Purinergic signalling mobilizes mitochondrial Ca²⁺ in mouse Sertoli cells

Non-technical summary

In mammalian testes, Sertoli cells play a key physiological role in germ cell development. Previous research has implicated local ATP release as a potential mechanism of Sertoli cell stimulation. We show that, in mouse Sertoli cells, two different receptor proteins are activated by ATP. Receptor activation, in turn, causes elevation of calcium ion levels inside the cells. By using a novel method to visualize such calcium signals, we identify mitochondria as essential elements of calcium regulation in the testis.

Abstract

Abstract

Intimate bidirectional communication between Sertoli cells and developing germ cells ensures the integrity and efficiency of spermatogenesis. Yet, a conceptual mechanistic understanding of the physiological principles that underlie Sertoli cell autocrine and paracrine signalling is lacking. Here, we characterize a purinergic Ca²⁺ signalling network in immature mouse Sertoli cells that consists of both P2X2 and P2Y2 purinoceptor subtypes, the endoplasmic reticulum and, notably, mitochondria. By combining a transgenic mouse model with a dedicated bioluminescence imaging device, we describe a novel method to monitor mitochondrial Ca²⁺ mobilization in Sertoli cells at subcellular spatial and millisecond temporal resolution. Our data identify mitochondria as essential components of the Sertoli cell signalling ‘toolkit’ that control the shape of purinergic Ca²⁺ responses, and probably several other paracrine Ca²⁺-dependent signals.

Introduction

The seminiferous tubule represents the functional unit of the testis. In the seminiferous epithelium, Sertoli cells are each intimately associated with ≥ 30 germ cells at all stages of the rodent spermatogenic cycle (Mruk & Cheng, 2004). Synchronized and bidirectional Sertoli–germ cell interactions at the molecular and biochemical level are, thus, imperative for spermatogenesis and reproduction (Cheng & Mruk, 2002; Griswold & McLean, 2006).

Endocrine control of spermatogenesis along the hypothalamic–pituitary–testicular axis functionally converges on Sertoli cells (Gorczynska-Fjalling, 2004). Though incompletely understood, the synergistic regulation of Sertoli cell function by follicle-stimulating hormone (FSH) and testosterone is based on a common mechanism – mobilization of cytosolic Ca²⁺ (Gorczynska-Fjalling, 2004). Recently, several groups suggested that purinergic signalling constitutes an additional important component of the testicular paracrine/autocrine communication system (Filippini et al. 1994; Foresta et al. 1995; Gelain et al. 2003).

In adult rat Sertoli cells, extracellular ATP triggers a rapid transient increase in the cytosolic Ca²⁺ concentration ([Ca²⁺]_c) that appears to be mediated by different P2 purinoceptors (Filippini et al. 1994; Foresta et al. 1995; Lalevee et al. 1999; Ko et al. 2003). Their molecular identity, however, remains controversial. The ATP-dependent Ca²⁺ signal counteracts various FSH-mediated effects and alters Sertoli cell oestradiol production, enzyme activity and secretory behaviour (Filippini et al. 1994; Meroni et al. 1998; Rossato et al. 2001; Gelain et al. 2005). Despite this putative role in testicular physiology, identification and a precise functional description of involved purinoceptor subtype(s) are lacking.

G-protein-coupled P2Y receptors form two distinct subgroups that either activate phospholipase C (PLC) via G_αq/G_α11 (P2Y1, P2Y2, P2Y4, P2Y6 and P2Y11) or couple to G_αi/o (P2Y12, P2Y13 and P2Y14). P2X receptors form homo- or heterotrimers that function as cationic ligand-gated ion channels and display a distinctive two transmembrane domain topology (Kawate et al. 2009). So far, six homomeric and several heteromeric channels have been described (Jarvis & Khakh, 2009).

In vivo, the polarized morphology and cytoplasmic compartmentalization of Sertoli cells pose a unique challenge to orchestrate discrete Ca²⁺-sensitive responses. Their speed, reliability and specificity not only depend on Ca²⁺ influx and release mechanisms, but also cytoplasmic buffer systems that limit Ca²⁺ diffusion. Bulk [Ca²⁺]_c is maintained at resting levels (∼100 nm) by the coordinated activity of the plasma membrane Ca²⁺ ATPase, the Na⁺/Ca²⁺ exchanger (NCX) and the sarco/endoplasmic reticulum Ca²⁺ ATPase (SERCA). In addition, mitochondria accumulate Ca²⁺ via the mitochondrial Ca²⁺ uniporter (mCU) (Rizzuto et al. 1998). Ca²⁺ sequestration in mitochondria serves a dual role by (a) meeting local energy demands via activation of electron transport chain (ETC) enzymes (Denton & McCormack, 1980) and (b) shaping the spatiotemporal dynamics of cytosolic Ca²⁺ signals. Therefore, the mitochondrial network is an essential determinant of both global and local patterns of Ca²⁺ mobilization and, thus, a key factor in numerous signalling pathways (Budd & Nicholls, 1996; Tang & Zucker, 1997).

At rest, the mitochondrial Ca²⁺ concentration ([Ca²⁺]_m) is close to [Ca²⁺]_c. During [Ca²⁺]_c elevations, however, the negative inner mitochondrial membrane (IMM) potential (ΔΨ_m; approximately −180 mV) drives massive Ca²⁺ influx into the mitochondrial matrix via the mCU (Kirichok et al. 2004). Ca²⁺ sequestration can be abolished pharmacologically by agents that cause a collapse of the IMM H⁺ gradient and dissipation of ΔΨ_m (Celsi et al. 2009). Undisturbed, however, the Ca²⁺ storage capacity of mitochondria even exceeds ER buffering (Feissner et al. 2009). Ca²⁺ clearance from the matrix is predominantly established by the mitochondrial Na⁺/Ca²⁺ exchanger (NCX_mito; Palty et al. 2010) and, to a lesser extent, by a H⁺/Ca²⁺ exchanger.

The physiological relevance of mitochondrial Ca²⁺ uptake has been established in a variety of cell types by expressing genetically targeted Ca²⁺-sensitive photoproteins in mitochondria (Rizzuto et al. 1992; Brini et al. 1994). Compared to most fluorescent probes, genetically targeted Ca²⁺-sensitive photoproteins exhibit a number of advantages for organelle-specific quantitative [Ca²⁺]_m recordings such as a broad dynamic range, steep Ca²⁺ dependence, low Ca²⁺ buffering capacity, irradiation-free imaging and pH insensitivity in the physiological range (Pozzan & Rudolf, 2009). In conventional assays, however, the poor photon emission of bioluminescent Ca²⁺ sensors renders single-cell [Ca²⁺]_m imaging at high spatio-temporal resolution virtually impossible (Fedrizzi & Brini, 2010). Here, we establish a live-cell bioluminescence [Ca²⁺]_m imaging approach that overcomes these limitations without compromising its numerous advantages. Using genetically engineered PhotoTopo mice that ubiquitously express the optically improved photoprotein c-Photina within the mitochondrial matrix (Cainarca et al. 2010) and a dedicated bioluminescence microscope, we record (sub)cellular mitochondrial Ca²⁺ signals at millisecond resolution. Combining both [Ca²⁺]_m and [Ca²⁺]_c imaging methods with whole-cell patch-clamp recordings, we describe here the key components of a Ca²⁺ mobilization network that underlies purinergic signalling in primary mouse Sertoli cells.

Methods

Chemicals

For tissue preparation, electrophysiological recordings and calibration of [Ca²⁺]_c values from ratiometric fura-2 imaging experiments, the following solutions were used: (a) HEPES-buffered extracellular solution (S₁) containing (in mm): 145 NaCl, 5 KCl, 1 CaCl₂, 0.5 MgCl₂, 10 HEPES; pH 7.3 (adjusted with NaOH); osmolarity, 300 mosmol l⁻¹ (adjusted with glucose); (b) Tyrode buffer solution (S₂) containing (in mm): 130 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, 5 NaHCO₃, 20 HEPES; pH 7.4; osmolarity, 300 mosmol l⁻¹; (c) HEPES-buffered pipette solution (S₃) containing (in mm): 120 potassium gluconate, 23 KCl, 2 KOH, 1 EGTA, 10 HEPES, 1 MgATP, 0.5 NaGTP; pH 7.1 (adjusted with KOH); osmolarity, 290 mosmol l⁻¹; (d) [Ca²⁺]_zero solution containing (in mm): 105 NaCl, 5 KCl, 0.5 MgCl₂, 10 HEPES, 20 EGTA; pH 7.3 (NaOH); osmolarity, 300 mosmol l⁻¹ (glucose); (e) Saturating Ca²⁺ solution containing (in mm): 125 NaCl, 5 KCl, 10 CaCl₂, 0.5 MgCl₂, 10 HEPES; pH 7.3 (NaOH); osmolarity, 300 mosmol l⁻¹ (glucose). Free Ca²⁺ concentrations were calculated using WEBMAXC STANDARD (available at http://www.stanford.edu/~cpatton/maxc.html). If not stated otherwise, all chemicals were purchased from Sigma (Schnelldorf, Germany). For compounds soluble in DMSO, final concentrations were ≤0.5%.

Animals

All animal procedures were in compliance with the European Union legislation on the protection of animals used for experimental purposes (Directive 86/609/EEC) and recommendations put forward by the Federation of European Laboratory Animal Science Associations (FELASA). PhotoTopo mice were housed in groups of both sexes at room temperature on a 12:12 h light–dark cycle with food and water available ad libitum. If not otherwise stated, experiments used 7-day-old males.

Sertoli cell culture

We recently described the generation of PhotoTopo mice (Cainarca et al. 2010). Briefly, c-Photina – a photoprotein derived from Clytin (Clytia gregaria) and modified by random mutagenesis – was optimized for mammalian codon usage and fused to a mitochondrial tag (human cytochrome C oxidase, subunit VIII). Stable expression of mito c-Photina in CHO cells allowed for functional characterization of construct stability. Transgenic PhotoTopo mice were then generated from electroporated mouse embryonic stem cells, blastocyst injection, germ line transmission, crossing of chimeric males and C57BL/6 females, and crossing of heterozygous progeny to obtain a homozygous population that showed no phenotypic abnormalities. In PhotoTopo mice, stable tissue-wide expression of mito c-Photina was confirmed by TaqMan qPCR analysis.

