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TYPE Original Research
PUBLISHED 05 August 2022
DOI 10.3389/falgy.2022.854038
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EDITED BY
Mary Prunicki,
Stanford University, United States
REVIEWED BY
Jan L. Ceuppens,
KU Leuven, Belgium
Padraic Fallon,
Trinity College Dublin, Ireland
*CORRESPONDENCE
Michelle M. Epstein
michelle.epstein@meduniwien.ac.at
SPECIALTY SECTION
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Determinants,
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Frontiers in Allergy
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PUBLISHED 05 August 2022
CITATION
Liu S-H, Kazemi S, Karrer G, Bellaire A,
Weckwerth W, Damkjaer J,
Homann O and Epstein MM (2022)
Influence of the environment on
ragweed pollen and their sensitizing
capacity in a mouse model of allergic
lung inflammation.
Front. Allergy 3:854038.
doi: 10.3389/falgy.2022.854038
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©2022 Liu, Kazemi, Karrer, Bellaire,
Weckwerth, Damkjaer, Homann and
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reproduction is permitted which does
not comply with these terms.
Influence of the environment on
ragweed pollen and their
sensitizing capacity in a mouse
model of allergic lung
inflammation
Shu-Hua Liu1, Sahar Kazemi1, Gerhard Karrer2, Anke Bellaire3,
Wolfram Weckwerth4,5, Jakob Damkjaer6, Oskar Homann7
and Michelle M. Epstein1*
1Laboratory of Experimental Allergy, Department of Dermatology, Medical University of Vienna,
Vienna, Austria, 2Institute of Botany, University of Natural Resources and Life Sciences, Vienna,
Austria, 3Department of Botany and Biodiversity Research, University of Vienna, Vienna, Austria,
4Department of Functional and Evolutionary Ecology, Molecular Systems Biology, University of
Vienna, Vienna, Austria, 5Vienna Metabolomics Center (VIME), University of Vienna, Vienna, Austria,
6ALK-Abelló A/S, Hørsholm, Denmark, 7Division of Pharmacology & Toxicology, Department of
Pharmaceutical Sciences, University of Vienna, Vienna, Austria
Common ragweed (Ambrosia artemisiifolia) is an invasive plant with allergenic
pollen. Due to environmental changes, ragweed pollen (RWP) airborne
concentrations are predicted to quadruple in Europe by 2050 and more
than double allergic sensitization of Europeans by 2060. We developed
an experimental RWP model of allergy in BALB/c mice to evaluate how
the number of RWP and how RWP collected from dierent geographical
environments influence disease. We administered RWP six times over 3 weeks
intranasally to the mice and then evaluated disease parameters 72 h later
or allowed the mice to recover for at least 90 days before rechallenging
them with RWP to elicit a disease relapse. Doses over 300 pollen grains
induced lung eosinophilia. Higher doses of 3,000 and 30,000 pollen grains
increased both eosinophils and neutrophils and induced disease relapses. RWP
harvested from diverse geographical regions induced a spectrum of allergic
lung disease from mild inflammation to moderate eosinophilic and severe
mixed eosinophilic-neutrophilic lung infiltrates. After a recovery period, mice
rechallenged with pollen developed a robust disease relapse. We found no
correlation between Amb a 1 content, the major immunodominant allergen,
endotoxin content, or RWP structure with disease severity. These results
demonstrate that there is an environmental impact on RWP with clinical
consequences that may underlie the increasing sensitization rates and the
severity of pollen-induced disease exacerbation in patients. The multitude
of diverse environmental factors governing distinctive patterns of disease
induced by RWP remains unclear. Further studies are necessary to elucidate
how the environment influences the complex interaction between RWP and
human health.
KEYWORDS
Ambrosia artemisiifolia, ragweed pollen, allergy, asthma, mice, environment, climate
change
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Liu et al. 10.3389/falgy.2022.854038
Introduction
Ambrosia artemisiifolia (short ragweed) releases pollen that
causes allergic rhinoconjunctivitis and asthma. Allergic disease
induced by RWP is on the rise in Europe because of a significant
increase in the number of plants and the release of high
concentrations of pollen. The underlying cause appears to be
related to climate change. For example, higher temperatures,
changes in precipitation, atmospheric gases like CO2and NO2
and air pollution prolong the growing season, increase the
spread of the plant, and the number of pollen grains in the air.
Computer models predict a quadrupling of pollen concentration
in Europe by 2050 and the spreading of plants and pollen to
previously unaffected countries like Scandinavia and the UK (1–
7). Based on climate change associated increases in airborne
pollen, experts predict that the number of ragweed-sensitized
Europeans will more than double from 33 to 77 million by
2060 (8).
Several studies have followed clinical allergies related
to RWP. One pan-European study showed that ragweed
sensitization rates were about 10% across Europe and up to
58% in Hungary (9), with a clear correlation between RWP
levels, symptoms of rhinitis or asthma, medication use, and
medical consultations (10–13). One study reported a particularly
strong association between sensitization in children and RWP
concentration at levels higher than 5,000 RWP grains m−3
year−1(14). The concentration of airborne pollen correlates
with allergic sensitization and disease severity but may also
relate to environmental changes that influence the quality of
the pollen. In other words, could the environment make the
pollen more allergenic? To address this question, we created an
experimental RWP mouse model of allergic lung inflammation
to test increasing doses of RWP and pollen from diverse
geographical and climatic environments to evaluate allergen
sensitization and disease severity.
Materials and methods
Animals
Female BALB/c mice (6–8-week old) purchased from
Charles River Laboratories Inc. (Sulzfeld, Germany) were
Abbreviations: AHR, airway hyperresponsiveness; AUC, area under
the curve; BAL, bronchoalveolar lavage; BSA, bovine serum albumin;
Cdyn, dynamic compliance; CO2, carbon dioxide; H2SO4, sulphuric
acid; H&E, hematoxylin and eosin; HDM, house dust mite; HRP,
horseradish peroxidase; i.n., intranasal; i.p., intraperitoneal; NH4NO3,
ammonium nitrate; NO2, nitrogen dioxide; OVA, chicken egg ovalbumin;
PBS, phosphate-buered saline; PAS, periodic-acid-Schi; RI, airway
resistance; RT, room temperature; RW, ragweed; RWE, ragweed extract;
RWP, ragweed pollen; SEM, standard error of the mean; TMB, 3,3′
,5,5′-
Tetramethylbenzidine.
housed in a specific pathogen-free facility at the Medical
University of Vienna in a temperature-controlled environment
with a 12-h dark/light cycle and water and food ad libitum. All
animal experiments were done according to the Animal Care
Committee of the Medical University of Vienna and approved
by the Austrian Ministry of Education, Science and Research.
