Misión

After extraction of total RNA, reverse transcription was used to generate cDNA. 5µg tem- plate RNA and 0.5µg oligo-dT primers were added to an RNAse-free tube. Nuclease-free water was added to a final volume of 12µL. The samples were heated to 70℃ for 5 min before chilling on ice. Subsequently, 1µL 10 mM dNTP mix, 4µL 5×reaction buffer (pro- vided with the BioScript enzyme), and 0.5µL RNAse inhibitor were added to each sample. The samples were mixed by pipetting before 1µL BioScript enzyme was added. The reverse transcriptase was thoroughly mixed into the sample by pipetting and the samples were in- cubated at 40℃for 60 min. The reaction was stopped by heating to 70℃for 10 min followed by chilling on ice. The cDNA was stored at−20℃without repeated freeze-thawing.

6.6.5

Testing of primers

Primers were ordered lyophilised and were reconstituted on arrival in water to a concentra- tion of 100µM. Pairs of primers were then diluted together to a working solution of 5µM of each primer in water. Before use, primers were tested to ensure that they amplified a spe- cific product of the correct mass. Reaction tubes for each primer containing 1µL template cDNA from the reverse transcription step, 1µL primer mix at 5µM in water, 8µL water and 10µL BioMix Red mix including DNA polymerase, dNTPs, buffers, salts and loading dye. The reaction tube was thoroughly mixed and cycled through the conditions shown in

Figure 6.14 in a PCR thermocycler.

Figure 6.14: BioMix Red PCR cycling conditions

The reaction products were separated on a 2% agarose gel. This gel was prepared by dissolving 2% w/v agarose in tris-acetate-EDTA (TAE) buffer (final concentrations 20 mM tris base, 10 mM acetic acid and 1 mM EDTA.Na2, pH 8.0) by careful heating in a microwave oven. Ethidium bromide was added from a 10 mg mL−1 stock to a final concentration of 0.5µg mL−1and the gel allowed to set before submerging in TAE buffer. 15µL of each 20µL PCR product was transferred to a well on the gel and a voltage of between 90–110 V was applied to the gel for 1–2 hours to separate the reaction products. The gel was said to be finished once the dye front had moved approximately 5 cm from the loading wells. The gel was viewed and photographed in a UV transilluminator (see Figure 6.15). Any primer pair which did not amplify with a single clean product was substituted for an alternative pair of primers for the same product. An additional technique to test primer quality not employed here would be to digest each product with a known restriction enzyme and to re-run the products on a second gel. The size ratio would indicate if the sequence of the amplified section is likely correct.

Figure 6.15: Primer quality control example gel image. Each lane represents a PCR reaction product. All lanes have produced a single product at the correct mass (as determined by including a molecular mass ladder on the gel) except lane 5 which has not amplified adequately.

6.6.6

Assay controls

All qPCR assays were thoroughly controlled to ensure reliability of the quantitative results. Water blank qPCR amplifications were performed in which the template was substituted for water and “RT−” amplifications were performed in which the template was prepared using a reverse transcription step with water substituting for the reverse transcriptase. Am- plification in the water blank would indicate contamination of the experiment by exogenous material whilst amplification in the RT− blank is usually due to amplification from the genomic DNA present in the sample.

Example traces for a properly conducted experiment are shown in Figure 6.16. In cases where the water blank amplified at all, it did so very late (usually past 35 cycles). The gap between the amplification of the target and the RT−control indicated the relative abundance of genomic DNA to cRNA. For example, amplification 7 cycles later represents 1_/27 _{or 0.7% of the target being present in the form of genomic DNA. Amplification 9 cycles}

later would represent 0.2%. At this stage, other errors become more significant and the effect of the genomic DNA can be ignored. It is not possible to perform a water blank amplification for every well so it was assumed that if the water blank on a plate did not amplify then the quality of the plate preparation was such that the rest of the plate was also free of contamination.

Figure 6.16: qPCR example control amplifications for THTR-1. This is an example of a properly conducted experiment. The RT+ sample crosses the threshold first at around 21 cycles. The RT− sample crosses much later at around 31 cycles meaning that it has an abundance of approximately 1_/21_0or 1_{/1000 of the RT+ sample. The water blank does not} cross the threshold indicating that there has been no contamination of the well.

In addition to these controls, a melt curve was prepared at least onece for each primer pair and target. This was achieved using the PCR machine to increase the temperature of the sample in small steps from a relatively low temperature (e.g. 65℃) to a relatively high temperature (e.g. 90℃) over a range that incorporated the melting temperature of each product. The fluorescence decreased gradually as the products melted. By plotting the first order differential of the fluorescence against temperature, peaks were derived that corresponded to products. Figure 6.17 is an example plot containing two peaks. The good qPCR reaction has only one sharp peak corresponding to a single product and the bad qPCR reaction contains a split peak indicating the presence of primer dimers or other artefacts of amplification. Reactions with poor peak shape had their primers re-designed.

