Using MAP-C, we show that inducible interchromosomal pairing between HAS1pr-TDA1pr alleles in saturated cultures of Saccharomyces yeast is mediated by three transcription factors, Leu3, Sdd4 (Ypr022c), and Rgt1. The coincident, combined binding of all three factors is strongest at the HAS1pr-TDA1pr locus and is also specific to saturated conditions. We applied MAP-C to further explore the biochemical mechanism of these contacts, and find they require the structured regulatory domain of Rgt1, but no known interaction partners of Rgt1. Altogether, our results demonstrate MAP-C as a powerful method for dissecting the mechanistic basis of chromosome conformation.The genome forms specific three-dimensional contacts in response to cellular or environmental conditions. However, it remains largely unknown which proteins specify and mediate such contacts. Here we describe an assay, MAP-C , that simultaneously characterizes the effects of hundreds of  Inside cells, genetic information is stored within molecules of DNA that are folded into three-dimensional structures known as chromosomes. Each fold in a chromosome forms when two points on a single DNA molecule link together to make a loop. DNA in two different chromosomes can also form links with each other (known as “contacts”). Many cells contain two copies of every chromosome and these copies are often able to make contacts with each other.HAS1pr-TDA1pr region stick together. This may help the budding yeast cells switch on genes that are needed to make use of alternative sources of food.DNA loops and contacts can change in response to the environment and this may help cells switch the right genes on and off at specific times. For example, in budding yeast cells that have used up most of their preferred food source – a sugar called glucose – the two copies of a region of DNA known as the Cells contain hundreds of proteins called transcription factors that can bind to specific locations on DNA and can also stick to each other. These proteins are thought to be responsible for anchoring bridges between the DNA at most loops and contacts. One way to find out which transcription factors form specific DNA loops and contacts is to generate many different genetic mutations in the DNA and identify precisely which mutations disrupt the links. However, current methods can only test one mutation at a time, so it remains unclear how and why many segments of DNA stick together.HAS1pr-TDA1pr region from forming contacts. This revealed that three transcription factors – Leu3, Sdd4, and Rgt1 – bridge contacts between the two copies of HAS1pr-TDA1pr.Now, Kim et al. have developed a new method known as MAP-C to test how hundreds of mutations in budding yeast affect a particular DNA contact, in a single experiment. The MAP-C method was used to test which mutations within either the DNA segment involved in the contact, or in genes encoding transcription factors, prevent copies of the Mutations that disrupt the three-dimensional structure of chromosomes can cause cancer, developmental disorders and other diseases. The MAP-C method will allow researchers to better understand which transcription factors control how DNA is folded inside the cell, and which mutations change this folding. Saccharomyces cerevisiae, a well-studied model of genome conformation, genes targeted to nuclear pores are activated  are attractive candidates for orchestrating such dynamic changes in chromatin conformation, given their site-specific DNA binding and changes in abundance or activity in response to differentiation and cellular signals . For manAmong chromosomal contacts and loops, interchromosomal contacts are less well-understood. This is in part due to the relative paucity of interchromosomal contacts in Hi-C and other 3C (chromosome conformation capture) data, which results from their greater contact distance  and chroMany known mechanisms for establishing 3D chromosome conformation may act on both intrachromosomal loops and interchromosomal contacts. The emerging consensus model for such DNA-DNA interactions involves loop extrusion by cohesin and other Structural Maintenance of Chromosomes (SMC) factors, which is thought to primarily mediate intrachromosomal loops . HoweverDrosophila (Mitotic (or somatic) homologous chromosome pairing is the preferential association of homologous pairs of loci in mitotically dividing cells. Homolog pairing occurs along the length of the genome in osophila , but is osophila . Fluoresosophila , but theosophila , in conjosophila . Howeverosophila  remain uHAS1pr-TDA1pr locus in diploid Saccharomyces yeasts , to characterize the molecular components mediating HAS1pr-TDA1pr pairing. We first perform saturating mutagenesis of the regulatory region that mediates the interchromosomal pairing (cis MAP-C) to identify sequence motifs required for pairing that potentially correspond to TF binding sites. We then test the effects of knocking out over one hundred TFs (trans MAP-C), and confirm that three—Leu3, Sdd4 (Ypr022c), and Rgt1—are necessary for inducible interchromosomal pairing. We verify their binding by chromatin immunoprecipitation, and find that HAS1pr-TDA1pr exhibits the strongest coincident, combined binding by all three factors across the genome in a condition-specific manner. We further use trans MAP-C to interrogate how interaction partners and domains of Rgt1 regulate pairing. Finally, we make an initial attempt to characterize the functional consequences of HAS1pr-TDA1pr pairing. Taken together, our results demonstrate how a combination of TFs can mediate inducible interchromosomal pairing. Furthermore, our study shows the utility of a pooled mutant approach to studying both the cis and trans dependencies of chromosome conformation.Here we describe a method that enables the simultaneous testing of hundreds of e.g. mutations) are needed. However, despite numerous advances in chromosome conformation capture (3C) technology over the last two decades  to discover this homolog pairing interaction, and furthermore identified a 1,038 bp noncoding region that was necessary and sufficient for pairing , in S. cerevisiae x S. uvarum hybrid yeast. We then performed cis MAP-C for pairing of the modified locus with the S. uvarum copy of HAS1pr-TDA1pr on a saturated culture of the pool, in two replicates . To confirm that this pattern of enrichment is specific to HAS1pr-TDA1pr homolog pairing rather than underlying all of its chromosomal contacts, we repeated the assay with a pair of primers amplifying a ~ 10 kb intrachromosomal contact , would induce pairing with the native HAS1pr-TDA1pr locus in saturated cultures of haploid S. cerevisiae  containing the minimal pairing region, with an average of 1.49 substitutions (range 0–14) per template . We inse pairing . We insp pairing  and 3. G pairing , the thi pairing , and the pairing . These m pairing , and all pairing . None of pairing , and all pairing . Interescis MAP-C, we identified a ~ 150 bp subsequence of HAS1pr-TDA1pr sufficient for pairing, containing five required TF motif occurrences. If these TF motifs are together sufficient for pairing, we would expect that 1) they are only observed in a cluster in the minimal pairing region and not elsewhere in the HAS1pr-TDA1pr, and 2) they are present in a cluster in the S. uvarum copy of this region, and potentially other Saccharomyces as well. Indeed, these motifs are clustered together only in the central region of HAS1pr-TDA1pr; remarkably, this pattern holds across all Saccharomyces species . With trans MAP-C, each mutation is uniquely associated with a short barcode sequence near a region involved in the chromosomal contact of interest, which is then assayed by 3C. Mutations that affect pairing frequency modulate the abundance of their corresponding barcodes in the 3C products, which can be readily quantified by deep sequencing. To study the trans requirements of HAS1pr-TDA1pr pairing, we ectopically integrated barcoded versions of the minimal pairing region, wherein the barcode identifies which TF is knocked out, and assayed their capacity to pair with the native copy of HAS1pr-TDA1pr , six had no substantial difference in pairing strength compared to their genomic neighbors ; however, its binding motif is not present in the HAS1pr-TDA1pr region, suggesting an indirect role. Two other knockouts, VHR1 and CBF1, appeared to also decrease pairing, but were at low abundances and might reflect noise  and YGR067C, were excluded in the initial screens due to their lack of annotations. Therefore, we repeated our fixed-locus TF knockout screen with a limited set of genes, including the two putative TFs and additional replicates for the hits MOT3, VHR1, and CBF1. We found that SDD4 is indeed required for pairing, suggesting that it is the trans-acting factor that binds the CCCCAC motif  and the DNA binding motifs of trans factors required for pairing (identified by trans MAP-C) suggests that Leu3, Sdd4, and Rgt1 bind the HAS1pr-TDA1pr to mediate pairing. To test this hypothesis, we performed chromatin immunoprecipitation (ChIP) for the tandem affinity purification (TAP) tagged versions of Leu3, Sdd4, and Rgt1  no loci other than HAS1pr-TDA1pr should have all three TFs bound on both the S. cerevisiae and S. uvarum copies, or 2) other loci that do have all three TFs bound should also exhibit pairing. Because DNA binding data was only available for the S. cerevisiae genome, we tested the first possibility by scanning the S. cerevisiae and S. uvarum genomes for clusters of the three motifs using permissive thresholds for motif matches and allowing up to 200 bp between motifs , and then assessed ChIP-seq data at these clusters. The promoters of four genes, TDA1 but also HXT3, YKR075C, and SKS1, harbored a motif cluster containing all three motifs in both S. cerevisiae and S. uvarum ; whether this plays a role in pairing is unclear. We infer that the combinatorial binding of Leu3, Sdd4, and Rgt1 on both homologs specifies inducible homolog pairing in saturated cultures.We next explored whether the combination of Leu3, Sdd4, and Rgt1 binding explains the uniqueness of . uvarum . We alsonditions , even whrevisiae . FurtherTDA1pr exhibits the strongest combinatorial TF binding, we wondered whether any other motif clusters pair inducibly like the HAS1pr-TDA1pr locus, albeit perhaps more weakly. To address this question, we leveraged our previously published Hi-C datasets  in saturated culture conditions  between HAS1pr-TDA1pr and HXT3pr. To test this possibility, we extracted the interaction frequencies among the four loci (i.e. two pairs) from our Hi-C data. In all saturated culture datasets where homolog pairing was present, inter-locus interactions were substantially weaker and similar in frequency to those in non-pairing conditions   . RGT1 anconstant . These r3EV estradiol induction system  and performed trans MAP-C to test the pairing function of the Rgt1 mutants , suggesting that the Q/N-rich domain inhibits pairing. Both the S88A and S758A mutations had a weak decrease in pairing, indicating that although these mutations are capable of disrupting Rgt1 activator function and intramolecular interactions  remains outstanding. Now that we have identified the precise DNA base pairs and proteins involved in HAS1pr-TDA1pr pairing, we are equipped to test whether Leu3, Sdd4, and Rgt1 play a role in transcription at HAS1 or TDA1 in saturated cultures. Furthermore, we can employ S. cerevisiae x S. uvarum hybrids, in which we can discriminate the two homologous copies of each gene, in order to distinguish the effects on transcription of disrupting the pairing DNA sequence in cis (presumably through impaired cis regulation) vs. in trans (presumably through impaired pairing). To this end, we generated mutant hybrid strains carrying a wild-type copy of S. uvarum HAS1pr-TDA1pr and a copy of S. cerevisiae HAS1pr-TDA1pr with either a wild-type (WT) genotype, mutated Leu3 binding site (leu3), or two mutated Rgt1 sites (rgt1 × 2)  . We thengt1 × 2) , as oppogt1 × 2) ; howevernditions , but hadany gene .rgt1∆  as well as the factors that are necessary to mediate the contact (trans MAP-C). Here we applied both versions of MAP-C to dissect the mechanism of inducible interchromosomal pairing between HAS1pr-TDA1pr alleles in budding yeast. Using a combination of gain-of-function and loss-of-function screens, we demonstrate that a trio of transcription factors—Leu3, Sdd4, and Rgt1—mediates pairing between clusters of binding sites.In summary, we developed Mutation Analysis in Pools by Chromosome conformation capture (MAP-C), a method to simultaneously test hundreds of mutations for their effects on a chromosomal contact of interest. MAP-C can be used to identify the precise sequences that are necessary for a contact  itsand INO1 . Howeverand INO1 , was alsand INO1 . More stHAS1pr-TDA1pr pairing unique? Based on Hi-C, we previously found HAS1pr-TDA1pr to exhibit the strongest interchromosomal interactions in saturated cultures of S. cerevisiae x S. uvarum hybrids, excluding centromeres, telomeres, and the chromosomes carrying the rDNA arrays  the introduced mutations or associated barcodes must be included in the 3C PCR product, and 2) the region of interest should not be digested by the restriction enzyme. As we have implemented this approach, the region targeted by saturation mutagenesis was limited to < 250 bp to allow for Illumina sequencing, but this could be extended using either barcode association, similar to our trans knockout screens, or long-read sequencing methods. Also, we focused on a single pair-wise interaction, but it is also possible to assay a mutant pool for multiple interactions, by using multiple primer pairs. MAP-C should be applicable to intrachromosomal contacts as well as interchromosomal ones, albeit with a potentially higher background of nonspecific contacts. It would be interesting to apply MAP-C to dissect the cis and trans regulators of enhancer-promoter loops in mammals, and thereby distinguish the contributions of cohesin and CTCF  to OD600 = 0.5 cultures grown in YPD  and grown for 2 hr. Galactose induction was performed by growth in synthetic complete medium (without uracil for selection for overexpression plasmid) with 2% v/v raffinose to OD600 = 0.75 followed by addition of 2% galactose and subsequent growth for 1.5 hr. For comparison, yeast were grown in synthetic complete medium with or without uracil with 2% glucose or 2% raffinose. S. cerevisiae x S. uvarum hybrids were generated by standard mating and auxotrophic or drug selection procedures. Yeast transformations were performed using a modified Gietz LiAc method  array-synthesized oligonucleotide pool . Transformants  were selected on G418 medium for a total of at least 4 days (including two rounds of scraping and replating a portion onto a fresh plate), and then used to inoculate a 50 ml culture in YPD .All 178 bp subsequences of the intergenic region between the TTCACCGCCTGCTATCATCC and GAATCGGCGGAATAACCTAACACG were used in a PCR reaction using the Agilent GeneMorph II kit, using as template 0.1 ng of a PCR product generated with the same primers from S. cerevisiae genomic DNA. The error-prone PCR products were then cloned, transformed, and selected as described above. The library contained ~ 63,000 transformants.The primers e.g. A->C) was included once at each position and so that no DpnII restriction sites (GATC) were created. These sequences were synthesized on an array (sequences included in For each set of 3 consecutive base-pairs between positions 532–675 (inclusive), we randomly chose three trinucleotide substitutions such that among them, each nucleotide change  deletions between amino acids 91 and 1040; we successfully created 87 of the 96 attempted deletions as plasmids. The wild-type RGT1-containing plasmid was amplified in two pieces, each using one primer in the ampicillin resistance gene and one within RGT1 so that the resulting pieces each contain homology for the other. These pieces were combined using NEBuilder HiFi, and then transformed into E. coli, extracted, and verified for the deletion junction by Sanger sequencing. Each deletion plasmid was amplified with the same primers used to create the fixed-locus knockout constructs, with a different chosen barcode for each deletion. These PCR products were pooled and transformed en masse into the rgt1 deletion strain from the MATa yeast deletion collection with the SGA reporter, in two replicates. Each pool was selected on synthetic complete medium without uracil including G418 and processed as described above.The RGT1-containing plasmid were amplified and cloned as for the 10 amino acid deletion scan of Rgt1, and then transformed using mixtures of 14 different barcodes per desired mutation and Sanger sequenced as in the Rgt1 interactor knockout screen. Five distinct barcoded versions of each mutant were pooled and processed as above, with positive and negative control strains included.Tested mutations included deletion of the zinc finger domain (amino acids 32–90), Q/N-rich domain (amino acids 293–459), C-terminal domain (amino acids 461–1163), and S88A (TCG->GCG) and S758A (TCC->GCC). Mutant versions of the wild-type cis experiments and galactose induction, 400U EcoRI-HF for trans experiments, and 100U NlaIII for estradiol inductions). The restriction digest was heat-inactivated at 65C for 20 min in the presence of 1.3% SDS, and then chilled on ice and added to a dilute ligation reaction in 4 ml volume with 1% Triton X-100, 1x T4 DNA Ligase Buffer (NEB), and 10,000U of T4 DNA ligase (NEB) and incubated at room temperature for 4 hr. The ligation products were reverse-crosslinked with proteinase K at 65C overnight, and then purified by phenol-chloroform extraction followed by clean-up on a Zymo DNA Clean and Concentrator-5 column . The resulting 3C DNA was quantified using a Qubit. Each technical replicate was processed separately beginning with cell lysis.Cells were crosslinked by addition of 37% formaldehyde to a final concentration of 1% (v/v) and incubation at room temperature for 20 min, quenched by addition of 2.5M glycine to a final concentration of 150 mM and incubation at room temperature for 5 min, and then washed in 1x Tris-buffered saline (TBS) and stored as a pellet at −80C in aliquots of 50–100 µl dry pellets. Cells were lysed by vortexing in lysis buffer ) with 500 µm acid-washed glass beads for 6 cycles of 2 min, with 2 min on ice between cycles. The lysate was collected by puncturing the bottom of each tube and then centrifuging the tube, stacked on top of an empty tube. The lysate was then washed in lysis buffer, then TBS, and finally resuspended in 10 mM Tris pH 8.0 to a volume of ~ 200 µl per 25 µl of starting dry pellet volume. A single 200 µl aliquot of lysate was then precleared by addition of 0.2% SDS and incubation at 65C for 10 min, cooled on ice, quenched by addition of 1% Triton X-100 (v/v), and then digested overnight with at least 100U of restriction enzyme  or a control off-target chromosomal contact (for off-target library), for 24–32 cycles. Genomic libraries were prepared by amplification of up to 4 reactions of 50 ng of either 3C DNA or genomic DNA using primers flanking the targeted mutations or barcodes, for 17–22 cycles. Reactions for each replicate were pooled, purified by Ampure XP beads , and then re-amplified with primers flanking the mutagenized or barcode region and including sequencing adapter sequences for 5–8 cycles, and then again with primers adding sample indices and Illumina flow-cell adapters for 6–9 cycles. All reactions were prepared with KAPA HiFi HotStart ReadyMix  with recommended thermocycling conditions, and included 0.5x SYBR Green I to monitor amplification by quantitative PCR and minimize the number of PCR cycles. The final libraries were sequenced on an Illumina MiSeq or Nextseq 500  using paired-end sequencing. See trans knockout experiments, in which only read one was used. These reads were then trimmed of the first 4 bp (corresponding to a randomized region for Illumina clustering purposes) and mapped using Bowtie 2 .Reads were mapped to all 192 possible barcode sequences, and the normalized fraction of reads mapping to a given barcode with a MAPQ ≥ 20 in the 3C library compared to the genomic control was calculated for each replicate.3C DNA was amplified using the same conditions as MAP-C libraries, in three replicates per primer pair. The pairing 3C products were normalized to the off-target  3C products, assuming 2-fold amplification per cycle.600 = 0.7–0.8) or 12 ml of saturated cultures of TAP-tagged strains . Cq values were calculated using the Bio-Rad CFX Manager 3.1 software, using the single threshold mode for Cq calculation and baseline subtracted curve fitting. Primer efficiencies were calculated using a 5-fold dilution series of genomic DNA from ChIP-seq libraries were prepared using Swift Accel-NGS 2S Plus  dual-indexed kits using 10 µl of IP samples or 1 ng of input samples, with 9 cycles of PCR for input samples and 12–15 cycles for IP samples. Libraries were sequenced to ~ 2.5–6 million read pairs per sample using 2 × 37 bp reads on an Illumina NextSeq 500.S. cerevisiae reference genome using Bowtie 2 (https://github.com/taoliu/MACS) (Sequencing reads were first pre-processed using cutadapt : reads wBowtie 2  with theiu/MACS) . Fold eniu/MACS) .Yeast strains were grown overnight in YPD in biological triplicate , pelleted and then stored at −80C. RNA was purified using acid phenol extraction, and then treated with Turbo DNase and purified with a Qiagen RNeasy Mini kit . Illumina libraries were prepared from 800 to 900 ng of total RNA, using the Illumina Truseq RNA Library Prep kit v2 (for TF binding site mutants) or the Illumina Truseq Stranded mRNA Library Prep kit (for TF knockouts). Libraries were sequenced to ~ 11–13 million read pairs per sample using 2 × 37 bp reads on an Illumina NextSeq 500.Reads were preprocessed and mapped as with the ChIP-seq libraries, but with the -X 500 option for Bowtie 2. Read pairs in which both reads had a mapping quality score of at least 30 were overlapped with annotated genes using HTSeq . Global 20 (13 µl reaction with 1 µl of 50 µM oligo(dT), 1 µl 10 mM dNTP mix) by incubating at 65C for 5 min and then on ice for 1 min. Reverse transcription was performed with SuperScript IV by adding 4 µl of 5x SuperScript IV buffer, 1 µl SUPERase In RNase inhibitor, 1 µl 100 mM DTT, and 1 µl of SuperScript IV enzyme, and then incubating at 50C for 10 min and then 80C for 10 min. For qPCRs, 2.5 µl of each reverse transcription reaction was used for each 25 µl PCR, using KAPA Robust 2G HotStart ReadyMix with standard cycling conditions  for 40 cycles on a Bio-Rad C1000 Touch thermal cycler with a CFX96 Real-Time System. Cq values were calculated using the Bio-Rad CFX Manager 3.1 software, using the regression mode for Cq calculation and baseline subtracted curve fitting. Primer efficiencies were calculated using a 5-fold dilution series of genomic DNA from S. cerevisiae BY4741 starting from 10 ng. Primers were specific to S. cerevisiae (at least two substitutions to S. uvarum). See Yeast were grown as described above in Yeast strains and culture, and then pelleted and stored at −80C. RNA was purified using acid phenol extraction, and then treated with Turbo DNase and purified with a Qiagen RNeasy Mini kit. For each sample, 1 ug of total RNA was annealed to oligo(dT)Predicted intrinsic disorder was calculated using IUPred2 long disorder . PredictSystematic scans of motifs were performed using YeTFaSCo  with theS. cerevisiae (version R64.2.1) and S. uvarum genomes .Code used to analyze data and generate figures are available at All sequencing data have been deposited in the Gene Expression Omnibus (GEO) under accession number GSE118118. Hi-C data from  