High Nutritional Conditions Influence Feeding Plasticity in Pristionchus pacificus and Render Worms Non‐Predatory

ABSTRACT

Developmental plasticity, the ability of a genotype to produce different phenotypes in response to environmental conditions, has been subject to intense studies in the last four decades. The self‐fertilising nematode Pristionchus pacificus has been developed as a genetic model system for studying developmental plasticity due to its mouth‐form polyphenism that results in alternative feeding strategies with a facultative predatory and non‐predatory mouth form. Many studies linked molecular aspects of the regulation of mouth‐form polyphenism with investigations of its evolutionary and ecological significance. Also, several environmental factors influencing P. pacificus feeding structure expression were identified including temperature, culture condition and population density. However, the nutritional plasticity of the mouth form has never been properly investigated although polyphenisms are known to be influenced by changes in nutritional conditions. For instance, studies in eusocial insects and scarab beetles have provided significant mechanistic insights into the nutritional regulation of polyphenisms but also other forms of plasticity. Here, we study the influence of nutrition on mouth‐form polyphenism in P. pacificus through experiments with monosaccharide and fatty acid supplementation. We show that in particular glucose supplementation renders worms non‐predatory. Subsequent transcriptomic and mutant analyses indicate that de novo fatty acid synthesis and peroxisomal beta‐oxidation pathways play an important role in the mediation of this plastic response. Finally, the analysis of fitness consequences through fecundity counts suggests that non‐predatory animals have an advantage over predatory animals grown in the glucose‐supplemented condition.

Some nematodes exhibit mouth‐form polyphenism resulting in different feeding strategies including predation. In this study, Piskobulu and co‐workers investigate the influence of glucose supplementation on mouth‐form expression and find that higher glucose levels render worms non‐predatory. Thus, nutritional status influences mouth‐form polyphenism in nematodes.

1. Introduction

Developmental plasticity allows organisms to alter their developmental trajectory to execute distinct phenotypes in response to environmental cues with polyphenisms representing the most remarkable form of plasticity by exhibiting environment‐sensitive alternative phenotypes without intermediate forms (West‐Eberhard ²⁰⁰³). In general, developmentally plastic responses can be observed at physiological, morphological, and behavioural levels, including life history traits of organisms. Many case studies have shown developmental plasticity to be ubiquitous in nature. It has been suggested to facilitate adaptation to novel or heterogenous environments and to play an important role in evolutionary diversification and the creation of novelty (Ghalambor et al. 2007; Pigliucci ²⁰⁰¹; West‐Eberhard ²⁰⁰³). Multiple environmental factors have been shown to influence polyphenisms including temperature, seasonal changes, and population density (see reviews; Simpson, Sword, and Lo ²⁰¹¹; Yang and Andrew Pospisilik ²⁰¹⁹). Specifically, nutrition is an important regulator of polyphenisms. Several studies in insects have highlighted the importance of nutritional conditions during development for the generation of discrete phenotypes, revealing genetic and epigenetic mechanisms involved (Casasa, Zattara, and Moczek ²⁰²⁰; Chandra et al. ²⁰¹⁸; Gotoh et al. ²⁰¹⁴; Kamakura ²⁰¹¹; Kucharski et al. ²⁰⁰⁸). For instance, caste polyphenism in honeybees and horn polyphenism in dung beetles represent some of the extreme cases of nutrition‐induced developmental plasticity (Emlen 1997; Haydak ¹⁹⁷⁰; Moczek ¹⁹⁹⁸). In addition, ant caste differentiation is long known to be regulated by nutrition and diet (Alvardo et al. 2014; Metzl, Wheeler, and Abouheif ²⁰¹⁸).

The nutritional composition of the diet can also influence plastic responses, such as mimicry in lepidopteran caterpillars (Greene 1989), migratory polyphenism in locusts (Cease et al. 2017), and wing polyphenism in aphids (Wang et al. 2024). Aside from insects, resource polyphenisms exhibited by rotifers and spadefoot toad tadpoles are also modulated by diet. For instance, trimorphism observed in Asplanchna species (Phylum Rotifera) can be regulated by dietary influences such as the type and size of prey (Gilbert 2017). Similarly, the developmental switch between the omnivore and carnivore morphs in spadefoot tadpoles in the genus Spea can be induced by the abundance of the prey (shrimp and other tadpoles) in the environment (Levis, de la Serna Buzón, and Pfennig ²⁰¹⁵). Moreover, the significance of the role of nutrition in developmental plasticity has also been investigated in mammals. In particular, studies in the agouti mice have revealed that nutritional supplementation‐induced changes in DNA methylation can regulate a developmentally plastic trait (fur colour), which has associated metabolic conditions (Dolinoy et al. 2006; Waterland and Jirtle ²⁰⁰³). Taken together, these studies indicate that investigations aimed at understanding the nutritional regulation of polyphenisms can elucidate novel mechanisms involved in developmental plasticity.

Besides the aforementioned organisms, nematodes provided substantial mechanistic insights into the regulation of polyphenisms in the last decade. In particular, the free‐living hermaphrodite Pristionchus pacificus has been a model system for the studies of developmental plasticity. Research in P. pacificus has provided insights into the genetic and epigenetic regulation as well as the ecological and evolutionary significance of plasticity (Schroeder 2021; Sommer ²⁰²⁰). In nature, P. pacificus can be found in soil and on scarab beetles (chafers, stag beetles, dung beetles), which, after their death, provide a nutritious environment for the nematode through proliferation of bacteria and fungi on the carcass (Herrmann, Mayer, and Sommer ²⁰⁰⁶; Meyer et al. ²⁰¹⁷; Ragsdale, Kanzaki, and Herrmann ²⁰¹⁵; Renahan et al. ²⁰²¹; Renahan and Sommer ²⁰²²). In contrast to Caenorhabditis elegans and most other free‐living nematodes, P. pacificus shows a remarkable mouth‐form dimorphism, which influences its dietary niche and behaviour. During postembryonic development, genetically identical P. pacificus worms form one of two, alternative morphs, the so‐called eurystomatous (Eu) and stenostomatous (St) mouth forms (Figure 1a) (Bento, Ogawa, and Sommer ²⁰¹⁰). While the Eu morph enables facultative predatory feeding on other nematodes by exhibiting two “teeth” and a wide mouth, the St morph leads to strict microbial feeding and forms only one tooth and a narrow mouth (Theska et al. 2020). Major components of the gene regulatory network that govern mouth‐form plasticity have been identified in this model (Bui, Ivers, and Ragsdale ²⁰¹⁸; Casasa, Katsougia, and Ragsdale ²⁰²³; Levis and Ragsdale ²⁰²³; Namdeo et al. ²⁰¹⁸; Ragsdale et al. ²⁰¹³; Serobyan et al. ²⁰¹⁶; Sieriebriennikov et al. ²⁰²⁰; Werner et al. ²⁰²³). For instance, a supergene locus with the sulfatase eud‐1 and two alpha‐N‐acetylglucosaminidase genes (nag‐1 and nag‐2), as well as the sulfotransferase, seud‐1/sult‐1, act as part of a main switch in mouth‐form determination (Bui, Ivers, and Ragsdale ²⁰¹⁸; Namdeo et al. ²⁰¹⁸; Ragsdale et al. ²⁰¹³; Sieriebriennikov et al. ²⁰¹⁸). They are involved in the regulation of the evolutionarily conserved nuclear hormone receptors nhr‐40 and nhr‐1 that have been co‐opted for the regulation of mouth‐form plasticity and act downstream of eud‐1 and seud‐1/sult‐1 (Sieriebriennikov et al. 2020; Theska and Sommer ²⁰²⁴). It is important to note that the understanding of the molecular regulation of feeding structure polyphenism is far from completion with many epigenetic aspects still to be identified (Brown et al. 2024; Werner et al. ²⁰²³).

