s2fieldsofstudy:
Biology
Environmental Science

text:
Teratological effect of 2,3,7,8-tetrachlorodibenzo- p -dioxin (TCDD): induction of cleft palate in the ddY and C57BL/6 mouse

2,3,7,8-Tetrachlorodibenzo- p -dioxin (TCDD), a highly toxic halogenated aromatic hydrocarbon, is a teratogen to induce cleft palate when exposed during the pregnancy. There are inter-strain differences in the sensitivity to cleft palate induced by TCDD and other chemicals including polychlorinated terphenyls (PCTs). The C57BL/6 mouse and the ddY mouse had been shown to be different in the induction of cleft palate following the treatment of PCTs, which attempts us to evaluate the TCDD-induced cleft palate in two mouse strains to understand the mechanism through which TCDD and PCTs induce cleft palate. This study evaluated the induction of cleft palate in the fetuses of ddY and C57BL/6 mice after subcutaneous treatment of TCDD on gestation day (GD) 10.5-14.5 or oral treatment on GD 8.5-13.5. Our results clearly showed that ddY mice, a susceptible strain to PCTs-induced cleft palate, are resistant to the induction of cleft palate by TCDD comparably to the high susceptibility of C57BL/6 mice, suggesting a different teratological mechanism between TCDD and PCTs. In addition, at the low doses, our study supported the concept of “window effect” of TCDD on around GD 12 for the induction of cleft palate in C57BL/6 and ddY mice. 2,3,7,8-tetrachlorod-ibenzo- p -dioxin

Introduction
2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD), a member of halogenated aromatic hydrocarbons, is a widely spread environmental contaminant [26]. TCDD has a variety of adverse biological effects including carcinogenesis, immune and hemopoietic dysfunction, neuronal cell damage, teratogenesis and reproductive toxicity [13,18,19,24]. The induction of cleft palate is known to be a sensitive teratological effect of TCDD when animals are exposed to TCDD during the pregnancy [10,21].
Many mouse strains have been used for toxicological and pharmacological studies. Sometimes, the use of mouse strains with different characteristics provides an important clue to approach the toxic and pharmacological mechanism of chemicals. In association with TCDD, DBA/2, a mouse strain with a mutation on the AhR locus of DNA, has been used to investigate the toxic mechanism of TCDD [8,19,22,23]. Compared with the TCDDsensitive C57BL/6 mice, the resistance of DBA/2 mice to TCDD-induced toxicity had suggested that AhR is involved in the toxic mechanism of TCDD. Later, it had been proved by AhR knock-out mice that the toxic effects of polyhalogenated aromatic compounds including TCDD are mediated by the AhR [20].
The polychlorinated terphenyls (PCTs) having a similar chemical structure to polychlorinated biphenyl (PCB) that is a member of polyhalogenated aromatic compounds has also been shown to induce cleft palate [17]. However, the mechanism by which PCTs induce cleft palate is still speculative. Kaneko and his college used C57BL/6 and ddY mice to investigate the teratological effect of PCTs and its mechanism [17]. It is interesting in their report that C57BL/7 mouse strain sensitive to TCDD-induced cleft palate was resistant to PCTs-induced cleft palate. On the other hand, ddY mice showed high incidence of cleft palate following PCTs treatment. On the basis of the previous study, we hypothesized that those two mouse strains would show different susceptibility to TCDDinduced cleft palate, of which the confirmation would be helpful to extend our understanding in the teratological mechanisms of TCDD and PCTs.
For that purpose, in the present study, we evaluated the induction of cleft palate in ddY and C57BL/6 mice after subcutaneous or oral treatment of TCDD during the pregnancy and compared. Our results clearly showed that, unlike the cleft palate induced by PCTs treatment, the ddY mouse was resistant to TCDD-induced cleft palate comparably to the high susceptibility of C57BL/6 mice, which strongly suggested that TCDD and PCTs give rise to their teratological effect by different mechanisms.

Materials and Methods
Chemicals 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) was purchased from Radian International, Cambridge Isotope Laboratories, Inc., Andover, MA, USA, and its purity was 98 %. TCDD was initially dissolved in a small volume of acetone and subsequently adjusted to a working concentration in olive oil.

Animals
Female and male C57BL/6 and ddY mice were obtained from Japan SLC Inc. (Hamamatsu, Japan) at 6-8 weeks of age and held for 2 weeks prior to mating. Two females were housed overnight with one male and checked the presence of a vaginal plug in the next morning, denoted as gestation day 0.5 (GD 0.5). The plug-positive females were maintained in a vinyl isolator established in the hazard room to prevent an environmental exposure. The room was kept under the conditions of 22 ± 1 o C in temperature, 50 ± 10% in humidity and 12/12 light/dark cycle. During the study, the mice were given food (CRF-1, Oriental Yeast Co. LTD) and water ad libitum.

Treatment and experimental design
For this study, two different administration routes were chosen, subcutaneous (SC) and oral (PO). The doses were selected on the basis of the results of previous studies [7] and our preliminary studies. C57BL/6 and ddY mice were respectively given a single dose of 0, 20, 40 and 80 µg TCDD/kg bw in 10 ml olive oil/kg bw by subcutaneous injection on GD10.5, 11.5, 12.5, 13.5 and 14.5. For the oral study, a single dose of 0, 10, 20 and 40 µg TCDD/kg bw for C57BL/6 mice and 0, 20, 40 and 80 µg TCDD/kg bw for ddY mice was given by gavage on GD8.5, 9.5, 10.5, 11.5, 12.5 and 13.5, respectively. Five pregnant mice per group were used, but the number was sometimes decreased because of non-pregnancy. On GD18, the dams were killed by decapitation. The number and position of all fetuses, live and dead, and of resorptions were noted. Live fetuses were grossly examined to evaluate the incidence of cleft palate, and then fixed in 10% neutral buffered formalin. For histological examination, the sections of craniofacial tissues were processed, embedded in paraffin and stained with hematoxylin and eosin (H&E).

Data analysis
The litter was considered the basic experimental unit. The Kruskal-Wallis test was used to assess the analysis of variance. The significance of the dose-response trend was determined using Jonckheere's test against ordered alternatives, and when this test indicated a significant trend, pairwise comparisons were made using the Mann-Whitney U test [14]. The magnitude of the right-left severity score difference for cleft palate was assessed using the Wilcoxon matched-pairs signed-ranks test [6].

Results
Fetal mortality and indcidence of cleft palate C57BL/6 mice ( Table 1, 3) : Four doses of TCDD (0, 20, 40 and 80 µg/kg bw) were singly injected subcutaneously on GD10.5, 11.5, 12.5, 13.5 and 14.5. No effects of TCDD at these concentrations, when injected subcutaneously, were seen on fetal mortality irrespective of the gestation days injected. The oral treatment of 20 µg TCDD/kg bw did not give any effect on the fetal mortality. However, when 40 µg/kg bw of TCDD was orally administered on GD8.5, the percentage of fetuses dying at the late stage was significantly high (31%). In C57BL/6 mice, TCDD clearly induced cleft palate, which was depending on the concentration and the gestation day when TCDD was injected. When 20 µg/kg bw of TCDD was subcutaneously injected, the incidence of cleft palate was observed in the fetuses exposed to TCDD only on GD 12.5 and 14.5 although its rate was very low. The incidence of cleft palate, when 40 µg/kg bw of TCDD was subcutaneously injected, was significantly high in the fetuses exposed to TCDD only on GD 12.5, indicating the "window effect" of TCDD on the induction of cleft palate. However, the subcutaneous treatment of 80 µg/kg bw of TCDD highly induced cleft palate at all TCDD-injected GD points except for GD 14.5.
When TCDD was administered orally, 20 µg/kg bw of TCDD failed to give an effect on fetal mortality in C57BL/ 6 mice. However, an increase in the number of fetuses dying at the late stage was noted when 40 µg/kg bw of TCDD was administered on GD 11.5. The teratological effect of TCDD was clear in the incidence of cleft palate when 10 µg/kg bw of TCDD was orally given on GD 11.5 and 12.5; 37.5 and 27.8%, respectively. The increase of dose to 20 µg/kg bw not only highly increased the incidence of cleft palate on GD11.5 and 12.5, but also induced cleft palate even on GD8.5, 9.5, 10.5 and 13.5. The oral treatment of 20 µg TCDD/kg bw on GD11.5 and 12.5 induced cleft palate in most of the fetuses (>94 %), and the incidence rates of cleft palate were respectively 20, 26.7, 69.1 and 35.3% when treated on GD8.5, 9.5, 10.5, and 13.5. The oral treatment of 40 µg TCDD/kg bw on GD8.5 -GD12.5 was enough to induce cleft palate in all of the fetuses, and on GD13.5 half of the fetuses were affected. ddY mice (Table 2, 4) : When TCDD was treated subcutaneously, there were no statistically significant effects of TCDD at the concentrations of 20, 40 and 80 µg /kg bw on fetal mortality irrespective of the gestation days injected. However, when 80 µg/kg bw of TCDD was orally administered on GD 10.5, 13 fetuses from two dams died at the late stage of gestation; 9/12 and 4/15,  respectively. Twelve fetuses from three dams treated with 80 µg/kg bw of TCDD on GD 13.5 died at the early stage of gestation (7/11, 2/11, and 3/14). Compared with C57BL/6 mice, ddY mice were very resistant to the teratological effect of TCDD in the induction of cleft palate. When TCDD was injected subcutaneously, cleft palate didn`t occur even at the concentration of 80 µg/kg bw. Only one fetus that 40 µg/ kg bw of TCDD was subcutaneously injected on GD 13.5 had cleft palate. The ddY mouse also showed a prominent resistance to the induction of cleft palate following the oral treatment of TCDD. In our study, while less than 10 µg/kg bw of TCDD clearly induced cleft palate in C57BL/6 mice, 20 µg/kg bw of TCDD was necessitated to induce cleft palate in ddY mice. The fetuses of ddY mice were affected when 20 and 40 µg/kg bw of TCDD were administered on GD 12.5, indicating a "window effect" of TCDD on the induction of cleft palate; the incidence rate were 9.52% and 4.48%, respectively. At the concentration of 80 µg/kg bw TCDD, the cleft palate was induced in the fetuses administered on GD10.5, 11.5 and 12.5, of which the incidence rates were 6.9, 10 and 18.6%, respectively. In the ddY mouse, GD12 was the most sensitive gestation day for the induction of cleft palate when TCDD was administered per oral.

Gross and histological morphology
The cleft palates induced in the fetuses of C57BL/6 and ddY mice treated with by TCDD were typical in their morphology, having normal sized palatal shelves in a vertical position (Figure 1). Two palatal shelves failed to meet and fuse each other, resulting in a wide gap between them (Figure 1, 2). Histologically, the cleft was lined by nasal epithelial cells, medial epithelial cells of two opposing prominences, and then connected with squamous epithelial cells of oral cavity (Figure 2).

