LAMP, a new imaging assay of gap junctional communication unveils that Ca2+ influx inhibits cell coupling


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Article

Nature Methods 2, 55 - 62 (2005)
Published online: 21 December 2004; | doi:10.1038/nmeth730

LAMP, a new imaging assay of gap junctional communication unveils that Ca²⁺ influx inhibits cell coupling

Kenneth Dakin, YuRui Zhao & Wen-Hong Li

Departments of Cell Biology and of Biochemistry, University of Texas Southwestern Medical Center at Dallas, 5323 Harry Hines Blvd., Dallas, TX 75390, USA.

Correspondence should be addressed to Wen-Hong Li wen-hong.li@utsouthwestern.edu

Using a new class of photo-activatible fluorophores, we have developed a new imaging technique for measuring molecular transfer rates across gap junction connexin channels in intact living cells. This technique, named LAMP, involves local activation of a molecular fluorescent probe, NPE-HCCC2/AM, to optically label a cell. Subsequent dye transfer through gap junctions from labeled to unlabeled cells was quantified by fluorescence microscopy. Additional uncagings after prior dye transfers reached equilibrium enabled multiple measurements of dye transfer rates in the same coupled cell pair. Measurements in the same cell pair minimized variation due to differences in cell volume and number of gap junctions, allowing us to track acute changes in gap junction permeability. We applied the technique to study the regulation of gap junction coupling by intracellular Ca²⁺ ([Ca²⁺]_i). Although agonist or ionomycin exposure can raise bulk [Ca²⁺]_i to levels higher than those caused by capacitative Ca²⁺ influx, the LAMP assay revealed that only Ca²⁺ influx through the plasma membrane store-operated Ca²⁺ channels strongly reduced gap junction coupling. The noninvasive and quantitative nature of this imaging technique should facilitate future investigations of the dynamic regulation of gap junction communication.

Intercellular communication through gap junctions formed by protein subunits called connexins plays an important role in maintaining cell homeostasis and synchronizing physiological functions of cells^1,^2,³. Understanding molecular mechanisms regulating cell junctional communication remains an outstanding biological challenge^4,⁵. The investigation of the gating of connexin channels by cellular biochemical changes has been hampered in part by the lack of a non-invasive and quantitative technique to assay cell coupling. Existing methods for measuring rates of junctional molecular transfer include microinjecting tracers such as Lucifer yellow or neurobiotin that cannot permeate the membrane, or following fluorescence recovery after photobleaching (FRAP; ref. 6). As microinjection disrupts intact cell membranes, this and other invasive methods such as scrape loading or electroporation may introduce uncertainties in quantifying dye transfer rates due to variations in cell recovery after dye loading. In gap junction-FRAP, cell permeable dyes such as carboxyfluorescein diacetate have been used. This offers the advantage of maintaining the integrity of cell membranes. However, as very strong laser light is required to rapidly photobleach fluorophores in a whole cell, the possible photo-damage of such intense laser illumination on cells needs to be assessed. In addition, the FRAP technique may be incompatible with multi-color imaging when other biochemical changes inside cells need to be monitored by fluorescent sensors. Another method, dual whole-cell patch clamping, is a sensitive method of high temporal resolution for monitoring junctional conductance⁷. The technique has been used to study ionic permeability and selectivity of connexin channels. However, intercellular current measurements do not necessarily correlate with the kinetics of molecular transfer through gap junctions^5,⁸, and quantification of junctional conductance can be especially problematic in well-coupled cells where contributions of series resistance can outweigh junctional resistance⁹.

