Investigation of the effects and underlying mechanisms of adenylate kinase 1 on inosine monophosphate deposition in Beijing-You chickens

Abstract

Inosine monophosphate (IMP) is a major flavor compound in meat. Given the observed positive correlation between adenylate kinase 1 (AK1) expression and IMP content in Beijing-You chickens (BJYs), we investigated the role of AK1 in regulating myoblast proliferation and differentiation. Cell counting kit-8 (CCK-8) assays, 5-ethynyl-2′-deoxyuridine (EdU) incorporation, flow cytometry, and Western blotting showed that AK1 promoted myogenic differentiation and suppressed the proliferative activity of myoblasts. During the proliferation of myoblasts, glutamine phosphoribosylpyrophosphate amidotransferase (GPAT) was suppressed by AK1 and adenylosuccinate synthase (ADSS) was enhanced, which led to reduced IMP accumulation and inhibited uric acid (UA) and adenosine triphosphate (ATP) production. During myoblast differentiation, the protein expression of de novo IMP synthesis genes, including GPAT, 5-aminoimidazole-4-carboxamide ribonucleotide formyltransferase/IMP cyclohydrolase (PurH), and adenylosuccinate lyase (ADSL), was upregulated, while ADSS was downregulated by AK1, resulting in enhanced IMP deposition and stimulated production of UA and ATP. In summary, this study elucidates the regulatory role of AK1 in coordinating the proliferation and differentiation of myoblasts, as well as its stage-specific modulation of IMP anabolism. These findings provide theoretical insights into the molecular mechanisms governing IMP deposition and offer potential guidance for improving meat flavor quality in chicken through molecular breeding strategies.

Introduction

Inosine monophosphate (IMP), a crucial intermediate in purine nucleotide metabolism, plays an important role in intracellular energy homeostasis and signal transduction (Pedley and Benkovic, 2017; Rybalka et al., 2023). In livestock and poultry meat, IMP also serves as a key flavor precursor, interacting with amino acids, carbohydrates, and other compounds to enhance umami and overall taste characteristics (Lee et al., 2011; Tu et al., 2021; Zhang et al., 2021; Belloir et al., 2025). IMP is synthesized through three distinct pathways in living organisms (Huang et al., 2022a). The de novo synthesis pathway of IMP is a metabolic process that uses multiple amino acids, including glycine, aspartate, and glutamine, as well as one-carbon units carried by tetrahydrofolate (THF) and small molecules such as CO₂. These precursors undergo a series of enzymatic reactions to produce IMP. (Lovászi et al., 2021). This pathway represents the predominant biosynthetic route in vivo, where 5-phosphoribosyl pyrophosphate (5-PRPP) undergoes a 10-step enzymatic cascade to ultimately yield IMP (Gai et al., 2023). In the formation of 5-PRPP, adenosine triphosphate (ATP) undergoes dephosphorylation to generate adenosine diphosphate (ADP) and adenosine monophosphate (AMP), releasing energy that drives the de novo synthesis of IMP (Yin et al., 2018). The salvage synthesis pathway refers to the metabolic mechanism whereby livestock and poultry organisms utilize endogenous free purines or purine nucleosides as substrates to synthesize IMP through relatively simple enzymatic reactions (Zhao et al., 2024). Under conditions where muscle glycogen and high-energy phosphate bonds are available, ADP is phosphorylated to ATP in muscle tissue, maintaining transient ATP homeostasis. When glycogen stores and high-energy phosphate bonds are depleted, ATP synthesis ceases. Anaerobic glycolysis predominates, generating lactate and reducing intracellular pH. This acidification induces sarcoplasmic reticulum damage and promotes Mg-ATP complex formation, thereby activating ATP kinase, which catalyzes ATP catabolism to yield IMP. This pathway is relatively slow, resulting in prolonged retention of IMP in livestock, which plays a crucial role in meat flavor development (Shen et al., 2024). These three biosynthetic pathways involve several key enzymes, including glutamine phosphoribosylpyrophosphate amidotransferase (GPAT), 5-aminoimidazole-4-carboxamide ribonucleotide formyltransferase/IMP cyclohydrolase (PurH), adenylosuccinate lyase (ADSL), adenosine monophosphate deaminase 1 (AMPD1), and adenylosuccinate synthase (ADSS) (Datta et al., 1994; Gavalas and Zalkin, 1995; Lushchak, 1996; Shu et al., 2008; Dasgupta et al., 2018; Tran et al., 2024; Wang et al., 2025).

