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BJU Int. Author manuscript; available in PMC 2008 Jun 1.
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PMCID: PMC1961637

Differentiation Potential of Urothelium from Patients with Benign Bladder Dysfunction



Benign dysfunctional bladder diseases encompass a number of poorly understood clinically-defined conditions, including interstitial cystitis (IC), idiopathic detrusor overactivity (IDO) and stress urinary incontinence (SUI). We developed a novel in vitro approach to test the hypothesis that failure of urothelial differentiation underlies the aetiopathology of IC, where there is evidence of compromised urinary barrier function.

Materials and Methods

Biopsy-derived urothelial cells from dysfunctional bladder biopsies were propagated as finite cell lines and examined for their capacity to undergo differentiation in vitro, as assessed by acquisition of a transitional cell morphology, a switch from a CK13lo/CK14hi to a CK13hi/CK14lo phenotype, expression of claudin 3, 4 and 5 proteins and induction of uroplakin gene transcription.


2/12 SUI cell lines showed early senescent changes in culture and were not characterised further; 1/7 IC, 1/5 IDO and a further 3 SUI cell lines displayed some evidence of senescence at passage 3. Of the IC-derived cell lines, 4/7 showed a near normal range of differentiation-associated responses, but the remainder of IC lines showed little or no response. A majority of IDO cell lines (4/5) showed a normal differentiation response, but at least 3/10 SUI cell lines showed some compromise of differentiation potential.


Our study supports the existence of a subset of IC patient in whom a failure of urothelial cytodifferentiation may contribute to the disease and provides a novel platform for investigating the cell biology of urothelium from SUI and other benign dysfunctional conditions.

Keywords: bladder, claudin, differentiation, idiopathic detrusor overactivity, interstitial cystitis, stress urinary incontinence, uroplakin


The urothelium is the highly-specialised transitional epithelium found lining the urinary bladder and associated urinary tract. Organised into basal, intermediate and terminally-differentiated superficial cell zones, the urothelium provides a highly effective barrier that prevents urine from penetrating the underlying tissues. The urinary barrier properties are primarily attributable to the specialisation of the terminally-differentiated superficial urothelial cells, which display molecular features that limit permeability via transcellular and paracellular routes.

The apical surface of the superficial cell membrane is covered with multiple thickened plaques of asymmetric unit membrane. The asymmetric unit membrane is a unique ultrastructural feature of urothelial cells that forms as a result of interactions between integral transmembrane proteins, known collectively as the uroplakins (UPK). There are four major uroplakin proteins that interact to form UPK1a/UPK2 and UPK1b/UPK3a pairs; disruption of asymmetric unit membrane formation, for example by germline deletion of the UPK3a gene(1), leads to major alterations in urothelial morphology and function, including increased transcellular permeability.

The intercellular tight junctions constitute the main paracellular barrier. The capacity for different epithelia to display a range of paracellular permeability functions and to modulate these in response to (patho)physiological signals is dictated by differential expression of the claudins, which form the primary seal-forming fibrils of the tight junction. Claudins constitute a family of some 24 proteins and in urothelium show a differentiation stage-related pattern of expression, with claudin 7 expressed in all but the superficial layer of the urothelium, claudins 4 and 5 expressed at basolateral junctions of superficial cells and claudin 3 restricted to the “kissing points” between adjacent superficial cells(2).

Interstitial cystitis (IC) is a chronic and often debilitating inflammatory disorder of the urinary bladder characterised by urinary urgency, frequency and bladder pain, in the apparent absence of any infectious agent. The aetiology and pathophysiological mechanisms of IC remain undetermined(3, 4) and it is considered unlikely that a single causal mechanism is responsible. A number of reports have suggested that a compromised urothelial barrier is a feature of the disease(5-7). However, it is unclear whether this is due to an inherent dysfunction of the urothelium itself, or an indirect influence of the local (e.g. cytokine) environment on urothelial differentiation and function.

