Biological clocks regulate a myriad of processes that occur periodically, from sleeping and waking to how cells use nutrients and energy. One such clock is the one that controls intestinal movements and defecation in the nematode worm Caenorhabditis elegans, which consists of three muscle contractions occurring every 50 seconds. This rhythm is controlled by calcium and proton signalling in the cells of the intestine.

The cells of the nematode intestine form a tube, through which gut contents pass. The inside of the tube is acidic, but acidity also plays a role on the outer face of the intestinal tube. In this area, nutrients are distributed and signals are conveyed to other tissues, such as muscles. In fact, acid – in the form of protons – secreted from the intestinal cells stimulates the muscles that contract in the biological clock that controls the worms’ defecation. However, it is poorly understood how the worms control the release of these protons.

Kaulich et al. identified two ion channels on the membranes of intestinal cells that become inhibited when the levels of acid surrounding them are high. These channels play distinct roles in controlling the contractions that move the contents of the roundworms’ intestines along. The first channel contains a protein called ACD-5, and it is in the membrane of the intestinal cells that faces the inside of the intestinal tube. The second channel is formed by three proteins: FLR-1, ACD-3 and DEL-5. This channel is found on the other side of the intestinal cells, the region where nutrients are distributed and signals are conveyed to the rest of the body.

To determine the role of each channel, Kaulich et al. genetically engineered the worms so they would not make the proteins that make up the channels, and imaged the live nematodes to see the effects of removing each channel. The inside of the intestines of worms lacking the ACD-5 containing channel was less acidic than that of normal worms, and the timing of the contractions that control defecation was also slightly altered. Removing the second channel (the one formed by three different proteins), however, had more dramatic effects: the worms were thin, developed more slowly, had less fat tissue and defecated very irregularly.

Kaulich et al. imaged live worms to show that the second channel plays a major role in regulating oscillations in acidity both inside and outside cells, as well as controlling calcium levels. This demonstrates that this channel is responsible for the rhythmicity in the contractions that control defecation in the nematodes. Their findings provide important insights towards better understanding proton signalling and the role of acid-sensing ion channels in cellular contexts and biological clocks.

Many physiological processes occur with a predictable periodicity. The maintenance of such rhythms can rely on fluctuating gene expression, hormone concentrations, or homeostatic oscillations in signalling molecules within cellular compartments, depending on the timescale of the clock (Hastings et al., 2018; Rensing et al., 2001; Branicky and Hekimi, 2006; Thomas, 1990). Disruption of biological clocks can have devastating consequences on homeostasis or behaviour (Marcheva et al., 2013). Here, we examine the physiological role of acid-sensing ion channels (ASICs) in the defecation motor program (DMP) of the nematode Caenorhabditis elegans, an ultradian rhythm that occurs about every 50 s (Thomas, 1990; Branicky and Hekimi, 2006). We take advantage of this short-period cellular oscillator to relate ion channel physiology in vivo to organismal metabolism and behaviour.

ASICs, the acid-sensing members of the Degenerin/Epithelial Sodium Channel (DEG/ENaC) superfamily of cation channels, are thought to be the main proton receptors in vertebrates (Soto et al., 2018; Du et al., 2017). They are an important drug target because acidosis is a feature of painful inflammatory and ischemic conditions. ASICs are widely expressed in the central and peripheral nervous system, where their roles in inflammation, ischemia, pain perception, and learning are extensively studied (Waldmann et al., 1997; Wemmie et al., 2002; Wemmie et al., 2003; Voglis and Tavernarakis, 2008; Kreple et al., 2014; Du et al., 2014). However, less is known about their physiological role in non-neuronal tissue such as the intestinal epithelium, despite cross-phyla evidence for gastrointestinal expression and the role of acidity therein (Dong et al., 2011; Schaefer et al., 2000; Take-uchi et al., 1998; Levanti et al., 2011). The extreme acidity of the stomach requires an elaborate system of mucosal protective mechanisms, failure of which results in conditions such as gastritis, ulceration, and dyspepsia. Acidification of the intestinal tract is also a hallmark of conditions such as irritable bowel syndrome and cystic fibrosis, so understanding the role of acid sensors, of which the ASICs are prime candidates, is of wide-ranging therapeutic importance (Holzer, 2007).

