Folding of the Voltage-gated K+ Channel T1 Recognition Domain*

Andrey Kosolapov and Carol DeutschDagger

From the Department of Physiology, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6085

Received for publication, September 13, 2002, and in revised form, October 31, 2002

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Voltage-gated K+ channels (Kv) are tetramers whose assembly is coordinated in part by a conserved T1 recognition domain. Although T1 achieves its quaternary structure in the ER, nothing is known about its acquisition of tertiary structure. We developed a new folding assay that relies on intramolecular cross-linking of pairs of cysteines engineered at the folded T1 monomer interface. Using this assay, we show directly that the T1 domain is largely folded while the Kv protein is still attached to membrane-bound ribosomes. The ER membrane facilitates both folding and oligomerization of Kv proteins. We show that folding and oligomerization assays can be used to study coupling between these two biogenic events and diagnose defects in assembly of Kv channels.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

K+ channels comprise a diverse and ubiquitous class of membrane proteins designed to facilitate the diffusion of K+ ions across the plasma membrane. Among this class of channels are voltage-gated K+ channels (Kv),1 which open and close in response to changes of membrane potential. Kv channels are formed by four subunits that surround the central permeation pathway. The monomeric Kv subunits assemble as tetrameric species in the ER membrane (1-7) and subsequently traffic to the plasma membrane. Once assembled, they do not dissociate in the plasma membrane (8), but it is not known whether they are in equilibrium with monomers in the ER membrane. Kv tetramerization likely occurs in a series of steps (6, 7, 9) involving different domains of the Kv protein and perhaps different biogenic intermediates. For example, the cytoplasmic T1 domain, an N-terminal sequence that is highly conserved among Kv channels and is responsible for subfamily-specific co-assembly of subunits (5, 10-12), can self-associate before the monomeric nascent Kv channel peptide detaches from the ribosome and exits from the protein-translocating channel in the ER (translocon) (9). This self-association occurs between folded T1 monomers (9, 13), giving rise to intersubunit interfaces that contain ~20 side chains that are involved in polar intersubunit interactions (14, 15). As is common for oligomeric proteins, the tertiary structure of individual subunits confers the proper interface for quaternary structure formation. Therefore, the aforementioned polar side chains are only in close proximity when the monomer is folded.

In this study, we focus on two biogenic events: folding of the T1 domain in the monomer and tetramerization of T1 domains. These two events must occur over the time frame of biogenesis. Folding into the tertiary structure could occur as soon as the T1 domain exits the ribosome tunnel into the cytosol, and tetramerization of T1 domains may be complete before Kv channels exit the ER membrane. The ability of T1 to form tetramers (tetramerization competence) could also begin when the nascent peptide first emerges from the ribosome. The interplay between T1 folding and tetramerization has not been explored, largely because of the unavailability of a folding assay. Previously, we suggested that ER membranes facilitate T1-T1 association in Kv1.3 (9), a feature that could underlie efficient ER assembly of oligomeric Kv channels. However, we do not know whether the ER membrane assists the T1 monomer folding to form a tetramerization-competent species or whether the monomer is already folded and tetramerization-competent before it reaches the ER membrane. Moreover, we do not know how completely folded the T1 domain is at any specific biogenic stage. Presumably, the tertiary structure of the T1 crystal structure represents one mature conformation.

To address these issues and directly study the folding of individual monomeric T1 domains, we have developed a biochemical microassay of formation of the folded intramolecular interface of monomeric Kv1.3. This assay is independent of a measurement of tetramer formation and uses 1) bismaleimides to cross-link pairs of cysteines engineered into an internally folded interface of the monomeric T1 domain and 2) a gel shift strategy that extends current pegylation techniques (16). This approach allows us to detect intramolecularly cross-linked Kv1.3 monomers. Specifically, we have introduced pairs of cysteines into the T1 Kv1.3 monomer, based on the crystal structure of the virtually identical T1 in Kv1.1a (14). To study protein folding at different stages of biogenesis, we have generated biogenic intermediates that remain attached to the ribosome. Our results demonstrate that T1 begins to fold prior to arrival of the nascent peptide at the ER membrane and that a point mutation reported to disrupt T1 tetramerization (15) and channel function has its primary effect at the oligomerization, not folding, stage.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Plasmid Constructs-- Standard methods of plasmid DNA preparation, restriction enzyme analysis, agarose gel electrophoresis, and bacterial transformation were used. The nucleotide sequences of all mutants were confirmed by automated cycle sequencing performed by the DNA Sequencing Facility at the School of Medicine on an ABI 377 sequencer using Big dye terminator chemistry (ABI). All mutant DNAs were sequenced throughout the entire coding region. Engineered cysteines were introduced into pSP/Kv1.3/cysteine-free (9) using QuikChange site-directed mutagenesis kit. Cysteine pairs were designed according to their proximity in the previously published crystal structure of T1 (14). Fig. 1 shows three such pairs.

In Vitro Translation-- Capped cRNA was synthesized in vitro from linearized templates using Sp6 RNA polymerase (Promega, Madison, WI). Linearized templates for Kv1.3 translocation intermediates were generated using KpnI or BstEII enzyme digestion to produce NH2 terminus-S1-S2-S3 (last amino acid is Gly292) or NH2 terminus-S1-S2-S3-S4-S5 (last amino acid is Val387), respectively. Proteins were translated in vitro with [35S]methionine (2 µl/25 µl translation mixture; ~10 µCi/µl Express; PerkinElmer Life Sciences) for 2 h at 22 °C in the presence of canine microsomal membranes and rabbit reticulocyte lysate (2 mM final [DTT]) according to the Promega Protocol and Application Guide.

Gel Electrophoresis and Fluorography-- Electrophoresis was performed using the NuPAGE system and precast Bis-Tris 10% or 4-12% gels and MOPS running buffer. Gels were soaked in Amplify (Amersham Biosciences) to enhance 35S fluorography, dried, and exposed to Kodak X-AR film at -70 °C. Typical exposure times were 16-30 h. Quantitation of gels was carried out directly using a PhosphorImager (Amersham Biosciences), which is very sensitive and detects cpm that are not necessarily visualized in autoradiograms exposed for 16-30 h.