Adenylate kinase 1 (AK1), a pivotal enzyme in energy metabolism, catalyzes the interconversion between ATP and AMP and may regulate IMP levels through nucleotide metabolism regulation (Zeleznikar et al., 1990; Janssen et al., 2000; Noma, 2005). Huang et al. (2023) employed proteomic and metabolomic approaches to identify key proteins associated with IMP deposition in different tissues of Jingyuan chickens. The differentially expressed protein AK1 showed higher expression in pectoralis muscle, where IMP content was significantly higher than thigh muscle. Our previous study, based on integrated transcriptomic and metabolomic analysis revealed a significant positive correlation between AK1 transcription levels and IMP content in Beijing-You chickens (BJYs), though the precise regulatory mechanisms remain unclear. In this study, we aimed to elucidate the role of AK1 in IMP deposition, thereby providing a theoretical foundation for deciphering the regulatory mechanisms of IMP accumulation and facilitating molecular breeding strategies to enhance IMP content in chicken.

Materials and methods

Ethics statement

Animal welfare practices and experimental procedures were performed in accordance with the Guide for the Care and Use of Laboratory Animals. All procedures were approved by the Animal Ethics Committee of the Beijing University of Agriculture (Approval No. BUA81240311).

Cell Isolation, culture and differentiation

Muscle-derived cells were isolated from the pectoral muscle of Beijing-You chickens embryos at 11–12 days of embryogenesis following the method described by Guo et al. (2022), with minor modifications. The isolated cells were subsequently identified as myoblasts as described in the following section. The extracted muscle tissue was digested with a pre-prepared collagenase mixture, and the enzymatic reaction was terminated by adding complete culture medium. The resulting cell suspension was sequentially filtered, centrifuged, and the supernatant was discarded to retain the cellular pellet. The pellet was resuspended in complete medium (10 % fetal bovine serum (FBS) + 2 % penicillin-streptomycin (PS) + 88 % DMEM/F12). Fibroblast contamination was minimized via differential adhesion, and the purified myoblasts were cultured under sterile conditions at 37°C with 5 % CO_2. When the density of myoblasts reached approximately 90 % confluence, the cells were digested with 0.25 % trypsin-EDTA, centrifuged, and resuspended, followed by uniform seeding into 6-well plates. Upon reaching 80 % confluence, the growth medium was replaced with differentiation medium (2 % horse serum (HS) + 2 % PS + 96 % DMEM/F12), which was refreshed every 48 h.

Immunofluorescence assay

The complete medium was discarded from the well plate, and the plate was gently washed three times with phosphate-buffered saline (PBS) for 5 min each. Cells were fixed with 4 % paraformaldehyde (200 μL per well) for 30 min. The plate was washed three times with 1 mL PBS per well. Subsequently, 0.2 % permeabilization solution was added to permeabilize the cells, and the plate was washed again three times with PBS. PBS-diluted bovine serum albumin (BSA) was added to block non-specific binding, and the plate was incubated at room temperature. Primary antibodies against the myogenic marker Desmin and the nuclear transcription factor paired box 7 (Pax7) were added, and the plate was incubated overnight at 4°C. Afterward, the plate was washed three times with PBS, and goat anti-rabbit secondary antibody was added and incubated for 1 h at room temperature in the dark. Following three PBS washes (5 min each), 200 μL Hoechst 33342 staining solution was added per well, incubated for 10 min at room temperature in the dark, and the cells were thoroughly washed three times with PBS. The myoblasts were observed and recorded using an Olympus inverted fluorescence microscope.

AK1 overexpression and siRNA-mediated knockdown in myoblasts

Four candidate siRNA sequences targeting AK1 were designed and synthesized by GenePharma Co., Ltd. (Suzhou, China). The sequences of the siRNAs and the negative control (NC) are listed in Supplementary Table S1. Their knockdown efficiencies were evaluated by RT-qPCR and Western blot analysis, and the siRNA with the highest silencing efficiency was selected for subsequent experiments. The sequence of the selected si-AK1 was: sense strand 5′-CCAUCAUGGAGAAGGGCGATT-3′ and antisense strand 5′-UCGCCCUUCUCCAUGAUGGTT-3′. Specific primers were designed based on the chicken AK1 gene transcript (NM_205109.3) obtained from the NCBI GenBank database to amplify the target AK1 gene fragment via polymerase chain reaction (PCR). The PCR product and pcDNA3.1(+) vector were double-digested with type II restriction enzymes BamHI and EcoRI. Following purification, the fragments were directionally ligated using T4 DNA ligase to construct the recombinant plasmid pcDNA3.1(+)-AK1 vector. The constructed plasmid was then confirmed by DNA sequencing (Sangon Biotech, Shanghai, China), and the sequence matched the intended AK1 coding sequence perfectly (Supplementary Fig. S1).

Cell Transfection

Myoblasts were cultured in 6-well plates until reaching 80 % confluence. According to the manufacturer's protocol of Lipofectamine 3000 (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA), cells were transfected with either si-AK1, pcDNA3.1(+)-AK1, or corresponding controls (including negative controls). For siRNA transfection, 100 pmol siRNA and 3.75 μL Lipofectamine 3000 were diluted in 250 μL Opti-MEM and incubated for 10–15 min at room temperature before being added to the cells. For plasmid transfection, 2.5 μg plasmid DNA, 3.75 μL Lipofectamine 3000, and 5 μL P3000 reagent were diluted in 250 μL Opti-MEM and incubated for 10–15 min prior to transfection according to the manufacturer’s instructions. Three biological replicates were performed for each treatment condition.

