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Originally published In Press as doi:10.1074/jbc.M504200200 on September 15, 2005

J. Biol. Chem., Vol. 280, Issue 46, 38193-38202, November 18, 2005
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An Energy-dependent Maturation Step Is Required for Release of the Cystic Fibrosis Transmembrane Conductance Regulator from Early Endoplasmic Reticulum Biosynthetic Machinery*

Jon Oberdorf1, David Pitonzo2, and William R. Skach3

From the Department of Biochemistry and Moleculor Biology, Oregon Health & Sciences University, Portland, Oregon 97239

Received for publication, April 18, 2005 , and in revised form, September 8, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Polytopic proteins are synthesized in the endoplasmic reticulum (ER) by ribosomes docked at the Sec61 translocation channel. It is generally assumed that, upon termination of translation, polypeptides are spontaneously released into the ER membrane where final stages of folding and assembly are completed. Here we investigate early interactions between the ribosome-translocon complex and cystic fibrosis transmembrane conductance regulator (CFTR), a multidomain ABC transporter, and demonstrate that this is not always the case. Using in vitro and Xenopus oocyte expression systems we show that, during and immediately following synthesis, nascent CFTR polypeptides associate with large, heterogeneous, and dynamic protein complexes. Partial-length precursors were quantitatively isolated in a non-covalent, puromycin-sensitive complex (>3,500 kDa) that contained the Sec61 ER translocation machinery and the cytosolic chaperone Hsc70. Following the completion of synthesis, CFTR was gradually released into a smaller (600–800 kDa) ATP-sensitive complex. Surprisingly, release of full-length CFTR from the ribosome and translocon was significantly delayed after translation was completed. Moreover, this step required both nucleotide triphosphates and cytosol. Release of control proteins varied depending on their size and domain complexity. These studies thus identify a novel energy-dependent step early in the CFTR maturation pathway that is required to disengage nascent CFTR from ER biosynthetic machinery. We propose that, contrary to current models, the final stage of membrane integration is a regulated process that can be influenced by the state of nascent chain folding, and we speculate that this step is influenced by the complex multidomain structure of CFTR.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The cystic fibrosis transmembrane conductance regulator (CFTR)4 is a polytopic membrane protein that is synthesized in the endoplasmic reticulum (ER) and expressed in the apical membrane of selected epithelial tissues (1). Inherited mutations that disrupt CFTR expression, trafficking, stability, or chloride channel activity cause clinical manifestations of cystic fibrosis (CF) (2). In >90% of CF patients the fundamental defect is failure of mutant CFTR to fold properly (3). Both wild-type and mutant proteins are initially synthesized in an immature, unstable conformation, and both can achieve functional activity in the ER (4). However, CFTR requires an ATP-dependent conformational change to complete its intracellular processing and delivery to the plasma membrane (5, 6). In most heterologous systems, ~70% of wild type and nearly 100% of common mutant variants (e.g. {Delta}F508) fail to complete this maturation step and are recognized and degraded via the ubiquitin-proteasome ER-associated degradation pathway (7–9). Recent studies have further indicated that maturation is dependent on cell type and that wild-type protein is efficiently trafficked in native epithelial cells under physiological growth conditions (10).

CFTR exhibits a modular, multidomain structure characteristic of the ABC transporter superfamily (1). It is comprises two membrane-spanning domains with six transmembrane (TM) segments each, two cytosolic nucleotide-binding domains, and a unique cytosolic regulatory (R) domain. A poorly understood, but important aspect of CFTR biogenesis is how folding and assembly of different domains are coordinated by cellular machinery in different cellular compartments. Topology of membrane-spanning domains is established during synthesis as the ribosome and nascent chain target to the ER membrane and bind a large protein-conducting channel (translocon) composed of Sec61{alpha} and its associated proteins (11–14). As translation proceeds, internal sequences within the nascent CFTR polypeptide interact with the ribosome and translocon where they initiate polypeptide translocation into the ER lumen, orient cytosolic loops, and direct transmembrane segment integration into the lipid bilayer (12, 15–17).

CFTR folding is facilitated by cytosolic and ER chaperones (Hsc70, Hsp90, and calnexin) (18–20) and co-chaperones (Hsp40 homologs and CHIP) (21, 22). Hsc70 and Hsp40 decrease nucleotide-binding domain aggregation and improve productive folding in vitro (23, 24). Similarly, Hsc70 and Hsp40 homologs bind cytosolic domains in vivo, and chaperone release correlates and co-chaperone with acquisition of a stable structure in the ER membrane (23, 25). In contrast, unstable conformations of CFTR remain bound to Hsc70 which, together with the co-chaperone/E3 ligase CHIP, can result in polyubiquitination (26, 27). The dynamic complement of chaperone and co-chaperone interactions therefore links CFTR biosynthetic and degradation pathways by facilitating both the acquisition of a trafficking-competent conformation (18, 23, 24, 28, 29) as well as the ubiquitination and degradation of misfolded conformers (26, 27, 30, 31). Although few steps of the in vivo CFTR folding pathway have been well characterized, studies predict that the early fate of CFTR is tightly regulated by protein-protein interactions that begin during translation and culminate as the mature protein is packaged for ER export (32).

Several lines of evidence indicate that proper formation of domain-domain contacts is the rate-limiting step for trafficking-defective CFTR mutants (25, 33–36). Thus, to understand how defects in CFTR biogenesis cause disease, it is necessary to determine how cellular folding machinery coordinates secondary and tertiary structure formation as the protein is synthesized and folded at the ER membrane. In this regard, some substrates are able to fold co-translationally such that N-terminal domains acquire functional activity even while C-terminal domains remain tethered to the translating ribosome (37, 38). Other membrane proteins complete their folding post-translationally only after the polypeptide has been fully synthesized and released from the ribosome-translocon complex (RTC) (39–41). In the case of CFTR, ribosome binding to the cytosolic face of the ER membrane likely constrains folding and association of the membrane-spanning domains, nucleotide-binding domains, and R domain. Thus, release from early biosynthetic machinery represents an important biosynthetic step toward conformational maturation. Although it has generally been assumed that polypeptides are spontaneously released from the RTC when translation is terminated, the precise mechanism by which membrane integration is coordinated for complex multidomain proteins such as CFTR remains unknown.

