Regular paper
Molecular characterization of central cytoplasmic loop in Aspergillus nidulans AstA transporter
Sebastian Piłsyk1*, Marzena Sieńko1, Jerzy Brzywczy1, Maciej P. Golan2,
Urszula Perlińska-Lenart1 and Joanna S. Kruszewska1
1Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland; 2Opta – Tech Co. Ltd., Warsaw, Poland
AstA (alternative sulfate transporter) belongs to a large, but poorly characterized, Dal5 family of allantoate permeases of the Major Facilitator Superfamily. The astA gene has been cloned from an IAM 2006 Japanese strain of Aspergillus nidulans by complementation of a sulfate permease-deficient mutant. In this study we show that conserved lysine residues in Central Cytoplasmic Loop (CCL) of the AstA protein may participate in anion selectivity, and control kinetic properties of the AstA transporter. A three-dimensional model containing four clustered lysine residues was created, showing a novel substrate-interacting structure in Major Facilitator Superfamily transporters. The assimilation constant (Kτ) of wild type AstA protein is 85 μM, while Vmax/mg of DW of AstA is twice that of the main sulfate transporter SB per mg of dry weight (DW) of mycelium (1.53 vs. 0.85 nmol/min, respectively). Amino acid substitutions in CCL did not abolish sulfate uptake, but affected its kinetic parameters. Mutants affected in the lysine residues forming the postulated sulfate-interacting pocket in AstA were able to grow and uptake sulfate, indicating that CCL is not crucial for sulfate transportation. However, these mutants exhibited altered values of Kτ and Vmax, suggesting that CCL is involved in control of the transporter activity.
Key words: Aspergillus nidulans, sulfate permease, alternative sulfate transporter, AstA, intracellular loop, mutagenesis
Received: 28 May, 2018; revised: 03 July, 2018; accepted:
20 September, 2018; available on-line: 14 November, 2018
*e-mail: Seba@ibb.waw.pl
Abbreviations: CCL, central cytoplasmic loop; MFS, major facilitator superfamily; SMR, sulfur metabolite repression; TM, transmembrane; Kt, assimilation constant; PCR, polymerase chain reaction; OE-PCR, overlap extension polymerase chain reaction
Introduction
Sulfate is taken up and utilized for the synthesis of organic sulfur compounds by bacteria, fungi and plants. In eukaryotes, sulfate uptake is carried out by sulfate permeases from the SulP family (Kertesz 2001; Loughlin et al., 2002; Piłsyk & Paszewski 2009; Alper & Sharma 2013). These proteins are assigned to the TC 2.A.53 class, according to the Transporter Classification (TC) system. In Aspergillus nidulans the main sulfate permease is encoded by the sB gene, however, an alternative sulfate transporter (AstA) is also functional in some A. nidulans strains. The astA gene has been cloned earlier from a genomic library of the Japanese IAM 2006 strain, as a suppressor complementing sulfate permease-deficiency of the sB mutant (Piłsyk et al., 2007). The astA gene is regulated at the transcriptional level by sulfur metabolite repression (SMR), being derepressed under sulfur limitation conditions. In the reference A. nidulans strains of Glasgow origin astA is non-functional, thus, SB is the only sulfate permease in these strains. Recently, it was shown that AstA transports also sulfite and choline sulfate (Holt et al., 2017). Orthologues of astA occur frequently in evolutionarily distant fungi belonging to the Dikarya phylum with often common feature of plant or animal pathogenicity. An astA orthologue identified from pathogenic Fusarium sambucinum fungus was found to be derepressed during potato tuber infection, where the AstA protein may efficiently take up sulfate (Piłsyk et al., 2015).
The A. nidulans AstA protein (GenBank® accession number ABA28286) is a member of the poorly characterized Dal5 allantoate permease family belonging to the Major Facilitator Superfamily (MFS, TC 2.A.1). The MFS transporters are localized in the cell membrane or in organellar membranes facilitating the transport of various substrates (ions, sugars, drugs, neurotransmitters, amino acids, or peptides). Contrary to ATP-dependent SulP family of sulfate permeases (Tweedie & Segel, 1970, Woodin & Wang, 1989), three different mechanisms of transport via MFS transporters are distinguished: uniporters transporting one type of substrate and energized by its gradient, symporters translocating simultaneously two or more substrates and drawing energy from the electrochemical gradient of one of them, and antiporters transporting two or more substrates in opposite directions (Law et al., 2008; Iancu et al., 2013; Ethayathulla et al., 2014; Patron et al., 2014). The AstA transporter can uptake sulfate in proton symport-driven energy (Holt et al., 2017). Until now, only one report presented experimental evidence for ATP-independence of MFS – the gene encoding Hxt13p hexose transporter was cloned while screening the yeast genomic library for resistance to antifungal drug miltefosine (Biswas et al., 2013). The authors have shown that Hxt13p acts as an ATP-independent efflux pump for miltefosine. On the other hand, it was shown that GLUT1 glucose transporter in human erythrocytes binds ATP, which triggers significant conformational changes in GLUT1 that lead to its inactivation (Blodgett et al., 2007). In this case ATP serves as low molecular weight effector rather than energy source.
Crystal structures of bacterial MFS transporters (FucP, LacY, GlpT, EmrD) have revealed the presence of 12 transmembrane (TM) alpha helices. These helices are clustered in the N- and C-terminal domains, each comprising six helices (Fig. 1) (Abramson et al., 2003; Lemieux et al., 2004; Yin et al., 2006; Dang et al., 2010). A similar structure of 12 TM helices has also been identified in another MFS phosphate transporter, the PiPT protein of the Piriformospora indica fungus (Pedersen et al., 2013). The substrate-binding residues are found in various TM helices: TM5 in the Candida albicans Mdr1p protein (Pasrija et al., 2007), TM7 in human GLUT-1 (Kasahara et al., 2009), TM1, TM5 and TM7 in E. coli GlpT and FucP (Lemieux et al., 2004; Dang et al., 2010), and TM5, TM 7, TM10 and TM11 in E. coli XylE (Sun et al., 2012). Between the N- and C-terminal domains of all MFS proteins, a third domain is present, called the Central Cytoplasmic Loop (CCL).
