Pulmonary arterial smooth muscle cells modulate cytokine- and LPS-induced cytotoxicity in endothelial cells

Bruce A. Johnson1, Bruce R. Pitt2, and Paul Davies3

1 Division of Pulmonary, Allergy and Critical Care Medicine, Department of Medicine, and 2 Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261; and 3 DuPont Pharmaceutical, Wilmington, Delaware 19880


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cytokines and lipopolysaccharide (LPS) are known to be injurious to vascular endothelial cells (ECs), but the influence of adjacent vascular smooth muscle cells (SMCs) on this injury is unknown. Exposure of cultured rat (RPMECs) or human (HPMECs) pulmonary microvascular ECs on tissue culture plastic to a mixture of cytokines (interleukin-1beta , tumor necrosis factor-alpha , and interferon-gamma ) and LPS (cytomix) resulted in a significant increase in 51Cr release to 35-40%. When unstimulated RPMECs were cocultured with cytomix-pretreated rat pulmonary microvascular SMCs (RPMSMCs) there was an increase in 51Cr release to 8.4%, which was nitric oxide dependent. However, when RPMECs or HPMECs were stimulated in direct contact with their respective SMCs, rather than a further increase in cytomix-induced injury (e.g., >35-40%), 51Cr release decreased to <10%. This cytoprotection was fully reproduced with fixed RPMSMCs, and partially reproduced by plating HPMECs on gelatin. These data show that the direct toxicity of a cytokine and endotoxin mixture on cultured ECs can be reduced by contact with vascular smooth muscle.

pulmonary microcirculation; pulmonary endothelium; pulmonary vascular smooth muscle; nitric oxide; human; rat; cytokines; endotoxin; lipopolysaccharide


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

CYTOKINES AND BACTERIAL products such as endotoxin are well known to have complex effects on endothelial cells (ECs). In addition to activation of various EC metabolic pathways, under certain conditions, tumor necrosis factor-alpha (TNF-alpha ) (26, 35), interleukin-1beta (IL-1beta ) (21), or combinations of these and other cytokines or lymphokines [interferon-gamma (IFN-gamma )] can be toxic to cultured ECs (6, 23, 43). Important roles for cytokines and endotoxin have been assigned in the pathogenesis of adult respiratory distress syndrome because these agents are known to cause injury to cultured pulmonary ECs (5, 9, 26-28) as well as to pulmonary ECs in vivo (18). Furthermore, recent evidence has shown that IL-1beta (41) and TNF-alpha (36) activate cultured human pulmonary artery ECs and endotoxin is cytotoxic to these cells (25).

The mechanisms underlying cytokine or endotoxin activation and/or cytotoxicity of pulmonary ECs are unclear, but several studies have identified important roles for partially reduced oxygen (29, 36, 46) and nitrogen (8, 9) species. Although ECs themselves can be the source of these toxic intermediates (12, 37), it is clear that other effector cells contribute to EC activation or injury. In this regard, most research has focused on the contribution of neutrophils to cytokine- and/or endotoxin-induced EC injury (43), including recent studies on human neutrophil and human pulmonary artery EC injury (14, 47). The human monocyte also appears to be an important source of reactive nitrogen species in mediating injury to human pulmonary endothelium (14), and sessile pulmonary intravascular macrophages are also important effector molecules in the response of the lung of various species to cytokines or endotoxin (40). In contrast, considerably less is known regarding an effector role of vascular smooth muscle, the other major residential cell near the endothelium. Previous work from our group has shown that rat pulmonary vascular smooth muscle synthesizes inducible nitric oxide synthase (iNOS)-derived nitric oxide (NO) (17, 32), superoxide anion (O-2), and peroxynitrite (3) in response to cytokines and endotoxin (e.g., cytomix) or IL-1beta alone in sufficient quantities to have an adverse effect on cocultures of rat pulmonary artery ECs (42).