We aimed to develop an imaging technique to assay molecular transfer rates across cellular gap junctions. Ideally, the technique should be non-invasive to cells, able to provide dynamic and quantitative information of dye transfer and allow multiple measurements of dye transfer in the same coupled cell pair to track changes of molecular permeability of connexin channels. To this end, we used a new class of photo-activatible fluorophores that were recently developed in our laboratory¹⁰. The assay involves loading cells with a caged and cell permeable fluorophore, NPE-HCCC2/AM. After loading, we locally uncaged one cell of a coupled cell pair to create an asymmetric fluorescent marking. Subsequent fluorescence imaging provides dynamic information of dye transfer from the donor cell to the recipient cell. Since NPE-HCCC2/AM can be loaded into cells to fairly high concentrations, we can carry out multiple measurements of dye transfer rates in the same coupled cell pair with additional uncagings after the initial dye transfer reaches equilibrium. Applying this new imaging assay to cultured primary human fibroblasts, we discovered that capacitative calcium influx across plasma membranes strongly inhibits cell coupling through gap junctions.

Results
LAMP assay development to image cell-cell communication
The fluorescent and photochemical properties of the caged dye, NPE-HCCC2/AM, are summarized in Fig. 1a. NPE-HCCC2/AM is a neutral and lipophilic molecule that can diffuse across cell membranes. Once inside cells, intracellular esterases hydrolyze the acetoxymethyl (AM) esters to generate NPE-HCCC2, which is trapped inside cells. The parent fluorophore of this coumarin cage is 7-hydroxy-6-chloro-coumarin 3-carboxamide (HCCC2). HCCC2 is highly fluorescent in aqueous solutions with an extinction coefficient () of 44,000 (408 nm) and a fluorescence quantum yield (Q_f) of 0.93. Masking the 7-hydroxy of HCCC2 with a 1-(2-nitrophenyl)ethyl (NPE) group quenches its fluorescence to a negligible level (Q_f = 0.0025). UV uncaging dose-dependently regenerated fluorescent HCCC2 (Fig. 1b−d). NPE-HCCC2 is remarkably sensitive to UV photolysis. Its uncaging cross section, which is a product of the uncaging quantum yield (0.33) and (20,000 M^-1 cm^-1), exceeds 6,000 at 365 nm, nearly two orders of magnitude higher than previous caged fluorophores¹⁰. This exceptional uncaging efficiency is desirable for in vivo imaging applications because we can efficiently photolyze the NPE cage while minimizing side effects of UV illumination on live specimens. The byproduct of photolyzing the NPE caging group is 2-nitrosoacetophenone, a relatively inert molecule in aqueous solutions. The molecule is small and lipophilic, so it can diffuse across cell membranes, further reducing its effective reactivity inside cells.

Figure 1. A new imaging assay of gap junctional communication based on a new generation of caged fluorophores.

	(a) Structures and fluorescent properties of a cell permeable and caged coumarin, NPE-HCCC2/AM, its intracellular hydrolysis product, NPE-HCCC2, and its parent fluorophore, HCCC2, after photo-uncaging. (b−d) Fluorescent images of primary human fibroblasts prior to UV (360 20 nm) uncaging (b), after 0.2 sec (c) and after 0.6 sec (d) of global UV exposures of all cells in the field of view. Human fibroblasts were loaded with 1 M NPE-HCCC2/AM for 1 h in HBS with 10 mM Hepes and 5.5 mM glucose, pH 7.35. After loading and washing, cells were imaged on an inverted fluorescence microscope (425 5 nm excitation, 460 10 nm emission). (e) Schematic of the LAMP technique for studying gap junction coupling. Black open circles and blue dots represent NPE-HCCC2 and HCCC2, respectively. Full Figure and legend (32K)

Because the molecular weight of HCCC2 (MW = 450) is well below the molecular passage limit ( $approx$ 1,000) of connexin channels, we expected that HCCC2 would transfer between cells through gap junctions. The new imaging assay of gap junction communication, LAMP, involves local uncaging of NPE-HCCC2 to generate fluorescent HCCC2 in one cell and monitoring of dye transfer to neighboring coupled cells by fluorescence microscopy (Fig. 1e).

