Samples conditions

Fifty blood samples were collected from children diagnosed with Acute Lymphoblastic Leukemia (ALL), while an additional 50 samples were obtained from healthy children, serving as the control group. All patients received appropriate medical care, supervision, and a precise diagnosis of ALL. These blood samples were subsequently analyzed for various indicators of ALL, including white blood cell count, blast cell count, C-reactive protein (CRP) levels, and interleukin-6 (IL-6) levels.

Counting white blood cells

A 50 µL sample of collected blood was added to 1000 µL of a 2% filtered acetic acid solution in a 12 × 75 labeled tube, resulting in a dilution factor of 20. The acetic acid lysed the red blood cells, leaving the white blood cells (WBCs) and nucleated red blood cells (NRBCs) intact. Methylene blue powder was then added to the diluent to stain the nuclei of the WBCs. A 10 µL aliquot of the diluted blood was placed on each side of the hemocytometer, and the cells were allowed to settle for 2 min. The count was performed using a 10X objective, and all WBCs in the four large corner squares on both sides of the hemocytometer were counted. The total WBC on each side was presented as WBC per microliter.

Platelet count assay

Platelet counting was performed using the BD Accuri™ C6 flow cytometer. In this method, 100 µL of EDTA-anticoagulated whole blood was labeled with antibodies targeting specific epitopes on the glycoprotein IIb/IIIa complex of both resting and activated platelets (anti-CD 41) at a 1:1000 dilution, followed by incubation at room temperature for one hour. The total sample volume was then adjusted to 500 µL with PBS. The samples were analyzed by flow cytometry at a flow rate of less than 4,000 events per second, which minimizes the need for adjustments related to coincident platelet/RBC or RBC/RBC events. To determine the RBC/platelet ratio, at least 1,000 platelet events and 50,000 RBC events were collected per sample. Events that displayed both RBC scatter signals and platelet fluorescence were considered as RBC-platelet coincidence events. The platelet count was then calculated using this ratio and the RBC concentration of the original blood sample, which was determined by impedance counting [17].

Immunotyping assay in ALL

Flow cytometry was used to assess CD10 expression as a marker for precursor B-cells in ALL. For each assay, 1 mL of peripheral whole blood was collected in 1.5 mL tubes. Samples were centrifuged at 1500 rpm for 3 min to separate plasma from red blood cells. A volume of 100 µL of plasma was transferred into clean tubes and processed following the manufacturer’s instructions, except for the lysis buffer, which was substituted. Within 1 h of collection, samples were incubated with mouse anti-human CD10-FITC–conjugated primary antibody (332775, BD Biosciences, CA, USA) for 30 min at room temperature in the dark. Following incubation, 400 µL of PBS was added, and the samples were gently inverted before acquisition on the Accuri C6 flow cytometer [18].

C reactive protein (CRP) assay

For the CRC detection protocol, an ELISA assay (SEA821Hu, Cloud-Clone Corp, USA) was conducted using blood serum, which was separated from the collected samples using standard laboratory methods with serum-separator Vacutainers. One milliliter of each sample underwent cooling centrifugation for 20 min at 10,000 × g to clarify any lipemic specimens. The resulting suspension was then sonicated with an ultrasonic cell disrupter until the solution became clear. Afterward, the homogenates were centrifuged for 5 min at 10,000 × g, and the supernatant was collected. The cells were precipitated by cooling centrifugation at 10,000 × g for 10 min and washed with PBS under the same centrifugation conditions. The cells were then resuspended in fresh lysis buffer at a concentration of approximately 10⁷ cells/mL. The lysed cells were subsequently centrifuged at 1,500 × g for 10 min at 4 °C to remove cellular debris, then resuspended in 500 µL PBS. Finally, the samples were loaded into an ELISA reader in four replicates. By using different dilutions of standard reagents, the final concentration of CRP was determined based on the standard curve, with measurements taken at a wavelength of 450 nm [19].

Enzyme-linked immunosorbent assay (ELISA)

The ELISA assay was used to measure the levels of IL-6, IL-27, and tumor necrosis factor alpha (TNF-α) by employing human ELISA kits from Invitrogen (USA). A 100 µL sample was added to each well of an ELISA plate and incubated for 2 h at room temperature (RT) with 100 µL of the control solution and 50 µL of a 1X biotinylated antibody. Following this incubation, 100 µL of a 1X streptavidin-HRP solution was added to each well and incubated in the dark for 30 min. Next, 100 µL of the TMB chromogen substrate solution was added, and the incubation continued for 15 min at RT, protected from light. The reaction was then stopped by adding 100 µL of stop solution. Absorbance was measured at 570 nm for each sample [20, 21].

