1 EMBL, Meyerhofstrasse1, D-69117 Heidelberg, Germany
2 Department of Developmental Biology, Utrecht University, Padualaan 8, NL-3584CH Utrecht, The Netherlands
* Present address: Division of Molecular Embryology, DKFZ, Im Neuenheimer Feld 280, D-69120 Heidelberg, Germany
Author for correspondence (e-mail: R.Zeller{at}bio.uu.nl)
Accepted June 6, 2001
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SUMMARY |
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Key words: Differentiation, EMT, FGF signalling, Process formation, Podocyte
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INTRODUCTION |
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Mature podocytes (or glomerular visceral epithelial cells) are highly differentiated cells within the kidney glomerulus that function in primary renal filtration (Saxén, 1987). During nephrogenesis podocytes originate as immature epithelial cells from the metanephric mesenchyme after induction by the ureteric bud (Saxén, 1987; Sorokin and Ekblom, 1992). During their subsequent differentiation, expression of epithelial markers ceases (Garrod and Fleming, 1990; Tassin et al., 1994), and mesenchymal markers such as vimentin and actin-associated synaptopodin are induced (Mundel et al., 1997a; Nagata et al., 1993). These changes coincide with extensive cytoskeletal reorganisation and extension of actin-based, branched cellular processes, the so-called foot processes (Saxén, 1987). Thus, podocytes appear to undergo EMT-like changes during their terminal differentiation. In support of this proposal, glomeruli of mice deficient for the homeobox transcription factor Pod-1 are nonfunctional due to a failure in terminal differentiation of podocytes (Quaggin et al., 1999), and morphological analysis showed that mutant podocytes were arrested in an epithelial state. Several FGFs and their receptors are expressed by nephrons (including podocytes [(Cancilla et al., 1999; Cauchi et al., 1996; Dono and Zeller, 1994; Mason et al., 1994; Ohuchi et al., 2000; Peters et al., 1992); and this study]) and are required for kidney morphogenesis (Celli et al., 1998; Ohuchi et al., 2000; Qiao et al., 1999; see also Discussion). Of relevance to the present study, FGF2 proteins are upregulated during podocyte differentiation and remain expressed in functional podocytes of the adult kidney (Dono and Zeller, 1994). However, the potential role of FGF2 signalling during podocyte differentiation and function remained unclear as no gross kidney defects were reported in mice carrying a Fgf2 loss-of-function mutation (Dono et al., 1998; Ortega et al., 1998).
To identify and study possible FGF signalling functions during podocyte differentiation and process formation, we used an established in vitro culture and cell differentiation system for mouse podocytes (Mundel et al., 1997b) based on conditional immortalisation (Jat et al., 1991). Conditionally immortalised mouse podocyte (MPC) cells cultured under permissive conditions grow as undifferentiated epithelial cells, whereas they develop mesenchymal characteristics similar to mature podocytes in vivo upon induction of post-mitotic differentiation. In particular, differentiating MPC cells reorganise their cytoskeleton and form branched actin-based processes, similar to foot processes (Majumdar and Drummond, 1999; Mundel et al., 1997b; Nagata and Watanabe, 1997). Here we have used the MPC cell culture system to study the differentiation potential of Fgf2 mutant podocytes in culture. Our studies indicate that FGF signalling is required for the EMT-like changes during differentiation, which results in actin cytoskeletal reorganisation and process formation.
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MATERIALS AND METHODS |
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Cell proliferation assays
Mitotic MPC cells were detected using the 5-bromo-2'-deoxy-uridine (BrdU) labelling and detection kit II (Boehringer Mannheim) as described (Mundel et al., 1997b). Proliferating cells in chicken embryonic mesonephric kidneys were labeled in organ culture for 48 hours with 100 µM BrdU. Mitotic and FGF2 expressing cells were detected on histological sections as described previously (Dono et al., 1998).