The DMP depends on inositol 1,4,5-trisphosphate (IP₃)-dependent Ca²⁺ oscillations in the intestinal epithelial cells. IP₃ signalling loss-of-function mutations cause dramatic increases in both period length and variability, whereas overexpression of the IP₃ receptor, ITR-1, reduces period length (Dal Santo et al., 1999; Walker et al., 2002; Espelt et al., 2005). Increased cytosolic Ca²⁺ results in the rhythmic release of protons from the basolateral membrane, into the pseudocoelom. This triggers the posterior body wall muscles to contract (pBoc), via activation of a proton-gated cation channel PBO-5/PBO-6 (Beg et al., 2008; Pfeiffer et al., 2008). Propagation of the Ca²⁺ wave from the posterior to anterior then initiates contraction of the body wall muscle at the anterior of the intestine (aBoc) a few seconds later, via activation of the motor neuron AVL, and then finally, GABA release from both AVL and DVB triggers the enteric muscle contraction (EMC), expelling gut contents (Beg and Jorgensen, 2003; Mclntire et al., 1993). Premature termination of the Ca²⁺ wave or pharmacologically blocking the Ca²⁺ wave propagation before it can reach the anterior disrupts aBoc and EMC (Teramoto and Iwasaki, 2006), further supporting the idea that Ca²⁺ is the master regulator.

In addition to Ca²⁺ and pseudocoelomic pH, proton oscillations in the intestinal lumen are also integral to the DMP. The luminal pH fluctuates between ~4.4 and 6.5 during the cycle and the pH transitions in a wave that travels from posterior to anterior (Beg et al., 2008; Allman et al., 2009; Pfeiffer et al., 2008; Bender et al., 2013; Chauhan et al., 2013), dependent on proton uptake and release at the apical membrane. Proton gradients are fundamental to nutrient absorption, and some DMP-affecting mutations can result in aberrant fat metabolism and deficient nutrient uptake (Sheng et al., 2015; Allman et al., 2009). Loss of proton regulation can also disrupt immunity and pathogen resistance, as seen in mutants for C. elegans CHP1 homolog, PBO-1, in which the lumen is constantly acidic (Benomar et al., 2020). Thus, dysregulation of components of this ultradian clock can have far reaching organismal consequences.

The precise relationship between luminal proton oscillations, intestinal Ca²⁺ and the DMP is unclear; but the mutant phenotypes of some of the known transporters and exchangers that control lumen pH provide some clues. Knockdown of VHA-6, a subunit of the apical proton-transporting V-type ATPase, leads to more neutral lumen pH and an increase in cycle length (Allman et al., 2009). For mutations in the Na⁺/H⁺ exchanger PBO-4, the effect on lumen pH is controversial, since measurement using KR35 dye (Bender et al., 2013; Benomar et al., 2020) reveals a defect in anterior acidification, whereas measurement using Oregon Green 488 (Pfeiffer et al., 2008) reveals that global pH oscillations are normal; cycle length and Ca²⁺ oscillations are unaffected (Thomas, 1990; Pfeiffer et al., 2008; Benomar et al., 2020; Bender et al., 2013; Beg et al., 2008). Mutants of pbo-1, on the other hand, have a highly acidic lumen and no proton wave, but Ca²⁺ oscillations and cycle length are largely unaffected (Wagner et al., 2011; Thomas, 1990; Pfeiffer et al., 2008; Benomar et al., 2020). So, there is some evidence that the periodicity of the DMP can be independent of luminal pH, and thus that proton dynamics would be downstream of the Ca²⁺ oscillator.

We and others hypothesized that a proton wave transitioning from the posterior to the anterior of the lumen could lead to sequential activation of proton sensors in the apical membrane (Chauhan et al., 2013; Bender et al., 2013; Benomar et al., 2020), and acid-sensing DEG/ENaCs are good candidates. Compared to mammals, the C. elegans genome encodes a vastly expanded array of 30 DEG/ENaCs. One of these, FLR-1 (FLuoride-Resistant phenotype), functions in DMP rhythmicity and execution (Kwan et al., 2008; Take-uchi et al., 1998), although, the mechanistic basis of this role is unclear. With the exception of ACD-1, the first acid-inactivated DEG/ENaC to be identified (Wang et al., 2008), the proton sensing properties of the C. elegans DEG/ENaCs remained uncharacterised.

