Study population
All donors and patients were of Chinese origin. Informed consent was obtained from all participants. Nine type I CD36-deficient blood samples and thirteen type II CD36-deficient blood samples from healthy blood donors, comprising 76.19% men and 23.81% women with an average age of 34.19 ± 7.13 years (range, 18–55 years), were randomly collected from the Guangzhou Blood Center CD36 Deficiency Bank. Seven normal blood samples from healthy blood donors (71.43% men and 28.57% women with an average age of 35.57 ± 7.51 years, range 18–55 years), were also randomly collected at the Guangzhou Blood Center. Four blood samples from patients with type I CD36 deficiency were collected randomly from FNAIT patients or PTR patients with anti-CD36 antibodies between 2015 and 2022 at the Clinical Testing Service Department of the Guangzhou Blood Center (Table 1)8,18,19. The samples were screened for antibodies against erythrocytes using the Coombs test, and all results were negative.
Primer design, amplification, library preparation, and SMRT sequencing
The primer panel was designed to amplify regions from 5’ UTR to the 3’ UTR of CD36 genes. Sequences from intronic regions were also included. Seven sets of primers were designed to amplify amplicons with more than 1 kb overlap between each amplicon, and all the amplicons are listed in Fig. 1. All primers used in this study are listed in Supplementary Material 1. Long-range polymerase chain reaction (LR-PCR) was performed with KOD FX Neo DNA polymerase (TOYOBO) according to the manufacturer’s instructions, using PCR cycling conditions consisting of a two-step cycle of 10 min each for 30 cycles. PCR products were detected by agarose gel electrophoresis and then purified with 0.6× Ampure PB beads (Pacific Biosciences).
Genetic structure of the CD36 gene locus. This delineates the human CD36 gene’s exon-intron structure, with long-read PCR amplicon locations and the exonic locations. Untranslated regions are in red, and coding sequences in blue.
Single-molecule real-time (SMRT) libraries were prepared using a one-step method according to the manufacturer’s instructions. In this method, DNA damage repair, end repair, and adapter ligation were performed simultaneously to produce pre-sequencing libraries containing unique barcode adapters. The final library was annealed with sequencing enzymes and primers using the Sequel II Binding Kit 2.2 (Pacific Biosciences) and the DNA Internal Control Kit 1.0 (Pacific Biosciences). Then, 150 pM DNA polymerase complexes were loaded and sequenced using the Sequel II platform (Pacific Biosciences) with a 20-hour run time.
Data analysis, DNA variant calling, and haplotype phasing
SMRT Link software (v10.1.0, Pacific Biosciences) was primarily used to analyze the output data. Raw reads were first demultiplexed and barcoded automatically at the end of the sequencing runs, and then subreads were analyzed to generate CCS reads using the ccs software v.3.0.0 (https://github.com/pacificbiosciences/unanimity/). Filtered CCS reads were aligned to the human reference genome (GRCh38) using pbmm2 (https://github.com/PacificBiosciences/pbmm2). The target CCS reads were realigned to the reference genome (GRCh38) using pbmm2. Variant calling was conducted with Google DeepVariant (v1.2.0) to identify SNVs and small indels. Finally, amplicon contig sequences were phased into individual haplotypes using the Flye assembler. Finally, the output data were compiled, and individual haplotypes, including gene crossover region haplotypes, were generated using an in-house sciprt module.
Linkage disequilibrium analysis of CD36, phylogenetic tree construction, and variant association analysis
To visualize pairwise linkage disequilibrium between genetic variants and identify linkage groups within CD36, we generated LD heatmaps using LDBlockShow (v1.40) with minor allele frequencies < 0.01. The maximum likelihood phylogenetic tree of haplotypes in LD blocks was constructed using MEGA11 (v11.0.13) with 1000 bootstrap repeats. The types of CD36 deficiency in the donors were classified as 0, 1, and 2. The associations between the genotypes of each SNV and the phenotypes were tested using Fisher’s exact test. A P value < 0.005 indicated a significant difference, and the odds ratios between variants and the CD36 phenotype were calculated.
