Airway obstruction due to bronchial vascular injury after sulfur mustard analog inhalation
Livia A Veress1,3, Heidi C. O’Neill2,3, Tara B. Hendry-Hofer3, Joan E. Loader3, Raymond C.
Rancourt3, Carl W. White1,2,3
1Department of Pediatrics, 2Department of Pharmaceutical Sciences, University of Colorado
Health Sciences Center, Denver, Colorado; 3Department of Pediatrics, National Jewish Health,
Corresponding author: Carl W. White, M.D., Phone: 303-322-4002, Fax: 303-270-2189, Email:
Descriptor number: 6.11 Inhalational Disasters Science and Health
This work was supported by the National Institutes of Health grant 3U54ES015678.
Correspondence and requests for reprints should be addressed to Carl W. White, M.D.,
Department of Pediatrics, Pulmonary Division, National Jewish Medical and Research Center,
1400 Jackson Street, Denver, CO 80206. Email: email@example.com
Short running head: Airway obstruction by sulfur mustard analog
Word count for body of manuscript: 6531
At a Glance Commentary
Scientific Knowledge on the Subject: Among the effects of sulfur mustard (SM) on the
respiratory tract, airway obstructive cast development after acute exposure remains poorly
What This Study Adds to the Field: We demonstrate a rat model for a SM surrogate agent-
induced bronchial cast formation, and show that bronchial vascular injury occurring early after
exposure results in extravasation of plasma proteins, and leads to the development of airway
This article has an online data supplement, which is accessible from this issue’s table of content
online at www.atsjournals.org.
Page 1 of 72
Media embargo until 2 weeks after above posting date; see thoracic.org/go/embargo
AJRCCM Articles in Press. Published on July 16, 2010 as doi:10.1164/rccm.200910-1618OC
Copyright (C) 2010 by the American Thoracic Society.
Rationale: Sulfur mustard (SM) is a frequently used chemical warfare agent, even in modern
history. SM inhalation causes significant respiratory tract injury, with early complications due to
airway obstructive bronchial casts, akin to those seen after smoke inhalation and in single
ventricle physiology. This process with SM is poorly understood because animal models are
Objectives: To develop a rat inhalation model for airway obstruction due to SM analog, 2-
chloroethyl ethyl sulfide (CEES), and investigate the pathogenesis of bronchial cast formation.
Methods: Adult rats were exposed to 0, 5 or 7.5% CEES in ethanol via nose-only aerosol
inhalation (15 min). Airway microdissection and confocal microscopy were used to assess cast
formation (4 and 18 h postexposure). Bronchoalveolar lavage fluid (BALF) retrieval and
intravascular dye injection were used to evaluate vascular permeability.
Measurements and Main Results: Bronchial casts, composed of abundant fibrin and lacking
mucus, occluded dependent lobar bronchi within 18 h after CEES exposure. BALF contained
elevated concentrations of IgM, protein, and fibrin. Accumulation of fibrin-rich fluid in
peribronchovascular regions (4 h) preceded cast formation. Monastral blue dye leakage
identified bronchial vessels as the site of leakage.
Conclusion: Following CEES inhalation, increased permeability from damaged bronchial vessels
underlying damaged airway epithelium leads to the appearance of plasma proteins in both
peribronchovascular regions and BALF. The subsequent formation of fibrin-rich casts within the
Page 2 of 72
airways then leads to airways obstruction, causing significant morbidity and mortality acutely
Keywords: fibrin; pseudomembrane; plastic bronchitis; vascular permeability; microdissection
Abstract word count: 236
Page 3 of 72
Sulfur mustard, bis (2-chloroethyl) sulfide (SM), is a chemical agent used in modern
warfare, most recently by Iraq in the 1983-1988 Iran-Iraq war (1-6). It is a vesicant, affecting
mainly the skin, eyes, and respiratory systems shortly after direct contact (3, 7, 8). In high doses,
SM exposure can lead to multiorgan involvement (3, 4, 8, 9) and can result in death (3, 6, 10-12).
There are 40-50,000 surviving victims of SM inhalation in Iran and Iraq alone, many with
permanent pulmonary disabilities such as bronchiolitis obliterans (13, 14).
Respiratory effects of SM exposure have long been investigated, focusing mainly on
chronic respiratory effects in survivors (1, 2, 13-16). Regarding the acute effects of SM on the
human respiratory tract, scarce data is available other than a few case reports (2-4, 6, 8, 10, 11,
17). Injury appears concentration-dependent (5, 7), with low level SM exposure affecting mainly
the upper respiratory tract. This can result in nasal mucosal injury, rhinorrhea, loss of smell and
taste, pharyngeal mucosal injury, and laryngitis (2, 3, 7, 10). Moderate SM exposure results in
varying degrees of tracheobronchial mucosal injury, leading to a painful and forceful cough (2, 3,
7, 15). By contrast, high level SM exposure can often lead to more severely disabling respiratory
lesions that may cause death (6, 8, 10, 12). These effects include severe airways edema and
ulceration, tracheobronchial mucosal sloughing, and airway occlusive pseudomembranes (2, 4-7,
10-12, 15, 17-19).
Although high dose SM can cause fatality from multiorgan failure alone (2, 6), numerous
case reports have also documented sudden deaths occurring from acute airways obstruction
due to pseudomembrane formation (6, 7, 10, 12, 20). The etiology of this airway occlusion
remains poorly understood, as human exposure in recent years has been fortunately infrequent,
Page 4 of 72
and no animal model has yet been developed to specifically mimic this phenomenon. Therefore,
no therapies have been evaluated in this process, and none exist to prevent or alleviate this
potentially fatal event. As SM continues to be a potential agent in bioterrorism (5, 21), an
improved understanding of this disorder could allow development of effective therapeutic
2-Chloroethyl ethyl sulfide (CEES) is a surrogate agent used to mimic SM injury in
laboratories (22, 23). It is less toxic than SM due to the absence of one of two terminal chloride
groups (that crosslink DNA and proteins), and due to its considerably shorter half-life in aqueous
solution. While safer to handle than SM, CEES possesses many of the same damaging properties
as an alkylating agent (22, 23) as does SM, making it suitable for use in experiments to mimic
SM-induced respiratory tract injury without the need for a specialized containment facility. In
this paper, we provide an animal model for potentially fatal airway obstruction seen acutely
after SM exposure using the SM surrogate agent, CEES. In addition, we examine the
composition of these pseudomembranous casts, and demonstrate the significant role that early
injury to the bronchial circulation plays in their formation.
Words: 442 (+ 51 numbers in references)
Materials and Methods
An expanded methods description can be found in the online supplement.
Page 5 of 72
2-chloroethyl ethyl sulfide (CEES, 8.41 M) was obtained from TCI America (Portland, OR). All
other chemicals were purchased from Sigma-Aldrich Chemical Co. (St. Louis, MO) unless
The Institutional Animal Care and Use Committee (IACUC) of National Jewish Medical and
Research Center approved this study. Adult male (275-350 g) Sprague-Dawley rats (Harlan Co.,
Indianapolis, IN) were used.
Inhalation Exposure to CEES
Rats were anesthetized with a cocktail of ketamine (75 mg/kg), xylazine (7.5 mg/kg), and
acepromazine (1.5 mg/kg), and placed in polycarbonate tubes with sealing plungers. Tubes
containing animals were mounted in a nose-only inhalation system (CH Technologies, NJ), and
were delivered compressed air with the aerosolized compound (ethanol, 5 or 7.5% CEES in
ethanol) for 15 minutes. Aerosolization was conducted via a Razel syringe pump (Razel
Scientific, St.Albans, VT) connected to a BioAerosol Nebulizing Generator (BANG; CH
Technologies, NJ). After 15 minutes of exposure, rats were removed from polycarbonate tubes,
and were observed in their cages until fully recovered from anesthesia.
