Gaete PS et al. (2020), A novel voltage-clamp/dye uptake assay reveals ...

XB-ART-57467

J Gen Physiol 2020 Nov 02;15211:. doi: 10.1085/jgp.202012607.

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A novel voltage-clamp/dye uptake assay reveals saturable transport of molecules through CALHM1 and connexin channels.

Gaete PS , Lillo MA , López W , Liu Y , Jiang W , Luo Y , Harris AL , Contreras JE .

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Large-pore channels permeable to small molecules such as ATP, in addition to atomic ions, are emerging as important regulators in health and disease. Nonetheless, their mechanisms of molecular permeation and selectivity remain mostly unexplored. Combining fluorescence microscopy and electrophysiology, we developed a novel technique that allows kinetic analysis of molecular permeation through connexin and CALHM1 channels in Xenopus oocytes rendered translucent. Using this methodology, we found that (1) molecular flux through these channels saturates at low micromolar concentrations, (2) kinetic parameters of molecular transport are sensitive to modulators of channel gating, (3) molecular transport and ionic currents can be differentially affected by mutation and gating, and (4) N-terminal regions of these channels control transport kinetics and permselectivity. Our methodology allows analysis of how human disease-causing mutations affect kinetic properties and permselectivity of molecular signaling and enables the study of molecular mechanisms, including selectivity and saturability, of molecular transport in other large-pore channels.

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Species referenced: Xenopus laevis
Genes referenced: calhm1 gja8 gjb1 gjb2 lrrc8a
GO keywords: transporter activity [+]

