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Cell Growth & Differentiation Vol. 10, 377-386, June 1999
© 1999 American Association for Cancer Research

The Wilms’ Tumor Suppressor, WT1, Inhibits 12-O-Tetradecanoylphorbol-13-acetate Activation of the Multidrug Resistance-1 Promoter1

Candice McCoy2, Sara B. McGee and Marilyn M. Cornwell3

Fred Hutchinson Cancer Research Center, Seattle, Washington 98109


    Abstract
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Overexpression of P-glycoprotein, the product of the multidrug resistance-1 (MDR1) gene, is associated with treatment failure in some hematopoietic tumors. Although expression of P-glycoprotein in normal hematopoietic cells is tightly regulated during hematopoietic differentiation, its aberrant overexpression in hematopoietic malignancies occurs at the transcriptional level. We have demonstrated that 12-O-tetradecanoylphorbol-13-acetate (TPA) increases transcription of the MDR1 gene and activates the MDR1 promoter, and that promoter activation by TPA requires binding of the zinc finger transcription factor EGR1 to specific MDR1 promoter sequences (C. McCoy and M. M. Cornwell, Mol. Cell. Biol., 15: 6100–6108, 1995). We demonstrate here that the Wilms’ tumor (WT) suppressor, WT1, a member of the EGR family, inhibits the response of the MDR1 promoter to TPA in K562 cells. Inhibition is likely a direct effect of WT1 binding to the MDR1 promoter because: (a) WT1 expression does not inhibit the increase in EGR1 after TPA treatment; (b) inhibition by WT1 requires the zinc finger domain; (c) WT1 binds to MDR1 promoter sequences that bind EGR1 and are responsive to TPA; and (d) there is an inverse correlation between WT1 protein expression and MDR1 expression and promoter activity. These results suggest that the MDR1 gene is a target for regulation by WT1 and suggest mechanisms by which MDR1 may be regulated by WT1 and EGR1 during normal and aberrant hematopoiesis.


    Introduction
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Chemotherapy resistance is the major cause of treatment failure in human tumors and expression of Pgp,4 the product of the MDR1 gene, is an important cause of such chemotherapy resistance. The exact mechanism by which Pgp confers resistance is unknown. However, it is thought to contribute to drug resistance, at least in part, by pumping hydrophobic chemotherapeutic agents out of the cell by an ATP-dependent mechanism (reviewed in Refs. 1 and 2 ). Its importance as a mechanism of resistance is underscored by the fact that several natural product-derived drugs are substrates for the Pgp pump; therefore, Pgp can confer resistance to many structurally different chemotherapeutic agents.

The best correlation of Pgp expression with treatment failure and clinical resistance to chemotherapy is observed in hematopoietic malignancies. These include acute myelogenous and lymphocytic leukemias, the blast phase of chronic myelogenous leukemia, multiple myeloma, and non-Hodgkin’s lymphoma (reviewed in Ref. 3 ). In AML, where Pgp expression occurs in 20% of de novo cases and {approx}75% of secondary cases (4, 5, 6, 7) , Pgp expression alone is a negative prognostic feature (Refs. 6, 7, 8, 9, 10, 11, 12 ; reviewed in Ref. 3 ). This is particularly true of AML in the elderly in which high MDR1 expression is associated with a poorer prognosis, independent of other poor prognostic features (13) . Pgp overexpression occurs more frequently in hematological tumors at relapse than at presentation (reviewed in Ref. 3 ), suggesting that Pgp-positive tumor cells are selected during chemotherapeutic treatment, that factors which control tumor progression regulate MDR1 gene expression, or both.

Pgp is also expressed in normal tissues, where its expression is largely limited to cells with a specialized secretory function. These include cells in renal proximal tubules, intestinal epithelia, biliary canaliculi, and pancreatic ducts, among others (reviewed in Refs. 1 and 2 ). Its exact function in these cells is unknown, although recent studies suggest that Pgp performs physiologically important secretory functions in some cells. The murine knockout of the MDR1 homologue mdr1a suggests that Pgp contributes to the blood-brain barrier (14) . A murine knockout of the MDR3 homologue, mdr2, suggests that Pgp has a secretory function in the biliary tract (15) . In support of the latter is that Pgp has recently been found to translocate a number of short-chain lipids, supporting its potential role as a lipid flippase (16) .

Pgp is also differentially expressed in subsets of normal hematopoietic cells (17, 18, 19, 20) . Levels of Pgp expression in normal hematopoietic cells are lower than those observed in hematopoietic malignancies, and its function in these cells is unknown. In the lymphoid lineage the highest Pgp expression is found in circulating CD8+ and CD56+ cells (18, 19, 20) . In the myeloid lineage, the highest expression is found in the CD34++, HLA-DR- subset with minimal expression in cells with lower CD34 expression or at later stages of differentiation (17, 18, 19, 20) . Although Pgp may play a role in in vitro lymphoid immune responses (21, 22, 23) , a role in in vivo lymphoid or myeloid function has not been established. Thus, although the functional role of Pgp in hematopoietic cells is not fully understood, its expression is tightly regulated during hematopoiesis.

