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1Dipartimento di Biologia Strutturale e Funzionale and 2Dipartimento di Scienze Biomediche Sperimentali e Cliniche, Università degli Studi dell'Insubria, Varese, Italy
Submitted 5 December 2005 ; accepted in final form 17 February 2006
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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PTD) was measured in five spontaneously breathing anesthetized rats. PTD was –1.93 cmH2O (SD 0.59) and –3.1 cmH2O (SD 0.82) at end expiration and at end inspiration, respectively, indicating the existence of a pressure gradient directed from the abdominal to the pleural cavity. Morphometrical analysis of the diaphragmatic lymphatic network was performed in the excised diaphragm of three additional rats euthanized with an anesthesia overdose. Optical and electron microscopy revealed that lymphatic submesothelial lacunae and lymphatic capillaries among the skeletal muscles fibers show the ultrastructural features of the so-called initial lymphatic vessels, namely, a discontinuous basal lamina and anchoring filaments linking the outer surface of the endothelial cells to connective tissue or to muscle fibers. Primary unidirectional valves in the wall of the initial lymphatics allow entrance of serosal fluid into the lymphatic network preventing fluid backflow, while unidirectional intraluminar valves in the transverse vessels convey lymph centripetally toward central collecting ducts. The complexity and anatomical arrangement of the two valves system suggests that, despite the existence of a favorable PTD, in the physiological condition no fluid bulk flow takes place between the pleural and peritoneal cavity through the diaphragmatic lymphatic network.
intraluminar lymphatic pressure; serosal fluid pressure; tissue fluid homeostasis
To verify whether a transdiaphragmatic fluid flux might actually occur under physiological conditions, we followed a double approach. On one hand we analyzed, by means of optical and electron microscopy, the morphological arrangement of the diaphragmatic lymphatic network, searching for possible transdiaphragmatic routes. On the other hand, we verified the existence of a transdiaphragmatic pressure gradient potentially supporting a fluid flux between the pleural and peritoneal cavities.
The results of this study, other than providing new information on the possible role of pleuroperitoneal lymphatic connections in the normal pleural and peritoneal fluid turnover, might also clarify some aspects of the development of clinically relevant phenomena related not only to extracellular fluid shift, like ascites or pleural effusion, but also involving cell spreading, like in metastatic tumor cells dissemination.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In a group of animals (n = 5), a heated pneumotachograph (model 8420, Hans Rudolph) equipped with a dedicated pneumotach amplifier (model 1110A, Hans Rudolph) was connected to the tracheal cannula for continuous recording of respiratory flow. The flow signal was then digitized with an analog-to-digital board, integrated through a dedicated LabView software (National Instruments) to obtain respiratory tidal volume and both flow and volume signals, and then displayed on a monitor screen. A blunted tip saline-filled plastic catheter was inserted into the right carotid artery and connected to a physiological pressure transducer (model P23 XL, Gould Electronics). The hydraulic pressure in the pleural and peritoneal space was measured through blunted saline-filled stainless steel cannulas (length: 5 cm; external diameter: 0.5 mm; internal diameter: 0.3 mm) inserted in the supradiaphragmatic and subdiaphragmatic region, respectively, according to a previously described procedure (10, 11). The cannulas were shaped to follow the diaphragmatic dome curvature, and their tip was perforated with three to four holes, whereas the other end was connected through a saline-filled plastic catheter to a physiological pressure transducer. After removal of the skin, the superficial tissues, and the external intercostals muscles at the right side of the thorax, the animals were placed supine, and the pleural cannula was inserted in the 7th-8th intercostal space by being gently pushed through the internal intercostal muscles to position the cannula tip medially over the diaphragmatic dome. The insertion was performed under a flush of saline to avoid air entrance into the pleural space.
Subsequently, the peritoneal cannula was inserted through the lateral wall of the abdomen, and it was driven along the cranial surface of the liver to position the cannula tip in the subdiaphragmatic medial region. The cannulas were placed horizontally at the same height, within 1 cm from the bottom part of the pleural and peritoneal space. The exact location of the recording cannula tip was assessed on opening the pleural and peritoneal cavity at the end of the experiment.
Systemic arterial, pleural, and peritoneal hydraulic pressures were monitored throughout the whole experiment by conveying the signal output from the pressure transducers to the corresponding amplifiers in a signal conditioner (model 6600, Gould Electronics); the pressure signals were then digitized and displayed on the monitor by using the dedicated LabView software. After all of the parameters were continuously recorded for up to 1 h from the insertion of the cannulas, the animals were euthanized with an anesthesia overdose.
