A dual role of EZH2 in regulating A-to-I RNA editing and mRNA stability through ADAR

Domain: neuroscience
Published: 2026-03-26
Editorial outcome: major_revision

Abstract
Adenosine-to-inosine (A-to-I) RNA editing, catalyzed by adenosine deaminases acting on
RNA (ADARs), is a widespread modification in metazoans. Cumulative evidence has revealed the
altered A-to-I editing profiles in cancers, but the underlying mechanism remains unclear. Here, we
discover the well-known histone lysine methyltransferase enhancer of zeste homologue 2 (EZH2)
as an unexplored ADAR interactor and editing regulator in prostate cancer (PCa). Through
competing with interleukin enhancer binding factor 2 (ILF2) for ADAR1 binding, EZH2 reshapes
the substrate selectivity of ADAR1 and thus exhibits a bidirectional role in editing regulation.
Moreover, EZH2 depletion induces the translational repression of transportin-1 (TRN1), which
further results in the accumulation of cytoplasmic ADAR1p110 isoform to protect many oncogenic
transcripts from degradation. Consistently, depletion of ADAR1 dramatically enhances the
sensitivity of cancer cells and tumors to EZH2 selective degraders. Collectively, our study sheds
new light on a link between two layers of epigenetic regulations at histone modification and RNA
editing levels, demonstrates a previously uncharacterized role of EZH2 in RNA editing and mRNA
stability independently of its lysine methyltransferase activity, and reveals the significance of
EZH2-ADAR1 cascade in governing RNA editing and mRNA stability, which may provide
additional perspectives for the advancement of EZH2-targeting cancer therapies.

Introduction
Adenosine deaminases acting on RNA (ADARs) are enzymes that catalyze the adenosine-to-
inosine (A-to-I) RNA editing in double-stranded RNA (dsRNA) substrates[1-3]. To date, millions
of A-to-I editing sites have been identified across the human transcriptome[4]. Since inosine (I) is
considered as guanosine (G) by the cellular machinery, these editing events have the potential to
generate diverse biological consequences such as amino acid substitution, splicing pattern
modification and change of noncoding RNA regulations[5]. In humans, ADAR1 (ADAR) and
ADAR2 (ADARB1) are the only two catalytically active ADAR family members. Unlike ADAR2
which is relatively abundant in the central nervous system (CNS), ADAR1 is widely expressed
throughout the body and contribute to the transcriptomic diversity of many cell types[6]. Notably,
aside from RNA editing, ADAR1 further regulates microRNA (miRNA) biogenesis[7, 8] and
mRNA stability[9, 10] independent of A-to- I enzymatic activity.

Recent genomic studies have revealed the aberrant A-to-I editing patterns in a broad range of
tumor types including prostate cancer (PCa), which could be largely attributed to the dysregulation
of ADAR1 in tumors[11-14]. However, how ADAR1 and A-to-I editing contribute to PCa
progression remains controversial. For instance, multiple previous works found that hypo-editing
events are more frequent than hyper-editing events in PCa, indicating a repressed ADAR1
activity[11, 15, 16]. In contrast, it has been reported that the long noncoding RNA (lncRNA)
prostate cancer antigen 3 (PCA3) could form a duplex with pre-mRNA of tumor suppressor
PRUNE2 in PCa, which subsequently downregulates PRUNE2 through ADAR1-mediated RNA
editing[17]. In this case, elevated ADAR1 expression may benefit PCa development through
elimination of PRUNE2. Furthermore, several missense mutations have been identified in
androgen receptor (AR) codons, which are introduced by RNA editing mechanisms[18].

Interestingly, inhibition of either ADAR1 or ADAR2 had minimal effect on the editing level of AR
transcripts, implicating the existence of an alternative pathway to mediate AR nucleotide
transitions in PCa cells. As such, it is of great importance to decipher the link between ADAR1
activity and altered A-to-I editing landscape in PCa and uncover the molecular basis by which
ADAR1 is modulated to provide selective advantages for PCa tumorigenesis.

As the catalytic subunit of polycomb repressive complex 2 (PRC2), EZH2 canonically directs
deposition of histone H3 lysine 27 trimethylation (H3K27me3) to maintain a repressive chromatin
state[19, 20]. Overabundance of EZH2 is closely correlated with aggressiveness and poor
prognosis of solid tumors, making it an attractive therapeutic target[21, 22]. However, regardless
of the massive reactivation of PRC2-repressed tumor suppressor genes, abolishing the enzymatic
activity of EZH2 alone has shown limited efficacy in treating EZH2-dependent malignancies such
as PCa[23, 24]. Meanwhile, compelling evidence including ours has proved that EZH2 could exert
multifaceted tumorigenic functions beyond H3K27me3 and PRC2[25-28].

In the present study, we disclose a dual mechanism whereby EZH2 remodels both the A-to-I
editome and mRNA turnover landscape in PCa via regulating both editing-dependent and -
independent functions of ADAR1. On these bases, we aim to delineate how the EZH2-ADAR1
cascade influences tumorigenesis and cancer therapy.

Results
EZH2 directly interacts with ADARs in the nucleus
To identify previously uncharacterized EZH2 binding partners in PCa, we queried our
previous co-immunoprecipitation/mass spectrometry (co-IP/MS) data[29] and the constitutively
expressed p110 isoform of ADAR1 (ADAR1p110) appeared as a potential EZH2-bound protein.
To confirm this observation, we performed co-IP assay using a PCa cell line C4-2. In accordance
with the co-IP/MS results, IP of EZH2 pulled down ADAR1p110 and the other two PRC2 core
components, EED and SUZ12 (Fig. 1a), while only EZH2 could be detected in the reciprocal
ADAR1 IP products (Fig. 1b). This interaction was further validated by IP-IB analysis using
LuCaP 35CR[30], a castration-resistant PCa (CRPC) patient-derived xenograft (PDX) model (Fig.
1c and 1d). Both EZH2 and ADAR1p110 proteins are known to be enriched in the nucleus. To
capture the in situ EZH2-ADAR1 interactions, proximity ligation assay (PLA) was next conducted
in C4-2 cells. As anticipated, PLA dots representing the endogenous EZH2-ADAR1 interaction
were accumulated in the nuclei of control cells, but almost completely abolished upon EZH2 or
ADAR1 knockdown (Fig. 1e). Although ADAR2 was not listed as an interacting partner of EZH2
by co-IP/MS, the structural and functional similarities of two ADARs prompted us to test the
potential of EZH2-ADAR2 interaction. Since ADAR2 expression is too low and hard to detect in
PCa model, we ectopically overexpressed Flag-tagged ADAR2 in C4-2 cells, followed by co-IP
assay using anti-Flag antibody. As predicted, EZH2, but not EED or SUZ12, was specifically
pulled down by exogenous ADAR2 (Supplementary Fig. 1a). Though both EZH2 and ADARs
bear critical RNA-binding capacities[31], the two EZH2-ADAR interactions were well preserved
following Ribonuclease A (RNase A) treatment, implying that EZH2 does not rely on RNA for
ADAR binding (Fig. 1f and Supplementary Fig. 1b). To further exclude the involvement of other
factors, such as DNA or histone, in the EZH2-ADAR interactions, purified recombinant proteins
of GST-tagged EZH2 and Flag-tagged ADAR1/2 were mixed and subjected to GST pull down
assay. In this context, EZH2 proteins were still able to pull down ADAR1/2 proteins (Fig. 1g and
Supplementary Fig. 1c), confirming that EZH2 and ADAR1/2 could directly interact with each
other. In addition, these findings were consolidated by AlphaLISA assays using purified EZH2 and
ADAR1/2 proteins, which revealed a strong binding affinity for both interactions (Fig. 1h and
Supplementary Fig. 1d-1f).

EZH2 mainly comprises two SANT protein-protein interaction domains, one cysteine-rich
CXC domain and an enzymatic SET domain (Fig. 1i). ADAR1p110 contains a single Z-DNA
binding domain, three dsRNA binding domains (dsRBDs) and a deaminase domain in the C-
terminal (Fig. 1j), while ADAR2 consists of two dsRBDs followed by one deaminase domain
(Supplementary Fig. 1g). To uncover the interacting domains that are required for the EZH2-
ADAR interactions, we co-transfected serially truncated mutants of Myc-tagged EZH2 or Flag-
tagged ADAR1/2 constructs with their full-length partner plasmids into HEK293T cells for co-IP
analyses. As revealed in Fig. 1k, except for EZH2ΔSANT2, all other truncated mutants and full-
length of EZH2 pulled down full-length ADAR1. Meanwhile, only ADAR1ΔdsRBD123 failed to
pull down full-length EZH2 (Fig. 1l). In terms of ADAR2, EZH2ΔSANT2 was also characterized
as the only EZH2 mutant which was unable to pull down full-length ADAR2 (Supplementary
Fig. 1h), while full-length EZH2 could be detected in all ADAR2 IP products excluding
ADAR2ΔdsRBD12 (Supplementary Fig. 1i). Collectively, these findings suggest that all dsRBDs
of both ADAR proteins retain the EZH2-binding capacity through EZH2’s SANT2 domain.

EZH2 reshapes the A-to-I editome in PCa cells primarily through ADAR1
The aforementioned interactions led us to investigate whether EZH2 could participate in the
RNA editing regulation. To this end, we perturbed the expression of EZH2 as well as two ADARs
in C4-2 cells by siRNAs, followed by RNA-seq to map the editing changes. To avoid the biases
introduced by single nucleotide polymorphisms (SNPs) or somatic mutations, we further
sequenced the entire genome of C4-2 cells as reference. The A-to-G (I) conversion was ranked as
the most frequent type of nucleotide changes, accounting for 73.6% of the changes observed across
the transcriptome of C4-2 cells (Fig. 2a). Notably, although EZH2 deficiency did not appear to
have any noticeable impact on ADAR expression, the overall A-to-I editing ratio was significantly
reduced in EZH2-deficient C4-2 cells when compared to the control group (Fig. 2b and
Supplementary Fig. 2a). As controls, a dramatic downregulation of global editing was also found
upon ADAR1 depletion, while further knockdown of already lowly expressed ADAR2 only led to
a slight editing decrease in C4-2 cells (Fig. 2b). Similar results were also found when using PCa
patient-derived organoids (PDOs) MDA-PCa-174 (Supplementary Fig. 2b and 2c). To
consolidate this observation, we first tested a series of public EZH2-knockdown RNA-seq datasets
in PCa and other cancer cell lines to reconfirm the decreased editing pattern (Supplementary Fig.
2d). Moreover, after dividing the Stand Up To Cancer (SU2C) metastatic PCa patient samples
based on EZH2 expression[32], the editing level in EZH2-high group was statistically higher than
the EZH2-low group without any change in terms of ADAR1 expression (Supplementary Fig.
2e). Interestingly, treatment of C4-2 cells with EZH2 enzymatic inhibitors of GSK126 and
EPZ6438 barely affected the global editing level (Supplementary Fig. 2f), indicating that EZH2
mainly regulates A-to-I editing in a way independently of its methyltransferase activity.

