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J Appl Physiol 102: 1255-1264, 2007. First published November 9, 2006; doi:10.1152/japplphysiol.00786.2006 Free Article
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INVITED REVIEW

HIGHLIGHTED TOPIC
Physiological Imaging of the Lung

Real-time lung microscopy

Wolfgang M. Kuebler,1 Kaushik Parthasarathi,2 Jens Lindert,2 and Jahar Bhattacharya2,3

1Lung and Circulatory Research Laboratory, Institute of Physiology, Charité-Universitätsmedizin, Berlin, Germany; and 2Lung Biology Laboratory, Department of Physiology and Cellular Biophysics, and 3Department of Medicine, College of Physicians and Surgeons, Columbia University, and St. Luke's-Roosevelt Hospital Center, New York, New York


   ABSTRACT
TOP
 ABSTRACT
INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Although proinflammatory cell signaling in the alveolo-capillary region predisposes to acute lung injury, key cell-signaling mechanisms remain inadequately understood. Alveolo-capillary inflammation is likely to involve coordinated signaling among cells of different phenotypes. For example, migration of inflammatory cells into the alveolus might entail coordinated signaling between adjoining alveolar epithelial and microvascular endothelial cells. The popular cultured cell experimental strategy fails to replicate this multicellular environment. Cultured lung cells, both alveolar and endothelial, undergo phenotypic transformations; hence they might inadequately reflect innate responses of native cells. Consequently, new approaches are required for the investigation of cell signaling in the native setting. Here we summarize new developments in classical intravital microscopy and discuss real-time fluorescence imaging as a novel technique for studying second-messenger mechanisms in the alveolo-capillary region.

calcium ion; mitochondria; mitochondrial calcium ion; reactive oxygen species; nitric oxide; P-selectin; surfactant


   INTRAVITAL MICROSCOPY
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Intravital microscopy (IVM) entails video microscopy of the bright-field illuminated lung surface. The alveolar-capillary region accessible to IVM consists of alveoli and associated juxta-alveolar arteriolar, septal, and venular microvessels. Divergent vs. convergent flows distinguish arteriolar from venular microvessels, respectively. In the rodent lung, the microvessels are anatomic "capillaries," since none exceed ~30 µm in diameter and none contain smooth muscle (80). Recent advances in IVM are attributable to window preparations for direct visualization of subpleural lung microvessels and alveoli in the intact thorax. Early preparations were established by Wearn and coworkers (85) and Terry (78). More recently, lung IVM was approached in dog by Wagner (81) and in rabbit by Kuhnle et al. (42) by implanting flanged steel windows in the thoracic wall. The implants stabilize the lung surface by means of suction applied through a system of boreholes, thereby enabling IVM in the presence of lung ventilation.

Window approaches have enabled quantitative analyses in lung microvasculature in vivo. Visualization of blood flow by fluorescent plasma markers or erythrocytes (Fig. 1A) has facilitated direct measurement of lung capillary transit times, which were found to be in the range of 12.7 ± 3.2 s and thus markedly exceeded prior estimates (83). Subsequent studies identified the dynamic regulation of capillary recruitment and derecruitment by changes in cardiac output, alveolar oxygen tension, and local rheological factors (41, 57, 82). Fluorescence labeling of circulating white blood cells (Fig. 1B) yielded important information on the size, site, and regulation of the marginated pool of leukocytes and helped to characterize kinetics and mechanisms of leukocyte sequestration in lung inflammation. Studies by Lien et al. (45) and Kuebler et al. (36) identified the alveolar capillary bed as the predominant site of physiological leukocyte margination in the lung. During their passage through the alveolar capillary network, >90% of leukocytes become retained at least once for varying periods of time, ranging from <1 s up to >20 min (34, 45). Leukocyte retention results in the accumulation of a marginated pool, which equals the total number of circulating leukocytes in blood (33). Whereas physical deformation of leukocytes from a spherical into an elongated shape suggests mechanical hindrance of leukocyte passage through lung capillaries as the predominant mechanism underlying leukocyte margination (19), marked acceleration of leukocyte transit through lung capillaries by a nonspecific selectin inhibitor, fucoidin, suggests an additional contribution of adhesion molecule-mediated cell/cell interaction (35).


Figure 1
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  		Fig. 1. Intravital microscopic images of pulmonary microvessels in vivo obtained in the ventilated rabbit (A and B) and mouse (C and D). Images obtained in rabbit lungs show FITC-labeled erythrocytes (A, arrow) and rhodamine 6G-stained leukocytes adherent to the vascular wall (B, arrowheads) in a pulmonary arteriole. Due to their high velocity, erythrocytes show with a fluorescent tail. Images obtained in mice show a pulmonary arteriole (C, arrow indicates direction of flow) and surrounding alveolar capillary networks visualized by the plasma marker FITC dextran and rhodamine 6G-stained leukocytes (D, arrowheads) in a small postcapillary venule. [Images C and D by courtesy of Dr. A. Tabuchi, Institute of Physiology, Charité Berlin.]

 
The availability of transgenic mouse or rat models has further propelled lung IVM in vivo (48, 61, 64). Recently, Tabuchi et al. (77) reported a new IVM model in mice that provides excellent images of subpleural pulmonary arterioles, venules, and capillary networks (Fig. 1C) and of dynamic cell-cell interactions in pulmonary microvessels (Fig. 1D). In this model, the predominant site of hypoxic pulmonary vasoconstriction could be localized in small arterioles >30 µm (77). The possible use of transgenic animals yields additional potential for the analysis of intra- and intercellular signaling pathways in these models. Transgenic expression of biofluorescent proteins, such as GFP under cell-specific promotors, furthermore facilitates the study of specific cell subsets and circumvents problems inherent to synthetic fluorophores (74).

Two notable IVM models have been proposed for chronic studies of inflammatory processes in the lung. Fingar and coworkers (18) implanted a lung chamber into the right thoracic wall of anesthetized rats. The chamber allows for microscopic investigation of the lung surface after the animals resume spontaneous respiration and showed progressive vascular leakage of fluorescent dyes from pulmonary capillaries into lung alveoli in oleic acid-induced lung injury. However, long-term observations in this model are yet lacking. Sikora and coworkers (70) chose a different approach by transplanting lung tissue into the dorsal skinfold chamber of nude mice (44). After revascularization, these lung allografts maintain characteristics of pulmonary vasoreactivity in that they undergo hypoxic vasoconstriction (71). Nicotine was shown to induce leukocyte rolling and adhesion via activation of MAPK and P-selectin in this model, suggesting that these mechanisms may be relevant in nicotine-induced airway inflammation (71).

In vivo preparations for IVM of leukocyte trafficking in the lung bear the advantage of closely reflecting complex hemodynamic, metabolic, and inflammatory conditions in the intact organism. Yet they pose manifold technical challenges and limitations, which arise from the need to create optical access to the lung, to minimize motion artifacts due to respiratory and cardiac movements, and to control global and local hemodynamics, which directly affect the kinetics of cell trafficking (34, 41).

Isolated lung preparations circumvent difficulties innate to intact preparations. Blocking antibodies and indirect immunofluorescence imaging of endothelial adhesion molecules have been used in the isolated lung to identify the role of adhesion molecules in leukocyte sequestration. These studies suggest that, in hyperoxic, bleomycin-, or ventilator-induced lung injury, ICAM-1 universally mediates leukocyte entrapment in lung capillaries and rolling in pulmonary venules (53, 56, 68). In contrast, selectins seem to contribute differentially to these effects in that P-selectin mediates leukocyte margination in response to mechanical stimuli, such as hydrostatic stress (25) or overventilation (53), whereas L-selectin regulates the response to hyperoxia (56).