Ragweed pollen
RWP were generously provided by Allergon AB, Ängelholm,
Sweden (AG1, AG2), ALK-Abelló, Hørsholm, Denmark (ALK1,
ALK2) and were collected by our group in two sites in Austria
(VA1, VA2). For details about the pollen samples used in
these experiments see Supplementary Table 1. We calculated the
number of pollen grains per sample by weight based on a
previous study demonstrating roughly 300 RWP grains per µg
(15). All pollen were stored at −20◦C until use.
Amb a 1 levels
One dried sample of each RWP underwent extraction
in ammonium bicarbonate (NH4HCO3, VWR Chemicals—
Søborg, Denmark), using 125 mM buffer at pH 8.3. The ratio of
pollen to the buffer was 4.0 grams pollen to 23 ml of buffer. The
extraction was performed for 2 h with continuous stirring with
the pH adjusted for ∼10, 30 and 60 min. The pollen was then
spun down, and the supernatant was filtered through a 0.22 µm
filter. The samples then underwent radial immunodiffusion
(16). Briefly, agarose gel 1% in TRIS-Veronal buffer (Litex
HSA 1000 Agarose) (Lonza—Basel, Switzerland; TRIS from
Merck—Barbital (Veronal) Sigma-Aldrich—St. Louis, Missouri,
USA) was heated under stirring and afterwards kept in a
water bath at ∼56◦C for at least 15 min before use. The
agarose gel was combined with the appropriate volume of
a monospecific rabbit polyclonal Amb a 1 antibody (ALK,
in-house antibody production), mixed, and then cast on a
glass plate. When the gel had solidified, wells were punched
at appropriate distances. An ALK in-house reference sample
(IHR) in 4 concentrations, and the pollen extract samples, were
applied to the wells, and the gel was placed in a humidity
chamber (to prevent the gel from drying out) for 48–72 h. Each
sample was tested using two volumes (5 and 8 µl), and each
volume was tested in quadruplicate. Then, the gel was pressed,
washed and stained using a Coomassie Blue R 250 stain (Merck,
Darmstadt, Germany). An electronic image of the gel was
generated using a flatbed scanner (Hewlett Packard—Palo Alto,
CA, Supplementary Figure 1), and image analysis with Image-
Pro Plus 6.3 Software (Media Cybernetics, Rockville, MD) was
used to quantify the stained area of the IHR samples. The stained
areas of the extract sample wells were used to calculate the
sample concentration in each well using interpolation on the
standard curve for the gel. The extract sample concentration
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Liu et al. 10.3389/falgy.2022.854038
was then calculated using a mean estimate calculation and is
expressed as Amb a 1 Units/gram (U/g) pollen.
Assessment of pollen structure
Scanning electron microscopy was performed with air-dried
pollen samples that were sputter-coated with gold, and images
were obtained in a scanning electron microscope (JSM-6390;
JEOL, Peabody, MA). For capturing the images, the software
Scanning electron microscopy CONTROLE USER INTERFACE
version 8.24 (JEOL, Peabody, MA) was used.
Induction of allergic disease
To induce experimental allergic disease, we anesthetized
6–8 week old female BALB/c mice with 50 mg/kg ketamine-
hydrochloride (Ketanest, Actavis, Italy S.p.A, Nerviano, Italy for
Pfizer) and 2 mg/kg xylazine-hydrochloride (Rompun, Bayer
AG, Vienna, Austria). Then, we intranasally (i.n.) administered
RWP suspended in 50 µl PBS without added adjuvant or
PBS alone once a day for 6 doses on days 0, 2, 4, 14,
16, and 18. On day 21, at least 3 mice were assessed for
disease parameters, and the rest were left to recover for more
than 100 days (termed “recovered” mice). For disease relapse
(immunological memory), we instilled a suspension of 10 µg
of RWP in 50 µl PBS i.n. to recovered mice and 72 h after the
last allergen challenge, and we evaluated disease parameters.
RWP were suspended 30 min before i.n. administration of the
whole suspension and a fresh suspension was prepared for
each administration.
Airway inflammation
To evaluate airway inflammation, bronchoalveolar lavage
(BAL) was carried out via tracheostomy. A cannula was
inserted into the trachea and then flushed with a total of 1 ml
of PBS. The leukocyte cells in the BAL fluid were counted
with a hemocytometer (Neubauer chamber). The BAL fluid
was cytocentrifuged (Cytospin-4, Shandon Instruments, UK)
and then stained with Kwik-Diff (Thermo Fisher Scientific
Inc., Pittsburg, PA). Macrophages, eosinophils, neutrophils and
lymphocytes were then enumerated based on morphological
examination. A total of at least 300 cells per sample
were counted.
Lung inflammation and mucus secretion
Following the BAL, the lungs were resected and perfused
with a 4% paraformaldehyde fixative solution. The lungs
were placed in histology cassettes and embedded in paraffin.
Using a microtome (HM400, Microm, Heidelberg, Germany),
sections of 3 µm thickness were prepared and stained with
hematoxylin and eosin (H&E) or Periodic-acid-Schiff reagent
(PAS). Inflammatory cell infiltration and mucus secretion were
assessed with a light microscope (Olympus BX41, Olympus
Corp., Tokyo, Japan) and evaluated blindly according to
semi-quantitative scoring systems. For lung inflammation,
H&E-stained lung sections were graded for the intensity and
extent of the inflammatory infiltrates. The histological grade
was calculated as the product of the intensity and extent
of lung tissue involved. For inflammation intensity: Grade
0: no inflammatory infiltrates; Grade 1: few inflammatory
cells around airways/blood vessels; Grade 2: thin layer (<2
cells) of inflammatory infiltrates around airways/blood
vessels; Grade 3: thick layer (>2 cells) of inflammatory cells
around airways/blood vessels. For the extent of inflammation:
Grade 0: no inflammatory infiltrates; Grade 1: inflammatory
infiltrates in central airways; Grade 2: inflammatory infiltrates
extending to the middle third of lung parenchyma; Grade
3: inflammatory infiltrates spreading to the lung periphery.
For the assessment of mucus-production, PAS-stained
tissue sections were graded: Grade 0: no mucus-producing
cells in airways; Grade 0.5: few mucus-producing cells in
the central airways; Grade 1: high mucus production in
the central airways; Grade 1.5: sparse mucus-producing
cells in the middle airways; Grade 2: abundant mucus
production in the middle airways; Grade 2.5: little mucus
production in the peripheral airways; Grade 3: many mucus-
producing cells in the periphery of the lungs. Representative
photomicrographs of stained lung sections were taken with
ProgRes R
CapturePro 2.9.0.1 software using a Progress Speed
XTCore 5 camera (Jenoptik, Jena, Germany) coupled to a
light microscope.
