Functional Structure of the Peritoneum as a Dialyzing Membrane

Gotloib, L.

doi:10.1007/978-0-387-78940-8_5

L. Gotloib³

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4 Citations

More than a century ago, Robinson [1], after summarizing more than two centuries of research, defined the diverse natural functions of the peritoneum as follows: a) to regulate fluid for nutrient and mechanical purposes: b) to facilitate motion; c) to minimize friction, and d) to conduct vessels and nerves to the viscera.

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The Evolution of Peritoneal Dialysis Solutions

Physiology of Peritoneal Dialysis

Monitoring the Functional Status of the Peritoneum

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More than a century ago, Robinson [1], after summarizing more than two centuries of research, defined the diverse natural functions of the peritoneum as follows: a) to regulate fluid for nutrient and mechanical purposes: b) to facilitate motion; c) to minimize friction, and d) to conduct vessels and nerves to the viscera.

Several medical and scientific developments that occurred during the 20th century originated a new approach for the peritoneum being used as a dialysing membrane for long-term life support [2–6]. These same developments created the need for a deeper understanding of peritoneal structure and function.

The peritoneum is a serous membrane embryologically derived from mesenchyma and composed of thin layers of connective tissue covered by a sheet of mesothelium [7]. When the membrane is folded, forming the omentum and the mesentery, both luminal surfaces are covered by mesothelium.

The anatomical peritoneal surface area for the human adult is considered to range between 2.08 [8] and 1.72 m² [9], with a ratio of area/body weight of 0.284. The intestinal mesothelium, together with that of mesentery, makes up to 49% of the total mesothelial area [10]. For infants having a body weight of 2,700–2,900 g, the total peritoneal surface was found to oscillate between 0.106 [10] and 0.151 m² [8], with an area to body weight ratio that fluctuates between 0.383 [10] and 0.522. In infants the contribution of intestine and mesentery to the total surface area is 67.5% [10].

However, from the functional point of view, vis-à-vis peritoneal dialysis, it may well be that the peritoneal area of contact with the dialysis solutions were substantially lower than the anatomical one. This concept, postulated by Krediet et al., was defined as the effective surface area [11]. This hypothesis finds strong support in the elegant study performed by Chagnac et al. [12] showing that the peritoneal surface actively involved in the dialytic process, estimated in six CAPD patients, was 0.55∼0.04 m², about one third of the area measured in anatomical studies. Interestingly, other investigators reached similar conclusions in experiments performed in rats [13].

Peritoneal thickness is not uniform and varies according to the area examined. Measurements are quite problematic in parietal and diaphragmatic peritoneum due to the considerable amount of connective tissue, and at times fat, intervening between the peritoneum itself and the underlying tissue (Fig. 5.1). The submesothelial connective tissue layer of visceral peritoneum is firmly bound to the fibrous tissue of the viscus. Therefore, the mesentery, having mesothelial lining on both surfaces and including its trabecular connective framework, appears to be the most appropriate peritoneal portion for estimation of membrane thickness which, in the rabbit, ranges between 30 and 38 μm [14, 15] (Figs. 5.2 and 5.3).

figure 5_1_978-0-387-78940-8 — **Fig. 5.1**

figure 5_2_978-0-387-78940-8 — **Fig. 5.2**

figure 5_3_978-0-387-78940-8 — **Fig. 5.3**

Normal Mesothelium

Electron microscopic studies performed on mouse embryo disclosed that the mesothelium is derived from mesenchymal cells that become flattened, form their own basement membrane, and develop tight junctions as well as desmosomes [16] (Fig. 5.4, inset). Both pinocytotic vesicles and rough endoplasmic reticulum were present. Yolk sac of human embryos at the 5th–7th week of gestation also exhibit flattened mesothelial cells lying on a hyaline, homogeneous basement membrane [17, 18].

figure 5_4_978-0-387-78940-8 — **Fig. 5.4**

The cell plasmalemma, when stained specifically, shows the typical trilaminar structure observed in all biological cell membranes [19]. The normal mesothelium occasionally shows macrophages implanted on the luminal peritoneal surface instead of mesothelial cells (Fig. 5.5).

figure 5_5_978-0-387-78940-8 — **Fig. 5.5**

The luminal aspect of the mesothelial cell plasmalemma has numerous cytoplasmic extensions: the microvilli (Figs. 5.2, 5.3, and 5.4), whose existence was originally reported by Kolossow [20] and many years later confirmed by electron microscopy on the serosa covering the rat oviduct [21, 22]. Even though microvilli are more frequently observed in visceral than in parietal peritoneum [23, 24], their distribution is variable and fluctuates from very numerous to completely absent [24, 25]. It should be taken into account, however, that microvilli are extremely sensitive to minor injury or even to dryness, and can therefore be lost from the cell surface if removal and handling of samples are not done with extremely careful techniques. On the other hand, loss of microvilli, as described in continuous ambulatory peritoneal dialysis (CAPD)patients [26] (Fig. 5.6), represent an early sign of impending apoptosis [27–29] that can be easily identified in mesothelial cell imprints (Fig. 5.7).

figure 5_6_978-0-387-78940-8 — **Fig. 5.6**

figure 5_7_978-0-387-78940-8 — **Fig. 5.7**

Light microscopy applied to the observation of resting mesothelium imprints [30] shows a continuous monolayer made up mostly of polygonal mononuclear cells (Fig. 5.8), showing, in mice visceral peritoneum, a density of about 300,000 cells/cm² [31]. The number of mesothelial cells per unit area seems higher on the visceral than on the parietal peritoneal surface. Of those cells, 1–2% are binucleated (Fig. 5.8, lower left inset), whereas cells showing three nuclei can be observed (Fig. 5.8).

figure 5_8_978-0-387-78940-8 — **Fig. 5.8**

Under normal circumstances the cell population of the monolayer is not stained by vital dyes such as Trypan Blue (Fig. 5.9). This is an indication of their viability. In perpendicular cuts observed under light microscopy, the resting normal mesothelium appears as a continuous layer formed by flattened cells that are apparently elongated, as a result of the angle of section (Fig. 5.10). The mesothelial sheet lies on a layer of connective interstitial tissue (Fig. 5.10), the thickness of which varies in the different portions of the peritoneum (Figs 5.1 and 5.3). The relevance of this point on peritoneal permeability will be discussed later.

figure 5_9_978-0-387-78940-8 — **Fig. 5.9**

figure 5_10_978-0-387-78940-8 — **Fig. 5.10**

Thickness of mesothelial cells in the rabbit ranges between 0.6 and 2 μm [14, 15] (Fig. 5.11).

figure 5_11_978-0-387-78940-8 — **Fig. 5.11**

The human omentum has not yet been studied in great depth. However, some ultrastructural investigations performed in mice and rats [22, 32] seem to indicate that there is little variation between species [33] and that, in mice, omental mesothelial cells can transiently increase their population of microvilli up to seven-fold, suggesting that their concentration in any given area could reflect functional adaptation rather than static structural variation [34].

The presence of pinocytotic vesicles in microvilli has been both reported [21, 23, 35] and denied [34].

Experimental studies done in mice and rats [35–37] using cationic tracers such as ruthenium red (MW 551 da) and cationized ferritin (MW 445 da) revealed the existence of anionic fixed charges on the luminal surface of the microvilli cytoplasmic membrane (Fig. 5.12, inset). This cell membrane coating, or glycocalyx, composed of fine fibers that are continuous with the membrane itself [38], furnishes the microvilli surface with electronegative charge, which most probably plays a significant role in the transperitoneal transfer of anionic macromolecules such as plasma proteins [36, 39], as well as in that of charged small molecules, as suggested by Curry and Michel [40] in their fiber matrix model of capillary permeability. This surface charge is substantially reduced in cells undergoing apoptosis [41]. The relevance of these charges upon peritoneal permeability will be discussed later.

figure 5_12_978-0-387-78940-8 — **Fig. 5.12**

Length of microvilli in rodents ranges between 0.42 and 2.7 μm, and their average diameter is 0.1 μm [14, 21, 23, 32]. We have observed a similar range in adult humans. However, mesothelial cells of human embryos (5th–7th week of gestation) showed microvilli up to 3.5 μm long [17].

It has been estimated that microvilli present in the striated border of intestinal epithelium increase the surface area of the intestine by a factor of 20 [42]. Consequently, it has been speculated that mesothelial microvilli could increase the actual peritoneal surface up to 40 m² [43].

Plasmalemma of mesothelial cells, like that of microvilli, shows electronegatively charged glycocalyx (Fig. 5.12) [35–37, 44].

Plasmalemmal vesicles, or caveolae, originally described by Lewis [45] in macrophages of rat omentum, are conspicuously present in mesothelial cells at both the basal and luminal borders, as well as in the paranuclear cytoplasm [21–23, 32, 46–48] (Fig. 5.2, inset). Their average diameter is approximately 0.717 μm [14]. At times, pinocytotic vesicles appear clustered together and communicating with each other (Fig. 5.2, inset). Occasionally they appear forming transcellular channels similar to those described in endothelial cells of blood capillaries [49, 50] (Fig. 5.13, inset), apparently communicating both aspects, luminal and abluminal, of the mesothelial cell. These channels can be formed by a chain of several vesicles (Fig. 5.13, inset) or just by two adjoining vesicles. Often pinocytotic vesicles appear to open through the plasma membrane into the luminal or abluminal aspect of the cell (Fig. 5.2, inset; Fig. 5.12), as well as into the intercellular space (Fig. 5.12), exhibiting a neck and a mouth whose respective average diameters are 0.176 and 0.028 μm [14]. With respect to the density distribution of these caveolae, it has been suggested that the parietal mesothelium is less well endowed than the visceral [47].

figure 5_13_978-0-387-78940-8 — **Fig. 5.13**

Palade [51] first proposed that a large part of the macromolecular transport across capillary walls could be attributed to exchange of pinocytotic vesicles between the internal and external surfaces of endothelial cells. This concept was repeatedly applied to the mesothelium. Several electron-dense tracers such as native ferritin [48], iron dextran [14, 32], and melanin [22] were found randomly distributed within pinocytotic vesicles of mesothelial cells after being injected intraperitoneally. Casley-Smith and Chin [47] calculated that the median transit time of vesicles through mesothelial cells ranges between 3 and 5 s, and that approximately 40% of the released vesicles reach the cytoplasmic membrane on the opposite side of the cell. It was even observed that metabolic inhibitors such as dinitrophenol, poisons (cyanide), or slow cooling to 0°C did not completely preclude the uptake of electrondense macromolecules by pinocytosis [48, 52]. This information, supporting Palade’s prediction [51] that vesicles could be the structural equivalent of the large pore [53], was challenged by stereological analysis of plasmalemmal vesicles. This study apparently showed that vesicles represent merely invaginations of the plasmalemma from both sides of the capillary wall in frog mesentery [54]. It was suggested that this organization of the vesicular system is incompatible with the concept that macromolecules could be transferred across cells by vesicular transport. The methodology followed in this study has been reviewed and criticized, and its conclusions have been refuted [55].

Furthermore, a huge body of scientifically based evidence indicates that endocytosis, transcytosis, as well as potocytosis (an endocytic pathway that utilizes phosphatylinositol anchored membrane proteins and plasmalemmal vesicles or caveolae to concentrate and internalize small molecules) are basic mechanisms used by cells to carry in, out, and through the cytoplasm a variety of substances [56]. The following part of our description applies to both mesothelium and endothelium.

Work done basically during the past decade shed new light on the intimal structure of pinocytotic vesicles. Even though their morphometric parameters are more or less homogeneous, differences in nature, function, and biochemical structure identified at least two kinds of vesicles showing distinctive characteristics.

Caveolae or plasmalemmal vesicles are membrane domains that represent a subcompartment of the plasma membrane [57], characteristic of all vascular endothelium [58]. In capillary endothelial cells, morphological studies indicate that caveolae are effectors of transcytosis of certain macromolecules across the microvascular endothelium: native as well as modified albumins [59–67], low-density lipoprotein (LDL) [68–71], protein hormones [72, 73], AGE [74], as well as orosomucoid [75], a 41 kDa glycoprotein that qualifies as a probe for the postulated large pore [76–78]. Furthermore, endocytosis and transcytosis of albumin–gold complexes have been observed in mice peritoneal mesothelium [79] (Fig. 5.14, inset; Fig. 5.15, inset).

figure 5_14_978-0-387-78940-8 — **Fig. 5.14**

figure 5_15_978-0-387-78940-8 — **Fig. 5.15**

Schnitzer and Oh have demonstrated that transendothelial transport of native albumin through caveolae in both experimental situations (in vivo as well as in the in vitro set-up) is dependent on the interactions of the probe with the endothelial cell surface protein albondin, a 60 kDa albumin binding protein formerly called gp60 [63]. Other binding proteins, gp30 and gp18, appear to mediate the attachment, endocytosis, and degradation of modified albumin. These vesicular carriers require key intracellular components that are sensitive to alkylation with N-ethylmaleimide. Indeed, this substance has been shown to substantially inhibit native albumin (MW: 67 kDa, r = 36 Å) and ferritin (MW ∼ 500 kDa, r = 100–110 Å) uptake, both transcytosed by caveolae [80]. Additional experiments have shown that transcytosis and capillary permeability of insulin and albumin are selectively inhibited by filipin, a complex of polyene antibiotics obtained from Streptomyces filipenensis, but does not affect endocytosis mediated by the clathrin-coated vesicles [81]. This concept that identifies two different vesicular pathways is completed with the recent discovery of caveolin, the major structural caveolar protein [82]. This substance is a 22 kDa integral protein that represents a subcompartment of the plasma membrane [57, 83]. Basically, it is a component of the coating covering the luminal aspect of caveolae [84] that, when specifically stained by immunocytochemical methods, serves as a useful marker to draw the diagnostic line between caveolae and other pinocytotic related structures, e.g., coated vesicles [85–87].