Nevertheless, some difficulties are that the isolated, perfused lung preparation may, per se, alter leukocyte kinetics because of cell activation by artificial surfaces and the need for anticoagulation (5). Moreover, leukocyte labeling, both in and ex vivo, carries problems. Ex vivo separation and labeling of cells allows for the study of specific leukocyte subsets, e.g., neutrophils or monocytes at the cost of possible cell activation. In vivo labeling by systemic infusion of nuclear or mitochondrial dyes, such as acridine orange or rhodamine 6G, avoids cell separation steps, but does not permit differentiation between different leukocyte subcategories and still cannot exclude dye-dependent alterations in cell trafficking (1, 21). Of note, it has proven difficult to track leukocyte transmigration into the lung interstitial and alveolar space. In vitro, the combination of phase contrast and fluorescence imaging yields important insights into cellular mechanisms regulating leukocyte transmigration (14, 76). In transilluminated preparations, transendothelial and interstitial migration of leukocytes can be clearly visualized and is not attenuated by fluorescent dyes such as rhodamine 6G (50). Yet in epi-illumination preparations, these techniques have proven difficult, potentially due to limitations by two-dimensional imaging techniques. As discussed later, recent advances in optical-sectioning microscopy yield the potential for subsequent three-dimensional image reconstruction and may thus provide a novel approach for the visualization of cell migration kinetics in three-dimensional tissue.

IVM has also been applied to micropuncture studies of lungs in vivo or in isolated, perfused lung preparations. Micropuncture measurements by the servo-null technique alone permit direct determinations of vascular and interstitial pressures in the lung. Micropuncture of sequential lung microvascular segments identified the alveolar septal capillary network as the major site of the total vascular pressure drop across the lung from pulmonary artery to left atrium (8, 22, 55, 69). Pulmonary vasoconstrictors, in contrast, tend to localize to either arterioles or venules as the predominant constriction site, with hypoxia or platelet-activating factor acting primarily in arteriolar segments (54, 63), while mediators such as histamine, endothelin-1, or thromboxane A2 constrict predominantly the venular compartment (62, 63). Micropuncture pressure measurements obtained in the interstititum surrounding alveolar septal junctions and in the adventitial space of 30- to 50-µm-diameter vessels revealed an interstitial pressure gradient that drives perimicrovascular fluid flux from the alveoli to the lung hilum (7, 20). Lung expansion abolishes this pressure gradient, suggesting that respiratory lung volume changes may cyclically vary interstitial liquid flow and thus generate a pumplike mechanism to facilitate interstitial fluid clearance (6, 43).


   REAL-TIME FLUORESCENCE IMAGING
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Real-time fluorescence imaging (RFI) methods, involving microscopic imaging of the isolated, perfused lung, have enabled the investigation of lung cellular and molecular responses in situ. The imaged field thereby provides an opportunity for obtaining optical data specifically from intact epithelial and endothelial cells of alveoli and microvessels, respectively.

Ca2+

Lung RFI enables determinations of cytosolic Ca2+ (Ca2+ cyt) levels in individual endothelial cells in intact capillaries. This approach led to the recognition of so-called "pacemaker" endothelial cells that are located at some vascular branch points. These branch-point endothelial cells regulate Ca2+ cyt at levels higher than midsegmental cells (Fig. 2A), and they generate intercellular Ca2+ cyt waves (87). In these in situ endothelial cells, increase of the microvascular pressure markedly increases the amplitude of Ca2+ cyt oscillation amplitude, while causing relatively small or no increases in the mean Ca2+ cyt levels (27, 39, 40). Microvascular injections of the proinflammatory agent TNF-{alpha} also increase Ca2+ cyt oscillations in branch-point endothelial cells (59). However, in contrast to the pressure-induced effects, TNF-{alpha} increases mean Ca2+ cyt levels. These findings reveal that a characteristic response of branch-point endothelial cells to proinflammatory stimuli is to increase Ca2+ cyt oscillations.


Figure 2
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  		Fig. 2. Real-time fluorescence imaging of lung capillaries. A: the image shows fluo 4 fluorescence in septal (SEP) and venular (VEN) capillaries surrounding alveoli (ALV). Note high fluorescence denoted by green pseudocolor occurs at the indicated branch point (white arrow). Direction of blood flow is indicated (black arrow). B: image obtained by confocal microscopy shows branch point (BR) region at the junction of a septal and a venular capillary. Note, fluorescence of mitochondrial dye, Mitotracker Green, is dominant in branch point cells (white arrows). An adherent leukocyte (*) shows uptake of fluorescence. C: concentration-dependent diaminofluorescein (DAF) fluorescence response in lung capillaries to infusion of nitric oxide donor S-nitroso-N-acetyl-penicillamine (SNAP). Fluorescence intensity (F) is expressed relative to its individual baseline (F0). D: nitric oxide response to increase of left atrial pressure (PLA). Vessel margins are depicted by line sketch, and direction of blood flow by arrow. Note response is dominant at branch points.

 
Mechanistic understanding of branch-point Ca2+ cyt oscillations was approached through Ca2+ determinations in the endoplasmic reticulum and mitochondria. Ca2+ uptake and release by the endoplasmic reticulum entails, respectively, endoplasmic reticulum Ca2+-ATPase pumps and inositol triphosphate receptors. Mitochondrial Ca2+ (Ca2+ mit) is regulated by Ca2+ entry through a Ca2+-sensitive uniporter (30) and by Ca2+ extrusion across a Na+/Ca2+ exchanger. Ca2+ levels in these organelles were determined by applying the dyes, fura 2-FF and rhod 2.

Fura 2-FF, which has low Ca2+-binding affinity, elicits Ca2+-sensitive fluorescence in the high-Ca2+ environment of the endoplasmic reticulum. In confirmation, Parthasarathi et al. (59) showed that thapsigargin, 2,5-di-(t-butyl)-1,4-hydroquinone, which depletes endoplasmic Ca2+ levels, decreases fura 2-FF fluorescence of lung capillaries. The fluorescence of cationic rhod 2, which accumulates specifically in the negatively charged mitochondrial matrix, reports Ca2+ mit levels. Through membrane permeablization studies, Ichimura et al. (27) confirmed the mitochondria-specific distribution of rhod 2 in lung capillary endothelium. These studies indicate that fura 2-FF, rhod 2, as well as the mitochondria-labeling dye Mitotracker Green, are most fluorescent at capillary branch points (Fig. 2B), attesting to high-density colocalization of the endoplasmic reticulum with mitochondria in branch-point endothelial cells.

TNF-{alpha} decreased fura 2-FF fluorescence, an effect that was blocked by the inositol triphosphate receptor blocker, xestospongin C, indicating that TNF-{alpha}-induced Ca2+ cyt oscillations occurred as a consequence of Ca2+ release from the endoplasmic reticulum. Importantly, both TNF-{alpha} and capillary pressure elevation increased Ca2+ mit. These Ca2+ mit responses were also xestospongin C inhibitable, indicating that, in branch-point endothelial cells, Ca2+ mit uptake occurs secondary to Ca2+ release from the endoplasmic reticulum.