Male PhotoTopo mice (postnatal day 7) were killed by CO₂ asphyxiation and decapitation using sharp surgical scissors. After testis isolation and removal of the tunica albuginea, seminiferous tubules were digested in 1 mg ml⁻¹ collagenase in MEM (Invitrogen, Darmstadt, Germany) (8 min; 37°C). The reaction was stopped by addition of serum-containing DMEM (Invitrogen) and subsequent centrifugation (8 min; 400 g). The supernatant was discarded and the pellet was resuspended in trypsin/EDTA (PAA Laboratories, Pasching, Germany). After incubation (5 min; 37°C), the cell suspension was centrifuged (10 min; 400 g) and the pellet was resuspended in DMEM (10% fetal calf serum; 100 U ml⁻¹ penicillin G; 100 μg ml⁻¹ streptomycin; PAA Laboratories). Cells were plated at a density of 5 × 10⁴ cells per 35 mm dish. Half of the medium was changed every other day. For 3 days in vitro (DIV), cells were kept at 37°C in 5% CO₂ in a humidified incubator. On day 4, cells were transferred to a humidified incubator providing a 34°C−5% CO₂ atmosphere. Experiments were performed on DIV 7. Sertoli cell identity/culture purity was routinely confirmed by immunochemical and morphological characteristics (see Results). Minor contamination by peritubular myoid cells was distinguished morphologically.

Sertoli cell transfection

For small interfering RNAs (siRNAs) experiments, Sertoli cells were transfected on DIV 5 using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions. Before transfection, the culture medium was replaced by antibiotic-free medium. siRNA (Applied Biosystems/Ambion, Darmstadt, Germany) was either co-transfected with a pIRES2-EGFP vector (Clontech, Mountain View, CA, USA) or with BLOCK-iT fluorescent oligo (FITC labelled) (Invitrogen). Opti-MEM I reduced serum medium (500 μl; Invitrogen) containing 6 μl Lipofectamine 2000, 167.5 ng siRNA (25 nm) and either 744 ng EGFP-plasmid or 0.625 μl BLOCK-iT fluorescent oligo (25 nm) was added to each culture dish. Cells were used for experiments 48 h after transfection. Transfected cells were identified by EGFP or FITC fluorescence. We used Silencer Select Pre-designed siRNA no. s71197 or s71196, respectively, to down-regulate P2Y2 protein expression. As negative control we used Silencer Select negative control no. 1 siRNA.

Single cell mitochondrial Ca²⁺ imaging

Sertoli cells were incubated in S₁ containing 10 μm coelenterazine (3 h; 37°C). Excessive substrate was not removed before the experiment. Cells were then transferred to the stage of a self-contained Olympus Luminoview LV200 luminescence microscope and constantly perfused (S₁). Depending on the experiment, cells were visualized at different magnifications (20× 0.4 NA Plan Apo objective, 60× 1.35 NA UPlanSApo objective and 100× 1.4 NA UPlanSApo objective) and images were captured by a cooled Hamamatsu ImageEM C9100-13 EM-CCD camera at 1 Hz frame rate. Stimuli were applied using a custom-made pressure-driven multi-barrel perfusion pencil that allows for instantaneous focal solution change (see Supplemental Fig. S2). Time-lapse experiments were analysed offline using Cell^M Live-Cell Imaging Software (Olympus). Fields of view were randomly chosen and 20 to 50 cells were analysed per experiment. Relative luminescence unit (RLU) values were calculated for each ROI/cell and processed as a function of time.

Mitochondrial bioluminescence Ca²⁺ imaging in Sertoli cell populations (FLIPR assay)

For population bioluminescence Ca²⁺ imaging, Sertoli cells were seeded in quadruplicates in 384-well white plates (1000 cells well⁻¹) and incubated for 72 h. Prior to tests, samples were incubated in S₂ (20 μl well⁻¹; 4 h) containing 10 μm coelenterazine (Pharma Tech International). Experiments were performed using a FLIPR^TETRA instrument (Molecular Devices, Sunnyvale, CA, USA) equipped with an enhanced ICCD camera. In each experiment, identical readout and injection parameters were selected (exposure time, 0.9 s; excitation intensity, 100%; gate, 100%; injection, 20 μl s⁻¹, up to 15 μl height). Stimulus-dependent light release was measured at 1 Hz for 65 s and expressed in RLU over time. Two consecutive injections ((1) 10 μl well⁻¹, 4× compound master plate; (2) 20 μl well⁻¹, 3× activator master plate) were performed at 5 min inter-stimulus intervals (ISI).

Single cell cytosolic fluorescence Ca²⁺ imaging

Sertoli cells, grown on 35 mm glass-bottom dishes for 7 days, were loaded with fura-2 AM (3 μm; 30 min; 37°C; S₁). Dye-loaded cells were washed in S₁ and transferred to the stage of an inverted microscope (Leica DMI4000B, Leica Microsystems, Wetzlar, Germany) equipped for ratiometric live-cell imaging with a 150 W xenon arc lamp, a motorized fast-change filter wheel illumination system for multi-wavelength excitation, a 12-bit 1376 × 1032 pixel monochrome CCD camera (DFC 360FX, Leica), and Leica LAS AF7000 imaging software. Ten to thirty cells in randomly selected fields of view were viewed at 20× magnification and illuminated sequentially at 340 nm and 380 nm (cycle time 1000 ms). The average pixel intensity within user-selected ROI was digitized and stored on a PC. Ca²⁺-dependent fluorescence signals at 510 nm were calculated as the F₃₄₀/F₃₈₀ intensity ratio. In experiments designed to calculate Ca²⁺ concentrations rather than relative changes, integrated fluorescence signals from individual ROIs were background subtracted and [Ca²⁺]_c was calculated according to Grynkiewicz et al. (1985) using the equation [Ca²⁺] = K_d× (F₀/F_s) ×[(R–R_min)/(R_max–R)], where R is the experimentally derived fluorescence intensity ratio (F₃₄₀/F₃₈₀), R_min and R_max are ratios measured using [Ca²⁺]_zero and saturating Ca²⁺ calibration solutions, respectively, K_d is the fura-2 dissociation constant (224 nm), and F₀ and F_s are proportionality coefficients for free and Ca²⁺-bound fura-2, respectively (measured at 380 nm excitation). R_min and R_max were determined in intact cells by applying 5 μm ionomycin in [Ca²⁺]_zero (20 mm EGTA) and saturating Ca²⁺ (10 mm Ca²⁺). Values for R_min, R_max and F₀/F_s were 0.038, 1.664 and 6.35, respectively. Stimuli were applied using a custom-made pressure-driven multi-barrel perfusion system that allows for almost instantaneous solution switch and focal application via an 8-to-1 manifold with microlitre dead-volume as assessed by employing different fluorescent dyes (data not shown; see Fig. S2).

RNA isolation, RT-PCR and quantitative PCR (qPCR)

Total RNA from cultured Sertoli cells and tissues (testis, brain/spinal cord) was isolated according to the instructions of the RNeasy Mini Kit (Qiagen, Hilden, Germany). cDNA was transcribed using the RevertAid™ H Minus M-MuLV Reverse Transcriptase (Fermentas, St. Leon-Rot, Germany) according to the manufacturer's instructions. Controls in which the reverse transcription step was omitted were routinely performed. PCR amplification was performed during 30 thermal cycles (94°C, 20 s; 54–56°C, 20 s; 72°C, 20 s). PCR products were visualized on an agarose gel via ethidium bromide staining to confirm the anticipated size of the product. The specific primer pairs used for PCR amplification are given in Table 1.

Table 1.

Primer pairs used for PCR amplification

Target	Accession no.	Forward primer 5′ to 3′	Reverse primer 5′ to 3′	Size
P2X2	NM_153400	TCCCTCCCCCACCTAGTCAC	CACCACCTGCTCAGTCAGAGC	149 bp
P2X3	NM_145526	CTGCCTAACCTCACCGACAAG	AATACCCAGAACGCCACCC	150 bp
P2X4	NM_011026	CCCTTTGCCTGCCCAGATAT	CCGTACGCCTTGGTGAGTGT	145bp
P2X5	NM_033321	GGATGCCAATGTTGAGGTTGA	TCCTGACGAACCCTCTCCAGT	81 bp
P2X6	NM_011028	CCCAGAGCATCCTTCTGTTCC	GGCACCAGCTCCAGATCTCA	152 bp
P2X7	NM_011027	GCACGAATTATGGCACCGTC	CCCCACCCTCTGTGACATTCT	171 bp
P2Y1	NM_008772	CGACAGGGTTTATGCCACTT	TCGTGTCTCCATTCTGCTTG	218 bp
P2Y2	NM_008773	CGTGCTCTACTTCGTCACCA	GACCTCCTGTGGTCCCATAA	206 bp
P2Y4	NM_020621	ACTGGCTTCTGCAAGTTCGT	AGGCAGCCAGCTACTACCAA	188 bp
P2Y6	NM_183168	CATTAGCTTCCAGCGCTACC	GCTCAGGTCGTAGCACACAG	187 bp
P2Y12	NM_027571	CATTGCTGTACACCGTCCTG	AACTTGGCACACCAAGGTTC	212 bp
P2Y13	NM_028808	CACTCTGGTGGCAGACTTGA	GTTTTCCTGAACGGCATGAT	204 bp
P2Y14	NM_133200	CAGTGCATGGAGCTCAAAAA	GCACAAAGCAGACGACAAAA	232 bp
β actin	NM_007393	GTCTTCCCCTCCATCGTGG	TGGATGCCACAGGATTCC	736 bp

With the exception of P2ry11 – a gene not found in rodents – P2ry coding sequences are intron-free (Abbracchio et al. 2006). To further test for genomic DNA contamination, all cDNA samples were additionally analysed using intron-spanning actin-specific primer pairs.

