Differential Regulation of Degradation and Immune Pathways Underlies Adaptation of the Ectosymbiotic Nematode Laxus Oneistus to Oxic-Anoxic Interfaces

doi:10.21203/rs.3.rs-1107288/v1

Download PDF

Research Article

Differential Regulation of Degradation and Immune Pathways Underlies Adaptation of the Ectosymbiotic Nematode Laxus Oneistus to Oxic-Anoxic Interfaces

https://doi.org/10.21203/rs.3.rs-1107288/v1

This work is licensed under a CC BY 4.0 License

You are reading this latest preprint version

Eukaryotes may experience oxygen deprivation under both physiological and pathological conditions. Because oxygen shortage leads to a reduction in cellular energy production, all eukaryotes studied so far conserve energy by suppressing their metabolism. However, the molecular physiology of animals that naturally and repeatedly experience anoxia is underexplored. One such animal is the marine nematode Laxus oneistus. It thrives, invariably coated by its sulfur-oxidizing symbiont Candidatus Thiosymbion oneisti, in anoxic sulfidic or hypoxic sand. Here, transcriptomics and proteomics showed that, whether in anoxia or not, L. oneistus mostly expressed genes involved in ubiquitination, energy generation, oxidative stress response, immune response, development, and translation. Importantly, ubiquitination genes were also highly expressed when the nematode was subjected to anoxic sulfidic conditions, together with genes involved in autophagy, detoxification and ribosome biogenesis. We hypothesize that these degradation pathways were induced to recycle damaged cellular components (mitochondria) and misfolded proteins into nutrients. Remarkably, when L. oneistus was subjected to anoxic sulfidic conditions, lectin and mucin genes were also upregulated, potentially to promote the attachment of its thiotrophic symbiont. Furthermore, the nematode appeared to survive oxygen deprivation by using an alternative electron carrier (rhodoquinone) and acceptor (fumarate), to rewire the electron transfer chain. On the other hand, under hypoxia, genes involved in costly processes (e.g., amino acid biosynthesis, development, feeding, mating) were upregulated, together with the worm’s Toll-like innate immunity pathway and several immune effectors (e.g., Bacterial Permeability Increasing proteins, fungicides).

In conclusion, we hypothesize that, in anoxic sulfidic sand, L. oneistus upregulates degradation processes, rewires oxidative phosphorylation and by reinforces its coat of bacterial sulfur-oxidizers. In upper sand layers, instead, it appears to produce broad-range antimicrobials and to exploit oxygen for biosynthesis and development.

General Microbiology

Immunology

Cellular Metabolism

cellular energy production

metabolism

energy generation

anoxic sulfidic conditions

Fluctuations that lead to a decrease in oxygen availability are common in nature (Hermes-Lima and Zenteno-Savin, 2002). The physiological and behavioral response to oxygen deprivation has been studied in animals that naturally experience oxygen deprivation, such as frogs, goldfish, and turtles (Hochachka et al., 1996; 1997, 2001; Hermes-Lima and Zenteno-Savin, 2002), as well as in invertebrate genetic models (Clegg 1997; Nystul et al., 2003; Teodoro and O’Farrell, 2003; Haddad 2006). When oxygen deprived, these organisms must face the challenge of a drastic drop in ATP (the energy-storing metabolite adenosine triphosphate) production, which leads to the failure of energy-demanding processes that are crucial for maintaining cellular homeostasis. Anoxia-tolerant organisms, however, are capable to save energy by stopping energy-costly cellular functions (e.g., protein synthesis, ion pumping, cell cycle progression), maintain stable and low permeability of membranes, and produce ATP by anaerobic glycolysis (Hochachka et al., 1996; Teodoro and O’Farrell, 2003; Liu & Simon, 2004; Liu et al., 2006; Galli et al., 2014).

When parasitic and free-living nematodes, including the model organism Caenorhabditis elegans, are experimentally exposed to anoxia (<0.001 kPa O₂), the intracellular ATP/ADP ratio drops dramatically and, within 10 h, they enter a state of reversible metabolic arrest called suspended animation. Namely, they stop to eat, move, develop or lay eggs, implying that oxygen deprivation affects their growth and behavior (Van Voorhies et al., 2000; Padilla et al., 2002; Nystul and Roth, 2004; Powell-Coffmann 2010; Fawcett et al., 2015; Kitazume et al., 2018). If these effects can be reversed upon oxygen reestablishment, the latter can also provoke a massive and sudden production of reactive oxygen species (ROS) that may overwhelm the organism’s antioxidant defense, and cause its death (reviewed in Hermes-Lima and Zenteno-Savin, 2002). Of note, an increase of mitochondrial ROS production was also observed in worms under hypoxia, because of the inefficient transfer of electrons to molecular oxygen (Nystul and Roth, 2004; Kim and Jin, 2015).

Because oxygen diffuses slowly through aqueous solutions, sharp concentration gradients of this electron acceptor may occur in marine environments and wet soil (Denny et al., 1993; Fawcett et al., 2015). It is at oxic-anoxic interfaces of marine sands that free-living nematodes coated with sulfur-oxidizing Gammaproteobacteria (Stilbonematinae) abound (Ott et al., 1989, 1991; Schiemer et al., 1990; Paredes et al., 2021). However, up to this study, the molecular mechanisms allowing symbiotic nematodes to withstand anoxia, and the inherent stress it is known to inflict upon metazoans, were unknown. Here, we incubated Laxus oneistus (Ott et al., 1995) in conditions resembling those it encounters in its natural environment (i.e. anoxic sulfidic or hypoxic), and applied comparative transcriptomics, proteomics and lipidomics, to understand how it copes with oxygen deprivation. Contrarily to our expectations, in anoxic sulfidic water Laxus oneistus did not appear to enter suspended animation. However, it upregulated genes required for ribosome biogenesis and energy generation, and for degradation pathways (e.g., ubiquitination-proteasome systems, autophagy) likely involved in recycling damaged cellular components and misfolded proteins into nutrients. Notably, under anoxic sulfidic conditions, it also upregulated putative symbiont-binding molecules such as lectins. In the presence of oxygen, on the other hand, the worm appeared to overexpress genes involved in energy-demanding processes (e.g., amino acid synthesis, development, feeding, and mating) and upregulated the synthesis of broad-range antimicrobials, likely via triggering the Toll/NF-kB pathway.

The nematode Laxus oneistus did not enter suspended animation upon 24 h anoxia

To survive anoxia, nematodes enter suspended animation to suppress metabolism and conserve energy. The most notorious sign of suspended animation is the arrest of motility (Nystul et al., 2003; Chan et al., 2010; Kitazume et al., 2018).

Surprisingly, although the whole population of four tested nematode species, including C. elegans, was reported to be in suspended animation upon 10 h in anoxia (Kitazume et al., 2018), L. oneistus kept moving not only after 24-h-long incubations, but also upon 6-day-long incubations in anoxic seawater (three batches of 50 worms were incubated under each condition). Additionally, the symbiotic nematodes appeared morphologically normal (Supplemental movies 1-4).

The fact that we could not observed suspended animation, led us to hypothesize that L. oneistus evolved different strategies to survive oxygen deprivation.

Stable transcriptional profile under hypoxic or anoxic sulfidic conditions

To understand the molecular mechanisms underlying L. oneistus response to oxygen, we subjected it to various oxygen concentrations. Namely, nematode batches were incubated under either normoxic (100% air saturation; O), hypoxic (30% air saturation; H) or anoxic (0% air saturation; A) conditions for 24 h. Additionally, given that L. oneistus thrives in reduced sand containing up to 25 µM sulfide (Ott and Novak., 1989; Paredes et al., 2021), we also incubated it in anoxic seawater supplemented with < 25 µM sulfide (anoxic sulfidic condition; AS).

While transcriptional differences of its symbiont (Candidatus Thiosymbion oneisti), incubated under normoxic (O) and hypoxic (H) conditions were negligible (Paredes et al., 2021), the expression profiles of nematode batches incubated under O conditions varied so much that they did not cluster (Figure S1). Consequently, there was no detectable differential expression between the transcriptomes of O nematodes and any of the other transcriptomes (H, A or AS; Figure S1B, C). We attribute the erratic transcriptional response of L. oneistus to normoxia to the fact that this concentration is not typically experienced by L. oneistus (Ott et al., 1989; Paredes et al., 2021).

As for the expression profiles of nematodes subjected to the H, A or AS conditions, replicates of each condition behaved more congruently (Figure S1B). While we did not find any significant difference between the A and AS nematodes, only 0.05% of the genes (8 genes; Data S1) were differentially expressed between the H and A nematodes and there was no significant difference between the H and A proteomes (t-test, FDR, Benjamini-Hochberg correction, p < 0.05; Figure S2A, Data S1). However, 4.8% of the expressed genes (787 out of 16,526) were differentially expressed between H and AS nematodes, with 434 upregulated under AS and 353 genes upregulated under H conditions (Figure S1C, Data S1).

Collectively, our data suggests that L. oneistus may be ill-equipped to handle normoxic sediment, but it maintains a largely stable physiological profile under both hypoxic and anoxic sulfidic conditions. Before discussing the subset of biological processes differentially upregulated in AS versus H nematodes and vice versa, we will present the physiological processes the worm appears to mostly engage with, irrespectively of the environmental conditions we experimentally subjected it to.

Top-expressed transcripts under all tested conditions

To gain insights on L. oneistus basal physiology, we treated all the 16 transcriptomes as biological replicates (i.e., O, H, A and AS transcriptomes were pooled) and identified the 100 most abundant transcripts out of 16,526 based on functional categories extracted from the UniProt database (2021) and comprehensive literature search (Figure 1, Data S2). Our manual classification was supported by automatic eggNOG classification (Data S2). Similarly, the H and A proteomes were pooled, and the 100 most abundant proteins out of 2,626 were detected (Figure S2).

Based on median gene expression values of the top 100 expressed genes, we found that some of the processes L. oneistus mostly engages with were ubiquitination (ubq-1, Stringham et al., 1992), energy generation (globin glb-1-like (Geuens et al., 2010), cytochrome c oxidase I subunit ctc-1 (UniProtKB P24893), nduo-4-like (UniProtKB P24892), stress response and detoxification (e.g., hsp-1, hsp-90, hsp12.2, and catalases ctl-1 and ctl-2; Birnby et al., 2000; Chávez et al., 2007), and immune defense (lysozyme-like proteins and lec-3) (Figure 1, Data S2).

Lastly, 48 out of the top 100 most expressed genes, were also detected among the top 100 proteins (Figure 1, Figure S2, and Data S2, Supplemental material). Despite the modest correlation between transcript and protein expression levels (r = 0.4) (Figure S3), there was an overlap in the detected biological processes (e.g., energy generation, stress response or detoxification categories, carbohydrate metabolism, cytoskeleton, locomotion, nervous system) (Figure S2).

All in all, except for those encoding for immune effectors, top-transcribed L. oneistus genes could not be ascribed to its symbiotic lifestyle. This differs to what observed for other chemosynthetic hosts, such as giant tubeworms and clams. Indeed, it is perhaps because these animals acquire their symbionts horizontally and feed on them as they are housed in their cells (and not on their surface) that they were found to abundantly express genes involved in symbiont acquisition, proliferation control and digestion (Sun et al., 2017, Hinzke et al., 2019; Yuen et al., 2019). Notably, we did observe a partial overlap of the most expressed gene categories (e.g., oxidative stress, energy generation, immune response), when L. oneistus was compared to the marine gutless annelid Olavius algarvensis. We ascribe the overlap to the fact that, albeit endosymbiotic, O. algarvensis also inhabits shallow water sand (Figure S4, Supplemental material) and, as hypothesized for L. oneistus, it may also acquire its symbionts vertically (Woyke et al., 2006; Dubilier et al., 2008; Wippler et al., 2016; Zimmermann et al., 2016).

To conclude, although both symbiont- (Paredes et al., 2021) and host-transcriptomics do not suggest a high degree of inter-partner metabolic dependence in the L. oneistus ectosymbiosis, the nematode seems well-adapted to both anoxic sulfidic (AS) and hypoxic (H) sand (Figure 2, Data S1). The transcriptional response of the worm to these two conditions is, however, significant (Figure 2, Data S1), and it will be reported next.

Genes upregulated in anoxic sulfidic (AS) nematodes

Chaperones and detoxification. The expression of chaperone-encoding (e.g., hsp12.2, grpE, dnaJ/dnj-2, pfd-1, pfd-6; Naylor et al., 1996; Lundin et al., 2008; Bar-Lavan et al., 2016), and ROS-detoxifying-related genes (e.g., superoxide dismutase sod-2 and a putative glutathione peroxidase, involved in the detoxification of superoxide dismutase and hydrogen peroxide, respectively; Suzuki et al., 1996; Margis et al., 2008) were higher in AS nematodes (Figures 2 and 3). Notably, transcripts encoding for the heme-binding cytochrome P450 cyp-13B1 were also more abundant in AS (Figure 3), perhaps to increase the worm’s capacity to cope with putative ROS formation (Oliveira et al., 2009). Indeed, as cells start being oxygen-depleted, mitochondrial ROS accumulate because of the inefficient transfer of electrons to molecular oxygen (Semenza, 1999; Nystul and Roth, 2004; Selivanov et al., 2009; Kim and Jin, 2015). Alternatively, the upregulation of antioxidant-related genes in AS worms could represent an anticipation response to an imminent reoxygenation. In animals alternating between anoxic and oxygenated habitats, the re-exposure to oxygen can be very dangerous, as it creates a sudden ROS overproduction that may overwhelm the organism’s oxidative defense mechanisms (Hermes-Lima and Zenteno-Savin, 2002; Hashimoto et al., 2004). Although it has not been reported for nematodes, overexpression of ROS-counteracting genes is consistent with what has been reported for vertebrates and marine gastropods which, just like L. oneistus, alternate between oxygen-depletion and reoxygenation (Hermes-Lima and Zenteno-Savin, 2002).

Mitochondrial and cytoplasmic ribosome biogenesis. In the cellular stress imposed by oxygen deprivation, mitochondria are central to both death and survival (Borutaite et al., 1995; Brookes et al., 2004; Brenner et al., 2012; Hawrysh et al., 2013; Galli et al., 2014). In this scenario, calcium regulation, the scavenging of ROS or the suppression of their production, and/or inhibition of the mitochondrial permeability transition pore (MPTP) opening, might help to preserve mitochondrial function and integrity (Horwitz et al., 1994; Murphy et al., 2008; Galli et al., 2014; Fanter et al., 2020). In addition, removal of specific mitochondrial components (mitochondrial-associated protein degradation, MAD), might also arise to maintain the overall mitochondrial homeostasis (Chatenay-Lapointe and Shadel, 2010; Heo et al., 2010). Perhaps as a response to anoxia-induced stress (reviewed in Galli et al., 2014), a gene involved in MAD (vms-1) (Chatenay-Lapointe and Shadel, 2010; Heo et al., 2010), was upregulated in AS worms (Figure 4). More abundant in this condition were also transcripts encoding for mitochondrial transmembrane transporters tin-44, slc-25A26 and C16C10.1 (UniProtKB O02161, Q18934, Q09461), putatively transporting, peptide-containing proteins from the inner membrane into the mitochondrial matrix, such as S-Adenosyl Methionine (Figure 6). Surprisingly, although the translation elongation factor eef-1A.2 (Tullet, 2015) was downregulated in AS worms, not only various mitochondrial ribosome structural components (28S: mrps, 39S: mrpl; Kaushal et al., 2014), and mitochondrial translation-related genes (e.g., C24D10.6 and W03F8.3; Sharika et al., 2018) were upregulated in AS nematodes, but also several cytoplasmic ribosome biogenesis (40S: rps, 60S: rpl; Melnikov et al., 2012) and subunit assembly genes (e.g., RRP7A−like, You et al., 2015) (Figure 4).

