Pulmonary lymphatics and edema accumulation after brief lung
injury
Dean E.
Schraufnagel1,
Narasimhan P.
Agaram1,
Aamir
Faruqui1,
Sajal
Jain1,
Leena
Jain1,
Karen M.
Ridge2, and
Jacob Iasha
Sznajder2
1 Departments of Medicine and Pathology, Section of
Respiratory and Critical Care Medicine, University of Illinois at
Chicago, Michael Reese Hospital, and Northwestern University, Chicago
60612; and 2 Pulmonary and Critical Care Medicine,
Northwestern University, Chicago, Illinois 60611
 |
ABSTRACT |
In a past study of hyperoxia-induced
lung injury, the extensive lymphatic filling could have resulted from
lymphatic proliferation or simple lymphatic recruitment. This study
sought to determine whether brief lung injury could produce similar
changes, to show which lymphatic compartments fill with edema, and to
compare their three-dimensional structure. Tracheostomized rats were
ventilated at high tidal volume (12-16 ml) or low tidal volume
(3-5 ml) or allowed to breathe spontaneously for 25 min. Light
microscopy showed more perivascular, interlobular septal, and alveolar
edema in the animals ventilated at high tidal volume (P < 0.0001). Scanning electron microscopy of lymphatic casts showed
extensive filling of the perivascular lymphatics in the group
ventilated at high tidal volume (P < 0.01), but
lymphatic filling was greater in the nonventilated group than in the
group that was ventilated at low tidal volume (P < 0.01). The three-dimensional structures of the cast interlobular and
perivascular lymphatics were similar. There was little filling and no
difference in pleural lymphatic casts among the three groups. More
edema accumulated in the surrounding lymphatics of larger blood vessels
than smaller blood vessels. Brief high-tidal-volume lung injury caused
pulmonary edema similar to that caused by chronic hyperoxic lung
injury, except it was largely restricted to perivascular and septal
lymphatics and prelymphatic spaces.
pulmonary edema; barotrauma; ventilator; corrosion casting; microscopy; scanning electron
 |
INTRODUCTION |
KEEPING THE
ALVEOLAR SPACE free of excess water is vital to normal pulmonary
gas exchange. The relation between lung edema and ventilation is
complex and depends on factors such as intravascular volume,
permeability of the microvascular endothelium, integrity of the
alveolar epithelium, cardiac status, and oncotic and osmotic forces
(5, 24, 25). The ability to remove fluids from the lung
also involves active sodium transport across the alveolar epithelium
via the interstitium and initial lymphatics (9, 11, 17,
28).
In previous studies of hyperoxic lung injury, an increased amount of
initial, saccular, and conduit lymphatics was observed after 7 days of
hyperoxia (8, 22). In those studies, the increased
lymphatic casting could have resulted from filling existing lymphatics
and potential tissue spaces or from lymphatic proliferation. This study
was undertaken to determine whether similar lymphatic filling could
take place in a model of injury over a brief period. Mechanical
ventilation with overinflation was chosen as a model to cause lung
damage and edema (6, 26). In addition, we sought to
determine the sites of this early edema formation and the structure of
these compartments.
With light and scanning electron microscopy of lymphatic casts, we
studied where fluid accumulates and the relations of the lymphatics to
the blood vessels and airways. The casting resin that is injected into
the pulmonary vasculature is viscous and hardens in ~1 min to show
where fluid goes transvascularly within that time. Vascular casting is
a sensitive method of detecting small amounts of lymphatic filling
(1, 8, 19, 22). Spontaneously breathing animals with
negative intrathoracic pressures were used in addition to animals
ventilated at low tidal volume and with positive pressure, because we
believed that the positive pressure may decrease focal fluid
collections that we could detect with this casting technique.
 |
METHODS |
Animal preparation.
The experiment was approved by the Animal Use and Care Committee at
Michael Reese Hospital. The animals were handled according to National
Research Council guidelines (14). Male pathogen-free Sprague-Dawley rats, weighing 289-304 g, were anesthetized
intraperitoneally with pentobarbital sodium (Nembutal, Abbott
Laboratories, North Chicago, IL), with an initial dose of 50 mg/kg and
termination when the animal did not respond to a tail pinch. After
anesthesia, a tracheostomy was performed with a 14-gauge plastic tube
placed and secured ~5 mm into the trachea. The tubing was attached to a small animal ventilator (model 683, Harvard Apparatus, South Natick,
MA). The rat was ventilated at a frequency of 70 breaths/min. The
low-tidal-volume animals were ventilated at 3-5 ml and peak airway
pressure of 8-10 cmH2O. The high-tidal-volume group
was ventilated at 12-16 ml and peak airway pressure of 35 cmH2O. Inspiratory pressure was monitored by a Honeywell
transducer attached to a strip chart recorder (model 2400, Gould,
Cleveland, OH). A third group of rats were anesthetized and
tracheostomized and allowed to breathe spontaneously for 25 min. All
animals breathed only room air.
