Motility of astrocytic mitochondria is arrested by Ca2+-dependent interaction between mitochondria and actin filaments

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Motility of astrocytic mitochondria is arrested by Ca2+ -dependent interaction between mitochondria and actin filaments Elena Kremneva 1 , Mikhail Kislin 1 , Xiaoying Kang, Leonard Khiroug ∗ Neuroscience Center, University of Helsinki, P.O. Box 56 (Viikinkaari 4), FI-00014 Helsinki, Finland

a r t i c l e

i n f o

Article history: Received 3 January 2012 Received in revised form 9 October 2012 Accepted 14 October 2012 Available online xxx Keywords: Mitochondria Astrocytes Ca2+ imaging Time-lapse Fluorescence microscopy Fluorescent protein Cytoskeleton Motility Velocity Tracking

a b s t r a c t Motility of mitochondria, as well as their activity-dependent immobilization (“trapping”), is essential for neuronal function, but its regulation by cytoskeleton and relevance for glial cell signalling are unknown. Using time-lapse fluorescence imaging in rat cultured astrocytes, we evaluated the role of microtubules and actin filaments in motility of mitochondria in resting cells and during physiological or pathological Ca2+ elevations. We found that mitochondria were significantly more aligned with microtubules than with actin filaments. Mitochondria were highly mobile under resting conditions at low intracellular free Ca2+ concentrations ([Ca2+ ]i ). Activation of a moderate increase in [Ca2+ ]i by either low-dose ionomycin or ATP immobilized mitochondria significantly but reversibly, without affecting mitochondrial morphology. A larger dose of ionomycin caused irreversible arrest and fragmentation of mitochondria. Disruption of microtubules completely arrested mitochondrial motility, while disruption of actin filaments had no effect on the basal mitochondrial motility at resting [Ca2+ ]i levels but significantly reduced mitochondrial immobilization during [Ca2+ ]i elevations. These results suggest that: (i) motility of astrocytic mitochondria is inversely related to [Ca2+ ]i , (ii) mitochondria require intact microtubules for their motility, and (iii) elevated [Ca2+ ]i immobilizes mitochondria by strengthening their interaction with actin filaments. © 2012 Elsevier Ltd. All rights reserved.

1. Introduction Mitochondria are semi-autonomous intracellular entities that play pivotal roles in a plethora of cellular processes ranging from basic metabolic support to cytoprotection or programmed cell death. In addition to being primarily responsible for ATP synthesis, these organelles act as potent Ca2+ buffers or, in other circumstances, as source of cytotoxic Ca2+ overload [1]. Mitochondria are dynamic organelles constantly undergoing fission/fusion and moving rapidly from one cellular region to another in a complex surveillance-like fashion [2,3]. Mitochondrial motility is mediated by motor proteins stepping along cytoskeletal elements [4]. In certain cases, e.g. in yeast, mitochondria move exclusively along actin filaments [5]. In neurons and many other cell types, microtubules seem to provide the primary path for mitochondrial movement, which is mediated by a variety of motor proteins [4,5]. However, both actin filaments and microtubules have been implicated in regulation of mitochondrial motility in neurons [6].

∗ Corresponding author. Tel.: +358 9 191 57644; fax: +358 9 191 57620; mobile: +358 45 635 2270. E-mail address: leonard.khirug@helsinki.fi (L. Khiroug). 1 These authors have contributed equally to this work.

Neuronal function crucially depends on motility of mitochondria [7]. Thus, axonal transport provides supply of mitochondria to synaptic terminals and mediates recycling of damaged mitochondria [5]. Dendritic mitochondria move bidirectionally along the dendrite shafts but become immobilized, or “trapped”, near active synapses in a Ca2+ -dependent manner [8,9]. In astrocytes, the major type of glial cells in the CNS, we had previously observed that mitochondria are “trapped” near plasma membrane during Ca2+ influx [10]. Besides plasma membrane, a Ca2+ -dependent immobilization of mitochondria was observed near endoplasmic reticulum (ER) during Ca2+ release from the ER Ca2+ stores [11–14]. “Arrest” of mitochondria at specific subcellular sites by high Ca2+ microdomains appears to be a general phenomenon with a range of functional consequences in neurons [9,12] and likely in astrocytes. In glial cells, however, neither the exact mechanism underlying mitochondrial “trapping” nor the implications of the latter have been elucidated. Using time-lapse imaging of fluorescently tagged mitochondria along with intracellular free Ca2+ concentration ([Ca2+ ]i ) in rat cultured astrocytes, we found that at resting [Ca2+ ]i mitochondrial motility required intact microtubules but did not depend on actin filaments. ATP- or ionomycin-induced [Ca2+ ]i rise resulted in a strong, reversible arrest of mitochondrial motility, which persisted as long as [Ca2+ ]i remained elevated and was by significantly reduced by manipulations on actin filaments.

