1 Central Technologies and 3 Respiratory Diseases Therapeutic Area, Novartis Pharma, CH-4002 Basel, Switzerland; and 2 Novartis Horsham Research Centre, Horsham RH12 5AB, United Kingdom
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Using magnetic resonance imaging (MRI), we detected a signal in the lungs of Brown Norway rats after intratracheal administration of endotoxin [lipopolysaccharide (LPS)]. The signal had two components: one, of diffuse appearance and higher intensity, was particularly prominent up to 48 h after LPS; the second, showing an irregular appearance and weaker intensity, was predominant later. Bronchoalveolar lavage fluid analysis indicated that generalized granulocytic (especially neutrophilic) inflammation was a major contributor to the signal at the early time points, with mucus being a major factor contributing at the later time points. The facts that animals can breathe freely during data acquisition and that neither respiration nor cardiac triggering is applied render this MRI approach attractive for the routine testing of anti-inflammatory drugs. In particular, the prospect of noninvasively detecting a sustained mucus hypersecretory phenotype in the lung brings an important new perspective to models of chronic obstructive pulmonary diseases in animals.
edema; goblet cell; lipopolysaccharide; chronic obstructive pulmonary disease; lung inflammation; magnetic resonance imaging
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
CHRONIC OBSTRUCTIVE PULMONARY DISEASE (COPD) is a complex, multicellular disease in which specific and nonspecific factors result in bronchial obstruction and chronic inflammation (for a recent review see Ref. 29). COPD is a major cause of mortality and a significant drain on health care resources (9, 40). A critical component of the disease is inflammation of the pulmonary mucosa and submucosa, characterized by an infiltration of the airways with neutrophils. The major issues concerning COPD are prevention of the disease, slowing its progression once diagnosis has been established, and prevention, as well as more effective treatment, of exacerbation (41).
Animal models have been established in an attempt to mimic and study
specific aspects of human respiratory disease (30). An
inflammation similar to that observed in COPD patients can be elicited
in animals with the administration of the endotoxin lipopolysaccharide
(LPS), a bacterial macromolecular cell surface antigen. LPS activates
mononuclear phagocytes through a receptor-mediated process, leading to
the release of a number of cytokines, including tumor necrosis
factor- (TNF-
) (42, 43). TNF-
increases the
adherence of neutrophils to endothelial cells, thus facilitating a
massive infiltration of neutrophils into the pulmonary spaces (2).
The majority of animal studies involving endotoxins has been carried out in mice (8, 13, 27, 28), due in part to the ease with which lung injury can be induced by systemic LPS administration in this species, but also because monoclonal antibodies to many mouse cytokines are available. However, the Brown Norway (BN) rat, which is used extensively in the investigation of the pathophysiology of allergic asthma (12, 17, 18), is also a suitable animal in which to study LPS-induced pulmonary injury (22, 24, 35, 39). Exposure of rats to LPS is characterized by infiltration of the alveolar and bronchiolar air spaces by neutrophils (34) and induction of mucous cell metaplasia (19).
Recently, magnetic resonance imaging (MRI) was used to investigate
noninvasively the development of an edematous signal in the lungs of
actively sensitized BN rats after intratracheal allergen challenge
(5). In the present study, a similar approach has been
used to detect and quantify the signal generated in the lungs of BN
rats after intratracheal instillation of LPS. Images were acquired at
regular intervals 16 days postchallenge. MRI results were compared
with the inflammatory status of the lung, represented by the degree of
cell infiltration into the alveolar space, and the mucus concentration,
determined by bronchoalveolar lavage (BAL) fluid analysis.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Animals. Male BN rats (Iffa-Credo, L'Arbresle, France) weighing ~250 g were used. They were housed in a temperature- and humidity-controlled environment and had free access to standard rat chow and tap water. All experiments were carried out according to the Swiss federal regulations for animal protection.
LPS exposure. Animals were anesthetized with 4% isoflurane (Abbott, Cham, Switzerland). LPS from Salmonella typhosa (0.03, 0.3 or 1 mg/kg dissolved in 0.2 ml saline; Sigma, Dorset, UK) or vehicle (0.2 ml of saline) was administered intratracheally, and the animals were allowed to recover.
BAL.
A detailed description of the bronchalveolar lavage (BAL) procedure and
the analysis of the parameters of inflammation is provided in Ref.
