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Department of Internal Medicine, Justus-Liebig University, 35385 Giessen; and Department of Anesthesia and Intensive Care Medicine, University Clinic Rudolf Virchow, 13353 Berlin, Germany
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
ACKNOWLEDGEMENTS
FOOTNOTES
REFERENCES
ABSTRACT 
Steudel, Wolfgang, Hans-Joachim Krämer, Daniela
Degner, Simone Rosseau, Hartwig Schütte, Dieter Walmrath, and
Werner Seeger. Endotoxin priming of thromboxane-related
vasoconstrictor responses in perfused rabbit lungs. J. Appl. Physiol. 83(1): 18-24, 1997.
In prior
studies of perfused lungs, endotoxin priming markedly enhanced
thromboxane (Tx) generation and Tx-mediated vasoconstriction in
response to secondarily applied bacterial exotoxins. The
present study addressed this aspect in more detail by employing
precursor and intermediates of prostanoid synthesis and performing
functional testing of vasoreactivity and measurement of product
formation. Rabbit lungs were buffer perfused in the
absence or presence of 10 ng/ml endotoxin. Repetitive intravascular
bolus applications of free arachidonic acid provoked constant pulmonary
arterial pressor responses and constant release reactions of
TxA2 and prostaglandin (PG)
I2 in nonprimed lungs. Within
60-90 min of endotoxin recirculation, which provoked progressive
liberation of tumor necrosis factor-
but did not effect any
hemodynamic changes by itself, both pressor responses and prostanoid
release markedly increased, and both events were fully blocked by
cyclooxygenase (Cyclo) inhibition with acetylsalicylic acid
(ASA). The unstable intermediate
PGG2 provoked moderate pressor
responses, again enhanced by preceding endotoxin priming and fully
suppressed by ASA. Vasoconstriction also occurred in response to the
direct Cyclo product PGH2, again amplified after endotoxin pretreatment, together with markedly enhanced
liberation of TxA2 and
PGI2. In the presence
of ASA, the priming-related increase in pressor responses and the
prostanoid formation were blocked, but baseline vasoconstrictor
responses corresponding to those in nonprimed lungs were maintained.
Pressor responses to the stable Tx analog U-46619 were not
significantly increased by endotoxin pretreatment, but some generation
of TxA2 and
PGI2 was also noted under these
conditions. We conclude that endotoxin priming exerts profound effects
on the lung vascular prostanoid metabolism, increasing the readiness to
react with Tx-mediated vasoconstrictor responses to various stimuli,
suggesting that enhanced Cyclo activity is an important underlying
event.
cyclooxygenase; endoperoxides; pulmonary arterial pressure; tumor
necrosis factor- INTRODUCTION SEPSIS AND SEPSIS-RELATED organ injury, including the
acute respiratory distress syndrome of the adult (ARDS), is the leading cause of mortality in critically ill patients (12). In
animal models, the clinical features of ARDS may be reproduced by
endotoxin, lipopolysaccharides (LPS) released from destroyed cell walls
of gram-negative bacteria (3, 8). The microcirculatory
disturbances with subsequent cellular injury and loss of organ function
induced by LPS have been related to the activation of inflammatory
cells and the induction of inflammatory mediator generation, including, in particular, cytokines such as tumor necrosis factor- The present study examined this hypothesis in greater detail by
employing various intermediates of prostanoid synthesis and performing
functional testing of vasoreactivity and measurement of product
formation. In particular, the vasculature of LPS-primed and nonprimed
perfused lungs were challenged with the precursor fatty acid
arachidonic acid (AA), the unstable endoperoxide intermediate prostaglandin (PG) G2, the
unstable cyclooxygenase (Cyclo) product PGH2, and the stable Tx analog
U-46619 (see Fig. 1). The results of these
studies suggest an important role for enhanced Cyclo activity in the LPS-priming phenomenon of the lung vasculature.
