Decreased expression of connective tissue growth factor in non-small cell lung cancer is associated with clinicopathological variables and can be restored by epigenetic modifiers

Goat polyclonal anti-CTGF antibody (Ab) (L-20), rabbit polyclonal anti-glyceraldehyde-3-phosphate (GAPDH) Ab (FL-335), rabbit anti-goat and goat anti-rabbit horseradish peroxidase (HRP)-conjugated Ab were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). TRI Reagent^®, 5-dAzaC, TSA, dimethyl sulfoxide (DMSO), ethanol, fetal bovine serum (FBS), cell culture antibiotics and media were provided by Sigma-Aldrich Co. (St. Louis, MO).

Primary lung cancerous and histopathologically unchanged lung tissues, located at least 10–20 cm away from the cancerous lesions, were obtained between March 2012 and December 2014 from 98 patients diagnosed with NSCLC, who underwent surgical resection at the Department of Thoracic Surgery, Poznan University of Medical Sciences, Poland (Tables 1, 2, 3; Supplementary tables 1 and 2). Among them, 18 patients were never smokers. None of the patients received any preoperative chemotherapy and/or radiation therapy. Histopathological classification was performed by an experienced pathologist. After surgical removal, tissue samples were immediately snap-frozen in liquid nitrogen and stored at −80 °C until further processing. All patients participated in this study had signed informed consent on the use of clinical specimens, and the study was approved by the Local Ethical Committee of Poznan University of Medical Sciences.

The human NSCLC cell lines—A549 and Calu-1, breast cancer cell line—T47D, and cervical carcinoma cell line—HeLa, were purchased from ATCC (Rockville, MD). Normal human bronchial epithelial cells—Beas-2B, were kindly provided by Dr M. Rusin from the Maria Skłodowska-Curie Memorial Cancer Center and Institute of Oncology, Gliwice Branch, Poland. A549, Calu-1 and T47D cells were routinely maintained in RPMI 1640 medium, Beas-2B cell line was cultured in DMEM/F12 medium, and HeLa cell line was maintained in DMEM. Cells were grown at 37 °C in humidified air with 5 % CO₂. All culture media were supplemented with 10 % heat-inactivated FBS, 2 mM glutamine and penicillin–streptomycin–amphotericin B solution (10,000 U penicillin, 10 mg streptomycin and 25 μg amphotericin B/ml).

The stock solutions of 5-dAzaC (10 mg/ml) and TSA (1 mg/ml) were prepared in DMSO and ethanol, respectively, aliquoted and stored at −20 °C until use. Prior to all experiments, cells were seeded and grown overnight. Next, the investigated compounds were diluted in the culture medium to the desired concentration and added to cell cultures. The same volume of DMSO or ethanol was used as a vehicle control, and their final concentration in culture medium never exceeded 0.1 %. All experimental media were exchanged every 24 h. To determine the effect of 5-dAzaC on CTGF transcript and protein levels in NSCLC cell lines, A549 and Calu-1 cells were cultured for 48, 72 and 96 h either in the absence or in the presence of 5-dAzaC at concentrations of 10 and 15 μM. In order to evaluate whether TSA may regulate CTGF transcript and protein contents in A549 and Calu-1 cell lines, cells were treated with different concentrations of TSA (30, 100, 300 nM) or vehicle control in a time-dependent manner for 12, 24, 48 and 72 h. Each experiment was performed in triplicate, and cells were used for protein and RNA isolation, Western blotting and RT-qPCR analysis. Additionally, genomic DNA was isolated from cells cultured in the absence or in the presence of 10 μM 5-dAzaC for 96 h in order to evaluate its impact on the methylation status of CTGF regulatory region.

A549 and Calu-1 cells were seeded into T-25-cm² flasks and treated with 5-dAzaC or TSA as described above. After each incubation period, cells were detached with Trypsin–EDTA solution, Sigma-Aldrich Co. (St. Louis, MO), centrifuged and resuspended in 8 ml of phosphate-buffered saline (PBS). Next, cell viability and proliferation were determined by a trypan blue staining. Fifty μl of cell suspension was taken and mixed with the same volume of trypan blue. Cells were counted with an EVE™ automatic cell counter, NanoEnTek Inc. (Seoul, Korea). For cell viability, results are expressed as the percentage of viable cells relative to respective controls (100 %), and for cell proliferation, the number of counted cells per 1 ml of solution is presented. All experiments were performed in triplicate, and data represent mean ± standard deviation (SD).

