PANC-1 cell line was purchased from National Infrastructure of Cell Line Resource (Beijing, China). PANC-1 cells were cultured in RPMI-1640 medium (Thermo Fisher Scientific, Inc., Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; HyClone; GE Healthcare Life Sciences, Logan, UT, USA), 1% L-glutamine and 0.5% gentamycin (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) at 37°C in an atmosphere of 5% CO₂. Hypoxic conditions were induced in hypoxia chamber in a humidified atmosphere with 94% N₂, 5% CO₂ and 1% O₂ for 24 h (15). To inhibit the mitochondrial fission, mitochondrial division inhibitor 1 (Mdivi1; 10 mM; Sigma-Aldrich; Merck KGaA) was used for 12 h at 37°C.

The miR-125a mimic (agmir-125a), miR-control (scramble miRNA), miR-125a inhibitor (antagomir), small interfering RNA (siRNA) targeting hypoxia-inducible factor 1 (HIF1) and negative control siRNA (siCtrl) were purchased from GenePharma Co., Ltd. (Shanghai, China). The oligonucleotides used in these studies are as follows: Agmir-125a, 5′-CCACAUGAACGCCCAGAGAUU-3′; scramble miRNA, 5′-GAACGGGAGUACAGAGAGAUU-3′; antagomir, 5′-UAACAAGACCAGAGAGCUGUU-3′; siHIF1, 5′-GAGGAAAAGGGAAAAUCUAUU-3′; siCtrl, 5′-AAUUCUUAAAUUGGGCUGGUU-3′. siRNA (1 µg/ml) and miRNA (2 µg/ml) were added to the media (per 3.5×10⁴ cells/well) supplemented with Lipofectamine^® 2000 (Thermo Fisher Scientific, Inc.) Media containing siRNAs were replaced with Dulbecco's modified Eagle's medium (DMEM) 12 h after transfection. Transfection was performed for 72 h and the knockdown of gene expression was assessed by western blot analysis.

To determine the protein levels, 1×10⁶ cells were lysed with radioimmunoprecipitation assay (RIPA) buffer (Thermo Fisher Scientific, Inc.) supplemented with phenylmethylsulfonyl fluoride. The protein concentration was analyzed using the bicinchoninic acid protein assay. Protein (50 µg) was separated by 10% SDS-PAGE and then transferred to polyvinylidene difluoride membranes. The membranes were blocked with 5% nonfat milk for 1 h at room temperature and then incubated with primary antibodies: Caspase-3 (cat. no. 9662; 1:2,000), caspase-9 (cat. no. 9508; 1:2,000), Bcl-2 (cat. no. 3498; 1:2,000), X-linked inhibitor of apoptosis (x-IAP; cat. no. 2042; 1:1,000) and Mfn2 (cat. no. 11925; 1:1,000) from Cell Signaling Technology, Inc. (Danvers, MA, USA); Bcl-2-associated agonist of cell death (Bad; cat. no. ab32445; 1:1,000), Bcl-2 associated X, apoptosis regulator (Bax; cat. no. ab32503; 1:2,000), HIF1 (cat. no. ab16066; 1:1,000), poly(ADP-ribose) polymerase (cat. no. 32064; 1:2,000), complex II (cat. no. ab110410; 1:1,000), complex IV subunit II (cat. no. ab110268; 1:1,000), complex I subunit NDUFB8 (cat. no. ab110242; 1:1,000), density-regulated protein 1 (Drp1; cat. no. ab56788; 1:1,000), mitochondrial fission 1 protein (Fis1; cat. no. ab71498; 1:1,000), G-actin (cat. no. ab123034; 1:1,000) and F-actin (cat. no. ab205; 1:1,000) from Abcam (Cambridge, MA, USA); and complex III subunit core (1:1,000; cat. no. 459220) from Invitrogen (Thermo Fisher Scientific, Inc.) overnight at 4°C. The membranes were washed in TBS Tween-20 for 15 min and then incubated with a horseradish peroxidase-conjugated secondary antibody (cat. nos. sc-2004 and sc-2005; Santa Cruz Biotechnology, Inc., Dallas, TX, USA) for 1 h at room temperature. Blots were detected via an enhanced chemiluminescence substrate kit (Thermo Fisher Scientific, Inc.), and were analyzed using Quantity One 4.6 software (Bio-Rad Laboratories, Inc., Hercules, CA, USA) (23).

