PACS-2 Ameliorates Tubular Injury by Facilitating Endoplasmic Reticulum–Mitochondria Contact and Mitophagy in Diabetic Nephropathy

Diabetic nephropathy (DN) is one of the common microvascular complications of diabetes and is the leading cause of end-stage renal disease (1). The pathogenesis of DN involves multiple mechanisms, among which the key role of tubular injury has been increasingly appreciated (2–4). Findings that show tubulointerstitial fibrosis correlates best with DN progression and that subtle tubular dysfunction may be detected in early DN, even preceding glomerular abnormalities, have challenged the classical “glomerulocentric” paradigm (5,6). Furthermore, the tubular view of hyperfiltration inspired by the renoprotective effects of SGLT2 inhibitors in patients with diabetes also highlights proximal tubular cells (PTCs) as a potential target of DN (7). However, the precise mechanisms underlying tubular injury under diabetic conditions remain obscure.

PTCs are rich in mitochondria and depend on oxidative phosphorylation within mitochondria to produce adequate ATP for maintaining renal function of reabsorption (8). On the other hand, PTCs are susceptible to mitochondrial dysfunction. Mitochondrial fitness requires an integrated quality control network formed by mitochondrial dynamics (fission and fusion) and mitophagy. Perturbation of either signaling pathway may induce mitochondrial damage and dysfunction (9). Tubular cells (TCs) in diabetic kidneys have excessive mitochondrial fission and defective mitophagy, leading to accumulation of dysfunctional mitochondria and increased production of mitochondrial reactive oxygen species (ROS), ultimately activating mitochondria-dependent apoptosis (2,10,11), but the precise mechanisms are largely unexplored. Mitochondria can form physical contacts with the endoplasmic reticulum (ER), namely mitochondria-associated ER membrane (MAM) (12). MAM is an important platform for maintaining cellular homeostasis through regulating mitochondrial dynamics, Ca²⁺ signaling, autophagy or mitophagy, and apoptosis (13–17). We have previously shown that the integrity of MAM is closely related to tubular injury in DN (18). However, the regulation of MAM integrity in states of hyperglycemia in tubules and the mechanism that links MAM to tubular pathobiology in DN remain to be elucidated.

Phosphofurin acidic cluster sorting protein 2 (PACS-2) is a key regulator of MAM and is associated with metabolic disease, such as obesity and insulin resistance (19). Deletion of the PACS-2 gene leads to increased mitochondrial fragmentation and mitochondria uncoupling from ER (20), and silencing PACS-2 also impairs mitophagy and increases cell apoptosis (21). Furthermore, we previously found that PACS-2 mainly is expressed in PTCs and is markedly decreased in the kidneys of patients with DN (22). However, there are few reports on the roles and related mechanisms of PACS-2 in various kidney diseases, including DN. Given that PACS-2 may play an important role in maintaining the integrity and function of MAM in TCs, we explored the role of PACS-2 in tubular injury in DN.

In this study, we demonstrate that the expression of PACS-2 was decreased in the kidneys of patients with DN and of mice with streptozotocin (STZ)-induced DN, which was associated with renal function deterioration and tubular atrophy as well as interstitial fibrosis. Furthermore, compared with control diabetic mice, diabetic mice with proximal tubule (PT)-specific Pacs-2 gene knockout (diabetic Pacs-2^ptKO mice) displayed aggravated albuminuria and tubular damage, which were accompanied by exacerbated mitochondrial fragmentation, disruption of MAM integrity, and impairment of mitophagy. In vitro studies showed that these changes induced by high glucose (HG) levels were rescued by overexpression of PACS-2 in HK-2 cells. Moreover, a PACS-2–dependent increase of BECN1 translocation to MAM through direct interaction was verified. To our knowledge, this is the first report of the role and mechanisms of PACS-2–mediated protection against tubular injury in DN.

Clinical samples were collected from the Second Xiangya Hospital of Central South University. A total of 25 patients with DN diagnosed by renal biopsy at the Department of Nephrology were enrolled in this study. Patients with minor glomerular lesions (n = 9) were selected as control participants, as described previously (23). Routine clinical parameters such as estimated glomerular filtration rate (eGFR) and pathological data, including interstitial fibrosis and tubular atrophy (IFTA), of these patients, were collected. The protocol was approved by the Ethics Committee of the Second Xiangya Hospital, Central South University. All the patients signed forms in advance indicating informed consent.