In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.eLife. Your article has been reviewed by has been reviewed by three peer reviewers, including Jeannie Lee as the Reviewing Editor and Reviewer #3, and the evaluation has been overseen by a Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Peter J Fraser (Reviewer #1); Hsueh ping Chu (Reviewer #2).Thank you for submitting your article "A combination of transcription factors mediates inducible interchromosomal contacts" for consideration by We find your study to be well-conducted and the findings to be broadly interesting to the community.Summary:The authors present the development of MAP-C  to study the regulation of chromosomal interactions through mutagenesis. Two flavors of the methods allow (i) identifying the local sequence required for interaction by cis MAP-C (ii) testing the requirement of other factors or sequences for interaction by trans MAP-C. The authors demonstrate the power of the method by applying it to the two HAS1pr-TDA1pr alleles, which they recently demonstrated to pair under defined biological conditions. The authors went on to identify the local genomic sequence and three transcription factors essential for allelic pairing of HAS1pr-TDA1pr. The authors suggest that a regulated interaction between the three factors controls both the specificity and physiological regulation of HAS1pr-TDA1pr pairing. The tools described will facilitate the study of pairing, an emerging epigenetic phenomenon that is still poorly understood.Essential revisions:Please focus on several of the key points raised by reviewers:1) The identification of 3 TFs for pairing suggests that transcriptional activity or RNA produced at the locus might play a role in the trans interaction. The authors should characterize the transcriptional status of the locus and its changes in the individual TFs knockouts and upon pairing.2) While the C-terminal region of Rgt1 seems required for pairing, the suggestion that it is regulated by phosphorylation is a stretch. We recommend testing the effect of mutating the regulatory S88 and S758 residues.3) Reviewer 1 would like to see either data or discussion of the biological purpose of localized pairing and why it is inducible. This is a fair question and I (reviewer 3) am also interested in the thinking behind it.4) We would require evidence that the factors actually bind the pairing locus. ChIP-seq of these three transcription factors to verify the binding of Leu3, Sdd4 and Rgt1 at the minimal pairing region would strengthen the paper.5) Data to address the differences in binding level of the three TFs at the minimal pairing region under the two different growth conditions. ChIP-seq is again recommended.6) As mentioned by reviewers below, there are other major cases of pairing in various organisms, including mammals. In the revision, please be sure to properly cite those studies to put the current study in perspective.7) Reviewer 2 also asked if homologous pairing at HAS1pr-TDA1pr regulates nearby gene expression in saturated cultures. If data are available to address this, please include them.Reviewer #1:eLife . In their previous publication, this group used diverged haploid yeast as a model (S. cerevisiae ×S. uvarum) and provided a global view of chromosome conformation in hybrid diploid yeasts. They found localized increases in homologous interactions between the HAS1-TAD1 alleles specifically under galactose induction and saturated growth conditions.This research is a follow-up study of this group's previous studies published in S. cerevisiae HAS1pr-TDA1pr locus with a library containing each of 861 tiling 178 bp subsequences of the 1,038 bp region. Following identification of the minimal pairing region, they used error-prone PCR to obtain a base-pair resolution map of the DNA sequences necessary for homologous pairing, and identified unique clusters of sequence motifs in the minimal pairing region, which correspond to specific TFs binding motifs. Next, they used trans MAP-C, to test trans knockouts of the TF candidates identified in the cis MAP-C and confirmed that three TFs  mediate the interchromosomal pairing. The method variants are interesting and useful, however there are a few points that could be addressed that could potentially make the manuscript more interesting for the readership of eLife.The current study aims to understand the regulatory mechanism underlying homologous pairing between the HAS1-TAD1 locus in diploid yeast. To identify the specific sequence elements that are necessary and sufficient to maintain homolog pairing between the HAS1-TAD1 alleles, the authors developed a method called MAP-C . The cis MAP-C method involves analyzing populations of cells with multiple mutations of specific regions suspected to mediate interchromosomal pairing followed by amplification of 3C products and sequencing. To find the minimal subsequence of the 1,038 bp HAS1pr-TDA1pr region that is essential for the homologous interactions, the authors replaced the native 1) Obviously, the homologous pairing between HAS1-TAD1 alleles is a localized event, and the pairing is inducible  and limited into a 1kb intragenic regions on chromosome XIII. However, this study did not attempt to answer key biological questions: what's the biological purpose or meaning of this localized pairing? Why it's inducible? There are no experimental data or discussions in the manuscript. This is an important question that could greatly improve the interest level and relevance of the findings and the manuscript.2) In the Introduction, the author described that "a motif match is insufficient to fully predict TF binding", which is true. Later on, this study identified binding of the three transcription factors binding to the "minimal pairing region" which are necessary and essential for homologous pairing between HAS1-TAD1 alleles. However, most of the Results section and Conclusion were based on motif scanning, motif searching, and mutations in this region. This study missed some very important experiments: ChIP-seq of these three transcription factors to verify the binding of Leu3, Sdd4 and Rgt1 at the minimal pairing region.3) Since the homologous pairing between HAS1-TAD1 alleles is significantly higher at growth saturation than at standard growth conditions, and this study also identified that three transcription factors  regulate this homolog pairing. What's the binding level of these three TFs at the minimal pairing region under the two different growth conditions? ChIP-seq of these three transcription factors under the two different growth conditions is very important to verify and solidify the conclusion.4) The authors claimed that HAS1pr-TDA1pr pairing is regulated by the expression level of TFs, particularly RGT1. In fact, using the mRNA expression level of TFs is not convincing way to convey this idea. Because increasing at mRNA level does not necessarily means increasing TFs' bindings at chromatin, DNA, or a specific loci (in the current study). The best experiment here would again be ChIP-seq of the three TFs to show differential binding.5) Last part of the Results section, there is no data to support that the "pairing domains" of Rgt1 needs to be phosphorylated for the purpose of homologous pairing. This is speculation without supporting data. If the authors believe that phosphorylation of the "pairing domains" of Rgt1 is one of the regulatory mechanisms of Rgt1 function, then they need to show that under growth saturation conditions, the "pairing domain" of Rgt1 is indeed phosphorylated.6) In the Introduction the authors claim that:"Although interchromosomal contacts, such as those between actively transcribed genes or homolog pairing in mitotically dividing yeast, have been observed by microscopy , many are not detectable using 3C (chromosome conformation capture) technologies. Some have argued that these contacts might be a result of fluorescent in situ hybridization (FISH) artifacts  or the insertion of binding site arrays often used for live-cell imaging . A more detailed understanding of the molecular components of interchromosomal contacts is needed to resolve this ambiguity."Drosophila and mouse cells. There are in fact many publications that have shown a correspondence between interchromosomal associations between transcribed genes by FISH and by 3C in mammalian cells going back over a decade, and more recently in live yeast cells with 3C technology verification. As always authors like to suggest novelty in order to get published, but there is no need to ignore the work of many other colleagues and many other publications. The author's method alone and its findings are powerful results. The authors should read and cite those original papers instead of making an erroneous claim that will only serve to irritate their colleagues.Although this passage is written to suggest they are only referring to yeast, the papers cited in fact refer to Reviewer #2:In the manuscript, the authors developed a method to study interchromosomal interactions to simultaneously analyze hundreds of cis or trans-acting mutations. This method will advance the dissection of the molecular mechanisms of the chromosomal pairing. The authors identified the minimal sequence that is essential for the homologous pairing at the HAS1pr-TDA1pr locus. The pairing event occurs particularly in the saturated cultures in budding yeast. They introduced a barcode to each gene knockout and tested hundreds of transcription factors knockouts. The authors found that LEU3, SDD4, and RGT1 are required for pairing. Finally, they identified a particular domain of RGT1 that is responsible for the pairing. This study demonstrated that transcription factors play an important role in chromosomal interactions using genetic approaches. In my opinion, this study was well done.1) The authors analyze the DNA binding sites for RGT1, LEU3, and SDD4 at the HAS1pr-TDA1pr locus . However, there is no evidence showing that these transcription factors directly bind to HAS1pr-TDA1pr locus in saturation cultures but not in exponentially growing cultures. Chromatin immunoprecipitation for RGT1 and LEU3 could answer this question.2) In Figure 3C legend), there is a typo in the sentence: {greater than or equal to}2 bins from a telomere. What does "bin" mean in the figure legend?3) Does the homologous pairing at HAS1pr-TDA1pr regulate nearby gene expression in saturated cultures? The authors could analyze the RNA-seq data to address this question.Reviewer #3:Kim et al., present the development of MAP-C , a new approach to study the regulation of chromosomal interactions by mutation. Two flavors of the methods allow (i) identifying the local sequence required for interaction by cis MAP-C (ii) testing the requirement of other factors or sequences for interaction by trans MAP-C. The authors demonstrate the power of the method by applying it to the two HAS1pr-TDA1pr alleles, which they recently demonstrated to pair under defined biological conditions.They identify the local genomic sequence and three transcriptions factors, which they demonstrate to be essential for allelic pairing of HAS1pr-TDA1pr. The authors suggest that a regulated interaction between the three factors controls both the specificity and physiological regulation of HAS1pr-TDA1pr pairing.Kim et al., describe a method allowing to probe the molecular mechanism of chromosomal pairing. They identify both the precise genomic sequence and the factors required for inter-chromosomal interaction. In the future, this new tool will facilitate the study of pairing, a widely observed but still poorly characterized biological enigma.HAS1pr-TDA1pr pairing” and Figure 1D: The authors claim to identify five clusters, however this choice seems arbitrary with additional depletions of substituted nucleotides (e.g regions upstream of E and between G-H).