A549, H1299, PC3, DU145 and U87-MG cell lines were purchased from the Cell Bank of the Type Culture Collection of the Chinese Academy of Sciences. A549 and U87-MG were cultured in complete Dulbecco’s modified Eagle’s medium (DMEM, cat. no. 12430-054), with 10% fetal bovine serum (cat. no. 10100-147) (both from Gibco, USA), 100 U/ml penicillin and 100 μg/ml streptomycin. DU145 and H1299 were cultured in RPMI-1640 medium (cat. no. 11875-093; Gibco) supplemented as above. PC3 cells were cultured in F-12 medium (cat. no. 21700-075; Gibco) supplemented as above. The glucose concentration in the medium was 25 mM, unless otherwise stated. For the glucose-starvation experiment, we mixed no-glucose DMEM (cat. no. 11966-025) or RPMI-1640 (cat. no. 11879-020) (both from Gibco) with the complete medium mentioned above in a certain proportion to make the glucose concentration 5 mM. We added sodium L-lactate (cat. no. 71718) or L-lactic acid (cat. no. L1750) (both from Sigma-Aldrich, USA) or HCl into the medium to create different culture environments with different lactate concentrations and pHs.

Cells (10⁴/well) were seeded in 24-well plates in complete medium with 25 mM glucose, 24 h before the experiment. The following day we changed the medium to one containing no glucose. Meanwhile, we added a different concentration of lactate into the medium at the start of the experiment. From day 2 to 12, we counted the live cells every 2 days; 3 wells/day for each culture environment.

A small interfering RNA (siRNA) targeting Bcl-2 (cat. no. GS596; Qiagen, Germany) was transfected into cells to block its function. We used Lipofectamine 2000 (cat. no. 11668019; Invitrogen, USA) as the transfection reagent and a negative control siRNA (cat. no. SI03650318; Qiagen). Forty-eight hours after transfection, we collected the mRNA and protein of the transfected cells. To assess knockdown efficiency, we analyzed the mRNA level using real-time PCR (LightCycler 480; Roche, Switzerland) and the level of protein by western blotting (Mini-PROTEAN^® 165–8004; Bio-Rad, USA). Both values were normalized to the expression of β-actin. Experiments were performed between 24 and 72 h after transfection.

The oxygen consumption rate (OCR) measurements of cells were performed using a Seahorse Bioscience XF24 instrument (Seahorse Bioscience, Billerica, MD, USA). Before running the experiment, the growth medium was removed and the cells were washed with PBS containing Ca²⁺/Mg²⁺ (pH 7.4), which was then aspirated and replaced with 700 ml of reduced serum (RS) buffer [CaCl₂ (1.8 mM), MgCl₂ (0.6 mM), KH₂PO₄ (0.5 mM), KCl (5.33 mM), Na₂HPO₄ (0.5 mM), NaCl (130 mM), glucose (5.6 mM)], glutamax, minimum essential medium (MEM) amino acid solution, MEM nonessential amino acids, MEM vitamin solution, penicillin/streptomycin, 1% bovine serum albumin (BSA, factor V fatty acid free), 1% FBS and insulin (100 nM). All components, except FBS and insulin, were combined before filter sterilization. Following the addition of FBS and insulin (usually 24–48 h pre-experiment), the RS buffer was warmed to 37°C and the pH adjusted to 7.4. The template for testing was set up as follows. Measurements were performed every 5 min, repeating metabolic measurements 3–4 times per condition for statistical analyses. The 5-min cycle included a 2-min mix period, a 1-min wait and a 2-min measuring time. After measurements of baseline activity, oligomycin (an ATP coupler) was injected, rates were measured, carbonylcyanide p-trifluoromethoxyphenylhydrazone (FCCP) (an electron transport chain accelerator) was injected, and finally rotenone and antimycin (mitochondrial inhibitors) were injected, followed by a final rate measurement.

Cells were seeded onto plates and allowed to grow in medium with 25 mM glucose for attachment overnight. The next day, we replaced the medium with fresh medium with 0 mM glucose at time zero. From time zero, every 24 h, we collected the medium samples and counted the number of cells in the same plate. The concentrations of lactate in the medium samples were measured with lactate reagent (cat. no. P0000024; CMA Microdialysis, Sweden). The results were normalized to the number of the cells in each sample.

