Reverse Transcription Applications

One of the most common applications of quantitative RT-PCR (RT-qPCR) is measurement of mRNA levels over time, across cells and tissues, or after an event (e.g., treatment). Due to higher sensitivity than RT-PCR, RT-qPCR is also widely used to examine the presence of retroviruses (RNA viruses). Similar to the RT-PCR workflow, RNA is first converted to cDNA, which is then amplified by PCR. The main difference, however, is that levels of amplified cDNA are measured by fluorescence in real time during the exponential phase of amplification. The amplification level is used as a basis to quantitate the original targets within the RNA population.

Learn more: Quantitative PCR
Blog post: Understanding C_t values in real-time PCR

The accuracy of the quantitation of gene expression by RT-qPCR depends heavily upon the quality and quantity of cDNA templates. Thus, the reverse transcription step is critical for success in RT-qPCR. The reverse transcription step should generate cDNA products that are representative of the original RNA population. The reverse transcriptase selected should therefore be able to synthesize cDNA efficiently, even with low-abundance genes and suboptimal and/or challenging RNA samples (e.g., high GC%, inhibitor presence, degradation).

Learn more: Reverse transcriptase properties
Application note:  Improved RT-qPCR analyses of plant samples
Application note:  Improved RT-qPCR analyses of whole-blood RNA samples

In addition to a highly efficient reverse transcriptase, there are a number of considerations in choosing reagents for the RT reaction. First, the dynamic range or linear amplification of cDNA over a broad range of input RNA is critical. The ability to obtain cDNA yields proportional to the amounts of input RNA ensures accurate quantitation of gene expression (Figure 2).

White paper:  Engineered reverse transcriptase

Figure 2. Linearity of qPCR results after using RT master mixes across a range of total RNA input, for detection of (A) high-abundance and (B) low-abundance RNA targets. RNA input, ranging from 10 pg to 1 μg, was reverse-transcribed and subsequently amplified by PCR. Both master mixes generated cDNA proportional to the input RNA, but a higher yield was obtained from Master Mix 1 as indicated by lower (i.e., earlier) C_t values, especially with the low-abundance gene target.

Furthermore, the reagents selected should produce abundant and consistent cDNA yields among replicates in order to obtain gene expression results with high sensitivity and little variability (Figure 3). A single-tube master mix containing all necessary components for reverse transcription helps minimize experimental variation, cross-contamination, and pipetting errors.

White paper:  Improved RT-qPCR master mix

Figure 3. Sensitivity and variability of qPCR results subsequent to using different RT master mixes, to detect (A) high-abundance and (B) low-abundance RNA targets. Among the reagents, Master Mix 1 produces qPCR results with the lowest average C_t and standard deviation from 30 experimental replicates, demonstrating the importance of reverse transcription reagent choice for reliable gene expression analysis.

One special procedure of RT-qPCR is direct reverse transcription from crude cell lysates without RNA isolation [1]. In experiments focusing on rare cells or events, using scarce samples, or selecting specific cells within populations, direct RT-qPCR may be considered to prevent potential sample loss and low RNA recovery. In the direct procedure, it is critical to inhibit endogenous RNases that would degrade RNA and to remove cellular genomic DNA during cell lysis. Sample preparation can be completed in as little as seven minutes while providing signals from only a single cell or cell lysates using kits optimized for low abundant targets. Highly processive reverse transcriptases are especially suited for reverse transcription of unpurified RNA extracts, because of their resistance to inhibitors and high sensitivity.

One of the first applications of reverse transcriptase in molecular biology was the construction of cDNA libraries [2–4]. A cDNA library consists of cDNA clones that represent the transcribed sequences within a specific sample. Therefore, a library provides information about the temporal and spatial expression of genes for a given cell type, organ, or developmental stage, for example. The cDNA library clones are used in the characterization of novel RNA transcripts, determination of gene sequences, and expression of recombinant proteins.

Essential in constructing cDNA libraries is the proper representation of RNAs in their full length and/or their relative abundance, making the selection of a reverse transcriptase extremely important. Highly processive reverse transcriptases can synthesize long cDNAs as well as capture low-abundance RNAs. Similarly, reverse transcriptases with increased thermostability are recommended for reverse-transcribing RNA with a high degree of secondary structure.

Learn more: DNA library construction
White paper:  Engineered reverse transcriptase

After reverse transcription, a number of approaches may be used to insert cDNA into a vector for cloning. The double-stranded cDNAs after second-strand synthesis often have blunt ends and can be cloned into blunt-ended vectors (Figure 4A). Although this approach involves fewer steps, blunt-end cloning may result in less efficient ligation and loss of directionality after insertion.

Learn more: Cloning workflow

Alternatively, cDNA ends may be modified to include additional nucleotides of known sequences. For example, to modify the 5′ end of cDNA, oligo(dT) primers with additional 5′ nucleotides can be used to initiate reverse transcription; to modify the 3′ end, short DNA oligos called linkers or adapters with desired sequences may be ligated (Figure 4B). In this manner, sites for directional insertion (e.g., restriction and homologous recombination), promoter binding (e.g., T3 and T7 sequences), and affinity purification (e.g., biotin and His tags) can be readily incorporated into the cDNA sequence.