Oocyte Expression and Electrophysiology-- Oocytes were isolated from Xenopus laevis females (Xenopus I, Michigan) as described previously (17). Stage V-VI oocytes were selected and microinjected with ~0.1 ng of cRNA encoding for wild-type or mutant Kv1.3. K+ currents from cRNA-injected oocytes were measured by two-microelectrode voltage clamping using a OC-725C oocyte clamp (Warner Instrument Corp., Hamden, CT) after 24-48 h, at which time currents at +50 mV were 2-10 µA. Electrodes (<1 MOmega ) contained 3 M KCl. The currents were filtered at 1 kHz. The bath Ringer solution contained 116 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 2 mM MgCl2, 5 mM Hepes (pH 7.6). The holding potential was -100 mV. For experiments in which inactivation kinetics were determined, we fit the data at +50 mV using the simplex algorithm (Clampfit; Axon Instruments). Activation and inactivation time constants are reported as mean ± S.E.

Cross-linking and Pegylation Assay-- Translation reaction (10-20 µl) was added to 500 µl of phosphate-buffered saline (PBS; calcium- and magnesium-free, pH 7.3, containing 2 mM dithiothreitol). The suspension was centrifuged at 50,000 rpm and 4 °C for 7 min (or 70,000 rpm, 15 min, and 4 °C for Kv1.3 translated in the absence of membranes), through a sucrose cushion (120 µl, containing 0.5 M sucrose, 100 mM KCl, 50 mM Hepes, 5 mM MgCl2, 2 mM dithiothreitol, pH 7.5). The pellet was resuspended in 50-500 µl of PBS. 0.5 mM ortho-phenyldimaleimide (PDM; Sigma) was added to those samples to be labeled while a control sample was treated identically but in the absence of PDM, at ~0 °C for 1 h. No reducing agent is present in these incubations. Samples containing PDM were quenched with 10 mM beta -mercaptoethanol at room temperature for 10 min. Control samples, untreated with PDM, were treated identically. A third sample was labeled with PDM but reserved for treatment with methoxy-polyethylene thiol (PEG-SH). Thiol-reducing agents must be avoided once PDM labels the protein; otherwise, free maleimides will be modified, and further assay with PEG-SH will be blocked. All samples were centrifuged at 50,000 rpm and 4 °C for 7 min, resuspended in 50 µl of PBS containing 1% SDS, and incubated at room temperature for 30 min. For membrane-free preparations, samples were centrifuged at 70,000 rpm, 4 °C, for 15 min. Those samples designated for pegylation with methoxy-polyethylene glycol maleimide (PEG-MAL; Mr 5000; Shearwater, Inc.) were treated with 10 mM beta -mercaptoethanol to prevent oxidation, which inhibits pegylation. Samples destined for pegylation with PEG-SH (Mr 5000; Shearwater, Inc.) received 50 µl of PBS containing only 1% SDS. All SDS-treated samples were diluted with either 50 µl of PBS containing PEG-MAL to give a final concentration of PEG-MAL of 20 mM and 5 mM beta -mercaptoethanol or 50 µl of PBS containing PEG-SH to give a final concentration of 20 mM PEG-SH. The pegylation reaction reached steady state by 1 h of incubation at 4 °C, and the PDM reaction reached steady state by 30 min.

Analysis of Pegylation Ladders-- For any given construct, radioactive protein incubated with PEG-MAL or PEG-SH was detected as distinct bands on NuPAGE gels and quantified using phosphorimaging. The data were analyzed as follows. For each lane, j, of the gel, the fraction of total protein molecules with exactly i pegylated cysteines was calculated as Wj(i) = cpm(i)/Sigma cpm(i), where cpm(i) is the counts per minute in the ith bin. For example, in Fig. 3, in which each Kv construct has 2 cysteines, i ranges from 0 to 2. If each cysteine is assumed to label to the same extent in the steady state, the fraction Fj of individual cysteines pegylated in the jth lane is Sigma iWj(i)/N, where N is the total number of cysteines in the protein molecule. For the gels in Figs. 3, F1 is the fraction of individual cysteines labeled by PEG-MAL (lane 1). F2 is the fraction of individual cysteines labeled by PEG-MAL after treatment with PDM (lane 2). From lane 3 the fraction of individual cysteines that has reacted with both PDM and PEG-SH is F3.

In most of our constructs, two cysteines were introduced. If these cysteines are sufficiently far from one another during PDM labeling, the probability of cross-linking them with PDM is very low. This analysis assumes that cross-linking by PDM did not occur after SDS denaturation. Our results will verify these assumptions either when the protein is denatured by SDS treatment or when the cysteines are sufficiently separated in the predicted crystal structure. By comparing the labeling in denatured versus nondenatured protein, we can estimate the cross-linking efficiency as follows. After SDS pretreatment (e.g. Fig. 3B, right gel) FPDM-SDS is the fraction of individual cysteines labeled with PDM, given by FPDM-SDS = (F1 - F2)/F1. From the same gel, F3 = FPDM-SDS PPEG-SH, where PPEG-SH is the probability that an individual cysteine labeled with PDM has reacted with PEG-SH. Thus, PPEG-SH = F3/((F1 - F2)/F1).