Figure 1.

Mouth‐form polyphenism of P. pacificus, and the experimental design for supplementation studies. (a) Differential interference contrast (DIC) images of the eurystomatous (Eu) and the stenostomatous (St) mouth morphs. The Eu morph has a wide buccal cavity, a hook‐like dorsal tooth (on the left, outlined), and a sub‐ventral tooth. The narrow‐mouthed stenostomatous (St) morph has a flint‐shaped dorsal tooth (on the left, outlined). Scale bar is 5 μm. (b) Illustration of the general experimental design using supplementation with monosaccharides and fatty acids.

Mouth‐form plasticity in P. pacificus is influenced by several environmental factors such as culture condition, population density, and temperature (Lenuzzi et al. 2023; Werner et al. ²⁰¹⁷, ²⁰¹⁸). Previous studies have also revealed prominent roles for nematode metabolites (e.g., ascarosides) and dietary bacteria in the mouth‐form decision and predatory feeding (Akduman et al. 2020; Bose et al. ²⁰¹²; Dardiry et al. ²⁰²³). In contrast, our knowledge of how changes to the nutritional status of worms affect mouth‐form plasticity is still scarce. Bento, Ogawa and Sommer (2010) have shown that starvation promotes the development of the Eu morph. More recent genetic work by Casasa and co‐workers identified the Mediator subunit MDT15/MED15 to regulate mouth‐form polyphenism, which is known to have a conserved role in metabolic responses to nutritional stress (Casasa, Katsougia, and Ragsdale ²⁰²³). These observations suggest that mouth‐form plasticity is sensitive to nutritional conditions in particular to low nutrition. However, mouth form and fitness consequences of a high nutritional condition have never been investigated in P. pacificus.

One approach to studying the effect of nutrition on nematode plasticity is to introduce supplements, such as monosaccharides and fatty acids, into the standard diet to increase fat storage in nematodes, which is a strong alteration to the nutritional status. This aided research in C. elegans to discover novel functions of molecular factors involved in lipid metabolism and associated biological processes (Alcántar‐Fernández et al. ²⁰¹⁸; Han et al. ²⁰¹⁷; Lee et al. ²⁰¹⁵; Nomura et al. ²⁰¹⁰; Wan et al. ²⁰²²). Note that while nutrient supplementation is well established in C. elegans, the role of direct and indirect effects of supplementation through the bacterial diet cannot be completely disentangled and is still controversial (Kingsley et al. 2024). Very likely, the effects observed in such experiments are due to both direct and indirect uptake of nutrients. Moreover, nutrient supplementation studies have prioritised investigating lipid storage as an indicator of nutritional status. Nematodes lack a specialised adipose tissue for lipid storage; however, they generate lipid droplets in many cells including the intestine to serve this function. In addition, storage lipids can be visualised through staining methods such as Oil Red O (ORO), which is a reliable post‐fix technique, providing a proxy for lipid storage when it is quantitatively measured (O'Rourke et al. ²⁰⁰⁹).

Here, we study the influence of nutrition on mouth‐form plasticity in P. pacificus by establishing experimental setups in which worms are grown in ‘high nutrition’ dietary conditions that promote fat storage. These conditions are obtained through the supplementation of monosaccharides and fatty acids (mainly glucose and oleic acid) into the nematode growth medium (NGM) (Figure 1b). We also examine the lipid storage in worms via ORO staining and imaging analysis. We show that fat storage‐promoting conditions, in particular glucose supplementation, render worms non‐predatory. Similarly, we test whether these dietary effects are mediated transgenerationally but find no such influence. In addition, we carry out transcriptomic and subsequent mutant analyses to explore associated molecular mechanisms. Results indicate that de novo fatty acid synthesis and peroxisomal beta‐oxidation pathways, which are involved in the storage and utilisation of lipids, are essential for nutrition‐induced mouth‐form plasticity. Finally, we examine worm fitness by using fecundity as a proxy to explore whether the mouth‐form response in the glucose‐supplemented dietary condition benefits P. pacificus. Our findings suggest that non‐predatory animals have a fitness advantage over predatory animals grown under the same nutritional conditions. Overall, this work establishes nutritional status as an additional factor influencing mouth‐form polyphenism and highlights the beneficial advantage of this plastic ability in P. pacificus.

2. Results

2.1. Monosaccharide and Fatty Acid Supplementations Render Worms Non‐Predatory in a Concentration Dependent Manner

For all experiments, we studied hermaphrodites of the PS312 strain, the wild type of P. pacificus that had its genome sequenced at the chromosome level (Dieterich et al. 2008; Rödelsperger et al. ²⁰¹⁷). This strain is highly Eu under standard lab conditions on an E. coli OP50 diet with more than 95% of animals expressing the Eu morph. First, we performed pilot experiments with monosaccharides ranging from 20 to 300 mM concentrations. Results revealed a concentration‐dependent effect on mouth‐form plasticity. Specifically, the number of St worms increased with increasing concentrations of monosaccharides (Figure S1a). Among all, we prioritised glucose as the main St‐inducing monosaccharide and performed further experiments by using 100 and 150 mM concentrations. Results indicated that 100 mM glucose is sufficient to render worms highly St without exerting observable adverse effects (Figure 2a). Note that even under glucose‐supplementation conditions, there is still an effect of population density on mouth‐form expression as pilot experiments starting from 10 adult worms (Figure S1a) revealed higher Eu ratios than similar glucose concentration experiments with three adult inoculations (Figure 2a).

Figure 2.

Effect of oleic acid and glucose supplementations on mouth‐form plasticity and lipid storage in P. pacificus. (a) Concentration‐dependent effect of oleic acid and glucose on mouth‐form plasticity. N ≥ 3 biological replicates per condition for each concentration. Each faint data point represents a replicate (plate), with 30 animals per replicate being scored for mouth‐form percentage (Eu%). Error bars represent SEM. (b) Oil Red O absorbance (RawIntDen/body area) obtained from worms grown in oleic acid, glucose, and respective control conditions. N = 15 per condition. Each faint data point represents an individual worm. P values are obtained from Welch two sample t‐test and two‐sample t‐test for oleic acid and glucose comparisons, respectively. Bars represent the mean values of all samples in each condition. Error bars represent s.d. (c) Representative images of ORO‐quantified worms, indicating lipid storage profile. Images are acquired from the blue channel in grayscale. Lipid droplets appear dark. Scale bar is 50 μm.