5. Briefly, animals challenged with 1 mg/kg of LPS or
saline were killed with an overdose of pentobarbital (250 mg/kg ip)
immediately after an MRI examination. The lungs were lavaged, and the
following parameters were assessed in the BAL fluid: number of
macrophages, eosinophils, and neutrophils; myeloperoxidase (MPO) and
eosinophil peroxidase (EPO) activities, and TNF-, and total protein
and mucus content.
Determination of rat TNF- in BAL fluid.
Fifty microliters of the samples and standards, in duplicate, were
added to the TNF-
antibody-coated wells. Fifty microliters of
biotinylated anti-TNF-
(Biotin Conjugate) solution were pipetted into each well except the chromogen blanks. The plates were incubated for 90 min at room temperature and then washed four times.
Streptavidin-horseradish peroxidase (HRP, 100 µl) was then added to
each well except the chromogen blanks. After incubation for 45 min at room temperature, the wells were washed four times, and 100 µl
of stabilized chromogen were added to each well. The plates were
incubated for 30 min in the dark at room temperature. The absorbance of
each well was read at 450 nm. Results were expressed as picograms per
milliliter, using a standard curve established with standards.
Determination of mucus in BAL fluid using the sandwich
enzyme-linked lectin assay.
Ninety-six-well, high-binding, flat-bottomed microtiter plates (Costar)
were coated with 100 µl of Ulex europaeus agglutinin-1 (UEA-1; Sigma) at 1.25 µg/ml in coating buffer (35 mM sodium
bicarbonate, 15 mM sodium carbonate, pH 9.5). Plates were
covered (with sticky back film) and incubated overnight at 4°C.
Excess lectin was removed by washing three times with 200 µl per well
of wash buffer (10 mM PBS, 0.05% Tween 20, and 0.05% gelatin;
PBS-T-G). Plates were tapped dry before the addition of blocking buffer
(10 mM PBS, 0.1% Tween 20; 150 µl) and incubation for 1 h at
37°C. Plates were washed three times as above with PBS-T-G and either
used immediately or stored at 80°C for up to 6 mo. Purified human mucus standard (100 µl; 24 µg/ml) was serially diluted in PBS over
nine wells in duplicate. Samples that had been stored frozen at
80°C were thawed and added to the plates (100 µl) in duplicate. PBS was substituted for samples in six wells on each plate to serve as
reagent controls. Plates were incubated for 1 h at 37°C, then
washed four times with 200 µl of PBS-T-G. HRP-conjugated UEA-1
(UEA-1/HRP; Sigma or EY Laboratories) was added at 1.25 µg/ml in PBS
(100 µl). Plates were incubated for 1.5 h at 37°C, and then
each plate was washed six times with 200 µl of wash buffer. Substrate
(0.05% orthophenylenediamine dihydrochloride; Sigma) in buffer (0.15 M
citrate phosphate buffer, pH 5.0, with 0.015% hydrogen peroxide added
immediately before use) was prepared in a foil-covered container and
added to the plate (150 µl). The color was allowed to develop in low
light conditions for 5-10 min at room temperature. The optical
density of each well was measured at a wavelength of 492 nm using a
SpectraMax 250 plate reader (Molecular Devices, Surrey, UK). A purified
human mucus sample from an otherwise healthy smoker was used as a
standard to convert optical densities to mucus concentrations. The
gravimetric weight was used to assign the concentration of mucus in the
standard. Approximately 80% of mucus consists of carbohydrate side
chains, of which substantial amounts are
-L-fucose,
which is detected by UEA-1 lectin. Because the epitope is present in
such abundance, it can be used as a generic marker of mucus concentration.
