New Results

Nucleotide binding is the critical regulator of ABCG2 conformational transitions

Zsuzsanna Gyöngy, Gábor Mocsár, Éva Hegedűs, Thomas Stockner, Zsuzsanna Ritter, László Homolya, Anita Schamberger, Tamás I. Orbán, Judit Remenyik, Gergely Szakács, Katalin Goda
doi: https://doi.org/10.1101/2022.10.19.512890

Abstract

ABCG2 is an exporter type ABC protein that can expel numerous chemically unrelated xeno- and endobiotics from cells. When expressed in tumor cells or tumor stem cells, ABCG2 confers multidrug resistance, contributing to the failure of chemotherapy.

Molecular details orchestrating substrate translocation and ATP hydrolysis remain elusive. Here we present methods to concomitantly investigate substrate and nucleotide binding by ABCG2 in cells. Using the conformation-sensitive antibody 5D3, we show that the switch from the inward-facing (IF) to the outward-facing (OF) conformation of ABCG2 is induced by nucleotide binding. IF-OF transition is facilitated by substrates, and hindered by the inhibitor Ko143. Direct measurements of 5D3 and substrate binding to ABCG2 indicate that the high-to-low affinity switch of the drug binding site coincides with the transition from the IF to the OF conformation. Low substrate binding persists in the post-hydrolysis state, supporting that dissociation of the ATP hydrolysis products is required to reset the high substrate affinity IF conformation of ABCG2.

1. Introduction

The human ABCG2 protein (also known as Breast Cancer Resistance Protein (BCRP), Mitoxantrone Resistance Protein (MXR), Placenta-Specific ABC transporter (ABCP)) is a primary active transporter that belongs to the ATP-binding cassette (ABC) protein superfamily (Doyle et al., 1998). ABCG2 can expel a large variety of chemically unrelated, mainly hydrophobic or amphipathic compounds from cells including chemotherapeutic drugs and toxic metabolic side-products, such as pheophorbide A (Jonker et al., 2002), estrone-3- sulphate (E3S) (Suzuki et al., 2003) and uric acid (Woodward et al., 2009). ABCG2 is expressed in tissues with barrier functions including the blood-brain barrier, blood-testis barrier, intestine, liver, placenta, kidneys and mammary glands (Doyle et al., 1998, Fetsch et al., 2006, van Herwaarden and Schinkel, 2006). Because of its tissue localization and its broad substrate spectrum ABCG2 plays an important role in the absorption, distribution, elimination, and toxicity (ADME-Tox) of chemotherapeutic drugs used for the treatment of various diseases (Giacomini et al., 2013, Sarkadi et al., 2006). In addition, the expression of ABCG2 in tumor tissues, especially in the so-called tumor stem cells or drug-tolerant persister cells correlates with an unfavorable prognosis of tumor chemotherapy (Ding et al., 2010). On the other hand, genetic polymorphisms leading to decreased expression and/or impaired function of ABCG2 may result in various pathological conditions (recently reviewed in (Homolya, 2021)). These include altered therapy responses, drug-related toxic reactions, as well as hyperuricemia and gout (Ishikawa et al., 2013), since ABCG2 is also a key player of the intestinal uric acid elimination pathway (Woodward et al., 2009, Kottgen and Kottgen, 2021).

ABCG2 is a “half-transporter” consisting of an N-terminal nucleotide binding domain (NBD) followed by a C-terminal transmembrane domain (TMD) that form homodimers in the plasma membrane to obtain a functional transporter (Kage et al., 2002). Although the substrate specificity of ABCG2 partially overlaps with that of other drug transporting ABC proteins, its sequence similarity is limited to the evolutionarily conserved NBDs. In all ABC transporters, ATP is bound by the Walker A and B motifs of one NBD and by the signature sequence of the other NBD, resulting in a close interaction of the two NBDs often referred as “sandwich-dimer” (Locher, 2009). The molecular mechanism of action of ABC transporters has intrigued scientists for many years. Recent publications of high resolution cryo-EM structures have allowed for interrogating of molecular details with atomic precision. According to the widely accepted idea, uphill transport of substrates is linked to conformational changes of the TMDs, which are regulated by the ATP-dependent formation and separation of the NBD “sandwich-dimer”. In absence of ATP, apo-ABCG2 adopts an inward-facing (IF) state, wherein the transmembrane cavity is open towards the cytosolic side, with the central four transmembrane helices (TH2, TH5, TH2’, and TH5’) forming a large drug-binding pocket (cavity 1) accessible for substrate binding. ATP-bound ABCG2 is in an outward-facing (OF) state, allowing the release of substrates to the extracellular side. The ATP-free IF state can be stabilized by the conformation-sensitive monoclonal antibody 5D3 (Taylor et al., 2017). This structure contains two substrate-binding cavities: a large central cavity at the cytoplasmic side (cavity 1) and an additional cavity (cavity 2) located toward the extracellular part of ABCG2. The two cavities are separated by L554 and L555, that form a tight hydrophobic seal at the top of cavity 1 preventing the entry of substrates to cavity 2 in the IF conformation (Taylor et al., 2017, Khunweeraphong et al., 2017). In other IF structures captured in the presence of substrates or inhibitors, cavity 1 is occupied by a substrate molecule or either one or two inhibitor molecules (Orlando and Liao, 2020). The OF conformer could be captured only by using the E211Q catalytic glutamate mutant ABCG2 variant. In the ATP-bound conformer of this mutant protein, cavity 1 is completely collapsed, the di-leucine gate is closed, while cavity 2 is open to the extracellular side of the plasma membrane (Manolaridis et al., 2018). However, mutation of the catalytic glutamate may affect the energetics of conformational changes in ABC transporters, thus, it may be argued that the OF state of the E211Q mutant ABCG2 observed in the presence of ATP does not represent a true catalytic intermediate (Manolaridis et al., 2018, Lusvarghi et al., 2021).

Despite the plethora of new ABCG2 structures, several long-standing questions concerning the intramolecular crosstalk between the substrate binding and the nucleotide binding sites remain unanswered, for example, the intramolecular mechanism by which transported substrates stimulate the ATPase activity has not been identified (Ozvegy et al., 2001, Sarkadi et al., 1992, Telbisz et al., 2012). Similarly, there is no universal agreement on how conformational changes induced by nucleotide binding and hydrolysis result in the decrease of drug binding affinity and lead to the release of the transported substrate. The prevailing view is that ATP binding leads to the formation of the closed NBD “sandwich-dimer”, switching the TMDs from the IF to the OF state, resulting in a concomitant decrease of substrate affinity (McDevitt et al., 2008). In that scenario, the role of ATP hydrolysis is to reset the transporter to the high substrate affinity IF state by initiating the dissociation of the closed NBD dimer (Khunweeraphong et al., 2019). However, an alternative model states that nucleotide binding is not sufficient, and the IF-OF transition requires ATP hydrolysis (Kapoor et al., 2018).

Since ABC transporters are very sensitive to the composition of their plasma membrane environment (Januliene and Moeller, 2020, Pal et al., 2007, Bordignon et al., 2020, Romsicki and Sharom, 1998), here we present fluorescence-based methods to study intramolecular crosstalk in live cells or semi-permeabilized cells, where ABCG2 molecules are present in their quasi-natural environment in an undisturbed context with other molecules (Barsony et al., 2016, Goda et al., 2020). To address how changes of TMD conformation and substrate binding are coupled to ATP binding and hydrolysis, we quantify binding of the conformation-sensitive 5D3 antibody (Ozvegy-Laczka et al., 2005). To characterize drug binding, we introduce confocal microscopy- and fluorescence correlation spectroscopy-based (FCS) assays. By measuring drug binding affinity and reactivity to 5D3, we show that nucleotide binding drives ABCG2 from a high to a low substrate-affinity state and find that this switch coincides with the flip from the IF to the OF conformation.

2. Results

2.1. MDCK II cells express fully functional ABCG2 and ABCG2-GFP

ABCG2 and its N-terminally GFP-tagged variant (ABCG2-GFP) were expressed at comparable levels in MDCK II cells (Figure 1A). In accordance with literature data (Orban et al., 2008, Haider et al., 2011), the transport activity of ABCG2 was not influenced by the GFP-tag (Figure 1B-D). ABCG2-GFP and ABCG2 exhibited comparably low 5D3-reactivity in the plasma membrane of untreated live cells, which could be significantly increased by treatment with the ABCG2 inhibitor Ko143 (Figure 1E-G).

Figure 1.
Figure 1. Functional characterization of human ABCG2 and ABCG2-GFP.

Western blot showing comparable expression of ABCG2 and ABCG2-GFP in MDCK II cells using the BXP-21 anti-ABCG2 mAb (A). ABCG2 and ABCG2-GFP expressing cells show decreased mitoxantrone (MX) accumulation, which is reversed to the level of the ABCG2-negative cells by 2 μM Ko143 (B-D). 2 μM Ko143 pre-treatment causes an increase in the 5D3-reactivity of ABCG2 (F) and ABCG2-GFP (G) expressing cells. ABCG2-negative cells show only a negligible 5D3 binding that is not affected by Ko143 (E).