Substrate uptake mechanism of MFS transporters involves a rocker-switch type of movement, followed by transient formation and breakage of a salt bridge between positively and negatively charged amino acids located opposite in the apical surface of the TM helices (Law et al., 2008). Thus, CCL transiently forms a latch-like structure in the outward-facing open conformation, while in the inward-facing open conformation, it unfolds into a partially disordered chain and swings away from the substrate translocation pathway (Wisedchaisri et al., 2014). Additionally, CCL acts as a constraint for movement of the N- and C-terminal domains and permits a relatively large interdomain movement, which has implications for the substrate translocation mechanism (Law et al., 2008). It has also been shown that CCL is critical for efficient substrate transport by the C. albicans Mdr1 multidrug transporter (Mandal et al., 2012) and likely participates in formation of a substrate translocation pore.
In this study, we demonstrate that CCL of the A. nidulans AstA protein controls sulfate transport. The role of four conserved lysine residues in the A. nidulans AstA protein was analyzed using site directed mutagenesis. Interestingly, four single amino acid substitutions (K260A, K263A, K263L and K264L), introduced in the substrate-interacting region of AstA, led to more selective uptake of sulfate compared to selenate.
Materials and Methods
Strains and media. The Aspergillus nidulans strains from our collection, carrying standard markers (Clutterbuck 1994; Martinelli 1994), together with the Escherichia coli strain used in this study, are listed in Table 1. The M111 strain, derived from Glasgow wild-type, bearing the sB43 mutation complemented with the sB gene and carrying a non-functional astA gene, was used as the A. nidulans reference strain.
Growth conditions. The following media (solid and liquid) were used: complete medium (CM) (Cove, 1966) for protoplasts or DNA isolation, minimal medium (MM) or minimal sulfur free medium (MM-S) (Lukaszkiewicz & Paszewski, 1976), the latter supplemented either with sulfate (1 mM, inorganic sulfur) or L-methionine (0.1 mM, organic sulfur). The minimal media were also supplemented according to the auxotrophic requirements of the cultured strain. Liquid cultures were grown at 37°C for 16 hours in a rotary shaker (200 rpm). The culture doubling times were calculated using the Doubling Time Computing web page (Roth 2006). Escherichia coli was grown in the standard LB medium supplemented with antibiotics as described before (Sambrook et al., 1989).
Nucleic acids manipulations, plasmid construction and mutagenesis. The plasmids used in this study are listed in Table 1. Plasmids were propagated and isolated according to the standard procedures (Sambrook et al., 1989). A. nidulans DNA was isolated using a salting out method. The frozen mycelia were disrupted by grinding in liquid nitrogen, followed by immediate suspension in warm STEN buffer (1% SDS, 100 mM Tris pH 7.5, 50 mM EDTA pH 8, 100 mM NaCl) (Sambrook et al., 1989). Polymerase chain reactions (PCR) were performed in a Techne Thermocycler. DNA was sequenced, and primers were synthesized by the DNA Sequencing and Oligonucleotide Synthesis Laboratory, Institute of Biochemistry and Biophysics, PAS. The sequences of the primers used are provided in supplementary Table S1. The kPMS11-52 plasmid carrying the wild-type astA gene (Piłsyk et al., 2007) was modified by an insertion of the Neurospora crassa pyr-4 selection cassette into the PvuII restriction site yielding the kPMS11-524 plasmid. The latter could be selected by uridine prototrophy and was used for construction of a series of plasmids bearing the mutated astA alleles. Selected lysine residues were changed to alanine or leucine using Overlap Extension PCR (OE-PCR) with Pfu polymerase (Thermo Scientific). Obtained PCR products were cleaved with the SnaBI and MunI restriction enzymes (Thermo Scientific) and ligated into the kPMS11-524 vector digested with the SnaBI and MunI restriction enzymes to replace the wild type astA. These mutations were verified by DNA sequencing and introduced into the A. nidulans sulfate permease-deficient M111 strain bearing the sB43 mutation (Table 1).
A 3.3 kb PstI fragment of the non-functional ΨastA gene was excised from the kPB6-7 plasmid (Piłsyk et al., 2007) and inserted into kPG23B yielding kPG23ΨastA plasmid. For reconstitution of the missing functional TM11-TM12 domains in ΨAstA, a 3.2 kb SpeI fragment from kPMS11-524 plasmid (last 586 nucleotides of astA 3’ ORF encoding TM11-TM12 together with N. crassa pyr-4 selection cassette) was ligated between SpeI restriction sites of kPB6-7 plasmid, yielding kPB611-12.
The Green Fluorescent Protein-labeled astA allele was constructed with OE-PCR of PCR-generated GFP with PCR-generated astA wild type allele using overlapping primers. The hybrid product was initially cloned into pGEM®-T Easy vector (Promega), then excised with MunI-Van91I, and inserted into MunI-Van91I sites of kPMS11-524, yielding the kPMS11-5243 plasmid. Similarly, the same MunI-Van91I fragment was introduced into a series of kPMS11-524 derivatives bearing the mutated astA alleles.
Transformation of A. nidulans strains. The mycelia for transformation were collected by filtration, washed with 0.6 M KCl and suspended in 0.6 M KCl buffered with 10 mM potassium phosphate pH 6.5 and containing the following lytic enzymes: 15–20 mg/ml of Glucanex® (Novozymes), 2 mg/ml of Driselase® (Sigma-Aldrich) and 1 mg/ml of snail acetone powder (Sigma-Aldrich). The protoplasts were prepared and transformed using the PEG method (Kuwano et al., 2008). To increase transformation efficiency, 5 μg of the HELp helper plasmid (Gems & Clutterbuck, 1993) was added. Transformants were selected for uracil prototrophy on MM-S medium supplemented with 2 mM sulfate and 1.2 M sorbitol.
Sulfate uptake assay. The sulfate uptake analysis was performed as described previously (Piłsyk et al., 2015) using 0.1 mM methionine – supplemented MM-S for mycelia growing under derepressing conditions. Ten ml aliquots were taken after 2, 3, 5, 10, 15 and 20 min of incubation, and the sulfate transport rates (nmol/mg dry weight) were calculated based on at least three measurements taken at different times.