A number of other benign dysfunctional conditions exist in the bladder, including urge urinary incontinence secondary to idiopathic detrusor overactivity (IDO) and stress urinary incontinence (SUI) associated with urodynamic stress incontinence. Although some cases of detrusor overactivity are neurogenic in origin, involving dysregulation of bladder function secondary to changes in the peripheral or central nervous system, in many cases the cause remains unidentified and the condition is classified as IDO(8). Due to its intimate association with the sub-urothelial afferent nerve fibres and a potential role in bladder sensation, it has been suggested that the urothelium may be involved in the aetiopathology of IDO(9), although urothelial barrier dysfunction has not been associated with the condition. SUI is a complaint of involuntary leakage on effort or exertion, or on sneezing or coughing. The urothelium has not been implicated in the aetiopathology of SUI, for which the principal mechanisms involve anatomical changes to the pelvic floor resulting in a loss of support at the bladder neck, and compromised neuromuscular function of the urethral sphincter(10).

We have developed methods to isolate and propagate normal human urothelial (NHU) cells as finite cell lines from surgical resection specimens(11, 12). In a low [0.09mM] calcium, serum-free culture system, NHU cells have a proliferative phenotype driven by autocrine/paracrine activation of the epidermal growth factor receptor (EGFR)(13) and can be induced to undergo differentiation in response to activation of the nuclear receptor peroxisome proliferator activated receptor gamma (PPARγ) when downstream signalling through EGFR is blocked(2, 14, 15). The in vitro cytodifferentiation of NHU cells is accompanied by specific changes in the transcription and/or translation of cytokeratins, claudins and uroplakins that relate to urothelial differentiation in situ and serve as objective markers of differentiation response(2, 14, 15). Thus, we have developed the tools to assess the differentiation potential of human urothelial cells in culture.

In the present study, we have established a novel method for the growth of finite urothelial cell lines from multiple (three) small cystoscopic cold-cut biopsies retrieved from patients with IC (diagnosed according to strict NIDDK guidelines(16)), IDO and SUI. In addition to investigating morphology and growth properties, we have examined whether urothelial cells from the different patient groups could be induced to undergo molecular differentiation in terms of a switch from a CK13lo/CK14hi to a CK13hi/CK14lo phenotype, expression of claudin 3, 4 and 5 proteins and induction of uroplakin gene transcription.

Materials and Methods

Pharmacological Reagents

The high affinity PPARγ ligand, troglitazone (TZ), was a kind gift from Parke-Davis Pharmaceutical Research (Ann Arbor, USA). The EGF receptor tyrosine kinase inhibitor, PD153035, was obtained from Calbiochem-Novabiochem Biosciences Ltd. (Nottingham, UK).


All urothelial specimens were collected with appropriate Local Research Ethics Committee approval and full informed patient consent. A bladder-derived NHU cell line established from a surgical resection specimen from a patient with no history of atypia or malignancy was included in all experiments as a positive control(11, 12). Four cold-cut biopsies were obtained from the non-trigone bladder of patients with IC diagnosed according to NIDDK specifications(17). Biopsies were similarly obtained from patients with clinically-proven urge urinary incontinence secondary to idiopathic detrusor overactivity (IDO) and stress urinary incontinence (SUI) secondary to urodynamic stress incontinence, as defined by the International Continence Society(18). There was no statistical difference in the age of patients for each group (IC mean age 51, range 25-67; IDO mean age 47, range 27-71; SUI mean age 52, range 38-80). Biopsies were collected in 10 ml Transport Medium consisting of sterile Hanks’ Balanced Salt Solution (HBSS) with Ca2+ and Mg2+ (Invitrogen Ltd, Paisley, UK) containing 10 mM HEPES (pH7.6) and 20 KIU/ml aprotinin (Trasylol; Bayer plc, Newbury, UK). One biopsy was fixed for 1 hour in 10% (v/v) formalin and processed into paraffin wax for immunohistology; the remaining three biopsies were pooled and used to establish primary urothelial cell cultures by scaling down and adapting methods developed for resection specimens (see below).

Haematoxilin and eosin-stained sections from all three patient groups were assessed to exclude carcinoma in situ and other inflammatory conditions causing identical symptoms(19), as well as to assess urothelial integrity for immunolabelling.


Cold cut biopsies were fixed in 10% (v/v) formalin in PBS for 1 hour, dehydrated in ethanol to isopropanol then xylene, before careful orientation and embedding in paraffin wax. This protocol was strictly adhered to in order to best preserve superficial cells and avoid potential processing artefacts caused by over-fixation. Haematoxylin and eosin-stained sections were screened for areas of full thickness urothelium for analysis of differentiation antigen expression.