Identifying the factors that regulate MDR1 gene expression in hematopoietic cells has been of interest because Pgp expression is regulated during hematopoiesis and because its association with treatment failure occurs primarily in hematopoietic tumors. Chemical agents, such as TPA, which induce phenotypic and morphological differentiation in hematopoietic cells, have also been shown to increase MDR1 gene expression and functional Pgp expression (24 , 25) . In K562 cells, TPA treatment increases MDR1promoter activity with a dose and time dependence that correlates with an increase in MDR1 gene expression (25) . We have shown previously that the zinc finger transcription factor EGR1 mediates TPA activation of the MDR1 gene (25) . EGR1 is a member of the EGR family of zinc finger transcription factors. EGR family members include EGR1, EGR2, EGR3, and EGR4 and the WT suppressor protein, WT1 (reviewed in Refs. 26 and 27 ). All EGR family members, including WT1, bind to the EGR consensus binding site (GCGGGGGCG; Refs. 26 and 27 ), although binding to alternative sites has been reported for both EGR1 (25 , 28, 29, 30, 31, 32) and WT1 (33, 34, 35, 36) . Both EGR1 and WT1 contain positive and negative regulatory elements (37 , 38) and are capable of activating or suppressing promoter activity. WT1 has been shown to repress in vitro transcription of a number of genes involved in growth promotion and differentiation (39, 40, 41, 42, 43, 44, 45, 46, 47, 48, 49) , and WT1 and EGR1 can exert opposing effects on promoter activity (44) . EGR1 binds to the MDR1 promoter TPA-responsive region, which contains sequences that are similar, but not identical, to the EGR consensus site (Ref. 25 ; Fig. 6Citation ). However, within this region, there are no sites identical to those reported previously to bind WT1.



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Fig. 6. The MDR1promoter overlapping the SP1/EGR site. Boxed area, the overlapping SP1/EGR site. Underlined area, sequence homology with the WTE site (36) . Mutant oligonucleotides and their ability to bind EGR1 in gel shift assays are illustrated. The wild-type oligonucleotides 59/58, 57/56, 54/53, and 49/47 are used as competitor oligonucleotides in Fig. 8Citation .

 
Both EGR1 and WT1 are expressed in hematopoietic cells and have been implicated in regulating hematopoietic differentiation. EGR1 is expressed in hematopoietic cells in response to growth factors (50 , 51) and restricts hematopoietic differentiation to the macrophage lineage in vitro (52 , 53) . WT1 gene expression in response to hematopoietic growth factors has not been well defined, but in vitro models of hematopoietic differentiation suggest that WT1 expression decreases under conditions that promote differentiation (54, 55, 56) . However, little is known about the genes regulated by WT1 during hematopoiesis.

We demonstrate in this work that WT1 inhibits the TPA-induced activation of the MDR1promoter in K562 cells. Inhibition is direct in that WT1 does not inhibit the increase in EGR1 after TPA treatment and inhibition requires the WT1 DNA binding domain. In the absence of TPA, WT1 activates the MDR1promoter in K562 cells, and activation also requires the zinc finger. Using EMSA, we have demonstrated that in vitro transcribed and translated WT1 binds to the MDR1promoter. WT1 RNA expression decreases after TPA treatment in K562 cells (54) . Here we demonstrate that WT1 protein expression decreases after TPA treatment. These findings support a model by which EGR1 and WT1 modulate MDR1promoter activity and suggest that EGR1 and WT1 may modulate Pgp expression during normal hematopoiesis and in hematopoietic tumors.


    Results
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
WT1 Inhibits TPA Activation of the MDR1 Promoter.
TPA activates a full-length MDR1promoter-luciferase reporter construct in K562 cells, and the time course of this activation corresponds to increases in steady-state MDR1 gene expression and Pgp expression (24 , 25) . We have demonstrated that activation by TPA of an MDR1promoter construct containing 69 bp upstream and 20 bp downstream of the start site is equal to that of the full-length promoter construct and is mediated by EGR1 binding to MDR1promoter sequences (25) . WT1 has been shown to inhibit EGR1-mediated transcriptional activation of promoter constructs containing EGR consensus sequences (44 , 57) . The TPA-responsive MDR1promoter region binds EGR1 in a GC-rich region that is similar to the EGR consensus site (Ref. 25 ; Fig. 6Citation ). Thus, we determined whether WT1 would inhibit activation of the MDR1promoter by TPA. K562 cells, which express low levels of WT1, were transiently cotransfected with the -69/+20-luciferase construct (Fig. 1A)Citation and WT1 expression vector or pCB6+ vector control and then treated with TPA or DMSO control. As described in "Materials and Methods," the pSV-ß-Gal construct was cotransfected with the other constructs as a control for transfection efficiency. Cells were then assayed for luciferase expression and ß-Gal expression at the time points indicated. Cotransfection of WT1 (Fig. 1C)Citation inhibited the response of -69/+20 to TPA, whereas cotransfection of pCB6+ vector did not (Fig. 1B)Citation . Inhibition of TPA activation by WT1 was dose dependent because TPA treatment resulted in decreasing luciferase activity as the amount of cotransfected CMV-WT1 was increased (Fig. 2)Citation .