Discussion
It has been well documented that the induction of cleft palate is a toxic effect of TCDD on fetal development [1-5, 7-10, 21, 24, 25, 28]. The normal development of palate is completed by a growth of opposing palatal shelves and their fusion through the programmed cell death of medial edge epithelial cells [12]. Therefore, cleft palate can be induced by inhibiting the growth of medial epithelial cells or by interfering with a fusion between two palatal shelves. The cleft palate induced by TCDD is considered to result from the poor development of palatal shelves [28] or an altered differentiation of medial cells to interfere with the programmed cell death [2,4,25].
Our study confirmed that TCDD is a teratogen to induce cleft palate and has a "window effect" at low dosages for the induction of cleft palate. Morphologically, the cleft palates induced by TCDD in C57BL/6 and ddY mice were typically composed of normal sized palatal shelves in a vertical position, resulting from the failure of fusion between two opposing palatal shelves (Figure 1, 2). The incidence was the most sensitive when TCDD was treated around GD12 in both C57BL/6 and ddY mice. In C57BL/6 mice, the cleft palate was, at the concentration of 40 µg/kg bw, clearly induced when TCDD was subcutaneously treated only on GD 12.5; the incidence was 25 % (Table 1). When TCDD was orally administered, the incidence of cleft palate was also limited on GD10.5 ñ GD12.5 at the concentration of 10 µg/kg bw, indicating that the incidence of cleft palate is the most sensitive when TCDD is treated around GD12 (Table 3). The dose-increase to 20 µg/kg bw   induced cleft palate on a wide range of gestation day (GD8.5−GD13.4), but the incidence was significantly high on GD11.5 and GD12.5 ( Table 3). The "window effect" of TCDD for the induction of cleft palate was also observed in ddY mice at the concentration of 20 and 40 µg/kg bw as cleft palate was clearly induced when TCDD was orally administered only on GD 12.5 ( Table 4). The incidence of cleft palate in TCDD-exposed embryos of C57BL/6 mice was in close agreement with that of the previous studies [7,9]. Table 3. Fetal mortality and incidence of cleft palate in C57BL/6 mice following oral treatment of TCDD during pregnancy Group (µg/kg) GD (day) GD 8.5 GD 9.5 GD 10.5 GD 11.5 GD 12.5 GD 13.  In our study, it was found that ddY mice were very resistant to the fetal mortality and the induction of cleft palate following TCDD treatment. In fetal mortality, when TCDD was orally administered, the effects of TCDD appeared at 40 µg/kg bw in C57BL/7 mice (Table 3), while at 80 µg/kg bw of TCDD in ddY mice (Table 4). In the induction of cleft palate, when TCDD is injected subcutaneously on GD12.5, 80 µg/kg bw of TCDD failed to induce cleft palate in ddY mice (Table 2), which was comparable to 27.3 % incidence of C57BL/6 mice ( Table  1). The resistance of ddY mice to the induction of cleft palate was also found when TCDD was orally administered. The oral treatment of 40 µg/kg bw of TCDD (a dose enough to affect all of fetuses in C57BL/6 mouse) to ddY mice on GD12.5 respectively induced cleft palate in only 18.6 % of fetuses ( Table 4). The strain difference in our study might be due to a difference in the expression of AhR in the craniofacial tissue between the two mouse strains, since AhR mediates the induction of cleft palate by TCDD and its level may determine the sensitivity of animals. C57BL/6 mice highly sensitive to TCDD-induced cleft palate have been known to have high-affinity AhR in craniofacial tissues, while DBA/2J mice, TCDD nonresponsive mice, have low-affinity AhR [23,27]. AKR/J mice are also known to be a relatively insensitive to the induction of cleft palate by TCDD, which is also assumed to be due to the low-affinity AhR of the strain [25]. Therefore, in our study, the low sensitivity of ddY mice to the induction of cleft palate by TCDD may be explained on the basis of the previous studies even if there is no report regarding to the expression of AhR in the craniofacial tissue of ddY mice. C57BL/6 and ddY mice were used to elucidate the mechanism through which PCTs induce cleft palate [17]. The previous study suggested that the cleft palate induced by PCTs be related with the up-regulation of corticosterone following PCTs treatment [17]. Nevertheless, the mechanism through which PCTs induce cleft palate is still unclear. In our study, the sensitivity of C57BL/6 and ddY mice to the TCDD-induced cleft palate was opposite to that of them to PCTs-induced cleft palate, indicating that the mechanism to induce cleft palate may be different between TCDD and PCTs. In addition, the increase of corticosterone level in plasma to have been observed after PCTs treatment in Kaneko`s study was not noted after TCDD treatment in our study (data not shown). It is also still unknown whether or not the toxicity of PCTs, like TCDD, is mediated by AhR.
Glucocorticoids (GC) are also teratogenic and induce cleft palate at pharmacological doses [11,15,16,24]. GC and TCDD are known to give rise to their effects through binding the respective receptors, GR and AhR [24]. It is still unclear whether there is any interaction between GR and AhR during the normal development of palate or in the incidence of cleft palate. However, Abbott et al.'s studies had shown there may be a cross-regulation of GR and AhR, since the synergistic interaction between TCDD and hydrocortisone for the induction of cleft palate was found [1,5]. According to their studies, TCDD treatment on GD14 induced up-regulation of GR and down-regulation of AhR, while the hydrocortisone exposure elevated the level of AhR and decreased the expression of GR. The treatment of both (TCDD + hydrocortisone) induced an increase of both receptors, followed by a synergistic increase of the incidence of cleft palate. The altered regulation of these receptors is followed by the altered expression of some growth factors [1,3], resulting in altered differentiation and proliferation of palatal epithelial cells. The mechanism of interaction cycle between GR and AhR is still speculative.
In summary, the present study showed that ddY mice, a susceptible strain to PCTs-induced cleft palate, were very resistant to the induction of cleft palate by TCDD, suggesting that the mechanisms through which TCDD and PCTs induce cleft palate may be different. In addition, we confirmed a "window effect" of TCDD for the induction of cleft palate in ddY mice.


extfieldsofstudy: Biology Medicine

text:
E-cadherin is a WT1 target gene.

The WT1 tumor suppressor gene encodes a transcription factor that can activate and repress gene expression. Transcriptional targets relevant for the growth suppression functions of WT1 are poorly understood. We found that mesenchymal NIH 3T3 fibroblasts stably expressing WT1 exhibit growth suppression and features of epithelial differentiation including up-regulation of E-cadherin mRNA. Acute expression of WT1 in NIH 3T3 fibroblasts after retroviral infection induced murine E-cadherin expression. In transient transfection experiments, the human and murine E-cadherin promoters were activated by co-expression of WT1. E-cadherin promoter activity was increased in cells overexpressing WT1 and was blocked by a dominant negative form of WT1. WT1 activated the murine E-cadherin promoter through a conserved GC-rich sequence similar to an EGR-1 binding site as well as through a CAAT box sequence. WT1 produced in vitro or derived from nuclear extracts bound to the WT1-response element within the murine E-cadherin promoter, but not the CAAT box. E-cadherin, a gene important in epithelial differentiation and neoplastic transformation, represents a downstream target gene that links the roles of the WT1 in differentiation and growth control.

The WT1 tumor suppressor gene is associated with Wilms' tumor, which can be considered a disease of disordered development, since the tumors exhibit the three elements of primordial kidney development including blastemal, stromal, and epithelial elements (1). The WT1 gene is predominantly expressed in the developing kidney, but WT1 expression is also detected in the fetal gonad, spleen, mesothelium, breast, and other tissues (reviewed in Ref. 2). During early nephrogenesis, WT1 expression is first detected in the metanephric mesenchyme. At the onset of epithelial differentiation, WT1 expression increases as the mesenchyme surrounding the ureteric bud condenses to form primary vesicles. These cells further differentiate into comma-and S-shaped bodies, and subsequently mature into glomeruli and proximal tubules. This phenomenon and the finding that mice null for WT1 fail to develop kidneys (3) suggest an essential role for WT1 in controlling the mesenchymal-epithelial transition of renal development. WT1 differs from other tumor suppressor genes such as Rb and p53 in that Rb and p53 are widely expressed in a variety of developing and adult tissues, while WT1 expression is restricted. The restricted pattern of WT1 expression suggests that its growthsuppressive properties may be linked to its role in organogenesis.
WT1 gene encodes a Cys-His zinc finger transcription factor, which functions as both a transcriptional repressor and activator (2, 4 -6). The DNA binding activity of WT1 was demonstrated on several different sequences including 5Ј-GCGGGG-GCG-3Ј, which is also recognized by EGR-1, EGR-2, and EGR-3 (4,(7)(8)(9)(10). Alternative splicing of the WT1 gene yields four major isoforms of the WT1 protein. Isoforms C and D include a 3-amino acid (KTS) segment between zinc fingers three and four of the protein, which alters the DNA binding specificity of the protein, while isoforms B and D include a 17-amino acid segment involved in transcriptional regulation. WT1(A) includes neither additional segment (2).
The tumor suppressor activity of WT1 was demonstrated by stable transfection of wild-type WT1 into a Wilms' tumor cell line harboring a mutant WT1 (11), as well as in heterologous osteosarcoma, CV1, and fibroblast cells (12)(13)(14)(15). WT1 can regulate expression of a number of growth and differentiationrelated genes in co-transfection assays including insulin-like growth factor II (16,17), insulin-like growth factor I receptor (18), platelet-derived growth factor A-chain (19,20), Pax-2 (21), syndecan-1 (22), Dax-1 (23), and amphiregulin (24) (reviewed in Refs. 2 and 25). Epidermal growth factor receptor downregulation by WT1 may contribute to growth suppression by inducing apoptosis (12), while syndecan activation (22) and amphiregulin (24) secretion may be important for renal differentiation. A problem in the interpretation of many of these studies has been the lack of a model for the action of WT1 in inducing both differentiation and growth control. We found that expression of WT1 in mesenchymal fibroblasts both suppresses cell growth (13) and induces features of epithelial differentiation (26). These WT1-expressing NIH 3T3 cells exhibit desmosome-like structures and increased expression of several epithelial marker genes such as E-cadherin, type IV collagen, and perlecan (heparin sulfate proteoglycan) along with reduced expression of ␣ 8 integrin and vimentin (26). Furthermore, we recently identified, by representational difference analysis, genes up-regulated in WT1-expressing cell lines which are expressed in the condensing metanephric mesenchyme of the developing mouse kidney. 1 This lends support to our hypothesis that expression of WT1 in this heterologous system mimics some of the effects of WT1 in kidney differentiation. Similarly, work by Haber's group identified amphiregulin as a gene induced by WT1 in an osteosarcoma cell line but also expressed in the condensing, developing human kidney (24). In this study we have focused on E-cadherin as a WT1 target gene, due to its close association with epithelial differentiation. E-cadherin is a calcium-dependent cell adhesion protein localized at the adherens junction that mediates cell-cell interactions. Expression of E-cadherin serves as a marker for epithelial differentiation (27,28). The intracellular portion of E-cadherin interacts with submembrane cytoskeletal proteins including the ␣and ␤-catenins (27)(28)(29)(30)(31). Loss of the E-cadherin expression is frequently seen in human tumors (28) and has been associated with loss of epithelial cell morphology and tumor invasion (32)(33)(34). The expression of E-cadherin and other cell adhesion proteins also play a key role in normal development during the mesenchymal-epithelial transition (35,36). E-cadherin appears in the induced metanephric mesenchyme once condensation into epithelia begins (37,38), the same time at which WT1 expression rises in the developing kidney (39,40).
In this report, we provide evidence that the WT1 protein can directly trans-activate E-cadherin gene expression. Both transient and stable expression of WT1 induced the endogenous E-cadherin gene. WT1 protein bound to a proximal GC-rich sequence in the E-cadherin gene promoter. This cis-acting element alone was sufficient for trans-activation of the E-cadherin promoter by WT1 in co-transfection experiments. A dominant negative mutant of WT1, WTAR inhibited the E-cadherin promoter in WT1 expressing NIH 3T3 cells. E-cadherin represents a downstream target gene that may link the functions of WT1 in epithelial differentiation and tumor suppression.

