Flavivirus in the Flaviviridae family) is capable of being sexually transmitted. Thirty to fifty percent of men with confirmed ZIKV infection shed ZIKV RNA in their semen, and prolonged viral RNA shedding in semen can occur for more than 6 months. The cellular reservoir of ZIKV in semen is unknown, although spermatozoa have been shown to contain ZIKV RNA and antigen. Yet, spermatozoa are not a requisite for sexual transmission, as at least one case of ZIKV sexual transmission involved a vasectomized man. To determine the cellular reservoirs of ZIKV in semen, an established animal model of sexual transmission was used. The majority of virus detected in the seminal fluid of infected mice during the peak timing of sexual transmission was from the supernatant fraction, suggesting cell-free ZIKV may be largely responsible for sexual transmission. However, some ZIKV RNA was cell-associated. In the testes and epididymides of infected mice, intracellular staining of ZIKV RNA was more pronounced in spermatogenic precursors (spermatocytes and spermatogonia) than in spermatids. Visualization of intracellular negative strand ZIKV RNA demonstrated ZIKV replication intermediates in leukocytes, immature spermatids and epididymal epithelial cells in the male urogenital tract. Epididymal epithelial cells were the principal source of negative-strand ZIKV RNA during the peak timing of sexual transmission potential, indicating these cells may be the predominant source of infectious cell-free ZIKV in seminal fluid. These data promote a more complete understanding of sexual transmission of ZIKV and will inform further model development for future studies on persistent ZIKV RNA shedding.While primarily a mosquito-borne virus, Zika virus  is primarily a mosquito-borne virus, there are now confirmed sexual transmission cases of ZIKV from infected males to their partners. Using a previously established mouse model of sexual transmission, ZIKV was herein demonstrated to infect the testes and epididymides concurrently, suggesting that testicular infection is not required to seed infection of the epididymides. Also, replication of ZIKV was visualized by staining for ZIKV negative-strand RNA. ZIKV replication in leukocytes, immature spermatids and epididymal epithelial cells correlated with the peak of sexual transmission potential. Spermatozoa were rarely observed to stain positive for ZIKV replicative RNA intermediates, demonstrating spermatozoa are likely not a major source of infectious virus in semen. In addition, a greater fraction of the infectious virus in seminal fluids was cell-free versus cell-associated, suggesting that cell-free virus is responsible for sexual transmission. Taken together, these data provide strong evidence for epididymal epithelial cells and leukocytes, and not spermatozoa, as the major sources of infectious ZIKV in semen. Flaviviridae) has been reported in thirteen countries . ZIKV irts, see . Furtherrts, see . These dZIKV shedding in seminal fluids has been modeled in Interferon α/β and–γ receptor knockout AG129 mice and in Rag1-/- deficient mice treated with monoclonal antibody to the Type I Interferon Receptor (IFNAR1) –6. ZIKV The source of ZIKV and/or ZIKV RNA in semen is unknown, though spermatozoa have been suggested as a potential candidate. Previous studies of ZIKV tropism within the murine male reproductive tract identified infectious ZIKV from cellular preparations from the epididymis and ejaculates , 8, 11. in situ hybridization and sequential branched-DNA amplification assay was used to detect intracellular ZIKV RNA molecules in epididymal and testicular samples from ZIKV-inoculated mice. Probes specific to ZIKV genomic RNA  and replicative intermediates of ZIKV RNA  identified epididymal epithelial cells as a major population in the epididymis that contained replicating ZIKV during the time period associated with the peak of sexual transmission in the AG129 mouse model. Leukocyte infiltration into the testes and the epididymides was tracked during acute ZIKV infection and replicating ZIKV was identified in a fraction of these leukocytes. As previously reported, mature spermatozoa harbored genomic ZIKV RNA; however, no mature spermatozoa in the epididymis were positive for negative-strand ZIKV RNA by RNA tissue ISH. Together, these data demonstrate that epididymal epithelial cells and leukocytes are sources of replicating ZIKV and likely play a crucial role in sexual transmission of ZIKV.This study was undertaken to identify cell populations within the murine male reproductive tract that could serve as a source/reservoir for ZIKV replication. To do so, ZIKV tropism for, and replication in, particular cell populations during the timing of peak sexual transmission was analyzed. A novel RNA 10 PFU of three different Asian genotype ZIKV strains , and three mice per group were euthanized at days 6, 8, 10 and 12 post-inoculation. Infectious virus was present in the testes, epididymides, and seminal vesicles of ZIKV-inoculated mice at all time points assessed, but the titers for each organ did not drastically increase over time .Splenomegaly was observed for all the ZIKV-inoculated mice . Signifi10 spermatozoa/mL vs. 6 log10 spermatozoa/mL (PRVABC59) and 6.1 log10 spermatozoa/mL (FSS 13205); p<0.05). P6-740-inoculated mice had lower mean spermatozoa counts over time, but a statistically significant decrease in mean spermatozoa counts was not observed by dpi 12 . Thus, during acute infection of AG129 mice with ZIKV, infection may result in decreased spermatozoa counts.Others have noted testicular pathological features in ZIKV-infected mice. To determine if spermatogenesis was impaired in ZIKV-inoculated AG129 mice, sperm counts were performed at dpi 8, 10 and 12. All ZIKV-inoculated mice had decreased spermatozoa counts from dpi 8 to 12, with a significant decrease on dpi 12 observed for PRVABC59- and FSS 13025-inoculated mice, compared to age-matched control mice  were stained with probes specific to ZIKV genomic RNA . The percent of Z+ mature spermatozoa ranged from 3.8% to 30% .Next, it was determined whether mature spermatozoa contained replicative intermediates of ZIKV RNA on dpi 10. To do this, probe sets specific to replicative intermediates of ZIKV RNA  were used. An average of 4.7% of mature spermatozoa stained positive for Z-, which was ~4-fold lower compared to spermatozoa that stained for Z+ . The stain situ hybridization (ISH) for both ZIKV genomic and replicative RNA at dpi 8 (n = 3) and dpi 10 (n = 3). Overall, ZIKV RNA was concentrated in the head of the epididymis, and minimally present in the testis, at dpi 8. By dpi 10, there was decreased staining in the epididymal head, with extensive staining in the body and tail of the epididymis, and extensive, patchy staining within seminiferous tubules of the testis  and mild, mixed, neutrophilic and mononuclear interstitial inflammation. Tubular lumens contained spermatozoa and degenerating round cells (sloughed spermatogenic precursors and/or leukocytes). The tail of the epididymis was without epithelial alterations and contained mature spermatozoa. RNA ISH detected ZIKV genomic RNA extensively within intact and necrotic EEC, intraluminal round cells, and immature spermatids in the head of the epididymis. ISH for ZIKV replicative RNA showed a similar distribution, with lesser amounts of staining than genomic RNA. Neither genomic nor replicative RNA was detected in the tail of the epididymis at dpi 8. ISH was also performed for protamine-2 (PRM2) mRNA, which encodes for a chromatin protein that replaces histones and is specifically expressed in only step 7 round spermatids through step 15 elongated spermatids . ISH forTestis exhibited mild neutrophilic to mixed interstitial inflammation and edema at dpi 8 and 10 . SeminifTaking into account that pathology showed a mixed neutrophilic and mononuclear interstitial inflammation in the epididymis and testes, infection induced leukocyte infiltration was monitored by flow cytometry. Testes samples from mice sacrificed at 6, 8, 10, and 12 dpi were co-stained with an antibody to the pan-leukocyte marker, CD45, and with Z+ probes see , and 82.Leukocytes with actively replicating ZIKV could be one mechanism through which ZIKV is trafficked to the testis or epididymal lumen. In the testes, leukocytes containing Z- were identified on both 8 and 10 dpi . LeukocySince recent work has implicated circulating monocytes as a reservoir for ZIKV, Z+ and Z- leukocytes were gated to distinguish monocytes from lymphocytes. As shown in To address whether the decrease in mature spermatozoa from 8 to 10 dpi was associated with a decrease in spermatids undergoing spermatogenesis, immature spermatids were quantified by flow cytometry. In the testes, ZIKV infection led to approximately a 50% decrease in PRM2+ round and elongating spermatids . Immatur10 PFU/ejaculate) was significantly higher than the mean titer in the cell pellet (1.4 and 1.5 log10 PFU/ejaculate)  were positive for infectious virus. Of the 15 ejaculates positive for infectious virus, 60% (9 ejaculates) had infectious virus only in the seminal plasma (and not in the cell fraction). All of the ejaculates with infectious virus in the cell pellet also contained infectious virus in the seminal plasma . In addition, the filtered seminal plasma contained higher titers of infectious virus. On dpi 10 and 11, the mean titer of infectious virus in the filtered seminal plasma . By tissue ISH, there was more intense staining for ZIKV RNA in the epididymis as compared to the testes at dpi 8. The more intense staining for ZIKV RNA in the head of the epididymis suggests that the testes and epididymis could be infected concurrently. In the testes, ZIKV positive and negative strand RNA was identified within interstitial leukocytes, peritubular myoid cells and immature spermatids. In the epididymis, ZIKV positive and negative strand RNA staining was found extensively in EEC. In the epididymal tubular lumens, replicating ZIKV was identified in leukocytes and immature spermatids. Consistent with other reports, 8–18% of mature spermatozoa from epididymal cell suspensions stained positive for genomic ZIKV RNA[In this study, a highly sensitive  ZIKV RNA, 17; howThe mechanism(s) by which ZIKV is sexually