Abstract
Mitochondrial RNA splicing 2 (MRS2) forms a magnesium (Mg2+) entry protein channel in mitochondria. Whereas MRS2 contains two transmembrane domains constituting a pore on the inner mitochondrial membrane, most of the protein resides within the matrix. Yet, the precise structural and functional role of this obtrusive amino terminal domain (NTD) in human MRS2 is unknown. Here, we show that the MRS2 NTD self-associates into a homodimer, contrasting the pentameric assembly of CorA, an orthologous bacterial channel. Mg2+ and calcium suppress lower and higher order oligomerization of MRS2 NTD, whereas cobalt has no effect on the NTD but disassembles full-length MRS2. Mutating-pinpointed residues-mediating Mg2+ binding to the NTD not only selectively decreases Mg2+-binding affinity ∼sevenfold but also abrogates Mg2+ binding–induced secondary, tertiary, and quaternary structure changes. Disruption of NTD Mg2+ binding strikingly potentiates mitochondrial Mg2+ uptake in WT and Mrs2 knockout cells. Our work exposes a mechanism for human MRS2 autoregulation by negative feedback from the NTD and identifies a novel gain of function mutant with broad applicability to future Mg2+ signaling research.
Introduction
Magnesium ions (Mg2+) are the most abundant divalent cations in eukaryotes, playing universal roles in myriad cell functions (Jahnen-Dechent & Ketteler, 2012). Within mitochondria, Mg2+ is an important protein-stabilizing cofactor, forms biologically functional Mg2+–ATP complexes, and regulates crucial enzymatic activities. Such roles are achieved through two unique properties of Mg2+: (i) the ability to form chelates with important intracellular anionic-ligands (i.e., small molecule or large biomolecule), and (ii) the capability to compete with calcium ions (Ca2+) for binding sites on proteins and membranes (Berridge et al, 2000; Moomaw & Maguire, 2008; Chaigne-Delalande et al, 2013). The effects of Mg2+ on Ca2+-handling proteins significantly influence intracellular Ca2+ dynamics and signaling (Gregan et al, 2001; Clapham, 2007; de Baaij et al, 2015).
Total cellular Mg2+ concentrations range between ∼17 and 30 mM; however, concentrations of free Mg2+ in the cytosol are estimated between ∼0.5 and 1.5 mM (Jung et al, 1990; Rutter et al, 1990; Romani, 2011). Intracellular Mg2+ concentrations are strongly buffered and regulated by the combined action of Mg2+-binding molecules, Mg2+ storage in organelles, and the action of Mg2+ channels and exchangers. Remarkably, Mg2+ can be mobilized from the ER in response to ligands such as L-lactate, moving into the mitochondria and dramatically modifying metabolism (Daw et al, 2020). Mg2+ can also alter the electrophysiological properties of ion channels such as voltage-dependent Ca2+ channels and potassium (K+) channels and affect the binding affinity of Ca2+ to EF-hand–containing proteins (Glancy & Balaban, 2012; Pilchova et al, 2017). All ATP-related biochemical reactions in cells are dependent on Mg2+ (Romani, 2007), and extracellular Mg2+ also regulates numerous channels such as glutamate receptors and N-methyl-D-aspartate receptors (Kumar, 2015). In addition, this divalent cation contributes to the maintenance of genome stability as a cofactor in DNA repair and protection (Hartwig, 2001).
Unsurprisingly, perturbations in intracellular Mg2+ concentrations can cause serious cellular dysfunction. For example, decreases in intracellular free Mg2+ lead to defective immune responses (Zhou & Clapham, 2009; Chaigne-Delalande et al, 2013; Kanellopoulou et al, 2019), mutations in the Na+/Mg2+ exchanger causing chronic intracellular Mg2+ deficiency trigger neuronal damage (Kolisek et al, 2013), and overexpression of Mg2+ channels is a hallmark of several types of cancer (Trapani & Wolf, 2019), to name a few. More specifically in terms of neuronal disease, the A350V mutation in SLC41A1 enhances Na+-dependent Mg2+ efflux by ∼70% in HEK293 cells (Kolisek et al, 2013), and low Mg2+ intake in rats and humans leads to loss of dopaminergic neurons (Oyanagi et al, 2006) and increased risk of idiopathic Parkinson’s disease, respectively (Aden et al, 2011). Indeed, decreased free cytosolic Mg2+ has been measured in the occipital lobes of Parkinson’s patients (Barbiroli et al, 1999). Transient receptor potential melastatin-6 mutations can also be pathophysiological, resulting in Mg2+ malabsorption and renal wasting, where affected individuals show seizures and muscle spasms during infancy (Schlingmann et al, 2002; Schaffers et al, 2018).
Mitochondria have an inner mitochondrial membrane (IMM), separating the mitochondrial matrix (MM) from the intermembrane space, and an outer mitochondrial membrane, enclosing the entire organelle. Unregulated, the highly negative IMM potential (∼−180 mV) (Marchi & Pinton, 2014) would drive catastrophically high concentrations of Mg2+ entry into the matrix; nevertheless, the matrix Mg2+ concentration is similar to the concentration in the cytoplasm, reinforcing that Mg2+ influx into the organelle is tightly controlled to maintain optimal mitochondrial function and bioenergetics (Marchi & Pinton, 2014; Pilchova et al, 2017).
Residing within the IMM in mammalian cells, mitochondrial RNA splicing 2 (MRS2) constitutes a major Mg2+ entry protein channel into the MM. Deletion of IMM-localized MRS2 abolishes Mg2+ influx into the MM, inducing functional defects in mitochondria and promoting cell death (Piskacek et al, 2009; Merolle et al, 2018). MRS2 belongs to the large heterogeneous CorA/Mrs2/Alr1 protein superfamily of Mg2+ transporters. This family is characterized by the highly conserved Gly-Met-Asn (GMN) motif at the end of the first transmembrane helix, essential for Mg2+ transporter function. Mutations of the GMN motif either completely abolish Mg2+ transport or profoundly change the ion selectivity of the channel (Knoop et al, 2005; Palombo et al, 2013; Merolle et al, 2018). Human MRS2 contains a large, amino terminal domain (NTD) oriented within the MM, corresponding to residues 58–333 and consisting of ∼71% of the mature polypeptide chain, two transmembrane (TM1 and TM2) domains connected by a highly conserved intermembrane space loop and a smaller, carboxyl terminal domain also oriented within the MM (Fig 1A).
