The C Terminus of Bax Inhibitor-1 Forms a Ca²⁺-permeable Channel Pore^*

Cell Culture

Mouse embryonic fibroblasts (MEF cells) were cultured at 37 °C in a 9% CO₂ incubator in DMEM/Ham's F-12 medium supplemented with 10% fetal calf serum, 3.8 mm l-glutamine, 85 units ml⁻¹ of penicillin, and 85 μg ml⁻¹ of streptomycin. All media were obtained from Invitrogen. DT40 cells were cultured as previously described (24).

For the ⁴⁵Ca²⁺ fluxes, MEF cells were seeded in 12-well clusters (Costar, MA, 4 cm²) at a density of ∼10⁴ to 2 × 10⁴ cells cm⁻². Experiments were carried out with confluent monolayers of cells (7.5 × 10⁴ cells cm⁻²) between the 5th and 7th day after plating. Lvec control cells were seeded in 12-well clusters (Greiner) at a density of 4 × 10⁴ cells/well, whereas IP₃R1-overexpressing L15 cells were seeded at a density of 6 × 10⁴ cells per well. These fibroblast L cell lines have been previously created and characterized (25, 26). Experiments were carried out on confluent cell monolayers between the 6th and 8th day after seeding.

Human cervical carcinoma cells HeLa T1 cells were cultured in DMEM containing 10% FBS, 2% Glutamax, 100 units ml⁻¹ of penicillin, and 100 units ml⁻¹ of streptomycin (Invitrogen). For studies of full-length BI-1, 3 × 10⁴ cells were seeded on 2-chamber borosilicate slides (Nunc) 1 day before transfection.

Peptides and Chemical Reagents

All synthetic peptides were obtained from ThermoFisher Scientific (Ulm, Germany) at a purity of more than 90%. The identity of the peptides was confirmed via mass spectrometry and the purifications were performed via HPLC. 2-Aminoethoxydiphenyl borate, ryanodine, and anisomycin were obtained from Sigma.

Topology Assay

Neuro2A cells were transfected with hemagglutinin-tagged (HA) BI-1 (HA-BI-1), BI-1-HA, ERp44-HA, or empty vector in 10-cm cell culture dishes at a confluence of 80–90% using the Attractene Transfection Reagent (Qiagen) according to the manufacturer's protocol. 24 h later, cells were harvested and the cell suspension was divided into three fractions. These fractions of suspended cells were permeabilized with either 20 μm digitonin or 0.1% Triton X-100 in PBS for 30 min on a rocking platform at room temperature or were kept unpermeabilized in PBS. For incubation with Triton X-100, cells were first fixed with 4% paraformaldehyde for 10 min at 37 °C. After permeabilization, cells were washed twice with either PBS containing 2 μm digitonin or 0.01% Triton X-100, or in PBS alone, and stained with Alexa Fluor 488-conjugated anti-HA tag antibody (Cell Signaling Technology) diluted 1:20 in wash buffer for 30 min on a shaker protected from light. After two wash steps cells were transferred to FACS tubes and analyzed on a FACS Calibur (BD Biosciences).

⁴⁵Ca²⁺ Fluxes

Unidirectional ⁴⁵Ca²⁺ flux experiments were performed essentially as previously described (27–29). The 12-well clusters were fixed on a thermostated plate at 25 °C on a mechanical shaker. The culture medium was aspirated, and the cells were permeabilized by incubating them for 10 min in a solution containing 120 mm KCl, 30 mm imidazole-HCl (pH 6.8), 2 mm MgCl₂, 1 mm ATP, 1 mm EGTA, and 20 μg ml⁻¹ of saponin. The non-mitochondrial Ca²⁺ stores were then loaded for 45 min in 120 mm KCl, 30 mm imidazole-HCl (pH 6.8), 5 mm MgCl₂, 5 mm ATP, 0.44 mm EGTA, 10 mm NaN₃, and 150 nm free ⁴⁵Ca²⁺ (28 μCi ml⁻¹). Then, 1 ml of efflux medium containing 120 mm KCl, 30 mm imidazole-HCl (pH 6.8), and 1 mm EGTA was added and replaced every 2 min. The indicated [IP₃] or 10 μm Ca²⁺ ionophore A23187 were added for 2 min, after 2, 10, or 20 min of efflux. At the end of the experiment, all ⁴⁵Ca²⁺ remaining in the stores was released by incubation with 1 ml of a 2% (w/v) SDS solution for 30 min. Concentration-response curves were fitted using Origin 7.0 (Northampton, MA) software using the Hill equation.

