American Journal of Respiratory and Critical Care Medicine

The effects of conventional mechanical ventilation (CMV) and high-frequency oscillatory ventilation (HFO) on intraalveolar expression of the tumor necrosis factor- α (TNF- α ) gene were studied in surfactant-depleted rabbits. After lung lavage with saline, 13 rabbits were administered either CMV (n = 6) or HFO (n = 7) for 1 h at an Fi O2 of 1.0 and a mean airway pressure of 13 cm H2O. Lung lavage was then repeated. The rabbits' RNA was extracted from the lavage cells, and mRNA for TNF- α was quantitated by reverse-transcription polymerase chain reaction using glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as an internal standard. At 1 h of ventilation, PaO2 was slightly lower with CMV than HFO, while lavage cell counts and cytology were similar between the two groups. The ratio of TNF- α mRNA to GAPDH mRNA increased with CMV (control, 0.48 ± 0.04 [SE] versus 1 h, 1.02 ± 0.14, p < 0.01) but did not change with HFO (0.55 ± 0.07 versus 0.73 ± 0.09). In a separate series of experiments, ten surfactant-depleted rabbits continued to be ventilated for 4 h either by CMV (n = 5) or HFO (n = 5). Conventional mechanical ventilation resulted in a progressive hypoxemia, decreased lung compliance, increased number of neutrophils in lung lavage fluid, and substantial morphological changes including hyaline membrane formation and neutrophil accumulation, whereas HFO was associated with minimal changes in such physiological and pathological abnormalities. These results suggest that activation of alveolar macrophages and production of proinflammatory cytokines may play a pivotal role in the early stage of ventilator-induced lung injury, and that ventilator mode (CMV or HFO) substantially modulates macrophage activation and hence the degree of lung injury.

Mechanical ventilation with intermittent positive pressure can by itself produce or worsen lung injury (1). A number of different ventilatory strategies have been developed in an attempt to reduce ventilator-induced lung injury (2). High-frequency oscillatory ventilation (HFO) is increasingly recognized as an effective alternative ventilator mode in neonatal and pediatric respiratory failure (3-5). Both clinical and experimental evidence suggests that HFO is associated with much less lung injury than conventional mechanical ventilation, or CMV (3, 5-9). However, the mechanism by which such different ventilator strategies affect progression of lung injury has not been well understood.

Various ventilator-dependent factors have been shown to be relevant in pathogenesis of ventilator-induced lung injury. These include high peak airway pressure (10-12), large tidal volumes (13, 14), and collapse and reopening of small airways with ventilation at low lung volumes (6, 8 15–17). A question that remains to be addressed is how such mechanical stresses and resultant insults on airway epithelium lead to diffuse lung damage, e.g., endothelial leakage, pulmonary edema, surfactant dysfunction, and hyaline membrane formation. Several studies suggest that the presence of neutrophils and their infiltration into the lung are essential in CMV-induced lung injury (6, 18). In contrast, HFO was found to be associated with less activation of neutrophils (19, 20) and less release of inflammatory chemical mediators such as platelet-activating factor (PAF) or prostaglandins (9, 21) in the lung. Thus, it seems a reasonable assumption that the inflammatory cells and mediators, which have been implicated in pathogenesis of adult respiratory distress syndrome (ARDS) (22), may also play an important role in progression of ventilator-induced lung injury. The differences in pulmonary inflammatory response may explain the marked differences in lung physiology and pathology observed between CMV and HFO (6-9).

The goal of the present study is to better understand the inflammatory aspects in the pathogenesis of ventilator-induced lung injury and evaluate the influence of different ventilator strategies, i.e., CMV versus HFO, on such pulmonary inflammatory response. We hypothesized that mechanical stress imposed on airway epithelium with CMV may lead to activation of alveolar macrophages, which then release proinflammatory cytokines and initiate an inflammatory cascade in the lung, including production of chemical mediators and neutrophil infiltration. On the other hand, oscillatory movement with HFO may produce less activation of alveolar macrophages, resulting in less pulmonary inflammatory response and less lung injury than CMV. With this general hypothesis, the present study investigated the role of alveolar macrophages in the early stage of ventilator-induced lung injury. Gene expression of one of the representative proinflammatory cytokines, tumor necrosis factor-α (TNF-α), was used as an index of macrophage activation, and we compared intraalveolar TNF-α mRNA levels in surfactant-depleted rabbit lungs ventilated by CMV or HFO.