Reverse transcription quantitative polymerase chain reaction (RT-qPCR)

Total ribonucleic acid (RNA) was extracted from cells using TRIzol® Reagent (Invitrogen, USA) according to the manufacturer’s instructions. The concentration and quality of RNA were determined using a multimode microplate reader (Thermo Fisher Scientific, USA) and samples with an A260/A280 ratio between 1.8 and 2.0 were considered suitable for downstream applications. Complementary DNA (cDNA) was synthesized from 1 μg of total RNA using a PCR thermal cycler (Eppendorf, DE). Quantitative real-time PCR was performed using Taq SYBR Green qPCR Premix (TransGen Biotech, Beijing, China) on an AriaMx Real-Time PCR System (Agilent, USA). The housekeeping gene GAPDH was used for normalization, and each reaction was performed in triplicate. Relative gene expression levels were calculated using the 2^−ΔΔCt method.

Quantification of IMP, uric acid (UA) and ATP

Following transfection, cells were cultured for 48 h in either complete medium or differentiation-inducing medium. Subsequently, the cells were washed twice with Dulbecco’s Phosphate-Buffered Saline (DPBS), detached using trypsinization, and centrifuged at 1,500 rpm for 5 min. The supernatant was carefully aspirated, and the cell pellet was resuspended in 200 µL of DPBS. IMP content was determined using an IMP Assay Kit (Enzyme Immunoassay, China; Cat. No. MM-6021001). ATP and UA levels were measured using commercial assay kits (ATP Assay Kit, Nanjing Jiancheng Bioengineering Institute, Nanjing, China; Cat. No. A095-1-1; UA Assay Kit, Nanjing Jiancheng Bioengineering Institute, Nanjing, China; Cat. No. C012-2-1) according to the manufacturers’ instructions. The reported recovery rate, precision, and detection limit of the kits meet the manufacturer’s validation standards. Finally, the absorbance of each test well was measured for quantitative analysis.

Western blot

Total proteins were extracted using a pre-prepared lysis buffer (Radio-Immunoprecipitation Assay (RIPA) buffer: phenylmethylsulfonyl fluoride (PMSF): 4 × Loading Buffer = 300:3:100). Following three washes with DPBS, cells were lysed in the buffer and subjected to three freeze-thaw cycles (10 min each). The lysates were centrifuged at 4°C, and then boiled at 100°C for 5 min. Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto polyvinylidene difluoride (PVDF) membranes. After blocking with 5 % non-fat milk, the membranes were incubated with primary antibodies overnight at 4°C, followed by incubation with species-matched horseradish peroxidase (HRP)-conjugated secondary antibodies. Immunoreactive bands were visualized using enhanced chemiluminescence (ECL) reagents and captured on X-ray films. The chemiluminescence intensity of target proteins was quantified using ImageJ software (NIH, USA) after background subtraction. The following primary antibodies were used for Western blot analysis: anti-AK1 (1:2000), anti-GPAT (1:5000), anti-ADSS (1:1000), anti-PurH (1:5000), anti-AMPD1 (1:2000), anti-GAPDH (1:10000), anti-MYOD (1:5000), and anti-MYHC (1:10000). All primary antibodies, except anti-ADSL (Bioss, 1:2000) and anti-Pax7 (Affinity, 1:1000), were purchased from Proteintech. The HRP-conjugated goat anti-rabbit IgG (H + L) secondary antibody (1:10000, Proteintech, SA00001-2) was used for detection.

Cell counting kit-8 (CCK-8) Cell viability assay

Myoblasts from BJYs were seeded in 96-well plates at a density of 1 × 10⁴ cells per well in 100 μL of complete medium. When cells reached 40-50 % confluence, they were transfected with either pcDNA3.1-AK1 or si-AK1, with three biological replicates per treatment group. At 24 h and 48 h post-transfection, 10 μL of CCK-8 solution was added to each well (1:10 dilution) under sterile conditions and protected from light. After incubation for 2 h at 37°C in a humidified incubator with 5 % CO₂, absorbance was measured at 450 nm using a microplate reader.

