PKCα: a versatile key for decoding the cellular calcium toolkit

Conventional protein kinases C (cPKCs) play an essential role in signal transduction and are believed to integrate both global Ca2+ transients and diacylglycerol signals. We provide evidence that PKCα is a ubiquitous readout sensor for the cellular Ca2+ toolkit, including highly restricted elementary Ca2+ release. Threshold stimulations of cells with Ca2+-mobilizing agonists resulted in PKCα translocation events with limited spatial spreads (<4 μm) comprising two groups of lifetimes; brief events (400–1,500 ms) exclusively mediated by Ca2+–C2 domain membrane interactions and long-lasting events (>4 s) resulting from longer DAG-C1a domain–mediated membrane interactions. Although upon uncaging NP-EGTA, which is a caged Ca2+ compound, WT-PKCα displayed rapid membrane translocations within <250 ms, PKCα constructs with C2 domains mutated in their Ca2+-binding region lacked any Ca2+-dependent translocation. Flash photolysis of diazo-2, a photosensitive caged Ca2+ buffer, revealed a biphasic membrane dissociation (slow and fast period) of WT-PKCα. The slow phase was absent in cells expressing PKCα-constructs containing mutated C1a-domains with largely reduced DAG binding. Thus, two groups of PKCα membrane interactions coexist; C2- and C1a-mediated interactions with different lifetimes but rapid interconversion. We conclude that PKCα can readout very fast and, spatially and temporally, very complex cellular Ca2+ signals. Therefore, cPKCs are important transducers for the ubiquitous cellular Ca2+ signaling toolkit.


Introduction
Ca 2+ is a versatile and ubiquitous messenger in all living cells. Increases in the intracellular Ca 2+ concentration occur as longlasting global oscillations, such as those that occur during clonal selection of white blood cells. In contrast, Ca 2+ increases also occur as transients with a very short lifetime and high spatial confi nement, so-called elementary Ca 2+ signals (Berridge, 2002;Berridge et al., 2003). All Ca 2+ signals require proteins to relay the Ca 2+ concentration downstream to different signaling networks. These specialized sensors contain Ca 2+ -binding domains and include such ubiquitous proteins as calmodulin, the structure of which comprises four highly conserved binding domains, the so-called EF hands (Vetter and Leclerc, 2003). Another very important Ca 2+ -binding domain is the C2 domain; e.g., during synaptic signaling, where it functions as the Ca 2+ -sensing domain of Munc-13 (Rettig and Neher, 2002).
A subfamily of the PKCs, the conventional PKCs (cPKCs), also contain C2 domains and, hence, have the principle ability to decode cellular Ca 2+ transients (Kohout et al., 2002). This family belongs to the AGC kinases (Manning et al., 2002) and comprises 12 isoforms, which are distinguished depending on their sensitivity toward signaling molecules.
The intrinsic function of PKC isozymes is regulated by multiple mechanisms (Newton, 2001), and they decode a variety of stimulation-induced intracellular messengers (Oancea and Meyer, 1998). PKCs are believed to translate messengers into the phosphorylation of many target proteins, such as ion transporters (e.g., Na + /Ca 2+ and Na + /H + exchangers; Philipson and Nicoll, 2000;Avkiran and Haworth, 2003) and proteins involved in the control of cell growth and proliferation (e.g., MAPK, Ras, and Raf;Nishikawa et al., 1997;Clerk and Sugden, 2001).
Phosphorylation assays with target proteins provide one possible way to detect PKC activity, but these approaches might be complicated by the fact that many signaling events occur only in confi ned subcellular spaces, the so-called signaling microdomains, and/or display a brief lifetime. Another approach PKCα: a versatile key for decoding the cellular calcium toolkit Gregor Reither, 1 Michael Schaefer, 2 and Peter Lipp 1 1 Institute for Molecular Cell Biology, Medical Faculty of the Saarland University, D-66421 Homburg/Saar, Germany 2 Institute for Pharmacology, Campus Benjamin Franklin, Charité-Universitätsmedizin Berlin, 14195 Berlin, Germany C onventional protein kinases C (cPKCs) play an essential role in signal transduction and are believed to integrate both global Ca 2+ transients and diacylglycerol signals. We provide evidence that PKCα is a ubiquitous readout sensor for the cellular Ca 2+ toolkit, including highly restricted elementary Ca 2+ release.
Although upon uncaging NP-EGTA, which is a caged Ca 2+ compound, WT-PKCα displayed rapid membrane translocations within <250 ms, PKCα constructs with C2 domains mutated in their Ca 2+ -binding region lacked any Ca 2+ -dependent translocation. Flash photolysis of diazo-2, a photosensitive caged Ca 2+ buffer, revealed a biphasic membrane dissociation (slow and fast period) of WT-PKCα. The slow phase was absent in cells expressing PKCα-constructs containing mutated C1a-domains with largely reduced DAG binding. Thus, two groups of PKCα membrane interactions coexist; C2-and C1a-mediated interactions with different lifetimes but rapid interconversion.
We conclude that PKCα can readout very fast and, spatially and temporally, very complex cellular Ca 2+ signals. Therefore, cPKCs are important transducers for the ubiquitous cellular Ca 2+ signaling toolkit.
is to follow translocation events in cells. Most of these experiments were performed by using long-lasting stimuli such as fatty acids (Shirai et al., 1998) or phorbolesters (e.g., phorbol 12-myristate 13-acetate [PMA]; Braz et al., 2002). These methods were typically combined with an in vitro readout at predefi ned time points during the stimulation process; thus, interpretation at the single-cell level was complicated by cellcell variability and limited temporal resolution.
