Rapid functional assays of recombinant IP3 receptors

Cell Calcium 38 (2005) 45–51

Rapid functional assays of recombinant IP3 receptors Alex J. Laude a , Stephen C. Tovey a , Skarlatos G. Dedos a , Barry V.L. Potter b , Sarah C.R. Lummis c , Colin W. Taylor a,∗ b

a Department of Pharmacology, Tennis Court Road, Cambridge CB2 1PD, UK Wolfson Laboratory of Medicinal Chemistry, Department of Pharmacy and Pharmacology, Claverton Down, University of Bath, Bath BA2 7AY, UK c Department of Biochemistry, Downing Site, Cambridge CB2 1QW, UK

Received 11 November 2004; received in revised form 25 April 2005; accepted 27 April 2005 Available online 15 June 2005

Abstract Functional assays of inositol 1,4,5-trisphosphate receptors (IP3 R) currently use 45 Ca2+ release methods, fluorescent Ca2+ indicators within either the ER or cytosol, or electrophysiological analyses of IP3 R in the nuclear envelope or artificial bilayers. None of the methods is presently amenable to the rapid, high-throughput quantitative analyses of IP3 R function needed to address the structural determinants of IP3 R behavior. We use a low-affinity Ca2+ indicator (Mag-fluo-4) to measure free [Ca2+ ] within the ER of permeabilized DT40 cells expressing only rat type 1 IP3 R, and establish that the indicator is capable of reliably reporting the Ca2+ release evoked by IP3 . A 96-well fluorescence plate reader equipped for automated fluid additions (FlexStation, Molecular Devices) is used to monitor IP3 -evoked Ca2+ release. The method allows quick and economical functional assays of recombinant IP3 R in small volumes (≤100 ␮l). © 2005 Elsevier Ltd. All rights reserved. Keywords: DT40 cell; IP3 ; Ca2+ mobilization; Screening; Luminal Ca2+ ; Ca2+ indicator

1. Introduction Fluorescent Ca2+ indicators are widely used to measure cytosolic [Ca2+ ] [1–3]. Their use has transformed our understanding of Ca2+ signalling [4] and they now provide a mainstay for high-throughput drug-screening assays [5]. The tools available to measure free [Ca2+ ] within specific subcellular compartments are less well developed and continue to present greater technical challenges. The methods rely either on fortuitous accumulation of conventional EGTA-derived indicators into intracellular organelles [6–8] or on specific targeting of Ca2+ -sensitive luminescent (aequorin, [9,10]) or fluorescent photoproteins. At present, the latter all use calmodulin or a modified calmodulin as the Ca2+ -sensor and the fluorescence (or FRET) from attached fluorescent proteins to report Ca2+ binding [2,11–13]. In general, the conventional indicators are simplest to use and they have the largest ∗

Corresponding author. Tel.: +44 1223 334062; fax: +44 1223 334040. E-mail address: [email protected] (C.W. Taylor).

0143-4160/$ – see front matter © 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.ceca.2005.04.001

dynamic range, but their subcellular targeting is imprecise. Ca2+ -sensitive proteins can be targeted with exquisite precision [2], but aequorin is both insensitive and difficult to calibrate, and the calmodulin-based indicators can be pHsensitive and they have a limited dynamic range [12]. Furthermore, AM-esters of the EGTA-derived indicators allow cell populations to be easily loaded for high-throughput assays, whereas transfection followed by selection of stable cell lines or high-efficiency infection with viral constructs would be required to load cell populations with Ca2+ -sensitive photoproteins. Analyses of cell populations is further compromised by pollution from those cells that accumulate significant amounts of mis-directed or mis-folded fluorescent proteins [13]. The aim of the present work was to develop a convenient, simple and inexpensive functional assay for recombinant inositol 1,4,5-trisphosphate receptors (IP3 R). Using DT40 cells, a chicken B lymphocyte cell line, Kurosaki and co-workers [14] successfully disrupted the genes for all three IP3 R subtypes and so provided the only available