BeyoClick™ 5-ethynyl-2′-deoxyuridine (EdU)-488 cell proliferation assay

After 48 h of post-transfection culture, EdU solution (2 μL of 10 mM stock, 1:1000 dilution) was added to each well and incubated for 3 h at 37°C. The culture medium was removed, and cells were fixed with 1 mL of 4 % paraformaldehyde for 15 min at room temperature. Following fixation, cells were washed three times with PBS (1 mL / well, 3-5 min per wash) and permeabilized with 0.3 % Triton X-100 in PBS for 15 min at room temperature. After permeabilization, cells were washed twice with PBS (1 mL / well, 3-5 min per wash). The Click reaction mixture (500 μL / well) was added and incubated for 30 min at room temperature, protected from light. The reaction solution was removed, and cells were washed three times with PBS (1 mL / well, 5 min per wash). Nuclei were counterstained with 1 mL of 1 × Hoechst 33342 solution per well for 10 min at room temperature in the dark, followed by three additional PBS washes (1 mL / well, 5 min per wash). Fluorescence was detected using an inverted fluorescence microscope: proliferating myoblasts were visualized by green fluorescence (EdU labeling), while cell nuclei were identified by blue fluorescence (Hoechst 33342 staining).

Flow cytometric analysis of cell cycle

After 48 h of post-transfection culture, cells were washed twice with DPBS and detached by trypsinization. The cell suspension was collected into 1.5 mL centrifuge tubes and centrifuged at 1500 rpm for 5 min. The pellet was resuspended in approximately 1 mL of ice-cold PBS and pipetted gently to ensure homogeneity. After centrifugation, the supernatant was removed, leaving approximately 60 μL PBS in the tube. The cell pellet was resuspended by gently flicking the tube to prevent clumping. For fixation, 500 μL of pre-chilled 70 % ethanol (−20°C) was added to each tube containing myoblasts, followed by gentle mixing. The samples were fixed at 4°C for 12 h. After fixation, cells were pelleted by centrifugation at 1500 rpm for 5 min. The propidium iodide (PI) staining solution was freshly prepared by mixing 500 μL staining buffer, 25 μL (20X) PI solution, and 10 μL (50X) RNase A per tube. For staining, 500 μL of PI staining solution was added to each sample, followed by gentle mixing and incubation at 37°C for 30 min in the dark. Cell cycle analysis was performed using flow cytometry to detect red fluorescence (PI). The DNA content distribution was analyzed using FlowJo software to determine the percentage of cells in the G0/G1, S, and G2/M phases.

Statistical analysis

All experiments were performed with at least three independent biological replicates. For cell-based experiments, biological replicates represent myoblasts independently isolated from different BJYs embryos. Data were normalized and presented as mean ± SEM. Statistical analyses were conducted using SPSS 27.0 software. Differences between two groups were analyzed using Student’s t-test, while comparisons among multiple groups were performed using one-way analysis of variance (ANOVA) followed by Duncan’s multiple range test. P < 0.05 was considered statistically significant. Different lowercase letters indicate significant differences between groups, whereas the same letters indicate no significant difference.

Results

Identification and characterization of myoblasts in BJYs

The biological identification of myoblasts isolated from Beijing You chickens was verified (Supplementary Fig. S4). During myogenic induction, myoblasts gradually exhibited morphological changes associated with myogenic differentiation. The cells transitioned from spindle-shaped mononuclear cells (0 d) to densely arranged myoblasts (2–4 d), and by 6 d, clearly visible multinucleated myotubes had formed (Fig. 1A). The expression levels of myogenic marker genes (Pax7, myogenic differentiation 1 (MYOD), and myosin heavy chain (MYHC)) during the early differentiation stage were measured using RT-qPCR. The time points (0, 1.5, and 3 days) were selected to capture the transcriptional changes during the transition from proliferation to differentiation, as key myogenic regulators are known to be upregulated within the first few days. As shown in Fig. 1B, Pax7 transcription gradually decreased (P < 0.05), whereas MYOD and MYHC transcription progressively increased (P < 0.05). These expression patterns are consistent with the typical transcriptional dynamics of myoblasts during early myogenic differentiation.

Fig. 1.

Identification and characterization of myoblasts in Beijing-You chickens (BJYs). (A) Morphological characteristics of myoblasts at different differentiation stages (0 d, 2 d, 4 d, 6 d). (B) Transcriptional levels of Pax7, MYOD, and MYHC genes during myoblast differentiation.

Dynamic changes of IMP levels and associated gene expression during myoblast differentiation

Myoblasts were cultured in differentiation medium for 2-6 days. The IMP content progressively increased during cell differentiation (P < 0.05, Fig. 2A). Western blot analysis revealed dynamic expression patterns of AK1 and enzymes involved in IMP metabolism, including de novo synthesis enzymes (GPAT, PurH, ADSL), the salvage synthesis enzyme (AMPD1), and the degradation enzyme (ADSS). The expression level of AK1 was significantly upregulated during differentiation (P < 0.05) and showed a strong positive correlation with IMP accumulation (Pearson correlation analysis, R² = 0.88, P < 0.05, Fig. 2B and Supplementary Fig. S5). The protein expression profiles of key metabolic enzymes displayed distinct temporal patterns: GPAT and ADSS peaked on day 2, PurH reached maximal expression on day 6, and ADSL showed sustained upregulation throughout differentiation. Notably, AMPD1 displayed a biphasic pattern, with initial downregulation followed by upregulation, reaching its peak on day 6. (Fig. 2C and Supplementary Fig. S6).