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Fig. 1. WT1 inhibits activation of -69/+20 by TPA. A, the -69/+20 MDR1 promoter-luciferase construct. Arrow, transcription start site. B, K562 cells were cotransfected with -69/+20 and pCB6+ vector. The ratio in micrograms of -69/+20 to pCB6+ was 1:4. Cells were allowed to recover, then treated with TPA or DMSO control (-TPA lanes)for 10 h, were harvested and lysed, and luciferase activity was measured with normalization for ß-Gal expression as described in "Materials and Methods." Three experiments are shown; bars, SD. C, K562 cells were cotransfected with -69/+20 and CMV-WT1 at a ratio of 1:4, allowed to recover, harvested, and lysed, and luciferase activity was measured as above. Three experiments are shown; bars, SD.

 


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Fig. 2. Dose response of inhibition by WT1. K562 cells were cotransfected with the -69/+20 reporter construct and increasing amounts of CMV-WT1. The amount of pCB6+ vector in each transfection was varied to keep the amount of DNA constant in each transfection. The amount of each construct transfected, in micrograms, is indicated below the bar graph. Cells were treated with TPA or DMSO control (-TPA lanes). Bars, SD.

 
WT1 Activates the MDR1 Promoter.
During the course of the above experiments, it was noted that basal luciferase activity in log phase K562 cells transfected with CMV-WT1 and -69/+20 was consistently higher than that in cells transfected with the empty vector pCB6+ and -69/+20 (Fig. 2Citation , -TPA lanes). To investigate whether WT1 activates the MDR1promoter in K562 cells, we transfected K562 cells with the -69/+20 MDR1 promoter construct and CMV-WT1 or pCB6+ vector control. Cells were allowed to recover for 24–48 h and assayed for luciferase activity and ß-Gal expression. This experiment has been repeated several times, and the results of three such experiments are shown in Fig. 3Citation . Luciferase activity was consistently 2–8-fold higher in cells transfected with WT1 compared with pCB6+ control. Although this effect is modest, it is comparable with the levels of WT1 transactivation observed for other promoters (49 , 58) .



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Fig. 3. Activation of -69/+20 by CMV-WT1. K562 cells were transfected with -69/+20 and the CMV-WT1 expression vector at a ratio of 1:4. Cells were allowed to recover for 24 (Exp 1.) to 48 h (Exp. 2 and Exp. 3), harvested, and lysed, and luciferase activity was measured with normalization for ß-Gal expression as described in "Materials and Methods." Bars, SD.

 
Expression of WT1 Does Not Inhibit the Increase in EGR1 Seen after TPA Treatment.
Because the EGR1 promoter contains an EGR1 consensus site, we thought it possible that WT1 inhibits TPA-induced EGR1 transcription and thereby has an indirect effect on MDR1induction by TPA. To investigate this possibility, we used Western analysis to measure EGR1 protein levels after TPA treatment of K562 cells. K562 cells were transiently transfected with -69/+20 and CMV-WT1 or pCB6+ vector control and then treated with TPA or DMSO control. Nuclear extracts were made from half of the cells, and aliquots of these nuclear extracts were electrophoresed using SDS-PAGE. Proteins were transferred to nylon membranes, and Western analysis was performed with anti-EGR1 or anti-WT1 antibodies to determine the level of expression of these proteins after TPA treatment. Luciferase assays were performed with the other half of the transfected cells to determine whether a correlation existed between protein expression and luciferase activity. As shown in Fig. 4ACitation , transient cotransfection of CMV-WT1 did not inhibit the increase in EGR1 protein expression seen after TPA treatment. However, activation of -69/+20 by TPA was inhibited, as shown in Fig. 4BCitation . This suggests that WT1 does not inhibit TPA activation of the MDR1promoter indirectly by inhibiting the increase in EGR1 protein expression.



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Fig. 4. WT1 does not inhibit MDR1promoter activation by TPA by inhibiting the increase in EGR1. K562 cells were cotransfected with -69/+20 and either pCB6+ vector control or CMV-WT1. The ratio in micrograms of -69/+20 to the cotransfected constructs was 1:6. Cells were allowed to recover and then treated with TPA or DMSO control (-TPA lanes) for 10 h. A, nuclear extracts were made from half of the cells, and aliquots of these extracts were used in Western analysis with anti-WT1 and anti-EGR1 antibodies. B, the other half of the cells were harvested and lysed, and luciferase activity was measured with normalization for ß-Gal expression as described in "Materials and Methods." Fold activation is shown below each transfection. Lanes 1 and 2 are control lanes containing 5 µg of nuclear extract from K562 cells transfected with pCB6+ vector or CMV-WT1, respectively. Lane 3 contains ITT-EGR1. Lanes 4–7 contain 10 µg of nuclear extract from K562 cells cotransfected with -69/+20 and the indicated construct, then treated with DMSO control or TPA. Lane 4, -69/+20 and pCB6+, DMSO control; Lane 5, -69/+20 and pCB6+, TPA; Lane 6, -69/+20 and CMV-WT1, DMSO control; Lane 7, -69/+20 and CMV-WT1, TPA.

