Cytokine RT-qPCR and ddPCR for immunological investigations of the endangered Australian sea lion (Neophoca cinerea) and other mammals

Real-time PCR primer design

Sequences for genes of interest (GOI) IL-4, IL-6, IL-10, IFNγ and TNFα of multiple mammal species were obtained from Genbank (http://www.ncbi.nlm.nih.gov/genbank) and aligned (Table 1).

Primers for qPCR (Macrogen, Seoul, South Korea) (Table 1) were designed within conserved regions using NCBI Primer-BLAST (Ye et al., 2012) and recommended parameters for designing SYBR^® Green primers (Thornton & Basu, 2011). Secondary structures and specificity against non-specific sequences for each primer set were assessed in silico using BeaconDesigner™ (http://www.premierbiosoft.com/qOligo/Oligo.jsp?PID=1) and NCBI Primer-BLAST (Ye et al., 2012), respectively. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was selected as a reference gene for this study given its stability in a variety of sample types and its validation in many species including domestic animals, marsupials and marine mammals (Beineke et al., 2004; Fonfara et al., 2008; Maher et al., 2014; Puech et al., 2015; Sharp et al., 2006). Primers for GAPDH (Macrogen, Seoul, South Korea) were selected from a previous publication that used Genbank sequences for Canis familiaris (Peters et al., 2004) and its performance was optimised to the study conditions.

Blood collection, RNA extraction and reverse transcription

Blood samples (0.5 mL) from domestic dog (C. familiaris, n = 4), cattle (Bos Taurus, n = 3) and sheep (Ovis aries, n = 3) were collected from brachial, tail and jugular veins, respectively into EDTA tubes (Sarstedt, Nümbrecht, Germany) and then centrifuged at 5,000×g for up to 3 min. Plasma was removed using a sterile disposable pipette, and the remaining red blood cells and buffy coat were resuspended in 1,300 µl RNAlater^™ (Applied Biosystems, Carlsbad, CA, USA) and then transferred into cryovials to approximate field storage conditions. Samples were initially stored at 4 °C for 2–4 days and then at −20 °C until RNA extraction was performed within 12 months of blood collection. RNA extractions from the same species were combined and used as pooled samples for further applications.

In 2010, four blood samples (0.5 mL) collected from the brachial vein from N. cinerea pups for a previous study (Marcus, Higgins & Gray, 2014) (Government of South Australia Department of Environment, Water and Natural Resources; Wildlife Ethics Committee approvals 3–2008 and 3–2011 and Scientific Research Permits A25088/4–5) were immediately transferred into EDTA tubes (Sarstedt, Nümbrecht, Germany) and processed as described above. Samples were initially stored at 4 °C for 2–4 days and then at −20 °C until RNA extraction was performed in 2018. RNA extractions were combined and used as pooled samples for further applications.

For RNA extractions, samples in RNAlater^™ were thawed at room temperature, centrifuged at 16,000×g for 1 min, and the supernatant of RNAlater^™ discarded from the cell pellet. Total RNA extraction was performed on the cell pellet using the RiboPure^™-Blood Kit (Ambion, Carlsbad, CA, USA) according to the manufacturer’s instructions, such that the equivalent of 200 µl of whole blood was used per extraction. The RNA concentration and purity were assessed (A₂₆₀/A₂₈₀) using a NanoDrop 1000, Thermo Scientific^™ (Waltham, MA, USA) and RNA stored as multiple aliquots at −80 °C for subsequent use.

For analysis, aliquots of RNA were thawed on ice, and two sequential DNase treatments were performed using the RNase-free DNase I (provided in the RNA extraction kit) to eliminate genomic DNA (gDNA). Reverse transcription was performed on 50–100 ng of RNA template in a 20 µl reaction with the RevertAid First Strand cDNA Synthesis Kit (Thermo Fisher^™, Carlsbad, CA, USA), with a combination (50:50) of random hexamer and oligo(dT)₁₈ primers to improve the sensitivity of cDNA synthesis (Ferrante, Hunter & Wellehan, 2018; Gallup, 2011). cDNA was stored as multiple aliquots at −20 °C for subsequent use.

Real-time PCR optimisation and validation

Optimisation and validation parameters were achieved following recommendations for qPCR assays from Bustin et al. (2009) (MIQE guidelines). All qPCR assays were performed using SYBR Green (SsoAdvanced^™ Universal SYBR^® Green Supermix, BioRad) on a CFX96 Real-Time cycler (Biorad, Hercules, CA, USA). Cycling conditions were optimised by annealing temperature (T_a) gradient and evaluation of three primer concentrations (100, 200 and 300 nM). Optimal parameters were determined based on those that yielded a single, sharp peak in the melt curve analysis with the lowest quantification cycle (Ct) for each primer pair (Table 1).

