Abstract
Extracellular vesicles (EVs), entities transporting a variety of cargo, are directly involved in many biological processes and intercellular communication, the characterization of which requires studying multi-tissue organisms. We previously demonstrated that the largest evolutionarily conserved EVs, exophers, are a component of the C. elegans maternal somatic tissue resource management system, and their formation is induced by the embryos developing in utero. In this study, we explored inter-tissue regulatory networks of exophergenesis. We found that exophergenesis activity is differentially modulated by sex-specific ascaroside (pheromones) signaling molecules, known to have multiple functions in development and behavior. While hermaphrodite-released pheromones down-regulate exophergenesis, male- released pheromones favor strong exopher production. This ascaroside-dependent regulation is fine-tuned by exopher-promoting olfactory neurons exposed to the environment and exopher- inhibiting sensory neurons exposed to the body cavity. Therefore, we uncovered critical control nodes for muscle exophergenesis in response to environmental and internal conditions. Our findings may imply the existence of an analogous mechanism regulating cardiomyocyte exophers, which contributes to the olfactory dysfunction-dependent risk of cardiovascular disease in humans.
Main
Extracellular vesicles (EVs) are lipid-bilayer-enclosed particles that most cell types release. Two major EV types can be distinguished based on their biogenesis: endosome-derived exosomes and membrane-derived ectosomes1. EVs can be employed by cells to remove unwanted biological material, such as misfolded proteins and damaged organelles, or to transport small molecules, including proteins and nucleic acids, enabling exchange and communication between cells. Therefore, they are critical in physiological processes and pathological states involving disrupted cellular homeostasis2–5. To study EVs’ genesis, content, and function, multicellular animal models are frequently employed. The nematode C. elegans has been successfully used to investigate the biology of EVs generated by various tissues, including neurons6, 7, muscles8, hypodermis9, and reproductive system10, 11. Here, as a model, we use a class of recently discovered EVs, termed exophers, to understand how ectosome biogenesis in somatic tissue is regulated at the whole organism level employing worms. Exophergenesis (i.e., exopher generation) is an evolutionarily conserved phenomenon found from invertebrates to mammals, including humans. Exophers were shown to play a significant role in cellular stress response, tissue homeostasis, and organismal reproduction7, 8, 12, 13. It was demonstrated that C. elegans neurons retain their regular activity in the face of proteotoxic stress by expelling protein aggregates, damaged mitochondria, and lysosomes into surrounding tissues via exophers7. Nicolás-Ávila et al. also demonstrated that mouse cardiomyocytes excrete defective mitochondria via exophers, which, in turn, restricts waste accumulation in the extracellular space and inflammasome activation, promoting metabolic homeostasis in the heart13. However, the biological roles of exophers extend beyond the elimination of superfluous cellular components. In our previous work, we showed that the body wall muscles (BWM) of C. elegans release exophers that can transport muscle-synthesized yolk proteins to support offspring development, increasing their odds of adapting to environmental conditions8. We do not, however, comprehend the mechanism of cell non-autonomous regulation of muscle exophergenesis nor how this maternal somatic tissue resource management system responds to environmental conditions.
