Hepatocellular carcinoma (HCC) is the most common type of primary liver cancer and has a poor prognosis. Pituitary tumor transforming gene 1 (PTTG1) is highly expressed in HCC, suggesting it could play an important role in hepatocellular carcinogenesis. Here, we evaluated the impact of PTTG1 deficiency on HCC development using a diethylnitrosamine (DEN)-induced HCC mouse model and a hepatitis B virus (HBV) regulatory X protein (HBx)–induced spontaneous HCC mouse model. PTTG1 deficiency significantly suppressed DEN- and HBx-induced hepatocellular carcinogenesis. Mechanistically, PTTG1 promoted asparagine synthetase (ASNS) transcription by binding to its promoter, and asparagine (Asn) levels were correspondingly increased. The elevated levels of Asn subsequently activated the mTOR pathway to facilitate HCC progression. In addition, asparaginase treatment reversed the proliferation induced by PTTG1 overexpression. Furthermore, HBx promoted ASNS and Asn metabolism by upregulating PTTG1 expression. Overall, PTTG1 is involved in the reprogramming of Asn metabolism to promote HCC progression and may serve as a therapeutic and diagnostic target for HCC.

Significance:

PTTG1 is upregulated in hepatocellular carcinoma and increases asparagine production to stimulate mTOR activity and promote tumor progression.

Hepatocellular carcinoma (HCC), the most prominent form of liver malignancy, is the second most common cause of cancer-related mortality worldwide (1). HCC is associated with high recurrence and metastasis, which lead to unsatisfactory 5-year survival rates (4%–17%), despite the application of various novel treatments (2, 3). Hepatitis B virus (HBV) infection is the most dominant risk factor for HCC, accounting for nearly half of all HCC cases (4). The HBV regulatory X protein (HBx), designated a “viral oncoprotein” (5), is encoded by the open reading frame (ORF) of the X region of HBV and plays a vital role in HBV-associated HCC, although the mechanism remains unclear. HBx has been reported to activate cell-cycle regulation, signaling pathways, DNA repair, and metabolism regulation (6–8). It has also been shown to sensitize hepatocytes to oncogenic cofactors.

Cancer cells can rewire their metabolism to meet the increased energetic and anabolic needs of sustained cell proliferation (9, 10). Amino acid (AA) metabolism occupies central positions in tumor metabolic reprogramming and is an important driver of tumor growth and malignant progression. Related therapeutic interventions targeting arginine, glycine, and glutamine (Gln) metabolism have shown promising anticancer effects (11–14). As a downstream metabolite of Gln, asparagine (Asn) also plays a crucial role in tumor mitochondrial respiration, cell survival, and tumor progression (15–17). Asn synthetase (ASNS) catalyzes the conversion of aspartate to Asn, fulfilling the demand of tumor cells for Asn (18). Current research indicates that ASNS is involved in some typical oncogenic pathways, such as the PI3K–AKT–mTOR and KRAS–ATF4 pathways (19, 20). ASNS can also be used as a biological marker for L-asparaginase) treatment in patients with acute lymphoblastic leukemia (ALL; ref. 21). However, the exact role of Asn and ASNS in HCC initiation/promotion has not yet been fully elucidated.

Pituitary tumor transforming gene 1 (PTTG1), a pituitary-derived transforming gene, has been reported to be a proto-oncogene involved in proliferation and metabolism (22–27). PTTG1 can inhibit sister chromatid separation, regulate the cell-cycle, and inhibit tumor cell apoptosis (28). Moreover, PTTG1, as a transcription factor, exerts its procancer effects by directly binding to several gene promoters or indirectly binding to transcription factors, such as p53, Sp1, and The peripheral blood film (PBF; refs. 29–32). Previous studies were preliminary and suggested that PTTG1 was associated with the proliferation of tumors (25, 33). Hamid and colleagues showed that PTTG1 overexpression in HCC affects tumor growth and progression by stimulating the expression and secretion of FGF-2 and VEGF (34). Moreover, PTTG1 expression has been reported to be regulated by HBx protein–mediated ubiquitination (35). Li and colleagues reported that HBV replication caused an obvious increase of PBF, which facilitates PTTG1 nuclear translocation (36). Our previous study verified that PTTG1 is overexpressed in HCC (37). PTTG1 appears to play a multi-faceted role in cancer promotion by regulating multiple different oncogenic processes. Therefore, it is necessary to further explore its mechanism in liver cancer, which can provide new insights into the diagnosis and treatment of HCC.

In this study, we assessed how PTTG1 promotes Asn metabolic growth pathways to affect HCC progression. First, we determined that PTTG1 deficiency significantly inhibited the occurrence and development of liver cancer. In addition, we found that HBx, as a crucial oncogenic factor, upregulated PTTG1 and promoted its nuclear localization. Furthermore, PTTG1/ASNS/mTOR promoted the proliferation of HCC. In conclusion, our study indicated that PTTG1 contributed to the development and progression of HCC through ASNS-mediated Asn metabolism. Thus, PTTG1 has the potential to be a biomarker and therapeutic target for HCC.

Tissue samples

In this study, 12 HCC tissues and paired nontumor tissues were used for Western blotting analysis. Eighty HCC and paired nontumor tissues (Supplementary Table S1) were used for the tissue microarray (TMA). All specimens were subjected to further pathologic confirmation. This study was approved by the Research Ethics Committee of The Third Affiliated Hospital of Sun Yat‐Sen University (Guangzhou, China). Before tissue acquisition (2014; refs. 2–7), written informed consent was obtained from each patient according to the policies of the committee. The transcriptome cases obtained from The Cancer Genome Atlas (TCGA) Liver Hepatocellular Carcinoma (LIHC) dataset (https://portal.gdc.cancer.gov) were used for survival analysis and downstream enrichment.

Mice and treatments

Wild-type (WT; HBx−/−) and HBx+/+ littermates on a mixed genetic background (C57BL/6 and CBA) were generated from HBx heterozygous transgenic male and female mice (kindly provided by Dr. D.Y. Yu, Korea Research Institution of Bioscience and Biotechnology, Korea). WT (PTTG1+/+) and PTTG1−/− littermates on a C57BL/6 background were derived from heterozygote intercrosses (provided by Saiye Biotechnology Co., Ltd.). HBx+/+ PTTG1−/− and HBx+/+ PTTG1+/+ littermates were generated from a cross between HBx+/+ mice and PTTG1−/− mice. Genotyping was performed as previously described (38). Male mice were used in all studies. The mice were housed in microisolator cages under a 12-hour dark–light cycle (lights on at 8:00 am) with food and water ad libitum. In the diethylnitrosamine (DEN; Sigma, N0756)-induced HCC model, 15 mg/kg DEN was intraperitoneally injected into 14-day-old WT (PTTG1+/+) and PTTG1−/− mice, and low-dose DEN (5 mg/kg) was administered to 14-day-old HBx+/+ PTTG1+/+ and HBx+/+ PTTG1−/− mice in the same manner. These mice were sacrificed after 9 months. The equivalent volume of physiologic saline was intraperitoneally injected as vehicle treatment. In the spontaneous HCC model, HBx+/+ PTTG1+/+ and HBx+/+ PTTG1−/− mice were sacrificed at 18 months. The number and size of surface tumor nodules in each liver lobe and the liver weight were measured and recorded. The sedative consisted of xylazine (15 mg/kg) and ketamine (50 mg/kg) was given intraperitoneally as the anesthesia. Carbon dioxide inhalation was used as the method of euthanasia. Animal research protocols were authorized by the Institutional Animal Care and Use Committee at the Third Affiliated Hospital of Sun Yat-Sen University.

Cell culture and treatment

Four human liver cancer cell lines (HepG2, Hep3B, Huh7, and Hep2.2.15) were maintained in DMEM (Gibco BRL) supplemented with 10% FBS (Gibco BRL) in an incubator with 5% CO2 at 37°C. HepG2 (HB-8065), Hep3B (HB-8064), and Huh7 (CRL-3216) cell lines were obtained from the ATCC. HepG2.2.15 cell lines were provided by the Guangdong Provincial Key Laboratory of Liver Disease Research, China. Short tandem repeat (STR) profiling was used to identify all cell lines. The cell lines were routinely tested for Mycoplasma and cultured for a maximum of 20 passages (∼ 2 months). Before performing experiments, all cell lines were allowed to recover for at least two passages. For drug intervention, cells were treated with 20 μmol/L rapamycin (MCE, HY-10219), 1 IU/ml L-asparaginase (Prospec, enz-287), and 0.1 mm L-Asn (Yuan Ye, S20385).

Transient transfection, generation of stable cell lines, and RNA interference

The 1.3-mer HBV genomic DNA and the HBV 1.3-mer X-null replicon plasmids were gifted from Wang-Shick Ryu (Addgene, 65459; Addgene, 65461). PTTG1-Flag, HBx-HA and ASNS were cloned into the pcDNA3.1 vector. siPTTG1 (5′-GTCACCAACAACACCAACA-3′) and siASNS (5′-TGTATGTTCAGAAGCT AAA-3′) were purchased from GenePhama. Cells were transfected using Lipofectamine 3000 (Invitrogen) according to the manufacturer's protocol. PTTG1-shRNA and shNC were cloned into the lentiviral vector GV248 (Shanghai Genechem). PTTG1-LvRNA and LvNC were cloned into the lentiviral vector GV367 (Shanghai Genechem). Stable transfections were selected with puromycin for 2 weeks before the next experiments.