For qPCR, we used the QuantiFast SYBR Green PCR Kit (Qiagen, Hilden, Germany) and an iQ5 thermal cycler (BIO-RAD, Hercules) according to the manufacturer's instructions. α1A-tubulin was used to normalize the amount of P2Y2 mRNA in the different cDNA populations tested. Relative mRNA levels were then calculated using the threshold cycle (Ct) value and the comparative Ct method. The primer pairs used for the qPCR are given in Table 2.

Table 2.

Primer pairs for qPCR

Target	Accession no.	Forward primer 5′ to 3′	Reverse primer 5′ to 3′	Size
P2Y2	NM_008773	CGTGCTCTACTTCGTCACCA	CATGACGGAGCTGTAAGCCAC	106 bp
α1A-tubulin	NM_011653.2	TCCCAAAGATGTCAATGCTG	CACAGTGGGAGGCTGGTAAT	115 bp

Immunohistochemistry

Testicular cryosections

Testes were fixed in 4% paraformaldehyde (PFA) in Ca²⁺−Mg²⁺-free phosphate-buffered saline (PBS^−/− (3 h; 4°C)) and subsequently cryoprotected in PBS^−/− containing 30% sucrose (≥24 h; 4°C). The dehydrated testis was embedded in Tissue Freezing Medium (Leica Microsystems) and sectioned at 15 μm on a cryostat (Leica Microsystems). Mounted sections were again fixed in ice-cold 4% PFA (15 min), washed several times in PBS^−/−, and then incubated in PBS^−/− (1 h, room temperature (RT)) containing 1% gelatin from cold water fish skin and 0.1% Triton X-100 (blocking solution).

Cultured Sertoli cells

For immunostainings, Sertoli cells were grown on coverslips in 24-well plates. After removal of culture medium, cells were washed (3×; PBS^−/−), fixed in ice-cold 4% PFA in PBS^−/− (15 min; RT), washed again (3×; PBS^−/−), and incubated in blocking solution (1 h, RT).

Primary antibodies (anti-P2X2 (Alomone Labs, Jerusalem, Israel), anti-DAZL (Abcam, Cambridge, UK), anti-Vimentin (Santa Cruz Biotechnology Inc., Santa Cruz, USA)) were diluted according to the manufacturer's recommendations. Sertoli cells/testicular cryosections were incubated with primary antibodies overnight (4°C) in a humidified chamber. After washing in PBS^−/− (3×), cells/sections were incubated in PBS^−/− (45 min; RT) containing Alexa Fluor-conjugated secondary antibodies (Molecular Probes, Karlsruhe, Germany). Excess antibodies were removed by washing. Fluorescent images were taken using an upright fixed stage scanning confocal microscope (Leica TCS SP5 DM6000 CFS, Leica Microsystems) equipped with a 20× 1.0 NA water immersion objective (HCX APO L, Leica Microsystems). To control for non-specific staining, experiments in which the primary antibodies were omitted were performed in parallel with each procedure. Digital images were uniformly adjusted for brightness and contrast using Adobe Photoshop CS3 (Adobe Systems, San Jose, CA, USA).

Electrophysiology

For electrophysiological recordings, Sertoli cells were transferred to the stage of an inverse video-microscope (DMI 4000B, Leica Microsystems). Cells were continuously superfused with solution S₁ (∼3 ml min⁻¹; gravity flow) at room temperature. Patch pipettes with a resistance of ∼5 MΩ were pulled from borosilicate glass capillaries with filament and fire-polished ends (1.50 mm OD/0.86 mm ID; Science Products, Hofheim, Germany) on a PC-10 vertical two-step micropipette puller (Narishige Instruments, Tokyo, Japan) and fire-polished using a MF-830 microforge (Narishige Instruments). Recording pipettes were filled with S₃ solution. An EPC-10 amplifier controlled by Patchmaster 2.50 software (HEKA Elektronik, Lambrecht/Pfalz, Germany) was used for data acquisition. Both pipette (C_fast) and cell membrane capacitance (C_slow) were monitored and automatically compensated throughout the experiment. Measured C_slow values served as an approximation of the cell surface area for normalization of current amplitudes (i.e. current density). Sertoli cells with unstable C_slow values were not considered for further analysis. Theoretical liquid junction potentials were calculated using JPCalcW software and automatically corrected online. In voltage-clamp experiments, leak currents were subtracted using a P/−4 procedure. Signals were low-pass filtered (analog 3- and 4-pole Bessel filters in series) with an effective corner frequency (−3 dB) automatically adjusted to a fifth to a third of the sampling rate (5–20 kHz, depending on protocol). Focal application of different stimuli and/or pharmacological agents was achieved by a software-controlled pressure-driven valve bank connected to a ‘perfusion pencil’. Between recordings, cells were kept at a holding potential (V_hold) of −50 mV. Individual voltage step protocols are described in the Results section.

Data analysis

All data were obtained from experiments performed on at least 2 days. Individual numbers of cells/experiments (n) are denoted in figure legends. If not stated otherwise, results are presented as means ± SEM and statistical analyses were performed using paired or unpaired t tests (as dictated by data distribution and experimental design).

FLIPR assay results were analysed using an implemented version of Spotfire Decision Site 9.1. Analysis and curve fitting was performed using the Hill equation. For each plate, the mean and standard deviation of quadruplicate well response values were calculated and used for curve fitting.

Ca²⁺ imaging and electrophysiological data were analysed offline using FitMaster 2.20 (HEKA Elektronik), IGOR Pro 6.03A (WaveMetrics, Lake Oswego, OR, USA) and Excel (Microsoft, Seattle, WA, USA) software. Ca²⁺ decay time constants (τ) were calculated by fitting individual traces to monoexponential functions I_(t) = I₁ (exp(−t/τ))+I₀. Dose−response curves were fitted by the Hill equation. The full duration at half-maximum (FDHM) was defined as the time period between the half-maximum of the Ca²⁺ rise and the half-maximum of the Ca²⁺ decay.

Results

In spermatogenesis, endocrine regulation pathways merge on Sertoli cell Ca²⁺ mobilization (Gorczynska-Fjalling, 2004), a signalling mechanism that is modulated by paracrine/autocrine purinergic communication (Filippini et al. 1994; Foresta et al. 1995; Rudge et al. 1995; Lalevee et al. 1999; Rossato et al. 2001; Ko et al. 2003). We therefore investigated the physiological mechanisms that underlie ATP-dependent Ca²⁺ dynamics in immature mouse Sertoli cells.

Parallel metabotropic and ionotropic P2 receptor-dependent Ca²⁺ mobilization pathways

We first recorded ATP-induced cytosolic Ca²⁺ signals in primary Sertoli cell cultures from pre-pubertal mice (P7). At this developmental stage, the seminiferous epithelia consist exclusively of immature highly proliferative Sertoli cells and type A spermatogonia (Bellve et al. 1977) (Fig. S1). When grown in a supplement-free medium, we obtained almost pure monolayer cultures of adherent Sertoli cells that displayed a flat and spread morphology (Fig. 1A). Analogous culture conditions were previously shown to provide the most reproducible endocrine Sertoli cell responses (Griswold & McLean, 2006). Cellular identity was confirmed by immunocytochemical labelling of vimentin, a cytoskeletal marker of Sertoli cells (Franke et al. 1979), and the characteristic abundance of cytoplasmic lipid droplets (Xiong et al. 2009) that were discernable from both DIC microscopy and specific labelling with a lipophilic fluorescent dye (Nile Red; Fig. 1A).

Figure 1. Ca²⁺ mobilization by P2X and P2Y purinoceptors.

A, representative DIC and confocal images of cultured Sertoli cell monolayers at 7 days in vitro (DIV). Cells are immunopositive for vimentin (green) and show characteristic lipid droplets (stained with Nile Red; red). B, [Ca²⁺]_c transients in response to ATP (100 μm; 10 s) are monitored by ratiometric fluorescence imaging in fura-2 AM-loaded Sertoli cells (grey scale image). Original trace depicts the integrated fluorescence ratio F₃₄₀/F₃₈₀ of a representative cell in a user-defined region of interest (ROI) as a function of time. Pseudocolour single frame images illustrate relative [Ca²⁺]_c at time points 1 – 4 (rainbow 256 colourmap; blue, low Ca²⁺; red, high Ca²⁺). C, representative traces (left; F₃₄₀/F₃₈₀versus time) of [Ca²⁺]_c transients recorded from a Sertoli cell challenged with increasing ATP concentrations (0.1–10 μm; 10 s; arrowheads) in presence (black) and absence (red) of external Ca²⁺. Sigmoid dose–response curves (right) were calculated from averaged data points (mean ± SEM) using the Hill equation (n = 344 (control, black); n = 370 (Ca²⁺-free, red)). D, representative [Ca²⁺]_c traces (F₃₄₀/F₃₈₀versus time) of ATP (100 μm; 10 s) responses in single Sertoli cells under control and pharmacologically modified conditions (n = 33 (‘Ca²⁺-free’); n = 163 (U73122); n = 106 (NMDG⁺); n = 154 (thapsigargin)). Treatment duration is indicated by red horizontal bars. U73343, an inactive analogue of the PLC inhibitor U73122, served as a control for drug specificity (red trace).

