Contents · 2019. 12. 16. · Contents Chapter 1 Introduction 5 Main types of PCR 6 What this book...

Contents

Chapter 1 Introduction 5Main types of PCR 6What this book will do for you 7

Chapter 2 Standard PCR 8No bands 9

Missing or non-optimized PCR reagents 9Gel electrophoresis 12DNA polymerase 14GC-rich targets 16Primers 17Sample quality 20

Weak bands 23Reaction components 23Non-optimal PCR program 24

Extra bands 26Primer dimer formation 27Unknown PCR product 27Bands appearing in negative controls 29

Alternative PCR methods 30Nested PCR 30Touchdown PCR and Stepdown PCR 33Multiplex PCR 35

General PCR troubleshooting guide 37Useful References 39

Chapter 3 Quantitative real-time PCR (qPCR) 41Hydrolysis (Taqman) probes 42Dual Hybridization (FRET) Probes 43SYBR Green Dye 44No fluorescent signal 45

qPCR assay set-up 45

Primer/probe design 47Plate preparation 48

Low sensitivity (high CT values) 49Biological samples 49Primer design 50qPCR assay set-up 51

Non-specific amplification 52Primer dimer formation 54Poor primer specificity 55Genomic DNA contamination 55

High degree of technical variation 56Program set-up 57Plate set-up 57

Poor standard curve 58Sample handling 59Program and plate set-up 60

Useful References 61

Chapter 4 Digital PCR 62Droplet digital PCR 63Low droplet (partition) count 65

Blocked micro-channels 65Empty wells 66Droplet damage/loss 66

Low fluorescence amplitude/threshold 67Poor assay optimization 67Inappropriate fluorescence threshold 68Template concentration 69

High degree of technical variation 71False negatives and false positives 71

Contamination 72PCR inhibitors 72Calibration control 73

A word on dMIQE 73Useful References 74

Conclusion 75

About the author: Kirsten Hogg 76

Chapter 1 Introduction

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Introduction

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The polymerase chain reaction (PCR) technique, first developed in the early 1980s by Kary Mullis, marked a ground-breaking

milestone for science. Put plainly, PCR enables us to synthesize DNA in a tube by combining a template, the basic building blocks of DNA (nucleotide bases) and polymerizing enzymes isolated from simple heat-stable organisms found in nature. The relatively effortless amplification of a tiny segment of genomic DNA into thousands of identical copies means that PCR has become a core laboratory tool for hundreds of different molecular and biotech applications. Over the last decade, the development of new PCR technologies has led to enormous advances in sensitivity, precision and throughput, paving the way for major new discoveries in dis-ease and enhancing our understanding of genome function.

Main types of PCR

The type of PCR you use in the lab will vary depending on the application or biological question. However, there are three main types of PCR that most of us use routinely:

Conventional PCR using a thermocycler revolutionized our ability to identify and screen for rare variants (mutations), perform vast swathes of genomic sequencing, analyze gene function through mutagenesis screening, find forensic evidence and (most commonly!) detect and quantify messenger RNA (mRNA) levels in biological tissues and cells.

The advent of real-time quantitative PCR (qPCR) elevated PCR technology to unquestionably higher levels. qPCR, which combines amplification and detection in one system, allows for the precise quantification of very small amounts of starting template so that gene expression can be accurately compared from sample to sample in ‘real-time’.

http://www.karymullis.com/pcr.shtml

Bitesize Bio Guide to PCR

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The most recent PCR technology to enter the stage is digital PCR (dPCR). dPCR provides the most ‘real’ form of absolute quantification yet, by physically counting each DNA or RNA molecule...one at a time. The sensitivity of dPCR goes far beyond what qPCR can achieve. In fact, dPCR can theoretically detect just one copy of a rare variant amongst thousands of wild-type templates!

What this book will do for you

While PCR is a well-established molecular biology technique and is routinely used in the lab, it does not come without its technical tribulations. In fact, it can be all too easy to go wrong with PCR – usually due to perfectly avoidable errors!

In addition, dPCR is currently in its infancy and is far less well-developed than qPCR, introducing a whole new set of potential technical difficulties.

This is where we come in!

This book will present the ins and outs of PCR experiments and design, from the very conceptual stages all the way through to the finished product. Each of the chapters delves deeply into the intricacies of PCR, providing detailed practical advice for the most common problems as well as the technical issues that you least expect. This book will mentor you in conventional PCR, qPCR and dPCR to provide a comprehensive trouble-shooting guide to fit the needs of most scientists working with PCR.

Chapter 2 Standard PCR

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PCR is the most routinely performed molecular biology technique and is a basic ‘bread and butter’ skill for any researcher. How-

ever, while PCR can be an easy and effective method of nucleic acid amplification, it also has a dark side, with multiple strata of complexity that can be problematic for even the most experienced of researchers. Fortunately, no matter whether the PCR problem originates with the starting material, primer design, PCR reagents, PCR thermal program or in the visualization of the PCR product itself, there are a number of simple strategies to overcome these challenges.

This chapter highlights common PCR problems and provides several tips and solutions for troubleshooting that tricky PCR experiment.

No bands

Most of us have experienced that heart-wrenching moment when you flick on the UV switch and don’t see any products light up on the gel. There are various reasons why the expected bands could be absent. Possibly, the PCR has not worked due to a primer failure or a mistake during the reaction set-up. Alternatively (and somewhat better!), the PCR has worked, but something has gone amiss during the gel electrophoresis step. In this section, we’ll discuss how to prevent or address the potential causes leading to no visible PCR product.

Missing or non-optimized PCR reagents

Missing PCR reagents

A common (and frustrating) reason behind a failed PCR experiment is the accidental omission of one of the PCR reagents. The simplest way to avoid this is to use a ‘check-list’ system – each

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time a reagent is added, tick it off. Also, adding each component in the same order every time will help to prevent mistakes and reduce the variation between experiments.

Non-optimized PCR reagents

The right concentration (and balance) of each PCR component is vital to achieving an effectively amplified and clean PCR product. Both magnesium and dNTP concentrations can be easily and inexpensively adjusted during PCR optimization to establish the most appropriate amounts for that assay.

Magnesium is an essential component of the PCR buffer mix, as it stabilizes the primer-template complex and acts as a co-factor for DNA polymerase. Too low levels of Mg2+ ions can prevent the PCR reaction from working at all; however, excessive levels can result in non-specific amplification due to the stabilization of poorly annealed primers.

The optimal magnesium range is between 1 mM and 4 mM per reaction. As a rule of thumb, 1.5 mM MgCl2 usually produces good results. If you have weak bands or non-specific products, then the Mg2+ concentrations can be adjusted easily. Most manufacturers offer PCR buffers that do not contain MgCl2 and supply additional vials of MgCl2 so you can optimize Mg2+ concentrations in the lab.

The tricky thing is that the direction of MgCl2 adjustment really depends on the assay. While increasing Mg2+can increase yield and lowering Mg2+ can increase specificity for some PCR products, the opposite can be true for other assays. Performing a MgCl2 gradient in 0.5 mM increments is the fastest way to identify the most optimal concentration! If this seems too tedious, you may


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want to try adjusting other PCR parameters first, such as annealing temperature (Ta) (see Decrease the annealing temperature).

Note that sometimes extra MgCl2 is needed if the DNA sample contains EDTA (a metal chelator) or is contaminated with proteins or other PCR inhibitors that sequester Mg2+ ions.

Deoxyribonucleotides (dNTPs) are essentially the individual bricks that build the DNA strand during each round of amplification. A final concentration of 0.04–0.2 mM of each dNTP is generally recommended for PCR.

If you are amplifying a particularly long PCR product, you may need to increase the dNTP concentration. Remember that magnesium can bind to dNTPs, so if you increase the concentration of Mg2+, you may also need to increase the dNTP concentration, and vice versa.

Avoid using a higher dNTP concentration than needed. Too many dNTPs in the reaction can reduce the fidelity of the DNA polymerase, increasing the error rate and resulting a less clean product. Very high dNTP concentrations can inhibit the DNA polymerase entirely.

Equal amounts of dNTP (dATP, dCTP, dGTP and dTTP) stock solutions can be pre-mixed to a working concentration (e.g. 10 mM) in nuclease-free water. It is not recommended to store dNTPs for extended times at concentrations of <10 mM, as this affects dNTP stability and can lead to hydrolysis of the nucleotide phosphate group. Hydrolysis of the dNTP will render it useless!

Working solutions of dNTPs should be stored in small aliquots at –20°C. It is important to avoid multiple freeze/thaw cycles,

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since freezing and thawing alters the pH and reduces dNTP stability.

Gel electrophoresis

The routine way to visualize a PCR product is to run it out on an agarose gel by electrophoresis (a 2% gel is sufficient for products of 50–1000 base pairs [bp]). Imaging this gel enables you to check the specificity and (to some extent) the efficiency of the primers and the PCR reaction.

The gel is prepared using agarose powder, a buffering solution (typically Tris-acetate-EDTA [TAE] or Tris-borate-EDTA [TBE]) and a staining compound such as ethidium bromide (0.5 μg/ml EtBr – there are safer alternatives on the market, yet EtBr is still commonly used). EtBr is a fluorescent stain which intercalates strongly with double-stranded DNA and is visualized upon exposure to ultraviolet (UV) light. PCR products are typically run next to a defined molecular weight (MW) DNA ladder so that the size of the stained bands can be estimated.

If you don’t see any PCR product when you image the gel, there could be an issue with the gel itself.

EtBr was not added to the gel

If you forget to add the EtBr, then the gel will be completely dark under UV light (no bright bands, including the ladder lane, will be visible). But all is not lost! The gel can be post-stained with EtBr (0.5 μg/ml in buffer or water). Remember that you will need to dispose of the EtBr solution safely after you’ve soaked your gel in it.


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The gel was run for too long or at too high a voltage

It is possible that the PCR products simply ran off the gel. If you can’t see the loading dye anymore, this may well be the case. Next time, run the gel for less time and/or turn down the voltage. If the product size is very small, increasing the percentage of the agarose in the gel can help. You may also want to consider switching to polyacrylamide gels instead.

It is also worth double-checking that the electrode leads were plugged into the power pack the correct way around! Often the wires will be color-coded to help prevent such a mistake (i.e. red-red; black-black). If the leads have gotten switched, then the products and ladder will very quickly run off the top of the gel into the buffer; a rookie error you will only make once!