Immunohistochemistry was performed on dewaxed 5 μm sections using the StreptABComplex/Horseradish Peroxidase system from Dako Cytomation (Ely, UK), as previously described(15). Primary antibodies are listed in Table 1. Blocking steps were included to neutralise any endogeneous peroxidase and avidin-binding activities and to prevent non-specific binding of secondary antibody. For most antibodies, sections were boiled for 10 minutes in 10 mM citric acid buffer, pH 6.0 to retrieve antigens lost during tissue processing. The exception was for retrieval of the CK20 antigen, where sections were digested for 10 minutes in 0.1% (w/v) trypsin in 0.1% (w/v) CaCl2, pH 7.6 at 37°C. Following overnight incubation in primary antibody at 4°C, slides were washed, incubated sequentially in biotinylated secondary antibodies and a streptavidin biotin horseradish peroxidase complex (Dako Cytomation) and visualised using a diaminobenzidine substrate reaction (Sigma-Aldrich Ltd, Poole, UK). The sections were counterstained with haemotoxylin, dehydrated and mounted in DPX (Sigma-Aldrich). Negative and positive antibody specificity controls were included in all experiments.

Table 1

Urothelial Biopsy Cell Culture

Biopsies were centrifuged in the original Transport Medium for 4 minutes at 61 × g in order to salvage any shed urothelial cells. To separate the urothelium from the stroma, pellets were resuspended in 3 ml of HBSS (without Ca2+ and Mg2+) containing 10 mM HEPES pH7.6, 20 KIU/ml aprotinin and 0.1% (w/v) EDTA and incubated for 3 hours at 37°C. Pellets were collected by centrifugation, flicked to resuspend and any stromal pieces removed with sterile forceps. The remaining urothelium was incubated in 1 ml collagenase type IV (200 U/ml, Sigma-Aldrich Ltd.) for 20 minutes at 37°C. After centrifugation at 61 × g for 4 minutes, media were aspirated and pellets resuspended in 2 ml Keratinocyte Serum-Free Medium complete (KSFMc) with bovine pituitary extract and epidermal growth factor at the manufacturer′s recommended concentrations (Invitrogen Ltd.) and 30 ng/ml cholera toxin (Sigma-Aldrich Ltd.). Cell suspensions were seeded into a 3 cm Primaria™-coated dish (BD Biosciences, Oxford, UK) and the medium was renewed every two or three days.

Biopsy-derived patient urothelial cell cultures were maintained at 37°C in a humidified atmosphere of 5% CO2 in air and passaged at just-confluence, exactly as detailed(11, 12). Cultures were transferred from a 3 cm Petri-dish (Passage 0) to a 1 × 25 cm2 flask (Passage 1), to 4 × 25 cm2 flasks (Passage 2), of which one flask was cryopreserved and the other three flasks transferred into 2 flasks each and used for analysis of differentiation potential (Passage 3). All experiments were performed with a bladder NHU cell line (Y607), which was established from a resection specimen and included as a positive control.

Analysis of Differentiation Potential

The differentiation assay was based on the optimised conditions for inducing uroplakin gene expression in NHU cell cultures we have described previously(14). Control NHU cell lines were grown to 70% confluence and treated with or without 1 μM TZ for 24 hours followed by incubation with or without 1 μM PD153035 for up to 6 days. Due to the limited availability of cells from biopsy-derived urothelial cell lines, experiments were constrained to a) 1 μM TZ for 24 hours followed by 1 μM PD153035 and b) no treatment control (0.01% v/v DMSO). Media were changed every 3 days and cells were harvested for RNA (day 3) or protein (day 6), as with the NHU cell controls. These time points were selected as optimal from our previous work(14). Cells seeded on 12-well Teflon®-coated slides and treated as above were used for immunofluorescence experiments.