For the estimation of the assimilation constant (Kτ), a cold sulfate was added to the final concentrations of 0.166, 0.2, 0.25, 0.33, 0.5 and 1 mM, and incubations were carried out for 1 h. Mycelia were treated as above and the initial rate of transport (nmol/min/mg dry weight) was calculated for each concentration of the sulfate, based on at least four measurements. These data were rendered as Hanes-Woolf plots, and linear regression was used for calculation of Vmax from the slope, and Kt from the ordinate axis intercept.
Confocal microscopy. The mycelia for microscopic observations were grown in liquid MM-S medium supplemented with 0.1 mM methionine (derepressing conditions) for 18 h at 37°C in a rotary shaker (200 rpm). The mycelia were harvested by filtration, suspended in fresh MM-S medium supplemented with 1 mM sulfate and kept at 37°C in a rotary shaker (200 rpm). The mycelial samples were collected at time of the medium shift (t 0) and 5 h after the shift, and examined under a Nikon Eclipse TE2000-E inverted microscope (Nikon, Tokyo, Japan).
Membrane protein isolation. For protein isolation, the mycelia were grown as above (Confocal microscopy), and the samples were harvested just before and 5 h after a shift to repressing conditions. Next, the collected samples were ground in TM isolation buffer (30 mM Tris-HCl pH 7.4, 50 mM EDTA pH 8, 1% sorbitol, 0.05% Tween-20, 200 mM NaCl, 20 mM dithiothreitol, 1 mM PMSF, protease inhibitors Complete™-Roche), and the homogenate was centrifuged at 4000×g for 10 min. The supernatants were centrifuged for 1 h at 50 000×g and the pelleted membrane fractions were used for Western dot-blots.
Immunodetection of AstA-GFP protein. Membrane proteins suspended in TM isolation buffer were subjected to the Immobilon P membrane (Milipore), and AstA-GFP protein was detected with rabbit anti-aGFP polyclonal antibodies (Life Technologies). Polyclonal rabbit antibodies raised against entire yeast Pma1p protein were used as an internal reference standard. Anti-rabbit IgG secondary antibodies conjugated to alkaline phosphatase (Sigma-Aldrich) were then applied as secondary antibodies. After washing, dots were visualized using BCIP/NBT (Promega).
Protein quantification. Protein concentration was estimated according to Lowry and others (Lowry et al., 1951).
Bioinformatics. Multiple alignments of protein sequences were generated using the MUSCLE software (URL: http://www.drive5.com/muscle/) (Edgar, 2004). Homology searches of the GenBank database (release 95.0) were carried out using the BLAST program (Altschul & Lipman, 1990). Transmembrane topology was predicted basing on four independent algorithms: the PredictProtein server ver. 10.20.04 (https://www.predictprotein.org/) (Rost et al., 2004), TMpred (https://embnet.vital-it.ch/software/TMPRED_form.html) (Hofmann & Stoffel 1993), PSIPRED (http://bioinf.cs.ucl.ac.uk/psipred/) (Buchan et al., 2013) and SMART (http://smart.embl-heidelberg.de/) (Letunic & Bork 2017). Modeling of 3D structure and ligand binding were performed with the I-TASSER metaserver (Zhang, 2008; Roy et al., 2010; Roy et al., 2012). The image of CCL was created with RasMol (Sayle & Milner-White, 1995).
Results
Verification of astA allele functionality
Our previous studies showed that sB mutants in A. nidulans could be complemented by the astA gene only when it was derived from the Japanese IAM2006 strain. Numerous attempts to complement the auxotrophic phenotype of sB mutants by a copy of astA derived from a reference wild-type strain of Glasgow origin, have failed (Piłsyk et al., 2007). Northern blot analysis of the astA expression in the reference strain showed no detectable signal. Moreover, comparison of the nucleotide sequences of astA ORFs in the Japanese and Glasgow strains suggested numerous mutations and deletions in the latter (Piłsyk et al., 2007), including lack of 187 bp sequence putatively encoding TM11 and TM12 (Suppl. Fig. S1 at www.actabp.pl). These mutations might abolish the function of the putative AstA protein in the reference strain. Hence, we concluded that this allele exists as a pseudogene (Ψ astA). However, RNA-seq results allowed for a distinction of transcriptional unit in AspGD, and transcriptomic data indicate that the astA locus (AN10387) in the reference strain is transcribed (Cerqueira et al., 2014; Sieńko et al., 2014).
To verify functionality of the Ψ astA gene, we generated two constructs bearing selectable N. crassa pyr-4 cassette and the Ψ astA gene or Ψ astA fused with functional sequence coding for TM11-TM12 from the Japanese astA allele. However, transformation of sB mutant with these constructs did not restore sulfate prototrophy (see: Suppl. Fig. S1 at www.actabp.pl). Thus, the lack of TM11 and TM12 helices was not the only reason for abolishing function of the astA gene in strains of Glasgow origin, but the other mutations of the astA ORF present in theses strains were sufficient to abolish protein activity. Hence, in further studies a copy of the astA gene derived from the Japanese IAM2006 strain was used as a reference.
In silico analysis of the Central Cytoplasmic Loop. A predicted topology of the A. nidulans AstA transporter revealed twelve transmembrane domains and a centrally located CCL (Fig. 1), the latter domain containing four lysine residues clustered into a tight bundle located in the middle of CCL – K260, K262, K263 and K264. These four lysine residues, together with the fifth basic amino acid (arginine or lysine), are conserved in AstA orthologs from various fungal pathogens (Fig. 2A). We previously suggested that the positively charged CCL of AstA could be responsible for interaction with a negatively charged sulfate anion during translocation (Piłsyk et al., 2007). The AstA region from D225 through A275 was modeled using the I-TASSER meta-server (Zhang, 2008; Roy et al., 2010; 2012) (Fig. 2B). Five similar models of the AstA CCL bound with sulfate were obtained, and the highest-scoring one (Fig. 2B) was used in further studies. This model exhibits a small ligand-binding region comprising amino acids 260 through 264 and resembling the calcium-binding pocket of human frequenin (PDB ID: 1G8I, Bourne et al., 2001). The predicted sulfate-interacting pocket contains the four conserved lysine residues (K260, K262, K263 and K264, see Fig. 2B), therefore, it seems likely that these lysine residues directly interact with the sulfate.
