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. 2015 Apr 1;20(3):431–440. doi: 10.1007/s12192-014-0568-6

Low-pH-induced apoptosis: role of endoplasmic reticulum stress-induced calcium permeability and mitochondria-dependent signaling

Vishal Sharma 1, Ramandeep Kaur 2, Archana Bhatnagar 3, Jagdeep Kaur 1,✉
PMCID: PMC4406939  PMID: 25823563

Abstract

The acidic microenvironment around tumor cells is a major determinant in cancer growth, metabolism, and metastasis. However, its role in cancer physiology is still not clearly understood. In the present investigation, an attempt has been made to explore the effect of acidic environment on physiology of cancer cells. Exposure of Raji cells to extracellular acidic environment was associated with enhanced cytosolic calcium level and endoplasmic reticulum stress response. X-box binding protein 1 (XBP1) splicing, CCAAT/enhancer-binding protein homologous protein (CHOP), and glucose-regulated protein 78 kDa (GRP78) upregulation suggested endoplasmic reticulum stress generation. On the other hand, real-time-based upregulation of Bax gene expression and flow cytometric analysis of cytochrome c release as well as enhanced active caspase-3 further confirmed mitochondrion-mediated events leading to induction of apoptosis. The expression of TP53 and p21 was upregulated. These observations collectively strongly suggest that both endoplasmic reticulum stress-mediated calcium release and Bax targeting might be altering mitochondrion membrane potential which in turn could induce secondary apoptotic signals; subsequently, endoplasmic reticulum stress can also lead to nuclear localization of Nuclear factor-κB (NF-κB) which in turn favors p53 mediated apoptotic signals.

Electronic supplementary material

The online version of this article (doi:10.1007/s12192-014-0568-6) contains supplementary material, which is available to authorized users.

Keywords: Acidic pH, Apoptosis, Calcium, Cytochrome c, Endoplasmic reticulum

Introduction

The body must maintain proper balance of acidity to alkalinity (pH) in the bloodstream in order to function properly. In cancer, exponentially growing cells receive poorly organized and only marginally functional vasculature. Due to increased glycolytic activity of malignant cells, insufficient vascular supply, and sluggish blood circulation, intratumor environment becomes acidic with pH less than 6.0 in certain tumors (astrocytomas and squamous cell carcinomas) as compared to a normal physiological pH of 7.3 (Schornack and Gillies 2003; Vaupel et al. 1989). In addition, low influx of metabolites and a low efflux of potentially toxic metabolic wastes further enhance the acidic conditions (Tannock and Rotin 1989). Acidic environment plays an important role in cell death during various pathophysiological states, including cancer (Williams et al. 1999). A shift to acidic pH within solid tumors can regulate proliferation, angiogenesis, immune suppression, invasion, and chemo resistance. Likewise, acidic environment or low pH can also influence oncogenesis, malignant transformation, and metastasis (Rofstad et al. 2006). The environmental acidity also greatly influences the response of cancer cells to various treatments. There are several contradictory reports in literature regarding the effect of acidic environment on physiology of cells; some studies suggest that low pH can promote tumor progression while others indicate that it can induce apoptosis in cancer cells (Hjelmeland et al. 2011; Park et al. 1999). It has been demonstrated that acidic stress can lead to the upregulation of Src activity, matrix metalloproteinases (MMPs), and VEGF expression, which in turn can promote tumor progression and metastasis (Chen et al. 2008; Kato et al. 1996, 2005; Xu et al. 2002; Fukumura et al. 2001). Studies have demonstrated that low pH has the ability to induce genes and pathways which collectively lead to progression of malignancy (Duggan et al. 2006). Bhat et al. (2012) have shown that acidic stress can promote oligomerization and membrane insertion of apoptosis repressor BclXL (Bhat et al. 2012). In contrast, Park et al. (1999) reported that acidic pH can promote the factors which can move the cancer cells physiology toward apoptosis; i.e., acidic pH can have therapeutic potential. Acidic pH-mediated induction of apoptosis was suggested to be a consequence of caspase activity (Park et al. 1999). Acidic environment could modulate growth as well as radiochemosensitivity of glioma cells (Reichert et al. 2002) and was demonstrated to have immuno-modulatory activity by promoting the expression of IL-8 in cervical cancer cells (Xu and Fidler 2000). Hypoxia and acidity represent two factors that might be exploited therapeutically to destroy such cells. Recently, resveratrol and many drugs like lovastatin and cantharidin have been shown to be more effective in a low pH environment than at normal pH (Shamim et al. 2012; Fukamachi et al. 2010). It has been demonstrated that intracellular acidification by exposure to acidic environment or by alteration in intracellular pH control mechanisms leads to cellular damage and sensitizes cells to chemotherapy or hyperthermia (Haveman 1979; Chu and Dewey 1988). But, even after extensive research in this area, there is a dearth of information about molecular mechanism involved in various physiological responses under acidic environment and the therapeutic potential of acidifying tumor microenvironment.