Analysis of cell morphology
MPC cell morphology was assessed by microscopic observation of living cells. Images were captured using a COHU CCD camera mounted on a Nikon eclipse TE 200 microscope in combination with the Scion Image software program. For indirect immunofluorescence, cells cultured on glass coverslips were fixed with either methanol/acetone (1:1) for 15 minutes at -20°C (ZO-1) or 4% PFA for 10 minutes at ambient temperature (all other markers). Following fixation, cells were washed with PBS, permeabilised with 0.3% Triton X-100 in PBS for 5 minutes (PFA fixed cells only) and blocked in 3% BSA, 3% FBS and 0.2% gelatine in PBS for 1 hour at room temperature (RT). Subsequently, cells were incubated for 1 hour at RT with either antibodies against FGF2 (Dono and Zeller, 1994), synaptopodin (Mundel et al., 1997a), paxillin (Transduction Laboratories), vimentin (C-20; Santa Cruz) or ZO-1 (Zymed) in blocking solution. After washing with PBS containing 0.2% gelatine and 1% Triton X-100, cells were incubated for 30 minutes at RT with the appropriate secondary antibodies. FITC-conjugated phalloidin (0.5 µg/ml, Sigma) was used to visualise actin filaments and DAPI (Boehringer) for nuclei. Images of immunofluorescent cells were captured with the OpenLab 1.7.6 software program (Improvision) using a Photonic-Science cooled CCD camera attached to a Zeiss Axioscope microscope. All illustrations were assembled and processed digitally using Adobe Photoshop 5.0.
Immunoblots and cell fractionation
MPC cells were harvested after mild trypsin digestion and washed with ice-cold PBS containing 1 mM PMSF. Cell pellets were resuspended in ice-cold extraction buffer (20 mM Tris, pH 7.5, 0.5 M NaCl, 1% Triton X-100, 1 µg/ml aprotonin, 1 µg/ml leupeptin, 0.5 µg/ml pepstatin and 0.5 mM PMSF; all Sigma) at a concentration of approximately 107 cells/100 µl. Suspensions were first homogenised, then sonicated on ice (20 Watts for 1 minute) and cleared by centrifugation (10,000 g for 30 minutes at 4°C). Separation of suspension into nuclear, cytoplasmic and membrane fractions was performed as described by (Dono and Zeller, 1994) with the following modifications: the crude cytoplasm was separated into membrane and cytoplasmic fractions by ultra-centrifugation (100,000 g for 30 minutes). The membrane pellets were dissolved in RIPA buffer (50 mM Tris-HCl, pH 7.4, 30 mM NaCl, 5 mM EDTA, 1% NP-40, 1% deoxychloate and 0.1% SDS). The protein contents of all total extracts and fractions were determined (Biorad kit) and extracts normalised. For FGF2 immunoblot analysis, 1 mg of total protein was heparin enriched. Antibodies against -tubulin (Sigma) and c-Jun/AP1 (Santa Cruz) were used as controls to assess the quality of the nuclear-cytoplasmic fractionation. Immune complexes were visualised by chemoluminescence according to manufacturers instructions (Amersham). Three independent experiments yielded results very similar to the ones shown in Fig. 1E,F.
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RESULTS |
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Fgf2 mutant MPC cells fail to undergo normal morphological differentiation and initiate process formation
To address the functional significance of FGF signalling during podocyte maturation, kidneys of mice carrying an Fgf2 loss-of-function mutation (Dono et al., 1998) were analysed. In a large population of adult mice housed under non-SPF conditions only a very small fraction of Fgf2 mutant mice (10 of 266) displayed symptoms of renal failure with glomerosclerosis and podocyte damage. Such defects were not observed in a similarly large group of wild-type mice (G.D., unpublished). The rare and stochastic nature of this phenotype in adult mice suggests that Fgf2 deficiency is at best a contributing factor to renal failure, which is normally compensated for in vivo. In an attempt to circumvent this potential functional compensation, we isolated and cultured wild-type and Fgf2 mutant podocytes for comparative molecular analysis. Conditionally immortalised MPC cells were isolated from apparently normal kidneys of Fgf2 mutant adult mice carrying the inducible and temperature-sensitive large T-antigen transgene (Jat et al., 1991). For all phenotypic analysis shown in Figs 2- 5, normalised pools of wild-type and Fgf2 mutant MPC cell clones were used to exclude clone-specific variation (see Materials and Methods). Wild-type and Fgf2 mutant MPC cells have similar proliferation and mitotic arrest kinetics (data not shown) as expected from conditional immortalisation (Jat et al., 1991). However, analysis of the growth characteristics under permissive conditions showed that Fgf2 mutant MPC cells tend to grow more as aggregates compared to their wild-type counterparts (compare Fig. 2A with Fig. 2B). Alterations in cell morphology became even more apparent upon shifting MPC cells to nonpermissive culture conditions. Post-mitotic, Fgf2 mutant cells remain tightly associated (Fig. 2D) and fail to form cellular processes (compare inset in Fig. 2C with inset in Fig. 2D) in contrast to the arborised cell morphology of differentiated wild-type cells (Fig. 2C).