2.2. Nucleotide binding is sufficient to trigger the switch from the 5D3-reactive IF conformation to a 5D3-dim OF conformation

To study nucleotide dependent conformation changes of ABCG2, we systematically changed the intracellular nucleotide concentrations in semi-permeabilized cells. In accordance with an ATP-regulated switch of the TMD conformation, increasing ATP/Mg2+ concentrations gradually decreased the 5D3-A647-reactivity of ABCG2-positive cells, with practically zero staining at high ATP/Mg2+ concentrations (Figure 2A and B). To prevent nucleotide hydrolysis, ATP was either added in the absence of Mg2+ (ATP+EDTA; Figure 2C and D), on ice (Figure 2E and F), or ATP was replaced with the non-hydrolysable ATP analogue AMP-PNP (Figure 2G and H). Interestingly, the conformational change driving ABCG2 into a 5D3-dim (i.e., 5D3 non-reactive) state also occurred in the absence of ATP hydrolysis and showed similar nucleotide concentration dependence (see Table 1). These results indicate that the 5D3-dim and 5D3-reactive conformations correspond to the OF and IF conformations as observed in ATP-bound and nucleotide-free crystal structures, respectively.

Figure 2.
Figure 2. Nucleotide dependence of 5D3-reactivity.

MDCK-ABCG2 cells were permeabilized to allow the titration of intracellular nucleotide concentrations. Dose-response curves of 5D3 binding with increasing concentrations of ATP/Mg2+ (panels A, B), ATP in the absence of Mg2+ (panels C, D),) AMP-PNP/Mg2+ (panels G, H) or ADP/Mg2+ (panels I, J) were obtained in the presence or absence of Vi (left panels) or BeFx (right panels). Samples were pre-treated with nucleotides at 37°C for 10 min, then further incubated with 5 μg/mL 5D3-A647 at 37°C for 20 min except for (panels E, F), where all the treatments were carried out on ice. In case of nucleotide trapping, permeabilized cells were co-treated with nucleotides and BeFx or Vi at 37°C for 30 min, then un-trapped nucleotides were washed out and 5D3 labeling was carried out on ice for 45 min. Representative curves are shown from 3-5 independent experiments.

View this table:
Table 1. Apparent nucleotide affinities (KA) determined in 5D3-reactivity experiments

By replacing the cleaved gamma phosphate following ATP hydrolysis, phosphate analogues, such as vanadate (Vi) or beryllium fluoride (BeFx), trap ABC transporters in a stable ternary complex (ABCG2-ADP-Vi/BeFx). Based on different geometries of myosin structures obtained with transition state analogs, the BeFx- or Vi-trapped post-hydrolytic complexes are believed to represent pre- and post-hydrolytic conformations, respectively (Smith and Rayment, 1996, Fisher et al., 1995, Szakacs et al., 2000). Co-treatment with Vi or BeFx increased the apparent nucleotide affinity of ABCG2 about 10-fold (Table 1) at conditions permitting ATP hydrolysis (Figure 2A and B), confirming that both phosphate analogues can form stable ADP-trapped complexes with ABCG2. When ATP hydrolysis was prevented, the phosphate analogues did not have any effect on the KA values (Figure 2C-H and Table 1).

Unexpectedly, ADP/Mg2+ was able to induce a conformational change that resulted in decreased 5D3 binding, albeit at slightly higher concentrations (KA = 7.38 ± 2.31 mM). Moreover, nucleotide trapping occurred in the presence of phosphate analogs and ADP/Mg2+ (Figure 2I-J). The KA values obtained with ADP/Mg2+ and Vi (KA = 0.44 ± 0.1 mM) or BeFx (KA = 0.57 ± 0.32 mM) did not differ from the KA values of trapping reactions starting from ATP/Mg2+. However, in the trapping reactions with ADP/Mg2+, about 30% of ABCG2 molecules remained in a 5D3-reactive state even in the presence of very high nucleotide concentrations. This observation may suggest that the ternary complex resulting from ADP/Mg2+ is different (i.e., less stable, possessing a shorter lifetime compared to the complex produced from ATP/Mg2+ in the hydrolytic cycle) and therefore the trapping reaction occurs with lower efficiency. Since in energized cells the cytosolic ATP concentration is more than 10-fold higher compared to ADP concentrations (Williams et al., 1993), these results indicate that in live cells, (i) the switch from the 5D3-reactive IF to a 5D3-dim OF conformation is induced by ATP binding; and (ii) resetting to the 5D3-reactive IF conformation can only occur after release of the hydrolysis products.

2.3. Substrates increase the rate of formation of the Vi- or BeFx-trapped species

Transported substrates increase the turnover rate of ATP hydrolysis in ABC transporters (Sarkadi et al., 1992, Telbisz et al., 2012). Progressive accumulation of the transporter molecules in the stable Vi- or BeFx-trapped post-hydrolytic states represents a partial reaction of the catalytic cycle in ABC transporters (Szabo et al., 1998). Accordingly, the accumulation of ABCG2 in the Vi- or BeFx-trapped post-hydrolytic complex was accelerated by substrates (Figure 3). Estrone-3-sulphate (E3S) and quercetin induced about a 5-fold increase in the rate of the trapping reaction (for t1/2 values, see Figure 3B), which is consistent with the extent of stimulations achieved by these compounds in the ATPase assay (Telbisz et al., 2012).

Figure 3.
Figure 3. Effect of transported substrates on the kinetics of the formation of the Vi- and BeFx-trapped complexes.

Permeabilized MDCK-ABCG2 cells were pre-treated or not with 10 μM quercetin or E3S for 10 min at 37°C and then further incubated with 0.5 mM ATP/Mg2+ or ADP/Mg2+ in the presence of Vi or BeFx. Samples taken at different time points were stained with 5μg/ml 5D3-A647 on ice for 45 min. Panel A shows a representative Vitrapping experiment in the absence or presence of E3S, while panel B summarizes the t1/2 values calculated from the exponential fit of the kinetic curves (see Materials and Methods). Mean ± SD of 3-5 independent experiments is shown. Significant differences compared to substrate untreated samples are shown by ***: P<0.001.

The relatively long t1/2 values compared to the total cycle time, which is in the order of 100 ms (as inferred from ATPase data (Yu et al., 2021)), suggest that formation of a stable post-hydrolysis complex by phosphate mimicking anions is a low-probability event. The increased rate of the trapping reaction in the presence of substrates may be explained by a higher turnover of the ATPase cycle, or a longer duration of the Vi- or BeFx-“sensitive” state, that is between the dissociation of the cleaved phosphate and the disassembly of the NBD dimer. Consistently, trapping reactions starting from ADP/Mg2+ were not accelerated by quercetin or E3S (Figure 3B), suggesting that substrates do not affect the overall stability or lifespan of the ADP-bound, phosphate analogue-sensitive ABCG2 conformer. This can happen, if substrates similarly accelerate the formation and the dissociation of the ADP-bound conformer, or alternatively, they do not have any effect on these processes.

2.4. Substrates accelerate the IF to OF transition of ABCG2

With the aim to pinpoint the transition that is accelerated by transported substrates, in the following experiments we studied how nucleotides and substrates affect the kinetics of the IF to OF transition detected by a shift in 5D3 binding. To align ABCG2 molecules in an IF state, semi-permeabilized (nucleotide-free) MDCK cells expressing ABCG2 were pre-labeled with 5D3-A647 antibody. Unbound 5D3-A647 molecules were removed, and cells were incubated at 37 °C in a sufficiently large volume to prevent rebinding of the antibody. Under these conditions, we observed a gradual decrease of the 5D3-A647 fluorescence of cells, which was completely prevented by Ko143 treatment, supporting the notion that the ABCG2 molecules are intrinsically dynamic, while Ko143 stabilizes them in the IF 5D3-reactive conformation (Figure 4A). The dissociation rate of the antibody was significantly enhanced in the presence of transported substrates or ATP/Mg2+ (Figure 4A and 4B). However, the largest (about 5-fold) decrease of the t1/2 values corresponding to the half-life of the 5D3-bound ABCG2 conformer was observed when substrates were co-administered with ATP/Mg2+ (Figure 4B).

Figure 4:
Figure 4: Effects of substrates and nucleotides on the kinetics of 5D3 dissociation when substrates were added simultaneously with nucleotides (A and B), or prior to nucleotides (C).

Permeabilized MDCK-ABCG2 cells were pre-labeled with 5D3-A647 in the absence (panels A and B) or presence (panel C) of substrates (10 μM quercetin or 10 μM E3S) for 20 min at 37°C. After removal of the unbound antibody, cells were further incubated with 3 mM ATP/Mg2+ or AMP-PNP/Mg2+ in the absence or presence of substrates. Panel A shows representative 5D3 dissociation curves in the indicated conditions, while panel B and C summarizes the t1/2 values calculated from the exponential fit of the dissociation curves (see Materials and Methods). Means ± SD of 3-5 independent experiments are shown. Significant differences compared to untreated samples (1st column) are shown by ###: P<0.001 or ##: P<0.01. Significant differences compared to only-nucleotide-treated samples are indicated by ***: P<0.001.

In further experiments, intracellular nucleotide pools were replenished with a non-hydrolysable ATP analogue AMP-PNP/Mg2+ (Figure 4B). In the absence of ATP hydrolysis, ABCG2 molecules undergo IF to OF transition (Figure 2G and H), and the backward transition to the IF state has an extremely low probability. The time dependence of the AMP-PNP/Mg2+-induced 5D3 dissociation was comparable to that obtained with ATP/Mg2+ supporting that the antibody dissociation kinetics observed either in the presence of ATP/Mg2+ or AMP-PNP/Mg2+ may reflect the first nucleotide-induced IF to OF transition of ABCG2 (Figure 4B). Strikingly, when co-administered with AMP-PNP/Mg2+, the same substrates did not increase further the dissociation rate of 5D3, suggesting that the NBD dimer formation induced by AMP-PNP/Mg2+ binding switches ABCG2 into the low drug binding affinity state (Figure 4B). However, similar decrease of the t1/2 values was observed when substrates were added before AMP-PNP/Mg2+ treatment (compare Figure 4B and 4C), indicating that transition to the OF state can also be accelerated by substrates with AMP-PNP/Mg2+ treatment. Collectively, the above data imply that substrate binding at the TMDs induces a structural change in the transporter that can facilitate the nucleotide-dependent NBD dimer formation and the concomitant IF to OF transition probably by reducing the energy barrier of the above conformational changes.