Central Cytoplasmic Loop mutants can grow. To check whether the lysine residues forming the predicted sulfate-interacting pocket (K260, K262, K263 and K264) take part in sulfate transport, they were replaced individually with alanine or leucine, and additionally, a quadruple mutant with all four lysine residues changed to alanine was constructed. Small alanine residues are preferentially chosen as a substitution in site-directed mutagenesis, however, substitutions of lysine with leucine are also tested because of comparable size of these amino acids. Altogether, nine mutated astA alleles were generated using site-directed mutagenesis (Table 1). The functionality of the mutated astA alleles was tested by transformation of the A. nidulans M111 recipient strain, which bears non-functional astA gene and the sB43 mutation, and is hence unable to grow on sulfate as the sole sulfur source (Arst 1968; Piłsyk et al., 2007).
The obtained transformants were named by combining the TR letters denoting a transformant with the name of the modified protein and the amino acid substitution. Wild type alleles of astA and sB were also introduced into the M111 recipient strain, yielding the TRAstAWT and TRSBWT transformants, respectively (Table 1).
Almost all the tested astA alleles complemented the sB43 mutation, rendering the transformants able to grow on sulfate, both on solid and in liquid minimal media (Fig. 3, Table 1). The TRAstAK262L mutant grew poorly after 48 h on solid minimal medium, however, after longer incubation time it exhibited growth rate comparable to that of the wild type TRAstAWT.
A. nidulans strains bearing wild type sB are sensitive to selenate, a toxic analogue of sulfate, under derepressing conditions (0.1 mM methionine), opposite to sulfate permease-deficient strains (e.g., sB43 mutants), which are resistant (Piłsyk et al., 2007). In our study, when compared to wild 1 type AstA, the K262A or K264A substitutions hardly affected growth in the presence of selenate, while the resistance to selenate of the K260A, K263A, K263L and K264L mutants was increased substantially, so that they could grow on MM-S medium containing 1 mM selenate (Fig. 3) even under derepressing conditions (0.1 mM methionine).
Central Cytoplasmic Loop mutants can take up sulfate
Sulfate uptake was measured in transformants of the M111 strain bearing the sB43 mutation and various mutated astA alleles (Table 1), and compared with the TRAstAWT and TRSBWT strains serving as positive controls. The SB transporter in TRSBWT was more effective in sulfate uptake than the AstA mutants or wild type AstA (Fig. 4, Table 1). The K262A, K263A and K260-264A mutations did not affect the uptake of sulfate significantly, while the K260A, K260L, K263L, K264A and K264L mutations increased the uptake by 38, 51, 71.6, 64.9 and 79.7%, respectively, compared to TRAstAWT (Fig. 4, Table 1). K262L decreased sulfate uptake by 16%. Altogether, the mutations affecting individual lysine residues in CCL did not abolish the sulfate uptake, moreover, most of the tested mutations led to even faster sulfate transport.
Mutations in Central Cytoplasmic Loop affect kinetics of sulfate uptake
Since the mutations affecting CCL did not impair sulfate uptake, it seemed possible that CCL could control its kinetics. Therefore, the effects of these mutations on the kinetic parameters of AstA were next studied. The Kτ values determined for wild type AstA and the main sulfate permease SB were 85 μM and 108 μM, respectively (Fig. 5, Table 1). Each of the four substitutions of a single lysine with alanine or leucine decreased the Kτ, with the most pronounced decrease observed in transformants bearing the K260A, K260L and K264L mutations, by 70.6, 57.6 and 56.4%, respectively (Table 1). Conversely, the quadruple substitution of the four lysine residues in CCL with alanine led to an increase of the Kt by almost 40%.
The Vmax for wild type transporters was 0.85 (nmol/min) for SB, and twice as much (1.53) for AstA, measured per mg of dry weight (DW) of mycelium (Fig. 5, Table 1). Each substitution of lysine with alanine in AstA led to a lowering of Vmax – a high decrease (by 48%) was observed in the K263A mutant. Meanwhile, K262L substitution led to the strongest decrease of transport velocity (by 57.5%), while two other mutants (TRAstAK263L and TRAstAK264L) had significantly increased velocity of transportation (by 82.4 and 142%, respectively). A single K260L mutation, or a quadruple substitution of all four lysine residues in CCL with alanine, did not affect Vmax significantly. In summary, the substitutions of the lysine residues located in CCL resulted in various changes of both Vmax and Kτ, suggesting that these residues are important for controlling activity of the AstA transporter. The highest increase of Kt (up to 161%) was observed in the quadruple TRAstAK260-264A mutant, in which all four lysine residues in CCL were replaced with alanine.
AstA protein localization and stability
To verify if the mutated AstA proteins were properly located in the cell membrane during the kinetic studies, each mutated AstA variant fused with GFP was expressed in the M111 recipient strain and examined under microscope. In all the transformants, both the GFP-tagged wild type AstA protein and its mutated variants were correctly located in the cytoplasmic membrane, both before and after the shift to repressing conditions (Suppl. Fig. S2A at www.actabp.pl). Additionally, the amount of AstA-GFP or its mutated variants was verified by Western dot-blot analysis (Suppl. Fig. S2B at www.actabp.pl), indicating that the AstA protein and its mutant versions were stable in the applied experimental conditions.
Sulfate uptake by AstA is ATP-independent, in contrast to SB (SulP) permease
In order to study AstA transport mechanism, we compared the effect of 100 mM sodium azide on the radioactive sulfate uptake by the TRSBWT or TRAstAWT strains (Fig. 6). Sulfate uptake by the SB permease was significantly decreased by azide while AstA was resistant to the azide treatment. Since azide inhibits ATP-synthase (Bowler et al., 2006), it leads to a decreased intracellular ATP level. Thus, azide-insensitive transport exhibited by the AstA transformant indicates an ATP-independent transport mechanism, contrary to the SB sulfate permease, which is ATP-dependent (Fig. 6 and earlier published results by Tweedie & Segel, 1970; Woodin & Wang, 1989).