General disruption of FGF signalling in Fgf2 mutant MPC cells
One possible explanation for this apparent discrepancy between intact kidneys and cultured MPC cells could be the loss of functional compensation by other FGFs in culture. Therefore, the levels of other FGFs and FGF receptors that are expressed in embryonic and/or adult kidneys (Cancilla et al., 1999; Ford et al., 1997; Mason et al., 1994; Ohuchi et al., 2000; Peters et al., 1992) were determined semi-quantitatively by RT-PCR (Fig. 3). The relative transcript levels in wild-type (+/+) and Fgf2 mutant (-/-) MPC cells were normalised and compared to the ones of dissected wild-type and mutant kidney cortex tissue (enriched in glomeruli). This analysis confirmed loss of Fgf2 mRNA in mutant MPC cells and kidneys (Fig. 3). Analysis of Fgf2 mutant kidney cortex tissue revealed no differences to wild-type tissue for any of the Fgf ligands and receptors analysed (Fig. 3, right panels). By contrast, significant changes in transcript levels were detected by comparing Fgf2 mutant (-/-) to wild-type (+/+) MPC cells (Fig. 3, left panels). In particular, Fgf1 transcript levels are upregulated in mutant MPC cells in comparison to wild-type cells (Fig. 3, upper panels). By contrast, the levels of both Fgf7 and Fgf10 transcripts are downregulated to low or undetectable levels in FGF2 mutant MPC cells (Fig. 3, upper panels), whereas Fgf8 expression remains normal (data not shown).
Fibroblast growth factor signals are transduced by high-affinity tyrosine kinase FGF receptors (FGFR) (Martin, 1998) and there is evidence that the IIIc isoforms of FGFR1 and FGFR2 have highest affinity for FGF2 (Ornitz et al., 1996). Although expression of the Fgfr1 isoform IIIc is not affected in mutant MPC cells, expression of the Fgfr2 isoform IIIc is downregulated compared with wild-type MPC cells (Fig. 3, lower panels). By contrast, the Fgfr1 and Fgfr2 IIIb isoforms are both upregulated in mutant MPC cells (Fig. 3, lower panels). In summary, the results in Fig. 3 show that expression of at least three Fgf ligands, which are normally upregulated upon MPC cell differentiation, are low or undetectable in mutant MPC cells. FGF signal reception is also altered in mutant cells, owing to changes in receptor isoforms. In agreement with this general alteration of FGF signalling, transient re-expression of FGF2 in FGF2 mutant MPC cells by DNA microinjection did not rescue the mutant phenotype (data not shown). It seems thus appropriate to refer to the phenotypes observed in mutant MPC cells (Figs 2; Fig. 4; Fig. 5) as defects caused by disrupting FGF rather than only FGF2 signalling.