Taken together, the maintenance of mitochondrial homeostasis, an anticipatory response to a potential upcoming ROS insult (see Chaperones and detoxification section) and/or their involvement in extra-ribosomal functions (Chen et al., 2010; Savada et al., 2014; Xu et al., 2016) might explain the upregulation of ribosomal biogenesis-related genes in AS nematodes. Although upregulation of ribosomal proteins has also been observed in anoxic gastropods (Larade et al., 2001), increased ribosomal biogenesis (which oftentimes directly correlates with an increase of protein synthesis) is not expected in animals that must repress their metabolism to cope with oxygen deprivation (Thomas et al., 2000; Hochachka and Lutz 2001; Shukla et al., 2012).

Energy generation. Equally surprising was the upregulation of all differentially expressed genes related to energy generation in AS nematodes (Figure 4). Namely, besides putative oxygen-binding globulin-like genes (e.g., glb-1, glb-14, Geuens et al., 2010), the following were upregulated in AS nematodes: key structural genes (e.g., atp-3, atp-5, Xu et al., 2018), assembly-related genes (H+-transport ATP synthase, Maglioni et al., 2016) of the mitochondrial ATP synthase (complex V), genes related to complex I (lpd-5, nuo-2, McKay et al., 2003; Rea et al., 2007), a subunit of the succinate dehydrogenase involved in complexII (mev-1, Hartman et al., 2001), a mitochondrial cytochrome C oxidase subunit II assembly gene related to complex IV (sco-1, Williams et al., 2005), and a mitochondrial gene (coq-5), involved in the synthesis of either ubiquinone (Q, aerobic) or rhodoquinone (RQ, anaerobic) electron carriers (Buceta et al., 2019) (Figure 4). This suggests that, under anoxia, the electron transfer chain (ETC) is rewired in such way that electrons still enter the ETC at complex I, but instead of reaching complex III and IV they are transferred to RQ. This, in turn, shuttles the electrons to succinate dehydrogenase. The latter enzyme uses fumarate as an alternative electron acceptor, reducing it to succinate. This mechanism would maintain the flow of electrons through the ETC, and, it would prevent mitochondrial ATP generation (complex V) from shutting down (Buceta et al., 2019; Del Borrello et al., 2019).

In short, under AS, similarly to what has been observed in other free-living and parasitic nematodes, complex I appears to be the sole proton pump in this truncated form of ETC (Buceta et al., 2019; Del Borrello et al., 2019). In accordance with this hypothesis, tryptophan (Trp) degradation-related genes (acsd-1, acsd-2) and the Trp RNA ligase (wars-1; Tsai et al., 2017) that might be required to synthesize RQ (Buceta et al., 2019; Del Borrello et al., 2017; Tan et al., 2020) were upregulated under AS. Intriguingly, upregulated was also an isocitrate dehydrogenase gene (idh-1). This produces reducing equivalent (NADPH) carrying electrons that may fuel complex I (Smolková et al., 2012; Martínez-Reyes et al., 2020), but it might also add to the stimulation of the antioxidant capacity or to the maintenance of redox homeostasis by regenerating reduced glutathione (Hermes-Lima and Zenteno-Savin, 2002; Penkov et al., 2015; Yang et al., 2019).

If glycolysis is a key process for ATP generation in anoxia (Lutz et al., 1997; Semenza et al., 2001; Hochachka et al., 2001; Huang et al., 2008; Larade et al., 2009) and if, consistently, hxk-2 was upregulated under this condition (Figure 6), based on the expression levels of transcripts encoding for alpha-amylases (see Carbohydrate metabolism in Figure 6), starch and/or glycogen (Jackson and McLaughlin, 2009) may be the prominent carbon sources under anoxic sulfidic conditions.

Ubiquitin-proteasome system and proteases. Proteolysis supplies amino acids or polypeptides to the cells, while impeding the accumulation of damaged or misfolded proteins. The two main mechanisms of cellular proteolysis are the lysosome-mediated intracellular protein degradation (autophagy) and the proteasome-mediated proteindegradation (ubiquitin-proteasome system, UPS). In the latter, ubiquitin-protein ligases covalently attach ubiquitin to proteins, allowing their recognition and further degradation by the proteasome (Lodish et al., 2008; Papaevgeniou and Chondrogianni, 2014).

As shown in Figure 1, transcripts encoding for polyubiquitin (ubq-1), had the highest median gene expression across all transcriptomes. However, all ubiquitination-related genes detected in the differential gene expression analysis between the AS and H conditions, were upregulated in AS worms (Figure 2 and 3, Data S1). For example, aos-1, encoding for a subunit of the ubiquitin-activating enzyme (E1) (Jones et al., 2001), two ubiquitin-protein ligases (E3s without detected cullin domains; Papaevgeniou and Chondrogianni, 2014), and kelch-like genes (e.g., kel-8-like and kel-20). The former are BTB-domain containing proteins known to interact with E3 enzymes, with kel-8 being involved in the degradation of glutamate neuroreceptors (Schaefer and Rongo 2006; Stogios et al., 2005; Kim et al., 2018). Additional ubiquitination-related genes upregulated in AS were csn-2, encoding for a component of the COP9 signalosome complex (Pintard et al., 2003; Brockway et al., 2014), and proteasome genes (pas-2 and pas-3; Fraser et al., 2000; Blumenthal et al., 2002).

Among the proteases that were upregulated in AS worms, aspartyl proteases have been involved in neurodegeneration (Syntichaki et al., 2002), whereas plasminogen and the zinc matrix metalloproteinase ZMP-2 were both reported to mediate degradation of extracellular matrix (ECM) (Vassalli et al., 1991; Altincicek et al., 2010; Fischer, et al., 2014) (Figure 3). C. elegans ZMP-2 was also shown to prevent the accumulation of oxidized lipoproteins (Fischer et al., 2014), and, therefore it may contribute to the enhanced antioxidant response observed in this condition.

Autophagy and amino acid degradation. Besides acting coordinately to withstand stress, autophagy cooperates with apoptotic UPS for the recovery and supply of nutrients when these are scarce (Vabulas et al., 2005; Scott et al., 2004; Huber and Teis, 2016; reviewed in Wang RC et al., 2010 and Russel et al., 2014). Transcripts of two autophagy-related genes, bec-1 (Liang et al., 1999) and the Ragulator complex protein LAMTOR4 (C7orf59-like) (Bar-Peled et al., 2012) were more abundant in AS nematodes (Figure 3). While the former positively regulates autophagy (Liang et al., 1999; Meléndez et al., 2003), the latter interacts with the mTOR Complex I (mTORC1), and tethers small GTPases (Rags and Rheb) to the lysosomal surface (Bar-Peled et al., 2012). When amino acid levels are low, mTORC1 is not translocated to the lysosomal surface (Wang et al., 2009; Bar-Peled et al., 2012), thereby favoring catabolic processes such as autophagy (Thompson et al., 2005). We propose that amino acid scarcity might result from the upregulation of genes involved in the degradation of lysin, glycin, tyrosin, cystein, leucin, isoleucin, valin or tryptophan (Figure 3, Data S1). This would decrease mTORC1 activity and, in turn, stimulates nutrient recycling via autophagy in AS worms.

Conversely, we hypothesize that in H worms, active mTORC1 interacts with the ribosomal protein S6 kinase (S6K), encoded by the rsks-1 gene which is also up in H worms (Ladevaia et al., 2014) (Figure 3). This direct interaction, upon a cascade of phosphorylation events, would stimulate translation, and ultimately cell growth and proliferation (Ma et al., 2009, Howell et al., 2011, and Ladevaia et al., 2014).

All in all, although it is currently unclear whether increased autophagy is beneficial or detrimental, under AS conditions, the upregulation of genes involved in self-digestion might play a protective role and foster recovery from starvation (Thompson et al., 2005), pathogens (Huber and Teis, 2016) or from neuronal and muscular degeneration induced by oxygen deprivation (Murphy and Steenbergen 2008).

Lectins and mucins. Given that symbiont attachment may be mediated by Ca²⁺-dependent lectins (Nussbaumer et al. 2004, Bulgheresi et al., 2006, 2011) and given that, under anoxia, the symbiont appeared to proliferate more (Paredes et al., 2021), we expected nematode lectins to be upregulated under this condition. Indeed, nine C-type lectin domain (CTLD)-containing proteins were upregulated in AS L. oneistus adults and only two (clec-78 and clec-78-like-2) were upregulated in the presence of oxygen (Figure 4). In addition to CTLD-containing proteins, mucins, a class of glycoproteins with more than 50% of its mass attributable to O-glycans, were also upregulated in AS nematodes. Considering that mucin glycans are used by vertebrate gut commensals for attachment, as well as a source of nutrients (Koropatkin et al., 2012), it is conceivable that their upregulation in anoxia (Figure 4), together with that of CTLD-containing proteins, would foster symbiont attachment.

We hypothesize that overexpression of two classes of putative symbiont-binding molecules, lectins and mucins, under conditions favoring symbiont proliferation (i.e., AS condition, Paredes et al., 2021) may mediate bacterial coat reinforcement.

Apoptosis. Mitochondria play an important role in apoptosis induction (Simon et al., 2000; Martínez-Reyes et al., 2020). Indeed, MPTP opening due to ROS (or the severe ATP decline imposed by the absence of oxygen) may cause cytochrome C release from mitochondria and this, in turn, triggers caspase activation (Martinou et al., 2000; Simon et al., 2000; Gogvadze et al., 2006; Galli et al., 2014). We observed that transcripts encoding for sco-1, a gene needed for the synthesis and assembly of mitochondrial cytochrome C (Williams et al., 2005) were more abundant in AS worms (Figure 4). Further, we observed upregulation of Caspase-3 (ced-3) which belongs to a family of cysteine proteases involved in apoptosis (Mangahas et al., 2005; Kaufmann et al., 2008) and which is activated upon mitochondrial cytochrome C release into the cytosol (Liu et al., 1996; Tafani et al., 2000; Kaufmann et al., 2008; Martínez-Reyes et al., 2020). Additional apoptosis-related genes that appeared to be upregulated in AS worms were: bec-1 (Figure 3), a gene that promotes autophagy and fine-tunes the Ced-3-mediated apoptosis (Liang et al., 1999; Takacs-Vellai et al., 2005); ttr-52, which mediates apoptotic cell recognition prior to engulfment (Wang, X. et al., 2010; Chen et al., 2013); a BAG family molecular chaperone regulator 1 (BAG1-regulator); a cell-death-related nuclease crn−2 (Parrish et al., 2003; Samejima et al., 2005) and phagolysosome forming arl-8 (Sasaki et al., 2013), and a tyrosine kinase Abl-1, (abl-1) that modulates apoptotic engulfment pathways (Hurwitz et al., 2009).

Lipid catabolism. Genes involved in lipid metabolism were similarly expressed between the AS and H conditions (Figure 2, Data S1).In accordance, lipidomes of nematodes incubated in the presence or absence of oxygen were not significantly different (Figure S5, Supplemental material). However, in line with the overall upregulation of degradation pathways, we observed upregulation of genes involved in FA beta-oxidation (kat-1; Berdichevsky et al., 2010), in lipid digestion (the lipase lipl-6; UniProtKB E2S7J2), and lipid degradation (a peripilin-2-like protein; Chughtai et al., 2015). Moreover, a gene that might be involved in oxidative-stress tolerance (a stearic acid desaturase fat-7 regulating the first step of the fatty acid desaturation pathway (Horikawa et al., 2009) was also upregulated in AS worms. Lipid degradation under anoxia might be a strategy to overcome starvation (Krivoruchko and Storey, 2015).

Notably, we also observed an upregulation of two genes involved in phosphatidylcholine (PC) synthesis (pmt-1, pmt-2, Brendza et al., 2007) (Figure 5). Intriguingly, PC was more abundant in the anoxic symbiont (Paredes et al., 2021), although the latter cannot synthetize it. Thus, their upregulation in AS worms suggests worm-symbiont lipid transfer.

GABA- and glutamate-mediated neurotransmission. Upregulated genes related to GABA synthesis were, unc-25, unc-104 and pdxk-1 (pyridoxal phosphate hexokinase) (Thomas et al., 1990; Mclntire et al., 1993; Jin et al., 1999; Gally et al., 2003; Nordquist et al., 2018; Risley et al., 2016) (Figure 5, Data S1). Consistent with an expected increase in glutamate requirement as a direct GABA precursor (Martin et al., 1993), we observed downregulation of two glutamine synthetases and a delta-1-pyrroline-5-carboxylate synthase (gln-3 and alh-13 respectively; van der Vos et al., 2012; Yen et al., 2021; Figure 6), known to convert glutamate to glutamine or to proline, respectively. Furthermore, an mgl-2 like gene encoding for a glutamate receptor, which is activated in the presence of glutamate (Tharmalingam et al., 2012), was up in AS worms. Note that, when oxygen is limited, glutamate may act as a neurotoxic amino acid (Baker et al., 1991; Lutz et al., 2003a). Therefore, increased GABA biosynthesis might, beneficially, prevent its accumulation (Milton et al., 2002; Mathews et al., 2003).

GABA-mediated neurotransmission has been documented for facultative anaerobic animals thriving in anoxic conditions (Lutz et al., 1997; Milton et al., 1998; Lutz et al., 2003a, b). Due to its inhibitory nature, it contributes to avoid membrane depolymerization (Nilsson et al., 1990; Milton et al., 1998). Moreover, given that it relaxes muscles, the increment of GABA may impact the movement of the animal (Mclntire et al., 1993; Schuske et al., 2004). Therefore, upregulation of GABA-mediated neuronal activity might explain why anoxic L. oneistus did not form tight worm clusters after 24h (Supplemental movie 3).

Dopamine-mediated neurotransmission. A gene encoding for the tyrosine hydroxylase Cat-2 (cat-2), which is needed for dopamine biosynthesis (Sawin et al., 2000) and two putative dopamine receptors (protein-D2-like and a G_PROTEIN_RECEP_F1_2 domain-containing protein (dop-5); Sanyal et al., 2004) were upregulated in AS worms. Moreover, a dat-1-like gene mediating dopamine reuptake into the presynaptic terminals was downregulated (Gainetdinov et al., 2002; McDonald et al., 2006) in AS worms (Figure 5).

Calcium-binding and -sensing proteins. Finally, in AS worms several calcium-binding or -sensing proteins (e.g., ncs-2, cex-2, and a calbindin-like (CALB1 homologue); Soontornniyomkij et al., 2012; Hobert et al., 2018; Figure 5), as well as calcium transporters (cca-1, Steger et al., 2005; Transport category, Figure 6) were upregulated. On the one hand, we hypothesize their involvement in the inhibitory neural signaling described above (for example, Ncs-2 mediates the cholinergic and GABAergic expression of C. elegans (Zhou et al., 2017). On the other, they may protect cells against the stress inflicted by anoxia, which involves calcium overload and consequent cellular acidification (Bickler et al., 1992; Dell'Anna et al., 1996; Galli et al., 2014).