- Subsection “A cluster of TF motifs is necessary and sufficient for - Figure 1—figure supplement 2C: While the intensity of ratios is clearly less intense in the control than in the 3C library, it shows a similar pattern (compare panel C to panel B).- Subsection “Three transcription factors are required for pairing”, second paragraph is difficult to understand. How does the position of deleted regions influence the interactions? This should be improved together with the explanation of trans MAP-C and Figure 2A.- The identification of three transcriptional factors (TFs) requird for HAS1pr-TDA1pr pairing suggests that transcriptional activity or RNA produced at the locus might play a role in regulating the interaction between the two regions. The authors should characterize the transcriptional status of the locus and its changes in the individual TFs knockouts and upon pairing.- Rgt1 domain analysis showed that the C-terminal region, largely structured, is required for pairing. However, there is quite a stretch from this observation to the suggestion that it is regulated by phosphorylation, as previously described . The authors should test the effect of mutating the regulatory S88 and S758 residues.eLife. We apologize for the additional time it has taken to render this decision. This is due to the fact that we waited for the remaining reviewer to respond. Although we never received their comments, we have decided to proceed on the basis of one review. Your revised article has been favorably evaluated by us and also by one of the original reviewers.Thank you for resubmitting your work entitled "A combination of transcription factors mediates inducible interchromosomal contacts" for further consideration at The manuscript has been improved but there is one remaining issue that needs to be addressed before acceptance. Reviewer 2 requests evidence that pairing has been successfully disrupted in order to validate interpretation of the results. Specifically, mutating Rgt1 binding sites results in a TDA1 upregulation but mutating Leu3 binding sites does not . It is important to determine by 3C-qPCR (for example) that pairing is lost in the mutants. The Log23C/genome data shown is not sufficient to make the claim. Reviewer 2 suggested that the authors compare HAS1pr-TDA1pr pairing with a non-pairing region or under exponential growth conditions. We believe that this experiment should not take too long to perform and would enable us to move forward with the manuscript, assuming all goes as planned.Reviewer #2:The revised manuscript includes ChIP-seq and RNA-seq data to investigate the mechanism of HAS1pr-TDA1pr pairing. The authors show that Leu3, Sdd4, and Rgt1 were more enriched at HAS1pr-TDA1pr locus in saturated culture conditions than in exponential growth condition, suggesting that these transcription factors are directly involved in the pairing. They performed RNA-seq for those TF mutants and TF binding site mutants and found that those mutations have little effects on transcription levels of HAS1 and TDA1 genes, and they haven't found the biological functions of the pairing in their manuscript. Several possible reasons why they have not detected a phenotypic effect from disrupting the pairing region or factors were discussed in the revised version. Although they have mostly addressed the questions I have, there are few concerns about whether they have successfully disrupted the pairing that may lead to a different interpretation of the results.HAS1 or TDA1 transcriptional regulation”). However, I don't see the measurement of the pairing events (such as 3C-qPCR) for those mutants . Do these mutants completely disrupt the pairing? They used Log23C/genome to show the decrease of pairing in those mutants, but it doesn't represent the level of pairing remained in those mutants. The authors should compare HAS1pr-TDA1pr pairing with a non-pairing region  as a control to show whether those mutants have entirely abolished the pairing or not.The authors found that the mutation of Rgt1 binding sites at the pairing locus results in a mild upregulation of TDA1 expression , but the mutation of Leu3 binding sites does not affect on gene expression . The authors claim that the disruption of pairing may not have any effect on transcription  The identification of 3 TFs for pairing suggests that transcriptional activity or RNA produced at the locus might play a role in the trans interaction. The authors should characterize the transcriptional status of the locus and its changes in the individual TFs knockouts and upon pairing.S. cerevisiae x S. uvarum hybrids, in order to detect potential trans effects that might result from disruption of pairing, as well as of TF knockouts in S. cerevisiae, all in the saturated culture conditions that produce HAS1pr-TDA1pr pairing. In addition, we analyzed existing microarray data of the same TF deletions in exponentially growing cells . As shown in Figure 5, we find that either mutating two Rgt1 binding sites in the HAS1pr-TDA1pr region or deleting the RGT1 gene result in mild (~1.5-2-fold) upregulation of TDA1, consistent with its known role as a repressor in low glucose conditions. However, these effects were limited to the copy of TDA1 adjacent to the mutations and indicate that pairing is not necessary for normal regulation of TDA1 under saturated conditions. Furthermore, none of the other perturbations had any significant effect on HAS1 or TDA1 expression. In the Discussion section, we describe the implications and possible reasons for a lack of a detected function for pairing.In response to this comment, we performed RNA sequencing of both TF binding site mutants in 2) While the C-terminal region of Rgt1 seems required for pairing, the suggestion that it is regulated by phosphorylation is a stretch. We recommend testing the effect of mutating the regulatory S88 and S758 residues.We have performed a MAP-C experiment with the Rgt1 S88A and S758A mutations described in Polish et al., 2005, alongside the ectopic wild-type Rgt1 control and deletions for the zinc finger, Q/N-rich, and C-terminal regulatory domains. As shown in Figure 4E and 4F, we do not find strong effects of preventing phosphorylation, only mild decreases in pairing. However, we suspect that Rgt1 would predominantly be in a dephosphorylated state under saturated culture conditions, and thus, our results do not rule out a role for phosphorylation in regulating pairing activity, i.e. preventing pairing in exponential growth conditions. But in light of these results, we have de-emphasized the possible role of phosphorylation in regulating pairing.3) Reviewer 1 would like to see either data or discussion of the biological purpose of localized pairing and why it is inducible. This is a fair question and I (reviewer 3) am also interested in the thinking behind it.HAS1pr-TDA1pr homolog pairing, or rather, why we fail to detect a molecular phenotype of disrupting either pairing TFs or their binding sites in the pairing region. As described in the revised Discussion section, there are various possible reasons why we have not detected a function: (1) even weak pairing  may suffice for its function; (2) pairing function may not act during saturated culture conditions, but instead upon exit from or re-entry into those conditions, as in epigenetic transcriptional memory; (3) pairing may regulate transcriptional dynamics rather than steady-state transcript levels. Finally, we speculate regarding the possibility that pairing has no function per se but is instead the result of conservation of homotypic intermolecular interactions among TFs that also mediate cooperative DNA binding.We have added extensive discussion of the function of 4) We would require evidence that the factors actually bind the pairing locus. ChIP-seq of these three transcription factors to verify the binding of Leu3, Sdd4 and Rgt1 at the minimal pairing region would strengthen the paper.S. cerevisiae grown to saturation and in exponentially growing cultures. As shown in Figure 2D and E, all three TFs bind much more strongly in saturated cultures, with only Leu3 showing detectable binding in exponential growth. We further use these data to extend our motif analyses assessing the uniqueness of HAS1pr-TDA1pr pairing and whether combinatorial TF binding specifies where homolog pairing occurs in saturated culture conditions . We find that HAS1pr-TDA1pr indeed exhibits the strongest coincident binding of all three TFs and conserved motif clusters in both S. cerevisiae and S. uvarum genomes. However, such coincident binding does not appear to be necessary for detectable inducible homolog pairing, as the copies of the HXT3 promoter exhibit pairing without Leu3 binding and only weak Sdd4 binding .We have performed ChIP-qPCR and ChIP-seq for Leu3, Sdd4, and Rgt1 using their TAP-tagged versions, both in haploid 5) Data to address the differences in binding level of the three TFs at the minimal pairing region under the two different growth conditions. ChIP-seq is again recommended.Please see our response to the last point.6) As mentioned by reviewers below, there are other major cases of pairing in various organisms, including mammals. In the revision, please be sure to properly cite those studies to put the current study in perspective.We would like to thank the reviewers for suggesting relevant literature. We have added discussion of other known cases of homolog pairing, including genome-wide pairing in flies and X chromosome pairing in mice, to the Introduction.7) Reviewer 2 also asked if homologous pairing at HAS1pr-TDA1pr regulates nearby gene expression in saturated cultures. If data are available to address this, please include them.HAS1 is strongly downregulated in saturated culture conditions, and TDA1 is mildly upregulated  and mention the expression of TDA1 in the Discussion section.Please see our response to point #1 above. We have also previously found  that Reviewer #1:[…] 1) Obviously, the homologous pairing between HAS1-TAD1 alleles is a localized event, and the pairing is inducible  and limited into a 1kb intragenic regions on chromosome XIII. However, this study did not attempt to answer key biological questions: what's the biological purpose or meaning of this localized pairing? Why it's inducible? There are no experimental data or discussions in the manuscript. This is an important question that could greatly improve the interest level and relevance of the findings and the manuscript.Please see our responses to main points #1, #3, and #7.2) In the Introduction, the author described that "a motif match is insufficient to fully predict TF binding", which is true. Later on this study identified binding of the three transcription factors binding to the "minimal pairing region" which are necessary and essential for homologous pairing between HAS1-TAD1 alleles. However, most of the Results section and Conclusion were based on motif scanning, motif searching, and mutations in this region. This study missed some very important experiments: ChIP-seq of these three transcription factors to verify the binding of Leu3, Sdd4 and Rgt1 at the minimal pairing region.Please see our responses to main points #4 and #5.3) Since the homologous pairing between HAS1-TAD1 alleles is significantly higher at growth saturation than at standard growth conditions, and this study also identified that three transcription factors  regulate this homolog pairing. What's the binding level of these three TFs at the minimal pairing region under the two different growth conditions? ChIP-seq of these three transcription factors under the two different growth conditions is very important to verify and solidify the conclusion.Please see our responses to main points #4 and #5.4) The authors claimed that HAS1pr-TDA1pr pairing is regulated by the expression level of TFs, particularly RGT1. In fact, using the mRNA expression level of TFs is not convincing way to convey this idea. Because increasing at mRNA level does not necessarily means increasing TFs' bindings at chromatin, DNA, or a specific loci (in the current study). The best experiment here would again be ChIP-seq of the three TFs to show differential binding.Please see our responses to main points #4 and #5.5) Last part of the Results section, there is no data to support that the "pairing domains" of Rgt1 needs to be phosphorylated for the purpose of homologous pairing. This is speculation without supporting data. If the authors believe that phosphorylation of the "pairing domains" of Rgt1 is one of the regulatory mechanisms of Rgt1 function, then they need to show that under growth saturation conditions, the "pairing domain" of Rgt1 is indeed phosphorylated.Please see our responses to main point #2.6) In the Introduction the authors claim that:"Although interchromosomal contacts, such as those between actively transcribed genes or homolog pairing in mitotically dividing yeast, have been observed by microscopy , many are not detectable using 3C (chromosome conformation capture) technologies. Some have argued that these contacts might be a result of fluorescent in situ hybridization (FISH) artifacts  or the insertion of binding site arrays often used for live-cell imaging . A more detailed understanding of the molecular components of interchromosomal contacts is needed to resolve this ambiguity."Although this passage is written to suggest they are only referring to yeast, the papers cited in fact refer to Drosophila and mouse cells. […] The authors should read and cite those original papers instead of making an erroneous claim that will only serve to irritate their colleagues.Please see our responses to main point #6.Reviewer #2:.