Thirty-five animals (divided into three groups) were used for
measurement of wet-to-dry lung weight ratio. After ventilation, the
rat's right upper lobe was tied off, removed, and placed in a
preweighed microcentrifuge tube. The centrifuge tube was then placed in
a lyophilizer and dried overnight. The tube was reweighed, and the tube
weight was subtracted from the final weight. The tube was then returned
to the vacuum for several more hours. The tube was again removed and
reweighed. This process was repeated until a constant weight, defined
as <0.001-g difference between the two measurements, was obtained.
Light microscopy.
Distribution of edema was assessed by light microscopy in 15 animals
(five in each group). After ventilation, or the 25 min without
ventilation, the lungs were removed en bloc and fixed. The trachea was
filled with a 10% formalin solution at 25 cmH2O pressure.
The lungs remained inflated with formalin for
24 h and then processed
for routine light microscopy with dehydration and paraffin embedding.
One block was made for each animal and positioned to give a coronal
section through the apex of the lung. The light-microscopic slides were
stained with hematoxylin and eosin and by the elastin-van Gieson method.
To measure the diameters of noncapillary vessels, the elastin-van
Gieson-stained slides were moved to select fields at random. Vessels
were identified as arteries or veins on the basis of their elastic
laminae and, to a lesser extent, their position near or distal to an
airway (10). The vessel diameter was the distance between
the outer boundaries of the adventitia. The cuffed diameter was the
outer border of the edematous areas that was distal to the outer
elastic lamina. Tangential cuts were avoided by taking the smallest
cylindrical diameter. The diameters were measured with a 10-mm reticule
with 0.1-mm markings with a ×10 magnification objective. The
percentage of cuffing was calculated as follows: 1
vessel
diameter
cuffed diameter. For each animal, measurements for 10 arteries and 10 veins were tabulated.
To assess damage to bronchial epithelia, we studied epithelial cell
sloughing severity by viewing the elastin-van Gieson-stained slides
with the ×10 magnification objective. No detectable epithelial abnormality was scored 0. If the epithelial uplifting from the basement
membrane was <100 µm long and no sloughed cells were found, the
score was 1. Epithelial uplifting >100 µm or >20 clumps of
epithelial cells was scored 2. More extensive damage of the epithelium,
with areas of denudation or sheets of bronchial cells in the airway or
alveoli, was scored 3.
Alveolar edema was graded by selecting random alveolar fields using the
×40 magnification objective to view the hematoxylin-and-eosin-stained slides. No evidence of edema was scored 0; a few strands of fibrin or
debris could be present in the whole field, and still the edema would
be scored 0. If fibrin strands or clumps were found in several, but
less than half, of the alveoli and at least one erythrocyte was present
per alveolus, then the score was 1. If pink, flocculent material or
clumps were present in more than half of the alveoli and erythrocytes
were present in most alveoli, the score was 2. If pink material or
blood was found in almost all (>80%) alveoli, the score was 3. If
dense proteinaceous material and erythrocytes flooded all alveoli, the
score was 4.
Interlobular septal widening from edema was identified by scanning the
entire slide using a ×4 magnification objective. The lengths of all
widened septa were measured with the reticule and the ×10
magnification objective and recorded as a total length in millimeters
of widened interlobular septa per slide per animal. Because no animal
had more than five widened septa, five entries were recorded for each
animal, with up to five zeros being assigned to those animals with no
widened interlobular septa.
Lymphatic casting and scanning electron microscopy.