Animal Preparation

The study protocol was approved by the Institutional Animal Care and Use Committee of the National Children's Medical Research Center in Tokyo. Twenty-three male Japanese white rabbits weighing 2.39 ± 0.10 (SE) kg were premedicated with intramuscular injections of ketamine hydrochloride (10 mg/kg). A plastic catheter was inserted into an ear vein, and the animal was anesthetized by intravenous infusion of ketamine (8 mg/h) and pancuronium bromide (0.3 mg/h) in 10 ml/h of 5% dextrose in Ringer's lactate solution. A 4.0 mm ID endotracheal tube was inserted via a tracheostomy and sutured in place while the animal was manually ventilated. The animal was placed on a piston pump HFO ventilator (Humming II; Senko Medical Instruments, Tokyo, Japan) at an inspiratory oxygen fraction (Fi O2 ) of 1.0, oscillatory frequency of 15 Hz, and mean airway pressure (MAP) of 7 cm H2O. The stroke volume of the sinusoidal oscillation was adjusted to maintain PaCO2 levels between 35 and 45 mm Hg. A fluid-filled catheter was introduced into the femoral artery for monitoring blood pressure and sampling arterial blood. Blood gas determinations were intermittently performed using a pH/blood gas analyzer (Model 178; Corning Medical, Medfield, MA). Body temperature was maintained between 37 and 39° C with a heating pad. The animals were then divided into two series of experiments.

Experimental Protocols

Series 1: Intraalveolar TNF-α mRNA levels with 1-hour ventilation. The purpose of this experiment was to compare the degree of activation of alveolar macrophages between CMV and HFO in the early stage of ventilator-induced lung injury. Once the 13 animals in this series were stabilized on HFO, lung lavage was performed three times with normal saline to produce surfactant depletion. This procedure in rabbits, when the animals are subsequently ventilated by CMV for several hours, produces a progressive ARDS-like lung injury characterized by diffuse microatelectasis, pulmonary edema, infiltration of neutrophils, and hyaline membrane formation (6, 18-21, 23). For each lavage, a 30 ml/kg aliquot of warmed saline was flushed into and out of the lung five times via tracheostomy tube and gently sucked out thereafter. Between each lavage, a sustained inflation of 30 cm H2O for 10 s was applied and MAP was raised by 2–3 cm H2O to minimize further airway collapse and progression of hypoxia. After the third lavage, the animal was restabilized on HFO with a MAP of 13 cm H2O. With several sustained inflation maneuvers, PaO2 of the animal soon returned to the prelavage level, i.e., more than 350 mm Hg. The drained lavage fluid was collected as the control lavage sample.

Each animal was then randomly assigned to receive either CMV (n = 6) or HFO (n = 7) at an Fi O2 of 1.0 and MAP of 13 cm H2O for 1 h. Conventional ventilation was performed by use of the CMV mode of the Humming II ventilator. Peak inspiratory pressure and positive end-expiratory pressure (PEEP) were set at 28 and 5 cm H2O, respectively. Inspiratory time and respiratory frequency were initially set at 1.0 s and 25 breaths/min, and later adjusted to achieve normocapnia while maintaining MAP at 13 cm H2O. In the other group, HFO was performed at an oscillatory frequency of 15 Hz throughout the experiment, and normocapnia was maintained by adjusting the stroke volume. Arterial blood gases were measured every 30 min. After 1 h of mechanical ventilation, lung lavage was again performed as described above to collect the “1-hour” lavage sample.

The control and 1-hour lavage fluid samples were analyzed for cytology and mRNA quantification. The samples were centrifuged at 400 × g for 5 min at 4° C, and the cell pellets were washed twice by phosphate-buffered saline. The number of total lavage cells was counted under light microscopy, and differential cytology was performed by use of Wright-Giemsa stained preparations. The isolated cell pellets were analyzed for TNF-α mRNA, as described later in detail.

