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Correspondence to Tung-Tien Sun: sunt01{at}med.nyu.edu
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P. Hu's present address is Procter & Gamble Corporate Research Biotechnology, Cincinnati, OH 45252.
Abbreviations used in this paper: 2D, two-dimensional; BUN, blood urea nitrogen; ES, embryonic stem; IVP, i.v. pyelogram; UP, uroplakin; VUR, vesicoureteral reflux.
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40% identical in sequence, and belong to the "tetraspanin" family (Yu et al., 1994). The tetraspanin family contains many cell surface molecules, including CD9, CD63, CD81, CD82, and CD151, that play key roles in cellular events involving membrane organization (Maecker et al., 1997; Levy et al., 1998; Berditchevski, 2001; Boucheix and Rubinstein, 2001; Hemler, 2001, 2003; Tarrant et al., 2003; Yunta and Lazo, 2003). Uroplakins II and III both have a single transmembrane domain, and they share a common stretch of 12 amino acid residues on the exoplasmic side of the transmembrane domain (Wu and Sun, 1993; Lin et al., 1994; Deng et al., 2002). Within the plaques, these four major uroplakins are organized into two heterodimers consisting of Ia/II and Ib/III, as demonstrated by chemical cross-linking (Wu et al., 1995) and protein isolation (Liang et al., 2001). In addition, transfection studies show that when individual uroplakins are expressed in 293T cells, they are retained in the ER; however, coexpression of uroplakins Ia plus II or Ib plus III permits the heterodimers to exit from the ER; these events suggest that the formation of specific heterodimers is a prerequisite for uroplakins to reach the cell surface (Tu et al., 2002).
To study the functional and disease implications of uroplakins, we previously generated uroplakin III–deficient mice using the gene-targeting approach (Hu et al., 2000). Such mice lack a typical urothelial umbrella cell layer. They have a reduced urothelial plaque size, compromised urothelial permeability barrier function, and retrograde flow of urine from the bladder into the ureters (vesicoureteral reflux [VUR]). In 70% of the mice, accumulation of urine in the renal pelvis (hydronephrosis) occurs (Hu et al., 2000, 2002). These results establish that uroplakin III is an integral subunit of urothelial plaques, which contribute to the permeability barrier function of the urothelial apical surface, and suggest that urothelial defects may play a role in VUR (Hu et al., 2000, 2002).
Some of the phenotypes of these UPIII-deficient mice remain hardly understood. For example, the observation that UPIII deficiency leads to the formation of small plaques seems incompatible with our current assumption that all four major uroplakins are required for the formation of the 16-nm particle (Sun et al., 1999). One possible explanation is that UPIIIb, a newly discovered, minor UPIII isoform (Deng et al., 2002), is responsible for the formation of the residual plaques observed in the UPIII-deficient urothelium. If it is, one would predict that knockout of uroplakin II, which does not have a known isoform, should completely abolish plaque formation. Another question has to do with the possible role of uroplakin mutation in VUR. Although knockout of the UPIII gene leads to VUR in mice, screening of human VUR patients revealed that polymorphism of uroplakin genes is only marginally associated with VUR (Giltay et al., 2004; Jiang et al., 2004). Moreover, although polymorphism was found in these VUR patients, no deletion, truncation, or frameshift mutations have been detected so far (Giltay et al., 2004; Jiang et al., 2004). The role of uroplakin defects in VUR and other lower urinary tract anomalies thus remains unclear.
To address these issues, we have genetically ablated the mouse UPII gene to perturb the other uroplakin pair consisting of UPIa/II. This led to the entrapment of UPIa in the ER, but allowed the remaining UPIb/III pair to reach the apical urothelial surface. No 16-nm particle or apical plaques were formed, however, which indicates that both uroplakin heterodimers were required for plaque formation. The small size of the superficial cells in both UPII- and UPIII-deficient urothelia suggested that the incorporation of fully assembled uroplakin plaques into the apical surface through fusion of the fusiform vesicles with the apical membrane played a role in the enlargement of umbrella cells. Because entire litters of UPII-deficient mice from some breeding pairs (originating from the inbred 129/SvEv and outbred Swiss Webster strains) reproducibly died 8–10 d postnatally, as a result of renal failure, it seems that major uroplakin defects such as gene deletion, frameshift, or truncation might cause urinary tract anomalies that, in severe cases, could lead to death. These results highlight the critical functional importance of uroplakins in the formation of a specialized urothelial apical surface, and shed light on the mechanisms of plaque formation, umbrella cell enlargement, and certain urinary tract abnormalities.
