In conditions of nutrient excess, adipocytes produce leptin to modulate CNS activity, promote satiety, and increase energy expenditure. Obesity is associated with hyperleptinemia and leptin resistance (53, 54). Indeed, leptin levels are 5-10x higher in obese patients compared to non-obese individuals (55, 56). Since the kidney is the primary organ responsible for leptin clearance (57), CKD is also associated with hyperleptinemia, the degree of which correlates with the CKD stage (18, 58, 59). Park et al. recently demonstrated a significant correlation between elevated serum leptin levels and the risk of CKD in men after adjusting for eGFR and age (60). Such associations are even more evident in females, probably owing to sex-specific differences in circulating leptin levels (61).

Numerous studies have shown leptin to induce glomerulosclerosis and hypertension, both risk factors to CKD (62). The short form of the leptin receptor (Lep-Ra) is the predominant leptin receptor expressed in the kidney compared to the long form (Lep-Rb). Glomerulosclerosis and renal fibrosis in obese mice have been linked to Lep-Rb-dependent JAK2-STAT signaling in renal mesangial cells (63). Leptin promotes TGFβ-1 release and type IV collagen and fibronectin production in the glomerulus, leading to proteinuria and glomerulosclerosis (62). This effect was initially found to be mediated via adenosine monophosphate-activated protein kinase (AMPK) activation (64), which paradoxically inhibits TGFβ-1 and protects against renal fibrosis in several mouse models (65). This discrepancy suggested that leptin-mediated fibrosis may additionally involve other signaling pathways in the kidney. Indeed, activation of the p38/MAPK signaling pathway is involved in leptin-mediated renal fibrosis (64). Leptin also induces endothelial dysfunction (ED) by upregulating vascular adhesion molecules such as intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) through AKT/GSK3β and Wnt/β-catenin signaling pathways, promoting renal inflammation and vascular remodeling (66, 67). Lastly, leptin promotes oxidative stress in renal tubular epithelial cells and stimulates monocytes to release IL-6 and TNF-α, promoting renal inflammation (68, 69).

Leptin is also considered a uremic toxin, contributing to many CKD complications including cachexia, protein-energy wasting (PEW), insulin resistance, hypertension, cardiovascular disease, and bone pathologies (70). PEW in CKD features anorexia, increased energy expenditure, decreased protein stores and muscle mass, and weight loss. Leptin suppresses food intake and increases energy expenditure through binding mineralocortin-4 receptors (MC4-R) in the hypothalamus, leading to uremic cachexia (71–73). Inhibiting MC4-R improves cachexia and reduces skeletal muscle wasting in preclinical models, but this needs further investigation in humans (73).

Leptin increases the risk of cardiovascular disease, the most common cause of mortality in CKD patients. Leptin induces hypertension by increasing sympathetic outflow, decreasing nitric oxide (NO) production, and increasing endothelin-1 in endothelial cells (74–76). Leptin promotes atherogenesis through endothelial cells, macrophages, and smooth muscle cells via the Lep-Rb receptor, which is reviewed elsewhere (77). Similarly, leptin binding to the Lep-Rb receptor on platelets enhances ADP signaling to induce platelet aggregation, which may cause the platelet dysfunction characteristic of uremia (78, 79). Lastly, hyperleptinemia decreases glucose-stimulated insulin release from pancreatic β-cells and impairs insulin signaling in hepatocytes (80, 81), although it should be noted that normal leptin levels enhance insulin release (80, 81).

Therefore, obesity-associated hyperleptinemia may contribute to renal pathology and CKD, mainly by causing secondary glomerulosclerosis. Furthermore, CKD-associated hyperleptinemia may contribute to numerous CKD complications. In agreement with these findings, leptin-deficient mice are significantly protected against albuminuria, glomerular crescent formation, macrophage infiltration, and glomerular thrombosis (82). Inhibiting leptin using specific antibodies or antagonists also substantially reduces blood pressure in mice with diet-induced obesity (83) and alleviates cachexia in CKD mice (84). Weight loss, either through lifestyle interventions, pharmacotherapies, or bariatric surgeries, is associated with significant decreases in leptin levels (85–89), but whether leptin normalization after weight loss directly improves renal function remains to be investigated.