RNA extraction and real-time PCR

Total RNA was extracted from tissues and cells by using TRIzol (Invitrogen, 15596–018) according to the manufacturer's instructions. RNA was transcribed into cDNA using a High Capacity cDNA Kit (TOYOBO, FSQ101). Then, qPCR analysis was performed using gene‐specific primers and ChamQ SYBR qPCR Master Mix (Vazyme, Q411) in a real‐time PCR system (Bio‐Rad). The sequences of the primers are listed as follows:

Human ASNS: 5′-CTGTGAAGAACAACCTCAGGATC-3′;

5′-AACAGAGTGGCAGCAACCAAGC-3′;

Human β-actin: 5′-CTGGAACGGTGAAGGTGACA-3′;

5′-AAGGGACTTCCTGTAACAATGCA-3′

Coimmunoprecipitation assay

For the coimmunoprecipitation (co-IP) assay, HepG2 cells transfected with HBx-HA and PTTG1-Flag or HBxmutant -HA and PTTG1-Flag cells were used to obtain protein lysates. Anti-HA magnetic beads and antiFlag magnetic beads (10 μL) were conjugated with antibodies against IgG (CST) at room temperature for 1 hour. The cell lysates were cleared, and preconjugated magnetic beads were added to the supernatant to pull the immune complexes by spinning them at 4°C overnight. The beads were washed three times with 0.1% PBS, soaked in lysis buffer and heated for 10 minutes at 70°C. The supernatant was subjected to Western blotting assay with Flag and HA-specific antibodies.

Isolation of nuclear-cytosolic fractions

The Nuclear and Cytoplasmic Protein Extraction Kit (Beyotime) was applied for nuclear and cytosolic fraction separation according to the manufacturer's instructions. The isolated proteins were quantified by a bicinchoninic acid detection kit (P0011, Beyotime Biotechnology) and prepared for Western blotting analysis. Histone 3 and β-actin were used as loading controls for the nuclear and cytoplasmic fractions, respectively.

Cell viability and colony formation assay

Cell viability assays were performed using Cell Counting Kit-8 (CCK-8). Cells were seeded into 96-well plates (1×103/well). In all, 20 μL CCK-8 (DOJINDO, CK04) was diluted to 10% with culture medium. After culturing for 2 hours in the incubator, the absorbance was measured at a wavelength of 450 nm. For the colony formation assay, cells were seeded into 6-well plates (2×103/well). After the colonies were identifiable, 4% paraformaldehyde (PFA; VETEC) and 0.1% crystal violet (Sigma-Aldrich) were used for fixation and staining, respectively.

Statistical analysis

GraphPad Prism version 8.0 (GraphPad Software) was used for performing statistical analysis. The significance of quantitative data was assessed by Student t test and one-way ANOVA. Pearson correlation analysis was used to assess the correlation between two parameters. The Kaplan‒Meier method with the log-rank test was performed to compare the overall survival (OS) and recurrence-free survival (RFS) rates. Every representative experiment was performed in triplicate. P < 0.05 was considered to be statistically significant.

Data availability statement

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. The transcriptome cases of HCC and nontumor tissues in this study were obtained from TCGA datasets (https://portal.gdc.cancer.gov).

PTTG1 is upregulated and associated with a poor prognosis in HCC

First, the differential expression of PTTG1 in HCC and nontumor tissues was revealed. Using the TCGA–LIHC dataset, PTTG1 was found to be significantly highly expressed in HCC (Fig. 1A). In 50 pairs of HCC and nontumor liver tissues, PTTG1 levels were elevated in HCC tissues (Fig. 1B and C). PTTG1 expression was correlated with advanced pathologic stage (Fig. 1D). According to the median PTTG1 expression, the TCGA–LIHC cohort was divided into two groups: the PTTG1 high (n = 187) and low-expression (n = 187) groups. Gene Set Enrichment Analysis indicated that the PTTG1 high-expression group was markedly enriched in the liver cancer proliferation pathway (normalized enrichment score; NES = 2.359, P < 0.05; Fig. 1E). The diagnostic efficiency of PTTG1 for HCC was 0.973 (Fig. 1F). Kaplan‒Meier survival curves revealed that high PTTG1 expression was associated with poor OS and RFS (Fig. 1G and H). We also collected 12 pairs of samples (HCC and corresponding nontumor tissues). The protein level of PTTG1 was further analyzed in the 12 paired samples. The results showed that PTTG1 protein was highly expressed in all HCC tissues (12 of 12) compared with corresponding nontumor tissues (Fig. 1I and J). To strengthen the illustrated expression differences and impact on prognosis of PTTG1, IHC staining and survival analysis were further conducted in the HCC TMA. A higher level of PTTG1 was found in HCC tumor tissues than in nontumor tissues (Fig. 1K; Supplementary Fig. S1A). High PTTG1 expression was remarkably associated with poor OS and RFS in patients with HCC, consistent with previous results (Fig. 1L and M). Moreover, the univariate and multivariate Cox proportional hazards analyses indicated that a high PTTG1 level was an independent prognostic factor for HCC patient prognosis (Supplementary Table S1; Supplementary Fig. S1B and S1C). Considering that HBV infection was still the most dominant risk factor for HCC, we further evaluated PTTG1 expression in HBV-related patients with HCC. The results showed that PTTG1 levels were also elevated in HBV related-HCC tissues, although the expression of PTTG1 showed almost no difference between HBV-related HCC and non–HBV-related HCC (Supplementary Fig. S1D and S1E). In addition, high PTTG1 expression was associated with poor survival, liver cancer cell proliferation in HBV-related patients with HCC (Supplementary Fig. S1F and S1I). In summary, PTTG1 was shown to be obviously upregulated in HCC, and upregulation of PTTG1 was predictive of a poor prognosis and advanced-stage disease in patients with HCC.

Figure 1.

PTTG1 is significantly upregulated in human HCC. A, Expression of PTTG1 in HCC and nontumor tissues in the TCGA dataset. B, Expression of PTTG1 in HCC tissues and paired nontumor tissues in the TCGA dataset. C, Expression of PTTG1 in tumor/nontumor tissues in the TCGA dataset. D, PTTG1 expression according to the pathologic stage of HCC in the TCGA dataset. E, Gene set enrichment analysis showed that high PTTG1 expression was associated with liver cancer proliferation in the TCGA dataset (NES = 2.359; P < 0.05). F, ROC analysis of the HCC diagnostic value of PTTG1 based on the TCGA dataset (AUC = 0.973). G and H, High PTTG1 levels predict worse OS and RFS based on the TCGA dataset. I and J, Western blot analysis and densitometry analysis of PTTG1 expression in HCC tissues and nontumor tissues (n = 12). K, Representative images of TMA staining for PTTG1 in HCC tissues and nontumor tissues. L and M, High PTTG1 levels predict worse OS (L) and RFS (M) on the basis of the TMA staining scores. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001; ns, nonsignificant.

Figure 1.

PTTG1 is significantly upregulated in human HCC. A, Expression of PTTG1 in HCC and nontumor tissues in the TCGA dataset. B, Expression of PTTG1 in HCC tissues and paired nontumor tissues in the TCGA dataset. C, Expression of PTTG1 in tumor/nontumor tissues in the TCGA dataset. D, PTTG1 expression according to the pathologic stage of HCC in the TCGA dataset. E, Gene set enrichment analysis showed that high PTTG1 expression was associated with liver cancer proliferation in the TCGA dataset (NES = 2.359; P < 0.05). F, ROC analysis of the HCC diagnostic value of PTTG1 based on the TCGA dataset (AUC = 0.973). G and H, High PTTG1 levels predict worse OS and RFS based on the TCGA dataset. I and J, Western blot analysis and densitometry analysis of PTTG1 expression in HCC tissues and nontumor tissues (n = 12). K, Representative images of TMA staining for PTTG1 in HCC tissues and nontumor tissues. L and M, High PTTG1 levels predict worse OS (L) and RFS (M) on the basis of the TMA staining scores. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001; ns, nonsignificant.

Close modal

PTTG1 aggravates hepatocellular carcinogenesis and proliferation of liver cancer cells

To further evaluate the role of PTTG1 in hepatocellular carcinogenesis, a PTTG1 knockout (KO; PTTG1−/−) HCC mouse model was established. WT and PTTG1−/− mice were treated with DEN (15 mg/kg) at 2 weeks and sacrificed at 9 months (Fig. 2A). Strikingly, PTTG1−/− mice (11%, 1/9) were protected against hepatocarcinogenesis, as a nearly complete reduction in liver tumors induced by DEN was observed in these mice compared with WT mice (89%, 8/9; Fig. 2B and C). We also found that tumor numbers and maximum tumor size were significantly reduced in PTTG1−/− mice compared with WT mice, even though there was no difference in the liver weight percentage (Fig. 2C). Tumor samples were confirmed by histopathologic analysis. IHC staining for proliferating cell nuclear antigen (PCNA), a proliferation marker, was markedly decreased in PTTG1−/− mice (Fig. 2D; Supplementary Fig. S2A). Then, we further observed the roles of PTTG1 in regulating human liver cancer cell proliferation. To obtain cell lines with stable knockdown and overexpression of PTTG1, HepG2, and Hep3B cells were infected with the shNC/shPTTG1 and LvNC/LvPTTG1 lentiviruses, respectively. We found that the efficient knockdown or overexpression of PTTG1 can accordingly alter the expression of PCNA (Fig. 2E and F; Supplementary Fig. S2B and S2C). Similarly, the results of the CCK-8 assay and colony formation assays demonstrated that with PTTG1 knockdown, cell proliferation, and the number of colonies formed were markedly decreased, while overexpression of PTTG1 promoted liver cancer cells proliferation and increased the number of colonies formed (Fig. 2GJ; Supplementary Fig. S2D and S2E). In addition, performing TUNEL staining and Western blot analysis of cleaved caspase-3, we found PTTG1 deficiency induces apoptosis in vivo (Supplementary Fig. S2F and S2G). The cleaved caspase-3 expression was also higher in shPTTG1 compared with shNC liver cancer cells (Supplementary Fig. S2H and S2I). These results demonstrated that PTTG1 deficiency apparently restrains chemically induced hepatocarcinogenesis by down regulating liver cancer cell proliferation.

