Major translocation of calcium upon epidermal barrier insult: imaging and quantification via FLIM/Fourier vector analysis
- First Online:
- Cite this article as:
- Behne, M.J., Sanchez, S., Barry, N.P. et al. Arch Dermatol Res (2011) 303: 103. doi:10.1007/s00403-010-1113-9
- 196 Views
Calcium controls an array of key events in keratinocytes and epidermis: localized changes in Ca2+ concentrations and their regulation are therefore especially important to assess when observing epidermal barrier homeostasis and repair, neonatal barrier establishment, in differentiation, signaling, cell adhesion, and in various pathological states. Yet, tissue- and cellular Ca2+ concentrations in physiologic and diseased states are only partially known, and difficult to measure. Prior observations on the Ca2+ distribution in skin were based on Ca2+ precipitation followed by electron microscopy, or proton-induced X-ray emission. Neither cellular and/or subcellular localization could be determined through these approaches. In cells in vitro, fluorescent dyes have been used extensively for ratiometric measurements of static and dynamic Ca2+ concentrations, also assessing organelle Ca2+ concentrations. For lack of better methods, these findings together build the basis for the current view of the role of Ca2+ in epidermis, their limitations notwithstanding. Here we report a method using Calcium Green 5N as the calcium sensor and the phasor-plot approach to separate raw lifetime components. Thus, fluorescence lifetime imaging (FLIM) enables us to quantitatively assess and visualize dynamic changes of Ca2+ at light-microscopic resolution in ex vivo biopsies of unfixed epidermis, in close to in vivo conditions. Comparing undisturbed epidermis with epidermis following a barrier insult revealed major shifts, and more importantly, a mobilization of high amounts of Ca2+ shortly following barrier disruption, from intracellular stores. These results partially contradict the conventional view, where barrier insults abrogate a Ca2+ gradient towards the stratum granulosum. Ca2+ FLIM overcomes prior limitations in the observation of epidermal Ca2+ dynamics, and will allow further insights into basic epidermal physiology.
KeywordsCalciumLifetime imagingEpidermisCalcium Green 5NDMSOPhasor analysis
Calcium is the ubiquitous second messenger system in cell biology (e.g., [17, 56]). In epidermis, it controls key events in epidermal barrier homeostasis and repair , neonatal barrier establishment , keratinocyte differentiation [32, 65], and signaling [73, 74], cell adhesion , and a range of pathologic states [49, 53]. Yet, tissue- and cellular-Ca2+ concentrations in physiologic and diseased conditions are only partially known.
To date, there are three well-established methods for calcium measurements in skin. Firstly, state of the art for complex tissues is calcium-PIXE (proton-induced X-ray emission). The publications employing this method have largely defined the current knowledge about the epidermal calcium gradient [9, 21, 46]. Nevertheless, this method measures total calcium concentrations over depth in a line-scan, irrespective of ionization-state or binding, and without information about cellular/subcellular (co-) localization. The resulting finding of a calcium gradient is therefore, in part, a mathematical artifact, i.e., the gradient might result less steep if the denominator were cellular layers, not tissue-depth. Further, calcium-PIXE requires tissue processing (sectioning followed by freeze drying), is only available in very specialized institutions, and is therefore limited to addressing only selected questions. Second, the other method that has been used frequently, a histochemical calcium-precipitation followed by transmission electron microscopy (TEM), requires extensive tissue fixation and processing, and is similarly not suitable for a close to in vivo assessment [49, 50, 78]. Third, calcium is measured in vitro using an array of fluorescent dyes (see below), or through transfection of targeted constructs (e.g., Aequorin) into cultured cells, measuring fluorescence or luminescence to quantify calcium in subcellular compartments [8, 63], creating detailed knowledge of intracellular calcium ranges and leading to insight on their roles .