For fatty acid supplementations, we used oleic acid (monounsaturated fatty acid) and linoleic acid (polyunsaturated fatty acid) to test 0.5 and 1 mM concentrations. Similar to monosaccharide supplementations, oleic acid effect on mouth‐form plasticity was mediated in a concentration‐dependent manner (Figure 2a). Observations based on morphology and developmental pace of worms at both concentrations suggested that 0.5 mM oleic acid would be ideal for further experiments. Results also indicated that the same concentrations were detrimental for worms in the linoleic acid condition. While worms did not populate the plates at 1 mM, they were developmentally slow and almost completely St at 0.5 mM linoleic acid (Figure S1b). Therefore, we performed another experiment for linoleic acid with reduced concentrations ranging from 0.01 to 0.2 mM. We found that 0.2 mM linoleic acid renders worms predominantly St, alleviating the detrimental effect observed at 0.5 mM concentration (Figure S1c). Overall, these results show that monosaccharides and fatty acid supplementations induce changes in the mouth‐form ratio towards St in P. pacificus. For further experiments, we selected oleic acid and glucose as main supplements at 0.5 and 100 mM concentrations, respectively.

2.2. Oleic Acid and Glucose Diets Do Not Induce a Transgenerational Effect on Mouth‐Form Plasticity

Some dietary and nutritional influences on nematodes are known to be transmitted transgenerationally (Beltran et al. 2020). Therefore, we propagated cultures on oleic acid and glucose conditions for five generations to explore the potential transgenerational effects of these diets on mouth‐form plasticity (Figure S2a,b). We found that worms exhibit similar mouth‐form ratios relative to their initial response (F1) through generations (Figure S2c,d). When we reverted worms from each generation back to the regular dietary condition without supplements, we did not observe any transgenerational memory of mouth form (Figure S2e,f). Note that over the course of these experiments, we observed more consistent and less variable mouth‐form responses in glucose condition (highly St, on average below 20% Eu) relative to oleic acid across different batches.

2.3. Oleic Acid and Glucose Supplementations Induce Fat Storage

Next, we performed ORO staining on worm populations reared on oleic acid and glucose diets to visualise and quantify their lipid storage (Figure 1b). Relative to control groups, oleic acid‐ and glucose‐fed worms showed an increase in ORO absorbance which signifies an increase in fat storage (Figure 2b). Whole‐body images of stained animals also demonstrated the distribution of storage lipids. Anterior regions of worms clearly indicated lipid accumulation in both oleic acid and glucose conditions (Figure 2c). In particular, the head region of the worm allows easy visualisation of the influence of supplementation on fat storage (Figure 2c inlets). Thus, both tested supplements induce fat storage in addition to their effect on mouth‐form expression.

2.4. Differential Gene Expression Analysis Points Out Pathways Involved in Lipogenesis and Lipolysis

To understand which molecular factors are involved in mediating this nutrition‐induced mouth‐form change, we carried out RNA‐seq on worm populations reared in supplemented and control conditions (Figure 3a). First, we performed differential gene expression analysis between each supplement and its control, and then a KEGG enrichment analysis of differentially expressed genes in both oleic acid and glucose conditions. KEGG enrichment analysis revealed pathways involved in lipogenesis (‘biosynthesis of unsaturated fatty acids’, ‘fatty acid elongation’) and lipolysis (‘peroxisome’, ‘fatty acid degradation’) (Figure 3b). This suggested that worms do not only store but also actively utilise lipids in supplemented conditions. Furthermore, we found genes which were upregulated and involved in peroxisomal beta‐oxidation and biosynthesis of unsaturated fatty acids. Therefore, we selected genes in these pathways as candidates to investigate whether they mediate the effect of nutrient supplementation on mouth‐form expression. We utilised CRISPR gene editing technology to introduce mutations in genetic components involved in both pathways as this method allows easy manipulation in P. pacificus (Han et al. 2020; Witte et al. ²⁰¹⁵). For these experiments, we only used a glucose‐supplemented diet to induce fat storage and test responses of mutants because of the higher reproducibility of results obtained through glucose supplementation and also the impracticality of fatty acid supplementations that require preventive measures from oxidation by light and air (see Materials and methods for more details).

Figure 3.

Transcriptome analysis. (a) An illustration of RNA extraction. (b) Overrepresented KEGG pathways of differentially expressed genes for oleic acid and glucose conditions. Statistical analysis by Fisher's exact test with multiple testing corrections (Bonferroni corrected p < 0.05).

2.5. Delta‐9 Desaturase Activity Is Essential for a Complete Mouth‐Form Response on Glucose‐Supplemented Diet

We targeted delta‐9 fatty acid desaturases for studying the role of the biosynthesis of unsaturated fatty acids and dhs‐28 and daf‐22 for investigating the peroxisomal beta‐oxidation pathway on nutritional plasticity of the P. pacificus mouth‐form. Delta‐9 desaturases facilitate lipid storage by acting early in the de novo fatty acid synthesis pathway, converting saturated fatty acids into monounsaturated fatty acids (Figure 4a). In C. elegans, for instance, simultaneous inhibition of the function of two delta‐9 desaturases (fat‐6 and fat‐7) results in the loss of endogenous unsaturated fatty acids and a significant reduction in lipid storage (Brock, Browse, and Watts ²⁰⁰⁷). Delta‐9 desaturases are activated by the transcription factor sbp‐1 (sterol regulatory element binding protein). Like C. elegans, P. pacificus has a single sbp‐1 gene (PPA37968), which is a one‐to‐one ortholog of Cel‐sbp‐1. First, we introduced frameshift mutations in this gene to block the activity of delta‐9 desaturases and explore its contribution to mouth‐form plasticity. It is known that loss‐of‐function mutations in sbp‐1 cause detrimental effects in C. elegans such as arrested development at the first larval stage (L1), and morphologically slim and pale worms (McKay et al. ²⁰⁰³). Frameshift mutations resulted in similar characteristics in P. pacificus. Specifically, homozygous mutants were sterile; therefore, we could not further our investigation with this gene. Next, we specifically targeted individual delta‐9 desaturases. Interestingly, this gene family has been amplified in P. pacificus relative to C. elegans (Markov, Baskaran, and Sommer ²⁰¹⁵). We constructed a phylogenetic tree based on protein sequences of all P. pacificus fatty acid desaturase domain‐containing genes and related C. elegans desaturases. We found a total of 10 P. pacificus genes which contain a delta‐9 fatty acid desaturase‐like protein domain (Figure S3). Note that unlike for sbp‐1, there is no one‐to‐one orthology relationship between delta‐9 fatty acid desaturase genes between P. pacificus and C. elegans, therefore requiring a new nomenclature. Among these genes, ppa_stranded_DN27845_c0_g3_i3, PPA1053, PPA40514, and ppa_stranded_DN27845_c0_g2_i1 showed highest protein sequence similarity with C. elegans delta‐9 desaturases (fat‐5, fat‐6 and fat‐7). In this order, we named these genes as Ppa‐pddl‐1 (Pristionchus delta‐9 desaturase‐like‐1), Ppa‐pddl‐2, Ppa‐pddl‐3 and Ppa‐pddl‐4 (Figure 4b; Figure S3). Among all, Ppa‐pddl‐1, Ppa‐pddl‐3 and Ppa‐pddl‐4 are expressed throughout the development of P. pacificus based on previous gene expression analysis (Baskaran et al. 2015). Therefore, we prioritised these three genes and introduced mutations through CRISPR. We obtained three frameshift alleles for both Ppa‐pddl‐3 (tu2033, tu2034, tu2035) and Ppa‐pddl‐4 (tu2030, tu2031, tu2032), and one frame shift allele (tu2028) and a 3 bp insertion allele (tu2029) for Ppa‐pddl‐1 (Table S2). Among these mutants, tu2028 and tu2029 were developmentally slow and had significantly reduced fat storage until late adulthood under standard conditions.

