Abstract

   Skeletal muscle atrophy is a prevalent complication of chronic kidney
   disease (CKD) and serves as an indicator of adverse prognosis and poor
   quality of life; however, the underlying mechanisms remain ambiguous.
   Emerging evidence has shown that long non-coding RNAs (lncRNAs) are
   involved in the pathogenesis of skeletal muscle atrophy. Using RNA
   sequencing (RNA-seq), we discerned elevated GAS5 expression in the
   muscles of CKD mice and verified these findings by real-time qPCR.
   Transmission electron microscopy confirmed morphological signs of
   pyroptosis, a potentially causal cellular death form. Additionally,
   elevated levels of pyroptosis markers, such as NLRP3, cleaved
   caspase-1, and GSDMD-N, were observed in CKD mouse models and
   lipopolysaccharide (LPS)/ATP-stimulated C2C12 myotubes. Intriguingly,
   the knockdown of GAS5 reduced these markers, alleviating pyroptosis and
   enhancing myofiber size, both in vitro and in vivo. Furthermore, we
   pinpointed an interaction between GAS5 and the mitochondrial
   translation elongation factor (TUFM) through RNA pull-down and mass
   spectrometry. This interaction amplified NLRP3 activity, contributing
   to pyroptosis and muscle atrophy. Notably, overexpressing TUFM
   counterbalanced this effect. Fundamentally, the interaction between
   GAS5 and TUFM appears to compromise the anti-pyroptosis capacity of
   TUFM. Consequently, this amplifies the activation of the NLRP3 pathway,
   which may underpin the crucial mechanism driving pyroptosis-mediated
   muscle atrophy. Our findings provide new evidence for GAS5’s role in
   regulating cellular pyroptosis in CKD-induced skeletal muscle atrophy.

   Keywords: MT: Non-coding RNAs, chronic kidney disease, CKD, skeletal
   muscle atrophy, long non-coding RNA, lncRNA, pyroptosis, NLRP3

Graphical abstract

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     __________________________________________________________________

   The expression of lncRNA GAS5 is elevated in CKD-induced skeletal
   muscle atrophy. An interaction between GAS5 and TUFM amplifies the
   activity of the NLRP3 pathway, contributing to cellular pyroptosis and
   muscle atrophy. GAS5 knockdown alleviates pyroptosis and attenuates
   muscle atrophy in CKD.

Introduction

   Chronic kidney disease (CKD) poses a significant public health
   challenge that affects >10% of the general population worldwide,
   amounting to >800 million individuals. CKD has emerged as one of the
   leading causes of death globally, with its mortality rate increasing
   over the past few decades. Importantly, CKD represents a large medical
   burden in many countries.[30]^1 Skeletal muscle atrophy is a common
   complication of CKD, characterized by a decline in skeletal muscle mass
   and diminished muscle functionality.[31]^2 As an independent
   risk factor for mortality, skeletal muscle atrophy is closely
   associated with the quality of life and adverse prognosis in patients
   with CKD. However, the mechanisms leading to muscle atrophy in patients
   with CKD remain unclear, and treatment options are very limited.

   Various molecular mechanisms contribute to the development of muscle
   atrophy. Evidence suggests that cell death may play a pivotal role in
   inducing muscle atrophy.[32]^3^,[33]^4 Pyroptosis, a novel form of
   programmed cell death that follows apoptosis and necrosis, is
   characterized by the production of oligomeric N-terminal fragments by
   the protein GSDMD. This process leads to the formation of pores on the
   host cell membrane (CM), resulting in cell lysis and death.[34]^5
   The most typical pyroptosis activation pathway is mediated by
   nucleotide-binding oligomerization domain (NOD)-like receptors
   (NLRs).[35]^6 The successive activation of NLRP3 and caspase-1
   facilitates the cleavage of gasdermin D, releasing inflammatory
   cytokines interleukin (IL)-1β and IL-18.[36]^7^,[37]^8^,[38]^9 It has
   been reported that NLRP3-mediated pyroptosis can lead to
   dexamethasone-induced skeletal muscle atrophy, and knocking down NLRP3
   can alleviate this response.[39]^10 Similarly, in AngII-induced
   skeletal muscle atrophy, the knockout of NLRP3 also demonstrated a
   protective effect against muscle atrophy.[40]^11 Recent studies have
   indicated that pyroptosis plays a key role in diabetic-induced skeletal
   muscle atrophy and muscle weakness.[41]^12 An increasing number of
   reports indicate that pyroptosis is involved in the mechanism of muscle
   atrophy. Still, few studies comprehensively reveal the role of
   pyroptosis in CKD-induced skeletal muscle atrophy.

   Long non-coding RNAs (lncRNAs) are a class of nucleotide RNAs that do
   not encode proteins and have a transcription length exceeding 200
   nt.[42]^13 Studies have shown that lncRNAs play vital regulatory roles
   in various biological processes, such as apoptosis, metabolism, and the
   cell cycle.[43]^14 An increasing body of evidence also indicates that
   lncRNAs exert crucial functions in various disease processes, such as
   inflammatory diseases, cancer, and cardiovascular diseases, by
   regulating pyroptosis.[44]^15^,[45]^16^,[46]^17 Accumulating evidence
   supports that lncRNAs may play direct or indirect roles upstream of the
   pyroptosis trigger pathway.[47]^18 Recent research showed that lncRNA
   TINCR could promote pyroptosis by positively regulating NLRP3 in
   diabetic cardiomyopathy, while METTL14 suppresses pyroptosis and
   diabetic cardiomyopathy by downregulating TINCR lncRNA.[48]^19 The
   aberrant expression of lncRNAs is associated with the pathogenesis of
   muscle atrophy. Our previous research demonstrated that lncRNA
   Atrolnc-1 is involved in the occurrence of muscle atrophy in
   CKD.[49]^20 However, the underlying mechanism of lncRNAs regulating the
   pyroptosis pathway in muscle atrophy with CKD is not clear.