With respect to the individual editing event, the majority of EZH2-affected A-to-I editing sites
reside in the three prime untranslated region (3′-UTR), followed by intergenic and intronic regions
(Fig. 2c). Intriguingly, both over-edited (editing ratio increased) and under-edited (editing ratio
decreased) sites (N =244 vs N =548) could be identified in EZH2-deficient C4-2 cells (Fig.
2d and Supplementary Data 1), indicating an opposing effect of EZH2 in controlling A-to-I
editing. In parallel, most sites altered upon ADAR1 depletion were under-edited (n =69 vs
n =4,326), while the number of over-edited sites was inversely higher than the under-edited
sites in ADAR2-deficient cells (n =502 vs n =312) (Fig. 2d and Supplementary Data 1).
We then defined the sites that showed significantly decreased editing frequency upon ADAR1/2
knockdown as the ADAR1/2-mediated editing sites to compare with our identified EZH2-affected
sites. As revealed in Fig. 2e and 2f, 70.4% (n=386) of under-edited and 20.5% (n=50) of over-
edited sites upon EZH2 deficiency could be classified as the ADAR1-mediated editing sites, while
only 17.5% (n=96) of under-edited and one single over-edited site upon EZH2 deficiency were
overlapped with the ADAR2-mediated editing sites. Moreover, among the 548 EZH2-regulated
under-edited sites, 80 sites are co-regulated by all three factors, while only 16 sites are shared
between EZH2 and ADAR2 without ADAR1 involvement (Supplementary Fig. 2g). In addition,
when all the ADAR1/2-shared editing sites were removed, we could still observe a highly
significant overlap between the ADAR1-regulated sites and EZH2-regulated sites
(Supplementary Fig. 2h). In contrast, only a comparatively weaker but still statistically
significant overlap could be found between EZH2 and ADAR2-regulated sites (Supplementary
Fig. 2i). All these data proved that EZH2 mainly modulates A-to-I editing in PCa cells through
ADAR1, while also exerting a modest but measurable influence via ADAR2.

Next, to test the accuracy of our sequencing results, 30 EZH2-affected sites (20 under-edited
and 10 over-edited) were randomly selected for validation using an RNA editing fingerprint
approach of RNA editing site-specific quantitative PCR (RESSq-PCR) [33]. As a result, 80% (16)
under-edited sites and 80% (8) over-edited sites were successfully validated to be regulated by
EZH2 in C4-2 cells (Fig. 2g, 2h and Supplementary Data 2). Consistently, most of the tested
EZH2-affected sites also underwent a drastic under-editing upon ADAR1 knockdown but
remained unchanged or even over-edited upon ANDAR2 depletion. Moreover, to confirm that these
EZH2-affected editing changes are not restricted in a certain prostate cell type, we overexpressed
EZH2 in human primary prostate epithelial cells (PrEC) which bear rare endogenous EZH2
expression (Supplementary Fig. 2j). RESSq-PCR results showed that the majority of our tested
sites could also been validated in EZH2-overexpressing PrEC model, in which circumstance the
editing ratios were changed towards an opposite direction as occurred in EZH2-knockdown C4-2
cells (Supplementary Fig. 2k, 2l and Supplementary Data 2). In summary, our data suggested
that, mainly through ADAR1, EZH2 reshapes the A-to-I editing profiles of PCa cells in a
bidirectional manner.

Identification of chr12: 69237519 at MDM2 transcript as a functional editing site affected by
EZH2 in PCa
To gain a deeper insight into the EZH2-remodeled RNA editing in PCa, we decided to perform
a comprehensive investigation on a specific editing site (chr12: 69237519) at the 3′-UTR of
MDM2 transcript, which was significantly under-edited upon EZH2/ADAR1 depletion
(Supplementary Data 1). By utilizing Sanger sequencing and RESSq-PCR assays, we first
validated this site in both C4-2 and PrEC models (Fig. 3a, 3b, Supplementary Fig. 3a, 3b).
Strikingly, conjoint analyses of the Cancer Genome Atlas (TCGA) and Genotype-Tissue
Expression (GTEx) databases observed a higher editing frequency of chr12: 69237519 in PCa
tissues relative to normal prostate controls, which was accompanied by the overexpression of
EZH2, but not ADAR1 (Fig. 3c). Moreover, instead of ADAR1, expression of EZH2 was
positively correlated with editing level of this site in PCa (Fig. 3d). Indeed, localized PCa is not
featured by abnormal ADAR1 expression, as immunohistochemistry (IHC) staining of ADAR1
proteins in a series of PCa specimens observed no significant difference between tumor and the
adjacent benign tissues (Supplementary Fig. 3c and 3d). Therefore, other than ADAR1, elevated
expression of EZH2 is more likely to be responsible for the increased editing level of chr12:
69237519 in PCa.

Of note, this specific site is harbored in the seeding sequences of miR-204 and -211 (Fig. 3e,
the upper panel). Consistently, endogenous MDM2 expression was markedly decreased in C4-2
cells transduced with mimics of both miRNAs (Supplementary Fig. 3e), confirming their
regulatory connections. To examine whether the A-to-I (G) conversion at chr12: 69237519 could
affect the binding of two miRNAs to MDM2 transcript, dual-luciferase reporters with either
wildtype (WT, unedited) or edited MDM2 3′-UTR were introduced for investigation (Fig. 3e, the
lower panel). After overexpression of miR-204 or -211 mimics, both WT and edited reporters
showed a decreased reporter activity (Fig. 3f). Remarkably, the reduction was much more evident
in WT group as compared to the edited group, suggesting that RNA editing of chr12: 69237519
enables MDM2 to partially evade targeting by miR-204 and -211. In keeping with our hypothesis
that EZH2 maintains the over-editing of chr12: 69237519, EZH2 suppression in C4-2 cells led to
the downregulation of MDM2 without affecting the expression of both miRNAs (Fig. 3g and
Supplementary Fig. 3f). In addition, EZH2 depletion failed to further decrease MDM2 expression
in ADAR1-deficient cells (Supplementary Fig. 3g), supporting a notion that EZH2 promotes
MDM2 expression through modulation of ADAR1-mediated RNA editing. Taking chr12:
69237519 as an example, we next investigated whether the methyltransferase activity is required
for EZH2 to regulate RNA editing. To achieve this, we overexpressed various RNAi-resistant
EZH2 mutants in EZH2-deficient C4-2 cells, followed by measurement of chr12: 69237519
editing ratio and MDM2 expression in each group. As shown in Fig. 3h and Supplementary Fig.
3h, re-expression of either WT or catalytically dead H689A-mutant EZH2, but not EZH2ΔSANT2,
could fully rescue the editing of chr12: 69237519. Meanwhile, the protein level of MDM2 was
coordinately changed (Fig. 3i). Together, these observations prove that interaction with ADAR1 is
essential for EZH2 to mediate RNA editing of MDM2 transcript in a methylation-independent
manner.

Selective binding to EZH2 or ILF2 determines the editing substrates of ADAR1
We next sought to discover the mechanism underlying EZH2-mediated site-specific editing
regulation. In accordance with our RNA-seq data, western blot analysis revealed no alteration in
ADAR1p110 expression upon EZH2 depletion (Supplementary Fig. 4a). Similarly, neither EZH2
expression nor H3K27me3 level was changed upon ADAR1 suppression (Supplementary Fig.
4b). Since EZH2 is capable of methylating proteins other than histone[34], we next wondered
whether ADAR1p110 is a distinct EZH2 substrate. ADAR1p110 proteins from control or EZH2-
deficient C4-2 cells were immunoprecipitated and subjected to western blot to measure the lysine
methylation levels. As depicted in Supplementary Fig. 4c, both mono-/di- and tri-methylation
levels of ADAR1p110 were barely changed after EZH2 depletion, suggesting that EZH2 is
unlikely to methylate lysine residues of ADAR1p110. Homodimerization of ADARs is critical for
their enzymatic activities[35]. However, co-IP results showed that EZH2 knockdown did not affect
the abundance of Myc-tagged ADAR1p110 coimmunoprecipitated with Flag-tagged ADAR1p110
(Supplementary Fig. 4d), excluding the possibility that EZH2 disturbs the formation of ADAR1
homodimer.

We then focused on the change of ADAR1 interactome since the preference of ADAR1 to
interact with other RNA-binding proteins (RBPs) could dictate its editing substrate specificity[36].
By conducting the quantitative co-IP/MS assay in C4-2 cells, we observed that, more interleukin
enhancer binding factor 2 (ILF2), a previously reported editing regulator[36], bound to
ADAR1p110 in EZH2-knockdown groups (Fig. 4a). To validate this proteomics result, we pulled
down endogenous ADAR1p110 along with its interacting partners from control or EZH2-deficient
C4-2 cells, followed by western blot to compare the abundance. Whereas EZH2 knockdown had
no impact on ILF2 expression, much more ILF2 proteins could be coimmunoprecipitated by
ADAR1p110 in EZH2-deficient cells than those in control cells (Fig. 4b). As compared, ILF3, a
related protein of ILF2 which also functions in editing regulation[36], showed no change in
ADAR1-binding capability upon EZH2 deficiency. We then hypothesized that EZH2 competes
with ILF2 to interact with ADAR1. To validate this, we first confirmed that ILF2 binds to the first
dsRBD (dsRBD1) of ADAR1p110 (Fig. 4c). Moreover, AlphaLISA assay revealed a gradually
reduced EZH2-ADAR1 interaction signals as the concentration of ILF2 proteins increased (Fig.
4d and Supplementary Fig. 4e), indicating that the binding of ADAR1p110 with EZH2 or ILF2
is mutually exclusive. To consolidate this finding, we pulled down endogenous ADAR1p110 along
with its interacting EZH2 proteins in the context of ILF2 deficiency. Interestingly, although ILF2
inhibition led to an unexpected EZH2 downregulation, more EZH2 proteins could still be
coimmunoprecipitated by ADAR1p110 in ILF2-knockdown groups (Supplementary Fig. 4f).
Consistently, more PLA dots representing the EZH2-ADAR1 interaction were also captured in
ILF2-deficient C4-2 cells (Supplementary Fig. 4g). Size-exclusion chromatography analyses
were then carried out using nuclear extracts from control and EZH2-deficient C4-2 cells. While
the elution profile of ADAR1p110 remained invariable upon EZH2 knockdown, EZH2
suppression led to an obvious shift of ILF2 elution profile towards ADAR1p110 fractions (Fig.
4e), reflecting a stronger ILF2-ADAR1 interaction.