Using this estimate of PPEG-SH, we now can determine the probability of a pair of cysteines being cross-linked by PDM in the absence of SDS pretreatment (e.g. left gel in Fig. 3B). As above, the fraction of individual cysteines labeled by PDM is FPDM = (F1 - F2)/F1. The fraction of available free maleimides after PDM labeling in this case is FfMAL = F3/PPEG-SH, where F3 is determined from lane 3 in the left gel of Fig. 3B, and PPEG-SH was estimated as described above from SDS-pretreated channels. Finally, the probability of a pair of cysteines being cross-linked by PDM is Pxlink = FPDM - FfMAL. One-way analysis of variance or a Student's t test was used to determine whether differences in Pxlink values are statistically significant.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

To investigate when and in which compartment T1 monomers fold into their tertiary structure, we used a cross-linking strategy. Pairs of cysteines were engineered into the folded monomer interfaces of Kv1.3/cysteine-free, similar to strategies used to study oligomerization of T1 (9, 13). Guided by the crystal structure of T1 in Kv1.1a (14), we devised three criteria to select appropriate pairs. First, the residues must be far enough apart in the primary sequence that they are proximate only when the protein is folded. Second, the pair must be within 5-10 Å in the folded T1 monomer and therefore within cross-linking distance. Third, the pair must be on the surface of the folded tetramer and therefore presumably nondisruptive and accessible to cross-linking reagents. Three pairs that fulfill these criteria are Arg83/Gln136, Gln72/Gly114, and Thr69/Gln112 (Fig. 1). The equivalent pairs in Kv1.1a are in layers 1 and 3 of the Kv1.1a T1 crystal structure for the first pair and layers 1 and 2 for the second and third pairs (see Fig. 7). Cysteine substitution of these residues in a cysteine-free background yields functional Kv1.3 channels, as determined by Xenopus oocyte expression and two-electrode voltage clamp (Fig. 3A). R83C/Q136C, Q72C/G114C, and T69C/Q112C each expressed current with activation and inactivation time constants of 5.7 ± 0.3 ms (n = 4) and 60.2 ± 4.9 ms (n = 7), 5.0 ± 0.1 ms (n = 4) and 46.1 ± 1.9 ms (n = 4), and 5.5 ± 0.8 ms (n = 4) and 66.0 ± 6.0 ms (n = 4), respectively. The inactivation time constants are comparable with that of cysteine-free Kv1.3 (70 ± 3.5 ms inactivation time constant, n = 3).


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Fig. 1.   Representation of Arg83/Gln136, Gln72/Gly114, and Thr69/Gln112 in the primary sequence, the folded tertiary structure of a T1 monomer (left), and the quaternary structure of the T1 tetramer (right), according to the crystal structure of the T1 domain of Kv1.1a (14) for the equivalent residues. For Kv1.3, residues Arg83, Gln72, and Thr69 are in red; Gln136, Gly114, and Gln112 are in blue. Structures were made in RasMol.

If the T1 monomer is folded, then we should be able to cross-link these engineered cysteines with bifunctional cysteine reagents. For this purpose, we have chosen o-phenyldimaleimide (o-PDM; Sigma), which has an intermaleimide distance of ~6 Å. We used this reagent previously to cross-link cysteines engineered into the T1-T1 intersubunit interface (9). Any reaction of our T1 monomers with PDM can result in a mixture of five different species (Fig. 2), each of which will appear at the parent molecular weight on a NuPAGE protein gel. Thus, we needed a way to distinguish these five species and, in particular, a means to detect the intramolecularly cross-linked species (bottom row, right). For this purpose, we have designed a mass-tagging strategy using pegylation with both PEG-MAL and PEG-SH (see Ref. 16 for details of the MAL-pegylation method). The addition of one of these PEG molecules shifts the protein molecular mass by >= 10 kDa. A free SH group can be labeled with PEG-MAL, and a free peptidyl-maleimide can be labeled with PEG-SH. Each species of PDM-modified protein will have a specific pattern of pegylation, but only the intramolecularly cross-linked protein will lack a gel shift with either PEG reagent and thus remain at the parent molecular weight on NuPAGE gel (Fig. 2). All other species will produce a gel shift with one or the other PEG reagent or both.


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Fig. 2.   Schematic diagram of assay for intramolecular cross-linking. Monomeric T1 is represented by the hairpin-shaped line. PDM is indicated by its chemical structure, and engineered cysteines are indicated by circles. The predicted results upon treating each monomeric T1 species with pegylating agents are shown below each monomeric T1 species.

To study when T1 folds and in which cellular compartment, we used biogenic intermediates that remain attached to the ribosome-translocon complex. In this case, the folded state of the Kv1.3 T1 domain can be assessed from the time of its emergence from the ribosome until its release into the bilayer. Using such intermediates, we have previously demonstrated that T1 can tetramerize prior to exit of membrane-bound Kv1.3 from the translocon, consistent with a folded T1 monomer at this stage (9). Our first task was to test whether the strategies shown in Fig. 2 could detect an intramolecularly folded T1 in biogenic intermediates, namely in a BstEII-cut Kv1.3 intermediate attached to a membrane-bound ribosome. This intermediate contains S1-S5 and half of the pore region (9). mRNA was made from truncated cDNA that lacks a stop codon in the coding region. Ribosomes halt when they reach the end of such a transcript, and the nascent peptide chain remains attached to the ribosome as a peptidyl-tRNA. We have previously demonstrated that this is true for Kv1.3 intermediates (9). Biogenic intermediates were made in a cysteine-free Kv1.3 for each of the pairs described above.