Although CorA and Mrs2, orthologues of MRS2 in bacteria and yeast, respectively, have been structurally resolved at high resolution (Eshaghi et al, 2006; Lunin et al, 2006; Payandeh & Pai, 2006; Guskov et al, 2012; Pfoh et al, 2012; Khan et al, 2013; Matthies et al, 2016; Johansen et al, 2022), low sequence similarity exists between CorA, Mrs2, and MRS2, especially in the NTDs (Zsurka et al, 2001). Specifically, the sequence similarity between human MRS2 and yeast (Saccharomyces cerevisiae) Mrs2 is 55.4% (and only 20.1% through the NTD), whereas the sequence similarity between human MRS2 and bacterial (Thermotoga maritima) CorA is 43.3% (and only 17.0% through the NTD) (Fig 1B). To reveal how the prominent MRS2 NTD governs the assembly and function of the full channel, we generated recombinant human NTD protein (MRS258–333; residues 58–333) and full-length human MRS2 (MRS258–443; residues 58–443). Using light scattering and chromatographic approaches, we find that in the absence of divalent cations, the NTD self-associates into a homodimer under dilute conditions, whereas both Mg2+ and Ca2+, but not cobalt (Co2+), suppress the self-association of the domain. In contrast, Co2+ disassembles full-length MRS2, whereas Mg2+ and Ca2+ have no effect on stoichiometry. 8-Anilino-1-naphthalene sulfonate (ANS) and intrinsic fluorescence measurements suggest that Mg2+ and Ca2+ bind to distinct sites on the NTD with ∼μM and mM affinity, respectively. Importantly, we identify the D216 and D220 as critical residues for Mg2+ coordination to the human MRS2 NTD, where mutating these residues decreases Mg2+ affinity ∼sevenfold, abrogates Mg2+ binding–induced increases in α-helicity and solvent accessible hydrophobicity, and suppresses the Mg2+-induced disassembly of the NTD. Finally, using both permeabilized and intact cell models, we show that Mg2+ binding to the NTD suppresses the rate of Mg2+ uptake into mitochondria as negative feedback. Collectively, our data reveal previously unknown mechanistic insights underlying human MRS2 autoregulation by the large NTD, which has important implications for understanding the crosstalk between MM Mg2+ concentrations, bioenergetics, and cell death.
Results
MRS2 NTD homodimer assembly is sensitive to divalent cations
Given that human MRS2 encodes only two putative TMs (Fig 1A) and must oligomerize to form a channel pore, we first evaluated the stoichiometry of the MRS2 NTD (i.e., MRS258–333) using size exclusion chromatography with in-line multi-angle light scattering (SEC-MALS). Recombinant MRS258–333 was successfully expressed and isolated with high yield (i.e., ∼8 g/l of culture) and purity (Fig 1C). Although the theoretical monomeric molecular weight of MRS258–333 is ∼32.2 kD, SEC-MALS revealed that, in the absence of divalent cations, MRS258–333 consistently self-associates into a homodimer with estimated molecular weights of 60.9 ± 1.8 kD and 61.3 ± 0.50 kD at 2.5 and 5.0 mg/ml, respectively (Fig 2A and B). Homodimer formation is apparently tight as single elution peaks and no protein concentration dependence on the molecular weight or elution volume in the 0.45–5 mg/ml range was observed (Table S1).
SEC-MALS was further used to assess the sensitivity of the homodimer assembly to Mg2+ and Ca2+ because earlier studies showed divalent cation binding to the CorA NTD regulates channel structure and function (Pfoh et al, 2012; Khan et al, 2013) (see also Discussion section). The presence of 5 mM MgCl2 in the elution buffer transitioned the molecular weight of MRS258–333 to monomer at 2.5 mg/ml, with a SEC-MALS molecular weight of 29.4 ± 6.0 kD (Fig 2C). In contrast, MRS258–333 remained dimeric at 5 mg/ml in the presence of 5 mM MgCl2, with a molecular weight of 60.06 ± 0.47 kD (Fig 2D). Adding 10 mM MgCl2 to the 5 mg/ml sample, however, resulted in a monomeric molecular weight of 32.6 ± 0.5 kD (Fig 2E). Remarkably, adding 5 mM CaCl2 to both the 2.5 and 5.0 mg/ml MRS258–33 samples robustly caused monomer formation, with measured molecular weights of 31.4 ± 1.0 and 29.0 ± 0.44 kD, respectively (Fig 2F and G). Supplementation with 5 mM MgCl2 to MRS258–333 samples at less than 2.5 mg/ml was sufficient to cause monomerization, and shifts to later elution volumes were consistent with all divalent cation-dependent disassembly observations (Table S1).
Divalent cations regulate MRS2 assembly in a domain-specific manner
Because SEC-MALS was performed at 10°C and is accompanied by a large column dilution (i.e., minimum ∼20-fold) that could affect assembly, we used dynamic light scattering (DLS) to assess the distribution of hydrodynamic radii (Rh) at 1.25 mg/ml in the absence of dilution and at higher temperature (i.e., 20 and 37°C). Bimodal distributions of Rh centered at ∼4 and 40 nm were observed for MRS258–333 in the absence of divalent cations at both 20 and 37°C (Fig 3A–F). The addition of either 5 mM CaCl2 (Fig 3A and B) or 5 mM MgCl2 (Fig 3C and D) at both temperatures eliminated the larger size distributions. The loss of larger Rh was supported qualitatively by earlier decays in the autocorrelation functions when compared with divalent cation-free protein samples (Fig 3A–D, insets). The bacterial orthologue of MRS2 and CorA can transport Co2+ and Mg2+, and Co2+ is found in trace levels in mammals (Tapiero et al, 2003; Guskov & Eshaghi, 2012; Czarnek et al, 2015); thus, we also assessed the sensitivity of the MRS2 NTD assembly to Co2+. The distribution of Rh was not affected by the addition of 5 mM CoCl2 at either 20 or 37°C (Fig 3E and F).
Next, we evaluated the effects of Mg2+, Ca2+, and Co2+ on the assembly of the full-length protein. Full-length MRS2, excluding the mitochondrial targeting sequence, (MRS258–443), was successfully expressed and isolated with high purity (Fig 1D). The experimental buffer for MRS258–443 included CHAPS and showed autocorrelation functions consistent with the presence of ∼1–1.5 nm micelles at 37°C (Fig 4A–C). MRS258–443 samples showed autocorrelation functions with later decay times compared with buffer alone, which were deconvoluted to Rh distributions centered at ∼4 and ∼20 nm at 37°C (Fig 4A–C). In contrast to the NTD data, changes in autocorrelation functions and size distributions were not observed with MRS258–443 when supplemented with either 5 mM MgCl2 or 5 mM CaCl2 (Fig 4A and B). Remarkably, 5 mM CoCl2 completely abrogated the larger Rh distributions, which was qualitatively supported by autocorrelation function shifts to earlier decay times (Fig 4C).