Animal-to-animal signals transmitted by pheromones in C. elegans have been shown to regulate maternal provisioning, development, and generation time 14–16. Since muscle exophers mediate the transfer of maternal resources to offspring supporting their development, we hypothesized that exophergenesis (Fig. 1a) is regulated by metabolites-mediated social cues generated within the worm population. To investigate this, we cultured hermaphrodites on plates under two conditions and quantified exopher production by BWM. As the first condition, we used hermaphrodites cultured individually on a plate, thereby eliminating social cues from other worms. As a second condition, we used worms raised in a ten-hermaphrodites population from the beginning of their development (Fig. 1b). We noted that animals grown together in a ten-hermaphrodite population released, on average, 44% fewer exophers than single-grown worms (Fig. 1c). Hermaphrodites from both experimental groups contain the same number of embryos in utero (Fig. 1d), demonstrating that signaling from other hermaphrodites can modulate exopher production independently from previously postulated embryo-maternal signaling8. Moreover, growing hermaphrodites on plates with different population densities indicate a dose-dependent effect (Extended Data Fig. 1a). To rule out the possibility that exophergenesis is substantially regulated by the molecules derived from the bacterial food source, which could indirectly influence animal to animal communication, we decided to test various bacterial strains effect on worm’s muscle exopher production. We directly compared Escherichia coli B strain OP50 and K-12 strain HT115, which are widely used in C. elegans culture17 and RNAi silencing experiments18, respectively. As a result, we observed a slight increase in exopher number upon the E. coli HT115 diet compared to the OP50 diet (Extended Data Fig. 1b). However, the number of eggs present in utero at adulthood day 2 was elevated upon the HT115 diet (Extended Data Fig. 1c). Whether or not the bacteria were metabolically active (PFA-killed prior to plate culture) was irrelevant (Extended Data Fig. 1b), confirming that exophergenesis is robustly activated in worm’s muscles regardless of the bacteria type used as a food source.
Next, we investigated if the presence of males influences exophers production by hermaphrodites similarly to the presence of other hermaphrodites. To verify this, we monitored the number of exophers in him-5 mutants characterized by a significant increase of males in the population (about 33% compared to 0.3% for wild type)19. Interestingly, him-5 animals grown with males until the L4 stage and then transferred to a male-free plate (Fig. 1e) produce approximately 2.5 times more exophers than wild-type hermaphrodites grown from L1 on a male-free plate (Fig. 1f). This increase appears to be mediated by the embryo-maternal signaling as him-5 mutant hermaphrodites contain 26% more embryos in utero than wild-type hermaphrodites (Fig. 1g). To rule out the possibility that an increase in the number of exophers released, may be a result of the him-5 mutation rather than the presence of males in the population, we grew wild-type hermaphrodites on a plate conditioned with males for 48 hours, which we then removed (Fig. 1h). Growing hermaphrodites on male-conditioned plates increased exopher production to the same degree as when hermaphrodites were grown with males until the L4 larvae stage (Fig. 1i), regardless of the bacteria strain used as a food source (Extended Data Fig. 1d). However, this increase in exophers production was associated with a rise in the number of in utero embryos (Fig. 1j), indicating that C. elegans male pheromones can also drive embryo retention in hermaphrodite’s uterus. Furthermore, adult hermaphrodites exposed to males’ secretions as larvae showed no further increase in exopher production (Extended Data Fig. 1e). Our data indicate that exophers generation by hermaphrodite BWM is modulated by signals released in response to pheromonal stimulation. Male pheromones act through embryo-maternal signaling, while pheromones released by hermaphrodites downregulate exophergenesis independently from this pathway.
To further investigate muscle exopher regulation by pheromones, we took advantage of several worm mutants with altered ascaroside pheromones side-chain biosynthesis20 (Fig. 2a). Mutants of the maoc-1 gene display a reduction in exopher production, whereas the daf-22 and acox-1 mutants show an increase in exopher production (Fig. 2b). However, changes in exophergenesis levels in mutants correlate with the number of embryos present in their uterus (Fig. 2c), therefore, could be dependent on the embryo-maternal signaling. To distinguish the change in exopher production from embryo-maternal signaling in the mutants mentioned above, we examined exophergenesis in wild-type animals maintained on plates with ascaroside biosynthesis mutants (one wild-type worm with nine mutant worms per plate) (Fig. 2d). The number of embryos in the uterus of wild-type animals reared in this way did not change (Fig. 2f), yet we observed a decrease in exophergenesis in wild-type worms grown together with maoc-1 mutants and an increase in the presence of acox-1 worms (Fig. 2e). Finally, we isolated C. elegans secretory products from starving N2 wild-type and maoc-1 mutant populations as previously described21. Next, we exposed growing worms to this isolate (Fig. 2g), which led to the increase in exophergenesis in a maoc-1-dependent manner (Fig. 2h). These results further confirm pheromone-based regulation of exopher formation with a critical role for ascarosides whose synthesis is mediated by MAOC-1.