Experimental data for the R83C/Q136C/BstEII-cut Kv1.3 intermediate are shown in Fig. 3B. Most of the free cysteines (80%) were pegylated to give two bands at ~ 55 and 70 kDa, equivalent to one and two PEG molecules per protein, respectively (Fig. 3B, left gel, lane 1). Pretreatment of the protein with PDM blocked ~93% of the free cysteines from reaction with PEG-MAL (Fig. 3B, left gel, lane 2). This result suggests that most cysteines reacted with PDM to produce either free peptidyl-maleimides (Fig. 2, bottom left) or peptides containing neither free maleimides nor free cysteines (bottom right, the cross-linked species). When PDM-treated protein was exposed to PEG-SH, little or no pegylation occurred (Fig. 3B, left gel, lane 3), indicating that almost no free maleimides were available in the protein. The probability that an individual cysteine has reacted with both PDM and PEG-SH is <0.1 (calculated from lane 3; see "Experimental Procedures"). Thus, intramolecular cross-linking had occurred. This conclusion is supported by the experiment shown in the right gel of Fig. 3B, in which the protein was first denatured with SDS (1%). Subsequently, the protein was treated as in the left gel in Fig. 3B with PEG-MAL (lane 1) or first with PDM and then PEG-MAL (lane 2) or first with PDM and then PEG-SH (lane 3). Lane 2 in the left gel is similar to lane 2 in the right gel, indicating that the reactivities of cysteines to PDM are not affected by SDS denaturation prior to PDM labeling (FPDM = 0.97, see "Experimental Procedures"). However, a dramatic gel shift was observed following PEG-SH treatment (cf. lanes 3), indicating that no cross-linking had occurred. Presumably, this is because SDS unfolded the T1 monomers, thus separating the two engineered cysteines. The probability that an individual cysteine has reacted with both PDM and PEG-SH is ~0.5 (calculated from lane 3, see "Experimental Procedures"). The data shown in Fig. 3B represent steady-state measurements, since no changes in pegylation were observed for incubations at 1, 3 (data shown), or 5 h with PEG reagents or 15, 30, or 60 (data shown) min with PDM. Moreover, these data represent only membrane-bound nascent peptides, since contamination with membrane-free ribosomes is <5% (data not shown). R83C/Q136C did not exhibit a gel shift when treated with PEG-SH alone, yet it could be pegylated with PEG-MAL (data not shown).


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Fig. 3.   Cross-linking of engineered constructs. A, functional assay of R83C/Q136C, Q72C/G114C, and T69C/Q112C. Xenopus oocytes were injected with cRNA for full-length (EcoRI-cut DNA templates) R83C/Q136C (trace 1), Q72C/G114C (trace 2), and T69C/Q112C (trace 3), and recordings were made 24-48 h postinjection. Peak current at +50 mV was measured to give the normalized current traces shown. B, cross-linking of R83C/Q136C/BstEII. Left gel, nascent peptide treated with PEG-MAL (lane 1) or first with PDM and then PEG-MAL (lane 2) or first with PDM and then PEG-SH (lane 3). Right gel, all samples were treated first with SDS. SDS samples were subsequently treated with PEG-MAL (lane 1) or with PDM and then PEG-MAL (lane 2) or with PDM and then PEG-SH (lane 3). C, cross-linking of Q72C/G114C/BstEII in the folded (left gel) and in the unfolded (SDS pretreatment, right gel) construct. D, cross-linking of T69C/Q112C/BstEII in the folded (left gel) and in the unfolded (SDS pretreatment, right gel) construct. For each gel, numbers on the left are molecular weight standards, and numbers on the right indicate singly pegylated (1), doubly pegylated (2), and unpegylated (0) protein. This applies to all pegylation gels. All samples were treated with strongly reducing conditions just prior to MAL-pegylation to minimize oxidation of cysteines and with RNase (20 µg/ml) prior to loading on the gel to remove the peptidyl-tRNA bands (9). All experiments were done in the presence of membranes. Unpegylated BstEII-cut intermediates appear as doublets at the predicted molecular mass of 43 kDa. The upper band of the doublet is due to ER core glycosylation; the lower one is unglycosylated Kv1.3 nascent peptide. Core glycosylation is not readily discerned at higher molecular weights.

For the data in Fig. 3B (right gel), the behavior of the single cysteine mutants alone predicts the observed probabilities. Each of the single mutants was pegylated (78 and 95% for R83C and Q136C, respectively), blocked with PDM (94 and 96%, respectively), and then pegylated following PDM-labeling (40-56%) (data not shown). If Gln136 was first denatured in SDS (unfolded) and then assayed for available cysteines and maleimides after PDM modification, the percentages of cysteines pegylated or blocked, or pegylated following PDM labeling, were relatively unchanged (75, 97, and 57%, respectively). Analysis of these results (see below) suggests that the presence of one cysteine in a pair does not affect the other cysteine (i.e. there is no cooperative pegylation, either positive or negative, between nearby cysteines). A similar lack of cooperativity was found for the intersubunit interface pair R118C/D126C in the Kv1.3 T1 domain (9).

We have evaluated other pairs of engineered cysteines as potential folding assays, namely Q72C/G114C and T69C/Q112C. These pairs are in layers 1 and 2 according to the T1 crystal structure (14, 15) (see Fig. 7). As shown in Fig. 3C, the Q72C/G114C construct exhibits strong intramolecular cross-linking. Lanes 3 are dramatically different for the two gels, the right gel representing prior SDS denaturation of the protein. The fraction of cysteines that reacted with both PDM and PEG-SH is 0.03 and 0.55 for the folded and unfolded peptide (lanes 3), respectively.

Although the folding assay is qualitatively suggestive, we quantified the results to obtain calculated probabilities of cross-linking, Pxlink. Such analysis permits a statistical evaluation. The probabilities may be calculated using the data obtained for pegylation, with and without SDS pretreatment, as described under "Experimental Procedures." For instance, for the two gels shown in Fig. 3B, F1 is the fraction of individual cysteines labeled by PEG-MAL (lane 1). F2 is the fraction of individual cysteines labeled by PEG-MAL after treatment with PDM (lane 2). F3 is the fraction of individual cysteines that has reacted with both PDM and PEG-SH. As described under "Experimental Procedures," using these fractions, F1-3 (Table I), we can calculate Pxlink, the probability of a pair of cysteines being cross-linked by PDM. Pxlink is 0.94 for Q72C/G114C. The T69C/Q112C construct has a lower cross-linking efficiency; Pxlink is 0.68 (Fig. 3D). In both cases, the single mutants (data not shown) account for the PEG-SH labeling shown in the double mutants. For comparison, Pxlink for R83C/Q136C is 0.78 ± 0.05 (n = 7).

                              
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Table I
Fraction of individual cysteines pegylated
Numbers are given as mean ± S.E. when n >=  3 and as mean ± average error when n = 2. Single constructs are considered as one group and include R83C, Q136C, Q72C, and T69C. +mm and -mm indicate translations done in the presence or absence of membranes.

