Pulmonary interstitial pressure and tissue matrix structure in acute hypoxia

Giuseppe Miserocchi1, Alberto Passi2, Daniela Negrini3, Massimo Del Fabbro3, and Giancarlo De Luca2

1 Department of Experimental and Environmental Medicine and Biotechnology, University of Milano-Bicocca, 20052 Monza; 3 Department of Medicine, Surgery and Dentistry, University of Milano, 20142 Milan; and 2 Department of Experimental and Clinical Biomedical Sciences, University of Insubria, 21100 Varese, Italy


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Pulmonary interstitial pressure was measured via micropuncture in anesthetized rabbits in normoxia and after breathing 12% O2. In normoxia [arterial PO2 = 88 ± 2 (SD) mmHg], pulmonary arterial pressure and pulmonary interstitial pressure were 16 ± 8 and -9.6 ± 2 cmH2O, respectively. After 6 h of hypoxia (arterial PO2 = 39 ± 16 mmHg), the corresponding values were 30 ± 8 and 3.5 ± 2.5 cmH2O (P < 0.05). Pulmonary interstitial proteoglycan extractability, evaluated by hexuronate assay after 0.4 M guanidinium hydrochloride extraction, was 12.3, 32.4, and 60.6 µg/g wet tissue in normoxia and after 3 and 6 h of hypoxia, respectively, indicating a weakening of the noncovalent bonds linking proteoglycans to other extracellular matrix components. Gel filtration chromatography showed an increased fragmentation of chondroitin sulfate- and heparan sulfate-proteoglycans during hypoxic exposure, accounting for a loss of extracellular matrix native architecture and basement membrane structure. Gelatin zymography demonstrated increased amounts of the proteolytically activated form of gelatinase B (matrix metalloproteinase-9) after hypoxic exposure, providing evidence that the activation of proteinases may play a role in hypoxia-induced lung injury.

high-altitude pulmonary edema; micropuncture; microvascular permeability; proteoglycans; metalloproteinases


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

RECENT WORK FROM OUR LABORATORY (12, 17, 18, 22, 23) was aimed at describing interstitial fluid dynamics in the lung parenchyma in physiological conditions and in the transition toward the development of hydraulic and lesional edema. These studies were carried out with a minimally invasive technique to measure pulmonary hydraulic interstitial pressure (Ppi) by preserving the integrity of the pleural space (13). This experimental approach implies that lung expansion is maintained in physiological conditions, namely with subatmospheric intrapleural pressure and atmospheric alveolar pressure. At a lung volume corresponding to the functional residual capacity of the respiratory system, Ppi at the heart level averages approximately -10 cmH2O in physiological conditions (13, 14). This value, compatible with a state of relative dehydration of the lung tissue, is maintained by the powerful draining action of the pulmonary lymphatics that, under steady-state conditions, balances the net microvascular fluid filtration (16).

In the transition from physiological conditions toward the development of pulmonary edema, an increase in microvascular filtration caused Ppi to shift from its control subatmospheric value up to ~2-4 cmH2O (12). This considerable Ppi change was accompanied by partial degradation of proteoglycans in the pulmonary interstitial matrix; indeed, at an early stage of development of hydraulic edema, the large chondroitin sulfate (CS) proteoglycan (versican) of the extracellular matrix (ECM) was fragmented (17, 22). Conversely, in the lesional edema model, the heparan sulfate (HS) proteoglycan of the basal membranes (perlecan) was at first affected (18). In both edema models, proteoglycan fragmentation was due to activation of matrix metalloproteinases (MMPs) (22).

In the present investigation, we focused our attention on the effect of hypoxic exposure on lung tissue. Hypoxia is a condition of high clinical relevance because it relates to pulmonary and cardiac diseases and has a profound influence on the pulmonary vasculature. Hypoxia is known to increase pulmonary arterial pressure and to trigger a marked remodeling of the vessel structure (27). High-altitude pulmonary edema is a well-known severe complication of hypoxic exposure (26). The present study was aimed at describing the alterations induced by acute hypoxic exposure on pulmonary and microvascular interstitial fluid dynamics and ECM structure in experimental animals. We studied, in particular, the proteoglycan component of ECM because it is largely responsible for the structural integrity and low compliance of the pulmonary interstitium. Integrity of the lung ECM depends on the balance between synthesis and degradation of its components, and ECM turnover and remodeling involves the activity of MMPs (11). In particular, the MMP subfamily of gelatinases [gelatinase A (MMP-2) and gelatinase B (MMP-9)] was recently found to be involved in the development of pulmonary edema and to recognize the core protein of lung interstitium proteoglycans as a substrate (23). On this basis, we also studied whether increased MMP activities are involved in the lung damage induced by hypoxic exposure.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