 
The WT1 Zinc Finger Is Necessary to Inhibit the TPA Response.
Like the other EGR family members, the WT1 zinc finger region is necessary for DNA binding. A truncated WT1 protein, WT1{Delta}297-429, lacks the entire zinc finger region and is unable to bind DNA (59) . In transiently transfected K562 cells, WT1{Delta}297-429 is expressed at levels comparable with wild-type WT1 (data not shown). To determine whether WT1 inhibited the response of the MDR1promoter to TPA directly by binding to the MDR1promoter, we determined whether WT1 mutants lacking the zinc finger would inhibit the TPA response. We transiently cotransfected K562 cells with -69/+20 and CMV-WT1, CMV-WT1{Delta}297-429, or vector control. Cells were then treated with TPA or DMSO control. The results of a representative experiment are shown in Fig. 5Citation . The luciferase activity in the samples cotransfected with vector control was approximately equal to that of samples cotransfected with CMV-WT1{Delta}297-429. These results suggest that DNA binding by WT1 is necessary for inhibition of the MDR1promoter response to TPA and are consistent with previous observations that the WT1 zinc finger domain is necessary for the inhibitory function of WT1 (57 , 59) . The WT1 zinc finger is also necessary for activation of -69/+20 (data not shown).



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Fig. 5. CMV-WT1{Delta}297-429 does not inhibit MDR1 promoter activation by TPA. K562 cells were cotransfected with -69/+20 and either pCB6+ vector control, CMV-WT1, or CMV-WT1{Delta}297-429. The ratio in micrograms of -69/+20 to the cotransfected constructs was 1:4. Cells were allowed to recover, then treated with TPA or DMSO control (-TPA lanes) for 10 h, were harvested and lysed, and luciferase activity was measured with normalization for ß-Gal expression as described in "Materials and Methods." The experiment was repeated three times, each time with different plasmid preparations, with similar results. One such experiment is shown. Bars, SD.

 
WT1 Binds to the MDR1Promoter.
The above results suggested that WT1 binds to MDR1promoter sequences between -69 and +20 bp of the transcription start site. Binding of SP1 and EGR1 to this region has been characterized previously by footprint and promoter mutation analysis (25 , 60 , 61) and is localized to sequences between -69 and -39 relative to the start site (Fig. 6)Citation . The high GC content and the fact that there exists some sequence similarity to the WTE site (Ref. 36 ; 5'-GCGTGGGAGT-3', Fig. 6Citation , underlined) suggested that WT1 might bind to this region. To determine whether WT1 bound to MDR1promoter TPA-responsive sequences, we performed EMSA with in vitro transcribed and translated WT1 and radioactively labeled -69/+20. As shown in Fig. 7ACitation , a complex was observed (left arrow) in Lane 3 containing ITT WT1. No binding was observed in Lane 2 containing unprogrammed reticulocyte lysate. The specificity of binding was confirmed using oligonucleotide inhibition. Binding was inhibited by -69/+20 (Fig. 7ACitation , Lanes 4 and 5) and by a double-stranded oligonucleotide corresponding to an EGR consensus site (Fig. 7Citation , Lanes 6 and 7).



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Fig. 7. WT1 binds to the MDR1 promoter. A, ITT WT1 was produced in a reticulocyte lysate system, and EMSA was performed, with (Lanes 4–7) or without (Lanes 1–3) unlabeled oligonucleotide competitor, as described in "Materials and Methods." Lane 1, free probe; Lane 2, unprogrammed reticulocyte lysate; Lane 3, ITT WT1; Lanes 4 and 5, ITT WT1 and a 10x or 100x molar excess of unlabeled -69/+20 probe, respectively; Lanes 6 and 7, 10x or 100x molar excess of unlabeled EGR1 consensus oligonucleotide, respectively. B, EMSA was performed in the absence (Lanes 1–3) or presence (Lanes 4–7) of anti-WT1 antibody as described in "Materials and Methods." All of the lanes were from the same polyacrylamide gel except that Lane 1 was separated from Lanes 2–7 by lanes containing information not relevant to the figure. Lane 1, free probe; Lane 2, unprogrammed reticulocyte lysate; Lane 3, ITT-WT1; Lanes 4 and 5, 2 or 4 µl of anti-WT1 antibody SC180, respectively; Lanes 6 and 7, 2 or 4 µl or anti-WT1 antibody C-19, respectively. Arrowsindicate WT1 bound to probe.

 
To determine whether the protein bound in Lane 3 was WT1, EMSA was performed in the presence of anti-WT1 antibodies sc180 and C19 (Fig. 7B)Citation . Binding of ITT WT1 was observed in Lane 3 (left arrow). Binding was not observed in the presence of antibody sc180, which recognizes the WT1 NH2 terminal domain (Fig. 7BCitation , Lanes 4 and 5). A more slowly migrating complex was observed in the presence of C19 antibody, which recognizes the WT1 COOH-terminal domain (Fig. 7BCitation , Lanes 6 and 7, right arrow). These data suggest that the complex observed in Fig. 7ACitation , Lane 3, and Fig. 7BCitation , Lane 3, is formed by WT1 binding to MDR1 promoter sequences between -69 and +20. The faint band observed in lanes containing unprogrammed lysate (Fig. 7BCitation , Lane 2) and those containing antibodies (Fig. 7BCitation , Lanes 4–7) is nonspecific. It was observed with some preparations of ITT WT1 and was dependent on the lot of reticulocyte lysate, not on the plasmid preparation.

