Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels are activated by membrane hyperpolarization and conduct an inward cation current, which contributes to rhythmic electrical activity of neural and cardiac pacemaker cells. HCN channels have been shown to undergo N-linked glycosylation, and the N-glycosylation has been shown to be required for membrane trafficking and possibly function. In this study, recombinant wild-type (WT) and glycosylation-defective N380Q HCN2 channels were individually or co-expressed in HEK-293 cells. We demonstrate that glycosylation is required for trafficking to the plasma membrane and for the stability of HCN channels in the cell. Interestingly, the heteromeric HCN2 channels of WT and glycosylation-defective N380Q have been observed on cell membranes, indicating that not all four subunits of a tetrameric HCN2 channel need to be glycosylated for HCN2 channels to traffic to plasma membranes. Subsequently, we investigate the effect of N-glycosylation on the function of HCN2 channels. We developed a fluorescence-based flux assay, which makes it possible to establish a negative potential inside liposomes to open HCN2 channels. Using this flux assay, we demonstrate that glycosylation-defective N380Q HCN2 channels reconstituted into liposomes function similarly to WT HCN2 channels. This suggests that N-glycosylation is not required for HCN2 channels to function.
The hyperpolarization-activated cyclic nucleotide-gated (HCN) ion channel is activated by hyperpolarization and is modulated by the binding of cyclic nucleotides (for reviews see [1,2]), throughout the nervous system and in the heart. When activated by membrane hyperpolarization, HCN channels conduct an inward cationic current termed Ih (or If in the heart) that is fundamentally involved in the control of pacemaking activity [3,4], resting membrane potential [5,6] and synaptic integration in dendrites [7,8]. Because these channels operate at the threshold of excitability, HCN channels have a significant impact on neuronal and cardiac electrical signalling . Besides the nervous system and the heart, HCN channels are also found in liver , kidney , ovary  and pancreatic cells . Although the physiological role of HCN channels in non-pacing cells is still uncertain, HCN channels have been suggested to control resting membrane properties, modulate synaptic transmission, mediate long-term potentiation and limit extreme hyperpolarization [13–15].
The HCN channel is a member of the six-transmembrane-segment (6TM) ion-channel family. The HCN channel is formed as a tetramer, and each subunit has six TM domains, denoted as S1–S6. Helices S5 and S6 and the pore loop form the ion conducting pore, and helices S1–S4 form the voltage sensor [16,17]. HCN channels also contain a cyclic nucleotide-binding domain (CNBD) in the large intracellular C-terminal region, which binds cytoplasmic cyclic nucleotides and is connected to the pore by a C-linker region that is conserved among HCN channels [18–20]. Binding of cyclic nucleotides to the CNBD shifts the voltage dependence of the HCN channel to more depolarized potentials, and the HCN channel activity is enhanced by the cAMP binding.
HCN channels were reported to be extensively N-glycosylated in the mouse brain and heart [21–23]. N-linked glycosylation in the endoplasmic reticulum (ER) and the Golgi complex is the most common post-translational protein modification in eukaryotic cells . Besides promoting protein folding, the glycans serve as recognition ‘tags’ allowing glycoproteins to interact with lectins, glycosidases and glycosyltranferases, thus affecting the fate of glycoproteins (e.g. trafficking and degradation). N-linked glycosylation promotes the proper folding, stability and oligomeric assembly of the channel proteins in the ER and facilitates transport and targeting to the plasma membrane [24,25]. In HCN channels, a highly conserved asparagine residue in the extracellular loop between helix S5 and the pore helix was identified to be glycosylated, and this post-translational channel modification was shown to be crucial for normal cell surface expression . The abrogation of N-glycosylation appears to abolish membrane expression of HCN channels in human embryonic kidney (HEK) cells  and CHO cells , and thus affects their function as measured by whole-cell patch clamp. It is unclear whether N-glycosylation of all four subunits of an HCN channel is required for its trafficking to the cell membrane. Additionally, there is no direct evidence showing whether glycosylation is crucial for HCN channels to function.