Table of reported/expected Ca2+ concentrations in skin
Concentration ranges reported
Low (comparable to SB); medium in lower SC, zero in outer SC; (very high)
Gradient rising towards SG; low values for proliferating (SB), and high values for differentiating cells (SS and SG)
Low, comparable to SG; serum levels
Together, these findings have defined the current view of calcium’s role in skin, while observation in vivo/in situ is still lacking. The need for a method to measure and localize Ca2+ in tissue is therefore evident. Recent reviews on the topic point to an apparent lack of experimental options [8, 63], although lifetime imaging was not considered.
Fluorescence imaging and measurement of Ca2+ concentrations have been reported in numerous papers, using a variety of indicator dyes in different modalities, mostly to assess intracellular concentrations of many different cell types [14, 23, 83], as well as keratinocytes and their organelles (e.g., ). Also, advantages and disadvantages comparing Ca2+-sensitive dyes and various methodologies for their use have been discussed extensively [35, 40, 41]. The general method of two-photon Ca2+ imaging has been described for various dyes . Also, fluorescence lifetime imaging (FLIM) to determine Ca2+ concentrations has been tested before [1, 36, 69, 85], and is further detailed conceptually . The first description for Calcium Green 5N (CaG5N)  discussed the advantages of its low Ca2+ affinity, avoiding the underestimations seen with other dyes , and thus indicating its usefulness for investigation in skin. In a parallel paper  we describe the heterogeneity of calcium distribution in human skin using FLIM, and a follow-up details the intracellular calcium-release following barrier perturbation at the SG–SC interface through pharmacologic manipulation .
This report is the continuation of our prior work where we successfully established FLIM to assess epidermal pH [5, 26], demonstrating that this method overcomes the specific limitations of fluorescence-based measurements in complex tissues. Here, we tested this novel approach on the known inducible perturbation of Ca2+ homeostasis, epidermal permeability barrier abrogation in rodent skin. We found a fast, highly dynamic response to an experimental barrier insult, where details partially contradict the view established through earlier methods.
Calcium Green 1, Calcium Green 5N, Rhod 5N and calcium-calibration buffer kits were purchased from Molecular Probes (Eugene, OR). Calcium Chloride dihydrate (C3306, molecular biology grade), and BSA (Bovine Serum Albumin, A7030) were from Sigma-Aldrich, Germany.
Male hairless mice (SKH1 h/hr, Charles River Laboratories, Wilmington, MA, USA) were fed Purina mouse diet and water ad libitum. Animals were 8–12 weeks old at time of experiments. Conventional surface pH measurements were performed using a flat glass surface electrode (Mettler-Toledo, Giessen, Germany) attached to a pH meter. The Stratum corneum (SC) was removed/stripped by several sequential strippings with D-squame disks (Acaderm, Menlo Park, CA, USA), inducing an increase in transepidermal water loss levels (TEWL) above base line (from ~0.2 to ~7–9 g/m2/h), and measured at 0, 2, and 18 h. Animal experiments were performed at the UIUC animal research facilities at the Urbana-Champaign campus, IL, USA and the subsequent FLIM experiments at the Laboratory for Fluorescence Dynamics, at the time of the experiments located at the Department of Physics, University of Illinois, IL, USA. Preliminary experiments were also performed at UCSF, and control experiments at the Dermatology Department, University Medical Center Hamburg-Eppendorf, and Anatomical Institute, University Hannover, both Germany.
Transmission electron microscopy (TEM)
Biopsies from mouse skin freshly obtained at time points matching the dye application protocol and earlier (untreated, 1,2,18 h) were fixed directly for 48 h in Karnovsky′s fixative , washed in PBS, and postfixed in buffered 1% osmium tetroxide . After careful dehydration in graded ethanol, the skin samples were embedded in Epon 812 (Serva)  and cut with a diamond knife on an ultramicrotome Ultracut E (Leica). Thin sections (<100 nm) were stained with methanolic uranyl acetate  and lead citrate , then viewed in an electron microscope EM10C (Zeiss) operated at 60 kV.