Heterozygote mice of Pacs-2 (Pacs-2^fl/+) mice (C57BL/6J background) were generated using CRISPR/Cas-9 gene editing technology by Shanghai Biomodel Organism Science & Technology Development Co., Ltd. (U.S. National Center for Biotechnology Information Gene ID: 217893). Ggt1-Cre mice were purchased from The Jackson Laboratory (24). Ggt1-Cre mice were on the mixed Balb/cJ/C57BL/6 background. The Pacs-2^ptKO mice and littermate control (Pacs-2^ctrl) mice were obtained by crossing Pacs-2^fl/fl mice with Ggt1-Cre mice. These mice were genotyped by PCR analysis of genomic DNA isolated from tail tissues using a Mouse Direct PCR kit (Bimake), as described previously (23). To confirm Cre-mediated recombination, genomic DNA was isolated from kidney cortex using a Universal Genomic DNA Kit (Beijing Cowin Biotech Co., Ltd.) according to the manufacturer’s instructions. The primer sequences used are listed in Supplementary Table 1.

Male mice aged 6 to 8 weeks were injected intraperitoneally with STZ (50 mg/kg body weight in citrate buffer, pH 4.5; Sigma-Aldrich) for 5 days to induce hyperglycemia, as described previously (11). A week after the injections, mice whose blood glucose levels were >16.7 mmol/L at random measurements were included in the experiment. The control mice were injected intraperitoneally with the same amount of citrate buffer. The mice were divided into the following four groups: Pacs-2^ctrl (n = 7), Pacs-2^ptKO (n = 5), Pacs-2^ctrl + STZ (n = 7), Pacs-2^ptKO + STZ (n = 7). These mice were followed up for an additional 12 weeks, and body weight and blood glucose levels were monitored every 2 weeks. Urine was collected at 20 weeks of age and kidney tissues were collected for various experiments. All the experiments were approved by the Medical Ethics Committee of Central South University and followed National Institutes of Health guidelines for the care and use of laboratory animals.

Urine albumin concentration was determined using a Mouse MAU (microalbuminuria) ELISA Kit (Sangon Biotech) according to manufacturer’s instructions. Urine creatinine levels were measured in the same samples using CicaLiquid-N CRE (Kanto Chemical Co., Inc.) following the manufacturer’s instructions.

Primary PTCs were isolated from mouse kidneys and cultured in medium containing DMEM/F12 culture medium with 5% FBS and 1% penicillin/streptomycin. Briefly, the kidney cortices were minced into pieces of ∼1 mm³, and these were rinsed with precooled Hanks’ balanced salt solution. The tissue pellets were digested with 2 mg/mL collagenase II (Sigma-Aldrich) prepared in DMEM/F12. Cells were then passed through 100-μm, 70-μm, and 40-μm sieves in turn. Filtrate under the 40-μm sieve was collected and centrifuged. Subsequently, the cell precipitates were resuspended in the aforementioned culture medium and grown in a cell incubator containing 5% CO₂ at 37°C. The isolated cells were identified by staining with an anti-megalin antibody, as previously described (25).

Total RNA of kidney cortices was extracted using RNAiso Plus (TaKaRa). Then, RNA was reverse transcribed to cDNA using PrimeScript Reagent Kit (TaKaRa). Subsequently, quantitative PCR was performed using a TB Green Premix Ex Taq II reagent (TaKaRa) with a 7300 Real-Time PCR System (Applied Biosystems), as previously described (11). All PCR primers are listed in Supplementary Table 1.

Total proteins of renal cortices or cultured HK-2 cells were extracted using radioimmunoprecipitation assay buffer containing protease inhibitors and phosphatase inhibitors (CWBIO), as previously described (11). A BCA Protein Assay Kit (Thermo Fisher Scientific) was used for quantifying the protein concentrations. Total proteins were subjected to SDS-PAGE and then transferred onto polyvinylidene fluoride membranes. After blocking, the membranes were incubated with primary antibodies at 4°C overnight. After washing, the membranes were incubated with corresponding secondary antibodies (Abcam). The membrane blots were detected using an enhanced chemiluminescence kit (Thermo Fisher Scientific). And the intensity of the bands was quantified by ImageJ software (National Institutes of Health) (11). The antibodies used are listed in Supplementary Table 2.