Series 2: Physiological changes with 4-hour ventilation. The purpose of this experiment was to confirm whether clinically relevant physiological and pathological changes indicating lung injury would take place if the surfactant-depleted rabbits were ventilated longer either by CMV or HFO. After removal of lung surfactant by saline lavage, the 10 animals in this series were randomized to receive either CMV (n = 5) or HFO (n = 5) at an Fi O2 of 1.0 and MAP of 13 cm H2O for 4 h. Exactly the same protocol as in Series 1 was used for lavage procedure and adjustment of ventilator settings. Only the duration of ventilation was different. Arterial blood gases were monitored every hour. Total respiratory system compliance (Crs) was measured before lung lavage and after 4 h of ventilation, using the passive expiratory flow-volume technique (24) with a Fleisch #0 pneumotachograph, a 3-way occlusion valve, and a personal computer–based pulmonary function testing system (Aivision Co., Tokyo, Japan), as previously described (25). After 4 h of mechanical ventilation, lung lavage was repeated and the lavage fluid samples were analyzed for cell counts and differential cytology. The animals were killed by pentobarbital overdose, their lungs were fixed by instillation of 10% buffered Formalin at a transpulmonary pressure of 15 cm H2O, and midsagittal cross sections were stained with hematoxylin and eosin for postmortem microscopic examination.

Reverse Transcription Polymerase Chain Reaction (RT-PCR)

Total RNA was extracted from the isolated lavage cell pellets of Series 1 animals by the acid guanidinium-phenol-chloroform technique, as described by Chomczynski and Sacchi (26). Two μg of total RNA was used for the synthesis of first-strand cDNA with reverse transcriptase (Superscript™; Gibco BRL, Gaithersburg, MD) and random hexanucleotides (pd [N]6; Pharmacia Biotech). The reaction was terminated by heating for 10 min at 95° C before PCR.

Oligonucleotide PCR primers were synthesized on a DNA synthesizer (392 DNA/RNA synthesizer; Applied Biosystem) from the published cDNA sequences of rabbit TNF-α (27, 28) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (29). The sequences of these primers are shown in Table 1. We used the GAPDH gene as an internal control to standardize PCR products. The PCR reaction mixture contained one-fortieth of each cDNA sample, the upstream (5′) and downstream (3′) primers for TNF-α and GAPDH (0.3 μM of each primer), dNTP mixture (200 μM of each dNTP), and 1.25 units of Thermus aquaticus DNA polymerase (TaKaRa Biomedicals, Tokyo, Japan). Then 32P-end-labeled 5′ primers (approximately 1 × 106 cpm for each primer) were added to the reaction mixture to quantify the amplified products. The amplification was performed by a thermal cycler (Perkin-Elmer Co., Norwalk, CT) for 27 cycles with denaturation at 95° C for 1 min, annealing at 60° C for 1 min, and extension at 72° C for 2 min. The PCR products were separated by electrophoresis using a 2% (wt/vol) agarose gel containing 0.5 μg/ml of ethidium bromide. Two clear bands, each corresponding to the predicted size of the amplified product for TNF-α mRNA (251 bp) or GAPDH mRNA (506 bp), were identified under ultraviolet transilluminator. The gels were dried, and radioactivity for each band was quantified by a radioanalytic imaging system (BAS 2000 Bio Image Analyzer; Fujix Film Co., Tokyo, Japan). The amount of TNF-α mRNA in each lavage sample was expressed by the radioactivity ratio of TNF-α mRNA to GAPDH mRNA.


Oligonucleotide SequencesAnneals toPredicted cDNA size
PCR primers
 TNF-α251 bp
  Upstream5′ AGCCCACGTAGTAGCAAACCC 3′bp 446-466
  Downstream5′ TTGATGGCAGAGAGGAGGTTGA 3′bp 676-697
 GAPDH506 bp
  Upstream5′ ATGTTTGTGATGGGCGTGAACC 3′bp 456-477
  Downstream5′ CCCAGCATCGAAGGTAGAGGA 3′bp 942-962
Probes for southern blotting

Base numbers refer to data described in References 27–29. Upstream and downstream PCR primers are taken from the sense and antisense sequences, respectively. Probes for southern blotting code antisense sequences.

