The GTPase Ras is a molecular switch engaged downstream of G-protein-coupled receptors and receptor tyrosine kinases that controls multiple cell-fate-determining signalling pathways. Ras signalling is frequently deregulated in cancer, underlying associated changes in cell phenotype. Although Ca2+ signalling pathways control some overlapping functions with Ras, and altered Ca2+ signalling pathways are emerging as important players in oncogenic transformation, how Ca2+ signalling is remodelled during transformation and whether it has a causal role remains unclear. We have investigated Ca2+ signalling in two human colorectal cancer cell lines and their isogenic derivatives in which the allele encoding oncogenic K-Ras (G13D) was deleted by homologous recombination. We show that agonist-induced Ca2+ release from the endoplasmic reticulum (ER) intracellular Ca2+ stores is enhanced by loss of K-RasG13D through an increase in the Ca2+ content of the ER store and a modification of the abundance of inositol 1,4,5-trisphosphate (IP3) receptor (IP3R) subtypes. Consistently, uptake of Ca2+ into mitochondria and sensitivity to apoptosis was enhanced as a result of K-RasG13D loss. These results suggest that suppression of Ca2+ signalling is a common response to naturally occurring levels of K-RasG13D, and that this contributes to a survival advantage during oncogenic transformation.
Ras proteins serve as molecular switches downstream of receptor tyrosine kinases and upstream of the Raf protein kinases (Cully and Downward, 2008; Downward, 2003a; Downward, 2003b; Schulze et al., 2004). This pathway is frequently de-regulated in cancer due to mutation in receptor tyrosine kinases (RTKs) (e.g. EGFR), Ras itself (∼20% of all human cancers) and B-Raf (Downward, 2003b). These mutations elicit significant consequences for cell fate owing to their position as upstream regulators of multiple pathways involved in the regulation of cell cycle, metabolism and cell death – hallmarks of the transformed phenotype (Hanahan and Weinberg, 2000; Hanahan and Weinberg, 2011). This central role of Ras and the downstream pathways it engages have been targeted by the pharmaceutical industry in the development of cancer therapeutics. Indeed, drugs targeting the B-Raf–Mek–Erk pathway have now been approved in the clinic (Belden and Flaherty, 2012; Little et al., 2013).
As Ras lies upstream of multiple cellular pathways, redundancy in function between these signal transduction cascades allows transformed cells to overcome drug targeting and develop resistance (Little et al., 2013). Many of these downstream pathways are also dysregulated in cancer (Wu et al., 2013). Understanding the nature of interactions between Ras and other major cellular signalling pathways is therefore essential for development of effective strategies for suppression of Ras-driven cancer (Wu et al., 2013). A major, but as yet undefined, signalling interaction in oncogenic transformation is that between Ras and Ca2+.
Ca2+ is a pleiotropic signalling messenger that, like Ras, plays key roles in life and death choices, including the decision to proliferate or die by apoptosis (Berridge et al., 2003) (Berridge et al., 1998). Oscillations in cytoplasmic Ca2+ are necessary to sustain the cell cycle, via calmodulin (CaM) (Cullen and Lockyer, 2002; Kahl and Means, 2003), whereas Ca2+ overload by the mitochondria is an initiator of the intrinsic apoptotic cascade (Rizzuto et al., 2003). Not surprisingly therefore, deregulation of Ca2+ homeostasis has been reported in diseases associated with overt or diminished proliferation and increased or insufficient cell death. Cancer cells are characterized in part by uncontrolled proliferation and apoptosis evasion (Hanahan and Weinberg, 2000; Hanahan and Weinberg, 2011); these characteristics have been proposed to be supported by a remodelling of their Ca2+ signalling toolkit (Roderick and Cook, 2008). Indeed, alterations in the expression of a number of Ca2+-handling proteins have been reported in various tumours (Monteith et al., 2007), but attempts to formulate general principles of Ca2+ signalling alterations in cancer have thus far failed. Little consistency in the alterations in Ca2+ protein expression is found among different tumours and between studies. Moreover, few studies have clarified whether altered Ca2+ signalling contributes to the cancer phenotype or is a consequence.
Nowhere is this more apparent than in studies linking Ras to Ca2+ signalling, which go back over 25 years and reveal a complex interplay between these pathways. For example, Ras was shown to enhance agonist-regulated inositol 1,4,5-trisphosphate (IP3) production (Hashii et al., 1993; Lang et al., 1991; Wakelam et al., 1986), a result that might be owing to the ability of Ras protein to bind phospholipase C (PLC) ε (Bunney et al., 2006; Bunney and Katan, 2006). Conversely, Ca2+ signalling can activate certain Ras guanine-nucleotide-exchange factors (GEFs) or Ras GTPase-activating proteins (GAPs) to promote or inhibit activation of Ras and Ras-dependent signalling (Cook and Lockyer, 2006; Roderick and Cook, 2008). The normal interplay between these events is complex and is made all the more so in cells expressing mutant oncogenic variants of Ras such as those harbouring missense substitutions at Gly12, Gly13 or Gln61 (Barbacid, 1987), which prevent the hydrolysis of GTP by GAPs, resulting in Ras being permanently active (Bollag and McCormick, 1991). These de-regulated Ras oncoproteins activate several effector pathways and contribute to virtually all of the hallmarks of the cancer cell (Hanahan and Weinberg, 2000; Hanahan and Weinberg, 2011), including promoting cell proliferation and survival. This pleiotropy of Ras undoubtedly contributes to some of the striking cell- and tissue-specific differences in the regulation of Ca2+ signalling seen in studies with oncogenic Ras mutants. In addition, however, it is apparent that mutant Ras proteins can elicit quite different effects depending on their expression level. Most strikingly, conditional overexpression of oncogenic K-Ras elicits cell cycle arrest and senescence in primary mouse embryo fibroblasts whereas conditional expression at endogenous levels causes cell proliferation and oncogenic transformation (Tuveson et al., 2004). Thus, although studies employing conditional overexpression of mutant Ras proteins have merits, it is important to confirm results in cell systems with native expression levels of mutant Ras to avoid artefacts arising from overexpression.