For casting and scanning electron microscopy, we used 15 additional
animals. After ventilation or tracheostomy-nonventilation, the abdomen
was opened and the caudal vena cava was cannulated. The vasculature was
rinsed with 50 ml of heparinized (5,000 U/l) normal saline warmed to
40°C. Fifteen milliliters of partially polymerized methyl
methacrylate (Mercox, Ladd Research Industries, Burlington, VT) mixed
with ~2 g of accelerator (50% benzoyl peroxide) were injected at a
constant flow rate into the caudal vena cava over 1 min with an
infusion pump (model 351, Sage Instruments, Cambridge, MA). The
methacrylate was allowed to harden for 1 h, and the lungs were
separated and placed in a sodium hydroxide solution until the tissue
was digested.
The casts were rinsed in detergent, water, and alcohol and cut into
~1 × 8 × 8-mm pieces. The specimens were fastened to
aluminum studs using double-sided tape. They were sputter coated with
palladium-gold and viewed with a scanning electron microscope (model
JSM-35C, JEOL). The accelerating voltage was 10 kV, and the working
distance was 15 mm (18, 18a).
From each animal, four studs of the cut surface and four studs of the
pleural surface were made. On the pleural surface, we recorded the
presence and type of lymphatics found on 10 randomly selected fields at
×200 magnification. Random field selections were carried out by
placing a numbered grid over the microscope's cathode ray tube with
the specimen image at the lowest magnification that allowed it to fill
the screen. A number was selected from a random-number table, and the
area under the number was moved to the center of the screen. We then
increased the magnification. On the cut surface, 10 arteries and 10 veins per stud were found at ×200 magnification. We recorded their
diameters and the presence of adjacent lymphatics. If we could not
determine whether a vessel was an artery or a vein, it was classified
as indeterminate and the same measurements were carried out (up to 10 indeterminate vessels per stud or until we had 10 arteries and 10 veins). If no lymphatic was present around a vessel, a score of 0 was
assigned. If the vessel was partially encircled by lymphatics, a score
of 1 was assigned, and if the perivascular lymphatics completely surrounded the vessel, a score of 2 was assigned.
On the basis of morphology, different types of lymphatics and
prelymphatic interstitial spaces can be cast by injecting a resin
through permeable pulmonary vasculature (19, 22) and air
space (8). The lymphatic casts around the large blood
vessels have been termed saccular and conduit lymphatics, and those on the pleural surface have been termed initial and conduit lymphatics (19).
Analysis.
The specimens were coded and examined in a blinded fashion. We used
analysis of variance to compare the three groups of animals for
continuous data. We used Scheffé's procedure (P < 0.05) to locate differences detected by the analysis of variance.
For discontinuous or rank data, we performed the nonparametric
Kruskal-Wallis test. For proportional data, such as the presence or
absence of lymphatics around vessels in the different tidal volume
groups, we used contingency table analysis to generate a
2 statistic. In addition to the rank order
(0-2) cast lymphatic data, we used the presence or
absence (0-1) of lymphatics. Values are means ± SE. Statistical analysis was performed with SPSS statistical software
(version 7.5, SPSS, Chicago, IL).
 |
RESULTS |
There were no external differences in the animals, except frothy
tracheal discharge during ventilation in two animals ventilated at high
tidal volume. The lungs of the group ventilated at high tidal volume
appeared more erythematous and edematous. The wet-to-dry lung weight
ratio was 5.97 ± 0.27 for animals ventilated at high tidal
volume, 4.98 ± 0.06 for animals ventilated at low tidal volume,
and 4.95 ± 0.04 for nonventilated animals. The difference between
the values of the animals ventilated at high tidal volume and the
others was significant (P < 0.0001; Fig.
1).

View larger version (7K):
[in this window]
[in a new window]
|
Fig. 1.
Wet-to-dry lung weight ratio. Ratio was greater in
animals ventilated at high tidal volume (VT) than in the
other 2 groups (*P < 0.01). There was no significant
difference between animals ventilated at low tidal volume and
spontaneously breathing (Spont) animals.
|
|
Light microscopy.
Light-microscopic viewing of the slides as unknowns did not distinguish
the groups, but the measurements showed that the arterial cuffing
(P < 0.0001; Figs. 2 and
3), venous cuffing (P < 0.01; Figs. 4 and
5), alveolar edema (P < 0.01; Fig. 6), and septal thickening
(P < 0.001; Figs. 7 and
8) were greater in the animals ventilated at high tidal volume than in the other groups. Septal widening was not observed in the group ventilated at low tidal volume,
but this was not different from the nonventilated group, in which
average septal widening was only 150 µm per animal. The epithelial
sloughing and damage were minimal in all animals and not different
among the groups. The scores were <0.5 in each group. The different
measures of edema were usually found together in most animals.