In contrast, real-time live-cell imaging overcomes many of these drawbacks. For this, fl uorescent proteins of various colors are coupled to the C termini of PKCs, and translocation is followed by video or confocal imaging (Mogami et al., 2003). With such an approach, Schaefer et al. (2001) presented evidence for diffusion-limited fast translocation kinetics of PKCα in histamine receptor-overexpressing human embryonic kidney (HEK) 293 cells (Lenz et al., 2002). To follow these events with appropriate temporal resolution, these authors used confocal linescanning by sacrifi cing one spatial dimension for a gain in acquisition speed. In contrast, Codazzi et al. (2001) used total internal refl ection microscopy imaging and found fast and robust responses of the neuronal cPKCγ. In addition, they could provide some initial evidence for spatially more complex translocation events, but these studies were restricted by their limited temporal resolution of several seconds per image and their confi nement to PKC measurements at the plasma membrane level.
We have used 2D real-time confocal microscopy of fl uorescent PKCα fusion constructs in two cell types to comprehensively characterize cPKC translocation dynamics. The α-isoform was chosen because it is the most ubiquitously expressed PKC isoform and, in contrast to other ubiquitous cPKCs (e.g., PKCβ), Figure 1. PKC translocation waves. HEK293 cells were transfected with PKCα-EGFP and stimulated with ATP. (A, a) Cells expressing the PKCα construct. The colored circles mark cellular locations from which fl uorescence was averaged for the corresponding fl uorescence tracings in A (b). For those time points marked with the vertical dashed lines, individual confocal images have been analyzed in B. The labels in A (b) correspond to B (a-e). For the left column in B, we calculated the relative fl uorescence and color coded the ratio according to the color wedge in B (e). 3D surface plots of the relative fl uorescence have been constructed for the middle and right columns, which represent two different viewing angles onto the same surface. Upward defl ections indicate relative increases and downward defl ections indicate relative decreases in the local PKCα-EGFP concentration. Similar results were obtained in 15 additional cells from 10 independent experiments.
there is only a single variant (Kofl er et al., 2002). We particularly concentrated on the question of whether and to what degree cPKCα can decode the spatially and temporally very complex Ca 2+ transients that occur during physiological Ca 2+ signaling in living cells (Berridge et al., 2003) into translocation signals and on PKCα membrane interactions via their C2 and C1 domains.

Results
Physiologically, PKCα is stimulated by diacylglycerol (DAG) through binding to its C1a domain (Medkova and Cho, 1999), which is produced, e.g., as a product of PLCβ-mediated hydrolysis of phosphatidyl-4,5-bisphosphate (PIP 2 ), and by Ca 2+ through its C2 domains. To activate these signaling networks, we used ATP stimulation of HEK293 and COS1 cells because they endogenously express P2Y receptors that couple to G q proteins. In the vast majority of cells stimulated with robust agonist concentrations, we found global oscillations of both Ca 2+ and PKCα-EYFP (Fig. S1, available at http://www.jcb.org/cgi/ content/full/jcb.200604033/DC1), and variations of the agonist concentration were readily translated into graded Ca 2+ signals and PKCα-EGFP translocations ( Fig. S1 and Video 1).
Stimulation of cells with lower concentrations of Ca 2+mobilizing agonists often leads to Ca 2+ transients with complex spatiotemporal properties, such as waves or spatially restricted Ca 2+ responses (for review see Berridge et al., 2000). Thus, we questioned to what degree PKCα is able to decode the complexity of Ca 2+ signals into translocation events. Fig. 1 depicts an analysis of the translocation process evoked by a Ca 2+ wave using rapid 2D confocal imaging. Regularly, linescans or slow 2D image acquisition can produce data that are diffi cult to interpret (Bootman et al., 1997a). The Ca 2+ wave underlying the response shown in Fig. 1 brought about a wave-like translocation of PKCα-EGFP that can be seen in the pseudocolored self-ratio images ( Fig. 1 B, left column). To selectively depict the spatial properties of such translocation waves on the whole-cell level, we have plotted the relative fl uorescence changes as increases or decreases in the height of 3D surface for the entire 2D plane ( Fig. 1 B, middle and right columns; and Video 2, available at http://www.jcb.org/cgi/content/full/jcb.200604033/DC1). From such data, we concluded that PKCα is able to reliably decode even fast Ca 2+ waves propagating through cells into translocation waves.
Low agonist concentrations provoke spatially complex Ca 2+ signals (Berridge et al., 2000). The investigation of subcellular Ca 2+ signals requires Ca 2+ indicators with a good signal-to-noise ratio, especially for small and spatially confi ned Ca 2+ signals (Thomas et al., 2000a). In PKCα-DsRed2-expressing HEK293 cells, we analyzed responses to the application of different lower ATP concentrations to provoke such Ca 2+ signals. Fig. 2 depicts the result of such a representative experiment. Four regions of interest (ROIs; Fig. 2 A) have been analyzed, and the time courses of the normalized fl uo-4 (green), plasma membrane (blue), and cytosolic (red) fl uorescence of the DsRed2 constructs have been replotted in Fig. 2 B. Application of 5 μM ATP caused a Ca 2+ signal that was largely restricted to the lower extension of the cell ( Fig. 2 B, compare traces 1 and 4). The subcellular distribution of this Ca 2+ transient can be seen in the self-ratio image displayed in Fig. 2 C (second column). Despite the fact that this Ca 2+ signal was brief and spatially restricted, the PKCα translocation refl ected that spatiotemporal property (compare traces in Fig. 2 B with the corresponding images in C). A detailed analysis of the fl uo-4 fl uorescence and the plasma membrane PKCα-DsRed2 fl uorescence revealed a small difference between the time courses in that the fl uo-4 fl uorescence increase preceded the increase in the plasma membrane PKCα-DsRed2 signal by ‫008ف‬ ms (i.e., four image pairs at a 5-Hz acquisition rate; in all nine experiments with the same stimulation regime the differences were always between three and four images at 5 Hz). In addition, we observed multiple  Fitting this dataset with a single Gauss distribution resulted in a signifi cantly higher χ 2 than fi tting with two Gauss distribution centered at ‫2.2ف‬ and ‫2.3ف‬ μm. For B, we measured the signal duration at 50% of their maximal amplitude and constructed the distribution by binning the data into 0.5-s time bins. Because some of these LTEs displayed very long durations, well exceeding 4 s, we combined them into one bin (>4 s). (C) The amplitudes of the LTEs were calculated as relative fl uorescence changes after background subtraction normalized to the average of at least fi ve resting images. In D, we plotted the relative amplitude of each LTE against its duration at 50% maximal amplitude. The two dashed lines depict the boundary conditions for LTEs. small local Ca 2+ fl uctuations after the initial spike ( Fig. 2 B, traces 1 and 2, arrows) that were not refl ected in translocation events. The application of 25 μM ATP caused a Ca 2+ wave covering the entire cell (Fig. 2, B and C, third and fourth columns). From these data we conclude that PKCα translocation is able to follow Ca 2+ signals with small amplitudes, brief lifetimes, and spatial restrictions.
