Viral Load Drives Disease in Humans Experimentally
Infected with Respiratory Syncytial Virus
John P. DeVincenzo1–3, Tom Wilkinson4, Akshay Vaishnaw5, Jeff Cehelsky5, Rachel Meyers5, Saraswathy Nochur5,
Lisa Harrison1,2, Patricia Meeking6, Alex Mann6, Elizabeth Moane6, John Oxford7, Rajat Pareek1, Ryves Moore1,
Ed Walsh8, Robert Studholme9, Preston Dorsett1,9, Rene Alvarez5, and Robert Lambkin-Williams6
1Departments of Pediatric and Department of Molecular Sciences, University of Tennessee College of Medicine,2Children’s Foundation Research
Center, and3Le Bonheur Children’s Medical Center, Memphis, Tennessee;4Southampton School of Medicine, Southampton, United Kingdom;
5Alnylam Pharmaceuticals, Cambridge, Massachusetts;6Retroscreen Ltd. and7Queen Mary’s School of Medicine and Dentistry, London, United
Kingdom;8University of Rochester, Rochester, New York; and9Viral Antigens/Meridian Life Science, Inc., Memphis, Tennessee
Rationale: Respiratory syncytial virus (RSV) is the leading cause of
childhood lower respiratory infection, yet viable therapies are
lacking. Two major challenges have stalled antiviral development:
ethical difficulties in performing pediatric proof-of-concept studies
and the prevailing concept that the disease is immune-mediated
rather than being driven by viral load.
infection model to address these challenges.
Methods: Healthy volunteers (n 5 35), in five cohorts, received
increasing quantities (3.0–5.4 log plaque-forming units/person) of
wild-type RSV-A intranasally.
shed virus. Infection rate, viral loads, disease severity, and safety were
similar between cohorts and were unrelated to quantity of RSV re-
in severity near when viral load peaked, and subsided as viral loads
(measured by real-time polymerase chain reaction) slowly declined.
Viral loads correlated significantly with intranasal proinflammatory
cytokine concentrations (IL-6 and IL-8). Increased viral load correlated
(symptoms, physical examination, and amount of nasal mucus).
with RSV infection. The observed parallel viral and disease kinetics
support a potential clinical benefit of RSV antivirals. This reproducible
model facilitates the development of future RSV therapeutics.
Keywords: RSV; pneumonia; bronchiolitis; pathogenesis; viral load
Respiratory syncytial virus (RSV) infects more than 68% of the
birth cohort annually (1) and is the most common cause of
lower respiratory infections in children less than 1 year of age
resulting in significant morbidity (2, 3). RSV also causes life-
threatening disease (4), with mortality rates approximately 10-
fold higher than for influenza in those less than 1 year of age (5).
Worldwide, as many as 1 million children may die annually of
RSV infection (6, 7). RSV also produces significant morbidity
and mortality in adult immunocompromised and debilitated
populations (8). Despite the high disease burden, there is no
vaccine for RSV and the only approved therapy (ribavirin) is
rarely used because of its potential teratogenicity and its limited
effectiveness (9–11). Although successful, passive monoclonal
antibody prevention strategies are currently applied to less than
5% of the at-risk childhood population (12, 13). Thus, there is
a major unmet medical need for effective therapies for RSV
There are two main reasons for the unavailability of RSV
therapeutics. The first is inadequate existing evidence of RSV
pathogenesis that might support the putative effectiveness of
RSV antivirals. Disease caused by RSV has long been thought
to be the result of the viral triggering of an exuberant, aberrant,
and long-lasting pathogenic immune response. This immunopa-
thogenic concept has been largely developed through experi-
mentation with rodent models of RSV infection (14–16). It is
thought to result from exaggerated helper T-cell type 2 (Th2)
AT A GLANCE COMMENTARY
Scientific Knowledge on the Subject
The pathogenesis of respiratory syncytial virus (RSV) in-
fection has been studied primarily in rodent systems and
indicates that disease is produced by an aberrant delayed
and pathogenic immune response. Studies of pathogenesis in
humans are largely lacking.
What This Study Adds to the Field
This study provides a human experimental infection model
that can be used to study the pathogenesis of the disease in
humans themselves, and that can also be used for evalua-
tion of therapeutics and vaccines. The pathogenesis of RSV
infection in humans shows that viral load is associated with
disease itself and that the viral load and disease track
together. This indicates that the previously held view of a
delayed and aberrant immune response causing the disease is
likely more applicable to animal models rather than to human
RSV infection. This research also supports the development
of antiviral therapeutics for RSV infection.
(Received in original form March 5, 2010; accepted in final form July 8, 2010)
Supported in part by the following grants to Dr. DeVincenzo: NIH RR 16187 from
the National Center for Research Resources, the Children’s Foundation Research
Center of Memphis, the University of Tennessee General Clinical Research Center
(GCRC) (UHPHS RR00211), the Le Bonheur Foundation, and Alnylam Pharma-
ceuticals, Cambridge, MA.
Disclaimer: The conduct of this research follows the human experimentation
guidelines of the U.S. Department of Health and Human Services. This study was
also conducted in accordance with ICH GCP guidelines (directive CPMP/ICH/
135/95), local regulatory requirements and the declaration of Helsinki, and all
relevant local laws and regulations. The protocol and informed consent docu-
ment were reviewed and approved by a properly constituted ethics committee.