In addition to measurements on unfolded (SDS-denatured) protein, we performed another control. We engineered a pair of cysteines in the same region of the T1 domain, but farther apart in the folded tertiary T1 monomer, and they should therefore not be cross-linked with PDM treatment (Fig. 4A). Gln72 and Gln136 (in layers 1 and 3 of the T1 crystal structure, respectively) are ~14 Å apart in the folded T1 tertiary structure and are on the surface of the quaternary structure, accessible to PDM. Fig. 4B indicates that the folded intermediates of BstEII-cut Q72C/Q136C were not cross-linked following PDM treatment. Pxlink is 0.16 and 0.82 for Q72C/Q136C (left gel) and R83C/Q136C (right gel), respectively. Moreover, Pxlink for Q72C/Q136C in the absence of membranes and in samples SDS denatured prior to PDM labeling, is 0.01 ± 0.02 (average of n = 2) and 0.00, respectively, similar to 0.03 ± 0.07 (n = 3) for Q72C/Q136C in the presence of membranes (see Fig. 5 and Table I). Thus, none of the incubation conditions per se lead to artifactual gel shifts.


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Fig. 4.   Cross-linking of Q72C/Q136C/BstEII-cut intermediate. A, representation of Q72/Q136 in the primary sequence, the folded tertiary structure of a T1 monomer (left), and the quaternary structure of the T1 tetramer (right) according to the crystal structure of the T1 domain of Kv1.1a (14) for the homologous residues. The residue numbers are for Kv1.3. Gln72 is in red; Gln136 is in blue. B, Q72C/Q136C/BstEII-cut intermediate (left gel) treated with PEG-MAL (lane 1) or first with PDM and then PEG-MAL (lane 2) or first with PDM and then PEG-SH (lane 3). Right gel, paired control R83C/Q136C/BstEII-cut intermediate, treated as in the left gel. Glycosylation and RNase treatment are as described in Fig. 3.


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Fig. 5.   Calculated probability of cross-linking, Pxlink. For each nascent Kv1.3 peptide, mRNA was translated either in the presence (left) or absence of membranes (right) and treated as described in Fig. 3. Double mutants are R83C/Q136C/BstEII-cut (black circles, n = 7), Q72C/G114C/BstEII-cut (black square, n = 1), T69C/Q112C/BstEII-cut (black triangle, n = 1), R83C/Q136C/KpnI-cut (open circle, n = 1), or Q72C/Q136C/BstEII-cut (open triangle, n = 3). Single mutants are grouped together and include T69C, Q72C, R83C, and Q136C (open hexagon, n = 5). The dotted line indicates the average Pxlink for Q72C/Q136C/BstEII-cut constructs (+mm, -mm), which cannot be cross-linked. Data, plotted as mean ± S.E. for n >=  3 and mean ± average error for n = 2, were obtained using the equations described under "Analysis of Pegylation Ladders."

Fig. 5 and Table I display the analyses for all of the constructs, both folded and unfolded (SDS-denatured). There is a statistically significant difference in Pxlink between folded constructs and all controls that cannot be cross-linked (i.e. those pairs that are too far apart (Q72C/Q136C) or single cysteine constructs) and denatured protein (p < 0.001 in all cases).

The results shown in Figs. 3-5 demonstrate that 1) the cross-linking/pegylation method can assay intramolecular folding events and 2) the T1 domain is folded in the nascent Kv1.3 protein before the protein exits from the translocon. To confirm these results, we performed an oligomerization assay, as described previously (9). This assay evaluates Kv tetramer formation in ER membranes by cross-linking engineered pairs of cysteines at putative interaction sites in the folded T1-T1 intersubunit interface of Kv1.3. Using R118C/D126C, we have previously shown close proximity between neighboring nascent subunits (9). This pair of residues lies in layer 3 (see Fig. 7) on the surface of the T1 tetramer, facing the membrane. In order to cross-link these cysteines, the T1 monomers must be correctly folded in the region of interest or the cysteines will not be close to each other in the quaternary structure. Likewise, if only certain regions of T1 are correctly folded while others are not, then some residues predicted by the crystal structure to come within a few angstroms of each other may be too far apart to be cross-linked. To confirm that T1 is correctly folded for oligomerization, we performed two oligomerization assays: one using the control R118C/D126C BstEII-cut intermediate and another using the R101C/D105C BstEII-cut intermediate. The former indicates folding and proximity in layer 3, and the latter indicates folding and proximity in layer 2. Both pairs indicate that tetramer formation occurs between ribosome-attached nascent Kv1.3 subunits. For R118C/D126C and R101C/D105C BstEII-cut intermediates, 54.8 ± 2.3% (n = 7) and 34.0 ± 2.4% (n = 3) of the protein, respectively, was cross-linked to give multimers (dimers, trimers, and tetramers; data not shown).

Application of the Folding Assay-- Can this approach be used to tell us when folding occurs and whether functional defects in mutant Kv proteins arise from folding and/or from oligomerization defects? To address this issue, we chose to study the nascent T1 peptide as it emerges from the ribosome, in both the presence and absence of microsomal membranes. Our previous studies suggest that the ER promotes tetramer formation (9). We do not know whether this is due to membrane-facilitated folding of T1 and/or oligomerization. It has been suggested for other membrane proteins that the ER membrane concentrates proteins, thereby facilitating oligomerization (18).

