Here we have investigated noncovalently bound complexes of an 8‐nucleotide RNA and six different ligands, all of which have a guanidinium moiety, by using electrospray ionization (ESI) and collisionally activated dissociation (CAD) mass spectrometry (MS). The order of complex stability correlated almost linearly with the number of ligand atoms that can potentially be involved in hydrogen‐bond or salt‐bridge interactions with the RNA, but not with the proton affinity of the ligands. However, ligand dissociation of the complex ions in CAD was generally accompanied by proton transfer from ligand to RNA, which indicated conversion of salt‐bridge into hydrogen‐bond interactions. The relative stabilities and dissociation pathways of [RNA+ All ligands investigated Table , that is22.1m RNA and 5–100 μm ligand (L) in 1:1 H2O/CH3OH at pH ≈7.5, adjusted by the addition of piperidine and imidazole (≈1 mm each). The additive mixture of piperidine and imidazole was used because it very efficiently suppresses the formation of sodium and potassium adducts without promoting formation of highly charged ions.http://rna.tbi.univie.ac.at/cgi‐bin/RNAWebSuite/RNAfold.cgi),m L−n H]n−, from ESI were observed for all ligands studied, as illustrated for guanidine in Figure n on the RNA–ligand complexes ranged from 2 to 5 , which is consistent with each protonated ligand compensating one of the negative charges of the RNA in the association reaction in solution [Reaction (I)]:Noncovalently bound complexes of the 8–nt RNA  and guanidine or guanidine derivatives Table  were prom; Figure S1), except for Gpa. At this concentration, RNA complex ions with up to 14 Gpa ligands were detected, along with abundant Gpa cluster ions. The latter were not observed for Gnd, meGnd, and tmeGnd, and were found in much lower abundance for aGpa and Arg at a concentration of 100 μm than for Gpa at 10 μm . A possible rationale for the different behavior of Gpa regarding cluster formation and binding to RNA is its net charge, which should be zero assuming that the guanidine and carboxylic acid groups are protonated and deprotonated, respectively, at the solution pH of ≈7.5 used. By contrast, Gnd, meGnd, tmeGnd, aGpa, and Arg should each carry a net charge of +1 at this pH assuming that both the guanidine and amino groups are protonated and the carboxylic acid moieties are deprotonated. In this case, Coulombic repulsion limits cluster formation and binding to RNA for all ligands studied, except for the overall neutral Gpa. Although the RNA complex and ligand cluster ions found in the ESI spectra do not necessarily reflect the species present in solution, these data suggest that the [RNA+m L−n H]n− ions predominantly originate from association reactions in solution.In agreement with previous studies of guanidinium derivatives binding to DNA,2.2m L−n H]n− complex ions with different numbers of ligand (m) and net charge (n) in the mass spectrometer prior to CAD ]:Products from Reaction (IV):m−1) L−(n+1) H]n+1)−(⋅[L+H]+ complexes into a hydrogen‐bond motif in [RNA+(m−1) L−n H]n−⋅L complexes by PT  depends on the PA values of the [RNA+(m−1) L−(n+1) H]n+1)− L−(n+1) H]n+1)−(⋅[L+H]+ and [RNA+(m−1) L−n H]n−⋅L  from protonated ligand to deprotonated RNA had occurred in the gaseous complex ions before ligand dissociation. The energy required for conversion of a salt‐bridge binding motif in n+1)− L−n H]n−⋅L ions, as a result of the far higher electrostatic interaction energies of the former.From the above PA values, PT from protonated ligand to dimethyl phosphate, that is, [L+H]m−1) L−(n+1) H]n+1)−(⋅[L+H]+ ions to produce [RNA+(m−1) L−n H]n−⋅L ions that can further dissociate into [RNA+(m−1) L−n H]n− and L n− ions were from successive losses of neutral ligand [Reaction (II)]. Moreover, no deprotonated ligand, [L−H]−, was detected for tmeGnd, meGnd, or Gnd irrespective of the net charge n and the CAD energy used, which suggests that the PAs of [tmeGnd−H]−, [meGnd−H]−, and [Gnd−H]− far exceed those of the [RNA+(m−1) L−n H]n− ions; a correspondingly high pKa value of 28.5 was reported for Gnd.m−1) L−(n−1) H]n−1)− ions with n=3 and 4. Moreover, time‐of‐flight differences in the transfer of ions with low and high m/z values (≈58 to ≈173 for [L−H]− versus ≈630 to ≈1261 for the corresponding RNA or RNA complex ions) from the collision to the ICR cells complicate quantitative detection of complementary ionic products from Reaction (III). We thus used only the signals of RNA (complex) ions for further data analysis.Consistent with the PAs of ligand anions comprising carboxylates, for example, ≈1385 kJ moln=4, the fraction of [RNA+(m−1) L−(n−1) H]n−1)−( and [RNA+(m−2) L−(n−2) H]n−2)−( ions from Reaction (III)  and (III)) generally increased with increasing energy used for CAD   n=4, theE1). At elevated energy (E2), an intermolecular PT between the guanidinium moiety of Gpa and the deprotonated RNA phosphodiester moiety becomes competitive and more products from Reaction (III) are observed −1),−1 over PT to a deprotonated phosphodiester moiety. Although the PAs of small model compounds generally differ from those of the corresponding sites in [RNA+m L−n H]n− ions, primarily as a result of hydrogen bonding and the presence of multiple charges,As discussed above, Gpa, aGpa, and Arg have zwitterionic structures at the solution pH of ≈7.5 used here, and probably bind to deprotonated RNA by the formation of salt bridges. Based on the energies in Scheme energy E, an inteE3), which makes another proton available for intermolecular PT and Gpa dissociation by Reaction (II). The latter PT reaction was negligible for m=2 to 3, which we attribute to higher energy requirements for structural transitions in the [RNA+2 Gpa−4 H]4− and [RNA+3 Gpa−4 H]4− ions that are stabilized by additional electrostatic interactions. Likewise, CAD of [RNA+Gpa−5 H]5− ions  showed only very few products from Reaction (II) at higher energy −1)E1′≈E1). At elevated energy (E2′≈E2), two protons are transferred to the RNA and [aGpa−H]− dissociates. At energy E3′, which is significantly higher than E3 because extension of the RNA structure requires more energy when the additional amino group also forms a hydrogen bond with the RNA, a proton is transferred from a remote site and neutral aGpa dissociates.The proposed interactions and PT reactions associated with aGpa dissociation from [RNA+aGpa−4 H]m L−4 H]4− ions of Gpa, aGpa, and Arg with m=2 and 3, very few products from Reaction (II) were observed at energies E3 and E3′  L−(n−1) H]n−1)−( and [RNA+(m−2) L−(n−2) H]n−2)−( ions from Reaction (III) at all energies used , which can be rationalized by the smaller number of protons available for Reaction (II).Consistent with a higher stability of [RNA+m L−n H]n− ions of tmeGnd, meGnd, and Gnd at n=2 to 5 and all energies used gave only products from loss of neutral ligand L [Reaction (II)] by PT from [L+H]+ to a deprotonated phosphodiester moiety and subsequent dissociation of the [RNA+(m−1) L−n H]n−⋅L complexes  does not account for the unintended loss of ligand discussed above; for example, in CAD of [RNA+4 Gnd−3 H]3− ions, the yield of free RNA ions, [RNA−3 H]3−, was constantly 1.816 % (±0.005 % standard deviation) at energies of up to 10 eV  at low energy and plateaus at 100 % (inverse 0 %) at high energy, and E50 value at the inflection point of the sigmoid of (31.42±0.05) eV instead of the energy value at 50 % yield of (30.98±0.13) eV from fitting the data with Equation (1). Thus the CAD data were fit by using Equation (2) unless the number of data points in the plateau region was too small to gauge potential offsets.accounts for the vertical offset of the sigmoid, and indicates an 3− ions produced mostly [RNA+1 Gnd−3 H]3− ions at energies of up to 15 eV, but above this energy, [RNA−3 H]3− ions became more abundant, consistent with sequential dissociation of Gnd ligands [Reaction (II)]. To determine the relative energies required for the dissociation of each Gnd ligand, the data were plotted by sequentially adding the yields of all species except that of free RNA. In this representation  and (19.40±0.07) eV), whereas for m=4, the first Gnd ligand dissociated at a far lower energy of (1.75±0.26) eV, and the second and third Gnd ligands dissociated at very similar energies, (22.40±0.04) and (23.80±0.03) eV, respectively. Moreover, there is no apparent systematic trend in the E50 values for the breakdown of [RNA+m Gnd−3 H]3− ions with m=1 to 4, that is, (15.86±0.08), (10.66±0.06), (18.08±0.06), and (1.75±0.26) eV for m=1, 2, 3, and 4, respectively  that was, within error limits, independent of the total number of ligands initially bound to the RNA except for m=1. Similar behavior was observed for all other ligands and complex net charges studied, although the E50 values generally increased in the order tmeGnd<meGnd<Gnd<Gpa<aGpa<Arg ,n=3 . The higher stability of Arg versus aGpa [RNA+m L−n H]n− complexes can be attributed to the longer alkyl chain of Arg that allows it to better adapt to the RNA structureHowever, the m Figure A. This mg Figure A. ImportE50 values are a relative measure of the minimum energy required for complex dissociation, we conclude that the contribution of [RNA−n H]n−⋅L binding energy to complex stability is significantly higher than that of ΔHPT,complex  that in turn decreases the binding energy of [RNA−n H]n−⋅L complexes , the linear‐fit functions in Figure m=0 but showed intercept energies as small as (−4.23±0.09) eV and as large as (26.03±2.82) eV. In other words, one of the m ligands (including that for m=1) can bind to the RNA more strongly than all others, that is, when E50(m=1)>Eslope or, for tmeGnd at n=3 for which E50(m=1)<Eslope, more weakly than all others  °C) are only slightly less stable than hairpin structures with the latter  °C).A possible RNA structure that agrees with all the experimental data from this study is the hairpin motif illustrated in Scheme m=1 can interact with all three phosphodiester moieties of the loop , in which the guanidine moiety of the ligands at p Scheme A, explaip Scheme B. Specif1 Figure C. Finalls Figure A becauses Figure . Any difn=2 to 4, and for tmeGnd, aGpa, and Arg at n=2 to 3  is far more affected by complex net charge than the binding of additional ligands (m>1). This is again consistent with the hairpin structures shown in Scheme n=5, all slope values were significantly smaller than those at n=4, which indicated that Coulombic repulsion limits overall complex ion stability at n=5. Moreover, both the E50(m=1) and Eslope values for aGpa and Arg at n=4 stand out, whereas those at n=2 and 3 follow the general trends discussed above  and Eslope ratio of close to one at n=4 is inconsistent with a unique binding site and instead suggests binding of up to three aGpa or Arg ligands to a largely extended RNA structure.The slopes of the linear fit functions in Figure 2.4E50 analysis are relative rather than absolute.As discussed by Rodgers and ArmentroutE50 values for sequential dissociation of individual ligands, summarized for tmeGnd, meGnd, Gnd, Gpa, aGpa, and Arg at n=2 to 4 and m=1 to 5 in Figure m, on the ΔE50 values  value of (20.09±0.16) eV, dissociation of the second ligand was observed at an additional ΔE50(2) value of (11.37±0.26) eV, and dissociation of the third ligand was observed at an additional ΔE50(3) value of (8.40±0.36) eV. By contrast, the values of ΔE50(1), ΔE50(2), and ΔE50(3) for aGpa at n=3 and m=4 were (20.51±0.23), (6.40±0.46), and (10.76±0.85) eV, respectively  value at m=4 (6.40 eV) was only 56 % of the ΔE50(2) value at m=3 (11.37 eV), whereas the ΔE50(3) value at m=4 (10.76 eV) was larger by 28 % than the ΔE50(3) value at m=3 (8.40 eV).However, the E50 were found for Gnd at m≥3  to 46.5 (tmeGnd), than those for the dissociation of all ligands . This indicates that PT to the RNA  and ligand (5–100 μm) in 1:1 CH3OH/H2O at pH ≈7.5 adjusted by the addition of ≈1 mm piperidine and imidazole each. Methanol was HPLC grade , H2O was purified to 18 MΩ cm−1 at RT by using a Milli‐Q system , and all ligand compounds  were purchased from Sigma–Aldrich . The 8‐nt RNA (5′‐GGCUAGCC‐3′ with HO termini) was prepared by solid‐phase synthesis using 2′‐O‐[(triisopropylsilyl)oxy]methyl (TOM) chemistry, purified by HPLC, and desalted by using Sep‐Pak C18 cartridges, washed with triethylammonium bicarbonate (0.1–0.15 m) in H2O, and eluted with 1:1 CH3CN/H2O.Eslope) or nonlinear (E50) least‐squares fitting procedures.Experiments were performed on a 7 T Fourier transform ion cyclotron resonance (FT‐ICR) mass spectrometer  equipped with an ESI source for [M−The authors declare no conflict of interest.As a service to our authors and readers, this journal provides supporting information supplied by the authors. Such materials are peer reviewed and may be re‐organized for online delivery, but are not copy‐edited or typeset. Technical support issues arising from supporting information (other than missing files) should be addressed to the authors.SupplementaryClick here for additional data file.