Discussion
Alternative sulfate transporter AstA, a member of the MFS transporters, is found in some fungi, particularly frequently in pathogenic species. Numerous mutations present in the astA locus (AN10387) in the A. nidulans reference strain abolish its functionality, as it was postulated earlier (Piłsyk et al., 2007) and confirmed in this study. Despite of detectable transcription of the ΨastA locus (Cerqueira et al., 2014, Sieńko et al., 2014), the protein product is either not produced or non-functional. We showed that truncated ΨastA was unable to complement the sulfate permease-deficient mutants, even when its ORF was reconstituted with a missing 3’ sequence encoding the last two TM helices 11 and 12. This result supports our earlier conclusion that the astA allele derived from the Japanese IAM2006 strain is functional, while A. nidulans strains of Glasgow origin contain pseudogene sequences.
Structural studies of various MFS transporters have shown the presence of two transmembrane domains, each one comprising six membrane-spanning helices. The two domains are connected by the central cytoplasmic loop (CCL) (Fig. 1), predictably localized in the way of the transported substrate (Law et al., 2008).
Looking for the role of CCL in AstA, we focused on a highly conserved motif of four lysine residues present in AstA orthologs from various fungi (Fig. 2A). These positively charged residues seemed to interact with a negatively charged sulfate ion during translocation, and this assumption was confirmed by the predicted structure of CCL bound with sulfate. Therefore, the role of these four lysine residues (K260, K262, K263 and K264) was subjected to analysis using site-directed mutagenesis.
Surprisingly, neither substitution of each individual lysine with alanine or leucine, nor substitution of all of them with alanine resulted in growth inhibition. Radiolabeling experiments confirmed that the mutated proteins possessed the ability to take up sulfate, albeit with significantly affected kinetics. Except of the quadruple TRAstAK260-264A mutant, all variants of AstA with a single lysine in CCL mutated had lower transportation constants (Kτ) for sulfate, which indicated enhanced affinity for sulfate. However, we did not find any similar potentially beneficial substitution in sequences of AstA orthologues from other fungi. This apparently surprising finding may be explained by AstA’s ability to transport sulfite and choline sulfate ester (Holt et al., 2017), which implies that under physiological conditions CCL of AstA should be flexible enough to conform to all these substrates. This versatility of AstA might also prevent substitution of K263 or K264 by a hydrophobic amino acid, which does not occur in nature. While all four single lysine substitutions into alanine resulted in the decreased maximum velocity of sulfate transport, unexpectedly, we found two substitutions among K to L mutants (K263L and K264L) leading to the increased velocity of transportation by 82.4 and 142%, respectively (Table 2). These changes in kinetic properties might result from enhanced CCL flexibility during MFS transporter movements. Regarding that the XylE transporter CCL forms transient latch-like structure (Wisedchaisri et al., 2014), it might be difficult to verify the transient hydrophobic enzyme-substrate interactions relying solely on solid crystal structures.
Among the single amino-acid substitution mutants, TRAstA262L had the most affected sulfate uptake rate and velocity of transportation (Vmax). Its growth on solid minimal media was also significantly affected after 48 h, however, in a longer time TRAstA262L reached growth rate similar to that of the wild type TRAstAWT strain. None of the K262 substitutions studied here altered Kt, indicating that this lysine 262 is not involved in direct interaction with the sulfate.
The quadruple TRAstAK260-264A mutant with all four lysine residues substituted, exhibited the most visibly altered substrate transportation, which could also be observed as a decreased selenate resistance on a minimal solid medium. This result suggests that negatively charged lysine residues in wild type CCL of AstA are necessary for the maximum efficiency of the transporter. However, the quadruple K260-264A mutation in CCL did not abolish AstA function completely, as opposed to mutations in CCL of C. albicans Mdr1 multidrug transporter (Mandal et al., 2012). Shortening of CCL in Mdr1 eliminated the drug resistance, suggesting an important role of the loop for the functioning of this transporter. Since insertional mutagenesis restoring the loop length did not result in a functional transporter, one can infer that the amino acid composition of CCL is more relevant to substrate transport than its mere length.
Altogether, our results indicate that CCL is not essential for the function of the entire transporter, but seems to be engaged in controlling of AstA activity. Interestingly, it has been found earlier that CCL of the E. coli lactose permease is involved in allosteric regulation by the glucose-specific enzyme IIA of the phosphoenolpyruvate: sugar phosphotransferase system (Hoischen et al., 1996, Seok et al., 1997). Similarly, CCL in the human GLUT1 transporter mediates ATP-dependent inhibition of glucose transport (Blodgett et al., 2007). In similar way, the AstA CCL could interact with the sulfate and form an allosteric site controlling the transporter’s activity. Since AstA is ATP-independent, as we have shown (Fig. 6), fungi bearing this gene may have a higher evolutionary fitness under the low oxygen conditions, when ATP synthesis is limited, for instance in a situation of the pathogens infecting host tissue.
The findings presented by Weinglass and Kaback (Weinglass & Kaback, 2000) show that the cytoplasmic loop between helices VI and VII functions as a temporal delay spacer upon insertion of the first six helices into the translocon, and their subsequent movement into the bilayer before insertion of the last six helices. Additionally, sequence alignments of LacY CCL indicate that this region is very poorly conserved (Weinglass & Kaback, 2000). Detailed studies of E. coli LacY by Frillingos and coworkers (Frillingos et al., 1998) and Weinglass and Kaback (Weinglass & Kaback, 2000) show that the bacterial LacY CCL plays no direct role in the substrate transportation. Bacterial MFS may be easily split into two halves discarding CCL, and the generated mutant exhibits expression and activity comparable to the wild type (Weinglass & Kaback, 2000). Even an introduction of the stretches of random hydrophilic amino acid into the poorly active D20 aa CCL mutant, partially restores its activity, up to the 30–50% of the wild-type level. Thus, in a case of bacterial MFS transporters, CCL seems not to be particularly important for the protein function. In contrast to what was observed for the bacterial LacY transporter and its CCL, our studies indicate that AstA CCL sequence has tightly conserved length and high amino acid homology (over 70%) among numerous fungal species (Fig. 2A).