Applying a cumulative deconvolution to extract one weight-averaged Rh from all autocorrelation functions reinforced the regularization/polydisperse deconvolution trends described above. Specifically, CoCl2 caused a robust decrease in the weight-averaged Rh for full-length MRS2, but not the NTD; MgCl2 and CaCl2 decreased Rh for the NTD, but not full-length MRS2 (Table S2). Collectively, these data suggest a domain-specific sensitivity to divalent cations, where NTD disassembly is promoted by Mg2+ and Ca2+, whereas Co2+ de-oligomerizes full-length MRS2 because of sensitivity outside the NTD.
Mg2+ and Ca2+ bind to distinct sites on the MRS2 NTD with disparate affinities
Given that both Mg2+ and Ca2+ dissociate MRS258–333, which contains 3×Trp and 7×Tyr residues, we next used changes in intrinsic fluorescence to evaluate divalent cation binding. Fluorescence emission spectra were acquired using an excitation wavelength of 280 nm as a function of increasing MgCl2, CaCl2, and CoCl2 concentrations. The intensities of the fluorescence emission spectra decreased as a function of increasing MgCl2 (Fig 5A) and CaCl2 (Fig 5B) concentrations. Both MgCl2 and CaCl2 effects were saturable; however, the intensity decreased by ∼32% with Mg2+ and only ∼5% with Ca2+, suggesting distinct structural effects and/or binding sites. In contrast, titration with CoCl2 caused small increases of ∼2% in fluorescence (Fig 5C). Fitting the binding curves to a one-site binding model that accounts for protein concentrations revealed apparent equilibrium dissociation constants (Kd)s of ∼0.14 ± 0.03, 1.01 ± 0.26, and 0.68 ± 0.30 mM for Mg2+, Ca2+, and Co2+ interactions, respectively (Table S3).
We next attempted to pinpoint the residues involved in Mg2+ coordination using the CorA crystal structure (4EED.pdb) (Pfoh et al, 2012) as a guide. Note that available yeast Mrs2 structures do not resolve any Mg2+ ions bound to the NTD. The T. maritima CorA crystal structure shows that two Asp residues, separated by three residues (i.e., DALVD) are involved in Mg2+ coordination at one site. Remarkably, human MRS2 contains the same DALVD sequence stretch in the homologous domain, which we posited could similarly coordinate Mg2+ (Fig 1B). However, sequence-based alignment of the bacterial and vertebrate DALVD regions are algorithm-dependent because of the poor sequence conservation: T-Coffee (Notredame et al, 2000) aligns the human and T. maritima DALVD stretches as conserved (Fig S1), whereas Clustal Ω (Sievers et al, 2011) does not (Figs 1B and S2). Moreover, the AlphaFold2 prediction for human MRS2 (see the Discussion section) agrees with the non-conserved Clustal Ω positioning of bacterial and vertebrate DALVD regions. Thus, our use of the term DALVD with respect to human MRS2 refers to the stretch of polypeptide chain containing the D216 and D220 residues and does not imply a motif that is conserved with T. maritima CorA DALVD. We use the term DALVD to highlight that the two Asp residues are at positions i and i+4 of this sequence stretch with no intervening helix-breaking residues, putatively orienting these side chains on the same side of an α-helix and adjacent in three-dimensional (3D) space and permitting both Asp to interact with the same Mg2+ ion.
After creating a D216A/D220A MRS258–333 double mutant, we reassessed divalent cation binding by intrinsic fluorescence. A double mutant was created because both Asp side chains coordinate the same Mg2+ ion in the CorA DALVD sequence, both Asp side chains are close in 3D space in the α-helix where most of the human MRS2 region is predicted to exist by AlphaFold2 (see the Discussion section), and the sub-mM Mg2+ Kd (Table S3) suggests both Asp are involved in the coordination. Not only did the D216A/D220A mutant show a small increase in fluorescence (Mg2+ causes a large decrease in the fluorescence intensity of WT MRS258–333; see above) but also a markedly suppressed intensity change as a function of increasing MgCl2, consistent with perturbation of Mg2+ binding (Fig 5D). In contrast, the CaCl2 and CoCl2 effects were similar to data acquired using WT MRS258–333 (Fig 5E and F). Indeed, fitting the datasets to one-site binding models revealed apparent equilibrium dissociation constants (Kd) of ∼0.98 ± 0.25, 0.74 ± 0.49, and 1.37 ± 0.51 mM (Table S3), consistent with disruption of the Mg2+ interactions but not Ca2+ or Co2+.
Taken together, these data suggest that Mg2+ and Ca2+ bind to distinct sites on the MRS2NTD, with D216 and D220 mediating interactions with Mg2+.
Mg2+ enhances whereas Ca2+ suppresses solvent exposed hydrophobicity of MRS2 NTD
Given the changes in stoichiometry observed by DLS and SEC-MALS, we next assessed the solvent-exposed hydrophobicity of MRS258–333 in the absence and presence of Mg2+ and Ca2+ by monitoring extrinsic ANS fluorescence. ANS binds to solvent accessible hydrophobic regions on biomolecules, resulting in a blue-shifted fluorescence emission maximum and increased intensity (Stryer, 1965). Baseline fluorescence emission spectra of ANS in the presence of buffer alone were insensitive to the addition of 5 mM MgCl2 or 5 mM CaCl2 (Fig S3A and B). Indeed, ANS binding was detected in the presence of 2.5 mg/ml MRS258–333, as evidenced by the blue-shifted fluorescence emission maximum and increased intensity compared with the buffer controls (Fig S3A and B). Supplementing the protein samples with 5 mM MgCl2 caused a small but significant increase in ANS fluorescence intensity, suggesting enhanced exposed hydrophobicity (Fig 6A and B). Conversely, supplementation with 5 mM CaCl2 caused a small but significant decrease in ANS fluorescence intensity, indicating decreased solvent exposed hydrophobicity (Fig 6C and D).
We next performed a similar set of experiments with the D216A/D220A MRS258–333 protein. Consistent with our observation that this double mutant disrupts Mg2+ but not Ca2+ binding to the NTD; ANS emission spectra in the presence of protein showed no differences with or without MgCl2 supplementation (Figs 6E and F and S3C), whereas CaCl2 supplementation caused a small but significant decrease in ANS fluorescence intensity (Figs 6G and H and S3D). This ANS binding data reinforces the notion of disparate Ca2+- and Mg2+-binding sites and suggests that these divalent cations may cause distinct MRS2 NTD conformational changes.