Total RNA from lung tumors and matched normal lung tissues obtained from the same patients, as well as from investigated cell lines, was isolated according to the method of Chomczyński and Sacchi (1987). RNA samples were quantified and reverse-transcribed into cDNA using M-MLV reverse transcriptase from Invitrogen, Life Technologies (Grand Island, NY), according to the manufacturer’s protocol. The RT-qPCR was conducted in a Light Cycler^®480 Real-Time PCR System, Roche Diagnostics GmbH (Mannheim, Germany), using SYBR Green I as a detection dye. The target cDNA was quantified by relative quantification method using a calibrator for primary tissues or respective controls for A549 and Calu-1 cells. For the calibrator, 1 µl of cDNAs from all tissue samples was mixed together. The quantity of CTGF transcript in each sample was standardized by the geometric mean of porphobilinogen deaminase (PBGD) and human mitochondrial ribosomal protein L19 (hMRPL19) cDNA levels. The PCR amplification efficiency for target and reference genes was determined by the different standard curves, created by consecutive dilutions of the cDNA template mixture, as provided in Relative Quantification Manual, Roche Diagnostics GmbH (Mannheim, Germany). For amplification, 1 µl of total (20 µl) cDNA solution was added to 9 µl of Light Cycler^®480 SYBR Green I Master mix (1 × concentrated) containing 2.5 mM MgCl₂ and 0.5 µM primers (Supplementary table 3). A sample of no reverse-transcribed RNA and a no-template control were included in each batch of samples to provide a negative control in subsequent PCR. Melting curve analysis and electrophoresis were applied to confirm the specificity of the amplified products. All experiments were performed in triplicate, and CTGF transcript levels in investigated tissues were expressed as the decimal logarithm of multiplicity of cDNA concentrations in the calibrator. CTGF transcript levels for A549 and Calu-1 cell lines treated with 5-dAzaC or TSA were presented as multiplicity of the respective controls.

Cells were harvested using trypsin–EDTA solution, washed twice with PBS and lysed using RIPA lysis buffer, Sigma-Aldrich Co. (St. Louis, MO). Tissue specimens were homogenized in liquid nitrogen and also treated with RIPA lysis buffer. In both cases, RIPA buffer was supplemented with protease inhibitor cocktail, Roche Diagnostics GmbH (Mannheim, Germany). Samples were incubated on ice for 30 min, with vortexing every 15 min, and then centrifuged at 10,000×g for 10 min at 4 °C in order to remove cell debris. Supernatant was collected for whole cell lysates. The concentration of total protein isolated from cell lines was determined by the bicinchoninic acid assay method (BCA) using BCA kit from Sigma-Aldrich Co. (St. Louis, MO). Next, proteins (30 µg for cell lines) were resuspended in a sample loading buffer, boiled at 95 °C for 10 min, rapidly cooled on ice and separated on 12 % Tris–glycine gel using SDS–PAGE. Gel proteins were transferred to a nitrocellulose membrane, which was then blocked with 5 % nonfat dry milk in 1× concentrated Tris-buffered saline/Tween 20 for 2 h at room temperature, on a shaker. After blocking, membranes were incubated overnight at 4 °C with goat polyclonal anti-CTGF Ab (L-20) at a dilution of 1:500, followed by 2-h incubation with rabbit anti-goat HRP-conjugated Ab (1:5000). To ensure protein loading control of the lanes, membranes were stripped and incubated with rabbit polyclonal anti-GAPDH Ab (FL-335; 1:3300), followed by incubation with goat anti-rabbit HRP-conjugated Ab (1:5000). Bands were revealed using SuperSignal West Femto Chemiluminescent Substrate, Thermo Fisher Scientific (Rockford, IL) and Biospectrum^® Imaging System 500, UVP Ltd. (Upland, CA). The amounts of analyzed proteins were presented as the CTGF-to-GAPDH band optical density ratio. For A549 and Calu-1 cells cultured in the absence of 5-dAzaC or TSA, the ratio of CTGF to GAPDH was assumed to be 1. Additionally, HeLa cell line lysate was used as a positive control for CTGF protein identification according to CTGF Ab datasheet.

