1 Department of Medicine, Surgery and Dentistry, University of Milano, 20133 Milan; 2 Department of Experimental and Clinical Biomedical Science and 3 Department of Clinical and Biological Sciences, University of Insubria, 21100 Varese, Italy; and 4 Department of Physiology, University of Bergen, N-5000 Bergen, Norway
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ABSTRACT |
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Interstitial fluid protein concentration (Cprotein) values in perivascular and peribronchial lung tissues were never simultaneously measured in mammals; in this study, perivascular and peribronchial interstitial fluids were collected from rabbits under control conditions and rabbits with hydraulic edema or lesional edema. Postmortem dry wicks were implanted in the perivascular and peribronchial tissues; after 20 min, the wicks were withdrawn and the interstitial fluid was collected to measure Cprotein and colloid osmotic pressure. Plasma, perivascular, and peribronchial Cprotein values averaged 6.4 ± 0.7 (SD), 3.7 ± 0.5, and 2.4 ± 0.7 g/dl, respectively, in control rabbits; 4.8 ± 0.7, 2.5 ± 0.6, and 2.4 ± 0.4 g/dl, respectively, in rabbits with hydraulic edema; and 5.1 ± 0.3, 4.3 ± 0.4 and 3.3 ± 0.6 g/dl, respectively, in rabbits with lesional edema. Contamination of plasma proteins from microvascular lesions during wick insertion was 14% of plasma Cprotein. In control animals, pulmonary interstitial Cprotein was lower than previous estimates from pre- and postnodal pulmonary lymph; furthermore, although the interstitium constitutes a continuum within the lung parenchyma, regional differences in tissue content seem to exist in the rabbit lung.
lung edema; lung fluid balance; regional pulmonary interstitial protein concentration
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INTRODUCTION |
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THE PULMONARY INTERSTITIUM is well structured to fulfill mechanical as well as gas exchange requirements of the lung. The thinness of the perialveolar interstitium and thus of the alveolocapillary barrier critically depends on tissue hydration, which, in turn, results from the balance between fluid filtration at the capillary level and lymphatic drainage of fluid and proteins from the pulmonary interstitium.
Pulmonary transcapillary fluid flux (Jv)
depends on the hydraulic (P) and colloid osmotic () pressures in the
capillary (Pcap and
cap, respectively) and
the surrounding interstitium (Ppi and
pi,
respectively) as described by the Starling equation
Jv = Lp · S · [(Pcap
Ppi)
(
cap
pi)], where Lp is the hydraulic conductivity, S is the surface area of the capillary
endothelium, and
is the reflection coefficient of the endothelium
to total plasma proteins.
pi is one of the components
determining transendothelial fluid flux and pulmonary tissue hydration
either in control conditions or during edema development.
In intact rabbit lungs physiologically expanded at negative pleural pressure and atmospheric alveolar pressure, direct measurements of pulmonary Pcap and perivascular Ppi showed that Ppi is very subatmospheric under control conditions (16), with a net gradient sustaining fluid filtration into the perivascular interstitial space along the whole microvascular network down to the venular end (18). However, because no direct measurements of perivascular pulmonary interstitial protein concentration (Cprotein) in control rabbits have been reported yet, the net transendothelial pressure gradient could be only indirectly estimated. Indeed, no direct samples of pulmonary interstitial fluid have ever been obtained from control lungs of rabbits or other mammalian species. In control and edematous lambs, sheep, and dogs, Cprotein in the pulmonary interstitial fluid has been considered similar to the Cprotein in fluid collected from lymphatic vessels (Clymph) efferent to the intrathoracic lymph nodes such as the tracheobronchial or caudal mediastinal nodes (postnodal lymph) (8, 10, 25, 29). However, 26% of the postnodal lymph from the caudal mediastinal node is of nonpulmonary intrathoracic origin (11). In addition, paired samples of pre- and postnodal lymph showed that prenodal Clymph more closely mirrors interstitial Cprotein, with lymph proteins being concentrated by ~20% in the passage through the node (14). In dog lungs, interstitial fluid has also been collected with glass pipettes (31), by sampling the perivascular or peribronchial cuff in frozen tissue (30), or with mediastinal wicks implanted in vivo in the peribronchial tissue layer running along the larger upper airways (22). However, these techniques were developed in severely edematous lungs and were not applicable to normally hydrated lungs. In addition, neither sampling of pre- or postnodal lung lymph nor the other mentioned techniques allowed simultaneous measurement of local pulmonary interstitial Cprotein in the same species and experimental conditions.