Leukemia is cancer of blood cells, and it has been shown that acidic microenvironment is a key feature of exponentially growing cells in bone marrow (Mortensen et al. 1998). Especially, acute leukemia is a rapidly progressing disease that results in the accumulation of immature, abnormal cells in the marrow and blood. The metabolic activity of rapidly growing cancer cells could alter environment around the cancer cells in soft tissue and bone marrow. Therefore, in the present investigation, acute lymphoid leukemia Raji cells were selected as a suitable model to study the effect of acidic environment on leukemic cell physiology. It was observed that Raji cells exposed to extracellular acidic environment underwent apoptosis without any contribution of reactive oxygen species (ROS). Further, the growth of Raji cells in a medium of low pH resulted in an augmentation in intracellular calcium level and generation of endoplasmic reticulum (ER) stress that induced apoptosis by Bax targeting, cytochrome c release, and caspase activation. Raji cells exposed to acidic stress also showed upregulation of TP53 and p21 and nuclear localization of Nuclear factor-κB (NF-κB). In conclusion, the study highlights that alteration (lowering of pH) of tumor microenvironment can be used as a therapeutic option to suppress tumor growth.

Materials and methods

Cell culture and treatment

Exponentially growing Raji cells (human acute lymphoid leukemia cells) were procured from the National Centre for Cell Science (NCCS), Pune. The cells were cultured under a humidified 5 % carbon dioxide and 95 % air atmosphere at 37 °C. The cell density was maintained at fewer than 3 × 105 cells/ml in 25-cm2 plastic tissue culture flasks with RPMI-1640 culture medium supplemented with 10 % (v/v) fetal calf serum. To determine the effect of low pH on Raji cells, cells were plated at a density of 0.5 × 106 cells/ml in culture media of pH 5.8 and 6.8 for 48 h and cells grown at physiological pH 7.3 were taken as control. pH values of 6.8 and 5.8 were selected based on the studies which suggested that tumor pH can vary from 6.8 to 5.8 (Song et al. 2006; Tannock and Rotin 1989).

Cellular proliferation assay

Cell viability was measured by the 3-(4,5-dimethythiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay. This method is based on the ability of viable cells to reduce MTT to form colored formazan product (Heckenkamp et al. 1999). Briefly, cells were cultured in 96-well plates at a density of 1 × 104 cells per well in media with different pH—pH 7.3, pH 6.8, pH 6.3, and pH 5.8. After incubation for 12, 24, and 48 h, MTT dissolved in phosphate-buffered saline (PBS) was added to each well at a final concentration of 5 mg/ml and then incubated at 37 °C and 5 % CO2 for 2 h. The water-insoluble dark blue formazan was dissolved in DMSO. Optical density was measured by a microplate reader (Bio-Rad) at a wavelength of 570 nm.