During experimental series in which we stimulated PKCα-EYFP-expressing COS1 cells with various lower ATP concentrations (5-25 μM), we found that ‫%51ف‬ of all COS1 cells gave spatially and temporally very complex responses, such as those exemplifi ed in Fig. 3. Fig. 3 B shows relative fl uorescence traces from six ROI pairs outlined in Fig. 3 A. These tracings illustrate very short-lived and highly confi ned increases in the PKCα-EYFP fl uorescence ( Fig. 3 B, traces 1-5). The individual "hot spots" were spatially independent of one another, but their activity increased, with a variable delay, after ATP application and ceased after ATP washout (unpublished data). As illustrated in the inset of Fig. 3 A (b), the confocal slice was positioned close to the bottom plasma membrane level of this COS1 cell; thus, we recorded PKCα-EYFP fl uorescence close to or at the lower plasma membrane. Therefore, the transient increases in the PKCα-EYFP fl uorescence were caused by spatially restricted translocation signals to the plasma membrane (local translocation events [LTEs]). LTEs were not restricted to the bottom plasma membrane; similar signals could also be observed at the upper plasma membrane (unpublished data). The detection of LTEs failed either when we increased the ATP concentration or when we moved the confocal section more toward the central region of the cell (unpublished data). Such a loss was not caused by the temporal resolution of our confocal microscope because with charge-coupled device camera-based fl uorescence acquisition we do not miss individual events; instead, they might only be blurred over time. Once a cell like the one displayed in Fig. 3 was found repetitive, stimulations with the same ATP concentration after 10-15-min resting periods evoked similar LTE patterns, although not all LTE locations could be restimulated and new ones were additionally triggered during the second ATP stimulation (Fig. 3 C).
In the following, we analyzed a larger population of LTE sites and signals (n = 160 from 60 locations; 18 cells). The results of this analysis are shown in Fig  Here, time runs into the paper plane, the preserved spatial dimension is represented in the horizontal direction, and the relative fl uorescence has been coded in both the color and the height of the 3D surface. The green and blue arrows identify the two LTEs analyzed in A, a. For B, we analyzed two LTE sites exhibiting short-lived LTE signals. The arrangement of B is similar to the one in A. For B c, a linescan image was constructed from lines acquired across the two LTE sites marked with the dashed line in B b 2. Here, time runs from left to right, and the preserved spatial dimension goes into the paper plane.
To investigate the relationship between these parameters, we plotted the amplitude of the LTEs against their individual lifetime ( Fig. 4 D). From this graph the following major fi ndings were identifi ed: (a) long and small events were absent, (b) the amplitudes of LTEs did not exceed 150% above resting values, and (c) the two LTE populations found in the individual distributions were also found in Fig. 4 D.
To visualize the spatiotemporal properties of these two populations of translocation signals, we plotted typical LTEs with long lifetimes (>6 s; Fig. 5 A and Video 3, available at http://www.jcb.org/cgi/content/full/jcb.200604033/DC1) and shorter durations (Fig. 5 B). Fig. 5 A exemplifi es two LTEs in close spatial proximity (Fig. 5 A, b). Despite continuous ATP stimulation, LTEs can display rather complex time courses (Fig. 5 A). For four selected time points, the PKCα-EYFP fl uorescence distributions were plotted as 3D surface plots in Fig. 5 A (b). The 3D surface representation of a pseudolinescan (Fig. 5 A, c) revealed that the spatial spread appears to be functionally limited, as depicted for the long-lasting LTE (black arrow). After an initial 3-s growth period, the spatial spread came to a standstill, despite ongoing translocation activity. This fi nding was typical for all long-lasting LTEs we investigated. From these fi ndings, and from those in Fig. 4 (i.e., two populations of LTEs), we conclude that long-lasting LTEs are characterized by limited amplitudes and spatial spreads.