Here, we address the physiological role of C. elegans acid-sensing DEG/ENaCs in the intestine. We explore how Ca²⁺ and proton oscillations maintain periodicity and execution of the DMP, and find that disruption of Ca²⁺ and proton homeostasis are detrimental to metabolism and organismal health. We show that four intestinal DEG/ENaC subunits form two acid-inactivated channels with distinct properties. The first contains ACD-5 and localises to the luminal face of the intestinal cells and acts as a proton sensor, contributing to the establishment of low luminal pH and thus enabling cyclic pH oscillations. The second channel, containing FLR-1, ACD-3, and/or DEL-5, localises to the basolateral membrane, facing the pseudocoelom, and is essential for the Ca²⁺ oscillations that control the timing and execution of the DMP and thus is a key component of this ultradian clock. We thus provide direct evidence linking the channel properties of these proton sensors to their roles in pH regulation and Ca²⁺ signalling in vivo, which can provide insight into potential cellular roles for other acid-sensing members of this important channel family.

Using a combination of electrophysiology, behavioural analysis, genetic manipulations, and pH and Ca²⁺ indicator imaging, we have obtained evidence for two acid-sensitive DEG/ENaC channels with distinct functions (Figure 9). One includes ACD-5, which localises to the luminal membrane and is the main proton sensing subunit of the channel. This proton sensor is required for maintenance and/or establishment of acidic luminal pH and yet only subtly influences the DMP, and the Ca²⁺ oscillator. The second, composed of FLR-1, ACD-3, and/or DEL-5, localises to the basolateral membrane and is essential for timing and execution of DMP muscle contractions, with consequences for fat metabolism and growth.

Figure 9

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ACD-5 and FLR-1-containing channels exhibit very different pH dependence and kinetics; ACD-5 is fully open at pH 6 and closed at pH 4, reflecting the range of pH oscillations in the lumen (Allman et al., 2009), whereas FLR-1-containing heteromers form a proton-inhibited cation leak channel, similar to ACD-1, TadNaC6 and vertebrate ASIC5 (Wang et al., 2008; Elkhatib et al., 2019; Wiemuth and Gründer, 2010). This presumably reflects their different physiological contexts, just as the pH dependence of human ENaCs reflects the pH range in epithelia where they are expressed (Collier and Snyder, 2009). Neither ACD-5 nor FLR-1 currents desensitise; the channel remains open for the duration of the pH stimulus (although we see some evidence of partial desensitisation of FLR-1 at certain pHs). This contrasts with the fast desensitisation of mammalian ASICs and is more analogous to the slower kinetics of ENaCs. It coincides well with our evidence that ACD-5 is actually required for establishment or maintenance of acidic luminal pH; that is, its activation ensures a return to the condition (pH~4) at which it is closed. By contrast, the FLR-1/ACD-3/DEL-5 heteromeric channel on the basolateral membrane is predicted to be constitutively open as the extracellular resting pH is around 7.35 (Allman et al., 2009).

All four intestinal DEG/ENaC subunits influence the DMP, but the degree of influence differs dramatically, providing us with mechanistic information about the relationships between extracellular and intracellular events. Research to date indicates that intestinal IP₃-dependent Ca²⁺ signalling act as the master oscillator and is responsible for cycle length and timing. Disrupting IP₃ signalling results in a dramatic lengthening and disruption of rhythmicity of the DMP, and a direct relationship is seen between the timing of Ca²⁺ fluctuations and the DMP (Kwan et al., 2008; Walker et al., 2002; Espelt et al., 2005; Teramoto and Iwasaki, 2006; Dal Santo et al., 1999). Our acd-5 mutant data supports this hypothesis; the underlying Ca²⁺ fluctuations, and DMP rhythmicity, were only subtly affected, despite dramatic disruption of luminal pH, underlining the dominance of the Ca²⁺ oscillator. Where detectable, the MAT interval length was also not affected, suggesting that ACD-5 is not responsible for the timing of the proton wave, but acts downstream. This is supported by the phenotype of animals deficient in the V-ATPase, VHA-6; lumen pH remained more neutral, but robust Ca²⁺ oscillations were maintained, with only a small increase in cycle length (similar to acd-5(ok2657)) (Allman et al., 2009). This suggests a functional link between the two; a pH~6-activated cation channel and a presumed H⁺ pump whose effect would be to acidify the lumen, although the vha-6 phenotype is more severe than acd-5, interfering with nutrient uptake and growth rate, indicating that there is not a complete overlap in their roles. Indeed, such a coupling has been proposed in Na⁺ uptake in the Zebrafish larva (Zimmer et al., 2018). We could speculate that the dominant mutation could result in interference with VHA-6 function, or alternatively it could be interfering with localisation or function of the other DEG/ENaC subunits (as we have shown that it can do for ACD-5 itself, when expressed in oocytes). Altering the subunit composition of the FLR-1-containing channels, rather than disrupting it completely, could result in such a subtle extension. Whatever the mechanism, it is clear that the Ca²⁺ oscillator is largely unperturbed. We do nevertheless see some influence; in particular, when the Ca²⁺ oscillator is disrupted, in itr-1 deficient animals, the very long cycles observed are dependent on acd-5, revealing an otherwise constraining role. The Ca²⁺ oscillator is thus upstream, and/or largely independent, of lumen pH, and is the dominant force driving DMP rhythmicity.