The EtBr ran off the gel

Over a long run time and/or at a high voltage, the EtBr can run off the top of the gel. This may also occur if you are re-using a gel. EtBr migrates towards the negative (cathode) electrode, and thus becomes depleted at the bottom of the gel, closer to the positive (anode) electrode. If you suspect that this has happened, then you can post-stain the gel with EtBr to visualize the product (see EtBr was not added to the gel above).

Tip: If you intend to re-use a gel, use the bottom portion first; then you can use the top portion later on without compromising EtBr stain.

The PCR sample did not remain inside the well

DNA molecules are not heavy enough to drop into the well of a gel and stay there without diffusing into the gel running buffer. You therefore need to “weigh down” your samples by mixing them

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with a loading dye (usually containing glycerol) that will allow the sample to sink down into the well. Some PCR buffers contain a loading dye, and these products can therefore be pipetted directly into the wells. Otherwise, you will need to combine your post-PCR samples with loading dye at a 1:1 ratio before loading them into the wells.

If the gel did not set for long enough, the wells can collapse when the comb is removed. Conversely, if the gel is left to set for too long, it can dry out and shrivel up! In both cases, the wells are likely to lose their shape, which makes pipetting solutions into them very tricky.

When preparing a gel, leave it to set for at least 45 minutes and don’t allow it to sit for more than 3 hours. If you make a gel ahead of time, either pop it in buffer until ready to use that day, or wrap it in paper towels soaked in buffer, then in plastic wrap and store at 4°C. It should keep well for 2–3 days in the fridge.

DNA polymerase

That tiny amount of enzyme that comes in the brand new vial from your favorite biotech company is the most expensive ingredient in the PCR reaction. If you have a problem with your PCR, it is cheaper to try optimizing other components of the PCR reaction first. However, if you have tried everything else and are still getting no PCR amplification, then this may be due to problems with the polymerase. Here are a few things to keep in mind when working with DNA polymerase:

Before use

Polymerases are very sensitive to temperature and require careful handling. DNA polymerase is stored in glycerol and will not


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freeze – it does not need to be thawed. Always place the enzyme vial ON ICE when in use, particularly if you are using a non-hot-start Taq. Before use, briefly centrifuge the tube and then mix gently using a pipette to eliminate any concentration gradient due to separation of the enzyme and glycerol. Never vortex the enzyme – this can cause it to denature!

During use

Ensure that the ratio of enzyme to reaction volume is at least 1:10; this prevents excess glycerol from interfering with the PCR reaction. Make sure that the enzyme is added to water containing PCR buffer (don’t add it straight to unbuffered water as this can denature it).

Get into the habit of adding the DNA polymerase to the master mix last; if there’s a slip-up along the way, expensive enzyme is not wasted! When adding the polymerase to the master mix, mix very gently by pipetting to minimize the risk of denaturing the enzyme.

If you are using a proof-reading polymerase such as Pfu, remember that it will take longer to elongate the DNA strand compared to Taq polymerase. Typically, you need to allow 2 minute per 1000 bp for elongation by Pfu. Pfu also has 3’–5’ exonuclease activity, which can result in the degradation of PCR primers. To avoid this, be sure to always add the enzyme to the master mix last.

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GC-rich targets

PCR products containing high levels (>70%) of G and C nucleotides can be difficult to amplify. These products can form unusual secondary structures (such as hairpin loops) that prevent the DNA strands from melting completely, therefore making it difficult for the DNA polymerase to proceed.

If you are working with a GC-rich region and are having trouble amplifying anything at all, try these options:

• Increase the time of the initial dissociation period at the beginning of the PCR run: while 5 minutes at 94–96°C is sufficient for most templates, increasing dissociation time to up to 15 minute can be helpful for GC-rich strands.

• Use a PCR additive such as 5% dimethyl sulfoxide (DMSO), 5% glycerol, 1–2 M betaine or 7-deaza-2’-deoxyguanosine (use at a 3:1 ratio relative to dGTP).

• Use Pfu or another high fidelity DNA polymerase: these polymerases are highly thermostable and tackle unstable or complex sequences better than Taq polymerase. The denaturing temperature can be increased up to 98°C.

• Increase the Ta and reduce the annealing time: this step will reduce the formation of inappropriate secondary structures by preventing semi-complementary regions from binding to one another.

• Try Touchdown PCR (see Touchdown PCR and Stepdown PCR), which starts off with a high Ta followed by incremental reductions in Ta, giving the primers opportunity to anneal at the most optimal temperature.


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• Nested PCR (see Nested PCR) may also be a good alternative, as this technique improves sensitivity by honing in on a specific genomic region first and then amplifying the desired product from that point on.

Primers

Primers are enormously critical to the success of a PCR experiment. These short sequences can cause major headaches when they don’t perform the way we expect. In addition to the sequence, the storage conditions and the PCR reagents we use with primers are fundamental to their function. Here are a few things you should know about primers: Primer melting and annealing temperature

The melting temperature (Tm) of both primers should be closely matched (i.e. no more than 1–2°C different) for optimal PCR amplification. If the Tm of your reverse primer varies widely from that of your forward primer, then the primers will not anneal to the template strand at the same time, so no product will be generated.

Usually the primer design software will compute the Tm for each primer for you (there are a number of formulae to do this), but a quick manual check can be performed using the following equation:

Tm (°C) = 4 x [C + G] + 2 x [A + T]*

The optimal Ta should be ~3–5°C below the calculated Tm, and is usually between 50°C and 65°C. The best way of precisely

*useful for primers <25 bp in length.

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identifying this is to assess multiple Ta temperatures in a range of 10°C under the Tm. If possible, use a PCR machine that has a temperature gradient function to save time. Testing in 2°C increments from this (i.e. 52°C, 54°C, 56°C, 58°C and 60°C) should provide a clear-cut optimal temperature.

SNPs in the primer sequence

Single nucleotide polymorphisms (SNPs) or mutations in a primer sequence (particularly at the 3’ end) should be avoided, as they can affect primer annealing. Depending on the position of the SNP, this won’t necessarily prevent amplification; however, a SNP can significantly affect overall PCR efficiency (and also specificity). Thus, PCR bias resulting from a high frequency SNP may account for apparent differences in gene expression levels. If you can’t avoid a SNP, then try to position it in the middle of the primer sequence to minimize effects on annealing.

Another solution is to design degenerate primers. For example, if the sequence contains a C/T SNP, the degenerate base Y can be inserted to give an equimolar mix of two possible primer sequences containing the C or the T base. However, this solution may still lead to some PCR bias, as the two slightly different primer sequences might have marginally different Tms and therefore Tas.

Depending on the study question (e.g. qualitative versus quantitative) the best solution might be to substitute the SNP base with an entirely different base (i.e. in the sample above, add an A or G). While this base won’t be complementary to the template, the primer will anneal with the same efficiency (reduced or not) in every individual. This eliminates potential PCR bias between individuals of a specific genotype.


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Primer storage

Short oligonucleotide sequences, like any nucleic acids, are vulnerable to multiple freeze/thaw cycles. The best practice is to re-suspend lyophilized stock primers to a concentration of 100 μM, store at –20°C and then dilute to your working concentrations in nuclease-free water for each new experiment. If you plan on using a primer set frequently (for instance for a reference gene), then you could prepare multiple aliquots of working solution to avoid excessive freeze/thaw cycles.

Polymerase-specific considerations (Pfu)

Pfu and other polymerases with 3’–5’ exonuclease activity can chew up primers during a PCR reaction. To counteract this, try increasing the primer concentration that you’re using in the assay. You could also increase the length of the primers (say to about 35 bp), so that they will still anneal efficiently even if a few bases are missing from the 3’ end. As a last resort, you may want to consider using nuclease-resistant primers (e.g. phosphorothioate primers), which will not be degraded by Pfu.

Sometimes primers just don’t work

However much we may strive to design ‘good’ primers, or spend hours playing around with the PCR reaction to improve primer specificity and efficiency, sometimes diligence just isn’t enough. The good news is, primers are cheap! So don’t be afraid to bin a ‘bad’ set of primers and start over – this may be the least expensive option in the long run and save you valuable lab hours!

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Sample quality

The quality of the sample (containing the DNA/RNA template) will significantly affect the success of the PCR experiment. It is crucial to take good care of DNA/RNA samples by reducing risk of contamination from PCR inhibitors and taking steps to avoid template degradation.

PCR inhibitors

PCR inhibitors are a common problem and can lead to failure of downstream PCR applications. Common sources include biological inhibitors found in the sample itself (such as blood or soil) or exogenous inhibitors that are introduced during the nucleic acid extraction (such as salts, ethanol, detergent or phenol). These compounds can bind to nucleic acids and will impede PCR amplification either directly or by interfering with DNA polymerase or co-factors.

If you suspect that your template solution contains PCR inhibitors, try these options:

• ‘Clean up’ the RNA/DNA using a commercially available nucleic acid purification kit, such as this one from Zymo Research, which specifically removes common PCR inhibitors from the sample.

• A PCR additive can be used in the PCR reaction itself to enhance the performance of the DNA polymerase and improve overall PCR kinetics. Examples include high quality bovine serum albumin (BSA; 0.1–0.8 μg/μL), betaine (1–2 M), or 5% formamide, glycerol or DMSO.

http://www.zymoresearch.com/rna/rna-clean-up/rt-pcr-inhibitor-removal/onestep-pcr-inhibitor-removal-kit

http://www.zymoresearch.com/rna/rna-clean-up/rt-pcr-inhibitor-removal/onestep-pcr-inhibitor-removal-kit


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• Simply diluting the DNA in nuclease-free water can attenuate the inhibitors and permit PCR amplification.

• If all else fails, reconsider your choice of DNA polymerase, as some types are more sensitive to inhibition than others.

Degraded template

A degraded template is one of the most likely causes for a failed PCR experiment. RNA can be a particularly challenging material to work with. Unlike double-stranded DNA, which is more stable and able to withstand lengthier processing times and varying storage temperatures, RNA can degrade rapidly. Poor quality or degraded RNA will be less efficiently transcribed to cDNA and lead to potential problems in amplification during the PCR reaction. When working with RNA and DNA templates, there are some fundamental ways to maintain nucleic acid integrity:

• Keep RNA samples ON ICE at all times when working at the bench (including during transfer from the bench to the freezer).