Genes upregulated in hypoxic (H) nematodes

Innate immune pathways and effectors. Animals recognize and respond to microbes by means of immunoreceptors including Toll-like receptors, conserved from sponges to humans (Akira et al., 2006). We identified almost all genes belonging to this pathway, including the one encoding for the NF-kB transcription factor. This came as a surprise given that, up to now, the has not been identified in any other nematode NF-kB (Pujol and Ewbank, submitted). As surprising, was the fact that not only two Toll-like receptors (tol-1 and tol-1-like), but also genes encoding for antimicrobial proteins such as a peroxisome assembly factor involved in defense against Gram- (prx-11-like, Wang, D. (2019), a putatively antifungal endochitinase (Dravid et al., 2015) and Bactericidal Permeability Increasing proteins (BPIs) were also more abundant in H worms. BPIs may bind LPS and perforate Gram- membranes and have shown to play a symbiostatic role in other invertebrates (Bruno et al., 2019; Krasity et al., 2015; Chen et al., 2017). However, it is unclear whether activation of the L. oneistus Toll pathway leads to the nuclear NF-kB switching on the expression of antimicrobial genes or whether, as shown in C. elegans, the Toll pathway mediates behavioral avoidance of pathogens (Pradel et al., 2007; Brandt et al., 2015).

Overall, the apparent oxygen stimulation of a central innate immunity pathway and, directly or indirectly, of broad range anti-defense mechanisms could be adaptations to the fact that in oxygenated environments (when crawling in superficial sand layers), L. oneistus is exposed to predation from bigger animals, but also to pathogenic members of the bacterioplankton. Overexpression of broad-range antimicrobials in response to oxygen might therefore help L. oneistus to avoid colonization by potentially deleterious, fouling bacteria (e.g., Vibrios, Roseobacters and Pseudoaltermonas/Alteromonadales) when crawling close to the water column (Dang and Lovell, 2016; M. Mussmann, personal communication).

Development. Although development-related genes were some of the most expressed under all conditions (Figure 1), many were upregulated in H nematodes (Figure 2 and 5). Among the development-related genes upregulated in H nematodes were those related to molting (e.g., nas-36, nas-38, chs-2, ptr-5, ptr-18, apl-1, myrf-1;Suzuki et al., 2004; Zhang et al., 2005; Zugasti et al., 2005; Hornsten et al., 2007; Russel et al., 2011), germ line establishment (e.g., ccm-3, rsks-1;Pan et al., 2007;Pal et al., 2017), oogenesis/spermatogenesis (crt-1,Park et al., 2001), embryonic development and yolk production (smp-1, cpna-1, plt-1, vit-6, crt-1, arrd-17, mlc-5; Clark et al., 1997;Goedert et al., 1996; Gatewood et al., 1997; Fuji et al., 2002; Gally et al., 2009; Zahreddine et al., 2010; Jee et al., 2012; Warner et al., 2013; Fisher et al., 2014; Perez and Lehner, 2019), and/or larval development (nmy-1, ifb-1; Ding et al., 2004;Osório et al., 2019), as well as male tip (Cdt1, plx-1, ver-3, ; Nelson et al., 2011; Dalpé et al., 2004; Dalpe et al., 2013), vulva morphogenesis (hda-1, unc-62), and a hermaphrodite-related gene (hda-1; Dufourcq et al., 2002; Choy et al., 2007) (Figure 5). Morever, transcripts encoding for a number of proteases shown to be involved in C. elegans molting (e.g., nas-38, nas-6-like; Park et al., 2010), development (e.g., teneurin-a-like; Topf and Drabikoswki, 2019), neuronal regrowth or locomotion (tep-1; Kim et al., 2018) and pharingeal pumping (e.g., neprilysin nep-1; Spanier et al., 2005) were also more abundant in H worms. Remarkably, vav-1, which, besides being involved in male tip and vulva morphogenesis (Nelson et al., 2011), may also regulate the concentration of intracellular calcium (Norman et al., 2005), was one of the few development-related genes to be downregulated in H nematodes (see previous section on Ca-binding proteins).

To sum up, and as expected, the host appears to exploit oxygen availability to undertake energetically costly processes, such as development and molting (De Cuyper and Vanfleteren 1982; Uppaluri and Brangwynne 2015).

Carbohydrate metabolism. If in AS nematodes, glycogen or starch appeared prominent carbon sources, H worms seemed to exploit trehalose and cellulose instead. Indeed, genes that degrade trehalose (tre-1, Pellerone et al., 2003) and cellulose (Ppa-cel-2, Schuster et al., 2012) were upregulated in H worms, as well as a putative ADP-dependent glucokinase (C50D2.7) involved in glycolysis (Yuan et al., 2012). The use of this pathway was supported by the overexpression of four genes encoding for sugar transporters (Slc2-A1, C35A11, K08F9.1, F53H8.3; Kitaoka et al., 2013; Bertoli et al., 2015), perhaps switched on by active mTOR (see above) (Figure 6) (Howell et al., 2011).

Additionally, L. oneistus appeared to exploit oxygen to synthesize complex polysaccharides, such as heparan sulfate (hst-1-like; Miyagawa et al. 1988; Bhattacharya et al., 2009) and glycan (Gcnt3-like) (Figure 6), as an ortholog of the N-deactetylase/N-sulfotransferase hst-1, related to heparin biosynthesis was also upregulated (Bhattacharya et al., 2009).

Although glycolysis seems to generate ATP in both AS and H worms, it is not clear why the latter would prefer to respire cellulose or trehalose instead of starch. Given its role as a membrane stabilizer, we speculate that AS worms might prioritize the storage of trehalose over its degradation to preserve membrane integrity (Figure 6) (Crowe et al 1987; Carpenter et al., 1988; Clegg et al., 1997; Chen et al., 2002; Haddad 2006). Of note, based on its genome draft, the symbiont may synthetize and transport trehalose, but it may not use it (Paredes et al., 2021). Therefore, we hypothesize symbiont-to-host transfer of trehalose under hypoxia. Consistently, the symbiont’s trehalose synthesis-related gene (otsB; Paredes et al., 2021), and the host trehalase (tre-1;Figure 6) were both upregulated under hypoxia and metabolomics could detect trehalose in both partners (Table S1). Metabolomics also detected sucrose in both the holobiont and the symbiont fraction (Table S1). Given that, based on transcriptomics and proteomics, the nematode can utilize sucrose but cannot synthesize it (Data S1), whereas the symbiont can (Paredes et al., 2021), as in the case for trehalose, we hypothesize symbiont-to-host sucrose transfer.

Acetylcholine-mediated neurotransmission. Instead of upregulating genes involved in inhibitory (GABA and dopamine-mediated) neurotransmission, hypoxicworms appeared to use excitatory acetylcholine-mediated neurotransmission as indicated by the upregulation of molo-1, acr-20, cup-4, lev-9, and sphingosine kinase sphk-1 that promotes its release(Mongan et al., 2002; Patton et al., 2005; Gendrel et al., 2009; Boulin et al., 2012; Chan et al., 2012) (Figure 5). On the one hand, acetylcholine-mediated neurotransmission might promote ROS detoxification in H worms (Sun et al., 2014). On the other hand, its downregulation in AS worms may beneficially decrease calcium influx (Hochachka and Lutz, 2001).

Feeding, mating, mechanosensory behavior and axon guidance and fasciculation. Transcripts related to the neuronal regulation of energy-demanding activities such as feeding, mating, motion, as well as nervous system development were more abundant in H nematodes (Figure 5, and Data S1). More precisely, upregulated genes were involved in pharyngeal pumping (nep-1, lat-2;Spanier et al., 2005; Guest et al., 2007), male mating behavior and touch (pdfr-1, tbb-4, ebax-1, Hurd et al., 2010; Wang, Z. et al., 2013), axon guidance and fasciculation (spon-1, igcm-1, ebax-1, tep-1; Kim et al., 2018; Woo et al., 2008; Schwarz et al., 2009;Wang, Z et al., 2013), mechanosensory behavior (e.g., mec-12, delm-2; Gu et al., 1996; Han et al., 2013). Additionally, we also observed the upregulation of a gene encoding for a glutamate receptor (glr-7) possibly involved in feeding facilitation (Li et al., 2012).

Amino acid biosynthesis. Transcripts of genes involved in the synthesis of glutamine and proline (gln-3 and alh-13, respectively),aspartate (L-asparaginases; Tsuji et al., 1999) and S-adenosyl-L-methionine (SAM) (sams-4; Chen et al., 2020) were all upregulated in H worms (Figure 6), as well as one encoding for the ornithine decarboxylase odc-1 which is involved in biosynthesis of the polyamine putrescin, and is essential for cell proliferation and tissue growth (Russell et al., 1968; Heby, 1981). Moreover, polyamines, with their high charge-to-mass ratio may protect against superoxide radicals, which, as mentioned, harm cell membranes and organelles, oxidize proteins, and damage DNA (Gilad et al., 1991; Longo et al., 1993).

Lipid biosynthesis. Genes upregulated in H worms mediate the biosynthesis of long chain fatty acids (acs-3, acs-14, elo-3 but not acs-5;Yuan et al., 2012; Ward et al., 2014; Wang et al., 2021), sphingolipids (a sphingosine kinase-1 (sphk-1) and egl-8, which controls egg laying and pharyngeal pumping in C. elegans (Bastiani et al., 2003). Notably, sphingolipids may be anti-apoptotic (Taha et al., 2006) or result in acetylcholine release (Chan et al., 2012).

On the other hand, ceramides, which have antiproliferative properties and who may mediate resistance to severe oxygen deprivation (Deng et al., 2008; Menuz, et al. 2009), appeared to be mainly synthesized in AS worms, as indicated by the upregulation of genes involved in ceramide biosynthesis (asm-3, ttm-5; Watts et al., 2017) (Figure 6).

Transport. As anticipated in the introduction, anoxia-tolerant animals switch off ATP-demanding processes such as ion pumping (Lutz et al., 1996; Galli et al., 2014). Indeed, transcripts encoding for proteins involved in cation channel activity (gtl-2, voltage gated H channel 1; Teramoto et al., 2010), sodium transport (delm-2-like; Han et al., 2013), chloride transport (anoh-1, best-13, best-14; Tsunenari et al., 2013; Wang, Y. et al., 2013; Goh et al., 2018), ABC transport (wht-2, pgp-2, slcr-46.3, F23F12.3, hmit-1.3; Currie et al., 2007; Schroeder et al., 2007; Kage-Nakadai et al., 2011) and organic transport (F47E1.2, oct-2; Pao et al., 1998) were all more abundant in H than AS worms (Figure 6).

Sulfur metabolism. The mpst-7 gene which is involved in organismal response to selenium and it is switched on in hypoxic C. elegans (Romanelli-Credrez et al., 2020) was upregulated in H nematodes (Figure 6). Given that the latter is thought to catalyze the conversion of sulfite and glutathione persulfide (GSSH) to thiosulfate and glutathione (GSH) (Filipovic et al., 2018), hypoxia-experiencing L. oneistus might express this enzyme to recharge the cells with GSH and hence, help to cope with oxidative stress (Hayes and McLellan, 1999; Mytilineou et al., 2002; Diaz-Vivancos et al., 2015). Also more abundant in H worms were transcripts encoding for the sulfatases 2 (sul-2) (Morimoto-Tomita et al., 2002) and a PAPS-producing pps-1 (3′-phospho-adenosine-5′-phosphosulfate (PAPS) considered the universal sulfur donor; Bhattacharya et al., 2009), as well as for the chaperones pdi-6 and protein-disulfide-isomerase-A5-like which require oxygen to mediate correct disulfide bond formation in protein folding (Teodoro and O’Farrell, 2003; Rose et al., 2017; Livshits et al., 2017) (Figure 6).

Conversely, a putative sulfide-producing enzyme (mpst-1) who protects C. elegans from mitochondrial damage (Qabazard et al., 2014; Ng et al., 2019; Kimura, 2020) was upregulated in AS nematodes. Notably, under AS, L. oneistus might detoxify sulfide by producing glutathione and taurine (Rose et al., 2017), as a persulfide dioxygenase (ethe-1) and a cysteine dioxygenase (cdo-1) which catalyzes taurine synthesis via cysteine degradation were upregulated. Sulfide detoxification via taurine accumulation is a common strategy in chemosynthetic animals (reviewed in Cavanaugh et al., 2006).

All in all, L. oneistus appeared to limit excess accumulation of free sulfide in anoxia and to free sulfate when oxygen was available.

Overall and irrespectively of the conditions it was subjected to, L. oneistus mostly expressed genes involved in degradation, energy generation, stress response and immune defense. Astonishingly, L. oneistus did not enter suspended animation when subjected to anoxic sulfidic conditions for days. We hypothesize that in the absence of oxygen, ATP production is supported by trehalose and cellulose catabolism, and by rewiring the ETC in such way as to use rhodoquinone (RQ) as electron carrier, and fumarate as electron acceptor. Moreover, the nematode activates several degradation pathways (e.g., ubiquitin-proteasome system (UPS), autophagy, and apoptosis) to gain nutrients from anoxia- or ROS-damaged proteins and mitochondria. Further, AS worms also upregulated genes encoding for ribosomal proteins and putative symbiont-binding proteins (lectins). Finally, as proposed for other anoxic-tolerant animals, the worm seems to upregulate its antioxidant capacity in anticipation of reoxygenation. When in hypoxic conditions (Figure 7, left), instead, we speculate that the worm uses starch for energy generation to engage in costly developmental processes such as molting, feeding, and mating, likely relying on excitatory neurotransmitters (e.g., acetylcholine), and it upregulates the Toll immune pathway and, directly or indirectly, the synthesis of broad range antimicrobials (e.g., fungicides, bactericidal permeability increasing proteins).

When looking at the Laxus-Thiosymbion symbiosis in light of what was recently published (Paredes et al., 2021), we could identify two signs of inter-partner metabolic dependence: in anoxia worms might transfer lipids to their symbionts, and in hypoxia the symbionts might transfer trehalose to their hosts.

Furthermore, we may conclude that, wherever in the sand the consortium is, one of the two partners is bound to be stressed: in anoxia, the symbiont appear to proliferate more, while its animal host engages in degradation of damaged proteins and mitochondria and in detoxification. In the presence of oxygen, the situation is inverted: the symbiont seems massively stressed, while the host can afford energy costly biosynthetic processes to develop and reproduce (Figure 7). It is therefore fascinating that, in spite of the dramatically different needs a bacterium and animal must have, the Laxus-Thiosymbion symbiosis evolved.

ACKNOWLEDGEMENTS

This work was supported by the Austrian Science Fund (FWF) grant P28743 (T.V., S.B., and L.K.), the FWF DK plus grant W1257: Microbial Nitrogen Cycling (G.F.P., L.K.), the FWF DOC 69 doc.fund (T.V). We thank Yin Chen for providing the facilities for lipidomics analysis, and Marvin Weinhold’s, Jana Matulla’s and Sebastian Grund’s excellent technical work during metabolite analysis, and protein sample preparation and MS analysis, respectively. We are grateful to the Carrie Bow Cay Marine Field Station, Caribbean Coral Reef Ecosystem Program, and Station Manager Zach Foltz and Scott Taylor for their continuous support during field work. We thank Nicole Dubilier for access to data on Olavius algarvensis, and Jonathan Ewbank and Marc Mussmann for insightful comments on the manuscript. Finally, we were inspired by insightful discussions with Monika Bright and Jörg A. Ott. This is contribution number XXX of the Carrie Bow Cay Marine Field Station, Caribbean Coral Reef Ecosystem Program.

