1. Structural Biology and Molecular Biophysics
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Screening of candidate substrates and coupling ions of transporters by thermostability shift assays

  1. Homa Majd
  2. Martin S King
  3. Shane M Palmer
  4. Anthony C Smith
  5. Liam DH Elbourne
  6. Ian T Paulsen
  7. David Sharples
  8. Peter JF Henderson
  9. Edmund RS Kunji  Is a corresponding author
  1. University of Cambridge, United Kingdom
  2. Macquarie University, Australia
  3. University of Leeds, United Kingdom
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Cite this article as: eLife 2018;7:e38821 doi: 10.7554/eLife.38821

Abstract

Substrates of most transport proteins have not been identified, limiting our understanding of their role in physiology and disease. Traditional identification methods use transport assays with radioactive compounds, but they are technically challenging and many compounds are unavailable in radioactive form or are prohibitively expensive, precluding large-scale trials. Here, we present a high-throughput screening method that can identify candidate substrates from libraries of unlabeled compounds. The assay is based on the principle that transport proteins recognize substrates through specific interactions, which lead to enhanced stabilization of the transporter population in thermostability shift assays. Representatives of three different transporter (super)families were tested, which differ in structure as well as transport and ion coupling mechanisms. In each case, the substrates were identified correctly from a large set of chemically related compounds, including stereo-isoforms. In some cases, stabilization by substrate binding was enhanced further by ions, providing testable hypotheses on energy coupling mechanisms.

https://doi.org/10.7554/eLife.38821.001

Introduction

Transport proteins, also called transporters, porters, carriers, translocases or permeases, encompass a diverse and ubiquitous group of membrane proteins that facilitate the translocation of ions and molecules across all types of biological membranes, linking biochemical pathways, maintaining homeostasis and providing building blocks for growth and maintenance. They comprise 5–15% of the genomic complement in most organisms. Transport proteins are classified into four major groups: primary active transporters, secondary active transporters, facilitative transporters, and channels. Human solute transport proteins have been divided into 65 subfamilies (based on sequence alignments or experimentally determined substrate specificities), which transport a diverse range of compounds, including amino acids, sugars, nucleotides, lipids, vitamins, hormones, inorganic and organic ions, metals, xenobiotics and drugs (http://slc.bioparadigms.org/). Of the 538 identified solute transport proteins in humans, more than a quarter have no assigned substrate specificity (http://slc.bioparadigms.org/). Consequently, their role in human physiology is ill-defined and opportunities for drug intervention are missed (Hediger et al., 2013). Currently, about half of the transport proteins have been associated with human disease (Rask-Andersen et al., 2013; Hediger et al., 2013; Lin et al., 2015), but only 12 classes of drugs target them directly, despite their inherent ‘druggability’ (Hediger et al., 2013; Lin et al., 2015; César-Razquin et al., 2015; Fauman et al., 2011). The situation is not better in other eukaryotic organisms and bacterial pathogens, currently on the WHO list (Supplementary file 1) (Elbourne et al., 2017). For most transporters in the sequence databases, the identifications are preliminary because they are based only on sequence homology without direct experimental evidence for the substrates, even though single amino acid residue variations in the substrate binding site can alter the specificity profoundly. Moreover, the substrate specificities of some transporters may have been incompletely or incorrectly assigned. Finally, there could be membrane proteins with unassigned function that belong to unidentified transporter families, which are not counted at all.

One of the major challenges is to find the correct substrates from a large number of potential candidates. There are approximately ~3,000 metabolites in Escherichia coli (Sajed et al., 2016), ~16,000 in Saccharomyces cerevisiae (Ramirez-Gaona et al., 2017), ~40,000 in Homo sapiens (Wishart et al., 2013), and plants must have even more as they carry out an extensive secondary metabolism. Some metabolites, such as vitamin B6, have several interconvertible species, each of which could be transported. The classical method to identify transport proteins monitors the accumulation of radiolabeled compounds in whole cells, membrane vesicles or proteoliposomes. However, these experiments can easily fail when the expressed transporters are not active due to targeting, insertion or folding issues, when they are unstable in purification and reconstitution experiments, or when substrate and coupling ion gradients are not setup correctly. Moreover, some compounds are not available in radioactive form or are prohibitively expensive, preventing large-scale identification trials. Given all these technical difficulties, it is often necessary to limit the number of candidate substrates first by using phylogenetic analysis, by analyzing phenotypic (patho)physiological data, by complementation studies or by metabolic analysis of knock-out or mutant strains.

Therefore, there is an unmet demand for the development of new methods to limit the number of potential substrates for identification of solute carriers. Here, we present a high-throughput screening method for the identification of substrates of transporters, which does not require radioactive compounds or prior knowledge. The method uses the simple concept that transporters recognize their substrates through specific interactions, enhancing their stabilization in thermostability shift assays. We verify the approach by defining the substrate specificity of three solute carriers from different bacterial and eukaryotic protein families and show that these experiments also provide valuable clues about the ion coupling mechanism.

Results

Principle of the method

Ligands, such as substrates or inhibitors, are recognized by transport proteins through specific interactions at the exclusion of other molecules. The formation of these additional bonds leads to an increase in the total number of interactions (Robinson and Kunji, 2006; Yan, 2017; Yamashita et al., 2005). Consequently, binding leads to an overall increase in the stability of the ligand-bound species compared to the unliganded species in the population of protein molecules. We have previously shown that the mitochondrial ADP/ATP carrier (AAC) from the thermophilic fungus Thermothelomyces thermophila (UniProt G2QNH0) when purified in dodecyl-maltoside is stabilized upon binding of its specific inhibitors carboxyatractyloside and bongkrekic acid in thermostability shift assays using the thiol-reactive fluorophore N-[4-(7-diethylamino-4-methyl-3-coumarinyl)-phenyl]-maleimide (CPM) (Crichton et al., 2015; King et al., 2016). In the assay, the apparent melting temperature Tm of a population of purified transporters is determined by monitoring the increase in fluorescence by CPM reacting with thiols that have become exposed due to thermal denaturation of the proteins (Figure 1a). The apparent melting temperature Tm is the temperature at which the rate of unfolding for a given population is highest. We tested whether transported substrates can also cause a shift in thermostability, as their properties differ quite substantially from those of inhibitors or other tight binders. Indeed, the thermostability of the AAC population was enhanced in the presence of ADP and ATP, but not in the presence of AMP, which reflects the known substrate specificity of the carrier well (Mifsud et al., 2013) (Figure 1b). This effect is only observed at high concentrations of the substrate, well above the apparent Km of transport (Figure 1—figure supplement 1). The higher the substrate concentration, the higher the likelihood that part of the population is prevented from unfolding by binding of the substrate, leading to the observed shift in thermostability. We reasoned that this approach could be applied as a screening method to find substrate candidates of uncharacterized transporters by using compound libraries. To verify the method, we have tested transporters from three different (super)families, which are distinct in structure and transport mechanism.