MATERIALS AND METHODS 
(TNF-). Low doses of LPS have been shown to prime
inflammatory cells in vitro and enhance responsiveness to a second
inflammatory stimulus such as lipid and peptide chemoattractants (1, 4,
7, 15, 24). Such priming phenomena have also been demonstrated in
intact lungs perfused with blood-free medium: after a preceding period
of intravascular LPS admixture, which did itself not affect pulmonary
hemodynamics or induce thromboxane (Tx) release, enhanced TxA2 generation and Tx-related
vasoconstriction were provoked by platelet-activating factor (18) and
the bacterial exotoxins Staphylococcus
aureus -toxin (26) and Escherichia
coli hemolysin (25, 27). The enhanced Tx-related
vasoconstrictor response in LPS-primed lungs was associated with severe
ventilation-perfusion mismatching (27). In the study performed with
E. coli hemolysin as a secondary
stimulus, the time (threshold ~60-90 min) and dose dependency
(threshold ~0.1-1 ng/ml LPS) of the LPS priming effect were
consistent with the hypothesis that de novo synthesis of enzymes
involved in Tx synthesis and/or Tx efficacy might represent the
event underlying the priming phenomenon.
Fig. 1.
Cyclooxygenase pathway of arachidonic acid (AA) and possible influence
of endotoxin and tumor necrosis factor- (TNF-). Endotoxin [lipopolysaccharide (LPS)] may activate cyclooxgenase and
facilitate secondary AA release.
PGG2,
PGH2, and
PGI2, prostaglandin
G2, H2, and
I2, respectively;
6-keto-PGF1,
6-ketoprostaglandin F1;
TxA2 and
TxB2, thromboxane
A2 and
B2, respectively; U46619, stable
Tx analog; ASA, acetlysalicylic acid; PLC, phospholipase C;
PIP2; phosphatidylinositol
4,5-bisphosphate; IP3, inositol 1,4,5-trisphosphate; DAG, 1,2-diacylglycerol.
[View Larger Version of this Image (19K GIF file)]
with established cross-reactivity with rabbit
TNF- was provided by J. C. Mathison (Scripps Research Institute, La
Jolla, CA); Salmonella abortus equii
endotoxin (LPS) was provided by C. Galanos (Freiburg, Germany). AA,
PGG2,
PGH2, and U-46619 were obtained
from Paesel and Lorei (Frankfurt, Germany), and
D,L-lysin-monoacetylsalicylate
(ASA) was from Bayer (Leverkusen, Germany). All other
chemicals were purchased from Merck (Darmstadt, Germany).
1 · kg
body wt1. With the use of
two perfusate reservoirs, the circulating buffer fluid could be
exchanged without interrupting the flow. Before the circulation was
closed, 1,000 ml of buffer fluid were discarded. Left atrial pressure
was set at 2 mmHg by adjusting the height of the left atrial catheter.
The pH of the circulating fluid was adjusted to 7.38 ± 0.4 by
varying the inhaled carbon dioxide concentration. Pulmonary arterial
pressure (PAP; mmHg), pulmonary venous pressure [left atrial
pressure (mmHg)], and lung weight gain (g) were recorded continuously.
Formaldehyde-sterilized and LPS-free perfusion circuit tubing and
sterile solutions were used throughout. Under these conditions, no LPS
could be detected in the recirculating perfusate under baseline
conditions [All values ranged below 10 pg/ml, the detection limit
of the photometric assay used (Kabi Vitrum, Munich, Germany)]. Lungs included in the study 1) had a
homogenous white appearance with no signs of hemostasis, edema, or
atelectasis; 2) had normal PAP and
airway pressure; and 3) were
isogravimetric (lung weight gain <0.3 g/h) during an initial
steady-state period of 30 min. Following these criteria, 15-20%
of all lung preparations were discarded before being used the study.
Biochemical measurements.