   In this study, we identified a novel CKD-associated lncRNA and
   clarified its function in muscle atrophy. The lncRNA GAS5 exhibited a
   markedly elevated expression in the CKD muscle atrophy model. We also
   found that after GAS5 interacts with TUFM, it enhances NLRP3 activity,
   activates the pyroptosis pathway, and leads to the occurrence of
   pyroptosis. These findings provide new evidence for GAS5’s role in
   regulating cellular pyroptosis in CKD-induced skeletal muscle atrophy
   and offer new insights for the prevention and treatment of muscle
   atrophy.

Results

Identification of differentially expressed lncRNAs in skeletal muscle atrophy
of CKD mice

   To probe the potential role of lncRNAs in CKD-associated skeletal
   muscle atrophy, we utilized RNA sequencing (RNA-seq) analysis on
   quadriceps femoris muscles from control and CKD mice. We induced CKD in
   12-week-old C57BL6 mice via a 5/6 nephrectomy procedure. Four weeks
   post-operation, when compared with sham-operated controls, CKD mice
   manifested elevated serum creatinine levels ([50]Figure 1A) and a
   significant decrease in both body and individual gastrocnemius muscle
   weights ([51]Figures 1B and 1C). Histological evaluation using
   hematoxylin and eosin (H&E) staining highlighted a pronounced reduction
   in the muscle cross-sectional area (CSA) in CKD mice compared to the
   results of controls ([52]Figure 1D, top). Further analysis of the CSA
   showcased a left-shifted muscle fiber distribution in the CKD group
   ([53]Figure 1D, bottom). Corresponding with these protein energy
   wasting (PEW) phenotypic changes, we observed elevated expressions of
   the atrophy-associated genes MuRF-1 and Atrogin-1 in the quadriceps
   femoris muscles of CKD mice, as evidenced by 6- and 5-fold increases,
   respectively, in the western blot analysis ([54]Figure 1E). After
   post-validation of muscle atrophy phenotypes in CKD mice, we delved
   into the lncRNA profiles within their muscles. The volcano plot
   provided an overview of all differentially expressed lncRNAs
   ([55]Figure 1F). Using criteria that required at least two counts per
   million (CPM) >0 in each group and a false discovery rate (FDR) < 0.01,
   we identified 13 lncRNAs differentially expressed in CKD mice muscles.
   Of these, 11 were upregulated, and 2 were downregulated, as illustrated
   in the heatmap ([56]Figure 1G). Prioritizing lncRNAs of potential
   clinical significance to humans, we focused on homologous molecules
   between human and mouse lncRNA pools ([57]Figure S1). This led to the
   exclusion of 11 non-human-specific lncRNAs. Among the remaining two
   lncRNAs, GAS5 emerged as the most significantly upregulated lncRNA
   shared between humans and mice, pinpointing it as our primary molecule
   of interest. To substantiate our RNA-seq findings, we validated GAS5
   expression levels using real-time qPCR in skeletal muscle tissues from
   both the CKD mouse models and patients with end-stage renal disease
   (ESRD). This confirmed the pronounced elevation of GAS5 expression in
   both groups ([58]Figures 1H and 1I). Collectively, our results
   underscore the elevated expression of GAS5 in the atrophic skeletal
   muscles of CKD mice. This suggests a potential regulatory interplay
   between GAS5 expression and CKD-induced muscle atrophy.

Figure 1.

   [59]Figure 1
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   Identification of differentially expressed lncRNAs in skeletal muscle
   atrophy of CKD mice

   (A) Serum creatinine levels in sham mice and CKD mice (n = 6,
   ∗∗p < 0.01). (B) Comparison of body weights between sham mice and CKD
   mice (n = 6, ∗∗p < 0.01). (C) Comparison of gastrocnemius muscle
   weights between sham mice and CKD mice (n = 6, ∗∗p < 0.01). (D) H&E
   staining shows a reduction in muscle cross-sectional area (CSA) in CKD
   mice (top). Scale bar: 50 μm. The distribution of myofiber sizes in CKD
   mice was shifted leftward (bottom). (E) Western blot confirmed the
   expression levels of MuRF-1 and Atrogin-1 in muscles of CKD mice (n =
   6, ∗∗p < 0.01). (F) The volcano plots display all differentially
   expressed lncRNAs. (G) Heatmap shows the top 13 differentially
   expressed lncRNAs in quadriceps muscles of CKD mice and sham mice. (H)
   Real-time qPCR confirmed the upregulation of GAS5 and VAULT in CKD mice
   (n = 5, ∗p < 0.05 and ∗∗p < 0.01). (I) Real-time qPCR confirmed the
   upregulation of GAS5 and VAULT in muscles of patients with ESRD (n = 5,
   ns: no statistically significant difference, ∗p < 0.05).