Figure 1 with 1 supplement see all
Substrate-induced stabilization of a mitochondrial ADP/ATP carrier.

(a) As protein molecules in a population unfold due to a gradual rise in temperature (25–90°C), buried cysteine residues become solvent-exposed and accessible to the thiol-specific probe CPM (blue stick representation) that becomes fluorescent upon reaction. (b) Typical unfolding curves of the mitochondrial ADP/ATP carrier of Thermothelomyces thermophila (2 μg) in the absence and presence of 2.5 mM AMP, ADP and ATP, shown in red, blue and green, respectively. (c) The apparent melting temperature (Tm) is the peak in the derivative of the unfolding curve (dF/dT), which is used as an indicator of thermal stability. The apparent melting temperatures reported in the text are from three independent protein purifications.

https://doi.org/10.7554/eLife.38821.002

Substrates cause specific thermostability shifts in different transporters

The galactose transporter GalP of E. coli (Henderson, 1977) is a prototypical member of the sugar porter family that belongs to the major facilitator superfamily (MFS) (Pao et al., 1998). Currently, the MFS contains 24 different transporter families and 320,665 sequence entries from 5224 species, but the substrates for the vast majority of them have not been formally identified (Pfam CL0015). In humans, 14 of the 65 solute carrier families belong to the MFS, and substrates are not known for around 12% of them (http://slc.bioparadigms.org/).

The structure of GalP has not been determined, but those of its mammalian homologs GLUT1 (Deng et al., 2014), GLUT3 (Deng et al., 2015), and GLUT5 (Nomura et al., 2015) are available (Figure 2a). GalP contains three cysteine residues, of which only one is readily accessible to reaction with N-ethylmaleimide (McDonald and Henderson, 2001). To evaluate the strategy, we measured the unfolding curves of purified GalP in dodecyl-maltoside in the presence of a large number of different sugars. We determined the temperature shift by subtracting the apparent Tm of unliganded GalP (57.6 ± 0.3°C) from the apparent Tm values observed in the presence of compounds (ΔTm). The GalP population was markedly stabilized by D-glucosamine (ΔTm; 5.7 ± 0.4°C), D-glucose (ΔTm; 4.2 ± 0.5°C), D-galactose (ΔTm; 2.1 ± 0.6°C) (Figure 2—figure supplement 1), and to a lesser extent by D-talose, 2-deoxy-D-glucose, 6-deoxy-D-glucose. Importantly, the related L-isoforms showed no significant shift (Figure 2b). These results match the known substrate specificity of GalP well (Henderson and Maiden, 1990; Walmsley et al., 1994). D-Glucosamine, the most stabilizing compound, had not been investigated previously, but transport assays showed that this compound is a new substrate (Figure 2—figure supplement 2), demonstrating that the assay can be used to discover as well as to extend the substrate specificity of transport proteins.

Figure 2 with 4 supplements see all
Validation of the method for determining substrate specificity using three unrelated proteins.

(a), (c) and (e) Structural models of GalP (based on GLUT5, PDB 4YBQ), Mhp1 (PDB 2JLN) and AAC (based on Aac2p, PDB 4C9G), respectively. The models are shown in rainbow cartoon and wheat surface representations. Cysteine residues are shown as spheres, except for Cys-19 of GalP, which could not be modeled. (b), (d) and (f) Thermostability screen of GalP (3 μg), Mhp1 (3 μg) and AAC (2 μg) against sugar (50 mM final concentration), nucleobase (2 mM) and nucleotide libraries (2.5 mM), respectively. The temperature shift (ΔTm) is the apparent melting temperature in the presence of compound minus the apparent melting temperature in the absence of compound. The data are represented by the standard deviation of five, three and three independent repeats, respectively. Two-tailed Student’s t-tests assuming unequal variances were performed for the significance analysis (0.05 < p-value: not significant; 0.01 < p-value<0.05: *; 0.001 < p-value<0.01: **; 0.0001 < p-value<0.001: ***; p-value<0.0001: ****). L-BH, 5-benzyl-L-hydantoin; BVH, 5-bromovinylhydantoin; L-IMH, 5-indolylmethyl-L-hydantoin, L-NMH 5- (2-naphthylmethyl)-L-hydantoin.

https://doi.org/10.7554/eLife.38821.004

The transport protein Mhp1 from Microbacterium liquefaciens (Uniprot D6R8X8) transports 5-substituted hydantoins in a sodium-coupled mechanism (Suzuki and Henderson, 2006; Shimamura et al., 2010) (Figure 2c), and is a member of the nucleobase-cation-symport family of homologous proteins that also transport purines, pyrimidines, vitamins and related metabolites. The transporter has the LeuT-fold (Shimamura et al., 2010; Yamashita et al., 2005) and belongs to the amino acid-polyamine-organocation superfamily (Västermark and Saier, 2014), which currently contains 20 families and 147,819 sequence entries, the substrates of which have mostly not been identified (Pfam CL0062). Mhp1 contains three cysteine residues, of which only one is readily accessible to reaction with N-ethylmaleimide (Calabrese et al., 2017). Purified Mhp1 in dodecyl-maltoside had an apparent Tm of 51.3°C ± 0.6°C and to test the binding specificity, the ΔTm was determined upon addition of different nucleobases and other compounds (Figure 2d). The only stabilizing compounds were 5-indolylmethyl-L-hydantoin (ΔTm; 14.1 ± 1.2°C), 5-benzyl-L-hydantoin (ΔTm; 13.6 ± 1.2°C) and 5-bromovinylhydantoin (ΔTm; 11.9 ± 1.1°C) (Figure 2—figure supplement 3). A chemically related inhibitor of Mhp1, 5-(2-naphthylmethyl)-L-hydantoin (L-NMH) also led to a thermostability shift (ΔTm; 15.6 ± 1.7°C) (Figure 2d). These results match the known substrate and inhibitor specificity of Mhp1 (Simmons et al., 2014), showing that the assay could identify substrates from a set of closely related compounds without false positives.

To validate the method further, we also screened the mitochondrial ADP/ATP carrier from T. thermophila (UniProt G2QNH0) (King et al., 2016) (Figure 2e). This transport protein belongs to the mitochondrial carrier family (MCF), which is involved in the translocation of chemically diverse compounds across the mitochondrial inner membrane, using uniport, symport or antiport modes of transport (Kunji, 2012). Currently, there are 89,340 different sequence entries from 831 different species in the database (Pfam PF00153). In humans, the MCF is the largest solute carrier family with 53 different members (SLC25), but the substrate specificity of only half of them has been defined (Kunji, 2012). We screened the thermostability of purified AAC using a library of mitochondrial compounds (Figure 2—figure supplement 4). In the presence of ATP, ADP, and dADP the population was stabilized, showing ΔTm values of 7.2 ± 0.2, 6.0 ± 0.5 and 1.8 ± 0.2°C, respectively (Figures 1b and 2f). Some other compounds, mostly structurally related nucleotides, also stabilized the protein, but with significantly smaller shifts (Figure 2f). For each type of nucleotide, the di- and tri-phosphate species showed larger shifts than the monophosphate forms, similar to the preference of ATP and ADP over AMP (Figures 1d and 2f), showing that the assay can also identify functional groups that are important contributors to substrate binding.