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24 cm of H2O (Fig. 6 c), >50% (11 out of 20) of the mice refluxed at a pressure lower than this (Fig. 6, d and e). Many of the (–/–) mice developed severe hydronephrosis with a greatly expanded renal pelvis (Fig. 7, b and d) and associated renal morphological changes (Fig. 7, a–f). To determine whether reflux caused hydronephrosis, we calculated the difference between the PM and the PR of each ureter (PK [hydrostatic pressure to the kidney] = PM – PR). For a normal mouse that had a PM of 25 cm of H2O and a PR of 50 cm, an intravesicular pressure of >25 cm of H2O would result in micturition and dissipation of pressure. However, for a UPII knockout mouse that had a PM of 25 cm of H2O but a lower PR of 18 cm of H2O, intravesicular pressures of 18–25 cm of H2O would lead to reflux, thus potentially transmitting up to 7 cm of hydrostatic pressure to the kidney (PK). We wanted to see whether this renal pressure correlated with the grade (G) of hydronephrosis—which we defined as G = D/T, where D was the internal diameter of the renal pelvis and T was the thickness of the remaining renal parenchyma (Fig. 7, c and d). If reflux were a main cause of hydronephrosis, there should have been a positive correlation between PK and G. However, we found that these two parameters were independent (P > 0.2), which suggests that reflux could not be the major etiology of hydronephrosis in this system (Fig. 7 g).
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Mechanism of uroplakin plaque formation
As mentioned earlier, the four major uroplakins can form two heterodimer pairs consisting of uroplakins Ia/II and Ib/III (Wu et al., 1995; Liang et al., 2001; Tu et al., 2002). Transfection studies indicated that the formation of correct heterodimer is required for uroplakins (except UPIb; see below) to exit from the ER and to reach the cell surface (Tu et al., 2002). Because 293T (a human embryonic kidney–derived cell line) cells were used in these transfection studies, the physiological significance of in vitro data obtained using nonurothelial cells might be questioned. Our in vivo data show that although UPIa is distributed throughout the cytoplasm of the UPII-deficient urothelium, suggesting ER entrapment (Fig. 3 d), uroplakins Ib and III (of the Ib/III pair) can reach the urothelial surface (Fig. 3, f and j). These in vivo results validate the transfection data obtained using the 293T cells (Tu et al., 2002), and further support the existence of the Ia/II and Ib/III heterodimers.
Although cultured bovine urothelial cells continue to synthesize large amounts of all the uroplakins, these cells do not form any mature-looking, uroplakin-delivering fusiform vesicles that are characteristic of the in vivo umbrella cells, and their apical cell surface is completely devoid of urothelial plaques (Surya et al., 1990; Chen et al., 2003). Based on what we have learned from cultured keratinocytes (Weiss et al., 1984; Schermer et al., 1989), cultured urothelial cells most likely mimic in vivo regenerating urothelium (Surya et al., 1990; Chen et al., 2003). The inability of regenerating urothelial cells to assemble plaques could be functionally important, because a cell surface laden with the rigid-looking plaques may be incompatible with cell migration and proliferation during wound repair. Therefore, although we can study certain early steps of uroplakin interaction in the currently available, cultured urothelial cells and transfected 293T cells, the final stages of uroplakin assembly and 2D crystal formation need to be studied using the in vivo urothelium. Our present finding that the uroplakin II–deficient urothelium lacks 16-nm particles (Fig. 4, c and d) indicates that, although the UPIb/III pair can be delivered to the apical surface (Fig. 3, f and j) using the small vesicles (150–200 nm in diameter) accumulated in the UPII-deficient superficial cells (Fig. 5, b, g, and h), the UPIb/III pair by itself cannot form the 16-nm particle, let alone the urothelial plaques (Fig. 4 d). Severs and Hicks (1979) have shown that, in normal urothelium, 16-nm particles can be detected in early vesicles that have just budded off the Golgi apparatus, which suggests that the two uroplakin heterodimers (Ia/II and Ib/III) interact before they leave the Golgi apparatus to form the 16-nm particle. Together, these data suggest that uroplakins first form, in the ER, heterodimers (Tu et al., 2002), and possibly heterotetramers; the heterotetramers then assemble into the 16-nm particles in the Golgi apparatus or the trans-Golgi network, and these particles, once reaching a sufficient density in the post-Golgi vesicles, can then aggregate to form small and, later, large 2D crystals.