To deeply dissect the consequence of competitive ADAR1 binding between EZH2 and ILF2,
we performed the enhanced crosslinking and immunoprecipitation-sequencing (eCLIP-seq) in C4-
2 cells to define the RNA targets of EZH2, ADAR1 and ILF2, respectively (Supplementary Data
3). As a result, the group of RNAs that are bound by each RBP was significantly overlapped with
each other (Fig. 4f). After integration with our aforementioned RNA-seq results, all three RBPs
showed evidence of binding at the A-to-I editing sites (Fig. 4g). Remarkably, the number of ILF2-
bound RNAs undergoing over-editing upon EZH2 knockdown was much higher than expected
(Fig. 4h), which accounts for 61% of the RNAs with EZH2 deficiency-induced over-edited sites
(Fig. 4i). In comparison, the number of EZH2-bound RNAs undergoing under-editing upon EZH2
knockdown was also higher than expected (Fig. 4j), which accounts for 59% of the RNAs with
EZH2 deficiency-induced under-edited sites (Fig. 4k). Further analysis suggested that, as
compared with ILF2, EZH2-binding regions preferentially overlap with the under-edited sites
upon EZH2 knockdown. In contrast, the ILF2-binding regions show greater overlap with the over-
edited sites following EZH2 depletion (Supplementary Fig. 4h). All these findings suggested that
the strengthened ADAR1-ILF2 interaction upon EZH2 suppression may change the editing
substrate selectivity of ADAR1, which subsequently results in the bidirectional alteration of
editing in EZH2-deficient cells. Therefore, for those EZH2-bound RNAs without ILF2-binding
capacity (e.g. MDM2, as depicted in Fig. 4l), their editing sites are prone to be under-edited upon
EZH2 deficiency (e.g. chr12: 69237519 of MDM2, as shown in Fig. 3). Meanwhile, for those
ILF2-bound RNAs lacking EZH2-binding capacity (e.g. SYTL1, as depicted in Supplementary
Fig. 4i), their editing sites are more likely to be over-edited upon EZH2 depletion (e.g.
chr1_27678712 of SYTL1, as presented in Fig. 2h).

EZH2 suppression induces cytoplasmic translocation of ADAR1p110 through translational
downregulation of TRN1
Since ADAR1p110 mainly edits RNA in the nucleus but further exerts editing-independent
functions in the cytoplasm[10, 37], we next examined the subcellular distribution of ADAR1p110
under EZH2 depletion. As expected, endogenous ADAR1p110 was found predominantly in the
nucleus of control C4-2 cells. However, the cytoplasmic fraction of ADAR1p110 was significantly
increased in the absence of EZH2 (Fig. 5a). To further validate this finding, we transfected GFP-
tagged ADAR1p110 plasmids into C4-2 cells and visualized by fluorescence microscopy along
with western blot analysis. As presented in Fig. 5b and Supplementary Fig. 5a, repression of
EZH2 also stimulated the cytoplasmic accumulation of GFP-tagged ADAR1p110, with a
phenotype similar to the previously reported stress-activated ADAR1p110 shuttling to the
cytoplasm[10].

To unveil the reason for the translocation of ADAR1p110 upon EZH2 suppression, we
checked the expression of transportin-1 (TRN1, also known as TNPO1) and exportin-5 (XPO5),
which controls the nuclear import and export of ADAR1p110 proteins, respectively[38]. As shown
in Fig. 5c, the protein level of TRN1, but not XPO5, was substantially reduced upon EZH2
knockdown. Intriguingly, TRN1 mRNA level was barely altered upon EZH2 suppression, albeit
the significant change of protein level (Supplementary Fig. 5b). This finding was reminiscent of
our recent report that EZH2 could exert a PRC2-independent role to facilitate the internal ribosome
entry site (IRES)-dependent translation initiation in cancer cells through fibrillarin (FBL) [27]. By
checking our ribosome profiling (Ribo-seq) data within this article, we found that TRN1 was
classified as a buffering down gene with minor decrease at mRNA level but significant reduction
at protein level upon EZH2 inhibition, which matched our current observations. To test whether
TRN1 is regulated by EZH2 at translational level, we first confirmed that TRN1 expression was
also decreased upon FBL knockdown (Supplementary Fig. 5c). In addition, treatment of C4-2
cells with EZH2 protein inhibitor DZNep which decreases EZH2 protein level, but not enzymatic
inhibitors of GSK126 and EPZ6438, led to the downregulation of TRN1 (Fig. 5d), indicating that
EZH2 controls TRN1 expression beyond its methyltransferase activity. A potential IRES element
has been predicted at the 5′-UTR of TRN1[39]. Next, we cloned this putative IRES sequence into
a bicistronic luciferase vector for analysis (Fig. 5e, the upper panel). In line with our hypothesis,
TRN1 IRES showed a reduced activity in both FBL- and EZH2-deficient cells (Fig. 5e, the lower
panel). To reinforce these observations in vivo, we first searched the TCGA database and
confirmed that mRNA level of TRN1 was similar between normal prostates and PCa tissues (Fig.
5f). In sharp comparison, IHC results of PCa tissue microarray (TMA) slide revealed that TRN1
proteins were hardly detected in benign prostate tissues but upregulated significantly with the
advancement of PCa (Fig. 5g). Meanwhile, IHC assay was conducted in serially sectioned PCa
TMA slides using both anti-EZH2 and anti-TRN1 antibodies. As a result, a strongly positive
correlation was found between expressions of EZH2 and TRN1 (Fig. 5h and 5i), further
supporting the EZH2-mediated translational regulation of TRN1.

Finally, we aimed to determine whether diminished TRN1 expression is responsible for the
enrichment of cytoplasmic ADAR1p110 upon EZH2 suppression. We first observed that inhibition
of TRN1 alone resulted in the dramatic retention of ADAR1p110 in the cytoplasm
(Supplementary Fig. 5d). More importantly, ectopic expression of TRN1 in EZH2-deficient cells
significantly restored the nuclear localization of ADAR1p110 (Fig. 5j and 5k). Above all, these
data indicate that EZH2-mediated regulation of TRN1 translation has a downstream effect on the
subcellular localization of ADAR1p110, which may contribute to the observed reduction of global
editing level in EZH2-deficient PCa cells (Fig. 2b and Supplementary Fig. 2b).

EZH2 influences the dynamic of mRNA stability through cytoplasmic ADAR1p110
Independent of its RNA editing function which is mostly restricted in the nucleus, the
cytoplasmic ADAR1p110 competes with staufen-1 (STAU1) to bind to a set of gene transcripts
with 3′-UTR dsRNA structures and thus antagonizes the STAU1-mediated mRNA decay[10]. To
investigate whether EZH2 depletion could affect the mRNA degradation process through
translocation of ADAR1p110, we treated control, ADAR1- and EZH2-deficient C4-2 cells with
Actinomycin D (ActD) to block nascent RNA synthesis, and collected samples at a series of time-
points for RNA-seq (Fig. 6a). As expected, loss of ADAR1 significantly reduced the RNA stability
g stability
globally, while an overall increased RNA half-life was observed in EZH2-deficient cells (Fig. 6b).
Further analysis revealed that genes with increased RNA half-lives upon EZH2 knockdown were
significantly overlapped with genes undergoing decreased RNA half-lives upon ADAR1
knockdown (Fig. 6c). In sharp contrast, no evident overlapping was found between genes with
decreased RNA stability upon EZH2 knockdown and those with increased RNA stability upon
ADAR1 suppression (Fig. 6d). Thus, these data suggested that the role of EZH2 in modulating
RNA stability is correlated with ADAR1.

We next focused on the 3810 gene transcripts with prolonged half-lives upon EZH2
deficiency while weakened stability upon ADAR1 suppression (Fig. 6c and Supplementary Data
4), which are deemed to be regulated by the EZH2-ADAR1 cascade in terms of RNA decay rate.
Unexpectedly, these genes were particularly enriched into a number of cancer-specific oncogenic
pathways (Fig. 6e), implying that elimination of EZH2 may not only release the epigenetic
silencing of key tumor suppressors at transcription level, but also inversely stabilize a set of
oncogenes post-transcriptionally. To further confirm this, we selected four gene transcripts of
CCNG1, ATM, YES1 and SMARCD1 for validation. All of these candidates are known ADAR1
binding targets with 3′-UTR dsRNA structures[10], and exhibit oncogenic properties during PCa
development[40-43]. RT-qPCR assay was firstly conducted to monitor the remaining mRNA levels
at each time-point after ActD treatment. In line with the sequencing results, mRNAs of all tested
oncogenes were stabilized upon EZH2 knockdown while destabilized in response to ADAR1
depletion (Fig. 6f). Simultaneously, the steady-state mRNA level of each candidate and the final
protein production were concordantly changed following the alterations in mRNA half-lives (Fig.
6g and Supplementary Fig. 6a). To explore whether EZH2 also controls the expression of these
oncogenes through conventional transcriptional repression, we labeled and captured nascent RNA
from control or EZH2-deficient C4-2 cells, followed by RT-qPCR analysis. As a positive control,
the well-known epigenetic EZH2 target CNR1 displayed an accelerated nascent transcription in
EZH2-deficient cells. With respect to the tested oncogenes, only SMARCD1 showed a similar
pattern with CNR1, whereas the rest candidates were basically unaffected in mRNA synthesis upon
EZH2 suppression (Supplementary Fig. 6b). Accordingly, these results confirmed mRNA
stabilization as a major contributor for the upregulation of these oncogenes in response to EZH2
depletion. The ADAR1p110-mediated inhibition of mRNA decay relies on its competitive binding
to dsRNAs against STAU1[10]. To this end, RNA immunoprecipitation-qPCR (RIP-qPCR) assay
was next performed to measure the enrichment of the selected oncogenic transcripts to
ADAR1p110 or STAU1 proteins. As presented in Supplementary Fig. 6c, EZH2 knockdown
strengthened the binding of ADAR1p110 to all test candidates. In comparison, the enrichment of
STAU1 to these candidates were substantially attenuated (Fig. 6h and 6i). Notably, this reduction
was markedly rescued upon additional ADAR1 knockdown, proving that EZH2 deficiency
stimulates cytoplasmic ADAR1 to protect these oncogenic transcripts from STAU1 binding and
degradation.