Animals were euthanized at 4, 18, or 72 h after exposure as per experimental design. If rats
became moribund, as demonstrated by weight loss >25% body weight, inability to eat or drink,
Page 6 of 72
etc., they were euthanized prior to the planned study termination as per IACUC protocol. Rats
within experimental groups were terminally anesthesized with pentobarbital (Sleepaway, Fort
Dodge Animal Health, Fort Dodge, IA), the tracheas were cannulated, and lungs were fixed at 20
cm H2O with either Karnovsky’s fixative or 4% paraformadehyde in phosphate-buffered saline
(PBS) for 10 and 30 min, respectively. Whole lungs were then removed by gross dissection.
The protocol for microdissection as described by Postlethwait et al (24, 25) was followed.
Microdissection was carried out on a petri dish. Beginning at the main lobar bronchus
(generation 3), the axial pathway was exposed by cutting the airway lumen at 3 and 9 o’clock
positions. Main daughter branches, or side branches, were also exposed during the
microdissection. A map of airway generations was drawn during microdissection (Figure 1).
Confocal Microscopy Using EthD-1 and YOPRO-1
Distribution of airway injury was assessed by using a three-dimensional imaging technique
modified after Postlethwait et al (24, 25). Main modifications to the previously described
protocol included the use of 20 ml/kg 6 µM ethidium homodimer-1 (EthD-1) (Molecular Probes
Inc., Eugene, OR) in phenol-red free RPMI medium used for vital staining of airways and their
contents. In addition, we also used 2 µM YOPRO-1 (Molecular Probes Inc., Eugene, OR) solution
in PBS for staining of right middle lobe airways after microdissection but prior to imaging. A
Zeiss Two Photon LSM 510 confocal microscope was used to obtain images and z-stacks. For
bronchial cast imaging, casts were removed via microdissection from their airway locations after
Page 7 of 72
EthD-1 labeling in situ, and then incubated with YOPRO-1 solution and imaged with the confocal
Differential Cell Counts in Bronchoalveolar Lavage Fluid
BALF was pooled and centrifuged, the pellet was washed in 2 ml PBS, then resuspended in 2 ml
PBS, and then centrifuged in a Cytospin (Shandon Scientific) followed by staining using a
modified Wright–Giemsa stain (Protocol Hema 3; Fisher Scientific, Fair Lawn, NJ, USA). Cell
counts were then obtained via hemacytometer, counting 200 cells minimum in three random
IgM and Total Protein Measurement in Bronchoalveolar Lavage Fluid
Two lavages with 5 ml of PBS each were instilled into the lungs via the tracheal cannula and
subsequently withdrawn and then pooled. IgM concentrations were quantitatively measured in
the BALF using standard ELISA protocol (Bethyl Laboratories, Inc., Montgomery, TX). Total
protein was measured in BALF using the bicinchoninic Acid (BCA) protein assay (Pierce, Rockville,
β-Fibrin Detection by Western Blot
β-Fibrin was detected in BALF by Western blot using polyclonal rabbit anti-human fibrinogen
(DAKO Cytomation, Denmark) or mouse monoclonal β-actin (Sigma, St. Louis, MO), followed by
an incubation with HRP-conjugated goat anti-rabbit IgG (Bio-Rad, Hercules, CA) or HRP-
conjugated rabbit anti-mouse IgG (Sigma), respectively. Rat fibrinogen (Sigma-Aldrich Chemical
Page 8 of 72
Co., St. Louis, MO) was used to generate fibrin standards. Results were normalized to protein
level within each sample.
Fibrinogen/β-Fibrin and Acetylated Tubulin Immunohistochemistry (IHC)
Avidin-biotin complex peroxidase methods were used on tissue sections for staining based on
VectaStain Elite ABC kit (Vector Laboratories, Burlingame, CA). The primary antibody used for
fibrin(ogen) IHC was polyclonal rabbit anti-human fibrinogen (DAKO Cytomation, Denmark), and
for acetylated tubulin IHC was monoclonal mouse acetylated alpha tubulin (Abcam, Cambridge,
MA), for 60 min incubations each. Secondary antibody used was biotinylated horse anti-mouse
IgG. Counterstaining was performed using hematoxylin.
Myeloperoxidase (MPO) activity assay
Snap-frozen lung tissue was homogenized, centrifuged, supernatant withdrawn, and steps
repeated until clear. The 1-ml reaction cuvette (with PBS, H2O2, and TMB) was followed for
3 min at 652 nm using a Beckman DU-64 spectrophotometer (Beckman Coulter, Fullerton, CA,
USA). Calculated milliunits of activity was normalized to milligrams of protein using the BCA
protein assay (Thermo Scientific).
Tissue Preparation for Histology
Paraffin embedded tissues were sectioned at 5 µm thickness, and then stained with hematoxylin
and eosin. Additional slides were also stained with a combined Alcian blue (AB) and periodic
acid Schiff (PAS) stain for the localization of acidic and neutral mucins, respectively, and
Page 9 of 72
counterstained with hematoxylin. In addition, Movat’s pentachrome staining was performed on
72 h lung sections for assessment of collagen deposition.
Evans Blue Dye Extravasation
Vascular permeability changes were assessed by monitoring the extravasation of Evans blue dye
(Sigma-Aldrich Chemical Co., St. Louis, Mo) using a method modified after Evans et al (26, 27).
Briefly, 45 min before they were killed and their lungs collected, the animals were injected via
tail vein with 30 mg/kg of Evans blue dye. After fixation, lungs were microdissected and airway
images obtained with a Nikon SMX 1500 camera (Nikon Instruments, Inc.).
Monastral Blue B-Labeling of Permeable Vessels
Thirty minutes before they were killed, the animals were injected via a tail vein with 30 mg/kg
Monastral blue B suspension (Sigma-Aldrich Chemical Co., St. Louis, Mo), a compound used to
label sites of vascular leak, at a concentration of 1 mg/ml (28). Animals were euthanized 4 h
after exposure, lungs were harvested, fixed, microdissected and imaged as described.
For statistical analysis, Prism 5.01 software (GraphPad, La Jolla, CA) was used. Results are
presented as mean ± SEM in the text and figures. Groups were subjected to one-way analysis of
variance (ANOVA), and when significance was found, Tukey’s post-hoc analysis was applied. A p
value < 0.05 was considered significant.
Page 10 of 72
Assessment of airway cast formation
At 18 h after CEES inhalation, we found bronchial casts within all lobes, especially within
dependent lung regions such as the right lower lobe, the lower portion of the left lobe, and the
accessory lobe (Figure 2B). Cast formation with 5% CEES was inconsistent in both location and
degree of obstruction. By contrast to 5% CEES, 7.5% CEES inhalation produced reliable cast
formation and a more severe injury, with larger bronchial casts that often caused complete
airways obstruction of some lobes. Again, this was particularly evident within the dependent
lobes. Complete occlusion of all lobes was incompatible with survival and was noted during
necropsy of several non-surviving animals. Mortality rate with 7.5% CEES was 25% at 18 h, and
67% at 72 h, while 5% CEES caused no mortality at all time points examined (Table 1). With
ethanol exposure alone, no cast formation was observed in any airways (Figure 2A). Detailed
mapping of bronchial casts within the airways revealed that such casts extended from the
tracheal bifurcation to, at most, airway generation 15 of the axial pathway (Figure 1). Major
daughter generations also contained extensions of same casts for up to an additional 4 distal
Composition of airway casts
Since bronchial cast composition is likely related to underlying mechanism(s) resulting in
their formation, we next sought to classify the casts formed after CEES exposure. After rats
were exposed to 5% CEES for 18 h, the lungs were fixed and microdissection was performed on
the right middle lobe. Bronchial casts were then carefully removed in their entirety, and
Page 11 of 72
processed for immunohistochemistry, histology, or confocal microscopy. Immunohistochemical
examination revealed fibrin(ogen) in great abundance as a component of these casts (Figure 3, A
and B). Periodic acid Schiff/Alcian blue staining of cast sections did not demonstrate mucus
staining at 18 h (Figure 3C), indicating that casts formed after CEES exposure were not mucin-
based. Hematoxylin and eosin staining of airway cast sections demonstrated scattered
inflammatory cells dispersed throughout entire casts. The inflammatory cells appeared to be
concentrated heavily along the edges of the casts, particularly by 72 h after exposure (Figure 3E).