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null-background (DT40KO cells) for IP3 R expression studies. We use these cells stably expressing rat IP3 R1 for our studies. Aequorin, calmodulin-based indicators and EGTA-based indicators have each been used to measure the luminal free [Ca2+ ] within the ER, where most IP3 R are expressed [15]. The resulting estimates of luminal free [Ca2+ ] vary widely, but most fall between about 100 and 500 ␮M [13,16]. The Ca2+ affinities of many of these indicators, whether proteins or EGTA-based, are too high to resolve [Ca2+ ] changes within this range. Nevertheless, our assay requires a luminal Ca2+ -indicator with appropriate sensitivity to Ca2+ . There are particular problems with expressing Ca2+ -sensitive photoproteins in DT40KO cells. The selection required to disrupt expression of two copies of each of the three IP3 R genes [14] provided the cells with resistance to most antibiotics so restricting opportunities for further selection of cells stably expressing photoproteins, and there are no viral vectors for high-efficiency infection of avian cells. The only practicable option at present is to use a low-affinity conventional Ca2+ -indicator to report [Ca2+ ] within the ER. The lowest affinity indicator that we succeeded in loading into the ER of DT40 cells is Mag-fluo-4 (KdCa = 22 ␮M) and we have now used this successfully to develop an inexpensive, medium-throughput functional assay for recombinant IP3 R.

2. Methods 2.1. Materials Media were from Invitrogen and sera were from Sigma. Cell culture plastics, 96-well assay plates with black sides and V-bottomed 96-well compound plates were from Greiner. FlexStation pipette tips were from Robbins Scientific (Solihull). IP3 (1) was from American Radiolabeled Chemicals (St. Louis, MO, USA), Mag-fluo-4 AM was from Molecular Probes, and ionomycin and 2-APB (2-aminoethoxydiphenyl borate) were from Calbiochem. Adenophostin A (2) [17], dimeric IP3 (4) [18] and l-(1,4,5)IP3 (3) were synthesized and purified as previously reported, the latter from d-1,2,4-tri-O-benzyl myo-inositol using a phosphite chemistry approach [19]. Molecular biology reagents were from Invitrogen and other materials, including heparin, TMB-8 (3,4,5-trimethoxybenzoic acid 8-(diethylamino)octyl ester), caffeine and dantrolene were from Sigma. 2.2. Cell culture and transfection DT40 cells, derived from chicken B lymphocytes, and cells in which both copies of each of the three genes encoding IP3 R have been disrupted (DT40KO cells) were provided by Kurosaki and co-workers [14]. Cells were cultured in RPMI 1640 medium supplemented with 10% foetal bovine serum, 1% heat-inactivated chicken serum, 2 mM glutamine

