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Pain Medicine  |   December 2001
Effects of Vaporized Perfluorocarbon on Pulmonary Blood Flow and Ventilation/Perfusion Distribution in a Model of Acute Respiratory Distress Syndrome
Author Affiliations & Notes
  • Matthias Hübler, M.D.
    *
  • Jennifer E. Souders, M.D.
  • Erin D. Shade
  • Nayak L. Polissar, Ph.D.
    §
  • Carmel Schimmel
  • Michael P. Hlastala, Ph.D.
  • * Staff Anesthesiologist, Department of Anesthesiology and Intensive Care Medicine, Technical University Dresden, Germany. † Assistant Professor, Department of Anesthesiology, ‡ Research Technician, Department of Medicine, ∥ Professor of Physiology and Biophysics and of Medicine, Departments of Physiology and Medicine, University of Washington School of Medicine, Seattle, Washington. § Statistical Consultant, Mountain-Whisper-Light Statistical Consulting, Seattle, Washington.
  • Received from the Department of Anesthesiology and Intensive Care Medicine, Technical University Dresden, Germany, and the Departments of Anesthesiology and Medicine, University of Washington School of Medicine, Seattle, Washington.
Article Information
Pain Medicine
Pain Medicine   |   December 2001
Effects of Vaporized Perfluorocarbon on Pulmonary Blood Flow and Ventilation/Perfusion Distribution in a Model of Acute Respiratory Distress Syndrome
Anesthesiology 12 2001, Vol.95, 1414-1421. doi:
Anesthesiology 12 2001, Vol.95, 1414-1421. doi:
A NUMBER of studies have demonstrated the ability of total liquid ventilation and partial liquid ventilation (PLV) with perfluorocarbon (PFC) liquids to improve gas exchange and pulmonary function in various animal models of acute respiratory failure. 1–4 These findings were confirmed in preliminary clinical trials. 5–7 Several, not mutually exclusive mechanisms of action of PFC are likely to be responsible for the observed improvements 8–11 : the high oxygen carrying capacity of PFC liquids, effects on surface tension at the air–liquid and liquid–tissue interfaces, antiinflammatory effects, and effects on pulmonary ventilation (V̇A) and perfusion (Q̇) distributions.
Vaporization is a new delivery technique for PFC, recently reported by Bleyl et al.  12 The authors found distinct positive effects on the arterial oxygen partial pressure (Pao2), lung compliance, and physiological shunt in an ovine model of adult respiratory distress syndrome (ARDS). Vaporization was achieved using perfluorohexane (PFX) in an anesthesia machine with modified bypass vaporizers. One advantage of this application technique compared with PLV is that the concentration of PFC delivered can be easily adjusted during administration without disconnecting the ventilator.
The aims of this study were twofold. First, we hoped to replicate the positive results obtained by Bleyl et al.  12 by vaporizing PFX in a sheep model of oleic acid (OA)–induced lung injury. Second, we hoped to shed light on the proposed mechanisms of the effects of vaporized PFC. Specifically, we hypothesized that vaporized PFX redistributes pulmonary blood flow to better ventilated lung regions, thus improving the matching of ventilation to perfusion.
Materials and Methods
The study was approved by the University of Washington Animal Care Committee (Seattle, Washington), and National Institutes of Health (Bethesda, Maryland) guidelines for animal use and care were followed throughout.