Targeting EZH2 and ADAR1 synergistically inhibit PCa progression
Regardless of the application of EZH2-targeting strategy in a number of clinical trials for
advanced PCa treatment[44], our data clearly suggested a limitation of this therapeutics: induces
the migration of ADAR1p110 into cell cytoplasm to stabilize a series of prostate oncogenic
transcripts. Therefore, it is desirable to evaluate whether targeting ADAR1 for depletion could
sensitize cancer cells to EZH2 inhibitors. MS8815, a recently discovered von Hippel-Lindau (VHL)
-recruiting EZH2 proteolysis targeting chimera (PROTAC) degrader[45], was employed for EZH2
targeting in our model. By utilizing EPZ6438 as the EZH2 binder while linked with an E3 ligase
ligand, MS8815 is capable of selectively targeting EZH2 for ubiquitination and degradation.
Treatment of C4-2 cells with MS8815 significantly reduced EZH2 and H3K27me3 levels in a
dose-dependent manner (Fig. 7a). Meanwhile, EED and SUZ12 protein levels were decreased
synchronously, indicating a disruption of the whole PRC2 complex. In contrast, MS177, another
PROTAC-based EZH2 degrader[26], showed a much weaker EZH2 elimination effect in PCa cells
(Fig. 7a); while EPZ6438, the parental inhibitor of MS8815, could only erase H3K27me3 marks
(Fig. 7b). Moreover, MS8815 exhibited a much lower half-maximal inhibitory concentration (IC50)
in C4-2 cells as compared with MS177 and EPZ6438 (Fig. 7c), indicating a much more powerful
antiproliferative activity. Given that the anti-tumor ability of MS8815 has never been tested in vivo
before, we next treated mice bearing C4-2 cells-derived xenograft with a range of MS8815 dosages
and monitored the effects. As shown in Supplementary Fig. 7a and 7b, intraperitoneal (i.p.)
injection of MS8815 as low as 25 mg per kg body weight daily was able to trigger tumor repression,
and the anti-tumor effect became more significant at the dose of 50 mg per kg body weight per day
or above. Meanwhile, all tested doses were well tolerated by mice, as no sudden death or evident
weight loss was displayed (Supplementary Fig. 7c). Consistent with our in vitro data, the apparent
EZH2 degradation effect was also observed in xenograft tumors (Supplementary Fig. 7d).
To assess the combinatorial effect of MS8815 with ADAR1 blockade, we first generated a
stable C4-2 cell line with inducible expression of ADAR1 shRNA for murine experiments
(Supplementary Fig. 7e). As presented in Fig. 7d-7f, expression of shADAR1 significantly
enhanced the efficacy of MS8815 in mitigating tumor growth. In comparison, EPZ6438 and
shADAR1 exhibited no synergistic effect on PCa cell growth, which is compatible with our above
conclusion that EZH2 controls ADAR1 shuttling and mRNA stability in a way independent of its
enzymatic activity.

To explore the clinical potential of this strategy, we further delivered EZH2/ADAR1 siRNA-
containing Nanoparticles (NPs) into LuCaP 35CR PDX-bearing mice for gene silencing
(Supplementary Fig. 7f). We found that combinatorial treatment of siEZH2 and siADAR1
significantly improves the clinical outcomes when compared with those utilizing one siRNA only
(Fig. 7g and 7h). In summary, these data suggest that targeting the EZH2-ADAR1 cascade may
serve as a more effective strategy than targeting EZH2 alone for PCa treatment.

Methods
Ethics statement
This study was conducted in accordance with all relevant ethical regulations. All animal
experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of
Northwestern University and were performed in accordance with institutional and national
guidelines. Publicly available datasets were accessed in compliance with their respective data
access policies.

Cell lines, PDX and PDO models
Human PCa cell line C4-2 was a generous gift from Dr. Leland Chung (Cedars-Sinai), while
PC-3 was purchased from ATCC. Both PCa cell lines were grown in RPMI 1640 medium
supplemented with 10% FBS. Human Prostate Epithelial Cells (PrEC) were purchased from Lonza
and cultured using PrEGM prostate epithelial cell growth medium bullet kit (Lonza). HEK293T
cells were purchased from ATCC and maintained in DMEM medium supplemented with 10% FBS.
All cell lines and primary cells were cultured at 37 °C and 5% CO2 in a humidified atmosphere,
and were authenticated and routinely screened for Mycoplasma.

The LuCaP 35CR patient-derived xenograft (PDX) was kindly provided by Eva Corey
(University of Washington).

For patient-derived organoid (PDO), PCa PDX MDA-PCa-174 was maintained in immune-
deficient, severe combined immunodeficiency (SCID) male mice[58]. Organoids were established
using the method described previously[59]. Briefly, tumors were dissected into small pieces and
digested with 5 mg/ml collagenase type II in base medium (Advanced DMEM/F12 with 10 mM
HEPES and 2 mM Glutamax) at 37°C for 1 hour with gentle shaking. A single tumor cell
suspension was obtained after red blood cell lysis. Single cells were then seeded into a 6-well plate
in growth factor-reduced Matrigel with complete human prostate organoid culture medium (B27,
N-Acetylcysteine 1.25 mM, recombinant human EGF 5 ng/ml, recombinant human Noggin 100
ng/ml, recombinant human R-spondin-1 500 ng/ml, A83-01 500 nM, recombinant FGF-2 5 ng/ml,
prostaglandin E2 1 µM, SB202190 10 µM, Nicotinamide 10 mM, DHT 1 nM, Y27632 10 µM in
base medium) to form organoids. Knockdown of EZH2, ADAR1, and ADAR2 were achieved by
introducing 100 nM of siRNA per well in a 6-well plate, followed by two consecutive knockdowns.
RNA samples were harvested 72 hours post-second knockdown. RNA was isolated using the
Qiagen miRNA isolation kit, and knockdown was confirmed by RT-qPCR before sending samples
for sequencing.

Transfection of siRNA or miRNA mimic
All silencer-select siRNAs used in this study were purchased from Thermo Fisher (siEZH2-
1: s4916, siEZH2-2: s4917; siADAR1-1: s1007, siADAR1-2: s1008; siADAR2-1: s1010,
siADAR2-2: s1011; siFBL-1: s4820, siFBL-2: s4821; siILF2-1: s7398, siILF2-2: s7399, siTRN1:
s7932). In addition, a siRNA targeting the 3′-UTR of endogenous EZH2 (sequence: 5′-
UUGCCUUCUCACCAGCUGC-3′) was synthesized by Thermo Fisher and used for the rescue
assay. Meanwhile, all mirVana miRNA mimics were purchased from Thermo Fisher (miR-204
mimic: MC11116, miR-211 mimic: MC10168).

Lipofectamine RNAiMAX (Invitrogen) were utilized for siRNA or miRNA mimic
transfection according to the procedure of manufacturer. Mediums were renewed after 24 h and
cell samples were collected at 3 days post-transfection.

Treatment of cells with inhibitors
For EZH2 inhibitors, MS8815 and MS177 were discovered and kindly provided by Dr. Jian
Jin (Mount Sinai); GSK126 was purchased from BioVision; DZNep and EPZ6438 were purchased
from Selleck Chemicals.

Western blot (WB)
To denature proteins, cell lysates were mixed with 2 × loading buffer (Bio-Rad) and heated
at 95 °C for 10 min. Protein samples were subjected to standard SDS-PAGE and semi-dry
transferred to PVDF membranes (Roche). After blocking for 45 min in Tris-buffered saline-Tween
20 (TBST) with 5% nonfat milk, membranes were incubated with primary antibodies for 2 h at
room temperature. Then, the membranes were washed and incubated with Clean-Blot IP Detection
Reagent (Thermo Fisher, 1:500 dilution, for co-IP WB) or goat anti-mouse/rabbit IgG (H+L)-HRP
secondary antibody (GAenDEPOT, 1:5000 dilution, for normal WB) for 1 h. The signals were
developed using western ECL substrate (Bio-Rad) and captured by a Bio-Rad imaging system.
The primary antibodies used for WB in this paper were listed in Supplementary Table 1.

Real-time (RT)-qPCR analysis
To detect mRNA level, the total RNA was isolated using RNeasy Plus Mini Kit (Qiagen),
followed by reverse transcription into cDNA using Maxima H Minus First Strand cDNA Synthesis
Kit (Thermo Fisher). The cDNA samples were amplified using Universal SYBR Green Supermix
(Bio-Rad) in a QuantStudio 6 Flex Real-time PCR System (GE Healthcare) following
manufacturer's instructions. The primers used for RT-qPCR analysis were summarized in
Supplementary Table 2. The relative RNA level was calculated using the 2-ΔΔCt method with the
Ct values normalized to GAPDH.

To measure miRNA expression, miRNeasy Mini Kit (Qiagen) was used to extract small RNA
from cells. The obtained small RNA was reverse transcribed into cDNA using TaqMan miRNA
Reverse Transcription Kit (Thermo Fisher). TaqMan PCR Assay was then performed following its
guideline from Thermo Fisher. The relative miRNA level was calculated using the 2-ΔΔCt method
with the Ct values normalized to U6 snRNA. All probes used for TaqMan qPCR analysis were
ordered from Thermo Fisher (miR-204: 000508, miR-211: 000514 and U6 snRNA: 001973).

Co-immunoprecipitation (co-IP) assay
The whole-cell lysate was prepared by lysing cells in the NP-40 lysis buffer (GenDEPOT)
containing protease and phosphatase inhibitor cocktail (Thermo Fisher). After sonication and
centrifugation to remove the insoluble materials, cell lysates were pre-incubated with Dynabeads
protein A/G (Invitrogen) to remove nonspecific binding. Then, antibodies were mixed into lysates
with new-added Dynabeads and incubated at 4 °C overnight. The immune complexes were
separated by a magnetic rack and washed three times with lysis buffer. To denature proteins, beads
were mixed with 2× loading buffer (Bio-Rad) and heated at 95 °C for 10 min. Protein samples
were subjected to western blot for further analysis. For the RNase A treatment, the whole-cell
lysates were incubated with RNase A (1 μg/ml, Thermo Fisher) at 37°C for 10 min before
immunoprecipitation.

Proximity ligation assay (PLA)
The Duolink PLA Fluorescence was performed according to the manufacturer’s protocol
(Sigma). Briefly, control, EZH2-deficient and ADAR1-deficient C4-2 cells grown on coverslips
were fixed by 4% paraformaldehyde for 10 min and permeabilized in phosphate-buffered saline
(PBS) containing 0.5% Triton X-100 for 10 min. Subsequently, cells were blocked by blocking
solution for 1 h at 37 °C and incubated with mouse anti-EZH2 and rabbit anti-ADAR1 antibodies
for 2 h. After washing, the slide was further incubated with PLUS and MINUS PLA probes for 1
h at 37 °C, followed by ligation and amplification. The slide was finally mounted using Duolink
In Situ Mounting Medium with DAPI (Sigma) and visualized by confocal microscopy.

GST pull down
To perform GST pull down assay, 1 μg recombinant GST-tagged EZH2 protein was mixed
with 1 μg Flag-tagged ADAR1/2 in 1 ml of NP-40 lysis buffer (GenDEPOT) with protease and
phosphatase inhibitor cocktail (Thermo Fisher). The mixture was incubated with pre-washed
Glutathione Sepharose 4B (GE Healthcare) for 1 h at room temperature. Then, the protein-attached
beads were pelleted by centrifugation and washed for three times. To prepare protein samples,
beads were added to 2× loading buffer (Bio-Rad) and heated at 95 °C for 10 min. Protein samples
were further analyzed by western blot. The recombinant proteins used were listed in
Supplementary Table 3.