Hence, in the present study, we proposed a new experimental approach to directly sample regional interstitial fluid from control or edematous rabbit lungs. To this aim, we developed a modified wick technique to collect interstitial fluid from loose connective tissue surrounding the lobar pulmonary veins and bronchi departing from the pulmonary hilum. A preliminary report of the present study has been presented elsewhere (19).
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METHODS |
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Experiments were performed on 38 adult rabbits (body weight 2.6 ± 0.14 kg) anesthetized with urethane diluted at 25% in saline plus pentobarbital sodium (10 mg/ml of urethane solution). To induce deep surgical anesthesia, 2.5 ml/kg body wt of the anesthetic were given in an ear vein. The animals were tracheotomized and left to breathe spontaneously while lying supine; a carotid artery and a jugular vein were cannulated with saline-filled catheters connected to three-way stopcocks.
Experimental Protocol
In 12 control rabbits, a blood sample was withdrawn, and the animals were killed with an anesthesia overdose for sampling of pulmonary interstitial fluid. After respiratory arrest, a bolus of saturated KCl was given intravenously to cause complete cardiac arrest.In another group of rabbits (n = 15), after a blood sample was withdrawn, pulmonary hydraulic edema was induced through intravenous saline infusion (10% body weight in 30 min). At the end of the infusion period, the animal was left to breathe spontaneously for 1 h to allow equilibration of the infused saline with the interstitial compartment. After a postinfusion blood sample was withdrawn, cardiac arrest was induced as described above.
To test the present methods of fluid tissue sampling against changes in protein levels in the perivascular and peribronchial pulmonary compartments, in a third group of animals (n = 4), lesional edema was induced through an intravenous single bolus of 200 µg (7 UI) of pancreatic elastase. A blood sample was withdrawn 180 min after the elastase injection, followed by an anesthesia overdose as described above. Pancreatic elastase is an omnivorous proteolytic enzyme that has been shown to induce lesional pulmonary edema by affecting the matrix architecture of the microvascular basement membrane (21, 26).
Sampling of Pulmonary Interstitial Fluid
Immediately after death, the tracheal cannula was occluded, the thorax was opened, and the ribs on both sides of the chest were removed. The inferior vena cava and esophagus were severed, and the mediastinal tissues and thymus were removed from the ventral surface of the pericardium; the latter was opened and partially displayed to allow exposure of the pulmonary hilar regions. The lung was slightly inflated with room air and covered with gauze wet with saline. Interstitial fluid was sampled from the perivascular and peribronchial connective tissue spaces of both lungs by using a wick method slightly adapted from that described by Aukland and Fadnes (1) and later modified for use in muscle by Wiig et al. (34). The rationale of the wick technique is to collect interstitial fluid by absorbing the free fluid and soluble solutes through dry wicks implanted in a given tissue, i.e., in the present study, the perivascular and peribronchial pulmonary connective layers. Three-strand nylon wicks (diameter ~390 µm, length ~2-3 cm) were inserted into siliconized 3-cm-long PE-50 plastic catheters. The wicks were prewashed in acetone and ethanol, rinsed twice in distilled water, dried, and kept in a humidified chamber before insertion into the catheters. The catheters filled with wicks were inserted into the connective tissue layers surrounding the main branches of the right and left pulmonary veins and the principal bronchi of the right and left caudal lung lobes. The catheter was introduced into the perivascular or peribronchial tissue via a small hole made with iridectomy forceps in the connective layer around the vessel or bronchus, medially with respect to the pulmonary hilum. Given the thinness of the pulmonary vein wall, this phase required extreme care to avoid lesion of the vessel wall. The catheter was gently advanced along the vessel or bronchus following the route of least tissue resistance until it was wedged in the pulmonary tissue; when the lobar vein was superficial, it was possible to check the proper lodging of the catheter in the perivascular interstitium. While the catheter was held with a small forceps, a thin solid steel wire (diameter 520 µm) was inserted into the catheter hole to hold the wick in place. The catheter was subsequently withdrawn over the wick; the wire was also removed, leaving the wick in place in the tissue. A typical example is presented in Fig. 1, showing a wick inserted along the lobar vein of the medium right lobe; the wick runs parallel to the vessels over a length of ~2 cm. In each of the right and left lungs, up to four wicks were inserted along the lobar veins and one to two along the main caudal bronchus. Care was taken not to perforate the lung parenchyma during catheter insertion. In cases in which the lung tissue was lesioned or the vessel was damaged, causing hemorrhage at the side of insertion, the catheter was withdrawn and implanted along a different branch. In four control rabbits, to check on the effect of lung tissue lesion on the protein content of the wick fluid, two to three extra wicks were purposely inserted directly into the lung tissue, clearly damaging the parenchyma.