In the current study, we pursued the role of pulmonary vascular smooth muscle-EC interaction and tested the following hypotheses: 1) cytokine stimulation is cytotoxic to human and rat lung ECs, 2) cytokine-stimulated production of reactive oxygen and nitrogen species by pulmonary vascular smooth muscle cells (SMCs) is toxic to adjacent ECs, and 3) direct EC-SMC interaction or matrix contact reduces the sensitivity of cultured ECs to cytokine-induced toxicity.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Adult male Sprague-Dawley rats were obtained from Charles River Breeding Laboratories (Wilmington, MA). Iron oxide-containing latex microspheres were custom-manufactured by Bangs Laboratories (Carmel, IN). Agarose was purchased from Bio-Rad (Richmond, CA). DMEM was supplied by JRH Biosciences (Lenexa, KS). Heat-inactivated low-endotoxin fetal bovine serum (FBS) was obtained from HyClone Laboratories (Logan, UT). Cell culture flasks, multiwell cell culture plates, and permeable 0.4-µm pore size, polyester cell culture inserts (Transwell Clear) were obtained from Costar (Cambridge, MA). Hanks' balanced salt solution (HBSS), trypsin, phenol, EDTA, and recombinant murine IFN-gamma were purchased from GIBCO BRL (Grand Island, NY). HEPES, penicillin, streptomycin, collagenase, MCDB-131 cell culture medium, mouse monoclonal anti-smooth muscle alpha -actin, mouse monoclonal anti-smooth muscle myosin, tetramethylrhodamine B isothiocyanate-conjugated rabbit anti-mouse IgG, superoxide dismutase (SOD), lipopolysaccharide (LPS) from Escherichia coli 0111:B4, N-(1-napthyl)ethylenediamine hydrochloride, NG-monomethyl-L-arginine (L-NMMA), N-sulfanilimide, Aspergillus nitrate reductase, sodium nitrite, sodium nitrate, NADPH, gelatin, BSA, Triton X-100, and rabbit serum were purchased from Sigma (St. Louis, MO). LabTek cell culture chamber slides were obtained from Nunc (Naperville, IL). DMSO, methanol, chloroform, isoamyl alcohol, and phosphoric acid were supplied by Fisher Scientific (Pittsburgh, PA). Paraformaldehyde was purchased from Polysciences (Warrington, PA). Recombinant murine TNF-alpha was supplied by Genzyme (Cambridge, MA). Acetylated low-density lipoprotein labeled with 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI-Ac-LDL) was purchased from Biomedical Technologies (Stoughton, MA). Sodium [51Cr]chromate was supplied by DuPont-NEN (Boston, MA). Recombinant human IL-1beta was manufactured by DuPont and distributed at no cost through the National Cancer Institute Biological Response Modifiers Program (Frederick, MD). Polyclonal anti-human platelet EC adhesion molecule (PECAM) antibody (1) was a gift from Dr. S. M. Albelda (Philadelphia, PA). A spontaneously transformed line of rat pulmonary microvascular ECs (RPMECs) (Ref. 7) was a gift from Dr. D. Langleben (Montreal, Quebec). Human pulmonary microvascular ECs (HPMECs) and human pulmonary artery SMCs (HPASMCs) and their respective media and subculture reagents were obtained from Clonetics (Walkersville, MD). Antibodies were obtained as follows: transforming growth factor-beta (TGF-beta ; R&D, Minneapolis, MN); L-selectin, very late antigen-4 (VLA-4), and beta 2-integrin (PharMingen, San Diego, CA); sialyl LewisX and sialyl LewisA (Kamiya, Seattle, WA); laminin, fibronectin, and collagen, (Chemicon, Temecula, CA).

Cell Culture

Cells cultured from the pulmonary microcirculation of rats were used in the experiments because the responses of both pulmonary ECs (26) and SMCs (17, 32) to cytokine stimulation differ depending on whether they are harvested from large conduit vessels or microvessels.

RPMECs were grown in MCDB-131 supplemented with 10% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin (EC medium). Cells were subcultured with trypsin (0.05%) and EDTA (0.02%) in HBSS and at a ratio of 1:5. EC identity was verified by the visualization under light microscopy of a contact-inhibited monolayer of cells with cobblestone morphology at confluence, uptake of DiI-Ac-LDL, anti-PECAM staining by indirect immunofluorescence microscopy as described (1), and lack of staining for smooth muscle-specific isoform of alpha -actin by indirect immunofluorescence microscopy. For experiments, RPMECs were plated at a density of 5 × 104 per 12-well dish

Rat pulmonary arterial microvascular SMCs (RPMSMCs) were grown from intra-acinar microvessel (<100-µm-diameter) explants as described (17) with the exception that a suspension of 22-µm-diameter paramagnetic latex microspheres was used to fill the pulmonary arterial circulation rather than iron oxide in suspension. In brief, adult male Sprague-Dawley rats were anesthetized with 50 mg/kg pentobarbital sodium, a thoracotomy was performed, and the pulmonary artery was flushed with PBS and subsequently filled with a suspension of paramagnetic microspheres in 0.5% agar. The air spaces are filled with agar, the subpleural margins were harvested and partially collagenase digested, and the microvessels were separated magnetically. The microvessels are then resuspended in 2-4 ml of culture medium (DMEM supplemented with 20% FBS, penicillin, streptomycin, and 20 mM HEPES), plated in 25-cm2 cell culture flasks at 2 ml of vessel suspension per flask, and incubated in humidified air with 5% CO2 at 37°C. A confluent layer was generally achieved by days 7-10, following which the cells were detached with trypsin (0.05%) and EDTA (0.02%) in HBSS, subcultured at a ratio of 1:3, and used for experiments between passages 3 and 10. Smooth muscle identity of the cells was verified by characteristic appearance on phase-contrast microscopy, indirect immunofluorescent antibody staining for smooth muscle-specific isoforms of alpha -actin and myosin, and lack of uptake of DiI-Ac-LDL. For experiments, RPMSMCs were plated at a confluent density of 2 × 105 per 12-well dish, which provided a RPMSMC-to-RPMEC ratio of 4:1.