Data availability. This Transcriptome Shotgun Assembly project has been deposited at DDBJ/EMBL/GenBank under the accession GJNO00000000. The version described in this paper is the first version, GJNO01000000. RNA-Seq data are available at the Gene Expression Omnibus (GEO) database and are accessible through accession number XXX.

Sample collection

Laxus oneistus individuals were collected on multiple field trips (2016-2019) at approximately 1 m depth from sand bars off the Smithsonian Field Station, Carrie Bow Cay in Belize (16°48′11.01″N, 88°4′54.42″W). The collection of the nematodes, the incubations set up for RNA sequencing, lipidomics, proteomics and metabolomics, as well as the RNA extraction, and library preparation are described in Paredes et al., 2021. Importantly, the nematodes had a bright white appearance and replicate incubations were started simultaneously. Note that the Supplemental material describes changes in the lipidomics and proteomics pipelines, as well as the metabolomics, and sequencing data of Olavius algarvensis.

Host transcriptome de novo assembly

In preparation for the assembly, reads from each sample were first mapped to the symbiont as described before (Paredes et al., 2021), and remaining rRNA reads from all domains of life were removed from unmapped reads using sortmerna v2.1 in combination with the SSURef_NR99_119_SILVA_14_07_14 and LSURef_119_SILVA_15_07_14 databases. Further, exact duplicate reads were removed using PRINSEQ lite’s derep option. Read files free of symbiont reads, rRNA reads and exact duplicates were used as input for transcriptome sub-assemblies via Trinity v2.6.6 with the strand-specific option (--SS_lib_type F) (Grabherr et al., 2011). Two sub-assemblies differing in the number and type of input read files were performed: (1) 9 input read files including biological triplicates from 3 incubation conditions (O, H, A) and (2) 4 input read files including a single replicate from 4 incubation conditions (O, H, A and hyper-O). Hyper-O refers to an incubation in which air was pumped directly into the exetainers for the entire incubation period to supersaturate the seawater (300 %O₂). However, as this incubation condition yielded an incongruous transcriptional response by the symbiont (data not shown), these read data were only used to extend the host transcriptome’s coding repertoire. The qualities of both sub-assemblies were assessed as described below.

We then performed an intra-assembly clustering step as described in (Cerveau and Jackson, 2016), during which identical transcripts were removed from the sub-assemblies using CD-HIT-EST (Fu et al., 2012). To further reduce redundant transcripts, only the longest isoform for each ‘gene’ identified by Trinity was kept using Trinity’s get_longest_isoform_seq_per_trinity_gene.pl utility. The remaining transcripts of each sub-assembly were then concatenated to produce a merged transcriptome assembly. The final assembly was created by applying another sequence clustering using CD-HIT-EST to avoid inter-assembly redundancy. Here, the identity parameter of 80% (-c 0.8) combined with a minimal coverage ratio of the shorter sequence of 80% (-aS 0.8) and minimal coverage ratio of the longest sequence of 0.005% (-aL 0.005) yielded the best-performing assembly in terms of number of transcripts (162,455) and contiguity (N50 value of 770) (data not shown).

Assembly completeness was assessed by estimating completeness via BUSCO nematode single-copy orthologs (Simão et al., 2015). Importantly, the merged assembly yielded a higher BUSCO-based completeness compared with the two sub-assemblies; 79.2% of the BUSCO nematode single-copy orthologs were found to be present and complete in the final assembly (636 single-copy/142 duplicated), whereas assembly (1) scored 77.8% (233 single-copy/531 duplicated) and assembly (2) was 76.2% complete (314 single-copy/434 duplicated). Further, assembled transcripts were filtered based on taxonomic classification. Transcripts were matched against the RefSeq protein database using blastx (E value 1E-3), and the output was then used as input for taxonomic assignment via MEGAN v5 (Huson et al., 2007). Only transcripts classified as belonging to ‘Eukarya’ were kept (MEGAN parameters: Min Score: 50, Max Expected: 1E-2, Top Percent: 2), which reduced the number of putative L. oneistus transcripts to 30,562. Assembled transcripts were also functionally annotated using Trinotate (Bryant et al., 2017). Briefly, predicted protein coding regions were extracted using TransDecoder (https://github.com/TransDecoder), both transcripts and predicted protein sequences were searched for protein homology via blastx and blastp, respectively, and predicted protein sequences were annotated for protein domains (hmmscan), signal peptides (signalP) and transmembrane domains (THMMM). 85,859 transcripts exhibited at least one functional annotation. Finally, only taxonomy-filtered transcripts with at least one functional annotation were kept, thereby further reducing the number of putative host transcripts to 27,984, with 22,072 thereof predicted to contain protein coding regions. BUSCO-based completeness for this filtered host transcriptome assembly was 78.8% (635 single-copy/139 duplicated).

Gene expression analysis

Raw sequencing reads quality assessment and preprocessing of data was followed as described in Paredes et al., 2021. Trimmed reads were mapped to the de novo transcriptome assembly and transcript abundance was estimated using RSEM v1.3.1 (Li and Dewey 2011) in combination with bowtie with default settings except for the application of strandedness (--strandedness forward). Read counts per transcript were used for differential expression analysis, and TPM (transcripts per kilobase million) values were transformed to log2TPMs as described in Paredes et al. 2021.

Gene and differential expression analyses were conducted using the R software environment and the Bioconductor package edgeR v3.28.1 (Gentleman et al., 2004; Robinson et al., 2010; R core Team, 2013), and as shown in Paredes et al., 2021. Here, we only describe the modifications that were made to the pipeline. Genes were considered expressed if at least ten reads in at least three replicates of one of the four conditions could be assigned. Excluding the replicates of the oxic condition, we found that 74.9% of all predicted nematode protein-encoding genes to be expressed (16,526 genes out of 22,072). Log₂TPM were used to assess sample similarities via multidimensional scaling based on Euclidean distances (R Stats package) (R core Team, 2013) (Figure S1B), and the average of replicate log₂TPM values per expressed gene and condition was used to estimate expression strength. Median gene expression of entire metabolic processes and pathways per condition was determined from average log₂TPM values.

Expression of genes was considered significantly different if their expression changed 1.5-fold between two treatments with a false-discovery rate (FDR) ≤ 0.05 (Rapaport et al., 2013). Throughout the paper, all genes meeting these thresholds are either termed differentially expressed or up- or downregulated. For the differential expression analyses between the AS, H and A conditions see Data S1. Heatmaps show mean-centered log₂TPM expression values to highlight gene expression change.

All predicted L. oneistus proteins were automatically annotated using eggNOG-mapper v2 (Cantalapiedra et al., 2021) against eggNOG 5.0 (Huerta-Cepas et al., 2019) using diamond v2.0.4 (Buchfink et al., 2021). All genes that are shown and involved in a particular process were manually curated by blasting them against both the NCBI BLASTP nr database (Altschul et al., 1990) and the WormBase (Harris et al., 2020; https://wormbase.org/tools/blast_blat).

