An unbiased genomic screen now revealed that the loss of either of two VRAC subunits, LRRC8A or LRRC8D, increased resistance against carboplatin and cisplatin. This finding can be largely explained by an unsuspected role of VRAC in drug transport. This transport required the obligatory channel subunit LRRC8A and also depended on LRRC8D, a subunit that strongly increased VRAC's permeability to cisplatin/carboplatin. Exposure to staurosporine revealed an additional, drug uptake‐independent, but possibly AVD‐related effect of LRRC8A on drug‐induced apoptosis. Moreover, LRRC8D increased VRAC's permeability to taurine, and cells lacking this subunit showed reduced cell volume regulation. Our work uncovers a novel role of VRAC in cisplatin and carboplatin uptake and resistance, identifies LRRC8A/LRRC8D‐containing VRACs as physiologically important organic osmolyte channels, and reveals differences in substrate selectivity and pharmacology between differently composed VRACs.

In cultured cells, the induction of apoptosis by cisplatin and other drugs such as staurosporine is characterized by an early cell shrinkage denominated apoptotic volume decrease (AVD) which is followed by other apoptotic hallmarks like the activation of caspases and DNA fragmentation. AVD and the induction of apoptosis could be inhibited by blockers of the ubiquitously expressed volume‐regulated anion channel VRAC (also known as VSOR or VSOAC) (Maeno et al , 2000 ; Ise et al , 2005 ; Okada et al , 2006 ; Lang & Hoffmann, 2012 ). Being closed under resting conditions, VRAC opens upon cell swelling and releases chloride and organic osmolytes in the course of regulatory volume decrease (RVD). The effect of VRAC blockers on drug‐induced apoptosis was attributed to a loss of VRAC‐dependent AVD (Maeno et al , 2000 ). However, the blockers used to substantiate a role of VRAC in apoptosis are all non‐specific, and the notion that AVD facilitates apoptosis is controversial (Orlov et al , 2013 ). VRAC has been characterized biophysically and physiologically for decades, but the failure to identify the underlying proteins (Okada, 1997 ; Pedersen et al , 2015 ) precluded conclusive genetic and biochemical studies. Only recently, LRRC8 heteromers were identified as essential VRAC components. LRRC8A is an obligatory VRAC subunit (Qiu et al , 2014 ; Voss et al , 2014 ), but it needs at least one other LRRC8 homolog (LRRC8B, LRRC8C, LRRC8D, or LRRC8E) to mediate volume‐activated anion currents (I Cl,vol ) or to release organic osmolytes like taurine (Voss et al , 2014 ). A weak homology to pannexins suggests that LRRC8 proteins may assemble to hexameric channels (Abascal & Zardoya, 2012 ). The formation of different LRRC8 heteromers indicates that there are several different VRACs that may vary in properties and tissue distribution. For instance, the subunit composition of LRRC8 heteromers might specify the channel's preference for chloride over taurine or vice versa. Such a scenario would support the contentious notion (Lambert & Hoffmann, 1994 ; Stutzin et al , 1999 ; Shennan, 2008 ) that VRAC is molecularly distinct from a postulated volume‐stimulated organic osmolyte/anion channel VSOAC (Jackson & Strange, 1993 ). However, the impact of specific LRRC8 subunits on VRAC's selectivity has not yet been investigated.

The platinum‐containing drugs cisplatin, carboplatin, and oxaliplatin are among the most successful drugs used for treating cancer (Kelland, 2007 ). Pt drugs introduce covalent adducts in DNA which eventually cause cell death. Tumor cells treated with therapeutic doses of Pt drugs probably die predominantly by mitotic catastrophe/necrosis (Borst et al , 2001 ; Brown & Attardi, 2005 ). Attempts to link Pt drug sensitivity to apoptosis remain prominent (Speirs et al , 2011 ), however, even though the induction of apoptosis by high drug concentrations has been shown to be due to a cytoplasmic off‐target effect of Pt drugs (Mandic et al , 2003 ; Berndtsson et al , 2007 ; Fayad et al , 2009 ).

