Slo2 channels are large-conductance potassium channels existing in mammals as well as invertebrates (Kaczmarek, 2013; Yuan et al., 2000). They are the primary conductor of delayed outward currents in many neurons examined (Budelli et al., 2009; Liu et al., 2014). Human and mouse each has two Slo2 channels (Slo2.1/Slick and Slo2.2/Slack) (Kaczmarek, 2013), whereas the nematode C. elegans has only one (SLO-2). These channels are abundantly expressed in the nervous system (Bhattacharjee et al., 2002; Bhattacharjee et al., 2005; Joiner et al., 1998; Liu et al., 2018; Rizzi et al., 2016), and play major roles in shaping neuronal electrical properties and regulating neurotransmitter release (Kaczmarek, 2013; Liu et al., 2014). Mutations of Slo2 channels cause epilepsies and severe intellectual disabilities in humans (Ambrosino et al., 2018; Cataldi et al., 2019; Evely et al., 2017; Gururaj et al., 2017; Hansen et al., 2017; Kawasaki et al., 2017; Lim et al., 2016; McTague et al., 2018; Rizzo et al., 2016), and reduced tolerance to hypoxic environment in worms (Yuan et al., 2003). Emerging evidence suggests that physiological functions of these channels depend on other proteins. For example, in mice, the fragile mental retardation protein (FMRP), a RNA binding protein, enhances Slack activity by binding to its carboxyl terminus (Brown et al., 2010). In worms, HRPU-2, a RNA/DNA binding protein, controls the expression level of SLO-2 through a posttranscriptional effect (Liu et al., 2018).

RNA editing is an evolutionally conserved post-transcriptional process catalyzed by ADARs (adenosine deaminases acting on RNA) (Gott and Emeson, 2000; Jin et al., 2009). ADARs convert adenosine (A) to inosine (I) in double-stranded RNA. Since inosine is interpreted as guanosine (G) by cellular machineries (Basilio et al., 1962), A-to-I RNA editing may alter the function of a protein by changing its coding potential, or regulate gene expression through altering alternative splicing, microRNA processing, or RNA interference (Deffit and Hundley, 2016; Nishikura, 2016). Human and mouse each has three ADARs: ADAR1, ADAR2 and ADAR3 (Chen et al., 2000; Kim et al., 1994; Melcher et al., 1996). ADAR1 and ADAR2 possess deaminase activity and catalyze the A-to-I conversion (Tan et al., 2017), whereas ADAR3 is catalytically inactive with regulatory roles in RNA editing (Nishikura, 2016). Millions of A-to-I editing sites have been detected in the human transcriptome through RNA-seq, with the vast majority of them found in non-coding regions (Nishikura, 2016). Biological effects of RNA editing at coding regions have been revealed for a variety of genes, including those encoding ligand- and voltage-gated ion channels and G protein-coupled receptors (Bhalla et al., 2004; Brusa et al., 1995; Burns et al., 1997; Gonzalez et al., 2011; Huang et al., 2012; Lomeli et al., 1994; Palladino et al., 2000; Rula et al., 2008; Sommer et al., 1991; Streit et al., 2011). However, little is known about the roles of RNA editing in non-coding regions (Nishikura, 2016).

In a genetic screen for suppressors of a sluggish phenotype caused by expressing a hyperactive SLO-2 in worms, we isolated mutants of several genes, including adr-1, which encodes one of two ADARs in C. elegans (ADR-1 and ADR-2). While ADR-2 has deaminase activity and plays an indispensable role in the A-to-I conversion, ADR-1 is catalytically inactive but can promote RNA editing by binding to selected target mRNA and tethering ADR-2 to RNA substrates (Ganem et al., 2019; Rajendren et al., 2018; Washburn et al., 2014). We found that loss-of-function (lf) mutations of adr-1 inhibit SLO-2 function through impairing RNA editing of scyl-1, which encodes an orthologue of human and mouse SCYL1. In adr-1(lf) mutants, a lack of A-to-I conversion at a specific site in scyl-1 3’-UTR causes reduced scyl-1 expression. Knockout of scyl-1 severely reduces SLO-2 current in worms whereas coexpression of SCYL1 with human Slack in Xenopus oocytes greatly augments channel activity. These results suggest that SCYL-1/SCYL1 proteins likely play an evolutionarily conserved role in physiological functions of Slo2 channels. Mutations or knockout mammalian SCYL1 may cause neural degeneration, intellectual disabilities, and liver failure, but the underlying mechanisms are unclear (Lenz et al., 2018; Li et al., 2019; Shohet et al., 2019; Spagnoli et al., 2019). The revelation of SCYL-1/SCYL1 as a protein important to Slo2 channels suggests a potential link between diseases caused by SCLY1 mutations and Slo2 channel functions.