Figure 4. Common methods to clone cDNA. (A) Double-stranded cDNA with blunt ends may be cloned directly into a blunt-end cloning vector. (B) For directional cloning, cDNA ends can be modified with unique sequences compatible with the vector. (C) Ligation-independent cloning may be performed with complementary terminal sequences to enhance the efficiency of insertion. (D) Gene-specific cloning via PCR may be considered when the insert’s sequence is known.

In another popular strategy, the 3′ ends of cDNA inserts and vectors are enzymatically extended with complementary homopolymeric tails. Using terminal deoxynucleotidyl transferase (TdT) and a single dNTP, a string of 20–30 nucleotides can be added to an insert, and a similar string of complementary nucleotides added to a vector (e.g., Cs on the insert and Gs on the vector), enabling the vector and insert tails to anneal to each other (Figure 4C). Ligation is not required because the gaps are repaired inside the bacteria after transformation.

When the target sequence is known, the insert may be generated by RT-PCR for cloning of a specific region of a cDNA (Figure 4D).

Learn more: PCR cloning

Rapid amplification of cDNA ends (RACE) is a PCR-based method for determining unknown sequences at the 5′ and 3′ ends of cDNA [5]. These methods are commonly known as 5′ RACE and 3′ RACE, respectively. The experimental goals of RACE include identification of 5′ and 3′ untranslated regions, investigation of heterogeneous transcriptional start sites, characterization of promoter regions, determination of complete cDNA sequences, and sequencing of complete open reading frames (ORFs) for protein expression.

PCR with single-sided specificity (also known as one-sided or anchored PCR [6,7]) is employed to amplify the unknown regions of cDNA as RACE products. 5′ RACE relies on extension of the 5′ end with an oligonucleotide for PCR primer binding, while 3′ RACE takes advantage of the poly(A) tail of mRNA as a generic priming site for PCR (Figure 5).

In 5′ RACE (Figure 5A), mRNA of a specific sequence or related family is reverse-transcribed into the first-strand cDNA using a gene-specific primer. The 3′ end of the cDNA is then extended with a homopolymeric tail (usually a string of Cs) by terminal deoxynucleotide transferase (TdT), or is ligated to an oligonucleotide adapter. Thereafter, two rounds of semi-nested PCR are performed to amplify the region with the 5′ unknown sequence. PCR also allows end-extension of amplicons via primers for downstream applications, such as restriction site introduction for directional cloning and universal sequencing primer binding sites for sequencing.

In 3′ RACE (Figure 5B), mRNA is reverse-transcribed to cDNA using an oligo(dT) primer with an adapter sequence. Two rounds of semi-nested PCR are then performed using primers specific to known upstream exon sequences and the adapter sequences introduced through the oligo(dT) primer. In this manner, unknown 3´ mRNA sequences between the exons and the poly(A) tail are amplified for further analysis.

The quality of input RNA and setup of the reverse transcription reaction are critical for successful RACE experiments. In 5′ RACE, first-strand cDNAs of any length (i.e., even those not reaching the 5′ end of the mRNA) will possess the added sequence (i.e., homopolymeric tail or adapter) and subsequently be amplified in PCR. To maximize full-length cDNA synthesis, reverse transcriptases with minimal RNase H activity, high processivity, and high thermostability should be selected.

Learn more: Reverse transcriptase properties

Development of DNA microarrays during the 1990s initiated large-scale profiling of gene expression without bias or prior hypothesis. Microarrays consist of thousands of chambers, known as “features” or “spots”, on glass or silicon wafers. Each feature contains, immobilized on its surface, identical copies of a single-stranded DNA sequence called a “probe”, which represents one gene. The probes hybridize to fluorescently labeled cDNA targets that are applied to the microarray, allowing simultaneous comparison of gene expression between two samples (Figures 6 and 7) [10–12].

Microarray probes are generated from known sequences of the genome or cDNA of an organism. For example, PCR can be used to make copies of every known gene, products of which are then denatured to single-stranded DNA and spotted onto a chip as immobilized probes. Alternatively, oligonucleotides of 20–60 nt can be synthesized directly on a chip as microarray probes [13].

Figure 7 gives an overview of how microarrays are used for differential gene expression analysis. First, total RNA or mRNA is isolated from two samples—experimental (also called “test” or “treated”) and control (also called “reference” or “normal”). The purified RNA samples are then converted to cDNA and labeled with different fluorescent dyes. Next, the labeled cDNA targets of both samples are mixed and allowed to hybridize to the probes on one microarray chip. After unbound targets are washed away, the microarray is scanned to detect the labeled fluorophores. The ratios of the two fluorescent signals are then analyzed to quantify expression of genes affected by the experimental conditions.

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cDNA targets can be labeled either during or after reverse transcription (Figure 8). In direct labeling, fluorescently labeled nucleotides are incorporated during cDNA synthesis. Alternatively, in indirect labeling, nucleotides modified to enable conjugation may be used in reverse transcription, and then the cDNAs are subsequently labeled with fluorophores. Although the indirect method involves a longer workflow, the fluorescence labeling tends to be more efficient.