Results

A genomewide screen identifies LRRC8A and LRRC8D as mediators of carboplatin and cisplatin resistance To identify genes affecting platinum drug sensitivity, we performed genomewide loss‐of‐function screens of haploid KBM7 cells that were subjected to insertional mutagenesis using a gene‐trap virus (Carette et al, 2009) (Fig 1A). Cells resisting exposure to 7 μM carboplatin for three weeks showed a significant enrichment of insertions in the LRRC8D and LRRC8A genes (Fig 1B and Appendix Table S1). Most sense insertions were localized in introns upstream of the ORF‐containing exon 3 (Fig 1C). These positions are consistent with gene‐inactivating mutations. Two LRRC8D‐deficient KBM7 clones, which display reduced blasticidin S uptake (Lee et al, 2014) and which lack the LRRC8D protein (Fig EV1), were indeed more resistant to both carboplatin and cisplatin, but not to the larger compound oxaliplatin (Fig 1D and Appendix Table S2). Likewise, KBM7‐derived haploid HAP1 cells (Carette et al, 2011) were resistant to carboplatin and cisplatin, but not to oxaliplatin, when LRRC8A or LRRC8D was disrupted (Fig EV2). Hence, two genes, LRRC8A and LRRC8D, affected carbo‐ and cisplatin sensitivity at drug concentrations that can be achieved in patients. Figure 1.Loss‐of‐function screen using haploid KBM7 cells identifies LRCC8A and LRRC8D as determinants of carboplatin resistance Outline of the loss‐of‐function screen. Genes enriched for gene‐trap insertions in a carboplatin‐selected cell population compared to unselected control cells. Circles represent genes and their size corresponds to the number of independent insertions identified in the carboplatin‐selected population. Genes are ranked on the x‐axis based on chromosomal position. Location of gene‐trap insertion sites (red arrowheads). White boxes indicate the 5′‐ and 3′‐untranslated regions, and the black boxes show the coding sequence in exons 3 and 4 (LRRC8A) and exon 3 (LRRC8D). Loss of LRRC8D causes resistance to carboplatin and cisplatin, but not to oxaliplatin. Survival of parental, vector‐transduced, or LRRC8D‐deficient GT1 and GT2 KBM7 cells exposed for 96 h to increasing concentrations of cisplatin, carboplatin, and oxaliplatin. The corresponding IC 50 values and 95% confidence interval (CI) are given in Appendix Table S2. Data are presented as mean ± SEM. Click here to expand this figure. Figure EV1.LRRC8 subunit expression in different cell lines A–C. Cl,vol at clamped voltages (Fig et al, 2014 LRRC8D1−/− and LRRC8D2−/− denote two independent HEK and HCT116 knockout clones. Western blot showing the expression of all LRRC8 subunits in HAP1 and KBM7 (A), HEK (B), and HCT116 (C) cell lines, including knockout cell lines. Tubulin or actin was used as loading control. Note that KBM7 cells virtually lack LRRC8E, explaining the lack of inactivation of their Iat clamped voltages (Fig 3 B). Notice that disruption of LRRC8A changes the apparent sizes of the other LRRC8 subunits (prominently seen for LRRC8D in HCT cells) because LRRC8B through E need LRRC8A to leave the ER (Voss) and are therefore not fully glycosylated in its absence.anddenote two independent HEK and HCT116 knockout clones. Click here to expand this figure. Figure EV2.Increased resistance of LRRC8A−/− and LRRC8D−/− HAP1 cells to carboplatin and cisplatin, but not to oxaliplatin A–C. Clonogenic growth of LRRC8A− and LRRC8D− or WT HAP1 cells treated with carboplatin, cisplatin, or oxaliplatin. Cells were exposed to the indicated concentrations of carboplatin (A), cisplatin (B), or oxaliplatin (C) for 7 days. Surviving colonies were formalin‐fixed and stained with crystal violet. The optical absorption was determined at 590 nm after extracting the dye with 10% acetic acid. Data are presented as mean ± SEM (n = 6). CI, confidence interval.