Histology. Challenged rats were killed by an overdose of pentobarbital (250 mg/kg ip). Lungs were perfused in situ via a cannula inserted into the pulmonary artery with 30 ml of modified Krebs solution (composition in mM: 118 NaCl, 4.8 KCl, 1.2 MgSO4, 2.5 CaCl2, 1.2 KH2PO4, 25 NaHCO3, and 11 glucose) and inflated with ~5 ml of 10% phosphate-buffered neutral formalin (BNF), pH 7.0, via the tracheal cannula. After being removed from the thorax, lungs were immersed in BNF for at least 24 h but not longer than 72 h. After 3 days of fixation, the lung tissue was dissected into 5-mm-thick slices and processed into paraffin wax overnight, before being embedded into a wax block. Sections of 4 µm thickness were cut and then stained with hematoxylin and eosin for general morphology or with Alcian blue-periodic acid Schiff for the detection of acid and neutral mucus and identification of goblet cells. For immunostaining, sections were placed in metal staining wax and passed through xylene and industrial methylated spirit. Slides were then treated with hydrogen peroxide in methanol to inhibit endogenous peroxidase and subsequently with 0.1% trypsin in 0.1% calcium chloride. Slides were then washed, and after draining, labeled lectin diluted with tri-buffered saline was applied, and slides were maintained overnight at 4°C. After washing, second-stage antibody (rabbit anti-FITC/HRP; DAKO) diluted in Tris · HCl (pH 7.6) was applied for 30 min. Slices were then again washed, and freshly prepared diaminobenzidine solution was applied for 10 min. After further washing, nuclei were counterstained in Coles hematoxylin. Goblet cells were quantified using a KS400 image analyzer (Imaging Associates, Thame, UK). The program produced a binary image of the microscopic field (from a video camera), detected the more darkly stained goblet cells from the negatively stained background, and created a 20-pixel-deep zone down from the apical surface of the epithelium (enough to include only the surface cells). All other areas of the field of view were ignored. The number and area of goblet cells present within this zone were then calculated.
MRI. Measurements were carried out with a Biospec 47/40 spectrometer (Bruker, Karlsruhe, Germany) operating at 4.7 T. A gradient-echo sequence (14) with repetition time 5.6 ms, echo time 2.7 ms, band width 100 kHz, flip angle of the excitation pulse ~15°, field of view 6 × 6 cm2, matrix size 256 × 128, and slice thickness 1.5 mm was used throughout the study. A single-slice image was obtained by computing the two-dimensional Fourier transformation of the averaged signal from 60 individual image acquisitions and interpolating the data set to 256 × 256 pixels. There was an interval of 530 ms between individual image acquisitions, resulting in a total acquisition time of 75 s for a single slice. The entire lung was covered by 28 consecutive slices. A birdcage resonator of 7 cm in diameter was used for excitation and detection. During MRI measurements, rats were anesthetized with 2% isoflurane in a mixture of O2-N2O (1:2), administered via a face mask, and placed in supine position. The body temperature of the animals was maintained at 37°C by a flow of warm air. Total examination time per animal, including positioning, was ~40 min. The examination protocol for each animal consisted of acquiring a set of baseline images before the LPS challenge. Then, images were acquired at 1, 6, 24, 48, 72, 96, 144, and 192 h after the challenge.
Magnetic resonance image analysis. The volume of signals appearing in the lung after LPS challenge was determined by a semiautomatic segmentation procedure implemented in the IDL (Interactive Data Language Research Systems, Boulder, CO) environment (version 5.1) on an SGI O2 (Silicon Graphics, Mountain View, CA) system. Images were first weakly low-pass filtered with a Gaussian profile filter and then transformed into a set of four gray level classes using adaptive Lloyd-Max histogram quantitation (21). This method avoided operator bias due to arbitrary choice of threshold levels on each image. Signals in response to LPS were represented by the highest gray level class in the transformed images. This class could be extracted interactively by use of a region grower. Because of the unknown extent of the signals detected in the lung, no morphology parameters were incorporated in the region growing process. Instead, a contour serving as a growing border was drawn to control region growing manually. The segmentation parameters were the same for all the analyzed images, chosen to segment regions corresponding to high-intensity signals. Because the signals from edema and vessels were of comparable intensities, the volume corresponding to the vessels was assessed on baseline images and then subtracted from the volumes determined on postchallenge images.
Statistics. Student's t-test with the Bonferroni correction was applied using the saline-treated rats as control group.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
MRI of rat lung after challenge with LPS.
Figure 1A shows representative
transverse sections through the thoracic region of a BN rat before and
at various times after intratracheal exposure to LPS (1 mg/kg). Clear
signal changes were present in the lung within a few hours after
application of the endotoxin. The signals in response to LPS had two
components. One was characterized by a diffuse signal and was
particularly prominent until ~48 h after LPS challenge. A second
component, characterized by an irregular appearance and much weaker
signal intensity, was present in the first hours after LPS challenge but predominated at the later time points. It was not possible to
differentiate the individual components, as they overlapped. Thus
results are presented as total signal volume. No signal changes were
seen in the lung at any time point after saline challenge (data not
shown). For comparison, an axial section through the thorax of an
actively sensitized BN rat, acquired 24 h after intratracheal challenge with ovalbumin (OA, 0.3 mg/kg), is shown in Fig.