Fig. 2.

Dynamic changes in IMP levels and related gene expression during myoblast differentiation. (A) Dynamic changes in IMP levels during myoblast differentiation. (B) Protein expression levels of AK1 at different differentiation stages of myoblasts. (C) Quantification of key enzymes involved in IMP metabolism, including de novo synthesis enzymes (GPAT, PurH, ADSL), the salvage pathway enzyme AMPD1, and the degradation enzyme ADSS, across different differentiation stages of myoblasts.

Regulatory role of AK1 in IMP metabolism

Myoblasts were transfected with si-AK1 or pcDNA3.1-AK1 to investigate the AK1-mediated regulation of IMP metabolism. The efficiency of AK1 knockdown (∼70 %) and overexpression (∼40-fold increase) was confirmed by Western blot analysis (Supplementary Fig. S2 and S3). Western blot analysis revealed distinct expression patterns of key metabolic enzymes (GPAT, PurH, ADSL, AMPD1, ADSS) during the proliferation and differentiation phases. During proliferation, AK1 knockdown significantly increased GPAT protein levels but decreased the expression of PurH, ADSL, AMPD1, and ADSS compared to controls (P < 0.05, Fig. 3A and Supplementary Fig. S7). Conversely, AK1 overexpression reduced GPAT but elevated PurH, ADSL, AMPD1, and ADSS levels (P < 0.05, Fig. 3B and Supplementary Fig. S8). During differentiation (48 h after the induction of differentiation), si-AK1 transfection downregulated GPAT, PurH, and ADSL while upregulated AMPD1 and ADSS (P < 0.05, Fig. 3C and Supplementary Fig. S9). The pcDNA3.1-AK1 group exhibited the inverse pattern, with elevated GPAT/PurH/ADSL and suppressed AMPD1/ADSS levels (P < 0.05, Fig. 3D and Supplementary Fig. S10). These results demonstrated that AK1 differentially regulated IMP deposition in myoblasts, suppressing accumulation during proliferation but promoting it during differentiation.

Fig. 3.

Regulatory role of AK1 in IMP metabolism. (A) Protein levels of key IMP metabolic enzymes (GPAT, PurH, ADSL, AMPD1, ADSS) after AK1 knockdown in proliferating myoblasts. (B) Protein levels of IMP metabolic enzymes following AK1 overexpression in proliferating myoblasts. (C) Protein levels of IMP metabolic enzymes after AK1 knockdown in differentiating myoblasts. (D) Protein levels of IMP metabolic enzymes following AK1 overexpression in differentiating myoblasts.

Impact of AK1 on IMP, UA, and ATP content

Myoblasts were transfected with si-AK1 or pcDNA3.1-AK1, cultured for 48 h, and then collected to measure intracellular IMP content. IMP measurement showed that AK1 knockdown significantly increased IMP content during the proliferation phase (P < 0.05) but decreased it during the differentiation phase (P < 0.05), whereas AK1 overexpression produced the opposite effects (P < 0.05, Fig. 4A). These results demonstrated AK1’s dual regulatory function, as it suppressed IMP accumulation during the proliferation phase and promoted its deposition during the differentiation phase. UA levels were determined enzymatically at 510 nm using a microplate reader. UA levels were significantly increased by si-AK1 during the proliferation phase (P < 0.05) but were markedly decreased during the differentiation phase (P < 0.05). In contrast, pcDNA3.1-AK1 markedly elevated UA levels during the differentiation phase (P < 0.05) but did not alter them during the proliferation phase (Fig. 4B). This indicated that AK1 inhibited UA production during proliferation but enhanced it during myogenic differentiation. ATP content analysis showed that AK1 knockdown increased ATP levels during the proliferation phase (P < 0.05) but decreased them during differentiation (P < 0.05), whereas AK1 overexpression produced the opposite effects (P < 0.05, Fig. 4C).

Fig. 4.

Impact of AK1 on IMP and energy metabolite homeostasis. (A) IMP levels in myoblasts following AK1 knockdown and overexpression. (B) UA levels in myoblasts following AK1 knockdown and overexpression. (C) ATP levels in myoblasts following AK1 knockdown and overexpression.