The C2 domain of cPKC is supposed to mediate the Ca 2+ -dependent portion of the cPKC membrane interaction (Medkova and Cho, 1998;Verdaguer et al., 1999), whereas the C1a and C1b domains are thought to be responsible for DAG and PMA binding, respectively (Medkova and Cho, 1999;Ananthanarayanan et al., 2003). To investigate the contributions of the C2 or the C1a domains to the translocation of cPKCs and their membrane interactions, we have constructed two PKCα-EGFP fusion proteins lacking a functional C2 and a C1a domain. Fig. 6 depicts results obtained with the wild-type (WT) PKCα-mRFP and the C2-mutated (C2-mut) PKCα-EGFP fusion protein. Although the distribution of the two constructs in resting cells was indistinguishable (Fig. 6 A, a), the ATP stimulation revealed protein translocation only for the WT construct ( Fig. 6 A, b; PMA provoked translocation of both proteins). Thus, the mutation we have introduced into the C2 domain specifi cally suppressed the ATP-induced, but not the PMA-induced, translocation. Because ATP application leads to the generation of two second messengers that interact with the cPKC, i.e., Ca 2+ (C2 domain) and DAG (C1a domain), we designed experiments that allowed us to distinguish between these two remaining possible membrane interactions. For Fig. 6 B, HEK293 cells coexpressing the two fusion protein constructs were loaded with NP-EGTA-AM, which is a membrane-permeable caged Ca 2+ . Upon a bright UV fl ash, NP-EGTA rapidly releases Ca 2+ (DelPrincipe et al., 1999) and, thus, specifi cally generates Ca 2+ signals while avoiding the production of DAG. Two UV fl ashes in succession triggered the translocation of the WT protein to the plasma membrane ( Fig. 6 B, c, red traces; compare top images in Fig. 6 B, b 1 and b 2), whereas the C2-mutated cPKC did not display any change in its subcellular distribution (Fig. 6 B, c, green traces; compare also the bottom images in Fig. 6 B,  b 1 and b 2). The ATP application at the end confi rmed the translocation properties already described for Fig. 6 A.
For the aforementioned LTEs with lifetimes of <1 s, the translocation speed of cPKC fl uorescence proteins has to be very fast. In Fig. 7, we addressed the question of how fast the translocation of the PKCα fusion proteins is in response to a stepwise increase in the intracellular Ca 2+ concentration. Cells were coloaded with NP-EGTA and fl uo-4, and a series of four consecutive UV fl ashes was delivered. The fi rst two fl ashes induced only small step changes in the Ca 2+ concentration (Fig.  7 A, a and b), causing no apparent or only a minute (second fl ash) translocation of the PKCα construct. Flashes 3 and 4 triggered substantial increases in the intracellular Ca 2+ concentration that were accompanied by rapid translocations toward the plasma membrane (Fig. 7 B, a and b). To characterize the speed of translocation, we analyzed consecutive images acquired with a frequency of 4 frame pairs/s. Fig. 7 C (2-5) depicts individual images of the fl uo-4 ratio (left) and the relative differential changes in the PKCα-mRFP fl uorescence (right) before ( Fig. 7 C, 2) and directly after the fourth UV fl ash (Fig.  7 C, 3-5; note that both the fl uo-4 and mRPF images were normalized to pre-fourth UV fl ash fl uorescence intensity). Within <250 ms, the cPKC fusion proteins accumulated at the plasma membrane level (see the green outline of the cell in the Fig. 7  C, 3, right). The Ca 2+ concentration had increased in a stepwise manner and remained constant from Fig. 7 C, 3-5 (left column). From this, we concluded that (a) cPKC translocation required a certain threshold Ca 2+ concentration and (b) that cPKC translocation to the plasma membrane can occur within <250 ms. Thus, cPKC translocation to the plasma membranes appeared fast enough for the LTEs recorded, but is the dissociation of the cPKC-membrane interaction also fast enough to cause the fast relaxation seen for the LTEs? What contribution do the C illustrates exemplifi ed images of the fl uo-4 (left) and mRFP (right) fl uorescence. The row labeled with 1 depicts the resting fl uorescence and the two lines used to generate the pseudolinescan images in A and B (dashed lines). Images in row 2 were taken immediately before the fourth UV fl ash, and the images in rows 3-5 were obtained at the time points indicated. The fl uo-4 and the mRPF images were ratioed against the fl uorescence just before the fourth UV fl ash. The color coding for the mRFP self-ratio images and the mRFP-derived pseudo linescan images in A is shown at the bottom of C. Similar results were obtained in nine additional cells recorded under similar experimental conditions. C2 and C1a domains make to the membrane residence time or dissociation kinetics?
To address these questions, we expressed WT PKCα-EGFP and coloaded the cells with Fura red and diazo-2, which is a caged Ca 2+ buffer (Lipp et al., 1996). For these experiments, we restricted the UV illumination during the UV fl ash to only part of the fi eld of view to simultaneously record cellular responses with and without the UV fl ash. For the cell depicted in Fig. 8 A (b), we applied a UV fl ash ‫01ف‬ s into the ATP stimulation. This caused a rapid release of a highaffi nity Ca 2+ buffer (K d = 73 nM) and, consequently, the Ca 2+ concentration quickly dropped to resting values (Fig.  8 A, b, red trace and Fig. 8 A, d). After this, the cytosolic PKCα-EGFP fl uorescence increased (Fig. 8 A, b, black trace), whereas the fl uorescence at the plasma membrane level decreased (Fig. 8 A, b, green trace). In Fig. 8 A (b), the black and orange arrows point to a fast and slow component of this decline, respectively. The transition between these two phases was present in all cells analyzed with the same experimental regime (15 cells in fi ve independent experiments), but the fractional contribution of each phase to the total PKCα dissociation process varied between 30 and 60%. From these experiments and from the two populations of LETs we concluded that, indeed, these two time constants of membrane dissociation represent two PKCα pools at the plasma membrane: (a) Ca 2+ -C2-bound PKCα causing the fast component (Fig. 8  A, b, black arrow) and (b) binding of PKCα via its Ca 2+ -C2 and DAG-C1 domains causing the second, slower component ( Fig. 8 A, b, orange arrow). C1 domain-mediated membrane interactions are much more stable and have a longer lifetime (according to in vitro data; Bittova et al., 2001). To test this hypothesis, we characterized the effect of a functional C1a mutation on the dissociation kinetics of PKCα from the plasma membrane (Fig. 8 B).