EXPERIMENTAL PROCEDURES
Cell Lines-The NIH 3T3 cells stably expressing WT1 (WR16 and WR35) and the Ras-transformed NIH 3T3 cells stably expressing WT1 (VW9) that were used in this study have been described previously (13,26). Ecotropic PHOENIX cells (gift of W. Pear, University of Pennsylvania, Philadelphia, PA) used to generate retroviral stocks were grown in Dulbecco's modified Eagle's medium with 10% fetal bovine serum.
Plasmid Construction-Expression vectors for wild-type WT1 isoforms (A, B, C, D) and the WT1 (AR) dominant negative mutant were described previously (13,(41)(42)(43). The promoter regions of the murine E-cadherin gene were amplified from murine genomic DNA by PCR 2 using various forward primers containing a BglII adaptor sequence to generate deletions or point mutations. Mutated bases are indicated in bold type. A reverse primer containing a HindIII adaptor sequence at ϩ113 of the murine E-cadherin promoter sequence was used (Sequences 1-17). The human E-cadherin promoter luciferase construct HECad (Ϫ368/ϩ125) was described previously (44). For the construction of HECad (Ϫ217/ϩ125) and HECad (DEL), E-cadherin promoter sequences were amplified by PCR and subcloned into SacI and HindIII of pGL2-Basic (Promega, Madison, WI). A bicistronic retroviral vector for WT1 was constructed by insertion of a Sau3A fragment of WT1(A) into the BglII site of the MIGR1 retroviral vector (gift of W. Pear), which contains the murine stem cell leukemia virus long terminal repeat.
A single reverse primer (R) containing a BglII adaptor was paired with each forward primer (F).
The polymerase chain reaction was performed at a denaturing temperature of 94°C for 30 s, annealing temperature of 60°C for 30 s, and extension for 1 min at 72°C for 40 cycles. After amplification, the PCR fragments were digested with BglII and HindIII and cloned into the pGL3-Basic luciferase reporter construct (Promega), and subjected to automated DNA sequencing.
Transfection Assay-For cotransfection assays involving the murine and human E-cadherin luciferase reporter and WT1 isoforms (A, B, C, D), NIH 3T3 cells and 293T cells were grown in Dulbecco's modified Eagle's medium containing 10% fetal calf serum and penicillin/streptomycin. At 18 h prior to transfection, NIH 3T3 cells were plated at a density of 3 ϫ 10 4 in 24-well dishes. Optimal quantities of effector (RSV-WT1(A, B, C, D), WTAR) and reporter (E-cadherin-Luc) used in transfection assays were empirically determined to be 0.5 g of effector (RSV-WT1(A, B, C, D) or RSV alone), 0.1 g of reporter, and 0.01 g of a thymidine kinase Renilla reporter gene, which serves as an internal control for transfection efficiency. WT1 modestly activated the thymidine kinase Renilla reporter, leading to an underestimate of activation by WT1 but did not alter the pattern of activation noted for E-cadherin promoter deletion constructs. The plasmids were transfected using the SuperFect transfection reagent (Qiagen, Valencia, CA) according to the manufacturer's directions. The NIH 3T3 cells were harvested 60 h after transfection and assayed for luciferase activity using the Dual-Reporter assay system (Promega). All results are shown as the average Ϯ standard deviation of three to six independent experiments. For preparation of nuclear extracts, 293T cells were plated at the density of 5 ϫ 10 6 cells/10-cm dish 18 h prior to transfection and 5 g of RSV-WT1(A) was used for transfection. The 293T cells were harvested 48 h after transfection, and nuclear extract was prepared as described below.
Generation of Retrovirus-A bicistronic vector harboring the green fluorescent protein and WT1(A) cDNAs was transiently transfected using Superfect (Qiagen) into PHOENIX cells. At 48 h after transfection, expression of the vector was monitored by phase contrast fluorescence microscopy (Nikon, Tokyo, Japan) and the supernatant of the packaging cells was collected, filtered, and layered in the presence of 5 g/ml Polybrene (Sigma) on a fresh culture of 60% confluent NIH 3T3 cells. At 24, 48, and 72 h after infection, the cells were monitored for green fluorescence and lysed for the preparation of total RNA using Trizol (Life Technologies, Inc.).
Analysis of mRNA Expression-Cytoplasmic RNAs were isolated 2 The abbreviations used are: PCR, polymerase chain reaction; GFP, green fluorescent protein; RSV, Rous sarcoma virus; GST, glutathione S-transferase; EMSA, electrophoretic mobility shift assay; DTT, dithiothreitol; PBS, phosphate-buffered saline. 5Ј-TAT AAG CTT CGC CGA GCA AAC ACT GAG CT-3Ј SEQUENCES 1-17 from cell lines as described previously (13,26). Cytoplasmic RNAs from each cell line (20 g) were subjected to Northern blot analysis for E-cadherin expression (45) and quantified by PhosphorImager analysis using ImageQuant software (Molecular Dynamics, Sunnyvale, CA). The E-cadherin probe was a gift from S. Mazur (Mount Sinai School of Medicine, New York, NY). E-cadherin expression in retrovirus-infected NIH3T3 cells was detected by reverse transcriptase-PCR. The reverse transcriptase reaction was performed with 1 g of total RNA using Superscript II, (Life Technologies, Inc.). The cDNA was then subjected to 35 rounds of PCR using E-cadherin-specific primers (E-cadherin forward PCR primer, 5Ј-TCA ACG ATC CTG ACC AGC AGT TCG-3Ј; reverse transcriptase primer, 5Ј-GGT GAA CCA TCA TCT GTG GCG ATG-3Ј; reverse nested PCR primer, 5Ј-ATC TTC TGA TCC ATG ACC GTG TCC G-3Ј). Polymerase chain reaction was performed at a denaturing temperature of 94°C for 30 s, annealing temperature of 60°C for 30 s, and extension for 1 min at 72°C for 35 cycles. An internal control for RNA integrity was simultaneously performed using mouse ␤-actinspecific primers (forward, GCC CAG AGC AAG AGA GGT AT; reverse, GGC CAT CTC TTG CTC GAA GT). The PCR products were visualized by agarose gel electrophoresis and ethidium bromide staining.
Electrophoretic Mobility Shift Assay (EMSA)-Double-stranded oligonucleotide probes were created by annealing complementary singlestranded oligonucleotides (Life Technologies, Inc.). The oligonucleotide sequences used were as follows. The putative WT1 binding site is underlined, and mutated position(s) are indicated in bold type.
The HindIII 5Ј overhanging ends of the duplex oligonucleotides were labeled with [␣-32 P]dCTP and the Klenow fragment of DNA polymerase I and gel-purified. In vitro coupled transcription/translations were performed with pSP64-WT1(A) (41) or an insertless pSP64 control vector using rabbit reticulocyte lysate (Promega). Nuclear extract from WT1(A)-expressing cell line VW9 (13), WT1(A)-transfected 293T, and a NIH 3T3 control cell line were extracted using a two-step method as follows. 5-10 ϫ 10 6 cells were grown to 80% confluence and resuspended in 10 mM HEPES (pH 7.9), 1.5 mM MgCl 2 , 10 mM KCl, 0.5 mM DTT. The cells were allowed to swell on ice for 10 min, vortexed, and briefly pelleted in a microcentrifuge at 4°C. The pellet was resuspended in 20 mM HEPES (pH 7.9), 25% glycerol, 420 mM NaCl, 1.5 mM MgCl 2 , 0.2 mM EDTA, and 0.5 mM DTT and incubated on ice for 20 min. The nuclear extract was then centrifuged for 2 min in a microcentrifuge at 4°C, and the supernatant was collected. Protein concentrations were determined using the Bio-Rad DC kit. To compare the binding activity of the alternative ϩKTS and ϪKTS forms of WT1 to the E-cadherin promoter WT1-response element, sequences encoding the zinc-finger domains of WT(A/ϪKTS), WT(C/ϩKTS), and WT(AR) (negative binding control) were cloned into the pGEX2TK GST-expression vector (Amersham Pharmacia Biotech), expressed in Escherichia coli strain BL21 and purified on glutathione beads (1).
DNA binding reactions using in vitro transcribed/translated WT1 were performed with 2.5 l of lysate programmed with pSP64-WT1(A) or the pSP64 control as described previously (46), except that the total volume of the reaction was 20 l and the amount of probe used was 0.2 ng (approximately 10 6 cpm/ng). Binding reactions utilizing nuclear extracts were performed by preincubating 3 g of extract in 20 mM HEPES (pH 7.5), 12% glycerol, 1 mg/ml bovine serum albumin, and 0.5 mM DTT with 2 g of d(I-C) and 1 g of DNase-free RNase A for 15 min at room temperature. Next, 0.2 ng (approximately 10 6 cpm/ng) of radiolabeled probe was added and the mixture was incubated for an additional 20 min at room temperature. Binding reactions utilizing GST-WT1 protein were performed by preincubating 200 ng of protein in 50 mM HEPES (pH 7.4), 50 mM KCl, 50 M ZnCl 2 , 5 mM MgCl 2 , 1 mM DTT, and 20% glycerol, with 1 g of d(I-C) for 10 min at room temperature. After this, 0.2 ng (approximately 10 6 cpm/ng) of radiolabeled probe was added and the mixture was incubated for 30 min at 4°C. The DNA-protein complexes were separated through a 5% non-denaturing polyacrylamide gel for 1.5 h at 300 V at room temperature. For Western blot analysis following the mobility shift assay, the assay was performed with an unlabeled oligonucleotide probe. The polyacrylamide gel was transferred to an Immobilon-P nylon membrane (Millipore, Bedford, MA), and immunoblotting analysis was performed as described below. The relative amount of DNA bound by WT1 protein was quantified utilizing a PhosphorImager and ImageQuant software (Molecular Dynamics, Sunnyvale, CA).
Immunoblotting-Cell lines were grown to 80% confluence in 10-cm plates, scraped into phosphate-buffered saline (PBS), and centrifuged at 1000 ϫ g. The cell pellets were resuspended in 200 l of PBS and solubilized in 200 l of 2ϫ Laemmli buffer (60 mM Tris, pH 6.8, 2% SDS, 10% glycerol) for 1 h at 4°C, The protein samples were then incubated at 100°C for 10 min and centrifuged in a microcentrifuge at 4°C for 15 min, and the resulting supernatant was collected. Protein concentrations were determined using the Bio-Rad DC kit. ␤-Mercaptoethanol was added to the samples to a final concentration of 0.5 M, and then heated at 85°C for 10 min prior to loading on a gel. Protein samples (40 g) were separated by electrophoresis through a 12% SDS-polyacrylamide gel and transferred to an Immobilon-P (Millipore, Bedford, MA) nylon membrane. The filters were blocked with 5% dry milk in PBS for 2 h, incubated with a rabbit polyclonal anti-WT1 antibody (C19, Santa Cruz Inc.) at 200 ng/ml, or with a rat monoclonal anti-mouse E-cadherin antibody (ECCD-2, Zymed Laboratories Inc., San Francisco, CA) at 1 g/ml in 0.5% dry milk Tris-buffered saline with 0.05% Tween 20 for 1 h, rinsed in Tris-buffered saline with Tween 20, and probed with a horseradish peroxidase conjugated anti-rabbit (Roche Molecular Biochemicals) (1:7500 dilution) or anti-rat (Santa Cruz Biotechnology) (1:1000 dilution) secondary antibody for 1 h. After rinsing in Trisbuffered saline with Tween 20, the immunoblots were developed by WT E-cad: Activates the E-cadherin Gene chemiluminescence (ECL, Amersham Pharmacia Biotech).

