Modulation of hexokinase association with mitochondria analyzed with quantitative three-dimensional confocal microscopy

Hexokinase isozyme I is proposed to be associated with mitochondria in vivo. Moreover, it has been suggested that this association is modulated in coordination with changes in cell metabolic state. To test these hypotheses, we analyzed the subcellular distribution of hexokinase relative to mitochondria in paraformaldehyde-fixed astrocytes using immunocytochemistry and quantitative three-dimensional confocal microscopy. Analysis of the extent of colocalization between hexokinase and mitochondria revealed that approximately 70% of cellular hexokinase is associated with mitochondria under basal metabolic conditions. In contrast to the immunocytochemical studies, between 15 to 40% of cellular hexokinase was found to be associated with mitochondria after fractionation of astrocyte cultures depending on the exact fractionation conditions. The discrepancy between fractionation studies and those based on imaging of distributions in fixed cells indicates the usefulness of using techniques that can evaluate the distributions of cytosolic enzymes in cells whose subcellular ultrastructure is not severely disrupted. To determine if hexokinase distribution is modulated in concert with changes in cell metabolism, the localization of hexokinase with mitochondria was evaluated after inhibition of glucose metabolism with 2-deoxyglucose. After incubation with 2-deoxyglucose there was an approximate 35% decrease in the amount of hexokinase associated with mitochondria. These findings support the hypothesis that hexokinase is bound to mitochondria in rat brain astrocytes in vivo, and that this association is sensitive to cell metabolic state

Ack1 is a dopamine transporter endocytic brake that rescues a trafficking-dysregulated ADHD coding variant

The dopamine (DA) transporter (DAT) facilitates high-affinity presynaptic DA reuptake that temporally and spatially constrains DA neurotransmission. Aberrant DAT function is implicated in attention-deficit/hyperactivity disorder and autism spectrum disorder. DAT is a major psychostimulant target, and psychostimulant reward strictly requires binding to DAT. DAT function is acutely modulated by dynamic membrane trafficking at the presynaptic terminal and a PKC-sensitive negative endocytic mechanism, or endocytic brake, controls DAT plasma membrane stability. However, the molecular basis for the DAT endocytic brake is unknown, and it is unknown whether this braking mechanism is unique to DAT or common to monoamine transporters. Here, we report that the cdc42-activated, nonreceptor tyrosine kinase, Ack1, is a DAT endocytic brake that stabilizes DAT at the plasma membrane and is released in response to PKC activation. Pharmacologic and shRNA-mediated Ack1 silencing enhanced basal DAT internalization and blocked PKC-stimulated DAT internalization, but had no effects on SERT endocytosis. Both cdc42 activation and PKC stimulation converge on Ack1 to control Ack1 activity and DAT endocytic capacity, and Ack1 inactivation is required for stimulated DAT internalization downstream of PKC activation. Moreover, constitutive Ack1 activation is sufficient to rescue the gain-of-function endocytic phenotype exhibited by the ADHD DAT coding variant, R615C. These findings reveal a unique endocytic control switch that is highly specific for DAT. Moreover, the ability to rescue the DAT(R615C) coding variant suggests that manipulating DAT trafficking mechanisms may be a potential therapeutic approach to correct DAT coding variants that exhibit trafficking dysregulation

Spontaneous transient outward currents arise from microdomains where BK channels are exposed to a mean Ca(2+) concentration on the order of 10 microM during a Ca(2+) spark

Ca(2+) sparks are small, localized cytosolic Ca(2+) transients due to Ca(2+) release from sarcoplasmic reticulum through ryanodine receptors. In smooth muscle, Ca(2+) sparks activate large conductance Ca(2+)-activated K(+) channels (BK channels) in the spark microdomain, thus generating spontaneous transient outward currents (STOCs). The purpose of the present study is to determine experimentally the level of Ca(2+) to which the BK channels are exposed during a spark. Using tight seal, whole-cell recording, we have analyzed the voltage-dependence of the STOC conductance (g((STOC))), and compared it to the voltage-dependence of BK channel activation in excised patches in the presence of different [Ca(2+)]s. The Ca(2+) sparks did not change in amplitude over the range of potentials of interest. In contrast, the magnitude of g((STOC)) remained roughly constant from 20 to -40 mV and then declined steeply at more negative potentials. From this and the voltage dependence of BK channel activation, we conclude that the BK channels underlying STOCs are exposed to a mean [Ca(2+)] on the order of 10 microM during a Ca(2+) spark. The membrane area over which a concentration \u3e or =10 microM is reached has an estimated radius of 150-300 nm, corresponding to an area which is a fraction of one square micron. Moreover, given the constraints imposed by the estimated channel density and the Ca(2+) current during a spark, the BK channels do not appear to be uniformly distributed over the membrane but instead are found at higher density at the spark site