Mag-Fluo4 Measurements in DT40 and TKO Cells

These experiments were performed as described (30, 31). Briefly, DT40 and triple IP₃R-knock out (TKO) cells were grown to a density of about 1 × 10⁶ cells/ml⁻¹ and loaded with Mag-Fluo4 AM (20 μm) for 60 min at 20 °C. Cells were permeabilized by incubation with saponin (10 μg·ml⁻¹) for 4 min at 37 °C. After centrifugation, cells were resuspended in Mg²⁺-free cytosol-like medium (140 mm KCl, 20 mm NaCl, 1 mm EGTA, and 20 mm PIPES, pH 7.0), supplemented with 375 μm CaCl₂ and 10 μm carbonyl cyanide p-trifluoromethoxyphenylhydrazone, which inhibits mitochondrial Ca²⁺ uptake. Cells were dispensed into a 96-well black-walled assay plate pre-coated with poly-l-lysine and centrifuged to spin the permeabilized cells to the bottom of the wells. Cells were incubated with 90 μl of fresh Mg²⁺-free cytosol-like medium, supplemented with 375 μm CaCl₂ and 10 μm carbonyl cyanide p-trifluoromethoxyphenylhydrazone. Mag-Fluo4 signals were measured with FlexStation3 with automated additions (settings: excitation 490 nm, emission 525 nm, reading sensitivity normal (five readings), and photomultiplier tube sensitivity high). At 30 s, 10 μl of 15 mm Mg-ATP (1.5 mm final) was added to initiate Ca²⁺ uptake. After reaching steady-state store loading, 20 μl of 6 μm thapsigargin (1 μm final) was added and 40 s later, 20 μl of peptides were added at 7× final concentration in the medium. This medium contained a final concentration of 1 μm thapsigargin to block Ca²⁺ pumps and 10 μm carbonyl cyanide p-trifluoromethoxyphenylhydrazone to block mitochondrial Ca²⁺ fluxes.

Fura-2 for Measuring Cytosolic [Ca²⁺]

Thirty thousand HeLa T1 cells were seeded on borosilicate 2-chamber slides (Nunc). The next day, cells were co-transfected with 1.5 μl of FuGENE 6 (Roche Diagnostics) and 0.25 μg of pcDNA3.1-mCherry and 0.25 μg of pcDNA3.1 (mock), pcDNA3.1 myc-BI-1, or myc-BI-1-D213R. Thirty-six hours post-transfection, cells were loaded with 1.25 μm Fura-2-AM (Invitrogen) diluted in Krebs-Ringer solution (11 mm HEPES, 125 mm NaCl, 6 mm KCl, 3 mm MgCl₂, 13 mm glucose, pH 7.5) and prepared for microscopy. For recording cytosolic [Ca²⁺], we used a Zeiss AxioObserver.Z1 microscope with a Fluor ×20/0.75 objective, Sutter Lambda DG-4 monochromator and filter sets 21 and 62 (Zeiss) for Fura-2 and mCherry, respectively. Images were recorded with an AxioCamHsM camera in the AxioVision version 4.6 software physiology module. mCherry-positive cells were preselected for the recordings, and Fura-2 fluorescence was recorded every 2 s. For the chelation of Ca²⁺, we used 3 mm 1,2-bis(2-aminophenoxyl)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) (Invitrogen), which rapidly chelates the extracellular Ca²⁺. Release of Ca²⁺ from intracellular stores was induced with 4 μm thapsigargin, which specifically inhibits the activity of sarco/endoplasmic reticulum Ca²⁺-ATPase pumps at low micromolar concentrations. Ca²⁺ traces were calibrated by incubating the cells for 10 min, first with Krebs-Ringer solution containing 50 mm EGTA and 2 μm ionomycin (LC Labs), and then with 250 mm CaCl₂ and 2 μm ionomycin. Traces were then converted into [Ca²⁺] using the formula,