General preparation. Experiments were done in rabbits (n = 31; body weight 2.5 ± 0.2 kg) anesthetized with 2.5 ml/kg of a mixture of urethane at 25% in a saline solution and 1.5 ml of pentobarbital sodium (60 mg/ml). This dose abolishes the corneal reflex, is long lasting, and does not substantially depress the respiratory activity. Subsequent doses of anesthetic were given throughout the experiment as judged by the arousal of ocular reflexes. The trachea was cannulated with a T tube, and a saline-filled catheter was placed in the carotid artery. In four rabbits, via a midsternal splitting incision, the chest was opened to expose the pericardium, leaving the pleural sacs intact. A saline-filled catheter was placed in the left atrium; a second catheter was inserted through the right ventricular wall and advanced into the pulmonary artery. Pulmonary arterial and left atrial pressures were measured under control conditions and during hypoxic exposure. Cardiac output was measured with an ultrasonic flowmeter (Transonic Systems model T201) secured around the aortic arch.

In the animals used for micropuncture, a "pleural window" was prepared on the right side of the chest at the level of the sixth or seventh intercostal space by resecting the external and internal intercostal muscles down to the endothoracic fascia. Under stereomicroscopic view and with fine tools, the fascia was stripped over a surface area of ~0.2 cm2, leaving intact the underlying parietal pleura that, being only ~30 µm thick, allows a neat observation of the lung surface. The prepared area was kept covered with Parafilm to avoid tissue dehydration before proceeding to micropuncture.

Experimental protocol. Measurements of Ppi were taken in 1) control animals in normoxia (room air breathing) at time 0 (n = 3) and 2) animals exposed for 3 (n = 11) or 6 (n = 7) h to 12% O2 in nitrogen. The gas mixtures were delivered as a "pass-through system" to the tracheal tube, generating a gas pressure at the tube inlet of ~1 cmH2O. The animals were left to breathe spontaneously up to the time of micropuncture. Blood samples (0.2 ml) were taken every 30 min to determine blood gases (Instrumentation Laboratories model 1620, Milan, Italy). The animals were killed after the micropuncture measurements were completed; the lungs and pieces of the intercostal muscles were removed, and lung tissue samples were taken and processed for biochemical analysis.

Micropuncture technique. For micropuncture, the animal was placed in the lateral decubitus position. Glass micropipettes with tips beveled to 2-3 µm and a taper of ~200 µm were used. The micropipettes were filled with a previously filtered 0.5 M NaCl solution (0.2-µm Millipore filter). Each pipette was calibrated (pressure range ±20 cmH2O) after being seated in a holder and connected to a pressure transducer (Gould P23XL) motor driven by a servo-null system (model 5A, Instrumentation for Physiology and Medicine). Before each measurement, the electrical zero was obtained by insertion of the pipette into a saline pool placed at the height of the pleural window and grounded to the animal. Before micropuncture, the animals were paralyzed with pancuronium bromide (2 mg/kg body wt) and mechanically ventilated through a ventilator delivering the same gas mixture provided during spontaneous ventilation. At the time of micropuncture, the ventilator was stopped, and the animal remained apneic for a period of ~5 min while delivery of the gas mixture (either room air or 12% O2) was continued through an intratracheal tube (outlet pressure approx 1 cmH2O). Micropuncture was performed by advancing the pipette through the pleural space at an angle of 45° relative to the lung surface. This angle allowed us to follow the pipette insertion on a color monitor (TM-150 PSN-K, JVC) connected through a video camera (Panasonic WV-F15E) to a stereomicroscope (Nikon SMZ-2T) set at a magnification of ×60 to ×100. Final magnification on the video screen was twofold higher. We aimed at micropuncturing the arteriolar perimicrovascular interstitial space, which is wide enough to host the micropipette tip. Height of the measurement was approx 60% of lung height relative to the lowermost point of the chest in the lateral decubitus position. Using a similar approach, we also micropunctured the internal intercostal muscles close to the site of pleural window. The animal was put back on the ventilator between successive attempts at micropuncture.

The criteria for accepting micropipette recordings were 1) an unchanged electrical zero on withdrawal of the pipette, 2) a stable recording for at least 1 min, and 3) similar values (within 2 cmH2O) obtained from the same site on successive recording attempts.

