Every weekday during a 2-mo period, all patients admitted during the preceding 24 h to the medical intensive care unit at Beth Israel Deaconess Medical Center were screened for study eligibility. Participants were identified by the parameters used to define sepsis [21]. Evidence of infection (e.g., positive cultures, typical clinical syndrome) must have been accompanied by two of the following four criteria: (1) temperature < 36 °C or > 38 °C; (2) heart rate > 90 bpm; (3) respiratory rate > 20 bpm or PaCO₂ < 32 mmHg; (4) white blood cell count > 12,000 cells/mm³, <4,000 cells/mm³, or >10% immature forms. In a prospective fashion, discarded serum samples and clinical data were collected on this cohort as well as from a randomly selected control group of patients hospitalized on the general medical service with a variety of illnesses that did not meet criteria for sepsis. All identifying information was removed; study specimens from patients and clinical data were encoded to protect the privacy of patients. Serum collected from the clinical laboratory was aliquoted in a sterile fashion and stored in cryo-vial tubes at −80 °C. The institutional review board of Beth Israel Deaconess Medical Center approved this study.

Ang-2 levels were measured in serum samples from patients by sandwich ELISA using the reagents and protocol supplied with the human Ang-2 ELISA kit (R&D Systems, Minneapolis, Minnesota, United States). Preliminary experiments confirmed the stability of Ang-2 in serum for 6–12 h at room temperature as well as its stability through several freeze–thaw cycles.

We purchased human recombinant Ang-1 and Ang-2 from R&D Systems. The RhoA-associated protein kinase inhibitor Y27632 and endothelial cell myosin light chain kinase (EC MLCK) inhibitor ML-7 were purchased from EMD Biosciences (San Diego, California, United States). Other reagents used in the experiments were obtained from Sigma (St. Louis, Missouri, United States).

Female FVB mice (obtained from Charles River, Wilmington, Massachusetts, United States) weighing 18–25 g were used throughout the study. All protocols were approved by the Beth Israel Deaconess Medical Center Animal Committee. They were acclimated to the animal facilities for at least 1 wk before the beginning of any experiment.

Human microvascular endothelial cells (HMVECs) from neonatal dermis (Cambrex Bio Science, Walkersville, Maryland, United States) were cultured in EBM-2 (Cambrex Bio Science) supplemented with 5% fetal bovine serum (FBS) and growth factors according to the manufacturer's instructions. Serum starvation was performed by incubation in 0.25% FBS/EBM-2 for 24 h.

Mice were injected with 10 μg of Ang-2 or vehicle intraperitoneal, and, after 16 h, were anesthetized with Avertin (2,2,2-Tribromoethanol). 2% Evans blue (50 μl) was then injected into the retro-orbital sinus. (In preliminary experiments with control mice, n = 6, we confirmed that the retro-orbital sinus provided a route of intravascular injection that allowed near-100% delivery of Evans blue in a reproducible fashion.) 10 min after Evans blue injection, mice were sacrificed and perfused with PBS with 2mM EDTA for 10 min through a cannula placed in the right ventricle. Blood and PBS were vented through an incision in the vena cava, thus allowing perfusate to pass through the pulmonary and systemic circulations. After 10 min of perfusing the right ventricle with PBS, the outflow from the vena cava was observed to be clear, confirming that blood (and intravascular Evans blue) had been flushed out of the circulation. Washout of intravascular contents was also confirmed histologically after 10 min of PBS perfusion. Organs were then harvested and homogenized in 1.5 ml of formamide. Evans blue was extracted by incubating the samples at 70 °C for 24 h, and the concentration of Evans blue was estimated by dual-wavelength spectrophotometer (620 nm and 740 nm). The following formula was used to correct optical densities (E) for contamination with heme pigments: E620_(corrected) = E620_(raw) − (1.426 × E740_(raw) + 0.030).

Mouse lung wet-to-dry weight ratio (W/D ratio) was used to measure lung water accumulation after Ang-2 injection. Lung wet weight was determined immediately after removal of the right lung. Lung dry weight was determined after the lung had been dried in an oven at 50 °C for 24 h. The W/D ratio was calculated by dividing the wet weight by the dry weight.

Adult mice were injected with 0, 10, or 20 μg of Ang-2 intraperitoneal and sacrificed at time zero, 3 h, and 48 h after injection. Lungs were harvested, fixed in 10% formalin, embedded in paraffin, sectioned, and stained with hematoxylin and eosin.

