Arabidopsis (Arabidopsis thaliana) plants were grown on soil at 24°C, 16 h light per day. The following lines were used: Landsberg erecta (Ler), Columbia-0 (Col-0), an-2 (Kim et al., 2002), an-1 (Kim et al., 2002), an-doq (Bai et al., 2013), 35S:PAB2-RFP [kindly provided by Prof. Julia Bailey-Serres, University of California, Riverside (Sorenson and Bailey-Serres, 2014)] and new lines established in this study including 35S:YFP-AN, 35S:AN-YFP, 35S:YFP-AN^DOQ1, 35S:YFP-AN^{GAD→V V A}, 35S:YFP-UBP1B, and 35S:YFP-EIF4E1. N-terminal YFP and C-terminal YFP fusions were created by recombination of the corresponding entry clones with the gateway destination vectors pENS-G-YFP and pEXS-G-YFP (Feys et al., 2005). Transformation of Arabidopsis thaliana was done by the Floral Dip method (Clough and Bent, 1998) and transgenic plants were selected in the T1 by Glufosinat (BASTA^®).

CDSs of all the genes were amplified from Col-0 cDNA and cloned through BP recombination in pDONR201 or pDONR207 (Invitrogen), sequenced and introduced into destination vectors by LR reactions (Invitrogen). List of constructs and primers used in this study with the AGI code are provided in the Supplementary Table S3. Mutant entry clones for AN^DOQ and AN^{GAD→V V A} were created by site-directed mutagenesis. AN^DOQ has a G170→D mutation mimicking the an-doq allele (Bai et al., 2013). AN^{GAD→V V A} has mutations at three sites important for NADH binding: G170→V; A175→V and D→193A. For Lumier, the destination vectors pcDNA3-Rluc-GW and pTREXdest30 (Invitrogen) were used which enable the N-terminal fusion of Renilla reniformis and Staphylococcus aureus proteins, respectively, and were described in detail before (Pesch et al., 2013; Pesch et al., 2015). Protein fusions were expressed in the human HEK293TN cells.

Dark grown Arabidopsis cell culture (wild-type Ler) was transfected with pENSG 35S:YFP-AN-strep transformed in Agrobacterium strain GV3101-pMP90RK as described before (Pesch et al., 2014) and grown at 22°C and 120 rpm in the dark for 5 days. The cells were harvested, pellets were crushed in liquid nitrogen, and protein crude extracts were prepared by homogenizing the cell pellets in 500 ml protein extraction buffer [50 mM Tris pH 7.5; 150 mM NaCl; 1 mM EDTA; 10% Glycerol; 5 mM DTT; 1% protease inhibitor cocktail (cOmpletes^TM, Roche); phosphatase inhibitor cocktail (PhosSTOP^TM, Roche); 1% Triton-X100]. After two centrifugation steps (12,000 rpm, 4°C, 20 min) the supernatant was incubated with Strep-Tactin^® MacroPrep (IBA) for 1 h followed by five washings (lysis buffer without Triton-X100 and Glycerol). The protein was eluted using 2.5 mM desthiobiotin containing elution buffer.

Luminescence Based Mammalian Interactome System (LUMIER) assays were done as described previously (Pesch et al., 2013, 2015). Luciferase activity was determined before (input) and after the pulldown. Cells transformed with empty vectors served as a negative control. Percentage pulldown efficiency was calculated by: [relative luminescence unit pulldown/(relative luminescence unit input)] × 100.