Animal Preparation and Experimental Protocol
The experiments were performed on nine adult sheep, ranging in weight from 18.2 to 33.6 kg. The animals were premedicated with 0.6 mg/kg xylazine (Phoenix Pharmaceutical, Inc., St. Joseph, MO), anesthetized with 20 mg/kg thiopental sodium (Abbott Laboratories, North Chicago, IL), intubated, and ventilated with a Servo 900 C ventilator (Siemens, Solna, Sweden). Volume-controlled intermittent positive pressure ventilation was initiated with a tidal volume of 12 ml/kg, a respiratory frequency of 26 breaths/min, a positive end-expiratory pressure (PEEP) of 5 cm H2O, an inspiratory oxygen concentration (Fio2) of 1.0, and an inspiratory/expiratory ratio of 1:1. Anesthesia was maintained using a continuous infusion of thiopental sodium (mean infusion rate between 24.9 ± 3.1 and 59.9 ± 8.1 mg · kg1· h1). A cuffed endotracheal tube (Rüsch; Waiblingen, Germany) was inserted via  a tracheotomy. A catheter was placed in a femoral artery to monitor systemic arterial pressure and to draw blood gas and inert gas samples. A No. 7 French Swan-Ganz thermodilution catheter (Baxter, Irvine, CA) was advanced into the pulmonary artery via  the right external jugular vein to measure pulmonary arterial pressure, pulmonary capillary wedge pressure, and temperature and for blood sampling. A femoral venous catheter was inserted for infusion of anesthetic drugs and maintenance fluids. A central venous catheter was inserted through the left external jugular vein and used as the injection port for OA and microspheres. After the end of instrumentation, the animals were turned to prone position. The animals were paralyzed thereafter with pancuronium (0.1 mg/kg) (Ohmeda PPD Inc., Liberty Corner, NJ), which was administered in regular intervals throughout the experiment. Airway pressures, arterial pressure, and pulmonary arterial pressure were measured continuously with amplifiers (Validyne, Northridge, CA) and recorded on a Mark12 data management system (Model DMS 1000; Western Graphtec, Irvine, CA). The end-tidal carbon dioxide partial pressure (PETco2) was measured with a mass spectrometer (medical gas analyzer Model MGA-1100; Perkin-Elmer, Pomona, CA). Cardiac outputs (thermodilution technique) and blood temperatures were measured with a cardiac output computer (Model Sat-2; Baxter Edwards Irvine, CA). Arterial and venous pH, oxygen partial pressures, and carbon dioxide partial pressures were measured with a blood gas analyzer (Radiometer Model ABL 330; Acid Base Laboratory, Copenhagen, Denmark) and corrected for temperature. The partial pressure of alveolar oxygen (PAo2) was derived fromMATHwhere PBis the barometric pressure. Physiologic oxygen-shunt fraction (Q̇sp/Q̇t) was calculated using the Berggren shunt equation,MATHwhere Q̇spis the physiologic shunt, Q̇tis the total cardiac output, Cc′o2is the capillary oxygen content (assuming equilibration with PAo2), CaO2is the arterial oxygen content, and Cv̄o2is the mixed venous oxygen content. The oxygen contents were estimated from the subroutines of Olszowka and Farhi. 13 Static lung-chest compliance (CRS) was calculated usingMATHwhere Vtis the tidal volume and Pplatis the end-inspiratory plateau pressure during pressure hold.
After an initial stabilization period during which the respiratory minute volume was adjusted to maintain Paco2between 36 and 46 mmHg, baseline measurements (tbase) were recorded. Lung injury was induced by injecting 0.1 ml/kg OA (C17H33COOH; Sigma Chemical Company, St. Louis, MO). During the injection of OA, 500 ml hydroxyethyl starch, 6%, was given to maintain blood pressure. Severe lung injury was considered established (tinj) when the clinical criteria for the definition from the American/European consensus conference on ARDS were fulfilled (ratio Pao2to Fio2< 200, pulmonary capillary wedge pressure < 19 mmHg). 14 If Pao2stabilized before reaching the criteria for ARDS, an additional 0.02 ml/kg OA was given. The mean amount of injected OA was 0.12 ± 0.02 (SD) ml/kg per animal with no differences between the groups (P  = 0.6). The lungs of animals in the treatment group (n = 4) were then ventilated for 30 min with approximately 20 vol% vaporized PFX (ABCR, Karlsruhe, Germany), whereas the ventilation protocol of the control animals (n = 5) was not changed. All animals were studied for 120 min after the injury was established.
Vaporized Perfluorocarbon
All experiments were performed using PFX with a purity of 95%. Its chemical and physical properties and similarity to volatile anesthetics are described elsewhere. 12 PFX was vaporized using a modified vaporizer for Servo ventilators (Isoflurane Vaporizer Model 952; Siemens-Elema, Solna, Sweden). The concentration of the vaporized PFX in volume percent was monitored continuously using an Amis Model 2000 mass spectrometer system (Innovision A/S, Odense, Denmark) with a sampling time and frequency of 8.33 ms and 40 kHz, respectively. The Amis Model 2000 system was calibrated for an atomic mass peak of 69.13 and 25 vol% using the vapor in the head space of a sealed PFX bottle at room temperature (22.5°C). This PFX vapor concentration was calculated with the Clausius-Clopeyron equation P = A · e−B·T, where P is vapor pressure in millimeters of mercury, T is temperature in Kelvin, and A and B are molecule-specific constants. A and B were derived using the vapor pressure (177 mmHg at 20°C) and boiling point (57°C) of PFX.