Cells were washed with ice-cold PBS three times and lysed with ice-cold RIPA buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, and 1 mM EDTA) supplemented with protease inhibitors (Roche Diagnostics, Indianapolis, Indiana, United States) and 1 mM NaF. Lysates were sonicated and centrifuged at 10,000 rpm for 10 min at 4 °C, and supernatants were collected. Protein concentrations were determined by Bradford protein assay with bovine serum albumin as a standard (Bio-Rad, Hercules, California, United States). A mixture of lysate, NuPAGE reducing agent, and NuPAGE sample buffer were heated at 70 °C for 10 min, electrophoresed in NuPAGE 4%–12% Novex Bis-Tris Gels (all from Invitrogen Life Technologies, Franklin Lakes, New Jersey, United States), transferred to polyvinylidene difluoride (PVDF) membrane, and immunoblotted with specific primary antibodies. Binding of primary antibodies was detected using horseradish peroxidase-conjugated secondary antibodies (Amersham Pharmacia Biotech, Piscataway, New Jersey, United States) and SuperSignal WestDura (Pierce Biotechnology, Rockford, Illinois, United States) reagents as chemiluminescence substrates. Primary antibodies were obtained from these suppliers: anti-Tie2 antibody (clone Ab33) was from Upstate Cell Signaling Solutions (Lake Placid, New York, United States); anti-GAPDH monoclonal antibody was from Chemicon International (Temecula, California, United States).

For immunoprecipitation of proteins from total cell lysates, we lysed cells with RIPA buffer, adjusting protein concentration by Bradford protein assay (Bio-Rad, Hercules, California, United States) and incubated 200 μg of total protein with anti-Tie2 antibody for 3 h, followed by incubation with protein A sepharose (Zymed, San Francisco, California, United States) for 2 h at 4 °C. After washing the beads, proteins were eluted by heating in SDS-sample buffer and detected by Western blot analysis with Anti-phospho-tyrosine (clone 4G10, Upstate Cell Signaling Solutions, Charlottesville, Virginia, United States) as described before.

After signal starvation with 0.25% FBS EBM-2 for 24 h, cells were treated with 100 ng/ml Ang-2 or vehicle for 0, 1, 3, and 6 h. MLC-p phospho-serine 19 and GAPDH were detected by Western blot analysis. For MLC-p assays conducted on HMVECs treated with serum from patients, serum Ang-2 concentration was first measured by ELISA. Then, serum was diluted to 5% with EBM-2 and filtered with low-protein-binding PVDF membrane (0.22 μm, Millipore, Bedford, Massachusetts, United States). Anti-MLC-phospho-serine-19 Ab was obtained from Abcam (Cambridge, Massachusetts, United States).

RhoA activity assay was performed and quantified using the RhoA activation assay kit according to the manufacturer's instructions (Cytoskeleton, Denver, Colorado, United States). After signal starvation with 0.25% FBS EBM-2 for 24 h, cells were treated with 100 ng/ml Ang-2 or vehicle for the indicated times (0, 30 min, 1 h, 3 h, and 6 h) and harvested into the lysis buffer. Following a brief centrifugation to remove cell debris, lysates from control and Ang-2-treated cells containing equivalent protein concentrations were rotated for 60 min with a 40-μl slurry of a GST-fusion protein composed of the Rho-binding domain of the specific RhoA effector rhotekin coupled to agarose beads. Beads were collected by centrifugation and washed three times with the lysis buffer. Beads were then re-suspended in NuPAGE reducing agent and NuPAGE sample buffer and heated at 70 °C for 10 min. The liquid phase was electrophoresed in NuPAGE 4%–12% Novex Bis-Tris Gels (Invitrogen Life Technologies). Whole-cell lysates from both control and Ang-2-treated cells were also run to determine baseline levels of total RhoA protein. Separated proteins were transferred to nitrocellulose and immunoblotted with a monoclonal antibody to RhoA (Santa Cruz Biotechnology, Santa Cruz, California, United States).