transmitted remain unclear, as the cell(s) that produce infectious ZIKV in seminal fluids are unknown. The cellular component of semen is a mixture of spermatozoa, white blood cells, immature germ cells, and sloughed epithelial cells . SeminalThe intense staining for ZIKV RNA (both genomic and replicative) in the head of the epididymis, as compared to the testes, may be a result of the increased susceptibility of the epididymis to leukocyte infiltration. Circulating ZIKV-infected blood cells could initiate infection in the head (caput) of the epididymis by traversing the tubuli recti or rete testis . Tissue For HIV, the epididymis is one potential reservoir for infectious virus in seminal fluid. Histopathology of the male reproductive tract, from both human autopsies and macaque studies, have identified HIV and Simian Immunodeficiency virus in the epithelium and connective tissue of the epididymis, in adherent myeloid, leukocyte and lymphocyte populations in the epididymis, and in infiltrating leukocytes in the epididymal lumen and stroma –26. SincA limitation of this model is that AG129 mice are highly susceptible to ZIKV infection. A less susceptible mouse model that does not show severe morbidity or mortality after ZIKV infection could be developed to assess long-term shedding of ZIKV RNA in seminal fluids. Such a model could also be used to study the potential for recrudescence of infectious ZIKV during long-term shedding. EECs express IFN-inducible antiviral proteins after activation of microbial sensors , and it in vitro fertilization treatments for women whose partner is an HIV-infected man include semen washing to separate spermatozoa from leukocytes and other non-spermatozoa cell types before use [The data reported here provide critical insight into potential route(s) of sexual transmission following ZIKV infection of the male murine reproductive tract. Identification of the tissue/cellular reservoir(s) of infectious ZIKV is required for better design and employment of therapeutic interventions, assessing the risk assessment of sexual transmissibility and determining the safety of seminal samples used for assisted reproductive technology. It is crucial to determine what cell populations in the male reproductive tract harbor persistent viral infection, and what the effects of immunosuppression are on the recrudescence of infectious virus production in these cell populations. Therapeutic interventions for fore use , 30. WheThe virus isolates used in this study were: PRVABC59 , P6-740 , and FSS13025 .10 PFU of ZIKV strain PRVABC59, P6-740, or FSS13025 diluted in PBS. Mice were euthanized after isoflurane-induced deep anesthesia followed by cervical dislocation. Tissues were collected at time of euthanasia and mice were not perfused prior to necropsy. For plaque assays on the testes and seminal vesicles, tissues were weighed and homogenized using a pestle in an equal weight/ volume of BA-1 medium, and then clarified by centrifugation and serially diluted for plaque assay on Vero cells to enumerate titers represented as plaque forming units (PFU per gram tissue). For plaque assays on the epididymides, a portion of the epididymal lumen single cell suspension prepared for flow cytometry was serially diluted for cell plaque assay to enumerate infectious titer (PFU per epididymides). The overlay for the plaque assay was added four days post-inoculation. A portion of the epididymal cell suspension was also used to enumerate mature spermatozoa. The cells were stained with trypan blue to increase contrast and observed under bright field microscopy. Intact, mature spermatozoa were counted on a hemocytometer.Mice deficient in interferon α/β and -γ receptors (AG129 mice) were bred in-house, and the receptor knockout genotype of the mice was confirmed as described previously . 12-to 1Seminal fluids from male AG129 mice were collected as previously described , 5. In bTo separate the ejaculates into cell pellet and seminal plasma fractions, ejaculates were spun at a 250 rcf, and the supernatant was diluted and filtered through a 0.2 μM filter to remove any remaining cells. The remaining cell pellet was washed three times with PBS and subsequent centrifugations to remove any extracellular virions before being sheared with a needle to release intracellular virions. Infectious ZIKV was titrated by Vero cell plaque assay.Single-cell suspensions of the testis and epididymides were generated by mechanical disruption. Disruption of the testis involved decapsulating each testis and homogenizing in 1X PBS and filtering the cells through a 70 μM filter (Sigma) followed by multiple washes of the filter with 1X PBS. The epididymides were minced in 1 mL of 1X PBS and incubated at 37°C for 10 minutes. The resulting cell suspension was filtered through a 70 μM filter (Sigma).For RNA staining analyses, cells were treated as per manufacturer’s instructions using a commercially available kit  and a probe set targeting either positive or negative strand ZIKV RNA . Briefly, cells were stained with fluorochrome-conjugated mAb against mouse anti-CD45  for 30 minutes at 4°C, and subsequently cells were fixed, permeabilized and stained with the ZIKV RNA, or PRM2 probe sets. Cells were collected at low pressure on an Accuri C6 flow cytometer (BD Biosciences) equipped with blue (488 nm) and red (635 nm) lasers. Single stained samples, UltraComp eBeads microspheres  stained with Alexa Fluor 488 or 647 RNA compensation controls or mouse anti-CD45 PE-Cy7, were used for compensation controls. FlowJo10.2  was used to analyze data, including the FlowJo automated compensation procedure. Gates for viral RNA positive cells were set using single cells suspensions from uninoculated mice that underwent the same staining procedures. Gates for ZIKV RNA staining were set with less than 0.2% background in uninoculated mice. See 10 μL aliquots of the epididymis single cell suspensions processed for PrimeFlow and stained with ZIKV RNA probes were dried onto a microscope slide. ProLong Diamond Antifade mountant was used to mount coverslips to the samples. Slides were visualized on a Zeiss LSM 800 confocal laser-scanning microscope using the 63x objective. Mature spermatozoa were counted as follows: sperm had to be clearly distinguished from one another, and only mature spermatozoa with a head attached to a midpiece and tail were counted. 50–100 sperm were counted for two mice from each ZIKV-inoculated group.in situ hybridization (ISH). The ViewRNA ISH tissue assay (ThermoFisher) with probe sets targeting either positive or negative strand ZIKV RNA or murine PRM2 mRNA (GenBank accession number X07626.1) were utilized. As a negative control, tissue sections from an uninfected male mouse were stained using the ZIKV (+) and (-) probe sets. As a positive control, tissues were stained using probes against the housekeeping murine mRNAs GAPDH, PIPB, and β-actin. Tissue sections were de-paraffinized and underwent an optimized View RNA ISH tissue assay, with pre-treatment and protease treatments lasting ten minutes each. Positive staining (red punctate staining) was detected using an alkaline-phosphatase label probe. Nuclei were counterstained with Gill’s Hematoxylin I.Tissues were placed into 10% neutral buffered formalin for 3 days and then stored in 70% ethanol prior to processing. Sections were cut at 4 microns and stained by ZIKV titers were compared using 2way ANOVA. The percent of Z+ and Z- cells were compared using 2way ANOVA or t-test. All tests were performed in Prism7.All experiments involving mice were approved by institutional animal care and use committee (IACUC) at the Division of Vector-Borne Diseases, Centers for Disease Control and Prevention (Protocol 16–013). All protocols and practices for the handling and manipulation of mice were in accordance with the guidelines of the American Veterinary Medical Association (AVMA) for humane treatment of laboratory animals.S1 FigEpididymal lumen cell suspensions were prepared and stained as described in the methods. An aliquot of the suspension was spun onto a glass microscope slide and a coverslip mounted using ProLong Diamond Anti-fade mountant. (A-C) are representative confocal images of epididymal lumen cells from ZIKV-inoculated mice. (A) Cells were labeled with an Alexa Fluor 488 label probe specific for the ZIKV RNA (+) strand. We observed many foci of ZIKV RNA (green) in round cells from the cell suspensions. Very little staining was observed in spermatozoa. (B) Cells were co-labeled with an Alexa Fluor 488 label probe specific for the ZIKV RNA (+) strand and with an Alexa Fluor 647 label probe specific for the ZIKV RNA (-) strand. Once again, the majority of staining for the ZIKV RNA (+) (green) and (-) (magenta) strands occurred in round cells. Very few spermatozoa stained positive for either the ZIKV RNA (+) or (-) strands. When staining was seen in spermatozoa, the foci were small and dim.(EPS)Click here for additional data file.S2 FigTestis and epididymides were harvested from 18–20 week-old AG129 male mice. Single testis cells and epididymis cells suspensions were prepared and stained as described in the methods. A gate to exclude debris was set first (1), followed by a gate to exclude aggregates (2). A Time vs. FSC-A gate was applied next (3). This gate is important to get rid of artifacts that occur when the cytometer pressurizes and de-pressurizes at the start and end of each run. If a live-dead stain was used, a gate for live cells was applied next (4). Since the PE channel was unused, any positive events in this region are not valid, and so a gate was set to exclude any PE+ events (5). This population was then analyzed for CD45 expression (x-axis) and ZIKV RNA events (y-axis). The ZIKV RNA+ events gate was set using an uninfected control mouse (6).(EPS)Click here for additional data file.S3 FigSpleens were harvested from 18–20 week old AG129 mice. A single cell suspension of the spleen was prepared and stained as described in the methods. The probe set for murine housekeeping mRNAs  were used for staining. This control was carried out each time the testis and epididymis single cells suspensions were stained with the ZIKV RNA probe sets. The splenic samples were gated as described in (EPS)Click here for additional data file.