Figure 6.17: qPCR example product melt curves. The green curve represents a good quality PCR reaction with a single product. The red curve represents the product obtained from a RT−control which is of poor quality with two discernible peaks.

6.6.7

Primer efficiency

Since only relative levels of expression were required it was not necessary to construct a standard curve of the type required for absolute quantitation. The traditional comparative Ctmethod or the ∆∆Ctmethod is an approximation method that assumes that each PCR

reaction is a perfect doubling. The ∆Ctbetween reference gene and target gene is calculated

for both the control condition and the test condition. The difference between these ∆Ct

reference gene is then given by 2∆∆Ct_{. I used a more accurate method for quantitation de-}

scribed by Pfaffl [2001] which used pre-determined efficiencies for PCR reactions to produce higher quality results.

Detemining efficiencies for the Pfaffel method was achieved by constructing a stan- dard curve based on serial dilutions of template DNA over several orders of magnitude. A typical curve extended from 10 to 0.01 ng RNA. The samples was amplified following the cycling conditions to be used for the assay and the Ct values were determined for each

concentration. The Ct values were plotted against the logarithm of the amount of RNA

(ng) and a straight line was fitted to the data. Figure 6.18 shows a typical standard curve obtained. The efficiency was determined according to Equation 6.1 wheremis the gradient of the efficiency plot. Convention varies but I used a value of 1 to mean a perfectly efficient reaction with a doubling of product every cycle. Efficiencies should be between 0.9 and 1.1 for properly functioning primers. Efficiencies that were outside this range were often due to off-target amplification or poor annealing and were solved by re-designing primers.

Figure 6.18: Example primer efficiency determination curve. This example curve was ob- tained for primers for the gene TXNIP. The gradient of the line can be used to determine efficiency using Equation 6.1.

Once primer efficiency had been determined, the ratio of relative expression levels of the genes of interest in treated versus control conditions was determined according to Equation 6.2, assumingβ-actin as the loading control [Pfaffl, 2001].

Ratio =(1 + Efficiencyβ-actin)

∆Ctβ-actin(control−treated)

(1 + Efficiency_target)∆Cttarget(control−treated) (6.2)

6.6.8

Design of experiments

Each primer pair was thoroughly tested prior to analysis of samples to determine efficiency and specificity by the methods described above. An RT−control was used with a typical template to determine whether the genomic DNA present would amplify at a similar rate to cDNA species. No cDNA species were sufficiently rare to amplify near the genomic DNA so further RT−controls for each primer pair were not performed to save space on plates. The loading control chosen for each experiment was β-actin, a commonly used house-keeping gene. Using a single reference gene is potentially problematic if its expression varies with experimental conditions. It would be beneficial to use multiple genes (e.g. GAPDH, 5S RNA) simultaneously to ensure that the normalisation is correct and appropriate.

Experiments were designed to fit onto the minimum number of plates possible. A given biological replicate was assayed for all genes of interest on one 96-well plate with three technical replicates for each gene.

6.6.9

Sample analysis

After plate layout design, the plates were constructed and run on a real-time cycler. To simplify preparation of the plates, a concentrated SYBR green-containing master mix was purchased (SensiMix SYBR Low-Rox kit). This master mix contained a hot-start Taq polymerase which was modified to cause complete inactivation before exposure to a high temperature (95℃for 10 min), thereby minimising primer-dimer formation and non-specific amplification which could potentially be caused during sample set-up and handling.

Each well contained a final volume of 20µL comprising 1µL primer solution at 5µM for each primer in water, 1µL template DNA, 8µL water and 10µL 2x SYBR green reaction master mix. To minimise pipetting errors, the primer solution and the SYBR green solution were combined into a single solution (11µL well−1_{) and the template DNA was diluted with} water to allow a pipetting volume of 9µL well−1_.

After dispensing reagents, the plates were sealed with transparent plate seals and mixed by vortexing. The plates were centrifuged briefly to sediment droplets of liquid

in the wells and to eliminate frothing and bubbling caused by mixing. The plates were transferred to the real-time cycler immediately if possible or stored at 4℃ for a maximum of 24 hours before analysis. Each plate was cycled according to Figure 6.19 and fluorescence measured each cycle during the extension phase. Fluorescence of the passive dye ROX was also recorded each cycle and used by the machine to normalise signal fluorescence. Not all plates were subjected to a dissociation stage since these are time-consuming. Instead, each primer pair was tested thoroughly beforehand to ensure that the products were of a known quality.

Figure 6.19: qPCR cycling conditions with SensiMix SYBR green reagents