Low LRRC8D expression correlates with reduced survival of Pt drug‐treated ovarian cancer patients To determine whether LRRC8A or LRRC8D expression might affect chemotherapy in patients, we examined The Cancer Genome Atlas (TCGA) data collection of ovarian cancer patients who were treated with platinum drugs. We analyzed the survival of patients with a low tumor expression of LRRC8A or LRRC8D versus the remaining patients. We used the lower tertile of the distribution of LRRC8A and LRRC8D expressions as cutoff. Whereas low LRRC8A expression had no influence on survival (Fig 2A), patients with a low LRRC8D gene expression in their ovarian cancers displayed a significantly reduced survival (Fig 2B). Most patients had also received taxane, but disruption of LRRC8A or LRRC8D did not provide resistance against docetaxel (Appendix Fig S1). To corroborate these results, we investigated the data derived from ovarian cancer patients that were recently published by Patch et al (2015). Although the available data are derived from fewer patients, also in this analysis a low expression of LRRC8D, but not LRRC8A, correlated with a modest, but significant decrease in survival (Fig 2C and D). Thus, LRRC8D might also affect platinum drug responses in cancer patients. Figure 2.Low expression of LRRC8D but not LRRC8A correlates with shorter survival of high grade serous ovarian cancer patients treated with platinum‐based drugs A–D. LRRC8A (A, C) or LRRC8D (B, D) gene expression as extracted from the TCGA database (et al ( 2015 P‐values were determined using the log‐rank test. Differential survival based on(A, C) or(B, D) gene expression as extracted from the TCGA database ( http://cancergenome.nih.gov/ ) (A, B) or using the data from Patch) (C, D). As cutoff the lower tertile of LRRC8A or LRRC8D gene expression was used.‐values were determined using the log‐rank test.

Pt drug resistance is not strictly correlated with lack of VRAC currents or volume regulation LRRC8A and LRRC8D are subunits of the volume‐regulated anion channel VRAC. The finding that the loss of LRRC8D conferred drug resistance was surprising because this subunit, unlike LRRC8A, is dispensable for VRAC Cl− currents (I Cl,vol ) in HCT116 cells (Voss et al, 2014), a finding we now confirmed for haploid HAP1 and KBM7 cells (Fig 3A and B, Appendix Fig S2). Hence, Pt drug resistance is not correlated with a loss of VRAC Cl− channel activity. Figure 3.Disruption of LRRC8A, but not of LRRC8D, abolishes I Cl,vol and blocks volume regulation A, B. Cl,vol ) of the HAP1 (A) and KBM7 (B) haploid cell lines. Left panels, example current traces of I Cl,vol fully activated by hypotonic cell swelling measured with the voltage‐clamp protocol shown in (A). Dashed lines indicate zero current. Right panels, averaged current/voltage relationships of maximally activated I Cl,vol . Consistent with VRAC currents, they needed hypotonic swelling for activation, displayed an I − > Cl − permeability sequence, and were blocked by DCPIB ( et al , 2014 n = 5–10. VRAC currents (I) of the HAP1 (A) and KBM7 (B) haploid cell lines. Left panels, example current traces of Ifully activated by hypotonic cell swelling measured with the voltage‐clamp protocol shown in (A). Dashed lines indicate zero current. Right panels, averaged current/voltage relationships of maximally activated I. Consistent with VRAC currents, they needed hypotonic swelling for activation, displayed an I> Clpermeability sequence, and were blocked by DCPIB ( Appendix Fig S2A–H ). The difference in current inactivation between HAP1 and KBM7 cells can be explained by the fact that KBM7 cells hardly express LRRC8E (Fig EV1 ) which accelerates VRAC inactivation (Voss). At potentials > +100 mV, also KBM7 currents inactivated ( Appendix Fig S2I ). Data are presented as mean ± SEM;= 5–10.