2.5. The nucleotide-free IF conformation of ABCG2 has higher substrate affinity compared to the Vi-trapped post-hydrolytic conformation

In the following experiments, we visualized the subcellular localization of ABCG2-GFP and the fluorescent ABCG2 substrate MX by using confocal microscopy. In accordance with previous observations (Homolya et al., 2011), at low concentrations, MX only stained poorly the MDCK ABCG2-GFP cells, while ATP depletion, Ko143 or Vi treatments increased the intracellular accumulation of MX (Figure 5A and B). Interestingly, ATP depleted cells exhibited strong plasma membrane staining by MX (Fig 5A). Plasma membrane staining by MX in both native and ATP depleted cells was abolished by treatment with the competitive inhibitor Ko143, suggesting that red fluorescence in the plasma membrane reflects MX binding to ABCG2 molecules. To quantify the fraction of MX-bound ABCG2 molecules, we calculated the Pearson’s correlation coefficients (PCC) between the MX and ABCG2-GFP signals in pixels representing the plasma membrane. Since the ABCG2-GFP signal was unchanged during the course of the different treatments (Figure 5C), the correlation coefficients depend mostly on MX binding to the transporter. In ATP depleted cells, the high correlation values indicate that the majority of ABCG2 molecules reside in a MX-bound conformation (PCC = 0.72 ± 0.12) (Figure 5D). The correlation between the two signals strongly decreased in the presence of Ko143 (PCC = −0.1 ± 0.18), suggesting displacement of MX from the substrate binding site of the transporter by the competitive inhibitor. Similar results were obtained when ATP depletion was combined with Ko143 treatment (PCC = −0.04 ± 0.17). Binding of MX to ABCG2 was also suppressed by Vi (PCC = 0.18 ± 0.13), which is consistent with the notion that the post-hydrolytic ABCG2 conformer possesses low substrate affinity. Interestingly, in untreated cells, we measured significantly higher co-localization between the MX and ABCG2-GFP signals (PCC = 0.3 ± 0.12) than in Ko143-treated cells, suggesting that in the plasma membrane of live cells, a significant subset of ABCG2 molecules resides in an MX-bound IF state.

Figure 5.
Figure 5. Cellular distribution of MX (red) in MDCK cells expressing ABCG2-GFP (green) in response to ATP depletion and/or Ko143 or Vi treatment.

(A). All treatments increased the intracellular MX fluorescence intensity (B), while the ABCG2-GFP fluorescence intensity in the plasma membrane remained unchanged (C). Pearson’s correlation coefficients between the intensity distributions of MX and ABCG2-GFP in the plasma membrane pixels reveal the stabilization of a high-affinity substrate binding ABCG2 conformation in energy-deprived cells (D). ATP depletion was induced by 15 min pretreatment of cells with 10 mM sodium azide and 8 mM 2-deoxy-D-glucose. Ko143, a non-fluorescent, competitive ABCG2 inhibitor, was added 10 min before MX staining. In panels B and C bars represent mean ± SD values, while panel D shows box and whisker plots. For each treatment group 150-200 cells were analyzed from 3-5 independent experiments. Significant differences compared to the untreated control are shown by ***: P<0.001.

2.6. MX binding to ABCG2 is confirmed by its reduced mobility using FCS measurements

As an independent approach to follow MX binding to ABCG2, we measured the mobility of MX in the plasma membrane by fluorescence correlation spectroscopy (FCS). MX molecules bound to ABCG2 are expected to show decreased diffusion compared to free MX (Horsey et al., 2020). We analyzed the fluorescence autocorrelation functions (ACFs) of MX (orange) and ABCG2-GFP (green) in the plasma membrane using a two-component model (Figure 6A). Upon ATP-depletion, the diffusion coefficient of MX decreased to the level obtained for ABCG2-GFP (Figure 6B), indicating that MX molecules readily bind to the nucleotide-free IF conformer of ABCG2. In accordance with the data obtained from cellular distributions (Figure 5A and D), the competitive inhibitor Ko143 prevented MX binding to ABCG2, resulting in the dominance of a high plasma membrane mobility MX population similar to the only-MX treated cells (Figure 6B).

Figure 6.
Figure 6. Fluorescence autocorrelation functions (A) and diffusion constants (B) of ABCG2-GFP (green) and MX (orange) in the plasma membrane of intact MDCK cells.

ABCG2-GFP expressing cells without pre-treatments or following ATP depletion or Ko143 pre-treatment were stained with 100 nM MX for 15 min at 37°C. Each bar shows the mean ± SD for n=50-100 cells from at least 3 independent measurements. Significant differences compared to the diffusion constant of ABCG2-GFP (green bar) in the plasma membrane of untreated cells are shown by ***: P<0.001.

2.7. AMP-PNP binding switches ABCG2 to a conformation that is unable to bind MX

As shown in Figure 2, permeabilization of cells with SLO synchronizes ABCG2 molecules in a nucleotide-free, 5D3-reactive IF conformation also observed in cryo-EM structures (Taylor et al., 2017). When permeabilized cells were treated with MX alone, we measured a strong co-localization between ABCG2-GFP and MX in the plasma membrane (PCC = 0.85±0.05), confirming the high substrate affinity of the IF conformation of ABCG2. Strikingly, pre-incubation of permeabilized cells with 5 mM AMP-PNP/Mg2+ strongly reduced the co-localization between MX and ABCG2-GFP in the plasma membrane (PCC = −0.1181 ± 0.2019), indicating that the conformational changes induced by AMP-PNP binding prevented MX binding to ABCG2 (Figure 7).

Figure 7.
Figure 7. Substrate affinity changes of ABCG2 in permeabilized cells.

SLO-permeabilized ABCG2-GFP expressing cells were pre-treated or not with 5 mM AMP-PNP/Mg2+ for 15 min and then stained with 500 nM MX for 15 min at 37°C. Permeabilized cells were identified by PI staining. Box and whisker plots show Pearson’s correlation coefficients of n>150 cells from 3 independent experiments. ***P<0.001 by Kolmogorov-Smirnov test.

3. Discussion

According to the alternating access model formulated by Jardetzky more than 50 years ago, membrane transporters alternate between IF and OF states, in which the centrally located substrate-binding site is accessible to only one side of the membrane at a time (Jardetzky, 1966). In active transporters, such as ABC transporters, accessibility changes are accompanied by a significant change of substrate affinity that is linked to binding and hydrolysis of ATP (Seeger and van Veen, 2009). Based on homology models and cryo-EM structures, ABCG2 is believed to alternate between a nucleotide-free IF and a nucleotide-bound OF conformation during its transport cycle. The switch between the NBD-dissociated IF and the NBD-associated OF states involves a series of conformational changes that finally result in the uphill transport of substrates. In agreement with cryo-EM studies, we show in nucleotide titration experiments (Figure 2) that 5D3 exclusively recognizes the nucleotide-free IF conformation of ABCG2 (Taylor et al., 2017), as the 5D3 recognized epitope of ABCG2 only forms in the IF conformation, while it is disrupted in the OF conformation (Figure 8).

Figure 8.
Figure 8. Two rotationally symmetric 5D3 epitopes are formed in the IF conformation (A, C) and are absent in the OF conformation of ABCG2 (B, D).

The two protomers of ABCG2 are colored in pink and green in cartoon backbone and transparent surface rendering. The two epitopes are highlighted in blue and orange, and include all residues that are in direct contact with the 5D3 Fab (white) in the 6ETI crystal structure. Panels A and B show an extracellular view of the 5D3-bound IF (PDB ID: 6ETI), and the OF (PDB ID: 6HBU) ABCG2 conformations, respectively. A side view of the IF (C) and the OF (D) ABCG2 are also shown. The membrane boundaries are indicated by dashed lines.

Here we used confocal microscopy and FCS technique to study MX binding to ABCG2 with the aim to better understand the intramolecular crosstalk between the nucleotide and the substrate binding domains. While the interactions of fluorescent substrates with purified ABC transporters have been studied in nanodiscs (Li et al., 2017) or styrene maleic acid lipid copolymer particles (Horsey et al., 2020), to our knowledge we have applied the FCS technique for the first time to study substrate binding to ABCG2 in its natural plasma membrane environment in live cells.

In accordance with structural studies, our confocal microscopy experiments carried out in live cells show that the apo form of ABCG2 possess high MX-affinity, while nucleotide binding and the concomitant dimerization of the NBDs induce conformation changes that prevent MX binding. The simultaneous drop in the 5D3- and MX-binding indicates that the high-to-low switch in drug binding affinity coincides with the transition from the IF to the OF conformation. Ko143, which can displace MX from the substrate binding site (see Figure 5 and 6) also prevents the transition of ABCG2 from the IF to the OF conformation (Figure 4A) in agreement with previous cryo-EM studies (Orlando and Liao, 2020, Yu et al., 2021, Jackson et al., 2018). Our data also directly indicate that high affinity binding of MX requires the IF conformation and that energizing the transporter by nucleotides is not required for substrate binding. The nucleotide titration experiments prove that nucleotide binding (ATP, AMP-PNP and ADP) and concomitant NBD dimer formation is sufficient to induce the conformational switch to the OF conformation, and that ATP hydrolysis is not essential for the nucleotide-dependent IF-OF transition (Figure 2) and the high-to-low switch in drug binding affinity (Figure 7) supporting the results of structural studies carried out using the catalytic glutamate mutant ABCG2 variant (Manolaridis et al., 2018).