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To avoid tissue dehydration, the sites of catheter insertion were wet with mineral oil, the lung was covered with gauze wet with saline, and the animal was moved into an incubator with 100% relative humidity. After 20 min of implantation, a time shown to be needed to reach fluid saturation of the wick strand (33) with the animal in the incubator, the wicks were withdrawn from the tissue and immediately immersed in mineral oil to avoid dehydration of the wick and/or loss of the sampled fluid. While still immersed in oil, both ends were cut over a length of ~0.5 cm, and the remaining wick length was placed under mineral oil in vials equipped with a funnel. Wicks that appeared contaminated with blood, as judged by visual inspection, were discarded.
The time interval from the induction of anesthesia to death was ~30 min; another ~60 min were required for wick insertion into the lung tissue and subsequent extraction after 20 min of implantation. Implantation and extraction of all wicks were completed within 1 h of the death of the animal.
Wicks from the perivascular interstitium of the right and left lungs were placed in different vials; conversely, to collect enough fluid for the subsequent analysis, wicks from the right and left peribronchial connective spaces were pooled together in the same vial.
After all the wicks were extracted, the lungs were excised, and each lung was cut into six parallel slices in the ventrodorsal direction. Tissue specimens were weighed after samples were taken and after they were oven-dried at 70°C for 24 h to measure the wet weight-to-dry weight ratio (W/D). A mean W/D was then calculated for each lung; data presented in the text for control and treated lungs are the averages calculated from the mean individual values.
Morphology
In one control rabbit, wicks were inserted in both the perivascular and peribronchial tissue layers in the right and left lungs as described in Sampling of Pulmonary Interstitial Fluid. Thereafter, the wicks were left in place while the whole lungs and upper airways were excised and fixed overnight in acetate-formalin. To show the wick position, the lungs were cut in frontal sections perpendicular to the hilum at intervals ranging from 0.3 to 0.5 cm. The specimens were routinely processed, paraffin embedded, cut into multiple sections (3/specimen), and stained with hematoxylin and eosin.Isolation of Fluid From the Wicks and Analysis Performed on the Interstitial Fluid
Fluid was isolated from the wicks and analyzed according to one of the following procedures.Biochemical determination of interstitial Cprotein and electrophoresis. Wicks from a group of rabbits (group A) that included seven control animals and five animals with hydraulic edema were submerged in mineral oil immediately after their removal from the tissue and were centrifuged at 17,000 g in a supercentrifuge (Beckman J-21 model) for 30 min. The liquid was recovered at the bottom of the centrifuge tubes and aspirated with a micropipette (Microman, Gilson). The typical volume recovered ranged between 1 and 2 µl. Fluid Cprotein was determined with the bicinchoninic acid method (Pierce, Rockford, IL) with BSA as a standard. SDS-PAGE was performed on samples of equal volume diluted 1:4 in reducing sample buffer (0.5 M Tris · HCl, pH 6.8, containing 10% glycerol, 2% SDS, 5% mercaptoethanol, and bromphenol blue as a tracer) and heated to 100°C for 1 min. The electrophoresis was carried out at 180 V for 1 h in a 10% polyacrylamide minigel (Bio-Rad Mini-PROTEAN II). The bands were detected after gel fixation in a methanol-acetic acid-water solution (50:10:40) and staining with a 0.2% Coomassie blue R-250 solution (methanol-acetic acid-water at 45:15:40). The gels were dried in a gel drier apparatus (Bio-Rad model 583) with cellophane membrane backing and then analyzed by computer-assisted densitometry (Bio-Rad model 780).
Direct measurement of of isolated interstitial fluid.
Wicks from the lungs of another group (group B) that
included five control rabbits, nine rabbits with hydraulic edema, and four rabbits with lesional edema were immersed in oil after removal from the tissue and spun at 3,000 rpm for 40 min. The wick fluid that
collected at the bottom of the centrifuge tube was aspirated into a
plain glass capillary and recentrifuged at 3,000 rpm for 5 min to
separate the sampled fluid from the mineral oil. The wick fluid volume,
measured through 0.5-µl glass microcapillaries, ranged between 0.5 and 3 µl. Wick fluid from seven control and nine edematous lungs was
transferred directly to a colloid osmometer (2) equipped
with a PM 30 membrane (molecular mass cutoff 30,000 Da; Amicon); the
osmometer was connected to a pressure transducer (model 4-327-1, Transamerica Delaval) that conveyed a pressure signal to a thermal
oscillograph (Gould 4600). To avoid evaporation of the fluid, the
osmometer sample chamber was covered with Parafilm. The fluid was put
in a microsyringe and poured onto the osmometer membrane surface
through a microneedle driven through the Parafilm layer into the sample
chamber. After each
measurement, the membrane was changed and the
zero reference pressure of the osmometer was checked.