We chose primary human fibroblasts isolated from foreskin as the biological system for initial studies. These cells express gap junction proteins and form gap junctions in culture (Supplementary Fig. 1 online). To achieve localized uncaging, we placed a field diaphram in the excitation light path to control the beam size of excitation light. During localized uncaging, we closed the iris of the diaphragm to a minimum to reduce the size of the UV beam to a fraction of a cell. This locally photolyzed caged HCCC2 and asymmetrically marked one cell of a coupled cell pair. We then rapidly opened the iris of the field diaphragm to its maximum and continued to image cells by digital fluorescence microscopy. Selected fluorescence images of cells at different time points illustrating asymmetric marking of the donor cell (cell 1) and subsequent HCCC2 transfer to the recipient cell (cell 2) are shown (Fig. 2a−d). We also plotted fluorescence intensities of HCCC2 from the coupled cell pair over the course of a dye transfer experiment (Fig. 2e). Three UV flashes were locally delivered to cell 1. UV flashes of increasing durations (0.2, 0.7, and 1.2 s) were needed to generate about the same increase of fluorescence in the donor cell because previous uncagings lowered the concentration of NPE-HCCC2 in the cell. Prior to the third uncaging, 18 alpha

-glycyrrhetinic acid ( alpha

-GA), a blocker of gap junction transmission¹¹, was added to cells. Subsequent local uncaging raised coumarin fluorescence of cell 1. However, dye transfer from cell 1 to cell 2 was blocked.

Figure 2. A non-invasive and quantitative imaging assay of gap junctional communication.

	(a−d) Fluorescence images of a pair of coupled primary human fibroblasts. Cells were loaded and imaged as in Figure 1. Whole-field view of cells (both cells were briefly uncaged by a global UV illumination so they could be identified by fluorescence) just prior to a local uncaging, (a); imaging during localized uncaging of cell 1 with the iris of the field diaphragm minimized, (b); whole-field view of cells immediately after the localized uncaging of cell 1, (c); transfer of HCCC2 from cell 1 (donor) to cell 2 (recipient) reached equilibrium about 400 s later, (d). (e) Time course of fluorescence intensities of HCCC2 (F_HCCC2, arbitrary units) in coupled human fibroblasts. Digital fluorescence images were taken every 15 s. -Glycyrrhetinic acid (-GA) was added prior to the third uncaging. Rates of HCCC2 diffusion are indicated under each episode of transfer. (f,g) Quantification of HCCC2 transfer rates using data from e. The slopes of the fitted lines (linear least-square fits) gave rates of dye transfer after first (k₁) and second (k₂) uncagings (r² values of linear fittings in parentheses). Full Figure and legend (22K)

To quantitatively analyze rates of intercellular coumarin transfer, we applied Fick's equation which describes the kinetics of dye passage in a system of two compartments separated by a membrane:

where C_e, C₀, and C_t are dye concentrations in the recipient cell (or donor cell) at equilibrium, zero time and time t, respectively. The rate constant k (in units of sec^-1) describes the kinetics of dye transfer between coupled cell pairs and is sufficient to assay gap junction permeability (P_j) in the same coupled cell pairs if the cell volume and gap junction surface area remain relatively constant in the course of an experiment⁵. A similar approach has been used to analyze FRAP data of gap junctions and nuclear pore complexes^6,¹² and to estimate permeability between cells following injection of dyes¹³. Assuming that the fluorescence signal measured under constant experimental conditions is proportional to dye concentrations, the above equation can be written as:

Where F_e, F₀, and F_t are cellular HCCC2 fluorescence intensity at equilibrium, zero time and time t, obtained from the measured cellular fluorescence after background subtraction. To apply this equation to analyze dye transfer rates across gap junctions, it would require that dyes diffuse much more rapidly in the cytosol than across connexin channels, so the rate-limiting step for intercellular dye passage is gap junction permeation. To confirm that HCCC2 diffuses rapidly inside cells, we locally uncaged NPE-HCCC2 at one end of a fibroblast (Supplementary Fig. 2 online). Within 10 s, the distribution of HCCC2 fluorescence in the cell reached equilibrium. Thus, by comparison with cell-cell transfer of HCCC2 through gap junctions which takes at least several minutes to equilibrate, intracellular diffusion of HCCC2 is not rate limiting.