We have previously characterized oligonucleotides containing mutations in the overlapping SP1/EGR binding region for their ability to bind SP1 and EGR1 (25 , 61) and have found only one mutation that binds SP1 but does not bind EGR1. EGR1 binding to these oligonucleotides is summarized in Fig. 6Citation . To determine whether WT1 binding sequences in the MDR1promoter overlap with those of EGR1, we determined whether the addition of cold mutant oligonucleotides to EMSA with -69/+20 would similarly alter WT1 binding. As shown in Fig. 8ACitation , ITT EGR1 binding was inhibited in a dose-dependent fashion by an oligonucleotide containing wild-type sequences from -69 to -39, the 59/58, 57/56 mutant oligonucleotides, and an EGR consensus oligonucleotide. EGR1 binding was only minimally inhibited by the 54/53 and 49/47 mutant oligonucleotides and was not inhibited by an AP1 consensus site oligonucleotide. Likewise, in Fig. 8BCitation , WT1 binding was inhibited by the wild-type, 59/58, 57/56, and the EGR consensus site oligonucleotides but was not inhibited by 54/53, 49/47, or the AP1 consensus oligonucleotide. Therefore, binding of WT1 to the MDR1promoter involves the region between -69 and -39, which also binds SP1 and EGR1. Mutations in this region that alter EGR1 binding similarly alter WT1 binding, suggesting that the binding site for WT1 overlaps that of EGR1.



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Fig. 8. WT1 binding specificity parallels that of EGR1. ITT EGR1 (A) or ITT WT1 (B) were made and used in EMSA as described in "Materials and Methods" in the presence (Lanes 4–10) or absence (Lanes 1–3) of unlabeled oligonucleotide competitors. The sequence of each competitor containing wild-type or mutated MDR1 promoter sequences is illustrated in Fig. 6.ACitation , ITT EGR1. Lane 1, free probe; Lane 2, unprogrammed reticulocyte lysate; Lane 3, ITT EGR1. Duplicate Lanes 4–10 contain 25x and 100x molar excess (with respect to -69/+20) of unlabeled oligonucleotide competitor. Note that in Lane 4, the 100x and 25x competitors are in reverse order. B, ITT WT1. Lanes 1 and 2, as in A; Lane 3, ITT WT1; Lanes 4–10, as in A. Arrowsindicate EGR1 (A) and WT1 (B) bound to probe.

 
Reciprocal Relationship between WT1 and MDR1 Expression after TPA Treatment.
It has been demonstrated by Phelan et al. (54) that WT1 RNA expression decreases in K562 cells after TPA treatment. However, that study did not address changes in WT1 protein expression after TPA treatment. To determine how WT1 contributes to regulating the TPA response in K562 cells, we used Western analysis of nuclear extracts from log phase or TPA-treated K562 cells to determine the level of endogenous WT1 protein expressed in K562 cells before and after TPA treatment. Although WT1 protein was expressed in log phase K562 cells, its expression diminished after TPA treatment (Fig. 9)Citation .



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Fig. 9. WT1 expression decreases in K562 cells after TPA treatment. Fifty µg of nuclear extract from untreated (log phase) K562 cells and K562 cells treated with TPA for 24 h or 10 µg of nuclear extract from K562 cells transfected with pCB6+ vector, CMV-EGR1, or CMV-WT1 were electrophoresed on a 10% SDS-polyacrylamide gel and transferred to nitrocellulose filters for immunodetection as described in "Materials and Methods." Exposure to film was 12 min for the anti-WT1 blot and 1 min for the anti-SP1 blot.

 

    Discussion
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Understanding the factors that regulate MDR1expression is important to better understand the role of MDR1in normal and abnormal hematopoiesis and to develop new means of modulating chemotherapy resistance in human tumors. We have demonstrated that WT1 inhibits the response of the MDR1promoter to TPA and that, in the absence of TPA, WT1 activates the MDR1promoter. Inhibition of the -69/+20 response to TPA and activation of -69/+20 requires the WT1 zinc finger, which suggests that DNA binding by WT1 is necessary for both. WT1 does not indirectly inhibit the MDR1 promoter response to TPA by inhibiting the increase in EGR1 after TPA treatment. Finally, WT1 binds to the MDR1promoter, suggesting that WT1 protein directly influences MDR1 promoter activity through DNA binding. In K562 cells, WT1 protein levels decrease after TPA treatment as EGR1 protein levels, EGR1 binding, steady-state MDR1expression (25) , and P-glycoprotein expression (24) increase. These findings suggest that in K562 cells, the up-regulation of MDR1expression after TPA treatment occurs in part by down-regulating WT1 expression, thereby overcoming the normally inhibitory effects of endogenous WT1. These findings also suggest that WT1 may suppress MDR1 gene transactivation in response to some cellular signals but may also activate the MDR1 gene, depending on the cellular context.