Figure 2.

PTTG1 aggravates hepatocellular carcinogenesis and proliferation of liver cancer cells. A, WT (n = 9) and PTTG1−/− (n = 9) mice were treated with DEN (15 mg/kg) at 2 weeks and were then sacrificed at 9 months. B, Liver photographs of WT and PTTG1−/− mice at 9 months. Scale bars, 5 mm. C, The tumor incidence in WT and PTTG1−/− mice at 9 months. Quantification of the percentage of liver weight/body weight. Quantification of the numbers of liver tumors in WT and PTTG1−/− mice at 9 months. Quantification of the maximal tumor size (diameter) in WT and PTTG1−/− mice at 9 months. D, Representative hematoxylin and eosin (H&E) and PCNA staining of liver tissues from WT and PTTG1−/− mice at 9 months. T, tumor. Scale bars, 100 μm. E and F, Western blot analysis of the effect of lentiviral infection and PCNA expression with PTTG1 knockdown and overexpression. G and H, Effect of PTTG1 knockdown and overexpression on HepG2/Hep3B cell growth by CCK-8 assay. I and J, Effect of PTTG1 knockdown and overexpression on HepG2/Hep3B cell proliferation by colony formation (n = 3). All values are the mean ± SD. **, P < 0.01; ns, no significance.

Figure 2.

PTTG1 aggravates hepatocellular carcinogenesis and proliferation of liver cancer cells. A, WT (n = 9) and PTTG1−/− (n = 9) mice were treated with DEN (15 mg/kg) at 2 weeks and were then sacrificed at 9 months. B, Liver photographs of WT and PTTG1−/− mice at 9 months. Scale bars, 5 mm. C, The tumor incidence in WT and PTTG1−/− mice at 9 months. Quantification of the percentage of liver weight/body weight. Quantification of the numbers of liver tumors in WT and PTTG1−/− mice at 9 months. Quantification of the maximal tumor size (diameter) in WT and PTTG1−/− mice at 9 months. D, Representative hematoxylin and eosin (H&E) and PCNA staining of liver tissues from WT and PTTG1−/− mice at 9 months. T, tumor. Scale bars, 100 μm. E and F, Western blot analysis of the effect of lentiviral infection and PCNA expression with PTTG1 knockdown and overexpression. G and H, Effect of PTTG1 knockdown and overexpression on HepG2/Hep3B cell growth by CCK-8 assay. I and J, Effect of PTTG1 knockdown and overexpression on HepG2/Hep3B cell proliferation by colony formation (n = 3). All values are the mean ± SD. **, P < 0.01; ns, no significance.

Close modal

PTTG1 deficiency significantly inhibits the oncogenic effect of HBx

Currently, HBV infection is one of the most important causative factors of hepatocarcinogenesis worldwide, and HBx, as one of the four coding frames of HBV, plays an important role in the carcinogenesis of HBV (8, 38, 39). To determine whether PTTG1 is critical for the HBx-mediated oncogenic effect, we first generated mice with HBx expression and PTTG1 knockout (HBx+/+ PTTG1−/−) by crossing HBx-TG (HBx+/+ PTTG1−/−) mice and single PTTG1 knockout (PTTG1−/−) mice. Male HBx+/+ and HBx+/+PTTG1−/− mice were treated with a single i.p. injection of low-dose DEN (5 mg/kg) at 2 weeks of age to accelerate the development of HBx-induced tumors. At 9 months, the mice were sacrificed (Fig. 3A). Surprisingly, we found that compared with HBx+/+ mice, HBx+/+ PTTG1−/− mice exhibited a lower incidence of tumors (HBx+/+ 100% 6/6 vs. HBx+/+ PTTG1−/− 33.3% 2/6), lower liver/body weights, fewer tumors, and smaller tumor sizes (Fig. 3B and C). Tumor samples were confirmed by histopathologic analysis. IHC staining of PCNA indicated less malignant cell proliferation in HBx+/+ PTTG1−/− mice than in HBx+/+ mice (Fig. 3D; Supplementary Fig. S3A). In addition, to further verify the role of PTTG1 in the oncogenic effect of HBx, a spontaneous HCC mouse model was established. HBx+/+ and HBx+/+ PTTG1−/− mice were raised for 18 months before being sacrificed (Fig. 3E). Similarly, HBx+/+ PTTG1−/− mice presented a lower incidence of tumors (HBx+/+ 87.5% 7/8 vs. HBx+/+ PTTG1−/− 33.3% 3/9), fewer tumors and smaller tumors (Fig. 3F and G). However, there was no difference in the liver weight percentage in the two types of mice (Fig. 3G). PCNA IHC staining also revealed a significantly lower level of tumor proliferation in HBx+/+ PTTG1−/− mice (Fig. 3H; Supplementary Fig. S3B). The results showed that hepatocarcinogenesis was markedly repressed in HBx+/+ PTTG1−/− mice.

Figure 3.

PTTG1 deficiency restrains HBx-induced hepatocellular carcinogenesis A,HBx+/+ (n = 6) and HBx+/+ PTTG1−/− (n = 6) mice were treated with DEN (5 mg/kg) at 2 weeks and were then sacrificed at 9 months. B, Liver photographs of HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Scale bars, 5 mm. C, The tumor incidence in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Quantification of the percentage of liver weight/body weight. Quantification of the numbers of liver tumors in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Quantification of the maximal tumor size (diameter) in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. D, Representative hematoxylin and eosin (H&E) and PCNA staining of liver tissues from HBx+/+ PTTG1−/− mice at 9 months. T, tumor. Scale bars, 100 μm. E,HBx+/+ (n = 8) and HBx+/+ PTTG1−/− (n = 9) mice were sacrificed at 18 months. F, Liver photographs of HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. Scale bars, 5 mm. G, The tumor incidence in HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. Quantification of the percentage of liver weight/body weight. Quantification of the number of liver tumors in HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. H, Representative hematoxylin and eosin and PCNA staining of liver tissues from HBx+/+ PTTG1−/− mice at 18 months. T, tumor. Scale bars, 100 μm. I, and J, Western blot analysis of PCNA expression in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids as indicated. Effect of transfection with vector, HBx siRNA, and/or PTTG1 plasmids on HepG2.2.15/Hep3B cell proliferation by CCK-8 assay. K and L, Effect of cell growth in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids by colony formation (n = 3). All values are the mean ± SD. **, P < 0.01; ***, P < 0.01; ns, no significance.

Figure 3.

PTTG1 deficiency restrains HBx-induced hepatocellular carcinogenesis A,HBx+/+ (n = 6) and HBx+/+ PTTG1−/− (n = 6) mice were treated with DEN (5 mg/kg) at 2 weeks and were then sacrificed at 9 months. B, Liver photographs of HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Scale bars, 5 mm. C, The tumor incidence in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Quantification of the percentage of liver weight/body weight. Quantification of the numbers of liver tumors in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. Quantification of the maximal tumor size (diameter) in HBx+/+ and HBx+/+ PTTG1−/− mice at 9 months. D, Representative hematoxylin and eosin (H&E) and PCNA staining of liver tissues from HBx+/+ PTTG1−/− mice at 9 months. T, tumor. Scale bars, 100 μm. E,HBx+/+ (n = 8) and HBx+/+ PTTG1−/− (n = 9) mice were sacrificed at 18 months. F, Liver photographs of HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. Scale bars, 5 mm. G, The tumor incidence in HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. Quantification of the percentage of liver weight/body weight. Quantification of the number of liver tumors in HBx+/+ and HBx+/+ PTTG1−/− mice at 18 months. H, Representative hematoxylin and eosin and PCNA staining of liver tissues from HBx+/+ PTTG1−/− mice at 18 months. T, tumor. Scale bars, 100 μm. I, and J, Western blot analysis of PCNA expression in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids as indicated. Effect of transfection with vector, HBx siRNA, and/or PTTG1 plasmids on HepG2.2.15/Hep3B cell proliferation by CCK-8 assay. K and L, Effect of cell growth in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids by colony formation (n = 3). All values are the mean ± SD. **, P < 0.01; ***, P < 0.01; ns, no significance.