Adiponectin secretion is decreased in obesity, promoting the development of obesity-related chronic complications. The development of adiponectin-KO animal models have allowed causal relationships to be drawn between adiponectin deficiency and several aspects of the metabolic syndrome. For example, adiponectin-KO mice develop hepatic steatosis, which is attenuated by transfecting the adiponectin gene (90, 91). In skeletal muscle, adiponectin stimulates beta-oxidation and reduces lipid deposition and consequent lipotoxicity (92). Furthermore, adiponectin inhibits lipolysis and stimulates triglyceride storage in subcutaneous adipose tissue. Adiponectin, therefore, promotes fat storage in AT and increases insulin sensitivity, with its decrease in obesity a causal factor in insulin resistance, lipotoxicity, and metabolic syndrome manifestations (92).

The renoprotective effects of adiponectin are well-documented (18). Two adiponectin receptor isoforms, ADIPOR1 and ADIPOR2, are expressed in the kidney. Stimulation of ADIPOR1 and ADIPOR2 activate AMPK and peroxisome-proliferator activated receptor-α (PPAR-α), respectively, which attenuate renal inflammation, fibrosis, glomerulosclerosis, podocyte effacement, and albuminuria (17). A rise in intracellular AMPK by ADIPOR1 activation in podocytes inhibits NADPH oxidase and reduces permeability to albumin (93). In this light, non-obese non-diabetic mice who are adiponectin-deficient still develop effacement of podocyte foot processes and albuminuria due to increased oxidative stress (93, 94). In mesangial cells, adiponectin increases AMPK to attenuate angiotensin-II-induced TGF-β1 production, decreasing renal fibrosis (95).

Adiponectin also exerts anti-inflammatory effects on the kidney. For example, MCP-1 binds to its cognate CCR2 receptor to promote macrophage infiltration into the kidneys and renal inflammation (96). Adiponectin-deficient CKD mice develop significant albuminuria, tubulointerstitial fibrosis, and inflammation characterized by high MCP-1, TNF-α, NADPH oxidase, and VCAM-1 upregulation (97). Adiponectin administration via an adenovirus vector significantly reduces albuminuria, tubulointerstitial fibrosis, glomerular hypertrophy, and inflammation by lowering TNF-α, NADPH oxidase, and VCAM1 (97). Adiponectin has also been shown to directly stimulate IL-10 production by macrophages and decrease IL-6 and TNF-α, suggesting polarization to an M2 phenotype (98).

Lastly, ceramides are a group of sphingolipids implicated in renal disease. Serum levels of several ceramides are independent risk factors for CKD development and stage progression (99), as well as insulin resistance and lipotoxicity (100). Ceramides also act at several levels of the insulin signal transduction pathway to impair insulin signaling. Notably, both ADIPOR1 and ADIPOR2 possess intrinsic basal ceramidase activity, which is enhanced by adiponectin binding (101). Elevated ceramidase activity by ADIPOR1 and ADIPOR2 overexpression increases insulin sensitivity and glucose utilization while opposing hepatic steatosis (102). Ceramidase metabolizes ceramides into sphingosine-1-phosphate, which has anti-apoptotic effects and may even induce proliferation (103). These studies indicate that the pleiotropic metabolic, anti-apoptotic, and insulin-sensitizing effects of adiponectin may at least partly involve amplifying receptor-associated ceramidase activity.

Serum adiponectin is lower in obese patients compared to lean individuals. ADIPOR1 and ADIPOR2 expression is also reduced in the kidneys of obese and diabetic mice (104). Therefore, kidneys from obese mice and humans showed reduced AMPK levels (105). Treatment with 5-aminoimidazole-4-carboxamide-1-β-D-furanoside (AICAR), which enhances adiponectin-mediated AMPK signaling, increases AMPK levels in obese kidneys and reduces mesangial expansion and albuminuria (106). The antioxidant resveratrol also restores ADIPOR expression in the kidney and increases AMPK activation in diabetic mice, associated with reductions in albuminuria, oxidative stress, and inflammation (104). The molecule berberine enhances adiponectin signaling through AMPK to ameliorate renal pathology in diabetic mice (107). In animal models of diabetic nephropathy and obesity, the AMPK agonist PF-06409577 and adiponectin receptor agonist AdipoRon reduce proteinuria, inflammation, and renal fibrosis (108–110). These results suggest that targeting adiponectin receptors or AMPK directly may be beneficial in obesity- and diabetes-related kidney disease.