Chem. Commun., 2018, DOI: ; 10.1039/c7md00593h.Correction for ‘The critical role of novel benzophenone analogs on tumor growth inhibition targeting angiogenesis and apoptosis’ by Yasser Hussein Eissa Mohammed  The following are the corrected names for compounds 5b: [2-chloro-6-fluoro-4-(4-fluorobenzoyl)phenoxy]acetic acid ethyl ester.5d: [2-chloro-6-fluoro-4-(4-methylbenzoyl)phenoxy]acetic acid ethyl ester.9a: N′-(2-(4-benzoylphenoxy)acetyl)-1-methyl-1H-imidazole-4-carbohydrazide.9b: N′-(2-(2-chloro-6-fluoro-4-(4-fluorobenzoyl)phenoxy)acetyl)-1-methyl-1H-imidazole-4-carbohydrazide.9c: N′-(2-(2-chloro-4-(4-chlorobenzoyl)-6-fluorophenoxy)acetyl)-1-methyl-1H-imidazole-4-carbohydrazide.9d: N′-(2-(2-chloro-6-fluoro-4-(4-methylbenzoyl)phenoxy)acetyl)-1-methyl-1H-imidazole-4-carbohydrazide.10a: N′-(2-(4-benzoylphenoxy)acetyl)-2-oxo-2H-pyran-5-carbohydrazide.10b: N′-(2-(2-chloro-6-fluoro-4-(4-fluorobenzoyl)phenoxy)acetyl)-2-oxo-2H-pyran-5-carbohydrazide.10c: N′-(2-(2-chloro-4-(4-chlorobenzoyl)-6-fluorophenoxy)acetyl)-2-oxo-2H-pyran-5-carbohydrazide.10d: N′-(2-(2-chloro-6-fluoro-4-(4-methylbenzoyl)phenoxy)acetyl)-2-oxo-2H-pyran-5-carbohydrazide.9a was correct in the main manuscript, but wrong in the ESI. The ESI has been corrected.In addition, the structure of The Royal Society of Chemistry apologises for these errors and any consequent inconvenience to authors and readers.


RNA from tissues of multiple varieties of cassava plants (Manihot esculenta Crantz). The resulting method is suitable for use in mini, midi, and maxi preparations and rapidly achieves high total RNA yields (170–600 μg·g−1) using low‐cost chemicals and consumables and with minimal contamination from polysaccharides, polyphenols, proteins, and other secondary metabolites. In particular, A260 : A280 ratios were > 2.0 for RNA from various tissues, and all of the present RNA samples yielded ribosomal integrity number values of greater than six. The resulting high purity and quality of isolated RNA will facilitate downstream applications (quantitative reverse transcriptase‐polymerase chain reaction or RNA sequencing) in cassava molecular breeding.We developed and modified a precise, rapid, and reproducible protocol isolating high‐quality  A260 : A280 ratios of only 1.7 to 2 and did not report A260 : A230 ratios. Similar to methods for species of the Euphorbiaceae family, such as Manihot esculenta, Hevea brasilensis, Jatropha curcas, and Ricinus communis, modified CTAB‐based protocols have been described as sources of better quality RNA. However, although A260 : A280 ratios were reportedly between 1.7 and 2.2, A260 : A230 ratios of 1.4 and −1.8 or less indicate sufficient contamination to preclude transcriptome analyses such as RNA sequencing Cassava (Multiple cassava tissues were collected from various genotypes of differing ages as follows:in vitro culture systems and were propagated in the field to 8–10 weeks of age by planting in pots with sterile soil containing a 3 : 1 ratio of sand: black soil (no clay topsoil). Leaf tissues of various ages were infested with A. socialis and were then collected at various times until the last nymph stage of the insect. Samples were also collected from storage roots at times that corresponded with differential expression studies of beta‐carotene contents in roots of cassava strains at 12 months of age. Whole tissues from all stages of the experiment were immediately placed in liquid nitrogen and were stored at −80 °C.Several cassava genotypes were collected, and resistances to whitefly were compared with respective controls. Study genotypes including ECU72, COL1468, COL2246, TMS60444, PER368, PER317, and PER608 were obtained from The cassava genotypes HMC‐1 and GM3500‐2 (Esparrago) were studied as representatives of early and late flowering genotypes. Plants were propagated from stakes in the field and, after cleaning, were placed in hydroponic solution for 2 weeks in vitro and in pots (stakes from the field) using growth chambers . Protocols for the preparation of stakes are detailed in part two. To prepare plants for in vitro experiments, tissue cultures were grown into plants and were then transferred to growth chambers for temperature treatments. After 3‐day acclimation, the youngest leaves, buds, and third and fourth leaves with their stems were sampled. Leaves, stems, and roots from in vitro plants were collected separately, and all collected tissues from all stages of the experiment were immediately placed in liquid nitrogen and stored at −80 °C.To analyze the effects of temperature on cassava flowering times, HMC‐1 and Esparrago strains were grown at 15 °C and 30 °C under LD conditions m Tris/HCl , sodium chloride solution 5 m (Sigma‐Aldrich), UltraPure™ 0.5 m ethylenediaminetetraacetic acid (EDTA), , sodium dodecyl sulfate , polyvinyl pyrrolidone , 2‐mercaptoethanol (Sigma‐Aldrich), UltraPure™ DNase/RNase‐free distilled water (Thermo Fisher Scientific), acid‐phenol: chloroform (pH 4.5) with isoamyl alcohol at a final ratio of 25 : 24 : 1 (Thermo Fisher Scientific), chloroform/isoamyl alcohol mixture (Sigma‐Aldrich), lithium chloride , ethanol absolute for analysis (Merck), chloroform for analysis (Merck), and RNaseZap™ RNase decontamination wipes (Invitrogen) were of molecular biology grade and were free of RNAses, DNAses, and pyrogens. All plastic supplies for the preparation of extraction buffer and the tubes used for extraction were disposable and were free of RNAses, DNAses, and pyrogens. We avoided the use of reagents with acute toxicity, such as diethyl pyrocarbonate, which is frequently used to inactivate RNAses.All stock reagents were purchased from commercial suppliers at appropriate stock concentrations, and only SDS and PVP40 were purchased as powders and were weighed and added directly to extraction buffers. The stock solutions UltraPure™ 1 The following RNA isolation methods were used for all RNA preparation scales, including mini preparations of up to 2 μg of total RNA for reverse transcription‐polymerase chain reaction (RT‐PCR) analysis and midi or maxi preparations of 20 μg of total RNA for RNA‐Seq analyses. Prior to extracting RNA, benchtop areas and pipettes with filter tips were cleaned using RNAse zap wipes to inactivate RNAses. All procedures were performed using nitrile or latex gloves without powder.m Tris/HCl, 100 mm NaCl, 25 mm EDTA, 1% SDS, 2% PVP40, and 2% 2‐mercaptoethanol was prepared daily in ultra‐pure water that was free of RNase and was then preheated to 65 °C.Extraction buffer containing 100 mTissues in liquid nitrogen were ground to a fine powder using a mortar and pestle for maxi preparations or a tissue lysis machine for mini‐scale preparations. Liquid nitrogen was continuously provided to avoid melting of frozen tissues and to prevent RNA degradation.g for 20 min at 4 °C, and supernatants were transferred to fresh 2‐mL tubes and were washed with chloroform two more times by shaking and inverting. Supernatants were then transferred to 2‐mL tubes with care not to contaminate with lower chloroform phases, and RNA was precipitated from supernatants (1.5 mL) by adding 0.33 volumes (0.5 mL) of 8 m lithium chloride and incubating at 4 °C overnight.Subsequently, for mini‐scale preparations, 100 mg samples of powdered tissues were added to 2.0‐mL tubes under liquid nitrogen, and 700 μL aliquots of preheated extraction buffer were then added and vortexed. After adding and vortexing of 700 μL aliquots of chloroform, mixtures were centrifuged at 15 000 g for 10 min at 4 °C, supernatants were discarded, and precipitates were dissolved in 500 μL of ultra‐pure water. Poor aqueous solubility of precipitates at this stage is indicative of high RNA quality. Subsequently, RNA was extracted by centrifuging at 15 000 g for 10 min at 4 °C in one volume of acidic phenol: chloroform: isoamyl alcohol (25 : 24 : 1) and then in one volume of chloroform: isoamyl alcohol (24 : 1). Supernatants (500 μL) were then transferred to fresh tubes, and RNA was precipitated using 0.25 volumes of 5 M NaCl (125 μL) and 2.5 volumes of ice‐cold (−20 °C) absolute ethanol (1250 μL) for ≥30 min at −80 °C. Mixtures were then centrifuged at 15 000 g for 20 min at 4 °C. After discarding supernatants, precipitated pellets were washed immediately with 70% ethanol and centrifuged again at 15 000 g for 10 min at 4 °C. Finally, supernatants were discarded, and RNA pellets were air‐dried for 5–20 min and then resuspended in 20–100 μL of ultra‐pure RNAse free water, depending on yield and preparation scale, and were stored at −80 °C.On the following day, mixtures were centrifuged at 15 000 Application of the above protocol for midi or maxi scale requires proportional scaling of sample, initial extraction buffer, chloroform volumes, and centrifuge speeds Table .A260 : A280 and A260 : A230 using a Nanodrop® ND‐1000 spectrophotometer (Thermo Fisher Scientific). RNA quality was further assessed electrophoretically on 1% denaturing agarose gels, which were prepared by adding 0.5 g of agar powder to 50 mL of ultra‐pure RNase free water and boiling until melted, followed by the addition of 8.75 mL of formaldehyde and 5 mL of 10 × MOPS buffer. Gels were run at 90 V for about 45 min. RNA integrity was assessed using an Agilent RNA 6000 Nano Kit with an Agilent 2100 Bioanalyzer  and GDPX9R (AAGCAATTCCAGCCTTGG). PCR mixtures were initially denatured at 94 °C for 5 min and were then subjected to 36 cycles of 