We used the Leica TCS-SP8/SPE confocal microscope equipped with the LCS software. Images were made using 20 or 40x water/oil immersion objectives. YFP was excited at 488 nm and detected at 510–560 nm. RFP was excited at 561 nm and detected in the 580–650 nm range. ImageJ was used for particle count/size analysis from maximum projections of Z series. Laser intensity, brightness and contrast settings were kept the same for an unbiased analysis for the particle count in wild-type and mutant backgrounds. For particle analysis, we considered granules between 0.185 and 5 μm² to exclude the background and nuclear signal from the images. Co-localization analysis was performed by calculating Pearson’s coefficient (Adler and Parmryd, 2010; Dunn et al., 2011) using the JACOP plugin of ImageJ. Z-stack images were obtained and merged to maximum projection. For quality control, the threshold was set for each image manually such that background noise, but not distinct aggregates were eliminated. Analysis was performed on the entire image. Microscopic settings for the localization of AN were chosen to enable a safe visualization of the weakly labeled granules allowing signal saturation in the big, strongly fluorescent AN granules that were not considered in our analysis.

Seeds were sterilized and plated on 1/2 MS medium and stratified for 4 days at 4°C in the dark and transferred to normal growth conditions. After 5 days, seedlings were transferred to medium supplemented with 150/175 mM NaCl for analyzing salt tolerance and 200/250 mM mannitol for analyzing osmotic stress tolerance. Six and nine days after transfer to NaCl or mannitol media, seedlings were analyzed for tolerance to stress conditions by analyzing cotyledon greening and root length under stress/non-stress conditions. Root length was measured using ImageJ.

Transfection of Arabidopsis leaves was performed by biolistic transformation (Mathur et al., 2003) and analyzed after 12–16 h by Confocal Laser Scanning Microscopy (CLSM).

pACT2 and pAS2 plasmids (Clontech) were used for the fusion of the proteins with GAL4 activation domain and GAL4 DNA binding domain respectively. Yeast two-hybrid assays were done as previously described (Gietz et al., 1995) and interactions were analyzed on synthetic dropout medium without leucine, tryptophan, and histidine supplemented with varying concentrations of 3-aminotriazole (3-AT).

Mass spectrometry analysis was performed as described previously (Borgal et al., 2014; Hosseinibarkooie et al., 2016). Sample analysis was performed on an LTQ Orbitrap Discovery mass spectrometer (Thermo Scientific) coupled to an EASYnLC II nano-LC system (Proxeon, part of Thermo Scientific).

AN (AT1G01510), PAB2 (AT4G34110), UBP1B (AT1G17370), EIF4E1 (AT4G18040).

In an attempt to understand the molecular function of AN in more detail, we searched for proteins interacting with AN using the yeast two-hybrid screens. Although we performed several screening rounds with the full length AN protein as a bait on different libraries, we isolated only two interactors, ANGUSTIFOLIA INTERACTING KINASE (AIK1, AT3g17750) and Asymmetric Leaf Enhancer 3 (AE3, AT5g05780). In parallel, we performed a pulldown experiment using an Arabidopsis cell culture expressing N-terminal YFP tagged AN with a Strep tag at the C-terminus. AN was affinity purified using Strep-tactin resin and co-purified proteins were analyzed by mass spectrometry. We identified 62 co-precipitated proteins from AN expressing cell cultures that were not found in the control cell culture (Supplementary Tables S1, S2). We noted that 12 co-purified proteins are predicted to be associated with RNA metabolism suggesting that AN might be involved in the regulation of the RNA metabolism (Table 1). To test this further, we studied the protein–protein interactions of AN with selected proteins in quantitative pulldown experiments in HEK cells (Table 1), using the LUMIER (Barrios-Rodiles et al., 2005; Pesch et al., 2013). In short, a Prot-A tagged protein is co-transfected with Renilla reniformis luciferase in HEK cells and the amount of the interacting protein is determined after co-immunoprecipitation by a quantitative analysis of the luminescence (Barrios-Rodiles et al., 2005). The AN protein was always used as a luciferase-fusion and the putative interactors as Prot-A fusions. We chose seven RNA associated proteins that we isolated in the AN pulldown and 16 well-established RBPs chosen from the literature for the interaction studies (Table 1). In addition, we included the yeast two-hybrid interactor AIK1 as it is homologous to dual-specificity tyrosine phosphorylation-regulated kinase 3 (DYRK3) which was shown to control the dissolution of SGs in mammals through its role in mTOR signaling (Wippich et al., 2013). As summarized in Table 1, we found direct interaction with several but not all RNA associated proteins including RNA BINDING PROTEIN 47B (RBP47B; AT3G19130), RNA BINDING PROTEIN 47C (RBP47C; AT1G47490), RNA BINDING PROTEIN 47C′ (RBP47C′; AT1G47500), EUKARYOTIC INITIATION FACTOR 4E1 (EIF4E1; AT4G18040), EIF(iso)4E (AT5G35620), TANDEM ZINC FINGER 3 (TZF3; AT4G29190), ARGONAUTE1 (AGO1; AT1G48410), POLY(A)BINDING PROTEIN 2 (PAB2; AT4G34110), ANGUSTIFOLIA INTERACTING KINASE (AIK1, AT3g17750).