All aspects of the study were explained in detail to prospective subjects and they
were informed of the voluntary nature of their participation. Written informed
consent was obtained from each subject. Portions of the data in this article have
been presented as Abstract V-1257 at the Interscience Conference on Antimi-
crobial Agents and Chemotherapy, Chicago, IL, September 18, 2007.
Correspondence and requests for reprints should be addressed to John DeVincenzo,
M.D., Children’s Foundation Research Center, Le Bonheur Children’s Medical
Center, Room 433, 50 North Dunlap Street, Memphis, TN 38103. E-mail:
This article has an online supplement, which is available from the issue’s table of
contents at www.atsjournals.org
Am J Respir Crit Care Med
Originally Published in Press as DOI: 10.1164/rccm.201002-0221OC on July 9, 2010
Internet address: www.atsjournals.org
Vol 182. pp 1305–1314, 2010
cellular immune responses (17, 18) through bystander killing
effects of activated cytotoxic T cells (19) and by RSV-induced
chemokine mimicry (20, 21). The failed RSV vaccine experi-
ence of the 1960s (22–24), which produced a prominent and
fatal eosinophilic pulmonary infiltrate after natural RSV in-
fection in the unfortunate vaccine recipients, further solidified
this immunopathogenic concept of RSV disease (25). In a dis-
ease process that begins days after the start of a short period of
viral replication, and that is followed by a prolonged immune-
mediated disease continuing after viral elimination, applying an
antiviral agent would likely be clinically ineffective. However,
the relevance of this immunopathogenic concept for human
RSV disease is not understood (26).
The second main reason for the current unavailability of
effective RSV therapeutics is the understandable reluctance to
study experimental therapies directly in the vulnerable pediatric
naturally infected population before proof-of-concept studies
can show antiviral effect in adults, who are able to understand
and provide informed consent for research (27). The naturally
infected immunocompromised adult population does not offer
a suitable alternative, as researchers must contend with widely
varying degrees of immunosuppression, major differences in
elapsed times between initial infection and RSV detection, the
difficulties produced by administration of multiple concomitant
medications, and uncontrollable occurrences of death and other
serious adverse events that may be rightly or wrongly attributed
to the drug being studied. Furthermore, RSV-infected immu-
nocompromised adult populations are small, geographically
dispersed, and difficult to recruit.
These two main reasons have served to inhibit the preclinical
and clinical development of RSV antiviral therapeutics. Other
than ribavirin, numerous small molecules and novel therapeu-
tics have been discovered with proven RSV antiviral activity
in submicromolar concentrations both in vitro and in vivo
(28–38). However, only one of these has been tested in a proof-
of-concept study within an RSV-infected population (39). This
trial, which involved adults undergoing hematopoietic stem
cell transplantation, failed to show an antiviral effect because
of many of these aforementioned logistic, immunologic, and
To reevaluate the prevailing immune-based model of RSV
pathophysiology and to provide a practical means to conduct
proof-of-concept RSV therapeutic trials, we sought to develop
a safe, reproducible, and well-characterized human experimental
RSV infection model in adult volunteers that paralleled natural
RSV infection and disease. Some of the results of these studies
have been previously reported in the form of abstracts (40, 41).
The study included healthy males and females 18–45 years of age.
Exclusion criteria included the following: any history during adulthood
of asthma of any etiology or any use of a bronchodilator within the past
year; chronic (or acute within the last 7 d) use of any medication or
other product (prescriptive or over the counter) for symptoms of
rhinitis or nasal congestion or for any chronic nasopharyngeal com-
plaint, or chronic use of any intranasal medication for any indication;
presence of any febrile illness or symptoms suggestive of viral re-
spiratory infection within the last 2 weeks; history of seasonal hay fever
or seasonal allergic rhinitis, including the use of symptomatic pre-
scriptive and nonprescriptive medication; contact with people at risk
for severe RSV infections; steroid use in the past month; chronic
sinusitis; and the presence of known immunosuppressive conditions.
All subjects’ nasal washes tested negative by real-time polymerase
chain reaction (PCR) on Study Day 21 for viral respiratory pathogens
(RSV-A and -B; influenza A and B; parainfluenza 1, 2, and 3; and
human metapneumovirus). Appropriate local regulatory, institutional
review board (University of Tennessee, Memphis, TN), and Ethics
Committee (East London and the City Local Research Ethics Com-
mittee 3, London, UK) approval was obtained as appropriate. All
subjects provided written informed consent.
Inoculating Virus (RSV)
RSV-A (Memphis 37 strain) was isolated and manufactured according
to Good Manufacturing Practice in Food and Drug Administration–
approved Vero cells from the respiratory secretions of an infant
hospitalized for bronchiolitis and who had known high viral loads of
RSV-A. The isolate was plaque-picked and passaged five more times to
prepare the inoculum. The identity of the inoculating virus (RSV) was
confirmed by an immunofluorescent antibody assay, electron micros-
copy, and N-gene sequencing. It was determined at several steps in the
selection and manufacturing process to be free of adventitial agents
and human pathogens by four methodologies: (1) 28-day culturing in
five indicator cell lines (MRC-5, Vero, MDBK, HeLa, and MEF) while
under specific RSV neutralization conditions (high concentrations of
RSV-specific monoclonal antibody). Routine cytopathic effect obser-
vations, hemabsorption, and hemagglutination at 14 days were per-
formed, followed by blind passage and identical repeat evaluations
after an additional 14 days (Bioreliance Inc., Rockville MD); (2)
a product-enhanced reverse transcriptase assay for the general de-
tection of retrovirus reverse transcriptase (Bioreliance Inc.); (3) an
expansive series of individual PCR assays designed to detect human
pathogens; and (4) electron microscopy. The Memphis 37 virus
preparation was diluted in 25% sucrose immediately before inoculation
of subjects to create the desired target inoculum for the specific cohort
(Figure 1b). Each volunteer in a cohort received an aliquot of the same
inoculum. Inoculation was by intranasal drops (0.5 ml/naris). To
calculate the amount of RSV inoculated into each volunteer within
a cohort, a quantitative culture (plaque assay in HEp-2 cells) was
performed in parallel on duplicate aliquots at the exact time of
inoculation of the first volunteer within a cohort and again in duplicate
at the exact time of inoculation of the last volunteer within a cohort.