Induction of E-cadherin Expression in NIH 3T3 Cells by Stable Transfection and Retroviral Infection-In previous studies, we created a number of normal (WR16 and WR35) and
Ras-transformed (VW9) NIH 3T3 cell lines stably expressing WT1 isoform A (Ϫ17 amino acids, ϪKTS). In these cells, a number of epithelial markers were consistently up-regulated, including the E-cadherin gene ( Fig. 1A and Ref. 26). To examine if the endogenous E-cadherin gene is directly and immediately regulated by WT1(A), NIH 3T3 cells were infected with a bicistronic retrovirus carrying WT1(A) and GFP (Fig. 1B). Ecadherin mRNA was induced in NIH 3T3 cells as early as 24 h after infection with a retrovirus harboring WT1(A) and continued to increase at 48 and 72 h after infection. No induction of E-cadherin was noted in a control culture infected with a virus containing only GFP. These results suggested that E-cadherin induction was a direct result of WT1 expression rather than an effect of clonal selection.
Transcription of the Mouse and Human E-cadherin Promoter Is Activated by WT1-In order to determine if WT1 directly regulated the E-cadherin gene, we tested the ability of WT1 to activate both the mouse and human E-cadherin promoters. Previous studies indicated that the proximal promoter region of murine E-cadherin (Ϫ178 to ϩ113) is required for epithelial expression (47-49) (Fig. 2). Within the proximal promoter there are three regions that contribute to epithelial expression ( Fig. 2A) (50,51): 1) a palindromic element (Pal/Ap2) located at (Ϫ98 to Ϫ78), which stimulates transcription in epithelial cell and represses E-cadherin expression in mesenchymal cells; 2) a CAAT box (Ϫ65 to Ϫ61); and 3) two G-C rich regions I (Ϫ52 to Ϫ44) and II (Ϫ41 to Ϫ32), which confer basic epithelial promoter activity. A WT1/EGR-1 binding site (CCGGGGGCG) was found at position Ϫ51 to Ϫ43 of the murine E-cadherin promoter ( Fig. 2A), suggesting that WT1 could directly regulate E-cadherin. Sixteen different murine E-cadherin promoter fragments were tested for their response to the WT1 protein (Fig. 2B). Initially, the full-length proximal (Ϫ178) reporter construct was co-transfected into NIH 3T3 cells with vectors expressing WT1 from the Rous sarcoma virus long terminal repeat (Fig. 3). All four major isotypes of WT1 (A, B, C, D) strongly activated the E-cadherin promoter. Particularly striking activation was noted in response to WT1(A) and WT1(B), the isoforms of WT1 that lack the KTS sequence between zinc fingers 3 and 4, while isoforms C and D (ϩKTS) stimulated the promoter to a lesser extent. Since our previous studies were performed using the WT1(A) protein, we continued analysis with this isoform.
Four deletion constructs of the E-cadherin promoter were utilized to test the importance of the three regions in E-cadherin promoter that contribute to epithelial expression. The E-cad(Ϫ178/ϩ113) construct contains wild-type proximal promoter elements which, as described previously, are sufficientspecific for epithelial expression (47). The E-cad(Ϫ74/ϩ113) and E-cad(Ϫ67/ϩ113) constructs are deleted for the palindromic element (E-pal), but retain a CAAT box and a GC-rich potential WT1 binding site consensus sequence. The E-cad(Ϫ61/ϩ113) construct is deleted for the E-Pal and CAAT box, but retains the GC sequence. The E-cad(Ϫ61/ϩ113[GCm]) construct harbors a point mutation in the GC/WT1 element. The (Ϫ30) construct is a complete deletion of the three Ecadherin epithelial responsive elements (Fig. 2).
As shown in Fig. 4A, the full-length construct (Ϫ178), was activated about 15-fold by WT1(A) in NIH 3T3 cells relative to a control transfection with the insertless RSV expression vector. The (Ϫ74) and (Ϫ67) constructs, which were deleted for the E-pal element, were still strongly activated by WT1(A) (25-and 15-fold, respectively), which showed that E-pal segment is not important for WT1-mediated activation. Deletion of the CAAT box and GC elements (Ϫ30) significantly abrogated the ability of WT1(A) to activate reporter gene activity. Therefore, the minimal set of cis-acting sequences responsive to WT1(A) included the GC-I box (Ϫ61). Consistent with this view, when the GC-I sequence was mutated from CCGGGGGCG to CCGGGT-GCG, the ability of the (Ϫ61) reporter construct to be activated by WT1 was abolished in a manner similar to deletion of the GC-I sequence (i.e. the Ϫ30 construct) (Fig. 4A). It must be noted that the deletion mutations of the E-cadherin promoter, in addition to eliminating the ability of WT1 to activate tran-scription, severely impaired the basal activity of the promoter. This may be due to elimination of GC-rich sequences, which can bind the widely expressed Sp1 and CAAT-binding transcription factors. Potentially WT1 may need to cooperate with such basal elements and transcription factors in order to activate gene transcription.
Constructs Ϫ74 and Ϫ67 had the strongest response to WT1, which suggested that both the CAAT box and the GC-I region were essential for full activation. Therefore, we next performed a fine mutational analysis of the region between Ϫ67 and Ϫ44 to pinpoint the WT1-responsive element(s). For convenience, we divided the region into three subregions (C is the CAAT box; I is the intermediate region between CAAT box and GC-I; and GC is the GC-I site) (Fig. 2B). Whereas the wild-type Our findings did not rule out the possibility that intermediate sequences, including nucleotides "GCGG" located between the CAAT box and GC-I site were able to substitute for the loss of the GC-I site and that the observed response of the CAAT element to WT1 was really due to a second cryptic WT1-response element. To test this hypothesis, the GCGG sequence was drastically altered to TTTT and co-transfection analysis with WT1 was performed (Fig. 4C). When the CAAT box and GC-I sites were intact, mutation of the intermediate sequence (C-Im-GC) alone did not affect the ability of the construct to be induced by WT1. Mutation of the intermediate sequence in combination with either a mutant CAAT box (Cm-Im-GC) or mutant GC-I site (C-Im-GCm, C-Im-GCmm) (Fig. 4C) did not drastically affect the reporter activity (Fig. 4, compare B and  C). When the CAAT, intermediate, and GC-I sequences were simultaneously mutated (Cm-Im-GCmm in Fig. 4C), response to WT1 was lost. Collectively, our findings indicate that WT1 mediates activation of the murine E-cadherin promoter through the GC-I element and the CAAT-box (see Table I). The GC element was sufficient to mediate significant induction of the promoter, but the combination of the two sites led to an additive effect on transcription in the presence of WT1.
The human and murine E-cadherin promoters are very similar (47,49), and the GC element and CAAT box are conserved.