1B (left). The intense and diffuse signal
appearing in the lung after OA instillation is related to edema
formation as has been described elsewhere (5). To
illustrate the differences between edema and mucus as detected by MRI,
test tubes containing 10 ml of either mucus (sputum obtained from a
heavy smoker) or water were imaged with the same acquisition parameters
as for the in vivo images (Fig. 1B, right). A
clear difference was evident, which reflected the in vivo findings.
Thus the sputum sample showed an irregular appearance, whereas the
water sample was diffuse and of higher intensity.
|
|
Comparison between signal changes in the lung detected by MRI and
BAL fluid parameters of inflammation after challenge with LPS.
The time course of the response in the airways induced by
intratracheal challenge with LPS (1 mg/kg) detected as an MRI signal in
the lung was compared with the inflammatory status of the lungs defined
by analysis of the BAL fluid (Table 1).
Animals were killed at each time point immediately after the MRI
acquisitions and the BAL fluid was recovered. In confirmation of the
observations summarized in Fig. 2, challenge with LPS led to an
extensive signal in the lung, with a peak of 0.79 ± 0.05 ml at
48 h. The signal declined by ~50% from this peak at 96 h
but was still detectable 16 days after the administration of LPS. BAL
fluid analysis revealed a marked increase in the number of neutrophils
at 24 h after LPS, which declined at later time points but was
still significantly elevated at 384 h. Significant and sustained
increases in macrophages, eosinophil number, and protein concentration
were observed up to 192 h after LPS challenge. MPO and EPO
activities were significantly elevated up to 48 h. TNF-
concentrations were significantly increased 24 h after challenge
but fell below the levels in saline-challenged rats at the later time
points.
|
Comparison between signal changes in the lung detected by MRI and
the mucus concentration in the BAL fluid after LPS-challenge.
The response in the airways induced by LPS challenge detected as an MRI
signal in the lung was compared with the mucus concentration in the BAL
fluid taken from the same animals killed immediately after the MRI
measurement (Fig. 3). Again, the time
course of the signal changes (Fig. 3A) was similar to that
of the earlier studies (Fig. 2 and Table 1). The changes in mucus
content of the BAL fluid followed a remarkably similar time course to
that of the MRI signal (Fig. 3B). Figure 3C shows
the changes in MRI signal plotted against BAL fluid mucus content for
individual animals; the correlation coefficient was 0.762 (P < 0.0001, n = 36).
|
Histology.
Figure 4 displays histological sections
of lungs from animals challenged with 1 mg/kg of LPS. A substantial and
sustained increase in goblet cell number was seen between 48 h and
16 days after challenge. Flocculent mucoid material was consistently
detected in LPS-treated rats, rarely in controls, close to the apical
surface of epithelial cells.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Noninvasive detection and quantification of edema in the rat lung after allergen challenge in sensitized animals have been demonstrated using MRI (5). The method interferes minimally with the well-being of the animals, because neither respiratory nor electrocardiogram triggering is necessary, and the rats respire freely during data collection. The same approach has been used in the present study to detect and quantify the signal in the lung after an intratracheal challenge with LPS. At the echo time used, 2.7 ms, the signal from lung parenchyma was too weak to be detected. To observe a parenchymal signal with a reasonable signal-to-noise ratio at 4.7 T, echo times on the order of 600 µs were required (6). The absence of a lung parenchymal signal in combination with a background free of artifacts provided a high contrast-to-noise ratio for the detection of signals in response to LPS challenge.
The effects of LPS were dose dependent and remarkably long lasting; after 1 mg/kg of LPS, for example, significant signals were detected up to 16 days after challenge. Qualitatively, the signals induced by LPS appeared to have two components: the first, characterized by a diffuse appearance and stronger intensity, was particularly prominent during the first 48 h after LPS challenge; the second component, characterized by an irregular appearance and much weaker signal intensity, was also evident in the first hours after LPS challenge but predominated at later time points. The predominance of the first component in the first 48 h after LPS challenge corresponds to the period during which neutrophil numbers, MPO activity, and protein concentration were markedly increased in the BAL fluid (Table 1). This suggests that edema resulting from generalized granulocytic (especially neutrophilic) inflammation is likely to have been the major contributor to the signal detected by MRI in the lung at the early time points.