The mean of these parallel aliquot quantitative culture results repre-
sents the cohort’s inoculum.
The study was conducted at a single quarantine unit. Thirty-six subjects
were enrolled through five sequential cohorts (A–E). Viral assays and
assessments were not completed on one subject because she voluntarily
withdrew from the study. Statistical analyses did not include this one
subject. Subjects were admitted to the quarantine unit for 13 days and
were observed for at least 1 day before RSV inoculation, which
occurred on Day 0. Nasal washes (5 ml of normal saline instilled and
withdrawn per naris) for viral assays and cytokine measurements were
obtained on the day of admission to the quarantine unit (Day 21) and
twice daily on Days 1 through 12. Pulmonary function tests were
performed daily on all volunteers, using a Viasis Micro-lab calibrated
spirometer. For assessment of upper respiratory signs and symptoms,
a physician’s daily directed physical examination (Days 21 to 12) and
a twice-daily subject-reported RSV symptom score card (Days 21 to
12) were completed. Details of these scoring instruments can be found
in the online supplement. Mucus weights for each 24-hour period were
recorded from Days 21 through 12. Adverse events were recorded
through the Day 28 follow-up visit.
Typical signs and symptoms of RSV infection captured in the
symptom or physical examination scores were not counted as adverse
events. No concomitant medications were allowed during quarantine
except acetaminophen for headache.
Nasal washes were collected into cold RSV stabilization medium
containing 25% sucrose, transported on ice, and placed onto HEp-2
cell monolayers within 30 minutes of collection. Parallel aliquots were
snap-frozen and stored at 2808C until use.
RSV quantitative cultures in HEp-2 cell plaque assays were performed
in 12-well plates with triplicate 10-fold dilutions of nasal wash as
1306AMERICAN JOURNAL OF RESPIRATORY AND CRITICAL CARE MEDICINE VOL 1822010
previously described (42). Plates were fixed and stained at 5 days with
hematoxylin and eosin. RSV quantitative standards (RSV-A Long)
(VR-26; American Type Culture Collection [ATCC], Manassas, VA)
were run in parallel with each plaque assay to ensure precision. Plaque-
counting rules were preestablished. Units are reported as log (base 10)
plaque-forming units per milliliter (log PFU/ml). The lower limit of
detection of the assay was set at 1.7 log PFU/ml.
The quantitative real-time reverse transcription-polymerase chain
reaction assay (qPCR) employed an ABI 7900ht sequence detection
system (Applied Biosystems International, Foster City, CA) amplifying
a portion of the N-gene as previously described and validated (43).
Each specimen was run in duplicate in 96-well plates with internal
standards of duplicate pairs of six 10-fold dilutions of RSV RNA
extracted from parallel aliquots containing a known quantity of RSV-A
Long (VR-26; ATCC) as defined by and as used in the plaque assay.
Results are presented as means of duplicates in log (base 10) plaque-
forming unit equivalents per milliliter (log PFUe/ml). The RSV
concentration in a single nasal wash collection is termed a viral load.
Viral loads below the level of detection were arbitrarily set at a value of
zero. Peak viral load is the highest viral concentration for a given
volunteer in any nasal wash. Area under the curve (AUC) viral load is
the area under the curve defined by a single volunteer’s multiple viral
loads collected twice daily. AUC was calculated by the trapezoidal
rule, using exact times of collection of each nasal wash.
Spin-enhanced cultures were produced, using 80% confluent
HEp-2 cell monolayers on coverslips within shell vials, and were
inoculated with 200 ml of fresh nasal wash and centrifuged at 700 3 g
for 60 minutes. Monolayers were acetone-fixed after 2 days. Coverslips
were evaluated for RSV by direct fluorescent antibody techniques
using RSV-specific mouse monoclonal antibodies (Bartels; Trinity
Biotech, Wicklow, Ireland).
Serum RSV-neutralizing antibodies were measured in an HEp-2
cell RSV 50% microneutralization assay as previously described (44)
but performed with the Memphis 37 strain. To maximize successful
infection, only volunteers with a relatively low serum RSV micro-
neutralization titer were included, representing approximately the
lower third of the normal distribution in healthy adults evaluated
during study screening (Figure 1a).
Cytokine and chemokine concentrations of IL-4, IL-6, IL-8, IL-10,
macrophage inflammatory protein (MIP)-1a, RANTES (regulated
upon activation, normal T-cell expressed and secreted), tumor necrosis
factor (TNF)-a, and IFN-g were measured in nasal washes from all
volunteers in cohorts B and C (n 5 15) on Days 1, 2, 4, 6, 8, 10, and 12.
Nasal wash aliquots were analyzed at neat, 1:10 and 1:50 dilutions using
chemiluminescent multiplexed sandwich enzyme–linked immunosor-
bent assay cytokine arrays (SearchLight, Aushon BioSystems, Billerica,
MA). Concentrations of cytokines/chemokines from an individual volun-
teer measured over all days were summed so as to represent the total
amount of cytokine/chemokine produced during the volunteer’s infection.