AlphaLISA assay
The AlphaLISA experiment was performed in the dark using white 96-well half area plate
(Greiner) according to the User’s Guide (PerkinElmer). In brief, protein cross-titration experiment
was firstly performed by testing multiple concentration combinations of two proteins in a matrix
to detect the optimal analyte association. Next, a third protein or an untagged version of one or the
other protein was introduced to the displacement assay to affect the existing association between
beads. The alpha signals indicating the binding affinity were measured in a Tecan plate reader
using the AlphaLISA detection mode. All the tagged or untagged recombinant proteins used for
this assay were summarized in Supplementary Table 3 and the corresponding Donor and
Acceptor beads were selected based on the tags of protein pairs.

RNA sequencing (RNA-seq) and whole genome sequencing (WGS) to identify editing events
Total RNA was isolated from C4-2 cells using RNeasy Plus Mini Kit (Qiagen) and subjected
to BGI for Poly(A) selection library preparation and sequencing using the DNBseq platform.
Meanwhile, the genomic DNA was extracted from C4-2 cells using Quick-DNA Kit (Zymo
Research) and subjected to BGI for human true PCR-free 30x coverage WGS.

For WGS, the raw fastq files were mapped to the human reference genome hg19 with BWA
(v0.7.17)[60]. Sorted bam file was then used as the corresponding genome reference of C4-2 for
identification of RNA editing events.

For RNA-seq, the raw reads with fastq format were aligned to the human reference genome
hg19 from iGenome (https://support.illumina.com/sequencing/sequencing_software
/igenome.html) using STAR aligner (v2.7.9a)[61]. The count number of mapped reads was
calculated by htseq-count (v0.11.2)[62]. After mapping, the software of REDItoolDnaRna.py from
REDItools (v1.3)[63] was applied to detect the RNA editing events using the parameters provided
by a published protocol[64]. The C4-2 genome information, database of SNPs (dbSNP) and several
stringent filters were utilized to minimize the detection of false RNA editing candidates and obtain
high confident editing events. These filtering processes include considering quality score >25,
mapping quality >30, base coverage >10× and removal of splicing sites and PCR duplicates. The
altered RNA editing events between siADAR1 or siEZH2 and siCTRL were determined using the
log likelihood ratio model REDIT-LLR in R package[65]. Only the A-to-I editing events were
further recruited for the differential analysis and functional annotations. An editing level cutoff of
≥5% was applied, and only sites with a P value < 0.05 were considered significant.

Similar analysis was also conducted on Stand Up To Cancer (SU2C) metastatic PCa samples
using transcriptome sequencing and matched whole exome sequencing data from the dbGAP
dataset (phs000915.v2p2[32]), as well as on PDO samples and C4-2 cells treated with EPZ-6438
and GSK-126 inhibitors. For SU2C data analysis, we divided PCa patient samples into EZH2-high
and -low groups using a previous method [66]. The EZH2-knockdown datasets of LNCaP[67],
LNCaP-abl[67], HL-60[68], MCF-7[69], and SUM159[70] cells were downloaded from GEO
database. The REDItools de novo analysis[63] was conducted to detect and quantify RNA editing
levels in these samples.

RNA editing analysis using TCGA samples
RNA-seq alignments stored as BAM files from 552 prostate adenocarcinoma (PRAD)
patients were downloaded through gdc-client v1.6.1. To associate the biospecimen data within
TCGA barcodes, the GenomicDataCommons package v1.22.1[71] was further used to map from
the universal unique identifier (UUID). For each biospecimen, variant calling was performed using
SAMtools v1.5 mpileup[72] and bcftools v 1.16 call[73]. Filtering was performed by discarding
SNVs or small indels in which the variant calling score at the QUAL field is lower than 20. The
editing ratio of chr12:69237512 was calculated using the VCF file of the TCGA primary prostate
dataset and the normal prostate dataset from RNA editing database[74].

RNA editing site-specific quantitative RT-PCR (RESSq-PCR) assay
The RESSq-PCR approach is a sensitive RNA editing fingerprint assay to detect RNA
recoding[33]. To conduct this, RNA editing site-specific primers that were compatible with SYBR
green RT-qPCR assay were firstly designed based on the principle of this strategy and were
presented in Supplementary Table 2. Then, the RT-qPCR analyses were carried out using the
cDNA samples from each group. The relative RNA editing level (Edited/WT ratio) was calculated
as 2(CtWT-CtEdited).

Sanger sequencing
The MDM2 cDNA fragment containing the interested editing site was PCR-amplified using
amifisure PCR Master Mix (GenDEPOT) and purified using QIAquick PCR Purification Kit
(Qiagen). The obtained PCR products were shipped to GENEWIZ for Sanger sequencing.
Sequencing chromatograms were visualized using SnapGene Viewer and the A-to-G editing
frequency was calculated as %G/(A+G) based on the raw sequencing data. The primer sequences
were presented in Supplementary Table 2.

MDM2 3′-UTR Reporter Assay
Both control (CmiT000001-MT06) and wildtype MDM2 3′-UTR (HmiT102206-MT06)
reporter clones were purchased from GeneCopoeia, while the edited MDM2 mutant was
constructed through A-to-G site-directed mutagenesis. HEK293T cells overexpressing control,
miR-204 or miR-211 mimics were seeded into 24-well plate and transfected with control, wildtype
or edited 3′-UTR reporter plasmids. Dual-luciferase assays were performed at 24 h post-
transfection of reporters using Luc-Pair Duo-Luciferase HS Assay Kit (GeneCopoeia) according
to the manufacturer’s instructions. A Tecan plate reader was used to measure the bioluminescence
intensity.

Label-free quantitative co-IP/MS
To quantitative comparison of proteins, duplicates of control and EZH2-deficient C4-2 cells
were subjected to nuclear isolation using NE-PER nuclear extraction kit (Thermo Fisher), followed
by co-IP assay to pull down ADAR1 along with its interacting proteins using the method as
mentioned above. IP eluate of each sample was then run into the stacking gel and submitted to
Northwestern Proteomics Core for MS analysis by following a previously established protocol[75].

Cell fractionation assays
For nuclear and cytoplasmic extraction, NE-PER nuclear extraction kit (Thermo Fisher) was
utilized to separate the nuclear and cytoplasmic fractions of C4-2 cells, followed by western blot
analysis.
For size-exclusion chromatography, nuclear extracts from control and EZH2-deficient C4-2
cells were prepared using NE-PER nuclear extraction kit (Thermo Fisher), and 5 mg of obtained
nuclear protein was concentrated into 200 μL using Ultra-0.5 Centrifugal Filter (Amicon). The
concentrated sample was then loaded into Pharmacia SMART system for size-exclusion. A total
of 48 fractions with a volume of 50 μL for each fraction were finally obtained and subjected to
western blot assay.

Enhanced Crosslinking and Immunoprecipitation-sequencing (eCLIP-seq)
The eCLIP-seq experiment was performed in duplicates following the method described
previously[76]. For each sample, a total of 50 million C4-2 cells were crosslinked, lysed and
partially digested by RNase I (Thermo Fisher). The resulting lysates were incubated with the
antibody of interested RBP and Protein A/G magnetic beads (Pierce) at 4 ºC overnight, followed
by washing, end repair and 3′ adaptor ligation. The protein-RNA complexes were then eluted from
the beads, resolved by denaturing gel electrophoresis and transferred to a nitrocellulose membrane.
Subsequently, the region of interest was cut from membrane and subjected to RNA extraction,
cDNA synthesis and library preparation. The cDNA library was further amplified using Q5 PCR
Master Mix (NEB) and sent to BGI for sequencing using the DNBseq platform.

For analysis, both STAR (v2.7.9a)[61] with default parameters and UCSC known genes were
firstly employed to map the genome hg19, followed by peak calling using CLIP-seq peak caller
Piranha with default parameters and bin size=20[77]. To define the RBP binding peaks in each
sample, a P-value cutoff of 0.01 was set up. The homer (v4.8.3) annotatePeaks.pl script[78] was
then applied to annotate the peaks from the eCLIP-seq data. A down sampling step was performed
using the samtools view function to match the sample with the smallest read count. The software
bamCoverage from deeptools (v3.5.5)[79] was used to generate a coverage track bigWig as output,
the bigwigCompare was used to generate the input subtracted bigwig signal.

Immunofluorescence
C4-2 cells were cultured on coverslips for 24 h before immunofluorescence staining. After
fixation in pre-cooled methanol for 15 min, cells were permeabilized in PBS containing 0.5%
Triton X-100 for 10 min at room temperature. Slides were rinsed thrice with PBS and blocked with
5% BSA in PBS for 30 min. Then, the slides were co-incubated with rabbit anti-EZH2 antibody
and mouse anti-GFP antibody at 4 ºC overnight. After rinsed thrice with PBS, the slides were
exposed to Alexa Fluor 488 conjugated goat anti-mouse and Alexa Fluor 555 conjugated goat anti-
rabbit antibodies (1:200, Invitrogen) for an additional hour at 37 ºC, followed by three washes with
PBS. Cell nuclei were stained in the dark with DAPI (Invitrogen) and mounted using ProLong
Diamond Antifade Mountant (Invitrogen). Immunostained cells were viewed and photographed
using Nikon A1R confocal microscope. The primary antibody information could be found in
Supplementary Table 1.

Dual-luciferase assay to detect IRES activity of TRN1
The bi-cistronic luciferase vector contain IRES element of TRN1 was constructed as
described previously[27], while the proposed TRN1 IRES sequence was determined based on a
previous report[39]. The dual-luciferase assay was performed at 24 h post-transfection using the
Dual-Glo luciferase reagent (Promega) according to the manufacturer’s instructions, followed by
signal capture through a Tecan plate reader.

Tissue microarrays (TMAs) and Immunohistochemistry (IHC) staining
Prostate tumor biopsies retrieved from Vancouver Prostate Centre tissue bank were recruited
to construct TMAs as published previously[80]. This protocol was approved by the office of
research ethics in the University of British Columbia. IHC was performed using Ventana Discovery
XT autostainer (Ventana) with the indicated antibodies by following its manual. All stained slides
were recorded by a Leica SCN400 scanner. Digital images were evaluated and scored by a
pathologist, Dr. Ladan Fazli. The histology score (H-score) of each stained protein was calculated
by the Aperio ImageScope software (Leica Biosystems) based on both intensity and percentage of
the IHC signals. All primary antibodies used for IHC were listed in Supplementary Table 1.

mRNA stability assay using actinomycin D (ActD)
To analyze the impact of EZH2 or ADAR1 suppression on mRNA decay rate, control, EZH2-
or ADAR1-deficient C4-2 cells were incubated with 5 μg/ml ActD (Sigma). Cell samples were
collected at 0, 3, 6, 12 and 24 h post-treatment, followed by total RNA isolation using RNeasy
Plus Mini Kit (Qiagen). The obtained RNA was either subject to RNA-seq analysis (n=2) in BGI
or RT-qPCR analysis (n=3) after reverse transcription.