• Minimize freeze/thaw cycles as much as possible. If you are working with the same DNA sample routinely, it is less damaging to store it at 4°C during this time than repeatedly freeze/thawing. The alternative is to aliquot RNA or DNA samples so that only some of the sample is compromised each time it is stored.

• Avoid RNase/DNase contamination. The lab environment contains abundant levels of RNases and DNases, which can chew up your template. Before working with nucleic acids, reduce this hazard by: wiping the bench thoroughly with RNase/DNase decontaminators; wearing clean gloves and

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changing them regularly; using RNase/DNase-free plastic; using autoclaved glassware and utensils; and diluting samples in nuclease-free water or nuclease-free TE buffer.

How can you check RNA/DNA quality? There are three common ways in which this can be done:

• Gel Electrophoresis. RNA with high integrity will appear as two strongly visible bands, each representing 28S and 18S ribosomal RNA. Degraded RNA will appear as a lower MW white streak. Genomic DNA contamination can be identified by the presence of high MW bands.

• Agilent Bioanalyzer. This is the gold standard method for assessing RNA quality. The Bioanalyzer separates RNA molecules by size, and also uses fluorescent dyes that intercalate with the RNA. The intensity of fluorescent output after separation is converted into peaks which are visualized on an electropherogram. An RNA Integrity Number (RIN; range 0–10) is assigned to each sample based on all of the measurements taken. Generally, an RIN of ≥7 represents high quality RNA that is acceptable for quantitative applications such as microarrays or qPCR.

• Spectrophotometry. A common method of measuring nucleic acid concentration and purity is using absorbance readings at 230 nm, 260 nm and 280 nm. Where absorbance at 260 nm measure RNA/DNA concentration, readings at 230 nm and 280 nm measure contaminants such as salt and protein. Thus, nucleic acid purity can be calculated by the 260 nm/230 nm and 260 nm/280 nm ratios which should ideally be >1.7 or between 1.8 and 2.2, respectively. Note, though, that this method cannot distinguish between RNA and


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DNA molecules and does not provide an indication of RNA integrity.

Weak bands

If you see very faint bands on your gel, then the good news is that the PCR has worked, although perhaps not optimally! Fortunately in this case you have a starting point which you can work with.

Reaction components

Increase the template concentration

An increased amount of input DNA will increase the initial number of template copies for DNA polymerase to synthesize, which may be especially useful if the template is of low quality or degraded. Thus, increasing the template concentration may result in improved amplification of a weak product. However, too much DNA can overload the reaction and lead to non-specific amplification.

So how much template is enough?

• Genomic DNA: As a rule, 100 ng is usually more than adequate for routine PCR. Depending on the experiment, you can vary this concentration between 1 ng and 1 μg of DNA. If you are performing multiplex PCR, you will need to use at least twice as much input DNA.

• cDNA: When amplifying RNA targets, you should add 5–40 ng cDNA reversed transcribed from 10–200 ng RNA for good results. However, if only a small amount of precious template is available and the PCR assay is well-optimized,

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you can get away with adding lower quantities of cDNA to the PCR.

Check the primer concentration

Primer concentrations between 0.1 and 0.5 μM usually provide optimal template binding efficiency and specificity. If the amplification efficiency is poor (i.e. you get only a weak product), you can increase the primer concentrations up to 1 μM; but remember that this can compromise specificity, as there will be less competition for binding to the intended target.

Increase the amount of DNA polymerase

If PCR amplification is low, increasing the concentration of DNA polymerase may improve this; however, adding too much enzyme can lead to synthesis of non-specific amplicons.

Use a PCR additive

PCR additives can reduce the formation of secondary structures, enhance the efficiency of DNA polymerase and reduce the effect of PCR inhibitors. Commonly used additives include BSA (0.1–0.8 μg/μL), betaine (1–2 M), or 5% formamide, glycerol or DMSO.

Non-optimal PCR program

In addition to optimizing the PCR reaction components, there are also several tweaks you can make to the PCR program to improve amplification of a weak product.


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Decrease the annealing temperature

Decreasing the Ta will increase primer binding efficiency and therefore increase the amount of product that is amplified. It is possible to obtain abundant amplification at 54°C when only a faint product is generated at 56°C, so it is worth taking the time to assess multiple Tas. However, this strategy is only useful if there is one specific weak band present in the first instance, as it will also increase the amplification of non-specific products.

Adjust the extension time

The length of time and temperature required for extension depends on the type of DNA polymerase used, and manufacturers usually recommend fairly conservative extension times. Taq polymerase has a relatively high processivity and thus the rule of thumb is to allow 1 minute per 1000 bp at 72°C.

Given that most amplicons in conventional PCR are ≤500 bp, 30 seconds is usually quite sufficient. If you are using a proof-reading enzyme such as Pfu, the extension time needed will be longer, and the program should be adjusted accordingly.

Remember that the extension rates recommended by polymerase manufacturers are probably an over-estimation. Therefore, reducing the extension time might avoid the amplification of non-specific larger MW products. For example, if your desired product is 150 bp but you also obtain erroneous amplicons of 1000 bp, you can reduce the extension time to 15–20 seconds to prevent the larger product from being amplified.

On the other hand, if you suspect that low levels of amplification are due to incomplete extension (particularly for longer length amplicons), then increasing the extension time

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may help. However, DNA polymerases are extremely efficient, and it is probable that alternative optimization efforts will be more fruitful.

Increase the number of PCR cycles

If you are amplifying low abundance targets (or a small amount of template DNA) then increasing the cycle number can increase PCR yield. That said, the PCR will generally max out (exit the exponential phase) and enter the plateau (linear) phase between 20–40 cycles. In addition, after 35 amplification cycles, the reaction components including DNA polymerase and primers will be exhausted and the PCR efficiency will become very low. Therefore, simply increasing the cycle number to obtain more PCR product might not be the best option. Increasing the cycle number can also lead to the amplification of non-specific products – check this by running the PCR product on a gel.

Despite our best efforts, sometimes these little tweaks to the conventional PCR program won’t lead to the amplification levels needed. By using an alternative PCR method, such as nested PCR (see Nested PCR), it may be possible to increase the sensitivity of your PCR experiment.

Extra bands

Additional, and unwanted, bands on the gel could be due to primer dimer formation, non-specific primer binding, specific (but unintended) primer binding or contamination from the lab environment. How might each of these arise and what can we do to combat it?


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Primer dimer formation

Primer dimer formation happens often in PCR – this is when the two primers anneal to each other due to complementarity at the 3’ termini. These tiny duplexes are then extended by DNA polymerase, resulting in a faint band that appears at around 30–50 bp.

The hallmark of a primer dimer “product” is that it appears in all lanes including the no template control (NTC). This band will typically be stronger in the NTC lane than the template-containing lanes, as there will have been no competition with the template for PCR reagents.

Checking primer complementarity in silico before ordering your primers (e.g. using IDT’s OligoAnalyzer tool) is a useful step, although it doesn’t guarantee that a primer dimer won’t form. If you do end up with a primer dimer, it can often be resolved by slightly increasing the Ta. Minimizing primer dimer formation will also free up the PCR reagents for a more specific and efficient reaction.

Unknown PCR product

Amplification of unknown, or non-specific products, is a common problem in PCR, and is virtually always due to lack of primer specificity. It is extremely worthwhile to determine the potential non-specific binding of the primers to the background genomic template or to other mRNA species, before purchasing the primer set. You can do this quite simply in silico using the NCBI Basic Local Alignment Search Tool (BLAST) tool.

Even if you do check primer specificity ahead of ordering, sometimes spurious bands will still appear on the gel (hence the

http://sg.idtdna.com/analyzer/Applications/OligoAnalyzer/

http://blast.ncbi.nlm.nih.gov/Blast.cgi


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necessity for in-lab optimization!). This could be due to the primers binding to undefined sequences or some type of contamination of your sample. There’s good news though: chances are you can still “rescue” your PCR by adjusting some of the PCR program parameters:

• Increase the Ta; this increases the stringency of primer binding, so they should only anneal to exact matches in the template.

• Decrease the primer concentration so there are fewer opportunities for these sequences to bind to the incorrect target (this will also reduce the chance of primer dimer formation).

• Reduce the extension period, thus decreasing the time the DNA polymerase has to extend non-specific products (if they are larger than the desired product) (see Adjust the extension time).

• Reduce the amount of DNA template added to the PCR reaction to achieve a cleaner product. Too much DNA can literally ‘get in the way’ of the polymerase, impeding its access to the target sequence.

• Reduce the number of PCR cycles – it is possible that the non-specific products are amplified after the desired amplicon. This tactic can be particularly effective in combination with reducing the amount of DNA template added to the reaction.

If none of the above work, using alternative PCR methods such as nested PCR (see Nested PCR) or touchdown PCR (see Touchdown PCR and Stepdown PCR) may be more successful.


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Bands appearing in negative controls

If you see one or more bands in the negative control lane, then you have contamination of some kind. You won’t be able to salvage your PCR if the controls are contaminated, but you can take steps to avoid encountering this problem in the future.

Environmental contamination

If you detect amplification in the no template control (NTC), this is most likely due to contamination of the PCR prep environment. PCR products are remarkably stable and can ‘float’ around in the air or on the bench, presenting a major source of potential contamination for the next PCR experiment.

If laboratory space allows, it is best to assign a separate pre-PCR and post-PCR area, or even better to set up the PCR in a separate room all together. This simple step may be the difference between a clean PCR and one containing a number of spurious products. Before starting the experiment clean the workspace with bleach, wipe down pipettes with ethanol and wear fresh gloves. It is a good idea to have clean ‘PCR-only’ lab coats for use in the assigned PCR area.

Another tip is to prepare the control samples (containing no sample) before you open a tube containing sample, thus lowering the risk of any cross-contamination. If you continue to see contamination in you PCR reactions, it’s best to throw away the old reagents and use fresh reagents – this will save you time and money in the long run!

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Genomic DNA contamination

When measuring mRNA expression levels, you should always include a reverse transcription control (RT NEG). This control indicates whether there is genomic contamination in the RNA (cDNA) sample. A spurious band in the RT NEG lane (but not the NTC lane) could indicate specific binding to the genomic equivalent, or non-specific (random) products. Increasing the Ta will favor more specific primer binding and may eliminate the RT NEG band.