Recovery Studies
The degree of recovery of proteins by the nylon wick was assessed by immersing the wicks in whole autologous plasma and in plasma diluted 1:2, 1:3, and 2:3 in a saline solution. After 20 min, the wicks were extracted from the standard solution in the incubator and processed for fluid isolation and measurement of Cprotein andEstimation of Traumatic Contamination of Wick Fluid by Plasma Proteins
The size of the catheter used for wick implantation largely exceeded the thickness of the connective perivascular and peribronchial tissue layers. Therefore, catheter insertion might have directly damaged the vessels and/or the surrounding parenchyma, leading to acute plasma protein extravasation into the pulmonary interstitium. To check for contamination of the wick fluid with proteins escaping from damaged vessels, in six control rabbits, a bolus of 0.08 MBq of 125I-human serum albumin (HSA; Institute for Energy Technology, Kjeller, Norway) was injected into the jugular vein and allowed to equilibrate for 5 min. Free 125I in the injectate was <1% as determined with trichloroacetic acid precipitation. Then, a blood sample was withdrawn, the animal was killed, and the wicks were inserted in both lungs according to the standard procedure. After 20 min of implantation, the wicks were extracted from the tissues and the wick fluid was isolated. The wick fluid volume was measured in 0.5-µl plain glass microcapillaries (Modulohm, Herlev, Denmark) that were subsequently placed in glass vials containing 1 ml of 0.02% azide saline for elution. Wick-containing tubes were capped and shaken vigorously to allow complete mixing of the sampled fluid in the glass microcapillary with the elution solution. Radioactivity in the wick fluid eluate and the plasma was then assayed in an LKB gamma counter (model 1282, Compugamma, Turku, Finland). After the radioactivity was counted, the albumin concentration in the eluate and plasma was measured with a fluorometric method with 1-anilinonaphthalene-8-sulfonic acid as described by Rees et al. (27) and modified by Aukland and Fadnes (1) to calculate the specific activity of albumin.Statistics
Data are means ± SD. Significance of the differences between mean values was tested with Student's t-test to compare perivascular and peribronchial Cprotein data in the same animal group (paired t-test) or between groups treated with the same experimental protocol (unpaired t-test). One-way analysis of variance (ANOVA) was performed with data from the groups treated with the different experimental protocols. The average data presented in Fig. 3 were tested with the all pairwise multiple comparison procedure (Tukey's test). Mean values were considered different when P < 0.05. ![]() |
RESULTS |
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Insertion of the catheter in the perivascular interstitium was difficult in some cases because 1) the size of the catheter exceeded the thickness of the connective perivascular layer, offering high resistance to sliding in the tissue and implying possible damage to the vessel wall and to the lung parenchyma; 2) sliding of the catheter along the vein was often hindered by lateral branches; and 3) it was possible to check for proper perivascular wick lodging only when the lobar vein was superficial. About 30% of the implanted wicks were discarded as a consequence of a local hemorrhage or evident parenchymal lesion. Peribronchial implantation never caused hemorrhage, but the insertion was more difficult than perivascular implantation due to a higher tissue resistance to catheter sliding. Fluid extracted from properly positioned perivascular and peribronchial wicks (Fig. 1) appeared straw colored.
Morphology
The microphotographs presented in Fig. 2A show a lung tissue section with a wick implanted in the perivascular connective layer near a small pulmonary vessel. Wick fibers are in close proximity to the vessel lined by clearly recognizable endothelial cells. None of the sections with perivascular wick implantation showed signs of vessel wall lesion. The section in Fig. 2B shows a wick implanted in the connective tissue surrounding a small bronchus that is clearly recognizable by the ciliated cells lining its inner wall. In both perivascular and peribronchial sections, the lung parenchyma surrounding the wick is slightly compressed, with small hemorrhagic areas. The thickness of the affected tissue around the wick never exceeded ~50 µm.