The above considerations predict that dynamic interactions of cellular proteins with nascent CFTR play a critical role in early folding events. Because of their transient nature, it has been difficult to capture proteinprotein interactions before polypeptide synthesis is completed. In the current study, we have overcome this limitation using cell-free and whole cell expression systems that enable us to visualize and manipulate actively translating CFTR biogenesis intermediates. Our results confirm that, even at the earliest stages of translation, CFTR interacts with highly dynamic protein complexes. Partially synthesized precursors were initially isolated with functional RTCs and cytosolic chaperones. CFTR was then gradually released into a smaller ATP-dependent 600- to 800-kDa complex after the completion of synthesis. Surprisingly, full-length CFTR exhibited prolonged interactions with early ER biosynthetic machinery, and release from the RTC was not spontaneous but required both NTPs and cytosolic factors. Thus, contrary to current models, the final step of CFTR integration into the membrane is an active rather than passive process that is regulated in a substrate-specific manner and likely reflects unique requirements of the CFTR folding pathway.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
In Vitro Transcription/Translation—WT and {Delta}F508 CFTR (plasmid pSPCFTR (42) and an identical plasmid missing Phe508), myc-tagged human P-gp (plasmid pSPMDR1 (42) containing LEQKLSSEEDLQ inserted after codon Lys-681), bovine prolactin (43), or myc-tagged AQP2 (44) were transcribed in vitro at 40°C for 1 h in reactions containing 0.2–0.4 mg/ml plasmid DNA, 40 mM Tris, pH 7.5, 6.0 mM MgOAc2, 2 mM spermidine, 0.5 mM each of ATP, CTP, UTP, 0.1 mM GTP, 0.5 mM GpppG (Amersham Biosciences), 10 mM DTT, 0.2 mg/ml bovine calf tRNA, 0.75 units/ml RNase inhibitor (Promega, Madison, WI), and 0.4 unit/ml SP6 RNA polymerase (Epicenter, Madison, WI). RRL was prepared from New Zealand White Rabbits as described (45, 46). Translation was performed at 25 °C for 1–2 h in reactions containing 20% transcript mixture, 40% nucleased rabbit reticulocyte lysate (45) plus 1 mM ATP, 1 mM GTP, 12 mM creatine phosphate, 40 µM each of 19 essential amino acids except methionine, 1 µCi/µl Tran[35S]-label (ICN Pharmaceuticals, Irvine, CA), 40 µg/ml creatine kinase, 0.1 mg/ml tRNA, 0.2 unit/µl RNase inhibitor (Promega, Madison WI), 10 mM Tris, pH 7.5, 100 mM KOAc, and 2 mM MgCl2. Canine pancreas microsomal membranes, prepared as described (47), were added at the start of translation (final concentration, 7.0 A280). 50–100 µM aurin tricarboxylic acid was added after 12–15 min to synchronize translation. Prior to solubilization, ER microsomes were isolated by pelletting through 0.5 M sucrose in Buffer A (20 mM Tris-HCl (pH 7.5), 50 mM KCl, 5 mM MgCl2, and 2 mM DTT) at 180,000 x g for 10 min as described (45, 48).

Chemical Cross-linking—Chemical cross-linking was performed by pelletting CFTR-containing microsomes through 0.5 M sucrose, 50 mM HEPES-KOH, (pH 7.5), 100 mM KOAc, and 5 mM Mg(OAc)2. Pellets were solubilized in 0.2% SDS, 1% Triton X-100 (Triton) or 2% digitonin in 20 mM HEPES-KOH (pH 7.5), 50 mM KCl (NaCl for SDS), 5 mM MgCl2, and 5% glycerol. Samples were incubated in 1 mM 3,3-dithiobis(sulfosuccinimidylproprionate) (DTSSP) for 30 min at 24 °C and quenched in 100 mM Tris-HCl, pH 7.5, for 10 min. Samples were then denatured in 1% SDS, 10 mM EDTA, and 100 mM Tris-HCl (pH 8.0) at 37 °C for 30 min prior to immunoprecipitation.

Xenopus Oocyte Expression—Oocytes were surgically removed from anesthetized Xenopus laevis and digested for 3 h at room temperature in ~4 vol. of calcium-free MBSH (88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 0.82 mM Mg SO4, 0.33 mM Ca(NO3)2, 0.41 mM CaCl2, 10 mM sodium-HEPES, pH 7.4) containing 0.2 unit/ml Blendzyme III (Roche Applied Sciences). Stage VI oocytes were collected and stored at 17 °C in MBSH containing 100 units/ml penicillin, 100 mM/ml streptomycin, and 50 µg/ml gentamicin. Oocytes were injected on an ice-cold stage with 50 nl of a 1:5 mixture (transcript:10x concentrated Tran[35S]-label) as described (12, 49). Incubations were carried out at 17 °C. Incubation times were determined empirically to capture both partial- and full-length CFTR protein at the time of initial homogenization. This ensured that active translation of radiolabeled CFTR was still ongoing and enabled us to distinguish different behaviors of these different populations of polypeptides. For gradient analysis, oocytes were homogenized in 5 volumes of 0.25 M sucrose in buffer A supplemented with 1x protease inhibitor I or protease inhibitor III (Calbiochem). Homogenate was centrifuged at 800 x g for 10 min. Supernatant was layered onto 0.1 ml of 1.8 M sucrose in buffer A and 0.4 ml of 0.5 M sucrose in buffer A and centrifuged at 250,000 x g for 10 min. Oocyte membranes were collected at the 0.5/1.8 M sucrose interface, diluted 1:5 with buffer A, and pelleted at 250,000 x g for 10 min. Except where indicated, homogenates and extracts were kept at 4 °C. For ex vivo release assays, membranes were resuspended in buffer A or freshly prepared oocytoplasm (supplemented with protease inhibitors) and ATP/GTP, puromycin, and cyclohexamide as indicated. When comparing different release conditions and sequential time points, samples were taken from the same batch of injected oocytes and processed and analyzed together. All experiments were repeated (two or more times) to verify reproducibility.