To overcome this issue, we have taken advantage of isogenic cell line pairs in which the KRAS allele encoding oncogenic K-Ras has been ablated by homologous recombination (Shirasawa et al., 1993). The parental cancer cell line harbouring the mutant allele can then be directly compared with an isogenic derivative that is identical apart from the lack of oncogenic Ras. This approach has the benefit of comparing the effects of a single copy of mutant KRAS rather than using supra-physiological expression. Using this approach, we show that oncogenic K-Ras inhibits Ca2+ release from the endoplasmic reticulum (ER), reduces ER Ca2+ levels and suppresses Ca2+ flux to the mitochondria. These results suggest that suppression of Ca2+ signalling is a common response to naturally occurring levels of K-RasG13D that contributes to a survival advantage during oncogenic transformation.
IP3-induced Ca2+ release is increased in cells deleted of K-RasG13D
Although alterations in expression of a number of proteins involved in Ca2+ regulation in various tumour types and tumour-derived cell lines have been described (Monteith et al., 2007), few studies have analysed how Ca2+ signalling is altered as a result of transformation. Moreover, the nature and role of the effect of the presence of natively expressed ‘driving’ oncogenes upon Ca2+ homeostasis has not been determined. As such, consensus regarding how Ca2+ signalling participates in cellular transformation is lacking (Roderick and Cook, 2008). Contributing to this great variability is the problem in identifying appropriate experimental controls for the cancer cells studied and the issues associated with use of experimental systems in which pleiotropic oncogenes are expressed at supra-physiological levels (Tuveson et al., 2004). In this study, we sought to analyse the effects of a single oncogenic allele at physiological expression levels. To this end, we compared the HCT116 colorectal cancer cell line (K-RasG13D/WT) with its isogenic derivative HKH2 (K-Ras−/WT) in which the mutated Ras allele has been deleted by homologous recombination. In contrast to HCT116 cells, HKH2 cells do not grow in soft agar and do not form tumours in nude mice (Shirasawa et al., 1993). We employed these cell lines to evaluate whether the presence of the endogenous oncogenic K-RasG13D allele modified the generation of Ca2+ signals. As shown in Fig. 1, as a consequence of loss of K-RasG13D in HKH2 cells, Ca2+ signals induced following stimulation of purinergic receptors with ATP were enhanced when compared to HCT116 cells (Fig. 1Ai). This difference was evident in the percentage of responding cells (Fig. 1Aii), in the amplitude (Fig. 1Aiii) and in the integral (area under the curve; AUC) of the Ca2+ transients (Fig. 1Aiv).
To isolate the contribution of Ca2+ influx to the agonist-induced Ca2+ transient, experiments were performed in the absence of extracellular Ca2+. As observed in Ca2+-containing buffer, the AUC and responsiveness to agonist (applied at a concentration where the greatest differences in agonist responses were observed in Ca2+-containing buffer) remained greater in HKH2 than HCT116 cells when Ca2+ was omitted from the imaging buffer. These data therefore indicated that K-RasG13D in HCT116 cells was acting to suppress Ca2+ release from the ER (Fig. 1B).
To further probe the interaction between Ras and Ca2+ signalling in HCT116, Ras expression was also suppressed by small interfering RNA (siRNA). Using this approach, Ras expression was reduced by 85% when compared to HCT116 cells transfected with control non-targeting siRNA (Fig. 2A). siRNA depletion of Ras in HCT116 cells resulted in a significant increase in ATP-stimulated Ca2+ signals in these cells (Fig. 2B). The increase in Ca2+ signalling was manifest as an increase in the percentage of responding cells and in the amplitude and AUC of the Ca2+ responses (Fig. 2Bii–iv). These data are consistent with that observed in HKH2 cells and support the conclusion that the difference between HCT116 and HKH2 cells is due to K-RasG13D ablation and not a phenotype developed as a result of HKH2 culture since their initial generation.
To test whether the enhanced Ca2+ responses observed as a result of K-RasG13D deletion in HCT116 cells was a general feature of G-protein-coupled receptor (GPCR) signalling in these cells, we examined histamine-induced Ca2+ responses, which proceed through a similar GPCR–Gq–PLC–IP3 pathway. In response to this agonist, a greater percentage of HKH2 cells exhibited Ca2+ transients, which were of a greater amplitude and AUC than those observed in HCT116 cells (Fig. 3A). These data indicated that the alteration in Ca2+ fluxes was not specific to differences in purinergic receptor signalling but was a more general effect, involving signals downstream of GPCR engagement.
To directly address whether an alteration in IP3 signalling contributed to the enhancement of ER Ca2+ release in cells lacking K-RasG13D, Ca2+ release was induced with a cell-permeant esterified form of IP3 [myo-inositol 1,4,5-trisphos-phate hexakis(butyryloxymethyl); IP3BM], which was perfused over the cells in Ca2+-free imaging buffer during the course of the experiment (Conway et al., 2006; Kasri et al., 2004; Thomas et al., 2000). Using this approach, IP3 receptors (IP3Rs) are directly engaged, bypassing GPCR, Gq, PLC and endogenous IP3. As in experiments using ATP and histamine, Ca2+ signals induced by IP3BM were also greater in HKH2 cells than in their HCT116 counterparts (Fig. 3B).
Given that K-Ras is frequently mutated in colorectal cancer, we investigated whether Ca2+ signalling was also remodelled as a result of loss of K-RasG13D in a second independent colorectal cancer cell line, DLD-1 (Shirasawa et al., 1993). As performed for HCT116 cells and their isogenic derivatives, experiments were carried out in Ca2+-free imaging buffer to restrict our analysis to Ca2+ release from the ER. ATP-induced Ca2+ fluxes were greater in the K-RasG13D-deleted DKO4 cell line (K-Ras−/WT) than in their parental isogenic DLD-1 cell line (K-RasG13D/WT) (Fig. 4). This was manifest as an increase in the percentage of responding cells and in the amplitude and AUC of the Ca2+ responses (Fig. 4ii–iv). Taken together, these data show that oncogenic K-RasG13D limits IP3-induced Ca2+ release (IICR) in both HCT116 and DLD-1 cells.