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Synaptopodin is a protein associated with actin filaments in differentiated podocytes and neurons (Mundel et al., 1997a). Both in vivo and in vitro, the expression of this differentiation marker is activated during actin-based foot process formation (Mundel et al., 1997a; Mundel et al., 1997b) (Fig. 4C). In accordance with the morphological alterations seen in mutant MPC cells (Fig. 2D) (Fig. 4B), synaptopodin expression is much lower in mutant MPC cells compared with wild-type cells after induction of differentiation (Fig. 4D,E). Interestingly, the podocyte lineage marker WT-1 (Wilms Tumour antigen) (Mundlos et al., 1993) is also markedly downregulated in proliferating and post-mitotic mutant MPC cells (Fig. 4E). Instead, podocalyxin and Pod-1, two additional podocyte lineage markers (Kerjaschki et al., 1984; Quaggin et al., 1999), remain expressed at levels similar to those seen in wild-type cells (Fig. 4E). Thus, although many properties of differentiated podocytes are disrupted, mutant MPC cells mostly maintain podocyte lineage characteristics.
Block of mesenchymal differentiation in mutant MPC cells
The results shown in Figs 2, Fig. 4 reveal a block in postmitotic differentiation of mutant MPC cells. To identify the underlying molecular defects causing this phenotype, we analysed genes implicated in the EMT-like transition that podocytes undergo during onset of differentiation. The intermediate filament protein vimentin is a marker of mesenchymal cells (Schmid et al., 1979) and is expressed by mature podocytes (Fig. 5A) (Holthofer et al., 1984; Yaoita et al., 1999). In agreement, vimentin expression is upregulated in differentiating wild-type MPC cells (Fig. 5A,E), whereas it remains at low levels in mutant MPC cells (Fig. 5B,E). This defect is paralleled by a marked upregulation of the two epithelial markers cytokeratin and desmocollin type-2 (Dsc2) (Franke et al., 1979; Koch et al., 1992) in mutant MPC cells (Fig. 5E; and data not shown). Furthermore, the tight junction protein ZO-1 (Stevenson et al., 1986) relocalises from an apical position to the slit-diaphragm forming area of foot processes during podocyte differentiation (Schnabel et al., 1990). Differentiated wild-type MPC cells express predominantly one of the two ZO-1 isoforms (Fig. 5E), which localises to discrete areas of the cellular processes (Fig. 5C, arrow). Instead, mutant MPC cells express both ZO-1 isoforms (Fig. 5E), which are uniformly distributed at cell-cell interfaces (Fig. 5D). Again, this ZO-1 expression profile is reminiscent of epithelial cells (Balda and Anderson, 1993). The zinc finger transcription factor slug has been implicated in FGF-induced EMT switches (Savagner et al., 1997) and marks mesenchymal cell state (Nieto et al., 1994). As expected, slug expression is upregulated in differentiated wild-type MPC cells, whereas it remains undetectable in mutant MPC cells (Fig. 5E). Taken together, these results indicate that the molecular pathway(s) leading to induction of mesenchymal differentiation are blocked in MPC cells with disrupted FGF signalling.
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DISCUSSION |
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FGF signalling induces EMT-like changes
Despite the fact that terminally differentiated podocytes retain a basolateral asymmetry, the morphological and molecular changes that take place during onset of differentiation, both in vivo and in vitro, are similar to an EMT (Holthofer et al., 1984; Mundel et al., 1997b; Schnabel et al., 1990). In particular, differentiating podocytes loose epithelial characteristics such as desmosomal adhesion plaques and cytokeratins, whereas they acquire mesenchymal features such as vimentin expression (Schmid et al., 1979) (this study). The inability of mutant MPC cells to develop such mesenchymal characteristics indicates that FGF signalling is required to induce EMT-like changes. In support of this, epiblast cells require FGF signalling to undergo the EMT during gastrulation (Ciruna et al., 1997) and epithelial cells in culture acquire mesenchymal properties upon FGF overexpression (Migdal et al., 1995). A downstream target of FGF-mediated effects on EMT is the transcription factor slug (Buxton et al., 1997; Savagner et al., 1997), whose expression is disrupted in mutant MPC cells. In fact, overexpression of slug has been shown to trigger an EMT and results in downregulation of desmosomal cadherins (Savagner et al., 1997). In agreement, the disruption of FGF signalling and failure to activate slug in mutant MPC cells is accompanied by a very significant upregulation of the desmosomal cadherin Dsc2. These studies show that FGF signalling acts upstream of slug activation and Dsc2 repression during initiation of an EMT or establishment of mesenchymal characteristics.