Glycerol Gradient Centrifugation—Microsomal membranes from in vitro translations or Xenopus oocytes were solubilized for 30 min in either 2% digitonin (Calbiochem), 1% Triton, or 0.2% SDS in Buffer B (50 mM KCl (or NaCl), 5 mM MgCl2, 20 mM Tris-HCl, pH 7.5), and 0.5% protease inhibitor mixture III (Calbiochem). Samples were clarified by centrifugation at 16,000 x g for 10 min and layered onto an 8% to 35% glycerol step gradient (8 x 110-µl steps 8.5, 12.5, 16.3, 20, 23.8, 27.5, 31.3, and 35%), in buffer B containing 0.1–0.2% of the same detergent used for solubilization. For SDS gradients, NaCl was substituted for KCl with no observable effect on the gradient profile. Gradients were centrifuged at 180,000 x g (maximum) for 2 h (Beckman TLS55 rotor). 12x 75-µl aliquots were collected. For cross-linking experiments, the solubilization and gradient buffers contained HEPES instead of Tris. The positions of chromatography standards bovine serum albumin (67 kDa), alcohol dehydrogenase (140 kDa), apoferritin (440 kDa), and thyroglobulin (670 kDa) were determined on parallel gradients.

Preparation of Oocytoplasm—100–200 injected (or uninjected) oocytes were rinsed several times in MBSH and centrifuged at 500 x g for 1 min to compress the oocytes and remove excess buffer. Oocytes were then lysed (in the presence of 50 µg/ml cytochalasin) by centrifugation at 19,000 x g (maximum) for 15 min (Beckman TLS55 rotor). The resulting grayish supernatant (middle layer) containing cytosol and cellular membranes was collected (oocytoplasm) and supplemented where indicated with an ATP generation system (10 mM creatine phosphate, 0.1 mg/ml creatine kinase), 50 mM KCl, 1 mM MgAc2, 2 mM glutathione (reduced), protease inhibitors, and 7.5 units/ml RNase inhibitor. Translation inhibitors were added as indicated. Extracts were incubated at 17 °C after which samples were diluted in 0.4 ml of 0.25 M sucrose in Buffer A, layered over 0.5 ml of 0.5 M sucrose in Buffer A, and sedimented at 250,000 x g (maximum) for 10 min. The membrane pellet was then extracted in buffer B containing 2% digitonin and prepared for glycerol gradient centrifugation as described above. In some experiments membranes containing radiolabeled CFTR were resuspended in a small volume of buffer B, 1 mM DTT, and translation inhibitors as indicated. They were then combined with fresh oocytoplasm (oocytoplasm:microsome ratio of 2:1), incubated at 17 °C, and prepared for glycerol gradient fractionation as described above.

Immunoprecipitation and SDS-PAGE—Gradient fractions from in vitro translations were either analyzed directly by SDS-PAGE or immunoprecipitated in Buffer C (0.1 M NaCl, 2 mM EDTA, 0.1 M Tris-HCl (pH 8.0), and 0.5 mM phenylmethylsulfonyl fluoride) containing 0.2% detergent. Samples from oocyte gradients were immunoprecipitated by denaturation in 100 µl of 1.0% SDS, 100 mM Tris HCl, pH 8.0, 10 mM EDTA, and 1x protease inhibitor mixture III. Extracts were incubated at 37 °C for 20–30 min, diluted with 1.0 ml 1% Triton in Buffer C, incubated on ice for an additional 20 min, and clarified by centrifuging at 16,000 x g for 10 min. 1 µl of antisera and 5 µl of Protein A-agarose (Bio-Rad) were added. The CFTR antibody is a polyclonal rabbit antisera raised against CFTR residues 45–65 (25). The anti-Hsc/Hsp70 polyclonal rabbit antisera was a generous gift of Dr. W. Welch. Samples were mixed overnight and washed 3x with 1 ml of 1% Triton in buffer C and 2x with 1 ml of 100 mM Tris-HCl (pH 8.0), 100 mM NaCl and subjected to SDS-PAGE. Gels were analyzed by autoradiography with or without En3Hance (PerkinElmer Life Sciences) fluorography and digitized with a UMax Powerlook III scanner and Adobe Photoshop software. Band signals were quantitated using a Bio-Rad Molecular PhosphorImager Fx (Kodak screens, Quantity-1 software).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
In Vitro Formation of CFTR Complexes—Identification of cellular proteins that participate in co-translational CFTR folding requires expression conditions where both full-length and partial-length polypeptides can be visualized. In vitro translation systems are well suited for such analysis and have been used to define mechanisms of CFTR topogenesis (11, 12, 15), identify early CFTR chaperone interactions (50), and characterize CFTR degradation pathways (48, 51). We previously showed that rabbit reticulocyte lysate (RRL) supplemented with ER-derived microsomal membranes faithfully reconstitutes CFTR synthesis, core glycosylation, and membrane integration (45, 51). In addition, because RRL lacks other cellular organelles, it provides a source of nascent polypeptide in which processing is limited to early events that occur within the ER compartment.



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  		FIGURE 1.
Nascent CFTR binds to large protein complexes in vitro. Microsomal membranes containing in vitro translated CFTR were solubilized in SDS, Triton, or digitonin and centrifuged through a glycerol gradient as described under "Materials and Methods." Gradient fractions top-to-bottom were collected and analyzed by SDS-PAGE. Migration of molecular weight standards are indicated above autoradiograms (downward arrows). Location of full-length CFTR (horizontal arrow) and partial-length translation products (brackets) are indicated on the right.