ER Ca2+ content is increased in cells deleted of K-RasG13D
Given that the magnitude of Ca2+ released from the ER is determined by its state of filling, we hypothesized that the enhanced Ca2+ release observed following K-RasG13D deletion in HKH2 cells was due to an increase in content of the IP3-releasable ER Ca2+ store. To assess ER Ca2+ levels, the magnitude of the Ca2+ mobilized from the ER by the SERCA pump inhibitor thapsigargin (Tg) was analysed. Through inhibition of SERCA, Tg reveals the non-specific Ca2+ leak from the ER causing Ca2+ accumulation in the cytosol. As store depletion with Tg also leads to Ca2+ influx across the plasma membrane, measurements were performed in Ca2+-free imaging buffer. Application of Tg induced an elevation in intracellular Ca2+ in both the HCT116 and HKH2 cell lines (Fig. 5A). The amplitude and AUC of the Tg-induced Ca2+ transient was, however, significantly greater in the HKH2 cell line compared to HCT116 cells (Fig. 5Aii,iii).
To complement these data and to accommodate for the indirect nature of using the Tg-induced elevation in cytosolic Ca2+ as a measure of ER luminal Ca2+ content, the free Ca2+ content of the ER was also measured directly using a genetically encoded GFP-based Ca2+ indicator targeted to the ER (known as D1ER) (Palmer et al., 2004). This indicator relies upon a Ca2+-dependent change in Förster resonance energy transfer (FRET) between cyan (CFP) and yellow (YFP) derivatives of GFP. D1ER was expressed in a reticular pattern and colocalized with the ER protein calnexin in both cell types, confirming its ER localization (Fig. 5Bi). In resting cells, greater FRET (the YFP:CFP ratio) was observed in the HKH2 cell line than in the HCT116 cell line, indicating higher basal Ca2+ in the ER of this cell line (Fig. 5Bii). Application of the Ca2+ ionophore ionomycin to fully deplete Ca2+ from the ER store resulted in a decline in FRET to a lower plateau, which was equivalent between the two cell types (Fig. 5Bii). The similar Ca2+-free FRET between both cell types indicated that D1ER was behaving equivalently in the two cell types. A ratio of basal FRET to Ca2+-free FRET was therefore used to normalize ER Ca2+ levels, which also indicated greater Ca2+ levels in the ER of HKH2 than HCT116 cells (Fig. 5Biii).
IP3R isoform expression is remodelled and SERCA2b expression is increased in cells deleted of K-RasG13D
Having identified that an increase in ER Ca2+ contributed to the enhanced Ca2+ signalling in K-RasG13D-deleted cells, an analysis of proteins involved in ER Ca2+ signalling was carried out. In these experiments, as we have employed elsewhere when analysing ER proteins of a high molecular mass (Drawnel et al., 2012; Harzheim et al., 2009), the ER membrane protein calnexin was used as a loading control for normalization of the protein of interest between cell types. Expression of calnexin was found to exhibit a similar expression profile between HCT116 and HKH2 cells as did two other proteins – GAPDH and β-actin – that are routinely used for normalization in immunoblotting (supplementary material Fig. S1). The expression of SERCA2b, which is primarily responsible for ER Ca2+ sequestration, was increased in HKH2 cells (Fig. 5C), whereas the expression of calreticulin, the major Ca2+ storage protein in non-excitable cells, was not altered between the two cell types (Fig. 5D). Given that SERCA3 upregulation has been reported in cancer (Brouland et al., 2005), its expression was also investigated but found to be unchanged in cells lacking K-RasG13D (supplementary material Fig. S1). Notably, the expression profile of IP3Rs was significantly different between HKH2 and HCT116 cells. Specifically, IP3R3 (also known as ITPR3) expression was increased and IP3R1 (also known as ITPR1) expression reduced in the HKH2 K-RasG13D-deleted cells when compared to HCT116 cells (Fig. 5E). IP3R2 (also known as ITPR2) expression was not detectable in either cell type (supplementary material Fig. S1).
Cells deleted of K-RasG13D exhibit increased mitochondrial Ca2+ uptake and sensitivity to apoptosis
Mitochondrial Ca2+ uptake is a low-affinity process that occurs in a privileged manner at microdomains of high Ca2+ generated by IP3Rs located at sites where the ER and mitochondria are in close proximity (Csordás et al., 2006; Duchen, 2000; Rizzuto et al., 1998; Rizzuto and Pozzan, 2006). Ca2+ flux from ER-localized IP3Rs to the mitochondria has been shown to play an important role in regulation of cell death and metabolism (Cárdenas et al., 2010; Pinton et al., 2001) – cell properties remodelled during oncogenic transformation (Hanahan and Weinberg, 2000; Hanahan and Weinberg, 2011). Given the enhancement in IP3-mediated Ca2+ signalling observed in K-RasG13D-deleted HKH2 cells, we hypothesized that mitochondrial Ca2+ uptake would also be enhanced and contribute to greater sensitivity to death-inducing stimuli in the K-RasG13D-negative cells. The effect of K-Ras deletion upon mitochondrial Ca2+ uptake during IP3-stimulated Ca2+ release from the ER was therefore analysed. To induce equivalent Ca2+ signals between cell types, and to restrict the source for mitochondrial Ca2+ sequestration to Ca2+ arising from IP3Rs, experimental conditions were used in which a maximal concentration of ATP was applied and cells were imaged in Ca2+-free imaging buffer. Mitochondrial matrix Ca2+ was measured by confocal imaging of mitochondrially compartmentalized rhod-2 AM (Fig. 6A). Cytoplasmic Ca2+ responses were detected by measuring the residual non-compartmentalized rhod-2 fluorescence in the nucleus. In this way, a mitochondrial-free region of the cell can be analysed and used as a surrogate for bulk cytosolic Ca2+ (Collins et al., 2001; Szado et al., 2008). ATP induced an increase in Ca2+ in the majority of mitochondria of both cell types, and this remained elevated for the duration of the recording (Fig. 6Bi). Although a minor difference in the percentage of responding mitochondria was observed between cell types (Fig. 6Bii), the integrated Ca2+ response of the K-Ras-deleted HKH2 cells was significantly greater than in HCT116 cells (Fig. 6Biii).
