Archives of Dermatological Research

, Volume 303, Issue 2, pp 103–115

Major translocation of calcium upon epidermal barrier insult: imaging and quantification via FLIM/Fourier vector analysis

Authors

    • Department of Dermatology and VenerologyUniversity Medical Center Hamburg-Eppendorf
  • Susana Sanchez
    • Laboratory for Fluorescence Dynamics, Department of Biomedical EngineeringUniversity of California
  • Nicholas P. Barry
    • Department of NephrologyUniversity of Colorado Health Science Center
  • Nina Kirschner
    • Department of Dermatology and VenerologyUniversity Medical Center Hamburg-Eppendorf
  • Wilfried Meyer
    • Anatomical InstituteUniversity Veterinary Medicine Foundation
  • Theodora M. Mauro
    • Veterans Affairs Medical Center Dermatology DepartmentsUniversity of California San Francisco
  • Ingrid Moll
    • Department of Dermatology and VenerologyUniversity Medical Center Hamburg-Eppendorf
  • Enrico Gratton
    • Laboratory for Fluorescence Dynamics, Department of Biomedical EngineeringUniversity of California
Original Paper

DOI: 10.1007/s00403-010-1113-9

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

Abstract

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.

Keywords

CalciumLifetime imagingEpidermisCalcium Green 5NDMSOPhasor analysis

Introduction

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 [48], neonatal barrier establishment [21], keratinocyte differentiation [32, 65], and signaling [73, 74], cell adhesion [2], 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 [7].

The combined view from these reports renders a rough “map” of calcium values reported and to be expected in skin (please refer to Table 1). From cell culture experiments, where regulation of proliferation and differentiation through calcium levels is standard procedure (e.g., [7, 28, 42]), intracellular concentration ranges are established. Virtually, all organelles are Ca2+ stores [51], and in general, low cytoplasmic values (0.1 μM) are distinct from several fold higher organelle values (e.g., ER 500 μM) [17]. Reports about Ca2+ concentrations in skin are somewhat divergent, hardly comparable qualitative [48, 49] and quantitative methods [25], with the generally accepted values from PIXE experiments [47], where calcium from serum-like levels in the SB roughly triple in the SG, to abruptly drop below serum levels in the SC.
Table 1

Table of reported/expected Ca2+ concentrations in skin

 

Concentration ranges reported

References

SC

Low (comparable to SB); medium in lower SC, zero in outer SC; (very high)

[47], [49], [24, 25]

SB–SG

Gradient rising towards SG; low values for proliferating (SB), and high values for differentiating cells (SS and SG)

[24, 25, 4749], [7, 28, 42]

Organelle

High

[17]

Dermis

Low, comparable to SG; serum levels

[47, 49], [71, 84]

The combined view of various reports renders a rough “map” of calcium values reported and to be expected in skin, with divergent findings in parentheses

In cell-culture experiments, regulation of keratinocyte proliferation and differentiation through calcium levels is standard procedure, and the concentration ranges are established for various mouse- and human-keratinocyte cell types and lines (e.g., [7, 28, 42])

Virtually, all organelles are Ca2+ stores [51], and in general, low cytoplasmic values (0.1 uM) are distinct from several fold higher organelle values (e.g., ER 500 uM) [17]

We exemplarily measured serum values for calcium (2.6 mmol/l), and inorganic phosphorous (3.3 mmol/l) in a male hairless mouse. These values are well in accordance with averages of 40 mouse strains displayed in The Mouse Phenome Database (http://aretha.jax.org/pub-cgi/phenome/mpdcgi?rtn=docs/home) [71]

Reports about Ca2+ concentrations in skin are somewhat divergent, hardly comparable qualitative [48, 49] and quantitative methods [25], with the generally accepted values from PIXE experiments [47], where calcium from serum-like levels in the SB roughly triple in the SG, to abruptly drop below serum levels in the SC

Although for skin a vacuum PIXE technique was applied [9], the values obtained there for dermis were quite similar to newer “air” measurement values for serum samples [31], which again compared well to standard serum electrolyte analyses [84], and their established reference values

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., [4]). 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 [37]. Also, fluorescence lifetime imaging (FLIM) to determine Ca2+ concentrations has been tested before [1, 36, 69, 85], and is further detailed conceptually [16]. The first description for Calcium Green 5N (CaG5N) [59] discussed the advantages of its low Ca2+ affinity, avoiding the underestimations seen with other dyes [68], and thus indicating its usefulness for investigation in skin. In a parallel paper [11] 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 [10].