Our LUMIER data show that AN binds to several proteins localizing to mRNP complexes. mRNP complexes are typically formed under stress conditions (Kedersha et al., 2005; Weber et al., 2008). We therefore investigated the localization of AN before and after different stress treatments. Previous reports have demonstrated that 35S:AN-GFP shows the same intracellular localization as ANp:AN-GFP and completely rescues the an mutant phenotype (Minamisawa et al., 2011). In this study, we established two lines expressing either AN-YFP or YFP-AN under the 35S promoter in the an-2 mutant background. These lines exhibit complete phenotypic rescue. Under normal conditions, both lines showed low cytoplasmic signal and one or few bright dots along with a few smaller dots showing weaker fluorescence, similar as reported before (Minamisawa et al., 2011). In a first experiment, we applied heat stress (39°C) to rosette leaves of AN-YFP plants for 40 min (Gutierrez-Beltran et al., 2015). When comparing leaves before and after the treatment we found the formation of many new cytosolic granules after heat stress (Figures 1A,B). The response to salt, osmotic and hypoxia was analyzed in the cotyledons of 5–7 days old seedlings grown on agar plates. Plants were transferred to depression slides and stress was induced by adding ½ MS solution containing 175 mM sodium chloride for salt stress conditions, 800 mM sorbitol (Wippich et al., 2013) for osmotic stress and 8 units/ml oxyrase for the induction of hypoxia (Sorenson and Bailey-Serres, 2014; Lokdarshi et al., 2016). All treatments caused the formation of many cytosolic granules (Figures 1C–H). Control experiments with ½ MS did not show a change in AN localization behavior (Supplementary Figure S1). We did not observe any differences in the recruitment behavior to granules between the C- and N- terminal fusions (AN-YFP and YFP-AN).

The stress dependent formation of AN-labeled granules combined with our finding that AN protein binds to RNA-associated proteins prompted us to test, whether AN localizes to mRNP granules or SGs. Toward this end, we co-expressed established XFP-labeled SG markers with XFP-AN in single epidermal cells in Arabidopsis rosette leaves by particle bombardment. We selected four established SG markers: PAB2 (Sorenson and Bailey-Serres, 2014), UBP1B (Weber et al., 2008; Sorenson and Bailey-Serres, 2014), RBP47C (Weber et al., 2008; Lokdarshi et al., 2016) and EIF4E1 (Weber et al., 2008). For the analysis of the co-localization of PAB2 (Sorenson and Bailey-Serres, 2014) and UBP1B (Weber et al., 2008; Sorenson and Bailey-Serres, 2014) we analyzed the same cell before and after 40 min heat stress (39°C) (Figure 2). After heat stress, we found a prominent co-localization of AN with both RNP-markers in small granules (Figures 2A,B). Similar results were obtained for the co-localization of AN with RBP47C (Supplementary Figure S2A; Weber et al., 2008; Lokdarshi et al., 2016) and EIF4E1 (Supplementary Figure S2B; Weber et al., 2008). Together these data indicate that AN protein can be recruited to SGs in a heat stress dependent manner. To determine whether AN localizes specifically to SGs or more generally to mRNP granules, we studied its co-localization with P-body marker DCP1. While SGs are thought to function as sites for transient RNA storage, P-bodies serve as platforms for RNA processing and decay (Anderson and Kedersha, 2008). Toward this end, we co-expressed mCHERRY-AN, YFP-UBP1B and the P-body marker CFP-DCP1 (Weber et al., 2008) (Supplementary Figure S3). While AN co-localized well with UBP1B, we found no convincing co-localization with DCP1 (Supplementary Figure S3). Occasionally we noted an association of AN and DCP1 labeled granules similarly as observed for DCP1a and TIA-1 (Kedersha et al., 2005).