Experiments were then performed to determine whether the differences in the mitochondrial Ca2+ uptake between the two cell types were a consequence of altered IP3 signalling in ER–mitochondrial microdomains or due to an intrinsic alteration in the Ca2+ uptake properties of the mitochondria. To this end, mitochondrial Ca2+ uptake during the increase in cytosolic Ca2+ associated with store-operated Ca2+ influx was also monitored. Ca2+ influx was initiated by re-addition of Ca2+ to the imaging buffer following depletion of intracellular ER Ca2+ stores with Tg in Ca2+-free imaging buffer (Collins et al., 2001; Giacomello et al., 2010; Hanson et al., 2008b). Under these conditions, a minor difference in the percentage of responding mitochondria was observed and no difference in the integrated Ca2+ response of mitochondria was observed between HCT116 and HKH2 cells (Fig. 6C). These observations are consistent with the reported properties of Ca2+ uptake from bulk cytosol rather than from the microdomains of high Ca2+ at the ER–mitochondrial interface (Collins et al., 2001; Giacomello et al., 2010; Hanson et al., 2008b). Together these data indicated that ER–mitochondrial Ca2+ flux is enhanced as a result of K-RasG13D deletion in a manner independent of an alteration in the intrinsic Ca2+ uptake properties of the mitochondria.
As a measure of the functional consequences of enhanced ER–mitochondrial Ca2+ flux following K-RasG13D deletion, the sensitivity of HCT116 and HKH2 cells to apoptosis induced by a stimulus that acts via Ca2+ was assessed. We and others have previously shown that menadione induces apoptosis through reactive oxygen species (ROS)-dependent activation of IP3Rs and an elevation in mitochondrial Ca2+ (Baumgartner et al., 2009; Szado et al., 2008). To this end, cytochrome c loss from mitochondria and DNA fragmentation were used as hallmarks of apoptosis. Cytochrome c distribution was assessed by confocal imaging of cells immunolabelled with antibodies against cytochrome c following exposure to menadione for 20 h (Fig. 7Ai). Although untreated HKH2 and HCT116 cells displayed a typical mitochondrial distribution of cytochrome c, this mitochondrial distribution was lost following treatment with menadione and became distributed diffusely throughout the cytosol (Fig. 7Ai). HKH2 cells however exhibited a greater sensitivity to menadione treatment than HCT116 cells with cytochrome c being lost from the mitochondria in a significantly greater number of cells at 50 µM menadione (Fig. 7Aii). DNA fragmentation was assessed as the percentage of the cell population with DNA content lower than that observed in the G1 phase of the cell cycle. DNA content was determined by flow cytometric analysis of cells stained with propidium iodide (PI) (Hanson et al., 2008a). Basal levels of cell death were detected in both HCT116 and HKH2 cells (Fig. 7B). Application of menadione induced apoptosis in both HCT116 and HKH2 cells. However, the percentage of the menadione-treated cell population with DNA content lower than in G1 phase was significantly greater in the K-RasG13D-deleted cells than in their HCT116 counterparts (Fig. 7B). Caspase activation, a further hallmark of apoptosis, was also observed following menadione treatment in HCT116 and HKH2 cells by imaging (Fig. 7Ai) and immunoblotting (supplementary material Fig. S2).
Taken together, these data indicate that decreased flux of Ca2+ to the mitochondria contributes to the oncogenic phenotype of HCT116 cells.
The impact of oncogenic K-Ras on Ca2+ signals, particularly in the context of oncogenic transformation is poorly understood. Here, we provide the first demonstration of an interaction of natively expressed oncogenic K-Ras with Ca2+ signalling and how this signalling crosstalk might affect cell fate. By comparing isogenic colon cancer cell line pairs expressing either a single copy of mutant K-RasG13D or no mutant K-Ras we have determined that K-RasG13D deletion enhances IP3-dependent Ca2+ signals and ER–mitochondrial Ca2+ flux and that this sensitizes cells to pro-apoptotic stimuli. From these data, we propose that suppression of IP3 signalling from the ER and Ca2+ uptake by the mitochondria contributes to the pro-survival properties of K-RasG13D associated with the oncogenic phenotype.
Cytosolic Ca2+ signals are generated by Ca2+ entry across the plasma membrane, Ca2+ release from intracellular stores or a combination of both (Berridge et al., 2003; Berridge et al., 2000; Bootman et al., 2003). Through manipulation of these Ca2+ signalling pathways in transformed cells, specific roles for each of these Ca2+ sources in controlling aspects of cancer cell biology including regulation of cell proliferation, migration and death have been described (Crépin et al., 2007; Humez et al., 2004; Legrand et al., 2001; Lipskaia et al., 2009; Szatkowski et al., 2010; Yoshida et al., 2012). Altered expression of a number of Ca2+-handling proteins in tumour tissue has also been determined (Arbabian et al., 2012; Korosec et al., 2006; Monteith et al., 2012; Monteith et al., 2007; Motiani et al., 2013). Although these findings are suggestive of an important role of certain Ca2+ signalling pathways in transformed cells, causality has not been demonstrated. However, somatic mutations in the gene encoding SERCA have been identified in patients with colon cancer leading to the proposal that altered Ca2+ signalling predisposes to oncogenic transformation (Korosec et al., 2006). Cancers arise through mutations in oncogenes such as KRAS or tumour suppressors that serve to promote or suppress the activity of the proteins they encode, respectively (Hanahan and Weinberg, 2000). How the native expression of a specific oncoprotein in transformed cells affects Ca2+ signalling and whether this contributes to the phenotype of the transformed cell is, however, not clear. A particular issue when investigating Ca2+ signalling pathways is that the analysis of Ca2+ signalling is only possible in live cells. The availability of appropriate controls for the cell line expressing the driving oncogene is also essential (Roderick and Cook, 2008). For analysing the effects of activated Ras isoforms, this is a particular concern because oncogenic K-Ras can induce senescence or cell proliferation depending on the level of overexpression (Serrano et al., 1997; Tuveson et al., 2004). These issues are minimized through the use of isogenic cell line pairs in which studies are performed upon a cancer cell line harbouring a single allele of an activating oncogene and a second cell line in which the driving oncogene has been deleted by homologous recombination (Shirasawa et al., 1993). By comparing hormone-agonist-induced Ca2+ signalling between such pairs of cell lines, we found that loss of K-RasG13D enhanced cytosolic Ca2+ signals. The elevated Ca2+ responses in K-RasG13D-deleted cells persisted in Ca2+-free imaging buffer, indicating that Ca2+ release from the ER was important in defining the properties of the Ca2+ response. Importantly, Ca2+ signals were greater in two different colorectal cancer cell lines in which K-RasG13D had been deleted (HKH2 and DKO4), as well as in HCT116 cells when K-Ras expression was reduced by siRNA, indicating that suppression of hormone-stimulated Ca2+ signalling is a common response to K-RasG13D in colorectal cell lines.