The RNA half-lives in each sample were calculated using a previously described method[81].
In brief, to minimize the gene expression noise caused by genes that showed higher expression
levels after the ActD treatment, we scaled the FPKM values based on the expression levels of the
top 10 upregulated genes. The RNA degradation rate k was calculated using the average ratio
decay
of the RPKM value at 0 h post-treatment and the RPKM values at the other time-points. The
pseudo-RNA half-life t was then determined by calculating ln(2)/k .
1/2 decay

Isolation and Detection of Nascent RNA
The newly minted RNAs were enriched using Click-iT Nascent RNA Capture Kit (Thermo
Fisher) by following manufacturer’s manual. In general, the nascent RNAs were firstly labeled by
incubation of live cells with 5-ethynyl uridine (EU) for 6 h. After incubation, the total RNA
containing EU-labeled nascent RNA was extracted using RNeasy Plus Mini Kit (Qiagen) and used
in a copper catalyzed click reaction with an azide-modified biotin. The captured nascent RNA
transcripts on streptavidin magnetic beads were reverse transcribed using the SuperScript VILO
cDNA synthesis kit (Thermo Fisher). The obtained cDNAs were then subjected to RT-qPCR
analysis with primers summarized in Supplementary Table 2. GAPDH was employed as an
internal control.

RNA immunoprecipitation (RIP)-qPCR analysis
The RIP assay was performed in triplicates using the EZ-Magna RIP kit (Millipore) using the
procedure provided by the manufacturer. Control or treated C4-2 cells were lysed using RIP lysis
buffer and incubated with antibody-magnetic beads mixture at 4 °C overnight to enrich targeted
proteins along with the associated mRNAs. These interacting mRNAs were collected by Proteinase
K digestion, followed by purification and reverse transcription into cDNA. Then, RT-qPCR assay
was performed to measure the %Input of RBP-binding mRNAs in each group using the primers
presented in Supplementary Table 2.

Cell viability assays
To measure IC50 of EZH2 inhibitors, C4-2 cells were seeded into 96-well plates at a density
of around 1,000 cells per well. After 24 h, cells were treated with DMSO or indicated serial
dilutions of each drug for 3 days. Cell viability was assessed using CellTiter-Glo Luminescent Cell
Viability Assay Kit (Promega) and was read on a Tecan plate reader. The cell proliferation curve
was drawn and fit by the bioluminescence to drug concentration and the IC50 was calculated with
non-linear fitting.

Generation of C4-2 cells stably expressing inducible ADAR1 shRNA
The ADAR1 shRNA oligos were synthesized in IDT with the sequences shown in
Supplementary Table 2. The oligos were annealed and ligated into pLKO-Tet-On vector
(Novartis). The obtained pLKO-Tet-On shADAR1 construct and the helper plasmids were then
co-transfected into HEK293T cells to pack lentiviruses, followed by infection into C4-2 cells. The
stable Dox-inducible shADAR1-expressing C4-2 cell line, along with the negative control cell line,
were finally obtained by selection with puromycin (Sigma) and verified by western blot analysis.

Murine PCa xenograft model
Five-week-old male SCID mice were purchased from Charles River. Animal care and use
conditions were followed in accordance with institutional and National Institutes of Health
protocols and guidelines, and all studies were approved by Northwestern University Animal Care
and Use Committee. The maximal permitted tumor burden was 1,000 mm³, as defined by the
approved protocol. Tumor volumes were monitored regularly, and animals were euthanized before
reaching this limit. The maximal tumor size /burden permitted by the IACUC was not exceeded in
any experiment.

MS8815 was dissolved in 5% (vol/vol) NMP, 5% (vol/vol) Solutol HS-15 and 90% (vol/vol)
normal saline; while EPAZ6438 was dissolved in ddH2O containing 0.5% NaCMC and 0.1% Tween-
80. Both drugs were administered twice a day.

For drug optimization assay, each mouse was injected subcutaneously in the left flank with
2×106 C4-2 cells suspended in 100 μL of PBS with 50% Matrigel (BD Biosciences). After tumor
volumes reached above 50 mm3, mice were randomized to 5 groups of 4 mice each: (1) vehicle
control; (2) 25 mg/kg/day MS8815; (3) 50 mg/kg/day MS8815; (4) 100 mg/kg/day MS8815; (5)
200 mg/kg/day MS8815. Drugs were intraperitoneally (IP) injected 5×/week for 4 weeks. Tumor
size was recorded twice a week over the course of the studies using calipers and tumor volume
was calculated as follows: MIN(a)2 × MAX(b) × 0.5. Animal body weights were measured at the
beginning and the end of the treatment. After 28 days of treatment, mice were euthanized, tumors
were excised and weighed. The effect of MS8815 treatment in degradation of EZH2 was further
examined by western blot.

For combination assay, each mouse was injected subcutaneously in the left flank with 2×106
Dox-inducible shCTRL or shADAR1 C4-2 cells suspended in 100 μL of PBS with 50% Matrigel
(BD Biosciences). When tumor volumes reached 100 mm3, doxycycline (Sigma) was added to the
drinking water (final concentration, 100 μg/ml) and the two sets of mice were further assigned into
6 groups of 8 mice each: (1) vehicle-shCTRL; (2) EPZ6438-shCTRL (150 mg/kg/day); (3)
MS8815-shCTRL (50 mg/kg/day); (4) vehicle-shADAR1; (2) EPZ6438-shADAR1 (150
mg/kg/day); (3) MS8815-shADAR1 (50 mg/kg/day). Drugs were delivered either by oral gavage
(EPZ6438) or IP injection (MS8815) 5×/week for 5 weeks. Tumor size was routinely recorded as
described above. After 35 days of treatment, mice were euthanized, tumors were excised and
weighed.

For the siRNA-loaded NP animal assay, the bulk EZH2 (sequence: 5′-
GCUGACCAUUGGGACAGUATT-3′) and ADAR1 (sequence: 5′-
CAGUGUUCCUGAAACCGCUTT-3′) siRNAs were synthesized in Dharmacon along with the
cy5-Luciferase control siRNAs. The lipid-polymer hybrid siRNA NPs were formulated by a self-
assembly nanoprecipitation method with modifications[82]. Briefly, 5 mg PLGA and 0.5 mg G0-
C14 were dissolved in 1 mL of organic solvent (e.g., DMF) respectively. 50 µL of siRNA (4 nM)
aqueous solution were mixed with the G0-C14 organic solution by pipetting to form a siRNA/G0-
C14 nanocomplex. Next, the organic solution with nanocomplexes and polymers was added
dropwise to 20 mL of aqueous solution containing 2 mg of lipid-PEG (e.g., DSG-PEG). 0.2 mL of
PBS was subsequently added to stabilize the formed NPs. The suspension was then stirred at room
temperature for 30 minutes. The hybrid nanoparticles were finally washed in Amicon tubes
(MWCO 100 kDa; Millipore), the remaining organic solvent and free compounds were removed
with ice-cold water, and concentrated in 1 mL of PBS solution.

LuCaP 35CR PDX bits were implanted subcutaneously into 6–8 weeks old pre-castrated male
athymic SCID mice. Mice were monitored for up to 12 weeks post implantation for initial growth.
When the tumors were approximately 200mm3 in size, mice were randomized into treatment
groups. Tumor size was measured three times weekly using the formula mentioned above.
Intratumoral delivery into the subcutaneous flank tumors were administered twice a week for 3
weeks in 50 μL volume with 28 Gauge needle syringe. Sonication is performed to achieve multiple
purposes including deagglomeration of nanoparticles every time before treatment. Mice were
euthanized when tumor volume exceeded 1000 mm3 or at the end of a predefined experimental
endpoint.

Visualization of protein domain structures
The domain structure of protein was illustrated by Domain Graph (DOG) 2.0 software[83].