Statistical methods were based on International Conference on Har-
monization guidelines. Continuous values below the lower limit of
quantification were set at zero. Normally distributed continuous vari-
ables were evaluated by two-sample t test or by linear regression as
appropriate. Continuous data that were not normally distributed, or
noncontinuous data, were analyzed via the Wilcoxon rank-sum test. All
statistical analyses were performed with SAS (SAS Institute, Cary, NC)
version 8.2 or higher for Windows. Linear regression graphs were
created with Prism version 4.0a for the Macintosh (GraphPad Soft-
ware, San Diego, CA).
Subjects and Safety
Thirty-six healthy volunteers without signs, symptoms, or
a history of recent upper respiratory infection and with low
serum RSV-neutralizing antibody titers were selected from 523
persons screened (Figure 1a). Volunteers were entered into
quarantine and confirmed to be clear of RSV-A and -B,
influenza A and B, parainfluenza virus 1, 2, and 3, and human
metapneumovirus, based on RT-PCR testing of nasal washes.
They were then inoculated with a wild-type strain of RSV-A
(Memphis 37). Volunteers, mean age 26.7 years (range, 19–43 yr;
16 of 36 [44%] women), were inoculated in five sequential
cohorts (A through E). Each volunteer in a cohort received the
same quantity of inoculum of RSV. Subsequent cohorts were
inoculated with increasing quantities of RSV (Figure 1b). One
volunteer withdrew from the study and virologic assessments
were stopped, although safety assessments continued on this
individual. Disease was restricted to the upper respiratory tract
with no medically significant pulmonary function changes oc-
curring. No serious adverse events occurred; all adverse events
were mild or moderate in severity and no trend toward in-
creasing frequency or severity of adverse events was noted with
increasing RSV inoculum. No medically significant ECG or
pulmonary function abnormalities occurred in any volunteers.
One volunteer developed bilateral otitis media. No conjunc-
tivitis, sinusitis requiring antibiotics, or clinical pneumonia
Characteristics of Viral Infection and Disease
RSV was quantified by two independent assays: a quantitative
viral culture assay (plaque assay) measured as log plaque-
forming units per milliliter (log PFU/ml) and a quantitative
reverse transcriptase-polymerase chain reaction technique
(qPCR) amplifying the RSV N-gene, measured as log PFU
equivalents per milliliter (log PFUe/ml). RSV was also detected
nonquantitatively in a spin-enhanced culture and by RSV
antigen detection. For two representative individual volunteers,
RSV infection and disease are shown in Figures 2a and 2b. A
volunteer was defined as infected if RSV was detected in
respiratory secretions at two or more sequential time points,
the first occurring between Days 2 and 8 (inclusive) after
inoculation. Of the 35 volunteers evaluated virologically, 27 of
35 (77%) were defined as infected by using the more sensitive
qPCR assay and 21 of 35 (60%) were found to be infected as
defined by quantitative culture. This infection rate was consis-
tent between cohorts and did not vary by quantity of inoculum
received (Figure 1b). As assayed by qPCR, the incubation
period was 3.1 (61.5) days and the mean duration of viral
shedding was 7.4 (62.5) days. Duration of shedding was lower
as assessed by quantitative culture and spin-enhanced culture
(Table 1). Of those infected as defined by qPCR, 40.7% were
still RSV qPCR positive on Study Day 12. All volunteer nasal
washes were negative for RSV by qPCR on follow-up evalua-
tion approximately 1 month after inoculation. There was no
correlation between amount of RSV inoculum received and any
quantitative measures of infection (Figure 1b), or between
inoculum and disease (data not shown). Because of the lack
of different outcomes between the cohorts, the rest of the
analyses were conducted on all cohorts combined. There were
35 fully evaluated volunteers in the combined cohorts.
Composite infection dynamics, with the incubation periods
arbitrarily normalized to Day 1, showed that viral dynamics
were significantly different when measured by qPCR as com-
pared with quantitative culture (Figures 2c and 2d). The time
elapsed until peak viral concentrations in respiratory secretions
(peak viral load) was similar between assays. The peak viral
loads occurred 3.5 days after RSV became detectable by qPCR
and 3 days after first detection by quantitative culture (Figures
2c and 2d). This corresponded to Postinoculation Days 7 and 6,
respectively. However, the rate of viral clearance after peak
viral loads had occurred was noticeably less rapid by qPCR as
compared with quantitative culture (Figures 2a–2d). Volunteers
DeVincenzo, Wilkinson, Vaishnaw, et al.: Human Experimental RSV Infection1307
defined as infected demonstrated significantly higher subjective
and objective measures of disease compared with those defined
as uninfected (Figures 3a–3f, P 5 0.021, P 5 0.18, and P 5
0.0056 for symptom scores, physical examination scores, and
mucus weights, respectively).
Relationships between Virus and Disease
The onset of infection as measured by culture and qPCR
coincided with the onset of subjects’ disease. Within the day
after the first detection of RSV (as defined by quantitative
culture) the symptom and physical examination scores had
significantly risen above their pre-RSV detection baselines
(P 5 0.0003 and P 5 0.015, respectively). This was also true
for symptom and physical examination scores if infection was
detected by qPCR (P 5 0.06 and 0.0052, respectively). The
timing of peak viral load also coincided closely with the timing
of peak disease severity. These temporal relationships can be
seen in Figure 2e. Statistically, the peak viral load (qPCR)
occurred 1.17 (95% confidence interval, 0.11–2.22) days after
peak symptom scores had occurred and 1.07 (95% confidence
interval, 0.07–2.08) days after peak mucus weights occurred. As
viral load (qPCR) declined, the severity of disease subsided in
parallel (Figures 2 and 3). Viral loads as assessed by quantita-
tive culture declined much more rapidly, nearly all becoming
abruptly culture negative on Day 6 after first culture detection.