The greater magnitude of hormone-induced Ca2+ release from the ER in K-RasG13D-deleted cells could have arisen through a number of mechanisms. For example, by increased GPCR expression or coupling to downstream effectors, modification in inositol phosphate metabolism, changes in IP3R expression or through a greater Ca2+ content of the ER. Given its pleiotropic nature, we speculated that Ras could interfere with any or all of these processes. Increased PLC activity and IP3 levels have been reported in a number of transformed cell lines and in breast, ovarian and colonic carcinoma, suggesting that basal signalling is enhanced as a result of transformation (Weber, 2005). Oncogenic K-Ras might also increase signalling activity through stimulating PLCε, which in turn, by increasing IP3 levels, would promote Ca2+ release from stores (Kelley et al., 2001). However, our observations that Ca2+ signals are negatively correlated with Ras abundance suggests that enhanced basal signalling possibly involving PLCε does not contribute to Ras-mediated regulation of Ca2+ signalling in cells harbouring mutated K-RasG13D. As Ca2+ signals induced by stimulation of the histamine receptor, another GPCR, were also greater in K-RasG13D-deleted cells, it is unlikely that modifications in purinergic receptor expression contributed to the effects of K-RasG13D deletion. Similarly, Ca2+ responses induced by a cell-permeant analogue of IP3 were greater in K-RasG13D-deleted cells than in the isogenic parental cell line. As this cell-permeant IP3 directly activates IP3Rs, circumventing GPCRs, G proteins and PLC, our data indicate that native levels of K-RasG13D has a direct effect on IP3R-mediated Ca2+ release from the ER.
Notably, although the combined expression of IP3R isoforms was unaltered by K-RasG13D deletion, the relative abundance of the expressed IP3R subtypes was altered in the K-RasG13D-deleted cells. In particular, in K-RasG13D-deleted cells, IP3R3 expression was increased and IP3R1 expression suppressed, indicating that in colorectal cancer cell lines, K-RasG13D represses IP3R3 expression. Changes in the expression of IP3Rs in cancer cells have been reported previously. Most notably, an increase in IP3R3 expression at the mRNA level has been detected in a recent microarray analysis of K-Ras-deleted cells lines (Vartanian et al., 2013). In gastric cancer cells, an increase in IP3R3 was observed in the ascites, but not in cancer cells established from primary tumours. In the ascites, IP3R3 inhibition by 2-aminoethoxydiphenyl borate (2-APB) induced apoptosis (Sakakura et al., 2003). IP3R3 expression is also increased in MCF-7 cells induced to proliferate with estradiol (Szatkowski et al., 2010). An increase in IP3R2 expression together with K-Ras has been observed in non-small cell lung cancer (NSCLC) cells (Heighway et al., 1996). Given the differing properties of each IP3R isoform, this change in isoform composition following K-Ras deletion might have important consequences. As different IP3R isoforms are regulated differently by IP3 and Ca2+, giving rise to distinct Ca2+ signalling fingerprints (Hattori et al., 2004; Miyakawa et al., 1999), the change in the relative abundance of each IP3R isoform could contribute to the differences in Ca2+ signalling observed between the two cell types. A notable feature of IP3R3 is that it is least sensitive to Ca2+-dependent inhibition (Hagar et al., 1998). As a result, whereas expression of IP3R1 supports regular Ca2+ oscillations, monophasic Ca2+ transients are observed in IP3R3-expressing cells (Almirza et al., 2010; Hattori et al., 2004; Miyakawa et al., 1999). Indeed, siRNA reduction in IP3R3 expression in MCF-7 breast cancer cells transformed Ca2+ responses from a peak–plateau to a more oscillatory profile (Szatkowski et al., 2010). These different Ca2+ signatures probably allow signalling from each receptor to participate in different cell fate choices. Ca2+ oscillations arising from IP3R1 might be optimized for controlling cytokinesis (Kittler et al., 2004) and gene expression (Dolmetsch et al., 1998), whereas the sustained Ca2+ signals arising from IP3R3 might promote cell death (Blackshaw et al., 2000; Khan et al., 1996; Mendes et al., 2005; Szatkowski et al., 2010). Thus, a potential outcome of these different Ca2+ signatures is that the Ca2+ oscillations that arise from IP3R1 in HCT116 cells sustain their rapid proliferation, whereas the reduction in IP3R3 protects the cells from cytotoxic Ca2+ signals.