To over-express Mfn2, the pDC316-mCMV-Mfn2 plasmid (pDC316-mCMV-Mfn2; NheI-forward, 5′-TATCTCATCAGATTGAGCTCGTCCA-3′ and HindIII-reverse, 5′-CGCCTTAGATCCACTCACTGTAGATTCGA-3′) was purchased from Vigene Biosciences, Inc. (Rockville, MD, USA). This pDC316-mCMV-Mfn2 plasmid (2.5 µg, per 3.5×10⁴ cells/well) was transfected into 293T cells (National Infrastructure of Cell Line Resource) in RPMI-1640 medium supplemented with 10% FBS using Lipofectamine^® 2000 according to the manufacturer's protocol. The viral supernatant was collected 48 h after transfection. Supernatant was acquired again and filtered through a 0.45-µm filter to obtain the adenovirus-Mfn2 (Ad-Mfn2). A total of 1×10⁵ cells/well were infected with 100 multiplicity of infection adenovirus in serum-free DMEM for 6 h at 37°C, following which the media was replaced with DMEM supplemented with 10% FBS.

Total RNA was extracted with TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.) and reverse transcribed with a One-step RT-PCR kit (TransGen Biotech Co., Ltd., Beijing, China) at 42°C for 3 min according to the manufacturer's instructions. The mRNA levels were determined by RT-qPCR in triplicate for each of the independently prepared RNAs and were normalized to the levels of GAPDH expression (24). Gene expression was determined 90 using cBNA SYBR-Green real-time PCR Master Mix (Takara Bio, Inc., Otsu, Japan) and qPCR. Primers for qPCR used were as follows: miR-125a reverse transcription primer, 5′-GTCGTATCCAGTGCAGGGTCCGAGGTATTCGCACTGGATACGACTCACAGG-3′; miR-125a PCR primers, sense, 5′-CTGGAGUCCCUGAGACCCUUUA-3′ and antisense, 5′-ACGCTTCACGAATTTGCGTGTC-3′; GAPDH, sense, 5′-TGAGTGCTGTCTCCATGTTTGA-3′ and antisense, 5′-TCTGCTCCCCACCTCTAAGTTG-3′. qPCR was performed at 94°C for 3 min and 94°C for 30 sec for 38 cycles, and finally 51°C for 30 sec. The mRNA ratio of the target genes to β-actin was calculated using the 2^−ΔΔCq formula (25).

A TUNEL assay was performed using a one-step TUNEL kit (Beyotime Institute of Biotechnology, Haimen, China) according to the manufacturer's instructions. TUNEL staining was performed with fluorescein-dUTP (Invitrogen; Thermo Fisher Scientific, Inc.) to stain apoptotic cell nuclei, and DAPI (5 mg/ml) was used to stain all cell nuclei at room temperature for 3 min. Cells in which the nucleus was stained with fluorescein-dUTP were defined as TUNEL positive. The slides were then imaged under a confocal microscope (26).

Hypoxic conditions were induced using fresh PBS solution with 94% N₂, 1% O₂ and 5% CO₂. The pH was adjusted to pH 6.8 with lactate (Sigma-Aldrich; Merck KGaA) to mimic ischemic conditions. The dishes were placed into a hypoxia incubator that was equilibrated with 94% N₂, 1% O₂ and 5% CO₂. Deferoxamine (DFX; 10 mg/ml) was used to treat cells for ~4 h to induce the hypoxia condition at 37°C.

Cell proliferation was measured with a Cell Counting Kit-8 (CCK-8) assay (Beyotime Institute of Biotechnology) (27). Briefly, 200 µl of the cell suspension was seeded in 96-well cell culture plates at a density of 1,000 cells/well and incubated at 37°C for 1–4 days as previously described (28).

Following treatments, PANC-1 cells were seeded at a density of 0.5×10⁶ cells/well in 6-well plates and then cultured overnight (90% confluence). A wound track was scored in each dish with a pipette head. Debris was removed by washing with PBS. After 0, 24 and 48 h of culturing, the migration distances were visualized and imaged (Olympus IX71; Olympus Corporation, Tokyo, Japan). Cell migration was also analyzed using a Transwell chamber assay (24 wells, 8-µm pore size with a polycarbonate membrane) as previously described (29).