As quantity of virus detected in respiratory secretions in-
creased, disease severity also increased in a statistically signif-
icant manner (Figures 4a–4f). This direct association was
consistent, being demonstrated across both viral quantification
assays (qPCR and quantitative culture) and all methods of
calculation of viral quantity (peak viral load, duration of
shedding, and area under the curve [AUC] viral load) and
was statistically significant (for all, P , 0.05). This direct
association was also significant when the evaluations were
restricted to include only those volunteers who were defined
as RSV-infected (Figures 4d–4f). A further evaluation of the
effect of viral load on disease severity was undertaken by
examining the viral loads and disease severity measurements
at individual time points within individual volunteers. Within
the individual 27 RSV-infected volunteers (defined by PCR), 22
of 27 (81%) showed a statistically significant association be-
tween their viral loads and at least one of their measures of
disease severity (symptom score, physical examination score, or
mucus weight). When all individual time points for all infected
individual volunteers were evaluated together, viral loads (as
measured either by qPCR or by quantitative culture) correlated
significantly with symptom score, physical examination score,
and mucus weight (P , 0.0001 for all disease severity markers).
Volunteers with shorter incubation periods had greater
quantities of virus (P 5 0.0001 and P , 0.0001 for viral AUC
cohorts. (A) Histogram of serum respiratory syncytial virus
(RSV)–neutralizing antibody titers to Memphis 37 strain in
screened and entered volunteers. (B) Each successive
cohort of volunteers was inoculated with increasing quan-
tities of RSV. The mean quantity of RSV administered to
volunteers within individual cohorts was measured as log
plaque-forming units (PFU) in an HEp2 cell quantitative
culture assay (plaque assay). A volunteer was defined as
infected if two successive respiratory secretion collections
contained detectable RSV by quantitative reverse tran-
scriptase-polymerase chain reaction (qPCR). The mean
amounts of RSV in each cohort are illustrated here in
two ways. The top row of bar graphs plots the mean area
under the curve (AUC) of the viral loads for each of the
volunteers in each cohort. All volunteers were inoculated
with RSV but only 71–86% in each cohort produced
detectable virus, meeting the definition of infection. The
viral AUC (calculated by including only volunteers who
were defined as infected) are represented by solid columns
and the viral AUC calculated from all volunteers in the
cohort are represented by open columns. The bottom row
of bar graphs plots the mean peak viral loads of the
volunteers in each cohort. These peak viral loads were
measured by quantitative culture (qCulture) (log PFU/ml;
open columns) or by qPCR (log PFU equivalents/ml [log
PFUe/ml]; solid columns). No significant differences in
mean viral AUC or peak viral loads existed between
cohorts. Error bars represent the SD.
Screening immunology and viral outcomes of
1308 AMERICAN JOURNAL OF RESPIRATORY AND CRITICAL CARE MEDICINEVOL 1822010
Figure 2. Viral load and disease over time in human volunteers. (a and b) Viral and disease measurements for selected individual volunteers over
time. Volunteers were inoculated with respiratory syncytial virus (RSV) on Day 0 and evaluated in quarantine until Day 12. Twice-daily symptom
scores (open columns), once-daily directed physical examination (DPE) scores (solid columns), and daily quantification of nasal mucus weight (striped
columns) are measures of RSV disease severity. Viral testing was performed on nasal washes collected twice daily. The nonquantitative twice-daily
spin-enhanced culture (SEC) and daily RSV antigen test results are indicated as either 1 or 2 in the top rows of graphs. Quantitative real-time reverse
transcriptase PCR (qPCR, dashed line) and quantitative culture (qCulture; plaque assay on HEp-2 cells, solid line) were measured twice daily. (c and d)
Mean daily viral loads (qPCR and quantitative culture) with normalized incubation periods. The timing of individual volunteer viral load curves was
normalized by inserting a common incubation period so that their first positive viral measurement occurred arbitrarily on Infection Day 1. (c) Viral
load measured by qPCR; (d) viral load measured by quantitative culture. Error bars represent the SD. (e) Timing of mean viral load and symptomatic
disease. Mean data from all infected volunteers from each collection time point starting from Day 1 postinoculation are shown. No adjustment for
the variable incubation periods is included in this panel. This makes the breadth of the curves wider and the magnitude of the curves lower than
when incubation periods are normalized [as in (c) and (d)]. Initial rises in mean viral loads (qPCR and quantitative culture) correlate with the timing
of onset of the rise in mean symptom scores. The timing of peak viral load correlates with the occurrence of peak symptom severity. log PFUe/ml 5
log plaque-forming unit equivalents per milliliter.
DeVincenzo, Wilkinson, Vaishnaw, et al.: Human Experimental RSV Infection1309
and peak viral load, respectively, by qPCR, and P 5 0.0021 and
P 5 0.0020 for AUC and peak viral load, respectively, as
measured by quantitative culture).