Hermes-Lima, M., & Zenteno-Savın, T. (2002). Animal response to drastic changes in oxygen availability and physiological oxidative stress. Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology, 133(4), 537-556.
Hochachka, P. W., Buck, L. T., Doll, C. J., & Land, S. C. (1996). Unifying theory of hypoxia tolerance: molecular/metabolic defense and rescue mechanisms for surviving oxygen lack. Proceedings of the National Academy of Sciences, 93(18), 9493-9498.
Hochachka, P. W., Land, S. C., & Buck, L. T. (1997). Oxygen sensing and signal transduction in metabolic defense against hypoxia: lessons from vertebrate facultative anaerobes. Comparative Biochemistry and Physiology Part A: Physiology, 118(1), 23-29.
Hochachka, P. W., & Lutz, P. L. (2001). Mechanism, origin, and evolution of anoxia tolerance in animals☆. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 130(4), 435-459.
Clegg, J. (1997). Embryos of Artemia franciscana survive four years of continuous anoxia: the case for complete metabolic rate depression. The Journal of Experimental Biology, 200(3), 467-475.
Nystul, T. G., Goldmark, J. P., Padilla, P. A., & Roth, M. B. (2003). Suspended animation in C. elegans requires the spindle checkpoint. Science, 302(5647), 1038-1041.
Teodoro, R. O., and O'Farrell, P. H. (2003). Nitric oxide‐induced suspended animation promotes survival during hypoxia. The EMBO Journal, 22(3), 580-587.
Haddad, G. G. (2006). Tolerance to low O₂: lessons from invertebrate genetic models. Experimental physiology, 91(2), 277-282.
Liu, L., & Simon, M. C. (2004). Regulation of transcription and translation by hypoxia. Cancer biology & therapy, 3(6), 492-497.
Liu, L., Cash, T. P., Jones, R. G., Keith, B., Thompson, C. B., & Simon, M. C. (2006). Hypoxia-induced energy stress regulates mRNA translation and cell growth. Molecular cell, 21(4), 521-531.
Galli, G. L., & Richards, J. G. (2014). Mitochondria from anoxia-tolerant animals reveal common strategies to survive without oxygen. Journal of Comparative Physiology B, 184(3), 285-302.
Van Voorhies, W. A., & Ward, S. A. M. U. E. L. (2000). Broad oxygen tolerance in the nematode Caenorhabditis elegans. Journal of Experimental Biology, 203(16), 2467-2478.
Padilla, P. A., Nystul, T. G., Zager, R. A., Johnson, A. C., & Roth, M. B. (2002). Dephosphorylation of cell cycle–regulated proteins correlates with anoxia-induced suspended animation in Caenorhabditis elegans. Molecular biology of the cell, 13(5), 1473-1483.
Nystul, T. G., & Roth, M. B. (2004). Carbon monoxide-induced suspended animation protects against hypoxic damage in Caenorhabditis elegans. Proceedings of the National Academy of Sciences, 101(24), 9133-9136.
Powell-Coffman, J. A. (2010). Hypoxia signaling and resistance in C. elegans. Trends in Endocrinology & Metabolism, 21(7), 435-440.
Fawcett, E. M., Hoyt, J. M., Johnson, J. K., & Miller, D. L. (2015). Hypoxia disrupts proteostasis in Caenorhabditis elegans. Aging Cell, 14(1), 92-101.
Kitazume, H., Dayi, M., Tanaka, R., & Kikuchi, T. (2018). Assessment of the behaviour and survival of nematodes under low oxygen concentrations. PloS one, 13(5), e0197122.
Kim, K. W., & Jin, Y. (2015). Neuronal responses to stress and injury in C. elegans. FEBS letters, 589(14), 1644-1652.
Denny, M. (1993). Air and Water: the Biology and Physics of Life’s Media. Princeton, NJ: Princeton University Press. 341pp.
Ott, J. A., & Novak, R. (1989). Living at an interface: Meiofauna at the oxygen/sulfide boundary of marine sediments.
Ott, J. A., Novak, R., Schiemer, F., . Hentschel, U., Nebelsick, M., & Polz, M. (1991). Tackling the sulfide gradient: a novel strategy involving marine nematodes and chemoautotrophic ectosymbionts. Marine Ecology, 12(3), 261-279.
Schiemer, F., Novak, R., & Ott, J. (1990). Metabolic studies on thiobiotic free-living nematodes and their symbiotic microorganisms. Marine Biology, 106(1), 129-137.
Paredes, G. F., Viehboeck, T., Lee, R., Palatinszky, M., Mausz, M. A., Reipert, S., ... & König, L. (2021). Anaerobic sulfur oxidation underlies adaptation of a chemosynthetic symbiont to oxic-anoxic interfaces. mSystems, 6(3), e01186-20.
Ott, J. A., Bauer-Nebelsick, M., & Novotny, V. (1995). The genus Laxus Cobb, 1984 (Stilbonematinae: Nematoda): description of two new species with ectosymbiotic chemoautotrophic bacteria. Proceedings of the Biological Society of Washington, 108(3), 508-527.
Chan, K., Goldmark, J. P., & Roth, M. B. (2010). Suspended animation extends survival limits of Caenorhabditis elegans and Saccharomyces cerevisiae at low temperature. Molecular biology of the cell, 21(13), 2161-2171.
UniProt: the universal protein knowledgebase in 2021. Nucleic Acids Research, 2021, 49. Jg., Nr. D1, S. D480-D489.
Stringham, E. G., Jones, D., & Candido, E. P. M. (1992). Expression of the polyubiquitin-encoding gene (ubq-1) in transgenic Caenorhabditis elegans. Gene, 113(2), 165-173.
Geuens, E., Hoogewijs, D., Nardini, M., Vinck, E., Pesce, A., Kiger, L., ... & Dewilde, S. (2010). Globin-like proteins in Caenorhabditis elegans: in vivo localization, ligand binding and structural properties. BMC biochemistry, 11(1), 1-15.
Birnby, D. A., Link, E. M., Vowels, J. J., Tian, H., Colacurcio, P. L., & Thomas, J. H. (2000). A transmembrane guanylyl cyclase (DAF-11) and Hsp90 (DAF-21) regulate a common set of chemosensory behaviors in Caenorhabditis elegans. Genetics, 155(1), 85-104.
Chávez, V., Mohri-Shiomi, A., Maadani, A., Vega, L. A., & Garsin, D. A. (2007). Oxidative stress enzymes are required for DAF-16-mediated immunity due to generation of reactive oxygen species by Caenorhabditis elegans. Genetics, 176(3), 1567-1577.
Sun, J., Zhang, Y., Xu, T., Zhang, Y., Mu, H., Zhang, Y., ... & Qian, P. Y. (2017). Adaptation to deep-sea chemosynthetic environments as revealed by mussel genomes. Nature Ecology & Evolution, 1(5), 1-7.
Hinzke, T., Kleiner, M., Breusing, C., Felbeck, H., Häsler, R., Sievert, S. M., ... & Markert, S. (2019). Host-microbe interactions in the chemosynthetic Riftia pachyptila symbiosis. Mbio, 10(6), e02243-19.
Yuen, B., Polzin, J., & Petersen, J. M. (2019). Organ transcriptomes of the lucinid clam Loripes orbiculatus (Poli, 1791) provide insights into their specialized roles in the biology of a chemosymbiotic bivalve. BMC genomics, 20(1), 1-14.
Woyke, T., Teeling, H., Ivanova, N. N., Huntemann, M., Richter, M., Gloeckner, F. O., ... & Dubilier, N. (2006). Symbiosis insights through metagenomic analysis of a microbial consortium. Nature, 443(7114), 950-955.
Dubilier, N., Bergin, C., & Lott, C. (2008). Symbiotic diversity in marine animals: the art of harnessing chemosynthesis. Nature Reviews Microbiology, 6(10), 725-740.
Wippler, J., Kleiner, M., Lott, C., Gruhl, A., Abraham, P. E., Giannone, R. J., ... & Dubilier, N. (2016). Transcriptomic and proteomic insights into innate immunity and adaptations to a symbiotic lifestyle in the gutless marine worm Olavius algarvensis. BMC genomics, 17(1), 1-19.
Zimmermann, J., Wentrup, C., Sadowski, M., Blazejak, A., Gruber‐Vodicka, H. R., Kleiner, M., ... & Dubilier, N. (2016). Closely coupled evolutionary history of ecto‐and endosymbionts from two distantly related animal phyla. Molecular ecology, 25(13), 3203-3223.
Naylor, D. J., Hoogenraad, N. J., & Høj, P. B. (1996). Isolation and characterisation of a cDNA encoding rat mitochondrial GrpE, a stress-inducible nucleotide-exchange factor of ubiquitous appearance in mammalian organs. FEBS letters, 396(2-3), 181-188.
Lundin, V. F., Srayko, M., Hyman, A. A., & Leroux, M. R. (2008). Efficient chaperone-mediated tubulin biogenesis is essential for cell division and cell migration in C. elegans. Developmental biology, 313(1), 320-334.
Bar-Lavan, Y., Shemesh, N., Dror, S., Ofir, R., Yeger-Lotem, E., & Ben-Zvi, A. (2016). A differentiation transcription factor establishes muscle-specific proteostasis in Caenorhabditis elegans. PLoS genetics, 12(12), e1006531.
Suzuki, N., Inokuma, K., Yasuda, K., & Ishii, N. (1996). Cloning, sequencing and mapping of a manganese superoxide dismutase gene of the nematode Caenorhabditis elegans. DNA research, 3(3), 171-174.
Margis, R., Dunand, C., Teixeira, F. K., & Margis‐Pinheiro, M. (2008). Glutathione peroxidase family–an evolutionary overview. The FEBS journal, 275(15), 3959-3970.
Oliveira, R. P., Abate, J. P., Dilks, K., Landis, J., Ashraf, J., Murphy, C. T., & Blackwell, T. K. (2009). Condition‐adapted stress and longevity gene regulation by Caenorhabditis elegans SKN‐1/Nrf. Aging cell, 8(5), 524-541.
Selivanov, V. A., Votyakova, T. V., Zeak, J. A., Trucco, M., Roca, J., & Cascante, M. (2009). Bistability of mitochondrial respiration underlies paradoxical reactive oxygen species generation induced by anoxia. PLoS computational biology, 5(12), e1000619.
Semenza, G. L. (1999). Perspectives on oxygen sensing. Cell, 98(3), 281-284.
Hashimoto, T., Yonetani, M., & Nakamura, H. (2004). Selective brain hypothermia protects against hypoxic-ischemic injury in newborn rats by reducing hydroxyl radical production. Kobe Journal of Medical Sciences, 49(3/4), 83-92.
Borutaite, V., Mildaziene, V., Brown, G. C., & Brand, M. D. (1995). Control and kinetic analysis of ischemia-damaged heart mitochondria: which parts of the oxidative phosphorylation system are affected by ischemia?. Biochimica et Biophysica Acta (BBA)-Molecular Basis of Disease, 1272(3), 154-158.
Brookes, P. S., Yoon, Y., Robotham, J. L., Anders, M. W., & Sheu, S. S. (2004). Calcium, ATP, and ROS: a mitochondrial love-hate triangle. American Journal of Physiology-Cell Physiology, 287(4), C817-C833.
Brenner, C., & Moulin, M. (2012). Physiological roles of the permeability transition pore. Circulation research, 111(9), 1237-1247.
Hawrysh, P. J., & Buck, L. T. (2013). Anoxia-mediated calcium release through the mitochondrial permeability transition pore silences NMDA receptor currents in turtle neurons. Journal of Experimental Biology, 216(23), 4375-4387.
Horwitz, L. D., Fennessey, P. V., Shikes, R. H., & Kong, Y. (1994). Marked reduction in myocardial infarct size due to prolonged infusion of an antioxidant during reperfusion. Circulation, 89(4), 1792-1801.
Murphy, E., & Steenbergen, C. (2008). Mechanisms underlying acute protection from cardiac ischemia-reperfusion injury. Physiological reviews, 88(2), 581-609.
Fanter, C. E., Lin, Z., Keenan, S. W., Janzen, F. J., Mitchell, T. S., & Warren, D. E. (2020). Development-specific transcriptomic profiling suggests new mechanisms for anoxic survival in the ventricle of overwintering turtles. Journal of Experimental Biology, 223(4), jeb213918.
Chatenay-Lapointe, M., & Shadel, G. S. (2010). Stressed-out mitochondria get MAD. Cell metabolism, 12(6), 559-560.
Heo, J. M., Livnat-Levanon, N., Taylor, E. B., Jones, K. T., Dephoure, N., Ring, J., ... & Rutter, J. (2010). A stress-responsive system for mitochondrial protein degradation. Molecular cell, 40(3), 465-480.
Tullet, J. M. (2015). DAF-16 target identification in C. elegans: past, present and future. Biogerontology, 16(2), 221-234.
Kaushal, P. S., Sharma, M. R., Booth, T. M., Haque, E. M., Tung, C. S., Sanbonmatsu, K. Y., ... & Agrawal, R. K. (2014). Cryo-EM structure of the small subunit of the mammalian mitochondrial ribosome. Proceedings of the National Academy of Sciences, 111(20), 7284-7289.
Sharika, R., Subbaiah, P., & Balamurugan, K. (2018). Studies on reproductive stress caused by candidate Gram positive and Gram negative bacteria using model organism, Caenorhabditis elegans. Gene, 649, 113-126.
Melnikov S, Ben-Shem A, Garreau de Loubresse N, Jenner L, Yusupova G, Yusupov M. One core, two shells: bacterial and eukaryotic ribosomes. Nat Struct Mol Biol. 2012;19(6):560–567
You, K. T., Park, J., & Kim, V. N. (2015). Role of the small subunit processome in the maintenance of pluripotent stem cells. Genes & development, 29(19), 2004-2009.
Chen, J., & Kastan, M. B. (2010). 5′–3′-UTR interactions regulate p53 mRNA translation and provide a target for modulating p53 induction after DNA damage. Genes & development, 24(19), 2146-2156.
Savada, R. P., & Bonham-Smith, P. C. (2014). Differential transcript accumulation and subcellular localization of Arabidopsis ribosomal proteins. Plant Science, 223, 134-145.
Xu, X., Xiong, X., & Sun, Y. (2016). The role of ribosomal proteins in the regulation of cell proliferation, tumorigenesis, and genomic integrity. Science China Life Sciences, 59(7), 656-672.
Larade, K., Nimigan, A., & Storey, K. B. (2001). Transcription pattern of ribosomal protein L26 during anoxia exposure in Littorina littorea. Journal of Experimental Zoology, 290(7), 759-768.
Thomas, G. (2000). An encore for ribosome biogenesis in the control of cell proliferation. Nature cell biology, 2(5), E71-E72.
Shukla, S. K., & Kumar, V. (2012). Hepatitis B virus X protein and c‐M yc cooperate in the upregulation of ribosome biogenesis and in cellular transformation. The FEBS journal, 279(20), 3859-3871.
Xu, C., Hwang, W., Jeong, D. E., Ryu, Y., Ha, C. M., Lee, S. J. V., ... & He, Z. M. (2018). Genetic inhibition of an ATP synthase subunit extends lifespan in C. elegans. Scientific reports, 8(1), 1-14.
Maglioni, S., & Ventura, N. (2016). C. elegans as a model organism for human mitochondrial associated disorders. Mitochondrion, 30, 117-125.
McKay, R. M., McKay, J. P., Avery, L., & Graff, J. M. (2003). C. elegans: a model for exploring the genetics of fat storage. Developmental cell, 4(1), 131-142.
Rea, S. L., Ventura, N., & Johnson, T. E. (2007). Relationship between mitochondrial electron transport chain dysfunction, development, and life extension in Caenorhabditis elegans. PLoS biology, 5(10), e259.
Hartman, P. S., Ishii, N., Kayser, E. B., Morgan, P. G., & Sedensky, M. M. (2001). Mitochondrial mutations differentially affect aging, mutability and anesthetic sensitivity in Caenorhabditis elegans. Mechanisms of ageing and development, 122(11), 1187-1201.
Williams, J. C., Sue, C., Banting, G. S., Yang, H., Glerum, D. M., Hendrickson, W. A., & Schon, E. A. (2005). Crystal structure of human SCO1: implications for redox signaling by a mitochondrial cytochrome c oxidase “assembly” protein. Journal of Biological Chemistry, 280(15), 15202-15211.
Buceta, P. M. R., Romanelli-Cedrez, L., Babcock, S. J., Xun, H., VonPaige, M. L., Higley, T. W., ... & Salinas, G. (2019). The kynurenine pathway is essential for rhodoquinone biosynthesis in Caenorhabditis elegans. Journal of Biological Chemistry, 294(28), 11047-11053.
Del Borrello, S., Lautens, M., Dolan, K., Tan, J. H., Davie, T., Schertzberg, M. R., ... & Fraser, A. G. (2019). Rhodoquinone biosynthesis in C. elegans requires precursors generated by the kynurenine pathway. Elife, 8, e48165.
Tsai, P. C., Soong, B. W., Mademan, I., Huang, Y. H., Liu, C. R., Hsiao, C. T., ... & Lee, Y. C. (2017). A recurrent WARS mutation is a novel cause of autosomal dominant distal hereditary motor neuropathy. Brain, 140(5), 1252-1266.
Tan, J. H., Lautens, M., Romanelli-Cedrez, L., Wang, J., Schertzberg, M. R., Reinl, S. R., ... & Salinas, G. (2020). Alternative splicing of coq-2 controls the levels of rhodoquinone in animals. Elife, 9, e56376.
Smolková, K., & Ježek, P. (2012). The role of mitochondrial NADPH-dependent isocitrate dehydrogenase in cancer cells. International journal of cell biology, 2012.
Martínez-Reyes, I., & Chandel, N. S. (2020). Mitochondrial TCA cycle metabolites control physiology and disease. Nature communications, 11(1), 1-11.
Penkov, S., Kaptan, D., Erkut, C., Sarov, M., Mende, F., & Kurzchalia, T. V. (2015). Integration of carbohydrate metabolism and redox state controls dauer larva formation in Caenorhabditis elegans. Nature communications, 6(1), 1-10.
Yang, H. C., Yu, H., Liu, Y. C., Chen, T. L., Stern, A., Lo, S. J., & Chiu, D. T. Y. (2019). IDH-1 deficiency induces growth defects and metabolic alterations in GSPD-1-deficient Caenorhabditis elegans. Journal of Molecular Medicine, 97(3), 385-396.
Lutz, P. L., & Nilsson, G. E. (1997). Contrasting strategies for anoxic brain survival--glycolysis up or down. The Journal of experimental biology, 200(2), 411-419.
Semenza, G. L. (2001). HIF-1, O2, and the 3 PHDs: how animal cells signal hypoxia to the nucleus. Cell, 107(1), 1-3.
Huang, S., Colmer, T. D., & Millar, A. H. (2008). Does anoxia tolerance involve altering the energy currency towards PPi?. Trends in plant science, 13(5), 221-227.
Larade, K., & Storey, K. B. (2009). Living without oxygen: anoxia-responsive gene expression and regulation. Current Genomics, 10(2), 76-85.
Jackson, A. D., & McLaughlin, J. (2009). Digestion and absorption. Surgery (oxford), 27(6), 231-236.
Lodish, H., Berk, A., Kaiser, C. A., Kaiser, C., Krieger, M., Scott, M. P., ... & Matsudaira, P. (2008). Molecular cell biology. Macmillan.
Papaevgeniou, N., & Chondrogianni, N. (2014). The ubiquitin proteasome system in Caenorhabditis elegans and its regulation. Redox biology, 2, 333-347.
Jones, D., Crowe, E., Stevens, T. A., & Candido, E. P. M. (2001). Functional and phylogenetic analysis of the ubiquitylation system in Caenorhabditis elegans: ubiquitin-conjugating enzymes, ubiquitin-activating enzymes, and ubiquitin-like proteins. Genome biology, 3(1), 1-15.
Schaefer, H., & Rongo, C. (2006). KEL-8 is a substrate receptor for CUL3-dependent ubiquitin ligase that regulates synaptic glutamate receptor turnover. Molecular biology of the cell, 17(3), 1250-1260.
Stogios, P. J., Downs, G. S., Jauhal, J. J., Nandra, S. K., & Privé, G. G. (2005). Sequence and structural analysis of BTB domain proteins. Genome biology, 6(10),
Kim, K. W., Tang, N. H., Piggott, C. A., Andrusiak, M. G., Park, S., Zhu, M., ... & Jin, Y. (2018). Expanded genetic screening in Caenorhabditis elegans identifies new regulators and an inhibitory role for NAD+ in axon regeneration. Elife, 7, e39756.
Pintard, L., Kurz, T., Glaser, S., Willis, J. H., Peter, M., & Bowerman, B. (2003). Neddylation and deneddylation of CUL-3 is required to target MEI-1/Katanin for degradation at the meiosis-to-mitosis transition in C. elegans. Current Biology, 13(11), 911-921.
Brockway, H., Balukoff, N., Dean, M., Alleva, B., & Smolikove, S. (2014). The CSN/COP9 signalosome regulates synaptonemal complex assembly during meiotic prophase I of Caenorhabditis elegans. PLoS genetics, 10(11), e1004757.
Fraser, A. G., Kamath, R. S., Zipperlen, P., Martinez-Campos, M., Sohrmann, M., & Ahringer, J. (2000). Functional genomic analysis of C. elegans chromosome I by systematic RNA interference. Nature, 408(6810), 325-330.