For this, HEK293 cells expressing C1a-mutated PKCα-EGFP were loaded with Fura red and diazo-2. The basic experimental design was the same as described for Fig. 8 A. Stimulation of the cells with 100 μM ATP resulted in a robust and reversible translocation to the plasma membrane without apparent differences to the WT construct (compare Fig. 8 A, c, and B, c). Illumination of the cells with a UV fl ash caused a rapid drop in the intracellular Ca 2+ concentration (Fig. 8 B, b, red trace) that induced an increase in the cytosolic PKCα(C1a-mut)-EGFP fl uorescence back to prestimulation levels ( Fig. 8 B, b, black trace). When we analyzed the time course of membrane fl uorescence, we found that the fl uorescence decay after the UV illumination was only monophasic (Fig. 8 B, b, black arrow), the kinetics of which was similar to the kinetics of the fast component of the WT PKCα (compare green traces in Fig.  8 A, b, and B, b). This lack of a slower decay phase constant was a consistent fi nding in all cells expressing the mutated cPKC (n = 7 cells in four independent experiments). From this we conclude that the slower dissociation time constant seen in the cells expressing the WT PKCα-EGFP was most likely caused by C1a-mediated membrane interactions, whereas the faster decay was dependent on a functional C2 domain in the protein (Fig. 8 B, d).

Discussion
Signaling cascades using the stimulation of PLCβ result in the production of the InsP 3 and DAG that activate InsP 3 receptors and PKCs, respectively, with Ca 2+ release from the ER triggered by InsP 3 . A tight coupling between intracellular Ca 2+ signals and PKCα translocation was proposed recently (Mogami et al., 2003). By using the physiological agonist ATP and fl ash photolysis of caged compounds, we were not only able to provide strong supporting evidence for such an idea but also revealed various pools of PKCα at the plasma membrane. Our fi ndings suggest fast and tight mechanisms for decoding incoming stimuli into translocation signals. Nevertheless, the molecular mass of conventional PKCs (PKCα, ‫28ف‬ kD) gives rise to principle doubts as to whether such a protein will be able to follow very fast and brief Ca 2+ signals (lifetime, <1 s) observed during cellular Ca 2+ signaling. Although the fl uorescence tags of the PKC constructs added another 27 kD to the molecule's molecular mass, the Stokes radius of the fl uorescent PKCα only increased slightly (from 42 to 46 Å; Schaefer et al., 2001).
We showed that fl uorescent PKCα fusion proteins display graded translocations, which are similar to those reported for glucose stimulation and PKCβ (Pinton et al., 2002). Further establishment of such a notion appears important in the context of PKC-mediated signal transduction because interpretation of amplitudes adds a new dimension to the parameter space that cPKCs can exploit and cells can use to decode incoming stimuli. A similar ability in decoding Ca 2+ signals has been suggested for calmodulin (Frey et al., 2000).
Cells do not only use the frequency and amplitude domains to decode incoming stimuli into cellular responses, but spatiotemporally complex Ca 2+ signals have also been described, ranging from global Ca 2+ waves to spatially restricted Ca 2+ waves and elementary Ca 2+ signals, such as Ca 2+ sparks and Ca 2+ puffs (for review see Berridge et al., 2000). The importance of such complex signaling processes for cellular signal transduction has been suggested only in an indirect way, by linking local Ca 2+ signaling to cellular responses or by electrophysiological measurements that allow direct readouts of, e.g., Ca 2+ -dependent membrane ion conductance in smooth muscle cells (Nelson et al., 1995;Bolton et al., 1999). Nevertheless, it appears important to present direct evidence and visualization of spatiotemporally very complex processes linking local Ca 2+ signaling to other signal transduction mechanisms. We provide such evidence in that the Ca 2+ signaling toolbox, from waves to spatially restricted Ca 2+ signals and elementary Ca 2+ release events, can indeed be decoded into PKCα translocations to the plasma membrane. Assuming similar properties for all cPKCs, it appears feasible to extend such behavior to the entire cPKC subfamily. An important requirement for such mechanisms is a rapid translocation, despite the substantial molecular mass of our constructs ‫911ف(‬ kD). With real-time imaging of PKCα translocations after UV fl ash-induced Ca 2+ increases, we demonstrated suffi ciently fast translocations kinetics (Fig. 7).
It should be noted that not every elementary Ca 2+ signal will necessarily result in local cPKC translocation because the k on and k off rates of Ca 2+ binding are very fast, and translocation also requires a certain Ca 2+ threshold concentration. Ca 2+ release deeper in the cell will lead to cPKC Ca 2+ binding, but unbinding of the ion will occur before the cPKC can reach the membrane and encounter PSs and DAG. Thus, only peripheral Ca 2+ signals or, in other words, only the peripheral component of Ca 2+ signals, will lead to a signifi cant cPKC translocation. Thus, this is another example of the versatility of Ca 2+ signals solely depending on their particular location inside cells. Although peripheral transients can induce, e.g., phosphorylation of membrane proteins by the PKC-signaling networks, perinuclear Ca 2+ release events have been suggested to result in specifi c nuclear Ca 2+ signals and trigger accompanying Ca 2+ -dependent transcription events (Lipp et al., 1997).