Quantitative analysis of dye transfer using equation (3) showed that rates of HCCC2 transfer changed little after the first and second uncagings (Fig. 2f,g). More extensive tests confirmed that rates of dye transfer of any given coupled cell pair remained constant over a period of at least several hours when cells were bathed in balanced saline, suggesting stable junctional communication in these primary human fibroblasts. However, the rate of dye transfer varied over a wide range from cell pair to cell pair, possibly reflecting cellular heterogeneities in connexin expression, the number of junctional channels assembled between cells and in cell volumes. Comparing dye transfer rates in single coupled cell pairs by multiple uncagings minimizes these variations, so that changes in dye transfer rates reflect changes in the molecular permeability of connexin channels.

Dual-color imaging of [Ca²⁺]_i regulation of cell coupling
As the first application of this new imaging assay of junctional coupling, we investigated how intracellular Ca²⁺ fluctuations regulate cell-cell communication in intact cells. The elevation of cytoplasmic Ca²⁺ concentration ([Ca²⁺]_i) has long been postulated to be involved in the gating of connexin channels¹⁴. A broad range of [Ca²⁺]_i elevations, usually controlled by dialyzing or perfusing ruptured cells with Ca²⁺ buffers of estimated Ca²⁺ concentrations, have been reported to inhibit cell coupling^15,^16,^17,¹⁸. However, few studies have examined how [Ca²⁺]_i gates connexin channels in fully intact cells. Furthermore, it has not been shown that cellular Ca²⁺ fluctuations within the physiological range can affect gap junction coupling⁵.

To assess how [Ca²⁺]_i fluctuations affect gap junction coupling in intact cells, we applied three protocols to raise [Ca²⁺]_i in human fibroblasts. They include activating cell surface receptors with agonists such as bradykinin or histamine, elevating [Ca²⁺]_i with a Ca²⁺ ionophore and stimulating capacitative Ca²⁺ influx by emptying intracellular stores. Bradykinin or histamine stimulates cell surface receptors, which activate phospholipase C to produce myo-inositol 1,4,5-trisphosphate, a second messenger that induces Ca²⁺ release from internal stores¹⁹. To activate Ca²⁺ influx, we applied thapsigargin (Tg), an inhibitor of the Ca-ATPase of the endoplasmic reticulum (ER), to cells bathed in Ca²⁺-free Hanks' balanced saline (HBS). This induced a transient [Ca²⁺]_i increase by emptying intracellular Ca²⁺ stores and initiated store-operated capacitative Ca²⁺ influx when Ca²⁺ is added back to the medium²⁰. By comparison with ionomycin, Tg-induced capacitative Ca²⁺ influx is mediated specifically through store-operated Ca²⁺ channels in the plasma membrane. Quantitative measurements of intracellular Ca²⁺ by ratiometric Fura-2 imaging²¹ showed that these procedures increased bulk [Ca²⁺]_i to micromolar concentrations (Supplementary Fig. 3 online). The peak levels of [Ca²⁺]_i approach 1.5

M upon stimulation with ionomycin or bradykinin.

We loaded human fibroblasts with NPE-HCCC2/AM and a long wavelength Ca²⁺ indicator, Fluo-3/AM²². The spectral separation of fluorescence excitation and emission of HCCC2 and Fluo-3 allowed us to image their individual fluorescent signals concurrently. Combining this dual-color imaging method with the LAMP technique allowed us to study Ca²⁺ gating of gap junctions with high spatiotemporal resolution.