In stark contrast to acd-5 mutants, flr-1 deficient animals exhibit a dramatic dysregulation of the DMP, with a combination of very long and very short cycles, underlining their importance for rhythmicity (as previous research showed; Kobayashi et al., 2011; Kwan et al., 2008; Katsura et al., 1994; Take-uchi et al., 1998), which is phenocopied by acd-3/del-5 double mutants. The lack of a clear phenotype for acd-3 and del-5 single mutants suggests that they act redundantly, in a heteromer, along with FLR-1. This correlates well with our observation that FLR-1 pH-sensitive currents in Xenopus oocytes were dependent on the presence of one or other of these subunits, and that acd-3/del-5 double mutant phenocopy other aspects of flr-1 loss of function: EMC and pBoC defects, developmental timing, growth, and fat accumulation. The severity of disruption to the DMP is reminiscent of that seen when Ca²⁺ signalling is disrupted (Kwan et al., 2008; Walker et al., 2002; Espelt et al., 2005; Dal Santo et al., 1999). Indeed, we found that in flr-1(syb3521) animals, the rhythmicity and magnitude of intestinal Ca²⁺ transients, measured in the posterior cells, where the Ca²⁺ wave initiates, were completely disrupted. This indicates that the FLR-1/ACD-3/DEL-5 channel(s) is upstream of the Ca²⁺ oscillator, perhaps forming part of the ‘master oscillator’, linking extracellular proton concentration to the initiation of the intracellular Ca²⁺ wave that controls periodicity and execution of the DMP (see Figure 9).

Interestingly, in contrast to itr-1, the flr-1 and acd-3/del-5 deficient animals also exhibit very short cycles, and ‘echos’ of the DMP as well as missed or weak EMC or pBoc muscle contractions which has previously been described for the unc-43 CaMK II mutant (Teramoto and Iwasaki, 2006) and cmd-1 (calmodulin) RNAi (Allman et al., 2013), which again links the phenotypes of these mutants to the disturbance in intestinal Ca²⁺ signalling. These shorter cycles, for which we also saw evidence in both intracellular pH and Ca²⁺ oscillations, indicate a role for calmodulin, CaMK II and the FLR-1 channel in retarding a shorter timescale oscillator. Cycle variability, the other effect of flr-1 or acd-3/del-5 mutations, presumably represents a loss of synchronicity between semi-independent oscillators, each of which has an innate homeostatic rhythm—but is checked by a delay in reaching permissive conditions provided by another. A channel that is open for most of the cycle is consistent with a role in maintaining cellular excitability, and thereby underpinning intracellular Ca²⁺ signalling; closing briefly following proton release into the pseudocoelom would serve to terminate the signal. This model fits well with the relatively low currents observed for the FLR-1 channel (e.g., compared with ACD-5) and the temporal relationship observed (Allman et al., 2009), where Ca²⁺ concentration precisely follows the decline in pseudocoelom pH.

Notwithstanding the challenges associated with imaging freely moving animals, two important barriers to dissecting the relationships between events in the intestinal cells and their extracellular environments are the dependent crosstalk between signalling events and the fact that some components are expressed on both the apical and basolateral membranes. PBO-1, for example, localises to both membranes, where it has distinct roles (Allman et al., 2016). Our data for ACD-3 and DEL-5 localisation to the basolateral membrane supports previous evidence for FLR-1 (Kobayashi et al., 2011). We did not detect apical localisation for ACD-3 or DEL-5 subunits. However, we did observe widespread, diffuse localisation within many cells, which might make any potential apical localisation difficult to distinguish, and Take-uchi et al., 1998 reported apical localisation (Take-uchi et al., 1998), so we cannot exclude the possibility of a dual role at the two membranes. Basolateral localisation fits well with the pH dependence of the channel. However, an important remaining question is how exactly the FLR-1 channel influences Ca²⁺ and calmodulin signalling, and the broad permeability of the channel does not help to shed light on this problem. Of key importance, and explaining why ACD-5 and FLR-1 have such distinct effects, may be the local nature of signalling events, with their activity thus specifically influencing components within their immediate vicinity.