DNA fragmentation assay

Along with control (pH 7.3), 0.5 × 106 cells/ml were cultured in acidic conditions (pH 6.8, pH 6.3 pH 5.8) for 48 h. DNA from treated cells was extracted according to Gong’s modified method (Gong et al. 1994). Briefly, cells were washed twice in cold PBS (10 mM) and resuspended in hypotonic lysis buffer (5 mM Tris HCl, 20 mM EDTA, pH 7.4) containing 0.5 % Triton X-100 for 30 min at 4 °C. The lysate was centrifuged at 13,000 g for 15 min at 4 °C. DNA was extracted from the supernatant with equal volume of phenol-chloroform-isoamylalcohol, precipitated by addition of 0.1 volume of 3 mM sodium acetate and two volumes of absolute ethanol. After treatment with RNAse A (500 U/ml) at 37 °C for 3 h, the pattern of DNA fragmentation was analyzed on 1.5 % agarose gel.

RT-PCR and quantification of mRNA expression levels

Total RNA was isolated from treated and untreated cells using TRIzol reagent (Sigma, USA). One microgram RNA was used for complementary DNA (cDNA) preparation using Verso cDNA kit (Thermo Scientific, USA). DNA contamination in total RNA isolated was avoided using reverse transcription (RT) enhancer available with kit. Real-time PCR analysis was performed in Eppendorf realplex system using the SYBR Green PCR Master mix (Thermo Scientific, USA). Real-time PCR was carried out for TP53 (Yang et al. 2013), p21 (Fields et al. 2005), Bax (Sharma et al. 2012), and β-actin (Jha et al. 2010) using gene-specific primers. β-Actin served as an internal control. Specificity of PCR products was analyzed using melting curve analysis, and delta CT method was used to quantify alteration in expression.

Detection of XBP1 mRNA splicing

Total RNA was extracted from treated and untreated cells and subjected to cDNA preparation as described above. The cDNAs were PCR amplified using specific primers for XBP1 (Zhu et al. 2012), CCAAT/enhancer-binding protein homologous protein (CHOP) (Wang et al. 2011), and glucose-regulated protein 78 kDa (GRP78) (Lin et al. 2012). The primers used were XBP1F (5′-GGTAAGGAACTGGGTCCTT-3′) and XBP1R (5′-GAGTTAAGACAGCGCTTGGG-3′). A 331-bp PCR product was amplified from the unspliced form of XBP1 messenger RNA (mRNA), which contains the 26-bp intron, and a 305-bp PCR product was amplified from the spliced form of XBP1 mRNA. The PCR products were separated on a 12.5 % polyacrylamide gel.

Cytochrome c release, active caspase-3, and p21 determination

Cells (0.5 × 106 cells/ml) were washed once in PBS and then fixed and permeabilized using the Cytofix/Cytoperm kit (BD Biosciences, USA) for 20 min on ice. Cells were pelleted, washed with Perm Wash Buffer (BD Biosciences, USA), and stained with fluorochrome-conjugated antibody cytochrome c mAb (Santa Cruz Biotechnology, USA), rabbit anti-human active caspase-3 fluorescein isothiocyanate (FITC) mAb (BD Biosciences, USA), or p21 PE mAb (Santa Cruz Biotechnology, USA) at 4 °C for 1 h. Cells were washed twice with Perm Wash Buffer and finally resuspended in Perm Wash Buffer for flow cytometry analysis. Flow cytometric analysis was performed on a BD FACS Canto II (BD Biosciences, USA) for a maximum cell count of 5000 and analyzed using BD FACS Diva software.

Cytosolic calcium and ROS determination

Cytosolic Ca2+ levels were determined using the fluorescent dye Fluo 3-AM (1 mM) (log mode in FITC setting). Treated and untreated cells were incubated with fluorescent dye for 15 min at 37 °C, washed with PBS containing 10 mM glucose, and analyzed immediately by flow cytometry. The intracellular accumulation of ROS was determined using 2′,7′ dichlorofluorescin diacetate (DCFH-DA) (Sigma, USA). After treatment, the cells were washed with PBS, stained with DCFH-DA for 20 min at 37 °C, and analyzed with flow cytometry for maximum cell count of 5000.

