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Microvascular membranes are heteroporous, so the mean osmotic reflection coefficient for a microvascular membrane (sigma d) is a function of the reflection coefficient for each pore. Investigators have derived equations for sigma d based on the assumption that the protein osmotic pressure gradient across the membrane (delta II) does not vary from pore to pore. However, for most microvascular membranes, delta II probably does vary from pore to pore. In this study, we derived a new equation for sigma d. According to our equation, pore-to-pore differences in delta II increase the effect of small pores and decrease the effect of large pores on the overall membrane osmotic reflection coefficient. Thus sigma d for a heteroporous membrane may be much higher than previously derived equations indicate. Furthermore, pore-to-pore delta II differences increase the effect of plasma protein osmotic pressure to oppose microvascular fluid filtration.

Many investigators have used the lymphatic protein washdown technique to estimate the pulmonary microvascular membrane reflection coefficient to protein (sigma d). With that technique, the investigator causes a high microvascular filtration rate then estimates sigma d from the lymph and plasma protein concentrations. However the lymph may contain protein washed from the lung tissue, and the tissue protein may cause investigators to underestimate sigma d. Plasma protein osmotic pressure (IIc) may cause investigators to underestimate sigma d because IIc opposes fluid filtration. To minimize the effect of IIc, we decreased IIc to 5.6 +/- 1.1 mmHg in five anesthetized sheep. We increased the microvascular filtration rate by increasing pulmonary microvascular pressure to 22 +/- 3 mmHg. Then we tagged plasma protein with Evans blue dye and estimated sigma d from the lymph and plasma dye concentrations. Because tissue protein was not tagged, it did not interfere with our sigma d estimate. Our sigma d estimate (0.79 +/- 0.08) was much higher than previous estimates in anesthetized animals.

Lymphatic vessels normally remove excess fluid from the lung tissue and thus prevent pulmonary edema. However, when lungs do become edematous, the edema fluid may be cleared via several pathways. Each clearance pathway has unique properties, and recent studies have helped to define those properties.

We used a new technique to estimate the pulmonary microvascular membrane reflection coefficient to plasma protein (sigma d) in anesthetized dogs. In five animals we continuously weighed the lower left lung lobe and used a left atrial balloon to increase the pulmonary microvascular pressure (Pc). We determined the relationship between the rate of edema formation (S) and Pc and estimated the fluid filtration coefficient (Kf) as delta S/delta Pc. From the S vs. Pc relationship and Kf, we estimated the Pc at which S/Kf = 10 mmHg for each dog. This pressure (P10) was 38.0 +/- 5.8 (SD) mmHg, and the plasma protein osmotic pressure (pi c) was 14.9 +/- 3.7 mmHg. In five additional dogs in which we decreased pi c to 2.9 +/- 1.7 mmHg, P10 = 27.2 +/- 2.6 mmHg. The P10 vs. pi c regression line fit to the data from all 10 dogs was P10 = 0.92 pi c +/- 24.4 mmHg (r = 0.88). We estimated sigma d from the slope of the regression line as sigma d = square root of delta P10/delta pi c. With this technique, we estimated that, with 95% probability, sigma d lies between 0.72 and unity. This is higher than most previous sigma d estimates.

We used lymph flow rate (QL) to lymphatic vessel outflow pressure (Po) relationships to analyze lymphatic flow in five unanesthetized sheep with liver lymphatic cannulas. The olecranon was the zero reference level for pressures. Increases in Po did not change QL until Po exceeded 19 +/- 4 (SD) cmH2O. However, for Po greater than 19 +/- 4 cmH2O, QL decreased linearly with increases in Po. We fit regression lines to the QL vs. Po data for Po greater than 19 cmH2O and estimated the effective lymphatic resistance (RL) as -delta Po/delta QL. The effective pressure driving lymph (PL) was the Po at which QL = 0. At baseline, RL = 0.18 +/- 0.10 cmH2O.min.microliter-1 and PL = 29.6 +/- 3.4 cmH2O. When we increased hepatic vein pressure by 5.7 +/- 1.7 cmH2O, QL increased to 6.2 +/- 3.2 times baseline, RL decreased to 0.050 +/- 0.015 cmH2O.min.microliter-1, and PL increased to 37.1 +/- 3.5 cmH2O (P less than 0.05). Thus 1) liver lymph flow is very sensitive to increases in hepatic vein pressure, 2) there is a substantial QL vs. Po plateau for liver lymphatics, and 3) after hepatic venous pressure elevations, liver lymph flow increases as if it were driven by a higher pressure through a lower resistance.