Coated pits and coated vesicles (Fig. 5.14, left and right insets) remain the most extensively characterized transport vesicles. They are involved in the intracellular transport of membrane proteins between a variety of membrane components, mediate endocytosis of transmembrane receptors, and transport newly synthesized lysosomal hydrolases from the trans-Golgi network to lysosomes [88]. The luminal coat contains at least six polypeptides, in addition to the above-mentioned 180 kDa polypeptide clathrin [89, 90]. This type of vesicle is also involved in receptor-mediated endocytosis. Cell surface mediators operate endocytosis clusters into clathrin-coated pits, which pinch off to form vesicles that transport the receptors and their ligand [91]. The complex process of invagination, constriction and budding of clathrin-coated vesicles employs the coordinated actions of several proteins. The best characterized of them is the expanding family of dynamin guanosine triphosphate phosphatases (GTPase), essential for receptor-mediated endocytosis [92, 93]. This enzyme appears to be assembled around the necks of clathrin-coated pits, and assists in pinching vesicles from the plasma membrane [93]. Recently published information suggests that dynamins mediate both clathrin-dependent endocytosis and the internalization of caveolae in microvascular endothelial cells [94, 95].

Some 70 years ago [96], it was suggested that junctions between capillary endothelial cells should be considered the main pathway for exchanges across the microvascular wall. This concept was later extended to the peritoneal blood microvessels and mesothelium, and extensively analyzed within the frame of the two [77, 78], and lately, the three [97] pore size model of capillary and/or peritoneal permeability. (Assumed pores size: large pore: > 150 Å; small pore: up to 40–45 Å; ultra-small pore: 2–5 Å.) To date, physiological studies and mathematical models have failed to convincingly identify the morphological equivalents of the hypothetical cylindrical water-filled pores [76]. On the other hand, however, this short review of the topic testifies that, at least for protein traffic, the vesicular carried hypothesis has been largely proven and accepted in the last few years [80]. Basically, that caveolae and transcellular channels function as a continuous operating conveyor belt, fusing with each other [49, 50, 98, 99], and moving through the cell [98, 99]. The source of energy fuelling vesicular movement remains one of the many questions still open [98–104].

Furthermore, the demonstrated presence of glucose transporters in mesothelial cells [105] furnish further support regarding the active role of the monolayer in solute’s transport from and to the peritoneal cavity. Specialized transporter proteins, which are the products of two closely related genes, UT-A (Sic 14a2) and UT-B (Sic 14a1), modulate the movement of urea across cell membranes. Up to date, five UT-A isoforms have been identified in most tissues [106]. It may be speculated that UT-A transporters are also present in the mesothelium. This hypothesis deserves to be explored.

Additionally, recently published evidence demonstrated not only the presence of aquaporin channels in mesothelial cells, but also that their expression can be modulated by both osmotic and non-osmotic stimulation [107]. The relevance of these channels for peritoneal permeability will be analyzed in the section dealing with peritoneal microvasculature. Their presence in mesothelial cells is one more indication giving support to Henle’s prediction that the essential anatomy and physiology of the peritoneum are located in its “endothelia” [108].

Simionescu et al. [109] showed the existence of differentiated microdomains on the luminal surface of capillary endothelium where they found a distinct and preferential distribution of electronegative fixed charges, also called anionic sites. Cationic tracers, which did not bind to caveolae or to transcellular channels, decorated the luminal glycocalyx, coated pits, and coated vesicles [98, 103, 109]. Recent studies applying cationic tracers such as ruthenium red and cationized ferritin in rat and mouse peritoneum also showed a preferential distribution of negative charges at the level of the mesothelial cells luminal surface [36, 37, 44] (Figs. 5.12 and 5.16). Density of these surface plasmalemmal charges is substantially reduced in cells undergoing apoptosis [41].

figure 5_16_978-0-387-78940-8 — **Fig. 5.16**

Mesothelial cell boundaries are tortuous, with adjacent cells often tending to overlap (Fig. 5.4, inset; Fig. 5.12). Tight junctions close the luminal side of the intercellular boundaries [14, 21, 32] (Figs. 5.4 and 5.12). When studied in the horizontal plane by using the freeze-fracture technique, these junctional contact areas were defined as cell extensions and finger-like processes, overlapping into the adjacent cell body. Cell processes were wedge-shaped and numerous, and the cell periphery appeared serrated [110]. Desmosomes have also been observed near the cellular luminal front [14, 23, 25, 32] (Fig. 5.17) and so have gap junctions [25]. The abluminal portions of cell interfaces usually show an open intercellular infundibulum. Completely open intercellular interphases have not been observed in normal, resting mesothelium [14, 21, 32]. Even desquamated mesothelial cells showing severe degenerative changes can keep their junctional system almost intact (Fig. 5.18). These junctional morphological features are, however, different from those observed between mesothelial cells covering the diaphragmatic lymphatic lacunae, which are more cuboidal and prominent than mesothelial cells observed in other areas of the peritoneal surface.

figure 5_17_978-0-387-78940-8 — **Fig. 5.17**

figure 5_18_978-0-387-78940-8 — **Fig. 5.18**

The existence of stomata (open intermesothelial communications between the abdominal cavity and the submesothelial diaphragmatic lymphatics), predicted by William Hewson [1] 100 years before being discovered by Von Recklinghausen [111], have been the subject of a long and rich controversy along the years. Accepted by some [112–114] and denied by others [115–117], it was not until the advent of electron microscopy that their existence was demonstrated [44, 118, 119]. Scanning electron microscopy disclosed the patent intermesothelial junctions forming gaps whose average diameter ranged between 4 and 12 μm [118, 119] and circumscribed by cuboidal mesothelial cells. These gaps open into submesothelial lymphatics [44] and have not been observed in diaphragmatic mesothelium covering nonlacunar areas [119]. Additional studies have shown the passage of particles from the abdominal cavity into the submesothelial diaphragmatic lymphatics [114, 120]. These studies also confirm the results of experiments performed by Allen [113], who demonstrated the passage of frog erythrocytes through stomata of the mouse diaphragmatic peritoneum, and their appearance within submesothelial lymphatics. This pathway paved the way for intraperitoneal blood transfusions that have been successfully performed in fetuses [121, 122], human adults [123], rats, mice, dogs, and lambs [124, 125]. On the other hand, intraperitoneal malignant cells [126] and bacteria [127] also leave the abdominal cavity on their way to the central venous circulation, through diaphragmatic stomata. The same pathway applies for absorption of albumin–gold complexes injected into the peritoneal cavity (Fig. 5.15) [79]. These structures can be found only between mesothelial cells overlying lacunae.

At the sites of stomata and their channels, mesothelial and lymphatic endothelial cells contain actin-like filaments [128] assumed to induce cell contraction, opening the stomatal pathway for the passage of macromolecules and cells. Cationized ferritin has been observed decorating the glycocalyx of mesothelial and lymphatic endothelial cells located along the stomata, as well as the coated pits and coated vesicles of both types of cells [44, 129]. It should be noted that the presence of stomata has been recently detected in mouse mesenteric mesothelium [130], in omental, ovaric, and pelvic peritoneum, as well as in that covering the anterior liver surface and the anterior abdominal wall [131, 132]. Therefore, it may be assumed that all these extradiaphragmatic openings contribute to the absorptive capacity of the entire peritoneal membrane. Albumin–gold complexes appear to be absorbed also from the peritoneal space through stomata [79] (Fig. 5.15), even though the capability of this pathway for the uptake of the probe did not seem to be much higher than that shown by nonstomatal mesothelial infundibular junctions that contained only 1% of the injected tracer.

Stomata have been ascribed the role of a preferential pathway for the output of fluids, cells, particles, and bacteria from the abdominal cavity [133]. However, the luminal surface of mesothelial cells (which limits the gaps), after staining with cationized ferritin, displayed dense labeling of their cytoplasmic plasmalemma as well as coated pits and coated vesicles. The same cationic tracer also decorated the lymphatic endothelial plasmalemma, which circumscribed the stomatal openings [44]. If this is so, the passage of solutes through stomata is most likely dependent not only on molecular weight, size, and shape, but also on electric charge [44].

Studies in rat and mouse perfused with ruthenium red revealed that intermesothelial cell junctions were, in general, stained just at the level of their infundibulum, even though the dye now and then decorated the junctional complex, staining approximately 50% of its length [37] (Fig. 5.12).

Nuclei are generally located in the central region of mesothelial cells, showing an elongated, oval, or reniform appearance with occasional irregularities in their outlines and sometimes protrusions and indentations (Fig. 5.11). The chromatin is fine, evenly distributed and forms a dense rim around the nuclear membrane (Fig. 5.11). In normal unexposed mesothelium, around 2% of cells are binucleated [31] (Fig. 5.8). Nucleoli have been reported both as present and absent [21, 32]. However, studies performed in imprints [31] showed that they are present and that their number ranges between 6 and 8 (Fig. 5.7). Rough endoplasmic reticulum and ribosomes are dispersed in the cytoplasm. Mitochondria and the Golgi complex are evident mainly in perinuclear areas (Fig. 5.6). Although seldom observed, isolated cilia may emerge from the luminal aspect of mesothelial cells, showing in their cytoplasmic part the axial microtubule as well as the attached basal body (Fig. 5.4). More frequently observed in splenic mesothelium [134], their functional significance is still unknown [135].

The submesothelial basement membrane, originally described by Todd and Bowman [136], and later reported as hyaline, homogeneous, one-layered, and continuous [112, 128], with an average thickness of approximately 40 nm for mouse and rabbit peritoneum [14, 22], normally appears lying under the mesothelial layer of visceral, parietal, and diaphragmatic peritoneum [137] (Figs. 5.11 and 5.17). As an exception, the functional significance of which is still unknown, the omental mesothelium of mice and humans lacks basement membrane [22, 138].

Submesothelial basement membrane of visceral, parietal, and diaphragmatic peritoneum of rat and mouse, perfused with the cationic tracer ruthenium red, consistently showed anionic charges periodically distributed along the lamina rara externa and interna, most of the time, forming double rows [36, 37] (Fig. 5.12).

The reported average diameter of ruthenium red–stained particles in the basement membrane was 2.7 nm, whereas the average distance measured between the one-row oriented basal lamina dye particles ranged between 65 and 90 nm, not far from the interval value of 60 nm observed using the same tracer in rats [139] and human kidney glomeruli [140].

The fact that these charges are, as stated above, distributed along both aspects of the basement membrane implies that the charge-free interval is actually smaller than the mean distance calculated for each membrane layer. The electric field of each particle of ruthenium is around 8–10 nm, and charge discrimination for negative tracers is effective for substances with a molecular radius around 1 nm, corresponding approximately to a globular molecule showing a molecular weight of 2 kDa [141]. In this sense it should be taken into account that the radius of macromolecular anionic albumin is 3.6 nm, whereas its molecular weight is 67 kDa.

It should be noted that the density distribution of these anionic fixed charges of the basement membrane almost disappears during the acute inflammatory reaction secondary to septic peritonitis [142] (Fig. 5.19), and is substantially reduced in rats, soon after 4 months of streptozotocin-induced, uncontrolled diabetes [143] (Fig. 5.16).

figure 5_19_978-0-387-78940-8 — **Fig. 5.19**

The relevance of the electronegative charge of the mesothelial monolayer upon the peritoneal permeability to anionic plasma proteins will be discussed in the section dealing with microvascular permeability.

Reduplicated submesothelial basement membrane has been observed in diabetic and nondiabetic chronic uremic patients treated by maintenance peritoneal dialysis [144, 145] (Fig. 5.20). It has been shown that perivascular basement membrane thickness increases with age [146, 147] as well as in the direction of head to foot [147, 148]. This same ultrastructural alteration has been observed in diabetics [147, 149]. It has been suggested that diabetes alone is not responsible for excessive accumulation of basement membrane associated with aging [150]. Therefore, it could be claimed that the reduplication of basement membrane observed in human mesothelium is a by-product of cell renewal regardless of the cause of cell death that triggers the process of repopulation [144, 151]. However, the fact that this phenomenon was also detected in the submesothelial basement membrane of diabetic rats suggests that a high glucose content in the extracellular fluid appears to be related to the mechanism(s) leading to these changes [143].

figure 5_20_978-0-387-78940-8 — **Fig. 5.20**

Interstitium

Connective tissue, which originates from mesenchyma, is composed of cells and fibers embedded in an amorphous substance. The main connective tissue cell is the fibroblast and the main fiber is collagen [152].