This study shows that both ADR-1 and SCYL-1 are critical to SLO-2 physiological function in neurons. While ADR-1 enhances SLO-2 function indirectly through promoting SCYL-1 expression, SCYL-1 regulates SLO-2 through physical interactions. These conclusions are supported by multiple lines of evidence, including the isolation of adr-1(lf) mutants as suppressors of SLO-2(gf), the inhibition of SLO-2 activities by either adr-1(lf) or scyl-1(lf), the reduction of scyl-1 transcript expression in adr-1(lf), the correlation between scyl-1 mRNA level and RNA editing at its 3’-UTR, the evidence of physical interactions between SCYL-1 and SLO-2, and the reduction of SLO-2 P_o due to changes in the dwell times of open and closed events in scyl-1(lf). Importantly, we found that the human Slo2.2/Slack is also regulated by SCYL1.

The biological significance of RNA editing at non-coding regions is only beginning to be appreciated. A recent study with C. elegans identified many neuron-specific A-to-I editing sites in the 3’-UTR of clec-41, and showed that elimination of these editing events by adr-2 knockout compromises clec-41 expression and impairs a chemotaxis behavior (Deffit et al., 2017). However, it remains to be determined how clec-41 expression is controlled by these editing events, and whether the chemotaxis defect of adr-2(lf) mutant is caused by decreased clec-41 expression. In the present study, we demonstrate that an A-to-I RNA editing event at the 3’UTR of scyl-1 controls its expression, and that SCYL-1 contributes to neuronal whole-cell currents through a direct effect on SLO-2. The results of these two studies have provided a glimpse of the biological roles of 3’-UTR RNA editing in gene expression and neuronal function.

Our results demonstrate that RNA editing at a single site in the 3’-UTR could have a profound effect on gene expression. The A-to-I conversion at the ADR-1-regulated editing site increases base pairing in the putative double-stranded structure of scyl-1 3’-UTR (Figure 10B). Increased base paring in a double-stranded RNA generally facilitates RNA degradation. It is therefore intriguing how such an editing event may cause increased gene expression. One possibility is that editing at this site helps maintain mRNA stability through recruiting other regulatory proteins to the 3’-UTR. The isolation of mutants exhibiting diminished GFP expression from a reporter construct containing the edited scyl-1 3’-UTR but not the unc-10 3’-UTR (Figure 10—figure supplement 1) indicates potential existence of such regulatory proteins. In combination with the observation that GFP signal was undetectable in transgenic worms expressing a reporter construct containing the non-edited scyl-1 3’-UTR, this result suggests that RNA editing at the ADR-1-dependent site in scyl-1 3’-UTR is likely important to recruiting the putative regulatory proteins to increase mRNA stability.

scyl-1 has four different transcripts with identical 5’-UTR and coding sequence but different 3’-UTRs of variable lengths (ranging from 167 to 1862 nucleotides) and sequences (www.wormbase.org). The ADR-1-dependent editing site exists only in the transcript with the longest 3’-UTR. The presence of an ADR-1-regulated editing site in this but not the other transcripts suggests that ADR-1 may regulate scyl-1 expression in a cell-specific manner depending on where the specific transcript is expressed. Interestingly, the 3’-UTR of human SCYL1 transcripts (NM_020680.4) also has a high probability of forming hair-pin structures based on software prediction (https://rna.urmc.rochester.edu/RNAstructureWeb/Servers). It remains to be determined whether human SCYL1 transcripts are also edited in the 3’-UTR, and if so, whether the editing regulates their expression.