HMVECs were grown to confluence on glass coverslips coated with 1% gelatin. The cells were fixed for 10 min in 4% paraformaldehyde in PBS, and incubated for 5 min in 0.5% Triton X-100 in PBS. After blocking, the monolayers were processed for staining with anti-VE-cadherin monoclonal antibody (BD Biosciences Pharmingen, San Diego, Califiornia, United States) and Alexa Fluoro 488 goat anti-mouse IgG, rhodamine phalloidin (Molecular Probes, Eugene, Oregon, California) for F-actin staining and TOPRO-3-iodine (Molecular Probes) for nuclear staining. Fluorescence images were obtained using a Bio Rad MRC confocal fluorescence microscope. For experiments using cells treated with serum from patients, serum Ang-2 concentration was first measured by ELISA. Then, patient serum was diluted to 10% with EBM-2 and filtered with low-protein-binding PVDF membrane (0.22 μm, Millipore) prior to application on EC monolayers.

HMVEC monolayer permeability was determined with the use of FITC-labeled bovine serum albumin (Sigma) as described elsewhere [22]. Coster Transwell membranes (Corning, Corning, New York, United States) were coated with fibronectin, and cells were grown until confluence. Vehicle or Ang-2 (400 ng/ml) with FITC-albumin (final concentration 1mg/ml) was added to the luminal chamber for 8 h, and samples were taken from both the luminal and abluminal chamber for fluorometry analysis. The readings were converted with the use of a standard curve to albumin concentration. These concentrations were then used in the following equation to determine the permeability coefficient of albumin (P_a):

where [A] is abluminal concentration; t is time in seconds; A is area of membrane in cm²; V is volume of abluminal chamber; and [L] is luminal concentration.

HMVECs were seeded on 10-cm dishes for Western blot (or on 1% gelatin-coated coverslips for immunohistochemistry experiments) 24 h before experiments. 20 μmol of validated, annealed short interfering RNA (siRNA) (Ambion, Austin, Texas, United States) directed to human Tie-2 was transfected using silentFect Lipid reagent (Bio-Rad) according to the manufacturer's instructions. Cells were used for experiments 3 d after transfection. Down-regulation of Tie-2 receptor was verified by Western blotting with anti-Tie-2 polyclonal antibody (Upstate Cell Signaling Solutions).

Results are reported as mean ± standard error of the mean. Comparisons between continuous variables were performed using an unpaired two-sided t-test.

Table 1 presents baseline characteristics of the 22 patients with sepsis enrolled in this initial case cohort over a 2-mo period (September 2004 to October 2004). A control cohort of 29 patients was also selected randomly from individuals hospitalized on the general medical service with a variety of illnesses over this same time period. Serum specimens were collected prospectively along with relevant clinical data (see Methods for enrollment criteria, specimen handling, reproducibility of measurements, and specimen storage). Circulating Ang-1 was not different between cases and controls (unpublished data), but Ang-2 at the time of enrollment (Figure 1) was 23.2 ± 9.1 ng/ml among patients with severe sepsis—defined by the presence of shock or multi-organ dysfunction (third bar, n = 17)—compared with only 3.5 ± 0.6 ng/ml among non-septic hospitalized controls (first bar, n = 29, p = 0.0071) and 4.8 ± 1.5 ng/ml among patients with sepsis but who did not have shock or multi-organ dysfunction (second bar, n = 5).

Serial measurements of Ang-2 revealed that the group of individuals with severe sepsis—the one that started with the highest circulating Ang-2—developed even higher peak Ang-2 during the course of hospitalization (32.4 ± 8.7 ng/ml), whereas hospitalized patients who did not have sepsis (controls) and patients who had sepsis but were without shock or multi-organ dysfunction (mild sepsis) had Ang-2 serum levels that generally remained ≤10 ng/ml. Our control values for serum Ang-2 concentration are consistent with data reported by others [23].

Serum from Patient CH (Figure 2, —▪—) shows a typical trend of Ang-2 values in a patient with septic shock who subsequently convalesced and was discharged. This patient grew enterococcus from her urine and initially required three vasoactive agents and ventilatory support to manage shock and hypoxemia. The high peak of Ang-2 followed by progressive decline to normal values mirrored the course of her illness. Patient AP's (Figure 2, —▴—) Ang-2 trend was typical for patients who were hospitalized without sepsis. Patient AP was an elderly woman admitted to the general medicine service with an infection, but without evidence of sepsis. On the other hand, Patient AG's (—○—) trend is illustrative of Ang-2 values both during a non-sepsis-related hospitalization (left of break line) as well as a later hospitalization (right of break line) during which the patient died from complications of severe sepsis. Patient AG was first hospitalized for hypotension following excessive fluid removal during dialysis without evidence of infection, systemic inflammatory response, or respiratory compromise (Figure 2, to the left of the broken line, depicted as hospital days 1–3). During this time, his Ang-2 remained flat and <10 ng/ml. However, 3 mo later, he was re-hospitalized for emergent lower leg amputation to treat gangrene and severe sepsis (to the right of the broken line, depicted as hospital days 6–8 for purposes of illustration). This second admission was associated with abruptly rising Ang-2—the last measurement was made on the day of his death despite aggressive measures in the intensive care unit. The rising Ang-2 curve associated with Patient AG's death was typical of patients with severe sepsis who died despite aggressive measures.