Postnodal intestinal lymphatic pressure increases during rapid intravenous infusions with Ringer solution in sheep. Part of the lymphatic pressure increase is due to increased venous pressure at the lymphatic outflow (the neck veins). We tested the hypothesis that other factors besides increased neck vein pressure may cause increased lymphatic pressure during intravenous infusions. We placed cannulas into postnodal lymphatic vessels in eight sheep. After the sheep recovered from the surgery, we infused Ringer solution [46 +/- 21 (SD) ml/kg body wt in 30 min] intravenously into the sheep and inflated a balloon in the inferior vena cava. We adjusted the balloon inflation to prevent any increase in neck vein pressure during the infusions. At baseline, the intestinal lymphatic pressure was 15.5 +/- 2.5 cmH2O. During the infusions, lymphatic pressure increased significantly, and for the last 10 min of the infusion period, intestinal lymphatic pressure was 24.0 +/- 6.1 cmH2O. These results are consistent with the hypothesis that factors in addition to increased neck vein pressure may cause increased intestinal lymphatic pressure during rapid intravenous infusions.

Intravenous fluid infusions cause increased venous pressure and increased lymph flow throughout the body. Together the increased lymph flow and increased venous pressure (the outflow pressure to the lymphatic system) should increase the pressure within the postnodal intestinal lymphatics. To test this, we measured the pressure in postnodal intestinal lymphatics and the neck vein pressure in five awake sheep. At baseline, the neck vein pressure was 1.2 +/- 1.5 (SD) cmH2O and the lymphatic pressure was 12.5 +/- 1.7 cmH2O. When we infused Ringer solution intravenously (10% body weight in approximately 50 min), the neck vein pressure increased to 17.3 +/- 0.9 cmH2O and the lymphatic pressure increased to 24.6 +/- 3.8 cmH2O (both P < 0.05). In two additional sheep, the thoracic duct lymph flow rate increased from 0.8 +/- 0.4 ml/min at baseline to 5.5 +/- 2.0 ml/min during the infusions. Our results show that postnodal intestinal lymphatic pressure may increase substantially during intravenous fluid infusions. This is important because increases in postnodal lymphatic pressure may slow lymph flow from the intestine.

The lymph from most organs drains through the thoracic duct and into veins in the neck. We hypothesized that increases in neck vein pressure (Pnv) are reflected through the thoracic duct to the lung lymphatic-thoracic duct junction. To test this, we cannulated the lung lymphatics in the direction of flow in four sheep. We advanced each cannula until it entered the thoracic duct. Thus the pressure at the tip of the lymphatic cannula (Px) was the pressure at the outflow of the lung lymphatics. We also placed a balloon into the superior vena cava. One to two days later, we measured Px in the awake sheep as we inflated the balloon and increased Pnv in steps to 25–45 cmH2O. We found no significant differences in Px and Pnv. Furthermore, Px closely followed Pnv after each step increase in Pnv. These results support our hypothesis that increases in Pnv cause increases in the outflow pressure to lung lymphatics.

Estimates of the pulmonary microvascular membrane reflection coefficient (sigma) and permeability-surface area product (PS) are frequently made with the assumption that a percent change in transmicrovascular fluid flux (Jv) will be represented by an equal percent change in the lymph flow rate (QL) from a single cannulated lung lymph vessel. To test this, we measured QL in seven anesthetized dogs with the outflow end of the lymph cannula set at several heights (H) above and below the lung hilus. The left atrial pressure was then elevated to increase Jv, and QL was again measured at several H's. The percent increase in QL at elevated left atrial pressure depended on H. We used the QL data and lymph and plasma protein concentrations to estimate sigma and PS with a modified form of the Kedem and Katchalsky equations. The calculated values varied considerably with H. Our results indicate that changes in Jv are not represented by equal changes in QL. Therefore, techniques for estimating permeability that depend upon QL as an estimate of Jv may lead to erroneous estimates of sigma and PS.