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Fig. 1. Co-staining of mitochondria and microtubules or actin filaments to reveal their co-localization and alignment. (A) A wide-field image of mitochondria labelled with mitoDsRed (red pseudocolor) appeared oriented along microtubules stained with Tubulin Tracker Green (green pseudocolor). Arrows indicate examples of such alignment. (B) A confocal image of mitochondria labelled by overexpression of mitoYFP (green pseudocolor) often appeared oriented perpendicular to actin filaments labelled by overexpressing LifeAct-RFP (red pseudocolor). Arrows indicate examples of such orthogonal positioning.

2. Results We visualized mitochondria by transfecting rat cultured astrocytes with mitochondria-targeted fluorescent proteins mitoDsRed (Fig. 1A) or mitoYFP (Fig. 1B). Either transfection resulted in brightly fluorescent mitochondria that exhibited a high level of motility under resting conditions in the absence of astrocyte stimulation (Suppl. Movie 1 available online). Mitochondrial shape and motility were similar between cells expressing fluorescent proteins and cells loaded with synthetic mitochondrial markers MitoTracker Red or MitoTracker Green (data not shown), suggesting that morphology and dynamics of mitochondria were not affected by either of these distinct visualization methods. Since the signal-to-noise contrast provided by mitochondria-targeted fluorescent proteins was superior to that offered by synthetic dyes, we chose the former approach for further investigation of mitochondrial motility mechanisms. To evaluate morphological relation between mitochondria and the major elements of cytoskeleton, dual labelling was used. We simultaneously visualized microtubules and mitochondria by Tubulin Tracker Green staining of live cells expressing mitoDsRed (microtubule colour-coded in green and mitochondria in red, Fig. 1A). In a separate group of experiments, actin filaments and mitochondria were visualized by astrocyte co-transfection with LifeAct-RFP and mitoYFP plasmids (actin colour-coded in red and mitochondria in green, Fig. 1B). Comparison of images obtained with these two approaches revealed that mitochondria were preferentially aligned with microtubules (Fig. 1A) rather than with actin filaments (Fig. 1B). Statistical analysis confirmed that the examples shown in Fig. 1 are representative: on average, 40.2 ± 4.1% of mitochondria were aligned with microtubules; in contrast, only 18.7 ± 3.0% of mitochondria were aligned with actin filaments. This difference was highly significant (P < 0.001; n = 10 and 14 for cells labelled with Tubulin Tracker Green and cells expressing LifeActRFP, respectively). These data suggest that astrocytic mitochondria are significantly more aligned with microtubules than with actin filaments. To address the role of microtubules and actin filaments in morphological dynamics of mitochondrial networks under resting and stimulated conditions, we used time-lapse imaging and established pharmacological tools that included nocodazole, Latrunculin B and ionomycin. We quantified the effects of these pharmacological agents on mitochondrial morphology using the automatic morphological subtyping method recently developed by C.N. Hsu and

co-workers [15]. We performed analysis of mitochondrial morphology by automated segmentation of mitochondria according to their shape by means of the MicroP software by Peng et al. [15]. Mitochondria were automatically classified in six groups: small globule, large globule, straight tubule, twisted tubule, branched tubule, and loops (Fig. 2A and B). We then pooled the first two groups, i.e. globules (both large and small), together and used their prevalence as an index of mitochondrial remodelling and fragmentation in response to stimulation with ionomycin (n = 17 for 1 ␮M ionomycin and n = 18 for 5 ␮M ionomycin). Application of 1 ␮M ionomycin for 2 min induced no change in the globule fraction (Fig. 2A and C), indicating that morphology of mitochondria remained unaffected by this moderate stimulation. In contrast, the stronger stimulus (5 ␮M ionomycin for 2 min) resulted in a highly significant (P < 0.001) increase in the globular fraction to as much as 0.9 (black bars in Fig. 2C). Absence of mitochondrial fragmentation in response to brief, low-dose application of ionomycin (1 ␮M) associated with transient [Ca2+ ]i elevations (see Fig. 5A below) is consistent with a physiological response of astrocytes [16]. Conversely, application of a high-dose of ionomycin (5 ␮M) induced remodelling and fragmentation of mitochondria similar to those caused by pathological [Ca2+ ]i elevations (see Fig. 6C below; cf. Tan et al. [16]). Next, we measured mitochondrial motility using time-lapse fluorescence imaging of mitoYFP-expressing cells. In particular, non-globular mitochondria (i.e. tubules and loops) had showed the most complicated motility patterns (Fig. 3 and Supplementary Movie 1 available online), ranging from directional back-and-forth movements (Fig. 3B) to turning along apparent curves (Fig. 3C) to branching, retraction or lateral swinging (Fig. 3D). For simplicity, we will here use the term “trains” to refer to the non-globular mitochondria (straight tubules, twisted tubules, branched tubules and loops). To obtain quantitative measurement of mitochondrial motility, we used two approaches: (i) a simple method of frame-byframe digital subtraction [10,11] and (ii) the single object tracking method. The former method provides a robust measurement of general motility of fluorescently labelled mitochondria, termed motility index [10]; the latter one allows semi-automatic tracking of individual mitochondria with Connected Components algorithm of Imaris (Bitplane) and calculating their average velocity (Fig. 4). We compared the mitochondrial motility index and average velocity under resting (unstimulated) and ionomycin-stimulated conditions (Fig. 4A–D, K–N versus E–H, O–S). Pretreatment with