Tissue samples. The wet weight-to-dry weight ratio (W/D) of tissue samples from the lung and intercostal muscles was determined by weighing the samples fresh and after drying in an oven at 70°C for 24 h.

Biochemical analysis. Total proteoglycan content of the lung tissue samples was evaluated by hexuronate assay after papain digestion of the tissue (24). After centrifugation of the digested material (15,000 g for 1 h at 4°C), hexuronate-containing glycosaminoglycans (GAGs), the polysaccharide component of proteoglycans, were precipitated with four volumes of ethanol at 4°C and assayed for hexuronate content.

Proteoglycans were extracted from small tissue slices with guanidinium hydrochloride (GuHCl), which breaks intermolecular noncovalent bonds (9), in 50 mM sodium acetate buffer, pH 6.0, containing protease inhibitors (17) at 4°C under mild stirring for 24 h. The denaturing effect and, therefore, the efficiency of the extraction depend on GuHCl concentration: 0.4 M GuHCl is the standard accepted low concentration that permits the formation of proteoglycan aggregates (9). On this basis, an increased proteoglycan recovery after 0.4 M GuHCl extraction of the tissue indicates a decrease in the strength of proteoglycan interactions with other ECM components.

The extracts were dialyzed against 8 M urea in 50 mM sodium acetate buffer, pH 5.8, 0.5% (vol/vol) Triton X-100, and protease inhibitors (buffer A) containing 0.15 M NaCl.

Proteoglycans were purified from a 0.4 M GuHCl extract on a DEAE-Sephacel column (1.6 × 50 cm, flow 12 ml/h), washed with 0.15 M and 0.3 M NaCl in buffer A, and eluted with 1.2 M NaCl in buffer A. Proteoglycans containing peaks were concentrated with a concentrator equipped with a PM 10 membrane. Concentrated samples were precipitated with nine volumes of ethanol at 4°C overnight and analyzed for hexuronate content. Aliquots of purified proteoglycans were analyzed by gel filtration chromatography with a Sepharose CL-4B column (1 × 50 cm, flow 6 ml/h) eluted with 50 mM sodium acetate buffer, pH 6.0, containing protease inhibitors and 4 M GuHCl to prevent nonspecific molecular interactions that may affect proteoglycan size analysis (17). Sepharose CL-4B is adequate for separation of molecular weights ranging from 105 to 108. The elution profiles were obtained by measuring proteoglycan content in the fractions by safranin method (7) and subsequently analyzing the spots with a densitometer (Bio-Rad GS700). The identification of the different proteoglycan populations was performed according to Negrini et al. (18) and Passi et al. (22); the type of GAGs linked to a proteoglycan core protein was determined by their sensitivity or resistance to different specific eliminases (chondroitinase ABC and heparanase plus heparitinase). The unsaturated disaccharides, released by chondroitinase ABC digestion from galactosamine-containing GAGs and by heparanase plus heparitinase digestion from glucosamine-containing GAGs, were fractionated and identified by capillary electrophoresis performed on a Biofocus 3000 apparatus.

Extraction and purification of gelatinases by affinity chromatography. Lung specimens were carefully washed, homogenized at 4°C in 50 mM Tris · HCl buffer, pH 7.5, containing 150 mM NaCl and 0.002% Tween 20 (buffer B) (10, 23), and cleaned by centrifugation at 12,000 g for 10 min. Aliquots of the crude enzyme extracts were incubated with gelatin-Sepharose resin (100 µg protein/100 µl resin) for 1 h at 4°C under gentle mixing to isolate gelatin-degrading enzymes (10). The resin was then recovered, carefully washed with buffer B containing 5 mM CaCl2 and 10 mM Na2-EDTA, resuspended in 4× Laemmli sample buffer, and used for electrozymograms.

SDS-PAGE zymography. SDS-PAGE was performed with a 10% (wt/vol) gel copolymerized with gelatin (1 mg/ml) (10) to identify proteins with gelatinolytic activities. After washing and overnight incubation of the gel at 37°C in an activating buffer (50 mM Tris · HCl buffer, pH 7.5, containing 200 mM NaCl and 5 mM CaCl2), the gel was stained with 0.5% (wt/vol) Coomassie brilliant blue R-250 (10). Proteolytic activities were detected as clear bands against the blue background, indicating areas where the gelatin was degraded by the enzymes. The area of the gelatinolytic bands was evaluated with a GS 700 densitometer (Bio-Rad) and showed a linear increase with increasing amounts of standard MMPs (from 0.2 to 1 µg, regression coefficient = 0.90 ± 0.04). The numerical range of gelatinolytic activity depends on the background of gel zymogram that may vary slightly. The molecular weights of gelatinolytic bands were estimated by comparing their electrophoretic migration to that of protein standards (Bio-Rad). To analyze metalloproteinase inhibition, the gels were incubated in the presence of 20 mM EDTA.