Despite adiponectin having numerous renoprotective effects, adiponectin levels are paradoxically increased in CKD and are positively correlated with albuminuria, CKD stage, and mortality, independent of body mass index (BMI) (58, 111). Adiponectin also predicts adverse cardiovascular outcomes in CKD patients (112). Unlike leptin, higher adiponectin levels in CKD cannot be simply explained by decreased renal clearance because the liver clears the high-molecular-weight form of adiponectin (113). Therefore, why adiponectin is elevated in CKD and is predictive of disease severity remains investigational.

Tian et al. induced CKD in non-obese mouse models with deoxycorticosterone acetate-salt (DOCA) and angiotensin II infusion (114). Transgenic adiponectin-overexpressing CKD mice showed significantly lower albuminuria, glomerular and interstitial fibrosis, and attenuated effacement of podocyte foot processes. Markers of tubular injury and inflammation were also lower in the transgenic models (114). These results are contrary to the unfavorable prognostic effect attributed to adiponectin in CKD patients. Yang et al. demonstrated that elevated adiponectin levels were associated with the presence of bone marrow-derived fibroblasts in kidneys with unilateral ureteral obstruction and ischemia/reperfusion injury (115). Adiponectin-deficient mice showed reduced renal fibroblast and M2 pro-fibrotic macrophage infiltration. The same study also showed adiponectin to activate AMPK on bone-marrow-derived monocytes, thereby increasing α-smooth muscle antigen (α-SMA) and production of extracellular matrix proteins. Therefore, the Yang et al. study suggested inhibiting the adiponectin/AMPK axis may ameliorate fibrotic renal disease (115). Similarly, Perri et al. reported that administration of lipopolysaccharide (LPS) induces adiponectin production by renal tubular epithelial cells to cause renal fibrosis (116). Numerous other studies have demonstrated the production of adiponectin by the kidney itself (117, 118). However, how kidney-derived adiponectin contributes to circulating adiponectin levels and any potential functional differences are not yet known.

PPAR-α is also known to exert renoprotective effects. Boor et al. demonstrated PPAR-α expression in the renal tubular epithelium but not the interstitium. PPAR-α levels decreased after fibrosis induction through unilateral ureteral obstruction and 5/6 nephrectomy (119). Treatment with the PPAR-α agonist BAY PP1 significantly increased PPAR-α expression, correlated with a reduction in tubulointerstitial fibrosis, inhibition of interstitial fibroblasts, lower TGF-β1 levels, and slowed down the progression of renal dysfunction. Therefore, PPAR-α in tubular epithelial cells attenuates fibrosis upon renal injury (119).

Senescence plays a crucial role in propagating age-related diseases (123). Senescent cells accumulate in most tissues and organs with aging, including in adipose tissue. Importantly, obesity increases the senescent cell burden in adipose tissue. The p53-dependent DNA damage response is the main inducer of senescence in adipose tissue (124). A study showed that DNA polymerase-η KO mice (to increase DNA damage) accumulate senescent cells in adipose tissue (125). SREBP1—a transcription factor involved in regulating the expression of genes encoding proteins involved in lipid metabolism—was recently found to also facilitate DNA repair in adipocytes (126). Deletion of SREBP1 increased DNA damage and accelerated senescence in adipocytes, followed by adipose tissue inflammation and consequent insulin resistance (126). Mice with senescent cell accumulation in adipose tissue are more prone to obesity and adipose tissue inflammation, even with a standard chow diet (125).

Oxidative stress-induced senescence in adipose tissue is linked to higher leptin, IL-6, and TNF-α production in the SASP, suggesting that adipocyte senescence may be causal in obesity-related chronic inflammation (124, 127). Depleting senescent cells in adipose tissue improves glucose homeostasis and insulin resistance (discussed below). Activin A is another SASP component which disrupts insulin signaling by decreasing the expression of insulin-dependent transcription factors including PPARγ and CCAAT-enhancer-binding protein α (C/EBPα) (128). These observations suggest that senescence in adipose tissue results in the production of cytokines and chemokines, leading to adipose tissue inflammation and insulin resistance. Adipose tissue is composed of many different cell types, each exhibiting varying susceptibility to senescence. The discussion herein focuses on the major cell types comprising adipose tissue and the causes and consequences of senescence induction in these cell types ( Figure 2 ).

Numerous studies report a higher senescent cell burden in diseased and aged kidneys. The regenerative potential of the kidney after injuries diminishes with aging and CKD. Cellular senescence is believed to impair regenerative mechanisms in aged and diseased kidneys, leading to maladaptive repair and fibrosis.