As the transformation of cells by particle bombardment may cause artificially high and thereby altered protein behavior, we confirmed the co-localization of the SG marker Poly-adenylate binding protein 2 (PAB2) with AN after stress in a stable transgenic line. PABP protein family is an integral SG component in mammals and is a universal marker for all SGs (Anderson and Kedersha, 2008). AtPAB2 has been shown to co-localize with another SG marker UBP1C (Sorenson and Bailey-Serres, 2014) and another member of PAB protein family, PAB8 has been used as a SG marker in Arabidopsis (Pomeranz et al., 2010). We crossed the YFP-AN line with the PAB2-RFP line (Sorenson and Bailey-Serres, 2014) and studied the protein co-localization in the F1 generation. As expected, YFP-AN was found in a big dot and few smaller dots and PAB2-RFP was cytoplasmic under normal conditions with (Figures 3A–C) and without YFP-AN (Supplementary Figure S4). We tested the response to heat stress in mature leaf cells and in the cotyledons of young seedlings. In both tissues we observed YFP-AN and/or PAB2-RFP positive granules (Figures 3D–I) after 40 min heat stress (39°C), which displayed a partial co-localization. The number of granules clearly showing both signals appeared to be lower than in the particle bombardment experiments (Figures 3G–I). This notion was confirmed by a lower Pearson coefficient of co-localization 0.30 ± 0.04 (n = 12 rosette leaves), 0.34 ± 0.4 (n = 8 cotyledons) in transgenic lines as compared to transient expression experiments (0.73 ± 0.12, n = 13). In our experiments we noted that both, the markers and AN can occur as single-labeled dots. Although we cannot resolve this at this stage, this can in principle result from different threshold levels for the detection of two proteins or be explained by qualitatively different granule populations. Taken together, these results show that heat stress dependent AN co-localization with SG markers occurs, although at different degrees, in transient expression assays as well as in transgenic lines.

In order to test, whether AN has a functional role in SG formation, we created transgenic marker lines expressing YFP-UBP1B and YFP-EIF4E1 under 35S promoter in an-2, an-doq and the respective wild-type backgrounds. In a first step, we quantified the number of YFP-UBP1B and YFP-EIF4E1 labeled SGs after 40 min heat stress (39°C). Cell sizes differ a lot in a given leaf and DNA content can range from 2C and 16C, it is therefore difficult to interpret the number of SGs per cell. For the comparison of wild-type and an mutants, we therefore studied the density per μm². This should reflect the physiologically relevant cytoplasmic density of SGs in the cells. For both markers, we found a statistically significant higher number of SGs in both an alleles as compared to the wild-type (Figure 4 and Supplementary Figure S5). This suggests that AN counteracts the formation of SGs. We additionally analyzed the size distribution of SGs in an mutants and wild-type. In both alleles we found a trend toward an increased number of smaller granules (Supplementary Figure S5). Thus, AN is required for the regulation of number and possibly the size of SGs.

Our finding that the an-doq mutant, which has a point mutation in the NAD(H) binding domain, displays an altered number and size of SGs, prompted us to study the function of the NAD(H) binding domain in more detail. Toward this end we used two mutant proteins: AN^DOQ, which is a mimic of an-doq mutant and AN^{GAD→V V A}, which carries three amino acid exchanges in the NAD(H) domain potentially disrupting the NADH binding.