Impact of AK1 on myoblast proliferation and differentiation

Myoblasts were transfected with either si-AK1 or pcDNA3.1-AK1 constructs. The CCK-8 assay showed that AK1 knockdown significantly promoted cellular proliferation compared with both the Control and si-NC groups (P < 0.05), whereas AK1 overexpression inhibited proliferation, with the most pronounced suppression observed at 48 h post-transfection (P < 0.05, Fig. 5A). The EdU incorporation assay further confirmed that AK1 silencing enhanced proliferation (P < 0.05) while its overexpression exerted inhibitory effects (P < 0.05, Fig. 5B). Flow cytometric analysis of the cell cycle revealed that AK1 interference increased the proportions of cells in the G1 and G2 phase (P < 0.05) while reducing the S-phase fraction (P < 0.05). Conversely, AK1 overexpression decreased the G1 phase population (P < 0.05) and increased S-phase the proportion (P < 0.05, Fig. 5C), indicating that AK1 may regulate cell cycle progression via the S/G2 checkpoint. Western blot analysis of the myoblasts marker Pax7 revealed that AK1 knockdown significantly elevated Pax7 protein levels (P < 0.05), whereas AK1 overexpression markedly reduced it (P < 0.05), collectively suggesting AK1 played a suppressive role in myoblast proliferation. Following 48 h differentiation induction, si-AK1 transfection significantly decreased the expression of AK1, MYOD and MYHC protein (P < 0.05), whereas pcDNA3.1-AK1 transfection significantly increased their expression levels (P < 0.05) compared with controls (Fig. 5D and Supplementary Fig. S11). These results demonstrated that AK1 was able to inhibit the proliferation and promote the differentiation of myoblasts.

Fig. 5.

Effects of AK1 on the proliferation and differentiation of myoblasts. (A) CCK-8 assay evaluating the proliferation capacity of myoblasts following AK1 interference or overexpression. (B) EdU incorporation assay showing the influence of AK1 modulation on DNA synthesis activity in myoblasts. Scale bar: 200 μm. (C) Flow-cytometric analysis of cell-cycle distribution in myoblasts after AK1 knockdown or overexpression. (D) Expression levels of myogenic marker genes/proteins in myoblasts subjected to AK1 interference or overexpression.

Discussion

BJY is a well-known indigenous chicken breed in China, characterized by its tender and flavorful meat. Among the flavor-enhancing compounds, IMP, contributes significantly to the unique taste of BJYs. In the present study, we demonstrated that AK1 significantly reduced IMP levels during the proliferative phase of myoblasts, while promoting IMP accumulation during myotube differentiation. These findings indicate that AK1 acts as a critical regulator of purine metabolism in myoblasts and exerts stage-dependent control over IMP deposition during myogenesis. The study provides new insights into the molecular mechanisms underlying IMP accumulation and muscle cell differentiation.

In our previous transcriptomic and metabolomic analyses, we observed a significant positive correlation between AK1 mRNA levels and IMP content during the developmental stages of BJYs. In this study, primary myoblasts were isolated from BJYs and cultured in vitro. During the differentiation process, both IMP content and AK1 mRNA expression gradually increased as the differentiation period was prolonged. These results indicated that AK1 expression was positively correlated with IMP synthesis during cell differentiation. This is consistent with previous reports in chickens, where AK1 was identified as a key differentially expressed protein associated with IMP deposition in Jingyuan chicken muscles (Zhang et al., 2020; Huang et al., 2022b), and integrative multi-omics analysis further confirmed that AK1 enzymatic activity correlates with IMP accumulation (Huang et al., 2023).

To explore the role of AK1 in IMP deposition, AK1 was silenced or overexpressed in myoblasts. Protein levels of key IMP metabolic enzymes (GPAT, PurH, ADSL, AMPD1, and ADSS) were determined by Western blotting. The results showed that in the proliferation phase, AK1 markedly increased the protein levels of PurH, ADSL, AMPD1, and ADSS, but significantly reduced GPAT expression, thereby inhibiting IMP deposition. De novo IMP synthesis proceeds through a series of consecutive and highly conserved enzymatic reactions, each of which is indispensable for maintaining the integrity of the pathway. Among these, glutamine phosphoribosyl amidotransferase (GPAT) catalyzes the conversion of phosphoribosyl pyrophosphate (PRPP) and glutamine to phosphoribosylamine (PRA), which represents the first committed and rate-limiting step of this pathway. This reaction is irreversible and is regulated by strict feedback inhibition from the end products IMP, AMP, and guanosine monophosphate (GMP), thereby exerting central control over purine nucleotide homeostasis (Becker and Kim, 1987). Recent studies have shown that AK1, together with AMPD1 and other purine metabolic enzymes, forms a coordinated regulatory network that determines IMP deposition efficiency in chicken muscles (Zhao et al., 2023). Consequently, overall IMP accumulation was reduced, suggesting that although the expression of downstream enzymes such as PurH and ADSL could potentially facilitate metabolic flux, this effect was insufficient to overcome the blockade at the rate-limiting step. This further highlights the pivotal role of GPAT in controlling flux through the de novo IMP synthesis pathway. IMP metabolism at the branch point of purine nucleotide biosynthesis is tightly regulated by key enzymes, and inosine 5′-monophosphate dehydrogenase (IMPDH), a rate-limiting enzyme for guanine nucleotide synthesis, exerts fine control over IMP flux via allosteric modulation of nucleotide binding and conformational switching (Buey et al., 2022). In contrast, during the differentiation phase, AK1 overexpression significantly upregulated GPAT, PurH, and ADSL, indicating an overall increase in purine nucleotide synthetic capacity and establishing the metabolic basis for IMP accumulation. However, steady-state IMP levels are not solely determined by the rate of synthesis but are also influenced by branching pathways. Our data demonstrated that AK1 overexpression concomitantly downregulated ADSS, which catalyzes the rate-limiting step of IMP-to-AMP conversion, and this reduction directly decreases IMP utilization (Balberova et al., 2021; Andres-Hernando et al., 2023). Thus, the combined effect of enhanced upstream synthesis and restricted downstream consumption accounts for the significant IMP accumulation observed during differentiation, underscoring the central role of AK1 in coordinating purine metabolism and cell fate determination.