The time course of the signal detected by MRI in the lung after LPS and that of mucus determined in the BAL fluid were highly correlated; in each case, a maximum was found 48 h postchallenge, followed by a decline. Histological analysis, on the other hand, revealed a sustained and significant increase in the number of goblet cells, as well as in the epithelial area stained for mucus from 48 h to 16 days after LPS challenge (Fig. 5). In other words, for time points later than 48 h, the volume of stored mucus remained elevated but the amount of mucus secreted decreased. Because the MRI signal elicited by LPS also decreased, the second and long-lasting component of the signal detected in the lung by MRI after LPS instillation is likely to have been due to secreted mucus. Furthermore, the fact that neither the number of goblet cells nor the mucus per unit length epithelium (Fig. 5) increased 24 h after challenge, while at the same time point significant amounts of mucus were secreted (Fig. 3), suggests that at 24 h post-LPS challenge, synthesis of mucus was keeping pace with secretion.
It is of interest to compare the results obtained with LPS with those from an earlier study (5) concerning OA challenge in actively sensitized BN rats. First, the dual aspect of the MRI signal elicited by LPS (Fig. 1A) was qualitatively different from that seen after allergen challenge (Fig. 1B), where strong and uniform diffuse signals were observed in the lungs up to 96 h after intratracheal instillation of OA (5). Ex vivo images of the sputum of a heavy smoker revealed exactly the same features as the signals seen in the lungs of LPS-challenged rats: the MRI signals of mucus were essentially discontinuous and completely different from the strong and uniformly diffuse signals of pure water (Fig. 1B). These data support a major contribution from mucus to the MRI signal in the lung seen after LPS. It bears emphasis that the images of sputum contained high-intensity signals, illustrating the difficulty in differentiating between "pure edema" and "pure mucus" using MRI. The signal detected by MRI in the lungs of OA-challenged rats, on the other hand, is considered to primarily reflect edema, based on the good correlation between the volume of MRI signal and the protein content determined by BAL fluid analysis (5, 36).
The volume of the signal after LPS instillation was also significantly
smaller (25-50%) than that observed after challenge with allergen
(5). The difference could possibly be accounted for by the
relative severity of the tissue eosinophilia. Thus after OA, the number
of eosinophils in the BAL fluid was some three times larger than the
number of eosinophils after LPS challenge (Ref. 5 and
Table 1). Tissue eosinophilia results in the liberation of a multitude
of proinflammatory mediators that cause smooth muscle contraction,
bronchial hyperresponsiveness, vasodilation, and increased vascular
leakage with the production of tissue edema (16, 23).
Administration of LPS, in contrast, results in a predominantly
neutrophilic infiltration orchestrated in large part by the release of
TNF- (3, 33, 37, 38). Macrophages, which are able to
secrete 1,000 times more TNF-
in response to LPS than any other cell
type (7, 26), were significantly elevated throughout the
observation period after LPS (Table 1) and less so after OA
(5). The fact that products of neutrophil (1), eosinophil (25), and macrophage
(32) activation are capable of stimulating an increase in
goblet cell number and/or an increase in mucus secretion would provide
a plausible explanation for the greater mucus contribution to the
signal after LPS than after OA.
In addition to edema and mucus, other mechanisms could potentially contribute to the signal changes described here after LPS instillation. For instance, spin-lattice (T1) and spin-spin (T2) relaxation times may reflect structural changes associated with experimental lung injury (15). Relaxation times have been measured in numerous models of lung injury, including pulmonary edema (caused, for example, by oleic acid and alloxan), bacterial and chemical inflammation, pulmonary hemorrhage, and other types of lung injury by various agents [reviewed by Shioya et al. (31)]. Cutillo et al. (11) showed an increase in T2 (20-100%) in excised, unperfused lungs removed from Sprague-Dawley rats 6 h after treatment with 10 mg/kg LPS ip. T1 was also significantly increased 6 and 9 h after endotoxin, although the changes were small (5-10%). A fivefold increase in T2* has also been observed 24 and 48 h after intratracheal challenge with LPS in regions where signals were present, whereas no significant change in T2* of parenchymal tissue was detected elsewhere in the lung (6). The good correlation between T2* assessments in edematous lung tissue and signal volumes indicates that the observed changes in T2* were primarily due to the increased water content at the sites of edematous lung tissue. A further effect that could potentially influence the signal intensity in the lung is atelectasis, which has been shown to induce changes in T2* (10). Our data do not support this concept since reduction in lung volume, a consequence of atelectasis, was observed in ~30% of saline- or LPS-challenged rats (data not shown), but signals were detected only in the lungs of LPS-treated animals. Finally, vasoreactivity and changes in vessel volume in response to LPS could potentially affect our results. However, the fact that most of the signal in the lung after the challenge appeared in regions where, in the baseline images, no vessels had been detected indicates that changes in vessel volume contributed only marginally to the postchallenge volume of MRI lung signals reported here.