Inert Gas Measurements
The multiple inert gas elimination technique was used to assess gas exchange 15,16 at five time points during the experiments: tbase, tinj, and 30, 60, and 120 min after injury (t30, t60, and t120, respectively). A dilute solution of six inert gases (sulfur hexafluoride, ethane, cyclopropane, halothane, diethyl ether, and acetone) dissolved in dextrose, 5%, was infused and allowed to equilibrate in the animal for at least 30 min before baseline samples were drawn. Inert gas partial pressures were measured in arterial (Pa) and mixed venous (Pv̄) blood and in mixed expired gas (PĒ). The mixed expired gas samples were collected from a heated Plexiglas mixing chamber that was fitted to the expiratory limb of the ventilator. All samples were collected simultaneously and in duplicate. Exhaled gas specimens were maintained at more than 40°C before analysis to avoid condensation and loss of highly soluble gases. The concentrations of inert gases in the gas samples were measured using a gas chromatograph (Varian, Walnut Creek, CA) equipped with a flame ionization detector and an electron capture detector. Because the high PFX concentration in the gas samples at t30influenced the measured concentrations of the inert gases, PĒ at this time point was calculated from Pa, Pv̄, Q̇t, and V̇E, where V̇Eis the minute ventilation. The double extraction method of Wagner et al.  17 was used to determine the concentration of the inert gases in the blood samples.
Gas exchange was assessed by changes in V̇Aand Q̇ distributions predicted by the 50-compartment model and by dispersion indices and arterial-alveolar difference ([a − A]D) areas derived from retention (R) and excretion (E) data. Inert gas shunt (Q̇S/Q̇T), inert gas dead space (V̇D/V̇T), and percentage of V̇Aand Q̇ to regions with a different V̇A/Q̇ ratio were calculated from the 50-compartment model. 15,16 Because OA lung injury primarily results in increases in shunt, and low V̇A/Q̇ units are infrequently observed, the percentage of Q̇ to shunt and low V̇A/Q̇ units was grouped together to indicate Q̇ to injured lung. 18 The 50-compartment distributions of V̇Aand Q̇ were divided into five regions for further analysis: (1) inert gas shunt (V̇A/Q̇ < 0.01), (2) low V̇A/Q̇ regions (V̇A/Q̇= 0.01–1), (3) midrange V̇A/Q̇ regions (V̇A/Q̇= 1–10), (4) high V̇A/Q̇ regions (V̇A/Q̇= 10–100), and (5) inert gas dead space (V̇A/Q̇ > 100). The R and E components of the inert gas ([a − A]D) area 19 and the parameters of dispersion (DISP) adapted from Gale et al.  20 (DISPR*− E*, DISPR*, and DISPE*) were calculated from direct analysis of R and E data. 21 The DISP parameters, corrected for shunt and dead space, and [a − A]D areas are derived from the difference between heterogeneous and homogeneous R and E curves. DISPR*and R[a − A]D are parameters of Q̇ distribution, and DISPE*and E[a-A]D are parameters of V̇Adistribution. Increases in any of the above parameters are indicative of increased V̇A/Q̇ heterogeneity.
Determination of Pulmonary Blood Flow
Details of the methods are described elsewhere. 22 Five fluorescent polystyrene microspheres (red, orange, blue green, yellow green, and crimson) of 15-μm diameter (Molecular Probes, Eugene, OR) were used in a randomized order to measure regional pulmonary blood flow. The time points of injection were tbase, tinj, t30, t60, and t120. The microspheres (1.5 × 106) were vortexed, then sonicated for 90 s and injected over a period of 30 s, followed by a saline flush.