Serum ragweed-specific antibodies
Blood was collected via cardiac puncture before the BAL.
Sera were acquired after coagulation of the blood samples and
centrifugation for 10 min at 15,000 rpm and frozen at −20◦C
until use. To measure ragweed-specific IgG1 and IgE antibodies,
96-well plates (Nunc Maxisorp, Thermo Fisher Scientific,
Roskilde, Denmark) were coated with 5 µg/ml of endotoxin-
free ragweed extract dissolved in PBS (Greer Laboratories,
Lenoir, NC) and stored overnight at 4◦C. On the following
day, plates were washed with washing buffer and blocked
with 2% bovine serum albumin (BSA) in PBS for 2 h at RT.
After washing, serial dilutions of sera were added, and the
plates were incubated for ∼24 h at 4◦C. Plates were washed
and incubated with biotinylated anti-IgG1 (Southern Biotech,
Birmingham, AL) or anti-IgE (BD Biosciences, San Jose, CA)
secondary antibodies for 2 h at 4◦C, followed by washing
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and incubation with streptavidin horseradish peroxidase (HRP,
Southern Biotech) for 1 h at RT. After washing, the plates were
incubated with 3,3′
,5,5′-Tetramethylbenzidine (TMB) substrate
solution in the dark for 10 min at RT and the reaction
was stopped by t he addition of a s top solution (0.18 N
H2SO4). Absorbance was measured with a spectrophotometer
at 450 nm.
Airway hyperresponsiveness
We measured airway hyperresponsiveness (AHR) as a
change in airway function at 24 h after RWP rechallenge in
response to increasing doses (0.39, 0.78, 1.56, 3.13, 6.25, 12.5
mg/ml) of aerosolized methacholine (Sigma-Aldrich, St. Louis,
MI). Anesthetized and tracheostomized mice were restrained
and ventilated using Fine PointTM Series Resistance/Compliance
equipment (Buxco Electronics Ltd.). Mice were exposed to
increasing doses of nebulized methacholine for 3 min with PBS
as the baseline. The FinePointTM (Buxco Electronics Ltd., NY)
software was used to calculate parameters of lung function:
Airway resistance (RI) and Compliance (Cdyn) of recovered and
relapsed mice in response to methacholine expressed as % of
PBS baseline.
Endotoxin measurements
Endotoxin was measured using the Pyrochrome Limulus
Amebocyte Lysate (LAL) test (Associates of Cape Cod, Inc.,
East Falmouth, MA) according to the manufacturer’s protocol.
Briefly, a 1:10,000 dilution for each pollen was prepared. The
samples and the control endotoxin standard (EC010; Associates
of Cape Cod, Inc.) (50 µl) were diluted with endotoxin-free
water (Associates of Cape Cod, Inc.) and mixed with 50 µl of the
LA-Lysate and chromogen substrate for 20 min at 37◦C followed
by 100 µl of a 50% acetic acid stop reagent. The Optical Density
(OD) was recorded at a wavelength of 405 nm.
Statistics
Data analyses were performed with GraphPad Prism
(Version 4.0a, GraphPad Software Inc., La Jolla, CA) applying
one-way ANOVA, Student’s t-test, Mann-Whitney test, and
chi-square test. All data are expressed as mean ±SEM,
and differences at p<0.05 were considered significant. For
AHR, the area under the curve (AUC) was calculated for
each experimental animal with resistance (y-axis) vs. the
methacholine concentration (mg/ml; x-axis) and analyzed with
Student’s t-test.
Results
RWP characterization
We sought to identify differences in allergenicity between
RWP samples obtained from distinct geographical locations
and environmental conditions (see Supplementary Table 1). We
selected cultivated ragweed plant populations from the USA
(from Allergon and ALK) and non-cultivated populations
collected in Austria. The pollen were from plant populations
grown in distinct environments. The AG and ALK populations
were cultivated, treated with fertilizers, and grown and harvested
in high temperatures with little precipitation. There were
differences in the climate between all the cultivated samples.
The collected VA1 and VA2 plants were not cultivated. The
VA1 plants grew in a rural meadow with some former soil
disturbance up to 2 m high, and t he VA2 pollen were from short,
stubby plants that were close to a heavily used highway and
were typically mowed several times a season and were mowed at
least once during the season when they were harvested. Neither
natural plant population was artificially fertilized or treated
with pesticides. Both sites offered relatively nutrient-rich soil
at comparable climatic conditions. The details of the climate
during the growing season up to the date of harvest were similar
without any expected interpretable differences. The AG1 pollen
differed from the others because it was the only one defatted
with acetone.
Based on the differences in the plant populations, we
sought to determine whether the pollen structure differed.
We examined the structure of RWP using scanning electron
microscopy (Figure 1C). The pollen at low power view (5,000×)
appeared similar with some slight differences within the
biological range. Minimal changes in the abortion rate for
pollen in sample VA1 compared to the other samples, and
some scattered debris in the AG2 sample was observed. We
did not detect fungi in the analyzed samples. However, fungal
contamination cannot be excluded from samples collected
from a natural environment. At high power magnification
(21,600×), all the pollen samples, except AG2, were similar
with intact surface and pollen kit. In contrast, the commercially
available defatted AG2 pollen was significantly different because
the acetone had removed the pollen kit exposing a porous
exine surface.
To further characterize the pollen, we measured the
concentration of the major Amb a 1 allergen because it is
possible that RWP from plants grown in distinct regions,
different seasons, and environmental conditions might differ in
the concentration of the allergens and lead to potential changes
in allergenicity. Figure 1A illustrates the calculated sample Amb
a 1 concentration obtained from the radial immunodiffusion
gels (Supplementary Figure 1). Indeed, RWP from plants grown
in distinct environments expressed variable amounts of Amb
a 1. AG1 and AG2 samples had 18 and 13 U/g of Amb a 1,
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Liu et al. 10.3389/falgy.2022.854038
FIGURE 1
Amb a 1 and endotoxin content, and structure of RWP from dierent geographical locations. The concentrations of (A) Amb a 1 in Units/gram
(U/g) and (B) Endotoxin in EU/ml in tested RWP samples. (C) Scanning electron microscopy photomicrographs were taken of AG1, AG2, VA1,
VA2 pollen to examine the structure at magnification of 5,000×in top row (scale bars =5µm) and 21,600×at bottom row (scale bars =1µm).
respectively; Amb a 1 content was 3 U/g in ALK1 and 21 U/g
in ALK2 and the samples collected in Austria had 8 and 5
U/g for VA1 and VA2, respectively. These data reveal that the
allergen content differed for all samples with AG1, AG2 and
ALK2 having the highest and ALK1 and the collected VA1 and
VA2 samples having the lowest concentrations.