Given our current assumption that both uroplakin pairs are required for the formation of the 16-nm particle, it was baffling that the UPIII-deficient urothelium still made 2D crystals, albeit small ones, from the 16-nm uroplakin particles (Hu et al., 2000). However, we now know that highly purified urothelial plaques contain a minor isoform of UPIII, which we recently characterized (Deng et al., 2002). This 35-kD protein was named uroplakin IIIb because it is urothelium specific, it shares a similar transmembrane topology and significant sequence homologies with UPIII (also known as UPIIIa), and it forms a heterodimer with UPIb, UPIIIa's partner (Deng et al., 2002). This newly found uroplakin IIIb, which is up-regulated in UPIIIa-deficient urothelium (Deng et al., 2002), should allow the formation of small amounts of heterotetramers containing heterodimers UPIb/UPIIIb and UPIa/UPII. The heterodimers can then be delivered by the 150–200-nm vesicles (Fig. 5, e and f) and account for the formation of small surface plaques in the UPIIIa-deficient urothelium. This interpretation is consistent with our finding that the ablation of UPII, which does not have a known isoform, greatly diminishes the amounts of small vesicle-associated uroplakins and completely abolishes the formation of the 16-nm particles (Fig. 4, c and d).
We showed previously that the ablation of UPIII led to abnormal synthesis and processing of UPIb, i.e., the level of UPIb mRNA was greatly increased, whereas the amount of UPIb protein was reduced, became hypoglycosylated, and was mistargeted to the basal/lateral cell surface (Hu et al., 2000). Because UPIII and UPIb were known to interact, we speculated that these UPIb changes were caused by the removal of its partner, UPIII (Hu et al., 2000). Our finding that UPII ablation led to a similar up-regulation of the UPIb mRNA level (Fig. 1 C) and hypoglycosylation of the UPIb protein (Fig. 1 E) indicates, however, that these UPIb changes represent a general response to a perturbed uroplakin assembly. This may explain why, in cultured human urothelial cells, the level of mRNA for UPIb is greatly elevated compared with those for other uroplakins (Varley et al., 2004). Uroplakin Ib is also unique in that it is the only uroplakin that, when expressed alone in 293T cells, can exit from the ER to reach the cell surface (although it is required for its partner, UPIII, to exit from the ER [Tu et al., 2002]); and in that it is the only uroplakin that is expressed in nonurothelial tissues, including corneal and conjunctival epithelia (Adachi et al., 2000) and, possibly, lung epithelium (Kallin et al., 1991; Olsburgh et al., 2003). These data indicate that uroplakin Ib is unique among the uroplakins in its regulation and function.
Uroplakin assembly and umbrella cell enlargement
Normal mouse urothelium is covered by large umbrella cells that have an average diameter of >100 µm (Hicks, 1965; Koss, 1969). The greatly enlarged superficial cells can minimize the intercellular space, thus contributing to the permeability barrier function of the apical urothelial surface (Negrete et al., 1996; Zeidel, 1996; Lewis, 2000; Apodaca, 2004). The mechanism by which these large umbrella cells, which are frequently tetraploid or octoploid (Farsund, 1975), are formed is unclear, although both arrested cytokinesis (Farsund, 1976; Farsund and Dahl, 1978) and cell fusion (Martin and Wong, 1981) have been suggested. Our finding that the superficial cells of both UPII- and UPIII-deficient urothelia failed to enlarge (Fig. 4, a and b; Hu et al., 2000) suggests that urothelial plaque formation may play a role in umbrella cell enlargement. It is possible that the continued insertion of fusiform vesicles into the apical surface not only expands (Porter and Bonneville, 1963; Hicks, 1965; Koss, 1969; Staehelin et al., 1972; Lewis and de Moura, 1982; Truschel et al., 2002; Apodaca, 2004), but also alters/stabilizes, the luminal surface, thus hindering cytokinesis; the fact that some umbrella cells can incorporate BrdU (Fig. 2, c and d; Erman et al., 2004) makes it plausible that arrested cytokinesis could contribute to polyploidy and cell enlargement. A combination of cell surface expansion and arrested cytokinesis may thus account for the formation of large polyploid umbrella cells, without resorting to cell fusion. This "surface alteration hypothesis" is supported by cell kinetic data suggesting arrested cytokinesis (Farsund, 1976; Farsund and Dahl, 1978); however, additional studies are needed to test this hypothesis.
