Blood samples are rich in biomolecules such as proteins, DNA and RNA, which can be used for disease diagnosis, and determining disease stage and prognosis . Compared with tissue samples, blood samples have advantages including being more readily accessible, suitable for continuous sampling, and amenable to numerous different tests. However, after blood samples are collected, they need to be processed, packed and stored in containers suitable for long-term cryopreservation. Otherwise, RNA in blood is prone to degradation under normal temperature transportation and daily storage conditions, which can affect the subsequent detection, analysis and development of molecular diagnostic biomarkers .
In clinical diagnostic laboratories, 60−70% of errors occur in the pre-processing process of specimen analysis, most of which are caused by specimen collection, specimen processing and specimen storage . As a basis for biomarker research and development, the acquisition of high quality blood samples is critical. The dipotassium ethylene diamine tetraacetic acid (EDTA-2K ) anticoagulant vessels are widely used in basic research and clinical laboratories to collect, transport and preserve blood samples, which is not conducive to protecting specific molecular biomarkers such as RNA molecules in blood. In the process of sample preservation, if a sample is immediately frozen at -20°C or -80°C without any protective measures, subcellular structures are severely damaged, and the structure and activity of RNA can be altered, often irreversibly . The quality of RNA in stored samples is directly affected by the method of sample storage . In the case of improper storage, RNA in blood is rapidly degraded by RNase enzymes present in blood or the external environment. Previous research has shown that although EDTA-K2 can effectively prevent hemagglutination, it cannot maintain the stability of RNA in blood cells. Following cryopreservation, RNA is degraded by about 80% in a short time . Therefore, the present work mainly compared the stability of messenger RNAs (mRNAs) and some non-coding RNAs (ncRNAs) in blood, explored the influence of relevant experimental operations on RNA degradation before and after RNA extraction, and quantitatively analysed on mRNA degradation in specimens and during experimental processes.
The miRNAs are short (~ 22 nucleotides), non-coding, RNA molecules that control diverse biological processes, including cell fate determination, cell proliferation, cell differentiation and cell death [7–9]. The miRNAs regulate gene expression post-transcriptionally by interacting with and down-regulating target mRNA molecules. There has been a growing interest in their use for clinical applications, largely due to their high stability and cell/tissue specificity. They are relatively stable and widely expressed in cells, tissues and organs throughout the body. In serum, plasma, saliva, urine, emulsions and other body fluids, miRNAs are highly stable extracellularly [10–12]. The miRNAs are associated with a variety of diseases and have been detected under a variety of pathological conditions . CircRNAs are a common type of non-coding RNA produced by reverse splicing. Reverse splicing is an unconventional splicing event characterized by non-coding RNA molecules with circular structure formed by covalent bonds without a 5 '-capped structure or a 3' -terminated polyadenylated tail. Since circRNAs are insensitive to nuclease, they are more stable than linear RNAs, but the mechanism by which cells eventually degrade circRNAs is still largely unknown compared to the mechanism of circRNA biogenesis [14, 15]. LncRNAs are non-coding RNAs with a length of more than 200 nucleotides.Studies have confirmed the existence of hundreds of lncRNAs in serum/plasma. These long-chain molecules are stable, abundant and non-degradable, easy to be quantitatively detected, and have significant disease specificity .
Although great progress has been made on studying ncRNAs biosynthesis and functions, there are few reports on their degradation . Due to the presence of nucleases in the blood, most researchers doubt that extracellular RNA can remain stable for long periods of time . For example, the miRNAs are generally considered to have a long half-life (T1/2), and can be stably present in serum and not easily degraded. Like other RNAs, miRNAs have a half-life, but exact data is rare in the literature [18, 19]. The regulation of miRNAs is critical to the definition of cell identity and behaviour in normal physiology and disease. However, the dynamics of miRNA degradation and the mechanisms involved remain largely obscure, particularly in higher organisms . Researchers developed a pulse-tracking method based on metabolic RNA markers to calculate the genome-wide miRNA attenuation rate in mammalian cells , and revealed the half-life of heterogeneous miRNAs, which are stable molecules in many species (T1/2 >24 h). However, other miRNAs, including passenger miRNAs and some guide miRNAs, are rapidly degraded (T1/2 = 4−14 h).
Insights into the mechanisms and relevance of miRNA degradation in physiology are just beginning to emerge. Some reports suggest that slow turnover might not be a general feature of miRNAs and, in some instances, miRNA decay may be relatively fast (a few hours), as demonstrated for miR-503 in fibroblasts, and miR-29b during mitosis [22, 23]. Degradation plays an important role in the regulation of intracellular ncRNA content, thereby regulating the growth and development of the body or stress responses and other processes. The determination of RNA half-life (T1/2) is important for understanding the regulatory mechanisms of gene expression and environmental changes that alter gene transcription levels .
Each method for measuring RNA levels has its advantages and disadvantages. The half-life of mRNA in tissues is generally determined by in situ hybridisation, which is time-consuming and labour-intensive . Plasma ncRNAs can be quantitatively detected by real-time fluorescence quantitative PCR and microarray. If higher sensitivity is required, real-time quantitative PCR is a good choice [9, 25]. The copy number of molecules in samples can be quickly and accurately determined by real-time quantitative PCR. The relative initial concentration is calculated based on the cycle threshold (Ct) value, and the half-life is calculated by establishing the regression linear equation over time. Due to the sensitivity of the method, the half-life of an RNA can be established even when it is expressed at low levels.
Therefore, a fast and reliable RNA half-life measurement method was established in the present study, based on real-time fluorescence quantitative PCR and supplemented by microspectrophotometry, to compare the half-life length of mRNAs and ncRNAs. Human-specific glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and β-actin genes were used as internal reference genes for mRNA stability, the U6 gene served as a reference for miRNA stability, and the hsa_circRNA_002532 and lncRNA GASL1 genes were used as reference for the stability of circRNAs and lncRNAs. Changes in RNA content in blood samples after different time periods were determined by fluorescence quantitative PCR, and the half-lives of the genes were established and assessed. Fresh, whole anticoagulant blood was collected and stored at room temperature, and RNA was extracted and measured every 12 h. Real-time quantitative PCR analysis of the genes was used to evaluate the concentrations of mRNAs and ncRNAs in each sample. We analysed various factors involved, from RNA extraction to gene amplification. We employed the orthogonal experimental method with three factors and three levels. The three factors were the storage time of fresh whole blood at room temperature, the storage time of RNA at room temperature, and the storage time of cDNA after reverse transcription at -20°C. The β-actin gene was amplified by real-time quantitative PCR and its Ct value was determined. The results were analysed by multivariate analysis of variance (ANOVA) and regression analysis.
In research on mRNA and ncRNAs molecular markers, standardisation of the entire testing process is very important for the stability and reproducibility of experimental results . Additionally, although ncRNAs have high stability, theoretically, they can still be affected by RNA-active enzymes . In clinical studies, blood samples are typically stored frozen to investigate the stability of miRNAs and establish optimal storage conditions . The present work focused on exploring the degradation of RNA at room temperature, using the same method to study the half-life of different RNAs and the influencing factors, and compare the degradation kinetics and characteristics of different RNAs in whole blood. The overall aim was to establish the optimal experimental and storage conditions, and thereby ensure the accuracy of experimental results.