After completion of the study, the animals were given 1,000 U/kg heparin (Elkins-Sinn, Inc., Cherry Hill, NJ) and 3 mg/kg papaverine hydrochloride (American Regent Laboratories, Inc., Shirley, NY), exsanguinated, and sternotomized. The lungs were flushed via  the main pulmonary artery with 50 ml/kg dextran, 2% (Sigma), removed, inflated to 30 cm H2O, punctured, and dried with warm air for 7 days. When dry, the lungs were coated with a one-component polyurethane foam (DAP Inc., Dayton, OH), suspended vertically in a square box, and embedded in rapidly setting urethane foam (2 pounds, Polyol and Isocyanate; International Sales, Seattle, WA). The foam block was cut into slices approximately 1.2 cm thick. With the use of a 12-mm-diameter core, the slices were sampled. Cores were obtained in a rigid X-Y grid system, with 2 cm between the centers of adjacent cores. The height of every core was measured using a caliper, and the volume was calculated. Two hundred one ± 47 (SD) samples were obtained from each sheep lung after discarding samples with airways occupying more than 25% of the core’s volume. The average volume of the samples was 1.35 ± 0.23 cm3(SD). The samples were soaked for 7 days in 3 ml 2-ethoxyethyl acetate (Aldrich Chemical Co., Milwaukee, WI). The fluorescence was read in a luminescence spectrophotometer (Model LS-50B; Perkin-Elmer, Beaconsfield, Buckinghamshire, UK) fitted with a flow cell and a red-sensitive photomultiplier tube. The volume-normalized relative blood flow (Q̇rel,i) of the piece i at every time point was calculated:MATHwhere xiis the obtained fluorescence divided by the volume of the piece (in cubic centimeters), and n is the number of pieces of the lung. Therefore, the mean normalized relative flow was 1.0.
Statistics
The values are reported as means ± SD. We used the unpaired t  test to compare physiological parameters between the control and treatment groups at tbase(preinjury) and at tinjbefore treatment. We used repeated measures analyses of covariance (RANCOVA) to determine whether values of a given variable differed between the treatment and control groups. The measurements taken after the treatment had begun (and after injury) were the repeated values of the dependent variable in the RANCOVA. For respiratory and cardiovascular variables, we used observations at 10, 20, and 30 min after tinjand every 15 min from 45 to 120 min after tinjas dependent variables in the RANCOVA analysis. For inert gas variables and pulmonary blood flow data, we used the observations at 30, 60, and 120 min after tinjas the dependent variables. For economy of space, table 1presents means only at selected times. For each response, the measurement taken after injury and before treatment had begun was used as a covariate (independent variable) in the RANCOVA. The use of the injury value as a covariate has the effect of comparing the groups on changes after injury, with injury considered as the starting point for each animal. Because the several animals start off after injury with varying values of the response variable, the treatment effect is best measured while controlling for this starting value. Without the covariate, treatment differences might simply be due to mean differences in the starting (injury) value of the response variable. The RANCOVA tests three null hypotheses of interest (H01, H02, H03) as follows. (1) The mean of the combined groups is constant after injury (H01). (2) The time patterns are parallel between groups (i.e.  , the mean difference between the two groups is constant over all time points after injury) (H02). In the RANCOVA model this hypothesis is tested using an interaction term between time and treatment group. (3) The two groups have the same overall mean, where the mean is taken across all observation times after injury, once treatment has been initiated (H03). In the RANCOVA model, this is the main effect of the treatment term. Only hypotheses H02 and H03 involve treatment effects and are reported. Throughout the present study, P  < 0.05 is used to designate statistical significance.
Results
The criteria for ARDS were attained after a mean time of 88 ± 41 (SD) min after the injection of OA with no differences between the groups (P  = 0.2).
Respiratory and Hemodynamic Data
The analysis of the hemodynamic parameters heart rate, cardiac output, systemic arterial pressure, and pulmonary arterial pressure did not yield any statistically significant differences between the groups at the time points tbase, tinj, and thereafter (data not shown). The respiratory data and the oxygen-shunt values are presented in table 1. The Pao2diverged significantly over time when comparing the two groups resulting in a significantly higher overall Pao2in the PFX group. The CR/Sdecreased in both groups over time (not significant). The time pattern in oxygen-shunt did not differ significantly between the groups, but the overall mean values were significantly lower in the PFX group when compared with control animals.
Table 1. Respiratory Data
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Table 1. Respiratory Data
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Inert Gas Exchange
In one animal of the control group, no inert gas data could be obtained because of a malfunction of the gas chromatograph. The V̇Aand Q̇ distributions to areas with a different V̇A/Q̇ ratio are shown in table 2. Analysis of the V̇Adistribution showed that the V̇Aincreased significantly in areas with a high V̇A/Q̇ ratio in the PFX group when compared with the control group. Treatment with PFX vapor also shifted the V̇Adistribution toward areas with a low V̇A/Q̇ ratio, although the comparison of the overall means of the two groups did not yield a statistically significant difference. The changes in inert gas shunt were comparable with the values obtained from the oxygen-shunt calculation.