Proximal tubular epithelial cells are particularly affected by senescence. Biopsies of transplanted kidneys and those with glomerular diseases stain positive for senescence markers p16^INK4a and p21^CIP1 in the proximal tubular epithelial cells (147, 148). Telomere attrition, elevated SA-β gal, p16^INK4a, and p21^CIP1 expression have also been directly correlated with IgA nephropathy progression (149). Baker et al. showed that senescent cells accumulate in aging kidneys in INK-ATTAC transgenic mice. An elevated renal senescent cell burden was associated with glomerulosclerosis, which was attenuated by depleting senescent cells in INK-ATTAC mice (150). Braun et al. demonstrated that transplanted kidneys in wild-type mice exhibit elevated senescence markers and incrementally undergo atrophy and fibrosis (151). By contrast, transplanted kidneys in p16^INK4A-KO mice show less atrophy and fibrosis after ischemia-reperfusion injury (151). Transplanting kidneys from senescence-depleted mice consistently show better longevity and proliferation of tubular epithelial cells. An elevated senescent cell burden may therefore contribute to long-term allograft kidney deterioration and CKD development in humans (151).

In vivo mouse models of aged and irradiated kidneys demonstrate elevated senescence markers, reduced proliferative repair after injury, and produce TGF-β as a part of their SASP to induce fibrosis. Using the senolytic ABT-263 to eliminate senescent proximal tubular epithelial cells improves these parameters (152). A recent study demonstrated renal tubular epithelial cell senescence – evidenced by higher p16^INK4a, p19, and p21^CIP1 expression – secondary to chronic ischemia from renal artery stenosis in mice and humans. A dasatinib and quercetin senolytic combination alleviated renal dysfunction in these mice, suggesting a causal relationship between chronic ischemia, cellular senescence, and kidney damage (153). Diabetic nephropathy is the major cause of CKD worldwide and a significant contributor to obesity-related kidney disease. Biopsies of kidneys from patients with type 2 diabetic nephropathy show elevated senescence markers SA-β gal and p16^INK4a in proximal tubular epithelial cells, mesangial cells, podocytes, and endothelial cells (154). WT diabetic kidney disease mouse models develop proteinuria and glomerular hypertrophy, and both effects are attenuated in p21^CIP1-KO mice (155). Dasatinib and quercetin combination also reduce AKI to CKD transition in murine models of cisplatin and radiation-induced kidney injury (156). Senescent markers SA-β gal, p16^INK4A, and p21^CIP1 are also elevated in the kidneys of rats and cats with CKD (157, 158). These data show cellular senescence underpins various renal pathologies that lead to CKD, and senolytics could mitigate this progression.

CKD is considered a systemic premature aging phenotype, known as uremia-associated aging (159). Uremic patients can develop age-related conditions, including osteoporosis, sarcopenia, frailty, impaired wound healing, infections, insulin resistance, cognitive dysfunction, hypogonadism, and vascular aging (160–163). Hence, certain uremic toxins expectedly accelerate biological aging hallmarks, including cellular senescence, to precipitate a premature aging phenotype (164). Uremia-induced senescenced was first demonstrated in the aortas of hypertensive rats, where indoxyl sulfate-related cellular senescence was correlated with aortic wall calcification and thickness, a sign of vascular aging (165). Uremia-induced senescence is mediated by oxidative stress and consequent DNA damage response signaling and upregulation of p21^CIP1 and p53 (166). A recent review by Huang et al. summarized the mechanisms behind CKD-induced senescence (167). The uremic toxins indoxyl sulfate and p-cresyl sulfate induce senescence in mesenchymal stem cells, evidenced by elevated p21^CIP1 expression (168, 169). A normocytic normochromic anemia is common in CKD pateints, and is mainly thought to be due to low erythropoietin production by the kidney. Mas-Oodi et al. recently demonstrated that indoxyl sulfate induced senescence in CD34+ hematopoietic stem cells, thereby arresting their proliferation and reducing erythropoiesis (170). Indoxyl sulfate also induces senescence in renal proximal tubular epithelial cells in CKD through ROS-dependent p53 expression (171). Senescent proximal tubular cells display a proinflammatory and profibrotic protein signature, with elevations in NFκB and TGF-β production, possibly contributing to further declines in renal function (171, 172).