SAMHD1 cause AGS; however, how SAMHD1-deficiency causes the type I interferon response in patients with AGS remains unknown. Here, we show that endogenous RNA substrates accumulated in the absence of SAMHD1 act as a major immunogenic source for the type I interferon response. Reconstitution of SAMHD1-negative human cells with wild-type but not RNase-defective SAMHD1 abolishes spontaneous type I interferon induction. We further identify that the PI3K/AKT/IRF3 signaling pathway is essential for the type I interferon response in SAMHD1-deficient human monocytic cells. Treatment of PI3K or AKT inhibitors dramatically reduces the type I interferon signatures in SAMHD1-deficient cells. Moreover, SAMHD1/AKT1 double knockout relieves the type I interferon signatures to the levels observed for wild-type cells. Identification of AGS-related RNA sensing pathway provides critical insights into the molecular pathogenesis of the type I interferonopathies such as AGS and overlapping autoimmune disorders.The autoimmune disorder Aicardi-Goutières syndrome (AGS) is characterized by a constitutive type I interferon response. SAMHD1 possesses both dNTPase and RNase activities and mutations in  While SLE is associated with more than 20 genes, AGS is caused by autosomal recessive mutations in one of several genes encoding enzymes involved in nucleic acid metabolism  or by gain-of-function mutations in the cytosolic RNA sensor IFIH17. Some children with AGS also display an early onset form of SLE. Given that the pathology of SLE is complex and heterogeneous, AGS could be an excellent model disease to study systemic autoimmunity and provide a clue to the pathogenesis of SLE.Aicardi-Goutières syndrome (AGS) is a monogenic autoinflammatory disorder that overlaps phenotypically with congenital viral infection and systemic lupus erythematosus (SLE), and is characterized by constitutive upregulation of type I interferon (IFN) in the serum and cerebrospinal fluidSAMHD1 was identified initially as the human ortholog of the mouse IFNγ-induced gene Mg118. Although SAMHD1 has been proposed to serve as a negative regulator of the innate immune response4, studies have focused overwhelmingly on the inhibitory mechanism of SAMHD1 against HIV-1 replication because of an unexpected identification of SAMHD1 as an HIV-1 restriction factor10. In mouse models, mouse SAMHD1 also seems to restrict the replication of certain retroviruses; however, Samhd1 knockout mice did not display AGS-like symptoms12. SAMHD1 possesses dual enzymatic activities: deoxynucleoside triphosphohydrolase (dNTPase) and phosphorolytic 3′-5′ exoribonuclease16. The physiological function of SAMHD1 under natural conditions remains poorly understood. In particular, the mechanism by which the mutations in SAMHD1 cause AGS needs to be determined. Recently, SAMHD1 was implicated in the DNA damage response and in preventing autoimmunity by maintaining genome integrity18. Considering that all of the AGS-related genes associated with nucleic acid metabolism and nucleic acid sensing dysfunction implicated in autoimmunity1, the elevated IFN signature observed in SAMHD1-related AGS patients might be caused by activation of the innate immune response against dysregulated endogenous nucleic acids. In this study, we explored the role of SAMHD1 in regulating nucleic acid-mediated type I IFN signaling to understand the molecular pathogenesis of AGS and overlapping autoimmune disorders.Samhd1-deficient mice displayed neither significant upregulation of IFN-stimulated genes (ISGs) in their sera nor other systemic autoimmune phenotypes, although Samhd1−/− mouse cells exhibited spontaneous induction of ISGs12. Therefore, we used human THP-1 cells, a well-established model of human monocytes, to study the molecular pathology of the SAMHD1-related AGS. We analyzed transcriptional signature of AGS using two independent SAMHD1-deficient THP-1 cells engineered by different clustered regularly interspaced short palindromic repeats (CRISPR) single guide RNAs (sgRNAs). Type I interferons including various subtypes of IFN-α and IFN-β, and ISGs  were upregulated in SAMHD1-deficient cells  was similar between SAMHD1-deficient and wild-type cells   Fig. . To invells Fig. . Intereslls Fig. , suggestSAMHD1-deficient cells showed distinct mRNA expression patterns among more than 1200 genes   to ascertain the IFN signatures of AGS. nes Fig. . Pathwaynes Fig. . Upstreanes Fig. . In partlls Fig. , which cCR) Fig. . These rSAMHD1-deficient cells activates IFN responses. We isolated both DNA and RNA from wild-type and SAMHD1-deficient cells and examined their abilities to stimulate the IFN response in phorbol 12-myristate 13-acetate (PMA)-differentiated THP-1 cells. Interestingly, only RNA derived from SAMHD1-deficient cells, but not that from wild-type cells, highly activated IFN-α expression. On the other hand, DNA isolated from wild-type and SAMHD1-deficient cells had comparable abilities to activate IFN-α expression  or melanoma differentiation associated gene 5 (MDA5)-dependent pathways could mask the IFN-β induction by RNA substrates of SAMHD1. To investigate the detailed features of IFN-stimulatory RNA species, we isolated small (<200 nt) and large (>200 nt) RNAs in two separate fractions from each of the wild-type and SAMHD1-deficient THP-1 cell lines and then stimulated PMA-differentiated THP-1 cells with these RNAs. The large RNAs (>200 nt) from SAMHD1-deficient cells activated IFN-α mRNA expression significantly  from wild-type cells  analysis with SAMHD1. Immunoprecipitates of SAMHD1 protein and SAMHD1-RNA complex were successfully detected by western blot and autoradiography Fig. . A totalSAMHD1-deficient cells. RIG-I-like receptors (RLRs), which include RIG-I and MDA5, and intracellular Toll-like receptors (TLRs) are well known cytosolic RNA sensors26. We assumed that one of these RNA sensors might be linked to the IFN signature in SAMHD1-deficient cells; therefore, depleted each of these sensors or downstream molecules independently using short interfering RNA (siRNA) treatment  were reduced to basal levels in cells treated with anti-IFNAR1, but not in those treated with the anti-IL10Rβ antibody. Co-treatment with both IFNAR1 and IL10Rβ antibodies did not show synergistic inhibitory effects on ISGs induction , and activation of the Janus kinase/signal transducers and activators of transcription (JAK/STAT) pathway is indispensable for ISGs induction27. JAK inhibitor treatment diminished ISG induction thoroughly  system. Phosphorylation of STAT1 and IRF3 induced by SAMHD1 knockout was markedly diminished upon SAMHD1/AKT1 double knockout  pathways, and the phosphoinositide 3-kinase (PI3K)/AKT pathwayway Fig. . Indeed,les Fig. . IRF3 waion Fig. . HoweverAT1 Fig. . Treatmeure Fig. . Consisture Fig. . To confout Fig. . Type I lls Fig. ; therefolls Fig. . Neutrallls Fig. . siRNA-mlls Fig. , suggestSAMHD1 knockout was antagonized by ectopic expression of wild-type SAMHD1 or RNase+/dNTPase− SAMHD1D137N. Expression of the RNase-defective mutants, SAMHD1D207N and SAMHD1Q548A, had little effect on AKT phosphorylation , are bound to SAMHD1 protein. snoRNAs are known to guide the modification (2′-O-methylation and pseudouridylation) of RNAs represented by rRNAs40 and the incorporation of 2′-O-methylated nucleosides or pseudouridine into RNA diminishes immunostimulatory activity42. Therefore, SAMHD1 would indirectly affect the immunostimulatory activity of endogenous RNAs by regulating RNA modification, accounting for a partial role for entire type I IFN response in SAMHD1-deficient cells. Considering the phenotypical overlap of AGS with SLE, the pathology of SLE patients, in some cases, might be associated with the accumulation of aberrantly metabolized endogenous nucleic acids. A greater understanding of the pathogenesis of type I interferonopathies is not only of scientific interest but might lead to new therapies for autoinflammatory diseases.Notably, SAMHD1 specifically targets retroviral RNA for degradationSAMHD1 were amplified by PCR from previously described plasmids15 and inserted into the EGFP-N3 vector. SAMHD1-deficient THP-1, HEK239T, and HeLa cells were generated using the CRISPR/Cas9 system. The SAMHD1/AKT double knockout THP-1 cell line was established by transfecting SAMHD1-deficient THP-1 cells with single guide RNAs (sgRNAs) specific for AKT1. Immunoblotting and genomic DNA sequencing confirmed all the knockout cell lines. Guide RNA sequences of SAMHD1 and AKT1 are listed in Supplementary Table SAMHD1-specific siRNA or control non-specific siRNA for two cycles to obtain efficient knockdown and incubated in RPMI media. After 48 h of incubation, PBMCs were analyzed by western blotting analysis to monitor the activation of STAT1 and AKT.Wild-type and mutants of The chemical reagents and antibodies used in this study were purchased from the following manufacturers: poly (I:C) and poly (dA:dT), Sigma; T4 Polynucleotide Kinase and T4 RNA ligase, Takara; T4 RNA Ligase 2 (truncated K227Q) and Antarctic Phosphatase, New England Biolabs (NEB); wortmannin and rapamycin, Sigma; MK-2206, A-674563, and Lapatinib, Selleckchem; Ruxolitinib, Invivogen; mouse monoclonal antibodies to IFNAR1 (Millipore) and SAMHD1 (OriGene); rabbit monoclonal antibodies to IRF3 pho-S386 (Abcam), RIG-I, STING, MyD88, TBK1, TBK1 pho-S172, Stat1 pho-Y701, and GSK-3β ; rabbit polyclonal antibodies to IRF3 (Santa Cruz), SAMHD1 , GAPDH (Ab Frontier), MAVS, TRIF, IRF7, STAT1, Akt, Akt pho-S473, and GSK-3β pho-S9 ; goat polyclonal antibody to hIL-10Rb (R&D systems).All siRNAs were purchased from Dharmacon as ON-Target plus. THP-1 cells were transfected by electroporation with indicated gene-specific siRNAs or non-specific siRNAs using Neon (Invitrogen). Lysates were collected 72 h after transfection for western blotting and qRT-PCR analysis. For nucleic acid stimulation, PMA-differentiated THP-1 cells were transfected with 5 μg/ml of poly (I:C), poly (dA:dT), or isolated DNA or RNA, using Lipofectamine 2000 transfection reagent (Invitrogen) and undifferentiated THP-1 cells were transfected by electroporation with 10 μg/ml of isolated RNA using Neon. Cells were harvested 4 h post transfection, followed by RNA isolation. For reconstitution of SAMHD1-deficient THP-1, THP-1 cells were transfected by electroporation with expression vectors for wild-type or mutant SAMHD1 using Neon. Isolated PBMCs were transfected using Interferin™ transfection reagent (Polyplus-transfection Inc.), according to the manufacturer’s instructions.Genomic DNA was purified using a Blood Mini Kit (Qiagen). Total RNA was isolated using the TRIzol reagent (Invitrogen). Fractionation and extraction of small and large RNAs were performed using the NucleoSpin miRNA system (Macherey-Nagel). Separation and purification of cytoplasmic and nuclear RNAs were carried out using a Cytoplasmic & Nuclear RNA purification