Although translocation of the cPKC molecule to its target, the plasma membrane, is an important step in the activation mechanism, the question still remains whether translocation itself can, in fact, indicate activation of cPKCs. This might be rather likely for long-lasting Ca 2+ transients, given the very high DAG affi nity of Ca 2+ -bound cPKC. In this study, we could provide strong direct evidence for a long-lasting membrane interaction of C1a and DAG in living cells; functionally mutating the C1a domain of PKCα rendered the dissociation process very fast (Fig. 8 B, d); thus, C1a-mediated membrane binding of PKCα represents a signifi cant pool of long-lasting PKCα membrane interactions out of which PKCα could have sufficient time to encounter and interact with target proteins and perform phosphorylation.
In contrast, this is not necessarily obvious for short-lived LTEs. If binding of the C1 and C2 domains to the plasma membrane occurs independently (Johnson et al., 2000), and the initial membrane affi nity is mainly determined by its membrane C2 interactions, ionic interactions between the Ca 2+ -occupied C2 domain and PS in the plasma membrane are the next step (Kohout et al., 2002). The C2 membrane interaction provides the fi rst part of the energy necessary for the allosteric release of the pseudosubstrate from the kinase domain (Newton, 2001). These processes have brought the C1 domain into the proximity of the plasma membrane for C1-membrane interactions that are believed to predominantly determine the residence time of the cPKC at the membrane (Bolsover et al., 2003). After C1a-DAG interactions, the pseudosubstrate is released from the kinase domain that is now ready for substrate interactions and phosphorylation of the target proteins (Nishikawa et al., 1997). Despite the fact that this sequence of events is, in principle, agreed on quite widely, some details are still open for discussion and might have to be reinterpreted in light of our fi ndings of very short-lived LTEs and the two pools of PKCα membrane interactions.
If DAG binding of Ca 2+ -cPKC is, indeed, the principle event linking the cPKC to the membrane, then how does this notion account for the two populations of lifetimes observed in this study? We propose that these distinct lifetimes refl ect different states of the cPKC membrane interactions. As we have previously described, rises in the Ca 2+ concentration usually "precede" cPKC translocation, and very low-amplitude signals are not necessarily translated into translocation events (Figs. 3  and 7), which is indicative of a certain Ca 2+ threshold concen-tration necessary for cPKC translocation. In fact, Ca 2+ binding to the C2 domain displays a surprisingly low affi nity (K d ) of ‫53ف‬ μM (measured in vitro in the absence of phospholipids; Kohout et al., 2002). Although we have not been able to provide calibrated Ca 2+ measurements for the experiments in Fig. 7 (resting Ca 2+ will be infl uenced by NP-EGTA), it appears feasible to assume that the Ca 2+ concentrations evoked by UV fl ash photolysis of NP-EGTA were much lower than the apparent K d of 35 μM (DelPrincipe et al., 1999).
Therefore, we suggest that the LTEs with the shorter lifetimes refl ect pure Ca 2+ -dependent binding of PKCα to the plasma membrane. With an apparent diffusion coeffi cient of 4.67 × 10 −7 cm 2 /s, a cytosolic viscosity of around 2.0-3.2 cP (Mastro et al., 1984;Schaefer et al., 2001), and an average lifetime of the Ca 2+ -PKCα complex of ‫-51-21ف‬ms molecules can only interact with the plasma membrane when Ca 2+ binding occurs ≤1 μm away from the plasma membrane (for an illustration see Fig. S2). Furthermore, even for long-lasting Ca 2+ increases, slower and longer lasting membrane interactions solely driven by Ca 2+ increases have been described at the global cellular level (Lenz et al., 2002;Sakai et al., 1997). When binding of the Ca 2+ -C2 domain to the PS of the plasma membranes occurs, the lifetime of the Ca 2+ -binding complex increases fi vefold and, thus, plasma membrane interaction times are signifi cantly prolonged (calculated for PKCα-C2 domain according to in vitro data; Nalefski and Newton, 2001;Kohout et al., 2002;Bolsover et al., 2003). With a putative phosphorylation turnover rate of 380-500 min −1 measured in vitro (Sando et al., 1998;Yeh et al., 2002), such brief membrane interactions might, in fact, be suffi cient to phosphorylate target proteins in case of an encounter. Elementary Ca 2+ signals, such as Ca 2+ puffs, have average lifetimes of ‫2-1ف‬ s (Yao et al., 1995;Bootman et al., 1997b;Lipp et al., 1997;Thomas et al., 2000a); thus, the shortest lifetimes of the LTEs observed here are compatible with a purely Ca 2+ -driven process. This notion is further substantiated in that, upon a stepwise decrease, the Ca 2+ concentration PKCα(C1a-mut) displayed a rapid membrane dissociation, very closely following the Ca 2+ decline (Fig. 8 B, d, red transparent trace).
Longer lifetimes of the cPKC membrane complex might indicate additional, and possibly more specifi c, interactions. Therefore, we propose that the membrane translocations with longer lifetimes represent interactions between the Ca 2+ -PKCα and DAG. This is particularly interesting because investigation of these lifetimes could yield additional information of that important interaction. Such extended membrane localizations could also indicate additional interactions of the activated cPKCs with receptors of activated kinases (Schechtman and Mochly-Rosen, 2001). Moreover, simultaneous cPKC interactions with several partners will increase the membrane-residence periods overproportionally because redistribution into the cytosol can only occur when the cPKCs unbind from both partners almost simultaneously. Thus, the observed long-lasting translocation events might represent a temporal "walking" of the molecule between dynamic binding/unbinding cycles to the various partners in the plasma membrane. Two additional possible schemes might give rise to such long-lasting translocation events, including highly repetitive or long-lasting Ca 2+ puffs, but these can most likely be excluded. Their frequency ought to be well above 2-3 Hz because we have visualized LTEs with lifetimes of around 300-500 ms. Such high-frequency Ca 2+ puffs have not been imaged yet and are, in fact, rather unlikely taking into account adaptive processes and refractory time periods of the InsP 3 R (Berridge et al., 1999). Our experiments (Fig. 8) do not seem to favor the notion of receptor of activated kinase binding because mutation of the C1a domain alone completely suppressed that PKCα pool with the longer membrane-residence time.