For recording of [Ca²⁺]_c, Sertoli cells were loaded with fura-2 AM and briefly (10 s) exposed to ATP at a concentration (100 μm) saturating for P2 purinoceptors (except P2X7; Burnstock, 2008). To achieve rapid and simultaneous solution exchange in the entire field of view, we used a custom-made perfusion system (Fig. S2). In most cells (96.6 ± 2.1%), ATP stimulation triggered an immediate (rise time = 6.5 ± 0.3 s; rising speed = 0.04 ratio units s⁻¹) and robust [Ca²⁺]_c signal that rapidly decayed (time constant (τ) = 8.1 ± 0.3 s) after stimulus cessation (n = 81; Fig. 1B). In both control (extracellular Ca²⁺ concentration [Ca²⁺]_e = 1 mm; n = 344) and ‘Ca²⁺-free’ medium ([Ca²⁺]_e = 30 nm; n = 370), ATP induced dose-dependent responses with a shared activation threshold of <100 nm (Fig. 1C; Movie S1). In situ calibration of indicator fluorescence in the presence of both [Ca²⁺]_zero and saturating [Ca²⁺]_c enables the conversion of fluorescence ratios into Ca²⁺ concentrations (Fig. S3; Grynkiewicz et al. 1985). Under control conditions, average [Ca²⁺]_c is 144 ± 10 nm, whereas a slightly reduced level is detected in ‘Ca²⁺-free’ medium (86 ± 7 nm). Saturating ATP concentrations caused average [Ca²⁺]_c elevations up to 636 ± 36 nm (control) and 380 ± 56 nm (‘Ca²⁺ free’), respectively. The linear dynamic range of the activation curve (Fig. 1C) spanned ∼2 log units under control conditions (EC₅₀ = 402 nm) and Ca²⁺ signals saturated at ∼10 μm[ATP]. In absence of extracellular Ca²⁺, however, the curve spanned only ∼1 log unit (EC₅₀ = 195 nm; saturation reached at ∼1 μm[ATP]). These data confirm that Sertoli cells express two functionally distinct populations of P2 purinoceptors (Foresta et al. 1995; Ko et al. 2003), i.e. metabotropic P2Y and ionotropic P2X receptors that show a relatively high versus low ATP affinity, respectively. At saturating concentrations (100 μm), ATP triggered pronounced, though significantly diminished, [Ca²⁺]_c transients when either P2X or P2Y signalling was impaired (Fig. 1D; Fig. S4A and C). Moreover, [Ca²⁺]_c signals were reduced by exchange of external cations for NMDG⁺. P2Y receptor signalling was disrupted by inhibition of either PLC-dependent phosphoinositide (PI) turnover (U73122, 10 μm) or Ca²⁺ release from the endoplasmic reticulum (ER; thapsigargin, 10 μm). Both treatments also significantly affected the kinetics of the cytosolic Ca²⁺ signal decay time constant (Fig. S4D). However, when the high-affinity P2 receptor population was selectively stimulated by reduced ATP concentrations (350 nm), PLC inhibition (U73122) or ER depletion (thapsigargin) essentially abolished Ca²⁺ signals (Fig. S4B). By contrast, ATP (350 nm) responses were unchanged in the absence of external Ca²⁺ or any permeable cations (Fig. S4E). Together, these data suggest that both high-affinity P2Y receptors, which signal via PI breakdown and Ca²⁺ release from storage organelles, and low-affinity P2X receptors mediate purinergic communication in immature mouse Sertoli cells.

To discriminate between P2Y and P2X receptors and specifically characterize the functional properties of Sertoli cell P2X receptors, we next recorded ATP-mediated whole-cell membrane currents (I_ATP) using the patch-clamp technique. Brief (5 s) stimulation with ATP (100 μm) triggered both robust inward currents and substantial membrane depolarization (Fig. 2A). I_ATP amplitudes increased dose-dependently over a broad concentration range (Fig. 2B). In a normalized dose–response plot (Fig. 2C), an activation threshold of ∼1 μm, an EC₅₀ of 12.7 μm, and saturating concentrations of ≥100 μm became apparent (n = 25).

Figure 2. Non-overlapping dynamic ranges of Sertoli cell P2Y and P2X receptors.

A, representative original patch-clamp recordings of (i) a typical ATP-induced inward current (top; voltage-clamp configuration; V_hold = −50 mV) and (ii) the corresponding ATP-dependent membrane depolarization in the same cell (bottom; current-clamp mode). Sertoli cells displayed an average membrane potential of −58.3 ± 3.8 mV (n = 15 cells). ATP (100 μm; 5 s) evoked an average current amplitude of 675 ± 150 pA (n = 25 cells). B and C, when challenged with increasing ATP concentrations, inward currents develop dose-dependently. The threshold concentration for activation is ∼1 μm, half-maximal activation (EC₅₀) is observed at 12.7 ± 1.8 μm, and stable saturated current levels are induced by ATP concentrations of ≥100 μm. A sigmoidal dose–response curve was calculated using the Hill equation. Individual data points show means ± SEM (n = 25). D, simultaneous measurements of [Ca²⁺]_c transients and I_ATP. Original traces illustrate cytosolic Ca²⁺ signals (red; F₃₄₀/F₃₈₀versus time) and inward currents (black; V_hold = −50 mV) induced by brief repetitive ATP stimulation (5 s). Inset shows repetitive stimulation with a constant ATP concentration (100 μm; n = 11; top), a phase contrast (bottom left) and fluorescence image (bottom right; 380 nm excitation) of a Sertoli cell diffusion-loaded with fura-2 (10 μm). E, P2X- (black) and P2Y- (red) specific dose−response curves. [Ca²⁺]_c was recorded in absence of external Ca²⁺. I_ATP amplitudes are plotted as current densities (pA pF⁻¹). Sigmoidal dose−response curves were calculated using the Hill equation. Individual data points depict means ± SEM (n = 25 cells (P2X); n = 370 cells (P2Y)).

By simultaneously performing patch-clamp and Ca²⁺ imaging recordings from cells diffusion-loaded with fura-2 via the patch pipette (Fig. 2D, inset), we confirmed that the linear dynamic ranges of P2Y and P2X receptors in Sertoli cells cover separated concentration ranges. P2Y receptor activation effectively mobilizes Ca²⁺ when exposed to nanomolar ATP concentrations, whereas micromolar ATP concentrations are required to recruit P2X receptors (Fig. 2E). At the latter concentrations, Sertoli cell P2Y receptors are saturated (n = 25 cells (P2X); n = 370 cells (P2Y)).

Identification of P2 purinoceptor subunits

Subunit identification as well as a detailed functional description of Sertoli cell P2X receptors is still lacking. So far, studies have focused on the adult rat testis and contradictory purinoceptor expression patterns have been proposed (MacKenzie et al. 1999; Glass et al. 2001; Rossato et al. 2001; Ko et al. 2003). Therefore, we investigated the biophysical/pharmacological characteristics of P2X receptors in whole-cell voltage-clamp recordings.

As P2X receptors are discriminated based on rapid versus slow desensitization (North, 2002), we first analysed I_ATP activation and inactivation kinetics (Fig. 3). When current amplitudes are plotted as a function of stimulus duration, I_ATP amplitudes reach a maximum level at exposure durations ≥1 s (Fig. 3A and B). Peak currents fully developed within a few seconds (average rise time t_10−90% = 1.7 ± 0.1 s). When cells were continuously challenged with ATP (100 μm; 2 min), currents slowly declined (τ = 37.9 ± 4.8 s). Even after two consecutive 2 min stimulations, substantial residual current was recorded (Fig. 3C). Moreover, in double-pulse experiments, inter-stimulus intervals (ISI) could be reduced to 2.8 s without receptor desensitization (Fig. 3D). These data show that immature mouse Sertoli cells express at least one member of the slowly desensitizing group of P2X receptors.

Figure 3. Activation and desensitization characteristics of Sertoli cell P2X receptors.

A, original voltage-clamp recording of I_ATP as a function of stimulus duration. When challenged with ATP (100 μm) for 100 ms, a robust inward current is observed. With prolonged stimulus exposure (300 ms; 1 s), I_ATP amplitude increases. B, plot of average I_ATP current density (mean ± SEM; n = 13 cells) versus exposure time. A short stimulus (100 ms) triggers a mean current of 2.7 ± 0.9 pA pF⁻¹. I_ATP amplitude increases to 4.8 ± 1.4 pA pF⁻¹ in response to 300 ms pulses and saturates at exposure times ≥1 s (6.2 ± 1.6 pA pF⁻¹). C, representative original I_ATP recording during two successive long-term ATP stimulations (100 μm; 2 min; ISI ∼1 s). During the recording period, slow and incomplete desensitization was observed in 4/4 cells. D, I_ATP amplitudes were unaffected by inter-stimulus intervals between 180 s (top) and 2.8 s (bottom; 5 s stimulation; n = 5 cells).

Next, we investigated the I_ATP current−voltage relationship (Fig. 4A and B). Brief ATP exposure (100 μm; 5 s) at different holding potentials (V_hold = −80 mV to +60 mV) revealed a reversal potential of ∼0 mV and strong inward rectification (n = 12). In addition, I_ATP was modulated by external protons and trace metals (Fig. 4C and D). At negative holding potential (V_hold = −50 mV), acidic pH (6.3; n = 5) strongly enhanced current amplitudes, whereas alkaline conditions (pH 8.3; n = 4) greatly reduced I_ATP. Copper and zinc ions significantly increased I_ATP amplitudes (n = 10). Finally, α,β-methylene-ATP (300 μm; n = 8), a selective P2X1 and P2X3 agonist (North, 2002), failed to elicit responses (Fig. 4E).

Figure 4. The biophysical fingerprint of ATP-induced currents is consistent with P2X2 expression.

A, representative ATP-dependent whole-cell current traces recorded at different membrane potentials (inset: pulse protocol). Voltage-dependent background currents were subtracted offline. B, ATP current density (mean ± SEM; n = 12) as a function of membrane potential. The I−V curve displays strong inward rectification. Current polarity reversed at ∼0 mV (E_ATP = 6.5 mV), suggesting a non-selective cation current (gluconate-based pipette solution (E_Cl = −47 mV)). C, brief ATP exposure (5 s; 10 μm; V_hold = −50 mV) triggers robust inward currents under control conditions (black). I_ATP amplitudes are either potentiated or diminished in presence of Cu²⁺, Zn²⁺, acidic and alkaline pH, respectively (red). D, bar chart illustrating I_ATP amplitudes (mean ± SEM; normalized to control). Currents are significantly (P < 0.05) potentiated by Cu²⁺ (134 ± 12%; n = 10), Zn²⁺ (139 ± 8 %; n = 10) and acidic pH (175 ± 16 %; n = 5). Alkaline pH causes strong inhibition (8 ± 2 %; n = 4). E, representative recording showing the insensitivity to α,β-methylene-ATP (5 s; 300 μm; V_hold = –50 mV). Subsequent ATP exposure (5 s; 10 μm) triggers substantial inward current (positive control; 8/8 cells).