and 50 ␮M 2-mercaptoethanol. Cells were grown at 37 ◦ C in an atmosphere of 95% air and 5% CO2 and were used or passaged when they reached a density of ∼2 × 106 cells/ml. The open reading frame of rat IP3 R1 was amplified by PCR from the expression vector pCMVI-9-IP3 R1 [20] using the following primers: 5 -AGGAATTCGCCACCATGTCTGACAAAATG-3 and 5 -CCGGTACCGAATTCTTAGGCTGGCTGCTGT-3 and then cloned as an EcoRI fragment into pENTR1a vector (Invitrogen). This construct was transferred into the Gateway-compatible expression vector, pcDNA3.2, to generate pcDNA3.2-IP3 R1. Sequencing of this construct confirmed that it was identical to that reported [21]. DT40KO cells were transfected by electroporation with linearized pcDNA3.2-IP3 R1 using a Gene Pulser apparatus (Bio-Rad Laboratories) at 330 V, 500 ␮F and 5 ␮g DNA/106 cells. G418 (2 mg/ml) was used to select and amplify clones of G418-resistant cells. 2.3. Measurement of free [Ca2+ ] within the intracellular stores of permeabilized DT40 cells A low-affinity Ca2+ indicator trapped within the ER was used to measure luminal free [Ca2+ ]. Cells were centrifuged (650 × g, 2 min) and resuspended (2 × 106 cells/ml) in Hepes-buffered saline (HBS) containing 1 mg/ml BSA, 0.2 mg/ml Pluronic F127 and 20 ␮M Mag-fluo-4 AM (from a 20 mM stock in anhydrous DMSO). HBS had the following composition: 135 mM NaCl, 5.9 mM KCl, 11.6 mM Hepes, 1.5 mM CaCl2 , 11.5 mM glucose, 1.2 mM MgCl2 , pH 7.3. After 60 min (20 ◦ C in dark with gentle shaking), cells were centrifuged (650 × g, 2 min) and suspended in Ca2+ free cytosol-like medium (50 ml, CLM) containing 10 ␮g/ml saponin. Cells were incubated with shaking at 37 ◦ C for 4 min and then, when all cells were permeable to Trypan blue (0.1%), they were centrifuged (650 × g, 2 min) and resuspended in CLM without Mg2+ , but supplemented with 375 ␮M CaCl2 buffered with EGTA to give a free [Ca2+ ] of 220 nM after addition of 1.5 mM MgATP (see below). CLM had the following composition:140 mM KCl, 20 mM NaCl, 1 mM EGTA, 2 mM MgCl2 , 375 ␮M CaCl2 , 20 mM Pipes, pH 7.0. After washing (650 × g, 2 min), cells were resuspended (5–10 × 106 cells/ml) in Mg2+ -free CLM supplemented with FCCP (10 ␮M). Cells (100 ␮l) were added to each well of a black-sided 96-well plate coated with polyl-lysine (0.01%), centrifuged (300 × g, 2 min) to spin cells onto the base of each well and then used for FlexStation analyses. Fluorescence was measured using a FlexStationTM (Molecular Devices, Sunnyvale, CA, USA, www. moleculardevices.com), a 96-well fluorescence spectrometer that allows up to three sequential additions to each well while recording fluorescence from a central spot 1.5 mm across in each of eight wells (one column of a 96-well plate). All experiments were performed at 20 ◦ C. Compounds were added from a 96-well reservoir plate, with the pipette heights, volumes of additions (usually 10 ␮l), rate of addition

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and mixing protocols optimized to minimize disturbance of the cells while ensuring rapid mixing. A Xe-lamp provided excitation light and monochromators were used to select excitation (485 nm) and emission (520 nm) wavelengths. At each time interval (1.5–2 s), five readings were taken from each well and the data stored for later analysis using SoftmaxPro (Molecular Devices). Mag-fluo-4 fluorescence was calibrated to free [Ca2+ ] from [Ca2+ ] = Kd (F − Fmin )/(Fmax − F) where KdCa = 22 ␮M (Molecular Probes; www.probes.com). Measurements of Fmin and Fmax are described later. 2.4. Analyses Concentration-effect relationships were fitted to fourparameter logistic equations using non-linear curvefitting (GraphPad Prism 4.0). Results are expressed as means ± S.E.M. for n independent experiments, with each performed in triplicate.

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3. Results 3.1. Loading of intracellular Ca2+ stores Most experiments used DT40 cells stably expressing rat IP3 R1 (DT40-IP3 R1 cells) (Fig. 1A). Our initial experiments used Mg2+ -containing CLM (with the free [Ca2+ ] buffered to 220 nM with EGTA), as used in previous studies of permeabilized cells [22]. Addition of ATP (1.5–10 mM) stimulated Ca2+ uptake but it first caused an immediate small decrease in Mag-fluo-4 fluorescence (not shown). The latter resulted from ATP chelating free Mg2+ , so reducing the amount of Mg2+ bound to residual Mag-fluo-4 in the medium. In all subsequent experiments, the problem was avoided by beginning the recording in Mg2+ -free CLM (see Section 2) and then initiating Ca2+ uptake by addition of 1.5 mM MgATP (to avoid any change in free [Mg2+ ]). Addition of ionomycin (1 ␮M) and BAPTA (5 mM) to cells prior to addition of ATP had no effect on the fluorescence (not shown) confirming

