1. Introduction to quantitative real- time PCR Veryan Codd vc15@le.ac.uk

2. Overview What is real-time PCR and what is it used for? Real time platforms available in CVS Basic Principles Experimental design, Controls and QC Quantification methods

3. What is real time PCR Ability to monitor PCR reaction in real time Accurate quantification of the amount of starting material (template) added to a reaction Increase dynamic range of detection over end point methods

4. Why do we use real-time PCR Quantification of gene expression levels Quantification of DNA (Telomere assays, mtDNA content, copy number) Virus titre Can be used as a quantitative step as part of another technique e.g. ChIP and DNA Methylation

5. Our facilities – ViiA7 Life Tech (Applied Biosystems) 96, 384 well plates and Taqman array cards Upgraded 7900 Advanced optics – 6 excitation and 6 emission wavelengths – 21 dye combinations Value ~75K

6. Our facilities – Rotorgene-Q Qiagen Unique rotary system – no well-to-well thermal variation – improved accuracy 0.2ml tubes (36), 0.1ml strip tube (72), rotordisc (72 or 100) Robotic set-up required but available for 100 well disc Additional quant methods in software 2-plex – Green (SYBR green, FAM) and Yellow (VIC, JOE)channels

7. Our facilities – Rotorgene-Q Value ~20K

8. Basic Principles PCR with a fluorescent dye that can be detected PCR is theoretically exponential – product doubles with each cycle The more starting material the faster the reaction will occur (lower cycle number)

9. Experimental design - reagents Primers – Specific to target – 18 – 24bp – Produce product 50 – 150 bp in length (200bp does work) – Product ~50% GC and avoid GC stretches and repetitive sequences – Check for primer dimers especially for SYBR green assays – F and R primers annealing temp within 5˚C – In separate exons or spanning exon boundaries Pre-designed assay available (taqman probes, sybr based assays) Array cards / expression sets

10. Experimental design – probe or SYBR? SYBR green Dye binds minor groove of all DNA Fluorescence much stronger when bound More product more dye bound Cheap Not specific At early stage check product on gel and perform dissociation analysis (melt curve) Probe Dye and quencher either end of probe Nuclease activity of polymerase removes dye and quencher during PCR More product produced, more fluorescence No melt curve Allows multiplexing

11. Experimental design- singleplex or multiplex Singleplex (simplex) One gene per PCR SYBR green or Probe – more flexible More variation Usually simpler to set up Uses more enzyme mix but can SYBR green with cheap primers instead of expensive probes More template? Multiplex Multiple genes per PCR Probe based – 2 dyes (VIC FAM) Minimises pipetting error Competition between reactions can be problematic Cheaper? Higher throughput – 2 measurements in one Less template?

12. Experimental design - reagents Enzyme or enzyme master mix governed by primer probe choice and by choice of platform (e.g. ViiA7 or rotorgene) 2x mastermixes widely available – Consistent – Pre-optimized (MgCl2 etc)

13. Experimental design - Template For gene expression analysis need to reverse transcribe mRNA into cDNA Good quality RNA Accurately quantified Variety of RT enzymes commercially available

14. Reverse transcription - Priming Three (or four) basic choices

15. Oligo-dT priming Favourite choice of many as specific to mRNA Total RNA – 80% rRNA – 15% tRNA – 5% mRNA Sensitive to secondary structure – incomplete reverse transcription Design primers at 3’ end of transcript Don’t use 18s as a control (it’s rRNA!)

16. Random Priming Uses random hexamer primers Bind throughout RNA molecule – Secondary structure less of a concern – Less sensitive to RNA degradation – Increase yield of cDNA Amplify ALL RNA – mRNA, tRNA and rRNA

17. Sequence specific priming Only gene of interest is reverse transcribed Limits use of cDNA, used up RNA stocks Only used in one-step reverse transcription- qPCR (one step qRT-PCR)

18. Option no 4 Mix of oligo-dT and random primers Best of both worlds

19. Good laboratory practice qPCR can be a sensitive technique if performed correctly and with care Keep everything clean Aliquot reagents to prevent freeze/thaw Accurate pipetting – Make master mix of everything except template then aliquot – Mix everything gently but thoroughly – 2x master mixes are viscous – Small volume pipetting less accurate – 22ul + 3ul or 23ul + 2ul works well (MM + Template) – Don’t use second stop on pipette to avoid “spraying” Perform triplicate PCRs for sample

20. Contamination Big problem for qPCR Most common cause of contamination comes from product of previous PCRs Don’t leave PCRs in machine indefinitely after completion If running product on get use separate pipettes to those you use for PCR set-up Use PCR dedicated Hood (genomics lab) to set up PCR MM’s – Keep primers and other reagents free from contamination – DNA/cDNA MUST NOT BE ADDED IN THE HOOD – pipette contamination