Careful primer design will help you avoid encountering this problem in the first place. By designing exon-exon (coding sequence) primers you know that your product must be synthesized from an mRNA and not a genomic template. Alternatively, exon-intron primers will amplify genomic template only.

Alternative PCR methods

Several alternative PCR techniques have been developed to help amplify tricky templates. In this section, we’ll discuss what applications these techniques are useful for, and how to troubleshoot any problems that may still arise.

Nested PCR

Nested PCR is a technique that efficiently eliminates non-specific amplification. This method is particularly useful when amplifying a sequence that is highly homologous to other gene regions (e.g. retroviral DNA). Nested PCR can also vastly improve PCR sensitivity, which is useful for poor quality starting templates, GC-rich regions and low abundance targets.


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Nested PCR uses two primer sets for two sequential rounds of PCR (Figure 1). The first set of ‘outer’ primers narrows down the broader region to be amplified, and this first round of PCR includes a moderate number of cycles (~20–25). A small amount (e.g. 1:100 dilution) of the PCR product generated from this first round of PCR is then used as the template for a second round of PCR, using an additional set of internal PCR primers (inner primers) that target a shorter region. The inner primers anneal to the first-round template and amplify this short product over ~20–25 cycles.

The use of a second set of primers and the reduced number of cycles help to discriminate against any non-specific binding that may have occurred during the first round of PCR. This results in more efficient amplification of a specific target product. Hemi-nested PCR, a variation on nested PCR, uses one inner primer in the second round of PCR rather than two, and thus still produces a shorter second round amplicon (Figure 1).

Nested PCR trouble-shooting:

1. Primer dimer formation. Avoid the design of primer dimer-prone oligonucleotides, i.e. avoid complementarity at the 3’ ends. You can also try increasing the Ta in the first and/or second round of PCR.

2. Presence of false positives after the second round of PCR. Avoid cross-contamination of PCR products (e.g. different samples/individuals) between first- and second-round PCR.

3. Presence of product after first-round PCR, but not second-round PCR. Ensure that the DNA from the first-round PCR was diluted (i.e. there is no DNA overload).

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FIGURE 1: Nested and hemi-nested PCR. In nested PCR, the region of interest and flanking DNA is amplified using a pair of outer primers that are complementary to the flanking DNA. This product is then used as the tem-plate for a second round of PCR in which the region of interest is amplified using a second set of inner primers. In hemi-nested PCR, a DNA fragment containing the region of interest and the flanking DNA is amplified as in nested PCR. The region of interest is then amplified using one of the outer primers and an inner primer that is complementary to the region of interest.


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4. Presence of multiple products after first-round PCR. Increase the first-round PCR Ta. Alternatively, run the first-round PCR out on an agarose gel, excise the band corresponding to the correct MW and gel-purify the product for use in second-round PCR.

Touchdown PCR and Stepdown PCR

Touchdown PCR enhances primer binding specificity in the early rounds of PCR amplification and then propagates continued synthesis of the specific PCR product in subsequent PCR rounds. In addition to being beneficial when primers are not target-specific, touchdown PCR may also be effective when the template presents difficulties for primer binding, for example, over GC-rich regions.

To perform touchdown PCR:

1. Start with as high a Ta as possible that still allows the primer to bind to the template (for example, 10°C above the calculated Tm). This yields a specific (and hopefully the desired) PCR product.

2. Next, decrease the Ta sequentially over subsequent rounds of PCR (e.g. 1°C/cycle for 10–15 cycles) to promote more efficient primer binding and take advantage of the 2-fold increase of the desired product over each cycle.

3. Once a Ta is reached that corresponds to the calculated Tm (or Tm minus 3–5°C), then perform a final block of PCR cycles (e.g. 20–25 cycles) using this Ta to increase the yield of the product.

Because the earliest generated products are very specific and subsequent products are generated from these early templates,

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this will discriminate against the amplification of non-specific regions.

Stepdown PCR is similar to touchdown PCR, except for the programming of one large drop in Ta following the initial high temperature annealing cycles. Stepdown PCR does avoid the time-consuming programming of the thermocycler to incremental changes in Ta, but is less likely to produce as optimal binding specificity and efficiency as touchdown PCR.

If you are having technical problems with touchdown/stepdown PCR, these troubleshooting tips may help!

• Weak product

- Increase the number of final stage (constant Ta) PCR cycles

- Try including a PCR additive (see Use a PCR additive)

• Multiple products or a DNA smear

- Adjust the initial Ta range upwards (e.g. increase the starting Ta by 5°C).

- Reduce the number of incremental stage (different Ta) cycles and/or final stage (constant Ta) cycles.

- Increase the number of PCR cycles per degree change during the incremental stage (e.g. 1°C change/2 cycles).


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Multiplex PCR

Multiplex PCR is a high-throughput tool used in basic and clinical (diagnostic) research settings to amplify two or more PCR products simultaneously in one reaction tube. It is commonly used in genomics to detect large gene duplications and deletions, as well as in microsatellite and SNP genotyping analyses. Multiplex PCR can also be used to detect infectious agents such as viruses and bacteria.

The obvious advantages of multiplex assays are the reduced costs in consumables, reagents and labor, and the increased amount of information obtained from one reaction with the added benefit of using up less precious sample. The data are also more uniform, as the starting template (DNA) is identical for each assay and there is no run-to-run bias.

The major challenges relating to multiplexing are the propensity for non-specific amplification and primer-primer annealing (primer dimer formation). These assays can therefore be extremely tricky to optimize.

While the PCR fundamentals are essentially the same for multiplex assays, combining multiple assays in one tube will alter reaction kinetics unpredictably. This will require some potentially tedious trial and error troubleshooting. Of course, most rules for standard PCR apply to multiplex PCR, but there are some technical features that are specific to this technology.

Multiplex PCR trouble-shooting:

1. Some multiplex products are weak and others are strong. Increase the primer concentration of the weak product and limit

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that of the strong product. This can help reduce the competition for reagents between the different assays. Use primer design software (e.g. PrimerPlex) for identifying primers that are most likely to behave optimally under multiplex conditions.

2. Reaction conditions. Before performing the multiplex PCR, optimize each PCR assay individually to obtain similar amplification efficiencies for each target using the same PCR conditions. This will provide a starting point for the multiplex reaction.

3. Template concentration. Remember to increase the amount of input DNA when performing a multiplex reaction. If you are amplifying two targets, then add twice as much template.

4. Increase the extension time. Since multiple amplicons are synthesized simultaneously, this will lead to competition for dNTPs and DNA polymerase. A longer extension time may be required to allow enough time for full length products to be obtained.

http://premierbiosoft.com/primerplex/index.html


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General PCR troubleshooting guide

To download a printable copy of this table, click here.

Conventional PCR problem

Possible solution

No product • Check that all of the reaction components were added – use a check-list!

• Check that the gel ran normally; if the ladder is not visible, post-stain with EtBr (this may have been omitted initially)

• Check that the primer sequence is complementary to the desired amplicon (using BLAST) and that the correct sequence was ordered

• Check that there are no SNPs at the 3’ termini of the primers

• Check that the thermocycler was programmed correctly

• Run a Ta temperature gradient with a range of at least 10°C

• Increase the extension time to 1 min/1000 bp• Increase the number of PCR cycles • Use a new aliquot of dNTPs (these are vulnerable to

multiple freeze/thaw cycles)• Increase the amount of input DNA• Check the quality of the starting material (RNA/

DNA)• Dilute the sample 1:10 or perform a DNA clean-up

(purification) to remove PCR inhibitors• If the target is GC-rich, add a PCR enhancer to

reduce formation of secondary structures e.g. 5% DMSO, glycerol or betaine (1–2 M), or try a 1:3 ratio of 7-deaza-2’-deoxyguanosine (a dGTP analogue)

• If none of these work: design new primers

http://bitesizebio.s3.amazonaws.com/wp-content/uploads/2014/10/Download-Table.pdf

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Possible solution

Weak product • Decrease the Ta• Run a Ta temperature gradient with a range of at

least 10°C• Increase extension time to 1 min/1000 bp• Increase the number of cycles• Use higher primer concentrations• Use higher DNA polymerase concentration• Adjust MgCl2 concentration• Check the quality of starting material (RNA/DNA)• Increase the amount of input DNA• Add a PCR additive, e.g. DMSO, formamide, glycer-

ol, BSA or betaine

Extra products: non-specific amplification

• Check primer specificity using BLAST • Increase the Ta• If the non-specific products are larger than the ex-

pected product, decrease the annealing and exten-sion times

• If the non-specific products are smaller than the expected product, increase the annealing and exten-sion times

• Decrease primer concentrations• Decrease DNA polymerase concentration• Adjust MgCl2 concentration• Use Hotstart Taq• Reduce the amount of input DNA• Reduce the number of PCR cycles• If none of these work: design new primers

Extra products: primer dimer for-mation

• Avoid 3’ complementarity when designing primers• Use lower primer concentrations• Increase the Ta


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Possible solution

Extra products: amplification of negative controls

• Check that the PCR parameters are strict enough (i.e., don’t use 50 cycles when 35 is enough)

• Set up the PCR reaction in a ‘clean area’ (don’t allow PCR products to come into contact with this area)

• Always wear fresh gloves and a clean lab coat• Use separate allocated centrifuges and pipettes for

pre- and post-PCR reactions• Use aerosol-resistant (barrier) tips• Prepare control reactions first and close tube lids

before bringing DNA to the work area• Use clean water that is stored only in pre-PCR area

and used only for PCR preparation• If none of these work: replace in-use reagents one

by one to determine if any are contaminated (many companies will give free samples which you can use to test if it is your Taq or other expensive reagent before you buy a new one/throw it out)

• If you see amplification in the RT negative control, treat the RNA with DNaseI

Useful References1. Dieffenbach CW, Lowe TM and Dveksler GS. (1993) General concepts for

PCR primer design. PCR Methods Appl. 3:S30–7.2. Elnifro EM, Ashshi AM, Cooper RJ, Klapper PE. (2000) Multiplex PCR: optimi-

zation and application in diagnostic virology. Clin Microbiol Rev. 13:559–70.3. Goode T, Ho WZ, O’Connor T, Busteed S, Douglas SD, Shanahan F, O’Con-

nell J. Nested RT–PCR: Sensitivity Controls are Essential to Determine the Biological Significance of Detected mRNA. In O’Connell J, editor. Methods in Molecular Biology, RT-PCR Protocols. Totowa, NJ: Humana Press Inc.; 2002. pp. 65–79.

