the cystic fibrosis transmembrane conductance regulator (CFTR) is a cAMP- and protein kinase-regulated chloride channel found on the apical membranes of polarized epithelial cells (10). Mutations within this gene lead to the CF phenotype that ultimately results in premature death primarily from a combination of chronic lung infections, loss of pulmonary function, and pancreatic insufficiency (5). This genetic disease is among the most prevalent in the human population, occurring in ∼1 of 2,500 Caucasian live births. CFTR is a member of the structurally related ATP-binding cassette family of transmembrane transport proteins whose other members include P-glycoprotein (Pgp, the MDR1 gene product), multidrug resistance-associated protein and other related multidrug resistance proteins, and the sulfonylurea receptor (21).

Of the more than 850 individual CFTR mutations leading to the CF phenotype that have been described, the most important is ΔF508 (phenylalanine deletion at amino acid position 508), which is seen in ∼70% of all CF patients. The ΔF508-CFTR mutation results in improper folding and trafficking of the CFTR protein, leading to its degradation in the endoplasmic reticulum by the 26S proteosome machinery of the cell (23). However, importantly, if this mutant protein is folded and expressed in the membrane by experimental cell culture techniques such as glycerol (20) or low temperature treatments (3), it functions normally as a chloride channel. Moreover, it is estimated that restoration of functional CFTR expression to ∼10% of normal levels would be sufficient to ameliorate the symptoms of the disease in vivo (19). Thus there is great interest in development of strategies that can enhance ΔF508-CFTR cell surface expression, which may be clinically useful in treatment of CF patients.

There is evidence of coregulation of Pgp and CFTR in epithelial cells, although the mechanism and level of this overlapping regulation is currently unknown (2). Previous work in our laboratory demonstrated both transcriptional and posttranscriptional effects of low-dose, noncytotoxic treatments with the cancer chemotherapy drugs mitomycin C (MMC) and doxorubicin (Dox) on expression of Pgp in various cell lines and in a mouse in vivo model (6, 7). We therefore hypothesized that CFTR expression might also be modified by these or related drugs under similar treatment conditions. We recently demonstrated that MMC increased total protein and cell surface functional expression of wild-type CFTR in HT-29 and T84 human colon cells (15a). In this study, we investigated the effects of Dox on CFTR and ΔF508-CFTR expression. We report here that low-dose Dox (0.25 μM) significantly increased functional cell surface expression of CFTR and ΔF508-CFTR in T84 and Madin-Darby canine kidney (MDCK) cells, respectively, and that this was primarily a result of posttranscriptional effects.

METHODS

Cell culture and treatment.

Human colon adenocarcinoma T84 cells (American Type Culture Collection, Manassas, VA) were maintained in DMEM/F-12 media (Life Technology, Rockville, MD) containing l-glutamine supplemented with 10% fetal bovine serum (FBS) and antibiotics (Life Technology). An isogenic line of MDCK/C-7 cells stably expressing green fluorescent protein (GFP) tagged to the NH₂ terminus of ΔF508-CFTR (MDCK/ΔF508) were created in a fashion similar to a cell line expressing GFP tagged to the NH₂ terminus of CFTR (MDCK/CFTR) described elsewhere (17, 18). This cell line and parental MDCK-C7 cells were maintained in minimum essential media (MEM, Life Technology) supplemented as above. Additionally, 0.3 mg/ml of G418 Sulfate (Sigma, St. Louis, MO) was added to the MDCK/ΔF508 media for maintaining the selection pressure. For immunoblot analysis and RT-PCR following drug treatment, cells were grown in six-well plates (Costar, Cambridge, MA) to confluence, washed with PBS, and treated with or without 0.25 μM Dox (Sigma). After this, completed media were replenished and cells were harvested at various time points. For short-circuit current measurements and cell-surface biotinylations, cells were cultured on Transwell-type filters (Corning, Ithaca, NY). The medium was completely changed every 24 h, and experiments were performed 14 days postseeding.

Relative RT-PCR.