Please cite this article in press as: E. Kremneva, et al., Motility of astrocytic mitochondria is arrested by Ca2+ -dependent interaction between mitochondria and actin filaments, Cell Calcium (2012), http://dx.doi.org/10.1016/j.ceca.2012.10.003

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Fig. 2. Automated analysis of changes in mitochondrial morphology induced by 1 or 5 ␮M ionomycin stimulation in untreated cell and in cells treated with Latrunculin B or nocodazole. (A and B) Mitochondrial subtypes in two representative untreated cells before and after application of ionomycin at 1 ␮M (A) or 5 ␮M (B). Individual mitochondria are colour-coded as follows: blue – small globules, yellow – large globules, green – straight tubules, orange – twisted tubules, purple – branched tubules, red – loops. (C) Ca2+ -dependent changes in the globular fraction of mitochondria that indicate mitochondrial remodelling and fragmentation in each group of cells. Statistical significance levels: *** corresponds to P < 0.001, non-significant difference is marked as n.s. and corresponds to P > 0.05.

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Fig. 3. Mitochondrial motility in cultured astrocytes under resting conditions. (A) Wide-field image of a cultured astrocyte expressing the mitochondria-targeted fluorescent protein mitoYFP. (B) Time-lapse sequence of zoomed-in images illustrating directional movement of mitochondrial “trains” indicated by arrows (zoom 2.5×; inter-image time interval 10 s). (C) Time-lapse image sequence illustrating a turning-like movement of the mitochondrial “train” (arrow) along a curved putative “track”. (D) Time-lapse image sequence illustrating branching behaviour of mitochondria that consisted in growth and retraction of mitochondria (arrow).

10 ␮M nocodazole resulted in disruption of microtubules and decreased the mitochondrial movement beyond detection by the image subtraction method (data not shown). Thus, general motility could not be estimated in nocodazole-treated cells. We therefore applied the MicroP-Imaris analysis to track individual mitochondria and quantify their velocity in cells treated with nocodazole. The treatment induced a decrease in mitochondrial velocity (1.42 ± 0.17 ␮m/min as compared to 2.91 ± 0.57 ␮m/min in untreated cells). Furthermore, the percentage of mitochondria which could be tracked and for which velocity could be calculated was only 3%. For comparison, in untreated and Latrunculin B-treated cells measurement of velocity was possible for as many as 15–20% of objects recognized as mitochondria. In contrast to microtubule disruption, depolymerization of actin filaments with 2.5 ␮M Latrunculin B affected neither the mitochondrial morphology (Fig. 2C) nor their motility (Fig. 5F) or velocity (Fig. 5G). Together, these observations are consistent with the notion that, in rat astrocytes mitochondrial “trains” use microtubules, rather than actin filaments, as “rails” for their directed movement. Similar observations have been made in neurons, as published previously [4,17,18].

Next, we explored the effects of selective manipulations with cytoskeleton on Ca2+ dynamics under stimulated and resting conditions. To measure intracellular Ca2+ , we loaded the mitoYFP-transfected astrocytes with the widely used Ca2+ sensitive indicator Fura-2 and monitored general motility of the mitoYFP-tagged mitochondria simultaneously with [Ca2+ ]i . To trigger [Ca2+ ]i elevations, we stimulated the cells through the bath perfusion application of ionomycin, the most selective Ca2+ ionophore [19]. When applied for 2 min at 1 ␮M, ionomycin triggered [Ca2+ ]i elevations similar in time course to those induced by the natural glio-transmitter ATP at 100 ␮M (Suppl. Fig. 1 available online). [Ca2+ ]i remained at a plateau level for the duration of the 2 min ionomycin application and then partly recovered (Fig. 5A). Ionomycin at 1 ␮M caused a significant reduction in the general mitochondrial motility index to 72 ± 2% of its pre-stimulus level (n = 11 cells; P < 0.001; Fig. 5D and F) and a decrease in mitochondrial velocity to 0.96 ± 0.08 ␮m/min (P < 0.001, Fig. 5G). Similarly, application of 0.1 mM ATP for 2 min produced a transient increase in [Ca2+ ]i (Suppl. Fig. 1A) and a decrease in the motility index to 83 ± 4% of the pre-stimulus level (n = 12 cells; P < 0.01; Suppl. Fig. 1B and F). This decrease in motility was partly reversible after