Formalin-fixed, paraffin wax-embedded tissue specimens were cut on 4-μm sections and mounted on adhesion microscope slides SuperFrost^®Plus (Menzel Gläser). Next, sections were dewaxed and rehydrated, and heat-induced antigen retrieval was carried out by cooking in low pH EnVision FLEX Target Retrieval Solution, Dako (Glostrup, Denmark), for 50 min at 97 °C. Endogenous peroxidase was blocked by EnVision FLEX Peroxidase-Blocking Reagent, Dako (Glostrup, Denmark), and sections were incubated in Novocastra Protein Block, Leica Biosystems (Wetzlar, Germany), for 10 min followed by overnight incubation with goat polyclonal anti-CTGF Ab (L-20; dilution 1:200) in a humid chamber at 4 °C. The primary antibody was diluted in EnVision FLEX Antibody Diluent, Dako (Glostrup, Denmark). Immunodetection was achieved using the LSAB System, Dako (Glostrup, Denmark), which is a two-step streptavidin–biotin–HRP method. Each layer was incubated for half an hour at room temperature. Between each staining steps slides were washed in EnVision FLEX Wash Buffer, Dako (Glostrup, Denmark). Visualization was achieved using 3-3′-diaminobenzidine tetrachloride (DAB, Leica Microsystems). Sections were counterstained with Mayer’s hematoxylin, dehydrated, cleared and mounted in DPX. Sections from formalin-fixed and paraffin-embedded normal human lung were used as positive control. Moreover, the presence of CTGF staining in histopathologically unchanged respiratory epithelium, macrophages and stromal fibroblasts within the tumor sections served as an internal positive control. Immunohistochemical staining was evaluated by experienced pathologist.

The position of CpG islands in the CTGF regulatory region was determined by online programs (UCSC Genome Bioinformatics Site and EMBOSS CpGPlot/CpGReport/Isochore). Methylation status of 106 CpG islands located at Chr6: 132271726–132272591 (according to UCSC GRCh37/hg19) was evaluated by sequencing of bisulfite-modified DNA fragment amplified by following primers (Supplementary table 3). Genomic DNA, from A549 and Calu-1 cells, cultured in the absence or in the presence of 10 μM 5-dAzaC for 96 h and from lung cancerous and histopathologically unchanged tissues of 98 NSCLC patients, was isolated using DNA Mammalian Genomic Purification Kit, Sigma-Aldrich Co. (St. Louis, MO). Next, 500 ng of genomic DNA was subjected to bisulfite conversion according to EZ DNA Methylation Kit™ protocol, Zymo Research Corporation (Orange, CA).

Methylation-sensitive high-resolution melting analysis (MS-HRM) was used as a screening method for the detection if there are any differences in DNA methylation patterns between lung cancerous and histopathologically unchanged tissues from NSCLC patients. Methylation level of DNA fragment located within the CpG island of CTGF gene was determined by RT-PCR amplification of bisulfite-modified DNA, followed by HRM profile analysis. The reaction was conducted in a Light Cycler^®480 Real-Time PCR System, Roche Diagnostics GmbH (Mannheim, Germany). The CTGF region containing 15 CpG dinucleotides located at Chr6: 132271312–132271581 (according to UCSC GRCh37/hg19) was amplified by a pair of primers complementary to the bisulfite-DNA-modified sequence (Supplementary table 3). For PCR amplification, 1 μl of the bisulfite-treated DNA from patients was added to 10 μl of 5× HOT FIREPol^® EvaGreen^® HRM Mix, Solis BioDyne Co. (Tartu, Estonia) with 0.2 μM primers. Each reaction was performed in triplicate. MS-HRM analysis was performed using Light Cycler^®480 Gene Scanning software and TM Calling software. The methylated and unmethylated DNA acquires different sequences after bisulfite treatment resulting in different melting profiles of PCR products. We analyzed the shape of achieved melting curves among lung cancerous and histopathologically unchanged tissues, and when differences were detected, samples were targeted for bisulfite sequencing.

For bisulfite sequencing, bisulfite-modified DNA fragment was amplified with FastStart Taq DNA Polymerase from Roche Diagnostics GmbH (Mannheim, Germany). The PCR products were separated on agarose gel, purified using Agarose Gel DNA Extraction Kit, Roche (Mannheim, Germany), and cloned into pGEM-T Easy Vector System I, Promega (Madison, WI). After overnight ligation, competent TOPO10 E. coli strain cells were transformed with plasmids. Plasmid DNA isolated from five positive bacterial clones was used for commercial sequencing. The results of bisulfite sequencing were presented using BiQ analyzer software and the Bisulfite sequencing Data Presentation and Compilation (BDPC) web server, respectively (Bock et al. 2005; Rohde et al. 2009).