FGF signalling and formation of actin-based cellular processes
During terminal differentiation, podocytes extend an ordered array of actin-based foot processes, which entwine glomerular capillaries and form an essential component of the filtration unit (Saxén, 1987). The inability of mutant MPC cells to initiate these EMT-like changes leads to a block in development of actin-base cellular processes. Interestingly, these defects are reminiscent of alterations occurring during onset of glomerular disease in vivo. Damaged podocytes retract their foot processes, detach from the glomerular basement membrane and frequently revert to a more epithelial morphology (Bariety et al., 1998; Whiteside et al., 1993). These changes in podocyte morphology are often accompanied by a significant expansion of their cytoplasm (Autio-Harmainen et al., 1981), a phenomenon seen often in mutant MPC cells. Together with the present study, these results indicate that FGF signalling might regulate differentiation or maintenance the intricate array of podocyte foot processes in interaction with neighbouring podocytes and the glomerular basement membrane (Saxén, 1987).
Finally, podocytes seem to share some similarity with neurons as both cell types extend complex actin-based processes during terminal differentiation (Reeves et al., 1978; Zigmond et al., 1999). Axonal path finding by neurons is controlled by (de)polymerisation of actin-based filopodia emerging from the growth cone located at the distal tip of the growing axon (Zigmond et al., 1999). Disrupting FGF signalling in cultured cerebellar neurons prevents axonal outgrowth (Saffell et al., 1997), whereas FGF stimulation of cultured retinal neurons promotes axonal outgrowth and target recognition (McFarlane et al., 1995). FGF signalling has also been shown to regulate actin filament reorganisation of endothelial cells during wound healing (Wang and Gotlieb, 1999), a process which is delayed in Fgf2 deficient mice (Ortega et al., 1998). Furthermore, it has recently been shown that embryonic cells extend cytonemes, which are long, actin-based cellular processes growing towards signalling centers (Ramirez-Weber and Kornberg, 1999). Interestingly, FGF signalling stimulates both outgrowth and orientation of cytonemes similar to FGF functions during axonal path finding (see above). In summary, FGF signalling seems to regulate aspects of growth and/or maintenance of actin-based processes during differentiation of various cell-types.
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ACKNOWLEDGMENTS |
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REFERENCES |
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Autio-Harmainen, H., Vaananen, R. and Rapola, J. (1981). Scanning electron microscopic study of normal human glomerulogenesis and of fetal glomeruli in congenital nephrotic syndrome of the Finnish type. Kidney Int. 20, 747-752.[Medline]
Balda, M. S. and Anderson, J. M. (1993). Two classes of tight junctions are revealed by ZO-1 isoforms. Am. J. Physiol. 264, C918-924.
Bariety, J., Nochy, D., Mandet, C., Jacquot, C., Glotz, D. and Meyrier, A. (1998). Podocytes undergo phenotypic changes and express macrophagic-associated markers in idiopathic collapsing glomerulopathy. Kidney Int. 53, 918-925.[Medline]
Blasco, M. A., Lee, H. W., Hande, M. P., Samper, E., Lansdorp, P. M., DePinho, R. A. and Greider, C. W. (1997). Telomere shortening and tumor formation by mouse cells lacking telomerase RNA. Cell 91, 25-34.[Medline]
Buxton, P. G., Kostakopoulou, K., Brickell, P., Thorogood, P. and Ferretti, P. (1997). Expression of the transcription factor slug correlates with growth of the limb bud and is regulated by FGF-4 and retinoic acid. Int. J. Dev. Biol. 41, 559-568.[Medline]
Cancilla, B., Ford-Perriss, M. D. and Bertram, J. F. (1999). Expression and localization of fibroblast growth factors and fibroblast growth factor receptors in the developing rat kidney. Kidney Int. 56, 2025-2039.[Medline]
Cauchi, J., Alcorn, D., Cancilla, B., Key, B., Berka, J. L., Nurcombe, V., Ryan, G. B. and Bertram, J. F. (1996). Light-microscopic immunolocalization of fibroblast growth factor-1 and -2 in adult rat kidney. Cell Tissue Res. 285, 179-187.[Medline]
Celli, G., LaRochelle, W. J., Mackem, S., Sharp, R. and Merlino, G. (1998). Soluble dominant-negative receptor uncovers essential roles for fibroblast growth factors in multi-organ induction and patterning. EMBO J. 17, 1642-1655.