To verify that the amplified products derived from our primer pairs were authentic, a separate set of PCR procedures was performed without radiolabeled primers. The amplified products were separated on a gel, transferred to a nylon filter membrane, and hybridized with a 32P-labeled oligonucleotide probe for either TNF-α or GAPDH. The sequences for these oligonucleotide probes were chosen from sequences midway between the upstream and downstream PCR primers (see Table 1). The radioactive probes hybridized only to the bands which corresponded in size to the ethidium bromide–stained gels, thereby confirming that the amplified PCR products consisted of correct TNF-α or GAPDH cDNA sequences.

The condition for PCR used in this study was carefully chosen to ensure that the amount of product synthesized was proportionally related to the amount of mRNA in the original preparation (30, 31). In a separate set of experiments, we analyzed the effects of PCR cycle numbers on the yield of amplified products for TNF-α and GAPDH under our experimental condition. At a cycle number of 27, our PCR assay was confirmed to be in the linear range during the amplification phase (Figure 1).

Data Analysis

Data are expressed as means ± SE. Changes in PaO2 , Crs, total and differential cell counts, and TNF-α mRNA expression over the course of mechanical ventilation were evaluated by paired t tests or one-way analysis of variance (ANOVA) for repeated measures with Scheffe's tests. Differences between CMV and HFO were analyzed either by unpaired t tests or two-way ANOVA for repeated measures. An alpha error of less than 0.05 was considered significant.

Intraalveolar TNF- α mRNA Levels with 1-Hour Ventilation (Series 1)

Figure 2 shows changes in PaO2 values in the Series 1 experiments. With CMV, PaO2 showed a small decrease at 1 h (p < 0.05) but still remained greater than 250 mm Hg. In contrast, PaO2 did not change with HFO. Changes in cell counts and cytology in the lung lavage samples are summarized in Figure 3. The total cell counts substantially decreased in the 1-hour lavage both with CMV and HFO (p < 0.05, Figure 3A). Cells detected in the lavage fluid consisted of alveolar macrophages, neutrophils, and a few epithelial cells. In the control lavage, almost all of the recovered cells were alveolar macrophages (> 97%). In the 1-hour lavage, the recovered cells included approximately 20% neutrophils with both CMV and HFO. There was no difference in neutrophil percentage, in either the control or 1-hour lavage, between CMV and HFO (Figure 3B).

Figure 4 illustrates typical PCR products obtained on gel electrophoresis by the radioanalytic imaging system. With CMV, mRNA for TNF-α showed a large increase in the 1-hour lavage compared with the control. In contrast, with HFO, the amount of TNF-α mRNA was almost similar between the control and 1-hour lavage. Changes in the relative amount of TNF-α message, as expressed by the mRNA ratio of TNF-α to GAPDH, are summarized in Figure 5. At 1 h of ventilation, TNF-α mRNA increased with CMV (p < 0.01) but not with HFO. The differences in changes in TNF-α mRNA between CMV and HFO were confirmed by a significant interaction p value (0.02) by two-way ANOVA.

Physiological Changes with 4-Hour Ventilation (Series 2)

Figure 6 shows changes in PaO2 values during the course of the Series 2 experiments. With CMV, PaO2 slightly decreased at 1 h and then showed a profound decrease to a level less than 100 mm Hg at 4 h (p < 0.01). In contrast, PaO2 with HFO did not show large changes and essentially remained at the prelavage level throughout the experiment. Changes in Crs and lavage cell cytology after 4-h mechanical ventilation are summarized in Figures 7 and 8. Compliance decreased with either mode of ventilation (p < 0.05), but the decrease in Crs was greater with CMV than HFO (two-way ANOVA interaction p = 0.01, Figure 7). The number of total cell counts in the 4-hour lavage were higher with CMV than HFO (p < 0.05, Figure 8A). The differential cytology demonstrated that the percentage of neutrophils was greater with CMV (95 ± 2%) than HFO (30 ± 7%) (p < 0.01, Figure 8B).

Figure 9 shows typical pathological findings obtained with 4 h of CMV or HFO. Extensive hyaline membrane formation and neutrophil accumulation in the terminal airways and alveoli were observed with CMV (Figure 9A). In contrast, the lungs ventilated with HFO showed much less hyaline membrane formation and neutrophil infiltration than CMV, and most of the alveoli were still well expanded with little evidence of degradation of normal lung architecture (Figure 9B).