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Perivascular and Peribronchial Interstitial Fluid Cprotein in Control Lungs
In control animals in which wick fluid Cprotein was measured directly (group A; n = 7) or was derived from measurements ofDirectly measured control peribronchial Cprotein
was 2.5 ± 1.4 g/dl (42.4% of plasma
Cprotein). In group B, peribronchial fluid was 11.2 ± 3.5 cmH2O, corresponding to
2.4 ± 0.7 g/dl, (34.2% of plasma
Cprotein), not different from the directly
measured peribronchial Cprotein. Given the
similarity between data from groups A and B,
perivascular and peribronchial Cprotein data were pooled and are presented in Fig. 3
together with the corresponding average plasma
Cprotein (6.4 g/dl; corresponding
calculated with Eq. 1 = 30.6 cmH2O). One-way ANOVA
performed on the data shown in Fig. 3 indicates that in control
animals, perivascular Cprotein (57.8% of
plasma Cprotein) was significantly higher than peribronchial Cprotein (37.5% of plasma value;
P < 0.05), and both were significantly lower than the
corresponding plasma Cprotein.
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When wicks were purposely inserted in the parenchyma to detect the effect of tissue lesion on wick fluid proteins, wick Cprotein was not significantly different (106.9 ± 33%) from the corresponding plasma value.
Perivascular and Peribronchial Interstitial Fluid Cprotein in Hydraulic and Lesional Edema
In animals receiving saline infusion (n = 15), plasma Cprotein and blood hematocrit averaged 6.2 ± 0.4 g/dl and 0.48 ± 0.054, respectively, before the onset of infusion and fell to 4.7 ± 0.7 g/dl (correspondingThe lung W/D was 4.9 ± 0.04 g H2O/g dry tissue
weight in control animals and increased to 5.4 ± 0.3 g
H2O/g dry tissue weight after the infusion, indicating
attainment of a condition of mild pulmonary edema (15,
20). In group A (n = 6 rabbits),
directly measured right and left perivascular wick fluid
Cprotein values were 2.3 ± 0.8 and 2.4 ± 0.9 g/dl, respectively. In group B (n = 9 rabbits), perivascular fluid was 11.7 ± 2.6 cmH2O, the corresponding Cprotein
being 2.5 ± 0.6 g/dl. After the infusion, peribronchial fluid was
2.3 ± 0.8 g/dl when directly measured in group A and 2.4 ± 0.4 g/dl when derived from the peribronchial wick fluid
(11.2 ± 3.5 cmH2O in group B). Because
Cprotein values did not differ when measured
directly or derived from the corresponding
values, direct and
derived perivascular and peribronchial Cprotein values were pooled and are presented in Fig. 3. ANOVA revealed that the
perivascular Cprotein was significantly reduced
with the saline infusion compared with the control value, whereas
peribronchial Cprotein was not affected by plasma
expansion. When expressed as a percentage of the corresponding plasma
value, postinfusion perivascular Cprotein
(47.3 ± 16.8% of plasma Cprotein) was not significantly different from the corresponding control value, indicating that endothelial sieving was unaffected.
In rabbits that developed interstitial lesional edema after receiving
pancreatic elastase (lung W/D = 5.3 ± 0.2; n = 4), plasma Cprotein before the anesthesia
overdose averaged 5.1 ± 0.3 g/dl (Fig. 3), a value ~20% lower
compared with the control value. directly measured on the wick
fluid extracted from the perivascular interstitial tissue averaged
20.4 ± 2.1 cmH2O, corresponding to a
Cprotein of 4.4 ± 0.4 g/dl, not
significantly different from perivascular control
Cprotein. However, when expressed as a percentage of the corresponding plasma Cprotein,
perivascular Cprotein was significantly higher
after elastase (86.6 ± 3.5%) compared with the control condition
(57.6 ± 3.7%), indicating an increased protein leakage in
lesional edema. Peribronchial
and the corresponding Cprotein averaged 15.6 ± 2.8 cmH2O and 3.3 g/dl (68.5 ± 7.2% of plasma value),
respectively. ANOVA indicated that peribronchial Cprotein after elastase was significantly
different from the corresponding plasma Cprotein
but not from the perivascular Cprotein.
Gel Electrophoresis of Perivascular Fluid Proteins
Protein gel electrophoresis (Fig. 4) of plasma and perivascular fluid under control conditions and in hydraulic edema clearly showed that the wick fluid contained only proteins normally present in plasma; the absence of foreign proteins suggested that no cell lesion occurred during acute wick insertion. The most represented protein in the control (58.9 ± 1.2% of total proteins) and edematous (64 ± 9%) perivascular interstitial fluids displays a molecular mass of ~70 kDa, likely corresponding to albumin. The plasma albumin fraction in the control fluid was 54.3 ± 16%.