Unraveling the protein interactome of GAS5 in CKD muscle

   After observing elevated levels of GAS5 in the muscles of CKD mice, we
   aimed to uncover its potential role in muscle atrophy. We first
   assessed the cellular distribution of GAS5 by fluorescence in situ
   hybridization (FISH) on C2C12 cells and mouse muscle tissues. Using a
   FAM-labeled Gas5 probe, we found that GAS5 is primarily localized in
   the cytoplasm, although a small fraction is also present in the
   nucleus. This dual localization suggests that GAS5 may have roles in
   both signaling and gene regulation ([61]Figure 2A). Given that lncRNAs
   are known to influence signaling pathways by interacting with specific
   proteins,[62]^21^,[63]^22 we hypothesized that GAS5 might function
   through its association with certain proteins. To explore this, we
   performed RNA pull-down experiments using C2C12 myotube extracts
   ([64]Figure 2B), followed by mass spectrometry (MS) to identify
   proteins interacting with the biotin-labeled GAS5 probe. Our analysis
   revealed that GAS5 is associated with over 45 proteins ([65]Figure 2C;
   [66]Table S1). Subsequent pathway enrichment analysis using KEGG
   highlighted a significant involvement of GAS5-associated proteins in
   the NLR signaling pathway, which is essential for inflammasome
   formation and chronic inflammatory responses ([67]Figure 2D). Among the
   proteins, TUFM caught our attention due to its known role in inhibiting
   NLRP3 inflammasome formation.[68]^23 To confirm this interaction, we
   used immunoblotting with an anti-TUFM antibody. A distinct protein
   band, corresponding to the expected molecular weight of TUFM, was found
   among the proteins associated with biotinylated GAS5 ([69]Figure 2E).
   Furthermore, RNA immunoprecipitation (RIP) experiments were performed
   with an antibody against TUFM or negative immunoglobulin (Ig)G, and
   immunoprecipitation using anti-TUFM antibodies from C2C12 myotube
   lysates followed by real-time qPCR showed significant enrichment of
   GAS5, whereas control (IgG) precipitates displayed undetectable levels
   of GAS5 ([70]Figure 2F). Collectively, these results demonstrated a
   direct interaction between GAS5 and TUFM and point to a potential role
   for GAS5 in regulating NLRP3-associated pathways, particularly in the
   context of CKD-induced muscle atrophy.

Figure 2.

   [71]Figure 2
   [72]Open in a new tab

   Unraveling the protein interactome of GAS5 in CKD muscle

   (A) Fluorescence in situ hybridization(FISH) analysis of C2C12 cells
   and muscles from mice. GAS5-positive signal manifests as green
   fluorescence. (B) SDS-PAGE displays the results of RNA pull-down assay.
   (C) Mass spectrometry (MS) identified proteins that specifically
   interact with GAS5. (D) KEGG pathway enrichment analysis indicated
   signaling pathways linked to GAS5 expression. (E) A representative
   immunoblotting shows a band at 50 kD using TUFM antibody in the
   proteins pulled down by biotin-labeled GAS5 probe. (F) Real-time PCR
   was used to quantify GAS5 in immunoprecipitates (top). Western blot
   shows a band at 50 kD in the immunoprecipitation (IP) product using
   TUFM antibody (bottom).

Linking GAS5 upregulation with pyroptosis and atrophic responses in C2C12
myotubes

   Given that activation of NLRP3-associated pathways frequently
   culminates in pyroptosis across various cell types, we aimed to devise
   an in vitro model of pyroptosis using C2C12 myotubes. The
   lipopolysaccharide (LPS)/ATP stimulation method is widely used in
   research to induce an inflammatory response, activate immune signaling
   pathways, and trigger relevant intracellular mechanisms, particularly
   through activation of the NLRP3 pathway to further enhance
   inflammation. Utilizing a well-documented approach, we treated C2C12
   myotubes with LPS (100 ng/mL; Sigma) for 2 h, followed by ATP (5.0 mM;
   Med Chem Express‌) for another 1 h to induce intracellular inflammatory
   responses.[73]^10 Subsequent analysis via western blotting revealed a
   marked elevation in the expression of pyroptosis-related proteins.
   Specifically, levels of NLRP3, cleaved caspase-1, GSDMD, GSDMD-N, and
   IL-1β showed significant increases, with GSDMD expression being the
   most prominent ([74]Figure 3A). To further corroborate the occurrence
   of pyroptosis, we employed Hoechst 33342/PI double staining, which
   confirmed significant pyroptotic activity in the LPS/ATP-stimulated
   C2C12 myotubes ([75]Figure 3B). Importantly, this LPS/ATP stimulation
   also led to a robust upregulation of GAS5 expression ([76]Figure 3C).
   Additionally, to assess the potential link between NLRP3 pathway
   activation and myotube atrophy, we observed a notable reduction in the
   diameter of C2C12 myotubes following LPS/ATP stimulation
   ([77]Figure 3D). Concurrently, there was a pronounced increase in the
   expression of muscle-atrophy-associated proteins, MuRF-1 and Atrogin-1
   ([78]Figure 3E). In summation, our established in vitro model
   effectively linked NLRP3 pathway activation to myotube atrophy.
   Furthermore, this model offers a valuable platform to further
   investigate the role of GAS5 in regulating both pyroptosis and myotube
   atrophy processes.

Figure 3.

   [79]Figure 3
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   Linking GAS5 upregulation with pyroptosis and atrophic responses in
   C2C12 myotubes

   (A) The protein levels of NLRP3, cleaved caspase-1, GSDMD, GSDMD-N, and
   IL-1β were detected in C2C12 myotubes by western blot (n = 6, ∗p < 0.05
   and ∗∗p < 0.01). (B) Detection of pyroptotic activity in the
   LPS/ATP-stimulated C2C12 myotubes by dual staining with Hoechst 33342
   and PI. (C) Real-time qPCR result shows increased GAS5 mRNA level in
   the LPS/ATP-stimulated C2C12 myotubes (n = 3, ∗∗p < 0.01). (D)
   Fluorescence images display notable reduction in the diameter of C2C12
   myotubes following LPS/ATP stimulation (top). Scale bar: 100 μm.
   Graphic presentation of myotube diameter in different groups (bottom).
   (E) Western blot analysis confirmed the upregulation of MuRF-1 and
   Atrogin-1 expression levels in the LPS/ATP-stimulated C2C12 myotubes
   (n = 6, ∗p < 0.05 and ∗∗p < 0.01).