Given our hypothesis that Ang-2 imbalance should preferentially affect lung permeability—where Tie-2 expression is highest [19]—we compared peak circulating Ang-2 value during hospitalization between individuals with very poor oxygenation versus those with less impaired oxygenation. PaO₂/FiO₂, the ratio of arterial blood's oxygen partial pressure (PaO₂) to the fraction of inspired air consisting of oxygen (FiO₂) [24], was used as a metric to assess the defect in oxygen absorption from the lung into the bloodstream. A cutoff of 200 in the PaO₂/FiO₂ ratio was used to segregate groups into more and less severe impairment in gas exchange because it is one of the diagnostic criteria for ARDS [5]. Indeed, Ang-2 did correlate strongly with nadir impairment in gas exchange—41.95 ± 12.44 ng/ml among those patients with PaO₂/FiO₂ < 200 versus 11.22 ± 2.44 ng/ml in the group of individuals with better oxygenation (PaO₂/FiO₂ > 200); p = 0.02 (Figure 3A). Ang-2 did not correlate with survival to discharge nor to level of critical illness at enrollment as defined by APACHE II (acute physiology and chronic health evaluation) (Figure 3B and 3C). Ang-2 also did not correlate with a history of congestive heart failure (Figure 3D). There was also no correlation of Ang-2 with an underlying history of lung disease, renal failure, or cancer (unpublished data). Of note, the one patient in our cohort who did meet full criteria for ARDS had Ang-2 = 139 ng/ml, the highest level measured in our group. Circulating Ang-2 elevation, therefore, appeared to correlate specifically with a low PaO₂/FiO₂ ratio.

Separation of adjacent ECs from one another leads to paracellular gap formation—a process driven by actin–myosin-based cell contraction [25–27]. Such gaps permit para-endothelial movement of macromolecules and, thus, represent a structural change that correlates with hyperpermeability. To test what structural effect human serum had on cultured ECs, we added serum from two patients, Patient CE4 (high Ang-2) and Patient CF1 (low Ang-2), to HMVECs and stained for F-actin and VE-cadherin, a structural protein that helps maintain intercellular junctions. Incubation of HMVECs with control medium (FBS/culture medium) resulted in a compact, confluent cell layer with thin actin filaments and localization of VE-cadherin to cell–cell junctions (Figure 4A–4C). However, addition of high Ang-2 serum (CE4, Ang-2 = 89 ng/ml) induced thick actin stress fibers and intercellular gap formation (Figure 4D–4F, arrows) whereas low Ang-2 serum (CF1, Ang-2 = 8.9 ng/ml) did not (Figure 4G–4I). The gap formation provoked by the serum from Patient CE4 was reversed with addition of recombinant human Ang-1 (Figure 4J–4L).

To address the potential biasing effect of unmeasured confounders between sera from two different patients, we repeated this experiment with serum from one patient taken at two time points during his hospitalization. Serum from Patient CG was collected on hospital day 2 (CG2, Ang-2 = 78 ng/ml), and on hospital day 16 (CG12, Ang-2 = 6.3 ng/ml). On hospital day 2, Patient CG had PaO₂/FiO₂ = 56, was in septic shock, and had ARDS; by hospital day 16, Patient CG was extubated, convalescing uneventfully and preparing for discharge. Serum from CG's hospital day 2 (Figure 4, CG2) induced gap formation and thick actin stress fibers (Figure 4M–4O), effects not seen with his serum at discharge (Figure 4P–4R, CG12); moreover, effects of high Ang-2 serum from hospital day 2 (Figure 4, CG2) were reversed with the addition of Ang-1 (Figure 4S–4U).

These results illustrate (a) the presence of a serum activity during severe sepsis that induces endothelial barrier disruption; (b) that clinical resolution correlates with falling Ang-2 and decreased barrier-disrupting activity; and (c) that this activity can be reversed with Ang-1, suggesting that Ang-2 in the serum of human patients is at least partially responsible for altering endothelial architecture in sepsis.