Indirect Immunofluorescence

Slides were fixed in methanol and acetone (vol:vol), air-dried and incubated with titrated primary antibody (Table 1), or no primary antibody as control, for 16 hours at 4°C. After washing in PBS, the slides were incubated with secondary antibodies conjugated to Alexa 488 (Molecular Probes, supplied by Invitrogen). To visualise nuclei, Hoechst 33258 (0.1 μg/ml; Sigma-Aldrich) was included in the penultimate wash. Slides were observed on an Olympus BX60 microscope under epifluorescence illumination.

Western Blot Analysis

Cells grown in 25 cm2 flasks were treated in situ with lysis buffer (25 mM Hepes pH 7.4, 125 mM NaCl, 10 mM NaF, 10 mM Na3VO4, 10 mM Na4P2O7, 0.2% (w/v) SDS, 0.5% (w/v) sodium deoxycholate, 1% (w/v) Triton X-100, 1 μg/ml aprotinin, 10 μg/ml leupeptin and 100 μg/ml phenylmethylsulfonyl fluoride) and the lysates were sheared by passing three times through a 21-gauge needle. After 30 minutes on ice, lysates were microcentrifuged at 10,000 × g for 30 minutes at 4°C before the protein concentrations of supernatants were determined by Coomassie assay (Pierce, supplied by Perbio Science UK Ltd, Cheshire, UK). Cell extracts were resolved on 12.5% SDS polyacrylamide gels, transferred electrophoretically onto nitrocellulose membranes and membranes were incubated with titrated primary antibodies (Table 1) for 16 hours at 4°C. Bound antibody was detected with anti-mouse Alexa Fluor® 680 (Molecular Probes) and anti-rabbit LI-COR IRDye™ 800 (Rockland, supplied by Tebu-Bio Ltd, Peterborough, UK) and quantified using the Odyssey™ Infrared Imaging System (LI-COR Biosciences UK Ltd., Cambridge).

Reverse-Transcribed Polymerase Chain Reaction (RT-PCR)

RNA was extracted from cell monolayers using Trizol™ (Invitrogen Ltd.) and isolated by chloroform extraction and isopropanol precipitation, as recommended by the manufacturer. The RNA was treated with DNase I to remove any DNA contamination (DNA-free™ kit, Ambion Europe Ltd., Huntingdon, UK) and cDNA was synthesised using 1 μg of total RNA and the Superscript™ first-strand synthesis system (Invitrogen Ltd.), as outlined by the manufacturer. PCR was performed as previously described using Surestart Taq polymerase (Stratagene Europe, Amsterdam, The Netherlands) and primer pairs designed to amplify specific claudin products, as described previously(2). Based on previous work(20), GAPDH was included as the internal transcript control using forward primer: 5’-GTCGGAGTCAACGGATTTGG-3’ and reverse primer: 5’-ATTGCTGATGATCTTGAGGC-3’. PCR reactions were performed as follows: 95°C for 12 minutes, then 35 cycles of 95°C for 1 minute, optimum annealing temp for 1 minute and 72°C for 1 minute, and a final extension at 72°C for 10 min in a PCR Express Thermal Cycler (Hybaid Ltd, Ashford, UK). All experiments were performed alongside no-template and no RT controls. PCR products were run on a 2% (w/v) agarose gel and visualised with ethidium bromide.

Real-Time RT-PCR

cDNA was synthesised as outlined above. Quantitative real-time PCR was performed using TaqMan real-time PCR primers and probes designed for uroplakin genes using the Primer Express Software (Applied Biosystems UK, Warrington) (Table 2). The reactions were performed in TaqMan Universal PCR master mix (Applied Biosystems) with 200 nM of probe and 300 nM of primers on an ABI Prism 7700 Sequence Detector system (Applied Biosystems) for 10 minutes at 95°C, followed by 40 cycles of 15 seconds at 95°C and 60 seconds at 60°C. The data were analysed using the ABI Prism 7700 SDS software (Applied Biosystems). Data were normalised against GAPDH, used as the internal control(20).

Table 2
Taqman PCR primers and probes

Statistical Methods

GraphPad InStat software (www.graphpad.com) was used for statistical analysis. Means and medians were used as descriptive statistics and plotted as solid and dashed lines, respectively. Non-parametric methods (2-tailed Mann Whitney U-test or the 2-tailed Wilcoxon matched-pairs signed-ranks test, as appropriate) were used for tests of statistical significance. On graphs, the level of significance is indicated between marked groups as * p ≤ 0.05; ** p ≤ 0.01; *** p ≤ 0.001.