An increase in ER Ca2+ store content was also observed in K-RasG13D-deleted cells. As the Ca2+ content of the ER is a dominant determinant of the magnitude of the IP3-stimulated Ca2+ transient (Berridge, 2006; Caroppo et al., 2003), it is likely that this alteration in ER Ca2+ store content contributes substantially to the enhancement in Ca2+ signalling observed in K-RasG13D-deleted cells. Our data are consistent with the view that a relatively low level of Ca2+ in the ER offers an advantage to the transformed cell (Bergner and Huber, 2008). Specifically, by limiting Ca2+ release, the effect of stimuli that serve to induce apoptosis is diminished. The cytotoxic effects of Ca2+ release from the ER are mediated through accumulation in mitochondria, which results in permeability transition and activation of apoptotic pathways (Pinton et al., 2008; Roderick and Cook, 2008). Elevated Ca2+ also activates DNA endonucleases, promotes phosphatidylserine exposure, leads to cellular ATP depletion, and increases ROS and ER stress (Orrenius et al., 2003). Despite this well-accepted view of the benefit of lower ER Ca2+ for survival of transformed cells, much of the supporting data has emerged through investigation into the mechanism of actions of proteins involved in regulation of cell death pathways that are dysregulated in cancer. For example, enhanced expression of Bcl-2 family members or knockout of BH3-only pro-apoptotic proteins (which result in increased abundance of the anti-apoptotic family members) result in a lowering of free ER Ca2+ levels and reduced flux of Ca2+ to the mitochondria (Pinton et al., 2008; Roderick and Cook, 2008). By increasing leak through the IP3R or suppressing their activity, Bcl-2 family members also reduce IP3-induced Ca2+ release (Chen et al., 2004; Oakes et al., 2005). No differences in the expression of Bak, Bax, Bcl-2, Bcl-XL or Mcl-1 was detected between the HCT116 and HKH-2 cell types analysed here (data not shown). Notably, when analysed in HCT116 cells, no prominent role for Bax in mitochondrial outer membrane permeabilization (MOMP) was detected (De Marchi et al., 2004), supporting the hypothesis that Ca2+ signalling remodelling by oncogenic K-Ras activation in HCT116 cells does not involve alterations in the expression of Bcl-2 family members. IP3R activity is also reduced by phosphorylation by the pro-survival kinase PKB/Akt (Khan et al., 2006; Marchi et al., 2012; Szado et al., 2008); a kinase that is increased in activity in many cancers.
A decrease in ER Ca2+ content has been detected in a subset of lung cancer cell lines when compared to normal human bronchial epithelial cells (Bergner et al., 2009). Consistent with the reduction in ER Ca2+ in cancer, a reduction in expression of SERCA pump has been observed in cancer-derived cell lines and in tumours. Mutations in SERCA that result in loss of expression or activity have also been detected in tumours (Monteith et al., 2007). The importance of SERCA expression is also demonstrated by the induction of squamous cell cancers in SERCA2b haploinsufficient mice (Prasad et al., 2005). In humans, however, loss of one SERCA allele results in Darrier's disease, which is characterized by a skin phenotype (Hovnanian, 2007). In line with these studies, SERCA2b expression was significantly increased in K-RasG13D-deleted cells, showing that a reduction in SERCA2b activity contributes to the phenotype of K-Ras-transformed cells. The increase in ER Ca2+ in K-RasG13D-deleted cells might also be explained by the observed reduction in IP3R1, which has been reported to contribute to the Ca2+ leak from the ER (Kasri et al., 2006; Oakes et al., 2005). A positive correlation between ER Ca2+ levels and proliferation has been observed in prostate cancer cell lines (Legrand et al., 2001). However, because depletion of the ER Ca2+ store inhibits proliferation and induces cell death in transformed cells, it would be important to correlate the absolute Ca2+ content of the ER and cell proliferation in these studies.
The mitochondria are an important target of Ca2+ released from the ER (Rizzuto et al., 2012). Mitochondrial Ca2+ uptake is a low-affinity process. As such, mitochondria preferentially accumulate Ca2+ at sites of close apposition with the ER called mitochondrial-associated membranes (MAMs), which are enriched in Ca2+ release channels including IP3Rs (Csordás et al., 2010; Hajnóczky and Csordás, 2010; Rizzuto et al., 2004). Through this preferred pathway, mitochondrial function, including metabolism and induction of apoptotic cell death, is acutely modulated by IP3-mediated Ca2+ signals. Here, we show the first evidence of modifications in mitochondrial Ca2+ uptake downstream of endogenous oncogenic K-Ras. The ablation of oncogenic K-RasG13D increased the accumulation of Ca2+ in the mitochondria following Ca2+ release from the ER. Notably, the difference between the HCT116 and HKH2 cells was lost when Ca2+ uptake during Ca2+ influx from the extracellular space was analysed. Under these conditions, Ca2+ uptake into the mitochondria is not restricted to the IP3R-containing MAMs. As such, microdomains of high Ca2+ at the ER–mitochondrial interface do not drive mitochondrial Ca2+ sequestration and the uptake observed is due to the properties of the mitochondrial uptake mechanisms alone (Collins et al., 2002; Rizzuto and Pozzan, 2006; Szabadkai and Duchen, 2008). Our data therefore suggest that the enhancement of ER–mitochondrial Ca2+ flux following K-RasG13D ablation is through modification of IICR. Consistent with increased IICR, this enhanced ER-to-mitochondria Ca2+ flux in HKH2 cells was mirrored by an increased sensitivity of these cells to Ca2+-induced cell death. The increase in the ER-to-mitochondria Ca2+ transfer observed in HKH2 cells indicates that the expression of K-RasG13D acts to reduce this flux in HCT116 cells. In line with their lower ER–mitochondrial Ca2+ flux and apoptosis, the expression of IP3R3, which has been proposed to specifically mediate pro-apoptotic Ca2+ fluxes at the MAM, is also reduced in HCT116 cells (Blackshaw et al., 2000; Khan et al., 1996; Mendes et al., 2005). More recently, the tumour suppressor PML, which is localized at the ER, has been reported to specifically mediate the dephosphorylation of IP3R3 by PP2a (Giorgi et al., 2010). Notably, the reintroduction of ER-targeted PML in PML−/− cells restored the sensitivity to Ca2+-dependent apoptosis (induced by Menadione and H2O2), but not that to Ca2+-independent apoptosis (induced by the DNA-damaging agent Etoposide) (Giorgi et al., 2010). This finding is consistent with the increased sensitivity to menadione observed in the K-RasG13D-ablated HKH2 cells.