In the present study, we show that N-linked glycosylation is crucial for HCN2 membrane trafficking as well as intracellular lifetime. However, not all four subunits of a tetrameric HCN2 channel need to be glycosylated for HCN2 channels to insert into cell membranes. In order to directly study the effect of N-glycosylation on the channel function, we overexpressed both wild-type (WT) and glycosylation-defective mutant (N380Q) HCN2 proteins in HEK-293 cells, and purified and reconstituted HCN2 channels into liposomes. In this way, trafficking to the plasma membrane and channel function were decoupled. A new fluorescence-based flux assay was developed to assess the function of the reconstituted HCN2 channels. The results showed that N-glycosylation is not required for HCN2 channel function.
Full-length mouse HCN2 (mHCN2) cDNA (Gene ID: 15166) was amplified via PCR, and cloned into the mammalian expression vector pCDNA3.1 (Invitrogen) with a FLAG tag fused to the C-terminus of HCN2. The N380Q mutant was created by Quick-change mutagenesis method using WT HCN2 as a template. For fluorescent fusion protein constructs, WT HCN2 and N380Q were inserted into pEGFP, pECFP and pEYFP plasmid vectors (BD Biosciences, Clontech). All constructs were verified by DNA sequencing.
Cell culture, transient transfection and confocal fluorescence microscopy
HEK-293 cells were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 100 units/ml penicillin and 100 μg/ml streptomycin and incubated at 37°C with 5% CO2. For confocal microscopic analysis, glass coverslips were placed in 24-well dishes and 200,000 HEK-293 cells were seeded. HEK-293 cells were transiently transfected using Lipofectamine® 2000 transfection reagent (Invitrogen) according to the manufacturer's instructions. The pEGFP-HCN2, pEGFP-HCN2-N380Q, pECFP-HCN2 and pEYFP-HCN2-N380Q plasmids were either individually transfected or co-transfected into HEK-293 cells. After 48 h, cells were washed once with PBS and then fixed with 4% paraformaldehyde in PBS for 10 min at room temperature. After washing twice with PBS, membranes were stained with Alexa Fluor® 594-conjugated Wheat Germ Agglutinin (Molecular Probes), and nuclei were stained with DAPI (Vector Laboratories, Inc.). The coverslips were transferred to a microscopic slide and mounted using Permafluor® mounting medium (Immunotech). Mounted cells were stored at 4°C before microscopic studies. Confocal examinations were performed with a LSM 510 META (Zeiss) at wavelengths specific for individual fluorescent protein tags [EGFP, excitation at 488 nm and emission at 507 nm; ECFP, excitation at 450 nm and emission at 477 nm; EYFP, excitation at 514 nm and emission at 527 nm; Alexa Fluor® 594 conjugate of wheat germ agglutinin (WGA), excitation at 590 nm and emission at 617 nm; DAPI, excitation at 345 nm and emission at 455 nm].
Isolation of plasma membranes
Cells were seeded in a 150 mm dish and incubated until they reached ~70% confluency. Cells were first washed twice with giant plasma membrane vesicle (GPMV) buffer (10 mM Hepes, 150 mM NaCl, 2 mM CaCl2, pH 7.4), then incubated with GPMV buffer containing 2 mM N-ethylmaleimide (NEM, a vesiculant required for vesicle formation) at 37°C . One hour later, culture supernatants were collected and centrifuged at 1000 g for 15 min to remove cell debris and intact cells. Supernatants were further centrifuged at 20000 g for 1 h at 4°C to pellet GPMV membranes. GPMV membrane pellets were subsequently solubilized in SDS loading buffer [125 mM Tris, pH 6.8, 20% (v/v) glycerol, 4% (w/v) SDS, 2% (w/v) 2-mercaptoethanol, 0.001% Bromophenol Blue], and were resolved by Western blotting with anti-FLAG antibody.