Cells were placed in 6-well plates and incubated for 24 h before the experiment. Cells in the control group were cultured in complete medium containing 25 mM glucose. Cells in the treated group were cultured in complete medium containing 0 mM glucose with different lactate contents. After 48 h of treatment, cells were trypsinized, centrifuged and resuspended in binding buffer with Annexin V-FITC and propidium iodide (PI) from the Dead Cell Apoptosis Kit #2 (cat. no. V13241; Life Technologies, USA). Stained cells were incubated for 15 min at room temperature in the dark. Flow cytometry was used to analyze the stained cells, measuring the fluorescence emission at 530 and 575 nm using 488 nm excitation.

Treated cells were lysed using RIPA lysis buffer with 1% phenylmethanesulfonyl fluoride (PMSF). Cell lysates were separated through a 10% SDS gel and blotted onto nitrocellulose membranes. The membranes were blocked with 5% non-fat dry milk in PBST at room temperature for 1 h and incubated separately with primary antibodies against Akt (cat. no. 4685), phospho-Akt (Thr308, cat. no. 13038) (both from Cell Signaling Technology, USA), phospho-Akt (Ser473, cat. no. ab66138), PTEN (cat. no. ab32199), Mcl-1 (cat. no. ab114016), Bcl-2 (cat. no. ab117115), Bcl-xL (cat. no. ab32370), phospho-mTOR (Ser2448, cat. no. ab84400) (all from Abcam, UK) or β-actin (cat. no. A1978; Sigma-Aldrich) overnight at 4°C. The next day, the membranes were washed 3 times with PBST and incubated with secondary antibodies for 1 h at room temperature. After washing the membranes 3 times with PBST, the membranes were scanned using an Odyssey Infrared Imaging System (Li-COR Biosciences, USA).

LY294002, perifosine and rapamycin were purchased from Selleck Chemicals, USA (cat. nos. S1105, S1037 and S1039). Insulin and resveratrol were purchased from Sigma-Aldrich (cat. no. R5010). Insulin-like growth factor-1 (IGF-1) was purchased from Invitrogen (cat. no. PHG0071).

Statistical analysis was performed using GraphPad Prism 5.0 and Instat 3.1 packages (GraphPad Software, Inc., San Diego, CA, USA). The results are expressed as the means ± standard error (SE). Statistical differences between the groups were compared using t-tests. P-values <0.05 were considered to indicate statistically significant results.

To determine the effect of lactate on glucose starvation, we exposed A549 cells to different culture conditions: complete DMEM containing different sodium lactate concentrations and without glucose. We found that in glucose-free conditions, adding sodium lactate into the medium effectively prolonged the survival times of the A549 cell line (Fig. 1A). At low doses, cell proliferation and survival time increased with the increment of sodium lactate concentration. The survival time of A549 cells in the presence of 20 mM sodium lactate was longer than that of any other concentrations of sodium lactate. Indeed, A549 cells grown with sodium lactate in glucose-free conditions survived for more than 2 weeks; even 1 month later there were still viable cells. At sodium lactate concentrations >20 mM, cell proliferation and survival time decreased. Without sodium lactate, in A549 cells onset of cell death started in 1–2 days and all the cells died after 4 days. However, high concentrations of sodium lactate (such as 80 mM) caused immediate death of A549 cells (Fig. 1B).

We also confirmed the above phenomena in other types of tumor cell lines. We cultured four human cancer cell lines derived from the brain (U87-MG), prostate (PC3 and DU145), and lung (H1299) in no-glucose cultivation conditions with various lactate concentrations. The addition of sodium lactate (20 mM) effectively prolonged the survival time of all 5 types of tumor cells. We then compared the proliferation and survival time between all 5 types of cancer cells in the presence of sodium lactate. We found that DU145, A549 and H1299 cell proliferation activity was stronger than PC3 and U87-MG cells (Fig. 1C), and their survival times were longer than those of the PC3 and U87-MG cell lines (Fig. 1D).

Some studies have reported that tumor cells have the ability to take up lactate and utilize it as an energy source via oxidative phosphorylation. Determination of dissolved oxygen in the culture of cells monitored in the Seahorse Bioscience XF24 instrument showed that, upon addition of lactate, oxidative phosphorylation levels did not change significantly (Fig. 1E). In addition, we examined the lactate concentrations in the cell culture; the extracellular lactate concentrations did not change significantly over the course of the experiment (Fig. 1F). These results suggest that under deprivation of glucose, lactate was not used as a substrate for energy metabolism.