Collagen deposition was noted by 72 h, with the appearance of spindle cells suggestive of
myofibroblasts or fibroblasts within the casts (Figure 3F). We also noted occasional clumps of
ciliated epithelial cells by 18 h (Figure 3, C and D), deeply embedded within the periphery of the
casts. The presence of ciliated epithelial cells was confirmed via immunohistochemical staining
for acetylated tubin present in cilia (see Figure E1 in the online data supplement).
In order to assess if the cells within these casts and their adjacent airways were still
viable, we employed a confocal microscopy double staining technique using YOPRO-1 and EthD-
1 nuclear dyes to indicate live (green) or dead (red) cells, respectively (24, 25) . YOPRO-1 nuclear
dye was used to stain all cell nuclei as a “background” stain by study design, and EthD-1 nuclear
dye was used to indicate dead or dying cells with compromised cell membrane integrity. We
found that the majority of cells within the casts were YOPRO-1-positive, or live cells, especially
those cells located at the periphery of the casts (see Figure E2 in the online data supplement).
Dead cells, which stained positively with EthD-1, were only sporadically noted, and were seen
mostly within the core of the cast, where cells identified by histology appeared to be mainly
inflammatory in origin.
Page 12 of 72
Plasma-derived protein quantitation in BALF
The presence of fibrin-rich casts within airways after CEES implied leakage of the fibrin
precursor protein, fibrinogen (340 kDa as dimer), from surrounding vasculature into the airway
lumen, since fibrinogen is normally found only within blood plasma. Therefore, we next sought
to quantify the amount of fibrin(ogen) within the BALF after different inhaled CEES
concentrations, using Western blotting and densitometry for quantitation. We also assessed the
concentration of total protein present in the BALF, as well as that of IgM, a high molecular
weight immunoglobulin normally confined to the circulation (700 kDa pentamer). At both 4 and
18 h, we found a significant dose-related increase in BALF protein, IgM and most notably β-fibrin
at both CEES concentrations tested. As compared to levels in ethanol-exposed rats, with 5%
CEES exposure we observed a 3-fold increase at 4 h and a 6-fold increase at 18 h in BALF total
protein concentration (Figure 4A). After 7.5% CEES inhalation, protein in BALF was increased 4-
fold at 4 h and 10-fold at 18 h over ethanol. BALF obtained from naïve rats contained total
protein at concentrations similar to those found after ethanol exposure, which were minimal.
When IgM concentrations were measured in BALF, there was a 3-fold increase with 5% CEES at 4
h, and a 19-fold increase at 18 h as compared to ethanol (Figure 4B). The BALF IgM levels
further increased with 7.5% CEES to 4-fold at 4 h and 31-fold at 18 h. IgM was not detected in
BALF from naïve rat lungs. The concentration of β-fibrin (normalized to protein content) also
was significantly increased in the BALF after CEES exposure (Figure 4C). Inhalation of 5% CEES
resulted in a 4-fold increase at 4 h and a 10-fold increase at 18 h of BALF β-fibrin concentration
compared to ethanol levels. After 7.5% CEES inhalation, this increase was 5-fold at 4 h and 12-
Page 13 of 72
fold at 18 h over ethanol levels. Again, β-fibrin concentrations in BALF from naïve rats were
comparable to those observed after diluent (ethanol) exposure.
Cell differential counts and myeloperoxidase activity
To assess the role of inflammation in cast formation, we analyzed differential cell counts
of inflammatory cells in BALF of both 5 and 7.5% CEES-exposed rat lungs at 4 and 18 h (see
Figure E3, in the online data supplement). Macrophage levels gradually declined over time and
with higher CEES concentrations. The BALF absolute macrophage counts with 7.5% CEES showed
a 2-fold (4 h) and a 3-fold (18 h) reduction over ethanol-exposed levels, while with 5% CEES no
change in macrophages was observed at 4 h, and only a modest decrease at 18 h (1.3-fold). In
contrast, the BALF percent and absolute polymorphonuclear leukocyte (PMN) count increased in
a time dependent fashion but without significant CEES dose-response (Figure 5A; see Figure E3
in the online supplement). Only a minimal increase in PMNs was detected in the BALF at the 4 h
time point with either 5 or 7.5 % CEES. However, at 18 h there was a very significant 15-fold
increase in percent BALF PMNs with both 5 and 7.5% CEES. Ethanol exposure caused no
measurable increase in BALF PMNs, and showed comparable macrophage levels to naïve (data
In order to assess whole lung inflammation, we next evaluated the levels of
myeloperoxidase (MPO) in lung homogenates after CEES exposure. MPO is a peroxidase enzyme
predominantly present in neutrophils, thereby serving as a useful marker for the presence of
these granulocytes. Relative to ethanol exposure, 5% CEES inhalation resulted in a 3-fold (4 h)
and a 19-fold (18 h) increase in MPO, while 7.5% CEES inhalation resulted in a 2-fold (4 h) and a
Page 14 of 72
14-fold (18 h) increase. Levels from naïve animals were comparable to ethanol exposure (data
Assessment of vascular permeability by Evans blue dye
As localization of fibrin within the airway implies vascular injury, we next sought to
examine vascular permeability after CEES inhalation by tracing the extravasation of Evans blue
dye from permeable vessels. Evans blue dye binds to serum albumin (66 kDa), and its leakage
implies that blood vessels are permeable to proteins of this size or greater. Since casts were
‘well formed’ by 18 h, and plasma proteins were a major component of the casts, we assumed
that vascular leakage must precede cast formation. Therefore, we assessed for increased
vascular permeability at 4 h via the Evans blue dye extravasation method, prior to appearance of
any casts. Animals were injected by tail vein with Evans blue dye (30 mg/kg) 45 minutes prior to
necropsy after exposure to CEES or ethanol. Microdissection of all lobes was then performed in
order to localize dye leakage. Following CEES exposure, we noted extravasation of Evans blue
dye around the distal trachea and central bronchi (see Figure E4, C and D, in the online data
supplement), but no dye was detected after ethanol-only exposure (see Figure E4, A and B, in
the online data supplement). This effect was CEES concentration-dependent, in that dye
extravasation was greater in the 7.5% CEES group (data not shown). In addition to peribronchial
and peritracheal staining, regions around the central pulmonary vessels (both arteries and veins)
also demonstrated increased Evans blue staining after CEES inhalation compared to both
ethanol and naïve controls (see Figure E5, A and B, in the online data supplement). No
parenchymal staining was noted at any concentration tested, indicating that increased
permeability did not occur in the pulmonary microcirculation.
Page 15 of 72
Histological assessment of vascular leakage
As Evans blue dye is albumin-bound, it is a nonspecific indicator of vascular permeability.
Nevertheless, it was useful in localizing vascular injury to peribronchial regions after CEES
exposure. Damage to the bronchial vasculature was further suggested by examination of
histological sections of accessory lobes stained with H&E, which demonstrated significant edema
formation within the peribronchovascular space, both at 4 and 18 h after 5% CEES exposure (see
Figure E6, C and D, respectively, in the online data supplement). While the eosinophilic staining
of this fluid appeared to be patchy in distribution, its presence probably indicates an elevated
protein content of the fluid. In some areas, eosinophilic staining of such fluid accumulation was
less intense, possibly due to post-fixation and/or processing artifact. In eosinophilic staining
regions, edema formation appeared to be particularly intense adjacent to the adventitial layer of
large pulmonary vessels, with relatively less edema noted within the immediate peribronchial
space. No edema was noted in any of these areas in lungs of either naïve or ethanol-exposed
animals (see Figure E6, A and B, respectively, in the online data supplement).