Substrate screening of a mitochondrial carrier from Tetrahymena thermophila

To see whether this method can be used to identify candidate substrates for a previously uncharacterized transporter, we performed a high-throughput screen on a purified mitochondrial carrier from the thermophilic ciliate Tetrahymena thermophila (UniProt I7M3J0). The carrier is phylogenetically related to the yeast mitochondrial carrier that transports inorganic phosphate in symport with a proton into the mitochondrial matrix for ATP synthesis (Runswick et al., 1987), but its substrates have not been identified experimentally. The population of carriers had an apparent Tm of 56.0 ± 0.8°C and was screened against a library of 132 different mitochondrial compounds at pH 6.0 (Figure 3). The highest shift in the Tm of the population was observed for phosphate (ΔTm; 4.6 ± 0.4°C), followed by glyoxylate (ΔTm; 3.6 ± 0.8°C), acetyl-phosphate (ΔTm; 3.2 ± 0.5°C) and phosphoenolpyruvate (ΔTm; 2.3 ± 0.2°C) (Figure 3—figure supplement 1), whereas small shifts were observed for nucleotides. These compounds either contain phosphate as a functional group or resemble the structure of phosphate, such as glyoxylate (Figure 3—figure supplement 2). Thus, this assay can be used to narrow down substantially the number of potential substrate candidates from a library of unlabeled compounds. Uptake assays using protein reconstituted into proteoliposomes confirmed that the mitochondrial carrier exchanges phosphate (Figure 3—figure supplement 3a). We also tested whether glyoxylate, phosphoenolpyruvate and acetyl phosphate, which also gave a shift in thermostability assays, were transported, but none of them did (Figure 3—figure supplement 3b). However, in competition assays glyoxylate abolished uptake, demonstrating that it competes effectively with [33P]-orthophosphate for binding when present in a 1000-fold excess (Figure 3—figure supplement 3c).

Figure 3 with 3 supplements see all
Identification of substrates for a mitochondrial phosphate carrier from the thermophilic ciliate Tetrahymena thermophila.

Purified carrier (1 μg) in lauryl maltose neopentyl glycol was incubated at pH 6.0 with 2.5 mM of each library compound separately and the ΔTm was determined, which is the difference between the apparent melting temperatures in the presence and absence of the tested compound. The data are represented by the average and standard deviation of three independent assays. The significance tests were performed as described in the legend to Figure 2.

https://doi.org/10.7554/eLife.38821.009

The effect of coupling ions

Secondary active transporters are widespread and often use the electrochemical gradient of protons or other ions to drive the uptake of substrates against their concentration gradient. Coupling ions are also recognized by transporters through specific interactions, often directly associated with the binding of substrates (Kunji and Robinson, 2010; Krishnamurthy et al., 2009).

5-Benzyl-L-hydantoin is transported by Mhp1 in symport with sodium ions (Suzuki and Henderson, 2006; Shimamura et al., 2010) and thus, we tested whether the temperature shifts of Mhp1 induced by substrate binding were ion-dependent. When different ions were tested, only sodium ions induced a large substrate stabilization, whereas relatively small stabilizations were observed in the presence of potassium, calcium and magnesium ions (Figure 4a).

The effect of coupling ions on the stabilization of transporters by substrate binding.

(a) Thermostability shifts of Mhp1 (3 μg) induced by binding of hydantoins in the presence of 140 mM NaCl, KCl, CaCl2 and MgCl2. L-BH: 5-benzyl-L-hydantoin, BVH: 5-bromovinylhydantoin, L-IMH: 5-indolmethyl-L-hydantoin. (b) Thermostability shifts of the phosphate carrier (1 μg) induced by binding of 2.5 mM phosphate-containing compounds at pH 6.0 and 8.0. Acetyl-Pi: acetyl-phosphate, Pi: phosphate. (c) Thermostability shifts of the (2 μg) induced by 2.5 mM ADP and ATP binding at pH 6.0 and 8.0, using AMP as control.

https://doi.org/10.7554/eLife.38821.013

Mitochondrial phosphate carriers are proton-driven, coupling phosphate translocation to proton symport (Kunji and Robinson, 2010). To investigate whether proton-dependency of substrate binding could be inferred from the assay, the shift in thermal stability by substrate binding was determined at pH 6.0 and 8.0. The pH can affect the reaction of maleimide groups of CPM with free thiols and the stability of the protein itself, but the ΔTm value shows the effect of pH on substrate binding only. In both cases, a substrate-induced stabilization was observed, but the effect was larger in pH 6.0 than in pH 8.0 (ΔTm 4.6 ± 0.4°C and 2.6 ± 0.4°C, respectively) (Figure 4b).

As a control, we carried out the same experiment with the mitochondrial ADP/ATP carrier, which is not a proton symporter (Figure 4c). The opposite effect was observed; substrate binding at pH 8.0 led to a bigger thermostability shift than at pH 6.0. The predicted substrate-binding site of AAC has three positively charged residues that interact with the negatively charged phosphate groups of ADP and ATP (Kunji and Robinson, 2006; Robinson and Kunji, 2006). ATP has pKa values of 0.9, 1.5, 2.3 and 7.7, whereas ADP has pKa values 0.9, 2.8 and 6.8. This means that the substrates are more negatively charged at pH 8.0 compared to pH 6.0, which will enhance their binding to the positively charged residues of the substrate binding site, leading to an increase in thermostability. In the case of the phosphate carrier, the situation is very different. Phosphate has pKa values of 2.2, 7.2 and 12.3, and the predicted binding site contains two positively charged residues to neutralize the two negative charges on phosphate. However, in addition, the binding site contains a negatively charged glutamate, which has a pKa of ~4.0. The coupling proton mediates bonding of the substrate and the carboxyl group of the glutamate residue (Kunji and Robinson, 2010), which occurs more readily at pH 6.0 than pH 8.0, explaining the difference in thermostability. In support of this notion the transport rates of the phosphate carrier are high at pH 6.0 and virtually absent at pH 8.0 (Stappen and Krämer, 1994). These results show that substrate binding stabilizes the transporters, but that this effect is enhanced further by the presence of coupling ions, providing testable hypotheses on energy coupling mechanisms.

Discussion

We have shown that thermostability assays can be used to study the interaction of substrates with transporters, which is not intuitive, as substrates only bind transiently. Still, for diverse types of transport proteins, which differ in structure and transport mechanism, a shift in thermostability was observed in the presence of specific substrates. The shift is observed at concentrations well above the apparent Km of transport, as under these conditions a larger part of the protein population is rescued from unfolding by binding of the substrate, increasing the overall number of interactions.