Calcium Green 5N was applied once (5 ul of a 0.1 mM solution in DMSO, to an approximate area of 5 mm diameter), and a biopsy was taken 18 h following dye application, in preliminary experiments also at 1 and 2 h, mounted for microscopy, and directly visualized. Control images to assess dye distribution in skin were taken on a Zeiss Axiophot, equipped with a Hamamatsu C7472 camera. For this purpose, freshly obtained hairless mouse skin biopsies in OCT compound were flash-frozen in liquid nitrogen, cut on a cryostat to 6 μm sections, coverslipped and viewed.
TEM, fluorescence microscopy, pH and TEWL measurements as well as the preliminary work to establish the choice of dye, were conducted as separate experiments. The functional experiments, imaging of calcium distribution via FLIM pre- and post-barrier abrogation were conducted as follows: Calcium Green 5N was applied once to one flank area of an anesthetized mouse. 18 h following dye application, a biopsy was taken without further treatment for one area. Barrier abrogation was performed on separate animals, treated otherwise identically, as the imaging process overall was too lengthy to conserve biopsies from one animal for both conditions.
Fluorescence lifetime imaging microscopy
In brief, two-photon FLIM to determine calcium was performed using a Millenia-pumped Tsunami titanium:sapphire laser system (Spectra-Physics) as the two-photon excitation source. Excitation of the sample was achieved by coupling the 800 nm output of the laser through the epifluorescence port of a Zeiss Axiovert microscope. The fluorescence was collected using a Hamamatsu (R3996) photomultiplier placed at the bottom port of the microscope. Scanning mirrors and a 40× infinity corrected oil objective (Zeiss F Fluar, 1.3 N.A.) were used. Z-slices (1.7 μm per slice) were obtained by adjusting the objective focus with a motorized driver (ASI Multi-Scan 4). Lifetime data were acquired using time-correlated single photon counting (TCSPC). Fluorescein was used as the reference lifetime standard (τf = 4.05 ns, pH 9.5). Background fluorescence was determined to be negligible, in accordance with our prior data . Additionally, DMSO was not expected to alter our measurements as its use in fluorescence is facilitated because of its optical transparency . Further, background fluorescence would be excluded via phasor analysis (see below). Data evaluation and visualization were performed directly with the in-house software SIM-FCS (http://www.lfd.uci.edu/globals/). Individual images were combined using Adobe Illustrator (Adobe Systems Incorporated, San Jose, CA, USA), but no further image processing was performed. Background fluorescence was measured in samples of unstained tissue, treated otherwise identically. To facilitate comparison of different experimental conditions, i.e., various dyes and skin pretreatments, we imaged morphologically similar sites, where the columnar arrangement in SC also, if to a lesser degree, is reflected in a regular arrangement of cells in layers underneath [12, 30, 82], as outlined in a prior publication .
Calibration via phasor plot
Briefly, in time-correlated single photon counting multiple lifetime components from different molecular species or different conformations of the same molecule are analyzed via exponential fitting of decay times, pixel by pixel for lifetime imaging. The phasor approach decomposes the decay into Fourier components for mathematical analysis in vector algebra, which is more easily computed than exponential fitting routines. Different proportions of the two fluorescent species expected for the indicator molecules used, here calcium-bound and free dye, arrange in one line in the geometrical display of such vectors, the phasor plot (Figs. 2, 3, panels e). Artifact components from background or other molecular species will not arrange along that same line, and are thus identified and excluded from the evaluation (for further detail please refer to ). Nevertheless, in the experiments presented here, we did not observe such effects. The images presented here were recalculated and are displayed calibrated based on the phasor plot derived from a calibration in a series of calcium-containing buffers (see “Results”).