In conclusion, our data describes for the first time the alterations in Ca2+ regulation driven by a single oncogenic K-RasG13D allele in colorectal cells. The enhancement of Ca2+ handling, mitochondrial sequestration and cell death as a result of loss of K-RasG13D in two isogenic models indicates that suppression of Ca2+ signalling is a common response to K-RasG13D. Owing to its pleiotropic actions, modulation by K-RasG13D in colorectal cells is likely also to contribute to other aspects of cell physiology that serve to promote cell transformation including enhancement of cell proliferation or modulation of metabolism. The importance of Ras activation and downstream pathways in cancers where mutated Ras is not the primary cause, such as mutations in EGFR or B-Raf, raises the possibility that the modulation of Ca2+ fluxes observed in this study due to K-RasG13D is a common feature of many cancers and is thus a target for intervention.
MATERIALS AND METHODS
HCT116 and DLD-1 cells (both K-RasG13D/WT) and their respective isogenic derivatives HKH2 and DKO4 (both K-Ras-/WT) were a kind gift of Senji Shirasawa (Fukuoka University, Japan) and have been previously described (Shirasawa et al., 1993). Cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Life Technologies, Carlsbad, CA, USA), containing 10% heat-inactivated fetal bovine serum (FBS) (Invitrogen), 1% penicillin/streptomycin solution (5 units penicillin, 55 µg streptomycin) (Sigma, Dorset, UK). Cells were maintained at 37°C under 5% CO2 in saturated humidity and were passaged upon reaching 80–90% confluency. Coverslips were coated with poly-L-lysine prior to seeding of cells.
siGENOME SMART Pool for K-Ras and siGENOME non-targeting control oligonucleotides (Dharmacon, Thermo) were reverse transfected using Dharmafect-2 transfection reagent (Dharmacon, Thermo) according to the manufacturer's instructions. Briefly, 2×105 cells in a 12-well dish or 4×105 cells in a 6-well dish were transfected with siRNAs at a final concentration of 25 nM. The medium overlying the cells was exchanged after 24 h, and protein expression measured and Ca2+ imaging experiments performed after a further 24 h.
Imaging of cytosolic Ca2+
Cytosolic Ca2+ was imaged as previously described (Peppiatt et al., 2003). Briefly, cells were seeded onto poly-L-lysine-coated coverslips at equivalent densities and imaged after 48 h. Prior to each experiment, coverslips were mounted into stainless steel imaging chambers and loaded with fura-2 AM (Life Technologies; 2 µM for 30 min, followed by de-esterification in imaging buffer for a further 30 min). Coverslips were imaged on the stage of a Nikon Eclipse TE200 inverted epifluorescence microscope equipped with a Nikon PlanFluor 20×/0.75 NA multi immersion objective (Nikon, Kingston Upon Thames, Surrey, UK). Excitation light at 340 and 380 nm was selected using a motorized filter wheel (Sutter Industries, Novato, CA, USA) at a frequency of 1 image pair every 3 s with an exposure of 200 ms, and emitted light was selected using a 400 nm dichroic mirror and filtered through a 460 nm long pass filter. Images were captured using a Hamamatsu ORCA ER charge-coupled device (CCD) camera. Three coverslips per cell type were imaged per day on three separate days, each coverslip containing at least 50 cells. Ca2+ concentration was calculated according to Grynkiewicz et al. (Grynkiewicz et al., 1985).
Imaging of ER Ca2+
The FRET-based, genetically-encoded D1ER Ca2+ indicator was a kind gift of Amy Palmer (University of Colorado, Boulder, USA). The affinity of the indicator for Ca2+ has been determined to be 60 µM, allowing its successful use to monitor resting and dynamic changes in ER luminal [Ca2+] (Palmer et al., 2004). In experiments involving D1ER, cells were seeded as indicated for ratiometric imaging but transfected with the D1ER construct after 24 h using JetPei (PolyPlus Transfection, Ilkirch, France) according to manufacturer's specification. Cells were imaged at 24 h post transfection. Coverslips were mounted in stainless steel chambers and imaged on the stage of an Olympus IX81 inverted microscope equipped with an Olympus UPlanSApo 20×/0.75 NA air objective. Excitation light at 435/10 nm was selected using a Polychrome V monochromator (Olympus, Southend-on-Sea, UK). Emitted fluorescence of CFP and YFP was simultaneously captured using a Cairn Optosplit II image splitter (Cairn Research Limited, Graveney Road, Faversham Kent). The image splitter unit was configured with a 515 nm dichroic mirror, which reflected the emitted fluorescence of CFP (further filtered through a 485/40 nm band-pass filter), and passed the emission of YFP (further filtered through a 535/30 nm band-pass filter).
Imaging of mitochondrial Ca2+
Mitochondrial Ca2+ was imaged as previously described using rhod-2 AM as a Ca2+ indicator (Szado et al., 2008). Prior to each experiment, cells were loaded with rhod-2 AM (4 µM for 30 min at room temperature followed by de-esterification in imaging buffer for a further 30 min). Cells were imaged using a VisiTech VoxCell Scan spinning disc confocal configured on a Nikon TE2000 microscope equipped with a Nikon 60×/1.25 NA oil immersion objective. Rhod-2 was excited by illumination with the 568 nm line of an argon/krypton laser. Emitted fluorescence was filtered through a 575/50 nm band-pass filter. Images were captured using a Hamamatsu ORCA ER CCD camera controlled by the Visitech Voxcell Scan software. Ca2+ concentration was calculated as previously described (Collins et al., 2001).