kit (Norgen). All the purification steps were processed according to the manufacturers’ instructions.g at 4 °C. anti-SAMHD1 and anti-rabbit IgG antibodies were incubated with Dynabeads for 1 h at room temperature. The conjugated beads were washed sequentially twice each with buffer A  and buffer B . Cell lysates were incubated with the prepared conjugated Dynabeads for 1 h at 4 °C. The immunoprecipitates were washed five times with buffer B, followed by an in vitro nuclease assay. The in vitro nuclease assay was performed as described previously15. In brief, immunoprecipitated proteins were incubated in 20 µl reaction mixtures containing phosphate-buffered saline (PBS) supplemented with 5 mM MgCl2, 2 mM dithiothreitol (DTT) and [γ−32P]-labeled RNA substrate (A20) at 37 °C for 30 min. The reactions were stopped by the addition of 20 µl of 2 × RNA loading buffer and then boiled at 95 °C for 5 min. RNA was separated on 8 M urea/15% polyacrylamide gels and analyzed using a BAS-2500 phosphorimager (Fujifilm).Pelleted THP-1 cells were lysed for 30 min at 4 °C in lysis buffer . After sonication, the lysates were centrifuged for 30 min at 13,000 × 43. Briefly, THP-1 cells were irradiated by 254 nm UV at total 600 mJ/cm2 for RNA-protein cross-liking (Spectroline) for SAMHD1-CLIP. The RNAs bound to SAMHD1 were immunoprecipitated with anti-SAMHD1 conjugated beads. After ligation with 3′ adaptors  using T4 RNA Ligase 2, truncated K227Q (NEB), RNAs were labeled with [γ-32P]ATP by T4 polynucleotide kinase (Takara). The RNA-protein complex was separated by SDS-PAGE electrophoresis, then transferred onto nitrocellulose membrane (Whatman). The membrane with RNAs was cut by a razor and RNAs were extracted by phenol/chloroform (Ambion), followed by ethanol precipitation for RNA isolation. 5′ adaptors  were ligated using T4 RNA ligase (Takara). For RNA-seq, total RNA was extracted from wild-type and SAMHD1-deficient THP-1 cells using the TRIzol reagent (Invitrogen), according to the manufacturer’s instructions. Ribosomal RNA was removed from the total RNA using a Ribo-Zero rRNA removal Kit (Epicentre). After RNA fragmentation by RNA fragmentation reagents (Ambion), RNAs of 30–60 nucleotides were purified and ligated with 3′ and 5′ adaptors using T4 RNA Ligase 2, truncated K227Q, and T4 RNA ligase, respectively. 5′ and 3′ adaptor ligated RNAs were reverse transcribed by using the RNA RT primer . PCR was performed to generate libraries for high throughput sequencing with the 5′ end Illumina RNA PCR Primer (RP1) and the 3′ end Illumina RNA PCR Primer with index sequences (Index 1–9). Sequencing was performed on a HiSeq. 2000/2500 (Illumina).CLIP-seq and RNA-seq was performed as previously described, with some modificationshttp://hannonlab.cshl.edu/fastx_toolkit/) version 0.0.13.2. We also discarded reads mapped to human rRNA or tRNA by Bowtie2 version 2.2.7 with -t - k 2–very-sensitive options. Tophat version 2.1.1 with–no-coverage-search–b2-very-sensitive  options was used to align these preprocessed sequencing reads against the human reference genome (GRCh37.p13) downloaded from the Reference Sequence (RefSeq) collection. In this alignment process, we also considered repetitive elements defined by the output of RepeatMasker that was downloadable from RefSeq. Based on these alignments, we calculated the read counts for each genomic position by using in- house software that uniformly divided each multi-mapped read to all of the positions it maps to. As previously described43, we applied Fisher’s exact test to detect significant peaks (CLIP-seq enriched regions over RNA-seq). One modification in this study was that we calculated p-values for every genomic position in the whole genome background. Then, the p-values were adjusted by qvalue package in R.For preprocessing, we removed adapter sequences and low-quality ends from CLIP-seq and RNA- seq reads by using Cutadapt version 1.10 with -m 17–match-read-wildcards -O 10 -e 0.1 -q 30,30 -g AATGATACGGCGACCACCGAGATCTACACGTTCAGAGTTCTACAGTCCGACGATC –a TGGAATTCTCGGGTGCCAAGGAACTCCAGTCAC options. Artifact reads were eliminated by the fastx_artifacts_filter command in FASTX-Toolkit  downloaded from the Reference Sequence (RefSeq) collection and estimate gene-level transcript abundances. DEGs were assessed by limma package in R45. We first filtered genes with total read counts less than 12 for 6 samples . We applied trimmed mean of M values (TMM) normalization to read counts for the estimation of scale factors among samples. Then, the voom transformation was applied to the filtered and normalized counts. After this, the usual limma procedure for differential expression analysis was followed. That is, we estimated the fold changes and standard errors by fitting a linear model for each gene and applied empirical Bayes smoothing to the standard errors. Finally, moderated t-statistics and corresponding p-values were computed. Benjamini-Hochberg method was used to adjust p-values for multiple testing.RSEMHuman blood samples were anonymously provided by the Blood Center of the Korean Red Cross, Seoul, under the approval of the Institutional Review Board of Korean Red Cross with consent for research use. Experiments involving human blood were approved by the Institutional Review Board at Seoul National University (SNUIRB No. E1512/001–004). All experiments were performed in accordance with Seoul National University guidelines.t-tests .Statistical analyses were carried out using GraphPad Prism version 5.0 software. Statistical significance was determined by two-tailed Student’s The datasets generated during or analyzed during the current study are available from the corresponding author on reasonable request.Supplementary Information


In eukaryotes, nascent RNA transcripts undergo an intricate series of RNA processing steps to achieve mRNA maturation. RNA editing and alternative splicing are two major RNA processing steps that can introduce significant modifications to the final gene products. By tackling these processes in isolation, recent studies have enabled substantial progress in understanding their global RNA targets and regulatory pathways. However, the interplay between individual steps of RNA processing, an essential aspect of gene regulation, remains poorly understood. By sequencing the RNA of different subcellular fractions, we examined the timing of adenosine-to-inosine (A-to-I) RNA editing and its impact on alternative splicing. We observed that >95% A-to-I RNA editing events occurred in the chromatin-associated RNA prior to polyadenylation. We report about 500 editing sites in the 3′ acceptor sequences that can alter splicing of the associated exons. These exons are highly conserved during evolution and reside in genes with important cellular function. Furthermore, we identified a second class of exons whose splicing is likely modulated by RNA secondary structures that are recognized by the RNA editing machinery. The genome-wide analyses, supported by experimental validations, revealed remarkable interplay between RNA editing and splicing and expanded the repertoire of functional RNA editing sites. The maturation of eukaryotic mRNAs involves an intricate series of molecular processes that modify newly synthesized RNA molecules, such as 5′ capping, splicing, RNA editing, and polyadenylation . These RAmong the various RNA processing steps, RNA editing and RNA splicing share the largest overlap in time and space during mRNA maturation . CatalyzAlu repeats enabled exonization of the noncoding sequences , suggesting that RNA editing may be an important means for exon evolution  that are required to form dsRNA substrates for editing . Thus, RDrosophila  splicing alteration by RNA editing in splice site sequences; and (2) splicing modulation by ADAR binding and editing of dsRNA structures. Supported by experimental validations, our study expands the repertoire of RNA editing sites that influence alternative splicing and provides new insights regarding the interplay between these two processes.Supplemental Fig. S1). We obtained RNA-seq data in triplicates, with two types of RNA (polyadenylated and nonpolyadenylated) in the Ch fraction and polyadenylated RNA in the Np and Cp fractions  . Read mapping was then carried out using our previously published method, which has superior performance in handling single-nucleotide variants in the reads . This result is consistent with the expectation that ChA− RNA mainly consists of nascent RNA molecules that are not yet fully spliced. The percentage of intronic reads in ChA+ was much larger than those in later stages, such as NpA+ and CpA+, indicating incomplete splicing in the ChA+ fraction, which is consistent with previous literature  and were thus combined for the RNA editing analysis described below.We observed that the ChAterature . In contSupplemental Table S2). It should be noted that ChA− had the most non-A-to-G or non-C-to-T types, which may reflect existence of unknown biology . Hereafter, we refer to this collection of sites as “editing sites” in general. It is important to note that these editing sites could be edited, unedited, or undetermined (without adequate read coverage) in a specific fraction . We further classified the editing sites into four groups based on their editing status in each fraction  were categorized into one of the four groups . The sites that could not be unambiguously classified into these groups were excluded , which could be due to two main reasons. The first reason is inadequate read coverage in at least one of the cell fractions. In general, if an editing site was “undetermined” due to low read coverage in any fractions prior to the fraction where it was observed with edited reads, this editing site cannot be categorized unambiguously. A total of 170,268 such editing sites were identified and excluded from the grouping. Second, an editing site was excluded if it was “unedited”  in a later fraction but “edited” in an earlier fraction. A total of 37,577 such sites were identified. These editing sites may be associated with complicated regulatory or functional mechanisms, which should be investigated in future studies.To obtain a detailed view of editing events across subcellular fractions, we enumerated all possible scenarios of an editing site being “edited,” “unedited,” or “undetermined” in each fraction (−). This observation suggests that most editing events occur cotranscriptionally in nascent RNA prior to polyadenylation. Since group 4 (CpA+-specific) consists of a very small number of editing sites , we excluded this group from further analyses.Among all categorized editing sites, more than 226,000 (97%) were included in group 1 (ChA− (NA−), nuclear poly(A)+ (NA+), and cytosolic poly(A)+ (CA+) data were obtained from human cell lines (HepG2 and K562) . Furthermore, we analyzed additional RNA-seq data that captured nascent RNA using 4sU