Table 2. Inert Gas Exchange (I)
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Table 2. Inert Gas Exchange (I)
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The dispersion indices and the [a − A]D areas derived from the R and E data are shown in table 3. Statistically, the values did not differ significantly between the groups at tbaseand at tinj, but were consistently somewhat higher in the PFX group than in the control group. All six parameters of heterogeneity continued to increase after tinjas is typically seen in pulmonary edema. RANCOVA analysis of the dispersion indices and the [a − A]D areas (from 10 min after tinjto t120) yielded significant higher overall means in the PFX group. The overall increase in heterogeneity in the PFX group was probably due to both the increase in V̇Aheterogeneity and the increase in Q̇ heterogeneity.
Table 3. Inert Gas Exchange (II)
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Table 3. Inert Gas Exchange (II)
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Pulmonary Blood Flow
The injection of OA changed significantly the pattern of Q̇relat tinjcompared with tbase. Q̇relwas redistributed from pieces that were high-flow at tbaseto pieces that were low-flow at tbase(fig. 1). The mean slope of the change in Q̇rel(Q̇relat tinjminus Q̇relat tbasevs.  Q̇relat tbase) was −0.50 ± 0.29 and significantly different from 0 (P  < 0.001; one-sample, two-tailed t  test). This slope was approximately the same in both groups (P  = 0.2). These changes in Q̇relare also substantially larger than random variation observed over time. 23 
Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
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Spatial analysis of the Q̇reldistributions after tinjshowed differences between the groups: In the control group, the mean slope of Q̇relversus  the dorsal-to-ventral axis became more negative, showing that Q̇relwas redistributed from dependent, ventral areas to nondependent, dorsal areas (mean slope at tinj:−0.031 ± 0.046; mean slope at t30:−0.06 ± 0.03; mean slope at t60:−0.081 ± 0.066; and mean slope at t120:−0.10 ± 0.057). In contrast, the mean slope of Q̇relversus  the dorsal-to-ventral axis remained almost unchanged in the PFX group (mean slope at tinj:−0.024 ± 0.079; mean slope at t30:−0.019 ± 0.068; mean slope at t60:−0.033 ± 0.071; and mean slope at t120:−0.012 ± 0.073). Statistical analysis yielded no differences in the overall means but a significantly different time pattern when comparing the two groups (P  = 0.048). Figure 2shows plots of Q̇relversus  the dorsal-to-ventral axis in one representative animal from each group.
Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus  the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus 
	the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus  the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
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Discussion
The important findings of our study are that after OA injury, treatment with PFX vapor leads to a higher overall V̇A/Q̇ heterogeneity in comparison with control animals. The injury caused by OA redistributed Q̇relfrom high-flow to low-flow pieces when compared with baseline values. The pattern of this redistribution, as measured with the microsphere method, was similar for both treatment and control groups. Short treatment with PFX vapor modified the further redistribution of Q̇rel.
Our microsphere data showed that Q̇relshifted from areas that were initially high-flow to areas that were initially low-flow after inducing the injury with OA. It may be reasonable to suppose that more OA was delivered to lung regions with a high initial Q̇rel. However, whether this correlates with a greater degree of injury to these lung areas is not known. The spatial analysis of the Q̇reldistribution after tinjshowed that, in the control group, Q̇relwas redistributed over time from dependent to nondependent lung areas, whereas the spatial Q̇reldistribution remained unchanged in the PFX group. Q̇relwas not redistributed from dependent to nondependent lung areas as described for PLV. The inspiratory PFX fraction was 0.2, which yields according to Dalton’s law to a partial pressure of PFX of approximately 142 mmHg inside the alveoli. Solving the Clausius-Clapeyron equation (P = A · e−B·T) for the animals’ body temperature yields a vapor pressure of 390 mmHg. Because of the difference in these pressures, we think that a thin film of PFX covers the bronchial tree but that a net accumulation of PFX with bulk deposition of condensed PFX does not occur. PLV redistributes V̇Afrom dependent to nondependent lung areas in OA-induced lung injury. 10 PLV in noninjured lungs changes the pulmonary blood flow distribution in the same direction. 24 Therefore, the better matching of V̇Aand Q̇ in the nondependent lung areas might be one reason, beside others, for the improved gas exchange during PLV. In our experiments, we did not measure the spatial V̇Adistribution but found an increase in V̇Aand Q̇ heterogeneity. However, at the same time, we observed improved oxygenation and a decrease in shunt. The later observation suggests that the overall matching of V̇Aand Q̇ improved after treatment with PFX vapor. These seemingly contradictory statements can be explained if, despite an increase in V̇Aand Q̇ heterogeneity, the regional V̇Aand Q̇ was better matched at each V̇A/Q̇ ratio. There may be a much wider range of V̇A/Q̇ ratios in the lung (increased heterogeneity), but the V̇Aand Q̇ plots might be almost identical (increased matching). Decreased alveolar V̇A(attributable either to atelectasis or decreased ventilation) leads to alveolar hypoxia and subsequent hypoxic pulmonary vasoconstriction. Probably, the PFX vapor prevents a further collapse of alveoli and V̇Adoes not decrease further. This is consistent with the proposed effect of PFX vapor on the surface tension at the air–liquid interface in the alveoli as one mechanism of action. 12 Alveolar expansion is facilitated and, consecutively, the hypoxic pulmonary vasoconstriction is either attenuated or even released, which leads to a better gas exchange. According to our data, this effect appears to be independent from gravitational factors.