21. Getting started For pre-designed assays have a test run to check for good amplification If designing own assay – Test to check product of correct size is formed – Optimise annealing temp (gradient PCR) – Test in real-time PCR that amplification looks reasonable – Titrate primer concs if/as necessary

22. Normalisation Housekeeping gene(s) – Accounts for pipetting error and differences in RT efficiencies between samples – Constantly expressed at a stable level – Put the same starting RNA in all RT-reactions when samples are to be compared – IT IS NOT THERE TO ACCOUNT FOR MASSIVE DIFFERENCES IN STARTING MATERIAL! Calibrator sample – Sample run on every assay T0 / Untreated – Compare everything to this sample

23. Controls NTC (No template control) – PCR contamination detection – Primer dimers no-RT control – Detection of genomic contamination and/or non- specific product – Especially important in transient over-expression experiments to check for carry over of plasmid DNA

24. Dissociation or Melt curve analysis To check for single specific product in SYBR green based assays Compare to product on gel in early stages of set-up Detection of primer dimers and non-specific products

25. Melt curve Example 1

26. Melt curve example 2 NTC Primer dimers Specific product

27. Melt curve example 3 Specific product Non-specific product In this case NTC’s showed no product, so no primer dimers, large non- specific product observed on gel

28. Standard curves Do I need a standard curve??? – YES! “My assay is predesigned and guaranteed to be efficient therefore I don’t need to waste my time doing a Standard curve” – Not true! But very common!!

29. What is a standard curve and why is it necessary? Real time PCR conducted across a series of serially diluted template/samples Tell you what the dynamic (linear) range of the assay is Allows calculation of efficiency

30. Cycle number Ct or take-off Log10 template concentration

31. Cycle number Ct or take-off Log10 template concentration

32. Cycle number Ct or take-off Log10 template concentration

33. Linear range 7 -10 cycles Therefore experimental samples with Ct’s outside of this should not be used for quantification Set the concentration of template in middle of linear range If input concentrations are highly variable then more likely to get samples falling outside of LR that cannot be accurately quantified

34. Efficiency Efficiency = 10 (-1/gradient) -1 Gradient of -3.32 = 100% efficient 90-110% considered ok (-3.1 to -3.6) For some, most commonly used, quantification methods it is important to have similar efficiencies between GOI and housekeeping genes

35. Quantification methods- Absolute Quantification Samples calculated against a standard curve of known concentration Allows quantification as copy number – Can be used when one condition contains no expression Takes some variability into account Requires more reagents Needs a known standard – Plasmid DNA containing product – no RT step so efficiencies may be different – In-vitro transcription to generate specific mRNA – RT, costly

36. Quantification methods- ΔCt, ΔΔCt Comparative quantification Threshold cycle number – Ct Slightly subjective Calibrator – i.e T0 or untreated Normalized to housekeeper (HK) ΔCt = Ct GOI -Ct HK Δ ΔCt = ΔCt sample - Δct calibrator Fold change = 2 - Δ ΔCt Assumes efficiency 100% for both GOI and HK Can substitute 2 for your calc efficiency but must be the same for GOI and HK

37. Example Efficiency is 88% GOI (red) Efficiency is 1.07% Housekeeper Calculate average DeltaCt for each dilution: 200-4.8 100-4.65 50-4.5 25-4.35 12.5-4.2 6.25-4.05 3.125-3.9 1.56-3.75 One end of range 1.05 cycles different to other end – equivalent to a two fold difference when it should be the same

38. Comparative quantification - Qiagen Method on rotorgene software Calculates efficiency for each individual sample based on rate of fluorescence change and mean efficiency across run Calculates a “take off” value – start of exponential phase, less subjective than setting threshold for Ct Set calibrator sample, T0 or untreated etc Relative amount calculated against calibrator Relative conc = MAE (calibrator takeoff-sample takeoff) Effectively a ΔCt but with real efficiency Then need to manually normalise to HK or a Δ ΔCt and fold change calculation

39. Example GOI efficiency = 96% HK efficiency = 104% Comparative quant each dilution for GOI and HK separately using one concentration as calibrator GOI/HK Δ ΔRc Δ Δ Ct (using Ct rather than TO) 2000.9990.9961.000 1001.0021.0000.749 501.0061.0010.620 251.0211.0030.488 12.50.9680.9970.357

40. Accuracy Singleplex reaction more prone to variation in pipetting accuracy Replica error – How consistent is Ct/take off value between triplicate measurements? – A 1 cycle difference across the triplicate is a 2-fold difference within the same sample! – Rotorgene 0.2 or 0.3 cycles easily achievable – ViiA7 and other block based ~0.5 Rotorgenes better at accurate quantification of small differences – No thermal variation – Take off calculation less prone to subjectivity

41. Further resources Real-time PCR handbook LifeTech Molecular Cloning, Green and Sambrook MIQE guidlines