Figure 4.

Significance of delta‐9 desaturase activity in glucose supplementation‐induced mouth‐form plasticity. (a) Schematic representation of the essential activity of delta‐9 desaturases, facilitating fat storage by converting saturated fatty acids into monounsaturated fatty acids. Sterol regulatory element binding protein 1 (SBP‐1) activates delta‐9 desaturases. (b) A simplified phylogenetic tree of Pristionchus delta‐9 desaturase‐like genes (Ppa‐pddl‐1, Ppa‐pddl‐2, Ppa‐pddl‐3, Ppa‐pddl‐4), and C. elegans delta‐9 desaturases (fat‐5, fat‐6 and fat‐7). Triangle denotes P. pacificus genes which are constitutively expressed during development (Baskaran et al. 2015). The phylogenetic tree is drawn based on the tree in Figure S3. (c) Eu percentages of CRISPR mutants of Ppa‐pddl‐1, Ppa‐pddl‐3, Ppa‐pddl‐4 and wild type animals in control and glucose conditions. N ≥ 2 biological replicates per strain for each condition. Bars represent mean values of all replicates. Error bars represent s.d. (d) Eu percentages of wild type, tu2028(Ppa‐pddl‐1), and CRISPR repaired‐tu2028 strains in control and glucose conditions. N ≥ 3 biological replicates per strain for each condition. (c,d) Each faint data point represents a replicate (plate), with 30 animals per plate being scored for mouth‐form percentage (Eu%). (e) ORO absorbance (RawIntDen/body area) obtained from wild type, tu2028(Ppa‐pddl‐1), and CRISPR repaired‐tu2028 strains grown on control and glucose conditions. N = 15 per strain for each condition. Each faint data point represents an individual worm. (d,e) Error bars represent s.e.m. (f) Representative images of ORO‐quantified strains from control (‐glucose) and glucose ( + glucose) conditions. Images are acquired from the blue channel in grayscale. Lipid droplets appear dark. Scale bar is 100 μm.

Next, we assessed mouth‐form responses of all mutants on glucose‐supplemented diets. Results revealed that the response of delta‐9 desaturase mutants to glucose is generally incomplete, that is, they have a higher Eu ratio than wild type animals (above 20% Eu) (Figure 4c). Compared to all other mutants, Ppa‐pddl‐1 mutants were more consistent in their mouth‐form response to the glucose‐supplemented diet (greater than 40% Eu on average) (Figure 4c). Therefore, we further studied Ppa‐pddl‐1 using tu2028 as reference allele. First, we measured the lipid storage profile of Ppa‐pddl‐1(tu2028) on standard diet by ORO staining. We found that Ppa‐pddl‐1(tu2028) exhibits significantly lower levels of ORO absorbance relative to wild type (Figure S4a,b). To confirm that the response of tu2028 to glucose‐supplemented diet is due to its mutational background in Ppa‐pddl‐1 and not to other mutations as a result of potential off target effects by CRISPR, we genetically reverted Ppa‐pddl‐1(tu2028) back to wild type using CRISPR editing via a repair template. Growing wild type, the repaired strain, and the original Ppa‐pddl‐1(tu2028) mutant on glucose‐supplemented diets revealed highly similar levels of mouth‐form ratios between wild type and repaired worms (Figure 4d). Specifically, both strains were highly St on glucose‐supplemented diets. Repaired and wild type strains also showed a similar trajectory of ORO absorbance between control and glucose conditions (Figure 4e). Among all, Ppa‐pddl‐1(tu2028) had the weakest ORO absorbance in both conditions (Figure 4e). In addition, we observed that Ppa‐pddl‐1(tu2028) mutant animals restored their growth on glucose‐supplemented diet and also regained their wild type physiological and morphological characteristics upon genetic repair (Figure 4f). Overall, these results highlight the significance of the function of Ppa‐pddl‐1 in both lipid storage and mouth‐form plasticity.

2.6. Peroxisomal Beta‐Oxidation Mutants Fail to Respond to Glucose Supplementation

Next, we studied mutants of two peroxisomal beta‐oxidation genes, dhs‐28 and daf‐22. P. pacificus has two copies of both genes in its genome (Markov et al. 2016) (Figure 5a). For Ppa‐dhs‐28.1, we induced mutation in the copy with the sterol carrier protein domain (PPA20393) and obtained two insertion and three deletion alleles (Table S2). All mutants exhibited the same observable characteristics and were morphologically smaller and developmentally slower relative to wild type animals. For daf‐22, we used the available double mutant Ppa‐daf‐22.1 Ppa‐daf‐22.2 (Markov et al. 2016). Note that mutants of Ppa‐dhs‐28.1 were generated in the PS312 wild type background, whereas the Ppa‐daf‐22.1 Ppa‐daf‐22.2 double mutant was generated in RS2333, which is a highly related derivative of PS312 that was used for several studies on dauer development (Falcke et al. 2018; Markov et al. ²⁰¹⁶). Therefore, we included both highly similar wild type strains in our experiments as independent controls. First, we assessed mouth‐form responses of three Ppa‐dhs‐28.1 mutant alleles (tu1855, tu1856, and tu1858) and the Ppa‐daf‐22.1 Ppa‐daf‐22.2 double mutant on glucose‐supplemented diets. Mutants of both genes had a consistently high Eu mouth‐form ratio between control and glucose conditions. Next, we simultaneously assessed mouth‐form plasticity and lipid storage of Ppa‐dhs‐28.1(tu1855) and the Ppa‐daf‐22.1 Ppa‐daf‐22.2 double mutant on glucose‐supplemented diets. Relative to both wild type strains, mutants remained highly predatory and showed no increase in ORO absorbance between control and glucose conditions (Figure 5b,c). We also observed that glucose‐supplemented peroxisomal beta‐oxidation mutants exhibit delayed development and reduced body size. Moreover, ORO‐stained peroxisomal beta‐oxidation mutants show a disruption in lipid storage integrity on glucose‐supplemented diets (Figure 5d). They also accumulate larger lipid droplets relative to wild type, indicating reduced lipid oxidation. Taken together, these results indicate that the peroxisomal beta‐oxidation pathway is required for the mouth‐form response to glucose.