Rats belonging to a separate group (n = 3) were euthanized with an overdose of the anesthetic cocktail. The rats used for the pressure recordings were not utilized for the morphological analysis, because the cannula insertion might have somehow damaged or scratched the diaphragmatic surface, thus affecting the result of the morphological analysis. The whole diaphragm was immediately excised and cut into eight large pieces of approximately the same size, belonging to four distinct diaphragmatic regions, as depicted in Fig. 1 : region 1, a ventrolateral portion corresponding to the peripheral muscular area of the ventral diaphragm; region 2, a medial ventral portion corresponding to the tendinous area of the ventral diaphragm; region 3, a dorsolateral muscular region; and region 4, a dorsal medial tendinous region.
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Light microscopy and transmission electron microscopy.
Small diaphragmatic blocks (
5 mm wide) were fixed in 2% glutaraldehyde in 0.1 M Na-cacodylate buffer (pH 7.2) for 2 h at room temperature and then washed in the same buffer and postfixed for 2 h with 1% osmic acid in 0.1 M Na-cacodylate buffer (pH 7.2) at room temperature. After standard steps of serial ethanol dehydration, specimen was embedded in an Epon-Araldite 812 mixture. Semithin sections (750 nm) and thin sections (80 nm) were obtained with a Reichert Ultracut S Ultratome (Leica, Nussloch, Germany) and stained by conventional methods by using, respectively, crystal violet and basic fuchsin or uranyl acetate and lead citrate. The observations were made with a light microscope Olympus BH2 or with a Jeol 1010 electron microscope (Jeol, Tokyo, Japan).
Three tissue samples were taken from each diaphragmatic region, and each tissue sample was sliced into 30 semithin sections for a total of 2,160 slices (270 slices/region) in the three rats used for morphological analysis. All semithin sections were carefully evaluated, and subsequent measurements were performed in 1 of 10 images, for a total of 27 pictures per each region (9 from each rat). The chosen images were selected on the basis of their clearness and of the simultaneous presence of as many as possible structures to be evaluated in the subsequent analysis. In each image the following parameters were measured: 1) the thickness of the whole diaphragm, when allowed by the section; 2) the total diaphragmatic surface area in the section; 3) the thickness of the mesothelial monolayers; 4) the thickness of the pleural and peritoneal submesothelial interstitial space; 5) the length, width, area, and number of pleural and peritoneal submesothelial lacunae; 6) the diameter and the number of the transverse lymphatic vessels; and 7) the number of transverse vessels showing intraluminar valves.
The measurements were performed on digital images of the sections by using an image analysis software (ImageJ software provided by NIH). Briefly, the one-dimensional parameters (thickness, length, width, and diameter) were manually highlighted by the operator on the personal computer monitor screen and then automatically measured by the software. The areas of the diaphragmatic surface and of each submesothelial lacunae appearing in the section were identified by drawing a line precisely along the object perimeter; at this point the object area was automatically measured by the software.
Histochemistry. Some of the diaphragmatic blocks from each of the four regions were immediately embedded in polyfreeze cryostat embedding medium (OCT, Polyscience Europe, Eppelheim, Germany) and stored in liquid nitrogen. Cryosections of diaphragm (10 µm) were obtained with a Reichert Jung Frigocut 2800; slides were immediately used or stored at –20°C. The sections were stained by utilizing histoenzymatic kit Masson Trichrome with aniline blue (Bio-Optica, Milan, Italy) to highlight the presence of collagen and Oil Red O solution for lipid metabolism.
Immunocytochemistry. Frozen sections (10 µm) of diaphragm were incubated for 30 min with 0.3% H2O2 in phosphate-buffered saline (PBS) to inhibit endogenous peroxidase. Successively, samples were washed twice with PBS and incubated for 30 min with PBS containing 3% bovine serum albumin (BSA) and then with primary antibody goat anti-mouse Lyve-1, working dilution 1:20, at room temperature for 1 h in a moist chamber. After incubation with the primary antibody, the specimens were washed and incubated for 1 h at room temperature with the secondary donkey anti-mouse antibody (dilution 1:100) conjugated with peroxidase (Jackson, Immuno Research Laboratories, West Grove, PA). Secondary antibody peroxidase conjugated was visualized by using 3,3'-diaminobenzidine tablet (DAB, Sigma, St. Louis, MO) or TrueBlue (KPL, Gaithersburg, MD). Nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI, working dilution 10 µg/ml). Control sections were incubated in PBS-BSA without the primary antibody.