Fig. 1. Ca2+ uptake by thapsigargin-sensitive Ca2+ stores. (A) Western Blot using an antibody selective for mammalian IP3 R1 [31] showing expression of rat IP3 R1 in DT40-IP3 R1 cells. (B) Addition of ionomycin (1 ␮M) and BAPTA (5 mM) to cells loaded with Ca2+ rapidly restores fluorescence to its initial value. (C) Addition (arrow) of MgCl2 (1.5 mM, i) or ATP alone (1.5 mM, ii) to permeabilized cells does not stimulate Ca2+ uptake. (D) After addition of MgATP (1.5 mM, first arrow, i), fluorescence increases rapidly to a plateau. The increase is completely prevented by thapsigargin (1 ␮M added at first arrow, followed by ATP at second arrow, ii). (E) In stores loaded with Ca2+ after addition of MgATP (arrow), addition of thapsigargin (1 ␮M, ii, solid bar; vs. CLM alone in i) causes the fluorescence to decline with no detectable latency. Traces show means ± S.E.M., n ≥ 6.

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that the stores were empty. Their addition after store-loading restored the fluorescence to its initial level (Fig. 1B). The affinity of the indicator is too low to be significantly affected by the low (220 nM) free [Ca2+ ] of CLM. These observations justify our use of the fluorescence recorded before addition of ATP as the Fmin value used in subsequent analyses. Both ATP (1.5 mM) and Mg2+ (1.5 mM) were required for Ca2+ uptake. Addition of ATP caused no increase in Mag-fluo-4 fluorescence (Fig. 1Cii), and addition of MgCl2 (1.5 mM) caused only the small increase expected from its binding to indicator in the medium (Fig. 1Ci). But after addition of MgATP the fluorescence increased (halftime = 8.5 ± 0.5 s, n = 30) to reach a plateau after <2 min (Fig. 1D). Thapsigargin (1 ␮M), a selective inhibitor of the ER/SR-Ca2+ -ATPase [23], abolished the MgATP-dependent increase in fluorescence (Fig. 1D). Addition of thapsigargin (1 ␮M) to cells that had reached a steady-state Ca2+ loading, caused the Mag-fluo-4 fluorescence to decrease towards its initial level and with no detectable latency (Fig. 1E). 3.2. Calibration of fluorescence signals In order to use fluorescence signals to measure the behavior of IP3 R, it is essential to establish whether the Ca2+ -indicator saturates during Ca2+ loading. The simplest means of achieving this, and of calibrating fluorescence to [Ca2+ ], is to use an ionophore, ionomycin, sequentially to equilibrate the indicator with Ca2+ -free medium and then with medium containing a saturating [Ca2+ ]. This simple approach fails in our FlexStation analyses because residual indicator in the medium contributes to the calibration, but not to the fluorescence signals generated by loading stores with Ca2+ . Hence, addition of CaCl2 (10 mM), even in the absence of ionophore, caused a large increase in fluorescence. Our methods were optimised to minimize this residual indicator, but we failed to eliminate it: additional washing or centrifugation steps caused loss of cells without improving the ratio of trapped to untrapped indicator. The problem, therefore, is to establish whether the fraction of Mag-fluo-4 trapped within intracellular stores, and so reporting Ca2+ uptake and release, is saturated when the stores are loaded. Because thapsigargin causes an immediate decrease in fluorescence when added to loaded stores (Fig. 1E), the stores need not lose substantial amounts of Ca2+ before the loss is reported by the indicator. It is unlikely, therefore, that the indicator is saturated. Results shown in Fig. 2 support this conclusion. CaCl2 (10 mM) was added alone or with Triton X-100 (0.1%, rapidly to permeabilize all membranes) to cells loaded with Ca2+ . The latter caused the fluorescence rapidly (≤5 s) to reach its maximal value, whereas addition of Ca2+ alone caused an equally rapid saturation of only a fraction of the fluorescence, followed by a slow (∼60 s) saturation of the remaining fluorescence (Fig. 2ii). The latter was similar whether thapsigargin was present or not (not shown). We interpret the rapid increase in fluores-