[…] 1) The authors analyze the DNA binding sites for RGT1, LEU3, and SDD4 at the HAS1pr-TDA1pr locus . However, there is no evidence showing that these transcription factors directly bind to HAS1pr-TDA1pr locus in saturation cultures but not in exponentially growing cultures. Chromatin immunoprecipitation for RGT1 and LEU3 could answer this question.Please see our responses to main points #4 and #5.2) In Figure 3C legend, there is a typo in the sentence: {greater than or equal to}2 bins from a telomere. What does "bin" mean in the figure legend?Bins refer to genomic segments used for Hi-C analysis, in this case of 32 kb in length. To clarify this, we now describe them as “genomic bins”.3) Does the homologous pairing at HAS1pr-TDA1pr regulate nearby gene expression in saturated cultures? The authors could analyze the RNA-seq data to address this question.Please see our responses to main points #1 and #7.Reviewer #3:[…] - Subsection “A cluster of TF motifs is necessary and sufficient for HAS1pr-TDA1pr pairing” and Figure 1D: The authors claim to identify five clusters, however this choice seems arbitrary with additional depletions of substituted nucleotides (e.g regions upstream of E and between G-H).We now selected a threshold of depletion of substitutions (log2 < -1.1) to highlight positions in Figure 1D based on the bimodal distribution of depletion scores . Based on this analysis, we have further highlighted a sixth cluster that also matches the Rgt1 motif . These highlighted motifs span all but one of the positions most depleted of mutations in the 3C library, and all clusters of two or more adjacent positions with extreme scores.- Figure 1—figure supplement 2C: While the intensity of ratios is clearly less intense in the control than in the 3C library, it shows a similar pattern (compare panel C to panel B).We note that while there are regions with weak enrichment or depletion in even the control 3C library, the positions with signal do not match those of the on-target 3C library.- Subsection “Three transcription factors are required for pairing”, second paragraph is difficult to understand. How does the position of deleted regions influence the interactions? This should be improved together with the explanation of trans MAP-C and Figure 2A.Apologies. We have sought to rewrite this section to explain these aspects more clearly.- The identification of three transcriptional factors (TFs) required for HAS1pr-TDA1pr pairing suggests that transcriptional activity or RNA produced at the locus might play a role in regulating the interaction between the two regions. The authors should characterize the transcriptional status of the locus and its changes in the individual TFs knockouts and upon pairing.Please see our responses to main points #1 and #7.- Rgt1 domain analysis showed that the C-terminal region, largely structured, is required for pairing. However, there is quite a stretch from this observation to the suggestion that it is regulated by phosphorylation, as previously described . The authors should test the effect of mutating the regulatory S88 and S758 residues.Please see our responses to main point #2.[…] The manuscript has been improved but there is one remaining issue that needs to be addressed before acceptance. Reviewer 2 requests evidence that pairing has been successfully disrupted in order to validate interpretation of the results. Specifically, mutating Rgt1 binding sites results in a TDA1 upregulation but mutating Leu3 binding sites does not . It is important to determine by 3C-qPCR (for example) that pairing is lost in the mutants. The Log23C/genome data shown is not sufficient to make the claim. Reviewer 2 suggested that the authors compare HAS1pr-TDA1pr pairing with a non-pairing region or under exponential growth conditions. We believe that this experiment should not take too long to perform and would enable us to move forward with the manuscript, assuming all goes as planned.S. cerevisiae x S. uvarum context in an effort to rigorously test for transcriptional effects of pairing, rather than effects in cis or other effects in trans. We believe that these mutations do not completely disrupt pairing as reviewer #2 suggests, based on our cis and trans MAP-C experiments in which disrupting a single TF binding site or TF gene results in ~4-10-fold weaker pairing , compared to the ~30-fold increase in ectopic pairing  conferred by the entire pairing sequence. Thus, as we have agreed upon, instead of performing the proposed experiment, we have edited the text (in the Results section and the Discussion section) to clarify our expectation that we have not fully disrupted pairing, but rather to ~10-25% of wild-type levels. We understand that reviewer #2 is concerned that this may have prevented us from finding transcriptional effects; however, this is just one of several potential reasons that we did not detect a phenotypic effect of disrupting HAS1pr-TDA1pr homolog pairing, as detailed in the Discussion section, and testing these possibilities is beyond the scope of this work.As we discussed, we introduced highly targeted Leu3 and Rgt1 binding site mutations in a hybrid Reviewer #2:The revised manuscript includes ChIP-seq and RNA-seq data to investigate the mechanism of HAS1pr-TDA1pr pairing. […] The authors should compare HAS1pr-TDA1pr pairing with a non-pairing region  as a control to show whether those mutants have entirely abolished the pairing or not.See above.


Drosophila. This protein contains acidic regions at both ends and a central cluster of 12 zinc finger domains, some of which are involved in the specific recognition of the binding site. One of the well-described in vivo function of Su(Hw) is the repression of transcription of neuronal genes in oocytes. Here, we have found that the same Su(Hw) C-terminal region (aa 720–892) is required for insulation as well as for promoter repression. The best characterized partners of Su(Hw), CP190 and Mod(mdg4)-67.2, are not involved in the repression of neuronal genes. Taken together, these results suggest that an unknown protein or protein complex binds to the C-terminal region of Su(Hw) and is responsible for the direct repression activity of Su(Hw).Suppressor of Hairy-wing [Su(Hw)] is a DNA-binding architectural protein that participates in the organization of insulators and repression of promoters in  In particular enhancers can activate target promoters at large distances (up to hundreds of kb in some cases), which raises the question of the mechanisms regulating such long-distance enhancer–promoter interactions. More than 25 years ago, a special class of regulatory elements, named insulators, was suggested to delimit the activity of enhancers12. Insulators are defined as regulatory elements that disrupt the communication between an enhancer and a promoter when inserted between them. Some insulator complexes contribute to higher-order organization of chromatin in topologically associated domains that are fundamental elements of the eukaryotic genomic structure14.High-resolution chromosome conformation capture techniques have provided evidence that regulatory elements form loops that are essential for gene regulation in higher eukaryotesgypsy retrotransposon, whose integration into genes often resulted in inactivation of enhancers that were separated from promoters by the gypsy insertion19. The phenotypes of the gypsy-induced mutations were suppressed by inactivation of the gene encoding the Suppressor of hairy wing protein [Su(Hw)]20. The gypsy insulator consists of 12 reiterated binding sites for Su(Hw)22. Today, Su(Hw) is one of the best characterized insulator proteins. It has been shown that artificial reiterated binding sites for Su(Hw) or gypsy insulator can block various enhancers at all stages of Drosophila development27.One of the first insulators was identified in the 2H2-type zinc finger domains, and the C-terminal region (aa 716–892) responsible for enhancer blocking activity32. Several Su(Hw) partners were identified, including Mod(mdg4), CP190, ENY2, Shep, Rump, and HIPP138. Mod(mdg4)-67.2 is one of the isoforms encoded by the mod(mdg4) locus40. The Mod(mdg4)-67.2 protein contains the N-terminal BTB/POZ domain and glutamine-rich (Q-rich) region which is common to all isoforms and the unique C-terminal region that is required for interaction with the 716–892 aa region of Su(Hw)34. The BTB and Q-rich domains of the Mod(mdg4)-67.2 interact with the M domain of CP190 and the N-terminal region of Su(Hw) (aa 1–238), respectively31. The functions of Mod(mdg4)-67.2 are as yet unknown, except for its participation in recruiting the Su(Hw) complex to specific chromatin sites and masking the repression activity of Su(Hw)42.The Su(Hw) protein contains the N-terminal region involved in the interaction with CP190, an array of 12 C46. The BTB domain interacts with two adjacent regions at the Su(Hw) N-terminus, which is essential for CP190 recruitment to Su(Hw) sites32. CP190 is involved in the recruitment of several transcriptional complexes to chromatin, including the NURF, dREAM, and SAGA complexes48. CP190 has major effects on chromatin, such as the depletion of nucleosomes and the prevention of heterochromatin expansion49. Similar to CP190, ENY2 is recruited to chromatin by interacting with Su(Hw) and participates in the boundary function of Su(Hw), which protects active genes from silencing36. Shep and Rump are the RNA binding proteins that function as negative regulators of the Su(Hw) enhancer blocking activity37. HIPP1 (CG3680) was identified as possible partner of Su(Hw) in a recent screen of proteins interacting with HP1, one of the main heterochromatin proteins33. Strong co-localization of HIPP1 and Su(Hw) sites suggests that these proteins are in the same complex33.The CP190 protein contains the N-terminal BTB domain that forms stable homodimers53. It has been shown that many SBS-O sites are involved in transcriptional repression54. The function of Su(Hw) as transcriptional repressor is required for female germline development54 and for sustained male fertility55.In previous genome-wide studies, three classes of the Su(Hw) binding regions have been identified, which are characterized by the binding