Close modal

Next, we further explored the role of PTTG1 in influencing the carcinogenic effect of HBx in vitro. We cotransfected siHBx and PTTG1 plasmids into HepG2.2.15 and Hep3B cells. We found that the inhibitory effect on PCNA protein levels, cell proliferation and colony formation induced by siHBx could be reversed by PTTG1 overexpression (Fig. 3IL; Supplementary Fig. S3C–S3F). These results suggest that PTTG1 is an important target for HBx in promoting tumorigenesis.

HBx promotes the expression and nuclear localization of PTTG1

We next explored the relationship between HBx and PTTG1. First, PTTG1 expression was found to be significantly higher in two HBx-expressing cell lines, HepG2.2.15 and Hep3B, but lower in HepG2 and Huh7 cells, which do not express HBx (Fig. 4A). In addition, we stimulated the HepG2–NTCP cell lines with normal human serum and hepatitis B patient serum. As a result, the expression of PTTG1 was remarkably higher after stimulation with hepatitis B patient serum (Fig. 4B). Furthermore, the HBV genomic sequence combined with HBx and HBx-null plasmids were transfected into HepG2 and Huh7 cells and PTTG1 was overexpressed only when HBx was expressed (Fig. 4C). To further verify the relationship between HBx and PTTG1, we transfected HBx-HA plasmids into HepG2 and Huh7 cells and found that efficient transfection increased the level of PTTG1 (Fig. 4D; Supplementary Fig. S4A). Accordingly, when HBx was effectively knocked down in HepG2.2.15 and Hep3B cells, the PTTG1 expression level was reduced (Fig. 4E; Supplementary Fig. S4B). In addition, the immunofluorescence (IF) results of PTTG1 in HepG2 and Hep2.2.15 cells showed that the presence of HBx notably upregulated the expression of PTTG1, especially in the nucleus (Fig. 4F and G). However, PTTG1 could not regulate the expression of HBx (Supplementary Fig. S4C and D).

Figure 4.

HBx promotes the expression and nuclear localization of PTTG1. A, Western blot analysis of PTTG1 protein levels in liver cancer cells. *, P < 0.05; **, P < 0.01 compared with the Huh7; #, P < 0.05 compared with the HepG2. B, Western blot analysis of PTTG1 protein levels in HepG2 cells with or without HBV infection. C, Western blot analysis of HBx and PTTG1 protein levels in cells transfected with the indicated plasmids. D, Western blot analysis of PTTG1 protein levels in HepG2 cells transfected with HBx–HA plasmids. E, Western blot analysis of PTTG1 protein levels in HepG2.2.15 cells transfected with HBx siRNA. F and G, Double IF staining of PTTG1 and HBx in transfected control and HBx–HA plasmid HepG2 cells and HepG2.2.15 cells transfected with control and HBx siRNA. 4',6-diamidino-2-phenylindole (DAPI) was used to stain nuclei. Scale bar, 10 μm. H, Western blot analysis of PTTG1 protein levels in the HepG2 cell cytosolic and nuclear fractions transfected with the indicated plasmids. I, Co-IP between exogenous HBx and PTTG1 in HepG2 cells transfected with PTTG1-Flag and HBx–HA plasmids. J, 3D structures of PTTG1/HBx. K, Co-IP between exogenous HBx and PTTG1 mutant in HepG2 cells transfected with PTTG1mutant- Flag and HBx–HA plasmids. L, Western blot analysis of HBx–HA and PTTG1 protein levels in cells transfected with the indicated plasmids. M, Western blot analysis of PTTG1 protein levels in the HepG2 cells cytosolic and nuclear fractions transfected with the indicated plasmids. N, Double IF staining for HBx/PTTG1 in 18 m WT and HBx+/+ mice. DAPI was used to stain nuclei. Scale bar, 80 μm. All values are the mean ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Figure 4.

HBx promotes the expression and nuclear localization of PTTG1. A, Western blot analysis of PTTG1 protein levels in liver cancer cells. *, P < 0.05; **, P < 0.01 compared with the Huh7; #, P < 0.05 compared with the HepG2. B, Western blot analysis of PTTG1 protein levels in HepG2 cells with or without HBV infection. C, Western blot analysis of HBx and PTTG1 protein levels in cells transfected with the indicated plasmids. D, Western blot analysis of PTTG1 protein levels in HepG2 cells transfected with HBx–HA plasmids. E, Western blot analysis of PTTG1 protein levels in HepG2.2.15 cells transfected with HBx siRNA. F and G, Double IF staining of PTTG1 and HBx in transfected control and HBx–HA plasmid HepG2 cells and HepG2.2.15 cells transfected with control and HBx siRNA. 4',6-diamidino-2-phenylindole (DAPI) was used to stain nuclei. Scale bar, 10 μm. H, Western blot analysis of PTTG1 protein levels in the HepG2 cell cytosolic and nuclear fractions transfected with the indicated plasmids. I, Co-IP between exogenous HBx and PTTG1 in HepG2 cells transfected with PTTG1-Flag and HBx–HA plasmids. J, 3D structures of PTTG1/HBx. K, Co-IP between exogenous HBx and PTTG1 mutant in HepG2 cells transfected with PTTG1mutant- Flag and HBx–HA plasmids. L, Western blot analysis of HBx–HA and PTTG1 protein levels in cells transfected with the indicated plasmids. M, Western blot analysis of PTTG1 protein levels in the HepG2 cells cytosolic and nuclear fractions transfected with the indicated plasmids. N, Double IF staining for HBx/PTTG1 in 18 m WT and HBx+/+ mice. DAPI was used to stain nuclei. Scale bar, 80 μm. All values are the mean ± SD. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Close modal

To further investigate the differential PTTG1 expression in cell structures under the promotion of HBx, we performed nucleus and cytoplasmic separation experiments in HepG2 cells. The results showed that PTTG1 increased more in the nucleus than in the cytoplasm when HBx was present (Fig. 4H; Supplementary Fig. S4E). Then, we found that PTTG1 and HBx proteins could bind to each other on the basis of co-IP experiments (Fig. 4I). A three-dimensional (3D) structural model of PTTG1 and HBx was generated. Docking simulation data demonstrated that AAs Glu-132, Leu130, HID134, and Ser142 of PTTG1, as well as AAs Cys143, His139, Ser64, and Phe151 of HBx, are responsible for PTTG1/HBx interactions (Fig. 4J). After randomly mutating those AAs in PTTG1, there was no interaction between mutated PTTG1 and HBx (Fig. 4K). Furthermore, after mutating those AAs in HBx, HBx could not promote PTTG1 expression and facilitate PTTG1 nuclear localization (Fig. 4L and M; Supplementary Fig. S4F and S4G). Next, HBx+/+ mice were used to further verify the relationship between HBx and PTTG1. Consistent with the results of liver cancer cells, the results of Western blot analysis and IHC/IF staining indicated that the expression of PTTG1 was apparently higher in HBx+/+ mice at 6 or 18 months than in WT mice (Fig. 4N; Supplementary Fig. S4H–S4K). In general, we revealed that HBx could promote PTTG1 expression and facilitate PTTG1 nuclear localization.

PTTG1 upregulates Asn metabolism by promoting ASNS transcription in HCC

To investigate the potential downstream pathway of PTTG1, we performed differential analysis in both the high and low PTTG1 expression groups in the TCGA–LIHC dataset and obtained 3,619 genes that were highly expressed in the high-expression group. These genes were then subjected to Gene Ontology- biological process (GO-BP) enrichment analysis and identified as significantly associated with AA metabolism (Fig. 5A). The untargeted metabolomics analysis showed clear differences in AAs between shNC and shPTTG1 in HepG2.2.15 cells (Supplementary Fig. S5A). Therefore, we next focused on AA metabolism. The results of enrichment analysis of AA metabolism–related pathways revealed that highly expressed PTTG1 could be significantly enriched in the Asn metabolism pathway (Fig. 5B; Supplementary Fig. S5B). Asn metabolism is the process by which ASNS catalyzes the formation of Asn as well as glutamate from Gln and aspartate (Fig. 5C). Then, from the TCGA dataset, we found a considerably positive correlation between PTTG1 and ASNS (Fig. 5D).

Figure 5.

PTTG1 positively regulates Asn metabolism through ASNS in HCC. A, The top 10 GO-BP enrichment results of metabolism-related pathways according to genes that were significantly elevated in the PTTG1 high-expression group. B, The top 10 Gene Set Variation Analysis results for AA metabolic signaling pathways according to the different PTTG1 mRNA expression levels in TCGA–LIHC. C, Schematic diagram describing Asn metabolism catalyzed by ASNS. D, The correlation between PTTG1 and ASNS in the TCGA dataset. E, Western blot analysis of ASNS protein levels in liver tissues from DEN-induced WT and PTTG1−/− mice at 9 months. n = 3 in each group. F, ASNS staining of liver tissues from DEN-induced WT and PTTG1−/− mice at 9 months. Scale bars, 100 μm. G, Intracellular amount of asparagine in shNC and shPTTG1 HepG2.215/Hep3B cells. H, Double immunofluorescent staining of ASNS and PTTG1 in shNC and shPTTG1 HepG2.215/Hep3B cells. DAPI was used to stain nuclei. Scale bars, 10 μm. I and J, qRT-PCR (I) and Western blot analysis (J) of ASNS in shNC and shPTTG1 HepG2.215/Hep3B cells. K, Luciferase reporter assay of ASNS promoter activity in HepG2.215/Hep3B cells transfected with PTTG1 plasmid. L and M, ChIP-qPCR assays (L) and agarose gel electrophoresis (M) were performed to analyze the interaction between PTTG1 and the ASNS promoter. N, Schematic diagram. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001. Glu, glutamate.