Application of bradykinin did not inhibit dye transfer either during the initial transient spike or during the later sustained elevation of Ca²⁺ (Fig. 3a,b). In cases where agonist (histamine) stimulation resulted in oscillation of intracellular Ca²⁺, the rate of dye transfer was also unaffected (Fig. 3c,d). Elevation of intracellular Ca²⁺ to even higher levels with up to 4

M ionomycin also did not inhibit dye transfer (Fig. 3e,f). Further attempts to measure dye transfer rates at even higher [Ca²⁺]_i by using more than 4

M of ionomycin turned out to be difficult. Dramatic cell shrinkage was observed shortly after ionomycin was added, probably due to gross cytotoxicity caused by the very high cellular Ca²⁺ level. These experiments suggested that neither transient Ca²⁺ spikes above 1.5

M, nor prolonged elevations of bulk [Ca²⁺]_i up to 0.3

M, nor Ca²⁺ oscillations were able to reduce gap junction coupling notably in fully intact human fibroblasts.

Figure 3. Agonist or ionomycin stimulated [Ca²⁺]_i rises did not reduce gap junction coupling.

	(a, c and e) HCCC2 transfer rates before and after [Ca²⁺]_i elevations in pairs of coupled human fibroblasts. Bradykinin (a), histamine (c) and ionomycin (e) were added to cells after the first episode of dye transfer. Dye transfer rates to recipient cells (shown on the graph below each episode of transfer, r² values in parentheses) were analyzed as in Fig. 2. (b,d, and f) Concurrent measurements of cytosolic Ca²⁺ fluctuations by Fluo-3 (Excitation 490 10 nm, Emission 525 10 nm, displayed as fold increase over the baseline fluorescence F₀) corresponding to experiments a, c and e, respectively. Grey and black lines represent [Ca²⁺]_i changes of donor and recipient cells, respectively. Dips correspond to decreases of measured Fluo-3 signal during the local uncaging when the iris of field diaphragm was reduced. Plots represent at least three measurements. Full Figure and legend (39K)

Cell coupling is srongly inhibited by capacitative Ca²⁺ influx
In contrast to [Ca²⁺]_i elevations induced by agonists or ionomycin, Ca²⁺ influx strongly inhibited gap junction coupling. After measuring basal dye transfer rates of coupled human fibroblasts bathed in HBS with 1 mM [Ca²⁺]_e, we changed the medium to nominally Ca²⁺-free HBS. After emptying intracellular Ca²⁺ stores with Tg, we added Ca²⁺ back to the medium to initiate Ca²⁺ influx. Continuous Ca²⁺ influx completely stopped dye transfer (Fig. 4a,b). Out of six pairs of coupled cells tested, continuous Ca²⁺ influx for about 8 min completely decoupled four pairs of cells, and it reduced the dye transfer rates by more than 50% in two other cell pairs. The dye transfer rate recovered slowly after Ca²⁺ influx was terminated (Fig. 4c,d). We uncaged the donor cell for the second time after Ca²⁺ influx had been continued for about 500 s. No dye transfer was observed at this time between a cell pair that had been nicely coupled prior to Ca²⁺ influx (Fig. 4c). The third uncaging further raised the HCCC2 concentration gradient between the cells. Still no dye transfer was seen. Later we washed out Tg and added LaCl₃ to block Ca²⁺ influx²³. About 10 min later, [Ca²⁺]_e was switched back to 1 mM. Dye transfer gradually recovered, suggesting that inhibition of gap junction coupling by Ca²⁺ influx is slowly reversible. In contrast, no changes in dye transfer rates were observed in control experiments using either Tg in Ca²⁺-free medium without switching to high [Ca²⁺]_e (Fig. 4e,f), or just changing [Ca²⁺]_e from 0 to 10 mM without prior administration of Tg (Fig. 4g,h).

Figure 4. Capacitative Ca²⁺ influx inhibits gap junction coupling.