 
To capture interactions with nascent CFTR polypeptides, ER microsomal membranes containing in vitro synthesized CFTR were solubilized in denaturing or non-denaturing detergents and analyzed by glycerol gradient centrifugation. In SDS, full-length CFTR sedimented with an apparent size of a 160-kDa monomer (Fig. 1, top panel). In contrast, non-denaturing detergents generated two distinct CFTR populations. The first was a very large and stable complex that pelleted through the gradient (Fig. 1, middle and bottom panels, lane 12). The second was a smaller heterogeneous complex that migrated with an apparent molecular mass of 300–400 kDa in Triton and 400–800 kDa in digitonin. This profile suggested that non-covalent CFTR-protein interactions are preserved in non-denaturing detergents. Differences in migration patterns for Triton and digitonin are consistent with milder properties of digitonin and its ability to better preserve membrane protein interactions (52–54). However, it is also possible these detergents bind with different stoichiometry to CFTR hydrophobic domains and/or associated membrane phospholipids. In contrast to full-length CFTR, partial-length translation intermediates were found in the lighter fractions in SDS, whereas they were almost exclusively recovered in the gradient pellet under non-denaturing conditions. This is consistent with their expected persistent binding to the RTC (>4,000 kDa).

Immature forms of CFTR bind Hsc70 in vivo (20). In vitro synthesized CFTR also co-immunoprecipitated with Hsc70 (Fig. 2A, lanes 1–3), and binding was disrupted when microsomes were pelleted in the presence of ATP (Fig. 2A, lanes 4–6). Thus Hsc70 undergoes physiological cycles of CFTR binding and release during in vitro ADP-ATP exchange. We therefore examined the effect of ATP on CFTR sedimentation. When microsomes were solubilized in the presence of ATP, CFTR that was released from the gradient pellet sedimented primarily in fractions 2–5 similar to the pattern observed for SDS (Fig. 2, B and D). In contrast, ATP depletion prior to solubilization resulted in a shift in sedimentation toward denser fractions (fractions 4–7) indicating stabilization of protein-protein interactions. Although ATP depletion yielded relatively small changes in CFTR migration, the shift in peak migration, representing an average mass difference of ~200 kDa, was highly reproducible (Fig. 2D). Immunoprecipitation of gradient fractions with Hsc/Hsp Hsp70 antisera further revealed that Hsc70-bound CFTR was recovered predominantly in fractions 6–9 and in the gradient pellet (fraction 12, Fig. 2, C and E). Thus in vitro synthesized CFTR exhibits ATP-dependent protein interactions that include Hsc70 and influence its sedimentation through glycerol gradients.



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  		FIGURE 2.
CFTR complexes are ATP sensitive and contain Hsc70. A, microsomes containing in vitro translated CFTR were pelleted through a sucrose cushion in the presence or absence of ATP as indicated. Membranes were solubilized in Triton and immunoprecipitated with non-immune sera (NIS), Hsc70 antisera, or CFTR antisera (51) and analyzed by SDS-PAGE and autoradiography. B, RRL translation mixtures were incubated with apyrase (0.1 unit/µl) or ATP (3 mM) for 10 min prior to microsome pelletting, Triton solubilization, and glycerol gradient centrifugation. Where indicated, buffers for pelletting also contained added ATP. Gradient fractions were analyzed directly by SDS-PAGE. C, microsomes from RRL were solubilized in Triton and analyzed on glycerol gradients. Fractions were collected and immunoprecipitated under non-denaturing conditions with Hsc/Hsp Hsp70 antisera (top panel) or analyzed directly by SDS-PAGE (bottom panel). D, autoradiograms were quantitated by phosphorimaging, and protein recovered in each fraction was normalized to total full-length CFTR recovered in fractions 1–11 (n = 4 ± S.E.). E, quantitated representative data from panel C.

 
Sec61{alpha} is a major translocon component that transiently interacts with nascent membrane proteins during translocation and integration (55–58). We therefore wished to identify the population of CFTR complexes (if any) that remained associated with the translocon. For these studies we used the bifunctional chemical cross-linking agent DTSSP that covalently reacts with free amino groups and contains a cleavable disulfide bond within its 12-Å spacer arm. Following Triton and digitonin solubilization, DTSSP converted CFTR into a large covalent complex that was readily detected by SDS-PAGE at the top of the gel. This material decreased and full-length CFTR (155 kDa) was again visualized, when the cross-linker was cleaved by DTT (Fig. 3A, compare lanes 3–4 and 6–7). Although in vitro synthesized CFTR exhibited relatively diffuse migration under non-reducing conditions, significant full-length protein was visualized in the absence of cross-linker and when solubilized in SDS prior to cross-linking (Fig. 3A, lanes 1 and 2). Some CFTR smearing was also observed after cleavage with DTT, presumably due to persistent modification of CFTR lysine residues and/or nonspecific aggregation. These findings provide independent evidence that in vitro synthesized CFTR is bound to large non-covalent complexes that can be stabilized by chemical cross-linking. To determine which population of CFTR interacted with Sec61{alpha}, gradient fractions prepared with nondenaturing detergents were similarly cross-linked with DTSSP and immunoprecipitated with Sec61{alpha} antisera. Co-immunoprecipitated CFTR was then analyzed by SDS-PAGE and autoradiography after cross-linker cleavage (Fig. 3B). These experiments confirmed that CFTR associates with Sec61{alpha} during early stages of its biogenesis and that complexes containing translocation machinery are recovered primarily at the bottom of the gradient.

CFTR Complexes Represent Distinct Steps of a Maturation Pathway— The RRL system faithfully reconstitutes early aspects of biogenesis. However, most CFTR is unable to reach a mature conformation in vitro and remains sensitive to ER-associated degradation (48, 51). We therefore used Xenopus oocytes to determine whether CFTR complexes observed in vitro were also formed in vivo. Oocytes have been widely used for CFTR synthesis and functional studies (59, 60). In addition, we previously demonstrated that they efficiently process wild-type CFTR (~90% efficiency) similar to that observed for native airway epithelia (10). However, they recognize and degrade CFTR trafficking mutants (25) and therefore also maintain an intact ER quality control system.

Synthesis was initiated by co-injecting mRNA and [35S]methionine to induce a rapid pulse of radiolabeled CFTR. Under our conditions, incorporation of radiolabel into full-length CFTR requires ~45–90 min, and little CFTR is trafficked out of the ER for 5–8 h (25). Solubilized oocyte membranes were analyzed on glycerol gradients, and CFTR was recovered by immunoprecipitation with N terminus-specific antisera. SDS solubilization again resulted in uniform sedimentation of ~160-kDa CFTR monomers (Fig. 4, top panel), whereas non-denaturing detergents generated two distinct CFTR populations, a small heterogeneous complex that sedimented with an apparent mass of ~350–800 kDa, and a larger complex that pelleted through the gradient. The migration of these complexes was very similar to those observed in vitro.