Figure 5.

PTTG1 positively regulates Asn metabolism through ASNS in HCC. A, The top 10 GO-BP enrichment results of metabolism-related pathways according to genes that were significantly elevated in the PTTG1 high-expression group. B, The top 10 Gene Set Variation Analysis results for AA metabolic signaling pathways according to the different PTTG1 mRNA expression levels in TCGA–LIHC. C, Schematic diagram describing Asn metabolism catalyzed by ASNS. D, The correlation between PTTG1 and ASNS in the TCGA dataset. E, Western blot analysis of ASNS protein levels in liver tissues from DEN-induced WT and PTTG1−/− mice at 9 months. n = 3 in each group. F, ASNS staining of liver tissues from DEN-induced WT and PTTG1−/− mice at 9 months. Scale bars, 100 μm. G, Intracellular amount of asparagine in shNC and shPTTG1 HepG2.215/Hep3B cells. H, Double immunofluorescent staining of ASNS and PTTG1 in shNC and shPTTG1 HepG2.215/Hep3B cells. DAPI was used to stain nuclei. Scale bars, 10 μm. I and J, qRT-PCR (I) and Western blot analysis (J) of ASNS in shNC and shPTTG1 HepG2.215/Hep3B cells. K, Luciferase reporter assay of ASNS promoter activity in HepG2.215/Hep3B cells transfected with PTTG1 plasmid. L and M, ChIP-qPCR assays (L) and agarose gel electrophoresis (M) were performed to analyze the interaction between PTTG1 and the ASNS promoter. N, Schematic diagram. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001. Glu, glutamate.

Close modal

Subsequently, we validated the relationship between PTTG1 and ASNS in DEN-treated WT and PTTG1−/− mice at 9 months. Western blot analysis and IHC staining of ASNS indicated that ASNS expression was markedly reduced in the PTTG1−/− group (Fig. 5E and F; Supplementary Fig. S5C). Asn levels were also reduced in PTTG1-knockdown cells (Fig. 5G). To further confirm the relationship between PTTG1 and ASNS in HCC, ASNS mRNA and protein levels were compared between shNC and shPTTG1 in HepG2.2.15 and Hep3B cells. Consistent with our expectations, IF staining of PTTG1 demonstrated that PTTG1-knockdown cells exhibited downregulated ASNS (Fig. 5H). qRT-PCR and Western blotting experiments confirmed the low expression of ASNS in PTTG1-knockdown HepG2.2.15/Hep3B cell lines (Fig. 5I and J; Supplementary Fig. S5D and S5E). Then, we further determined how PTTG1 affects the expression of ASNS. Luciferase assays revealed that PTTG1 overexpression promoted ASNS transactivation (Fig. 5K). Moreover, chromatin IP (ChIP) assays further revealed that PTTG1 could bind to the ASNS promoter (Fig. 5L and M). These findings indicated that PTTG1 could promote the transcription of ASNS by binding to the promoter of ASNS and enhancing Asn metabolism (Fig. 5N).

HBx upregulates ASNS-mediated Asn metabolism via PTTG1 in HCC

We previously revealed that PTTG1 could bind to the ASNS promoter to accelerate ASNS transcription and subsequently upregulate Asn metabolism. Next, we further investigated whether HBx could regulate this process. High ASNS expression was detected in the liver tissues of 6/18-month-old HBx+/+ mice compared with those of WT mice. As expected, ASNS was clearly reduced by PTTG1 deletion in both 6- and 18-month-old mice (Fig. 6AF). In addition, 9-month-old mice treated with DEN showed that ASNS protein levels were also decreased in HBx+/+PTTG1−/− mice tissues compared with HBx+/+ mice tissues (Fig. 6GJ). To further explore the relationship between HBx, PTTG1, and ASNS, we transfected the HBx–HA plasmid accompanied by shNC/shPTTG1 into HepG2 and Huh7 cells. In addition, HBx siRNA and PTTG1 overexpression plasmids were cotransfected into HepG2.2.15 and Hep3B cells. As anticipated, the Western blotting analysis results indicated that HBx could participate in upregulating ASNS-mediated Asn metabolism through PTTG1 (Fig. 6K and L; Supplementary Fig. S6A–S6D).

Figure 6.

HBx promotes ASNS-mediated Asn metabolism through PTTG1 in HCC. A, C, and D, ASNS staining of liver tissues from WT, HBx+/+, and HBx+/+PTTG1−/− mice at 6 and 18 months. n = 3 in each group, Scale bars, 100 μm. B, E, and F, Western blot analysis of ASNS protein levels in liver tissues from WT, HBx+/+, and HBx+/+PTTG1−/− mice at 6 and 18 months. n = 3 in each group. G and H, ASNS staining of liver tissues from DEN-induced HBx+/+ and HBx+/+PTTG1−/− mice at 9 months. n = 3 in each group, Scale bars, 100 μm. I and J, Western blot analysis of ASNS protein levels in liver tissues from HBx+/+ and HBx+/+PTTG1−/− mice at 9 months. n = 3 in each group. K, Western blot analysis of ASNS expression in shNC or shPTTG1 HepG2 and Huh7 cells transfected with vector or HBx–HA plasmids. L, Western blot analysis of ASNS expression in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids as indicated. All values are the mean ± SD. **, P < 0.01; ***, P < 0.01.

Figure 6.

HBx promotes ASNS-mediated Asn metabolism through PTTG1 in HCC. A, C, and D, ASNS staining of liver tissues from WT, HBx+/+, and HBx+/+PTTG1−/− mice at 6 and 18 months. n = 3 in each group, Scale bars, 100 μm. B, E, and F, Western blot analysis of ASNS protein levels in liver tissues from WT, HBx+/+, and HBx+/+PTTG1−/− mice at 6 and 18 months. n = 3 in each group. G and H, ASNS staining of liver tissues from DEN-induced HBx+/+ and HBx+/+PTTG1−/− mice at 9 months. n = 3 in each group, Scale bars, 100 μm. I and J, Western blot analysis of ASNS protein levels in liver tissues from HBx+/+ and HBx+/+PTTG1−/− mice at 9 months. n = 3 in each group. K, Western blot analysis of ASNS expression in shNC or shPTTG1 HepG2 and Huh7 cells transfected with vector or HBx–HA plasmids. L, Western blot analysis of ASNS expression in HepG2.2.15 and Hep3B cells transfected with vector, HBx siRNA, and/or PTTG1 plasmids as indicated. All values are the mean ± SD. **, P < 0.01; ***, P < 0.01.

Close modal

PTTG1 promotes HCC proliferation through ASNS-mediated Asn metabolism

Through Western blotting and IHC staining, ASNS expression was upregulated in HCC tissues (Fig. 7A and B). Using the TCGA–LIHC dataset, ASNS was found to be highly expressed in HCC (Fig. 7C). In 50 pairs of HCC tissues and nontumor tissues, ASNS was markedly upregulated in tumor tissues (Supplementary Fig. S7A and S7B). ASNS expression was correlated with advanced pathologic stage (Supplementary Fig. S7C). However, the expression of ASNS was not different between HBV-related HCC samples and non–HBV-related HCC samples (Supplementary Fig. S7D). In addition, ASNS could provide potential assistance for the diagnosis of HCC (Supplementary Fig. S7E). Survival analysis showed that high ASNS expression was associated with poorer prognosis in patients with HCC (Fig. 7C). To further investigate the role of ASNS in the oncogenic effect of PTTG1, PTTG1 was knocked down in HepG2.2.15 and Hep3B cells overexpressing ASNS. From the results of PCNA expression and CCK-8 assays, the inhibitory effect of shPTTG1 on proliferation could be reversed by overexpression of ASNS (Fig. 7D and E; Supplementary Fig. S7F and S7G). Because Asn is the product of Asn synthetase (11, 18, 40), we determined whether Asn could mediate the effect of PTTG1 on cell proliferation. Fortunately, similar to ASNS, Asn could eliminate the inhibitory effect of shPTTG1 on the proliferation of liver cancer cells, according to the results of Western blot analysis and CCK-8 assays (Fig. 7F and G; Supplementary Fig. S7H and S7I). Asparaginase, a drug targeting ASNS, has been applied to treat many tumors (11, 17, 41, 42). However, there are few studies on the use of asparaginase to treat HCC. Next, we found that asparaginase could inhibit the proliferation of HepG2.2.15 and Hep3B cells and could inhibit the proliferative effect of overexpressing PTTG1 on liver cancer cells (Fig. 7H and I; Supplementary Fig. S7J and S7K). In summary, PTTG1 could promote the proliferation of liver cancer cells through Asn metabolism mediated by ASNS. Asparaginase might be a potential treatment for HCC.

Figure 7.