labeling, a method that is completely different from cell fractionation . Together, our analyses showed widespread cotranscriptional RNA editing in human and mouse cells, which is consistent with the previous finding in Drosophila nd K562)  and anotnd K562) . These aionation . These dosophila . Further+ and CpA+) were generally higher than those from fractions of more nascent RNAs (ChA− and ChA+). To exclude the possibility that the observed editing change is due to loss of low-level editing sites in introns in later fractions, we carried out the same analysis but excluding intronic editing sites and observed the same pattern of higher editing ratios in the later fractions . Indeed, a “monotonicity Z-score” (MZ score) . Among the known recoding sites in the literature . All of these recoding sites had positive MZ scores.Next, we carried out a comparative analysis of A-to-I editing in different subcellular fractions. For this purpose, we included editing sites with a total read coverage of at least five in all fractions. The levels of RNA editing differ greatly for different editing sites C. In eacZ score)  calculatZ score) E. Consisr groups F. The interature , five weSupplemental Fig. S9). Thus, the “earlier” editing sites, those that are edited early in nascent RNA, tend to occur in longer introns.For all kinetic groups, the majority of editing sites were located in the intronic regions A. The leAlu regions .All kinetic groups were mostly composed of editing sites located in  regions C. Notablng sites D, consis regions C. This o regions E. EvolutADAR1 knockdown (KD) in U87MG cells  consist of ADAR-regulated editing events. Yet, the differences among these groups indicate that additional regulatory mechanisms may exist to impact RNA editing at different stages of RNA maturation, a topic of future investigation.+ fraction and ≥0.95 in the NpA+ fraction, and coTS exons as those with coSI ≥ 0.95 in the ChA− fraction. In total, we identified 6104 coTS exons and 3304 postTS exons. We then compared the splicing difference between these two groups of exons upon ADAR1 KD in U87MG cells. The results showed that the change in exon inclusion in postTS exons was larger than that in coTS exons , suggesting that RNA editing imposes a larger impact on exons undergoing post-transcriptional splicing.Because our data suggested that most editing sites occurred soon after the RNA was transcribed and likely before some introns were spliced out, we hypothesized that RNA editing affects post-transcriptionally spliced exons more than cotranscriptionally spliced ones. Note that this hypothesis does not preclude the possibility that editing may affect cotranscriptionally spliced exons, depending on the relative kinetics of these two processes on specific exons. To test this hypothesis, we adopted the “completed splicing index” (coSI) metric that quantifies the splicing completeness around an internal exon . We calc− fraction that were located in the splice site consensus. Specifically, a 9-mer sequence was defined as the 5′ splice site (5′ss) with 3 nt in the exon and 6 nt in the intron. A 23-mer sequence was defined as the 3′ss with 3 nt in the exon and 20 nt in the intron, based on previous literature  had PSI changes in the same direction as expected based on the alteration in the 3′ss sequence by the A-to-G change , resulting in a total of 498 such unique editing sites if combining data from the three cell lines. To verify their impact on splicing, we analyzed the splicing patterns of the associated exons as described above, using the ENCODE ADAR1 KD RNA-seq of the respective cell lines . Altogether, our analyses of multiple data sets led to identification of an unexpectedly large number of editing sites located in splice site regions that can potentially alter splicing.As an independent analysis, we searched for splice site–altering editing sites in the NAll lines . Among tAlu repeats. About 25% of these exons overlapped Alu sequences, a much higher percentage than what was observed for control exons that do not have known editing sites in the 3′ss . This alteration in 3′ss is opposite to the previously reported example, in which A-to-I editing creates a 3′ss (changing AA to AG) and enables Alu exonization  in Alu-containing exons as a proxy for sequence conservation. We observed that Alu-containing exons with 3′ss editing had significantly lower SNP density than control exons  . The Fay and Wu's H values . Thus, it is likely that edited Alu-containing exons are under selection to preserve their sequences more than random controls.The aforementioned nization . Thus, iH values  were notAlu-containing exons with 3′ss editing sites across 46 vertebrates spanning primates to fish . Together, both Alu-containing and non-Alu exons with 3′ss editing showed evidence of higher conservation than their corresponding controls.Next, we examined the sequence conservation of non- to fish . Since t to fish B and bec to fish , the 3′sSupplemental Methods), the genes that contain 3′ss-edited exons (Alu or non-Alu) were enriched in functional categories related to protein trafficking, cellular metabolism (protein or energy-related), RNA processing, or cytoskeletal structures . Thus, consistent with the highly conserved nature of the exons, these genes are potentially implicated in critical cellular functions.Based on Gene Ontology (GO) analysis . Note that the endogenous editing levels are generally low, and these five candidates had an average editing level of 0.36. Thus, the endogenous splicing changes of these exons upon ADAR1 KD are not expected to be high. Nevertheless, all five exons demonstrated splicing changes in the expected direction  (FBXL4 and DENND4A) were alternatively skipped exons (SEs), with increased exon inclusion levels in ADAR1 KD cells. This observation is consistent with the expectation that the edited version of the 3′ss (GG) weakens the splicing signal and causes exon skipping. The other three exons each had an alternative 3′ss (A3SS). In these cases, given the editing event, the original 3′ss was partially or completely abolished. Instead, an alternative 3′ss in the downstream  or upstream (RAP1GDS1) region was used, which created a shorter or longer form of the original exon, respectively.To experimentally confirm the impact of RNA editing on splicing, we randomly picked five 3′ss-edited exons and tested whether their splicing patterns were altered upon ficance) B. Two exSupplemental Table S6). We created two versions of the constructs for each target, one harboring the unedited (A) and the other for the edited (G) alleles in the 3′ss sequence. For the SE events , the exons had significantly reduced inclusion levels in the G version of the construct  or upstream  region was used in the presence of the G allele. Taken together, our experimental results support that RNA editing could alter splicing by modifying the splice site sequences.As a complementary experiment, we performed minigene reporter assays on the aforementioned five exons and two additional examples, all of which had 3′ss editing  passed a stringent criterion for dsRNA formation  that showed a PSI change (in either direction) of at least 10% upon seq data . The splseq data   of the 33 exons had increased PSI values upon ADAR1 KD . Endogenous splicing inclusion levels of these exons were estimated via RT-PCR followed by gel electrophoresis. Three exons demonstrated a trend of higher splicing levels upon ADAR1 KD, consistent with the predictions via RNA-seq (C17orf67) failed this experiment due to existence of multiple PCR amplification products indicating likely unspecific primers or complex splicing patterns .To provide experimental validation of the ADAR1-dependent splicing changes, we tested the splicing patterns of four exons in U87MG cells of  RNA-seq C. Note tThese results suggest that ADAR1 may bind to dsRNA structures and repress splicing of the neighboring exons. Since we have shown that RNA editing occurs early in the nascent pre-mRNA, and it is known that splicing is also cotranscriptional , this obcis-regulatory sequences or by changing the stability of dsRNA structures, which may affect the interaction of splicing machineries with RNA. In the second category, the editing machinery (ADAR proteins), instead of editing sites, influences splicing either by competitively binding to dsRNA regions and precluding access of splicing machineries to the RNA or by recruiting trans-acting factors that regulate splicing.RNA editing and splicing are both RNA processing steps that may significantly diversify gene expression. An increasing number of studies have examined the intricate crosstalk between these processes . In geneThe impact of RNA editing on splicing has been demonstrated in a number of example genes, although the underlying mechanisms sometimes remain unknown . Despite−) RNA-seq is essential to this study. In identifying RNA editing sites using standard polyadenylated RNA-seq data, it is a general practice to exclude RNA–DNA mismatches located in the splice site regions from the list of predicted editing sites , reflecting incomplete splicing  on ice for 4 min after homogenization by flicking. The sample was centrifuged for 10 min at 4°C, 15,000g, after loading the lysate onto 24% sucrose cushion (24% RNase-free sucrose in plasma membrane lysis buffer) using large orifice tips. The supernatant was extracted as the cytoplasmic fraction. After washing with 1× PBS/1 mM EDTA, the pellet was resuspended in 100 µL of prechilled glycerol buffer  by gentle flicking of the tube. An equal volume (100 µL) of nuclei lysis buffer  was added. The sample was then vortexed vigorously (highest setting on vortex) for 2 × 2 seconds. After incubation for 10 min on ice, the nucleoplasm and chromatin fraction were then separated by centrifugation at 4°C, 15,000g, for 2 min. Fractionation efficiency was validated by Western blotting using antibodies specific to marker proteins for each fraction: β-tubulin  for cytoplasm, U1-70k (a kind gift from Dr. Douglas Black) for nucleoplasm, and Histone 3  for chromatin.U87MG cells were fractionated as previously described  with som25 Dynabeads  following the instructions provided by manufacturer. The flow-through was collected, and the RNA was extracted using TRIzol LS as nonpolyadenylated RNA. Three biological replicates (independent cell culture) were obtained for each fraction.Total RNA was extracted from each fractionated sample using TRIzol LS , and ribosomal RNA was depleted using the RiboMinus Transcriptome Isolation Kit  according to the manufacturer's instructions. Polyadenylated RNA was then selected by Oligo (dT)To prepare for RNA-seq libraries, we started with 50 ng RNA extracted from each sample. Paired-end strand-specific RNA-seq was performed with the NEBNext Ultra Directional RNA Library Prep Kit  with the following adjustments to the manufacturer's