of Su(Hw) alone (SBS-O), both Su(Hw) and CP190 (SBS-C), or three proteins Su(Hw)/CP190/Mod(mdg4)-67.2 (SBS-CM)Here, we made an attempt to identify Su(Hw) domains involved in the repression of several neuronal genes in the ovary. We found that the same C-terminal region (aa 720–892) in Su(Hw) is required for enhancer blocking and for promoter repression.56. These genes contain binding sites for Su(Hw) in promoter regions, suggesting the involvement of this protein in direct repression of transcription. To further characterize Su(Hw) domains involved in repression, we selected five representative neural genes whose promoters are bound by Su(Hw) alone , by Su(Hw) and CP190 (Syn2), or by Su(Hw), CP190, and Mod(mdg4)-67.2 (mAcR-60C)  isoform was recruited to other promoters by an unknown DNA-bound protein. In general, the observed patterns of Su(Hw), CP190, and Mod(mdg4)-67.2 recruitment to the tested promoters in the ovaries and heads were similar, which is in agreement with the previous observation that these proteins stably bind to their sites in different Drosophila tissues and cell lines54.As shown previously, Su(Hw) is involved in the repression of many neural genes in the ovary0C) Fig. . Using C0C) Fig. . Interessu(Hw)− background. In the absence of the Su(Hw) protein, transcription from the tested genes proved to be increased 10- to 20-fold in the ovaries, while its level in the heads remained unchanged  binding sites led to an increase in transcription, compared to the wild-type level, by a factor of 4 for mAcR-60C and by a factor of 6 for Rph  protein can repress the promoters: (1) Su(Hw) competes for the binding site with a transcription factor that activates transcription; (2) Su(Hw) recruits a tissue-specific complex that is responsible for repression. To test these alternative models, we mutated the Su(Hw) binding sites in the promoters of the nes Fig. . The lacRph Fig. . At the hsp70 promoter and two-time heat shock treatment for 2 hours at the embryonic and larval stages. The results were confirmed by Western blot analysis  gene transcription, compared to the wild-type level  can affect transcription of the model genes in the heads. To induce Su(Hw) overexpression, we used the sis Fig.  and RT-qvel Fig. . Despiteyn2 Fig. . This re54 that the CP190 and Mod(mdg4)-67.2 partners of Su(Hw) are not usually essential for repression, since Su(Hw) alone is present at many repressed promoters. As noted above, the mAcR-60C promoter is bound by Su(Hw), CP190, and Mod(mdg4)-67.2, while the Syn2 promoter, only by Su(Hw) and CP190. Hence, the question has arisen whether CP190 and Mod(mdg4)-67.2 are involved in the regulation of these promoters. According to our previous data32, the BTB domain of CP190 binds to two regions located between aa 88 and 202 at the N-terminus of Su(Hw)  complex. We compared the recruitment of Su(Hw) and CP190 proteins to the promoters of model genes in pupae of transgenic lines expressing full-length Su(Hw)+ and Su(Hw)Δ114 in the absence or presence of the mod(mdg4)u1 mutation inactivating the Mod(mdg4)-67.2 isoform /CP190/Mod(mdg4)-67.2) and Syn2 (Su(Hw)/CP190) promoters. In the Su(Hw)Δ114 mutants, CP190 did not bind to the mAcR-60C promoter, confirming the role of Su(Hw) in the recruitment of this protein. In contrast, CP190 still bound to the Syn2 promoter in these mutants, suggesting that CP190 is recruited to this promoter independently of Su(Hw).It has been shownHw) Fig. . The delorm Fig. . UnexpecΔ114 mutation did not lead to derepression of model genes  alone) and the mAcR-60C promoter (Su(Hw)/CP190/Mod(mdg4)-67.2) this mutation resulted in a noticeable decrease of transcription, suggesting the positive role of the corresponding Su(Hw) domain in stimulation of transcription from the promoters. Taken together, these results confirm that the Mod(mdg4)-67.2 and CP190 proteins are not involved in repression.The RT-qPCR analysis in the ovaries and heads showed that the Su(Hw)nes Fig. . In cont30. To map the domains involved in transcriptional repression, we generated transgenes expressing different variants of Su(Hw) under control of the ubiqitin63-E promoter − (su(Hw)v/su(Hw)e04061trans-heterozygous) background. Western blot analysis showed that all Su(Hw) variants were expressed at similar levels Δ283 line expressed the protein lacking 16 aa (from 760 to 778) that are critical for the enhancer-blocking activity of Su(Hw)30. In addition, we generated two transgenes mimicking the su(Hw)J and su(Hw)e7 mutations described previously29. The Su(Hw)e7 and Su(Hw)J transgenes produced truncated proteins lacking 223 and 150 aa, respectively. Both deletions affect the domain that is essential for interaction with the C-terminal domain of Mod(mdg4)-67.234. ChIP with pupae showed that all Su(Hw) variants bound to the tested promoters at a level comparable to that of wild-type Su(Hw)  protein that contains the acidic and enhancer-blocking regions ter Fig. . The traels Fig. . The Su(Hw) Fig. .Figure 5J, Su(Hw)e7, or Su(Hw)Δ283 females, their expression level in the ovaries proved to be increased up to 5-fold, as in su(Hw)− females, while that in the heads expectedly remained unchanged. In contrast, deletion of the C-terminal acidic domain had no effect on the repression activity of the mutant Su(Hw) protein. The expression of different Su(Hw) variants directly correlated with female fertility: Su(Hw)J, Su(Hw)e7, and Su(Hw)Δ283 females were sterile, while Su(Hw)ΔC females were fertile. These results show that the enhancer-blocking domain mediates the Su(Hw) dependent repression of the model promoters.We then compared the effect of Su(Hw) mutants on the expression of model genes in the ovaries and heads Fig. . In Su(Hgypsy insulator fails to insulate the eye enhancer in the transgenic lines25. The Su(Hw) protein in the head fails to function either as an insulator or as a repressor. This provides a basis for the hypothesis that the same unknown complex bound to the 720–892 aa region of Su(Hw) is responsible for the promoter repression and enhancer blocking. It seems likely that this complex is bound to Su(Hw) in many tissues, with the exception of cells attributed to the nervous system. This hypothesis is confirmed by the observation that the gypsy insulator cannot block communication between the iab cis-regulatory domains and Abd-B promoter in the CNS58. In addition, the RNA-binding protein Shep has been shown to antagonize Su(Hw) activity in the CNS but not in other tissues35. It is possible that Shep competes with the hypothetical enhancer-blocking/repression complex for binding to the Su(Hw) protein.We have mapped the 720–892 aa region of the Su(Hw) protein, which is responsible for tissue-specific repression of neural genes in the ovaries. This region overlaps with the Su(Hw) enhancer-blocking domain (aa 716–892) characterized previously. Our results show that Su(Hw) overexpression does not lead to increased repression of promoters in the head, suggesting that Su(Hw) itself is not involved in this process. The same deletions in Su(Hw) affect insulation, transcriptional repression and lead to female sterility. As found previously, one copy of the 56. At the same time, Mod(mdg4)-67.2 affects the enhancer-blocking activity of Su(Hw)-dependent insulators60. Mod(mdg4)-67.2 can interact with proteins responsible for long-distance communication between enhancers and promoters. For example, Mod(mdg4)-67.2 interacts with Zeste, which is essential for long-range interactions between the eye enhancer and white promoter61.The Mod(mdg4)-67.2 protein is not required for female fertility and is not involved in the Su(Hw)-mediated repression of the promoters34. In the y2 model system63, the Su(Hw) complex acts as an insulator, while in the absence of Mod(mdg4)-67.2 it becomes a repressor64, suggesting that Mod(mdg4)-67.2 competes with the hypothetical enhancer-blocking/repression complex. This hypothesis is consistent with the observation that the genomic sites that bind the Su(Hw) protein in the absence of Mod (mdg4)-67.2 act as negative regulators of transcription54.Interestingly, Mod(mdg4)-67.2 protein interacts with the 716–892 aa region of Su(Hw) that includes a leucine zipper motif and completely overlaps with the enhancer blocking/repression domain46. Moreover, CP190 protein is critical for the functioning of the Fub insulator element that topologically separates the Ubx gene from the posterior abd-A gene60. CP190 is also involved in the recruitment of transcriptional complexes that open chromatin and stimulate transcription65. We have demonstrated that deletion of the Su(Hw) N-terminal domain responsible for the interaction with CP190 leads to a significant decrease in the expression of model genes, suggesting that CP190 is involved in transcription activation rather than in Su(Hw)-dependent repression.As shown previously, CP190 mediates protein–protein interactions necessary for the establishment of long-term contacts between insulator complexesgypsy insulator42. Su(Hw) binds to the mAcR-60C promoter in complex with CP190 and Mod(mdg4)-67.2. However, deletion of the Su(Hw) N-terminal domain even in combination with the mod(mdg4)u1 mutation has no effect either on Su(Hw) binding or on promoter repression. Thus, Su(Hw) recruitment to the mAcR-60C promoter does not depend on the presence of Mod(mdg4)-67.2 or CP190. It is noteworthy in this context that CP190 is recruited to the Syn promoter independently of Su(Hw). Our previous data show that multiple interactions between CP190, Mod(mdg4)-67.2, and Su(Hw) are involved in the formation of a stable DNA-bound chromatin complex32. Based on this fact, we hypothesize that only the Su(Hw)/Mod(mdg4)-67.2/CP190 complex or Su(Hw) in combination with as yet unknown proteins are bound to chromatin, while SBS-C sites are formed by independent binding of the Su(Hw) protein and CP190 in complex with another architectural protein.The CP190 and Mod(mdg4)-67.2 proteins facilitate the binding of Su(Hw) to many SBS-CM sites and, in particular, to the 54. This transcriptional repressor complex of Drosophila contains three subunits: CoREST, lethal (3) malignant brain tumor protein (L(3)mbt), and L(3)mbt interacting protein 1 (dLint-1)67. As shown previously, over a third of CNS-enriched repressed target genes contain SBSs that colocalize with L(3)mbt and half of them contain SBSs that colocalize with dLint-154. Based on this finding, we examined whether L(3)mbt colocalized with the promoters of our model genes mbt on the tested promoters: this proteint was either enriched in the larval CNS and Kc cells (Rph) or absent in both cases . It was only on the cg32017 promoter that L(3)mbt was enriched in Kc cells and not detected in the CNS. Apparently, L(3)mbt is not directly recruited to SBSs by Su(Hw). This assumption is confirmed by results of yeast two hybrid assay (Suppl. Table 33. We have shown that HIPP1 dirrectly interacts with Su(Hw) region responsible for repression (unpublished data). As demonstrated redcently, however, HIPP1 is dispensable for the repression activity of Su(Hw)69. Taken together, these results suggest that additional, as yet unidentified complex is responsible for tissue-specific repression of neural genes in the ovaries. Further studies are needed to identify components of this complex and mechanism of its formation on SBSs.It was previously speculated that Su(Hw) repression might depend on the recruitment of the LINT complexnes Fig. . We analThe transgenic constructs are described in the Supplementary Materials.y1M{vas-int.Dm}ZH-2A w*; M{3xP3-RFP.attP′}ZH-38D genotype57. The generation of transgenic lines and construct introduction into the Su(Hw)v/Su(Hw)e04061 background were performed as described41.All flies were maintained at 25 °C on the standard yeast medium. To obtain transgenic flies with insertion in 38D, the DNA of reporter constructs was injected into preblastoderm embryos of 52. After