Total RNA from T84 cells was isolated using TRIAZOL reagent (Life Technology). One microgram of total RNA from T84 cells was reverse-transcribed using the RNA PCR Core Kit (Perkin Elmer, Foster City, CA) in 20 μl total volume. The cDNAs were divided into two equal aliquots and amplified using 0.5 μl of AmpliTaq DNA polymerase (5 U/μl), 20 pM upper and lower primers specific for human CFTR (upper: 5′-CCT GAA CCT GAT GAC ACA CT-3′, lower: 5′-TAA AAC TGC GAC AAC TGC TA-3′) or 1.25 μl of 5 μM 18S RNA specific primers and Competimers (Ambion, Austin, TX) in a ratio 3.5:6.5. Prior titration experiments ensured that amplification of both CFTR and 18S RNA products were within linear range. The cDNA products were separated by electrophoresis on 1% agarose gels (Life Technology) and stained with ethidium bromide to confirm proper product sizes.

Immunoblotting.

Cells were treated with solvent or Dox, washed once with cold PBS, and scraped in a small volume of lysis buffer (50 mM Tris · HCl, pH 6.8, 150 mM NaCl, 1% Nonidet P-40) containing a protease inhibitor cocktail (Roche Biochemicals, Indianapolis, IN). Protein was extracted from the samples by a 30-min incubation on ice with intermediate vortexing and frozen at −70°C. Protein in each sample was quantified by the bichinchoninic acid assay (Pierce, Rockford, IL). SDS-PAGE was performed on 4–15% minigels using 50 μg of protein lysate per lane. Proteins were separated and transferred to polyvinylidene difluoride membrane (Millipore, Bedford, MA) at 120 mA for 1 h in Towbin's transfer buffer (25 mM Tris, 192 mM glycine, 12% methanol). After transfer, the blots were blocked and probed sequentially, first with polyclonal A2 rabbit anti-CFTR antisera (25) (generously provided by Dr. W. Skach, Oregon Health Sciences Institute) overnight at 4°C and then with a horseradish peroxidase (HRP)-labeled anti-rabbit secondary polyclonal antibody (Amersham, Piscataway, NJ). Membranes were washed six times in PBS + 0.3% Tween 20 (PBST) at room temperature for 10 min each in between. The blots were developed by enhanced chemiluminescence (ECL) Blaze substrate (Pierce) and exposed to film.

Cell-surface biotinylation and immunoprecipitation.

T84 cells were biotinylated on the apical plasma membrane with a commercially available biotinylation kit (Roche Biochemicals). After treatment at 4°C with LC-(+)-biotin-hydrazide, cells were washed with cold PBS and harvested in lysis buffer on ice, and total protein in each sample was quantified as described above. Two hundred micrograms of protein from each sample was immunoprecipitated using polyclonal A2 rabbit anti-CFTR antisera (1:1,000) and Protein A-agarose (Roche Biochemicals). The beads were washed in PBST, and immunoreactive proteins were separated by SDS-PAGE and detected by streptavidin-HRP and ECL as described above. The ΔF508-CFTR-expressing MDCK cells were immunoprecipitated similarly but immunoblotted using a monoclonal anti-GFP antibody (Clontech Laboratories, Palo Alto, CA). In other experiments, CFTR was immunoprecipitated with A2 as described above and detected with the M3A7 monoclonal anti-CFTR antibody (12), a generous gift of Dr. J. Riordan (Mayo Institute, Scottsdale, AZ).

Measurement of short circuit current.

Short-circuit current (I_sc) was measured across monolayers of T84 cells grown on filters. Cells were treated with Dox (0.25 μM, 24 h) beginning at 14 days after seeding, and the media were changed to remove residual drug beforeI_sc measurement. I_sc was measured in cells in the presence of amiloride (10⁻⁵ M) in the apical solution to block the potential contribution of sodium transport to I_sc. In the presence of amiloride, 8-(4-chlorophenylthio)-cAMP (CPT-cAMP)-stimulatedI_sc in T84 cells is equivalent to electrogenic Cl⁻ secretion (1). The CFTR antagonist diphenylamino carboxylic acid (DPC) and the Cl⁻/HCO${}_3^{-}$ exchange inhibitor 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (DIDS) were added as previously described (1, 17). Bath solutions were maintained at 37°C and were stirred by bubbling with 5% CO₂-95% air. Current output from the voltage clamp was digitized by a TL-1 DMA interface analog-to-digital converter (Axon Instruments, Foster City, CA). Data collection and analysis were done with Axotape 2.0 software (Axon Instruments). During experiments, cells were bathed in a FBS-free MEM solution containing (in mM) 116 NaCl, 116 NaHCO₃, 3 KCl, 2 MgCl₂, 0.5 CaCl₂, 3.6 Na-HEPES, 4.4 H-HEPES (pH 7.4), and 10 glucose.