FIG. 3. Transcription of the murine E-cadherin promoter by the different isoforms of WT1.
A murine E-cadherin promoter/luciferase construct (0.1 g) was transiently co-transfected into NIH3T3 cells along with an expression vector for each of the four major WT1 spliced isoforms (WTA, WTB, WTC, WTD) (0.01, 0.1, 0.5, and 1 g) and a Renilla luciferase internal control vector (0.01 g). At 60 h after transfection, a dual luciferase assay was performed. Normalized luciferase expression Ϯ S.D. of triplicate experiments is plotted. White bar, 0.01 g; pale gray bar, 0.1 g; dark gray bar, 0.5 g; black bar, 1 g.
The human E-cadherin promoter responded to WT1 in a pattern qualitatively similar to the murine promoter (Fig. 5). Expression from a reporter containing sequences from Ϫ368 to ϩ125 of human promoter HE-cad (Ϫ368/ϩ125) was activated about 3-fold by WT1(A) relative to RSV control transfection, while in this set of experiments the murine promoter was activated 5-fold.
HE-cad (Ϫ217/ϩ125), which still contains the essential elements of the human E-cadherin promoter, was activated to a similar extent as the full proximal promoter. HE-cad(DEL), a construct in which all three critical elements of the promoter (Pal/Ap2, CAAT, and GC-I) were deleted, was not responsive to WT1. The CAAT box region and GC-I are well conserved between human and mouse, but the Pal/Ap2 region is not, which suggests the importance of CAAT and GC-rich region in Ecadherin gene regulation ( Fig. 2A) and the potential evolutionary conservation of a WT1/E-cadherin regulatory circuit.
Disruption of E-cadherin Expression by a Dominant Negative Form of WT1-Dominant negative form of WT1 mutants were previously identified both in tumor specimens as well as in patients with the Denys Drash syndrome (2). We showed that one particular mutant, WT1(AR) (43), which is deleted for exon 9 sequences encoding the third zinc finger motif of WT1, inhibited the ability of wild-type WT1 to activate transcription in a cotransfection assay (41). Therefore, we determined if WTAR could disrupt the activation of E-cadherin in cells stably and transiently expressing wild-type WT1. In a WT1-expressing NIH 3T3 cell line (WR16), E-cadherin promoter activity was approximately 5 times higher than in control NIH 3T3 cells that do not express WT1 (Fig. 6A). However, when WR16 cells were transiently transfected with the E-cadherin promoter along with a vector expressing the WT1(AR) mutant allele, E-cadherin promoter activity decreased by over 80% to a level similar to that obtained in control NIH 3T3 cells (Fig. 6B). When WTAR was transiently cotransfected with WT1(A) in the NIH 3T3 cells, WTAR blocked the WT1A-mediated activation of the E-cadherin promoter by more than 50% (Fig. 6C).
WT1 Binds to the GC-I Element of the E-cadherin Promoter-As shown above, deletion analysis of the murine E-cadherin promoter indicated that a GC-rich, EGR-1-like element and the CAAT box were required for WT1-mediated activation. To demonstrate a direct interaction between WT1 protein and these promoter sequences, a electrophoretic mobility shift assay (EMSA) was performed. An oligonucleotide corresponding Activates the E-cadherin Gene to nucleotides Ϫ62 to Ϫ24 of the E-cadherin promoter, which contains the GC-I/WT1 binding site, was incubated with in vitro transcribed and translated WT1 and subjected to electro-phoresis. A DNA-protein complex was formed when the oligonucleotide was incubated with WT1-programmed rabbit reticulocyte lysate (Fig. 7A, lane 2, lower arrow), but not with unprogrammed lysate (lane 1). This complex was specifically retarded by a WT1 antibody (Fig. 7A, lane 4, upper arrow) but was unaffected by a control antibody, G4 (lane 6). No WT1/DNA complex was formed on a mutant oligonucleotide representing the same GCm sequence that disabled the response of the murine E-cadherin promoter to co-expressed WT1 (data not shown). Unlabeled GC oligonucleotides efficiently competed for WT1 binding to the wild-type probe, while the GCm sequence could not effectively compete for binding even when present in the reaction at a 2000-fold molar excess (Fig. 7B).
Since all four isoforms of WT1 could transactivate the Ecadherin promoter (Fig. 3), we tested whether both the ϩ and Ϫ KTS forms of WT1 could bind to the GC-I/WT1-response element. GST fusion proteins containing the four WT1(A/ϪKTS) zinc fingers, WT1(C/ϩKTS) zinc fingers, and three zinc fingers of the dominant negative/DNA binding-deficient WT(AR) mutant were allowed to bind an oligonucleotide containing the GC and CAAT elements of the E-cadherin promoter (Fig. 8). As expected (43), GST-WT1(AR) did not bind to DNA. GST-WT1(A/ϪKTS) bound relatively strongly to the promoter sequence, while WT1(C/ϩKTS) yielded 1/10 the amount of DNAprotein complex. This is consistent with transfection data, which showed that WT1(C) and WT1(D) have a reduced but definite capacity to activate the E-cadherin promoter (Fig. 3).
To map the WT1 binding element within the E-cadherin promoter, oligonucleotides containing both the CAAT box and the GC-I/WT1 binding site (Sequences 18 -27) as well as mutant versions of these sites were utilized in an EMSA with in vitro translated WT1(A) (Fig. 9). Since the CAAT box and the GC-I site recognize many widely expressed transcription factors, it was necessary to add a WT1 antibody to the reaction to more clearly identify the WT1/DNA complex (Fig. 9, A-D, lane  1, top arrow). A WT1 antibody supershift complex was observed only when oligonucleotides containing the wild-type GC-I/WT1 binding site were used as probes (Fig. 9, A (lanes 5 and 9), and B (lane 5)). Mutations in the CAAT box or the intermediate sequence still allowed the WT1 protein to bind to GC-I site (Fig.  9, A (lane 9) and B (lane 5)). An oligonucleotide containing the CAAT box alone did not have any WT1 binding activity (Fig.  9B, lanes 7-10). Oligonucleotides with a wild-type CAAT box and mutant GC-I site (Fig. 9, C and D, lanes 3-6) did not bind to WT1 even though the CAAT box could mediate transactivation in response to WT1. To show that WT1 produced in tran- FIG. 5. Activation of the human Ecadherin promoter by WT1. NIH 3T3 cells were transfected with 0.5 g of a RSV control expression vector (white bar) or 0.5 g of RSV-WT1(A) (black bar) along with 0.1 g of the indicated human Ecadherin promoter construct. The HECad (Del) is devoid of all three major promoter elements (Pal/Ap2, CAAT, and GC-I) and is similar to Ϫ30 murine E-cadherin promoter construct. A full-length murine Ecadherin promoter construct (MECad Ϫ178) was used as a positive control. A Renilla luciferase expression vector (0.01 g) was used to normalize for transfection efficiencies. The average normalized luciferase activity Ϯ S.D. of triplicate experiments is presented.

FIG. 6. A dominant negative WT1 mutant inhibits the E-cadherin promoter.
A, the E-cadherin promoter is stimulated in WT1expressing NIH 3T3 cells. The E-cadherin luciferase construct (0.1 g) was transfected into parental NIH 3T3 cells or WT1-expressing WR16 cells along with 0.01 g of the Renilla luciferase control expression vector to normalize transfection efficiency. B, the E-cadherin promoter (0.1 g) was transfected into WR16 cells along with an insertless RSV expression vector or RSV-WT1(AR) (0.5 g) encoding a dominant negative form of WT1. C, the E-cadherin promoter (0.1 g) was co-transfected into NIH 3T3 cells with RSV-WTA (0.5 g) in the presence or absence of RSV-WT1(AR) (0.5 g). In all experiments, normalized luciferase activity was determined 60 h after transfection. The average normalized luciferase activity Ϯ S.D. of triplicate experiments is presented. sient transfections could bind the GC element, we attempted mobility shift assays utilizing extracts from transfected cells but a large number of background complexes were noted, potentially representing proteins such as SP1 and Ap2 that can also bind to GC-rich elements. To determine if WT1 was among these complexes, we performed an EMSA with extract from 293T cells transfected with RSV-WT1(A). followed by transfer of the gel for immunoblot analysis (Fig. 10). A WT1-DNA complex signal was detected only in 293T cell nuclear extracts transfected with WT1 (lanes 3 and 4) and not in NIH 3T3 cell nuclear extract (Fig. 10, lanes 1 and 2) or mock-transfected 293T cell extract (Fig. 10, lanes 5 and 6). A summary of DNA binding activity and transcriptional response of the mutant promoter constructs is presented in Table I. DISCUSSION Although it had been shown that WT1 protein could activate or repress the expression of a number of promoters in co-transfection experiments (2), then endogenous forms of many of these putative target genes were later shown not to be regulated by WT1 in biological systems (25,52). Much of this discrepancy is due to the mode by which WT1 target genes were identified, namely by searching the promoters of promising candidate target genes for sequences matching the EGR1/WT1 binding site. Given the relatively GC-rich nature of many promoters, this approach identified a number of putative target gene. True WT1 target genes should have a specific binding site for the protein, respond to WT1 expression in transient assays, and be activated or repressed by both transient and stable co-transfection. However, in addition, the expression of the endogenous gene, not just a promoter construct should be affected by expression of WT1. Furthermore, a WT1 target gene should be expressed in the developing and adult organism in a temporal and spatial pattern that is concordant with WT1 expression. The transcriptional activity of WT1 required for growth control has been controversial. Recently, we demonstrated that missense mutants of WT1 defective for transcriptional activation, but competent for transcriptional repression, failed to suppress cell growth (42). This indicates that genes whose transcription is activated by WT1 may be critical for the biological activity of this tumor suppressor.
We have attempted to identify WT1 target genes using a model system in which engineered expression of WT1 converts the NIH 3T3 fibroblast cell line into cells with many features of epithelia (26). This system was used to survey the expression of many epithelial markers without prior knowledge of the nature of their promoter sequences. E-cadherin was among those genes up-regulated in WT1-expressing cells. Here we report that the E-cadherin gene meets criteria for a WT1 target gene potentially relevant for the roles of WT1 in both inhibiting cell growth and inducing differentiation. Though up-regulation of E-cadherin in the original cell lines could possibly be attributed to clonal selection, we found that E-cadherin mRNA expression was induced within 24 h of infection of NIH 3T3 cells with a retrovirus expressing WT1, consistent with a direct effect of WT1.
Deletions and point mutational analysis of the murine and human E-cadherin promoters and mobility shift assays indicated that an EGR-1-like cis-acting sequence was necessary and sufficient for WT1-mediated transactivation. We then demonstrated that the GC element and the CAAT box were required for full and robust WT1-mediated transactivation of E-cadherin in mesenchymal NIH 3T3 fibroblasts (Fig. 4). Furthermore, a dominant negative mutant of WT1 WT1(AR) inhibited E-cadherin promoter activity in cells expressing wild-  1-6), or a mutant oligonucleotide (lanes 7-12) containing a G to T transversion was incubated with control reticulocyte lysates (C) or in vitro translated WT1 (W). Antibodies used for supershift assay were anti-WT1 (C19) (lanes 3 and 4) and an anti-Gal4 (G4) antibody as a negative control (lanes 5 and 6). B, competition EMSA showing specificity of WT1 binding. In vitro translated WT1 was allowed to bind to the GC element in the absence of competitor or in the presence of the indicated -fold excess of wild-type or mutant GC sequence from the E-cadherin promoter. type WT1 (Fig. 6). Finally, we found that WT1 protein derived from nuclear extracts could bind to the WT1-response element in the E-cadherin promoter (Fig. 10).
It must be noted that detection of WT1 DNA binding activity in nuclear extracts has been reported in only a few publications (53)(54)(55). Curiously, WT1 also appeared to mediate activation of the E-cadherin promoter through a CAAT box, because mutation of this element, but not sequences between the CAAT box and the GC-I element, significantly impaired the ability of WT1 to activate the E-cadherin promoter. Furthermore, a construct with a mutation in the GC box, but with an intact CAAT box, still supported transcriptional activation by WT1 (summarized in Table I). This is consistent with prior information that WT1 can affect transcription through protein-protein interactions. WT1 was shown to interact with p53 and modulate the transcription of reporter genes containing only p53 binding sites (56). WT1 also affects the transcription of the HSP70 promoter, presumably by sequestration of HSP70 from the heat shock transcription factor, thus leading to the activation of the heat shock transcription factor (57). The CAAT box can bind to the CTF/NF1 factors as well as the heteromeric CP1/CP2 factor. Whether WT1 directly binds to these factors to augment their transcriptional effect or sequesters negative regulators that interact directly or indirectly with the E-cadherin CAAT box is currently under investigation. However, the finding that a CAAT box may act as a WT1-responsive element suggests a novel mechanism of WT1 action and a broader potential range of target genes.
It was previously reported that epithelial-specific transcription of E-cadherin required a palindromic sequence as well as two tandem AP-2 transcription factor binding sites within the proximal GC-rich portion of the promoter, which we have demonstrated can also bind to WT1 (47,50,51). Rb and Myc were later shown to activate E-cadherin gene expression via interaction with AP-2, through the E-Pal and GC-rich region in epithelial cells, but not in mesenchymal NIH 3T3 cells (58). Consistent with these data, we found that transfection of Rb did not activate the E-cadherin promoter and co-transfection of WT1 and Rb did not yield synergistic or additive activation on E-cadherin expression in NIH 3T3 cells (data not shown). However, unlike Rb or Myc, WT1 could transactivate the E-cadherin gene in non-epithelial NIH 3T3 cells. This suggests an independent pathway for epithelial differentiation exerted by WT1 and is consistent with the notion that WT1 might upregulate E-cadherin expression in mesenchymal cells during the mesenchymal-epithelial transition. However, our studies have not excluded the possibility of inteaction between WT1 and AP-2 or other proteins bound to the E-Pal site. WT1mediated activation was most efficient when the E-Pal element was deleted (Figs. 3 and 4). This finding suggests the presence of a negative regulator bound to the E-Pal element in mesenchymal cells, consistent with previous findings (47,50,51). Such a protein might act to counter WT1-mediated activation of E-cadherin in certain tissues.
E-cadherin expression is characteristic of epithelia and is critical for epithelial development (27). E-cadherin is required for embryogenesis, as targeted disruption of this gene leads to failure of early murine development due to a defect in embryo compaction (59,60). More specifically, in the kidney, E-cad- Oligonucleotides representing nucleotides Ϫ67 to Ϫ24 of the E-cadherin promoter, which contains both CAAT box and GC-I WT1 binding site and its mutant versions, were incubated with in vitro translated WT1 (W) or unprogrammed lysate (C). Antibodies used for supershift assay were anti-WT1 C-19 antibody (C19), while a anti-Gal4 antibody (G4) was used as a negative control. A, probes C-I-GC and Cm-Im-GC. B, probes Cm-I-GC and CAAT box alone. C, probes C-Im-GCmm and Cm-Im-GCmm. D, probes C-I-GCmm and Cm-I-GCmm.
FIG. 10. EMSA/Western analysis of WT1 protein bound to GC-I/WT1 binding site. An EMSA assay was performed allowing an unlabeled Ϫ62/Ϫ24 oligonucleotide to interact with nuclear extracts derived from NIH 3T3 cells (lanes 1 and 2), mock-transfected 293T cells (lanes 5 and 6), and 293T cells transfected with a WT1 expression plasmid (lanes 3 and 4). The DNA-WT1 complex was separated by non-denaturing gel electrophoresis and transferred to a nylon membrane, followed by immunoblot with an anti-WT1 antibody. herin expression is induced in the condensing metanephric mesenchyme during the mesenchymal-epithelial transition (37,38,61) at the same time that WT1 is up-regulated in this tissue (39,40). This fact, along with the other evidence outlined, makes a compelling case for E-cadherin as a direct WT1 target gene. Direct transactivation of E-cadherin by WT1 demonstrates an evident link between the tumor suppressive activity of WT1 and a gene that plays a major role in differentiation. It confirms our previous report that WT1 plays a direct role in both the induction of the epithelial phenotype and inhibition of tumorigenesis by controlling the expression of a subset of genes encoding molecules that control epithelial characteristics (26). E-cadherin might mediate many facets of growth suppression by WT1. Homotypic interactions mediated by E-cadherin lead to increased cell adhesion and decreased invasiveness. Furthermore, E-cadherin was recently demonstrated to have an anti-proliferative effect of its own, correlated with the up-regulation of the CDK inhibitor, p27 (62). We checked for p27 up-regulation after infection of 3T3 cells with the WT1 retrovirus and found that p27 was expressed at a basal level in 3T3 but was not up-regulated by WT(A) (Data not shown). This may be due to the relatively low level of Ecadherin induced after transient expression and does not preclude the possibility that E-cadherin expression in the developing kidney may affect the expression of p27 or other cyclindependent kinase inhibitors.
The E-cadherin gene is frequently inactivated in human cancer, and its down-regulation is particularly associated with the acquisition of an invasive tumor phenotype (28,34,63). Allelic loss of heterozygosity of E-cadherin has been reported in cervical, prostrate, breast, hepatocellular, and colorectal carcinomas as well as in Wilms' tumor itself (28, 64 -68). Point mutations of the E-cadherin gene were found in gastric, renal, lung, colorectal, and breast cancers (28, 66, 69 -74). Defects in trans-acting pathways regulating E-cadherin expression have been suggested in breast cancer (44), and epigenetic inactivation of E-cadherin gene was also implicated by changes in chromatin structure and the methylation status of CpG sites within of the E-cadherin promoter (50). Recently, E-cadherin expression within Wilms' tumors was found only in a limited set of cells resembling renal tubules (75). These tumors coexpressed cytokeratin and vimentin, underscoring the aberrant differentiation of the tumor. Altered E-cadherin expression as a result of lost or mutated WT1 could play a role in the pathogenesis of Wilms' tumors. Furthermore, E-cadherin complexes with ␤-catenin, a protein involved in transduction of wnt signaling to the nuclear TCF/LEF transcription factors (76). Activating mutations of ␤-catenin were recently identified in Wilms' tumor lending more evidence to the potential importance of disruption of the E-cadherin/catenin axis in this tumor (77).
In summary, E-cadherin is a gene coordinately regulated with WT1 during kidney development, whose expression can be activated in response to stable or transient expression of WT1 and whose promoter can be bound and activated by WT1. Hence, E-cadherin is a likely a bona fide target gene of WT1.