The mechanism by which WT1 inhibits MDR1promoter activation by TPA is likely by binding directly to MDR1promoter sequences. In support of this hypothesis: (a) the MDR1 promoter response to TPA is mediated by EGR1 binding to promoter sequences; (b) the WT1 zinc finger is required for inhibition of the TPA response; and (c) WT1 binds to the MDR1promoter sequences, which also bind EGR1. We have not directly addressed here whether WT1 binding inhibits MDR1promoter activation by TPA by interfering with EGR1 binding, although competition for binding between EGR1 and WT1 and between SP1 and WT1 has been reported (43 , 49) . Another way to address this is to use mutant MDR1 promoter constructs in transient transfection assays to determine which mutations abrogate the effect of WT1 on the TPA response. However, the mutations we have identified that inhibit WT1 binding also inhibit EGR1 binding. If a mutation inhibits EGR1 binding, it inhibits the MDR1promoter response to TPA (25) ; therefore, an effect of WT1 on this response cannot be measured.

The results presented here do not fully explain the mechanism by which WT1 on the one hand inhibits activation of the MDR1promoter by TPA and on the other hand has a modest stimulatory effect in the absence of TPA. Although promoter architecture has been shown to modulate the effect of WT1 on transcription (34 , 38 , 62) , our results are consistent with those of others demonstrating that cellular context can also modify the effect of WT1 on transcription without modification of promoter architecture (49) . One possible explanation for this is that one or both of the effects we observed of WT1 on MDR1promoter activity requires interaction by WT1 with another protein, the expression of which is modulated by TPA. In support of this is that WT1 interacts with other proteins such as p53 and par-4 (63, 64, 65) , and such interactions can modulate the effect of WT1 on transcriptional regulation (63 , 64) . In our preliminary experiments, neither the presence nor absence of functional p53, nor the expression of par-4, had any effect on WT1 transactivation of the MDR1promoter in transfected cells.5 Thus, the functional consequences of these particular interactions may not be observed in all cell types or with all promoters.

Another possible explanation for the differential action of WT1 on the MDR1promoter is TPA-induced posttranslational modification of an interacting protein or of WT1 itself. Phosphorylation of WT1 occurs after activation of protein kinase C (66) . Such phosphorylation inhibits WT1 binding to DNA in vitro, inhibits its suppression of a Krox-24 promoter construct, and is associated with its sequestration in the cytoplasm (66) . It is unlikely that phosphorylation of WT1 modulates its effect on the MDR1promoter: (a) the results presented here suggest that WT1 binding to the MDR1promoter is necessary for suppression of the TPA response, whereas in the previous study, phosphorylation inhibited WT1 binding to DNA; (b) binding to the MDR1promoter requires nuclear localization, whereas in the previous study, phosphorylation of WT1 was associated with cytoplasmic, not nuclear, localization; and (c) we did not detect a shift in the molecular weight of WT1 after TPA treatment (Fig. 4Citation , Lane 7), whereas such a shift was observed in the previous study. Taken together, our results are not consistent with phosphorylation mediating the differential effects of WT1 on the MDR1promoter.

Correlative studies of Pgp and WT1 expression in normal hematopoietic cells may ultimately contribute to a better understanding of how WT1 regulates MDR1gene expression in vivo. However, such studies will be technically difficult, and it is not presently known whether a relationship between WT1 and MDR1expression exists during normal hematopoiesis: (a) whether the WT1 gene is expressed in normal hematopoietic cells is controversial (56 , 67, 68, 69, 70, 71) ; (b) WT1 gene expression may be heterogeneous within the CD34+ population (72) , may vary over time, and/or may be limited to CD34+ subpopulations; and (c) whether WT1 protein is expressed in hematopoietic progenitors is controversial (72 , 73) , with conflicting results potentially due to the use of differing CD34+ cell sources and experimental methods. Thus, because Pgp expression can also vary within the CD34+ population (17 , 19) , studies examining the coexpression of Pgp and WT1 will need to focus on protein, not gene, expression and single cells, not populations.

In some acute leukemias, a high level of WT1 transcript expression has been observed (71 , 73, 74, 75, 76, 77, 78, 79) . In addition, the expression of mutant WT1 alleles encoding a truncated WT1 protein with impaired DNA binding has been reported (80) . The expression of such truncated WT1 protein may exert a dominant-negative effect on gene expression mediated by wild-type WT1 expressed from the other allele (81 , 82) . Thus, studies that attempt to correlate WT1 and Pgp expression in leukemias will need to examine whether truncated, potentially dominant-negative WT1 proteins are expressed. Only one study has examined the frequency of mutant WT1 gene expression in sporadic acute leukemia (80) . The WT1 gene was expressed in >70% of the 36 cases examined, and five WT1 mutations, predicted to produce a nonbinding WT1 protein, were found in 4 patients (80) . Three of these patients had AML and failed to respond to standard chemotherapy, despite favorable prognostic features. The expression of many genes is likely to be disrupted in these leukemias by expression of a dominant-negative WT1 protein. However, we would like to speculate that high MDR1 gene expression would be facilitated in this setting, thus contributing to the poor response to chemotherapy observed in these patients.