HPMECs and HPASMCs were propagated as per the manufacturer's instructions. The media are proprietary, and the precise constituents are not defined, but the HPMEC growth medium consists of a basal medium, which is a modified MCDB-131 plus antibiotics, 5% FBS, and unstated amounts of hydrocortisone, vascular EC growth factor, epidermal growth factor, fibroblast growth factor, EC growth supplement, ascorbate, insulin-like growth factor, and heparin. For cytotoxicity experiments, HPMECs were plated at 2.5 × 105 cells/well and stimulated in EC basal medium with 0.1% BSA.

Nitrite/Nitrate Analysis

To verify NO production, nitrite and nitrate, the stable end products of spontaneous NO oxidation were measured by the Griess reaction after nitrate was first reduced to nitrite with the enzyme nitrate reductase (13). In brief, 100 µl of cell culture supernatant were mixed with a 40-µl solution of nitrate reductase, NADPH, and FAD in potassium phosphate buffer, and incubated for 30 min in the dark at room temperature. Specifically, 100 µl of cell culture supernatant or 140 µl of reduced supernatant mixture were mixed with an equal volume of Griess reagent [0.1% N-(1-napthyl)ethylenediamine hydrochloride freshly mixed 1:1 (vol/vol) with 1% sulfanilimide in 5% phosphoric acid] with the development of a colored product whose absorbance was read immediately at 550 nm with distilled water as a blank. Nitrite and nitrate concentrations were calculated from sodium nitrite and nitrate standard curves.

51Cr Release EC Cytotoxicity Assay

RPMEC cytotoxicity was measured by 51Cr release from RPMECs preloaded before coculture with RPMSMCs or cytokine stimulation. In brief, confluent monolayers of RPMECs were incubated overnight in EC medium with 200 µCi of 51Cr per 25-cm2 cell culture flask, rinsed three times with HBSS, trypsinized, and plated at 5 × 104 per well in EC medium either on permeable supports or directly in cell culture dishes described below. ECs were allowed 3 h to attach and spread, after which the EC medium was aspirated and the cell cocultures were rinsed and incubated a further 24 h with the medium conditions described in Experimental Protocols. The medium was then collected and gamma emission was detected in a LKB-Wallac 1272 gamma counter. Each set of experiments included control wells with untreated cells to measure the spontaneous rate of 51Cr release and a maximum release well in which cells were lysed by the addition of 50 µl of 20% (vol/vol) Triton X-100 per ml of cell culture supernatant (1% final Triton X-100 concentration). An injury index, which reflects the percent of 51Cr released after subtracting the spontaneous release (usually 25-30% of the total), was calculated as follows
injury index

= <FR><NU>(experimental<SUP> 51</SUP>Cr release − spontaneous<SUP> 51</SUP>Cr release)</NU><DE>(maximum<SUP> 51</SUP>Cr release − spontaneous<SUP> 51</SUP>Cr release)</DE></FR>

× 100
To control for any possible differences in plating efficiency, a separate maximum 51Cr release determination was made for each set of experiments and each type of plating surface. By the definition of the formula, the injury index in control cells is zero.

Experimental Protocols

Cells for 51Cr release cytotoxicity were stimulated as described below for 24 h either separately or in coculture with a mixture of the following cytokines and LPS referred to as "cytomix": human IL-1beta (50 U/ml), LPS from E. coli 0111:B4 (10 µg/ml), rat IFN-gamma (100 U/ml), TNF-alpha (500 U/ml) in DMEM supplemented with penicillin (100 U/ml), streptomycin (100 µg/ml), and 0.1% BSA.

Cytomix stimulation of RPMECs and HPMECs on tissue culture plastic. To verify that cytokine and LPS stimulation cause toxicity and cell lysis in cultured rat and human lung ECs, ECs were 51Cr loaded, plated into 12-well plastic dishes, and stimulated 3 h later with cytomix. Injury index was calculated from the 51Cr release after 24 h.