Ciruna, B. G., Schwartz, L., Harpal, K., Yamaguchi, T. P. and Rossant, J. (1997). Chimeric analysis of fibroblast growth factor receptor-1 (Fgfr1) function: a role for FGFR1 in morphogenetic movement through the primitive streak. Development 124, 2829-2841.
Deng, C. X., Wynshaw-Boris, A., Shen, M. M., Daugherty, C., Ornitz, D. M. and Leder, P. (1994). Murine FGFR-1 is required for early postimplantation growth and axial organization. Genes Dev. 8, 3045-3057.[Abstract]
Dono, R. and Zeller, R. (1994). Cell-type-specific nuclear translocation of fibroblast growth factor-2 isoforms during chicken kidney and limb morphogenesis. Dev. Biol. 163, 316-330.[Medline]
Dono, R., Texido, G., Dussel, R., Ehmke, H. and Zeller, R. (1998). Impaired cerebral cortex development and blood pressure regulation in FGF-2-deficient mice. EMBO J. 17, 4213-4225.
Eckes, B., Dogic, D., Colucci-Guyon, E., Wang, N., Maniotis, A., Ingber, D., Merckling, A., Langa, F., Aumailley, M., Delouvee, A. et al. (1988). Impaired mechanical stability, migration and contractile capacity in vimentin-deficient fibroblasts. J. Cell Sci. 111, 1897-1907.
Ford, M. D., Cauchi, J., Greferath, U. and Bertram, J. F. (1997). Expression of fibroblast growth factors and their receptors in rat glomeruli. Kidney Int. 51, 1729-1738.[Medline]
Franke, W. W., Appelhans, B., Schmid, E., Freudenstein, C., Osborn, M. and Weber, K. (1979). Identification and characterization of epithelial cells in mammalian tissues by immunofluorescence microscopy using antibodies to prekeratin. Differentiation 15, 7-25.[Medline]
Garrod, D. R. and Fleming, S. (1990). Early expression of desmosomal components during kidney tubule morphogenesis in human and murine embryos. Development 108, 313-321.[Abstract]
Hogan, B. L. (1999). Morphogenesis. Cell 96, 225-233.[Medline]
Holthofer, H., Miettinen, A., Lehto, V. P., Lehtonen, E. and Virtanen, I. (1984). Expression of vimentin and cytokeratin types of intermediate filament proteins in developing and adult human kidneys. Lab Invest. 50, 552-559.[Medline]
Jat, P. S., Noble, M. D., Ataliotis, P., Tanaka, Y., Yannoutsos, N., Larsen, L. and Kioussis, D. (1991). Direct derivation of conditionally immortal cell lines from an H-2Kb-tsA58 transgenic mouse. Proc. Natl. Acad. Sci. USA 88, 5096-5100.[Abstract]
Kerjaschki, D., Sharkey, D. J. and Farquhar, M. G. (1984). Identification and characterization of podocalyxin the major sialoprotein of the renal glomerular epithelial cell. J. Cell Biol. 98, 1591-1596.[Abstract]
Koch, P. J., Goldschmidt, M. D., Zimbelmann, R., Troyanovsky, R. and Franke, W. W. (1992). Complexity and expression patterns of the desmosomal cadherins. Proc. Natl. Acad. Sci. USA 89, 353-357.[Abstract]
Majumdar, A. and Drummond, I. A. (1999). Podocyte differentiation in the absence of endothelial cells as revealed in the zebrafish avascular mutant, cloche. Dev. Genet. 24, 220-229.[Medline]
Martin, G. R. (1998). The roles of FGFs in the early development of vertebrate limbs. Genes Dev. 12, 1571-1586.
