The C. elegans N2 Bristol wild-type strain was acquired from the CGC (Caenorhabditis Genetics Centre), funded by the NIH National Center for Research Resource in Minnesota, USA. Standard methods as described by Brenner (1974) (20) were utilized for the cultivation and observation of C. elegans. N2 worms were cultured and maintained at 20°C on Nematode Growth Media (NGM) plates, utilizing Escherichia coli strain OP50 as the primary food source, which served as the control in all assays and analyses. Diabesity in worms was induced by supplementing glucose (100 mM) + cholesterol (25 µM) into nutrient-rich substrate on NGM agar plates. The survival assay was conducted following the protocol outlined by Mir et al. (2020) (21). Synchronized adult-stage organisms were obtained by treating adult-stage hermaphrodites with a mixture of commercial bleach and 5 M KOH in a 1:1 ratio (22).

Age-synchronized L4 stage worms were subsequently transferred to 24-well plates for three biological replicates containing fluorodeoxyuridine (FUDR). Survival assessments began on the first day of adulthood, with each condition represented by a population of 40 or more worms. E. coli OP50 was added to the experimental setup at a cell density of approximately 0.2 OD at 600 nm in the wells. The worms were monitored at 12-hour intervals to record complete survival times. Worms unresponsive to external stimuli such as tapping and gentle touches using a worm picker were classified as dead. Mean and maximum lifespans were calculated using GraphPad Prism, and statistical analyses, including P-values determined using the log-rank (Mantel-Cox) statistic, were performed.

To assess the frequency of pharyngeal pumping in both control and diabesity-induced C. elegans, a total 25 number of worms per biological replicate were placed onto their respective NGM plates seeded with E. coli OP50. The quantification of pharyngeal pumping, measured as the number of contractions per 10 seconds, was observed at specific time intervals using a Stereo microscope, following the method outlined by Durai et al. (2013) (23).

This study utilized N2 nematode strains under two different conditions: Diabesity and standard conditions. The nematodes were cultured on NGM plates containing FUDR regularly. Motility assessments were conducted on the first, fourth, and seventh days of adulthood. Locomotion data were collected and analyzed following the methodology outlined by Rollins et al. (2017) (24). An hour before video recording, the animals were transferred to fresh NGM plates. Young adult to aged synchronized animals were cultured at 20°C for the motility assay. Individual nematodes were tracked, and continuous 1-minute videos were recorded. Wormlab software version 4.1 (MBF Bioscience) (24, 25) was used to analyze the videos. For each strain studied, 10 videos were captured for subsequent analysis. Differences in motility were assessed using the log-rank test in Graph Pad Prism 6.0 software (26–28).

To quantify triglyceride accumulation, Nile Red was employed to stain C. elegans under both diabesity and standard conditions. Adult worms underwent a series of procedures: initially, they were washed in M9 buffer followed by phosphate-buffered saline (PBS). Subsequently, the worms were fixed in 1% paraformaldehyde at 4°C for 1 hour, following the methodology outlined by Escorcia et al. (2018) (29). Further, the worms were dehydrated using 60% ethanol at room temperature for 15 minutes. The final step involved incubating the worms with a Nile Red staining solution, which consisted of 6 volumes of 1 mg/ml Nile Red in ethanol mixed with 4 volumes of ethanol. This staining process continued for 12 hours as detailed in the protocol by Peng et al. (2016) (30)

The ROS was determined utilizing 2’,7’-Dichlorofluorescein diacetate (DCF-DA). Nematodes were collected and suspended in 50 µl of PBS. Subsequently, 10 μl of a 100 mM DCF-DA solution was introduced to the samples, and incubated in darkness for 15 minutes. Following this, samples were washed with M9 buffer. The worms were paralyzed by adding a 1:100 ratios of 100 mM of sodium azide and subsequently the samples were observed and documented using a Nikon fluorescent microscope (31).

For protein and fatty acid analysis, an equal number of control and diabesity worms were homogenized with 100 mg of potassium bromide. The resulting mixture underwent vacuum drying to form a solid pellet. Using FTIR an infrared spectrum ranging from 400 to 4000 cm^-1 was captured. The obtained data points were plotted, represented intensity against wavenumber, following the methodology detailed by Sethupathy et al. (2017) (32).