In conclusion, challenge with endotoxin induced the appearance of signals in the lungs of BN rats that differed qualitatively and quantitatively from those seen after allergen challenge in sensitized animals (5). The signals after application of LPS had two components: one of diffuse appearance and a second showing an irregular appearance and weaker intensity. BAL fluid analysis indicated that edema resulting from generalized granulocytic (especially neutrophilic) inflammation could be the major contributor to the signal at the early time points (up to 48 h), with mucus being the major contributing factor at the later time points. The facts that animals can breathe freely during data acquisition and that neither respiration nor cardiac triggering was applied renders this MRI approach attractive for the routine testing of anti-inflammatory drugs (4). In particular, the prospect of noninvasively detecting sustained mucus hypersecretion (20) in the lung brings an important new perspective to models of COPD in animals.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Nigel Sansom for helpful discussions concerning the histology. The technical support of Elisabeth Schaeublin and René Borer is gratefully acknowledged.
![]() |
FOOTNOTES |
---|
Address for reprint requests and other correspondence: N. Beckmann, Novartis Pharma AG, Central Technologies, Analytics & Imaging Sciences Unit, Lichtstr. 29, WSJ-386.2.09, CH-4002 Basel, Switzerland (E-mail: nicolau.beckmann{at}pharma.novartis.com).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published February 1, 2002;10.1152/ajplung.00373.2001
Received 19 September 2001; accepted in final form 22 January 2002.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Agusti, C,
Takeyama K,
Cardell LO,
Ueki I,
Lausier J,
Lou YP,
and
Nadel JA.
Goblet cell degranulation after antigen challenge in sensitized guinea pigs. Role of neutrophils.
Am J Respir Crit Care Med
158:
1253-1258,
1998
2.
Albelda, SM,
Smith CW,
and
Ward PA.
Adhesion molecules and inflammatory injury.
FASEB J
8:
504-512,
1994
3.
Arreto, C-D,
Dumarey C,
Nahori M-A,
and
Vargaftig BB.
The LPS-induced neutrophil recruitment into rat air pouches is mediated by TNF: likely macrophage origin.
Mediator Inflamm
6:
335-343,
1997[ISI].
4.
Beckmann, N,
Mueggler T,
Allegrini PR,
Laurent D,
and
Rudin M.
From anatomy to the target: contributions of magnetic resonance imaging to preclinical pharmaceutical research.
Anat Rec
265:
85-100,
2001[ISI][Medline].
5.
Beckmann, N,
Tigani B,
Ekatodramis D,
Borer R,
Mazzoni L,
and
Fozard JR.
Pulmonary edema induced by allergen challenge in the rat: noninvasive assessment by magnetic resonance imaging.
Magn Reson Med
45:
88-95,
2001[ISI][Medline].
6.
Beckmann, N,
Tigani B,
Mazzoni L,
and
Fozard JR.
MRI of lung parenchyma in rats and mice using a gradient-echo sequence.
NMR Biomed
14:
297-306,
2001[ISI][Medline].
7.
Beutler, BA,
Mulsak IW,
and
Cerami A.
Cachectin/tumor necrosis factor: production, distribution and metabolic fate in vivo.
J Immunol
135:
3972-3977,
1985
8.
Brandolini, L,
Asti C,
Ruggieri V,
Intilangelo A,
Pellegrini L,
Chiusaroli R,
Caselli GF,
and
Bertini R.
Lipopolysaccharide-induced lung injury in mice. II. Evaluation of functional damage in isolated parenchyma strips.
Pulm Pharmacol Ther
13:
71-78,
2000[ISI][Medline].
9.
Calverley, PM.
COPD: early detection and intervention.
Chest
117 Suppl 2:
365S-371S,
2000
10.
Case, TA,
Durney CH,
Ailion DC,
Cutillo AG,
and
Morris AH.