Western blotting

Cells were harvested and cell pellet was washed with cold phosphate-buffer saline. Cells were lysed in lysis buffer (20 mM Tris HCl (pH 7.5), 150 mM NaCl, 1 % NP-40, 1 mM ethylene glycol-bis(β-aminoethyl ether)-tetraacetic acid, 1 mM EDTA, 50 mM NaF, 1 mM β-glycerophosphate, 2.5 mM sodium pyrophosphate, 1 mM orthovanadate, one protease inhibitor cocktail, 1 mM PMSF), and protein content in the supernatant was determined by protein estimation kit (Bangalore Genie). Equal amount of cell lysate (200 μg/well) from both treated (pH 6.8, pH 5.8) and untreated (pH 7.3) was resolved on 10 % SDS-PAGE followed by transfer onto a PVDF membrane. Membranes were probed with anti p53 (1:1000), Bax (1:1000), and NF-κB (1:1000) antibodies (Sigma, USA). Horseradish peroxidise-conjugated secondary antibodies were used to detect immune-reactive bands using 3,3′-diaminobenzidine as substrate.

Subcellular fractionation

After treatment, Raji cells were suspended in hypotonic buffer A (250 mM sucrose, 20 mM Hepes, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 100 μM PMSF) on ice and incubated for 30 min followed by pipetting for 30 min. Cell lysate was centrifuged at 8700 × g for 10 min at 4 °C to remove unlysed cells and nuclei. The supernatant was centrifuged at 43,000 × g for 5 min at 4 °C. The pellet was saved as mitochondrion fraction followed by four times washing of pellet with lysis buffer. With the Pasteur pipette, supernatant was recovered and centrifuged at 110,000 × g for 1 h at 4 °C. The supernatant was saved as cytosolic fraction and the pellet as ER/microsome fraction. To further purified ER/microsome fractions, the post-mitochondrial supernatant was passed through stepwise sucrose gradients (1.5–0.25 M) and sample-containing tube was centrifuged at 280,000 × g for 1 h. For quality assessment of mitochondrion fraction, succinic dehydrogenase activity assay was carried out as described previously (Jones et al. 2013). For ER/microsome fraction, cytochrome p450 reductase assay was carried out for quality assessment as described previously (Yim et al. 2005).

Statistical analysis

The data was analyzed statistically using SPSS 13.0. The results were analyzed using one-way ANOVA to determine the significance of the mean between groups. Values of P < 0.005 were considered to be significant. The means of the data are presented together with standard error of mean (SEM) from multiple independent experiments.

Results and discussion

It has been a well-known fact that microenvironment in exponentially growing cancer is generally acidic as compared to normal tissue and low extracellular pH has been documented to induce coordinate regulation of gene expression in cells (Duggan et al. 2006).

Acidic environment-induced apoptosis

Anti-proliferative effect of acidic pH on Raji cells was studied by MTT assay. Cells were grown in media of pH 6.8 and 5.8, and cell viability was analyzed. After 12 h of incubation, no significant variation was observed at different pH. After 24 h of incubation at pH 6.8, no variation in cell viability was observed while nearly 89 % cells were viable at pH 5.8 after 24 h. A drastic change was observed after 48 h of incubation. At pH 5.8, the survival of cells decreased to nearly 2.5-fold when compared with cell viability at pH 6.8. To examine the effect of acidic stress treatment on cancer cells, Raji cells were cultured in different pH media (7.3, 6.8, 6.3, and 5.8) for 48 h and analyzed for apoptosis by DNA laddering assay. In comparison to control (pH 7.3) and cells grown in medium of pH 6.8, the DNA of Raji cells grown in pH 5.8 medium demonstrated markedly increased fragmented DNA (Fig. 1). These results indicate the involvement of apoptosis in low (acidic)-pH-mediated cell cytotoxicity in Raji cells. These results were in agreement with the earlier studies which strongly suggested that acidic stress can induce DNA damage and apoptosis (Jolly et al. 2004). Flow cytometric evaluation of cell morphology at low pH showed a shift in cellular granularity, which further suggested unhealthy or cells prone to apoptosis at the acidic pH as compared to control (pH 7.3) (Fig. 2).

Fig. 1.