To ascertain the glucose concentration, the worms were homogenized using the Mini homogenizer MT-13K and centrifuged at 7,500 rpm for 5 minutes to remove the debris. An equivalent concentration of sample was mixed with 1000 μl of Autospan Liquid Gold Glucose reagent, facilitating the quantification of glucose levels through the enzymatic oxidation of glucose. This oxidation process yields gluconic acid and hydrogen peroxide as products. Following a 10-minute incubation period, the color change resulting from the hydrogen peroxide reaction was evaluated using a chromogen. The assessment was performed using a Semi auto analyzer, measuring absorbance at a wavelength of 505 nm (33).

To evaluate the triglyceride level, an equal concentration of sample was mixed with 1000 μl of triglyceride reagent Liqui CHECK™. The enzymatic measurement of triglycerides involved a calorimetric quantification process, where the reaction’s intensity was measured at 546/630 nm wavelengths. This assessment was performed using a Semi Auto Analyser system following the methodology outlined by Sullivan et al. (1985) (34).

Approximately 50 x 10⁵ worms were used to extract lipids from worms. For each 200 μl sample aliquot, 1.5 ml of methanol was added into glass tubes and vigorously vortexed. Subsequently, 5 ml of MTBE (methyl tert-butyl ether) was added to the mixture, and incubated for one hour incubation at room temperature. Following this, 1.25 ml of MS-grade water was added to the mixture to induce phase separation. The mixture was incubated for 10 minutes at room temperature and then centrifuged at 1,000 g for 10 minutes. The upper (organic) phase was collected, while the lower phase underwent re-extraction as described by Matyash et al., 2008 (35).

The lipid extract analysis using Thin-Layer Chromatography (TLC) was conducted on Merck Silica Gel 60 HPTLC plates measuring 10 × 10 cm. The development of silica gel plates was prepared using a solvent mixture comprising n-hexane, diethyl ether, and acetic acid in a ratio of 75:13:9 (v/v/v). To visualize the lipid bands, a spray solution consisting of 20% H₂SO₄ in ethanol was applied. Subsequently, the plates were charred at a temperature of 200°C to visualize the lipid bands, following the method described by Touchstone et al. (1995) (36).

Tissue sections from frozen blocks of both control and diabesity samples were cut at a thickness ranging from 2 to 4 μm to ensure optimal resolution and placed on microscope slides. Subsequently, 10% neutral buffered formalin (NBF) was applied onto the slides and allowed to incubate for approximately 10 to 15 minutes. Heating at 58°C for one hour was conducted for proper preparation of the sections. Following this, the sections underwent deparaffinization and hydration steps. The immunohistochemical process was initiated by fixing the sections with a 4% v/v paraformaldehyde solution in 1X PBS at room temperature. After fixation, the sections were washed twice and then permeabilized using a 0.2% Triton X-100 solution in 1X PBS for 20 minutes at room temperature. Subsequent to permeabilization, the sections were blocked using 2% BSA. After another round of washing, the sections were left to incubate overnight with a primary antibody at a dilution of 1:50, maintained at 4°C. Following incubation with the primary antibody, the sections underwent three washes with PBS and were then incubated with secondary antibodies labeled with Alexa Fluor 488 (Invitrogen) for 1 hour at room temperature. A washing step using diluted 1% BSA in 1X PBS for 5 minutes followed this incubation. Digital images were acquired using a fluorescence microscope from Nikon, Japan, and the process was facilitated using Nis-Elements D software (37)

The LC-MS analysis was conducted based on the methodology outlined in (38), with minor adjustments. Initially 200 mM freshly prepared DTT was combined with 200 µg of protein per sample in a volume of 100 µl. This mixture was incubated in darkness for 1 hour at room temperature. Subsequently, 20 µl of 200 mM IAA was added and incubated for 1 hour at a temperature of 56°C. Following the addition of 200 mM DTT, the sample underwent an in-solution trypsin digestion using trypsin in 55 mM acetic acid at a 1:50 ratio. The samples were then left to incubate for 16 hours at 37°C. For the subsequent LC-MS analysis, an Agilent 6545 ESI-LC/qToF-MS/MS instrument was utilized. Data acquisition was performed using Mass Hunter Workstation software, version B.08.00, while data processing employed Spectrum Mill software, version 06.00. Regarding taxonomic assignment, C. elegans was designated as the organism of interest. A mass tolerance of 20 ppm was set for the precursor mass, while the product mass tolerance was established at 600 mDa (39).