The submesothelial connective tissue normally has a low cell population surrounded by high-molecular-weight intercellular material. Fibroblasts, mast cells in the proximity of blood microvessels (Fig. 5.21), occasional monocytes and macrophages (Fig. 5.2) are frequently observed.

figure 5_21_978-0-387-78940-8 — **Fig. 5.21**

Substantial amounts of quite compact bundles of collagen are usually interposed between the blood microvessels and the mesothelial layer (Figs. 5.1, 5.2, and 5.3). The collagen density distribution in the different regions of visceral peritoneum is quite variable [146].

The macromolecular common denomination of connective tissues is a broad molecular class of polyanions: the tissue polysaccharides. They form a gel-like structure with the collagen fibers [153], which, when stained with ruthenium red, shows the presence of anionic fixed charges [37].

Thickness of the interstitial layer is extremely variable in the different portions of the peritoneum. This heterogeneity can also be applied to the distances separating the submesothelial blood vessels from the peritoneal cavity, ranging between 1–2 μm to ≥ 30 μm (Fig. 5.1). It should be noted that restriction of molecular movement through the interstitial tissue and its progression from or to the microvasculature is affected not only by their molecular weight, shape, and electric charge, but also by the length of the pathway. According to Fick’s law of diffusion, it is the difference in concentration per unit of distance (the concentration gradient) that determines the rate of movement of the solute. If we double the distance over which the same concentration difference occurred, the gradient and, therefore, the rate of transfer, would be cut in half. Therefore, the relevance of the interstitial compartment thickness in the transperitoneal transfer of solutes may well be critical [78, 154], and diffusion of solutes coming out from capillaries far from the abdominal cavity could be rendered useful only in long-dwell exchanges, like those performed in CAPD.

The question of the interstitium in terms of plasma to lymph traffic of macromolecules has been basically investigated in lung interstitial tissue [155], which, at physiological pH, has the properties of a negatively charged membrane [156]. Therefore, the polyanionic glycosaminoglycans (mainly hyaluronan) and glycoproteins located in the interstitial ground substance have the capability of influencing the interstitial distribution of volumes of plasma proteins coming out from the intravascular compartment, according to their molecular charge [157]. It has been suggested that these glycosaminoglycans restrict free diffusion through the interstitium [158] and can both reduce the interstitial distribution volume of anionic plasma proteins, and retard the plasma-lymph traffic of cationic macromolecules [155]. Anyway, the effect of protein charge on the interstitial hydraulic conductivity has been only partly clarified.

The extremely low and, at times, negative interstitial pressure (0 to –4 mm Hg) [159–161] represents, together with the capillary permselectivity and the lymphatic drainage, one of the three key factors modulating the plasma-to-lymph fluid traffic, therefore, preventing the formation of interstitial edema [162, 163]. Specifically, during peritoneal dialysis, studies by Flessner [164] have shown that transfer of small solutes through the tortuous interstitial pathways is primarily by diffusion, and that convection may contribute to overall transport in parietal tissue. As stated above, in normal conditions the interstitium has a hydrostatic pressure near 0 [159, 164]. During clinical peritoneal dialysis the intra-abdominal pressure ranges between 4 and 10 cm H₂O [165, 166], thus creating a pressure gradient that drives fluid as well as solutes out of the peritoneal cavity to the interstitium. Thus, fluid loss from the abdominal cavity to the periperitoneal interstitial space is directly proportional to intra-abdominal pressures higher than 2 cm H₂O [166].

Blood Microvessels

Capillaries of human and rodent parietal [167] and visceral peritoneum [43, 168] have been reported to be of the continuous type (Figs. 5.1, 5.3, 5.22–5.24), according to the classification of Majno [169]. However, the existence of fenestrated capillaries in human parietal and rabbit diaphragmatic peritoneum (Figs. 5.1 and 5.25), as well as in mouse mesentery [170–172] has been reported. The incidence of fenestrated capillaries in human parietal peritoneum (Fig. 5.22, inset) appears to be low (1.7% of the total number of capillaries) [172]. The reported density of fenestrated microvessels in mouse mesentery and rabbit diaphragmatic peritoneum ranged between 26 and 29% of the observed capillaries, whereas their presence in parietal peritoneum of nondialyzed uremic patients was only 1.7%. It should be noted, however, that the anterior abdominal wall of humans comprises less than 4% of the peritoneal surface area [10]. Diameter of fenestrae, which ranged between 60 and 90 nm, is well within the range of fenestrae observed in other capillary beds: 40–70 nm in renal peritubular capillaries [173], glomerular capillaries [169], and rabbit submandibular gland [174]. The reported density of fenestrae counted along the capillary circumference of mice mesenteric microvessels is 3.4 fenestrae/micron [175]. This value is quite close to that of 3 fenestrae/micron of capillary circumference observed in renal glomerular capillaries [169]. Density of fenestrae per square micron of endothelial surface is 45–60/μm² in renal peritubular capillaries [176] and 20/μm² in renal glomerular capillaries [169], whereas their frequency in mouse mesenteric capillaries is approximately 12/μm² [171].

figure 5_22_978-0-387-78940-8 — **Fig. 5.22**

figure 5_24_978-0-387-78940-8 — **Fig. 5.24**

figure 5_25_978-0-387-78940-8 — **Fig. 5.25**

The density distribution of submesothelial microvessels along the different portions of the peritoneum is variable. In the rabbit the mesentery appears to be the most vascularized peritoneal segment (contributing 71.1% of the total number of observed capillaries). The reported diaphragmatic and parietal contributions to the total microvascular bed examined were 17.9 and 10.9%, respectively [177].

In rabbit mesentery the main population of continuous blood microvessels is represented by:

1.
True capillaries (without perithelial cells), the mean luminal diameter of which is 7.2 μm and whose mean wall thickness is 0.4 μm (Figs. 5.11, 5.22, 5.23, and 5.24).
2.
Venous capillaries usually formed by the confluence of two or three capillaries. These show a thin endothelial layer, occasional peripheral perithelial cells, and have a mean luminal diameter of 9.2 μm.
3.
Postcapillary venules whose luminal diameter ranges between 9.4 and 20.6 μm [43].

figure 5_23_978-0-387-78940-8 — **Fig. 5.23**

With increasing luminal diameter there is a proportional increase in wall thickness due to the presence of more perithelial cells encircling the endothelial layer [178] (Fig. 5.23). The average ratio of luminal diameter to wall thickness is approximately 10/1 [178]. All aforementioned exchange vessels present at their luminal aspect a limiting area that separates the endothelial cell from the circulating blood and is formed by the plasmalemma with its trilaminar structure [19] (Fig. 5.24) and the glycocalyx (Fig. 5.24). The latter, originally described by Luft [38] in other vascular beds, has also been observed at the luminal aspect of peritoneal microvessels [39, 179] The presence of sialoconjugates, proteoglycans, and acidic glycoproteins organized as a fibrous network provides the plasmalemmal glycocalyx with electronegative charge [180] (Fig. 5.26).

figure 5_26_978-0-387-78940-8 — **Fig. 5.26**

There is evidence that anionic plasma proteins (albumin and IgG) are adsorbed to the glycocalyx of microvascular endothelial cells [181]. The fiber-matrix model of capillary permeability envisages the glycocalyx as a meshwork of glycoprotein fibers that, after adsorbing circulating proteins, would tighten its mesh, thereby rendering the underlying endothelium less accessible to water and other water-soluble molecules [40]. Furthermore, it has been shown that the adsorption of circulating anionic plasma proteins to the glycocalyx renders the underlying endothelium relatively impermeable to large, electron-dense, anionic tracers such as native ferritin (MW ∼ 450 kDa) [181].

The mean endothelial cell width of rabbit mesenteric capillaries is 0.4 μm, unless the cytoplasm bulges up to more than 1 μm at the site of the nucleus (compare Figs. 5.3 and 5.22 with Fig. 5.24). The cytoplasm includes the usual cell organelles: mitochondria, rough endoplasmic reticulum and free ribosomes [14, 169].

The mitochondrial content of vascular endothelial cells in frog mesentery decreases gradually from arterioles towards venous capillaries and subsequently increases toward venules [182].

The Golgi complex displays variable degrees of development in biopsies taken from different patients. This same variability was observed when comparing different peritoneal microvascular endothelial cells present in a single sample.

The cytoplasmic matrix of endothelial cells shows long filaments, parallel to the longitudinal cellular axis. Their diameter ranges between 20 and 100 Å [169], and at times they appear in bundles. These intermediate-size filaments seem to be a common component of the cytoplasmic matrix of vascular endothelial cells showing, however, a lower density distribution than that observed in other cell types [135].

Nuclei are generally oval, elongated (Fig. 5.22), or occasionally kidney-shaped with focal surface irregularities (Fig. 5.11). Their mean short-axis width in rabbit mesentery is 0.957 ± 0.417 μm [14].

Plasmalemmal vesicles, which can be found in most cell types, are particularly common in capillary endothelia [183], where they occupy approximately 7% of the cell volume [101] (Fig. 5.23). Their outer diameter is approximately 700 Å (it ranges between 500 and 900 Å) [14, 36, 50] and they have a round or oval shape surrounded by a three-layered membrane of 80 Å thickness (Fig. 5.24, inset).

According to their location in the cytoplasmic matrix, vesicles can be classified into three groups: a) vesicles attached to the plasmalemma limiting the blood front of the endothelial cell; b) free vesicles within the cytoplasmic matrix; and c) attached vesicles, but this time to the tissue front of the endothelial cell plasmalemma [50] (Fig. 5.23). The density population of plasmalemmal vesicles varies considerably from one vascular segment to another, even within the same microvascular territory [49, 50]. In the mouse diaphragm, arterioles show 200 vesicles/μm², true capillaries 900 μm², venular segments of capillaries 1,200 μm², and postcapillary venules 600 μm² [49].

Most vesicles that open to the extracellular medium have necks whose diameter can be as small as 100 Å [50]. Transendothelial channels formed by a chain of vesicles opening simultaneously on both fronts of the endothelium have been described in capillaries of mouse diaphragmatic muscle [49] as well as in postcapillary venules of rabbit and rat mesentery (Figs. 5.23 and 5.27) [14]. The relative frequency of transendothelial channels has been found to be higher in true capillaries than in arterioles and venules, with the highest density in the venular segment of capillaries [103]. Microvessels of frog mesentery showed a density distribution of three transendothelial channels for every 400 vascular profiles examined [182]. Just as in observations made on mesothelial cells, plasmalemmal vesicles and transendothelial channels do not bind cationic electron-dense tracers that, on the other hand, decorate the luminal aspect of coated pits and coated vesicles [98, 109] in the peritoneal microvasculature [37].

figure 5_27_978-0-387-78940-8 — **Fig. 5.27**

The functional significance of plasmalemmal vesicles or caveolae, vesicles, transendothelial channels, coated pits, and coated vesicles has been discussed in the section on normal mesothelium.

As stated above, in fenestrated capillaries, endothelial cells are pierced by fenestrae closed by a diaphram [184] (Fig. 5.22, inset; Figs. 5.25 and 5.28). Fenestrae are not static structures. It has been shown that their prevalence can be increased under the effect of vitamin A metabolites [185], the influence of sexual hormones [168], thrombocytopenia [186], and by the acute inflammatory reaction [169]. In this sense a microvascular bed (capillaries and postcapillary venules), supplied with a continuous endothelium, can rapidly develop endothelial fenestrations under the influence of vascular endothelial growth factor (VEGF), a 34–42 kDa cytokine, released by different cell types (eosinophils, neutrophils, and others) during the acute inflammatory reaction [187–190]. This effect has been demonstrated in vivo [191, 192], after acute and chronic exposure of different microvascular beds.

figure 5_28_978-0-387-78940-8 — **Fig. 5.28**

High concentrations of negative fixed charges (heparin and heparan sulfate) have been found on the blood front of fenestral diaphragms in several microvascular beds [98, 109, 184, 193–196]. They are expected to discriminate against anionic macromolecules, essentially anionic plasma protein. Similarly, mesenteric fenestrated capillaries of mice perfused with the cationic tracer ferritin showed densely packed anionic fixed charges on the endothelial cell glycocalyx, on the luminal aspect of fenestral diaphragms, as well as along both sides of the subendothelial basement membrane [175] (Fig. 5.25).

What is the role of fenestrae and intercellular junctions in the still ill-defined mechanisms related to capillary permeability? For more than 25 years the fenestral pathway was ascribed a major role in the permeability capabilities of fenestrated capillaries [197]. Some investigators suggested that, while open fenestrae could represent the ultrastructural equivalent of the large pore [77], fenestrae, closed by diaphragms (Fig. 5.25), could also provide a diffusive pathway for water- and lipid-soluble substances [198]. However, fenestral openings of 60–90 nm diameter are too large to be considered the structural equivalent of the hypothetical large pores, the radii of which range between 11 and 35 nm [77, 199]. Furthermore, the density of these pores, estimated at one every 20 μm² [50], is substantially lower than the density of fenestrae per square micron observed in microvascular beds.