where K_d = 225 nm (32).

Western Blot

Cells from 2-chamber slides were trypsinized and lysed in RIPA buffer (20 mm Tris, 150 mm NaCl, 1.5 mm MgCl₂, 0.5 mm DTT, and 1% Triton X-100, pH 7.5) with 1× protease inhibitor cocktail (Roche Diagnostics). Protein concentration was determined with the Bradford method and 10 μg of total protein lysate was prepared in urea sample buffer (200 mm Tris, pH 6.8, 5% SDS, 0.1 mm EDTA, 8 m urea, 0.1% bromphenol blue, and 100 mm DTT). Samples were heated for 10 min at 37 °C before loading on a precasted NuPAGE 4–12% BisTris gel in MOPS buffer (Invitrogen). Proteins were transferred in running buffer (25 mm Tris, 190 mm glycine, 10% methanol) on a PVDF membrane. The membrane was blocked in TBS (10 mm Tris, 150 mm NaCl, pH 7.5) supplemented with 5% milk powder and 0.1% Tween 20. Expression of the Myc-BI-1 fusion protein was detected using an anti-Myc antibody (Sigma). The membrane was stripped with a mild stripping buffer (ThermoFisher Scientific) and reblotted with an antibody against glyceraldehyde-3-phosphate dehydrogenase (Sigma) as a loading control.

Electrophysiological Recordings from Planar Lipid Bilayers

Planar lipid bilayer were formed (3:1 mixture of phosphatidylethanolamine/phosphatidylserine lipids (Avanti Polar Lipids) dissolved in decane) on a 200-μm diameter aperture separating cis and trans chambers containing: (cis) 95 mm Tris-OH, 200 mm HEPES (pH 7.3, ∼300 mosmol; [Ca²⁺]_free = 10 nm) and (trans) 50 mm Ba(OH)₂, 55 mm HEPES (pH 7.3, ∼285 mosmol) solutions, respectively. Peptides dissolved in dimethyl sulfoxide were added to the cis chamber at gradually increasing concentrations, and stirred continuously between the periods of membrane current recordings that were done using the Axopatch 1 amplifier and pClamp6 software (Axon Instruments). The cis chamber was grounded. Experiments were done at room temperature. Holding potential (V_h) was 0 mV for all experiments except when indicated. Off-line current-trace filtering (500 Hz), calculations of the mean current passing by lipid bilayer (I_mean), and histogram calculations and statistics analysis were done using pClamp10 and OriginPro 7.5 (OriginLab) software.

The Seventh Transmembrane Domain of BI-1 Resembles a Pore

The attempt to attribute a Ca²⁺-channel function to BI-1 is hampered by unresolved exact topology of the BI-1 in the ER membrane. We have previously shown that the C terminus of BI-1 resides in the cytosol (16), which results in two different models (Fig. 1A). In model A, which is supported by TMpred, the N terminus is located in the ER lumen. In model B, which is bioinformatically supported by TMHMM, the N terminus is cytosolic and the semi-hydrophobic seventh transmembrane domain, which bears some homology to the loop domain of the voltage-gated sodium channel SCN5A, would dip into the membrane, creating a loop domain (Fig. 1B). Unfortunately, it is not possible to use the enhanced green fluorescent protein protease-protection assay used previously (16) to distinguish between these two models, because the addition of an N-terminal enhanced green fluorescent protein alters the function of BI-1,⁴ indicating that the bulky enhanced green fluorescent protein moiety at the N terminus might alter the topology or intracellular localization of BI-1.