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Fig. 4. Ionomycin induced [Ca2+ ]i rise and arrest of mitochondrial motility estimated in the same cell using two complementary methods of frame-by-frame subtraction (A–H) and object tracking (K–S). (A–D, K–N) Shape and motility of mitochondria prior to application of 1 ␮M ionomycin. (E–H, O–S) Ionomycin application affects mitochondrial motility (G) but not their overall shape or brightness (E and F). Epifluorescence image of an astrocyte expressing mitoYFP (A), zoomed-in (B). (C, G) The “trace” image produced by digital subtraction of an epifluorescence image from the one immediately following it. (D, H) Colour-coded overlay of the original epifluorescence image (green) with the “trace” image (red). (K, L, O, P) Image segmentation with MicroP (K, O), zoomed-in (L, P). (M, R) Surfaces created in Imaris. (N, S) Tracked mitochondria (grey) with their tracked trajectories (red lines).

stimulation with 1 ␮M ionomycin and fully reversible after stimulation with ATP. Both stimulation protocols selectively affected the motility of mitochondria (Fig. 5F and Suppl. Fig. 1F) and had no effect on their shape or brightness (cf. Fig. 4B versus F), ruling out such adverse effects as deregulation of mitochondrial fission/fusion balance or irreversible intracellular acidification that would have quenched YFP fluorescence. We then addressed the role of actin filaments in Ca2+ dependent arrest of mitochondria. Disruption of actin filaments with Latrunculin B completely abolished the Ca2+ -dependent arrest of mitochondrial motility both upon low-dose ionomycin stimulation (Fig. 5B versus D, summarized in Fig. 5F) and ATP stimulation (Suppl. Fig. 1B versus D). Importantly, Latrunculin B incubation had no effect on the size of ionomycin- or ATP-induced [Ca2+ ]i rise (P > 0.8 for ionomycin and P > 0.6 for ATP; Suppl. Fig. 1E) and did not change the baseline pre-ionomycin motility of mitochondria (P > 0.05; Fig. 5F). In contrast to untreated cells, low-dose ionomycin stimulation did not change the mitochondrial velocity in Latrunculin B-treated cells (P > 0.05; Fig. 5G). In the presence of 1 ␮M ionomycin, mitochondrial velocity remained at a significantly higher level in Latrunculin B-treated than in untreated cells (P < 0.05). These data strongly suggest that interaction with actin filaments is required for mitochondrial immobilization associated with the transient [Ca2+ ]i elevations induced by 1 ␮M ionomycin and 100 ␮M ATP. They further indicate that the effects of actin filament disruption were downstream of the mechanisms responsible

for the Ca2+ rise, presumably at the level of Ca2+ -dependent interaction between mitochondria and actin. The actin-dependent mitochondrial immobilization induced by ATP or low-dose ionomycin was not associated with mitochondrial remodelling. We therefore asked whether 5 ␮M ionomycin, which efficiently triggered mitochondrial remodelling (Fig. 2C), would trigger mitochondrial immobilization and whether this immobilization would be dependent on actin. Stimulation with 5 ␮M ionomycin induced a [Ca2+ ]i elevation which was more sustained than that induced by 1 ␮M ionomycin application (Fig. 6C versus Fig. 5A). In the majority of experiments, 5 ␮M ionomycin eventually caused cell death, presumably due to the toxic effects of persistently high [Ca2+ ]i . In untreated cells, shortly before the loss of cell integrity we did observe a highly significant decrease in general motility of mitochondria (by as much as 75%; Fig. 6D and E) and in their velocity (Fig. 6F). The arrest of mitochondria was followed by remodelling and fragmentation (see Fig. 2C). In Latrunculin B-treated cells, we observed a significant drop in mitochondrial velocity induced by high-dose ionomycin stimulation (P < 0.05; Fig. 6F). Moreover, in the presence of 5 ␮M ionomycin there was no significant difference in mitochondrial velocity between Latrunculin B-treated and untreated (P > 0.05; Fig. 6F). In contrast, the low-dose ionomycin (1 ␮M) did not change the velocity of mitochondrial movements in Latrunculin B-treated cells (P > 0.05; Fig. 5G). In terms of general mitochondrial motility index, the effect of 5 ␮M ionomycin was similar to 1 ␮M in that the significant