We used a gravimetric technique to test for increased pulmonary capillary permeability after Escherichia coli endotoxin infusion in unanesthetized sheep. The sheep were chronically prepared with cannulas placed into the left atrium and pulmonary artery 1–2 wk before the experiments. We estimated pulmonary capillary pressure (Pc) as the average of pulmonary arterial and left atrial pressures, and used the modified method of Pierce to estimate the ratio of extravascular fluid weight (EVF) to blood-free dry weight. In 15 sheep we inflated a left atrial balloon to raise Pc to -10.7, 5, 10, or 15 mmHg above plasma oncotic pressure (IIc) for 3 h, then measured EVF. EVF averaged 4.0 +/- 0.2 (base line), 4.3 +/- 0.1, 4.5 +/- 0.1, and 5.1 +/- 0.5 (SD), respectively, for the four levels of Pc - IIc. We gave seven additional sheep 1 microgram/kg of E. coli endotoxin (0127:B8) and measured EVF after 3 h of stable Pc. Endotoxin increased Pc in each sheep. EVF was higher than control for the endotoxin sheep with Pc - IIc greater than -1. This finding is consistent with an increase in pulmonary capillary permeability caused by endotoxin. However, EVF was not elevated in the endotoxin sheep with Pc - IIc less than 1 mmHg. This shows that the increased permeability was insufficient to cause edema unless Pc was elevated. Thus endotoxin may cause edema by two mechanisms, 1) an increase in capillary permeability, and 2) an increase in Pc.

Lung weight is a useful indicator of increases in lung extravascular volume. In addition, the lung lymph flow rate (QL) is an important factor in lung fluid balance. We have studied the weight and QL responses to elevations in capillary pressure (Pc) in intact dog lung lower left lobes. We measured lobe weight continuously. We also measured QL from small lymph vessels from the same lobes. The base-line QL was 1.7 +/- 1.5 microliter/min, and the weight was constant. After we increased Pc by 8-20 cmH2O, both weight and QL increased transiently. In most lungs the weight reached a new steady state. When we increased Pc further, weight increased continuously; however, QL reached a plateau. The continuous weight gain was due to edema. These results show that weight and QL respond similarly in nonedematous lungs; however, the weight and QL responses in edematous lungs may be different.

The preparation for collecting lung lymph from sheep caudal mediastinal lymph node (CMN) efferent vessels is widely used to study the effects of endotoxin on lung microvascular permeability. However, there are nonpulmonary lymph vessels that drain into the CMN along with the afferent lymph vessels from the lung. Thus CMN lymph is a mixture of lymph from the lung and diaphragm lymph vessels as well as from other nonpulmonary lymph vessels. We studied the effect of 0.5–1.0 microgram/kg Escherichia coli endotoxin on the flow rates in diaphragm and CMN efferent lymph vessels (Qdi and QCMN, respectively) in unanesthetized sheep. For the time period between 2 and 5.5 h after endotoxin QCMN was increased from its base line of 7.2 +/- 4.4 (SD) to 17.3 +/- 10.6 ml/h and the lymph-to-plasma protein concentration ratio (L/PCMN) had increased from 0.68 +/- 0.11 to 0.81 +/- 0.06. During the same time period, Qdi was 4.5 +/- 3.1 ml/h compared with 1.0 +/- 0.8 ml/h at base line and the diaphragm lymph-to-plasma protein concentration ratio (L/Pdi) was 0.92 +/- 0.07 (base line = 0.74 +/- 0.15). The increases in flow rate and protein concentration were significant for each type of vessel (P less than 0.05). We conclude that the period of increased QCMN and L/PCMN after endotoxin is associated with an increase in Qdi and L/Pdi. Thus, it is difficult to determine how much of the CMN lymph response comes from the lungs and how much comes from diaphragm lymph vessels.