We expressed both mutant proteins as N-terminal YFP fusions in the an-2 mutant under the control of the 35S promoter. Although an-doq mutant shows a null phenotype, overexpression of YFP-AN^DOQ rescued the an-2 mutant morphological phenotypes suggesting that AN^DOQ protein has some residual activity. In contrast, YFP-AN^{GAD→V V A} overexpression showed weak rescue of the leaf length/width ratio and no rescue of the trichome phenotype (Supplementary Figure S6). Both proteins were cytoplasmic (Figures 5A,D). In contrast to wild-type AN protein (Figure 1), we found no dots under normal conditions. Upon heat stress, YFP-AN^DOQ and YFP-AN^{GAD→V V A} labeled granules were formed (Figures 5B,C,E,F). This suggests that the recruitment of both proteins to SGs can occur in the absence of wild-type AN protein.

In a next step, we analyzed the co-localization of YFP-AN^DOQ and YFP-AN^{GAD→V V A} with the SG marker PAB2 in double transgenic lines. Without stress, we observed only cytoplasmic signals (Figures 6A–C and Supplementary Figures S7A–C). After heat stress, PAB2-RFP and YFP-AN^DOQ and YFP-AN^{GAD→V V A} positive granules were found (Figures 6D–I and Supplementary Figures S7D–I). Strikingly, YFP-AN^DOQ and YFP-AN^{GAD→V V A} signals appeared to almost completely co-localize with PAB2-RFP in contrast to the partial co-localization observed in case of wild-type protein (Figure 3). This visual impression was confirmed by a much higher Pearson coefficients of YFP-AN^DOQ (0.75 ± 0.06) and YFP-AN^{GAD→V V A} (0.77 ± 0.09) with PAB2-RFP as compared to wild-type AN (Pearson coefficient: 0.30 ± 0.04).

Previous reports on the role of the NAD(H) domain of CtBP in regulating its protein–protein interactions (Balasubramanian et al., 2003; Fjeld et al., 2003) prompted us to study the dimerization and interaction ability of AN^DOQ and AN^{GAD→V V A} proteins. In yeast two-hybrid assays, the dimerization of wild-type AN was observed at 3AT concentrations up to 30 mM whereas we detected no dimerization between AN^DOQ proteins (Supplementary Figure S8). To independently confirm this result, we used the LUMIER assay. Here we found a marked reduction of the interaction between AN^DOQ proteins (Figure 7A). Similar results were obtained for the AN^{GAD→V V A} protein (Figure 7A). In order to determine, whether the NAD(H) domain mutant proteins are generally impaired in protein–protein interactions or whether only specific protein interactions depend on this domain, we studied the interaction of AN^DOQ with five AN-interactors including AIK1, AGO1, RBP47B, PAB2, and EIF(iso)4E. We found a drastically reduced interaction of AN^DOQ with AIK1 (Figure 7B) (reduced to 1.6% of wild-type AN). However, we did not detect significant changes in interaction ability with AGO1, RBP47B, PAB2 and EIF(iso)4E (Figure 7B). This indicates that the NAD(H) binding domain of AN is relevant for some but not all interactions.

Our findings that AN is recruited to SGs upon stress treatment and its role in regulating SG number and size raised the question whether AN has, in addition to the well-reported cell morphogenesis phenotypes, also a role in stress responses. First evidence for a role of AN in biotic and abiotic stress tolerance was reported by Gachomo et al. (2013). In this study, we chose salt and osmotic stress: two conditions under which we observed AN recruitment to SGs and for which standardized treatments are well-established. an mutants and wild-type plants were grown on ½ MS plate supplemented with different concentration of salt or mannitol. After 6–9 days of growth, the primary root length and the greening of the cotyledons was compared between non-treated and stress treated seedlings. At 150 and 175 mM NaCl concentrations an mutants displayed hyposensitivity to salt stress (Figure 8). an mutants showed longer roots and greener cotyledons as compared to wild-type. Similarly, an mutants were less sensitive than wild-type to 200 and 250 mM mannitol treatments (Figure 8).