SCYL1 proteins are evolutionarily conserved proteins that share an N-terminal pseudokinase domain (Manning et al., 2002; Pelletier, 2016). Results of previous studies with cultured cells suggest that SCYL1 may regulate intracellular trafficking between the Golgi apparatus and the ER (Burman et al., 2008; Burman et al., 2010), and facilitate nuclear tRNA export by acting at the nuclear pore complex (Chafe and Mangroo, 2010). Mutations of SCYL1 in humans are associated with a variety of disorders, including neurodegeneration, intellectual disabilities, and liver failure (Lenz et al., 2018; Li et al., 2019; Schmidt et al., 2015; Shohet et al., 2019; Spagnoli et al., 2019). Mice with SCYL1 deficiency develop an early onset and progressive neurodegenerative disorder (Pelletier et al., 2012). However, it is unclear whether the documented mutant phenotypes of SCYL1 are related to its roles in intracellular trafficking and nuclear tRNA export (Pelletier, 2016). This study brings into view a new potential mechanism for SCYL1 mutation-associated disorders: impairing Slo2 channel function. What might be the molecular mechanism through which SCYL-1 enhances SLO-2 activity? Since SCYL-1 physically associates with SLO-2, and enhances SLO-2 single-channel P_o by altering the open and closed states, it likely regulates channel function either directly or through a closely associated protein. Although the exact mechanism remains to be determined, the similar effects of SCYL-1 and SCYL1 on Slo2 channels suggest that they likely play an important role in Slo2 physiological functions across animal species.

The expression patterns of scyl-1 and slo-2 largely do not overlap. Although they are coexpressed in ventral cord motor neurons, most other neurons expressing slo-2 do not express scyl-1, suggesting that the regulatory effect of SCYL-1 on SLO-2 is cell- and tissue-specific. The expression of scyl-1 in cells not expressing slo-2 suggests that SCYL-1 physiological functions are not limited to regulating SLO-2. In mouse, SCYL1 and Slo2.2 are both expressed in the hippocampus and cerebellum but their expression patterns do not completely overlap (Joiner et al., 1998; Schmidt et al., 2007). Conceivably, the regulation of hSlo2.2 by SCYL1 might also occur in a cell- and tissue-specific manner, and SCYL1 likely performs other physiological functions. The latter possibility is supported by the pleiotropic phenotypes observed in patients and mice with SCYL1 mutations (Lenz et al., 2018; Li et al., 2019; Pelletier et al., 2012; Schmidt et al., 2015; Shohet et al., 2019; Spagnoli et al., 2019).

In summary, this study demonstrates that ADAR-mediated RNA editing controls the expression of SCYL-1, which interacts with SLO-2 to allow SLO-2 perform its physiological functions. Moreover, this study shows that this regulatory mechanism is conserved with mammalian SCYL1 and Slo2. Our findings reveal a new molecular mechanism of Slo2 channel regulation, and provide the bases for investigating how physiological functions of human Slo2 are regulated by SCYL1, and whether the neurodegeneration and intellectual disability phenotypes of SCYL1 mutations are related to Slo2 channel dysfunction.

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 1, 2, 4, 5, 7, 8, 10,and 11. Sequencing data have been deposited in GEO under accession code GSE141316.

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Thank you for submitting your article "Slo2 potassium channel function depends on a SCYL1 protein" for consideration by eLife. Your article has been reviewed by Richard Aldrich as the Senior Editor, a Reviewing Editor (Oliver Hobert), and two reviewers. The following individuals involved in review of your submission have agreed to reveal their identity: Leonard Kaczmarek (Reviewer #1); Thomas Boulin (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. As you will see in the reviews below, there is general agreement about the general interest and importance of the study. However, a set of important clarifications and experiments need to be conducted to make this paper become acceptable for publication in eLife.