Blumenthal, T., Evans, D., Link, C. D., Guffanti, A., Lawson, D., Thierry-Mieg, J., ... & Kim, S. K. (2002). A global analysis of Caenorhabditis elegans operons. Nature, 417(6891), 851-854.
Syntichaki, P., Xu, K., Driscoll, M., & Tavernarakis, N. (2002). Specific aspartyl and calpain proteases are required for neurodegeneration in C. elegans. Nature, 419(6910), 939-944.
Vassalli, J. D., Sappino, A. P., & Belin, D. (1991). The plasminogen activator/plasmin system. The Journal of clinical investigation, 88(4), 1067-1072.
Altincicek, B., Fischer, M., Fischer, M., Lüersen, K., Boll, M., Wenzel, U., & Vilcinskas, A. (2010). Role of matrix metalloproteinase ZMP-2 in pathogen resistance and development in Caenorhabditis elegans. Developmental & Comparative Immunology, 34(11), 1160-1169.
Fischer, M., Fitzenberger, E., Kull, R., Boll, M., & Wenzel, U. (2014). The zinc matrix metalloproteinase ZMP-2 increases survival of Caenorhabditis elegans through interference with lipoprotein absorption. Genes & nutrition, 9(4), 414.
Vabulas, R. M., & Hartl, F. U. (2005). Protein synthesis upon acute nutrient restriction relies on proteasome function. Science, 310(5756), 1960-1963.
Scott, R. C., Schuldiner, O., & Neufeld, T. P. (2004). Role and regulation of starvation-induced autophagy in the Drosophila fat body. Developmental cell, 7(2), 167-178.
Huber, L. A., & Teis, D. (2016). Lysosomal signaling in control of degradation pathways. Current opinion in cell biology, 39, 8-14.
Wang, R. C., & Levine, B. (2010). Autophagy in cellular growth control. FEBS letters, 584(7), 1417-1426.
Russell, R. C., Yuan, H. X., & Guan, K. L. (2014). Autophagy regulation by nutrient signaling. Cell research, 24(1), 42-57.
Liang, X. H., Jackson, S., Seaman, M., Brown, K., Kempkes, B., Hibshoosh, H., & Levine, B. (1999). Induction of autophagy and inhibition of tumorigenesis by beclin 1. Nature, 402(6762), 672-676.
Bar-Peled, L., Schweitzer, L. D., Zoncu, R., & Sabatini, D. M. (2012). Ragulator is a GEF for the rag GTPases that signal amino acid levels to mTORC1. Cell, 150(6), 1196-1208.
Meléndez, A., Tallóczy, Z., Seaman, M., Eskelinen, E. L., Hall, D. H., & Levine, B. (2003). Autophagy genes are essential for dauer development and life-span extension in C. elegans. Science, 301(5638), 1387-1391.
Wang, X., & Proud, C. G. (2009). Nutrient control of TORC1, a cell-cycle regulator. Trends in cell biology, 19(6), 260-267.
Thompson, A. R., & Vierstra, R. D. (2005). Autophagic recycling: lessons from yeast help define the process in plants. Current opinion in plant biology, 8(2), 165-173.
Ladevaia, V., Liu, R., & Proud, C. G. (2014, December). mTORC1 signaling controls multiple steps in ribosome biogenesis. In Seminars in cell & developmental biology (Vol. 36, pp. 113-120). Academic Press.
Ma, X. M., & Blenis, J. (2009). Molecular mechanisms of mTOR-mediated translational control. Nature reviews Molecular cell biology, 10(5), 307-318.
Howell, J. J., & Manning, B. D. (2011). mTOR couples cellular nutrient sensing to organismal metabolic homeostasis. Trends in Endocrinology & Metabolism, 22(3), 94-102.
Nussbaumer, A.D., Bright, M., Baranyi, C., Beisser, C.J., and Ott, J.A. (2004) Attachment mechanism in a highly specific association between ectosymbiotic bacteria and marine nematodes. Aquat Microb Ecol 34:239–246
Bulgheresi S, Schabussova I, Chen T, Mullin NP, Maizels RM, Ott JA. A new C-type lectin similar to the human immunoreceptor DC-SIGN mediates symbiont acquisition by a marine nematode. Appl Environ Microbiol. 2006;72:2950–2956
Bulgheresi S, Gruber-Vodicka HR, Heindl NR, Dirks U, Kostadinova M, Breiteneder H, Ott JA. Sequence variability of the pattern recognition receptor Mermaid mediates specificity of marine nematode symbioses. ISME J. 2011;5:986–998
Koropatkin, N. M., Cameron, E. A., & Martens, E. C. (2012). How glycan metabolism shapes the human gut microbiota. Nature Reviews Microbiology, 10(5), 323-335.
Simon, H. U., Haj-Yehia, A., & Levi-Schaffer, F. (2000). Role of reactive oxygen species (ROS) in apoptosis induction. Apoptosis, 5(5), 415-418.
Martinou, J. C., Desagher, S., & Antonsson, B. (2000). Cytochrome c release from mitochondria: all or nothing. Nature cell biology, 2(3), E41-E43.
Gogvadze, V., Orrenius, S., & Zhivotovsky, B. (2006). Multiple pathways of cytochrome c release from mitochondria in apoptosis. Biochimica et Biophysica Acta (BBA)-Bioenergetics, 1757(5-6), 639-647.
Mangahas, P. M., & Zhou, Z. (2005, April). Clearance of apoptotic cells in Caenorhabditis elegans. In Seminars in cell & developmental biology (Vol. 16, No. 2, pp. 295-306). Academic Press.
Kaufmann, S. H., Lee, S. H., Meng, X. W., Loegering, D. A., Kottke, T. J., Henzing, A. J., ... & Earnshaw, W. C. (2008). Apoptosis-associated caspase activation assays. Methods, 44(3), 262-272.
Liu, X., Kim, C. N., Yang, J., Jemmerson, R., & Wang, X. (1996). Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell, 86(1), 147-157.
Tafani, M., Schneider, T. G., Pastorino, J. G., & Farber, J. L. (2000). Cytochrome c-dependent activation of caspase-3 by tumor necrosis factor requires induction of the mitochondrial permeability transition. The American journal of pathology, 156(6), 2111-2121.
Takacs-Vellai, K., Vellai, T., Puoti, A., Passannante, M., Wicky, C., Streit, A., ... & Müller, F. (2005). Inactivation of the autophagy gene bec-1 triggers apoptotic cell death in C. elegans. Current biology, 15(16), 1513-1517.
Wang, X., Li, W., Zhao, D., Liu, B., Shi, Y., Chen, B., ... & Xue, D. (2010). Caenorhabditis elegans transthyretin-like protein TTR-52 mediates recognition of apoptotic cells by the CED-1 phagocyte receptor. Nature cell biology, 12(7), 655-664.
Chen, Y. Z., Mapes, J., Lee, E. S., Skeen-Gaar, R. R., & Xue, D. (2013). Caspase-mediated activation of Caenorhabditis elegans CED-8 promotes apoptosis and phosphatidylserine externalization. Nature communications, 4(1), 1-9.
Parrish, J. Z., & Xue, D. (2003). Functional genomic analysis of apoptotic DNA degradation in C. elegans. Molecular cell, 11(4), 987-996.
Samejima, K., & Earnshaw, W. C. (2005). Trashing the genome: the role of nucleases during apoptosis. Nature reviews Molecular cell biology, 6(9), 677-688.
Sasaki, A., Nakae, I., Nagasawa, M., Hashimoto, K., Abe, F., Saito, K., ... & Kontani, K. (2013). Arl8/ARL-8 functions in apoptotic cell removal by mediating phagolysosome formation in Caenorhabditis elegans. Molecular biology of the cell, 24(10), 1584-1592.
Hurwitz, M. E., Vanderzalm, P. J., Bloom, L., Goldman, J., Garriga, G., & Horvitz, H. R. (2009). Abl kinase inhibits the engulfment of apopotic cells in Caenorhabditis elegans. PLoS biology, 7(4), e1000099.
Berdichevsky, A., Nedelcu, S., Boulias, K., Bishop, N. A., Guarente, L., & Horvitz, H. R. (2010). 3-Ketoacyl thiolase delays aging of Caenorhabditis elegans and is required for lifespan extension mediated by sir-2.1. Proceedings of the National Academy of Sciences, 107(44), 18927-18932.
Chughtai, A. A., Kaššák, F., Kostrouchová, M., Novotný, J. P., Krause, M. W., Saudek, V., ... & Kostrouchová, M. (2015). Perilipin-related protein regulates lipid metabolism in C. elegans. PeerJ, 3, e1213.
Horikawa, M., & Sakamoto, K. (2009). Fatty-acid metabolism is involved in stress-resistance mechanisms of Caenorhabditis elegans. Biochemical and biophysical research communications, 390(4), 1402-1407.
Krivoruchko, A., & Storey, K. B. (2015). Turtle anoxia tolerance: biochemistry and gene regulation. Biochimica et Biophysica Acta (BBA)-General Subjects, 1850(6), 1188-1196.
Brendza, K. M., Haakenson, W., Cahoon, R. E., Hicks, L. M., Palavalli, L. H., Chiapelli, B. J., ... & Jez, J. M. (2007). Phosphoethanolamine N-methyltransferase (PMT-1) catalyses the first reaction of a new pathway for phosphocholine biosynthesis in Caenorhabditis elegans. Biochemical Journal, 404(3), 439-448.
Thomas, J. H. (1990). Genetic analysis of defecation in Caenorhabditis elegans. Genetics, 124(4), 855-872.
Mclntire, S. L., Jorgensen, E., Kaplan, J., & Horvitz, H. R. (1993). The GABAergic nervous system of Caenorhabditis elegans. Nature, 364(6435), 337-341.
Jin, Y., Jorgensen, E., Hartwieg, E., & Horvitz, H. R. (1999). The Caenorhabditis elegans gene unc-25 encodes glutamic acid decarboxylase and is required for synaptic transmission but not synaptic development. Journal of Neuroscience, 19(2), 539-548.
Gally, C., & Bessereau, J. L. (2003). GABA is dispensable for the formation of junctional GABA receptor clusters in Caenorhabditis elegans. Journal of Neuroscience, 23(7), 2591-2599.
Nordquist, S. K., Smith, S. R., & Pierce, J. T. (2018). Systematic functional characterization of human 21st chromosome orthologs in Caenorhabditis elegans. G3: Genes, Genomes, Genetics, 8(3), 967-979.
Risley, M. G., Kelly, S. P., Jia, K., Grill, B., & Dawson-Scully, K. (2016). Modulating behavior in C. elegans using electroshock and antiepileptic drugs. PLoS One, 11(9), e0163786.
Martin, D. L., & Rimvall, K. (1993). Regulation of γ‐aminobutyric acid synthesis in the brain. Journal of neurochemistry, 60(2), 395-407.
van der Vos KE, Coffer PJ. Glutamine metabolism links growth factor signaling to the regulation of autophagy. Autophagy. 2012;8:1862–1864
Yen, C. A., & Curran, S. P. (2021). Incomplete proline catabolism drives premature sperm aging. Aging cell, 20(2), e13308.
Tharmalingam, S., Burns, A. R., Roy, P. J., & Hampson, D. R. (2012). Orthosteric and allosteric drug binding sites in the Caenorhabditis elegans mgl-2 metabotropic glutamate receptor. Neuropharmacology, 63(4), 667-674.
Baker, A. J., Zornow, M. H., Scheller, M. S., Yaksh, T. L., Skilling, S. R., Smullin, D. H., ... & Kuczenski, R. (1991). Changes in extracellular concentrations of glutamate, aspartate, glycine, dopamine, serotonin, and dopamine metabolites after transient global ischemia in the rabbit brain. Journal of neurochemistry, 57(4), 1370-1379.
Lutz, P. L., Nilsson, G. E., & Prentice, H. M. (2003a). The brain without oxygen: causes of failure-physiological and molecular mechanisms for survival. Springer Science & Business Media.
Milton, S. L., Thompson, J. W., & Lutz, P. L. (2002). Mechanisms for maintaining extracellular glutamate levels in the anoxic turtle striatum. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, 282(5), R1317-R1323.
Mathews, G. C., & Diamond, J. S. (2003). Neuronal glutamate uptake contributes to GABA synthesis and inhibitory synaptic strength. Journal of Neuroscience, 23(6), 2040-2048.
Milton, S. L., & Lutz, P. L. (1998). Low extracellular dopamine levels are maintained in the anoxic turtle (Trachemys scripta) striatum. Journal of Cerebral Blood Flow & Metabolism, 18(7), 803-807.
Lutz, P. L., Prentice, H. M., & Milton, S. L. (2003b). Is turtle longevity linked to enhanced mechanisms for surviving brain anoxia and reoxygenation?. Experimental Gerontology, 38(7), 797-800.
Nilsson, G. E. (1990). Long-term anoxia in crucian carp: changes in the levels of amino acid and monoamine neurotransmitters in the brain, catecholamines in chromaffin tissue, and liver glycogen. Journal of Experimental Biology, 150(1), 295-320.
Schuske, K., Beg, A. A., & Jorgensen, E. M. (2004). The GABA nervous system in C. elegans. Trends in neurosciences, 27(7), 407-414.
Sawin, E. R., Ranganathan, R., & Horvitz, H. R. (2000). C. elegans locomotory rate is modulated by the environment through a dopaminergic pathway and by experience through a serotonergic pathway. Neuron, 26(3), 619-631.
Sanyal, S., Wintle, R. F., Kindt, K. S., Nuttley, W. M., Arvan, R., Fitzmaurice, P., ... & Van Tol, H. H. (2004). Dopamine modulates the plasticity of mechanosensory responses in Caenorhabditis elegans. The EMBO journal, 23(2), 473-482.
Gainetdinov, R. R., Sotnikova, T. D., & Caron, M. G. (2002). Monoamine transporter pharmacology and mutant mice. Trends in pharmacological sciences, 23(8), 367-373.
McDonald, P. W., Jessen, T., Field, J. R., & Blakely, R. D. (2006). Dopamine signaling architecture in Caenorhabditis elegans. Cellular and molecular neurobiology, 26(4), 591-616.
Soontornniyomkij, V., Risbrough, V. B., Young, J. W., Soontornniyomkij, B., Jeste, D. V., & Achim, C. L. (2012). Hippocampal calbindin-1 immunoreactivity correlate of recognition memory performance in aged mice. Neuroscience letters, 516(1), 161-165.
Hobert, O. (2018). The neuronal genome of Caenorhabditis elegans. WormBook: The Online Review of C. elegans Biology
Steger, K. A., Shtonda, B. B., Thacker, C., Snutch, T. P., & Avery, L. (2005). The C. elegans T-type calcium channel CCA-1 boosts neuromuscular transmission. Journal of Experimental Biology, 208(11), 2191-2203.
Zhou, K., Cherra III, S. J., Goncharov, A., & Jin, Y. (2017). Asynchronous cholinergic drive correlates with excitation-inhibition imbalance via a neuronal Ca2+ sensor protein. Cell reports, 19(6), 1117-1129.
Bickler, P. E. (1992). Cerebral anoxia tolerance in turtles: regulation of intracellular calcium and pH. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, 263(6), R1298-R1302.
Dell'Anna, E., Geloso, M. C., Magarelli, M., & Molinari, M. (1996). Development of GABA and calcium binding proteins immunoreactivity in the rat hippocampus following neonatal anoxia. Neuroscience letters, 211(2), 93-96.
Akira, S., Uematsu, S., & Takeuchi, O. (2006). Pathogen recognition and innate immunity. Cell, 124(4), 783-801.
Ott et al., 2021. BOOK CHAPTER
Wang, D. (2019). Epidermal Barrier for Nematodes Against Toxicity of Environmental Toxicants or Stresses. In Target Organ Toxicology in Caenorhabditis elegans (pp. 97-122). Springer, Singapore.
Dravid, P., Kaushal, D. C., Saxena, J. K., & Kaushal, N. A. (2015). Isolation and characterization of endochitinase and exochitinase of Setaria cervi. Parasitology international, 64(6), 579-586.
Krasity, B. C., Troll, J. V., Lehnert, E. M., Hackett, K. T., Dillard, J. P., Apicella, M. A., ... & McFall-Ngai, M. J. (2015). Structural and functional features of a developmentally regulated lipopolysaccharide-binding protein. MBio, 6(5), e01193-15.
Chen, F., Krasity, B. C., Peyer, S. M., Koehler, S., Ruby, E. G., Zhang, X., & McFall-Ngai, M. J. (2017). MBio.
Pradel, E., Zhang, Y., Pujol, N., Matsuyama, T., Bargmann, C. I., & Ewbank, J. J. (2007). Detection and avoidance of a natural product from the pathogenic bacterium Serratia marcescens by Caenorhabditis elegans. Proceedings of the National Academy of Sciences, 104(7), 2295-2300.
Brandt, J. P., & Ringstad, N. (2015). Toll-like receptor signaling promotes development and function of sensory neurons required for a C. elegans pathogen-avoidance behavior. Current Biology, 25(17), 2228-2237.
Dang, H., & Lovell, C. R. (2016). Microbial surface colonization and biofilm development in marine environments. Microbiology and molecular biology reviews, 80(1), 91-138.
Suzuki, M., Sagoh, N., Iwasaki, H., Inoue, H., & Takahashi, K. (2004). Metalloproteases with EGF, CUB, and thrombospondin-1 domains function in molting of Caenorhabditis elegans.
Zhang, Y., Foster, J. M., Nelson, L. S., Ma, D., & Carlow, C. K. (2005). The chitin synthase genes chs-1 and chs-2 are essential for C. elegans development and responsible for chitin deposition in the eggshell and pharynx, respectively. Developmental biology, 285(2), 330-339.
Zugasti, O., Rajan, J., & Kuwabara, P. E. (2005). The function and expansion of the Patched-and Hedgehog-related homologs in C. elegans. Genome research, 15(10), 1402-1410.
Hornsten, A., Lieberthal, J., Fadia, S., Malins, R., Ha, L., Xu, X., ... & Li, C. (2007). APL-1, a Caenorhabditis elegans protein related to the human β-amyloid precursor protein, is essential for viability. Proceedings of the National Academy of Sciences, 104(6), 1971-1976.
Russel, S., Frand, A. R., & Ruvkun, G. (2011). Regulation of the C. elegans molt by pqn-47. Developmental biology, 360(2), 297-309.
Pan, K. Z., Palter, J. E., Rogers, A. N., Olsen, A., Chen, D., Lithgow, G. J., & Kapahi, P. (2007). Inhibition of mRNA translation extends lifespan in Caenorhabditis elegans. Aging cell, 6(1), 111-119.
Pal, S., Lant, B., Yu, B., Tian, R., Tong, J., Krieger, J. R., ... & Derry, W. B. (2017). CCM-3 promotes C. elegans germline development by regulating vesicle trafficking cytokinesis and polarity. Current Biology, 27(6), 868-876.
Park, B. J., Lee, D. G., Yu, J. R., Jung, S. K., Choi, K., Lee, J., ... & Ahnn, J. (2001). Calreticulin, a calcium-binding molecular chaperone, is required for stress response and fertility in Caenorhabditis elegans. Molecular Biology of the Cell, 12(9), 2835-2845.
Clark, S. G., Shurland, D. L., Meyerowitz, E. M., Bargmann, C. I., & Van Der Bliek, A. M. (1997). A dynamin GTPase mutation causes a rapid and reversible temperature-inducible locomotion defect in C. elegans. Proceedings of the National Academy of Sciences, 94(19), 10438-10443.
Goedert, M., Baur, C. P., Ahringer, J., Jakes, R., Hasegawa, M., Spillantini, M. G., ... & Hill, F. (1996). PTL-1, a microtubule-associated protein with tau-like repeats from the nematode Caenorhabditis elegans. Journal of cell science, 109(11), 2661-2672.
Gatewood, B. K., & Bucher, E. A. (1997). The mup-4 locus in Caenorhabditis elegans is essential for hypodermal integrity, organismal morphogenesis and embryonic body wall muscle position. Genetics, 146(1), 165-183.
Fujii, T., Nakao, F., Shibata, Y., Shioi, G., Kodama, E., Fujisawa, H., & Takagi, S. (2002). Caenorhabditis elegans PlexinA, PLX-1, interacts with transmembrane semaphorins and regulates epidermal morphogenesis.
Gally, C., Wissler, F., Zahreddine, H., Quintin, S., Landmann, F., & Labouesse, M. (2009). Myosin II regulation during C. elegans embryonic elongation: LET-502/ROCK, MRCK-1 and PAK-1, three kinases with different roles. Development, 136(18), 3109-3119.
Zahreddine, H., Zhang, H., Diogon, M., Nagamatsu, Y., & Labouesse, M. (2010). CRT-1/calreticulin and the E3 ligase EEL-1/HUWE1 control hemidesmosome maturation in C. elegans development. Current Biology, 20(4), 322-327.
Jee, C., Choi, T. W., Kalichamy, K., Yee, J. Z., Song, H. O., Ji, Y. J., ... & Lee, S. K. (2012). CNP-1 (ARRD-17), a novel substrate of calcineurin, is critical for modulation of egg-laying and locomotion in response to food and lysine sensation in Caenorhabditis elegans. Journal of molecular biology, 417(3), 165-178.
Warner, A., Xiong, G., Qadota, H., Rogalski, T., Vogl, A. W., Moerman, D. G., & Benian, G. M. (2013). CPNA-1, a copine domain protein, is located at integrin adhesion sites and is required for myofilament stability in Caenorhabditis elegans. Molecular biology of the cell, 24(5), 601-616.