A caspase-3 activity kit and a caspase-9 activity kit (Beyotime Institute of Biotechnology) were used to detect the activity of caspase-3 and caspase-9, respectively, according to the manufacturer's protocol (30). Following the appropriate treatments, cultured cells were lysed with RIPA lysis buffer (Beyotime Institute of Biotechnology) for 30 min and centrifuged at 14,000 × g for 30 min at 4°C. The absorbance was measured at a wavelength of 405 nm using a microplate reader (BioTek Instruments, Inc., Winooski, VT, USA). The data are expressed as the ratio of the optical density (OD) value of the treated group to the OD of the control group. The extracellular lactate was measured using the cell culture medium with lactate assay kit (cat. no. K607-100; BioVision, Inc., Milpitas, CA, USA). Intracellular glucose was measured using cell lysates with glucose assay kit (cat. no. K606-100; BioVision, Inc.). The uptake of glucose, the production of lactate and the levels of ATP were all measured according to the manufacturer's instruction. The assay was repeated three times.

mtDNA strand breaks were detected based on methods described previously (11). Briefly, a 200 µl cell (1×10⁶) suspension was centrifuged at 15,000 × g at 4°C for 20 min. The supernatant was discarded and 400 μl solution (0.25 mmol/l inositol, 10 mmol/l Na₃PO₄ and 1 mmol/l MgCl₂, pH 7.2) was added at 4°C for 30 min (31). The relative amounts of mtDNA and nuclear DNA content were used to assess the mtDNA copy numbers via qPCR, which was performed as described above. The mtDNA and nuclear ampli-cons were generated from a complex IV sequence and GAPDH segment, respectively. The mtDNA primers were 5′-CTATGTCGTGTCCAGAG-3′ and 5′-CATGTTGTCCCGTGTCATG-3′. The GAPDH primers, chosen as the internal standards, were 5′-CTCAGTCGTATTCGAGTGGTCCT-3′ and 5′-CCTGTGGAAGTCCACAACATGTC-3′. The transcript level of mtDNA was reflected by two different components: NADH dehydroge-nase subunit 1 (ND1) and cytochrome c oxidase subunit I. The primers for cytochrome c oxidase subunit I were 5′-ATCGTTCGGTGAGGTCGTG-3′ and 5′-CGCCGGTGTCATTATCGTATA-3′. The primers for ND1 were 5′-TTGCCGTATATTCAGTATC-3′ and 5′-ATCCTGTTGCCCAGTCCAGT-3′. GAPDH was selected as the internal standard (32).

The cellular ATP levels were measured using a firefly luciferase-based ATP assay kit (Beyotime Institute of Biotechnology). The opening of mPTP was visualized as a rapid dissipation of tetramethylrhodamine ethyl ester fluorescence as described in a previous study (33). Mitochondrial respiration was initiated by adding glutamate/malate to final concentrations of 5 and 2.5 mmol/l for 5 min, respectively. State 3 respiration was initiated by adding ADP (150 nmol/l) for 5 min; state 4 was measured as the rate of oxygen consumption after ADP phosphorylation. The respiratory control ratio (state 3/state 4) and ADP/O ratio (number of nmol ADP phosphorylated to atoms of oxygen consumed) were calculated as previously described (34). Mitochondrial depolarization was evaluated using MitoProbe™ JC-1 assay kit (Thermo Fisher Scientific Inc.), according to the manufacturer's protocol.

To determine cytochrome c (cyt-c) localization and mitochondrial division, immunofluorescence staining was used. Cells were fixed in 3.7% paraformaldehyde for 10 min at room temperature and permeabilized in 100% pre-chilled acetone (Sinopharm Chemical Reagent Co., Ltd., Shanghai, China). Following blocking with 5% bovine serum albumin (Sigma-Aldrich; Merck KGaA) in PBS for 1 h at room temperature, the cells were incubated with primary antibodies for 4 h at room temperature. Subsequently, the cells were incubated with Alexa-Fluor 116 488 donkey anti-rabbit secondary antibody (1:1,000; cat. no. A-21206; Invitrogen; Thermo Fisher Scientific, Inc.) at 37°C for 1 h in the dark. Images were captured using a laser confocal microscope (TcS SP5; Leica Microsystems, Inc., Buffalo Grove, IL, USA). The primary antibodies used for cell immunofluorescence were cyt-c (1:500; cat. no. ab90529), translocase of outer mitochondrial membrane 20 (1:500; cat. no. ab56783) and F-actin (1:500; cat. no. ab205) from Abcam. DAPI (5 mg/ml; Sigma-Aldrich; Merck KGaA) was used to stain the nucleus at room temperature for 3 min (35).