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  		FIGURE 3.
CFTR cross-linking to Sec61{alpha}. A, in vitro translated CFTR was subjected to DTSSP cross-linking after solubilization in the indicated detergent and analyzed under non-reducing (lanes 1–4) or reducing (lanes 5–7) conditions. *, DTSSP-dependent CFTR complexes. B, CFTR gradients were prepared as in Fig. 1, and gradient fractions were cross-linked with DTSSP prior to immunoprecipitation with Sec61{alpha} antisera. Immunoprecipitates of each fraction were analyzed by SDS-PAGE under reducing conditions. The arrow at right indicates full-length CFTR translation product. Downward arrows show position of molecular weight standards on glycerol gradients.

 
To ensure that the cohort of CFTR recovered in the gradient pellet did not represent an off-pathway aggregate or degradation intermediate, in vivo pulse-chase metabolic labeling studies were carried out. These and subsequent experiments were performed using digitonin solubilization to better preserve CFTR interactions. At early time points after microinjection, >95% of immunoreactive CFTR sedimented through the gradient (Fig. 5A). Some of this material migrated as full-length protein (160 kDa) but most consisted of a mixture of polypeptides (60–150 kDa) representing partial-length nascent chains that were captured during intermediate stages of synthesis (Fig. 5A, lane 12). When oocytes were chased in unlabeled media prior to homogenization, these partial-length polypeptides were converted into full-length CFTR protein that was gradually released from the gradient pellet and recovered in fractions 4–8 (Fig. 5, A–C). Thus the large CFTR complex (in fraction 12) represents a true physiological intermediate in the CFTR maturation pathway. However, there appeared to be a significant delay in the release of full-length CFTR into the lighter gradient fractions, suggesting that this transition was temporally uncoupled from the completion of protein synthesis (discussed further below).



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  		FIGURE 4.
Formation of CFTR-protein complexes in vivo. A, CFTR mRNA and [35S]methionine were co-injected into in X. laevis oocytes. Membranes were collected, solubilized in the detergent indicated, and analyzed by glycerol gradient centrifugation as in Fig. 1. Each fraction was denatured in SDS, immunoprecipitated with CFTR antisera, and analyzed by SDS-PAGE and autoradiography. Migration of molecular weight standards in the gradient is indicated at top. Full-length CFTR is indicated by horizontal arrows. B, CFTR bands from A were quantitated by phosphorimaging. Protein in each fraction was expressed as percentage of total protein recovered.

 
Similar gradient profiles were observed for both WT and {Delta}F508 CFTR (Fig. 5, D–F). Translational precursors of {Delta}F508 were recovered solely in the gradient pellet at early time points, and full-length protein was gradually released from this complex into fractions 4–8 during the 3-h chase. At later time points than those examined here, less {Delta}F508 CFTR was recovered because the mutant protein is degraded via the ER-associated degradation pathway (Ref. 51 and data not shown). Because both proteins were able to undergo this release step with similar efficiency, subsequent experiments focused on WT protein.

Reconstitution of CFTR Complex Maturation ex Vivo—Requirements for CFTR maturation were further examined using a hybrid in vivo/ex vivo expression system that allowed us to manipulate final conditions of CFTR synthesis. Following co-injection of mRNA and [35S]methionine, oocytes were gently crushed by centrifugation, and translation of pre-programmed ribosomes was completed ex vivo in the collected oocytoplasm. By carefully controlling the time of homogenization, most immunoreactive CFTR could be initially captured as partial-length translation intermediates (faint 80- to 150-kDa bands, better visualized in Fig. 6B). These intermediates efficiently chased into full-length protein in both intact oocytes and isolated oocytoplasm (Fig. 6A). Full-length CFTR generated in oocytoplasm did not result from de novo translation, because it appeared very quickly and because oocytoplasm was unable to efficiently initiate translation (61).5 To determine whether maturation of CFTR complexes was also completed ex vivo, membranes were collected at each time point (from oocytoplasm shown in panel A), solubilized, and independently analyzed on glycerol gradients (Fig. 6B). For each gradient, CFTR recovered in fractions 5–8 was pooled and compared with fractions 11 and 12. As we observed for intact oocytes, CFTR precursors (translation intermediates) were recovered exclusively in fractions 11 and 12 (Fig. 6B, lanes 5–7). Full-length CFTR also initially appeared in these fractions and was then slowly and selectively released into fractions 5–8. At the end of the chase period, >90% of CFTR precursors had completed synthesis, and nearly 70% of CFTR had been released into the smaller, more mature protein complex.



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  		FIGURE 5.
CFTR complexes are dynamic. A–C, oocytes expressing WT CFTR were incubated at 17 °C for 1.5 h and chased in media containing 1 mM unlabeled methionine. Membranes were collected 1.5 h (A), 3 h(B), and 4.5 h (C) after injection, solubilized in digitonin, and subjected to glycerol gradient centrifugation as in Fig. 4. Migration of full-length CFTR (horizontal arrow) and partial-length CFTR intermediates (bracket) are indicated on the right. During the incubation period, translation intermediates chase into full-length protein, and full-length protein is released from fraction 12 into fractions 4–8. D–F, oocytes expressing {Delta}F508 CFTR were injected and processed as in panels A–C except that the initial homogenization (panel D) was performed 1 h after injection.