Cell secretion of lactic acid would affect the value of pH in the extracellular environment. To confirm whether the acidic environment plays a role in tumor cell survival in a glucose-free environment, we cultured A549 cells separately in the following 4 types of environment: no glucose, no glucose + 20 mM sodium lactate (pH 7.4), no glucose + 20 mM lactic acid (pH 6.8), no glucose + 20 mM hydrochloric acid (pH 6.8). After 72 h of culture, we found that adding hydrochloric acid to the tumor cells in glucose-free environment had no significant effect on survival, adding lactic acid had some effect, and adding lactate had the most significant effect (Fig. 1G). We concluded that tumor cell survival in a glucose-free environment was mainly a function of lactate itself, and had little to do with the acidic environment.

In the above experiments, we found that sodium lactate had different effects on the survival of the different cell lines in glucose-free conditions. Accordingly, we divided the five cancer cell lines into 2 groups: a lactate-sensitive group, including DU145, A549 and H1299; and a lactate-insensitive group, including PC3 and U87-MG. To explore why lactate had different effects on the survival of the 2 groups of cells in glucose-free conditions, we compared the genetic background of the 2 groups of cells. We found that the 2 lactate insensitive cell lines, PC3 and U87-MG, had lost PTEN activity (phosphatase and tensin homolog deleted on chromosome 10) (Fig. 2A). PTEN is a tumor-suppressor gene with phosphatase activity, which is involved in the negative regulation of the Akt signaling pathway, where it blocks the activation of Akt and its downstream effector molecules (13). The Akt signaling pathway plays an important role in cell proliferation and survival (14). Therefore, to confirm the existence of activation of the Akt signaling pathway in PC3 and U87-MG cell lines with PTEN deletion, we examined the phosphorylation status of Akt in the 2 cell lines (Fig. 2B). The results showed that, compared with the lactate-sensitive group, Akt phosphorylation in the lactate-insensitive group increased, which would result in Akt signaling pathway activation. We also compared the reactions of the 2 groups of cells to insulin, an activator of the Akt signaling pathway (Fig. 2C). Insulin caused a dose-dependent increase in cell numbers in the cells of the lactate-sensitive group. By contrast, the growth of the cells in the lactate-insensitive group was not affected by insulin. The above results suggest that the state of the Akt signaling pathway was a key factor that distinguished the role of lactate in cancer cell proliferation and survival time in glucose-free conditions.

To analyze Akt signaling induced by lactate, A549 cells were cultured overnight in DMEM without serum, and the cells were stimulated with sodium lactate for 30 min. Lysates of these cells were analyzed by western blotting, and Akt activation was measured by detection of Thr308 and Ser473 phosphorylated forms of Akt. We used insulin to stimulate the serum-deprived cells as a positive control for the activation of Akt. Under this environment, we observed that Akt Thr308 and Ser473 phosphorylation increased in the presence of lactate to a similar extent to that obtained with insulin (Fig. 3A and C). We then used perifosine (an inhibitor of Akt) to suppress the activation of Akt. We found that Akt Thr308 and Ser473 phosphorylation by lactate or insulin were suppressed by perifosine (Fig. 3C). We obtained the same result when we used different cell lines, such as H1299 and DU145, indicating that the differences in Akt phosphorylation were not specific to the A549 cell line (Fig. 3B). Time course experiments showed that Akt phosphorylation at Thr308 and Ser473 phosphorylation sites were both rapid (1–5 min after incubation with lactate), with maximum levels being achieved between 10 and 30 min (Fig. 3D). To estimate whether lactate induced Akt phosphorylation downstream of PI3K, we used LY294002 (an inhibitor of PI3K). Akt Thr308 and Ser473 phosphorylation in the presence of lactate were substantially reduced in the presence of LY294002 (Fig. 3E). These results suggest that lactate induced the activation of Akt significantly and rapidly through Thr308 and Ser473 phosphorylation mediated by PI3K.