A mathematical model of diamagnetic line broadening in lung tissue and similar heterogeneous systems: calculations and measurements.
J Magn Reson
73:
304-314,
1987[ISI].
11.
Cutillo, AG,
Chan PH,
Ailion DC,
Watanabe S,
Albertine KH,
Durney CH,
Hansen CB,
Laicher G,
Scheel RF,
and
Morris AH.
Effects of endotoxin lung injury on NMR T2 relaxation.
Magn Reson Med
39:
190-197,
1998[ISI][Medline].
12.
Elwood, W,
Lotvall JO,
Barnes PJ,
and
Chung KF.
Characterization of allergen-induced bronchial hyperresponsiveness and airway inflammation in actively sensitized brown-Norway rats.
J Allergy Clin Immunol
88:
951-960,
1991[ISI][Medline].
13.
Faffe, DS,
Seidl VR,
Chagas PS,
Gonçalves de Moraes VL,
Capelozzi VL,
Rocco PR,
and
Zin WA.
Respiratory effects of lipopolysaccharide-induced inflammatory lung injury in mice.
Eur Respir J
15:
85-91,
2000
14.
Frahm, J,
Haase A,
and
Matthaei D.
Rapid NMR imaging of dynamic processes using the FLASH technique.
Magn Reson Med
3:
321-327,
1986[ISI][Medline].
15.
Ganesan, K,
Ailion DC,
Hackmann A,
Laicher G,
Chan P,
and
Cutillo AG.
NMR relaxation and water self-diffusion mechanisms in lung.
In: Application of Magnetic Resonance to the Study of Lung. Armonk, NY: Futura, 1996, p. 115-139.
16.
Giembycz, MA,
and
Lindsay MA.
Pharmacology of the eosinophil.
Pharmacol Rev
51:
213-340,
1999
17.
Haczku, A,
Moqbel R,
Jacobson M,
Kay AB,
Barnes PJ,
and
Chung KF.
T-cells subsets and activation in bronchial mucosa of sensitized Brown-Norway rats after single allergen exposure.
Immunology
85:
591-597,
1995[ISI][Medline].
18.
Hannon, JP,
Tigani B,
Williams I,
Mazzoni L,
and
Fozard JR.
Mechanism of airway hyperresponsiveness to adenoside induced by allergen challenge in actively sensitized Brown Norway rats.
Br J Pharmacol
132:
1509-1523,
2001
19.
Harkema, JR,
and
Hotchkiss JA.
In vivo effects of endotoxin on intraepithelial mucosubstances in rat pulmonary airways. Quantitative histochemistry.
Am J Pathol
141:
307-317,
1992[Abstract].
20.
Jackson, AD.
Airway goblet-cell mucus secretion.
Trends Pharmacol Sci
22:
39-45,
2001[ISI][Medline].
21.
Jain, AK.
Fundamentals of Digital Image Processing. Englewood Cliffs, NJ: Prentice Hall, 1989, chapter 4, pp. 88-102.
22.
Jesch NK, Dorger M, Messmer K, and Krombach F. Formation of nitric
oxide by rat and hamster alveolar macrophages: an interstrain and
interspecies comparison. Toxicol Lett 96-97:
47-51, 1998.
23.
Kroegel, C,
Virchow JC, Jr,
Luttmann W,
Walker C,
and
Warner JA.
Pulmonary immune cells in health and disease: the eosinophil leucocyte (Part I).
Eur Respir J
7:
519-543,
1994
24.
Leal-Berumen, I,
Conlon P,
and
Marshall JS.
IL-6 production by rat peritoneal mast cells is not necessarily preceded by histamine release and can be induced by bacterial lipopolysaccharide.
J Immunol
152:
5468-5476,
1994
25.
Lundgren, JD,
Davey RT, Jr,
Lundgren B,
Mullol J,
Marom Z,
Logun C,
Baraniuk J,
Kaliner MA,
and
Shelhamer JH.
Eosinophil cationic protein stimulates and major basic protein inhibits airway mucus secretion.
J Allergy Clin Immunol
87:
689-698,
1991[ISI][Medline].
26.
Matthews, N.
Tumour-necrosis factor from the rabbit. II. Production by monocytes.
Br J Cancer
38:
310-315,
1978[ISI][Medline].
27.
Nick, JA,
Young SK,
Brown KK,
Avdi NJ,
Arndt PG,
Suratt BT,
Janes MS,
Henson PM,
and
Worthen GS.