These studies highlight the different contributions that these assays can make to studying the properties of transport proteins. First, they can be used to limit the number of potential substrate candidates from libraries of unlabeled compounds, providing important clues about the substrate specificity (Figures 2 and 3). In the tested cases, the assay correctly identified the substrates from a set of chemically related compounds, including stereo-isoforms. Still, these candidates need to be tested by transport or other types of assays, as the compounds could potentially be inhibitors, competitors or regulators. However, no prior knowledge is required and the number of radioactive compounds that need testing is highly reduced, meaning that it is easier and cheaper to establish a robust transport assay. Second, the assays can be used to extend the substrate specificity of known transport proteins, as was shown for GalP with D-glucosamine (Figure 2b). Third, they can be used to identify functional groups that are key to substrate binding, as shown for AAC, where nucleotide di- and tri-phosphates bound more tightly than nucleotide monophosphates (Figure 2f). In the case of mitochondrial phosphate carriers, the common properties of compounds, such as the phosphate group, may lead directly to the most probable candidates. Fourth, this assay can provide important clues on the involvement of coupling ions in substrate binding, as shown for cases where transport is driven by sodium ion or proton symport (Figure 4).

This method has major advantages over the traditional methods of identification. It is highly reproducible and can be performed in a high-throughput manner, allowing screening of about 100 compounds per machine per hour. The assay requires a relatively small amount of protein per assay (micrograms), depending on the number of buried cysteine residues. Another advantage is that the unfolding curves are themselves important quality checks, as they indicate that the protein is folded and therefore competent to bind substrate. Compounds which are poorly soluble in water can be added dissolved in dimethyl sulfoxide, provided that the final concentration of the solvent is kept low. Another advantage is that most compounds of the library can be used as internal controls, providing a reliable discrimination between positive and negative results. Furthermore, generic stabilizing and destabilizing compounds can be identified as more proteins of the same family are tested. For example, the polyamines spermine and spermidine have a destabilising effect on all purified mitochondrial carriers tested in our laboratory with the exception of AAC. These observations will reduce false positives for members of the same protein family. We have also observed that proteins incorporated into nanodiscs or amphipols can be successfully assayed using this approach, which is potentially useful for membrane proteins that are unstable in detergents.

There are also some limitations, as only proteins that have buried thiols can be used, although accessible cysteines can be tolerated as they only raise the base line of the assay without interfering with the unfolding curve. Thiol-containing compounds, such as cysteine or glutathione, cannot be screened as the probe reacts with them directly. However, changes in the fluorescence of environmentally sensitive dyes or endogenous tryptophans can be used as alternatives to CPM. Furthermore, the assay requires purified and stable transporter in detergent, although partially purified protein might be sufficient. Advances in bacterial (King et al., 2016; Ward et al., 1999), yeast (Routledge et al., 2016), insect (Contreras-Gómez et al., 2014), and mammalian (Goehring et al., 2014; Andréll and Tate, 2013) expression systems have allowed many transporters to be expressed in folded and functional states. Also, robust purification methods and novel classes of ‘stabilizing’ detergents, such as the neopentyl glycol maltoside detergents (Chae et al., 2010), have allowed even poorly expressed transporters to be purified in the small quantities required for these studies.

Even though the main purpose of this study was to identify potential substrates of transporters, the assays may also be used in drug discovery or in the identification of substrates, inhibitors and regulators of any other soluble or membrane proteins with occluded cysteines.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Chemical
compound, drug
Dodecyl-β-D-maltosideGlycon Biochemicals
GmbH
Glycon:D97002
Chemical
compound, drug
Dodecyl maltose
neopentyl glycol
AnatraceAnatrace:NG310
Chemical
compound, drug
N-[4-(7-diethylamino-4-methyl-3-
coumarinyl)phenyl]
maleimide (CPM)
SigmaSigma:C1484
Chemical
compound, drug
Complete Mini EDTA-free
protease inhibitor tablets
RocheRoche:5056489001
Chemical
compound, drug
Nickel Sepharose
(High Performance)
GE HealthcareGE Healthcare:17526802
Chemical
compound, drug
Nickel-NTA SuperflowQiagenQiagen:30430
Chemical
compound, drug
Factor Xa proteaseNEBNEB:P8010L
Chemical
compound, drug
Tetraoleoyl cardiolipin (TOCL)Avanti Polar LipidsAvanti Polar
Lipids:710335C
Chemical
compound, drug
E. coli polar lipid extractAvanti Polar LipidsAvanti Polar
Lipids:100600C
Chemical
compound, drug
[14C]-galactoseAmerican Radiolabeled
Chemicals
American Radiolabeled
Chemicals:ARC0117
Chemical
compound, drug
[14C]-glucosamineAmerican Radiolabeled
Chemicals
American Radiolabeled
Chemicals:ARC0118A
Chemical
compound, drug
[33P]-orthophosphateHartmann AnalyticHartmann Analytic:FF-01
Chemical
compound, drug
BioBeadsBioRadBioRad:152–3920
Chemical
compound, drug
C10E5SigmaSigma:76436
Chemical
compound, drug
Hydantoin compoundsOtherA gift from Marta Sans,
Maria Kokkinidou and
Arwen Pearson,
University of Hamburg
Software,
algorithm
PrismGraphPad
Strain, strain
background
(S. cerevisiae)
Saccharomyces
cerevisiae W303-1B
ATCCATTC:201238
Strain, strain
background
(S. cerevisiae)
Saccharomyces
cerevisiae WB12
PMID:9878703A gift from Dr H. Terada,
Tokyo University of Science
Strain, strain
background (E. coli)
Escherichia coli JM1100PMID:15558
Recombinant
 DNA reagent
Modified pYES3 vectorPMID:26453935

An additional key resources table is presented in Supplementary file 2, and details the compounds in the libraries.

Materials

Chemicals were obtained from Sigma Aldrich and Thermo Fisher Scientific (USA). Nickel NTA and sepharose beads were purchased from Qiagen (USA). All enzymes were provided by New England BioLabs (USA). Lipids were purchased from Avanti Polar Lipids (USA) and detergents from Anatrace (USA). The hydantoin compounds were a kind gift of Marta Sans, Maria Kokkinidou and Arwen Pearson (University of Hamburg).

Protein expression

For expression of mitochondrial ADP/ATP carrier of Thermothelomyces thermophila and the putative phosphate carrier of Tetrahymena thermophila in yeast mitochondria, gene constructs were designed to contain an N-terminal tag composed of eight histidine residues followed by a Thr-Ser-Glu-Asp linker and an Ile-Glu-Gly-Arg Factor Xa protease cleavage site. The genes were cloned into pYES3/CT vector (Invitrogen, USA) with a constitutively active promoter (pMIR promoter of the S. cerevisiae phosphate carrier). The plasmids were transformed into S. cerevisiae WB.12 (MATα ade2-1 trp1-1 ura3-1 can1-100 aac1::LEU2 aac2::HIS3) and W303-1B (MATα leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15) strains, respectively. Cells were grown in a 50-l fermenter after which mitochondria were prepared (Kunji and Harding, 2003).