Knockdown of GAS5 improves LPS/ATP-induced pyroptosis and atrophy in C2C12
myotubes

   To investigate the regulatory role of GAS5 in myotube atrophy and
   pyroptosis, we employed an adeno-associated virus 1 (AAV1) vector
   bearing shGAS5 to knock down GAS5 in C2C12 myotubes. We first ensured
   the transfection efficiency by using an AAV1 vector with GFP (AAV1-GFP)
   and observed that nearly 100% of the myofibers expressed GFP
   ([81]Figure S2). Following this, we transfected C2C12 myotubes with
   either control AAV1 (AAV1-CTL) or AAV1-shGAS5 for 48 h. Subsequent
   treatment with LPS/ATP for 3 h was employed to induce pyroptosis. Using
   real-time qPCR, we observed that LPS/ATP-induced upregulation of GAS5
   was significantly inhibited in the AAV1-shGAS5 transfected myotubes,
   with GAS5 levels decreasing by approximately 90% compared to the
   AAV1-CTL-transfected myotubes ([82]Figure 4A). Further investigation
   using immunoblot analysis revealed that the LPS/ATP-induced expression
   of both atrophy-related proteins (MuRF-1 and Atrogin-1) and
   pyroptosis-associated proteins, including NLRP3, cleaved caspase-1,
   GSDMD, GSDMD-N, and the inflammatory cytokine IL-1β, was markedly
   suppressed upon GAS5 knockdown ([83]Figure 4B). We further found that
   LPS/ATP-induced reduction in the diameter of C2C12 myotubes was
   partially improved by GAS5 knockdown ([84]Figure 4C). In addition,
   Hoechst 33342/PI double staining highlighted a significant reduction in
   LPS/ATP-induced pyroptosis in the GAS5-knockdown C2C12 cells compared
   to the controls ([85]Figure 4D). In conclusion, our findings
   demonstrate that GAS5 knockdown effectively ameliorates both cellular
   pyroptosis and myotube atrophy induced by LPS/ATP in C2C12 myotubes.

Figure 4.

   [86]Figure 4
   [87]Open in a new tab

   Knockdown of GAS5 improves LPS/ATP-induced pyroptosis and atrophy in
   C2C12 myotubes

   (A) Real-time qPCR confirmed the downregulation of GAS5 in C2C12
   myotubes transfected with AAV1-shGAS5 (n = 3, ∗∗p < 0.01). (B) The
   protein levels of MuRF-1, Atrogin-1, NLRP3, cleaved caspase-1, GSDMD,
   GSDMD-N, and IL-1β were measured by western blot under different
   treatment conditions (n = 3, ∗∗p < 0.01 vs. control+AAV1-CTL; #p < 0.05
   and ##p < 0.01 vs. LPS+ATP+AAV1-CTL). (C) Fluorescence images display
   an improvement in the diameter of LPS/ATP-stimulated C2C12 myotubes
   transfected with AAV1-shGAS5 (top). Scale bar: 100 μm. Graphic
   presentation of myotube diameter under different treatment conditions
   (bottom). (D) Detection of pyroptotic activity in C2C12 myotubes under
   different treatment conditions by dual staining with Hoechst 33342 and
   PI.

TUFM overexpression counteracts GAS5-driven NLRP3 activation and pyroptosis
in C2C12 myotubes

   To understand the intricate role of TUFM in regulating NLRP3 activity,
   we employed an AAV1 to overexpress the TUFM gene (AAV1-TUFM). Given
   existing literature highlighting TUFM’s inhibitory effects on
   NLRP3-mediated pyroptosis and our observations of GAS5’s interaction
   with TUFM, we hypothesized that GAS5 might negate this inhibition.
   Following transfection of C2C12 myotubes with AAV1-TUFM, we noted an
   11-fold surge in TUFM mRNA levels ([88]Figure 5A). Immunoblot analysis
   revealed that the expression of TUFM was markedly increased upon TUFM
   overexpression, indicating effective transfection ([89]Figure 5B). This
   process resulted in diminished pyroptosis-related protein levels in the
   AAV1-TUFM transfected myotubes; concurrently, MuRF-1 and Atrogin-1
   expression levels also showed a decline ([90]Figure 5C). Complicating
   the narrative, our data revealed diminished TUFM levels in both CKD
   mice and LPS/ATP-stimulated C2C12 myotubes when compared to their
   respective controls ([91]Figure 5D). Concomitant with these results, an
   LPS/ATP-induced reduction in the diameter of C2C12 myotubes could be
   improved by TUFM overexpression ([92]Figure 5E). The interaction
   between GAS5 and TUFM may present dual outcomes. On one hand, GAS5
   seems to counteract the inhibitory effect of TUFM. On the other hand,
   GAS5 might compromise the stability of TUFM. In aggregate, these
   impacts underscore the central role of the GAS5-TUFM interaction in
   catalyzing the NLRP3-mediated pyroptosis pathway. Overall, our findings
   shed light on the intricate interplay between GAS5 and TUFM,
   elucidating their potential roles in modulating NLRP3-mediated
   pyroptosis, a possible factor in CKD-induced muscle atrophy.

Figure 5.