Together, these data are most consistent with functional expression of homomeric P2X2 receptors (Ding & Sachs, 1999; Khakh & North, 2006). To address this directly, we designed specific intron-spanning primers for the different P2rx genes and performed RT-PCR experiments on total RNA extracted from primary mouse Sertoli cell cultures, immature testis (P7), adult testis, and brain–spinal cord tissue (positive control; Burnstock, 2008). P2X2-encoding cDNA was amplified from Sertoli cells, immature and adult testes (Fig. 5A). A similar expression pattern was observed for P2rx7. Transcripts coding for P2X4 were detected in whole testis samples (P7), but not in Sertoli cells, indicating P2rx4 gene expression by other testicular cell types. None of the remaining P2rx genes appears to be expressed in mouse testes. On the protein level, P2X2 immunofluorescence was detected in cultured Sertoli cells (Fig. 5C) as well as in the seminiferous epithelia of immature and adult testes (data not shown). P2X2 immunoreactivity in primary Sertoli cell culture was largely confined to perinuclear regions (Fig. 5C). P2X7 immunopositive cells showed a more diffuse staining (data not shown). In the adult testis, however, P2X7 antibody staining was almost exclusively observed in extratubular cells (data not shown). In concert with the described biophysical hallmarks (Figs 2–4), these results suggest that P2X2 receptor subunits predominantly, if not exclusively, mediate ionotropic ATP signalling in immature mouse Sertoli cells.

Figure 5. P2X/P2Y receptor expression profiling.

A, RT-PCR reveals transcripts for P2rx2 and P2rx7 in cultured Sertoli cells, P7 and adult testis. P2X4-encoding cDNA was amplified from whole testis samples (P7), but not Sertoli cells. Primer pairs were designed to generate amplificates of 100–250 bp, respectively. B, P2ry2, P2ry6 and P2ry14 transcripts were amplified from Sertoli cell cDNA. In addition, cDNA coding for P2Y1, P2Y2 and P2Y12 was detected in immature and adult testicular samples. By contrast, P2ry6 and P2ry14 transcripts were found in P7, but not in adult testis. Expected PCR products were 150–250 bp, respectively. cDNA from whole brain/spinal cord served as positive control (A and B). C, immunocytochemical expression analysis of P2X2 receptors in Sertoli cell cultures. Confocal DIC and fluorescence images show representative cells. Primary anti-P2X2 antibodies are fluorescently labelled (green) by secondary antibodies conjugated to Alexa488. In the merged image, a predominantly perinuclear P2X2 staining pattern becomes apparent.

To obtain a complete P2 purinoceptor expression profile, we additionally analysed P2Y receptor transcription (Fig 5B). In Sertoli cell cultures, bands of the expected size were detected for P2ry2 and P2ry14 transcripts. A considerably weaker band was additionally found for P2ry6 (Fig. 5B). For P2ry2, mRNA was also amplified from immature and (to a lesser extend) adult testes. P2ry6 and P2ry14 transcripts − encoding for ATP-insensitive receptors (Ralevic & Burnstock, 1998) − were only detected in P7, but not in adult testes, suggesting developmentally regulated expression. In addition, P2ry1 and P2ry12 mRNA was amplified from P7 and adult tissue, but not Sertoli cells, indicating expression by other testicular cell types. While we cannot unequivocally exclude genomic DNA contamination, our results indicate that P2Y2 receptors are critical for metabotropic ATP signalling in immature mouse Sertoli cells.

To verify this proposed role of P2Y2, we assessed the effect of selective P2Y2 gene silencing (knockdown) on metabotropic Sertoli cell ATP signalling by administration of small interfering RNAs (siRNA). Therefore, cells were transiently transfected with a fluorescent marker and either of two targeting siRNA constructs or a non-targeting negative control siRNA containing a non-specific sequence (Fig. 6A). Under conditions that selectively trigger Ca²⁺ signals mediated by P2Y receptors (i.e. stimulation with a low ATP concentration (350 nm) in Ca²⁺-free medium), both response amplitude (Fig. 6B and C) and probability (Fig. 6B and D) are strongly reduced relative to non-transfected cells as well as non-targeting siRNA controls. Viability of siRNA-treated cells was confirmed by P2X receptor activation using an elevated ATP concentration (10 μm) in regular S₁ solution ([Ca²⁺]_e = 1 mm; Fig. 6B). Effective P2Y2 knockdown by either targeting siRNA was independently confirmed by quantitative real-time PCR (Fig. 6E). Relative to non-targeting siRNA controls, P2Y2 transcript levels were reduced to 34 ± 13% (RNAi 1) and 37 ± 6% (RNAi 2), respectively. Together, these results show that metabotropic Ca²⁺ signalling in Sertoli cells is predominantly, if not exclusively, mediated by P2Y2 receptors.

Figure 6. Post-transcriptional gene silencing reveals a crucial role of P2Y2 receptors in metabotropic Sertoli cell ATP signalling.

A, fura-2 AM-loaded Sertoli cells (grey scale image; 380 nm excitation; top left) were transfected with one of two siRNA constructs to knockdown P2Y2 protein expression and FITC-labelled BLOCK-iT™ fluorescent oligos to enable identification of co-transfected cells by their fluorescent nuclei (top middle). In merged images (top right), transfected (encircled by red lines; ROI 2 and 4) and non-transfected Sertoli cells (delimited by white lines; ROI 1 and 3) are readily identified. Bottom: [Ca²⁺]_c is monitored by ratiometric (F₃₄₀/F₃₈₀) fluorescence imaging. Pseudocolour single frame images illustrate relative [Ca²⁺]_c at time points 1–3 (rainbow 256 colourmap; black, low Ca²⁺; pink, high Ca²⁺). B, original traces (F₃₄₀/F₃₈₀versus time) comparing pairs of transfected (red traces) and non-transfected (black traces) Sertoli cells. ROIs and indicated time points correspond to cells shown in A. In Ca²⁺-free medium (grey horizontal bars), [Ca²⁺]_c elevations in response to ATP (350 nm) are detected in non-transfected cells, whereas administration of P2Y2 siRNA essentially abolished ATP signals in transfected Sertoli cells. Note that ATP responses are observed in all cells when stimulated with an increased ATP concentration (10 μm) under control conditions. C and D, bar charts depicting the mean maximal response amplitude (C) and the average response rate (D) in cells exposed to ATP (350 nm) in Ca²⁺-free medium (means ± SEM). Sertoli cells transfected with either P2Y2 siRNA 1 (n = 8) or P2Y2 siRNA 2 (n = 36) are represented by yellow and red bars, respectively. Cells treated with non-targeting negative control siRNA (neg. ctr. RNAi; n = 15) are shown by grey, untransfected cells by white bars (n = 38). Black bars represent randomly chosen cells that were treated with siRNA, but were not transfected (i.e. not fluorescent; n = 96). Compared to untransfected, not fluorescent, and non-targeting siRNA controls, transfection with either P2Y2 targeting siRNA results in both a significantly diminished ATP response amplitude (C; P < 0.001) and a significantly reduced response probability (D; P < 0.001). No significant differences were observed between controls. E, knockdown of P2Y2 receptor expression in cultured RNAi-treated Sertoli cells was confirmed by quantitative real-time PCR. Relative expression levels (mean ± SEM) of P2Y2 transcripts are shown in a bar graph. α1A-tubulin served as an internal normalization standard. Transcript quantities were normalized to P2Y2 mRNA levels in cells treated with non-targeting siRNA controls. Administration of both targeting siRNAs effectively reduced P2Y2 transcription (siRNA 1 (P < 0.01); siRNA 2 (P < 0.001)).

Mitochondria control purinergic Ca²⁺ mobilization

We next investigated the function of a potentially critical determinant of [Ca²⁺]_c– the Sertoli cell mitochondrial network. First, we analysed mitochondrial distribution and morphology in cultured Sertoli cells by using MitoTracker Deep Red to label functional mitochondria (Fig. 7A). Mitochondria are preferentially localized in perinuclear regions and display a rather tubular morphology. As evident from time-lapse microscopy, however, mitochondrial network distribution is highly dynamic (data not shown). Measuring Ca²⁺ mobilization in the mitochondrial matrix is challenging. Here, based on the combination of (i) genetically engineered PhotoTopo mice that express an optically improved photoprotein (c-Photina) within the mitochondrial matrix (Cainarca et al. 2010), (ii) a dedicated bioluminescence microscope, and (iii) a fast EM-CCD camera with single photon sensitivity, we devised a strategy to enable live-cell bioluminescence [Ca²⁺]_m imaging in single Sertoli cells with both subcellular spatial and millisecond temporal resolution.

Figure 7. Mitochondrial Ca²⁺ can be directly monitored in real-time at single CHO cell resolution.

A, confocal fluorescence images of cultured Sertoli cells after Ca²⁺ dye loading (fluo-4 AM; green) and MitoTracker labelling of energized mitochondria (red). B, [Ca²⁺]_c transients in response to focal ATP application (50 μm; 10 s) as monitored by ratiometric fluorescence imaging in fura-2-loaded Mito c-Photina CHO cells (grey scale image). Original trace depicts the integrated fluorescence ratio F₃₄₀/F₃₈₀ of a representative ATP-sensitive cell in a user-defined ROI (white ellipse) as a function of time. Pseudocolour single frame images illustrate the relative cytosolic Ca²⁺ concentration at time points 1–4 (rainbow 256 colourmap; blue, low Ca²⁺; red, high Ca²⁺). C, representative original trace and corresponding single frame images of a live-cell [Ca²⁺]_m bioluminescence recording from cultured Mito c-Photina CHO cells stimulated with ATP (50 μm; 10 s). RLUs in a user-defined ROI (red ellipse) are plotted as a function of time (grey scale intensity map; black, low [Ca²⁺]_m; white, high [Ca²⁺]_m). On an expanded scale (inset; dotted rectangle), the transient and directly time-locked (black application bar) nature of the bioluminescence signal becomes apparent.