Following treatment, cells were collected and fixed with 3% glutaraldehyde in 100 mM cacodylate buffer at 4°C overnight, post-fixed in 1% cacodylate-buffer osmium tetroxide for 2 h at room temperature, and dehydrated in a graded series of ethanol (50, 70, 90 and 100% for 20 min each). Then, cells were embedded in EponAradite. Ultrathin sections (60 nm) were cut with a diamond knife on a Leica EM UC6rt (Leica Microsystems GmbH, Wetzlar, Germany) and double-stained with uranyl acetate and lead citrate. The ultrastructure of cells was observed with a Hitachi H7650 transmission electron microscope (TEM; Hitachi, Ltd., Tokyo, Japan) at 80 kV. Three slides were used in each experiments and the TEM assay was repeated three times (36).

Wild-type Mfn2 3′-UTR (WT) and mutant Mfn2 3′-UTR (MUT) containing the putative binding site of miR-125a were chemically synthesized and cloned downstream of the firefly luciferase gene in a pGL3-promoter vector (Promega Corporation, Madison, WI, USA). PANC-1 cells were placed on a 48-well plate and cultures until 80% confluence (37). Cells were then co-transfected with luciferase plasmids (2.5 µg per 3.5×10⁴ cells/well) and miR-125a or control miRNA in DMEM medium supplemented with 10% FBS using Lipofectamine^® 2000 according to the manufacturer's protocol. The pRL Renilla control reporter vectors (Promega Corporation) was used as an internal control to normalize the values of the experimental reporter gene. At 48 h after transfection, the intensities were measured with a Luciferase Reporter Assay System (Promega Corporation).

Mito-ROS levels were measured using flow cytometry with a MitoSOX red mitochondrial superoxide indicator (Molecular Probes; Thermo Fisher Scientific, Inc.). Cells (3.5×10⁶ cells/well) were plated in 6-well culture plates. Subsequently, cells were incubated with MitoSOX (25 µM) in PBS at 37°C for 30 min, washed twice with PBS, and detached by treatment with trypsin-EDTA. The detached cells were collected and resuspended in PBS, and the fluorescence intensity of cells (3.5×10⁶ cells/well) was measured using flow cytometry (BD FAcSVerse) and analyzed via BD Paint-A-Gate™ Pro software (version 4.2) (both from BD Biosciences) (38). ETCx activities were analyzed via ELISA according to the manufacturer's protocol. The ELISA assay kits for ETCx I, II, and V were purchased from Beyotime Institute of Biotechnology (cat. nos. S0052, S0101 and S0052) (39).

All analyses were performed with SPSS 20.0 software (IBM Corporation, Armonk, NY, USA). All experiments were repeated three times. The data are presented as the mean ± standard deviation and statistical significance for each variable was estimated by a one-way analysis of variance followed by Tukey's test for the post hoc analysis. P<0.05 was considered to indicate a statistically significant difference.

Initially, to verify the role of miR-125a in regulation of the physiological processes of PANC-1 cells, a mimic (agmir-125a) and inhibitor (antagomir) of miR-125a were used. A CCK-8 assay was used to evaluate the growth capacity of PANC-1 cells. As shown in Fig. 1A, agmir-125a reduced the viability of PANC-1 cells, whereas application of antagomir marginally promoted cellular growth, as evidenced by higher OD values in the CCK-8 assay. Furthermore, expression of cleaved caspase-3 and its substrate, indicators of cellular apoptosis, were also significantly elevated in response to agmir-125a treatment in PANC-1 cells (Fig. 1B). To further understand whether miR-125a modifies cellular survival, TUNEL staining was used (Fig. 1C). Similarly, introduction of miR-125a via agmir-125a augmented the ratio of TUNEL positive cells. By contrast, inhibition of miR-125a reduced the percentage of TUNEL positive cells. These data indicated that miR-125a is a pro-apoptotic factor in PANC-1 cells.