In addition to allergen concentration, endotoxins adhered
to the pollen could also contribute to the immune response
by inducing a TLR4-dependent neutrophilic inflammatory
response (17). However, in Figure 1B, we show that the
endotoxin concentration in all samples was approximately the
same, indicating that the relative differences in reactivity to the
pollen would not result from endotoxin.
RWP dose and allergic airway
inflammation
To establish the model of experimental RWP-induced
allergic lung inflammation, we selected the untreated RWP
AG1 provided by Allergon, which was of high quality and
allergenicity with known growth conditions. We administered
a dose range of 0.1–100 µg (∼30–30,000 pollen grains/dose)
RWP suspension into the nose of mice once a day for 6
days over 3 weeks (Figure 2A). Three days after the last
pollen instillation, we harvested BAL fluid to assess airway
inflammation. We found that mice receiving PBS had 6.06 ±
0.63 x 104airway inflammatory cells/ml, which contained 95%
macrophages, few lymphocytes and almost no eosinophils or
neutrophils. Administration of as little as 0.1 µg or ∼30 RWP
grains boosted airway cell infiltration to 9.89 ±0.67 ×104
cells/ml with an increase of 11.62 ±1.5% eosinophils, 2.95 ±
0.5% neutrophils, and 3.38 ±0.4% lymphocytes. Administered
doses of 1 µg (∼300 grains), 10 µg (∼3,000 grains) and
100 µg (∼30,000 grains) induced increasingly higher total
airway cell numbers with increases in macrophages, eosinophils,
neutrophils and lymphocytes (Figure 2B), illustrating a dose-
dependent inflammatory response. The highest dose (100 µg)
lead to 7 times more inflammatory cells than in PBS control mice
with 31.58 ±5.1% eosinophils, 13.95 ±1.7% neutrophils, and
11.03 ±1.4% lymphocytes. The resulting mixed eosinophil and
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FIGURE 2
Dose-dependent acute onset of airway inflammation in
response to RWP. (A) Protocol scheme illustrating administration
of PBS alone and increasing doses of AG1 RWP 0.1, 1, 10, 100
µg per 50 µl PBS without added adjuvant over 21 days. (B) Total
inflammatory cell count and dierential counts in BAL 72h after
the last allergen instillation. Data are presented as mean ±SEM
and are representative of at least two experiments; n=8.
Asterisks indicate significant dierences between PBS and RWP
challenged groups, for total cells: *p<0.05, **p<0.01, ***p<
0.001, and for eosinophils: +p<0.05, +++p<0.001,
one-way ANOVA.
neutrophil infiltrate demonstrates that RWP induced a severe
form of allergic lung inflammation.
RWP dose and allergic lung inflammation
To evaluate the inflammatory infiltrates within the lung
tissue, we examined H&E-stained lung tissue sections. Naïve,
healthy lungs in untreated mice had no inflammation, while
intranasal challenge with RWP induced inflammatory infiltrates
(Figure 3A). With the lowest dose (0.1 µg), there were sparse
infiltrates predominantly near the central airways with few
eosinophils. In contrast, with 1 µg, there were infiltrates
in the central and peripheral airways with more infiltrating
eosinophils than with the lowest dose. At doses of 10 and 100
µg, there was extensive inflammation throughout the lungs,
with many infiltrates in the periphery containing numerous
eosinophils. Semi-quantitative scores for the severity and
extent of inflammation mirrored the dose-dependent response
observed in the airways with an increasing trend that correlated
with the dose and statistically significant differences for the two
highest doses (Figure 3A).
RWP dose and mucus hypersecretion
Another feature of allergic asthma is the high production
of mucus in the airways. We stained lung sections with PAS
to observe the mucus produced in the goblet cells and found
that PBS-treated mice have almost no cells producing mucus
(Figure 3B). In contrast, mice treated with the lowest pollen dose
(0.1 µg) had a few mucus containing cells present in the central
airways. At higher doses, we observed more mucus increasingly
in the central and peripheral airways, with the highest RWP
doses inducing substantially more mucus throughout the lungs.
Semi-quantitative scores for the extent of mucus in the lungs was
dose-dependent (Figure 3B).
Lung inflammation in response to diverse
RWP
To determine whether pollen from distinct environments
altered the level of sensitization and disease severity, we tested
the capacity of each pollen sample to induce allergic lung
inflammation. Based on the dose-dependent response in the
airways (Figure 2B), we selected the 10 µg dosing schedule.
Instilling about 3,000 pollen grains per dose induced a more
moderate inflammatory response, and we reasoned that it would
be optimal for comparing the pollen samples. We assessed
the extent of airway inflammation and found a large range
from 11.5 to 40.1 ×104cells/ml BAL fluid (PBS control
subtracted) of infiltrating inflammatory cells in the airways;
AG1 (40.1 ±6.2 ×104cells/ml) >VA2 (38.5 ±5.7 ×
104cells/ml) >AG2 (24.9 ±3.8 ×104cells/ml) >ALK2
(17.4 ±3.0 ×104cells/ml) >VA1 (16.3 ±3.7 ×104
cells/ml) >ALK1 (11.5 ±2.7 ×104cells/ml) (Figure 4).
When we evaluated the composition of the airway infiltrates,
we found that AG1 and AG2 had a mixed infiltrate consisting
of 57–60% eosinophils and 9–11% neutrophils, respectively.
In contrast, ALK2 and VA2 had a Th2-type inflammatory
infiltrate with between 54 and 60% eosinophils and <4%
neutrophils and ALK1 and VA1 had 39–44% eosinophils and
<5% neutrophils. RWP collected from different environments
qualitatively altered the magnitude and type of allergic response
in the airway.
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FIGURE 3
Dose-dependent lung inflammation and mucus production in response to increasing doses of AG1 RWP. Representative photomicrographs of
H&E- and PAS- stained lung tissue sections from PBS alone, 0.1, 1, 10, and 100 µg per 50 µl PBS AG1 RWP doses without added adjuvant over
21 days at a magnification of 40×.(A) In H&E-sections, PBS-treated mice had normal tissue without evidence of inflammation, whereas RWP
administration induced inflammatory infiltrates containing eosinophils (insets 400×). Arrows indicate inflammatory infiltrates. (B) In PAS-stained
lung tissue sections, naïve mice had no mucus in the goblet cells but RWP immunized mice had mucus in the goblet producing cells (arrows).