Figure 5.

Response of peroxisomal beta‐oxidation mutants to glucose‐supplemented diet. (a) Schematic of peroxisomal beta‐oxidation pathway, indicating duplicate copies in P. pacificus for dhs‐28 and daf‐22 genes relative to C. elegans. (b) Eu percentages of peroxisomal beta‐oxidation mutants and wild type strains in control and glucose conditions. N ≥ 2 biological replicate per strain for each condition. Each faint data point represents a replicate (plate), with 30 animals per plate being scored for mouth‐form percentage (Eu%). (c) ORO absorbance (RawIntDen/body area) obtained from peroxisomal beta‐oxidation mutants and wild type strains grown on control and glucose conditions. N = 15 per strain for each condition. Each faint data point represents an individual worm. (b,c) Error bars represent s.e.m. (d) Representative images of ORO‐quantified strains from control (‐glucose) and glucose ( + glucose) conditions. Images are acquired from the blue channel in grayscale. Lipid droplets appear dark. Scale bar is 100 μm.

2.7. Mouth‐Form Plasticity Switch Genes Are Required for Phenotypic Response in Glucose Diet

Next, we tested whether the influence of glucose‐supplemented diets on mouth‐form plasticity is mediated through the central gene regulatory network of mouth‐form plasticity (Figure 6a). For that, we utilised mutants of plasticity switch genes which are St‐form defective. Specifically, we used sult‐1(tu1061) and the double mutant nag‐1(tu1142) nag‐2(tu1143) and assessed their mouth form and lipid storage responses on glucose‐supplemented diets. Both mutants were unable to switch from Eu to St on glucose‐supplemented diets, indicating that the nutritional effect acts upstream of the plasticity switch module (Figure 6b). However, plasticity switch mutants were able to increase their lipid storage when fed on glucose‐supplemented diets (Figure 6c). This suggests that the plasticity switch genes are essential for mouth‐form response but not for lipid storage.

Figure 6.

Role of plasticity switch genes in glucose supplementation‐induced mouth‐form plasticity. (a) Schematic of the mouth‐form gene regulatory network, indicating different modules with genetic and epigenetic components. (b) Eu percentages of plasticity switch mutants and wild type strains in glucose and control conditions. N = 3 biological replicates per strain for each condition. Each faint data point represents a replicate (plate), with 30 animals per plate being scored for mouth‐form percentage (Eu%). (c) ORO absorbance (RawIntDen/body area) measured in plasticity switch mutants and wild type strains grown on control and glucose conditions. N = 15 per strain for each condition. Each faint data point represents a worm. (b,c) Error bars represent s.e.m.

2.8. Non‐Predatory Worms Exhibit a Fitness Advantage Over Predatory Ones

Our final aim was to explore whether there is a fitness advantage of favouring the development of one morph over the other in a fat storage‐inducing condition. For this, we chose fecundity as proxy for fitness to evaluate the hermaphrodites' reproductive success by counting viable progeny produced by isolated individuals. First, we reduced the concentration of glucose from 100 to 80 mM to increase the likelihood of obtaining Eu animals. Then, we isolated animals of both morphs grown in the glucose‐supplemented diets and measured their fecundity on the standard diet (Figure 7a). Results revealed that St worms on average exhibit higher daily and total fecundity than Eu worms (Figure 7b,c). These findings suggest that promoting the development of the St morph under such a dietary influence confers a fitness advantage to P. pacificus.

Figure 7.

Fecundity of different morphs after glucose supplementation. (a) Illustration for the experimental design to study fecundity of Eu and St animals obtained from the glucose‐supplemented condition. (b) Daily fecundity of Eu and St animals. Error bars represent s.e.m. (c) Overall fecundity of Eu and St animals. Bars represent mean values of all individuals for each morph. Error bars represent s.d. P value is obtained from Wilcoxon rank sum test. (b,c) N = 34 per morph. Each faint data point represents a worm.

3. Discussion

This study highlights the significance of nutrition in mouth‐form plasticity in P. pacificus. First, we showed that fat storage‐promoting conditions induce the non‐predatory morph. This suggests that growing in an ‘overly satiated’ nutritional state increases the likelihood of worms developing as non‐predatory. These findings are consistent with previous studies, which indicated that opposite dietary condition, i.e., low nutrition, can lead to the predatory morph in P. pacificus (Bento, Ogawa, and Sommer ²⁰¹⁰); and even a cannibalistic novel predatory mouth form in another diplogastrid nematode, Allodiplogaster sudhausi (Wighard, Witte, and Sommer ²⁰²⁴). In addition, results obtained from transgenerational experiments suggest that worms must constantly be kept at overly satiated state during development to obtain and maintain a mouth‐form response, at least in the wild type P. pacificus PS312 strain.

Supplementing NGM agar plates with glucose and oleic acids has been an effective method for delivering monosaccharides and fatty acids to worms via dietary ingestion (Alcántar‐Fernández et al. ²⁰¹⁸; Deline, Vrablik, and Watts ²⁰¹³; Nomura et al. ²⁰¹⁰). However, it is important to note that this method does not address any potential indirect effect caused by the interaction between the supplement and the bacteria (Kingsley et al. 2021), which can be a research interest on its own. In our experiments, we did not monitor fatty acid and glucose uptake or content in worms, instead we utilised ORO staining, to monitor the outcome of changes in lipid storage. Our results are concordant with previous findings. For instance, glucose supplementation to NGM plates does not only result in the accumulation of glucose but also triacylglycerols in C. elegans (Alcántar‐Fernández et al. ²⁰¹⁸). Oil Red O stains neutral lipids, mainly triacylglycerols. Studies in C. elegans have shown an increase in ORO levels in worms grown in oleic acid‐ and glucose‐supplemented conditions (Choi et al. 2021; Lee et al. ²⁰¹⁵). Therefore, ORO does not only provide a way of measuring the nutritional status of worms but also validate the uptake of supplementations. Moreover, our results showed that glucose supplementation consistently induces the St morph in higher proportions relative to oleic acid. Note that we could not rule out the cause of inconsistency and the high variability of mouth‐form ratios in oleic acid supplementations. Regardless, we selected glucose‐supplemented diet as the main fat storage‐promoting condition to induce the non‐predatory morph.