Relationships between Immunity and Viral Loads
The mean preinoculation (Day 22) RSV microneutralization
titer of the volunteers was 7.11 MU/ml, and all but one titer fell
within the range of 5.29–8.79 MU/ml (Figure 1a). Within this
relatively narrow range of systemic immunity evaluated there
were no statistically significant correlations between preexisting
RSV microneutralization titers and the percentage of volun-
teers who became infected, the quantity of virus, the duration of
shedding, or any disease measures. However, greater quantities
of RSV detected in the volunteers did produce significantly
greater rises in their RSV microneutralization titer (DDay 22 to
Day 28, P , 0.0001, r25 0.61; P , 0.0001, r25 0.65 for viral
AUC by qPCR and viral AUC by quantitative culture, re-
spectively). Longer durations of shedding and higher peak viral
loads also correlated significantly with the production of
a greater rise in RSV microneutralization titer (for all, P <
0.007; data not shown).
Cytokines and Chemokines as Biomarkers
Cytokine and chemokine concentrations were measured in
parallel aliquots of nasal washes. The concentrations of these
individual cytokines and chemokines at all collected time points
MIP-1a,RANTES, IL-8,andTNF-aconcentrations appearedto
function as biomarkers of disease, because their concentrations
correlated most significantly and consistently with all disease
severity measures (P , 0.05 for symptom score, physical exam-
ination score, and mucus weight; Figures 5a and 5b). Cytokine
production appeared to be driven by the viral loads in these
nasal secretion concentrations correlated with quantitative viral
measures (P 5 0.004, r25 0.486, and P 5 0.04, r25 0.284,
respectively; Figure 5c). The concentrations of the Th2 cytokine
IL-4 did not correlate with disease severity and the concentra-
tions of this cytokine actually tended to be lower with greater
degrees of illness.
This experimental model contains features that mimic natural
infection: being initiated by a relatively low inoculum of a clinical
strain, having a defined incubation period, generating sustained
viral replication in an appropriate anatomic site, producing typ-
ical disease manifestations, and then self-resolving in immuno-
competent adults without sequelae. Limited prior experiences
with experimental RSV infection models primarily using labora-
tory-adapted RSV strains were safe in healthy adults (45–47).
TABLE 1. MEAN VIRAL INFECTION MEASURES THROUGHOUT
STUDY: COHORTS A–E COMBINED
AUC viral load,* log10PFUe/ml
Duration of shedding,* d
Incubation period,* d
N/A 10.1 (67.2)
Definition of abbreviations: AUC 5 area under the curve; log10PFUe/ml 5 log
(base 10) plaque-forming unit equivalents per milliliter; N/A 5 not applicable;
qPCR 5 quantitative reverse transcriptase-polymerase chain reaction.
Numbers in parentheses represent the SD.
* Calculated only for volunteers who met the definition of infection as
determined by the specific assay listed.
Figure 3. Disease measures over time in infected and uninfected volunteers. Volunteers were defined as infected if two successive samples of nasal
washes, collected between Study Days 2 and 8 inclusive, contained detectable respiratory syncytial virus (RSV). (a–c) Volunteers who became
infected; (d–f) volunteers who failed to reach the definition of infected. Infected volunteers showed significantly higher measures of disease than did
uninfected volunteers (all days combined). Mean total daily symptom scores, P 5 0.021 (a vs. d), mean total daily directed physical examination
(DPE) scores, P 5 0.181 (b vs. e) and mean daily nasal mucus weights, P 5 0.0056 (c vs. f). Error bars represent the SD.
1310AMERICAN JOURNAL OF RESPIRATORY AND CRITICAL CARE MEDICINE VOL 182 2010
The model reported here, using wild-type RSV, is safe and is also
highly reproducible. Five separate cohorts (A through E) of
volunteers studied over a 4-month period and inoculated with
a range of about 500-fold different viral quantities, produced
essentially the same quantitative viral and disease outcomes.
Faithfully reproducing natural RSV infection and disease
makes this model useful in proving the antiviral concept that
reducing RSV can reduce human disease and in evaluating the
effectiveness of specific RSV therapeutics before progressing to
studying such experimental therapeutics in the vulnerable
Animal models of RSV have several features that are likely
not parallel to human disease. Mouse models require large
nonphysiologic inoculums of RSV, approximately log 6–7 PFU
per animal, aspirated directly into their lungs, and respiratory
disease is not reliably produced. Conversely, the features of
Figure 4. Relationships between quantitative viral and disease measures. All volunteers were inoculated with respiratory syncytial virus (RSV) and
are represented in (a–c). (d–f) Analyses restricted to those volunteers who met the definition of infected. Disease measures (total symptom scores,
total directed physical examination [DPE] scores, and mucus weights) for individual volunteers are plotted against their viral area under the curve
(AUC). Viral AUC is the area under the viral load curves calculated for individual volunteers. P values represent the probability that the slopes of the
regression lines do not include a slope of zero. The dashed curved lines indicate the 95% confidence interval of the slopes of the regression line (solid
line). Similar statistically significant direct relationships were observed between viral and disease measures when viral AUC was measured by
quantitative real-time reverse transcriptase-polymerase chain reaction. qCulture 5 quantitative culture.