In surfactant-depleted rabbits, we found that CMV for only 1 h produced large increases in TNF-α mRNA in the intraalveolar cells, presumably the alveolar macrophages, while HFO of the same duration produced minimal increases in TNF-α gene transcripts. With 4 h of mechanical ventilation, rabbits undergoing CMV developed substantial physiological and pathological changes indicating lung injury, whereas those on HFO did not. These findings suggest that activation of alveolar macrophages and production of proinflammatory cytokines such as TNF-α may play a pivotal role in the early stage of CMV-induced lung injury, and that HFO may produce less activation of alveolar macrophages, thereby leading to less lung injury than CMV.

In the present study we used saline-lavaged ventilated rabbits, because that is an established, extensively investigated model of ventilator-induced lung injury. A number of investigators have used this model to evaluate the effects of different ventilator strategies on lung injury, particularly the effects of HFO as compared with CMV (6, 8, 18-21, 23). Physiological and pathological profiles of the progressive ARDS-like lung injury in these rabbits have been well characterized and found experimentally reproducible. The Series 2 experiments showed that CMV of 4 h in these rabbits produced a progressive hypoxemia, decreased lung compliance, increased number of neutrophils in the lung lavage fluid, and substantial morphological changes in the lungs, including hyaline membrane formation and neutrophil accumulation. In contrast, HFO of 4 h was associated with minimal changes in lung physiology and pathology. These findings, essentially consistent with the previous studies in the literature, demonstrated that with the experimental protocol employed in the present study, the different ventilator strategies (CMV versus HFO) actually produce clinically significant differences in the degree of lung injury in these rabbits.

With only 1 h of mechanical ventilation in the Series 1 experiments, however, there were virtually no large differences in blood gas or lung lavage cytology between CMV and HFO. The PaO2 at 1 h was slightly lower with CMV, but the animal was still well oxygenated. The cell counts decreased at 1 h with both CMV and HFO, presumably because the control lavage removed most of the alveolar macrophages, and a 1 h period was not long enough to recruit cells from the blood. The neutrophil percentage in the 1-hour lavage increased but remained relatively small (about 20%) in both groups, without any difference between CMV and HFO. Thus, after 1 h of mechanical ventilation, the pathological process of ventilator- induced lung injury in these rabbits had not been apparent or fully initiated with either CMV or HFO.

Despite the lack of significant abnormalities in the physiological parameters, the RT-PCR technique demonstrated a clear difference in the amount of TNF-α mRNA in the lavage cells between CMV and HFO at 1 h. This implies that the activation of intraalveolar cells occurs very shortly after the initiation of CMV, before any physiological signs of lung injury become apparent. Moreover, HFO substantially attenuated the degree of the cell activation and cytokine mRNA expression. Although the levels of rabbit TNF-α protein were not directly measured in this study, the enhanced TNF-α gene expression at the transcriptional level would soon lead to actual production of TNF-α protein with CMV. The increased TNF-α protein should then initiate inflammatory cascade in the lung, resulting in production of lipid mediators such as PAF or prostaglandins and neutrophil infiltration and activation in the alveolar space, as observed in the prior studies by ourselves (19, 21) and others (6, 20). Once all of the above inflammatory processes are fully activated in the lung, significant physiological and pathological abnormalities would take place, as in the Series 2 animals with CMV. Our findings thus provide a molecular basis for how different ventilator patterns influence the initial pulmonary inflammatory response in the early stage of ventilator-induced lung injury. The degree of activation of intraalveolar cells and cytokine gene expression may later lead to the marked differences in the degree of lung injury between CMV and HFO.