	(a,c,e and g) Transfer of HCCC2 between pairs of coupled human fibroblasts. Changes of [Ca²⁺]_e were indicated by bars above the plots. (b,d,f and h) Concurrent measurements of cytosolic Ca²⁺ fluctuations by Fluo-3 corresponding to experiments a, c, e and g, respectively. Full Figure and legend (20K)

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Discussion
We have developed a new imaging assay, LAMP, to study dynamic cell-cell communication through gap junctions. By comparison with other techniques that are currently available for monitoring junctional coupling, our imaging method offers at least three major improvements. First, the LAMP assay is truly non-invasive. NPE-HCCC2/AM can be loaded into fully intact cells and NPE-HCCC2 can be photolyzed with a small dose of UV light because of its extraordinary uncaging cross section¹⁰. In contrast, techniques such as microinjection or patch clamping breach cell membranes and may cause loss or dilution of cytosolic factors involved in the gating of connexin channels. Maintaining cell membrane integrity is especially important for studying the regulation of gap junction communication by cellular biochemical changes. As there is a huge concentration gradient (> 10⁵) of Ca²⁺ across cell membranes and because there are Ca²⁺ channels and Ca-ATPases in plasma membranes mediating Ca²⁺ fluxes across cell membranes, any invasive manipulation that disrupts cell lipid bilayers may perturb cellular Ca²⁺ homeostasis and complicate interpretations of subsequent experiments involving changes of [Ca²⁺]_i.

The second improvement is that the LAMP assay can reliably detect changes in molecular permeability of connexin channels in coupled cell pairs. This important feature mainly results from two desirable properties of the newly developed caged fluorophore: the cell loading efficiency of NPE-HCCC2/AM and the small molecular size of HCCC2. As NPE-HCCC2/AM can be loaded into cells to fairly high concentrations, we can apply the LAMP technique several times in a coupled cell pair once the previous episodes of dye diffusion reach equilibrium. This allows multiple measurements of kinetics of dye transfer in cell pairs and thus detects changes in cell coupling. This type of experiment is difficult to carry out using techniques such as microinjection or gap junction-FRAP because of the invasiveness of the manipulation or the potential photo-damage to cells from the high-intensity illumination required by gap junction-FRAP. The small molecular size of HCCC2 ensures that the fluorophore can quickly diffuse across connexin channels to reach equilibrium in coupled cells. This rapid dye transfer facilitates quantitative analysis of kinetics of junctional diffusion using Fick's equation, and it improves the sensitivity of the detection when there are changes in the permeability of connexin channels. Because the number of connexin channels and cell volumes remain relatively constant within a short time span, the method is ideal for studying how acute biochemical or physiological changes such as Ca²⁺ influxes regulate the gating of gap junctions.

The last major improvement is that the LAMP assay can be used in conjugation with other fluorescent sensors for multi-color imaging. Because HCCC2 emits blue fluorescence, it spectrally complements a variety of fluorescence indicators that emit green or red light. We have demonstrated the simultaneous monitoring of gap junction coupling and cytoplasmic [Ca²⁺]_i changes using the LAMP assay and Fluo-3 (Figs. 3 and 4). Future explorations of combinatory uses of the LAMP assay and other long-wavelength fluorescent sensors should further expand the imaging windows. This may provide new experimental approaches for dissecting how cellular biochemical changes affect cell-cell communication with high spatiotemporal resolution.