instructions: Fragmentation was performed for 12 min at 94°C, and PCR cycling was set to 12 cycles. Unique index was ligated to each sample RNA library construct using the NEBNext Multiplex Oligos for Illumina Index Primers Sets 1 and 2 .Supplemental Material). Reads mapped in the hyperediting alignment step were pooled with those uniquely mapped in the first round of alignment. Triplicates of the same experiment were combined for RNA editing detection to increase read coverage. Mismatches identified in the final mapped reads were examined. Final RNA editing sites were identified after removing likely false positives due to sequencing errors, inaccurate mapping to repetitive sequences or spliced junctions, or technical biases (such as strand bias of reads) (− fraction but were excluded from the list of editing sites identified in the other fractions.The RNA-seq data sets were mapped to the hg19 genome and Ensembl (Release 75) transcriptome  using a f reads) . In addi−, ChA+, NpA+, and CpA+ samples to represent the approximate kinetic order of RNA maturation. RNA editing sites identified from all fractions were pooled together to obtain the overall set of editing sites considered in this study. For each editing site, we then examined whether it was edited, unedited, or undetermined in each fraction as follows: (1) A site was deemed “edited” in one fraction if it had ≥1 read with the edited G nucleotide, regardless of the total number of reads; (2) a site was deemed “unedited” in one fraction if its total read coverage was at least k (see below), but no read contained the edited G; or (3) a site was deemed “undetermined” if its read coverage was inadequate (<k), and no read contained the edited G. Given these three possibilities, an editing site may fall into one of 65 possible categories, considering its status in the four fractions . In this study, we focused on the categories that have unambiguous interpretation regarding the kinetic timing of an editing site's occurrence and further organized them into four groups  editing sites in this study is relatively conservative.To unambiguously determine whether an editing site was “unedited” in a specific fraction, the parameter ed reads . One excractions A. For th− (NA−), nuclear poly(A)+ (NA+), and cytoplasmic poly(A)+ (CA+) RNA CA+) RNA . Using tP > 0.05). Intermediate group consisted of editing sites with ER > 0 in both 4sU-RNA and Total-180 min, but significant increase of ER was observed in Total-180 min relative to 4sU-RNA . Late editing refers to those editing sites with ER = 0 in 4sU-RNA, and ER > 0 in Total-180 min RNA.The editing kinetic groups of A-to-I editing sites in mouse dendritic cells were derived from published time-course 4sU-labeled RNA-seq data sets . 4sU wasADAR1 KD and control samples. This method examines spliced junction reads and read coverage in putative exonic and intronic regions to call exons. These putative exons were compared with those in the reference annotation to identify novel exons. We required a novel exon to have a read density of ≥10  and ≥2 exon junction reads supporting each exon boundary. The PSI values of annotated and novel exons were calculated following a previously developed protocol  based on the RNA-seq data of three human cell lines . We focused on the AS exons with exonic or intronic editing sites within 500 nt. AS exons without editing sites in these regions were used as controls. To identify candidate dsRNA structures encompassing the AS exons, we ran BLASTN  were amplified using genomic DNA from HEK293 cells. Primers used in this study are listed in us study .https://www.ncbi.nlm.nih.gov/geo/) under accession number GSE105773. Custom scripts are available in Supplemental_Code.zip.RNA-seq data derived from subcellular fractions of the U87MG cells from this study have been submitted to the NCBI Gene Expression Omnibus (GEO;


Conserved among animal mRNAs, IRE-RNA structures are noncoding and bind Fe2+ to regulate biosynthesis rates of the encoded, iron homeostatic proteins. IRP1 protein binds IRE-RNA, inhibiting mRNA activity; Fe2+ decreases IRE-mRNA/IRP1 binding, increasing encoded protein synthesis. Here, we observed heat, 5 °C to 30 °C, increased IRP1 binding to IRE-RNA 4-fold (FRT IRE-RNA) or 3-fold (ACO2 IRE-RNA), which was enthalpy driven and entropy favorable. Mn2+  increased IRE-RNA/IRP1 binding (Kd) 12-fold (FRT IRE-RNA) or 6-fold (ACO2 IRE-RNA); enthalpic contributions decreased ~61% (FRT) or ~32% (ACO2), and entropic contributions increased ~39% (FRT) or ~68% (ACO2). IRE-RNA/IRP1 binding changed activation energies: FRT IRE-RNA 47.0 ± 2.5 kJ/mol, ACO2 IRE-RNA 35.0 ± 2.0 kJ/mol. Mn2+ (50 µM) decreased the activation energy of RNA-IRP1 binding for both IRE-RNAs. The observations suggest decreased RNA hydrogen bonding and changed RNA conformation upon IRP1 binding and illustrate how small, conserved, sequence differences among IRE-mRNAs selectively influence thermodynamic and kinetic selectivity of the protein/RNA interactions.Comparison of kinetic and thermodynamic properties of IRP1 (iron regulatory protein1) binding to FRT (ferritin) and ACO2 (aconitase2) IRE-RNAs, with or without Mn IRPs are ubiquitously expressed, with IRP1 expression dominant in liver, kidney and brown fat, while IRP2 expression is dominant in the central nervous system6. IRP1 and IRP2 share significant sequence homology8 and bind the same family of IRE-mRNAs, but are regulated by iron through different mechanisms. IRP2 also has an additional cysteine rich 73 amino acid domain, the function of this domain is yet unknown10. IRP1 is a bifunctional protein, having activity as the aforementioned IRE-RNA binding protein in its apo-form, and as cytosolic aconitase upon assembly of a [4Fe-4S] cluster3. Interconversion between these mutually exclusive activities, favoring the aconitase when iron is replete and the apo IRE-RNA binding form in low iron is the basis for gene regulation by IRP1. On the other hand, IRP2 is mono-functional, having activity only as an IRE-RNA binding protein. Iron controls IRP2 activity by regulating IRP2 protein stability. IRPs play an important role in maintaining iron level. Iron homeostasis is closely linked to many diseases such as Alzheimers’s, Parkinson’s, Diabetes, Cancer and Tuberculosis12.Cellular iron homeostasis is accomplished by the coordinated and balanced expression of iron storage protein ferritin and the iron-uptake protein transferrin receptor. Post-transcriptional regulation of iron homeostasis primarily occurs through the action of iron regulatory proteins, IRP1 and IRP2. IRPs coordinate iron-related gene expression by binding to iron responsive elements (IREs) located in the 5′ or 3′ untranslated regions of mRNAs encoding proteins for iron transport, storage, or utilization; binding blocks either mRNA translation  or degradation 13. Binding of IRE-RNA with IRP1 occurs mainly through bonds made to the terminal loop and stem interrupting C through two separate binding sites2. Approximately two dozen RNA/protein bonds are distributed between the binding sites, giving high affinity and specificity to the interaction. It appears likely that IRP1 binds all naturally occurring IRE-RNA utilizing the same bonding pattern. The structure of the IRE-RNA bound to IRP1 differs from the predominant structure of the RNA in solution, underscoring the importance of conformational flexibility for this high affinity interaction, and raising the possibility that affinity differences observed among these RNA/protein interactions depend in part on the IRE-RNA’s conformational flexibility.Canonical IRE-RNAs are 28–30 nucleotide long stem-loop structures, with a conserved terminal loop sequence, 5′-CAGUGX-3′ , and an unpaired C residue interrupting the stem 5-nucleotides to the 5′ side of the terminal loop sequence. The C and second G residues of the terminal loop sequence form a base pair creating a pseudo-triloop –AGU- in IRE-RNAsFacile conformation change is also important to IRP1 function, particularly its interconversion between cytosolic aconitase and IRE-RNA binding protein. This interconversion involves large scale domain repositioning and localized conformational changes to form the two RNA binding pockets. Notably, coordination of the iron-sulfur cluster prevents key local protein rearrangements necessary to form the terminal loop binding pocket. Iron regulation of IRP1 function is based on such structural constraints.15, rRNA16, ribozymes21, riboswitches in bacterial mRNAs, where metals contribute to RNA function and metal sensing25. Changes in translation of the iron responsive messenger RNAs, dependent on noncoding structures (IRE), are currently ascribed to iron effects on the IRP1 and IRP2, repressor protein, and translation initiation factor (eIF4F), that bind IRE-RNAs and which are modified or degraded by increases in cellular iron concentration26. IRE-RNA binds metal ions (Mg2+) at specific sites27 as do tRNAs, rRNAs, ribozymes and riboswitches. We have previously shown29 that metal ions directly affect the IRE-RNA/IRP1 stability and the binding affinity favors ferritin IRE-RNA over aconitase IRE-RNA, illustrating the effects of phylogenetically conserved differences30 between the two RNAs. Further, Fe2+  could bind to IRE-RNA and act as a ligand to reduce IRP1 binding28. This ligand has a further positive effect by increasing binding of eIF4F31. Metal ion changes the conformation of ferritin IRE-RNA based on nuclear magnetic resonance32 and IRE-RNA containing fluorescence 2-amino purine31. Conformational changes of the IRP1 after binding to IRE-RNA has been explain using crystal structure33. However, metal ion induced conformational change of IRP1/IRE-RNA complex is not known.Metal ions directly regulate the function of many RNA classes, e.g., tRNA2+; Mn2+ is an oxygen-stable analogue” for Fe2+. Kd values increased with an increase in temperature. Thermodynamic studies showed a lowering of enthalpy and free energy for the IRE-RNA/IRP1 complex formation in the presence of Mn2+, suggesting reduced hydrogen bonding and overall conformational rearrangement during IRE-RNA/IRP1 complex formation. It has been shown from the 3-dimentional crystal structure that hydrogen-bonding and hydrophobic interactions contribute to the stabilization of IRP1/IRE-RNA complexes2. Hydrogen bonding contributes to the conformational stability of the molecule; change in the number of hydrogen bonds  can cause to conformational changes in the IRE-RNA/IRP1 complex. In order to understand more fully changes in IRE-RNA/IRP1 interactions, we investigated the effect of temperature on the kinetics of IRE-RNA binding with IRP1 in the absence and presence of Mn2+. Temperature dependent rate constant values were used to determine the activation energies for FRT and ACO2 IRE-RNA binding to IRP1. Significant decrease in activation energy for binding of FRT and ACO2 