 



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  		FIGURE 6.
Ex vivo CFTR complex maturation. A, microinjected oocytes were incubated for 45 min and chased in unlabeled methionine (lanes 1–4), or crushed by centrifugation to generate oocytoplasm (lanes 5–8). At times indicated (after injection) homogenates were immunoprecipitated with CFTR antisera and analyzed by SDS-PAGE and autoradiography. B, pelleted membranes from oocytoplasm (obtained from panel A) were solubilized in digitonin and subjected to glycerol gradient centrifugation as in Fig. 5. Fractions 5–8 and 11–12 were pooled prior to immunoprecipitation. C, oocytoplasm was prepared from crushed oocytes 45 min after microinjection and incubated in the absence (lanes 1–3) or presence (lanes 4–6) of 1 mM puromycin prior to CFTR immunoprecipitation. D, membranes isolated from aliquots of oocytoplasm (C) were collected, solubilized in digitonin, and analyzed by glycerol gradient centrifugation. Indicated gradient fractions were pooled, and CFTR was immunoprecipitated. In the absence of puromycin, only full-length CFTR was released from fractions 11 and 12 during the ex vivo chase (lanes 1–6). In contrast, both partial- and full-length CFTR polypeptides were released following addition of puromycin (lanes 7–12). Migration of full-length CFTR (arrow) and partial-length CFTR intermediates (bracket) are shown. All time points were measured from time of injection.

 
CFTR complexes recovered in gradient fractions 5–8 are too small to contain the ribosome-translocon machinery. This suggests that time-dependent changes in migration reflect abrupt changes in protein-protein interactions that involve the release of nascent chains from early ER biosynthetic machinery. We therefore tested whether the transition of CFTR into the smaller complex would be mimicked by the aminoacyl-tRNA analog puromycin, which results in premature chain termination and cleavage of the peptidyl-tRNA bond. When oocytoplasm was incubated in the absence of puromycin, translation continued to generate full-length polypeptides that were selectively released into gradient fractions 5–8 (Fig. 6, C and D). Consistent with our hypothesis, puromycin inhibited protein synthesis and facilitated the premature release of both full-length and CFTR precursors out of the gradient pellet (Fig. 6D).

CFTR Complex Maturation Requires NTPs and Cytosolic Components—Nascent membrane proteins remain attached to the RTC during synthesis and are usually spontaneously released as the terminal peptidyl-tRNA bond is cleaved upon translation termination (56–58). However, a significant proportion of full-length CFTR was consistently recovered at the bottom of the gradient (fractions 11 and 12), suggesting that the final stage of integration into the lipid bilayer did not occur immediately after translation was completed. Indeed, when oocyte membranes were resuspended in buffer containing puromycin, cyclohexamide and/or supplemental ATP/GTP, neither full-length nor partial-length CFTR polypeptides were released into the lighter gradient fractions (Fig. 7A). However, when membranes were resuspended in oocytoplasm collected from uninjected oocytes, puromycin released the majority of CFTR-reactive polypeptides into the smaller complex (Fig. 7B). Although puromycin might have failed to catalyze peptidyl-tRNA cleavage due to conformational-specific ribosomal stalling, this seems unlikely because similar effects were observed for a relatively broad range of CFTR polypeptides (~100–160 kDa) in which the ribosome would be positioned on different portions of the mRNA. In addition, we and others have shown that puromycin efficiently cleaves peptidyl-tRNA bonds and releases polypeptides from ribosomes under the buffer conditions tested (52, 53).

It remained possible, however, that delayed release of so called "full-length" CFTR might be caused by translational stalling at or near the very end of the coding sequence. If so, then polypeptides near the end of synthesis would appear to migrate as full-length by SDS-PAGE but could remain tethered to the ribosome if the stop codon had not yet been reached. To rule out this possibility, oocyte membranes were isolated and incubated with cyclohexamide together with exogenous oocytoplasm and NTPs. Under these conditions, partial-length CFTR precursors remained in gradient fractions 11 and 12 (Fig. 8A). Thus CFTR release cannot occur if the peptidyl-tRNA bond remains intact. When CFTR was homogenized at a slightly later time point to allow full-length CFTR to accumulate, full-length polypeptides were selectively released into fractions 5–8 in the presence of ATP supplemented oocytoplasm but not in ATP-depleted oocytoplasm (Fig. 8B). Because protein synthesis was inhibited throughout the incubation, CFTR must have completed translation prior to homogenization. Yet these full-length polypeptides remained associated with ER biosynthetic machinery during membrane isolation and were released only upon further incubation. These results demonstrate that disassociation of CFTR from early ER biosynthetic machinery involves a novel maturation step that requires both energy and cytosolic components.



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  		FIGURE 7.
CFTR release from the RTC is cytosol dependent. A, oocytes were homogenized 1 h after RNA injection, and membranes were pelleted, resuspended in buffer, and supplemented with cyclohexamide (lanes 1–2 and 5–6), puromycin (lanes 3–4 and 7–8) and Mg-ATP/GTP (lanes 5–8). Incubation was carried out at 17 °C for an additional 30 min, and membranes were repelleted, solubilized in digitonin, and subjected to glycerol gradient centrifugation. Fractions 5–8 (A) and 11–12 (B) were pooled and immunoprecipitated as in Fig. 6. B, oocyte membranes were collected as in panel A and resuspended in either buffer (lanes 1–4) or freshly prepared oocytoplasm (lanes 5 and 6). All samples were supplemented with Mg-ATP/GTP or an energy regenerating system. Cyclohexamide (lanes 3 and 4) or puromycin (lanes 5 and 6) was added to block protein synthesis. Locations of full-length (right arrow) and partial-length (bracket) CFTR are indicated.

 
Conditions of RTC Release Are Substrate-specific—To determine whether requirements for RTC release were unique to CFTR, we examined several control proteins, including a secretory protein (preprolactin, 26 kDa), a small polytopic protein with six transmembrane segments (aquaporin-2, 29 kDa), and a related ABC transporter human P-gp. Prolactin has been widely used as a model substrate to investigate nascent chain-translocon interactions in vitro and has been shown by cross-linking studies to spontaneously release from the translocon upon cleavage of the peptidyl-tRNA bond (13, 62, 63). Consistent with this, greater than 95% of newly synthesized prolactin was released from the RTC at the earliest time points that gradient fractions could be visualized (30 min after injection) (Fig. 9B). Similarly, ~90% of the transmembrane protein aquaporin-2 was also released from the RTC coincident with the completion of synthesis (Fig. 9D). Because radiolabeled prolactin and aquaporin are only first detected 15–20 min after microinjection (Fig. 9, A and C, respectively), these newly synthesized proteins must be release from the RTC almost immediately after the completion of translation. Thus RTC interactions with simple secretory and transmembrane proteins differ substantially from those of CFTR.