Mason, I. J., Fuller-Pace, F., Smith, R. and Dickson, C. (1994). FGF-7 (keratinocyte growth factor) expression during mouse development suggests roles in myogenesis, forebrain regionalisation and epithelial-mesenchymal interactions. Mech. Dev. 45, 15-30.[Medline]
McFarlane, S., McNeill, L. and Holt, C. E. (1995). FGF signaling and target recognition in the developing Xenopus visual system. Neuron 15, 1017-1028.[Medline]
Migdal, M., Soker, S., Yarden, Y. and Neufeld, G. (1995). Partial EMT/actin reorganisation by FGF signaling in MDCK cells. J. Cell Physiol. 162, 266-276.[Medline]
Mundel, P., Heid, H. W., Mundel, T. M., Kruger, M., Reiser, J. and Kriz, W. (1997a). Synaptopodin: an actin-associated protein in telencephalic dendrites and renal podocytes. J. Cell Biol. 139, 193-204.
Mundel, P., Reiser, J., Zuniga Mejia Borja, A., Pavenstadt, H., Davidson, G. R., Kriz, W. and Zeller, R. (1997b). Rearrangements of the cytoskeleton and cell contacts induce process formation during differentiation of conditionally immortalized mouse podocyte cell lines. Exp. Cell Res. 236, 248-258.[Medline]
Mundlos, S., Pelletier, J., Darveau, A., Bachmann, M., Winterpacht, A. and Zabel, B. (1993). Nuclear localization of the protein encoded by the Wilms tumor gene WT1 in embryonic and adult tissues. Development 119, 1329-1341.
Nagata, M. and Watanabe, T. (1997). Podocytes in metanephric organ culture express characteristic in vivo phenotypes. Histochem. Cell Biol. 108, 17-25.[Medline]
Nagata, M., Yamaguchi, Y. and Ito, K. (1993). Loss of mitotic activity and the expression of vimentin in glomerular epithelial cells of developing human kidneys. Anat. Embryol. (Berl.) 187, 275-279.[Medline]
Nieto, M. A., Sargent, M. G., Wilkinson, D. G. and Cooke, J. (1994). Control of cell behavior during vertebrate development by Slug, a zinc finger gene. Science 264, 835-839.[Medline]
Ohuchi, H., Hori, Y., Yamasaki, M., Harada, H., Sekine, K., Kato, S. and Itoh, N. (2000). FGF10 acts as a major ligand for FGF receptor 2 IIIb in mouse multi-organ development. Biochem. Biophys. Res. Commun. 277, 643-649.[Medline]
Ornitz, D. M., Xu, J., Colvin, J. S., McEwen, D. G., MacArthur, C. A., Coulier, F., Gao, G. and Goldfarb, M. (1996). Receptor specificity of the fibroblast growth factor family. J. Biol. Chem. 271, 15292-15297.
Ortega, S., Ittmann, M., Tsang, S. H., Ehrlich, M. and Basilico, C. (1998). Neuronal defects and delayed wound healing in mice lacking fibroblast growth factor 2. Proc. Natl. Acad. Sci. USA 95, 5672-5677.
Peters, K. G., Werner, S., Chen, G. and Williams, L. T. (1992). Two FGF receptors gene are differentially expressed in epithelial and mesenchymal tissues during limb formation and organogenesis in the mouse. Development 114, 233-243.[Abstract]
Qiao, J., Uzzo, R., Obara-Ishihara, T., Degenstein, L., Fuchs, E. and Herzlinger, D. (1999). FGF-7 modulates ureteric bud growth and nephron number in the developing kidney. Development 126, 547-554.
Quaggin, S. E., Schwartz, L., Cui, S., Igarashi, P., Deimling, J., Post, M. and Rossant, J. (1999). The basic-helix-loop-helix protein pod1 is critically important for kidney and lung organogenesis. Development 126, 5771-5783.