Cytomix-induced cytotoxicity in RPMECs on plastic with NO and O-2 inhibition. To evaluate the role of NO and O-2 after cytokine stimulation on tissue culture plastic in the absence of SMCs, RPMECs were plated onto untreated cell culture plastic and stimulated 3 h later with cytomix or control medium and the NO synthase inhibitor 1.0 mM L-NMMA, 100 U/ml SOD, or both. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

Cytotoxicity to unstimulated RPMECs after coculture with cytomix-prestimulated RPMSMCs and inhibition of NO, O-2, or both. Knowing that cytokine-stimulated SMCs generate NO and O-2, we wanted to test the hypothesis that cytokine-activated RPMSMCs might injure adjacent RPMECs by producing toxic amounts of NO. Because cytokines are directly toxic to ECs, we separated the effect of cytokine activation of RPMECs from that of cytokine activation of RPMSMCs by first cytomix stimulating a monoculture of RPMSMCs for 24 h, rinsing off the cytokines, and then plating unstimulated RPMECs onto the activated layer of RPMSMCs. The RPMECs were allowed 3 h to attach and spread, after which the EC medium was replaced with DMEM with 0.1% BSA (control medium) with and without 1.0 mM L-NMMA, 100 U/ml SOD, or both. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

Cytotoxicity to unstimulated RPMECs after coculture with cytomix-prestimulated RPMSMCs and physical separation of the cells. Because NO, O-2, and other related reactive oxygen species (ROS) have relatively short diffusion distances, we suspected that any cytotoxic effect of activated RPMSMCs on RPMECs would be attenuated by increasing the distance between the two cells. Accordingly, RPMECs were plated onto a permeable support suspended 1.0 mm above cytokine-prestimulated RPMSMCs. When RPMECs are separated from RPMSMCs, they should no longer be NO sensitive, so a series was included with and without L-NMMA. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

Cytomix stimulation of previously established RPMEC-RPMSMC cocultures with inhibition of NO, O-2, or both. To test whether concomitant cytokine activation of RPMECs alter their response to RPMSMC-derived NO, O-2, or both, RPMEC-RPMSMC cocultures were first established by plating RPMECs onto unstimulated RPMSMCs. After 3 h were allowed for RPMECs to attach, the cocultures were stimulated with cytomix or control medium and 1.0 mM L-NMMA, 100 U/ml SOD, or both. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

RPMEC cytotoxicity after culture in conditioned medium from cytomix-stimulated RPMSMCs. To determine whether RPMEC cytoprotectivity conferred by RPMSMCs was due to the elaboration of stable soluble mediators, RPMSMCs were stimulated with control media, cytomix, cytomix plus L-NMMA, or L-NMMA alone for 24 h. The media were then aspirated, any detached cells were pelleted, and the media were applied to ECs plated onto plastic 3 h earlier. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

Cytomix-induced cytotoxicity in RPMECs cultured on RPMSMC extracellular matrix or fixed layers of RPMSMCs. To determine whether RPMEC cytoprotectivity conferred by RPMSMCs was mediated by RPMSMC matrix or surface antigens, matrix and fixed layers of RPMSMCs were prepared as below, RPMECs were plated at 5 × 104 per well, allowed 3 h to adhere, and then stimulated with cytomix or control media. Supernatants were harvested 24 h later for nitrite and nitrate analysis and gamma counting.

RPMSMC extracellular matrix (ECM) was prepared as described (7). RPMSMCs were plated into 12-well dishes at confluent density and returned to the incubator for 48 h. The dishes were then rinsed with PBS, the cells were removed with 0.5% Triton X-100 in MilliQ water for 15 min at room temperature, and the residual ECM was rinsed three times with sterile MilliQ water. The dishes with ECM were then either used immediately or stored for up to 4 h in PBS at 4°C.

Paraformaldehyde-fixed RPMSMC layers were prepared by plating and rinsing RPMSMCs as described in Cell Culture, followed by application of freshly prepared 4% paraformaldehyde in PBS for 15 min at room temperature. The layers were then rinsed with PBS and incubated with 0.1% glycine in PBS for 5 min at room temperature to quench unreacted paraformaldehyde, following which the layers were rinsed three times with PBS and either used immediately or stored for up to 4 h in PBS at 4°C.

Methanol-fixed RPMSMC layers were prepared by plating and rinsing RPMSMCs as described in Cell Culture followed by fixing with 4°C methanol for 10 min. The layers were then rinsed three times with PBS and either used immediately or stored for up to 4 h in PBS at 4°C.

To try and identify cell surface receptor/ligands mediating the cytoprotection, the fixed cell matrix studies were repeated with antibodies directed against TGF-beta , L-selectin, sialyl LewisX, sialyl LewisA, beta 2-integrin, VLA-4, laminin, fibronectin, and collagen. Antibodies were applied at the minimum dilution recommended by the vendor or empirically at 100:1 if no suggestions were given in PBS with 0.1% BSA overnight at 4°C. The antibody solutions were then aspirated, the layers were rinsed once with PBS, and 51Cr-loaded RPMECs were plated onto them and stimulated with cytomix as in Cytomix stimulation of RPMECs and HPMECs on tissue culture plastic. The antibodies were chosen based on commercial availability and vendor reported cross-reactivity with rat antigens, with the exception of the Lewis antibodies where rat binding had not been specifically documented.