View larger version (11K):
[in this window]
[in a new window]
|
Fig. 2.
Periarterial edema shown as percentage of entire diameter
taken up by edematous cuff. Periarterial edema was greater in animals
ventilated at high tidal volume than in the other 2 groups
(*P < 0.01).
|
|

View larger version (117K):
[in this window]
[in a new window]
|
Fig. 3.
Light-microscopic view of a bronchovascular bundle from
an animal ventilated at high tidal volume. Artery is surrounded by a
fluid cuff, the boundaries of which are marked by arrows. Note scant
edema in nearby larger bronchiole (B).
|
|

View larger version (11K):
[in this window]
[in a new window]
|
Fig. 4.
Perivenous edema shown as percentage of entire diameter
taken up by edematous cuff. Perivenous edema was greater in animals
ventilated at high tidal volume than in the other 2 groups
(*P < 0.01).
|
|

View larger version (98K):
[in this window]
[in a new window]
|
Fig. 5.
Light-microscopic view of edema around a pulmonary vein
in an animal ventilated at high tidal volume.
|
|

View larger version (11K):
[in this window]
[in a new window]
|
Fig. 6.
Alveolar edema scores. Alveolar edema was greatest in
animals ventilated at high tidal volume (P < 0.01).
|
|

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 7.
Interlobular septa widened in animals ventilated at high
tidal volume. This index was a summation of the length of all widened
interlobular septa in the lung slice. Animals ventilated at low tidal
volume had no widened interlobular septa, but this was not different
from the nonventilated group in which total length of septa was only
0.15 mm.
|
|

View larger version (134K):
[in this window]
[in a new window]
|
Fig. 8.
Low-power light-microscopic view of a widened
interlobular septum in an animal ventilated at high tidal volume.
|
|
Scanning microscopy of casts.
The group ventilated at high tidal volume had more arterial (Figs.
9 and
10) and venous cuffing (Figs.
11 and
12) than the others. The cuffing
involved the prelymphatic space and saccular and conduit lymphatics
(19). Lymphatics in the interlobular septa were cast in
the group ventilated at high tidal volume. Lymphatics of the pleural
surface were scant in all animals.

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 9.
Cast lymphatics around arteries. Lymphatics were
increased most in animals ventilated at high tidal volume
(*P < 0.01).
|
|

View larger version (138K):
[in this window]
[in a new window]
|
Fig. 10.
Scanning electron micrograph of cast lymphatics (arrows)
in space around a pulmonary artery (PA). Alveolar blood capillaries
(arrowheads) are outside bronchovascular bundle.
|
|

View larger version (9K):
[in this window]
[in a new window]
|
Fig. 11.
Cast lymphatics around veins. Number of cast lymphatics
was greatest in animals ventilated at high tidal volume
(*P < 0.01).
|
|

View larger version (157K):
[in this window]
[in a new window]
|
Fig. 12.
Scanning electron micrograph of cast lymphatics (arrows)
in space around a pulmonary vein (V).
|
|
There were lymphatic casts around ~18% of the arteries of the
unventilated animals, 5% of the arteries of the animals ventilated at
low tidal volume, and 29% of the arteries of the animals ventilated at
high tidal volume (P < 0.01; Fig. 9). For this
parameter, the groups differed from each other (P < 0.05; Table 1). The animals ventilated at
high tidal volume and the unventilated animals had more saccular and
conduit lymphatics than the animals ventilated at low tidal volume
(P < 0.01) but were not different from each other. In
the group ventilated at high tidal volume, more lymphatics completely
surrounded the arteries. A similar pattern of lymphatics was observed
around veins (Fig. 11; P < 0.01) and indeterminate vessels (P < 0.001) in all the groups.
The larger vessels generally had more cast lymphatics than smaller
vessels. Cast lymphatics around arteries were not different from those
around veins when all groups were taken together. Nonventilated animals
had more periarterial than perivenous casts (18 vs. 8%, P < 0.01).