At least from a theoretical point of view, the presence of anionic fixed charges at the level of fenestral diaphragms, as well as in the subendothelial basement membrane (Fig. 5.25), is a strong argument against the transfenestral passage of macromolecular anionic proteins [139, 200]. Indeed, previously reported physiological studies have demonstrated the selectivity and restriction of the fenestrated microvascular wall to the passage of electronegatively charged macromolecules [201–203]. Moreover, the permeability of fenestrated capillaries to anionic macromolecules is not higher than that of capillaries of the continuous type [204]. In this context Fig. 5.28 offers a descriptive account of the problem, observed in our laboratory (Gotloib L, unpublished observations). Rat mesentery was perfused (in vivo) through the arterial tree with negatively charged albumin–gold complexes for a period of 30 min. As can be seen, substantial amounts of albumin–gold particles appear contacting the luminal aspect of the endothelial cell plasmalemma, as well as free into the capillary lumen. Particles of the tracer can be observed in close apposition to the luminal front of fenestral diaphragms. However, albumin–gold particles were not seen in the subendothelial space, even 30 min after perfusion. These observations support the hypothesis that fenestrae are not permeable to anionic plasma proteins. Consequently, it appears that fenestral openings are unrelated to the theoretically predicted large pore system [197, 200, 204, 205]. On the other hand, fenestrated endothelia have higher hydraulic conductivity, and are more permeable to small ions and molecules than continuous endothelia [206].

Capillary endothelial cells are linked to each other by tight junctions (zonula occludens), originally described by Farquhar and Palade [207–209] (Fig. 5.29). Communicating or gap junctions have been observed in arteriolar endothelium [208]. Postcapillary venules have loosely organized junctions with discontinuous ridges and grooves of which 25–30% appear to be open with a gap of 20–60 Å [50]. The presence of gap junctions has also been documented [14].

figure 5_29_978-0-387-78940-8 — **Fig. 5.29**

Cytoplasmic plasmalemma bordering both sides of junctions also shows anionic fixed charges [37]. Their functional significance in relation to the passage of charged molecules will be discussed later.

Research performed during the past 10–15 years revealed that interendothelial tight junctions appear as a set of long, parallel, linear fibrils that circumscribe the cell, with short fibrillar fragments interconnecting the main parallel array. The number of fibers correlates with junctional permeability: the more densely packed the fibrillar mesh, the lower the junctional permeability [210]. Therefore, the tight junction is not a simple fusion between the outer plasmalemmal leaflets of neighboring cells [211]; rather, it consists of protein molecules such as occludins and cadherins in tight junctions, desmoleins and desmocolins in desmosomes, and connexins in gap juctions [212, 213].

Occludin is an integral membrane protein, exclusively localized at tight junctions in both epithelial and endothelial cells [212], and is directly involved in sealing the cleft, creating the primary barrier to the diffusion of solutes through the paracellular pathway as well as regulating, according to the modulation of occludin expression, the permeability properties of different microvascular beds [214]. Occludin is bound on the endothelial cytoplasmic surface to ZO-1, a 220 kDa membrane-associated protein likely to have both structural and signaling roles [215, 216].

Vascular endothelial cadherin, in turn, is an endothelial-specific cadherin that regulates cell to cell junction organization in this cell type, and provides strength and cohesion to the junction [217]. Cadherins are also implicated in junctional permeability, basically under the effect of inflammatory mediators such as tumor necrosis factor and histamine, which have been shown to induce a redistribution of these adhesion molecules to nonjunctional regions and junctional disassembly [218, 219].

The role of tight junctions in the permeability capabilities of the microvasculature, during the situation of normal physiology, has been a topic for intensive research and controversy through the years. Whereas some groups considered the intercellular cleft as the main pathway for water, as well as for small and large solutes and electrolytes [220, 221], other groups developed the concept that tight junctions create a regulated paracellular barrier to the movement of water, solutes, and immune cells between the microvascular compartment and the interstitial space, enabling the endothelial monolayer to create compositionally different fluid compartments [210, 222–224]. Recently published information indicates that the presence of tight junctions does not imply a foolproof seal of the intercellular cleft. Instead, this structure contains discrete ion-selective pathways through the extracellular portion of the junction, regulated, at least in part, by the activity of the cytoskeleton [210, 225]. As stated above, the transmembrane protein occludin is an excellent candidate for the sealing protein. Understanding the mechanisms involved in junction permeability will require both a more detailed molecular characterization of tight junction proteins and the regulation by the endothelial cells of their attachment to the perijunctional cytoskeleton [222]. As stated by Renkin [204] in 1977, identification of the tight junction with the diffusional pathway for macromolecular plasma proteins, in a situation of normal physiology, still remains questionable. Their role in capillary permeability is still debated.

As for blood cells, recent investigations showed that neutrophils preferentially migrate by crossing at tricellular corners, rather than passing through tight junctions that lie between two adjacent endothelial cells [226].

The basement membrane of true capillaries is normally a thin sheet at the interface between the abluminal aspect of the endothelial cell and the connective tissue (Figs. 5.13 and 5.22). In postcapillary venules it is interposed between the endothelial and the periendothelial cell (Fig. 5.23). Generally uniform for a given structure, its thickness varies among the different parts of the body. True capillaries of normal rabbit mesentery have a mean basal membrane thickness of 0.234 ± 0.095 μm [14]. As described for the submesothelial basement membrane, that of human capillaries also exhibits a significantly increasing thickness in the direction of head to foot [147]. It has been suggested that these regional variations are secondary to differences in venous hydrostatic pressure effective on the capillary bed [147]. Diabetic and nondiabetic patients on long-term peritoneal dialysis, showing reduplicated submesothelial basement membrane, had similar alterations on the capillary basement membrane of parietal peritoneum [144]. These changes were also observed in postcapillary venules and small arterioles of parietal peritoneum taken from diabetic uremics on CAPD (Figs. 5.30 and 5.31), as well as in skin capillaries (Fig. 5.32). Additionally, reduplication of mesenteric subendothelial capillary basement membrane has been recently reported in streptozotocin-induced diabetic rats, as early as after 4 months of uncontrolled hyperglycemia [144], whereas thickening was seen in the same animals 6 months after induction of the disease. These structural alterations of diabetic basement membranes seem to be derived from a substantial increased presence of collagen IV [149, 227–231] which, according to in vitro studies, appears to derive from extended exposure of cells to high concentrations of glucose [232, 233].

figure 5_30_978-0-387-78940-8 — **Fig. 5.30**

figure 5_31_978-0-387-78940-8 — **Fig. 5.31**

figure 5_32_978-0-387-78940-8 — **Fig. 5.32**

In rats, both thickening and layering of microvascular basement membrane can also develop as a consequence of aging [151, 234], but not before completing the first year of life [234].

So far, it may be speculated that reduplication or layering of submesothelial and peritoneal microvascular basement membranes in nondiabetics on CAPD could result from their continuous and long exposure to high glucose concentrations.

Subendothelial basement membranes of both continuous and fenestrated capillaries (Figs. 5.24 and 5.25), have regularly distributed anionic fixed charges along both aspects of the membrane [39, 175]. Their density distribution in continuous capillaries ranges between 31 and 34 μm of basement membrane [39, 143]. These values, shown in the section devoted to the mesothelial basement membrane, are not far from those detected in other microvascular basement membranes.

The chemical composition of the fixed electronegative charges linked to the subendothelial basement membrane has been explored in several microvascular beds. Studies have shown that their main structural components are glycosaminoglycans such as heparan sulfate and chondroitin sulfate [109, 235–237]. This is at variance with the biochemical and histochemical observations made on the glycocalyx cell surface charges, the main component of which is sialic acid and sialo conjugate. This pattern has been detected in microvascular endothelium [238–242], pleural, pericardial, and peritoneal mesothelial cells [243], as well as in macrophages [244], erythrocytes [245], and platelets [246].

What is the functional significance of these electronegative charges? A strong body of literature supports the concept that the permselectivity of capillary walls to anionic macromolecules is basically dependent on molecular charge, besides size and shape [139, 141, 167, 205, 236, 247–253]. Investigations performed in in vivo, whole organ studies [155], in isolated perfused frog capillaries [254], and in isolated rat hindquarters [248] have demonstrated their presence evaluating permeability of different endogenous proteins in a variety of microvascular beds. Indeed, similar results were observed in patients on CAPD comparing dialysate to plasma concentrations of amino acids, having almost the same molecular weight but quite different charge [255], as well as in rat peritoneum, using charged dextrans [256]. The fact that endogenous proteins of graded size are heterogeneous with respect to their molecular charge [257] lead to some conflicting results [258]. The key to this problem was found investigating clinical and experimental situations, where the permselectivity was substantially reduced or neutralized, enabling the observer to evaluate changes in permeability, derived from the absence of the normally present fixed electronegative charges. In the clinical set-up, type I diabetes [259], congenital [260, 261], and acquired nephrotic syndrome [262, 263] have been shown to expose the association of depleted glomerular negative charges and loss of the permselectivity of the capillary wall, leading to massive proteinuria. Experimental interventions performed in laboratory animals confirmed, in turn, the aforementioned findings. Enzymatic removal of sulfated (heparan sulfate) or nonsulfated (hyaluronic acid) glycosaminoglycans from the glomerular basement membrane resulted in a substantially increased permeability to bovine serum albumin [264]. Rats with streptozotocin-induced diabetic nephropathy showed reduced glycosaminoglycan contents in the glomerular basement membrane [265], decreased presence of their heparan sulfate-associated anionic sites [266], as well as significantly increased proteinuria [143]. Further observations made in the streptozotocin diabetic rat have shown a substantial reduction in the submesothelial and capillary subendothelial density distribution of anionic fixed charges (from 31 ± 2 to 12 ± 2 ruthenium red-decorated anionic sites/μm of basement membrane) and, at the same time, a significant increase of albumin losses in the peritoneal dialysis effluent, indicating a marked decrease of the permselective capabilities of the charged components of the peritoneal membrane (Fig. 5.16) [143]. Similar observations were made in intact rats after neutralization of the peritoneal negative charges with protamine sulfate [267].

The acute inflammatory reaction is the most spectacular experimental set-up to demonstrate the permeability changes derived from an acute reduction of the microvascular negative charge. This situation has been classically defined by the development of acute low hydrostatic pressure, high capillary permeability, and albumin-rich interstitial edema [163, 268]. In this sense the generalized acute inflammatory reaction derived from abdominal sepsis promotes a major erosion of the density distribution of the anionic fixed charges in several microvascular beds, diaphragmatic and mesenteric peritoneum (showing values as low as six anionic sites/micron of basement membrane) [269], myocardium [270], skeletal muscle, pancreas, renal peritubular capillaries [271], as well as in the submesothelial basement membrane of rat diaphragmatic and mesenteric peritoneum [142] (Figs. 5.19 and 5.26). Additional studies in the same experimental model of abdominal sepsis in rats demonstrated abnormally increased albumin content in mesenteric, diaphragmatic, and pancreatic interstitial fluid [272]. This drastic loss of the permselectivity of the capillary wall derives from a massive liberation and reduced inactivation of a host of mediators of inflammation triggered by acute inflammation [273–275], including tumor necrosis factor alpha, interleukins, platelet-activating factor, leukotrienes, thromboxane A2, activators of the complement cascade, kinins, transforming growth factor B, vascular endothelial growth factor, as well as many others already known, or still waiting to be identified [185, 276–279].

The role of intercellular junctions in macromolecular leakage during acute inflammation is still controversial. Some groups pointed to the endothelial tight junction as the main pathway for extravasation of macromolecular plasma protein. It was postulated that inflammatory mediators such as histamine, serotonin, bradykinin and leukotriene E4 induced junctional openings (Fig. 5.23, inset), by means of endothelial cells contraction [280–283] or by a loss of occludin and cadherin from the junctional complex [284, 285]. Unpublished observations from our laboratory (Gotloib L.) made in intact rats, as well as in rats with E. coli peritonitis, by means of intra-arterial injection of albumin–gold complexes, showed that most particles of the tracer cross the endothelial barrier transcellularly, via plasmalemmal vesicles. In both experimental situations, intact and infected rats, the tracer was not seen beyond the junctional infundibulum (Figs. 5.33 and 5.34); just the opposite: the tracer was present in plasmalemmal vesicles (Fig. 5.34, inset), and reached the subendothelial space in areas far from intercellular junctions (Figs. 5.33 and 5.34). This concept of transcellular transport of albumin through the capillary wall, also during acute inflammation, is supported by recently published evidence postulating a significant role for plasmalemmal vesicles and even for fenestrations induced by vascular endothelial growth factor [286–288].

figure 5_33_978-0-387-78940-8 — **Fig. 5.33**

figure 5_34_978-0-387-78940-8 — **Fig. 5.34**

Water transport across endothelium of continuous capillaries was classically thought to occur almost completely via the paracellular pathway through intercellular junctions. Transcellular transport was considered to be nil.