Fluorescent chloride efflux assay.

Cells were grown to confluence in six-well plates and loaded with 10 mMN-(ethoxycarbonylmethyl)-6-methoxyquinolinium bromide (MQAE, Molecular Probes, Eugene, OR) overnight as described in detail previously (24). Briefly, cells were incubated in a chloride-containing, nitrate-free buffer for chloride concentration equilibration inside and outside of the cells, and background fluorescence was recorded over 3 min. After this, the buffer was changed to a chloride-free, nitrate-containing buffer. Both buffers contained 100 μM CPT-cAMP (Roche Biochemicals) to stimulate CFTR. In the presence of chloride, MQAE is caged and the probe's fluorescence is quenched. Upon changing over to a chloride-free, nitrate-containing buffer, a chloride gradient is established, which results in the rapid exchange of chloride for nitrate in the cells. Over the first few minutes, the rate of increase in free MQAE fluorescence is then proportional to the number of chloride channels at the membrane. Increase in MQAE fluorescence was monitored over 20 min after buffer exchange. The rates of chloride efflux were calculated from the slopes of the fluorescence curves over the first 3 min. A Cytofluor II plate reader (PerSeptive Biosystems, Framingham, MA) equipped with a 360 nm excitation/460 nm emission filter set was used to measure MQAE fluorescence in the cells.

Software.

Densitometric quantification and image processing were carried out using Adobe Photoshop (Adobe Software, San Jose, CA) and NIH Image (National Institutes of Health, Bethesda, MD) software packages. All statistical analyses were performed using Instat and Prism software (Graphpad Software, San Diego, CA).

RESULTS

Effects of Dox on CFTR expression in T84 cells. In all experiments described below (except Fig. 2C), cells were treated with 0.25 μM Dox for up to 24 h, which caused no overt cytotoxicity or change in cell viability as measured by trypan blue exclusion assay. This dose is 20-fold lower than the half-maximal lethal concentration (LC₅₀) for Dox in these cells (J. W. Hamilton et al., unpublished results). The effects of Dox on CFTR mRNA expression were examined in T84 cells by a semiquantitative RT-PCR assay. Treatment of cells for up to 24 h with 0.25 μM Dox had little or no effect on steady-state CFTR mRNA expression (Fig.1). In contrast, Dox treatment significantly altered CFTR protein levels. A 24-h Dox treatment increased CFTR protein by approximately twofold (Fig. 2,A–C), particularly the “C” band, which represents the mature, fully glycosylated form of CFTR (Fig.2A). Dose-response and time course studies indicated that the optimal treatment was 0.25 μM Dox for 24 h (Fig. 2, B and C). An increase in CFTR protein was also observed after immunoprecipitation with the A2 antibody and subsequent immunoblotting with a second CFTR-specific monoclonal antibody, M3A7 (Fig. 2, D and E).

To assess whether Dox specifically increased membrane CFTR expression, we performed selective cell-surface biotinylation of apical membrane proteins followed by immunoprecipitation with a CFTR-specific polyclonal antibody and detection using HRP-conjugated streptavidin. As demonstrated in Fig. 2, F and G, Dox-treated cells, compared with control cells, had an approximately twofold increase in levels of plasma membrane-associated CFTR that was detected as a single 170-kDa band (mature “C” band) on the immunoblots (Fig. 2E). Taken together, these experiments indicate that Dox treatment of T84 cells leads to enhanced CFTR protein expression at the cell surface.

Experiments were then conducted to assess whether the increased plasma membrane expression of CFTR resulted in enhanced chloride secretion in T84 cells. When we used an MQAE fluorescence-based assay, a 24-h Dox treatment increased chloride permeability approximately twofold relative to control, as shown in Fig.3B. Electrogenic chloride secretion was also measured across monolayers of T84 cells (Fig. 3D) in an Ussing chamber. Dox treatment increased CFTR-associated chloride secretion approximately twofold compared with control. Thus, when we used two different assays for CFTR function, Dox significantly increased CFTR-mediated chloride permeability in association with increased levels of membrane-associated CFTR in these cells.