We also observed that in the presence of physiological ATP concentrations, in the context of the plasma membrane of live cells, a small, but significant fraction of ABCG2 molecules reside in the MX-bound state (Figure 5D) ready to initiate a productive transport cycle. The Vi-trapped post-hydrolysis state possesses low substrate affinity (Figure 5A and D), confirming that dissociation of the hydrolysis products is required to switch the transporter from the OF conformation back to the IF conformation and to reset its high substrate affinity.

Previous studies observed analogous changes of the substrate binding affinity in ABCB1 (Martin et al., 2001), ABCC1 (Rothnie et al., 2006) and ABCG2 (McDevitt et al., 2008) using radio ligand binding assays. A similar, bidirectional interdomain cross-talk between the NBDs and TMDs was observed in human ABCB1 using the UIC2-reactivity assay (Barsony et al., 2016, Goda et al., 2020). The fact that the antibody binding site and the NBDs of the transporters are ~80 Å apart and on opposite sides of the membrane suggests that long range conformational couplings between the TMD and NBD motions are conserved features among human ABC exporters. However, the similarity in coupling of the ATPase activity and the transport cycles of ABCB1 and ABCG2 is remarkable in view of the differences in the conformational changes of the TMDs and the substrate binding cavities (Arana and Altenberg, 2019, van Wonderen et al., 2014, Locher, 2016), and the different modes of the nucleotide dependent NBD motions (Stockner et al., 2020).

Like several other ABC transporters, ABCG2 possesses significant basal ATPase activity, which is generally increased 2 to 5-fold by transported substrates (Telbisz et al., 2012). The increased catalytic turnover is reflected by the enhanced steady-state ATP hydrolysis rate (Telbisz et al., 2012, Ozvegy et al., 2001) and the increased rate at which ABCG2 becomes trapped in the Vi- or BeFx-bound post-hydrolytic states (Figure. 3).

Our experiments demonstrated that 5D3 binding to ABCG2 is reversible, and addition of substrates, ATP/Mg2+ or their combination can accelerate the kinetics of antibody dissociation. The reversible nature of 5D3 binding also explains previous observations that treatments with saturating concentration of the antibody did not induce complete inhibition of transport and of ATPase activity (Ozvegy-Laczka et al., 2005).

Importantly, 5D3 dissociation experiments carried out in the absence of nucleotides clearly showed that substrate binding alone has an effect on the conformation of ABCG2 (Figure 4A and 4B). When substrates are presented prior to the nucleotides, the IF to OF transition is accelerated. In contrast, when the nucleotide analogue AMP-PNP is allowed to bind first, substrates cannot promote the transition (Figure 4B and C). Thus, substrates accelerate the cumulative step of nucleotide binding, NBD “sandwich-dimer” formation and the concomitant IF to OF transition, but only when having access to the substrate binding site as present in the IF state. Accordingly, in recent cryo-EM studies carried out at turnover conditions (in the presence of substrate and ATP/Mg2+) two conformers representing the transition from the IF state to a semi-closed state were identified. These structures revealed that the accessibility of the substrate binding site is gradually decreasing upon the closure of the NBD dimer (Yu et al., 2021), and therefore, substrates should bind to the IF conformer.

In the presence of saturating concentrations of nucleotides added either at hydrolysing conditions (ATP/Mg2+ at 37°C) or at non-hydrolysing conditions (ATP/Mg2+ at 4°C, Mg2+-free ATP or AMP-PNP/Mg2+) the majority of ABCG2 molecules populate the low substrate affinity OF conformation, suggesting that similar to ABCC1 (Wang et al., 2020) and ABCB1 (Barsony et al., 2016), ABCG2 can transition from the IF to OF state, and this transition is not the rate limiting step of the whole catalytic cycle. Therefore, acceleration of the IF to OF transition by substrates does not decrease the total cycle time considerably. The channel-type ABC protein CFTR (ABCC7) is different in this respect, as the pore opening is believed to correspond to the IF to the OF transition and to represent the slowest step of the gating cycle (Vergani et al., 2013). Detailed kinetic analysis of several ABC transporters will be needed to establish differences and similarities of the rate limiting step within the superfamily. Since substrates can no longer bind when ABCG2 is switched to the OF, low-substrate-affinity conformation, substrates should dissociate before ATP hydrolysis and the concomitant NBD dissociation. Taken together, substrates increase the steady-state ATPase activity by selectively accelerating the IF to OF transition without affecting the backward transitions to the IF conformation induced either by ATP hydrolysis or spontaneous NBD dissociation. Consistently, a similar mechanism of substrate stimulation was observed in ABCC1 using single-molecule fluorescence spectroscopy (Wang et al., 2020).

In conclusion, our results indicate that nucleotide binding is the major regulator of TMD conformation in ABCG2. When ABCG2 is flipped into the OF conformation, substrates can no longer bind, which implies that substrate-stimulation of the steady-state ATPase activity is realized by the acceleration of ATP-induced IF to OF transition without affecting the rate of the backward transitions and thus resulting in an increased chance of ATP hydrolysis. The nucleotide induced IF to OF transition coincides with the high-to-low switch of substrate affinity, and this event precedes ATP hydrolysis. The cumulative half cycle consisting of ATP hydrolysis, the release of the hydrolysis products and the OF to IF transition is the rate limiting step of the whole transport cycle. In live cells, only a small fraction of ABCG2 molecules is in a transient substrate-bound conformation ready to harvest the energy of ATP for transport, while depletion of ATP increases this fraction prolonging the waiting time before the IF to OF transition. ABCG2 molecules that have reached the NBD sandwich dimer without substrate binding should undergo a futile ATP hydrolysis cycle to reset the transporter into an IF state with high drug binding affinity. In the future, single molecule-based approaches (Catipovic et al., 2019, Josts et al., 2020, Wang et al., 2020) may provide further insights into the kinetics of the above transitions.

4. Materials and Methods

View this table:

4.1. Chemicals

Cell culture media, supplements, and chemicals were purchased from Sigma-Aldrich (Budapest, Hungary). Alexa 647 succinimidyl ester (A647) was purchased from Life Technologies, Inc. (Carlsbad, CA, USA). The 5D3 anti-ABCG2 mAb was prepared from hybridoma supernatants by affinity chromatography. Purity (> 97%) was verified by SDS/PAGE. The 5D3 hybridoma cell line was a kind gift from Brian P. Sorrentino (Division of Experimental Hematology, Department of Hematology/Oncology, St. Jude Children’s Research Hospital, Memphis, Tennessee, USA). 5D3 antibody was labeled with A647 (5D3-A647) and was separated from the unconjugated dye by gel filtration using a Sephadex G-50 column. The dye to protein ratio was approximately 3 for each antibody preparation. Stock solutions of nucleotides were prepared in distilled water at pH 7 (by Tris-Base).

4.2. Cell lines

The MDCK II Madin-Darby canine kidney cell line was a kind gift from Gerrit van Meer, (Dept. of Membrane Enzymology, Centre for Biomembranes and Lipid Enzymology Utrecht, The Netherlands). The MDCK II cell lines stably expressing ABCG2 or its N-terminally green fluorescent protein (GFP) tagged variant (Orban et al., 2008) were established using the Sleeping Beauty transposon-based gene delivery system (Erdei et al., 2018). Cells expressing the transgene at high level were selected based on their 5D3-A647 or GFP fluorescence by repeated flow cytometry sorting using a Becton Dickinson FACSAria III Cell Sorter (Becton Dickinson, Mountain View, CA, USA). Cells were grown as monolayer cultures in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 0.1 mg/ml penicillin-streptomycin cocktail, 10% heat-inactivated fetal calf serum and 2mM L-glutamine. Cells were maintained at 37°C in a 5% CO2 atmosphere and were grown to approximately 80% confluency.

4.3. Western blot analysis

Cells (2 × 105) were lysed in 100 μl reducing Laemmli sample buffer (6X) for 10 min at 95 °C. Afterwards the lysates were subjected to SDS-polyacrylamide gel electrophoresis using an 8% polyacrylamide gel and then electroblotted onto a nitrocellulose membrane with a pore size of 0.45 μm (GE Healthcare Life Sciences, Little Chalfont, Buckinghamshire, UK). ABCG2 expression was detected by the BXP-21 mouse mAb, while actin was labeled with the C-2 mouse mAb (both from Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA). As a secondary antibody, a goat anti-mouse HRP-conjugated IgG (Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA) was applied. All antibodies were used at 1:2,500 dilution. Bands were visualized with SuperSignal™ West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific, Waltham, MA) using the FluorChem Q gel documentation system (Alpha Innotech Corp., San Leandro, CA).

4.4. Mitoxantrone accumulation test

The transport activity of ABCG2 and ABCG2-GFP was studied using a mitoxantrone (MX) accumulation assay (Tarapcsak et al., 2017). Cells (0.5 × 106ml-1 in PBS containing 7 mM glucose (gl-PBS)) were pre-incubated in the presence or absence of 2μM Ko143 for 15 min at 37 °C and then stained with 5 μM MX for 30 min. Samples were washed three times with ice-cold gl-PBS containing 0.5% fetal bovine serum (FBS) and stored on ice until flow cytometry measurement. To exclude dead cells from the analysis, samples were stained with propidium iodide (PI).