To determine whether folding occurs prior to membrane targeting, we performed this assay in the absence of membranes. R83C/Q136C BstEII-cut (contains the N terminus through the pore region) was translated in the absence of membranes and then cross-linked and pegylated to determine folding. As shown in Fig. 5, the mean Pxlink for BstEII-cut intermediates in the presence of membranes is 0.78 ± 0.05 (n = 7), significantly different from 0.34 ± 0.08 (n = 4) in the absence of membranes (p = 0.001, Student's t test). Furthermore, a KpnI-cut intermediate, a nascent peptide containing only the N terminus through the end of S3 was also cross-linked and pegylated (Fig. 5 and Table I). Because ~40 amino acids are required for a nascent peptide to traverse the ribosomal exit tunnel (19), hydrophobic transmembrane segments comprise only a small proportion (~15 and 20%, respectively) of the aqueous-exposed protein of KpnI-cut and BstEII-cut intermediates. Pxlink for the KpnI-cut intermediate is 0.77 and 0.47 in the presence and absence of membranes, respectively. Therefore, in the absence of membranes, T1 is folded, albeit less efficiently, in these constructs. Moreover, values of Pxlink for the KpnI-cut intermediates are similar to those for BstEII-cut intermediates, suggesting that the difference in length of these two classes of intermediates was not a determinant in folding of the T1 domain. Pxlink for Q72C/G114C/BstEII-cut and T69C/Q112C/BstEII-cut are likewise decreased in the absence of membranes but significantly greater than zero. These results suggest that T1 folding begins in the cytosol soon after emergence of the nascent peptide from the ribosome. We do not yet know whether the partially folded structure is tetramerization-competent. Perhaps tetramerization competence is achieved only when the complex arrives at the ER membrane.

To begin to identify determinants of T1 folding, we have examined a T1 residue that affects the ability of T1 to tetramerize (15). Specifically, we made T65D in a R83C/Q136C Kv1.3 background. This residue is in the T1-T1 intersubunit interface. T65D does not express current and cannot be cross-linked in an oligomerization assay, whereas T65V is functional and can be cross-linked to give multimers (Fig. 6, A and B). The activation and inactivation time constants for T65V are 5.0 ± 0.4 ms (n = 2) and 60.0 ± 3.9 ms (n = 3), respectively, consistent with that of cysteine-free Kv1.3. The fraction of cross-linked multimers in the oligomerization assay is 0.49 ± 0.05, 0.40 ± 0.07, and 0.03 ± 0.03 (average ± average error, n = 2) for Thr65, T65V, and T65D, respectively, suggesting that T65D does not form multimers. The analogous substitutions in Kv1.2 also alter structure and function (15), the Asp mutant leading to anomalous aggregation behavior, the Val mutant strengthening the T1-T1 interaction in the soluble T1 domain. The behavior of T65D may be due to impaired folding, impaired tetramerization, or both.


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Fig. 6.   Engineered cysteine constructs containing T65V and T65D. A, functional assays. Xenopus oocytes were injected with cRNA for full-length (EcoRI-cut DNA templates) T65V/R83C/Q136C (trace 1) or T65D/R83C/Q136C (trace 2), and recordings were made 24-48 h postinjection. Peak current at +50 mV was measured to give the current trace shown. Similar current records were obtained for T65V and T65D in a R118C/D126C Kv1.3 background (data not shown). B, tetramerization assay. T65/R118C/D126C, T65V/R118C/D126C, and T65D/R118C/D126C (lanes 1-3, respectively) were translated in the presence of membranes and treated with 0.5 mM PDM. For T65/R118C/D126C and T65V/R118C/D126C, monomers, dimers, trimers, and tetramers were detected as bands at 43, 90, 130, and 170 kDa, respectively. For T65D/R118C/D126C, only trace amounts of multimers and mostly monomers were detected. These results are typical of four experiments. C, assay of folding of T1 domains containing T65V and T65D. T65V/R83C/Q136C and T65D/R83C/Q136C BstEII-cut intermediates were each translated in the presence (+mm) or absence (-mm) of membranes and treated as described in Fig. 3. Data, plotted as mean ± S.E. for n >=  3, were obtained using the equations described under "Analysis of Pegylation Ladders." Data for T65/R83C/Q136C (black circles; n = 7 in presence of membranes; n = 4 in the absence of membranes) are taken from Fig. 5. In the presence of membranes, Pxlink for T65V/R83C/Q136C (gray circles; n = 3) is significantly different from that for T65D/R83C/Q136C (open circles; n = 3); p = 0.01 (Student's t test). Note that the S.E. is within the size of the symbol for T65V in the presence of membranes.

For the folding assay, our choice of indicator pairs (R83C/Q136C) was based on placing the cysteine pair far enough from the Thr65 site to avoid electrostatic consequences of a nearby negative charge (the carboxylate of aspartate) on ionization of SH to thiolate ions for the reaction with PDM. The validity of this strategy is shown by the complete block of R83C/Q136C/T65D by PDM, average FPDM = 0.97 ± 0.02 (n = 3). Thus, any effect on folding manifest at R83C/Q136C represents a large scale disruption of folding and not an effect on PDM labeling. The other indicator pairs (Q72C/G114C and T69C/Q112C) are in close proximity to Thr65 and thus are subject to both a charge effect on the cross-linking reaction and local misfolding. Both T65V and T65D in R83C/Q136C/BstEII-cut are translated equally, and both are targeted to ER membranes (data not shown); however, T65V folds, as determined by a Pxlink of 0.78 ± 0.01 (n = 3), whereas T65D does not fold very efficiently, Pxlink being 0.46 ± 0.07 (n = 3) (Fig. 6C). Pxlink for T65V is virtually identical to Thr65. Both mutated residues are available to cross-linking reagent, since the fraction of individual cysteines labeled by PDM is 0.97 ± 0.02 (n = 3) and 0.98 ± 0.01 (n = 3) for T65V and T65D, respectively, similar to the fraction obtained for the Thr65 construct (0.95 ± 0.02, n = 7). In the absence of membranes, neither Thr65, T65V, nor T65D fold completely. For all three intermediates, independent of the residue at position 65, Pxlink clusters between 0.16 and 0.34, with no significant difference (p = 0.17, one-way analysis of variance). The inability to fold completely in the presence of membranes correlates with the apparent inability to form tetramers. These results suggest that 1) folding and oligomerization are coupled, 2) oligomerization promotes completion of folding, and 3) lack of T65D folding is probably due to an oligomerization, not a folding, defect. Moreover, the ER membrane facilitates folding of Thr65, T65V, and T65D (Fig. 6).