Since many olfactory neurons detect ascarosides22 (Fig. 3a), we examined whether genetic ablation of all ciliated sensory neurons would abolish pheromone-regulated exopher induction. Indeed, as shown in Fig. 3b, che-13 mutants, which do not form proper cilia and are incapable of pheromone detection23, produce a minimal number of exophers. Furthermore, exposure of the che-13 mutant to metabolites secreted by starving wild-type worms did not affect exopher production (Fig. 3c). To determine which sensory neurons mediate pheromone-dependent modulation of exophergenesis, we examined its level in animals with genetically ablated sensory neurons previously shown to be capable of pheromone detection22. The removal of ASK, ADL, or AWC neurons inhibited exopher production comparable to that observed in che-13 mutants, whereas the ablation of ASH led to a slight decrease in exophergenesis. Notably, hermaphrodites with the addition of male-specific, pheromone-sensing CEM neurons (via ceh-30 gain-of-function mutation24) did not show alterations in exopher production (Fig. 3d).
To identify neurons critical for the exophergenesis downregulation in response to the presence of other hermaphrodites, we grew hermaphrodites with genetic ablations of different classes of olfactory neurons either as a single animal or in a ten-worms population (Fig. 1b scheme). Our analyses using mutant strains with impaired olfaction showed that ASH and ASI neurons play a crucial role in exophergenesis, with a small modulation coming from ASK neurons (Fig. 3e). Finally, masculinization of a hermaphrodite olfactory circuit by the addition of CEM male- specific neurons leads to a decrease in exopher production in solitary animals (Fig. 3e).
Next, we investigated which olfactory neurons are necessary to detect male pheromones responsible for exophergenesis upregulation. To this end, we grew hermaphrodites with genetic ablations of different classes of olfactory neurons on male-conditioned plates (as shown in Fig. 1h scheme). Our results demonstrate that the removal of ASK, AWB, or ADL neurons abolishes an increase in exopher production induced by male-released pheromones (Fig. 3f). These results align with previously described roles for ASK, AWB, and ADL neurons in male pheromones sensing25–28.
Since genetic ablation of AWC thermo-responsive neurons29, 30 has a prominent effect on exophergenesis (Fig. 3d), we also investigated whether this process could be temperature- dependent. To assess this, we grew worms at 15, 20, or 25°C and measured the number of exophers produced by hermaphrodites that were at the peak of progeny production for each temperature31. We observed that the generation of muscle exophers increased as worm incubation temperature rises (Fig. 3g). Moreover, the removal of AWC aggravates this response, demonstrating temperature-dependent regulation of exophergenesis by these neurons (Fig. 3g).
The binding of signaling molecules to the relevant receptor is the first step in transducing neuron chemosensory signals. More than 1,300 G protein-coupled receptors (GPCRs) mediate this communication in C. elegans32. Internal states and environmental conditions can modulate GPCRs expression to affect worm behavior33–36. To identify the receptor(s) responsible for ascaroside-mediated alterations in exopher formation, we performed RNA-sequencing of animals grown either as a single animal or in a ten-hermaphrodites population (Extended Data Fig. 2a). On the transcriptional level, neither group differed markedly (Extended Data Fig. 2b), and none of the 7TM receptors was significantly up- or down-regulated (Supplementary Table 1). However, str-173 receptor transcript, which shows among all of the 7TM receptors one of the most evident trends between growth conditions (2.3 fold change) is, according to single- cell RNA-seq data37, expressed almost exclusively in ASK neurons (Extended Data Fig. 2c). Since ASK is crucial for pheromone-mediated modulation of exopher formation, we investigated the str-173 role in this pathway.
The wrmScarlet CRISPR/Cas9-mediated knock-in for the str-173 gene confirmed its strong expression in ASK neurons and revealed additional expression in OLQ neurons, the pharynx, vulva muscles, and the tail (Extended Data Fig. 3a-b). Next, we created str-173 null mutants again using CRISPR/Cas9 editing (Extended Data Fig. 3c) and observed that the basal level of exophergenesis was comparable to the wild-type control (Extended Data Fig. 3d). However, exophergenesis was lower for str-173 null mutants grown as a single animal on their own plate compared to the wild-type control (Extended Data Fig. 3e). Moreover, str-173 mutants exhibit a smaller increase in exopher production in response to male pheromones (Extended Data Fig. 3f). This finding suggests that STR-173 plays a role in the signal processing that occurs in response to the metabolites secreted by other worms.