Protein expression, purification and reconstitution
The FLAG-tagged WT HCN2 or N380Q mutant was transfected into HEK-293 cells, and the stable cell lines expressing either WT-HCN2 or N380Q-HCN2 were established. HCN2 protein was purified as previously described  with minor modifications. Briefly, cells were broken by sonication for 1 min; and sonication was repeated twice. Cell debris and nuclei were removed by spinning down at 1000 g for 15 min at 4°C. Supernatants were spun down at 40000 g for 1 h at 4°C. Subsequently, pellets were mixed with the extraction buffer [16 mM n-dodecyl β-D-maltoside (DDM), 200 mM KCl, 50 mM Tris/HCl, pH 7.4, 5 mM EDTA, 1× Protease Inhibitor Cocktail (P8340; Sigma)], and rotated for 2 h at 4°C. Then detergent-resistant membranes were spun down at 17000g for 40 min at 4°C. Supernatants were collected and mixed with FLAG-beads (anti-FLAG M2 affinity gel A2220; Sigma) and rotated at 4°C for 2 h for binding. After washing three times with 10 bead volumes of washing buffer [200 mM KCl, 50 mM Tris, pH 7.4, 5 mM EDTA, 4 mM DDM, 1× Protease inhibitor (Sigma P8340)], HCN2 proteins were eluted with 0.5 mg/ml FLAG peptide (F3290; Sigma) and concentrated. The purified HCN2 channels were reconstituted into liposomes as previously described  with some modifications. Briefly, the POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine) and POPG (1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1′-rac-glycerol)) lipid mixture (3:1 molar ratio) in chloroform was dried under nitrogen for 30 min, and rehydrated in buffer A (50 mM Tris/HCl, pH 7.4, 150 mM KCl, 30 mM DM) to a final concentration of 5 mM. The resulting solution was mixed with the purified HCN2 channels to a final protein concentration of 0.1 mg/ml (1:5000 protein:lipid molar ratio). The detergent was removed by serial-dialysis against the buffer containing 20 mM Tris, pH 7.4, 150 mM KCl and 2 mM/0.5 mM/0 mM DM at 4°C for 3 days (one day for each buffer). Empty liposomes were prepared in the same manner without the addition of protein prior to dialysis.
Fluorescence based flux assay for functional study of reconstituted HCN2 channels
Empty liposomes or proteoliposome were 100-fold diluted into a buffer containing 150 mM NaCl, 20 mM Tris/HCl, pH 7.4, and 2 μM 9-amino-2-methoxy-6-chloroacridine (ACMA; Sigma–Aldrich). This yielded a final K+ concentration of 1.5 mM outside liposomes. Fluorescence intensity was measured every 1 s using a SpectraMax fluorometer (Molecular Devices) for a total of 180 s with excitation at 395 nm and emission at 490 nm. The proton ionophore carbonyl cyanide m-chlorophenyl hydrazone (CCCP; Sigma–Aldrich) was added to a final concentration of 1 μM after 60 s, and the sample was gently mixed with a pipette. Valinomycin (Sigma–Aldrich) was added to a final concentration of 20 nM at 120 s. For blocking experiments, before the addition of ACMA, CsCl or ZD7288 was added to a final concentration of 10 mM or 50 μM, respectively.
HEK-293 cells stably expressing FLAG-tagged WT HCN2 proteins were transiently transfected with pEGFP-HCN2-N380Q using Lipofectamine® 2000 transfection reagent (Invitrogen) according to the manufacturer's instructions. After 48 h cultivation, GFP positive cells were isolated by FACS in an Aria III sorter and collected in DMEM. The sorted HEK-293 cells were cultured in DMEM supplemented with 10% fetal bovine serum, 100 units/ml penicillin, and 100 μg/ml streptomycin and 500 μg/ml of G418 disulfate salt (Sigma) for another 72 h. Cells were harvested and membrane fraction was obtained as described before. The detergent-solubilized membrane proteins were purified with 100 μl of anti-FLAG M2 affinity agarose beads and washed five times with 1 ml of washing buffer [200 mM KCl, 50 mM Tris/HCl, pH 7.4, 5 mM EDTA, 4 mM DDM, 1× protease inhibitor (Sigma P8340)]. Then, proteins were removed from beads with SDS loading buffer and analysed by Western blotting.