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.

Methods

Materials

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.

Animal experiments

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 [33], washed in PBS, and postfixed in buffered 1% osmium tetroxide [52]. After careful dehydration in graded ethanol, the skin samples were embedded in Epon 812 (Serva) [45] and cut with a diamond knife on an ultramicrotome Ultracut E (Leica). Thin sections (<100 nm) were stained with methanolic uranyl acetate [67] and lead citrate [61], then viewed in an electron microscope EM10C (Zeiss) operated at 60 kV.

Fluorescence microscopy

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.

Experimental procedure

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 [26]. Additionally, DMSO was not expected to alter our measurements as its use in fluorescence is facilitated because of its optical transparency [87]. 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 [5].

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 [16]). 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”).

Results

Preliminary experiments

Dye application and distribution, tissue morphology, and epidermal function

In preliminary experiments, we tested dyes from the Calcium Green- and Rhod-series, and settled on CaG5N as its use resulted most advantageous for its quantum yield and range of calcium sensitivity (14 μM, Invitrogen/Molecular Probes), as suggested by a number of prior publications [13, 39, 43, 59, 76]. This dye is only poorly soluble in the recommended pH >6 aqueous medium (Invitrogen/Molecular Probes), or alcoholic solution and did not penetrate epidermis in these solvents. Fluorescence could therefore be detected at the SC surface only (not shown). We next injected CaG1 in DMSO intraepidermally and followed dye distribution/fluorescence over time. Within 30 min, we could observe the regular aspect where epidermal cells could easily be distinguished (data not shown). Nevertheless, the dye injection and the following intraepidermal distribution resulted poorly reproducible. We therefore established topical application of a dye–DMSO solution to the skin of hairless mice as routine for our experiments. Dye distribution across the epidermis following topical application was exemplarily assessed with standard fluorescence microscopy, showing a distribution gradient, but nevertheless dye penetration across the entire epidermal depth (Fig. 1c). We next performed a number of control experiments to assess whether the application of DMSO left a discernible effect on epidermal permeability barrier function. Macroscopically, application of DMSO leads to the known wheal [34] (Fig. 1a), and after its disappearance (at the earliest between 60 and 90 min post application) when a dye–DMSO solution was applied, a slightly stained spot remained (Fig. 1b), which was biopsied at 18 h for lifetime experiments. By electron microscopy (EM) of dye–DMSO-exposed skin, compared to untreated, gross morphology, as well as nuclei/ER/SC appeared normal at all time points (1, 2, 18 h) (Fig 1d, 18 h). At 1 h, occasional vacuoles in granular layer cells could be observed (not shown); further ultrastructural alterations were not detected and therefore we did not routinely perform further EM controls.
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Fig. 1

Topical dye/DMSO application and distribution, and gross and tissue morphology. Panel a wheal reaction from DMSO only. Panel b CaG5N spot remaining at 18 h post application in DMSO. Panel c fluorescence microscopy image of a histologic section of mouse skin, CaG5N distribution 18 h following application in vivo. Panel d electron micrograph of mouse skin at 18 h post DMSO application

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 [15], side effects of anesthesia, and circadian rhythm [18]. 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

For in vitro calibration, we used established buffer kits [72], in the highest concentrations on occasion extended with solutions prepared from a calcium chloride stock. We repeatedly performed single-point fluorescence measurements, using different buffer batches to exclude dilution errors. We found very little variation in the buffer measurements (at n = 4 an SEM between 0.00 and 0.20), with the resulting Kd of 4.9 μM, well in range with other reported calibrations (see Table 2). Such buffer systems [72] address correction for viscosity [58], ionic strength [27], and presence of proteins [55, 62]. Also, in our prior work on lifetime measurements of pH in SC, we established that lipids, i.e., a saturated solution of cholesterol, did not alter Kd [5]. Nevertheless, for the purposes of this study, measurements in the protein-rich epidermal layers rather than SC, it appeared imperative to ascertain the influence of protein on the calibration. To test whether presence of protein affected Kd, we added BSA in increasing concentrations (9.1–20%) to the buffer system. As the BSA used contained calcium (according to specifications between 0.004 and 0.007%), a shift corresponding to less than 0.005 mM calcium was to be expected. Instead, we found an attenuation of sensitivity, introduced in both the low and high concentration ranges; similar dye behavior has been reported earlier for CaG1 [70]. Yet, most importantly, the resulting Kd underwent only little change (less than 0.6 μM between buffer and highest BSA concentration), and these measurements were as invariant as the buffer-only measurements. Consequently, and for reasons of reproducibility, we based further calculations on the Kd derived from measurements in the prefabricated calcium-buffer kit without protein additions. This process is further validated through the use of the phasor plot, which requires only a Kd to compute images (ref. to “Methods” and [11]).
Table 2