dissection, the flies were collected into a 50-mL Falcon tube, frozen in liquid nitrogen, and vigorously vortexes for 1 min. The contents of the tube were poured onto a sheet of white paper, and the heads of flies were separated from the bodies with a bird feather.For each replicate, approximately 100 ovaries from 4- to 6-hour-old virgin females were dissected as described previouslyras64B as an internal control. Primer sequences used in real-time PCR analysis are listed in Suppl. Table For RT-qPCR experiments, total RNA from the heads and ovaries of females was isolated with TRIzol reagent (Invitrogen). Genomic DNA was removed by treatment with DNase I  followed by purification with a QIAGEN RNeasy kit. RNA was reverse transcribed into cDNA with a RevertAid H Minus RT Revert Transcriptase (Fermentas) following the manufacturer’s instructions. The resulting cDNA was analyzed by quantitative PCR (Bio-Rad CFX 96 Cycler) using SYBR Green. Relative steady-state mRNA levels were determined from the threshold cycle for amplification using the ΔΔCT method. Each experiment was performed in two to three independent biological replicates, and the results were averaged. The expression level of each gene was determined using 2, 0.5% Triton X100, 0.5 mM DTT, 10 mM sodium butyrate, EDTA-free protease inhibitors cocktail, and 1.8% formaldehyde) at room temperature using first a Potter homogenizer and then a Dounce homogenizer with type A pestle (three strokes). The protease inhibitors cocktail  was used following the manufacturer’s instructions. After 15 minutes , glycine solution was added to a concentration of 225 mM, and the mixture was stirred, incubated for 5 minutes, and then centrifuged at 4000 g and 4 °C for 5 minutes. The pellet was washed three times with buffer A at 4 °C and resuspended in 0.5 mL of lysis buffer . The mixture was incubated on a rotary shaker at 4 °C for 10 minutes, sonicated on ice with a Branson Sonifier 450 , rotated for another 10 minutes, and then centrifuged at 15,000 g for 5 minutes. The supernatant was transferred to a new tube. The pellet was resuspended in 0.5 mL of lysis buffer, incubated on a rotary shaker for 10 minutes, and centrifuged again. The first and second supernatants were pooled and centrifuged twice at 15,000 g for 10 minutes. The supernatant was diluted with ten volumes of ChiP Dilution Buffer  and, to reduce nonspecific background, pre-cleared by incubation with protein A or protein G agarose beads for 30 min at 4 °C, with constant stirring. Agarose was pelleted by brief centrifugation, and the supernatant was collected for chromatin immunoprecipitation with appropriate antibodies (see below). After overnight incubation at 4 °C on a rotary shaker, protein A or protein G agarose beads were added to collect the precipitated complexes, and incubation was continued for 2 hours under the same conditions. Agarose was pelleted by centrifugation , the supernatant was carefully removed, and the pellet was washed with the following buffers : Low Salt Wash Buffer , High Salt Wash Buffer , LiCl Wash Buffer , and two portions of TE Buffer. The complex was removed from the agarose by two rounds of incubation with 250 μL of elution buffer  at room temperature for 15 minutes, with rotation. The eluates were pooled, supplemented with 20 μL of 5 M NaCl, and heated at 65 °C for 4 hours to reverse the complex–DNA crosslinks. Then 20 μL of 1 M Tris-HCl (pH 6.5), 10 μL of 0.5 M EDTA, and 2 μL of Proteinase K solution (10 mg/mL) were added, and the mixture was incubated at 45 °C for 1 hour. DNA was recovered by phenol/chloroform extraction and ethanol precipitation and solubilized in water for PCR. PCR products were amplified from at least two separate immunoprecipitates from three different chromatin preparations. Primer sequences used in PCR for ChIP analysis are shown in Suppl. Table Chromatin was prepared either from the heads and ovaries of female flies or from middle pupae. The material (150–200 mg) was homogenized in 5 mL of buffer A -67.2 (1:500), mouse anti-Mod-common (1:500), and rabbit anti Su(Hw) N-terminal domain (1:200) raised in our lab70; rabbit antibodies against the C-terminal domain of Su(Hw) (1:200) kindly provided by M. Erokhin32.Specific antibodies and working dilutions were as follows: rat anti-CP190 (1:500), rabbit antiDataset 1


The thioredoxin domain containing proteins are a group of proteins involved in redox regulation and have been recently reported to be associated with tumor progression. However, the role of thioredoxin proteins in hepatocellular carcinoma (HCC) remains largely unknown. Here in our study, we demonstrated that thioredoxin domain containing protein 9 (TXNDC9) was over-expressed in HCC and promoted HCC progression. We found that TXNDC9 expression was amplified in HCC tissues and associated with an advanced grade of HCC. And, we demonstrated that overexpression of TXNDC9 was correlated with poor prognosis of HCC. Furthermore, by using CRISPR-Cas9 mediated TXNDC9 knockout and RNA-seq analysis, we found that TXNDC9 accelerated HCC proliferation regulation. Moreover, we demonstrated that TXNDC9 directly interacted with MYC and knockout/knockdown of TXNDC9 decreased the protein levels of MYC and inhibited MYC-mediated transcriptional activation of its targets. Besides, we identified that TXNDC9 was trans-activated by FOXA1, JUND, and FOSL2 in HCC. Taken together, our study unveiled an oncogenic role of TXNDC9 in HCC and provided a mechanistic insight into the TXNDC9 mediated gene regulation network during HCC development. HCC is characterized by a high and increasing incidence, a late diagnosis when curative-intent treatments are not feasible, a low resectability rate, a high recurrence after a curative-intent surgery, a poor response to medical treatments, and finally a grave prognosis4. The mortality rate due to HCC in most countries is nearly equal to the incidence rate, indicating the lack of effective therapies at diagnosis5. Hence, it is critical to explore novel biomarkers for HCC diagnosis, prediction of patient and treatment outcomes, and individualization of targeted therapies7.Hepatocellular carcinoma (HCC) is an increasingly serious health problem, with over 700,000 new cases every year worldwide9. This superfamily of proteins has roles in various functions in the regulation of redox homeostasis, oxidative stress, disulfide-containing proteins, and transcription11. Emerging evidence on cancer development has shown that the thioredoxin system contributed greatly to tumorigenesis and that aggressive tumors expressed high levels of thioredoxin14. Thioredoxin domain containing 9 (TXNDC9), also known as phosducin-like protein 3 (PHLP3), is a distinct member of thioredoxins, which have ATP binding protein activity. It modulates the ATPase activity of chaperonin TCP1 complex, a key complex for protein folding, and diminishes actin and tubulin folding15. Interestingly, recent studies have demonstrated that TXNDC9 was overexpressed in colorectal cancer, indicating a potential oncogenic function of TXNDC9 in malignant diseases16. However, the role of TXNDC9 in HCC and the mechanism by which TXNDC9 may exert an oncogenic function remains largely unknown.Thioredoxin (TRX) domain containing proteins are a group of redox proteins that catalyze the reversible oxidation of cysteine thiol to disulfideHere, we demonstrated that TXNDC9 was highly expressed in HCC and that the high expression of TXNDC9 was a poor prognosis factor of HCC. The knockout/knockdown of TXNDC9 resulted in a significant inhibition of the growth of HCC cells. Moreover, we observed that TXNDC9 directly interacted with MYC and stabilized the MYC protein, and the knockout of TXNDC9 impaired the binding of MYC to chromatin. Finally, we found that FOXA1, JUND, and FOSL2 bound at the promoter of TXNDC9 and led to TXNDC9 overexpression in HCC.A total of 208 patients with HCC who had undergone surgical resection at Jiangyin People’s Hospital, School of Medicine, Southeast University from January 2010 to December 2014 were enrolled in this study. This study included 176 males and 32 females, with a mean age of 50.9 years old (from 25 to 75 years old). The follow-up period in this study ranged from 2 to 86 months after surgery. This research was performed in accordance with the ethical standards of Jiangyin People’s Hospital, School of Medicine, Southeast University and the Declaration of Helsinki and its amendments. Written consents were acquired from all participants.TMAs were constructed using HCC and paired adjacent noncancerous liver tissue specimens from each patient. The slides were immunolabeled by a primary rabbit monoclonal antibody against human TXNDC9 . The staining intensity for TXNDC9 was graded as 0 (negative), 1 (weak), or 2 (strong). The staining extent was graded as 0 (0%), 1 (1–25%), 2 (26–50%), 3 (51–75%), or 4 (76–100%). Specimens were classified into negative (0–1), weakly positive (2–4), or strongly positive (5–6), based on the sum of the staining intensity and staining extent scores. Weakly and strongly positive were considered as positive. The slides were analyzed by two independent researchers.® ep realplex  using an iQTM SYBR® Green Supermix kit . The mRNA levels were normalized to those of β-Actin. The following specific primers for qPCR were used: TXNDC9-F: 5′- CTGCTTCAGACTACCAAACTGG-3′, TXNDC9-R: 5′- CTCTGTAGAAATGGCAAACCACA-3′; SRM-F: 5′-GTGGTGGCCTATGCCTACTG-3′, SRM-R: 5′-CTCCTGGAAGTTCGTGCTCG-3′; DUSP2-F: 5′-GGGCTCCTGTCTACGACCA-3′, DUSP2-R: 5′-GCAGGTCTGACGAGTGACTG-3′; WDR74-F: 5′-CCTGGGGTGTGTAGGATGC-3′, WDR74-R: 5′-CAAGTCCAGCCAGTCATTCCG-3′; PES1-F: 5′-GGCCACCAACTACATCACCC-3′, PES1-R: 5′-AGAATGCACAGCCGCCTAAA-3′; β-Actin-F: 5′- CATGTACGTTGCTATCCAGGC-3′, β-Actin-R: 5′-CTCCTTAATGTCACGCACGAT-3′. The relative mRNA expression levels were evaluated using the 2−ΔΔCT comparative method.Total RNA of frozen tissues was extracted using an All-Prep DNA/RNA Mini kit . cDNA was reverse transcribed with the A3500 RT-PCR System  from 1 μg of total RNA. The relative TXNDC9 mRNA levels were detected by qPCR with MastercyclerTotal protein, cytoplasmic and nuclear protein were isolated and quantified by a bicinchoninic acid protein assay kit . Then, aliquots of proteins were separated on a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to polyvinylidine difluoride filter membranes . The membranes were blocked with 5% nonfat milk and probed by a primary rabbit monoclonal antibody against human TXNDC9 , GAPDH  or rabbit polyclonal antibody MYC . Following incubation with a horseradish peroxidase-conjugated secondary antibody , blots were visualized by chemiluminescence . GAPDH was used as the loading control.17. The sgRNA sequences are listed below. 5′-GTTGAGCTTTCCTTAGTGCC-3′ or 5′-TCTGAAGCAGCTGATGCTCC-3′. Lentivirus was produced as previously described17.The HepG2 and Hep3B cell lines were purchased from the American Type Culture Collection (ATCC) and cultured in DMEM containing 10% of fetal bovine serum. The CRISPR-Cas9 system was used to create the TXNDC9 knockout. The sgRNA sequences, targeting the first exon of TXNDC9, were inserted to the lentiCRISPRv2 plasmid (Addgene Plasmid #52961)siRNAs were purchased from the Genepharm co. ltd. siRNA primers are listed below. si-FOXA1-sense: 5′-GAGAGAAAAAAUCAACAGCdTdT-3′, si-FOXA1-antisense: 5′-GCUGUUGAUUUUUUCUCUCdTdT-3′; si-JUND-sense: 5′- CCCUCAAGAGUCAGAACACdTdT-3′, si-JUND-antisense: 5′- GUGUUCUGACUCUUGAGGGdTdT-3′; si-FOSL2-sense: 5′- GGAUUAUCCCGGGAACUUUdTdT-3′, si-FOSL2-antisense: 5′-AAAGUUCCCGGGAUAAUCCdTdT-3′. siRNAs were transfected into HepG2 cells using lipofectamine 2000  according manufactures’ instructions. shRNA targeting TXNDC9 were constructed in plvx-sh2 vector with the following primer. shTXNDC9-F: 5′- GATCCGCGGTTCTTCTGACATTCTTAATTCAAGAGATTAAGAATGTCAGAAGAACCGTTTTTTG-3′, shTXNDC9-R: 5′- AATTCAAAAAACGGTTCTTCTGACATTCTTAATCTCTTGAATTAAGAATGTCAGAAGAACCGCG-3′.