The mouse kidney tissues were fixed in 4% paraformaldehyde and embedded in paraffin. Sections 4-μm thick were subjected to hematoxylin-eosin (HE), periodic acid Schiff (PAS), and Masson staining, as previously described (26). A semiquantitative scoring system was used to evaluate the degree of tubular interstitial damage, as described by Tervaert et al. (27).

The MAM integrity and the morphological change of mitochondria in renal TCs of mice were observed by transmission electron microscopy (TEM) (11,28). Mitochondrial length was measured using ImageJ software (National Institutes of Health). The morphology of at least 180 mitochondria was determined for each experimental group. Mitochondria were then divided into three different categories, based on length, as fragmented (mitochondrial length <1 μm), tubular (1–2 μm), and elongated (>2 μm), as described previously (29). The length of mitochondrial outer membrane associated with ER (distance, <50 nm) and mitochondrial perimeter were traced and measured using ImageJ software. Data were reported as the proportion of mitochondrial surface adjacent to ER, as previously described (18).

The MAMs in paraffin-embedded kidney sections were detected and quantized using in situ proximity ligation assay (PLA; Sigma-Aldrich) by measuring VDAC1 and IP3R1 proximity, as previously described (18). Briefly, after dewaxing, hydrating, antigen repairing, and blocking, the kidney sections were incubated with anti-VDAC1 antibody and anti-IP3R1 antibody at 4°C overnight. Then, the sections were incubated with the PLA probes, followed by ligation and amplification, and then the sections were incubated with polymerase. After washing, the sections were incubated with the substrate solution, followed by hematoxylin to stain nuclei. The sections were examined by light microscopy.

Briefly, 4-μm–thick paraffin-embedded kidney sections were dewaxed, hydrated, and antigen repaired. Then, the sections were blocked with 5% BSA and subsequently incubated with primary antibodies (listed in Supplementary Table 2) at 4°C overnight. After washing, the sections were incubated with secondary antibody, reacted with diaminobenzidine (Servicebio), and stained with hematoxylin. The sections were examined by light microscopy. Quantification of the expression level of Pacs-2 and fibronectin was determined using Image-Pro Plus 6.0 (Media Cybernetics).

Briefly, the kidney tissue sections were dewaxed, hydrated, and antigen repaired, and then were permeabilized and subsequently blocked with 5% BSA. The sections were incubated with primary antibodies at 4°C overnight. Primary antibodies used are listed in Supplementary Table 2. After washing with PBS, sections were incubated with Alexa Fluor-488– or Alexa Fluor-594–labeled secondary antibodies (Abcam) at 37°C for 1 h. The nuclei were stained with DAPI (SouthernBiotech), as previously described (11).

The HK-2 cell line was obtained from ATCC and cultured as previously described (11). In this study, HK-2 cells were treated with different concentrations of d-glucose (5 mmol/L or 30 mmol/L). Furthermore, cells were transfected with pcDNA3 hPACS-2 flag plasmid (a gift from Professor Gary Thomas, Department of Microbiology and Molecular Genetics, University of Pittsburgh, Pittsburgh, PA) using a Lipofectamine 3000 reagent (Invitrogen) according to the manufacturer’s instructions.

Cell immunofluorescence (IF) was carried out as described previously (11). Briefly, HK-2 cells were seeded onto confocal dishes, grew to a nearly suitable state, and then were subjected to various treatments. The cells were incubated with MitoTracker Red (500 nmol/L; Invitrogen) solution. After washing, the cells were fixed, permeabilized, and blocked. Then, the cells were incubated with various antibodies at 4°C overnight. After washing, the cells were incubated with secondary antibodies conjugated with Alexa-Fluor, then were stained with DAPI. For monitoring mitophagy, HK-2 cells were transfected with COX8-mCherry-GFP plasmid, a gift from Dr. Zheng Dong (plasmid no. 78520; Addgene). Cells were visualized by an LSM 780 META laser scanning microscope (Zeiss). Colocalization analysis were performed using ImageJ Colocalization Finder.

HK-2 cells were transfected with flag-tagged PACS-2 or control vector for 48 h. Cells were lysed with immunoprecipitation lysis buffer (catalog 87787; Thermo Fisher Scientific) containing protease/phosphatase inhibitors and incubated with anti-flag magnetic beads (catalog 580022; Bimake) overnight at 4°C. The precipitated materials were used for Western blot analysis with anti-flag and anti-BECN1 antibodies.