Further, mutant analyses revealed that genes involved in lipogenesis and lipolysis play a significant role in glucose supplementation‐induced mouth‐form plasticity. First, we showed that delta‐9 desaturase activity, particularly Ppa‐pddl‐1, is required for a complete mouth form and lipid storage response. The spatial transcriptome data, published by Rödelsperger et al. (2021), indicates that Ppa‐pddl‐1 is enriched in anatomical regions that suggests expression in the intestine. Inactivation of this gene causes a drastic reduction in lipid storage and developmental rate; such phenotypic effect is obtained through simultaneous inhibition of fat‐6 and fat‐7 in C. elegans (Brock, Browse, and Watts ²⁰⁰⁷). However, Ppa‐pddl‐1 mutant still exhibits a partial ability to respond to glucose‐supplemented diet. This suggests that remaining functional delta‐9 desaturases may have compensated for the loss of Ppa‐pddl‐1. Therefore, further functional characterisation of delta‐9 desaturases is required to completely elucidate potential role of this gene family in mouth‐form plasticity. Furthermore, growing peroxisomal beta‐oxidation mutants on glucose‐supplemented diet revealed that these mutants have disrupted lipid storage integrity, which resulted in a weaker ORO absorbance relative to wild type strains. Additional observations in these mutants, such as developmental delay and reduction in body size, suggest that they are not able to nutritionally benefit from the glucose‐supplemented diet. Taken together, these findings suggest that pathways associated with storage and utilisation of lipids carry out essential metabolic processes, mediating mouth‐form plasticity in a high nutrition environment.

Importantly, the nutritional sensitivity of the mouth‐form polyphenism never resulted in intermediate phenotypes, consistent with the idea that mouth‐form plasticity represents a developmental switch. This observation goes back to the first genetic analysis of P. pacificus mouth formation (Ragsdale et al. 2013) and has been continuously observed (for recent review see Wighard and Sommer ²⁰²⁴). Additionally, our findings suggest that the dietary effect of glucose supplementation acts upstream of the plasticity switch module. However, how this dietary effect is mediated to influence downstream components, affecting mouth‐form decision remains subject of future research. Of note, de novo fatty acid synthesis and peroxisomal beta‐oxidation pathways produce signalling molecules which have diverse functions (Artyukhin et al. 2018; Watts ²⁰¹⁶). For example, polyunsaturated fatty acids produced by the de novo fatty acid synthesis pathway can be integrated into storage lipids or processed further into signalling molecules such as eicosanoids, which can function as ligands for transcription factors, affecting gene expression. Cytochrome P450 enzymes are involved in the production of eicosanoids from polyunsaturated fatty acids (Kulas et al. 2008) and their activity have been associated with several biological functions such as development, dauer formation, and lipid metabolism in C. elegans (for review, see Larigot et al. ²⁰²²). Cytochrome P450‐related pathways are enriched (KEGG) in our differential gene expression data for both oleic acid and glucose, suggesting a potential role. Moreover, peroxisomal beta‐oxidation pathway produces ascarosides, extracellular signalling molecules which can influence dauer formation and mouth‐form plasticity (Butcher 2017; Butcher et al. ²⁰⁰⁷; Markov et al. ²⁰¹⁶; Werner et al. ²⁰¹⁸). Intriguingly, a transcriptomic‐based study in Spea multiplicata (Mexican spadefoot toad) has also revealed a strong association between the resource polyphenism and the peroxisome function (Levis et al. 2021). Levis and colleagues have indicated a promising role of lipid metabolism in the development of the carnivore toads, suggesting peroxisomes as important organelles that mediate developmental switch between the omnivore and carnivore morph in this organism. Furthermore, lipid‐derived signalling mediators, such as hormones, have been implicated in resource polyphenisms in nematodes (Bento, Ogawa, and Sommer ²⁰¹⁰) and spadefoot toads (Ledón‐Rettig, Pfennig, and Crespi ²⁰⁰⁹; Pfennig ¹⁹⁹²), and in insect polyphenisms (Moczek and Nijhout 2002; Nijhout ¹⁹⁹⁹). In particular, Bui and Ragsdale (2019) have speculated that a satiety‐signalling molecule may get activated by sulfotransferase seud1/sult‐1 to repress the activity of NHR‐40, resulting in the expression of the St morph in P. pacificus. Taken together, lipid‐mediated signalling may have potential functions not only in the regulation of mouth‐form plasticity but also in other forms of polyphenisms. Therefore, further research is required to elucidate associated mechanisms.

Finally, we sought to explore whether there is an adaptive value of facilitating the development of the St morph in a high nutrition environment. In P. pacificus, while the Eu morph exhibits a fitness advantage associated with its predatory ability (Serobyan, Ragsdale, and Sommer ²⁰¹⁴), the St morph claims this advantage through a faster development and a higher fecundity (Dardiry et al. 2023; Serobyan et al. ²⁰¹³). In addition, highly non‐predatory natural isolates of P. pacificus have higher fecundity and exhibit faster development on standard dietary conditions relative to highly predatory strains, which were isolated from similar localities in nature (Dardiry et al. 2023). Growing worms on a glucose‐supplemented diet allowed the separation of morphs; then assessing their fecundity revealed a fitness disadvantage for the predatory mouth form. Intriguingly, our findings suggest that glucose supplementation does not offset the fitness cost of producing the Eu phenotype or promote its development to begin with. Hence, the fitness advantage associated with the non‐predatory morph suggests that favouring its development, under such dietary effects, can be beneficial for the population. It is also important to note that glucose can interact with E. coli to produce reactive oxygen species and advanced glycation end products, resulting in detrimental effects in nematodes such as reduced lifespan and health span (Kingsley et al. 2021, ²⁰²⁴). High levels of glucose also induce a transgenerational decrease in fecundity in C. elegans (Alcántar‐Fernández et al. ²⁰¹⁸; Tauffenberger and Parker ²⁰¹⁴). In P. pacificus, the development of the St individuals may circumvent the adverse influences of secondary effects of glucose supplementation on worms' fitness.

In summary, this study signifies the nutritional sensitivity of mouth‐form polyphenism in P. pacificus and adds nutritional status as an important environmental factor influencing mouth form. We introduced glucose supplementation as a novel environmental condition to induce non‐predatory morph. Our findings suggest that the nutritional status of the worm can potentially dictate its mouth‐form fate. We also found a strong association between lipid metabolism and mouth‐form plasticity. Lastly, glucose‐supplementation helped understand the fitness consequences of the mouth‐form determination, indicating an advantage for the non‐predatory morph.

4. Materials and Methods

4.1. Nematode Stock Maintenance

All P. pacificus strains were cultured on 6 cm NGM plates with 300 μL E. coli OP50 provided as food at 20°C as previously described (Sommer et al. 1996). The wild‐type strain PS312 was used for experimentation unless otherwise noted. Stocks were maintained by passing three young adult hermaphrodites with eggs every 5 days. Mutant P. pacificus strains were similarly cultured and are available from the SommerLab.