As a proof-of-concept experiment, we comparatively recorded [Ca²⁺]_c and [Ca²⁺]_m transients in CHO cells stably expressing mitochondrial c-Photina (mito c-Photina; Cainarca et al. 2010). In CHO cells, endogenous P2Y receptors couple to the PLC–IP₃ pathway and induce Ca²⁺ release from internal stores (Iredale & Hill, 1993). Accordingly, we recorded robust [Ca²⁺]_c transients in fura-2 AM-loaded mito c-Photina CHO cells in response to ATP exposure (50 μm; 10 s; n = 80; Fig. 7B). When monitoring [Ca²⁺]_m via luminescence microscopy under the exact same experimental conditions, we observed a strong transient ATP-dependent bioluminescence signal (n = 79; Fig. 7C), directly corresponding to a rise in matrix Ca²⁺. The full duration at half-maximum (FDHM) of the [Ca²⁺]_m response was 2.4 ± 0.1 s, whereas the cytosolic Ca²⁺ signal was prolonged (FDHM = 24.8 ± 1.4 s). Mitochondrial origin of the bioluminescence signal was confirmed in pharmacological assays. Dissipation of ΔΨ_m by the ETC complex III inhibitor antimycin A (2 μm) or the protonophore carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP; 10 μm), respectively, abolished [Ca²⁺]_m bioluminescence signals (6/7 experiments, antimycin A; 10/10 experiments, FCCP), whereas ATP still evoked [Ca²⁺]_c transients (n = 80; data not shown). Amplitudes and kinetic parameters of cytosolic signals, however, were significantly altered when mitochondrial Ca²⁺ uptake was impaired (data not shown), indicating that mitochondria are involved in CHO cell Ca²⁺ regulation.

To address whether mitochondria also constitute a component of the purinergic Ca²⁺ signalling machinery in immature Sertoli cells, we adopted the bioluminescence [Ca²⁺]_m imaging approach to primary Sertoli cell cultures from P7 PhotoTopo mice (Fig. 8A). Brief exposure to saturating ATP concentrations (100 μm, 10 s) triggered [Ca²⁺]_m transients in 84% of cells (250/298). At resting conditions, no ‘background bioluminescence’ was detected (Fig. 8A, image 1 and 4; Movie S2). Subcellular resolution at fast frame rates (up to 25 Hz) was routinely achieved with only a minor trade-off in signal-to-noise ratio (Fig. 8A, right inset; Movie S3). When cells were stimulated at different ATP concentrations (n = 31), mitochondrial Ca²⁺ response amplitudes increased dose-dependently (Fig. 8A, left inset). The fitted [Ca²⁺]_m dose–response curve reveals a threshold ATP concentration of <200 nm, an EC₅₀ of 1.87 μm and saturation at ATP levels ≥100 μm (Fig. 8B). From calibrated [Ca²⁺]_c data (Fig. 8B; Fig. S3), it becomes apparent that minute changes in average [Ca²⁺]_c are sufficient to trigger mitochondrial Ca²⁺ uptake. Compared to P2Y-mediated [Ca²⁺]_c signals and P2X-dependent inward currents, dynamic mitochondrial Ca²⁺ buffering is effective at an intermediate range of stimulus concentrations. These data show that P2Y receptor activation alone ([ATP]≤ 1 μm) is sufficient to initiate mitochondrial Ca²⁺ uptake. At stimulus concentrations that exceed the dynamic range of P2Y signalling ([ATP]≥ 3 μm), additional P2X-dependent Ca²⁺ influx is also partly buffered by mitochondria.

Figure 8. Mitochondrial Ca²⁺ uptake in Sertoli cell purinergic signalling.

A, representative trace and corresponding single frame images of a live-cell [Ca²⁺]_m bioluminescence recording from Sertoli cells stimulated with ATP (100 μm, 10 s). RLUs are plotted as a function of time. Insets show a dose-dependent (left) and a fast frame rate (25 Hz; right) [Ca²⁺]_m recording. B, ATP dose–response relationship for [Ca²⁺]_m signals (red triangles; EC₅₀ = 1.87 μm) in comparison to P2X-mediated inward currents (grey rectangles, EC₅₀ = 16.7 μm) and P2Y-dependent [Ca²⁺]_c transients (black dots, EC₅₀ = 0.2 μm). Sigmoidal curves were fitted from average data points (mean ± SEM) using the Hill equation.

Distinct pharmacological profile of purinoceptor-dependent mitochondrial Ca²⁺ signalling

We next assessed the pharmacological profile of purinoceptor-dependent [Ca²⁺]_m signalling by bioluminescence screening of Sertoli cell populations in a plate reader assay. Using a broad spectrum of agonists and antagonists, we investigated the hypothesis that both P2X2 and P2Y2 represent the predominant purinoceptor subtypes in mouse Sertoli cells. When challenged with increasing concentrations of the P2X agonists α,β-methylene ATP, β,γ-methylene ATP, 3′-O-(4-benzoyl)benzoyl-ATP (Bz-ATP) or ATPγS, both Bz-ATP and ATPγS elicited dose-dependent [Ca²⁺]_m transients (n = 12; Fig. 9A), with EC₅₀ values close to (14.8 μm, Bz-ATP) or even lower (1.5 μm, ATPγS) than recorded for P2X-dependent membrane currents (12.7 μm). By contrast, α,β-methylene ATP and β,γ-methylene ATP proved essentially ineffective (n = 12). In fura-2-based control experiments, we confirmed that both Bz-ATP and ATPγS dose-dependently elevate [Ca²⁺]_c at comparably low concentrations (Fig. S5). Mitochondrial Ca²⁺ uptake in response to Bz-ATP (100 μm) was exemplified in high-resolution bioluminescence recordings from single Sertoli cells (Fig. 9B; Movie S4). Activation of P2Y receptors by UTP triggered robust and dose-dependent [Ca²⁺]_m signals (n = 12; EC₅₀ = 0.9 μm; Fig. 9C). Purinoceptor antagonists blocked ATP-evoked [Ca²⁺]_m transients with different efficiency (n = 16; Fig. 9D). Diinosine pentaphosphate (IP5I) and suramin both inhibited ATP responses at concentrations >10 μm (IC₅₀ = 40.8 μm (IP5I) and 38.6 μm (suramin)). Reactive blue-2 (RB-2) and 2′,3′-O-(2,4,6- trinitrophenyl) adenosine 5′-triphosphate (TNP-ATP) acted as more potent antagonists. Inhibition of [Ca²⁺]_m signals became apparent at concentrations ≥3 μm and half-maximal effects were observed at 23.3 μm (RB-2) and 11.9 μm (TNP-ATP), respectively. Together, this pharmacological fingerprint strongly supports the proposed role of P2X2 and P2Y2 in pre-pubertal mouse Sertoli cells (see Discussion).

Figure 9. Pharmacological profile of ATP-dependent [Ca²⁺]_m signals.

A, agonist-specific dose–response curves for ATPγS, Bz-ATP, α,β-methylene ATP and β,γ-methylene ATP. B, example of a high-resolution bioluminescence imaging recording (2 Hz frame rate) from single Sertoli cells stimulated with Bz-ATP (100 μm; 5 s). Individual frames correspond to the indicated time points (application start, 0.0 s; grey scale intensity map). Note: initial [Ca²⁺]_m signals are detected in the cell periphery, whereas perinuclear [Ca²⁺]_m transients display some delay. C, UTP evokes dose-dependent [Ca²⁺]_m signals. By contrast, neither AMP nor adenosine trigger [Ca²⁺]_m increase (Ko et al. 2003). D, dose–response relation for different purinoceptor inhibitors. Half-maximal inhibitory concentrations (IC₅₀) range from 11.9 μm (TNP-ATP) to 40.8 μm (IP5I). Data points represent means ± SEM.

To address the individual impact of either metabotropic or ionotropic ATP signalling on [Ca²⁺]_m, we next performed single Sertoli cell bioluminescence imaging experiments in which we selectively impaired either P2X-dependent Ca²⁺ influx (‘Ca²⁺-free’ medium; n = 101; Fig. 10A) or P2Y-mediated Ca²⁺ signalling (inhibition of SERCA (n = 37; Fig. 10B) or PLC (n = 30; Fig. 10C)). Neither removal of external Ca²⁺ nor PLC inhibition affected the incidence of [Ca²⁺]_m signals (50/55 experiments (‘Ca²⁺-free’ medium) and 14/15 experiments (PLC inhibition)). Depletion of endoplasmic Ca²⁺ stores by thapsigargin, however, significantly reduced the occurrence of ATP-induced [Ca²⁺]_m transients (25/42 experiments). The strength of the [Ca²⁺]_m signal was not significantly affected by removal of external Ca²⁺, whereas pharmacological disruption of the PLC–IP₃ pathway resulted in significantly decreased [Ca²⁺]_m response amplitudes (Fig. 10D; P < 0.001). Analogous results were obtained from [Ca²⁺]_m rising speed analysis. In ‘Ca²⁺-free’ medium, [Ca²⁺]_m increased at a similar rate (243 ± 30 relative luminescence units (RLU) s⁻¹; n = 101) as under control conditions (251 ± 24 RLU s⁻¹; n = 108). By contrast, [Ca²⁺]_m responses developed significantly slower when P2Y signalling was impaired (62 ± 9 RLU s⁻¹, n = 35 (thapsigargin); 124 ± 35 RLU s⁻¹, n = 30 (U73122), P < 0.001). Together, these data suggest that mitochondria sequester Ca²⁺ that is released from internal stores more efficiently than P2X-dependent Ca²⁺ influx across the plasma membrane.

Figure 10. Mitochondria modulate Sertoli cell purinergic signalling.

A–C, original bioluminescence [Ca²⁺]_m recordings in response to ATP (100 μm; 10 s). RLUs in single cell ROIs are plotted versus time. Traces are representative for Ca²⁺-free medium (A), endoplasmic store depletion (thapsigargin; B), and PLC inhibition (U73122; C), respectively. For comparison, insets show traces (F₃₄₀/F₃₈₀versus time) of [Ca²⁺]_c under the exact same conditions. D, bar chart depicting [Ca²⁺]_m signal strength (mean ± SEM) under different ionic/pharmacological conditions relative to control (red horizontal line; RLU = 552; n = 205). [Ca²⁺]_m amplitude was essentially unaffected by removal of external Ca²⁺ (RLU = 580; n = 101). By contrast, both drugs that disrupt P2Y receptor signalling significantly reduced [Ca²⁺]_m response amplitudes (P < 0.001; RLU = 268, n = 37 (thapsigargin); RLU = 247, n = 30 (U73122)).