Perfusate concentrations of TxB2
and 6-ketoprostaglandin F1
(6-keto-PGF1), the stable
metabolites of TxA2 and
PGI2, respectively, were assayed
by solid-phase extraction and post-high-performance liquid
chromatography (HPLC) enzyme-linked immunosorbent assay (ELISA), as
previously described (10). In vitro control studies ascertained that neither U-46619,
PGG2, nor
PGH2 cross-reacted with the
monoclonal antibody employed for
TxB2 quantification in the
post-HPLC fractions in this assay. Perfusate concentrations of TNF-
were determined by a cytolytic cell assay in the mouse fibrosarcoma
cell line, WEHI 164 clone 13 (donated by T. Espevik), as previously
described (6). The WEHI cells (2 × 104) were
incubated with serial dilutions of perfusate in microtiter wells (Nunc,
Wiesbaden, Germany). After 18 h,
3-(4,5-dimethyl-thiiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT)
(5 mg/ml in phosphate-buffered saline; 100 µl/well) was added. The
reaction was stopped after 4 h by the addition of 5% formic acid in
2-propanol, and the content of reduced MTT was measured in a
micro-ELISA autoreader (570 nm). The amount of rabbit
TNF- was expressed in picograms per milliliter, as measured against
a recombinant murine TNF- standard. To establish the nature of the
cytolytic activity as TNF-, an antiserum directed against human
TNF- with established cross-reactivity with rabbit TNF- (13) was
added to the perfusate. This antiserum neutralized all cytolytic
activity measured with the WEHI cells.
Experimental protocol (Fig. 2).
All studies were performed in buffer-perfused lungs to selectively
study the effects of LPS on lung-resident cells and to exclude effects
on circulating blood cells. Experiments were begun after a steady-state
period of 30 min (time 0). AA,
U-46619, PGG2, or
PGH2 was injected via an injection
port into the pulmonary artery. For each compound, the concentration
(related to the recirculating perfusate) required to induce a pressor
response of 5-15 mmHg under LPS-free conditions was determined in
pilot experiments [for decreased reactivity to
PGG2 see
PAP pressor responses (Fig. 3)]. In a standard
protocol, the following concentrations were used: 30 nM AA
(group 1), 0.3 nM U-46619 (group
2), 3 nM PGG2 (group 3), and 3 nM
PGH2 (group
4). The timing of repetitive bolus applications is given in Fig. 2. Each challenge was applied under three
conditions (subgroups; each subgroup including 4-6 experiments): 1) in LPS-free, sterile perfusate;
2) after pretreatment with 10 ng/ml
LPS, added to the perfusate at time 0;
and 3) after pretreatment with 10 ng/ml LPS and 1 µM ASA, both added to the perfusate at time 0.
and prostanoid analysis were drawn
intermittently. Timing of sampling is indicated by vertical lines below
time axis.
) and in lungs pretreated with LPS (
) or LPS and ASA
(
). Range of initial PAP and PAP between injections, which was
similar in all 3 groups of lungs, is indicated by dotted lines. Data
points represent means ± SE of at least 4 independent experiments.
* Significant difference among all 3 subgroups,
P < 0.05. x Significant difference
between experiments with LPS pretreatment compared with LPS and ASA
pretreatment, P < 0.05.
Evaluation of data and statistical analysis. PAP values taken for evaluation were 1) those directly before each bolus application of the different agents (baseline) and 2) the maximum values within 5 min after stimulus application (peak). Changes in lung weight were compared with the initial baseline value (time 0). Samples for TNF-
were drawn at
time 0 and immediately before each
stimulus application. Prostanoids were analyzed from samples taken at
time 0, immediately before, and 1, 2, 3, 4, and 5 min after each stimulus application. The five
poststimulation values were averaged. All data are reported as means ± SE. Differences among times and groups were assessed by analysis
of variance with repeated measures followed by the Mann-Whitney test
for unpaired samples (NCSS Statistics and Graphics, 5.X
series). Differences were considered to be significant at a P value <0.05.
RESULTS
concentration was
1.1 ± 0.1 ng/ml. At this time point, no TNF- was detectable. In
lungs not undergoing challenges with the different agents, the PAP
values did not change over the subsequent 190-min observation period,
and this was also true for lungs being pretreated with LPS or LPS plus
ASA (data not shown).
PAP pressor responses (Fig. 3).