In this paper, we demonstrated that eukaryotic MFS may have different kinetics than bacterial transporters. In contrast to E. coli LacY, the fungal AstA mutants: single substituted K260, K264 or quadruple K260-264A led to a competitive inhibition of sulfate transportation by altering Kt.
Since the lysine substitutions in CCL resulted in the altered sulfate transportation constants (Kτ) in the AstA transporters, it seems likely that CCL limits the AstA affinity for substrate under the conditions of high intracellular sulfate concentration. Consequently, the substitutions of the lysine residues in CCL could abolish that allosteric regulation, resulting in an altered sulfate affinity of the mutated AstA variants. The most pronounced shift in Kτ was observed for quadruple TRAstAK260-264A mutant, which was also susceptible to selenate, a toxic analog of sulfate. Since an alteration of a substrate binding constant usually refers to a competitive inhibition of the enzyme binding site (Strelow et al., 2012), our results indicate that CCL indeed interacts with the sulfate. Whether the proposed binding of sulfate by CCL would be assisted by interaction with other proteins, as it is in the case of the E. coli lactose permease, remains an open question.
The observed physiological effects of the mutations affecting CCL could be alternatively explained, if one assumed that CCL formed a proofreading centre (or a checkpoint), controlling the parameters of the incoming substrate. Such possibility is supported by our observation that the K260A, K260L, K263A and K264L mutants of AstA with decreased Kt (Table 2) were more resistant to selenate (Fig. 3), suggesting that these mutations could affect interaction with selenate rather than with the sulfate. Hence, CCL could be a flexible goalkeeper of the AstA transporter, forming a transient substrate-interacting pocket, acting as a checkpoint for the molecules entering the cell.
Numerous biochemical studies characterizing the MFS transporters were focused on substrate translocation pore, located within TM helices. However, Kτ or Km values are still poorly studied, and CCL has not been well investigated. The mutations introduced in substrate binding site of MFS transporters usually lead to a pronounced 10-fold Km alteration (Will et al., 1998; Sahin-Tóth et al., 1999). Among some previously described enzymes, S-1,2-propanediol oxidoreductase (FucO) alters its substrate specificity substantially upon just a single point mutation L259V in the substrate-binding site, while its Km increases just two-fold (Blikstad et al., 2014). In case of the higher eukaryotes, some mutated enzymes, it was shown that human galactokinase, point-mutated in its substrate-binding site (A198V), has Km only weakly altered (30% decrease) and its kinetic parameters are almost comparable to the wild type (Timson & Reece, 2003). However, the A198V mutation in galactokinase is severely manifested on the cellular and organism level, with a clinical phenotype of a tendency to develop cataracts (Okano et al., 2001). Since there is no data referring to kinetic parameters of eukaryotic CCL, it was not possible to compare our results to published values.
Crystal studies of the bacterial XylE transporter revealed that CCL can act as a structural linker between the N- and C-terminal domains of MFS proteins, transiently forming a latch-like structure (Wisedchaisri et al., 2014). Till now, CCL was regarded as a structural linker only, but we have shown that this loop can affect the substrate uptake. Considering that some mutations located within the linker resulted in the significant Kτ shifts, CCL may also participate in substrate recognition. Summarizing, the presented results suggest that the four lysine residues located within the CCL linker region are not necessary for the sulfate transport, but they select for the incoming substrate and facilitate its uptake.
Acknowledgements
We are indebted to Dr. Małgorzata Lichocka for her help in confocal microscopy and to Prof. Andrzej Paszewski for his careful reading of the manuscript. We are also indebted to Dr. Joanna Kamińska for a gift of the anti-aPma1p antibodies.
Acknowledgements of Financial Support
This work was supported by the State Committee for Scientific Research/National Center of Science (NCN), grant number N N303 814040 to S.P. and by The European Regional Development Fund and Innovative Economy, Poland, grant number UDA-POIG.01.03.01-14-038/09 to J.S.K. The equipment used was sponsored in part by the Centre for Preclinical Research and Technology (CePT), a project co-sponsored by The Innovative Economy, The National Cohesion Strategy of Poland.
References
Abramson J, Smirnova I, Kasho V, Verner G, Kaback HR, Iwata S (2003) Structure and mechanism of the lactose permease of Escherichia coli. Science 301: 610–615. https://doi.org/10.1126/science.1088196
Alper SL, Sharma AK (2013) The SLC26 gene family of anion transporters and channels. Mol Aspects Med 34: 494–515. https://doi.org/10.1016/j.mam.2012.07.009
Altschul SF, Lipman DJ (1990) Protein database searches for multiple alignments. Proc Natl Acad Sci USA 87: 5509–5513
Arst HN Jr (1968) Genetic analysis of the first steps of sulphate metabolism in Aspergillus nidulans. Nature 219: 268–270. https://doi:10.1038/219268a0
Biswas C, Djordjevic JT, Zuo X, Boles E, Jolliffe KA, Sorrell TC, Chen SC (2013) Functional characterization of the hexose transporter Hxt13p: an efflux pump that mediates resistance to miltefosine in yeast. Fungal Genet Biol 61: 23–32. https://doi.org/10.1016/j.fgb.2013.09.005