Statistics and reproducibility
Statistical analysis was performed using GraphPad Prism (version 6.0) or R and presented as
means ± SD. Unless otherwise specified, the P values were obtained using two-tailed Student’s t-
tests for comparison of two datasets or by analysis and variance (ANOVA) where appropriate.
Statistical data were considered significant if P < 0.05. The results were reproducible and
conducted with established internal controls. When feasible, experiments were repeated three or
more times and yielded similar results. We have indicated the n values used for each analysis in
the figure captions.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_4": "Overall I find this to be a difficult to follow study and mechanism and one that has not revealed itself in physiological conditions based on many mouse models and cell lines models of ADAR1 and EZH2 biology.\nSpecific comments:\n- all westerns should be quantitated and statistically assessed.\n- no analysis of the transcriptional changes that occur with knock-down of Ezh2 compared to ADAR1 - are the same pathways being regulated or is there little overlap? They are likely to be significantly different between siADAR1 and siADAR2 so teh comparison to siEzh2 would be enlightening as to how much of the transcriptional signature of siADAR1 is shared.\n- what type of RNA-seq was done - it says BGI undertook it but this doesnt explain whether it was ribosome depleted or polyA libraries. Based on the distribution of editing sites I assume it is most likely polyA as the relative proportions of repeat editing are generally much higher that shown in Fig 2c.\n- The authors must specify which ADAR1 isoform they are talking about where. This is a profound deficiency in the current text.\n- related to ADAR1 isoforms - it is not clear that any of the reagents used can specifically delineate between ADAR1p110 and ADAR1p150 - this applies to the siRNAs and the antibody used for localisation of the suggested \"cytoplasmic\" ADAR1. If the reagent cant discern between the ADAR1 isoforms then it is hard to substantiate that its p110 (as implied) that is in the cytoplasm.\n- 8-Azaadenosine is not an ADAR1 inhibitor - https://pubmed.ncbi.nlm.nih.gov/35586115/\n- Data don’t support the conclusion in lines 559-561 - this work presents very limited and a potentially subtle effect of Ezh2 on ADAR1 activity.\n- it is incorrect to describe the editing of a specific single adenosine as characterised for the editing chr12: 69237519 as hyperediting (Hyperediting is most commonly used in the field to refer clusters of editing in the same transcript not increases in editing at a specific site - see https://www.nature.com/articles/ncomms5726).\nComment on the authors’ response to the previous Reviewers\n-the adjusted Venn diagrams in the response to reviewers comments should be presented in the main figure.\n-I find the text presented i response to reviewer 1 point 3 unconvincing and the eCLIP data as presented show broad enough binding patterns that overlaps may be circumstantial;\n- the response to reviewer 1 point 4 should be conducted with the same window and preferably a smaller one as eCLIP should give residue level information about the RNA binding footprint of each when done correctly (200nt for ADAR1 and EZH2 or 1000nt for both - it is not clear why a different window of binding would be appropriate or justified).\n-The model proposed involving all three dsRBD of ADAR1 should be supported with Alphafold predictions if possible - I find it hard to see how ADAR1 could commit all three dsRBD to interacting with EZH2 and there be both increased and decreased editing. The alternative model could be that the interaction occurs separately from editing by ADARs.\n-There are multiple examples of unexpected grammar which also impair readability (e.g. line 529 - \" Accordingly, it is esteemed that the landscape of ...\" - esteemed is a strange choice)"}, "author_response": "Point-by-point response to Reviewer’s comments\nDear Reviewer,\nWe would like to thank you for taking the time to provide critical and insightful\ncomments on our manuscript. Your suggestions are very helpful for us to improve our\nresearch. We have carefully considered the comments and done our best to address\nevery one of them. We hope you will appreciate our strong new supporting results and\nefforts in this revised version of manuscript.\nAt the following, all points mentioned by the reviewer will be discussed.\nReviewer #4:\nOverall, I find this to be a difficult to follow study and mechanism and one that has\nnot\nrevealed itself in physiological conditions based on many mouse models and cell lines\nmodels of ADAR1 and EZH2 biology.\nResponse: We appreciate the reviewer’s thoughtful comment and the opportunity to\nclarify our study. In this work, we have evaluated the proposed mechanism across\nmultiple in vitro and in vivo systems, including cancer cell lines, primary normal cells,\npatient-derived xenografts (PDXs), and patient-derived organoids (PDOs). We\nrecognize that the mechanistic interaction between ADAR1 and EZH2 is complex and\ncannot be fully delineated within a single study. Accordingly, we have carefully revised\nthe manuscript to temper our claims and to focus on the findings that are directly\nsupported by our data.\nSpecific comments:\n- all westerns should be quantitated and statistically assessed.\nResponse: Thanks for pointing out this issue. In the revised manuscript, we have\nquantitated and statistically assessed the key western blot data in Fig. 3i, Fig. 4b, Fig.\n5a, Fig. 5j, Fig. S4f and Fig. S5a. We have updated all related figures to show these\nquantification results.\n- no analysis of the transcriptional changes that occur with knock-down of Ezh2\ncompared to ADAR1 - are the same pathways being regulated or is there little overlap?\nThey are likely to be significantly different between siADAR1 and siADAR2 so teh\ncomparison to siEzh2 would be enlightening as to how much of the transcriptional\nsignature of siADAR1 is shared.\nResponse: Following the reviewer’s instructive suggestion, we performed a\ncomparative analysis of differentially expressed genes (DEGs) upon EZH2, ADAR1,\nor ADAR2 knockdown in C4-2 cells. As shown in Response Letter Figure 1, both\nupregulated and downregulated genes in these groups exhibit significant overlap and\nare enriched in multiple biological pathways that are either shared or unique. This\nobservation is expected, as RNA editing can modulate mRNA targets through diverse\nmechanisms that do not always correspond directly to transcriptional changes.\nMeanwhile, EZH2 is a well-established master transcriptional regulator in cancer cells,\nand its depletion can cause extensive transcriptomic reprogramming. Therefore, it is\nchallenging to meaningfully assess the shared transcriptional signatures between\nsiADAR1/2 and siEZH2 conditions.\nResponse Letter Figure 1\n(A, B) Venn diagram to show the overlap of upregulated (A) and downregulated (B)\ngenes upon EZH2, ADAR1, or ADAR2 knockdown in C4-2 cells. P values were\ncalculated by one-tailed Fisher’s exact test.\n(C, D) Gene Ontology enrichment analysis of the upregulated (C) and downregulated\n(D) genes upon EZH2, ADAR1, or ADAR2 knockdown in C4-2 cells.\n- what type of RNA-seq was done - it says BGI undertook it but this doesn’t explain\nwhether it was ribosome depleted or polyA libraries. Based on the distribution of editing\nsites I assume it is most likely polyA as the relative proportions of repeat editing are\ngenerally much higher that shown in Fig 2c.\nResponse: We appreciate the reviewer’s question. Yes, we performed Poly(A)\nselection-based RNA-seq. We have revised the Methods section to clarify this point.\n- The authors must specify which ADAR1 isoform they are talking about where. This\nis a profound deficiency in the current text.\nResponse: Thanks for pointing out this potential weakness. In the revised manuscript,\nwe have carefully specified the ADAR1 isoform being referred to throughout the text\nand figures. In this study, we mainly studied the ADARp110 isoform since this is the\nmajor isoform expressed in prostate cancer cell lines and tissues (please see the details\nbelow).\n- related to ADAR1 isoforms - it is not clear that any of the reagents used can\nspecifically delineate between ADAR1p110 and ADAR1p150 - this applies to the\nsiRNAs and the antibody used for localization of the suggested \"cytoplasmic\" ADAR1.\nIf the reagent can’t discern between the ADAR1 isoforms then it is hard to substantiate\nthat its p110 (as implied) that is in the cytoplasm.\nResponse: We thank the reviewer for this thoughtful comment. We agree that most\ncommercially available siRNAs and antibodies, including those used in our study,\nrecognize both ADAR1 isoforms. However, several lines of evidence from our study\nindicate that the cytoplasmic ADAR1 species we focused on corresponds to the p110\nisoform:\n1- In prostate cancer (PCa) cells, we consistently observed a single ADAR1 band\ncorresponding to the p110 isoform (as shown in all related uncropped western blot\nimages). The p150 isoform was only detectable in normal PrEC cells (Fig. S2j).\n2- Our western blot analyses (Fig. 5a and Fig. 5j) demonstrated that upon EZH2\ndepletion, only the ADAR1p110 isoform, not p150, was redistributed from the nucleus\nto the cytoplasm.\n3- Overexpression of GFP-tagged ADAR1p110 in PCa cells further confirmed that this\nisoform was translocated to the cytoplasm following EZH2 knockdown (Fig. 5b), and\ncan be rescued by re-expression of TRN1 (Fig. 5k).\nTogether, these results support our conclusion that the cytoplasmic ADAR1 observed\nin this study corresponds to the p110 isoform.\n-8-Azaadenosine is not an ADAR1 inhibitor-\nhttps://pubmed.ncbi.nlm.nih.gov/35586115/\nResponse: We thank the reviewer for raising this important point. We agree with the\nconclusion of the cited study that 8-aza is not a specific ADAR1 inhibitor. Therefore,\nwe only used this compound in certain cell line-based assays and relied on ADAR1-\nspecific shRNA/siRNA approaches to validate our findings in vivo. Indeed, to date, no\nselective ADAR1 inhibitor is available. Recently, one group reported the development\nof a small-molecule ADAR1 inhibitor, ZYS-1[1]. However, multiple follow-up studies\nhave shown that ZYS-1 treatment neither reduces cellular A-to-I editing nor\ndownregulates ADAR1 expression[2, 3]. Consistent with these reports, when we tested\nZYS-1 in our A-to-I editing reporter C4-2 cell line, we found that 8-aza, but not ZYS-\n1, significantly decreased the editing level (Response Letter Figure 2A and 2B).\nBased on these results, we believe that 8-aza remains the most suitable compound for\nin vitro studies at this stage. To avoid misunderstanding, we have revised our\ndescription of 8-aza from “ADAR1 inhibitor” to “RNA editing repressor.”\nResponse Letter Figure 2 8-aza is better than ZYS-1 in RNA editing suppression.\n(A) Dual luciferase reporter monitoring A-to-I editing. The reporter utilizes the natural\nGluA2 editing substrate in which the R/G editing site has been modified into a stop\ncodon (UAG) that upon editing is recoded into a tryptophan codon (UGG). A firefly\nluciferase reporter gene upstream of the edited site monitors translation and a\nnanoluciferase reporter gene downstream is used to measure read-through after editing.\nEditing is measured as the ratio between luminescence from nanoluciferase and firefly\nluciferase. The principle of the design was inspired by a prior study[4].\n(B) Dual-Luc assay to measure the change of RNA editing level upon ZYS-1 (20 μM)\nor 8-aza (1 μM) treatment in A-to-I editing reporter C4-2 cell line.\n- Data don’t support the conclusion in lines 559-561 - this work presents very limited\nand a potentially subtle effect of Ezh2 on ADAR1 activity.\nResponse: We have tempered the related claims in the manuscript into “In summary,\nour findings indicate a possible mechanism through which EZH2 modulates both the\nRNA editing-dependent and -independent activities of ADAR1 (Supplementary Fig.\n8), although additional studies will be required to fully establish this relationship and\nits therapeutic implications” to address this concern.\n- it is incorrect to describe the editing of a specific single adenosine as characterised for\nthe editing chr12: 69237519 as hyperediting (Hyperediting is most commonly used in\nthe field to refer clusters of editing in the same transcript not increases in editing at a\nspecific site - see https://www.nature.com/articles/ncomms5726).\nResponse: Thanks for pointing out this error. In the revised manuscript, we have\nchanged “hyperediting” into “increased editing level” or “over-editing” in all related\ntext.\nComment on the authors’ response to the previous Reviewers\n-the adjusted Venn diagrams in the response to reviewer’s comments should be\npresented in the main figure.\nResponse: Thanks for the suggestion. Due to the size limit of Fig. 2, we have moved\nthese adjusted Venn diagrams to Supplementary Fig. 2g-2i and revised the text\naccordingly.