Scale bars are 500 µm. Graphs illustrate the quantification of the lung sections using a blinded semi-quantitative scoring system. Data are
presented as mean ±SEM and are representative of at least two experiments; n=8. Asterisks indicate significant dierences vs PBS, *p<0.05,
***p<0.001, chi-square test.
Inhalation of each of the RWP samples induced
inflammation in the central and peripheral airways (Figure 5A).
As in the BAL, mice immunized with AG1, AG2, VA2, and
ALK2 had extensive inflammation in the central and peripheral
lung parenchyma and contained many eosinophils and some
neutrophils. Similar to the airway response, ALK1 and VA1
induced far less lung inflammation with only sparse infiltrates
mostly confined to the central airways with few eosinophils.
Scores for the severity and extent of inflammation illustrated
these differences in a semi-quantitative fashion (Figure 5C).
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FIGURE 4
Airway inflammation during the onset of acute allergic disease
with diverse RWP from dierent geographical locations without
added adjuvant over 21 days. Total number of cells in BAL with
the distribution of macrophages, eosinophils, neutrophils and
lymphocytes. Data are presented as mean ±SEM minus PBS
control and are representative of at least two experiments; n=
8. Asterisks show significant dierences in total cell numbers: *p
<0.05, t-test. Significant dierences in eosinophils: ++p<0.01,
and in neutrophils: #p<0.05, Mann-Whitney test.
Mucus hypersecretion in response to
diverse RWP
In lung sections stained with PAS, all RWP induced mucus
hypersecretion in central and peripheral airways except for
ALK1 pollen, which caused less mucus production and was
predominantly in the central airways (Figures 5B,D).
Immunological “allergic” memory in
response to RWP
Because allergic asthma in patients is usually a relapsing-
remitting disease caused by a repeated encounter with an
allergen, we sought to mimic naturally occurring allergen-
induced relapse during the ragweed season. Figure 6A depicts
the protocol we used to induce disease relapse. We selected
three RWP samples with varying Amb a 1 content (AG1,
VA1, and VA2) to investigate allergic memory responses by
administering RWP intermittently over 3 weeks to induce
acute allergic lung inflammation (Figure 2A). The mice, then
recovered from the initial onset of disease for at least 90 days,
were once rechallenged intranasally with the same sensitizing
RWP (Figure 6A). Recovered mice had mainly macrophages
in their airways with few lymphocytes and neutrophils but
no eosinophils, illustrating that the mice had recovered from
acute allergic airway inflammation. Furthermore, they had
small, scattered infiltrates throughout the lung parenchyma
containing lymphocytes, macrophages and very few eosinophils
(Figure 7A), a pattern that differentiates recovered mice with
acute inflammation and naïve, healthy mice. In contrast,
RWP rechallenge induced a significant increase in airway
inflammation with a mixed eosinophilic and neutrophilic
infiltration compared to recovered mice. The relative increase
in airway inflammation at relapse was greater than at the
initial acute onset of disease illustrating a more robust
memory response. RWP similarly increased the total number
of airway inflammatory cells at relapse, though, eosinophilia
was significantly greater for VA2 than VA1 (Figure 6B) and
an intense mixed eosinophilic and neutrophilic inflammatory
response in the lung parenchyma which was similar for all
the tested samples (Figures 7A,C). Upon close examination
of the stained lung sections, there was no evidence of lung
remodeling which is similar to our OVA-induced allergic lung
inflammation model (18), in which recovered mice maintain
quiescent inflammatory infiltrates without remodeling unless
they are repeatedly rechallenged (19). Notably, RWP rechallenge
also boosted mucus production in airway goblet cells, which
added to the persisting mucus observed during recovery from
acute disease (Figures 7B,D).
Systemic immune responses in response
to RWP
A systemic RWP-specific IgG1 antibody response was
observed in the sera of mice immunized with 10 and 100
µg of RWP (Figure 8A,Supplementary Figure 2A). However,
immunization with 0.1 and 1 µg of RWP did not induce
specific IgG1, despite evidence of inflammation and mucus
hypersecretion in the lungs (Figures 2B,3A,B). These data show
that a dose equal to and lower than 300 pollen grains could not
elicit a systemic antibody response that doses over 3,000 pollen
grains were able to induce.
Although allergen-specific IgG1 and IgE in mice are Th2
class antibodies, we observed IgG1 but not IgE even in
undiluted sera (data not shown), suggesting that there could
be local IgE production but not systemic IgE. However,
we could also not detect IgE in the BAL fluid (data not
shown). Sera from mice immunized with RWP from different
sources had similar titers at the initiation of disease, except
for a lower titer in ALK1-immunized samples (Figure 8B,
Supplementary Figure 2B), which correlates with a low Amb
a 1 content, and other disease parameters, including lung
inflammation and mucus secretion.
During recovery, serum IgG1 was highest in mice
immunized with VA1 and VA2 and lowest in mice immunized
with AG1 (Figure 8C,Supplementary Figure 2C), which does
not correlate with Amb a 1 content or inflammation during
disease initiation. At disease relapse, IgG1 antibody titers
increased in AG1, were almost the same in VA2, and lower in
VA1 immunized sera. Several possibilities could explain these
differences; one is that the antibody titers were not measured
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FIGURE 5
Lung inflammation and mucus secretion during the onset of acute allergic disease with diverse RWP from dierent geographical locations.
Representative photomicrographs of H&E- and PAS- stained lung tissue sections from 10 µg per 50 µl PBS RWP without added adjuvant over 21
days at a magnification of 400×.(A) In H&E-sections, all RWP samples induced acute inflammatory infiltrates containing eosinophils (arrows),
and (B) in PAS-stained lung tissue se ctions, all RWP samples had mucus in the goblet producing cells (arrows). Scale bars are 50 µm. Graphs
illustrate the quantification of the lung sections using a blinded semi-quantitative scoring system for lung inflammation (C) and mucus
production (D). Data are presented as mean ±SEM and are representative of at least two experiments; n=8. Asterisks indicate significant
dierences *p<0.05 in the chi-square test.