CaG5N calibrations reported

References

Tissue/target

Kd (μM)

Calibration

Comments

Combettes [13]

Rat hepatocyte IP3

35

In vivo/in vitro

in vivo on hepatocytes; identical values in vitro and in vivo

Eilers [19]

Rat cerebellar Purkinje neurons

17

In vitro

Use of CaG5 N to reduce buffering of Ca2+

Escobar [22]

Frog skeletal muscle fibers

45

In vitro

Calibration via Fmin/Fmax; patch-clamp controlled

Hixon [29]

Recombinant cytosolic PLA2

50

In vitro

 

Kubitscheck [38]

Erythrocytes

2

In vivo

Calibration on erythrocyte ghosts

Llano [43]

Rat cerebellar basket cell axons

20

In vitro

Rationale for choice of dye

Naraghi [54]

in vitro only

23.1

In vitro

 

Peretz [57]

Drosophila photoreceptors

25

Not mentioned

 

Rajdev [59]

Rat brain neurons

4.3

In vivo/in vitro

Additionally in vivo calibration determining Fmin/Fmax

Tucker [75]

Turtle hair cells

25

In vitro

 

32

In vivo

 

Ukhanov [76]

Limulus ventral photoreceptor

67

In vitro

Fluorescence ratio measurements using ANTS, Ca2+-sensitive electrode controlled, Mg2+ had no effect

Vergara [77]

Skeletal muscle fibers

85

In vitro

 

Walz [79]

Honeybee drone photoreceptors

74

In vitro

 

Wang [80]

Aging mouse skeletal muscle fibers

31

In vivo

Calibration on muscle fibers

Wang [81]

Adult skeletal muscle fibers

33

In vitro

 

Yao [86]

Xenopus oocytes

10–12

In vitro

No effect of CaG5N on oscillatory membrane current responses

Zhao [88]

Frog skeletal muscle fibers

  

Protein binding of dyes affected response time, not sensitivity; extensive dye comparison

156

In vivo

Absorbance in muscle fibers

63

In vitro

Absorbance measurements

An extensive literature search yielded a number of publications where CaG5N was calibrated. The table lists reports with original determinations of Kd values for Ca2+ binding, with the tissues investigated, and method of calibration. In vitro calibrations generally were performed using a defined calcium buffer series, as used for the experiments reported in this manuscript, or similar buffer systems. Values in this table were obtained using the method of fractional changes in fluorescence (ΔF) above the resting baseline (ΔF/F)

Given the fact that different methods and setups were used, the wide span of values is not surprising. Our resulting Kd of 4.9 μM fits well into the range of most values (refer to “Results”)

The phasor plot offers a number of advantages for the purpose of imaging ionic concentrations; our recent paper [11] further details this method beyond the already published concept [16]. In short, this approach displays the whole of calcium-values obtained in the image series presented here (Figs. 2, 3, phasor plots in panels e) in form of a cloud, superimposed with the calibration-curve obtained in buffers of distinct calcium concentrations. The distribution of experimentally obtained values along this calibration graph therefore justifies the choice of dye, as values are evenly spread along and around the calibration plot. Had we chosen an indicator dye with a sensitivity range not suited for our samples, experimental values could be expected to arrange towards the low or high end of the calibration graph, or not be covered by the calibration altogether. Different fluorescent species than the two forms expected from the indicator dye (e.g., from other, interfering ions) would likewise be evident in the phasor plot, but were not observed. In different approaches to lifetime-analysis such species would be difficult to discern, but, when present, complicate mathematical fittings. Further, a shift of pH would only affect the SC-extracellular domain, as shown in our prior data [5, 6, 26]. Thus, we concluded that our measurements were suitable for rodent skin, and artifactual lifetime components were not present in our images.
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Fig. 2