In the first report about the use of PFX vapor, vaporization was achieved using an anesthesia machine with modified bypass vaporizers. 12 In our experiments, we used a Servo 900 C ventilator equipped with a modified Isoflurane vaporizer Model 952 and measured the PFX vapor concentration with a mass spectrometer. The aimed inspiratory PFX concentration was reached within 2 min after opening the vaporizer. When the vaporizers were closed at the end of the treatment period, the mass spectrometer was only able to detect PFX for approximately 3 min. One advantage of this type of setup (compared with an anesthesia machine) is that it is easy to use in an intensive care unit. However, it is known that PFC vapor may affect pneumotachometers and that the measured tidal volumes are falsely high. 25 Therefore, it is mandatory that the ventilator is adjusted appropriately to deliver the correct tidal volume to the patient or the animal, and to get the correct numbers for the calculation of the compliance. In our experimental setting, we had to increase the preset tidal volume of approximately 20% during vaporization to correct for this error.
Our study clarifies some issues of vaporized PFX, but the mechanism of action still remains poorly understood. It is unlikely that the oxygen solubility in PFX is a major mechanism of action because molecules in the gaseous state of matter diffuse freely. It was shown that PLV attenuates lung injury when compared with conventional mechanical ventilation in endotoxin, 11 as well as in OA-induced models of ARDS. 10 Previous studies have suggested that PFC may have antiinflammatory properties 26,27 and that alveolar macrophage function may be impaired after exposure to PFC. 28 In a recent in vitro  study it was shown that PFX reduces the expression and the release of proinflammatory and procoagulant mediators, 29 which is further evidence of direct antiinflammatory properties of PFCs on the cellular level. Therefore, it is possible that PFX vapor might have an effect in a less simple mechanistic manner than it appears.
In summary, we have shown in an animal model of ARDS that short-term treatment with PFX vapor increased V̇A/Q̇ heterogeneity. This increase was independent from changes in shunt and in dead space as shown by the dispersion indices. The relative blood flow distribution as measured by the microsphere method did not yield a gravitational effect of the vapor. Our results are consistent with an effect of PFX vapor on the surface tension, thus preventing further alveolar collapse and improving gas exchange over time.
The authors thank Wayne Lamm, M.A., Research Scientitst, Ian Starr, Research Technician, Susan Bernard, D.V.M., Research Scientist, and Dowon An, Research Technician, Department of Medicine, University of Washington School of Medicine, Seattle, Washington, for expert technical help.
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Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
Fig. 1. Oleic acid–induced change in Q̇reldistribution. Plot shows pooled lung pieces of all animals. Q̇reldecreased in pieces with initial high flows and increased in pieces with initial low flows, suggesting that high-flow pieces were more injured. Q̇rel(tbase) = relative pulmonary blood flow at baseline; Q̇rel(tinj) = relative pulmonary blood flow at injury.
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Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus  the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus 
	the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
Fig. 2. Relative pulmonary blood flow (Q̇rel) distribution versus  the dorsal-to-ventral axis in one representative animal from each group. The numbers of pieces per lung were 236 for the control animal and 233 for the treatment animal, respectively. The slopes remained unchanged in treatment animals, whereas the slopes became more negative over time in control animals showing that Q̇relwas redistributed from dependent (ventral) to nondependent (dorsal) lung areas. CTL = control group; PFX = treatment group; tinj= established injury; t30= 30 min after established injury; t60= 60 min after established injury; t120= 120 min after established injury.
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Table 1. Respiratory Data
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Table 1. Respiratory Data
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Table 2. Inert Gas Exchange (I)
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Table 2. Inert Gas Exchange (I)
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Table 3. Inert Gas Exchange (II)
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Table 3. Inert Gas Exchange (II)
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