Perez, M. F., & Lehner, B. (2019). Vitellogenins-yolk gene function and regulation in Caenorhabditis elegans. Frontiers in physiology, 10, 1067.
Ding, M., Woo, W. M., & Chisholm, A. D. (2004). The cytoskeleton and epidermal morphogenesis in C. elegans. Experimental cell research, 301(1), 84-90.
Osório, D. S., Chan, F. Y., Saramago, J., Leite, J., Silva, A. M., Sobral, A. F., ... & Carvalho, A. X. (2019). Crosslinking activity of non-muscle myosin II is not sufficient for embryonic cytokinesis in C. elegans. Development, 146(21), dev179150.
Nelson, M. D., Zhou, E., Kiontke, K., Fradin, H., Maldonado, G., Martin, D., ... & Fitch, D. H. (2011). A bow-tie genetic architecture for morphogenesis suggested by a genome-wide RNAi screen in Caenorhabditis elegans. PLoS genetics, 7(3), e1002010.
Dalpé, G., Zhang, L. W., Zheng, H., & Culotti, J. G. (2004). Conversion of cell movement responses to Semaphorin-1 and Plexin-1 from attraction to repulsion by lowered levels of specific RAC GTPases in C. elegans.
Dalpe, G., Tarsitano, M., Persico, M. G., Zheng, H., & Culotti, J. (2013). C. elegans PVF-1 inhibits permissive UNC-40 signalling through CED-10 GTPase to position the male ray 1 sensillum. Development, 140(19), 4020-4030.
Dufourcq, P., Victor, M., Gay, F., Calvo, D., Hodgkin, J., & Shi, Y. (2002). Functional requirement for histone deacetylase 1 in Caenorhabditis elegans gonadogenesis. Molecular and cellular biology, 22(9), 3024-3034.
Choy, S. W., Wong, Y. M., Ho, S., & Chow, K. L. (2007). C. elegans SIN-3 and its associated HDAC corepressor complex act as mediators of male sensory ray development. Biochemical and biophysical research communications, 358(3), 802-807.
Park, J. O., Pan, J., Möhrlen, F., Schupp, M. O., Johnsen, R., Baillie, D. L., ... & Hutter, H. (2010). Characterization of the astacin family of metalloproteases in C. elegans. BMC developmental biology, 10(1), 1-13.
Topf, U., & Drabikowski, K. (2019). Ancient function of teneurins in tissue organization and neuronal guidance in the nematode Caenorhabditis elegans. Frontiers in neuroscience, 13, 205.
Spanier, B., Stürzenbaum, S. R., Holden-Dye, L. M., & Baumeister, R. (2005). Caenorhabditis elegans neprilysin NEP-1: an effector of locomotion and pharyngeal pumping. Journal of molecular biology, 352(2), 429-437.
Norman, K. R., Fazzio, R. T., Mellem, J. E., Espelt, M. V., Strange, K., Beckerle, M. C., & Maricq, A. V. (2005). The Rho/Rac-family guanine nucleotide exchange factor VAV-1 regulates rhythmic behaviors in C. elegans. Cell, 123(1), 119-132.
De Cuyper, C., & Vanfleteren, J. R. (1982). Oxygen consumption during development and aging of the nematode Caenorhabditis elegans. Comparative Biochemistry and Physiology Part A: Physiology, 73(2), 283-289.
Uppaluri, S., & Brangwynne, C. P. (2015). A size threshold governs Caenorhabditis elegans developmental progression. Proceedings of the Royal Society B: Biological Sciences, 282(1813), 20151283.
Pellerone, F. I., Archer, S. K., Behm, C. A., Grant, W. N., Lacey, M. J., & Somerville, A. C. (2003). Trehalose metabolism genes in Caenorhabditis elegans and filarial nematodes. International journal for parasitology, 33(11), 1195-1206.
Schuster, L. N., & Sommer, R. J. (2012). Expressional and functional variation of horizontally acquired cellulases in the nematode Pristionchus pacificus. Gene, 506(2), 274-282.
Yuan, Y., Kadiyala, C. S., Ching, T. T., Hakimi, P., Saha, S., Xu, H., ... & Feng, Z. (2012). Enhanced energy metabolism contributes to the extended life span of calorie-restricted Caenorhabditis elegans. Journal of Biological Chemistry, 287(37), 31414-31426.
Kitaoka, S., Morielli, A. D., & Zhao, F. Q. (2013). FGT-1 is a mammalian GLUT2-like facilitative glucose transporter in Caenorhabditis elegans whose malfunction induces fat accumulation in intestinal cells. PLoS One, 8(6), e68475.
Bertoli, S., Neri, I. G., Trentani, C., Ferraris, C., De Amicis, R., Battezzati, A., ... & Tagliabue, A. (2015). Short-term effects of ketogenic diet on anthropometric parameters, body fat distribution, and inflammatory cytokine production in GLUT1 deficiency syndrome. Nutrition, 31(7-8), 981-987.
Miyagawa, K., Sakamoto, H., Yoshida, T., Yamashita, Y., Mitsui, Y., Furusawa, M.... & Terada, M. (1988). hst-1 transforming protein: expression in silkworm cells and characterization as a novel heparin-binding growth factor. Oncogene, 3(4), 383-389.
Bhattacharya, R., Townley, R. A., Berry, K. L., & Bülow, H. E. (2009). The PAPS transporter PST-1 is required for heparan sulfation and is essential for viability and neural development in C. elegans. Journal of cell science, 122(24), 4492-4504.
Crowe, J. H., Crowe, L. M., Carpenter, J. F., & Wistrom, C. A. (1987). Stabilization of dry phospholipid bilayers and proteins by sugars. Biochemical Journal, 242(1), 1.
Carpenter, J. F., & Crowe, J. H. (1988). Modes of stabilization of a protein by organic solutes during desiccation. Cryobiology, 25(5), 459-470.
Chen, Q., Ma, E., Behar, K. L., Xu, T., & Haddad, G. G. (2002). Role of trehalose phosphate synthase in anoxia tolerance and development in Drosophila melanogaster. Journal of Biological Chemistry, 277(5), 3274-3279.
Mongan, N. P., Jones, A. K., Smith, G. R., Sansom, M. S., & Sattelle, D. B. (2002). Novel α7‐like nicotinic acetylcholine receptor subunits in the nematode Caenorhabditis elegans. Protein Science, 11(5), 1162-1171.
Patton, A., Knuth, S., Schaheen, B., Dang, H., Greenwald, I., & Fares, H. (2005). Endocytosis function of a ligand-gated ion channel homolog in Caenorhabditis elegans. Current biology, 15(11), 1045-1050.
Gendrel, M., Rapti, G., Richmond, J. E., & Bessereau, J. L. (2009). A secreted complement-control-related protein ensures acetylcholine receptor clustering. Nature, 461(7266), 992-996.
Boulin, T., Rapti, G., Briseño-Roa, L., Stigloher, C., Richmond, J. E., Paoletti, P., & Bessereau, J. L. (2012). Positive modulation of a Cys-loop acetylcholine receptor by an auxiliary transmembrane subunit. Nature neuroscience, 15(10), 1374-1381.
Chan, J. P., Hu, Z., & Sieburth, D. (2012). Recruitment of sphingosine kinase to presynaptic terminals by a conserved muscarinic signaling pathway promotes neurotransmitter release. Genes & development, 26(10), 1070-1085.
Sun, L., Zang, W. J., Wang, H., Zhao, M., Yu, X. J., He, X., ... & Zhou, J. (2014). Acetylcholine promotes ROS detoxification against hypoxia/reoxygenation-induced oxidative stress through FoxO3a/PGC-1α dependent superoxide dismutase. Cellular Physiology and Biochemistry, 34(5), 1614-1625.
Guest, M., Bull, K., Walker, R. J., Amliwala, K., O’Connor, V., Harder, A., ... & Hopper, N. A. (2007). The calcium-activated potassium channel, SLO-1, is required for the action of the novel cyclo-octadepsipeptide anthelmintic, emodepside, in Caenorhabditis elegans. International journal for parasitology, 37(14), 1577-1588.
Hurd, D. D., Miller, R. M., Núñez, L., & Portman, D. S. (2010). Specific α-and β-tubulin isotypes optimize the functions of sensory cilia in Caenorhabditis elegans. Genetics, 185(3), 883-896.
Wang, Z., Hou, Y., Guo, X., van der Voet, M., Boxem, M., Dixon, J. E., ... & Jin, Y. (2013). The EBAX-type Cullin-RING E3 ligase and Hsp90 guard the protein quality of the SAX-3/Robo receptor in developing neurons. Neuron, 79(5), 903-916.
Woo, W. M., Berry, E. C., Hudson, M. L., Swale, R. E., Goncharov, A., & Chisholm, A. D. (2008). The C. elegans F-spondin family protein SPON-1 maintains cell adhesion in neural and non-neural tissues.
Schwarz, V., Pan, J., Voltmer-Irsch, S., & Hutter, H. (2009). IgCAMs redundantly control axon navigation in Caenorhabditis elegans. Neural development, 4(1), 1-15.
Gu, G. U. O. Q. I. A. N. G., Caldwell, G. A., & Chalfie, M. (1996). Genetic interactions affecting touch sensitivity in Caenorhabditis elegans. Proceedings of the National Academy of Sciences, 93(13), 6577-6582.
Han, L., Wang, Y., Sangaletti, R., D'Urso, G., Lu, Y., Shaham, S., & Bianchi, L. (2013). Two novel DEG/ENaC channel subunits expressed in glia are needed for nose-touch sensitivity in Caenorhabditis elegans. Journal of Neuroscience, 33(3), 936-949.
Li, Z., Li, Y., Yi, Y., Huang, W., Yang, S., Niu, W., ... & Xu, T. (2012). Dissecting a central flip-flop circuit that integrates contradictory sensory cues in C. elegans feeding regulation. Nature communications, 3(1), 1-8.
Tsuji, N., Morales, T. H., Ozols, V. V., Carmody, A. B., & Chandrashekar, R. (1999). Identification of an asparagine amidohydrolase from the filarial parasite Dirofilaria immitis. International journal for parasitology, 29(9), 1451-1455.
Chen, C. C., Lim, C. Y., Lee, P. J., Hsu, A. L., & Ching, T. T. (2020). S-adenosyl methionine synthetase SAMS-5 mediates dietary restriction-induced longevity in Caenorhabditis elegans. PloS one, 15(11), e0241455.
Russell, D., & Snyder, S. H. (1968). Amine synthesis in rapidly growing tissues: ornithine decarboxylase activity in regenerating rat liver, chick embryo, and various tumors. Proceedings of the National Academy of Sciences of the United States of America, 60(4), 1420.
Heby, O. (1981). Role of polyamines in the control of cell proliferation and differentiation. Differentiation, 19(1-3), 1-20.
Gilad, G. M., & Gilad, V. H. (1991). Polyamines can protect against ischemia-induced nerve cell death in gerbil forebrain. Experimental neurology, 111(3), 349-355.
Longo, L. D., Packianathan, S., McQueary, J. A., Stagg, R. B., Byus, C. V., & Cain, C. D. (1993). Acute hypoxia increases ornithine decarboxylase activity and polyamine concentrations in fetal rat brain. Proceedings of the National Academy of Sciences, 90(2), 692-696.
Ward, J. D., Mullaney, B., Schiller, B. J., He, L. D., Petnic, S. E., Couillault, C., ... & Yamamoto, K. R. (2014). Defects in the C. elegans acyl-CoA synthase, acs-3, and nuclear hormone receptor, nhr-25, cause sensitivity to distinct, but overlapping stresses. PloS one, 9(3), e92552.
Wang, F., Dai, Y., Zhu, X., Chen, Q., Zhu, H., Zhou, B., ... & Pang, S. (2021). Saturated very long chain fatty acid configures glycosphingolipid for lysosome homeostasis in long-lived C. elegans. Nature Communications, 12(1), 1-14.
Bastiani, C. A., Gharib, S., Simon, M. I., & Sternberg, P. W. (2003). Caenorhabditis elegans Gαq regulates egg-laying behavior via a PLCβ-independent and serotonin-dependent signaling pathway and likely functions both in the nervous system and in muscle. Genetics, 165(4), 1805-1822.
Taha, T. A., Kitatani, K., El‐Alwani, M., Bielawski, J., Hannun, Y. A., & Obeid, L. M. (2006). Loss of sphingosine kinase‐1 activates the intrinsic pathway of programmed cell death: modulation of sphingolipid levels and the induction of apoptosis. The FASEB journal, 20(3), 482-484.
Deng, X., Yin, X., Allan, R., Lu, D. D., Maurer, C. W., Haimovitz-Friedman, A., ... & Kolesnick, R. (2008). Ceramide biogenesis is required for radiation-induced apoptosis in the germ line of C. elegans. Science, 322(5898), 110-115.
Menuz, V., Howell, K. S., Gentina, S., Epstein, S., Riezman, I., Fornallaz-Mulhauser, M., ... & Martinou, J. C. (2009). Protection of C. elegans from anoxia by HYL-2 ceramide synthase. Science, 324(5925), 381-384.
Watts, J. L., & Ristow, M. (2017). Lipid and carbohydrate metabolism in Caenorhabditis elegans. Genetics, 207(2), 413-446.
Lutz, P. L., Nilsson, G. E., & Peréz-Pinzón, M. A. (1996). Anoxia tolerant animals from a neurobiological perspective. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 113(1), 3-13.
Teramoto, T., Sternick, L. A., Kage-Nakadai, E., Sajjadi, S., Siembida, J., Mitani, S., ... & Lambie, E. J. (2010). Magnesium excretion in C. elegans requires the activity of the GTL-2 TRPM channel. PloS one, 5(3), e9589.
Tsunenari, T., Sun, H., Williams, J., Cahill, H., Smallwood, P., Yau, K. W., & Nathans, J. (2003). Structure-function analysis of the bestrophin family of anion channels. Journal of Biological Chemistry, 278(42), 41114-41125.
Wang, Y., Alam, T., Hill-Harfe, K., Lopez, A. J., Leung, C. K., Iribarne, D., ... & Choe, K. P. (2013). Phylogenetic, expression, and functional analyses of anoctamin homologs in Caenorhabditis elegans. American Journal of Physiology-Regulatory, Integrative and Comparative Physiology, 305(11), R1376-R1389.
Goh, K. Y., & Inoue, T. (2018). A large transcribed enhancer region regulates C. elegans bed-3 and the development of egg laying muscles. Biochimica et Biophysica Acta (BBA)-Gene Regulatory Mechanisms, 1861(5), 519-533.
Currie, E., King, B., Lawrenson, A. L., Schroeder, L. K., Kershner, A. M., & Hermann, G. J. (2007). Role of the Caenorhabditis elegans multidrug resistance gene, mrp-4, in gut granule differentiation. Genetics, 177(3), 1569-1582.
Schroeder, L. K., Kremer, S., Kramer, M. J., Currie, E., Kwan, E., Watts, J. L., ... & Hermann, G. J. (2007). Function of the Caenorhabditis elegans ABC transporter PGP-2 in the biogenesis of a lysosome-related fat storage organelle. Molecular biology of the cell, 18(3), 995-1008.
Kage-Nakadai, E., Uehara, T., & Mitani, S. (2011). H+/myo-inositol transporter genes, hmit-1.1 and hmit-1.2, have roles in the osmoprotective response in Caenorhabditis elegans. Biochemical and biophysical research communications, 410(3), 471-477.
Pao, S. S., Paulsen, I. T., & Saier Jr, M. H. (1998). Major facilitator superfamily. Microbiology and molecular biology reviews, 62(1), 1-34.
Romanelli-Credrez, L., Doitsidou, M., Alkema, M. J., & Salinas, G. (2020). HIF-1 Has a Central Role in Caenorhabditis elegans Organismal Response to Selenium. Frontiers in genetics, 11, 63.
Filipovic, M. R., Zivanovic, J., Alvarez, B., & Banerjee, R. (2018). Chemical biology of H2S signaling through persulfidation. Chemical reviews, 118(3), 1253-1337.
Hayes, J. D., & McLellan, L. I. (1999). Glutathione and glutathione-dependent enzymes represent a coordinately regulated defense against oxidative stress. Free radical research, 31(4), 273-300.
Mytilineou, C., Kramer, B. C., & Yabut, J. A. (2002). Glutathione depletion and oxidative stress. Parkinsonism & related disorders, 8(6), 385-387.
Diaz-Vivancos, P., de Simone, A., Kiddle, G., & Foyer, C. H. (2015). Glutathione–linking cell proliferation to oxidative stress. Free Radical Biology and Medicine, 89, 1154-1164.
Morimoto-Tomita, M., Uchimura, K., Werb, Z., Hemmerich, S., & Rosen, S. D. (2002). Cloning and characterization of two extracellular heparin-degrading endosulfatases in mice and humans. Journal of Biological Chemistry, 277(51), 49175-49185.
Rose, P., Moore, P. K., & Zhu, Y. Z. (2017). H₂S biosynthesis and catabolism: new insights from molecular studies. Cellular and Molecular Life Sciences, 74(8), 1391-1412.
Livshits, L., Chatterjee, A. K., Karbian, N., Abergel, R., Abergel, Z., & Gross, E. (2017). Mechanisms of defense against products of cysteine catabolism in the nematode Caenorhabditis elegans. Free Radical Biology and Medicine, 104, 346-359.
Qabazard, B., Li, L., Gruber, J., Peh, M. T., Ng, L. F., Kumar, S. D., ... & Moore, P. K. (2014). Hydrogen sulfide is an endogenous regulator of aging in Caenorhabditis elegans. Antioxidants & redox signaling, 20(16), 2621-2630.
Ng, L. F., Ng, L. T., van Breugel, M., Halliwell, B., & Gruber, J. (2019). Mitochondrial DNA damage does not determine C. elegans lifespan. Frontiers in genetics, 10, 311.
Kimura, H. (2020). Hydrogen sulfide signaling in the central nervous system-Comparison with nitric oxide. Authorea Preprints.
Cavanaugh, C. M., McKiness, Z. P., Newton, I. L., & Stewart, F. J. (2006). Marine chemosynthetic symbioses. The prokaryotes, 1, 475-507.
Grabherr, M.G., Haas, B.J., Yassour, M., Levin, J.Z., Thompson, D.A., Amit, I., et al. (2011) Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nat Biotechnol 29: 644–52
Cerveau, N., and Jackson, D.J. (2016) Combining independent de novo assemblies optimizes the coding transcriptome for nonconventional model eukaryotic organisms. BMC Bioinformatics 17: 525
Fu, L., Niu, B., Zhu, Z., Wu, S., and Li, W. (2012) CD-HIT: Accelerated for clustering the next-generation sequencing data. Bioinformatics 28: 3150–3152
Simão, F.A., Waterhouse, R.M., Ioannidis, P., Kriventseva, E. V., and Zdobnov, E.M. (2015) BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs. Bioinformatics 31: 3210–3212
Huson, D.H., Auch, A.F., Qi, J., and Schuster, S.C. (2007) MEGAN analysis of metagenomic data. Genome Res 17: 377–386.
Bryant, D.M., Johnson, K., DiTommaso, T., Tickle, T., Couger, M.B., Payzin-Dogru, D., et al. (2017) A Tissue-Mapped Axolotl De Novo Transcriptome Enables Identification of Limb Regeneration Factors. Cell Rep 18: 762–776
Li, B., & Dewey, C. N. (2011). RSEM: accurate transcript quantification from RNA-Seq data with or without a reference genome. BMC bioinformatics, 12(1), 1-16.
Gentleman, R. C., Carey, V. J., Bates, D. M., Bolstad, B., Dettling, M., Dudoit, S., ... & Zhang, J. (2004). Bioconductor: open software development for computational biology and bioinformatics. Genome biology, 5(10), 1-16.
Robinson, M. D., McCarthy, D. J., & Smyth, G. K. (2010). edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics, 26(1), 139-140.
Team, R. C. (2013). R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria.
Rapaport, F., Khanin, R., Liang, Y., Pirun, M., Krek, A., Zumbo, P., ... & Betel, D. (2013). Comprehensive evaluation of differential gene expression analysis methods for RNA-seq data. Genome biology, 14(9), 1-13.
Cantalapiedra, C. P., Hernández-Plaza, A., Letunic, I., Bork, P., & Huerta-Cepas, J. (2021). eggNOG-mapper v2: Functional Annotation, Orthology Assignments, and Domain Prediction at the Metagenomic Scale. Molecular Biology and Evolution. msab293.
Huerta-Cepas, J., Szklarczyk, D., Heller, D., Hernández-Plaza, A., Forslund, S. K., Cook, H., ... & Bork, P. (2019). eggNOG 5.0: a hierarchical, functionally and phylogenetically annotated orthology resource based on 5090 organisms and 2502 viruses. Nucleic acids research, 47(D1), D309-D314.
Buchfink, B., Reuter, K., & Drost, H. G. (2021). Sensitive protein alignments at tree-of-life scale using DIAMOND. Nature methods, 18(4), 366-368.
Altschul, S. F., Gish, W., Miller, W., Myers, E. W., & Lipman, D. J. (1990). Basic local alignment search tool. Journal of molecular biology, 215(3), 403-410.
Harris, T. W., Arnaboldi, V., Cain, S., Chan, J., Chen, W. J., Cho, J., ... & Sternberg, P. W. (2020). WormBase: a modern model organism information resource. Nucleic Acids Research, 48(D1), D762-D767.