4.2. Monosaccharide Supplementation Experiments

Monosaccharides were purchased from Sigma‐Aldrich as follows: glucose (D‐(+)‐Dextrose, product number: D9434), galactose (D‐(+)‐Galactose, product number: G0750), fructose (D‐(−)‐Fructose, product number: F0127). All monosaccharides were dissolved in purified and autoclaved water to obtain 1 M stock concentrations. Stock solutions were filtered through 0.22 μm sterile filters before storage. Plate pouring was carried out under a sterile hood. Each monosaccharide stock was diluted into liquid NGM media to obtain the desired final concentrations. Monosaccharide‐added NGM was then stirred for approximately 1 min to obtain a homogenous media. Media was then poured into 6 cm plates using a sterile 10 mL serological pipette, obtaining a volume of 9 mL in each plate. Control plates with the same volume were poured from the main NGM media without monosaccharides. Plates were allowed to dry for 2 days and were then seeded with 300 μL overnight grown E. coli OP50. After 2 days, three young hermaphrodites with eggs (from stocks grown under standard conditions) were added onto each plate for each strain, condition, and replicate. After adding worms, plates were incubated at 20°C until F1 adults emerged. Plates were then used for mouth‐form phenotyping and ORO staining.

4.3. Fatty Acid Supplementation Experiments

Fatty acid supplementation was carried out as described by Deline, Vrablik and Watts (2013); however, cholesterol was added to the NGM media after autoclaving. Sodium oleate (Sigma‐Aldrich, product number: O7501) and sodium linoleate (Sigma‐Aldrich, product number: L8134) were used for oleic acid and linoleic acid supplementation, respectively. For each experiment, a 0.1 M fatty acid working stock was prepared fresh in purified and autoclaved water. Once completely dissolved, the fatty acid working stocks were purged with nitrogen gas to prevent oxidation by air. Plate pouring was carried out as in monosaccharide supplementation except the volume of the NGM in each plate was 8 mL. Both supplemented and control plates contained 0.1% Tergitol (NP‐40, Sigma‐Aldrich, product number: NP40S) to facilitate fatty acid absorption. Once poured, plates were covered with a box to avoid light oxidation. The same steps as in monosaccharide supplementation were followed for seeding the plates with E. coli OP50, culturing nematodes, mouth‐form phenotyping and ORO staining.

4.4. Mouth‐Form Phenotyping

Mouth‐form phenotypes were scored using Zeiss Discovery V.20 stereomicroscope based on the mouth width of individual worms as previously described (Ragsdale et al. 2013; Theska et al. ²⁰²⁰). Each plate was taken as a biological replicate and 30 worms were scored for their mouth‐form, unless otherwise mentioned. The percentage of EU animals on each plate was then calculated. Mouth‐form percentage (Eu%) is graphed as a mean of percentages of all plates for each strain in a particular dietary condition.

4.5. Transgenerational Experiments

For testing potential transgenerational effects, lines were established by picking three young (day 1) adult worms with eggs from standard conditions onto oleic acid, glucose, and control plates, respectively. Ten lines were established for each condition and each line was propagated by transferring three young adult worms for five generations on the same condition. In case of contamination, the line was discontinued and not used for counting mouth form or reversal. At every generation, 6–7 lines (among the already established 10 lines) were selected and reverted back to standard culture conditions. For each condition, mouth‐form phenotypes were scored for all plates at every generation and reversal.

4.6. Oil Red O Staining

Post‐fix ORO staining method was modified by Li et al. (2016). Oil Red O stock solution was prepared by dissolving 0.5 g ORO (Sigma‐Aldrich, product number: O0625) in 100 mL isopropanol. This stock solution was shaken on a see‐saw rocker for several days. To prepare the ORO working solution, the required amount was taken from the stock and centrifuged for 5 min at 4500 rpm to pellet all ORO‐related precipitates. Working ORO solution was prepared fresh by diluting precipitate‐free ORO (supernatant) in purified water to obtain 60% ORO. This solution was also shaken on a see‐saw rocker for 10 min and centrifuged for 5 min at 4500 rpm before staining worms. For worm fixation, a 1% formaldehyde solution was used by diluting 1 mL 16% formaldehyde solution (Thermo Scientific, catalogue number: 28906) in 15 mL 1X PBS (phosphate buffered saline). For dehydrating worms, 60% isopropanol was prepared fresh by diluting isopropanol in purified water. First, worms were washed off the plates with 1X PBS and collected in a 15 mL Falcon tube for each sample. Note that, after mouth‐form phenotyping, several biological replicates (plates) were washed into one sample tube for each strain for a particular dietary condition. Once worms formed a sediment, extra volume was removed, leaving only 1 mL. This volume containing worms was then transferred to a new 1.5 mL tube. To pellet worms after transfer or wash, samples were briefly centrifuged in a quick‐spin centrifuge. Worms were fixed with 1% formaldehyde (1 mL) for 30 min by shaking on a see‐saw rocker. Then, samples were frozen in liquid nitrogen ( ~ 8 s) and thawed under running tap water, three times in total. Samples were washed with 1X PBS three times and were dehydrated in 60% isopropanol (1 mL) for 2 min. Finally, samples were stained with 0.5 mL 60% ORO working solution for 30 min on a see‐saw rocker. The staining solution was applied through a 0.22 μm sterile filter. After staining, samples were washed three times with 1X PBS. The supernatant of each sample was removed, leaving approximately 0.1 mL sample of stained worms. Two drops of Vectashield (Biozol, catalogue number: H‐1000) were then added to each sample. Samples were gently mixed by pipetting up and down before mounting onto microscope slides. Samples were then imaged for ORO quantification.

4.7. ORO Quantification and Analysis

Oil Red O quantification was performed as described in Feldhaus, Piskobulu and 2024 (2024 preprint). Briefly, we imaged worms on an AxioImager.Z1 (Zeiss, Oberkochen, Germany) with a 10x/0.3 objective and an AxioCam 506 mono. For ORO quantification, images were obtained with filter sets for DAPI and Texas Red in transmission mode. The filter‐based imaging with a monochrome camera was chosen to remove any potential bias from white balance settings of the camera. Oil Red O absorbance per worm was semi‐automatically calculated by manually outlining the worm area and then obtaining the difference between the Texas Red channel (where ORO appears transparent) and the DAPI channel (where ORO absorbs) after correcting for differences in the spectral response for both channels. In total, 15 adult worms were quantified per strain and condition. For each worm, raw integrated density (RawIntDen), from channel 4 of the processed image, was taken as a measure of ORO absorbance. To compare individuals with different body size (e.g. mutants), ORO absorbance (RawIntDen) was normalised by the body area (μm²) for each worm (RawIntDen/body area). For calculating body area, images of worms from the blue channel (DAPI) were saved in grayscale. The body area was then calculated for each worm by ‘Threshold’ and ‘Analyse Particles’ functions on Fiji (Schindelin et al. 2012). Therefore, the final ORO absorbance value was determined as RawIntDen divided by the body area for each worm.