Dye application and distribution, tissue morphology, and epidermal function
In control experiments parallel to dye application, we assessed changes in TEWL and surface pH following DMSO application. Changes in surface pH over time were observed, i.e., an initial rise of about 0.5 pH units over the first 2 h, and restoration to initial values by 18 h. As treated and untreated mice (i.e., mice which had been anesthetized to measure pH and TEWL) displayed identical behavior, we attributed these changes to stress of manipulation , side effects of anesthesia, and circadian rhythm . TEWL underwent similar, although transitory larger changes, which were to be expected for the solvent properties of DMSO. Initially, in parallel to the wheal reaction, there was a steep rise of TEWL in DMSO treated skin, which subsided, i.e., returned to within less than 15% difference of starting value, 18 h following DMSO application (data not shown). Further, in initial experiments we compared the Ca2+ distribution in biopsies taken at 2 and 18 h following application of CaG5N in DMSO. We did not discern any major differences between these time points, and for reasons of practicability chose the latter, overnight time point thus also excluding lingering DMSO effects. Finally, none of the mice treated in this manner ever displayed signs of discomfort, or scratching at the site of dye–DMSO application.
Calibration of CaG5N in lifetime experiments
CaG5N calibrations reported
Rat hepatocyte IP3
In vivo/in vitro
in vivo on hepatocytes; identical values in vitro and in vivo
Rat cerebellar Purkinje neurons
Use of CaG5 N to reduce buffering of Ca2+
Frog skeletal muscle fibers
Calibration via Fmin/Fmax; patch-clamp controlled
Recombinant cytosolic PLA2
Calibration on erythrocyte ghosts
Rat cerebellar basket cell axons
Rationale for choice of dye
in vitro only
Rat brain neurons
In vivo/in vitro
Additionally in vivo calibration determining Fmin/Fmax
Turtle hair cells
Limulus ventral photoreceptor
Fluorescence ratio measurements using ANTS, Ca2+-sensitive electrode controlled, Mg2+ had no effect
Skeletal muscle fibers
Honeybee drone photoreceptors
Aging mouse skeletal muscle fibers
Calibration on muscle fibers
Adult skeletal muscle fibers
No effect of CaG5N on oscillatory membrane current responses
Frog skeletal muscle fibers
Protein binding of dyes affected response time, not sensitivity; extensive dye comparison
Absorbance in muscle fibers
Calcium distribution in epidermis at steady state
We initially tried to establish whether typical concentration ranges for distinct areas of epidermis existed, in parallel to the reported calcium gradient across epidermis (please also compare to Table 1). We rather found that within certain value-ranges Ca2+ distribution stayed identical. Therefore, the individual images displayed in Figs. 2a, b, 3a, b were set to a low, intermediate and high range (corresponding to columns b, c, and d), at the same time exploiting the sensitivity range of the indicator dye, and in the process resulting to be typical for the extracellular, intracellular, and possibly the bound or calcium-store compartments.
We then used this protocol to ascertain calcium distribution in undisturbed rodent skin, baseline condition for our comparisons. We found low Ca2+ concentrations in the extracellular compartment at the surface, i.e., in the interstitial areas of the SC, extending to, but not beyond the SC/SG interface (Fig. 2, first and second rows, column b).
Ca2+ concentrations in a medium range can be found throughout the epidermis, restricted to the intracellular compartment of the SC, quasi delineating the cell membranes and intracellular compartment in the SG, with occasional areas of increased Ca2+ concentration indicating intracellular compartments (arrow 1), and possibly extending to the extracellular compartment in lower epidermal layers (Fig. 2, first through fourth rows, column c, arrow 2).
High Ca2+ concentrations were not found within the SC, but throughout the epidermis and limited to the intracellular compartment only (Fig. 2, second through fourth rows, column d).
Calcium distribution in epidermis following barrier disruption
Finally, we tested this imaging approach on the known, inducible perturbation of Ca2+ homeostasis, epidermal permeability barrier abrogation in rodent skin.