We therefore modified the assay by using cells overexpressing proteins tagged with the very short HA tag and selective membrane permeabilization. Digitonin makes epitopes in the cytosol, but not in the ER lumen, accessible to fluorescent antibodies, which can be quantified by flow cytometry. Treatment with Triton X-100, in contrast, permeabilizes all membranes and leads to staining of both luminal and cytosolic epitopes. We overexpressed HA-BI-1 or BI-1-HA in Neuro2a cells and used the luminal endoplasmic-reticulum protein (ERp) 44-HA as negative control (Fig. 1C). Both HA-BI-1 and BI-1-HA resulted in a similar increase in immunoreactivity after digitonin treatment, whereas ERp44 staining never exceeded the level apparent in cells transfected with an empty vector. Permeabilization with Triton X-100, on the other hand, dramatically increased HA immunoreactivity in ERp44-HA-transfected cells and had no further significant effect on HA-BI-1 and BI-1-HA. These results favor model B, although it cannot be completely excluded that the fusion with HA itself may alter the topology of BI-1.

A C-terminal Peptide Representing the Putative Pore Domain of BI-1 Causes Ca²⁺ Release from the ER Ca²⁺ Stores

The model supported by the experiments in Fig. 1 suggested a possibility of the presence of a pore domain right before the proposed C-terminal domains of BI-1 interacting with IRE1α (12), G-actin (13), and NADPH-P450 reductase (15), and forming the pH-sensor domain (7). Hence, we developed a set of two overlapping peptides named CTP1 (HGDQDYIWHCIDLFLDFITV) and CTP2 (CIDLFLDFITVFRKLMMILAMNEKD) covering the residues following the 6th transmembrane domain, but excluding the pH-sensor domain of BI-1 (Fig. 1B).

The Ca²⁺-flux properties of these peptides were examined in unidirectional ⁴⁵Ca²⁺-flux assays in saponin-permeabilized MEF cells. This assay provides an accurate assessment of Ca²⁺-flux properties of channels under unidirectional conditions (28, 29, 33, 34). In short, after permeabilization of the MEF cells, ⁴⁵Ca²⁺ was loaded to a steady-state in the non-mitochondrial Ca²⁺ stores, which correspond largely to the ER. After irreversible inhibition of the Ca²⁺ pumps by thapsigargin, the Ca²⁺ leak from these stores can be measured under unidirectional conditions. The Ca²⁺ content of the stores gradually decreased with time under these conditions (Fig. 2A). The released Ca²⁺ divided by the amount of Ca²⁺ that was present in the store at that time (= fractional loss), was determined and plotted (Fig. 2B). This analysis provides an accurate quantitative analysis of the Ca²⁺ leak from the ER. In control conditions, the Ca²⁺ content of the stores gradually decreased (Fig. 2A, vehicle), whereas addition of 60 μm CTP1, but not of CTP2, accelerated the decrease in store Ca²⁺ content (Fig. 2A) and resulted in an increased Ca²⁺ leak from the ER (Fig. 2B). The concentration-dependent effects of CTP1 on the Ca²⁺ leak were obtained by adding different concentrations of CTP1 (5, 10, 20, 40, and 80 μm) after 10 min of efflux (Fig. 2C). These effects were normalized for the maximal releasable Ca²⁺, obtained by the addition of 10 μm A23187, a Ca²⁺ ionophore. The dose-response curve (Fig. 2D) was fitted by a logistic curve fitting, yielding an EC₅₀ value of about 30 μm and a power (steepness of the curve) of 3. This indicates that CTP1-induced Ca²⁺ release is highly cooperative, which is compatible with an oligomerization-dependent process and suggests that at least three CTP1 peptides are required to form a functional Ca²⁺-channel pore.