Bolus application of AA provoked brief increases of the
PAP. Values reached a maximum within 1 min and subsequently decreased rapidly to baseline values. In nonprimed lungs, the height of the
pressure peaks in response to repetitive challenge with AA, as well as
the baseline PAP values, did not change over the entire observation
period. Within 90 min, pretreatment with LPS changed the pulmonary
vascular reactivity to AA injections, resulting in a time-dependent
enhancement of pressor responses, again with unchanged baseline PAP
values. In the presence of ASA, any PAP increase in response to AA
bolus application was suppressed. U-46619 injections elicited constant
pressor responses that were not significantly augmented by LPS
pretreatment and were not suppressed by ASA. PGG2 provoked only very moderate
initial PAP responses that slightly increased over time in the absence
of LPS. This increase was markedly amplified within 90 min in
LPS-pretreated lungs. Any pressor response to
PGG2 was fully blocked by ASA.
PGH2 injections elicited pressor responses that were reproducible over the entire observation period. Within 90 min of LPS pretreatment, a marked increase in vascular reactivity was again noted, with more than threefold amplification of
the pressor responses. In the presence of ASA and LPS, repetitive PGH2 injections still elicted
constant pressure responses, but the time-dependent increased
vasoreactivity, as demonstrated after repetitive
PGH2 injections in lungs treated
only with LPS, was suppressed.
TxB2 concentrations (Fig.
4).
TxB2 release increased
consistently in response to bolus administration of AA in LPS-free
lungs. The TxB2 release progressively increased
after LPS pretreatment and was suppressed by ASA. U-46619 injections
did not induce major changes of TxB2
concentrations from baseline values in the absence of LPS, but
increased TxB2 liberation was noted after LPS
pretreatment and U-46619 administration. This increase was blocked by
ASA. TxB2 release in response to PGG2
injections was very low and increased moderately over time. TxB2 release in response to PGG2
injections did not differ in LPS-pretreated and nonpretreated lungs and
was unaffected by ASA treatment. Similar to PGG2,
PGH2 provoked moderate TxB2 release, which increased over time. TxB2 release was increased
nearly 10-fold after LPS pretreatment and was fully blocked by ASA.
) and in lungs pretreated with LPS () or LPS and ASA
(). After each injection, 5 samples were drawn from perfusate, and
measured concentrations were averaged. Data points represent means ± SE of at least 4 experiments. * Significant difference
among all 3 subgroups, P < 0.05. x Significant difference
between experiments with LPS pretreatment compared with LPS and ASA
pretreatment, P < 0.05.
6-keto-PGF1
concentrations (Fig. 5).
All the agents tested induced some release of
6-keto-PGF1, which was largely constant over the
190-min observation period. LPS pretreatment enhanced the
6-keto-PGF1 release in response to AA, U-46619, and
PGH2, whereas the release provoked by
PGG2 was not signicantly amplified. In the presence of
ASA, all LPS-related increases of 6-keto-PGF1
liberation were blocked.
concentrations in isolated lungs after repetitive injections of 30 nM
AA (A), 0.3 nM U-46619
(B), 3 nM
PGG2
(C), or 3 nM
PGH2
(D) into pulmonary artery of sterile lungs () and in lungs pretreated with LPS () or LPS and ASA (). After each injection, 5 samples were drawn from perfusate, and
measured concentrations were averaged. Data points represent means ± SE of at least 4 experiments. * Significant difference among all 3 subgroups, P < 0.05. x Significant difference
between experiments with LPS pretreatment compared with LPS and ASA
pretreatment, P < 0.05.
TNF-
release (Fig. 6).
In LPS-free experiments, only a very moderate liberation of
TNF- was measured. LPS pretreatment was associated with marked TNF- release, with concentrations approaching 800-1,100 pg/ml. This cytokine response was not significantly affected by ASA.
concentrations of sterile lungs () and in lungs
pretreated with LPS () or LPS and ASA ().
A: 30 nM AA. B: 0.3 nM U-46619.
C: 3 nM
PGG2.
D: 3 nM
PGH2. Samples were drawn 5 min
before compound injections. Data points represent means ± SE of at
least 4 experiments. Analysis of variance revealed a significant
difference between subgroups of lungs stimulated with AA
(P < 0.05), U-46619
(P < 0.05), and
PGG2
(P < 0.001).