The limitations of earlier approaches to calcium imaging were outlined in the introduction, and from a biologic perspective, the time delay between a given physiologic state and its microscopic observation is the largest artifact introduced with an experiment, especially as fast intra- and extracellular calcium signaling is firmly established from cell-culture experiments. We therefore aimed at an early time point for our experiments, and reproducibly could start acquiring images 30 min past tapestrip and preparation of a biopsy for microscopy.
At this early time point, we found low Ca2+ concentrations only extracellularly at the surface and SC/SG interface (Fig. 3, first and second rows, column b, arrow 1).
Medium concentrations were found, now more pronounced, intracellulary in the SC to SC/SG interface (arrow 2), while below the SC/SG-interface medium concentrations are shifted to or outline the cell membranes and extracellular compartment of the epidermis (arrow 3), and in the stratum basale (SB) are confined again to intracellular areas (Fig. 3, first through fourth rows, column c, arrow 4).
High Ca2+ concentrations post-tapestrip can now be found more evenly distributed throughout the cells in all upper layers (Fig. 3, first through third row, column d), occasionally even intracellularly in the SC (Fig. 3, first row, column d) and increasingly downwards to the SB, at the same time shifting from strictly limited and delineating the intracellular compartment in the SG (arrow 5) to delineating the cell membranes and cell periphery in the SB (Fig. 3, first through fourth rows, column d, arrow 6). Together, we find a rapid shift of Ca2+ towards higher concentrations and apical epidermal layers, including SC. As a summary of our findings please refer to Fig. 4.
Technical considerations and limitations
First, we could not detect calcium below the epidermis, which we attribute to the anatomy of rodent skin where very little dermal structure and therefore no label acquiring, i.e., calcium-binding or -retaining structures could be observed. In other species, the dermal compartment might serve as an internal reference, as values from serum should equal concentrations found in dermis (see legend to Table 1). Thus, in rodent skin only basal-layer values may be compared to serum levels. In our data, the intermediate range values to be found intra- and extracellularly in the basal layer of non-disrupted skin appears as the next best reference, and again fits with these findings.
Second, of concern are also the physico-chemical characteristics of DMSO; it is a hygroscopic, amphiphilic, aprotic, small size solvent for organic and inorganic compounds as well as macromolecules , which readily penetrates biological membranes and skin. There is a large body of work on the use of DMSO as a cryoprotectant, and since the initial description and speculation on possible mechanisms , there was little progress in elucidating the mechanisms involved. A recent and detailed review on the use of DMSO  lists multiple pharmacological, cellular, and molecular aspects. Transient, rather rapid effects on calcium, observed in cells and isolated organs also were reported. From our preliminary experiments to establish the method used here, we concluded that the 18-h delay between application of DMSO and the functional experiments had a compensatory effect, leading to complete absorption of the minor amount from skin and elimination from the living animal. Accordingly, our control experiments demonstrated only an early and transiently affected epidermal function and no further discernible untoward effects of DMSO. As corroborating observation might serve the fact that we did not find an accumulation of CaG5N fluorescence inside corneocytes in undisturbed epidermis (Fig. 1, panel c), where the most direct or intense DMSO exposition occurred. Also, the distinct FLIM signal from this compartment is in accordance with the underlying layers (compare first and second rows, Figs. 2, 3). Based on these reports and our preliminary experiments to establish CaG5N labeling, we considered the use of DMSO safe and non-interfering in our experimental protocol.
Third, in respect to the indicator dye, there is the additional, general caveat that there may be more-than-single-site Ca2+ binding at low Ca2+ concentration [76, 88]. A different, to date unpublished explanation of this effect postulates two fluorescent species at low-to-zero Ca2+, where free Calcium green-1 has a closed form with short lifetime, and an open, bound to Ca2+ form with longer lifetime (personal communication; David Jameson, University of Hawaii). For its chemical structure, this concern should apply to CaG5N also, but is also resolved through the phasor analysis, which eliminates a host of artifacts common to fluorescence based methods (see Preliminary Experiments).