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Extravasation of Plasma Proteins
125I-HSA recovery from wick fluid in control samples indicated that there was no significantly different contamination in perivascular (13.9 ± 3.3% of total wick fluid proteins) compared with peribronchial (14.5 ± 3.6%) wicks. The similarity of isotope recovery in perivascular and peribronchial fluids indicates that contamination by plasma proteins derived from a microvessel lesion rather than from rupture of larger vessels during catheter insertion.With the assumption of a constant interstitial fluid volume, actual
interstitial Cprotein was calculated from the
average perivascular and peribronchial values as actual interstitial
Cprotein = experimental interstitial
Cprotein 0.14 · plasma
Cprotein. After the correction, control
perivascular and peribronchial fluid Cprotein
values were 2.7 ± 0.4 (
calculated with Eq. 1 = 12.5 cmH2O) and 1.5 ± 0.8 g/dl (
= 7 cmH2O), respectively; the perivascular interstitial-to-plasma Cprotein ratio
(Ci/Cp) was 0.42 ± 0.03, significantly
higher than the corresponding peribronchial
Ci/Cp of 0.23 ± 0.11 (by one-way ANOVA).
The corrected perivascular and peribronchial
Cprotein values after saline infusion were
1.8 ± 0.5 (
= 8.4 cmH2O) and 1.7 ± 0.3 (
= 7.9 cmH2O) g/dl, respectively, yielding
Ci/Cp values of 0.37 ± 0.11 and 0.35 ± 0.07, respectively. Hence, after saline infusion, the peribronchial
Ci/Cp was not significantly different from the
corresponding perivascular value but was significantly higher than the
control peribronchial Ci/Cp. Saline infusion
caused a small, although not significant, decrease in perivascular
Ci/Cp compared with the control value,
suggesting that the development of local tissue edema was essentially
due to plasma protein dilution without lesion of the endothelial layer. Tissue washout did not seem to affect the peribronchial interstitium where Ci/Cp increased as a consequence of
plasma protein dilution in the face of an unchanged tissue
Cprotein.
In lesional edema, perivascular and peribronchial
Cprotein values corrected for plasma protein
extravasation amounted to 3.7 ± 0.4 ( = 17.5 cmH2O) and 2.7 ± 0.5 (
= 12.7 cmH2O) g/dl, respectively, with the corresponding
Ci/Cp values being 0.73 ± 0.03 and
0.53 ± 0.07. Both the perivascular and peribronchial ratios
were significantly higher than those in the control condition,
indicating a greater escape of plasma proteins into the
surrounding tissue; yet the Ci/Cp was still
lower in peribronchial than in perivascular fluid.
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DISCUSSION |
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Validation of the Methods
Tissue fluid has been sampled from subcutis, muscle, and skin with saline-soaked wicks implanted in vivo or dry wicks implanted postmortem (1, 12, 17, 33, 34). In the present study, we adopted the postmortem dry wick implantation technique to sample pulmonary interstitial fluid; due to lung morphology, the wick catheters could be lodged only in the connective tissue layers surrounding the lobar pulmonary vein and the bronchi coming from the pulmonary hilum. An intrinsic limit of this approach is that the fluid sample is necessarily restricted to the superficial lobar veins and might not satisfactorily represent the actual situation around the vast majority of the smaller pulmonary vessels. On the other hand, filtration of pulmonary interstitial fluid occurs at the level of the smallest arterioles and of the capillary bed down to the net pressure gradients set by the Starling forces, depending on the permeability properties of the microvessel wall. No significant differences have been reported so far between the vascular pressure profile (18) or the permeability properties of the superficial compared with the deeper pulmonary microvascular network. Hence perivascular interstitial protein composition might not substantially differ in the superficial compared with the deeper parenchyma.Catheter insertion induced mechanical damage to the pulmonary tissue as seen in Fig. 2. Local lesion of the vascular bed wall was seen also by recovery of the intravascular 125I-HSA in the wick fluid. However, damage seemed to be limited to a thin layer of parenchyma and to local capillary lesion. Indeed, because isotope leakage from the blood was similar in perivascular and peribronchial wick fluids, blood contamination cannot derive from large blood vessel rupture. Extensive capillary lesion along the wick path was observed only after direct insertion of the catheter in the parenchyma, leading wick fluid Cprotein to approximate plasma Cprotein. Contamination of wick fluid by 125I-HSA during wick insertion was higher than that observed in rat hindlimb or back muscles [1-2% (34)] in experiments in which a similar wick technique was used. This might depend on a much greater capillary surface area per unit of interstitial tissue volume (24) in the lung (~3,000 cm2/g tissue) compared with that in muscle (70 cm2/g tissue). However, the isotope extravasation experiments enabled us to account for this source of error.