Fig. 2. Calibration of fluorescence signals. Cells were loaded to steady-state with Ca2+ before addition of CaCl2 (10 mM) alone (ii) or with Triton-X-100 (0.1%, i). The vertical bar denotes the component of fluorescence attributed to Mag-fluo-4 in the medium and the histogram compares the amplitude of this fluorescence detected in the experiment (c) with that detected by addition of Ca2+ (10 mM) to an identical volume of medium removed from the cells (m).

cence to reflect Ca2+ binding to untrapped indicator and the slower increase to binding to luminal indicator as Ca2+ leaks down its concentration gradient into the stores. Consistent with this interpretation, when medium was removed from the wells and Ca2+ then added to it, the increase in fluorescence was similar to that observed during the rapid phase of the response to Ca2+ addition to cells (Fig. 2). From this analysis, we estimate that 26 ± 2% (n = 12) of the maximal fluorescence comes from untrapped indicator. Because Mag-fluo-4 has such low-affinity for Ca2+ (Kd = 22 ␮M), this indicator does not contribute significantly to the fluorescence signals recorded during an experiment. With 74 ± 2% of indicator trapped within stores, it is clear that the steady-state fluorescence signal (Fig. 2) causes saturation of 69 ± 2% of that trapped indicator (i.e. 69% of indicator has Ca2+ bound and 31% is free). Hence, when the stores have loaded to steadystate with Ca2+ , the luminal free [Ca2+ ]/Kd = 0.69/0.31 = 2.2. If the affinity of Mag-fluo-4 within the stores is similar to that determined in solution (KdCa = 22 ␮M, but see below), this would suggest a steady-state luminal free [Ca2+ ] of ∼48 ␮M. We conclude that under the conditions used for our assays, where cells are bathed in medium designed to mimic the cytosolic [Ca2+ ] of unstimulated cells, the luminal [Ca2+ ] within the thapsigargin-sensitive Ca2+ store is ∼2.2× the Kd of luminal Mag-fluo-4. We can be less confident of the absolute luminal [Ca2+ ] because the Kd of the indicator may not be the same within the stores and in solution. The Ca2+ -affinities of each of a range of Ca2+ indicators are lower (by factors of two to six) in cytosol or nucleoplasm relative to their published affinities measured in solution [24]. A similar disparity for luminal and free Mag-fluo-4, could easily increase our estimate of luminal free [Ca2+ ] to >100 ␮M. The Km of SERCA for Ca2+ is <1 ␮M [25]: active Ca2+ uptake into thapsigargin-sensitive stores is therefore saturated once the cytosolic free [Ca2+ ] exceeds ∼10 ␮M. By allowing stores actively to accumulate Ca2+ from normal CLM, and then adding additional Ca2+ , we established that there was

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Fig. 3. IP3 -evoked Ca2+ release. (A) Fluorescence changes evoked by addition of IP3 to DT40-IP3 R1 cells. (B) Addition of 10 ␮M IP3 to DT40KO cells causes no change in fluorescence. (C) Concentration-dependent release of Ca2+ by IP3 . Results show means ± S.E.M., n ≥ 3.

a further slow increase in fluorescence (as shown in Fig. 2) only when the medium free [Ca2+ ] exceeded ∼110 ␮M (suggesting that luminal free [Ca2+ ] must exceed 110 ␮M) and then reached a plateau (presumably reflecting saturation of luminal indicator) when the medium free [Ca2+ ] exceeded 180 ␮M. This result is consistent with our conclusion that the luminal indicator is not saturated, that the indicator probably has a reduced affinity for Ca2+ within cells, and with published estimates of luminal free [Ca2+ ] (typically 100–500 ␮M) [13,16]. 3.3. Ca2+ release mediated by IP3 R1 The time course of Ca2+ loading was similar for DT40KO and DT40-IP3 R1 cells, but only the latter responded to IP3 with a decrease in Mag-fluo-4 fluorescence (Fig. 3A and B). Because the free [Ca2+ ] within fully loaded stores is only 2.2× the KdCa of the indicator (see above), we assume a linear relationship between fluorescence and free [Ca2+ ] when measuring concentration–effect relationships. The errors arising from this assumption are small. The effects of agonists of IP3 R are, therefore, expressed as percentages