4. Grunenwald H. Optimization of Polymerase Chain Reactions. In Bartlett JMS, Stirling D, editors. Methods in Molecular Biology, PCR Protocols. 2nd ed. Totowa, NJ: Humana Press Inc.; 2003. pp. 89–99.

5. Haff LA. (1994) Improved quantitative PCR using nested primers. PCR Meth-ods Appl. 3:332–7.

http://genome.cshlp.org/content/3/3/S30.full.pdf

http://genome.cshlp.org/content/3/3/S30.full.pdf

http://cmr.asm.org/content/13/4/559.full

http://cmr.asm.org/content/13/4/559.full

http://www.ncbi.nlm.nih.gov/pubmed/7920237

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6. Henegariu O, Heerema NA, Dlouhy SR, Vance GH, Vogt PH. (1997) Multiplex PCR: critical parameters and step-by-step protocol. Biotechniques. 23:504–11.

7. Korbie DJ and Mattick JS. (2008) Touchdown PCR for increased specificity and sensitivity in PCR amplification. Nat Protoc. 3:1452–6.

8. Mamedov TG, Pienaar E, Whitney SE, TerMaat JR, Carvill G, Goliath R, Subra-manian A, Viljoen HJ. (2008) A Fundamental Study of the PCR Amplification of GC-Rich DNA Templates. Comput Biol Chem. 32: 452–7.

9. Mülhardt C. The Polymerase Chain Reaction. In Mülhardt C, Beese E. Mo-lecular Biology and Genomics. Waltham MA: Academic Press; 2007. pp. 65–94.

10. Roux KH. (2009) Optimization and troubleshooting in PCR. Cold Spring Harb Protoc. 4:pdb.ip66.







http://cshprotocols.cshlp.org/content/2009/4/pdb.ip66.full

Chapter 3 Quantitative real-time PCR

(qPCR)

Image source: Stuart Caie. Licenced under CC BY 2.0.

https://www.flickr.com/photos/kyz/3340435836

http://creativecommons.org/licenses/by/2.0

Quantitative real-time PCR (qPCR)

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Quantitative real-time PCR (qPCR) is a highly sensitive tech-nique that has revolutionized our ability to examine gene

expression dynamics. qPCR is most commonly used for mRNA detection. However, qPCR is also useful for a variety of other appli-cations, including studying miRNA expression, determining copy number and genotype variation between individuals, or detecting viral or microbial loads in clinical or agricultural samples.

The key strength of qPCR is its ‘real-time’ function. By measuring the amount of PCR product present during the log-linear phase (when the PCR product theoretically doubles with each cycle) instead of at the very end of the reaction, it is possible to directly and quantitatively compare amplification between experimental samples.

Hundreds of sophisticated commercial products and kits are available for qPCR-based applications, enormously increasing qPCR data output and reducing the time-consuming optimization needed for each experiment. However, it is still essential to know the qPCR basics to equip yourself with the expertise to troubleshoot an experiment when it doesn’t go to plan.

Hydrolysis (Taqman) probes

Hydrolysis probes, universally known as Taqman probes, are widely used in qPCR. Taqman qPCR is extremely sensitive due to the high specificity that comes from using a probe in addition to forward and reverse PCR primers. A fluorogenic probe is designed that contains a reporter dye (fluorophore) at the 5’ end and a quencher at the 3’ end. Because of the short (20–30 bp) distance between the fluorophore and quencher, most of the fluorescent signal is quenched when the probe is intact. During annealing and extension, the primers and probe bind to the complementary DNA


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sequence, and the DNA polymerase extends the strand towards the probe. Once the DNA polymerase reaches the probe, its 5’–3’ exonuclease activity cleaves the fluorophore attached to the probe, removing it from the proximity of the quencher and permitting excitation. The amount of fluorescent signal emitted is proportional to the amount of PCR product present. The probe is degraded from the strand, and DNA synthesis continues towards the reverse primer to generate a full length product.

Taqman qPCR is highly sensitive and specific, and is useful for working with transcripts that are hard to detect. In addition, Taqman qPCR can be used in multiplexing or SNP-based assays to detect multiple products using different fluorescent dyes. Another advantage of Taqman qPCR is that amplicon length does not influence amount of fluorescence emitted. However, this technique is more expensive than non-probe-based assays.

Dual Hybridization (FRET) Probes

Dual hybridization qPCR is another PCR method that combines probes and sequence-specific primers. This system takes advantage of fluorescence resonance energy transfer (FRET) between two back-to-back fluorophore-labeled probes. The first probe contains a 3’ donor fluorophore, and the second probe contains a 5’ acceptor fluorophore. When the probes anneal to the complementary DNA template, the donor and acceptor are ‘parked’ very close to each other (within 1–5 bases), resulting in energy transfer between the two fluorophores and fluorescent emission from the acceptor. This is when the fluorescence (proportional to the amount of DNA template) is measured. During extension, the probes are degraded from the strand and the forward and reverse primers extend the full length PCR product for use in next PCR round.


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Dual hybridization qPCR shares many of the same advantages as Taqman qPCR: it is highly sensitive and specific, it can be used in multiplexing or SNP-based assays to detect multiple products using different fluorescent dyes and it can accommodate a variety of amplicon lengths without influencing the amount of fluorescence emitted. However, like Taqman qPCR, dual hybridization PCR can be quite expensive compared to non-probe-based assays. In addition, dyes that are suitable for FRET probes may only be compatible with a limited number of qPCR systems. Finally, this technique requires very careful probe design.

SYBR Green Dye

SYBR Green-based qPCR eliminates the need for a probe, as it only uses forward and reverse primers, similar to conventional PCR. SYBR Green is a fluorescent dye that binds indiscriminately to double-stranded DNA. During extension, the dye incorporates into the growing double-stranded PCR amplicon and emits a fluorescent signal that is directly proportional to the amount of PCR product generated. However, because there is no probe, the primer pairs need to be optimized stringently before using them in a qPCR experiment (e.g. by assessing via conventional PCR, optimizing Ta and so on). In addition, a melt curve analysis (see Non-specific amplification) should always be included post-qPCR (this step is programmed into the PCR machine at the start of the run and cannot be added once the PCR run has finished!). This step identifies the melting (or dissociation) temperature at which the double-stranded amplicon denatured, revealing whether one or more products are present based on the number of derivative peaks generated.


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The advantages of SYBR Green-based qPCR are that it is low cost relative to probe-based assays and can be used on any qPCR system. However, it is less sensitive and less specific due to the lack of a probe and the fact that the SYBR Green dye binds to all double-stranded DNA. In addition, the amount of fluorescence emitted is proportional to the length of the amplicon, so if you want to compare the absolute quantity of different genes (e.g. at a specific developmental time point), you will need to design amplicons with very similar amplicon lengths. Finally, SYBR Green-based qPCR is not suitable for duplex or multiplex reactions, as the products are indistinguishable.

In this chapter, we discuss the most common problems that arise during qPCR assay design, reaction preparation, qPCR system set-up and post-run analysis.

No fluorescent signal

If there is absolutely no fluorescent signal at the end of the qPCR run, there is likely a simple technical explanation. To check if any there was any amplification at all, run the post-PCR products out on an agarose gel. This could help to get to the bottom of whether there is a primer issue or another problem with the assay, such as probe design or qPCR set-up. In this section, we’ll discuss a variety of factors that could result in no amplification, and thus no fluorescent signal.

qPCR assay set-up

There are a number of things that can go wrong very easily when setting up your qPCR experiment. Fortunately, most of them are also easy to fix!


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If you see no fluorescent signal at all at the end of your run, ask yourself the following questions:

1. Was the machine set to the correct (corresponding) reporter dye for the qPCR chemistry or assay (for example, ‘SYBR’ for SYBR Green qPCR, or ‘VIC’/’FAM’ etc. for probe-based qPCR)? If this is incorrect, then the fluorescent excitation will be measured at the wrong wavelength and no signal will be captured.

2. Likewise, does the probe type or ‘quencher’ ordered match the programmed qPCR settings? For example, if you order a probe containing a TAMRA (fluorescent) quencher vs. a NFQ-MGB (non-fluorescent) quencher, ensure that the corresponding quencher setting is programmed correctly on the machine.

3. Does the machine support, and is it calibrated for, the chosen reporter dye? Dual hybridization FRET probes for example, can only be used on a limited number of qPCR instruments (e.g. the LightCycler from Roche).

4. Was ‘data collection’ turned on? Check that the ‘Run Method’ program is set for data collection at the end of each cycle. On some Applied Biosystems qPCR systems there is a small square icon below each data-entry slot for program cycling. This icon must be active (bold) at the end of the extension step. If this icon is accidently clicked off (not bold), then data collection at the end of each cycle will be inactivated and there will be no retrievable data at the end of the run.


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Primer/probe design

If you’ve checked all of the qPCR program parameters and everything seems to be in order, there may be a problem with your primer and/or probe design.

SNPs in the primer sequence can affect PCR amplification. A single base polymorphism, particularly near the 3’ terminus of a primer, may disrupt annealing, leading to poor or no amplification.

Conversely, a SNP within a probe can impede annealing to the template and prevent the generation of a fluorescent signal, although PCR amplification will be unaffected (this is why checking the post-PCR product on a gel can be helpful).

To avoid these problems, always check for SNPs in silico using the publicly available NCBI dbSNP database, which can be easily accessed through genome bioinformatics sites such as the UCSC or Ensembl genome browsers. Finally, verify primer products on a gel prior to qPCR to ensure amplification.

Another consideration that will affect the fluorescent signal (although not affecting amplification itself) is the Tm of the probe. For example, the Tm of the probe should be approximately 10°C higher than that of the primers. Typically the Tm of the primer set is around 60°C and the Tm of the probe is around 70°C. Note that the type of quencher used will also determine the probe Tm. Non-fluorescent quenchers containing a minor groove binding (MGB) moiety will have an increased Tm compared to a TAMRA probe.

http://www.ncbi.nlm.nih.gov/projects/SNP/

http://www.genome.ucsc.edu

http://uswest.ensembl.org/index.html


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Plate preparation

The PCR plate set-up procedure is another good place to check for possible problems. Proper handling is crucial for producing good qPCR results, so run through the following questions:

1. Were all the necessary and correct reagents added to the reaction mix? If one of the reagents is missing, the PCR will fail – using a check-list system can ensure that this does not happen.