s2fieldsofstudy: Biology Chemistry Medicine

text:
Potentiation of Rho-A-mediated Lysophosphatidic Acid Activity by Hyperinsulinemia*

We have shown previously that insulin promotes phosphorylation and activation of farnesyltransferase and geranylgeranyltransferase (GGTase) II. We have now examined the effect of insulin on geranylgeranyltransferase I in MCF-7 breast cancer cells. Insulin increased GGTase I activity 3-fold and augmented the amounts of geranylgeranylated Rho-A by 18%. Both effects of the insulin were blocked by an inhibitor of GGTase I, GGTI-286. The insulin-induced increases in the amounts of geranylgeranylated Rho-A resulted in potentiation of the Rho-A-mediated effects of lysophosphatidic acid (LPA) on a serum response element-luciferase construct. Preincubation of cells with insulin augmented the LPA-stimulated serum response element-luciferase activation to 12-fold, compared with just 6-fold for LPA alone (p < 0.05). The potentiating effect of insulin was dose-dependent, inhibited by GGTI-286 and not mimicked by insulin-like growth factor-1. We conclude that insulin activates GGTase I, increases the amounts of geranylgeranylated Rho-A protein, and potentiates the Rho-A-dependent nuclear effects of LPA in MCF-7 breast cancer cells.

We have shown previously that insulin promotes phosphorylation and activation of farnesyltransferase and geranylgeranyltransferase (GGTase) II. We have now examined the effect of insulin on geranylgeranyltransferase I in MCF-7 breast cancer cells. Insulin increased GGTase I activity 3-fold and augmented the amounts of geranylgeranylated Rho-A by 18%. Both effects of the insulin were blocked by an inhibitor of GG-Tase I, GGTI-286. The insulin-induced increases in the amounts of geranylgeranylated Rho-A resulted in potentiation of the Rho-A-mediated effects of lysophosphatidic acid (LPA) on a serum response element-luciferase construct. Preincubation of cells with insulin augmented the LPA-stimulated serum response element-luciferase activation to 12-fold, compared with just 6-fold for LPA alone (p < 0.05). The potentiating effect of insulin was dose-dependent, inhibited by GGTI-286 and not mimicked by insulin-like growth factor-1. We conclude that insulin activates GGTase I, increases the amounts of geranylgeranylated Rho-A protein, and potentiates the Rho-A-dependent nuclear effects of LPA in MCF-7 breast cancer cells.
Small molecular weight GTPases of the Ras superfamily play an important role in cell proliferation, differentiation, structural organization, and vesicular trafficking (1)(2)(3)(4). The members of this superfamily (including Ras, Rho, and Rab proteins) are activated by GTP loading in response to guanine nucleotide exchange factors (5). Post-translational modification of these proteins by prenylation appears to be a prerequisite for their subsequent activation (6,7).
Prenylation of these GTPases is catalyzed by farnesyltransferase (FTase) 1 or geranylgeranyltransferases I or II (GGTase I and II), which promote the attachment of either a farnesyl or a geranylgeranyl moiety, respectively, to conserved cysteine residues located at the C termini of Ras, Rho, and Rab proteins (8 -10). Whether a protein is farnesylated or geranylgeranylated is determined by its specific C-terminal sequence. Ras proteins contain serine, methionine, or glutamine at the X position of the terminal CAAX box and are farnesylated, whereas Rho proteins have leucine at this position and are geranylgeranylated. The C terminus of the Rab proteins contains either a CC or CXC sequence, which is double geranylgeranylated by GGTase II.
We have recently demonstrated that insulin promotes the phosphorylation of the ␣-subunit of FTase, increases its enzymatic activity, and augments the amounts of farnesylated p21 ras in 3T3-L1 fibroblasts, 3T3-L1 adipocytes, and vascular smooth muscle cells (11)(12)(13). Furthermore, tissues of hyperinsulinemic animals display both increased activity of FTase and increased amounts of farnesylated p21 ras (14). Because FTase and GGTase I share the same ␣-subunit of the heterodimeric structure (15), we decided to evaluate the potential effect of insulin on GGTase I and the amount of geranylgeranylated Rho proteins. This has not been tested experimentally to date.
Furthermore, because Rho-A mediates transcriptional activation of the serum response element (SRE)-dependent genes by lysophosphatidic acid (LPA), we also examined whether the insulin-induced increases in the amounts of geranylgeranylated Rho-A potentiate the transcriptional activity of LPA in MCF-7 breast cancer cells. These cells possess insulin receptors and represent a good model to study insulin's mitogenic influence.