We performed the RT-PCR analysis for TNF-α on the total cell component of each lavage sample, without separating the macrophages from neutrophils. This was because the rabbit alveolar macrophages stimulated by ventilation had diverse and nonhomogeneous specific gravities such that ordinary Ficoll-Hypaque centrifuge methods failed to reliably separate them from the neutrophils. In addition, we did not wish to purify the macrophages by use of any adherence techniques, which may result in inadvertent induction of cytokine mRNA expression. We were therefore unable to draw a definite conclusion regarding the cell source for the increased TNF-α gene transcripts. However, there is ample evidence in the literature that the predominant sources of TNF-α in the lung are the alveolar macrophages (32). Xing and colleagues found in rat lungs that the neutrophils can produce TNF-α protein in response to lipopolysaccharide stimulation, but the alveolar macrophages serve as the predominant source for TNF-α in the early stage of lung inflammation (33). Moreover, we found in two of the CMV-treated rabbits that almost all of the cells (> 95%) recovered in the 1-hour lavage were the macrophages, yet TNF-α mRNA substantially increased similarly to the other CMV-treated animals. Thus, it seems reasonable to speculate that the major source of the increased TNF-α message in this study were the alveolar macrophages, not neutrophils.

Among the numerous cytokines and chemical mediators produced by alveolar macrophages, we focused on TNF-α and used its gene transcripts as an index of macrophage activation. Our main goal was to quantitatively evaluate the level of macrophage activation in the early stage of ventilator-induced lung injury. The relative impact of TNF-α as compared with those of other proinflammatory mediators in pathogenesis of lung injury was not directly tested and hence remains to be further investigated. However, TNF-α has been shown to be a principal mediator in the early phase of various inflammatory processes, with substantial effects and stimulations on other inflammatory cells and mediators (32). Tumor necrosis factor-α is now recognized as the key proinflammatory cytokine to initiate and amplify the process of multiorgan failure during sepsis, and it has also been implicated in progression of ARDS (22, 32, 34-37). Taken together, these findings make it likely that TNF-α may also play an important role in the initial stage of ventilator-induced lung injury. The similar TNF-α–triggered inflammatory cascade, as observed in sepsis/ARDS syndrome, may explain a large part of the progression of lung injury with CMV, though the first triggering signal is not chemical (lipopolysaccharide) but mechanical insult (stress on airway epithelium).

Since the advent of mechanical ventilation, clinicians and scientists have realized that intermittent positive-pressure ventilation per se exacerbates lung injury. Experimental studies have clarified the relative importance of various ventilator-dependent factors in the progression of lung injury (1). High peak inspiratory pressures (10-12) or large tidal volumes (13, 14) are generally accepted as the key factors, imposing overdistending stresses on airway epithelium and producing epithelial and endothelial damages. Recent studies further demonstrated that moderate levels of PEEP may actually protect the lung from ventilator-induced damage (13, 16, 17), consistent with the notion made by Robertson that intermittent collapse and reopening of small airways or alveolar ducts and the resultant high shear stresses on airway epithelium are responsible for CMV-induced lung injury (15). Bryan and Froese utilized this concept to explain why HFO with appropriate volume-recruitment protocols does not produce epithelial damage (6, 8). Based on these findings the currently accepted strategies for preventing lung injury focus mainly on optimization of such mechanical factors, i.e. maintaining the lowest possible inspiratory pressures or tidal volumes while volume-recruiting the lung with active application of PEEP (1, 2) or using newly developed, less invasive modes of ventilation such as HFO in pediatrics (3-5).

The results of this study, together with the previous reports clarifying the role of neutrophils (6, 18-20) and chemical mediators (9, 21), suggest that the inflammatory events are also involved from the very early stage of ventilator-induced lung injury. Both the mechanical and inflammatory processes appear to be essential for progression of diffuse lung damage during mechanical ventilation. Therefore, in addition to the use of less invasive ventilator strategies such as HFO, pharmacological or immunological modulation of the inflammatory cells and mediators may provide a useful way to attenuate ventilator-induced lung injury in neonatal and adult respiratory distress syndrome.

The authors thank Doctors Y. Imai, T. Kawano, and K. D. Bloch for critical reviews of this work.

Supported by Grant H8-PK8-02, Ministry of Health and Welfare, Japan.