Alveolar lining fluid might have been absorbed into the wick after
lesion of the lung parenchyma. For an approximate wick catheter length
of 2 cm (see Fig. 1) and a width of the damaged parenchyma of ~50
µm (Fig. 2A), damaged tissue volume would be ~4 × 104 cm3 that, assuming a tissue density of
~0.3 g/cm3, would weigh ~1 × 10
4 g.
Assuming an alveolar liquid volume of 0.37 ml/kg body wt
(28) and a lung weight of ~10 g in a 2.6-kg rabbit, the
alveolar fluid volume from lesioned parenchyma would be ~0.012 µl,
accounting for ~1% of the average fluid volume extracted by the
wick. The total Cprotein of alveolar lining fluid
[3.7 g/dl (28)] is higher than that in the perivascular
and peribronchial fluids, so that contamination with alveolar fluid
might lead to an overestimation of the Cprotein
of the pulmonary interstitial liquid. However, in the epithelial lining
fluid collected through bronchoalveolar lavage, the albumin fraction
was ~20%, the majority being proteins of a higher molecular mass
(7). Because the fraction of proteins larger than albumin
was negligible in the control perivascular wick fluid (Fig. 4), we
conclude that contamination with alveolar fluid would not significantly
affect our present interstitial Cprotein.
As observed by Wiig and colleagues (33, 34) in the rat hindlimb, needle wick insertion would cause the appearance in the wick fluid of a substantial fraction of high molecular mass globulins, not corresponding to plasma components but mostly derived from cell lesion. Albumin was the greater protein fraction in perivascular fluid, with no presence of foreign proteins not appearing in plasma (Fig. 4); the albumin percentage was similar to that observed in catheter-inserted wick fluid in the hindlimb but higher than that observed in needle-inserted wicks, suggesting that cell disruption did not significantly affect perivascular interstitial Cprotein.
The above considerations led us to conclude that although one cannot exclude a certain degree of pulmonary tissue lesion, the fluid sampled with the present technique is a reliable method of collection of pulmonary perivascular and peribronchial interstitial fluids. In addition, to test the reliability of our methods, we sampled interstitial fluid in three different conditions (control, hydraulic edema, and lesional edema) to assess whether the methods were sensitive to changes in interstitial fluid protein composition. The experimental protocols adopted to develop hydraulic and lesional edema have previously been used extensively (15, 20, 21, 26) to induce a condition of mild interstitial edema as also indicated by a moderate increase in the W/D observed in both edema conditions. In particular, elastase treatment has been shown to increase microvascular permeability by affecting the molecular integrity of heparan sulfate proteoglycans of the perimicrovascular basement membrane (21, 26). The choice of inducing a mild rather than a severe hydraulic or lesional edema was made to modify the parameters dealing with transvascular fluid bulk flow, i.e., the net pressure gradients and/or microvascular permeability, without causing massive alveolar flooding. The modified wick technique allowed us to detect pulmonary perivascular Cprotein variations depending on the sieving properties of the capillary network and/or plasma Cprotein. Indeed, when expressed as a percentage of the corresponding plasma value, perivascular Cprotein decreased after saline infusion, a condition characterized by a low plasma Cprotein and a normal endothelial sieving. On the other hand, the percentage of perivascular Cprotein significantly increased after elastase treatment, suggesting a damaged endothelial basement membrane layer, whereas the corresponding absolute perivascular Cprotein was not significantly different compared with the control value (see Fig. 3). This apparent contradiction can be explained by the observation that plasma Cprotein was reduced by ~20% after elastase administration. A decrease in plasma Cprotein in this model of lesional edema may be expected, considering that pancreatic elastase is known to be an omnivorous proteolytic enzyme that likely disrupts the molecular structure of several plasma as well as tissue proteins.