Luciferase assay
A luciferase reporter assay was conducted to investigate the potential regulatory role of the c.-132A > C mutation in the 5’ UTR on CD36 transcriptional activity. The transcriptional activity of CD36 was assessed using luciferase activity driven by the c.-132 A > C mutation. HEK293T cells were cotransfected with pGL3-basic or pGL3-CC and empty pcDNA3.1 vector plasmids in 24-well plates using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s instructions. Renilla luciferase activity was measured using the Luciferase Reporter Assay System (Promega) and a BioTek Synergy H1 microplate reader (Agilent) to determine the firefly-to-Renilla luciferase ratio.
FNAIT 1
At 23 weeks of gestation, a Chinese fetus was diagnosed with hydrops fetalis (HF) by ultrasound. An increase in the middle cerebral artery peak systolic velocity (MCA-PSV) to 58 cm/s was detected in the fetus. His mother, a 29-year-old (Luo, the number is 23N00901, Supplementary Material 2) with no history of blood transfusion or transplantation, had experienced three pregnancy losses due to intrauterine fetal death (IUD) or hydrops fetalis (HF). Direct and indirect antiglobulin assays were used to test for irregular antibodies against erythrocytes in the maternal serum. Targeted analysis test results for pathogenic variants in the thalassemia genes (HBA1, HBA2, HBB) were also negative. The long-read sequencing of maternal CD36 gene reveals that both of her alleles carried nonsense SNVs or in/dels (c.329_330delAC on one allele and c.1006 + 2 T > G on the other). Both alleles also carried the c.-132 A > C mutation in the 5’-UTR. FACS analysis of CD36 surface antigen expression on platelets and monocytes revealed type I CD36 deficiency in this patient (Table 2).
FNAIT 2
The 27-year-old mother with type I CD36 deficiency had no history of blood transfusions or transplants and experienced a miscarriage detected by ultrasound at 23 weeks of gestation due to HF. The middle cerebral artery peak systolic velocity (MCA-PSV) at 22 weeks of gestation during the second pregnancy was 60 cm/sec. Both direct and indirect antiglobulin assay results were negative for abnormal antibodies against red blood cells in the maternal serum. Targeted analysis test results for pathogenic variants in the thalassemia genes (HBA1, HBA2, and HBB) were also negative. Each allele of her CD36 gene carried a nonsense SNVs or in/dels (c.329_330delAC on one and c.430-1G > C on the other one ). Both alleles also carried the c.-132 A > C mutation. Anti-CD36 antibodies were detected in both fetal and maternal blood serum (Table 2). The patient gave birth to a premature baby girl at 32 weeks of gestation.
Flow cytometry analysis of anti-CD36 antibodies from fetal serum using transfected HEK293T cells
The Flow cytometry analysis assay was conducted utilizing transfected HEK293T cells as previously described4. Aliquots comprising 10^5 cells and 25 µL of fetal serum were incubated at 37℃ for 30 min. Unbound antigen-antibody complexes were eliminated through washing, Then, 50 µL of anti-human IgG (or IgM or IgG + IgM + IgA)-FITC was added and analyzed by Flow cytometry.
Pairwise linkage disequilibrium between variants and linkage groups in CD36 with LD patterns. The approximately 35 kb region of the CD36 gene, spanning from the 5’ UTR to the 3’ UTR, is divided into the 5’ block (front part) and the 3’ block (tail part). These blocks are separated by a breakpoint around intron 3 to intron 4. The above points indicated the Fisher’s exact test between genetic variants and observed phenotypes (Type I/II or healthy). A stronger association was found between the variants in the 5’ block and type I deficiency.
Ethics approval and consent to participate
Informed consent was obtained from all enrolled subjects and protocol was approved by the Ethics Committee of Guangzhou Blood Center (ethics approval No. GZBC-2022-052). All experiments were performed in accordance with relevant guidelines and regulations.
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- Source: https://www.nature.com/articles/s41598-024-74917-0