The combined advantages of the LAMP assay outlined above are key to our success in identifying a cellular Ca²⁺ signaling pathway that exerts a strong inhibition effect on cell coupling. Few studies have examined how [Ca²⁺]_i changes gate connexin channels in fully intact cells with normal ATP supplies and Ca²⁺ homeostasis. Previous studies of Ca²⁺ gating using Ca²⁺ buffers in ruptured cells usually led to the conclusion that large fluxes of Ca²⁺ up to millimolar concentrations are needed to uncouple cells^5,^17,¹⁸. As such high levels of bulk [Ca²⁺]_i can only be achieved in injured or dying cells with compromised membranes, it has been thought that Ca²⁺ gating may serve as a protection mechanism to isolate damaged cells from their healthy neighbors. Our findings raise the possibility that the process of Ca²⁺ influx through a particular type of plasma membrane Ca²⁺ channel may play physiological roles in modulating the gating properties of connexin channels among healthy cells with intact plasma membranes.

The bulk [Ca²⁺]_i changes caused by bradykinin or ionomycin were of substantially higher amplitudes than that caused by Tg induced Ca²⁺ influx (Supplementary Fig. 3 online). However, local Ca²⁺ elevations immediately beneath plasma membranes during Ca²⁺ influx through membrane Ca²⁺ channels are reported to be orders of magnitude higher than the rise in bulk [Ca²⁺]_i, possibly reaching tens or hundreds of micromolar^24,²⁵. As capacitative Ca²⁺ influx was much more effective in uncoupling cells, this suggests that very high local Ca²⁺ concentrations near the sites of Ca²⁺ influx are probably involved in gating connexin channels. Electron microscopy structures of rat liver gap junctions in the presence of 50

M Ca²⁺ or the absence of Ca²⁺ with 5 mM EGTA have suggested that Ca²⁺ could induce a small cooperative rearrangement of connexin subunits²⁶. Recent studies by atomic force microscopy showed that the cytoplasmic surface of isolated gap junction plaques changes its appearance when 0.5 mM Ca²⁺ was injected²⁷. Such structural changes have been hypothesized to play a role in mediating the gating of connexin channels by high levels of Ca²⁺.

In addition to [Ca²⁺]_i concentrations, cellular acidification is reported to decrease junctional cell communication^5,²⁸ and some studies suggest that there is an apparent interaction between the effects of Ca²⁺ and pH on gap junction coupling^5,^16,¹⁸. Future studies will address this issue by measuring cellular pH during Ca²⁺ influx and examine if there is a synergy of [Ca²⁺]_i increase and cellular pH drop in reducing gap junction coupling when capacitative Ca²⁺ influx is activated. Finally, Ca²⁺ may gate connexin channels indirectly through cytosolic factors such as calmodulin²⁹. [Ca²⁺]_i increases above 1

M from ionomycin or bradykinin stimulation did not inhibit cell coupling, indicating that either the Ca²⁺ affinity of calmodulin is altered³⁰ when it binds to connexins, or high subplasmalemmal Ca²⁺ is required to first induce a structural change in connexins prior to engaging connexin-calmodulin interactions. Whether capacitative Ca²⁺ influx modulates cell coupling directly by local high Ca²⁺ activities or indirectly through cytosolic factors, our results strongly indicate that there are store-operated Ca²⁺ channels localized in close proximity to connexin channels in the plasma membrane. Such a colocalization may allow capacitative Ca²⁺ influx to exert a much stronger gating effect on gap junction coupling than [Ca²⁺]_i elevations from other sources.

Regardless of the detailed mechanisms of inhibiting cell coupling by capacitative Ca²⁺ influx, our results may help to reconcile a dilemma of Ca²⁺ gating of gap junction channels and intercellular Ca²⁺ signaling. If elevated levels of [Ca²⁺]_i reduce the permeability of connexin channels, how do intercellular Ca²⁺ waves spread in confluent cell populations? Perhaps during the propagation of Ca²⁺ waves, Ca²⁺ activities subjacent to the plasma membrane may never reach the threshold level required to close connexin channels. Recent developments in sensor development and cellular imaging have highlighted the importance of distinguishing spatiotemporal aspects of the same biochemical changes in modulating cell responses. Local Ca²⁺ activities from different sources may carry out distinct functions from global Ca²⁺ fluctuations.