2. Has the expiry date of the Taqman or SYBR Green buffer passed? These items to tend to have a limited shelf-life. Over-exposure to light will also result in degradation of the SYBR Green dye, so make sure it is stored in the dark at 4°C between uses.

3. Was the correct qPCR buffer used? PCR buffers vary by application and are often specific to particular real-time kinetics. For example, different PCR buffers are used for FAST vs. Standard runs.

4. Was the correct type of PCR plate used? Different 96-well PCR plates are required for FAST and Standard PCR runs, although 384-well PCR plates can be used for either type of run.

5. How soon after preparation was the PCR plate run? It is unlikely that high abundance transcripts will degrade rapidly, so storing the plate overnight should not affect PCR performance. However, if you consistently get an absent or low fluorescent signal, then avoid delays between setting up the reaction and running it. If the plate is not run straight after it is prepared, store it at 4°C away from light.


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Low sensitivity (high CT values)

Low template amplification (indicated by a high cycle number at which the fluorescence threshold is met – the CT value) is a fairly typical qPCR problem and can occur due to low assay sensitivity. High CT values can occur for either biological or technical reasons. Below is a guide for identifying why your qPCR is exhibiting low sensitivity.

Biological samples

Multiple biological factors can contribute to the low sensitivity of your assay. If you suspect that this is this case, try addressing one or more of the following parameters:

1. Did you include a positive control? This will quickly establish whether a high CT value in the experimental sample is due to an inefficient PCR reaction (i.e. the positive control is also exhibiting high CT values), or if there were simply low levels of target template to begin with (i.e. the positive control is showing much lower CT values than the unknown sample).

2. Is the nucleic acid sample degraded? If the reference genes are being amplified late in the reaction, e.g. after 25 cycles, this may indicate that the sample is degraded or of poor quality. It is likely that the amplicon of interest won’t be amplified until after 30 cycles, when quantification is less accurate. qPCR is of little value when working with poor quality samples, as low levels of template reduce the precision of the assay. See Sample quality for a number of ways to improve the nucleic acid quality.

3. Was the appropriate amount of template added? The dynamic range of the assay is the amount of DNA or cDNA that is amplifiable without overloading or underloading the


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reaction. The dynamic range will therefore vary depending on the assay and the expression level of the target amplicon. Between 1 ng and 100 ng of input material is usually adequate. A good starting point is to add 100 ng RNA per 10 μL of reverse transcription reaction (or 200 ng RNA in a 20 μL reaction, and so on). You would therefore add 10 ng template in 1 μL cDNA to the PCR reaction. If the target gene (reference gene or amplicon of interest) is very highly expressed and the CT values are <10 cycles, then you should add less template (e.g. dilute the cDNA). Conversely, if CT values are coming up very late (>30 cycles), more template can be added (e.g. start off with a higher RNA concentration in the reverse transcription reaction).

Primer design

Primer design can affect the sensitivity of your qPCR assay. The optimal amplicon length for both SYBR Green and probe-based qPCR assays is between 60–150 bp. While larger amplicons of up to 500 bp can be amplified, this may negatively impact the efficiency of the PCR reaction. Although this is not likely to lead to absolutely no signal, amplicon length should be a consideration during primer design.

Primer specificity is important in probe-based, as well as non-probe-based, qPCR. While non-specific amplicons will not contribute to the fluorescent signal in Taqman/FRET systems, amplification of these non-specific products will still compete for PCR reagents (e.g. DNA polymerase, dNTPs), therefore reducing sensitivity and efficiency. This will be particularly problematic if the expression of the amplicon of interest is low to begin with.


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qPCR assay set-up

Errors in setting up the qPCR program can lead to an apparent reduction in the sensitivity of the assay. Potential problems with the program include:

1. The number of cycles. Usually the default qPCR software settings are for 40 cycles, but it is worth checking that this is not mistakenly set to a lower number. Increasing the cycle number beyond 40 is unlikely to improve PCR amplification and will likely lead to higher levels of noise and non-specific amplification.

2. Setting the correct baseline. The baseline should include the initial/early cycles in which the background noise is high, before the appearance of the amplification curve. It is important that this is set correctly, because the baseline value will be subtracted from the normalized reporter fluorescence to accurately determine the CT value. After the run, the analysis software will automatically set the baseline using pre-defined mathematical parameters. However, in real life this can often deviate (possibly due to experimental error), so the baseline should be checked and manually adjusted if necessary. Normally, the baseline should be set to between 3 and 15 cycles. If there is a large deviation from this in either direction, this indicates that too much or too little input template was added.

3. The qPCR cycling conditions. Many labs are now choosing to use FAST instead of Standard (universal) PCR conditions to halve the overall run time and increase assay throughput. However, for some assays, using FAST parameters may reduce sensitivity (and increase variability). If your PCR experiment is exhibiting low sensitivity, consider using Standard parameters:


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this will give the DNA polymerase more time to activate, increase the length of each PCR cycle to allow more time for strand denaturing, and provide a longer annealing/extension period. Remember to use the correct qPCR buffer for the chosen conditions.

If you are working with a region that is difficult to amplify or with an unavoidably long amplicon, it can also help to separate the annealing and extension steps and select more optimal PCR conditions, such as adjusting the step times and temperatures (e.g. by programming a specific Ta). These conditions can be optimized using conventional PCR and applied to the qPCR program to improve the sensitivity and precision of the experiment.

Non-specific amplification

Amplifying extra products in addition to the desired amplicon is also a common problem in qPCR. In probe-based assays, non-specific primer-template binding results in lowered sensitivity, as the PCR reagents face competition from the wrong product. In SYBR Green assays, non-specific amplification renders the results completely unusable, as the fluorescent signal does not distinguish between multiple products.

For SYBR Green assays, a dissociation or melt curve must always be included on the SYBR Green qPCR ‘Run Method’ at the end of the PCR cycling. This will determine if more than one product is present in the post-PCR reaction based on the dissociation Tm of the amplicon. If there is a small shoulder to the left of the melt curve plot (Figure 2), this is most likely a primer dimer (the derived Tm is usually ~5°C lower than the desired product). If you are targeting a low abundance transcript, it is even more important to avoid designing primer dimer-prone


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FIGURE 2. Melt curve plots. When performing SYBR Green PCR, it is essential to include a melt curve to ensure that only one PCR product has been generated. If there is only one product in the post-PCR reaction, then only one peak will appear in the melt curve. A small “shoulder” to the left of the peak most likely represents primer dimer formation. Two or more peaks indicate that multiple PCR products were generated, suggesting a lack of primer specificity or contamination with genomic DNA.


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oligonucleotides. If very subtle changes in gene expression are expected, the presence of a primer dimer (which contributes to the overall fluorescent signal) may mask any biological findings.

If two distinct dissociation peaks are present (Figure 2), this signifies that you have amplified two products: either the primers are not specific and are binding another mRNA, or there is genomic contamination leading to inadvertent amplification of the target DNA.

For both SYBR Green and probe-based assays, primer specificity can be determined post-qPCR by running the products out on an agarose gel. This way you can visualize primer dimer and/or non-specific products and determine their MW, which will be useful if primer re-design is required.

Primer dimer formation

Similar to standard PCR, primer dimer formation can also occur in qPCR. To minimize the chance of primer dimer formation interfering with your qPCR assay, during primer design, check that the 3’ end of the forward and reverse primer are not complementary to one another.

If you’ve already ordered your primers and suspect that they may by forming duplexes, use a qPCR buffer containing hot-start Taq polymerase to prevent elongation of the primer dimer before the PCR run starts. Preparing reaction mixes on ice can also minimize primer binding before the PCR program begins.


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Poor primer specificity

Most commonly, extra products in your qPCR run are due to the primers binding to other unexpected targets. It is important to always check primers for specificity on an agarose gel before trying to optimize for qPCR, as this will save you a lot of time and expense. It is also a good idea to check primer specificity in silico using the NCBI BLAST tool. While this won’t guarantee specificity, it is helpful to eliminate oligonucleotides that are likely to have non-specific binding, particularly if you are working with a low complexity region.

If you suspect that your primer pair is amplifying an off-target product, try increasing the temperature of the combined annealing/extension step (the default is usually 60°C) to increase the stringency of primer binding. Alternatively, you can program separate annealing and extension steps and alter the Ta only. Be aware that altering these parameters will also affect the kinetics of other assays such as amplification of the reference gene, so be sure to monitor the efficiency of these reactions as well.

You may also wish to consider using FAST instead of Standard PCR conditions. This will result in stricter PCR parameters (e.g. shorter time for primer annealing and extension), and so may reduce the amplification of unwanted products. Remember to use the correct qPCR buffer for the chosen conditions.

Genomic DNA contamination

Contamination from residual genomic DNA is another frequent cause of extra products in your qPCR run. To avoid this possibility from the outset, design exon junction-spanning primers when



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possible. These primers completely span an intronic region and therefore cannot amplify the genomic equivalent.

If you suspect that your template contains genomic DNA, then you can treat the RNA sample with DNase I to chew up the contaminating DNA.

High degree of technical variation

The level of between-replicate standard deviation (SD) that can be tolerated will depend on the biological parameters of the experiment. If you are amplifying a highly abundant transcript and expect large fold-changes in gene expression between test groups, running duplicate reactions with a SD threshold of 0.3 SD points may be satisfactory. However, there are times when a more conservative approach is needed. A large SD can particularly be a problem when working with low abundance transcripts, such as transcripts that amplify at a high CT (between 30 and 35 cycles). Due to skewed statistical distribution at low copy numbers, there will inevitably be higher levels of between-replicate variation for these targets. In these cases, setting the SD threshold to 0.2 SD points and increasing the number of replicates would be more appropriate.

Likewise, if a very subtle change in gene expression is expected (e.g. less than a 2-fold change in expression), a more stringent approach will generate more reliable and precise results. In addition, increasing the number of replicates (i.e. running up to 3–5 replicates for highly variable samples) will help improve your statistics.


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There are a number of other steps that you can take to reduce between-replicate variation by changing the program set-up and the plate set-up as discussed below.

Program set-up

If you are consistently seeing unacceptably high levels of variability, make sure that you have set the correct threshold (ΔRn level) in log-linear (exponential) phase. Although the analysis software will automatically set the threshold using an algorithm, this can sometimes be inappropriate and should be adjusted manually.