This study demonstrated that AK1 suppressed UA synthesis during the proliferative phase of myoblasts. Conversely, AK1 promoted UA accumulation during differentiation. The energy metabolic states differ markedly between the proliferative and differentiation phases. AK1, as a key adenylate kinase and regulator of purine metabolism, plays distinct roles at each stage. During proliferation, myoblasts undergo rapid DNA replication and protein synthesis, while maintaining an overall steady-state energy metabolism. AK1 catalyzes the reversible reaction 2ADP ↔ ATP + AMP to maintain the ATP/AMP balance, thereby ensuring intracellular energy homeostasis (Dzeja and Terzic, 2009). As a cytosolic AK isozyme highly expressed in skeletal muscle, AK1 plays a pivotal role in muscle energetic metabolism by regulating adenine nucleotide turnover, and its deficiency disrupts muscle energy homeostasis despite compensatory rearrangements of alternative phosphotransfer pathways (Fujisawa, 2023). At this stage, AK1 activity prevents excessive AMP accumulation, limiting its conversion to IMP and ultimately reducing UA production (Traut, 1994). Consequently, proliferation is characterized by relatively stable ATP levels and low concentrations of IMP and UA. Upon permanent withdrawal from the cell cycle, myoblasts shift toward myotube formation via enhanced transcriptional and translational programs, leading to a marked increase in energy and purine nucleotide demand. During differentiation, AK1 overexpression elevates AMP levels, thereby activating the AMP-activated protein kinase (AMPK) pathway (Hardie et al., 2012). Unlike its inhibitory role during proliferation, AMPK activation during differentiation does not suppress purine biosynthesis; instead, it promotes IMP synthesis and utilization through metabolic reprogramming. In addition, AK1 contributes to the regulation of extracellular ATP signaling, which facilitates differentiation-related intercellular communication. Therefore, the differentiation phase is characterized by enhanced ATP generation, elevated IMP accumulation, and increased UA production to meet both metabolic and signaling requirements. The physiological relevance of this metabolic remodeling is underscored by recent findings in human skeletal muscle. Smith et al. reported that IMP formation during muscle contractions acts as a vital mechanism to prevent excessive ADP accumulation and safeguard the cellular energy state (ΔG∼ATP∼) (Smith et al., 2025). In the context of our study, the enhanced IMP deposition driven by AK1 during myoblast differentiation may reflect a similar, albeit chronic, adaptation to meet the high energy demands of myotube formation. By promoting IMP synthesis, AK1 may help maintain a favorable energy balance that supports the extensive protein synthesis and cytoskeletal reorganization required for myogenesis, thus linking nucleotide metabolism to cell fate determination.

The proliferation and differentiation of myoblasts are essential for muscle growth and regeneration. However, the role of AK1 in these processes remains unclear. In this study, during proliferation, AK1 overexpression caused S-phase arrest and reduced Pax7 expression, indicating that AK1 inhibits myoblast proliferation through energy-dependent checkpoint regulation. As a key enzyme maintains adenine nucleotide balance (2ADP ↔ ATP + AMP), excessive AK1 activity may alter the ATP/ADP/AMP ratio and activate AMPK, which in turn phosphorylates p53 to trigger replication stress and S-phase arrest (Jones et al., 2005; Zervou et al., 2021). Furthermore, decreased Pax7 expression suggested a shift from self-renewal toward differentiation, consistent with its known role in maintaining satellite cell identity (Olguin and Olwin, 2004; Von Maltzahn et al., 2013). Interestingly, despite its reported pro-proliferative role in non-muscle and tumor cells (De Bruin et al., 2004), AK1 inhibited proliferation in myoblasts. This discrepancy may stem from cell type–specific differences in metabolic programming, signaling networks, or purinergic receptor distribution, suggesting that AK1 exerts a pronounced context-dependent function (Klepinin et al., 2020). Moreover, whether the proliferation-inhibitory effect of AK1 was primarily mediated by AMPK activation or by AMP deamination into IMP and subsequent UA production remains to be clarified. Future studies employing integrated metabolic and signaling analyses could help to disentangle these mechanisms.