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 20 s, and a final extension of 72 °C for 5 min. Amplified products were then separated on 1% agarose gels and were stained with SYBR™ Safe DNA Gel Stain (Invitrogen). Finally, 75 ng cDNA specimens were subjected to quantitative reverse transcriptase‐polymerase chain reactions (qRT‐PCRs) in triplicate using Brilliant II SYBR Green PCR kits (Agilent Technologies) as described in the Rotor‐Gene Q (Qiagen) protocol with the primers 18S‐Fw (ATGATAACTCGACGGATCGC) and 18S‐Rv (CTTGGATGTGGTAGCCGTTT), which amplified a 169‐bp 3ʹ sequence of the 18S gene. A reverse transcription non‐template negative control was included to confirm the absence of genomic DNA, primer‐dimers, and other unfavorable reactions in RT‐PCR and qRT‐PCR analyses.Total RNA  were isolated from tissues of HMC‐1 and Esparrago (ESP) cassava genotypes and were treated with 1 U of DNase I (Promega). First‐strand cDNA were then synthesized in reaction mixtures containing total RNA, anchored oligo‐(dT)20 primer, and SuperScript™ III Reverse Transcriptase (Invitrogen) as described in the manufacturer's instructions. To confirm the absence of DNA contamination, a 412‐bp region of the M. esculenta).Initially, we extracted RNA from cassava leaves using TRIZOL reagent according to previously described methods et al. A260 : A280 ratios increased to 2.02–2.2 indicating very low protein contamination, and A260 : A230 ratios of 2.03–2.29, except in one sample  and Pure link RNA Reagent (Thermo Fisher), are available for RNA extraction, these kits are not designed for analyzing plant tissues containing high starch concentrations, such as cassava plant tissues. The use of these kits without any additional purification steps for such samples dramatically reduces the quality and quantity of extracted RNA. Our past experience with the RNaesy Plant Mini Kit revealed its applicability only for fresh leaf samples or young tissues and not for old leaf tissues or cassava leaves, which are used for treating conditions such as infestation and other stresses, owing to a dramatically reduced quantity and quality of extracted RNA Table .Furthermore, it is obvious that research cost is an extremely critical aspect of a research project. Most of these kits are expensive, and the cost drastically increases with an increased number of samples to be analyzed or with a high concentration of high‐quality RNA requirement. Table . Utsumi et al. Colletotrichum gloeosporioides. However, there is no information regarding the efficiency of the RNaesy Plant Mini Kit for RNA extraction from cassava plants. Wilson et al. et al. 260/A280 ratio was not 2, the 26S rRNA concentration (except on day 1) was low, and no information was obtained regarding the RIN value and the yield of the extracted RNA. In addition, our method has demonstrated applicability not only for tuber tissues but also for other tissues such as leaf and stem.Moreover, some of these kits require specific conditions for certain plant tissues, whereas our method is suitable for various tissues of differing ages under different experimental conditionset al. 1993, we added high concentrations of lithium chloride to samples prior to overnight incubation after the first day to limit DNA contamination Although the extraction buffer used here contained SDS, Tris/HCl, EDTA, sodium chloride, PVP, and β‐mercaptoethanol, as in former protocols, we determined optimal concentrations of these constituents for Euphorbiaceae family plants and confirmed that 2% SDS, 2% PVP, and 1–2% β‐mercaptoethanol Table  produce A260 : A280 ratios of our RNA from various cassava tissues and varieties were all > 2.0, indicating high purity of isolated RNA. Although this ratio is an important indicator of sample quality, the best indicator of RNA integrity is functionality in subsequent applications. Thus, we evaluated RNA integrity according to RIN using a 2100 Bioanalyzer instrument with Agilent RNA 6000 Nano Assay kits (Agilent Technologies). These analyses showed high RIN values in all cases , which extracted by Trizol Method.Table S2. Yields according to A260 : A280 and A260 : A230 ratios of isolated total RNA from various cassava tissues.Table S3. The cost of extracted RNA based on our methodology and different RNA extraction kits.Click here for additional data file.


We designed a series of RNA and DNA constructs with a 1 × 2 loop sequence contained in the ISL, and its modifications, to measure the thermodynamic effects of protonation and magnesium binding using UV-visible thermal denaturation experiments. We show that the wild-type RNA construct gains 0.43 kcal/mol in 1 M KCl upon lowering the pH from 7.5 to 5.5; the presence of magnesium ions increases its stability by 2.17 kcal/mol at pH 7.5 over 1 M KCl. Modifications of the helix closing base pairs from C–G to U•G causes a loss in protonation-dependent stability and a decrease in stability in the presence of magnesium ions, especially in the C68U construct. A79G single-nucleotide bulge loop construct showed the largest gain in stability in the presence of magnesium ions. The DNA wild-type construct shows a smaller effect on stability upon lowering the pH and in the presence of magnesium ions, highlighting differences in RNA and DNA structures. A U6 RNA 1 × 2 loop sequence is rare in the databases examined.U6 RNA contains a 1 × 2-nt internal loop that folds and unfold during spliceosomal assembly and activation. The 1 × 2 loop consists of a C Eukaryotic translation of proteins requires excision of introns from pre-mRNA to form mature messenger RNA. Nuclear pre–messenger RNA excision, termed splicing, is catalyzed by the spliceosome. The spliceosome is a multimegadalton complex that functions as a ribozyme to excise introns and ligate exons in the process of creating mature messenger RNA . It is ccis Watson–Crick/Watson–Crick, cWW), with one-hydrogen bond between residues C67•A79, and an adjacent unpaired uracil, U80 (numbering according to yeast Saccharomyces cerevisiae)  . This 1 evisiae) . The intgen bond . The NMRgen bond . At a pHgen bond . NMR perU80 pair . AnokhinP nonbridging phosphate oxygen abolishes the first step of splicing but can be rescued by thiophilic ions such as cadmium  of RNA and DNA constructs , with C66U showing 1.99 kcal/mol and C68U showing 2.39 kcal/mol lowered stability than wild-type construct in pH 5.5. This indicates that the closing U•G base pairs are changing the structural dynamics of the internal loop.Changing the closing base pairs on the stems from C–G to U•G base pairs, in C66U  and C68U  constructs, decreases the stability of RNA by 1.15 and 1.97 kcal/mol relative to the wild-type construct in pH 7.5. Neither C66U or C68U show an increase in stability in 1 M KCl in pH 5.5 over pH 7.5 , creates an all-pyrimidine 1 × 2 internal loop with stability that is 0.44 kcal/mol lower than the wild type at pH 7.5, indicating that the wild-type construct has at least a single hydrogen bond between Cormation ; Fig. 3.67•A79 with a canonical C67–G79 base pair and converts the construct to a single-nucleotide (U80) bulge loop; this construct shows a 4.83 kcal/mol gain in stability over the wild-type construct at pH 7.5. Altering the 3′ and 5′ neighbor of the newly formed G–C base pair from a C–G to a U•G wobble pair (A79G_C66U and A79G_C68U) decreases the stability of the RNA by 2.6 and 2.9 kcal/mol, respectively; these constructs still have greater stability than the wild-type construct.Changing the A79 to G (A79G) replaces the CThe RNA helix was designed by eliminating the internal loop and has a stability of −17.02 kcal/mol at pH 7.5; the change in pH did not affect its stability ; Fig. 3.Go37 of −7.57 kcal/mol, with a gain of 0.28 kcal/mol in stability upon lowering the pH to 5.5  had ΔH to 5.5 . Subsequ as well . The DNA2 buffer, the RNA wild-type construct gained an additional 1.36 and 2.17 kcal/mol in stability over 1 M KCl in pH 5.5 and 7.5, respectively  of −1.25 kcal/mol (pH 7.5) and −0.56 kcal/mol (pH 5.5) over 1 M KCl   see . When U•cWW) see . No stru67•A79 base pair likely has a single hydrogen bond at pH 7 with energetics of 0.45 kcal/mol. Protonation of A79 allows for another hydrogen bond formation that increases RNA stability by 0.43 kcal/mol. Previous studies show that adenine protonation contributes 0.7 ± 0.3 kcal/mol to the overall stability of the construct in this RNA and is within the range measured here , the pH-dependent gain in stability is lost, indicating that there is no Cse pairs .67•A+79 base pair. The C66U and C68U constructs did not gain stability upon lowering the pH, implying that the modifications to both the 5′ and 3′ base pairs flanking the noncanonical A•C base pair have an impact on the RNA structure of the loop and formation of the hydrogen bonds between C67•A79. A significant decrease in stability is seen for U•G substitutions at positions 66 and 68, with a greater impact on RNA stability seen for the C68U construct. It is possible that the closing U•G base pair alters the loop structural dynamics in the C68U construct. In single-nucleotide bulge loops, closing G•U base pairs destabilized the RNA by 0.5–4 kcal/mol and showed non–nearest neighbor effects (In RNA constructs that contain C–G to U•G modifications (C66U and C68U), we examined the role of C–G closing base pairs in the formation of the C effects .2+ binding pocket in the U6 ISL. Upon protonation of A79, U80 gets flipped into the