IRE-RNA with IRP1 was observed in the presence of Mn2+. Decreases in the enthalpic contribution, and lower activation energy for the IRE-RNA/IRP1 complex in the presence of Mn2+ provide a path with lower energy barrier.The stability of the IRP1/IRE-mRNA complex is dictated by free energy change that involves both enthalpic and entropic contributions, and thus requires a thermodynamic approach. In this study, we show the effects of temperature on the equilibrium of IRE-RNA binding with IRP1 in the absence and presence of MnKd) for the interactions of the two IRE-RNAs with IRP1 increased with an increase in temperature. Kd values for the binding of IRP1 to FRT IRE-RNA increased from 4.6 ± 0.2 nM to 19.2 ± 0.4 nM, and for ACO2 IRE-RNA from 55.9 ± 3 nM to 155 ± 4 nM at temperatures ranging from 5 °C to 30 °C (Table 2+). The dissociation constant for the FRT IRE-RNA-Mn2+/IRP1 complex increased from 83.3 ± 4 to 193 ± 5 nM while the dissociation constant of ACO2 IRE-RNA-Mn2+/IRP1 complex increased from 535 ± 33 nM to 801 ± 36 at temperatures ranging from 5 °C to 30 °C , entropy (ΔS) and free energy (ΔG) obtained from van’t Hoff plots of - ln Keqversus reciprocal of temperature (T−1) are shown in Fig. 2+) significantly changed the enthalpic and entropic contributions for the binding of FRT IRE-RNA and ACO2 IRE-RNA with IRP1. The ∆H and ∆S values for the binding of the FRT IRE-RNA with IRP1 were −38.4 ± 2.4 kJ/mol and 21.1 ± 1.3 J/mol/K, respectively, whereas for binding of FRT IRE-RNA to IRP1 in presence of Mn2+ the values were −23.6 ± 1.3 kJ/mol and 51.0 ± 3.3 J/mol/K, respectively. The thermodynamic analyses in the absence of Mn2+ showed that the binding of FRT IRE-RNA to IRP1 is 88% enthalpically driven and 15% entropy favorable with a 73% greater enthalpic contribution to ∆G at 25 °C. Similarly, the binding of ACO2 IRE-RNA with IRP1, showed 71% enthalpically and 29% entropically favorable with a 42% greater enthalpic contribution to ∆G at 25 °C. Addition of Mn2+ decreases the enthalpic contribution to about 61% and increases the entropic contribution to about 39% for FRT IRE-RNA binding to IRP1. Further, Mn2+ decreases the enthalpic contribution to about 32% and increases the entropic contribution to about 68% for ACO2 IRE-RNA binding to IRP1. These data suggest that metal ion induces a conformational change in FRT and ACO2 IRE-RNA/IRP1 complexes resulting in reduced hydrogen bonding. The ∆G values at 25 °C were calculated from Eq. 2+ further decreased in ΔG value for FRT IRE-RNA and ACO2 IRE-RNA binding to IRP1  experiments; they were conducted using high concentration of IRP1 and limiting concentrations of FRT IRE RNA and ACO2 IRE RNA to ensure that the bimolecular combination of IRP1 and FRT and ACO2 IRE RNA was pseudo-first order. Stopped-flow data for a typical reaction of IRP1 binding to FRT or ACO2 IRE-RNA were plotted as the anisotropy change versus time are shown in Fig. 29. Analyses of the data using a double exponential components did not improve the fitting results (not shown). The values of the observed rate constants for complexes of IRP1 with FRT or ACO2 IRE-RNA increased with an increase in temperature (Table k2 = 397 ± 8.5 s−1) and ACO2 IRE-RNA/IRP1 binding rate (k2 = 53 ± 2.4 s−1) at 25 °C were ~4.0- and 2.5-fold faster than at 5 °C  of the FRT and ACO2 IRE-RNA binding to IRP1 in the absence and presence of Mn2+, the temperature dependent rate constant  contributes to the conformational stability of the molecule. IRP1 contains two binding pockets that make 22 bonds with IRE-RNA. The conversion of IRP1 from an aconitase to an IRE-RNA/IRP1 complex requires extensive changes in protein and RNA conformation2. The conformational change of the IRE-RNA/IRP1 complex may also be controlled thermodynamically as reported previously for RNA-protein interactions for U1A protein binding to stem/loop II of U1 snRNA38. This conformational change most likely also plays a role in kinetics of the FRT and ACO2 IRE-RNA with IRP1 complexes.In this study, we have examined the detailed temperature dependent equilibria and kinetics for complex formation of ferritin and mitochondrial aconitase IRE-RNAs binding to IRP1. Previously39. In order to respond rapidly to changes in the cellular Fe2+ concentrations, repression and de-repression of IRE-mRNA must occur on a relatively rapid time scale and modulation of these rates can have important consequences for iron homeostasis. The kinetic rates for the interaction of FRT and ACO2 IRE-RNA with IRP1 follow a single, bimolecular binding step mechanism such as a fast binding followed by a conformational change. Kinetic mechanism is consistent with our previously determined rates of reaction for FRT IRE-RNA and ACO2 IRE-RNA with IRP129. Using stopped-flow kinetics, we have determined the kinetic rate at different temperature in the absence and presence of metal ion for the binding of FRT and ACO2 IRE-RNA with IRP1, which has not previously been measured. Metal ion affected both the association rate and the dissociation constant for IRP1 interaction with FRT and ACO2 IRE-RNA. These results suggest the interactions with metal ions affect the intermediate conformational change in complex formation and reflect a direct sensing of the ion by IRE-RNA, analogous to a riboswitch. Metal ion destabilization of messenger IRE-RNA/protein repressor complexes competes with the stabilization conferred by the very large number of bonds between the protein and the IRE-RNA and the stability of the RNA fold. Selective, metal induced destabilization of IRE-RNA/repressor protein complexes as well as other selective, metal-RNA interactions28, emphasize the sensitivity of RNA structure/function to the environment.The mechanism of the iron effect in cells is unknown; when cells or animals are treated with iron, usually as iron salts or heme, IRE-mRNAs are shifted to polysomes (e.g. ferritin) or degraded (e.g. transferrin receptor) along with degradation of RNA-free repressor proteins, and increased insertion of [4Fe-4S] clusters into the RNA- free form of one of the protein repressors (IRP1), which becomes cytoplasmic aconitase40. The concentration of protein was determined by a Bradford assay with bovine serum albumin as standard41 using a Bio-Rad protein assay reagent . RNA oligonucleotides labeled with fluorescein at the 5′ terminus (FIIRE-RNA) were synthesized by Gene Link, Inc. Hawthorne, New York, U.S.A. The ferritin IRE-RNA  and mitochondrial aconitase IRE-RNA  were used for equilibrium and kinetic studies as described previously29. FRT IRE-RNA or ACO2 IRE-RNA was dissolve in buffer containing 40 mM HEPES/KOH, pH 7.2 and RNA was melted and reannealed by heating to 85 °C for 15 min with slow cooling to 25 °C42. The concentrations of FRT IRE-RNA and ACO2 IRE-RNA were determined spectrophotometrically using the absorbance at 260 of 40 µg/ml RNA as 1. The purity of synthesized RNAs were checked by measuring the absorbance ratio, A260/280 nm, and the absorbance ratio was 2.1.The recombinant rabbit IRP1 protein was expressed in yeast and isolated using methods as described previously2+ (an O2-resistant surrogate) in the titration buffer, 40 mM HEPES/H+ , pH 7.2, 100 mM KCl. We use Mn2+ as an oxygen-resistant surrogate for Fe2+; in addition, Mn2+ has effects similar to Fe2+ for the IRP1/IRE-RNA complex formation28, to facilitate experiments in the presence of air43. All samples were pre-incubated for at least 15 minutes prior to the titration experiments. The FRT IRE-RNA/IRP1 and ACO2 IRE-RNA/IRP1 interaction experiments were performed at temperatures, 5, 10, 15, 20, 25 and 30 °C; sample temperatures were measured using a thermocouple device inside the cuvette. Interactions of FRT IRE-RNA/IRP1 and ACO2 IRE-RNA/IRP1 complexes were measured by the enhancement in FIIRE-RNA anisotropy. The anisotropy data were fitted to Equation  was used to obtain the Kd values and standard errors for parameters using nonlinear least square fitting of the data. For all equilibrium measurements, three independent titration experiments were performed, and the value reported is the average.Temperature dependent fluorescence anisotropy experiments for the binding of FRT and ACO2 IRE-RNA with IRP1 protein were performed using an L-format detection configuration of a Spex Fluorolog τ2 Spectrofluorimeter equipped with excitation and emission polarizers. The anisotropy values for each sample were monitored at an excitation wavelength of 490 nm and emission wavelength of 520 nm. The excitation and emission slits were 4 and 5 nm, respectively. The excitation slits were chosen to avoid photo-bleaching, and the absorbance of the sample at the excitation wavelength was less than 0.02 to minimize the inner-filter effect. Emission spectra were corrected for the wavelength dependent lamp intensity and monochromator sensitivities. In order to study the temperature dependence of the FRT IRE-RNA/IRP1 and ACO2 IRE-RNA/IRP1 complexes, samples were thermostatically adjusted at different temperatures , using a 10-mm path length quartz cuvette. 100 nM of 5′ fluorescein labeled FRT IRE-RNA or ACO2 IRE-RNA was incubated with varying concentrations of IRP1 (0.0–1 mM) in the absence and presence of 50 µM Mn2+, the samples were thermostatically adjusted at different temperatures, 5, 10, 15, 20, 25 and 30 °C, respectively. The temperature dependent Keq was used to determine enthalpy (ΔH), entropy (ΔS), and free energy (ΔG) of FRT and ACO2 IRE-RNA binding to IRP1 in the absence and presence of Mn2+. The thermodynamic parameters of FRT IRE-RNA and ACO2 IRE-RNA binding to IRP1 were analyzed according to the van’t Hoff equation.R and T are the universal gas constant and absolute temperature, respectively. Keq, the equilibrium constant, was determined as a function of temperature. ΔH and ΔS were obtained from the slope and intercept of the plot of lnKeqversus 1/T (kelvin). ΔG was determined at 25 °C using Equation In order to study the temperature dependence of FRT IRE-RNA and ACO2 IRE-RNA interaction with IRP1 in the absence and presence of 50 µM MnFIIRE-RNA anisotropy. After rapid mixing of 0.1 µM  FRT or ACO2 FIIRE-RNA with 1 µM  of IRP1 protein, the time course of the anisotropy change was recorded by computer data acquisition. Samples were degassed prior to loading into the syringes. All measurements were performed in titration buffer containing 40 mM HEPES/H+, pH 7.2, 100 mM KCl. The stopped-flow traces shown under “Results” are the average of 5–7 individual shots to improve the signal-to-noise ratio. Each averaged set of stopped-flow anisotropy data was then fitted to nonlinear analytical equations using GlobalWorksTM analysis software (OLIS).Temperature dependent kinetic measurements of FRT IRE-RNA and ACO2 IRE-RNA interactions with IRP1 protein at temperatures, 5, 10, 15, 20 and 25 °C were performed on an OLIS RSM 1000 stopped-flow spectrometer with polarizers. The dead time of