D216A/D220A mitigates Mg2+-dependent disassembly of the MRS2 NTD
Next, we tested whether the D216A/D220A double mutation could abolish the Mg2+-dependent monomerization and decreased Rh observed with WT MRS2 NTD. In the absence of the divalent cation, SEC-MALS revealed that D216A/D220A MRS258–333 elutes as a homodimer with a molecular weight of 59.7 ± 2.0 kD when injected at 2.5 mg/ml (Fig 7A), similar to WT MRS258–333 (Table S1). In contrast to WT evaluated at 2.5 mg/ml, however, the SEC-MALS–determined molecular weight of the double mutant remained dimeric (i.e., 59.0 ± 2.4 kD) after the addition of 5 mM MgCl2 (Fig 7B). We also assessed whether Mg2+ could alter Rh of the D216A/D220A MRS258–333 by DLS. Addition of 5 mM MgCl2 neither altered the distribution of Rh nor the autocorrelation function compared with samples evaluated in the absence of the cation (Fig 7C). It is to be noted that a bimodal distribution of Rh centered at ∼4 and ∼40 nm was observed with the D216A/D220A MRS258–333 protein (Fig 7C), similar to WT.
Together, these light scattering analyses demonstrate that Mg2+-dependent disassembly of the MRS2 NTD requires the D216 and D220 residues, where double mutation to Ala abrogates quaternary structure sensitivity to the cation.
D216A/D220A abrogates increased α-helicity and thermal stability in the MRS2 NTD caused by Mg2+ binding
Having observed that Mg2+ binding affects the quaternary and tertiary levels of MRS2 NTD structure, we next used far-UV circular dichroism (CD) spectroscopy to assess the secondary structure. At 37°C, MRS258–333 displayed well-defined mean residue ellipticity minima at ∼208 and ∼222 nm, indicating high levels of α-helicity (Fig 8A). Remarkably, addition of 5 mM MgCl2 directly to the cuvette resulted in an increase in α-helicity, evidenced by more intense negative ellipticity at ∼208 and 222 nm (Fig 8A and C). Similar results were observed for MRS258–333 at 20°C (Fig S4A).
To gain further evidence that D216 and D220 play a critical role in Mg2+ binding to the NTD, we also acquired far-UV CD spectra using D216A/D220A MRS258–333. The far-UV CD spectrum of the double mutant showed a similar level of negative ellipticity as WT with two well-defined minima at ∼208 and ∼222 nm (Fig 8A and B), suggesting that secondary structure folding was not perturbed by the D216A/D220A substitutions. Unlike WT, adding 5 mM MgCl2 directly to the cuvette did not significantly alter the ellipticity for the double mutant (Fig 8D). Unchanging spectra after 5 mM MgCl2 addition were also observed for the double mutant at 20°C (Fig S4B).
We next evaluated thermal stability by monitoring the change in far-UV CD ellipticity at 222 nm as a function of increasing temperature. The thermal melts of MRS258–333 acquired in the absence of Mg2+ exhibited a mean Boltzmann sigmoidal–fitted midpoint of temperature denaturation (Tm) of 51 ± 0.62 °C (Fig 8E). Protein samples supplemented with 5 mM MgCl2 were stabilized by ∼7°C as the mean Tm shifted to 58 ± 0.36°C (Fig 8E). Thermal melt experiments with the D216A/D220A MRS258–333 protein revealed similar mean Tm values of 52 ± 0.70°C and 52 ± 0.64°C in the presence and absence of Mg2+, respectively (Fig 8F).
Collectively, these data reveal that Mg2+ binding stabilizes the MRS2 NTD, consistent with an observed increase in α-helicity. Furthermore, the structural and stability augmentation is dependent on D216 and D220 as mutation of these residues renders the NTD insensitive to Mg2+, reinforcing the importance of these sites to coordinating Mg2+.
Mg2+ binding to the MRS2 NTD negatively regulates mitochondrial Mg2+ uptake
To link our in vitro observations with MRS2 function, we monitored Mg2+ dynamics using Mag-Green in HeLa cells overexpressing WT and D216A/D220A MRS2. HeLa cells were incubated with the membrane-permeant Mag-Green-AM to cytosolically load the cells with the Mg2+ sensitive dye. After washing and bathing the cells with intracellular buffer (IB), the plasma membrane (PM) was permeabilized with 5 μM digitonin, and 3 mM MgCl2 was added to the bath. Mitochondrial Mg2+ uptake rates were inferred from the clearance of extramitochondrial Mg2+, measured as the decrease in Mag-Green fluorescence, as previously done (Daw et al, 2020). After MgCl2 addback, digitonin-permeabilized HeLa cells transfected with empty pCMV vector (control), pBSD-MRS2 (WT), and pBSD-MRS2 D216A/D220A (mutant), all showed increases in Mag-Green fluorescence followed by a decay associated with Mg2+ clearance (Fig 9A). Fitting the data to single exponential decays indicated greater extramitochondrial Mg2+ clearance rates for WT MRS2-expressing cells compared with control cells and mutant MRS2-expressing cells compared with control and WT MRS2-expressing cells (Fig 9B and C). Addition of 10 mM or 30 mM NaCl to similarly permeabilized cells caused no change in the Mag-Green signal, suggesting minimal influence of osmolarity on our Mag-Green measurements (Fig S5A and B).
Collectively, these data suggest that MRS2 overexpression enhances extramitochondrial Mg2+ clearance, and Mg2+ interactions with the MRS2 NTD act as a negative feedback switch to temper Mg2+ uptake into the mitochondria.
Gain of function D216K/D220K mutant relieves negative feedback on MRS2 activity
To probe whether mutation of the Mg2+-binding site causes a bona fide gain of function, we reconstituted human WT and D216K/D220K MRS2 in WT and Mrs2 knockout (Mrs2−/−) hepatocytes. Primary murine hepatocytes were transfected with empty vector, human MRS2-mRFP, or MRS2 D216K/D220K-mRFP plasmids. 24 h post-transfection, a genetically encoded, mitochondrially targeted Mag-FRET biosensor (i.e., mito-Mag-FRET) was transduced into the cells to directly measure mitochondrial Mg2+ uptake. This mito-Mag-FRET sensor was previously shown to localize to hepatocyte mitochondria, reporting reciprocal lactate-induced Mg2+ responses compared with an ER-targeted/retained Mag-FRET sensor (Daw et al, 2020). Murine Mrs2-mRFP under the control of a CMV promoter, similar to the construct used in the present study, was also shown to properly co-localize with dihydrorhodamine-123 in hepatocyte mitochondria (Daw et al, 2020). Here, confocal images of the transfected/transduced WT hepatocytes show strong co-expression and co-localization of MRS2 and MRS2 D216K/D220K with the cerulean and citrine fluorescence of the mito-Mag-FRET biosensor, indicating mitochondrial localization of the human WT and mutant MRS2 in murine cells (Figs 10A and S6A). The pixel intensity profiles of the mito-Mag-FRET citrine and human MRS2-mRFP (WT and mutant) signals exhibit coincident peak maxima, consistent with this co-localization (Fig S7A and B). Human WT and mutant MRS2 also strongly co-localized with the mito-Mag-FRET fluorophores in the Mrs2−/− hepatocytes (Figs 10B and S6B).