major groove allowing the RNA to become close to an A-form helix, causing a loss of the binding pocket. Hence, Mg2+ binding was suggested to be lost upon A79 protonation , likely indicating that the G•U base pair at the end of the helix causes significant changes to the loop, perhaps leading to a potential 2 × 3 internal loop, for which ΔGo37,loop of 3.5 kcal/mol is utilized in prediction models  or due to site-specific binding, various studies in our laboratory show a trend that RNA containing bulge loops and internal loops is additionally stabilized in the presence of magnesium ions . ExperimS. cervisiae U6 ISL . For TLC purification, a Baker Si500 silica plate and running buffer of 6:3:1 (v/v/v) 1-propanol, 30% ammonium hydroxide, and water was used. Oligomers were extracted from silica with water and then spin filtered to remove excess silica. Samples were dried by lyophilization using SPD1010 SpeedVac System (Savant). Samples were deprotected with 100 mM acetic acid adjusted to pH 3.8 with TEMED . Samples were desalted with C18 Sep-Pak column (Waters WAT020515) using 5 mM sodium bicarbonate loading buffer (pH 6). Elution was performed twice with 2 mL of 30% acetonitrile and once with 100% acetonitrile. Oligomers were collected, dried, and suspended in a designated buffer. Concentrations were calculated using extinction coefficient derived from pairwise values for nearest neighbors.2 at pH of 7.5, 7, or 5.5. The concentrations of divalent ions were established by complexometric titrations or atomic absorption spectroscopy . Thermodynamic analyses performed here are similar to those in previous work and allow for determining changes in RNA stabilities with respect to its own stability under different ionic conditions, allowing for a comparative analysis between different RNA constructs ] plot for non-self-complementary RNA using the following equations . Thermodynamic data reported is from the van't Hoff analysis, with the exception of the C68U_A79G construct where curve fit data was used.Thermodynamic data were collected as previously described . The conGo37,loop parameters. For a single-nucleotide bulge (Go37 of the RNA control helix was used for ΔGo37, helix. Constructs containing the A79G modification have an additional C–G base pair in their helix and are predicted to contribute −3.3 kcal/mol to the helix (Go37, helix value was used to calculate the helix stability by changing the appropriate nearest neighbor parameters (Disrupting a helical structure destabilizes the RNA. The changes to RNA stability upon the incorporation of internal loops are represented as Δde bulge B, the asde bulge  and, henhe helix . For conrameters .Go37, loop parameter were performed with the energetics of disrupted base stacking considered (Go37, loop parameter were performed with the energetics for new helical parameters that included modifications of appropriate nearest neighbor parameters for G•U base pairs.For 1 × 2 internal loops, the calculations for the Δnsidered :ΔG37,loThe nearest neighbor parameters from Appendix 1The 7-nt sequence shown below was searched using FR3D and FRABase 2.0.5′C C C3′        3′GUAG5′


The fact that structural RNA motifs can direct RNAs to fold and self-assemble into predictable pre-defined structures is an attractive quality and driving force for RNA’s use in nanotechnology. RNA’s recognized diversity concerning cellular and synthetically selected functionalities, however, help explain why it continues to draw attention for new nano-applications. Herein, we report the modification of a bifurcated reporter system based on the previously documented Spinach aptamer/DFHBI fluorophore pair that affords the ability to confirm the assembly of contiguous RNA strands within the context of the previously reported multi-stranded RNA nanoring. Exploration of the sequence space associated with the base pairs flanking the aptamer core demonstrate that fluorescent feedback can be optimized to minimize the fluorescence associated with partially-assembled RNA nanorings. Finally, we demonstrate that the aptamer-integrated nanoring is capable of assembling directly from transcribed DNA in one pot. RNA nanotechnology leverages the formation of programmable base pairs and regular three-dimensional folding patterns of structural RNA moieties to construct materials with precise, predefined shapes ,2,3,4,5.A promising strategy that has been developed to visualize RNA in recent years involves the use of light-up RNA aptamer/fluorophore pairs ,15,16,17Given its unique structure and demonstrated ability to be functionalized with a variety of RNA-based functional groups, we chose to integrate the split-Spinach aptamer into the previously reported RNA nanoring ,37. The taq DNA polymerase  via polymerase chain reaction (PCR), and isolated by a DNA purification kit . Transcription of amplified DNA was accomplished using T7 RNA polymerase  in vitro. The resulting transcripts were purified by polyacrylamide gel electrophoresis (PAGE) . Excised gel fragments containing RNA were placed in Crush and Soak buffer , shook overnight at 5 °C, and the RNA was isolated the next day via ethanol precipitation. The fluorophore, 3,5-difluoro-4-hydroxybenzylidene imidazolinone (DFHBI), was synthesized as previously reported according the protocol of the Paige research group . Ge. Ge41]. After evaluating stem stabilities and linker contributions we identified several sequence combinations that provided at least a 10-fold increase in signal between the split-aptamer in the presence of all the ring components compared to split-aptamer strands incubated by themselves with the highest ratio reaching nearly a 20-fold fluorescence gain. Following these results we evaluated the ability for the split-aptamer system to distinguish between fully- and partially-assembled nanorings. Partially-assembled rings (composed of five out of the six nanoring struts) were expected to produce a higher fluorescent signal than the split-aptamer dimers alone because, like the fully-assembled nanorings, they provide a physical conduit for the split-aptamer strands to be brought together. Previous evaluation of the nanoring however showed that partially-assembled rings are not as thermodynamically stable as fully-assembled rings and so we postulated that the split-aptamer system should be able to show some level of discrimination between the two  In orderEvaluation of the partially-assembled rings revealed a number of interesting insights . In the A distinct advantage of RNA-based nanoparticles relates to their ability to self-assemble isothermally directly from their transcription via DNA templates ,18,43. T2) were used to compare the resulting RNA assemblies of the transcription mixture with an RNA ladder prepared via snap-cooled assembly of the corresponding nanoring. Native PAGE reveals that the fully-assembled nanorings constitute the primary assembly product  require careful testing. This suggests that full-scale automation of the design process remains elusive and that the design of new sytems will undoubtedly contain their own particular parameters which will also require their own experimental optimization. Simply put, the overall performance of variants is and remains context dependent and very difficult to predict. In this regard, even though in vitro studies provide a baseline for assessing the behavior of the a self-assembling nanoparticle/split-aptamer/fluorophore system, evaluation in increasingly complex environments remain necessary for validation and further refinenment. Modeling and design provide an essential starting point but due to myriad of parameters, experimental refinement remains absolutely necessity. Our work also shows that the destablization of stems surrounding the aptamer core generally provide increased sensitivity between fully- and partially assembled nanoparticles. The same is likely to be true for future developments with different nanoparticles—where the goal or purpose of nanoparticle assembly functions, at least in part, to restore and/or compensate the intentionally diminished stabilities of the altered stems.The split-aptamer/nanoparticle system demonstrates the ability, not only to monitor the assembly of multiple strands, but the ability to monitor the assembly of RNA strands not directly linked to the aptamer units. To our knowledge this is the first system developed that demonstrates the ability to detect the formation of tertiary interactions on contiguous strands of RNA. We demonstrate further that fluorescent signaling associated with fully-assembled nanorings can be maximized over the partially assembled split-aptamer system associated with two or five strand mixtures. Given its ability to assemble isothermally from DNA templates, we propose that the co-transcriptional assembly of the split-aptamer/nanoring system provides a robust platform for the further development of a variety of automated and/or high-throughput applications. The nanoring’s four other struts are readily functionalizable and because the nanoring’s helical struts can be increased a full helical turn to accommodate a larger interior space, it is possible that more complex aptamer systems may be able to be incorporated (such as the addition of secondary modular aptamer domain). Finally, fluorescent reporting of fully-formed nanorings provides instant feedback for the previously proposed automated assembly of therapeutic RNA nanoparticles .