Amplifications were performed in white 8-strip PCR tubes (BioRad, California, USA), following manufacturer’s instructions for the SYBR Green Supermix in a 20 µl reaction with each primer pair at optimised concentration and 4 µl of cDNA template. In addition, “no-reverse transcription” controls (NRT), and “no template” controls (NTC) of RNase-DNase free water, were included in each run to check for gDNA contamination and the formation of primer-dimers, respectively. Under identical qPCR cycling conditions, reactions were validated across cDNA templates from N. cinerea, C. familiaris, B. taurus and O. aries. Amplification conditions were 95 °C for 1 min (1 cycle); 95 °C for 10 s and 60 °C for 20 s (40 cycles). After each cycling protocol, a melt curve analysis was generated by heating from 65 °C to 95 °C with 0.5 °C increments for 5 s to confirm the absence of non-specific products or primer dimers and define melting temperatures (T_m) for each amplicon. The size of each qPCR product was confirmed by 2% agarose gel electrophoresis and identity of the amplicon was further confirmed by DNA sequence analysis (Macrogen, Seoul, South Korea) and comparison with nucleotide sequences of terrestrial and marine mammals using the NCBI BLAST programme (Altschul et al., 1990).

Confirmed qPCR products were removed from the plates and diluted for further use as a template in standard curves. Amplification efficiencies for each gene of interest (GOI) and the reference gene (GAPDH) were determined for each species through a standard curve using serial dilutions of qPCR product with a minimum of five standards with the dilution extending at least to that producing a Ct of 34. The efficiency (E) and linearity (R²) of each primer pair were calculated on these curves using the Bio-Rad CFX Maestro software 1.1 (BioRad, Hercules, CA, USA) with automatic threshold settings. Linear regression of the qPCR standard curves was recalculated with Microsoft Excel (Microsoft, Redmond, WA, USA; 2016). Primer pairs with efficiencies of 100% ± 10% and R² value > 0.96 were considered optimised for qPCR (IL-4, IL-6, IL-10, IFNγ and TNFα) following MIQE recommended ranges. The limit of quantification (LoQ) was based on the linear operating range of each assay (Berdal & Holst-Jensen, 2001). Assays that showed linear and efficient amplification but that produced Ct values above the dynamic range of qPCR (Ct alues >35) when applied to N. cinerea pup blood samples, were selected for subsequent development of novel ddPCR. As the assays were developed for use in relative expression studies using the delta Ct method, absolute quantification and limits of detection (LOD) were not derived using quantified standards. Quantified standards were, however used for direct comparison of sensitivity of ddPCR vs qPCR in IL-4 and IFNγ assays.

ddPCR assays for Neophoca cinerea

qPCR products obtained from the amplification of N. cinerea samples with the IL-4 and IFNγ qPCR primers formerly described in this study were sequenced and aligned using the CLC Main Workbench 6.9.1 (Qiagen, Redwood City, CA, USA). NCBI Primer-BLAST programme (Ye et al., 2012) was used to design N. cinerea primer-probe pairs for ddPCR following recommended parameters from the Droplet Digital^™ PCR Applications Guide (www.bio-rad.com). The primers and probes sequences (Macrogen, Seoul, South Korea) are listed in Table 2. The in silico tool ‘PCR Primer Stats’ (http://www.bioinformatics.org/sms2/pcr_primer_stats.html) was used to evaluate primer-probe pairs melting temperature, GC content and secondary structures (Stothard, 2000). Probes were labelled with FAM (F) fluorophore and quenched with non-fluorescent black hole quenchers number 1 (BHQ-1).

Instruments, reagents and consumables for the ddPCR workflow were supplied by Bio-Rad (Bio-Rad, Hercules, CA, USA). Optimal ddPCR annealing temperatures for IL-4 and IFNγ assays were defined by performing a temperature gradient in the annealing/extension step of the thermal cycling protocol as suggested by ddPCR guidelines (Huggett et al., 2013). All ddPCR optimisation assays were performed using the ddPCR^™ Supermix for Probes (no dUTP) master mix in a C1000 Touch^™ Thermal Cycler. The fluorescence difference between NTC and positive samples within a single run was used to set a threshold between negative and positive droplets. Positive droplets show increased fluorescent amplitude when compared to the negative droplets and contain at least one copy of the target per sample. The optimal annealing temperature for these assays was defined as the one giving the largest difference in fluorescence between negative and positive droplets (Table 2; Fig. 1) (Huggett et al., 2013).