Interdependent cytosolic and mitochondrial Ca²⁺ dynamics

How do mitochondria and other components of the Sertoli cell Ca²⁺ signalling ‘toolkit’ correlate? Does mitochondrial Ca²⁺ sequestration directly regulate functional parameters of purinergic signalling? To address these questions, we first analysed [Ca²⁺]_m transients that were selectively mediated by P2X receptors (Fig. 11A). Surprisingly, inhibition of P2Y receptor signalling either by impairing PLC-dependent PI turnover (U73122; n = 24), or by depletion of ER Ca²⁺ stores (thapsigargin; n = 32), affected [Ca²⁺]_m response kinetics differently. The efficiency of Ca²⁺ clearance from the mitochondrial matrix was not altered when PLC activity was inhibited. By contrast, SERCA block significantly decelerated [Ca²⁺]_m decay (τ = 2.8 ± 0.1 s (control; n = 154) versus 5.4 ± 0.3 s (thapsigargin)) and, thus, prolonged the [Ca²⁺]_m signal (FDHM = 3.7 ± 0.2 s (control; n = 155) versus 7.0 ± 0.2 s (thapsigargin)). These results indicate that a working ER Ca²⁺ buffer facilitates Ca²⁺ clearance from the mitochondrial matrix. As shown in Fig. S4A and B, SERCA inhibition led to a slow gradual [Ca²⁺]_c increase that lasted for several minutes. However, thapsigargin treatment did not induce a detectable [Ca²⁺]_m elevation (Fig. 10B), suggesting that only rapid [Ca²⁺]_c changes trigger substantial Ca²⁺ sequestration by mitochondria.

Figure 11. Mitochondria shape the purinergic Ca²⁺ response.

A, P2X-mediated [Ca²⁺]_m dynamics depend on the Ca²⁺ sequestration capacity of the ER. [Ca²⁺]_m decay kinetics remain unaffected by PLC inhibition (U73122, 10 μm; τ = 2.2 ± 0.4 s; n = 24), but are strongly delayed when Ca²⁺ uptake by the ER is blocked (thapsigargin, 10 μm; τ = 5.4 ± 0.3 s; n = 32; P < 0.001). Accordingly, average FDHM is not significantly altered by U73122 treatment (3.0 ± 0.1 vs.; n = 23), but significantly prolonged in presence of thapsigargin (7.0 ± 0.2 s; n = 99; P < 0.001). B, CGP37157-dependent block of NCX_mito accelerated mitochondrial Ca²⁺ uptake (n = 89; P < 0.001) and increased [Ca²⁺]_m signal amplitude (n = 140; P < 0.001). C, representative [Ca²⁺]_m recording (RLU versus time). FCCP (10 μm) completely abolishes ATP signals. After washout, ATP (100 μm; 10 s) responses rapidly recover. Oligomycin (5 μm) was added to block ATP synthase. When applied alone, oligomycin had no effect (data not shown). Insets: representative traces (F₃₄₀/F₃₈₀versus time) showing a gradual (left) or more rapid (right) [Ca²⁺]_c increase upon administration of antimycin A (2 μm; left) or FCCP (10 μm; right). D, FCCP did not significantly change Sertoli cell [Ca²⁺]_c response frequency (n = 83), whereas mitochondrial Ca²⁺ uptake was essentially abolished (n = 121). E, bar diagram of Sertoli cell [Ca²⁺]_c decay time constants when Ca²⁺ sequestration by mitochondria is inhibited. Compared to control conditions (red horizontal line), both FCCP and antimycin A significantly decelerated cytosolic Ca²⁺ clearance (τ = 17.1 ± 1.9 s; n = 81; P < 0.001 (FCCP) τ = 17.1 ± 1.2 s; n = 122; P < 0.001 (antimycin A)).

Mitochondrial Ca²⁺ extrusion can be pharmacologically separated from net uptake by the selective NCX_mito inhibitor chloro-5-(2-chlorophenyl)-1,5-dihydro-4,1-benzothiazepin-2(3H)-one (CGP37157; Cox & Matlib, 1993). When NCX_mito-dependent Ca²⁺ shuttling was impaired (CGP37157, 10 μm), we observed a significant increase in both [Ca²⁺]_m signal amplitude and rise time (Fig. 11B; P < 0.001). Qualitatively similar results were obtained using the less specific NCX blocker KB-R7943 (10 μm; data not shown). These results show that Sertoli cell mitochondria function as effective Ca²⁺ shuttling organelles that orchestrate both rapid Ca²⁺ uptake and release.

Agents that cause dissipation of ΔΨ_m inhibit mitochondrial Ca²⁺ uptake (Celsi et al. 2009). We therefore used either FCCP, a rapidly reversible protonophore shown to preferentially target mitochondria (Rizzuto et al. 1992; Herrington et al. 1996), or the respiratory chain blocker antimycin A (Lai et al. 2005). Both drugs were administered in conjunction with oligomycin (5 μm; Penefsky, 1985) to prevent ATP hydrolysis in the reversed mode of ATP synthase. In the presence of FCCP (10 μm), ATP-dependent [Ca²⁺]_m signals in cultured Sertoli cells were completely abolished (28 experiments; Fig. 11C and D). The occurrence of [Ca²⁺]_c transients in response to ATP, however, was not significantly changed. These data confirm the efficacy of the drug as well as the specificity of both mitochondrial c-Photina expression and [Ca²⁺]_m-dependent bioluminescence signalling in Sertoli cells from PhotoTopo mice. Interestingly, dissipation of ΔΨ_m induced an increase in [Ca²⁺]_c (Fig. 11C, insets), suggesting that mitochondria also participate in the homeostatic control of resting [Ca²⁺]_c which, under control conditions, is ∼150 nm in cultured Sertoli cells (Fig. S3). Upon ATP stimulation (100 μm; 10 s), [Ca²⁺]_c recovery was significantly decelerated when mitochondrial Ca²⁺ uptake was blocked, indicating that mitochondrial Ca²⁺ sequestration represents an important mechanism of cytosolic Ca²⁺ clearance (Fig. 11E).

Discussion

Sertoli cells have been the subject of intense research and considerable advances in understanding their biochemistry have been made (Griswold & McLean, 2006; Hermo et al. 2010). Yet, a conceptual mechanistic understanding of the physiological principles that underlie Sertoli cell autocrine and/or paracrine signalling is lacking. In this study, we describe an ATP-dependent Ca²⁺ signalling pathway in immature mouse Sertoli cells that consists of both P2X and P2Y purinoceptors, the ER and, notably, mitochondria. By combining a transgenic mouse model with a dedicated bioluminescence imaging device, we establish a novel method to monitor mitochondria-specific Sertoli cell Ca²⁺ mobilization at subcellular spatial and millisecond temporal resolution. Our data identify the Sertoli cell mitochondrial network as a critical component of the testicular purinergic Ca²⁺ signalling system.

Purinoceptor-induced Ca²⁺ mobilization has been shown to affect the endocrine control of Sertoli cell physiology (Meroni et al. 1998; Rossato et al. 2001; Gelain et al. 2005). In general, ATP-mediated communication gains its versatility and specificity from the functional heterogeneity of the P2 receptor population. Mouse Sertoli cell purinoceptor subtypes, however, have not been identified and a functional description of ATP-dependent Ca²⁺ signalling in these cells is lacking. So far, few studies (exclusively in the adult rat) have addressed this issue at all and controversial data have been reported (Glass et al. 2001; Rossato et al. 2001; Ko et al. 2003). Our results show that pre-pubertal mouse Sertoli cells express both metabotropic and ionotropic purinoceptor subtypes. Molecular, immunocytochemical and functional studies strongly suggest that P2X2 and P2Y2 represent the predominant, if not exclusive, ATP receptor subunits in these cells. The high ATP affinity (EC₅₀ = 0.2 μm) together with the only slightly reduced sensitivity for UTP (EC₅₀ = 0.9 μm) are reminiscent of the G_αq-coupled P2Y2 subtype, a finding confirmed on the mRNA and protein level. The rat P2Y2 receptor has previously been proposed to mediate PI turnover in adult Sertoli cells (Filippini et al. 1994; Rudge et al. 1995), however, in response to considerably higher ATP concentrations (EC₅₀∼20 μm). The biophysical and pharmacological fingerprint of the mouse Sertoli cell P2X receptor matches the characteristic properties reported for recombinant P2X2 channels from various species (Table 3). Our data are thus inconsistent with previous reports of mutually exclusive Sertoli cell expression/function of P2X1 and P2X3 (Rossato et al. 2001) or P2X4 and P2X7 (Ko et al. 2003), respectively. In a recent immunohistochemical analysis, coexpression of P2X2, P2X3 and P2X7 subunits in Sertoli cells of the adult rat testis was reported (Glass et al. 2001). Interestingly, P2X2 and P2X3 expression appeared restricted to stages I–VIII of the spermatogenic cycle. At stage VIII, spermiation and blood-testis barrier restructuring take place simultaneously (Hess & Renato de Franca, 2008) and Sertoli cells mediate the androgen-dependent initiation of meiosis (Griswold & McLean, 2006). It is tempting to speculate that developmentally regulated transient expression of P2X2 could provide a functional basis for stage-specific purinergic communication within the seminiferous epithelium. In other systems, purinergic signalling in embryonic and postnatal development is an emerging concept (Abbracchio et al. 2009) and transient purinoceptor expression appears to regulate cellular proliferation and differentiation.

Table 3.