Immunoblot analysis was performed after SDS-PAGE, with Western transfer to polyvinylidene difluoride membrane done with a semidry transfer cell (Bio-Rad) at 15 V/cm2 for 20 min. The identification was then performed with specific polyclonal sheep antibodies (1:5,000), which reacted with both the zymogen latent forms and the proteolytically activated forms of these enzymes. The positive bands were revealed by chemiluminescence (Super Signal Pierce) after incubation with anti-sheep donkey antibodies conjugated with horseradish peroxidase.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Figure 1 shows that pulmonary arterial pressure roughly doubled over 3 h of hypoxic exposure relative to that in normoxia, whereas left atrial pressure increased from a control value of approx 5 to approx 8 cmH2O. Arterial PO2, PCO2, and pH were 88 ± 2 mmHg, 36 ± 4 mmHg and 7.45 ± 0.04, respectively, at the onset of the experiment; after 30 min of hypoxia, the corresponding values were 38 ± 16 mmHg, 26 ± 5 mmHg, and 7.38 ± 0.11, and remained essentially steady up to 6 h of hypoxia.


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Fig. 1.   Time course of pulmonary arterial pressure and left atrial pressure in control normoxia (time 0) and during 12% O2 exposure. Data are means ± SD.

The time course of Ppi and intercostal muscle interstitial pressure (Pmus) is shown in Fig. 2. Ppi significantly increased (P < 0.05) from a control normoxic value of -9.6 ± 2 to 3.5 ± 2.5 cmH2O after 6 h of hypoxic exposure (P < 0.05). Within the same time frame, Pmus also significantly increased from -1.9 ± 1.2 (normoxic value) to approx 0 cmH2O at 3 h (P < 0.05), remaining steady thereafter.


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Fig. 2.   Time course of pulmonary interstitial pressure (Ppi) and intercostal muscle pressure (Pmus) in control normoxic conditions (time 0) and during 12% O2 exposure. Data are means ± SD.

The lung W/D was 4.9 ± 0.1 in normoxia and 4.9 ± 0.2 and 5.0 ± 0.2 (~ 2% increase; not significant) after 3 and 6 h, respectively, of hypoxic exposure. The corresponding W/D values of the intercostal muscles were 3.3 ± 0.2, 3.4 ± 0.3, and 3.8 ± 0.2, the latter value representing a significant 13% increase with respect to the control value.

Biochemical determinations. Figure 3 presents the data of hexuronate recovery from lung samples after proteoglycan extraction with 0.4 M GuHCl and clearly shows that proteoglycan extractability increased with increasing time of hypoxic exposure. After 6 h of hypoxic exposure, proteoglycan extraction approached ~51% of total proteoglycan content of lung tissue (118 ± 4 µg/g wet tissue), indicating a weakening of noncovalent bonds linking proteoglycans to other ECM components.


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Fig. 3.   Hexuronate recovery after 0.4 M guanidinium hydrochloride (GuHCl) extraction of lung tissues in control normoxia and after 3 and 6 h of hypoxia. Line on top, total hexuronate content of tissue samples determined after papain digestion. Data are means ± SD.

Figure 4 shows the elution patterns of gel filtration chromatography of 0.4 M GuHCl lung extracts performed on a Sepharose CL-4B column under dissociative conditions. The various peaks correspond to proteoglycan families of different size and different GAG composition, in agreement with previous experiments carried out under the same experimental conditions (22, 23). In normoxic lungs (Fig. 4, top), large CS proteoglycan (molecular mass > 0.5 MDa) accounted for ~25% of total extracted proteoglycans; HS proteoglycan (0.5-0.1 MDa) for ~35%, and, finally, peptidoglycans (PDGLs; <100,000 Da) for 40%. After hypoxic exposure (3 and 6 h; Fig. 4, middle and lower, respectively), the relative content of CS proteoglycan and HS proteoglycan was markedly lower with respect to normoxia, indicating a fragmentation of both proteoglycan families. The fragmentation products were likely recovered in the PDGL fraction, which indeed increased relative to the other fractions. In Fig. 5A, the data of hexuronate recovery after 0.4 M GuHCl extraction were plotted as a function of time during hypoxic exposure, considering both total proteoglycans and each proteoglycan family. With respect to the normoxic conditions (time 0), the data show a progressively larger recovery of the PDGL family with increasing time. In Fig. 5B, the hexuronate recovery data were plotted as a function of Ppi values, the control value in normoxia corresponding to a Ppi of approximately -10 cmH2O (see data presented in Fig. 2). These relationships indicate a clear functional correlation between increased proteoglycan extractability and increased hydraulic interstitial fluid pressure.