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FIGURE 6
Airway inflammation during disease relapse elicited by AG1, VA1 and VA2 pollen. (A) Protocol scheme of disease relapse which illustrates the
induction of acute disease with the administration of six RWP doses over 21 days, a recovery period of at least 90 days, followed by a challenge
with a single 10 ug intranasal RWP dose before evaluating the mice at 72h. (B) Total and dierential BAL cell counts of mice recovered from
AG1, VA1, or VA2 pollen-induced disease and at relapse. Data are presented as mean ±SEM minus PBS control and are representative of at least
two experiments; n=4–8. Significant dierence in eosinophils: +p<0.05,t-test.
in sera from the same mice pre- and post-allergen exposure
and were from different groups of mice, which could lead to
variability between groups (Supplementary Figure 2C). Another
possibility is that the relapse antibody measurements were
done at 72 h after pollen challenge early after relapse and
would continue to rise over the following days. Nevertheless, the
relative differences between groups did not correlate consistently
with Amb a 1 allergen content or disease parameters.
AHR in response to methacholine during
disease relapse
We tested lung function during disease relapse as a
particularly important feature of exacerbations in patients
with allergic asthma. We found that pollen exposure induced
AHR in the presence of increasing doses of methacholine in
the RWP-rechallenged compared to the recovered mice, with
higher airway resistance (RI) (Figure 9A) and lower dynamic
compliance (Cdyn) (Figure 9B). Although, we attempted to
measure AHR in mice with acute onset of disease, the mice did
not survive the procedure.
Discussion
Here we show novel RWP-induced mouse models
resembling clinically-relevant allergic lung inflammation.
Mice administered whole un-manipulated RWP in the
absence of added adjuvants develop RWP-specific airway and
lung inflammation, mucus hypersecretion, antibodies and
allergen-specific immunological memory. A key observation
of this study is that the severity of the allergic response to
the pollen differed between samples collected in distinct
geographical locations as well as on the amount of pollen
administered. There was no apparent correlation between
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Liu et al. 10.3389/falgy.2022.854038
FIGURE 7
Lung inflammation and mucus secretion during disease relapse elicited by AG1, VA1, and VA2 pollen. Representative photomicrographs of H&E-
and PAS- stained lung tissue sections from AG1, VA1, and VA2 pollen (10 µg per 50 µl PBS) at a magnification of 400×.(A) In H&E-sections,
recovered mice have infiltrates without eosinophils, in contrast to relapse with inflammation containing eosinophils (arrows). (B) In PAS-stained
lung tissue sections, recovered mice have some mucus in goblet cells (arrows) compared with an increase in the rechallenged mice. Graphs
illustrate the quantification of the lung sections using a blinded semi-quantitative scoring system for lung inflammation (C) and mucus
production (D). Scale bars are 50 µm. Data are presented as mean ±SEM and are representative of at least two experiments; n=8.
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FIGURE 8
RWP-specific IgG1 serum titres. Allergen-specific antibody
responses against (A) titrated RWP (AG1) doses during acute
onset disease, (B) AG1, AG2, ALK1, ALK2, VA1 and VA2 pollen
from diverse geographical locations during acute disease onset,
and (C) AG1, VA1, VA2 pollen during relapse induction. Data are
presented as mean ±SEM [minus PBS control for (B) and (C)]
and are representative of at least two experiments; n=4–8.
disease severity and the concentration of Amb a 1, the major
allergen, endotoxin content, or alterations in the structure of the
pollen. Our pollen-specific allergic mouse models demonstrate
that both the quantity and quality of RWP influence allergic
responsiveness and disease severity, suggesting that climate and
other environmental factors affect the allergenicity of RWP. We
hypothesize that there is an environmental impact on RWP
with clinical consequences, which may underlie the increasing
sensitization rates and the severity of pollen-induced disease
exacerbations, which are likely to worsen in the future.
The natural approach to sensitization underscores the
importance of these experimental models of RWP allergic
lung inflammation. We instilled whole un-manipulated pollen
intranasally in the absence of systemic immunization (e.g.,
intraperitoneal immunization) or added adjuvants (e.g., alum).
Thus, the pollen includes the matrix and potential intrinsic
or adhered adjuvants, such as endotoxin, as in a natural
environment. Since the first mouse models of allergic lung
inflammation in the 1990s, they have become the most
frequently used species in allergy research (20–24). Many
experimental protocols induce disease with chicken egg
ovalbumin (OVA), house dust mite (HDM), purified or
recombinant allergens, and often added adjuvant or systemic
immunization is required to facilitate disease induction (25–
29). Most pollen-induced allergic disease protocols utilize
ragweed extract with or without adjuvants (e.g., alum) with
systemic sensitization (i.p.) followed by a respiratory challenge
(30–34) or with high RWP doses (e.g., 1 mg/dose i.n.) and
mainly with commercial ragweed samples without comparing
different lots (35–38). We focused on the natural sensitization
process occurring in the human respiratory tract. We developed
clinically relevant models of allergic airway disease to study
the allergenic potential of RWP from different locations
and environments.
Using our experimental model, we addressed the dose of
RWP necessary for sensitization. We found that 0.1 µg (∼30
pollen grains) of RWP instilled i.n., six times intermittently
within 3 weeks for a total of ∼180 pollen grains was sufficient to
sensitize mice and induce mild inflammation in the airways and
mucus secretion. Our data illustrate that the dose necessary for
sensitization in the mice is low and imply that only a few pollen
grains could sensitize individuals. However, it is very difficult
to compare the doses necessary to elicit a response in mice
in an experimental setting with the concentration humans are
exposed to over a whole pollen season, where clinical findings
show that a threshold of >5,000 RWP grains/ m3/year is
necessary for sensitizing patients (14). In an often-cited study,
the number of RWP grains required to elicit hay fever symptoms
was calculated (39). The authors applied RWP directly to the
nasal mucosa and found that 20,000 to 30,000 grains instilled
into the nose produced symptoms and then they estimated that
environmental ragweed-sensitive patients will have symptoms
with a concentration of 25 pollen granules/yard3based on a
person inhaling 20 cubic yards of air in 24 h resulting in a total of
500 pollen grains inhaled/day with the caveat that not all would
land on the mucous membranes. Typically, RWP sensitization
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FIGURE 9
Airway hyperresponsiveness of AG1-immunized mice during recovery and disease relapse 24 h after RWP challenge. (A) Airway resistance (RI)
and (B) Compliance (Cdyn) of mice during recovery and disease relapse (24 h after RWP challenge) in response to methacholine expressed as %
of baseline in response to increasing doses of aerosolized methacholine. Data are presented as mean ±SEM and are representative of at least
two experiments; n=3–4. Asterisks indicate a significant dierence between recovered and RWP rechallenged groups, **p<0.01,t-test of the
area under the curve (AUC).
occurs after 3–5 RW seasons and 5,000 pollen grains/m3/year,
an individual would need a total of ∼9,000–25,000 pollen
grains/m3for sensitization in 3–5 years, respectively. These
observations illustrate the capacity of RWP to cause disease
in mice at low concentrations compared to the dose needed
for humans.