Summary of biophysical and pharmacological properties of recombinant mammalian P2X receptor subtypes (homo- and heteromeric channels)

	P2X1	P2X2	P2X3	P2X4	P2X5	P2X6	P2X7	P2X2/3	P2X1/5	P2X2/6	P2X4/6
ATP EC50	0.1 μm	10–30 μm	1 μm	10 μm	10 μm	10 μm	>100 μm	1–3 μm	0.1 μm	30 μm	3–10 μm
αβ-meATP EC50	0.3–1 μm	»100 μm	1 μm	»100 μm	»100 μm	»100 μm	»100 μm	3 μm	3 μm	»100 μm	10–30 μm
Bz-ATP EC50	3–30 nm	3–30 μm	0.1 μm	3–10 μm	»100 μm		>100 μm			10 μm	1 μm
βγ-meATP EC50	10 μm	»100 μm	»100 μm				»100 μm	»100 μm		»100 μm
ATPγS EC50	1 μm	1–10 μm	1–3 μm	10–100 μm				10 μm		10 μm
TNP-ATP IC50	5 nm	1–10 μm	1 nm	15 μm	»10 μm		>30 μm	3 nm	0.7 μm	1 μm	15 μm
Suramin IC50	1 μm	3–30 μm	3 μm	no effect	3–10 μm	»100 μm	»300 μm	3 μm	1 μm	2 μm	no effect
Acidic pH (6.3)	inhibition	potentiation	inhibition	inhibition	inhibition		inhibition	potentiation	inhibition	potentiation	inhibition
Alkaline pH (8.3)	no effect	inhibition	no effect	no effect	no effect			inhibition	inhibition	inhibition	no effect
Zn2+ (10–100 μm)	inhibition	potentiation	potentiation	potentiation	potentiation		inhibition	potentiation		potentiation	potentiation
Cu2+ (100 μm)		potentiation		no effect			inhibition			potentiation	no effect
τ (S)	300 ms	»10 s	<100 ms	−1 s	»10 s	»10 s	»10 s	»10 s		»10 s	∼1 s

Shown are EC₅₀ values for P2X receptor agonists, IC₅₀ values for antagonists, general pharmacological effects, and desensitization time constants (τ). Properties that match the characteristics of Sertoli cell P2X receptor currents are shown in italics. Values are taken from the following references: Gever et al. 2006; Alexander et al. 2008; Donnelly-Roberts et al. 2009; Jarvis & Khakh, 2009.

Given the non-overlapping dynamic ranges of the P2Y2 and P2X2 receptor, Sertoli cells are endowed with two parallel, yet functionally distinct signalling pathways that allow for hierarchical concentration-dependent decoding of purinergic information. Moreover, both receptors hardly desensitize and substantial Ca²⁺ mobilization is already observed in response to brief ATP stimulations (≤100 ms). Sertoli cells are thus equipped to respond to both very short and persistent ATP signals in a time-locked fashion. In addition, P2X2-mediated currents trigger substantial and potentially regenerative (e.g. by voltage-dependent Ca²⁺ channels; Lalevee et al. 1997) membrane depolarization, show strong inward rectification, and are modulated by protons and trace elements. Both zinc and copper potentiate P2X2 currents. Interestingly, elevated testicular zinc levels are important for male reproduction and zinc deficiency leads to gonadal dysfunction and impaired spermatogenesis (Bedwal & Bahuguna, 1994).

Purinergic Ca²⁺ mobilization may, thus, provide an effective route for situational control of Sertoli cell Ca²⁺ homeostasis and developmental stage. In this study, we identified the Sertoli cell mitochondrial network as a key component of its Ca²⁺ signalling ‘toolkit’. A dense tubular network of branched elongated mitochondria extends from perinuclear regions to distal cellular areas. The predominant perinuclear localization could reflect a strategic position as a mitochondrial ‘firewall’ that might prevent uncontrolled spread of Ca²⁺ signals to the nucleus (Rizzuto & Pozzan, 2006). Upon ATP stimulation, a rapid and transient increase in [Ca²⁺]_m is detected in the great majority of cells. High-resolution imaging indicates that mitochondrial Ca²⁺ accumulation initially occurs in more distal areas, whereas Ca²⁺ buffering in perinuclear regions is detected with some delay (Fig. 9B). Whether this phenomenon reflects privileged localization of some mitochondria in close proximity to local Ca²⁺‘hot spots’ (Giacomello et al. 2010) remains to be investigated.

What is the physiological role of [Ca²⁺]_m transients in Sertoli cells? Ca²⁺ accumulation could exert a positive feedback on ATP synthesis by activating ETC dehydrogenases, thereby meeting increased cellular energy demands (McCormack et al. 1990). As spermatocytes and spermatids are uncoupled from blood sugar supply after blood-testis barrier passage, they rely on metabolite breakdown by Sertoli cells (Boussouar & Benahmed, 2004). Our data, however, argue for a more complex function of [Ca²⁺]_m signalling in Sertoli cells. Collapse of the IMM H⁺ gradient and dissipation of ΔΨ_m induced an increase in [Ca²⁺]_c, indicating that a low resting level of cytosolic Ca²⁺ is, in part, maintained by mitochondria. Our results, thus, support the notion that, despite the low K_d (∼10 μm) and high activation threshold of the mCU (Kirichok et al. 2004), mitochondria are recruited as Ca²⁺ storage organelles even at rest, a concept that is controversially discussed (Rizzuto & Pozzan, 2006; Contreras et al. 2010).

Sertoli cell mitochondria sequester cytosolic Ca²⁺ regardless of its origin (i.e. ER release versus influx) at relatively low threshold concentrations. Mitochondrial Ca²⁺ uptake and release could thus bear important consequences for the spatio-temporal ‘fingerprint’ of Sertoli cell [Ca²⁺]_c responses, including, but not limited to, purinergic signals. Intramitochondrial calcium tunneling could connect cytosolic microcompartments and accomplish ER Ca²⁺ replenishment (Malli et al. 2005). Close apposition (≤100 nm) of mitochondria to ER Ca²⁺ uptake/release sites has been described (Rizzuto et al. 1998; Csordas et al. 2006). Our results support a concept of Ca²⁺ signalling cross-talk between mitochondria and the ER in Sertoli cells. Ca²⁺ clearance from the matrix is significantly delayed when SERCA proteins are irreversibly blocked, an effect we did not observe after PLC inhibition. Given the reduced overall [Ca²⁺]_m amplitude under conditions of disrupted PLC–IP₃–ER signalling, the decelerated [Ca²⁺]_m decay and increased FDHM were surprising and argue for facilitated Ca²⁺ shuttling between mitochondria and the ER.

We propose that Sertoli cell mitochondria buffer a substantial amount of the ATP-dependent cytosolic Ca²⁺ load. This is illustrated by the significantly decelerated [Ca²⁺]_c extrusion that we measured when mitochondrial Ca²⁺ uptake was inhibited. Matrix Ca²⁺ accumulation might therefore be particularly important for restoring [Ca²⁺]_c base levels under conditions of repeated stimulation. Vice versa, Ca²⁺ release from loaded mitochondria could extend the effective duration of individual [Ca²⁺]_c responses (Pizzo & Pozzan, 2007) and, thus, mediate fundamental physiological functions.

In recent years, the physiological relevance of [Ca²⁺]_m accumulation has been demonstrated in a variety of tissues (Rizzuto & Pozzan, 2006). Currently, however, the major methodological obstacle in live-cell analysis of [Ca²⁺]_m signalling is the lack of organelle-specific Ca²⁺ probes that (a) provide a broad/linear dynamic range and (b) generate sufficient light to measure [Ca²⁺]_m transients in real-time at cellular resolution. Targeting specificity of synthetic fluorescent Ca²⁺ dyes is based on variable physicochemical differences between cellular compartments. Thus, a considerable amount of cytosolic dye remnant ‘blurs’[Ca²⁺]_m measurements. Genetically targeted Ca²⁺ sensor proteins provide superior organelle specificity (Pozzan & Rudolf, 2009). Here, we establish a novel live-cell bioluminescence [Ca²⁺]_m imaging approach that combines the use of primary cells from PhotoTopo mice with a dedicated bioluminescence imaging set-up and, thus, allows recording of (sub)cellular [Ca²⁺]_m signals at millisecond resolution. Irradiation-free [Ca²⁺]_m imaging prevents phototoxic damage and measurements are essentially free of background, considerably increasing the signal-to-noise ratio. Our experimental approach should thus be directly applicable to the analysis of [Ca²⁺]_m in a variety of primary cells and tissues.

In summary, our data identify mitochondria as essential components of the Sertoli cell signalling ‘toolkit’ that control the shape of purinergic Ca²⁺ responses mediated by both P2X2 and P2Y2 receptors.

Glossary

Abbreviations

Bz-ATP

3′-O-(4-benzoyl)benzoyl-ATP

[Ca²⁺]_c

cytosolic Ca²⁺ concentration

[Ca²⁺]_m

mitochondrial Ca²⁺ concentration

CGP37157

chloro-5-(2-chlorophenyl)-1,5-dihydro-4,1-benzothiazepin-2(3H)-one

DAZL

deleted in azoospermia-like

DIC

differential interference contrast

DIV

days in vitro

ER

endoplasmic reticulum

ETC

electron transport chain

FCCP

cyanide-p-trifluoromethoxyphenylhydrazone

FDHM

full duration at half-maximum

FSH

follicle-stimulating hormone

I_ATP

ATP-mediated whole-cell current

IMM

inner mitochondrial membrane

IP5I

diinosine pentaphosphate

ISI

inter-stimulus interval

mCU

mitochondrial Ca²⁺ uniporter

NCX

Na⁺/Ca²⁺ exchanger

PI

phosphoinositide

RB-2

reactive blue-2

RLU

relative luminescence unit

ROI

region of interest

SERCA

sarco/endoplasmic reticulum Ca²⁺ ATPase

siRNA

small interfering RNA

ΔΨ_m

mitochondrial membrane potential

Author contributions

The experiments were performed in the laboratories of H.H., S.L. and M.S. Original concept of research: S.V., H.H., S.L., E.M.N. and M.S. The research was designed by S.V., T.V., S.Ca., S.Co., J.S., E.M.N. and M.S. Data were collected by S.V., T.V., S.Ca., D.F., C.H.E., J.S. and analysed by S.V., T.V., S.Ca., C.H.E. and J.S. The manuscript was written by S.V., E.M.N. and M.S. with assistance from T.V., S.Ca., S.Co., D.F. and J.S. All authors have read and approved the final version of the manuscript.

Supplementary material

Supplementary Figure 1

Supplementary Figure 2

Supplementary Figure 3

Supplementary Figure 4

Supplementary Figure 5

Supplementary Movie S1-4

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