No competing interests reported.

DataS1.xlsx
Data S1. Ca. T. oneisti genes, functional annotations, transcript and protein expression.
DataS2.xlsx
Data S2. Top 100 expressed genes (RNA-Seq) and detected proteins (proteomics data).
Supplementalmovie16daysanoxia.mov
Supplemental video 1. A batch of 50 Laxus oneistus after 6 days in anoxic seawater.
Supplementalmovie2anoxiaT0.mov
Supplemental video 2. A batch of 50 Laxus oneistus at the beginning (T0) of the incubations.
Supplementalmovie3anoxiaT24.mov
Supplemental video 3. A batch of 50 Laxus oneistus after 1 day (T24 h) in anoxic sulfidic seawater (0 % air saturation, 25 µM H2S).
Supplementalmovie4oxicT24.mov
Supplemental video 4. A batch of 50 Laxus oneistus after 1 day (T24 h) in oxic seawater (87 % air saturation, 0 µM H2S).
FigureS1Experimentalsetup070721.pdf
Figure S1. Experimental conditions, sample similarity and differential expression. (A) Experimental setup was previously described (Paredes et al. 2021). Briefly, nematodes were subjected to different oxygen concentrations for 24 h: anoxic with sulfide (AS: 0mM O2, 25mM sodium sulfide added), anoxic without sulfide (A, 0mM O2), hypoxic (H, 60mM O2 after 24 h), and oxic (O, 100mM O2 after 24 h). The box around the anoxic incubation vials illustrates that these incubations were carried out in a polyethylene glove bag. (B) Similarity between transcriptome samples based on Euclidean distances between expression values (log2TPM), and visualized by means of multidimensional scaling (C) Differential gene expression (DE) analysis between incubations showed that the number of DE genes was low (maximum value was 4.8% of all expressed genes for the H vs AS conditions). Genes were considered differentially expressed if their expression changed 1.5-fold with a false-discovery rate (FDR) of ≤ 0.05.
FigureS2Top100Proteome070721.pdf
Figure S2. Statistical analysis, relative transcript abundance and expression levels of the top 100 detected proteins of L. oneistus across all conditions. (A) Relative protein abundance (%) of the top 100 detected proteins present in a particular manually curated functional category. The top 100 proteins were collected by averaging the expression values across all replicates of all incubations (Figure S1A, Data S2). Functional classifications were extracted from the curated database UniProt and from comprehensive literature search focused mainly on C. elegans, and confirmed with the automatic annotated eggNOG classification (Data S1). (B) Median gene expression levels of selected L. oneistus manually annotated functional categories of the top 100 expressed proteins. Each dot represents the average %cOrgNSAF per protein across all replicates of all incubations. Notice that some categories were created with genes of overlapping functions (e.g., cytoskeleton/locomotion/nervous system). All protein names (or locus tags for unidentified protein names) are listed in Data S2.
FigureS3ProteomevstranscriptomeFunctionalcategories.pdf
Figure S3. Transcriptomics vs proteomics comparison. Pearson correlation between all transcripts and proteins (Data S1) automatically classified based on their functional category. The Pearson correlation between all expressed transcripts and all detected proteins (r = 0.4) was found to be low (Figure S3).
FigureS4top100Olavius.png
Figure S4. Relative transcript abundance and expression levels of the top 100 expressed genes of O. algarvensis across all conditions. (A) Relative transcript abundance (%) of the top 100 expressed genes with a manually curated functional category. The top 100 expressed genes were collected by averaging the expression values (log2TPM) across all replicates of all incubations (see Supplemental material). Functional classifications were extracted from the curated database UniProt and from comprehensive literature search focused mainly on C. elegans). (B) Median gene expression levels of selected O. algarvensis manually annotated functional categories of the top 100 expressed genes. Metabolic processes include both differentially and constitutively expressed genes. Each dot represents the average log2TPM value per gene across all replicates of all incubations.
FigureS5.L.oneistuslipidcomposition.pdf
Figure S5. L. oneistus lipid composition in anoxic and oxic conditions after 24 h. Major lipid classes and their abundance relative to all lipids detected showed no statistical difference between both conditions. For details on methodology see Supplemental material.
TableS1.HostpaperMetabolomicsdata.xlsx
Table S1. Metabolites detected in at least two biological replicates of either the holobiont fraction (Laxus oneistus and its ectosymbiont) or in the symbiont fraction (see Supplemental material). RT: retention time. Area: area of a peak from a specific compound detected in the GC-MS chromatograms. Grey boxes: no metabolites detected. Blank boxes: Unknown metabolites that are either below the detected threshold (< 700) or might be products of derivatization reagents. Note that cholestane and ribitol were used as internal standards.

Download PDF

Editorial decision: Major revision
15 Mar, 2022
Reviews received at journal
14 Mar, 2022
Reviews received at journal
05 Feb, 2022
Reviewers agreed at journal
18 Jan, 2022
Reviewers agreed at journal
18 Jan, 2022
Reviewers invited by journal
18 Jan, 2022
Editor assigned by journal
18 Jan, 2022
Editor invited by journal
27 Nov, 2021
Submission checks completed at journal
27 Nov, 2021
First submitted to journal
23 Nov, 2021

You are reading this latest preprint version

Differential Regulation of Degradation and Immune Pathways Underlies Adaptation of the Ectosymbiotic Nematode Laxus Oneistus to Oxic-Anoxic Interfaces

Status:

Version 1

Abstract

Figures

Introduction

Results And Discussion

Conclusions

Declarations

ACKNOWLEDGEMENTS

MATERIALS AND METHODS

References

Additional Declarations

Supplementary Files

Status:

Version 1