CTP1-induced Ca²⁺ Release Is Independent of the Presence of BI-1, IP₃Rs, RyRs, Presenilins, and Translocon

To elucidate whether the Ca²⁺ release mediated by CTP1 was caused by modulation of BI-1 or other documented Ca²⁺-release mechanisms, we examined the Ca²⁺-flux properties of CTP1 in the absence of BI-1, IP₃R, RyR, presenilin, and translocon activity. First, we used primary BI-1 knock-out MEF cells and found that the Ca²⁺ release mediated by CTP1 did not differ between primary wild-type and BI-1^−/− MEF cells (Fig. 3A). Second, using inhibitors of IP₃Rs (100 μm 2-aminoethoxydiphenyl borate), RyRs (100 μm ryanodine), and the translocon (100 μm anisomycin), we found that the Ca²⁺-release properties of CTP1 were not affected by these inhibitory compounds (Fig. 3B). Third, a recent study proposed presenilins as important Ca²⁺-leak channels from the ER (35), although direct modulation of IP₃R activity by presenilins may also contribute to the effect (36–38). However, we excluded the contribution of presenilins by examining the Ca²⁺-release properties of CTP1 in presenilin-1/2 double knock-out MEF cells (Fig. 3C). CTP1 mediated similar increases in the Ca²⁺ flux from the ER of wild-type and presenilin-1/2 double knock-out MEF cells, indicating that the Ca²⁺-release properties of CTP1 were not due to modulation or activation of presenilin Ca²⁺-channel activity. Fourth, the contribution of IP₃Rs in mediating CTP1-induced Ca²⁺ release was further directly assessed by using L15 fibroblasts overexpressing IP₃R1 and DT40 TKO cells. By analyzing the CTP1-induced Ca²⁺ release from the ER of permeabilized fibroblasts either expressing an empty vector (Lvec) or overexpressing IP₃R1 (L15), we found that CTP1 mediated similar increases in the Ca²⁺-leak rate from the ER of control fibroblasts and IP₃R1-overexpressing fibroblasts (Fig. 3D). Next, we monitored the (Ca²⁺)_ER by using ER-trapped Mag-Fluo4 in permeabilized TKO cells (Fig. 3E) and TKO cells ectopically overexpressing IP₃R1 using an automated Ca²⁺ assay (Fig. 3F). After loading the ER Ca²⁺ stores to steady-state by adding ATP for active Ca²⁺ uptake, an unidirectional Ca²⁺ leak from the ER was initiated by adding thapsigargin, and was measured by monitoring the ER Ca²⁺ concentration using the low-affinity Ca²⁺ dye Mag-Fluo4. Then, CTP1 was added and the change in the Ca²⁺-leak rate was measured. Consistent with our previous experiments, we found that CTP1 enhanced the Ca²⁺-leak rate in TKO cells and in TKO cells ectopically expressing IP₃R1. These results rule out that the effect of CTP1 is mediated through IP₃R Ca²⁺-release channels. Taken together, these experiments indicate that the mechanism of CTP1-induced Ca²⁺ release does not involve BI-1 or other documented Ca²⁺-leak or release mechanisms, like IP₃Rs, RyRs, translocon, or presenilins.

Source of CTP1-induced Ca²⁺ Release Is IP₃-sensitive Ca²⁺ Store

The ER, the main intracellular Ca²⁺ store, is IP₃ sensitive. Hence, we investigated whether depleting the IP₃-sensitive Ca²⁺ store by adding an activating concentration of IP₃ would abolish the CTP1-induced Ca²⁺ release. Importantly, the ability of CTP1 to mediate Ca²⁺ release was almost completely abolished after depletion of the IP₃-sensitive stores (Fig. 4A), indicating that the main source of the CTP1-induced Ca²⁺ release largely corresponds to the IP₃-sensitive store. This was further supported by experiments in which we compared the CTP1-induced Ca²⁺ release relative to the Ca²⁺ release induced by a maximal concentration (50 μm) of IP₃ (Fig. 4B). This analysis showed that the maximal amount of releasable Ca²⁺ by a saturating concentration of CTP1 (80 μm) was >80% of the maximal amount of releasable Ca²⁺ by 50 μm IP₃.