Bombus terrestris are characterized by wide phenotypic variability among genetically similar full-sister workers, suggesting a major role for epigenetic processes. Here, we report a high level of ADAR-mediated RNA editing in the bumblebee, despite the lack of an ADAR1-homolog. We identify 1.15 million unique genomic sites, and 164 recoding sites residing in 100 protein coding genes, including ion channels, transporters, and receptors predicted to affect brain function and behavior. Some edited sites are similarly edited in other insects, cephalopods and even mammals. The global editing level of protein coding and non-coding transcripts weakly correlates with task performance (brood care vs. foraging), but not affected by dominance rank or juvenile hormone known to influence physiology and behavior. Taken together, our findings show that brain editing levels are high in naturally behaving bees, and may be regulated by relatively short-term effects associated with brood care or foraging activities.Colonies of the bumblebee  Bombus terrestris and its weak correlation to task performance.Bumblebee workers are genetically highly similar but they show different behaviors such as brood care and foraging. Here the authors report a high level of ADAR-mediated RNA editing in the bumblebee  Variation in behavioral and physiological state in many species is not associated with genetic variation (which is often fairly low), but rather with differential expression of transcription factors, histone acetylation, DNA methylation, and non-coding RNA regulation5. Thus, much attention has been given to epigenetic regulatory processes that may contribute to the regulation of gene expression and increase the functional diversity among genetically similar individuals. These processes include chemical alterations of DNA, chromatin modifiers, non-protein coding RNAs, as well as modifications to the sequence or chemistry of coding and non-coding RNAs3. Previous studies have indeed suggested that methylation mediates changes in brain function in response to even small changes in the environment or  in social role7. However, the rich behavioral repertoire of social insects is complex and cannot be fully accounted for by methylation or any other of the abovementioned processes alone8.Social insects such as ants, termites, wasps and bees, provide excellent model systems for studying how a single genome can generate substantial functional diversity. Individual social insects typically show remarkable plasticity, switching between behavioral and physiological states along with the social environment they experience, during development, or even as fully formed adult insects10. Three ADAR enzymes are encoded by the mammalian genome: double-stranded RNA (dsRNA)-specific adenosine deaminase , dsRNA-specific editase 1  and the catalytically inactive ADAR3 (or ADARB2). Insects are believed to encode a single ADAR enzyme, orthologous to mammalian ADAR211. Given that inosine is read as guanosine by the translation machinery, RNA editing can lead to changes in protein sequences and function12. A-to-I editing of protein coding genes may contribute to behavioral plasticity12 by modifying the amino acid sequence (“recoding”), altering splice sites13, or affecting the regulation of gene expression. Recoding of even a single amino acid may affect protein function. For example, modifications in the catalytic site of an enzyme in the pore forming regions of an ion channel, can profoundly affect specificity, level of activity or ion permeability15. ADAR can also affect gene expression patterns by editing non-coding RNA such as microRNAs and long non-coding RNAs17. The fraction of edited molecules among the RNA copies of a given editing target varies between 0–100% and may be tissue or even cell-dependent18. Editing levels are dynamically regulated, and were shown to depend on developmental stages, various disease states, as well as environmental conditions19. Thus, RNA editing is well positioned to increase phenotypic variability by providing functional heterogeneity across tissues, brain regions or even among cells within the same tissue. The temporal regulation of RNA editing and its sensitivity to varying conditions and environments  may contribute to behavioral or developmental plasticity.Here, we study RNA editing, an epigenetic process that modifies nucleotides in pre-mRNA sequences in a social bumblebee. We focus on RNA editing, mediated by the ADAR (adenosine deaminase acting on RNA) enzymes, the most common type of editing known in animals. ADAR enzymes are evolutionary conserved, predominantly active in the nervous system, and act through selective deamination of specific adenosines to inosines (“A-to-I editing”) within the target RNA20). For example, in the nematode Caenorhabditis elegans ADAR mutants show impaired chemotaxis and abnormal behavior21. In the fruit fly Drosophila melanogaster, ADAR mutants are morphologically normal but exhibit extreme behavioral deficits that include temperature-sensitive paralysis, impaired locomotion coordination, abnormal courtship behavior, weaker circadian rhythms, and aberrant sleep patterns24. Mice mutants with homozygotic deletions of ADAR2 show frequent seizures and die prematurely. Given the substantial effects on behavior and physiology, and its dynamic regulation, we wondered whether RNA editing also plays a role in the social regulation of behavior, a noted hallmark of insect societies. The importance of RNA editing in social insects has so far only been investigated in the leaf-cutting ant (Acromyrmex echinatior), which lives in complex societies with morphologically distinct castes. Approximately 800 transcripts expressed in the ant head showed evidence of A-to-I RNA editing. These transcripts were functionally enriched for genes involved in neurotransmission, circadian rhythms, temperature response, RNA splicing and carboxylic acid biosynthesis. Some editing sites were conserved across ant subfamilies. Moreover, the level of RNA editing of some transcripts was associated with caste differentiation25. Caste differentiation in leaf-cutting ants occurs early in development and produces very distinct queen and worker phenotypes that differ greatly in their behavior and physiology. It remains unknown whether RNA editing also contributes to dynamic, more subtle, changes in behavior of adult social insects.Given the broad spectrum of ADAR activity, it is not surprising that A-to-I RNA editing has been shown to affect physiology and behavior  of genetically highly related “full-sister” worker bees . Although genetic variation among full-sisters is low, they are yet remarkably diverse in terms of body size, physiology, reproductive state, sensory sensitivity, circadian rhythmicity, learning capacity, and behavior . Unlike the caste system of ants, task performance of bumblebee workers is much less rigid over time, and individual bees commonly switch between brood care and foraging activities within the same day28. Here, we study the association between RNA editing and behavioral variation among genetically related bumblebee workers. Combining RNA sequencing and behavioral observations, we show that A-to-I RNA editing in the bumblebee brain is extensive, and weakly correlates with task performance.The bumblebee B. terrestris genome. These predicted proteins are close homologs of the Drosophila ADAR and the mammalian ADAR2. We verified that the available B. terrestris genome (see Methods) does not encode for additional ADAR genes. We also found two ADAR splice variants in the honeybee A. mellifera  edited reads supporting clusters of editing sites, and the second takes advantage of all RNA-seq reads aligned to a genomic location in search of isolated editing events.30. This approach looks for multiple consecutive mismatches of the same type between the reference genome and the RNA-seq data , which are unlikely to result from genomic polymorphisms12. We analyzed RNA-seq data of brain samples originating from 58 individual bumblebee workers, consisting of ~4 billion 2 × 100 bp reads. Among these, we identified 4,040,095 hyper-edited reads harboring 24,314,377 editing events, located at 1,150,394 unique genomic sites that cluster to 51,930 distinct genomic regions (see Methods). Specificity was very high, with noise level lower than 1%  for the control . Furthermore, putative dsRNAs were found for only 1.6% (73/4510) of random regions of similar length. Taken together, the remarkably high signal-to-noise ratio, the depletion of G in the -1 position, and the enrichment of dsRNA structures, strongly support the premise that the sites detected represent genuine A-to-I editing sites.The sequence context of the putative editing sites is consistent with the known preference of ADARs (both ADAR1 and ADAR2) in metazoans33 Fig. . MoreoveB. terrestris brain (65.1) was ~30-fold higher than in the fruit fly (Drosophila melanogaster) head33 (2.57), the honeybee (A. mellifera) brain6 (28.5), the leaf-cutting ant (Acromyrmex echinatior) head25 (28.7), and the human brain33 (26.5)  for .5) Fig. . GeneralAR1 Fig. .While the magnitude of hyper-editing is impressive, one expects much of the functional impact of differential editing to come from sites in coding regions. Among the sites identified by the hyper-editing method, we found 208,089 sites  more highly expressed and more highly edited than those found by the hyper-editing scheme. The genetic background of the four bees was relatively homogenous as expected from their genetic relatedness  of the sites in coding region are non-synonymous (“recoding”) sites. The sites are well-covered (reads per site per sample: average 107 ± 102 (1 std), median 85), and the editing levels (averaged over the 58 bee samples) vary, with an average of 26% ± 27% (1 std) and median of 16%. Editing was overall lower in protein coding sequences (CDS) relative to all other sites  and exhibit similar editing levels in both species36  and conserved in Drosophila melanogaster and other insects .The BtADAR transcript contains two recoding sites  and at this stage, should not be interpreted as a sign for increased editing activity. Yet, we cannot reject the possibility that autoediting modifies the functionality of ADAR and affects the global editing level. This interesting possibility deserves further research.ADAR autoediting in the bumblebee targets different amino acids than in Drosophila and mice (Supplementary Note To study the correlation between editing and task performance (brood care vs. foraging activities), we observed 15 individual worker bees for 3 successive days, and found 8 of them to perform mostly foraging activities (termed “foragers”), and the other 7 to mostly tend brood . We looked at the editing profile for these 15 bees, considering all sites residing in coding regions with a median coverage of 50 reads or more  differential task-related edit  with a small (up to 11%) but statistically significant . Accordingly, the distribution is significantly shifted towards positive values . We therefore, defined a task-specialization index as the number of observed nursing events minus the number of observed foraging trips, and correlated this index with the editing levels observed across the 15 bees for each of the 149 editing events in coding sites. These site-specific measured correlation coefficients are inherently noisy, due to the rather small number of bees, the limited observation time, the rather large error in editing level quantifications and the presumed small effect of task-specialization on editing levels in bumblebees. Indeed, we observe a rather wide distribution of correlation coefficients, none of which is individually significant. Nevertheless, under the null hypothesis of editing activity independent of task performance, one would expect the observed coefficients to be symmetrically distributed around zero, with a similar number of positive and negative values. In contrast, we found that the majority of correlation coefficients are positive; 68.5% [102/149] of the sites in coding sequences, and 68.1% [1779/2611] of all well-covered sites  or for which we manipulated JH levels .