Among the 118 classes of neurons in C. elegans, only four are directly exposed to the pseudocoelomic cavity38. Three classes of these neurons, AQR, PQR, and URX regulate social feeding in worms39. Given that exophers are released to the worm’s pseudocoelomic cavity and are regulated by social cues, we hypothesized that AQR, PQR, and URX might play a role in exophergenesis regulation. To test for that, we first investigated the effect of genetic ablation of AQR, PQR, and URX neurons on exopher production. Indeed, the removal of these neurons leads to a substantial increase in exophergenesis (Fig. 4a). Notably, the increased number of exophers generated by worms with genetically ablated AQR, PQR, and URX neurons is not the result of embryo-maternal signaling as these animals contain an even lower number of eggs in utero (Fig. 4b) and have a smaller brood size (Extended Data Fig. 4a-b) than wild-type control.
To further validate the role of AQR, PQR, and URX neurons in the regulation of exophergenesis, we optogenetically inactivated or activated them using ArchT40 or ReaChR41, respectively, and compared the number of exophers before and after the stimulus. We observed that 60 min of AQR, PQR, and URX neuron inactivation leads to a significant increase in exopher release (Fig. 4c and Extended Data Fig. 4c). On the other hand, 60 min of AQR, PQR, and URX neuron activation resulted in a significant decrease in exopher release after the stimulus was completed (Fig. 4d and Extended Data Fig. 4d).
Our data also demonstrates that opposing exophergenesis phenotypes in ASK-ablated animals and in worms with the genetic elimination of AQR, PQR, and URX are counterbalanced in worms with all four neuron classes removed (Fig. 4e). Finally, we show that AQR, PQR and URX participate in mediating response to hermaphrodite pheromones (Fig. 4f) but not to male pheromones (Fig. 4g) and their high activity is not sufficient to overcome the critical role of embryo-maternal signaling in exopher production (Fig. 4h-i). Altogether our data indicate that the volatile signals lowering exopher levels secreted by hermaphrodites act through the olfactory system and partly via AQR, PQR, and URX neurons. The male secretions detected by olfactory neurons, in turn, potentiate exophergenesis by promoting embryo accumulation in utero, and this triggers pro-exopher signals independent of the activity of AQR, PQR, and URX neurons (Fig. 4j).
Discussion
Sensory neurons, through various types of metabolites, tune the functionality of somatic tissues to environmental conditions. We demonstrated that this also applies to the production of exopher vesicles by muscles. Our data show that, depending on the type of ascaroside pheromone, exophergenesis can be up- or down-regulated within the C. elegans hermaphrodite population. This communication system enables individuals to optimally utilize their muscle resources for reproductive purposes depending on the biological and environmental context. Accordingly, the higher level of exophers in hermaphrodites exposed to male metabolites or mated with males is consistent with the idea that male pheromones promote resource allocation to the germline42, and exophers may contribute to oocyte/embryo quality. However, increasing exophergenesis accelerates the age-related deterioration in worm muscle function8. This is also in accordance with Aprison and Ruvinsky’s observations that hermaphrodites hasten development and somatic aging in the presence of males43. As such, exophers probably act as biological executors and carriers of information in inter-animal communication. This is also consistent with observations that Drosophila males secrete extracellular vesicles that are important for mating behavior44, and the exchange of information between male and female flies leads to increased EV release from sperm secretory cells, which promotes fertility45. Therefore, this bidirectional regulation of physiological processes related to EV-mediated communication between animals appears to be evolutionarily conserved. In this regard, sensory neurons exposed to the body cavity are also anticipated to play a role. We showed that C. elegans AQR, PQR, and URX neurons, which monitor the metabolic state of the animal, transmit neuroendocrine signals to downregulate exophergenesis. Interestingly, these neurons belong to the class of ciliated neurons and their counterparts exposed to the environment are capable of extracellular vesicle production6. Therefore, it is tempting to speculate that AQR, PQR, and URX could transmit biological information within the worm body using extracellular vesicles released to the pseudocoelomic cavity, which could assist classical neurotransmitters or neuropeptides. However, additional research is required to establish how this class of ciliated neurons suppresses exophers formation/release by BWM.