Cryo-EM imaging of HCN2 proteoliposome
A thin layer of carbon was floated on to C-Flat holey carbon TEM grids (EMS, PA). Six microlitres of proteoliposome solution was applied to the glow-discharged grid immediately after 3-day dialysis. After 1 min, grids were side-blotted by a piece of filter paper, and then plunged into liquid ethane, which was cooled by liquid nitrogen. Cryo-EM samples were stored in liquid nitrogen till observation. Cryo-EM images were recorded on an Eagle 4k × 4k camera (FEI) using a Tecnai G2 F20 microscope (FEI) at 200 kV with a 40 μm objective aperture. The electron dose for each exposure was approximately 2000 e−/nm2. Cryo-EM images were taken at −4 μm defocus, and the effective pixel size was 0.22 nm. The size of liposomes was analysed using a homemade Matlab program.
HCN2 channel is heterogeneously glycosylated
To study the function of the HCN2 channel, we overexpressed HCN2 protein in HEK-293 cells. There were always two bands of HCN2 protein observed before and after purification (Figure 1A). Previous studies have shown that the HCN2 protein is predominantly N-glycosylated in brain tissue and cardiac tissue [21,22,27]. To test whether the higher molecular weight band corresponds to the glycosylated form of HCN2, we used peptide N-glycosidase F (PNGF) to remove the N-linked glycan and monitored the migration of the HCN2 protein in Western blots (WB). As shown in Figure 1(B), the majority of the glycan was removed and the higher molecular weight band became invisible. The remaining major band was the lower molecular weight band, which corresponds to the non-glycosylated form of HCN2. Endoglycosidase H (EndoH) is widely used to monitor posttranslational modification in the Golgi apparatus. It cleaves asparagine-linked mannose rich oligosaccharides from glycoproteins in Golgi apparatus, until the enzyme Golgi alpha-mannosidase II removes two mannose subunits from glycoproteins. Thus EndoH is not able to cleave highly processed complex oligosaccharides from glycoproteins (i.e. glycoproteins translocated to the plasma membrane are resistant to EndoH cleavage). As shown in Figure 1(B), EndoH could not cleave HCN2 proteins, which indicates that HCN2 proteins were correctly processed through the ER and Golgi and were resistant to EndoH cleavage. After replacement of the asparagine at position 380 with a glutamine, which abolishes the N-glycosylation, the N380Q mutant showed only one band corresponding to the non-glycosylated HCN2 protein (Figure 1C). As a further confirmation, we used WGA-conjugated lectin beads to purify the glycosylated HCN2. As expected, the higher molecular weight band (i.e. glycosylated HCN2 protein) was enriched (Figure 1D). Surprisingly, there was always a weak band corresponding to the non-glycosylated HCN2 protein in the lectin-purified HCN proteins. As HCN2 channels are formed as tetramers, it is possible that some non-glycosylated subunits in hetero-tetrameric HCN2 channels were co-purified with glycosylated subunits by lectin beads. This observation raised the question of whether the HCN2 channels trafficked to the plasma membrane were homogeneously glycosylated. To answer this question, the HCN2 proteins trafficked to cell plasma membrane were purified from GPMVs, which was isolated by chemical vesiculant NEM from HEK-293 cells expressing WT HCN2 channels (Figure 2A). As shown in Figure 2(B), the non-glycosylated HCN2 band still existed in the plasma membrane fraction isolated from cells expressing WT HCN2. This suggests that not all four subunits in a HCN2 tetramer that is transported to cell membranes are glycosylated.
HCN2 protein expressed in HEK-293 cells is N-linked glycosylated.