How can these novel findings be integrated to the current concepts on the molecular function of AN? Function and localization of AN has been controversial since its discovery. Some reports suggest it to be involved in transcriptional control similar as known for its mammalian homolog CtBP (Chinnadurai, 2007). In support of this, microarray analysis revealed differential gene expression between wild-type and an mutants in Arabidopsis (Kim et al., 2002; Fulton et al., 2009; Bai et al., 2013). This view is challenged by the findings that AN cannot rescue drosophila CtBP mutants (Stern et al., 2007) and functions outside the nucleus where it partially localizes to the trans-Golgi network (Minamisawa et al., 2011). Consistent with this, we found various membrane trafficking proteins in our pulldown experiments that are associated with AN (Supplementary Table S1) such as VPS34 and KEULE. Thus, AN appears to be associated with both membrane trafficking components as well as with mRNP components suggesting a dual function in the cytoplasm.

A possible role of AN in SGs is suggested by its interaction with various SG components and its localization to SGs after various stress treatments. The interactions of AN to SG components is also seen under non-stress conditions in our assays. This suggests that AN might generally interact with SG components under non-stress and stress conditions also in plant cells. Given that various SG proteins are involved in translational processes, it is possible that AN functions in the translational machinery. This would provide an elegant explanation for its broad range of functions in development, morphogenesis and biotic/abiotic stress responses. Nevertheless, our finding that SG formation is altered in an mutants indicates that AN has a relevant function in SGs (Figure 4 and Supplementary Figure S5).

A major mechanism by which CtBPs regulate transcriptional events is by their ability to bind NAD(H) which results in a conformational change in CtBPs, in turn effecting its binding to interaction partners (Balasubramanian et al., 2003; Kuppuswamy et al., 2008; Nardini et al., 2009; Zhang and Arnosti, 2011). CtBPs are proposed to act as redox sensors (Zhang et al., 2002; Fjeld et al., 2003) due to their reported higher affinity for reduced NADH dinucleotide over NAD+, hence linking cellular redox state to the transcriptional output. Also, the disruption of the NAD(H) binding site in CtBPs lead to reduced or abolished dimerization rates (Nardini et al., 2009; Madison et al., 2013) and dimerization in turn is important for the regulation of co-transcriptional activity of CtBP (Madison et al., 2013). A functional role of the AN NADH binding domain is suggested by the finding that a point mutation in the NAD(H) domain results in a complete lack of function phenotype in an-doq allele (Bai et al., 2013). In this study, we found the same cellular phenotype for the stress dependent formation of SGs in an-2 and an-doq allele (Figure 4 and Supplementary Figure S5) and observed hyposensitivity of an-2 and an-doq alleles to salt and osmotic stress (Figure 8). This suggests that the NAD(H) binding domain is also important for AN function in stress responses.

As experimental evidence for a binding of NAD(H) to AN is lacking, we cannot conclude whether the AN^DOQ/AN^{rmGAD→V V A} mutations lead to the predicted reduced or no binding of NAD(H) to AN or whether they disturb the functions of the domain otherwise (e.g., stability, conformation, interaction ability). As the point mutations specifically target the positions predicted to be relevant for NAD(H) binding, it is reasonable to assume that AN activity is regulated by binding to NAD(H). As an-doq mutants display a null phenotype we propose that the NAD(H) bound form is the active form. The findings that overexpression of the AN^DOQ protein can fully rescue the an mutant trichome and leaf phenotypes and that overexpression of the AN^{GAD→V V A} protein can partially rescue the leaf length/width phenotype (Supplementary Figure S6) can be explained by some residual activity of the mutant proteins and that this reduced activity can be compensated by providing excess protein levels.