As expected, a 10-mM MgCl2 bolus increased the mito-Mag-FRET signal in WT cells (Fig 10C). Although WT hepatocytes expressing human MRS2 showed a similar mito-Mag-FRET response to the MgCl2 bolus, cells transfected with human MRS2 D216K/D220K exhibited highly potentiated mitochondrial Mg2+ uptake compared with controls (Fig 10D). Human MRS2 was fully capable of functionally reconstituting the Mg2+ channel in Mrs2−/− hepatocyte mitochondria. Remarkably, the MRS2 D216K/D220K formed channels that greatly enhanced mitochondrial Mg2+ uptake compared with WT human MRS2 (Fig 10E). Note that the mRFP fluorescence intensities of human WT MRS2 and human MRS2 D216K/D220K in WT and Mrs2−/− hepatocytes were similar, suggesting comparable expression levels across all groups (Fig S6C). Given the striking potentiation of Mg2+ uptake in hepatocytes co-expressing the D216K/D220K mutant but not WT human MRS2 with endogenous Mrs2, our data suggest that the Mg2+ binding–deficient MRS2 mutant dominantly mediates a gain of mitochondrial Mg2+ uptake function.
Discussion
Human MRS2 belongs to the heterogeneous CorA/Mrs2/Alr1 superfamily of Mg2+ transporters, where CorA, Alr1, and Mrs2/MRS2 comprise the principal Mg2+ uptake systems in bacteria, yeast PM, and mitochondria, respectively. Bacterial CorA has been the most extensively studied family member, yielding mechanistic and functional insights on these channels (Franken et al, 2022; Jin et al, 2022). Nevertheless, given the low sequence similarity between human MRS2 and these homologues, there remains a major knowledge gap concerning the precise structural, functional, and regulatory mechanisms of human MRS2. Here, we isolated and biophysically characterized the largest domain of human MRS2, corresponding to the matrix-oriented NTD. We found that MRS2 NTD forms a homodimer under dilute conditions, which may be a building block to higher order oligomers. Remarkably, Mg2+ and Ca2+ disassembled both higher order MRS2 NTD oligomers and homodimers but not full-length MRS2 assemblies. In contrast, Co2+ disassembled full-length MRS2 oligomers but not MRS2 NTD. We estimated the Kd of Mg2+ binding to be ∼0.14 mM, and a D216A/D220A MRS2 NTD double mutant disrupted this Mg2+ binding but had no effect on Ca2+ binding, indicating disparate binding sites for these two divalent cations. Remarkably, this D216A/D220A double mutant abrogated the enhanced solvent exposed hydrophobicity, α-helicity, and thermal stability mediated by Mg2+ binding. Furthermore, MRS2 NTD oligomers and homodimers harboring this double mutation remained intact in the presence of Mg2+. Finally, we showed that reconstitution of D216A/D220A or D216K/D220K MRS2 mutants in mammalian cells greatly increased mitochondrial Mg2+ uptake compared with WT MRS2-expressing cells.
Several CorA crystal and cryoelectron microscopy structures have been elucidated in the presence of divalent cations, revealing a pentameric assembly (Eshaghi et al, 2006; Payandeh & Pai, 2006; Guskov et al, 2012; Pfoh et al, 2012; Nordin et al, 2013; Cleverley et al, 2015; Matthies et al, 2016; Johansen et al, 2022). The first TM, which lines the channel pore, and second TM orient the intervening GMN motif for ion binding and selectivity at the pore entrance (Pfoh et al, 2012). Upstream of TM1, a large intracellular domain of CorA, analogous to the matrix-oriented human MRS2 NTD, fans out into the cytoplasm and is composed of eight α-helices and a six-stranded β-sheet (T. maritima; 4EED.pdb) (Fig 11A). For T. maritima CorA, two Mg2+-binding sites (M1 and M2) have been identified per intracellular domain (Pfoh et al, 2012). M1 is made up of D89 and D253, whereas M2 is comprised of D175 and D179. Whereas earlier studies indicated that a symmetrizing of the pentameric intracellular domain assembly upon Mg2+ binding to the NTD closes the channel (Pfoh et al, 2012; Matthies et al, 2016), more recent work indicates both symmetric and asymmetric assemblies are formed in the presence and absence of Mg2+, and channel conductance is dependent on lowered symmetric state population and coupled with a reduced energy barrier to an ensemble of open states in low Mg2+ (Kowatz & Maguire, 2019; Johansen et al, 2022). Nevertheless, it is evident that Mg2+ binding increases the rigidity/decreases the dynamics of the CorA intracellular domain (Chakrabarti et al, 2010; Pfoh et al, 2012; Rangl et al, 2019; Johansen et al, 2022).
Source Data for Figure 11[LSA-2022-01742_SdataF11.pdb]
The M1 Mg2+-binding site of T. maritima CorA does not appear conserved in human MRS2 based on multiple sequence alignments (Figs S1 and S2) or 3D superposition of the CorA crystal (Pfoh et al, 2012) and human AlphaFold2 (Jumper et al, 2021) MRS2-predicted structures. Furthermore, the AlphaFold2 model of human MRS2 orients the D216 and D220 Mg2+-binding residues, which we experimentally validated, six helical turns closer to the membrane domains compared with the CorA M2 DALVD residues (Fig 11A and B). Interestingly, a superposition of T. maritima CorA and S. cerevisiae Mrs2 (Khan et al, 2013) crystal structures structurally aligns the bacterial CorA DALVD with an INVMS sequence in S. cerevisiae Mrs2 (Fig S8A), suggesting the bacterial M2 Mg2+-binding site is not conserved in yeast. However, a superposition of the human AlphaFold2 model with the S. cerevisiae crystal structure suggests a structural conservation between human D216 and D220 and yeast D203 and E207 (of DLENE) (Fig S8B), not apparent from the multiple sequence alignments (Figs S1 and S2).