Reactive Oxygen Species

Lung RFI has revealed new understanding of mechanisms underlying endothelial reactive oxygen species (ROS) production in lung capillaries. The dyes, dichlorofluorescein (DCF) and hydroethidene, detect capillary fluorescence of different ROS species, namely H2O2 and superoxide, respectively (2, 27, 59). Since DCF fluorescence might be attributable to nonspecific factors, such as the excitation light, DCF responses must be interpreted against appropriate controls. By slowing down the imaging acquisition rate, Parthasarathi et al. (59) and Ichimura et al. (27) obtained stable baselines of DCF fluorescence lasting more than 20 min, indicating that progressive light- and self-induced effects (10, 65) were absent. Using this imaging protocol, they showed that H2O2 caused dose-dependent increases in DCF fluorescence and that polyethylene glycol-catalase blocked DCF responses, while nitric oxide (NO) inhibitors were without effect. Inclusion of these controls provided a measure of security in interpreting the DCF data.

Al-Mehdi et al. (2) showed that nonhypoxic ischemia increases ROS production. Findings by Brueckl et al. (9) and Zhang et al. (88) indicate that hyperoxia and ischemia also increase lung capillary ROS production. In the studies by Brueckl et al. (9), hyperoxia increased ROS in two successive phases that were, respectively, mitochondria and NADPH oxidase dependent. The studies by Minamiya and coworkers revealed direct evidence for the neutrophil respiratory burst resulting from endotoxin (52) and apoptosis and ROS production in lung endothelial and epithelial cells in a model of reexpansion injury (67). Findings by Parthasarathi et al. (59) and Ichimura et al. (27) indicate that branch-point endothelial cells generate H2O2 entirely by mitochondrial mechanisms and that mitochondrial H2O2 increases in response to the TNF-{alpha}- and pressure-induced Ca2+ mit increases. These findings indicate that the physiological significance of augmented Ca2+ cyt oscillations is to increase Ca2+ mit that, in turn, increases mitochondrial ROS production.

In Situ Immunofluorescence

Detection of signaling proteins by in situ immunofluorescence has been applied in conjunction with RFI determinations. In lung microvessels, immunofluorescence revealed expression of the leukocyte adhesion receptor, P-selectin (34), and NADPH-oxidase activation by the small GTPase, Rac1 (9). In alveoli, the approach was used to distinguish type 1 from type 2 cells (25) and to detect activation of the cytosolic phospholipase A2 (37).

These approaches have revealed the mitochondrial role in proinflammatory signaling. Endothelial P-selectin is held in apical vesicles called Weibel-Palade bodies (WPB). WPB exocytosis causes P-selectin expression on the endothelial surface, thereby marking proinflammatory endothelial activation. Although increase in Ca2+ cyt levels is thought to stimulate WPB exocytosis, Parthasarathi et al. (59) and Ichimura et al. (27) showed that, in lung capillaries, H2O2 is the critical exocytosis stimulus, since blocking H2O2 with polyethylene glycol-catalase blocked P-selectin expression but not Ca2+ cyt oscillations.

Taking together these findings from branch-point endothelial cells, it appears that the major function of Ca2+ cyt oscillations is to modulate Ca2+ mit. Increases in Ca2+ cyt oscillations increase Ca2+ mit, thereby increasing mitochondrial H2O2 production. Mitochondrial H2O2 diffuses to cytosolic targets, namely WPB, to cause P-selectin exocytosis and, subsequently, leukocyte adherence (25). Hence endothelial Ca2+ cyt oscillations and mitochondria are revealed as critical regulators of proinflammatory signaling in lung capillaries.

Alveolo-capillary Cross Talk

RFI has led to major new understanding of alveolar-capillary signaling. Airway instillation of bacteria, such as P. aeruginosa, induces alveolar leukocyte migration in ~4 h (72). However, relevant signaling mechanisms are inadequately understood. Since epithelial and endothelial barriers restrict and possibly inhibit direct diffusion of chemokines, the chemotactic signal has to be vectorially transmitted from alveoli to capillaries by cross-compartmental signaling.

Addressing this question, Kuebler et al. (37) noted that alveolar microinjection of TNF-{alpha} increases Ca2+ cyt in both the alveolar epithelium and the adjacent capillary endothelium, revealing the presence of alveolar-capillary signal transmission. The endothelial response occurred in ~5 min, indicating that signal transmission initiates rapidly. Intra-alveolar injection of an antibody against TNF-{alpha} receptor-1 blocked the capillary Ca2+ cyt response, while intracapillary antibody injection had no effect. These findings rule out alveolar TNF-{alpha} leakage as an experimental artifact. Intracapillary TNF-{alpha} injection failed to increase alveolar epithelial Ca2+ cyt, pointing to the vectorial specificity of the TNF-{alpha}-induced alveolar-capillary signal transmission.

Characteristically, the alveolar epithelial response consists of high-amplitude Ca2+ oscillations that are absent in the capillary endothelial response, attesting to different patterns of Ca2+ mobilization in the two cell types. The oscillations reflect endoplasmic Ca2+ release, while the damped endothelial response indicates the presence of direct Ca2+ entry as induced by arachidonate (51). Accordingly, alveolar pretreatment with cytosolic phospholipase A2 inhibitors blocks the TNF-{alpha}-induced alveolar-capillary Ca2+ transmission (37). These findings indicate that epithelial and endothelial cells of the alveolo-capillary membranes effectively coordinate vectorial transmission of proinflammatory signals.

NO

To allow for direct detection of the diatomic free-radical NO in living cells and tissue, Kojima and coworkers (31) designed and synthesized a new group of fluorescent indicators, the diaminofluoresceins (DAFs). Due to their reactive aromatic vicinal diamines, the weakly fluorescent DAFs are rapidly nitrosated in the presence of NO and O2, yielding strongly fluorescent benzotriazole derivatives. DAF assays yield quantitative data on NO production, since the fluorescence intensity increases linearly with the concentration of exogenous NO donors (28), whereas NO scavengers attenuate the reaction in a concentration-dependent manner (47). Since nitrosation of DAF is an irreversible reaction, actual NO production is not directly given by fluorescence intensity, but by the first derivative of the fluorescence response over time. The detection limit of the assay has been estimated at 5 nmol/l in living cells for DAF-2 and as low as 3 nmol/l in the more recently developed derivative 4-amino-5-methylamino-2',7'-difluorescein (32). Of note, DAF is probably not nitrosated by NO itself, but rather by dinitrogen trioxide (N2O3), the product of NO autoxidation. Moreover, the fluorescence yield is affected by the stoichiometry between O2– and NO due to peroxynitrite-dependent nitrosylation of DAF (17). Hence, the potential influence of oxidative stress should be addressed in DAF bioassays, e.g., by the use of antioxidants or superoxide dismutase.

DAFs have been successfully applied to monitor NO production in isolated, perfused rat lungs (3, 27, 38). After loading of lung microvascular endothelial cells with the membrane-permeable dye diacetate via a microcatheterization technique, DAF imaging yields a dose-dependent fluorescence response to an exogenous NO donor, S-nitoso-N-acetylpenicillamine (Fig. 2C). DAF imaging in intact lung microvessels presents a powerful tool for in vivo studies of endothelial NO regulation, as well as for the analysis of endothelial dysfunction, its underlying mechanisms, and potential treatment options under various pathophysiological conditions. Previously, this technique has allowed the unraveling of cellular signaling pathways underlying endothelial NO responses to mechanical stimuli, such as microvascular pressure (Fig. 2D) or shear stress (3, 38). Of note, fluorescent NO cheletropic traps were introduced a few years ago for direct fluorescence detection of NO with high sensitivity (49). Yet a broader application of this interesting group of functional dyes has not yet been carried out.