There was no difference in cast lymphatics on the pleural surface
between the groups. Of the 10 fields selected randomly and viewed at
×200 magnification, <10% had cast lymphatics in each of the groups
(9, 8, and 6% in the nonventilated, low-tidal-volume, and
high-tidal-volume groups, respectively). Cast conduit lymphatics were
present on the pleural surface in
3% of the ×200 magnification fields (2, 3, and 1% in the nonventilated, low-tidal-volume, and high-tidal-volume groups, respectively).
 |
DISCUSSION |
Mechanical ventilation with high tidal volumes, even for a short
period of time, causes increased edema formation (6, 27) and impairs clearance of fluid from alveoli (25). Using
this ventilated rodent model, we studied the structure and filling of
different lymphatic forms in rat lungs as they underwent acute changes
(25 min) caused by high tidal ventilation. The short duration of the
ventilation was chosen to show the changes in early edema formation,
which contrasts with our previous studies of hyperoxic lung injury,
where an increase in initial, saccular, and conduit lymphatics was
observed after 7 days of hyperoxia (8, 22). Undiluted
Mercox is a viscous material with a viscosity of ~27 cP before
addition of the benzoyl peroxide accelerator (2). After
the accelerator is added, the viscosity increases logarithmically with
time. The increased viscosity with our method makes extrusion of the
resin through the 20-gauge needle difficult after a little more than 1 min. In <5 min the resin hardens to become rigid (20). This method, therefore, identifies lymphatics that fill within this
short time interval. The ventilation was stopped just before the lungs
were cast, so that mechanical factors from ventilation could not affect
intralymphatic and interstitial pressure, which in turn could affect
the filling.
The structure of these lymphatic casts as viewed by electron microscopy
was identical to that in models of chronic hyperoxic lung injury
(22) and other models of acute pulmonary edema
(19). The edema fluid moves from capillaries into the
interstitium and from there into characteristic tissue spaces
(prelymphatics) that are intimately connected to the initial lymphatics
and conduit lymphatics.
With ventilator-induced lung injury, perivascular lymphatics become
more engorged than other types of lymphatics (Figs. 2, 4, 5, and
10-12). The perivascular lymphatic filling is not associated with
significant pleural lymphatic filling, in contradistinction to our
chronic hyperoxic model (8, 22). In the group ventilated at high tidal volume, edema formation was most apparent in the periarterial, perivenous, and interlobular septa and alveoli. There was
more lymphatic filling in the larger vessels than in the smaller
vessels, which could mean that the stress and edema-forming forces were
greater in the proximal vessels or that the large vessels had more
lymphatic capacity and greater ability to be recruited. It could also
be that smaller lymphatics feed into the larger lymphatics and the
brief time was sufficient for them to fill more completely.
In a model of chronic lung injury caused by exposure of rats to 85%
oxygen for 7 days, all lymphatic beds were greatly increased (8,
22). It is well known that this hyperoxic stimulus causes significant proliferation of alveolar epithelial cells and lung fibroblasts (3). These studies raised the question whether the increase in lymphatic capacity was the result of the formation of
new lymphatics caused by cell proliferation in response to chronic
hyperoxia or the recruitment of preexisting lymphatics. This short-term
study provides strong evidence to support the notion that prelymphatics
exist, ready to be rapidly recruited in situations where edema develops.
The ready casting of lymphatics by injection of a viscous, partially
polymerized methacrylate into the pulmonary vasculature shows that the
model of ventilator-induced lung injury produces a permeability edema
for this high-molecular-weight resin. Webb and Tierney
(29) observed alveolar and perivascular edema when they
ventilated rats for a short duration with high peak inspiratory pressure. The eosinophilic character of the edema suggested that it was
protein rich. This was confirmed by several investigators (6, 7,
12, 16). However, the mean light-microscopic alveolar edema
score for our animals was only 0.24, and no slide had a score of 3 or
4. A score of 3 or 4 might be expected with severe proteinaceous and
hemorrhagic pulmonary edema, which we have seen with neurogenic
pulmonary edema (21, 23) and prolonged (
60-min)
mechanical ventilation (12). Light microscopy was unable
to detect major airway damage in this model.
There were more perivascular cast lymphatics in the spontaneously
breathing, anesthetized tracheostomized rats than in rats ventilated at
low tidal volume. This may have resulted from the positive-pressure
ventilation, which is known to shift intrapulmonary fluid
(15). It is also possible that breathing efforts of the unventilated animals and the greater negative intrathoracic pressure could have increased vascular transudation. Negative-pressure pulmonary
edema has been reported in patients with upper airway obstruction
(4), but lymphatic filling was only mildly increased in
our spontaneously breathing animals, as detected by scanning electron
microscopy of the casts. This method may be more sensitive than the
wet-to-dry weight ratio of the lung. Although the nonventilated animals
were anesthetized, breathing movements were observed as the resin was injected.