To explore the mechanism by which miR-125a regulates PANC-1 cell death, mitochondrial damage was investigated. Recent studies have suggested that mitochondrial fission is an early event that triggers mitochondria-related apoptosis pathways (40,41). Therefore, the change of mitochondrial morphology was evaluated. As shown in Fig. 2A, compared to spindle mitochondria in the control group, agmir-125a markedly increased the amount of fragmented mitochondria, as evidenced by more round and mitochondrial debris. However, application of the mitochondrial fission inhibitor Mdivi1 blocked the effects of agmir-125a on mitochondria fragmentation. By contrast, antagomir treatment led to more mitochondria with a longer length compared with the control group. These data indicated that miR-125a activates mitochondrial fission. Additionally, to investigate whether mitochondrial fission was associated with apoptosis, proteins associated with mitochondrial damage were evaluated. Introduction of agmir-125a increased Bax, Bad and caspase-9 expression, and reduced Bcl-2 and x-IAP expression, suggesting activation of mitochondria-associated apoptosis pathways (Fig. 2B–H). However, the mitochondrial fission inhibitor blocked the pro-apoptotic effects of agmir-125a. These data indicated that the excessive fission activated by miR-125a contributes to the initiation of mitochondria-associated apoptosis pathways. Furthermore, mitochondrial apoptosis is induced due to mitochondrial membrane potential dissipation, mPTP opening and subsequent cyt-c leakage into the cytoplasm, which activates caspase-9 and caspase-3. Therefore, we observed upstream changes in the context of miR-125a-mediated fission. As shown in Fig. 2I, agmir-125a treatment destroyed the membrane potential with evidence of decreased red fluorescence, but increased green fluorescence. However, the mitochondrial fission inhibitor reversed these changes. Furthermore, the mPTP opening rate was increased upon application of agmir-125a, but decreased when treated with fission inhibitors (Fig. 2J). The agmir-125a also caused more cyt-c leakage from mitochondria into the cytoplasm, and some cyt-c even migrated into the nucleus (Fig. 2K). However, these changes were blocked by the mitochondrial fission inhibitor. Together, these data indicate that miR-125a activates mitochondrial fission, which induces mitochondrial potential collapse, mPTP opening and cyt-c leakage into the cytoplasm and nucleus, leading to activation of caspase-9-dependent mitochondria apoptosis pathways.

In addition to activating apoptosis, the central role of mitochondria is to generate energy and regulate metabolism, which are fundamental for tumor development and progression (42,43). Therefore, we observed energy or metabolism alterations upon miR-125a stimulation. As shown in Fig. 3A, agmir-125a reduced the contents of double-stranded mtDNA (Fig. 3A), mtDNA copy number (Fig. 3B), mtDNA transcripts (Fig. 3C) and ATP generation (Fig. 3D) in PANC-1 cells, which were blocked by the fission inhibitor. Electron transport chain complexes (ETCx) are mainly encoded by mtDNA and primarily responsible for ATP generation via using hydrion, electron and oxygen. ETCx expression and activity is vital to delivery hydrion and electron to oxygen, favoring to the mitochondrial oxidative phosphorylation. Accordingly, the downregulation and inactivation of ETCx would predispose the energy undersupply. Based on this, the influence of miR-125a on ETCx expression and activity was evaluated. The results shown in Fig. 3E–J demonstrate that agmir-125a treatment significantly reduced the ETCx concentration and activity, which are coupled to the decrease of the state 3 respiratory rate. Furthermore, agmir-125a-treated PANC-1 cells took up less glucose and therefore produced less lactate. However, the fission inhibitor abolished the inhibitory effects of agmir-125a on PANC-1 cells. Additionally, the inhibitor of miR-125a slightly elevated ETCx activity (P>0.05) and the subsequent mitochondrial respiratory function, which were also associated with increased glucose consumption and lactate production. Similar results were observed with mitochondrial reactive oxygen species (Fig. 3K). These data suggest that mitochondrial fission activated by miR-125a impairs PANC-1 cell energy metabolism.