In brief, the requested experiments are:

1) Independent validation of interactions (reviewer #1, comment 2)

2) Engineering the A>G mutation into the genome (reviewer #2)

3) Improved expression pattern analysis (reviewer #2)(the in vivo recombineering technique has by now been shown to be inadequate).

There are also a number of very important clarifications that are required (e.g. reviewer #1, comment 1)

Reviewer #1:

This manuscript presents some very interesting work demonstrating that Slo2 channels in C-elegans interact with SYCL^-1 to alter open probability of these channels. The effects of SYCL^-1 on the channels is, in turn, influenced by ADR-2, which is required for A-to-I RNA editing of a site in the 3' UTR of the scyl-1 gene. The work is provocative and points the way to future work to unravel how this potential interaction affects the biology of neurons in the nematode and in mammalian systems.

1) My major comment is on the work that extrapolates the nematode findings to the human Slack channel. The latter has previously been shows to differ from the C-elegans channel in several ways. This interaction with the human channel is an important aspect of the presented work. Examination of the data in Figure 11, however, suggest that the gating of the human channel is quite different from the Slo2 channel records shown in Figure 8. Specifically (if the time bars shown in Figure 11A are correct, the overall open probability of the human channel is dominated by the presence of very long closed states (lasting many seconds). Examination of the X-axis scale bars in Figure 11B and C suggest these may have been excluded from the analyses. The conclusions on the effects of SCYL1 on open probability stated in the last paragraph of the Results section may not be completely valid until these are taken into account.

2) The bimolecular fluorescence complementation assay presented does make a case that there may be a physical interaction of SCLY-1 with SLO-2 but is not completely definitive. It would be good to have some other indicator of physical interaction to support this claim. A more conventional coimmunoprecipitation experiments would be a good addition, one that could perhaps be carried out using the co-expression Xenopus oocyte heterologous expression system.

Reviewer #2:

In this study Niu et al., describe a striking and entirely unsuspected regulatory cascade that controls SLO-2/Slo2 potassium channel activity. They report the role of SYCL^-1, a novel physical interactor of the SLO-2 potassium channel, and describe how syCl^-1 expression is controled by ADAR-dependent RNA editing in the non-coding sequence this gene. After dissecting this mechanism in worms, they proceed to directly demonstrate the conservation of this novel regulatory interaction with the human SLO-2 ortholog. Given the importance of this class of potassium channel in health and disease, identifying this modulatory mechanism is a very important finding in my opinion.

Essentiaol revisions:

- Please clarify the functional relationship between adr-1 and adr-2. Indeed, in the Introduction the authors seem to indicate that ADAR proteins could compete ("altering the accessibility") for certain binding sites. In this model, wouldn't one expect that adr-1(lf) would "free" access to the scyl-1 site, which would be inconsistent with the similarity of adr-1 and adr-2 mutant phenotypes?

Do the authors think that ADR-1 promotes the recruitement of ADR-2 to the scyl-1 3'UTR?

- I was very intrigued by the results described in Figure 10, and specifically Figure 10D/E. The result is not what I had intuitively expected. Since it was performed in an adr-1/2 wild-type background as far as I could determine from the Materials and methods section, I would have thought that ADAR activity would have edited the wild-type sequence and I would have expected to see GFP in both cases.

I would have performed this experiment in an adr-1(lf) background. Have the authors attempted this experiment in this background? How many independent lines were tested with the wp1923 construct? Is expression seen anywhere else in the nervous system (outside de VNC) to make sure that GFP can indeed be expressed from this transgene?

- Following on this previous point, a direct way to demonstrate the functional importance of this editing site would be to generate the following genotype by engineering the A>G mutation by CRISPR/Cas9 gene editing:slo-2(gf); adr-1(0); scyl-1(A>G)

My prediction would be that the mutation in scyl-1 would restore the slo-2 gain-of-function locomotor impairement, by bypassing the adr-1 requirement. To me this experiment would very strongly support this new and exciting functional regulation and I would encourage the authors to perform this rather simple experiment.