Ca²⁺-flux Properties of CTP1 Depend on Its α-Helical Structure and Hydrophobic Residues Clustered in the Peptide

First, we tested whether shorter fragments of CTP1 were able to promote Ca²⁺ release from the ER (Fig. 5A). Importantly, shortening the CTP1 peptide to smaller fragments, CTP1–1 (YIWHCIDLFLDFITV), CTP1–2 (HGDQDYIWHCIDLFL), and CTP1–3 (YIWHCIDLFL), completely abolished the Ca²⁺-release properties of CTP1, indicating that the entire CTP1 sequence conveys its functional activity.

Second, because the predicted secondary structure of the CTP1 peptide displays an α-helical structure (Fig. 5B) with a high hydrophobicity index (second half of CTP1; Fig. 5C), we examined whether the α-helical properties and residues in the hydrophobic stretch were important to promote Ca²⁺ release (Fig. 5D). Hence, we analyzed the double mutant CTP1-L210A,F211A in the unidirectional ⁴⁵Ca²⁺-flux assay. This peptide contains two altered residues in its hydrophobic stretch, but retains its α-helical properties (Fig. 5B). The mutated residues slightly reduce the hydrophobicity score of the second half of CTP1 (Fig. 5C). We found that CTP1-L210A,F211A was not able to induce Ca²⁺ release. Similar results were found with CTP1-L210G,F211G, which does not possess an α-helical structure (Fig. 5B). To examine whether the α-helical structure of CTP1 is essential for its activity, we created CTP1-C207G,I208G. This peptide contains the essential Leu-210 and Phe-211 residues, but it has a largely altered secondary structure (Fig. 5B). Importantly, this peptide was also not able to promote Ca²⁺ release. Finally, we developed a control peptide CTP1-C207A, which retains its α-helical structure (Fig. 5B) and its hydrophobicity score (Fig. 5C). As a consequence, this peptide was able to mobilize Ca²⁺ (Fig. 5D). These data indicate that both residues in the hydrophobic stretch as well as the α-helical structure of CTP1 are essential for its activity. Because the CTP1-L210A,F211A peptide still retains a very high hydrophobicity score, we hypothesize that the Leu-210 and Phe-211 residues may contribute to the oligomerization of the peptide in the membrane, which may be required for CTP1-mediated Ca²⁺ mobilization from the ER.