PAP peak decay after AA and U-46619 challenge (Fig. 7). To compare the duration of the vasoconstrictor responses, the decrease of PAP after peak pressure was reached was analyzed for bolus applications of AA and U-46619 in nonprimed lungs. After challenges with AA, PAP decreased to 49.5 ± 3.8% of the maximal response within 2 min. After challenges with U-46619, PAP decreased to 50.4 ± 5.5% of the maximal response within 3 min. The differences between AA and U-46619 were significant at these time points (P < 0.05).
) or AA () in
nonprimed lungs. Note that PAP after AA injections returned to its half
maximum within 2 min, whereas half maximum after U-46619 injections was
reached after 3 min. Maximum increase of PAP during first minute after
a bolus injection was considered to represent 100%. Each data point
represents mean ± SE of at least 24 peaks in each group (4-6
experiments with 6 stimulations each). 
, Change. * Significant
differences between corresponding values,
P < 0.05.
Weight gain. All lungs gained weight during the maneuvers of repetitive stimulus-evoked pulmonary vascular pressure elevations. At the end of experiments, the total weight gain ranged between 3 and 8 g in LPS-free lungs, 7 and 21 g in LPS-pretreated lungs, and 3 and 10 g under conditions of ASA pretreatment. No attempts were undertaken to determine whether the greater weight gain in LPS-primed lungs was due to increased vascular permeability or to increased in pressure-induced fluid filtration as a result of the enhanced pressor responses.
DISCUSSION
The results obtained with direct administration of the precursor fatty
acid AA clearly point to an enhancement of Cyclo activity in LPS-primed
lungs and thus support a suggestion of increased Cyclo activity in
earlier studies in endotoxin-exposed dog lungs (22). After an initial
delay of 60-90 min, which corresponds to the time dependency
observed for LPS priming of
Staphylococcal -toxin- and
E. coli hemolysin-elicited pressor
responses (25, 27), the PAP increased two- to threefold after AA
administration. This increase was accompanied by enhanced
TxB2 and
6-keto-PGF1 release. Both the
increased vasoreactivity and enhanced prostanoid generation were
completely blocked in the presence of the Cyclo inhibitor ASA.
Moreover, because the PAP increase in response to the stable Tx analog
U-46619 was not enhanced after LPS priming, the augmentation of pressor
responses to AA may not be attributed to alterations of signaling
events downstream to the occupany of the
TxA2-PGH2 receptor. Augmentation
of the vasoconstrictor response occurs with increased synthesis of both
agents because the vasoconstrictive potency of
TxA2 is greater than the
vasodilatory effects of PGI2 (20).
The cellular source of Tx released into the buffer medium of perfused
lungs is not established. Lung macrophages and endothelium-adherent
monocytes (5) represent likely candidates. Preincubation of macrophages
with LPS in vitro primes these cells for accelerated and enhanced AA
metabolite release in response to various stimuli (1, 2,
16).
Increase of lung Cyclo activity might also be responsible for the ASA-inhibitable enhancement of pressor responses to PGG2, which is converted to PGH2 by peroxidase activity of the Cyclo complex as a subsequent enzymatic step (14). But this increase in vasoreactivity was not accompanied by significant increases of TxA2 or PGI2 synthesis in our study, which would be expected to accept such an explanation. It has to be kept in mind, however, that PGG2 is very unstable and probably undergoes rapid decomposition after injection into the pulmonary artery. The moderate response to PGG2 in our study may be due to only a small fraction of intact PGG2 gaining access to Cyclo in different cell types. PGG2 may, therefore, not be an ideal probe for the peroxidase step of the Cyclo complex in an intact organ. In contrast to PGG2, PGH2 provoked marked pressor responses in LPS-primed lungs. These were accompanied by very high quantities of both TxA2 and PGI2 release. Clearly, this finding may not solely be explained by enhanced Cyclo activity in the LPS-pretreated lungs, because PGH2 is a product and not a substrate of this enzyme complex. The amplification of PGH2-provoked prostanoid formation in LPS-primed lungs might be due to increased the activities of Tx synthase and PGI2 synthase (24). However, the data with coapplication of ASA, which inhibits Cyclo but not Tx and PGI2 synthase (11), strongly argue against such a mechanism. In the presence of ASA, some baseline pressor response to PGH2 was maintained, which corresponds well to the PAP elevation by PGH2 in nonprimed lungs and may well be related to direct occupancy of the TxA2-PGH2 receptor by this agent. Any extra PAP increase and TxA2 and PGI2 synthesis were blocked by ASA. Thus events upstream of the site of ASA interference must be involved in the exaggerated responsiveness to PGH2 in LPS-primed lungs. These events must necessarily include the liberation of endogenous AA. The extensive prostanoid formation in response to PGH2, related to an extra increase in PAP, could be explained by a secondary loop of PGH2-effected liberation of endogenous AA via pathways facilitated by LPS priming.