For expression of GalP in E. coli the promoter region and the galP gene, which was modified to encode six histidine residues at the C-terminus of the protein, were cloned into plasmid pBR322 to form plasmid pGP1, which was transformed into the galactose/glucose transport-deficient host strain JM1100 (ptsG ptsM ptsF mgl galP Hfr Δhis gnd thyA galK) (Henderson et al., 1977). Cells were grown in basal salts medium supplemented with 30 mM glycerol, 20 µg/ml thymine, 80 µg/ml histidine and 15 µg/ml tetracycline in a fermenter (30- or 100-liter scale). The gene hyuP from M. liquefaciens, modified to encode six histidine residues at the C-terminus, was cloned into plasmid pTTQ18 (Stark, 1987; Suzuki and Henderson, 2006). The His6-tagged Mhp1 hydantoin transport protein was expressed in E. coli BL21 (DE3) grown in M9 medium supplemented with 20 mM glycerol, 20 mM NH4Cl, 100 μg/ml of carbenicillin, 0.2% w/v casamino acids, 2 mM MgSO4.7H2O, 0.4 mM CaCl2.2H2O, using induction by IPTG in 100-l fermenter cultures. In all cases after harvesting the intact cells preparations were made of inner membranes (Ward et al., 1999), which were stored at −80°C in Tris-HCl buffer pH 7.5 until used for purification of each individual protein.

Protein purification

The mitochondrial ADP/ATP carrier and phosphate carrier were purified using the detergents dodecyl-maltoside and lauryl maltose neopentyl glycol, respectively, using established procedures (King et al., 2016).

For the purification of His-tagged GalP, E. coli membranes were solubilized in buffer containing 20 mM Tris-HCl pH 8, 20 mM imidazole, 300 mM NaCl, 20% glycerol and 1% dodecyl-maltoside for 1 hr at 4°C with gentle mixing to a final protein concentration of 2.5 mg/ml. After centrifugation (108,000 x g, 1 hr, 4°C), the supernatant was mixed with pre-washed nickel-NTA resin and purified by immobilized nickel affinity chromatography (1 ml resin per 37 mg of total protein; Superflow, Qiagen) for batch-binding affinity chromatography for 1 hr at 4°C. Non-specific proteins were removed with buffer containing 20 mM Tris-HCl pH 8, 20 mM imidazole, 150 mM NaCl, 10% glycerol and 0.02% dodecyl-maltoside, after which His-tagged GalP was eluted with buffer containing 20 mM Tris-HCl pH 8, 200 mM imidazole, 100 mM NaCl, 5% glycerol and 0.02% dodecyl-maltoside. Imidazole was then removed by passing the sample through pre-equilibrated PD-10 desalting column (GE Healthcare). Protein samples were snap-frozen and stored in liquid nitrogen.

The purification of Mhp1 followed the same procedure as the GalP purification with the following modifications. Membranes were solubilized for 2 hr and incubated with Nickel NTA resin for 2 hr. The wash buffer contained 10 mM Tris-HCl pH 8, 20 mM imidazole, 10% glycerol and 0.05% dodecyl-maltoside and the elution buffer contained 10 mM Tris-HCl pH 8, 200 mM imidazole, 2.5% glycerol and 0.05% dodecyl-maltoside.

Reconstitution

Twelve milligrams of total polar lipid extract (Avanti Polar Lipids, Alabaster) was dried under a stream of nitrogen, washed with methanol, and dried as before. Lipids were re-hydrated in 20 mM MES pH 6.5, 50 mM NaCl and, when required, compound was added to a final concentration of 20 mM from a 200 mM stock. Lipids were solubilised by vortexing in the presence of 1.5% (v/v) pentaethylene glycol monodecyl ether, and 50 μg protein was added. The samples were incubated on ice for 5 min, and pentaethylene glycol monodecyl was removed to form liposomes by addition of bio-beads (Bio-Rad, Hemel Hempstead, UK). Eight additions of bio-beads were made to the sample: the first four used 60 mg, and the final four used 120 mg. Between additions, the samples were incubated with inversion at 4˚C for 20 min. Following the final bio-bead addition, the samples were incubated overnight at 4˚C with inversion. Bio-beads were removed by passage of the sample through empty micro-bio spin columns (Bio-Rad, Hemel Hempstead, UK). The external buffer was replaced with 20 mM MES pH 6.5, 50 mM NaCl using a PD10 desalting column (GE Healthcare, Little Chalfont, UK). When required, competitors were added externally to a final concentration of 20 mM.

Transport assays

Robotic transport assays were performed using a Hamilton MicroLab Star robot (Hamilton Robotics Ltd, UK). For GalP transport assays, E. coli cells (GalP-expressing strain pGP1/JM1100 and JM1100 control strains) were diluted to OD600 of 20 in MES buffer (5 mM MES pH 6.6, 150 mM KCl) and incubated at room temperature with 20 mM glycerol for 10 min to be energized prior to being loaded into the wells of a MultiScreenHTS +Hi Flow-FB (pore size = 1.0 μm, Millipore). Transport assays were initiated by addition of 100 μl of buffer containing 5 μM [14C]-galactose (American Radiolabeled Chemicals, 0.2 GBq/mmol) or 5 μM [14C]-glucosamine (American Radiolabeled Chemicals, 0.2 GBq/mmol). For the competition assay, radiolabeled substrate was mixed with a 4000-fold excess of the competitor compound prior to the assay. For phosphate transport assays, 100 μl proteoliposomes were loaded into a MultiScreenHTS-HA 96-well filter plate (pore size, 0.45 µm; Millipore, Billerica), and transport was initiated by the addition of 100 μl of buffer containing 20 μM [33P]-orthophosphate (Hartmann Analytic, 0.5 GBq/mmol). For competition assays, transport was initiated in the presence of 20 mM compound (1000-fold excess). Transport was stopped after 0, 10, 20, 30, 45, 60, 150 s and, 5, 7.5 and 10 min incubation times with 200 μl ice-cold buffer and the samples were filtered and washed as above. Levels of radioactivity were measured by adding 200 μl MicroScint-20 (Perkin Elmer) and measured with a TopCount scintillation counter (Perkin Elmer). Initial rates were determined from the linear part of the uptake curves.