Role of p38 mitogen-activated protein kinase in a murine model of pulmonary inflammation.
J Immunol
164:
2151-2159,
2000
28.
O'Malley, J,
Matesic LE,
Zink MC,
Strandberg JD,
Mooney ML,
De Maio A,
and
Reeves RH.
Comparison of acute endotoxin-induced lesions in A/J and C57BL/6J mice.
J Hered
89:
525-530,
1998
29.
Sethi, S.
Bacterial infection and the pathogenesis of COPD.
Chest
117 Suppl 1:
286S-291S,
2000
30.
Shapiro, SD.
Animal models for COPD.
Chest
117, Suppl 1:
223S-227S,
2000
31.
Shioya, S,
Haida M,
and
Fukuzaki M.
NMR study of lung water compartments.
In: Application of Magnetic Resonance to the Study of Lung. Armonk, NY: Futura, 1996, p. 227-286.
32.
Sperber, K,
Gollub E,
Goswami S,
Kalb TH,
Mayer L,
and
Marom Z.
In vivo detection of a novel macrophage-derived protein involved in the regulation of mucus-like glycoconjugate secretion.
Am Rev Respir Dis
146:
1589-1597,
1992[ISI][Medline].
33.
Steinberg, KP,
Milberg JA,
Martin TR,
Maunder RJ,
Cockrill BA,
and
Hudson LD.
Evolution of bronchoalveolar cell populations in the adult respiratory distress syndrome.
Am J Respir Crit Care Med
150:
113-122,
1994[Abstract].
34.
Tesfaigzi, J,
Johnson NF,
and
Lechner JF.
Induction of EGF receptor and erbB-2 during endotoxin-induced alveolar type II cell proliferation in the rat lung.
Int J Exp Pathol
77:
143-154,
1996[ISI][Medline].
35.
Tesfaigzi, Y,
Fischer MJ,
Martin AJ,
and
Seagrave J.
Bcl-2 in LPS- and allergen-induced hyperplastic mucous cells in airway epithelia of Brown Norway rats.
Am J Physiol Lung Cell Mol Physiol
279:
L1210-L1217,
2000
36.
Tigani, B,
Schaeublin E,
Sugar R,
Jackson AD,
Fozard JR,
and
Beckmann N.
Pulmonary inflammation monitored noninvasively by MRI in freely breathing rats.
Biochem Biophys Res Commun
292:
216-221,
2002[ISI][Medline].
37.
Ulich, TR,
Watson LR,
Yin S,
Guo K,
and
del Castillo J.
The intratracheal administration of endotoxin and cytokines. I: characterization of LPS-induced TNF- and IL-1 mRNA expression and the LPS-, TNF-, and IL-1-induced inflammatory infiltrate.
Am J Pathol
138:
1485-1496,
1991[Abstract].
38.
Ulich, TR,
Yin S,
Remick DG,
Russell D,
Eisenberg SP,
and
Kohno T.
Intratracheal administration of endotoxin and cytokines. IV. The soluble tumor necrosis factor receptor type I inhibits acute inflammation.
Am J Pathol
142:
1335-1338,
1993[Abstract].
39.
Van Helden, HP,
Kuijpers WC,
Steenvoorden D,
Go C,
Bruijnzeel PL,
van Eijk M,
and
Haagsman HP.
Intratracheal aerosolization of endotoxin (LPS) in the rat: a comprehensive animal model to study adult (acute) respiratory distress syndrome.
Exp Lung Res
23:
297-316,
1997[ISI][Medline].
40.
Voelkel, NF.
Raising awareness of COPD in primary care.
Chest
117 Suppl 2:
372S-375S,
2000
41.
Voelkel, NF,
and
Tuder R.
COPD: exacerbation.
Chest 117 Suppl
2:
376S-379S,
2000.
42.
Watson, RW,
Redmond HP,
and
Bouchier-Hayes D.
Role of endotoxin in mononuclear phagocyte-mediated inflammatory responses.
J Leukoc Biol
56:
95-103,
1994[Abstract].
43.
Yang, RB,
Mark MR,
Gray A,
Huang H,
Xie MH,
Zhang M,
Goddard A,
Wood WI,
Gurney AL,
and
Godowski PJ.
Toll-like receptor-2 mediates lipopolysaccharide-induced cellular signalling.
Nature
395:
284-288,
1998[ISI][Medline].