MATERIALS AND METHODS
Tissue culture media, gentamicin, and the LipofectAMINE/Plus Reagent transfection kit were from Life Technologies, Inc. Fetal calf serum was from Gemini Bio-Products, Inc. (Calabasas, CA). Bovine serum albumin and other biochemicals were from Sigma. Insulin was from Elli Lilly (Indianapolis, IN). IGF-1, L-␣-lysophosphatidic acid and acid molybdate reagent spray were from Sigma. Anti-Rho-A mouse monoclonal antibody was from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Protein G-Plus/Protein A-agarose immunoprecipitation reagents were from Oncogene Science, Inc. (Cambridge, MA). Bicinchoninic acid (BCA) protein assay kit was from Pierce. [ 3 H]Geranylgeranyl pyrophosphate and [ 32 P]orthophosphate were from PerkinElmer Life Sciences. SDS-PAGE supplies and reagents were from Bio-Rad, TLC (thin layer chromatography) Silica gel 60 plates were from EM Separations Technology (Darmstadt, Germany), enhanced chemiluminescence (ECL) kit was from Amersham Pharmacia Biotech, the Enhanced Luciferase Assay kit was from Analytical Luminescence Laboratory (Sparks, MD), and the Luminescent ␤-Galactosidase Detection Kit II was a product of CLONTECH Laboratories, Inc. (Palo Alto, CA). The serum response element-luciferase (SRE-Luc) construct, the luciferase construct without the SRE (PGL 3 ), and the cytomegalovirus-␤-galactosidase (CMV-␤Gal) construct were all gifts from Dr. Songzhu An (University of California, San Francisco, CA). MCF-7 breast cancer cells were a gift from Dr. Kathryn B. Horwitz (University of Colorado Health Science Center, Denver, CO). Geranylgeranyltransferase inhibitor-286 (GGTI-286) was from Calbiochem (San Diego, CA). Recombinant human Ras-CVLL was from Calbiochem (San Diego, CA) and was used as a sub-* This work was supported by a Veterans Affairs Research Service career development award (to J. C.), by the Foundation for Biomedical Education and Research, by an American Heart Association fellowship (to M. G.), and by the American Diabetes Association. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
GGTase I Activity-MCF-7 cells were grown to 80% confluence at 37°C, 5% CO 2 (Eagle's minimal essential medium (MEM) (Life Technologies, Inc.) ϩ 5% heat-inactivated fetal bovine serum, nonessential amino acids, L-glutamine (200 mM), and insulin (60 pM). The cells were serum-and insulin-starved for 24 h and then preincubated with insulin (100 nM) for 1 h with and without 3 M GGTI-286. The cells were washed with phosphate-buffered saline and lysed using a Triton X-100based lysis buffer (50 mM HEPES, pH 7.5, 150 mM NaCl, 15 mM MgCl 2 , 1 mM phenylmethylsulfonyl fluoride, 1 mM Na 2 HPO 4 , 1% Triton X-100, 1 mM dithiothreitol, 1 mM sodium vanadate, 0.05% SDS, 10 g/ml aprotinin, 10 g/ml leupeptin). After pelleting the cellular debris, the supernatant was normalized for protein using the BCA protein assay. A 5-l aliquot of the lysate was incubated with a reaction assay solution containing recombinant Ras-CVLL (a Rho analog) and [ 3 H]geranylgeranyl pyrophosphate for 30 min at 30°C. The reaction was stopped using 1 N HCl in ethanol and filtered through Whatman GF/C filter paper. The quantity of [ 3 H]geranylgeranyl incorporation into recombinant human Ras-CVLL per milligram of protein was determined by liquid scintillation counting. 32 P Phosphorylation of ␣-Subunit of Geranylgeranyltransferase I-MCF-7 cells were grown to confluence at 37°C, 5% CO 2 (as described above). Cells were then incubated for 60 min at 37°C in serum-and phosphate-free MEM ϩ nonessential amino acids, and L-glutamine (200 mM), then preincubated overnight with 250 Ci of [ 32 P]orthophosphate (10 mCi/mmol). Cells were then incubated for 60 min with insulin (100 nM) or IGF-1 (13 nM). Cells were lysed as described previously (11)(12)(13)16), sonicated, centrifuged, and protein concentrations diluted to 0.5 mg/ml. FTase protein was immunoprecipitated, using antibodies to the ␤-subunit of farnesyltransferase. Geranylgeranyltransferase was then immunoprecipitated from the post-FTase immunoprecipitaiton lysates with antibodies to the ␣-subunit of GGTase I and analyzed by 12% SDS-PAGE. Amounts of phosphorylation were visualized by autoradiography, and amounts of protein were determined by Western blotting.
Measurements of Geranylgeranylated Rho-A-MCF-7 cells were grown to confluence at 37°C, 5% CO 2 (as described above). The cells were serum-and insulin-starved for 24 h, and then preincubated with insulin (100 nM) for 1 h and 24 h (with and without 3 M GGTI-286). The cells were washed with phosphate-buffered saline and lysed as described above. After pelleting cellular debris, the supernatant was standardized to 1 mg/ml protein using BCA protein assay and then separated into an aqueous and detergent phases using Triton X-114 as described previously (11)(12)(13). Using anti-Rho antibodies, non-prenylated Rho-A and prenylated Rho-A were immunoprecipitated from the aqueous and detergent phase, respectively (11)(12)(13)16). Immunoprecipitates were resolved by PAGE, determined by Western blot, and quantitated by densitometry. Previous experiments with prenylated Ras and Rab proteins determined that 100% of the non-prenylated and prenylated proteins are recovered from their respective phases (11)(12)(13)16). We use the Triton X-114 methodology because it allows quantitation of the changes in the amounts of prenylated proteins. The mobility shift approach to identify prenylated proteins is strictly qualitative and, therefore, was not utilized.
GTP Loading of Rho-A-MCF-7 cells were grown to confluence at 37°C, 5% CO 2 (as described above). Cells were then incubated for 24 h at 37°C in serum-and phosphate-free MEM ϩ nonessential amino acids, and L-glutamine (200 mM). The cells were then preincubated for 24 h with insulin (1 nM) Ϯ 3 M GGTI-286. 500 Ci of [ 32 P]orthophosphate (10 mCi/mmol) was added, and the cells were incubated another 4 h. 10 M LPA was added, and the cells were then incubated for an additional 60 min. Cells were lysed as described previously (11)(12)(13)16), sonicated, centrifuged, and protein concentrations diluted to 0.5 mg/ml. Rho-A was immunoprecipitated and resolved by thin layer chromatography (TLC Silica gel 60 plates) using a solution containing 0.75 M ammonium formate, 0.5 M LiCl, and 0.57 M HCl. Gels were dried overnight, then sprayed with acid molybdate reagent, and allowed to dry. The position of labeled GTP and GDP on the TLC plates was visualized by autoradiography. The areas of the gel corresponding to the GTP and GDP bands of the autoradiogram were excised from the plates, and the amounts of labeled nucleotide were assessed by scintillation counting. The amounts of GTP were expressed in percentage above control of total GTP/GDP binding to Rho-A.
Effect on SRE-dependent Gene Transcription-MCF-7 cells were grown to confluence at 37°C, 5% CO 2 (as described above). Using LipofectAMINE/Plus reagent, the cells were co-transfected with either a SRE-Luc construct or a luciferase construct minus the SRE (PGL 3 ), and a CMV-␤Gal construct, which was used as a control for transfection efficiency and processing. The cells were serum-starved for 24 h, preincubated with insulin (100 nM) in the absence or presence of GGTI-286 (3 M) for 24 h, and then challenged with LPA (10 M) for 16 h. Control cells were incubated with or without insulin for 40 h. The cells were washed with phosphate-buffered saline and lysed by a freeze/thaw method using a Triton X-100-based lysis buffer provided in the Enhanced Luciferase Assay kit. Luciferase activity was then measured on a Monolight 2010 luminometer (Analytical Luminescence Laboratory, Sparks, MD) using the Enhanced Luciferase Assay kit and corrected for by ␤-galactosidase activity (determined by chemiluminescence). Efficiency is equal to the luciferase activity divided by the ␤-galactosidase activity. overnight. The cells were then challenged with insulin (100 nM) or IGF-1 (13 nM) for 1 h. Cell lysates were normalized for protein, followed by immunoprecipitation and clearance of FTase protein, using antibodies to the ␤-subunit of FTase. GGTase I was then immunoprecipitated from the residual supernatant, using antibodies to the ␣-subunit of GGTase I, and analyzed by SDS-PAGE. Relative amounts of phosphorylation were determined by autoradiography. A representative experiment is shown.
Statistical Analysis-Statistics were analyzed by Student's paired or unpaired t test, with p Ͻ 0.05 considered significant.