   [93]Figure 5
   [94]Open in a new tab

   TUFM overexpression counteracts GAS5-driven NLRP3 activation and
   pyroptosis in C2C12 myotubes

   (A) Real-time qPCR result shows increased TUFM mRNA levels after the
   overexpression of TUFM in C2C12 myotubes (n = 3, ∗∗p < 0.01). (B)
   Western blot confirmed the protein level of TUFM was increased after
   the overexpression of TUFM in C2C12 myotubes (n = 3, ∗∗p < 0.01). (C)
   The protein levels of MuRF-1, Atrogin-1 NLRP3, cleaved caspase-1,
   GSDMD, GSDMD-N, and IL-1β were measured by western blot under different
   treatment conditions (n = 3, ∗p < 0.05 and ∗∗p < 0.01 vs.
   control+AAV1-CTL; #p < 0.05 and ##p < 0.01 vs. LPS+ATP+AAV1-CTL). (D)
   The downregulation of TUFM was confirmed by immunoblots in CKD mice
   (left) and LPS/ATP-treated C2C12 myotubes (right) (n = 3, ∗p < 0.05 and
   ∗∗p < 0.01). (E) Fluorescence images display an improvement in the
   diameter of LPS/ATP-stimulated C2C12 myotubes by TUFM overexpression
   (top). Scale bar: 100 μm. Graphic presentation of myotube diameter in
   different groups (bottom).

Evidence linking pyroptosis to muscle atrophy in CKD mice

   Previous studies have implicated inflammation, particularly pyroptosis,
   in the progression of muscle atrophy.[95]^24 To investigate the
   presence of pyroptosis in skeletal muscles of CKD mice, we employed
   transmission electron microscopy (TEM). Our observations revealed
   marked alterations in the skeletal muscles of CKD mice compared to the
   sham control group. Specifically, we noted increased occurrences of
   membrane rupture and mitochondrial damage; large sections of the
   skeletal muscle CM displaying discontinuities, forming extensive holes
   (highlighted with red arrows in [96]Figure 6A); swelling and
   enlargement of organelles, partial rupture of muscle fibers (denoted by
   yellow arrows in [97]Figure 6A), and areas of muscle fiber fusion and
   thickening; symmetrically distributed sarcomeres, with the majority of
   myofibrillar filaments exhibiting dense structures and minor areas
   displaying ruptures; and mitochondria (M) interspersed among muscle
   fibers, characterized by significant swelling, matrix dissolution, and
   vacuolation ([98]Figure 6A, left). We found that the nuclei (N) in
   muscle cells are irregularly shaped and that the nuclear membrane
   remains intact in the CKD group; we also noticed the condensation of
   chromatin ([99]Figure 6A, blue arrows), while in the sham control
   group, the nuclear chromatin is evenly distributed ([100]Figure 6A,
   right). These morphological features are characteristic of pyroptosis.
   To corroborate our morphological findings, we probed for molecular
   markers of pyroptosis in CKD mouse muscles. Western blot analysis
   demonstrated elevated expression levels of pyroptosis-associated
   proteins, including NLRP3, cleaved caspase-1, GSDMD, GSDMD-N, and
   IL-1β, in CKD mice when compared to sham controls ([101]Figure 6B).
   Similarly, immunofluorescence (IF) staining underscored heightened
   expression of NLRP3 and cleaved caspase-1 in CKD mouse skeletal muscles
   ([102]Figure 6C). Taken together, our results provide both
   morphological and molecular evidence for the occurrence of pyroptosis
   in the skeletal muscle atrophy instigated by CKD.

Figure 6.

   [103]Figure 6
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   Evidence linking pyroptosis to muscle atrophy in CKD mice

   (A) TEM confirmed the morphological characteristics of pyroptosis in
   muscles of CKD mice. Large sections of cell membrane (CM) displaying
   discontinuities, forming extensive holes (red arrows), with partial
   rupture of muscle fibers (yellow arrows, left). Irregularly shaped
   nuclei (N) and chromatin condensation were found in the CKD group (blue
   arrows, right). (B) The protein levels of NLRP3, cleaved caspase-1,
   GSDMD, GSDMD-N, and IL-1β in quadriceps muscles of mice were measured
   by western blot (n = 7, ∗p < 0.05 and ∗∗p < 0.01). (C)
   Immunofluorescence staining underscored heightened expression of NLRP3
   (red) and cleaved caspase-1 (green) in CKD mice muscles.

Knockdown of GAS5 alleviates cellular pyroptosis and muscle atrophy in CKD
mice

   Given the observed regulatory effect of GAS5 on LPS/ATP-induced
   pyroptosis and skeletal muscle atrophy in C2C12 myotubes, we
   hypothesized that GAS5 might play a similar role in an in vivo context.
   We injected either AAV1-shGAS5 or AAV1-CTL into the quadriceps of sham
   control and CKD mice via intramuscular injection ([105]Figure S3). As
   illustrated in [106]Figure 7A, GAS5 mRNA levels in CKD mice injected
   with AAV1-shGAS5 were significantly lower than those in CKD mice
   injected with AAV1-CTL, indicating a successful GAS5 knockout in the
   CKD model. Immunoblot analysis revealed that in comparison to the
   CKD+AAV1-CTL group, the CKD+AAV1-shGAS5 group exhibited significantly
   reduced expression of MuRF-1, Atrogin-1, and pyroptosis-related
   proteins such as NLRP3, cleaved caspase-1, GSDMD, GSDMD-N, and IL-1β
   ([107]Figure 7B). These findings were further corroborated by IF
   staining of muscles from CKD mice, which consistently displayed reduced
   expression levels of NLRP3 and cleaved caspase-1 ([108]Figure 7C).
   Notably, the muscle CSA in the CKD+AAV1-shGAS5 group showed significant
   improvement compared to the CKD+AAV1-CTL group, suggesting that GAS5
   knockout can attenuate the loss of skeletal muscle mass in CKD mice
   ([109]Figure 7D). This assertion was further supported by the shift in
   the distribution of muscle fiber CSA observed after GAS5 knockout
   ([110]Figure 7E). In summary, our results demonstrate that GAS5
   knockout can mitigate both cellular pyroptosis and muscle atrophy in
   CKD mice.

Figure 7.