Histology and Immunohistochemistry

The urothelial thickness on biopsies ranged from two to six cell layers. Focal denudation of the urothelium was not accompanied by underlying granulation tissue or oedema and was regarded as artefactual. The IC cases revealed variable dense non-specific chronic inflammation in the lamina propria. No granulomatous or eosinophilic inflammation was seen in any patient group.

Assessment of the in situ differentiation status of urothelium from patient biopsies was limited to biopsies that upon histological examination, displayed an intact, full thickness urothelium. Sections from IC, IDO and SUI biopsy specimens were screened with a panel of differentiation-associated antigens (Figure 1). In keeping with normal adult human urothelium (ureter control tissue), the urothelia from all patients were mitotically quiescent, with very few cells in the cell cycle, as assessed by an absence of Ki67 (MIB-1) labelling.

Figure 1

Two cytokeratin isotypes were investigated: CK7, which is expressed in all urothelial cell layers and CK20, which labels superficial and occasional intermediate cells in normal urothelium(reviewed(21)). In most biopsies, CK7 showed a heterogeneous labelling pattern, with patches of intense full thickness CK7 labelling, interspersed by patches of weak labelling; this phenomenon was not related to clinical derivation and has been described previously in normal bladder urothelium(22). CK20-positive cells were identified in all specimens, primarily located at the superficial edge, but often with the morphology of late intermediate cells, suggesting either a loss or absence of mature superficial cells. UPK3a was localised discretely along the apical edge of superficial urothelial cells in intact urothelium, but the extent of UPK3a labelling in patient biopsies varied from near normal to absent and, as with the CK20 labelling, reflected either the absence or artefactual shedding of superficial cells.

Expression and localisation of tight junction-associated proteins, occludin and the claudins 4, 5 and 7, was very similar to ureteric urothelium(2), once allowance was made for the absence or loss of some superficial cells (Figure 1). Claudins 4 and 5 were most intensely expressed at basolateral junctions of superficial cells and were useful discriminatory markers for the presence or absence of superficial cells (Figure 1). One IC sample showed absence of claudin 5 expression from the urothelium, which was despite claudin 5 expression in the vascular endothelium and confirmed presence of UPK3a-positive superficial cells in that sample (Figure 1). Claudin 3 is restricted to the terminal junction between superficial urothelial cells, but appeared negative in all biopsy specimens, although this interpretation was confounded by the loss/absence of superficial cells.

The results from the immunohistochemistry studies indicated no gross differences between IC urothelium and the other benign conditions (IDO and SUI), with the possible exception of claudin 5 down-regulation in one IC specimen. However, the differentiated phenotype could only, by definition, be assessed in areas of full thickness urothelium. In order to determine more specifically the differentiation and function potential of urothelium from IC patients, a cell culture system was developed.

Growth of bladder urothelial cells from biopsy specimens

Urothelial cells isolated from each of three biopsies were used to establish successful primary urothelial cell cultures from six IC, five IDO and twelve SUI patients (Table 3). In a further two cases of IDO and one case of SUI, primary culture was unsuccessful due to the isolation of too few urothelial cells. One additional IC cell line was established from tissue obtained at cystectomy (referred to as IC1). In all cases, primary cultures displayed a typical epithelioid pavement morphology, comparable to NHU cell cultures established from bladder resection specimens (Figure 2). In the majority of cases, the primary cultures grew to confluence and were successfully sub-cultured as finite cell lines through a minimum of three passages, generating adequate cell numbers to assess differentiation and functional potential (Table 3 and see below). The exceptions were two SUI cell lines, which were growth compromised and failed to grow beyond passage 2; these were not characterised further. By passage 3, it was noted that cell lines from one IC, one IDO and three further cases of SUI had begun to show some evidence of senescence, with reduced cell growth and the appearance of larger, mitotically-quiescent cells (Table 3 and Figure 2).