Since bronchial casts were found to be composed of abundant fibrin, we next used
immunohistochemistry to more precisely localize fibrin(ogen) in the peribronchovascular
regions. Accessory lobes of ethanol-exposed and 5% CEES-exposed lungs were assessed 4, 18,
and 72 h after exposure. By 4 h, we detected increased fibrin(ogen) staining within the
peribronchovascular space, which was particularly intense within the perivascular regions
(Figure 6B). As expected, ethanol-exposed rat lungs did not demonstrate fibrin(ogen) staining
outside blood vessels (Figure 6A). By 18 h after CEES exposure, when casts were well formed,
we noted fibrin(ogen) staining persisting within the peribronchovascular space (Figure 6C),
Page 16 of 72
similar to that seen after 4 h, both as to location and intensity. By 72 h, fibrin staining within
peribronchovascular spaces diminished (Figure 6D), but strong fibrin(ogen) staining remained
within airway casts. Within peribronchovascular spaces where fibrin(ogen) staining was evident
earlier, increased collagen deposition was now detected by pentachrome staining. Both mature
(yellow staining) and immature (blue staining) collagen was noted, particularly within the
thickened subepithelial interstitium 72 h after CEES exposure (Figure 6E). No increased collagen
deposition was noted with ethanol exposure in any regions (Figure 6F).
Detection of increased vascular permeability by Monastral blue pigment
Although we could localize both edema and fibrin(ogen) staining within the
peribronchovascular space using histology and Evans blue dye labeling, we were unable to
identify which specific vessel(s) had increased permeability after CEES exposure by these
methods. Therefore, we employed Monastral blue pigment as a tracer to label vessels with
increased permeability (30 mg/kg, IV). Monastral blue pigment is unique in that it readily
crosses the endothelium of abnormally permeable blood vessels, but it can then become
trapped within the basal lamina of these vessels, thereby labeling these sites of extravasation.
We assessed rat lungs for vascular injury at 4 and 18 h after CEES or ethanol exposures, 30
minutes after Monastral blue pigment injection. With the aid of microdissection, we observed
numerous Monastral blue-labeled vessels immediately beneath the airway epithelium at 4 h
after CEES exposure (Figure 7, B and E), consistent with the anatomic location of the airway
bronchial vascular plexus (29). This intensity of Monastral blue labeling of bronchial vessels
persisted at 18 h (Figure 7, C and F), indicating a sustained increase in permeability, and
therefore injury, of bronchial vessels. Monastral blue-labeled vessels extended from the distal
Page 17 of 72
1/3 of trachea (see Figure E7B in the online data supplement) with deposition distally to airway
generation 12 of each lobar bronchus. This labeling was consistent with the location of the
bronchial circulation. In the trachea, we observed increased Monastral blue staining within the
intercartilagenous mucosal regions, with less intense staining within the cartilage rings
themselves at both 4 and 18 h (see Figure E8, B and D, respectively, in the online data
supplement). Monastral blue staining was concentration-related with respect to CEES
inhalation, with increased vascular staining intensity after 7.5% CEES exposure. Following
ethanol exposure alone, no Monastral blue labeling was detected in any lung regions at either of
these time points (Figure 7, A and D; and see Figure E7A, and Figure E8, A and C, in the online
data supplement). No pulmonary artery or pulmonary vein labeling was noted at any
concentration of CEES tested. In addition, Monastral blue labeling was never observed within
the pulmonary parenchymal microcirculation, again indicating a lack of pulmonary microvascular
injury often seen in many forms of inhalation insult associated with acute lung injury.
Confocal microscopic airway analysis
With use of Monastral blue pigment labeling, we were able to identify injured bronchial
vessels with increased permeability in subepithelial regions. If the vessels in subepithelial
regions were most affected after inhalation of this noxious agent, we hypothesized that the
epithelial surface itself was also potentially altered. Therefore, we next sought to identify the
distribution, type and intensity of epithelial surface injury after CEES inhalation exposure using
confocal microscopy. Microdissection was used to expose axial and daughter pathways of the
right middle lobe of each rat lung within the naïve, 5% CEES-exposed, and ethanol-exposed
groups at 18 h after inhalation. We evaluated the distribution of EthD-1-positive cells on the
Page 18 of 72
costal surface of each right middle lobe. Again, double staining was performed with YOPRO-1
and EthD-1 nuclear dyes to assess for cell death. YOPRO-1 nuclear dye was used to stain all cell
nuclei as a “background” stain by study design, and EthD-1 nuclear dye was used to indicate
dead or dying cells with compromised cell membrane integrity. We found that, after 5% CEES
exposure, casts were present within airway lumens by 18 h. After careful removal of these casts,
large collections of EthD-1 positive cells were noted at the site of cast attachment to the
epithelial layer at generation 3 of the main lobar bronchus (see Figure E9 in the online data
supplement). Distal to generation 3, no EthD-1-positive cells were seen within either axial or
daughter pathways (Figure 8C). More importantly, numerous gaps or voids were noted within
the epithelial layer after CEES exposure, where a lack of YOPRO-1 staining was evident. These
defects were approximately 1-10 cell diameters in size. Although they were distributed
throughout all daughter and axial generations, they appeared more prominently within the
proximal axial pathways. After ethanol exposure, we detected EthD-1-positive cells throughout
all pathways and generations (Figure 8B). Along the axial pathway, ethanol exposure resulted in
EthD-1-positive cells in a linear pattern of distribution (see Figure E10 in the online data
supplement), while bifurcation sites off the axial pathway into major daughter branches (areas
of greatest airflow turbulence) showed the greatest amount of EthD-1 staining. Daughter
branches also showed diffusely scattered EthD-1-positive staining after ethanol exposure. Gaps
or voids within the epithelial layer were never found after ethanol exposure. When naïve rat
airways were examined, we found occasional single EthD-1-positive cells within all distal
pathways (generations 22-25), but none within the proximal pathways (Figure 8A). No defects
within the epithelial layer were seen in naïve rats. When assessing the trachea, diffuse red
Page 19 of 72
staining was noted within the mid-tracheal epithelium after 5% CEES exposure at 18 h, indicating
cell death (EthD-1-positive) without detachment at this stage (see Figure E11C in the online data
supplement) compared to naïve or ethanol-exposed tracheas (see Figure E11, A and B,
respectively, in the online data supplement).
Our study demonstrated that airway casts produced after inhalation of the SM analog
CEES were composed of abundant fibrin, a plasma-derived protein, and that the casts were
consistently found only within the proximal half of the conducting airways (generation 2-15 from
the total of 25 airway generations microdissected routinely). In addition, we showed that other
plasma-derived proteins such as IgM also appeared within the airway lumen, indicating the
presence of an early vascular insult leading to increased permeability. With use of Monastral
blue pigment, we identified the bronchial circulation to be directly involved in the vascular
leakage and, thereby, formation of casts. Injured bronchial vessels caused extravasation of
plasma components into adjacent regions, which then appeared within airways. Once within
airways, plasma components organized into casts capable of causing obstruction and
compromising respiratory function.
Obstructive cast formation, often referred to as plastic bronchitis or cast bronchitis, is
not unique to sulfur mustard exposure. Airway casts have been found in children with
congenital heart diseases particularly after Fontan procedures (30, 31), as well as in patients
with asthma (32-34), cystic fibrosis (35), sickle cell disease with acute chest syndrome (36),
Page 20 of 72
allergic and infectious pulmonary states (37), and after burns or smoke inhalation injury (38-43).
While plastic bronchitis is not common, it is commonly fatal (44). Mortality rates reported for
congenital heart disease associated casts is 15 - 50 % due to complete airways occlusion (45).