    Materials and Methods
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Reagents and Antibodies.
TPA was purchased from Sigma Chemical Co. and stored at -20 at 10 mg/ml in DMSO. Before adding it to cultured cells, this TPA solution was diluted further in RPMI or Iscove’s medium so that DMSO concentrations were <0.01%. Poly(deoxyinosinic-deoxycytidylic acid) was purchased from Pharmacia. [32P]ATP (6000 Ci/mmol) was purchased from DuPont NEN. Polyclonal antibodies specific for human SP1, EGR1, and WT1 were purchased from Santa Cruz Biotechnology. Mouse monoclonal antibody to WT1 was the kind gift of F. Rauscher III (Wistar Institute, Philadelphia, PA). This antibody detects an epitope within the first 80 amino acids of WT1.

Cell Culture.
The K562 cell line was the kind gift of S. J. Collins (Fred Hutchinson Cancer Research Center). K562 cells were maintained in RPMI 1640 supplemented with 10% calf serum, 1 mM glutamine, and 25 µg (each) streptomycin and penicillin per ml (complete media) in a humidified incubator containing 5% CO2.

Expression Vectors and Oligonucleotides.
The -434-MDR1-luciferase and -69/+20 reporter plasmids have been described previously (25) . The CMV-WT1 construct containing the full-length WT1 cDNA under control of the CMV promoter has been described previously (39) and was the kind gift of F. Rauscher III (Wistar Institute). The CMV-WT1{Delta}297–429 construct has been described previously (59) and was the kind gift of T. Deuel (Beth Israel Hospital, Boston, MA). The pos8WT1 construct was the kind gift of T. Deuel (Beth Israel Hospital) and contains the full-length WT1 cDNA cloned into the EcoRI site of the pTM1-derived vector, pos8. ITT WT1 was expressed from this construct using the TNT coupled in vitro transcription-translation system with T7 RNA polymerase (Promega). ITT EGR1 was made from a construct containing the full-length EGR1 cDNA as described previously (25) .

Oligonucleotides used in Fig. 8Citation are illustrated in Fig. 6Citation . The oligonucleotides contain mutations in the wild-type MDR1 promoter nucleotides between -69 and -47. All mutations are illustrated in Fig. 6Citation and replace a wild-type G with T, or C with A at the residues specified in the name of the mutant oligonucleotide. The oligonucleotides containing EGR and AP1 consensus sequences (underlined) are: EGR, 5'-GCCAACGCCCCCGCAACCG-3' and AP1, 5'-GGATGTTATAAAGCATGAGTCAGACACCTCTGGT-3'.

Transient Transfection.
Transient transfection of K562 cells was performed by electroporation as follows. A DNA mixture containing reporter and effector plasmids, 2–4 µg of pSV-ß-Gal plasmid as an internal control for transfection efficiency, 5 µg/ml DEAE dextran, and TE (10 mM Tris HCl, 2 mM EDTA, pH 8.0) was made to bring the volume to 50 µl/transfection. The amount of luciferase reporter construct transfected was 10 µg unless otherwise specified. Each transfection was performed in duplicate, and each experiment was performed at least three times with different plasmid preparations. K562 cells growing in logarithmic phase were collected by brief centrifugation and resuspended in complete media at 107 cells/ml. Fifty µl of DNA mixture and 500 µl of cells were added to each electroporation cuvette (BTX), incubated for 5 min at room temperature, and placed in a Gene Pulsor apparatus (Bio-Rad). Current was applied at 260 V and 960 µF capacitance. Cuvettes were then immediately put on ice, and contents were transferred to a 60-mm culture dish containing 5 ml of Iscove’s medium with 10% calf serum, 1 mM glutamine, and 25 µg (each) of penicillin and streptomycin. After a recovery period of 12–24 h, cells were treated with a 1:1000 dilution of DMSO or 16 nM (10 µg/ml) TPA, and cells were harvested after incubation at 37°C for the period of time indicated. Cells were harvested by centrifugation and assayed for luciferase activity and ß-Gal activity as described previously (25) , with ß-Gal activity being used to normalize the luciferase activity for differences in transfection efficiency.

Nuclear Extracts.
Nuclear extracts from cells transfected with pCB6+ vector or CMV-WT1 in Figs. 4Citation and 9Citation were made as per Andrews and Faller (83) , with minor modifications. After transfection, K562 cells were allowed to recover at 37°C for 48 h. Approximately 107 cells were quickly centrifuged at 4°C, washed once in PBS, and centrifuged again. Cells were resuspended in ice-cold buffer A [10 mM HEPES-KOH (pH 7.9) at 4°C, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT, 0.2 mM phenylmethylsulfonyl fluoride, 2.5 µg/ml leupeptin, and 2.5 µg/ml trypsin inhibitor], allowed to swell on ice, vortexed, and centrifuged. The nuclei were then suspended in buffer C [20 mM HEPES-KOH (pH 7.9) at 4°C, 25% glycerol, 42 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 0.2 mM phenylmethylsulfonyl fluoride, 2.5 µg/ml leupeptin, and 2.5 µg/ml trypsin inhibitor], incubated on ice for 20 min, and centrifuged for 2 min. Supernatants were stored in aliquots at -70°C. The nuclear extracts from log phase or TPA-treated K562 cells in Fig. 9Citation were made as described previously (25) .