Although the stoichiometry of the human MRS2 channel remains unknown, we generated a homopentamer in homology to CorA, using AlphaFold-Multimer (Evans et al, 2022 Preprint) to model how Mg2+ binding to D216 and D220 may cause matrix domain disassembly (Fig 11B). PDBsum analysis (Laskowski et al, 2018) of the multimer model indicates that D216 and D220 do not participate in interprotomer H-bonding, salt-bridges, or other nonbonded contacts. However, P221 H-bonds and forms other nonbonded contacts with R228 of an adjacent subunit (Fig 11C). Furthermore, the loop following D220 exits into a 32-residue helix that contains many additional interprotomer contacts (Fig 11C). We showed that Mg2+ binding to D216 and D220 increases α-helicity, which could rearrange the adjacent loop that contains P221 and the position of the immediate downstream helix, leading to subunit dissociation. The human MRS2 homopentamer model also reveals clusters of negatively charged residues across interfaces (i.e., E243, D247, D305, and E312), which could mediate additional divalent cation–binding sites (Fig 11C). Ultimately, experimentally determined high-resolution structures of human MRS2 are needed to reveal channel stoichiometry, the basis for assembly and mechanisms for Mg2+-induced disassembly.
Mg2+ increased α-helicity and stability of the human MRS2 NTD, consistent with past NMR data, showing decreased backbone dynamics of CorA in the presence of high Mg2+ (Johansen et al, 2022). The far-UV CD spectra reported here resemble previous data from our laboratory, where we found no effect by MgCl2, likely due to variability in protein concentration measurements (Daw et al, 2020). Here, we applied MgCl2 addback to the same sample to expose the secondary structure change. Several lines of evidence suggest distinct Ca2+- and Mg2+-binding sites on the MRS2 NTD. First, the change in intrinsic fluorescence caused by the two cations was different; second, whereas Mg2+ increased, Ca2+ decreased solvent accessible hydrophobicity; third, D216A/D220A double mutant increased the Mg2+ Kd ∼sevenfold, whereas having no effect on the Ca2+ Kd; finally, the Mg2+-dependent solvent accessible hydrophobicity change was abrogated, whereas the Ca2+ response was maintained by the D216A/D220A double mutant. Given the MM has a free Mg2+ concentration of ∼0.5–1.5 mM (Jung et al, 1990; Rutter et al, 1990), the Mg2+ Kd of ∼0.14 mM reported here would suggest the MRS2 structure, stability, and oligomerization would be sensitive to physiologically relevant fluctuations in Mg2+ levels within the matrix.
A broad range of free MM Ca2+ concentrations in mammalian cells have been reported, dependent on cell type, stimulus, and indicator; moreover, most estimates are < 100 µM (reviewed in Fernandez-Sanz et al [2019]), much lower than our MRS2 NTD Ca2+ Kd estimate of ∼1 mM. Hence, the MRS2 NTD would have to be positioned close to a Ca2+ channel pore to be affected, where local Ca2+ concentrations may approach the ∼mM range (Chad & Eckert, 1984; Bauer, 2001; Tadross et al, 2013). Directly assessing how Ca2+ binding affects MRS2 activity in cellulo is problematic due to the weak Ca2+ Kd of 1 mM. For example, 100 μM matrix Ca2+ would occupy < 10% of the MRS2 Ca2+-binding sites and perturb mitochondrial membrane potential. Nevertheless, indirectly, mitochondrial Ca2+ uniporter KO studies show unaltered lactate-stimulated mitochondrial Mg2+ uptake (Daw et al, 2020), suggesting Ca2+ may not play a crucial role in MRS2 regulation.
Interestingly, although Co2+ dissociated larger full-length MRS2 assemblies, we observed no effect on MRS2 NTD by DLS. In contrast, Mg2+ and Ca2+ did not alter the assembly of full-length MRS2 but dissociated the MRS2 NTD. We posit that MRS258–333 (NTD) within MRS2 full-length undergoes disassembly in the presence of Mg2+ and Ca2+, whereas the C-terminal domain and/or TM regions remain interacting. Such a change would be undetectable by DLS, as the complex size would be unaffected. Furthermore, we believe Co2+-mediated disassembly occurs via binding to a region outside the NTD. Because estimates for the mitochondrial concentration of Co2+ range from ∼50 to 90 nM (Tapiero et al, 2003; Czarnek et al, 2015), the precise physiological significance of Co2+ interactions with any human MRS2 domain remains unclear.
Using permeabilized and intact cells, our data show that Mg2+ binding to the MRS2 NTD negatively regulates the channel. Permeabilized cells overexpressing the D216A/D220A double-mutant MRS2 cleared extramitochondrial Mg2+ at increased rates compared with WT MRS2-expressing cells. Furthermore, human WT and D216K/D220K MRS2 were fully capable of reconstituting functional MRS2 channels in intact primary murine Mrs2−/− hepatocytes, with the double mutant causing highly potentiated Mg2+ uptake in Mrs2−/− and WT mitochondria, indicative of gain of function activity. We do not believe that osmolarity changes because of MgCl2 addition influenced these trends since construct-specific responses were observed and 10 mM or 30 mM NaCl had no effect on the Mag-Green responses (Fig S5). Interestingly, a study using CorA harboring mutations aimed at disrupting Mg2+ binding to M1 showed WT-like 63Ni2+ transport (Kowatz & Maguire, 2019). Here, we focused on an M2-like cluster of residues because M1 does not appear to be conserved in human MRS2, discovering a robust, dominantly increased mitochondrial Mg2+ uptake upon disruption of Mg2+ binding to the NTD.
In conclusion, our work reveals the large NTD functions as a negative feedback regulator of human MRS2 channel function. We propose Mg2+ binding to the MRS2 NTD, contributed by D216 and D220, disrupts NTD:NTD interactions without disassembly of the channel (Fig 11D). Mg2+ binding to the MRS2 NTD increases α-helicity, stability, and solvent exposed hydrophobicity but dissociates NTD:NTD complexes, which we believe underlie key structural changes that propagate to the pore and/or crucial gating residues to inhibit the channel (Fig 11D). These data distinguish human MRS2 from bacterial CorA observations, where Mg2+ binding to the analogous intracellular domain shields electrostatically repulsive interfaces, promoting and bridging a symmetric interaction between intracellular domains (Pfoh et al, 2012; Matthies et al, 2016).
Because D216 and D220 are predicted to be near H232, Q236, and K239 of an adjacent protomer (Fig 11C), the D216A/D220A mutation could potentially perturb inter–protomer interactions involving these residues. In this scenario, destabilization of inter-domain interactions could lead to increased human MRS2 channel open probability, similar to the Mg2+ dissociation-dependent mechanism recently articulated in detail for T. maritima CorA by the A. Guskov group (Nemchinova et al, 2021). However, this scenario appears to be inconsistent with the Mg2+-binding–induced monomerization we observed for the human MRS2 matrix domain and our observations that D216A/D220A does not alter the dimer stoichiometry of the human MRS2 matrix domain or the higher order full-length human MRS2 assembly in CHAPS micelles.