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Fig. 5. Time course of low-dose ionomycin-induced Ca2+ rise (upper traces) and transient decrease in general motility of mitochondria (lower traces). (A) Averaged Fura-2 signal induced by 1 ␮M ionomycin application (horizontal bar) in untreated cells. (B) Averaged plot of motility index recorded in the same cells in response to the same ionomycin stimulation (horizontal bar). (C and D) Averaged Fura-2 and motility index changes in Latrunculin B-treated cells treated in response to 1 ␮M ionomycin. Note lack of significant difference in Fura-2 traces between control (A) and Latrunculin B-treated (C) cells, and a strong depression of ionomycin-induced changes in motility index (D and G) upon treatment with Latrunculin B. (E–G) Effect of low-dose ionomycin on [Ca2+ ]i (E), general motility of mitochondria (F) and average velocity (G) in untreated versus Latrunculin B-treated cells. Statistically significant differences are indicated by asterisks (*** corresponds to P < 0.001, ** to P < 0.01, * to P < 0.05), non-significant difference is marked as n.s. and corresponds to P > 0.05. Note that Latrunculin B treatment did not affect the amplitude of the ionomycin-induced [Ca2+ ]i rise (E), but completely abolished the Ca2+ -dependent mitochondrial arrest induced low-dose ionomycin (F,G).

mitochondrial immobilization observed in untreated cells (P < 0.01) was eliminated by Latrunculin B treatment (P > 0.05; compare Fig. 6E to Fig. 5F). Taken together, these results suggest that, during pathological [Ca2+ ]i elevations that eventually lead to mitochondria remodelling and fragmentation, the Ca2+ -induced immobilization of mitochondria is (at least in part) actin-independent. Finally, an interesting observation was made when time-lapse imaging of mitochondrial motility and immobilization was performed in cells transfected with the mCherry-actin plasmid (Suppl. Fig. 2 available online). When we applied 5 ␮M ionomycin to the cells expressing mCherry-actin, we found that the degree of mitochondrial immobilization by ionomycin-induced [Ca2+ ]i rise was reduced to an insignificant level (P > 0.05), while the baseline (preionomycin) motility was unaffected by mCherry-actin expression (P > 0.05). General motility of mitochondria in the presence of ionomycin was significantly higher in mCherry-expressing cells than in non-transfected cells (P < 0.05). This observation indicates that not only disruption but also modification of actin filaments may result in a loss of Ca2+ -dependent immobilization of mitochondria in stimulated astrocytes. 3. Discussion Our main observations are as follows: (i) mitochondria were highly mobile at low [Ca2+ ]i , whereas during a [Ca2+ ]i rise their motility was temporarily and partly arrested; (ii) intact

microtubules, but not actin filaments, were required for baseline mitochondrial motility at low [Ca2+ ]i ; (iii) astrocytic mitochondria were significantly more aligned with microtubules than with actin filaments; (iv) intact actin filaments were essential for Ca2+ -dependent immobilization of mitochondria. Based on these findings, we propose a model where, at low [Ca2+ ]i , mitochondrial “trains” move using microtubules as their “railroad tracks” without detectible interaction with actin filaments (“railroad ties”). During [Ca2+ ]i elevation, motility of mitochondria is halted as they become “anchored” to actin filaments due to their Ca2+ -dependent interaction with the latter. The Ca2+ -dependent “go/no-go” dichotomy with the “go” role assigned to microtubules and the “no-go” role for actin filaments, as outlined above for astrocytes, is essentially similar to the emerging picture in neurons. Indeed, the general consensus is that mitochondrial motility in neuronal processes is mediated primarily by microtubules but regulated by primarily actin filaments [4,5,8,20]. In an elegant study employing RNAi-mediated knockdown of myosin V and VI, Pathak and co-workers demonstrated that myosin-mediated interaction with actin opposes anterograde and/or retrograde movements of mitochondria along microtubules [21]. This conclusion is fully consistent with earlier studies where actin, along with neurofilaments, has been implicated in mitochondrial trapping (or anchoring) in both dendrites and axons. Thus, disruption of actin cytoskeleton increases mitochondrial motility [20] and NGF-dependent docking of axonal mitochondria is

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Fig. 6. Effect of high-dose (5 ␮M) ionomycin-induced Ca2+ rise on mitochondrial motility, average velocity in untreated cells and in cells with disrupted actin filaments. (A and B) Tracked mitochondria in a representative Latrunculin B-treated cell before (A) and after (B) 5 ␮M ionomycin application. Trajectory (red line) and surface (grey) obtained with Imaris form mobile mitochondria in the mitoYFP-expressing astrocyte. (C) Averaged Fura-2 signal induced by 5 ␮M ionomycin application in untreated cells. (D) Averaged plot of motility index recorded in the same cells in response to the ionomycin treatment. (E and F) Ca2+ -dependent changes in general motility (E) and average velocity (F) of mitochondria induced by 5 ␮M ionomycin application. Ca2+ -dependent arrest of mitochondrial motility becomes insignificant after actin filaments are disrupted. Statistical significance levels: *** corresponds to P < 0.001, ** to P < 0.01, * to P < 0.05.