the instruments was 1-ms. Excitation wavelength was 490 nm and emission wavelength was 520 nm for fluorescein labeled FRT IRE-RNA and ACO2 IRE-RNA. In each experiment, 1000 pairs of data points were collected throughout the reaction. Anisotropy changes were examined up to 150 ms. In order to determine the temperature dependent association rate constants, the temperature of the flow-cell and solution reservoir were maintained using a temperature controlled circulating water bath. IRP1 protein binding induced an increase in 2+ on the temperature dependent FRT and ACO2 IRE-RNA binding with IRP1, 50 µM Mn2+ was added to both IRE-RNA and protein solutions, at the same concentration, and the solutions were incubated separately for 15 min before adding to titration buffer containing the same metal ion concentration as the IRE-RNA and protein solutions (50 µM Mn2+). Experiments were performed as described above. Kinetic data were evaluated by fitting to the single-exponential functions as described elsewhere49 and further analyzed as described below.In order to measure the effect of Mn51. Data were fit to the single-exponential functions. Fitted curves correspond to the following single-exponential equation,(t) is the observed anisotropy at any time, t, and rf is the final value of anisotropy, and A is the amplitude. kobs is the observed first order rate constant. The reaction was consistent with a single-exponential process. An assessment of each fit was made from the residuals, which measure the difference between experimental data and the calculated fit. The program KaleidaGraph version 2.1.3 (Synergy Abelbeck Software) was used for least-squares fitting of data with linear equations and determination of standard errors for parameters obtained from the fits.Temperature dependent stopped-flow traces representing binding of FRT IRE-RNA or ACO2 IRE-RNA with IRP1 protein were analyzed using a curve-fitting program, Global analysis software as described previously2+, the fitted rate constants were used to construct Arrhenius plots according to the equationEa is the activation energy, k is the rate constant, R is the universal gas constant, T is the absolute temperature, and A is the Arrhenius pre-exponential factor. The activation energies were calculated using the slopes of the fitted linear plot of ln k versus 1/T (kelvin).To determine the activation energy of the complexes, FRT or ACO2 IRE-RNA with IRP1 protein in the absence and presence of Mn


Symbiodinium spp. from the coral Porites astreoides originating from an inshore and offshore reef in the Florida Keys. The acclimatization potential of Symbiodinium spp. to changing temperature regimes was also measured via a two-year reciprocal transplant between the sites. Offshore coral fragments displayed significantly higher expression in Symbiodinium spp. genes PCNA, SCP2, G3PDH, PCP and psaE than their inshore counterparts (p<0.05), a pattern consistent with increased bleaching susceptibility in offshore corals. Additionally, gene expression patterns in Symbiodinium spp. from site of origin were conserved throughout the two-year reciprocal transplant, indicating acclimatization did not occur within this multi-season time frame. Further, laboratory experiments were used to investigate the influence of acute high temperature (32°C for eight hours) and disease (lipopolysaccharide of Serratia marcescens) on the five metabolically related symbiont genes from the same offshore and inshore P. astreoides fragments. Gene expression did not differ between reef fragments, or as a consequence of acute exposure to heat or heat and disease, contrasting to results found in the field. Gene expression reported here indicates functional variation in populations of Symbiodinium spp. associated with P. astreoides in the Florida Keys, and is likely a result of localized adaptation. However, gene expression patterns observed in the lab imply that functional variation in zooxanthellae observed under conditions of chronic moderate stress is lost under the acute extreme conditions studied here.Scleractinian coral are experiencing unprecedented rates of mortality due to increases in sea surface temperatures in response to global climate change. Some coral species however, survive high temperature events due to a reduced susceptibility to bleaching. We investigated the relationship between bleaching susceptibility and expression of five metabolically related genes of  Symbiodinium spp. commonly referred to as zooxanthellae) declines severely in the host coral allowing the white skeleton below to show [A decline in global coral populations as high as 50% has been recorded over the last 30–40 years, with strong links to coral bleaching –3. Bleac to show , the los to show . Given t to show –11.Symbiodinium spp. clade type harbored by the host [et al. [Stylophora pistillata colonies was determined by compositional differences at the subclade level within Symbiodinium spp. of clade C. In the Florida Keys, several studies found variation in bleaching susceptibility between inshore and offshore communities of P. astreoides [P. astreoides of the Florida Keys is relatively unknown [Coral susceptibility to bleaching is influenced by the the host . For exa [et al.  demonstrtreoides –16. The treoides . However unknown .in vivo under acute, extreme stress most commonly associated with mass mortality events. Here, chronic moderate stress refers to elevated levels of a stressor experienced over a lengthy period of time (i.e. weeks or months). Chronic moderate stress will produce a response such as bleaching, but may not result in death [Coral diseases are a second leading stressor threatening the mortality of coral reefs , and canin death . Acute, in death . Herein,Serratia marcescens [S. marcescens is derived from the gut of animals, including humans, and was identified in wastewater of the Florida Keys [Acropora palmata species in the Florida Keys [Scleractinian coral have recently experienced up to 80% mortality in the Florida Keys as a result of the separate and combined effects of bleaching and disease . Acroporrcescens . S. marcida Keys . Acroporida Keys . The conida Keys ,20.via gene expression analysis, which targets genes related to the production of metabolically important proteins. Common examples include, proliferating cell nuclear antigen (PCNA), a protein involved in DNA synthesis and replication [SCP2), which mediates the transfer of all common lipids [G3PDH), an enzyme involved in glycolysis [PCP) and photosystem I reaction center subunit IV (psaE). PCP is involved in the capture of solar energy [psaE stabilizes interactions within the photosystem 1 complex [Changes in metabolic processes as markers of bleaching susceptibility have a long-standing history in the study of zooxanthellae response to stress ,26–29. Clication , non-spen lipids , Glyceraycolysis , peridinr energy  and psaE complex . Variati complex –37.P. astreoides by evaluating symbiont gene expression in reciprocal transplants of P. astreoides in the Florida Keys. The expression of zooxanthellae genes PCNA, SCP2, G3PDH, PCP and psaE was measured in coral sub-samples from inshore and offshore reefs differing in thermal and irradiance regimes [S. marcescens in order to determine the synergistic effects of acute (8-hour) heat and LPS exposure. Lipopolysaccharides are large molecules found on the outer membrane of gram-negative bacteria [Here, we studied the role of zooxanthellae in the bleaching susceptibility of  regimes ,17. Diffbacteria . When prbacteria . VariatiSymbiodinium spp. from P. astreoides reciprocally transplanted from an inshore and offshore site in the lower Florida Keys were analyzed for changes in gene expression over two years. Study sites, coral collection, reciprocal transplant design and sample collection for analyses were completed as outlined in Hauff et al. [P. astreoides were collected from Birthday reef (inshore: 24.57917N -81.49693W) and an additional ten 16x16 cm fragments were obtained from Acer24 reef (offshore: 24.55268N -81.43741W). Birthday reef and Acer24 reef are patch reefs separated by Hawk Channel and have notably distinct temperature and turbidity regimes, but are otherwise similar . Relative to Acer24 reef, Birthday reef has higher turbidity, lower light, higher annual average temperatures (~1°C) and lower bleaching .In this study, f et al. . BrieflyP. astreoides fragments taken from Birthday reef and Acer24 reef were sectioned into nearly equal halves. These small fragments were approximately 8 x 16 cm each. In a 2x2 design, corals from each of the two reefs were transplanted either back to their reef of origin or to the other reef (i.e. fragments from offshore transplanted to inshore site) (n = 40). Each smP. astreoides fragments originating and replaced at Birthday (n = 6) and Acer24 (n = 6) reef were brought back to Mote Marine Tropical Laboratory (MMTL). Upon arrival, all 12 colony fragments were sectioned into nine, approximately 2x2 cm fragments each, for a total of 108 samples (54 per site) and allowed to recover for 72 hours [At the end of the field experiment, 72 hours  in a shaS. marsescens lipopolysaccharide were applied  ; the latter application was induced to mimic the immune systems recognition of bacteria via polysaccharide. Purified LPS is a common reagent used to elicit immune responses in human and animal studies [S. marsescens colonies to minimize potential cross contamination of S. marsescens into the surrounding environment, as MMTL’s flow-through system is connected directly to an adjacent public waterway. Additionally, LPS was not added to flow-through sections of the experimental design, see below. Control fragments were maintained in experimental tanks at ambient temperature (28°C) reflective of temperatures experienced in water tables, without exposure to LPS. Each treatment was replicated in two tanks, as outlined below, for six independent units of analysis.The laboratory experiment consisted of a one-way design with three experimental levels: a control temperature , a temperature known to cause bleaching  and a treatment in which both the bleaching temperature and  studies . We chosvia the adjacent waterway to aid in temperature maintenance.Experiments were conducted by placing coral fragments in custom made acrylic boxes (2.5 x 7.5 x 7.5 cm) set within 38-liter fish tanks. The acrylic boxes included three distinct subdivisions each of which housed a single coral fragment. Six 38-liter tanks containing five acrylic boxes each were used in the experiment for n = 90 experimental units. The experimental designs for the six tanks included two controls and two tanks for each of the two treatments. The six 38-liter tanks were filled with artificial sterile seawater  and placed in flow through water tables adjacent to one another. Water tables were 680 liters, fiberglass troughs with seawater pumped