Fig. 4. Effects of different ligands of IP3 R. (A) Structures of the ligands. (B) Concentration-dependent effects of each ligand on Ca2+ release: (
) l(1,4,5)IP3 , (䊉) d-(1,4,5)IP3 , () adenophostin A, () dimeric IP3 . Results show means ± S.E.M., n ≥ 6.

of the Floaded − Fmin fluorescence signals. In DT40-IP3 R1 cells, a maximal concentration of IP3 (10 ␮M) caused the fluorescence to decrease by 73 ± 3%, the EC50 was 44 ± 9 nM and the Hill slope was 0.9 ± 0.1 (Fig. 3C). The potencies of additional ligands of IP3 R1 derived using this method are consistent with previous work using conventional 45 Ca2+ flux assays (Fig. 4; Table 1). l-(1,4,5)IP (≤10 ␮M) 3 failed to stimulate Ca2+ release and adenophostin A and a dimer of IP3 released the same fraction of the stores as IP3 , but both were more potent than IP3 . Table 1 Ca2+ release by ligands of IP3 R1

IP3 l-(1,4,5)IP3 Adenophostin A IP3 dimer

EC50 (nM)

Hill coefficient

Maximal release (%)

44 ± 9 Inactive at 10 ␮M 1.7 ± 0.2 5.4 ± 0.7

0.9 ± 0.1 – 1.2 ± 0.1 1.0 ± 0.1

73 ± 3 – 75 ± 1 75 ± 1

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Fig. 5. Effects of antagonists of IP3 R. (A) Ca2+ uptake into intracellular stores in the presence of MgATP (added at arrow) and the indicated concentrations of 2-APB. (B) Effects of IP3 on Ca2+ release in the presence of the indicated concentrations of 2-APB. (C and D) Effects of heparin on IP3 -evoked Ca2+ release, and Schild analysis of the data (D). (E and F) Effects of caffeine (E) and TMB-8 (F) on IP3 -evoked Ca2+ release. All antagonists (except caffeine, which was present throughout the Ca2+ loading) were added once the stores had loaded with Ca2+ and then for 30 s before addition of IP3 . All results are means ± S.E.M., n ≥ 3.

3.4. Antagonists of IP3 R1 Fig. 5 summarises results examining the effects of antagonists of intracellular Ca2+ channels. 2-APB, which is widely used as an antagonist of IP3 R because it is membrane-permeant, inhibited responses to IP3 (Fig. 5B) but at concentrations >10 ␮M, it both inhibited Ca2+ uptake (Fig. 5A) and caused non-parallel rightward shifts of the IP3 concentration–effect relationships (Fig. 5B). These results are consistent with reports suggesting that 2-APB has many additional effects and that it is of limited utility as an antagonist of IP3 R [26]. Responses to IP3 were competitively inhibited by heparin (Sigma Mr ∼ 3000, Fig. 5C and D), which is membrane-impermeant. From Schild analysis of the data (slope = 0.95 ± 0.1) the affinity (Kd ) of IP3 R1 for heparin was 5.7 ␮g/ml (∼2 ␮M, Fig. 5D). Although caffeine activates ryanodine receptors, it is also used as a very lowaffinity membrane-permeant antagonist of IP3 R. Our results are consistent with caffeine being a competitive antagonist (slope of Schild plot = 1.2 ± 0.1) with Kd = 7 mM. Neither