Data analysis. Data are reported as means (SD). Absolute values were compared by one-way ANOVA using pairwise multiple comparison procedures (Bonferroni t-test). Differences between mean values were considered significant at P < 0.05. Analysis of data variability was performed by calculating the Fisher ratio (F), i.e., the ratio of the variance between different groups and the residual variance. The variability of the mean values is attributable to the variability between animals (or to the low number of animals) in case F is significantly >1. The significance is tested by a P test of the ratio considered significantly different from 1 at P < 0.05.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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PTD) calculated as PTD = Ppl – Pabd. The PTD values attained at end expiration [PTD-E, –1.93 cmH2O (SD 0.59); t = –4.40, P < 0.05], at end inspiration [PTD-I, –3.1 (SD 0.82) cmH2O; t = –10.24, P < 0.01], and over the entire respiratory cycle [–2.29 cmH2O (SD 0.79); t = –5.5, P < 0.01], as well as the PTD pressure swing [1.17 cmH2O (SD 0.22); t = 11.1, P < 0.01] were all significantly different from zero. These data support the existence, throughout the respiratory cycle, of a net pressure gradient directed from the peritoneal to the pleural space.
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Ultrastructural analysis of lymphatic network. Deepening from the pleural and peritoneal mesothelial surfaces into the diaphragmatic tissue (Fig. 3), the lymphatic network is organized in: 1) submesothelial lacunae located within the interstitial space beneath the mesothelial monolayer (Figs. 3, a–c, i); 2) lymphatic capillaries surrounded by a continuous endothelium and located among the skeletal muscle fibers (Fig. 3e); 3) transverse lymphatic ducts that depart perpendicularly from the submesothelial lacunae and connect them to the deep central collectors (Fig. 3, a and f). The wall of transverse ducts, like that of submesothelial lacunae, mostly lacks a continuous endothelium (Fig. 3f); and 4) central collectors located in the deep interstitial space and surrounded by a continuous endothelial layer (Fig. 3, a and f).
The superficial and deep diaphragmatic lymphatic vessels are immersed in connective tissue mainly formed by collagen, as evidenced by the Masson Trichrome reaction (Fig. 3, h and i). Oil red O reaction is used to clearly differentiate lymphatic vessels from adipose tissue cells seldom found in the deep interstitium around the central collectors (Fig. 3, l and m). At the electron microscope, the lymphatic vessels located underneath the mesothelium (Fig. 4 a), the capillary vessels dispersed among muscle fibers (Fig. 4b), and those in the deep interstitium (Fig. 4c) show the same ultrastructural features of the submesothelial vessels belonging to the so-called initial lymphatic vessels. Indeed, the basal lamina is discontinuous and anchoring filaments link endothelial cells to connective tissue or to muscle fibers (Fig. 4, a–c). Endothelial cells are joined by tight junctions (Fig. 4, a and b) or by overlapping junctions (Fig. 4c).
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The F ratio calculated through ANOVA for each of the parameters reported in Table 1 was not significantly different from 1, indicating that the high degree of variability of the mean values reflects a random sampling rather than variability between different rats.
As indicated by the data presented in Table 2, the density of the deeper transverse lymphatic ducts is rather homogenous throughout the diaphragm. However, the transverse ducts draining from the tendinous region of the peritoneal diaphragm display a much higher diameter compared with the other ducts. Independent of their size, density, and origin, transverse lymphatic ducts were almost invariably characterized by the presence of intraluminar unidirectional valves (Table 2 and Fig. 6) conveying lymph centripetally toward the central collecting ducts (Fig. 3). The latter had an average diameter of 24.8 µm (SD 6.6) and run in the deeper diaphragmatic tissue surrounded by a thick [46.2 µm (SD 18.3)] interstitial layer.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The present study allows one to recognize, within the diaphragmatic lymphatic network, various types of vessels that may be classified judging from the morphology of their wall and from the presence of the primary or secondary unidirectional valves. These features, ultimately, identify the vessel types in relation to their absorptive-propulsive function.