Using C. elegans as a model, we showed that exopher production in one tissue is controlled by signals from another. Similar programs that enable fine-tuning of cell-to-cell communication based on exophers facilitating efficient use of resources in multiple tissues may work in humans, as confirmed by studies on other classes of EVs46. Moreover, our results demonstrate the worm’s olfactory system’s importance in regulating exopher formation in muscles. Notably, olfactory dysfunction in humans has been shown to lead to an increased likelihood of cardiovascular disease (CVD)47, 48. This could result from faulty regulation of exophergenesis in the heart, which helps maintain homeostasis during cardiac stress. However, additional research is required to determine whether mammals possess a comparable exopher control system depending on the olfactory network.
Figures and figure legends
Materials and Methods
Reagents and Tools table
Methods and Protocols
Worm maintenance and strains
Worms were maintained on nematode growth medium (NGM) plates seeded with bacteria Escherichia coli OP50 or HT115 at 20°C unless stated otherwise17 A list of all C. elegans strains used in the study is provided in the table at the beginning of the “Materials and Methods” section.
Scoring exophers and fluorescence microscopy
For the assessment of exophers number, the protocol described in Turek et al. and Banasiak et al. (Bio-Protocol, detailed, step-by-step guide) was applied. Briefly, exophers were scored using stereomicroscope Zeiss Axio Zoom.V16 with filter sets 63 HE and 38 HE. For each exopher scoring assay, worms were age-synchronized from pretzel-stage embryos, L1 larvae, or L4 larvae. On adulthood day-2, animals were visualized on NGM plates, and the number of exophers was counted in each freely moving worm.
The representative pictures presented in the manuscript were acquired with an inverted Zeiss LSM800 laser-scanning confocal microscope with a 40x oil immersion objective. To excite the GFP and RFP fluorescent proteins, 488- and 561-nm lasers were used. For visualization, animals were immobilized on 3% agarose pads with 6µl of PolySciences 0.05 µm polystyrene microspheres or 25 µM tetramisole.
RNA interference assay
RNA interference in C. elegans was performed using the standard RNAi feeding method and RNAi clone49, previously described in Turek et al., 20218. Briefly, NGM plates, supplemented with 12,5µg/ml tetracycline, 100µg/ml ampicillin and 1mM IPTG, were seeded with HT115 E. coli bacteria containing dsRNA against the gene of interest. Control group animals were fed with bacteria containing the empty vector. Age-synchronized pretzel-stage embryos or L1 larvae were placed on freshly prepared plates and cultured until day 2 of adulthood. The age of the worms was verified at the L4 larvae stage, either younger or older worms were removed from the experiment.
Pre-conditioning of the plates with C. elegans males
50 males from the strain with him-5(e1490) mutation were transferred on a fresh, 35 mm NGM plate with E. coli OP50 or HT115 at the L4 developmental stage and older. After 48 hours, males were removed from the plates. L1 larvae were transferred to plates pre-conditioned with males in a group of 10 animals per plate. For the control group, L1 larvae were transferred to fresh, 35 mm NGM plates seeded with E. coli OP50 or HT115 without pre-conditioning with males. Worms were cultured up to adulthood day 2 when muscle exophers were counted in each animal.
Egg retention assay
Firstly, single worms were immobilized with 25 μM tetramizole on an NGM, bacteria-free plate. Secondly, using the stereomicroscope, muscle exophers were counted in each worm. In the following step, hermaphrodites were exposed to 1.8% hypochlorite solution. When they dissolved, embryos retained in the uterus were counted.