Preparation of the mitochondrial compound library

To identify mitochondrial metabolites every enzyme in the KEGG metabolic database (Kanehisa et al., 2017) was evaluated for mitochondrial localization using the MitoMiner database of mitochondrial localization evidence (Smith and Robinson, 2016). A wide range of data were considered including large-scale experimental evidence from GFP tagging and mass-spectrometry of mitochondrial fractions, mitochondrial targeting sequence predictions, immunofluorescent staining from the Human Protein Atlas (Thul et al., 2017), and annotation from the Gene Ontology Consortium (The Gene Ontology Consortium, 2017). Ensembl Compara allowed these data to be shared across human, mouse, rat and yeast homologs (Zerbino et al., 2018). Once a list of enzymes with probable mitochondrial localization was collated, KEGG was used to produce a corresponding list of potential mitochondrial compounds. Additional candidates were taken from a computer model of mitochondrial metabolism that manually partitioned metabolites on either side of the mitochondrial inner membrane (Smith and Robinson, 2011). Compounds were dissolved in PIPES buffer (10 mM PIPES pH 7.0, 50 mM NaCl) to a final concentration of 25 mM or in dimethyl sulfoxide to a final concentration of 100 mM. pH was adjusted if necessary and the stocks were frozen at −80°C.

Analysis of the total number of identified transport protein substrates in different species

The TransportDB database contains a large number of organisms for which the transporter complement has been identified via a bioinformatic pipeline, along with substrate predictions for the transporters characterized (Elbourne et al., 2017). To acquire the figures in Supplementary file 1, a Perl script driving an SQL query of the underlying MySQL database to TransportDB was developed to quantify those transporters where no prediction could be made for the substrate for the listed species. The number of transporters with unassigned specificity represents a minimal number, as the search procedure uses sequence similarity to characterised transporters as a criterion. However, the substrate specificity needs to be determined experimentally, as a single mutation in the binding site can profoundly change substrate recognition.

Thermostability shift assay and library screening

To determine stability, purified protein (typically 1–3 μg) was mixed with 20 μg/ml of the thiol-reactive fluorophore 7-diethylamino-3-(4'-maleimidylphenyl)−4-methylcoumarin (CPM) and the fluorescent intensity was measured over a 25–90°C range of temperature using a rotatory qPCR multi sample instrument (Rotor-Gene Q, Qiagen, the Netherlands). Following an initial 18°C pre-incubation step of 90 s, the temperature was ramped 1°C every 15 s, with a 4 s wait between readings, which is equal to a ramp rate of 5.6 °C/min. The excitation and emission wavelengths were 460 nm and 510 nm, respectively. Five mg/ml stocks of CPM prepared in dimethyl sulfoxide were diluted to 0.1 mg/ml and equilibrated in assay buffer for 1 hr at room temperature in the dark before addition to the protein sample. The assay buffer was usually the buffer in which the protein was purified or in a similar buffer with a different pH (MES for pH 6.0, HEPES for pH 8.0) or a different concentration of other ions. Data analyses and apparent melting temperatures (Tm, the inflection point of a melting temperature) were determined using software supplied with the instrument.

For GalP experiments, 500 mM sugar stocks were made in MilliQ water (Merck-Millipore, USA) as 10 times stocks. For Mhp1 experiments, 100 mM compound stocks were made in dimethyl sulfoxide as 50 times stocks.

Data analyses and representation

Statistical analyses were performed using Microsoft Excel with the inbuilt function of two-tailed, two-sample unequal variance Student’s t-test. The average apparent Tm of ‘no ligand’ control samples (three technical repeats within each Rotor-Gene Q run) was subtracted from the apparent Tm measured for each compound addition in the same run. This assay was performed with three or five biological repeats using independent batches of purified protein. The null hypothesis of the t-test was that the observed ΔTm for each compound was not significantly different from zero.

References

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    Comprehensive Biophysics
    1. ERS Kunji
    (2012)
    Structural and Mechanistic Aspects of Mitochondrial Transport Proteins, Comprehensive Biophysics, Elsevier.
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    Cysteine residues in the D-galactose-H+ symport protein of Escherichia coli: effects of mutagenesis on transport, reaction with N-ethylmaleimide and antibiotic binding
    1. TP McDonald
    2. PJ Henderson
    (2001)
    The Biochemical Journal 353:709–717.
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    Major facilitator superfamily
    1. SS Pao
    2. IT Paulsen
    3. MH Saier
    (1998)
    Microbiology and Molecular Biology Reviews : MMBR 62:1–34.
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    Kinetic mechanism of phosphate/phosphate and phosphate/OH- antiports catalyzed by reconstituted phosphate carrier from beef heart mitochondria.
    1. R Stappen
    2. R Krämer
    (1994)
    The Journal of Biological Chemistry 269:11240–11246.
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    8-Anilino-1-naphthalenesulfonate is a fluorescent probe of conformational changes in the D-galactose-H+ symport protein of Escherichia coli
    1. AR Walmsley
    2. GE Martin
    3. PJ Henderson
    (1994)
    The Journal of Biological Chemistry 269:17009–17019.
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Decision letter

  1. Volker Dötsch
    Reviewing Editor; J.W. Goethe-University, Germany
  2. Richard Aldrich
    Senior Editor; The University of Texas at Austin, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Screening of candidate substrates and coupling ions of transporters by thermostability shift assays" for consideration by eLife. Your article has been reviewed by three peer reviewers including Volker Dötsch as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Richard Aldrich as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Dirk Slotboom (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. Given that we consider this work to be more of a technical than a conceptual advance, we feel the contribution would be better placed in the category of a Tools and Resources paper than a Research Article.

Summary:

The substrates of many transporters, including those belonging to the human SLC family of transporters are unknown. In vitro transport assays are the gold standard, but for technical reasons these can be hard to establish. The Kunji lab nicely demonstrates how thermal shift assays with the CPM dye can be used in a high-throughput format to screen for potential substrates. The premise is simple. Only interacting ligands that show increased (high) thermostabilization are potential substrates. They show the usefulness of the CPM assay to identify the correct substrates and ligands for three different transport proteins and also show how it is able to also pick-up ion dependent (Na+) or (H+) substrate binding. They further show the use of the assay to identify potential substrates for an unknown mitochondrial carrier.

Essential revisions:

1) The de-orphanizing of the mitochondrial carrier is a very important result of this study and should be confirmed by a classical transport assay to validate the main conclusion that this is a phosphate transporter. Do other negatively charged anions, such as sulphate, show a change in its melting temperature?

2) The discussion of the data should be expanded. For instance, spermidine and spermine destabilize the phosphate transporter, but have no effect on the ADP/ATP carrier (AAC). Such observations should be discussed. Similarly, for the stabilizing effect of zinc chloride on AAC. Unfortunately, all substrates shown in Figure 2—figure supplement 4 were also tested for the phosphate transporter (Figure 3), apart from zinc. Why? Zinc ions seem interesting.

3) Related to this question, the authors claim: "Transported substrates bind only transiently and relatively weakly, leading to conformational change that in turn causes the release…" This is not strictly true. In some cases, transported substrate bind very tightly (aspartate to GltPh for instance). In addition, in detergent solution transporters are simply binding proteins that are characterized by an apparent Kd, regardless of conformational changes. This part must be phrased more carefully.