χ2 = 19.867, df = 3, p = 0.00018; Fig. P-value = 0.86, data not shown), or in an independent experiment in which we also compared bees developing in queenless or queenright colonies . In addition, brain BtADAR expression did not differ between foragers and nurses in the samples used for the RNA editing analyses , as well as in an independent experiment with bees from different source colonies . Consistent with this analysis, there is a significant difference between age groups  but statistically significantly higher than in both normal age (7-day-old) nurses and normal age (21-day-old) foragers , is surprising considering that we show that bees, like other insects, have lost ADAR1, and express only a single ADAR enzyme11. It is generally accepted44 that the primary role of the widely expressed ADAR1 in vertebrates is to edit endogenous long dsRNAs, in order to prevent an innate immune response triggered by such endogenous transcripts. On the other hand, mammalian ADAR2 is expressed primarily in the brain and arteries45, with a primary role of targeting isolated editing sites within coding sequence. Consistent with this view, Drosophila species that express a single ADAR enzyme homologous to ADAR2, exhibit a relatively low level of hyper-editing , but do show hundreds of recoding sites. Our transcript abundance analyses show that BtADAR expression is enriched in brain tissue. Thus, the comparison of our findings for the bumblebee with other species challenges the common view by providing an example of extensive editing of non-coding sequences in an insect lacking an ADAR1 homolog. These findings suggest that the bumblebee ADAR enzyme (and probably that of other insects) can take on some of the functions performed by ADAR1 in mammals, and partially compensate for the lack of a second editing enzyme. Two-hundred and nineteen of the editing sites reside in protein coding sequences, 164 of which are recoding sites that can potentially modify protein function. Remarkably, some of these sites are similarly edited in other insects and even in cephalopods and mammals. Given these prevailing nonsysnonymous substitutions (recoding) we further compared brain editing and ADAR transcript abundance in bees differing in their behavior, physiology, or social environment.We studied A-to-I RNA editing in naturally behaving animals in an ecologically relevant setting. We found extensive RNA editing in 1.15 million unique genomic sites, mostly within non-coding regions. The extent of hyper-editing in the bumblebee, higher than that reported for brains of humans and other mammals Fig. ; ref. 33ing Fig. ; ref. 3346. Social insects are specifically interesting in this context because they show substantial behavioral, physiological and morphological plasticity that is largely regulated by their social environment. In many species, including bumblebees, considerable phenotypic variation is exhibited by genetically similar individuals, suggesting that genetic variation plays a minor role, if any. The only previous study on RNA editing in social insects compared morphologically distinct castes of the highly social leaf-cutting ant Acromyrmex echinatior25. This study reported a consistent editing pattern for each of the three castes considered , but these sites recoded overall only 34 protein coding genes, only three of these are also edited in other ant species that they tested. The differences between queen and worker ants are considerable, many of which are determined during development, and many are fixed for life. Here, we asked whether RNA editing can be associated with relatively short-term changes during the adult stage of bumblebee workers. We found that brain RNA editing was not influenced by dominance rank (which is associated with aggressiveness) or JH (which regulates fertility and aggression in bumblebees) manipulation, but was correlated with the task the worker bee performs. Similarly to the ants, while the global editing pattern could be correlated with behavior, very few individual sites could be pinpointed as being modified. Admittedly, the association between task performance and RNA editing in the brain is rather weak. However, task-specialization in bumblebees is also relatively weak and flexible, with many workers switching between brood care and foraging activities within the same day. Moreover, we analyzed the whole brain, which is expected to mask significant task-related differences in RNA editing that are restricted to specific brain regions. The notable enrichment for ion channels and other genes involved in neurotransmission among the RNA edited transcripts lend credence to the hypothesis that A-to-I editing in the bumblebee affect neuronal functions.Recoding by RNA editing is a relatively subtle regulation mechanism, enabling dynamic fine-tuning of the transcriptome and proteome in response to changing external, internal, or developmental conditionseag, Shab, Shal, Shaw, and subfamily KQT member 1(XM_003396133.1), calcium-activated potassium channel (slowpoke), voltage-dependent calcium channels, and sodium channels. Some of these ion channels were edited in multiple sites . This is noteworthy because relatively small changes in the ion channel amino acid sequence, such as those created by non-synonymous RNA editing, can have significant influence on the ion channel permeability or regulation. Unlike mammals, insects typically have only a single paralog for each subunit of a given ion channel gene48, and functional diversity is achieved by means of alternative splicing and RNA editing that generate variants differing in their physiology and pharmacological properties49. Indeed, transgenic manipulation analyses in model organisms have shown that editing in orthologs of the ion channels that are edited in the bumblebee affect the channel biophysical and physiological properties, which in turn can alter behavior54. This evidence lends credence to the premise that RNA editing in bumblebee is also functionally significant. We further focus on Shab because there is much detail on the functional outcome of editing its transcript in similar voltage-activated potassium Kv channels54. Additionally, we found 14 different edited sites in the B. terrestris Shab protein, two of which were differentially edited between worker bees performing nursing and foraging activities.We found editing events in many ion channels, including five potassium voltage-gated channel transcripts , each with six transmembrane domains. The first four transmembrane domains form the voltage-sensing domain, while the last two form the pore domain . Incipient colonies contained a queen, approximately ten to fifteen workers and brood at all stages of development. “Mini-Colonies” were obtained about 2 weeks after first worker emergence and contained a queen, ~20 workers and brood at all stages of development. Each colony was housed in wooden observation boxes (30 × 23 × 20cm) with a transparent Plexiglas cover. All the colonies were placed in environmental chambers ) at the Bee Research Facility at the Edmond J. Safra campus of the Hebrew University of Jerusalem, Givat Ram, Jerusalem, Israel. Bees were fed ad libitum with commercial sugar syrup and a processed pollen (fresh pollen mixed with sugar syrup), all of which were purchased from Pollination Services . All observations and treatments were conducted under dim red light, to which bees are not visually sensitive. As an index for body size, we measured the length of the Marginal cell in the front wings28. Bees collected for molecular analyses were flash-frozen in liquid nitrogen (−196 °C), immediately transferred to dry ice, and then stored in an ultra-low freezer (−75 °C) until further analysis.TM). We placed the nesting boxes with the bees in an environmental chamber  under constant darkness, in the Bee Research Facility, and fed them as described above. For the RNA-seq experiments we used workers from 12 colonies; eight for the juvenile hormone and dominance rank experiments; and four for the worker division of labor experiment. In order to minimize genetic variation among treatments, all colonies used for these experiments were headed by full-sister queens that are genetically very similar . These full-sister queens were crossed with unrelated drones and induced to found colonies according to standard industry protocols.For the RNA sequencing experiments, we purchased incipient colonies  from BioBee Biological System . Each incipient colony was placed in a wooden nesting box (21 × 21 × 12 cm) with a front wall and cover made of transparent acrylic plastic (PlexiglasApis mellifera) were collected from freely foraging colonies maintained according to standard beekeeping techniques at our Bee Research Facility (for details see ref. 57). The honeybees were derived from a mixture of European races , which are typical to Israel.Honeybees (Bombus terrestris (version Bter_1.0) and Apis mellifera (version Amel_4.5). Details of the primers and amplicons of these transcripts are summarized in Supplementary Table We used tBLASTX to search for ADAR genes in the genomes of We first separated the heads and bodies on dry ice, and then opened the head capsule cuticle and partially lyophilized the head for 60 min in SpeedVac (Heto DryWinner) to facilitate the dissection of the brain. Following freeze-drying, we opened the head capsule and removed the brains. We performed the dissection on deeply frozen aluminum stage immersed in dry ice such that the tissue remained frozen during the entire procedure. We extracted total brain RNA using the Invisorb Spin Tissue RNA Mini Kit . The RNA was stored in an ultra-low freezer until shipped on dry ice for sequencing at the Carver Biotechnology Center at the University of Illinois at Urbana-Champaign, IL (UIUC).RNA quality was assessed at UIUC with Agilent Bioanalyzer . We prepared RNA-seq libraries with unique barcodes using Illumina’s TruSeq RNA-seq Sample Prep kit. Libraries were quantified by qPCR and pooled before sequencing. We sequenced the pooled libraries on eight lanes for 101 cycles from each end using a TruSeq SBS sequencing kit (version 3) on an Illumina HiSeq2500. We used Casava 1.8.2. to generate Fastq files.For the DNA sequencing we extracted DNA from leg and wing tissues of four bees using the MOBIO power soil DNA extraction kit (MOBIO Laboratories (now Qiagen), Carlsbad, CA). Shotgun genomic libraries were prepared using the Kapa Hyper-Prep kit . Libraries were quantified by qPCR and sequenced for 161 cycles from each end on an Illumina HiSeq2500 using TruSeq sequencing kit version 1. We generated and demultiplexed Fastq files with the bcl2fastq v1.8.4 Conversion Software .Elongation Factor 1a (EF1a) as the housekeeping control gene as it was previously shown not to vary between the brains of bees showing different behaviors . For experiments comparing mRNA levels across different tissues we normalized ADAR mRNA levels relative to two control genes, RPL13 and S16, selected based on a series of preliminary studies  according to the manufacturer’s instructions. Two-hundred nanograms of total RNA was reverse transcribed with (Bioline cat# 27036) to obtain cDNA template. To measure RNA levels, we used real-time quantitative RT-PCR (qPCR) with an ABI’s STEP-ONE Real-Time PCR sequence analyzer using a fast SYBR green detection protocol (ThermoFisher cat# 4385612). For each biological sample, we measured three technical replicates. For brain RNA analyses we used 073 accor60), six newly emerged bees were collected (easily identified based on the lack of yellow pigmentation). We dissected the collected bees on dry ice to obtain the following body parts: head capsule (without the brain), brain, thorax, and abdomen.We housed two incipient colonies in nest boxes that we placed in an environmental chamber. Colonies were connected to the outside with a transparent plastic tube allowing worker bees to forage for nectar and pollen outside; we supplemented the colonies with pollen or nectar as necessary. One week after connection to the outside, when the colonies were self-supported and before the Competition Point , and randomly distributed to queenless (QL) or queenright (QR) treatments. QR bees developed in a colony with a queen, workers (up to 30), and brood at all stages of development; QL workers developed in triplets without a queen. Each QL group was housed in a small wooden cage with transparent walls (10 × 11 × 5 cm). We placed the QR colonies and the QL groups in the same environmental chamber with pollen and nectar ad libitum. We used bees of similar body size within each cage in order to avoid the influence of body size