However, the relevance of the transcellular pathway also for water has recently been brought to the forefront by the immunohistochemical identification of aquaporin-1 channels in peritoneal microvascular endothelium [289, 290], as well as in rat peritoneal mesothelium [107]. Expression of this transmembrane water channel protein in the endothelial cell surface of continuous capillaries appeared to be basically localized in plasmalemmal vesicles, and its concentration is quantitatively comparable to that seen in erythrocyte plasma membrane [291]. The presence of aquaporin-1 in microvascular endothelium provides a molecular explanation for the water permeability of some capillary beds [292], as well as a low-energy cost pathway [293] for almost 70% of the transmembrane transport of water [289]. Furthermore, this evidence confirms the predicted concept that, also during peritoneal dialysis, not less than 50% of the transperitoneal water flow occurs through postulated ultra-small transcellular pores [97] that appear to be metabolically driven pathways rather than just holes located in the microvascular wall.

Summarizing the information obtained from ultrastructural and physiological studies, it can be stated that the microvascular endothelial cell should be considered a highly active structure, serving not only as a permeability barrier and an effective thromboresistant surface, but also as the location of important synthetic and other metabolic activities [182, 209].

Continuous capillaries are more permeable to larger molecules than are fenestrated capillaries [199]. Coated pits and coated vesicles are involved in receptor-mediated endocytosis, whereas the uncharged pinocytotic vesicles and transcellular channels are involved in the transfer of proteins and fluid-phase pinocytosis. The transcellular pathway plays a relevant role in the transmembrane transport of water and macromolecular plasma proteins. Additionally, all the resistances described by Predescu et al. [75] along the pathway leading from the microvascular lumen to the abdominal cavity, are negatively charged [37, 39].

Lymphatics

The lymphatic system serves to drain, from the interstitial compartment, a range of materials such as water, proteins, colloid materials and cells [294], all elements included in the interstitial fluid. Under normal conditions fluid crosses the microvascular endothelial membrane at a rate whose magnitude depends on the Starling forces acting at each aspect of the capillary membrane, as well as on the permeability properties of the endothelial microvascular monolayer. The local autoregulation of interstitial volume is provided by automatic adjustment of the transcapillary Starling forces and lymphatic drainage [161]. Therefore, an alteration in the aforementioned forces results in interstitial accumulation of fluid that will eventually be removed by the lymphatic flow that, in situations of high capillary permeability edema occurring during acute inflammation, can increase by a factor of ten [295]. In the abdominal cavity, lymphatics have a relevant role in the prevention of ascites [296].

Work during the last 20 years has revealed relevant evidence characterizing lymphatic structure and organization. The first stage of lymph collection occurs through a system of interstitial nonendothelial channels, or low-resistance pathways known as pre-initial lymphatics [297, 298], which have been seen in the cat and the rabbit mesentery [299].

This most peripheral part of the lymph vessel system is a completely open net of tissue channels which drain, at least in the cat mesentery, mainly along the paravascular area of the venous microvasculature, into a network of 0.5 mm long, irregularly shaped endothelial tubes, approximately 20–30 μm width [300]. By the time these tubes are completely filled they can reach a maximal diameter of up to 75 μm [301]. A single endothelial layer (Fig. 5.35, inset) forms these endothelial tubes defined as initial lymphatics or lymphatic capillaries [299, 301, 302].

figure 5_35_978-0-387-78940-8 — **Fig. 5.35**

The subendothelial area is most of the time devoid from basement membrane, as well as from a smooth muscle layer present in larger lymphatic collectors [303]. The sporadically observed patches of basement membrane have, as in blood capillaries, anionic fixed charges that can be decorated by cationic tracers [304]. Due to the absence of muscular layer, lymphatic capillaries lack the capability for spontaneous contractility [305]. However, the fact that lymphatic endothelial cells contain an abundant supply of fine actin-like filaments, 40–60 Å in diameter, arranged in bundles parallel to the long axis of the cell, led some investigators to postulate that these filaments could function as a contractile element of the lymphatic capillary wall [306, 307].

Anchoring filaments, having histochemical and ultrastructural characteristics similar to those observed in elastin-associated microfibrils, form a uniform population of fibrous elements, leading to the development of structural and functional continuity between the abluminal aspect of the lymphatic capillary endothelial cell and the elastic network of the adjacent connective tissue [308]. The main role of these anchoring filaments is the prevention of capillary collapse, when the interstitial pressure gains strength as a consequence of expanded fluid content of the interstitial compartment [309]. This simple element enables the lymphatic system to launch a mechanism of fluid drainage that accounts for 25% of the safety factors that can prevent formation of interstitial edema [308]. In this context it has been proposed that initial lymphatics directly sense and regulate the interstitial fluid volume [310].

The total surface area of pre-initial and initial lymphatics seems to be smaller than the total exchange area of blood microvessels [198]. Other studies on cat and rabbit mesentery showed the additional presence of flat, blind saccular structures up to 40 μm wide, with a wall made up of a simple layer of thin endothelial cells, devoid of basement membrane [198, 300].

Lymphatic endothelial cells are flat and elongated, showing an average thickness of 0.3 μm in non-nuclear areas [46, 306]. The luminal aspect of lymphatic endothelium, when exposed to cationic tracers such as cationized ferritin or ruthenium red, shows a high density of anionic fixed charges that, at times, can also be detected labeling the luminal aspect of the intercellular cleft (Fig. 5.36) [307, 44]. These charges prevent the adhesion of electronegatively charged blood cells to the endothelial luminal surface and may play a significant role in the movement of charged solutes from the interstitial compartment to the capillary lumen [44]. Furthermore, the absence of subendothelial and negatively charged basement membrane (Fig. 5.36) points at the asymmetry of the lymphatic capillary wall that is at variance with the electric symmetry characteristic of blood capillaries.

figure 5_36_978-0-387-78940-8 — **Fig. 5.36**

Nuclei of endothelial cells are flattened and, on electron microscopy, appear elongated. Their irregular outline shows a thin peripheral rim of dense chromatin. Plasmalemmal vesicles [46, 311] and transendothelial channels, similar to those described for blood microvessels, are commonly observed [307]. Plasmalemmal vesicles have been shown to participate in the transcellular movement of albumin–gold complexes, from the submesothelial interstitial space to the lumen of capillary lymphatics [79] (Fig. 5.37). Furthermore, the endocytotic pathway has been shown up by the presence of the same tracer into cytoplasmic endosomes (Fig. 5.38).

figure 5_37_978-0-387-78940-8 — **Fig. 5.37**

figure 5_38_978-0-387-78940-8 — **Fig. 5.38**

Several types of interendothelial junctions have been described. Approximately 2% of the whole junctional system consists of open junctions, showing gaps up to 100 nm width, that can, at times, be as wide as 1,000 nm [307, 312]. These openings serve as a way in for macromolecular solutes such as gold-labeled albumin (Fig. 5.38) or cationized ferritin (Fig. 5.39). At times two adjoining endothelial cells overlap each other, forming a kind of valvular junction that can be easily opened by an eventual increase of interstitial pressure (Fig. 5.36, inset). Junctional infundibuli show anionic fixed charges similar to those observed in the luminal endothelial glycocalyx (Fig. 5.36). Around 10% of junctions are zonula adherens, whereas the rest are tight junctions [301, 313]. It has been proposed that, in addition to the organized prelymphatic system [314], a small percentage of open junctions (1–6%) can account for a substantial proportion of the lymphatic pathway for fluid as well as small and large molecule drainage [301].

figure 5_39_978-0-387-78940-8 — **Fig. 5.39**

The diaphragmatic lymphatic capillary net is organized as a plexus along the submesothelial surface [315], which drains, through an intercommunicating microvascular system, into a plexus on the pleural side of the diaphragm [133]. The distribution of the whole diaphragmatic network is irregular, and varies in different species.

A prominent feature of diaphragmatic lymphatics is the presence of flattened, elongated cisternae or lacunae, approximately 0.3–0.6 cm length, with a long axis that is parallel to the long axis of the muscle fibres [315–317] (Fig. 5.40). The monolayer endothelial lining of the lymphatic lacunae is thin and shows no tight junctions. Adjacent cells usually overlap, forming valve-like processes, leaving an open interface that can be as wide as 12 μm. The cytoplasm of endothelial cells, basement membrane, and anchoring filaments of lymphatic lacunae are similar to those structures described for lymphatic capillaries. While anionic sites have not been observed, the glycocalyx of cisternal endothelium, when exposed to ferritin, is heavily decorated by the cationized tracer, which also appears along the open intercellular clefts [44, 318].

figure 5_40_978-0-387-78940-8 — **Fig. 5.40**

Diaphragmatic lymphatic lacunae, regularly connected by transverse anastomosis [114], and capillaries from the whole peritoneal lymphatic network, including the rich omental plexus [319], drain into a system of precollector, small-caliber lymph vessels that have a poorly developed smooth muscle layer underlying the endothelium. These vessels, which have semilunar valves [1, 320], drain, in turn, into the larger collecting vessels, whose diameter ranges between 40 and 200 μm [321]. The luminal aspect of the endothelial layer shows a sequence of valvular segments, with a semilunar bicuspidal valve at the distal end of each [133, 321]. The smooth muscle layer underlying the subendothelial basement membrane shows a spiral arrangement around the endothelial tube that becomes more pronounced towards the downstream end of the intervalvular segment [305, 322, 323]. Distances between adjacent valves range between 0.1 and 0.6 mm [323]. Thereby, the anatomical and functional unit (lymphangion) is established, consisting of one valve and the following intervalvular segment, which measures 2–3 mm in length [324]. This collecting segment, limited by two one-way valves and an intrinsic smooth muscle layer, compresses the lymphatic lumen driving the intravascular fluid centrally into the next compartment, making up an escalated system of drainage that has, in the proximal part of each lymphangion, a valve that prevents retrograde flow [305]. The presence of valves also enables this part of the system to reach differential intraluminal pressures around 1–2 cm of water [325].

Capillary lymph that flows from the interstitium slowly moves downstream (average velocity for particles with diameter up to 5 μm = 1 μm/min) [303], drains into large collecting channels (40–200 μm diameter), and proceeds in the direction of the central venous circulation, propelled by peristaltic and rhythmic contractions of consecutive lymphangions [292, 303, 321, 326], with frequencies ranging between 4 and 12 contractions/minute [321, 326]. Within each lymphangion, hydrostatic pressure increases to a threshold of approximately 12 cm of water, after which the proximal valve is closed, and the downstream valve is opened. The cycle is repeated in the following segment. Contractility of lymphangions is modulated by a pacemaker site of spontaneous activity, apparently located, at least in bovine mesenteric lymphatics, in the vessel wall near the inlet valve of the unit. Activity propagates at a speed of 4 mm/s, and the ejection fraction was evaluated at 45–65% [327].

Contractions are generated by myogenic stimuli (hydrostatic pressure of 5–7 cm water) [327], and influenced by activation of α- and β-adrenoreceptors [328–330], histamine, leukotriene C4 and D4, platelet-activating factor [331, 332], PGF2 alpha, PGA2, PGB2 [333], bradykinin [334], and vasoactive intestinal peptide [335]. It should be noted that all the aforementioned vasoactive substances are mediators of inflammation present in high concentrations in blood and tissues during the localized or the generalized acute defense reaction [276].

Lymph flows from collectors to the thoracic duct and the right lymph duct, and finally drains into the subclavian veins. Lymphangions join larger collecting lymphatic vessels, forming a dichotomous tree that drains entire tissue regions. This arrangement has been described in the diaphragm as well as in mesentery [305, 325, 336].

Innervation of lymph vessels has been studied in the dog and cat mesentery, by means of silver stains [320]. It was shown that large lymphatic collectors have myelinated nerves that remain on the adventitial area, and nonmyelinated nerve fibers that penetrate into the region of valve attachment and are considered to be the motor supply to the smooth muscle. Bovine mesenteric lymphatics show adrenergic nerve fibers in the media, as well as in the adventitia. Human mesenteric lymph collector neurotransmitters are both adrenergic and cholinergic, the former being prevalent. Lymphatic capillaries are devoid of innervation [337].

Since Starling [338, 339], it has been accepted that, besides the removal of excess interstitial tissue, the lymphatic system has the special function of absorbing protein. Normally, blood capillaries leak protein, which will not re-enter the blood vessels unless delivered by the lymphatic system [340]. It is generally accepted that the rate of lymph formation is equal to the net capillary efflux under normal physiological conditions, in order for the interstitial fluid volume to remain constant [310]. However, the mechanisms involved in the formation of lymph, at the level of the most peripheral part of the lymphatic system, are still controversial. According to Allen and Vogt [341], who formulated the hydraulic theory, lymph formation is the end-result of hydraulic forces acting across initial lymphatics. Assuming that the interstitial hydrostatic pressure is zero, or even negative [159, 342], any rise will also increase the initial lymphatic flow, and edema will eventually develop if and when the lymphatic drainage capabilities are exceeded [343–345]. This concept of increased hydrostatic pressure as the main factor in the process of lymph formation was extrapolated to the lymphatic absorption from the peritoneal cavity [346, 347]. In this context it was postulated that, during peritoneal dialysis, the intra-abdominal pressures [348, 349] modulated lymphatic drainage from the abdominal cavity well within the range of values observed in dialyzed patients [166, 350]. This concept has been substantially challenged by a series of elegant studies performed by Flessner et al. [166, 351], who showed that a significant proportion of the intra-abdominal fluid is lost to the abdominal wall, the rate of which is also dependent on the intra-abdominal pressure. This fluid, after being incorporated to the tissues surrounding the abdominal cavity, will be drained through the lymphatic circulation [352]. The eventual influence of the intrathoracic negative pressure upon the lymphatic downstream circulation [353] may well be an additional component of the hydrostatic forces involved in lymph progression to the venous-blood compartment.