1. Parker J. C., Hernandez L. A., Peevy K.Mechanisms of ventilator-induced lung injury. Crit. Care Med211993131143
2. Marini, J. J. 1993. Mechanical ventilation and newer ventilatory techniques. In R. C. Bone, D. R. Dantzker, R. B. George, R. A. Matthay, and H. Y. Reynolds, editors. Pulmonary and Critical Care Medicine, Vol. 3, Part R, Chapter 5. Mosby-Year Book, Inc., St. Louis. 1–26.
3. Clark R. H., Gerstmann D. R., Null D. J., deLemos R. A.Prospective randomized comparison of high-frequency oscillatory and conventional ventilation in respiratory distress syndrome. Pediatrics891992512
4. Ogawa Y., Miyasaka K., Kawano T., Imura S., Inukai K., Okuyama K., Oguchi K., Togari H., Nishida H., Mishina J.A multicenter randomized trial of high frequency oscillatory ventilation as compared with conventional mechanical ventilation in preterm infants with respiratory failure. Early Hum. Dev321993110
5. Arnold J. H., Hanson J. H., Toro F. L., Gutierrez J., Berens R. J., Anglin D. L.Prospective, randomized comparison of high-frequency oscillatory ventilation and conventional mechanical ventilation in pediatric respiratory failure. Crit. Care Med22199415301539
6. Hamilton P. P., Onayemi A., Smyth J. A., Gillan J. E., Cutz E., Froese A. B., Bryan A. C.Comparison of conventional and high-frequency ventilation: oxygenation and lung pathology. J. Appl. Physiol551983131138
7. Delemos R. A., Coalson J. J., Gerstmann D. R., Null D. J., Ackerman N. B., Escobedo M. B., Robotham J. L., Kuehl T. J.Ventilatory management of infant baboons with hyaline membrane disease: the use of high frequency ventilation. Pediatr. Res211987594602
8. McCulloch P. R., Forkert P. G., Froese A. B.Lung volume maintenance prevents lung injury during high frequency oscillatory ventilation in surfactant-deficient rabbits. Am. Rev. Respir. Dis137198811851192
9. Meredith K. S., deLemos R. A., Coalson J. J., King R. J., Gerstmann D. R., Kumar R., Kuehl T. J., Winter D. C., Taylor A., Clark R. H., Null D. M. J.Role of lung injury in the pathogenesis of hyaline membrane disease in premature baboons. J. Appl. Physiol66198921502158
10. Egan E. A., Nelson R. M., Olver R. E.Lung inflation and alveolar permeability to non-electrolytes in the adult sheep in vivo. J. Physiol. (Lond.)2601976409424
11. Parker J. C., Townsley M. I., Rippe B., Taylor A. E., Thigpen J.Increased microvascular permeability in dog lungs due to high peak airway pressures. J. Appl. Physiol57198418091816
12. Dreyfuss D., Basset G., Soler P., Saumon G.Intermittent positive-pressure hyperventilation with high inflation pressures produces pulmonary microvascular injury in rats. Am. Rev. Respir. Dis.1321985880884
13. Dreyfuss D., Soler P., Basset G., Saumon G.High inflation pressure pulmonary edema: respective effects of high airway pressure, high tidal volume, and positive end-expiratory pressure. Am. Rev. Respir. Dis.137198811591164
14. Hernandez L. A., Peevy K. J., Moise A. A., Parker J. C.Chest wall restriction limits high airway pressure-induced lung injury in young rabbits. J. Appl. Physiol.66198923642368
15. Robertson B.Current and counter-current theories on lung surfactant. Scand. J. Respir. Dis571976199207
16. Sandhar B. K., Niblett D. J., Argiras E. P., Dunnill M. S., Sykes M. K.Effects of positive end-expiratory pressure on hyaline membrane formation in a rabbit model of the neonatal respiratory distress syndrome. Intensive Care Med141988538546
17. Muscedere J. G., Mullen J. B., Gan K., Slutsky A. S.Tidal ventilation at low airway pressures can augment lung injury. Am. J. Respir. Crit. Care Med.149199413271334
18. Kawano T., Mori S., Cybulsky M., Burger R., Ballin A., Cutz E., Bryan A. C.Effect of granulocyte depletion in a ventilated surfactant-depleted lung. J. Appl. Physiol6219872733