We used a gravimetric method to determine the ratio (gamma) of pulmonary venous to total pulmonary vascular resistance in intact dog lungs. From this ratio, pulmonary capillary pressure (Pc) can be calculated. The average value of gamma was 0.50 +/- 0.06 (mean +/- SD) in 10 dogs. We found no correlation between gamma and PO2, PCO2, pH, or hematocrit in the narrow ranges of these experiments. Over the capillary pressure range of 22.4--35.2 mmHg we found no correlation between gamma and Pc. The value of gamma found in this study is not significantly different from the value found in isolated perfused lungs.

Many investigators have used the chronic sheep lung lymph preparation to collect caudal mediastinal node (CMN) efferent lymph. These investigators have assumed that the lymph collected with the preparation is almost pure lung lymph. We examined 17 sheep for possible systemic contamination to the lymph, and in each sheep we found one to five lymph vessels that ran from the diaphragm to the CMN. Contamination from these vessels would not be eliminated in the chronic sheep preparation. We estimated the flow rate from these vessels to be 3.0 +/- 2.6 (SD) ml/h in anesthetized sheep. This represents 25-60% of the lymph flow rate in the chronic lymph preparation. In five sheep, we also located 1-4 esophageal lymph vessels that entered th CMN. These results show that lymph collected with the chronic sheep lung lymph preparation contains a significant nonpulmonary contamination.

The liver lymphatic system plays an important role in removing excess fluid from the hepatic tissue. A complete analysis of the liver lymphatic system would be difficult. However, we used a simple circuit-analysis technique to represent the intrahepatic portion of the lymph system as a single pressure source (PL) pushing lymph through a single resistance (RL). Liver lymphatic vessels were cannulated in nine halothane-anesthetized dogs. The lymphatic vessel outflow pressure (PO) was varied by raising the outflow end of the cannula. Lymph flow from the cannula (QL) decreased linearly with PO, and we calculated RL as -delta PO/delta QL and PL as the extrapolated PO at which QL = 0. At base line, PL = 8.5 +/- 2.9 cmH2O, and RL = 0.05 +/- 0.03 cmH2O.min/microliter. After we increased inferior vena caval pressure from 5.8 +/- 2.7 to 15.2 +/- 2.5 cmH2O, PL increased significantly to 13.7 +/- 3.4 cmH2O, and RL decreased to 0.02 +/- 0.02 cmH2O.min/microliter (P less than 0.05). The results indicate that increases in QL occur because the effective pressure pushing lymph from the liver (PL) increases, and the effective resistance of the intrahepatic lymph vessels (RL) decreases.

Lysophosphatidylcholine (lyso-Pc) is a lysophospholipid normally found in low concentrations in the lung. At high concentrations lyso-Pc, instilled into the airways, causes pulmonary edema. We tested the hypothesis that the edema caused by lyso-Pc was due to an increase in pulmonary microvascular membrane permeability. In 11 anesthetized dogs we continuously weighed the left lower lobes (LLL) while instilling lyso-Pc (20 mM) into the LLL airways. After 30 min we determined the microvascular membrane fluid filtration coefficient (Kf) from the relationship between the rate of LLL weight gain and the pulmonary microvascular pressure. Kf was not significantly different between the lyso-Pc-treated lobes (0.048 +/- 0.018 ml.min-1.mmHg-1) vs. control lobes (0.067 +/- 0.031 ml.min-1.mmHg-1). Our data do not support the hypothesis that lyso-Pc, instilled into the airways, causes an increase in pulmonary microvascular permeability.

We estimated the pulmonary capillary membrane filtration coefficient (Kf) and the maximum capillary pressure (PCcritical) at which the lung could maintain a constant weight in 1) 5 control experiments in anesthetized open-chested dogs, 2) 7 experiments in which the dogs were given 3.6-8.3 microgram . kg-1 . min-1 of histamine phosphate, and 3) in 6 experiments after 75-100 mg/kg of alloxan. In additional experiments, pulmonary lymph flow (QL) and protein concentration (CL) were measured during the infusion of histamine and alloxan. After histamine, Kf averaged 0.045 +/- 0,008 ml . min-1mmHg-1 (SE) and PCcritical was 22.1 +/- 1.1 mmHg. These values were not significantly different from the control Kf and PCcritical (0.036 +/- 0.006 and 22.5 +/- 2.3, respectively). After alloxan, Kf (1.43 +/- 0.69) was larger and PCcritical (12.4 +/- 1.3) was significantly less than control (P less than 0.05). Histamine caused no significant change in QL or CL; however, both were increased after alloxan. These results show that Kf, PCcritical, QL, and CL are all changed by an increase in capillary membrane permeability caused by alloxan. Because none of these factors as significantly affected by histamine, dog lung capillary membrane permeability may not be affected by histamine.