Figure 2
Phase contrast micrographs
Table 3
Summary of Results for Individual Cell Lines

Response to activation of PPARγ

We have demonstrated previously that cultured NHU cells undergo a programme of differentiation-associated gene expression changes in response to activation of PPARγ and that this response is enhanced when signalling downstream of the EGF receptor is blocked(14). Immunofluorescence labelling of the cystectomy-derived IC1 cell line confirmed that PPARγ and RXRα showed an equivalent localisation and intensity to that seen in the NHU bladder control cell line, with both antigens showing a predominantly nuclear localisation (Figure 3).

Figure 3
Localisation of PPARγ and RXRα in NHU and IC urothelial cells

To determine whether urothelial cells from IC, IDO and SUI patients retained the capacity to differentiate, cells were treated with the PPARγ agonist, TZ, and the EGFR-specific tyrosine kinase inhibitor, PD153035. The NHU bladder cell line control and a proportion of cultures from all three patient groups showed morphological evidence of response to treatment with TZ and PD153035, forming rosettes of tear-shaped cells within 3 days of treatment (Table 3 and Figure 2 arrows), as has been described previously for NHU cell cultures(14).

Treatment with TZ and PD153035 has been shown previously to switch NHU cells from a squamous to a transitional differentiation programme, as assessed by the shift in cytokeratin expression from a CK13lo/CK14hi to a CK13hi/CK14lo profile(15). This was confirmed in the bladder-derived NHU cell line by immunoblotting (Figure 4 and Table 3). Characterisation of IC-derived cell lines revealed induction of CK13 expression and decrease in CK14 expression in response to TZ/PD153035 in 3/6 lines, but a much reduced (n=2) or absent (n=1) response in the remaining three cell lines tested. Of the IDO cell lines tested, 2/4 showed an induction of CK13 and decrease of CK14. In the majority of SUI-derived cell lines (7/10), expression of CK13 was either not induced or induced only marginally, although in 8/10 cases, there was a reduction in CK14 expression in response to treatment (Figure 4 and Table 3).

Figure 4
Influence of differentiation on CK13 and CK14 expression in cultured IC, IDO and SUI urothelial cells

The induction of differentiation in NHU cells with TZ and PD153035 is associated with de novo expression of genes associated with urothelial terminal differentiation, including the uroplakins(14). The expression of uroplakin UPK1a, UPK1b, UPK2 and UPK3a transcripts was quantified by real-time PCR in the IC, IDO and SUI cell lines and compared to the control bladder NHU cell line (Figure 5). In all three patient groups, there was a significant increase in uroplakin gene expression in the differentiated versus non-differentiated conditions, and there was also a significantly higher expression of UPK1b in differentiated SUI versus IC groups (Figure 5). When assessed in terms of individual samples, two of the IC cell lines showed a consistent lack of induction of UPK1a, UPK2 or UPK3a gene expression in response to TZ and PD153035, whereas the remaining IC lines showed induction of UPK1a and UPK2 to at least 50% of the NHU cell line control (Figure 5 and Table 3). All of the IDO cell lines responded to some degree, but 3/9 SUI cell lines showed only limited up-regulation of UPK1a, UPK2 and UPK3a expression after treatment (Figure 5).

Figure 5
Effect of TZ and PD153035 on uroplakin expression in cultured IC, IDO and SUI urothelial cells

Development of tight junctions by cultured IC, IDO and SUI cells

One of the features of urothelial cytodifferentiation is the formation of intercellular tight junctions associated with changes in the transcription and protein stability of the claudins(2). In agreement with our previous study using ureter-derived NHU cells, the bladder-derived NHU cell line expressed claudins 1, 2, 4, 5 and treatment with TZ and PD153035 resulted in induction of claudin 3 (Figure 6A). Claudin transcript expression was examined by RTPCR in three IC and two IDO cell lines. All showed constitutive expression of claudins 1, 2 and 4, and baseline expression of claudin 3 was detected in 2/3 IC and 2/2 IDO cell lines. There was some evidence of upregulation of claudin 3 in response to treatment with TZ and PD153035 in 1/2 IC cell lines and in 2/2 IDO cell lines examined by RTPCR (Figure 6A).