The osmotic theory of lymph formation [354] postulates the existence of a protein-concentrating mechanism at the level of the initial lymphatics, the main result of which would be that only 10–40% of the fluid initially entering within the lymphatic network would flow downstream, back to the blood compartment, and the remaining fluid would be filtered out from the lymphatics as a protein-free solution. This process would eventually cause a high protein concentration, and an oncotic gradient between the contents of the initial lymphatics and the surrounding interstitial fluid. Other investigators have proposed a vesicular theory of lymph formation, which holds that plasmalemmal vesicles provide the major route for transendothelial transport of protein, thereby creating the oncotic gradient needed for further fluid flow between adjacent cells or through transendothelial channels [355, 356]. This hypothesis is supported by recent studies that have shown active transendothelial transport of albumin in plasmalemmal vesicles [60, 357]. Figure 5.37 shows gold-labeled albumin transported by plasmalemmal vesicles of a mouse diaphragmatic lymphatic capillary.

On the other hand, it has been suggested that the postulated mechanisms may not necessarily be exclusive in the sense that some or all could function simultaneously. However, the relative influence of each one could vary in different areas of the initial lymphatic network [358].

The relevance of peritoneal lymphatics, as well as their impact upon ultrafiltration during peritoneal dialysis, is addressed in Chapter 6. Omental milky spots have, relatively recently, attracted the attention of investigators working in the field of peritoneal dialysis. First described by Von Recklinghausen in pleura of rabbits [359], and later on in humans [360], these structures are part of the peritoneal lymphatic system [361]. Been described as submesothelial lymphoid structures essential for the maturation of resident peritoneal macrophages, they are actively involved in peritoneal defense reactions under a diversity of inflammatory conditions [362]. Macroscopically, milky spots appear as small (up to 1 mm diameter), white bodies, most commonly detected in perivascular areas of the greater omentum. Their structure, as seen under light microscopy, has been thoroughly reviewed by Di Paolo et al. [362, 363] in rats, rabbits as well as in patients on long-term peritoneal dialysis. In all the abovementioned species, they are located in the submesothelial tissue, showing blood capillaries surrounded by lymphocytes and macrophages and, at times, even lymphatic microvessels that can be identified within the frame of the same structure. The cell population of milky spots is made up by 400–600 cells, including macrophages (45–70%), lymphocytes (14–29%), and a low number of plasma cells (around 6%). Occasionally, megakariocytes and adipocytes can also be detected. Work done in experimental animals showed that number and size of milky spots, as well as their cell population, substantially increase after introduction of a peritoneal catheter, at the time of infection and inflammation, or after repeated exposure of the peritoneum to dialysis solutions containing 1.5 or 4.25% glucose [363]. Observations made in patients depicted changes not far from those seen in experimental animals, since both, size and cellularity of milky spots increased with the time-span of maintenance term peritoneal dialysis [363].

Peritoneal Innervation

The first report announcing the presence of nerves in the peritoneal interstitium was made by Haller in 1751 [364] and confirmed during the 19th century by Ranvier and Robin who, using osmic acid and silver nitrate, described nerve trunks, branches, and nerve endings accompanying arteries and veins. Robinson [1] described the peritoneum as being richly supplied with myelinated and nonmyelinated nerves (Fig. 5.41).

figure 5_41_978-0-387-78940-8 — **Fig. 5.41**

In rat mesentery, networks of adrenergic axons innervate the principal and small arteries and arterioles. Precapillary arterioles, collecting venules and small veins are not innervated, and are most likely under the influence of humoral vasoactive substances [365]. Lymphatic innervation was described in the previous section.

In 1741, Vater observed that the submesothelial connective tissue of cat mesentery contained oval corpuscles with a diameter of approximately 1–2 mm. In 1830, Paccini rediscovered and gave a systemic description of this corpuscle, known as the Vater–Paccini corpuscle [1]; it takes the form of a nonmyelinated nerve ending, which, in transverse section, appears as a sliced onion. In humans it has been observed in the peritoneum of mesentery and visceral ligaments, functioning as the main receptor for perception of pressure.

Cytology of the Peritoneal Fluid

The peritoneal fluid of laboratory animals has classically been a favored site for experiments dealing with the inflammatory response [366], as well as for those designed to analyse the biological reaction to infection [367].

More than 50 years ago, Josey and Webb realized that fluid shifts into and out of the peritoneal cavity could change the concentration of cells without affecting their absolute number [368–371]. The methodological answer to this question was given by Seeley and colleagues, who weighed peritoneal fluid and measured the cellular concentrations, and so were able to estimate the absolute number of cells [372]. Padawer and Gordon [370], after analyzing the cellular elements present in peritoneal fluid of eight different normal mammals, concluded that the most frequently observed cells were eosinophils, mast cells and mononuclears (including lymphocytic and macrophagic elements). Total cell numbers, as well as percentages of the different cells, varied greatly among the species examined. Neutrophils were never observed in normal animals. Total absolute counts were higher for females than for males, as well as for older animals compared with younger ones. In the individual animal, under normal conditions, the number of cells present within the abdominal cavity was constant [370].

Observation of peritoneal fluid obtained from healthy women showed that macrophages and mesothelial cells contributed more than 70% of the whole cell population, whereas lymphocytes and polymorphonuclears contributed to a lesser extent (18 and 7%, respectively) [373]. Other investigators observed that at the midphase of the menstrual cycle, macrophages, which comprised 82–98% of the peritoneal cells, showed morphological as well as biochemical heterogeneity and were seen to be involved in phagocytosis of erythrocytes [374] (Fig. 5.42). However, other studies showed up to four different types of cytological patterns in peritoneal fluid of women during the course of the menstrual cycle, in all of which mesothelial cells contributed substantially to the total cell counts. The paramenstrual type was in most cases hemorrhagic and highly cellular [375]. Ciliocytophtoria, anucleated remnants of ciliated mesothelial cells, can be occasionally observed in effluent dialysate, basically in young women. Inability to identify these structures can mislead the laboratory team as well as the physician to search for parasitic or fungal contamination [376].

figure 5_42_978-0-387-78940-8 — **Fig. 5.42**

The apparently puzzling effect of intraperitoneal saline inducing substantial influx of neutrophils into the abdominal cavity, which was observed long ago [367], was not confirmed when the experiments were carried out using sterile techniques. Bacterial lipopolysaccharides proved to be very effective in producing intraperitoneal exudate rich in cells [371]. This phenomenon was inhibited by prior intraperitoneal injection of cortisone [377].

In humans, sterile inflammatory effusions are characterized by a rich cellular content including neutrophils, lymphocytes, macrophages, mesothelial cells, eosinophils, and basophils, usually in that order of frequency [378]. The presence of macrophages, mesothelial cells, lymphocytes, eosinophils, and even plasma cells has been confirmed by electron microscope studies [379–381].

Peritoneal eosinophilia (eosinophils >10–50%) has been experimentally induced by intraperitoneal injection of iodine, chalk, nucleic acids, pilocarpine, hemoglobin or red blood cells, egg albumin, gold salts, mineral and vegetable oils, hydatidic fluid, and saline [378, 382]. On the other hand, intraperitoneal injection of bacteria and/or bacterial endotoxins induces a massive migration of neutrophils and monocytes into the peritoneal cavity [370, 383, 384].

The information presented above suggests that the cell content of effluent peritoneal dialysate is likely to be modified by so many factors that a concise description of a standardized cytological pattern becomes extremely difficult. There are, however, a few aspects of peritoneal effluent dialysate that have been defined: a) Patients on CAPD have total cell counts up to 50 cells/mL [383]. (b) The population of resident peritoneal cells observed in patients on long-term peritoneal dialysis is basically made up by macrophages (around 50% of the population), and lower prevalence of lymphocytes, mast cells, and mesothelial cells [385–388]. (c) During infection there is a substantial increase in total cell number [389], as well as in the proportion of neutrophils [385–388]. (d) Fluid eosinophilia is a basic component of the still ill-defined eosinophilic peritonitis [390–392].

Besides, the unphysiological situation of PD derives in substantial micro environmental changes that significantly affect the life cycle of the exposed and still attached monolayer. As a result, the population of cells recovered from dialysate effluent from a new patient is considerably different from that of patients on long-term PD. As shown in Fig. 5.43, cells detected in the first effluent of a new patient show remarkable mitotic activity (Figs 5.43a and 5.43b), a quite modest proportion of non viable cells (Fig. 5.43c) as well as a low prevalence of cells demonstrating positive expression to beta galactosidase, denoting the low prevalence of cells undergoing terminal replicative senescence (Fig. 5.43d). On the other hand, mesothelial cells isolated from effluent of patients on long-term PD (Fig. 5.44) show a quite different phenotype: they appear mostly as nonviable (Fig. 5.44a), the mitotic activity is nil (Fig. 5.44b), the vast majority are positively stained by beta galactosidase (Figs 5.44c and 5.45d), whereas a high proportion are undergoing apoptosis (Fig. 5.46). As it will be discussed later, this senescent phenotype of mesothelium was also detected in the monolayer of experimental animals exposed to high glucose concentration dialysis fluids. So, it may be hypothesized that careful and sequential observation of mesothelial cells recovered from patients' effluent, using the aforementioned staining technique, may well give quite representative information regarding the regenerative capabilities of the mesothelial monolayer still dressing the cavitary aspect of the peritoneal membrane of patients undergoing long-term PD.

figure 5_43_978-0-387-78940-8 — **Fig. 5.43**

figure 5_44_978-0-387-78940-8 — **Fig. 5.44**

figure 5_45_978-0-387-78940-8 — **Fig. 5.45**

figure 5_46_978-0-387-78940-8 — **Fig. 5.46**

Ultrastructure of Peritoneal Fluid Cells

Free-floating mesothelial cells are round or oval in shape and show a central, round nucleus (Fig. 5.47). Occasionally, binucleated mesothelial cells can be observed (Fig. 5.48). Nuclear chromatin is quite evenly distributed (Fig. 5.49, inset) and a small nucleolus may be observed. Numerous slender and sometimes branching microvilli emerge from the cytoplasmic membrane [379, 381, 393–395]. Branching microvilli, similar to those observed in human embryos [14], can be quite crowded in some cells, whereas in others they are scarce [381] (Fig. 5.49, inset). The glycocalyx covering the luminal aspect of the plasmalemma is endowed with electronegative fixed charges as shown in preparations exposed to the cationic tracer ruthenium red. Mitochondria, numerous cisternae of rough endoplasmic reticulum, and free ribosomes are mainly located in the outer part of the cytoplasm, and so are pinocytotic vesicles [379, 395]. The presence of intermediate-size filaments, perinuclear or irregularly scattered along the cytoplasm, has been documented in young free-floating mesothelial cells [378, 380], as well as in those recently implanted on the peritoneal surface. These free-floating mesothelial cells should be distinguished from desquamated, degenerating mesothelial cells wandering in the peritoneal fluid (Fig. 5.48) [396].

figure 5_47_978-0-387-78940-8 — **Fig. 5.47**

figure 5_48_978-0-387-78940-8 — **Fig. 5.48**

figure 5_49_978-0-387-78940-8 — **Fig. 5.49**

Macrophages, which can be observed in large numbers, usually show an irregular and, at times, kidney-shaped nucleus with distorted masses of chromatin concentrated along the nuclear membrane (Figs. 5.42 and 5.47). The cytoplasmic outline of macrophages is irregular, with thin processes of variable length which, at times, engulf degenerated cells (Fig. 5.42) or take the form of signet-ring macrophages (Fig. 5.5, inset). Mitochondria, a small Golgi complex and phagolysosomes are more evident when the cell is involved in phagocytic activity (Fig. 5.42).

The ultrastructural aspect of inflammatory cells that eventually appear in the peritoneal fluid is similar to that classically described for other tissues.