4.5. 5D3-reactivity assay

Cells (0.5 × 106 ml-1 in gl-PBS) were pre-incubated with or without 2 μM Ko143 for 10 min and then further incubated with 5 μg/ml 5D3-A647 monoclonal anti-ABCG2 antibody at 37 °C. After 30 min of incubation, samples were washed 2 times with ice-cold gl-PBS and centrifuged for 5 min at 435 ×g at 4 °C. The 5D3-A647 fluorescence intensity of the cells was measured by flow cytometry.

4.6. Permeabilization of cells with streptolysin-O toxin

Streptolysin-O ((SLO) (Sigma-Aldrich, Budapest, Hungary)) is a pore-forming exotoxin from Streptococcus pyogenes. The SLO pores formed in the membrane are permeable to small water-soluble molecules including nucleotides (Yang et al., 2006). SLO is an oxygen-labile toxin that is reversibly activated by dithiothreitol (DTT). Cell suspensions (1 × 107 cells/ml) were treated with 250 U/ml SLO in the presence of 1 mM DTT, Protease Inhibitor Cocktail (2 mM AEBSF, 0.3 μM aprotinin, 116 μM bestatin, 14 μM E-64, 1 μM leupeptin), 0.5 mM PMSF, and 1% FBS in gl-PBS at 37°C for 30min, which allowed permeabilization of approximately 50% of cells, as it was verified by PI staining. The reaction was stopped with 20 ml PBS containing 1% FBS and the cells were centrifuged at 635×g for 5 min at room temperature. Unbound toxin was removed by washing the cells 3 times with PBS and the cell pellet was resuspended in PBS (Goda et al., 2020). The applied 1 mM DTT concentration did not affect the 5D3-reactivity of ABCG2.

Cells grown in eight-well chambered coverslip plates (ibidi GmbH, Gräfelfing, Germany) for confocal microscopy experiments were permeabilized using 62.5 U/ml SLO in the presence of 1 mM DTT and Protease Inhibitor Cocktail at 37 °C for 15 min, in HEPES solution (20 mM HEPES, 123mM NaCl, 5 mM KCl, 1.5 mM MgCl2, 1 mM CaCl2) containing 1% FBS.

4.7. Determination of apparent affinity of nucleotide binding

Apparent affinity of nucleotide binding (KA) was determined as described previously (Barsony et al., 2016, Goda et al., 2020). Permeabilized cells (1 × 106 ml-1) were pre-treated with nucleotides added at different concentrations in the presence of equimolar concentrations of Mg2+ at 37°C for 10min and then further incubated with 5 μg/ml 5D3-A647 at 37°C for 20 min. To prevent ATP hydrolysis, ATP was added without Mg2+ in the presence of 5 mM EDTA or the whole experiment was carried out on ice. In nucleotide trapping experiments nucleotide treatments were applied together with 0.5 mM sodium orthovanadate (Vi) or BeFx (200 μM BeSO4 and 1 mM NaF) at 37 °C for 30 min. Subsequently, cells were labeled with 5 μg/ml 5D3-A647 on ice for 45 min after removal of the un-trapped nucleotides by washing them twice with ice-cold PBS. After antibody labeling samples were washed again 3 times with ice-cold PBS and centrifuged for 5 min at 635× g at 4 °C. The mean 5D3-A647 fluorescence intensity of the cells was determined by flow cytometry and plotted as a function of the nucleotide concentration. To determine the apparent affinity of ABCG2 for nucleotides (KA), data points were fitted with the four-parameter Hill function, where the Fmin and Fmax values represent the minimum and maximum fluorescence intensities, respectively: Embedded Image

4.8. Studying the kinetics of nucleotide trapping by Vi or BeFx

Permeabilized cells (1 × 106 ml-1) were incubated with 0.5 mM ATP/Mg2+ or ADP/Mg2+ and 0.5 mM Vi or BeFx (200 μM BeSO4 and 1 mM NaF) in the presence or absence of ABCG2 substrates (10 μM quercetin or 10 μM estrone-3-sulfate (E3S)) in PBS at 37 °C. To follow the kinetics of the trapping reaction, 500 μl aliquots were taken at different time points and washed twice with 5 ml ice-cold PBS. After washing the samples were resuspended in 500 μl ice-cold PBS and labeled with 5 μg/ml 5D3-A647 at 4 °C for 45 min. The 5D3-A647 fluorescence intensity of the samples (F) was plotted as a function of time (t). The t1/2 values, representing the half-life of the 5D3-reactive ABCG2 conformation, were calculated from an exponential fit of the data points according to the following equation: Embedded Image where in F0 is the difference between the zero and infinite time point of the curve and c is the background fluorescence intensity of cells.

4.9. 5D3 dissociation

Permeabilized MDCK-ABCG2 cells (1 × 106 ml-1) were pre-labeled with 5D3-A647 in the presence or absence of 10 μM quercetin or 10 μM E3S for 20 min at 37°C. After removing the unbound 5D3-A647, cells (1 × 105 ml-1) were further incubated with 3 mM ATP/Mg2+ or AMP-PNP/Mg2+ in the absence or presence of the above substrates at 37 °C. To study the kinetics of 5D3 dissociation, 500 μl aliquots were taken at regular intervals and washed twice with ice-cold PBS. The 5D3-A647 fluorescence intensity of the cells was measured by flow cytometry and plotted as a function of time (t). The t1/2 values, representing the half-life of the 5D3-reactive ABCG2 conformation, were calculated from an exponential fit of the data points using Equation 2.

4.10. Flow cytometry

Flow cytometry analysis was carried out using a Becton Dickinson FACS Array flow cytometer (Becton Dickinson, Mountain View, CA, USA). A 635 nm laser was used for the excitation of MX and A647 and their fluorescence was detected in the red channel through a 661/16nm bandpass filter, while a 532nm laser was used for the excitation of PI and the emitted light was detected using a 585/42nm bandpass filter. Cell debris was excluded from analysis on the basis of FSC and SSC signals. Fluorescence signals of 2×105 cells/sample were collected in logarithmic mode, and the cytofluorimetric data were analyzed using the Flowing software (Cell Imaging Core, Turku Centre for Biotechnology, Turku, Finland).

4.11. Confocal laser scanning microscopy and fluorescence co-localization analysis

To assess the co-localization of ABCG2-GFP and the fluorescent ABCG2 substrate MX in the plasma membrane of MDCK cells, we carried out confocal laser scanning microscopy (CLSM) experiments. Measurements were performed in eight-well chambered coverslip plates (ibidi GmbH, Gräfelfing, Germany). ATP depletion of intact cells was induced by a 15 min pre-treatment with 8 mM 2-deoxy-D-glucose and 10 mM sodium azide in glucose-free medium. ATP depleted or non-ATP depleted cells were pre-treated with 2 μM Ko143 or 0.5 mM Vi for 15 min, stained with 500 nM MX for 15 min at 37°C and then washed three times with HEPES solution. SLO-permeabilized cells were pre-stained with 6 μg/ml PI, then further incubated with 500 nM MX for 15 min at 37°C in the presence or absence of 5 mM AMP-PNP and subsequently washed three times with HEPES solution.

Fluorescence images were acquired with a Nikon A1 Eclipse Ti2 Confocal Laser-Scanning Microscope (Nikon, Tokyo, Japan) using a Plan Apo 60× water objective [NA=1.27]. Laser lines of 488 nm and 647 nm were used for the excitation of ABCG2-GFP and MX, while fluorescence emissions were detected through band pass filters of 500-550 nm and 660-740 nm, respectively. All the images were recorded with the same settings of the equipment, such as same high voltages, laser powers and pinhole. Images were acquired in sequential mode to minimize the crosstalk between channels. Images of approximately 1 μm thick optical sections, each with 512×512 pixels, and a pixel size of approximately 200 nm, were acquired. A spatial averaging filter with a 3×3 mask was used to denoise the images. Colocalization analysis was carried out by calculating the Pearson’s correlation coefficients between the pixel intensities of the two detection channels in pixels representing the plasma membrane (Vamosi et al., 2004). Only pixels where at least one of the intensities was above the threshold (2× the average auto fluorescence intensity) were included in the analysis. Image analysis methods and routines were implemented in MATLAB scripts (Mathworks Inc., Natick, MA) (Volko et al., 2019).

4.12. Fluorescence correlation spectroscopy (FCS)

To distinguish free and ABCG2-bound MX molecules based on their different diffusion properties, fluorescence correlation spectroscopy (FCS) measurements were performed. FCS measurements were carried out using a Nikon A1 Eclipse Ti2 Confocal Laser-Scanning Microscope (Nikon, Tokyo, Japan), equipped with a Plan Apo 60× water objective [NA=1.27] and a PicoQuant time-correlated single photon counting FCS (TCSPC-FCS) upgrade kit (PicoQuant, Berlin, Germany).

FCS measurements were carried out on live MDCK cells expressing ABCG2-GFP in eight-well chambered coverslip plates (ibidi GmbH, Gräfelfing, Germany). Cells were stained with 100 nM MX for 15 min at 37°C in the presence or absence of 2 μM Ko143 or after ATP depletion. Fluorescence of ABCG2-GFP and MX was excited with a 488 nm and a 647 nm laser, respectively. The fluorescence signals emitted by ABCG2-GFP and MX were detected in the spectral ranges of 500-550 nm and 660-740 nm using single photon counting detectors (PicoQuant, Berlin, Germany). Measurements of 10×10 second runs were taken at three selected points in the cross-section of the plasma membrane of each selected cell. Fluorescence autocorrelation curves were calculated using SymPhoTime64 software (PicoQuant, Berlin, Germany) at 200 time points from 300 ns to 1 s with a quasi-logarithmic time scale.