Fig. 1

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Acidic stress-induced apoptosis in Raji cells. a Percentage viability of Raji cells grown in media of pH 7.3, 6.8, 6.3, and 5.8 after 12, 24, and 48 h. b Detection of apoptosis by DNA laddering assay. Raji cells were grown in acidic environment (pH 6.8, pH 6.3, and pH 5.8) for 48 h, and the cells grown at pH 7.3 were taken as control

Fig. 2.

Fig. 2

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Increase in intracellular calcium at acidic pH. Raji cells were cultured in normal (pH 7.3) and acidic media pH 6.8 and pH 5.8 for 48 h followed by incubation with the fluorescent dye Fluo 3-AM. Representative FACS analysis histograms of Fluo 3-AM stained Raji cells in control and after acidic stress treatment. Five thousand events were counted. Increase in calcium at pH 5.8 was found statically significant (p value <0.005) with respect to control. Cells at physiological pH (pH 7.3) were taken as control

Acidic stress-affected calcium homeostasis

To interrogate the role of acidic stress on Ca2+ homeostasis, we selectively examined cytosolic Ca2+ in Raji cells in normal and acidic stress conditions. The increase of ER membrane permeability and calcium release from ER are related with the induction of apoptosis in tumor cells (Boya et al. 2002). In comparison to control (pH 7.3), approximately 8-fold enhanced cytosolic Ca2+ was detected in Raji cells when pH of the growth medium was shifted to 5.8 (Fig. 2). Thus, acid stress-mediated dysregulation of Ca2+ homeostasis might indicate ER stress which in turn triggers apoptosis.

Acidic stress-induced endoplasmic reticulum stress

Considering the enhanced cytosolic Ca2+ after acidic stress, we further studied the effect of acidic stress on endoplasmic reticulum, the major cellular store of calcium. Factors and conditions that disrupt protein folding in the endoplasmic reticulum, such as a chemical, nutrient, and environmental stress, activate stress signaling pathway jointly known as unfolded protein response (UPR). UPR activation ultimately moves the cell physiology toward cell-cycle arrest and the induction of apoptosis. Endoplasmic reticulum stress induces activation of the IRE1, an endonuclease that causes the unconventional splicing of mRNA encoding a UPR-specific transcription factor, called X-box binding protein 1 (XBP1) (Schadewijk et al. 2012). UPR is mediated by upregulation of chaperones such as GRP78, and apoptotic pathways are activated and mediated by the induction of CHOP. Induction of stress response genes XBP1, GRP78, and CHOP in response to growth of cells in acidic environment was studied by RT-PCR. To evaluate the induction of IRE1α/ XBP1 pathway in the cells grown under acidic environment, XBP1 splicing was examined by RT-PCR. XBP1 mRNA splicing, indicative of ER stress, was detected in the cells grown at pH 5.8 (Fig. 3). In contrast, XBP1 splicing was not induced in the cells grown at pH 6.8 and 7.3. Similarly, upregulation of GRP78 and CHOP mRNA in cells exposed to acidic pH provides further evidence of generation of ER stress in these cells. Thus, the exposure of Raji cells to extracellular acidic environment resulted in activation of endoplasmic reticulum stress response.

Fig. 3.

Fig. 3

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Effect of acidic stress on XBP1 splicing, CHOP, and GRP78 in Raji cells. Raji cells were grown at pH 7.3, pH 6.8, and pH 5.8 (acidic stress) for 48 h. Total RNA was extracted from the cells, and the mRNA of XBP1 was detected and analyzed by RT-PCR. The RT-PCR products of XBP1 were analyzed on 12.5 % PAGE, and GRP78 and CHOP were analyzed on 2 % agarose gel. β-Actin was used as internal control. Cells grown at physiological pH 7.3 were taken as control

Acidic stress-mediated cell death associated with Bax translocation to mitochondrion