dantrolene (≤100 ␮M, not shown), a selective antagonist of type 1 ryanodine receptors, nor TMB-8 (Fig. 5F), another antagonist of ryanodine receptors [27], affected IP3 -evoked Ca2+ release at concentrations more than sufficient to inhibit ryanodine receptors. 3.5. Conclusions To date, assays based on measuring 45 Ca2+ fluxes have provided the simplest means of quantifying IP3 R function [22]. Single channel recording [28,29], and luminal [6,9] or cytosolic [30] Ca2+ indicators have each made important contributions but the existing methods are not amenable to rapid, high-throughput analysis of the function of recombinant IP3 R. Using DT40KO cells as an expression system for mammalian IP3 R, Mag-fluo-4 reliably to report changes in luminal free [Ca2+ ], and a FlexStation to provide rapid and semi-automated recording and fluid additions, we have developed a method that allows quick and economical functional assays of recombinant IP3 R in small volumes (≤100 ␮l).

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Acknowledgements This work was supported by grants from the Wellcome Trust to CWT (072084), BVLP (060554) and SCRL. References [1] R.Y. Tsien, Fluorescent probes of cell signalling, Annu. Rev. Neurosci. 12 (1989) 227–253. [2] R.Y. Tsien, Imagining imaging’s future, Nat. Rev. Mol. Cell Biol. 4 (2003) SS16–SS21. [3] R. Rudolf, M. Mongillo, R. Rizzuto, T. Pozzan, Looking forward to seeing calcium, Nat. Rev. Mol. Cell Biol. 4 (2003) 579–586. [4] M.J. Berridge, M.D. Bootman, H.L. Roderick, Calcium signalling: dynamics, homeostasis and remodelling, Nat. Rev. Mol. Cell Biol. 4 (2003) 517–529. [5] P.A. Johnston, P.A. Johnston, Cellular platforms for HTS: three case studies, Drug Discovery Today 7 (2002) 353–363. [6] A.M. Hofer, T.E. Machen, Technique for in situ measurement of calcium in intracellular inositol 1,4,5-trisphosphate-sensitive stores using the fluorescent indicator Mag-fura-2, Proc. Natl. Acad. Sci. U.S.A. 90 (1993) 2598–2602. [7] L. Combettes, T.R. Cheek, C.W. Taylor, Regulation of inositol trisphosphate receptors by luminal Ca2+ contributes to quantal Ca2+ mobilization, EMBO J. 15 (1996) 2086–2093. [8] T. Miyakawa, A. Maeda, T. Yamazawa, K. Hirose, T. Kurosaki, M. Iino, Encoding of Ca2+ signals by differential expression of IP3 receptor subtypes, EMBO J. 18 (1999) 1303–1308. [9] P. Pinton, T. Pozzan, R. Rizzuto, The golgi apparatus is an inositol 1,4,5-trisphosphate-sensitive Ca2+ store, with functional properties distinct from those of the endoplasmic reticulum, EMBO J. 17 (1998) 5298–5308. [10] P. Pinton, D. Ferrari, E. Rapizzi, F. Di Virgilio, T. Pozzan, R. Rizzuto, The Ca2+ concentration of the endoplasmic reticulum is a key determinant of ceramide-induced apoptosis: significance for the molecular mechanism of Bcl-2 action, EMBO J. 20 (2001) 2690–2701. [11] A. Miyawaki, J. Llopis, R. Heim, J.M. McCaffery, J.A. Adams, M. Ikura, R.Y. Tsien, Fluorescent indicators for Ca2+ based on green fluorescent proteins and calmodulin, Nature 388 (1997) 882–887. [12] T. Nagai, S. Yamada, T. Tominaga, M. Ichikawa, A. Miyawaki, Expanded dynamic range of fluorescent indicators for Ca2+ by circularly permuted yellow fluorescent proteins, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 10554–10559. [13] N. Demaurex, M. Frieden, Measurements of the free luminal ER Ca2+ concentration with targeted “cameleon” fluorescent proteins, Cell Calcium 34 (2003) 109–119. [14] H. Sugawara, M. Kurosaki, M. Takata, T. Kurosaki, Genetic evidence for involvement of type 1, type 2 and type 3 inositol 1,4,5trisphosphate receptors in signal transduction through the B-cell antigen receptor, EMBO J. 16 (1997) 3078–3088.