\n-I find the text presented in response to reviewer 1 point 3 unconvincing and the eCLIP\ndata as presented show broad enough binding patterns that overlaps may be\ncircumstantial;\nResponse: Thank you for this important question. Inspired by your subsequent\ncomment regarding the overlap between eCLIP peaks and RNA editing sites, we\nanalyzed the relationship between ADAR1/EZH2/ILF2 eCLIP peaks and EZH2-\nmediated editing sites. We found that, as compared with ILF2, EZH2-binding regions\npreferentially overlap with the under-edited sites. In contrast, the ILF2-binding regions\nshow greater overlap with over-edited sites following EZH2 depletion (Response\nLetter Figure 3, also included in the revised manuscript as Supplementary Fig. 4h).\nThese findings suggest that EZH2 binding tends to coincide with editing sites\ndownregulated upon EZH2 knockdown, while ILF2 binding tends to coincide with\nediting sites upregulated upon EZH2 knockdown.\nResponse Letter Figure 3 Overlap of the EZH2-mediated editing sites with ADAR1-,\nEZH2-, and ILF2-binding mRNA regions.\nBar plots showing the observed overlap counts between the under-edited sites (left) /\nover-edited (right) sites upon EZH2 knockdown and the binding regions of ADAR1,\nEZH2, and ILF2 (±200 nt) determined by eCLIP-seq. For each panel, the “Random”\nbar represents the null distribution generated from random genomic regions matched in\nnumber and size (permutation-based sampling), shown as mean ± SD. The colored bars\nrepresent the observed overlap counts for each RBP. P-values were calculated by\nPermutation test.\n- the response to reviewer 1 point 4 should be conducted with the same window and\npreferably a smaller one as eCLIP should give residue level information about the RNA\nbinding footprint of each when done correctly (200nt for ADAR1 and EZH2 or 1000nt\nfor both - it is not clear why a different window of binding would be appropriate or\njustified).\nResponse: We appreciate the reviewer’s insightful comment regarding the choice of\nwindow size around eCLIP peaks. In the revised analysis, we have applied a uniform\n±200 nt window for both ADAR1 and EZH2 datasets to ensure methodological\nconsistency. As depicted in Response Letter Figure 4, a significant overlap could still\nbe observed between ADAR1-bound regions and EZH2-regulated editing sites using\nthe same window.\nResponse Letter Figure 4 Graph showing the overlap count between ADAR1-bound\nregions (±200 nt) and EZH2-regulated editing sites (±200 nt). Permutation test P = 0.\n-The model proposed involving all three dsRBD of ADAR1 should be supported with\nAlphafold predictions if possible - I find it hard to see how ADAR1 could commit all\nthree dsRBD to interacting with EZH2 and there be both increased and decreased\nediting. The alternative model could be that the interaction occurs separately from\nediting by ADARs.\nResponse: We thank the reviewer for this great advice and performed Alphafold 3\npredictions as suggested. As depicted in Response Letter Figure 5, dsRBD1 and\ndsRBD3 domains from ADAR1 are scored moderately well to be plausible candidates\nto interact physically with the SANT2 domain of EZH2, whereas dsRBD2 was scored\nlow. Worth to note that, Alphafold 3 recognizes ipTM ≥ 0.75 for a strong interface\nplausibility, 0.5-0.75 for possible interface and < 0.5 for weak interface. We had cellular\ndata suggesting that all three dsRBD domains from ADAR1 are critical for the\ninteraction with EZH2 and it is possible that dsRBD1 and dsRBD3 provides direct\nphysical interface and dsRBD2 serves as a connecting region to coordinate this\ninteraction, so that all of them are indispensable.\nResponse Letter Figure 5 AlphaFold3-predicted structural models of EZH2-SANT2\ndomain in complex with individual ADAR1 dsRBD domains.\nProtein-protein interaction predictions between EZH2-SANT2 (cyan) and ADAR1\ndsRBD1 (green), dsRBD2 (yellow), or dsRBD3 (red) were generated using AlphaFold3\nand visualized in PyMOL. The three panels show the top-ranked models for each\ndomain pair. Interface predicted TM-scores (ipTM) are shown below each panel to\nindicate the confidence of the predicted interaction. Higher ipTM values are consistent\nwith more confident interface predictions.\n-There are multiple examples of unexpected grammar which also impair readability (e.g.\nline 529 - \" Accordingly, it is esteemed that the landscape of ...\" - esteemed is a strange\nchoice)\nResponse: We thank the reviewer for pointing out this issue. We have carefully revised\nthe manuscript to correct grammatical errors and improve readability throughout.\nSpecifically, we have replaced “esteemed” in line 529 with “suggested”."}, {"round": 2, "reviewer_comments": {"reviewer_4": "The authors have made efforts to amend the manuscript in response to the comments raised during review. It remains a very dense and complex manuscript with the chosen figure presentation not helping the reader access the important findings given how dense each figure panel is.\nBased my initial reviews and considering both the response to reviewers from the authors and the changes to the text itself I would recommend removal of the 8-Aza data. It has been demonstrated not to be an ADAR inhibitor so why the authors have decided to persist as if it is is not clear to me at all. The moderated text is the the manuscript wants the reader to conclude that 8-Aza is an in cell ADAR editing regulator when this is clearly not established or supported by the data available from cell line treatments with this agent (which notably are not cited by the present authors). The current text is disingenuous when the authors acknowledge in the rebuttal that 8-Aza is not an inhibitor and then make a superficial change to the text.\nThe authors should clearly state the level of editing change considered significant for the analysis (ie what cut off was used to call a site edited and then what change (5%, 10% etc) was considered significant and how many of the replicates were required to support the changes).\nWhy has Figure 4a have statistical analysis when it says n=2?\nThe meaning of “edited/Wt ratio” for the analysis of the editing in Figure 2g-2h needs to be more clearly defined. A change from 1% editing to 2% editing is a 2 fold difference but unlikely to be meaningful – the way the data has been chosen to be displayed removes the important information about the absolute level of editing (as shown in Fig 3a) of a given site from the analysis.\nLine 361 – no primary reference to support editing-independent p110 functions in the cytoplasm? Cited reference is a general review\nLine 206-208 – no evidence genetically for compensation in vivo – this claim is not required."}, "author_response": "Point-by-point response to Reviewer’s comments\nDear Reviewer,\nWe would like to thank you again for taking time to provide additional comments\non our manuscript. We have carefully considered your comments and do our best to\naddress every one of them as follows.\nReviewer #4:\nThe authors have made efforts to amend the manuscript in response to the comments\nraised during review. It remains a very dense and complex manuscript with the chosen\nfigure presentation not helping the reader access the important findings given how\ndense each figure panel is.\nResponse: We thank the reviewer for acknowledging our efforts in the first-round\nrevision. In the current revision, we have made additional changes to further improve\nclarity and readability, including revising the presentation of figures and associated text\nto help readers more easily access the key findings.\nBased my initial reviews and considering both the response to reviewers from the\nauthors and the changes to the text itself I would recommend removal of the 8-Aza data.\nIt has been demonstrated not to be an ADAR inhibitor so why the authors have decided\nto persist as if it is is not clear to me at all. The moderated text is the manuscript wants\nthe reader to conclude that 8-Aza is an in cell ADAR editing regulator when this is\nclearly not established or supported by the data available from cell line treatments with\nthis agent (which notably are not cited by the present authors). The current text is\ndisingenuous when the authors acknowledge in the rebuttal that 8-Aza is not an\ninhibitor and then make a superficial change to the text.\nResponse: We agree with the reviewer on this point and have removed all 8-Aza-related\ndata (the original Fig. 7d and 7e) from our manuscript.\nThe authors should clearly state the level of editing change considered significant for\nthe analysis (ie what cut off was used to call a site edited and then what change (5%,\n10% etc) was considered significant and how many of the replicates were required to\nsupport the changes).\nResponse: We thank the reviewer for requesting clarification of the criteria used to\ndefine RNA editing sites, the thresholds for significant changes, and the number of\nreplicates analyzed. We have now explicitly stated these details in the figure legends\nand/or Methods section and summarized below:\nFor Fig. 2b and Fig. 2d, RNA editing sites were identified using REDItoolDnaRna.py\nfrom REDItools (v1.3) with default parameters[1]. SNPs and splicing sites were filtered\nout, and sites with a read coverage greater than 10 were considered as candidate editing\nsites. An editing level cutoff of ≥5% was applied, and only sites with a P value < 0.05\nwere considered significant.\nIn Fig. 2b, the global A-to-I editing ratios were compared across groups using a two-\ntailed unpaired Wilcoxon test, with n=4 biological replicates for siCTRL, n=4 for\nsiEZH2, n=6 for siADAR1, and n=5 for siADAR2. In Fig. 2d, scatter plots show\ncomparisons of individual editing sites between control and knockdown conditions in\nC4-2 cells using the same replicate numbers and statistical criteria.\nFor Fig. 3c, editing levels were retrieved directly from VCF files generated from the\nTCGA dataset, including 152 normal and 496 cancer samples.\nFor Fig. 3d, samples were stratified into EZH2-Low, EZH2-High, ADAR1-Low, and\nADAR1-High (n=50 for each) groups based on expression levels.\nFor Fig. 4i, a total of n=131 genes were analyzed.\nFor Fig. 6b, analyses were performed using two independent RNA-seq biological\nreplicates.\nFor Supplementary Fig. 2c, RNA editing sites were identified using\nREDItoolDnaRna.py (REDItools v1.3) with default cutoffs after filtering SNPs and\nsplicing sites, retaining sites with coverage greater than 10. Each organoid sample\nincludes two biological replicates.\nFor Supplementary Fig. 2d, due to the relatively low sequencing depth of normal\npoly(A) RNA-seq data (15-20 Mb per sample), two or three replicates from public\ndatasets were merged for analysis. In this case, REDItools de novo analysis was applied,\nand only editing sites with editing levels between 5% and 40% were retained, as the de\nnovo model is more prone to false-positive calls compared with REDItoolDnaRna.py.\nFor Supplementary Fig. 2e, n=10 samples were analyzed per group.\nFor Supplementary Fig. 2f, analyses were conducted using 4 RNA-seq biological\nreplicates.\nWhy has Figure 4a have statistical analysis when it says n=2?\nResponse: We are sorry for this mistake and have removed the statistics in Fig. 4a.\nThe meaning of “edited/WT ratio” for the analysis of the editing in Figure 2g-2h needs\nto be more clearly defined. A change from 1% editing to 2% editing is a 2 fold difference\nbut unlikely to be meaningful – the way the data has been chosen to be displayed\nremoves the important information about the absolute level of editing (as shown in Fig\n3a) of a given site from the analysis.\nResponse: We thank the reviewer for this important question. We agree that fold\nchanges in the edited/WT ratio alone do not convey the absolute level of RNA editing,\nand that a 2-fold increase from a very low baseline may not be biologically meaningful.\nIn principle, RESS-qPCR is designed to measure relative changes in the abundance of\nedited versus wild-type transcripts between experimental conditions, rather than\nabsolute editing frequency[2]. In this study, RESS-qPCR was primarily used to\nexperimentally validate RNA-seq-identified editing changes. Accordingly, the\nedited/WT ratios shown in Fig. 2g and 2h are intended to reflect directional and\ncomparative changes in editing efficiency, not absolute editing percentages.\nFor downstream functional characterization of a specific EZH2-mediated editing site,\nwe employed sequencing-based approaches, including Sanger sequencing, to directly\nquantify absolute RNA editing levels (Fig. 3a, 3b, and 3h).\nTo address the reviewer’s concern and improve clarity, we have now explicitly defined\nthe meaning of the “edited/WT ratio” in the Fig. 2g and 2h figure legend as: The\nedited/WT ratio reflects relative differences in editing efficiency between conditions,\nnot absolute editing percentages.\nLine 361 – no primary reference to support editing-independent p110 functions in the\ncytoplasm? Cited reference is a general review.\nResponse: In the current manuscript, the original article which reported the editing-\nindependent p110 functions in the cytoplasm has also been cited in Line 361[3].\nLine 206-208 – no evidence genetically for compensation in vivo – this claim is not\nrequired.\nResponse: Following the reviewer’s suggestion, we have deleted this sentence in the\nupdated manuscript."}, {"round": 3, "reviewer_comments": {"reviewer_4": "I have no additional comments on the manuscript."}, "author_response": ""}]

Decision letter