PTTG1 exacerbates the proliferation of liver cancer cells through ASNS-mediated Asn metabolism. A, Western blot analysis and densitometry analysis of ASNS expression in HCC liver tissues and nontumor tissues. n = 3 in each group. B, ASNS staining of liver tissues from HCC tissues and nontumor tissues. n = 3 in each group. Scale bars, 100 μm. C, ASNS level of HCC in the TCGA dataset. High ASNS levels predict worse OS based on the TCGA dataset. D and E, Western blot analysis of PCNA expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids as indicated. Effect of cell growth in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids by CCK-8 assay. F and G, Western blot analysis of PCNA expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells treated with control and Asn as indicated. Effect of cell growth in shNC and shPTTG1 HepG2.2.15/Hep3B cells treated with control and Asn by CCK-8 assay. H and I, Western blot analysis of PCNA expression in LvNC and LvPTTG1 HepG2.2.15/Hep3B cells treated with control and asparaginase as indicated. Effect of cell growth in LvNC and LvPTTG1 HepG2.2.15/Hep3B cells treated with control and asparaginase by CCK-8 assay. All values are the mean ± SD. ***, P < 0.001.

Figure 7.

PTTG1 exacerbates the proliferation of liver cancer cells through ASNS-mediated Asn metabolism. A, Western blot analysis and densitometry analysis of ASNS expression in HCC liver tissues and nontumor tissues. n = 3 in each group. B, ASNS staining of liver tissues from HCC tissues and nontumor tissues. n = 3 in each group. Scale bars, 100 μm. C, ASNS level of HCC in the TCGA dataset. High ASNS levels predict worse OS based on the TCGA dataset. D and E, Western blot analysis of PCNA expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids as indicated. Effect of cell growth in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids by CCK-8 assay. F and G, Western blot analysis of PCNA expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells treated with control and Asn as indicated. Effect of cell growth in shNC and shPTTG1 HepG2.2.15/Hep3B cells treated with control and Asn by CCK-8 assay. H and I, Western blot analysis of PCNA expression in LvNC and LvPTTG1 HepG2.2.15/Hep3B cells treated with control and asparaginase as indicated. Effect of cell growth in LvNC and LvPTTG1 HepG2.2.15/Hep3B cells treated with control and asparaginase by CCK-8 assay. All values are the mean ± SD. ***, P < 0.001.

Close modal

ASNS-mediated Asn metabolism promotes the proliferative effects of liver cancer cells through activation of the mTOR pathway

mTOR is activated in many kinds of tumors and has an important role in regulating tumor proliferation and metabolism (43). Our study found that knockdown of PTTG1 inhibited mTOR, p-mTOR, pAKT, and downstream targets (p4EBP1, pRPS6, and pS6K) expression (Supplementary Fig. S8A–S8C). As an mTOR inhibitor, rapamycin could inhibit the proliferative effect of overexpressing PTTG1 on HepG2.2.15 and Hep3B (Supplementary Fig. S8D). ASNS restored the inhibition of mTOR and downstream targets (p4EBP1, pRPS6, and pS6K) by knockdown of PTTG1 (Fig. 8A and B; Supplementary Fig. S8E and S8F). In addition, ASNS also activated mTOR and downstream targets through Asn (Fig. 8C and D; Supplementary Fig. S8G and S8H). Given that Asn occupied important positions in activating the mTOR pathway, we next investigated whether the mTOR pathway could mediate the proliferative effect of Asn on liver cancer cells. Thus, we treated both HepG2.2.15 and Hep3B cells with Asn and rapamycin, an mTOR pathway inhibitor. On the basis CCK-8 assay, the proliferative effect of Asn on liver cancer cells could be largely eliminated by rapamycin (Fig. 8E and F). Together, the proliferative effect of ASNS-mediated Asn metabolism on liver cancer cells was promoted by activating the mTOR pathway.

Figure 8.

ASNS-mediated Asn metabolism facilitates proliferation via mTOR signaling. A and B, Western blot analysis of mTOR, p-mTOR, p-p70 S6K (Thr389/412), p-4E-BP1 (Thr37/46), and pRPS6 (Ser235) expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids as indicated. C and D, Western blot analysis of mTOR and p-mTOR, p-p70 S6K (Thr389/412), p-4E-BP1 (Thr37/46), and pRPS6 (Ser235) expression in HepG2.2.15/Hep3B cells transfected with ASNS siRNA and/or treated with Asn as indicated. E and F, Effect of cell growth of HepG2.2.15/Hep3B cells treated with vector, Asn (0.1 mm), and/or rapamycin (20 μmol/L) by CCK-8 assay. G, Schematic diagram. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001.

Figure 8.

ASNS-mediated Asn metabolism facilitates proliferation via mTOR signaling. A and B, Western blot analysis of mTOR, p-mTOR, p-p70 S6K (Thr389/412), p-4E-BP1 (Thr37/46), and pRPS6 (Ser235) expression in shNC and shPTTG1 HepG2.2.15/Hep3B cells transfected with vector or ASNS plasmids as indicated. C and D, Western blot analysis of mTOR and p-mTOR, p-p70 S6K (Thr389/412), p-4E-BP1 (Thr37/46), and pRPS6 (Ser235) expression in HepG2.2.15/Hep3B cells transfected with ASNS siRNA and/or treated with Asn as indicated. E and F, Effect of cell growth of HepG2.2.15/Hep3B cells treated with vector, Asn (0.1 mm), and/or rapamycin (20 μmol/L) by CCK-8 assay. G, Schematic diagram. All values are the mean ± SD. **, P < 0.01; ***, P < 0.001.

Close modal

In summary, HBx, as a crucial oncogenic factor, upregulated PTTG1 expression and promoted its nuclear localization. Furthermore, PTTG1 could bind to the promoter of ASNS, promote its transcription, and further increase Asn metabolism. The upregulated ASNS and Asn activated the mTOR pathway and promoted the proliferation of HCC (Fig. 8G).

In this study, we demonstrated that PTTG1 expression was significantly elevated in HCC. A DEN-induced PTTG1−/− HCC mouse model was established, and it was verified that PTTG1 deficiency could inhibit hepatocarcinogenesis. We also established DEN-induced and spontaneous HBx+/+PTTG1+/+ and HBx+/+PTTG1−/− HCC mouse models and demonstrated for the first time that PTTG1 deficiency significantly inhibited the development of HCC induced by HBx. In addition, we found that PTTG1, whose expression was upregulated and whose nuclear location was promoted by HBx, could facilitate ASNS transcription by binding to the ASNS promoter. ASNS acts as a key activator of Asn metabolism. Subsequently, the upregulation of ASNS and Asn activates the mTOR pathway to promote HCC.

PTTG1 is upregulated in pituitary, gliomas, thyroid, breast, and gastrointestinal tumors (44). PTTG1 plays an important role in various biological processes, such as cell replication, DNA damage repair, and metabolic regulation (22–27). In this study, we found that PTTG1 expression was significantly upregulated in HCC and that high levels of PTTG1 were associated with poorer prognosis in patients with HCC. As mentioned previously, PTTG1 might be significant for tumor growth. Our previous study showed that PTTG1 promoted the proliferation of liver cancer cells through c-myc (37). In this study, PTTG1 deficiency markedly inhibited the development of HCC was confirmed by innovatively constructing a DEN-induced PTTG1−/− HCC mouse model. At present, HBV infection remains the most important cause of HCC, and HBx, as one of the HBV coding frames, plays an important role in HBV carcinogenesis. We generated mice with HBx expression and PTTG1 knockout (HBx+/+ PTTG1−/−) by crossing HBx-TG (HBx+/+ PTTG1−/−) mice and single-PTTG1 knockout (PTTG1−/−) mice. The DEN-induced and spontaneous HCC mouse models in HBx+/+ PTTG1+/+ and HBx+/+ PTTG1−/− mice both showed that PTTG1 deletion significantly inhibited HBx-induced malignant progression of HCC. Our study showed that HBx could interact with PTTG1, upregulate PTTG1 expression and promote PTTG1 nuclear localization. Similarly, Meng reported that HBV replication inhibited miR-122 and caused an obvious increase in PBF levels. PBF could interact with PTTG1 and facilitate PTTG1 nuclear translocation (36). Molina-Jiménez and colleagues’ research has shown that HBx could interact with PTTG1 and Cul1, disrupting the interaction between PTTG1 and the Skp1–Cul1–F-box ubiquitin ligase complex (SCF), which suppresses PTTG1 ubiquitination and subsequent degradation (35). In our study, when the AAs of HBx responsible for the interaction with PTTG1 were mutated, the ability of HBx to promote PTTG1 nearly disappeared. It is consistent with Molina-Jiménez and colleagues’ research.

Recent studies indicated that metabolic changes could directly participate in tumor growth and metastasis (45, 46). By bioinformatics and untargeted metabolomics analysis, we found that PTTG1 could be notably enriched in the Asn metabolic pathway. Asn, a nonessential AA, has long been accepted as a metabolite to be targeted in cancer (47). Recent research has revealed that Asn promotes tumor growth by modulating survival, growth, and the mTOR pathway (16, 47–49). ASNS, which is widely expressed in mammalian cells, catalyzes the synthesis of Asn using aspartic acid and Gln as substrates (18). The level of ASNS expression is directly correlated with its ability to synthesize Asn by the cell (16). ASNS expression is upregulated in many tumors, such as colon cancer, lung cancer, and pancreatic cancer. Overexpression of ASNS promotes tumor proliferation, cell apoptosis, and the cell-cycle (20, 40, 50, 51). Our study revealed that Asn and ASNS expression were downregulated when PTTG1 was deficient. PTTG1 could act as a transcription factor-binding to the promoter of ASNS and upregulate the expression of ASNS. In our study, ASNS expression was considerably elevated in HCC tissues, and high ASNS expression was associated with poor prognosis in patients with HCC. We found that the inhibitory effect of PTTG1 knockdown on liver cancer cells could be reversed by overexpression of ASNS and Asn. Asparaginase acts as an inhibitor of ASNS. Recent studies have shown that asparaginase affects the level of reactive oxygen species (ROS), the cell-cycle, autophagy, and apoptotic cell death (52, 53). It also inhibits some important signaling-pathways in cell growth and survival, such as the Akt/mTOR and extracellular signal-regulated kinase (ERK) signaling pathways (54). The proliferation of breast cancer cells is remarkably inhibited via the cytotoxic effect of isolated asparaginase from microbial sources (55). However, the value of asparaginase in HCC needs to be further developed. We found that asparaginase could inhibit the proliferation of liver cancer cells. The proliferation-promoting effect of PTTG1 overexpression could also be largely inhibited by asparaginase. Human asparaginase might provide an effective anticancer enzyme for clinical therapy.