To identify shared proteins within control and diabesity samples protein identification from the LC-MS analysis was subjected to the Venny 2.1.0 tool (http://bioinfogp.cnb.csic.es/tools/venny/), an online platform for creating Venn diagrams. The Gene Ontology (GO) classification of the proteins identified through mass spectrometry was performed using the UniProt database (http://www.uniprot.org/uploadlist/) in accordance with the methodology outlined in (38). To explore the interactions among the identified proteins, the STRING Version 10.5 online tool (http://string-db.org/) was utilized. A medium confidence score of 0.400 was employed to assess protein-protein interactions, as described by Balasubramaniam et al. (2020) (39). Furthermore, the pathways regulated by the identified proteins and gene enrichment analyses were carried out using the KEGG Mapper - search pathway online tool (http://www.genome.jp/kegg/tool/map_pathway1.html). This tool, as described by Pooranachithra et al. (2021) (40), allowed for the exploration of pathways and gene enrichment associated with the identified proteins.

All the experiments were performed independently in triplicates. Results are represented as mean ± standard errors of the mean (SEM). The significance of statistical difference was analyzed using one-way ANOVA with Dunnett’s multiple comparisons test (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001) in SPSS software version 27.0.1 and GraphPad Prism 9 (41).

To understand the mechanistic basis of diabesity, C. elegans were raised in NGM containing high glucose and cholesterol rich diet (see Methods). It has been reported that growing C. elegans on NGM plates containing 40 mM glucose resulted in an intracellular concentration of 14 mM, a range observed in poorly controlled diabetic patients (12, 14). In order to understand the impact of diabesity condition generated by high glucose and cholesterol-rich diets in C. elegans, the changes were observed in lifespan. The introduction of high glucose and cholesterol-rich diets resulted in a significant reduction in both mean lifespan—from 10 ± 0.6 to 5.5 ± 0.4 days (P < 0.0001)—and maximum lifespan—from 21 ± 0.4 to 10 ± 0.4 days (P < 0.0001)—when compared to the control group fed with E. coli OP50 ( Figure 1A ).

For the analysis of intracellular glucose and cholesterol concentrations at the mean lifespan (day 5 of adulthood), whole-body extracts from C. elegans were examined. Culturing C. elegans on agar plates containing 100 mM glucose and 25 μM cholesterol under diabesity conditions resulted in intracellular concentrations of 48 mM glucose and 15 μM cholesterol (P < 0.001, n=50), notably higher than the control worms’ levels at 11 ± 4 mM glucose and 2 ± 1 μM cholesterol (P < 0.0001, n=50) ( Figure 1B ).Under standard conditions, no supplementary glucose is introduced into the nematode growth medium. However, in all subsequent experiments, the high glucose and cholesterol conditions for C. elegans refer to concentrations of 100 mM glucose and 25 μM cholesterol, respectively. This distinction was crucial in assessing the consequences of diabesity conditions induced by these dietary components in C. elegans. In addition, when compared to the control group, the pharyngeal pumping of diabesity C. elegans progressively decreases starting from day 2 of adulthood ( Figure 1C ). After day 6^th of adulthood, all other physiological activities also progressively decreased until C. elegans becomes immobile. Compared to the control, diabesity worms exhibit other defective physiological characteristics such as a fattened body and apathetic behavior ( Figure 1D ). Each experiment was conducted in biological triplicate, and the error bars represent the mean ± SEM (p < 0.005).

Due to the absence of adipocytes, C. elegans stores its fat in small droplet-like organelles referred to as lipid droplets, predominantly located in the intestine and hypodermis (42). To investigate the impact of high glucose and cholesterol on lipid droplets accumulation in C. elegans, Nile red staining was conducted. This staining method targets neutral lipids such as triglycerides or cholesterol esters. The fluorescence from Nile red staining was quantified using ImageJ software. The supplementation of high glucose to the NGM resulted in a significant increase in lipid accumulation in diabesity samples compared to the control ( Figures 2A, B ), consistent with previous findings that used glucose as an obesogenic stimulus in C. elegans (43, 44). Intracellular fat droplets in C. elegans were visualized using fluorescence microscopy (45), and the present study revealed a notable increase in fat and lipid storage within adipose tissues in the diabesity condition and no significant fluorescence was observed in control ( Figure 2C ).

Furthermore, the quantification of triglycerides indicated a substantial elevation in diabesity samples compared to the control. Triglyceride levels were assessed at day 2, 4, and 6 of adulthood. At day 2 of adulthood, diabesity animals exhibited approximately 369 ± 1 ng of triglycerides compared to 170 ± 3 ng in the control (P< 0.0001). By day 4 of adulthood, diabesity animals showed 520 ± 5 ng compared to 188 ± 5 ng in the control (P< 0.0001, n=500). Finally, at day 6 of adulthood, diabesity animals displayed 740 ± 1 ng of triglycerides, whereas the control exhibited 300 ng (P< 0.001, n=500) ( Figure 2D ). In this study, we have also conducted the separation of lipid extracts through thin-layer chromatography following the standard procedure (see methods). Our results demonstrate a significant increase of triglycerides, cholesterol esters, and notably higher amounts of glycolipid species (Sphingomyelin and Phospholipids) in diabesity animals compared to the control group ( Supplementary Figure S1 ). The proportional content analysis from each lane of the TLC plates was performed by densitometry of digital photographs using ImageJ software ( Figure 2E ). Each experiment was conducted in biological triplicates, and the error bars represent the mean ± SEM (p < 0.001).