Our data suggest that the recruitment of AN to SGs does not depend on the NAD(H) domain as YFP-AN^DOQ and YFP-AN^{GAD→V V A} mutant proteins localize to SGs in the absence of wild-type protein (Figure 5). Both mutant proteins are evenly distributed in the cytoplasm (Figure 5). Big dots as observed with wild-type AN were not found, possibly because they represent non-physiological aggregates that do not form because the dimerization is strongly reduced in the mutant AN proteins (Figure 7A). The increased number of SGs in an mutants suggests that binding of the active form of AN counteracts the formation of SGs. It is conceivable that binding of the active form of AN is very transient and that this population of AN molecules cannot be seen in our co-localization experiments. As a consequence we would see only the inactive form of AN at SGs. These considerations would explain that AN^DOQ and AN^{GAD→V V A} show close to 100% co-localization with PAB2 labeled SGs while wild-type AN exhibits partial co-localization (Figure 6 and Supplementary Figure S7). Along the same line, this scenario can explain the higher co-localization levels of wild-type AN with PAB2 in bombardment experiments as compared to transgenic lines as the high levels of AN reached in bombardment experiments may result in a larger population of inactive AN molecules, due to the limited availability of cellular NADH pools. What could be the role of AN in SGs? The moderately increased number of SGs in an mutants indicates that AN can counteract their formation not as a key regulator but rather as a modulator. One attractive possibility is that AN could fine tune SG formation in a redox dependent manner by its ability to monitor the NAD⁺/NADH ratio similar as shown for CtBPs.

A molecular basis for the functionality of NAD(H) binding domain is offered by the differential interaction of AN^DOQ with itself and other proteins. While some interactions are not effected by point mutations in the NAD(H) domain, the dimerization and its interaction with AIK1 is severely reduced (Figure 7). This suggests that the NADH binding domain mediates specific binding of AN to target proteins, thereby regulating their function in RBPs. Biotic and abiotic stresses can lead to an increase in reactive oxygen species (ROS) levels within a cell, leading to redox imbalance and oxidative stress (Hossain et al., 2015) that in turn could be sensed by the NADH binding domain of AN. We therefore speculate that AN function is governed by the redox status of the cell which in turn controls its interaction with specific SG associated proteins and thereby, the function of SGs. This is analogous to the function of Enhancer of decapping 3 (EDC3) and Apoptosis inducing factor (AIF) in mammals, which has been postulated to control mRNP granule formation in a redox dependent manner (Candé et al., 2004; Walters et al., 2014). A structural feature known to mediate aggregation of proteins into RNA granules is the presence of an intrinsically disordered/low complexity region (Kedersha et al., 2013). Using IUPred¹ we found an intrinsically disordered region in AN between amino acid 342 and 487 at the N-terminus. It is therefore possible that AN can associate with SGs through such intrinsically disordered regions.

Is the association of AN with mRNP granules and its role in mRNP granule formation functionally significant? AN is involved in different stress responses as an mutants are more resistant than wild-type to dehydration stress and pathogen attack (Gachomo et al., 2013). More specifically, we observed a hyposensitivity of an mutants as compared to wild-type for two stress treatments triggering a re-localization of AN- salt and osmotic stress (Figure 8). These data suggest that a role of AN in stress responses can be explained by a function as a component of SGs, which are typically formed in response to stress. Examples for a correlation between SG formation and stress response phenotype include the SG components UBP1B and Tudor staphylococcal nuclease (TSN). UBP1B has been shown to be required for SG formation in plants (Weber et al., 2008) and ubp1b mutants are hypersensitive to salt and osmotic stress responses (McCue et al., 2012). Similarly, tsn1tsn2 mutants show reduced number of SGs and are hypersensitive to heat stress (Gutierrez-Beltran et al., 2015). The novel findings hence suggest a role of AN as a post-transcriptional regulator, rather than a transcriptional repressor. An attractive scenario would be a role of AN in balancing growth and metabolism by sensing the cellular redox states.