You should also ensure that ROX is selected as a passive reference during assay set-up. The ROX dye acts as an internal passive fluorescent reference to normalize the fluorescent signal of the reporter dye based on various technical parameters, which may include PCR volume, bubbles (dislodged during the run) or pipetting error.

Plate set-up

Many problems with high variability can be resolved with careful handling of the samples and plate during qPCR assay set-up. First of all, check that all the PCR reagents are completely thawed and thoroughly mixed before setting up the reactions to ensure that the reagents are distributed equally.

To reduce pipetting error, use a multi-dispenser pipette to dispense the reaction mix and a multichannel pipette to dispense the nucleic acid template. Eyeball the volumes drawn into the pipette tips each time to ensure consistency during use. It also helps to have the pipettes regularly calibrated. If you have access

http://www.youtube.com/watch?v=qq0l_wyhFZo

http://www.youtube.com/watch?v=qq0l_wyhFZo


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to a robotic liquid handler, which can set up the plate for you, even better!

Carefully seal the plate with the plastic adhesive cover. There should be no visible wrinkles around the edges of the plate, and the neighboring wells should be sealed off from one another. If the plate is not properly sealed, this can result in evaporation, leading to inaccurate and highly variable data.

Finally, ensure that there are no bubbles settled at the bottom of the well. Immediately before running the qPCR plate, spin it briefly at low speed to dislodge any bubbles and bring down any splashes from the well wall.

Poor standard curve

It is common to include a standard curve in a qPCR experiment. The genomic sample (template) you use to generate the curve will depend on the quantification analysis method (e.g. absolute vs. relative).

Absolute quantification measures the exact amount of DNA present in a sample. Standard curves for absolute quantification are prepared from a nucleic acid sample with a known quantity. The curve is used to extrapolate the exact copy number of the target gene present in the sample.

For routine gene expression studies, such as those comparing the expression of a certain target gene before and after treatment, it is common to use relative quantification, where the absolute concentration of the standards is not known.


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For the standard curve method of relative quantification, a standard curve is included for both the target gene and a reference gene. The relative amounts of the target and reference genes are calculated from their respective standard curves, and the expression level of the target gene is quantified by normalizing to the expression level of the reference gene.

The delta delta (ΔΔ)CT method is probably the most widely used method for relative quantification. For this method, standard curves are not run on the plate each time (one of the main reasons researchers like it so much!). The CT value of the target gene is normalized to the CT value of the reference gene to yield ΔCT, and changes in gene expression are measured by comparing to a calibrator (e.g. untreated control sample) to generate the ΔΔCT value. The relative gene expression value of the normalized sample is calculated using the formula 2–ΔΔCT.

If you are consistently obtaining a poor standard curve, this is most likely due to operator error, as standard curves are very sensitive to how the samples are handled. It is also possible to improve your standard curves by changing how the plate and/or program are set up.

Sample handling

For all standard curves, good laboratory practice is paramount, or the precision of the curve will be compromised. For a standard curve to be reproducible and provide accurate results, scrupulous pipetting is essential!


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For best practice, use the following guidelines to set up your standard curve:

1. Use the serial dilution method to dilute stock template.

2. Mix each standard thoroughly (you can vortex them briefly) and refresh the tip between each new standard.

3. Use a consistent pipetting strategy. For example, use the same type of tips each time, pre-rinse the tip in the template solution each time and prepare all the standards in one sitting.

If you continue to have problems, try using an alternative cDNA sample. If the starting template is degraded or the expression of the target is low, then the standard curve will be poor. Keep in mind that efficiencies above 100% are an artifact. If the efficiency is very high (e.g. 125%), this could indicate the presence of PCR inhibitors in the sample. Try ‘cleaning up’ the RNA and starting over.

Program and plate set-up

When designing your standard curve, include at least three replicates for each point in the standard curve. You can increase the quality of your curve by increasing the number of points on the standard curve.

It’s also worth checking that the qPCR software is properly programmed, e.g. that the correct serial factor is selected or the correct dilutions have been input into the program.


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Useful References

1. Bustin SA, Benes V, Garson JA, Hellemans J, Huggett J, Kubista M. (2009) The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem. (2009) 55:611–22.

2. Caplin BE, Rasmussen RP, Bernard PS and Wittwer CT. (1999) LightCycler hybridisation probes - the most direct way to monitor PCR amplification and mutation detection. Biochemica 1:5–8.

3. Didenko VV. (2001) DNA Probes Using Fluorescence Resonance Energy Transfer (FRET): Designs and Applications. Biotechniques 31:1106–21

4. Hilscher C, Vahrson W and Dittmer DP. (2005) Faster quantitative real-time PCR protocols may lose sensitivity and show increased variability. Nucleic Acids Res. 33:e182.

5. Hruz T, Wyss M, Docquier M, Pfaffl MW, Masanetz S, Borghi L. (2011) Ref-Genes: identification of reliable and condition specific reference genes for RT-qPCR data normalization. BMC Genomics 12:156.

6. Livak KJ and Schmittgen TD. (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2-ΔΔCT Method. Methods 25:402–8.



http://www.fluoresentric.com/documents/HybProbe.pdf







http://www.biomedcentral.com/1471-2164/12/156





Chapter 4 Digital PCR

Image source: ddPCR workflow. Image courtesy of BioRad Laboratories.


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Digital PCR (dPCR) is an extremely sensitive method that is used to detect very small amounts of a specific template in a biolog-

ical sample. While various dPCR methods have been developed, they all involve the central approach of isolating a single nucleic acid molecule (DNA or RNA) and PCR amplifying each molecule separately. This method permits the enrichment of very rare variant templates that would otherwise be ‘drowned out’ by the high back-ground level of common wild-type templates.

Early dPCR methods involved fractionation of the sample into thousands of partitions via a microfluidic chip that separates molecules into individual chambers, followed by quantification using real-time PCR. A more recent method is BEAMing (Beads, Emulsification, Amplification and Magnetics), in which PCR primers are pre-coated with magnetic beads, emulsion PCR is performed, and the clonal template copies are pulled out and purified using a magnet. Fluorescent probes are used to distinguish between different alleles, and the amount of each template is quantified by flow cytometry.

Droplet digital PCR

The most cutting-edge form of dPCR is droplet digital PCR (ddPCR). ddPCR has a greater dynamic range than previous digital methods and increases throughput at a lower overall cost. ddPCR involves partitioning the sample into thousands of individual droplets and using fluorescence to quantify the ‘absolute’ concentration of template. In ddPCR, absolute quantification is determined by the proportion of positive droplets in the sample relative to the total number and volume of droplets generated.

QuantaSoft (now owned by Bio-Rad Laboratories) was the first to develop and manufacture a system for this newer droplet-based

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digital PCR method, although other systems (e.g. the RainDrop dPCR system by RainDance Technologies) are now available. Given that the Bio-Rad system is currently the most widely applied third-generation ddPCR system, in this chapter we will provide specific advice for this system, in addition to giving more general pointers for ddPCR.

The Bio-Rad ddPCR system uses a ‘water-in-oil’, or emulsion, approach. A PCR mastermix (Taqman assay and DNA/RNA template) and droplet generation oil are pipetted into separate wells of a multichannel cartridge. The two solutions are then sucked though a channel into a single chamber by a vacuum to generate up to 20,000 tiny (~1 nL) droplets. Each droplet will contain either no target DNA molecules or one or more target DNA molecules, approximating to a Poisson distribution. The individual droplets are then transferred from the cartridge into a 96-well PCR plate and amplified over 35–50 PCR cycles according to the pre-optimized PCR cycling conditions.

At the end of the PCR reaction (end-point/plateau phase), the plate is transferred to a droplet reader, which streams droplets through a channel in single file. The fluorescent signal is detected in each individual droplet, yielding a binary readout – negative (0) or positive (1) – and this information, combined with the known volume of the droplet and the partition number (the number of droplets generated), is used to calculate the concentration of the target DNA. A Poisson correction is applied to the calculation to correct for the presence of more than one target molecule in some droplets.

One of the main advantages of ddPCR, compared to other quantification methods such as qPCR, is that there is no need to generate a calibration curve to quantify the amount of template.

http://en.wikipedia.org/wiki/Poisson_distribution


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Also, ddPCR is more sensitive, and can therefore detect much smaller fold-changes in nucleic acid concentrations than qPCR. Under optimal assay conditions, this makes ddPCR more precise and repeatable than qPCR. However, as ddPCR is still in its infancy compared with the qPCR alternative, deciding on good experimental design (and trouble-shooting problems that arise during set-up and analysis) may be more challenging. In this chapter, we’ll discuss a variety of technical pointers for designing and troubleshooting ddPCR experiments.

Low droplet (partition) count

At the end of every ddPCR run, the reader will provide a droplet count. Since the dynamic range of ddPCR is established by the number of droplets that are generated in an experiment, low droplet counts can reduce sensitivity. Proper handling of the samples and the plates will ensure that the droplet count reaches adequate levels.

Blocked micro-channels

If the channels of the ddPCR system become blocked, this will noticeably reduce the droplet count. Before use, inspect the bottom of the cartridge wells for dust particles, which can easily block channels.

Secondly, add the PCR mastermix to the cartridge before adding the droplet generation oil. Adding the oil first can result in the micro-channels becoming blocked and will reduce the number of droplets that are generated.

Tip: Prepare PCR mastermix (buffer, primers and nucleic acid) to in-

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clude a 10% error volume; e.g. rather than preparing 20 μL sample volume make up a 22 μL volume.

Once you have added the mastermix and the oil, inspect the bottom of the cartridge wells again to see if there are any air bubbles. Remove any bubbles before generating droplets.

Empty wells

Use all of the wells/sample spaces in the droplet generation cartridge. If there are more wells than available samples, then use a commercially available (low-cost) mastermix substitute for non-sample wells. This ensures that the droplets will be partitioned with more even volumes. This is important, as only uniform droplets will be accepted by the analysis software, so oddly-sized droplets will not be counted.

Droplet damage/loss

Damaging the droplets will change their size, again resulting in a low droplet count, so pipette the droplets slowly and with care. If you are using an electronic multichannel pipette, check that the automation speed is not set too fast – keep the speed settings low for optimal control.

Tip: Use good quality and reliable tips that are suitable for the pipette. When transferring the droplets from the cartridge to the PCR plate, use a P200 volume pipette instead of a P20 pipette, as the larger diameter of the 200 μL tips will help prevent droplet damage/loss.