19. Matsuoka T., Kawano T., Miyasaka K.Role of high-frequency ventilation in surfactant-depleted lung injury as measured by granulocytes. J. Appl. Physiol761994539544
20. Sugiura M., McCulloch P. R., Wren S., Dawson R. H., Froese A. B.Ventilator pattern influences neutrophil influx and activation in atelectasis-prone rabbit lung. J. Appl. Physiol77199413551365
21. Imai Y., Kawano T., Miyasaka K., Takata M., Imai T., Okuyama K.Inflammatory chemical mediators during conventional and high frequency oscillatory ventilation. Am. J. Respir. Crit. Care Med150199415501554
22. Bone, R. C. 1993. The acute respiratory distress syndrome (ARDS). In R. C. Bone, D. R. Dantzker, R. B. George, R. A. Matthay, and H. Y. Reynolds, editors. Pulmonary and Critical Care Medicine. Vol. 3, Part R, Chapter 3. Mosby-Year Book, Inc., St. Louis. 1–9.
23. Lachmann B., Robertson B., Vogel J.In vivo lung lavage as an experimental model of the respiratory distress syndrome. Acta Anaesthesiol. Scand241980231236
24. LeSouef P. N., England S. J., Bryan A. C.Passive respiratory mechanics in newborns and children. Am. Rev. Respir. Dis1291984552556
25. Hu H., Takata M., Kusakawa I., Fujita M., Miyasaka K.Intratracheal administration of phosphodiesterase III inhibitor attenuates bronchoconstriction in cats: a preliminary report. Pediatr. Pulmonol191995360364
26. Chomczynski P., Sacchi N.Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem1621987156159
27. Ito H., Yamamoto S., Kuroda S., Sakamoto H., Kajihara J., Kiyota T., Hayashi H., Kato M., Seko M.Molecular cloning and expression in Escherichia coli of the cDNA coding for rabbit tumor necrosis factor. DNA51986149156
28. Ito H., Shirai T., Yamamoto S., Akira M., Kawahara S., Todd C. W., Wallace R. B.Molecular cloning of the gene encoding rabbit tumor necrosis factor. DNA51986157165
29. Applequist S. E., Keyna U., Calvin M. R., Beck-Engeser G. B., Raman C., Jack H. M.Sequence of the rabbit glyceraldehyde-3-phosphate dehydrogenase-encoding cDNA. Gene1631995325326
30. Abe J., Forrester J., Nakahara T., Lafferty J. A., Kotzin B. L., Leung D. Y.Selective stimulation of human T cells with streptococcal erythrogenic toxins A and B. J. Immunol146199137473750
31. Choi Y. W., Kotzin B., Herron L., Callahan J., Marrack P., Kappler J.Interaction of Staphylococcus aureus toxin “superantigens” with human T cells. Proc. Natl. Acad. Sci. U.S.A.86198989418945
32. Ulich, T. R. 1993. TNF. In J. Kelley, editor. Cytokines of the Lung. Marcel Dekker, New York. 307–332.
33. Xing Z., Kirpalani H., Torry D., Jordana M., Gauldie J.Polymorphonuclear leukocytes as a significant source of tumor necrosis factor-alpha in endotoxin-challenged lung tissue. Am. J. Pathol143199310091015
34. Tracey K. J., Lowry S. F., Cerami A.Cachetin/TNF-alpha in septic shock and septic adult respiratory distress syndrome. Am. Rev. Respir. Dis138198813771379
35. Hyers T. M., Tricomi S. M., Dettenmeier P. A., Fowler A. A.Tumor necrosis factor levels in serum and bronchoalveolar lavage fluid of patients with the adult respiratory distress syndrome. Am. Rev. Respir. Dis1441991268271
36. Suter P. M., Suter S., Girardin E., Roux L. P., Grau G. E., Dayer J. M.High bronchoalveolar levels of tumor necrosis factor and its inhibitors, interleukin-1, interferon, and elastase, in patients with adult respiratory distress syndrome after trauma, shock, or sepsis. Am. Rev. Respir. Dis.145199210161022
37. Tran Van Nhieu J., Misset B., Lebargy F., Carlet J., Bernaudin J. F.Expression of tumor necrosis factor-alpha gene in alveolar macrophages from patients with the adult respiratory distress syndrome. Am. Rev. Respir. Dis.147199315851589
Correspondence and requests for reprints should be addressed to Masao Takata, M.D., Ph.D., Pathophysiology Research, National Children's Medical Research Center, 3-35-31 Taishido, Seyagaya, Tokyo 154, Japan.


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