The normality of the observed patient data distribution was assessed by Shapiro–Wilk test. Parametric unpaired, two-tailed t-test was used to consider statistically significant differences of CTGF mRNA and protein levels between lung cancerous and histopathologically unchanged tissues (p < 0.05). Multivariate regression was performed to detect association between histological type of cancer, smoking history and CTGF mRNA and protein levels in cancerous tissues. Parametric unpaired, two-tailed t-test and ANOVA with post hoc RIR Tukey’s test were used to compare normally distributed variables between groups. Otherwise, Kruskal–Wallis test was performed. Data groups for cell lines were assessed by ANOVA to evaluate whether there was significance (p < 0.05) between the groups. For all experimental groups, which fulfilled the initial criteria, individual comparisons were made by post hoc Tukey’s test with the assumption of two-tailed distribution. Data are expressed as the mean ± SD. Statistical analysis was performed with STATISTICA 12 software.

In the present study, employing RT-qPCR and Western blotting, we found significantly lowered levels of CTGF transcript (p < 0.0000001) and protein (p < 0.0000001) in lung cancerous tissues compared with adjacent histopathologically unchanged tissues obtained from 98 patients with NSCLC (Fig. 1a–c). Of the 98 pairs, only 9 pairs (~9 %) presented higher amount of CTGF mRNA level in tumor tissue, 13 (~13 %) pairs presented higher CTGF protein content in cancerous tissue, and in 6 pairs (~6 %), CTGF mRNA level did not correspond to the protein level. To clarify the clinical significance of CTGF expression in NSCLC, we showed the differences in CTGF mRNA and protein levels between lung cancerous and histopathologically unchanged tissues including various clinicopathological features (Tables 1, 2; Supplementary tables 1 and 2). A substantial decrease in CTGF mRNA levels was detected in lung cancerous tissues, regardless of gender (males and females: p < 0.0000001), patients age (age ≤ 60 years: p < 0.000001; age > 60 years: p < 0.0000001) and histological type of NSCLC (ADC: p < 0.0000001; SCC: p < 0.000001; LCC: p = 0.0007; carcinoid: p = 0.005; Table 1). Those differences were statistically significant in all LC stages except stage 0 (p = 0.08). As shown in supplementary table 1, lowered CTGF transcript levels in lung cancerous tissues compared with matched normal specimens were associated with almost all tumor sizes, with the exception of Tis and T4 and with all grades of lymph node metastasis. Western blotting results showed that significantly lower amount of CTGF protein in lung cancerous tissues was detected at both genders (males: p < 0.0000001; females: p = 0.02) and in ADC (p = 0.005), SCC (p = 0.00001) and LCC (p = 0.01) histological subtypes of NSCLC (Table 2). Moreover, the decreased amount of CTGF protein was associated with T1b (p = 0.02), T2a (p = 0.000002) and T3 (p = 0.0007) tumor size and with all grades of lymph node metastasis (N0: p = 0.000004; N1: p = 0.01; N2: p = 0.03; Supplementary table 2). However, the difference in CTGF protein level between cancerous and histopathologically unchanged tissues did not reach statistical significance in the group of patients under 60 years of age, in patients with recognized carcinoid and with LC stage 0 and 2B (Table 2). We cannot consider the association of CTGF expression level with distant metastasis as 95 patients presented no distant metastasis (M0). We performed multivariate analysis to investigate the association of CTGF mRNA and protein levels with a history of smoking and histological type of LC. All models were weak, with an adjusted R ² < 0.7. We did not find a significant association of CTGF expression level with smoking status. However, we observed that in the group of cancerous tissues CTGF protein level was associated with histological type of LC (p = 0.013). The highest amount of CTGF protein was detected in carcinoid tumors and the lowest amount in ADCs. We also analyzed the relationship between CTGF transcript and protein levels in cancerous tissues and different clinicopathological parameters (Table 3). We found that there was significant difference in CTGF protein levels among histological types of LC (p = 0.047). The amount of CTGF protein was significantly decreased in ADCs compared with carcinoid tumors (p = 0.026). We did not identify any statistically significant difference between CTGF transcript and protein levels and other clinicopathological variables such age, sex, LC stage, tumor size, lymph node metastasis or smoking (Table 3).