Surfactant Secretion

A notable success has been the application of RFI for the direct quantification of alveolar surfactant secretion. Traditionally, lung surfactant secretion is determined indirectly through quantification of surfactant phospholipid content in the bronchoalveolar lavage, although bronchoalveolar lavage recovery procedures may stimulate surfactant secretion (24). The RFI approach uses the LysoTracker dyes (Molecular Probes), which are fluorophore-linked weakly basic compounds that become protonated and thereby sequestered in acidic organelles, such as surfactant-containing lamellar bodies (LBs). In LysoTracker-loaded alveoli, LB fluorescence of type 2 cells dominates over the weaker fluorescence of smaller lysosomal inclusions and is conveniently detected as brightly lit fluorescent spots (Fig. 3). Loss of type 2 cell fluorescence denotes LB exocytosis.


Figure 3
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  		Fig. 3. Real-time confocal microscopy of pulmonary alveoli. Left: image shows alveoli (ALV) containing type 2 cells loaded with lamellar body localizing dye, Lysotracker Red (arrows). Note density of type 2 cells is 1–2/alveolus. Right: the region enclosed by the dotted circle in the left image is shown at higher magnification. Red Lysotracker Red fluorescence (arrows) denotes lamellar body content and distribution.

 
Using RFI, Ashino et al. (4) addressed cell-signaling mechanisms underlying lung inflation-induced surfactant secretion (24). This mechanical effect is modeled in cultured type 2 cells by cell-stretch protocols (13, 46, 60, 86) that identify Ca2+ cyt increase as the major determinant of stretch-induced surfactant secretion (46, 86). However, the studies by Ashino et al. (4) reveal important differences in the stretch-induced Ca2+ cyt response between cultured and intact alveolar cells. In cultured type 2 cells, cell stretch induces external Ca2+ entry (46), thereby markedly increasing mean Ca2+ cyt (86). However, in the intact alveolus, the major Ca2+ cyt response to lung inflation is a prolonged increase in Ca2+ cyt oscillations, but no increase in mean Ca2+ cyt (4).

Gap junctional inhibitors blocked Ca2+ cyt oscillations in type 2 cells, but not in type 1 cells, indicating that the oscillations are transmitted by gap junctional communication from type 1 to type 2 cells. This transmission is critical, since inhibiting the oscillations inhibits LB exocytosis (4). The new understanding gained from these studies is that type 1 cells act as alveolar strain sensors. Thus, during inflation, type 1 cells respond to alveolar stretch by transmitting Ca2+ cyt oscillations to type 2 cells, causing surfactant secretion.


   NEW DIRECTIONS
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Photolytic Uncaging

This method enables optically induced intracellular Ca2+ release (15), avoiding nonspecific effects of soluble ligands or mechanical stimuli. For photolytic uncaging of Ca2+, cells are loaded with membrane-permeable, nitrophenyl (NP)-EGTA AM that deesterifies intracellularly to NP-EGTA. Because of its high Ca2+ affinity (Kd 80 nM), NP-EGTA acts as a "cage," sequestering free Ca2+ (16). Photo-excitation with high-intensity UV light (320 nm) dissociates NP-EGTA into iminodiacetic products of low-Ca2+ affinity (Kd 3 mM), thereby releasing the caged Ca2+ and increasing Ca2+ cyt. Uniquely, photolytic uncaging affords Ca2+ release in specific photo-targeted cells.

The application of photolytic release of caged Ca2+ in conjunction with RFI has enabled the detection of cell-cell communication in alveoli and lung capillaries. Ichimura et al. (26) loaded the Ca2+ cage in a cluster of alveoli (acinus) and determined Ca2+ cyt and surfactant secretion rates by the approaches reviewed above. Photolytic Ca2+ uncaging increased alveolar Ca2+ cyt and induced surfactant secretion, not only in the photo-targeted alveolus, but also in neighboring alveoli, indicating that Ca2+ is intercommunicated between adjacent alveoli. Pretreating the alveoli with the connexin 43 (Cx43)-inhibiting peptides blocked these conducted responses.

In the experiments by Parthasarathi et al. (58), in which NP-EGTA was loaded in the lung microvascular bed, photolytic uncaging revealed conduction of the evoked Ca2+ signal from septal capillaries to distances of ~150 µm from the uncaging site. Ca2+ increases induced in the alveolar capillary activated P-selectin expression in venules. As in the alveolar experiments, Cx43 peptides blocked capillary Ca2+ conduction. In addition, no conduction occurred in Cx43-deficient mice. These findings attest to a novel role for Cx43-mediated gap junctions as conduits for the spread of proinflammatory signals in the lung capillary bed. Taking together these uncaging studies in alveoli and capillaries, it appears that gap-junction-mediated Ca2+ conduction plays an important role in establishing defensive cell signaling in the lung.

Optical-Sectioning Microscopy

Confocal or multiphoton microscopy of optical sections eliminates blurring due to out-of-focus fluorescence that is inherent in standard, wide-angle microscopy (principles reviewed in Refs. 12, 23). Safdar et al. (66) applied confocal microscopy to determine F-actin distribution in lung venular capillaries, as determined by the fluorescence of rhodamine-phalloidin. Optical sections taken along a vertical axis revealed consistently greater rhodamine fluorescence at branch points, especially around ostia of tributary vessels (Fig. 4A, top). As tributaries progressively merged with the mainstream capillary, the intensity of actin fluorescence progressively decreased (Fig. 4A, middle and bottom). In general, actin fluorescence is higher at tributary branch points than at midsegmental locations (Fig. 4B), indicating that actin distribution is spatially nonuniform within a capillary segment.


Figure 4
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  		Fig. 4. Real-time confocal microscopy of a lung venular capillary. The capillary was loaded with actin dye, rhodamine-phalloidin, and Ca2+ dye fluo 4. E-cadherin was determined by immunofluorescence. Images show predominance of actin fluorescence at branch points (Brp) (A and B) and colocalization of E-cadherin and actin at the endothelial junction (C). Mid, midsegmental.

 
In the same paper, Safdar et al. (66) determined endothelial cell geometry in venular capillaries in terms of fluo 4 fluorescence, which was bright in the nuclear region, but became attenuated toward the cell periphery (Fig. 4C). These peripheral sites were evidently cell junctions at which E-cadherin colocalized with actin. Interestingly, the capillaries failed to stain for VE-cadherin, the cadherin isoform detected in large-vessel endothelia. These findings indicate the usefulness of optical sectioning to access spatial data from three-dimensional anatomic structures.

In an interesting application of confocal microscopy, Carter et al. detected fluorescence emission from an optically fixed volume to quantify alveolar liquid flow (11). By filling the isolated, perfused mouse lung with buffer solution containing FITC-dextran, they achieved fluorescent labeling of the alveolar lumen. Images acquired at a series of z-positions were used to reconstruct the three-dimensional alveolar shape. To validate their measurements, they changed the osmolality of the vascular perfusate to induce water movement between the alveolar and perialveolar space. Perfusion with hyperosmolar buffer increased alveolar fluorescence compared with isosmolar conditions, indicating concentration of the fluorescent tracer, hence liquid removal from the alveolus. Accordingly, hyposmolar perfusate reduced alveolar fluorescence, consistent with liquid inflow into the alveolus.

The same group later extended this approach by filling the air space of mouse lung with fluorescent indicators of Na+ and Cl– concentrations to determine unidirectional ion flux. The flux rates deduced by measurement of pleural surface fluorescence were validated by analysis of air space fluid samples. These findings have led to the understanding that cAMP-dependent Cl– and Na+ transport occurs across the distal air space barrier into the alveolar lumen (29).