For casts due to inhalation burn injury, mortality reaches 20-30% (43) . Reports of fatal plastic
bronchitis in children with asthma have also been reported (32), but with lesser frequency. The
composition of casts (fibrin versus mucin) has been used as a guide to their prognosis and
treatment. The Seear classification system (30) was developed to classify casts into two types:
Type I (fibrin casts with abundant inflammatory cells, particularly eosinophils); and Type II
(mucin casts with minimal cellularity). The presence of type I, or predominantly fibrin casts,
results in a more ominous course, as seen with congenital heart disease, inhalational burns, and
also in asthma. In our model, casts contained abundant fibrin. While some airway epithelial
cells and scant inflammatory cells were present in the “early casts” at 18 h, no eosinophils were
found. By 72 h, increased numbers of neutrophils appeared, with the addition of newly
deposited collagen and with the appearance of spindle cells (fibroblasts or myofibroblasts). A
potential explanation for the relative paucity of inflammatory cells within the “early casts” in our
model was that 18 h may be an insufficient amount of time for an extensive inflammatory
response to be fully manifested within the airway lumen. Indeed, analysis of BALF at 18 h
revealed an only minimal increase in absolute number of neutrophils over ethanol, further
demonstrating a present but subdued inflammatory response within the airways. While casts
from asthma, congenital heart disease or cystic fibrosis can take several days or weeks to form,
cast formation in our model was very rapid (by 18 h) after inhalation exposure. This timing of
cast formation in the SM model resembles most closely that seen after burns and smoke
Page 21 of 72
inhalation (39, 46, 47), extensively studied in sheep (41), where casts were formed within 24 h
after injury (39). While that model is similar to ours, it differs in three distinct aspects. First,
casts after burns and smoke inhalation contain not only fibrin but also eosinophils and mucus
(39). Second, casts after burns and smoke inhalation contain extensive sloughed epithelium
(39, 48), which was much less extensive in our model, appearing as distinct voids within the
epithelial surface. And third, the distribution of casts throughout the tracheobronchial tree in
burns and smoke injury is from the very proximal trachea down to the level of the most distal
bronchioles (47), which was not the case with CEES inhalation wherein tracheal casts were
Intrapulmonary conducting airways spanning from immediately past the carina to the
level of the terminal bronchioles at airway generation 15 (49) were the only locations in which
casts were found after SM analog exposure. Microdissection of both fixed and unfixed lungs
showed this similar distribution, where immobile casts were firmly attached to surrounding
epithelium in several locations along the bronchi. The respiratory bronchioles, alveolar ducts,
and alveoli were free of casts, as was the trachea. While the aerosol particle size (mass median
diameter of 0.6 – 0.7 µm) would predict strong deposition even to the level of the alveoli, there
was remarkably little evidence of injury to distal lungs noted by light microscopy, with no
evidence of fibrin deposition within the lung parenchyma. By contrast, fibrin deposition was
noted via IHC methods within the peribronchovascular space of airways containing plastic casts.
Patchy epithelial sloughing was also observed via confocal microscopy, mainly within the
mainstem bronchi and terminal bronchioles, but also along the distal trachea and respiratory
bronchioles where neither casts nor fibrin deposition were seen. While the exact mechanism
Page 22 of 72
for this epithelial sloughing has not yet been elucidated, we recently demonstrated that CEES
inhalation injury is at least in part dependent upon reactive oxygen and/or nitrogen species (50).
In addition, direct damage to the epithelial cytoskeleton may occur shortly after contact with the
inhaled CEES compound (unpublished data), which may not be visible via light microscopy. While
sloughed epithelium was noted throughout the entire airway at 18 h, fibrin cast formation and
peribronchial fibrin deposition were not. Therefore, loss of epithelial integrity is likely not the
sole cause of cast formation, albeit it may be a contributing one. Any degree of damage to the
epithelium, gross or microscopic, will expose the underlying submucosal and bronchial blood
vessels, potentially facilitating entry of CEES or its downstream reactive species into the local
circulation, thereby facilitating injury and increased vascular permeability. Indeed, when we
employed Monastral blue labeling to survey for highly permeable injured vessels, we
successfully localized vascular injury to the entire bronchial plexus. Interestingly, the
distribution of the bronchial circulation corresponded to the distribution of cast formation and
peribronchovascular fibrin deposition (noted via IHC) along the tracheobronchial tree. Together,
these findings suggest that airway epithelial injury may extend to involve the underlying
bronchial circulation leading to extravasation of plasma contents such as fibrin, and activation of
the clotting cascade within the airways. While adaptive benefits of increased vascular
permeability might include recruitment of inflammatory mediators and coagulation factors
designed to help in repair, we believe that the increased vascular permeability occurring after
SM analog inhalation is excessive, potentially contributing to deleterious acute and chronic
respiratory effects seen after exposure. Therefore, we next focused on the effects of this
Page 23 of 72
increased bronchial vascular permeability, particularly as it relates to leakage of fibrin and other
plasma-derived proteins into airways prior to cast formation.
The significant increase of plasma-derived proteins within the BALF of rats exposed to
the SM analog CEES indicated a vascular injury occurring early after exposure. A concentration
and time-related increase was seen between groups in BALF fibrin, total protein and large
molecular weight IgM. While significant increases were noted, we suspected that the actual
levels of these components in the CEES-exposed BALF were likely underestimated, as casts were
not included in the BALF analyzed. The increased levels of these proteins detected relative to
the diluent-exposed group indicated a substantial vascular alteration allowing the leakage of
these components out of the intravascular compartment. Evans blue dye injection in vivo
allowed us to confirm the vascular leakage by noting (via microdissection) blue-labeled albumin
extravasation into the peribronchovascular region from distal trachea to terminal bronchiole
(airway generation 17), corresponding to the bronchial circulation. Within these regions,
histological examination revealed edema formation and immunohistochemistry revealed fibrin
deposition. By 72 h after exposure, substantial spindle cells (potentially fibroblast or
myofibroblast) were noted within the peribronchovascular regions, where a large amount of
fibrin was noted earlier at 18 h. Both mature and immature collagen deposition was seen in
these areas (51), particularly within the subepithelial interstitium of the submucosa, as well as
within the surrounding regions. The pattern of collagen deposition seen 72 h after CEES injury,
together with the fibrinous obstructive cast formation with spindle cell invasion, resembles that
seen in the early stages of bronchiolitis obliterans (52). While bronchiolitis obliterans is a
known late complication after SM injury (1, 13, 16), its pathogenesis remains unclear. We
Page 24 of 72
believe that our findings of acute epithelial, submucosal, peribronchial and bronchial vascular
injury with subsequent obstructive cast formation and collagen deposition within the
intrapulmonary conducting airways may play an integral part in development of bronchiolitis
obliterans after SM inhalation. Further studies focusing on chronic lung injury model after SM
exposure will be necessary to understand the evolution of this process.
The extensive involvement of the bronchial circulation in cast formation after inhalation
injury has been seen and studied extensively in the sheep burn and smoke inhalation model (38,
40). While SM injury differs in several respects, the similarities of early bronchial circulation
involvement in cast formation seen in both models lead us to speculate that some of the
underlying mechanisms could be similar. It has been proposed that three main events occur
after burn and smoke inhalation which lead to early cast formation (38). First, abundant nitric
oxide production appears to lead to increased blood flow into the bronchial circulation (38, 53),
followed by an increase in activated neutrophils that bind to the endothelium of the bronchial
vessels. Thus, reactive oxygen and/or nitrogen species might be involved. Activation of adherent
neutrophils could then lead to increased vessel wall permeability (38, 54), leading to
extravasation of plasma components involved in coagulation within the airway lumen.