actin-dependent [22]. Furthermore, it has been shown that actin mediates anchoring of mitochondria at the base of those dendritic spines that are actively involved in synaptic transmission [8]. In addition to the “no-go” role of actin, mitochondrial immobilization at neuronal compartments (e.g. synapses) has also been attributed to Ca2+ -dependent uncoupling of mitochondria from microtubules and/or microtubule-associated motor proteins [18]. The adaptor proteins Mitlon/TRAK and Miro1 play a key role in mitochondrial trafficking, and their Ca2+ -sensitive interactions with each other and/or microtubules are thought to be involved in immobilization of mitochondria in neurons [9,17,23]. Although our data are consistent with the rail track – railroad tie scheme, there is an alternative mechanism by which F-actin disruption can interfere with Ca2+ -triggered mitochondrial immobilization. Indeed, Ca2+ -triggered formation of F-actin may by itself reduce the mobility of mitochondria (and other organelles) merely by creating a physical constrain that hampers their movements, as reported recently for pancreatic duct epithelial cells [24]. This study is in agreement with our findings and reinforces the notion that Ca2+ -induced F-actin restructuring and/or its interaction with mitochondria play a central role in activity-dependent immobilization of mitochondria within a high [Ca2+ ]i microdomain. In the present study, we found that a pathological stimulus (5 ␮M ionomycin) may trigger an actin-independent mitochondrial arrest while milder, more physiological stimuli (ATP or 1 ␮M ionomycin) cause the mitochondrial immobilization that requires intact and unmodified actin filaments. Future experiments may clarify relative contributions by the actin-dependent and

actin-independent mechanisms to Ca2+ triggered arrest of mitochondria. It seems plausible however, that both mechanisms coexist and may engage in complex interactions depending on specific pathophysiological conditions as well as on sources and spatio-temporal profiles of Ca2+ elevations in neuronal compartments. In astrocytes, the mitochondrial motility and remodelling mechanisms remained largely unstudied until the recent work published by the Rintoul group [16]. In their study, Tan and co-workers applied varied doses of a Ca2+ ionophore to cultured cortical astrocytes and observed a graded response in terms of mitochondrial remodelling (also referred to as “rounding”) and fission. The authors concluded that large pathophysiological increases in [Ca2+ ]i induce mitochondrial rounding and immobilization, while the more modest increases result in remodelling but do not affect mitochondrial motility [16]. Our present results may appear somewhat inconsistent with those of Tan et al. [16]: indeed, we observed that both large and moderate rise in [Ca2+ ]i (induced by 1 ␮M and 5 ␮M ionomycin, respectively) resulted in decreased mitochondrial motility (Figs. 5 and 6), while remodelling was induced only by large but not moderate [Ca2+ ]i increases (Fig. 2). This discrepancy may be attributed to one or more of the following differences in our experimental setups: (i) use of two different ionophores (ionomycin versus 4Br-A23187); (ii) difference in extracellular Ca2+ concentration (2 mM in our study versus 1.4 mM in Tan et al.); (iii) possible differences in ionophore-induced [Ca2+ ]i elevations, which cannot be directly compared because Tan and colleagues used Fura-2 as a ratiometric dye, while we were constrained to its use as a