ATP levels were determined via an ATP Assay Kit (Beyotime Biotechnology) used according to the manufacturer’s instructions.

The mitochondrial membrane potential of HK-2 cells was assessed by using tetramethylrhodamine, ethyl ester (Molecular Probes), as previously described (26).

The level of ROS, which represents oxidative stress in the kidney tissues, was evaluated by staining with dihydroethidium (DHE; Invitrogen) (11). The mitochondrial ROS production in HK-2 cells was measured by staining with MitoSox Red (5 μmol/L; Invitrogen) as previously described (11).

Briefly, for kidney tissues, the paraffin-embedded kidney sections were dewaxed, hydrated, antigen repaired, and blocked, and then the apoptosis was assessed by TUNEL (KeyGen Biotech Co., Ltd.) following the manufacturer’s instructions. TUNEL assay was also used to detect apoptosis in HK-2 cells.

Data were analyzed using SPSS, version 23.0, software (IBM Corp.) and GraphPad Prism, version 8.0, software. The values are presented as mean ± SEM and analyzed by Student t test or one-way ANOVA. Pearson correlation analysis was used to test the correlations between two numerical variables. P < 0.05 was considered statistically significant.

The experimental data sets generated and/or analyzed during this study are available from the corresponding author upon reasonable request. No applicable resources were generated or analyzed in the current study.

An STZ-induced DN mice model was established to investigate the expression of Pacs-2. As indicated in Fig. 1A, Pacs-2 was mainly expressed in renal tubules and was significantly downregulated in diabetic kidney in comparison with the control. The findings were confirmed by Western blot analyses (Fig. 1B and C). The expression of PACS-2 was also decreased in patients with DN compared with expression in control participants, and more pronounced reduction was observed in advanced DN as detected by immunohistochemical analyses (Fig. 1D and Supplementary Table 3). Furthermore, the expression of PACS-2 was positively correlated with eGFR (Fig. 1E) and negatively correlated with degrees of IFTA (Fig. 1F) in all participants.

To gain insight into the role of PACS-2 in the progression of DN, we generated mice lacking the Pacs-2 gene specifically in the PT by crossing Pacs-2 exon 6,7-floxed mice (Pacs-2^fl/fl mice) with Ggt1-Cre mice (Fig. 2A). Pacs-2^fl/fl; Ggt1-cre⁺ mice (i.e., Pacs-2^ptKO mice) were phenotypically normal and born at the expected mendelian ratio (Fig. 2B). PCR analysis of genomic DNA from renal cortex indicated the expected 458-bp band amplified from the null allele in Pacs-2^ptKO mice, whereas the 1,549 bp from the wild-type allele of the Pacs-2 gene was absent, suggesting successful recombination mediated by Ggt1-Cre (Fig. 2C). The 458-bp band signal representing the null allele was not detected in Pacs-2^Ctrl (Pacs-2^fl/fl; Ggt1-cre⁻) mice.

The deletion of the Pacs-2 gene was also confirmed by quantitative PCR of kidney cortices and Western blot analysis of primary PTCs isolated from Pacs-2^Ctrl mice and Pacs-2^ptKO mice (Fig. 2D and E). Compared with control mice, PTC-specific Pacs-2 deficiency did not elicit a significant increase in the of urinary albumin to creatinine ratio (ACR) and obvious histopathologic consequences (Fig. 2I and J). These findings are consistent with previous literature in which researchers reported Pacs-2 global knockout mice does not have overt phenotype unless being stressed (30).

DN was then induced in Pacs-2^Ctrl mice and Pacs-2^ptKO mice by administration of STZ (Fig. 2F). Both groups developed hyperglycemia after STZ treatment. There was no significant difference in body weight and blood glucose level between STZ-induced Pacs-2^Ctrl mice and Pacs-2^ptKO mice (Fig. 2G and H). However, urinary ACR was higher in diabetic Pacs-2^ptKO mice than in control diabetic mice (Fig. 2I).

Although there were no significant differences between the groups in terms of serum creatinine concentration (Supplementary Fig. 1), there were strong differences histologically. HE, PAS, and Masson staining of diabetic mouse kidneys revealed dilated cortical PTs with loss of brush borders, as well as tubular basement-membrane thickening and interstitial fibrosis when compared with control mouse kidneys. Interstitial fibrosis in diabetic kidneys was also substantiated by immunostaining of fibronectin. These lesions were further aggravated in diabetic Pacs-2^ptKO mice (Fig. 2J–L), which suggested that PACS-2 is a key regulator of tubular injury in DN.