Materials and Methods
MRS2 expression and purification
The human MRS2 NTD was identified as residues 58–333 using bioinformatic identification of the mitochondrial targeting sequence (Fukasawa et al, 2015; Almagro Armenteros et al, 2019; Buchan & Jones, 2019), and TM1 and TM2 (Krogh et al, 2001), after the comparison with these predictions with the annotations in UniProt (Accession Q9HD23). Human MRS258–333 was subcloned out of the BDS vector into pET-28a (Novagen) using PCR and NdeI and XhoI restriction sites. Overnight protein expression at 37°C from the pET-28a-MRS258–333 vector was done using BL21 (DE3) Escherichia coli cells cultured in Luria broth, induced with 0.4 mM IPTG. Protein was purified under native conditions using HisPur (Thermo-Fisher Scientific) nickel–nitrilotriacetic acid beads as per the manufacturer guidelines. The wash and elution buffers contained 20 mM Tris (pH 8.0), 150 mM NaCl, 1 mM DTT, 20 mM Tris (pH 8.0), 150 mM NaCl, 1 mM DTT, and 300 mM imidazole. After dialysis in 20 mM Tris (pH 8.0), 150 mM NaCl, and 1 mM DTT buffer using a 3,500 D MWCO membrane (Thermo Fisher Scientific), the N-terminal hexa-histidine tag was cleaved with ∼2 U of bovine thrombin (Sigma-Aldrich) per 1 mg of protein. A final SEC step through an S200 10/300 Gl column (Cytiva), achieved >95% protein purity as assessed by SDS–PAGE and Coomassie blue staining.
D216A and D220A mutant were introduced into MRS258–333 by PCR-mediated site-directed mutagenesis and expression and purification for this construct were performed as described for WT-MRS258–333. The complementary mutagenic primers were 5′-CCTTGAGACCTTGGCTGCTTTGGTGGCCCCCAAACATTCTTC-3′ and 3′-GAAGAATGTTTGGGGGCCACCAAAGCAGCCAAGGTCTCAAGG-5′.
Full-length human MRS2 taken as residues 58–443 (MRS258–443) was cloned and expressed using the same approach described for MRS258–333. Purification was performed as described for the NTD, except with the addition of 10 mM CHAPS to both the elution and SEC buffers.
SEC with in-line multi angle light scattering
SEC-MALS was performed using a Superdex 200 Increase 10/300 Gl column (Cytiva) connected to an AKTA pure FPLC system (Cytiva). A DAWN HELEOS II detector (Wyatt) and an Optilab TrEX differential refractometer (Wyatt) were used to estimate the molecular weight of MRS258–333 under various experimental conditions. The entire in-line FPLC/MALS system was housed in cold cabinet maintained at ∼10°C. Data were obtained for four different protein concentrations: 0.45 mg/ml, 0.90 mg/ml, 2.5 mg/ml, and 5 mg/ml in 20 mM Tris (pH 8), 150 mM NaCl, and 1 mM DTT, using 100 µl injections of sample at each concentration. MALS molecular weights were determined in the accompanying ASTRA software (version 7.1.4; Wyatt) using Zimm plot analysis and a protein refractive index increment (dn/dc) = 0.185 L/g. Divalent cation containing experiments were performed by supplementing the running buffers and protein samples with 5 or 10 mM MgCl2 and CaCl2, as indicated.
DLS
DLS measurements were made with a DynaPro NanoStar (Wyatt) instrument using a scattering angle of 90°. After centrifugation at 15,000g for 5 min at 4°C, 5 µl of supernatant was loaded into a JC-501 microcuvette, and measurements were collected as 10 consecutive acquisition scans with each acquisition being an average of 5 s. MRS258–333 protein samples were assessed at 1.25 mg/ml in 20 mM Tris (pH 8), 150 mM NaCl, and 1 mM DTT in the absence or presence of 5 mM MgCl2, CaCl2, or CoCl2. Similarly, MRS258-443 protein samples were assessed at 0.5 mg/ml in 20 mM Tris (pH 8), 150 mM NaCl, 10 mM CHAPS, and 1 mM DTT, in the absence or presence of 5 mM MgCl2, CaCl2, or CoCl2. For both proteins, data were acquired at 20 and 37°C, as indicated. All autocorrelation functions were deconvoluted using the regularization algorithm to extract the polydisperse distribution of hydrodynamic radii (Rh) and cumulants fit for monodisperse weight-averaged Rh using the accompanying Dynamics software (version 7.8.1.3; Wyatt).
Intrinsic fluorescence measurements for cation binding
A Cary Eclipse spectrofluorimeter (Agilent/Varian) was used to acquire intrinsic fluorescence emission spectra. Spectra were acquired for 0.1 mg/ml MRS258–333 in 20 mM Tris (pH 8), 150 mM NaCl, and 1 mM DTT, using a 600-µl quartz cuvette. The fluorescence emission intensities were recorded at 22.5°C from 300 to 450 nm, using a 1 nm data pitch and an excitation wavelength of 280 nm. Excitation and emission slit widths were set to 5 and 10 nm, respectively, and the photomultiplier tube detector was set to 650 V. Emission spectra were obtained before and after supplementation with increasing concentrations of CaCl2, MgCl2, or CoCl2, added directly to the cuvette. A total of 15 emission spectra were acquired with increasing concentrations of divalent cation between 0 and 5 mM. Spectral intensities at 330 nm were corrected for the dilution associated with the volume change upon each addition to the cuvette, and resultant curves were fit to a one site binding model that takes into account protein concentration using R (version 4.2.1) to extract apparent equilibrium dissociation constants (Kd).
Extrinsic 8-anilinonapthalene-1-sulfonic acid fluorescence
Extrinsic ANS fluorescence measurements were performed using a Cary Eclipse spectrofluorometer (Agilent/Varian). Spectra were acquired at 15°C for 30 µM MRS258–333 in 20 mM Tris (pH 8), 150 mM NaCl, 1 mM DTT, and 0.05 mM ANS, using a 600-µl quartz cuvette. The excitation wavelength was set to 372 nm, and the extrinsic ANS fluorescence emission spectra were acquired from 400 to 600 nm, with the photomultiplier tube detector set at 750 V. Excitation and emission slit widths were set to 10 and 20 nm, respectively, for all ANS experiments. To monitor divalent cation-induced changes in exposed hydrophobicity of MRS258–333, 5 mM CaCl2 or 5 mM MgCl2 was added directly into the cuvette. Negligible effects of these cations on free ANS fluorescence were confirmed by acquiring similar spectra in the absence of protein.
Far-UV CD spectroscopy
Far-UV CD spectra were acquired using a Jasco J-810 CD spectrometer with electronic Peltier temperature regulator (Jasco). Each spectrum was taken as an average of 3 accumulations, recorded at 37°C using a 1-mm path length quartz cuvette in 1-nm increments, 8-s averaging time, and 1 nm bandwidth. To eliminate technical variability in magnitude signals, after acquiring divalent cation-free spectra, 5 mM MgCl2 was added to the same samples, and spectra were re-acquired.