Calcium distribution and concentrations in undisturbed rodent epidermis, two figures from separate experiments. Fluorescence lifetime imaging of hairless mouse skin, stained with CaG5N overnight to image areas of 107 μm2. Experiments were performed in triplicate (n ≥ 3), and typical images are shown here. Column a: fluorescence-intensity images were adjusted to enhance structural features and to visualize progression of the microscopic focus in z-direction into the epidermis. SC to SG and following are continuous images, two further optical sections until reaching the SB were omitted Columns bd: images identical to column a, overlaid with the red signal indicating calcium distribution at the concentrations indicated at the top of the columns. Panel e: phasor plot of the cloud of calcium values in the universal phasor circle. The cloud represents all values of the panels in columns bd, overlaid with the calibration line resulting from the series of calcium-containing buffers used to calculate the calcium signal in columns bd. Arrows to indicate specific concentrations, in Fig. 2a, please also refer to “Results”: intracellular compartments (arrow 1). Extracellular compartment in lower epidermal layers (arrow 2). Mutually exclusive subcellular compartments for medium and high concentrations (arrows 3). Intracellular-submembrane accumulation for the intermediate concentration (arrow 4)

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Fig. 3

Calcium distribution and concentrations in barrier-abrogated epidermis, two figures from separate experiments. Samples were taken form separate animals than the undisturbed, treated otherwise identically. Conditions and order of panels matching Fig. 2. Note that the cloud of calcium-values in panel e is left-shifted, representing slightly higher values overall. Arrows to indicate specific concentrations, in Fig. 3a, please also refer to “Results”: low Ca2+ concentrations extracellularly at the surface and SC/SG interface (arrow 1). Medium concentrations intracellulary (arrow 2), cell membranes and extracellular compartment (arrow 3), and in the stratum basale (SB) in intracellular areas (arrow 4). High Ca2+ concentrations delineating the intracellular compartment in the SG (arrow 5) and delineating the cell membranes and cell periphery in the SB (arrow 6)

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).

As the method and dye application protocol does not allow counterstaining, the only approach to attributing distinct Ca2+ concentrations to cellular layers and structures is the comparison of the specific, calibrated calcium signal to the separately displayed fluorescence-intensity images (Figs. 2, 3, columns a), and their progression with the microscopic depth focus. It was therefore impossible to identify specific structures beyond cellular outlines over depth; nevertheless, the outlines generated through the Ca2+ signal occasionally appear as mutually exclusive subcellular compartments for medium and high concentrations (e.g., Fig. 2, second row, columns c and d, arrows 3). The membrane delineation via the Ca2+ signal may indicate an intracellular-submembrane accumulation for the intermediate concentration, consistent with a signaling role for this concentration range (Fig. 2, column c, second and third rows, arrow 4). Additionally, although an unequivocal in- or exclusion cannot be made at the current resolution, the images are also suggestive of distinct Ca2+ concentrations within the extracellular spaces in epidermis. Taken together, we find a specific pattern of Ca2+ concentrations at baseline, also summarized in Fig. 4.
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Fig. 4

Schematic summary of results. View of epidermis pre- (left panel) and post-tapestrip (right panel), summarizing the details of changes in calcium distribution induced by acute barrier disruption. Note the increase in concentrations, from intracellular stores to the cell periphery and extracellular space, then upward in epidermis. The concentration ranges used match the actual data displayed in Figs. 2, 3, i.e., low (0.1 μM), medium (1 μM), and high (10 μM)

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.

Discussion

Technical considerations and limitations

The comparison to calcium values reported elsewhere or obtained from different experiments fits well with our findings (refer to Tables 1, 2), with several technical caveats:

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 [87], 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 [44], there was little progress in elucidating the mechanisms involved. A recent and detailed review on the use of DMSO [64] 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 [5] 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 [51].

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., [66]). In epidermis, especially in the case of psoriasis this also holds true, although further roles and functions remain to be explored (e.g., [60]).

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 [20], 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.

Acknowledgments

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.

Copyright information

© Springer-Verlag 2010