C. Dependence of regulatory volume decrease (RVD) of HEK cells on LRRC8 genes. Cells were exposed to hypotonic medium starting at t = 0, and intracellular calcein fluorescence was followed over ˜1 h as semiquantitative measure of cell volume. Data are presented as mean values ± SEM from sixteen wells. In view of the postulated role of VRAC activation in cisplatin‐induced cell shrinkage and subsequent apoptotic cell death (Maeno et al, 2000; Okada et al, 2006; Lang & Hoffmann, 2012), we explored the role of LRRC8A and LRRC8D in regulatory volume decrease (RVD) that can be determined more reliably than AVD. Like I Cl,vol (Voss et al, 2014), RVD depended on LRRC8 heteromers because it was similarly impaired in LRRC8A−/− and LRRC8(B,C,D,E)−/− cells that lack LRRC8 isoforms B through E (Fig 3C). In LRRC8D−/− cells, notwithstanding unchanged I Cl,vol amplitudes (Fig 3A and B; Voss et al, 2014), RVD was significantly reduced but not abolished (Fig 3C). Hence, it seems unlikely that LRRC8D disruption protects cells against cisplatin toxicity by impairing VRAC‐ and AVD‐dependent apoptosis.

LRRC8‐dependent induction of apoptosis by cisplatin and staurosporine We next measured drug‐induced activation of caspase‐3 to test whether the induction of apoptosis by staurosporine or high concentrations of cisplatin depends on LRRC8 subunits. Cisplatin‐induced caspase activation was indeed suppressed in LRRC8A−/−, LRRC8D−/−, and LRRC8−/− cells that lack all LRRC8 subunits (Fig 4A). To explore whether activation of VRAC by hypotonic swelling facilitated cisplatin‐induced apoptosis, we exposed cells during 1 h to 200 μM cisplatin in hypotonic medium (−25%) and measured caspase activity after 1 to 3 days. This procedure drastically increased caspase activation in WT cells (Fig 4B). Swelling‐enhanced caspase induction by cisplatin depended on VRAC as it was abolished in LRRC8A−/− and LRRC8−/− cells. It was strongly reduced, but not abolished, in LRRC8D−/− cells (Fig 4B). Figure 4.LRRC8 subunit‐ and osmolarity‐dependent caspase induction in HCT116 cells A, B. Cisplatin‐induced caspase activity in the continuous presence of 200 μM cisplatin under isotonic conditions (A), or after 1 h exposure to 200 μM cisplatin under iso‐ and hypotonic conditions (B), was followed over time in WT, LRRC8A −/− , LRRC8D −/− , and LRRC8 −/− HCT116 cells. Results from LRRC8A −/− and LRRC8D −/− were obtained with two different clonal cell lines each and averaged.