   [111]Figure 7
   [112]Open in a new tab

   Knockdown of GAS5 alleviates cellular pyroptosis and muscle atrophy in
   CKD mice

   (A) Real-time qPCR confirmed the expression of GAS5 mRNA in quadriceps
   muscles transfected with AAV1-shGAS5 or AAV1-CTL (n = 3, ∗∗p < 0.01).
   (B) Protein levels of MuRF-1, Atrogin-1, NLRP3, cleaved caspase-1,
   GSDMD, GSDMD-N, and IL-1β in quadriceps muscles of mice were determined
   by western blot (n = 3, ∗p < 0.05 and ∗∗p < 0.01). (C)
   Immunofluorescence staining determined the expression of NLRP3 (red)
   and cleaved caspase-1 (green) in quadriceps muscles transfected with
   AAV1-shGAS5 or AAV1-CTL. (D) H&E staining shows the CSAs of mice
   muscles transfected with AAV1-shGAS5 or AAV1-CTL (scale bars: top,
   50 μm and bottom, 25 μm; magnified images). (E) The distribution of
   myofiber sizes in CKD mice transfected with AAV1-shGAS5 was shifted
   rightward compared with CKD mice transfected with AAV1-CTL.

Discussion

   This study discovered that GAS5 regulates skeletal muscle atrophy in
   CKD through the activation of the NLRP3 signaling pathway. First, we
   observed a marked increase in the expression of GAS5 in atrophied
   skeletal muscles in CKD. GAS5 interacts with the pyroptosis inhibitory
   factor TUFM, mediated by the NLRP3 signaling pathway. This interaction
   reduces the inhibitory effect of TUFM on NLRP3, leading to an enhanced
   activity of the NLRP3 signaling pathway, resulting in increased
   pyroptosis and skeletal muscle atrophy ([113]Figure 8). Apart from
   demonstrating that GAS5 is one of the molecules involved in regulating
   muscle atrophy, we also proved that knocking out GAS5 can alleviate
   pyroptosis and muscle atrophy, while overexpressing TUFM can inhibit
   pyroptosis and improve skeletal muscle atrophy.

Figure 8.

   [114]Figure 8
   [115]Open in a new tab

   A summative diagram of GAS5 regulates pyroptosis in skeletal muscle
   atrophy with CKD

   GAS5 interacts with TUFM, and this interaction promotes NLRP3
   activation, consequently activating the pyroptosis pathway and leading
   to the occurrence of pyroptosis and muscle atrophy in CKD.

   Pyroptosis mediated by NLRP3 is an essential mechanism for the
   occurrence of muscle atrophy.[116]^25 Several studies in
   non-CKD-induced muscle atrophy models have confirmed this notion. For
   instance, Yan et al. reported that treating ischemia-induced skeletal
   muscle cell pyroptosis with exosomes containing circHIPK3 can reduce
   NLRP3 activation and caspase-1 cleavage and improve mouse muscle
   strength.[117]^26 Similarly, in dexamethasone-induced muscle atrophy,
   extracts from the leaves of Ageratum conyzoides could inhibit muscle
   degradation by blocking the nuclear factor κB (NF-κB)/NLRP3-mediated
   pyroptosis pathway.[118]^27 Further studies indicated that the
   upregulation of glycolysis in the muscle tissues of patients with
   idiopathic inflammatory myopathies activated NLRP3, causing muscle cell
   pyroptosis and leading to muscle weakness.[119]^28 It has been reported
   that IL-1β, an activation product of the pyroptosis pathway, can act
   directly on muscle cells, resulting in skeletal muscle atrophy. By
   knocking out NLRP3, the activation of IL-1β can be inhibited, thus
   preventing muscle atrophy in critically ill patients.[120]^29 As we
   know, the inflammatory signaling pathway is an important cause of organ
   damage.[121]^30 Our study further confirmed that, in skeletal muscle
   atrophy caused by CKD, pyroptosis and the NLRP3 signaling pathway are
   activated, with IL-1β secretion playing a role in the onset and
   progression of skeletal muscle atrophy.

   Increasing evidence suggests that lncRNAs regulate NLRP3-mediated
   pyroptosis through various mechanisms.[122]^31 In addition to directly
   interacting with specific proteins, lncRNAs can also form sponges by
   binding with microRNAs (miRNAs), influencing the expression of NLRP3
   mRNA. Luo et al. reported that lncRNA NLRP3 acts as a sponge
   interacting with miR-138-5p, thereby upregulating the expression of
   NLRP3.[123]^32 Moreover, lncRNAs can regulate the activation of the
   NLRP3 inflammasome by modulating the NF-κB signaling pathway.[124]^33
   Yi and colleagues found that lncRNA Gm4419 upregulates pro-inflammatory
   cytokines in mesangial cells of diabetic nephropathy through the
   NF-κB/NLRP3 pathway.[125]^34 Research has shown that lncRNA Neat1
   promotes the assembly of the NLRP3 inflammasome, thus activating the
   inflammatory response.[126]^35 Like the majority of studies, our
   research indicated that lncRNA has a positive regulatory effect on the
   NLRP3 pathway. We found that in CKD-induced muscle atrophy, pyroptosis
   is activated by lncRNA GAS5 through the NLRP3 signaling pathway. We
   revealed that GAS5 knockdown downregulated the expression of
   atrophy-related proteins such as MuRF-1 and Atrogin-1. We also
   identified that pyroptosis-associated proteins, including NLRP3,
   cleaved caspase-1, GSDMD, GSDMD-N, and the inflammatory cytokine IL-1β,
   were markedly suppressed upon GAS5 knockdown. lncRNA does not always
   exert positive regulation on NLRP3; conversely, lncRNAs can also reduce
   the expression of NLRP3. For example, lncRNA XLOC_000647 can inhibit
   the development of pancreatic cancer by suppressing the promoter
   activity of NLRP3, thus negatively regulating the expression of
   NLRP3.[127]^36

   This study confirmed that lncRNA GAS5 interacts with TUFM and activates
   the NLRP3 signaling pathway, leading to skeletal muscle atrophy,
   suggesting a new mechanism of lncRNA regulation of the NLRP3 signaling
   pathway. Importantly, using the AAV1-shGAS5 muscle injection method, we
   demonstrated that inhibiting GAS5 can significantly alleviate cell
   pyroptosis and muscle atrophy in CKD mice, highlighting the
   significance of GAS5 in the pathogenesis of muscle atrophy.