Having observed this effect of serum from human patients on cultured ECs, we next tested whether Ang-2 alone could reproduce disruption of endothelial architecture. Recombinant human Ang-2 (100 ng/ml) was added to HMVECs, which were subsequently stained for F-actin and VE-cadherin. As suspected, Ang-2 induced the formation of thick actin stress fibers and intercellular gaps (Figure 5Ad–Figure 5Af, arrows), effects not seen with vehicle incubation (Figure 5Aa–Figure 5Ac). This experiment confirmed the hypothesis raised by the results in Figure 4—namely that Ang-2 alone could provoke potentially pathologic structural changes in endothelium.

After establishing that Ang-2 could promote pathologic structural changes in endothelial monolayers, we next asked whether Ang-2 could modulate barrier function. To determine the effect of Ang-2 on monolayer permeability, we monitored the clearance of FITC-labeled albumin across an HMVEC monolayer with and without Ang-2 stimulation. Figure 5B shows that Ang-2 stimulation for 8 h increased permeability by approximately 20% compared with the control (p < 0.01).

Since Ang-2 appeared to be a likely mediator of endothelial barrier disruption in serum from humans with sepsis, we next sought to understand the intracellular mechanism through which Ang-2 could distort endothelial shape and cell–cell contacts.

Endothelial barrier function is known to be tightly regulated by myosin-driven cellular contraction [8,28–30]. For contraction to occur, MLC must be phosphorylated at Ser-19 by EC MLCK, and phosphorylation of Ser-19 by EC MLCK is needed to activate actomyosin ATPase function [31,32]. We therefore hypothesized that Ang-2 up-regulated MLC-p.

Initially, we tested the effect of serum from humans with sepsis on MLC-p. Serum was taken from the same patients used for immunohistochemistry in Figure 4D–4I—CE2 (Ang-2 = 77 ng/ml) and CF5 (Ang-2 = 7.9 ng/ml)—and added at 1:20 dilution to 24-h serum-starved HMVECs. The high Ang-2 serum (CE2) caused MLC-p that was inhibited by addition of Ang-1, whereas the low Ang-2 serum led to markedly less MLC-p (Figure 6A).

Next, we tested the effect of Ang-2 alone (100 ng/ml) on 24-h serum-starved HMVECs at serial time points. We observed that MLC-p was elevated at 3 h and 6 h of stimulation (Figure 6B) and persisted for 24 h (unpublished data).

Rho-GTPases play a pivotal role in the control of cellular actin rearrangement and cell shape [33]. Rho-kinase, a downstream target of RhoA, stimulates stress fiber formation by up-regulating MLC-p through two mechanisms: activation of EC MLCK and inhibition of myosin phosphatase activity [34,35]. Given this 2-fold effect of RhoA on MLC, one would predict that a RhoA inhibitor would be even more potent than an MLCK inhibitor at blocking MLC-p. We asked if Ang-2-induced MLC-p required activated RhoA. We tested this possibility by treating 24-h-serum-starved HMVECs with Ang-2 (100 ng/ml) for different time periods (0, 30 min, 1 h, 3 h, and 6 h) and then by measuring RhoA activation. Indeed, Ang-2 induced an increase in the active form of RhoA (Rho-GTP) (Figure 6C) that peaked between 30 min and 1 h. Pre-treatment of Ang-2-stimulated HMVECs with a specific inhibitor of Rho-kinase (Y27632, 10 μM) completely abolished Ang-2-induced MLC-p (Figure 6D, third column) while an EC MLCK inhibitor (ML-7, 10μM) partially inhibited MLC-p (Figure 6D, fourth column). These results demonstrate that Ang-2 mediates MLC-p in a RhoA-dependent fashion in human microvascular ECs.

Given that Ang-2 induces gap formation between ECs (Figure 5) and that Ang-2 stimulates MLC-p through Rho-kinase (Figure 6B–6D), we next asked whether Ang-2′s gap-formation effect occurred through Rho-kinase and MLCK. Y27632 (10 μM) completely reversed the formation of thick actin stress fibers and paracellular gaps induced by Ang-2 (Figure 6Ea–Figure 6Ef). ML-7 (10 μM) partially reversed the Ang-2-induced structural changes on actin and cell junctions (Figure 6Eg–Figure 6Ei), consistent with the less potent effect of ML-7 versus Y27632 on MLC phosphorylation (Figure 6D). These results demonstrate that the deleterious structural effects of Ang-2 on ECs are mediated through Rho-kinase and MLCK.