We have shown that two acid-sensing DEG/ENaC channels with distinct properties play very different roles in the interplay between extracellular pH and intracellular Ca²⁺. ACD-5 is required to maintain acidic lumen pH and has a subtle impact on the Ca²⁺ oscillations that control DMP timing. In contrast, FLR-1, ACD-3, and/or DEL-5-containing channel is the essential link between pseudocoelomic pH and Ca²⁺, and thus timing and execution of the DMP. DEG/ENaCs are conserved across the animal kingdom, and the ASICs and ENaC members are relevant to a broad range of medical conditions, and thus of significant therapeutic importance. Our data provide an insight into the relationship between the acid-sensing properties of these channels and their physiological role in epithelia. However, the family contains a diversity of members with distinct properties, and vastly expanded numbers in some species. Better understanding the relevance of channel properties to function will help us to better understand their role, and the role of acid-sensing, in other cellular contexts.

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Decision letter after peer review:

Thank you for submitting your article "Distinct roles for two Caenorhabditis elegans acid-sensing ion channels in an ultradian clock" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Reviewing Editor and Piali Sengupta as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another and share their enthusiasm about your work. However, their reviews highlight multiple gaps in your study that require some extensive and essential revisions. The Reviewing Editor has drafted this to help you prepare a revised submission.

Please find below a detailed list of all reviewer comments. Among these, we found most essential to:

(i) Provide complete tissue specific rescue- and subcellular localization studies of the respective channels (R1#1, R2#9-10),

(ii) Perform more detailed pH measurements in both intestine and the pseudocoelom (R2#9-10),

(iii) Address the discrepancy between the genetic effects on intestinal calcium waves and apparently normal pBoc and Exp contractions (R2#11), including an improved discussion of the differential effects you observe (R3#19),

(iv) Provide a biophysical characterization of the putative FLR-1/ACD-3/DEL-5 complex, like you nicely did for ACD-5 (R2#12, R3#18).

We believe that the remaining points can be easily addressed with standard analyses and text revisions. We are looking forward to reading a revised manuscript.

Reviewer #1 (Recommendations for the authors):

(1) Is expression of flr-1, acd-3 and del-5 restricted to the intestine as partially suggested by Figure 4A-B. The authors mention a previous study that acd-3 and del-5 are also expressed in neurons implicated in the defecation rhythm; can this be confirmed with the reported marker line here? And how about flr-1? I think the study could be improved by tissue specific rescue experiments of all reported phenotypes (incl. Figure 7) the of these three genes.

(2) The authors report for some of their assays "excessive" long cycles in some mutants (e.g. Figure 3D,E); these findings should explicitly quantified with the adequate comparison groups; for example using K-S test complimentary cumulative distribution or any appropriate test.

Reviewer #2 (Recommendations for the authors):

(1) Figure 2: The authors show that acd-5 mutants have higher pH in the anterior intestinal lumen during MAT, compared to wild type controls, as revealed by KR35 imaging. Is pH only increased in the anterior intestine during MAT, or are pH levels lower in other parts of the intestine and at other times of the cycle in acd-5 mutants? In other words it would be helpful to know whether acd-5 regulates global pH throughout the intestine or whether it only affects pH during MAT.

(2) Figure 4A: the authors conclude that the FLR-1 channels function at the basolateral surface of the intestine based on the localization shown in Figure 4A. In the images shown, it is not obvious that this is the case. They appear more diffuse throughout the intestine. Are the fusion proteins functional? Do they co-localize with known basolateral markers? Do mutants in the channel affect pseudocoelom pH? These questions should be addressed if the authors want to make conclusions regarding the subcellular site of action.

A similar analysis looking at the effect of flr-1 mutations on pseudocoelom pH would really help here (a phlourin reporter has been used for this purpose), especially since they claim that flr-1 functions on the pseudocoelom surface of the intestine, but they show no data to support this claim.