Overexpression of ASNS and increased Asn promote activation of the mTOR pathway, a key sensor of nutrition. In tumors, activated mTOR-signaling promotes tumor cell proliferation, metastasis, and drug resistance (43). To date, some mTOR inhibitors have been applied in clinical trials for the treatment of cancers, including HCC. Our results suggest that PTTG1 activates the mTOR pathway by upregulating ASNS-mediated Asn metabolism. In contrast, the mTOR inhibitor rapamycin inhibited PTTG1 and Asn-induced proliferation of liver cancer cells.

In conclusion, our study showed that PTTG1 deficiency significantly suppressed hepatocellular carcinogenesis. PTTG1 upregulated ASNS-mediated Asn metabolism, thereby activating the mTOR pathway and promoting hepatocarcinogenesis. PTTG1 is promising as a marker and therapeutic target for HCC.

No disclosures were reported.

Q. Zhou: Conceptualization, resources, data curation, software, formal analysis, validation, investigation, visualization, methodology, writing–original draft, project administration, writing–review and editing. L. Li: Conceptualization, resources, data curation, supervision, methodology, writing–original draft, writing–review and editing. F. Sha: Conceptualization, data curation, software, formal analysis, visualization, and methodology. Y. Lei: Resources, investigation, writing–original draft, project administration, writing–review and editing. X. Tian: Resources, formal analysis, investigation, visualization, and methodology. L. Chen: Conceptualization, resources, data curation, and software. Y. Chen: Conceptualization, visualization, and writing–original draft. H. Liu: Conceptualization, validation, and methodology. Y. Guo: Conceptualization, resources, formal analysis, supervision, funding acquisition, investigation, methodology, writing–original draft, project administration, writing–review and editing.

This work was supported by the Guangzhou Science and Technology Program key projects (201803010018) and the National Natural Science Foundation of China (82170604, H0308). The authors thank Professor D.Y. Yu at the Korea Research Institution of Bioscience and Biotechnology for providing the HBx-transgenic mice.

The publication costs of this article were defrayed in part by the payment of publication fees. Therefore, and solely to indicate this fact, this article is hereby marked “advertisement” in accordance with 18 USC section 1734.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