Bax, a proapoptotic Bcl-2 family member, is involved in mitochondrial apoptotic signaling pathway. Bax is also known to be induced by p53, and Bax redistributes both to mitochondria and ER upon apoptosis induction (Smith and Deshmukh 2007). Therefore, the expression of Bax after exposure of cells to acidic environment was evaluated by quantitative RT-PCR. The expression of Bax was upregulated (4.2-fold) in cells grown under acidic stress, i.e., pH 5.8 as compared to control (Fig. 4). These results suggested that acidic stress-mediated induction of apoptosis is associated with Bax. Further, to confirm the role of Bax in apoptosis, cytosolic, endoplasmic reticulum, and mitochondrion fractions were isolated and analyzed for Bax distribution. In cytosolic fractions, no significant changes in the level of Bax were observed while in endoplasmic reticulum fraction, a significant decrease was observed at pH 5.8. In contrast to it, a significant increase in Bax level was observed in mitochondrion fraction at pH 5.8 as compared to control (pH 7.3). These results are in agreement with earlier studies, which strongly suggested that endoplasmic reticulum stress causes robust activation, translocation, and N-terminal exposure of Bax in the mitochondria (Zhang and Armstrong 2007).

Fig. 4.

Fig. 4

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Bax induction and translocation to mitochondrion by low pH. a Raji cells were incubated in acidic environment pH 6.8 and pH 5.8 for 48 h. Expression analysis was carried out by quantitative real-time PCR. Real-time PCR was carried out using gene-specific primers, and specificity of product was confirmed by melting curve analysis. The data was normalized to β-actin. b Subcellular fractionation was performed to obtain the fractions for cytosol, mitochondria, and ER/microsome followed by western blot analysis using anti Bax-specific monoclonal antibody. Under acidic environment, a significant increase in Bax expression at pH 5.8 was observed in mitochondrion fraction while in contrast to it, Bax expression decreased in endoplasmic fractions. Cells grown at physiological pH (pH 7.3) were taken as control

Acidic stress caused cytochrome c release, enhanced active caspase-3 expression, and PARP cleavage

Both Bax and calcium have been documented to alter mitochondrial permeability. To reveal the role of mitochondria in acidic stress-induced apoptosis, the status of cytosolic cytochrome c in normal and cells grown in acidic medium was analyzed (Fig. 5a). Culturing of Raji cells in acidic environment (pH 5.8) leads to nearly four times higher release of cytochrome c when compared to control. Cytochrome c is an intermediate in apoptosis. A major consequence of cytochrome c release is the induction of caspase cascade mediated apoptosis induction. To verify caspase induction, the expression of active caspase-3, which is a downstream target of caspase-9, was analyzed by flow cytometry (Fig. 5b). Raji cells, cultured at acidic pH 5.8, demonstrated nearly 2.3-fold increase in expression of active caspase-3 as compared to control. Incubation of Raji cells in acidic media resulted in cleavage of poly(ADP-ribose) polymerase (PARP) to 85-kDa fragment. PARP cleavage was significant in pH 6.8 medium, and it further increased in pH 5.8 medium (Fig. 5c). These results suggested that acidic stress-induced apoptosis was accompanied by cytochrome c release which in turn leads to the induction of caspase-3 mediated apoptotic pathway.

Fig. 5.

Fig. 5

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Acidic stress treatment leads to enhanced cytochrome c release and caspase-3 activation. a Representative FACS analysis histograms of anti-cytochrome c stained Raji cells after acidic stress. Bar graph showing cytochrome c expression after acidic stress treatment expressed as mean florescence intensity. b Representative FACS analysis histograms of anti-active caspase-3 stained Raji cells after acidic stress. Bar graph showing caspase-3 expressions after acidic stress treatment expressed as mean florescence intensity. A significant (P value <0.005) upregulation in active caspase-3 was observed with respect to control. Cells grown at physiological pH (pH 7.3) were taken as control. Five thousand events were counted per tube. c Illustrated PARP cleavage leads to generation of 85-kDa fragments as compare to control

Acidic stress mediated ROS independent apoptosis

In order to investigate the effect of acidic pH medium on ROS-mediated induction of apoptosis, we performed flow cytometric analysis on Raji cells after treatment at pH 6.8 and pH 5.8. We did not observe any significant alteration in ROS after acidic stress treatment (Fig. 6) after 48 and 24 h (supplementary Fig. 1). The result suggests that ROS is not involved in inducing apoptosis in Raji cells in acidic growth conditions.

Fig. 6.