Effect of N-linked glycosylation on HCN2 channel membrane trafficking and stability
N-glycosylation enhances protein stability
Compared with WT HCN2, the expression level of the N380Q mutant was much lower. Previous studies have shown that the inhibition of N-linked glycosylation reduced the stability of the acetylcholine receptor and Shaker K+ channel proteins [30–32]. We then sought to examine the stability of WT HCN2 and N380Q mutant in HEK-293 cells. Analyses of cells by Western blotting following treatment with the protein synthesis inhibitor cycloheximide showed an almost complete loss of the N380Q mutant after two hours (Figure 2C). In contrast, no significant decrease in WT HCN2 was observed after cycloheximide treatment. The finding that WT HCN2 was more stable when overexpressed in HEK-293 cells than the N380Q mutant suggests that N-glycosylation stabilizes HCN2 and protects it from degradation.
Some but not all of the four subunits in a tetrameric HCN2 channel need to be glycosylated for trafficking to cell membranes
N-glycosylation on proteins has been shown to affect membrane trafficking . By confocal microscopy, we observed that WT-GFP HCN2 resides in the cell plasma membrane (Figure 3A), whereas the glycosylation-defective N380Q mutant did not traffic to the plasma membrane (Figure 3B). This was consistent with the analysis of proteins purified from isolated plasma membranes from HEK-293 cells expressing WT HCN2 and the N380Q mutant. As shown in Figure 2(B), WT HCN2 can be detected in the plasma membrane isolated from cells expressing WT HCN2, but not from cells expressing N380Q mutant alone. However, the proteins purified from the plasma membrane isolated from WT HCN2 expressing cells always showed a non-glycosylated HCN2 band (Figure 2B), which suggests that not all four subunits in a HCN2 tetramer need to be glycosylated for membrane trafficking. To further investigate this, we constructed fusions of WT HCN2 with CFP and N380Q mutant with YFP, and co-expressed these two versions of the HCN2 proteins to probe the localization of hetero-tetrameric HCN2 channel. As shown in Figure 4, the N380Q mutant co-localized with WT HCN2 in both plasma membrane and cytosolic compartments. To further verify the co-localization of WT HCN2 and N380Q mutant on cell plasma membrane, we co-expressed WT HCN2 with FLAG tag and N380Q mutant with GFP tag, and carried out the co-immunoprecipitation experiment. As shown in Figure 2(D), WT HCN2 can pull down N380Q mutant from the membrane fraction indicating mutant N380Q can form hetero-tetrameric channels with WT HCN2 and traffic to the plasma membrane. All together, these data showed that WT HCN2/N380Q hetero-tetrameric channels could traffic to the plasma membrane, and that glycosylation of all four subunits in a tetrameric HCN2 channel is not required for trafficking to cell membranes.
Cellular localization of WT HCN2 and the N380Q mutant in HEK-293 cells
Co-expression of HCN2 WT and N380Q mutant in HEK-293 cells
N-glycosylation may not affect the function of HCN2 channel
It is still under debate whether N-glycosylation of HCN2 affects the function of the channel. In the present study, we initially tested the conductance of WT HCN2 and N380Q mutant channels by whole cell patch clamp. A strong Ih current for the cells expressing WT HCN2 channels was observed, whereas very low or no current was observed for N380Q channels when cells were hyperpolarized (results not shown). The lack of or low conductance from N380Q mutant channels might be due to the low expression and improper plasma membrane localization. To rule out the differences in cell expression that hindered probing the function of N380Q mutant channels, we developed a flux assay to assess the function of HCN2 channels reconstituted into liposomes.
It has been reported that the fluorescent dye ACMA could be used to assess the function of reconstituted potassium channel TRAAK and K2P1 [33,34]. K+ channels open at zero membrane potential, and K+ ions flow down the K+ gradient, establishing a negative potential inside liposomes (Figure 5A). Then H+ flows down the electrical gradient via the proton ionophore, CCCP, and quenches the fluorescence signal of the pH sensitive dye, ACMA. However, starting near zero membrane potential does not work for HCN channels as HCN channels are activated by membrane hyperpolarization (i.e. negative membrane potential). Instead, we developed a protocol to obtain a negative membrane potential to open HCN channels, as shown in Figure 5(A). When K+ ions flowed out of liposomes via valinomycin (K+ ionophore), a negative membrane potential was established inside liposomes. The negative membrane potential opened HCN2 channels, which allowed Na+ ions flow into liposomes. The influx of Na+ ions compensated the negative membrane potential, thus no quenching of the fluorescence signal occurred when functional HCN2 channels were present.