PACS-2, as an integral MAM protein, has previously been reported to be involved in maintaining ER–mitochondria tethering through preventing BAP31-mediated mitochondria fragmentation and uncoupling from ER (20). By TEM, we first examined the mitochondrial morphology in PTCs. As shown in Fig. 3A and B, mitochondria from tubule cells in Pacs-2^ptKO mice were rounder and smaller compared with the elongated and tubular mitochondria observed in Pacs-2^Ctrl mice. In addition, in diabetic Pacs-2^Ctrl mice, 36% of mitochondria were fragmented, and more extensive fragmentation (50%), along with partial cristolysis, was detected in diabetic Pacs-2^ptKO mice.

Consistent with morphological changes, Western blot analysis of kidney cortex (the bulk of which is made up of PTs) protein revealed significantly increased expression of mitochondrial fission protein, including dynamin-related protein-1 (Drp1), mitochondrial fission protein 1 (Fis1), and decreased expression of mitochondrial fusion protein, including mitofusion 1 (Mfn1) and mitofusion 2 (Mfn2) in diabetic Pacs-2^Ctrl mice compared with nondiabetic control. These changes were more pronounced in diabetic Pacs-2^ptKO mice (Fig. 3E–J, Supplementary Fig. 2). Accompanied by increased mitochondrial fission and decreased fusion, fragmented mitochondria uncoupled from ER in TCs in Pacs-2^ptKO mice and diabetic Pacs-2^Ctrl mice, whereas the fewest ER–mitochondria contacts were observed in diabetic Pacs-2^ptKO mice (Fig. 3A and C). We also used an in situ PLA as an additional strategy to detect and quantify the number of MAMs and similar results were obtained (Fig. 3A and D). These observations from independent means of detection and analysis indicate that Pacs-2 ablation leads to alterations in mitochondrial structure and decreased MAM formation in PTCs.

Mitochondrial fission yields asymmetric daughter mitochondria. The daughter mitochondria with normal membrane potential could play a role in mitochondrial biogenesis, while unhealthy daughter mitochondria with loss of potential undergo segregation and subsequent degradation by mitophagy (31). Because autophagosomes form at MAM (16), we next attempted to figure out whether mitophagy was affected when PACS-2 was deleted in PTCs.

Kidney sections were costained for LC3, an autophagosome marker, and TOM20, a mitochondrial outer membrane protein. Colocalization between punctate LC3 and TOM20, which delineated activation of mitophagy, was largely reduced in tubules from Pacs-2^ptKO mice and diabetic Pacs-2^Ctrl mice compared with tubules from nondiabetic control mice, and the formation of mitophagosomes was further decreased in tubules from diabetic Pacs-2^ptKO mice (Fig. 4A). The expression of BECN1, PINK1, and PARKIN also displayed parallel changes as assessed by Western blot (Fig. 4B and C). Furthermore, we also evaluated the advanced steps of the mitophagic process by determining the degradation of mitochondrial proteins residing in the outer and inner membranes and the matrix (TOM20, TIM23, and COX4, respectively), and the autophagy substrate p62. In accordance with the aforementioned findings, mitochondrial clearance was further blocked in diabetic Pacs-2^ptKO mice compared with diabetic Pacs-2^Ctrl mice, as evidenced by more remarkable increase of TOM20, TIM23, and COX4, along with the accumulation of p62 (Fig. 4D and E).

Accumulated dysfunctional mitochondria have been recognized as a major source of ROS and proapoptotic factors. Here, we observed ROS overproduction and increased apoptosis in tubules of STZ-induced Pacs-2^Ctrl mice as detected by DHE and TUNEL staining, respectively, both of which were further aggravated in diabetic Pacs-2^ptKO mice (Fig. 4F–H). Taken together, these observations suggest that Pacs-2 is required for mitophagosome formation in tubules, and loss of Pacs-2 results in oxidative stress and cell death.

We next carried out a series of in vitro experiments to investigate whether overexpression of PACS-2 could rescue HG-induced effects in HK-2 cells and potntial mechanisms. First, we checked expression and subcellular localization of PACS-2 in TCs. Overall, the expression level of PACS-2 was decreased in HK-2 cells under HG conditions. Costaining with mitochondrial and ER markers demonstrated the MAM localization of PACS-2 in PTCs, whereas the amount of PACS-2 protein associated with mitochondria and MAM was largely diminished in response to HG stimulation (Fig. 5A).