Uroplakins and urothelial growth
A striking phenotype of both uroplakin II– and uroplakin III–deficient urothelia is that they become hyperplastic, with a BrdU-labeling index of >10%, versus a normal value of 0.1% (Fig. 2, c and d) (Hu et al., 2000). It is possible that the permeability barrier function of the uroplakin-deficient urothelia is compromised (Hu et al., 2002) so that the EGF-related mitogens that are known to be present in high concentrations in the mouse urine (Lakshmanan et al., 1990; Parries et al., 1995) can reach urothelial basal cells to stimulate cell proliferation (Rebel et al., 1994). Alternatively, fully assembled urothelial plaques can trigger certain growth-inhibiting signals, which are now absent in the uroplakin-deficient urothelia, leading to hyperplasia. Additional studies are needed to distinguish these possibilities.
Roles of uroplakin defects in urinary tract diseases
Our earlier observation that inactivation of the mouse uroplakin III gene resulted in VUR and hydronephrosis raised the possibility that uroplakin defects may cause VUR in humans (Hu et al., 2000; Mak and Kuo, 2003). By surveying 76 VUR patients, we identified 18 single-nucleotide polymorphisms; 7 of them are missense, with no truncation or frameshift mutations (Jiang et al., 2004). Only two of the missense polymorphisms (an Ala7Val change in the UPIa gene and a Pro154Ala change in the UPIII gene) were found to be marginally associated with VUR (both P = 0.08). These results, as well as those by Giltay et al. (2004), suggest that the uroplakin point mutations that have been detected so far do not play a major role in causing VUR (Jiang et al., 2004). There are two possible explanations for this apparent inconsistency between the human and mouse data. First, the results of mouse studies cannot be directly extrapolated to humans because of minor, but still significant, differences in the development of the lower urinary tracts. Perhaps for this reason, inactivation of angiotensin II receptor (AT2) gene can cause VUR in mice (Nishimura et al., 1999), but polymorphisms in this gene do not seem to be involved in causing primary familial VUR in humans (Hohenfellner et al., 1999; Yoneda et al., 2002). Second, and perhaps more importantly, no truncation or frameshift mutations have been detected in any of the VUR patients that have been studied so far, and some breeding pairs of the UPII and UPIII knockout mice reproducibly die postnatally around days 8–10, as a result of renal failure (Fig. 10; Hu et al., 2000), suggesting that certain uroplakin defects (such as deletion, truncation, or mutations in some structurally crucial amino acid residues), in combination with certain genetic backgrounds (Sanford et al., 2001; Wolfer et al., 2002), may be embryologically or postnatally lethal and are thus undetected in VUR patients. It is interesting to note that a recent survey showed that urinary tract abnormalities account for 20% of all congenital anomalies detected during pregnancies and up to one year after birth (Scott, 2002). Of the 560 cases of prenatal or neonatal deaths resulting from congenital abnormalities, 58% were because of renal anomalies, including a significant number with obstruction (Scott, 2002).
An important finding of this study is that urothelial abnormalities caused by uroplakin defects can lead to renal failure and death (Fig. 10). There are several possible mechanisms by which this could occur. First, defects in the uroplakin-positive renal pelvis urothelium, as well as in the simple epithelium that covers the renal papilla expressing uroplakin prenatally (Fig. 10 e; Jiang et al., 2004), may lead to renal dysfunction and failure. Second, urothelial hyperplasia can cause obstruction of the ureteral lumen (Fig. 8, f and h). Although it has been well-established that mechanical obstruction can induce urothelial hyperplasia (Monson et al., 1992; Curhan et al., 2000), it is unclear whether urothelial hyperplasia, per se, can cause obstruction. In our system, urothelial hyperplasia seems to be the cause, rather than the effect, of obstruction because (a) urothelial hyperplasia precedes obstruction and hydronephrosis; and (b) although only 70–80% of the mice develop hydronephrosis, 100% of the animals show urothelial hyperplasia throughout the urinary tract, thus ruling out obstruction as a cause of urothelial hyperplasia (but see Gamp et al., 2003). Third, defects in the ureteral urothelium may interfere with the development of its underlying smooth muscle (Master et al., 2003), leading to abnormal peristalsis or functional obstruction (Santicioli and Maggi, 1998)—a possibility supported by our recent finding that uroplakin defects lead to abnormal bladder muscle contractility and/or function (Christ, G., personal communication). These possible mechanisms, or a combination thereof, may cause a functional/structural obstruction of the urinary tract, leading to abnormal renal development (Chung and Chevalier, 1996), obstructive nephropathy, hydronephrosis, and, in severe cases, renal failure and death.