In order to complement our Western blotting results and to define the distribution and localization of CTGF protein in investigated patients’ material, representative NSCLC and non-tumoral lung specimens were immunohistochemically stained with anti-CTGF Ab (Fig. 2). Positive CTGF staining was found in normal lung tissues, with strong immunoreactivity in macrophages and in respiratory epithelium (Fig. 2a, b). SCC cells showed no or very weak cytoplasmic staining (Fig. 2c, d), whereas ADC cells were all negative for CTGF staining (Fig. 2e, f). However, the presence of CTGF protein was detected in tumor-infiltrating macrophages, stromal fibroblasts and normal bronchial epithelial cells located in the tumor field and was used as an internal positive control (Fig. 2c–f). Representative hematoxylin- and eosin-stained slides are shown in Supplementary figure 1.

Basal expression patterns of CTGF in investigated cells were determined by RT-qPCR, and the amount of CTGF protein was detected by Western blotting, respectively (Fig. 3a). A549 and Calu-1 cells presented definitely lowered content of CTGF mRNA in comparison with Beas-2B cell line. We observed that CTGF mRNA levels were about 7.5 times lower in A549 cells and 3.4 times lower in Calu-1 cells than in Beas-2B cells. The amounts of CTGF protein were consistent with the RT-qPCR results (Fig. 3a). However, the content of CTGF transcripts and protein in NSCLC cells was higher than in T47D, which is known to express CTGF at very low levels (Chen et al. 2007b). CTGF mRNA and protein levels detected in A549 cells were reduced more than twofold in comparison with those from Calu-1 cells. As expected, these results confirmed that the expression level of CTGF in NSCLC cells is diminished.

In order to determine whether the expression of CTGF can be epigenetically regulated in NSCLC cells, we investigated whether DNMTs inhibition by 5-dAzaC would exert any effect on the CTGF mRNA level. After 96 h of incubation with 5-dAzaC at concentrations of 10 and 15 μM, we observed an elevation of CTGF transcript levels in A549 and Calu-1 cells (Fig. 3b). However, this induction was statistically significant only for Calu-1 cell line (p < 0.001). We noticed an approximately twofold increase in CTGF mRNA levels in A549 cells and 2.5-fold increase in Calu-1 cells. The effect was not dose dependent as both concentrations of 5-dAzaC up-regulated CTGF expression to the similar level in each cell line. Next, we examined whether RT-qPCR results would correspond to CTGF protein content. The densitometric analysis of Western blotting bands revealed that 5-dAzaC (10 μM) was able to raise the CTGF protein amount in investigated cells after 72 and 96 h of incubation as compared to respective controls (Fig. 3c). The double bands (36 and 38 kDa) of full length, intact CTGF were detected (Zarrinkalam et al. 2003), and both were markedly increased after 5-dAzaC treatment. Moreover, we investigated whether the same effect, exerted by 5-dAzaC, would be observed in Beas-2B cells. We revealed that 5-dAzaC (10 uM) was also able to elevate CTGF mRNA and protein levels in Beas-2B cell line after 48, 72 and 96 h of treatment (Supplementary figure 2).

To determine whether concentrations of 5-dAzaC used in previous experiments do not exert a pronounced cytotoxic effect in investigated cell lines, we applied trypan blue exclusion assay for the measurement of cell viability. We showed that 5-dAzaC at concentrations of 10 and 15 µM reduced cell viability in a dose-dependent manner in both A549 and Calu-1 cell lines. The results were statistically significant at each time point (Fig. 4a, b). However, after 48, 72 and 96 h of incubation with 5-dAzaC (10 µM), A549 and Calu-1 cell viability was higher than 90 % compared with respective controls. The second dose of 5-dAzaC (15 µM) was more effective in reducing cell viability, although the amount of viable A549 and Calu-1 cells was still not lower than 80 %. On the contrary, both concentrations of 5-dAzaC were extremely potent in reducing the proliferation rate of A549 and Calu-1 cells. The cell growth inhibition was statistically significant and occurred after 72 h of incubation, for both cell lines, at both concentrations of 5-dAzaC (p < 0.001; Fig. 4c, d). Cell number counting by trypan blue assay revealed that after 96 h, 5-dAzaC (10 and 15 µM) reduced the proliferation of A549 cells by approximately 51 and 70 %, respectively (Fig. 4c). Both concentrations of investigated compound exhibited the same effect in Calu-1 cells at indicated time point, diminishing the proliferation rate by 54 % compared with untreated controls (Fig. 4d).