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Release the droplets into the PCR plate using an even speed and avoid touching the bottom of the well with the tip – don’t poke the fragile bubbles with the sharp end of the tip! And finally, when all of the samples have been added to the plate, transfer it gently to the machine… do not vortex or centrifuge the PCR plate!

If the reaction components and set-up have been checked and double-checked and there are no obvious explanations for low droplet counts, it may simply be that the cartridge is defective. The only option is to try again, using a new batch of cartridges.

Low fluorescence amplitude/threshold

Poor ddPCR performance will reduce both the sensitivity and precision of the experiment. While the droplet count affects performance, there are various other factors that can also affect performance, resulting in inefficient PCR amplification.

Poor assay optimization

Like all PCR assays, ddPCR assays must be adequately optimized. The positive and negative droplets should be easily distinguishable by the intensity of the fluorescent signal post-PCR: you should see distinct clouds of positive and negative samples on the read-out (Figure 3). The presence of ‘rain’ between the clouds indicates poor assay optimization, as it is uncertain where some droplets belong. In fact, the formation of compact clouds reflects true output values more so than the maximum number of positive droplets.

Consider the following points when optimizing the ddPCR reaction:

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1. During primer design, check primers for specificity (e.g. by using NCBI BLAST) and avoid sequences that could result in a primer dimer or form secondary structures, or sequences that include a SNP. Primer design for ddPCR should follow the same general rules as for qPCR assays (see Primer/probe design and Primer design).

2. Validate ddPCR assays using conventional PCR (check for specificity), and optimize the Ta using a temperature gradient. You can also optimize Tas (within a narrower range) in the ddPCR experiment itself to find the Ta that provides the best separation of positive and negative droplets. In the example shown (Figure 3), the second Ta in the gradient (57°C) provides the best distinction between positive and negative clouds.

3. If the amplification remains low after optimization, consider increasing the number of PCR cycles.

If you are running a duplex assay, you should first optimize the assays individually and then run them together in the same reaction. There should be no major changes in performance for either assay when run in duplex. If you note a discrepancy in amplitude between duplex and singleplex runs (i.e. one template is preferentially amplified), then further optimization of the duplex reaction will be required, (for example, adjusting the primer concentrations).

Inappropriate fluorescence threshold

The analysis software will automatically set a threshold to distinguish between positive and negative fluorescent signals. At the end of each run, visually inspect the threshold to ensure that the detection level for positive droplets is well above the



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FIGURE 3. ddPCR droplet analysis. Post-PCR analysis of a ddPCR reaction should reveal distinct clouds of positive and negative droplets. The presence of “rain” between the clouds indicates the need for additional optimization. This figure was modified with permission from Dr. Irina Manokhina.

background fluorescence of the negative droplets. This background fluorescence occurs when the fluorescent dye is not completely quenched. If there is not much leeway between the background signal (negatives) and the positive signals, you may be able to increase the amplitude by increasing the number of PCR cycles.

Template concentration

The precision of a ddPCR assay is determined by the mean number of molecules that are incorporated into each droplet. The optimal number is estimated to be an average of 1.59 molecules per droplet. It’s important to remember that generating far more positive droplets than negative droplets will affect the ability of the Poisson statistical adjustment to correctly predict the number of positive droplets containing >1 target copy. Therefore, adding

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either too much template (saturation) or not enough template (no amplification) may significantly affect assay precision.

The amount of optimal starting material for a particular assay can be established by running a series of template dilutions. This will give you a ballpark idea of the optimal amount of template needed to add to achieve good precision. This is nicely illustrated in the example shown (Figure 3); the second highest DNA input provided optimal separation of positive and negative clouds without compromising amplitude and maintaining an adequate droplet count.

Another thing to note is that a high concentration of intact DNA will increase the viscosity of the solution and change the volume of the droplet. As droplets of different volumes will not pass software quality checks and will lower the number of droplets read, you may want to use a restriction enzyme to chop up the DNA into smaller fragments to facilitate uptake into the droplets. First check how intact the DNA sample is by running out on an agarose gel (look for high MW bands), and then treat with restriction enzymes if required. Of course, make sure that the restriction sites are not located within the sequence you want to amplify!

If you are measuring extremely rare variants, you can use an in silico merging format, which will assess one sample across multiple wells and merge the data. For example, if one sample is run across 5 wells, this will theoretically increase the number of partitions to 100,000, thus substantially increasing the dynamic range for that sample.


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High degree of technical variation

In principle, you can measure samples by ddPCR in a single reaction, without the need for duplicates or triplicates, since absolute quantities are ‘counted’. However, in the early days of assay optimization and/or to control for any pipetting inaccuracies, it might be prudent to run each sample as a duplicate. It may also be necessary to include replicates if the droplet count is consistently low or if very rare variants are being detected, to ensure the reliability of the data. High standard deviation between replicates may indicate a problem somewhere in the experimental set-up. If this is the case, try:

1. Running the samples in triplicate.

2. Ensure accurate and consistent pipetting during set-up, and ensure that the sample is well mixed into the mastermix.

3. Check that the foil plate cover is properly heat-sealed and no crinkles are present that might allow sample evaporation.

4. When loading the plate into the PCR machine, lower the lid carefully onto the foil and screw the lid down so that it just touches the foil seal.

False negatives and false positives

Similar to qPCR, it is important to carefully consider what the most appropriate controls for ddPCR are. Including the proper controls will help you rule out false negative and false positive results, which can arise due to a variety of reasons.

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Contamination

False positives may occur due to carry-over contamination from a previous run or sample cross-contamination. To eliminate the possibility of reading these as authentic positive results, always include a basic negative control, which consists of the ddPCR reaction without nucleic acid (the volume is replaced with water). The droplets generated from this sample should produce a compact cloud of negative read-outs (below the threshold for a positive count).

To avoid accidentally generating false positives due to contamination, designate separate pre- and post-PCR areas. Wear a clean lab coat and change gloves regularly to prevent transferring samples unknowingly. If possible, use a multichannel pipette to reduce the chance of inserting a tip into the wrong well.

PCR inhibitors

If you are concerned about the presence of false negatives, be sure to include a positive control for every ddPCR experiment. A positive control can be any sample that highly expresses the target of interest. The droplets generated from this sample should produce both positive and negative read-outs; however, the number of positive droplets should be high.

The presence of false negatives could indicate that your samples contain PCR inhibitors that are impeding the reaction. Try to minimize the presence of PCR inhibitors during nucleic acid extraction steps. If you suspect that your samples are already contaminated, you can dilute them 1:10 to alleviate the effect of the inhibitors, or purify the DNA or RNA sample using a ‘clean-up’ kit.


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Calibration control

Using a calibrator sample with a known concentration can control for the accuracy of ddPCR data, although a calibration curve is not required for quantification. A calibrator sample (can be the same as the positive control) can be run on multiple plates to measure repeatability between runs or to ensure reproducibility if the assay is subsequently run in another laboratory. Including a calibrator is especially important if you are measuring RNA levels, as there is more room for technical variance when using RNA templates.

For further detailed information on using the BioRad ddPCR sys-tem, check-out their Droplet Digital PCR User’s Guide.

A word on dMIQE

As a follow-up to the published Minimum Information for Publication of Quantitative real-time PCR Experiments (MIQE) for qPCR users, Digital (d)MIQE Guidelines have now been established. These guidelines recommend both ‘desirable’ and ‘essential’ data that should be reported on when publishing dPCR data. Adhering to these recommendations will provide a benchmark for high reporting standards for dPCR.

Some of the essential reporting recommendations include:

• Number of droplets (partitions) generated.

• Individual and total droplet volume.

• Mean number of nucleic acid molecules per droplet.

http://www.gene-quantification.de/bio-rad-ddpcr-app-guide-6407.pdf

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• Information on template structure (e.g. single- or double-stranded DNA), where the template originates from and how the sample was processed.

• Intra-assay variation.

• Details of all controls included.

• Inclusion of supplemental data to show the clear demarcation of positive and negative droplets.

Also, don’t forget to include all of the other essential information outlined in both MIQE and dMIQE pertaining to nucleic acid extractions, assay design, PCR optimization and so on!

Useful References1. Hindson BJ, Ness KD, Masquelier DA, Belgrader P, Heredia NJ, Makarewicz

AJ et al. (2011) High-throughput droplet digital PCR system for absolute quan-titation of DNA copy number. Anal Chem. 83:8604–10

2. Huggett JF, Foy CA, Benes V, Emslie K, Garson JA, Haynes R, et al. (2013) The digital MIQE guidelines: Minimum Information for Publication of Quantita-tive Digital PCR Experiments. Clin Chem. 59:892–902

3. Pohl G, Shih IeM. (2004) Principle and applications of digital PCR. Expert Rev Mol Diagn. 4:41–47

4. Mazaika E and Homsy J. (2014) Digital Droplet PCR: CNV Analysis and Other Applications. Curr Protoc Hum Genet. 82: 7.24.1–13.









The PCR journey can be a long one – dealing with troublesome PCR assays can be a tortuous endeavor, requiring a lot of

thought and time. As we have seen over the course of this book, there are multiple points during the PCR set-up that can lead to a failed PCR experiment, and identifying these is not always straight forward. This book is your comprehensive guide to where your PCR has gone wrong, why and how to fix it. In real life it is not always possible to start out with the ‘perfect’ sample or design nucleic acid primers to the most permissive of genomic regions. But, whatever the experiment, this PCR trouble-shooting guide is here to aim you in the right direction and get the most out of your precious time in the lab.

Here’s to a happy, hassle-free PCR life!

Conclusion

About the author: Kirsten Hogg

Kirsten Hogg is a reproductive biologist with a special interest in epigenetic inheritance and fetal programming. Kirsten re-

ceived her PhD in Edinburgh, Scotland before moving to Canada to post-doc and is now based in Melbourne, Australia where she is researching germ cell development. Kirsten finds scientific writing refreshing and fun, loves developing skills in this area and has con-tributed a number of bite-sized articles to BiteSize Bio.

© 2014 Science Squared Ltd, UKImage source: Wordle image created by Kirsten Hogg via http://www.wordle.net.

http://bitesizebio.com/profile/kirsten-hogg/

http://www.wordle.net

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Contents · 2019. 12. 16. · Contents Chapter 1 Introduction 5 Main types of PCR 6 What this book...

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