Fig. 6

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Low pH caused ROS independent cell death. Raji cells exposed to acidic stress were incubated with DCFH-DA after and 48 h followed by flow cytometry analysis. Cells grown at physiological pH (pH 7.3) were taken as control. Acidic stress-mediated induction of apoptosis was found to be independent of ROS

Acidic stress caused upregulation of TP53, p21, and nuclear localization of NF-κB

Recently, studies have suggested that endoplasmic reticulum stress involves the key role of NF-κB in regulation of p53 and its association with endoplasmic reticulum (Lin et al. 2012; Yang et al. 2013). Therefore, the effect of acidic stress on localization of NF-κB along with expression of TP53 and its downstream effector p21 was investigated. The p21 (CIP1/WAF1) protein that is upregulated by p53 in response to various stimuli binds to and inhibits the activity of cyclin-CDK2 or CDK1 complexes and thus functions as a regulator of cell cycle progression. In response to extracellular acidic stress, the level of TP53 mRNA was enhanced up to about 7-fold and p21 mRNA was approximately 14-fold higher as compared with the control (Fig. 7a). These findings are in accordance with the earlier reports which suggested the activation of TP53 and p21 in acidic environment (Williams et al. 1999). The elevated level of p53 and p21 transcripts correlated with the elevated expression of p53 and p21 proteins in acidic stress condition (Fig. 7c, b). Although the induction of p53 in response to acidic environment exposure is correlated with upregulation of p21 expression, it is not conclusive that mutated p53 in Raji cells can upregulate p21 expression. It is possible that p53 in Raji cell line which harbors two p53 alleles each having a point mutation resulting in changed amino acids (Duthu et al. 1992) retains some of its wild-type function to regulate p21 expression. Further, NF-κB translocation to nucleus in response to acidic environment (Fig. 7c) also indicated that p53 might be regulated by NF-κB in Raji cells. It suggested that under acidic pH, the cumulative effect of calcium and Bax induction along with p53 induction might contribute to apoptosis.

Fig. 7.

Fig. 7

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Acidic environment upregulated mRNA level of TP53 and p21 and elevated the expression of NF-κB, p53, and p21 in Raji cells. a Quantitative real-time PCR-based analysis of TP53 and p21 expression. RNA was isolated from control and acidic stress (pH 5.8 and pH 6.8) exposed Raji cells. Real-time PCR was carried out using gene-specific primers, and specificity of product was confirmed by melting curve analysis. A significant (P value <0.005) increase in p21 mRNA was found at pH 6.8 and pH 5.8 with respect to control. b Representative FACS analysis histograms of anti-p21 stained Raji cells after acidic stress. Bar graph showing p21 expression after acidic stress treatment expressed as mean florescence intensity. c Induction of p53 and nuclear localization of NF-κB by acidic environment in Raji cells. The data was normalized to β-actin for real-time analysis and GAPDH for Western blot analysis. Cells at pH (pH 7.3) were taken as control

Conclusion

The findings from the present study suggest that acidic stress causes apoptosis of Raji cells via mitochondrion-dependent and ER stress-triggered signaling pathway. This is evident from acidic stress-mediated enhancement of cytosolic Ca+2, stimulation of XBP1 splicing, induction of Bax, and release of cytochrome c in response to acidic stress (Fig. 8). Therefore, it is proposed that acidic stress could activate distinct apoptotic events, which involve ER stress generation and mitochondrial membrane permeability transition. To the best of our knowledge, it is the first report showing induction of endoplasmic reticulum stress under acidic stress in Raji cells

Fig. 8.

Fig. 8

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Proposed hypothesis or pathway

Electronic supplementary material

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Acknowledgments

This study was supported by the Council of Scientific and Industrial Research (CSIR), New Delhi, India.

Contributor Information

Vishal Sharma, Email: Sharmavishal_biotech@yahoo.com.

Ramandeep Kaur, Email: gndu.ramandeep@gmail.com.

Archana Bhatnagar, Email: bhatnagar.archana@gmail.com.

Jagdeep Kaur, Email: jagsekhon@yahoo.com.

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