P-gp is similar in size to CFTR (140 kDa) and like CFTR contains two six-spanning membrane domains and two large cytosolic nucleotide binding domains (64). It also contains three N-linked glycosylation sites that undergo carbohydrate processing and is efficiently trafficked in oocytes with somewhat faster kinetics than CFTR (data not shown). We therefore expressed a myc-tagged P-gp construct in pulse-chase labeled oocytes to analyze RTC release. At early time points following injection, partial-length P-gp fragments were quantitatively recovered in fraction 12 as expected (Fig. 10A, top panel). Full-length P-gp was also initially recovered in fraction 12 and fractions 4–8. After 90 min of chase, nearly all P-gp (full-length and larger glycosylated species) were released from the RTC (Fig. 10A, middle panel). Therefore P-gp maturation resembles that of CFTR, although there appeared to be subtle differences. In contrast to CFTR most full-length P-gp was already released from the RTC when partial-length intermediates were still plainly visible (compare Fig. 5, A and B, with Fig. 10A). Consistent with this, we were unable to capture sufficient P-gp associated with the RTC for ex vivo release assays, because the majority of full-length protein was consistently released into the lighter fractions at early time points examined (Fig. 10B). Taken together, these findings suggest that the rate of RTC release depends upon both the complexity of domain structure and unique properties of the substrate protein.



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  		FIGURE 8.
CFTR release from the RTC requires cytosol and NTPs but not protein synthesis. Oocytes were homogenized 1 h (A) or 1.5 h (B) after RNA microinjection. Membranes were collected and incubated in buffer supplemented with 1 mM ATP/GTP (lanes 1 and 2), in freshly prepared oocytoplasm treated with apyrase (lanes 3 and 4) or with an ATP regeneration system (lanes 5 and 6). Cyclohexamide was present in all samples to prevent protein synthesis. Membranes were pelleted after 30 min, solubilized, and subjected to glycerol gradient centrifugation. Fractions 5–8 (A) and 11–12 (B) were pooled, and CFTR was immunoprecipitated prior to SDS-PAGE and autoradiography. Location of full-length (horizontal arrow) and partial-length (bracket) CFTR is shown.

 



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  		FIGURE 9.
RTC release of secretory and TM control proteins. A, oocytes were co-injected with preprolactin mRNA and [35S]methionine and homogenized at times indicated. Radiolabeled prolactin was recovered by immunoprecipitation and analyzed by SDS-PAGE and autoradiography. B, oocytes were injected as in panel A, and membranes were collected after 30 min and analyzed on glycerol gradients as in Fig. 5. Gradient fractions were immunoprecipitated with prolactin antisera. C, myc-tagged AQP2 mRNA was expressed in oocytes as in panel A, and AQP-2 was recovered at times indicated with myc monoclonal antibody (9E10). D, oocytes expressing myc-AQP2 were processed and analyzed on glycerol gradients 30 min after injection as in panel B. A minor band of glycosylated AQP2 is indicated (arrow).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, cell free and Xenopus expression systems were used to capture partial-length nascent CFTR polypeptides and examine the dynamic behavior of early protein complexes in the ER membrane. CFTR solubilization in non-denaturing detergents revealed that both partial- and full-length polypeptides are initially bound to a very large protein complex containing ER biosynthetic machinery. Following the completion of synthesis, CFTR was gradually released into a smaller (~600 kDa), ATP-dependent complex and continued to bind cellular chaperones. This transition represented a distinct step in the maturation pathway, occurred in a time-dependent manner in Xenopus oocytes and isolated oocytoplasm, and required either the completion of protein synthesis or premature cleavage of the peptidyl-tRNA bond. Although we do not know the complete composition of the large protein complex, its size, sensitivity to puromycin, and quantitative binding to partial-length translation intermediates strongly indicate that it contains the ribosome and ER translocon complex. Consistent with this, CFTR recovered in gradient pellets cross-linked and co-immunoprecipitated with Sec61{alpha}, the major component of the translocation channel. Surprisingly, release of full-length CFTR from the RTC both in vivo and in vitro did not occur spontaneously upon the completion of protein synthesis. Rather, this critical step required both NTPs and cytosolic components. Our results therefore identify a novel, energy-dependent step in the early CFTR maturation pathway that is required for newly synthesized polypeptide to be released from ER biosynthetic machinery.

CFTR folding begins co-translationally while the nascent polypeptide is bound to the RTC (11, 12, 16, 65). The ribosome and Sec61 translocon play a critical role in these early stages of folding by translocating polypeptide into the ER lumen, orienting peptide loops into the cytosol, and integrating TM segments into the lipid bilayer (reviewed in Refs. 13, 17, and 55). During this process, cytosolic domains must be localized near the interface between the translocon and the membrane-bound 60 S ribosomal subunit (13, 54, 66, 67). Given the limited space available, it is unlikely that final steps of CFTR folding and domain-domain association could be completed until TM segments are released laterally from the translocon into the ER membrane. In this regard, TM segments from some bitopic proteins passively partition into the ER membrane during translation (68, 69), whereas others can remain within the translocon for significant periods of time (57, 58, 70). In the latter case, integration is proposed to be triggered upon translation termination and/or cleavage of the peptidyl-tRNA bond as evidenced by rapid loss of contact with translocon components (56–58, 63, 71). For polytopic proteins, integration patterns are more complex, and TM segments can integrate into the membrane individually (72), in pairs (43, 73), or in groups (74, 75). However, it has generally been assumed that these polypeptides are still spontaneously released from the RTC into the lipid bilayer once the peptidyl-tRNA bond is severed (13).