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single-wavelength indicator (see Section 4); (iv) temperature differences (37 ◦ C in our experiments versus room temperature in those of Tan et al. [16]), which must have had a significant effect on ATP-dependent, motor protein-mediated motility of mitochondria. Despite these differences, the main conclusions from our present work and that of Tan et al. [16] are congruent in that astrocytic [Ca2+ ]i controls mitochondrial motility and remodelling in a dosedependent manner. The general significance of actin-dependent immobilization of mitochondria in the regions of elevated Ca2+ (a.k.a. high Ca2+ microdomains; [25]) becomes particularly evident if one considers mitochondria themselves as mobile signalling microdomains. Indeed, interactions between mitochondria and other cellular organelles, such as ER or plasma membrane, have been shown to require close co-localization with these compartments [12]. The high Ca2+ microdomains, which are formed during Ca2+ influx from extracellular space [26] and efflux from ER [27–29], trigger “trapping” of mitochondria in the vicinity of plasma membrane [10,30,31] or ER [11]. Functional consequences of such trapping (or lack of it) must be profound, as mitochondrial Ca2+ uptake (which only can be activated if a mitochondrion is located in close proximity of the Ca2+ source; [10,32]) has been strongly implicated in cytoand neuro-protection [33,34]. In the CNS, inability of mitochondria to translocate towards high Ca2+ microdomains, get immobilized therein and take up excessive Ca2+ is likely to result in apoptotic cell death thus giving rise to neurological disorders [7,35,36]. Thus, the actin-dependent mechanism of mitochondrial immobilization by elevated Ca2+ characterized in the present study represents an attractive target for drug development aimed at a variety of unmet clinical needs including traumatic, neurodegenerative and neurodevelopmental disorders. 4. Materials and methods 4.1. Astrocyte cultures Primary hippocampal and cortical astrocytes were obtained from 27 neonatal (P2–P4) Wistar rat pups. Cells were dissociated with enzymatic treatment (0.5 mg/ml papain for 10 min) and plated on poly-l-lysine coated MatTek glass-bottom Petri dishes (100,000–120,000 cells per dish) in DMEM supplemented with 10% of foetal calf serum, penicillin 100 units/ml and streptomycin 100 ␮g/ml. Astrocytes were cultured for 1–1.5 weeks up to forming a continuous monolayer. During the imaging experiments, cells were continuously perfused using a peristaltic pump with a standard solution containing (in mM): NaCl 127, KCl 3, CaCl2 2, MgCl2 1.3, HEPES 20, glucose 10; pH was adjusted to 7.4 with NaOH. 4.2. Fluorescence imaging Cultured astrocytes were investigated using the CellR imaging system (Olympus Europe, Hamburg, Germany). Images were collected either in (i) wide-field epifluorescence mode using a 60X oil-immersion objective (Olympus, Japan) and a CCD camera (Orca, Hamamatsu, Japan), or in (ii) confocal mode using the HCX PL APO 63x/1.2 water-immersion objective and a confocal laser scanning microscope (Leica SP2 AOBS) with an argon (488 nm), DPSS (561 nm) lasers. To achieve physiological temperature and improve focus stability, the microscope frame and the culture dish mounted on it were maintained at a constant temperature of 37 ◦ C by means of an incubator (Solent Scientific, Segensworth, UK). Images were acquired every 10 s in a time-lapse mode. To visualize mitochondria and actin cytoskeleton, astrocytes were transfected either with the mitochondria-targeted yellow fluorescent protein (mitoYFP), the mitochondrial-targeted red

fluorescent protein (mitoDsRed) or with actin labelled with a red fluorescent protein (LifeAct-RFP) using standard Lypofectamine 2000 technique (Invitrogen Corporation) and imaged 12–18 h after transfection. To visualize microtubules, astrocytes were loaded with tubulin-Tracker-Green by a 30 min incubation with 100 nM dye at 37 ◦ C. For [Ca2+ ]i measurements, astrocytes were loaded by a 45 min incubation with 5 ␮M Fura-2 AM (Molecular Probes, USA) supplemented with 0.02% Pluronic F-127 (Sigma). Fura-2 fluorescence was excited at 380 nm and images were collected at 510 nm. Ratiometric Fura-2 imaging was not feasible due to poor transparency of the high magnification 60× objective at 340 nm. 4.3. Manipulations with microtubules, actin filaments and [Ca2+ ]i For disruption of microtubules, astrocytes were first incubated for 1 hour on ice, then with 10 ␮M nocodazole for 40 min at +37 ◦ C. Disruption of actin filaments was achieved by incubation with 2.5 ␮M Latrunculin B for 1 hour at +37 ◦ C prior to the imaging. To stimulate an increase in [Ca2+ ]i , astrocytes were perfused for 2 min with the standard solution with the addition of 0.1 mM ATP or ionomycin at either 1 ␮M or 5 ␮M. 4.4. Data analysis To analyze morphological relation between mitochondria and either microtubules or actin filaments, the number of aligned mitochondria was calculated manually in astrocytes stained with Tubulin Tracker Green or those expressing LifeAct-RFP, respectively. A given mitochondrion was ranked as aligned when at least half of its length was parallel to either a microtubule or an actin filament. To quantify general motility of mitochondria, a sequential image subtraction protocol was used [10,11]. The fluorescence changes for each pixel were calculated by subtraction of sequential images and application of median filter (3 × 3 pixels kernel). The number of pixels that exhibited a change (either positive or negative) greater than a fixes threshold (25% of the maximum fluorescence intensity/pixel) was calculated for each time point and normalized to the total area of mitochondria. The motility index obtained from such frame subtraction and averaging can range between 0 (corresponding to no motility) and 1 (maximal motility, i.e. translocation of all fluorescent mitochondria to a different position compared to the preceding time point). The motility index for each group of cells was normalized to the total area of mitochondria (to allow comparison between cells) and averaged over 1 min before ionomycin and over 1 min after. Morphological analysis of mitochondria was performed using the MicroP software written in Matlab kindly donated by Dr. J.Y. Peng and colleagues [15]. The software automatically classifies mitochondria into six subtypes: small globule, large globule, straight tubule, twisted tubule, branched tubule, and loops. Ratios of mitochondrial subtypes were calculated for n = 42 individual cells. Differences in the distribution of mitochondrial morphological features for each subtype were investigated to define effects of [Ca2+ ]i rise in each group of cells and whether the shape of each mitochondrial subtype is affected by the pretreatments. To quantify mitochondrial velocity, the images after final segmentation in MicroP program were used to create surfaces in Imaris (Bitplane AG). These surfaces were semiautomatically tracked over time with the connected components algorithm. Obtained tracks were used to calculate averaged velocity for individual mitochondria. The tracking approach relies on the clear recognition of individual mitochondria and has some limitations in accurately representing mitochondria population. In particular, rapidly moving, elongated and branched mitochondria are unsuitable for such analysis.