(3) The issue of why flr-1 mutants have apparently absent intestinal calcium waves, yet have near normal pBoc and Exp contractions needs to be sorted out. In Figure 6 reports a calcium wave defect, but only show some representative traces are shown, with no quantification. However, the text states that calcium oscillations are "completely disrupted". Either calcium waves aren't very important for the pBoc and Exp contractions, which does not agree with previous research or their model, or more likely, calcium waves are not "completely disrupted", but are more intact than they report. It is further unclear if the traces represent fluorescence in the posterior intestine only or of the entire intestine. Is there a defect in the anterior propagation of calcium waves? Is there a correlation in animals that do not exhibit calcium transients and a lack of pBoc, or EMC? If calcium transients are eliminated in flr-1 mutants, why do animals exhibit pBoc in 90% of cycles and EMC in 50% of cycles? Is there another oscillator, like pH in the intestine that drives these steps? Can the authors measure pH changes in the intestine during EMC to address this?

(4) Figure 4. Can the authors present an analysis of which ions pass through FLR-1/ACD-3/DEL-5, as they did for ACD-5? Knowing this, particularly if these pass calcium, may help the authors to formulate a mechanism by which FLR-1 regulates intestinal calcium dynamics.

Reviewer #3 (Recommendations for the authors):

(1) The authors have done a nice job in characterizing the biophysical properties of ACD-5. But their characterization of FLR-1/ACD-3/DEL-5 is a bit cursory, and the data from FLR-1/ACD-3/DEL-5 recordings is not nearly as convincing. Is the FLR-1/ACD-3/DEL-5 heteromeric channel sensitive to amiloride? The authors claim that it is a leaky cation channel, but does not provide much evidence to support this claim. For example, is it impermeable to NMDG like ACD-5 (to show it is a cation channel)? Is it a monovalent channel like ACD-5 or also permeable to divalent ions? Is it sensitive to ca²⁺ inhibition like ACD-5?

(2) The authors have proposed a nice model in Figure 8, but it is still not so clear to me why FLR-1/ACD-3/DEL-5 affects ca²⁺ wave but ACD-5 does not. Both channels are inhibited by acidic pH, though ACD-5 is also inhibited by high pH; and both channels are non-desensitizing. Their distinct subcellular localization patterns do not seem to explain it. The authors need to put up a better explanation to this interesting phenomenon in the discussion.

Reviewer #1 (Recommendations for the authors):

(1) Is expression of flr-1, acd-3 and del-5 restricted to the intestine as partially suggested by Figure 4A-B. The authors mention a previous study that acd-3 and del-5 are also expressed in neurons implicated in the defecation rhythm; can this be confirmed with the reported marker line here? And how about flr-1? I think the study could be improved by tissue specific rescue experiments of all reported phenotypes (incl. Figure 7) the of these three genes.

Apologies for omitting the transcriptional fusion data, showing that, with our constructs, we only see expression in the intestine. We have added Figure 4—figure supplement 1 – transcriptional fusions, together with a description in the text (page 10 line 258). We have also added this information where we refer to the CENGEN data (page 12 line 331)

We agree that tissue-specific rescue is an important question. We have inserted two additional figures (Figure 5—figure supplement 3 and Table 1), showing that intestinal expression of acd-3 or del-5 rescues the cycle length, cycle variability, missed and weak pBocs, missed EMCs and echos phenotypes of the acd-3/del-5 double mutant, and text describing it (page 12 line 334), as well as adding that Take-Uchi et al., (1998) previously reported that an intestinally-expressed fusion rescues flr-1.

(2) the authors report for some of their assays "excessive" long cycles in some mutants (e.g. Figure 3D,E); these findings should explicitly quantified with the adequate comparison groups; for example using K-S test complimentary cumulative distribution or any appropriate test.

Thank you very much for this excellent suggestion. We have added Figure 3—figure supplement 4, showing the cumulative distribution and results of a K-S test, and text referring to it (page 9 line 229).

Reviewer #2 (Recommendations for the authors):

(1) Figure 2: The authors show that acd-5 mutants have higher pH in the anterior intestinal lumen during MAT, compared to wild type controls, as revealed by KR35 imaging. Is pH only increased in the anterior intestine during MAT, or are pH levels lower in other parts of the intestine and at other times of the cycle in acd-5 mutants? In other words it would be helpful to know whether acd-5 regulates global pH throughout the intestine or whether it only affects pH during MAT.

We have added a better description in the text (page 7 line 183). As shown in the Figure 2 example pictures, the posterior part of the intestine is often too low in fluorescence to quantify. The KR35 dye is fully fluorescent at pH <~2, with a pKa of 3.5, and is non-fluorescent at a pH of ~5.5. See Figure 2H, Bender et al., 2013 for a full characterization of the pH/fluorescence relationship. Thus, observing animals with low fluorescence indicates a more neutral pH during the resting phase.