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Fig. 4.   Gel filtration chromatography of proteoglycans isolated from 0.4 M GuHCl extracts in control normoxia (top) and after 3 (middle) and 6 (bottom) h of 12% O2. CS-PG, chondroitin sulfate proteoglycan; HS-PG, heparan sulfate proteoglycan; PDGL, peptidoglycans. Dashed lines, range of molecular masses for the 3 proteoglycan families.



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Fig. 5.   Hexuronate recovery data for total proteoglycan extraction and CS-PG, HS-PG, and PDGL families from lung tissue samples in control normoxia and during hypoxic exposure. A: hexuronate recovery data vs. time of hypoxic exposure (normoxia corresponds to time 0). B: hexuronate recovery as a function of Ppi. Control normoxia corresponds to Ppi of approximately -10 cmH2O.

Gelatin zymography revealed different gelatinolytic bands in the purified enzyme extract from normoxic and hypoxic lungs (Fig. 6). The bands at ~72 (band A) and 92 kDa (band B) corresponded to standards of latent proMMP-2 (progelatinase A) and proMMP-9 (progelatinase B), respectively, and the identification was confirmed by specific antisera. The total area of bands A and B was enlarged in hypoxic compared with normoxic lungs. The minor, faster-migrating band (band B) was recognized by the antibody against MMP-9 and likely represented the proteolytically activated form of the enzyme. This band was clearly more evident in hypoxic samples.


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Fig. 6.   Photodensitometric representation of gelatin zymography performed on purified enzyme extracts from control normoxic and hypoxic lungs (6-h exposure). Transmittance is expressed in arbitrary units; the numerical range of gelatinolytic activity depends on the background of gel zymography. The molecular mass of main gelatinolytic bands was estimated by standard molecular mass markers. A and B, peaks of 72 and 92 kDa, corresponding to matrix prometalloproteinases (proMMP)-2 and proMMP-9, respectively; C and D, multiple matrix metalloproteinase (MMP)-9 forms; b, proteolytically activated form of MMP-9.

The antibody against MMP-9 also reacted with the bands of higher molecular mass (>200 kDa; bands C and D), which probably consisted of multiple MMP-9 forms (23). Also, band C was more evident in hypoxic compared with normoxic lungs.

In animals exposed to 3 h of hypoxia, the gelatin zymography showed a pattern similar to that found after 6 h of exposure.

No gelatinolytic bands were detectable when the gels were incubated in the presence of 20 mM EDTA, a typical inhibitor of MMPs.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The aim of the present investigation was to extend our knowledge of the pathophysiology of pulmonary interstitial fluid dynamics during hypoxic exposure. We took advantage of an experimental approach that allows us to detect early changes in Ppi (as measured by micropuncture in in situ lung) relative to the physiological conditions.

The results of the present study indicate that 6 h of hypoxic exposure in anesthetized animals led to an increase in interstitial fluid content as witnessed by the progressive increase in Ppi (Fig. 2); this, in turn, indicates that a few hours at an arterial PO2 of ~40 mmHg (comparable to an altitude exposure of ~6,000 m) causes an increase in microvascular filtration. One cannot exclude that anesthesia may account, in part, for the observed effects; however, anesthesia per se is a cause of mismatch between ventilation and perfusion and can therefore lead to hypoxia. Conversely, correction of anesthesia-dependent hypoxia through an enriched oxygen mixture may lead to the opposite complication represented by hyperoxia. The increased interstitial volume in hypoxia may reflect 1) loss of the "tissue safety factor," 2) augmented water and solute permeability of the microvascular barrier, 3) increased capillary hydraulic pressure, and 4) increased exchange surface area secondary to capillary recruitment. We shall review these factors in terms of the present results and the available evidence on the effects of hypoxia on microvascular fluid exchange.