DNA isolation and methylation analysis

Genomic DNA was extracted from blood samples of ALL patients and healthy children using a DNA isolation kit (Qiagen, USA). Methylation changes in the promoter regions of LAMP1 and LAMP2 were analyzed in all samples using a sodium bisulfite conversion method. Specific primers were designed for each promoter region: wild type primers that detect methylated regions and modified primers that detect unmethylated regions. One microgram of purified DNA was treated with 1 M sodium bisulfite and 10 µL of DNA-protecting buffer, bringing the total volume to 25 µL, using RNase- and DNase-free water. The mixture was denatured at 95 °C for 5 minutes, then incubated at 50 °C for 7 hours to convert unmethylated cytosines to uracil. This conversion process was repeated three times using a thermal cycler (BIO-RAD, USA) to ensure complete conversion. The treated DNA was then used for conventional PCR amplification of the promoter regions, employing specific primers to detect both methylated and unmethylated fragments. The wild type primers used were: LAMP1-F-5’-AACGCCAGCCCTTGGCGCCCGC-3’ and LAMP1-R-5’-ATGGCGCGAGGCGGCCGGGTACG-3’. The modified primers used were: LAMP1-M-F-5’-ACGCCAGCCCTTGGCGCCC-3’ and LAMP1-M-R-5’-ATGGAGCGAGGAGGCAGGGT-3’. The PCR conditions were as follows: an initial denaturation at 95 °C for 10 min, followed by 40 cycles of 95 °C for 30 s, 60 °C for 30 s, and 72 °C for 45 s. The PCR products were separated on a 1% agarose gel in 1X-TBE buffer, and visualized under UV light (320 nm) using a gel documentation system. Methylated fragments appeared as a single band at 80 bp, while unmethylated fragments were represented as a single band at 90 bp [4, 22].

To further evaluate the methylation status of the promoter regions in the selected genes, the genomic DNA was subjected to digestion with the methylation-sensitive restriction enzyme HpaII. HpaII cleaves unmethylated cytosine residues within its restriction site (CCGG). The digestion was performed using 5 units of enzyme per 200 ng of DNA for 4 h at 37 °C. After digestion, the DNA was used for quantitative RT-PCR (qRT-PCR) analysis of the LAMP1 and LAMP2 gene promoter regions, employing the previous wild-type specific primers for each region. The qRT-PCR was carried out using the QuantiTect SYBR Green PCR Kit (Qiagen, USA), with GAPDH promoter primers as a housekeeping gene for normalization. The PCR conditions were as follows: an initial denaturation at 95 °C for 5 min, followed by 35 cycles of 95 °C for 30 s, 60 °C for 30 s, and 72 °C for 30 s, with a final extension at 72 °C for 10 min to complete the amplification [5, 23].

qRT-PCR procedure for gene expression

Total RNA was extracted from the derived samples using the TriZol reagent (Invitrogen, USA), along with chloroform and isopropanol, following the standard protocol. The extracted RNA was then purified using an RNA purification kit (Invitrogen, USA) according to the manufacturer’s instructions. The purified RNA was utilized to synthesize complementary DNA (cDNA) using the QuantiTech Reverse Transcription Kit (Qiagen, USA) following the provided protocol. The relative expression levels of DNMT1, DNMT3, methionine synthases (MS), ten-eleven translocation (TET1), LAMP1, LAMP2, ATG5, and LC3B were quantified in patient samples and normalized to their expression in control samples using specific primers (refer to Table 1). PCR amplification of the targeted gene fragments was carried out using the QuantiTect SYBR Green PCR Kit (Qiagen, USA), along with GAPDH, which served as the housekeeping gene control. The PCR machine was programmed with the following conditions: 94 °C for 5 min, followed by 40 cycles of 94 °C for 30 s, 60 °C for 30 s, and 72 °C for 45 s. The relative expression of each gene was calculated by normalizing the cycle threshold (Ct) value of each target gene to GAPDH and adjusting between patient and control samples. The relative gene expression levels were then determined using the delta-delta Ct method and are presented as fold changes [24, 25].

Table 1 Oligonucleotide sequences employed for the quantitative analysis of the specified genes using qRT-PCRFlow cytometric protein profiling

To assess the kinetic expression of LAMP1, LAMP2, ATG5, and LC3B in the collected samples, a flow cytometry-based analysis was performed. Initially, blood samples were centrifuged at 1500 rpm for 3 min to separate plasma from red blood cells. The plasma was then transferred to clean tubes and subjected to a second centrifugation at 1500 ×g for 5 min. Following this, the supernatant was discarded, and the resulting cell pellet was washed with phosphate-buffered saline (PBS), centrifuged again, and subsequently fixed in 2% formaldehyde prepared in PBS. For cell permeabilization, the fixed cells were incubated for 3 min in PBS containing 0.1% Triton X-100. Primary antibody staining was carried out by incubating the permeabilized cells overnight at 4 °C in PBS with 1% bovine serum albumin (BSA). The primary antibodies used were: rabbit monoclonal anti-LAMP1 (MA5-29385, ThermoFisher Scientific, USA), rabbit polyclonal anti-LAMP2 (BS-2379R, ThermoFisher Scientific, USA), and rabbit polyclonal anti-LC3B (ab51520, Abcam, USA), each applied at a 1:100 dilution. After overnight incubation, cells were washed and then incubated for 2 h at room temperature with a secondary antibody, Alexa Fluor 488-conjugated goat anti-rabbit IgG (ab150077, Abcam, USA), diluted 1:500 in PBS containing 1% BSA. Finally, the cells were resuspended in 1 mL of PBS and analyzed using a BD Accuri C6 Plus flow cytometer. Protein expression levels were detected using the FITC channel. LAMP1 and LAMP2 signals primarily appeared in the lower-right quadrant of the flow cytometry plots, represented by blue fluorescence. Similarly, LC3B expression was detected using the same fluorescence settings and visualized as red signals in the corresponding quadrant. Cells with low fluorescence intensity consistently appeared in the lower-left quadrant, indicating low or negligible expression of the target proteins [26, 27].

Statistical analysis

Delta-delta Ct analysis was used to evaluate relative gene expression, represented as fold changes in steady-state mRNA levels. Ct values for the housekeeping gene GAPDH served as the reference for normalization. A two-tailed t-test was conducted to assess differences in the data, with P < 0.05 indicating statistical significance (*) and P < 0.01 denoting high statistical significance [28, 29].