CTP1 Displays Channel-like Activity with Permeability for Divalent Cations

Our experiments indicated that CTP1 induces Ca²⁺-channel activity in ER membranes. Hence, to unequivocally demonstrate that CTP1 is capable to form divalent cation-permeable channel pores, we examined, using Ba²⁺ as a permeant ion, whether the CTP1 incorporation into planar lipid bilayers would initiate ionic currents transmitted by divalent cations. In a separate set of experiments, either CTP1 or CTP1-L210A,F211A mutant peptides were added at 10 μm increment concentrations to the cis compartment of the bilayer chamber containing a HEPES/Tris solution (free [Ca²⁺] buffered at 10 nm), whereas the opposite trans chamber contained 50 mm Ba²⁺. The exposure of the lipid bilayer to CTP1 incrementing from 10 to 30 μm for several minutes for each concentration was without effect, but the further increment to 40 μm resulted in the appearance of a progressively growing Ba²⁺ current (Fig. 6A, top). The current developed from zero to more than 20 pA within 3–8 min, with longer exposure resulting in disintegration of the bilayer, which prevented careful measurements of the concentration-response relationship at higher concentrations due to a continuously increasing nonstationary current. Such a current was not observed when the CTP1-L210A,F211A mutant peptide was tested at concentrations up to 50 μm (Fig. 6A, bottom). The threshold-like appearance of the CTP1-induced current might be explained by a model where CTP1 accumulation to a critical concentration within a lipid bilayer initiates a chain reaction of formation of ion-conducting channel assemblies. The biophysical properties of the CTP1-forming channels were not studied in detail due to the limited period of observation of clearly distinguishable single-channel openings before appearance of the complicated fluctuating pattern of the CTP1-multichannel activity (Fig. 6B). The current amplitudes analyzed during the first minute of quasi-stationary observation of CTP1 channel activity yielded a broadly distributed shallow peak on a histogram compared with a zero-centered peak for the CTP1-L210A,F211A mutant peptide, which revealed no channel activity for the latter (Fig. 6C). To obtain a meaningful quantitative parameter, a mean current recorded during the first minute of observation of the channel activity in the presence of 40 μm CTP1 was calculated in multiple bilayers and compared with the mean currents recorded in the presence of 50 μm CTP1-L210A,F211A (Fig. 6D), showing a significant ion-transporting capability of the CTP1-forming channels. To confirm the identity of the current-carrying ions in our asymmetric ionic conditions, varying the polarity of the membrane voltage during an interval of a relatively stable level of the CTP1-induced (40 μm) currents revealed a voltage-dependent current rectification, indicating a high permeability for Ba²⁺ and negligible permeability for large cations (Tris) or for large anions (HEPES) by CTP1-formed ion channels (Fig. 6E). The observation of a threshold concentration and time dependence of these bilayer experiments suggest that accumulation in the membrane and subsequent oligomerization of the peptide may be required for its pore properties. To underpin these observations, we performed additional biochemical experiments, showing that N-terminal biotinylated CTP1 was able to accumulate in ER microsomal membranes from HeLa cells within 5 min (supplemental Fig. 1A). The Ca²⁺-flux properties of biotin-CTP1 were confirmed in ⁴⁵Ca²⁺-flux assays (supplemental Fig. 1B). As a control, we used a biotin-CTP1 mutant peptide where four hydrophobic residues were replaced by arginines and found that this peptide did not accumulate in ER microsomal membranes and did not provoke ER Ca²⁺ release. Importantly, whereas biotin-CTP1 D209A did not cause ER Ca²⁺ flux, it potently accumulated in ER microsomal membranes, indicating that its lack of Ca²⁺-flux properties is not caused by deficient accumulation in ER membranes, but rather due to its lack to form Ca²⁺-permeable channel pores. Finally, we also found that biotin-CTP1 peptides may form oligomers in ER microsomal membranes of HeLa cells (supplemental Fig. 1C). These observations correlate well with the high cooperativity (steepness coefficient = 3) found in the CTP1 concentration-response curve in the ⁴⁵Ca²⁺-flux assay (Fig. 2, C and D).

Ca²⁺-flux Properties of CTP1 Are Evolutionarily Conserved in Animal but Not in Plant or Yeast Orthologs of BI-1

Because BI-1 is a highly evolutionarily conserved protein with orthologs in yeast, plants, and animals, we examined the Ca²⁺-flux properties of CTP1 obtained from its yeast, plant, and animal BI-1 orthologs (Fig. 7A). Using unidirectional ⁴⁵Ca²⁺-flux assays, we found that all CTP1 peptides derived from animal BI-1 orthologs were able to induce Ca²⁺ release, whereas CTP1 peptides derived from plant or putative yeast BI-1 orthologs were not (Fig. 7B). Interestingly, CTP1 from the zebrafish Danio rerio was less potent than CTP1 from other vertebrates. This may be due to the fact that the predicted propensity of zebrafish CTP1 to form an α-helix was reduced compared with mammalian CTP1 (supplemental Fig. S2). These observations again underpin the importance of the α-helical properties to form functional Ca²⁺-channel pores. In addition, we also analyzed the Arabidopsis thaliana CTP1 and found a high predicted probability for α-helical structure formation. This may indicate that its lack to induce Ca²⁺ release is not due to structural defects but presumably to the lack of the critical Asp residue at position 12 of the peptide. The CTP1 region of the animal BI-1 orthologs seems not to be conserved in the yeast BI-1 ortholog and a recent phylogenetic analysis also suggested that this protein is in fact closer related to the BI-1 paralog TMBIM4 (16). In plant BI-1 orthologs, the CTP1 region is ∼65% conserved, but CTP1 plant orthologs were, nevertheless, not able to provoke Ca²⁺ release. Hence, the Ca²⁺ channel-like activity of the CTP1 domain of BI-1 seems to be a property acquired later in evolution and is only present in animals.