Second, we examined whether PACS-2 overexpression could reverse the effects of HG on mitochondrial dynamics and MAM formation (Fig. 5B). HK-2 cells exposed to HG conditions had a higher percentage of fragmented mitochondria that less frequently attached to ER than those cells cultured at normal glucose (NG) levels (Fig. 5C–E). However, HK-2 cells transfected with PACS-2 plasmid cloning DNA subjected to HG conditions rescued the mitochondrial morphology changes, characterized as an interconnected network of tubular and elongated structures, which resembled NG conditions (Fig. 5C and D). This was accompanied by decreased expression of DRP1 and FIS1 and increased expression of MFN2, whereas the levels of MFN1 were not changed significantly (Fig. 5F). Furthermore, the integrity of MAM was also restored by PACS-2 overexpression (Fig. 5C and E).

Last, we tested the translocation of DRP1 to mitochondria in our experimental conditions, because it has been speculated, but not validated, that PACS-2 deficiency promotes mitochondrial fission by regulating DRP1 translocation to mitochondria in response to calcium influx. Here, we found that mitochondrial recruitment of DRP1 in HK-2 cells exposed to HG conditions was significantly blocked by PACS-2 overexpression (Fig. 5G and H).

A reduction of autophagy and mitophagy was noted in HK-2 cells treated with HG compared with NG, as reflected by the reduction of punctuate LC3 and puncta associated with mitochondria, both of which were partially recovered by overexpressing PACS-2 (Fig. 6A). This finding was further substantiated by Western blot analyses that showed increased LC3 cleavage and enhanced degradation of p62 and mitochondrial protein TIM23 and TOM20 (Fig. 6B and C). Notably, while maintained in medium with NG levels, HK-2 cells transfected with PACS-2 plasmid also displayed a substantial increase of LC3-II in comparison with control cells (Fig. 6B and C). We further monitored the delivery of mitochondria to lysosomes, using a mitochondrial matrix–localized EGFP-mCherry reporter (32,33). Mitochondria within lysosomes where EGFP fluorescence becomes quenched due to the acidic microenvironment show red-only fluorescence. When compared with HK-2 cells cultured in NG conditions, cells under the HG condition had fewer red-only mitochondria, and these were restored by PACS-2 overexpression (Fig. 6D and E).

Next, we sought to determine the potential mechanisms by which PACS-2 regulates mitophagy. PINK1 and BECN1 relocalize at MAM after mitophagic stimuli and promote the formation of mitophagosome (34). Considering that PACS-2 also functions as a membrane-trafficking protein that identifies and transfers cargoes with acidic cluster to corresponding organelles (35), we asked whether PACS-2 could play a role in the redistribution of PINK1 and BECN1 to MAM. By analyzing the protein sequence of the two molecules, we discovered that BECN1 contains a CK2-phosphorylatable acidic cluster, ET⁵⁶QEEE, a typical domain frequently found in PACS-2 client proteins. Moreover, this domain is highly conserved across species (Fig. 7A and B). To verify the interaction between PACS-2 and BECN1, we performed immunoprecipitation with HK-2 cells expressing FLAG-tagged PACS-2 and found significant enrichment of BECN1 (Fig. 7C). The interaction between PACS-2 and BECN1 was further validated by IF showing colocalization of the two molecules, which was significantly reduced under HG condition (Fig. 7D and E). Furthermore, the BECN1 levels at MAM were decreased in HK-2 cells exposed to the HG condition and were partially recovered by PACS-2 overexpression, suggesting that PACS-2 is required for BECN1 accumulation at MAM (Fig. 7F and G).

To link mitochondrial quality control improvement to mitochondrial fitness, we first examined mitochondrial ATP production. We found that overexpression of PACS-2 in HK-2 cells under HG conditions led to an ∼33% increase in ATP content, compared with cells transfected with empty control (Fig. 8A). Overexpression of PACS-2 in HK-2 cells also prevented HG-mediated decreases in mitochondrial membrane potential (Fig. 8B and C). Consistent with the improvement of mitochondrial function, mitochondrial ROS production and apoptosis in HK-2 cells exposed to HG conditions, as revealed by MitoSox and TUNEL staining, were also inhibited by overexpression of PACS-2 (Fig. 8B, D, and E). These changes were accompanied by a decreased BAX to BCL2 ratio and reduced expression of cleaved caspase-3 (Fig. 8F–H). To summarize, these observations indicated that PACS-2 could normalize the mitochondrial quality control and lead to mitigated ROS production and cell apoptosis in renal TCs treated with HG.