Such additional LPS priming of pathways resulting in enhanced liberation of endogenous AA might also explain the finding that the stable Tx analog U-46619 effected significant liberation of both TxA2 and PGI2 in LPS-pretreated but not in control lungs. In contrast to the experiments with PGH2, however, such additional prostanoid and in particular Tx generation caused only insignificant increases of the U-46619-elicited pressor responses. Two explanations are possible for this observation: 1) the U-46619-evoked secondary TxA2 synthesis was severalfold less than that elicited by PGH2 and 2) the occupancy of the TxA2-PGH2 receptor by U-46619 has a longer duration than that of the native products TxA2 and PGH2, as reflected by the longer PAP decay time on application of U-46619 compared with AA (Fig. 6). This might interfere with the access of additionally formed endogenous Tx to the common receptor. Together, these differences may well explain the observation that secondary prostanoid generation was stimulated by both PGH2 and U-46619 subsequent to LPS pretreatment but significantly contributed to the pressor responses in LPS-primed lungs only in the case of PGH2.
The LPS exposure of the blood-free vasculature of the perfused lungs
was accompanied by progressive liberation TNF-. Besides being an
independent indicator of LPS activity, this cytokine might also be
involved in the observed changes of AA metabolism. TNF- was
previously reported to enhance lung vasoconstrictor responses to
secondarily applied vasoactive agents such as
N-formyl-L-methionyl-L-leucyl-L-phenylalanine (9), angiotensin II (23), and U-46619 (21). It has to be kept in mind,
however, that the maximum TNF- generation occurred after 90 min, in
other words at a time when the priming of the Tx-related
vasoconstrictor response was already fully evident.
In conclusion, LPS priming of lung Tx generation and Tx-related vasoconstrictor responses was demonstrated for direct application of the precursor fatty acid AA. The time course of this event, the enhanced generation of both TxA2 and PGI2, the susceptibility of the response to Cyclo inhibition, and the relative constancy of the pressor responses to the direct vasoconstrictor agent U-46619 support the notion that an increase of lung Cyclo activity is the predominant underlying event (Fig. 1). In addition, the capability of the PG intermediate PGH2 and the stable Tx analog U-46619, which act downstream of the Cyclo complex, to effect secondary generation of TxA2 and PGI2 in LPS-pretreated lungs indicates that secondary AA release is facilitated by LPS priming, which may then provide endogenous AA for further Cyclo metabolism (Fig. 1). Overall, LPS priming of perfused rabbit lungs, although devoid of direct hemodynamic effects, exerts a profound impact on the prostanoid homeostasis in this vasculature and results in markedly enhanced readiness to react with Tx-related vasoconstrictor responses to various stimuli.
ACKNOWLEDGEMENTS
We thank Carmen Schmidt and Florian Michnazs for excellent technical assistance and Ralf Czimek for support in the performance of the cytotoxicity assays.
FOOTNOTES
Address for reprint requests: W. Seeger, Dept. of Internal Medicine, Justus-Liebig Univ. Giessen, Klinikstrasse 36, 35392 Giessen, Germany.
Received 14 May 1996; accepted in final form 26 February 1997.
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