The resistance of tissue matrix to an increase in extravascular water can be expressed on mechanical grounds as interstitial compliance, given by the ratio of the change in interstitial volume to the corresponding change in Ppi; in physiological conditions, pulmonary tissue compliance is as low as 0.2 ml · mmHg-1 · 100 g-1, the corresponding value for the muscle being 8.8 ml · mmHg-1 · 100 g-1, ~40-fold higher. Clearly, a low tissue compliance of the pulmonary parenchyma acts as a strong tissue safety factor, preventing the development of edema; in fact, hypoxic exposure did not cause a significant increase in the lung W/D. Conversely, the W/D significantly increased in the intercostal muscles after 6 h of hypoxic exposure. Despite the fact that extravascular lung water was still within normal values after 6 h of hypoxia, the biochemical analysis of the tissue matrix revealed a progressive alteration of the native properties of ECM structure as indicated by the significantly greater extractability of ECM proteoglycans (Fig. 3); this reflected, in turn, the progressive weakening of the noncovalent bonds linking proteoglycans to other ECM macromolecules. Furthermore, gel chromatography data show an increase in the fraction that includes small molecular mass fragments of proteoglycans extracted from the lung tissue, the obvious interpretation being that high molecular mass proteoglycans undergo a degradation process. This process involves both interstitial CS proteoglycan (versican) and HS proteoglycan (perlecan). Versican, a large proteoglycan forming aggregates with hyaluronan, is primarily responsible for tissue hydration and biomechanical properties (25). The gel chromatography profiles (Fig. 4) indicate that the fragmentation of extracted versican increases with hypoxic exposure, yet the nonextracted fraction was still providing a low mechanical tissue compliance as proved by the fact that Ppi was positive. Interestingly, peak Ppi was similar to that observed in other conditions, implying a different pathogenic mechanism leading to edema (17, 18, 22), suggesting that this pressure represents an upper limit of mechanical tolerance of the ECM during interstitial fluid loading. The development of severe edema was, in fact, shown to be primarily related to the massive fragmentation of versican, with complete loss of the tissue safety factor and a decrease in Ppi toward zero (17, 18, 22, 23). A possible cause for an increase in microvascular filtration during hypoxic exposure might involve HS proteoglycan (perlecan) degradation as suggested by the data of Fig. 5B that show that HS proteoglycan degradation positively correlates to the increase in Ppi. Perlecan is present in the endothelial and epithelial basal membranes and, due to the high negatively charged sulfate moieties in its native form, contributes to the selective barrier properties of basal membranes (8). The fragmentation of perlecan could therefore cause an increase in microvascular permeability, accounting for increased microvascular filtration.

Extreme hypoxia down to ~10 mmHg has been reported to cause an increase in microvascular permeability in isolated and perfused dog lungs within 60 min (21). The hypoxia-dependent increase in membrane permeability was observed in other microvascular districts such as myocardial capillaries (1) and in cultured endothelial cells (19). Furthermore, acute hypoxia (inspired O2 fraction = 0.10) was shown to cause an increase in the pulmonary lymph flow rate associated with an unchanged lymph-to-plasma protein concentration ratio, these data being consistent with an increased pulmonary endothelial permeability (4). The increased permeability to albumin was also confirmed in a recent study on humans acutely exposed to an altitude of 4,350 m (6).

Gelatin zymography analysis indicated that the fragmentation process of proteoglycans, accompanied by the loss of their interactive properties with other ECM macromolecules, is likely to depend on the activation of tissue MMPs, in particular MMP-9. MMP-9 is produced by macrophages and alveolar epithelial cells (20) and was recently shown to cleave proteoglycan core protein (23). Our results suggest that an upregulation of tissue MMPs may largely contribute to the damage of pulmonary interstitial proteoglycans during hypoxic exposure, leading to a disturbance of the dynamic equilibrium between degradation and synthesis of proteoglycans. An increase in MMPs was found in rat pulmonary arteries exposed to 10% O2 for up to 10 days (28); furthermore, an increase in the proMMP-2 was demonstrated in the coronary effluent of isolated perfused heart after 20 min of ischemia (2).