During differentiation, AK1 overexpression upregulated MYOD and MYHC expression, thereby promoting myoblast differentiation, whereas AK1 knockdown impaired this process and hindered myotube maturation. This effect may be attributed to the role of AK1 in sustaining cellular energy homeostasis and facilitating ATP regeneration required for myogenic differentiation. AK1-derived AMP can activate AMPK, which promotes mitochondrial biogenesis and oxidative metabolism—both are essential for terminal differentiation of muscle cells (Fu et al., 2015). Furthermore, enhanced ATP turnover by AK1 supports the biosynthetic and cytoskeletal remodeling processes necessary for myotube fusion and maturation (Dzeja and Terzic, 2009). Taken together, these findings suggest that AK1 promotes myogenic differentiation by coupling energy metabolism with the transcriptional activation of muscle-specific genes. During C2C12 myogenesis, AK1 levels increased, leading to a significant enhancement of cellular ATP synthesis (Choo et al., 2008). These findings are highly consistent with our observations during the differentiation phase. Additionally, AK1-deficient mice exhibited reduced muscle energy efficiency and showed disrupted nucleotide homeostasis (Janssen et al., 2000), further supporting the role of AK1 as a key regulator of energy metabolism. Collectively, these findings indicate that AK1 functions as a molecular switch that couples cellular energy metabolism with the transition from proliferation to differentiation. By regulating adenine nucleotide balance and AMPK activation, AK1 provides a metabolic basis for skeletal muscle regeneration and plays a dual role in determining the fate of myoblasts. Based on these findings, a schematic model illustrating the stage-specific regulatory role of AK1 in myoblast proliferation, differentiation, and IMP deposition is proposed (Fig. 6).

Fig. 6.

Proposed regulatory schematic of AK1 during the proliferation and differentiation of chicken myoblasts. The diagram illustrates the stage-specific regulatory effects of AK1 on key enzymes (ADSS and GPAT) and the metabolite IMP. The left panel represents the proliferation phase, while the right panel represents the differentiation phase. Arrows indicate the regulatory relationships among AK1, enzymes, and metabolites, where “→” denotes promotion and “—|” denotes inhibition. This schematic summarizes the potential role of AK1 in modulating IMP deposition during different stages of myogenic development.

In addition to its mechanistic insights, this study has potential implications for broiler molecular breeding and meat quality improvement. AK1 may serve as a key regulatory gene for balancing breeding strategies aimed at improving both growth performance and meat flavor in chickens. Furthermore, understanding the regulatory network of IMP deposition may inform nutritional or management strategies to enhance muscle flavor compounds. In this study, we primarily used in vitro-cultured chicken myoblasts to explore the role of AK1 in IMP deposition, which may not entirely reflect the situation in vivo, because in vitro systems lack the complex physiological environment, including hormonal regulation, cell–cell interactions, and systemic metabolic processes present in living organisms. To address this limitation, future studies could employ gene-edited chicken models, such as conditional knockout or overexpression systems, to validate the regulatory role of AK1 in IMP deposition at the in vivo level. In addition, large-scale population analyses across different chicken breeds (e.g., fast-growing commercial broilers and high-quality indigenous yellow-feathered chickens) could be conducted to screen SNPs within the AK1 gene and its promoter region, with the aim of identifying advantageous haplotypes associated with high IMP deposition without compromising growth performance. Furthermore, the identified favorable AK1 markers could be integrated into molecular breeding chips to facilitate marker-assisted selection for improved meat flavor in chickens.

Conclusion

In conclusion, this study revealed the regulatory role of AK1 in IMP deposition in chicken myoblasts and further demonstrated its dual function in inhibiting proliferation while promoting differentiation. These findings suggest that AK1 exhibits a pleiotropic effect during chicken muscle development and indicate a potential link between cellular metabolism and cell fate regulation. These findings also offer a theoretical basis for understanding the regulatory mechanisms of IMP deposition and offer support for future breeding strategies aimed at improving IMP content. Notably, the function of AK1 appears to be cell type-dependent, highlighting the need for further mechanistic studies to clarify its pathway-specific effects and potential roles in skeletal muscle regeneration and IMP deposition.

Declaration of generative AI and AI-assisted technologies in the writing process

The authors did not use any artificial intelligence assisted technologies in the writing process.

CRediT authorship contribution statement

Jingyan Hou: Writing – original draft, Visualization, Formal analysis, Data curation. Yao Zhang: Visualization, Methodology, Data curation. Liyang Zhu: Writing – review & editing, Visualization. Xiaolong Qi: Writing – review & editing. Xiangguo Wang: Writing – review & editing. Zili Lin: Writing – review & editing. Longfei Xiao: Writing – review & editing. Cheng Long: Writing – review & editing. Jinhuan Dou: Writing – review & editing. Yaxi Xu: Writing – review & editing. Xihui Sheng: Methodology, Funding acquisition, Conceptualization.

Disclosures

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.