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	Figure S1. Effect of antisense for Cx38 on ionic currents and dye uptake. (A) Representative traces of the ionic current observed in control noninjected oocytes or in oocytes injected with an antisense oligonucleotide to reduce the expression of endogenous Cx38 (ASCx38). Currents were recorded in a divalent cation-free Ringer solution at room temperature. (B) I-V relationship from data shown in A. (C) Quantification of currents recorded at +60 mV reveals that differences are statistically significant. (D) Time course of the ethidium uptake recorded in noninjected oocytes and oocytes injected with ASCx38. Ethidium uptake was evaluated at resting membrane potential in oocytes perfused with a divalent cation-free Ringer solution in the presence of 50 µM ethidium bromide. (E) Quantification of ethidium uptake rates is taken from data shown in D. All the experiments shown in this figure were performed in oocytes that did not express exogenous channels. *, P < 0.05 versus noninjected by unpaired Student’s t test. Error bars are SEM.
	Figure S2. DAPI does not saturate the number of binding sites available for ethidium intercalation. (A) Left: Representative recordings of fluorescence emitted by a drop (100 µl) of water (control; black trace), 1 µM ethidium bromide (EtBr; dissolved in water; red trace), 1 µM EtBr + 0.5 mg/ml salmon DNA (magenta trace), or 1 µM EtBr + 0.5 mg/ml salmon DNA + 100 µM DAPI (blue trace). Fluorescence was detected by a photomultiplier tube in a dark room at 20–22°C. The horizontal arrow indicates the baseline where the LED is turned off. The horizontal bar indicates the period when samples are excited (3 s; excitation [ex] filter, 530 nm; emission [em] filter, 600 nm). Right: Analysis of the fluorescence shown in the left panel. (B) Fluorescence detected in a microplate reader in a fluorescence spectrophotometer (Cary Eclipse). Plates containing 0.5 mg/ml salmon DNA were evaluated in the absence and presence of 100 µM DAPI. Fluorescence was analyzed before (0 EtBr) and after the addition of 1 µM EtBr. n.s., nonsignificant by unpaired Student’s t test. Error bars are SEM.
	Figure 1. A centrifugation procedure turns oocytes translucent and improves visualization of polynucleotide-dependent fluorescent dyes. (A) Centrifugal force renders animal pole translucent. NC, noncentrifuged. (B) Representative fluorescent signal (excitation filter, 350 nm; emission filter, 460 nm) detected in noncentrifuged oocytes injected with DNA plus DAPI or water (as a control). The fluorescent signal detected in centrifuged oocytes is shown in the right panel. Asterisks indicate the translucent zone in the animal pole. (C) Quantification of the fluorescent signal in centrifuged oocytes shown in B. , P < 0.05; **, P < 0.001 by one-way ANOVA plus Newman-Keuls post hoc test; n.s., nonsignificant. Error bars are SEM. (D) Representative pictures of YO-PRO or ethidium uptake in centrifuged oocytes. To allow unimpeded flux of dyes into the cytosol, oocytes were permeabilized with 0.01% Triton X-100 and then incubated 3 min with YO-PRO or ethidium (7.5 µM). The vehicle of YO-PRO and ethidium is also shown. Asterisks indicate the translucent zone in the animal pole. (E) Scheme of the TEVC/dye uptake assay. Oocytes were centrifuged to create a translucent zone in the animal pole. Then, oocytes were microinjected with 0.5 mg/ml salmon DNA to enhance the detection of polynucleotide-dependent fluorescent dyes. DAPI (100 µM) was coinjected to visualize the exogenous DNA. Finally, oocytes were placed in a chamber with the translucent zone (located in the animal pole) facing the objective of an inverted epifluorescence microscope. In parallel, the oocyte was impaled using the TEVC configuration to measure and control the membrane potential, as well as to determine the expression of exogenous large-pore channels by measuring the magnitude of ionic currents. Small fluorescent dyes were incubated or perfused using a gravity-dependent system. Dye uptake was recorded along the time course using a photomultiplier tube (PMT). (F) Concentration-response curve of ethidium uptake in translucent oocytes permeabilized with Triton X-100. Dye uptake experiments shown in D and F were performed using oocytes that did not express exogenous channels, at resting membrane potential (i.e., oocytes were not voltage-clamped), and perfused with a divalent cation-free Ringer solution.
	Figure S3. The gain of function of Cx30G11R channels is not associated with an increase in protein expression. (A) Representative Western blot of biotinylated wild-type Cx30 (Cx30WT) and Cx30G11R expressed in Xenopus oocytes. In addition to the biotinylated Cx30, we examined the total Cx30. Membrane containing total protein was stained with Ponceau to show the total protein loaded in each lane. (B) Densitometric analysis of biotinylated Cx30. (C) Densitometric analysis of total Cx30. Expression of Cx30WT and Cx30G11R was evaluated 24 h after cRNA microinjection. In these experiments, oocytes were microinjected with the same amount of cRNA (10 ng) for Cx30WT and Cx30G11R. n.s., nonsignificant by unpaired Student’s t test. Error bars are SEM.
	Figure 2. Centrifugation and DNA injection do not affect the biophysical properties of CALHM1 channels and connexin hemichannels. (A) Resting membrane potentials recorded in noncentrifuged (Non-cent) and centrifuged oocytes. Values were obtained from oocytes perfused with Ringer solution in the presence of 1.8 mM extracellular Ca2+. Membrane potentials were measured in noninjected oocytes and in oocytes expressing CALHM1, Cx26, or Cx30. (B) Representative current traces recorded during a voltage pulse (step) from −80 mV (holding potential) to 0 mV in CALHM1- or Cx26-expressing oocytes. The dashed line indicates 0 µA. Currents were measured in a Ringer solution containing 1.8 mM Ca2+. (C) Effect of extracellular Ca2+ on voltage-activated currents mediated by CALHM1 channels or Cx26 hemichannels. Tail currents were recorded using voltage-step pulses from −80 mV (holding potential) to 0 mV; duration step, 40 s. For experiments shown in B and C, oocytes were microinjected with BAPTA. Data in C were fit to a Hill equation to obtain Kd values. Error bars are SEM.