Simultaneously, the extrinsic coagulation cascade may become activated with an increase in
expression of tissue factor on pulmonary epithelial cells and alveolar macrophages (38, 55),
allowing cast formation to be initiated. In our model, markers of inflammation appear after cast
formation has begun, and only modestly. At 4 h after CEES exposure, when first signs of cast
formation become evident, BALF neutrophils were scant and lung homogenate myeloperoxidase
levels were virtually unchanged from control levels. At the same time, highly permeable
Page 25 of 72
bronchial vessels were abundant (as per Monastral blue labeling), the presence of fibrin within
the peribronchovascular space was pronounced, and BALF β-fibrin and protein levels were
significantly elevated. By 18 h after CEES exposure, when casts were fully formed a more
pronounced inflammatory response was evident. For this reason, we believe that while
inflammation was most likely a potentiator of CEES-induced airway injury, it was not the primary
cause of cast formation. We have recently shown that oxidant injury is an important feature of
CEES-induced airway injury, demonstrating strong attenuation of BALF plasma-protein levels
(IgM, protein) after 5% CEES by administering the catalytic antioxidant compound, AEOL 10150
(50). The role of oxidant injury in cast formation after CEES inhalation will require further
Several therapeutic strategies to limit cast formation have been studied in other causes
of plastic bronchitis, especially in the burn and smoke inhalation model. The main target for
these therapies has been the coagulation cascade, using anticoagulant (i.e. heparin) and
fibrinolytic (i.e. tissue plasminogen activator) strategies to reduce morbidity and mortality (31,
32). These same studies have yet to be performed after inhalation injury from SM or its
surrogate, CEES. Our rat model using the SM analog CEES is a useful model for conducting such
therapeutic and mechanistic studies.
In summary, we have shown that inhalation of high concentrations of the SM analog
CEES in rats reliably produces airway occlusive casts similar to those noted in the literature to
cause fatal obstruction in patients exposed to SM. We demonstrated increased permeability of
the bronchial circulation developing early after CEES exposure, causing leakage of plasma
proteins, including abundant fibrin(ogen) into airways. The resulting airway luminal casts
Page 26 of 72
formed then organize within 3 days after exposure, with the appearance of early histopathologic
changes suggestive of bronchiolitis obliterans. This study demonstrates and confirms the
usefulness of an approach combining airways microdissection and confocal microscopy using
dual vital dye staining, as originally described in ozone injury (24, 25), for localizing and
characterizing airways injury due to toxic inhalation and for determining disease pathogenesis.
In this study, we demonstrated a rat model for airway cast formation after inhalation of the SM
surrogate, CEES. This model will be useful in future studies further assessing mechanisms of cast
formation, potential progression to bronchiolitis obliterans, and therapeutic interventions to
limit both processes. (50)
Page 27 of 72
The authors thank the following people from National Jewish Research and Medical Center for
their contributions to our study: Brian Day, PhD from the Department of Medicine for his expert
advice on inhalation exposures; Steve Groshong, M.D. from the Department of Pathology for his
assistance with histological methods and interpretation; Tara N. Jones and Xiaoling Guo for their
expert technical assistance. The authors would also like to thank Todd Carpenter, M.D. from
University of Colorado Health Sciences Center Department of Pediatric Critical Care Medicine for
his generous gift of the Monastral blue pigment used in our study, as well as Edward M.
Postlethwait, PhD, and Michelle Fanucchi, PhD, both from the University of Alabama at
Birmingham, for their assistance and advice which allowed initiation of microdissection and
confocal microscopy techniques within our laboratory, and to Dallas Hyde, PhD, who
recommended that we take this approach in the model. The research is supported by the
CounterACT Program, National Institutes Of Health Office of the Director, and the National
Institute of Environmental Health Sciences, Grant Number U54 ES015678.
Page 28 of 72
1. Beheshti J, Mark EJ, Akbaei HM, Aslani J, Ghanei M. Mustard lung secrets: Long term
clinicopathological study following mustard gas exposure. Pathol Res Pract 2006;202:739-744.
2. Balali-Mood M, Hefazi M. Comparison of early and late toxic effects of sulfur mustard in iranian
veterans. Basic Clin Pharmacol Toxicol 2006;99:273-282.
3. Somani SM, Babu SR. Toxicodynamics of sulfur mustard. Int J Clin Pharmacol Ther Toxicol
4. Kehe K, Balszuweit F, Emmler J, Kreppel H, Jochum M, Thiermann H. Sulfur mustard research-
strategies for the development of improved medical therapy. Eplasty 2008;8:e32.
5. Sidell FR UJ, Smith WJ, Hurst CG. Vesicants. In medical aspects of chemical and biological warfare
textbook of military medicine. Washington, DC: Borden Institute; 1997. p. 197-228.
6. Zilker Th FN. S-mustard gas poisoning - experience with 12 victims. Clin Toxicol 2002;40:251.
7. Kehe K, Szinicz L. Medical aspects of sulphur mustard poisoning. Toxicology 2005;214:198-209.
8. Sinclair DC. The clinical features of mustard-gas poisoning in man. Br Med J 1948;2:290-294.
9. Hassan ZM, Ebtekar M, Ghanei M, Taghikhani M, Noori Daloii MR, Ghazanfari T.
Immunobiological consequences of sulfur mustard contamination. Iran J Allergy Asthma Immunol
10. Eisenmenger W, Drasch G, von Clarmann M, Kretschmer E, Roider G. Clinical and morphological
findings on mustard gas [bis(2-chloroethyl)sulfide] poisoning. J Forensic Sci 1991;36:1688-1698.
11. Kehe K, Thiermann H, Balszuweit F, Eyer F, Steinritz D, Zilker T. Acute effects of sulfur mustard
injury-munich experiences. Toxicology 2009.
12. Koch W. Direckte kriegserkrankung durch einwirkung chemischer mittel. In: Aschoff L, editor.
Pathologische anatomie. Leipzig, Germany: JA Barth; 1921. p. 526-536.
Page 29 of 72
13. Ghanei M, Mokhtari M, Mohammad MM, Aslani J. Bronchiolitis obliterans following exposure to
sulfur mustard: Chest high resolution computed tomography. Eur J Radiol 2004;52:164-169.
14. Khateri S, Ghanei M, Keshavarz S, Soroush M, Haines D. Incidence of lung, eye, and skin lesions as
late complications in 34,000 iranians with wartime exposure to mustard agent. J Occup Environ Med
15. Freitag L, Firusian N, Stamatis G, Greschuchna D. The role of bronchoscopy in pulmonary
complications due to mustard gas inhalation. Chest 1991;100:1436-1441.
16. Emad A, Rezaian GR. The diversity of the effects of sulfur mustard gas inhalation on respiratory
system 10 years after a single, heavy exposure: Analysis of 197 cases. Chest 1997;112:734-738.
17. Papirmeister B, Feister, AJ., Robinson, SI; Ford, RD. Medical defense against mustard gas: Toxic
mechanisms and pharmacological implications. Boca Raton, Florida, USA: CRC Press; 1991.
18. Prakash UB. Chemical warfare and bronchoscopy. Chest 1991;100:1486.
19. Pant SC, Vijayaraghavan R. Histomorphological and histochemical alterations following short-
term inhalation exposure to sulfur mustard on visceral organs of mice. Biomed Environ Sci 1999;12:201-
20. Willems JL. Clinical management of mustard gas casualties. Ann Med Milit Belg 1989;3S:1-61.
21. Borak J, Sidell FR. Agents of chemical warfare: Sulfur mustard. Ann Emerg Med 1992;21:303-308.
22. McClintock SD, Hoesel LM, Das SK, Till GO, Neff T, Kunkel RG, Smith MG, Ward PA. Attenuation of
half sulfur mustard gas-induced acute lung injury in rats. J Appl Toxicol 2006;26:126-131.
23. McClintock SD, Till GO, Smith MG, Ward PA. Protection from half-mustard-gas-induced acute lung
injury in the rat. J Appl Toxicol 2002;22:257-262.
24. Postlethwait EM, Joad JP, Hyde DM, Schelegle ES, Bric JM, Weir AJ, Putney LF, Wong VJ, Velsor
LW, Plopper CG. Three-dimensional mapping of ozone-induced acute cytotoxicity in tracheobronchial
airways of isolated perfused rat lung. Am J Respir Cell Mol Biol 2000;22:191-199.
Page 30 of 72
25. Joad JP, Bric JM, Weir AJ, Putney L, Hyde DM, Postlethwait EM, Plopper CG. Effect of respiratory
pattern on ozone injury to the airways of isolated rat lungs. Toxicol Appl Pharmacol 2000;169:26-32.