Effects of Dox on ΔF508-CFTR expression in MDCK cells.

The effects of Dox treatment on functional cell surface expression of the CFTR folding mutant ΔF508-CFTR were then assessed with these same techniques. CFTR-associated chloride secretion was measured in an Ussing chamber across monolayers of parental MDCK-C7 cells and two isogenic MDCK cell lines stably expressing either GFP-tagged human CFTR or human GFP-ΔF508-CFTR. Dox treatment had no effect on chloride secretion in MDCK-C7 parental cells (Fig.4A). However, in MDCK cells expressing the mutant GFP-ΔF508-CFTR, Dox treatment caused an ∼2.3-fold increase in CFTR-mediated chloride secretion compared with untreated control cells (Fig. 2A). The increase in GFP-ΔF508-CFTR expression was confirmed by Western blotting of these cells (Fig. 4C). It is important to note that the control level of chloride secretion in these cells was similar to that of the parental MDCK cells expressing the canine CFTR (Fig. 4A), suggesting that there was essentially very little or no functional ΔF508-CFTR expression in the plasma membrane of these cells before Dox treatment.

Interestingly, Dox had no effect on chloride secretion in the MDCK cell line expressing the wild-type human GFP-CFTR (Fig.4A). There was also no effect on expression as determined by Western blotting (Fig. 4B). The reason for the lack of effect on either canine or human wild-type CFTR in these experiments is not clear, since wild-type CFTR expression had been affected in the T84 cells (Figs. 2 and 3). This may be due to the already high level of expression of CFTR in the MDCK cells as demonstrated by the fivefold increase in their chloride secretion compared with the parental MDCK cells expressing only the canine form (Fig. 4A). Conversely, it may be that in these MDCK cells, there are two distinct pathways for normal and mutant CFTR protein trafficking, the latter of which is more Dox sensitive. Several previous experimental cell culture procedures, such as high concentrations of glycerol, low-temperature incubation, or treatment with the xanthine drug, CPX, have been shown to be able to increase functional cell surface ΔF508-CFTR expression (3, 4,14). Interestingly, these same treatments did not appear to affect wild-type CFTR expression in these cells, suggesting that these agents are targeting an alternative pathway specific for the mutant, misfolded protein rather than general trafficking of CFTR or other proteins (3, 4, 14). Thus there may be differences in the efficiency of wild-type CFTR expression in these cells, e.g., more CFTR may follow the alternative pathway in T84 cells, and therefore be available for “rescue” by Dox treatment compared with the MDCK cells. In any event, Dox's lack of effect on functional expression of endogenous canine CFTR in the parental MDCK cells or on expression of the stably expressed normal human GFP-CFTR suggests that the change in chloride current in the GFP-ΔF508-CFTR MDCK cells was principally a result of substantially increased cell surface expression of GFP-ΔF508-CFTR from an essentially undetectable functional level.

To further confirm the I_sc and Western results, we then examined changes in GFP-ΔF508-CFTR expression in the MDCK cells by immunoprecipitation. As shown in Fig. 4D, Dox increased the expression of ΔF508-CFTR protein in these cells. Large glycoproteins typically migrate at a reduced rate on denaturing gels and, as a result, the mature “C” band of GFP-ΔF508-CFTR is detected at ∼240 kDa rather than at 210 kDa as predicted. Our laboratory had previously demonstrated a similar migration pattern for wild-type GFP-CFTR (17). Taken together, these results demonstrate that a noncytotoxic dose of Dox can substantially increase the functional cell surface expression of ΔF508-CFTR in epithelial cells.

DISCUSSION

We report that a single, noncytotoxic dose of Dox increased the amount of functional CFTR in the plasma membrane of T84 cells and the amount of functional ΔF508-CFTR in the plasma membrane of MDCK cells. In the T84 cells, this appeared to be principally a posttranscriptional effect because there was little effect on CFTR mRNA expression. In both cell models, Dox caused a significant increase in total cellular CFTR protein levels and a proportionate increase in CFTR-associated chloride secretion. These results suggest that analogs may be identified that will be clinically useful in ameliorating the CF disease phenotype in human CF patients.