On the other hand, long-lasting Ca 2+ puffs with extended lifetimes (>3-5 s) are also unlikely as the responsible mechanism. Such long-lasting Ca 2+ puffs have not been described yet, and local depletion mechanisms in the ER might actually terminate local Ca 2+ release much earlier, as has been proposed for Ca 2+ sparks (DelPrincipe et al., 1999).
Nevertheless, this cannot easily account for the limited spatial spread of such long-lasting LTEs (Fig. 6 A). For this to occur, at least one of the binding partners has to exhibit a restricted lateral diffusion in the plane of the plasma membrane ‫1ف(‬ μm 2 /s; unpublished data). The likely candidates for such a mechanism are phosphorylation target proteins that have been anchored in the plasma membrane or membrane lipid domains that were rich in DAG after receptor stimulation. In these situations, residence times of the cPKCs are primarily determined by C1a-DAG binding or target protein interactions in the plasma membrane, and the extent of membrane location will be increased largely.
In conclusion, we propose that the two LTE lifetimes described here represent visual correlates of distinct states of cPKCs during their activation/deactivation cycle. K off rates seem to be predominantly regulating the duration of cPKC membrane interactions with purely C2 domain-dependent membrane interactions being the most dynamic process. C1a domain-mediated membrane or cPKC target protein interactions are more stable, and, consequently, LTEs display a longer lifetime. These notions are supported by our mutation and UV fl ash experiments, in which we illustrated that C2-mediated membrane interactions were very fast in both directions and that association and dissociation and functional C1a domains largely increased membrane residence times, i.e., decay times were signifi cantly extended. In addition, we provided fi rst evidence that two PKCα populations coexist in the membrane (C2-and C1a-bound proteins), which most likely undergo rapid interconversion.
Our results, thus, foster the understanding of cellular signaling via PKCα (possibly also for the other cPKCs). At resting Ca 2+ concentrations, the C2 domain repels the PKCα from the membrane because of electrostatic interactions. As soon as the Ca 2+ concentration is increasing, subplasmamembrane PKCα molecules with a Ca 2+ -C2 domain bind to the PS in the plasma membrane and the PS-Ca 2+ -C2 complex is stabilized. Failing to interact with PS causes rapid dissociation from the membrane. This is an essential step that can be imaged as the association of fl uorescently tagged PKCα molecules to the membrane. We thus propose that during cellular Ca 2+ signals Ca 2+ -C2-PKCα in fact "tiptoes" on the membrane until it "sticks" to a PS. This stabilization step gives the PKCα suffi -cient time to "probe" for DAG in the membrane. Binding to the DAG (via its C1a domain) further stabilizes the membrane interactions and further increases membrane-residence times. Such a stabilized complex is now given enough time to phosphorylate target proteins.
From this we conclude that, at least for cPKCs, spatial and temporal targeting of cPKC-mediated signaling is mostly, if not solely, driven by the spatiotemporal properties of the underlying Ca 2+ -signaling machinery.
We believe that these fi ndings put cPKCs in at least the same position as the classic Ca 2+ signal decoder calmodulin. We have visualized that a downstream signaling molecule, PKCα, is able to decode cellular and subcellular Ca 2+ signals in all their complexity into translocation and, thus, is an essential linker between Ca 2+ transients and downstream signal transduction networks (e.g., RhoA, Rac1 or ERK/MAPK; Nakashima, 2002), but also directly phosphorylates membrane proteins such as ion channels and transporters. To our knowledge we illustrated for the fi rst time in living cells that PKCα rapidly converts between various states of membrane interactions and that there are at least two pools of PKCα in the membrane after agonist stimulation.

PKC mutagenesis
In line with the published results of Cho (1998, 1999) on PKCα mutants, we performed site-directed mutagenesis (Kirsch and Joly, 1998)  Cell culture and transfection HEK293 and COS1 (CRL-1573 and CRL-1650; American Type Culture Collection) cells were grown in DME (Invitrogen) supplemented with 5% FBS (PANBiotech), 50 U/ml penicillin, and 50 U/ml streptomycin (both from Invitrogen). Cells were seeded on 20-mm glass coverslips in 12-well plates and transfected the following day at 20-30% confl uency with Lipofectamine 2000 (Invitrogen) and appropriate amounts of plasmid DNA per well. All experiments were performed in both cell types and the results shown are representative of both cell types. It must be mentioned that for all the fi ndings we verifi ed in COS and HEK cell lines, we never found qualitative differences between the cell types.