Lipid peroxidation has been observed to be higher in obese individual mice compared to non-obese (46, 47). Hyperglycemia results in elevated levels of reactive oxygen species (ROS) production through the polyol pathway, leading to the formation of advanced glycation end products (AGEs) and causing significant oxidative stress (48). Assessing ROS levels serves as an effective means to quantify oxidative damage resulting from hyperglycemia. In this study, we measured ROS levels using H₂DCFDA staining. ROS induction was examined at day 5^th of adulthood. We found a higher generation of ROS in worms exposed to diabesity conditions, whereas no significant fluorescence was observed in the control sample ( Figures 3A, B ).

The application of FT-IR spectroscopy has proven to be a valuable tool for investigating biomolecular complex structures within cells (49). In the context of examining the impact of oxidative stress induced by diabesity on C. elegans lipids and proteins, FT-IR analysis revealed significant differences in intensity peaks between diabesity samples and the control group. Specifically, the analysis of C. elegans’ fatty acids exhibited distinct intensity peaks at various wavelengths: 3000-2800 cm^-1, 3100 cm^-1, and 2930 cm^-1. These peaks correspond to fatty acids, N-H stretching of proteins associated with lipids, and CH₂ anti-symmetric stretching of lipids, respectively (50–52) ( Figure 3C ). Additionally, the protein FT-IR spectroscopy analysis depicted features such as carbonyl stretching (1800 - 1500 cm^-1), vibration of Amide I and Amide II bands of proteins (1740 cm^-1), and broadening of the peak at 1650 cm^-1, suggesting protein oxidation (53–55). Furthermore, noticeable differences in intensity were observed in the 1200-900 cm^-1 range, associated with carbohydrate bands (56–58). These spectral differences in the C. elegans fatty acid and protein region imply the impact of stress induced by high diabesity conditions on cellular molecules. Moreover, differences in protein FT-IR spectroscopy indicate variations in protein glycosylation. Changes in protein glycosylation have been linked to various physiological and pathological conditions, such as tumor invasion, cell growth differentiation, host-pathogen interaction, cell trafficking, and transmembrane signaling (59).

High glucose causes susceptible to cognitive impairment, progression of Alzheimer’s disease and decreases the locomotor activity (60–64). The motility of C. elegans is regulated by acetylcholine receptors, which plays important role in synaptic functions (65). The ability to move relies on neuronal signaling, maintenance of muscle mass, and integrity of connective tissue (66–69). Consequently, motility serves as a valuable indicator for evaluating the neurotransmission and is a useful marker to assess health span (70). The motility assays conducted in our study were devised to investigate the intricate connections between high glucose and locomotor behavior. Our study involved the examination of changes in motility at two distinct time points: day-3 and day-6 of adulthood. We conducted these assessments in both under standard conditions and diabesity condition. We utilized MATLAB worm tracking software to analyze quantitative data obtained from video recordings of motility (see Methods). At both day 3 and day 6 of adulthood, nematodes experiencing the diabesity condition displayed decreased motility compared to those in standard conditions ( Figure 4A ). Additionally, worms under standard conditions displayed a notably higher frequency of thrashing movements than those in the diabesity condition ( Figure 4B ). The dopamine signaling in C. elegans is responsible for the gait transition from swim to crawl (71). Studies using ontogenetic confirmed D1-like dopamine receptors switch animals, away from the bacterial food source (72–74). In our studies diabesity condition, nematodes move off from the food source E. coli OP50 bacteria, compared to those under standard conditions (data not shown).