Ramirez-Weber, F. A. and Kornberg, T. B. (1999). Cytonemes: cellular processes that project to the principal signaling center in Drosophila imaginal discs. Cell 97, 599-607.[Medline]
Reeves, W., Caulfield, J. P. and Farquhar, M. G. (1978). Differentiation of epithelial foot processes and filtration slits: sequential appearance of occluding junctions, epithelial polyanion, and slit membranes in developing glomeruli. Lab Invest. 39, 90-100.[Medline]
Saffell, J. L., Williams, E. J., Mason, I. J., Walsh, F. S. and Doherty, P. (1997). Expression of a dominant negative FGF receptor inhibits axonal growth and FGF receptor phosphorylation stimulated by CAMs. Neuron 18, 231-242.[Medline]
Savagner, P., Yamada, K. M. and Thiery, J. P. (1997). The zinc-finger protein slug causes desmosome dissociation, an initial and necessary step for growth factor-induced epithelial-mesenchymal transition. J. Cell Biol. 137, 1403-1419.
Saxén, L. (1987). Organogenesis of the Kidney. In Developmental and Cell Biology Series (ed. P. W. Barlow, P. B. Green and C. C. Wylie), Cambridge: Cambridge University Press.
Schmid, E., Tapscott, S., Bennett, G. S., Croop, J., Fellini, S. A., Holtzer, H. and Franke, W. (1979). Differential location of different types of intermediate-sized filaments in various tissues of the chicken embryo. Differentiation 15, 27-40.[Medline]
Schnabel, E., Anderson, J. M. and Farquhar, M. G. (1990). The tight junction protein ZO-1 is concentrated along slit diaphragms of the glomerular epithelium. J. Cell Biol. 111, 1255-1263.[Abstract]
Sorokin, L. and Ekblom, P. (1992). Development of tubular and glomerular cells of the kidney. Kidney Int. 41, 657-664.[Medline]
Stevenson, B. R., Siliciano, J. D., Mooseker, M. S. and Goodenough, D. A. (1986). Identification of ZO-1: a high molecular weight polypeptide associated with the tight junction (zonula occludens) in a variety of epithelia. J. Cell Biol. 103, 755-766.[Abstract]
Szebenyi, G. and Fallon, J. F. (1999). Fibroblast growth factors as multifunctional signaling factors. Int. Rev. Cytol. 185, 45-106.[Medline]
Tassin, M. T., Beziau, A., Gubler, M. C. and Boyer, B. (1994). Spatiotemporal expression of molecules associated with junctional complexes during the in vivo maturation of renal podocytes. Int. J. Dev. Biol. 38, 45-54.[Medline]
Valles, A. M., Boyer, B., Badet, J., Tucker, G. C., Barritault, D. and Thiery, J. P. (1990). Acidic fibroblast growth factor is a modulator of epithelial plasticity in a rat bladder carcinoma cell line. Proc. Natl. Acad. Sci. USA 87, 1124-1128.[Abstract]
Wang, D. I. and Gotlieb, A. I. (1999). FGF2 induces actin filament reorganisation of endothelial cells in wound healing. Ex. Mol. Pathol. 66, 179-190.
Whiteside, C. I., Cameron, R., Munk, S. and Levy, J. (1993). Podocytic cytoskeletal disaggregation and basement-membrane detachment in puromycin aminonucleoside nephrosis. Am. J. Pathol. 142, 1641-1653.[Abstract]
Yamaguchi, T. P., Harpal, K., Henkemeyer, M. and Rossant, J. (1994). fgfr-1 is required for embryonic growth and mesodermal patterning during mouse gastrulation. Genes Dev. 8, 3032-3044.[Abstract]
Yaoita, E., Franke, W. W., Yamamoto, T., Kawasaki, K. and Kihara, I. (1999). Identification of renal podocytes in multiple species: higher vertebrates are vimentin positive/lower vertebrates are desmin positive. Histochem. Cell Biol. 111, 107-115.[Medline]
Zigmond, M. J., Bloom, F. E., Landis, S. C., Roberts, J. L. and Squire, L. R. (1999). In Fundamental Neuroscience (ed. M. J. Zigmond, F. E. Bloom, S. C. Landis, J. L. Roberts and L. R. Squire), pp. 519-527. San Diego, CA: Academic Press.