Asp-209 and Asp-213 Are Key Residues in CTP1 Ca²⁺-Channel Pore

CTP1 contains four negatively charged residues (Asp-200, Asp-202, Asp-209, and Asp-213). Importantly, CTP1-D202R,D213R and CTP1-D200R,D209R did not display Ca²⁺-flux properties (Fig. 8A). To have a systematic assessment of the importance of each aspartate residue for the Ca²⁺-mobilizing properties of CTP1, we analyzed four single CTP1 mutant peptides (40 μm), in which each respective aspartate residue was changed into an alanine residue (Fig. 8, B and C). Remarkably, D209A and D213A mutations in the CTP1 peptide completely abolished its Ca²⁺-flux properties. D202A only partially inhibited the Ca²⁺-flux properties of CTP1, whereas Asp-200 did not contribute to the Ca²⁺-flux properties of CTP1. This indicates that the negative side chains of Asp-209 and Asp-213 are directly involved in the Ca²⁺ flux through the ER membranes, whereas Asp-202 might serve as an acceptor residue at the cytosol/ER membrane interface. In fact, these models correlate well with the predicted BI-1 structure, in which Asp-209 and Asp-213 are located in the loop domain inside the ER membrane, whereas Asp-202 may be at the interface of the cytosol with the ER membrane and Asp-200 at the cytosolic face.

Importantly, comparing animal CTP1 orthologs with plant CTP1 orthologs, we found that Asp-209, which is essential for the Ca²⁺-flux properties of human CTP1, is conserved among the animal CTP1 orthologs, but not among plant CTP1 orthologs, where it corresponds to a threonine residue. Therefore, we assessed human CTP1-D209T and found that this peptide also lacked Ca²⁺-flux properties (Fig. 8, B and C). Hence, these data indicate that the appearance of Ca²⁺-flux properties of animal CTP1 orthologs is probably due to the change of a critical plant threonine to an aspartate residue.

In Contrast to Full-length Wild-type BI-1, BI-1-D213R Is Not Able to Reduce ER Ca²⁺ Content

Next, we assessed the importance of the critical Asp-213-residue in the CTP1 region of full-length BI-1 for its Ca²⁺ leak and cell death-inhibiting function. For Ca²⁺ analysis, we co-transfected mCherry plasmid with empty vector, myc-BI-1, or myc-BI-1-D213R in HeLa cells loaded with Fura-2-AM. Only Ca²⁺ signals in cells containing the mCherry plasmid as a marker for successful transfection were analyzed. The Fura-2 ratio (F₃₄₀/F₃₈₀) was calibrated to obtain [Ca²⁺]. We also verified similar expression levels of Myc-BI-1 and Myc-BI-1-D213R using Western blotting with anti-Myc antibodies (Fig. 9A). In agreement with previous observations (6, 7), wild-type Myc-BI-1 significantly reduced the amount of thapsigargin-releasable Ca²⁺ (p < 0.005; n = 68 cells), confirming that BI-1 lowers [Ca²⁺]_ER by acting as a Ca²⁺-leak channel (Fig. 9, B and C). Remarkably, similarly to mock transfected cells (n = 61 cells), Myc-BI-1-D213R did not affect the thapsigargin-releasable Ca²⁺ (n = 64). These data indicate that Asp-213 is an essential residue for the Ca²⁺ leak property of BI-1.