(2) Figure 4A: the authors conclude that the FLR-1 channels function at the basolateral surface of the intestine based on the localization shown in Figure 4A. In the images shown, it is not obvious that this is the case. They appear more diffuse throughout the intestine. Are the fusion proteins functional? Do they co-localize with known basolateral markers? Do mutants in the channel affect pseudocoelom pH? These questions should be addressed if the authors want to make conclusions regarding the subcellular site of action.

A similar analysis looking at the effect of flr-1 mutations on pseudocoelom pH would really help here (a phlourin reporter has been used for this purpose), especially since they claim that flr-1 functions on the pseudocoelom surface of the intestine, but they show no data to support this claim.

Getting better localisation data has proven problematic; where detectable, we see a diffuse distribution, as was seen for FLR-1 (in the paper from Take-uchi et al., 1998), making membrane localisation difficult to distinguish. We have tried several different fusions for each, and different concentrations, as well as CRISPR insertions (which were not detectable).

We have added additional images to figure 4A and a longer description to better emphasise that, while we can detect basolateral localisation, we cannot exclude that the channel is also elsewhere (page 10 line 265). In the discussion (page 17 line 500), we had qualified our assumption that the channel is basolateral, referring to a previous report that FLR-1 may also be localised to the luminal membrane (Take-Uchi et al., 1998). We have expanded this to also say that we cannot exclude that ACD-3 and DEL-5 are also luminal, since, in cells where expression appears to be higher, widespread diffuse localisation could prevent us from distinguishing apical localisation. We have also added another supplementary figure (Figure 4—figure supplement 5) exploring potential electrophysiological interactions of the subunits with the luminal ACD-5. We did not detect any changes in pH-sensitivity when we co-expressed any of the subunits together with ACD-5.

We have added figure 5—figure supplement 3 and table 1, showing that the fusions rescue the double mutant

We have added a new figure (figure 7), showing that flr-1 RNAi does indeed disrupt pseudocoelomic pH oscillations.

We have also added Figure 4—figure supplement 5, showing that co-expression of FLR-1, ACD-3 or DEL-5 does not alter the properties of ACD-5, which helps to support our hypothesis that these are functionally separate channels.

(3) The issue of why flr-1 mutants have apparently absent intestinal calcium waves, yet have near normal pBoc and Exp contractions needs to be sorted out. In Figure 6 reports a calcium wave defect, but only show some representative traces are shown, with no quantification. However, the text states that calcium oscillations are "completely disrupted". Either calcium waves aren't very important for the pBoc and Exp contractions, which does not agree with previous research or their model, or more likely, calcium waves are not "completely disrupted", but are more intact than they report. It is further unclear if the traces represent fluorescence in the posterior intestine only or of the entire intestine. Is there a defect in the anterior propagation of calcium waves? Is there a correlation in animals that do not exhibit calcium transients and a lack of pBoc, or EMC? If calcium transients are eliminated in flr-1 mutants, why do animals exhibit pBoc in 90% of cycles and EMC in 50% of cycles? Is there another oscillator, like pH in the intestine that drives these steps? Can the authors measure pH changes in the intestine during EMC to address this?

Apologies, we have described our observations poorly. We have inserted additional descriptions (page 12 line 343) to better explain that calcium transients are not eliminated, but their rhythmicity is severely disrupted We see a mix of traces completely lacking calcium peaks, traces in which very small, shorter interval (< 17 second) peaks are discernible, and traces where larger magnitude, but extremely noisy and variable, calcium fluctuations occur. The variability, and particularly the noisy fluctuations, make further meaningful quantification difficult. We cannot satisfactorily identify all pBoc events in these videos, particularly the weaker ones, so cannot correlate this information with the calcium transients. However, the mix of transients observed appears to correlate well with the mix of long, short and “normal” cycle intervals, weak and missed pBocs, and missed EMCs observed.

We have included 2 additional supplemental figures (Figure 6—figure supplements 1 and 2) to show all of the remaining ca²⁺ traces, illustrating this diversity, along with wild type controls.

We have also updated the legends for these figures, to make clear that we are imaging in the posterior cells only.

We have added an additional figure (figure 7), where we show that flr-1 RNAi disrupts both intracellular and pseudocoelomic pH oscillations, together with a description in the text (page 13 line 358). The dip in pseudocoelomic pH that is thought to trigger pBoc is severely attenuated, agreeing well with the pBoc phenotype. Likewise, intracellular pH exhibits a similar basal level to wild type, but the acidification at pBoc is less extensive.