EMSA.
The -69/+20 double-stranded probe was made by PCR and end labeled with [{gamma}-32P]ATP and T4 polynucleotide kinase by standard methods. EMSA was performed by incubating unprogrammed reticulocyte lysate or ITT protein with 1.5 ng (0.0263 pmol) of 32P-labeled -69/+20 at room temperature in a reaction mix containing 12 mM HEPES (pH 7.5), 42 mM KCl, 3 mM MgCl2, 60 µM ZnSO4, 0.3 mM DTT, 50 µg poly (dI-dC), 0.03% NP40, and 7.2% glycerol. Samples were electrophoresed on a 5% acrylamide gel (19:1 bis) in 0.5x Tris-borate-EDTA. Gels were dried, and bands were visualized by autoradiography (Figs. 7Citation and 8ACitation ) or phosphorimaging (Fig. 8B)Citation . In Fig. 7BCitation , reactions, including -69/+20 probe, were incubated with or without the addition of anti-WT1 antibodies for 1 h at room temperature. In Fig. 8Citation , ITT protein was coincubated with or without competitor oligonucleotide for 10 min before the addition of probe, followed by a 20-min incubation at room temperature.

SDS-PAGE and Immunoblotting.
Nuclear extracts were loaded on 10% SDS-polyacrylamide gels (Bio-Rad), electrophoresed, and transferred to nitrocellulose membranes (Schleicher and Schuell). Membranes were incubated at room temperature in TBS containing 5–6% (w/v) nonfat dry milk. In Fig. 4Citation , the blot was incubated with anti-WT1 antibody diluted 1:500 in 6% dry milk in TBS, washed 2 h in TBS-1% Tween, then incubated for 2 h in a 1:500 dilution of horseradish peroxidase-conjugated anti-mouse IgG in 6% dry milk in TBS-0.5% Tween. Final wash was in TBS-1% Tween overnight. Detection was with enhanced chemiluminescence reagents (Amersham International) and Hyperfilm ECL (Amersham). The blot was then stripped for 30 min at 50°C in 100 mM ß-mercaptoethanol, 2% SDS, and 62.5 mM Tris-Cl (pH 6.7) and blocked in TBS-6% dry milk for 2 h before being incubated with anti-EGR1 antibody C-19 (Santa Cruz Biotechnology) at 1 µg/ml for 1.5 h. Blot was then washed for 1 h in TBS-1% Tween, incubated with horseradish peroxidase-conjugated anti-rabbit IgG, washed 1.5 h as above, and detected as above. The blots in Fig. 9Citation were duplicate blots and were blocked in TBS-5% dry milk overnight before being incubated for 4 h in anti-WT1 antibody diluted 1:500 or anti-SP1 antibody diluted 1:1000 in TBS-5% dry milk. Blots were washed in TBS-0.5% Tween, followed by TBS-1% Tween, and then incubated for 2 h in a 1:500 dilution of horseradish peroxidase-conjugated anti-mouse IgG (anti-WT1) or anti-rabbit IgG (anti-SP1) in 5% dry milk in TBS-0.5% Tween. Final wash was in TBS-1% Tween overnight. Detection was as above except with Biomax MR film (Kodak).


    Acknowledgments
 
We thank F. Rauscher III for the CMV-WT1 construct and vector control, as well as the monoclonal anti-WT1 antibody. We thank T. Deuel for providing the CMV-WT1 297-429 and pos8WT1 constructs. We also thank M. Ihnat and S. Kane for comments on the manuscript.


    Footnotes
 
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.

1 Supported in part by NIH Clinical Investigator Development Award K08 CA 68003 (to C. M.) and NIH Grant R01 CA 51728 (to M. M. C.). Back

2 Present address: Immunex Corporation, 51 University, Seattle, WA 98101. Back

3 To whom requests for reprints should be addressed, at Fred Hutchinson Cancer Research Center, 1100 Fairview Avenue North, D2-100, Seattle, WA 98109. Phone: (206) 667-6166; Fax: (206) 667-6600. Back

4 The abbreviations used are: Pgp, P-glycoprotein; MDR, multidrug resistance; WT, Wilms’ tumor; AML, acute myelogenous leukemia; TPA, 12-O-tetradecanoylphorbol-13-acetate; EGR, early growth response; ITT, in vitro transcribed and translated; EMSA, electrophoretic mobility shift assay; ß-Gal, ß-galactosidase; CMV, cytomegalovirus. Back

5 C. McCoy, unpublished observations. Back

Received for publication 1/ 9/98. Revision received 1/29/99. Accepted for publication 4/ 5/99.


    References
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 

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