The purpose of these studies was to obtain proof of principle for this class of drugs. Although the exact mechanistic basis behind this effect is currently being investigated, one possibility is that Dox increases the stability of the ΔF508-CFTR nascent forms, allowing them to fold properly. Dox may alter the kinetics of ΔF508-CFTR biogenesis influencing the interaction of the nascent protein with cellular chaperones. Previous reports indicate that heat shock cognate protein (HSC70) facilitates the early steps of ΔF508-CFTR biogenesis that appear to be important for maturation of the protein (22), and compounds that are capable of altering this association facilitate ΔF508-CFTR (9) expression and chloride secretion. Another possibility is that Dox directly interacts with and alters the folding, stability, or trafficking of ΔF508-CFTR during its biogenesis. For example, mutations in Pgp that alter its folding and plasma membrane trafficking, including a mutation that is comparable to ΔF508-CFTR, can be corrected by treatment of cells with Pgp substrate drugs (15). The authors proposed that such drugs acted as “chemical chaperones” that interact with Pgp and allow proper folding of the nascent molecule to the functional, mature form. A similar phenomenon could occur for ΔF508-CFTR, although this same study indicated no effect of these treatments on ΔF508-CFTR expression indicating that the effect was Pgp specific. However, anthracycline analogs were not tested in that study. Other drugs have also been described that are capable of altering both CFTR and Pgp protein levels. For example, butyrate compounds have been shown to be capable of stimulating both Pgp (16) and CFTR (18) protein expression. We found that another cancer chemotherapy drug, mitomycin C, also increased functional cell surface expression of CFTR in T84 and HT-29 cells, although the effects were more modest than those of Dox (R. Maitra et al., unpublished observations).

Alternatively, Dox may inhibit degradation of misfolded ΔF508-CFTR, which might favor proper folding and maturation. However, previous studies by Kopito and colleagues (11, 23) demonstrated that inhibition of the 26S proteosomal degradation pathway per se does not enhance ΔF508-CFTR folding, but rather leads to aggregation. Because other proteolytic pathways are involved in CFTR degradation (8), this possibility cannot be completely ruled out. These and other possibilities remain to be explored in future studies.

Perhaps most interestingly, a recent clinical observation was made (13) in a CF patient with ΔF508-CFTR mutation in one allele and G673X stop mutation in the other allele who had fibrosarcoma. This patient demonstrated significantly improved lung function and inhibition of his Pseudomonas aeruginosainfection after combination chemotherapy with cyclophosphamide and the anthracycline epirubicin. This is an intriguing clinical observation that should be further investigated systematically in CF cancer patients receiving chemotherapy.

In this study, the Dox treatment regimen at which significant ΔF508-CFTR effects were observed (0.25 μM) is more promising from a clinical pharmacology perspective. This Dox concentration is ∼20-fold lower that its LC₅₀ in these cell systems, suggesting that one might also be able to achieve these effects in vivo at a comparably low dose that is well below those used in cancer chemotherapy and that can elicit significant “nontarget” toxicity. Dox itself is unlikely to be useful in a CF clinical setting due to its cumulative systemic toxicity. However, it is likely, given that there are thousands of anthracyclines and their derivatives that have been developed and characterized over the past 50 years, that analogs can be discovered or developed that share the beneficial properties of Dox in increasing ΔF508-CFTR expression while avoiding its cumulative toxicities. The long-term goal of this research is to develop clinically useful drugs for the systemic treatment of CF patients with the aim of ameliorating their disease phenotype.

FOOTNOTES

• This work was supported by grants from the National Institutes of Health to B. A. Stanton, and from the Cystic Fibrosis Foundation to B. A. Stanton and J. W. Hamilton. J. W. Hamilton was also partially supported by the Norris Cotton Cancer Center, and R. Maitra was partially supported by a research grant to J. W. Hamilton from Bristol-Myers Squibb.

• Address for reprint requests and other correspondence: J. W. Hamilton, Dept. of Pharmacology & Toxicology, Dartmouth Medical School, 7650 Remsen, Hanover, NH 03755–3835 (E-mailjosh.[email protected]edu).

• The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.