Comparison With "In Vivo" Interstitial Fluid Protein Content
Postmortem composition of the interstitial fluid might differ from the actual in vivo situation. After cardiac arrest, functional capillary pressure drops to zero, nullifying transcapillary filtration; in addition, lymphatic drainage is abolished. Hence, no net movement of fluid and/or proteins is expected to occur postmortem between the vascular and the extravascular compartments.A possible source of postmortem modification of interstitial fluid is fluid shift from the extracellular to the intracellular compartment down osmotic pressure gradients, potentially leading to cell swelling and increased interstitial Cprotein. However, this phenomenon ought to be negligible in the lung; indeed, given the high degree of vascular supply to the pulmonary parenchyma, an increase in tissue Cprotein would likely be offset by a capillary-to-tissue water shift. Experiments done in rat hindlimb and back muscles showed that interstitial Cprotein was stable up to 80 min of wick implantation time (34), ruling out the possibility, in this frame time, of both cell swelling and/or a possible interstitium-to-capillary fluid shift due to colloid osmotic forces. Because we implanted and withdrew all wicks within 60 min from cardiac arrest, we expect that the present interstitial fluid Cprotein mirrors the actual control value; this consideration was strengthened by the similarity observed between interstitial fluid Cprotein measured in skin (33), muscles (33, 34) and extrapleural interstitium (17) with postmortem dry wicks or with in vivo saline-soaked wicks.
In normal sheep, lambs, and dogs, the average in vivo postnodal Clymph ranged between 3 and 3.8 g/dl (8, 10, 25, 29), with no significant interspecies differences. A more reliable estimate of interstitial Cprotein would derive from prenodal Clymph, ranging between 2.5 and 3.8 g/dl, with an average Clymph-to-Cprotein ratio of ~0.7 (9, 14, 23). The present perivascular Cprotein values after extravasation correction are in the lower range of prenodal values; however, as pointed out by Aukland and Reed (3), prenodal Clymph may be higher than interstitial Cprotein due to tissue manipulation during lymph vessel cannulation.
Differences Between Perivascular and Peribronchial Fluid Cprotein
Under control conditions, Cprotein is higher in the perivascular than in the peribronchial interstitium (Fig. 3); in addition, in the face of what was observed in the perivascular region, no significant washout of proteins occurred in the peribronchial adventitial connective tissue with saline infusion. In lesional edema, a greater capillary permeability to proteins resulted in an increase in both perivascular and peribronchial Cprotein, even though the effect was more evident in the perivascular compartment where Cprotein was not significantly different from the plasma value.This would confirm the suggestion (23) that although the pulmonary interstitium may be regarded as a continuum connecting the septal perialveolar interstitium to the hilar connective tissue (32), differences exist in local tissue fluid dynamics.
Although the alveolar interstitial space is supplied by the pulmonary circulation, the connective tissue surrounding the main vessels and airways may be partially supplied by the bronchial systemic circulation, accounting for 20-90% of total canine airway blood flow (13). Systemic-to-pulmonary vascular anastomoses were observed at pre- and postcapillary levels in dogs, sheep, horses, and humans (4). In dogs and sheep, even though the bronchial veins empty into the pulmonary veins, the hydraulic pressure is higher in the bronchial than in the pulmonary capillaries; in addition, unlike pulmonary capillaries in which the endothelium is of the continuous type, the bronchial endothelium is fenestrated (13). At variance with what was observed in dogs and sheep, bronchopulmonary anastomoses are not common in rabbits and the microvascular network around the airways are poorly developed (6). These morphological features and limited development of the bronchial microvascular network to the peribronchial adventitial tissue might account for the low peribronchial Cprotein as well as for the lack of local tissue washout.
In summary, in the present study, we developed an implanted wick approach to extract fluid from the perivascular and peribronchial interstitial fluids, allowing, for the first time, a direct measurement of pulmonary interstitial Cprotein as well as a simultaneous comparison between perivascular and peribronchial connective tissues in rabbits. Further utilization of this technique will allow more careful investigation of the interstitial fluid composition and transcapillary fluid dynamic in normal and edematous mammalian lungs.
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ACKNOWLEDGEMENTS |
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We are grateful to Prof. Giuseppe Miserocchi for useful discussion of the data.
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FOOTNOTES |
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This study was supported in part by the Italian Ministry of University and Scientific and Technological Research, the L. Meltzers Fund (University of Bergen, Bergen, Norway), and Training and Mobility Research Grant ERBFMRXCT980219 from the European Community Grant.
Part of this research was performed while D. Negrini was a guest researcher at the University of Bergen.
Address for reprint requests and other correspondence: D. Negrini, Dipartimento di Medicina, Chirurgia e Odontoiatria, Sezione Fisiologia Umana, Via Mangiagalli 32, 20133 Milan, Italy (E-mail: daniela.negrini{at}unimi.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 25 August 2000; accepted in final form 28 November 2000.
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