Figure 5. Relationships between IL-6 concentration, disease severity, and quantity of respiratory syncytial virus (RSV). Concentrations of IL-6 were
measured in respiratory secretions of volunteers and were compared with disease measures and viral quantities. (a) The cumulative sum of the IL-6
concentrations versus disease severity as measured by individual volunteer cumulative symptom scores. (b) The cumulative sum of IL-6
concentrations versus disease severity as measured by individual volunteer cumulative nasal mucus weight. (c) Comparison of the cumulative
sum of IL-6 concentrations versus area under the curve (AUC) viral load (quantitative real-time reverse transcriptase-polymerase chain reaction,
qPCR). P values represent the probability that the slopes of the regression lines do not include a slope of zero. The dashed curved lines indicate the
95% confidence interval of the slopes of the regression line (solid line). Similar statistically significant direct relationships were observed when viral
AUC was measured by quantitative culture.
DeVincenzo, Wilkinson, Vaishnaw, et al.: Human Experimental RSV Infection 1311
natural human RSV infection (upper respiratory tract inocula-
tion, appropriate incubation period, and typical upper respira-
tory symptomatic disease) are replicated in this experimental
infection model of wild-type RSV. Studying RSV infection
directly in humans reveals several important points, especially
the quantitative and temporal correlation between viral repli-
cation and disease. Viral load has been shown to be an
independent risk factor for RSV disease severity in children
experiencing their first RSV infection (48, 49), but this associ-
ation has been controversial and difficult to replicate (50, 51).
Here we convincingly demonstrate the relationship between
viral load and disease severity in this human experimental RSV
infection model. Increased viral load (peak, duration of shedding,
and viral AUC) is consistently and significantly associated with
both subjective and objective measures of increased disease
severity (Figure 4). Importantly, once infection is established in
a volunteer, increased viral replication is significantly associated
with both subjective (symptom scores) and objective (mucus
weight) measures of increased disease severity (Figure 4).
not support this Th2-mediated disease hypothesis. The two Th2
cytokines evaluated, IL-4 and IL-10, did not correlate with
disease severity, and IL-4 actually showed a trend toward
correlating inversely with disease severity. Conversely, other
non-Th2 proinflammatory cytokines and chemokines correlated
strongly with disease severity and also with viral loads. This
implies that Th2-driven immune responses may not be as im-
portant in human RSV disease as animal models have suggested.
The correlation between viral load and proinflammatory cyto-
kines IL-6 and IL-8 suggests that if viral load were reduced by
a robust antiviral, then disease severity might also be decreased
even if disease were mediated by these proinflammatory cyto-
kines, but studies employing an effective antiviral therapy in
humans need to be performed to directly prove this.
We have shown a close temporal association between onset,
peak, and clearance of viral replication and the onset, peak, and
resolution of the disease. This clearly argues against a patho-
genesis hypothesis involving infectious triggering of a delayed
pathogenic immune response and conversely supports the po-
tential clinical effectiveness derived by achieving a robust re-
duction in viral load through antiviral compounds.
Human experimental wild-type RSV infection described
here can inform us well regarding the pathogenesis of natural
human infections, even those occurring in infants and children.
However, when extrapolating these results to natural disease in
the pediatric population, certain potential differences should be
considered. First, infants are immune-naive with respect to RSV
whereas the adults studied here were not. Because of the
ubiquitous nature of RSV in all human populations, there is
no such thing as an immunologically RSV-naive adult. We
studied those adults with relatively low RSV-neutralizing an-
tibodies against Memphis 37 (Figure 1a), but it is likely that
even these adults had virologically significant serum neutraliz-
ing antibodies. The correlates of protection for RSV in adults
have not been determined, but serum neutralizing antibodies (in
the form of IgG) provide significant concentrations to the
epithelial lining fluid of the lower respiratory tract, but not
the upper respiratory tract, thus preferentially protecting the
lower respiratory tract (52). We observed essentially an upper
respiratory tract disease in this model, and this may, in part, be
due to these neutralizing antibodies preventing infection and
consequent disease in the lower respiratory tract.
Cell-mediated immune function, triggered and sustained by
multiple natural exposures to RSV throughout life, also likely
plays a role in limiting RSV replication and disease in the adults
studied. Infants with natural RSV infection likely have limita-
tions in both humoral and cell-mediated aspects of immune
control of RSV. Therefore we expect that infants would have
more extensive viral replication involving both upper and lower
respiratory tracts. Indeed, using the same qPCR assay employed
here, these adults had mean peak nasal viral loads of about 4.5
log PFUe/ml compared with an extrapolated mean peak of
RSV-A in naturally infected infants of more than 6 log PFUe/ml
(43). Extrapolation of the timing of disease from this model to
infants also bears discussion. Pathology studies and studies of
RSV-specific cell-mediated immune function in adults have not
been performed. It is unlikely that preexisting RSV-specific cell-
mediated immune function would produce earlier symptoms in
adults than in RSV-naive infants. This is because even severe
RSV disease (leading to respiratory failure and death) in infants
is produced despite the relative absence of such immune
effector cells in the lungs. Although difficult to perform, studies
of natural infections of children should be done to verify these
early viral and disease dynamics.
This study reveals a striking difference between the magni-
tude and duration of RSV infection quantified by traditional
approaches (quantitative culture and spin-enhanced culture) as
compared with real-time qPCR. The degree of prolonged RSV
shedding evidenced by qPCR in our study has not been
previously evaluated or described. It is possible that the pre-
viously accepted relatively short duration of culture-defined
viral shedding is not reflective of true viral replication in the
human respiratory tract. RSV has been shown to be neutralized
by antibodies, including IgA, and the disappearance of cultur-
able RSV corresponds to the onset of detectable RSV-specific
IgA in respiratory secretions of naturally infected infants (53).