Please cite this article in press as: E. Kremneva, et al., Motility of astrocytic mitochondria is arrested by Ca2+ -dependent interaction between mitochondria and actin filaments, Cell Calcium (2012), http://dx.doi.org/10.1016/j.ceca.2012.10.003

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Images were quantified and processed using Imaris (Bitplane), AnalySIS (Olympus Biosystems), ImageJ (NIH) and ImagePro 5.1 (Media Cybernetics, Silver Spring, MD) software. Background level was subtracted prior to calculations. Plots were constructed using Origin 6.0 software (Microcal). Data from regions of interest delineating individual cells were pooled and presented as mean ± S.E.M. For analysis of normally distributed data, Student’s t-test was used. Otherwise, we used ANOVA. Acknowledgements The mitoYFP and mitoDsRed plasmids were a gift from Dr. A. Miyawaki, and the mCherry-actin plasmid was kindly donated by Dr. P. Lappalainen. The authors are grateful to Dr. A. Surin for his critical comments and suggestions. This work was supported by the Academy of Finland (grants number 135222 and 126321) and Center for International Mobility (CIMO), Finland. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.ceca.2012.10.003. References [1] M.R. Duchen, Mitochondria and Ca(2+)in cell physiology and pathophysiology, Cell Calcium 28 (5–6) (2000) 339–348. [2] M. Karbowski, R.J. Youle, Dynamics of mitochondrial morphology in healthy cells and during apoptosis, Cell Death and Differentiation 10 (8) (2003) 870–880. [3] I.R. Boldogh, L.A. Pon, Mitochondria on the move, Trends in Cell Biology 17 (10) (2007) 502–510. [4] R.L. Morris, P.J. Hollenbeck, Axonal transport of mitochondria along microtubules and F-actin in living vertebrate neurons, Journal of Cell Biology 131 (5) (1995) 1315–1326. [5] I.R. Boldogh, L.A. Pon, Interactions of mitochondria with the actin cytoskeleton, Biochimica et Biophysica Acta 1763 (5–6) (2006) 450–462. [6] L.A. Ligon, O. Steward, Role of microtubules and actin filaments in the movement of mitochondria in the axons and dendrites of cultured hippocampal neurons, Journal of Comparative Neurology 427 (3) (2000) 351–361. [7] G.L. Rintoul, I.J. Reynolds, Mitochondrial trafficking and morphology in neuronal injury, Biochimica et Biophysica Acta 1802 (1) (2010) 143–150. [8] Z. Li, K. Okamoto, Y. Hayashi, M. Sheng, The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses, Cell 119 (6) (2004) 873–887. [9] A.F. Macaskill, J.E. Rinholm, A.E. Twelvetrees, et al., Miro1 is a calcium sensor for glutamate receptor-dependent localization of mitochondria at synapses, Neuron 61 (4) (2009) 541–555. [10] J. Kolikova, R. Afzalov, A. Giniatullina, A. Surin, R. Giniatullin, L. Khiroug, Calcium-dependent trapping of mitochondria near plasma membrane in stimulated astrocytes, Brain Cell Biology 35 (2006) 75–86. [11] M. Yi, D. Weaver, G. Hajnoczky, Control of mitochondrial motility and distribution by the calcium signal: a homeostatic circuit, Journal of Cell Biology 167 (2004) 661–672. [12] R. Rizzuto, M.R. Duchen, T. Pozzan, Flirting in little space: the ER/mitochondria Ca2+ liaison, Science’s STKE 2004 (215) (2004) re1.

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Please cite this article in press as: E. Kremneva, et al., Motility of astrocytic mitochondria is arrested by Ca2+ -dependent interaction between mitochondria and actin filaments, Cell Calcium (2012), http://dx.doi.org/10.1016/j.ceca.2012.10.003