Lastly, we here present data early following barrier disruption, as such observations were heretofore not possible. A series of experiments covering the complete time-course of epidermal permeability barrier recovery will hopefully provide more insight. At the same time, our prior work described the changes of pH within the extracellular compartment of the SC  in response to barrier disruption, which we now show similarly for calcium within epidermis. Whether and how both calcium and pH are mechanistically connected is unclear, but such a connection might follow also from our prior work [3, 20, 21]; both aspects are topic of an ongoing project in our lab.
Findings in skin
As compared to earlier reports on Ca2+ in skin, our images display Ca2+ distribution and their dynamics in epidermis in detail in close to in vivo conditions. Intra- and extracellular compartments across epidermal depth can mostly be distinguished, and although subcellular compartments cannot be identified in our approach, a number of images hint at intracellular structures, and at least suggest the existence of intracellular stores of higher Ca2+ concentration, which are firmly established through other methods .
Through previous observations only a uniform calcium gradient across epidermal depth could be distinguished (see "Introduction"), and the redistribution upon barrier abrogation was known, although with limited detail. The experimental data shown here depict the dynamic, cellular redistribution of calcium in epidermis following a barrier insult, as summarized in Fig. 4 and thus partially contradicts the previous concept. In undisturbed skin, low concentrations are found from surface to SC/SG interface, and medium concentrations below these layers. Upon barrier disruption, we found a fast change of Ca2+ distribution. The lowest concentrations almost disappear, while medium and higher concentrations are now present in SG and SB, respectively. The pattern emerging from this more detailed view now shows a shift of calcium upon barrier insult, from deeper-layer compartments, moved to the cell membranes and upward through the epidermis, a dynamic shift within cellular localizations and epidermal layers. This same shift, although at the resolution of our images not entirely distinguishable, can be found in the extracellular spaces. Overall, calcium is shown in a differential distribution, which still may be viewed as a gradient of sorts, albeit barrier disruption, with the time-wise closer look provided in the method described here, shows no abrogation of the Ca2+ gradient. Our findings are summarized schematically in Fig. 4. The membrane orientation of calcium upon barrier insult is consistent with a signaling role of calcium. The upward shift of higher concentrations following barrier disruption indicates a calcium-loss when barrier function is defective, but also its involvement in the biochemistry of barrier repair. Probably temporarily, these shifts involve an extracellular rise in calcium, for the upward shift and eventual loss in case of a defective barrier. Furthermore, a rapid barrier restoration may even hint at a calcium-conserving strategy, which is mostly known from bone metabolism and the interplay of vitamin D and parathyroid hormone to maintain calcium-levels (e.g., ). In epidermis, especially in the case of psoriasis this also holds true, although further roles and functions remain to be explored (e.g., ).
With FLIM, we now visualize close to in vivo biologically relevant changes of Ca2+ distribution over epidermal depth, at cellular resolution, without tissue processing, minimizing artifact. Specifically, TCSPC offers precision at low light intensities, and in conjunction with the novel phasor analysis, visual control over the exclusion of artifactual contributions in the mathematical analysis is provided. We view this approach as key to further insight into regulation, coordination and orchestration of barrier repair and other calcium-dependent processes in skin. Future experiments, using advanced equipment currently being established, will allow us to gain further insight into the time-dependent processes of reestablishing barrier function, and will have to show whether it is barrier status per se which regulates the formation of the epidermal calcium gradient, as earlier data shows , or if calcium itself contributes to the regulation of barrier status.
Finally, with increasing dissemination of two-photon-equipment, the method used here should become more frequently utilized and help to extend this approach beyond rodent epidermis into deeper layers of mammal epidermis at subcellular resolution.
This work was supported through a grant from the European Community’s Marie-Curie-Program, MC-IRG 6675 (to MB). We thank Mrs. Monika Thiel for expert assistance in preparing graphics
Conflict of interest
The authors declare that they have no conflict of interest.