Thus, it is possible that the previously recognized short course
of RSV infection in humans is a reflection of the virus being
neutralized within respiratory secretions despite its persistent
replication as shown by qPCR. There are alternative explana-
tions for the observed prolonged high levels of RSV detectable
by qPCR as compared with quantitative culture. It is unlikely
that the prolonged higher levels of RSV-specific nucleic acid are
caused simply by increased molecular detection thresholds
provided by PCR. If prolonged detectability were simply an
improved molecular detection phenomenon, then the quantita-
tive viral load curves for an infected patient over the time
course of the infection would be expected to have the same
ratio of culturable virus compared with virus quantified by PCR.
That is, the viral load curves (qPCR vs. quantitative culture)
would simply run in parallel (qPCR being similarly higher
quantitatively than quantitative culture at all time points).
Observations of the data do not support this (Figures 2a and
2b). Rather, the viral load curves of qPCR and quantitative
culture appear to be parallel during the upswing of the in-
fection, but then the two curves dissociate after peak viral load
is achieved in an individual infected volunteer (Figures 2a and
2b). The ratios of N-gene copy number compared with cultur-
able viral particles for in vitro cultures of RSV-A, using the
same assays as employed here, has been measured as approx-
imately 150:1 (43). Another explanation for the prolonged viral
loads as quantified by PCR is that culturable (replication-
competent) RSV ceases to be produced in these volunteers at
a relatively early time point (before approximately Day 6 of
infection) and that after being produced, this virus becomes
replication incompetent or is somehow cleared faster from the
nasal washes compared with the clearance of components of the
virus detectable by nucleic acid amplification techniques (PCR).
These and other explanations deserve to be evaluated as they
have major pathogenesis, diagnostic, and transmission implica-
1312AMERICAN JOURNAL OF RESPIRATORY AND CRITICAL CARE MEDICINE VOL 182 2010
tions for RSV and other respiratory viruses. If true, this newly
measured prolongation of likely RSV replication has implica-
tions for pathogenesis and adds impetus for the development of
In conclusion, a safe and reproducible human experimental
modelof RSV infection and disease has been developed.It alters
establishing a significant direct relationship between viral repli-
cation and resultant clinical disease in humans and a parallel
effectiveness before being tested in vulnerable RSV-infected
pediatric populations. These findings and the future use of this
wild-type experimental human infection model relieve major
obstacles in the development of therapeutics for this important
Author Disclosure: J.P.D. has received consultancy fees from AstraZeneca,
Novartis, Retroscreen, Tibotec, Gilead Sciences, Arrow Pharma, MicroDose
Therapeutx, Roche (all $1,001–$5,000), and MedImmune ($5,001–$10,000);
he has received lecture fees from MedImmune and Abbott International (both
$5,001–$10,000); he has received industry-sponsored grants from Alnylam
Pharmaceuticals (more than $100,000, grants to the University of Tennessee
for sponsored research and contract research), and ADMA Pharmaceuticals
($50,001–$100,000, grants to LeBonheur Children’s Hospital for sponsored
research); he owns stock in Alnylam ($10,001–$50,000); he has received
sponsored grants from the NIH (more than $100,000). T.W. has no financial
relationship with a commercial entity that has an interest in the subject of this
manuscript. A.V. is employed by, and owns stock in, Alnylam Pharmaceuticals.
J.C. is employed by Alnylam Pharmaceuticals; he owns stock in Alnylam
Pharmaceuticals ($50,001–$100,000). R.M. is employed by Alnylam Pharma-
ceuticals; she holds patents along with Alnylam; she holds stock in Alnylam (more
than $100,000); she has received sponsored grants from the NIH (more than
$100,000). S.N. is employed by Alnylam Pharmaceuticals; he owns stock in
Alnylam ($50,001–$100,000). L.H. has no financial relationship with a commer-
cial entity that has an interest in the subject of this manuscript. P.M. has no
financial relationship with a commercial entity that has an interest in the subject
of this manuscript. A.M. has no financial relationship with a commercial entity
that has an interest in the subject of this manuscript. E.M. has no financial
relationship with a commercial entity that has an interest in the subject of this
manuscript. J.O. has received lecture fees from GSK, Roche, MedImmune, Sanofi
Pasteur, Baxter Healthcare (all $1,001–$5,000); he owns stock in Retroscreen
Virology. R.P. has no financial relationship with a commercial entity that has an
interest in the subject of this manuscript. R.M. has no financial relationship with
a commercial entity that has an interest in the subject of this manuscript. E.W.
has received sponsored grants from the NIH and CDC (both more than
$100,000). R.S. has no financial relationship with a commercial entity that has
an interest in the subject of this manuscript. P.D. is employed part-time by
Meridian Life Science, Inc.; he owns stock in Meridian Life Science, Inc. (more
than $100,000). R.A. is employed by Alnylam Pharmaceuticals; he owns stock in
Alnylam Pharmaceuticals (up to $1,000). R.L. has no financial relationship with
a commercial entity that has an interest in the subject of this manuscript.
Acknowledgment: The authors acknowledge and thank Obus Opute and Lynne
Batty for help with and inside the quarantine unit; Shobana Balasingam,
Stephanie Blanc, Maryanne Formica, David Konys, Sayda Elbashir, Ivanka
Toudjarska, and Svetlana Shulga Morskaya for laboratory and logistical expertise;
Karen Foote, Angela Ning, and Sally Garnet for their attention to detail; Nigel
Dodd, Leonid Zeitlin, and Vladimir Mats for statistical expertise; the altruism of
the subject volunteers themselves, and all for their dedication toward the goal of
improving the care of children.
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