Metabolic inactivation of bacterial food source
The preparation of plates with a metabolically inactive food source for the worms to determine its effects on exophergenesis was done according to the protocol described in Beydoun et al., 2021. Briefly, a single colony of E. coli HT115 or OP50 was inoculated overnight and then the bacterial culture was split into two flasks. Next, paraformaldehyde (PFA) was added to a final concentration of 0.5% in only one of them, and the flask was placed in the 37°C shaking incubator for 1 h. Afterward, the aliquots were transferred to 50 mL tubes and centrifuged at approximately 3000 × g for 20 min and washed with 25 mL of LB five times. Control and PFA- treated bacteria were later concentrated accordingly and seeded on the NGM plates.
As control of PFA treatment, new bacterial cultures in LB were set in the 37°C shaking incubator overnight to make sure the replication was blocked and there was no bacterial growth. Plates were used for experiments at least 5 days after their preparation to make sure there are no bacteria-derived metabolites left on plates with PFA-treated E. coli. A new batch of bacterial food source was prepared for each biological repetition.
Brood size quantification
Age synchronized worms were transferred on fresh, 60mm NGM plates with E. coli OP50 or HT115 at the L4 developmental stage, single or two hermaphrodites on each of ten plates per biological repeat. Worms were transferred to fresh plates every day since adulthood day 1. The number of eggs laid over the worms’ reproductive lifetime was counted manually every day. The data is presented as the total number of eggs laid by each animal or an average of the total number of eggs laid by each pair.
Quantifying number of exophers in worms grown in different temperatures
Worms were confronted with a range of physiological temperatures: low – 15°C, optimal –20°C, and high – 25°C throughout their development until exophers were scored. To compare the corresponding stages of development at various temperatures, worms were additionally sorted at the L4 stage. The exopher number was assessed based on the timepoint of maximal egg laying, approx. 140 h or 78 h from egg hatching at low or high temperatures of maintenance, respectively. Calculations of timing were based on the C. elegans development timeline at different temperatures31.
Culturing worms in different population sizes
L1 larvae were transferred to 35 mm Petri dishes seeded with E. coli OP50 or HT115 bacteria strains (100 μL of bacteria from 10 mL overnight culture grown in 50 mL Erlenmeyer flask) immediately after hatching as a single larva or in a group of 5, 10 or 100 animals. In total, thirty plates with single worms, six plates with 5 worms, three plates with 10 worms, and one plate with 100 worms were used per biological repeat.
Quantifying the influence of molecules released by ascaroside biosynthesis mutant on exopher production in wild-type worms
Nine freshly hatched L1 larvae from selected ascaroside biosynthesis mutant strains (maoc-1(ok2645), daf-22(ok693), or acox-1(ok2257)) or wild-type hermaphrodites (as a control) were transferred to fresh NGM plates. Next, one L1 larva of a reporter strain expressing RFP in BWM was added to each plate and 10 worms in total were grown together.
Isolation of excretory-secretory (ES) metabolites containing ascarosides
Liquid-culture protocol for synchronous dauer formation from Cell Press STAR PROTOCOLS was applied step-by-step (Hibshman et al., 2020). Larvae enter dauer diapause by day 4-5 and dauers can be maintained for min. 40 days without a decrease in viability or the ability to recover. For the convenience of experimental conduct, wild-type and maoc-1(ok2645) larvae were maintained in liquid culture for 11 days before the isolation of excretory-secretory products.
On day 11 of liquid culture, worms were washed several times with S-medium to remove any possibly remaining debris and finally concentrated to approx. 30,000 dauers/mL. They were then rinsed three times with water and incubated for one hour on an orbital shaker at room temperature. After one hour they were centrifuged as described in Kaplan et al.21 and the liquid with worm excretory-secretory products was filtered on ice through a 0.22 µm filter using a sterile 1 mL syringe. Extracts were immediately aliquoted per 20 µL, frozen and stored at -80°C.