Cells were harvested 48 h post seeding and protein lysate was quantified using a bicinchoninic acid (BCA) protein assay kit (Thermo Scientific). An equivalent amount of each sample (15 to 50 µg) was loaded onto 7% self-poured polyacrylamide gels or onto 4–12% gradient pre-cast gels (NUPAGE; Life Technologies). Proteins were transferred from the gels onto polyvinilidene fluoride (PVDF) membranes (for IP3Rs and SERCAs and their loading controls) or nitrocellulose. Non-specific protein-binding sites were first blocked by incubation for 1 h in TBS containing 0.05% Tween 20 (TBS-T) and 5% milk. Membranes were subsequently probed with primary antibodies (diluted as indicated below in TBS-T milk) for 1 h at room temperature or 4°C overnight. Details of primary antibodies are as follows: anti-K-Ras (dilution 1∶500, AbdSerotec); anti-SERCA2b [dilution 1∶1000, kind gift of Frank Wuytack, University of Leuven (Wuytack et al., 1989)]; anti-IP3R3 (dilution 1∶1000, BD Biosciences); anti-IP3R1 [dilution 1∶1000, in-house generated (Kasri et al., 2004)]; anti-calnexin (dilution 1∶20,000, Sigma); anti-SERCA3 [dilution 1∶1000, gift of Frank Wuytack (Wuytack et al., 1994)]; anti-IP3R2 [dilution 1∶500, in-house generated (Harzheim et al., 2009)]; anti-calreticulin (1∶1000; Roderick et al., 1997); anti-active caspase 3 (1∶1000, BD BioSciences); anti-GAPDH (1∶5000, Sigma) and anti-β actin (1∶5000, AbCam). Excess antibodies were removed by washing with TBS-T. Membranes were then probed with either horseradish peroxidase (HRP)-conjugated (Jackson Immunoresearch; 1∶10,000 dilution) or fluorescently-labelled secondary antibodies (Life Technologies and LI-COR; both at 1∶5000). All membranes were then washed in five exchanges of TBS-T and one of TBS before band detection. HRP-conjugated antibodies were detected by chemiluminescence (ECL) (Thermo Scientific) and subsequent exposure to film, and fluorescently-labelled secondary antibodies (Life Technologies and LI-COR) were detected by digital scanning (LI-COR Odyssey). For band quantification, intensity values were obtained either through analysis of digitized film (ImageJ, for ECL detection) or Image Studio (for LI-COR detection). Bands of the protein of interest were normalized against the corresponding loading control band. Following normalization, protein abundance in the experimental conditions of interest was normalized to the control.
Immunofluorescence was performed as previously described (Higazi et al., 2009). Briefly, at 48 h post seeding, cells were fixed with fixation buffer (2% paraformaldehyde, 0.05% glutaraldehyde in PBS) and permeabilized with 0.2% Triton X-100 in PBS. After incubation in blocking solution (0.1% Triton X-100, 5% donkey goat serum diluted in PBS), cells were probed with primary antibodies (diluted in blocking solution) for 1 h at room temperature. Details of primary antibodies are as follows: anti-cytochrome c (dilution 1∶200, Santa Cruz Biotechnology); and anti-active caspase 3 (1∶200, BD BioSciences); anti-calnexin (1∶500, Sigma). After removal of excess antibodies by washing in 0.1% Triton X-100 in PBS, cells were incubated with Alexa-Fluor-labelled secondary antibodies (Life Technologies) for 1 h at room temperature. Excess antibody was subsequently removed with five washes in 0.1% Triton X-100 in PBS and two washes in PBS. Coverslips were mounted in Vectashield containing DAPI, which also counterstained nuclei. Cells were imaged by point-scanning confocal microscopy using appropriate laser lines for excitation of the dyes (Olympus FV1000 confocal configured on an Olympus IX81 inverted microscope using a 60×/1.35 NA oil immersion objective for calnexin and YFP imaging, and a Nikon A1R confocal configured on a Nikon Ti inverted microscope and using 60×/1.4 NA oil immersion objective for imaging of cytochrome c and activated caspase). Images were processed and analysed using Image J.
Cell cycle analysis
Cells in the medium were collected and then pooled with cells that remained attached to their substrate that were harvested by trypsinization. After washing in PBS, cells were fixed with 70% ethanol prior to RNase treatment and staining with propidium iodide (PI). Stained cells were analysed with a Becton Dickinson FACSCalibur flow cytometer (Oxford, UK). Single cells in suspension were excited at 488 nm by an argon laser and analysed according to the intensity of emitted fluorescence through a 585/42 band pass filter (Hanson et al., 2008a).
Menadione treatment of cells
At 24 h post seeding, cells were exposed to menadione diluted in culture medium at concentrations between 25 and 100 µM. Control samples cultured in parallel were also analysed. Cells were harvested 20 h after exposure to menadione and processed for flow cytometric analysis or immunofluorescence.
Where data was compared to normalized control a one-sample Student’s t-test was employed. Other experiments were analysed by Student's t-test or two-way ANOVA. Significance was accepted at P<0.05.
We are grateful to Stuart Conway, University of Oxford for the IP3BM, Senji Shirasawa, Fukuoka University, Japan for the cell lines, Amy Palmer, University of Colorado, USA for the D1ER FRET construct and Frank Wuytack, KU Leuven for the SERCA antibodies. We would also like to acknowledge Anne Segonds-Pichon for assistance with statistical analysis, Rachel Walker for flow cytometry and Simon Walker for imaging.
C.P., T.C.F. and H.L.R. performed experiments and analysed data. C.P., M.D.B., S.J.C. and H.L.R. conceived of the study and interpreted data. C.P., S.J.C. and H.L.R. prepared the manuscript.
C.P. was funded by the Aldobrandini Studentship of St John's College, Cambridge, UK. The H.L.R. and S.J.C. laboratories are supported by the Babraham Institute, the Biotechnology and Biological Sciences Research Council (BBSRC) [Epigenetics and Signalling ISPGs (Institute Strategic Programme Grant)]. T.C.F. was funded by a Research Experience Placement grant from the University of Cambridge BBSRC Doctoral Training Programme DTP. H.L.R. was also supported by a Research Fellowship from the Royal Society during the course of this work. Deposited in PMC for immediate release.
The authors declare no competing interests.