Multiphoton microscopy provides a potentially promising approach for lung RFI studies. As different from conventional single-photon excitation, multiphoton fluorescence is excited by the combined energy of two or more low-energy (long wavelength) photons. This excitation is limited to the focal plane, since the probability of simultaneous absorption is negligible at other planes in the light path. This approach has several advantages over conventional confocal microscopy. First, the bulk of the fluorophore in the light path is unperturbed, although photobleaching might occur at the focal plane. Second, the incorporation of tunable lasers in the technology allows combined detection of UV and visible-light excited fluorophores (Fig. 5). Consequently, cells of different phenotypes are accurately localized in the alveolo-capillary region.


Figure 5
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  		Fig. 5. Real-time multiphoton microscopy of the lung. Top: images of the alveolo-capillary region showing fluorescence of the UV-excited nuclear dye Hoechst 33342 (left), the Ca2+ dye fluo 4 (middle), and the corresponding overlay (right). Localization of nuclei allows identification of fluorescence profile at different locations of the capillary, as for example at branch points (arrow). Bottom: second harmonic generation (SHG) in the subpleural interstitium (left) and in alveolar septae (right, arrow). Note SHG of collagen fibers is considerably greater in the subpleural than in the alveolar interstitium.

 
Third, the long wavelength excitation light used for multiphoton excitation provides a unique opportunity to detect lung connective tissue without the need for fluorescent staining. Light polarization in specific tissues causes second harmonic generation (SHG), which is a light signal at exactly one-half the excitation wavelength (44). SHG is typically generated by collagen in biological samples (89). Initial studies indicate that SHG can be applied for detection of collagen fibers on the lung surface and in alveolar septae (Fig. 5, bottom panels). This approach may find useful application in dynamic studies of lung fibrosis.

Fluorescence Energy Resonance Transfer

Fluorescence energy resonance transfer (FRET) occurs in the presence of two appropriate fluorophores (donor and acceptor) lying in close proximity, as for example when bound to different protein domains. Under these conditions, excitation of the donor results in partial energy transfer to the acceptor. If proximity of the donor to the acceptor changes, as for example by a conformational change in the protein, the amount of transferred energy changes, resulting in change in the ratio of the fluorescence emissions of the donor and acceptor. However, the ratio might also be determined by changes in protein dipole moment. To distinguish between these possibilities, donor fluorescence lifetimes are separately determined in the presence and absence of the acceptor. The true interfluorophore distance is determined by insertion of fluorescence lifetimes in the Förster equation (75). FRET pairs consisting of genetically encoded fluorescent proteins can be expressed in living cells and targeted to subcellular regions by specific promoters.

St. Croix et al. (73) applied this approach to transfect vascular endothelium of mouse lungs with cDNA encoding for a NO-sensitive metallothionein protein containing a cyan fluorescent protein/yellow fluorescent protein FRET pair. Transfection was confirmed after 48 h in isolated lungs, and the FRET response recorded by confocal microscopy indicated conformational changes of the metallothionein in response to a NO donor. Clearly, the application of FRET might enlarge the scope of RFI to address intracellular signaling at subcellular locations.


   CONCLUDING REMARKS
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
We summarize here some of the main findings from real-time lung microscopy. In general, the microscopic approach to in vivo and isolated, perfused lungs has evolved a novel understanding of lung proinflammatory signaling. The dynamic determinations with RFI portray the alveolo-capillary region as a field that is highly signaling sensitive. Rapid second-messenger responses incorporate different intracellular compartments and coordinate interactions among different cells. Importantly, for the first time, mitochondria are recognized as initiators of lung inflammation. New directions in lung RFI being developed are likely to reveal other presently unknown mechanisms. Wider application of real-time lung microscopy might establish a novel understanding of cell signaling underlying different processes of lung disease.


   APPENDIX: NOTES ON METHODS
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Fluorescent Dyes

RFI determinations have been obtained using cell-permeable fluorescent indicators. For example, the Ca2+-sensitive dye fura 2 is membrane permeable in the AM form. In the cytosol, hydrolases cleave the AM moiety, leaving the dye membrane impermeable, hence cell locked. However, dyes may be extruded from the cytosol by plasma membrane transporters, an effect best evident for the ROS-detecting dye DCF. To ensure steady cellular delivery of intracellular DCF, infusions of the precursor agent dichlorofluorescein diacetate are given throughout. For fura 2, dye content is indicated by the Ca2+-insensitve fluorescence that is excited at wavelength of 360 nm.

Fluorescence indicators are either of the single- or dual-wavelength varieties. Dual-wavelength dyes that include fura 2 allow ratiometric imaging. Ratiometric quantification corrects for variability of cell volume and light absorption, which potentially introduces errors in quantitative interpretations of fluorescence emissions from single-wavelength dyes. For fura 2, in situ calibration allows direct interpretation of the imaged data in terms of the Ca2+ cyt concentration. However, dyes such as fura 2 are excited at or near the UV range, excluding their application where use of UV is not indicated.

The advantage of single-wavelength dyes such as fluo 4 is that the dyes are excited in the range of visible light wavelengths and are, therefore, suitable for non-UV applications. Although the fluorescence emissions of these dyes do not provide a direct readout of the Ca2+ cyt concentration, calibration against the concentration-calibrated fluorescence of a ratiometric dye such as fura 2 is possible by coloading both the single wavelength and the ratiometric dyes in the same cell (26). Alternatively, the cell volume problem associated with single-wavelength dyes may be suitably addressed by fluorescence determinations by optical sectioning microscopy.

Dye Loading

For lung microvascular RFI, one approach is to inject the dyes through a venous microcatheter wedged in the microvascular bed. Such microcatheter injections infuse a delimited microvascular region, protecting the bulk of the lung vasculature from dye exposure, hence reducing nonspecific fluorescence (background). While targeting capillary fluorescence, nonspecific fluorescence from extravascular cells may be eliminated by maintaining absorptive conditions in the vasculature during dye loading. Absorptive conditions are established by maintaining low vascular pressures and by controlling the solution colloid osmotic pressure of the dye at physiological levels.

For alveolar RFI, dyes are microinfused to fill seven to eight alveoli through a glass micropipette introduced in a single alveolus. Air withdrawal through the micropipette confirms correct micropipette tip placement in the alveolar lumen. Since alveolar microinfusions are rapidly (~2–4 s) removed (84), presumably by alveolar surface-active forces, dye loading requires prolonged microinfusions (20–30 min). Cells reseal spontaneously after micropipette removal (79); hence alveolar dye leakage does not occur from the micropuncture site. Faulty micropuncture that might tear the alveolar wall causes interstitial dye loading, the diffuse fluorescence of which is distinct from the discrete fluorescence of dye-loaded alveolar cells.