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The Folding Assay-- If a protein cannot achieve its correct secondary, tertiary, and quaternary structure, it will fail to function properly. The journey from a nascent peptide chain to a mature protein passes through several stages of folding and oligomerization. What is the nature of this process, which involves coupled and causally related events? Reasonable approaches exist to probe this issue for soluble proteins that can be synthesized in milligram quantities; however, the physiologically relevant question is how does this occur inside the cell. For membrane proteins, this question is more difficult to investigate. A membrane protein is synthesized initially from mRNA in a ribosome-tRNA-protein complex in the cytosol. The entire complex is targeted to the ER membrane where synthesis continues. Our first task in elucidating mechanisms of secondary, tertiary, and quaternary Kv structure formation is to develop methods with which to assess each biogenic event. Toward this end, we designed a new biochemical folding assay that can be used on various biogenic species, from nascent peptide emerging from the ribosome to mature protein integrated into the membrane. For this purpose, we used an in vitro translation system, with the assumption that such a system recapitulates in vivo biogenesis (20-22). Vis à vis an electrophysiological approach, which only evaluates the functional status of the mature channel in the plasma membrane, the biochemical approach permits detection of electrically silent residues that are involved in folding and oligomerization of biogenic intermediates. Among the biochemical methods, we have opted for cross-linking and pegylation strategies to detect intramolecular folding. Alternative approaches were considered, namely the use of enzymatic or chemical cleavage of intervening loops between cross-linked cysteines. Neither Lys-peptidase nor furin worked under our experimental conditions. Chemical cleavage with hydroxylamine, which recognizes Asn-Gly bonds, also gave inefficient cleavage. These strategies require engineering cleavage sites, which may perturb the secondary and tertiary structure of the Kv1.3 protein and are context-dependent. Cysteine substitution may also affect folding, although this is likely to be less disruptive than introduction of cleavage sites.

The results presented in this paper validate the scheme shown in Fig. 2 for detecting intramolecular cross-linking of a folded T1 domain. This assay is based on detection of free cysteines and free maleimides in the protein. The intensities of the bands on NuPAGE gels can be quantified and probabilities can be calculated and statistically evaluated. Folded protein yields high, quite reproducible cross-linking probabilities for neighboring residues, whereas unfolded protein or distant pairs of cysteines yield low cross-linking probabilities.

In the majority of experiments, we get nearly complete pegylation with PEG-MAL and complete block with PDM but incomplete reaction with PEG-SH. This is true even for single cysteines and for a native cysteine (Cys71), which is exposed in the cytosol at a protein-aqueous interface (16) (data not shown). There are three possible explanations. First, PDM attached to protein could react with another SH group (e.g. a free small molecular weight protein or with the free amino acid cysteine) and thus be unavailable to react with PEG-SH. Second, the unbound maleimide of PDM could be partially hydrolyzed, rendering it unreactive to PEG-SH. Third, the purity of PDM may not be 100% bifunctional. In each of these cases, we would expect incomplete gel shifts of the Kv1.3-PDM-labeled protein with PEG-SH. However, we can quantitatively account for these fractions using single cysteine constructs and SDS experiments (see "Analysis of Pegylation Ladders"). Furthermore, a similar pegylation efficiency of Kv1.3 containing PDM-labeled Cys71 (16) indicates that the SDS micelle does not limit efficiency of pegylation, but rather one of the causes cited above is responsible.

Measurements were carried out at steady state, ensuring that when low probabilities of PDM labeling or PEG-SH pegylation were observed, they were not due to kinetic differences between folded and unfolded protein but represent true differences in the number of available peptidyl-maleimides.

Folding of T1-- For some proteins, oligomerization requires prefolded structures (e.g. influenza hemagglutinin protein and vesicular stomatitis virus G protein (23, 24)), whereas others oligomerize before synthesis of the protein is complete (e.g. thrombospondin (25)). Indeed, quaternary structure may catalyze or assist folding, as is true for procollagen (26), where oligomerization is required for complete folding. Finally, some oligomeric proteins may alternately use both strategies along the assembly pathway, as suggested by Papazian and co-workers for Shaker potassium channels (6). Previously, we have shown that T1 domains can tetramerize while still attached to ribosomes (9) and suggested that T1-T1 association may serve to globally restrict nascent peptide chains of individual subunits (27, 28). Similarly, Zerangue et al. (29) have proposed that T1 tetramerization promotes a high concentration of monomers for speedy tetramerization of the transmembrane parts of the channel. Consistent with this hypothesis is the observation that T1-deleted Kv mutants require high mRNA concentrations and long times to form channels (30, 31). We might even speculate that the T1 tetramer acts like a chaperone to assist folding of the transmembrane domains during tetramerization of the rest of the channel. Good precedents for such assisted folding of mature peptide are hormone and enzyme propeptides. Furthermore, it is possible that T1 aligns the nascent subunits in proper spatial arrangement for transmembrane intersubunit interactions. Thus, defects in T1 could cause misalignment due to inhibited T1 tetramerization.

Our pegylation assays directly demonstrate that nascent T1 monomers still attached to ribosomes are folded in ER membranes, which is consistent with previous results showing that self-associated T1 tetramers exist between ribosome-attached nascent subunits in ER membranes (9). Thus far, the oligomerization and folding assays suggest that much of the T1 domain is folded while the nascent peptide is attached to membrane-bound ribosomes (Fig. 7), without requiring complete synthesis of the Kv channel protein. Several specific regions, as defined by Kreusch et al. (14), are detected by these assays. For example, intramolecular cross-linking occurs between residues in layer 1 and layer 3 in the T1 monomer (R83C/Q136C) and between layer 1 and layer 2 (Q72C/G114C and T69C/Q112C). Intermolecular cross-linking occurs within layer 3 (R118C/D126C) and within layer 2 (R101C/D105C), consistent with proper folding of these regions of the T1 domain (9, 13). However, is folding prior to, or concomitant with, oligomerization of T1? To address this issue, we assessed whether T1 monomers could be intramolecularly cross-linked in the absence of membranes. The probability of cross-linking R83C/Q136C in the absence of membranes was increased compared with cross-linking of the predenatured protein (Fig. 5), consistent with T1 folding in the nascent monomer before it targets to ER membranes. However, the cross-linking efficiency is less than that observed for membrane-bound ribosome complexes.