The new imaging assay described here for studying intercellular communication may have broad applications for studying gap junction biology in different systems. A comparison of our new imaging technique with previous methods of studying gap junction communication is presented in Table 1. Other caged fluorophores homologous to NPE-HCCC2 can be prepared by conjugating 6-chloro-7-hydroxy-coumarin 3-carboxylate with various amines. This provides a facile synthetic route for making new caged fluorophores for probing the molecular selectivity of connexin channels. In addition to intercellular gap junction coupling, the LAMP assay may also be adapted to study the regulation of connexin hemi-channels (connexons) in live cells. Recent studies have detected cell surface connexons that connect cytoplasm to extracellular medium³¹. How these hemi-channels are activated and what physiological functions they may play are questions of substantial biological interest and have been under intense investigation³². To track the molecular transfer across hemi-channels, most studies have relied on assaying dye uptake by incubating cells in media containing relatively high concentrations (typically around millimolar) of fluorophores such as lucifer yellow. Cells are then washed extensively before cellular fluorescence is measured. Limitations of such an assay include a lack of dynamic information of dye transfer and interference from other cellular processes such as pinocytosis or endocytosis³¹. Moreover, the assay only monitors molecular influx, but not efflux, through connexons. Future development of new assays of hemi-channel communication using cell-permeable and caged fluorophores such as NPE-HCCC2/AM may overcome some of these limitations. Finally, the noninvasive cellular delivery together with remarkable two photon excitation cross sections¹⁰ of this new class of photo-activitable fluorophores may allow us to follow intercellular dye transfer with high three-dimensional resolution in intact tissues or organs by using two-photon uncaging and imaging techniques. This should greatly enhance our capability to examine the dynamic regulation of gap junction coupling and its physiological consequences in vivo.

Table 1. A comparison of methods for studying gap junction communication

	Full Table

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Methods
Cells and reagents.
Human primary fibroblasts (generously supplied by F. Grinnell) isolated from the foreskin of newborns were cultured in high glucose DMEM medium (Gibco) containing 10% fetal bovine serum and 1% penicillin/streptomycin. Cells were used until they reached12−15 doublings. One day prior to an imaging experiment, cells were passed on to 35 mm petri dishes with glass bottoms (MatTek) at about 30% confluence. NPE-HCCC2/AM was synthesized as described¹⁰. Fura-2/AM and Fluo-3/AM were purchased from Molecular Probes.

Dye loading and fluorescence imaging.
To load dyes into cells, cells were washed twice with HBS (Gibco) containing 10 mM Hepes (pH 7.35) and 5.5 mM glucose. Dyes (typically 1

M final loading concentration) in HBS were added to cells. We usually loaded cells for 40 min before washing away excess dyes. Cells were incubated for another 15 min in HBS to allow complete hydrolysis of AM esters.

Fluorescence microscopy was carried out on an inverted fluorescence microscope (Axiovert 200, Carl Zeiss) with a 40 $times$ oil-immersion objective (NA 1.3). Epifluorescence was detected with an ORCA-ER cooled CCD camera (Hamamatsu). Cells were excited with light from a 175 W xenon lamp after passing through appropriate bandpath filters. Switching of excitation wavelengths between UV uncaging and image acquisition was controlled by the Lambda DG-4 high-speed wavelength switcher (Sutter Instrument). Image acquisition and analysis were done using the Openlab integrated imaging software (Improvision).

Note: Supplementary information is available on the Nature Methods website.

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Received 10 September 2004; Accepted 22 November 2004; Published online: 21 December 2004.

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Acknowledgments
This research was supported by a research grant (I−1510) from the Welch Foundation and a Career Development Award from the American Diabetes Associations to W.-H. Li. We thank F. Grinnell for providing primary human fibroblasts. We also thank K. Luby-Phelps, R. Anderson and F. Grinnell for critical comments on the manuscript.

Competing interests statement: The authors declare that they have no competing financial interests.

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