26. Dallal MM, Chang SW. Evans blue dye in the assessment of permeability-surface are product in
perfused rat lungs. J Appl Physiol 1994;77:1030-1035.
27. Evans TW, Rogers DF, Belvisi MG, Rohde JA, Chung KF, Barnes PJ. Endotoxin-induced plasma
exudation in guinea-pig airways in vivo and the effect of neutrophil depletion. Eur Respir J 1990;3:299-
28. Carpenter TC, Reeves JT, Durmowicz AG. Viral respiratory infection increases susceptibility of
young rats to hypoxia-induced pulmonary edema. J Appl Physiol 1998;84:1048-1054.
29. Charan NB, Baile EM, Pare PD. Bronchial vascular congestion and angiogenesis. Eur Respir J
30. Seear M, Hui H, Magee F, Bohn D, Cutz E. Bronchial casts in children: A proposed classification
based on nine cases and a review of the literature. Am J Respir Crit Care Med 1997;155:364-370.
31. Costello JM, Steinhorn D, McColley S, Gerber ME, Kumar SP. Treatment of plastic bronchitis in a
fontan patient with tissue plasminogen activator: A case report and review of the literature. Pediatrics
32. Wagers SS, Norton RJ, Rinaldi LM, Bates JH, Sobel BE, Irvin CG. Extravascular fibrin, plasminogen
activator, plasminogen activator inhibitors, and airway hyperresponsiveness. J Clin Invest 2004;114:104-
33. Perez-Soler A. Cast bronchitis in infants and children. Am J Dis Child 1989;143:1024-1029.
34. Matthay MA, Clements JA. Coagulation-dependent mechanisms and asthma. J Clin Invest
35. Waring WW, Brunt CH, Hilman BC. Mucoid impaction of the bronchi in cystic fibrosis. Pediatrics
Page 31 of 72
36. Manna SS, Shaw J, Tibby SM, Durward A. Treatment of plastic bronchitis in acute chest syndrome
of sickle cell disease with intratracheal rhdnase. Arch Dis Child 2003;88:626-627.
37. Brogan TV, Finn LS, Pyskaty DJ, Jr., Redding GJ, Ricker D, Inglis A, Gibson RL. Plastic bronchitis in
children: A case series and review of the medical literature. Pediatr Pulmonol 2002;34:482-487.
38. Murakami K, Traber DL. Pathophysiological basis of smoke inhalation injury. News Physiol Sci
39. Cox RA, Burke AS, Soejima K, Murakami K, Katahira J, Traber LD, Herndon DN, Schmalstieg FC,
Traber DL, Hawkins HK. Airway obstruction in sheep with burn and smoke inhalation injuries. Am J Respir
Cell Mol Biol 2003;29:295-302.
40. Barrow RE, Morris SE, Basadre JO, Herndon DN. Selective permeability changes in the lungs and
airways of sheep after toxic smoke inhalation. J Appl Physiol 1990;68:2165-2170.
41. Soejima K, Schmalstieg FC, Sakurai H, Traber LD, Traber DL. Pathophysiological analysis of
combined burn and smoke inhalation injuries in sheep. Am J Physiol Lung Cell Mol Physiol
42. Phillips AW, Cope O. Burn therapy. Ii. The revelation of respiratory tract damage as a principal
killer of the burned patient. Ann Surg 1962;155:1-19.
43. Fidkowski CW, Fuzaylov G, Sheridan RL, Cote CJ. Inhalation burn injury in children. Paediatr
Anaesth 2009;19 Suppl 1:147-154.
44. Dabo L, Qiyi Z, Jianwen Z, Zhenyun H, Lifeng Z. Perioperative management of plastic bronchitis in
children. Int J Pediatr Otorhinolaryngol;74:15-21.
45. Madsen P, Shah SA, Rubin BK. Plastic bronchitis: New insights and a classification scheme.
Paediatr Respir Rev 2005;6:292-300.
Page 32 of 72
46. Murakami K, McGuire R, Cox RA, Jodoin JM, Bjertnaes LJ, Katahira J, Traber LD, Schmalstieg FC,
Hawkins HK, Herndon DN, et al. Heparin nebulization attenuates acute lung injury in sepsis following
smoke inhalation in sheep. Shock 2002;18:236-241.
47. Pietak SP, Delahaye DJ. Airway obstruction following smoke inhalation. Can Med Assoc J
48. Enkhbaatar P, Murakami K, Cox R, Westphal M, Morita N, Brantley K, Burke A, Hawkins H,
Schmalstieg F, Traber L, et al. Aerosolized tissue plasminogen inhibitor improves pulmonary function in
sheep with burn and smoke inhalation. Shock 2004;22:70-75.
49. Plopper CG, Chu FP, Haselton CJ, Peake J, Wu J, Pinkerton KE. Dose-dependent tolerance to
ozone. I. Tracheobronchial epithelial reorganization in rats after 20 months' exposure. Am J Pathol
50. O'Neill HC, White CW, Veress LA, Hendry-Hofer TB, Loader JE, Min E, Huang J, Rancourt RC, Day
BJ. Treatment with the catalytic metalloporphyrin aeol 10150 reduces inflammation and oxidative stress
due to inhalation of the sulfur mustard analog 2-chloroethyl ethyl sulfide. Free Radic Biol Med;48:1188-
51. Cool CD, Groshong SD, Rai PR, Henson PM, Stewart JS, Brown KK. Fibroblast foci are not discrete
sites of lung injury or repair: The fibroblast reticulum. Am J Respir Crit Care Med 2006;174:654-658.
52. Estenne M, Maurer JR, Boehler A, Egan JJ, Frost A, Hertz M, Mallory GB, Snell GI, Yousem S.
Bronchiolitis obliterans syndrome 2001: An update of the diagnostic criteria. J Heart Lung Transplant
53. Murakami K, Enkhbaatar P, Yu YM, Traber LD, Cox RA, Hawkins HK, Tompkins RG, Herndon D,
Traber DL. L-arginine attenuates acute lung injury after smoke inhalation and burn injury in sheep. Shock
Page 33 of 72
54. Basadre JO, Sugi K, Traber DL, Traber LD, Niehaus GD, Herndon DN. The effect of leukocyte
depletion on smoke inhalation injury in sheep. Surgery 1988;104:208-215.
55. Idell S. Extravascular coagulation and fibrin deposition in acute lung injury. New Horiz
Page 34 of 72
Gross specimen of a microdissected right middle lobe following exposure (18 h) to CEES (7.5%).
Bronchial cast material (arrow) is present within the exposed central airway to axial generation
15. Daughter branches show extension of cast formation for an additional 4-8 generations.
Gross specimen of cross-sectioned accessory lobe main bronchi following aerosol exposure (18
h) to diluent (ethanol; ‘A’) or CEES (5%; ‘B’). Complete airway occlusion from cast material is
noted (arrow) with CEES.
Photomicrographs of central airways casts removed from central airways following 5% CEES
aerosol exposure, at 18 h (A-D) or 72 h (E-F), which were then processed for
immunohistochemistry (IHC), or stained with periodic acid-schiff/Alcian blue (PAS/AB) or
hematoxylin and eosin (H&E). (A) IHC staining for fibrin within the cast. (B) IHC staining for the
non-specific anti-rat IgG rather than antibodies directed against fibrinogen/fibrin (nonspecific
controls). (C) PAS/AB staining of cast at 18 h, showing no mucin staining. A ciliated epithelial
cell clump incorporated within the cast is seen (arrow), as well as adherent inflammatory cells.
(D) H&E staining of cast at 18 h, with a notable incorporated epithelial cell clump (arrow). (E)
H&E staining of cast at 72 h, with a large increase of inflammatory cells noted. (F) H&E staining
Page 35 of 72
of cast at 72 h, with the appearance of spindle cells suggestive of fibroblasts/ myofibroblasts
within the cast matrix (thick arrows). All pictures were taken at 40x magnification.