Submesothelial lacunae and lymphatic capillaries. The submesothelial lacunae display a discontinuous endothelium adjacent to the mesothelial structures and are characterized, on their mesothelial side, by several discontinuities through the mesothelial layer, presumably corresponding to the so-called stomata (18, 26) connecting the pleural and/or the peritoneal cavities to the lymphatic lumen. The structure of the lacunae wall, delimited by a discontinuous endothelium and apparently devoid of smooth muscle cells, suggests that these structures do not possess the spontaneous vasomotion that characterizes, for instance, the lymphatic mesenteric vessels (3, 7, 8, 27). Hence, fluid entry is supported by a passive hydraulic pressure gradient developing between the pleural and/or peritoneal cavities and the lumen of the submesothelial lacunae. Direct measurements performed on rabbits and rats (13, 17) have demonstrated that the hydraulic pressure in lymphatic diaphragmatic vessels (Plymph), likely corresponding to the pleural submesothelial lacunae, is subatmospheric, amounting to about –5 mmHg; it is worth noting that Plymph is even more negative when the chest is intact at a lung volume corresponding to the functional residual capacity (13).
Whereas it has been shown that during spontaneous breathing, rat intercostal Plymph was significantly lower at end inspiration compared with the end-expiratory value (16), no data on the diaphragmatic Plymph changes associated to the respiratory activity are so far available. In fact, because of obvious experimental difficulties, diaphragmatic Plymph has never been measured during spontaneous breathing. In the micropuncture experiments so far performed on the diaphragmatic lymphatics, the animals were paralyzed, and the chest was necessarily open to allow pipette insertion into the lymphatic vessels or adjacent interstitial space (13, 17); in these conditions the recorded diaphragmatic Plymph values reflected the mechanical stress of the diaphragmatic tissue and its cardiogenic motion (17) but not the respiratory activity. Because of the anatomical arrangement of the diaphragm, it is quite difficult to infer what the diaphragmatic Plymph behavior might be during spontaneous breathing. Indeed, whereas the peripheral skeletal muscle fibers (Fig. 1) contract and shorten on inspiration, they exert an outer stress on the medial tendinous fibers. Hence, one might expect Plymph in the lymphatics of pleural tendinous zone, which display a greater arealacunae-to-areatissue ratio compared with the muscular zone (Table 1), to drop to more subatmospheric values during spontaneous inspiration. With consideration that diaphragmatic Ppl also decreases on increasing lung volume (10, 11), the Plymph drop during inspiration would be very important to guarantee the maintenance of a positive (Ppl – Plymph) gradient and thus of a continuous removal of pleural fluid into the diaphragmatic lymphatics throughout the whole respiratory cycle.
Whereas it has been possible to measure Plymph in the pleural diaphragm, the concavity of the diaphragmatic dome has not yet allowed to directly measure Plymph on its abdominal surface. With consideration that the average radius of curvature of the in situ diaphragm (2 cm in rat) is much larger than its average thickness (about 750 µm, see RESULTS), one might expect the mechanical stress exerted on the diaphragmatic tissue fibers be comparable on the two surfaces of the diaphragm. Because diaphragmatic Plymph depends on tissue stress (13, 16, 17), one might thus expect regional Plymph on the pleural and peritoneal sides to be similar. It is, however, worth noting that, as described in RESULTS, there is a notable difference in the construction of the mesothelial interstitium between the pleural and peritoneal sides. The arrangement of collagen fibers may be a major component for developing the mechanical environment. A difference in arrangement of the fibers from loosely connected bundles to tighter bundles aligned in a parallel fashion (see RESULTS) could create a significant difference in mechanical stresses on pleural versus peritoneal lacunae. In any case, one might observe that under physiological conditions, fluid turnover and corresponding lymphatic removal are greater in the peritoneal compared with the pleural cavity (15). This difference might be compatible with a greater distribution of the peritoneal lymphatic network and/or a higher driving pressure gradients promoting lymph formation. The present data in fact describe a more developed initial lymphatic network, at least in terms of vessel density and arealacunae-to-areatissue ratio, over the peritoneal compared with the pleural side of the diaphragm (Table 1), confirming the previous observation on the distribution of diaphragmatic lacunae (14) and stomata (18) in normal rabbits. Concerning the pressure gradients arising between the peritoneal cavity and the diaphragmatic lumen, at this state of the art one may only comment that because Pabd > Ppl during the whole respiratory cycle (see RESULTS), Pabd – Plymph might also be greater than Ppl – Plymph provided that Plymph is similar on the two sides of the diaphragm.