Figure 6Figure 6
Influence of TZ and PD153035 on claudin expression in cultured NHU, IC and IDO urothelial cells

At the protein level, treatment of control bladder NHU cells with TZ and PD153035 resulted in large increases in claudins 4 and 5, with a small induction of the late differentiation marker claudin 3 (Figure 6B and C); this was in agreement with our previous study of ureteric NHU cell lines(2). In addition, claudins 1 and 7 showed some change in expression, which were non-significant in ureteric NHU cell lines(2). A similar pattern was seen in the patient groups, with a significant increased expression of claudins 4 and 5 in response to differentiating conditions, although claudin 5 was poorly induced in some individual cell lines (Tables (Tables33 and and4).4). Induction of claudin 3 was more variable and was absent in 3/6 IC, 1/4 IDO and 6/10 SUI cell lines (Figure 6B and 6C; Table 3). It was noted that in samples where there was a failure to induce claudin 3, the induction of claudin 5 was correspondingly weak.

Table 4
Changes in Claudin Expression following Induction by TZ and PD153035

In the cystectomy-derived IC cell line (IC1), which showed a normal claudin expression pattern, response to TZ and PD153035 was associated with re-localisation of tight junction-associated proteins to intercellular borders, as seen in the control bladder NHU cells (Figure 7).

Figure 7
Immunofluorescence showing the effect of differentiation on the expression and localisation of CK13 and claudin proteins in cultured urothelial cells


This is the first report of the application of an in vitro cell culture system for normal human urothelium to the study of dysfunctional bladder syndromes. Because clinical considerations limit the amount of urothelium that can be harvested by cystoscopic biopsy, a modified method for the isolation and in vitro expansion of urothelial cells from very small amounts of starting material has been developed. We considered it critical that control groups for the study should be derived from similarly harvested tissues and due to ethical constraints preventing the biopsy of normal bladders, this led us to a decision to compare IC to two other benign dysfunctional bladder syndromes of IDO and SUI.

Objective comparison of the in vitro differentiation potential of urothelial cells from different patient groups, especially when selected by stringent clinical criteria, may reveal disease-specific differences and hence open a path to identifying the underlying mechanisms. However, diseases such as IC are defined primarily by symptomatology, and may reflect convergent progression pathways from more than one causal mechanism. Several causal mechanisms for IC can be postulated. A leaky urinary barrier giving rise to symptoms of cystitis could be caused by an inherent failure of urothelial cytodifferentiation, but alternatively could reflect a response to exogenous factors derived from, for example, the immunological microenvironment, having an influence on urothelial tissue integrity. In order to determine the influence of intrinsic factors, we used an in vitro approach wherein extrinsic factors are absent, or introduced experimentally under controlled conditions. Although our study has not addressed all questions and will need to be extended to include larger sample populations, nevertheless, we believe it offers a new platform for understanding the pathogenesis of benign bladder diseases and will open the door to identifying biomarkers to differentiate disease subsets based on mechanism. This in turn will lead to development of therapies aimed at causal rather than symptom relief.

We have described previously the culture of NHU cells from resection specimens. Despite being derived from a mitotically-quiescent tissue in situ, cultures of NHU cells have a remarkable, albeit finite, proliferative capacity, reflecting the regenerative capacity of native urothelium(13). Although the biopsies were derived from diseased bladders, there was little evidence of proliferation being driven in situ, as seen from the lack of Ki67-positive cells by immunohistochemistry. This was even the case in the IC-derived specimens, where chronic inflammatory-mediated tissue damage might be expected to initiate regenerative field changes. The small starting cell population obtainable from biopsies did limit ultimate expansion capacity compared to either the bladder resection-derived NHU cell line included as control, or the cystectomy-derived IC1 cell line. Nevertheless, urothelial cells derived from a majority of biopsies showed a proliferative phenotype in culture, enabling adequate expansion through limited serial passage to enable controlled differentiation studies. An unexpected finding was that cell lines showing premature senescence were mostly derived not from the IC, but from the SUI group. This difference was not due to donor age per se, as there was no age-related pattern of senescence (mean age of donors for non-senescent and senescent cultures was 50 and 51 years, respectively).

Our previous studies have identified a PPARγ-mediated pathway that, in the absence of EGFR signalling, initiates the programmed expression of gene and protein changes associated with urothelial cytodifferentiation, including characteristic changes to the cytokeratins(15), claudins(2) and uroplakins(14). From this programme, we selected several components to use as objective markers of differentiation in response to PPARγ activation: a) morphology, b) switch from a CK14 squamous to a CK13 transitional phenotype; c) induction of claudin 3, 4 and 5 protein expression; and d) induction of uroplakin gene transcription.