Figure 8. Direct and indirect cDNA labeling.

When lower amounts of input RNA are available (e.g., 10–100 ng), RNA may be reverse-transcribed to double-stranded cDNA using T7-oligo(dT) promoter primers. The subsequent cDNAs are then amplified by in vitro transcription (Figure 9). During in vitro transcription, RNA may be labeled directly or indirectly using modified ribonucleotides. Alternatively, amplified RNA may be reverse-transcribed and labeled to generate the cDNA targets.

In selecting a reverse transcriptase for preparation of cDNA targets for microarray experiments, the ability to obtain full-length cDNAs in high yields, even when RNA sequences have high GC content or secondary structure, is critical for good coverage of the RNA populations. Equally important, the reverse transcriptase must be able to incorporate modified nucleotides efficiently in order to ensure high signal-to-background ratios that enable accurate and unbiased detection of the input RNA populations.

Learn more: Reverse transcriptase properties
White paper:  Engineered reverse transcriptase

RNA sequencing, or RNA-Seq, is commonly performed to gain insight into RNAs transcribed from the genome and their regulation. With the advent of next-generation sequencing (NGS), RNA-Seq has become a high-throughput approach for analysis of the whole transcriptome (i.e., coding and long noncoding RNA species that have been transcribed), determination of gene expression, discovery of splice variants and fusion transcripts, and detection of low-abundance genes [14,15]. Advantages of RNA-Seq over microarrays include greater dynamic range, higher sensitivity, and the ability to characterize RNA sequences without prior genomic information.

Reverse transcription is involved in the preparation of templates for RNA-Seq, since most sequencing platforms are designed for DNA. It is desirable that the resulting cDNA population represent the original RNA population, including the low-abundance transcripts, with minimum bias. Full-length cDNA synthesis is also important, to capture all RNA sequences in the sample. The error rate of reverse transcription may be critical, depending on the sequencing library size and data quality. Therefore, the reverse transcriptase should be selected with careful consideration.

Learn more: Reverse transcriptase properties

Research goals and sequencing technologies will dictate the order and method of RNA-Seq template preparation [16,17]. Nevertheless, a typical workflow for generating a library for sequencing includes enrichment of the RNA of interest, fragmentation of RNA or cDNA, reverse transcription, addition of sequencing adapters (and indices or barcodes, if multiplexing), and optional PCR amplification of the library (Figure 10).

Figure 10. Traditional workflow of RNA sequencing.

Learn more: RNA sequencing

RT-LAMP is a fast, simple, and sensitive solution for RNA detection, with several methods for evaluating results. Due to its simple workflow and fast reaction time, it is especially useful in research or field settings for the detection and surveillance of viral pathogens.

The LAMP method relies on DNA polymerase with a strong strand-displacement activity, and specifically designed inner and outer primers as well as loop primers. For amplification of RNA targets, a one-step reaction can be carried out by simply adding a reverse transcriptase and RNase inhibitors to a LAMP reaction (RT-LAMP).

LAMP occurs at a constant temperature (60–65°C) and is classified as isothermal amplification. The LAMP technique requires 4 or 6 specially designed primers that bind to two distinct target regions (~300 bp apart). LAMP was originally developed using 4 primers, but subsequent addition of two loop primers reduced reaction time in half. Currently, target RNA or DNA using LAMP method can be amplified in less than 15 minutes. Primers needed for LAMP include two outer (F3 and B3) primers, two inner primers (forward inner primer (FIP) and backward inner primer (BIP), and loop primers (loop forward (Loop F) and loop backward (Loop B).

LAMP occurs in two steps—noncyclic and auto-cyclic. The first step is primer extension from the inner primer (FIP), which hybridizes to the target DNA and starts complementary strand synthesis. This is followed by strand invasion extension from the outer primer (F3), releasing single-stranded DNA that serves as a template for the backward primers. The converted inner sequence forms a stem-loop structure at the F-linked end. The same process is repeated on the other end with BIP and B3 primers, resulting in a dumbbell structure with stem-loops on both the 3′ and 5′ ends as they become complementary to sequences further inwards (enabling the formation of a stem-loop DNA structure). This structure contains multiple sites for repeated amplification initiation and facilitates DNA amplification by auto-cycling, resulting in multiple lengths and cauliflower-like structures of amplified DNA (Figure 11).

• Doesn’t require thermal cycler (only heating block)
• Fast turnaround time (15–60 minutes)
• Multiple options for detection
• Simplified workflow
• Higher tolerance to inhibitors
• Adaptability for field testing

RT-LAMP requires only low quantities of RNA, is tolerant of inhibitors, and offers easy handling as well as good specificity and sensitivity. Amplification under isothermal conditions removes the need for a thermal cycler and offers higher amplification efficiency, as there is no need to wait for temperature changes. Because of these qualities, LAMP technology has undergone exponential growth in its applications since its discovery. RT-LAMP is used in laboratories for faster detection of pathogens (bacteria, parasites, and viruses). Due to its simplicity, it is also a key method for adaptability to field or point-of-care settings.

Explore: LAMP solutions