RESULTS
GGTase I Activity and Phosphorylation-In the initial experiments, we examined the effect of insulin on GGTase I activity in MCF-7 cells. Insulin activated GGTase I, and its effect on GGTase I activity was inhibited by a specific inhibitor of this prenyltransferase, GGTI-286 (Fig. 1). The effect of insulin was evident by 10 min of incubation. Although the time course shown in Fig. 1 is limited to 120 min, the effect of insulin on all of the prenyltransferases remains for as long as insulin is present (data not shown).
We then examined the effect of insulin on the phosphorylation of the GGTase I ␣-subunit. Because FTase and GGTase I share a common ␣-subunit (15), we eliminated FTase by immunoprecipitation with antibody to the ␤-subunit of FTase. We then used the anti-␣-subunit antibody to immunoprecipitate and analyze the ␣-subunit remaining with GGTase I. Insulin increased phosphorylation of the GGTase I ␣-subunit (Fig. 2) without affecting the total amount of GGTase I protein (data not shown). Insulin's ability to activate GGTase I was consistent with its effects on the phosphorylation of the ␣-subunit and similar to its influence on FTase (11)(12)(13) and GGTase II (16). We also tested the effect of IGF-1 on the phosphorylation of the GGTase I ␣-subunit and found that the ability to enhance the phosphorylation of the GGTase I ␣-subunit was specific to insulin (Fig. 2).
Since activation of the prenyltransferases result in increases in the amounts of prenylated proteins, we investigated the effect of insulin on the amounts of geranylgeranylated Rho-A, a substrate of GGTase I. The amount of geranylgeranylated Rho-A was determined by Triton X-114 extraction, as described for prenylated Ras and Rab proteins (11,16). The detergent phase completely extracts prenylated proteins from the cell lysate, while non-prenylated proteins remain in the aqueous phase (11,16). The amount of geranylgeranylated Rho-A is then expressed as the percent of the total cellular Rho-A. In accord with its stimulatory effect on GGTase I, insulin significantly increased the amounts of geranylgeranylated Rho-A from 38 Ϯ 3% to 56 Ϯ 4% by 1 h (p Ͻ 0.01). These increases were completely blocked by GGTI-286 (Fig. 3). The total amount of cellular Rho-A were not influenced by insulin and remained constant in all experiments (data not shown).
GTP Loading of Rho-A-Because insulin increased the amount of prenylated Rho-A available for activation by GTP loading, we were interested in determining the levels of Rho- Amounts of GTP and GDP bound to Rho-A were determined as described under "Materials and Methods," visualized by autoradiography, and quantitated by liquid scintillation. The amounts of Rho-A-GTP were expressed as percentage above control, which ranged between 31% and 35%, *, p Ͻ 0.02.
A-GTP in cells preincubated with insulin and then challenged with LPA. Insulin and LPA alone stimulated loading of Rho-A with GTP above control by 32% and 45%, respectively. Preincubation with insulin (1 nM) and subsequent challenge with LPA increased Rho-A GTP loading by 57% (Fig. 4). Because incubation of cells in the presence of GGTI-286 resulted in essentially complete inhibition of Rho-A geranylgeranylation, we observed only a negligible effect of insulin with LPA on Rho-A GTP loading in the presence of GGTI-286.
Effect of Insulin on LPA-mediated SRE-dependent Gene Transcription-Geranylgeranylated Rho-A plays an important role in mediating the transcriptional activation of SRE-dependent genes by LPA (17). If insulin increases the amounts of geranylgeranylated Rho-A available for activation, conceivably this effect might enhance the transcriptional activity of LPA. To test this hypothesis, we co-transfected MCF-7 cells with either an SRE-Luc construct or a luciferase construct without the SRE (PGL3), and a constitutively active CMV-␤Gal construct. The cells were then incubated with insulin alone, LPA alone, or insulin for 24 h followed by LPA. After appropriate corrections for the transfection efficiency (using ␤-galactosidase) and background (using PGL3) we found that 100 nM insulin increased the luciferase reporter activity 3-fold, whereas LPA increased the luciferase reporter activity 6-fold (Fig. 5A). Preincubation of the cells with insulin increased LPA effect to 12-fold. This "priming" effect of insulin was blocked by GGTI-286 and was not mimicked by IGF-1 (Fig. 5A). The effect of GGTI-286 was specific for insulin because no effect was noted in the presence of LPA alone (data not shown). Furthermore, the ability of insulin to potentiate the effect of LPA was dosedependent (Fig. 5B). Preincubation of MCF-7 cells with 1, 10, or 100 nM insulin increased LPA's effect by 5%, 40%, and 102%, respectively. Although the data in Fig. 5B show that the LPA effect is significant only in the presence of 100 nM insulin, potentiation by 10 nM insulin approached significance in the number of experiments run. DISCUSSION The salient feature of this investigation is that insulin activates GGTase I and increases the cellular amounts of gera- nylgeranylated Rho-A. Physiologically, this action of insulin results in the augmentation of the Rho-A-mediated transcriptional activation of SRE-dependent genes by LPA. Cells preincubated with insulin and then followed by LPA exhibited a 12-fold increase in luciferase activity, which was greater than stimulation by insulin alone (3-fold), LPA alone (6-fold), or the addition of these two results. Thus, the response to insulin plus LPA was synergistic and gave a greater than additive effect in luciferase activity, clearly showing the ability of insulin to potentiate LPA's action on SRE-dependent genes. Synergy is defined as a result greater then that of either A or B alone and greater than could be expected from simple addition of the individual effects (18). The effect of insulin alone was not significantly different from the control, but the dose response to insulin showed a greater "priming" effect with increasing doses of insulin (Fig. 5B).
We have previously shown that insulin is a potent activator of FTase and GGTase II in a variety of cell types and tissues (11,13,16,19). Insulin-stimulated FTase activity results in significant increases in the amounts of farnesylated p21 ras and augmentation of DNA synthesis in response to other growth factors, as measured by incorporation of bromodeoxyuridine (13,20). Thus, IGF-1, epidermal growth factor, and plateletderived growth factor elicit greater incorporation of bromodeoxyuridine into DNA of 3T3-L1 fibroblasts preincubated with insulin than in its absence. In vascular smooth muscle cells, insulin potentiates the platelet-derived growth factor-induced vascular endothelial growth factor gene expression and thymidine incorporation (12). These "priming" effects of insulin are blocked by an inhibitor of FTase, ␣-hydroxyfarnesylphosphonic acid, suggesting that the effects of insulin are mediated by increased activity of FTase (12,20). In 3T3-L1 adipocytes, insulin also promotes increases in the activity of GGTase II and the amounts of geranylgeranylated Rab-3 and Rab-4 (16). The physiological relevance of the effect of insulin on geranylgeranylation of Rab proteins remains to be determined.
The mechanism of insulin's effect on the prenyltransferases involves the phosphorylation of the ␣-subunit of FTase (19) and GGTase II (16). We now demonstrate that insulin also promotes the phosphorylation of the ␣-subunit of GGTase I in MCF-7 cells. Inhibition of the phosphorylation of the ␣-subunit of FTase results in diminution of FTase activity (19). The ability of insulin to promote the phosphorylation of the ␣-subunit and activate these prenyltransferases is insulin-specific and is not mimicked by IGF-1, epidermal growth factor, or platelet-derived growth factor (13). This effect requires the presence of the intact insulin receptor, as cells derived from insulin receptor knock-out mice did not respond to insulin in terms of FTase activation (13). Furthermore, cells with a chimeric insulin-IGF-1 receptor also fail to activate FTase in response to insulin (13). Even though IGF-1 is a more potent mitogen than insulin, IGF-1 alone showed no significant increase in the amounts of farnesylated p21 ras , indicating no effect on the activity of farnesyltransferase (13). The present data are in agreement and demonstrate that IGF-1 alone exerted a greater influence on SRE-Luc than insulin, yet had no effect on either phosphorylation of the GGTase I ␣-subunit (Fig. 2) or potentiation of the nuclear effect of LPA (Fig. 5A). Because GGTase I shares the ␣-subunit with FTase (15) and IGF-1 has no effect on its phosphorylation (Fig. 2), it would not, therefore, be expected to have an effect on GGTase I activity. Furthermore, the inability of IGF-1 to prime the effect of LPA is consistent with the lack of IGF-1 effect on prenylation.
Investigation of insulin signaling to GGTase I was beyond the scope of this study. Previously, we have shown that insulin's effect on the phosphorylation of the ␣-subunit of FTase and GGTase II was blocked by an inhibitor of the mitogenactivated protein kinase kinase (16,19) and by transfection with a dominant negative mutant of Ras (19). These data suggest that insulin activates FTase in a "positive feedback" fashion using the Ras-mitogen-activated protein kinase pathway to promote the phosphorylation and activation of the enzyme. Interestingly, however, other growth factors, which activate the Ras-mitogen-activated protein kinase pathway, fail to influence FTase (13), indicating the specificity of insulin action and that other signaling intermediates may be involved for insulin to activate prenyltransferases.
The ␣-subunit of FTase also belongs to GGTase I (15). Because this subunit is phosphorylated in response to insulin, one could propose that insulin might activate GGTase I similarly to FTase. We have demonstrated that insulin increases the activity of GGTase I and consequently, the amounts of geranylgeranylated Rho-A. The question that presents itself is, what could be the physiological or pathophysiological consequences of this aspect of insulin action? Rho-A is a member of the Ras superfamily of small (20)(21)(22)(23)(24)(25) GTPases. Currently at least 14 mammalian Rho family proteins have been identified and have been shown to share 50 -90% amino acid sequence homology (2). Similar to Ras, Rho proteins function as GTP/GDP-regulated binary switches, which control signaling pathways responsible for numerous important cellular functions. Rho proteins are active when they are in the GTP-bound state. Guanine nucleotide exchange factors (of the Dbl oncogene family) catalyze the release of bound GDP, thus allowing GTP to bind and activate these GTPases (21). GTPase-activating proteins, on the other hand, stimulate the low intrinsic GTPase activity of the Rho proteins, hence acting as negative regulators of Rho protein function (22).
Rho-A is an important cellular GTPase that appears to mediate the effects of LPA on the phosphorylation of myosin light chain (23), multiple nuclear responses including cell cycle progression (2,3,24), and possibly malignant transformation (21). At least five Rho family proteins have been implicated as critical regulators of Ras-mediated oncogenic transformation (22), and recent observations indicate that aberrant function of Rho family proteins may also contribute to malignant transformation (21). An important aspect of the transformed phenotype is the ability of Rho-A to promote cell motility (25,26) and invasion. Thus, Rho protein function was found to be required for the LPA-induced invasion of hepatoma cells through a me- sothelial cell monolayer (24). Rho proteins are known to play an important role in the cytoskeletal processes that involve filamentous actin (27,28), including focal adhesion assembly and integrin-mediated signaling. In addition, Rho proteins play a role analogous to Ras in activating gene transcription. Rho, along with Rac and Cdc42, has been shown to activate SREdependent transcription (17), activate the transcription factor NFB (29,30), and facilitate expression of cyclins D-1 and E (31). The mechanism whereby extracellular stimuli, including LPA, activate Rho proteins remains largely unknown and may involve an interaction of the heterotrimeric G proteins with Rho guanine nucleotide exchange factors (32). Rho proteins presumably signal downstream via a family of Rho kinases (33,34), and, like the Ras proteins, they must be prenylated in order to activate their effectors (35).
Enhanced function of Rho proteins can result either from its excessive loading with GTP by guanine nucleotide exchange factors or from increased availability of geranylgeranylated Rho. The latter step is stimulated by hyperinsulinemia, which promotes the phosphorylation and activation of GGTase I (Fig.  6). The effect of insulin on GGTase I can be blocked by the GGTase I inhibitor, GGTI-286 (which was not due to a nonspecific effect of GGTI-286 as shown by the lack of its effect on LPA alone), or by inhibitors of hydroxymethylglutaryl-coenzyme A reductase (statins), which block cholesterol synthesis prior to the formation of the geranylgeranyl moiety. Insulin-induced increases in the availability of geranylgeranylated Rho lead to hyperactivation of Rho in response to its usual stimuli, resulting in enhanced nuclear responses of cell and tissues exposed to hyperinsulinemia.
Our current experiments in MCF-7 breast cancer cells indicate that insulin significantly increases GGTase I activity, the amounts of prenylated Rho proteins, and augments the Rhomediated transcriptional activity of LPA. Even though the effect of insulin on the amounts of prenylated Rho-A is modest, it translates into a 57% increase in the GTP loading of Rho-A and doubling (from 6-fold to 12-fold) of the nuclear effect of LPA (as well as nuclear effects of angiotensin II and advanced glycation end products in vascular smooth muscle cells). 2 Conceivably, insulin may also augment the amounts of prenylated Rac, another GTPase that is geranylgeranylated by GGTase I. Since Rac can also activate SRE, a portion of the "priming" effect of insulin may be Rac-mediated. In any event, this would still be a prenylation-dependent influence of hyperinsulinemia. Taken together, these results suggest that hyperinsulinemia can augment the Rho-dependent proliferative responses of cancer cells and, thereby, contribute to their progression.
In summary, our present and previous observations strongly suggest that insulin is a major activator of all three prenyltransferases (11,16,20). There is strong evidence that hyperinsulinemia may not be an innocent bystander, but contributes significantly to the progression of atherosclerosis and certain cancers. The "priming" effect of insulin on the Ras-and Rhodependent signaling pathways may be critical for the prolifer-ative responses of various tissues. The ability of insulin to potentiate the action of other growth factors may provide biochemical evidence and an explanation of the association of hyperinsulinemia with an increased incidence of atherosclerosis and cancer of the breast, colon, prostate, and endometrium.