4.8. RNA Extraction

For RNA extraction, three young adults with eggs, from standard condition, were placed on oleic acid, glucose, and control conditions, respectively. For each condition, we scanned through 5–9 plates and collected 150 F1 young adults without eggs in two separate tubes for RNA extraction (two technical replicates). RNA extraction was carried out using a Direct‐zol RNA Microprep Kit (Zymo Research, R2060). The quality of RNA was assessed using NanoDrop. The concentration of RNA was assessed using a Qubit 2.0 Fluorometer (Invitrogen). Samples were then diluted to the required concentration and sent to NovoGene Co Ltd for sequencing.

4.9. RNA‐Seq and Kegg Enrichment Analysis

Raw reads were aligned to the reference P. pacificus genome (El Paco) with STAR (version 2.7.1a) and quantified with featureCounts from the Subread R package (version 2.0.1) based on the latest gene annotations (Athanasouli et al. 2020; Dobin et al. ²⁰¹³; Liao, Smyth, and Shi ²⁰¹⁴). The count matrix was filtered by removing genes with less than 10 reads, reducing the number of genes to be examined from 28,896 to 18,174 and 18,049 for oleic acid and glucose respectively. Differential gene expression analysis was performed for each of the two conditions with DESeq. 2 (FDR‐corrected p < 0.05, version 1.42.0) (Love, Huber, and Anders ²⁰¹⁴). The differentially expressed genes from each condition were tested for overrepresentation of KEGG pathways using Fisher's exact test with multiple testing correction (Bonferroni corrected p < 0.05) in R (version 4.3.1), based on the existing KEGG annotations for the P. pacificus genes (Athanasouli et al. 2023).

4.10. Phylogenetic Tree

All C. elegans fatty acid desaturase protein sequences were obtained from WormBase (www.wormbase.org). Longer isoforms were picked for genes with different isoforms. Each C. elegans fatty acid desaturase protein was then blasted against P. pacificus on the protein database (www.pristionchus.org, El Paco annotation v3, 2020). From each blast result, all P. pacificus hits were collected and analysed on InterProScan (www.ebi.ac.uk) for conserved protein domains. Note that a few of the Pristionchus hits are transcript‐derived gene predictions with a different name format that starts with ‘ppa_stranded’. These gene predictions always match a genome annotation but they are not assigned with a “PPANNNNN” accession by WormBase. We found two genes (PPA03289 and PPA05783) which did not contain desaturase domains, which were discarded. The rest of the protein sequences were aligned by ClustalW on MEGA11 software (Tamura, Stecher, and Kumar ²⁰²¹). Based on this alignment, a maximum likelihood phylogenetic tree was constructed with the LG model, and 100 bootstrap replications. The phylogenetic tree was visualised and edited on FigTree software (version 1.4.4).

4.11. CRISPR/Cas9 Mutagenesis and Identification of Mutants

For the generation of mutants by CRISPR, the protocols of Witte et al. (2015) and Han et al. (2020) were followed with minor modifications. All components of the CRISPR/Cas9 complex were purchased from Integrated DNA Technologies (IDT). Each CRISPR RNA (crRNA) was designed to target 20 bp upstream of the protospacer adjacent motif (PAM). To construct the CRISPR/Cas9 complex, 5 μL CRISPR RNA (crRNA) from 100 μM stock was mixed with 5 μL trans‐activating CRISPR RNA (tracrRNA) obtained from 100 μM stock (catalogue number 1072534). This mixture was first denatured at 95°C for 5 min, and then cooled down at room temperature for annealing (5 min). Hybridised crRNA:tracrRNA product (5 μL) was mixed with 1 μL Cas9 protein from 62 μM stock (catalogue number: 1081058) and incubated at room temperature for 5 min. Tris‐EDTA buffer was then added into the CRISPR/Cas9 mix to obtain a final concentration of 18.1 μM for crRNA:tracrRNA hybrid and 2.5 μM for Cas9. Coinjection marker, with Ppa‐eft‐3 promoter and TurboRFP sequences, was also integrated into the injection mix (Han et al. 2020). Injected worms were allowed to lay eggs for 24 h. Fluorescent marker‐positive plates were used to isolate emerging F1 progeny. From each positive plate, 8 ‐10 F1 worms were isolated. Each F1 worm was allowed to lay eggs and then was lysed in single worm lysis buffer (10 mM Tris–HCl at pH 8.3, 50 mM KCl, 2.5 mM MgCl₂, 0.45% NP‐40, 0.45% Tween 20, 120 μg/mL Proteinase K) in a thermal cycler programme with 65°C for 1 h; 95°C for 10 min. Lysates were used to carry out polymerase chain reactions to amplify the target region. Mutants in F1 generation (predominantly heterozygotes) were first identified through Sanger sequencing (GENEWIZ Germany GmbH). Subsequent re‐isolation and sequencing of the following generation (F2) allowed capturing homozygous mutants. All crRNAs, and related primers for target genes are provided in Table S1.

4.12. Genetic Repair of Ppa‐Pddl‐1(Tu2028)

To genetically revert Ppa‐pddl‐1(tu2028) back to wild type through CRISPR editing, a crRNA (5’‐ TACTATCCTCTATCCTATAG‐3’) was designed targeting 20 bp upstream of the TGG PAM site at exon 4 based on the mutant sequence including 7 bp insertion. A 100 bp wild‐type repair template, covering the target region, was also used (synthesised by IDT). The repair template was as follows: 5’‐ CATGAATAGTTCTAACCACTTCTTCTCCTTCAGACATTACTATCCTCTAGTGGCCCTTCTCTGTTTCCTCATGCCAACTGTCGTTCCTGTCTATTATTGG‐3’. The repair template was integrated into the CRISPR injection mix at a concentration of 10μM. Homozygous wild‐type worms were isolated in F2 generation post‐injection, and validated via Sanger sequencing.

4.13. Fecundity Experiment

To study the fecundity of Eu and St morphs, young adult worms without eggs (F1) were isolated from glucose‐supplemented diets (80 mM concentration) based on their mouth morph and subsequently transferred to standard dietary conditions with 20 μL E. coli OP50. Worms were passed onto new plates for 8 consecutive days and were removed at the end of 8th day. After 5 days from inoculation, viable progeny was counted on each plate.

4.14. Data Visualisation and Statistical Analysis

All plots were generated in RStudio (version 1.2.5042) using ggplot2 and ggpubr packages. Illustrations were produced in Affinity Designer (version 1.9.2). Statistical significance testing was carried out by comparing two independent groups, e.g., control versus treatment, mutant versus wildtype, Eu versus St, for ORO absorbance and total fecundity. Briefly, data was first tested for normal distribution mostly via Shapiro–Wilk normality test with additional visual observations. In case of normal distribution, either a sample t‐test (equal variance) or Welch two sample t‐test (unequal variance) was used. In case of nonnormal distribution, the Wilcoxon rank sum test was used as the nonparametric equivalent. All statistics were performed in RStudio (version 1.2.5042).

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The data that support the findings of this study are openly available in ENA at http://www.ebi.ac.uk/embl/index.html, reference number PRJEB76787.

1. Peer Review

The peer review history for this article is available at https://www.webofscience.com/api/gateway/wos/peer-review/10.1002/jez.b.23284.

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