through via pumps inside the tanks but external to acrylic boxes. Heaters placed external to the acrylic box maintained temperatures characteristic of bleaching (32°C±0.5) and were monitored with HOBO tags . For the disease response treatment, LPS from S. marsescens  [Coral fragments were rinsed in ASSW prior to placement in acrylic box subsections. ASSW held within the tanks was circulated 5 μg/mL)  was addevia air infusion using disposable pipettes. Water was aerated  taking care not to disturb the fragments. At the end of each experiment, coral fragments were removed from their individual tanks using sterile tongs, wrapped in combusted foil and placed in freezer bags. Coral fragments were then immediately flash frozen in liquid nitrogen and stored at -80°C until processing.Coral fragments were exposed to treatments for eight hours. Within that period of time, individual subdivisions were aerated each hour P. astreoides were stored at -80°C and crushed to a fine powder in liquid nitrogen using a pre-chilled mortar and pestle. Between samples, mortar and pestles were cleaned with liquid detergent and RNase AWAY  and rinsed in ultrapure water.All sub-samples of TRI Reagent  was used to extract RNA from 0.1 g coral powder following a modified manufacturer’s protocol. Extracted RNA was quantified and two-50 μL samples were stored at -80°C. Prior to reverse transcription, samples of RNA were diluted to 5 ng/μL and examined on a bioanalyzer to check for RNA integrity and re-extracted if necessary. Once proper integrity was obtained, 2.5 ng of RNA was treated with DNase I  and reverse transcribed into cDNA following manufacturer instructions using the SuperScript III First-Strand Synthesis SuperMix for qRT-PCR .2O, 1μL cDNA, 0.5μL primer . Cycling conditions for all reactions were: 50°C-2 minutes (1x), 95°C-10 minutes (1x), 95°C-15 seconds and 60°C-1 minute (40x). Primers were designed using Primer-BLAST (NCBI). Primer specificity parameters ignoring targets with six or more mismatches was ignored to insure primer specificity. More detailed primer information for all genes can be found in Tables Fully transcribed cDNA was diluted 1:16 for qRT-PCR. Reaction volumes for qRT-PCR were 10 μL and as follows: 5 μL Power SYBR Green PCR Master Mix, 3.5 μL HCox) and proliferating cell nuclear antigen (PCNA) have previously been identified as viable housekeeping genes in Symbiodinium spp. [In this study, six zooxanthellae-specific genes were analyzed across all sub-samples, and include one housekeeping gene and five experimental genes . The houium spp. ,43. TechMCMC.qpcr [via simultaneous analysis of all genes in one model, accounts for low amplification targets by using molecule counts, and eliminates the need for control genes [et al.[All analyses were performed in R (v3.1.3) using the specialized package CMC.qpcr . In thisol genes . For this [et al..Cox, For analysis of field samples, a two-way model was fit using “treatment” (i.e. transplant versus non-transplant) and “time” (i.e. winter and summer) as fixed factors, as well as their interaction, “treatment:time”. This resulted in a model describing the individual effect of each factor, as well as the effect of their interaction (i.e. the treatment dependent effect of sampling time). Two models were run in order to report all relevant comparisons. The first model used fragments originating from, and replaced at, Acer24 reef as a baseline comparison, meaning all other fragment conditions were compared to fragments originating from and replaced at, Acer24 reef. The second model used fragments originating and replaced at Birthday reef for baseline comparison. Three Markov Chain Monte Carlo chains were run for 25,000 iterations with the first 4,000 discarded as burn-in. Additionally, the model was “informed” using qRT-PCR data generated here from one previously established control gene  and “treatment”  as fixed factors, as well as their interaction term, “reef:treatment”. This resulted in a model describing the individual effects of each factor, in addition to the interaction effect (i.e. the site dependent effect of treatment). Three Markov Chain Monte Carlo chains were run for 25,000 iterations with the first 4,000 discarded as burn-in. The model was “informed” using qRT-PCR data generated here from one previously established housekeeping gene  and usePCNA, . DiagnosSymbiodinium spp. from Acer24 reef that were transplanted to Birthday reef were not significantly different from patterns of zooxanthellae that were from, and held, at Acer24 reef  , S1 Filer24 reef .P. astreoides sub-samples originating from Birthday reef exhibited a significant down regulation relative to symbionts originating from Acer24 reef. These included PCP (p = 0.002), PCNA (p = <0.001) and psaE (p = 0.044) and G3PDH (p = 0.052) (Symbiodinium spp. originating from Birthday reef and transplanted to Acer24 reef exhibited significant down-regulation of the genes PCP (p<0.001), PCNA (p<0.001), psaE (p = 0.018), and G3PDH (p = 0.024)(Symbiodinium spp. originating at Acer24 reef and transplanted to Birthday reef exhibited significant up-regulation of genes PCP (p<0.001), PCNA (p<0.001), psaE (p = 0.002), G3PDH (p<0.001) and SCP2 (p = 0.028) , Table 3 = 0.024), Table 3 = 0.028), Table 3In contrast to above, no significant effect of site was found when comparing zooxanthellae gene expression from sub-samples originating at Acer24 reef and zooxanthellae originating from Acer24 reef sub-samples transplanted to Birthday reef p>0.05, . FurtherG3PDH and PCP experienced slight decreased expression in winter compared to summer, while psaE experienced no change in expression. Finally, the interaction between time and transplant was not significant .No significant effect of site or treatment was found on the expression of  p>0.05, , S2 File p>0.05, . For insCox), an increase in temperature resulted in a decrease in expression  displayed significantly higher expression in all offshore Acer24 reef sub-samples compared to inshore Birthday reef sub-samples , reflects elevated expression in zooxanthellae from Acer24 sub-samples relative to those at Birthday reef. PsaE expression increases in the cyanobacterium Synechocystis spp. during light stress [psaE expression counteracts the effects of reactive oxygen species (ROS), whose presence breaks down photosystem II machinery [Symbiodinium spp. to bleaching conditions [psaE in Acer24 reef samples may represent an initial attempt to counteract degradation of photosynthetic machinery repair as reported in McGinley et al. [Symbiodinium spp. A13. Increase in the expression of psaE also demonstrates the higher stress environment offshore.Rates of photosynthesis increase in response to moderate stress in cultured ium spp. . The inct stress . The incachinery . ROS pronditions . Increasy et al.  for SymbP. astreoides from Acer24 and Birthday reefs in response to reciprocal transplant [et al. [P. astreoides in the Florida Keys are functionally variable and the lower expression of stress response genes relative to the offshore reef likely reflects better adaptation to bleaching susceptibility inshore.The lack of an effect of transplantation and the retention of zooxanthellae gene expression patterns from reef of origin, demonstrates that environmental change does not influence the expression patterns of zooxanthellae genes investigated in this study. If acclimatization were an important means of counteracting bleaching susceptibility, zooxanthellae gene expression patterns would have likely changed in the transplant experiment. Instead, gene expression may be genetically determined by adaptation, as was also proposed for the coral host . The lacansplant . Differe [et al.  reflect P. astreoides demonstrated functional variation in the field, the continued success of coral reefs will likely be determined by the ability of the coral-zooxanthellae association to survive exposure to chronic moderate stress and acute extreme bleaching events [S. marsescens; a treatment that represents a primary stress coupled with a secondary stress. Although data was not statistically significant  evaluated the genetic response of the coral host. Coral host genes associated with LPS recognition increased under acute heat and LPS stress. This suggests that coral, rather than zooxanthellae, are the first responders to acute stress. Additionally, as the current study modeled the response of zooxanthellae to LPS exposure, and not a bacterial infection, it is possible the 8-hour sampling period was too long to observe a proper response. The response of the innate immune system to LPS molecules is very rapid, and may not have been captured after eight hours [We speculate the lack of a symbiont gene response is likely due to the coral host influencing zooxanthellae gene expression by providing protection to zooxanthellae. This possibility is suggested by the observation that nditions . Becauseht hours . Decipheht hours  will be P. astreoides as a consequence of chronic stress was established under similar or less stressful conditions in the field. The role of the host as a protective mechanism may explain the apparent lack of variability reported. Alternatively, the short time period may have also skewed the results, indicating longer sampling intervals are required to induce a significant stress response in zooxanthellae. The ecological implications of the field and lab findings suggest that the response of the coral holobiont to stress is dynamic. As such, the future persistence of P. astreoides is dependent on a myriad of environmental and biological parameters, as well as a complex interplay between the host and symbiont. To the authors’ knowledge, this study marks the first study of zooxanthellae gene expression in response to synergistic temperature as a primary stress coupled to LPS as a secondary stress.Short-term perturbations such as acute stress are difficult to capture in the field. As a counterpart to field studies, laboratory studies are important as they allow for manipulation and the capture of acute stress perturbations. This study demonstrated an absence of statistically significant functional variability to acute stress, even though functional variation in zooxanthellae populations from S1 FileCt values for corresponding genes are listed in columns 1–6. Transplant lists reef or origin and reef of transplant, respectively. Time refers to the season in which the fragment was sampled, i.e. Winter or Summer.(CSV)Click here for additional data file.S2 FileCt values for corresponding genes are listed in columns 1–6. Sample refers to the coral fragment the gene corresponds to with duplicate internal replicates, while individual refers to parent coral colonies. Transplant corresponds to reef or origin (i.e. Birthday Reef or Acer Reef). Treatment refers to whether the coral experienced control,(CSV)Click here for additional data file.