Autocorrelation curves of the doubly labeled cells were fitted to a triplet state model with two diffusion components to describe the 3D-diffusion of free MX (fast component) and the 2D diffusion of ABCG2-bound MX in the x-z plane of the plasma membrane (slow component). The laser beam was positioned in a region of the cell membrane parallel to the long axis of the ellipsoidal laser volume. Embedded Image

In equation (3), N is the average number of fluorescent molecules in the detection volume, T is the fraction of molecules in the triplet state, τtrip is the triplet correlation time. The diffusion rate is characterized by the diffusion time τD, which is the average time spent by a molecule in the illuminated volume. τD1 and τD2 are the diffusion times of the fast and slow components, ρ is the fraction of the first component and 1-ρ is the fraction of the second component. The diffusion coefficients (D) of the fast and slow components were determined from the following equation: Embedded Image

Wherein, ωxy is the lateral e-2radius of the detection volume. ωxy was measured by determining the diffusion time of 100 nM A647 dye (dissolved in 10 mM Tris, 0.1 mM EDTA-containing buffer, pH 7.4) with known diffusion coefficient (DA647 = 330 μm2/s, at T = 22.5°C) (Weidemann and Schwille, 2013) and substituting it into Equation 4 that corresponds to the aspect ratio of the ellipsoidal confocal volume, defined as the ratio of its axial and radial dimensions. This parameter was estimated by fitting the autocorrelation curves of a 100 nM A647 dye solution.

4.13. Statistical analysis and curve fitting

For the statistical analysis of data, SigmaPlot (version 14, SSI San Jose, CA, USA) was used. For the comparison of two samples from normally distributed populations with equal variances, Student’s t-test was performed, while in case of unequal variances a Kolmogorov–Smirnov test was applied. Multiple comparisons were performed with analysis of variance (ANOVA) applying the Holm–Sidak test for post hoc pair-wise comparison of the data. In the case of unequal variances, the Dunnett T3 post hoc pair-wise comparison method was used. Differences were considered significant at p < 0.05.

All curve fitting was carried out by SigmaPlot (version 14, SSI San Jose, CA, USA) except for fitting of autocorrelation curves that was performed by using the QuickFit 3.0 software developed in the group B040 (Prof. Jörg Langowski) at the German Cancer Research Center (DKFZ).

Statements & Declarations

Data availability

All data generated or analysed during this study are included in this published article.

Funding

We are grateful for the financial supports by the Hungarian National Research, Development and Innovations Office (NKFIH; https://nkfih.gov.hu/english), grant number K124815 to KG, GINOP-2.2.1-15-2017-00079 to JR and KG and HunProtEx 2018-1.2.1-NKP-2018-00005 to GS. GS was supported by a Momentum Grant of the Hungarian Academy of Sciences. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing Interests

The authors have no relevant financial or non-financial interests to disclose.

Author contributions

Conceptualization: Katalin Goda, Gergely Szakács, Thomas Stockner; Methodology: Katalin Goda, Gábor Mocsár, Tamás I Orbán, László Homolya, Judit Remenyik, Formal analysis and investigation: Zsuzsanna Gyöngy, Gábor Mocsár, Zsuzsanna Ritter, Éva Hegedűs, Anita Schamberger; Writing - original draft preparation: Zsuzsanna Gyöngy, Katalin Goda, Gergely Szakács, Thomas Stockner; Writing - review and editing: Katalin Goda, Gergely Szakács, Thomas Stockner, László Homolya; Funding acquisition: Katalin Goda, Gergely Szakács, Judit Remenyik; Supervision: Katalin Goda

All authors read and approved the final manuscript.

Acknowledgments

The authors thank László Csanády for the critical reading of the manuscript and for helpful discussions and Adél Vezendiné Nagy for her skilful technical assistance.

Abbreviations

A647
Alexa 647 succinimidyl ester
ABCP
Placenta-Specific ABC transporter
ADME-Tox
absorption, distribution, metabolism, excretion and toxicity
AMP-PNP
adenylyl-imidodiphosphate
BCRP
Breast Cancer Resistance Protein
E3S
estrone-3-sulfate
cryo-EM
cryogenic electron microscopy
CFTR
Cystic Fibrosis Transmembrane Conductance Regulator (ABCC7)
FCS
fluorescence correlation spectroscopy
FBS
foetal bovine serum
GFP
green fluorescent protein
IF
inward-facing
mAb
monoclonal antibody
MX
mitoxantrone
MXR
Mitoxantrone Resistance Protein
NBD
nucleotide binding domain
OF
outward-facing
PI
propidium iodide
SLO
streptolysin-O
TMD
transmembrane domain
Vi
vanadate