- In wormbase, the annotation of the scyl-1 locus shows a rather sizable 3'UTR. I was curious whether the authors have any comments on that point, and whether this is a common feature of ADAR-edited 3'UTRs.

Does the human SCYL1 3'UTR have a similar hair-pin structure, which could suggest a similar regulation mechanism?

- I was a bit surprised about the strategy used to generate the scyl-1 expression pattern. Why was the in vivo recombination approach used? What happens when the 0.5kb promoter-GFP construct is injected alone?

The previous gene is only approx 2kb upstream and there is significant sequence conservation to C. remanei and C. briggsae DNA less than 1kb upstream of the ATG (see UCSC genome browser for example). Did the authors test such a 2kb promoter fragment?

- Figure 6: I'm not convinced that scyl-1::GFP labels vm1 or vm2 muscles based on this image. I could be wrong, but higher magnification images would need to be checked.

Reviewer #1:

1) My major comment is on the work that extrapolates the nematode findings to the human Slack channel. The latter has previously been shows to differ from the C-elegans channel in several ways. This interaction with the human channel is an important aspect of the presented work. Examination of the data in Figure 11, however, suggest that the gating of the human channel is quite different from the Slo2 channel records shown in Figure 8. Specifically (if the time bars shown in Figure 11A are correct, the overall open probability of the human channel is dominated by the presence of very long closed states (lasting many seconds). Examination of the X-axis scale bars in Figure 11B and C suggest these may have been excluded from the analyses. The conclusions on the effects of SCYL1 on open probability stated in the last paragraph of the Results section may not be completely valid until these are taken into account.

We thank the reviewer for the comment. The open probability in the original Figure 11 was calculated from all the open and closed events. As the reviewer pointed out, events of the very long closed state had a major impact on the open probability. However, they had little contribution to the dwell time histograms in the original figure because these events account for a tiny (nearly negligible) portion of the total event counts and the majority of them fell outside of the x-axis displayed in the figure. In the revised manuscript, we added a new panel (panel D) to Figure 11 to show dwell time histograms of the very long closed events (>30 ms, chosen arbitrarily as the threshold) for both groups. Because these events were scarce, our approach used for comparing τ and A values in panels B and C of this figure and of Figure 8 could not be applied to them. Therefore, we compared the average duration and frequency of these very long closed events between hSlo2.2 and hSlo2.2+SCYL1 (Figure 11D, bottom). We also have replaced the traces in Figure 11A with longer and more representative ones. In addition, the dwell time histograms in panels B and C of both Figures 8 and Figure 11, which were based on a single sample recording trace in the original figures, have been revised to include events from all the recordings to be consistent with the histogram in the new panel D. In the revised manuscript, we describe the presence of the very long closed state and its impact on the open probability (–subsection “scyl-1 expression depends on RNA editing at a specific 3’-UTR site”). The legends of Figure 8 and Figure 11 have also been updated accordingly.

2) The bimolecular fluorescence complementation assay presented does make a case that there may be a physical interaction of SCLY-1 with SLO-2 but is not completely definitive. It would be good to have some other indicator of physical interaction to support this claim. A more conventional coimmunoprecipitation experiments would be a good addition, one that could perhaps be carried out using the co-expression Xenopus oocyte heterologous expression system.

We performed coimmunoprecipitation assays with transgenic worms expressing HA-tagged SCYL-1 and GFP-tagged SLO-2. Worms instead of a heterologous expression system were used for this experiment because SLO-2 is poorly expressed in both Xenopus oocytes and HEK293 cells even with a codon-optimized version (our unpublished observation). Our results of the co-IP assays were in agreement with those of the BiFC assays (Figure 9), which provides further evidence for physical interactions between SLO-2 and SCYL-1.