The study could not distinguish a significant difference in lymphatic
compartments around the arteries and veins. Heavy resin accumulation
around these blood vessels may make it difficult or impossible to
distinguish the blood vessel type, which is dependent on the
characteristic surface features seen under the electron microscope
(13). The presence of indeterminate vessels may have masked an arteriovenous difference. When the vessel wall is visible, small veins can be distinguished from small arteries much better by
electron microscopy of vascular casts than by light microscopy (18).
This study provides evidence that brief ventilation with high tidal
volumes produces edema and that the perivascular and septal lymphatics
are readily available to accept the increased amount of fluid. The
structure of the lymphatics in brief lung injury is identical to that
in chronic edema. Edema can be scored by light-microscopic measurements
and visualized and scored by casting the lymphatics through the
vascular space with an appropriate resin. The casting technique
provides a "snapshot" of the lymphatic filling that occurs within
~60 s.
 |
ACKNOWLEDGEMENTS |
This work was supported by National Heart, Lung, and Blood
Institute Grant HL-48129 (J. I. Sznajder), a National Institutes of Health National Research Service Award (K. M. Ridge), and the James E. Liston ARDS Research Fund (D. E. Schraufnagel).
 |
FOOTNOTES |
Address for reprint requests and other correspondence:
D. E. Schraufnagel, Dept. of Medicine M/C 719, University of
Illinois at Chicago, 840 S. Wood St., Chicago, IL 60612-7323 (E-mail:
schrauf{at}uic.edu).
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 January 24, 2003;10.1152/ajplung.00333.2002
Received 3 October 2002; accepted in final form 26 November 2002.
 |
REFERENCES |
1.
Aharinejad, S,
Böck P,
Firbas W,
and
Schraufnagel DE.
Pulmonary lymphatics and their spatial relationship to venous sphincters.
Anat Rec
242:
531-544,
1995[ISI][Medline].
2.
Aharinejad, S,
and
Lametschwandtner A.
Microvascular Corrosion Casting in Scanning Electron Microscopy. Techniques and Applications. New York: Springer, 1992.
3.
Crapo, J,
Barry BD,
Foscue HA,
and
Shelburne J.
Structural and biochemical changes occurring during exposure to lethal and adaptive doses of oxygen.
Am Rev Respir Dis
122:
123-143,
1980[ISI][Medline].
4.
Deepika, K,
Kenaan CA,
Barrocas AM,
Fonseca JJ,
and
Bikazi GB.
Negative-pressure pulmonary edema after acute upper airway obstruction.
J Clin Anesth
9:
403-408,
1997[ISI][Medline].
5.
Dematte, JE,
and
Sznajder JI.
Mechanisms of pulmonary edema clearance: from basic research to clinical implication.
Intensive Care Med
26:
477-480,
2000[ISI][Medline].
6.
Dreyfuss, D,
and
Saumon G.
State of the art: ventilator-induced lung injury. Lessons learned from experimental studies.
Am J Respir Crit Care Med
157:
294-323,
1998[ISI][Medline].
7.
Dreyfuss, D,
Soler P,
and
Saumon G.
Spontaneous resolution of pulmonary edema caused by short periods of cyclic overinflation.
J Appl Physiol
72:
2081-2089,
1992[Abstract/Free Full Text].
8.
Hainis, K,
Sznajder JI,
and
Schraufnagel DE.
Lung lymphatics cast from the air space.
Am J Physiol Lung Cell Mol Physiol
267:
L199-L205,
1994[Abstract/Free Full Text].
9.
Havill, AM,
and
Gee MH.
Role of interstitium in clearance of alveolar fluid in normal and injured lungs.
J Appl Physiol
57:
1-6,
1984[Abstract/Free Full Text].
10.
Kay, JM.
Blood vessels of the lung.
In: Comparative Biology of the Normal Lung, edited by Parent RA.. Boca Raton, FL: CRC, 1991, p. 163-172.
11.
Lauweryns, JM,
and
Baert JH.
State of the art: alveolar clearance and the role of the pulmonary lymphatics.
Am Rev Respir Dis
115:
625-683,
1977[ISI][Medline].
12.
Lecuona, E,
Saldias F,
Comellas A,
Ridge K,
Guerrero C,
and
Sznajder JI.