Spontaneous Transient Outward Currents Arise from Microdomains Where BK Channels Are Exposed to a Mean Ca2+ Concentration on the Order of 10 μM during a Ca2+ Spark

Ca2+ sparks are small, localized cytosolic Ca2+ transients due to Ca2+ release from sarcoplasmic reticulum through ryanodine receptors. In smooth muscle, Ca2+ sparks activate large conductance Ca2+-activated K+ channels (BK channels) in the spark microdomain, thus generating spontaneous transient outward currents (STOCs). The purpose of the present study is to determine experimentally the level of Ca2+ to which the BK channels are exposed during a spark. Using tight seal, whole-cell recording, we have analyzed the voltage-dependence of the STOC conductance (g(STOC)), and compared it to the voltage-dependence of BK channel activation in excised patches in the presence of different [Ca2+]s. The Ca2+ sparks did not change in amplitude over the range of potentials of interest. In contrast, the magnitude of g(STOC) remained roughly constant from 20 to −40 mV and then declined steeply at more negative potentials. From this and the voltage dependence of BK channel activation, we conclude that the BK channels underlying STOCs are exposed to a mean [Ca2+] on the order of 10 μM during a Ca2+ spark. The membrane area over which a concentration ≥10 μM is reached has an estimated radius of 150–300 nm, corresponding to an area which is a fraction of one square micron. Moreover, given the constraints imposed by the estimated channel density and the Ca2+ current during a spark, the BK channels do not appear to be uniformly distributed over the membrane but instead are found at higher density at the spark site

Using Total Fluorescence Increase (Signal Mass) to Determine the Ca2+ Current Underlying Localized Ca2+ Events

The feasibility of determining localized Ca2+ influx using only wide-field fluorescence images was explored by imaging (using fluo-3) single channel Ca2+ fluorescence transients (SCCaFTs), due to Ca2+ entry through single openings of Ca2+-permeable ion channels, while recording unitary channel currents. Since the image obtained with wide-field optics is an integration of both in-focus and out-of-focus light, the total fluorescence increase (ΔFtotal or “signal mass”) associated with a SCCaFT can be measured directly from the image by adding together the fluorescence increase due to Ca2+ influx in all of the pixels. The assumptions necessary for obtaining the signal mass from confocal linescan images are not required. Two- and three-dimensional imaging was used to show that ΔFtotal is essentially independent of the position of the channel with respect to the focal plane of the microscope. The relationship between Ca2+ influx and ΔFtotal was obtained using SCCaFTs from plasma membrane caffeine-activated cation channels when Ca2+ was the only charge carrier of the inward current. This relationship was found to be linear, with the value of the slope (or converting factor) affected by the particular imaging system set-up, the experimental conditions, and the properties of the fluorescent indicator, including its binding capacity with respect to other cellular buffers. The converting factor was used to estimate the Ca2+ current passing through caffeine-activated channels in near physiological saline and to estimate the endogenous buffer binding capacity. In addition, it allowed a more accurate estimate of the Ca2+ current underlying Ca2+ sparks resulting from Ca2+ release from intracellular stores via ryanodine receptors in the same preparation

Imaging Ca2+ Entering the Cytoplasm through a Single Opening of a Plasma Membrane Cation Channel