Functional study of reconstituted WT HCN2 and N380Q channels
Subsequently, we used this flux assay protocol to assess the function of reconstituted HCN2. The success of reconstitution of HCN2 was verified by Western blotting (Figure 5B). The resulting proteoliposomes are unilamellar vesicles (Figure 5C), and the size ranges from 10 to 100 nm in radius with an average radius of 30±14 nm (Figure 5D). In the empty liposomes experiencing a 100-fold K+ gradient across the lipid membrane, the addition of valinomycin induced an efflux of K+, which produced a negative potential inside the liposomes. The negative potential drove H+ influx via CCCP, resulting in a significant decrease in the fluorescence signal of the pH-sensitive dye ACMA (Figure 5E). In the presence of reconstituted WT HCN2 channels, the negative potential opened HCN2 channels, which caused an influx of Na+ and compensated the negative potential. Thus, there was no pH change inside the liposomes, and no quenching was observed (Figure 5E). The fluorescence quenching profile of reconstituted N380Q mutant channels was similar to that of the WT HCN2 channels (Figure 5E). This indicates that reconstituted N380Q mutant channels are able to open. To further verify the function of reconstituted WT HCN2 and N380Q mutant channels, we used the HCN2 blocker Cs+ to block the channel. With the addition of CsCl, an ACMA fluorescence quenching was observed for both WT HCN2 and N380Q mutant channels (Figure 5F). Similar results were obtained by using a HCN2 specific blocker, ZD7288 (Figure 5G). These results indicate that reconstituted WT and N380Q mutant HCN2 channels are functional and the N-glycosylation is not required for the opening of HCN2 channels.
Membrane proteins are usually glycosylated, and glycosylation plays important roles in protein folding, oligomerization, stability and trafficking [24,25]. Previous studies indicated that glycosylation could facilitate ion channel expression in cells, membrane trafficking [21,23,27], and the inhibition of N-linked glycosylation could alter the voltage dependence of ion channel gating in Kv1.1 and Kv-LQT1/mink K+ channels [35,36] and Na+ channels . It has been reported that HCN2 was glycosylated in mouse brain and other tissues, and when expressed in mammalian cells [21,22,27]. One study tested the co-opera-tion of heteromeric HCN family subunits and found co-transfection of N380Q HCN2 with HCN4 could rescue N380Q HCN2 to the plasma membrane . However, there is no direct or quantitative data to show how the N-glycosylation influences the HCN2 channel membrane surface localization. In the present report, we found that HCN2 channels were heterogeneously glycosylated. When the glycosylation-defective N380Q mutant was co-expressed with WT HCN2, the N380Q mutant could be targeted to the plasma membrane. Not all four subunits of a tetrameric HNC2 channel are required to be glycosylated for membrane trafficking to the cell membrane.
We noticed that when HCN2 was expressed in HEK-293 cells, the N380Q mutant produced less protein compared with WT HCN2, as observed by Hegle et al. . The origin of this difference has not yet been delineated. In this report, we monitored the half-life of WT HCN2 and the N380Q mutant after the protein synthesis was inhibited by cycloheximide treatment. We found that the half-life of N380Q was significantly shorter than that of WT HCN2. Although N-linked glycosylation is generally believed to protect proteins from proteolytic degradation, the specific proteolytic system involved in the rapid degradation of non-glycosylated proteins is still not fully understood. The accelerated degradation of non-glycosylated acetylcholine receptor involved an intracellular lysosomal proteolytic system , whereas the rapid degradation of non-glycosylated Shaker K+ channel protein involved cytoplasmic proteasomes . The origin of the low expression level of N380Q mutant may lie in the quality-control processes on newly synthesized protein or an enhancement of proteolysis resistance by glycosylation. The quality-control processes that newly synthesized proteins undergo en route to the plasma membrane is still awaiting for further investigation.