Ventilator-associated lung injury decreases lung ability to clear edema in rats.
Am J Respir Crit Care Med
159:
603-609,
1999[Abstract/Free Full Text].
13.
Miodonski, A,
Hodde KC,
and
Bakker C.
Rasterelektronenmikroskopie. Morphologische Unterschiede zwischen Arterien und Venen.
Beitr Elektronenmikroskop Direktabb Oberfl
9:
435-442,
1976.
14.
National Research Council.
Guide for the Care and Use of Laboratory Animals. Washington, DC: National Academy Press, 1996.
15.
Pare, PD,
Warriner B,
Baile EM,
and
Hogg JC.
Redistribution of pulmonary extravascular water with positive end-expiratory pressure in canine pulmonary edema.
Am Rev Respir Dis
127:
590-593,
1983[ISI][Medline].
16.
Parker, JC,
Townsley MI,
Rippe B,
Taylor AE,
and
Thigpen J.
Increased microvascular permeability in dog lungs due to high peak airway pressures.
J Appl Physiol
57:
1809-1816,
1984[Abstract/Free Full Text].
17.
Saldias, FJ,
Lecuona E,
Comellas AP,
Ridge KM,
Rutschman DH,
and
Sznajder JI.
-Adrenergic stimulation restores rat lung ability to clear edema in ventilator-associated lung injury.
Am J Respir Crit Care Med
162:
282-287,
2000[Abstract/Free Full Text].
18.
Schraufnagel, DE.
Microvascular casting of the lung: a state-of-the-art review.
Scanning Microsc
1:
1733-1747,
1987[ISI][Medline].
18a.
Schraufnagel, DE.
Ranking corrosion efficiency: a Latin square study on rat lung microvascular corrosion casts.
Scanning Microsc
3:
299-304,
1989[ISI][Medline].
19.
Schraufnagel, DE.
Forms of lung lymphatics: a scanning electron microscopic study of casts.
Anat Rec
233:
547-554,
1992[ISI][Medline].
20.
Schraufnagel, DE,
and
Ganesan DP.
Tracers in vascular casting resins enhance backscattering brightness.
Scanning
24:
121-126,
2002[ISI][Medline].
21.
Schraufnagel, DE,
Kurtulus M,
and
Patel TH.
Effect of age on the contraction of pulmonary venous sphincters in rats.
Am J Respir Crit Care Med
149:
227-231,
1994[Abstract].
22.
Schraufnagel, DE,
Llopart Basterra L,
Hainis K,
and
Sznajder JI.
Lung lymphatics increase after hyperoxic lung injury: an ultrastructural study of casts.
Am J Pathol
144:
1393-1402,
1994[Abstract].
23.
Schraufnagel, DE,
and
Patel KR.
Sphincters in pulmonary veins: an anatomic study in rats.
Am Rev Respir Dis
141:
721-726,
1990[ISI][Medline].
24.
Staub, NC.
Alveolar flooding and clearance.
Am Rev Respir Dis
127:
544-551,
1983.
25.
Sznajder, JI.
Strategies to increase alveolar epithelial fluid removal in the injured lung.
Am J Respir Crit Care Med
160:
1441-1442,
1999[Free Full Text].
26.
Sznajder, JI,
Ridge KM,
Saumon G,
and
Dreyfuss D.
Lung injury induced by mechanical ventilation.
In: Pulmonary Edema, edited by Mathay R.. New York: Dekker, 1998, p. 413-430.
27.
Trembley, L,
Valenza F,
Ribeiro SP,
Li J,
and
Slutsky AS.
Injurious ventilatory strategies increase cytokines and c-fos mRNA expression in an isolated rat lung model.
J Clin Invest
99:
944-952,
1995.
28.
Ware, LB,
and
Matthay MA.
Alveolar fluid clearance is impaired in the majority of patients with acute lung injury and the acute respiratory distress syndrome.
Am J Respir Crit Care Med
163:
1376-1383,
2001[Abstract/Free Full Text].
29.
Webb, HH,
and
Tierney DF.
Experimental pulmonary edema due to intermittent positive-pressure ventilation with high inflation pressures. Protection by positive end-expiratory pressure.
Am Rev Respir Dis
110:
556-565,
1974[ISI][Medline].
Am J Physiol Lung Cell Mol Physiol 284(5):L891-L897
1040-0605/03 $5.00
Copyright © 2003 the American Physiological Society