The results obtained from physiological assays prompted further investigation into the primary molecular mechanisms underlying C. elegans diabesity. LC-MS-based proteomic analyses were conducted in biological triplicates to ensure the robustness and reproducibility of the results. Following Post-MS Data analysis for protein identification and expression, a total of 627 differentially regulated proteins were detected. We set the relative expression ratio for downregulated and upregulated proteins between control and treated samples at ≥-1.5 and ≥1.5, respectively, across all biological replicates (p < 0.05). Out of these, 18 proteins were common with 10 being down regulated and 8 being up regulated significantly. Detailed information on these differentially regulated proteins can be found in Supplementary Tables S1 , S2 . Subsequently, the high-throughput protein profile and expression data underwent comprehensive bioinformatics analysis. To assign potential physiological functions to the regulated proteins, a Gene Ontology (GO) annotation and functional enrichment analysis were performed using the UniProtKB tool. The differentially regulated proteins were categorized into molecular functions, biological processes, and cellular components. Based on the GO functional annotation, the major group of regulated proteins in diabesity is associated with response to stimuli and cellular processes. Additionally, genes related to locomotion, localization, immune system processes, responses to stimuli, and signaling pathways exhibit regulatory changes. The primary biological processes predominantly affected among the differentially regulated proteins include metabolic processes and cellular activities ( Figures 5A, B ).

The functional annotation and gene enrichment of all regulated proteins were conducted using the DAVID tool. Functional annotation delineates the role of individual proteins across various biological processes. Among the biological functions exhibiting the highest numbers of differentially regulated proteins between the control and diabesity conditions were binding and catalytic processes. Notably, transporter activity genes showed an upregulation in diabesity compared to the control, as illustrated in ( Figure 5C ). In diabesity, proteins responsible for signaling receptor activity, nutrient reservoir activity, fusogenic activity, cytoskeletal motor activity, and transcription coactivator activity were observed to be downregulated when compared to the control.

The interactome network was constructed using the STRING tool to analyze regulated proteins, investigating their interactions and predicting functional associations. The resulting interaction map of regulated proteins illustrates the relationships among molecular players involved in Toll-like receptor (TLR) activation by endogenous ligands, plasma lipoprotein remodeling, serine biosynthesis, LDL clearance, lipid transporter activity, and unblocking of NMDA receptors ( Figure 5D ). These proteins are interconnected, either directly interacting with one another or indirectly through their partners. Moreover, employing the STRING tool helped in identifying potential dysregulated KEGG pathways associated with diabesity. Subsequently, KEGG pathway analysis unveiled that differentially regulated proteins were predominantly associated with pathways such as the insulin/IGF pathway, Alzheimer’s disease pathway, nicotinic acetylcholine receptor signaling pathways, and the P53 pathway. The pathways most significantly regulated in diabesity are depicted in ( Figure 5E ).

In mice lacking collagen, the development of insulin resistance and glucose intolerance occurs, accompanied by elevated serum triglycerides and fat accumulation (75, 76). In C. elegans, deficiency in ATGL-1 results in elevated lipid content and shortened lifespan. Despite ATGL-1 being identified as a cytosolic lipase, nuclear morphology deterioration is not rescued upon upregulation of ATGL-1 (77). In our studies, we observed a noteworthy increase in lipid droplets and triglycerides within diabesity models ( Figure 2 ). The accumulation of triglycerides during diabesity is linked to ATGL-1 deficiency, preventing their hydrolysis and leading to their encapsulation in lipid droplets. Consequently, organelles undergo enlargement due to this accumulation.

In this study, one of the focuses was isolation and estimation of C. elegans collagen (see methods). Our investigation into diabesity revealed a significant reduction in collagen levels at the 4^th and 6^th days of adulthood compared to the control group. Specifically, control specimens exhibited 80 ng/50 mg (4^th day) and 120 ng/50 mg (6^th day), while diabesity-afflicted animals demonstrated 50 ng/50 mg and 70 ng/50 mg respectively ( Figure 6A ). Notably, no substantial changes were observed at day 2 of adulthood. These findings emphasize the influence of diabesity on collagen production within the C. elegans model, indicating a potential interplay between metabolic disorders and collagen regulation. The reduced collagen levels could lead to disruptions in anatomical structure and its integrity, due to sustained elevated levels of glucose and triglycerides. To validate these abnormalities, our microscopic analysis of control sections exhibited well-defined epidermal and cuticle layers, including specialized worm body structures such as the hemidesmosomes layer. Conversely, in diabesity, these specialized features like cuticles and hemidesmosomes were less distinct and displayed compromised mechanical strength ( Figures 6B, C ). Immunofluorescence staining was conducted to visualize the distribution and subcellular localization of ATGL-1 (see Methods). The results revealed a predominant presence of ATGL-1 in the epidermal region and hemidesmosomes in the control group. Conversely, in diabesity-induced conditions, its presence showed inconsistency across the cuticle layer, hypodermal areas, and distal gonad. These findings confirm the inhibition of ATGL-1’s lipolytic activity in diabesity-induced worms ( Figure 6D ).