Akt plays an important role in cell apoptosis and survival in response to extracellular stimuli, such as insulin or growth factors. Our previous experiments showed that the state of the PI3K/Akt signaling pathway was a key factor to distinguish the effect of lactate on cancer cell proliferation and survival in glucose-free conditions, where lactate induces the activation of Akt by Thr308 and Ser473 phosphorylation. We hypothesized that lactate helps tumor cells to survive by activating Akt in glucose-free conditions. To test the hypothesis, we cultured A549 cells with perifosine, an Akt inhibitor. We found that treatment with perifosine inhibited the cell survival induced by lactate in glucose-free conditions. In addition, treatment with an Akt activator, IGF-1, increased A549 cell survival in glucose-free conditions, but less effectively compared with lactate (Fig. 4A). These results showed that the lactate increased tumor cell survival in glucose free conditions through Akt activation.

Many studies have demonstrated that under the metabolic stress of no glucose, tumor cells undergo significant apoptosis. Thus, we investigated metabolic stress resistance to apoptosis in tumor cells incubated with lactate (Fig. 4B). We cultured A549 cells in high glucose (25 mM glucose), glucose-free and glucose-free with added lactate conditions. Twenty-four hours later, cells were trypsinized, centrifuged and resuspended in binding buffer with Annexin V-FITC and PI. The stained cells were analyzed by flow cytometry. We found that compared with high glucose conditions, there were many apoptotic cells in the glucose-free environment. There was an ~10.1- fold (P<0.001) increase in early apoptosis and an ~12-fold (P<0.001) increase in late apoptosis in the population of cells without glucose compared to the cells with high glucose. The apoptosis rate in the culture with added lactate culture was greatly reduced, and was close to that under the high glucose cultivation. The above results showed that under glucose-free conditions, a large number of A549 underwent apoptosis, and the addition of lactate could prevent this apoptosis, the mechanism of which will be further explored below.

We checked the apoptosis rate of A549 cells and then added perifosine or IGF-1. We found that treatment with perifosine prevented the decrease in apoptosis induced by lactate in glucose-free conditions, and that IGF-1 reduced the cell apoptosis caused by no glucose (Fig. 4C). We concluded that lactate prevented apoptosis in glucose-free conditions through Akt activation.

Several studies have pointed out that the anti-apoptotic Bcl-2 family is pivotal for cell survival under metabolic stress. We found lactate can reduce tumor cell apoptosis caused by the lack of glucose. We, therefore, examined whether lactate affected the expression levels of anti-apoptotic Bcl-2 family proteins such as Mcl-1, Bcl-2 and Bcl-xL. A549 cells were cultured with added lactate in glucose-free conditions. After 48 h, lysates of these cells were analyzed by western blotting. Treatment with lactate increased Bcl-2 levels, while Mcl-1 or Bcl-xL levels were not affected by treatment with lactate (Fig. 4D). To confirm whether Bcl-2 plays a major role in cell survival, we used a Bcl-2 targeting siRNA to lower its protein expression. The results showed that treatment with siBcl-2 prevented resistance to apoptosis induced by lactate in glucose-free conditions. Furthermore, we observed that resveratrol, a Bcl-2 activator, reduced the apoptosis of A549 cells in glucose-free conditions, similarly to lactate (Fig. 4E). These findings revealed that the upregulation of Bcl-2 by lactate is important for tumor cell resistance to apoptosis in glucose-free conditions.

Akt plays a core role in promoting cell survival, through activation of anti-apoptotic substances. Therefore, we aimed to confirm whether Bcl-2 upregulation is mediated via activation of the Akt/mTOR signaling pathway upon treatment with lactate in glucose-free conditions. First, we found that in A549 cells treated with lactate, Akt was activated and the expression of the Bcl-2 protein was increased, accompanied by mTOR activation (Fig. 4F). To demonstrate that the increase in the expression levels of Bcl-2 and mTOR were dependent on the activation of Akt, we added perifosine and found that the expression levels of Bcl-2 and mTOR were restored to the original levels. Furthermore, the expression levels of Bcl-2 and mTOR were upregulated in the presence of IGF-1 (Fig. 4G). Addition of rapamycin, an mTOR inhibitor, blocked the increased expression of Bcl-2 (Fig. 4H). These results demonstrated that Akt activation and mTOR upregulation by lactate in glucose-free conditions led to the upregulation of Bcl-2.

Taking all the above results into consideration, we conclude that lactate, through activation of Akt by phosphorylation mediated by PI3K, activates mTOR and further increases the expression of anti-apoptotic protein Bcl-2, to help tumor cells resist apoptosis caused by glucose starvation.