From the data presented in Fig. 1 and also reported in the literature (5), hypoxia determines an increase in pulmonary arterial pressure. It remains unclear to what extent the increase in pulmonary vascular pressures is transmitted to the capillary bed. It has been shown that precapillary vascular resistances are increased in hypoxia (5), an important reflex to avoid a rise in capillary pressure and, consequently, in microvascular filtration rate (3). Furthermore, there is evidence that in humans pulmonary wedge pressure remains normal despite an increase in pulmonary arterial pressure (5). However, it has also been proposed that the precapillary vasoconstriction during hypoxia is not homogeneously distributed so that some capillaries are not protected against an increase in hydraulic pressure that, in turn, may lead to a stress failure of the endothelial wall (30). Hypoxic exposure as short as ~15 min has been shown to induce a fourfold increase in the extension of the pulmonary capillary network (29), leading to an increase in exchange surface area. In the transition toward edema formation, capillary recruitment has been invoked as a mechanism buffering an increase in pulmonary capillary pressure (15).

The results from the present investigation concerning the pulmonary interstitial matrix are new and of interest if one considers that the protocol implied a low PO2 but a relatively short exposure time. We think, therefore, that it appears important to highlight that it nevertheless caused damage to the pulmonary interstitium. High-altitude pulmonary edema is characteristically related to a marked hypoxic level and longer exposure time; our data suggest that this form may develop as a consequence of a progressive increase in microvascular permeability due to degradation of basal membranes and to a loss of tissue safety factor due to disruption of ECM architecture.


    ACKNOWLEDGEMENTS

This study was funded by the Italian Ministry of University and Scientific and Technological Research (MURST).


    FOOTNOTES

Address for reprint requests and other correspondence: G. Miserocchi, Istituto di Fisiologia Umana, Via Mangiagalli 32, 20133 Milano, Italy (E-mail: giuseppe.miserocchi{at}unimib.it).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 21 June 2000; accepted in final form 15 December 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Al-Haboubi, H, Tomlinson D, and Ward B. The influence of hypoxia on transvascular leakage in the isolated rat heart: quantitative and ultrastructural studies. J Physiol (Lond) 482: 157-166, 1995[Abstract].

2.   Cheung, PY, Sawicki G, Wozniak M, Wang W, Radomski M, and Schulz R. Matrix metalloproteinase-2 contributes to ischemia-reperfusion injury in the heart. Circulation 101: 1833-1839, 2000[Abstract/Free Full Text].

3.   Glazier, J, and Murray J. Sites of pulmonary vasomotor reactivity in the dog during alveolar hypoxia and histamine infusion. J Clin Invest 50: 2550-2558, 1971[ISI][Medline].

4.   Grimbert, FA, Martin D, Parker JC, and Taylor AE. Lymph flow during increases in pulmonary blood flow and microvascular pressure in dogs. Am J Physiol Heart Circ Physiol 255: H1149-H1155, 1988[Abstract/Free Full Text].

5.   Groves, B, Reeves J, Sutton J, Wagner P, Cymerman A, Malconian M, Rock P, Joung P, and Huston C. Operation Everest II: elevated high-altitude pulmonary resistance unresponsive to oxygen. J Appl Physiol 63: 521-530, 1987[Abstract/Free Full Text].

6.   Hansen, J, Olsen N, Feldt-Rasmussen B, Kanstrup L, Dechaux M, Dubray C, and Richalet J. Albuminuria and overall capillary permeability of albumin in acute altitude hypoxia. J Appl Physiol 76: 1922-1927, 1994[Abstract/Free Full Text].

7.   Hascall, V, Calabro A, Midura R, and Yanagishita M. Isolation and characterization of proteoglycans. Methods Enzymol 230: 390-417, 1994[ISI][Medline].

8.   Hascall, V, and Hascall G. Proteoglycans. In: Cell Biology of Extracellular Matrix, edited by Hay ED.. New York: Plenum, 1981, p. 39-63.

9.   Heinegard, D, and Sommarin Y. Isolation and characterization of proteoglycans. Methods Enzymol 144: 319-371, 1987[ISI][Medline].

10.   Mazzieri, RL, Masiero L, Zanetta L, Monea S, Onisto M, Garbisa S, and Mignatti P. Control of type IV collagenase activity by components of the urokinase-plasmin system: a regulatory mechanism with cell-bound reactants. EMBO J 16: 2319-2332, 1997[Abstract/Free Full Text].

11.   McElvaney, NG, and Crystal RG. Proteases and lung injury. In: The Lung: Scientific Foundations, edited by Crystal RG, West JB, and Barnes PJ.. Philadelphia, PA: Lippincott-Raven, 1997, vol. 2, p. 2205-2218.

12.   Miserocchi, G, Negrini D, Del Fabbro M, and Venturoli D. Pulmonary interstitial pressure in intact in situ lung: the transition to interstitial edema. J Appl Physiol 74: 1171-1177, 1993[Abstract].