References

  1. ↵
    Arana, M. R. & Altenberg, G. A. 2019. ATP-binding Cassette Exporters: Structure and Mechanism with a Focus on P-glycoprotein and MRP1. Curr Med Chem, 26, 1062–1078.doi:10.2174/0929867324666171012105143
    OpenUrlCrossRef
  2. ↵
    Barsony, O., Szaloki, G., Turk, D., Tarapcsak, S., Gutay-Toth, Z., Bacso, Z., Holb, I. J., Szekvolgyi, L., Szabo, G., Csanady, L., Szakacs, G. & Goda, K. 2016. A single active catalytic site is sufficient to promote transport in P-glycoprotein. Sci Rep, 6, 24810.doi:10.1038/srep24810
    OpenUrlCrossRefPubMed
  3. ↵
    Bordignon, E., Seeger, M. A., Galazzo, L. & Meier, G. 2020. From in vitro towards in situ: structure-based investigation of ABC exporters by electron paramagnetic resonance spectroscopy. FEBS Lett, 594, 3839–3856.doi:10.1002/1873-3468.14004
    OpenUrlCrossRef
  4. ↵
    Catipovic, M. A., Bauer, B. W., Loparo, J. J. & Rapoport, T. A. 2019. Protein translocation by the SecA ATPase occurs by a power-stroke mechanism. EMBO J, 38.doi:10.15252/embj.2018101140
    OpenUrlAbstract/FREE Full Text
  5. ↵
    Ding, X. W., Wu, J. H. & Jiang, C. P. 2010. ABCG2: a potential marker of stem cells and novel target in stem cell and cancer therapy. Life Sci, 86, 631–7.doi:10.1016/j.lfs.2010.02.012
    OpenUrlCrossRefPubMedWeb of Science
  6. ↵
    Doyle, L. A., Yang, W., Abruzzo, L. V., Krogmann, T., Gao, Y., Rishi, A. K. & Ross, D. D. 1998. A multidrug resistance transporter from human MCF-7 breast cancer cells. Proc Natl Acad Sci U S A, 95, 15665–70.doi:10.1073/pnas.95.26.15665
    OpenUrlAbstract/FREE Full Text
  7. ↵
    Erdei, Z., Schamberger, A., Torok, G., Szebenyi, K., Varady, G., Orban, T. I., Homolya, L., Sarkadi, B. & Apati, A. 2018. Generation of multidrug resistant human tissues by overexpression of the ABCG2 multidrug transporter in embryonic stem cells. PLoS One, 13, e0194925.doi:10.1371/journal.pone.0194925
    OpenUrlCrossRef
  8. ↵
    Fetsch, P. A., Abati, A., Litman, T., Morisaki, K., Honjo, Y., Mittal, K. & Bates, S. E. 2006. Localization of the ABCG2 mitoxantrone resistance-associated protein in normal tissues. Cancer Lett, 235, 84–92.doi:10.1016/j.canlet.2005.04.024
    OpenUrlCrossRefPubMedWeb of Science
  9. ↵
    Fisher, A. J., Smith, C. A., Thoden, J. B., Smith, R., Sutoh, K., Holden, H. M. & Rayment, I. 1995. X-ray structures of the myosin motor domain of Dictyostelium discoideum complexed with MgADP.BeFx and MgADP.AlF4. Biochemistry, 34, 8960–72.doi:10.1021/bi00028a004
    OpenUrlCrossRefPubMed
  10. ↵
    Giacomini, K. M., Balimane, P. V., Cho, S. K., Eadon, M., Edeki, T., Hillgren, K. M., Huang, S. M., Sugiyama, Y., Weitz, D., Wen, Y., Xia, C. Q., Yee, S. W., Zimdahl, H., Niemi, M. & INTERNATIONAL TRANSPORTER, C. 2013. International Transporter Consortium commentary on clinically important transporter polymorphisms. Clin Pharmacol Ther, 94, 23–6.doi:10.1038/clpt.2013.12
    OpenUrlCrossRefPubMed
  11. ↵
    Goda, K., Donmez-Cakil, Y., Tarapcsak, S., Szaloki, G., Szollosi, D., Parveen, Z., Turk, D., Szakacs, G., Chiba, P. & Stockner, T. 2020. Human ABCB1 with an ABCB11-like degenerate nucleotide binding site maintains transport activity by avoiding nucleotide occlusion. PLoS Genet, 16, e1009016.doi:10.1371/journal.pgen.1009016
    OpenUrlCrossRef
  12. ↵
    Haider, A. J., Briggs, D., Self, T. J., Chilvers, H. L., Holliday, N. D. & Kerr, I. D. 2011. Dimerization of ABCG2 analysed by bimolecular fluorescence complementation. PLoS One, 6, e25818.doi:10.1371/journal.pone.0025818
    OpenUrlCrossRefPubMed
  13. ↵
    Homolya, L. 2021. Medically Important Alterations in Transport Function and Trafficking of ABCG2. Int J Mol Sci, 22.doi:10.3390/ijms22062786
    OpenUrlCrossRef
  14. ↵
    Homolya, L., Orban, T. I., Csanady, L. & Sarkadi, B. 2011. Mitoxantrone is expelled by the ABCG2 multidrug transporter directly from the plasma membrane. Biochim Biophys Acta, 1808, 154–63.doi:10.1016/j.bbamem.2010.07.031
    OpenUrlCrossRefPubMed
  15. ↵
    Horsey, A. J., Briggs, D. A., Holliday, N. D., Briddon, S. J. & Kerr, I. D. 2020. Application of fluorescence correlation spectroscopy to study substrate binding in styrene maleic acid lipid copolymer encapsulated ABCG2. Biochim Biophys Acta Biomembr, 1862, 183218.doi:10.1016/j.bbamem.2020.183218
    OpenUrlCrossRef
  16. ↵
    Ishikawa, T., Aw, W. & Kaneko, K. 2013. Metabolic Interactions of Purine Derivatives with Human ABC Transporter ABCG2: Genetic Testing to Assess Gout Risk. Pharmaceuticals (Basel), 6, 1347–60.doi:10.3390/ph6111347
    OpenUrlCrossRef
  17. ↵
    Jackson, S. M., Manolaridis, I., Kowal, J., Zechner, M., Taylor, N. M. I., Bause, M., Bauer, S., Bartholomaeus, R., Bernhardt, G., Koenig, B., Buschauer, A., Stahlberg, H., Altmann, K. H. & Locher, K. P. 2018. Structural basis of small-molecule inhibition of human multidrug transporter ABCG2. Nat Struct Mol Biol, 25, 333–340.doi:10.1038/s41594-018-0049-1
    OpenUrlCrossRefPubMed
  18. ↵
    Januliene, D. & Moeller, A. 2020. Cryo-EM of ABC transporters: an ice-cold solution to everything? FEBS Lett, 594, 3776–3789.doi:10.1002/1873-3468.13989
    OpenUrlCrossRef
  19. ↵
    Jardetzky, O. 1966. Simple allosteric model for membrane pumps. Nature, 211, 969–70.doi:10.1038/211969a0
    OpenUrlCrossRefPubMedWeb of Science
  20. ↵
    Jonker, J. W., Buitelaar, M., Wagenaar, E., Van Der Valk, M. A., Scheffer, G. L., Scheper, R. J., Plosch, T., Kuipers, F., Elferink, R. P., Rosing, H., Beijnen, J. H. & Schinkel, A. H. 2002. The breast cancer resistance protein protects against a major chlorophyll-derived dietary phototoxin and protoporphyria. Proc Natl Acad Sci USA, 99, 15649–54.doi:10.1073/pnas.202607599
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Josts, I. Gao, Y., Monteiro, D. C. F., Niebling, S., Nitsche, J., Veith, K., Grawert, T. W., Blanchet, C. E., Schroer, M. A., Huse, N., Pearson, A. R., Svergun, D. I. & Tidow, H. 2020. Structural Kinetics of MsbA Investigated by Stopped-Flow Time-Resolved Small-Angle X-Ray Scattering. Structure, 28, 348–354 e3.doi:10.1016/j.str.2019.12.001
    OpenUrlCrossRef
  22. ↵
    Kage, K., Tsukahara, S., Sugiyama, T., Asada, S., Ishikawa, E., Tsuruo, T. & Sugimoto, Y. 2002. Dominant-negative inhibition of breast cancer resistance protein as drug efflux pump through the inhibition of S-S dependent homodimerization. Int J Cancer, 97, 626–30.doi:10.1002/ijc.10100
    OpenUrlCrossRefPubMedWeb of Science
  23. ↵
    Kapoor, P., Horsey, A. J., Cox, M. H. & Kerr, I. D. 2018. ABCG2: does resolving its structure elucidate the mechanism? Biochem Soc Trans, 46, 1485–1494.doi:10.1042/BST20180145
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Khunweeraphong, N., Stockner, T. & Kuchler, K. 2017. The structure of the human ABC transporter ABCG2 reveals a novel mechanism for drug extrusion. Sci Rep, 7, 13767.doi:10.1038/s41598-017-11794-w
    OpenUrlCrossRef
  25. ↵
    Khunweeraphong, N., Szollosi, D., Stockner, T. & Kuchler, K. 2019. The ABCG2 multidrug transporter is a pump gated by a valve and an extracellular lid. Nat Commun, 10, 5433.doi:10.1038/s41467-019-13302-2
    OpenUrlCrossRef
  26. ↵
    Kottgen, A. & Kottgen, M. 2021. A novel mouse model of hyperuricemia expressing a human functional ABCG2 variant. Kidney Int, 99, 12–14.doi:10.1016/j.kint.2020.10.021
    OpenUrlCrossRef
  27. ↵
    Li, M. J., Nath, A. & Atkins, W. M. 2017. Differential Coupling of Binding, ATP Hydrolysis, and Transport of Fluorescent Probes with P-Glycoprotein in Lipid Nanodiscs. Biochemistry, 56, 2506–2517.doi:10.1021/acs.biochem.6b01245
    OpenUrlCrossRefPubMed
  28. ↵
    Locher, K. P. 2009. Review. Structure and mechanism of ATP-binding cassette transporters. Philos Trans R Soc Lond B Biol Sci, 364, 239–45.doi:10.1098/rstb.2008.0125
    OpenUrlCrossRefPubMed
  29. ↵
    Locher, K. P. 2016. Mechanistic diversity in ATP-binding cassette (ABC) transporters. Nat Struct Mol Biol, 23, 487–93.doi:10.1038/nsmb.3216
    OpenUrlCrossRefPubMed
  30. ↵
    Lusvarghi, S., Durell, S. R. & Ambudkar, S. V. 2021. Does the ATP-bound EQ mutant reflect the pre-or post-ATP hydrolysis state in the catalytic cycle of human P-glycoprotein (ABCB1)? FEBS Lett, 595, 750–762.doi:10.1002/1873-3468.14054
    OpenUrlCrossRef
  31. ↵
    Manolaridis, I., Jackson, S. M., Taylor, N. M. I., Kowal, J., Stahlberg, H. & Locher, K. P. 2018. Cryo-EM structures of a human ABCG2 mutant trapped in ATP-bound and substrate-bound states. Nature, 563, 426–430.doi:10.1038/s41586-018-0680-3
    OpenUrlCrossRef
  32. ↵
    Martin, C., Higgins, C. F. & Callaghan, R. 2001. The vinblastine binding site adopts high-and low-affinity conformations during a transport cycle of P-glycoprotein. Biochemistry, 40, 15733–42.doi:10.1021/bi011211z
    OpenUrlCrossRefPubMed
  33. ↵
    Mcdevitt, C. A., Crowley, E., Hobbs, G., Starr, K. J., Kerr, I. D. & Callaghan, R. 2008. Is ATP binding responsible for initiating drug translocation by the multidrug transporter ABCG2? FEBS J, 275, 4354–62.doi:10.1111/j.1742-4658.2008.06578.x