ddPCR master mix reactions included 10 µl of the Supermix, one µl of each primer and probe, six µl of cDNA from pooled N. cinerea samples and RNase-DNase free water to complete a 20 µl total volume reaction (Table 2). PCR mix and Droplet Generation oil for Probes were added to corresponding wells in the Droplet Generator DG8^™ Cartridge and covered with DG8^™ Gaskets following the manufacturer’s instructions. Droplets were generated by using the QX100^™ Droplet Generator, gently transferred to a clean 96-well PCR plate and sealed using a PCR Plate Sealer. Amplification was performed using the following cycling conditions: an initial enzyme activation period of 10 min at 95 °C, followed by 40 cycles consisting of denaturation at 95 °C for 30 s and annealing/extension step of 59 °C for 1 min, and followed by an enzyme deactivation period of 10 min at 95 °C and a 4 °C indefinite hold. The overall ramp rate was set at 2 °C s⁻¹. Droplets were read as positive or negative with a QX200 Droplet Reader, and further analysis was performed using the Quanta-Soft Analysis Pro^™ software (Bio-Rad). The software, based on fluorescence amplitude, establishes a threshold between positive and negative droplets. Positive droplets are converted to copy numbers in the PCR mix based on Poisson algorithms (Droplet Digital^™ PCR Applications Guide).

To assess linearity (R²), efficiency (E) and to compare the limits of detection (LoD) and quantification (LoQ) of qPCR and ddPCR, IL-4 and IFNγ consensus sequences were selected to synthesise double-stranded DNA standards (gBlocks^® gene fragments; Integrated DNA Technologies, Singapore). Each DNA standard was resuspended in Tris EDTA buffer (The Bosch Institute, Faculty of Medicine, The University of Sydney) to reach a final concentration of 10 ng µl⁻¹ according to the manufacturer’s instructions with subsequent storage at −20 °C. For qPCR, a calibration curve (regression line) was performed with 10-fold serial dilutions of the standard ranging from 10⁶ to one target copies µl⁻¹ and six technical replicates for each dilution. For ddPCR, 5-fold serial dilutions ranged from 200 to 0 target copies µl⁻¹ with three replicates for each dilution. LoD and LoQ were calculated with Microsoft Excel (2016) as 3.3 and 10 times, respectively, the standard deviation of the y-intercept of the regression line, divided by the slope of the corresponding calibration curve (FDA, 1996; Mohamad, 2018).

Real-time qPCR design and validation

The primer pairs designed in this study measured cytokine gene expression in mammalian species in separate evolutionary clades, representing terrestrial (domestic dog, sheep and cattle) and the Australian sea lion. mRNA sequences from qPCR products showed high homology to available mammalian sequence data, with values ranging from 86 to 100% (Table 3).

The integrity of isolated RNA was demonstrated in all blood samples from every species by the amplification of GAPDH mRNA (Ct 25 ± 3.1, mean ± SD). Optimal qPCR parameters allowed amplification of IL-4 (Australian sea lion and dog), IL-6 (Australian sea lion and dog), IL-10 (all four targeted species), TNFα (all four targeted species), IFNγ (Australian sea lion and dog) and GAPDH (all four targeted species), using the same cycling protocol, with a combined annealing and extension step at 60 °C. The defined optimal parameters for the assays are shown in Table 1. In addition, the presence of a single specific product was confirmed by melt curve analysis (Figs. S1–S4), agarose gel electrophoresis and sequencing. The absence of non-specific products and primer-dimers was confirmed (Figs. S1–S4). No amplification was detected in NRT controls and NTC. Two consecutive DNAse treatments were required to eliminate evidence of gDNA in NRT controls. All assay efficiencies ranged between 87% (Ovis aries TNFα) and 111% (Bos taurus IL-10) and had strong linearity (R²) (Table 4). Although sample sizes were too small for comparison, the low expression of IL-4 and IFNγ in N. cinerea was consistent with levels in canine samples that we assessed (Ct 32 ± 2.5, n = 4).

ddPCR assays for Neophoca cinerea

The ddPCR primers and probe assays designed in this study effectively amplified N. cinerea blood mRNA. The forward-reverse and probe sequences of each assay are listed in Table 2. Gene sequence data obtained for both primer-probe pairs was BLASTn compared against similar mammalian species genes, and the results show 98% homology with marine mammals (Table 5). No amplification was detected in NTC.

The optimal ddPCR annealing/extension temperature for IL-4 and IFNγ defined by a temperature gradient was 60°C and 58°C, respectively (Fig. 1). The defined optimal parameters for the assays are shown in Table 2. LoD and LoQ were confirmed to be lower in ddPCR than in qPCR (Table 6; Fig. 2). Quantification of IFNγ and IL-4 in the pooled N. cinerea samples indicated 69 and 13 copies per reaction, respectively (Table 6).