on dominance hierarchy61. Tagging and assignment of bees to treatments occurred over a 1-week period. We collected all the tagged bees 2 weeks after the first tagging day. Thus, the bees in this experiment were 6–12 days of age. Dominance rank was determined as in previous studies . The most dominant bee in each QL group was designated as the ‘α bee’, the second ranked ‘β’, and the most subordinate ‘γ’. We compared QL and QR bees, as well as QL bees differing in the dominance rank. Given that dominance behavior is correlated with ovarian state , we also compared worker bees with developed and undeveloped ovaries. We used the length of the largest terminal oocyte as an index for ovarian state.We performed a series of experiments to test whether brain ADAR transcript abundance is influenced by the social environment or behavior of worker bees. In the first experiment we tested the influence of the queen presence and dominance rank on brain BtADAR mRNA levels. We used four “mini” colonies; two of which were marked for observational purposes, and two were used as “Competition Point’ (refs. 63). All bees were collected on a single day, when they were 4–9 days of age. For the analyses of task influence on ADAR mRNA levels we included only bees which showed either foraging or nursing, but not both.In the second experiment, we tested the influence of task performance on brain BtADAR mRNA levels. We used two incipient colonies that were connected to the outside. Four additional incipient colonies were used as sources for newly emerging bees (“donor colonies”). Starting 1 week from connection to the outside and throughout the following week, newly emerged callow bees were collected from the experimental  and donor colonies. Each collected bee was marked with a colored plastic number tag  and immediately introduced randomly into one of the experimental colonies. Two days after tagging we started sessions of behavioral observations. Foraging observations were performed at peak activity times of the day, i.e., 4 h in the morning (06:00 to 10:00 a.m.), and two in the afternoon (4:00 to 6:00 p.m.). Foraging activity was recorded when a bee left the colony in a direct line of flight. This flight pattern is distinct from ‘orientation flights’ that are characterized by circular maneuvers in front of the hive entrance. We recorded the number tags of foraging bees returning to the colony with conspicuously visible pollen loads on their hind legs. We performed additional observations inside the nest box during a session between 11:00 a.m. and 1:00 p.m. in which we recorded brood-feeding behavior as a proxy for nursing behavior. Observations and collections of bees for RNA analyses were carried out when the colonies were already self-supported and before the ‘Apis mellifera) colonies that were maintained using standard beekeeping procedures in our Bee Research Facility. In order to obtain newly emerged worker honeybees, we removed three honeycomb frames with sealed brood, each from a different field source colony. We brushed all the adult bees, placed each frame in a wooden box, and transferred it to an incubator (32 ± 1 °C and 55 ± 5% humidity). Bees emerging over the night were color marked (a distinct color for each source colony), and reintroduced back into their source colony. Given that task activity in honeybees relates to age, we repeated this procedure two times: first tagging bees for foraging observations, and 2 weeks later, for nursing observations. Three weeks from the first tagging day, and 1 week following the second tagging, we collected the tagged bees. Three-week-old bees captured when returning to the hive with pollen were classified as ‘foragers’; 7-day-old bees inserting their heads into brood cells were classified as ‘nurses’. On the same day, we also collected newly emerged bees.For comparison we also tested the influences of age and task performance on brain AmADAR levels in honeybees, in which the division of labor relates to age. We used two honeybees  in sites with genomic adenosines (A), RNA sequences with multiple editing sites are widely different from their corresponding DNA. The computational screen overcomes this obstacle by masking potential editing sites in unaligned reads30. As an input we used 58 RNA-seq datasets (paired-end 100 bp reads) and the Bombus terrestris genome assembly version Bter1.0. As the B. terrestris genome draft is still rather fragmented two paired-end reads may be mapped to different scaffolds, alignment that would be considered improper. We thus treated each paired-end read as independent single-end reads. We searched for high-quality sequencing reads (Phred score ≥ 30) containing at least five A-to-G mismatches per 100 bp sequence, and for which the A-to-G mismatches comprise > 60% of the total number of DNA–RNA mismatches. To validate the specificity of the pipeline, we used the same procedure to search for all other possible types of mismatches . We could not distinguish between a given mismatch and its complementary one, since the RNA-seq reads could be either sense or antisense (non-stranded library). Thus, although there are 12 possible single-nucleotide mismatches, we report results for only six categories of editing events: A-to-G, G-to-A, A-to-T, A-to-C, G-to-C, and C-to-A and each category represents both the forward and complementary mismatch. For genes and coding region annotations we downloaded the NCBI RefSeq annotations from the BeeBase site (http://hymenopteragenome.org/beebase) on 7 May, 2014.Putative hyper-editing sites were identified as described in Porath et al.30. Overlapping clusters were merged to create a hyper-edited region, maintaining a maximum distance of 20 bp between clusters. The main targets of ADAR enzymes are long dsRNA structures64. To test whether the hyper-edited regions occur within such structures, we aligned the DNA sequence of each hyper-edited region (found in one representative sample) to the genomic sequence spanning 2 kbp upstream and 2 kbp downstream of the region, looking for reverse-complement matches that may indicate a possible dsRNA structure. As a control, we looked for same-strand matches . As an additional control, we repeated the search for random regions of similar lengths. For these alignments we used bl2seq65 with parameters -F F -W 7-r 2, keeping only alignments with at least 65% complementarity along 60% of the hyper-edited region length.Hyper-edited clusters were defined as the genomic sequence between the first to the last A-to-G mismatch in each hyper-edited readB. terrestris genome with the STAR 2-pass aligner, using the default parameters66. We then used Picard (http://broadinstitute.github.io/picard/) and GATK67 software packages with the following tools ‘picard MarkDuplicates’, ‘picard ReorderSam’, ‘GATK SplitNCigarReads’, ‘GATK RealignerTargetCreator’, ‘GATK IndelRealigner’, and ‘Picard FixMateInformation’. We further used REDItools68 with parameters: -v 1 -n 0.01 -c 1 -T 6–6 to estimate the level of A-to-I editing. Editing index over sites was measured as the total number of Gs divided by the total number of Gs + As69.To estimate editing levels for each hyper-edited site, we first aligned the RNA-seq datasets to the 70 with default parameters and STAR 2-pass66 to align the DNA-seq, and the RNA-seq to the published B. terrestris genome. Alignments were improved by picard (http://broadinstitute.github.io/picard/) and GATK67 tools. Detection of putative editing sites was done by MuTect version 1.1.4 (using default parameters), an algorithm that was developed for identifying somatic mutations in cancer samples by comparing the normal genomic DNA sequence with the matched tumor DNA sequence of the same individual34. As the “normal” genomic reference sequence we used the consensus sequence generated from aligning the four DNA-seq datasets that we generated. The RNA sequence of each bee was entered in the “tumor sample” option of the MuTect program. We have also attempted detection by the well-used REDItools DNA–RNA tool68, with different combinations of parameters. However, the resulting A-to-G fraction for all genomic sites was only 40–43%, substantially lower than the 64% obtained by MuTect.In a separate set of analyses, we detected putative RNA editing sites by comparing DNA and RNA sequences. To get a DNA genomic reference for our bumblebee population, we sequenced the genome of four representative bees (out of the 58 for which we sequenced the brain transcriptome). We used BWADrosophila flies, we aligned the bee protein sequences harboring recoding sites to the sequences of the proteins harboring recoding sites previously identified as conserved across Drosophila species35. We identified seven sites conserved between flies and bees, where recoding occurs at the homologous amino acid in both species, resulting in the same non-synonymous alteration. Editing levels of those conserved sites in the fly were computed using data from Duan et al.36.To detect recoding sites conserved between bumblebees and 46, one bee was removed for each two bees introduced60. We marked and introduced a total of 35 bees into each free-foraging colony over 2 successive days. When the tagged bees were 3–4 days of age, we started behavioral observations that continued over 2 days. We observed the colonies for 3 h in the morning (07:00–10:00 a.m.), and 2 h during late afternoon . We focused on foraging and brood care activities because these behaviors are ubiquitous, well characterized, crucial for the division of labor and colony performance, and easy to identify. Each observation session was divided between 1 h in which we recorded foraging activity next to the nest entrance, and 1 h in which we recorded larval feeding behavior inside the room. Larval feeding behavior was composed of a sequence of acts starting with the worker bee opening a brood cell, inserting her mouthparts into the open cell, and regurgitating food. Food regurgitation is characterized by the typical contraction of the abdomen and takes a few seconds56. RNA was sequenced from four foragers and four nurses from each colony. The foragers were defined as bees that were observed conducting at least three foraging trips and were not observed feeding larvae (with the exception of one forager that performed two feeding behaviors and eight foraging trips). Nurses were defined as bees observed performing at least three feeding acts, and were not seen foraging at all. We flash freezed the focal bees in liquid nitrogen following the second day of observations when they were 5 to 6 days of age. We transferred the frozen bees to marked tubes immersed in dry ice and stored them in an ultra-low freezer until further analysis.We performed three experiments testing the influence of behavior or physiological state on brain RNA editing. In Experiment 1 we tested the effect of task performance focusing on workers differing in the levels of brood care (“nursing”) and foraging activities. We placed four colonies in an environmental control room . Two of the colonies were connected to the outside with a 1 m long clear plastic tube, and the remaining two were used as donor colonies. We collected newly emerged bees from the free-foraging colonies, tagged them with individual number tags (as above), and returned them to their mother colony for 2 days. We similarly collected and tagged bees from the two donor colonies, and introduced them randomly into the two foraging colonies. In order to avoid increasing the total numbers of bees in the colonies, which can affect colony developmentχ2-tests with Benjamini–Hochberg multiple-testing correction, setting the false discovery rate to 5%.In Experiment 2 we compared queenless  workers differing in dominance rank (dominant vs. subordinate). In Experiment 3 we compared bees with experimentally manipulated JH levels. The details of Experiments 2 and 3 are summarized in the Supplementary Methods. For each individual RNA-seq dataset, editing levels in 219 sites located within protein coding sequences were quantified and compared. We limited the analysis to sites with a median coverage of at least 50 reads per sample. We used Further information on experimental design is available in the Supplementary InformationPeer Review FileReporting SummaryDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5Supplementary Data 6Supplementary Data 7Supplementary Data 8Supplementary Data 9Source Data