P-cresyl sulfate activates NADPH oxidase and induces oxidative stress in mouse 3T3-L1 adipocytes. Exposure to p-cresyl increases TNF-α and IL-6 production by 3T3-L1 adipocytes and increases ATM infiltration, suggesting that this uremic toxin is a mediator of CKD-induced adipose tissue inflammation (172, 173). In agreement with these findings, Koppe et al. reported that administering p-cresyl sulfate to normal mice for 4 weeks triggered lipotoxicity and insulin resistance (174). Although senescence markers were not evaluated in these studies, it is conceivable that the intracellular accumulation of p-cresyl sulfate may induce adipocyte senescence, since oxidative stress is the major pathway of adipose tissue senescence. In the context of the adipose-renal axis, these studies suggest that the accumulation of uremic toxins in CKD trigger senescence in adipose tissue, amplifying the inflammaging seen in CKD pateints. Further studies are needed to determine which uremic toxins induce adipose tissue senescence, the mechanisms involved, which subpopulations of cells in adipose tissue are affected, the important SASP components by which uremia-induced senescent adipose tissue exerts pathologic systemic effects, and whether senolytic or other senescent-targeting strategies are effective in ameliorating uremia-induced senescence. Another pathway to consider in the adipose-renal axis in CKD is the gut microbiome. CKD alters the symbiotic relationship between the intestinal microbiome and the body (i.e., gut microbiome dysbiosis), leading to the fermentation of macronutrients and production of various uremic toxins, including indoxyl sulfate, p-cresyl sulfate and others. CKD-related impairment of the intestinal epithelial barrier allows for the spillover of these toxins into the bloodstream, which drive systemic oxidative stress and inflammation (175, 176). Whether and how certain dietary modifications lifestyle interventions such as exercise, restore host-enterobiome symbiosis and alleviate senescence in the context of the adipose-renal axis is an important topic for future studies to address.

Targeting senescent cells pharmacologically can alleviate numerous age-related diseases. Baker et al. initially demonstrated that depleting senescent cells prevents the development of age-related changes in adipose tissue, skeletal muscle, and eyes of INK-ATTAC transgenic mice (150). Numerous strategies have emerged to deplete senescent cells or mitigate their harmful effects. Briefly, senolytic drugs inhibit SCAPs characteristic of senescent cells, allowing for the selective depletion of senescent cells. Dasatinib, quercetin, and fisetin are the most studied senolytics in preclinical animal models thus far, and numerous clinical trials testing their efficacy in age-related disorders are underway (177). Senomorphic drugs inhibit various SASP components without inducing senescent cell death. Most senomorphics target transcriptional regulators of the SASP, including ATM, p38 MAPK, JAK/STAT, NFκB, and mTOR pathways. Other strategies are also emerging, recently reviewed by Zhang et al. (178, 179). Many preclinical studies have shown senolytics to alleviate aging phenotypes, including cancer, chemotherapy- and radiation-induced premature aging, diabetes, osteoarthritis, neurodegeneration, glaucoma, age-related macular degeneration, idiopathic pulmonary fibrosis, heart failure, and CKD (180).

The impact of senescent cell depletion has also been investigated in obesity and kidney disease. Palmar et al. cleared senescent cells either by senolytic combination dasatinib+quercetin or by selective depletion of p16^INK4a-expressing cells and observed reduced adipose tissue inflammation and improved glucose tolerance and insulin sensitivity (181). Since DNA damage is the major inducer of senescence in adipose tissue, interventions such as exercise, N-acetylcysteine, and senolytics that reduce oxidative stress in adipose tissue decrease adipose tissue SC burden and attenuate ATM infiltration and adipose tissue inflammation (125). Another study showed that senescent cell clearance in adipose tissue of HFD obese mice by dasatinib+quercetin and navitoclax improved insulin sensitivity and increased plasma adiponectin levels (182). Consistent with higher adiponectin levels, senescent cell clearance anormalizes microalbuminuria and podocyte barrier integrity (181). An HFD increases senescent cell burden in mouse kidneys—detected by p16^INK4a, p19, and p53 expression and SASP upregulation—linked to renal fibrosis and functional impairment (144). Quercetin administration reduced senescent cell burden in the kidney, attenuated renal fibrosis, increased renal cortical oxygenation, and lowered plasma creatinine levels (144).

These findings suggest that depleting adipose tissue-resident senescent cells by senolytics restores adipogenesis, reduces adipocyte hypertrophy, improves glucose tolerance and insulin sensitivity, reduces macrophage infiltration into adipose tissue, and increases adiponectin secretion.