Thermal melts were recorded using a 1-mm path length quartz cuvette by monitoring the change in the CD signal at 222 nm from 20–95°C. A scan rate of 1°C min−1, 1 nm bandwidth, and 8-s averaging time was used during data acquisition. Mg2+-free and Mg2+-supplemented data were fit using a Boltzmann sigmoidal equation to estimate the midpoint of temperature denaturation (Tm) using R (version 4.2.1).
Mitochondrial Mg2+ uptake experiments using Mag-Green
HeLa cells were cultured in DMEM with high glucose (Wisent), 10% (vol/vol) FBS (Sigma-Aldrich), 100 μg/ml penicillin, and 100 U/ml streptomycin (Wisent) at 37°C in a 5% CO2, 95% (vol/vol) air mixture. Cells cultured in 35-mm dishes were transfected with PolyJet transfection reagent (SignaGen) according to the manufacturer guidelines. After ∼12 h, cells were incubated with 0.725 µM of the Mg2+ indicator Mag-Green for 30 min at 37°C. Cells were subsequently washed in divalent cation-free PBS, pH 7.4, and suspended in 2 ml of IB composed of 20 mM HEPES (pH 7), 130 mM KCl, 2 mM KH2PO4, 10 mM NaCl, 5 mM succinate, 5 mM malate, and 1 mM pyruvate. A 20% (vol/vol) cell suspension in IB was created in a quartz cuvette. Mag-Green fluorescence was monitored using a PTI QuantMaster spectrofluorimeter (Horiba) equipped with electronic temperature control using excitation and emission wavelengths of 506 and 531 nm, respectively, and excitation and emission slit widths of 2.5 and 2.5 nm, respectively. After a 30-s Mag-Green baseline fluorescence measurement, 2 mM EDTA plus 5 µM digitonin was added to permeabilize the PM. After 300 s, 3 mM MgCl2 (or 10–30 mM NaCl for osmolarity controls) was added to the cuvette and the Mag-Green signal was measured for 600 s. The first three intensity values recorded immediately after the cation addback were not included in any trace because of potential mixing and light artefact contributions. Mitochondrial Mg2+ uptake was correlated with the clearance of Mg2+, taken as the decrease in Mag-Green fluorescence after re-introduction of Mg2+ into the system, as previously done (Daw et al, 2020). The rates of Mag-Green fluorescence decrease were extracted by fitting the traces after the Mg2+ addback to a single exponential decay in R (version 4.2.1).
Mito-Mag-FRET measurements in primary murine hepatocytes
Primary murine hepatocytes isolated from WT and Mrs2−/− (Daw et al, 2020) mice grown on 25-mm collagen-coated glass coverslips were transfected with empty vector, human MRS2-mRFP (Hu-MRS2-mRFP), or Hu-MRS2 D216K/D220K-mRFP plasmids. 24 h post-transfection, hepatocytes were transduced with adenoviral mito-Mag-FRET (Daw et al, 2020) (20 MOI) for an additional 48 h. The subcellular localizations of ectopically expressing MRS2 and mito-Mag-FRET were visualized using a Leica SP8 confocal microscope. The cells were excited using the 405-nm laser line, and the emission was collected using the hybrid detector (HyD). The cerulean channel, 460–490 nm, and citrine channel, 510–550 nm, served to detect the emissions from the fluorescence resonance energy transfer (FRET). FRET emissions were acquired following donor and acceptor excitation sequences. Selected region of interests (ROIs) were drawn, and the acquired sequences were background corrected for acceptor cross excitation cross-talk, acceptor cross excitation, and FRET cross-talk (α = A/C; γ = B/C; δ = A/B). Time-lapse imaging was performed using the above-described acquisition mode, and the corresponding FRET efficiencies were analyzed. Selective ROIs focused on mitochondrial-targeted mito-Mag-FRET sensor signals, and the captured FRET sensitized emissions (relative FRETSE) were plotted.
To compare Hu-MRS2-mRFP and Hu-MRS2 D216/KD220K-mRFP protein abundance, the pixel intensities of mRFP signals were evaluated by selecting multiple ROIs for the following conditions: WT + Hu-MRS2-mRFP, WT + Hu-MRS2D216K/D220K-mRFP, Mrs2−/− + Hu-MRS2-mRFP, and Mrs2−/− + Hu-MRS2D216K/D220K-mRFP.
Structure modeling and visualization
The human MRS2 (UniProt accession Q9HD23) homopentamer model was generated using AlphaFold-Multimer (v2.2.0) (Evans et al, 2022 Preprint) on the Shared Hierarchical Academic Research Computing Network (SHARCNET:www.sharcnet.ca) of the Compute Canada/Digital Research Alliance of Canada. A total of 25 predictions were made (five seeds per model), using a maximum template date of 2022-11-01 and all available genetic databases. The highest confidence homopentamer model based on predicted LDDT was taken for visualization and analysis. All structure images were generated using PyMOL (Version 2.4; Schrödinger, LLC.).
Statistics
Unpaired t test was used when comparing two independent groups, paired t test was used when comparing outcomes of the same group before and after treatment, and one-way ANOVA after Tukey’s post hoc test was used for multiple means comparisons between three or more groups. All nonlinear regression fitting and statistical analyses were done in GraphPad Prism (4.03) or R (4.2.1).
Data Availability
The AlphaFold-Multimer generated human MRS2 homopentamer coordinates have been included as Source Data, and all other data are available from the corresponding author upon request.
Acknowledgements
This work was supported by a Canadian Institutes of Health Research Project Scheme Grant 438225 (to PB Stathopulos) and National Institutes of Health (R01GM109882, R01HL086699, R01HL142673, and R01GM135760) and DOD/DHP-CDMRP PR181598P-1 grants (to M Madesh).
Author Contributions
S Uthayabalan: conceptualization, formal analysis, investigation, visualization, methodology, and writing—original draft.
N Vishnu: conceptualization, formal analysis, investigation, visualization, methodology, and writing—original draft.
M Madesh: conceptualization, resources, data curation, supervision, funding acquisition, validation, and writing—review and editing.
PB Stathopulos: conceptualization, resources, data curation, funding acquisition, validation, visualization, project administration, and writing—review and editing.
Conflict of Interest Statement
The authors declare that they have no conflict of interest.
- Received September 27, 2022.
- Revision received January 25, 2023.
- Accepted January 26, 2023.
- © 2023 Uthayabalan et al.
This article is available under a Creative Commons License (Attribution 4.0 International, as described at https://creativecommons.org/licenses/by/4.0/).