Reviewer #2:

Comments

1) Please clarify the functional relationship between adr-1 and adr-2. Indeed, in the Introducution the authors seem to indicate that ADAR proteins could compete ("altering the accessibility") for certain binding sites. In this model, wouldn't one expect that adr-1(lf) would "free" access to the scyl-1 site, which would be inconsistent with the similarity of adr-1 and adr-2 mutant phenotypes?

Do the authors think that ADR-1 promotes the recruitement of ADR-2 to the scyl-1 3'UTR?

We thank the reviewer for the comment. Indeed, our description about the functional relationship between ADR-1 and ADR-2 in the Introduction was inaccurate. ADR-1 promotes RNA editing by binding to target mRNAs and recruiting ADR-2 to some specific editing sites. Our results with adr-1(lf) and adr-2(lf) mutants are compatible with this knowledge. In the revised manuscript, a sentence in the Introduction has been changed to “ADR-1 is catalytically inactive but can promote RNA editing by binding to selected target mRNA and tethering ADR-2 to RNA substrates (Ganem et al., 2019; Rajendren et al., 2018; Washburn et al., 2014).” (Introduction)

2) I was very intrigued by the results described in Figure 10, and specifically Figure 10D/E. The result is not what I had intuitively expected. Since it was performed in an adr-1/2 wild-type background as far as I could determine from the Materials and methods section, I would have thought that ADAR activity would have edited the wild-type sequence and I would have expected to see GFP in both cases.

I would have performed this experiment in an adr-1(lf) background. Have the authors attempted this experiment in this background? How many independent lines were tested with the wp1923 construct? Is expression seen anywhere else in the nervous system (outside de VNC) to make sure that GFP can indeed be expressed from this transgene?

We obtained five wp1923 lines and three wp1924 lines. While GFP signal was observed in all the wp1924 lines, no GFP signal was detected anywhere in any of the wp1923 lines. We were also surprised by the lack of GFP expression in the wp1923 lines. As suggested, we performed the same experiments as those in Figure 10E with the adr-1(zw96) mutant. Specifically, adr-1(zw96) mutant strains expressing the wp1923 transgene were made by injecting the wp1923 plasmid into the mutant whereas that expressing the wp1924 transgene by first integrating the wp1924 transgene into the wild-type genome and then crossing it into adr-1(zw96). As expected, GFP signal was not detected in the adr-1(zw96) mutant worms harboring the wp1923 transgene (not shown). Interestingly, GFP signal from the wp1924 transgene was much weaker (by ~50%) in the mutant strain than the wild-type strain (Figure 10 F and G). Because scyl-1 3’-UTR had already been “edited” at the ADR-1-dependent editing site in the wp1924 transgene, the difference in GFP signal between wild-type and adr-1(zw96) strains suggests that ADR-1 can also regulate scyl-1 mRNA level through a post-editing effect, perhaps through interacting with some other proteins. In agreement with this possibility, we were able to isolate several mutants showing decreased GFP signal from the wp1924 transgene in a pilot genetic screen (Figure 10—figure supplement 1).

3) Following on this previous point, a direct way to demonstrate the functional importance of this editing site would be to generate the following genotype by engineering the A>G mutation by CRISPR/Cas9 gene editing:slo-2(gf); adr-1(0); scyl-1(A>G)

My prediction would be that the mutation in scyl-1 would restore the slo-2 gain-of-function locomotor impairement, by bypassing the adr-1 requirement. To me this experiment would very strongly support this new and exciting functional regulation and I would encourage the authors to perform this rather simple experiment.

As suggested by the reviewer, we tried to engineer the A>G mutation at the ADR-1-dependent editing site in scyl-1 3’-UTR in the worm genome by CRISPR/Cas9. Unfortunately, our attempt was unsuccessful, probably because this editing site is located within an inverted sequence (Figure 10D). We then tried to address the reviewer’s comment by expressing Prab-3::scyl-1::unc-10 3’-UTR in slo-2(gf);adr-1(lf) double mutant and wild type (as a control) in the hope that expression of this transgene in the double mutant would restore the slo-2(gf) sluggish locomotion by bypassing the ADR-1 requirement. However, expression of the transgene caused larval arrest in the double mutant and a sluggish phenotype in wildtype worms, which prevented us from performing further analysis. Finally, we built a scyl-1(zw99);adr1(zw96) double mutant, and recorded whole-cell currents from VA5 motor neuron. We observed similar VA5 outward currents between the double mutant and the scyl-1(zw99) single mutant (Figure 7A), suggesting that SCYL-1 and ADR-1 likely contribute to SLO-2 function through a common pathway. This result provides further evidence for the putative role of ADR-1 in regulating SCYL-1 expression.