Here, we report a critical role of PACS-2 in protection of tubular injury in DN. PT-specific Pacs-2 deficiency exacerbates mitochondrial fragmentation, MAM integrity disruption, and mitophagy insufficiency in TCs of diabetic mice, resulting in worsening albuminuria and renal fibrosis. Our results also highlight the MAM mechanism as the important mediator that links PACS-2 to mitochondrial quality surveillance, thus preventing oxidative stress and cell apoptosis (Fig. 8I).

PACS-2 is a known MAM resident protein that can regulate ER–mitochondria contact formation and interorganelle communication (19,20). As the multifaceted contributions of MAM to cell homeostasis have been increasingly appreciated in different disease models, PACS-2 has been associated with insulin resistance in the liver and with vascular smooth muscle cell apoptosis through regulating MAM (21,36). However, PACS-2 expression and functions in the kidney have not been described to date to our knowledge. In this study, we demonstrate that PACS-2 mainly distributes in TCs in the kidney. Importantly, PACS-2 is significantly downregulated both in tubules from STZ-induced diabetic mice and patients with DN. In addition, the level of PACS-2 in tubules correlates well with tubulointerstitial fibrosis and kidney function in patients with DN, suggesting a potential role of PACS-2 in the pathophysiology of DN.

To verify the role of PACS-2 in DN, we generated PT-specific Pacs-2 knockout mice and induced the DN model by administration of STZ. Nondiabetic Pacs-2^ptKO mice did not have albuminuria or gross histological abnormalities, although mitochondrial fragmentation and MAM reduction were observed in tubules, suggesting that in a healthy animal, those changes were either tolerated or compensated and were insufficient to disrupt tubular structure and function. Similar observations were previously noted in the liver, where disturbance of MAM did not cause gross morphological changes and metabolic phenotype in lean mice until they were stressed with high-fat diet (30,36). In the present study, conditional Pacs-2 deletion in PTCs in STZ-induced diabetic mice induced a further decrease of the MAM amount, leading to significantly increased albuminuria and aggravated tubulointerstitial lesions. Overall, these data indicate that PACS-2 may be a new target molecule for tubular injury induced by hyperglycemia, by maintaining MAM integrity in TCs.

A more tubulocentric view of DN has challenged the conventional view of DN as a microvascular complication over the years. TCs have a high density of mitochondria, the homeostasis of which is critical for the proper functioning of tubules, especially in response to stressful stimuli. We and others previously provided a series of evidence that identified mitochondrial dysfunction as an important contributor to tubular damage in DN (2,18,26,37,38). In the present study, we show that Pacs-2 deficiency caused mitochondrial fragmentation and subsequent MAM uncoupling in TCs from both control and diabetic mice. In fact, it has been reported that loss of MAM integrity, in turn, induces mitochondrial fission and compromises mitochondrial respiration (18,28,39). Mechanistically, fragmentation of mitochondria that detached from ER in the context of PACS-2 deletion was reported to be BAP31/p20–dependent (20). We contributed to this hypothesis by verifying the downstream signaling by which PACS-2 could regulate mitochondrial recruitment of DRP1, a central step to drive division of mitochondria. Taken together, PACS-2 deficiency induces the cleavage of BAP31 to p20, which can promote Ca²⁺ transmission from ER to mitochondria and activate calcineurin-dependent dephosphorylation of DRP1, followed by DRP1 translocation to mitochondria and initiation of fission (20,40).