We also found that increasingly higher doses of pollen
correlated with the mice developing more intense inflammation.
When we increased the dose to 1 µg (∼300 pollen grains), 10
µg (∼3,000 pollen grains) and 100 µg (∼30,000 pollen grains),
the number of airway inflammatory cells increased, most notably
the percentage of eosinophils and neutrophils, with 100 µg
leading to the most severe disease. Thus, titrated doses of RWP
caused increasing severity of acute airway and lung disease, as
previously shown in mice and patients (10, 11, 13, 14, 40, 41).
Because endotoxin is present in the samples, increasing pollen
dose also increases the endotoxin content and together they
could induce the more intense, mixed inflammatory response
that we observed. However, endotoxin is ubiquitous in a natural
environment and its content was similar between samples, it is
likely that it contributes equally to the immune response against
the tested pollen.
In addition to local lung responses, we also evaluated the
systemic reaction to inhaled RWP by measuring serum allergen-
specific antibodies. Surprisingly, we detected IgG1 antibodies in
mouse sera upon administration of 10 and 100 µg RWP, but not
with of 0.1 and 1 µg. These results suggest that the lower doses
of ∼300 pollen grains could induce local lung inflammation
and mucus hypersecretion without allergen-specific IgG1. It is
possible that the ELISA used in our experiments was unable to
detect low IgG1 titers with low RWP doses using our ELISA or
that after the last allergen challenge, there was a delay that could
enable us to detect IgG1 at a later time.
More puzzling is that we could not detect allergen-specific
IgE in any ragweed-immunized mice, which contradicts other
ragweed models in which both serum allergen-specific IgG1
and IgE were detected (37, 42, 43). However, these models
were done with substantially different protocols in which RWP
with alum were administered by intraperitoneal injection (37),
or RWP extract were instilled intranasally (42) or RWP were
administered intranasal over 5 weeks followed by an intranasal
challenge with Amb a 1 (43). Our results, however, support
previous findings in which immunized animals developed
allergic lung inflammation without the IgE (29, 38, 44). Another
possibility is that the IgG1 titers are so high that they mask
IgE. To address this issue is straightforward with single protein
allergens, like ovalbumin. Antigen-specific IgE can be unmasked
by excluding IgG1 with an ELISA in which the plate is
coated with anti-IgE, followed by the addition of biotinylated
antigen and then a conjugated streptavidin. However, this
approach is problematic for ragweed because the pollen are
a complex mixture of allergens, making the biotinylation
step more complicated. Notably, it is also possible that
our protocol induced IgE-independent disease, as previously
reported (19, 45).
After establishing the dose-dependent effect on disease, we
selected a moderate dose of 10 µg (∼3,000 pollen grains) for
testing differences between RWP from diverse environments.
At the same RWP doses, we observed significant differences in
the ability of the pollen samples to induce allergic disease. We
found that exposure to AG2 and AG1 induced extensive mixed
eosinophilic and neutrophilic inflammatory cell infiltration
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in the airway and lungs, which is remarkably similar to
inflammation in patients with severe asthma (27, 46, 47).
In contrast, VA2 and ALK2 induced a predominantly Th2
eosinophilic inflammatory response, whereas VA1 and ALK1
were both less aggressive and induced mild inflammation
compared to AG1 and AG2. ALK1 also induced the lowest
allergen-specific IgG1 titer. These data demonstrate that RWP
at the same dose but originating from distinct sources or
locations markedly altered disease severity and the composition
of inflammatory cells in the airways, suggesting that along with
the quantity, there are additional pollen properties that can make
the initiation of disease worse.
To further identify potential properties of the pollen samples
that influenced disease induction, we measured the content of
the major allergen Amb a 1, a pectate lyase recognized by over
90% of ragweed-sensitized patients and induces high antibody
levels in mice and humans (48–50). We found that the Amb a 1
content ranged from 3 to 21 U/g depending on the sample and
found no apparent correlation between allergen concentration
and disease severity. AG1 pollen induced the most severe disease
and had a high Amb a 1 content while the acetone-treated AG2
pollen samples had lower levels of Amb a 1 and induced less
severe disease. In contrast, ALK2 had high Amb a 1, but did not
cause as severe disease as AG1. The VA2 pollen were also highly
allergenic but had a low Amb a 1 content, suggesting that despite
being the immunodominant allergen in RWP, Amb a 1 content
does not appear to influence disease severity. This is supported
by studies showing that i.n. administration of Amb a 1 alone
does not cause airway inflammation or increase specific IgG1
(32, 44). However, we found significantly higher scores of tissue
inflammation as well as mucus production and RWP-specific
IgG1 with higher Amb a 1 levels in ALK2 compared to lower
levels in ALK1, suggesting that there is a partial impact of Amb
a 1 content.
The differences in Amb a 1 content, even though Amb a
1 does not correlate with disease severity could be attributed
to a multitude of environmental factors like location, weather,
timing of sampling and changes occurring during the storage or
processing, e.g., defatting which reduced Amb a 1 content and
may be related to acetone disruption of the pollen membranes.
Indeed, the AG2 pollen caused less intense inflammation but
a similar mixed eosinophilic, neutrophilic pattern compared to
untreated pollen, suggesting that other allergenic components
may be partially removed from the pollen upon acetone
treatment, but it is likely that the samples differed because they
were from different seasons and areas.
Environmental factors might also underlie our observed
disease-enhancement with specific RWP samples. A myriad
of factors could explain the differences including weather
conditions during the growing season, e.g., amount of rain,
temperature etc., soil properties, air pollution, use of fertilizers
or pesticides, repeated mowing or other mitigation approaches,
the timing of the harvest, time since collection, conditions for
pollen storage after collection, e.g., temperature and humidity.
For example, the VA2 pollen collected from the highway
roadside might induce more severe disease because of the
repeated mowing of the plants due to roadside maintenance
or heavy highway pollution. Pollution has been a factor
previously reported to alter pollen (51). Lectins, and other
contaminating air and ground pollutants (e.g., diesel fuel
particles, NH4NO3), or other particulate matter on the pollen
could potentially influence its allergenicity. Carbon dioxide
(CO2) increases the content of the major allergen Amb a 1
(52–54), and NO2up-regulates Amb a 1 encoding transcript
levels (55), increases Amb a 1 isoforms and other allergens,
with enhanced overall nitrosylation and increased Amb a
1 allergenicity (56). Furthermore, the increasing number of
ragweed allergic patients appears to correlate to high airborne
pollen concentration, which is associated with climate change,
i.e., elevated temperatures and CO2levels (3–6). The impact of
environmental factors was observed with allergenic birch pollen,
with ozone, increased temperatures, and pollution enhancing
the allergenicity by Bet v 1-nitration (57–60). Additionally,
urban birch pollen samples differed in protein expression
and chemotactic activity on human neutrophils (61), grass
pollen from plants exposed to cadmium, ozone, or an urban
environment had higher allergenic potential, and increased
Cupressaceae pollen from polluted areas had higher allergen
content (62–66). Although, it is tempting to speculate that
differences between the effects of VA1 and VA2 pollen are due
to pollution, testing of more replicates of rural or urban area
pollen sources would be needed. Furthermore, controlling the
environment during the growing of the ragweed plants will be
essential for determining the conditions that alter the pollen and
subsequently influence their allergenicity.