4) – In wormbase, the annotation of the scyl-1 locus shows a rather sizable 3'UTR. I was curious whether the authors have any comments on that point, and whether this is a common feature of ADAR-edited 3'UTRs.

Does the human SCYL1 3'UTR have a similar hair-pin structure, which could suggest a similar regulation mechanism?

In Wormbase, there are four different scyl-1 transcripts that differ only in the 3’-UTR sequence and length (167 nt, 169 nt, 564 nt, and 1862 nt). The identified ADR-1-dependent editing site exists only in the variant with the longest 3’UTR, which we speculate could be a mechanism for cell-specific expression of scyl-1. It is difficult to comment on whether RNA editing by ADARs at 3’-UTRs is a common mechanism of regulating mRNA expression based on the rather limited literatures. The 3’-UTR of human SCYL1 transcripts (NM_020680.4) also has a high probability of forming hair-pin structures based on software prediction (https://rna.urmc.rochester.edu/RNAstructureWeb/Servers). It remains to be determined whether human SCYL1 transcripts are also edited at the 3’-UTR, and if so, whether the editing regulates SCYL1 transcript expression. We added these comments to the Discussion section in the revised manuscript –.

5) I was a bit surprised about the strategy used to generate the scyl-1 expression pattern. Why was the in vivo recombination approach used? What happens when the 0.5kb promoter-GFP construct is injected alone?

The previous gene is only approx 2kb upstream and there is significant sequence conservation to C. remanei and C. briggsae DNA less than 1kb upstream of the ATG (see UCSC genome browser for example). Did the authors test such a 2kb promoter fragment?

We initially tried to examine scyl-1 expression pattern using a 2-kb Pscyl-1 because, as the reviewer pointed out, another gene (lap-2) resides approximately 2 kb upstream of scyl-1. However, GFP signal was not detected in transgenic worms expressing the Pscyl-1(2 kb)::gfp transcriptional fusion. We therefore resorted to the commonly used in vivo homologous recombination approach to include potential distant upstream regulatory elements in the Pscyl-1::gfp transcriptional fusion. A description about the unsuccessful experiment with the 2-kb Pscyl-1 has been added to the manuscript (–subsection “ADR-1 regulates SLO-2 function through SCYL-1”). As suggested by the reviewer, we also created transgenic worms expressing only the Pscyl-1(0.5 kb)::gfp transcriptional fusion but were unable to detect any GFP signal from them (not shown).

6) Figure 6: I'm not convinced that scyl-1::GFP labels vm1 or vm2 muscles based on this image. I could be wrong, but higher magnification images would need to be checked.

We thank the reviewer for pointing out our mistake. In Figure 6, Pscyl-1::GFP labels uterine ventral cells rather than vulval muscle cells.

Author details

1. Long-Gang Niu

   Department of Neuroscience, University of Connecticut Health Center, Farmington, United States

   Contribution

   Data curation, Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

2. Ping Liu

   Department of Neuroscience, University of Connecticut Health Center, Farmington, United States

   Contribution

   Data curation, Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

3. Zhao-Wen Wang

   Department of Neuroscience, University of Connecticut Health Center, Farmington, United States

   Contribution

   Conceptualization, Resources, Supervision, Funding acquisition, Writing - review and editing

   Competing interests

   No competing interests declared

4. Bojun Chen

   Department of Neuroscience, University of Connecticut Health Center, Farmington, United States

   Contribution

   Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   bochen@uchc.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1141-9101

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