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  		FIGURE 10.
P-gp release from the RTC. A, oocytes expressing myc-tagged human P-gp were labeled for 1.5 h and then chased in media containing 1 mM methionine. Membranes were harvested 90 min, 3 h, and 4.5 h after injection (top, middle, and bottom panels, respectively) and analyzed on glycerol gradients as in Fig. 5. Position of full-length P-gp (black arrow) and a larger species, presumably representing a higher molecular weight glycosylated form (gray arrow), are indicated on right. Partial length translation intermediates are bracketed (right). B, oocyte membranes containing myc-P-gp were collected 1.5 h after injection and incubated for 30 min in buffer supplemented with puromycin, cyclohexamide, and ATP/GTP as indicated. Membranes were then pelleted, solubilized, and analyzed on glycerol gradients. Fractions 5–8 (A) and 11–12 (B) were pooled and immunoprecipitated with myc antibody as in Fig. 7. Under all conditions, full-length P-gp was released from the pellet and recovered in lighter fractions.

 
We now show that, for CFTR, RTC release is temporally dissociated from the completion of protein synthesis and is subject to regulatory (energy-dependent) influences. Thus our results raise new questions regarding how and where such polypeptides might persistently interact with RTC components. Recent cross-linking studies have indicated that TM segments, including those from polytopic proteins, occupy and move through specific environments within the translocon during membrane integration (58, 76–78). Detailed analysis of AQP4 integration has further shown that the translocon can accommodate multiple TMs simultaneously, suggesting that early tertiary folding of transmembrane domains could begin in the immediate vicinity of Sec61{alpha} (78). It is therefore plausible that final integration of polytopic proteins might require formation of a stable structure within the membrane. In this regard, CFTR contains an unusual number of charged residues in its TM segments (4 Arg, 2 Lys, 3 Glu, 1 Asp, and 1 His) that play important functional roles and might influence the stability of isolated TM segments within the lipid bilayer (79–81). Indeed, preliminary studies from our group have demonstrated that certain individual TM segments continue to cross-link Sec61{alpha} even after puromycin release.6 Alternatively, regions of CFTR might also remain associated with peripheral translocon components such as ribophorin I as was recently demonstrated for fragments of the amyloid precursor protein and opsin (82).

It is difficult to explain a direct role for NTPs in TM segment integration, because no core translocon components have been demonstrated to bind or hydrolyze nucleotides. On the other hand, ATP plays a key role in the function of several chaperones implicated in CFTR folding (18, 20, 23, 27). The ATP dependence observed here raises the possibility that RTC release might be coordinated or perhaps influenced by the folding of cytosolic domains or perhaps initiation of domain-domain interactions. Limited proteolysis experiments have revealed that ATP is also required to convert CFTR into a mature structure prior to exit from the ER (6, 20). Our data demonstrate that Hsc70 is co-translationally recruited to CFTR while the nascent chain is still attached to the RTC and continues to bind after RTC release. Thus maturation that occurs within the RTC likely represents early stages of folding that take place before a mature, trafficking-competent structure is acquired. This is supported by our results that {Delta}F508 CFTR is released from the RTC to nearly the same extent as WT protein even though it fails to mature and is ultimately subject to ER-associated degradation (25). Similar results have been reported in mammalian cells in which solubilized WT and {Delta}F508 CFTR are both found in a heterogeneous smaller complex similar to that observed here, but only WT is able to complete the maturation process (19). Taken together, the maturation step required for RTC release appears to occur before the structural change required for CFTR intracellular trafficking (6). CFTR therefore undergoes at least two separate energy-dependent steps as it matures in the ER compartment. Understanding how these two processes are related represents an important area for CF and other disorders of membrane protein folding.

It has been particularly difficult to examine RTC-nascent chain interactions in mammalian cells, because they are transient and highly dependent on the rate of protein synthesis, stability of interactions, and rate of RTC release. Xenopus oocytes provide a distinct advantage in this regard. They have been widely used for heterologous expression studies (59, 60, 83) and efficiently process CFTR at levels similar to those observed in native epithelial cells (10). Moreover, their reduced rate of protein synthesis at physiological temperatures allowed us to identify transient populations of full- and partial-length nascent CFTR polypeptides prior to RTC release. Importantly, oocyte RTCs do not indiscriminately exhibit prolonged interactions with substrates. Simple secretory and transmembrane control proteins were released at a rate too fast to be detected. In contrast, partial- and full-length intermediates of human P-gp were released only slightly faster than CFTR. Unfortunately, we were unable to determine whether P-gp release from the RTC also required ATP and cytosol, because insufficient full-length protein remained associated with RTCs during the post-homogenization studies. Because P-gp and CFTR exhibit a similar size, domain architecture and topological profile, it is likely that RTC interactions are influenced by both the complexity of domain organization as well as key features of the CFTR folding pathway. Given the diversity of native membrane proteins and the high degree of conservation in biosynthetic machinery, it is likely that the biogenesis requirements observed here for CFTR will also apply to other substrates and expression systems.


   FOOTNOTES
 
* This work was supported by the Cystic Fibrosis Foundation Therapeutics Inc. and by National Institutes of Health Grants DK51818 and GM53457 (to W. R. S.) and T32 DK07674 (to J. O.). 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. Back

1 Present address: Division of Hematology and Oncology, Portland Veterans Administration Medical Center, Portland, OR 97239. Back

2 Present address: Dept. of Physiology and Pharmacology, Oregon Health & Sciences University, Portland, OR. Back

3 To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology, Mail code L-224, Oregon Health & Sciences University, 3181 SW Sam Jackson Park Road, Portland, OR 97239. Tel.: 503-494-7322; Fax: 503-494-7368; E-mail: skachw{at}ohsu.edu.

4 The abbreviations used are: CFTR, cystic fibrosis transmembrane conductance regulator; DTSSP, 3,3-dithiobis(sulfosuccinimidylproprionate); DTT, dithiothreitol; ER, endoplasmic reticulum; TM, transmembrane; RRL, rabbit reticulocyte lysate; RTC, ribosome-translocon complex; Triton, Triton X-100. Back

5 J. Oberdorf, D. Pitonzo, and W. R. Skach, our unpublished observations. Back

6 D. Pitonzo and W. Skach, unpublished observations. Back


   ACKNOWLEDGMENTS
 
We thank Dr. W. Welch and K. Matlack for providing Hsc/Hsp70 and Sec61{alpha} antisera, respectively, and J. Eledge for excellent technical assistance.



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