediatric parechovirus infections. J. Clin. Virol. 60,84–89 (2014).
34. Selvaraju, S. B., Nix, W. A., Oberste, M. S. & Selvarangan, R. Optimization of a
combined human parechovirus-enterovirus real-time reverse transcription-PCR
assay and evaluation of a new parechovirus 3-specific assay for cerebrospinal
fluid specimen testing. J. Clin. Microbiol. 51, 452–458 (2013).
35. Aizawa, Y., Koyama, A., Ishihara, T., Onodera, O. & Saitoh, A. Performance of a
realtime PCR-based approach and droplet digital PCR in detecting human par-
echovirus type 3 RNA. J. Clin. Virol. 84,27–31 (2016).
36. She, R. C., Hymas, W. C., Taggart, E. W., Petti, C. A. & Hillyard, D. R. Performance of
enterovirus genotyping targeting the VP1 and VP2 regions on non-typeable
isolates and patient specimens. J. Virol. Methods 165,46–50 (2010).
37. Wei, D. et al. The molecular chaperone GRP78 contributes to toll-like receptor
3-mediated innate immune response to hepatitis C virus in hepatocytes. J. Biol.
Chem. 291, 12294–12309 (2016).
38. Peng, Y. et al. Naturally occurring deletions/insertions in HBV core promoter tend
to decrease in hepatitis B e antigen-positive chronic hepatitis B patients during
antiviral therapy. Antivir. Ther. 20, 623–632 (2015).
39. LeMessurier, K. S. et al. Influenza A virus directly modulates mouse eosinophil
responses. J. Leukoc. Biol. 108, 151–168 (2020).
40. Hanson, K. E. & Couturier, M. R. Multiplexed molecular diagnostics for respiratory,
gastrointestinal, and central nervous system infections. Clin. Infect. Dis. 63,
1361–1367 (2016).
41. Leber, A. L. et al. Multicenter evaluation of biofire filmarray meningitis/ence-
phalitis panel for detection of bacteria, viruses, and yeast in cerebrospinal fluid
specimens. J. Clin. Microbiol. 54, 2251–2261 (2016).
42. Vetter, P., Schibler, M., Herrmann, J. L. & Boutolleau, D. Diagnostic challenges of
central nervous system infection: extensive multiplex panels versus stepwise
guided approach. Clin. Microbiol. Infect. 26, 706–712 (2020).
43. BioFire FilmArray Meningitis/Encephalitis Panel: package insert (2023).
44. Liesman, R. M. et al. Evaluation of a commercial multiplex molecular panel for
diagnosis of infectious meningitis and encephalitis. J. Clin. Microbiol. 56, e01927-
17 (2018).
45. Liu, B. Universal PCR primers are critical for direct sequencing-based enterovirus
genotyping. J. Clin. Microbiol. 55, 339–340 (2016).
46. Wilson, M. R. et al. Clinical metagenomic sequencing for diagnosis of meningitis
and encephalitis. N. Engl. J. Med. 380, 2327–2340 (2019).
47. Simner, P. J. et al. Development and optimization of metagenomic next-
generation sequencing methods for cerebrospinal fluid diagnostics. J. Clin.
Microbiol. 56, e00472-18 (2018).
48. van Houten, C. B. et al. A host-protein based assay to differentiate between
bacterial and viral infections in preschool children (OPPORTUNITY): a double-
blind, multicentre, validation study. Lancet Infect. Dis. 17, 431–440 (2017).
49. Liu, B. M., Martins, T. B., Peterson, L. K. & Hill, H. R. Clinical significance of mea-
suring serum cytokine levels as inflammatory biomarkers in adult and pediatric
COVID-19 cases: a review. Cytokine 142, 155478 (2021).
50. Liu, B. M. & Hill, H. R. Role of host immune and inflammatory responses in COVID-
19 cases with underlying primary immunodeficiency: a review. J. Interferon
Cytokine Res. 40, 549–554 (2020).
51. Nolte, F. S. et al. Evaluation of a rapid and completely automated real-time
reverse transcriptase PCR assay for diagnosis of enteroviral meningitis. J. Clin.
Microbiol. 49, 528–533 (2011).
52. Capaul, S. E. & Gorgievski-Hrisoho, M. Detection of enterovirus RNA in cere-
brospinal fluid (CSF) using NucliSens EasyQ Enterovirus assay. J. Clin. Virol. 32,
236–240 (2005).
53. Kuypers, J. et al. Comparison of the Simplexa HSV1 & 2 Direct kit and laboratory-
developed real-time PCR assays for herpes simplex virus detection. J. Clin. Virol.
62, 103–105 (2015).
54. Trovato, M., Sartorius, R., D’Apice, L., Manco, R. & De Berardinis, P. Viral emerging
diseases: challenges in developing vaccination strategies. Front. Immunol. 11,
2130 (2020).
55. Soman Pillai, V., Krishna, G. & Valiya Veettil, M. Nipah virus: past outbreaks and
future containment. Viruses 12, 465 (2020).
56. Oranges, T., Dini, V. & Romanelli, M. Skin physiology of the neonate and infant:
clinical implications. Adv. Wound Care 4, 587–595 (2015).
57. Liu, B. et al. Development and evaluation of a fully automated molecular assay
targeting the mitochondrial small subunit rRNA gene for the detection of
Pneumocystis jirovecii in bronchoalveolar lavage fluid specimens. J. Mol. Diagn. 22,
1482–1493 (2020).
AUTHOR CONTRIBUTIONS
B.M.L. drafted the article, Figs. 1–4, and Table 1. S.B.M., J.M.C. and R.L.D. revised the
manuscript.
FUNDING
B.M.L. was supported by the William L. Roberts Memorial Fund (553-Liu-11.20.20 and
553-Mehta-08.31.20), ARUP Institute for Experimental Pathology. This publication
resulted, in part, from research supported by the District of Columbia Center for AIDS
Research, an NIH funded program (P30AI117970), which is supported by the
following NIH Co-Funding and Participating Institutes and Centers: NIAID, NCI, NICHD,
NHLBI, NIDA, NIMH, NIA, NIDDK, NIMHD, NIDCR, NINR, FIC, and OAR. Research
reported in this work was also supported by the National Center for Advancing
Translational Sciences and the NIAID of the NIH under award number U54AI150225.
The content is solely the responsibility of the authors and does not necessarily
represent the official views of the NIH.
COMPETING INTERESTS
The authors declare no competing interests.
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