Besides being involved in the cycle with mitochondrial fission, MAM alteration also exerts effects on other intracellular events, including ER stress, lipid synthesis, autophagy and mitophagy, and apoptosis (16,20,34,41), which are implicated in the pathogenesis of DN. We have particular interest in the mitophagy process because mitophagy can eliminate depolarized mitochondria resulting from fission to sustain a population of healthy mitochondria. Studies have shown that mitophagy is inhibited in diabetic kidney, and pharmacologic activation of mitophagy can protect against the progression of DN (2,10). However, the mechanism underlying the regulation of mitophagy in TCs remains largely unexplored. Authors of a recent study suggested that MAM represents sites of mitophagosome formation (34). More recently, Moulis et al. (21) showed that PACS-2 knockdown impaired mitophagy and potentiated oxidized lipoprotein–induced apoptosis in vascular smooth muscle cells. Here, we found that the disruption of MAM integrity induced by PACS-2 deficiency blocked mitophagosome formation and mitophagy in PTCs in vivo, which was consistent with the results of these two studies (21,34). Moreover, overexpression of PACS-2 in vitro restored mitophagy and thus mitigated mitochondrial ROS production and apoptosis in HK-2 cells under HG conditions. These data indicate that PACS-2 plays a key role in removal of accumulated dysfunctional mitochondria by enhancing mitophagy through maintenance of MAM integrity.

We also explored the more precise mechanisms by which PACS-2 regulates mitophagy. Current knowledge of how MAMs orchestrate mitophagy is quite limited beyond knowing MAM is the membrane source of mitophagosome. Manganelli et al. (42) described the presence of cardiolipin within MAM as a signaling platform in mitophagosome formation. It has also been reported that BECN1 accumulated in the MAM fraction and enhanced the formation of autophagosome precursors in SH-SY5Y cells upon treatment of mitophagic stimuli in a PINK1-dependent fashion (34), which shed a new light on the role of PINK1 and BECN1 in mitophagy. Interestingly, we found that the expression level of PINK1 and BECN1 was decreased in tubules of diabetic mice and was even lower in tubules of diabetic Pacs-2^ptKO mice. Furthermore, PACS-2 directly interacted with BECN1 and mediated its translocation toward MAM. Hence, our results suggest that PACS-2 identifies BECN1 as a cargo protein and transports it from ER to MAM, where it enhances ER–mitochondria tethering and the formation of mitophagosome.

Although we did not explore the mechanism of downregulation of PACS-2 in renal tubules under diabetic conditions, we speculate that PACS-2 likely is modulated at the transcriptional level because both mRNA and protein levels were downregulated. Thus, using ChIPBase, version 2.0, we analyzed the coexpression data for Pacs-2 and transcriptional factors with binding sites within 1 kb upstream of the Pacs-2 promoter. Some transcriptional factors known to respond to HG were predicted to regulate the transcription of Pacs-2, such as CREB, FOXA2, and FOXO1 (43–45). More studies are warranted to investigate whether Pacs-2 downregulation in diabetic kidneys is associated with these transcriptional factors. Moreover, researchers have reported posttranslational regulation of PACS-2 by proteasomal degradation (46). It would also be interesting to explore whether diabetes induces ubiquitination of PACS-2 and results in its degradation.

We did not observe a correlation between HbA_1c and PACS-2 expression levels in kidney biopsy samples from control participants and patients with DN (data not shown), although results of in vitro experiments implied that HG per se could regulate PACS-2 expression. We speculate that these findings mainly were due to the limited sample size of the control group. Most participants in the control group lacked HbA_1c data because HbA_1c is not routinely tested in patients without diabetes. Thus, these observations need to be confirmed in a larger cohort with a prospective design.

Collectively, our study reveals a critical but previously unrecognized role of PACS-2 in maintaining mitochondrial homeostasis in TCs during DN. We also demonstrate that PACS-2 regulation of MAM integrity and BECN1 redistribution to MAMs are required for an elegant mitochondrial quality control network in TCs. Thus, understanding how PACS-2 protects against DN provides new insights into the mechanisms of tubular injury in DN and a novel potential target for the treatment of DN.

Acknowledgments. The authors thank Professor Gary Thomas (Department of Microbiology and Molecular Genetics, University of Pittsburgh, Pittsburgh, PA) for providing pcDNA3 h PACS-2 flag plasmid.

Funding. This work was supported by grants from the National Natural Science Foundation of China (grant 81730018) and the National Key R&D Program of China (grant 2018YFC1314002).

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. C.L. and L.L. performed the experiments and wrote the manuscript. M.Y., J.Y., C.Z., Y.H., H.Z., N.J., L.W., Y. Xiao, Y.L., X.X., Y. Xi, S.L., and F.D. provided technical support for this study and participated in discussions about this study. W.C., S.Y., X.Z., L.X., and L.S. designed this study and edited this manuscript. All authors approved the final version of the manuscript. L.S. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Prior Presentation. Parts of this study were presented at the annual congress of the Chinese Society of Nephrology, Zhuhai, China, 15–19 December 2020.