4) What is the reliable working range of thermostabilization? In this setup it seems that Tm shifts as low as 2 degrees are significant. Is this empirically-based and/or are there recommendations based on other studies for using thermal shift assays for identifying potential ligands?

5) Some membrane proteins will be destabilised in DMSO. If the protein is too unstable in DMSO would this not make it difficult to measure a shift in thermostabilization? In other words, what is the working range of thermostability of the protein required for accurately measuring ligand binding by this method? Indeed, all of the transporters tested in this study had melting temperatures higher than 50 °C, which seem high and many membrane proteins, especially mammalian membrane proteins, will have lower melting temperatures.

6) Related, could this assay also be carried out with the membrane protein incorporated into nanodiscs or into bicelles obtained through detergent titration from nanodiscs (thus avoiding complications from MSP melting)? This would make it a useful assay for membrane proteins of non-thermophilic organisms, in particular mammalian proteins. Without this possibility this method would not change the current situation for mammalian transporters of guessing the ligand based on homology to bacterial transporters. Since the authors use human and eukaryotic transporters as justification for their method development in the Introduction, this issue should at least be discussed.

7) Data representations and statistics should be improved. For instance, in Figure 1—figure supplement 1, the authors state that data represent averages and SDs of 3 biological replicates. Yet, for the AMP data 4-5 data points, which is more than 3 were used, and there are no error bars with SD.

8) The explanation for the stabilizing effect of low pH (pH 6 compared to pH 8) on the phosphate transporter seems questionable (subsection “The effect of coupling ions). Protonation of the glutamate residue in the binding site (pKa of 4) is less likely than protonation of the substrate pKa of 7.2.

9) The stock concentrations of ligands are very high (either 25 mM in buffer or 100 mM in DMSO). What was the final concentration actually used for library screening? Along these lines, how can one eliminate false positives due to non-specific interactions with such high concentrations of ligands? In particular hydrophobic compounds will easily become incorporated into the micelles and could cause effects.

https://doi.org/10.7554/eLife.38821.020

Author response

Essential revisions:

1) The de-orphanizing of the mitochondrial carrier is a very important result of this study and should be confirmed by a classical transport assay to validate the main conclusion that this is a phosphate transporter. Do other negatively charged anions, such as sulphate, show a change in its melting temperature?

We have now demonstrated that the orphan mitochondrial carrier from Tetrahymena thermophila transports phosphate; we have included the data as (Figure 3—figure supplement 3) and we have submitted the data to the Dryad repository. We also tested the other compounds that gave a large shift in thermostability (glyoxylate, phosphoenolpyruvate and acetyl phosphate) for transport; none of these compounds are transported by the phosphate carrier. In competition assays, glyoxylate did inhibit phosphate uptake, showing that it can directly compete with phosphate transport when present in excess. We have included a brief discussion in the manuscript.

We have tested the effect of sulphate on the stability of both the mitochondrial phosphate carrier (Figure 3) and ADP/ATP carrier (Figure 2—figure supplement 4). There is a very small, but significant, stabilising effect of 0.3 °C for the phosphate carrier, but no statistically significant change in the melting temperature of the mitochondrial ADP/ATP carrier.

2) The discussion of the data should be expanded. For instance, spermidine and spermine destabilize the phosphate transporter, but have no effect on the ADP/ATP carrier (AAC). Such observations should be discussed. Similarly, for the stabilizing effect of zinc chloride on AAC. Unfortunately, all substrates shown in Figure 2—figure supplement 4 were also tested for the phosphate transporter (Figure 3), apart from zinc. Why? Zinc ions seem interesting.

Spermine and spermidine have a generic destabilising effect on all tested mitochondrial carrier family members (e.g. mitochondrial phosphate carrier, aspartate/glutamate carrier, YMC1 and ATP-Mg/Pi carriers) with the exception of the mitochondrial ADP/ATP carrier. Polyamines in general bind to surface charges of proteins and alter the solubility and stability of proteins (that is why they are additives in crystallisation screens). We have included a brief discussion in the manuscript.

We have also tested the effect of zinc on the phosphate carrier, but unfortunately the unfolding curves are uninterpretable and irreproducible (see Author response image 1), as we suspect that zinc addition leads to aggregation of the carrier. This is quite a special case, as we have not seen this effect with other metal ions.

3) Related to this question: the authors claim: "Transported substrates bind only transiently and relatively weakly, leading to conformational change that in turn causes the release…" This is not strictly true. In some cases, transported substrate bind very tightly (aspartate to GltPh for instance). In addition, in detergent solution transporters are simply binding proteins that are characterized by an apparent Kd, regardless of conformational changes. This part must be phrased more carefully.

The purpose of this statement was to emphasise that substrates of transport proteins often have orders of magnitude lower binding affinities (Kd of ADP for AAC is ~10 μM) than inhibitors (Kd of CATR for AAC is ~10 nM), but the reviewer is right that this statement might not be universal. It is true that transporters in solution do not actually transport as there are no compartments. However, we have found that they do cycle through conformational changes in detergent micelles, similar to those observed in the membrane. Still, we have removed this sentence, as it is not pertinent to the contents of the manuscript.

4) What is the reliable working range of thermostabilization? In this setup it seems that Tm shifts as low as 2 degrees are significant. Is this empirically-based and/or are there recommendations based on other studies for using thermal shift assays for identifying potential ligands?

The working range of thermostabilisation assays varies widely, as it is protein-dependent and needs to be determined empirically. One of the advantages of this method is that the library contains compounds that will not bind and therefore will not stabilise the target protein, which can then act as internal controls, allowing even small positive results to be identified. We have added comments along these lines.

5) Some membrane proteins will be destabilised in DMSO. If the protein is too unstable in DMSO would this not make it difficult to measure a shift in thermostabilization? In other words, what is the working range of thermostability of the protein required for accurately measuring ligand binding by this method? Indeed, all of the transporters tested in this study had melting temperatures higher than 50 °C, which seem high and many membrane proteins, especially mammalian membrane proteins, will have lower melting temperatures.

Indeed, some membrane proteins might be destabilised in DMSO, but this would be immediately apparent from the absence of an unfolding curve. To avoid these complications, we prepare compound stocks in high concentrations (typically 100 mM as a 50-times stock in DMSO), therefore minimising the concentration of DMSO introduced into the assay. Control reactions can be carried out in the presence of the same amount of DMSO to verify the effect on protein stability. We have added a line to that effect in the Discussion.