Cytomix-induced cytotoxicity in HPMECs cultured on HPASMCs or a collagen matrix. To investigate the hypothesis that EC-SMC contact or ECM contact would decrease cytokine-stimulated toxicity, additional ECs were plated onto confluent layers of SMCs or gelatin-coated dishes (HPMECs only). For gelatin coating of 12-well dishes, 0.5 ml of 2% gelatin solution was allowed to adhere in the incubator overnight, after which the solution was aspirated and the cells were plated as in Cytomix stimulation of RPMECs and HPMECs on tissue culture plastic.

Statistical Analysis

Results are expressed as the means ± SE. In each experiment, gamma counting was performed in duplicate and nitrite plus nitrate was analyzed in triplicate. Each data point reflects the mean of at least three separate experiments. Group means were compared by ANOVA. If the overall F statistic was significant at the 0.05 level, then subsequent intergroup significance testing was accomplished by Scheffé's method, with P < 0.05 considered significant. Inspection of the frequency histograms for some data groups gave cause to question the assumption of normality, and in these cases, overall comparison of multiple group means was accomplished with Kruskal-Wallis ANOVA by ranks. If the overall H statistic was significant at the 0.05 level, subsequent intergroup significance testing was performed as described (24), with P < 0.05 considered significant. For comparison of experimental injury index with untreated controls, which have an injury index of zero by definition, one-sample t-tests were performed, with P < 0.05 considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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Cytomix Stimulation Is Toxic to Rat and Human Pulmonary ECs

Direct cytomix stimulation of isolated RPMECs resulted in toxic changes as evident by increased numbers of rounded or floating cells and contraction of remaining adherent monolayer (Fig. 1). Similar structural changes were noted in isolated HPMECs exposed to cytomix (data not shown). Exposure to cytomix caused a significant increase in injury index as determined by 51Cr release to 35 ± 3 and 41 ± 2% in RPMECs (n >=  4) and HPMECs (n = 3), respectively. Cytomix-induced injury to RPMECs was not sensitive to 1 mM L-NMMA, 100 U/ml SOD, or the combination (Fig. 2).


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Fig. 1.   Phase-contrast appearance of control and cytomix-stimulated rat pulmonary microvascular endothelial cells (RPMECs). RPMECs were plated onto cell culture plastic and subsequently exposed to control medium (A) or 50 U/ml cytomix [CTM; human interleukin-1beta (IL-1beta ), 10 µg/ml lipopolysaccharide (LPS) from E. coli 0111:B4, 100 U/ml rat interferon-gamma (IFN-gamma ), and 500 U/ml tumor necrosis factor-alpha (TNF-alpha )] (B). Magnification ×80.



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Fig. 2.   Cytomix-stimulated cytotoxicity in monocultured RPMECs was not changed by NG-monomethyl-L-arginine (L-NMMA) or superoxide dismutase (SOD). Unstimulated RPMECs were 51Cr loaded, plated into culture dishes, and stimulated with CTM, CTM + L-NMMA, CTM + SOD, or CTM + SOD and L-NMMA. Injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. control; ** P < 0.05 vs. CTM; n >=  4.

Cytomix-Prestimulated Cultured RPMSMCs Are Cytotoxic to Cocultured, Unstimulated RPMECs

Plating unstimulated RPMECs directly onto a cytomix-prestimulated layer of RPMSMCs increased their injury index to 8.4% at 24 h (Fig. 3). The injury was eliminated when the RPMECs were separated from the RPMSMCs by 1.0 mm in a Transwell coculture configuration (Fig. 3). Inhibition of NO synthesis with L-NMMA decreased injury index, and the degree of injury was directly proportional to the magnitude of cytomix-induced NO synthesis (Fig. 4). Exogenous SOD, however, did not affect cytomix-induced injury index nor interact with L-NMMA to affect the residual injury (Fig. 5).


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Fig. 3.   Cytomix-prestimulated rat pulmonary microvascular smooth muscle cells (RPMSMCs) were cytotoxic to unstimulated, cocultured RPMECs. Unstimulated RPMECs were 51Cr loaded and plated either directly (direct culture, solid bars) onto RPMSMCs prestimulated with control medium or CTM with and without L-NMMA (1.0 mM) or onto permeable supports suspended 1.0 mm above correspondingly stimulated RPMSMC layers (open bars). Injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. control; ** P < 0.05 vs. CTM direct culture; n >=  4.