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Morphological studies
Mouse urothelium was examined by scanning EM (model JSM-840; JEOL) and transmission EM (model 200CX; JEOL) (Hu et al., 2000). Quick-freeze deep etch was performed as described previously (Kachar et al., 1999). Cell proliferation was assessed based on the nuclear incorporation of BrdU (Sigma-Aldrich). 19-d-old mice received five intraperitoneal injections of BrdU (100 mg/kg of body weight, in PBS), 1.5 h apart, and were killed 2 h after the final injection. Various tissues, including esophagus and urinary bladder tissues, were removed, fixed in 10% formalin overnight, and processed for immunostaining with a horseradish peroxidase–conjugated anti-BrdU monoclonal antibody (CHEMICON International). Samples were visualized with a microscope (Axiophot; Carl Zeiss MicroImaging, Inc.) with 10x/0.32 and 20x/0.60 (Plan-Apochromat) or 40x/0.75 (Plan-Neofluar) objective lenses. Images were captured with a digital camera (model DKC-5000; Sony) at room temperature. The images were processed in size and contrast/brightness with Adobe Photoshop 6.0.
Isolation of detergent-insoluble proteins from mouse urothelium
Mouse urothelial cells were homogenized in buffer A (10 mM Hepes, pH 7.5, 1 mM EDTA, 1 mM EGTA, and 1 mM PMSF), loaded onto a 1.6-M sucrose cushion (in buffer A), and centrifuged at 16,000 rpm for 25 min at 4°C (SW41; Beckman Coulter). The crude membranes concentrated at the interface were isolated, washed with buffer A, treated with 2% Sarkosyl in buffer A for 10 min at 25°C, and pelleted. The detergent-insoluble membranes, in which the urothelial plaques were highly enriched, were washed with buffer A before they were solubilized in SDS-PAGE sample buffer and analyzed by SDS-PAGE, followed by silver nitrate staining, or by immunoblotting (Wu et al., 1994; Liang et al., 1999; Zhou et al., 2001).
Determination of reflux pressure and hydronephrosis
The reflux pressure was recorded as the hydrostatic pressure (i.e., cm of H2O) at which an India ink suspension in PBS backflowed from the bladders into the ureters of anesthetized mice (Hu et al., 2000). At the end of the experiment, the kidneys were dissected longitudinally to confirm the presence of the ink in the renal pelvis (Fig. 6 b).
Determination of the micturition pattern and urine/blood chemistry
The micturition pattern of the mice was recorded using special cages, as described previously (Hu et al., 2000). A total of 500 µl of mouse urine was collected from each mouse and used for assaying the concentrations of uric acid, creatinine, potassium, sodium, and chloride ions (Tufts Veterinary Diagnostic Laboratory). Mouse sera were used for assaying the BUN level (Anilytics).
Determination of obstruction by IVP and serial sectioning of the mouse ureter
IVP was performed in anesthetized 4-mo-old wild-type (n = 2) and uroplakin II–deficient (n = 4) mice. The contrast Omnipaque (Iohexol; 3 µl/g of body weight) was injected into the orbital sinus, and serial radiographs were made at 0, 5, 15, 30, 45, 60, and 90 min. After lethal anesthesia, the urinary tracts were dissected and fixed in 10% formalin overnight. The ureters were embedded in paraffin and serially sectioned. Sections were numbered from superior to inferior orientation and stained with hematoxylin and eosin.
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This work was funded by National Institutes of Health grants DK39753, DK52206, and DK66491.
Submitted: 7 June 2004
Accepted: 9 November 2004
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