A morphological change from a homogeneous squamous pavement morphology to the formation of rosettes of “tear-shaped” cells of characteristic transitional morphology provided an indicator of PPARγ-mediated response and occurred in 50% of biopsy-derived cultures that showed no senescent change. However, it was not an absolute predictor of response to PPARγ, as there was at least one cell line (IC5) that did not show a morphological change, but still showed responses at the molecular level. A majority of biopsy-derived cell lines showed some molecular evidence of response to PPARγ, although this response was limited in the case of three of the IC-derived cell lines (IC4, IC6 and IC7).

As markers, the claudins provided particular insight. In situ, the claudins show a stage-related pattern of expression in urothelium, with claudins 4 and 5 expressed at a later stage of differentiation than claudin 7 and occludin, but prior to claudin 3, which, together with ZO-1, is localised in the terminal junction between adjacent superficial cells(2). In vitro, PPARγ activation results in de novo expression of claudin 3 and the stabilisation of claudins 4 and 5 proteins(2). With the exception of one IC and one SUI cell line, all biopsy-derived cell lines showed upregulation of claudins 4 and 5, but detection of claudin 3 was limited to cases where there was a strong upregulation of claudin 5 protein. Claudin 3 and ZO1 can be difficult to locate in tissue sections and the absence/loss of superficial cells from biopsies restricted the use of these markers. However, at least one of the informative IC cases was negative for claudin 5 in situ. Given the probability that IC has no single cause, we suggest that there may be a subset of IC cases in which there is aberrance in the formation of tight junction structures during urothelial differentiation, which would be predicted to affect paracellular permeability. Future studies using a more sophisticated 3D differentiation model (23) are proposed, as these could combine functional paracellular permeability assessment with claudin expression profiling and hence may provide functional evidence to support claudin dysregulation in the aetiopathology of a subset of IC. However, we also emphasise a note of caution for follow-up studies, as we have found many tissue blocks from hospital archives to show artefactual loss of claudin antigenicity, possibly due to poorly controlled fixation conditions (unpublished observations).

Although urothelial dysfunction has been implicated in IC and IDO, there has been no prior suggestion of urothelial involvement in the aetiopathology of SUI, which was therefore presumed to represent a “normal” urothelium. The unexpected finding from our study was that urothelial cells from a number of patients with SUI showed poor growth and differentiation characteristics. Previous research has mainly focused on the diagnosis and treatment of SUI, however, the role of nerve damage in the aetiology of the disorder is attracting interest as it offers a potential therapeutic target. It has been suggested that the purinergic signalling pathway may be involved in functional motor as well as sensory bladder disorders(24). It is reasonable to hypothesise that interaction between the neuromuscular elements of the bladder wall and the urothelium may play a role in more than just bladder dynamics and bladder sensation and that dysregulation of this intimate relationship may underlie the pathophysiology of bladder dysfunction syndromes that do not appear to include a primary sensory element. Further work to characterise the purinergic receptors present in human urothelium in normal and diseased states would assist in defining the precise relationship between urothelial sensation and function.

In summary, a novel in vitro approach has been developed to investigate the growth and differentiation potential of urothelium from dysfunctional bladder syndromes and provides a platform for investigating the causal mechanisms in IC, IDO and SUI. Our results suggest that there may be a subset of IC cases in which the differentiation capacity of the urothelium is compromised, possibly through derangement of tight junction structure, which would give rise to a leaky urothelium and associated symptoms of chronic cystitis.


The authors would like to thank Professor JJ Walker for helpful discussions. They are extremely grateful to Drs C Ramage, L Rogerson and G Urwin who assessed patients and provided clinical specimens for the study. JS holds a Research Chair funded by York Against Cancer.

Contract grant sponsor: NIH

Contract grant number: IR21 DK066075


EGFRepidermal growth factor receptor
IDOidiopathic detrusor overactivity
ICinterstitial cystitis
NHUnormal human urothelial
PPARγperoxisome proliferator activated receptor gamma
SUIstress urinary incontinence
TERtransepithelial resistance


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