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Fig. 4.   Nitric oxide (NO) production by cytomix-stimulated RPMSMCs was positively correlated with cytotoxicity in cocultured RPMECs. Unstimulated RPMECs were 51Cr loaded and plated directly onto cytomix-prestimulated RPMSMCs, and 24 h later nitrate plus nitrite was measured and injury index was calculated from 51Cr release. Cytomix-stimulated nitrate plus nitrite values for individual experiments varied spontaneously and were plotted against corresponding injury index. Regression equation was y = 0.122x + 3.43, r2 = 0.936, and P < 0.001.



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Fig. 5.   RPMEC cytotoxicity from cocultured RPMSMCs prestimulated with cytomix was decreased by L-NMMA but not by SOD. Unstimulated RPMECs were 51Cr loaded and plated onto RPMSMC layer prestimulated with CTM, CTM + L-NMMA, CTM + L-NMMA + SOD (100 U/ml), or CTM + SOD. Injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. control; ** P < 0.05 vs. CTM; n >=  4.

Contact With RPMSMCs Protects Cultured RPMECs Against Cytomix-Induced Injury

We assumed that exposure of ECs simultaneously to cytokine mixture and cytomix-stimulated vascular SMCs would be more toxic than either stimulus alone. In contrast, we noted that coculture of RPMECs with either prestimulated RPMSMCs or simultaneous exposure of a coculture of RPMECs and RPMSMCs significantly decreased the cytotoxicity of cytomix to RPMECs alone (Fig. 6). Cytomix-stimulated NO production by prestimulated RPMSMCs continued in the 24 h after removal of the cytokines and was not significantly different from NO production when RPMSMC-RPMEC cocultures were stimulated simultaneously (Table 1).


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Fig. 6.   Cocultured RPMSMCs protected RPMECs against cytomix-induced cytotoxicity. Unstimulated RPMECs were 51Cr loaded and plated onto CTM-prestimulated RPMSMCs or onto cell culture plastic or unstimulated RPMSMCs. Cells plated onto plastic or unstimulated RPMSMCs were subsequently CTM stimulated. Injury index was calculated from 51Cr release 24 h later. Comparison shown by arrows was not significant (ns). * P < 0.05 vs. either coculture; n >=  7.


                              
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Table 1.   Supernatant nitrate plus nitrite concentration for cytomix-stimulated RPMSMC-RPMEC cocultures and RPMECs cultured alone for 24 h

RPMECs do not produce significant increases in Griess-detectable amounts of nitrite and nitrate following cytomix stimulation. Indeed, NO itself did not appear to contribute to this protection because the reduction in injury index from 35 to 40% left an L-NMMA (and SOD)-insensitive residual injury of 6-8% (Fig. 7). Furthermore, the reduction in cytomix-induced RPMEC injury when these cells were cocultured with RPMSMCs was not apparent when conditioned medium only was used from cytomix-stimulated RPMSMCs (with or without L-NMMA) as shown in Fig. 8. In marked contrast, the RPMEC cytoprotection conferred by intact RPMSMCs was entirely reproduced when RPMECs were plated onto RPMSMCs fixed with either methanol or paraformaldehyde and subsequently cytomix stimulated as shown in Fig. 9. The cytoprotective effect of coculture was also observed for HPMECs and HPASMCs as well as shown in Fig. 10. A significantcomponent of the cytoprotective effect was obtained by plating HPMECs on tissue culture plastic coated with collagen (Fig. 10). Nonetheless, in preliminary experiments, we could not reverse the cytoprotection of fixed SMCs for cocultures of RPMECs exposed to cytomix by adding antibodies raised against TGF-beta , L-selectin, sialyl LewisX, sialyl LewisA, beta 2-integrin, VLA-4, laminin, fibronectin, or collagen (data not shown).


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Fig. 7.   Simultaneous cytomix stimulation of RPMECs and RPMSMCs in coculture abolished NO-dependent RPMEC cytotoxicity. Unstimulated RPMECs were 51Cr loaded and plated onto an RPMSMC layer after which the coculture was stimulated with CTM, CTM + L-NMMA, CTM + L-NMMA + SOD, or CTM + SOD. Injury index was calculated from 51Cr release 24 h later. No significant intergroup differences were found by ANOVA, P < 0.05 for all groups vs. control; n >=  3.



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Fig. 8.   Cytomix-stimulated RPMEC cytotoxicity was not decreased by conditioned medium from cytomix-stimulated RPMSMCs. Unstimulated RPMECs were 51Cr loaded, plated into culture dishes, and stimulated with cell culture supernatants harvested from RPMSMCs stimulated for 24 h with CTM, CTM + L-NMMA, or L-NMMA alone. Controls included RPMECs plated onto cell culture plastic or RPMSMCs and subsequently CTM stimulated, and injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. CTM-stimulated RPMECs plated onto cell culture plastic; n = 3.