   Although our study mainly investigated the function of GAS5 in the
   cytoplasm, given the multifunctionality of lncRNA, we cannot exclude
   the possibility that GAS5 regulates gene expression in the nucleus. The
   findings of this study have not been validated in human skeletal muscle
   samples yet. Our future work will validate the regulatory mechanism of
   GAS5 on pyroptosis and skeletal muscle atrophy in human muscles, which
   will make our research results more clinically relevant and practically
   valuable. Additionally, we need to further study the specific mechanism
   of pyroptosis in regulating CKD-induced skeletal muscle atrophy,
   especially the interaction mechanism between GAS5 and TUFM. lncRNAs
   have many unconfirmed essential physiological functions, and our study
   provides a basis for identifying the role of GAS5 in the pathogenesis
   of CKD-induced muscle atrophy.

   In conclusion, we have demonstrated that GAS5 interacts with TUFM,
   promoting the activation of pyroptosis, leading to skeletal muscle
   atrophy. Inhibiting GAS5 in CKD muscles can suppress the activation of
   pyroptosis and alleviate muscle atrophy. Our research results offer new
   insights for the prevention and treatment of CKD-induced muscle
   atrophy.

Materials and methods

Animal and CKD model

   All experiments were approved by the Ethics Committee of Xinhua
   Hospital, Shanghai Jiao Tong University School of Medicine. C57BL/6J
   mice (12 weeks old) were obtained from Shanghai Slac Laboratory Animal,
   the previously described two-step 5/6 nephrectomy was used to establish
   CKD mouse models, and the sham control group underwent anesthesia
   induction without surgical removal of the kidney mass.[128]^37 Four
   weeks after 5/6 nephrectomy, CKD mice and sham controls received an
   intermuscular injection of AAV1 into the quadriceps femoris muscles and
   were pair fed for another 4 weeks. AAV1 carrying sh-GAS5 (AAV1-shGAS5)
   and shRNA-scramble (AAV1-CTL) were synthesized by Hanbio Biotechnology,
   China. The preparation and purification processes of AAV1 vectors were
   as follows: first, the pHBAAV-U6-MCS-CMV-Zs Green vector was designed,
   restriction enzyme digestion and ligation were used to construct the
   plasmid vector, then we confirmed the accuracy of sequence. After virus
   packaging, using lysis buffer solution to lysis cell. Further
   purification of the virus was performed after initial purification by
   ultracentrifugation. The concentrations of the stock solutions of
   AAV1-shGAS5 and AAV1-CTL were 1.58 × 10^10 and 2.51 × 10^10
   plaque-forming unit (PFU)/mL, respectively, and the injection dose for
   both groups was 50 μL. Sham control mice ate the same amount of food as
   the CKD mice during paired feeding, and this sequence was repeated for
   the duration of the experiment.

Cell culture and transfection

   The C2C12 mouse myoblasts (ATCC; Manassas, VA, USA) were cultured in
   Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum
   and 1% penicillin/streptomycin. For differentiating myotubes, C2C12
   myoblasts were treated with 2% horse serum for at least 72 h. In the
   follow-up experiments, C2C12 myotubes were incubated with LPS
   (100 ng/mL; Sigma) for 2 h and ATP (5.0 mM; Med Chem Express‌) for an
   additional 1 h to cause inflammation.[129]^10 Viral infection was
   performed at a multiplicity of infection (MOI) of 50 for 24 h. We
   transfected AAV1-shGAS5 and AAV1-CTL into C2C12 myotubes with
   concentrations of 2 and 1.25 μL/mL for 48 h. RT-qPCR was performed to
   measure transfection efficiency. Target sequences of these short
   hairpin RNAs (shRNA) were as follows: sh-GAS5:
   5'-AGAAGATGGTGTCAGATATAT-3' and shRNA-scramble: 5'-TTCT CCG
   AACGTGTCACGTAA-3'.

RNA-seq analysis

   The mirVana miRNA Isolation Kit (Ambion, USA) was used to extract the
   total RNA of quadriceps femoris muscles from mice, in accordance with
   the manufacturer’s protocol. RNA libraries were generated using TruSeq
   Stranded Total RNA prep kit with Ribo-Zero Gold (Illumina, USA). The
   DESeq software (R package) was employed to analyze the differentially
   expressed lncRNAs in CKD mouse models.

Histological examination

   After being dissected, the gastrocnemius muscles were rinsed with PBS
   and fixed for 24 h in 4% paraformaldehyde (PFA). Paraffin-embedded
   muscle tissues were desiccated to make 5-μm slices, H&E staining was
   performed to examine changes in the CSAs of muscles, and the CSAs was
   measured by ImageJ.

TEM

   Gastrocnemius muscles were isolated from mice, cut into pieces no
   larger than 1 mm^3 in volume, and then incubated in electron microscope
   fixative for 2–4 h at 4°C and rinsed in 0.1 M phosphate buffer (PB; pH
   7.4) 3 times. The tissues were fixed in 1% osmium acid-0.1 M PB (pH
   7.4) at room temperature (20°C) for 2 h. For dehydration, tissues were
   sequentially placed in 50% alcohol, 70% alcohol, 80% alcohol, 90%
   alcohol, 95% alcohol, 100% alcohol, 100% alcohol, 100% acetone, 100%
   acetone for 15 min each time. The sections were dried with uranium-lead
   double staining at room temperature overnight after embedding, and a
   transmission electron microscope was used to analysis the images.