13.   Miserocchi, G, Negrini D, and Gonano C. Direct measurements of interstitial pulmonary pressure in in situ lung with intact pleural space. J Appl Physiol 69: 2168-2174, 1990[Abstract/Free Full Text].

14.   Miserocchi, G, Negrini D, and Gonano C. Parenchymal stress affects interstitial and pleural pressures in in situ lung. J Appl Physiol 71: 1967-1972, 1991[Abstract/Free Full Text].

15.   Negrini, D. Pulmonary microvascular pressure profile during development of hydrostatic edema. Microcirculation 2: 1-8, 1995[Medline].

16.   Negrini, D, Gonano C, and Miserocchi G. Microvascular pressure profile in intact in situ lung. J Appl Physiol 72: 332-339, 1992[Abstract/Free Full Text].

17.   Negrini, D, Passi A, De Luca G, and Miserocchi G. Pulmonary interstitial pressure and proteoglycans during development of pulmonary edema. Am J Physiol Heart Circ Physiol 270: H2000-H2007, 1996[Abstract/Free Full Text].

18.   Negrini, D, Passi A, De Luca G, and Miserocchi G. Proteoglycan involvement during development of lesional pulmonary edema. Am J Physiol Lung Cell Mol Physiol 274: L203-L211, 1998[Abstract/Free Full Text].

19.   Ogawa, S, Koga S, Kuwabara K, Brett J, Morrow B, Morris S, Billezikian J, Silverstein S, and Stern D. Hypoxia-induced increased permeability of endothelial monolayers occurs through lowering of cellular cAMP levels. Am J Physiol Cell Physiol 262: C546-C554, 1992[Abstract/Free Full Text].

20.   Pardo, A, and Selman M. Matrix metalloproteinases and lung injury. Braz J Med Biol Res 29: 1109-1115, 1996[ISI][Medline].

21.   Parker, R, Granger D, and Taylor AE. Estimates of isogravimetric capillary pressures during alveolar hypoxia. Am J Physiol Heart Circ Physiol 241: H732-H739, 1981[ISI][Medline].

22.   Passi, A, Negrini D, Albertini R, De Luca G, and Miserocchi G. Involvement of lung interstitial proteoglycans in development of hydraulic- and elastase-induced edema. Am J Physiol Lung Cell Mol Physiol 275: L631-L635, 1998[Abstract/Free Full Text].

23.   Passi, A, Negrini D, Albertini R, Miserocchi G, and De Luca G. The sensitivity of versican from rabbit lung to gelatinase A (MMP-2) and B (MMP-9) and its involvement in the development of hydraulic lung edema. FEBS Lett 456: 93-96, 1999[ISI][Medline].

24.   Radhakrishnamurthy, B, Jeansonne N, Smart F, and Berenson G. Proteoglycans from lungs treated with pronase and cadmium chloride. Am Rev Respir Dis 131: 855-861, 1985[ISI][Medline].

25.   Roberts, CR, Weight TN, and Hascall VC. Proteoglycans. In: The Lung: Scientific Foundations, edited by Crystal RG, West JB, and Barnes PJ.. Philadelphia, PA: Lippincott-Raven, 1997, vol. 1, p. 757-767.

26.   Schoene, R, Hackett P, Henderson W, Sage E, Chow M, Roach R, Mills W, and Martin T. High-altitude pulmonary edema. Characteristics of lung lavage fluid. JAMA 256: 63-69, 1986[Abstract].

27.   Stenmark, K, Durmowicz A, and Demsey E. Modulation of vascular wall cell phenotype in pulmonary hypertension. In: Pulmonary Vascular Remodelling, edited by Bishop J, Reeves J, and Laurents G.. London: Portland, 1995, p. 171-212.

28.   Thakken-Varia, S, Tozzi CA, Poiani GJ, Babiarz JP, Tatem L, Wilson FJ, and Riley DJ. Expression of matrix-degrading enzymes in pulmonary vascular remodeling in the rat. Am J Physiol Lung Cell Mol Physiol 275: L398-L406, 1998[Abstract/Free Full Text].

29.   Wagner, W, Latham L, and Kapen R. Capillary recruitment during airway hypoxia: role of pulmonary artery pressure. J Appl Physiol 47: 383-387, 1979[Abstract/Free Full Text].

30.   West, J, Tsukimoto K, Mathieu-Costello O, and Prediletto R. Stress failure in pulmonary capillaries. J Appl Physiol 70: 1731-1742, 1991[Abstract/Free Full Text].


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