Because 5-dAzaC was able to up-regulate CTGF mRNA and protein levels in A549, Calu-1 and Beas-2B cells, we decided to investigate whether those changes were caused by direct demethylation of the CTGF CpG-rich regulatory region. For that purpose, we incubated cells either in the absence or in the presence of 5-dAzaC (10 µM) for 96 h and then we examined the methylation status of 106 CpG islands within CTGF regulatory region (Chr6: 132271726–132272591) by bisulfite sequencing. Surprisingly, for both A549 and Calu-1 cells, no methylation in investigated region was detected (Fig. 5). The same result was obtained for Beas-2B cell line (Supplementary figure 3). These data suggest that 5-dAzaC is able to restore CTGF expression in NSCLC cells and in normal bronchial epithelial cells; however, it does not act through direct demethylation of CTGF regulatory region. We also evaluated the DNA methylation status of CTGF in lung cancerous and in matched normal tissues from 28 patients with NSCLC. We used MS-HRM analysis as a screening method in order to detect whether there are any differences in methylation status in CTGF regulatory region between lung cancerous and matched normal tissues in the group of 98 patients. We found 56 matched samples with different melting profiles of PCR products (Supplementary figure 4), and we further analyzed them by bisulfite sequencing. We detected different patterns of DNA methylation in examined region (Chr6: 132271726–132272591) between histopathologically unchanged tissues and lung cancerous tissues in the group of 28 NSCLC patients (Fig. 6a, b). Among those patients, 16 had SCC, 9 had ADC, 2 had LCC and 1 was diagnosed with carcinoid tumor. Changes in DNA methylation patterns between normal and cancerous tissues were not statistically significant (p = 0.08); however, this result is preliminary and further studies, on the larger group of patients, are needed to estimate whether DNA methylation plays any role in CTGF silencing in NSCLC.

Next, we investigated whether other epigenetic mechanism—histone acetylation, could restore the expression of CTGF in NSCLC cell lines. We chose one of the most potent HDACs inhibitor—TSA, and tested its impact on A549 and Calu-1 cell viability and proliferation, in order to estimate appropriate concentrations for further experiments. Different doses of TSA (30, 100, 300 and 900 nM) were applied to cell cultures for 12, 24, 48 and 72 h, and cell viability was evaluated by trypan blue staining and compared with ethanol-treated controls. Treatment with 900 nM of TSA markedly decreased A549 cell viability (41 % of controls after 48 h and 24 % of controls after 72 h) as well as Calu-1 cell viability (77 % of controls after 48 h and 53 % of controls after 72 h), and this concentration was excluded from further analysis (Fig. 7a, b). Moreover, we observed that A549 cell line is more vulnerable to TSA as significant cell morphology changes were detected after exposure to 900 nM of this compound. Other doses of TSA did not reduce cell viability to such extent and were used in other experiments. After 72 h of incubation with 300 nM of TSA, A549 cell viability was higher than 55 % and Calu-1 cell viability was higher than 80 % (Fig. 7a, b). Moreover, TSA appeared to be a highly potent inhibitor of NSCLC cells proliferation. The growth rate of A549 and Calu-1 cells after TSA treatment was significantly diminished in a dose and time-dependent manner compared with respective controls (Fig. 7c, d).

We showed that TSA is able to regulate the expression of CTGF in A549 and Calu-1 cells. After treatment with increasing doses of TSA, we observed a progressive elevation of CTGF transcript levels in A549 cell line at each time point (Fig. 8a). Although the upward trend was visible for each concentration of TSA, statistically significant differences in CTGF mRNA levels between A549 treated and control cells were detected at 300 nM of TSA (p < 0.001; Fig. 8a). On the contrary, in Calu-1 cells only the highest tested dose of TSA (300 nM) resulted in an increase in CTGF transcript levels at each time of incubation (p < 0.001; Fig. 8b). Changes in CTGF transcript levels, induced by TSA, in both NSCLC cell lines were correlated with an elevation of CTGF protein amount (Fig. 7c, d). The densitometric analysis of Western blotting bands showed a progressive increase in CTGF protein levels after TSA treatment in A549 cells, in a dose-dependent manner at 12, 24, 48 and 72 h of incubation (Fig. 8c). The CTGF protein content in Calu-1 cells incubated with 300 nM of TSA was significantly elevated as compared to the respective controls at each time of incubation (Fig. 8d). Taken together, we showed that although A549 and Calu-1 cells vary in the sensitivity to TSA, this compound is able to restore the expression level of CTGF in those cell lines.