Fluid accumulation within the lungs is a potentially fatal complication in critically ill patients. Sepsis and increased microvascular permeability are often implicated as the cause. This article shows that edema can be prevented by lowering systemic venous pressure (to permit pulmonary lymph to drain), by lowering pulmonary arterial pressure, and by maintaining plasma colloid osmotic pressure. It points out the importance of understanding the basic physiology behind pulmonary edema and therapeutic measures.

Pulmonary lymph drains into the thoracic duct and then into the systemic venous circulation. Since systemic venous pressure (SVP) must be overcome before pulmonary lymph can flow, variations in SVP may affect lymph flow rate and therefore the rate of fluid accumulation within the lung. The importance of this issue is evident when one considers the variety of clinical interventions that increase SVP and promote pulmonary edema formation, such as volume infusion, positive-pressure ventilation, and various vasoactive drug therapies. We recorded pulmonary arterial pressure (PAP), left atrial pressure (LAP), and SVP in chronic unanesthetized sheep. Occlusion balloons were placed in the left atrium and superior vena cava to control their respective pressures. The superior vena caval occluder was placed above the azygos vein so that bronchial venous pressure would not be elevated when the balloon was inflated. Three-hour experiments were carried out at various LAP levels with and without SVP being elevated to 20 mmHg. The amount of fluid present in the lung was determined by the wet-to-dry weight ratio method. At control LAP levels, no significant difference in lung fluid accumulation could be shown between animals with control and elevated SVP levels. When LAP was elevated above control a significantly greater amount of pulmonary fluid accumulated in animals with elevated SVP levels compared with those with control SVP levels. We conclude that significant excess pulmonary edema formation will occur when SVP is elevated at pulmonary microvascular pressures not normally associated with rapid fluid accumulation.

We studied the effect of left atrial pressure (LAP) elevation on the formation of pleural effusion in unanesthetized sheep. We prepared the animals by placing catheters in the left atrium, pulmonary artery, femoral artery, and vein. We also placed a balloon catheter in the left atrium. After a recovery period of at least 1 wk, we measured LAP, pulmonary artery pressure (PAP), systemic arterial pressure, systemic venous pressure, cardiac output, plasma protein concentration, and plasma colloid osmotic pressure (pi c). We calculated capillary pressure (Pc) as 0.5(PAP - LAP). We then elevated LAP such that Pc-pi c was between -10 and 19.5 mmHg for 6-24 h. At the end of the experiment, we killed the sheep and measured the volume and protein concentration of the right pleural effusion. We also determined the extravascular fluid to blood free dry weight of the right lung. We found that pleural effusions and pulmonary edema formed when Pc-pi c greater than 5 mmHg. We also found that the pleural effusion volume correlated with the amount of pulmonary edema. Our data show that elevated LAP may cause pleural effusions, but only after pulmonary edema has developed.

Several equations to calculate plasma oncotic pressure (pi) from the total protein concentration (C) have been previously described. These equations were derived empirically from samples with a wide range of C obtained by diluting or concentrating normal plasma samples. To test these equations over a range of naturally occurring C, we measured C and pi of plasma samples from 40 dogs. C ranged from 5.3 to 8.7 g/dl and averaged 6.5 +/- 0.1 (mean +/- SE) and pi averaged 17.9 +/- 0.3 mmHg. The regression equation was pi = 78.14 + 1.67 C (r = 0.74). pi increased with C much less than predicted with the commonly used equations. The albumin-to-globulin concentration ratios (A/G), determined in 27 of the dogs, decreased with increasing C (A/G = 1.56-0.128 C, r = 0.62). The lower A/G at the higher C's could cause the lower than predicted increase in pi with C, because the equations were developed from data in which A/G was constant.