Discrete localized fluorescence transients due to openings of a single plasma membrane Ca2+ permeable cation channel were recorded using wide-field digital imaging microscopy with fluo-3 as the Ca2+ indicator. These transients were obtained while simultaneously recording the unitary channel currents using the whole-cell current-recording configuration of the patch-clamp technique. This cation channel in smooth muscle cells is opened by caffeine (Guerrero, A., F.S. Fay, and J.J. Singer. 1994. J. Gen. Physiol. 104:375–394). The localized fluorescence transients appeared to occur at random locations on the cell membrane, with the duration of the rising phase matching the duration of the channel opening. Moreover, these transients were only observed in the presence of sufficient extracellular Ca2+, suggesting that they are due to Ca2+ influx from the bathing solution. The fluorescence transient is characterized by an initial fast rising phase when the channel opens, followed by a slower rising phase during prolonged openings. When the channel closes there is an immediate fast falling phase followed by a slower falling phase. Computer simulations of the underlying events were used to interpret the time course of the transients. The rapid phases are mainly due to the establishment or removal of Ca2+ and Ca2+-bound fluo-3 gradients near the channel when the channel opens or closes, while the slow phases are due to the diffusion of Ca2+ and Ca2+-bound fluo-3 into the cytoplasm. Transients due to short channel openings have a “Ca2+ spark-like” appearance, suggesting that the rising and early falling components of sparks (due to openings of ryanodine receptors) reflect the fast phases of the fluorescence change. The results presented here suggest methods to determine the relationship between the fluorescence transient and the underlying Ca2+ current, to study intracellular localized Ca2+ handling as might occur from single Ca2+ channel openings, and to localize Ca2+ permeable ion channels on the plasma membrane

A Close Association of RyRs with Highly Dense Clusters of Ca2+-activated Cl− Channels Underlies the Activation of STICs by Ca2+ Sparks in Mouse Airway Smooth Muscle

Ca2+ sparks are highly localized, transient releases of Ca2+ from sarcoplasmic reticulum through ryanodine receptors (RyRs). In smooth muscle, Ca2+ sparks trigger spontaneous transient outward currents (STOCs) by opening nearby clusters of large-conductance Ca2+-activated K+ channels, and also gate Ca2+-activated Cl− (Cl(Ca)) channels to induce spontaneous transient inward currents (STICs). While the molecular mechanisms underlying the activation of STOCs by Ca2+ sparks is well understood, little information is available on how Ca2+ sparks activate STICs. In the present study, we investigated the spatial organization of RyRs and Cl(Ca) channels in spark sites in airway myocytes from mouse. Ca2+ sparks and STICs were simultaneously recorded, respectively, with high-speed, widefield digital microscopy and whole-cell patch-clamp. An image-based approach was applied to measure the Ca2+ current underlying a Ca2+ spark (ICa(spark)), with an appropriate correction for endogenous fixed Ca2+ buffer, which was characterized by flash photolysis of NPEGTA. We found that ICa(spark) rises to a peak in 9 ms and decays with a single exponential with a time constant of 12 ms, suggesting that Ca2+ sparks result from the nonsimultaneous opening and closure of multiple RyRs. The onset of the STIC lags the onset of the ICa(spark) by less than 3 ms, and its rising phase matches the duration of the ICa(spark). We further determined that Cl(Ca) channels on average are exposed to a [Ca2+] of 2.4 μM or greater during Ca2+ sparks. The area of the plasma membrane reaching this level is <600 nm in radius, as revealed by the spatiotemporal profile of [Ca2+] produced by a reaction-diffusion simulation with measured ICa(spark). Finally we estimated that the number of Cl(Ca) channels localized in Ca2+ spark sites could account for all the Cl(Ca) channels in the entire cell. Taken together these results lead us to propose a model in which RyRs and Cl(Ca) channels in Ca2+ spark sites localize near to each other, and, moreover, Cl(Ca) channels concentrate in an area with a radius of ∼600 nm, where their density reaches as high as 300 channels/μm2. This model reveals that Cl(Ca) channels are tightly controlled by Ca2+ sparks via local Ca2+ signaling

Micro-Meta App: an interactive tool for collecting microscopy metadata based on community specifications

For quality, interpretation, reproducibility and sharing value, microscopy images should be accompanied by detailed descriptions of the conditions that were used to produce them. Micro-Meta App is an intuitive, highly interoperable, open-source software tool that was developed in the context of the 4D Nucleome (4DN) consortium and is designed to facilitate the extraction and collection of relevant microscopy metadata as specified by the recent 4DN-BINA-OME tiered-system of Microscopy Metadata specifications. In addition to substantially lowering the burden of quality assurance, the visual nature of Micro-Meta App makes it particularly suited for training purposes