Airway protein components appearing 4 h and 18 h following CEES exposure. (A) Effect of CEES
on total protein concentration in BALF . Analysis done by BCA binding assay. (B) Effect of CEES
on IgM concentration in BALF. Analysis by ELISA. (C) Effect of CEES on β-fibrin in BALF.
Analysis done via Western blotting, with densitometry used to obtain concentration values.
Values are mean ± SEM (n=6 per group, except in the β-fibrin study for the CEES group (n=3).
*** p <0.0001, ** p <0.001, * p <0.05 for comparison of both CEES-exposed groups vs. both
naïve and ethanol groups.
BALF polymorphonuclear leukocyte (PMN) and whole lung myeloperoxidase (MPO) levels
following inhalation of CEES. (A) Effect of CEES (5 and 7.5%) on percent PMNs at both 4 and 18
h after exposure. (B) Levels of MPO activity in homogenized whole lung after CEES (4 and 18 h; 5
and 7.5%) is shown normalized to protein levels. Values are mean ± SEM (n=6 per group), with
*** P <0.0001, ** P<0.01, * P<0.05 for comparison of both CEES-exposed groups vs. ethanol
Fibrin deposition detected by immunohistochemistry staining in lung sections of accessory lobe
main bronchi in (A) diluent (ethanol; 18 h), (B) CEES (5%; 4 h), (C) CEES (5%; 18 h), and (D) CEES
Page 36 of 72
(5%; 72 h), as well as Movat’s pentachrome staining obtained 72 h after diluent (F) or CEES (E,
5%) exposure. Intense fibrin staining is illustrated in peribronchovascular space (arrow heads)
after CEES, as well as around large pulmonary vessels (thick arrows) adjacent to the main
bronchi (asterisks). Panel ‘C’ and ‘D’ show airway cast (thin arrow) with strong fibrin staining
(magnification 10x). Panel ‘E’ shows peribronchial and subepithelial (large arrows) deposition of
both immature (blue) and mature (yellow) collagen, as well as denudated ciliated epithelium
(arrowhead) and airway luminal cast (small arrow), neither of which is seen in lungs exposed to
diluent alone (panel ‘F’).
Gross specimen of accessory lobe main stem bronchi after Monastral blue pigment injection 4
and 18 h following exposures to CEES. Control rat airway (diluent exposed) shown in cross-
section (A) and en face (D) 4 h after Monastral blue injection. CEES-exposed (5%) rat airway
shown in cross section and en face 4 h (B and E, respectively) and 18 h (C and F, respectively)
after pigment injection. Note the diffuse Monastral blue-labelling of bronchial vessels (yellow
arrows) within the peribronchial space at both 4 and 18 h after CEES exposure. A bronchial cast
(black arrow) is present at 18 h (Panel ‘C’) within luminal air bubbles present.
Confocal microscopic analysis of central airways (axial pathway generation 5) utilizing double
staining for live (green) and dead (red) cells, as described in ‘Methods,’ following CEES inhalation
(18 h). Rats were (A) naïve, (B) diluent (ethanol)-exposed, or (C) CEES (5%)-exposed. “Gaps” in
Page 37 of 72
green staining of airway epithelium (arrow) were routinely noted in CEES-exposed proximal and
Page 38 of 72
TABLE 1. MORTALITY FROM CEES IN RATS
CEES, 2-chloroethyl ethyl sulfide
Ethanol 5% CEES 7.5% CEES
0 % (0/14) 0 % (0/14) 0 % (0/8)
0 % (0/14) 0 % (0/11) 25 % (7/28)
67 % (4/6)
0 % (0/8)
0 % (0/10)
Page 39 of 72
147x158mm (72 x 72 DPI)
Page 40 of 72
117x197mm (72 x 72 DPI)
Page 41 of 72
168x188mm (72 x 72 DPI)
Page 42 of 72
158x257mm (72 x 72 DPI)
Page 43 of 72
153x228mm (72 x 72 DPI)
Page 44 of 72
192x218mm (72 x 72 DPI)
Page 45 of 72
232x148mm (72 x 72 DPI)
Page 46 of 72
115x345mm (72 x 72 DPI)
Page 47 of 72
Airway obstruction due to bronchial vascular injury after sulfur mustard analog inhalation
Livia A Veress, Heidi C. O’Neill, Tara B. Hendry-Hofer, Joan E. Loader, Raymond C. Rancourt,
Carl W. White
Online Data Supplement
Page 48 of 72
Online Data Supplement Methods
The Institutional Animal Care and Use Committee (IACUC) of National Jewish Medical and
Research Center approved this study. The guidelines of the National Institutes of Health for the
care and use of experimental animals were carefully followed. Adult male (275-350 g),
pathogen-free Sprague-Dawley rats (Harlan Co., Indianapolis, IN) were used in these studies.
Animals were housed in cages in pairs, were provided free access to food and water, and were
subjected to a similar day-night light cycle. After arrival to Denver altitude (1600 m), rats were
allowed to acclimate for 1 week prior to use in experimental studies.
Inhalation Exposure to CEES
Rats were randomly allocated to either the naïve group (unexposed), the diluent-exposed group
(absolute ethanol) or the CEES-exposed groups (5%, equivalent to 420 mM, in diluent; or 7.5%,
equivalent to 630 mM, in diluent). Animals were anesthesized for exposure for humane reasons,
and to counteract any potential hypoventilation induced by this irritant. Rats were anesthetized
with a cocktail of ketamine (75 mg/kg), xylazine (7.5 mg/kg), and acepromazine (1.5 mg/kg), and
placed in polycarbonate tubes with sealing plungers. Tubes containing animals were mounted in
a nose-only inhalation system (CH Technologies, NJ), and were delivered compressed air with
the aerosolized compound (ethanol or CEES in ethanol) at 12 L/min for 15 minutes (with the
[CEES] delivered for 5% CEES being 13.9mg/L/min/port, and for 7.5% CEES being 20.8
mg/L/min/port). Respiratory rates were monitored for calculation of estimated CEES dose
delivered, with mean respiratory rate of 54 (range 46 – 64). Assuming a physiologic tidal volume
Page 49 of 72
of 2ml per breath (6.7 ml/kg) (1), the mean minute ventilation was calculated to be 108 ml/min. Download full-text
Therefore, the estimated [CEES] delivered during exposure over 15 minutes with 5% CEES was
1.5 mg, and with 7.5% CEES was 2.2 mg. Aerosolization was conducted via a Razel syringe pump
(Razel Scientific, St.Albans, VT) connected to a BioAerosol Nebulizing Generator (BANG; CH
Technologies, NJ), delivering the ethanolic CEES solution or ethanol at a rate of 12.7 ml/hr to the
animals. After 15 minutes of exposure, rats were removed from polycarbonate tubes, and were
observed in their cages until fully recovered from anesthesia.
Rats were terminally anesthesized with pentobarbital (Sleepaway, Fort Dodge Animal Health,
Fort Dodge, IA) at 4, 18 or 72 h post-exposure as per experimental design. Time points
correspond to the first clinical signs of respiratory distress (4 h), the first occurrence of mortality
and peak of biomarkers (18 h), and the first signs of recovery from respiratory distress (72 h). If
rats became moribund, as demonstrated by weight loss >25% body weight, inability to eat or
drink, etc., they were euthanized prior to the planned study termination as per IACUC protocol.
Necropsies were performed on some animals that died prior to planned euthanasia. Rats within
experimental groups were terminally anesthesized with pentobarbital (Sleepaway, Fort Dodge
Animal Health, Fort Dodge, IA), the tracheas were cannulated, and lungs were fixed at 20 cm
H2O with fixative. For confocal analysis, the fluorescent dye used was first instilled as described
below, followed by intratracheal fixation with Karnovsky’s fixative (1.1% glutaraldehyde, 0.9%
paraformaldehyde, cacodylic buffer 0.1M, pH 7.4) at 20 cm H2O pressure for 10 minutes. For
non-confocal studies, the lungs were fixed via tracheal instillation (20 cm H2O pressure) with 4%
Page 50 of 72