®  according to the manufacturer’s protocol. Sequencing was performed using the Illumina Hiseq Xten platform.RNA was extracted from TXNDC9 knockout or wild-type HepG2 cells using TRIzol Reagent  according to the manufacturer’s protocol. Polyadenylated RNA was isolated from 5 μg total RNA with the mRNA Capture Beads. An RNA-seq library was constructed with VAHTSTM mRNA-seq V2 Library Prep Kit for Illumina18 with a reference annotation from the Ensembl database. No-novel-juncs options were used to ignore de novo gene and transcript identification. A gene expression profile of each sample, as well as differentially expressed genes between wild-type and TXNDC9 knockout HepG2 cells, were identified using the Cuffdiff suite from Cufflinks19. Genes with two-fold changes between two groups, q value < 0.01 and average expression > 1 were identified as TXNDC9 regulated targets. Raw data and processed data were deposit at the GEO database with following links: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE113400 with token key: gzmjeceqpnohler.Reads were mapped to human genome (hg38) using TopHat2ChIP experiments were performed with the ChIP-IT high sensitivity kit  in TXNDC9 knockout and wild-type HepG2 cells according to the manufacture’s instruction. Antibodies against MYC  were used. ChIP-qPCR primers were used as list below. EIF3B- ChIP-q-F: GGTCAATGCTCCCATGAGTT; EIF3B-ChIP-q-R: AGGACTCGCATCCACAAGAC; EIF4A1-ChIP-q-F: GGGATGTGGATTGTTCATGG; EIF4A1-ChIP-q-R: GGGGAGATTGCACAACACTT; CDK4-ChIP-q-F: TGGGAAGGGACTGCACTTAC; CDK4-ChIP-q-R: GAGGGGGCCTCTCTAGCTT; Negative-F: TTAGACGGAGTCGGAGCTGT; Negative-R: TCTTCAGGTCGGGCATTATC.http://encodeqt.simple-encode.org) was used for transcription enrichment analysis. In all, −2000 to 200 bp of the transcription start site of genes regulated by TXNDC9, identified by RNA-seq analysis, were used for transcription factor enrichment.The ENCODE ChIP-seq Significance tool  was evaluated by the Kaplan–Meier method using the log-rank test. Independent risk factors for OS were investigated by Cox regression analysis. Statistical analysis was conducted with SPSS 21.0 . All tests were two-sided, and a statistically significant difference was considered when p < 0.05.Correlation of TXNDC9 expression with clinicopathologic characteristics was assessed by Pearson’s p < 0.001, Fig. To determine the effect of TXNDC9 expression on HCC progression, we examined TXNDC9 expression in HCC tissues and paired adjacent noncancerous liver tissues. We performed immunohistochemical staining of TMAs containing 208 paired samples of HCC and normal liver tissues, and observed that the frequency of TXNDC9-positive staining in cancerous tissues 70.7%, 147/208) was statistically significantly higher than that observed in normal liver tissues   database and seven clinical cohorts from GEO database. In accordance with our findings, the expression of TXNDC9 in HCC tissues were significantly higher than those in normal liver tissues Figs. , suggestp = 0.035). This was further supported by the TCGA datasets, as illustrated in Fig. p = 0.013) and poor tumor differentiation (p = 0.022). However, there was no significant correlation between TXNDC9 expression and gender, age, hepatitis B virus DNA, alpha-fetoprotein, cirrhosis, tumor number, or portal vein invasion . Thus, TXNDC9 is up-regulated in HCC with advanced grade and might serve as an oncogene during HCC development and progression.Because TXNDC9 expression was significantly increased in HCC, we sought to determine whether TXNDC9 expression according to the TMAs immunohistochemical staining results was associated with the clinicopathologic characteristics of HCC patients Table . High TXp < 0.010, Fig. p < 0.010). Furthermore, the association between 11 clinicopathologic characteristics and survival rate of HCC patients after surgical resection was investigated by Cox regression analysis. Univariate Cox regression analysis indicated that multiple tumors (p = 0.014), tumor size > 5 cm (p = 0.008), portal vein invasion (p = 0.007), poor tumor differentiation (p = 0.018), TNM stage III–IV (p = 0.001), and positive TXNDC9 expression (p = 0.011) were predictors for a lower OS (Table p = 0.003) and positive TXNDC9 expression  may predict a lower OS independently . As shown in Fig. To determine how TXNDC9 affected the progression of HCC, we performed RNA-seq analysis in TXNDC9 knockout and wild-type HepG2 cells. We identified 1758 genes were up-regulated while 1783 genes were down-regulated upon TXNDC9 knockout Table . We then4bTo determine the function associated with TXNDC9 regulation, we performed gene set enrichment analysis using the 50 hallmark gene sets in the GSEA database. Interestingly, we found that MYC targets were significantly enriched at genes down-regulated upon TXNDC9 knockout Fig. . To valiFurthermore, to evaluate the functional importance of MYC in TXNDC9 mediated regulation, we restored the expression of MYC and determined the cell proliferation and apoptosis upon TXNDC9 knockout. As shown in Fig. To further determine the role of TXNDC9 in gene regulation, we analyzed the distribution of TXNDC9 in HCC cells. As shown in Figure To better understand the role of TXNDC9 in the MYC-mediated transcriptional program, we performed ChIP-qPCR using antibodies against MYC in TXNDC9 knockout and wild-type HepG2 cells. Interestingly, we found the binding of MYC on its target genes was significantly reduced in the TXNDC9 knockout cells Fig. , indicatNext, we performed ChIP-seq enrichment analysis of the TXNDC9 regulated genes using the ChIP-seq data from the Encyclopedia of DNA Elements (ENCODE) project to validate the finding that TXNDC9 regulates MYC targets. Indeed, we found that the promoter of genes down-regulated upon TXNDC9 knockout were significantly enriched at genome-wide binding sites of c-MYC as well as its partner MXI1, MAZ, and MAX Fig. . This fuThe mechanism by which TXNDC9 is overexpressed in HCC remains unanswered. To address this issue, we first investigated DNA methylation of the TXNDC9 promoter using the DNA methylation array results from TCGA database Figure . In HCC Next, we scanned the binding of transcription factors on the regulatory element of TXNDC9 in HepG2 cells. We found that FOXA1, JUND, and FOSL2 binding sites were significantly enriched at the promoter of TXNDC9, indicating a direct regulation of TXNDC9 by these transcription factors in HepG2 cells Fig. . To confThioredoxins have been reported to be involved in many malignant cancers, but their role in HCC remains unclear. In this study, we have demonstrated that TXNDC9, a member of thioredoxin superfamily, functioned as an oncogene in HCC. TXNDC9 was overexpressed in HCC and a high level of TXNDC9 was correlated with poor prognosis of HCC. Ablation of TXNDC9 significantly reduced the cancer phenotype of HCC. TXNDC9 directly interacted with MYC and stabilized the MYC protein and TXNDC9 knockout decreased the binding of MYC on chromatin. In addition, we demonstrated that TXNDC9 overexpression occurred via transcriptional activation by FOXA1, JUND, and FOSL2. Taken together, we have shown that the FOXA1/JUND /FOSL2-TXNDC9-MYC regulatory network promoted cancer progression of HCC Fig. .16. Although this study didn’t highlight the mechanism underlying TXNDC9 mediated oncogenesis, it emphasized the possibility of the important role of TXNDC9 in tumorigenesis. In addition, studies on TXNDC5, another member of the thioredoxin family, have revealed that it played a vital role in the proliferation and migration of tumor cells, acting as a tumor-enhancing gene20. Moreover, the tumorigenic effects of TXNDC5 have been described in cervical carcinoma21, lung cancer22, and liver cancer risk23. Taken altogether, the results from this and other studies demonstrate that thioredoxin overexpression might represent a signature in cancer, and that over-activated thioredoxin signaling might be a prognostic marker in cancer, and might also serve as potential therapeutic targets in cancer therapy.In this study, we also demonstrated that TXNDC9 was over-expressed in HCC and associated with a larger tumor, poor histological type, and higher TNM stage. In addition, overexpressed TXNDC9 was associated with lower survival, suggesting that higher TXNDC9 expression could increase the progression and metastasis of HCC. Of note, another study has also observed overexpression in TXNDC9 in colorectal tumor tissues25. Amplification of MYC directly drives the tumorigenesis of many cancers by regulation of genes necessary for cell proliferation, cell invasion, and metastasis26. Studies on peroxiredoxins, another super-family of redox catalysts, demonstrated that Peroxiredoxin 1 promoted cell proliferation via direct interaction with MYC and alteration of the transcription of c-MYC targets27, suggesting a direct association between a redoxin domain containing protein and c-MYC. Generally, the thioredoxin super family is mostly involved in redox and oxidative progress regulation28. The thioredoxin domain of TXNDC9 could affect the oxidative stress and generate redox imbalance in HCC cells. In addition, many studies on MYC regulation have demonstrated a direct link between redox stress and MYC activation29. Thus, it may be that TXNDC9 overexpression directly alters the cellular redox state of HCC, and thus affects the transcriptional activity of MYC, which leads to HCC cell proliferation. Future studies need to be done to understand the association between thioredoxins and MYC function.In our study, we found TXNDC9 directly interacted with MYC and the knockout of TXNDC9 directly decreased the expression of MYC and its binding on chromatin, indicating that the importance of TXNDC9 in the protein levels of MYC and the transcription network governed by MYC. MYC is a key oncogene in gene regulation30. Here we observed that the FOSL2, a member of Fos family, and JUND, a key member of Jun family, co-localized at the promoter of TXNDC9 and activated its transcription. FOXA1, JUND, and FOSL2 have previously been demonstrated to be involved in HCC32, but their association with thioredoxin have not been discussed. Our demonstration that FOXA1, JUND, and FOSL2 directly regulate TXNDC9 suggests that their role in tumorigenesis may be based on the regulation of thioredoxin domain containing proteins.In addition to downstream factors involved in TXNDC9 mediated oncogenesis in HCC, we also observed that the upstream regulator of TXNDC9 was FOXA1, JUND, and FOSL2. We demonstrated that FOXA1, JUND, and FOSL2 co-localized at the promoter of TXNDC9. All three of these transcription factors contribute mainly to transcriptional activation in cell. Indeed, we observed that knockdown of the expression of these factors led to decreased expression of TXNDC9 in HepG2 cells. The Fos gene family members dimerize with the Jun family forming the transcription factor AP-1 complex, which trans-activates the transcription of its downstream targetsIn conclusion, the current study demonstrated that the TXNDC9 was over-expressed in HCC and the overexpressed TXNDC9 was associated with poor prognosis of HCC. In addition, the FOXA1/JUND/FOSL2-TXNDC9-MYC axis is the basis of TXNDC9 mediated oncogenesis in HCC. Finally, the potential implication of TXNDC9 as a therapeutic target in HCC should be explored in future studies.Gene set enrichment analysis on biological process invovled in TXNDC9 knockoutSupplementary Figure LegendsTXNDC9 was overexpressed in HCCImpact of TXNDC9 on the mRNA expression of MYC partnersMG132 induced the resorted expression of genes down-regulated upon TXNDC9 knockout/knockdownOverexpression of MYC abolished the TXNDC9-knockout induced cell apoptosisTXNDC9 localized in the cytoplasm in HCC cellsDNA methylation of TXNDC9 were similar and rare in HCC and adjacent normal tissuesGenes differentially expressed upon TXNDC9 knockoutGene set enrichment analysis on molecular function invovled in TXNDC9 knockout