51

[15] C.W. Taylor, A.A. Genazzani, S.A. Morris, Expression of inositol trisphosphate receptors, Cell Calcium 26 (1999) 237–251. [16] F.L. Bygrave, A. Benedetti, What is the concentration of calcium ions in the endoplasmic reticulum? Cell Calcium 19 (1996) 547– 551. [17] R.D. Marwood, V. Correa, C.W. Taylor, B.V.L. Potter, Synthesis of adenophostin A, Tetrahedron: Asymmetry 11 (2000) 397–403. [18] A.M. Riley, A.J. Laude, C.W. Taylor, B.V.L. Potter, Dimers of dmyo-inositol 1,4,5-trisphosphate: design, synthesis, and interaction with Ins(1,4,5)P3 receptors, Bioconjugate Chem. 15 (2004) 278– 289. [19] A.M. Cooke, B.V.L. Potter, Synthesis of dl-myo-inositol 1,4,5trisphosphate, Tetrahedon Lett. 28 (1987) 2305–2308. [20] G.A. Mignery, C.L. Newton, B.T. Archer, T.C. S¨udhof, Structure and expression of the rat inositol 1,4,5-trisphosphate receptor, J. Biol. Chem. 265 (1990) 12679–12685. [21] T.J.A. Cardy, C.W. Taylor, A novel role for calmodulin: Ca2+ independent inhibition of type-1 inositol trisphosphate receptors, Biochem. J. 334 (1998) 447–455. [22] C.W. Taylor, J.S. Marchant, Measuring inositol 1,4,5-trisphosphateevoked 45 Ca2+ release from intracellular stores, in: G. Milligan (Ed.), Signal Transduction. A Practical Approach, IRL Press, Oxford, 1999, pp. 361–384. [23] O. Thastrup, P.J. Cullen, B.K. Drøbak, M.R. Hanley, A.P. Dawson, Thapsigargin, a tumor promoter, discharges intracellular Ca2+ stores by specific inhibition of the endoplasmic reticulum Ca2+ -ATPase, Proc. Natl. Acad. Sci. U.S.A. 87 (1990) 2466–2470. [24] D. Thomas, S. Tovey, T.J. Collins, M.D. Bootman, M.J. Berridge, P. Lipp, A comparison of fluorescent Ca2+ indicator properties and their use in measuring elementary and global Ca2+ signals, Cell Calcium 28 (2000) 213–223. [25] J. Lytton, M. Westlin, S.E. Burk, G.E. Shull, D.H. MacLennan, Functional comparisons between isoforms of the sarcoplasmic or endoplasmic reticulum family of calcium pumps, J. Biol. Chem. 267 (1992) 14483–14489. [26] F. Wissing, W.P. Nerou, C.W. Taylor, A novel Ca2+ -induced Ca2+ release mechanism mediated by neither inositol trisphosphate nor ryanodine receptors, Biochem. J. 361 (2002) 605–611. [27] P. Palade, C. Dettbarn, D. Brunder, P. Stein, G. Hals, Pharmacology of calcium release from sarcoplasmic reticulum, J. Bioenerg. Biomembr. 21 (1989) 295–320. [28] I. Bezprozvanny, J. Watras, B.E. Ehrlich, Bell-shaped calciumresponse curves for Ins(1,4,5)P3 - and calcium-gated channels from endoplasmic reticulum of cerebellum, Nature 351 (1991) 751–754. [29] D.-O. Mak, J.K. Foskett, Single channel inositol 1,4,5-trisphosphate receptor currents revealed by patch clamp of isolated Xenopus oocyte nuclei, J. Biol. Chem. 269 (1994) 29375–29378. [30] T. Meyer, D. Holowka, L. Stryer, Highly cooperative opening of calcium channels by inositol 1,4,5-trisphosphate, Science 240 (1988) 653–656. [31] T.J.A. Cardy, D. Traynor, C.W. Taylor, Differential regulation of types 1 and 3 inositol trisphosphate receptors by cytosolic Ca2+ , Biochem. J. 328 (1997) 785–793.