Apart from cellular survival and energy metabolism, cellular migration is another factor that contributes to tumorigenesis (44). Therefore, whether mitochondrial fission has a role in cellular migration was investigated. Initially, a wound-healing assay was used to evaluate the influence of miR-125a on the mobilization of PANC-1 cells. As shown Fig. 4A, exogenously administered agmir-125a reduced the migration of PANC-1 cells, while the mitochondrial fission inhibitor blocked such changes. These data indicated that miR-125a has a role in cellular migration via fission. As F-actin is the key stress fiber that directly regulates cellular mobilization, it was hypothesized that the blunted migration was derived from mitochondrial fission-involved F-actin dyshomeostasis. Fluorescence was also used to obverse F-actin changes. As shown in Fig. 4B, agmir-125a significantly reduced the fluorescence intensity of F-actin, but increased the amount of mitochondrial debris, in PANC-1 cells, which were reversed by the mitochondrial fission inhibitor and F-actin depolymerization inhibitors. As F-actin is composed of G-actin, it was hypothesized that there was an imbalance between F-actin and G-actin. As expected, agmir-125a treatment accelerated F-actin division into G-actin, as evidenced by increased G-actin and decreased F-actin protein expression in the agmir-125a group (Fig. 4C and D). Mdivi1 blocked these conformation alterations, suggesting that mitochondrial fission is responsible for F-actin depolymerization to G-actin. Furthermore, inhibition of F-actin degradation via Jasplakinolide reversed migration under agmir-125a treatment, which is similar to the results in the Mdivi1 group (Fig. 4A–D). Similar results were observed in Transwell assays (Fig. 4E). Together, these data indicated that miR-125a impairs cellular migration by inducing F-actin depolymerization into G-actin by activating mitochondrial fission.

To determine the mechanism by which miR-125 regulates mitochondrial fission, the change of Mfn2, which prevents excessive mitochondrial fission was investigation. Treatment with agmir-125a reduced the contents of Mfn2 in PANC-1 cells. However, introduction of miR-125a antagomir reversed expression of Mfn2 in PANC-1 cells (Fig. 5A). The similar results were observed for other mitochondrial fission makers, including Fis1 and Drp1. These data indicated that miR-125a negatively regulated the expression of Mfn2. Furthermore, through analysis of the gene sequences, miR-125a was matched to Mfn2, which is highly conserved among rat, human and mouse (Fig. 5B). To test whether miR-125a directly targeted the 3′-UTR of Mfn2, luciferase assays were performed using 3′-UTR sequence fragments containing the predicted target of Mfn2 and its mutated version inserted downstream of a luciferase reporter (Fig. 5C). The results demonstrated that the luciferase activity was downregulated in cells co-transfected with miR-125a mimics and the wild-type Mfn2 3′-UTR compared with the mutated type, suggesting that Mfn2 is a target gene of miR-125a. These data indicate that miR-125a directly regulates transcription and expression of Mfn2.

To confirm whether Mfn2 was responsible for fission, Mfn2 was overexpressed using an adenovirus (Ad-Mfn2) in PANC-1 cells. The transfection efficiency is shown in Fig. 5D. Mfn2 overexpression via Ad-Mfn2 blocked the promotive effects of agmir-125a on mitochondrial fission, as demonstrated by less mitochondrial fragmentation or debris in PANC-1 cells (Fig. 5E). These data confirmed the hypothesis that miR-125a triggers mitochondrial fission by inhibiting Mfn2 expression. Additionally, transfection with Ad-Mfn2 also ablated caspase-9 activity, which was upregulated in response to agmir-125a stimulation (Fig. 5F), suggesting that Mfn2 is also associated with mitochondrial fission-mediated mitochondrial apoptosis.

Finally, to elucidate the mechanism by which miR-125a is downregulated in PANC-1 cells, HIF1 was investigated. Several studies have reported that miR-125a is negatively modified by HIF1 and that hypoxic conditions further repress expression of miR-125a (45,46). Thus, it was hypothesized that HIF1 is the upstream regulator of miR-125a. siRNA was used to knockdown expression of HIF1 in PANC-1 cells, and the silencing efficiency of the siRNA is shown in Fig. 6A. Following HIF1 knockdown, mRNA expression of miR-125a was significantly increased. Additionally, hypoxic conditions were induced using a hypoxia chamber with 1% oxygen to enhance the HIF1 signals. Under hypoxic conditions, miR-125a was further downregulated. Furthermore, DFX was also used to mimic hypoxic conditions, and the results were in agreement with the above findings. These data indicated that miR-125a is negatively regulated by HIF1. To provide further evidence regarding the role of HIF1 in fission, the influence of HIF1 on Mfn2 expression was evaluated. Knockdown of HIF1 also reduced expression of Mfn2, which was increased under hypoxic conditions, suggesting a regulatory role for HIF1 in Mfn2 (Fig. 6B and C). Furthermore, the TEM results indicated that more fragmented mitochondria appeared in response to HIF1 silencing. However, upon exposure to hypoxic conditions, mitochondrial fission was inhibited, as demonstrated by longer mitochondria (Fig. 6D). Collectively, these data confirm that HIF1 inhibits mitochondrial fission via the miR125a/Mfn2 pathways.