Blikstad C, Dahlström KM, Salminen TA, Widersten M (2014) Substrate scope and selectivity in offspring to an enzyme subjected to directed evolution. FEBS J 281: 2387–2398. https://doi.org/10.1111/febs.12791
Blodgett DM, De Zutter JK, Levine KB, Karim P, Carruthers A (2007) Structural basis of GLUT1 inhibition by cytoplasmic ATP. J Gen Physiol 130: 157–168. https://doi.org/10.1085/jgp.200709818
Bourne Y, Dannenberg J, Pollmann V, Marchot P, Pongs O (2001) Immunocytochemical localization and crystal structure of human frequenin (neuronal calcium sensor 1). J Biol Chem 276: 11949–11955. https://doi.org/10.1074/jbc.M009373200
Bowler MW, Montgomery MG, Leslie AG, Walker JE (2006) How azide inhibits ATP hydrolysis by the F-ATPases. Proc Natl Acad Sci U S A 103: 8646–8649. https://doi.org/10.1073/pnas.0602915103
Buchan DW, Minneci F, Nugent TC, Bryson K, Jones DT (2013). Scalable web services for the PSIPRED Protein Analysis Workbench. Nucleic Acids Res 41 (Web Server issue): W340-W348. https://doi.org/10.1093/nar/gkt381
Cerqueira GC, Arnaud MB, Inglis DO, Skrzypek MS, Binkley G, Simison M, Miyasato, SR, Binkley J, Orvis J, Shah P, Wymore F, Sherlock G, Wortman JR (2014). The Aspergillus Genome Database: multispecies curation and incorporation of RNA-Seq data to improve structural gene annotations. Nucleic Acids Res 42: D705–D710. https://doi.org/10.1093/nar/gkt1029
Clutterbuck AJ (1994) Linkage map and locus list. Prog Ind Microbiol 29: 791–824
Dang S, Sun L, Huang Y, Lu F, Liu Y, Gong H, Wang J, Yan N (2010) Structure of a fucose transporter in an outward-open conformation. Nature 467: 734–738. https://doi.org/10.1038/nature09406
Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res 32: 1792–1797. https://doi.org/10.1093/nar/gkh340
Ethayathulla AS, Yousef MS, Amin A, Leblanc G, Kaback HR (2014) Structure-based mechanism for Na+/melibiose symport by MelB. Nature Commun 5: 3009. https://doi.org/10.1038/ncomms4009
Frillingos S, Sahin-Tóth M, Wu J, Kaback HR (1998) Cys-scanning mutagenesis: a novel approach to structure function relationships in polytopic membrane proteins. FASEB J 12: 1281–1299. https://doi.org/10.1096/fasebj.12.13.1281
Gems DH, Clutterbuck AJ (1993) Co-transformation with autonomously-replicating helper plasmids facilitates gene cloning from an Aspergillus nidulans gene library. Curr Genet 24: 520–524
Hirai T, Heymann JA, Shi D, Sarker R, Maloney PC, Subramaniam S (2002) Three-dimensional structure of a bacterial oxalate transporter. Nat Struct Biol 9: 597–600. https://doi.org/10.1038/nsb821
Hofmann K, Stoffel W (1993) TMbase – A database of membrane spanning proteins segments. Biol Chem Hoppe-Seyler 374: 166
Hoischen C, Levin J, Pitaknarongphorn S, Reizer J, Saier MH Jr (1996) Involvement of the central loop of the lactose permease of Escherichia coli in its allosteric regulation by the glucose-specific enzyme IIA of the phosphoenolpyruvate-dependent phosphotransferase system. J Bacteriol 178: 6082–6086
Holt S, Kankipati H, De Graeve S, Van Zeebroeck G, Foulquié-Moreno MR, Lindgreen S, Thevelein JM (2017) Major sulfonate transporter Soa1 in Saccharomyces cerevisiae and considerable substrate diversity in its fungal family. Nat Commun 8: 14247. https://doi.org/10.1038/ncomms14247
Iancu CV, Zamoon J, Woo SB, Aleshi, A, Choe JY (2013) Crystal structure of a glucose/H+ symporter and its mechanism of action. Proc Natl Acad Sci U S A 110: 17862–17867. https://doi.org/10.1073/pnas.1311485110
Kasahara T, Ishiguro M, Kasahara M (2006) Eight amino acid residues in transmembrane segments of yeast glucose transporter Hxt2 are required for high affinity transport. J Biol Chem 281: 18532–18538. https://doi.org/10.1074/jbc.M602123200
Kasahara T, Maeda M, Bole,s E, Kasahara M (2009) Identification of a key residue determining substrate affinity in the human glucose transporter GLUT1. Biochim Biophys Acta 1788: 1051–1055. https://doi.org/10.1016/j.bbamem.2009.01.014
Kertesz MA (2001) Bacterial transporters for sulfate and organosulfur compounds. Res Microbiol 152: 279–290
Kuwano T, Shirataki C, Itoh Y (2008) Comparison between polyethylene glycol- and polyethylenimine-mediated transformation of Aspergillus nidulans. Curr Genet 54: 95–103. https://doi.org/10.1007/s00294-008-0204-z
Law CJ, Maloney PC, Wang DN (2008) Ins and outs of major facilitator superfamily antiporters. Annu Rev Microbiol 62: 289–305. https://doi.org/10.1146/annurev.micro.61.080706.093329
Lemieux MJ, Huan, Y, Wang DN (2004) The structural basis of substrate translocation by the Escherichia coli glycerol-3-phosphate transporter: a member of the major facilitator superfamily. Curr Opin Struct Biol 14: 405–412. https://doi.org/10.1016/j.sbi.2004.06.003
Letunic I, Bork P (2017) 20 years of the SMART protein domain annotation resource. Nucleic Acids Res 46(D1): D493–D496. https://doi:10.1093/nar/gkx922
Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193: 265–275
Łukaszkiewicz Z, Paszewski A (1976) Hyper-repressible operator-type mutant in sulphate permease gene of Aspergillus nidulans. Nature 259: 337–338. https://doi.org/10.1038/259337a0
Mandal A, Kumar A, Singh A, Lynn AM, Kapoor K, Prasad R (2012) A key structural domain of the Candida albicans Mdr1 protein. Biochem J 445: 313–322. https://doi.org/10.1042/BJ20120190
Martinelli SD (1994) Gene symbols. Prog Ind Microbiol 29: 825–827