Assay with usage of excretory-secretory (ES) metabolites containing ascarosides
Approx. 40 animals were transferred to 60 mm NGM plates seeded with E. coli HT115 immediately after hatching. Then, 40 μL of excretory-secretory metabolites were applied on the bacterial lawn and wrapped with parafilm right afterwards. In control plates, water was used instead. Worms’ synchronous growth was monitored at the L4 stage when an additional 20 μL of ES metabolites or water was administered.
Generation of str-173 mutant strains
The str-173 gene mutants (str-173(wwa1) and str-173(wwa2)) were generated using CRISPR/Cas9 method as previously described50. The crRNA sequence used was ATAATTGGTGGATATACAAATGG. The str-173 gene locus was sequenced and deletions were mapped to the first exon (Extended Data Figure 3d). Both mutations cause frame shifts, therefore, are most likely molecular null alleles.
Generation of optogenetic strains
Optogenetic strains created for this paper contain red-shifted Channel Rhodopsin (ReaChR) or archaerhodopsin from Halorubrum strain TP009 (ArchT). To generate these strains, firstly, mKate2-unc-54 3’UTR was amplified from the template and cloned into pCG150 to create pAZ03 plasmid. Next, gcy-36 promoter was amplified from pMH389, ReaChR and ArchT were amplified from respective templates. The gcy-36 promoter was then cloned into pAZ03 plasmid with ReaChR and ArchT separately. As a result two plasmids were created: gcy-36 promoter::ReaChR::mKate2-unc-54 3’UTR in pCG150 and gcy-36 promoter::ArchT::mKate2-unc-54 3’UTR in pCG150. To verify the correct sequence of the cloned constructs, plasmids were sequenced. All constructs generated for this study were made using the SLiCE method51.
Transgenic strains with extrachromosomal arrays were generated by microinjection. DNA was injected into exopher reporter strain worms with muscle exopher RFP and mitochondrial GFP marker (ACH93). For injection, DNA was prepared as follows: construct 90 ng/µL and co- injection marker 10 ng/µL. Positive transformants were selected according to the presence of co-injection markers (myo-2 promoter::mNeonGreen).
The constructs created for this project and primers used for amplification are listed in the table at the beginning of the “Materials and Methods” section.
Optogenetics assay
For optogenetic activation or inhibition, 35-mm NGM plates seeded with HT115 E. coli bacteria were covered with 0.2 µM all-trans retinal (ATR). Control plates were not covered with ATR. Ten age-synchronized worms from optogenetic strains (expressing ReaChR or ArchT in AQR/PQR/URX neurons) were picked per plate at adulthood day 1. After 24-hour incubation at 20°C and darkness, muscle exophers extruded by worms were counted. Next, experimental plates were placed on the stereomicroscope and illuminated for 1 hour with green light (HXP 200C illuminator as a light source, band-pass filter Zeiss BP 572/25 (HE), the green light intensity measured at 561 nm = 0.07 mW/mm2). Immediately after illumination muscle exophers were counted. Subsequently, exophers scorings were performed in 15 minutes intervals, and worms were kept in the darkness in between counts. The control group was not illuminated. To provide similar environmental conditions control plates were placed next to the experimental plate but shielded from light. Control and treated groups were randomized before the start of the experiment.
FUdR assay
Age-synchronized young adult animals (day 0) were placed on NGM plates containing 25 μM fluorodeoxyuridine (FUdR) or control NGM plates without FUdR. Exophers number were scored when worms reached adulthood day 2 using a stereomicroscope.
Transcriptome analysis
RNA extractions, library preparations, and sequencing were conducted at Azenta US, Inc (South Plainfield, NJ, USA) as follows:
RNA Extraction
Total RNA was extracted using Qiagen RNeasy Plus mini kit following the manufacturer’s instructions (Qiagen, Hilden, Germany).
Library Preparation with polyA selection and Illumina Sequencing
Extracted RNA samples were quantified using Qubit 2.0 Fluorometer (Life Technologies, Carlsbad, CA, USA) and RNA integrity was checked using Agilent TapeStation 4200 (Agilent Technologies, Palo Alto, CA, USA).