Multiple lines of evidence suggest that Ang-1 and Ang-2 are an agonist–antagonist pair at the Tie-2 receptor [36]. Ang-1 activates Tie-2, leading to receptor phosphorylation and subsequent signal transduction that promotes EC survival and vessel assembly. Ang-2, on the other hand, is believed to act as a Tie-2 ligand that competitively binds the receptor and interferes with agonistic Ang-1/Tie-2 functions—i.e., Ang-2 binding of Tie-2 blocks Tie-2 phosphorylation. To further delineate the pathway through which Ang-2 promotes EC contraction and gap formation, we first confirmed that Ang-2 (100 ng/ml) inhibited phosphorylation of Tie-2 receptor in HMVECs (Figure 7A).

We then assessed the effect of Tie-2 signaling on MLC-p by using siRNA against Tie-2 receptor (Tie-2-siRNA). Tie-2-siRNA induced robust MLC-p (Figure 7B, middle blot), recalling the effect seen with Ang-2 treatment (Figure 6B). Tie-2-siRNA caused a spindle phenotype (Figure 7Ce), thick actin stress fibers and paracellular gap formation (Figure 7Cf–Figure 7Ch, arrows), effects not observed with negative control siRNA transfection (Figure 7Ca–Figure 7Cd). These morphologic changes are on the same spectrum as, but even more severe than, those seen with the addition of Ang-2 (Figure 5Ad–Figure 5Af). These results suggest that Tie-2 signaling is constitutively active in this system. Addition of Ang-2 blocks Tie-2 signaling, leading, in turn, to Rho-kinase activation and MLC-p, with the end result being EC contraction, gap formation, and disruption of barrier integrity.

So far, we had identified Ang-2 as an important circulating factor in serum from humans with sepsis that distorts the endothelial barrier, and we had determined that this action was carried out by Tie-2 blockade and activation of Rho-kinase and MLCK. To establish the functional importance of maintaining this barrier, we hypothesized that systemic administration of Ang-2 would provoke pulmonary vascular hyperpermeability and congestion.

Evans blue avidly binds to serum albumin and can therefore be used as a tracer for trans-capillary flux of macromolecules. The extravasation of Evans blue has frequently been employed to quantify in vivo vascular permeability [37,38]. Given the several limitations of in vitro permeability assays—e.g., lack of flow and variable hydrostatic pressure, use of cultured cells, absence of microenvironment, and absence of interacting cell types such as neutrophils [39]—we felt that it was important to confirm leak across intact blood vessels in an in vivo setting. Adult mice were pre-treated with either vehicle or Ang-2 (10 μg) injected intraperitoneally prior to Evans blue dye injection. Although no mice in either group exhibited any signs of sickness (e.g., motionlessness, piloerection, diarrhea) 16 h after treatment, spectrophotometric quantification of extravasated dye showed enhanced leakage, with a 3-fold increase in lungs and a 2-fold increase in livers of Ang-2-treated mice compared with vehicle-treated mice (Figure 8A) (p < 0.01). Intestines were also tested and showed a statistically non-significant trend toward increased permeability (Figure 8A).

After washout of intravascular Evans blue by perfusing PBS through the right ventricle and venting from the vena cava, lungs of vehicle-treated mice were blanched-appearing (Figure 8B, left panel); however, lungs of Ang-2-treated mice appeared more congested and purple-tinted (Figure 8B, right panel), suggesting retention of dye in the extravascular space. In addition, the lung W/D weight ratio increased from 5.01 ± 0.26 to 6.13 ± 0.03 with Ang-2 treatment (Figure 8C) (p < 0.01), suggesting increased lung water accumulation following Ang-2 administration.

Lung sections were also taken for histologic characterization. Figure 9A shows lung from a control mouse injected with vehicle—alveolar septa form a fine, thin network (inset on lower right). 3 h after systemic Ang-2 administration (10 μg), there is an increase in cellularity, congestion, and early extravasation into air spaces (Figure 9B). These changes are even more pronounced at 48 h (Figure 9C) following a total Ang-2 dose of 20 μg. These results establish an in vivo causative role for Ang-2 in pulmonary vascular leak and further substantiate the in vitro permeability effects observed earlier with Ang-2 stimulation (Figure 5). Moreover, progressive lung injury appears to occur with increasing amount and duration of systemic Ang-2 exposure.