1.
Maluccio
M
,
Covey
A
.
Recent progress in understanding, diagnosing, and treating hepatocellular carcinoma
.
CA Cancer J Clin
2012
;
62
:
394
9
.
2.
Thomas
MB
,
Jaffe
D
,
Choti
MM
,
Belghiti
J
,
Curley
S
,
Fong
Y
, et al
.
Hepatocellular carcinoma: consensus recommendations of the National cancer institute clinical trials planning meeting
.
J Clin Oncol
2010
;
28
:
3994
4005
.
3.
Ren
TM
,
Ting
EWY
,
Hei
SW
,
Wei Tian
AN
,
Lit-Hsin
L
,
Chow
PK
, et al
.
Global epidemiology and genetics of hepatocellular carcinoma
.
Gastroenterology
2023
;
164
:
766
82
.
4.
Akinyemiju
T
,
Abera
S
,
Ahmed
M
,
Alam
N
,
Alemayohu
MA
,
Allen
C
, et al
.
The burden of primary liver cancer and underlying etiologies from 1990 to 2015 at the global, regional, and national level: results from the global burden of disease study 2015
.
JAMA Oncol
2017
;
3
:
1683
91
.
5.
Neuveut
C
,
Wei
Y
,
Buendia
MA
.
Mechanisms of HBV-related hepatocarcinogenesis
.
J Hepatol
2010
;
52
:
594
604
.
6.
Andrisani
OM
,
Barnabas
S
.
The transcriptional function of the hepatitis B virus X protein and its role in hepatocarcinogenesis (Review)
.
Int J Oncol
1999
;
15
:
373
9
.
7.
Bouchard
MJ
,
Schneider
RJ
.
The enigmatic X gene of hepatitis B virus
.
J Virol
2004
;
78
:
12725
34
.
8.
Tang
H
,
Oishi
N
,
Kaneko
S
,
Murakami
S
.
Molecular functions and biological roles of hepatitis B virus x protein
.
Cancer Sci
2006
;
97
:
977
83
.
9.
Boroughs
LK
,
DeBerardinis
RJ
.
Metabolic pathways promoting cancer cell survival and growth
.
Nat Cell Biol
2015
;
17
:
351
9
.
10.
Ward
PS
,
Thompson
CB
.
Metabolic reprogramming: a cancer hallmark even warburg did not anticipate
.
Cancer Cell
2012
;
21
:
297
308
.
11.
Tardito
S
,
Chiu
M
,
Uggeri
J
,
Zerbini
A
,
Da Ros
F
.,
Dall'Asta
V
, et al
.
L-Asparaginase and inhibitors of glutamine synthetase disclose glutamine addiction of β-catenin-mutated human hepatocellular carcinoma cells
.
Curr Cancer Drug Targets
2011
;
11
:
929
43
.
12.
Satriano
L
,
Lewinska
M
,
Rodrigues
PM
,
Banales
JM
,
Andersen
JB
.
Metabolic rearrangements in primary liver cancers: cause and consequences
.
Nat Rev Gastroenterol Hepatol
2019
;
16
:
748
66
.
13.
Huang
Q
,
Tan
Y
,
Yin
P
,
Ye
G
,
Gao
P
,
Lu
X
, et al
.
Metabolic characterization of hepatocellular carcinoma using nontargeted tissue metabolomics
.
Cancer Res
2013
;
73
:
4992
5002
.
14.
Liu
Z
,
Wang
J
,
Liu
L
,
Yuan
H
,
Bu
Y
,
Feng
J
, et al
.
Chronic ethanol consumption and HBV induce abnormal lipid metabolism through HBx/SWELL1/arachidonic acid signaling and activate tregs in HBV-Tg mice
.
Theranostics
2020
;
10
:
9249
67
.
15.
Balasubramanian
MN
,
Butterworth
EA
,
Kilberg
MS
.
Asparagine synthetase: regulation by cell stress and involvement in tumor biology
.
Am J Physiol Endocrinol Metab
2013
;
304
:
E789
99
.
16.
Krall
AS
,
Mullen
PJ
,
Surjono
F
,
Momcilovic
M
,
Schmid
EW
,
Halbrook
CJ
, et al
.
Asparagine couples mitochondrial respiration to ATF4 activity and tumor growth
.
Cell Metab
2021
;
33
:
1013
26
.
17.
Deng
L
,
Yao
P
,
Li
L
,
Ji
F
,
Zhao
S
,
Xu
C
, et al
.
p53-mediated control of aspartate-asparagine homeostasis dictates LKB1 activity and modulates cell survival
.
Nat Commun
2020
;
11
:
1755
.
18.
Richards
NG
,
Kilberg
MS
.
Asparagine synthetase chemotherapy
.
Annu Rev Biochem
2006
;
75
:
629
54
.
19.
Toda
K
,
Kawada
K
,
Iwamoto
M
,
Inamoto
S
,
Sasazuki
T
,
Shirasawa
S
, et al
.
Metabolic alterations caused by KRAS mutations in colorectal cancer contribute to cell adaptation to glutamine depletion by upregulation of asparagine synthetase
.
Neoplasia
2016
;
18
:
654
65
.
20.
Gwinn
DM
,
Lee
AG
,
Briones-Martin-Del-Campo
M
,
Conn
CS
,
Simpson
DR
,
Scott
AI
, et al
.
Oncogenic KRAS regulates amino acid homeostasis and asparagine biosynthesis via ATF4 and alters sensitivity to L-asparaginase
.
Cancer Cell
2018
;
33
:
91
107
.
21.
Kawedia
JD
,
Rytting
ME
.
Asparaginase in acute lymphoblastic leukemia
.
Clin Lymphoma Myeloma Leuk
2014
;
14 Suppl
:
S14
7
.
22.
Huang
JL
,
Cao
SW
,
Ou
QS
,
Yang
B
,
Zheng
SH
,
Tang
J
, et al
.
The long non-coding RNA PTTG3P promotes cell growth and metastasis via up-regulating PTTG1 and activating PI3K/AKT signaling in hepatocellular carcinoma
.
Mol Cancer
2018
;
17
:
93
.
23.
Puri
R
,
Tousson
A
,
Chen
L
,
Kakar
SS
.
Molecular cloning of pituitary tumor transforming gene 1 from ovarian tumors and its expression in tumors
.
Cancer Lett
2001
;
163
:
131
9
.
24.
Shibata
Y
,
Haruki
N
,
Kuwabara
Y
,
Nishiwaki
T
,
Kato
J
,
Shinoda
N
, et al
.
Expression of PTTG (pituitary tumor transforming gene) in esophageal cancer
.
Jpn J Clin Oncol
2002
;
32
:
233
7
.
25.
Chen
L
,
Puri
R
,
Lefkowitz
EJ
,
Kakar
SS
.
Identification of the human pituitary tumor transforming gene (hPTTG) family: molecular structure, expression, and chromosomal localization
.
Gene
2000
;
248
:
41
50
.
26.
Heaney
AP
,
Singson
R
,
McCabe
CJ
,
Nelson
V
,
Nakashima
M
,
Melmed
S
.
Expression of pituitary-tumour transforming gene in colorectal tumours
.
Lancet
2000
;
355
:
716
9
.
27.
Wen
CY
,
Nakayama
T
,
Wang
AP
,
Nakashima
M
,
Ding
YT
,
Ito
M
, et al
.
Expression of pituitary tumor transforming gene in human gastric carcinoma
.
World J Gastroenterol
2004
;
10
:
481
3
.
28.
Zou
H
,
McGarry
TJ
,
Bernal
T
,
Kirschner
MW
.
Identification of a vertebrate sister-chromatid separation inhibitor involved in transformation and tumorigenesis
.
Science
1999
;
285
:
418
22
.
29.
Chien
W
,
Pei
L
.
A novel binding factor facilitates nuclear translocation and transcriptional activation function of the pituitary tumor-transforming gene product
.
J Biol Chem
2000
;
275
:
19422
7
.
30.
McCabe
CJ
,
Boelaert
K
,
Tannahill
LA
,
Heaney
AP
,
Stratford
AL
,
Khaira
JS
, et al
.
Vascular endothelial growth factor, its receptor KDR/Flk-1, and pituitary tumor transforming gene in pituitary tumors
.
J Clin Endocrinol Metab
2002
;
87
:
4238
44
.
31.
Tong
Y
,
Tan
Y
,
Zhou
C
,
Melmed
S
.
Pituitary tumor transforming gene interacts with Sp1 to modulate G1/S cell phase transition
.
Oncogene
2007
;
26
:
5596
605
.
32.
Yu
R
,
Heaney
AP
,
Lu
W
,
Chen
J
,
Melmed
S
.
Pituitary tumor transforming gene causes aneuploidy and p53-dependent and p53-independent apoptosis
.
J Biol Chem
2000
;
275
:
36502
5
.
33.
Huang
S
,
Liao
Q
,
Li
W
,
Deng
G
,
Jia
M
,
Fang
Q
, et al
.
The lncRNA PTTG3P promotes the progression of CRPC via upregulating PTTG1
.
Bull Cancer
2021
;
108
:
359
68
.
34.
Hamid
T
,
Malik
MT
,
Kakar
SS
.
Ectopic expression of PTTG1/securin promotes tumorigenesis in human embryonic kidney cells
.
Mol Cancer
2005
;
4
:
3
.
35.
Molina-Jiménez
F
,
Benedicto
I
,
Murata
M
,
Martín-Vílchez
S
,
Seki
T
,
Antonio Pintor-Toro
J
, et al
.
Expression of pituitary tumor-transforming gene 1 (PTTG1)/securin in hepatitis B virus (HBV)-associated liver diseases: evidence for an HBV X protein-mediated inhibition of PTTG1 ubiquitination and degradation
.
Hepatology
2010
;
51
:
777
87
.
36.
Li
C
,
Wang
Y
,
Wang
S
,
Wu
B
,
Hao
J
,
Fan
H
, et al
.
Hepatitis B virus mRNA-mediated miR-122 inhibition upregulates PTTG1-binding protein, which promotes hepatocellular carcinoma tumor growth and cell invasion
.
J Virol
2013
;
87
:
2193
205
.
37.
Lin
X
,
Yang
Y
,
Guo
Y
,
Liu
H
,
Jiang
J
.,
Zheng
F
, et al
.
PTTG1 is involved in TNF-α-related hepatocellular carcinoma via the induction of c-myc
.
Cancer Med
2019
;
8
:
5702
15
.
38.
Yu
DY
,
Moon
HB
,
Son
JK
,
Jeong
S
,
Yu
SL
,
Yoon
H
, et al
.
Incidence of hepatocellular carcinoma in transgenic mice expressing the hepatitis B virus X-protein
.
J Hepatol
1999
;
31
:
123
32
.
39.
Levrero
M
,
Zucman-Rossi
J
.
Mechanisms of HBV-induced hepatocellular carcinoma
.
J Hepatol
2016
;
64
(
1 Suppl
):
S84
s101
.
40.
Xu
Y
,
Lv
F
,
Zhu
X
,
Wu
Y
,
Shen
X
.
Loss of asparagine synthetase suppresses the growth of human lung cancer cells by arresting cell cycle at G0–G1 phase
.
Cancer Gene Ther
2016
;
23
:
287
94
.
41.
Hinze
L
,
Labrosse
R
,
Degar
J
,
Han
T
,
Schatoff
EM
,
Schreek
S
, et al
.
Exploiting the therapeutic interaction of WNT pathway activation and asparaginase for colorectal cancer therapy
.
Cancer Discov
2020
;
10
:
1690
705
.
42.
Williams
RT
,
Guarecuco
R
,
Gates
LA
,
Barrows
D
,
Passarelli
MC
,
Carey
B
, et al
.
ZBTB1 regulates asparagine synthesis and leukemia cell response to L-asparaginase
.
Cell Metab
2020
;
31
:
852
61
.
43.
Szwed
A
,
Kim
E
,
Jacinto
E
.
Regulation and metabolic functions of mTORC1 and mTORC2
.
Physiol Rev
2021
;
101
:
1371
426
.
44.
Domínguez
A
,
Ramos-Morales
F
,
Romero
F
,
Rios
RM
,
Dreyfus
F
,
Tortolero
M
, et al
.
hpttg, a human homologue of rat pttg, is overexpressed in hematopoietic neoplasms. Evidence for a transcriptional activation function of hPTTG
.
Oncogene
1998
;
17
:
2187
93
.
45.
Hoxhaj
G
,
Manning
BD
.
The PI3K-AKT network at the interface of oncogenic signalling and cancer metabolism
.
Nat Rev Cancer
2020
;
20
:
74
88
.
46.
Ogretmen
B
.
Sphingolipid metabolism in cancer signalling and therapy
.
Nat Rev Cancer
2018
;
18
:
33
50
.
47.
Liu
Y
,
Birsoy
K
.
Asparagine, a key metabolite in cellular response to mitochondrial dysfunction
.
Trends Cancer
2021
;
7
:
479
81
.
48.
Knott
SRV
,
Wagenblast
E
,
Khan
S
,
Kim
SY
,
Soto
M
,
Wagner
M
, et al
.
Asparagine bioavailability governs metastasis in a model of breast cancer
.
Nature
2018
;
554
:
378
81
.
49.
Krall
AS
,
Xu
S
,
Graeber
TG
,
Braas
D
,
Christofk
HR
.
Asparagine promotes cancer cell proliferation through use as an amino acid exchange factor
.
Nat Commun
2016
;
7
:
11457
.
50.
Pathria
G
,
Lee
JS
,
Hasnis
E
,
Tandoc
K
,
Scott
DA
,
Verma
S
, et al
.
Translational reprogramming marks adaptation to asparagine restriction in cancer
.
Nat Cell Biol
2019
;
21
:
1590
603
.
51.
Ye
J
,
Kumanova
M
,
Hart
LS
,
Sloane
K
,
Zhang
H
,
De Panis
DN
, et al
.
The GCN2-ATF4 pathway is critical for tumour cell survival and proliferation in response to nutrient deprivation
.
EMBO J
2010
;
29
:
2082
96
.
52.
Song
P
,
Wang
Z
,
Zhang
X
,
Fan
J
,
Li
Y
,
Chen
Q
, et al
.
The role of autophagy in asparaginase-induced immune suppression of macrophages
.
Cell Death Dis
2017
;
8
:
e2721
.
53.
Costa-Silva
TA
,
Costa
IM
,
Biasoto
HP
,
Lima
GM
,
Silva
C
,
Pessoa
A
, et al
.
Critical overview of the main features and techniques used for the evaluation of the clinical applicability of L-asparaginase as a biopharmaceutical to treat blood cancer
.
Blood Rev
2020
;
43
:
100651
.
54.
Dhankhar
R
,
Gupta
V
,
Kumar
S
,
Kapoor
RK
,
Gulati
P
.
Microbial enzymes for deprivation of amino acid metabolism in malignant cells: biological strategy for cancer treatment
.
Appl Microbiol Biotechnol
2020
;
104
:
2857
69
.
55.
Mazloum-Ravasan
S
,
Madadi
E
,
Mohammadi
A
,
Mansoori
B
,
Amini
M
,
Mokhtarzadeh
A
, et al
.
Yarrowia lipolytica L-asparaginase inhibits the growth and migration of lung (A549) and breast (MCF7) cancer cells
.
Int J Biol Macromol
2021
;
170
:
406
14
.