Other potential environmental factors including
contamination with endotoxin and fungi could potentially
explain our observations. However, we did not detect fungal or
spore contamination and the endotoxin content was similar for
all pollen samples, suggesting that these factors do not underlie
the differences between the pollen. Taken together, our findings
and previous reports suggest that changes in the environment
where ragweed plants grow or where their pollen are released
and transported may alter them in a way that could increase the
prevalence and severity of disease onset.
Allergic asthma in patients is a relapsing-remitting disease
with allergen inducing disease exacerbations. To determine
whether RWP-immunized animals also developed disease
relapse and to test the magnitude and character of the response
to different pollen samples, we elicited an allergen-induced
relapse based on an immunological ’allergic’ memory model
with purified OVA protein (18, 67). Disease relapse followed
acute disease onset and a recovery period of at least 90 days
before rechallenging the mice with one i.n. dose of 3 selected
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Liu et al. 10.3389/falgy.2022.854038
RWP samples. We found that the secondary challenge induced
a significant inflammatory response in the airways and lungs for
the tested pollen. The disease relapse was severe for each pollen
sample irrespective of the initial onset of disease or the Amb a 1
content. AG1, VA1, and VA2 significantly increased the numbers
of eosinophils and neutrophils, demonstrating that the disease
relapse was more severe than the acute disease. Furthermore,
the VA2 pollen caused even more inflammation than VA1 with
significantly higher airway eosinophil counts, similar to acute
onset disease. RWP induced AHR in response to methacholine
compared to recovered mice illustrating the full spectrum of
memory responses in the model. AHR was also tested in mice
with acute onset of disease, but the mice did not survive the
procedure, implying a remarkable, unexplained sensitivity to
methacholine during disease initiation.
RWP-induced disease relapse mimics disease exacerbations
in patients. But can the RWP dose for eliciting a response in
humans compare with mice? Using the findings of Feinberg
and Steinberg again, we note that 20,000–30,000 RWP grains
elicited symptoms in humans, while it only takes 3,000 RWP
grains to induce a robust relapse in mice. There are possible
explanations for disparate eliciting doses between mouse and
patient. Firstly, there are expected differences of the inhaled
RWP grain deposition in the lung based on the total fraction of
inhaled aerosol that deposits in the respiratory tract including
the nose and mouth, the respiratory tract deposited particle
dose rate, which is expressed as the concentration of particles
in the inhaled air by the minute ventilation which equals the
deposition in the respiratory tract over time and depends on the
morphology of the lungs and respiratory parameters including
respiratory rate and volume. Secondly, disease readouts differ.
In the experiment with humans, the values were for eliciting
hay fever symptoms, not the allergic lung inflammation, mucus
hypersecretion and AHR we tested in the mice. Thirdly,
differences could be attributed to the timing of symptoms.
There is a significant lag of 3–18 days after the first day
of pollen exposure for the development of ocular, nose and
lung symptoms (68). Despite differences in the RWP quantity
required to elicit a relapse, RWP-induced disease relapse in
mice mimics disease exacerbations in patients and provides a
valuable model for further study of diverse RWP and potential
therapeutic intervention.
In summary, our findings illustrate that inflammatory
cell recruitment is dependent on the quantity and quality
of RWP and differs greatly depending on environmental
factors of distinct geographical regions affecting the
RWP, which could lead to different sensitization rates
and disease exacerbation severity. Further investigation
is necessary to elucidate the contribution of pollen
characteristics (e.g., proteomics, metabolomics) and the
underlying mechanisms.
Data availability statement
The raw data supporting the conclusions of this article will
be made available by the authors, without undue reservation.
Ethics statement
This study was carried out in strict accordance with the
guidelines for the care and use of laboratory animals of the
Austrian Ministry of Science. The protocol was approved by the
Committee on the Ethics of the Austrian Ministry of Science
(Number: GZ: 66.009/0330-II/3b/2013). All painful procedures
were performed under anesthesia, and all efforts were made to
minimize suffering.
Author contributions
S-HL designed and performed experiments, analyzed the
samples and contributed to the manuscript preparation. SK
performed experiments and contributed to the manuscript
preparation. GK assisted in the collection of RWP. AB
acquired the scanning electron microscopy photomicrographs
and evaluated them. WW provided knowledge on pollen
characteristics. JD performed Amb a 1 measurements. OH
was responsible for the endotoxin assay. ME supervised
experiments, analyzed the data, and contributed to the
manuscript preparation. All authors read and approved the final
version of the manuscript.
Funding
This project was supported by the European Community
Framework Programme 7, ATOPICA (Atopic diseases in
changing climate, land use and air quality), grant agreement
no. 282687.
Acknowledgments
The authors would like to thank Markus Debiasi and
Ciprian B. Anea for their assistance in collecting the RWP in
Austria. We are grateful for the donation of the pollen samples
and advice provided by Magdalena Rahl from Allergon AB,
Välingevägen 309, SE-262 92, Ängelholm, Sweden, Lisa A. Myers
from Thermo Fisher Scientific, Carthage, MO, USA and Jay
Chism and Ed Browning from the University of Missouri, USA
for information on the AG pollen, and Jacob Illeman from
ALK-Abelló in Denmark.
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Liu et al. 10.3389/falgy.2022.854038
Conflict of interest
The authors declare that the research was conducted in the
absence of any commercial or financial relationships that could
be construed as a potential conflict of interest.
Publisher’s note
All claims expressed in this article are solely those of the
authors and do not necessarily represent those of their affiliated
organizations, or those of the publisher, the editors and the
reviewers. Any product that may be evaluated in this article, or
claim that may be made by its manufacturer, is not guaranteed
or endorsed by the publisher.
Supplementary material
The Supplementary Material for this article can be
found online at: https://www.frontiersin.org/articles/10.3389/
falgy.2022.854038/full#supplementary-material
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