IF

   The gastrocnemius muscles were isolated, washed with PBS, and fixed in
   4% PFA for 24 h. After paraffin-embedded muscle tissues were
   desiccated, the sections were subjected to an antigen-repair cassette
   containing EDTA antigen repair buffer (pH 8.0) and placed in a
   microwave oven. The slides were shaken dry, and an autofluorescence
   quencher was added, and then they were incubated with anti-mouse NLRP3
   (Proteintech, USA, 1:200) and anti-mouse cleaved caspase-1 (Cell
   Signaling Technology, USA, 1:200) at 4°C overnight after being blocked
   with 5% goat serum for 30 min. Subsequently, the sections were
   incubated with secondary antibodies for 1 h at room temperature in the
   dark. 4-6-diamidino-2-phenylindole was used to stain the nuclei, the
   changes were examined with a fluorescent microscope (NIKON ECLIPSE C1),
   and images were captured using the NIKON DS-U3 imaging system.

Western blotting and antibodies

   The proteins were extracted from C2C12 myotubes and quadriceps muscles
   of mice. The concentrations of immunoreactive proteins were measured
   using ImageJ and normalized to the concentrations of β-actin or GAPDH.
   The primary antibodies used were as follows: anti-MuRF-1 (Santa Cruz,
   USA, sc-398608), anti-Atrogin-1 (ECM Biosciences, USA, AP2041),
   anti-NLRP3 (Cell Signaling Technology, USA, 15101S), anti-GSDMD (Abcam,
   USA, ab209845), anti-GSDMD-N (Abcam, USA, ab209845), anti-cleaved
   caspase-1 (Cell Signaling Technology, USA, 89332S), anti-IL-1β (Abcam,
   USA, ab2105), anti-GAPDH (Proteintech, USA, ag16692), and anti-β-actin
   (Servicebio, China, GYW00011).

FISH

   The FISH assay was conducted in C2C12 myotubes and gastrocnemius
   muscles of mice. The sections were digested with proteinase K
   (20 μg/mL) for 5 min after being fixed in 4% PFA for 20 min, rinsed
   with PBS, and incubated at 37°C for 1 h; we dropwise added the
   pre-hybridization solution and hybridization solution (containing a
   probe GAS5 concentration of 1 μM) and hybridized the sections at 42°C
   overnight. After washing off the hybridization solution, we performed
   DAPI staining for 8 min. Subsequently, an anti-fluorescence quenching
   sealer was added to seal the sections. A Nikon orthomosaic fluorescence
   microscope was used to observe the images. The probe information of
   GAS5: 5′-CCATCACAGAGGTCCACACTGCCAT TCCTGCT-3' FAM labeling.

RNA pull-down assay and MS

   PCR amplification was performed to screen for suitable PCR primers for
   the sense and antisense strands, and the PCR products were used as
   templates and then transcribed in vitro to obtain pure lncRNA GAS5
   using the MAXIscript kit (Thermo Fisher Scientific). The transcribed
   lncRNA GAS5 was mixed with biotinylated cytidine bisphosphate and
   incubated at −20°C overnight to obtain biotin-labeled lncRNA GAS5. The
   labeled lncRNA GAS5 was incubated with streptavidin magnetic beads for
   30 min at room temperature and incubated with pre-prepared C2C12
   cytoplasmic lysates and nucleus lysates at 4°C for 1 h. Following
   incubation, the beads were collected and washed once by adding 200 μL
   1× washing buffer. 200 μL 1× washing buffer was added and mixed with
   the beads again. We put the mixture into a magnetic stand, discarded
   the supernatant, and then added 100 μL elution buffer and vortexed to
   mix it well, followed by incubation for 15 min at room temperature.
   Subsequently, we added 20 μL of elution buffer and boiled it for 5 min,
   placed the centrifuge tube on a magnetic stand for 2 min, and removed
   the supernatant. Silver staining was used to detect proteins that were
   separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis
   (SDS-PAGE). The primer sequences were as follows: lncRNA GAS5 sense F2,
   5'-TAATACGACTCACTATAGGGGTCTTTTCGAGGTAGGAGTCGACTCCTGTG-3'; lncRNA GAS5
   sense R2, 5'-TTTTTTTTTTTTTTTTTTTTTTTTGATGGGATTGCA-3'; lncRNA GAS5
   antisense F2, 5'-TAATACGACTCACTATAGGGTTTTTTTTTTTTTTTTTTGATGGATTGCA-3';
   and lncRNA GAS5 antisense R2, 5'-GTCTTTTCGAGGTAGGAGTCGACTCCTGTG-3'. The
   RNA-binding protein complexes were washed, eluted, and analyzed by MS.
   The MS was performed by LC-BIO Biotech (Hangzhou, China).

RIP

   RIP experiments were conducted using RIP kit (BersinBio, China,
   Bes5101), complying with the manufacturer’s instructions. RNA was
   co-immunoprecipitated using anti-TUFM antibody (1:20, ABclonal, USA,
   A6423). Anti-TUFM (Proteintech, USA, 67802-1-Ig) was used for western
   blotting, and RT-PCR was performed to measure the expression of GAS5.

Statistical analysis

   SPSS 25.0 was used for the statistical analysis, and all data are
   presented as mean ± SD. To evaluate the differences in the two groups,
   we used two-tail unpaired Student’s t tests, while two-way ANOVA was
   performed for more than two groups. ∗p < 0.05 and ∗∗p < 0.01 were
   considered statistically significant.

Data and code availability

   The data generated in this study are available upon request from the
   corresponding author.

Acknowledgments