The Origin of the New Mesothelial Cells

It has been experimentally shown that small and large mesothelial wounds heal at the same rate within 7–10 days after injury [397]. The basal, normally observed mitotic rate of mesothelial cells, as measured in the rat by ³H-thymidine incorporation, ranges between 1 and 2% day (Fig. 5.50). This concept is supported by the fact that the steady state of the cell population is clearly defined by the following parameters: the proportion of cells passing through the G1 checkpoint, indicated by PCNA expression (proliferative cell nuclear antigen) that is also around 1–2%, as well as prevalence of non viable (Trypan Blue stained) senescent (positively stained with beta galactosidase at pH 6) (Fig. 5.45) and apoptotic mesothelial cells (Figs 5.46 and 5.51) that are within the same range [398, 399]. This rate of renewal is significantly increased during peritonitis, reaching maximal values of up to 19% between 1 and 3 days after injury, and returning to the basal activity on the 4th or 5th day [400]. It should be noted, however, that proliferations of fibroblasts, as well as mesothelial cell regeneration, are substantially inhibited in experimental uremic animals [400–402].

figure 5_50_978-0-387-78940-8 — **Fig. 5.50**

The origin of the new mesothelial cells repopulating denuded areas of injury is still controversial. Four different hypotheses have been proposed:

1.
The repopulating cells originate from the bone marrow [102]. Other experimental studies showed, however, that whole-body irradiation sufficient to depress peripheral white blood cell count as well as cell replacement by the bone marrow did not prevent mesothelial healing [403]. Therefore, the existence of a circulating mesothelial precursor originating from the bone marrow seems unlikely.
2.
Free-floating cells of the serosal cavity settle on the injured areas and gradually differentiate into new mesothelial cells [283, 404–406], (Fig. 5.49). Research done in order to settle this proposal, exposed evidence indicating that, after injury, free floating mesothelial cells, exfoliated from noninjured areas or from omental milky spots (12), settle on the injured areas, being instrumental to the healing of the monolayer [399, 407, 408]. Even though some investigators have not accepted this hypothesis [401, 406, 409], this approach finds support in other studies demonstrating the feasibility of mesothelial cells transplantation in humans, rabbits and rats as reported by several investigators [410, 411], as well as by our group (Fig. 5.52a).
3.
Other studies [393, 401, 412] suggested the sequence of a two-stage process; during the first 24 h, macrophages forming the first line of defense [413] and coming from the peritoneal fluid, repopulate the wound surface (Fig. 5.5, inset). Later, during the second stage, new mesothelial cells, arising from metaplasia of mesenchymal precursors located in the interstitial tissue well below the site of injury, migrate to the surface and differentiate into mature mesothelial cells (Fig. 5.53). This hypothesis has not been universally accepted [25, 283, 399, 404, 414]. It has also been suggested that the early implanted macrophages are gradually transformed into mesothelial cells [414]. However, Raftery [415], after labeling peritoneal macrophages with polystyrene spheres, presented strong evidence against the hypothesis that peritoneal macrophages could be transformed into mesothelial cells.

On the other hand, elongated, fish-like mesothelial cell precursors coming up from the submesothelial connective tissue were also observed under the damaged areas. The nuclear and cytoplasmic aspects of these cells were identical to that shown by new mesothelial cells already implanted on the peritoneal surface (Fig. 5.52b).
4.
Mature mesothelial cells from adjacent areas migrate and proliferate to repopulate a depopulated area [407, 416]. This approach is supported by in vitro studies [417, 418] showing early migration and increased bromodeoxyuridine (BrdU) incorporation 24 h after injury, the latter showing values ranging between 20 and 26% of the observed cells. It should be noticed that these figures, representing the proportion of cells in S phase, imply an underestimation of the actual number of cells undergoing mitosis. Indeed, the proportion of cells that passed the G1 checkpoint and supposed to reach S phase after some 10 h may well be similar, or even higher, than that observed during incorporation of BrdU. In addition, in vivo studies have been performed evaluating sequentially the dynamics of mesothelial repopulation, after a local mechanical exfoliation creating a doughnut-like area of undressed peritoneal surface [419]. This study demonstrated that repopulation also takes place by replication and centripetal migration of mature mesothelial cells located in in the monolayer bordering the injured area. (Figs 5.52c and 5.52d, Figs 5.53 and 5.54). Here again, the prevalence of cells undergoing mitosis is several times higher than that observed in intact, unexposed mesothelium.

figure 5_52_978-0-387-78940-8 — **Fig. 5.52**

figure 5_53_978-0-387-78940-8 — **Fig. 5.53**

figure 5_54_978-0-387-78940-8 — **Fig. 5.54**

All this evidence suggests that, most likely, mesothelial cell regeneration takes place through three different processes occurring simultaneously: implantation of young wandering mesothelial cells, migration of mesothelial cell precursors coming from the underlying connective tissue, and mitosis and migration of mature mesothelial cells bordering the injured area. The individual contribution of each mechanism cannot still be evaluated, as suggested by the presence of isolated new mesothelial cells repopulating denuded areas of the peritoneal surface (Fig. 5.55). These cells could eventually derive from by any of the already mentioned pathways of regeneration. According to studies done using the doughnut model of mesothelial regeneration, complete repopulation of the monolayer occurs after a recovery period of 15 days (Fig. 5.56).

figure 5_55_978-0-387-78940-8 — **Fig. 5.55**

figure 5_56_978-0-387-78940-8 — **Fig. 5.56**

The Price of a Failing Regeneration

The normal density population of mesothelial cells dressing the cavitary aspect of the peritoneum is around 300,000/cm² in mice and rats [31, 420].

This number remains basically unchanged in the intact, unexposed animal, showing a minimal and non significant variability between samples of the same animal, as well as between those obtained from different mice or rats. This situation of no numerical change results from a continuous replacement of dying cells by means of cell replication. This mechanism has also been detected in peritoneal biopsies taken from human patients on long-term peritoneal dialysis [421], suggesting that the therapeutic procedure induced a situation of continuous mesothelial injury, coupled to an also continuous process of regeneration. This working hypothesis found support in the original observation of Di Paolo et al. [26] regarding the absence of mesothelial microvilli in peritoneal biopsies taken from patients on CAPD, later on identified as a sign of impending apoptosis [422] (Fig. 5.51).

figure 5_51_978-0-387-78940-8 — **Fig. 5.51**

A tight regulation of the rates of cell growth and cell death is critical for maintaining a normally populated monolayer. In this sense, a decreased rate of mesothelial cells growth, an increased proportion of dying cells, or both could eventually lead to a depopulated monolayer that, in turn, would result in repair by means of connective tissue [399, 423, 424], the thickness of which can be as high as 100 μ.

So far, during the situation of steady state, new mesothelial cells continuously replace the dying ones [425]. This steady state is broken when: a) the magnitude of cell injury overwhelms the regenerating capabilities of the monolayer; b) the cell cycle of the mesothelial cells is blocked or departs from its normal course; and c) both developments occurring simultaneously. When the balance between regeneration and injury is broken, proliferative mechanisms are required to relieve the structural alterations represented by a depopulated monolayer. This process of repopulation and regeneration is dependent on the presence of a resilient cell population that has retained the potential for proliferation and differentiation. Failure of this mechanism leads to repair by means of connective tissue, which, in turn, becomes the first step toward peritoneal fibrosis and sclerosis [425] (Fig. 5.57). At this point it is pertinent to remind that submesothelial peritoneal sclerosis is an extremely frequent complication of long term peritoneal dialysis. It has been detected in about one half of dialyzed patients during the first year on peritoneal dialysis [426], whereas its prevalence reaches an 80% level after only 2 years on the aforementioned technique of renal replacement therapy [427, 428]. Actually, a variable degree of diffuse peritoneal fibrosis has been documented in all patients who have been on long-term peritoneal dialysis [429]. And, in turn, this development paves the way to membrane failure [430], a situation in which, at least from the point of view of its dialytic capabilities, the peritoneum is no more peritoneum.

figure 5_57_978-0-387-78940-8 — **Fig. 5.57**

Mesothelial cells are extremely vulnerable to minor injury. Mild drying or wetting of rat cecal peritoneum for 5 min induced mesothelial cell degeneration and detachment, and severe interstitial edema [393, 394]. This fragility of the monolayer is somehow compensated by the remarkable regenerative capabilities mentioned before. Evidence of this property is brought to light by the almost complete repopulation of the mesothelium 15 days after its massive exfoliation resulting from exposure to a 0.125 mg. Trypsin solution during a period of 10 min [431] (Fig. 5.58). This enzyme is commonly used in order to harvest mesothelial cells from human omentum or from experimental animals [432].

figure 5_58_978-0-387-78940-8 — **Fig. 5.58**

However, as stated in the title of this section, exfoliation always demands a price. Meticulous observation of peritoneal biopsies showed, in the same animals, small domains were depopulation was repaired by connective tissue, launching at the local level the mechanisms involved in peritoneal sclerosis (Fig. 5.58). Recently published investigations have shown that new fibroblastic cells can develop from native mesothelial cells by a mechanism of epithelial-to-mesenchymal transition, launched by injury resulting from the use of poorly biocompatible dialysis solutions [433]. So far, these findings coincide with previously reported observations [420, 425] postulating that the mesothelial cell plays a key role in the preservation of the peritoneum as an effective dialysis membrane, as well as in its structural break down and final functional failure. In addition, it has been postulated that submesothelial myofibroblasts, actively involved in the reaction to the persistent tissue injury derived from exposure to PD solutions, take part in the inflammatory response leading to extracellular matrix accumulation and angiogenesis. Probably, these cells may also arise from mesothelial cells through epithelial to mesenchymal transition [433].

Peritoneal sclerosis goes along with a marked increase in the density of microvessels of neoformation. This phenomenon of neoangiogenesis has been detected in peritoneal biopsies of patients in long term peritoneal dialysis [433–436], as well as in rats after experimental induction of the fibrous reaction [431]. Those studies showed that the thicker the peritoneal tissue, the higher the number of vessels/surface area unit.

Besides, analysis of the microvascular alterations seen in humans on long-term peritoneal dialysis led Williams et al. [436] to systemize and define four sequential degrees of pathology, that go from presence of subendothelial hyaline material with thickness lower than 7 μm (degree 1); same changes but with thickness over 7 μm (degree 2); additional luminal distortion or narrowing (degree 3); and luminal obliteration (degree 4). It is interesting to remark that in this same study [436], 87% of patients treated with PD for periods of 6 years, exhibited clear signs of microvasculopathy, and that in 66% of them, changes reached degree 4. Therefore, two thirds of microvessels appeared closed. From this information it may be deduced that, in the long range, development of neovascularization does not automatically imply increase of blood flow. Indeed, having such a high proportion of occluded microvessels, not few areas of the peritoneal tissue become, with time, underperfused. This point should be taken into consideration in order to analyze the pathophysiology of permeability changes commonly detected in cases of membrane failure. This, in addition to the increased thickness of the peritoneal membrane that, per se, will substantially affect the transit time of solute’s molecules between the still permeable capillaries and the peritoneal cavity.

All osmotic agents present in commercially available solutions for peritoneal dialysis share, regarding the long-term exposed monolayer, at least three basic effects: a substantial reduction of the cell population density of around 50%, a mitotic index near zero, and a significantly increased prevalence of nonviable cells [423].

This information suggested that the monolayer became depopulated under the influence of dialysis solutions (Figs. 5.59 and 5.60). But, going from bad to worse, the regenerative capabilities of the mesothelial cells still dressing the cavitary surface of the peritoneum appeared substantially reduced. In addition, the mechanisms leading to regeneration are substantially restrained, at least in experimental grounds, by the continuous exposure to the commonly used osmotic agents. This concept finds support in experimental observations done in rats using the “Doughnut” model of mesothelial regeneration in the rat [419]. After a ring of the monolayer was exfoliated, animals were exposed to either 4.25% glucose or 7.5 icodextrin dialysis solution for a period of 30 consecutive days. Macroscopic observation of the abdominal cavity at the end of the observation period showed that most animals developed scarring and fibrous adhesions at the level of the injured areas. Imprints and biopsies taken from the affected domains confirmed that not only repopulation failed, but that the missing mesothelial dressing was replaced by a thick layer of fibrous tissue, containing numerous microvessels of neoformation (Fig. 5.61). Consequently, these observations support the contention that both osmotic agents, 4.25% glucose and 7.5 icodextrin, substantially restrain the normal process of mesothelial repopulation expected to take place during and after the experimental exfoliation. This development launched the repair mechanisms leading to peritoneal sclerosis.

figure 5_59_978-0-387-78940-8 — **Fig. 5.59**

figure 5_60_978-0-387-78940-8 — **Fig. 5.60**

figure 5_61_978-0-387-78940-8 — **Fig. 5.61**

Several studies have shown evidence indicating that different cell types exposed to hydrogen peroxide display a reduced rate of proliferation, premature senescence, and, consequently, higher prevalence of apoptosis [437]. Within this context, additional experiments exposed to view the existence of a dose-related effect. Indeed, low levels of oxidants potentiate growth signals and enhance proliferation as long as the specific cell type can initiate new rounds of mitosis (Fig. 5.50), whereas higher concentrations of oxidants can block cell proliferation, which, in turn, derives in premature senescence and the consequent activation of the mechanisms leading to apoptotic cell death [438] (Figs. 5.45 and 5.46).

Higher degrees of oxidative injury lead to cell death by nonphysiological, necrotic pathways that, in turn, put in motion the local inflammatory reaction derived from extravasation of the cytoplasmic contents into the interstitial tissue [439] (Fig. 5.62). Both glucose-enriched solutions and icodextrin have the intrinsic capabilities of inducing different degrees of oxidative stress upon the exposed mesothelium. Glucose acts through products derived from its nonenzymatic degradation [440] and the irreversible formation of AGE products [441] by glucose autoxidation [442] and/or by oxidative mitochondrial DNA damage [443]. Icodextrin, in turn, induces substantial lipid peroxidation of mesothelial cell’s membrane almost immediately after being infused into the abdominal cavity [444, 445].

figure 5_62_978-0-387-78940-8 — **Fig. 5.62**

This injury derives, at least in part, from the intra-abdominal formation of carbonyl compounds during the dwell time [446]. This phenomenon of carbonyl compounds liberation during the dwell time has been shown by the same group of investigators, using amino acid–based dialysis solutions.