Okano Y, Asada M, Fujimoto A, Ohtake A, Murayama K, Hsiao KJ, Choeh K, Yang Y, Cao Q, Reichardt JK, Niihira S, Imamura T, Yamano T (2001) A genetic factorfor age-related cataract: identification and characterization of a novel galactokinase variant, ‘Osaka’, in Asians. Am J Hum Genet 68: 1036–1042. https://doi.org/10.1086/319512
Pasrija R, Banerjee D, Prasad R (2007) Structure and function analysis of CaMdr1p, a major facilitator superfamily antifungal efflux transporter protein of Candida albicans: identification of amino acid residues critical for drug/H+ transport. Eukaryot Cell 6: 443–453. https://doi.org/10.1128/EC.00315-06
Patron M, Checchetto V, Raffaello A, Teardo E, Reane DV, Mantoan M, Granatiero V, Szabo I, De Stefani D, Rizzuto R (2014) MICU1 and MICU2 finely tune the mitochondrial Ca2+ uniporter by exerting opposite effects on MCU activity. Mol Cell 53: 726–737. https://doi.org/10.1016/j.molcel.2014.01.013
Paulsen IT, Brown MH, Skurray RA (1996) Proton-dependent multidrug efflux systems. Microbiol Rev 60: 575–608
Pedersen BP, Kumar H, Waight AB, Risenmay AJ, Roe-Zurz, Z., Chau BH, Schlessinger A, Bonomi M, Harries W, Sali A, Johri AK, Stroud RM (2013) Crystal structure of a eukaryotic phosphate transporter. Nature 496: 533–536. https://doi.org/10.1038/nature12042
Piłsyk S, Natorff R, Sieńko M, Paszewski A (2007) Sulfate transport in Aspergillus nidulans: a novel gene encoding alternative sulfate transporter. Fungal Genet Biol 44: 715–725. https://doi.org/10.1016/j.fgb.2006.11.007
Piłsyk S, Paszewski A (2009) Sulfate permeases – phylogenetic diversity of sulfate transport. Acta Biochim Pol 56: 375–384
Piłsyk S, Natorff R, Gawińska-Urbanowicz H, Kruszewska JS (2015) Fusarium sambucinum astA gene expressed during potato infection is a functional orthologue of Aspergillus nidulans astA. Fungal Biol 119: 509–517. https://doi.org/10.1016/j.funbio.2015.02.002
Rost B, Yachdav G, Liu J (2004) The PredictProtein Server. Nucleic Acids Res 32 (Web Server issue): W321–W326. https://doi.org/10.1093/nar/gkh377
Roth V (2006) Doubling Time Computing, Available from: http://www.doubling-time.com/compute.php
Roy A, Kucukural A, Zhang Y (2010) I-TASSER: a unified platform for automated protein structure and function prediction. Nature Protocols 5: 725–738. https://doi.org/10.1038/nprot.2010.5
Roy A, Yang J, Zhang, Y (2012) COFACTOR: an accurate comparative algorithm for structure-based protein function annotation. Nucleic Acids Res 40 (Web Server issue): W471-7. https://doi.org/10.1093/nar/gks372
Sahin-Tóth M, le Coutre J, Kharabi D, le Maire G, Lee JC, Kaback HR (1999) Characterization of Glu126 and Arg144, two residues that are indispensable for substrate binding in the lactose permease of Escherichia coli. Biochemistry 38: 813–819. https://doi.org/10.1021/bi982200h
Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press. https://doi.org/10.1016/0092-8674(90)90210-6
Sayle RA, Milner-White EJ (1995) RASMOL: biomolecular graphics for all. Trends Biochem Sci 20: 374. https://doi.org/10.1016/S0968-0004(00)89080-5
Seok YJ, Sun J, Kaback HR, Peterkofsky A (1997) Topology of allosteric regulation of lactose permease. Proc Natl Acad Sci U S A 94: 13515–13519. https://doi.org/10.1073/pnas.94.25.13515
Sieńko M, Natorff R, Skoneczny M, Kruszewska J, Paszewski A, Brzywczy J (2014) Regulatory mutations affecting sulfur metabolism induce environmental stress response in Aspergillus nidulans. Fungal Genet Biol 65: 37–47. https://doi.org/10.1016/j.fgb.2014.02.001
Strelow J, Dewe W, Iversen PW, Brooks HB, Radding JA, McGee J, Weidner J (2012) Mechanism of Action Assays for Enzymes, In Assay Guidance Manual, Sittampalam GS, Coussens NP, Nelson H et al, eds, Eli Lilly & Company and the National Center for Advancing Translational Sciences, 2004
Sun L, Zeng X, Yan C, Sun X, Gong X, Rao Y, Yan N (2012) Crystal structure of a bacterial homologue of glucose transporters GLUT1-4. Nature 490: 361–366. https://doi.org/10.1038/nature11524
Timson DJ, Reece RJ (2003) Functional analysis of disease-causing mutations in human galactokinase. Eur J Biochem 270: 1767–1774. https://doi.org/10.1046/j.1432-1033.2003.03538.x
Tweedie JW Segel IH (1970) Specificity of transport processes for sulfur, selenium, and molybdenum anions by filamentous fungi. Biochim Biophys Acta 196: 95–106. https://doi.org/10.1016/0005-2736(70)90170-7
Weinglass AB, Kaback HR (2000) The central cytoplasmic loop of the major facilitator superfamily of transport proteins governs efficient membrane insertion. Proc Natl Acad Sci U S A 97: 8938–8943. https://doi.org/10.1073/pnas.140224497
Will A, Grassl R, Erdmenger J, Caspari T, Tanner W (1998) Alteration of substrate affinities and specificities of the Chlorella Hexose/H+ symporters by mutations and construction of chimeras. J Biol Chem 273: 11456–11462. https://doi.org/10.1074/jbc.273.19.11456
Wisedchaisri G, Park MS, Iadanza MG, Zhen H, Gonen T (2014) Proton-coupled sugar transport in the prototypical major facilitator superfamily protein XylE. Nat Commun 5: 4521. https://doi.org/10.1038/ncomms5521
Woodin TS Wang JL (1989) Sulfate permease of Penicillium duponti. Exp Mycol 13:380-391. https://doi.org/10.1016/0147-5975(89)90034-0
Yin Y, He X, Szewczyk P, Nguyen T, Chang G (2006) Structure of the multidrug transporter EmrD from Escherichia coli. Science 312: 741–744. https://doi.org/10.1126/science.1125629
Zhang Y (2008) I-TASSER server for protein 3D structure prediction. BMC Bioinformatics 9: 40. https://doi.org/10.1186/1471-2105-9-40