When the N380Q mutant was co-expressed with WT HCN2, the glycosylation-defective N380Q mutant could be targeted to the plasma membrane. This indicates the tetrameric channel assembly occurs in the cytosolic compartments. As long as there are some glycans attached to the tetrameric HCN2 channel, the HCN2 channel could traffic to plasma membrane. This has been confirmed by three types of experiments: (1) there are non-glycosylated HCN2 proteins in lectin-purified HCN2 proteins (Figure 1D); (2) there are non-glycosylated HCN2 proteins in proteins purified from isolated plasma membranes (Figure 2B); (3) WT-HCN2/N380Q hetero-tetrameric HCN2 channels could traffic to the plasma membrane (Figures 2D and 4). It will be intriguing to investigate how many HCN2 subunits are glycosylated in the native HCN2 channel and to what extent glycosylation is required for the membrane trafficking of HCN2 channel.
To investigate the effect of glycosylation on HCN2 channel function, whole-cell patch clamping was employed, and a severe reduction in current for glycosylation-defective N380Q mutant was observed. This might be due to the observation by us and Much et al.  that the N380Q mutant could not be targeted to the plasma membrane. In the present study, we studied the effect of glycosylation on HCN2 channel function using reconstituted HCN2 channels. The use of reconstituted channels bypassed the issues of membrane trafficking. There are a few ways to measure the function of reconstituted ion channels. One way is to form giant unilamellar proteoliposomes and carry out patch-clamping measurements. This method usually involves dehydration of lipid membranes, which may be detrimental to membrane associated proteins. Another method is to carry out black lipid membrane measurements by fusing proteoliposomes with a pre-formed lipid bilayer. This method requires a lot of practice and can measure the function of a few copies of reconstituted ion channels. In the present study, we developed a flux assay to monitor the function of reconstituted HCN2 channels in a defined condition. This method was based on a method used to detect the function of the reconstituted potassium channels TRAAK and K2P1. In contrast with most voltage-gated potassium channels (e.g. TRAAK and K2P1) [33,34], HCN2 channels are not open at zero voltage. Thus, we designed a protocol to establish a negative potential inside liposomes to open HCN2 channels. The effectiveness of this method was verified by the observation that the amplitude of fluorescence quenching depends on the K+ gradient (i.e. negative potential) across the lipid membrane (results now shown). The flux assay data of the reconstituted HCN2 channels clearly showed that N-glycosylation-defective N380Q channels could be functional. This is the first direct experiment showing that N-linked glycosylation is not required to open the HCN2 channel.
We thank Dr William Zagotta (University of Washington) for mHCN2 cDNA and for use of the fluorometer. We also thank Dr Dale Hailey for assistance with confocal microscopy and Dr David Raible for the use of the confocal microscope.
Mo Li and Liguo Wang designed the experiments, analysed the data and co-wrote the paper. Mo Li generated HCN2 N380Q mutant, purified recombination proteins, carried out glycosylation assays and co-localization of WT HCN2 and N380Q mutant channels and chased the proteins half-life in vivo. Mo Li and Lige Tonggu reconstituted proteins into liposomes and co-developed the flux assay method. Lige Tonggu collected all negative stain and cryo-EM images of proteoliposomes. Lan Tang cultured HEK-293 cells, purified HCN2 proteins and carried out the budding experiment.
The present work was supported by the US National Institutes of Health (NIH) [grant number R01GM096458 (to L.W.)].
carbonyl cyanide m-chlorophenyl hydrazone
cyclic nucleotide-binding domain
Dulbecco’s modified Eagle’s medium
giant plasma membrane vesicle
hyperpolarization-activated cyclic nucleotide-gated
human embryonic kidney
wheat germ agglutinin