Development of oxidative injury is facilitated by the demonstrated capability of mesothelial cells in culture to generate hydrogen peroxide [447]. This interpretation of the above-mentioned chain of events has been substantiated by a recent study, showing that acute and severe in vivo oxidative stress applied to the mesothelial monolayer results in extensive fibrosis, adhesions, and permeability changes similar to those observed in clinical ultrafiltration failure [448] (Fig. 5.63).

figure 5_63_978-0-387-78940-8 — **Fig. 5.63**

Peritoneal sclerosis has been induced in rodents by in vivo exposing the membrane to a variety of experimental interventions: asbestos [449], 0.1% chlorexidine [450], iron dextran [451], glucose degradation products [452], AGE deposits derived from uremia per se [453], sodium hypochlorite [454], lipopolysaccharide [455], low pH of around 3.8 [456], pure water combining low pH and hypo-osmolarity [457], silica [458], and zymosan [459].

It should be noticed at this point of the analysis that, with a few exceptions (pure water, chloroxidine, and low pH), the other substances quoted as used to experimentally induce peritoneal sclerosis operate setting out different degrees of oxidative stress [460–471]. So far, after evaluating the aforementioned offered evidence, it may be concluded that addition of antioxidant agents to the currently used peritoneal dialysis solutions seems to be a quite rational and wanted development [472, 473].

We cannot complete this review without mentioning the enigmatic problem of sclerosing encapsulating peritonitis (SEP), currently also named encapsulating peritoneal sclerosis (EPS). This fearful syndrome leads to a situation in which a thickened, fibrous sheet of tissue envelops the small intestine [474–477], liver, and stomach, as well as pelvic organs. This complication covers a wide range of morphological alterations starting from peritoneal opacification, passing through the tanned peritoneum syndrome, and finally reaching replacement of the serosal layer by fibrous tissue. Fibrous bands may be present compromising mesentery, gallbladder, spleen, liver, and stomach, as well as pelvic organs and even the cavitary aspect of the peritoneal tissue. The most affected areas configure, at times, a mass of fibrous tissue packaging abdominal viscera, conforming the cocoon that usually includes loops of small intestine as well as pockets of encapsulated ascites. Light microscopy reveals serosal fibrosis and total absence of the mesothelial monolayer, replaced by a thick layer of connective fibrous tissue, the thickness of which can reach 4 cm [478, 479] (Fig. 5.64 ). Neovascularization is also seen, even though these blood vessels show major structural alterations: sclerosis of the whole vascular wall, at times occlusion of the lumen, and even hyaline changes of the blocked microvessels (Fig. 5.65 ). Peritoneal calcifications and formation of bone and even bone marrow have been detected. All this may be combined with wide areas of inflammatory infiltrates [480, 481].

figure 5_64_978-0-387-78940-8 — **Fig. 5.64**

figure 5_65_978-0-387-78940-8 — **Fig. 5.65**

Changes detected in patients with SEP/EPS appear far away from those described in the commonly observed peritoneal sclerosis. Besides, its prevalence in patients on peritoneal dialysis is, fortunately, extremely low, whereas the impact of each condition is absolutely different. SEP/EPS carries a quite poor prognosis, with a mortality rate ranging between 26 and 93% [480], whereas simple peritoneal sclerosis basically leads patients to switch to other ways of renal replacement therapy, usually hemodialysis. These differences support the concept postulated by Di Paolo and Garosi [478, 479], who concluded that both conditions represent different nosological entities. In patients on peritoneal dialysis, the origin of this complication is still ill-defined. Many possible factors have been invoked (acetate, hyperosmolarity, recurrent peritonitis, glucose, antiseptics, intraperitoneal antibiotics, bacterial endotoxins), even though there is no available evidence clearly demonstrating specific relevance for any of them [474, 482]. Besides, at least one case has been reported on a chronic uremic patient having renal replacement therapy by means of only hemodialysis [483].

It should be noticed that sclerosing peritonitis has been reported in not few clinical situations unrelated to both, chronic uremia and peritoneal dialysis as: idiopathic [484, 485], associated with the use of some β-blockers such as practolol [486], propranolol [487], or timolol [488], as well as to metoprolol [489], oxprenolol [490], and intraperitoneally administered antibiotics such as tetracycline [491]. Besides, the literature mentions cases of sclerosing peritonitis associated to intra-abdominal tumors like gastric carcinoma, carcinoma of pancreas, familial polyposis of colon, renal carcinoma, lymphoma, ovarian teratoma or thecoma, and even in patients affected by liver cirrhosis as well as after liver transplantation [492–502]. So far, our understanding of SEP/EPS is still blurred as a result of the complexity of the problem, and namely, by its multiple ethiopathogenesis.

The Potential Use of the Mesothelium as a Source of Mesenchymal Stem Cells

As stated in the previous paragraphs, the existence of pluripotent mesenchymal cells as precursors of the mesothelium has been already proposed. Even though a mesothelial stem cell has not yet been definitely identified, the existence of pluripotent mesenchymal cells in the mesothelial monolayer as well as in the submesothelial connective tissue, has been repeatedly considered in the literature [401], opening the way in order to consider their use, as an actual option, in regenerating therapies [503].

Some observations performed in human pathology as well as in animal experimentally induced tissue reactions lend strong support to this contention. As a living proof, it may be mentioned that differentiation towards cartilage and bone has been described in a primary tumour of pleura, suggesting that mesothelial cells are pluripotent. In this sense, being mesenchyma, they may well retain the potential to differentiate along embryonic developmental lines, including cartilage and bone [504, 505].

Besides, cartilaginous differentiation of the peritoneum not associated with intra-abdominal malignancy has been already detected [506], whereas bone and cartilaginous formation has been reported in both, human patients and experimentally induced mesothelioma [507, 508]. Bone formation was also seen in four cases of sclerosing peritonitis observed in patients treated by means of peritoneal dialysis, whereas in two of them, islands of bone marrow were also detected [481].

Interestingly, glomerular-like structures in fibrous tissue of biopsies of visceral peritoneum have been detected in biopsies taken from rats with experimentally induced peritoneal sclerosis [505] (Fig. 5.66 ). This information emphasizes the capability of the mesothelium to differentiate in other cell lines in response to injury [504].

figure 5_66_978-0-387-78940-8 — **Fig. 5.66**

Additional research identified mesenchymal cells in the adult human synovial membrane. These cells showed the capability to differentiate in chondrocytes and osteocytes [509, 510]. Synovium, also of mesenchymal origin, is considered one more type of serous membrane like peritoneum, pericardium and pleura [511]. Therefore, this common embryological origin opens the possibility of eventual therapeutic interventions in the course of joint diseases that could be performed using peritoneal mesothelial cells [505].

Recent published evidence exposed to view the fact that, when placed under the appropriate biophysical and/or biochemical conditions mesothelial cells demonstrate a remarkable degree of plasticity. This property supports the concept, that mesothelial cell progenitors are endowed with the capability to switch between different cell types, according to the conditions of their microenvironment [504]. Within this context, it is illustrating to remind the observed myofibroblastic conversion of human adult mesothelial cells in culture, under the influence of transforming growth factor (TGF)-β-1 [512, 513].

The relationship between blood cells and mesothelium represents one more exciting aspect of this topic. As mentioned before, a proposed mechanism of mesothelial healing postulated that progenitor cells, originally located in the bone marrow, migrate and convert into mesothelial cells [380]. Although additional research concluded that this hypothesis looks unlikely [403], a link between both cell lines seems to be possible. A quite strong point is that hemangioblasts, a proposed progenitor of the endothelial and hematopoietic cell lineages, derive from the embryonic splanchnic mesothelium. This structure, in turn, has been proposed as the embryonary source of the endothelium-lined vascular system, pointing at a specialization of the phylogenetically older celomic cavities. Within this context, the origin of the hematopoietic cells might be related to differentiation of celomocytes derived from the celomic epithelium. So far, endothelial and blood cells appear to derive from a common mesothelial-derived progenitor [514].

In addition, morphological and immunohistochemical evidence for a translocation of cells from the celomic mesothelium to the ventral wall has been observed during development of the quail embryos. Consequently, the concurrence of translocation of mesothelial cells and the appearance of aortic smooth muscle cell progenitors point at a link between the former and the latter cells lineages [515].

This ontogenetic relationship between mesothelium, blood, and blood vessels is substantiated by two excellent studies performed applying the tools of tissue bioengineering. Donna et al. [516] presented evidence demonstrating that cultured adult human mesothelial cells have the capabilities to generate hematopoietic cells, similar to those of the bone marrow. This conversion was confirmed by morphological analysis as well as by cell immunoreactivity toward specific antibodies directed to antigens of the hematopoietic cell lines, at various stages of differentiation. The experiment was performed culturing mesothelial cells in collagen sponges. This is one more suggestion of the remarkable plasticity of the mesothelium, as well as of the relevance of the microenvironment hosting the cultured cells. Regarding blood vessels, a key study is that reported by Campbell et al. [517], who succeeded in creating an artificial blood conduct by inserting a Silastic tubing into the peritoneal cavity of rats. Two weeks after the surgical intervention, a new laparotomy showed that the implanted silicon rubber tubing was covered by several layers of fibroblasts, collagen matrix, and a single layer of mesothelium. This new “blood vessel” was everted and successfully grafted by end-to-end anastomosis, in arteries of the same animal in which it was grown. These observations have been confirmed by Moldovan and Haveman [518], as well as by our group [505].

So far, it seems evident that peritoneal mesothelial cells are endowed with a degree of plasticity that shapes their capability of generating other cell lines, if placed in the appropriate micro-environment. Investigative steps will have to define the best conditions that will eventually lead to the use of mesothelium in stem cell therapy as well as in tissue engineering, taking also in account that harvesting of large numbers of cells from patients having an unexposed monolayer, may well be unlimited.

Final Remarks

It was not the purpose of the author merely to deliver a cold and tedious description of anatomical structures. On the contrary, the goal has been to offer the reader a comprehensive and balanced analytical approach of structure and function covering, at least in part, their interactions. It is evident that the function of the peritoneum as a dialysis membrane cannot be evaluated only within the frame of passive diffusion through water-filled, cylindrical pores [204] and/or mathematical models [519], based on assumptions that, at times, lose sight of the formidable barrier of the living cell membrane as well as the structural organization of the tissues.

Research during the last two decades provided enough evidence to characterize the peritoneum not as an inert dialyzing sheet, but as a living and reusable membrane for dialysis [271], as predicted more than 25 years ago [520].

It becomes evident that the mesothelial monolayer continuously exposed to dialysis solutions in vivo is structurally and functionally different, at least from the histochemical point of view, from that observed in unexposed–intact cells, or in those growing in the in vitro set-up of culture and later exposed to experimental incubation [521]. Therefore, I have the feeling that a good deal of creative thinking is required to integrate data obtained during 50 years of physiological studies and mathematical models, with the realities of tissue structure and cell biology.

Introduction of peritoneal dialysis as a therapeutic tool to fight chronic uremia shaped a kind of chain reaction that went well beyond the expectations of the early years. Actually, observations made in clinical settings showed the way to a new window open to the fascinating world of cell biology. Within this specific field, there are still more questions than answers.

It is the author’s hope that this chapter will serve to stimulate the imagination of young scientists as a catalytic element for further research.

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Ha’Emek Medical Center, Department of Nephrology, Afula, Israel
L. Gotloib

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School of Medicine Department of Medicine Division of Nephrology, MA436 Health Sciences Center, Columbia, MO 65212, USA
Ramesh Khanna
Academic Medical Center Dept. Medicine, University of Amsterdam, Meibergdreef 9, 1105 AZ Amsterdam, The Netherlands
Raymond T. Krediet

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Gotloib, L. (2009). Functional Structure of the Peritoneum as a Dialyzing Membrane. In: Khanna, R., Krediet, R.T. (eds) Nolph and Gokal's Textbook of Peritoneal Dialysis. Springer, Boston, MA. https://doi.org/10.1007/978-0-387-78940-8_5

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Functional Structure of the Peritoneum as a Dialyzing Membrane

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The Evolution of Peritoneal Dialysis Solutions

Physiology of Peritoneal Dialysis

Monitoring the Functional Status of the Peritoneum

Keywords

Normal Mesothelium

Interstitium

Blood Microvessels

Lymphatics

Peritoneal Innervation

Cytology of the Peritoneal Fluid

Ultrastructure of Peritoneal Fluid Cells

The Origin of the New Mesothelial Cells

The Price of a Failing Regeneration

The Potential Use of the Mesothelium as a Source of Mesenchymal Stem Cells

Final Remarks

References

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Functional Structure of the Peritoneum as a Dialyzing Membrane

Similar content being viewed by others

The Evolution of Peritoneal Dialysis Solutions

Physiology of Peritoneal Dialysis

Monitoring the Functional Status of the Peritoneum

Keywords

Normal Mesothelium

Interstitium

Blood Microvessels

Lymphatics

Peritoneal Innervation

Cytology of the Peritoneal Fluid

Ultrastructure of Peritoneal Fluid Cells

The Origin of the New Mesothelial Cells

The Price of a Failing Regeneration

The Potential Use of the Mesothelium as a Source of Mesenchymal Stem Cells

Final Remarks

References

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