Whereas the pleural and peritoneal fluid enters the diaphragmatic lymphatic network through the stomata and the submesothelial lacunae, interstitial tissue fluid and solutes enter directly into the initial lymphatics observed along the perimeter of the skeletal muscle cells. Like the lacunae, the initial lymphatics display anchoring filaments on their outer wall (Fig. 3); these collagen filaments are highly sensitive to interstitial stress and exert radial tension on the vessel increasing their luminal volume (1, 22). In addition, the electron microscopy analysis revealed the existence, only in the smallest submesothelial lymphatic lacunae (1–20 µm diameter) and in the initial lymphatic (1–10 µm diameter, Fig. 6) dispersed among the muscle fibers, of the so-called primary valves (9, 23) formed by two adjacent endothelial cells that display overlapping cytoplasmic extensions at their junction. The presence of these primary valves has been already demonstrated in the cremaster muscle of rat by microsphere injection (24), but it was never observed before in the diaphragmatic tissue. In conjunction with the secondary intralymphatic valves, the primary valves facilitate the unidirectional transport of fluids and solutes from the interstitium into initial lymphatic vessels preventing in the mean time fluid backflow (9). The absence of smooth muscle cells from the wall of most of the diaphragmatic lymphatic vessels suggests that in the diaphragm lymph formation and propulsion are promoted by tissue motion (extrinsic mechanism) rather than by intrinsic contraction of the vessel wall (intrinsic mechanism).
Collecting transport system. The morphological evidences provided in the present study indicate that the transverse lymphatic ducts and the central collectors receive the newly formed lymph from the submesothelial lacunae and from the internal and deeper interstitium, functioning as a collector system for the lymph to be carried out of the diaphragm through extra diaphragmatic collectors, mostly the right and partly the left lymphatic duct.
The present evidence indicates that lymphatic smooth muscles cells, albeit previously detected in some diaphragmatic vessels (20), are not ubiquitously distributed in the diaphragmatic vessel, as instead observed in other tissues for vessels of similar size and function (1, 22). Hence, not only lymph formation, but also its progression, is presumably mostly dependent on tissue motion, including cardiogenic oscillations (17). In analogy with what is shown for the intercostal lymphatics the function of which is promoted mainly by the active contraction of the inspiratory intercostal muscles rather than by the tissue stress related to the passive change in lung volume (16), the major contribution to diaphragmatic extrinsic mechanisms likely consists in the rhythmic contraction of the diaphragmatic skeletal muscle fibers during inspiration.
It is worth noting that the density of the submesothelial lacunae (Table 1) and of the transverse vessels (Table 2) is, notwithstanding some variability, rather homogeneous in the muscular and tendinous regions. However, because of the radial disposition of the muscular fibers (Fig. 1), the mechanical stress elicited by the muscular contraction during the respiratory cycle is likely very far from being uniform. Indeed, the medial tendinous fibers are radially stressed outward when the muscular fibers contract during inspiration and vice versa during expiration. A thorough evaluation of the stress arising in the diaphragm during a respiratory cycle is far too complex to be addressed in the present study. However, the uniform distribution of the lymphatic structures suggests that lymph formation and progression could be guaranteed, in either the muscular or the tendinous regions, throughout the whole respiratory cycle.
Pleuroperitoneal lymphatic communications.
The existence of a net transdiaphragmatic pressure gradient (PTD) potentially driving fluid from the peritoneal to the pleural cavity might lead one to hypothesize that a net transfer of fluid might take place from the peritoneal to the pleural space through direct transdiaphragmatic lymphatic pathways. However, fluid entrance into the submesothelial lacunae is driven by the pressure difference (Pabd – Plymph and Ppl – Plymph) between the serosal compartment and the lymphatic lumen. In normal conditions, diaphragmatic Plymph is significantly lower than both Ppl and Pabd (10, 13, 15, 17). Hence, under physiological conditions, fluid backflow from the lacunae into the pleural cavity would be prevented both by the Ppl – Plymph pressure gradient and by the double unidirectional valve system.
The situation might be different in some pathological conditions, like the development of hydrothoraces as clinical complication of peritoneal dialysis or ascitis, that have been regarded as secondary to the existence of direct transdiaphragmatic lymphatic pathways allowing exchange of fluid, solutes, and even cells between the pleural and peritoneal cavities (4, 25). A peritoneal fluid volume expansion increases Pabd and thus Pabd – Plymph, resulting in an increased lymph flow rate. In these instances, direct pleuroperitoneal connections might form through dilated transverse lymphatics as a result of valve system incontinence, abnormalities of the lymphatic structure and/or lymph flow saturation, leading to failure of the lymphatic drainage.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
* A. Grimaldi and A. Moriondo equally contributed to the research. 
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REFERENCES |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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