(4) Figure 4. Can the authors present an analysis of which ions pass through FLR-1/ACD-3/DEL-5, as they did for ACD-5? Knowing this, particularly if these pass calcium, may help the authors to formulate a mechanism by which FLR-1 regulates intestinal calcium dynamics.

We agree that this is an important question. We have added panels (Figure 4E, F, G) to show the ion selectivity, as well as the lack of amiloride sensitivity, along with a description (page 11 line 288).

Reviewer #3 (Recommendations for the authors):

(1) The authors have done a nice job in characterizing the biophysical properties of ACD-5. But their characterization of FLR-1/ACD-3/DEL-5 is a bit cursory, and the data from FLR-1/ACD-3/DEL-5 recordings is not nearly as convincing. Is the FLR-1/ACD-3/DEL-5 heteromeric channel sensitive to amiloride? The authors claim that it is a leaky cation channel, but does not provide much evidence to support this claim. For example, is it impermeable to NMDG like ACD-5 (to show it is a cation channel)? Is it a monovalent channel like ACD-5 or also permeable to divalent ions? Is it sensitive to ca²⁺ inhibition like ACD-5?

We absolutely agree that it is important to include this data. We have added panels (Figure 4E, F, G) to show the ion selectivity and lack of amiloride sensitivity, along with a description (page 11 line 288).

We have also added Figure 4 —figure supplement 5, showing that co-expression of FLR-1, ACD-3 or DEL-5 does not alter the properties of ACD-5.

(2) The authors have proposed a nice model in Figure 8, but it is still not so clear to me why FLR-1/ACD-3/DEL-5 affects ca²⁺ wave but ACD-5 does not. Both channels are inhibited by acidic pH, though ACD-5 is also inhibited by high pH; and both channels are non-desensitizing. Their distinct subcellular localization patterns do not seem to explain it. The authors need to put up a better explanation to this interesting phenomenon in the discussion.

We agree that there are questions remaining, particularly how precisely the FLR-1 channel exerts its effect on ca²⁺ and calmodulin signalling. We have added additional discussion on this topic, including that the low currents observed for the FLR-1 channel would be more consistent with a “leak” channel that is open for much of the cycle, and discussion of the likely role of spatially confined signalling events (page 58).

Author details

1. Eva Kaulich

   Neurobiology Division, MRC Laboratory of Molecular Biology, Cambridge, United Kingdom

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Performed all experiments except calcium and pH imaging, Resources, Visualization, Writing - original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-0868-3702

2. Trae Carroll

   Department of Pathology and Lab Medicine, University of Rochester Medical Center, Rochester, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Performed and analysed intracellular and pseudocoelomic pH experiments, Writing – review and editing

   Competing interests

   No competing interests declared

3. Brian D Ackley

   Department of Molecular Biosciences, University of Kansas, Lawrence, United States

   Contribution

   Conceptualization, Data curation, Funding acquisition, Investigation, Methodology, Performed luminal pH experiments, Resources, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1257-2407

4. Yi-Quan Tang

   1. Neurobiology Division, MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
   2. State Key Laboratory of Medical Neurobiology and MOE Frontiers Center for Brain Science, Institutes of Brain Science, Fudan University, Shanghai, China

   Contribution

   Methodology, Supervision, Writing – review and editing, Provided significant technical guidance for electrophysiology

   Competing interests

   No competing interests declared

5. Iris Hardege

   Neurobiology Division, MRC Laboratory of Molecular Biology, Cambridge, United Kingdom

   Contribution

   Methodology, Supervision, Writing – review and editing, Provided significant technical guidance for electrophysiology

   Competing interests

   No competing interests declared

6. Keith Nehrke

   Department of Medicine, Nephrology Division, University of Rochester Medical Center, Rochester, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Performed and analysed intracellular and luminal pH measurements, Resources, Supervision, Writing – review and editing

   Competing interests

   No competing interests declared

7. William R Schafer

   1. Neurobiology Division, MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
   2. Department of Biology, KU Leuven, Leuven, Belgium

   Contribution

   Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Writing – review and editing

   For correspondence

   wschafer@mrc-lmb.cam.ac.uk

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6676-8034

8. Denise S Walker

   Neurobiology Division, MRC Laboratory of Molecular Biology, Cambridge, United Kingdom

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Performed calcium imaging experiments, Supervision, Writing - original draft, Writing – review and editing

   For correspondence

   dwalker@mrc-lmb.cam.ac.uk

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1534-1679

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