   GRANTS
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
The studies reported were supported by the following: National Heart, Lung, and Blood Institute (NHLBI) Grants HL-57556, HL-64896, and HL-69514 to J. Bhattacharya; NHLBI Grant HL-075503 to K. Parthasarathi; and research grants of the Medical Faculty of the University of Munich and the Deutsche Forschungsgemeinschaft KU1218/4 and GRK 865 to W. M. Kuebler.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
REFERENCES
 
Dr. Sunita Bhattacharya read the manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Bhattacharya, St. Luke's-Roosevelt Hospital Center, AJA 509, 1000 10thAve., New York, New York 10019 (e-mail: jb39{at}columbia.edu)


   REFERENCES
TOP
 ABSTRACT
 INTRAVITAL MICROSCOPY
 REAL-TIME FLUORESCENCE IMAGING
 NEW DIRECTIONS
 CONCLUDING REMARKS
 APPENDIX: NOTES ON METHODS
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 

  1. Abbitt KB, Rainger GE, Nash GB. Effects of fluorescent dyes on selectin and integrin-mediated stages of adhesion and migration of flowing leukocytes. J Immunol Methods 239: 109–119, 2000.[CrossRef][Web of Science][Medline]
  2. Al-Mehdi AB, Shuman H, Fisher AB. Intracellular generation of reactive oxygen species during nonhypoxic lung ischemia. Am J Physiol Lung Cell Mol Physiol 272: L294–L300, 1997.[Abstract/Free Full Text]
  3. Al-Mehdi AB, Song C, Tozawa K, Fisher AB. Ca2+- and phosphatidylinositol 3-kinase-dependent nitric oxide generation in lung endothelial cells in situ with ischemia. J Biol Chem 275: 39807–39810, 2000.[Abstract/Free Full Text]
  4. Ashino Y, Ying X, Dobbs LG, Bhattacharya J. [Ca2+]i oscillations regulate type II cell exocytosis in the pulmonary alveolus. Am J Physiol Lung Cell Mol Physiol 279: L5–L13, 2000.[Abstract/Free Full Text]
  5. Baufreton C, Kirsch M, Loisance DY. Measures to control blood activation during assisted circulation. Ann Thorac Surg 66: 1837–1844, 1998.[Abstract/Free Full Text]
  6. Bhattacharya J, Gropper MA, Shepard JM. Lung expansion and the perialveolar interstitial pressure gradient. J Appl Physiol 66: 2600–2605, 1989.[Abstract/Free Full Text]
  7. Bhattacharya J, Gropper MA, Staub NC. Interstitial fluid pressure gradient measured by micropuncture in excised dog lung. J Appl Physiol 56: 271–277, 1984.[Abstract/Free Full Text]
  8. Bhattacharya S, Glucksberg MR, Bhattacharya J. Measurement of lung microvascular pressure in the intact anesthetized rabbit by the micropuncture technique. Circ Res 64: 167–172, 1989.[Abstract/Free Full Text]
  9. Brueckl C, Kaestle S, Kerem A, Habazettl H, Krombach F, Kuppe H, Kuebler WM. Hyperoxia-induced reactive oxygen species formation in pulmonary capillary endothelial cells in situ. Am J Respir Cell Mol Biol 34: 453–463, 2006.[Abstract/Free Full Text]
  10. Burow S, Valet G. Flow-cytometric characterization of stimulation, free radical formation, peroxidase activity and phagocytosis of human granulocytes with 2,7-dichlorofluorescein (DCF). Eur J Cell Biol 43: 128–133, 1987.[Web of Science][Medline]
  11. Carter EP, Matthay MA, Farinas J, Verkman AS. Transalveolar osmotic and diffusional water permeability in intact mouse lung measured by a novel surface fluorescence method. J Gen Physiol 108: 133–142, 1996.[Abstract/Free Full Text]
  12. Conchello JA, Lichtman JW. Optical sectioning microscopy. Nat Med 2: 920–931, 2005.
  13. Correa-Meyer E, Pesce L, Guerrero C, Sznajder JI. Cyclic stretch activates ERK1/2 via G proteins and EGFR in alveolar epithelial cells. Am J Physiol Lung Cell Mol Physiol 282: L883–L891, 2002.[Abstract/Free Full Text]
  14. Cuvelier SL, Paul S, Shariat N, Colarusso P, Patel KD. Eosinophil adhesion under flow conditions activates mechanosensitive signaling pathways in human endothelial cells. J Exp Med 202: 865–876, 2005.[Abstract/Free Full Text]
  15. Diaz ME, Trafford AW, Eisner DA. The effects of exogenous calcium buffers on the systolic calcium transient in rat ventricular myocytes. Biophys J 80: 1915–1925, 2001.
  16. Ellis-Davies GC, Kaplan JH. Nitrophenyl-EGTA, a photolabile chelator that selectively binds Ca2+ with high affinity and releases it rapidly upon photolysis. Proc Natl Acad Sci USA 91: 187–191, 1994.[Abstract/Free Full Text]
  17. Espey MG, Thomas DD, Miranda KM, Wink DA. Focusing of nitric oxide mediated nitrosation and oxidative nitrosylation as a consequence of reaction with superoxide. Proc Natl Acad Sci USA 99: 11127–11132, 2002.[Abstract/Free Full Text]
  18. Fingar VH, Taber SW, Wieman TJ. A new model for the study of pulmonary microcirculation: determination of pulmonary edema in rats. J Surg Res 57: 385–393, 1994.[CrossRef][Web of Science][Medline]
  19. Gebb SA, Graham JA, Hanger CC, Godbey PS, Capen RL, Doerschuk CM, Wagner WW Jr. Sites of leukocyte sequestration in the pulmonary microcirculation. J Appl Physiol 79: 493–497, 1995.[Abstract/Free Full Text]
  20. Glucksberg MR, Bhattacharya J. Effect of dehydration on interstitial pressures in the isolated dog lung. J Appl Physiol 67: 839–845, 1989.[Abstract/Free Full Text]
  21. Hansell P, Berger E, Chambers JD, Arfors KE. Influence of acridine tracer dyes on neutrophil function. J Leukoc Biol 56: 464–468, 1994.[Abstract]
  22. Haworth ST, Rickaby DA, Linehan JH, Dawson CA. Subpleural pulmonary microvascular pressures in the dog lung. J Appl Physiol 79: 615–622, 1995.[Abstract/Free Full Text]
  23. Helmchen F, Denk W. Deep tissue two-photon microscopy. Nat Med 2: 932–940, 2005.
  24. Hildebran JN, Goerke J, Clements JA. Surfactant release in excised rat lung is stimulated by air inflation. J Appl Physiol 51: 905–910, 1981.[Abstract/Free Full Text]
  25. Ichimura H, Parthasarathi K, Issekutz AC, Bhattacharya J. Pressure-induced leukocyte margination in lung postcapillary venules. Am J Physiol Lung Cell Mol Physiol 289: L407–L412, 2005.[Abstract/Free Full Text]
  26. Ichimura H, Parthasarathi K, Lindert J, Bhattacharya J. Lung surfactant secretion by inter-alveolar Ca2+ signaling. Am J Physiol Lung Cell Mol Physiol 291: L596–L601, 2006.[Abstract/Free Full Text]
  27. Ichimura H, Parthasarathi K, Quadri S, Issekutz AC, Bhattacharya J. Mechano-oxidative coupling by mitochondria induces proinflammatory responses in lung venular capillaries. J Clin Invest 111: 691–699, 2003.[CrossRef][Web of Science][Medline]
  28. Itoh Y, Ma FH, Hoshi H, Oka M, Noda K, Ukai Y, Kojima H, Nagano T, Toda N. Determination and bioimaging method for nitric oxide in biological specimens by diaminofluorescein fluorometry. Anal Biochem 287: 203–209, 2000.[CrossRef][Web of Science][Medline]