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Fig. 9.   Cytomix-stimulated RPMEC cytotoxicity was prevented by contact with intact or fixed RPMSMCs but not by RPMSMC matrix. Unstimulated RPMECs were 51Cr loaded and plated onto cell culture plastic, intact RPMSMC layers, Triton X-100-extracted RPMSMC layers, or RPMSMC layers fixed with paraformaldehyde or methanol and subsequently CTM stimulated. Injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. RPMECs stimulated on plastic; n = 3.



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Fig. 10.   Cytomix-stimulated human pulmonary microvascular endothelial cell (HPMEC) cytotoxicity was prevented by contact with human pulmonary artery smooth muscle cells (HPASMCs) and decreased by contact with gelatin. Unstimulated RPMECs were 51Cr loaded and plated onto untreated cell culture plastic, HPASMC monolayers, or gelatin-coated plastic, and subsequently CTM stimulated. Injury index was calculated from 51Cr release 24 h later. * P < 0.05 vs. HPMECs stimulated on plastic; dagger  P < 0.05 vs. RPMECs stimulated on gelatin; n >=  3.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In the current study, we observed significant cytokine- and endotoxin (cytomix)-induced toxicity to cultured RPMECs (Figs. 1 and 2) and HPMECs (Fig. 10). This direct effect of cytomix on RPMECs was not due to NO because these cultured cells do not synthesize large amounts of NO in response to this mixture and the injury was not affected by either L-NMMA, SOD, or the combination (Fig. 2). It is possible that the transformed nature of RPMECs resulted in a different phenotype from a primary culture of proximal or distal rat pulmonary endothelium that has been noted to make measurable levels of nitrite after stimulation with various combinations of cytokines (12). It appears that after LPS (19) or 3 wk of hypoxia (33), there is immunoreactive iNOS in the rat pulmonary endothelium in situ but its longitudinal distribution (large vs. small vessel) remains to be determined. The mixture of cytokines and endotoxin represents several of the critical molecules that were identified in the medium of coculture experiments that originally identified a role for L-arginine-dependent NO synthesis in Kupfer cell-mediated hepatotoxicity (39). Although we did not attempt to identify contributions of any single agent in this mixture with respect to EC toxicity, it is clear from numerous other studies that endotoxin (15, 16, 25, 27, 46) or TNF-alpha (5, 9, 26) alone is sufficient to cause EC injury. The mechanism underlying such toxicity remains unclear but considerable data exist to suggest a role for cytokine- and LPS-induced intracellular oxidants in cultured pulmonary artery or microvascular ECs from calf (4), rat (12, 31), sheep (27, 46), and human (25) sources. The combination of cytokines and endotoxin used in these experiments did not result in apoptosis as determined by DNA laddering on gel electrophoresis (data not shown) in either cultured rat or human pulmonary ECs. This is different from our previous report in endotoxin-exposed primary cultures of sheep pulmonary artery endothelium (15, 16, 46). It is likely that there are significant species differences (25) in response to endotoxin or cytokines, as well as heterogeneity between large- and small- vessel endothelium (27).

The effect of cytokines and endotoxin on the pulmonary circulation has been studied extensively because these are important components of the pathogenesis of acute lung injury. The contribution of the L-arginine biosynthetic pathway in this pathophysiology has been the focus of recent attempts to understand the mechanism of pulmonary EC injury. Although the concept of cell-cell interactions is well developed in these experimental paradigms, it generally has centered around neutrophil-EC interactions. In this regard, a recent model system showed that low levels of NO protect cultured human pulmonary artery ECs against neutrophil-mediated injury, whereas high levels of NO released by activated monocytes exacerbate such injury (14). Considerably less attention has been devoted to the interaction of endothelium and pulmonary vascular smooth muscle despite the fact that iNOS is readily expressed in pulmonary vascular smooth muscle (17, 19, 32, 33). Our group previously reported that rat pulmonary artery SMC iNOS-derived NO is capable of decreasing adherent rat pulmonary artery ECs in a coculture system using Transwell configuration (42). In the current study, we show that RPMSMC iNOS-derived NO is cytotoxic to RPMECs in an L-NMMA-sensitive fashion when they are in direct contact during coculture (Fig. 3). The cytotoxicity was directly proportional to the magnitude of NO biosynthesis (Fig. 4). Separation of these cells by 1 mm via Transwell plates actually decreased this cytotoxicity (Fig. 3) consistent with the labile nature of the toxic substance. Thus assuming a diffusion constant of 3.3 mm2 × s-1 and an estimated half-life of 4 s in medium, steady-state concentrations of NO would have been reduced to approximately 1% from its source (20, 22). Presumably our previous report quantifying toxicity by measuring adherent ECs may have provided a more sensitive index in this coculture system. The impact of diffusion of NO may have contributed to another report showing that IL-1beta -stimulated vascular SMC-derived NO was not toxic to cocultures of ECs when they were separated by a permeable support (11). In this latter report, IL-1beta was toxic to cultured vascular smooth muscle, causing the release of smooth muscle-derived basic fibroblast growth factor that appeared to account for a stable component of smooth muscle-conditioned medium capable of promoting EC growth. We have not noted smooth muscle toxicity with either the cytokine and endotoxin combination of the current study or individual components of this mixture over a limited range of concentrations in primary rat pulmonary vascular SMCs (32, 45). Accordingly, it remains possible that cytokine-stimulated pulmonary vascular SMCs in situ may serve as a source of potentially damaging levels of NO or a secondary nitrogen monoxide to underlying vascular endothelium.