Confocal imaging
Localization and translocation of the fl uorescent fusion proteins, as well as changes in the fl uorescence of Ca 2+ indicators such as fl uo-4 and Fura red (Invitrogen), were analyzed by laser scanning confocal microscopy. We either used a Nipkow disk-based QLC100 (VisiTech International) equipped with a dual-port adaptor for monitoring two emission wavelengths simultaneously (ORCA-ER cameras; Hamamatsu) or the kilobeam 2D-array scannerbased VTinfi nity (VisiTech International) equipped with an ORCA-ER camera for single emission recordings. Both confocal systems were mounted to either an inverted microscope (TE-2000U;Nikon) or an upright microscope (E600; Nikon) and controlled by VoxCellScan software (VisiTech International). Excitation of the fl uorescent fusion proteins was performed with the 488-nm line (EGFP or EYFP) or the 568-nm line (for DsRed2) of an Ar/Kr multiline laser (LaserPhysics). Emission was imaged at wavelengths >500 nm for EGFP and EYFP, and at wavelengths >580 nm for DsRed2. In experiments where we used fusion proteins and Ca 2+ indicators, we used two different setups. For the fi rst setup, EGFP or EYFP together with Fura red were only excited at 488 nm, while the emissions were imaged at 520 and 600 nm, respectively. For the second setup, fl uo-4 and DsRed2 required excitation at two wavelengths (488 and 568 nm), and the emissions were imaged at 520 and 600 nm. Individual images or image pairs were recorded at acquisition speeds ranging from 1 to 10 Hz through 40× (PlanFluor, NA 1.3, DIC H), 60× (PlanApo, NA 1.4, DIC H), or 100× oil immersion objectives (PlanApo, NA 1.4, Ph3 DM; all Nikon). All experiments were performed at room temperature (20-22°C) 2-4 d after transfection. In experiments with Ca 2+ indicators, the dyes (1 μM fl uo-4 AM or 3 μM Fura red AM) were loaded into the cells for 30 min, with an additional 20 min after wash-out for deesterifi cation.

UV fl ash photolysis of caged compounds
For rapidly increasing or decreasing the intracellular Ca 2+ concentration, we loaded the cells with 7.5 μM NP-EGTA-AM or diazo-2-AM (Invitrogen), respectively. A UV fl ash II (Till Photonics) was attached to an upright microscope (E600; Nikon) via an additional epifl uorescence input. The UV was directed into the excitation light path via a UV dichroic mirror (HQ450LP; OptiLab). We used a water immersion objective (40×, NA 0.8; Nikon). This allowed for simultaneous recording of the cellular fl uorescence and application of the UV fl ash. The timing between imaging and the fl ash was programmed within the VoxCellScan confocal software. Because both caged compounds decreased resting Ca 2+ to an unknown degree, quantitative measurements of Ca 2+ were not possible. In addition, the amount of Ca 2+ photoreleased from NP-EGTA largely depends on the amount of (a) photolysed compound and (b) Ca 2+ loading of NP-EGTA. The latter effect explains the different Ca 2+ steps in Fig. 8 caused by UV fl ashes of identical amplitude. Before application of the fi rst fl ash, the resting Ca 2+ was very low (K d for uncaged NP-EGTA 80 nM; upon UV illumination K d is increased to 1 mM); thus, the amount of photoreleased Ca 2+ was rather low, but growing with increasing "resting" Ca 2+ before the subsequent fl ashes (higher occupancy of NP-EGTA by Ca 2+ ). With AM loading, such effects are largely unavoidable because we could not "preload" the caged compound as is possible when perfusing the cell in the whole-cell confi guration of the patch clamp technique used elsewhere (DelPrincipe et al., 1999). Diazo-2 is a caged Ca 2+ buffer (caged BAPTA) and its K d is increased upon UV illumination from 2.2 to 73 nM.

Solutions
All experiments were performed in a Hepes (Merck)-buffered salt solution (extracellular medium) composed of (in mM): 135 NaCl, 5.4 KCl, 2 MgCl 2 , 10 glucose, 2 CaCl 2 , and 10 Hepes adjusted to pH 7.35 with NaOH. ATP (Sigma-Aldrich) and PMA (Calbiochem) were dissolved as stock solutions in the appropriate solvents and diluted in the external salt solution at the given concentrations before each experiment. Rapid solution switch (exchange time <1 s) was achieved by means of a solenoid-driven custom-made perfusion system. With this system a laminar fl ow of ‫002ف‬ μm in diameter was applied to the entire fi eld of view; thus, concentration changes of agonists display global changes occurring simultaneously over the fi eld of view.

Data analysis
After storage of the image, we analyzed the data using ImageJ (W. Rasband, National Institutes of Health, Bethesda, MD). ROI fl uorescence over time data were transferred into IGOR software (Wavemerics) and further processed for data display. To account for inhomogeneous dye distribution, we calculated self-ratio images as indicated. For this, we divided subsequent images of a video by a mean image obtained after averaging the 5-10 resting images of each movie (Bootman et al., 1997a,b). Colorcoded 2D images were constructed in ImageJ as 8-bit grayscale images to which the given color look-up table was applied. When given in the fi gures, Ca 2+ concentrations were calculated as described earlier, assuming a resting concentration of 100 nM (Thomas et al., 2000b). 3D surface representations of individual images or entire movies were calculated with Imaris4 software (Bitplane). All results of our live-cell imaging were analyzed in cells displaying a wide range of expression levels. For the quantitative analysis of LTEs presented in Fig. 4, we had identifi ed criteria that local fl uorescence changes had to meet to be treated as LTEs. We identifi ed fl uorescence changes as LTEs when their spatial spread was <6 μm, their duration was at least 2 images (or 250 ms) above twofold the standard deviation of the resting signal, and their amplitude was at least two times the standard deviation of the resting fl uorescence noise (horizontal dashed line in Fig. 4 D) for at least 2 images (vertical dashed line in Fig. 4 D). Fig. S1 exemplifi es HEK and COS cells expressing PKCα fusion proteins showing Ca 2+ and PKCα oscillations and graded Ca 2+ and PKCα responses. Fig. S2 shows a model of the mechanisms possibly leading to the generation of LTEs. Video 1 shows the tight relationship between Ca 2+ oscillations (Fura-Red) and PKCα-EYFP translocation. Video 2 displays the spatiotemporal properties of a PKCα translocation wave propagating through HEK cells stimulated with ATP. Video 3 illustrates the spatiotemporal properties of a long-lasting LTE in COS1 cells upon stimulation with threshold concentrations of ATP. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200604033/DC1.