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Fig. 7.   Folded regions of the T1 domain. Shown is a ribbon representation (in RasMol) of the T1 domain of Kv1.1a taken from the crystal structure of Kreusch et al. (14). Three distinct layers are indicated by red (layer 1), blue (layer 2), and yellow (layer 3) as originally assigned by Kreusch et al. Pairs of residues at the T1-T1 interface (labeled tetramerization) or the folded T1 monomer interface (labeled folding) are depicted as space-filling molecules and correspond, respectively, to Kv1.3 residues R118C/D126C and R101C/D105C, which were cross-linked in the tetramerization assay, and to R83C/Q136C, Q72C/G114C, and T69C/Q112C, which were cross-linked in the folding assay.

Four reasons could account for less efficient cross-linking. First, the absolute distance between cross-linkable cysteine pairs could be different in the absence of membranes. Second, the orientation of the cysteines could be less optimal for cross-linking than in the presence of membranes, especially given the possibility that the membranes might orient or restrict T1. Third, an auxiliary protein (e.g. a chaperone) could interact with membrane-free nascent peptide to prevent premature folding and tetramerization before targeting the nascent peptide-ribosome complex to the ER membrane. Fourth, the nascent peptide could be partially aggregated in the absence of membranes. The last possibility is unlikely because, for BstEII- and KpnI-cut intermediates, most of the protein is composed of the soluble NH2 terminus, and some hydrophobic transmembrane segments (S5 and S3, respectively) are buried inside the ribosome (19). Furthermore, despite additional transmembrane segments in the BstEII-cut intermediate, the cross-linking is similar to that for the KpnI-cut intermediate. Nonetheless, we conclude that T1 begins to acquire its folded tertiary structure in the nascent Kv1.3/ribosome complex before targeting to ER membranes. T1s partially fold but do not tetramerize until the membrane surface is involved (or possibly until a chaperone is removed). It is possible that tetramerization and further folding are coupled, cooperative events in the ER membrane. We have previously suggested that the ER membrane itself facilitates oligomerization (9). But what is the mechanism? Nascent peptide could interact with ER membrane protein translocating machinery or be partially integrated into the bilayer, or the N terminus could adsorb onto the lipid surface. Each scenario leads to concentration or restricted orientation of T1 domains and thus speedier oliogmerization.

One final consideration is that the folding assay provides insight into why Kv1.3/T65D fails to form a channel. The tetramerization assay indicates that this nonfunctional channel protein does not form tetramers, consistent with the failure of an isolated, soluble T1 from Kv1.2 containing the analogous mutation to form tetramers (as indicated by gel filtration chromatography) (15) and to fold correctly (as indicated by circular dichroism experiments) (15). Based on the T1 crystal structures (14, 15), several residues in the folded T1 monomer are expected to be within 3-5 Å of Thr65 and could contribute steric or electrostatic barriers when aspartate is substituted for threonine. The accommodation of isosteric valine at this position causes only a minimal change in structure of the isolated, folded T1 domain of Kv1.2 (15) and folds correctly in the nascent Kv1.3 peptide, yet substitution with aspartate prevents correct folding in both cases. In the absence of membranes, both T65V and T65D partially fold, as does the wild-type protein.

It is possible that when T1 is tethered to the rest of the Kv protein being synthesized on the ribosome, it is constrained and only able to fold partially. Neither complete folding nor oligomerization occur until it arrives at the ER membrane. This may be particularly limiting for mutations at residue 65. Chaperone factors and even the ribosome itself could alter T1 folding and oligomerization. We further speculate that only certain folded states are achieved at this biogenic stage, protecting against premature oligomerization before the channel protein reaches the ER membrane. Events at the translocon may continue folding the N terminus and/or allosterically induce an oligomeric-competent T1. Precedents exist in nonchannel proteins (32). It is also possible that folding and oligomerization are coordinated, cooperative events and that folding requires some degree of oligomerization. Nonetheless, our results eliminate the possibility that T65D malfunction is due to a pure oligomerization defect of a completely folded T1. Rather, T65D is arrested at a partially folded stage that fails to oligomerize. More complete folding may be coupled to oligomerization, which in turn is facilitated by the ER membrane. The membrane may even directly contribute to folding, as is suggested by the improved folding of T65D in the presence of membranes despite its inability to tetramerize.

Deliberate design of engineered pairs of cysteines might enable mapping of specific subdomains of the T1 monomer to determine which regions fold first or are capable of folding and which are more sensitive to intramolecular packing. Moreover, using the strategies outlined in this paper, we can investigate biogenic Kv intermediates of various lengths and their tertiary structures. This approach, including the development of a new assay for intramolecularly cross-linked monomers, not only will provide insight into the rules governing T1 folding but also is applicable to the study of folding of virtually any other protein.

    ACKNOWLEDGEMENTS

We thank Dr. LiWei Tu for contributing the electrophysiology experiments and John M. Robinson for the tetramerization assay of Thr65 mutants. We thank Dr. R. Horn for critical reading of the manuscript and extensive comments.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant GM 52302 and National Research Service Award HL-07027.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.

Dagger To whom correspondence should be addressed: Dept. of Physiology, University of Pennsylvania, Philadelphia, PA 19104-6085. Tel.: 215-898-8014; Fax: 215-573-5851; E-mail: cjd@mail.med.upenn.edu.

Published, JBC Papers in Press, November 12, 2002, DOI 10.1074/jbc.M209422200

    ABBREVIATIONS

The abbreviations used are: Kv, voltage-gated K+ channel; ER, endoplasmic reticulum; MOPS, 4-morpholinepropanesulfonic acid; PDM, ortho-phenyldimaleimide; PEG, polyethylene glycol; PEG-MAL, methoxy-polyethylene glycol maleimide; PEG-SH, methoxy-polyethylene thiol; PBS, phosphate-buffered saline.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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