Nonetheless, the modest toxic effect of prestimulated RPMSMCs on cocultured nonexposed RPMECs (Fig. 3) may be contrasted to the cytoprotection that rat (Fig. 6) and human (Fig. 10) pulmonary vascular SMCs afforded cytokine-stimulated ECs in coculture. Under these conditions, we might have expected a summation of the direct effects of cytomix on monocultures of ECs (35-40%; Fig. 2) and the NO-derived EC toxicity (8-10%; Fig. 4) of the stimulated cocultured vascular SMC. Rather cytotoxicity was reduced to less than 10% (Fig. 6). It is unlikely that the decrease in cytotoxicity was due to a molecular effect of NO because the residual toxicity was L-NMMA (and SOD) independent (Fig. 7). Most compelling in understanding this toxicity was that it did not include a stable product in SMC-conditioned medium (Fig. 8) but rather could be reproduced even in paraformaldehyde- or methanol-fixed SMCs (Fig. 9). Cytoprotection could also be partially reproduced by contrasting the response of cytomix HPMECs grown on plastic to those grown on a collagen matrix (Fig. 10). Collectively, these data suggest a cytokine-activated EC surface molecule that interacts with either the SMC surface itself or an elaborated ECM. Candidates for this molecule might include those known to be induced on ECs, including P-selectin, E-selectin, or intercellular adhesion molecule (34). Corresponding ligands on SMCs could include sialyl LewisX, sialyl LewisA, L-selectin, and beta 1- or beta 2-integrins. TGF-beta can also be activated from its latent form by an EC-SMC interaction (10) that requires direct contact (2). Our initial antibody blocking studies with antibodies raised against TGF-beta , L-selectin, sialyl LewisX, sialyl LewisA, beta 1- and beta 2-integrins, VLA-4, laminin, fibronectin, and collagen did not alter cytoprotection (data not shown). These negative results, however, should be interpreted with caution because we did not test for the blocking or neutralizing effects of these antibodies. In this regard, a more telling observation was that gelatin partially reproduced the cytoprotection, suggesting an arginine-glycine-aspartic acid related moiety.

This interaction of ECs with surrounding cells and/or matrix may be of special importance in cytokine-induced acute lung injury where EC dysfunction and interstitial edema are early findings (18). Our studies suggest that a loss of this interaction in vivo could accelerate EC injury and/or death and are consistent with the observation that most of the vascular leak occurs at the alveolar level where SMCs are absent. If the receptors involved in the EC-SMC interaction can be identified, EC injury may be ameliorated by supplementation with synthetic analogs. For instance, administration of sialyl LewisX analogs reduces mortality and lung injury in rat models of acute lung injury (30) and trauma (38). The role of cytokine-activated EC expression of P-selectin in binding neutrophils is well appreciated but P-selectin also enhances EC attachment and spreading and decreases superoxide anion production in adherent neutrophils (44). Comparable EC interactions with adjacent vascular smooth muscle could decrease cytokine-induced EC injury or death.

In summary, we found that cytokine- and LPS-stimulated pulmonary vascular SMCs are capable of injuring adjacent ECs in an NO-dependent manner. This interaction, however, may be offset by the discovery of a novel SMC-EC interaction, which stabilizes both human and rat ECs against cytokine- and LPS-induced toxicity.


    ACKNOWLEDGEMENTS

We thank Drs. Ahmad Boota and William Stark for help in completing these studies. We also gratefully acknowledge Karla Wasserloos excellent technical assistance.


    FOOTNOTES

This work was supported in part by National Institutes of Health Grants HL-32154, GM-53789, and HL-03018; an American Heart Association Grant-in-Aid-Beginning, and The George Love Fund.

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. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: B. A. Johnson, Div. of Pulmonary, Allergy and Critical Care Medicine, Dept. of Medicine, Univ. of Pittsburgh School of Medicine, 3550 Terrace St., Pittsburgh, PA 15261 (E-mail: johnsonba2{at}msx.upmc.edu).

Received 18 June 1999; accepted in final form 20 September 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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