C. Caspase activation after 1‐h exposure to 4 μM staurosporine under iso‐ or hypotonic conditions of WT, LRRC8A−/−, LRRC8D−/−, and LRRC8−/− HCT116 cells. Data information: Data are presented as mean ± SEM, n = 3–6. *P < 0.05; **P < 0.01; and ***P < 0.001. Similar results were obtained in three independent experiments. Fold change in (A) refers to t = 0. Control experiments indicated that hypotonicity per se had no effect ( Data information: Data are presented as mean ± SEM,= 3–6. *< 0.05; **< 0.01; and ***< 0.001. Similar results were obtained in three independent experiments. Fold change in (A) refers to= 0. Control experiments indicated that hypotonicityhad no effect ( Appendix Fig S4 ). By contrast, staurosporine‐induced caspase activation was not enhanced by hypotonic swelling and was suppressed in LRRC8A−/−, but not in LRRC8D−/− cells (where it was rather increased for unknown reasons) (Fig 4C). Since both I Cl,vol and caspase induction by staurosporine depend on LRRC8A but not LRRC8D, these results are compatible with the hypothesis that VRAC activation‐dependent AVD facilitates the progression of apoptosis (Maeno et al, 2000; Shimizu et al, 2004). VRAC activation by pro‐apoptotic stimuli was monitored by the quenching of YFP fluorescence by externally added iodide (Voss et al, 2014). Iodide permeates VRAC efficiently and hence VRAC opening increases YFP quenching rates. The slow emergence of an LRRC8A‐dependent quenching component upon incubation with 200 μM cisplatin (Fig 5A–C) or 4 μM staurosporine indicated that both drugs slowly activated VRAC (Fig 5E and F). Drug‐induced VRAC activation was small compared to that elicited by acute hypotonic cell swelling (Fig 5D). A direct comparison of quenching slopes, however, underestimates the difference in activation because quenching rates after acute exposure to hypotonicity (but not with drug preincubation) also reflect the time course of cell swelling and ensuing VRAC opening (Appendix Fig S2A and E). Figure 5.Activation of LRRC8 channels by pro‐apoptotic stimuli A–C. Cisplatin‐induced iodide influx into WT (black ■), but not LRRC8A −/− (green ▼) HEK cells indicates VRAC halide current activation during apoptosis. Cells expressing an iodide‐sensitive YFP variant were exposed to 200 μM cisplatin for periods of 0.5 h (A), 4.5 h (B), or 8.5 h (C) before adding extracellular I − (50 mM final). The difference in slopes of YFP fluorescence quenching between control and cisplatin‐treated cells semiquantitatively reflects VRAC current activation. Note that increased YFP quenching with cisplatin preincubation is not due to large non‐specific leaks resulting from cell morbidity. Such leaks should lead to a fast component of YFP quenching in WT, but not LRRC8A −/− cells after the pipetting artifact that immediately follows addition of iodide (indicated by arrows).

D. Swelling‐induced iodide influx into WT (black ■) and LRRC8A −/− (green ▼) HEK cells for comparison. Iodide (50 mM final) was added in isotonic or hypotonic (230 mOsm final) solution at the time indicated by arrow.

E, F. Time course of VRAC activation by 200 μM cisplatin (E) or 4 μM staurosporine (F) determined as in (A–C). Averaged maximal slopes of YFP quenching from eight wells (E) or 16 wells (F) each were evaluated to estimate iodide influx rates. WT (black ■) and LRRC8A−/− (green ▼). Data are presented as mean ± SEM.

VRAC Cl− currents are inhibited by cisplatin–DMSO complexes in a LRRC8D‐dependent manner In a high‐throughput drug screen for VRAC transport inhibitors, in which compounds were routinely dissolved in DMSO, we identified cisplatin as putative VRAC blocker (data not shown). However, suppression of I Cl,vol required DMSO, with which cisplatin forms adducts like Pt(NH 3 ) 2 (Cl)(DMSO) (Fischer et al, 2008). Cisplatin–DMSO complexes inhibited native I Cl,vol of HEK cells in a dose‐dependent manner (Fig 7A), but had no effect on currents of LRRC8D−/− cells (Fig 7B). Hence, differently composed LRRC8 channels can also differ in their pharmacology, and cisplatin‐conducting VRACs might be specifically targeted without abolishing I Cl,vol and cell volume regulation. Figure 7.Cisplatin–DMSO inhibition of I Cl,vol depends on the LRRC8D subunit Upper panel, example current traces (as in Fig 3A) of fully activated I Cl,vol in HEK cells exposed to hypotonic solution containing vehicle (0.3% DMSO) or 200 μM cisplatin in 0.3% DMSO. Dashed lines indicate zero current. Lower panel, I Cl,vol current densities (at −100 mV and 100 mV) of WT HEK cells treated with different cisplatin concentrations. No effect of 200 μM cisplatin/DMSO on I Cl,vol in LRRC8D−/− HEK cells. Data information: Data are presented as mean ± SEM; the number of experiments is given for each bar; *P < 0.05; **P < 0.01. Data information: Data are presented as mean ± SEM; the number of experiments is given for each bar; *< 0.05; **< 0.01.