RNA sequencing libraries were prepared using the NEBNext Ultra II RNA Library Prep Kit for Illumina following the manufacturer’s instructions (NEB, Ipswich, MA, USA). Briefly, mRNAs were first enriched with Oligo(dT) beads. Enriched mRNAs were fragmented for 15 minutes at 94 °C. First strand and second strand cDNAs were subsequently synthesized. cDNA fragments were end repaired and adenylated at 3’ends, and universal adapters were ligated to cDNA fragments, followed by index addition and library enrichment by limited-cycle PCR. The sequencing libraries were validated on the Agilent TapeStation (Agilent Technologies, Palo Alto, CA, USA), and quantified by using Qubit 2.0 Fluorometer (Invitrogen, Carlsbad, CA) as well as by quantitative PCR (KAPA Biosystems, Wilmington, MA, USA).
The sequencing libraries were multiplexed and loaded on the flowcell on the Illumina NovaSeq 6000 instrument according to the manufacturer’s instructions. The samples were sequenced using a 2x150 Pair-End (PE) configuration v1.5. Image analysis and base calling were conducted by the NovaSeq Control Software v1.7 on the NovaSeq instrument. Raw sequence data (.bcl files) generated from Illumina NovaSeq was converted into fastq files and de- multiplexed using Illumina bcl2fastq program version 2.20. One mismatch was allowed for index sequence identification.
Sequencing Data Analysis
After investigating the quality of the raw data, sequence reads were trimmed to remove possible adapter sequences and nucleotides with poor quality using Trimmomatic v.0.36. The trimmed reads were mapped to the Caenorhabditis elegans reference genome available on ENSEMBL using the STAR aligner v.2.5.2b. The STAR aligner is a splice aligner that detects splice junctions and incorporates them to help align the entire read sequences. BAM files were generated as a result of this step. Unique gene hit counts were calculated by using feature Counts from the Subread package v.1.5.2. Only unique reads that fell within exon regions were counted.
After the extraction of gene hit counts, the gene hit counts table was used for downstream differential expression analysis. Using DESeq2, a comparison of gene expression between the groups of samples was performed. The Wald test was used to generate p-values and Log2 fold changes. Genes with adjusted p-values < 0.05 and absolute log2 fold changes > 1 were called as differentially expressed genes for each comparison.
Data analysis and visualization tools
The exophers were scored at adulthood day 2 using a stereomicroscope unless stated otherwise. Data analysis was performed using Microsoft® Excel® and GraphPad Prism 9 software. Graphical representation of data was depicted using GraphPad Prism 9.
Statistical analysis
No statistical methods were used to predetermine the sample size. Worms were randomly allocated to the experimental groups for all the data sets and experiments were performed blinded for the data sets presented in the following figures: Fig. 2b, e, f, h; Fig. 3b, c, f; Fig. 4f; Extended Data Fig. 3d, e, f. Non-Gaussian distribution of residuals was assumed and therefore nonparametric statistical tests were applied: Mann–Whitney (in comparison between two groups) or Kruskal-Wallis test with Dunn’s multiple comparisons test (in comparison between more than two groups). P-value < 0.05 is considered significant.
Acknowledgements
Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). We thank Henrik Bringmann and Lukas Kapitein for plasmids; Frank Schroeder for expert advice; Peter Askjaer, Henrik Bringmann, and Antonio Miranda Vizuete for discussions and comments on the manuscript; Zofia Olszewska, Marta Niklewicz, and Monika Woźniak for assistance with worms maintenance. Work in the M. T. Laboratory was mainly funded by a National Science Centre SONATA grant (2019/35/D/NZ3/04091) and additionally supported by a National Science Centre SONATA BIS grant (2021/42/E/NZ3/00358). Work in the W. P. Laboratory was funded by the Foundation for Polish Science co-financed by the European Union under the European Regional Development Fund (grant POIR.04.04.00-00-5EAB/18-00 to K.B., and W.P.), and additionally supported by the European Molecular Biology Organization (EMBO Installation Grant No. 3916 to K.B., and W.P.), and the Norwegian Financial Mechanism 2014-2021 operated by the Polish National Science Centre, Poland (project contract number 2019/34/H/NZ3/00691 to W.P.).