The apparent melting temperature must not be confused with the half-life of a protein in detergent at that temperature. Mitochondrial carriers are among the most unstable eukaryotic membrane proteins with a half-life of less than 3h in dodecyl-maltoside buffers at 4 oC (Bamber et al., 2006). They consist of three domains with few polar interactions between them and their transport mechanism involves six moving elements, making them the most dynamic transport proteins characterised to date. We have tested a range of other membrane proteins from different protein families and from different species (including mammalian and human membrane proteins heterologously expressed in S. cerevisiae) using our thermostability assay. We have found that the apparent melting temperatures of unliganded proteins typically varies from 40 – 55 °C. Temperature ramping using our program starts at 25 °C; in theory, as long as the protein is in a folded conformation at the start of the assay, unfolding curves can be generated with an apparent melting temperature as low as 30 °C. This is a broadly applicable method.

6) Related, could this assay also be carried out with the membrane protein incorporated into nanodiscs or into bicelles obtained through detergent titration from nanodiscs (thus avoiding complications from MSP melting)? This would make it a useful assay for membrane proteins of non-thermophilic organisms, in particular mammalian proteins. Without this possibility this method would not change the current situation for mammalian transporters of guessing the ligand based on homology to bacterial transporters. Since the authors use human and eukaryotic transporters as justification for their method development in the Introduction, this issue should at least be discussed.

We have successfully assayed proteins incorporated into both nanodiscs and amphipols. We have added a sentence to the text accordingly.

7) Data representations and statistics should be improved. For instance, in Figure 1—figure supplement 1, the authors state that data represent averages and SDs of 3 biological replicates. Yet, for the AMP data 4-5 data points, which is more than 3 were used, and there are no error bars with SD.

We apologise for this oversight, and have corrected the figure legend for Figure 1—figure supplement 1. We decided to plot each individual data point on this graph, rather than an average and standard deviation, which has now been changed.

8) The explanation for the stabilizing effect of low pH (pH 6 compared to pH 8) on the phosphate transporter seems questionable (subsection “The effect of coupling ions). Protonation of the glutamate residue in the binding site (pKa of 4) is less likely than protonation of the substrate pKa of 7.2.

The point is that the proton binding site, a glutamate residue, and phosphate binding sites are located close together, as we have observed in many other proton-coupled mitochondrial carriers, such as the mitochondrial aspartate/glutamate carrier and citrate carrier. The premise is that the proton is shared between the negatively charged substrate and the carboxyl group of the glutamate residue, forming proton-mediated bonds that help to increase the interaction energy of phosphate binding. In support of this notion the transport rates are highest at pH 6 and virtually absent at pH 8 (Stappen and Kramer, 1994). We have added a better description and the appropriate reference to explain this better.

9) The stock concentrations of ligands are very high (either 25 mM in buffer or 100 mM in DMSO). What was the final concentration actually used for library screening? Along these lines, how can one eliminate false positives due to non-specific interactions with such high concentrations of ligands? In particular hydrophobic compounds will easily become incorporated into the micelles and could cause effects.

We have listed the stock concentrations of substrate in the Materials and methods. For clarity, we have added the final working concentrations of each library in the figure legends.

An advantage of this method is that generic stabilising and destabilising compounds will be identified as more proteins are tested. For example, spermine and spermidine destabilise the phosphate, aspartate/glutamate, ATP-Mg/Pi and YMC1 carriers. We have added a sentence in the Discussion.

References:

Bamber, L., Harding, M., Butler, P. J. G., and Kunji, E. R. S. (2006) Yeast mitochondrial ADP/ATP carriers are monomeric in detergents. Proc. Natl. Acad. Sci. U.S.A.103, 16224-16229

Stappen, R., and Kramer, R. (1994) Kinetic mechanism of phosphate/phosphate and phosphate/OH- antiports catalyzed by reconstituted phosphate carrier from beef heart mitochondria. J. Biol. Chem. 269, 11240-11246

https://doi.org/10.7554/eLife.38821.021

Article and author information

Author details

  1. Homa Majd

    Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, United Kingdom
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Contributed equally with
    Martin S King
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2048-1839
  2. Martin S King

    Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, United Kingdom
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Contributed equally with
    Homa Majd
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6030-5154
  3. Shane M Palmer

    Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, United Kingdom
    Contribution
    Resources, Large-scale Fermentation
    Competing interests
    No competing interests declared
  4. Anthony C Smith

    Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, United Kingdom
    Contribution
    Resources, Data curation, Formal analysis
    Competing interests
    No competing interests declared
  5. Liam DH Elbourne

    Department of Molecular Sciences, Macquarie University, Sydney, Australia
    Contribution
    Data curation, Software, Formal analysis
    Competing interests
    No competing interests declared
  6. Ian T Paulsen

    Department of Molecular Sciences, Macquarie University, Sydney, Australia
    Contribution
    Data curation, Software, Formal analysis
    Competing interests
    No competing interests declared
  7. David Sharples

    1. Astbury Centre for Structural Molecular Biology, University of Leeds, Leeds, United Kingdom
    2. School of Biomedical Sciences, University of Leeds, Leeds, United Kingdom
    Contribution
    Resources, Methodology, Large-scale fermentation
    Competing interests
    No competing interests declared
  8. Peter JF Henderson

    1. Astbury Centre for Structural Molecular Biology, University of Leeds, Leeds, United Kingdom
    2. School of Biomedical Sciences, University of Leeds, Leeds, United Kingdom
    Contribution
    Formal analysis, Supervision, Writing—original draft, Project administration, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9187-0938
  9. Edmund RS Kunji

    Medical Research Council Mitochondrial Biology Unit, University of Cambridge, Cambridge, United Kingdom
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    ek@mrc-mbu.cam.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0610-4500

Funding

Medical Research Council (MC_UU_00015/1)

  • Homa Majd
  • Martin S King
  • Shane M Palmer
  • Anthony C Smith
  • Edmund RS Kunji

Cambridge Commonwealth, European and International Trust

  • Homa Majd

Leverhulme Trust (EM-2014 -045)

  • Peter JF Henderson

Biotechnology and Biological Sciences Research Council (MPSI BBS/B/14418)

  • David Sharples

Wellcome (JIF 062164/Z/00/Z)

  • David Sharples

University of Leeds

  • David Sharples

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by grant MC_UU_00015/1 of the Medical Research Council, UK. HM gratefully acknowledges the Cambridge Commonwealth, European and International Trust for support of her PhD studies. PJFH thanks the Leverhulme Trust for an Emeritus Research Fellowship (Grant number EM-2014–045). The fermenters and allied equipment for protein production in Leeds were funded by the BBSRC (MPSI BBS/B/14418), the Wellcome Trust (JIF 062164/Z/00/Z) and the University of Leeds. The hydantoin derivatives were a kind gift from Marta Sans, Maria Kokkinidou and Arwen Pearson (University of Hamburg).

Senior Editor

  1. Richard Aldrich, The University of Texas at Austin, United States

Reviewing Editor

  1. Volker Dötsch, J.W. Goethe-University, Germany

Publication history

  1. Received: July 11, 2018
  2. Accepted: October 11, 2018
  3. Accepted Manuscript published: October 15, 2018 (version 1)
  4. Version of Record published: November 1, 2018 (version 2)

Copyright

© 2018, Majd et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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