    OpenUrlCrossRefPubMed
  34. ↵
    Orban, T. I., Seres, L., Ozvegy-Laczka, C., Elkind, N. B., Sarkadi, B. & Homolya, L. 2008. Combined localization and real-time functional studies using a GFP-tagged ABCG2 multidrug transporter. Biochem Biophys Res Commun, 367, 667–73.doi:10.1016/j.bbrc.2007.12.172
    OpenUrlCrossRefPubMed
  35. ↵
    Orlando, B. J. & Liao, M. 2020. ABCG2 transports anticancer drugs via a closed-to-open switch. Nat Commun, 11, 2264.doi:10.1038/s41467-020-16155-2
    OpenUrlCrossRef
  36. ↵
    Ozvegy-Laczka, C., Varady, G., Koblos, G., Ujhelly, O., Cervenak, J., Schuetz, J. D., Sorrentino, B. P., Koomen, G. J., Varadi, A., Nemet, K. & Sarkadi, B. 2005. Functiondependent conformational changes of the ABCG2 multidrug transporter modify its interaction with a monoclonal antibody on the cell surface. J Biol Chem, 280, 4219–27.doi:10.1074/jbc.M411338200
    OpenUrlAbstract/FREE Full Text
  37. ↵
    Ozvegy, C., Litman, T., Szakacs, G., Nagy, Z., Bates, S., Varadi, A. & Sarkadi, B. 2001. Functional characterization of the human multidrug transporter, ABCG2, expressed in insect cells. Biochem Biophys Res Commun, 285, 111–7.doi:10.1006/bbrc.2001.5130
    OpenUrlCrossRefPubMedWeb of Science
  38. ↵
    Pal, A., Mehn, D., Molnar, E., Gedey, S., Meszaros, P., Nagy, T., Glavinas, H., Janaky, T., Von Richter, O., Bathori, G., Szente, L. & Krajcsi, P. 2007. Cholesterol potentiates ABCG2 activity in a heterologous expression system: improved in vitro model to study function of human ABCG2. J Pharmacol Exp Ther, 321, 1085–94.doi:10.1124/jpet.106.119289
    OpenUrlAbstract/FREE Full Text
  39. ↵
    Romsicki, Y. & Sharom, F. J. 1998. The ATPase and ATP-binding functions of P-glycoprotein--modulation by interaction with defined phospholipids. Eur J Biochem, 256, 170–8.doi:10.1046/j.1432-1327.1998.2560170.x
    OpenUrlCrossRefPubMedWeb of Science
  40. ↵
    Rothnie, A., Callaghan, R., Deeley, R. G. & Cole, S. P. 2006. Role of GSH in estrone sulfate binding and translocation by the multidrug resistance protein 1 (MRP1/ABCC1). J Biol Chem, 281, 13906–14.doi:10.1074/jbc.M600869200
    OpenUrlAbstract/FREE Full Text
  41. ↵
    Sarkadi, B., Homolya, L., Szakacs, G. & Varadi, A. 2006. Human multidrug resistance ABCB and ABCG transporters: participation in a chemoimmunity defense system. Physiol Rev, 86, 1179–236.doi:10.1152/physrev.00037.2005
    OpenUrlCrossRefPubMedWeb of Science
  42. ↵
    Sarkadi, B., Price, E. M., Boucher, R. C., Germann, U. A. & Scarborough, G. A. 1992. Expression of the human multidrug resistance cDNA in insect cells generates a high activity drug-stimulated membrane ATPase. J Biol Chem, 267, 4854–8
    OpenUrlAbstract/FREE Full Text
  43. ↵
    Seeger, M. A. & Van Veen, H. W. 2009. Molecular basis of multidrug transport by ABC transporters. Biochim Biophys Acta, 1794, 725–37.doi:10.1016/j.bbapap.2008.12.004
    OpenUrlCrossRefPubMedWeb of Science
  44. ↵
    Smith, C. A. & Rayment, I. 1996. X-ray structure of the magnesium(II).ADP.vanadate complex of the Dictyostelium discoideum myosin motor domain to 1.9 A resolution. Biochemistry, 35, 5404–17.10.1021/bi952633+
    OpenUrlCrossRefPubMedWeb of Science
  45. ↵
    Stockner, T., Gradisch, R. & Schmitt, L. 2020. The role of the degenerate nucleotide binding site in type I ABC exporters. FEBS Lett, 594, 3815–3838.doi:10.1002/1873-3468.13997
    OpenUrlCrossRef
  46. ↵
    Suzuki, M., Suzuki, H., Sugimoto, Y. & Sugiyama, Y. 2003. ABCG2 transports sulfated conjugates of steroids and xenobiotics. J Biol Chem, 278, 22644–9.doi:10.1074/jbc.M212399200
    OpenUrlAbstract/FREE Full Text
  47. ↵
    Szabo, K., Welker, E., BAKOS, Muller, M., Roninson, I., Varadi, A. & Sarkadi, B. 1998. Drug-stimulated nucleotide trapping in the human multidrug transporter MDR1. Cooperation of the nucleotide binding domains. J Biol Chem, 273, 10132–8.doi:10.1074/jbc.273.17.10132
    OpenUrlAbstract/FREE Full Text
  48. ↵
    Szakacs, G., Ozvegy, C., Bakos, E., Sarkadi, B. & Varadi, A. 2000. Transition-state formation in ATPase-negative mutants of human MDR1 protein. Biochem Biophys Res Commun, 276, 1314–9.doi:10.1006/bbrc.2000.3576
    OpenUrlCrossRefPubMedWeb of Science
  49. ↵
    Tarapcsak, S., Szaloki, G., Telbisz, A., Gyongy, Z., Matuz, K., Csosz, E., Nagy, P., Holb, I. J., Ruhl, R., Nagy, L., Szabo, G. & Goda, K. 2017. Interactions of retinoids with the ABC transporters P-glycoprotein and Breast Cancer Resistance Protein. Sci Rep, 7, 41376.doi:10.1038/srep41376
    OpenUrlCrossRef
  50. ↵
    Taylor, N. M. I., Manolaridis, I., Jackson, S. M., Kowal, J., Stahlberg, H. & Locher, K. P. 2017. Structure of the human multidrug transporter ABCG2. Nature, 546, 504–509.doi:10.1038/nature22345
    OpenUrlCrossRefPubMed
  51. ↵
    Telbisz, A., Hegedus, C., Ozvegy-Laczka, C., Goda, K., Varady, G., Takats, Z., Szabo, E., Sorrentino, B. P., Varadi, A. & Sarkadi, B. 2012. Antibody binding shift assay for rapid screening of drug interactions with the human ABCG2 multidrug transporter. Eur J Pharm Sci, 45, 101–9.doi:10.1016/j.ejps.2011.10.021
    OpenUrlCrossRefPubMed
  52. ↵
    Vamosi, G., Bodnar, A., Vereb, G., Jenei, A., Goldman, C. K., Langowski, J., Toth, K., Matyus, L., Szollosi, J., Waldmann, T. A. & Damjanovich, S. 2004. IL-2 and IL-15 receptor alphasubunits are coexpressed in a supramolecular receptor cluster in lipid rafts of T cells. Proc Natl Acad Sci U S A, 101, 11082–7.doi:10.1073/pnas.0403916101
    OpenUrlAbstract/FREE Full Text
  53. ↵
    Van Herwaarden, A. E. & Schinkel, A. H. 2006. The function of breast cancer resistance protein in epithelial barriers, stem cells and milk secretion of drugs and xenotoxins. Trends Pharmacol Sci, 27, 10–6.doi:10.1016/j.tips.2005.11.007
    OpenUrlCrossRefPubMed
  54. ↵
    Van Wonderen, J. H., Mcmahon, R. M., O’mara, M. L., Mcdevitt, C. A., Thomson, A. J., Kerr, I. D., Macmillan, F. & Callaghan, R. 2014. The central cavity of ABCB1 undergoes alternating access during ATP hydrolysis. FEBS J, 281, 2190–2201.doi:10.1111/febs.12773
    OpenUrlCrossRefPubMed
  55. ↵
    1. Roberts, G. C. K.
    Vergani, P., Gadsby, D. C. & Csanády, L. 2013. CFTR, an Ion Channel Evolved from ABC Transporter. In: Roberts, G. C. K. (ed.) Encyclopedia of Biophysics. Berlin, Heidelberg: Springer Berlin Heidelberg.doi:10.1007/978-3-642-16712-6_364
    OpenUrlCrossRef
  56. ↵
    Volko, J., Kenesei, A., Zhang, M., Varnai, P., Mocsar, G., Petrus, M. N., Jambrovics, K., Balajthy, Z., Muller, G., Bodnar, A., Toth, K., Waldmann, T. A. & Vamosi, G. 2019. IL-2 receptors preassemble and signal in the ER/Golgi causing resistance to antiproliferative anti-IL-2Ralpha therapies. Proc Natl Acad Sci U S A, 116, 21120–21130.doi:10.1073/pnas.1901382116
    OpenUrlAbstract/FREE Full Text
  57. ↵
    Wang, L., Johnson, Z. L., Wasserman, M. R., Levring, J., Chen, J. & Liu, S. 2020. Characterization of the kinetic cycle of an ABC transporter by single-molecule and cryo-EM analyses. Elife, 9.doi:10.7554/eLife.56451
    OpenUrlCrossRefPubMed
  58. ↵
    Weidemann, T. & Schwille, P. 2013. Dual-color fluorescence cross-correlation spectroscopy with continuous laser excitation in a confocal setup. Methods Enzymol, 518, 43–70.doi:10.1016/B978-0-12-388422-0.00003-0
    OpenUrlCrossRefPubMed
  59. ↵
    Williams, S. P., Fulton, A. M. & Brindle, K. M. 1993. Estimation of the intracellular free ADP concentration by 19F NMR studies of fluorine-labeled yeast phosphoglycerate kinase in vivo. Biochemistry, 32, 4895–902.doi:10.1021/bi00069a026
    OpenUrlCrossRefPubMed
  60. ↵
    Woodward, O. M., Kottgen, A., Coresh, J., Boerwinkle, E., Guggino, W. B. & Kottgen, M. 2009. Identification of a urate transporter, ABCG2, with a common functional polymorphism causing gout. Proc Natl Acad Sci U S A, 106, 10338–42.doi:10.1073/pnas.0901249106
    OpenUrlAbstract/FREE Full Text
  61. ↵
    Yang, W. S., Park, S. O., Yoon, A. R., Yoo, J. Y., Kim, M. K., Yun, C. O. & Kim, C. W. 2006. Suicide cancer gene therapy using pore-forming toxin, streptolysin O. Mol Cancer Ther, 5, 1610–9.doi:10.1158/1535-7163.MCT-05-0515
    OpenUrlCrossRefPubMed
  62. ↵
    Yu, Q., Ni, D., Kowal, J., Manolaridis, I., Jackson, S. M., Stahlberg, H. & Locher, K. P. 2021. Structures of ABCG2 under turnover conditions reveal a key step in the drug transport mechanism. Nat Commun, 12, 4376.doi:10.1038/s41467-021-24651-2
    OpenUrlCrossRef
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Posted October 21, 2022.
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Nucleotide binding is the critical regulator of ABCG2 conformational transitions
Zsuzsanna Gyöngy, Gábor Mocsár, Éva Hegedűs, Thomas Stockner, Zsuzsanna Ritter, László Homolya, Anita Schamberger, Tamás I. Orbán, Judit Remenyik, Gergely Szakács, Katalin Goda
bioRxiv 2022.10.19.512890; doi: https://doi.org/10.1101/2022.10.19.512890
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