	Figure 3. The TEVC/dye uptake assay reveals that YO-PRO permeation through CALHM1 channels is saturable and Ca2+ sensitive. (A) Representative current traces recorded in oocytes expressing CALHM1 incubated with Ringer solution containing 100 µM Ca2+. The effect of 20 µM ruthenium red (Rut red), a CALHM1 blocker, is shown. The dashed line indicates 0 µA. NI, noninjected. (B) Representative time course of YO-PRO uptake in oocytes expressing CALHM1. Recordings were performed in a divalent cation-free Ringer solution and at resting membrane potential. (C) Analysis of YO-PRO uptake rates in NI oocytes (black squares) or in oocytes expressing CALHM1 (circles). YO-PRO uptake was evaluated in a divalent cation-free Ringer solution or in the presence of 1 mM extracellular Ca2+. All measurements were performed at resting membrane potential. (D) The uptake of 25 µM YO-PRO at resting membrane potential is inhibited by 20 µM Rut red. *, P < 0.05 versus control by one-way ANOVA plus Newman-Keuls post hoc test. (E) YO-PRO uptake rate is correlated to CALHM1 expression (determined as the magnitude of the tail current measured before dye uptake recording). †, P < 0.05 by t test. (F) YO-PRO uptake rates shown in C were normalized by CALHM1 expression (see Materials and methods). Data in F were fit to a Michaelis-Menten equation to obtain Vmax and Km. Error bars are SEM.
	Figure 4. Voltage dependence of YO-PRO uptake reveals that permeability of molecules and ionic currents in CALHM1 channels are differentially affected. (A) I-V relationship of CALHM1 channels. Oocytes were clamped at a holding potential of −80 mV, and depolarizing-step pulses from −40 mV to +70 mV were applied. The ionic current was evaluated in a divalent cation-free Ringer solution (red circles) or in the presence of 1 mM extracellular Ca2+ (black circles). For these experiments, oocytes were microinjected with BAPTA. (B) Voltage dependence of YO-PRO uptake (triangles). The uptake of 7.5 µM YO-PRO was evaluated at different controlled membrane potentials in the presence of 1 mM extracellular Ca2+. Dye uptake was recorded in clamped oocytes during 10 min to calculate dye uptake rates. The current shown in A was normalized to the current recorded in divalent cation-free Ringer solution. (C) Analysis of the effect of 1 mM extracellular Ca2+ on the ionic currents (atomic ion permeation) and the maximum flux of YO-PRO (molecular permeation), both measured at resting membrane potential. *, P < 0.05 versus 0 Ca2+ by paired Student’s t test. (D) Change in YO-PRO flux and ionic currents after Ca2+ removal from the extracellular bath illustrates the differential effect of Ca2+ on atomic ion permeation versus molecular permeation. Numbers indicate the fold change value. Error bars are SEM.
	Figure 5. Ethidium permeation through Cx30 hemichannels is saturable and has quantifiable kinetic properties. (A) Representative current traces recorded in oocytes expressing Cx30 incubated with a divalent cation-free Ringer solution. The effect of 200 µM La3+, a connexin hemichannel blocker, is shown. The dotted line indicates 0 µA. NI, noninjected. (B) Ethidium uptake rates detected in NI oocytes or in oocytes expressing Cx30 and Cx26. In these experiments, dye uptake was measured in oocytes incubated with a divalent cation-free Ringer solution plus 50 µM ethidium bromide. Ethidium uptake was evaluated at resting membrane potential in the absence or presence of 200 µM La3+. *, P < 0.05 by one-way ANOVA plus Newman-Keuls post hoc test. (C) Ethidium uptake rates measured in oocytes stimulated with a divalent cation-free Ringer solution at resting membrane potential. Ethidium uptake was evaluated in oocytes expressing Cx30 (green squares) and in NI oocytes, which do not express Cx30 (black squares). (D) Ethidium uptake is correlated to Cx30 expression (determined as the magnitude of the tail current measured before dye uptake recording). (E) Kinetic parameters of ethidium uptake after normalization of data shown in C by Cx30 expression (see Materials and methods). Data were fit to a Michaelis-Menten equation to obtain Vmax and Km. Error bars are SEM.
	Figure 6. The N-terminal region is a major determinant of the permselectivity of ethidium in connexin hemichannels. (A) Comparison of Cx30 and Cx26 monomers shows differences in the N-terminal domain (NT) and the intracellular loops (ICLs). (B) Molecular reconstruction of Cx30 hemichannels reveals high similarity to hemichannels formed by Cx26. Cx30 and Cx26 hemichannels are colored with blue and red, respectively. (C) Assessment of pore diameter of MD-equilibrated Cx26 and Cx30 hemichannels indicates similar pore width. (D) Voltage-activated current in oocytes expressing wild-type Cx30 (black trace) or Cx30G11R (red trace). The horizontal dashed line indicates 0 µA. (E) Ethidium uptake rates measured in oocytes expressing different levels of wild-type Cx30 (black circles) or Cx30G11R (red circles). (F) Voltage-activated current in oocytes expressing wild-type Cx26 (black trace) or Cx26N14K (red trace). The horizontal dashed line indicates 0 µA. (G) Ethidium uptake rates obtained from oocytes with different expression levels of wild-type Cx26 (black circles) or Cx26N14K (red circles). Wild-type Cx30 is permeable to ethidium, but the mutation G11R eliminates permeability to this dye. In contrast, wild-type Cx26 is not permeable to ethidium, but the mutation N14K turns the channel permeable to this molecule. The ionic currents shown in D and F were recorded from oocytes perfused with a Ringer solution containing 1 mM Ca2+, using a voltage-step pulse from −80 mV to 0 mV. For experiments shown in D and F, oocytes were injected with the same amount of cRNA. Dye uptake experiments were performed at resting membrane potential, using 7.5 µM ethidium bromide in the bath solution, and at zero extracellular Ca2+. Tail currents shown in E and G were obtained using a voltage-step pulse from −80 mV to 0 mV (40 s at zero extracellular Ca2+) before ethidium uptake measurements (see Materials and methods).

References [+] :

Best, Optimization of the additive CHARMM all-atom protein force field targeting improved sampling of the backbone φ, ψ and side-chain χ(1) and χ(2) dihedral angles. 2012, Pubmed