  29. Jiang J, Song Y, Bai C, Koller BH, Matthay MA, Verkman AS. Pleural surface fluorescence measurement of Na+ and Cl– transport across the air space-capillary barrier. J Appl Physiol 94: 343–352, 2003.[Abstract/Free Full Text]
  30. Kirichok Y, Krapivinsky G, Clapham DE. The mitochondrial calcium uniporter is a highly selective ion channel. Nature 427: 360–364, 2004.[CrossRef][Medline]
  31. Kojima H, Nakatsubo N, Kikuchi K, Kawahara S, Kirino Y, Nagoshi H, Hirata Y, Nagano T. Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal Chem 70: 2446–2453, 1998.[Medline]
  32. Kojima H, Urano Y, Kikuchi K, Higuchi T, Hirata Y, Nagano T. Fluorescent indicators for imaging nitric oxide production. Angew Chem Int Ed Engl 38: 3209–3212, 1999.[CrossRef][Medline]
  33. Kuebler WM, Goetz AE. The marginated pool. Eur Surg Res 34: 92–100, 2002.[CrossRef][Web of Science][Medline]
  34. Kuebler WM, Kuhnle GE, Goetz AE. Leukocyte margination in alveolar capillaries: interrelationship with functional capillary geometry and microhemodynamics. J Vasc Res 36: 282–288, 1999.[CrossRef][Web of Science][Medline]
  35. Kuebler WM, Kuhnle GE, Groh J, Goetz AE. Contribution of selectins to leucocyte sequestration in pulmonary microvessels by intravital microscopy in rabbits. J Physiol 501: 375–386, 1997.[Abstract/Free Full Text]
  36. Kuebler WM, Kuhnle GE, Groh J, Goetz AE. Leukocyte kinetics in pulmonary microcirculation: intravital fluorescence microscopic study. J Appl Physiol 76: 65–71, 1994.[Abstract/Free Full Text]
  37. Kuebler WM, Parthasarathi K, Wang PM, Bhattacharya J. A novel signaling mechanism between gas and blood compartments of the lung. J Clin Invest 105: 905–913, 2000.[Web of Science][Medline]
  38. Kuebler WM, Uhlig U, Goldmann T, Schael G, Kerem A, Exner K, Martin C, Vollmer E, Uhlig S. Stretch activates nitric oxide production in pulmonary vascular endothelial cells in situ. Am J Respir Crit Care Med 168: 1391–1398, 2003.[Abstract/Free Full Text]
  39. Kuebler WM, Ying X, Bhattacharya J. Pressure-induced endothelial Ca2+ oscillations in lung capillaries. Am J Physiol Lung Cell Mol Physiol 282: L917–L923, 2002.[Abstract/Free Full Text]
  40. Kuebler WM, Ying X, Singh B, Issekutz AC, Bhattacharya J. Pressure is proinflammatory in lung venular capillaries. J Clin Invest 104: 495–502, 1999.[Web of Science][Medline]
  41. Kuhnle GE, Kuebler WM, Groh J, Goetz AE. Effect of blood flow on the leukocyte-endothelium interaction in pulmonary microvessels. Am J Respir Crit Care Med 152: 1221–1228, 1995.[Abstract]
  42. Kuhnle GE, Leipfinger FH, Goetz AE. Measurement of microhemodynamics in the ventilated rabbit lung by intravital fluorescence microscopy. J Appl Physiol 74: 1462–1471, 1993.[Abstract/Free Full Text]
  43. Lai-Fook SJ. Perivascular interstitial fluid pressure measured by micropipettes in isolated dog lung. J Appl Physiol 52: 9–15, 1982.[Abstract/Free Full Text]
  44. Lehr HA, Leunig M, Menger MD, Nolte D, Messmer K. Dorsal skinfold chamber technique for intravital microscopy in nude mice. Am J Pathol 143: 1055–1062, 1993.[Abstract]
  45. Lien DC, Wagner WW Jr, Capen RL, Haslett C, Hanson WL, Hofmeister SE, Henson PM, Worthen GS. Physiological neutrophil sequestration in the lung: visual evidence for localization in capillaries. J Appl Physiol 62: 1236–1243, 1987.[Abstract/Free Full Text]
  46. Liu M, Xu J, Tanswell AK, Post M. Inhibition of mechanical strain-induced fetal rat lung cell proliferation by gadolinium, a stretch-activated channel blocker. J Cell Physiol 161: 501–507, 1994.[CrossRef][Web of Science][Medline]
  47. Lopez-Figueroa MO, Caamano C, Marin R, Guerra B, Alonso R, Morano MI, Akil H, Watson SJ. Characterization of basal nitric oxide production in living cells. Biochim Biophys Acta 1540: 253–264, 2001.[Medline]
  48. McCormack DG, Mehta S, Tyml K, Scott JA, Potter R, Rohan M. Pulmonary microvascular changes during sepsis: evaluation using intravital videomicroscopy. Microvasc Res 60: 131–140, 2000.[CrossRef][Web of Science][Medline]
  49. Meineke P, Rauen U, de Groot H, Korth HG, Sustman R. Nitric oxide detection and visualization in biological systems. Applications of the FNOCT method. Biol Chem 381: 575–582, 2000.[CrossRef][Web of Science][Medline]
  50. Mempel TR, Moser C, Hutter J, Kuebler WM, Krombach F. Visualization of leukocyte transendothelial and interstitial migration using reflected light oblique transillumination in intravital video microscopy. J Vasc Res 40: 435–441, 2003.[CrossRef][Web of Science][Medline]
  51. Mignen O, Shuttleworth TJ. I(ARC), a novel arachidonate-regulated, noncapacitative Ca2+ entry channel. J Biol Chem 275: 9114–9119, 2000.[Abstract/Free Full Text]
  52. Minamiya Y, Tozawa K, Kitamura M, Saito S, Ogawa J. Platelet-activating factor mediates intercellular adhesion molecule-1-dependent radical production in the nonhypoxic ischemia rat lung. Am J Respir Cell Mol Biol 19: 150–157, 1998.[Abstract/Free Full Text]
  53. Miyao N, Suzuki Y, Takeshita K, Kudo H, Ishii M, Hiraoka R, Nishio K, Tamatani T, Sakamoto S, Suematsu M, Tsumura H, Ishizaka A, Yamaguchi K. Various adhesion molecules impair microvascular leukocyte kinetics in ventilator-induced lung injury. Am J Physiol Lung Cell Mol Physiol 290: L1059–L1068, 2006.[Abstract/Free Full Text]
  54. Nagasaka Y, Bhattacharya J, Nanjo S, Gropper MA, Staub NC. Micropuncture measurement of lung microvascular pressure profile during hypoxia in cats. Circ Res 54: 90–95, 1984.[Abstract/Free Full Text]
  55. Negrini D, Gonano C, Miserocchi G. Microvascular pressure profile in intact in situ lung. J Appl Physiol 72: 332–339, 1992.[Abstract/Free Full Text]
  56. Nishio K, Suzuki Y, Aoki T, Suzuki K, Miyata A, Sato N, Naoki K, Kudo H, Tsumura H, Serizawa H, Morooka S, Ishimura Y, Suematsu M, Yamaguchi K. Differential contribution of various adhesion molecules to leukocyte kinetics in pulmonary microvessels of hyperoxia-exposed rat lungs. Am J Respir Crit Care Med 157: 599–609, 1998.
  57. Okada O, Presson RG Jr, Godbey PS, Capen RL, Wagner WW Jr. Temporal capillary perfusion patterns in single alveolar walls of intact dogs. J Appl Physiol 76: 380–386, 1994.[Abstract/Free Full Text]
  58. Parthasarathi K, Ichimura H, Monma E, Lindert J, Quadri S, Issekutz A, Bhattacharya J. Connexin 43 mediates spread of Ca2+-dependent proinflammatory responses in lung capillaries. J Clin Invest 116: 2193–2200, 2006.[CrossRef][Web of Science][Medline]
  59. Parthasarathi K, Ichimura H, Quadri S, Issekutz A, Bhattacharya J. Mitochondrial reactive oxygen species regulate spatial profile of proinflammatory responses in lung venular capillaries. J Immunol 169: 7078–7086, 2002.[Abstract/Free Full Text]
  60. Patel AS, Reigada D, Mitchell CH, Bates SR, Margulies SS, Koval M. Paracrine stimulation of surfactant secretion by extracellular ATP in response to mechanical deformation. Am J Physiol Lung Cell Mol Physiol 289: L489–L496, 2005.[Abstract/Free Full Text]
  61. Presson RG Jr, Fisher A, Glenny RW, Lamm WJE. Pulmonary microvascular reserve in the intact rat (Abstract). Proc Am Thorac Soc 3: A682, 2006.
  62. Raj JU, Anderson J. Pulmonary venous responses to thromboxane A2 analogue and atrial natriuretic peptide in lambs. Circ Res 66: 496–502, 1990.[Abstract/Free Full Text]
  63. Raj JU, Toga H, Ibe BO, Anderson J. Effects of endothelin, platelet activating factor and thromboxane A2 in ferret lungs. Respir Physiol 88: 129–140, 1992.[CrossRef][Web of Science][Medline]
  64. Razavi HM, Wang L, Weicker S, Quinlan GJ, Mumby S, McCormack DG, Mehta S. Pulmonary oxidant stress in murine sepsis is due to inflammatory cell nitric oxide. Crit Care Med 33: 1333–1339, 2005.[CrossRef][Web of Science][Medline]
  65. Rota C, Fann YC, Mason RP. Phenoxyl free radical formation during the oxidation of the fluorescent dye 2',7'-dichlorofluorescein by horseradish peroxidase. Possible consequences for oxidative stress measurements. J Biol Chem 274: 28161–28168, 1999.[Abstract/Free Full Text]
  66. Safdar Z, Wang P, Ichimura H, Issekutz AC, Quadri S, Bhattacharya J. Hyperosmolarity enhances the lung capillary barrier. J Clin Invest 112: 1541–1549, 2003.[CrossRef][Web of Science][Medline]
  67. Saito S, Ogawa J, Minamiya Y. Pulmonary reexpansion causes xanthine oxidase-induced apoptosis in rat lung. Am J Physiol Lung Cell Mol Physiol 289: L400–L406, 2005.[Abstract/Free Full Text]
  68. Sato N, Suzuki Y, Nishio K, Suzuki K, Naoki K, Takeshita K, Kudo H, Miyao N, Tsumura H, Serizawa H, Suematsu M, Yamaguchi K. Roles of ICAM-1 for abnormal leukocyte recruitment in the microcirculation of bleomycin-induced fibrotic lung injury. Am J Respir Crit Care Med 161: 1681–1688, 2000.[Abstract/Free Full Text]
  69. Shepard JM, Gropper MA, Nicolaysen G, Staub NC, Bhattacharya J. Lung microvascular pressure profile measured by micropuncture in anesthetized dogs. J Appl Physiol 64: 874–879, 1988.[Abstract/Free Full Text]
  70. Sikora L, Johansson AC, Rao SP, Hughes GK, Broide DH, Sriramarao P. A murine model to study leukocyte rolling and intravascular trafficking in lung microvessels. Am J Pathol 162: 2019–2028, 2003.[Abstract/Free Full Text]
  71. Sikora L, Rao SP, Sriramarao P. Selectin-dependent rolling and adhesion of leukocytes in nicotine-exposed microvessels of lung allografts. Am J Physiol Lung Cell Mol Physiol 285: L654–L663, 2003.[Abstract/Free Full Text]
  72. Skerrett SJ, Martin TR, Chi EY, Peschon JJ, Mohler KM, Wilson CB. Role of the type 1 TNF receptor in lung inflammation after inhalation of endotoxin or Pseudomonas aeruginosa. Am J Physiol Lung Cell Mol Physiol 276: L715–L727, 1999.[Abstract/Free Full Text]
  73. St. Croix CM, Pitt BR, Watkins SC. The use of contemporary fluorescent imaging technologies in biomedical research. Med Sci 10: 16–29, 2005.
  74. Stein JV, Rot A, Luo Y, Narasimhaswamy M, Nakano H, Gunn MD, Matsuzawa A, Quackenbush EJ, Dorf ME, von Andrian UH. The CC chemokine thymus-derived chemotactic agent 4 (TCA-4, secondary lymphoid tissue chemokine, 6Ckine, exodus-2) triggers lymphocyte function-associated antigen 1-mediated arrest of rolling T lymphocytes in peripheral lymph node high endothelial venules. J Exp Med 191: 61–76, 2000.[Abstract/Free Full Text]
  75. Stryer L. Fluorescence energy transfer as a spectroscopic ruler. Annu Rev Biochem 47: 819–846, 1978.[CrossRef][Web of Science][Medline]
  76. Su WH, Chen HI, Huang JP, Jen CJ. Endothelial [Ca2+]i signaling during transmigration of polymorphonuclear leukocytes. Blood 96: 3816–3822, 2000.[Abstract/Free Full Text]
  77. Tabuchi A, Kuppe H, Pries AR, Kuebler WM. Intravital microscopy of the murine pulmonary microcirculation (Abstract). J Vasc Res 43, Suppl 1: 7, 2006.
  78. Terry RJ. A thoracic window for observation of the lung in a living animal. Science 90: 43–44, 1939.[Free Full Text]
  79. Togo T, Krasieva TB, Steinhardt RA. A decrease in membrane tension precedes successful cell-membrane repair. Mol Biol Cell 11: 4339–4346, 2000.[Abstract/Free Full Text]
  80. Tsukada H, Ying X, Fu C, Ishikawa S, McKeown-Longo P, Albelda S, Bhattacharya S, Bray BA, Bhattacharya J. Ligation of endothelial alpha v beta 3 integrin increases capillary hydraulic conductivity of rat lung. Circ Res 77: 651–659, 1995.[Abstract/Free Full Text]
  81. Wagner WW Jr. Pulmonary microcirculatory observations in vivo under physiological conditions. J Appl Physiol 26: 375–377, 1969.[Free Full Text]
  82. Wagner WW Jr, Latham LP. Pulmonary capillary recruitment during airway hypoxia in the dog. J Appl Physiol 39: 900–905, 1975.[Abstract/Free Full Text]
  83. Wagner WW Jr, Latham LP, Gillespie MN, Guenther JP, Capen RL. Direct measurement of pulmonary capillary transit times. Science 218: 379–381, 1982.[Abstract/Free Full Text]
  84. Wang PM, Ashino Y, Ichimura H, Bhattacharya J. Rapid alveolar liquid removal by a novel convective mechanism. Am J Physiol Lung Cell Mol Physiol 281: L1327–L1334, 2001.[Abstract/Free Full Text]
  85. Wearn JT, Ernstene AC, Bromer AW, Barr JW, German WJ, Zschiesche LJ. The normal behavior of the pulmonary blood vessels with observations on the intermittence of the flow of blood in arterioles and capillaries. Am J Physiol 109: 236–256, 1934.[Free Full Text]
  86. Wirtz HR, Dobbs LG. Calcium mobilization and exocytosis after one mechanical stretch of lung epithelial cells. Science 250: 1266–1269, 1990.[Abstract/Free Full Text]
  87. Ying X, Minamiya Y, Fu C, Bhattacharya J. Ca2+ waves in lung capillary endothelium. Circ Res 79: 898–908, 1996.[Abstract/Free Full Text]
  88. Zhang Q, Matsuzaki I, Chatterjee S, Fisher AB. Activation of endothelial NADPH oxidase during normoxic lung ischemia is KATP channel dependent. Am J Physiol Lung Cell Mol Physiol 289: L954–L961, 2005.[Abstract/Free Full Text]
  89. Zipfel WR, Williams RM, Christie R, Nikitin AY, Hyman BT, Webb WW. Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation. Proc Natl Acad Sci USA 100: 7075–7080, 2003.[Abstract/Free Full Text]



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